Current Protocols in Protein Science Edited by: John E. Coligan (Bethesda, Maryland); Ben M. Dunn (University of Florida); David W. Speicher (The Wistar Institute); Paul T. Wingfield (Bethesda, Maryland); Past Editor: Hidde L. Ploegh Series Editor: Gwen Taylor
Copyright © 2007 by John Wiley and Sons, Inc. ISBN: 978-0-471-11184-9 0-471-11184-8 - Looseleaf 0-471-14098-8 - CD-ROM Last updated: 26 Apr 2007
PREFACE roteins are like people: they are highly diverse, with very few traits shared by the entire population. Even close relatives often have very different structural (physical) characteristics and behave quite differently. This heterogeneity presents challenges for scientists interested in studying protein structure and function. The need for a readily accessible, updatable reference for protein methods has grown in tandem with the ongoing protein science renaissance driven by advances in molecular biology, genetics, structural biology, and related disciplines. Using recombinant methods, it is now practical to overexpress both normal and reengineered forms of a protein and to create chimeric proteins with unique new properties. The ability to express proteins usually found in low abundance at high levels and to manipulate their amino acid sequences provides unprecedented opportunities to study a vast array of proteins and their associated biological processes (both in vitro and in vivo). The combination of biochemical, biophysical, and high-resolution structural analyses of proteins with manipulation of the associated genes (in cultured cells or an entire mammalian organism) has already led to spectacular advances in many areas of biotechnology and molecular medicine.
P
Because proteins are so diverse, a wide range of techniques must often be considered and tested empirically for a particular protein, and each researcher’s arsenal must include both recent innovations and techniques that have been used for many years. Moreover, methods appropriate for purifying and initially characterizing the protein(s) associated with a biological activity of interest are often not suitable for analyzing the corresponding overexpressed recombinant protein. This manual includes both specific detailed protocols as well as strategies for adapting methods to particular projects at hand. It is designed to meet the needs of scientists with little prior experience of protein isolation and characterization, including graduate students and scientists trained in other biological disciplines. At the same time, the broad range of techniques presented here should ensure that even seasoned experts will find new and useful approaches and will benefit from the manual’s convenient compilation of standard information and methods that typically must be culled from many sources. The quarterly updating service will allow ongoing expansion of the core volume to include chapters covering critical topics that were deferred due to initial space limitations and to keep pace with the “cutting edge” of developments in this fast-moving field.
HOW TO USE THIS MANUAL Format and Organization This publication is available in looseleaf, CD-ROM and online formats. For looseleaf subscribers, binders are provided to accommodate the growth of the manual via the quarterly updates. The looseleaf binder format allows easy insertion of new pages, units, and chapters that are added. The index and table of contents are updated with each supplement. CD-ROM subscribers receive a completely new disc every quarter and should dispose of their outdated discs. The online version is available through institutional license and offers many useful features such as downloadable PDF files of units, enhanced text searching for contents from all Current Protocols titles, real-time updating, and superior color graphics. The material covered in all three versions is identical. Subjects in this manual are organized by chapters and sections, and protocols are contained in units. Units generally describe a method and include one or more protocols with
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listings of materials, steps and annotations, recipes for unique reagents and solutions, and commentaries on the “hows” and “whys” of the method; there are also “overview” units containing theoretical discussions that lay the foundation for subsequent protocols. Page numbering in the looseleaf version reflects the modular arrangement by unit; for example, page 2.3.5 refers to Chapter 2 (Computational Analysis), UNIT 2.3 (Protein Secondary Structure Prediction), page 5 of that particular unit. Many reagents and procedures are employed repeatedly throughout the manual. Instead of duplicating this information, cross-references among units are used extensively. Crossreferencing helps to ensure that lengthy and complex protocols are not overburdened with steps describing auxiliary procedures needed to prepare raw materials and analyze results. Certain units that describe commonly used techniques and recipes (e.g., gel electrophoresis, ion-exchange chromatography) are cross-referenced in other units that describe their application. Thus, whenever it is necessary to isolate or identify a protein band in a protocol, the appropriate unit in Chapter 10—describing various procedures for gel electrophoresis—is cross-referenced (i.e., UNIT 10.1). For some widely used techniques (such as dialysis), readers are cross-referenced to APPENDIX 3.
Introductory and Explanatory Information Because this publication is first and foremost a compilation of laboratory techniques in protein science, we have not provided extensive instructive material. We have, however, included explanatory information where required to help readers gain an intuitive grasp of the procedures. Chapter 1 provides an overview of strategies for protein purification and characterization, to help the reader plan for all aspects of an experimental study. Some subsequent chapters begin with special overview units that describe the state of the art of the topic matter and provide a context for the procedures that follow. Chapter and unit introductions describe how the protocols that follow connect to one another, and annotations to the actual protocol steps describe what is happening as a procedure is carried out. Finally, the Commentary that closes each protocol unit describes background information regarding the historical and theoretical development of the method, as well as alternative approaches, critical parameters, troubleshooting guidelines, anticipated results, and time considerations. All units contain cited references and many indicate key references to inform users of particularly useful background reading, original descriptions, or applications of a technique. Protocols Many units in the manual contain groups of protocols, each presented with a series of steps. The basic protocol is presented first in each unit and is generally the recommended or most universally applicable approach. Alternate protocols are provided where different equipment or reagents can be employed to achieve similar ends, where the starting material requires a variation in approach, or where requirements for the end product differ from those in the basic protocol. Support protocols describe additional steps that are required to perform the basic or alternate protocols; these steps are separated from the core protocol because they might be applicable to other uses in the manual, or because they are performed in a time frame separate from the basic protocol steps.
Preface
Reagents and Solutions Reagents required for a protocol are itemized in the materials list before the procedure begins. Many are common stock solutions, others are commonly used buffers or media, while others are solutions unique to a particular protocol. Recipes for the latter solutions are provided in each unit, following the protocols (and before the commentary) under
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the heading Reagents and Solutions. It is important to note that the names of some of these special solutions might be similar from unit to unit (e.g., SDS sample buffer) while the recipes differ; thus, make certain that reagents are prepared from the proper recipes. On the other hand, recipes for commonly used stock solutions and buffers are provided once in APPENDIX 2E. These universal recipes are cross-referenced parenthetically in the materials lists rather than repeated with every usage.
Commercial Suppliers Throughout the manual, we have recommended commercial suppliers of chemicals, biological materials, and equipment. In some cases, the noted brand has been found to be of superior quality or it is the only suitable product available in the marketplace. In other cases, the experience of the author of that protocol is limited to that brand. In the latter situation, recommendations are offered as an aid to the novice protein science experimenter in obtaining the tools of the trade. Experienced investigators are therefore encouraged to experiment with substituting their own favorite brands. Addresses, phone numbers, and facsimile numbers of all suppliers mentioned in this manual are provided in the SUPPLIERS APPENDIX.
Safety Considerations Anyone carrying out these protocols may encounter the following hazardous or potentially hazardous materials: (1) radioactive substances, (2) toxic chemicals and carcinogenic or teratogenic reagents, and (3) pathogenic and infectious biological agents. Readers are referred to APPENDIX 2A and APPENDIX 2B for guidelines regarding use, storage, and disposal of these materials. Although cautionary statements are included in the appropriate units with cross-references to this appendix, we emphasize that users must proceed with the prudence and precaution associated with good laboratory practice, and that all materials must be used in strict accordance with local and national regulations. Reader Response Most of the protocols included in this manual are used routinely in our own laboratories. These protocols work for us; to make them work for you we have annotated critical steps and included critical parameters and troubleshooting guides in the commentaries to most units. However, the successful evolution of this manual depends upon readers’ observations and suggestions. Consequently, a self-mailing reader-response survey can be found at the back of the manual (and is included with each supplement); we encourage readers to send in their comments. Feedback can also be submitted online from the Current Protocols homepage at http://www.currentprotocols.com using the “Contact Us” link. ACKNOWLEDGMENTS This manual is the product of dedicated efforts by many of our scientific colleagues who are acknowledged in each unit and by the hard work by the Current Protocols editorial staff at John Wiley and Sons. The publisher’s commitment and continuing support for a protein science manual, particularly through the efforts of Brian Crawford and Paul Majchrzyk, was essential for realizing this ambitious project. We are extremely grateful for the critical contributions by Virginia Chanda (Series Editor) who kept the editors and the contributors on track and played a key role in bringing the entire project to completion. Other skilled members of the Current Protocols staff who contributed to the project include Janet Blair, Hazel Chan, Elizabeth Konkle, Alice Ro, and Kathy Wisch. The extensive copyediting required to produce an accurate protocols manual was ably handled by Rebecca Barr, Karen Hopkin, Arline Keithe, Kathy Morgan, Connie Parks, and Allen Ranz, and electronic illustrations were prepared by Gae Xavier. Finally, we
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would like to thank two people who played important roles in initiating this project. Sarah Greene initially conceived this project and had a critical role in developing it. John A. Smith made important contributions during the early planning phases of the project. Helpful advice and criticisms of the project during early planning stages were provided by Fred Ausubel, Charles S. Craik, Bruce A. Cummingham, Mark Hermodson, Tony E. Hugli, Robert Kingston, Albert Light, Paul Matsudaira, David Moore, George Rose, and Kevin Struhl. Mark Hermodson read and critiqued the entire manual and individual chapters were thoughtfully reviewed by John Freed and John Kay. Finally, we are especially grateful to our many colleagues, who contributed material to this manual, for sharing their procedures and experiences.
RECOMMENDED BACKGROUND READING Branden, C. and Tooze, J. 1999. Introduction to Protein Structure, 2nd edition. Garland Publishing, New York. Easy-to-read overview of basic structural principles of proteins with extensive illustrations. Fersht, A. 1998. Structure and Mechanism in Protein Science: A Guide to Enzyme Catalysis and Protein Folding. 3rd edition. W. H. Freeman. Well established and clear review of protein structure and function, with an emphasis on enzymes. Hunte, C., von Jagow, G. and Schagger, H., editors. 2003. Membrane Protein Purification and Crystallization: A Practical Guide (2nd edition). Academic Press. Deals with specific problems encountered when purifying membrane-associated proteins. Link, A.J. (editor). 1998. 2-D Proteome Analysis Protocols. Humana Press. Extensive treatment of 2-D gel analysis and its role in proteomics, including preparations of samples from diverse sources, alternative 2-D gel analysis methods and data analysis/interpretation. Petsko, G.A. and Ringe, D. 2004. Protein Structure and Function. New Science Press Ltd., London. An updated look at protein structure in the genomics age. Includes discussion of sequence-to-structure and structure-to-function correlations. Speicher, D.W. (editor). 2004. Proteome analysis: Interpreting the Genome. Elsevier. Overview of most current proteomics methods and supporting technologies. Voet, D. and Voet, J.G. 2005. Biochemistry, 3rd edition. John Wiley & Sons. An updated biochemistry textbook whose protein-related chapters provide a comprehensive background in biochemistry.
John E. Coligan, Ben M. Dunn, Hidde L. Ploegh, David W. Speicher, and Paul T. Wingfield
Preface
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FOREWORD
O
ver the past five to ten years we have witnessed a revolution in biological research which is fueled by our increasing capacity to decipher biological information of three types: the digital information embedded in DNA with its four-letter alphabet; the three-dimensional information represented by proteins, the major executors of biological function; and the four-dimensional information of complex biological systems and networks representing the temporal and spatial interaction of multiple components. The analysis of complex systems and networks in their entirety is essential for scientists to understand the molecular basis of fascinating processes such as growth, development, and differentiation, and to extract the biological meaning of emergent properties such as consciousness, memory, and the ability to learn which evolved over billions of years. Not surprisingly, the linear nucleotide sequence in DNA has been the first type of information to be deciphered. Large-scale genetic mapping and DNA sequencing rapidly generate enormous amounts of biological information. In many ways, programs such as the human genome project represent some of the earliest attempts to understand biological complexity. Successful completion of the human genome and similar projects will pose the challenge of interpreting the information contained in billions of nucleotides and of explaining how the interplay of the products of perhaps 100,000 genes results in the myriad of biological phenotypes. Meeting these challenges will require new interdisciplinary strategies which draw from the expertise of scientists from disciplines as different as molecular biology, biochemistry, engineering, chemistry, applied mathematics, and computer science. Protein science is central to such integrated strategies. Separation of proteins from complex mixtures, followed by structural and functional analyses, control of protein function, post-translational processing, and modification and formation of macromolecular complexes of proteins and other biomolecules are but a few examples of topics which are essential for all those scientists who attempt to interpret the linear DNA sequence in terms of the three- and four-dimensional information of complex biological systems and networks. The following chapters detail the necessary protocols for this endeavor, provided by experts in experimental protein science. Two attractive features set this manual apart from other collections of research protocols. First, Current Protocols in Protein Science will be continuously updated and expanded by quarterly additions to the core edition. Secondly, this volume as well as the updates are also available on CD-ROM. Numerous cross-references within the manual by hypertext links, context based searching, and provisions for making individualized notebooks containing frequently used protocols are attractions of the CD-ROM version which are in tune with the increasing dependence on computers in biological research laboratories.
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Protein science is a diverse, experimentally challenging, and rapidly evolving discipline. This book provides expert guidance in experiment design and execution and promises to remain up-to-date, in contents and in format, for many years to come. Leroy E. Hood1 and Ruedi Aebersold1 1 Seattle, Washington
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Table of Contents
Preface Foreword
Chapter 1 Strategies of Protein Purification and Characterization Unit 1.1 Overview of Protein Purification and Characterization Unit 1.2 Strategies for Protein Purification Unit 1.3 Protein Purification Flow Charts Unit 1.4 Purification of Glutamate Dehydrogenase from Liver and Brain Unit 1.5 Overview of the Physical State of Proteins within Cells
Chapter 2 Computational Analysis Introduction Unit 2.1 Computational Methods for Protein Sequence Analysis Unit 2.2 Hydrophobicity Profiles for Protein Sequence Analysis Unit 2.3 Protein Secondary Structure Prediction Unit 2.4 Internet Basics Unit 2.5 Sequence Similarity Searching Using the BLAST Family of Programs Unit 2.6 Protein Databases on the Internet Unit 2.7 Protein Tertiary Structure Prediction Unit 2.8 Protein Tertiary Structure Modeling Unit 2.9 Comparative Protein Structure Prediction Unit 2.10 Protein Charge Determination Unit 2.11 Basic Protein Sequence Analysis
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Unit 2.12 Minimotif Miner: A Computational Tool to Investigate Protein Function, Disease, and Genetic Diversity
Chapter 3 Detection and Assay Methods Introduction Unit 3.1 Spectrophotometric Determination of Protein Concentration Unit 3.2 Quantitative Amino Acid Analysis Unit 3.3 In Vitro Radiolabeling of Peptides and Proteins Unit 3.4 Assays for Determination of Protein Concentration Unit 3.5 Kinetic Assay Methods Unit 3.6 Biotinylation of Proteins in Solution and on Cell Surfaces Unit 3.7 Metabolic Labeling with Amino Acids Unit 3.8 Analysis of Selenocysteine-Containing Proteins Unit 3.9 Solid-Phase Profiling of Proteins
Chapter 4 Extraction, Stabilization, and Concentration Introduction Unit 4.1 Overview of Cell Fractionation Unit 4.2 Purification of Organelles from Mammalian Cells Unit 4.3 Subcellular Fractionation of Tissue Culture Cells Unit 4.4 Desalting, Concentration, and Buffer Exchange by Dialysis and Ultrafiltration Unit 4.5 Selective Precipitation of Proteins Unit 4.6 Long-Term Storage of Proteins Unit 4.7 Extraction of Proteins from Plant Tissues
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Chapter 5 Production of Recombinant Proteins Introduction Unit 5.1 Production of Recombinant Proteins in Escherichia coli Unit 5.2 Selection of Escherichia coli Expression Systems Unit 5.3 Fermentation and Growth of Escherichia coli for Optimal Protein Production Unit 5.4 Overview of the Baculovirus Expression System Unit 5.5 Protein Expression in the Baculovirus System Unit 5.6 Overview of Protein Expression in Saccharomyces cerevisiae Unit 5.7 Overview of Protein Expression in Pichia pastoris Unit 5.8 Culture of Yeast for the Production of Heterologous Proteins Unit 5.9 Overview of Protein Expression by Mammalian Cells Unit 5.10 Production of Recombinant Proteins in Mammalian Cells Unit 5.11 Overview of the Vaccinia Virus Expression System Unit 5.12 Preparation of Cell Cultures and Vaccinia Virus Stocks Unit 5.13 Generation of Recombinant Vaccinia Viruses Unit 5.14 Characterization of Recombinant Vaccinia Viruses and Their Products Unit 5.15 Gene Expression Using the Vaccinia Virus/ T7 RNA Polymerase Hybrid System Unit 5.16 Choice of Cellular Protein Expression System Unit 5.17 Use of the Gateway System for Protein Expression in Multiple Hosts Unit 5.18 Wheat Germ Cell-Free Expression System for Protein Production
Chapter 6 Purification of Recombinant Proteins Introduction Unit 6.1 Overview of the Purification of Recombinant Proteins Produced in Escherichia coli Unit 6.2 Preparation of Soluble Proteins from Escherichia coli
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Unit 6.3 Preparation and Extraction of Insoluble (Inclusion-Body) Proteins from Escherichia coli Unit 6.4 Overview of Protein Folding Unit 6.5 Folding and Purification of Insoluble (Inclusion Body) Proteins from Escherichia coli Unit 6.6 Expression and Purification of GST Fusion Proteins Unit 6.7 Expression and Purification of Thioredoxin Fusion Proteins Unit 6.8 Automated Large-Scale Purification of a Recombinant G-Protein-Coupled Neurotensin Receptor
Chapter 7 Characterization of Recombinant Proteins Introduction Unit 7.1 Overview of the Characterization of Recombinant Proteins Unit 7.2 Determining the Identity and Purity of Recombinant Proteins by UV Absorption Spectroscopy Unit 7.3 Determining the Identity and Structure of Recombinant Proteins Unit 7.4 Transverse Urea-Gradient Gel Electrophoresis Unit 7.5 Analytical Ultracentrifugation Unit 7.6 Determining the CD Spectrum of a Protein Unit 7.7 Determining the Fluorescence Spectrum of a Protein Unit 7.8 Light Scattering Unit 7.9 Measuring Protein Thermostability by Differential Scanning Calorimetry Unit 7.10 Characterizing Recombinant Proteins Using HPLC Gel Filtration and Mass Spectrometry Unit 7.11 Rapid Screening of E. coli Extracts by Heteronuclear NMR
Chapter 8 Conventional Chromatographic Separations Introduction Unit 8.1 Overview of Conventional Chromatography Unit 8.2 Ion-Exchange Chromatography
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Unit 8.3 Gel-Filtration Chromatography Unit 8.4 Hydrophobic-Interaction Chromatography Unit 8.5 Chromatofocusing Unit 8.6 Hydroxylapatite Chromatography Unit 8.7 HPLC of Peptides and Proteins Unit 8.8 Expanded-Bed Adsorption Chromatography
Chapter 9 Affinity Purification Introduction Unit 9.1 Lectin Affinity Chromatography Unit 9.2 Dye Affinity Chromatography Unit 9.3 Affinity Purification of Natural Ligands Unit 9.4 Metal-Chelate Affinity Chromatography Unit 9.5 Immunoaffinity Chromatography Unit 9.6 Purification of Sequence-Specific DNA-Binding Proteins by Affinity Chromatography Unit 9.7 Purification of DNA-Binding Proteins Using Biotin/Streptavidin Affinity Systems Unit 9.8 Immunoprecipitation Unit 9.9 Overview of Affinity Tags for Protein Purification
Chapter 10 Electrophoresis Introduction Unit 10.1 One-Dimensional SDS Gel Electrophoresis of Proteins Unit 10.2 One-Dimensional Isoelectric Focusing of Proteins in Slab Gels Unit 10.3 One-Dimensional Electrophoresis Using Nondenaturing Conditions Unit 10.4 Two-Dimensional Gel Electrophoresis
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Unit 10.5 Protein Detection in Gels Using Fixation Unit 10.6 Protein Detection in Gels Without Fixation Unit 10.7 Electroblotting from Polyacrylamide Gels Unit 10.8 Detection of Proteins on Blot Membranes Unit 10.9 Capillary Electrophoresis of Proteins and Peptides Unit 10.10 Immunoblot Detection Unit 10.11 Autoradiography Unit 10.12 Overview of Digital Electrophoresis Analysis Unit 10.13 Capillary Electrophoresis of Peptides and Proteins Using Isoelectric Buffers
Chapter 11 Chemical Analysis Introduction Unit 11.1 Enzymatic Digestion of Proteins in Solution Unit 11.2 Enzymatic Digestion of Proteins on PVDF Membranes Unit 11.3 Enzymatic Digestion of Proteins in Gels for Mass Spectrometric Identification and Structural Analysis Unit 11.4 Chemical Cleavage of Proteins in Solution Unit 11.5 Chemical Cleavage of Proteins on Membranes Unit 11.6 Reversed-Phase Isolation of Peptides Unit 11.7 Removal of N-Terminal Blocking Groups from Proteins Unit 11.8 C-Terminal Sequence Analysis Unit 11.9 Amino Acid Analysis Unit 11.10 N-Terminal Sequence Analysis of Proteins and Peptides Unit 11.11 Determination of Disulfide-Bond Linkages in Proteins
Chapter 12 Post-Translational Modification: Glycosylation Introduction
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Unit 12.1 Overview of Glycoconjugate Analysis Unit 12.2 Metabolic Radiolabeling of Animal Cell Glycoconjugates Unit 12.3 Inhibition of N-Linked Glycosylation Unit 12.4 Endoglycosidase and Glycoamidase Release of N-Linked Oligosaccharides Unit 12.5 Detection of Glycophospholipid Anchors on Proteins Unit 12.6 Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins Unit 12.7 Determining the Structure of Glycan Moieties by Mass Spectrometry Unit 12.8 Detection and Analysis of Proteins Modified by O-Linked N-Acetylglucosamine
Chapter 13 Post-Translational Modification: Phosphorylation and Phosphatases Introduction Unit 13.1 Overview of Protein Phosphorylation Unit 13.2 Labeling Cultured Cells with 32Pi and Preparing Cell Lysates for Immunoprecipitation Unit 13.3 Phosphoamino Acid Analysis Unit 13.4 Detection of Phosphorylation by Immunological Techniques Unit 13.5 Detection of Phosphorylation by Enzymatic Techniques Unit 13.6 Preparation and Application of Polyclonal and Monoclonal Sequence-Specific AntiPhosphoamino Acid Antibodies Unit 13.7 Assays of Protein Kinases Using Exogenous Substrates Unit 13.8 Permeabilization Strategies to Study Protein Phosphorylation Unit 13.9 Phosphopeptide Mapping and Identification of Phosphorylation Sites Unit 13.10 Use of Protein Phosphatase Inhibitors
Chapter 14 Post-Translational Modification: Specialized Applications Introduction Unit 14.1 Analysis of Disulfide Bond Formation
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Unit 14.2 Analysis of Protein Acylation Unit 14.3 Analysis of Protein Prenylation and Carboxyl-Methylation Unit 14.4 Analysis of Oxidative Modification of Proteins Unit 14.5 Analysis of Protein Ubiquitination Unit 14.6 Analysis of Protein S-Nitrosylation Unit 14.7 Tyrosine O-Sulfation Unit 14.8 Analysis of Protein Sumoylation Unit 14.9 Identifying and Quantifying Sites of Protein Methylation by Heavy Methyl SILAC
Chapter 15 Chemical Modification of Proteins Introduction Unit 15.1 Modification of Cysteine Unit 15.2 Modification of Amino Groups
Chapter 16 Mass Spectrometry Introduction Unit 16.1 Overview of Peptide and Protein Analysis by Mass Spectrometry Unit 16.2 Matrix-Assisted Laser Desorption/Ionization Time-of-Flight Mass Analysis of Peptides Unit 16.3 Sample Preparation for MALDI Mass Analysis of Peptides and Proteins Unit 16.4 In-Gel Digestion of Proteins for MALDI-MS Fingerprint Mapping Unit 16.5 Searching Sequence Databases Over the Internet: Protein Identification Using MS-Fit Unit 16.6 Searching Sequence Databases Over the Internet: Protein Identification Using MS-Tag Unit 16.7 Enzymatic Approaches for Obtaining Amino Acid Sequence: On-Target Ladder Sequencing Unit 16.8 Introducing Samples Directly into Electrospray Ionization Mass Spectrometers Using a Nanospray Interface Unit 16.9 Introducing Samples Directly into Electrospray Ionization Mass Spectrometers Using Microscale Capillary Liquid Chromatography
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Unit 16.10 Protein Identification Using a Quadrupole Ion Trap Mass Spectrometer and SEQUEST Database Matching Unit 16.11 De Novo Peptide Sequencing via Manual Interpretation of MS/MS Spectra
Chapter 17 Structural Biology Introduction Unit 17.1 Overview of Protein Structural and Functional Folds Unit 17.2 Electron Microscopy and Image Processing: An Essential Tool for Structural Analysis of Macromolecules Unit 17.3 Principles of Macromolecular X-Ray Crystallography Unit 17.4 Crystallization of Macromolecules Unit 17.5 Introduction to NMR of Proteins Unit 17.6 Probing Protein Structure and Dynamics by Hydrogen Exchange–Mass Spectrometry Unit 17.7 Introduction to Atomic Force Microscopy (AFM) in Biology Unit 17.8 Raman Spectroscopy of Proteins Unit 17.9 Crystallization of Integral Membrane Proteins
Chapter 18 Preparation and Handling of Peptides Introduction Unit 18.1 Introduction to Peptide Synthesis Unit 18.2 Synthesis of Multiple Peptides on Plastic Pins Unit 18.3 Synthetic Peptides for Production of Antibodies that Recognize Intact Proteins Unit 18.4 Native Chemical Ligation of Polypeptides Unit 18.5 Synthesis and Application of Peptide Dendrimers As Protein Mimetics Unit 18.6 Disulfide Bond Formation in Peptides Unit 18.7 Guide for Resin and Linker Selection in Solid-Phase Peptide Synthesis
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Chapter 19 Identification of Protein Interactions Introduction Unit 19.1 Analysis of Protein-Protein Interactions Unit 19.2 Interaction Trap/Two-Hybrid System to Identify Interacting Proteins Unit 19.3 Phage-Based Expression Cloning to Identify Interacting Proteins Unit 19.4 Detection of Protein-Protein Interactions by Coprecipitation Unit 19.5 Imaging Protein-Protein Interactions by Fluorescence Resonance Energy Transfer (FRET) Microscopy Unit 19.6 High-Throughput Screening for Protein-Protein Interactions Using Yeast Two-Hybrid Arrays Unit 19.7 Identification of Protein Interactions by Far Western Analysis Unit 19.8 Scintillation Proximity Assay (SPA) Technology to Study Biomolecular Interactions Unit 19.9 Identifying Protein Interactions by Hydroxyl-Radical Protein Footprinting Unit 19.10 Visualization of Protein Interactions in Living Cells Using Bimolecular Fluorescence Complementation (BiFC) Analysis Unit 19.11 Production and Use of Trimeric Isoleucine Zipper Fusion Proteins to Study Surface Receptor Ligand Interactions Unit 19.12 Fluorescence Quenching Methods to Study Lipid-Protein Interactions Unit 19.13 Overview of Biacore Systems and Their Applications Unit 19.14 Using Biacore to Measure the Binding Kinetics of an Antibody-Antigen Interaction Unit 19.15 Identifying Small-Molecule Modulators of Protein-Protein Interactions Unit 19.16 Determination of Protein Contacts by Chemical Cross-Linking With EDC and Mass Spectrometry
Chapter 20 Quantitation of Protein Interactions Introduction Unit 20.1 Overview of the Quantitation of Protein Interactions Unit 20.2 Measuring Protein Interactions by Optical Biosensors Unit 20.3 Analytical Centrifugation: Equilibrium Approach
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Unit 20.4 Titration Microcalorimetry Unit 20.5 Reduced-Scale Large-Zone Analytical Gel-Filtration Chromatography for Measurement of Protein Association Equilibria Unit 20.6 Size-Exclusion Chromatography with On-Line Light Scattering Unit 20.7 Analytical Ultracentrifugation: Sedimentation Velocity Analysis Unit 20.8 Spectroscopic Methods for the Determination of Protein Interactions Unit 20.9 Application of Amide Proton Exchange Mass Spectrometry for the Study of Protein-Protein Interactions Unit 20.10 Circular Dichroism to Study Protein Interactions Unit 20.11 Quantitative Determination of Protein Stability and Ligand Binding by Pulse Proteolysis
Chapter 21 Peptidases Introduction Unit 21.1 Proteases Unit 21.2 Papain-like Cysteine Proteases Unit 21.3 Overview of Pepsin-like Aspartic Peptidases Unit 21.4 Metalloproteases Unit 21.5 Purification and Characterization of Proteasomes from Saccharomyces cerevisiae Unit 21.6 Purification of the Eukaryotic 20S Proteasome Unit 21.7 Serpins (Serine Protease Inhibitors) Unit 21.8 Caspases Unit 21.9 Use of GFP as a Reporter for the Analysis of Sequence-Specific Proteases Unit 21.10 An Overview of Serine Proteases Unit 21.11 Over-Expression and Purification of Active Serine Proteases and Their Variants from Escherichia coli Inclusion Bodies Unit 21.12 Assaying Proteases in Cellular Environments Unit 21.13 Expression, Purification, and Characterization of Caspases
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Unit 21.14 Expression, Purification, and Characterization of Aspartic Endopeptidases: Plasmodium Plasmepsins and “Short” Recombinant Human Pseudocathepsin Unit 21.15 Zymography of Metalloproteinases Unit 21.16 Monitoring Metalloproteinase Activity Using Synthetic Fluorogenic Substrates Unit 21.17 Applications for Chemical Probes of Proteolytic Activity
Chapter 22 Gel-Based Proteome Analysis Introduction Unit 22.1 Overview of Proteome Analysis Unit 22.2 Protein Profiling Using Two-Dimensional Difference Gel Electrophoresis (2-D DIGE) Unit 22.3 Laser Capture Microdissection for Proteome Analysis Unit 22.4 Preparing Protein Extracts for Quantitative Two-Dimensional Gel Comparison Unit 22.5 Isolation of Organelles and Prefractionation of Protein Extracts Using Free-Flow Electrophoresis Unit 22.6 Protein Profiling by Microscale Solution Isoelectrofocusing (MicroSol-IEF)
Chapter 23 Non-Gel-Based Proteome Analysis Introduction Unit 23.1 Analysis of Protein Composition Using Multidimensional Chromatography and Mass Spectrometry Unit 23.2 Quantitative Protein Profile Comparisons Using the Isotope-Coded Affinity Tag Method Unit 23.3 Proteomic Analysis Using 2-D Liquid Separations of Intact Proteins From Whole-Cell Lysates Unit 23.4 Quantitative Protein Analysis Using Proteolytic [18O]Water Labeling Unit 23.5 Protein Microarrays
Chapter 24 Targeted Proteomics Introduction Unit 24.1 Human Serum and Plasma Proteomics
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Unit 24.2 Organelle Proteomics Unit 24.3 Glycoproteomics Using Chemical Immobilization
Chapter 25 Proteome Bioinformatics Introduction Unit 25.1 Bioinformatics Analysis for Interactive Proteomics
Chapter 26 Protein Engineering Introduction Unit 26.1 Misincorporation Proton-Alkyl Exchange (MPAX): Engineering Cysteine Probes into Proteins Unit 26.2 Combinatorial Recombination of Gene Fragments to Construct a Library of Chimeras Unit 26.3 Incorporation of Isotopically Enriched Amino Acids
Appendix 1 Useful Data 1A Characteristics of Amino Acids 1B Commonly Used Detergents 1C Conversion Factors and Half-Life Information for Radioactivity 1D Common Conversion Factors
Appendix 2 Laboratory Guidelines, Equipment, and Stock Solutions 2A Laboratory Safety 2B Safe Use of Radioisotopes 2C Centrifuges and Rotors 2D Standard Laboratory Equipment 2E Commonly Used Reagents
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Appendix 3 Commonly Used Techniques 3A Use of Protein Folding Reagents 3B Dialysis 3C Techniques for Mammalian Cell Tissue Culture 3D Importing Biological Materials 3E Silanizing Glassware 3F Protein Precipitation Using Ammonium Sulfate 3G Statistics: Detecting Differences Among Groups 3H Analyzing Radioligand Binding Data
Appendix 4 Molecular Biology Techniques 4 Molecular Biology Techniques 4A Media Preparation and Bacteriological Tools 4B Growth in Liquid or Solid Media 4C Preparation of Plasmid DNA 4D Introduction of Plasmid DNA into Cells 4E Purification and Concentration of DNA from Aqueous Solutions 4F Agarose Gel Electrophoresis 4G Southern Blotting 4H Hybridization Analysis of DNA Blots 4I Digestion of DNA with Restriction Endonucleases 4J The Polymerase Chain Reaction 4K Quantitation of Nucleic Acids with Absorption Spectroscopy 4L Growth and Manipulation of Yeast
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Appendix 5 Biophysical Methods: Data Analysis 5A Theoretical Aspects of the Quantitative Characterization of Ligand Binding Suppliers Selected Suppliers of Reagents and Equipment
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Overview of Protein Purification and Characterization AIMS AND OBJECTIVES Protein purification has an over 200-year history: the first attempts at isolating substances from plants having similar properties to “egg albumen,” or egg white, were reported in 1789 by Fourcroy. Many proteins from plants were purified in the nineteenth century, though most would not be considered pure by modern standards. A century later, ovalbumin was the first crystalline protein obtained (by Hofmeister in 1889). The year 1989 may not go down in history as a milestone in protein chemistry, but since then there has been a resurgence of interest in proteins after more than a decade of gene excitement. The aims of protein purification, up until the 1940s, were simply academic. To then, even the basic facts of protein structure were not fully appreciated, and pure proteins were needed just to study structure and test the rival theories of the pre-DNA days. During the Second World War, an acute need for blood proteins led to development of the Cohn fractionation procedure for purification of albumin and other proteins from serum (Cohn et al., 1946). This was the inception of large-scale protein purifications for commercial purposes; Cohn fractionation continues to be used to this day. As more proteins, and particularly enzymes, were purified and crystallized, they started to be used increasingly in diagnostic assays and enzymatic analyses, as well as in the largescale food, tanning, and detergent industries. Many enzymes used in industry are not in fact very pure, but as long as they do the job, that is sufficient. “Process” enzymes such as α-amylase, proteases, and lipases are pro-duced in ton quantities, mainly as secretion products in bacterial cultures, and may undergo only limited purification processes to mini-mize costs. At the other extreme, enzyme products for research and analysis require a high degree of purification to ensure that contaminating activities do not interfere with the intended use. Anyone familiar with molecular biology enzymes will appreciate how minute levels of contamination of DNase or RNase can completely destroy carefully planned experiments. The 1960s and 1970s could be described as the peak years for protein and enzyme research, and most of the methods used in protein puri-
Contributed by R.K. Scopes Current Protocols in Protein Science (1995) 1.1.1-1.1.6 Copyright © 2000 by John Wiley & Sons, Inc.
fication were established by then, at least in their principles. More recent developments have been mainly in instrumentation designed to optimize the application of each methodology. Developments in instrumentation have been stimulated by the rapid progress in molecular biology, because gene isolation has often been preceded by isolation of the gene product. Because such products can now be characterized sufficiently (i.e., partially sequenced) using minute amounts of protein, the need for large-scale or even moderate-scale procedures has decreased. Hence there has been an explosive development of modern equipment designed specifically for dealing with amounts of protein in the milligram to microgram range. On the other hand, structural studies using X-ray crystallography and nuclear magnetic resonance (NMR) require hundreds of milligrams of pure protein, so larger-scale equipment and procedures are still needed in the research laboratory. The nature of the proteins studied has also changed substantially. Whereas enzymes were once the most favored subjects, they have now been superceded by nonenzymatic proteins such as growth factors, hormone receptors, viral antigens, and membrane transporters. Many of these occur in minute amounts in the natural source, and their purification can be a major task. Heroic efforts in the past have used kilogram quantities of rather unpleasant starting materials, such as human organs, and ended up with a few micrograms of pure product. It is now more usual, however, to take the genetic approach: clone the gene before the protein has been isolated or even properly identified, then express it in a suitable host cell culture or organism. The expression level may be orders of magnitude higher than in the original source, which will make purification a relatively simple task. It can be useful to know beforehand some physical properties of the protein, to facilitate the development of a suitable purification protocol from the recombinant source. On the other hand, there are now several ways of preparing fusion proteins, which can be purified by affinity techniques without any knowledge of the properties of the target protein. Moreover, there are ways of modifying the expressed product to simplify purification further.
UNIT 1.1
Strategies of Protein Purification and Characterization
1.1.1 CPPS
Overview of Protein Purification and Characterization
Thus the approach to protein purification must first take into account the reason it is being done, as the methods will vary greatly with different requirements. At one extreme is the one-of-a-kind purification, in a well-financed and equipped laboratory, that is carried out to obtain a small amount of product for sequencing so that gene isolation can proceed. In this case, expense of equipment and reagents may be no problem, and a very low overall recovery of product can be acceptable, provided it is pure enough. At the other extreme are the requirements of commercial production of a protein in large amounts on a continuing basis, where high recovery and economy of processing are the chief parameters to be considered. There are many intermediate situations as well. Many publications in the area of protein research are entitled “Purification and characterization of…,” and describe a purification procedure in sufficient detail that it can be reproduced in another laboratory. The characterization section may include structural, functional, and genetic information, and carrying out such studies is likely to require at least milligram quantities of pure protein. Ideally the purification should involve a small number of steps, with good recovery at each step. If the recovery is poor (<50% at any step), however, there should be some indication of what happened to the missing activity. Has it been discarded in the other fractions for the sake of purity, or does it represent a true loss of activity? If the latter, then the end-product may be less than fully active despite apparent homogeneity indicated by standard analysis. The choice between recovery and purification at each step can be problematical; taking a narrow cut of a chromatographic peak may provide a very pure fraction, at the expense of losing a good deal of less pure active component on either side. In making such decisions, the objective of the exercise must be kept in mind: if yield is not important, then the choice of poor yield for the sake of purity may be logical. By far the most important requirement of a publication is reproducibility of the method reported. It is not sufficient to have carried out the process only once if it is expected that other investigators will want to repeat it. There are always factors that influence the process that may be overlooked at first, and which if varied slightly can have a major effect on the purification procedure. The reported process should always be repeated exactly as described before submitting the manuscript for publication. There is one exception, namely, the case where
purification was conducted simply to obtain enough protein for sequencing and gene isolation; if that was achieved, there should be no need to provide instructions for repetition.
SOURCES OF MATERIAL FOR PROTEIN PURIFICATION For many people embarking on a protein purification project, there is no choice of material. They are studying a particular biological tissue or organism, and the objective is to purify a protein from that source. However, there may be approaches that can make the project simpler. If, for instance, the source is difficult to obtain in large amounts, it may be best to carry out at least preliminary trials on a source species more readily obtained. The most obvious and relevant example is when the species being studied is Homo sapiens, and tissue samples are not readily available for practical or ethical reasons, or both. In this case, it is usual to go to where mammalian tissue is readily available (i.e., an abattoir) and work with bovine, ovine, or porcine sources. Alternatively, if quantity of tissue is not a problem, the humble laboratory rat may suffice. Once a protocol for purifying the protein from substitute sources has been worked out, it will be much easier to develop one using human material—the identical procedure may work satisfactorily. Proteins differ to a fairly small extent between species that have diverged within about 100 million years, a time frame that groups together most higher mammals. Thus the behavior of proteins derived from different animals with respect to the various fractionation procedures is likely to be similar, and a protocol worked out for pig tissues is likely to need only minor adjustments for application to human tissues. A second example is where the interest is mainly on the function of a protein, especially an enzyme, for which functions and actions have generally been strongly conserved through evolution. In that case, a preliminary screening of potential sources, or, better still, the literature, should come up with a raw material that is best suited to the investigator’s purposes. Considerations should include the following: (1) What functions are required of the end product? For instance, an enzyme having a low Km may be needed, so selecting the source with the highest activity may not suffice. (2) How convenient is it to grow or obtain the raw material, and are there problems concerning pathogenicity or extractability? (3) Does the quantity of the protein vary with growth conditions or age, and does it deteriorate in situ
1.1.2 Current Protocols in Protein Science
if left too long? Obviously one requires a source that reliably produces the highest amount of the desired protein per unit volume to maximize the chances of developing a good purification procedure. (4) What storage conditions are required for the raw material? It is important to consider that fresh raw material may not be immediately available whenever a purification is attempted. The above considerations are relevant to the traditional situation for commencing a protein purification project. It is becoming increasingly common, however, for proteins to be purified as recombinant products using techniques in which the gene is expressed in a host organism or in cultured cells. This of course requires the gene encoding the protein of interest to be available. Until the mid-1980s, such material was usually obtained by hybridization of an oligonucleotide synthesized according to amino acid sequence information. This required the protein to have been purified first, so the initial task of protein purification still needed to be done at least once. More recently, genetic techniques have permitted the isolation of many genes encoding known proteins, even though the proteins may never have been studied directly. Moreover, with the expansion of the Human Genome Project and related DNA sequencing efforts, many genes for both known and unknown proteins will become available and will be able to be expressed in recombinant form without ever being purified from the host species. As a result some completely new considerations for protein purification come into play, including the possibility of modifying the gene structure not only to increase expression level and alter the protein product itself to enhance a desired function, but equally importantly to aid in purification. Recombinant proteins may be expressed in bacteria, yeasts, insect cells, and animal tissue cultures. Further details may be found in UNIT 1.2.
DETECTION AND ASSAY OF PROTEINS During a protein purification procedure there are two measurements that need to be made, preferably for each fraction. Measurements both of the total protein and of the amount (usually bioactivity) of the desired protein must be made. Details of the most commonly used assay methods are given in Chapter 3. It is not possible to isolate a protein without a method of determining whether it is present; an assay, either quantitative or at least
semiquantitative, indicating which fraction contains the most of the desired protein is essential. Assays may range from the quick-and-easy type (e.g., instantaneous spectrophotometric measurement of enzyme activity) to long and tedious bioassays that may take days to produce an answer. The latter situation is very difficult, because by the time one knows where the protein is, it may be “was,” owing to degradation or inactivation. Moreover, this may not become clear until the next step has been completed and its products assayed. Any assay that is quick is therefore advantageous, even if it means a sacrifice of accuracy for speed. Measurement of total protein is useful, as it indicates the degree of purification at each step. However, unless the next step critically depends on how much protein is present, total protein measurement is not extremely important: a small sample can be put aside and measured later, when the purification is complete. It is, however, very important to know how much protein is present in the final, presumed pure sample, as this will indicate the specific activity (if the protein has an activity), which can be compared with other preparations. The general object is to obtain as high a specific activity as possible (taking into account recovery considerations), which means retaining as much of the desired protein as possible while ending up with as little total protein as possible.
METHODS FOR SEPARATION AND PURIFICATION OF PROTEINS The methods available for protein purification range from simple precipitation procedures used since the nineteenth century to sophisticated chromatographic and affinity techniques that are constantly undergoing development and improvement. Methods can be classified in several alternative ways—perhaps one of the best is based on the properties of the proteins that are being exploited. Thus the methods can be divided into four distinct but interrelated groups depending on protein characteristics: surface features, size and shape, net charge, and bioproperties.
Methods Based on Surface Features of Proteins Surface features include charge distribution and accessibility, surface distribution of hydrophobic amino acid side chains, and, to a lesser extent, net charge at a given pH (see discussion of net charge). Methods exploiting surface fea-
Strategies of Protein Purification and Characterization
1.1.3 Current Protocols in Protein Science
tures mainly depend on solubility properties. Differences in solubility result in precipitation by various manipulations of the solvent in which the proteins are solubilized. Methods for obtaining an extract containing the desired protein in soluble form are given in Chapter 4. The solvent, nearly always water containing a low concentration of buffer salts, can be treated to alter properties such as ionic strength, dielectric constant, pH, temperature, and detergent content, any of which may selectively precipitate some of the proteins present. Conversely, proteins may be selectively solubilized from an insoluble state by manipulation of the solvent composition. The surface distribution of hydrophobic residues is an important determinant of solubility properties; it is also exploited in hydrophobic chromatography, both in the reversed-phase mode (UNIT 11.6) and in aqueousphase hydrophobic-interaction chromatography (UNIT 8.4). Also included in this group is the highly specific technique of immunoaffinity chromatography, in which an antibody directed against an epitope on the protein surface is used to pull out the desired protein from a mixture.
Methods Based on Whole Structure: Size and Shape Although the size and shape of proteins can have some influence on solubility properties, the chief method of exploiting these properties is gel-filtration chromatography (UNIT 8.3). In addition, preparative gel electrophoresis makes use of differences in molecular size. Proteins range in size from the smallest classified as proteins rather than polypeptides, around 5000 Da, up to macromolecular complexes of many million daltons. Many proteins in the bioactive state are oligomers of more than one polypeptide (see UNIT 1.2), and these can be dissociated, though normally with loss of overall structure. Thus many proteins have two “sizes”: that of the native state, and that (or those) of the polypeptides in the denatured and dissociated state. Gel-filtration procedures normally deal only with native proteins, whereas electrophoretic procedures commonly involve separation of dissociated and denatured polypeptides.
Methods Based on Net Charge
Overview of Protein Purification and Characterization
The two techniques that exploit the overall charge of proteins are ion-exchange chromatography (by far the most important) and electrophoresis (Chapter 10). Ion exchangers bind charged molecules, and there are essentially only two types of ion exchangers, anion and
cation. The net charge of a protein depends on the pH—positive at very low pH, negative at high pH, and zero at some specific point in between, termed the isoelectric point (pI). It should be stressed that at the pI a protein has a great many charges; it just happens that at this pH the total negatives exactly equal the total positives. The most charged state (disregarding the charge sign) is in the pH range 6.0 to 9.0. This is the most stable pH range for most proteins, as it encompasses common physiological pH values. Ion exchangers consist of immobilized charged groups and attract oppositely charged proteins. They provide the mode of separation that has the highest resolution for native proteins. High-performance reversedphase chromatography has equivalent or even better resolution, but it generally involves at least partial denaturation during adsorption and so is not recommended for sensitive proteins such as enzymes. Protein purification using ion-exchange chromatography has mainly employed positively charged anion exchangers, for the simple reason that the majority of proteins at neutral pH are negatively charged (i.e., have a low isoelectric point). Details of methodology are found in UNIT 8.2.
Methods Based on Bioproperties (Affinity) A powerful method for separating the desired protein from others is to use a biospecific method in which the particular biological property of the protein is exploited. The affinity approach is limited to proteins that have a specific binding property, except that proteins are theoretically able to be purified by immunoaffinity chromatography (UNIT 9.1), which is the most specific of all affinity techniques. Most proteins of interest do have a specific ligand: enzymes have substrates and cofactors, and hormone-binding proteins and receptor molecules are designed to bind specifically and tightly to particular hormones and other factors. Immobilization of the ligand to which the protein binds (or of antibody to the protein) enables selective adsorption of the desired protein in the technique known as affinity chromatography (Chapter 9). There are also nonchromatographic modes of exploiting biospecific interactions.
CHARACTERIZATION OF THE PROTEIN PRODUCT Once a pure protein is obtained, it may be employed for a specific purpose, such as enzymatic analysis (e.g., glucose oxidase and lac-
1.1.4 Current Protocols in Protein Science
tate dehydrogenase), or as a therapeutic agent (e.g., insulin and growth hormone). However, it is normal, when a protein has been isolated for the first time, to characterize it in terms of structure and function. Several features are generally expected in characterization of a new protein. These include molecular weight, or at least the size of the subunit(s), determined by SDS-polyacrylamide gel electrophoresis (Chapter 10) and/or gel filtration (UNIT 8.3). Spectral properties such as the UV spectrum (Trp and Tyr content), circular dichroism (CD) spectrum (secondary structure), and special characteristics of proteins with prosthetic groups (e.g., quantitation and spectra) may be presented. The quantity and nature of carbohydrates on glycoproteins should be determined (Chapter 12). Also, if the gene has not already been reported, some amino-terminal sequence analysis should be given, if at all possible, along with the results of a database search for similar sequences (UNIT 2.1). Functional proteins should be demonstrated to have the appropriate function, and for enzymes detailed kinetic characterization is appropriate. Ultimately the full three-dimensional structure of the protein may be determined, which will require crystals: any successful crystallization attempts should be reported.
THE PROTEIN PURIFICATION LABORATORY The requirements for a protein purification laboratory cannot be exactly formulated because they depend greatly on the types and amounts of proteins being isolated. To cover all eventualities, it would be necessary to have one set of equipment to deal with submicrogram quantities and another set to deal with multigram quantities—a range of around 108! One laboratory dedicated to protein purification may not need small-scale equipment if, for example, it works with plasma proteins that are always available in large quantities. Another may have all the latest in high-performance equipment but not be able (nor need) to handle quantities of protein in excess of a few milligrams. If it is assumed that neither extreme in quantity is to be attempted, and that the laboratory is handling a variety of protein types and sources, then certain basic pieces of equipment are needed. Obtaining the starting material and making an extract of it require homogenization equipment and centrifuges to remove insoluble residues. Preliminary fractionation, when starting with a crude extract of tissue or cells,
requires equipment and materials that will not become clogged by particulates. Adsorbents and similar materials used at the first step should be relatively inexpensive so that when performance falls off after a few uses, owing to intransigent impurity buildup, they can be discarded. It is also relevant that a larger amount is handled at the initial step than later steps; therefore, reagent expense can be an important consideration. After the first one or two steps, the sample should be sufficiently clean and clear to enable use of high-performance equipment. High-performance liquid chromatography, or HPLC, is a term with a variety of meanings. To some it refers exclusively to reversed-phase chromatography; to others it includes all sorts of chromatography provided that the equipment is fully automated and high-performance adsorbents are used. A high-performance system designed specifically for proteins—introduced by Pharmacia Biotech (see SUPPLIERS APPENDIX) called Fast Protein Liquid Chromatography, or FPLC—uses standard protein chromatographies such as ion exchange, hydrophobic interaction, and gel filtration. Scaleup is possible with larger equipment based on the FPLC design, so that laboratory development can be quickly translated to large-scale production. FPLC is designed to separate proteins in their native active configuration, whereas reversed-phase HPLC often causes at least transient denaturation during adsorption and elution. Reversed-phase HPLC has a high resolving power, but it is best suited to peptides and proteins smaller than ∼30 kDa. Chromatography run with older-style low-pressure adsorbents is sometimes referred to as “low-performance” or “open-column” chromatography; neither of those descriptions is necessarily accurate. Simple fraction collector and monitoring equipment is needed. This equipment will be used for larger-scale operations (tens of milligrams of protein and upward), probably at an earlier stage in the protocol than with HPLC. Various columns, both prepacked with proprietary adsorbents and empty for self-packing, will be needed, with the sizes and types depending on the scale of operations. Several anionexchange columns (different sizes), one or two cation-exchange columns, and gel-filtration media are essential, along with a range of alternative adsorbents such as hydrophobic interaction materials, dyes, hydroxyapatite, and chromatofocusing and specialist affinity media. Fully equipped protein purification laboratories should also have preparative electropho-
Strategies of Protein Purification and Characterization
1.1.5 Current Protocols in Protein Science
resis and isoelectric focusing apparatuses for rare occasions when other techniques fail to give sufficient separation. In addition to equipment used in the actual fractionation processes, a variety of other items are needed. In particular it should be possible to change buffers quickly and to concentrate protein solutions with ease. These operations require such things as dialysis membranes (APPENDIX 3B), ultrafiltration cells, and gel-exclusion columns of various sizes (UNIT 8.3). Finally, equipment for assaying and analyzing the preparations is needed. Most such equipment is fairly standard in biochemical laboratories and includes spectrophotometers, scintillation counters, analytical gel and capillary electrophoresis apparatuses, immunoblotting materials, and immunochemical reagents. A listing of standard equipment is found in APPENDIX 2D.
Literature Cited Cohn, E.J., Strong, L.E., Hughes, W.L., Mulford, D.J., Ashworth, J.N., Melin, M., and Taylor, H.L. 1946. Preparation and properties of serum and plasma proteins. IV. A system for the separation into fractions of the proteins and lipoprotein components of biological tissues and fluids. J. Am. Chem. Soc. 68:459-475.
Key References Deutscher, M.P. (ed.) 1990. Guide to protein purification. Methods Enzymol. 182:1-894. Extensive collection of purification methods with some general protocols and examples. Janson, J.-C. and Ryden, L.G. 1989. Protein Purification: Principles, High Resolution Methods, and Applications. VCH Publishers, New York. A useful collection of methods and examples. Kennedy, J.F. and Cabral, J.M. (eds.) 1993. Recovery Processes for Biological Materials. John Wiley & Sons, New York. A useful introduction to the problems of large-scale methods. Kenny, A. and Fowell, S. (eds.) 1992. Practical protein chromatography. Methods Mol. Biol. 11:1-327. Extensive descriptions of affinity chromatographic techniques with protocols and recipes. Scopes, R.K. 1993. Protein Purification, Principles and Practice, 3rd ed. Springer-Verlag, New York and Heidelberg. General principles of all the main techniques used in purifying proteins. A useful laboratory handbook; does not include recipes or procedures for specific proteins.
Contributed by R.K. Scopes La Trobe University Bundoora, Australia
Overview of Protein Purification and Characterization
1.1.6 Current Protocols in Protein Science
Strategies for Protein Purification CLASSIFICATION OF PROTEINS As with most heterogeneous collections of things, proteins can be classified in several different ways, such as by function, by structure, or by physicochemical characteristics. Each protein species consists of identical molecules with exactly the same size, amino acid sequence, and three-dimensional shape. In this way a solution of a mixture of proteins differs from a solution of synthetic polymers or sheared DNA, both of which contain a complete spectrum of possible sizes centered around the average. The protein mixture has only discrete sizes of molecules corresponding to each type of protein present. Although we could classify proteins by size, it would be of limited use, as there is usually no obvious relationship between size and function. A more useful structural classification takes into consideration shape and oligomeric structure (Table 1.2.1). In part, structure reflects biological location and origin. Simple, fairly rigid protein molecules occur in the extracellular environment, more complex and readily deactivated molecules are found intracellularly, and hydrophobic proteins are associated with membranes. Classification by function is more relevant (Table 1.2.2). Proteins can be simply stores of amino acids, can be structural, or can have specific binding functions. The most “funcTable 1.2.1
UNIT 1.2
tional” proteins are enzymes, which have both binding and catalytic roles. In part, this reflects the degree to which the detailed structure is a requirement for the protein’s function, which in turn relates to conservation of structure through evolution. But as with every attempt at classification, there are going to be examples that do not fit the pattern well. Most proteins of interest to the pharmaceutical industry belong to the general class of binding proteins, for instance, hormones [e.g., insulin and bovine somatostatin (BST)], viral antigens (e.g., hepatitis B antigen), growth factors [e.g., interferons, interleukins and colony-stimulating factors (CSFs)], and antibodies.
STRATEGIES FOR PROTEIN PURIFICATION Soluble Extracellular Proteins The source of soluble extracellular proteins is the extracellular medium, whether it be an animal source such as blood or spinal fluid, or a culture medium in which bacterial, fungal, animal, or plant cultures have been grown. Generally these do not contain a large number of different proteins (blood is an exception), and the desired protein may be a major component, especially if produced as the result of recombinant expression. Nonetheless, the protein in the starting material may be quite dilute,
Classification of Proteins by Structural Characteristics
Structural characteristic
Examples
Comments
Monomeric
Lysozyme, growth hormone
Usually extracellular; often have disulfide bonds
Glyceraldehyde-3-phosphate dehydrogenase, catalase, alcohol dehydrogenase, hexokinase Aspartate carbamoyltransferase, pertussis toxin
Mostly intracellular enzymes; rarely have disulfide bonds
Mitochondrial ATPase, alkaline phosphatase
Readily solubilized by detergents
Integral
Porins, cytochromes P450, insulin receptor
Require lipid for stability
Conjugated
Glycoproteins, lipoproteins, nucleoproteins
Many extracellular proteins contain carbohydrate
Oligomeric Identical subunits
Mixed subunits Membrane-bound Peripheral
Contributed by R.K. Scopes Current Protocols in Protein Science (1995) 1.2.1-1.2.4 Copyright © 2000 by John Wiley & Sons, Inc.
Allosteric enzymes; different subunits have separate functions
Strategies of Protein Purification and Characterization
1.2.1 CPPS
Table 1.2.2
Classification of Proteins by Function
Function
Examples
Amino acid storage
Seed proteins (e.g., gluten), milk proteins (e.g., casein)
Structural Inert With activity Binding Soluble Insoluble With activity
Collagen, keratin Actin, myosin, tubulin Albumin, hemoglobin, hormones Surface receptors (e.g., insulin receptor), antigens (e.g., viral coat proteins) Enzymes, membrane transporters (e.g., amino acid uptake systems, ion pumps)
and a large volume may therefore need to be processed. The starting fluid may also contain many compounds other than proteins, whose behavior must be taken into account. The first stage should aim mainly to reduce the volume and get rid of as much nonprotein material as possible; some protein-protein separation is also useful, but not essential. No general rules can be given, but a batch adsorption method using an inexpensive material such as hydroxyapatite, ion-exchange resin, immobilized metal affinity chromatography (IMAC) medium, or affinity adsorbent is best, if feasible. Following the first step, the sample should be in a form that is amenable to standard purification processes such as precipitation and column chromatography (Chapter 8).
Intracellular (Cytoplasmic) Proteins
Strategies for Protein Purification
To obtain soluble intracellular proteins (which are mainly enzymes), cells must be broken open or lysed to release their soluble contents. The ease with which cell disruption can be accomplished varies considerably; animal cells are readily broken, as are many bacteria, but plants and fungi have tough cell walls. Methods for obtaining cell extracts are given in Chapter 4. The macromolecular soluble contents of cells are mainly proteins, with nucleic acids as a minor but significant component. Bacterial extracts may be viscous unless DNase is added to break down the long DNA molecules. Although chromatographic procedures can be applied to crude extracts, valuable highperformance materials should not be employed in the first step, as there are always compounds, including unstable proteins, that may bind to them and be difficult to remove.
Membrane-Associated Proteins There are two approaches to isolating a membrane-associated protein. In one method, the relevant membrane fraction can first be prepared and then used to isolate the protein. Alternatively, whole tissue can be subjected to an extraction that solubilizes the membranes and releases the cytoplasmic contents as well. The former is much better in that purification is accomplished by isolating the membranes: the specific activity of the solubilized membrane fraction will be much higher than in the second method. However, the process of purifying the membrane fraction may lead to substantial losses, and it may be difficult to scale up. If total recovery of the protein is more important than purity, a whole-tissue extract is likely to be more appropriate. Although this means that a greater degree of purification is needed, the fact that membrane proteins have, by definition, properties somewhat different from those of cytoplasmic proteins permits some very effective purification steps (e.g., hydrophobic chromatography or fractional solubility separation). Peripheral membrane proteins are only loosely attached and may be released by gentle conditions such as high pH, EDTA, or low (nonionic) detergent concentrations. Once in solution, some peripheral proteins no longer require the presence of detergent to maintain their solubility. Integral membrane proteins are much more difficult—they require high concentrations of detergent for solubilization (i.e., complete solubilization of the membrane is needed to release them) and generally are neither soluble nor stable in the absence of detergent. It is sometimes necessary to maintain
1.2.2 Current Protocols in Protein Science
natural phospholipids in association with the proteins in order to maintain activity. Even though the final objective may not require activity (e.g., amino acid sequencing), there is a need for some sort of assay during the purification process to determine where the protein is. If a particular band on a gel is known to be the desired protein, then no other assay is needed and loss of bioactivity can be allowed. Purification processes may be affected by the presence of detergents. The problem of association with detergent micelles makes purifying integral membrane proteins difficult; the close association of the different proteins originating from membranes often results in very poor separation in conventional fractionation procedures.
Insoluble Proteins Natural proteins that are insoluble in normal solvents are generally structural proteins, which are sometimes cross-linked by posttranslational modification. The first stage of purification is obvious—it involves extracting and washing away all proteins that are soluble, leaving the residue containing the desired material. Further purification in a native state, however, may be impossible; extracting away other proteins using more vigorous solvents or attempting to solubilize the target protein may destroy the natural structure. Cross-linked proteins such as elastin or aged collagen cannot be dissolved without breaking the cross-links, and the individual proteins may even be crosslinked together.
Insoluble Recombinant Proteins (Inclusion Bodies) A major new class of insoluble proteins are recombinant proteins expressed (usually in Escherichia coli) as inclusion bodies. These are dense aggregates found inside cells that consist mainly of a desired recombinant product, but in a nonnative state. Inclusion bodies may form for a variety of reasons, such as insolubility of the product at the concentrations being produced, inability to fold correctly in the bacterial environment, or inability to form correct, or any, disulfide bonds in the reducing intracellular environment. Their purification is simple, since the inclusion bodies can be separated by differential centrifugation from other cellular constituents, giving almost pure product; the problem is that the protein is not in a native state, and is insoluble. Some methods for obtaining an active product from inclusion bodies are described in UNIT 6.3.
Soluble Recombinant Proteins Recombinant proteins that are not expressed in inclusion bodies either will be soluble inside the cell or, if using an excretion vector, will be extracellular (or, if E. coli is the host, possibly periplasmic). They can be purified by conventional means. In some systems, expression is so good that the desired product is the major protein present and its purification is relatively simple. In systems where the expression level is low, the purification process can be tedious, though easier, it is hoped, than isolation from the natural source. It should be remembered that a procedure developed for isolating a protein from natural sources may not work successfully with the recombinant product, because the nature of the other proteins present influences many fractionation procedures. Because of the difficulties often experienced in purifying recombinant proteins, a variety of vector systems (see UNIT 6.1 and Sassenfeld, 1990, for some examples) have been developed in which the expressed prod-uct is a fusion protein containing an N-terminal polypeptide that simplifies purification. Such “tags” can be subsequently removed using a specific protease. A further advantage is that the expression level is dictated mainly by the transcription and translation signals for the fusion portion of the protein, which are optimized. Tags used include proteins and polypeptides for which there is a specific anti- body, binding proteins that will interact with columns containing a specific ligand, polyhistidine tags with affinity to immobilized metal columns, sequences that result in biotinylation by the host and enable purification on an avidin column, and sequences that confer insolubility under specified conditions (UNIT 6.1 & UNIT 6.5). Unstable proteins may be modified by the molecular biological technique of site-directed mutagenesis to remove the site of instability— for instance, an oxidizable cysteine. Such techniques are appropriate for commercial production of proteins, but may of course alter natural functioning parameters. Increased thermostability can be one modification, although it is not easy to predict mutations that will improve that parameter. Thermostable proteins originating from thermophilic bacteria do not need structural modification and, if expressed in large amounts, can be purified satisfactorily in one step by simply heat-treating the extract at 70°C for 30 min, which denatures virtually all the host proteins (e.g., see Oka et al., 1989). The host bacteria used for production of recombinant proteins are usually Escherichia
Strategies of Protein Purification and Characterization
1.2.3 Current Protocols in Protein Science
coli, or Bacillus subtilis; they may express proteins at 1% to over 50% of the cellular protein, depending on such variables as the source, promoter structure, and vector type. Generally the proteins are expressed intracellularly, but leader sequences for excretion may be included. In the latter case, the protein is generally excreted into the periplasmic space, which limits the amount that can be produced. Excretion from gram-positive species such as B. subtilis sends the product into the culture medium, with little feedback limitation on total expression level.
Literature Cited Oka, M., Yang, Y.S., Nagata, S., Esaki, N., Tanaka, M., and Soda, K. 1989. Overproduction of thermostable leucine dehydrogenase of Bacillus stearothermophilus and its one-step purification from recombinant cells of Escherichia coli. Biotechnol. Appl. Biochem. 11:307-316. Sassenfeld, H.M. 1990. Engineering proteins for purification. Trends Biotechnol. 8:88-93.
Key References Deutscher, M.P. (ed.) 1990. Guide to protein purification. Methods Enzymol. 182:1-894.
Janson, J.-C. and Ryden, L.G. 1989. Protein Purification: Principles, High Resolution Methods, and Applications. VCH Publishers, New York. A useful collection of methods and examples. Kennedy, J.F. and Cabral, J.M. (eds.) 1993. Recovery Processes for Biological Materials. John Wiley & Sons, New York. A useful introduction to the problems of large-scale methods. Kenny, A. and Fowell, S. (eds.) 1992. Practical protein chromatography. Methods Mol. Biol. 11:1-327. Extensive descriptions of affinity chromatographic techniques with protocols and recipes. Scopes, R.K. 1993. Protein Purification, Principles and Practice, 3rd ed. Springer-Verlag, New York and Heidelberg. General principles of all the main techniques used in purifying proteins. A useful laboratory handbook; does not include recipes or procedures for specific proteins.
Contributed by R.K. Scopes La Trobe University Bundoora, Australia
Extensive collection of purification methods with some general protocols and examples.
Strategies for Protein Purification
1.2.4 Current Protocols in Protein Science
Protein Purification Flow Charts Protein purification flow charts are presented to give a broad outline of the methods used for different types of proteins. They cannot give any detail, as the process appropriate for each protein will have its own variations at each stage. In most cases, the first stage is to obtain a solution containing the desired protein, after which it can be dealt with by the many separation techniques described in the following chapters. In some cases the insolubility of the desired protein can be exploited by removing soluble fractions. Purification procedures are commonly divided into three stages: (a) the primary steps, which deal with crude mixtures of proteins and other molecules present in the raw material; (b) the secondary processing, which generates a product near to homogeneity; and (c) the polishing steps, which remove minor contaminants, a process that is especially important for therapeutic proteins.
SOLUBLE RECOMBINANT PROTEINS Proteins expressed in a recombinant manner may be (1) soluble in the cytoplasm, (2) insoluble as inclusion bodies (see section on Insoluble Recombinant Proteins), (3) excreted from the cells into the culture medium, (4) excreted into the periplasmic space (e.g., in gram-negative bacteria), or (5) associated with organelles or membrane fractions. In addition they may be expressed (6) as the normal, mature, naturally occurring protein, (7) containing a natural leader peptide that would normally be processed, (8) as a fusion protein with a peptide that is not natural to the protein, or (9) lacking glycosylation or other post-translational modification, or incorrectly modified. Possibilities (1) to (5) affect the method of extraction used to obtain the starting material for purification. Cases (6) to (9) can affect the methods used for purification. The scheme for purifying soluble recombinant proteins is outlined in Figure 1.3.1. The first stage is to obtain a clarified solution containing the desired protein, with as little in the way of unwanted proteins as possible. For soluble cytoplasmic proteins, case (1), it is not normally possible to exclude any significant amount of unwanted soluble proteins, but in cases (2) to (5) the compartmentalization away from the cytoplasm allows such separation in the initial stage. It may be necessary to carry out a concenContributed by R.K. Scopes Current Protocols in Protein Science (1995) 1.3.1-1.3.7 Copyright © 2000 by John Wiley & Sons, Inc.
UNIT 1.3
tration step before proceeding, especially if the protein has been excreted into the culture medium. Normally ultrafiltration is used, although other techniques are possible, especially if the extract contains particulates that block ultrafiltration membranes. Recombinant expression in the cytoplasm of bacteria, followed by extraction via total cell disruption, results in large amounts of nucleic acids being solubilized with the protein. A number of treatments to remove nucleic acids are possible. Streptomycin is used to precipitate ribosomal material, and cationic polymers such as protamine (a basic protein) and polyethylenimine will form insoluble complexes (at low ionic strength) with nucleic acids. In addition, viscosity caused by DNA can be reduced by adding small amounts of DNase.
INSOLUBLE RECOMBINANT PROTEINS It has been found that many proteins expressed in bacteria (mainly in Escherichia coli) do not fold correctly, and as a result aggregation occurs, leading to large insoluble inclusion bodies within the cytoplasm of the cells (see Chapter 6). Although this creates major difficulties in obtaining satisfactory amounts of active native product, it greatly simplifies the initial stage of purification. The purification scheme for recombinant insoluble proteins is outlined in Figure 1.3.2. After cell disruption, inclusion bodies can be obtained in a fairly pure state by differential centrifugation. They must now be solubilized, however, and the active protein generated by encouraging correct folding. Solubilization is usually accomplished with guanidine hydrochloride and/or urea, and thiols such as 2-mercaptoethanol or glutathione are included to disrupt any disulfides that have formed and prevent more from forming. Folding the protein correctly may require a variety of additions to the solution, as well as slow removal of the denaturant. The latter can be carried out by simple dilution or by dialysis. Folding occurs best at low protein concentrations, so dilution may be adequate. If the native protein does contain disulfides, then it is important to create redox conditions such that some (but not excessive) oxidation of thiols can occur. A combination of oxidized and reduced glutathione is commonly used. In addition, the action of the enzyme protein disulfide isomerase, which can
Strategies of Protein Purification and Characterization
1.3.1 CPPS
cells with protein soluble in cytoplasm
cells with protein excreted to medium
cells with protein in periplasmic space
cells with protein organelleassociated
break up cells; centrifuge or ultrafilter to remove insolubles
centrifuge or filter to remove cells
gently treat with lysozyme to minimize cell lysis
separate organelles; perform differential centrifugation
treat to remove nucleic acids
remove protoplasts and cell debris
extract to solubilize proteins, remove insolubles
perform concentration step
starting material: soluble fraction containing the protein
perform purification steps use salt fractionation, ion-exchange chromatography, hydrophobic chromatography, affinity and pseudoaffinity chromatography, gel filtration, or other less conventional steps (i.e., preparative electrophoresis)
for tagged fusion protein, use affinity column chromatography and polishing steps
purified protein
Figure 1.3.1 Purification scheme for soluble recombinant proteins, which may be excreted or located in the periplasm, in the membrane fraction, or most commonly the cytoplasm. The first step is to obtain an extract containing the desired protein in soluble form. After this, conventional purification steps may be carried out, or affinity purification of tagged fused proteins can be performed.
Protein Purification Flow Charts
make and unmake disulfides by exchange reactions, has been found to be beneficial in many cases. If the native protein is of intracellular origin, it probably will not contain disulfides; it will, however, contain cysteines, so a full reducing potential should be maintained. Specific methodology is discussed in UNIT 6.5. Not all proteins can fold unassisted by other cellular components. Chaperonins are proteins whose role is to refold denatured proteins such as those that form during heat shock (Zeilstra-
Ryalls et al., 1991). The most studied, and just becoming commercially available as of 1995, are the E. coli chaperonins GroEL and GroES, both of which are needed, together with ATP, to renature many proteins. Proline residues can adopt two isomeric conformations in proteins, and the wrong conformation is switched to the correct one by the enzyme prolyl isomerase, aiding the process of protein folding. At present these are not large-scale prospects, both because of the cost of the chap-
1.3.2 Current Protocols in Protein Science
cells containing protein
disrupt cells perform differential centrifugation
wash inclusion bodies dissolve in denaturing agents
dilute with buffer or dialyze to dilute denaturing agents add appropriate reducing agents and/or folding factors concentrate perform purification procedure: remove unwanted proteins and incorrectly folded species (e.g., by ion-exchange chromatography, immunoaffinity methods, gel filtration)
purified protein
Figure 1.3.2 Purification scheme for insoluble recombinant proteins that are produced as inclusion bodies in the cytoplasm of host cells. The cells must be broken open, and then the insoluble inclusion bodies are separated by differential centrifugation. Solubilization is achieved by the use of denaturing solvents, and renaturation of the dissolved protein occurs on removal of the denaturant. Further polishing steps will be needed to remove small amounts of contaminating proteins as well as incorrectly folded species. Additional information can be found in UNIT 6.3-6.5.
eronins and because the agents operate best in vitro at very low protein concentrations. Once the proteins are folded, the purification process consists of removing small amounts of still incorrectly folded protein plus any other host proteins that were trapped with the original inclusion bodies. The former may be difficult, since incorrectly folded species have a size and charge similar to those of the correct product. However, subtle differences arising from the folded conformation can be exploited by chromatographic techniques. In ideal cases immunoaffinity techniques using antibodies specific for either the incorrectly folded form or the correct one can be used to resolve the mixture.
SOLUBLE NONRECOMBINANT PROTEINS There are so many sources of soluble proteins that it is not possible to give a complete overview of methods used to obtain starting extracts from which a desired protein can be isolated. The sources can be classified as either microorganisms, plants, or animals, as shown in Figure 1.3.3, but these in turn should be subdivided according to how the starting extract is obtained. In particular there is a distinction between extracellular and intracellular proteins. With the latter it is necessary to disrupt the cells and release the proteins, whereas with the former, if the extracellular fluid can be obtained directly, there need be no contamination with intracellular proteins. Extracellular
Strategies of Protein Purification and Characterization
1.3.3 Current Protocols in Protein Science
microorganisms
plants
animals
disrupt cells, e.g., using French press, bead mill, or sonication with 5-10 vol buffer
disrupt cells, e.g., using French press or by grinding with sand using 1-2 vol buffer
homogenize with 2-5 vol buffer or mince and stir with buffer
treat to remove nucleic acids
treat to remove phenols
centrifuge
starting extract (5-20 mg/ml protein)
extracellular proteins
perform initial fractionation: use salt fractionation, precipitation with organic solvents, affinity methods, and/or two-phase partitioning perform secondary fractionation: use ion-exchange chromatography, hydrophobic chromatography, affinity methods, other adsorbents, and/or gel filtration perform polishing step: use HPLC– reversed phase, HPLC– ion exchange, or isoelectric focusing
purified protein
Figure 1.3.3 Purification scheme for soluble proteins present in their natural host cells. Cells must be disrupted to release the proteins, usually in the presence of 2 to 10 ml of a suitable buffer per gram weight. After removal of insoluble material, the process will generally require several steps, using various standard fractionation procedures in a suitable order. For production of highly pure protein, a final polishing step may be required to remove final trace contaminants. Additional information can be found in UNIT 6.2.
Protein Purification Flow Charts
sources include microorganism culture medium, plant and animal tissue culture medium, venoms, milk, blood, and cerebrospinal fluids. Soluble proteins may also occur within organelles such as mitochondria; these may be best obtained by first isolating the organelle, then disrupting it to release the contents. The starting extract normally contains be-
tween 5 and 20 mg protein per milliliter, though lesser concentrations can be dealt with, especially if working on a small scale. It may be necessary to include a concentration step before starting the purification process in order to approach that level. There are exceptions to every rule, however, and very high protein concentrations can be handled, for example,
1.3.4 Current Protocols in Protein Science
whole tissue homogenize without detergent; centrifuge
supernatant
disrupt cells; gently isolate organelles
residue homogenize with detergent; centrifuge
supernatant containing membrane-associated proteins perform primary fractionation: use hydrophobic chromatography, precipitation with organic solvents, ion-exchange chromatography, and/or phase partitioning perform secondary fractionation: use ion-exchange chromatography, affinity methods, and/or gel filtration
residue containing insoluble proteins
suspend and extract with other solvents (e.g., ethanol, urea, or SDS)
purified protein
insoluble residue
perform polishing step: use HPLC or isoelectric focusing
purified protein
Figure 1.3.4 Purification scheme for membrane-associated and poorly soluble proteins (nonrecombinant). An initial purification can be achieved by isolation of organelles containing the desired protein. Membrane proteins are normally solubilized with a nonionic detergent, although loosely associated proteins may be extracted without detergent at high pH, with EDTA, or with small amounts of an organic solvent such as N-butanol. Normal fractionation procedures may need some modification if the detergent is required throughout to maintain the integrity of the protein.
with two-phase partitioning (Walter and Johansson, 1994). When isolating proteins on a large scale, the volumes being manipulated become of increasing concern, so maximizing protein concentration can be an important aim. The starting extract should be clarified, usually by centrifugation; on a large scale, ultrafiltration methods are becoming more widely used. Pretreatment of certain extracts to remove ex-
cessive amounts of nucleic acids, phenolics, and lipids may be necessary in order to obtain an extract that is amenable to standard fractionation procedures. Fractionation procedures can somewhat arbitrarily be divided into three steps: initial fractionation, secondary fractionation, and polishing. Initial processing, which deals with a large amount of extract that is not all protein, may
Strategies of Protein Purification and Characterization
1.3.5 Current Protocols in Protein Science
involve materials that become soiled and unable to be used many times. Consequently, preferred methods do not require expensive reagents or adsorbents such as are used in highperformance liquid chromatography (HPLC). Classic salt fractionation and the less-used organic solvent fractionation can achieve, if not a high degree of purification, a useful level of concentration and removal of much unwanted nonproteinaceous material. Alternatively, a highly selective affinity procedure may be used as the first step, but only if the affinity material is inexpensive to make and/or the extract is a simple, clear solution, as opposed to a turbid whole-cell homogenate. Secondary processing achieves the main purification, and may involve two or more steps in difficult situations. Ion-exchange and hydrophobic-interaction chromatography, gel filtration, and affinity techniques (see Chapter 8 and Chapter 9) are the main procedures. Finally, it may be necessary to remove traces of contaminants by “polishing,” using high-resolution procedures such as reversed-phase HPLC (UNIT 11.6) and isoelectric focusing (IEF; UNIT 10.2 & UNIT 10.4). Because every protein has unique characteristics, it is impossible to make general statements about procedures to be followed.
MEMBRANE-ASSOCIATED AND INSOLUBLE NONRECOMBINANT PROTEINS
Protein Purification Flow Charts
Proteins that are not physiologically soluble can be purified after extracting and removing soluble proteins, thereby achieving a substantial degree of purification at the extraction step (Fig. 1.3.4; also see UNITS 6.1 & 6.2). To carry out a purification it is nearly always necessary to obtain the desired protein in a soluble form, which will often require the addition of solubilizing agents such as detergents. Some proteins remain insoluble even with detergent treatment, and so can be substantially purified by removing the soluble fractions. Some membrane-associated proteins become partly solubilized during breaking up of the tissue, and recovery in the particulate fraction may be poor. In such cases it may be best to solubilize the whole tissue by including detergent in the homogenizing buffer. Extraction of insoluble residues using detergents can be done differentially; some proteins are released at low detergent concentration, whereas others require complete solubilization of the membrane fraction. Suitable detergents include nonionic (e.g., Triton) and weakly acidic types (e.g., cholic acid deriva-
tives). Strongly acidic detergents such as sulfate esters (e.g., sodium dodecyl sulfate) usually result in denaturation. Detergents can be removed either by adsorption of the protein on a column, and subsequent elution without detergent, by use of special detergent-adsorbing beads, or even by extraction with nonmiscible organic solvents in which the detergent partitions. On the other hand, many membrane proteins require the presence of detergent at all times in order to remain in solution and in a native conformation. These include most integral membrane proteins, for example, cytochrome: P450, transmembrane receptors, and transporters. The most sensitive proteins require a particular combination of natural lipids (in addition to the detergent) to maintain structural integrity. Purification methods include most of those used for soluble proteins, but some techniques are not recommended if detergent is needed at all times. For instance, ammonium sulfate precipitation will often cause a detergent-protein complex to come out of solution and float rather than sink on centrifugation—this can be useful, but the “floatate,” when redissolved, may have a high detergent content. Hydrophobic chromatography can be very useful, as membrane proteins are naturally hydrophobic. Integral membrane proteins that are completely insoluble in normal detergents may be solubilized by denaturation using compounds such as sodium dodecyl sulfate and guanidine hydrochloride. Some cross-linked proteins such as elastin are not soluble without disruption of the covalent linkages.
Literature Cited Walter, H. and Johansson, G. (eds.) 1994. Aqueous two-phase systems. Methods Enzymol. 228:1725. Zeilstra-Ryalls, J., Fayet, O., and Georgopoulos, C. 1991. The universally-conserved GroE (Hsp60) chaperonins. Annu. Rev. Microbiol. 45:301-325.
Key References Deutscher, M.P. (ed.) 1990. Guide to protein purification. Methods Enzymol. 182:1-894. Extensive collection of purification methods, with some general protocols and examples. Janson, J.-C. and Ryden, L.G. 1989. Protein Purification: Principles, High Resolution Methods, and Applications. VCH Publishers, New York. A useful collection of methods and examples.
1.3.6 Current Protocols in Protein Science
Kennedy, J.F. and Cabral, J.M. (eds.) 1993. Recovery Processes for Biological Materials. John Wiley & Sons, New York. A useful introduction to the problems of large-scale methods. Kenny, A. and Fowell, S. (eds.) 1992. Practical protein chromatography. Methods Mol. Biol. 11:1-327. Extensive descriptions of affinity chromatographic techniques with protocols and recipes.
Scopes, R.K. 1993. Protein Purification, Principles and Practice, 3rd ed. Springer-Verlag, New York and Heidelberg. General principles of all the main techniques used in purifying proteins. A useful laboratory handbook; does not include recipes or procedures for specific proteins.
Contributed by R. K. Scopes La Trobe University Bundoora, Australia
Strategies of Protein Purification and Characterization
1.3.7 Current Protocols in Protein Science
Purification of Glutamate Dehydrogenase from Liver and Brain
UNIT 1.4
Because of the wide variety of procedures that may be used to purify any given enzyme, it is not possible to provide a single protocol that includes them all. In this unit two protocols for the purification of glutamate dehydrogenase (GDH)—also referred to as + L-glutamate-NAD(P) oxidoreductase (deaminating; EC 1.4.1.3:GDH)—have been selected as examples because they present two very different approaches to the purification of the enzyme. In the first (see Basic Protocol 1), affinity chromatography using a column consisting of the allosteric inhibitor guanosine 5′-triphosphate (GTP) bound to Sepharose is described. Support protocols detail the preparation of the GTP-Sepharose needed for this procedure (see Support Protocol 1), assessment of incorporation of GTP into the GTP-Sepharose gel, if necessary (see Support Protocol 2), the activation of Sepharose 4B with CNBr, if necessary (see Support Protocol 3), and reuse and equilibration of the chromatography media (see Support Protocols 4 and 5). An alternative purification procedure (see Alternate Protocol) uses a bifunctional ligand composed of two NAD+ molecules linked together by a spacer arm to precipitate the enzyme. Protocols for the synthesis of the bis-NAD compound (see Support Protocol 7), resolution of NAD+ derivatives (see Support Protocol 8), and instructions for a pilot affinity study (see Support Protocol 9) complement this alternative approach. Of value to both studies are methods for concentrating the protein product (see Support Protocol 6), determining the protein concentration (see Basic Protocol 2), and assaying for GDH activity (see Basic Protocol 3). Although some of these techniques can be found elsewhere in this manual, instructions are included in this unit that are specific to the systems mentioned here. As discussed below, some tissues, including the brain but not the liver, contain an additional isoenzyme of glutamate dehydrogenase (GLUD2) as a relatively minor component. The procedures described here are for purifying the major form (GLUD1). Since this unit is concerned with the purification of mammalian glutamate dehydrogenase, a detailed review of its properties and functions is not appropriate. However, some of the essential properties are briefly summarized below. Further details can be found in the references cited. References to earlier work on the enzyme can be found in Tipton and Couée (1988) and are not repeated here. GDH catalyzes the reversible oxidative deamination of glutamate to 2-oxoglutarate (α-ketoglutarate). Either NAD+ or NADP+ can serve as cofactor, although the kinetic and regulatory behavior depend on which is used. The native enzyme is a hexamer of six identical subunits, each consisting of 505 amino acids, with approximate relative molecular mass of 56,700. However, at high protein concentrations, the hexamers may aggregate further to form high-molecular-weight polymers. The crystal structure of the enzyme has been recently determined (Peterson and Smith, 1999; Smith et al., 2001). The enzyme activity is allosterically modulated by a range of metabolites, including adenosine, guanosine, and inosine di- and trinucleotides, as well as L-leucine (see Couée and Tipton 1989a). The exact behavior of these compounds depends on the conditions under which activity is measured and whether NAD+ or NADP+ is used as cofactor. The kinetic behavior in the direction of glutamate oxidation is complex. Substrate inhibition by high concentrations of 2-oxoglutarate occurs in the reaction leading to NADPH oxidation, but this is not apparent in the reaction pathway leading to NADH oxidation. For that reason, activity determinations are usually performed in the direction of 2-oxoglutarate amination with NADH as the cofactor. Phospholipids, such as cardiolipin, also
Strategies of Protein Purification and Characterization
Contributed by Martha Motherway, Keith F. Tipton, Alun D. McCarthy, Ivan Couée, and Jane Irwin
1.4.1
Current Protocols in Protein Science (2002) 1.4.1-1.4.34 Copyright © 2002 by John Wiley & Sons, Inc.
Supplement 29
affect enzyme activity (Couée and Tipton, 1989b) and the enzyme is also inhibited by antipsychotic drugs such as perphenazine (Couée and Tipton, 1990a,b; Yoon et al., 2001). GDH is localized in the mitochondrial matrix, although small amounts of the activity have been detected associated with microtubules (Rajas and Rousset, 1993) and endoplasmic reticulum (Lee et al., 1999). A nuclear-associated activity has also been reported (McDaniel, 1995) and the enzyme has been found to be capable of functioning as an RNA-binding protein (Preiss et al., 1993). In addition to the ubiquitously distributed GDH isoenzyme (GLUD1), which is encoded by a gene on chromosome 10, brain, retina, and testis tissue also contain a second form of the enzyme (GLUD2), encoded on the X chromosome (see Shashidharan et al., 1994). This isoenzyme differs in its kinetic and regulatory properties from the major form of GDH (see e.g., Cho et al., 1995; Shashidharan et al., 1997). The possible functions of these two GDH isoenzymes in brain have been discussed by Plaitakis and Zaganas (2001). NOTE: Unless otherwise stated, all required chemicals can be obtained from SigmaAldrich or BDH. They should be of the highest purity available. BASIC PROTOCOL 1
PURIFICATION OF OX BRAIN OR LIVER GLUTAMATE DEHYDROGENASE BY AFFINITY CHROMATOGRAPHY ON GTP-SEPHAROSE The initial steps in this protocol involve the classical procedures of salt precipitation and ion-exchange chromatography. These are then followed by affinity chromatography on GTP Sepharose. GTP is an allosteric inhibitor of glutamate dehydrogenase when NAD+ is used as the substrate (see McCarthy and Tipton, 1985; Couée and Tipton, 1989a). The procedure described here is that of McCarthy et al. (1980). The steps involved are summarized in Figure 1.4.1. Materials Fresh ox (beef) brain (∼400 g) or liver (∼50 g) from slaughterhouse 0.32 M sucrose (optional; if tissue samples are to be stored before use) Homogenization buffer (see recipe), 4°C Ammonium sulfate [(NH4)2SO4] 20 mM and 200 mM sodium phosphate buffer, pH 7.4 (APPENDIX 2E), 4°C Affinity chromatography buffer (see recipe) 400 mM KCl in affinity chromatography buffer (see recipe for buffer) Glycerol
Purification of Glutamate Dehydrogenase from Liver and Brain
Scissors Waring blender or kitchen liquidizer Refrigerated centrifuge (e.g., Sorvall RC-5C) Dounce (hand-held) homogenizer with loosely-fitting pestle (clearance, ∼0.03 cm) 250 ml or 500 ml polycarbonate centrifuge tubes Dialysis tubing, 12,000 molecular weight cut-off (MWCO) Conductivity meter Gradient mixer (e.g., UNIT 8.2) Chromatography columns (see Support Protocol 5 for equilibration; column lengths may be varied somewhat according to availability): 40 × 3–cm packed to height of 30 cm with DEAE-cellulose (DE-52, Whatman) and equilibrated with 20 mM sodium phosphate buffer, pH 7.4 (see APPENDIX 2E for buffer) 30 × 2.5–cm packed to height of 22 cm with Sephadex G-25 (Pharmacia Biotech) and equilibrated with affinity chromatography buffer (see recipe for buffer)
1.4.2 Supplement 29
Current Protocols in Protein Science
ox brain (400 g)
homogenize for 1 min in 3 volumes (w/v) of 4 mM sodium phosphate buffer, pH 7.4, containing 0.5mM EDTA and 0.1 mM PMSF
ox liver (50 g)
homogenize for 1 min in 500 ml of 4 mM sodium phosphate buffer, pH 7.4, containing 0.5 mM EDTA and 0.1 mM PMSF
stage 1 (homogenate) add solid (NH4)2 SO4 (113 g/liter), stir for 20 min
centrifuge discard pellet
stage 2 (NH4)2 SO4 supernatant
add solid (NH4)2 SO4 (188 g/liter), stir for 20 min
stage 3 (dialyzed pellet)
centrifuge discard supernatant dialyse against 20 mM sodium phosphate buffer, pH 7.4 and apply to DEAE-cellulose column elute with 20 to 200 mM sodium phosphate, pH 7.4, gradient
stage 4 (DEAE eluate)
gel-filter into affinity chromatography buffer; pass through hydrazide-Sepharose; and apply to GTP Sepharose column
stage 5 (GTP-Sepharose eluate)
elute with 0 to 15 mM KCl gradient in affinity-chromatography buffer concentrate by ultrafiltration dialyze against 20 mM sodium phosphate buffer, pH 7.4 purified glutamate dehydrogenase
Figure 1.4.1 Flow chart for the purification of ox liver and brain glutamate dehydrogenase according to the Basic Protocol 1. Full details of the steps involved are given in the text.
6 × 2–cm packed to height of 4 cm with ∼10 ml of hydrazide-Sepharose 4B and equilibrated with affinity chromatography buffer (see Support Protocol 1, steps 1 to 3 for resin; see recipe for buffer) 6 × 2–cm packed to height of 4 cm with GTP-Sepharose 4B and equilibrated with affinity chromatography buffer (see Support Protocol 1, steps 1 to 7 for resin; see recipe for buffer)
Strategies of Protein Purification and Characterization
1.4.3 Current Protocols in Protein Science
Supplement 29
Additional reagents and equipment for dialysis (UNIT 4.4 and APPENDIX 3B), determining protein concentration (see Basic Protocol 2 and UNIT 3.4), assaying GDH activity (see Basic Protocol 3), and concentrating protein solutions by ultrafiltration (see Support Protocol 6) NOTE: Unless indicated otherwise, all steps are performed at 0° to 4°C in a cold room or refrigerated cabinet. Prepare and homogenize the tissue 1. Obtain one ox brain (∼400 g) or liver (∼50 g) from a freshly slaughtered animal transported to the laboratory on ice to be used as the starting material for the purification. The brain may be used fresh or stored at −70°C, using the procedure as follows: Place the fresh tissue in ice-cold 0.32 M sucrose solution and keep it in a −20°C freezer until it has frozen. Transfer to a −70°C freezer until required. Before use place the vessel containing the frozen material in a water bath at 37°C and allow it to thaw, with occasional agitation. The larger amount of enzyme in ox liver permits the use of a smaller amount of starting material (e.g., 50 g in the example purification shown here) to be used.
2a. For brain tissue: Divide the brain tissue into three equal portions, weigh, and suspend each portion in 3 volumes (w/v) of cold homogenization buffer. 2b. For liver tissue: Weigh the liver tissue and suspend in 10 volumes (w/v) of cold homogenization buffer. 3. Chop the tissue into smaller pieces using scissors, then homogenize in a Waring blender at full speed for 1 min. Retain a small aliquot of this homogenate for determining protein concentration by the biuret procedure (see Basic Protocol 2) and assaying GDH activity (see Basic Protocol 3). A kitchen liquidizer can be used instead of a Waring blender. However, extra care is then needed with any biohazard material (see Critical Parameters) since kitchen liquidizers can release aerosols. These are generally less powerful than the Waring blender and it is thus necessary to increase the homogenization time (e.g., 1.5 min using a Kenwood liquidizer at full speed). The volumes given in the subsequent steps refer to the preparation of GDH from brain. The corresponding values for liver are given in Table 1.4.6 (see Critical Parameters and Troubleshooting).
Precipitate with ammonium sulfate 4. Add solid (NH4)2SO4 slowly in four approximately equal amounts to the homogenates while they are still in the blender, homogenizing for 1 min between each addition, to yield a final 20% saturated solution (113 g/liter). Continue homogenization for another 30 sec at full speed after the final addition. For brain tissue, repeat this procedure with the other two tissue homogenates. 5. Combine the ammonium sulfate–treated homogenates in a beaker on ice and stir the mixture for 20 to 30 min. Centrifuge 30 min at 14,000 × g, 4°C. Carefully decant the supernatants and discard the pellets. Retain a small aliquot of this material for determining protein concentration by the biuret procedure (Basic Protocol 2) and assaying GDH activity (Basic Protocol 3). Purification of Glutamate Dehydrogenase from Liver and Brain
6. Add 188 g/liter of solid (NH4)2SO4 slowly with continuous stirring. Stir the mixture for an additional 20 to 30 min, then centrifuge again as described in step 5. This results in 50% saturation with ammonium sulfate.
1.4.4 Supplement 29
Current Protocols in Protein Science
7. Decant and discard the supernatants. Resuspend the pellets in 20 mM sodium phosphate buffer, pH 7.4, using a Dounce homogenizer, to yield a total volume of 200 to 250 ml (160 ml for liver). Retain a small aliquot of this material for determining protein concentration by the biuret procedure (Basic Protocol 2) and assaying GDH activity (Basic Protocol 3). The ammonium sulfate pellet can be resuspended by careful stirring if a Dounce homogenizer is not available.
8. Dialyze (UNIT 4.4 and APPENDIX 3B) for 18 hr against 20 volumes (v/v) of cold 20 mM sodium phosphate buffer, pH 7.4, with at least two changes of buffer, using an MWCO 12,000 dialysis membrane. 9. Centrifuge the material from the dialysis bag 30 min at 40,000 × g, 4°C, and retain the supernatants. Resuspend the pellets in the 20 mM sodium phosphate buffer, pH 7.4, and centrifuge again for 30 min at 40,000 × g, 4°C. Combine the supernatants with those from the first centrifugation and discard the pellets. Retain a small aliquot of this material for determining protein concentration by the biuret procedure (Basic Protocol 2) and assaying GDH activity (Basic Protocol 3). Extraction by resuspension and recentrifugation of the pellet after dialysis and centrifugation increases the yield of enzyme, since otherwise some glutamate dehydrogenase activity is lost in the pellet. As is common with salt precipitation in enzyme purification, this step does not give a high degree of purification of glutamate dehydrogenase. However, it is an effective way of reducing the volume of material for the subsequent steps, and it also removes some contaminants.
Perform chromatography on DEAE-cellulose 10. Pour the supernatant carefully onto the top surface of a 40 × 3–cm chromatography column packed to 30 cm with DEAE-cellulose, equilibrated with 20 mM sodium phosphate buffer, pH 7.4 (Support Protocol 5). After the protein solution has run into the column, continue to pass buffer through it and determine the absorbance of aliquots of the effluent at 280 nm (A280). Continue washing the column until the A280 of the effluent is <0.1. 11. Pass a 2-liter linear gradient from 20 to 200 mM sodium phosphate buffer, pH 7.4, through the column. Collect fractions (∼10 ml) and determine their A280 (see Basic Protocol 2), conductivity, and enzyme activities (see Basic Protocol 3). Pool the fractions containing glutamate dehydrogenase activity. Retain a small aliquot of this pooled material for determining protein concentration by the biuret procedure (Basic Protocol 2) and assaying GDH activity (Basic Protocol 3). The combined active fractions require concentration before the next step; this can be done either by ultrafiltration or ammonium sulfate precipitation. The authors have used both of these procedures and they work equally well.
12a. Concentrate protein by ultrafiltration: Concentrate the combined fractions containing GDH activity in an Amicon ultrafiltration cell using either an Ultracel Amicon YM 30 or Ultracel PLTK membrane, as described in Support Protocol 6, until the volume remaining in the cell is reduced to ∼20 ml. 12b. Concentrate protein by ammonium sulfate precipitation: Add 312 g/liter solid (NH4)2SO4 slowly, over a period of 30 min with continuous stirring (this brings the saturation to 50%). Continue stirring for a further 20 min, then centrifuge 20 min at 14,000 × g, 4°C. Discard the supernatant and resuspend the pellet in ∼20 ml of 20 mM sodium phosphate buffer, pH 7.4.
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Perform chromatography on GTP-Sepharose 13. Desalt the sample by passing the concentrated enzyme solution through a 30 × 2.5–cm chromatography column packed to 22 cm with Sephadex G-25, equilibrated with affinity chromatography buffer. The next chromatographic steps should be performed as quickly as possible, since the enzyme is slightly unstable in the affinity chromatography buffer.
14. Apply the enzyme solution to a 6 × 2–cm chromatography column packed to 4 cm with hydrazide-Sepharose 4B equilibrated with affinity chromatography buffer. Elute by passing an additional 50 ml of affinity chromatography buffer through the column. Glutamate dehydrogenase activity is not appreciably retarded by this material, and essentially complete recovery of the applied activity is obtained when the applied volume plus an additional 50 ml of buffer has been passed through the column. This procedure removes any components which bind nonspecifically to the Sepharose or to the spacer arm.
15. Apply the enzyme solution eluted from the above column to a 6 × 2–cm chromatography column packed to 4 cm with GTP-Sepharose 4B, equilibrated with affinity chromatography buffer. Use a flow rate of not more than ∼1 ml/min. Pass affinity chromatography buffer through the column until the A280 of the effluent has fallen to <0.01 (∼150 ml of buffer should be required). Once the enzyme solution has passed into the column the flow rate may be increased to 1.5 ml/min, and this flow rate should be used for step 16.
16. Elute the enzyme activity with a linear 400-ml gradient from 0 to 400 mM KCl in affinity chromatography buffer. Collect fractions (∼10 ml) and monitor the A280 (see Basic Protocol 2), conductivity, and enzyme activity (see Basic Protocol 3) in each. 17. Combine fractions containing glutamate dehydrogenase activity and concentrate by ultrafiltration, followed by osmotic dehydration, as described in Support Protocol 6, to yield a protein concentration of at least 3 mg/ml. 18. Dialyze (UNIT 4.4 and APPENDIX 3B) overnight against 3 liters of 20 mM sodium phosphate buffer, pH 7.4, using MWCO 12,000 dialysis membrane. Add glycerol to the dialyzed enzyme solution to yield a final concentration of 30% (v/v) and store in a sealed container, which has been flushed with N2, at 4°C. There may be some small variations between different preparations of the GTP-Sepharose in the sharpness of the peak of activity that emerges and in the salt concentration necessary to elute the enzyme. This may reflect varying concentrations of bound ligand, but does not affect the degree of purification obtained. SUPPORT PROTOCOL 1
PREPARATION OF GTP-SEPHAROSE Although GTP-agarose is available from Sigma, the preparation described in this protocol generally yields more satisfactory and reproducible results than commercial preparations. The steps involved are outlined in Figure 1.4.2. GTP is coupled to Sepharose 4B by the method of Jackson et al. (1973) as modified by Godinot et al. (1974), where L-glutamic acid γ-methyl ester is used as the spacer instead of ε-aminohexanoic acid methyl ester. After the spacer is coupled to the activated Sepharose and the ester group is treated with hydrazine (Godinot et al., 1974), a portion of the gel (∼10 ml) is retained and used as the hydrazide gel in the purification procedure.
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OH OH Sepharose CNBr
O C O
NH CH2OPOPOP
HO L-glutamic acid γ -methyl ester
COOH
GTP
O
NHCHCH2 CH2COCH3 OH
G
NaIO4
H2NNH2 COOH
O
HO
CH2OPOPOP
O
O
NHCHCH2 CH2CNHNH2 OH hydrazine derivative
O
O G
CH2OPOPOP
COOH
O
NHCHCH2 CH2CNHN OH GTP-Sepharose
O G
Figure 1.4.2 The chemical processes involved in the synthesis of GTP-Sepharose according to the method described in Support Protocol 1. The activation of Sepharose by cyanogen bromide (Support Protocol 3) is also shown in simplified form; alternative activated products are formed in this step.
Materials Cyanogen bromide–activated Sepharose 4B (Pharmacia Biotech, or see Support Protocol 3). Sodium bicarbonate buffer: prepare 0.1 M NaHCO3 and adjust pH to 9.0 with NaOH L-glutamic acid γ-methyl ester Hydrazine hydrate Guanosine 5′-triphosphate (GTP) 0.1 M disodium hydrogen phosphate adjusted to pH 5.0 with 1 M citric acid Sodium periodate Ethylene glycol
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Nitrogen source 50 mM sodium phosphate buffer, pH 6.8 (APPENDIX 2E) containing 5 mM EDTA Butanol Sintered-glass filter funnel (Pyrex, porosity G2) 70°C water bath Prepare the hydrazide gel 1. Wash ∼4.5 g of cyanogen bromide–activated Sepharose 4B in a sintered-glass filter funnel with ice-cold water, then wash with 100 ml ice-cold sodium bicarbonate buffer. Finally, suspend the resin in sodium bicarbonate buffer to a volume of 16 ml. The cyanogen bromide–activated Sepharose 4B contains dextran and lactose as stabilizers. It is necessary to remove these by washing. It is possible to activate Sepharose with cyanogen bromide in the laboratory as described in Support Protocol 3. This is a less expensive alternative, but cyanogen bromide is toxic and should be handled with great care.
2. Dissolve 1.12 g of L-glutamic acid γ-methyl ester in 4 ml sodium bicarbonate buffer. Add this solution to the ∼16 ml of the gel suspension and stir this mixture for 16 to 18 hr at 4°C. Finally, filter the mixture and wash it with cold water in a sintered-glass filter funnel before suspending it in water to a volume of 30 ml. The following steps should be performed in a fume hood.
3. Add 28 ml of hydrazine hydrate (99% to 100%) slowly to 30 ml of the suspended gel and heat the reaction mixture to 70°C in a water bath for 15 min. Allow the mixture to cool and then filter and wash with cold water on a sintered-glass filter. This material, the hydrazide gel, may be overlaid with butanol to impede bacterial growth, and stored at 4°C. Retain a 10-ml portion of this hydrazide gel for use in the purification procedure. After use, the hydrazide gel can be recycled as described in Support Protocol 3.
Prepare GTP-Sepharose 4. Dissolve 14.2 mg of GTP in 5 ml of 0.1 M disodium hydrogen phosphate that has been adjusted to pH 5.0 with 1 M citric acid. Dissolve 8.6 mg of sodium periodate in 0.4 ml of the same buffer. Mix these two solutions and incubate in the dark for 30 min at room temperature. Add 8 µl of ethylene glycol to remove the excess periodate, and incubate the mixture for a further 15 min. Bubble nitrogen through the solution for 5 to 10 min. Bubbling with N2 is effected by connecting a Pasteur pipet to a nitrogen cylinder by plastic tubing. The Pasteur pipet is placed in the tube containing the reacted mixture and the nitrogen is allowed to bubble through slowly. This procedure removes any formaldehyde produced during the reaction, which would compete with the oxidized GTP for reaction with the hydrazide gel.
5. Dilute the mixture from step 4 (oxidized GTP solution) to 10 ml with 0.1 M disodium hydrogen phosphate that has been adjusted to pH 5.0 with 1 M citric acid. If necessary, remove the butanol layer from the hydrazide gel (from step 3) by aspiration and wash the gel with cold water, then suspend it in 20 ml of cold water. Mix 10 ml of this gel suspension with the oxidized GTP solution and stir the mixture at room temperature for 2 hr. Purification of Glutamate Dehydrogenase from Liver and Brain
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6. Filter the gel on a sintered-glass funnel and wash with at least 100 ml of cold 50 mM sodium phosphate buffer, pH 6.8, containing 5 mM EDTA. Retain washings if step 8 is to be performed. Store the resultant GTP-Sepharose at 4°C with an overlay of butanol to retard bacterial growth. Remove this butanol layer by aspiration and wash the gel with buffer prior to use of the gel. After use, the GTP-Sepharose gel can be recycled as described in Support Protocol 4.
ASSESSMENT OF INCORPORATION OF GTP INTO THE HYDRAZIDE GEL The following modification to Support Protocol 1 may be applied if there are any problems using the GTP-Sepharose medium prepared in Support Protocol 1 in the affinity purification steps (see Basic Protocol 1, steps 13 to 16). It is generally not necessary to resort to this procedure.
SUPPORT PROTOCOL 2
CAUTION: When working with radioactivity, take appropriate precautions to avoid contamination of the experimenter and the surroundings. Carry out the experiment and dispose of wastes in an appropriately designated area, following the guidelines provided by the local radiation safety officer (also see APPENDIX 2B). Additional Materials (also see Support Protocol 1) [8-14C]GTP (Amersham Pharmacia Biotech) 250:500:1 (v/v/w) Triton X-100/toluene/2,5-diphenyloxazole (PPO) Carry out Support Protocol 1 with the following modifications at the indicated steps. 4a. Add 2.7 µCi of [8-14C]GTP to the solution of unlabeled GTP before periodate oxidation. 7b. Determine radioactivity in a sample of the gel washings by scintillation counting in a mixture by scintillation counting in 250:500:1 Triton X-100/toluene/PPO. About 35% of the added GTP should be found to be tightly bound to the gel, corresponding to 0.6 ìmol of GTP per ml of settled gel (see Table 1.4.1).
Table 1.4.1 Recovery of [8-14C]GTP After Coupling to Sepharose 4B
Aqueous phase
Radioactivity (cpm)a,b
Total radioactivity (cpm)a,b
10 µM of initial 14C-GTP solution + 1 ml H2O 1 ml gel washings (from 475 ml) 10 µM of initial 14C-GTP solution + 1 ml gel washings
14760 5167 20100
3.77 × 106 2.44 × 106 —
a9 ml of 250:500:1 (v/v/w) Triton X-100:toluene:PPO were mixed with 1 ml of the aqueous phase and the
radioactivity determined in a liquid scintillation counter. bSince the cpm obtained with the 10 µl 14C-GTP solution + 1 ml gel washings equals the sum obtained for the
two solutions separately, there is no significant difference in the quenching for the two solutions and therefore it is unnecessary to convert the results to dpm.
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SUPPORT PROTOCOL 3
ACTIVATION OF SEPHAROSE 4B WITH CYANOGEN BROMIDE This protocol is given as an alternative to the purchase of preprepared cyanogen-bromideactivated Sepharose. Materials Sepharose 4B (Pharmacia Biotech) Cyanogen bromide Acetonitrile 5 M sodium hydroxide Sodium bicarbonate buffer: prepare 0.1 M NaHCO3 and adjust pH to 9.0 with NaOH CAUTION: Because cyanogen bromide is very toxic, it should be handled with extreme care. See APPENDIX 2A for general precautions. 1. Wash 10 ml Sepharose 4B with water and suspend in 40 ml water. 2. Stir this mixture slowly in an ice-bath in a fume-hood. Add 2.4 g cyanogen bromide dissolved in 1.7 ml acetonitrile and monitor the pH of the resultant mixture constantly, maintaining it at ≥pH 11 by the addition of 5 M sodium hydroxide. Use extreme caution as hydrocyanic acid may be formed in this step.
3. After hydrogen ion release has subsided (i.e., the pH ceases to fall), filter the gel on a sintered-glass funnel. Wash with at least 100 ml of ice-cold sodium bicarbonate and resuspend in 16 ml of that buffer. SUPPORT PROTOCOL 4
REUSE OF CHROMATOGRAPHY MEDIA Hydrazide-Sepharose (prepared as in Support Protocol 1, steps 1 to 3) Wash the resins after use with 500 ml of 2 M KCl, followed by 1 liter of water. Store at 4°C with an overlay of butanol, to retard bacterial growth. Remove the butanol layer by aspiration and equilibrate with affinity chromatography buffer (see recipe) immediately before use. GTP Sepharose (prepared as in Support Protocol 1, steps 1 to 7) Follow the same procedure as described for the hydrazide-Sepharose, above. Over a period of several months, the properties of the GTP-Sepharose change and the enzyme activity is eluted earlier in the gradient, probably as a result of instability of the bound GTP. The gel should therefore be discarded after 6 months. DEAE-Cellulose The manufacturers provide full instructions for recycling this material for reuse. After use, remove tightly bound material from the ion-exchange material by passing 2 column volumes of 2 M NaCl through the column. Equilibrate the column with the buffer to be used and then pass 1 column volume of the same buffer containing 0.02% sodium azide through the column. Before use, wash the column with azide-free buffer.
Purification of Glutamate Dehydrogenase from Liver and Brain
If a small portion at the top of the column becomes discolored as a result of tightly bound material, that portion may be removed and replaced by fresh DEAE-cellulose. Removal of the top few centimeters from the column may also be effective if the flow rate through the column becomes excessively slow. After removing the discolored top resin, transfer the remaining resin from the column to a beaker. Add the appropriate amount of fresh
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DEAE-cellulose from which the fines have been removed and which has been equilibrated with 20 mM sodium phosphate buffer, pH 7.4 (see Support Protocol 5), and repack the column. If the characteristics of the DEAE-cellulose change after protracted use, it may be regenerated by the following procedure. 1. Empty the contents of the column into a beaker. Stir with 15 volumes of 0.5 M HCl for 30 min. Stirring can be continued for up to, but no longer than, 2 hr if convenient.
2. Filter the DEAE-cellulose on a sintered-glass funnel and wash it with water until the pH of the effluent reaches 4. 3. Stir the DEAE-cellulose with 15 volumes of 0.5 N NaOH for 30 min. 4. Allow to settle, remove the supernatant by decantation, and repeat step 3. 5. Filter on a sintered glass funnel and wash with water until the pH of the effluent is close to neutrality. The DEAE-cellulose is then ready for re-equilibration and reuse, although it may be necessary to remove the fines (see Support Protocol 5). For longer-term storage, remove the DEAE-cellulose from the column and store it in a saturated solution of NaCl, in a tightly-sealed container. Sephadex G-25 Sephadex G-25 is supplied in dry form and must be swollen before use. Full instructions for swelling and recycling this material are provided in the literature from the manufacturers. EQUILIBRATION OF CHROMATOGRAPHIC MEDIA The chromatographic medium must never be allowed to dry. When using it in columns there should always be a layer of the appropriate buffer above the surface.
SUPPORT PROTOCOL 5
Equilibration of DEAE-Cellulose DEAE-cellulose (Whatman DE-52) may contain small amounts of relatively small-sized particles (“fines”) which can slow down the rate of flow through a column if not removed. The following steps are appropriate for the DEAE-cellulose chromatography performed in Basic Protocol 1 and the Alternate Protocol. For the DEAE-cellulose chromatography performed in Support Protocol 7 (i.e., step 15 of that protocol), the procedure is the same as that described below except that 1 M NH4HCO3 may be used for removing the fines. A total of 10 column volumes of 1 M NH4HCO3 should then be used to equilibrate the column, which is then washed with 8 liters of water. Materials DEAE-cellulose (Whatman DE-52) 200 mM and 20 mM sodium phosphate buffer, pH 7.4 (APPENDIX 2E) Chromatography column; 40 × 3–cm (length × diameter) Conductivity meter (e.g., Linton, type WPA CMD80) 1. Suspend the DEAE-cellulose resin in 5 volumes of 200 mM sodium phosphate buffer, pH 7.4. Allow the mixture to settle and remove the buffer and fines by decantation. Repeat this process at least three times, after which the supernatant, upon settling, should be clear.
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2. Pour a suspension of this material into a 40 × 3–cm chromatography column to produce a final bed volume of 30 × 3 cm. Allow 20 mM sodium phosphate buffer, pH 7.4, to flow through the column until the effluent has the same pH and conductivity as the in-flowing buffer. The column is now ready for use.
Equilibration of Dowex AG 1X2 Dowex AG 1X2 medium is used in Support Protocol 7. Suspend the Dowex AG 1X2 (200 to 400 mesh, Cl− form) in 3 M HCl and pour into a column of 4 cm diameter to give a settled bed volume of 10 cm. Wash the column before use with 1 liter of 3 M HCl, followed by exhaustive washing with at least 20 liters of water, until the pH of the effluent reaches neutrality. Equilibration of Sephadex G-25 Pass at least 2.5 column volumes of the appropriate buffer through the packed column before use. SUPPORT PROTOCOL 6
CONCENTRATION OF PROTEIN-CONTAINING SOLUTIONS Some of the purification procedures used in Basic Protocol 1 and the Alternate Protocol result in some degree of concentration. Precipitation with ammonium sulfate followed by resuspension of the pellet in a smaller volume, as well as ion-exchange chromatography, can also be effective concentration procedures. In contrast, however, dialysis and gel filtration tend to dilute samples. The procedures described below are specifically for concentration purposes. For additional information, see UNIT 4.4. It is important to note that in both of the procedures described below, inattention may result in the concentration proceeding until all the fluid has been removed. It is essential to monitor the progress of the procedure and stop when the desired volume has been reached, because attempts to recover enzyme activity from material that has been taken to dryness by either of these procedures will result in significant losses. Ultrafiltration The Amicon ultrafiltration cell uses pressure of nitrogen gas to force the solution through a membrane that retains molecules above a pre-determined size. Prepare the apparatus according to the manufacturer’s instructions using the ultrafiltration cell from Amicon either an Amicon Ultracel YM 30 or Ultracel PLTK membrane. Slowly stir the contents during the concentration process, which is carried out at 4°C. Continue until the required volume remains in the cell. Earlier work used Amicon Diaflo XM-50 membranes for this procedure, but these are no longer readily available. The Ultracel Amicon YM 30 or Ultracel PLTK membranes are appropriate substitutes.
Purification of Glutamate Dehydrogenase from Liver and Brain
Centriplus concentrators (Amicon) may be an alternative for concentrating the enzyme from volumes of less than 60 to 70 ml. They rely on centrifugation to force solvent through a membrane (e.g., Centriplus 50, MWCO 50 kDa). The sample containing enzyme is retained by the membrane. The authors have found this procedure to be successful with several other dehydrogenases.
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Osmotic Dehydration Although ultrafiltration is a highly effective procedure for concentrating relatively large volumes, it may be less effective and convenient with small volumes and high protein concentrations. Osmotic dialysis relies on the use of a high concentration of material outside a dialysis tube to draw water and small-molecule solutes from its contents by osmosis, leaving a more concentrated protein solution within the dialysis tubing. Either sucrose or polyethylene glycol (PEG; Mr = 15,000 to 20,000) may be used. However the purity of PEG can be variable and it is important to use preparations that are as pure as possible to avoid contamination of the enzyme solution from low-Mr impurities that may be inhibitory. Put the enzyme solution in a length of dialysis tubing (see APPENDIX 3B for preparation) that has been knotted at the lower end, and then tie a knot in the other end of the tubing. Surround the resulting tube with solid sucrose or PEG. This may be done by heaping the sucrose or PEG on a sheet of cooking foil, embedding the dialysis tubing in the sucrose, and then folding the foil to form a parcel around the sucrose or PEG-covered dialysis tube. Alternatively, the dialysis tubing may be buried in the material within a plastic box of appropriate dimensions. Leave at 4°C until the solution in the dialysis bag has reached the required volume; this will take several hours. Rinse the outside of the dialysis tube with water or buffer to remove adhering sucrose or PEG. This also dampens the dialysis membrane, which becomes brittle during the concentration procedure. Whereas with ordinary dialysis it is necessary to leave an empty space in the tubing to allow for the osmotic volume increase, this need not be done for osmotic dehydration where a decrease in volume occurs. In Basic Protocol 1 and the Alternate Protocol, this concentration procedure is followed by dialysis. In order to avoid too great a degree of dilution occurring at this stage, reknot the dialysis tube to confine the enzyme solution in a smaller volume of the tubing. DETERMINATION OF PROTEIN CONCENTRATION The first two methods presented under this heading are the Biuret and Bio-Rad protein assays. A spectrophotometer and 1-cm light-path cuvettes (either glass or plastic) will be required for each of these determinations. In the case of the third method presented below, determination of protein concentration by absorbance at 280 nm (A280), the cuvettes must be quartz or some other UV-transmitting material.
BASIC PROTOCOL 2
Other procedures for determining protein concentration may also be used, including the Lowry-Folin method, which is described in adequate detail in Dawson et al. (1989). UNIT 3.4 also provides additional information on determination of protein concentration. Biuret Protein Assay Materials Biuret reagent (see recipe). 1.5% (w/v) sodium deoxycholate (1.5 g sodium deoxycholate in 100 ml distilled H2O) 50 mg/ml bovine serum albumin (BSA), or alternative protein calibration standard 1. Prepare a series of standards in triplicate according to the scheme described in Table 1.4.2.
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Table 1.4.2
Preparation of Standard Curve for Biuret Protein Assay
mg protein
50 mg/ml BSA (µl)
1.5% (w/v) sodium deoxycholate (µl)
H2O (µl)
Biuret reagenta (ml)
0 0.25 0.5 0.75 1.0 1.5 2.0
0 5 10 15 20 30 40
100 100 100 100 100 100 100
900 895 890 885 880 870 860
4 4 4 4 4 4 4
aSee recipe in Reagents and Solutions.
2. For enzyme protein determination use 20 to 100 µl of appropriately diluted sample and prepare as described in Table 1.4.2. 3. Mix each sample and allow the reaction to proceed at room temperature for ∼30 min. 4. Measure absorbance at 540 nm using the samples containing no protein as blanks. 5. Prepare a standard curve by plotting absorbance at 540 nm against the protein concentration. 6. Use the standard curve to determine the protein concentration of the enzyme sample from its absorbance. Determine the absorbance of the sample against a blank containing the same medium. Dilute, if necessary, to ensure that the absorbance is <1.0. Bio-Rad Protein Assay Materials Bio-Rad Dye Reagent Concentrate 0.1 mg/ml bovine serum albumin (BSA), or alternative protein calibration standard (this can be conveniently prepared by diluting some of the stock solution prepared for the biuret assay; see above) Method 1. Prepare a series of standards in triplicate according to the scheme described in Table 1.4.3. 2. For enzyme protein determination use 20 to 100 µl of appropriately diluted sample and prepare as described in Table 1.4.3.
Purification of Glutamate Dehydrogenase from Liver and Brain
Table 1.4.3
Preparation of Standard Curve for Bio-Rad Protein Assay
µg protein
0.1 mg/ml BSA (µl)
H2O (µl)
Bio-Rad Dye Reagent Concentrate (µl)
0 2 5 10 15 20
0 20 50 100 150 200
800 780 750 700 650 600
200 200 200 200 200 200
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3. After mixing, leave for a period of 5 to 60 min, then determine the absorbance at 595 nm using the samples containing no protein as blanks. 4. Prepare a standard curve by plotting absorbance at 595 nm against the protein concentration. 5. Use the standard curve to determine the protein concentration of the enzyme sample from its absorbance. Determine the absorbance of the sample against a blank containing the same medium. Dilute, if necessary, to ensure that the absorbance is <1.0. Note that the Bio-Rad assay involves determination of the absorbance of 1 ml of mixture. The optical arrangements of some spectrophotometers may not permit this in standard 1-cm2 cross-section cuvettes. In such cases microcuvettes should be used. However, some older spectrophotometers may not focus the light beam to pass through the narrower path of a microcuvette. It is a good idea to check the instructions supplied with the spectrophotometer in regard to this. If all else fails, the volumes shown for this procedure can all be increased proportionally.
Absorbance at 280 nm Using quartz or other non-UV-absorbing cuvettes, determine the absorbance at 280 nm of the sample against a blank composed of the medium in which the sample is dissolved. A UV-Vis spectrophotometer is required. Use the rough approximation that a 1 mg/ml protein solution has an absorbance of 1.0 at this wavelength. ASSAY OF GLUTAMATE DEHYDROGENASE ACTIVITY Glutamate dehydrogenase catalyzes the reaction:
BASIC PROTOCOL 3
2-oxoglutarate + NH 3 + NADH + H + U L-glutamate + H 2 O + NAD +
The assay described here determines the decrease of absorbance at 340 nm as NADH is oxidized to NAD+ concomitant with the reductive amination of 2-oxoglutarate to L-glutamate. Materials Enzyme preparation (see appropriate steps of Basic Protocol 1 or Alternate Protocol) 0.4% (v/v) Triton X-100 in 50 mM sodium phosphate buffer, pH 7.4 100 mM potassium cyanide (KCN) 50 mM sodium phosphate buffer, pH 7.4 (APPENDIX 2E) 2.5 M ammonium sulfate in 50 mM sodium phosphate buffer, pH 7.4 4 mM NADH in 50 mM sodium phosphate buffer, pH 7.4 (prepared fresh on day of use) 0.25 M 2-oxoglutarate (α-ketoglutarate), in 50 mM sodium phosphate buffer, pH 7.4 Spectrophotometer with cuvette holder set at 37°C 1-cm path-length cuvettes (quartz or plastic with good light transmittance at 340 nm) NOTE: All GDH enzyme assays are performed at 37°C. In order to ensure rapid temperature equilibration, the assay buffer should be prewarmed to that temperature.
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Other hints on the performance and validation of enzyme assays can be found in Tipton (2002). 1. For crude enzyme preparations (see Basic Protocol 1, step 1): Incubate 50 µl of the enzyme solution with 2.68 ml of 0.4% Triton X-100 in 50 mM sodium phosphate buffer, pH 7.4, and 30 µl of 100 mM KCN (which will give a final KCN concentration of 1 mM in the complete assay) for 2 min at 37°C before proceeding to step 2. For assaying the enzyme at later steps of the purification, begin with step 2. Triton X-100 serves to disrupt particulate material, and KCN decreases any contaminating NADH oxidase activity. It has been shown that neither Triton X-100 nor KCN affect the activity of purified GDH at these concentrations. These assays are performed in phosphate buffer. Do not attempt to perform them in Tris buffer since the enzyme is unstable in Tris and either no activity or a rapid decrease in activity may be observed.
2. Add reagents to the cuvettes in the order listed, in a final volume of 3 ml: 2.71 ml 50 mM sodium phosphate buffer, pH 7.4 60 µl 2.5 M ammonium sulfate (final concentration 50 mM) 120 µl 4 mM NADH (final concentration 160 µM) 50 µl enzyme preparation. 3. Cover the cuvette with Parafilm, mix the contents by inversion and incubate in the spectrophotometer for 3 min. Initiate the reaction by the addition of 60 µl of 0.25 M 2-oxoglutarate (final concentration 5 mM), invert to mix and allow the reaction to proceed for 10 min in the spectrophotometer. 4. Determine the initial (linear) rate of decrease in absorbance as NADH is oxidized to NAD+, subtracting any blank rate occurring before the addition of the 2-oxoglutarate (see Critical Parameters). The initial absorbance resulting from the NADH present will be ∼1.0, although impurities in the enzyme solution may add to this. The assay has been found to proceed in a linear fashion until the absorbance has decreased by 0.15 absorbance units. In all cases it will be necessary to dilute the enzyme preparation in order to obtain a rate that is sufficiently slow to measure with accuracy. See Critical Parameters for a detailed discussion of dilution considerations.
5. Calculate the specific activity. Since the molar absorbance coefficient of NADH in a 1-cm path-length cuvette is 6220 M-1cm-1, the production of 1 ìmol of glutamate, or the reductive amination of 1 ìmol 2-oxoglutarate in an assay volume of 3.0 ml will give an absorbance change of 2.073. Thus, for a 1-cm path-length cuvette, the specific activity of the enzyme (ìmol product formed.per min per mg protein) will be related to the rate of decrease in absorbance per min (∆A/min), which will be given by:
specific activity =
∆A /min assay volume (ml) × 6.22 protein (mg)
It is most common to express enzyme activities in these terms. 1 unit (U) of enzyme activity is defined as the amount catalyzing the production of 1 ìmol of product (or the disappearance of 1 ìmol of substrate) in 1 min under stated conditions. Purification of Glutamate Dehydrogenase from Liver and Brain
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PURIFICATION OF RAT OR OX LIVER GDH INVOLVING AFFINITY PRECIPITATION WITH bis-NAD+
ALTERNATE PROTOCOL
Affinity precipitation of enzymes was developed by Mosbach and co-workers (see Flygare et al., 1983; Larsson et al., 1984). The bifunctional ligand bis-NAD+ (N2, N2′-adipodihydrazido-bis-(N6-carbonyl-methyl)NAD+) comprises two molecules of NAD+ joined by a spacer arm (see Fig. 1.4.5). This separates the NAD+ groups by a sufficient distance (∼1.7 nm) to allow them to bind to the coenzyme-binding sites of two different molecules of glutamate dehydrogenase. Mammalian GDH is a polymer, so extensive cross-linking of the enzyme molecules occurs and hence they precipitate from solution, as illustrated in Figure 1.4.3. Since there are many oligomeric dehydrogenases
+ glutamate dehydrogenase
bis-NAD+
Figure 1.4.3 A diagrammatic view of the cross-linking of glutamate dehydrogenase hexamers by bis-NAD+, which results in the formation of an insoluble lattice. The “locking-on” in the presence of glutarate is not shown. Further details of the process are given in the Alternate Protocol.
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in the cell, this process might be expected to be somewhat nonspecific, causing the precipitation of a number of enzymes containing binding sites for NAD+. However, the procedure can be made specific for the precipitation of a selected dehydrogenase by the use of a compound that is a competitive inhibitor of the enzyme with respect to its other substrate. Thus, in the case of GDH, glutarate, which is a competitive inhibitor towards glutamate, is used. This strengthens the binding between the bis-NAD+ and glutamate dehydrogenase, allowing it to be used at concentrations below those that would be necessary to precipitate other dehydrogenases.
ox liver (46 g)
rat liver (10g)
homogenize for 1 min in 10 volumes (w/v) of 4 mM sodium phosphate buffer, pH 7.4, containing 0.5 mM EDTA and 0.1 mM PMSF
Stage 1 (homogenate) add (NH4)2 SO4 (113 g/liter), stir for 20 min
centrifuge and discard pellet add (NH4)2 SO4 (188 g/liter), stir for 20 min centrifuge and discard pellet dialyze against 20 mM sodium phosphate buffer, pH 7.4
Stage 2 [dialyzed (NH4)2 SO4 pellet] apply to DEAE-cellulose column
elute with 20 to 200 mM sodium phosphate, pH 7.4, gradient and concentrate to ~10 ml
Stage 3 (DEAE eluate)
affinity precipitate with the appropriate amount determined from pilot precipitation experiment (Support Protocol 9) of bis-NAD+ in the presence of 79 mM glutaric acid and keep on ice overnight
centrifuge discard supernatant redissolve pellet by stirring for 6 hr in 1 ml of buffer containing 0.6 mM NADH
Stage 4 (purified enzyme)
dialyze against 20 mM sodium phosphate buffer, pH 7.4 purified glutamate dehydrogenase
Purification of Glutamate Dehydrogenase from Liver and Brain
Figure 1.4.4 Outline scheme for the purification of ox and rat liver glutamate dehydrogenase according to the Alternate Protocol. Full details of the steps involved are given in the text.
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This behavior, which was termed the “locking-on” effect by O’Carra (see O’Carra et al., 1996; O’Flaherty et al., 1999; Mulcahy and O’Flaherty, 2001), was originally developed for increasing the strength of enzyme binding to an immobilized ligand. It depends on the analog of the substrate that is oxidized by NAD+ binding to the enzyme⋅NAD+ binary complex and thus displacing the coenzyme-binding equilibrium by ternary complex formation: +
NAD glutarate + ZZZZZX Z GDH < NAD+ YZZZZZ ZZZZZZ X Z GDH YZZZZ Z Z Z GDH< NAD < glutarate
This “locking-on” behavior occurs only if the oxidizable substrate, or a competitive inhibitor that is an analog of the oxidizable substrate, binds to the enzyme⋅NAD+ binary complex and not to the free enzyme. The basic concept is that, although several enzymes bind NAD+, only the GDH⋅NAD+ complex binds glutarate. The majority of dehydrogenases have ordered sequential kinetic mechanisms in which the coenzyme binds before the second substrate. To date, the technique has been found to work well for the purification of yeast alcohol dehydrogenase and lactate dehydrogenase (see Irwin and Tipton, 1996), and the synthesis of the corresponding bis-ATP analog has extended the application of bis-coenzymes to kinases (Beattie et al., 1987). The precipitated dehydrogenase can then be dissolved by incubation in the presence of NADH, which effectively competes with bis-NAD+ for binding to the enzyme. The procedure described here is that of Graham et al. (1985), as modified by Irwin and Tipton (1996). The steps involved are summarized in Figure 1.4.4. Additional Materials (also see Basic Protocol 1) Bis-NAD+ (see Support Protocol 7) 0.7 M glutaric acid, adjusted to pH 7.0 with NaOH Additional reagents and equipment for pilot-scale precipitation (see Support Protocol 9) NOTE: Unless indicated otherwise all steps are performed at 0° to 4°C. 1. Homogenize the sample and perform ammonium sulfate precipitation and DEAEcellulose chromatography steps (see Basic Protocol 1, steps 1 to 12), concentrating the eluate from the DEAE-cellulose column to ∼10 ml at step 12a or b. Although it is possible to perform affinity precipitation with preparations that have not been subjected to chromatography on DEAE-cellulose, the yield is much lower and the precipitated material is not completely pure.
2. Dialyze the enzyme preparation overnight at 4°C against 200 ml of sodium phosphate buffer, pH 7.4, with at least one change of buffer. Carry out a pilot-scale precipitation as described in Support Protocol 9 to determine the optimal amount of bis-NAD+ to use. Add that amount of bis-NAD+ to the remaining dialyzed enzyme preparation and add 0.7 M glutaric acid (preadjusted to pH 7) to a final concentration of 79 mM glutarate. Keep the mixture on ice overnight.
3. Centrifuge 15 min at 10,000 × g, 4°C. Redissolve the pellet by stirring for 6 hr in 1 ml of 20 mM sodium phosphate buffer, pH 7.4, containing 0.6 mM NADH. Dialyze the redissolved pellet overnight (using an MWCO 12,000 dialysis membrane) against 200 ml of sodium phosphate buffer, pH 7.4. Add glycerol and store as described in Basic Protocol 1, step 18.
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SUPPORT PROTOCOL 7
SYNTHESIS OF N2, N2′-ADIPODIHYDRAZIDO-bis(N6-CARBONYLMETHYL)NAD+ (bis-NAD) This reagent is used in the Alternate Protocol. The steps involved are outlined in Figure 1.4.5. Materials Iodoacetic acid (fresh) 2 M and 1 M lithium hydroxide (LiOH) NAD+ (98%, free acid) 2 M and 6 M HCl 96% (v/v) ethanol 0.24 M NaHCO3 1 M NaOH Nitrogen source Sodium dithionite 2 M Tris (base) Acetaldehyde, redistilled Yeast alcohol dehydrogenase (crystalline, Sigma-Aldrich or Roche Diagnostics) 5 mM CaCl2, pH 2.7, and 50 mM CaCl2, pH 2.0 2 M Ca(OH)2 Adipic acid dihydrazide dichloride (Sigma-Aldrich) 0.5 M 1-ethyl-3-(3-dimethylaminopropyl)-carbodiimide hydrochloride (EDC; Sigma-Aldrich) 0.25, 1, and 2 M ammonium bicarbonate (NH4HCO3) Sintered-glass funnel (≥10 cm diameter) Rotary evaporator Vacuum desiccator 75°C water bath 4 × 10–cm Dowex AG 1X2 anion-exchange column (Support Protocol 5) Gradient mixer (e.g., UNIT 8.2) 30 × 2.5–cm chromatography column packed with Whatman DE-52 resin and equilibrated (see Support Protocol 5) with 1 M NH4HCO3, pH 8.0 Additional reagents and equipment for resolution of NAD+ derivatives (see Support Protocol 8) Synthesize N(1)-carboxymethyl-NAD+ 1. Dissolve 9 g fresh iodoacetic acid in ∼10 ml of water, neutralize the solution with 2 M LiOH, and add 3 g of NAD+. Adjust the pH to 6.5 with 2 M HCl (the total volume should be ∼30 ml) and leave the solution in darkness for 7 days at room temperature (∼20°C), or alternatively, at 37°C for 2 days. Check the pH daily (or every 4 to 6 hr at the higher temperature) and readjust to 6.5 with 2 M HCl as required. The progress of the reaction should also be followed by TLC and/or HPLC (see Support Protocol 8). The solid iodoacetic acid should be white; if it is discolored as a result of decomposition it should not be used. Solutions of iodoacetic acid should be prepared fresh.
2. When reaction is complete, adjust pH to 3.0 with 6 M HCl and add 2 volumes of 96% (v/v) ethanol to produce a milky, pink-tinged suspension. Add a further 10 volumes of cold 96% ethanol to complete the precipitation. Purification of Glutamate Dehydrogenase from Liver and Brain
The water/ethanol ratio is important; using a wet vessel can give rise to the formation of a brown, sticky substance which is water-soluble.
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3. Filter the crude N(1)-carboxymethyl-NAD+ on a sintered funnel, wash with ethanol and diethyl ether, and dry in a rotary evaporator at 35°C under vacuum or by lyphilization (average yield 2.9 g). Store the product at −20°C under vacuum. Carry out rearrangement of N(1)-carboxymethyl-NAD+ to N6-carboxymethyl-NAD+ 4. Dissolve the crude N(1)-carboxymethyl-NAD+ in 0.24 M NaHCO3 (90 ml), which gives a pale orange solution, and adjust the pH to 8.5 with 1 M NaOH. 5. Deoxygenate the solution by bubbling N2 gas through it for 2 min, add 1.5 g of sodium dithionite, and leave the solution in the dark until maximum reduction is achieved. Monitor progress of reduction by taking samples, diluting them 1:50 or 1:100, and measuring the increase in absorbance at 340 nm (A340). 6. Terminate the reaction by stirring vigorously for 10 min to oxygenate the solution and then bubble N2 gas through it for 2 min. Adjust the pH to 11.5 with 1 M NaOH and leave in a water bath at 75°C to allow the rearrangement from the N(1)- to the N6-substituted derivative to occur (Dimroth rearrangement; see Engel, 1975). Monitor the rearrangement by HPLC or TLC (see Support Protocol 8). 7. Cool the reaction mixture to room temperature and add 6 ml of 2 M Tris (base) and 1.5 ml of redistilled acetaldehyde. Adjust the pH to 7.5 with 1 M HCl and add 8 to 24 mg (2500 to 7500 U) of yeast alcohol dehydrogenase. 8. Monitor the reaction at 340 nm as in step 5. When a minimum A340 is reached, add 1 volume of 96% ethanol and pour the milky flocculent precipitate into 10 volumes of vigorously stirred 96% ethanol. Leave for 30 min (or overnight, if desired) and collect the precipitate by filtration. Store the crude N6-carboxymethyl-NAD+ at −20°C in a vacuum desiccator for up to a month (yield ∼2.6 g). Purify the crude N6-carboxymethyl-NAD+ 9. Dissolve the crude powder (2.6 g, from step 7) in 30 ml water, adjust the pH to 8.0 with 1 M LiOH, and apply this solution to a 2.5 × 30–cm Dowex AG 1X2 column. See Support Protocol 5 for details of packing and equilibrating the Dowex column.
10. After applying the coenzyme solution, wash the column with 0.5 liter of water, followed by 1 liter of 5 mM CaCl2, pH 2.7, until the pH of the effluent is 2.8. Apply a 2-liter linear gradient from 5 mM CaCl2, pH 2.7, to 50 mM CaCl2, pH 2.0. Collect 10 to 20 ml fractions and monitor the absorbance at 260 nm. The composition of the fractions can be monitored by TLC (see Support Protocol 8).
11. Pool the fractions containing the N6 derivative, adjust the pH to 7.0 with 2 M Ca(OH)2, and concentrate to 5 to 10 ml by rotary evaporation at 40°C. 12. Precipitate with 96% ethanol as in step 2, above, and lyophilize. This gives a pale yellow compound (yield 0.82 g, 25%); ε = 19,300 M−1cm−1 at 266 nm.
Synthesis of bis-NAD+ 13. Dissolve 0.82 g of purified N6-carboxymethyl-NAD+ and 105 mg of adipic acid dihydrazide dihydrochloride in 20 ml water, to produce a brownish solution. 14. Adjust the pH to 4.6 with 1 M HCl and then add 0.5 M EDC at 0°C in 15- to 100-µl aliquots over a period of 35 min. Monitor the pH and readjust it to 4.6 before each addition. 15. Add water (2 liters) to dilute the solution 100-fold, adjust the pH to 8.0 with 2 M NH4HCO3, and apply the solution to a 30 × 2.5–cm DEAE-cellulose (DE-52) column equilibrated with 1 M NH4HCO3, pH 8.0 (Support Protocol 5). Wash the column with
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water until the A260 is <0.1 and apply a 2-liter linear gradient from 0 to 0.25 M NH4HCO3, pH 8.0. 16. Monitor the Rf values of the fractions by TLC (Support Protocol 8). The yield from pure N6-carboxymethyl-NAD+ is ∼14%. ε= 42,800 M−1 cm−1 at 266 nm.
17. Concentrate the product-containing fractions to ∼3 to 5 ml by rotary evaporation at 35°C and store at –20°C. For longer-term storage (>7 to 10 days) the material may be lyophilized and stored in a vacuum desiccator over silica gel at –20°C.
NH2
NH2
N
N
N
N
R
NAD+
ICH2COO–
N
pH 6.5, 25°C, 5 to 7 days
N
+ N
CH2COO–
N N(1) carboxymethyl-NAD+
R
Sodium dithionite pH 8.5, 2 to 4 hr NHCH2 COO–
N N
NH2 Dimroth rearrangement
N
pH 11.5, 75°C, 2 hr
N
RH N 6-carboxymethyl-NADH
alcohol dehydrogenase, acetaldehyde, pH 7.5 NHCH2 COO–
N N R
+ N
N
CH2COO–
N
N RH N(1) carboxymethyl-NADH
CONH2 + N H
O–
OH
H
CH2O P O P OCH2 O O O O H OH OH H N O H H H H OH OH N 6-carboxymethyl-NAD+
R=
N
H2NOC(CH2)4 CONH2 + EDC pH 4.6, 0°C, 35 min 0 HN
N N R
CH2
0
0
C NH NH C (CH2)4 C NH NH C CH2 NH
N
N
N
N
N
bis-NAD+
EDC= CH3CH2-N Purification of Glutamate Dehydrogenase from Liver and Brain
0
N
R
C N
(CH2)3 NH(CH2)2 Cl-
1-ethyl-3-(3)-dimethylaminopropyl) carbodiimide hydrochloride
Figure 1.4.5 Outline of the steps involved in the synthesis of bis-NAD+. Full details of the steps involved are given in Support Protocol 7.
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RESOLUTION OF NAD+ DERIVATIVES
SUPPORT PROTOCOL 8
Thin-layer Chromatography (TLC) The techniques of TLC are not described in detail here since they are adequately described in many textbooks (e.g., Wilson and Walker, 2000; Hahn-Deinstrop, 2000). Two different TLC solvent systems have been used for monitoring the products of the synthesis of bis-NAD+: System A: 60:100:2 (w/v/v) (NH4)2SO4/0.1 M potassium phosphate, pH 6.8/1-propanol. System B: 5:3 (v/v) isobutyric acid/1 M aqueous NH3, saturated with disodium EDTA. Apply the samples as spots to TLC plates (aluminium backed, silica gel 60, fluorescent indicator F254, layer thickness 0.2 mm; Merck), dry, and develop by ascending chromatography in a TLC developing tank. After developing the plates and allowing them to dry, the spots can be visualized by their fluorescence under UV light (wavelength 254 nm). They appear purple on a green background. The Rf values of bis-NAD+ and the compounds involved as intermediates in its synthesis are shown in Table 1.4.4. High-Performance Liquid Chromatography (HPLC) Inject the samples onto the HPLC column (4 mm × 30-cm C18 reversed-phase; e.g., Waters µBondapak C18) and elute isocratically with 0.1 M KH2PO4, pH 6.0, containing 10% methanol, at a flow rate of 0.7 to 1.0 ml/min. The elution profile is determined by continuously monitoring the absorbance of the eluate at 259 to 266 nm using a UV detector flow cell. Good separation between NAD+ and N(1)-carboxymethyl-NAD+ can be achieved, and N6-carboxymethyl-NAD+ and bis-NAD+ can be well separated, but this system does not resolve N(1)-carboxymethyl-NAD+ and N6-carboxymethyl-NAD+ satisfactorily. Retention times for NAD+ derivatives are given in Table 1.4.4. The retention times are decreased if buffer containing 20% methanol is used. A gradient of 0% to 20% methanol may be used to separate the N(1)- and N6-substituted NAD+ derivatives. The advantage of HPLC over TLC is that it is easier to quantify the levels of any impurities present in the product.
Table 1.4.4
Some Properties of bis-NAD+ and its Precursors
Rfa (TLC)
Compound
Retention timeb Molar absorbance (M-1cm-1) Solvent Ac Solvent Bd (HPLC)
NAD+ N(1)-carboxymethyl-NAD+ N6-carboxymethyl NAD+ Bis- NAD+
0.41 0.31 0.22 0.09
0.44 0.27 0.22 0.05
4.6 min 3.7 min 3.6 min 6.9 min
16,900 (at 259 nm) 17,100 (at 259 nm) 19,300 (at 266 nm) 42,800 (at 266 nm)
aThe R values given for TLC of coenzyme derivatives are not exact. They may vary by up to ±0.05. This may result from f
slight changes in the solvent composition over time, possibly as a result of evaporation. bThe HPLC retention times were obtained for samples chromatographed on a reversed-phase µBondapak C-18 column
(4 mm × 30 cm) and eluted with 0.1 M KH2PO4 containing 10% methanol. cSolvent A: 60:100:2 (w/v/v) (NH ) SO ;/0.1 M potassium phosphate, pH 6.8/1-propanol. 42 4 d5:3 (v/v) isobutyric acid/1 M aqueous NH , saturated with disodium EDTA. 3
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SUPPORT PROTOCOL 9
PILOT-SCALE AFFINITY PRECIPITATION STUDY In order to obtain optimal results from the purification procedure described in the Alternate Protocol, it is necessary to perform a small-scale (pilot) study to determine the optimum concentrations of protein and bis-NAD+ to use (see Fig. 1.4.6). This will vary somewhat between different enzyme preparations, owing to differences in enzyme concentration and the presence of other contaminants that may bind the bis-coenzyme. The optimum ratio of NAD+ equivalents/enzyme subunit may vary with the preparation used; for example, with a preparation of beef liver enzyme that had a specific activity of 2.7 U/mg (see Basic Protocol 3), about half the activity was found to precipitate at a ratio of 2 NAD+ equivalents/GDH subunit. However, a preparation with the lower specific activity of 0.8 U/mg required ∼8 NAD+ equivalents/GDH subunit to precipitate half the enzyme activity. The precipitation yield also depends on the protein concentration, as shown, for example, in Figure 1.4.7. Materials Sample of the enzyme preparation (Alternate Protocol) Bis-NAD+ (see Support Protocol 7) 700 mM glutaric acid, pH 7.0 Additional reagents and equipment for assaying protein concentration (see Basic Protocol 2) and GDH activity (see Basic Protocol 3) 1. Take an aliquot of the enzyme-containing solution from the DEAE-cellulose purification step (see Alternate Protocol and Basic Protocol 1) and assay it for GDH activity (see Basic Protocol 3) and protein concentration (see Basic Protocol 2) to determine its specific activity.
Precipitation (%)
100
50
0 0
5
10
15
20
[bis-NAD+ ] (mol/mol GDH subunit) Purification of Glutamate Dehydrogenase from Liver and Brain
Figure 1.4.6 An example of a pilot-scale affinity precipitation of ox liver glutamate dehydrogenase purified according to steps 1 to 3 of the Alternate Protocol. The experimental details are described in Support Protocol 9. The precise behavior will vary between preparations. The affinity precipitation is expressed as a percentage of the activity remaining in the control sample.
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Precipitation (%)
100
50
0
0
0.05 0.1 0.2 0.3 0.4
1
5
10
[Protein] (mg/ml)
Figure 1.4.7 An example of the effects of protein concentration on the pilot-scale affinity precipitation of ox liver glutamate dehydrogenase by bis-NAD+. These results are from a sample experiment performed as described in Support Protocol 9 in which the enzyme-containing sample was diluted to the indicated protein concentration before precipitation with a constant amount of bis-NAD+.
2. Calculate the approximate concentration of enzyme, based on the specific activity and subunit relative molecular mass. Assuming each subunit to have a relative molecular mass of 56,700 and the pure enzyme to have a specific activity of 40 ìmol/min/mg protein, this corresponds to ∼2267 mol product/min/mol enzyme subunit. Thus:
mol of subunit =
activity (µmol/min ) 2267 × 10 6
3. Measure effects of bis-NAD+ concentration as follows: a. Add a 100-µl sample of the enzyme preparation to each of 10 microcentrifuge tubes. b. To each tube, add 12.7 µl of 700 mM glutaric acid, pH 7.0, and varying amounts of bis-NAD+ within the range 0 to 10 coenzyme equivalents/enzyme subunit. c. Also prepare sample containing no bis-NAD+ (to which water is added in place of the coenzyme derivative) as well as a control tube that contains no bis-NAD+ or glutarate, so that the percentage inhibition resulting from the presence of the glutarate can be calculated. The concentration of bis-NAD+ can be calculated from the absorbance at 266 nm and the molar absorbance coefficient given in Table 1.4.4. A 1 M solution of bis-NAD+ will give an absorbance, in a 1-cm light-path cuvette, of 42,800, and each mole of bis-NAD+ contains 2 NAD+ equivalents. Thus:
2( A266 ) = coenzyme equivalent concentration in mol/liter 42,800
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4. Allow the tubes to stand at 4°C for at least 30 min (or overnight, if required), then microcentrifuge 5 min at maximum speed (∼13,000 × g), 4°C. 5. Assay the GDH activity remaining in the supernatant (see Basic Protocol 3) of each sample and express it as a percentage of the activity remaining in the control sample. The maximum affinity precipitation has occurred in the tube with the minimal residual activity and thus gives the appropriate concentration of bis-coenzyme to add to obtain maximum affinity precipitation. These experiments should be performed at least in duplicate.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Affinity chromatography buffer 100 mM Tris acetate 1 mM KH2PO4 0.1 mM EDTA Adjust pH to 7.15 at 4°C Store at 0° to 4°C up to 1 week; check pH before use and adjust if necessary Biuret reagent Dissolve 1.5 g copper sulfate (CuSO4⋅5H2O) and 6.0 g sodium potassium tartrate (C4H4KNaO6⋅4H2O) in 500 ml distilled H2O. Add 300 ml of 10% (w/v) sodium hydroxide and dilute the mixture to 1 liter with distilled water. Homogenization buffer 4 mM sodium phosphate buffer, pH 7.4 (APPENDIX 2A) 0.5 mM EDTA Store with above components at 0° to 4°C up to 1 week; check pH before use and adjust if necessary 0.1 mM phenymethanesulfonyl fluoride (PMSF; add immediately before use as described below) PMSF is unstable in aqueous solution and must be added to the buffer immediately before use. This is done by adding a suitable volume of a 100 mM PMSF solution in acetone to a small quantity (approximately one-tenth of the final volume) of boiling buffer, and immediately mixing this solution with the remainder of the ice-cold buffer.
COMMENTARY Background Information
Purification of Glutamate Dehydrogenase from Liver and Brain
There are several factors that must be considered in the choice of a purification procedure. The major factors are how much enzyme is needed for the planned studies, how long the procedure will take, and how much it will cost. To these should be added the question of whether the purified enzyme is different from the material that was in the tissues to start with. As discussed below, GDH purified by some older procedures has been shown to have undergone limited proteolysis, which alters its kinetic and regulatory behavior. Clearly, such preparations can give misleading information about the behavior of the enzyme.
Several alternative procedures have been used to purify GDH from different mammalian sources. The procedures described in this unit are preferable to the earlier procedures that involved precipitation with ethanol or acetone and/or detergent extraction followed by fractional precipitation with ammonium sulfate and/or sodium sulfate (see e.g., Fahien et al., 1969). Although that procedure resulted in a crystalline preparation of the enzyme, it has been shown that limited proteolysis results in a loss of a tetrapeptide from the amino-terminal end of the protein (McCarthy et al., 1980). This modification, which results in a slightly faster mobility on SDS (0.1% w/v) PAGE in 13%
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polyacrylamide gels (McCarthy et al., 1980), significantly affects the kinetic behavior of the enzyme (McCarthy and Tipton, 1984) and its in teraction s with allosteric effectors (McCarthy and Tipton, 1985; Couée and Tipton, 1989a) and with antipsychotic drugs such as perphenazine (Couée and Tipton, 1990a,b). An obvious alternative to the hard work of purifying an enzyme is to buy it from a commercial source. Purified preparations of ox (beef) liver GDH have been available from several commercial sources: BoehringerMann heim (n ow Ro ch e Diagn ostics, Mannheim), Calbiochem, PL Biochemicals (now Pharmacia P-L Biochemicals), Sigma, and United States Biochemical (USB), although some of these sources no longer supply purified preparations of GDH. However, the authors have found the enzyme from all these sources to have suffered the same, or more, proteolytic modification as that suffered by the enzyme prepared by the earlier procedures discussed above (see McCarthy et al., 1980). Cloning and expression of GDH is an alternative source of enzyme (Shashidharan et al., 1994; Plaitakis et al., 2000). An expression system has the advantage that it permits site-directed mutagenesis studies (see Fang et al., 2002; Zaganas and Plaitakis, 2002). It is also, of course, useful in cases where there are rather small quantities of enzyme in the chosen source material. This is not the case with the major isoform of GDH (GLUD1) in liver, but it is the case for the so-called “nerve tissue–specific” form (GLUD2) which is present in much lower abundance in the brain (see Zaganas and Plaitakis, 2002). It is still necessary to purify such an expressed enzyme if one needs to work with a homogeneous preparation. In some cases, this can be facilitated by the insertion of a tag into the gene to be expressed. Such tags range from oligohistidine tails to another protein, such as glutathione-S-transferase. It is common not to remove a tag after purification, and indeed in some cases it is not easy to do so. The assumption that the presence of such a tag will not affect the behavior of the expressed enzyme may sometimes be true, but it is certainly not always so (see e.g., Halliwell et al., 2001; Hiser et al., 2001; Ledent et al., 1997). In the case of GDH, as already discussed, modification at the amino-terminal end results in alterations in its behavior, and the C-terminal region appears to be involved in its allosteric regulation (Peterson and Smith, 1999). This suggests that expression of the protein with a tag at either end would be
likely to produce artefactual results. Indeed, it is necessary to use conventional procedures, involving a GTP affinity column, to purify the enzyme after cloning and expression (see Fang et al., 2002; Zaganas and Plaitakis, 2002). To assess the purity of the protein, polyacrylamide gel electrophoresis (PAGE) in the presence and absence of sodium dodecyl sulfate (SDS) remains the accepted standard for assessing protein purity (UNITS 10.2-10.4). The hexameric glutamate dehydrogenase from ox and some other mammalian species forms higher aggregates at higher protein concentrations (see, e.g., McCarthy et al.,1981). Although this process is reversible, with a dissociation constant of ∼1 mg/ml (Cohen et al., 1976), it hampers attempts to determine the behavior on nondenaturing PAGE. However PAGE in the presence of SDS reveals that the enzyme prepared as described here contains only a few minor impurities (see McCarthy et al., 1980).
Critical Parameters and Troubleshooting
A critical aspect of protein purification procedures is to have everything that will be needed prepared in advance. For example, it is a recipe for disaster to finish one step and then find that the column needed for the next has not yet been equilibrated. Many enzymes are relatively unstable and susceptible to degradation by contaminating peptidases. Generally, to avoid losses of enzyme activity, the procedure should be completed as rapidly as possible. The essence of any robust purification procedure is that any minor problems with one step are corrected by the others. Thus, for example, a somewhat lower degree purification on the DEAE-cellulose column may be compensated for in the affinity-chromatographic step. Some problems that are common to many purification procedures are listed below. Failure to use the highest purity reagents. Analytical grade chemicals should be used for such procedures as preparing buffers and ammonium sulfate precipitation. Less pure materials may contain compounds that inhibit enzyme activity. Working at the wrong temperature. Most of the procedures involved should be executed at 0° to 4°C. This is most easily accomplished in a cold room, although cooling the material in an ice bucket is satisfactory for some steps. Performing those steps at higher temperatures will result in enzyme inactivation. Overzealous stirring. This can cause frothing of protein solutions, which results in some
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Purification of Glutamate Dehydrogenase from Liver and Brain
denaturation. Stirring speeds should be fast enough to ensure adequate mixing, but no faster; if frothing does occur the stirring is too fast. Note that some frothing does occur during steps 1 and 2 of Basic Protocol 1, but this does not result in significant losses of GDH activity, probably because of the high protein concentrations at those stages. Failure to equilibrate chromatographic columns. Equilibration can be judged to have occurred when the pH and conductivity of the out-flowing buffer is the same as that of the in-flowing equilibration buffer. To use a column before complete equilibration has occurred will usually result in poor resolution of the sample components. Using a chromatographic column that is too old. The degradation of GTP-Sepharose after ∼6 months of storage is discussed in Support Protocol 4. DEAE-cellulose is quite stable if stored under the conditions described in Support Protocol 4. However, with time, the accumulation of tightly-bound material may affect the performance of columns, in which case the recycling procedure outlined in Support Protocol 4 should be used, or, alternatively, fresh DEAE cellulose can be used. Failure to check that the pH of buffers is correct. The pH should be adjusted at the temperature at which it will be used. Some buffers contain additional components, such as EDTA, and the pH should be checked and, if necessary, adjusted after all ingredients have been added. Failure to desalt samples before chromatographic steps. The presence of high concentrations of salt may prevent compounds of interest from binding to the column. In all cases where the enzyme is meant to bind to the column, it is important to retain the material that passes unretarded through the column and to assay it for enzyme activity as soon as possible. If material that should have bound passes straight through the column, it may be possible to rescue it by further desalting and application to a freshly equilibrated column. If in doubt about effective desalting, check the conductivity. The flow rate through the column is very slow, or there is apparently no flow at all. Do not load the enzyme-containing material onto a column that is running more slowly than specified. Very slow flow rates may occur if the material in the column has become compacted. This can arise if the column is run at too high a pressure. Always follow the manufacturers’ instructions about the maximum hydrostatic pressure that can be applied, and do not exceed that value. The accumulation of denatured material at the top of
a column can also slow the flow rate. This may be rectified by removing some of the chromatographic material from the top of the column, which is often identifiable by discoloration, adding an equivalent amount of fresh material, and repacking the column (see Support Protocol 4). In the case of DE-52 DEAE-cellulose, failure to remove the “fines” before using the resin (see Support Protocol 5) can result in inadequate flow rates. Flow rates through columns that are flowing well are likely to slow considerably if the sample applied contains appreciable amounts of particulate material. If particulate material is present, it should be removed by centrifugation, or, where appropriate, filtration, before the sample is applied to the column. Air bubbles or cracks are present in columns. This can result in the applied material not coming into adequate contact with the chromatographic material. Air bubbles can occur all too easily in gel-filtration media (e.g., Sephadex and Sepharose, including GTP-Sepharose) unless the manufacturer’s instructions for packing the material into columns are heeded. Cracks can occur if the material in the column is allowed to dry out. If that occurs, the only solution is to remove the material from the column, rehydrate and reequilibrate it, and then repack the column. The following subsections discuss only those steps where difficulties may be encountered through causes other than those discussed above. Should the enzyme lose activity or the final product have a specific activity significantly lower than expected, comparison of the data for each step with those shown in Tables 1.4.5 and 1.4.6 for Basic Protocol 1, or Tables 1.4.7 and 1.4.8 for the Alternate Protocol, should indicate where the problem occurred. Basic Protocol 1: purification of GDH involving affinity chromatography on GTP-Sepharose Ammonium sulfate precipitation. Too rapid addition of solid ammonium sulfate can result in high local concentrations of the salt, causing excess precipitation of material that does not all redissolve. In the present procedure, homogenization between additions minimizes this problem. If the solid ammonium sulfate salt contains lumps, break these up using a pestle and mortar before using it. It is possible to perform the precipitation by continuously stirring the enzyme solution in a beaker and adding the ammonium sulfate to it. If this approach is used, the salt should be added very slowly, over a period of 1 to 2 hr, to minimize the above problem. In either proce-
1.4.28 Supplement 29
Current Protocols in Protein Science
Table 1.4.5 Protocol 1
Purification of Glutamate Dehydrogenase From Ox Brain According to Basic
Stage (see Fig. 1.4.1)
Volume (ml)
Total protein Total activity (mg) (µmol/min)
1500 1100 250 600 90
75,000a 23,000a 6200a 320b 12.5b
1 2 3 4 5
2280 1150 925 775 500
Specific activity Purification (µmol/min/mg) (x-fold) 0.03 0.05 0.15 2.4 40
1 2 5 80 1300
Yield (%) 100 50 41 33 22
aProtein concentration determined by the biuret procedure. bProtein concentration determined from the absorbance at 280 nm.
Table 1.4.6 Purification of Glutamate Dehydrogenase From Ox Liver According to Basic Protocol 1
Stage (see Fig. 1.4.1) 1 2 3 4 5
Volume (ml)
Total protein Total activity (mg) (µmol/min)
500 450 160 500 90
6000a 2500a 880a 150b 25b
1800 15000 1410 1200 1000
Specific activity Purification (µmol/min/mg) (x-fold) 0.3 0.6 1.6 8.0 40
1 2 5 25 120
Yield (%) 100 85 75 67 55
aProtein concentration determined by the biuret procedure. bProtein concentration determined from the absorbance at 280 nm.
Table 1.4.7 Protocol 1
Purification of Glutamate Dehydrogenase From Ox Liver According to Alternate
Stage (see Fig. 1.4.4)
Volume (ml)
Total protein Total activity (mg)a (µmol/min)
370 75 8.4 1.2
8580 2010 139 9.4
1 2 3 4
1920 1640 408 376
Specific activity Purification (µmol/min/mg) (x-fold) 0.2 0.3 2.9 40
1 4 13 180
Yield (%) 100 86 21 20
aProtein concentration determined by the Lowry method.
Table 1.4.8 Purification of Glutamate Dehydrogenase From Rat Liver According to Alternate Protocol 1
Stage (see Fig. 1.4.4) 1 2 3 4
Volume (ml)
Total protein Total activity (mg)a (µmol/min)
90 35 9.8 1.0
2720 980 130 9.4
760 505 195 190
aProtein concentration determined by the Lowry method.
Specific activity Purification (µmol/min/mg) (x-fold) 0.3 0.5 1.5 37
1 2 5 140
Yield (%) 100 67 26 25
Strategies of Protein Purification and Characterization
1.4.29 Current Protocols in Protein Science
Supplement 29
dure, it is important to continue to stir for 20 to 30 min after the last ammonium sulfate addition, to ensure that any overprecipitated material has a chance to redissolve and that any material that should precipitate at that salt concentration does so. The use of a homogenizer during this stage has proven to be effective, but it is not recommended for ammonium sulfate fractionation of purer material, since excessive frothing can result in denaturation. Addition of ammonium sulfate will result in a decrease in pH. For GDH purification, it is not necessary to adjust the pH to the starting value between each addition of ammonium sulfate, although such readjustment is necessary for many other enzymes. Dialysis and concentration. Provided that the dialysis tubing is pretreated as described in UNIT 4.4 and APPENDIX 3B and that it has been tested for leaks, there should be no difficulties with these procedures. It should be stressed again that an air-free space should be left at the top of the knotted dialysis tube, because volume increases during dialysis can otherwise cause the bag to rupture. The concentration procedures should not cause difficulties if the directions given in Support Protocol 6 are followed. Chromatography on DEAE cellulose. Common difficulties with this step have been discussed in the general points at the beginning of this section. Chromatography on GTP-Sepharose. If the GDH enzyme does not bind to the column and the troubleshooting points at the beginning of this section have been taken into account, the likely causes are either that the affinity resin has degraded (see Support Protocol 4) or that there was a problem with its preparation. In first case, the GTP-Sepharose should be replaced by fresh material. If there has been a problem with its preparation, a fresh batch should be prepared using the procedure described in Support Protocol 1 to check for effective GTP coupling. Choice of reaction buffer. As indicated in Basic Protocol 1, the enzyme is unstable in the Tris/acetate buffer and losses of activity can occur if the enzyme is left in that buffer for extended periods.
Purification of Glutamate Dehydrogenase from Liver and Brain
Basic Protocol 3: assay of glutamate dehydrogenase activity In all cases, it will be necessary to dilute the enzyme preparation in order to obtain a rate that is sufficiently slow to measure with accuracy. The amount of dilution will depend on the state of purification of the enzyme, its protein concentation, and also the scale expansion used on
the spectrophotometer. For example, the crude homogenate of liver GDH has a specific activity of 0.3 µmol/min/mg protein (see Table 1.4.6). Since the production of 1 µmol of glutamate in the assay will give an absorbance change of 2.073 (see Basic Protocol 3, annotation to step 5) , 1 mg of this material will give an absorbance decrease, in 1 min, of 0.3 × 2.073 ≈ 0.62. The protein concentration of this crude homogenate is ∼12 mg/ml (see Table 1.4.6); therefore, 50 µl of this would contain 0.6 mg protein, and would be expected to give an absorbance change of ∼0.373 absorbance units per min. This would mean that the reaction would be finished in <3 min, which would be too fast for accurate initial-rate determinations. See, for example, Tipton (2002) for a fuller treatment of enzyme assays and initial-rate determinations. Dilution of this material 20× with the assay buffer before adding 50 µl to the assay would give an absorbance change of 0.01865 absorbance units per min, which would mean that it would take a little over 8 min for the absorbance to fall by 0.15 absorbance units. Similar calculations from the values in Table 1.4.6 for the purified liver GDH show that a dilution factor of 600 would be necessary so that a 50-µl volume added to the assay would give a rate similar to that given by the crude enzyme. The appropriate dilution factors for the other stages of purification and for the brain enzyme can be estimated from the date given in Tables 1.4.6 and 1.4.5, respectively. It should be noted that these numbers are only a rough guide, since the degree of purification and protein concentration may vary for individual steps of the purification. The addition of smaller volumes of the enzyme preparation to the assay will reduce the degree of dilution necessary. Alternate Protocol: purification of GDH involving affinity precipitation with bis-NAD+ The initial steps in this procedure are the same as those of Basic Protocol 1. Affinity precipitation. A pilot-scale precipitation, as described in Support Protocol 9, should always be carried out to determine the optimum concentration of bis-NAD+ to use. Figure 1.4.6 shows an example of a concentration-precipitation relationship, but it should be remembered that the optimum bis-NAD+ concentration may vary with protein concentration. Failure to obtain affinity precipitation could result from: 1. A fault in the synthesis of the bifunctional ligand. Monitoring the success of each step in the preparatory procedure using the pro-
1.4.30 Supplement 29
Current Protocols in Protein Science
cedures outlined in Support Protocol 8 and the data in Table 1.4.2, should reveal any problems in this respect and where they occur. 2. Attempting to precipitate from a solution containing too low a concentration of GDH (see Fig. 1.4.7). Further concentration of the enzymecontaining sample should deal with this problem. Stability The enzyme, stored as described in Basic Protocol 1, is stable for at least 6 months without significant loss of activity or alteration of its mobility on a 13% polyacrylamide gel in the presence SDS. This is important in view of the relatively large amounts of purified GDH that can be prepared. However, changes can occur in its responses to allosteric effectors when the enzyme is stored in an atmosphere of air (Couée and Tipton, 1991). These changes, which may only become apparent after 2 to 3 months, are prevented by storing the enzyme solution under an atmosphere of nitrogen, and tubes containing stored GDH preparations should be routinely flushed with N2 each time they are opened. Safety All personnel involved should have received training in safe laboratory procedures and in the use of radioactive materials if Support Protocol 2 (involving radiolabeled GTP) is to be used. National and local regulations for the disposal of such materials as radiochemical waste and or-
Anticipated Results Basic Protocol 1: purification of ox brain or liver glutamate dehydrogenase involving affinity chromatography on GTP-Sepharose Tables 1.4.5 and 1.4.6 summarize the results obtained from typical purifications of the ox brain and liver enzymes using this procedure.
A280 nm ( ) conductivity ( ) GDH Activity ( )
6
8 4 6 2
GDH activity (µmol/min/ml)
Conductivity (mS)
10
ganic solvents should be followed. Several of the compounds used in these procedures are highly toxic and should be handled with great care. Toxicity information on these compounds is available from the manufacturers (e.g., Sigma-Aldrich at https://www.sigma-aldrich. com) and should be consulted in advance. For the use of animal materials, precautions against infectious and transmissible diseases are necessary. It is advisable that personnel be immunized against such diseases where appropriate. Although the ox tissue samples that the authors have used recently have been certified as being BSEfree, the initial steps of the purification are still performed under biohazard containment, protection and disposal conditions. It is essential that those steps indicated in Support Protocols 1 and 3 be performed under an adequately functioning, biohazard-containment level II fume hood. Safety glasses should be worn, and, in connection with the TLC procedure in Support Protocol 8, these should provide protection from UV light.
4
0 50
100 Fraction number
150
Figure 1.4.8 Chromatography of ox liver glutamate dehydrogenase at step 3 of Basic Protocol 1. A typical elution profile is shown. However the exact behavior may vary between preparations. The fraction size was ∼10 ml. In the left-hand y-axis label, mS stands for millisiemens.
Strategies of Protein Purification and Characterization
1.4.31 Current Protocols in Protein Science
Supplement 29
A280 nm ( ) conductivity ( ) GDH Activity ( )
12
10
2.0
8 30
A280
1.5
1.0
20
10
0.5
Conductivity (mS)
6
4
2
Glutamate dehydrogenase activity (µ mol/min/ml)
2.5
0 15
30 Fraction number
45
Figure 1.4.9 A typical elution profile of affinity chromatography of ox liver glutamate dehydrogenase on GTP-Sepharose at step 4 of Basic Protocol 1. The exact chromatographic behavior may vary between preparations. The fraction size was ∼10 ml. In the label for the inset scale, mS stands for millisiemens.
Examples of the elution profiles from DEAE-cellulose and GTP-Sepharose are shown in Figures 1.4.8 and 1.4.9.
Time Considerations
Day 3: Gel filtration into the GTPSepharose buffer, passage through the hydrazide gel column, and affinity chromatography on GTP-Sepharose. This schedule excludes the time taken to prepare the GTP-Sepharose. This should take 3 days or less, starting from CNBr-activated Sepharose. However much of this time is taken up by incubations, such as the 16- to 18-hr reaction of L-glutamic acid γ-methyl ester with the activated gel (see Support Protocol 1). The affinity resin, once made, can be used several times.
Basic Protocol 1: purification of ox brain or liver glutamate dehydrogenase involving affinity chromatography on GTP-Sepharose The entire purification procedure can easily be performed in 3 days. Day 1: Extraction and ammonium sulfate precipitation, followed by overnight dialysis. Day 2: DEAE-cellulose fractionation and concentration of appropriate column fractions.
Alternate Protocol: purification of rat or ox liver GDH involving affinity precipitation with bis-NAD+ This is a somewhat quicker procedure than Basic Protocol 1 because it avoids the final column chromatography steps. However, it still extends into several days. The first two of these are the same as those for Basic Protocol 1, and the third is for the pilot-scale precipitation study and the affinity precipitation itself. The preparation of bis-NAD+ can be completed in 4 days, provided that the reaction
Alternate Protocol: purification of rat or ox liver GDH involving affinity precipitation with bis-NAD+ Tables 1.4.7 and 1.4.8 summarize the results obtained from typical purifications of the ox and rat liver enzymes using this procedure. The time estimates given below do not include the time involved in pouring and equilibrating the chromatographic columns, which should be done before the purification procedure is started.
Purification of Glutamate Dehydrogenase from Liver and Brain
1.4.32 Supplement 29
Current Protocols in Protein Science
between iodoacetate and NAD+ is performed at 37°C (see Support Protocol 7). Once prepared, the bis-NAD+ may be used for several GDH preparations, as well as for the purification of some other dehydrogenases (see Irwin and Tipton, 1996)
Literature Cited
Beattie. R.E., Buchanan, M.,. and Tipton K.F. 1987. The synthesis of N2, N2-adipodihydrazido-bis(N-carboxymethyl-ATP) and its use in the purification of phosphofructokinase. Biochem. Soc. Trans. 15:1043-1044. Cho, S.W., Lee, J., and Choi, S.Y. 1995. Two soluble forms of glutamate dehydrogenase isoproteins from bovine brain. Eur. J. Biochem. 233:340-346. Cohen, R.J., Jedziniak, J.A., and Benedek, G.B. 1976. The functional relationship between polymerization and catalytic activity of beef liver glutamate dehydrogenase. II. Experiment. J. Mol. Biol. 108:179-199. Couée, I. and Tipton, K.F. 1989a. Activation of glutamate dehydrogenase by L-leucine. Biochim. Biophys. Acta 995:97-101. Couée, I. and Tipton, K.F. 1989b. The effects of phospholipids on the activation of glutamate dehydrogenase by L-leucine. Biochem. J. 261:921-925. Couée, I. and Tipton, K.F. 1990a. The inhibition of glutamate dehydrogenase by some antipsychotic drugs. Biochem. Pharmacol. 39:827-832. Couée, I. and Tipton, K.F. 1990b. Inhibition of ox brain glutamate dehydrogenase by perphenazine Biochem. Pharmacol. 39:1167-1173. Couée, I. and Tipton, K.F. 1991. The sulphydryl groups of ox brain and liver glutamate dehydrogenase preparations and the effects of oxidation on their inhibitor sensitivities.. Neurochem. Res. 16:773-780. Dawson, R.M.C., Elliott, D.C., Elliott ,W.H., and Jones, K.M. 1989. Data for Biochemical Research, 3rd ed. Clarendon Press, Oxford. Engel, J.D. 1975. Mechanism of the Dimroth rearrangement in adenine. Biochem. Biophys. Res. Commun. 64:581 - 585. Fahien, L.A., Strmecki, M., and Smith, S. 1969. Studies of gluconeogenic mitochondrial enzymes. I. A new method of preparing bovine liver glutamate dehydrogenase and effects of purification methods on properties of the enzyme. Arch. Biochem. Biophys. 130:449-455. Fang, J., Hsu, B.Y., MacMullen, C.M., Poncz, M., Smith, T.J., and Stanley, C.A. 2002. Expression, purification and characterization of human glutamate dehydrogenase (GDH) allosteric regulatory mutations. Biochem J. 363:81-87. Flygare, S., Griffin, T., Larsson, P.O., and Mosbach, K. 1983. Affinity precipitation of dehydrogenases. Anal. Biochem. 133:409-416. Godinot, C, Julliard, J.H., and Gautheron, D.C. 1974. A rapid and efficient new method of purification of glutamate dehydrogenase by affinity
chromatography on GTP-sepharose. Anal. Biochem. 61:264-270. Graham, L.D., Griffin, T.O., Beattie, R..E., McCarthy, A..D., and Tipton, K.F. 1985. Purification of liver glutamate dehydrogenase by affinity precipitation and studies on its denaturation. Biochim. Biophys. Acta 828:266-269. Hahn-Deinstrop, E. 2000. Applied Thin-Layer Chromatography: Best Practice and Avoidance of Mistakes. Wiley-VCH, Weinheim, Germany. Halliwell, C.M., Morgan, G., Ou, C.P., and Cass, A.E. 2001. Introduction of a (poly)histidine tag in L-lactate dehydrogenase produces a mixture of active and inactive molecules. Anal. Biochem. 295:257-261. Hiser, C., Mills, D.A., Schall, M., and FergusonMiller, S. 2001. C-terminal truncation and histidine-tagging of cytochrome c oxidase subunit II reveals the native processing site, shows involvement of the C-terminus in cytochrome c binding, and improves the assay for proton pumping. Biochemistry 40:1606-1615. Irwin, J.A., and Tipton, K.F. 1996. Affinity precipitation methods. In Methods in Molecular Biology 59: Protein Purification Protocols (S. Doonan, ed.) pp. 217-238. Humana Press, Totowa, N.J. Jackson, R.J., Wolcott, R.M., and Shiota, T. 1973. The preparation of a modified GTP-sepharose derivative and its use in the purification of dihydroneopterin triphosphate synthetase, the first enzyme in folate biosynthesis. Biochem Biophys. Res. Commun. 51:428-435. Larsson, P.O., Flygare, S., and Mosbach, K. 1984. Affinity precipitation of dehydrogenases. Methods Enzymol. 104:364-349. Ledent, P., Duez, C., Vanhove, M., Lejeune, A., Fonze, E., Charlier, P., Rhazi-Filali, F., Thamm, I., Guillaume, G., Samyn, B., Devreese, B., Van Beeumen, J., Lamotte-Brasseur, J., and Frere J.M. 1997. Unexpected influence of a C-terminal-fused His-tag on the processing of an enzyme and on the kinetic and folding parameters. FEBS Lett. 413:194-196. Lee, W.K., Shin, S., Cho, S.S., and Park, J.S. 1999. Purification and characterization of glutamate dehydrogenase as another isoprotein binding to the membrane of rough endoplasmic reticulum. J. Cell. Biochem. 76:244-253. McCarthy, A.D. and Tipton, K.F. 1984. The effects of magnesium ions on the interactions of ox brain and liver glutamate dehydrogenase with ATP and GTP. Biochem. J. 220:853-855. McCarthy, A.D. and Tipton, K.F. 1985. Ox liver glutamate dehydrogenase: Comparison of the kinetic properties of native and proteolysed preparations. Biochem. J. 230:95-99. McCarthy, A.D., Walker, J.M., and Tipton, K.F. 1980. Purification of glutamate dehydrogenase from ox brain and liver: Evidence that commercially available preparations of the enzyme have suffered proteolytic cleavage. Biochem. J. 191:605-611.
Strategies of Protein Purification and Characterization
1.4.33 Current Protocols in Protein Science
Supplement 29
McCarthy, A.D., Johnson, P., and Tipton, K.F. 1981. Sedimentation properties of native and proteolysed preparations of ox glutamate dehydrogenase. Biochem. J. 199:235-236. McDaniel, H.G. 1995. Comparison of the primary structure of nuclear and mitochondrial glutamate dehydrogenase from bovine liver. Arch. Biochem. Biophys. 319:316-321. Mulcahy, P. and O’Flaherty, M. 2001. Kinetic locking-on strategy and auxiliary tactics for bioaffinity purification of NAD(P)(+)-dependent dehydrogenases. Anal. Biochem. 299:1-18. O’Carra, P., Griffin, T., O’Flaherty, M., Kelly, N., and Mulcahy, P. 1996. Further studies on the bioaffinity chromatography of NAD(+)-dependent dehydrogenases using the locking-on effect. Biochim. Biophys. Acta 1297:235-243. O’Flaherty, M., O’Carra, P., McMahon, M., and Mulcahy, P. 1999. A “stripping” ligand tactic for use with the kinetic locking-on strategy: Its use in the resolution and bioaffinity chromatographic purification of NAD(+)-dependent dehydrogenases. Protein Expr. Purif. 16:424-431.
Purification of Glutamate Dehydrogenase from Liver and Brain
Approach (R. Eisenthal and M.J. Danson, eds.) pp. 1-47. Oxford University Press, Oxford, U.K. Tipton, K.F. and Couée, I. 1988. Glutamate dehydrogenase. In Glutamine and Glutamate in Mammals. (E. Kvamme, ed.) pp.81-100. CRC Press, Baton Roca, Fla. Wilson, K. and Walker, J. (eds.) 2000. Principles and Techniques of Practical Biochemistry, 5th ed.. Cambridge University Press, Cambridge, U.K. Yoon, H.Y., Hwang, S.H., Lee, E.Y., Kim, T.U., Cho, E.H., and Cho, S.W. 2001. Effects of ADP on different inhibitory properties of brain glutamate dehydrogenase isoproteins by perphenazine. Biochimie 83:907-913. Zaganas, I. and Plaitakis, A. 2002. A single amino acid substitution (Gly456Ala) in the vicinity of the GTP binding domain of human housekeeping glutamate dehydrogenase markedly attenuates GTP inhibition and abolishes the enzyme’s cooperative behavior. J. Biol. Chem. 277:2642226428.
Key References
Peterson, P.E. and Smith, T.J. 1999. The structure of bovine glutamate dehydrogenase provides insights into the mechanism of allostery. Structure Fold. Des. 15:769-782.
Glutamate dehydrogenase
Plaitakis, A. and Zaganas, I. 2001. Regulation of human glutamate dehydrogenases, implications for glutamate, ammonia and energy metabolism in brain. J. Neurosci. Res. 66:899-908.
There have been few comprehensive reviews of glutamate dehydrogenase in recent years. The references cited above cover some fundamental aspects.
Plaitakis, A., Metaxari, M., and Shashidharan, P. 2000. Nerve tissue-specific (GLUD2) and housekeeping (GLUD1) human glutamate dehydrogenases are regulated by distinct allosteric mechanisms: Implications for biologic function. J. Neurochem. 75:1862-1869.
Affinity chromatography
Preiss, T., Hall, A.G., and Lightowlers, R.N. 1993. Identification of bovine glutamate dehydrogenase as an RNA-binding protein. J. Biol. Chem. 268:24523-24526.
Affinity precipitation
Rajas, F. and Rousset, B. 1993. A membrane-bound form of glutamate dehydrogenase possesses an ATP-dependent high-affinity microtubule-binding activity. Biochem. J. 295:447-455.
Describes the theory underlying affinity precipitation and reviews its applications.
Shashidharan, P., Michaelides, T.M., Robakis, N.K., Kretsovali, A., Papamatheakis, J., and Plaitakis, A. 1994. Novel human glutamate dehydrogenase expressed in neural and testicular tissues and encoded by an X-linked intronless gene. J. Biol. Chem. 269:16971-16976
Contributed by Martha Motherway and Keith F. Tipton Department of Biochemistry Trinity College Dublin, Ireland
Shashidharan, P., Clarke, D.D., Ahmed, N., Moschonas, N., and Plaitakis, A. 1997. Nerve tissue-specific human glutamate dehydrogenase that is thermolabile and highly regulated by ADP. J. Neurochem. 68:1804-1811.
Alun D. McCarthy GlaxoSmithKline Middlesex, United Kingdom
Smith, T.J., Peterson, P.E., Schmidt, T., Fang, J., and Stanley, C.A. 2001. Structures of bovine glutamate dehydrogenase complexes elucidate the mechanism of purine regulation. J. Mol. Biol. 307:707-720.
Plaitakis, A. and Zaganas, I. 2001. See above. Tipton, K.F. and Couée, I. 1988. See above.
Matejtschuk, P. (ed.) 1997. Affinity Separations: A Practical Approach. Oxford University Press, U.K. A collection of chapters on the theory and practice of different aspects of affinity chromatography.
Irwin, J.A. and K.F. Tipton, K.F. 1995. Affinity precipitation: A novel approach to protein purification. Essays Biochem. 29:137-156.
Ivan Couée Centre National de la Recherche Scientifique Rennes, France Jane Irwin University College Dublin Dublin, Ireland
Tipton K.F. 2002. Principles of enzyme assay and kinetic studies. In Enzyme Assays: A Practical
1.4.34 Supplement 29
Current Protocols in Protein Science
Overview of the Physical State of Proteins Within Cells The word protein comes from the Greek word proteios, meaning primary. And, indeed, proteins are of primary importance in the study of cell function. It is difficult to imagine a cellular function not linked with proteins. Almost all biochemical catalysis is carried out by protein enzymes. Proteins participate in gene regulation, transcription, and translation. Intracellular filaments give shape to a cell while extracellular proteins hold cells together to form organs. Proteins transport other molecules, such as oxygen, to tissues. Antibody molecules contribute to host defense against infections. Protein hormones relay information between cells. Moreover, protein machines, such as actin-myosin complexes, can perform useful work, including cell movement. Thus, studying proteins is a prerequisite in understanding cell structure and function. The physical characterization of proteins began well over 150 years ago with Mulder’s characterization of the atomic composition of proteins. In the latter half of the nineteenth century Hoppe-Seyler (1864) crystallized he-
UNIT 1.5
moglobin and Kühn (1876) purified trypsin. A variety of physical methods have been developed over the years to increase convenience and precision in the characterization and isolation of proteins. These include ultracentrifugation, chromatography, electrophoresis, and others. In many instances our understanding of cell proteins parallels the introduction and use of new techniques to examine their structure and function.
PROTEIN CLASSIFICATIONS All proteins are constructed as a linear sequence(s) of various numbers and combinations of ∼20 α-amino acids joined by peptide bonds to form structures from thousands to millions of daltons in size. Proteins are the most complex and heterogeneous molecules found in cells, where they account for >50% of the dry weight of cells and ∼75% of tissues. Proteins can be classified into three broad groups: globular, fibrous, and transmembrane (Fig. 1.5.1; Table 1.5.1). Globular proteins are, by definition, globe-shaped, although in prac-
globular
fibrous
Figure 1.5.1 General classifications of proteins. In these schematic representations of globular, fibrous, and transmembrane proteins, hydrophobic regions are shaded. Note that the disposition of hydrophobic residues often reflects the protein class.
transmembrane
Strategies of Protein Purification and Characterization Contributed by Howard R. Petty Current Protocols in Protein Science (2002) 1.5.1-1.5.10 Copyright © 2002 by John Wiley & Sons, Inc.
1.5.1 Supplement 31
Table 1.5.1
Broad Classifications for Proteinsa
Type
Location/type
Examples
Globular
Intracellular
Hemoglobin, lactate dehydrogenase, cytochrome c Serum albumin, immunoglobulins, lysozyme
Extracellular Fibrous
Intracellular Extracellular
Transmembrane
Single pass Multipass
Intermediate filaments, tropomyosin, lamins Collagen, keratin, elastins Insulin receptor, glycophorin, HLAsb Glucose transporter, rhodopsin, acetylcholine receptor
aAdditional information regarding fibrous and transmembrane proteins can be found in Squire
and Vibert (1987) and Petty (1993). Information concerning globular proteins can be found in numerous books on proteins and enzymes such as Schultz and Schirmer (1979). bHuman histocompatibility leukocyte antigens.
tice they can be spherical or ellipsoidal. Globular proteins are generally soluble in aqueous environments. Examples of globular proteins are hemoglobin, serum albumin, and most enzymes. Fibrous proteins are elongated linear molecules that are generally insoluble in water and resist applied stresses and strains. Collagen is a physically tough molecule of connective tissue. Just as collagen gives strength to connective tissues, intermediate filaments linked to desmosomes give strength to cells in tissues. The third general class of proteins, transmembrane proteins, contain a hydrophobic sequence buried within the membrane; these proteins are discussed more fully below (see Membrane Proteins). These protein categories are not mutually exclusive. For example, the nominally fibrous intermediate filament proteins also have globular domains. Similarly, transmembrane proteins almost always possess globular domains. Thus, these definitions serve as a useful guide but should not be rigidly applied.
HYDROPATHY PATTERNS OFTEN REFLECT A PROTEIN’S CLASSIFICATION
Overview of the Physical State of Proteins Within Cells
A key physical feature of proteins is their hydropathy pattern (i.e., the distribution of hydrophobic and hydrophilic amino acid residues). Indeed, hydrophobic interactions provide the primary net free energy required for protein folding. Figure 1.5.1 illustrates the disposition of hydrophobic amino acids in proteins. In an intact globular protein, hydrophobic
amino acids are generally shielded from the aqueous environment by coalescing at the center of the molecule, with the more hydrophilic residues exposed at its surface. However, the linear arrangement of hydrophobic residues fluctuates in an apparently random fashion. The α helices within globular proteins may express a hydrophobic face oriented toward the center of the protein. (Within these helices hydrophobic residues are nonrandomly positioned every three or four amino acids to yield a hydrophobic face.) For coiled-coil α helix–containing fibrous proteins, such as tropomyosin and αkeratin, hydrophobic residues at periodic intervals allow close van der Waals contact of the chains and potentiate assembly as hydrophobic residues are removed from the aqueous environment (Schulz and Schirmer, 1979; Parry, 1987). Secondarily, regularly spaced charged groups can also contribute to the shape of fibrous proteins (Schulz and Schirmer, 1979; Parry, 1987). Transmembrane proteins provide a rather different physical arrangement of hydrophobic residues in which hydrophobic residues are collected primarily into a series of amino acids that is embedded within a cell membrane. One important means of analyzing the hydropathy of a sequenced protein is a hydropathy plot (Kyte and Doolittle, 1982). In this method, each amino acid residue is assigned a hydropathy value, an ad hoc measure that largely reflects its relative aqueous solubility; these values are plotted after being averaged. The successful interpretation of hydropathy plots
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depends on the parameters chosen for averaging. The parameters are the number of residues averaged (amino acid interval or “window”) and how many amino acids are skipped when calculating the next average (step size). Using this approach with a window of ∼10 residues, it is often possible to find the positions of hydrophobic residues coalescing near the interior of globular proteins. The method is particularly useful in predicting transmembrane domains of proteins, generally with a window of ∼20 amino acids. To detect the repetitious pattern of coiled-coil fibrous proteins, however, windows smaller than the repeat length would be required.
MEMBRANE PROTEINS In addition to their presence in the extracellular and intracellular milieus, proteins are also found in association with biological membranes. Proteins constitute one-half to threequarters of the dry weight of membranes. Membrane proteins perform a broad variety of functions including intermembrane and intercellular recognition, transmembrane signaling, most energy-harvesting processes, and biosynthesis in the endoplasmic reticulum (ER) and Golgi complex. Membrane proteins have been traditionally characterized as integral (or intrinsic) or peripheral (or extrinsic) on the basis of operational criteria. Peripheral membrane proteins are associated with membrane surfaces and can be dislodged from membranes using hypotonic or hypertonic solutions, pH changes, or chela-
Table 1.5.2
tion of divalent cations. Components of the erythrocyte membrane skeleton, for example, are peripheral membrane proteins. Although most peripheral proteins are removed by washing a sample with buffers, integral proteins cannot be removed by such treatments. To isolate integral membrane proteins, which are embedded within the lipid bilayer, one must use detergents that disrupt the bilayer and bind to the proteins, thus solubilizing them. In general, integral membrane proteins have a portion of their peptide sequence buried in the lipid bilayer whereas peripheral proteins do not. However, the discovery of glycosylphosphatidylinositol (GPI)-linked membrane proteins added to the ambiguity of the situation. GPI-linked proteins are globular proteins with no membrane-associated peptide sequence, yet they require harsh conditions for solubilization. As the technology for studying membrane proteins improved, it became necessary to develop a more precise vocabulary to describe membrane proteins. Transmembrane integral membrane proteins have at least one stretch of amino acids spanning a membrane. Membrane proteins are classified as type I, II, III, or IV depending on the nature of their biosynthesis and topology in membranes (Spiess, 1995; Table 1.5.2 and Fig. 1.5.2). The biosynthetic insertion of these proteins in membranes is, in turn, dependent on the presence or absence of a cleavable signal peptide, the relative positions of the hydrophobic transmembrane domain and positively charged topogenic signals, and/or
Definitions of Integral Transmembrane Proteins
Type
Definition
I
An N-terminal–cleavable signal peptide is removed at the luminal face yielding a luminal N terminal during biosynthesis. (Positive charges are found on C-terminal side of first long hydrophobic sequence after the signal peptide.) An N-terminal–uncleaved signal peptide leads to a cytoplasmic N terminus. (Positive charges are generally found on N-terminal side of first long hydrophobic sequence.) A long N-terminal hydrophobic sequence is followed by a sequence of positive charges. This leads to a luminal N terminus in the absence of a cleavable signal peptide. A short C terminus is present at the luminal side of membrane. A large N terminus is exposed at the cytoplasmic face.
II
III
IV
Examples LDL receptor, insulin receptor, glycophorin A, thrombin receptor
Transferrin receptor, sucrase/isomaltase, band 3
β-Adrenergic receptor, cytochrome P450
Synaptobrevin, UBC6
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transmembrane
nontransmembrane type I N
type II C
type III
type IV
N C
bilayer
exterior or lumen
cytosol
C
N
+ + +
C
+ + +
N
+ + +
C
N
Figure 1.5.2 Membrane proteins containing hydrophobic anchors. A nontransmembrane or monotopic membrane protein is anchored to the membrane via a hydrophobic amino acid sequence. Transmembrane proteins are classified as types I, II, III, and IV (Table 1.5.2). The first transmembrane segment of a multispanning membrane protein can be inserted as in type I, II, or III proteins. This segment functions as a start-transfer peptide. Subsequent transmembrane segments will function as stop-transfer and start-transfer sequences, resulting in a multispanning membrane topology.
Overview of the Physical State of Proteins Within Cells
the mechanism of nascent protein delivery to the ER. Type I membrane proteins are synthesized with an amino-terminal signal sequence that is inserted into the ER membrane. When the signal sequence is proteolytically removed in the ER lumen, a new luminal amino terminus is exposed. A series of positively charged residues at the C-terminal side of the first hydrophobic transmembrane domain following the signal sequence generally denotes the end of the first transmembrane domain (von Heijne and Gavel, 1988). Although a hydrophobic transmembrane domain followed by a positive sequence of amino acids is sufficient to act as a stop-transfer signal, this motif is not required for stoptransfer events and other, less well-understood regulatory mechanisms are also involved (Andrews and Johnson, 1996). Membrane proteins types I, II, and III are delivered to the ER membrane via a signal recognition particle (SRP)-dependent mechanism. In contrast to type I proteins, type II and III membrane proteins do not have a cleavable N-terminal signal sequence. Instead, they have an internal hydrophobic signal that acts as both a signal sequence for ER delivery and a transmembrane domain in the mature protein. Type II proteins have a cytoplasmic amino terminus and a luminal (or extracellular) carboxyl terminus. In this case a positively charged sequence
of amino acids at the N-terminal side of the first hydrophobic sequence causes the amino terminus to be retained at the cytoplasmic face of the ER membrane. Thus the internal uncleaved signal peptide becomes the transmembrane domain of the mature protein. Type III membrane proteins have the same overall topology as type I proteins, but they are inserted into membranes by a different mechanism. In type III proteins the first hydrophobic sequence of amino acids is immediately followed by a series of positively charged amino acids. Thus, the first hydrophobic sequence becomes the transmembrane domain of the protein, with the amino terminus at the luminal face of the membrane. Type IV membrane proteins are characterized by a large, cytoplasmically exposed amino-terminal domain and a short carboxylterminal domain facing the lumen. Importantly, these proteins are delivered to the ER by an unknown SRP-independent mechanism. In addition to the single-pass membrane proteins just described, integral membrane proteins can display zero, two, three, or more transmembrane domains. Some membrane proteins, such as cytochrome b5, have protein segments buried in the hydrophobic core of membranes but do not cross the membrane. Membrane proteins with multiple membranespanning domains are classified as type I, II, or
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III depending on the topogenic signals in the first transmembrane domain. For example, a multispan membrane protein with a cleavable signal sequence, luminal amino terminal, and a positively charged sequence following the first transmembrane domain from the amino terminal, such as the thrombin receptor, is a type I membrane protein. The remaining transmembrane domains are inserted into the bilayer depending on the orientation of the first transmembrane domain. Multispanning type II and III proteins are similarly defined according to the properties of their single-spanning counterparts. In addition to hydrophobic protein sequences acting as membrane anchors, membrane proteins may also carry bilayer-associated hydrophobic lipid components. These hydrophobic lipid anchors define three broad groups of lipid-modified proteins: fatty acylated, isoprenoid-linked, and GPI-linked (Fig. 1.5.3). Several cytosolic transmembrane proteins have been identified that contain a covalently attached hydrophobic fatty acyl residue. For example, fatty acids, including palmitic, palmitoleic, cis-vaccenic, and cyclopropylene-
hexadecanoic, are covalently linked to the amino terminus and the amino-terminal glycerylcysteine of E. coli lipoprotein. Moreover, palmitate- and myristate-labeled transmembrane proteins have been observed in eukaryotic cells (e.g., Schlesinger et al., 1980). In both isoprenoid-linked and GPI-linked proteins, globular proteins become membranebound due to the addition of a hydrophobic lipid moiety. Certain proteins containing conserved cysteine residues at or near the C-terminus are modified by prenylation, in which a farnesyl or geranylgeranyl isoprenoid tail is added (Zhang and Casey, 1996). This hydrophobic moiety promotes protein association with the cytoplasmic face of cell membranes. Notably, cytosolic G proteins and protein kinases that participate in signal transduction are prenylated. GPI-linked proteins are a major class of membrane proteins (Cardoso de Almeida, 1992; Englund, 1993). In contrast to isoprenoid-modified proteins, GPI-linked proteins are attached to the luminal or extracellular face of membranes via a glycosylphosphatidylinositol anchor of variable structure (e.g.,
EtN P
P exterior or lumen bilayer
P
cytosol
S
C O
fatty acylated
S GPI-linked
Cys-CH3
isoprenoid-linked
Figure 1.5.3 Membrane proteins containing lipid moieties. In the simplest case, fatty acids can be covalently attached to transmembrane proteins. Hydrophobic tails are also attached to proteins to form isoprenoid-linked proteins. A third class of lipid-attached proteins are the GPI-linked proteins. Hydrophobic regions are shaded.
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Fig. 1.5.3). Well over 100 GPI-linked proteins have been identified in cells, where they perform numerous functions including acting as enzymes and receptors. The ability of GPIlinked proteins, which possess no transmembrane or cytosolic sequences, to elicit transmembrane signals seems paradoxical. However, studies have suggested that interactions with other proteins, including transmembrane integrins (Petty et al., 1996), contribute to transmembrane signaling of these proteins. In addition, GPI-linked proteins can collect in microdomains called lipid rafts within cell membranes (Rietveld and Simons, 1998). Although GPI-linked proteins must collaborate with other membrane proteins to elicit signals, they do possess certain functional advantages. First, GPI-linked proteins (and isoprenoid-linked proteins as well) diffuse in membranes much faster than transmembrane proteins and thus relay information faster. Second, certain cells, such as leukocytes, can rapidly shed their GPIlinked proteins, thus altering their functional properties in seconds. Although the importance of lipid-linked membrane proteins has only recently been appreciated, the impact of these structures on our understanding of cell properties is growing rapidly.
ADDITIONAL FACTORS AFFECTING THE PHYSICAL HETEROGENEITY OF PROTEINS
Overview of the Physical State of Proteins Within Cells
Additional factors contributing to the physical-chemical heterogeneity of proteins are size, charge, chemical modifications, and assembly. A typical amino acid has a molecular mass of ∼110 Da, and a small protein has a molecular mass of a few thousand daltons (e.g., for insulin, Mr = 5733). Large proteins have molecular masses of several hundred thousand daltons. When proteins are assembled to form large multiprotein complexes such as ribosomes, molecular masses are well into the millions. The diameters of these structures range from 4 Å for an individual amino acid to ∼30 nm for a ribosome. Electrostatic charge is of major importance in protein structure and function. Charged proteins are more soluble than uncharged proteins. The large number of positive charges on histones allow them to bind DNA. The spatial arrangement of charges on cytochrome c allows it to bind the complementary charges of its oxidase and reductase, thereby orienting the proteins prior to electron transfer. Similarly, the arrangement of charges on the apoprotein and receptor for low-density lipoprotein (LDL) al-
lows for lock-and-key–like interactions (Petty, 1993). In addition to structural and binding considerations, electrostatic interactions play a regulatory role. For example, the phosphorylation and dephosphorylation of insulin receptors alter electrostatic interations between the active site and a regulatory loop of the kinase domain, thereby changing its three-dimensional shape (Hubbard et al., 1994). This changes the Vmax of the kinase, thus triggering intracellular signals. In addition to the types of physical heterogeneity listed above, >100 distinct chemical modifications of proteins have been observed. These include, for example, glycosylation, ubiquitin attachment, phosphorylation, acetylation, and hydroxylation (Table 1.5.3). Thus, proteins undergo extensive physical-chemical modification.
PROTEIN ASSEMBLIES Proteins can be assembled in a variety of states in both aqueous media and within membranes. Protein assembly into complex supramolecular structures plays vital roles in enzyme regulation, cell skeleton formation, and transmembrane signaling. Both covalent bonds and noncovalent bonds participate in protein assembly. One frequently encountered covalent mechanism of protein assembly is the formation of disulfide bonds. These covalent linkages often form during protein maturation. They can link two separate proteins together or two portions of the same protein. For example, the two chains of insulin molecules are held together by disulfides, as are the two chains of its membrane receptor. However, disulfide bond formation is mostly limited to oxidative environments such as the ER lumen and the exterior face of the cell surface. One of the best-known examples of noncovalent assembly is the formation of hemoglobin tetramers. Polymerization is another frequently encountered mechanism for protein assembly in cells. The globular protein actin polymerizes to form microfilaments in the absence of covalent bond formation. Intermediate filaments are formed by the polymerization of fibrous proteins. Under certain circumstances transmembrane proteins polymerize as well; bacteriorhodopsin, for example, forms two-dimensional pseudocrystals called purple membranes. Protein assemblies formed from various numbers of similar units are homodimers, homooligomers, and homopolymers. Assembly of protein structures from dissimilar subunits is more common than assem-
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Table 1.5.3
Common Physical-Chemical Modifications of Proteinsa
Modification
Example
Homodimerization Homooligomerization Homopolymerization Heterodimerization Heterooligomerization Heteropolymerization Proteolytic cleavage Prosthetic group addition Oxidation-reduction Glycosylation Phosphorylation
Transferrin receptor S. typhimurium glutamine synthetase Actin Integrins Histones, proteasomes Ribosomes Signal peptide cleavage in ER Heme addition to cytochromes and hemoglobin Disulfide bond formation in ER Glycoprotein maturation Regulation of protein function, such as the tyrosine kinase activity of insulin receptors Blockage of N-termini of certain membrane proteins Ubiquitin-dependent proteolysis via proteasomes, histones Proline hydroxylation on collagen Insulin receptors, E. coli lipoprotein G proteins Alkaline phosphatase, urokinase receptors
Acetylation Ubiquitination Hydroxylation Fatty acylation Isoprenylation GPI addition
aFor details, see Freedman and Hawkins (1980, 1985), Schlesinger et al. (1980), Englund (1993), and Zhang and
Casey (1996).
bly from identical subunits. For example, heterodimers are formed from the α and β chains of integrins within cell membranes. Complex heterooligomeric and heteropolymeric structures vary from relatively small structures such as histone octamers, which bind to DNA in the nucleus, to large particles such as ribosomes, found both in the cytosol and attached to nuclear and ER membranes. The signal recognition particle is a relatively small heterooligomeric structure, composed of one RNA subunit and six proteins, that potentiates the delivery of secretory and most membrane proteins to the ER membrane. Membrane-associated heterooligomeric structures have also been observed. One of the best examples of such structures is the components of the electron transport systems in chloroplasts and mitochondria (Petty, 1993). For example, the ubiquinone-cytochrome c reductase is composed of eleven different subunits. Thus, proteins can be assembled in a variety of manners within cells. Although some protein assemblies, such as intermediate filaments, are static structures, many are dynamic structures which provide functional flexibility. For example, microfilaments can rapidly assemble and disassemble. In addition to the physical changes in assembly state, compositional dynamics is also observed.
For example, interferon γ treatment alters the composition of proteasomes. Developmental changes in protein composition are also observed. As an example, fetal and newborn forms of a component of cytochrome c reductase are expressed in humans. Thus, protein assemblies can be characterized by both physical and compositional dynamics.
ALTERING THE SOLUBILITY OF PROTEINS: PROTEIN EXTRACTION The in vitro characterization of cellular proteins begins with their extraction from tissues or cells into a buffer. With the exception of globular secretory proteins, such as those found in plasma, proteins are generally not easily accessible for experimental manipulation. For example, many fibrous proteins are not soluble in aqueous buffers. Cellular proteins are entrapped within or on a cell and therefore must be extracted from the cell in a soluble form. A variety of methods including osmotic lysis, enzyme digestion, homogenization using a blender or mortar and pestle, and disruption by French press and sonication have been employed to disrupt cells. For a cytosolic protein such as hemoglobin, no further extraction from the sample is necessary. However, many important cellular proteins, such as those associated
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(CCl3COO−). Hydrophobic interactions are also reduced by exposure to organic solvents and low salt concentrations. Electrostatic interactions are reduced by high salt conditions; this decreases the Debye-Hückel screening length and coulombic attraction. To disrupt hydrogen bonds, high concentrations of urea or guanidine are often employed. More vigorous methods of sample denaturation using very low pH or harsh detergents such as sodium dodecyl sulfate are also used to diminish intermolecular contacts. Once proteins are extracted, their size can be characterized by ultracentrifugation on sucrose gradients (UNIT 4.2), gel filtration chromatography (UNIT 8.3), SDS-PAGE (UNIT 10.1), and other methods (Table 1.5.5). The charge characteristics of proteins can be assessed using isoelectric focusing and ion-exchange chromatography. Specific interactions, such as antigen-antibody and biotin-avidin interactions, can also be employed in the characterization and isolation of proteins. These are useful in immunoblotting (UNIT 10.10) and affinity chromatography methods (Chapter 9).
with membranes, cytoskeletal components, and DNA, remain insoluble. To further solubilize cell proteins, both nonionic (e.g., Triton X-100) and ionic (e.g., sodium dodecyl sulfate) detergents are often employed. Detergents are small amphipathic molecules that interact with both nonpolar and polar environments. Detergents disrupt membranes. They also bind to hydrophobic regions of proteins, such as their transmembrane domains, thereby replacing the unfavorable contacts between hydrophobic protein regions and water with the more favorable hydrophilic domains of the detergent. Thus, instead of the hydrophobic regions of the insoluble protein forming an aggregate in the bottom of a test tube, the protein becomes soluble and can be employed in most in vitro analyses. In addition to detergents, several other solubilization strategies are useful for the extraction and in vitro characterization of proteins (Table 1.5.4). Chaotropic agents enhance the transfer of nonpolar molecules to aqueous environments by their disrupting influence on water structure. Chaotropic agents are generally large molecular ions such as thiocyanate (SCN−), perchlorate (ClO4−), and trichloroacetate
Table 1.5.4
Physical Bases of Common Protein Extraction and/or Elution Methods
Physical property perturbed
Agents
Hydrogen bonds Ion pair interactions Hydrophobic interactions
Urea or guanidine⋅HCl, pH changes High salt, pH changes Detergents, chaotropic agents, organic solvents, low salt
Table 1.5.5 Physical Bases of Common Protein Characterization and Isolation Methods
Overview of the Physical State of Proteins Within Cells
Physical property
Method
References to other units
Solubility
Extraction with salts, detergents, and enzymes
Chapter 4, Racker (1985)
Size
Ultracentrifugation on sucrose gradients Gel filtration SDS-PAGE
UNIT 4.2 UNIT 8.3 UNIT 10.1
Charge
UNIT 10.2 Isoelectric focusing Ion-exchange chromatography UNIT 8.2
Biospecific interaction
Immunoblotting Immunoaffinity chromatography
Hydrophobicity
UNIT 10.10
Chapter 9
Hydrophobic chromatography UNIT 8.4 Reversed-phase HPLC UNIT 8.7
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LIMITATIONS OF THE IN VITRO MANIPULATION OF PROTEINS The very act of isolating proteins perturbs their physical environment. Although this is not often a major problem, a few cautionary notes should be made. The most primitive compartment of a cell, the cytosol, is a chemically reducing environment. Consequently, free sulfhydryl groups are observed in the cytosol; in fact, multiple cytosolic pathways help in preserving the proper redox conditions. On the other hand, the extracellular milieu and the luminal side of the ER are oxidative environments. The oxidizing condition within the ER is presumably due to the unidirectional transport of glutathione and cystine. Consequently, disulfides are frequently observed in the ER and extracellular environments. Thus, to prevent disulfide formation during manipulation, sulfhydryl blocking reagents such as iodoacetamide are included in extraction buffers. The cytosol is also a K+-rich and Ca2+-poor solution. These parameters should be considered in designing physiologically relevant experiments. The experimental manipulation of membrane proteins is decidedly more difficult. The exterior face exists in a high Na+ and Ca2+ solution that is oxidative; just the opposite is true for the cytoplasmic face. Since no appropriate solvent exists for such isolated proteins, experimental questions can be directed at properties associated with just one face of the molecule. A second limitation common to all in vitro studies of transmembrane proteins is that they must be solubilized using detergents. In addition to solubilizing a transmembrane protein, detergents can also bind to hydrophobic regions in the globular domain(s) of the protein, thus affecting the properties under study. One means of countering this problem is to test several detergents in the hope of finding one that retains the full biological activity of the purified protein. Protein solubilization can also lead to loss of physiologically relevant protein-protein interactions. This can occur by simple dilution or by disruption of noncovalent interactions among proteins. For example, hemoglobin exists as a supersaturated solution in vivo which cannot be duplicated in vitro. Furthermore, protein-protein associations are generally stronger in the restricted confines of membranes than after solubilization into a buffer. Thus, protein assemblies found in cells may disappear during solubilization. One means of countering these potential difficulties is to co-
valently cross-link protein assemblies prior to disruption and to solubilize proteins using mild detergents (e.g., Brij-58).
CONCLUSIONS Structural motifs, especially stretches of hydrophobic amino acids, contribute to the shape of a protein and its classification as globular, fibrous, or transmembrane. Proteins are heterogeneous at many different levels including physical attributes, covalent modifications, and supramolecular assembly. The physical properties of proteins are used to characterize and isolate these molecules. For example, the size of a protein is examined by sedimentation on sucrose gradients (UNIT 4.2), gel filtration (UNIT 8.3), and polyacrylamide gel electrophoresis (UNIT 10.1). Its charge is the key physical parameter in isoelectric focusing and ion-exchange chromatography. The units that follow contain detailed protocols describing the characterization of cellular proteins.
LITERATURE CITED Andrews, D.W. and Johnson, A.E. 1996. The translocon: More than a hole in the ER membrane? Trends Biochem. Sci. 21:365-369. Ausubel, F.M., Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G., Smith, J.A., and Struhl, K. (eds.) 1998. Current Protocols in Molecular Biology. John Wiley & Sons, New York. Cardoso de Almeida, M.L. 1992. GPI Membrane Anchors. Academic Press, New York. Englund, P.T. 1993. The structure and biosynthesis of glycosyl phosphatidylinositol protein anchors. Annu. Rev. Biochem. 62:121-138. Freedman, R.B. and Hawkins, H.C. 1980. The Enzymology of Post-Translational Modifications of Proteins, Vol. 1. Academic Press, New York. Freedman, R.B. and Hawkins, H.C. 1985. The Enzymology of Post-Translational Modifications of Proteins, Vol. 2. Academic Press, New York. Hoppe-Seyler, F. 1864. Über die chemischen und optischen Eigenschaften des Blutfarbstoffs. Virchows Arch. 29:233-235. Hubbard, S.R., Wei, L., Ellis, L., and Hendrickson, W.A. 1994. Crystal structure of the tyrosine kinase domain of the human insulin receptor. Nature 372:746-754. Kühn, W. 1876. Über das Verhalten verschiedner organisirter und sogenannter ungeformter Fermete. Über das Trypsin (Enzym des Pankreas) [Reprint, FEBS Lett. 62:E3-E7 (1976).]. Kyte, J. and Doolittle, R.F. 1982. A simple method for displaying the hydrophobic character of a protein. J. Mol. Biol. 157:105-132. Strategies of Protein Purification and Characterization
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Parry, D.A.D. 1987. Fibrous protein structure and sequence analysis. In Fibrous Protein Structure (J.M. Squire and P.J. Vibert, eds.) pp. 141-171. Academic Press, New York. Petty, H.R. 1993. Molecular Biology of Membranes: Structure and Function. Plenum, New York.
KEY REFERENCES Racker, 1985. See above. A wonderful little book on membrane protein manipulation which disproves the hypothesis that scientists can’t write.
Petty, H.R., Worth, R.G., and Todd, R.F. III. 2002. Interactions of integrins with their partner proteins in leukocyte membranes. Immunol. Res. 25:75-95.
Tanford, C. 1961. Physical Chemistry of Macromolecules. Academic Press, New York.
Racker, E. 1985. Reconstitutions of Transporters, Receptors, and Pathological States. Academic Press, New York.
Tanford, C. 1980. The Hydrophobic Effect. John Wiley & Sons, New York.
Rietveld, A. and Simons, K. 1998. The differential miscibility of lipids as the basis for the formation of functional membrane rafts. Biochim. Biophys. Acta. 1376:467-479. Schlesinger, M.J., Magee, A.I., and Schmidt, M.F.G. 1980. Fatty acylation of proteins in cultured cells. J. Biol. Chem. 255:10021-10024.
A rigorous introduction to the physical properties of proteins, which remains useful several decades later.
A very readable introduction to the hydrophobic effect.
INTERNET RESOURCES http://www.expasy.ch A user-friendly protein database including two-dimensional PAGE data and 3D protein structures.
Schultz, G.E. and Schirmer, R.M. 1979. Principles of Protein Structure. Springer-Verlag, New York.
ftp://ftp.pdb.bnl.gov/
Spiess, M. 1995. Heads or tails—what determines the orientation of proteins in the membrane. FEBS Lett. 369:76-79.
http://ww.ncbi.nlm.nih.gov
Contains protein crystallography data.
An important protein database.
Squire, J.M. and Vibert, P.J. (eds.) 1987. Fibrous Protein Structure. Academic Press, New York. von Heijne, G. and Gavel, Y. 1988. Topogenic signals in integral membrane proteins. Eur. J. Biochem. 174:671-678.
Contributed by Howard R. Petty Wayne State University Detroit, Michigan
Zhang, F.L. and Casey, P.J. 1996. Protein prenylation: Molecular mechanisms and functional consequences. Annu. Rev. Biochem. 65:241-269.
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CHAPTER 2 Computational Analysis INTRODUCTION
T
he development of the techniques of modern molecular biology, including PCR, cloning, and nucleotide sequencing, has revolutionized the practice of protein chemistry. Whereas the initial phase of detailed study of a new protein activity once involved chemical analysis of the amino acid sequence by Edman degradation of multiple fragments, it is more common today to establish a protein’s linear sequence from the nucleotide sequence of the gene that encodes it. While the number of new sequences reported based solely on classical protein sequence analysis has drastically declined, the molecular biology revolution has generated a rapidly expanding collection of new sequence data. The current efforts to sequence specific genomes, including the human genome, in their entirety will undoubtedly lead to further expansion of this information. Because linear sequence information is so readily available, computer methods for analyzing these data have also developed rapidly. This chapter will present methods for deriving important information on possible familial, structural, and functional relationships using only the linear sequence information as input.
As soon as any sequence information is obtained—for example, by running a newly isolated protein through an N-terminal gas-phase sequencer or by dideoxy sequencing a piece of cloned DNA—searches for similarity to other proteins can begin. If the new sequence can be identified as related to that of a known enzyme or gene from another organism, this provides excellent initial confirmation that the protein or gene under study is worthy of further investigations. Conversely, if a match is found to the sequence of a known protein that could only plausibly arise from sample contamination, the study can be halted immediately at considerable savings of time, energy, and resources. UNIT 2.1 describes procedures for searching sequence databases, provides theoretical background to aid in interpreting the resulting output, and discusses resources for further exploration of sequence relationships. Once substantial sequence information (particularly a whole gene sequence) is obtained, the most pressing question is what structure it is likely to have. A first step toward answering this question is the analysis of the corresponding amino acid sequence for hydrophobic and hydrophilic regions. Of particular interest are patterns or groupings of hydrophobic residues—typically transmembrane helices, leucine zippers, and amphipathic segments—which can indicate structural and even functional features of the protein. Hydropathy scales, indices designed to quantify the relative hydrophobicity of amino acid residues within a protein, and their application to protein sequences are reviewed in UNIT 2.2. More sophisticated forms of analysis can be used to predict overall protein secondary structure based on linear sequence. The procedures developed for this purpose utilize statistical data that has been compiled over the past 25 years as more and more three-dimensional protein structures have become available. Although these methods are not completely accurate at present, the success rate for predictions is now in the 60% to 80% range. This information is fully adequate to develop hypotheses, design experimental tests, and suggest relationships to known tertiary structures. The prediction of protein secondary structure and folding class are reviewed in UNIT 2.3. As well as providing Contributed by Ben M. Dunn Current Protocols in Protein Science (2000) 2.0.1-2.0.2 Copyright © 2000 by John Wiley & Sons, Inc.
Computational Analysis
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theoretical background and detailing the different predictive approaches, the discussion includes an analysis of the strengths and weaknesses of each approach with respect to different types of proteins (using human hemoglobin β, interleukin 1β, and profilin I as examples) and gives some guidance to obtaining the necessary software. To perform any method of computer analysis, especially comparisons with other known protein sequences, it is necessary to have facile methods for transfer and capture of a variety of files. Fortunately, standard methodology now exists to perform many of these functions. UNIT 2.4 provides detailed instructions for the initial steps of getting connected, finding information, and transferring data between computers around the world. Although each university or company will have their own local protocols and systems, and although individuals will need to obtain help from their local computer wizards, UNIT 2.4 is written to provide directions that are generally applicable to most systems. Specific recommendations for utilizing various forms of the BLAST programs for sequence searching and comparison are provided in UNIT 2.5. Protocols for submission of search commands and search sequence are provided in detail, and examples of search results are presented. The ultimate goal of any protein chemist should be to understand the relationship between the three-dimensional structure and the functional properties. Once a new sequence has been discovered, many scientists inevitably begin to speculate on the possible three-dimensional structure. To accomplish this, one must proceed through several steps. An important beginning is the search for proteins of related sequence, as one may find important clues through family relations. The appropriate databases that can be utilized for this purpose are collected in UNIT 2.6. There are several general methods to approach the prediction of structure: homology modeling, fold recognition, and ab initio prediction. The differences between these methods are described in UNIT 2.7. Comparative protein modeling can be extremely successful if there are structurally-defined proteins with a significant degree of sequence relatedness. The definitions and steps necessary for conducting comparative modeling using the SwissModel Web server are detailed in UNIT 2.8. Future updates to this chapter will present alternative methods to complete the sequence to structure algorithm. Ben M. Dunn
Introduction
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Current Protocols in Protein Science
Computational Methods for Protein Sequence Analysis Despite continued advances since the first applications of computers in protein sequence analysis in the late 1960s, computer-assisted biological sequence analysis is still undergoing rapid changes and improvements. With so much activity and growth in the scope and importance of genomic research, the need for efficient biological sequence analysis has become more urgent. Nevertheless, users of computational tools should be aware of the limitations of particular methods. Good results require the user to be able to select the right tools on the basis of some theoretical knowledge and to interpret all results with biological correctness in mind. This unit is presented as a guide to addressing the issue of what to do with a protein sequence once it is obtained. The practical approach depends on how far the investigation has progressed. If only a short part of the protein (e.g., ∼10 to 20 amino acids) has been determined, initial scans of databases to see if there are any significant matches with a known sequence are appropriate. If a complete protein sequence is in hand, it is time to start identifying regions of similarity that resemble functional sites of known proteins.
THEORETICAL BACKGROUND FOR PROTEIN SEQUENCE ANALYSIS Computer representations of protein sequences are of two basic types: artificial and natural. An example of an artificial or symbolic protein sequence is the common 24-letter residue code sequence (ABCDEFGHIKLMNPQRSTVWXYZ*) that represents each amino acid as suggested by the Nomenclature Committee of the International Union of Biochemistry (IUB). This code allows for ambiguities and unknown residue modifications. A natural protein sequence, on the other hand, is directly representative of the physical properties of the individual amino acids; for example, in a natural protein sequence the individual elements might be vectors representing the physical characteristics of the amino acid (Robson and Greaney, 1992; Rohde and Bork, 1993). Most molecular sequence analysis algorithms have been designed for the IUB 24-character amino acid representation. A sequence is an ordered list of elements. Contributed by George Michaels and Robert Garian Current Protocols in Protein Science (1995) 2.1.1-2.1.18 Copyright © 2000 by John Wiley & Sons, Inc.
Sequences can vary in length, composition, and pattern. The number of possible different protein sequences possible of a particular length is 20L, where L is the length. The patterns in such sequences are said to constitute the primary structure of the protein. In general, proteins have nonrandom sequences that are presumed to be related in the sense that all proteins have evolved from other proteins (Doolittle, 1981, 1986, 1989, 1990). Protein sequences are also subject to sequence convergence: different sequences may represent the same native structure of a protein. Some terms that deal with comparisons of protein sequence data are often used and misused in the literature. Similarity is generally used to describe two sequences or structures that resemble one another due to the order of identical residues. Homology means that similar sequences or structures resemble one another due to descent from a common evolutionary ancestor. It is important to realize that the sequence analysis tools used to search data bases detect character patterns that resemble portions of the query sequence. This means that they are finding similarities. Homologies are identified through the recognition of correlated structural and functional data in addition to similarity data. There are three basic types of sequence analyses: listings, traces, and alignments (Kruskal, 1983; Kruskal and Sankoff, 1983). Let a = (a1,..., am) and b = (b1,..., bn) be two sequences and let E be a set of elementary operations including insertion, deletion, and substitution. An indel is either an insertion or a deletion. A listing is a finite sequence of operations taken from E that transform a into b. A trace is a one-to-one correspondence of a subsequence of a and a subsequence of b. An alignment of a and b is a one-to-one correspondence such that each element of one sequence corresponds either to an element of the opposite sequence or to a null element indicating the presence of a gap. The best alignments are obtained when the lengths of a and b are similar. The correspondence between two nonnull elements is called an identity or a continuation if the two elements are identical and a substitution otherwise. Lines representing the correspondence between two sequences should not cross.
UNIT 2.1
Computational Analysis
2.1.1 CPPS
Computational Methods for Protein Sequence Analysis
In the comparison of two different alignments, it is not necessarily the one with the larger number of identities that is better. An alignment with fewer indels (<5%) might be preferable; therefore, some alignment methods set a limit to the number of indels allowed in a given alignment. Alignments are the most frequently used type of sequence analysis. There are two basic types of alignments: local and global. Local alignments are used to identify the most similar common subregions of two sequences. They are especially useful in identifying distantly related sequences. A global alignment, on the other hand, attempts a comparison of two sequences in their entirety and, therefore, often misses small subregions of similarity (Pizzi et al., 1991). Global alignments are most useful for comparing highly similar or homologous sequences that are close to one another in length. Two sequences are said to be homologous if it can be inferred on the basis of all available information that the sequences have a common ancestor. There are no degrees of homology. Divergent sequences are those that can be derived from the same sequence, usually by applying the operations in E. One way that divergent sequences are identified is by applying scoring matrices. There are several scoring matrices available based on allowable amino acid substitutions found in protein alignments of related proteins (see discussion of scoring matrices that follows). The most common matrices used are the protein alignment matrix (PAM; Dayhoff, 1978); the mutation data matrix (MDM; George, et al., 1990), which is a log-odds version of PAM; and BLOSUM (Henikoff and Henikoff, 1993), which incorporates the consideration of the conversation of function data between aligned proteins. It should be noted that the restriction of global alignments to operations in E would fail to align two sequences containing transpositions, in which segments of the protein sequence have been reordered within the sequence (Burks, 1990; States and Boguski, 1990). The similarity of two biological sequences is usually specified by either a similarity measure or a distance measure. Similarity measures typically compute the score for an alignment using an optimizing function that rewards identities and penalizes nonidentities and gaps. Distance measures are usually defined as functions that satisfy the axioms for a metric function:
Distance(a,b) = 0 if and only if a = b; Distance(a,b) = distance(b,a); Distance(a,b) ≤ distance(a,c) + distance(c,b).
Distance functions can be thought of as minimizing differences to find the best alignments, whereas similarity methods search for the best alignments by maximizing similarities. “All problems known to be solvable with distance methods can be solved with similarity methods” (Waterman, 1989). The metric approach appeals to the geometric intuition: sequences can be thought of as points in space, and any three distinct points a, b, and c form a triangle ABC that satisfies the rules above. The smaller the distance between two points, the greater the similarity of the sequences. There are many factors to be considered in protein sequence comparison (Hodgman, 1992). Amino acids are not equally abundant in protein sequences. Moreover, proteins fold into structures having well-conserved hydrophobic interior residues and poorly conserved exterior residues. Indeed, there is a descending order of residue conservation: active-site residues > crucial folding residues > internal packing residues > surface residues. Some residues have preference for position, secondary structure, and substitution; for example, active-site residues are often found in loops that are part of a supersecondary unit and may be partially buried. An identity of >25% implies similarity of function, whereas 18% to 25% identity may indicate similarity of structure or function. The latter range is within the “Twilight Zone” (15% to 25% identity) that, according to Doolittle, arises in the theory of protein evolution (Doolittle, 1986). Alignment methods usually have a specific region of applicability (Taylor, 1988). For example, pairwise alignment is most applicable at high levels of identity (∼70%); consensus methods (∼45%), template methods (∼25%), and structure prediction methods are applicable at even lower levels of identity. These ranges take into account the fact that even two unrelated or random sequences over 100 residues long may show identity in the range of 20%. Two complementary properties of sequence comparison methods are sensitivity and specificity (States and Boguski, 1990). Sensitivity is the ratio of false negatives to the sum of true positives and false negatives; specificity is the ratio of true positives to the sum of true positives and false positives. The goal of this unit is to provide some
2.1.2 Current Protocols in Protein Science
starting points for the analysis of protein sequence data. A theoretical review of principles is complemented by practical examples of computer methods for protein sequence analysis that were done using the GCG package of programs (Genetics Computer Group, 1994). There are many other commercial packages for IBM-compatible computers, Macintoshes, and UNIX workstations that have similar capabilities. There are also many public domain software packages available over the Internet that have identical or enhanced functionalities (see Table 2.1.1). When starting to assemble programs for protein sequence analysis, one is strongly advised to browse the Internet before purchasing. The CD-ROM version of Current Protocols in Protein Science also includes the Entrez and MACAW multiple sequence alignment programs from the National Center for Biotechnology Information (NCBI) at no additional charge. UNIT 2.2 describes the analysis of linear protein sequence with respect to hydrophobic/hydrophilic characterization that can suggest structural or functional segments. UNIT 2.3 presents methods for the prediction of secondary structural features of linear protein sequences.
MATRIX METHODS FOR SEQUENCE COMPARISON: DOT PLOTS Many computer-based sequence comparison methods rely on matrix methods (Fitch, 1966, 1969; Gibbs and McIntyre, 1970; McLauchlan, 1971, 1972; Maizel and Lenk, 1981; Argos, 1987; Landau et al., 1988, 1990). Pairwise comparison of two sequences can be represented in its simplest form by a matrix that is the logical product of the two sequences: if a corresponding pair of elements is identical, then the matrix element at the intersection of the row and column is marked true (dot); if the pair is different, then the matrix element is marked false (blank). The result is a visualization of the similarity of the two sequences. In dot-plot matrix displays, a run of identities will be seen as a continuous diagonal segment within the matrix. Discontinuities along diagonals with horizontal shifts to parallel lines are interpreted as indels and indicate the need for gaps in any alignment. If the two sequences being compared are identical, the matrix will reveal any internal repeats and the main diagonal will be filled in. Self-comparisons always result in symmetrical dot plots. Another commonly used dot-matrix method compares fixed-length segments (windows) of
the sequences instead of individual residues. See Staden (1994a-d) for a description of how to use SIP, a program based on dot-matrix plots. A palindrome is a subsequence that remains the same after being reversed. On a dot plot, palindromes are displayed as segments perpendicular to the main diagonal of the matrix. Palindromes are not usually relevant to protein sequences. In RNA and DNA sequences, a palindrome is defined as an inverted, self- co mp lementary region such as ACTGATATCAGT. More sophisticated comparisons have been defined on dot matrices that filter out isolated dots or small segments as noise. The stringency of the similarity determines the amount of data that will be filtered out before the matrix is displayed or undergoes further processing. In this case, the matrix elements are filled in with weighted numerical scores instead of simple identity values, and the result is plotted as a surface to visualize the regions of greatest similarity. Dot-matrix methods remain an active area of research (Streletc et al., 1991; Landes et al., 1993; Nedde and Ward, 1993). Dot plots are the simplest form of sequence comparison using a matrix display. These graphical methods allow quick observation of the level and extent of similarity between two sequences. For example, a comparison of horse and rat serum albumin protein sequences from the SWISS-PROT protein database is summarized in Figure 2.1.1. Dot-plot analysis is done in two phases: pairwise sequence comparison to generate a matrix of similarity values is followed by generation of the graphic display. The GCG package implements this process by using two programs—COMPARE and DOTPLOT—to generate the dot-matrix plot. COMPARE does a pairwise analysis to define positions of similarity between the two sequences. The parameters to be varied are (1) window size, or the number of residues to be compared, and (2) stringency, or the minimal number of matches per window to score a dot. DOTPLOT generates the graphic matrix display of the COMPARE analysis (Fig. 2.1.1).
SEQUENCE SIMILARITY SEARCHING Once a small portion of protein sequence data from an unknown sample has been generated, the immediate impulse is to ask what other proteins in the database does the unknown resemble, and how similar are the sequences. Several available tools can help answer these
Computational Analysis
2.1.3 Current Protocols in Protein Science
600
Albu_Horse.Sw ck: 307, 1 to 607
1, 1995 17:05
Points: 4,481 Stringency: 18.0
February
400
600
Window: 60
Density: 692.05
200
400
200
COMPARE
DOTPLOT of: albu_rat.pnt
0
0 Albu_Rat ck: 6,355, 1 to 608
Figure 2.1.1 Dot plot generated from comparison of rat and horse albumin protein sequences. The continuous diagonal line of identity traverses the middle of the matrix. Regions of similarity between the two sequences appear as lines parallel and offset to the line of identity. Axis label takes the form filename ck: checksum_value start_position to end_position.
questions. Most commercial sequence analysis packages contain programs for performing pairwise alignments of protein sequences. The most popular tools for searching databases, FASTA (FAST All) and BLAST (the Basic Local Alignment Search Tool), are included in commercial sequence analysis suites and are also freely available to be downloaded to a local computer to be used in a stand-alone manner or as network services.
BLAST Database Searching
Computational Methods for Protein Sequence Analysis
The program BLAST, which is available from the NCBI, conducts rapid searching of sequence databases (Altschul et al., 1990). The program searches for similarity between a query sequence and sequences in a nucleotide database (using BLASTN) or peptide database (using BLASTP). Two versions of the program are available: (1) a version that will run on a local workstation and (2) client software that connects a local computer via the Internet to a high-speed search engine located at the NCBI.
The latter accesses the most recent versions of sequence databases available. BLAST uses a word/dictionary approach to compare fixed-length “words” from the query sequence against the dictionary of words from the database searched. This program is an excellent choice for quickly screening for exact matches—for example, when the end of a new protein has been sequenced. A word of caution is, however, in order: BLAST can create enormous files, especially if the query sequence has repeated residues or is similar to a repeated motif in the databases. The BLAST output file contains (1) the sorted match statistics, probability measure of a random match and the number of matched regions; (2) the alignments generated for the query and the matched sequence region (shown in Fig. 2.1.2); and (3) a copy of the search parameters (not shown). The example in Figure 2.1.2 shows output from a BLAST search conducted using a sequence fragment from an unknown mammalian blood protein as a query against the nonredundant
2.1.4 Current Protocols in Protein Science
Similarity Table
Sequences producing High-scoring Segment Pairs: sp|P35747|ALBU_HORSE sp|P08835|ALBU_PIG gp|M36787|PIGALBA_1 gp|X17055|OASERALB_4 sp|P14639|ALBU_SHEEP sp|P02769|ALBU_BOVIN pir|A47391|A47391 gp|M73993|BOVALBUMIN_1 gp|V01222|RNALBU_4 sp|P02770|ALBU_RAT pir|S17599|S17599 pir|S29749|S29749 gp|A15293|A15293_1 gp|A00279|A00279_1 gp|A16010|A16010_1 gp|A03758|A03758_1 gp|M12523|HUMALBGC_1 gp|A09561|A09561_1 sp|P02768|ALBU_HUMAN pir|A93743|ABHUS
SERUM ALBUMIN PRECURSOR. >pir|S340.. SERUM ALBUMIN PRECURSOR (FRAGMENT).. albumin [Sus scrofa] Sheep mRNA for serum albumin. [Ovi.. SERUM ALBUMIN PRECURSOR. >pir|S069.. SERUM ALBUMIN PRECURSOR. >pir|A388.. serum albumin prepropeptide - rhes.. albumin [Bos taurus] Messenger RNA for rat preproalbumi.. SERUM ALBUMIN PRECURSOR. >pir|A938.. albumin - human serum albumin - dog Mature HSA [Homo sapiens] Human serum albumin [Unknown] synthetic mature HSA [Unknown] serum albumin [Homo sapiens] Human serum albumin (ALB) gene, co.. human serum albumin [Unknown] SERUM ALBUMIN PRECURSOR. >gp|M1252.. serum albumin precursor - human >g..
High Score 101 98 98 96 96 95 94 94 92 92 79 79 85 85 85 85 85 85 85 85
Smallest Sum Probability P(N) N 6.8e-08 1.8e-07 1.8e-07 3.4e-07 3.5e-07 4.8e-07 6.6e-07 6.6e-07 1.3e-06 1.3e-06 1.8e-06 4.0e-06 1.2e-05 1.2e-05 1.2e-05 1.2e-05 1.2e-05 1.2e-05 1.2e-05 1.2e-05
1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1
Alignment >sp|P35747|ALBU_HORSE SERUM ALBUMIN PRECURSOR. >pir|S34053|ABHOS serum albumin precursor - horse >gp|X74045|ECSERALB_1 preproalbumin [Equus caballus] Length = 607 Score = 101 (47.0 bits), Expect = 6.8e-08, P = 6.8e-08 Identities = 22/26 (84%), Positives = 22/26 (84%) Query: Sbjct:
1 TIKSEIAHVFNDLGEPVFKGLVLVAF 26 T KSEIAH FNDLGE FKGLVLVAF 26 THKSEIAHRFNDLGEKHFKGLVLVAF 51
Figure 2.1.2 BLAST output table and alignment for an unknown mammalian blood protein. The Query is the sequence of interest and the sbjct is the sequence region found in the database that matches the query. Note that in searches of the nonredundant databases at the NCBI, matches are found between the test sequence and the translation of mRNAs in GenBank. Important similarities have small probability scores, i.e., P(N) < 10−3. This analysis suggests that the unknown is an albumin, but it is not identical to any known sequence.
protein sequence database maintained at the NCBI. This database contains a compilation of protein sequences found in all of the molecular sequence databases with duplicate sequences removed. Figure 2.1.2 presents a table of the top 20 highest similarity scoring sequences and the best alignment.
FASTA Database Searching FASTA conducts a Lipman and Pearson (1985) search for similarity between a query sequence and any FASTA-formatted database or group of sequences collected in a database by the user. FASTA is a program developed for fast searching using an algorithm that allows for the insertion of gaps during the alignment
phase. This approximates evolutionary insertions and deletions during divergence. However, the gaps are generated to maximize the number of aligned residues. FASTA does a good job of generating a global alignment for a sequence. Figure 2.1.3 and Figure 2.1.4 present results from a FASTA search of the SWISS-PROT database with a short sequence fragment from an unknown mammalian blood protein. The FASTA output contains (1) a histogram that shows the distribution of relevant word matches found between the query sequence and the database searched; (2) the match statistics; and (3) the alignments generated for the query sequence and the matched sequence region. There
Computational Analysis
2.1.5 Current Protocols in Protein Science
Score Init1 Initn (-) (+) < 2 533 533:================================================== 4 4 4:== 6 134 134:================================================== 8 222 222:================================================== 10 1964 1964:================================================== 12 5289 5289:================================================== 14 7609 7609:================================================== 16 9434 9434:================================================== 18 2834 2834:================================================== 20 4914 4914:================================================== 22 2686 2686:================================================== 24 1808 1808:================================================== 26 1241 1241:================================================== 28 721 721:================================================== 30 375 374:================================================== 32 261 259:================================================== 34 138 137:================================================== 36 57 57:============================= 38 27 27:============== 40 18 18:========= 42 4 5:==+ 44 6 7:===+ 46 0 0: 48 0 1:+ 50 3 4:== 52 1 1:= 54 1 1:= 56 2 2:= 58 0 0: 60 0 0: 62 0 0: 64 0 0: 66 0 0: 68 0 0: 70 0 0: 72 0 0: 74 0 0: 76 0 0: 78 0 0: 80 0 0: > 80 6 6:=== Mean score calculations exclude scores greater than 72 mean initn score: 16.3 (s.d. 5.20) mean init1 score: 16.3 (s.d. 5.19) 1838 scores better than 26 saved, joining threshold: 27
Figure 2.1.3 FASTA histogram from a search of the SWISS-PROT database for sequences similar to a fragment from an unknown mammalian blood protein. Numbers of windows at each init1 score are plotted. Note that there are six highly significant alignments, as well as several sequences (those with init1 scores in the 48 to 56 range) that represent protein sequences from more distantly related species.
Computational Methods for Protein Sequence Analysis
are three similarity scores presented in the output from FASTA: init1, the highest similarity score allowing conservative substitutions; initn, the highest score generated by linking nonoverlapping regions; and opt, the score for the best alignment presented. In practice the histogram would be used to determine how many sequences have highly significant levels of similarity. The table of scores allows one to decide quickly how many and which sequence alignments should be viewed.
ALIGNMENT METHOD CONSIDERATIONS Dynamic programming is a different matrix method that is used to find the optimal global alignment of two sequences (Needleman and Wünsch, 1970; Sellers, 1974; Pearson and Miller, 1992; Panjukov, 1993). Let a be a sequence of length m and let b be a sequence of length n. Let D be the working matrix. Then, given a predefined scoring function, the basic steps of the alignment algorithm (Doolittle, 1986) are as follows. Assign the scores for all exact matches to the corresponding matrix el-
2.1.6 Current Protocols in Protein Science
Alignment Score Table The best scores are: init1 initn opt Sw:Albu_Horse P35747 equus caballus (horse). serum album... 99 99 100 Sw:Albu_Rat P02770 rattus norvegicus (rat). serum albumi... 99 99 99 Sw:Albu_Pig P08835 sus scrofa (pig). serum albumin precu... 99 99 101 Sw:Albu_Bovin P02769 bos taurus (bovine). serum albumin ... 98 98 99 Sw:Albu_Sheep P14639 ovis aries (sheep). serum albumin p... 95 95 96 Sw:Albu_Human P02768 homo sapiens (human). serum albumin... 93 93 93 Sw:Albu_Chick P19121 gallus gallus (chicken). serum albu... 56 56 77 Sw:Albu_Mouse P07724 mus musculus (mouse). serum albumin... 55 55 55
Best Scoring Alignment SCORES
Init1: 99 Initn: 99 Opt: 84.6% identity in 26 aa overlap
100
10 20 TIKSEIAHVFNDLGEPVFKGLVLVAF | |||||| |||||| ||||||||| Albu_H MKWVTFVSLLFLFSSAYSRGVLRRDTHKSEIAHRFNDLGEKHFKGLVLVAFSQYLQQCPF 10 20 30 40 50 60 Pc12_P
Figure 2.1.4 Partial FASTA alignment table and best scoring alignment for the same search illustrated in Figure 2.1.3. The table shows the best alignment scores sorted by highest init1 score.
ements. Transform the matrix by beginning with element Dmn and adding its value (less any gap penalty) to Dm−1, n−1 and all the elements directly above it or to its left. Next, select the matrix element in column m − 2 and above row m − 1 that contains the largest score and repeat the summing procedure. Repeat the previous step until the entire matrix is transformed. Next, determine the path from the largest score in the matrix back toward the bottom right-hand corner. The path, with any gaps as indicated by off-diagonal moves, is then used to align a and b. This matching algorithm is used in most global alignment programs, which vary primarily in the sophistication of the scoring functions (similarity or distance). Note that this method assumes (incorrectly) that each residue has equal importance, and it ignores sequential constraints. Local alignments are usually found using some version of the Smith-Waterman method (1981), which applies when common regions are being sought in two sequences whose similarity is unknown or limited to a few regions, as is the case in database searches. The algorithm usually produces a single highest-scoring alignment with respect to the operations in E, although it has been successfully extended to produce the second-best and other alignments (Barton, 1990, 1993; Waterman and Eggert, 1991). In its simplest form (Barton, 1993), the local alignment method is as follows. Given two
sequences a and b, and a working (comparison) matrix H, the matrix elements of H are filled in by applying a scoring rule: the score assigned to a matrix element Hij is either zero or the maximum value of any of the three scores of its neighbors and predecessors that lie directly above, diagonally to the left, or directly to the left of Hij, adjusted for gaps or substitution of residues. The maximum element in H is found and traced back through the matrix to find the alignment.
SCORING MATRICES Scoring matrices (George et al., 1990; States and Boguski, 1990), such as the PAM, BLOSUM (Henikoff and Henikoff, 1993), and related MDM matrices (Dayhoff, 1978), are based on the compilation of frequencies of amino acid substitution in a set of homologous sequences. The data are used as an amino acid transition probability matrix. Increasing powers of the matrix correspond to increasing divergence. Lower powers are used for sequences that seem closely related and higher powers for those that are not so obviously related. Thus, PAM250 represents a matrix with 250 accepted protein mutations per 100 amino acids. The scoring matrix method has greatly improved the quality and sensitivity of alignments; however, it does have the deficiency that it ignores structure and the corresponding sequence constraints which are significant for predicting structure. Not all sequence comparison methods are
Computational Analysis
2.1.7 Current Protocols in Protein Science
Computational Methods for Protein Sequence Analysis
matrix methods. Programs such as FASTP and FASTN are representative of hash-coding methods (Dumas and Nunio, 1982; Wilbur and Lipman, 1983; Lipman and Pearson, 1985; Pearson, 1990, 1994). The methods are based on fast table lookups of subsequences of a target sequence using hash functions. Hash functions transform strings in numbers that serve as pointers into a table containing further information. Thus, a fixed-length subsequence (or k-tuple) of one sequence can quickly be checked for occurrence in another sequence after a hash table of k-tuples and their positions in the sequence is constructed for each sequence. FASTP focuses on regions having the highest density of identities and uses the PAM250 matrix. Readers are referred to Pearson (1994) for an introduction to using FASTA for comparing both DNA and protein sequences. The RDF2 and RSS programs for testing the statistical significance of FASTA alignments are also discussed. Other methods for sequence comparison include finite state machines that are capable of recognizing regular expressions, and various statistical approaches (States and Boguski, 1990; Tyler et al., 1991; Allison et al., 1992; Brendel et al., 1992). Regular expressions are formulas such as (a + b*)c, which represents the set of all strings that begin with the letter a or zero or more b’s and followed by a single c. Complicated patterns can be constructed and detected in sequences using the finite state method. Statistical approaches include the database-searching program BLAST (see above section on BLAST database searching), which approximates gapless alignments that optimize local similarity on the basis of maximal segment pairs. The BLAST strategy is to search for only those segment pairs that contain a word pair whose comparison score exceeds some threshold. It uses the PAM120 scoring matrix to allow for conservative substitutions of residues in a comparison. Statistical methods for sequence alignment (Karlin et al., 1988, 1989, 1991; Brendel, 1992; Pevzner, 1992; Mrazek and Kypr, 1993; Staden, 1994a-d) consider the statistical significance of a variety of protein sequence properties. Such properties include the relative composition of a sequence with respect to residue types; anomalous clusters, runs, and periodic variations of charge; internal repeats; local periodicities that might indicate regularity of structure; and the relative spacings of certain residue types. For a description of routines for
plotting hydrophobicity, charge, and hydrophobic moments using the program PIP, see Staden (1994a-d). In addition to the methods described above, interest in the development of theory and tools for linguistic (formal grammar) analysis of biological sequences is growing (Collado-Vides, 1991; Michaels et al., 1993; Searls, 1993). The next level of sequence analysis is the simultaneous comparison of more than two sequences, as described in the following section on Multiple Alignments.
MULTIPLE ALIGNMENTS Once a group of potentially homologous sequences has been identified, the next step could be to do a multiple alignment that compares the new sequence with the group of sequences determined to have highly significant similarities. There are several programs that will do multiple alignments. PILEUP from the GCG package does a good job with closely related sequences in an automated fashion. MACAW from NCBI, LINEUP from GCG, and DSCE (De Rijk and De Wachter, 1993) allow the investigator to generate an alignment manually. (The CD-ROM version of Current Protocols in Protein Science includes MACAW and Entrez from NCBI.) Figure 2.1.5 demonstrates a multiple alignment generated using PILEUP from five highscoring albumin sequences in the SWISSPROT database identified from the FASTA analysis (Fig. 2.1.4). PILEUP performs a cluster analysis based on pairwise comparisons of each sequence to be aligned. As well as the multiple alignment (Fig. 2.1.5), PILEUP produces a dendrogram (not shown) that displays the clustered relationships between the sequences. The dendrogram is not a phylogenetic tree, but a graphical representation of the clustered similarity statistics. After the multiple alignment is generated, one can begin to infer relationships between conserved regions and function or structure. The program Pro_Anal (Eroshkin et al., 1993) and the program for fast protein block searches (Fuchs, 1994) can help in this process. Programs like OBSTRUCT (Heringa et al., 1992) are useful in identifying clusters of protein sequences on the basis of structural and sequence similarity criteria. Kanaoka et al. (1989) have described a process of aligning protein sequences on the basis of hydrophobicity. Livingstone and Barton (1993) describe a strategy to generate multiple alignments for hierarchical analysis of residue conservation.
2.1.8 Current Protocols in Protein Science
Albu_Bovin.Sw Albu_Chick.Sw Albu_Horse.Sw Albu_Human.Sw Albu_Mouse.Sw
1 MKWVTFISLL MKWVTLISFI MKWVTFVSLL MKWVTFISLL EADKSEIADR
LLFSSAYSRG FLFSSATSRN FLFSSAYSRG FLFSSAYSRG YNDLGEIHFK
VFRRDTHKSE LQRFARDAEH VLRRDTHKSE VFRRDAHKSE CAIPNLRENY
IAHRFKDLGE KSEIAHRYND IAHRFNDLGE VAHRFKDLGE GELADCCTKQ
50 EQFKGLVLIA LKEETFKAVA KHFKGLVLVA ENFKALVLIA EPERNECFLQ
Albu_Bovin.Sw Albu_Chick.Sw Albu_Horse.Sw Albu_Human.Sw Albu_Mouse.Sw
51 FSQYLQQCPF MITFAQYLQR FSQYLQQCPF FAQYLQQCPF HKDDNPSLPP
DEHVKLVNEL CSYEGLSKLV EDHVKLVNEV EDHVKLVNEV FERPEAEAMC
TEFAKTCVAD KDVVDLAQKC TEFAKKCAAD TEFAKTCVAD TSFKENPTTF
ESHAGCEKSL VANEDAPECS ESAENCDKSL ESAENCDKSL MGHYLHEVAR
100 HTLFGDELCK KPLPSIILDE HTLFGDKLCT HTLFGDKLCT RHPYFYAPEL
Albu_Bovin.Sw Albu_Chick.Sw Albu_Horse.Sw Albu_Human.Sw Albu_Mouse.Sw
101 VASLRETYGD ICQVEKLRDS VATLRATYGE VATLRETYGE LYYAEQYNEI
MADCCEKQEP YGAMADCCSK LADCCEKQEP MADCCAKQEP LTQCCAEADK
ERNECFLSHK ADPERNECFL ERNECFLTHK ERNECFLQHK ESCLTPKLDG
DDSPDLPKLK SFKVSQPDFV DDHPNLPKLK DDNPNLPRLV VKEKALVSSV
150 PDPNTLCDEF QPYQRPASDV PEPDAQCAAF RPEVDVMCTA RQRMKCSSMQ
Albu_Bovin.Sw Albu_Chick.Sw Albu_Horse.Sw Albu_Human.Sw Albu_Mouse.Sw
151 KADEKKFWGK ICQEYQDNRV QEDPDKFLGK FHDNEETFLK KFGERAFKAW
YLYEIARRHP SFLGHFIYSV YLYEVARRHP KYLYEIARRH AVARLSQTFP
YFYAPELLYY ARRHPFLYAP YFYGPELLFH PYFYAPELLF NADFAEITKL
ANKYNGVFQD AILSFAVDFE AEEYKADFTE FAKRYKAAFT ATDLTKVNKE
200 CCQAEDKGAC HALQSCCKES CCPADDKLAC ECCQAADKAA CCHGDLLECA
Albu_Bovin.Sw Albu_Chick.Sw Albu_Horse.Sw Albu_Human.Sw Albu_Mouse.Sw
201 LLPKIETMRE DVGACLDTKE LIPKLDALKE CLLPKLDELR DDRAELAKYM
KVLASSARQR IVMREKAKGV RILLSSAKER DEGKASSAKQ CENQATISSK
LRCASIQKFG SVKQQYFCGI LKCSSFQNFG RLKCASLQKF LQTCCDKPLL
ERALKAWSVA LKQFGDRVFQ ERAVKAWSVA GERAFKAWAV KKAHCLSEVE
250 RLSQKFPKAE ARQLIYLSQK RLSQKFPKAD ARLSQRFPKA HDTMPADLPA
Albu_Bovin.Sw Albu_Chick.Sw Albu_Horse.Sw Albu_Human.Sw Albu_Mouse.Sw
251 FVEVTKLVTD YPKAPFSEVS FAEVSKIVTD EFAEVSKLVT IAADFVEDQE
LTKVHKECCH KFVHDSIGVH LTKVHKECCH DLTKVHTECC VCKNYAEAKD
GDLLECADDR KECCEGDMVE GDLLECADDR HGDLLECADD VFLGTFLYEY
ADLAKYICDN CMDDMARMMS ADLAKYICEH RADLAKYICE SRRHPDYSVS
300 QDTISSKLKE NLCSQQDVFS QDSISGKLKA NQDSISSKLK LLLRLAKKYE
Albu_Bovin.Sw Albu_Chick.Sw Albu_Horse.Sw Albu_Human.Sw Albu_Mouse.Sw
301 CCDKPLLEKS GKIKDCCEKP CCDKPLLQKS ECCEKPLLEK ATLEKCCAEA
HCIAEVEKDA IVERSQCIME HCIAEVKEDD SHCIAEVEND NPPACYGTVL
IPENLPPLTA AEFDEKPADL LPSDLPALAA EMPADLPSLA AEFQPLVEEP
DFAEDKDVCK PSLVEKYIED DFAEDKEICK ADFVESKDVC KNLVKTNCDL
350 NYQEAKDAFL KEVCKSFEAG HYKDAKDVFL KNYAEAKDVF YEKLGEYGFQ
Albu_Bovin.Sw Albu_Chick.Sw Albu_Horse.Sw Albu_Human.Sw Albu_Mouse.Sw
351 GSFLYEYSRR HDAFMAEFVY GTFLYEYSRR LGMFLYEYAR NAILVRYTQK
HPEYAVSVLL EYSRRHPEFS HPDYSVSLLL RHPDYSVVLL APQVSTPTLV
RLAKEYEATL IQLIMRIAKG RIAKTYEATL LRLAKTYETT EAARNLGRVG
EECCAKDDPH YESLLEKCCK EKCCAEADPP LEKCCAAADP TKCCTLPEDQ
400 ACYSTVFDKL TDNPAECYAN ACYRTVFDQF HECYAKVFDE RLPCVEDYLS
Figure 2.1.5 Multiple alignment of mammalian blood proteins using PILEUP. The first 400 residues of five high-scoring sequences identified from FASTA analysis (Fig. 2.1.4) are aligned.
Similarly, the program PIMA (Smith and Smith, 1992) uses an algorithm that takes advantage of secondary structure features to determine gap penalties during the multiple alignment process. Depiereux and Feytmans (1991) have used simultaneous multivariate multiple alignment methods to reveal a correspondence between physiochemical profiles and structurally conserved regions. The multiple alignment programs described above take advantage of pairwise comparison
algorithms and are subject to their limitations. Alignment algorithms that use a rigorous consideration of more than three sequences at once are computationally infeasible for most labs. Although some speed improvements have been obtained for the optimal alignment of three sequences simultaneously (Murata et al., 1985; Gotoh, 1986), new methods have to be developed for multiple sequence alignment. The earliest methods were sequence editors for interactive alignments. See Pearson and Miller
Computational Analysis
2.1.9 Current Protocols in Protein Science
Computational Methods for Protein Sequence Analysis
(1992) for a discussion of considerably improved dynamic programming algorithms for sequence analysis. Multiple sequence alignment tends to be order-sensitive. A sequence of pairwise comparisons may not lead to the best overall alignment for a given set of sequences because the sequences are not independent (States and Boguski, 1990)—that is, they are not necessarily equidistant from a common ancestor. The consensus sequence approach to multiple sequence alignment (Waterman, 1989, 1990; Waterman and Jones, 1990; Day and McMorris, 1993) is based on the construction of a single sequence that can be used to align all other given sequences; the sequence is determined by an iterative process of alignment followed by consensus sequence modification. A consensus residue can be stringently defined from a multiple alignment when the content of a column is represented by a single amino acid at that position. There are many multiple alignment algorithms—too many to describe here. A list of available sequence similarity and alignment software can be accessed via the Internet as described in the section below on Internet Resources. Once sequences have been aligned, the quality of the alignment should be assessed. It is clearly important to distinguish between chance similarities and true sequence similarities. One method for estimating the quality or statistical significance of an alignment (Doolittle, 1986) is to compare the alignment score with the mean score of a set of randomized sequences that are permutations of the original sequences and which preserve their length and relative composition. The further the alignment score is from the mean of the randomized set, the more confident we can be in the alignment. A possible objection to this data shuffling or permutation method is that the permuted sequences are not like real protein sequences (i.e., permuted sequences are unlikely to fold like real proteins). Other methods (States and Boguski, 1990) include the following: McLachlan’s (1972) “double matching probability” function, which is used to compute the frequency scores in infinite random sequences of the same composition as the given sequences; Reich and Meiske’s stochastic random sequence model for determining the significance of window scores in dot-matrix analysis (Reich and Meiske, 1987); and Argos’ significance tests (Argos and Vingron, 1990), based on actual protein sequences
instead of the randomized sequences used by McLachlan (1972). Despite efforts to determine the quality of alignments, it remains true that low statistical significance does not always imply a lack of biological significance (Collins and Coulson, 1990). Another important aspect of alignments is the accuracy of the original molecular sequence (States, 1992). Errors can be introduced in molecular sequence data in random or biased ways. The translation of nucleic acid sequences into amino acid sequences is very sensitive to indel errors: an error rate of 1% can lead to a frameshifting error in reading frames longer than 24 amino acids. Empirical estimates of sequence accuracy in current databases have revealed a rate of 3 to 5 errors per 1000 bases.
CLUSTER METHODS AND TREES Once a multiple alignment has been performed and regions of common similarity have been identified among the proteins, the question of evolutionary distance will arise. There are several programs for determining the level of divergence between a group of sequences. Some are available over the Internet via the World Wide Web (WWW; see also Internet Resources section). The programs DISTANCES and GROWTREE from the GCG package (Genetics Computer Group, 1994) calculate pairwise distance measures and plot a dendrogram of those measures. The programs depend on an accurate multiple alignment of the sequences involved. This means that the multiple alignments should be examined carefully and then edited as necessary to incorporate additional knowledge that might be available about the structure or function of the proteins. Figure 2.1.6 shows the table of pairwise phylogenetic distances calculated by DISTANCES for the top nine albumins identified from the FASTA analysis (Fig. 2.1.4), and the phylogram produced by GROWTREE is shown in Figure 2.1.7. The purpose of protein sequence comparison is to learn more about the structure, function, and evolution of proteins. It is known that different proteins and different parts of proteins change at characteristic but unequal rates in the course of evolution (Doolittle, 1986). Binding sites and catalytic units are conserved independently of their position, whereas surface regions change faster than buried regions. In addition, entire domains of a genome are subject to a number of editing operations such as domain joining, swapping, and reordering (States and Boguski, 1990). Thus, homologies
2.1.10 Current Protocols in Protein Science
Matrix 1, dimension: 9 Key for column and row indices: 1 2 3 4 5 6 7 8 9
Albu_Bovin.Sw Albu_Chick.Sw Albu_Horse.Sw Albu_Human.Sw Albu_Mouse.Sw Albu_Pig.Sw Albu_Ranca.Sw Albu_Rat.Sw Albu_Sheep.Sw
1 2 3 4 5 6 7 8 9 __________________________________________________________________________ .. | 1 | 0.00 999.99 31.75 192.01 999.99 999.99 999.99 237.70 8.19 | 2 | 0.00 999.99 999.99 999.99 999.99 999.99 999.99 999.99 | 3 | 0.00 190.56 999.99 999.99 999.99 222.61 29.05 | 4 | 0.00 999.99 999.99 999.99 32.75 193.48 | 5 | 0.00 999.99 999.99 999.99 999.99 | 6 | 0.00 999.99 999.99 999.99 | 7 | 0.00 999.99 999.99 | 8 | 0.00 237.70 | 9 | 0.00
Figure 2.1.6 Evolutionary distance table for mammalian blood proteins calculated using the GCG DISTANCES program. Corrections to the calculations were done using the Kimura protein method as described in the GCG manual (Genetics Computer Group, 1994). The matrix values represent the number of matches between each pair of sequences divided by its length (999.9 = identity). These data were used to generate the GROWTREE phylogram presented in Figure 2.1.7.
Growtree Phylogram of: Albu_Bovin.Distances, Tree Tree_ February 3, 1995 15:02 Albu_Ranca.Sw Albu_Pig.Sw Albu_Mouse.Sw Albu_Chick.Sw Albu_Rat.Sw Albu_Human.Sw Albu_Horse.Sw Albu_Sheep.Sw Albu_Bovin.Sw
Figure 2.1.7 GROWTREE phylogram of Albu_Bovin.Distances, Tree Tree_1. Results of graphical cluster analysis of the distance data presented in Figure 2.1.6 are shown. In this representation sequences with the least divergence have the shortest branches.
Computational Analysis
2.1.11 Current Protocols in Protein Science
might be found between entire proteins or just between certain parts of proteins.
IDENTIFICATION OF FUNCTIONAL SITES
Computational Methods for Protein Sequence Analysis
Sequence comparisons are used to test for homology and to find functional sites and motifs. The goal is to establish structure and function for parts of an unknown sequence using known homologous or analogous (i.e., similar, but not homologous) sequences as sources of additional information. The goal is to obtain answers to questions of what functions the protein has and what its structure is. There are several paths the analysis can take. One approach that will reveal much about potential functional sites in the protein is to look for motifs. Motifs are defined in three ways: as consensus patterns of amino acids that correspond to regions of proteins containing the common residues; as the excised alignment of several sequences that contain the motif for use as a database discriminator; or as a single representative sequence (States and Boguski, 1990; Parry-Smith and Atwood, 1992). Motif searches are usually done using a program that takes advantage of a dictionary of sequence patterns which have been identified with specific protein activities. Many programs will do this analysis, and most use the PROSITE database of motifs found through analysis of the SWISS-PROT protein database. Figure 2.1.8 shows an example using the GCG MOTIFS program to analyze an albumin sequence. Note that the output file contains an alignment of the motif found and an extensive annotation describing the motif and listing the associated bibliography. The PIPL program developed by Staden (1994a-d) searches libraries of sequences to find patterns, where a pattern is a set of motifs with variable spacing. Patterns are constructed using logical operators to form a search term that is then used to scan a database such as PROSITE (Bairoch, 1992). The MOTIFS program of the Genetics Computer Group (1994) uses PROSITE as a dictionary of motifs. Weight matrices are used in scanning for motifs and developing a motif pattern. The profile analysis method (Gribskov et al., 1988, 1990; Gribskov, 1994) also involves weight matrices and is related to the consensus method. It is used for sensitive similarity matching, for homology seeking, and for scanning databases for motifs. It measures the similarity between a given sequence, called the target, and a group of aligned sequences called
the probe. The alignment of the probe uses a modified MDM78 mutational matrix. The matrix values of the profile are computed columnwise as the weighted average of the score for each residue at that position in the aligned sequences and the residue represented by the column of the profile; a row of the profile is the weighted average of the rows in the MDM78 table corresponding to the aligned residues at that position. The result is the set of properties that tend to be conserved at particular positions along the sequence. For detailed information on using profile analysis software to scan for homologies and motifs, see Gribskov (1994). A relatively new homologous sequence retrieval program (using the BLOSUM250 matrix) is dFLASH, which is available by E-mail server (Rigoutsos and Califano, 1994). There are many ways to proceed in computational analysis of new sequences. Each will provide a wealth of information to guide further laboratory experimentation. The following procedures illustrate two approaches to protein sequence analysis and can be adapted as required. Hodgman (1992) suggests the following protocol for protein function determination: 1. Split sequences into overlapping 200- to 300-residue segments. 2. Use a program such as FASTA to search a protein database for sequences similar to those in the segments. 3. If a match having >25% identity over 80 residues is found, then the search has been successful. 4a. For weaker matches, use dot-matrix plots to verify similarity. 4b. Alternatively, search the database again with the matching sequence to find related sequences for alignment to detect conserved positions. 5. Conserved regions can be used to search further with PIPL (Staden, 1994a-d), PROFILESEARCH (GCG), or SCRUTINEER (EMBL). 6. Use any biological information about the conserved regions to determine an analogous function for the test protein. 7. If the procedure has not been successful, use other resources such as PROSITE. 8. Check that all results are biologically credible. 9. If all else fails, (a) look for repeats within the sequence itself: if there are only two or three, it usually indicates that the protein has a repeated functional domain; (b) examine the distribution of particular amino acids or classes
2.1.12 Current Protocols in Protein Science
Motifs Analysis Atp_Gtp_A
(A,G)x4GK(S,T) (G)x{4}GK(S) 425: HIVLS GESYSGKS TNARL
the motif from the Prosite database the pattern matched in this sequence the acutal region of sequence containing the motif
Attached annotation ***************************************** * ATP/GTP-binding site motif A (P-loop) * ***************************************** From sequence comparisons and crystallographic data analysis it has been shown [1,2,3,4,5] that an appreciable proportion of proteins that bind ATP or GTP share a number of more or less conserved sequence motifs. The best conserved of these motifs is a glycine-rich region, which probably forms a flexible loop between a betastrand and an alpha-helix. This loop interacts with one of the phosphate groups of the nucleotide. This sequence motif is generally referred to as the 'A' consensus sequence [1] or the 'P-loop' [5]. There are numerous ATP- or GTP-binding proteins in which the P-loop is found.We list below a number of protein families for which the relevance of the presence of such motif has been noted:
A list of proteins containing the motif has been omitted for brevity. Not all ATP- or GTP-binding proteins are picked-up by this motif. A number of proteins escape detection because the structure of their ATP-binding site is completely different from that of the P-loop. Examples of such proteins are the E1-E2 ATPases or the glycolytic kinases. In other ATP- or GTP-binding proteins the flexible loop exists in a slightly different form; this is the case for tubulins or protein kinases. A special mention must be reserved for adenylate kinase, in which there is a single deviation from the P-loop pattern: in the last position Gly is found instead of Ser or Thr. -Consensus pattern: [AG]-x(4)-G-K-[ST] -Sequences known to belong to this class detected by the pattern: a majority. -Other sequence(s) detected in SWISS-PROT: in addition to the proteins listed above, the 'A' motif is also found in a number of other proteins. Most of these proteins probably bind a nucleotide, but others are definitively not ATP- or GTP-binding (as for example chymotrypsin, or human ferritin light chain). -Expert(s) to contact by email: Koonin E.V.
[email protected] -Last update: June 1994 / Text revised. [ 1] Walker J.E., Saraste M., Runswick M.J., Gay N.J. EMBO J. 1:945-951(1982). [ 2] Moller W., Amons R. FEBS Lett. 186:1-7(1985). [ 3] Fry D.C., Kuby S.A., Mildvan A.S. Proc. Natl. Acad. Sci. U.S.A. 83:907-911(1986). [ 4] Dever T.E., Glynias M.J., Merrick W.C.
Figure 2.1.8 MOTIFS analysis of an albumin sequence. This example shows results for a single motif found in the test protein. If the query sequence contains many motifs, the output can be quite extensive. Note that a long table of sequences which contain the motif and some of the references have been deleted due to space constraints.
for suggested functional sites; or (c) apply biological and biochemical knowledge to evaluate the composition of the sequence (e.g., regions rich in basic residues suggest the binding of nucleic acids, and acidic regions suggest generalized protein-protein interactions). Structure determination can begin with the ChouFasman (Chou and Fasman, 1974) or GarnierOsguthorpe-Robson algorithm (Garnier et al., 1978); both are discussed in UNIT 2.3. States and Boguski (1990) offer another scheme for the analysis of protein sequences:
1. Find all intrasequence repeats using dotmatrix methods, and, if repeats are found, split the sequence into pieces accordingly. 2. Use FASTA to find similar sequences or use PROFILESEARCH for a more sensitive search. 3. Compare the sequence to known motifs, for example, in the PROSITE database. 4. Search for hydrophobic segments using the Kyte-Doolittle (1982) method and for α helices and β strands using the hydrophobic moment (see UNITS 2.2 & 2.3).
Computational Analysis
2.1.13 Current Protocols in Protein Science
5. If step 1 or 2 was successful, then search for helices using the variation moment. 6. If step 1 or 2 was successful, calculate the hydrophobic correlation to estimate similarity. 7. Keeping in mind that the results will be only 55% to 65% accurate, compare the outputs of several secondary structure prediction methods with information from all available homologous sequences. 8. For antibodies, identify possible antigenic regions using hydrophilicity plots and antigenicity index plots. 9. Look for unusual features such as leucine zippers; this can be done with any program, such as PROSITE, that takes advantage of a dictionary of motifs. 10. Determine potential sites of cleavage by various chemicals; cleavage sites can be predicted by computer methods and verified experimentally.
entific and medical researchers. A few are presented here. For a listing of molecular sequence and related software, get the Catalogue of Molecular Biology from Genethon by connecting via ftp ftp.genethon.fr and entering cd pub/resig/catalogu then mget bio.catal*. Amos Bairoch’s bibliographic database on sequence analysis (seqanalref), the Listing of Molecular Biology Databases (LiMB), a directory of researchers who apply artificial intelligence (AI) techniques to problems in molecular biology (aimbdb), and many other valuable files can be found using anonymous FTP to ncbi.nlm.nih.gov. Bairoch’s lists of molecular biology e-mail, FTP, and BBO servers are also available using anonymous FTP: type ftp expasy.hcuge.ch, then cd databases/info, then mget serv_ema.txt serv_ftp.txt serv_bbo.txt. For a copy of the BIOSCI FAQ, a collection of Frequently Asked Questions, use ftp net.bio.net, then cd pub/BIOSCI/doc and get biosci.FAQ.
INTERNET RESOURCES
Computational Methods for Protein Sequence Analysis
The Internet has become an important resource in bioinformatics. There are new tools that make it easy to locate the latest software and databases. Since 1994 a new Internet-based network of hypertext tools that takes advantage of the World Wide Web (WWW) has emerged. To enter the WWW, you will need a network hypertext browser such as MOSAIC. Versions of MOSAIC, which was originally developed by the National Center for Supercomputer Applications (NCSA), are now freely available for most commonly used laboratory computers. If your local computer is connected to the Internet and your system has file transfer protocol (FTP) capability, connect to ftp.ncsa.uiuc.edu and log in under the username anonymous to retrieve a copy of MOSAIC for the computer you use most often. Once you have MOSAIC or another WWW browser running, we recommend connecting to the Australian National University’s Bioinformatics server (http://life.anu.edu.au) and selecting “Molecular biology” from the menu. Table 2.1.1 reproduces the list of resources available. Another current resource of hyperlinks is available through the author using MOSAIC or Netscape at the George Mason Comp uta tion al B iolo gy Resource (http://www.science.gmu.edu/~michaels/ comp_bio). As indicated by that list, there is a tremendous number of resources available on the Internet. In addition, there are many useful text and program files available by FTP that are of interest to molecular biologists and other sci-
LITERATURE CITED Allison, L., Wallace, C.S., and Yee, C.N. 1992. Finite-state models in the alignment of macromolecules. J. Mol. Evol. 35:77-89. Altschul, S.F., Gish, W., Miller, W., Myers, E.W., and Lipman, D.J. 1990. Basic local alignment search tool. J. Mol. Biol. 215:403-410. Argos, P. 1987. A sensitive procedure to compare amino acid sequences. J. Mol. Biol. 193:385396. Argos, P. and Vingron, M. 1990. Sensitivity comparison of protein amino acid sequences. Methods Enzymol. 183:352-365. Bairoch, A. 1992. PROSITE: A dictionary of protein sites and patterns. Nucl. Acids Res. 19:22412245. Barton, G.J. 1990. Protein multiple sequence alignment and flexible pattern matching. Methods Enzymol. 183:403-428. Barton, G.J. 1993. An efficient algorithm to locate all locally optimal alignments between two sequences allowing for gaps. CABIOS 9:729-734. Brendel, V. 1992. PROSET—A fast procedure to create nonredundant sets of protein sequences. Math. Comput. Modeling 16(6/7):37-43. Brendel, V., Bucher, P., Nourbaksh, I.R., Blaisdell, B.E., and Karlin, S. 1992. Methods and algorithms for statistical analysis of protein sequences. Proc. Natl. Acad. Sci. U.S.A. 89:20022006. Burks, C. 1990. The flow of nucleotide sequence data into data banks: Role and impact of largescale sequencing projects. In Computers and DNA, Santa Fe Institute (G. Bell and T. Marr, eds.) pp. 35-45. Addison-Wesley, Reading, Mass.
2.1.14 Current Protocols in Protein Science
Table 2.1.1
Australian National University (ANU) Molecular Biology Resources
Databases French WWW genomic service, including Caenorhabditis elegans Database BLAST Database Searches at NCBI CompoundKB database—981 metabolic intermediate compounds Codon usage tables (major species) & EMBL mirror DNA Data Bank of Japan EC enzyme database EMBL data (Heidelberg) ESTDB—Expressed Sequence Tag Database (TIGR) FASTA Database search program (Virginia) Genbank searches (Indiana) Genbank/Swiss-Prot/Protein/PIR (via NIH) GENETHON Human Genome Centre Mendelian Inheritance in Man (index) Metabolic intermediate compounds Microbial germplasm Miscellaneous, e.g. codon usage, profiles (Weizmann) PIR (Houston) Prosite (via NIH) Protein databank (Brookhaven) REF52 2D Gel Protein Database REBASE—Restrict. Enzyme Data Base (NEB) REBASE restriction enzymes Swiss-Prot: EMBNet (Heidelberg) and ExPASy (Geneva) SWISS-2DPAGE—Two-dimensional Polyacrylamide Gel Electrophoresis Database (Geneva) Various (ICGEB, Italy) Biologists’s Control Panel Bibliographies/Tutorials Biosequence Comparison Sequence analysis (search index) Biocomputing bibliography Biological Journals—current titles Molecular biology journals References to molecular biology algorithms Periodical references to journals in molecular biology Software ANU software (gopher link) SIMPLE34—Detection of sequence repetition in nucleic acid MELANIE software packages for 2D PAGE computer analysis (Geneva) NCBI repository
Sequence Analysis Tools (Trieste) IUBio Biology Software and Database Archive (Indiana) Quest II software for 2D gel proteins CODA—Conservation Options and Decision Analysis RAPDistance (gopher link) RAPDistance (WWW link) News groups BioNet—various biological topics The Scientist (biweekly news with a biotech slant) BIOSCI mailing lists and newsgroup archives Special topics The Australian National Genomic Information Service (ANGIS) Human Genome Mapping Project (UK) Artificial Intelligence in molecular biology Database of molecular biologists working in AI Agricultural Genome World Wide Web server A Caenorhabditis elegans DataBase (ACeDB) Other sites Australian National Genomic Information Service (ANGIS) CAMIS Centre for Advanced Medical Informatics (Stanford) Caenorhabditis Genetics Centre (CGC) Chlamydomonas Genetics (Duke) EMBNet ExPASy (Geneva) Harvard Biological Laboratories Human Genome Center at Lawrence Berkeley Laboratory Johns Hopkins University, BioInformatics National Center for Biotechnology Information (NCBI) QUEST Protein Database Center Ribosomal Database Project (Argon Natl. Laboratory) Molecular biology archives (list) NIH GenoBase server Othersites (gopher links) Yeast Genome Information Server (Stanford) WAIS—EC enzymes WAIS—Molecular Biology WAIS—indexes sorted alphabetically (ADFA)
Computational Analysis
2.1.15 Current Protocols in Protein Science
Chou, P.Y. and Fasman, G.D. 1974. Prediction of protein conformation. Biochemistry 13:222-244. Collado-Vides, J. 1991. The search for a grammatical theory of gene regulation is formally justified by showing the inadequacy of context-free grammars. CABIOS 7:321-326. Collins, J.F. and Coulson, A.F.W. 1990. Significance of protein sequence similarities. Methods Enzymol. 183:474-487. Day, W.H.E. and McMorris, F.R. 1993. A consensus program for molecular sequences. CABIOS 9:653-656. Dayhoff, M.O. 1978. Atlas of Protein Sequence and Structure. National Biomedical Research Foundation, Washington, D.C. Depiereux, E. and Feytmans, E. 1991. Simultaneous and multivariate alignment of protein sequences: Correspondence between physicochemical profiles and structurally conserved regions (SCR). Protein Eng. 4:603-613. De Rijk, P. and De Wachter, R. 1993. DCSE, an interactive tool for sequence alignment and secondary structure search. CABIOS 9:735-740. Doolittle, R.F. 1981. Similar amino acid sequences: Chance or common ancestry? Science 214:167339. Doolittle, R.F. 1986. Of URFs and ORFs: A Primer on How to Analyze Derived Amino Acid Sequences. University Science Books, Ann Arbor, Mich. Doolittle, R.F. 1989. Redundancies in protein sequences. In Prediction of Protein Structure and the Principles of Protein Conformation (G.D. Fasman, ed.) pp. 599-623. Plenum, New York.
Computational Methods for Protein Sequence Analysis
Doolittle, R.F. 1990. What we have learned and will learn from sequence databases. In Computers and DNA, Santa Fe Institute (G. Bell and T. Marr, eds.) pp. 21-31. Addison-Wesley, Reading, Mass. Dumas, J.P. and Nunio, J. 1982. Efficient algorithm for folding and comparing nucleic acid sequences. Nucl. Acids Res. 10:197-206. Eroshkin, A.M., Zhilkin, P.A., and Fomin, V.I. 1993. Algorithm and computer program: Pro_Anal for analysis of relationship between structure and activity in a family of proteins or peptides. CABIOS 9:491-497. Fitch, W.M. 1966. An improved method of testing for evolutionary homology. J. Mol. Biol. 16:916. Fitch, W.M. 1969. Locating gaps in amino acid sequences to optimize the homology between two proteins. Biochem. Genet. 3:99-108. Fuchs, R. 1994. Fast protein block searches. CABIOS 10:79-80. Garnier, J., Osguthorpe, D.J., and Robson, B. 1978. Analysis of the accuracy and implications of simple methods for predicting the secondary structure of globular proteins. J. Mol. Biol. 120:97-120.
Genetics Computer Group. 1994. GCG Program Manual for the Wisconsin Package, Version 8, September 1994. Genetics Computer Group Inc., Madison, Wis. George, D., Hunt, L.T., and Barker, W.C. 1990. Mutation data matrix and its uses. Methods Enzymol. 183:333-351. Gibbs, A.J. and McIntyre, G.A. 1970. The diagram, a method for comparing sequences. J. Biochem. 16:1-11. Gotoh, O. 1986. Alignment of three biological sequences with an efficient traceback procedure. J. Theor. Biol. 121:327-337. Gribskov, M., Homyak, M., Edenfield, J., and Eisenberg, D. 1988. Profile scanning for three-dimensional structural patterns in protein sequences. CABIOS 4:61-66. Gribskov, M., Luethy, R., and Eisenberg, D. 1990. Profile analysis. Methods Enzymol. 183:146159. Gribskov, M. 1994. Profile analysis. Computer analysis of sequence data. Methods Mol. Biol. 24:247-266. Henikoff, S. and Henikoff, J.G. 1993. Proteins Struct. Funct. Genet. 17:49-61. Heringa, J., Sommerfeldt, H., Higgins, D.G., and Argos, P. 1992. OBSTRUCT: A program to obtain the largest cliques from a protein sequence set according to structural resolution and sequence similarity. CABIOS 8:599-600. Hodgman, T.C. 1992. Nucleic acid and protein sequence management. In Microcomputers in Biochemistry: A Practical Approach (C.F.A. Bryce, ed.) pp. 131-158. IRL Press, Oxford. Kanaoka, M., Kishimoto, F., Ueki, Y., and Umeyama, H. 1989. Alignment of protein sequences using the hydrophobic core scores. Protein Eng. 2:347-351. Karlin, S.P., Morris, M., Ghandour, G., and Leung M.-Y. 1988. Algorithms for identifying local molecular sequence features. CABIOS 4:41-51. Karlin, S.P., Ost, F., and Blaisdell, B.E. 1989. Patterns in DNA and amino acid sequences and their statistical significance. In Mathematical Methods for DNA Sequences (M.S. Waterman, ed.) pp. 133-157. CRC Press, Boca Raton, Fla. Karlin, S., Bucher, P., and Brendel, V. 1991. Statistical methods and insights for protein and DNA sequences. Annu. Rev. Biophys. Chem. 20:175203. Kruskal, J.B. 1983. An overview of sequence comparison. In Time Warps, String Edits, and Macromolecules: The Theory and Practice of Sequence Comparison (D. Sankoff and J.B. Kruskal, eds.) pp. 1-44. Addison-Wesley, Reading, Mass. Kruskal, J.B. and Sankoff, D. 1983. An anthology of algorithms and concepts for sequence comparison. In Time Warps, String Edits, and Macromolecules: The Theory and Practice of Sequence Comparison (D. Sankoff and J.B. Kruskal, eds.) pp. 265-310. Addison-Wesley, Reading, Mass.
2.1.16 Current Protocols in Protein Science
Kyte, J. and Doolittle, R.F. 1982. A simple method for displaying the hydrophobic character of a protein. J. Mol. Biol. 157:105-132. Landau, G.M., Vishkin, U., and Nussinov, R. 1988. Locating alignments with k differences for nucleotide and amino acid sequences. CABIOS 4:19-24. Landau, G.M., Vishkin, U., and Nussinov, R. 1990. Fast alignment of DNA and protein sequences. Methods Enzymol. 183:487-502.
Pearson, W.R. and Miller, W. 1992. Dynamic programming algorithms for biological sequence comparison. Methods Enzymol. 210:576-610. Pevzner, P.A. 1992. Statistical distance between texts and filtration methods in sequence comparison. CABIOS 8:121-127. Pizzi, E.M., Attimonelli, M., Liuni, S., Frontali, C., and Saccone, C. 1991. A simple method for global sequence comparison. Nucl. Acids Res. 20:131-136.
Landes, C., Henaut, A., and Risler, J.-L. 1993. Dotplot comparisons by multivariate analysis (DOCMA): A tool for classifying protein sequences. CABIOS 9:91-196. Lipman, D.J. and Pearson, W.R. 1985. Rapid and sensitive protein similarity searches. Science 227:1435-1441. Livingstone, C.D. and Barton, G.F. 1993. Protein sequence alignments: A strategy for the hierarchical analysis of residue conservation. CABIOS 9:745-756.
Reich, J.G. and Meiske, W. 1987. A simple statistical significance test of window scores in large dot matrices obtained from protein or nucleic acid sequences. Comput. Appl. Biosci. 3:25-30. Rigoutsos, I. and Califano, A. 1994. Searching in parallel for similar strings. IEEE Computatl. Sci. Eng. 60-75. Robson, B. and Greaney, P.J. 1992. Natural sequence code representations for compression and rapid searching of human-genome-style databases. CABIOS 8:283-289.
Maizel, J.V. and Lenk, R.P. 1981. Enhanced graphic matrix analysis of nucleic acids and protein sequences. Proc. Natl. Acad. Sci. U.S.A. 78:76657669. McLachlan, A.D. 1971. Test for comparing related amino acid sequences: Cytochrome c and cytochrome c-551. J. Mol. Biol. 61:409-424. McLachlan, A.D. 1972. Repeating sequences and gene duplication in proteins. J. Mol. Biol. 72:417-437. Michaels, G.S., Taylor, R., Hagstrom, R., Price, M., and Overbeek, R. 1993. Searching for genomic organizational motifs: Explorations of the E. coli chromosome. Comp. Chem. 17:209-217.
Rohde, K. and Bork, P. 1993. A fast, sensitive pattern-matching approach for protein sequences. CABIOS 9:183-189.
Mrazek, J. and Kypr, J. 1993. UNIREP: A microcomputer program to find unique and repetitive nucleotide sequences in genomes. CABIOS 9:355-360. Murata, M., Richardson, J.S., and Sussman, J.L. 1985. Simultaneous comparison of three protein sequences. Proc. Natl. Acad. Sci. U.S.A. 82:3073-3077. Nedde, D.N. and Ward, M.O. 1993. Visualizing relationships between nucleic acid sequences using correlation images. CABIOS 9:331-335. Needleman, S.B. and Wünsch, C.D. 1970. A general method applicable to the search for similarities in the amino acid sequence of two proteins. J. Mol. Biol. 48:443-453. Panjukov, V.V. 1993. Finding steady alignments: Similarity and distance. CABIOS 9:285-290. Parry-Smith, D.J. and Atwood, T.K. 1992. ADSP— A new package for computational sequence analysis. CABIOS 8:451-459. Pearson, W.R. 1990. Rapid and sensitive comparison with FASTP and FASTA. Methods Enzymol. 183:63-98. Pearson, W.R. 1994. Using the FASTA program to search protein and DNA sequence databases. Methods Mol. Biol. 24:365-389.
Searls, D. 1993. The computational linguistics of biological sequences. In Artificial Intelligence and Molecular Biology (L. Hunter, ed.) pp. 47120. MIT Press, Cambridge, Mass. Sellers, P.H. 1974. On the theory and computation of evolutionary distances. SIAM J. Appl. Math. 26:787-793. Smith, R.F. and Smith, T.F. 1992. Pattern-induced multisequence alignment (PIMA) algorithm employing secondary structure–dependent gap penalties for use in comparative protein modeling. Protein Eng. 5:35-41. Smith, T.F. and Waterman, M.S. 1981. Comparative biosequence metrics. J. Mol. Evol. 18:38-46. Staden, R. 1994a. Statistical and structural analysis of protein sequences. Methods Mol. Biol. 24:125-130. Staden, R. 1994b. Searching for motifs in protein sequences. Methods Mol. Biol. 24:131-139. Staden, R. 1994c. Using patterns to analyze protein sequences. Methods Mol. Biol. 24:141-154. Staden, R. 1994d. Comparing sequences. Methods Mol. Biol. 24:155-170. States, D.J. 1992. Molecular sequence accuracy: Analyzing imperfect data. Trends Genet. 8:5255. States, D.J. and Boguski, M.S. 1990. Sequence Analysis Primer. Stockton Press, New York. Streletc, V.B., Shindyalov, I.N., Kolchanov, N.A., and Lim, H.A. 1991. Fast, statistically based alignment of amino acid sequences on the base of diagonal fragments of dot matrices. CABIOS 8:529-534. Taylor, W.R. 1988. Pattern matching methods in protein sequence comparison and structure prediction. Protein Eng. 2:77-86. Computational Analysis
2.1.17 Current Protocols in Protein Science
Tyler, E., Horton, M.R., and Krause, P.R. 1991. A review of algorithms for molecular sequence comparison. Computers Biomed. Res. 24:72-96. Waterman, M.S. 1989. Sequence alignments. In Mathematical Methods for DNA Sequences (M.S. Waterman, ed.) pp. 53-90. CRC Press, Boca Raton, Fla. Waterman, M.S. 1990. Consensus patterns in sequences. In Mathematical Methods for DNA Sequences (M.S. Waterman, ed.) pp. 93-115. CRC Press, Boca Raton, Fla. Waterman, M.S. and Eggert, M. 1991. A new algorithm for best subsequence alignments with application to tRNA-rRNA comparisons. J. Mol. Biol. 197:723-728.
Waterman, M.S. and Jones, R. 1990. Consensus methods for DNA and protein sequence alignment. Methods Enzymol. 183:221-237. Wilbur, W.J. and Lipman, D.J. 1983. Rapid similarity searches of nucleic acid and protein data banks. Proc. Natl. Acad. Sci. U.S.A. 80:726-730.
Contributed by George Michaels and Robert Garian George Mason University Fairfax, Virginia
Computational Methods for Protein Sequence Analysis
2.1.18 Current Protocols in Protein Science
Hydrophobicity Profiles for Protein Sequence Analysis INTRODUCTION Seminal work by Anfinsen et al. (1961) showed that the three-dimensional structure of a protein is determined by its amino acid sequence. Thus, significant effort has been aimed at unraveling the complex relationship between sequence and structure. Although there is no immediate prospect of a general solution to the protein folding problem, stepwise empirical methods have been developed that analyze protein sequences and successfully provide predictive results concerning protein structure. The work of Kauzmann (1959) determined that hydrophobic interactions are a major force in protein folding. Accounting for the hydrophobicity of amino acids within a protein is thus the first step toward understanding protein folding (Tanford, 1980). Hydrophobicity studies have resulted in a wide range of hydropathy scales, indices that attempt to quantify the relative hydrophobicity of the amino acids. The actual hydrophobic content attributed to each amino acid is dependent on the methodology used to estimate it and may vary from scale to scale. Some scales are based solely on physicochemical measurements such as the free energy of side-chain transfer (Nozaki and Tanford, 1971; Wolfenden et al., 1981). Other scales are based on statistical analysis of threedimensional structures (Chothia, 1976; Rose et al., 1985). In addition, scales can be a combination of both procedures (Kyte and Doolittle, 1982). The general observation that hydrophobic residues tend to be buried in the interior of the protein whereas hydrophilic residues prefer to be exposed to solvent is predominant in all scales. Generating a hydropathy profile requires two things: (1) the primary structure and (2) the hydropathy scale. The primary structure can be obtained by translating the cDNA or by sequencing the protein. Hydropathy profiles for a protein are obtained by averaging or combining the hydrophobicity or hydrophilicity of an individual amino acid (taken from a particular hydropathy scale) with the values of several neighboring residues along the protein sequence, then plotting the local averages or sums against the amino acid sequence. In such hydrophobicity plots, regions of hydrophobicity and hydrophilicity appear as maxima and minima, respectively. A fundamental use of hy-
dropathy profiles, therefore, is predicting the overall hydropathy (hydrophobicity or hydrophilicity) of a protein from its amino acid sequence. The distribution of local hydropathy values can provide insight into the overall folding pattern of the protein (see Applications). Numerous procedures have been developed for calculating the hydropathy profile of a protein based on the free energy of side-chain transfer from water or vacuum to an organic solvent (Nozaki and Tanford, 1971; Tanford, 1980; Wolfenden et al., 1981; Fauchere and Pliska, 1983). The procedures differ primarily in the hydropathy scale used, and the different hydropathy scales have arisen, in part, to address specific applications. For example, the hydrophilicity scale of Hopp and Woods (1981) was developed to locate antigenic segments; the Goldman, Engleman, and Steitz (GES) scale (Engleman et al., 1986) was developed to identify transmembrane regions in a protein. Although individual scales were developed for different purposes or by alternative procedures, significant similarity exists in the resultant profiles. Most hydropathy plots show similar distributions of minima and maxima in the sequence profiles. In Figure 2.2.1, the result of applying the most widely used hydrophobicity scale, that of Kyte and Doolittle (1982), is compared to results of applying the scales of Hopp and Woods (1981) and Eisenberg (1984) to hen egg white lysozyme. The similarity in profiles is partially due to the fact that maxima and minima often reflect secondary structure elements; regions of hydrophobicity and hydrophilicity can be correlated with β sheets and β turns, respectively (Kuntz, 1972; Roy and Rose, 1980). Hydropathy profiles can be used to examine the surface features of proteins in order to generate hypotheses that can be confirmed experimentally. For example, such analyses received widespread use in the prediction of antigenic and other interaction sites on proteins. Furthermore, when used in conjunction with other secondary structure prediction algorithms (see UNIT 2.3), hydropathy profiles have been shown to correlate well with α-helical, turn, and β-sheet propensities. As such, they have proved to be invaluable for predicting protein secondary structure. The most extensive use of hydropathy profiles has been for
Contributed by Stanley R. Krystek, Jr., William J. Metzler, and Jiri Novotny Current Protocols in Protein Science (1995) 2.2.1-2.2.13 Copyright © 2000 by John Wiley & Sons, Inc.
UNIT 2.2
Computational Analysis
2.2.1 CPPS
20
Hydropathy value
10
0
–10
– 20
–30 0
50
100
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Sequence position
Figure 2.2.1 Hydropathy profiles of hen egg white lysozyme. The three profiles were created with the Kyte and Doolittle, the Hopp and Woods, and the Eisenberg scales using a scanning window of 7 amino acids. The Hopp and Woods scale was modified as described in the text to match other hydropathy scales.
deducing transmembrane regions in membrane proteins. Previous reviews describe systematic evaluations of hydropathy procedures and provide criteria for selecting the most appropriate amino acid hydropathy scale (Engleman et al., 1986; Cornette et al., 1987; Banghan, 1988; Hopp, 1989; Rose and Dworkin, 1989). This unit describes the application of hydrophobicity plots to typical problems and provides suggested uses for a few selected scales.
METHODOLOGY Selecting Hydropathy Scales
Hydrophobicity Profiles for Protein Sequence Analysis
The first step in generating a hydropathy profile (or profiles) for an individual amino acid sequence is selecting the appropriate scale(s). Table 2.2.1 lists several widely used scales. This unit focuses primarily on four hydropathy scales: the Hopp and Woods, the Kyte and Doolittle, the Eisenberg, and the GES scales. Differences among the hydropathy indices for each amino acid in the four scales (Table 2.2.2) arise primarily from differences in the experimental data from which the scales were derived and from the methodologies by
which each scale was optimized. The hydrophilicity scale derived by Hopp and Woods is based on solubility values of hydrophobic and hydrophilic amino acids (Nozaki and Tanford, 1971; Levitt, 1976). In the Kyte and Doolittle and the Eisenberg scales, the hydropathy index is a measure of the relative affinity of an amino acid for hydrophobic phases. The GES scale is based on physicochemical considerations of helical structure and several energetic factors that reflect partitioning of an amino acid side chain between aqueous solution and a membrane bilayer. Both the Kyte and Doolittle and the Eisenberg scales are combinations of earlier scales. The Kyte and Doolittle scale combines values from the water-to-vapor transfers and the internal/external distributions of amino acid residues from known structures as determined by Chothia (1976). The Eisenberg scale is a consensus of the Wolfenden (Wolfenden et al., 1981), Chothia (Chothia, 1976), Janin (Janin, 1979), and von Heijne (von Heijne and Blomberg, 1979) scales and is designed to minimize effects of the outlying values observed for some amino acids in previous scales.
2.2.2 Current Protocols in Protein Science
Table 2.2.1
Hydropathy Scales
Reference scale
Literature
Derivationa
Bigelow Tanford Chothia Levitt Janin von Heijne and Blomberg Roy and Rose Hopp and Woods Wolfenden Argos Fraga Kyte and Doolittle Fauchere and Pliska Eisenberg Hopp Rose GES Parker, Guo, and Hodges Esposti et al.
Bigelow, 1967 Nozaki and Tanford, 1971 Chothia, 1976 Levitt, 1976 Janin, 1979 von Heijne and Blomberg, 1979 Roy and Rose, 1980 Hopp and Woods, 1981 Wolfenden et al., 1981 Argos et al., 1982 Fraga, 1982 Kyte and Doolittle, 1982 Fauchere and Pliska, 1983 Eisenberg, 1984 Hopp, 1984 Rose et al., 1985 Engleman et al., 1986 Parker et al., 1986 Esposti et al., 1990
Experimental Experimental Theoretical Experimental Theoretical Theoretical Experimental Experimental Experimental Theoretical Theoretical Combination Experimental Combination Theoretical Theoretical Theoretical Experimental Theoretical
aExperimental, largely original data from physicochemical experiments; theoretical, derivation based on
analysis of structural data; combination, consensus of multiple scales.
There are several other interesting scales that deserve mention. The first hydropathy scale for membrane proteins was developed and revised by von Heijne (von Heijne and Blomberg, 1979) and corresponds rather closely to the GES scale described above (Engleman et al., 1986). A hydrophilicity scale by Parker, Guo, and Hodges (1986) provides new hydrophilicity values that are determined from HPLC retention times of model synthetic peptides. Another important set of scales includes the statistical scales developed by Argos and co-workers for prediction of membraneburied helices (Argos et al., 1982; Rao and Argos, 1986). These scales are derived from the statistical preference of specific amino acid residues to form the integral regions in transmembrane proteins, and they have been shown to correlate with highly resolved crystal structures much better than the physicochemical hydropathy scales (Esposti et al., 1990). Because all hydropathy scales are conceptually similar, the strategy of applying them to a given problem should be analogous regardless of the particular scale chosen. Moreover, although there may always be debates over which of the scales (methods) will prove to be best, experience has shown that application of
several different hydropathy schemes and alternative predictive rules will tend to build a consensus picture of protein hydrophobicity. For this unit the values of the Hopp and Woods and GES scales are multiplied by −1 so that these scales comply with others having peak maxima corresponding to hydrophobicity and peak minima corresponding to hydrophilicity (the scale values were not modified in Table 2.2.2). The Hopp and Woods and GES plots shown are therefore inverse plots.
Applying the Scanning Window Averaging Technique After the appropriate scale(s) has been selected, the next stage is generating the hydropathy profile for a protein. To create the profile, the numerical values derived from the hydropathy scales for each residue in the protein are repetitively averaged over the length of protein sequence. This procedure is commonly referred to as the scanning window averaging technique (Hopp and Woods, 1981; Kyte and Doolittle, 1982). In practice, the hydropathy is obtained by summing the individual hydropathy values (hydrophobicities or hydrophilicities) for a given number of contiguous amino acids along the protein chain. The sum of hy-
Computational Analysis
2.2.3 Current Protocols in Protein Science
Table 2.2.2
Hydrophobicity Indices for Selected Scales
Amino acid
H & Wa
K & Db
Eisenbergc GESd
Neutral, hydrophobic, aliphatic −0.4 Gly 0.0 −0.5 Ala 1.8 −1.5 Val 4.2 −1.8 Ile 4.5 −1.8 Leu 3.8 −1.3 Met 1.9
0.16 0.25 0.54 0.73 0.53 0.26
−1.0 −1.6 −2.6 −3.1 −2.8 −3.4
Neutral, hydrophobic, aromatic −2.5 Phe 2.8 −2.3 −1.3 Tyr −3.4 −0.9 Trp
0.61 0.02 0.37
−3.7 0.7 −1.9
Neutral, hydrophilic Ser 0.3 −0.4 Thr Asn 0.2 Gln 0.2
−0.8 −0.7 −3.5 −3.5
−0.26 −0.18 −0.64 −0.69
−0.6 −1.2 4.8 4.1
Acidic, hydrophilic Asp 3.0 Glu 3.0
−3.5 −3.5
−0.72 −0.62
9.2 8.2
Basic, hydrophilic −0.5 His Lys 3.0 Arg 3.0
−3.2 −3.9 −4.5
−0.40 −1.1 −1.8
3.0 8.8 12.3
Thiol-containing −1.0 Cys
2.5
0.04
−2.0
−1.6
−0.07
0.2
Imino acid Pro
0.0
aHopp and Woods, 1981. bKyte and Doolittle, 1982. cEisenberg, 1984. dEngleman et al., 1986.
Hydrophobicity Profiles for Protein Sequence Analysis
dropathy values is calculated progressively through the sequence, with each amino acid being the start of a new sum (Fig. 2.2.2). The resultant sums (or averages) are plotted versus the sequence position at the position of the central (or first) amino acid in the segment being summed. The choice of the center versus the first residue has no effect on the calculated profiles but shifts the sequence axis by half the window size. We prefer to plot versus the center position as it seems more intuitive. Similarly, the choice of summing or averaging has no effect on the profile but modifies the relative values plotted on the y-axis. We use sums for the profiles presented in this unit. This approach
displays the general hydropathic character of a protein. The ability to discern structurally interpretable features in the calculated hydropathy profile is critically dependent on the width of the scanning window, that is, the number of amino acids included in the summation. Figure 2.2.3 shows the effect of altering the size of the scanning window on hydropathy profiles generated for lysozyme. In Figure 2.2.3A, where the window size is set to 1 amino acid, the resultant hydropathy profile is featureless, providing essentially no structural information. On introducing a window size of 7 amino acids, well-resolved peaks appear in the hydropathy
2.2.4 Current Protocols in Protein Science
10 20 protein sequence . . . RTYFCDEQASQDWLVNAR . . . sliding window (size of 7) average hydropathy . . . 4 5 6 7 8 9 10 11 12 13 14 . . . value mapped to center residue of window
Figure 2.2.2 Schematic depicting the scanning window averaging technique.
profile (Fig. 2.2.3B). Increasing the window size to 11 amino acids (Fig. 2.2.3C) results in a smoothing or broadening of observed peaks in the profile, and with window sizes of 15 or 19 amino acids (Fig. 2.2.3D,E), the profiles again have few well-resolved features. From the family of profiles in Figure 2.2.3, it is readily apparent that window sizes exceeding 10 amino acids result in a loss of local information. This decrease in local information leads to increased errors in predicting hydrophobic regions in globular proteins (Kyte and Doolittle, 1982). In contrast, significant correlations do exist between hydropathy profiles generated with window sizes of 5 to 9 amino acids and known structural elements (Roy and Rose, 1980; Kyte and Doolittle, 1982; Rao and Argos, 1986; Cornette et al., 1987). In selecting the appropriate window size for a particular problem, the length of the structural property being investigated should be used as a guide. For example, transmembrane helices are roughly 20 residues in length, but the helices of globular proteins tend to be much shorter. One might therefore opt for a larger window size if searching for transmembrane helices, whereas a length of 7 or 9 amino acids is more appropriate for predicting surface sites or secondary structure. In cases of uncertainty, analysis of profiles generated with various window sizes will afford the most reliable results.
Threshold or Cutoff Values To facilitate the analysis of hydropathy profiles, it is convenient to define a threshold value, that is, the amplitude above which the hydropathy value must remain for a particular amino acid to be considered part of a contiguous region. Although somewhat arbitrary, the actual choice of threshold value for a particular profile is often derived empirically or from
experience. For example, if one were interested in analyzing the sequences of known membrane proteins for transmembrane helices, then an appropriate threshold value might be one which delineates peaks in the hydropathy profile to approximately 20 amino acids (see section on Predicting Transmembrane Regions in Proteins). As with selecting scanning window size, analyzing profiles with different threshold values should yield more reliable results until experience is gained.
APPLICATIONS Predicting Interaction Sites Hydropathy profiles have found widespread use in investigations aimed at elucidating the antigenic structure and surface features of proteins for which there are no three-dimensional structures (Hopp and Woods, 1981; Krystek et al., 1985a,b). Antigenic sites tend to occur on protein surfaces, where predominantly hydrophilic residues are both exposed to solvent and accessible for interaction with antibody. One would therefore anticipate good correlation between antigenic sites and regions of high hydrophilicity. A test of this hypothesis carried out by Tanaka et al. (1985) reported a success rate of 56%. Figure 2.2.4 depicts the Hopp and Woods profiles for myoglobin generated with a window size of 7 amino acids. Regions of hydrophilicity are detected as peak minima in the profile. The lines at the bottom of Figure 2.2.4 indicate the locations of five major antigenic sites determined experimentally (Atassi, 1984). All of the antigenic sites are associated with regions of high hydrophilicity. In addition, there are two regions of high hydrophilicity (peaks at positions 40 to 45 and 78 to 84) that are not associated with antigenicity. It is plau-
Computational Analysis
2.2.5 Current Protocols in Protein Science
A
10 5 0 –5 –10 20
B
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C Hydrophobicity value
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D
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Hydrophobicity Profiles for Protein Sequence Analysis
Figure 2.2.3 Effect of altering the scanning window size on the resolution of the hydropathy profile of lysozyme. All profiles were created with the Kyte and Doolittle scale. Window sizes were set to (A) 1, (B) 7, (C) 11, (D) 15, and (E) 19 amino acids. Note that the scale in A is reduced to show the profile more clearly.
2.2.6 Current Protocols in Protein Science
15
Hydrophilicity value
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Figure 2.2.4 Hopp and Woods profile for sperm whale myoglobin using a scanning window of 7 amino acids. Lines below the profile indicate the locations of five experimentally determined antigenic sites (Atassi, 1984).
sible that these correspond to antigenic sites that have yet to be identified experimentally, or they may point to an alternate, functionally significant property of myoglobin. Whatever the case, these results serve to emphasize the point that hydropathy profiles should be used only as a guide for predicting antigenicity. Additional features of protein surfaces can be highlighted by hydropathy profiles. As an example, consider human hemoglobin, whose three-dimensional structure is known (Fermi, 1975). Hemoglobin is found as a tetramer in its native state, being composed of two α- and two β-subunits. In the native protein the β- and α-subunits are folded against one another, forming contact regions that more closely resemble the interior of a globular protein. Because these contact regions are hydrophobic, one might examine hydropathy profiles for regions of hydrophobicity as a means of identifying potential subunit interacting surfaces. The Hopp and Woods profile for human hemoglobin is shown in Figure 2.2.5. The regions of contact in the β-subunit of human hemoglobin (Yoshioka and Atassi, 1986) are indicated as bars above the profile. The two largest hydrophobic peaks correspond to regions of the β-subunit in contact with each of
the α-subunits. Analogous analysis of hydropathy profiles has been successfully used to identify the subunit interacting surfaces for the α-subunits of glycoprotein hormones lutropin and follitropin (Krystek et al., 1991, 1992). Human hemoglobin also provides a demonstration of the dangers of using hydropathy profiles for analyzing antigenic sites of proteins that possess significant quaternary structure. In Figure 2.2.5 the lines below the profile indicate the regions of five major continuous antigenic determinants. Although two sites correlate with regions of hydrophilicity, the others do not. In such cases the amino acid sequence alone is not sufficient for predictability. This may be due in part to the unique nature of proteins composed of multiple subunits.
Correlating Profiles with Secondary Structure An increasingly important application of hydropathy profiles is their use in conjunction with secondary structure prediction tools (see UNIT 2.3). Figure 2.2.6 shows the hydropathy profiles for interleukin 1β (IL-1β) generated with the Eisenberg scale (Fig. 2.2.6A) and the Kyte and Doolittle scale (Fig. 2.2.6B), using a scan window of 7 amino acids. The solid bars
Computational Analysis
2.2.7 Current Protocols in Protein Science
15
Hydrophilicity value
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Figure 2.2.5 Hopp and Woods profile for human hemoglobin β-subunit using a scanning window size of 7 amino acids. Bars above the profile mark the locations of β-subunit surfaces in contact with α-subunits (Yoshioka and Atassi, 1986). Lines below the profile indicate the locations of antigenic sites.
Hydrophobicity Profiles for Protein Sequence Analysis
above the profiles indicate the locations of β strands found in the crystal structure of this all-β protein (Finzel et al., 1989). Although there is a difference in scale magnitude between the Kyte and Doolittle and Eisenberg methods, both profiles show exceptionally good correlation of hydrophobicity (peak maxima) with the location of β strands. The correlation of hydrophobicity with β-sheet structural elements is a direct reflection of the fact that sheets are most often located in the hydrophobic interior of a protein; thus, they comprise residues most compatible with the hydrophobic environment. There is also good correlation between hydrophilicity (peak minima) and the location of turns in proteins. For interleukin 1β essentially all of the turns identified in the structure correspond to minima. Figure 2.2.7 shows the hydropathy profiles for myoglobin, a protein composed solely of α helices. The larger α helices appear to correlate with peak maxima. Like β sheets, helices are frequently found in regions of hydrophobicity in the hydrophobic core of proteins. Because of the interspersed nature of hydrophilic and hydrophobic residues of amphipathic helices, however, hydropathy profiles are less useful in
predicting the location of these helices. Moreover, as both helices and sheets result in peak maxima, the hydropathy profile is not sufficient to discriminate between them. Therefore, correlations of secondary structure with protein hydropathic character are most successful when used in conjunction with protein secondary structure prediction algorithms (see UNIT 2.3).
Predicting Transmembrane Regions in Proteins Perhaps the most popular use of hydropathy profiles is predicting transmembrane regions in proteins. In addition to hydropathy scales, there are several membrane preference scales. Such scales have been developed independently from the hydropathy scales and are based on statistical distributions of each amino acid in membrane regions of proteins. The statistical scales have been previously compared to the hydropathy scales (Esposti et al., 1990); in general, they are equally efficient at predicting the location of the transmembrane and amphipathic helices. Examining the structure of known membrane proteins has enabled the predictive rules
2.2.8 Current Protocols in Protein Science
6
A
4 2 0 –2
B
Hydropathy value
–4 –6 –8 20 10 0 –10 –20 –30 0
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Figure 2.2.6 Hydropathy profiles for interleukin 1β using a scanning window size of 7 amino acids. Bars above the profile indicate locations of the β strands, and lines below the profile mark the locations of turns (Finzel et al., 1989). (A) Profile generated using the Eisenberg scale. (B) Profile generated using the Kyte and Doolittle scale.
for the various scales to be optimized. In addition, estimates of the hydrophobic thickness of the bilayer have provided guides to the minimum number of helical amino acids necessary to span the bilayer: the minimum number is estimated to be 20 (Engleman et al., 1986). This greatly facilitates the prediction of putative transmembrane helices by adding an additional criterion to the analysis of hydropathy profiles for transmembrane regions, namely, maxima in the profiles must span ∼20 residues. Figure 2.2.8A shows the Kyte and Doolittle profile for bacteriorhodopsin, a membrane protein whose structure is known (Henderson et al., 1990). For Kyte and Doolittle profiles, the best resolution is seen for window scans of 7 or 9 amino acids and threshold values of 0.7 × window size. For a window scan of 7, the
threshold value is 4.9. There are seven major peaks with threshold values greater than 4.9 that span at least 20 amino acids (indicated by brackets above the profile in Fig. 2.2.8A). Comparison of the predicted transmembrane regions with the location of the actual helices identified in the structure (bars above the profile) indicates that the prediction is quite good. Similar agreement is seen when the method of Engleman et al. (1986) is used to predict the transmembrane regions for bacteriorhodopsin (Fig. 2.2.8B). Although the location of the helices can be predicted reliably, neither the Kyte and Doolittle nor the GES method is able to predict accurately the boundaries of the transmembrane regions. Fortunately, visual inspection of the sequence, as well as general features of the
Computational Analysis
2.2.9 Current Protocols in Protein Science
30
Hydrophobicity value
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Figure 2.2.7 Kyte and Doolittle hydropathy profile for sperm whale myoglobin generated with a scanning window size of 7 amino acids. Bars above the profile mark the locations of the helices in myoglobin.
Hydrophobicity Profiles for Protein Sequence Analysis
transmembrane helices, can frequently be used to delineate transmembrane regions and the interfacial residues at each end of the helices. Some general features common to transmembrane regions to consider are as follows. (1) Approximately 75% of the amino acids in transmembrane regions are hydrophobic (Leu, Ile, Val, Met, Phe, Trp, Tyr, Cys, Ala, Pro, and Gly). (2) Proline residues are found in ∼20% of the transmembrane helices but in only 3% of globular proteins (Barlow and Thornton, 1988). (3) Aromatic residues are clustered near the interface of the transmembrane helix and bulk water. Positional preferences for amino acids in transmembrane regions have been identified for single-spanning membrane proteins (Landolt-Marticorena et al., 1993) and can serve as a further guide for delineating the ends of transmembrane regions. It can be expected that these preferences will be similar for polytopic membrane proteins. The positional preferences define a motif with the following characteristics: an extracellular terminal flanking region (Asp, Ser and Pro); an extracellular-interfacial region (Trp); a transmembrane domain composed primarily of hydrophobic residues (Leu, Ile, Val, Met, Phe, Trp, Cys, Ala, Pro, and Gly); an
intracellular-interfacial region (Tyr, Trp, and Phe); and an intracellular terminal flanking region (Lys and Arg). Application of these sequence analysis rules for transmembrane proteins greatly improves the results of using hydropathy profiles for predicting transmembrane regions.
CONCLUSIONS 1. Virtually all hydropathy scales are able to reveal structural information in an equivalent manner. Selection of the appropriate hydropathy scale is therefore a practical and/or personal decision. 2. Window scanning procedures should use an appropriate window size for the structural element being examined: 7 or 9 amino acids for predicting surface sites or secondary structure and 19 amino acids for predicting membranespanning regions. 3. Identification of interaction sites can be accomplished using peak minima (hydrophilicity). 4. Correlation of protein secondary structure with hydropathy profiles may be useful for protein structure prediction methodology (see UNIT 2.3).
2.2.10 Current Protocols in Protein Science
A
30
Hydrophobicity value
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Free energy of transfer to water
B
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300
Sequence position
Figure 2.2.8 Hydropathy profiles for bacteriorhodopsin. Bars above the profiles mark the locations of the helices. (A) Kyte and Doolittle profile generated using a scanning window size of 7 amino acids. Brackets indicate the helices predicted on the basis of the profile alone. (B) GES profile generated using a scanning window size of 19 amino acids.
5. Prediction of transmembrane regions of integral membrane proteins can be made using a variety of scales and protocols; analysis of the hydropathy profiles should be augmented with the applicable sequence analysis rules.
rithms can also easily be encoded into programming languages or spreadsheets and interfaced with numerous graphical software packages for plotting the hydropathy profiles (if graphics are not included in the particular prediction program). These are also discussed in UNIT 2.1.
ACCESSIBILITY OF SOFTWARE Most commercial personal computer or computational packages (Genetics Computer Group, 1994; Biosym Technologies, 1991; Tripos Associates, 1993) contain algorithms for hydropathy profile generation along with other secondary structure prediction tools (see Table 2.3.6). In addition, the authors of each scale or protocol have published analysis programs (Table 2.2.1), of which many are available from public domain sources on the Internet. Algo-
LITERATURE CITED Anfinsen, C.B., Haber, E., Sela, M., and White, F.H., Jr. 1961. The kinetics of formation of native ribonuclease during oxidation of the reduced polypeptide chain. Proc. Natl. Acad. Sci. U.S.A. 47:1309-1314. Argos, P., Rao, J.K.M., and Hargrave, P.A. 1982. Structural prediction of membrane-bound proteins. Eur. J. Biochem. 128:565-575. Atassi, M.Z. 1984. Antigenic structure of proteins. Eur. J. Biochem. 145:1-20.
Computational Analysis
2.2.11 Current Protocols in Protein Science
Banghan, J.A. 1988. Data-sieving hydrophobicity plots. Anal. Biochem. 174:142-145. Barlow, D.J. and Thornton, J. 1988. Alpha helices in proteins. J. Mol. Biol. 210:601-619. Bigelow, C.C. 1967. On the average hydrophobicity of proteins and the relation between it and protein structure. J. Theor. Biol. 16:187-211. Biosym Technologies. 1991. Insight II/Homology Modules. Biosym Technologies, San Diego. Chothia, C. 1976. The nature of the accessible and buried surfaces of proteins. J. Mol. Biol. 105:114. Cornette, J.L., Cease, K.B., Margalit, H., Spouge, J.L., Berzofsky, J.A., and DeLisa, C. 1987. Hydrophobicity scales and computational techniques for detecting amphipathic structures in proteins. J. Mol. Biol. 195:659-685. Eisenberg, D. 1984. Three-dimensional structure of membrane and surface proteins. Annu. Rev. Biochem. 53:595-623. Engleman, D.M., Steitz, T.A., and Goldman, A. 1986. Identifying nonpolar transbilayer helices in amino acid sequences of membrane proteins. Annu. Rev. Biophys. Chem. 15:321-353. Esposti, M.D., Crimi, M., and Venturoli, G. 1990. A critical evaluation of the hydropathy profile in membrane proteins. Eur. J. Biochem. 190:207219. Fauchere, J.L. and Pliska, V. 1983. Hydrophobic parameters P of amino acid side chains from the partitioning of N-acetyl-amino acid amides. Eur. J. Med. Chem. 18:369-375. Fermi, G. 1975. Three-dimensional Fourier synthesis of human deoxyhaemoglobin at 2.5 angstroms resolution, refinement of the atomic model. J. Mol. Biol. 97:237-256. Finzel, B.C., Clancy, L.L., Holland, D.R., Muchmore, S.W., Watenpaugh, K.D., and Einspahr, H.M. 1989. Crystal structure of recombinant human interleukin-1β at 2.0 angstroms resolution. J. Mol. Biol. 209:779-791. Fraga, S. 1982. Theoretical prediction of protein antigenic determinants from amino acid sequences. Can. J. Chem. 60:2606-2610. Genetics Computer Group. 1994. GCG Program Manual for the Wisconsin Package. Genetics Computer Group, Inc., Madison, Wis. Henderson, R., Baldwin, J.M., Ceska, T.A., Zemlin, F., Beckmann, E., and Downing, K.H. 1990. Model for the structure of bacteriorhodopsin based on high resolution electron cryomicroscopy. J. Mol. Biol. 213:899-929. Hopp, T.P. 1984. Protein antigen conformation: Folding patterns and predictive algorithms; selection of antigenic and immunogenic peptides. Ann. Sclavo 2:47-60.
Hydrophobicity Profiles for Protein Sequence Analysis
Hopp, T.P. 1989. Use of hydrophilicity plotting procedures to identify protein antigenic segments and other interaction sites. Methods Enzymol. 178:571-585.
Hopp, T.P. and Woods, K.R. 1981. Prediction of protein antigenic determinants from amino acid sequences. Proc. Natl. Acad. Sci. U.S.A. 78:3824-3828. Janin, J. 1979. Surface and inside volumes in globular proteins. Nature 277:491-492. Kauzmann, W. 1959. Some factors in the interpretation of protein denaturation. Adv. Prot. Chem. 14:1-63. Krystek, S.R., Jr., Dias, J.A., Reichert, L.E., and Andersen, T.T. 1985a. Prediction of antigenic sites in follicle stimulating hormones: Difference profiles enhance antigenicity prediction methods. Endocrinology 117:1125-1130. Krystek, S.R., Jr., Reichert, L.E., and Andersen, T.T. 1985b. Analysis of computer generated hydropathy profiles for human glycoprotein and lactogenic hormones. Endocrinology 117:11101117. Krystek, S.R., Jr., Dias, J.A., and Andersen, T.T. 1991. Identification of subunit contact sites on the α-subunit of lutropin. Biochemistry 30:18581864. Krystek, S.R., Jr., Dias, J.A., and Andersen, T.T. 1992. Identification of a subunit contact site on the α-subunit of follitropin. Pept. Res. 25:165168. Kuntz, I.D. 1972. Protein folding. J. Am. Chem. Soc. 94:4009-4012. Kyte, J. and Doolittle, R.F. 1982. A simple method for displaying the hydropathic character of a protein. J. Mol. Biol. 157:105-132. Landolt-Marticorena, C., Williams, K.A., Deber, C.M., and Reithmeier, A.F. 1993. Nonrandom distribution of amino acids in the transmembrane segments of human type I single span membrane proteins. J. Mol. Biol. 229:602-608. Levitt, M. 1976. A simplified representation of protein conformations for rapid simulation of protein folding. J. Mol. Biol. 104:59-107. Nozaki, Y. and Tanford, C. 1971. The solubility of amino acids and two glycine peptides in aqueous ethanol and dioxane solutions. J. Biol. Chem. 246:2211-2217. Parker, J.M.R., Guo, D., and Hodges, R.S. 1986. New hydrophobicity scale derived from highperformance liquid chromatography peptide retention data: Correlation of predicted surface residues with antigenicity and X-ray-derived accessible sites. Biochemistry 25:5425-5432. Rao, J.K.M. and Argos, P. 1986. A conformational preference parameter to predict helices in integral membrane proteins. Biochim. Biophys. Acta 869:197-214. Rose, G.D. and Dworkin, J.E. (1989). The hydrophobicity profile. In Prediction of Protein Structure and the Principles of Protein Conformation (G.D. Fasman, ed.) pp. 625-633. Plenum, New York. Rose, G.D., Geselowitz, A.R., Lesser, G.J., Lee, R.H., and Zehfus, M.H. 1985. Hydrophobicity of amino acid residues in globular proteins. Science 229:834-838.
2.2.12 Current Protocols in Protein Science
Roy, S. and Rose, G.D. 1980. Hydrophobic basis of packing in globular proteins. Proc. Natl. Acad. Sci. U.S.A. 8:4643-4647. Tanaka, T., Slamon, D.J., and Cline, M.J. 1985. Efficient generation of antibodies to oncoproteins by using synthetic peptide antigens. Proc. Natl. Acad. Sci. U.S.A. 82:3400-3404. Tanford, C. 1980. The Hydrophobic Effect. John Wiley & Sons, New York. von Heijne, G. and Blomberg, C. 1979. Transmembrane translocation of proteins. The direct transfer model. Eur. J. Biochem. 97:175-181. Tripos Associates. 1993. Biopolymer/Composer Module. Tripos Associates, Inc., St. Louis, Mo. Wolfenden, R., Andersson, L., Cullis, P.M., and Southgate, C.C. 1981. Affinities of amino acid side chains for solvent water. Biochemistry 20:849-855.
Yoshioka, N. and Atassi, M.Z. 1986. Subunit interacting surfaces of human haemoglobin. Biochem. J. 234:457-461.
KEY REFERENCES Engleman et al., 1986. See above. Provides applications of hydropathy profiles to transmembrane region prediction. Kyte and Doolittle, 1982. See above. Describes the fundamental development and application of hydropathy profiles.
Contributed by Stanley R. Krystek, Jr., William J. Metzler, and Jiri Novotny Bristol-Myers Squibb Pharmaceutical Research Institute Princeton, New Jersey
Computational Analysis
2.2.13 Current Protocols in Protein Science
Protein Secondary Structure Prediction It is widely accepted that knowledge of the structure of a protein can provide important information regarding its function and mechanism of action. However, determining the threedimensional structure of a protein, whether by X-ray crystallography (Wyckoff et al., 1985; Blundell and Johnson, 1976) or nuclear magnetic resonance (NMR) spectroscopy (Oppenheimer and James, 1989; Wuthrich, 1986), remains a formidable task, as it requires relatively large amounts of pure protein (generally greater than milligram quantities). Thus, the structures of many proteins will remain out of reach. At the same time, continual advances in molecular biology provide protein sequence information (primary structures) at a pace that far exceeds the speed with which higher-order protein structures can be determined, thus making the need for methods to derive structural information even more pressing. This unit describes procedures developed for predicting protein structure from the amino acid sequence. On the basis of function and location in the cell, proteins can be loosely divided into three groups: globular, structural, and membrane. To date, the majority of protein structural data has been derived from studies of globular (soluble) proteins because of the ease of chemical handling of globular proteins. Thus, a wide disparity has developed in the number of three-dimensional structures that have been experimentally determined in the three protein groups. As a result, current secondary structure prediction schemes apply almost solely to globular proteins. In the native structures of globular proteins, the hydrophobic residues are generally buried in the interior and the hydrophilic residues are exposed to the surrounding solvent (Kauzmann, 1959; Fisher, 1964; Dill, 1990). Disulfide bonds are often present and stabilize the structure. Sections of amino acid sequence rich in hydrophobic residues can be correlated with secondary structural elements such as α helices and β strands (Roy and Rose, 1980). Those rich in polar residues correspond to surface loops and β turns (Kuntz, 1972; UNIT 2.2). These secondary structural elements (Pauling and Corey, 1951; Pauling et al., 1951) are characterized by hydrogen bonding between the amino and carboxyl groups of the polypeptide backbone (Schulz and Schirmer, 1979). Several reviews of protein structure exist, and readers are referred to books by Schulz and
Schirmer (1979) and Branden and Tooze (1991) for thorough discussions of protein structure. Protein secondary structure elements associate to form compact domains. Levitt and Chothia (1976) and Richardson (1981) have classified protein domains according to the types of arrangements of secondary structures that form them. Illustrations for the four classes, referred to as all-α, all-β, α/β, and α + β, are available (Richardson, 1981; Lesk, 1991). This unit focuses on examining secondary structure predictions using specific examples from different protein classes; methods of predicting folding class are also discussed. This unit is divided into four sections. The first section is an overview and brief history of structure prediction schemes. The second section describes four distinct prediction schemes, with emphasis on their differences. In the third part each prediction scheme is used to evaluate three proteins that have different folding patterns. The final section is a comparison of the prediction results and suggestions for secondary structure prediction.
OVERVIEW OF PREDICTION SCHEMES Early Predictive Schemes Two of the earliest secondary structure prediction methods were developed by Guzzo (1965) and Prothero (1966). By analyzing a database of protein structures and corresponding sequences, they generated a set of rules for predicting structure. Because of the limited number of structures available at the time, their prediction methods were restricted to helices, the most abundant conformation in the structures available. Soon after, Schiffer and Edmunson (1967) developed the helical wheel method, a pictorial tool that is still widely used. The helical wheel is a circular plot that represents the view one obtains when looking down the helix axis (Fig. 2.3.1). The “spokes” of the wheel are precisely positioned on the basis of the Corey-Pauling α-helical geometry, having 3.6 residues per turn, which results in an angular shift of two neighboring residues of 100° (Pauling et al., 1951). A helical wheel is constructed by placing successive amino acids at the spokes of the wheel. Helical wheels have been particularly valuable for identifying proteins and peptides that contain amphipathic helices. In an amphipathic
Contributed by Stanley R. Krystek, Jr., William J. Metzler, and Jiri Novotny Current Protocols in Protein Science (1995) 2.3.1-2.3.20 Copyright © 2000 by John Wiley & Sons, Inc.
UNIT 2.3
Computational Analysis
2.3.1 CPPS
T
K
11
7
P
A 4
14 G
A
15
3
T 10
8
17
1
I
L
6
V
G
12 G
5
13 L
2 I
9 L
16
V
L
Figure 2.3.1 Helical wheel for a portion of the amphipathic helix of melittin. Residues in bold type form a hydrophobic side. Note that this is not a perfect amphipathic helix as the polar face includes Ala-4, Ala-15, and Val-8. This helix also contains several glycine residues, which are not commonly found in helices (Table 2.3.1).
Protein Secondary Structure Prediction
helix, hydrophobic residues will cluster to one side of the wheel and polar residues will be found on the opposite side. Figure 2.3.1 shows a helical wheel for the amphipathic helix of melittin. Helical wheels have often been used for analyzing helical dimerization proteins such as leucine zippers (Krystek et al., 1991; O’Shea et al., 1992), for designing helical multimers (Harbury et al., 1993; Kamtekar et al., 1993; Lovejoy et al., 1993), and for predicting and analyzing membrane proteins which contain helices that span the lipid bilayer (Baldwin, 1993). Other early predictive schemes include those developed by Dunhill (1968) and Lim (1974). Dunhill’s “helical net” is a pictorial method that is similar to the helical wheel and is still used in protein design and helix prediction (DeGrado, 1988; Goodman and Kim, 1991). Lim’s method relies on an extensive set of rules derived from the physicochemical properties of amino acids in conjunction with the analysis of a structural database. The large number of detailed rules, however, makes the Lim method unwieldy. When several methods were compared in a blind test organized by Schulz et al. (1974), the methods of Ptitsyn and Finkelstein (1970) and Chou and Fasman (1974a,b) performed better than others (Burgess et al., 1971; Lewis et al., 1971; Robson and Pain, 1971; Nagano, 1973).
Statistical Prediction Schemes The statistical methods developed by Chou and Fasman (1974a,b) and by Garnier et al. (1978) are the most prominent of the class of predictive schemes based on statistical amino acid preference rules. These secondary structure prediction schemes tacitly assume that only local sequence interactions influence the formation of local structure. This is almost certainly an oversimplification. Significant interactions occur between protein secondary structural elements (tertiary structure); these interactions may involve hydrophobic and/or electrostatic interactions between structural units. Therefore, the success and accuracy of prediction schemes that rely on empirical algorithms are necessarily limited and are unlikely to exceed a level of 70% to 75%. Reports of secondary structure prediction accuracy levels of ∼70%, however, are not uncommon using these methods (Fasman, 1985).
Other Predictive Schemes Two additional methodologies for structure prediction not based on statistics of amino acid occurrence have evolved. Of the neural network methods (Quinn and Sejnowski, 1988; Rost and Sander, 1993a,b; Kneller et al., 1990), that of Rost and Sander claims a level of predictive accuracy exceeding 70% for all proteins tested and is publicly available through the
2.3.2 Current Protocols in Protein Science
Internet computer network (for details, see discussion of Applying the PHD Scheme and Table 2.3.6 therein). The method of Stultz et al. (1993) is based on state-space modeling with a set of predefined structural class models.
Selecting Prediction Tools Secondary structure prediction proceeds as follows. First, it should be confirmed that the three-dimensional structure has not yet been determined. This can be done by searching known X-ray and NMR structures in the Protein Data Bank (Brookhaven, 1994) for an amino acid sequence identical to that of interest. Next, identification of proteins with significantly similar sequences whose structures are already known should be attempted. Numerous software packages are available for performing similarity searches (see UNIT 2.1). If proteins of similar sequence (>20% identity) are found, the Protein Data Bank should again be searched to determine whether the structures of any of those proteins are known. If so, then comparative homology modeling (Greer, 1990; Bajorath et al., 1993) can be used to develop a three-dimensional model of the desired sequence. Only if there are no proteins in the structural database of sufficient similarity should protein secondary prediction schemes be used. It should be pointed out, however, that proteins may have the same fold without a detectable sequence similarity (Kabsch and Sander, 1984, 1985). In the future, sequence threading through known three-dimensional structures will be helpful (Jones et al., 1992; Sippl and Weitckus, 1992). It is recommended that protein secondary structure prediction be performed in conjunction with other sequence analysis approaches, such as hydrophobicity profiling, flexibility analysis, surface exposure experiments, charge distribution mapping, and motif searches (UNIT 2.2; Fasman, 1989b; Genetics Computer Group, 1994). For investigators using methods to predict protein secondary structure, at least two questions should be addressed: (1) which predictive scheme(s) should be used, and (2) what are the advantages of one method over others? This unit attempts to find answers to these questions by studying three proteins in detail: human hemoglobin β-subunit, human interleukin 1β (IL-1β), and human profilin I (see Application of Secondary Prediction Methods and Fig 2.3.2).
METHODS FOR SECONDARY STRUCTURE PREDICTION The Chou and Fasman Method The most popular secondary structure prediction scheme is the statistical method of Chou and Fasman (1974a). Using a database of 15 protein structures determined by X-ray crystallography, Chou and Fasman tabulated the number of occurrences of a given amino acid in α helix, β strand, or coil. From the tabulation, a set of conformational preference parameters for each amino acid type was calculated. The preference parameters were based on three functions: (1) the relative frequency of a specific amino acid type within each protein, (2) the occurrence of that amino acid in a given type of secondary structure, and (3) the fraction of amino acid residues occurring in each type of secondary structure. With the conformational preference parameters in hand, the authors then derived a set of empirical rules for predicting protein secondary structure (Chou and Fasman, 1974b). Chou and Fasman later published an extension of their analysis using a larger set of X-ray structures, totaling 29 proteins and over 4700 amino acids. In addition to refining the values for α helices and β strands, they included a conformational preference parameter for β turns as well (Chou and Fasman, 1977, 1978a,b). For β turns, a positional preference for residues was identified; for example, proline was found to occur more frequently in position 2 of the turn than in positions 1, 3, or 4. A further refined set of conformational preference parameters for helical and sheet residues was later derived from an expanded data set of 64 protein structures (Chou, 1989). These parameters are similar to those based on the 29 protein structures (differences are noted in Chou, 1989). An interesting addition to secondary structure prediction using the conformational preference parameters is the prediction of protein class. Chou presented a computerized algorithm that assigns a structural class to proteins on the basis of amino acid composition; however, these predictions have not been widely used. In the initial studies, Chou and Fasman used the conformational preference parameters to describe amino acids as being formers, breakers, or indifferent for helical and strand regions (Chou and Fasman, 1974a). Their analysis showed that residues with the highest prefer-
Computational Analysis
2.3.3 Current Protocols in Protein Science
A T α, β
HB
B T α, β
HB
C T α, β
HB Residue number Figure 2.3.2 Secondary structure prediction using the method of Chou and Fasman as implemented by Novotny and Auffray (1984). Predicted secondary structure for (A) human hemoglobin β-subunit, (B) interleukin 1β, and (C) human profilin I. Below the amino acid sequence are given the turn prediction (T), the α helix (thin line) and β strand prediction (thick line), the location of positive (upward spikes) or negative charges (downward spikes) as derived specifically from amino acid side chains, and a hydrophobicity profile (HB).
Protein Secondary Structure Prediction
ence for helices resided near the helix center, whereas residues with low helix preference could be found clustered at the helix termini. For helix and strand, residues were independently classified as strong formers, formers, weak formers, indifferent, breakers, and strong breakers. Table 2.3.1 contains a list of the conformational preference parameters and refined conformational assignments derived from the analysis of 64 proteins (Chou, 1989). These
values are combined with the turn preferences shown in Table 2.3.2, or values from the 29protein study (Chou and Fasman, 1978a,b; Prevelige and Fasman, 1989), to carry out sequence prediction as described below. An important contribution to helix prediction strategies is the recognition that helix start and stop signals in the amino acid sequence grammar are quite distinct (Presta and Rose, 1988; Richardson and Richardson, 1988). The
2.3.4 Current Protocols in Protein Science
Table 2.3.1 β Stranda
Conformational Preference and Assignments for α Helix and
α helix
β strand
Residue
Pα
Assignmentb
Glu Ala Met Leu Lys His Gln Phe Asp Trp Arg Ile Val Cys Thr Asn Tyr Ser Gly Pro
1.44 1.39 1.32 1.30 1.21 1.12 1.12 1.11 1.06 1.03 1.00 0.99 0.97 0.95 0.78 0.78 0.73 0.72 0.63 0.55
Hα Hα Hα Hα hα hα hα hα hα Iα Iα iα iα iα iα iα bα bα Bα Bα
Residue
Pβ
Assignmentc
Val Ile Thr Tyr Trp Phe Leu Cys Met Gln Ser Arg Gly His Ala Lys Asp Asn Pro Glu
1.64 1.57 1.33 1.31 1.24 1.23 1.17 1.07 1.01 1.00 0.94 0.94 0.87 0.83 0.79 0.73 0.66 0.66 0.62 0.51
Hβ Hβ hβ hβ hβ hβ hβ hβ Iβ Iβ iβ iβ iβ iβ iβ bβ bβ bβ Bβ Bβ
aValues taken from Chou, 1989. bHα, strong helix former; hα, helix former; Iα, weak helix former; iα, indifferent; bα, helix
breaker; Bα, strong helix breaker. cHβ, strong β-strand former; hβ, β-strand former; Iβ, weak β-strand former; iβ, indifferent; bβ,
β-strand breaker; Bβ, strong β-strand breaker
α-helical structures in proteins are commonly stabilized by reciprocal hydrogen bond formation between the unpaired main-chain amino and carboxyl groups near the helix termini and the side chains of amino acids flanking the termini (Harper and Rose, 1993). The helix-terminal residues are referred to as the N-cap and C-cap. Because of the unique geometry of proline, this amino acid is frequently found at the N-cap + 1 position and has been referred to as being an α-helix initiator (Richardson and Richardson, 1988). Residues that typically occupy the N-cap and C-cap positions of helices are listed in Table 2.3.3.
The Garnier, Osguthorpe, and Robson Method The Garnier, Osguthorpe, and Robson (GOR) method (Garnier et al., 1978; Garnier and Robson, 1989) is a widely available secondary structure prediction method that has been incorporated into several computer packages,
such as that from Genetics Computer Group (GCG). Although the GOR method incorporates essentially all the elements of the ChouFasman statistics, it also uses information theory to reach the final decision. A detailed description of the GOR method can be found elsewhere (Garnier and Robson, 1989). In principle, the GOR method assigns the conformation of a given residue by considering the contribution of residues relatively distant to it in the sequence. On the basis of studies showing the interdependence of residue conformations up to eight amino acids away, the GOR method assigns single-residue information values to each amino acid along the polypeptide chain. The predicted conformational state for an amino acid is the state with the highest positive information value. An extension of this procedure, the GORIII method, has been developed to use pair information instead of directional information. The GORIII method extracts both single-residue information values
Computational Analysis
2.3.5 Current Protocols in Protein Science
Table 2.3.2
Conformational Preference and Assignments for β Turnsa
Positional preferences for β turnsb
Residue Preference fi Asn Gly Pro Asp Ser Cys Tyr Lys Gln Thr Trp Arg His Glu Ala Met Phe Leu Val Ile
1.56 1.56 1.52 1.46 1.43 1.19 1.14 1.01 0.98 0.96 0.96 0.95 0.95 0.74 0.66 0.60 0.60 0.59 0.50 0.47
Asn Cys Asp His Ser Pro Gly Thr Tyr Trp Gln Arg Met Val Leu Ala Phe Glu Lys Ile
0.161 0.149 0.147 0.140 0.120 0.102 0.102 0.086 0.082 0.077 0.074 0.070 0.068 0.062 0.061 0.060 0.059 0.056 0.055 0.043
fi+2
fi+1 Pro Ser Lys Asp Thr Arg Gln Gly Asn Met Ala Tyr Glu Cys Val His Phe Ile Leu Trp
0.301 0.139 0.115 0.110 0.108 0.106 0.098 0.085 0.083 0.082 0.076 0.065 0.060 0.053 0.048 0.047 0.041 0.034 0.025 0.013
Asn Gly Asp Ser Cys Tyr Arg His Glu Lys Thr Phe Trp Gln Leu Ala Pro Val Met Ile
0.191 0.190 0.179 0.125 0.117 0.114 0.099 0.093 0.077 0.072 0.065 0.065 0.064 0.037 0.036 0.035 0.034 0.028 0.014 0.013
fi+3 Trp Gly Cys Tyr Ser Gln Lys Asn Arg Asp Thr Leu Pro Phe Glu Ala Ile Met His Val
0.167 0.152 0.128 0.125 0.106 0.098 0.095 0.091 0.085 0.081 0.079 0.070 0.068 0.065 0.064 0.058 0.056 0.055 0.054 0.053
aValues taken from Chou and Fasman (1977, 1978a,b). bThe positional preference f is defined as the relative occurence of a specific amino acid found in position i of a β turn. i
Table 2.3.3 Amino Acid Positional Preferences in α Helicesa
Effect
Residuesb
Amino-terminal stabilizing (N-cap)
Gly Ser, Thr Asp, Glu Asn, Gln His Gly Asn His(+), Lys, Arg His(+), Lys, Arg Val, Leu, Ile, Met Ser Asp Val, Leu, Ile, Met
Carboxyl-terminal stabilizing (C-cap)
Amino-terminal destabilizing Carboxyl-terminal destabilizing
aAmino acid preferences were compiled from Fersht and Serrone (1993),
Presta and Rose (1988), and Richardson and Richardson (1988).
Protein Secondary Structure Prediction
bResidues listed in order of increasing magnitude of the effect.
2.3.6 Current Protocols in Protein Science
and pair information values for each possible pair and for amino acids within eight amino acids of a central residue. The predictive accuracy of the GOR methods is similar to that of the Chou and Fasman method (Garnier et al., 1978). Thus, the GOR methods may serve as a check of the results obtained with the Chou and Fasman method.
The EMBL Profile Neural Network Method An important feature of the EMBL Profile Neural Network (PHD) prediction scheme is the use of multiple sequence alignments (Rost and Sander, 1993a,b). Previous studies have shown that homologous proteins (generally with at least 20% to 30% sequence identity but often less) frequently have identical three-dimensional folds (tertiary structure). Thus, multiple sequence alignments are grouped by structural family and used as input. By considering many aligned sequences, the algorithm makes use of more information about structure than is available from just the individual protein sequence for which the secondary structure is to be predicted. To determine which protein sequences are to be used in the prediction, the PHD method uses a weighted dynamic programming method, MaxHom (Sander and Schneider, 1991), in searching a database of sequences. A family profile for the input sequence is generated and the amino acid frequencies at each alignment position are calculated for use in the prediction. Results have shown that the multiple sequence alignments improve secondary structure prediction >6% compared to singlesequence predictions (Rost and Sander, 1993a,b). The PHD method uses a reference network or neural network that has been trained and tested on a database of nonhomologous protein structures. In a typical database there is an uneven distribution of secondary structure types, such that there are many more examples of turns than of helices or sheets (Rost and Sander, 1993a,b). Because of the uneven distribution, empirically based predictions of loops are more accurate than those of helices, which in turn are more accurate than those of sheets. The neural network approach of the PHD method eliminates this bias: Rost and Sander developed a network that is trained with each type of secondary structure in equal proportion, rather than the proportions present in any database of protein structures. This results in a more balanced prediction.
The network system for structure prediction consists of three layers. In the first layer, secondary structure is predicted on the basis of the multiple aligned sequences. The results are passed to the second layer, which collectively considers secondary structure elements that are adjacent in the protein sequence, thereby allowing nearest neighbor structure to influence the structure of a particular region of the protein. The third layer of the network averages the results of the independently trained networks. The output from the third layer is determined by a “winner take all” prediction of the secondary structure at a given sequence position (Rost and Sander, 1993a,b).
The PSA Method The Protein Sequence Analysis (PSA) method (Stultz et al., 1993) computes the probability that a given amino acid belongs to a particular secondary structure element when there are no homologous protein sequences or structural data. To do this, mathematical models were developed for 15 single-domain protein superclasses or macroclasses. These superclasses, patterned after the protein class categorization of Richardson (1981), are groups of proteins with similar secondary and tertiary structures. The mathematical models represent constraints on the patterns of secondary structure elements for a given protein class. To perform structure prediction, the PSA method uses the amino acid sequence to calculate the probability that a given residue is contained in each of the modeled secondary structure elements. Also calculated is the probability that the submitted sequence is a member of one of the predefined superclasses. Thus, in addition to evaluating the secondary structure prediction, the PSA method provides the bonus of the potential tertiary structural information contained in the predicted structural superclass.
The Helical Wheel The helical wheel is a plot of the amino acid residues around a potentially helical segment (see Fig. 2.3.1 and discussion in Early Predictive Schemes). The method was developed to find helices with a hydrophobic face buried away from a polar solvent, with the graphical representation showing the clustering of polar and/or nonpolar residues toward one face of a helix. In its original use the helical wheel facilitated the identification of potential helical segments in protein sequences (Schiffer and Edmundson, 1967), but its applicability has since been expanded to include designing proteins (e.g.,
Computational Analysis
2.3.7 Current Protocols in Protein Science
leucine zipper proteins; Harbury et al., 1993; Kamtekar et al., 1993; Lovejoy et al., 1993) and studying transmembrane proteins (e.g., G-protein-coupled receptors; Baldwin, 1993).
APPLICATION OF SECONDARY STRUCTURE PREDICTION METHODS Applying the Chou and Fasman Method The wide use of the Chou and Fasman method is due in part to the simplicity of application, ease of understanding, and potential interactive nature of the analysis. It also ranks favorably in accuracy of prediction relative to other methods. Application of the method entails the following steps. 1. Conformational preference parameters for each type of secondary structure are locally smoothed by averaging techniques (see UNIT 2.2 for an example of the scanning window averaging method) in order to calculate α-helix, β-sheet, and turn probabilities along the polypeptide backbone. 2. The α-helix, β-strand, and turn probabilities are plotted versus sequence position for interpretation. Basically, the conformation is assigned according to the highest curve (Fig. 2.3.2, Fig. 2.3.3, and Fig. 2.3.4). 3. On the basis of conformational preference parameters, amino acids are assigned as formers, breakers, or indifferent for α-helical and β-sheet regions (Fig. 2.3.3A). 4. Program options usually include full predictions that are based on the original Chou and Fasman rules (Table 2.3.4). Predictions can also be made by the user in an interactive environment by combining the Chou and Fasman rules with sequence analysis using other empirical methods such as hydrophobicity profiling, flexibility prediction, charged residue mapping, and surface probability calculation (Fig. 2.3.3 and Fig. 2.3.4; see also Comparison of Prediction Schemes). We find that an interactive approach allows for the best interpretation of the prediction results (Fasman, 1989a; Schulz, 1988). Several other correlations can be used in this analysis and are listed in Table 2.3.5. Many programs can also generate sketches of predicted peptide structures (Fig. 2.3.5).
Applying the GOR Scheme Protein Secondary Structure Prediction
The GOR scheme is coded into many commercial and academic protein structure analysis packages. This method provides estimated
probabilities that a given residue is in a particular conformational state, and it predicts the conformation of an amino acid without any input from the user other than the sequence of interest. There are two outputs from the GOR method. The first is a plot of the turns, α helices, and β strands as predicted from the method. The second is a summary of the full protein structure prediction similar to that produced by the Chou and Fasman method. Unlike the Chou and Fasman method, assignment of each amino acid to a structural element is made automatically. Because many prediction schemes can be printed in the same output format, it is possible to print results of Chou and Fasman, GOR, and other structural predictions together for a more interactive analysis of the sequence (Fig. 2.3.4). Use of a combination of structural prediction schemes and the secondary structure correlations presented in Table 2.3.5 should increase the accuracy of secondary structure prediction.
Applying the PHD Scheme Access to the PHD scheme is available only through electronic mail or the WWW (World Wide Web). The Internet address for this service is given in Table 2.3.6. To perform sequence analysis with the PHD scheme, one merely needs to supply the sequence to the EMBL Predictprotein server. Additionally, one can submit prealigned sequences or a defined set of homolog sequences to be used in the predictive scheme. If a single sequence is submitted, the scheme will search the sequence database, and, if homologs are found, the sequence prediction will be performed on the multiple sequence alignments (Fig. 2.3.6). The output from the scheme consists of several pages describing the network and analysis, including the list of any homologous proteins along with the sequence alignment used in the analysis, the definition of accessibility, and a reliability index for the accuracy of prediction (Fig. 2.3.7). This method provides prediction of solvent accessibility as well as secondary structure. A comparison of this analysis to others described in this unit is described in the section on Comparison of Prediction Schemes.
Applying the PSA Scheme The PSA scheme, like the PHD scheme, has been established on the Internet (Table 2.3.6). The electronic mail server, called Protein Sequence Analysis (PSA) System, accepts as input the amino acid sequence of a protein. After sequence analysis the server returns four
2.3.8 Current Protocols in Protein Science
A
PEPPLOT
0 A G W N A Y I D N L M A D G
50 T C Q D A A I V G Y K D S P S V W A A V P G K T F
V N I T P A E V G V L V G K D R S S F Y V N G L
100 T L G G Q K C S V I R D S L L Q D G E F S M D L
R T K S T G G A P T F N V T V T K T D K T L V L L
M G K E G V H G G L I N K K C Y E M A S H L R R
S Q Y
Basic Acidic
1.5
HPhobic Hphilic Beta Forming Beta Breaking
Beta Chou & Fasman
1.0 Alpha 0.5 Alpha Forming 15 Alpha Breaking Beta Alpha 0 Beta Alpha 5
NH2 End
15 COOH End 0 Turn Hydrophobic Moment
0
1
Beta Alpha
0
3
Goldman et al. Kyte-Doolittle
0
HPhobic HPhilic
–3
0
50
100
B
Alpha Alpha Beta Beta Alpha Beta Pos Res Stat Ave Stat Ave NH2 COOH NH2 COOH Turn ------------------------------------------------------------------ .. 1 A 1.42 0.94 0.83 0.96 0.43 0.17 0.29 0.56 0.30 2 G 0.57 0.94 0.75 0.96 0.73 0.04 2.33 0.42 0.15 3 W 1.08 0.97 1.37 1.14 0.57 0.25 0.47 0.76 0.28 4 N 0.67 0.97 0.89 1.20 1.08 0.78 0.75 1.41 0.78 5 A 1.42 1.05 0.83 1.11 0.51 1.25 0.16 1.75 0.04 6 Y 0.69 0.86 1.47 1.13 0.25 0.31 1.24 1.27 0.45 7 I 1.08 0.99 1.60 1.08 0.58 0.08 2.44 0.52 0.63 8 D 1.01 1.09 0.54 0.95 2.19 0.12 0.56 1.08 0.24 9 N 0.67 1.19 0.89 1.02 1.98 0.95 0.09 2.40 0.03 10 L 1.21 1.27 1.30 0.93 0.82 3.18 0.04 2.33 0.14 11 M 1.45 1.11 1.05 0.79 0.31 1.52 0.31 0.63 1.41 12 A 1.42 0.96 0.83 0.83 0.84 0.17 2.28 0.22 0.99 13 D 1.01 0.78 0.54 0.92 2.45 0.13 1.21 0.25 1.04 14 G 0.57 0.80 0.75 1.06 3.18 0.13 1.15 0.46 1.26 15 T 0.83 0.91 1.19 1.01 1.79 0.29 0.45 0.39 0.14 16 C 0.70 1.06 1.19 0.92 0.53 0.19 2.91 0.18 1.52 17 Q 1.11 1.24 1.10 0.83 0.27 0.13 2.86 0.10 0.17 18 D 1.01 1.23 0.54 0.95 0.15 0.40 1.29 0.25 0.22 19 A 1.42 1.25 0.83 1.24 0.12 0.71 0.30 0.76 0.03 20 A 1.42 1.03 0.83 1.22 0.14 3.45 0.11 2.52 0.09 21 I 1.08 0.85 1.60 1.38 0.75 0.78 0.15 1.95 0.49 22 V 1.06 0.87 1.70 1.17 1.03 0.56 0.18 4.19 0.57 23 G 0.57 0.86 0.75 0.88 4.24 0.87 0.31 3.57 0.39 24 Y 0.69 0.91 1.47 0.88 1.08 1.11 0.49 4.63 1.79 25 K 1.16 0.88 0.74 0.65 1.39 0.50 1.50 0.74 0.51
HPhob –0.56 –0.33 –0.08 0.22 0.22 –0.02 0.04 –0.18 –0.18 –0.18 –0.40 –0.41 –0.11 0.74 0 .74 0 .68 –0.03 –0.03 0.27 –0.11 –0.40 –0.43 –1.00 –0.76 –0.41
Figure 2.3.3 Secondary structure prediction for human profilin I using the method of Chou and Fasman as implemented by GCG:PepPlot algorithm. (A) Predicted secondary structure and (B) numerical output for residues 1 through 25.
output files to the requester: a text file summarizing the tertiary class probabilities and most probable superclass; a file containing plots of the class and superclass probabilities; a file containing the secondary structure probability distributions displayed on a contour plot; and a file containing the secondary structure probability distributions on x-y plots. The secondary structure and protein class are easily deter-
mined from the combination of plots (Fig. 2.3.8) and can be compared with those suggested by other structural prediction methods.
COMPARISON OF PREDICTION SCHEMES The secondary structures of hemoglobin (an all-α protein), interleukin 1β (an all-β protein), and profilin (an α/β protein) were predicted
Computational Analysis
2.3.9 Current Protocols in Protein Science
A
PLOTSTRUCTURE
50
100
50
100
5.0 KD Hydrophilicity –5.0 10.0 Surface Prob. 0.0 1.2 Flexibility 0.8 1.7 Jameson-Wolf (Antigenic Index) –1.7 CF Turns CF Alpha Helices CF Beta Sheets GOR Turns GOR Alpha Helices GOR Beta Sheets Glycosyl Sites
B
PEPTIDESTRUCTURE Hydrophilicity (Kyte-Doolittle) averaged over a window of: 7 Surface Probability according to Emini Chain Flexibility according to Karplus-Schulz Secondary Structure according to Chou-Fasman Secondary Structure according to Garnier-Osguthorpe-Robson Antigenicity Index according to Jameson-Wolf Pos 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35
Protein Secondary Structure Prediction
AA A G W N A Y I D N L M A D G T C Q D A A I V G Y K D S P S V W A A V P
GlycoS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
HyPhil
0.750 0.240 0.417 –0.286 0.471 0.914 0.243 –0.529 –0.529 –0.214 0.486 0.086 -0.771 0.271 1.043 1.043 0.286 –0.414 –1.114 –0.700 –1.014 –0.957 –0.200 0.171 1.043 1.757 1.100 1.043 0.229 –0.529 –1.243 –1.243 –1.300 –0.143 –0.171
SurfPr
FlexPr
0.633 0.500 0.613 0.426 0.718 1.099 0.563 0.552 0.356 0.848 0.502 0.451 0.293 0.513 0.848 0.513 0.523 0.254 0.352 0.201 0.189 0.374 0.618 1.181 2.460 3.331 1.578 0.830 0.502 0.378 0.182 0.210 0.279 0.531 0.759
1.000 1.000 1.000 1.000 0.921 0.927 0.946 0.962 0.959 0.958 0.960 0.983 1.016 1.039 1.049 1.036 1.018 0.994 0.963 0.938 0.921 0.927 0.949 0.989 1.030 1.056 1.068 1.042 1.000 0.951 0.908 0.906 0.929 0.975 1.029
CF-Pred GORPred . . . . . . h h h h h h t t t t T T B B B B B B T T . T T B B B B B T
. . . . . B B B B B T T T T T B B B B B B B B T T . . . . B B B B B .
AI-Ind .. 0.750 0.450 0.450 –0.150 0.300 0.750 0.300 –0.600 –0.600 –0.300 0.700 0.700 0.150 1.050 1.350 0.950 0.850 –0.200 –0.600 –0.600 –0.600 –0.600 –0.300 0.850 1.700 1.300 0.900 1.150 0.850 –0.600 –0.600 –0.600 –0.600 –0.300 0.250
Figure 2.3.4 Secondary structure prediction for human profilin I using the methods of Chou and Fasman and GOR as implemented by GCG: PeptideStructure algorithm and plotted using PlotStructure utility. (A) Predicted secondary structure and (B) numerical output for residues 1 through 35.
2.3.10 Current Protocols in Protein Science
with the Chou and Fasman, GOR, PHD, and PSA methods (Fig. 2.3.9). Analysis of the Chou and Fasman profiles was enhanced by displaying smoothed hydrophobicity profiles and consulting the amino acid positional preferences of Table 2.3.3. An example of the complete
analysis of profilin is presented in Figure 2.3.3, Figure 2.3.4, and Figure 2.3.5 (or Fig. 2.3.2C). The GOR predictions were made with the GCG commercial package (Fig. 2.3.4). The PHD and PSA predictions were obtained from the electronic mail response to a structure query; ex-
Abbreviated Chou and Fasman Prediction Rulesa
Table 2.3.4
Feature
Rule
Helix
Helix is initiated by a cluster of four helix-promoting residues out of six amino acids along the polypeptide sequence. Helical segment is extended in both directions until sets of tetrapeptide breakers are reached. Prolines can occur only within the first three residues of the N-terminal end. A segment length of six or more amino acids with α probability greater than β probability is predicted as helical. A cluster of three β formers or a cluster of three β formers out of five amino acids along the polypeptide sequence will initiate β-strand formation. The β strand is extended in both directions until terminated by a set of tetrapeptide breakers. A segment of three or more amino acids with β probability greater than α probability is predicted as β strand. Turn probability is calculated as peaks of probability that are greater than α or β probability. The β turns are also identified by Pt > 0.75 × 10−4, where Pt = fi × fi+1 × fi+2 × fi+3. For a region containing overlapping α helix and β strand, the prediction would be for helix if the α probabilities are greater than β probabilities, and strand if β probabilities exceed α probabilities.
Strand
Turns
Overlap
aFor the complete set of prediction rules, see Chou and Fasman (1974a,b, 1978a,b).
Table 2.3.5
Secondary Structure Correlations
Structure
Characteristics
β strands
Often correspond to peaks of hydrophobicity β-branched amino acids (Val, Ile, Thr) are typical Positively charged residues are relatively abundant Negatively charged residues (Glu) virtually never occur Average residue length is 7, resulting in short and acute peaks in hydrophobicity plots (see UNIT 5.2) β bulges may occur in the edge strands. Their signature is small amino acids and glycines (e.g., Gly-Xxx-Gly sequence in immunoglobulin V fold, strand G). Hydrophobicity peaks are longer, with a fine structure (teeth). Ideally, fine structure shows n + 4 periodicity. Glu, Ala, Leu, are typical Amino acid preferences exist at the N-cap and C-cap positions (see Table 2.3.3) Prolines can occur in the first turn of helices in globular proteins, and most frequently occupy the N-cap + 1 position Prolines, which “kink” the helices, are overrepresented in transmembrane helices Glycines frequently occur at the C-cap position Hydrophobic residues often occupy positions N-cap + 4 and C-cap − 4 Helices may be completely buried and not amphipathic
α helices
Computational Analysis
2.3.11 Current Protocols in Protein Science
PLOTSTRUCTURE NH2
50 0
100 0
COOH
Figure 2.3.5 Sketch of the human profilin secondary structure as predicted in Figure 2.3.3 and Figure 2.3.4 by Chou-Fasman method.
Protein Secondary Structure Prediction
amples of the output for the profilin secondary structure prediction are provided in Figure 2.3.6, Figure 2.3.7, and Figure 2.3.8. Figure 2.3.9A summarizes the results of secondary structure prediction for the β subunit of human hemoglobin. Immediately below the amino acid sequence is the location of the eight helices found in the three-dimensional structure of hemoglobin (Brookhaven Protein Data Bank entry 2hhb; Fermi, 1975). The Chou and Fasman scheme predicts most of the helical residues in helices 1, 2, 3, 4, 6, 7, and 8. Helices 6 and 7 are predicted to be longer than they actually are, and helices 2 and 3 are not distinguished as independent helices. A turn is predicted in place of the fifth helix. The GOR method predicts most of the residues for helices 1, 2, 5, 6, and 8, but it predicts a turn for helix 3 as well as a turn and a strand for helix 7. The PHD program found over 400 sequence homologs of hemoglobin-β, with one of the sequences being the submitted sequence. The PHD prediction identified a homolog with a known three-dimensional structure in the protein database and also suggested structure prediction by homology modeling. The prediction by PHD did quite well, accounting for seven of the eight helices and missing only the small helix 4. The PSA procedure suggested the α/β class as the most probable. Most of the helical residues in helices 2, 3, 5, 6, and 8 and part of helix 7 were correctly predicted. Helix 1 was missed by the PSA method, however, and the
C- and N-terminal residues of helices 2 and 3, respectively, were predicted to be in sheet structure. Figure 2.3.9B summarizes the results of secondary structure prediction for interleukin 1β. IL-1β contains 12 β strands connected by intervening turns (Brookhaven Protein Data Bank entry 1i1b; Finzel et al., 1989). The Chou and Fasman method predicted most of the residues in 8 of the 12 strands in the β sheet (strands 1, 2, 4, 5, 6, 8, 11, and 12). Helical regions were predicted for β strands 3 (with the adjacent turn) and 7. The Chou and Fasman method correctly predicted several of the turns. The GOR method correctly predicted 4 of the 12 β strands, but it predicted helical regions for β strands 3 (with the adjacent turn), 5, 6, and 8. Interestingly, both the Chou and Fasman and GOR methods predict helix for the third strand. This may be due to the negatively charged glutamate, which is generally considered a strand breaker (see Table 2.3.1). For IL-1β, the PHD output listed the 10 sequence homologs that were used in the structure prediction and also identified a known three-dimensional structure in the protein database. The PHD method correctly predicted most of the residues in the 12 β strands; however, a deficiency in the method is that it does not predict the location of possible turns. The PSA method did not successfully predict either the class or the secondary structure of IL-1b: the method pre-
2.3.12 Current Protocols in Protein Science
Table 2.3.6
Commercial Packages for Sequence Analysis
Software
Platforma
Vendor or Internet locationb
Sybyl/Biopolymerc InsightII/Homologyc GCG MacVector Geneworks PC/Gene DNASTAR
SGId SGId VAX Macintosh Macintosh IBM-compatible IBM-compatible or Macintosh
Tripos Associates Biosym Technologies Genetics Computer Group International Biotechnologies IntelliGenetics/Betagen IntelliGenetics/Betagen DNASTAR
Internet locations for sequence analysis PHD methode
PSA methode
EMBL Predictprotein Server
[email protected] [email protected] http://www.embl-heidelberg.de/predict protein/predict protein.html Protein Sequence Analysis Server
[email protected]
aSoftware may run on other platforms in addition to these listed here. For a comprehensive list, contact
the appropriate vendor. bFor suppliers addresses, see SUPPLIERS APPENDIX. cBiopolymer and Homology are optional modules that can be purchased to run with Sybyl and InsightII, respectively. dSilicon Graphics Inc. ePHD and PSA methods are currently accessible only through the Internet; thus, results can be accessed by any platform.
dicted the protein fold class to be all-α rather than all-β, and therefore incorrectly predicted α structure rather than β structure. It is noteworthy that several of the predicted α helices correspond to the location of β strands, namely, those of β strands 2, 3, 4, 5, 7, 9, 11, and 12. Further testing is required to determine whether there are any inherent problems with the predictions of all-β proteins by the PSA method. Figure 2.3.9C summarizes the results of secondary structure prediction for human profilin, an α/β protein whose high-resolution structure (Metzler et al., 1993, 1995) was not publicly known at the time of testing. Profilin is composed of a central seven-stranded, antiparallel β sheet and four helices (Fig. 2.3.10). Seven turns that connect the helical and sheet structural elements in the structure have been identified. Figure 2.3.2C, Figure 2.3.3A, and 2.3.4A show typical Chou and Fasman profiles as determined with the programs of Novotny (Novotny and Auffray, 1984), PepPlot (GCG, 1994), and Plotstructure (GCG, 1994). Figure 2.3.3B shows the numerical output from the modified Chou and Fasman prediction pro-
grams (PepPlot), and Figure 2.3.4B shows the automated prediction by the Peptidestructure program using the Chou and Fasman and the GOR methods. A schematic representation of the predicted protein secondary structure is shown in Figure 2.3.5. The Chou and Fasman method predicted six of the seven β strands and the two terminal helices of profilin (Fig. 2.3.8C). This method also predicted correct locations of several of the turns located between the strands. In comparison, the GOR method performed significantly worse, predicting only four of seven β strands and neither of the terminal helices. The PHD method identified six sequence homologs of profilin (Fig. 2.3.6) that it used in the secondary structure prediction (Fig. 2.3.6). The method correctly predicted all seven β strands and the two terminal helices. It did, however, predict additional β strands for the interior helices but did not predict the locations of any turns (owing to a deficiency in the method, as stated previously). The PSA method predicted the profilin sequence to be a member of the all-β protein fold class (Fig. 2.3.8). The method did
Computational Analysis
2.3.13 Current Protocols in Protein Science
## PROTEINS : EMBL/SWISSPROT identifier and alignment statistics NR. ID STRID %IDE %WSIM IFIR ILAS JFIR JLAS LALI NGAP PROTEIN 1 : pro1_human 1.00 1.00 1 139 1 139 139 0 PROFILIN I. 2 : prof_mouse 0.96 0.97 1 139 1 139 139 0 PROFILIN. 3 : prof_bovin 0.95 0.97 1 139 1 139 139 0 PROFILIN. 4 : pro2_human 0.62 0.71 1 139 1 139 139 0 PROFILIN II. 5 : prof_varv 0.32 0.48 1 137 2 131 130 2 PUTATIVE PROFILIN. 6 : prof_vaccv 0.31 0.47 1 137 2 131 130 2 PUTATIVE PROFILIN. ## ALIGNMENTS 1 6 SeqNo PDBNo AA STRUCTURE BP1 .4....:....5....:....6....:....7 1 A U 0 2 G U 0 3 W U 0 4 N U 0 5 A U 0 6 Y U 0 7 I U 0 8 D U 0 9 N U 0 10 L U 0 11 M U 0 12 A U 0 13 D U 0 14 G U 0 15 T U 0 16 C U 0 17 Q U 0 18 D U 0 19 A U 0 20 A U 0 21 I U 0 22 V U 0 23 G U 0 24 Y U 0 25 K U 0
BP2 ACC 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0
0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0
NOCC 6 6 6 6 6 6 6 6 6 6 6 6 6 6 6 6 6 6 6 6 6 6 6 6 6
VAR 0 30 0 35 50 41 5 15 29 21 53 56 24 33 51 47 24 6 0 0 0 0 24 0 27
LGAP LSEQ2 ACCNUM 0
139
P07737
0
139
P10924
0
139
P02584
0
139
P35080
7
133
P33828
7
133
P20844
....:....1....:....2....:....3....:... AAAAAA GGGGEE WWWWWW NNNQHH AAASKK YYYYII IIIVII DDDDEE NSNNDD LLLLII MMMMSS AAACKK DDDDNN GGGGNN TTTCNK CCCCFF QQQQEE DDDEDD AAAAAA AAAAAA IIIIII VVVVVV GGGGDD YYYYYY KKKCKK
Figure 2.3.6 Sequence homologs and a portion of the sequence alignment statistics used in secondary structure prediction for human profilin I with PHD, the EMBL neural network method. Secondary structure prediction given in Figure 2.3.7. For description of column labels see Rost and Sander (1993a,b) and EMBL Predictprotein Server (address given in Table 2.3.6).
Protein Secondary Structure Prediction
correctly predict the location of five of the β strands, but it predicted additional β strands for the amino and carboxyl termini and the second interior helices. The PSA method correctly predicted several of the turns located between the β strands, as well. In summary, both the PHD and the interactive Chou and Fasman method performed well, correctly predicting most secondary structural elements (see Table 2.3.7). The interactive Chou and Fasman method generally outperformed the GOR method. However, it must be
pointed out that in predicting the secondary structure with the Chou and Fasman method, the correlations contained in Table 2.3.3 and Table 2.3.5 and the Chou and Fasman rules contained in Table 2.3.4 facilitated both locating helix ends and discriminating between helix and strand when ambiguity existed. In contrast, a strict interpretation of the GOR results was used. It is also possible to interpret the GOR results interactively; this may equalize the predictive power of the two methods. In testing the PHD method, we were unable
2.3.14 Current Protocols in Protein Science
Table 2.3.7 Prediction
Summary of Secondary Structure
Structure Hemoglobin Interleukin 1β Profilin
Method (%)a C-F
GOR
PHD
PSA
78 53 60
62 41 44
86 67 87
60 — 48
aNumber of amino acids correctly predicted, divided by the total
number of amino acids known to be in helices, sheets, or turns. No value is provided for the PSA analysis of interleukin 1β because the class predicted was all-α.
protein:
2a20
length
139
....,....1....,....2....,....3....,....4....,....5....,....6 AA |AGWNAYIDNLMADGTCQDAAIVGYKDSPSVWAAVPGKTFVNITPAEVGVLVGKDRSSFYV| PHD sec | HHHHHHHHH EEEEE EEEEEE EEEEE EEEEEEEE EEEE| Rel sec |965679988733797356089964258635986257448665481777999726874787| detail: prH prE prL subset: SUB
sec sec sec sec
ACCESSIBILITY 3st: P_3 acc 10st: PHD acc Rel acc subset: SUB acc
|026778888763100210000000000110000000000000001000000000000000 |000000010000001211489876421156887521268777304788989752116788 |972110001136897567410013568622012368631222684111000147883111 |LLHHHHHHHH..LLL.LL.EEEE..LLL.EEEE.LL..EEEE.L.EEEEEEE.LLL.EEE |
|eebeebbebbbeeeebebbbbbbbee eebbbbbbeebbbeb be bbbbbeee eebbb| |980670060007777060000000775770000007700070507400000777576000 |842231810404525012897740440647597304515146213084247355141324 |ee....b..b.ee.e...bbbbb.ee.eebbbb..ee.b.eb....bb.bb.ee.e...b|
....,....7....,....8....,....9....,....10...,....11...,....1 AA |NGLTLGGQKCSVIRDSLLQDGEFSMDLRTKSTGGAPTFNVTVTKTDKTLVLLMGKEGVHG| PHD sec |EEEEE EEEEEEEEE EEEEEE EEEEEE EEEEEE | Rel sec |324627717999975224589315997625799988313999975525888882678767| detail: prH prE prL subset: SUB
sec sec sec sec
ACCESSIBILITY 3st: P_3 acc 10st: PHD acc Rel acc subset: SUB acc
|000000000000000111000011000001000000000000001121100000011100 |656751148999986542200346887741100011345899982126888884100011 |342247851000012336788542001156898988653000016741000005788877 |...E.LL.EEEEEEE...LLL..EEEEE.LLLLLLL...EEEEELL.EEEEEE.LLLLLL |
|bbbebbeeebbbb bbbbeebebbbeb beeeeebbbbebbbbebbebbbbbbbeeebee| |000600776000050000770700060507779700006000060070000000776077 |201100442808311640440542725137454511011738111055997684450044 |......ee.b.b...bb.ee.eb.b.b..eeeee.....b.b....ebbbbbbbee..ee |
2...,....13...,....14...,....15...,....16...,....17...,....1 AA |GLINKKCYEMASHLRRSQY| PHD sec | HHHHHHHHHHHHHH | Rel sec |8424899999999972589 detail: prH prE prL subset: SUB
sec sec sec sec
|0146889999999985200 |0210000000000000000 |8632100000000014689 |L...HHHHHHHHHHH.LLL|
ACCESSIBILITY 3st: P_3 acc 10st: PHD acc Rel acc subset: SUB acc
|eebeeeb ebbeeb eeee| |7606770570076057789 |4131359142931413469 |e....eb.e.b..b..eee|
Figure 2.3.7 Secondary structure prediction for human profilin I with PHD, the EMBL neural network method. Sequence homologs and a portion of the sequence alignment used in the analysis are given in Figure 2.3.6.
Computational Analysis
2.3.15 Current Protocols in Protein Science
Superclass Probabilities for Sequence p using vpp6.m, 15-Dec-94 (10:50:50)
Probability
1 0.8 0.6 0.4 0.2 0 alpha-beta
alpha
irregular
beta
Probability of being in a strand for Sequence p (psp2.m), 15-Dec-94 (10:53:40)
1 0.5 0 Probability of being in a turn for Sequence p (psp2.m), 15-Dec-94 (10:53:40)
Probability
1 0.5 0 Probability of being in a helix for Sequence p (psp2.m), 15-Dec-94 (10:53:39)
1 0.5 0 0
20
40
60
80
100
120
130
Residue position
Figure 2.3.8 Class and secondary structure predictions for human profilin I with the PSA method.
to identify any protein whose structure is included in the Brookhaven Protein Data Bank for which there are no sequence homologs in sequence databases. If such a sequence and structure were available, the comparison of the predictive methods might have been less biased. However, this point highlights an advantage of the PHD method. Sequence homologs with known three-dimensional structures often help achieve an accurate prediction of secondary structure. The availability of homologous amino acid sequences imparts a significant predictive advantage to the PHD method in many cases.
Protein Secondary Structure Prediction
analysis (which requires interpretation of the output data) and PHD neural network predictions (which are automatic) is recommended as the most accurate and preferable procedure. As three-dimensional protein structural databanks grow, the future of protein structure prediction may lie with methods that “thread” the polypeptide sequence onto known motifs and evaluate the resulting model by pseudoenergetic analysis (e.g., PROSA; Sippl and Weitckus, 1992). Such programs currently exist, and successful fold identifications are being reported. However, the probability of a successful sequence-structure match is currently 10% (Sippl and Weitckus, 1992).
RECOMMENDATIONS AND THE FUTURE
CONCLUSIONS
On the basis of the examples presented here and our experience in general, a combination of the Chou and Fasman interactive sequence
1. Empirically derived rules can aid sequence prediction using the GOR and Chou and Fasman statistical methods. Summaries of
2.3.16 Current Protocols in Protein Science
A
1 Hem. Structure C-F GOR PHD PSA Hem. Structure
B
C
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VHLTPEEKSAVTALWGKVNVDEVGGEALGRLLVVYPWTQRFFESFGDLSTPDAVMGNPKVKAHGKKVLGAFSDGLA
80
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HLDNLKGTFATLSELHCDKLHVDPENFRLLGNVLVCVLAHHFGKEFTPPVQAAYQKVVAGVANALAHKYH
C-F GOR PHD PSA 1
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Il-1b. Structure C-F GOR PHD PSA
APRVRSLNCTLRDSQQKSLVMSGPYELKALHLQGQDMEQQVVFSMSFVQGEESNEKIPVALGLKEKNLYLSCVLKDD
Il-1b. Structure C-F GOR PHD PSA
KPTLQLESVDPKNYPKKKMEKRFVFNKIEINNKLEFESSAQFPNWYISTSQAENMPVFLGGTKGGQDITDFTMQFVSS
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Profilin. Structure C-F GOR PHD PSA
AGWNAYIDNLMADGTCQDAAIVGYKDSPSVWAAVPGKTFVNITPAEVGVLVGKDRSSFYVNGLTLGG
Profilin. Structure
QKCSVIRDSLLQDGEFSMDLRTKSTGGAPTFNVTVTKTDKTLVLLMGKEGVHGGLINKKCYEMASHLRRSQY
80
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C-F GOR PHD PSA
Figure 2.3.9 Summary of the secondary structure predicted for (A) human hemoglobin β-subunit, (B) human interleukin 1β, and (C) human profilin I. Each panel lists, from top to bottom, the amino acid sequence, the secondary structure determined experimentally, and the secondary structure predicted by each of the following: C-F, the Chou and Fasman method; GOR, the GOR method; PHD, the EMBL neural network method; and PSA, the state-space method. The location of helices (solid bars), turns (open bars), and sheet structure (hatched bars) are indicated.
these correlations (rules) are contained in Tables 2.3.4 and 2.3.5. 2. Methods that are open to interpretation do not guarantee the same result when applied by different users. 3. The PHD and PSA methods allow no input from the user, other than the sequence of interest, and thus are the easiest to use. For the other methods described here, the user can interactively participate in the prediction process. 4. If sequence homologs and, in particular, three-dimensional data are available, the PHD method gives reliable secondary structure predictions.
5. The PSA method was found to be less accurate than the other predictive schemes tested.
ACCESSIBILITY OF SOFTWARE Several prediction schemes have been incorporated into commercial packages (e.g., GCG) that also contain algorithms for additional sequence analysis. Other packages have interactive graphics modules that combine sequence analysis with molecular displays (Tripos and Biosym); however, these packages do not easily incorporate additional user input. There have been numerous reports in the literature describing computerized implementation of various
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2.3.17 Current Protocols in Protein Science
Figure 2.3.10 Schematic representation of the three-dimensional structure of human profilin I (Metzler et al., 1995) as displayed by Molscript (Kraulis, 1991). A seven-stranded antiparallel β sheet bisects the molecule. The N- and C-terminal α helices are on the left-hand side, and the two interior α helices are on the right-hand side in the view shown.
prediction schemes (Fasman, 1989b). Several of these programs have been published in a variety of programming languages. There are also several sites available on the Internet for sequence analysis. A listing of some of the commercial and Internet programs is contained in Table 2.3.6.
Literature Cited Bajorath, J., Stenkamp, R., and Aruffo, A. 1993. Knowledge-based model building of proteins: Concepts and examples. Protein Sci. 2:17981810. Baldwin, J.M. 1993. The probable arrangement of helixes in G protein–coupled receptors. EMBO J. 12:1693-1703. Blundell, T.L. and Johnson, L.N. (eds.) 1976. Protein Crystallography. Academic Press, London.
Protein Secondary Structure Prediction
acid residues and prediction of backbone topography in proteins. Isr. J. Chem. 12:239-286. Chou, P.Y. 1989. Prediction of protein structural class from amino acid compositions. In Prediction of Protein Structure and the Principles of Protein Conformation (G.D. Fasman, ed.) pp. 549-586. Plenum, New York. Chou, P.Y. and Fasman, G.D. 1974a. Conformational parameters for amino acids in helical, beta sheet and random coil regions calculated from proteins. Biochemistry 13:211-222. Chou, P.Y. and Fasman, G.D. 1974b. Prediction of protein conformation. Biochemistry 13:223-245. Chou, P.Y. and Fasman, G.D. 1977. β-turns in proteins. J. Mol. Biol. 115:135-175. Chou, P.Y. and Fasman, G.D. 1978a. Empirical predictions of protein conformation. Annu. Rev. Biochem. 47:251-276.
Branden, C. and Tooze, J. 1991. Introduction to Protein Structure. Garland Publishing, New York.
Chou, P.Y. and Fasman, G.D. 1978b. Prediction of the secondary structure of proteins from their amino acid sequence. Adv. Enzymol. 47:45-148.
Brookhaven. 1994. Protein Data Bank. Brookhaven National Laboratory, Upton, N.Y.
DeGrado, W.F. 1988. Design of peptides and proteins. Adv. Protein Chem. 39:51-124.
Burgess, A.W., Ponnuswamy, P.K., and Scheraga, H.A. 1971. Analysis of conformations of amino
Dill, K.A. 1990. Dominant forces in protein folding. Biochemistry 29:7133-7155.
2.3.18 Current Protocols in Protein Science
Dunhill, P. 1968. The use of helical net diagrams to represent protein structures. Biophys. J. 8:865875.
Jones, D.T., Taylor, W.R., and Thornton, J.M. 1992. A new approach to protein fold recognition. Nature 358:86-89.
Fasman, G.D. 1985. A critique of the utility of the prediction of protein secondary structure. J. Biosci. 8:15-23.
Kabsch, W. and Sander, C. 1984. On the use of sequence homologies to predict different conformations. Proc. Natl. Acad. Sci. U.S.A. 81:10751078.
Fasman, G.D. 1989a. The development of the prediction of protein structure. In Prediction of Protein Structure and the Principles of Protein Conformation (G.D. Fasman, ed.) pp. 193-316. Plenum, New York. Fasman, G.D. (ed.) 1989b. Prediction of Protein Structure and the Principles of Protein Conformation. Plenum, New York. Fermi, G. 1975. Three-dimensional Fourier synthesis of human deoxyhaemoglobin at 2.5 angstroms resolution, refinement of the atomic model. J. Mol. Biol. 97:237-256. Fersht, A.R. and Serrone, L. 1993. Principles of protein stability derived from protein engineering experiments. Curr. Opin. Struct. Biol. 3:7583. Finzel, B.C., Clancy, L.L., Holland, D.R., Muchmore, S.W., Watenpaugh, K.D., and Einspahr, H.M. 1989. Crystal structure of recombinant human interleukin-1β at 2.0 angstroms resolution. J. Mol. Biol. 209:779.
Kabsch, W. and Sander, C. 1985. Identical pentapeptides with different backbones. Nature 317:207. Kamtekar, S., Schiffer, J.M., Xiong, H., Babik, J.M., and Hecht, M.H. 1993. Protein design by binary patterning of polar and non-polar amino acids. Science 262:1680-1685. Kauzmann, W. 1959. Some factors in the interpretation of protein denaturation. Adv. Protein Chem. 14:1-63. Kneller, D.G., Cohen, F.E., and Largridge, R. 1990. Improvements in protein secondary structure prediction by an enhanced neural network. J. Mol. Biol. 214:171-182. Kraulis, P.J. 1991. MOLSCRIPT: A program to produce detailed and schematic plots of protein structures. J. Appl. Cryst. 24:946-950. Krystek, Jr., S.R., Bruccoleri, R.E., and Novotny, J. 1991. Stabilities of leucine zipper dimers estimated by an empirical force energy method. Int. J. Pept. Protein Res. 38:229-236.
Fisher, M.F. 1964. A limiting law relating the size and shape of protein molecules to their composition. Proc. Natl. Acad. Sci. U.S.A. 51:1285-1291.
Kuntz, I.D. 1972. Protein folding. J. Am. Chem. Soc. 94:4009-4012.
Garnier, J. and Robson, B. 1989. The GOR method for predicting secondary structure in proteins. In Prediction of Protein Structure and the Principles of Protein Conformation (G.D. Fasman, ed.) pp. 417-466. Plenum, New York.
Levitt, M. and Chothia, C. 1976. Structural patterns in globular proteins. Nature 261:552-558.
Garnier, J., Osguthorpe, D.J., and Robson, B. 1978. Analysis of the accuracy and implications of simple methods for predicting the secondary structure of globular proteins. J. Mol. Biol. 120:97-120. Genetics Computer Group. 1994. (GCG) Wisconsin Sequence Analysis Package Program Manual. Genetics Computer Group, Inc., Madison, Wis. Goodman, E.M. and Kim, P.S. 1991. Periodicity of amide proton exchange rates in a coiled-coil leucine zipper peptide. Biochemistry 30:1161511620. Greer, J. 1990. Comparative modeling: Application to the family of the mammalian serine proteases. Proteins Struct. Funct. Genet. 7:317-334. Guzzo, A.V. 1965. The influence of amino acid sequence on protein structure. Biophys. J. 5:809822. Harbury, P.B., Zhang, T., Kim, P.S., and Alber, T. 1993. A switch between two-, three-, and fourstranded coiled coils in GCN4 leucine zipper mutants. Science 262:1401-1407. Harper, E.T. and Rose, G.D. 1993. Helix stop signals in proteins and peptides: The capping box. Biochemistry 32:7605-7609.
Lesk, A.M. 1991. Protein architecture. IRL Press, Oxford.
Lewis, P.N., Momany, F.A., and Scheraga, H.A. 1971. Folding of polypeptide chains in proteins: A proposed method of folding. Proc. Natl. Acad. Sci. U.S.A. 68:2293-2297. Lim, V.I. 1974. Algorithms for prediction of α helices and β-structural regions in globular proteins. J. Mol. Biol. 88:873-894. Lovejoy, B., Choe, S., Cascio, D., McRorie, D.K., DeGrado, W.F., and Eisenberg, D. 1993. Crystal structure of a synthetic triple-stranded α-helical bundle. Science 259:1288-1293. Metzler, W.J., Constantine, K., Friedrichs, M.S., Bell, A., Ernst, E., Lavoie, T.B., and Mueller, L. 1993. Characterization of the three-dimensional structure of human profilin: 1H, 13C and 15N assignments and global folding pattern. Biochemistry 32:13818-13829. Metzler, W.J., Farmer, B.T., II, and Mueller, L. 1995. Refined solution structure of human profilin I. Protein Sci. 4:450-459. Nagano, K. 1973. Logical analysis of the mechanism of protein folding. I. Prediction of helices, loops and β-structures from primary structure. J. Mol. Biol. 75:401-420.
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Novotny, J. and Auffray, C. 1984. A program for prediction of protein secondary structure from nucleotide sequence data: Application to histocompatability antigens. Nucl. Acids Res. 12:243255. Oppenheimer, N.J. and James, T.L. (eds.) 1989. Nuclear magnetic resonance (Part B). Methods Enzymol. Volume 177.
Roy, S. and Rose, G.D. 1980. Hydrophobic basis of packing in globular proteins. Proc. Natl. Acad. Sci. U.S.A. 8:4643-4647. Sander, C. and Schneider, R. 1991. Database of homology-derived structures and the structural meaning of sequence alignment. Proteins Struct. Funct. Genet. 9:56-68.
O’Shea, E.K., Rutkowski, R., and Kim, P.S. 1992. Mechanism of specificity in the Fos-Jun oncoprotein heterodimer. Cell 68:699-708.
Schiffer, M. and Edmundson, A.B. 1967. The use of helical wheels to represent the structures of proteins and to identify segments with helical potential. Biophys. J. 7:121.
Pauling, L. and Corey, R.B. 1951. The pleated sheet, a new layer of configuration of polypeptide chains. Proc. Natl. Acad. Sci. U.S.A. 37:251-256.
Schulz, G.E. 1988. A critical evaluation of methods for prediction of protein secondary structures. Annu. Rev. Biophys. Chem. 17:1-21.
Pauling, L., Corey, R.B., and Branson, H.R. 1951. The structure of proteins: Two hydrogen-bonded helical configurations of the polypeptide chain. Proc. Natl. Acad. Sci. U.S.A. 37:205-211.
Schulz, G.E., Barry, C.D., Friedman, J., Chou, P.Y., Fasman, G.D., Finkelstein, A.V., Lim, V.I., Ptitsyn, O.B., Kabat, E.A., Wu, T.T., Levitt, M., Robson, B., and Nagano, K. 1974. Comparison of predicted and experimentally determined secondary structure of adenyl kinase. Nature 250:140-142.
Presta, L.G. and Rose, G.D. 1988. Helix signal in proteins. Science 240:1632-1641. Prevelige, P., Jr. and Fasman, G.D. 1989. Chou-Fasman prediction of the secondary structure of proteins. In Prediction of Protein Structure and the Principles of Protein Conformation (G.D. Fasman, ed.) pp. 391-416. Plenum, New York. Prothero, J.W. 1966. Correlation between the distribution of amino acids in alpha helices. Biophys. J. 6:367-370. Ptitsyn, O.B. and Finkelstein, A.V. 1970. Biofizika 15:757-768. Quinn, N. and Sejnowski, T.J. 1988. Predicting the secondary structure of globular proteins using neural network models. J. Mol. Biol. 202:865884. Richardson, J.S. 1981. The anatomy and taxonomy of protein structure. Adv. Protein Chem. 34:167339. Richardson, J.S. and Richardson, D.C. 1988. Amino acid preferences for specific locations at the ends of α-helices. Science 240:1648-1652. Robson, B. and Pain, R.H. 1971. Analyses of the code relating sequence to conformation in proteins: Possible implication for the mechanism of formation of helical regions. J. Mol. Biol. 58:237-259. Rost, B. and Sander, C. 1993a. Improved prediction of protein secondary structure by use of sequence profiles and neural networks. Proc. Natl. Acad. Sci. U.S.A. 90:7558-7562. Rost, B. and Sander, C. 1993b. Prediction of protein structure at better than 70% accuracy. J. Mol. Biol. 232:584-599.
Schulz, G.E. and Schirmer, R.H. 1979. Principles of Protein Structure. Springer-Verlag, New York and Heidelberg. Sippl, M.J. and Weitckus, S. 1992. Detection of native-like models for amino acid sequences of unknown three-dimensional structure in a data base of known protein conformations. Proteins Struct. Funct. Genet. 13:258-271. Stultz, C.M., White, J.V., and Smith, T.F. 1993. Structural analysis based on state-space modeling. Protein Sci. 2:305-314. Wuthrich, K. 1986. NMR of Proteins and Nucleic Acids. John Wiley & Sons, New York. Wyckoff, H.W., Hirs, C.H.W., and Timasheff, S.N. (eds.) 1985. Diffraction methods for biological molecules (Parts A and B). Methods Enzymol. Volumes 114 and 115.
Key Reference Fasman, 1989b. See above. Provides extensive descriptions and a historical perspective describing methods for protein structure prediction.
Contributed by Stanley R. Krystek, Jr., William J. Metzler, and Jiri Novotny Bristol-Myers Squibb Pharmaceutical Research Institute Princeton, New Jersey
Protein Secondary Structure Prediction
2.3.20 Current Protocols in Protein Science
Internet Basics
UNIT 2.4
With the explosion of sequence and structural information available to researchers, the field of bioinformatics is playing an increasingly large role in the study of fundamental biomedical problems. The challenge facing computational biologists will be to aid in gene discovery and in the design of molecular modeling, site-directed mutagenesis, and experiments of other types that can potentially reveal previously unknown relationships with respect to the structure and function of genes and proteins. This challenge becomes particularly daunting in light of the vast amount of data that has been produced by the Human Genome Project and other systematic sequencing efforts to date. Before embarking on any practical discussion of computational methods in solving biological problems, it is necessary to lay the common groundwork that will enable users to both access and implement the algorithms and tools discussed in this book. We begin with a review of the Internet and its terminology, also discussing major classes of Internet protocols, without becoming overly engaged in the engineering minutiae underlying these protocols. A more in-depth treatment on the inner workings of these protocols may be found in a number of well-written reference books intended for the lay audience (Krol and Klopfenstein, 1996; Rankin, 1996; Kennedy, 1999). This unit will also discuss matters of connectivity, ranging from simple modem connections to digital subscriber lines (DSL). Finally, we will address one of the most common problems that has arisen with the proliferation of Web pages throughout the world—i.e., finding useful information on the World Wide Web. INTERNET BASICS Despite the impression that the Internet is a single entity, it is actually a network of networks, composed of interconnected local and regional networks in more than 100 countries. While work on remote communications began in the early 1960s, the true origins of the Internet lie with a research project on networking at the Advanced Research Projects Agency of the U.S. Department of Defense in 1969 named ARPANET. The original ARPANET connected four nodes on the West Coast, with the immediate goal of being able to transmit information on defense-related research between laboratories. A number of different network projects subsequently surfaced, with the next landmark developments coming over ten years later. In 1981, BITNET (“Because It’s Time”) was introduced, providing point-to-point connections between universities for the transfer of electronic mail and files. In 1982, ARPA introduced the Transmission Control Protocol (TCP) and Internet Protocol (IP); TCP/IP allowed different networks to be connected to and communicate with one another, creating the system that is in place today. A number of references chronicle the development of the Internet and communications protocols in detail (Quarterman, 1990; Froehlich and Kent, 1991; Krol and Klopfenstein, 1996). Most users, however, are content to leave the details of how the Internet works to their systems administrators; the relevant fact to most is that it does work. Once the machines on a network are connected to one another, there needs to be some way to unambiguously specify a single computer so that messages and files find their intended recipient. To accomplish this, all machines directly connected to the Internet have an IP number. IP numbers are unique, identifying one and only one machine. The IP number is made up of four numbers separated by periods; for example, the IP number for the main file server at the National Center for Biotechnology Information (NCBI) at the National Institutes of Health (NIH) is 130.14.25.1. The numbers themselves represent, Contributed by Andreas D. Baxevanis and B.F. Francis Ouellette Current Protocols in Protein Science (2000) 2.4.1-2.4.16 Copyright © 2000 by John Wiley & Sons, Inc.
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from left to right, the domain (130.14 for the NIH), the subnet (.25 for the National Library of Medicine at NIH), and the machine itself (.1). While the use of IP numbers aids the computers in directing data, they are obviously very difficult for users to remember. Therefore, an IP number often has a fully-qualified domain name (FQDN) associated with it, which is dynamically translated in the background by a domain name server. Going back to the NCBI example, instead of using 130.14.25.1 to access the NCBI computer, a user could instead use ncbi.nlm.nih.gov and achieve the same result. Reading from left to right, the IP number goes from least to most specific, while the FQDN equivalent goes from most specific to least. The name of any given computer can then be thought of as taking the general form computer.domain, with the top-level domain (the portion coming after the last period in the FQDN) falling into one of the six broad categories shown in Table 2.4.1. Outside the United States, the top-level domain names may be replaced with a two-letter code specifying the country where the machine is located (e.g., .ca for Canada and .uk for the United Kingdom). In an effort to anticipate the needs of Internet users in the future, as well as to try to erase the arbitrary line between top-level domain names based on country, the now-dissolved International Ad Hoc Committee (IAHC) was charged with developing a new framework of generic top-level domains (gTLD). The new, recommended gTLDs were set forth in a document entitled The Generic Top Level Domain Memorandum of Understanding (gTLD-MOU); these gTLDs are overseen by a number of governing bodies and are also shown in Table 2.4.1. The most concrete measure of the size (and, thereby, the success) of the Internet lies in actually counting the number of machines physically connected to it. The Internet Table 2.4.1
Top-Level Domain Names
Top-level domain names Inside U.S. .com .edu .gov .mil .net .org
Commercial site Educational site Government site Military site Gateway or network host Private (usually not-for-profit) organizations
Examples of top-level domain names used outside the United States Canadian site .ca Academic site in the United Kingdom .ac.uk Commercial site in the United Kingdom .co.uk Generic top-level domains proposed by IAHC Firms or businesses .firm Businesses offering goods to purchase (stores) .shop Entities emphasizing activities relating to the World Wide Web .web Cultural and entertainment organizations .arts Recreational organizations .rec Information sources .info Personal names (e.g., yourlastname.nom) .nom
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Software Consortium conducts an Internet Domain Survey twice each year to count these machines, otherwise known as hosts. In performing this survey, ISC considers not only how many hostnames have been assigned, but how many of those are actually in use; a hostname might be issued, but the requestor may be holding the name in abeyance for future use. To test for this, a representative sample of host machines are sent a probe (a “ping”), with a signal being sent back to the originating machine if the host was indeed found. The rate of growth of the number of hosts has been phenomenal; from a paltry 213 hosts in August 1981, the Internet has in excess of 60 million hosts now. The doubling time for the number of hosts is on the order of 18 months. Most of this growth has come from the commercial sector, capitalizing on the growing popularity of multimedia platforms for advertising and communications such as the World Wide Web. CONNECTING TO THE INTERNET Of course, before being able to use all of the resources that the Internet has to offer, one needs to actually make a physical connection between one’s own computer and “the information superhighway.” For purposes of this discussion, the elements of this connection have been separated into two discrete parts: the actual, physical connection (meaning the “wire” running from one’s computer to the Internet backbone) and the service provider, who handles issues of routing and content once connected. Keep in mind that, in practice, these are not necessarily treated as two separate parts—for instance, one’s service provider may also be the same company that will run cables or fibers right into one’s home or office. Copper Wires, Coaxial Cables, and Fiber Optics Traditionally, users who were attempting to connect to the Internet away from the office have had one and only one option—a modem, which uses the existing copper twisted-pair cables carrying telephone signals to transmit data. Data transfer rates using modems are relatively slow, allowing for data transmission in the range of 28.8 to 56 kilobits per second (Kbps). The problem with using conventional copper wire to transmit data lies not in the copper wire itself, but in the switches that are found along the way that route information to their intended destinations. These switches were designed for the efficient and effective transfer of voice data, but were never intended to handle the high-speed transmission of data. While most people still use modems from their homes, a number of new technologies are already in place and will become more and more prevalent for accessing the Internet away from hard-wired Ethernet networks. The maximum speeds at which each of the services that are discussed below can operate are shown in Figure 2.4.1. The first of these “new solutions” is the integrated services digital network, or ISDN. While the advent of ISDN was originally heralded as the way to bring the Internet into the home in a speed-efficient manner, it required that special wiring be brought into the home. It also required that users be within a fixed distance from a central office, on the order of 20,000 feet or less. The cost of running this special, dedicated wiring, along with a per-minute pricing structure, effectively placed ISDN out of reach of most individuals. While ISDN is still available in many areas, this type of service is quickly being supplanted by more cost-effective alternatives. In looking at alternatives that did not require new wiring, cable television providers began to look at ways in which the coaxial cable already running into a substantial number of households could be used to also transmit data. Cable companies are able to use bandwidth that is not being used to transmit television signals (effectively, unused channels) to push Computational Analysis
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Telephone modem 0.056 Cellular wireless 0.128 ISDN 0.128 0.4 Satellite T1 1.544 Cable modem ADSL Ethernet 0
2
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4 6 Maximum speed (Mbps)
10
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10
Figure 2.4.1 Performance of various types of Internet connections, by maximum throughput. The numbers indicated in the graph refer to peak performance; often the actual performance of any given method may be on the order of one-half slower, depending on configurations and system conditions.
data into the home at very high speeds, up to 4.0 megabits per second (Mbps). The actual computer is connected to this network through a cable modem, which uses an Ethernet connection to the computer and a coaxial cable to the wall. Homes in a given area all share a single cable, in a wiring scheme very similar to that by which individual computers are connected via the Ethernet in an office or laboratory setting. While this branching arrangement can serve to connect a large number of locations, there is one major disadvantage—as more and more homes connect through their cable modems, service effectively slows down as more signal attempts to pass through any given node. One way of circumventing this problem is the installation of more switching equipment and reducing the size of a given “neighborhood.” Since the local telephone companies were the primary ISDN providers, they quickly turned their attention to ways in which the existing, conventional copper wire already in the home could be used to transmit data at high speed. The solution here is the digital subscriber line, or DSL. By using new, dedicated switches that are designed for rapid data transfer, DSL providers can circumvent the old voice switches that slowed down transfer speeds. Depending on the user’s distance from the central office and whether a particular neighborhood has been wired for DSL service, speeds are on the order of 0.8 to 7.1 Mbps. The data transfers do not interfere with voice signals, and users can use the telephone while connected to the Internet; the signals are “split” by a special modem that passes the data signals to the computer and a microfilter that passes voice signals to the handset. There is a special type of DSL called asynchronous DSL, or ADSL. This is the variety of DSL service that is becoming more and more prevalent. Most home users download much more information than they send out, so systems are engineered to provide super-fast transmission in the “in” direction, with transmissions in the “out” direction being 5 to 10 times slower. Using this approach maximizes the amount of bandwidth that can be used without necessitating new wiring. One of the advantages of ADSL over cable is that ADSL subscribers effectively have a direct line to the central office, meaning that they do not have to compete with their neighbors for bandwidth. This, of course, comes at a price; at the time of this writing, ADSL connectivity options were on the order of twice as expensive as cable Internet. Internet Basics
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Some of the newer technologies involve wireless connections to the Internet. These include using one’s own cell phone or a special cell phone service (such as Ricochet) to upload and download information. These cellular providers can provide speeds on the order of 28.8 to 128 Kbps, depending on the density of cellular towers in the service area. Fixed-point wireless services can be substantially faster, since the cellular phone does not have to “find” the closest tower at any given time. Along these same lines, satellite providers are also coming on-line. These providers allow for data download directly to a satellite dish with a southern exposure, with uploads occuring through traditional telephone lines. While the satellite option has the potential to be amongst the fastest of the options discussed, current operating speeds are only on the order of 400 Kbps. Content Providers versus ISPs Once an appropriately fast and price-effective connectivity solution is found, users will then need to actually connect to some sort of service that will enable them to traverse the Internet space. The two major categories in this respect are on-line services or Internet service providers (ISPs). On-line services such as America Online (AOL) and CompuServe offer a large number of interactive digital services, including information retrieval, electronic mail (e-mail), bulletin boards, and “chat rooms” where users who are on line at the same time can converse with each other about any number of subjects. While the on-line services now provide access to the World Wide Web (see discussion of The World Wide Web), most of the features and services available through these systems reside within a proprietary, closed network; once a connection is made between the user’s computer and the on-line service, accessing the special features, or content, of these systems does not require ever leaving the on-line system’s host computer. Specialized content can range from access to on-line travel reservation systems to encyclopedias that are constantly being updated—items that would not be available to anyone unless they subscribed to that particular on-line service. Internet service providers, or ISPs, take the opposite tack. Instead of focusing on providing content, the ISPs provide the tools necessary for users to send and receive e-mail, upload and download files, and navigate around the World Wide Web to find information at remote locations. The major advantage of ISPs is connection speed; ISPs often provide faster connections than the on-line services. Most ISPs charge a monthly fee for unlimited use. The line between on-line services and ISPs has already begun to blur. AOL’s now monthly flat fee pricing structure allows users to obtain all of the proprietary content found on AOL as well as all of the Internet tools available through ISPs, often at the same cost as a simple ISP connection. The extensive AOL network puts access to AOL as close as a local phone call in most of the United States, providing access to e-mail no matter where the user is located—a feature that small, local ISPs cannot match. Not to be outdone, many of the major national ISP providers now also provide content through the concept of portals. Portals are Web pages that can be customized to the needs of the individual user and which serve as a jumping-off point to other sources of news or entertainment on the Net. In addition, many national firms such as Mindspring are able to match AOL’s ease of connectivity on the road, and both ISPs and online providers are becoming more and more generous in providing users the capacity to publish their own Web pages. Developments such as this, coupled with the move of local telephone and cable companies into providing Internet access through new, faster fiber optic networks, foretell major changes in how people will access the Net in the future, changes that should favor the end-user both in price and performance.
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ELECTRONIC MAIL Most people are introduced to the Internet through the use of electronic mail (e-mail). The use of e-mail has become practically indispensable in many settings owing to its convenience as a medium for sending, receiving, and replying to messages. Its advantages are many: It is much quicker than postal, or “snail mail.” Messages tend to be much clearer and more to the point than is the case in typical telephone or face-to-face conversations. Recipients have more flexibility in deciding whether a response needs to be sent immediately, relatively soon, or at all, giving individuals more control over workflow. It provides a convenient method by which messages can be filed or stored. There is little or no cost involved in sending an e-mail message. While these and other advantages have pushed e-mail to the forefront of interpersonal communication in both industry and the academic community, users should be aware of several major disadvantages. First is the issue of security. As mail travels towards its recipient, it may pass through a number of remote nodes. The message could be intercepted and read at any one of those nodes by someone with high-level access, such as a systems administrator. Second is the issue of privacy. In industrial settings, e-mail is often considered to be an asset of the company for use only in official communication and, as such, is subject to monitoring by supervisors. The opposite is often true in academic, quasi-academic, or research settings; for example, National Institutes of Health policy encourages personal use of e-mail within the bounds of certain published guidelines. The key words here are “published guidelines”; no matter what the setting, users of e-mail systems should always be informed as to their organization’s policy regarding the appropriate use and confidentiality of e-mail so that they may use the tool properly and effectively. An excellent, basic guide to the effective use of e-mail is highly recommended (Rankin, 1996). Sending E-Mail E-mail addresses take the general form
[email protected], where user is the name of the individual user and computer.domain specifies the actual computer that the e-mail account is located on. Like a postal letter, an e-mail message is comprised of an envelope or header, showing the e-mail addresses of the sender and recipient, a line indicating the subject of the e-mail, and information about how the e-mail message actually travelled from the sender to the recipient. The header is followed by the actual message, or body, analogous to what would go inside the postal envelope. Figure 2.4.2 illustrates all the components of an e-mail message. E-mail programs vary widely, depending on both the platform and the needs of the users. Usually the characteristics of the local area network (LAN) dictate what types of mail programs can be used, and the decision is often left to systems administrators rather than individual users. Among the most widely used e-mail packages with a graphical user interface are Eudora for the Macintosh and both Netscape Messengerand Microsoft Exchange for Macintosh, Windows, and UNIX platforms. Text-based e-mail programs, which are accessed by logging into a UNIX-based account, include Elm and Pine.
Internet Basics
Bulk E-Mail As with postal mail, there has been an upsurge in “spam” or “junk e-mail,” where companies compile bulk lists of e-mail addresses for use in commercial promotions. Since most of these lists are compiled from on-line registration forms and similar sources, the
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Header Body
From:
[email protected] (PredictProtein) To:
[email protected] Subject: PredictProtein
Delivery details (Envelope)
Received: from dodo.cpmc.columbia.edu (dodo.cpmc.columbia.edu [156.111.190.78]) by members.aol.com (8.9.3/8.9.3) with ESMTP id RAA13177 for <
[email protected]>; Sun, 2 Jan 2000 17:55:22 -0500 (EST) Received: (from phd@localhost) by dodo.cpmc.columbia.edu (980427.SGI.8.8.8/980728.SGI.AUTOCF) id RAA90300 for
[email protected]; Sun, 2 Jan 2000 17:51:20 -0500 (EST) Date: Sun, 2 Jan 2000 17:51:20 -0500 (EST) Message-ID: <
[email protected]>
Sender, Recipient, and Subject
PredictProtein Help PHDsec, PHDacc, PHDhtm, PHDtopology, TOPITS, MaxHom, EvalSec Burkhard Rost Table of Contents for PP help 1. Introduction 1. What is it? 2. How does it work? 3. How to use it?
Figure 2.4.2 Anatomy of an e-mail message, with relevant components indicated. This message is an automated reply to a request for help file for the PredictProtein E-mail server.
best defense for remaining off of these bulk e-mail lists is to be selective as to whom e-mail addresses are provided. Most newsgroups keep their mailing lists confidential; if in doubt and this is a concern, one should ask. E-mail Servers Most often, e-mail is thought of as a way to simply send messages, whether it be to one recipient or many. It is also possible to use e-mail as a mechanism for making biological predictions or retrieving records from biological databases. Users can send e-mail messages in a predefined format, defining the action to be performed for remote computers known as servers; the servers will then perform the desired operation and e-mail back the results. While this method is not interactive (in that the user cannot adjust parameters or have control over the execution of the method in real time), it does place the responsibility of hardware maintenance and software upgrades on the persons maintaining the server, allowing users to concentrate on their results instead of on programming. For most servers, sending the message help to the server e-mail address will return a detailed set of instructions for using that server, including the way in which queries need to be formatted. Aliases and Newsgroups In the example in Fig. 2.4.2, the e-mail message is being sent to a single recipient. One of the strengths of e-mail is that a single piece of e-mail can be sent to a large number of people. The primary mechanism for doing this is through aliases; a user can define a group of people within their mail program and give the group a special name, or alias. Instead of using the individual e-mail addresses for all of the people in the group, the user can just send the e-mail to the alias name, and the mail program will handle broadcasting the message to each person in that group. Setting up alias names is a tremendous time-saver even for small groups; it also ensures that all members of a given group actually receive all e-mail messages intended for the group.
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The second mechanism for broadcasting messages is through newsgroups. This model works slightly differently in that the list of e-mail addresses is compiled and maintained on a remote computer through subscriptions, much like magazine subscriptions. For example, the BIOSCI newsgroups are amongst the most highly-trafficked, offering a forum for discussion or the exchange of ideas in a wide variety of biological subject areas. To begin receiving the messages posted to the automated sequencing discussion group within BIOSCI, a user would send a message to [email protected] with the wording subscribe autoseq in the body of the message. The user would then receive all future postings to that group and be able to participate in the discussions. If a user wished to be removed from the group, a message would be sent to the same address, but this time, the body of the message would read unsubscribe autoseq. For more information on BIOSCI, including a complete list of discussion groups, an e-mail message can be sent to [email protected]; in this case the subject line should be left blank and the words info faq typed in the body of the message. The BIOSCI server will then return a copy of the Frequently Asked Questions (FAQ) in response, with detailed information on each newsgroup overseen by BIOSCI. It is also possible to participate in newsgroups without having each and every piece of e-mail flood into one’s private mailbox. Instead, interested participants can use newsreading software, such as NewsWatcher for the Macintosh, which provides access to the individual messages making up a discussion. The major advantage is that the user can pick and choose which messages to read by scanning the subject lines; the remainder can be discarded by a single operation. NewsWatcher is an example of what is known as a client-server application—the client software (here, NewsWatcher) runs on a client computer (a Macintosh), which in turn interacts with a machine at a remote location (the server). Client-server architecture is interactive in nature, with a direct connection being made between the client and server machines. Once NewsWatcher is started, the user is presented with a list of newsgroups available to them (Fig. 2.4.3); this list will vary, depending on the user’s location, as systems administrators have the discretion to allow or block certain groups at a given site. From the rear-most window in the figure, the user double-clicks on the newsgroup of interest (here, bionet.genome.arabidopsis), which spawns the window shown in the center. At the top of the center window is the current unread message count, and any message within the list can be read by double-clicking on that particular line. This, in turn, spawns the last window (in the foreground), showing the actual message. If a user decides not to read any of the messages, or is done reading individual messages, the balance of the messages within the newsgroup (center) window can be deleted by first choosing Select All from the File menu, then selecting Mark Read from the News menu. Once the newsgroup window is closed, the unread message count is reset to zero. Every time NewsWatcher is restarted, it will automatically poll the news server for new messages that have been created since the last session. As with most of the tools that will be discussed in this unit, news-reading capability is built into Web browsers such as Netscape Navigator and Microsoft Internet Explorer. FILE TRANSFER PROTOCOL
Internet Basics
Despite the many advantages afforded by e-mail in transmitting messages, experienced e-mail users have no doubt experienced frustration in trying to transmit files (attachments) along with an e-mail message. The mere fact that a file can be attached to an e-mail message and sent does not mean that the recipient will be able to detach, decode, and actually use the attached file. While more cross-platform e-mail packages such as
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Figure 2.4.3 Using NewsWatcher to read postings to newsgroups. The list of newsgroups that the user has subscribed to is shown in the Subscribed List window (left). The list of new postings for the highlighted newsgroup (bionet.genome.arabidopsis) is shown in the center window. The window in the foreground shows the contents of the posting selected from the center window.
Microsoft Exchange are being developed, the use of different e-mail packages by people at different locations means that sending files via e-mail is not an effective, foolproof method, at least in the short term. One solution to this problem is through the use of a file transfer protocol (or FTP). The workings of FTP are quite simple—a connection is made between a user’s computer (the client) and a remote server, and that connection remains in place for the duration of the FTP session. File transfers are very fast, at rates on the order of 5 to 10 kilobytes per second, with speeds varying with time of day, distance between the client and server machines, and overall traffic on the network. In the ordinary case, making an FTP connection and transferring files requires that a user have an account on the remote server. However, there are many files and programs that the academic community makes freely available, and access to those files does not require having an account on each and every machine where these programs are stored. Instead, connections are made using a system called anonymous FTP. Under this system, the user connects to the remote machine, and instead of entering a username/password pair, types anonymous as the username and enters an e-mail address in place of a password. Providing one’s e-mail address allows the server’s systems administrator to compile
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$ ftp ftp.bio.indiana.edu Connected to magpie.bio.indiana.edu. 220 iubio.bio.indiana.edu FTP server ready. Name: anonymous 331 Guest login ok, send your complete e-mail address as password. Password: ******** 230-
Welcome to IUBio archive!
230230-
This is a user-supported archive for biology software and data.
230230-
See the file Archive.Doc for details of this archive.
230230-
See IUBio Bio-Mirror archive of large data sets at
230230-
ftp to iubio.bio.indiana.edu, user: iubio, password: iubio This includes GenBank, EMBL and DDBJ and other biosequence data.
230230-
Report problems, uploads and other matters via e-mail to
230-
[email protected].
230230 Guest login ok, access restrictions apply. Remote system type is UNIX. Using binary mode to transfer files. ftp> cd /molbio/align/clustal 250 CWD command successful. ftp> get clustalw1.75.unix.tar.Z local: clustalw1.75.unix.tar.Z remote: clustalw1.75.unix.tar.Z 200 PORT command successful. 150 Opening BINARY mode data connection for clustalw1.75.unix.tar.Z (230379 bytes). 226 Transfer complete. 230379 bytes received in 0.45 seconds (500.75 Kbytes/s) ftp> quit 221-You have transferred 230379 bytes in 1 files. 221-Total traffic for this session was 231859 bytes in 1 transfers. 221-Thank you for using the FTP service on iubio.bio.indiana.edu. 221 Goodbye.
Figure 2.4.4 Using UNIX FTP to download a file. An anonymous FTP session is established with the molecular biology FTP server at the University of Indiana to download the ClustalW alignment program. The user inputs are shown in boldface.
access statistics which may, in turn, be of use to those actually providing the public files or programs. An example of an anonymous FTP session using UNIX is shown in Figure 2.4.4. Although FTP occurs within the UNIX environment, Macintosh and PC users can employ programs that utilize graphical user interfaces (GUI, pronounced “gooey”) to navigate through the UNIX directories on the FTP server. Users need not have any knowledge of UNIX commands to download files; they can instead rely on pop-up menus and the ability Internet Basics
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Figure 2.4.5 Using Fetch to download a file. An anonymous FTP session is established with the molecular biology FTP server at the University of Indiana (top) to download the ClustalW alignment program (bottom). Notice the difference between this GUI-based program and the UNIX equivalent illustrated in Figure 2.4.4.
to point-and-click their way through the UNIX file structure. The most popular FTP program on the Macintosh platform for FTP sessions is Fetch. A sample Fetch window is shown in Figure 2.4.5 to illustrate the difference between using a GUI-based FTP program and the equivalent UNIX FTP in Figure 2.4.4. In the figure, notice that the Automatic radio button (near the bottom of the second window under the Get File button) is selected, meaning that Fetch will determine the appropriate type of file transfer to perform. This may be manually overridden by selecting either Text or Binary, depending on the nature of the file being transferred. As a rule, text files should be transferred as Text, programs or executables as Binary, and graphic format files such as PICT and TIFF files as Raw Data. Computational Analysis
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THE WORLD WIDE WEB While FTP is of tremendous use in the transfer of files from one computer to another, it does suffer from some limitations. When working with FTP, once a user enters a particular directory, they can only see the names of the directories or files. In order to actually view what is within the files, it is necessary to physically download them onto one’s own computer. This inherent drawback led to the development of a number of distributed document delivery systems (DDDS), interactive client-server applications that allowed information to be viewed without having to perform a download. The first generation of DDDS development led to programs like Gopher, which allowed plain text to be viewed directly through a client-server application. From this evolved the most widely known and widely used DDDS, namely the World Wide Web. The Web is an outgrowth of research performed at the European Nuclear Research Council (CERN) in 1989 that was aimed at sharing research data between several locations. That work led to a medium through which text, images, sounds, and videos could be delivered to users on demand, anywhere in the world. Navigation on the World Wide Web Navigation on the Web does not require advance knowledge of the location of the information being sought. Instead, users can navigate by clicking on specific text, buttons, or pictures. These clickable items are collectively known as hyperlinks. Once one of these hyperlinks is clicked, the user is taken to another Web location, which could be at the same site or halfway around the world. Each document displayed on the Web is called a Web page, and all of the related Web pages on a particular server are collectively called a Web site. Navigation strictly through the use of hyperlinks has been nicknamed “Web surfing.” Users can take a more direct approach to finding information by entering a specific address. One of the strengths of the Web is that the programs used to view Web pages (appropriately termed browsers) can be used to visit Gopher and FTP sites as well, somewhat obviating the need for separate Gopher or FTP applications. As such, a unified naming convention was introduced to indicate to the browser program both the location of the remote site and, more importantly, the type of information at that remote location so that the browser could properly display the data. This standard-form address is known as a uniform resource locator (URL), and takes the general form protocol://computer. domain, where protocol specifies the type of site and computer.domain specifies the location (Table 2.4.2). The http used for the protocol in World Wide Web URLs stands for hypertext transfer protocol, the method used in transferring Web files from the host computer to the client.
Table 2.4.2 Uniform Resource Locator (URL) Format for Each Type of Transfer Protocol
Site
URL format (example)
General form FTP site
protocol://computer.domain ftp://ftp.ncbi.nlm.nih.gov
Gopher site Web site
gopher://gopher.iubio.indiana.edu http://www.nhgri.nih.gov
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Browsers Browsers, which are used to look at Web pages, are client-server applications that connect to a remote site, download the requested information at that site, display the information on the user’s monitor, then disconnect from the remote host. The information retrieved from the remote host is in a platform-independent format called hypertext markup language (HTML). HTML code is strictly text-based, and any associated graphics or sound for that document exist as separate files in a common format. For example, images may be stored and transferred in GIF format, developed by CompuServe for the quick and efficient transfer of graphics; while GIF format is most commonly used for graphics, other formats such as JPEG and BMP may also be used. Because of this, a browser can display any Web page on any type of computer, whether it be a Macintosh, IBM-compatible, Linux, or UNIX machine. The text is usually displayed first, then the remaining elements are placed on the page as they are downloaded. With minor exceptions, a given Web page will look the same when the same browser is used on any of the above platforms. The two major players in the area of browser software are Netscape, with their Communicator product, and Microsoft, with Internet Explorer. As with many other areas where multiple software products are available, the choice between Netscape and Internet Explorer comes down to one of personal preference. While the computer literati will debate the fine points of difference between these two packages, for the average user, both packages perform equally well and offer the same types of features, adequately addressing the Web-browser needs of most users. It is worth mentioning that, while the Web is by definition a visually based medium, it is also possible to travel through Web space and view documents without the associated graphics. For users limited to line-by-line terminals, a browser called Lynx is available. Developed at the University of Kansas, Lynx allows users to use their keyboard arrow keys to highlight and select hyperlinks, using their return key the same way that Netscape and Internet Explorer users would click their mouse. Internet versus Intranet The World Wide Web is normally thought of as a way to communicate with people at a distance, but the same infrastructure can be used to connect people within an organization. Such intranets provide an easily accessible repository of relevant information, capitalizing on the simplicity of the Web interface. It also provides another channel for broadcast or confidential communication within the organization. Having an intranet is of particular value when members of an organization are physically separated, whether it be in different buildings or different cities. Intranets are protected in such a way that people who are not on the organization’s network are prohibited from accessing the internal Web pages; additional protections through the use of passwords are also common. Finding Information on the World Wide Web Most people find information on the Web the old fashioned way—by word of mouth either by using lists such as Table 2.4.3 or by simply following hyperlinks put in place by Web authors. Continuously clicking from page to page can be a highly ineffective way of finding information, especially when the information sought is of a very focused nature. One way of finding interesting and relevant Web sites is to consult virtual libraries, which are curated lists of Web resources arranged by subject. Virtual libraries of special interest to biologists include the WWW Virtual Library, maintained by Keith Robison at Harvard and the EBI BioCatalog, based at the European Bioinformatics Institutes. The URLs for these sites can be found in Table 2.4.3.
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Table 2.4.3
World Wide Web Sites of Interest
Site
URL
Domain names gTLD-MOU Internet Software Consortium
http://www.gtld-mou.org http://www.isc.org
Electronic mail and newsgroups BIOSCI Newsgroups http://www.bio.net/docs/biosci.FAQ.html Eudora http://www.eudora.com Microsoft Exchange http://www.microsoft.com/exchange/ NewsWatcher ftp://ftp.acns.nwu.edu/pub/newswatcher/ File Transfer Protocol Fetch 3.0/Mac FTP Voyager
http://www.dartmouth.edu/pages/softdev/fetch.html http://ftpvoyager.deerfield.com
Internet access America Online AT&T Bell Atlantic Bell Canada CompuServe MCI Ricochet Telus
http://www.aol.com http://www.att.com/worldnet http://www.bellatlantic.net http://www.bell.ca http://www.compuserve.com http://www.mci.com http://www.ricochet.net http://telus.com
Virtual libraries EBI BioCatalog Amos’ WWW Links Page NAR Database Collection WWW Virtual Library
http://www.ebi.ac.uk/biocat/biocat.html http://www.expasy.ch/alinks.html http://www3.oup.co.uk/nar/Volume_28/Issue_01/introduction/ http://mcb.harvard.edu/BioLinks.html
World Wide Web browsers Internet Explorer http://www.microsoft.com/insider/ie5/default.htm Lynx ftp://ftp2.cc.ukans.edu/pub/lynx Netscape Navigator http://home.netscape.com World Wide Web search engines AltaVista http://www.altavista.com Excite http://www.excite.com HotBot http://hotbot.lycos.com Infoseek http://infoseek.go.com Lycos http://www.lycos.com Northern Light http://www.northernlight.com World Wide Web meta-search engines MetaCrawler http://www.metacrawler.com Savvy Search http://www.savvysearch.com
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Table 2.4.4 Number of Hits Returned for Four Defined Search Queries on Some of the More Popular Search and Meta-Search Engines
Search term genetic mapping human genome positional cloning prostate cancer
Search engine
Meta-search engine
HotBot
Excite
Infoseek
Lycos
478 13,231 279 14,044
1,040 34,760 735 53,940
4,326 15,980 1,143 24,376
9,395 19,536 666 33,538
MetaCrawler SavvySearch 62 42 40 60
58 54 52 57
It is also possible to directly search the Web by using search engines such as Alta Vista and Excite, among others. A search engine is simply a specialized program that can perform full-text or keyword searches on databases that catalog Web content. The result of a search is a hyperlinked list of Web sites fitting the search criteria from which the user can visit any or all of the found sites. However, search engines use slightly different methods in compiling their databases. One variation is the attempt to capture most or all of the text of every Web page that the search engine is able to find and catalog (“Web crawling”). Another technique is to catalog only the title of each Web page rather than its entire text. A third is to consider words that must appear next to each other or only relatively close to one another. Because of these differences in search-engine algorithms, the results returned by issuing the same query to a number of different search engines can produce wildly different results (Table 2.4.4). It may also be noted from Table 2.4.4 that most of the numbers are exceedingly large, reflecting the overall size of the World Wide Web. Unless a particular search engine ranks its results by relevance (for example, by scoring words in a title higher than words in the body of the Web page), the results obtained may not be particularly useful. It is also necessary to keep in mind that, depending on the indexing scheme that the search engine is using, the found pages may actually no longer exist, leading the user to the dreaded “404 Not Found” error. Compounding this problem is the issue of coverage—the number of Web pages that any given search engine is actually able to survey and analyze. A comprehensive study by Lawrence and Giles (1998) indicates that the coverage provided by any of the search engines studied is both small and highly variable. For example, the HotBot engine produced 57.5% coverage of what was estimated to be the size of the “indexable Web,” while Lycos had only 4.41% coverage, a full order of magnitude less than HotBot. The most important conclusion from this study was the extent of coverage increased as the number of search engines are increased and the results from those individual searches are combined. Combining the results obtained from the six search engines examined in this study produced coverage approaching 100%. To address this point, a new class of search engines called meta-search engines have been developed. These programs will take the user’s query and poll anywhere from five to ten of the “traditional” search engines. The meta-search engine will then collect the results, filter out duplicates, and return a single, annotated list to the user. One big advantage is that the meta-search engines take relevance statistics into account, returning much smaller lists of results. Even though the hit list is substantially smaller, it is much more likely to contain sites that directly address the original query. Since the programs must poll a number of different search engines, searches conducted this way obviously take longer to perform, but the higher degree of confidence in the compiled results for a given query outweighs the extra few minutes (sometimes only seconds) of search time. Reliable and easy-to-use meta-search engines include SavvySearch and MetaCrawler (see Table 2.4.3).
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LITERATURE CITED Froehlich, F. and Kent, A. 1991. ARPANET, the Defense Data Network, and Internet. In Encyclopedia of Communications. Marcel Dekker, New York. Kennedy, A.J. 1999. The Internet: Rough Guide 2000. Rough Guides, London. Krol, E. and Klopfenstein, B.C. 1996. The Whole Internet User’s Guide and Catalog. O’Reilly and Associates, Sebastopol, Calif. Lawrence, S. and Giles, C.L. 1998. Searching the World Wide Web. Science 280:98-100. Rankin, B. 1996. Dr. Bob’s Painless Guide to the Internet and Amazing Things You Can Do with E-mail. No Starch Press, San Francisco. Quarterman, J. 1990. The Matrix: Computer Networks and Conferencing Systems Worldwide. Digital Press, Bedford, Mass.
Contributed by Andreas D. Baxevanis National Human Genome Research Institute, NIH Bethesda, Maryland B.F. Francis Ouellette Centre for Molecular Medicine and Therapeutics University of British Columbia Vancouver, British Columbia
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Sequence Similarity Searching Using the BLAST Family of Programs
UNIT 2.5
Database sequence similarity searching is carried out thousands of times each day by researchers worldwide. Scientists in traditional laboratories use search results, for example, to infer the functions of newly discovered cDNAs, predict new members of gene families, and explore evolutionary relationships between sequences. In turn, they populate sequence databases with biochemically and/or genetically characterized sequences. With the advent of whole-genome sequencing, a new breed of scientist is now using these characterized sequences to predict the location and function of coding and regulatory regions in large segments of genomic DNA. Their contribution to the sequence databases is the submission, at a rapid pace, of generally uncharacterized mRNA and genomic DNA sequences. These data can then be used by the first group of scientists to enhance the understanding of their sequences. Both types of sequencing efforts have greatly increased the size and quality of the sequence databases in recent years. Thus, sequence similarity searching has become a valuable research tool for all molecular biologists (Altschul et al., 1994; Schuler, 1998; also see UNIT 2.1). Over the years, a number of algorithms have been implemented that allow searching of sequence databases. The most useful of these tools should share the following characteristics: (1) Speed. Because today’s databases are so large, the programs must be fast in order to process megabases of sequence in seconds. (2) Sensitivity. The programs must report all potentially interesting similarities. (3) Rigorous statistics. The programs must provide a way to evaluate the significance of the results. (4) Ease of use. Scientists with no formal training in sequence-analysis algorithms should understand how to use the programs and interpret the results. Advanced users should have the option to tailor the programs to their needs. (5) Access to up-to-date databases. The doubling time of GenBank is currently ∼16 months. It is important to search the most recent version of the database. The BLAST (Basic Local Alignment Search Tool) family of sequence similarity search programs satisfies the above criteria. In short, users input either a nucleotide or amino acid query sequence, and search a nucleotide or amino acid sequence database. The program returns a list of the sequence “hits,” alignments to the query sequence, and statistical values. This unit describes how to choose an appropriate BLAST program and database, perform the search, and interpret the results. ACCESSING BLAST PROGRAMS AND DOCUMENTATION The National Center for Biotechnology Information (NCBI) currently supports two versions of BLAST free of charge: BLAST 2.0 (gapped BLAST) and Position-Specific Iterated BLAST (PSI-BLAST). BLAST 2.0 is the standard version of BLAST, which allows a user to search a sequence database with a nucleotide or protein sequence of interest. BLAST 2.0 places gaps into the query and target sequences so that separate areas of similarity between the two sequences can be returned as one hit. PSI-BLAST is an iterative BLAST search, which is optimized for finding distantly related sequences. Other BLAST programs are also available from the NCBI Web page. “BLAST 2 sequences” uses the BLAST search engine to produce an alignment of two sequences entered by the user. On the Specialized BLAST pages, researchers can use the BLAST engine to search sequences that are not in GenBank. At present, these databases include unfinished microbial genomes, P. falciparum (the human malaria parasite), and tentative Contributed by Tyra G. Wolfsberg and Thomas L. Madden Current Protocols in Protein Science (1999) 2.5.1-2.5.29 Copyright © 1999 by John Wiley & Sons, Inc.
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human consensus (THC) sequences from The Institute for Genomic Research (TIGR). The content of these pages is, however, subject to change. The easiest and most popular way to access the BLAST suite of programs is through the NCBI World Wide Web site, at http://www.ncbi.nlm.nih.gov/BLAST/. All versions of BLAST are accessible from this site, and can be used to query all sequence databases available at the NCBI. Documentation, which includes an overview of BLAST, BLAST frequently asked questions (FAQs), a “What’s New” page, the BLAST manual, and a list of references, is also available here. For users who want to run BLAST against private local databases or downloaded copies of NCBI databases, the NCBI offers a stand-alone version of the BLAST program. BLAST binaries and documentation are provided for the latest versions of IRIX, Solaris, DEC OSF1, and Win32 systems. BLAST 2.0 executables may be found on the NCBI anonymous FTP server at ftp://ncbi.nlm.nih.gov/blast/executables/. BLAST can also be run as a client-server program, in which the user installs client software on a local machine that communicates across the network with a server at NCBI. This setup is useful for researchers who run large numbers of searches on NCBI databases, because they can automate the process to run on their local computer. The BLAST client may be found on the NCBI anonymous FTP server at ftp://ncbi.nlm.nih.gov/blast/network/netblast. The NCBI BLAST e-mail server is the best option for people without convenient access to the Web. A similarity search can be performed by sending a properly formatted e-mail message containing the nucleotide or protein query sequence to [email protected]. The query sequence is compared against a specified database and the results are returned in an e-mail message. For more information on formulating e-mail BLAST searches, please send a message consisting of the word HELP to the same address, [email protected]. This unit concentrates on the BLAST searches which can be performed from the NCBI Web site. The other implementations are mainly for advanced users of BLAST, or those with special needs. For any of these services, questions should be directed to [email protected]. INTRODUCTION TO BLAST Basic Versus Advanced BLAST Searches BLAST 2.0 is available in a basic or advanced version. In both versions, the user can select the type of BLAST program and the database to be searched, and choose whether to filter the query sequence to mask low-complexity regions (see below). The advanced version allows the user to change parameters as well. For most researchers, the Basic version, which uses the default parameters, is adequate. For a discussion of BLAST parameters, see Appendix A at the end of this unit. BLAST Programs Five types of the BLAST program have been developed to support sequence similarity searching using a variety of nucleotide and protein sequence queries and databases. These programs are listed and described in Table 2.5.1. The type of BLAST search to be carried out is dependent on the type of information that is desired. Sequence Similarity Searching Using BLAST
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Table 2.5.1
BLAST Search Programs
Program
Query sequence Database sequence Comments
BLASTP
Protein
Protein
BLASTN
Nucleotide (both strands)
Nucleotide
BLASTX
Nucleotide (six-frame translation)
Protein
TBLASTN Protein
Nucleotide (six-frame translation)
TBLASTX Nucleotide (six- Nucleotide (sixframe translation) frame translation)
Can be run in standard mode or in a more sensitive iterative mode (PSI-BLAST), which uses the previous search results to build a profile for subsequent rounds of similarity searching. Parameters optimized for speed, not sensitivity; not intended for finding distantly related coding sequences. Automatically checks complementary strand of query. Very useful for preliminary data containing potential frameshift errors (ESTs, HTGs, and other “single-pass” sequences). Essential for searching protein queries against EST database. Often useful for finding undocumented open reading frames or frameshift errors in database sequences. Should be used only if BLASTN and BLASTX produce no results. Restricted for search against EST, STS, HTGS, GSS, and Alu databases.
NCBI Databases One frequent mistake in sequence similarity searching is failure to search an up-to-date database. The NCBI produces GenBank (Benson et al., 1998; ftp://ncbi.nlm.nih.gov/genbank/gbrel.txt) and updates it daily. It also shares data on a daily basis with the DNA Data Bank of Japan (DDBJ; Tateno et al., 1998) and the European Molecular Biology Laboratory (EMBL; Stoesser et al., 1998). A search of the NCBI databases using the BLAST Web page, client, or e-mail server guarantees access to the most recent database. The NCBI supports a number of databases for sequence similarity searching. These databases are subject to change, and a current list and description are available on the NCBI Web site at http://www.ncbi.nlm.nih.gov/BLAST/blast_databases.html. This section describes some of the commonly used NCBI databases. Examples demonstrating the utility of various BLAST programs and NCBI databases are given in the next section. Peptide sequence databases for BLASTP and BLASTX nr. The nr (nonredundant) database is the most comprehensive. GenBank, DDBJ, and EMBL are nucleotide sequence databases. If any nucleotide sequence in any of the databases is annotated with a coding sequence (CDS), this CDS appears in nr. nr also contains protein sequences obtained from PDB (sequences associated with 3-dimensional structures in the Brookhaven Protein Data Bank), Swiss-Prot (a curated database of protein sequences; Bairoch and Apweiler, 1998), PIR (Protein Identification Resource, a comprehensive collection of protein sequences; Barker et al., 1998), and PRF (Protein Research Foundation). Although nr may contain multiple copies of similar sequences, identical sequences are merged into one entry. To be merged, two sequences must have identical lengths and every residue at every position must be the same. There are nr databases for both peptide and nucleotide sequences. The peptide database is automatically selected for BLASTP and BLASTX searches.
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month. The month database receives its sequences from the same sources as nr, but contains only those sequences released within the last 30 days. All sequences in month are also present in nr. Alu. The Alu database contains six-frame translations of representative Alu repeats from all Alu subfamilies (Claverie and Makalowski, 1994). If a query sequence containing an Alu repeat is used in a BLAST search of nr or month, many of the resulting high-scoring hits will also contain Alu sequences. It may be useful, especially with a genomic sequence query, to perform a search of the Alu database to identify the location of any Alu repeats that might produce high-scoring and potentially misleading hits in queries of other databases. Nucleotide sequence databases for BLASTN, TBLASTN, and TBLASTX nr. The nr (nonredundant) database contains all nucleotide sequences present in GenBank, EMBL, and DDBJ. It also contains nucleotide sequences obtained from PDB (sequences associated with 3-dimensional structures in the Brookhaven Protein Data Bank). nr comprises only sequences that are normally well annotated, so it does not contain expressed sequence tag (EST), sequence-tagged site (STS), genome survey sequence (GSS), or high-throughput genomic (HTG) sequences. Although nr may contain multiple copies of similar sequences, identical sequences are merged into one entry. To be merged, two sequences must have identical lengths and every nucleotide at every position must be the same. month. The month database contains all nucleotide sequences present in GenBank, EMBL, DDBJ, and PDB that were released within the last 30 days. Unlike nr, it also contains EST, STS, GSS, and HTG sequences released within the last month. EST. EST accesses a nonredundant copy of all ESTs present in GenBank, EMBL, and DDBJ (Boguski et al., 1993). ESTs are short sequences, a few hundred nucleotides in length, which are derived by partial, single-pass sequencing of inserts of randomly selected cDNA clones (Adams et al., 1991). Since the number of ESTs is increasing rapidly, it is an important database to search for novel cDNAs. As of August, 1998, ∼70% of the sequences in GenBank were ESTs; of these, 61% were from human, 20% from mouse. STS. STS contains a nonredundant copy of all STSs present in GenBank, EMBL, and DDBJ. An STS is a short unique genomic sequence that is used as a sequence landmark for genomic mapping efforts (Olson et al., 1989). As of August, 1998, 83% of the sequences in the STS database were from human.
Sequence Similarity Searching Using BLAST
HTGS. HTGS contains “unfinished” DNA sequences generated by the high-throughput sequencing centers (Ouellette and Boguski, 1997). A typical HTG record might consist of all the first-pass sequence data generated from a single cosmid, BAC, YAC, or P1 clone. The record is composed of two or more sequence fragments that have a total length of ≥2 kb and contain one or more gaps. The sequences are normally updated by the sequencing centers as more data become available. A single accession number is assigned to this collection of sequences. The accession number does not change as the record is updated, and only the most recent version of the record remains in GenBank. Phase 1 HTG sequences are unordered, unoriented contigs with gaps. Phase 2 HTG sequences are ordered, oriented contigs with or without gaps. All HTG records contain a prominent warning that the sequence data is unfinished and may contain errors. When a record is considered finished, it becomes a Phase 3 HTG and is moved to the nr database with the same accession number. HTGS is a valuable source of new genomic sequences not yet in nr.
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GSS. GSS includes short, single-pass genomic data identified by various means (Smith et al., 1994). Many of the sequences have been mapped. As of August, 1998, 80% of the sequences in GSS were from human, 14% from Arabidopsis thaliana. Alu.The Alu database contains representative Alu repeats from all Alu subfamilies (Claverie and Makalowski, 1994). If a query sequence containing an Alu repeat is used in a BLAST search of the above nucleotide databases, many of the resulting high-scoring hits will also contain Alu sequences. It may be useful, especially with a genomic sequence query, to perform a search of the Alu database to identify the location of any Alu repeats that might produce high-scoring and potentially misleading hits in queries of other databases. Vector. The Vector database contains nucleotide sequences of a number of standard cloning vectors. New sequences should be screened against the Vector database to assure that they do not contain any vector contamination. Mito. The Mito database contains representative mitochondrial sequences from many families. Nuclear-derived sequences may be screened against the Mito database to assure that they do not contain any mitochondrial contamination. Formatting the Query Sequence Users of the BLAST Web page can initiate a search either by entering the sequence itself, or, if the sequence is already in the sequence database, by entering the accession number or gi (see Appendix B at the end of this unit). The preferred format for entering new sequences is the so-called FASTA format; however, if the sequence is not in FASTA format or the sequence is interspersed with numbers and spaces, BLAST will still accept the query. A sequence in FASTA format begins with a single-line description, followed by lines of sequence data. The description line is distinguished from the sequence data by a > symbol (greater than) in the first column. An example sequence in FASTA format is: >aaseq Human choroideremia protein MADNLPTEFDVVIIGTGLPESILAAACSRSGQRVLHIDSRSYYGGNWASFSFSGLLSWLKEYQQNNDIGE ESTVVWQDLIHETEEAITLRKKDETIQHTEAFPYASQDMEDNVEEIGALQKNPSLGVSNTFTEVLDSALP EESQLSYFNSDEMPAKHTQKSDTEISLEVTDVEESVEKEKYCGDKTCMHTVSDKDGDKDESKSTVEDKAD EPIRNRITYSQIVKEGRRFNIDLVSKLLYSQGLLIDLLIKSDVSRYVEFKNVTRILAFREGKVEQVPCSR ADVFNSKELTMVEKRMLMKFLTFCLEYEQHPDEYQAFRQCSFSEYLKTKKLTPNLQHFVLHSIAMTSESS CTTIDGLNATKNFLQCLGRFGNTPFLFPLYGQGEIPQGFCRMCAVFGGIYCLRHKVQCFVVDKESGRCKA IIDHFGQRINAKYFIVEDSYLSEETCSNVQYKQISRAVLITDQSILKTDLDQQTSILIVPPAEPGACAVR VTELCSSTMTCMKDTYLVHLTCSSSKTAREDLESVVKKLFTPYTETEINEEELTKPRLLWALYFNMRDSS GISRSSYNGLPSNVYVCSGPDCGLGNEHAVKQAETLFQEIFPTEEFCPPPPNPEDIIFDGDDKQPEAPGT NNVVMAKLESSEESKNLESPEKHLQN
Alternatively, the user can enter the database accession number or gi. The above sequence is in fact already in the database with a gi identifier of 116365 and a Swiss-Prot accession number of P26374. Either of these sequence identifiers can be entered on the BLAST page. The type of identifier (for nucleotide or protein sequence) must match the type of query sequence used in that search (Ostell and Kans, 1998; Ouellette, 1998). Sequences in manuscripts are often referred to by their GenBank accession number; however, this accession number refers to a nucleotide (not protein) sequence, and cannot be used to initiate a BLASTP or TBLASTN search, both of which require a protein sequence query. The identifier of the protein sequence encoded by a nucleotide accession number can be obtained by searching the NCBI’s Entrez nucleotide database, at http://www.ncbi.nlm. nih.gov/Entrez. Computational Analysis
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Filtering Sequences Both nucleotide and protein sequences may contain regions of low complexity, i.e., regions with homopolymeric tracts, short-period repeats, or segments enriched in one or only a few residues. Such low-complexity regions commonly give spuriously high BLAST scores that reflect compositional bias rather than significant position-by-position alignment (Altschul et al., 1994). For example, two protein sequences that contain low-complexity regions rich in the same amino acids may produce high-scoring alignments in those regions even though other parts of the proteins are entirely dissimilar. Because these alignments do not reflect a common ancestry, no functional inference is justified despite their high statistical significance. Filtering the query sequence (that is, replacing the repeated sequence with strings of n for nucleotide sequence or X for protein sequence) can eliminate potentially confounding matches, such as hits to low-complexity, proline-rich regions or poly(A) tails present in the database. By default, all searches performed through the NCBI BLAST Web page automatically filter the query sequence, as do the BLAST clients, e-mail server, and stand-alone programs. However, filtering can be turned off, even on the Basic BLAST page. Filtered sequence is represented in the final BLAST report as a string of n or X (e.g., nnnnnnnnnn or XXXXXXXXX). BLASTN queries are filtered with DUST (R.L. Tatusov and D.J. Lipman, pers. comm.). Other BLAST queries use SEG (Wootton and Federhen, 1993, 1996). Viewing the BLAST Results The NCBI BLAST Web page can return results in one of three ways. The default is to display the results in the browser window from which the user initiated the search. The document will have hypertext links that make it easier to analyze the results. BLAST queries are processed in the order in which they are received, except that less computationally intensive jobs (e.g., BLASTN, BLASTP, and smaller databases such as month) are given priority. In the middle of the afternoon, the BLAST server may be busy. Thus, it is sometimes more efficient to receive the BLAST results by e-mail. E-mail results are sent either as plain text or in HTML format. The HTML-formatted results must be opened in a Web browser, and the resulting document contains hypertext links. EXAMPLES OF BLAST SEARCHES In this section, interpretation of BLAST results is explained using examples for BLASTP, BLASTX, TBLASTN, BLASTN, and PSI-BLAST. An example is not shown for TBLASTX, as it is a tool of last resort and not useful for the majority of users. The first example, for BLASTP, contains general information, and should be read by all BLAST users. GenBank increases in size by thousands of sequences every week; thus, results obtained from running the same searches again will differ from those shown in the examples. BLASTP The BLASTP program compares a protein query to a protein database. A typical search is shown in Figure 2.5.1. The Swiss-Prot database has been selected and the program has been changed to BLASTP. A query in FASTA format has been entered in the input box. The query is the human choroideremia protein, implicated in hereditary blindness (Seabra et al., 1993). Sequence Similarity Searching Using BLAST
The top of the results page for this search is shown in Figure 2.5.2. It begins with some header information about the type of program (BLASTP), the version (2.0.5), and a release
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Figure 2.5.1 Submitting a BLASTP search using the NCBI’s World Wide Web interface.
Figure 2.5.2 Example of the top portion of a BLASTP report.
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Figure 2.5.3 Example of the graphical view of a BLASTP report. The bars are color coded by the strength of the database match. The strongest matches (those with a bit score >200) are red, followed by pink (bit score 80 to 200), green (50 to 80), blue (40 to 50), and black (<40).
date. The version and release date will change as the program is updated. Also listed are a reference, the query definition line, and a summary of the database used. Figure 2.5.3 presents a graphical overview of the results. The database hits are shown aligned to the query, which is the numbered bar at the top. The next three bars show high-scoring database matches that align to the query sequence throughout its length. The next nine bars show lower-scoring matches that align to two regions of the query—one from residue 3 or 4 of the query up to about residue 60, and the other from about residues 220 to 500. For these nine bars the solid segments (on the left and right) indicate aligned regions; the cross-hatched segment indicates that the two alignments (i.e., the solid portions) are from the same database sequence, but that there is no alignment where the bar is cross-hatched. If there is no cross-hatching between two or more bars in the same row (i.e., the thirteenth row), this denotes different alignments with unrelated database sequences. These bars are presented on the same line to conserve space in the graphical view. Moving the cursor over a bar in the graphic (“mousing over”) causes the identifier and definition line of the database match to be shown in the window. If the results of a BLAST search are sent by e-mail, this graphical view is not included.
Sequence Similarity Searching Using BLAST
The hit list produced by the BLASTP search is shown in Figure 2.5.4. Each line of the hit list is composed of four fields. The first field contains the database designation, accession number, and locus name for the matched sequence, separated by vertical bars (see Appendix B at the end of this unit for more information). The second field contains a brief textual description of the sequence, the definition line. The content of the definition line varies between and within databases, but usually includes information on the organism from which the sequence was derived, the type of sequence (e.g., mRNA or DNA), and some information about function or phenotype. The definition line is often truncated in the hit list to keep the display compact, but is fully displayed in association with the local alignments (see below). The third field contains the alignment score in bits. Higher scoring hits are found at the top of the list. In very general terms, this score is calculated from a formula that takes into account the alignment of similar or identical
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Figure 2.5.4 Example of the hit list from a BLASTP report.
residues, as well as any gaps that must be introduced in order to align the sequences. A key element in this calculation is the “substitution matrix,” which assigns a score for aligning any possible pair of residues. The BLOSUM-62 matrix is the default for BLAST, and it works well for most searches. The fourth field contains the Expect (E) value, which provides an estimate of statistical significance. E values reflect how many times one expects to see such a score occur by chance. A statistician would consider an E value <0.05 to be significant. However, an inspection of the alignments (see below) is required to determine biological significance. The maximum number of hits displayed in the hit list is set at a default value of 500. This number can be changed on the advanced BLAST page with the descriptions option. For the first hit in the list, the database designation is sp (for Swiss-Prot), the accession number is P26374, the locus name is RAE2_HUMAN, the definition line begins RAB PROTEINS, the score is 1223, and the E value is 0.0. The higher the score and the lower the E value, the more statistically significant is the hit. Note that the first thirteen hits all have very low E values (<10−14) and are either RAB proteins or GDP dissociation inhibitors. The next eighteen database matches have much higher E values (≥0.35), meaning that roughly one match with that score would be expected by chance. Furthermore, the definition lines no longer show the same consistency. Given the combination of inconsistent definition lines and high E values, caution should be used in assuming an evolutionary relationship between the query and the eighteen lower-scoring database hits. It would be necessary to examine the alignments in order to make a final determination. The pairwise alignments between the query and the database hits are displayed below the hit list. Figure 2.5.5 shows one sequence alignment between the query sequence and a
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Figure 2.5.5 Example of a BLASTP alignment.
Sequence Similarity Searching Using BLAST
database hit; other alignments are omitted for brevity. The alignment is preceded by the sequence identifier, the full definition line, and the length, in amino acids, of the database sequence. Next comes the score of the match in bits (the raw score is in parentheses), the Expect value (E value) of the hit, the number of identical residues in the alignment (identities), the number of conservative substitutions (positives) according to the scoring system (e.g., BLOSUM-62), and, if applicable, the number of gaps in the alignment. Finally the actual alignment is shown, with the query on top and the database match labeled as subject (Sbjct). Since both the query and the match are amino acid sequences, the numbers at left and right refer to the position in the amino acid sequence. The center line between the two sequences indicates the similarities between the sequences. If the residue is identical between the query and the subject, the residue itself is shown; conservative substitutions, as judged by the substitution matrix, are indicated with a + sign. One or more dashes (−) within a sequence indicates insertions or deletions. Query residues masked for low complexity are replaced by Xs (see fourth and eleventh blocks, Fig. 2.5.5). By default, a maximum of 500 pairwise alignments are displayed. This number can be changed on the advanced BLAST page with the alignments option.
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Results generated on the NCBI BLAST Web page contain text hyperlinks that assist the user in navigating around the page. Clicking on a colored bar in the graphical view (Fig. 2.5.3) moves the user to the sequence alignment (Fig. 2.5.5). The database sequence identifiers, visible both in the one-line descriptions of the database matches (Fig. 2.5.4) and in the sequence alignment (Fig. 2.5.5), link to the Entrez record that describes the sequence. Entrez may provide more information about the sequence, including links to relevant publication abstracts in PubMed. The score (to the right of the one-line description; Fig. 2.5.4) provides a link within the page to the alignment between that hit and the query. BLASTX BLASTX searches involve a six-frame translation of a nucleotide sequence queried against a protein database (Gish and States, 1993). Researchers most often use a BLASTX search if they have sequenced a DNA molecule and do not know the reading frame of the coding sequence, the beginning or end of the open reading frame, or the function of the encoded protein. It is also useful for analyzing preliminary sequence data containing potential sequencing or frameshift errors, such as ESTs, HTGs, or GSSs. A BLASTX search of the nr protein database can quickly highlight any similarities between all the open reading frames in a nucleotide sequence and any characterized proteins. In the example illustrated here, the mRNA sequence encoding human ataxia telangiectasia (ATM; accession number U33841; Lavin and Shiloh, 1997) was queried against the nr protein database. The output is similar to that from BLASTP. At the top (not shown) is information about the search, including the query, database, and type of BLAST program used. Next is the graphical overview representing the alignment of the database hits to the query sequence (Fig. 2.5.6), followed by a list of the hits, including their definitions, scores, and E values (Fig. 2.5.7 shows the top portion of the list). Pairwise alignments between the query and hit sequences follow (selected alignments are shown in Fig. 2.5.8). The top-scoring seven hits, which have scores >2600, are to other orthologs of the ATM gene (Fig. 2.5.7). Most of these hits align with the query sequence over its entire length (in Fig. 2.5.6, the first five bars span the entire length of the query sequence; note also the longer bars in the sixth and seventh lines). The 3′ end of the ATM mRNA (carboxy terminal end of the resulting protein translation), aligns with the catalytic domain of a number of phosphatidylinositol 3-kinases or similar proteins—such as the yeast TEL1 gene (a phosphatidylinositol kinase involved in controlling telomere length), the yeast ESR1 gene (a putative phosphatidylinositol kinase required for mitotic cell growth, DNA repair, and meiotic recombination), and the yeast TOR1 protein (a phosphatidylinositol kinase required for cell cycle progression). The alignment with these proteins starts in different places depending on the protein; one group starts around nucleotide 4700 of ATM (lines 9, 10, and 15 in Fig. 2.5.6), and other groups start around nucleotide 5800 (lines 17 to 19), 6800 (lines 7, 8, and 12 to 14), and 8000 (lines 20 to 60). Such similarities have provided insight into the role of ATM in response to DNA damage and cell cycle control (Lavin and Shiloh, 1997). The ATM gene also shares similarity with viral capsid proteins around nucleotides 2500 and 6000 (short bars on lines 6 to 14). However, these hits have low scores and high E values (not shown) and are not likely to be significant. The hits to the orthologs of ATM reveal the location of the coding sequence within the query nucleotide sequence. In a BLASTX output, position in the query sequence (the top line of a pairwise alignment) is described as a nucleotide position, while the position in the database hit (the bottom line of the alignment) is described as an amino acid position (Fig. 2.5.8). For example, amino acids 1 to 3066 of the mouse ortholog of ATM, gi 1469394, align with the translation of nucleotide positions 190 through 9357 of the query
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Figure 2.5.6 Example of the graphical view of a BLASTX report.
sequence. The exact location of the ATM coding sequence cannot be easily determined from the hits to the yeast sequences, as the similarity does not extend throughout the length of the protein (Fig. 2.5.8, bottom). However, the hits to the yeast sequences do provide confirmation about the phase of the reading frame, and possibly the location of specific domains conserved between humans and yeast. TBLASTN
Sequence Similarity Searching Using BLAST
TBLASTN searches involve a protein sequence queried against the six-frame translation of a nucleotide sequence database. TBLASTN searches are especially useful for finding novel similarities to a protein sequence in the open reading frames from uncharacterized, sometimes lower-quality nucleotide sequences, such as those found in EST, STS, HTGS, and GSS databases. The translations of these sequences are not present in nr because the sequences are not annotated. The most common use of TBLASTN is to search for ESTs that are similar to a protein of interest. These ESTs might represent orthologs of the protein from a different species, or just additional members of a gene family.
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Figure 2.5.7 Example of the hit list from a BLASTX report.
The following example shows a TBLASTN search of HTGS, a database that contains unfinished and uncharacterized genomic sequences generated by genome sequencing centers. Since HTGs are genomic and often contain introns, the results can be more difficult to interpret than those from an EST search. However, HTGs can also provide insight into the chromosomal organization of genes. In this case, the six-frame translation of HTGS is searched with the amino acid sequence of the disintegrin-like domain of mouse ADAM 1 (Wolfsberg et al., 1995). TBLASTN output is similar to that of BLASTP and BLASTX. Figure 2.5.9 shows the graphical overview of the alignment, Figure 2.5.10 shows the list of database hits, and Figure 2.5.11
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Figure 2.5.8 Example of selected BLASTX alignments.
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Figure 2.5.9 Example of the graphical view of a TBLASTN report.
Figure 2.5.10 Example of the hit list from a TBLASTN report.
shows selected pairwise alignments between the query sequence and the hits. For TBLASTN searches, the top line (Query) of the alignment contains protein sequence, and the location refers to amino acids. The bottom line (Sbjct) of the alignment contains a translated nucleotide sequence, and the location refers to the nucleotide location. Disintegrin-like domains contain a characteristic pattern of cysteines, so it is usually easy to identify the true hits by checking for conservation in the number of cysteine residues, and the spacing between them. The first hit, a 627-kb C. elegans clone, aligns with the
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Figure 2.5.11 Example of a TBLASTN alignment.
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Figure 2.5.12 Example of the graphical view of a BLASTN report.
query sequence in two regions. Both alignments encompass the full length of the query sequence, and both contain a characteristic alignment of cysteine residues. The first region (the first aligned block) comprises nucleotides 322651-322911 of the C. elegans clone. The second region consists of the second and third alignment blocks. The first 55 amino acids of the disintegrin domain align with nucleotides 102880-103107 of the clone, and the last 25 amino acids align with nucleotides 103155-103232 of the clone. Thus, this C. elegans clone appears to encode two distinct disintegrin-like domains, and these two sequences are likely linked in the C. elegans genome. The region between nucleotides 103107-103155 of the clone may contain an intron. The second database hit is to a shorter C. elegans clone that, from the sequence of its encoded disintegrin domain, appears to contain some of the same sequence as the first C. elegans clone. The third database hit is to a human clone. As amino acids 1 to 16 of the query aligns with nucleotides 53178 to 53225 of the hit, and amino acids 24 to 49 align with nucleotides 53474 to 53551, the human sequence may contain an intron between nucleotides 53225 and 53474. There is a likely second intron between nucleotides 53551 and 57012. BLASTN Figure 2.5.12 presents the graphical overview of a BLASTN search of accession number U93237 against the human EST database. U93237 is the 9.2-kb genomic sequence of the human menin (MEN1) gene for multiple endocrine neoplasia type I. This example demonstrates the power of a BLAST search to confirm exons in a genomic sequence, as well as the danger of accepting results without further investigation. The matches on the
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Figure 2.5.13 Example of the hit list from a BLASTN report.
right half of the diagram (base pairs 4000 to 9000) confirm the locations of exons 2 to 10 of the MEN1 gene (Chandrasekharappa et al., 1997; Zhang and Madden, 1997). The large number of matches on the left side of the figure (base pairs 500 to 1000 and 3000 to 3600) appear at first glance to be interesting matches, but are actually caused by the presence of Alu elements in the query and EST sequences. One can determine this by checking the definition lines of the matches in this region, as many of these ESTs are annotated as being similar to Alu elements. It is important to remember, though, that the definition line summaries (e.g., Fig. 2.5.13) contain truncated definition lines, and that it is necessary to look at the full definition line (provided in the alignment section, Fig. 2.5.14) to see the Alu annotation. One can also perform a BLASTN search of U93237 against the Alu database. Fig. 2.5.13 shows the top portion of the list of database hits for the BLASTN search of U93237 against human ESTs. Fig. 2.5.14 presents one alignment of the query sequence with an EST containing an Alu repeat. Vertical bars connect bases that are identical in the query and subject. Since both the query and the subject are nucleotide sequences, the numbers shown refer to nucleotide sequence locations. Nucleotides in the query sequence that have been masked for low complexity are replaced by the letter n (e.g., nnnnnnn in the third block). BLASTN checks the plus and minus strand of the query against the plus strand of the database sequence. The alignment shown here presents the plus strand of the query to the minus strand of the database sequence, which is equivalent to showing the minus strand of the query against the plus strand of the database sequence.
Sequence Similarity Searching Using BLAST
BLASTN is optimized for speed, not sensitivity. The algorithm uses a simple system to assess the alignments; matches are given a positive score, mismatches a negative score. BLASTN should only be used when a direct comparison of nucleotide sequences is desired. BLASTN results should not be used to make any predictions about the functions of the encoded proteins. Generally, searches that involve protein comparisons are more sensitive, owing to the larger alphabet (twenty residues versus four nucleotides) and the
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Figure 2.5.14 Example of a BLASTN alignment.
degeneracy of the genetic code (two codons, with different third bases often coding for the same protein). The more sophisticated scoring system used with protein comparisons also takes into account similarities between different residues. PSI-BLAST A major new feature of the BLAST version 2.0 is the iterative search. Position-specific iterated BLAST (PSI-BLAST) first performs a normal BLASTP run, but then internally computes a profile using the most significant hits (Altschul et al., 1997). A profile may be understood as a table that lists the probability of finding a residue at each position in a conserved protein domain. Conserved regions between the query and the most-significant matches are used to compute the profile. These regions are presumably important to the function of the protein. The profile is then used to perform another search (or iteration), at which point the new hits are also added into the profile. The cycle may then be repeated, with a new profile computed from the results of the last iteration. Each PSI-BLAST iteration takes into account the most significant sequences found in the previous round (i.e., those below the threshold for inclusion in the profile). The algorithm is more sensitive than BLASTP, and more likely to lead to the discovery of distantly related sequences. At present, such iterative searching is only available with the BLASTP program. The PSI-BLAST output is different from the other programs. Figure 2.5.15 shows the one-line summaries from the first PSI-BLAST iteration (i.e., the first search using the profile) of a mouse adenosine deaminase (Swiss-Prot accession P03958) against the nr database. The one-line summaries are divided into two groups, those with an E value better (lower) than that used to build the profile (Fig. 2.5.15; the default value is 0.001), and those with a worse E value (not shown). Matches better than the threshold, which will be included in the profile, are checked. The user may check (or uncheck) matches, causing them to be included (or excluded) from the profile used in the next iteration. Matches found on a previous iteration are marked with a green ball. New sequences are
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Figure 2.5.15 Example of the hit list from a PSI-BLAST report.
marked with a yellow “new”. The page also alerts the user when convergence is achieved (i.e., no new sequences below the threshold for inclusion in the profile were found in that iteration). In the hit list in Figure 2.5.15, the first iteration of PSI-BLAST finds many adenosine deaminase sequences (marked with a green ball) found in the previous (standard) BLASTP search. In addition, PSI-BLAST identifies a number of AMP deaminase sequences (marked with “new”). Indeed, there is a well-known homology between adenosine deaminase and AMP deaminase (Chang et al., 1991; Holm and Sander, 1997) that would not normally be found by BLASTP, but is found by PSI-BLAST. Sequence Similarity Searching Using BLAST
If one wants to report the E value for a hit found with a PSI-BLAST search, one should use the E value calculated during the first iteration that the match is below the threshold
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for inclusion in the profile (i.e., when the hit is marked with a “new” flag). However, if the hit is found in the first BLASTP run, then the reported E value should be from this run. In either case, in subsequent iterations, the hit itself is used to compute the profile, and the reported E value is not representative of the true value. SEARCHING STRATEGIES Filtering for Low-Complexity Regions Low-complexity regions tend to result in misleading database search results. The residue frequencies within such regions differ dramatically from the database as a whole, disrupting the statistics used by BLAST. This can produce alignments that are based purely on compositional bias rather than a significant position-by-position alignment. By default, low-complexity sequences are filtered out of query sequences and replaced by strings of n (nucleotide) or X (protein). A BLASTP search of a proline-rich, DNA-directed RNA polymerase (Swiss-Prot accession P11414) against the nr database illustrates the differences in output resulting from a filtered or unfiltered query sequence. Twenty four database matches with an E value better than 10 are found if filtering is enabled; 4873 are found if filtering is disabled. The top-scoring database matches for the unfiltered search are shown in Figure 2.5.16; high-scoring matches that are not found in a filtered search are marked. Note that the definition lines of most of the new matches indicate that the sequences are proline-rich, like the query, but do not provide information about the function of the protein. Other hits to RNA polymerases, found near the top of the list in a filtered search, are buried much deeper in the list of matches. The match to “sp|P14248,” apparently a good match based on the definition line, does not appear in a filtered BLAST search. An examination of the alignment (not shown) indicates that the match is based on solely on proline-rich regions. Reporting BLAST Results The most reliable indicator of the importance of a BLAST alignment is the Expect (E) value (the number of chance database matches one expects to see at the same score). This number takes into account the length of the query and size of the database as well as the scoring system. Neither the bit nor the raw score is a reliable indicator, as their significance cannot be judged independently of the information used to calculate the E value. Normal intuition fails when one is faced with databases of the current size (e.g., the EST database contains ∼700 million base pairs at this time), as chance alignments are likely and it is impossible for a user to know at what score matches become significant. Statistical significance also varies from database to database (e.g., a hit with a certain score may be statistically significant in a search of the relatively small month database, but not in a search of nr), so that reporting a certain score without the context of the database size can be misleading. The percentage identity is also a poor indicator of statistical significance for the same reasons; additionally, there is really no way to estimate an E value or even a score from a percentage identity, making it impossible to even guess whether a hit could be due to chance or not. Analyzing mRNA Sequences 1. Identify the open reading frame to obtain the protein sequence. Perform a BLASTX search of the cDNA sequence against the nr protein database. Any resulting hits may identify the correct reading frame for the protein sequence and provide information about its function. Computational Analysis
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Figure 2.5.16 Example of a hit list from a BLASTP report in which the query sequence was not filtered. Black squares, added manually by the authors, indicate hits that would not appear if the query had been filtered.
2. Determine whether the protein sequence is similar to that of any previously identified proteins. Perform a BLASTP and a PSI-BLAST (iterative) search against the nr protein database using the protein sequence as a query. 3. Determine whether the protein sequence is similar to the translation products of any uncharacterized DNA sequences whose translations are not in nr. Perform a TBLASTN search of the EST, STS, GSS, and HTGS databases. Sequence Similarity Searching Using BLAST
4. After the initial search, perform a monthly search of the month nucleotide and protein sequence databases to provide information about newly added sequences. A BLASTP
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search of the month protein database will reveal newly characterized protein sequences. A TBLASTN search of the month nucleotide database will reveal new open reading frames in nr, EST, HTGS, STS, and GSS. PSI-BLAST searches of nr should also be conducted occasionally, as new sequences may be added to the profile that will, in turn, help identify more distantly related sequences. 5. If the protein sequence contains multiple functional domains, it may be useful to perform the searches with each of these domains individually. Analyzing Genomic DNA Sequences 1. Identify a potential transcribed segment. Perform a BLASTN search of the nucleotide sequence against the DNA sequences in the EST database to determine if any mRNAs are derived from the genomic sequence. EST hits will highlight the location of exons. A BLASTN search against nr may identify transcripts or previously characterized genomic segments. 2. Identify potential open reading frames. Perform a BLASTX search against the nr protein database. Any resulting hits may identify the correct reading frame of a protein sequence and provide information about its function. If no results are found with the previous methods, a TBLASTX search (translated nucleotide query versus a translated nucleotide database) against EST, GSS, HTGS, and STS may identify other open reading frames. 3. After the initial search, a monthly search of the month nucleotide (with BLASTN) and protein (with BLASTX) sequence databases will provide information about newly added sequences. 4. Any potential transcribed sequences can be analyzed as per the above instructions for mRNA sequences. Searching Short Sequences Short sequences can only produce short alignments. Such short alignments need to be relatively strong (i.e., have a high percentage of matching residues) to rise above the background noise. Short, but strong, alignments are more easily detected using a matrix with a higher relative entropy than that of the default BLOSUM-62 (Altschul, 1991). Relative entropy is basically the average information available per position to distinguish the alignment from chance. The BLOSUM series of matrices does not include any that are suitable for the shortest queries, so it is recommended to use the PAM matrices instead (Dayhoff et al., 1978; Schwartz and Dayhoff, 1978). These matrices may be chosen from a menu on the Advanced BLAST page. Suggested matrices for different query lengths are shown in Table 2.5.2 (S.F. Altschul, pers. comm.). Table 2.5.2 Substitution Matrices for Short Sequences
Query length (amino acids)
Matrix
<35 35-50 50-85 >85
PAM-30 PAM-70 BLOSUM-80 BLOSUM-62 Computational Analysis
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SEQUENCE ALIGNMENT ALGORITHMS There are two types of pairwise sequence alignments, global and local. Global strategies attempt to align along the entire length of both sequences, while local alignments focus on sequence similarity over shorter regions. Historically, local alignments have been very successful at identifying similarities between proteins. Subsequences of similarity, surrounded by unrelated sequences, can identify, for example, active sites that are conserved between proteins or exons in genomic sequence. Smith and Waterman (1981) were the first to develop a local alignment algorithm. However, such rigorous alignment algorithms are often quite slow and impractical for searching current databases on a routine basis. Heuristic algorithms implement certain shortcuts to allow a comparison against an entire database within a reasonable time. They are not exhaustive searches in the sense that they do not explore possibilities that seem unlikely to be interesting. Some programs allow the user to adjust parameters that increase the sensitivity of the search (i.e., explore more possibilities), but this must be balanced against the slower speed of the search. Studies have shown that heuristic algorithms, with default settings, approach the sensitivity of exhaustive searches (Altschul et al., 1997). BLAST (Altschul et al., 1990, 1997) and FASTA (Pearson, 1990; Pearson and Lipman, 1988) are two popular heuristic programs. The rest of this discussion will focus on the newest version (2.0) of BLAST. BLAST calculates a statistical significance for every alignment it produces. Significance is expressed in terms of the E value, which is the number of matches with a given score (the calculation of the score is described below) that one would expect by chance. The E value depends on the size of the database and the length of the query, as increasing either of these parameters increases the number of chance matches. Calculation of the E value is performed with Karlin-Altschul statistics (Karlin and Altschul, 1990, 1993). The raw score of an alignment is calculated with a scoring system, which is a table of residue substitution scores and penalties for the existence and extension of a gap. For normal protein/protein comparisons, a matrix (i.e., table of residue substitution scores) provides a score for each possible alignment of two amino acids. For example, L aligned with I gets a positive score, while L aligned with E gets a negative score. BLOSUM-62 is the default matrix, and experimentation has shown that it is among the best for detecting weak protein similarities (Henikoff and Henikoff, 1992). PSI-BLAST uses a table that lists the frequency of finding a certain residue at each position in a conserved protein domain. This table is calculated “on the fly” for each iteration from the best matches to the database. BLASTN uses a simple scheme where matches have a positive score and mismatches have a negative score. Gaps are opened or extended when one sequence is longer than another over a certain region. BLAST charges the score −a for the existence of a gap and −b for each letter in a gap, so that a gap of length k letters is penalized −(a + bk). Gap costs of this form are known as affine gap costs. These gap costs may be changed by the user. The scoring system determines the raw score of an alignment. A bit score can be calculated from the raw score using the following formula: Sbit = [λ × Sraw − ln(K)]/ln(2)
Sequence Similarity Searching Using BLAST
where λ and K are Karlin-Altschul parameters that depend on the scoring system. The statistical significance corresponding to the resulting bit score is independent of the scoring system used. This allows one to compare the bit scores obtained from two different BLAST runs, each performed using different matrices or gap extension and existence values. Nevertheless, it is more meaningful to compare the results of different BLAST
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runs using their E values rather than their bit scores, as the E value takes into account the size of the database. A word-based approach is used to find matches in BLAST. An ungapped search, with BLASTN, may be used to demonstrate the concept of a word. First a list is made of all the subsequences in the nucleic acid query sequence that are of a certain length (called the word size), and the positions are recorded. The database is then scanned, looking for words that are in the list. If a match is found between a query and a database word, the alignment is extended (without allowing gaps) until the corresponding score drops a certain amount below the maximum found, at which point that maximum score (and the corresponding alignment) is used. The aligned region is a local alignment. With BLASTN, only exact matches are extended and the default word size is 11 bp. The procedure is more complicated for the other programs (BLASTP, BLASTX, TBLASTN, and TBLASTX). Exact word matches are not required and the default word size is three residues. A match between a query word and a database word is based on the scoring matrix, which specifies whether two residues result in a positive, negative, or neutral match. If three residues in a query word are compared with three residues in a database sequence, and the score is above a certain threshold, this word warrants further consideration. The default behavior in BLAST version 2.0 is that two words (on the query) must match with two words (on the database sequence) within a window of 40 residues before the match is extended, as described for BLASTN. The effect of looking for two word matches is to eliminate random matches, increasing the speed of BLAST by a factor of three. Ungapped alignments are used as a starting point for the gapped alignments that BLAST produces. The eleven highest-scoring letters are determined and the center letter is used as the start of the gapped alignment. The alignment is then extended, with gaps allowed. This extension is not exhaustive; if the score drops by more than a certain amount from the maximum found so far, that maximum (and the corresponding alignment) is used (Altschul et al., 1997; Zhang et al., 1998). The default BLAST parameters are tuned for a balance of speed and sensitivity. It is possible that significant matches may be missed, especially if small databases are searched. In that case, it may be advisable to use a shorter word size for BLASTN or a lower-threshold word for the other programs. It is important to remember that BLAST calculates similarity. Homology is an evolutionary relationship that must be proved by further analysis. LITERATURE CITED Adams, M.D., Kelley, J.M., Gocayne, J.D., Dubnick, M., Polymeropoulos, M.H., Xiao, H., Merril, C.R., Wu, A., Olde, B., Moreno, R.F., et al. 1991. Complementary DNA sequencing: Expressed sequence tags and human genome project. Science 252:1651-1656. Altschul, S.F. 1991. Amino acid substitution matrices from an information theoretic perspective. J. Mol. Biol. 219:555-565. Altschul, S.F., Gish, W., Miller, W., Myers, E.W., and Lipman, D.J. 1990. Basic local alignment search tool. J. Mol. Biol. 215:403-410. Altschul, S.F., Boguski, M.S., Gish, W., and Wootton, J.C. 1994. Issues in searching molecular sequence databases. Nature Genet. 6:119-129. Altschul, S.F., Madden, T.L., Schäffer, A.A., Zhang, J., Zhang, Z., Miller, W., and Lipman, D.J. 1997. Gapped BLAST and PSI-BLAST: A new generation of protein database search programs. Nucl. Acids Res. 25:3389-3402. Bairoch, A. and Apweiler, R. 1998. The SWISS-PROT protein sequence data bank and its supplement TrEMBL in 1998. Nucl. Acids Res. 26:38-42.
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Barker, W.C., Garavelli, J.S., Haft, D.H., Hunt, L.T., Marzec, C.R., Orcutt, B.C., Srinivasarao, G.Y., Yeh, L.S.L., Ledley, R.S., Mewes, H.W., Pfeiffer, F., and Tsugita, A. 1998. The PIR-International Protein Sequence Database. Nucl. Acids Res. 26:27-32. Benson, D.A., Boguski, M.S., Lipman, D.J., Ostell, J., and Ouellette, B.F. 1998. GenBank. Nucl. Acids Res. 26:1-7. Boguski, M.S., Lowe, T.M., and Tolstoshev, C.M. 1993. dbEST—database for “expressed sequence tags”. Nature Genet. 4:332-333. Chandrasekharappa, S.C., Guru, S.C., Manickam, P., Olufemi, S.E., Collins, F.S., Emmert-Buck, M.R., Debelenko, L.V., Zhuang, Z., Lubensky, I.A., Liotta, L.A., et al. 1997. Positional cloning of the gene for multiple endocrine neoplasia-type 1. Science 276:404-407. Chang, Z.Y., Nygaard, P., Chinault, A.C., and Kellems, R.E. 1991. Deduced amino acid sequence of Escherichia coli adenosine deaminase reveals evolutionarily conserved amino acid residues: Implications for catalytic function. Biochemistry 30:2273-2280. Claverie, J.M. and Makalowski, W. 1994. Alu alert. Nature 371:752. Dayhoff, M.O., Schwartz, R.M., and Orcutt, B.C. 1978. A model of evolutionary change in proteins. In Atlas of Protein Sequence and Structure, Vol. 5, suppl. 3. (M.O. Dayhoff, ed.) pp. 345-352. Natl. Biomed. Res. Found., Washington, D.C. Gish, W. and States, D.J. 1993. Identification of protein coding regions by database similarity search. Nature Genet. 3:266-272. Henikoff, S. and Henikoff, J.G. 1992. Amino acid substitution matrices from protein blocks. Proc. Natl. Acad. Sci. U.S.A. 89:10915-10919. Holm, L. and Sander, C. 1997. An evolutionary treasure: Unification of a broad set of amidohydrolases related to urease. Proteins 28:72-82. Karlin, S. and Altschul, S.F. 1990. Methods for assessing the statistical significance of molecular sequence features by using general scoring schemes. Proc. Natl. Acad. Sci. U.S.A. 87:2264-2268. Karlin, S. and Altschul, S.F. 1993. Applications and statistics for multiple high-scoring segments in molecular sequences. Proc. Natl. Acad. Sci. U.S.A. 90:5873-5877. Lavin, M.F. and Shiloh, Y. 1997. The genetic defect in ataxia-telangiectasia. Annu. Rev. Immunol. 15:177-202. Olson, M., Hood, L., Cantor, C., and Botstein, D. 1989. A common language for physical mapping of the human genome. Science 245:1434-1435. Ostell, J.M. and Kans, J.A. 1998. The NCBI data model. In Bioinformatics: A Practical Guide to the Analysis of Genes and Proteins (A.D. Baxevanis and B.F.F. Ouellette, eds.) pp. 121-144. John Wiley & Sons, New York. Ouellette, B.F.F. 1998. The GenBank sequence database. In Bioinformatics: A Practical Guide to the Analysis of Genes and Proteins (A.D. Baxevanis and B.F.F. Ouellette, eds.) pp. 16-45. John Wiley & Sons, New York. Ouellette, B.F. and Boguski, M.S. 1997. Database divisions and homology search files: A guide for the perplexed. Genome Res. 7:952-955. Pearson, W.R. 1990. Rapid and sensitive sequence comparison with FASTP and FASTA. Methods Enzymol. 183:63-98. Pearson, W.R. and Lipman, D.J. 1988. Improved tools for biological sequence comparison. Proc. Natl. Acad. Sci. U.S.A. 85:2444-2448. Schuler, G.D. 1998. Sequence alignment and database searching. In Bioinformatics: A Practical Guide to the Analysis of Genes and Proteins (A.D. Baxevanis and B.F.F. Ouellette, eds.) pp. 145-171. John Wiley & Sons, New York. Schwartz, R.M. and Dayhoff, M.O. 1978. Matrices for detecting distant relationships. In Atlas of Protein Sequence and Structure, Vol. 5, suppl. 3. (M.O. Dayhoff, ed.) pp. 353-358. Natl. Biomed. Res. Found., Washington, D.C. Seabra, M.C., Brown, M.S., and Goldstein, J.L. 1993. Retinal degeneration in choroideremia: Deficiency of rab geranylgeranyl transferase. Science 259:377-381. Smith, T.F. and Waterman, M.S. 1981. Identification of common molecular subsequences. J. Mol. Biol. 147:195-197. Smith, M.W., Holmsen, A.L., Wei, Y.H., Peterson, M., and Evans, G.A. 1994. Genomic sequence sampling: A strategy for high resolution sequence-based physical mapping of complex genomes. Nature Genet. 7:40-47. Sequence Similarity Searching Using BLAST
Stoesser, G., Moseley, M.A., Sleep, J., McGowran, M., Garcia-Pastor, M., and Sterk, P. 1998. The EMBL nucleotide sequence database. Nucl. Acids Res. 26:8-15.
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Tateno, Y., Fukami-Kobayashi, K., Miyazaki, S., Sugawara, H., and Gojobori, T. 1998. DNA Data Bank of Japan at work on genome sequence data. Nucl. Acids Res. 26:16-20. Wolfsberg, T.G., Straight, P.D., Gerena, R.L., Huovila, A.P., Primakoff, P., Myles, D.G., and White, J.M. 1995. ADAM, a widely distributed and developmentally regulated gene family encoding membrane proteins with a disintegrin and metalloprotease domain. Dev. Biol. 169:378-383. Wootton, J.C. and Federhen, S. 1993. Statistics of local complexity in amino acid sequences and sequence databases. Comput. Chem. 17:149-163. Wootton, J.C. and Federhen, S. 1996. Analysis of compositionally biased regions in sequence databases. Methods Enzymol. 266:554-571. Zhang, J. and Madden, T.L. 1997. PowerBLAST: A new network BLAST application for interactive or automated sequence analysis and annotation. Genome Res. 7:649-656. Zhang, Z., Berman, P., and Miller, W. 1998. Alignments without low-scoring regions. J. Comput. Biol. 5:197-210.
Contributed by Tyra G. Wolfsberg and Thomas L. Madden National Center for Biotechnology Information National Library of Medicine, NIH Bethesda, Maryland
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APPENDIX A: BLAST PARAMETERS The Advanced BLAST page offers the possibility to change a number of BLAST parameters. The most important options supported are listed below. Descriptions (pull-down menu) restricts the number of one-line descriptions to the number specified. The default is 500. Alignments (pull-down menu) restricts the number of database hits for which alignments are shown. Several alignments may be associated with one database sequence, so the number of alignments may actually be larger than this number. The default is 500. Expect value (pull-down menu; also known as the E value) is the statistical significance threshold for reporting matches. The default value is 10, such that ten alignments are expected to be found merely by chance, according to the stochastic model of Karlin and Altschul (1990). Lower E thresholds are more stringent, leading to fewer chance matches being reported. Fractional values are acceptable. Filtering (pull-down menu) masks off segments of the query sequence that have low compositional complexity, as determined by the SEG program (Wootton and Federhen, 1993, 1996) or, for BLASTN, by the DUST program (R.L. Tatusov and D.J. Lipman, pers. comm.). Filtering is only applied to the query sequence (or its translation products), not to database sequences. Default filtering is DUST for BLASTN, SEG for other programs. Genetic code (pull-down menu) selects the genetic code to be used for a BLASTX translation of the query. The default setting is the universal code. The choice of code is determined by the source of the query sequence (e.g., species, mitochondrial). Graphical overview (checkbox) selects whether an overview of the database sequences aligned to the query sequence will be shown. The score of an alignment is indicated by one of five different colors, which divides the range of scores into five groups. If more than one alignment is displayed for a database sequence, the two alignments are connected by a cross-hatched bar. Mousing over a match causes the definition and score to be shown in the window at the top; clicking on a bar takes the user to the associated alignments. The overview is on by default. Matrix (menu) allows one to change the matrix from the default of BLOSUM-62. This is useful for searches against short sequences (see Searching Short Sequences). It is also possible to change the gap existence and extension penalties, but the defaults for a given matrix are recommended. Organism (pull-down menu) limits the search by organism. The menu provides a list of the most common organisms; others may be entered in a text box.
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APPENDIX B: SEQUENCE IDENTIFIER SYNTAX The syntax of sequence header lines used by the NCBI BLAST server depends on the database from which each sequence was obtained. Table 2.5.3 lists the identifiers for the databases from which the sequences were derived. For example, an identifier might be gb|M73307|AGMA13GT, where the gb tag indicates that the identifier refers to a GenBank sequence, M73307 is its GenBank accession number, and AGMA13GT is its GenBank locus. NCBI assigns gi identifiers for all sequences contained within NCBI’s sequence databases (Ostell and Kans, 1998). The gi identifier provides a uniform and stable naming convention whereby a specific sequence is assigned its unique gi identifier. If a nucleotide or protein sequence changes, a new gi identifier is assigned, even if the accession number of the record remains unchanged. Thus, gi identifiers provide a mechanism for identifying the exact sequence that was used or retrieved in a given search. For searches of the nr protein database where the sequences are derived from conceptual translations of sequences from the nucleotide databases the gi syntax is gi|gi_identifier. An example would be gi|451623 (U04987) env [Simian immunodeficiency..., where 451623 is the gi identifier and U04987 is the accession number of the nucleotide sequence from which it was derived. Users may select the -gi option for BLAST output, which will produce a header line with the gi identifier concatenated with the database identifier of the database from which it was derived. For example, gi|176485|gb|M73307|AGMA13GT would be used for a match from a nucleotide database, and gi|129295|sp|P01013|OVAX_CHICK for a protein database. The gnl (general) identifier allows databases not listed in Table 2.5.3 to be identified with the same syntax. An example here is the PID identifier gnl|PID|e1632. PID stands for Protein-ID, and the e in e1632 indicates that this ID was issued by EMBL. As mentioned above, use of the -gi option produces the NCBI gi (in addition to the PID), which users can also use to retrieve sequences of interest. Table 2.5.3
Identifier Syntax for Sequence Databases
Database
Identifier syntax
GenBank EMBL Data Library DDBJ (DNA Data Bank of Japan) NBRF PIR Protein Research Foundation Swiss-Prot Brookhaven Protein Data Bank Patents GenInfo Backbone Id General database identifier
gb|accession|locus emb|accession|locus dbj|accession|locus pir||entry prf||name sp|accession|entry name pdb|entry|chain pat|country|number bbs|number gnl|database|identifier
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Protein Databases on the Internet
UNIT 2.6
Protein databases have become a crucial part of modern biology. Huge amounts of data for protein structures, functions, and particularly sequences are being generated. These data cannot be handled without using computer databases. Searching databases is often the first step in the study of a new protein. Without the prior knowledge obtained from such searches, known information about the protein could be missed, or an experiment could be repeated unnecessarily. Comparison between proteins and protein classification provide information about the relationship between proteins within a genome or across different species, and hence offer much more information than can be obtained by studying only an isolated protein. In this sense, protein comparison through databases allows one to view life as a forest instead of individual trees. In addition, secondary databases derived from experimental databases are also widely available. These databases reorganize and annotate the data or provide predictions. The use of multiple databases often helps researchers understand evolution, structure, and function of a protein. Protein databases are especially powered by the Internet. Unlike traditional media, such as the CD-ROM, the Internet allows databases to be easily maintained and frequently updated with minimum cost. Researchers with limited resources can afford to set up their own databases and disseminate their data quickly. Notably, many small databases on specific types of proteins, such as the EF-Hand Calcium-Binding Proteins Data Library (http://structbio.vanderbilt.edu/cabp_database/) and O-GlycBase (http://www.cbs.dtu. dk/databases/OGLYCBASE/), became widely available. Users worldwide can easily access the most up-to-date version through a user-friendly interface. Most protein databases have interactive search engines so that users can specify their needs and obtain the related information interactively. Many protein databases also allow submitters to deposit data interactively, and allow database servers to check the format of the data and provide immediate feedback. Although some protein databases are widely known, they are far from being fully utilized in the protein science community. This unit provides a starting point for readers to explore the potential of protein databases on the Internet. Databases for different aspects of proteins are discussed, with the focus on sequence, structure, and family. The strengths and weaknesses of the databases will be addressed. For Web addresses of the databases discussed in this unit, see Internet Resources and Table 2.6.1. From hundreds of on-line protein databases, several major databases are discussed as examples to illustrate their features and how they can be used effectively. Most other protein databases can be explored in a similar way. PROTEIN SEQUENCE DATABASES Thanks to the Human Genome Project and other sequencing efforts, new sequences have been generated at a prodigious rate. A large number of DNA sequences, which are equivalent to hundreds of new proteins, are determined each day for the human genome alone. It is expected that 10,000 genomes will be sequenced in the next decade. These sequences provide a rich information source and the core of the revolutionary movement toward “large-scale biology.” Various databases contain protein sequences with different focuses. SwissProt (Bairoch and Apweiler, 1999) provides more annotations than any other sequence database, with a minimal level of redundancy, through human input or integration with other databases. However, the labor-intensive enhancement process prevents many sequences from being added to SwissProt quickly. As a supplement of SwissProt, the TrEMBL database Contributed by Dong Xu and Ying Xu Current Protocols in Protein Science (2003) 2.6.1-2.6.15 Copyright © 2003 by John Wiley & Sons, Inc.
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Table 2.6.1
Web Addresses and Sizes of Selected Protein Databases
Web site
Sizea
DDBJ DAtA GeneCards Genome Channel KEGG nr OWL PEDANT PIR PRF STACK SwissProt SYSTERS TIGR Gene Indices TrEMBL UniGene Structure:
http://www.ddbj.nig.ac.jp/ http://luggagefast.stanford.edu/group/arabprotein/ http://bioinfo.weizmann.ac.il/cards/ http://compbio.ornl.gov/channel/ http://www.genome.ad.jp/kegg/ http://www.ncbi.nlm.nih.gov/BLAST/ http://www.bioinf.man.ac.uk/dbbrowser/OWL/ http://pedant.mips.biochem.mpg.de http://pir.georgetown.edu http://www.genome.ad.jp/htbin/www_bfind?prf http://www.sanbi.ac.za/Dbases.html http://www.expasy.ch/sprot/sprot-top.html http://systers.molgen.mpg.de/ http://www.tigr.org/tdb/tgi/ http://www.expasy.ch/sprot/sprot-top.html http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=unigene
25,149,821 sequences 25,846 sequences 14,519 genes NA 433,051 genes 129,6341 sequences 279,796 sequences NA 1,173,204 sequences 216,752 sequences NA 123,192 sequences 290,811 sequences NA 829,760 sequences NA
3Dee BioMagResBank Decoys ’R’ Us EBI-MSD Enzyme Structures GRASS Image Library LPFC MolMovDB PDB PDBsum PRESAGE STING WPDB Sequence family:
http://www.compbio.dundee.ac.uk/3Dee/ http://www.bmrb.wisc.edu http://dd.stanford.edu/ http://msd.ebi.ac.uk http://www.biochem.ucl.ac.uk/bsm/enzymes/ http://trantor.bioc.columbia.edu/GRASS/ http://www.imb-jena.de/IMAGE.html http://www-smi.stanford.edu/projects/helix/LPFC/ http://bioinfo.mbb.yale.edu/MolMovDB/ http://www.rcsb.org/pdb/ http://www.biochem.ucl.ac.uk/bsm/pdbsum/ http://presage.berkeley.edu/ http://trantor.bioc.columbia.edu/STING/ http://www.sdsc.edu/pb/wpdb/
NA 2494 structures NA NA 10208 structures NA NA NA NA 20,473 structures 21,361 structures NA NA >6000 structures
BLOCKS COG DOMO InterPro Pfam PRINTS ProClass ProDom PROSITE SBASE VIDA Structure family:
http://www.blocks.fhcrc.org/ http://www.ncbi.nlm.nih.gov/COG/ http://www.infobiogen.fr/services/domo/ http://www.ebi.ac.uk/interpro/ http://www.sanger.ac.uk/Pfam/ http://www.bioinf.man.ac.uk/dbbrowser/PRINTS/ http://pir.georgetown.edu/gfserver/proclass.html http://protein.toulouse.inra.fr/prodom.html http://www.expasy.ch/prosite/ http://hydra.icgeb.trieste.it/~kristian/SBASE/ http://www.biochem.ucl.ac.uk/bsm/virus_database/VIDA.html
8656 blocks 104,101 sequences 83,054 sequences 6383 families 5193 families 1750 sites 155,868 sequences 481,952 sequences 1614 sites 338,655 sequences NA
CATH CE
http://www.biochem.ucl.ac.uk/bsm/cath/ http://cl.sdsc.edu/ce.html
36,480 domains NA
Database Sequence:
continued
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Table 2.6.1
Web Addresses and Sizes of Selected Protein Databases, continued
Database
Web site
Sizea
CL Dali Domain Dictionary FSSP HOMSTRAD HSSP PartsList SCOP SLoop Function family:
http://cl.sdsc.edu/cl1.html http://columba.ebi.ac.uk:8765/holm/ddd2.cgi
NA 771 domain classes
http://www2.ebi.ac.uk/dali/fssp/ http://www-cryst.bioc.cam.ac.uk/~homstrad/ http://swift.embl-heidelberg.de/hssp/ http://bioinfo.mbb.yale.edu/align/ http://scop.mrc-lmb.cam.ac.uk/scop/ http://www-cryst.bioc.cam.ac.uk/~sloop/
3242 families 1033 families NA NA 17406 structures NA VAST
http://rose.man.poznan.pl/aars/index.html http://ir2lcb.cnrs-mrs.fr/ABCdb/ http://www.eez.csic.es/arac-xyls/ http://wwwmgs.bionet.nsc.ru/mgs/gnw/aspd/ http://condor.bcm.tmc.edu/ermb/bcgd/ http://www.chemie.uni-marburg.de/~csdbase/ http://www.helicase.net/dexhd/dbhome.htm
NA NA NA NA NA NA NA
http://structbio.vanderbilt.edu/cabp_database/ http://ecocyc.org/ http://us.expasy.org/enzyme/ http://geneontology.org/ http://www.gpcr.org/7tm/ http://www.biosci.ki.se/groups/tbu/homeo.html http://merops.sanger.ac.uk/ http://wehih.wehi.edu.au/mhcpep/ http://npd.hgu.mrc.ac.uk/
NA NA 4173 enzymes NA NA NA NA 13,000 peptides 1227 proteins
http://www.cbs.dtu.dk/databases/OGLYCBASE/ http://www-lecb.ncifcrf.gov/PDD/ http://www.biochem.ucl.ac.uk/bsm/PROCAT/PROCAT.html http://pkr.sdsc.edu/html/index.shtml
242 glycoproteins NA NA 7588 clusters
http://www.mbio.ncsu.edu/RNaseP/home.html http://www-wit.mcs.anl.gov/sentra/ http://66.93.129.133/transporter/wb/index2.html http://condor.bcm.tmc.edu/oncogene.html http://wit.mcs.anl.gov/WIT2/ http://www.stanford.edu/~rnusse/wntwindow.html
NA NA NA >300 genes NA NA
http://binddb.org http://dip.doe-mbi.ucla.edu http://biodata.mshri.on.ca/grid/ http://daisy.bio.nagoya-u.ac.jp/golab/hetpdbnavi.html http://surya.bic.nus.edu.sg/mpid/
15,141 interactions 22,229 interactions 13,819 interactions NA 90 complexes
AARSDB ABCdb AraC/XylS database ASPD Breast Cancer Gene CSDBase DExH/D Family Database EF-hand CaBP EcoCyc ENZYME Gene Ontology (GO) GPCRDB (receptors) Homeobox Page MEROPS (peptidase) MHCPEP (peptides) Nuclear Protein Database (NPD) O-GlycBase PDD PROCAT Protein Kinase Resource (PKR) RNase P Sentra TransportDB Tumor Gene WIT2 Wnt gene Homepage Binding: BIND DIP GRID Het-PDB Navi MHC-Peptide Interaction Database
continued
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Table 2.6.1
Web Addresses and Sizes of Selected Protein Databases, continued
Database
Web site
Sizea
Protein-Protein Interface ReLiBase Energetics: ProTherm References:
http://www-lmmb.ncifcrf.gov:80/~tsai/
NA
http://relibase.rutgers.edu/
11,938 proteins
http://www.rtc.riken.go.jp/jouhou/Protherm/protherm.html
13046 entries
http://www.infobiogen.fr/services/dbcat/
511 databases
http://www.ncbi.nlm.nih.gov/PubMed/ http://www.expasy.ch/seqanalref/
>12,000,000 citations NA
http://www.rtc.riken.go.jp/jouhou/3dinsight/3dinsight.html http://www.ncbi.nlm.nih.gov/Entrez/ http://srs.ebi.ac.uk
NA NA 264 databases
DBCAT (Catalog of Databases) MEDLINE SeqAnalRef Combined: 3DinSight Entrez SRS
aNA, size not available at time of printing. The data are as of March, 2003.
(Bairoch and Apweiler, 1999) contains all the translations of EMBL nucleotide sequence entries (Rodriguez-Tom et al., 1996) that have not been integrated into SwissProt. Another protein sequence database, the Protein Identification Resource (PIR; Barker et al., 1999), attempts to build a complete and nonredundant database from a number of protein and nucleic acid sequence databases. Identical and highly similar sequences from the same species are merged into a single entry. Each entry in PIR provides bibliographic and annotated information for the protein. The nr protein database is used for BLAST searching (Altschul et al., 1997), which is described in UNIT 2.5 of this book. It includes entries from the nonredundant GenBank (Benson et al., 1999) translations, SwissProt, PIR, Protein Research Foundation (PRF) in Japan, and the Protein Data Bank (PDB). Only entries with absolutely identical sequences are merged. Most of the sequence databases have a sequence search tool and cross-references to entries of other protein and gene databases. Many sequence databases, such as SwissProt and PIR, also provide text searching using, for instance, protein names or key words. To study a new protein, the authors recommend first performing a sequence search using BLAST in nr if the protein sequence is available. The search often gives entry names in the protein databases included in nr. Even when the protein is not found in nr, it is likely that a homologous protein will be hit, which can often lead to some useful information, such as the function of the query protein. If the sequence of the query protein is unavailable, doing a text search in SwissProt or PIR usually identifies the protein. SwissProt is probably the place to obtain the most information about a protein if it can be found in SwissProt. However, some additional information may be found by checking other sequence databases. For example, PIR provides some useful information on protein family classification for each entry, and the Kyoto Encyclopedia of Genes and Genomes (KEGG; Ogata et al., 1999) annotates some gene entries with information about metabolic and regulatory pathways. If the query protein is not available in nr, one may find a gene model (predicted protein sequence) by searching genetic sequence databases that contain predicted genes from nucleotide sequences (e.g., the Genome Channel; Mural et al., 1999). Although predicted sequences generated by computational gene-finding tools may contain errors, such databases cover a large number of proteins and are often reliable enough to provide useful information. Protein Databases on the Internet
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SwissProt, as a curated protein sequence database, offers a wide range of annotations, covering areas such as function, domain parsing, post-translational modifications, and variants. SwissProt can be accessed at http://www.expasy.ch/sprot/sprot-top.html (home server, Switzerland), http://us.expasy.org/sprot/sprot-top.html (USA), http://ca.expasy. org/sprot/sprot-top.html (Canada), http://cn.expasy.org/sprot/sprot-top.html (China), http://kr.expasy.org/sprot/sprot-top.html (Korea), and http://tw.expasy.org/sprot/sprot-top. html (Taiwan). SwissProt is integrated into many other databases, such as SRS (see SRS below). Human vitronectin is used here as an example for searching protein sequence databases. To locate the SwissProt entry for this protein, one can search either the entry name (VTNC_HUMAN) or the accession number (P04004) obtained from a BLAST search. Alternatively, one can use the full-text search at the SwissProt Web page to search by protein name (human vitronectin) or key words (e.g., serum spreading, as vitronectin is also called serum spreading factor s-protein). A combination of several entries can be used in a search. The entry name in SwissProt has the general format X_Y, where X is a mnemonic code of up to four characters indicating the protein name (in this case, VTNC), and Y is a mnemonic species identification code of up to five characters for the biological source of the protein. Some codes used for Y are full English names, e.g., HORSE, HUMAN, MAIZE, MOUSE, PIG, RAT, SHEEP, YEAST (baker’s yeast, Saccharomyces cerevisiae), and WHEAT. Some are abbreviations, including BOVIN (bovine), CHICK (chicken), ECOLI (Escherichia coli), PEA (garden pea, Pisum sativum), RABIT (rabbit), SOYBN (soybean, Glycine max), and TOBAC (common tobacco, Nicotina tabacum). An entry name may have several accession numbers if they have been merged. An accession number is always conserved from release to release, and therefore allows unambiguous citation. Each entry contains the following items shown in table format in the NiceProt View layout: (1) general information about the entry, (2) name and origin of the protein, (3) references providing the protein sequence, (4) comments (e.g., annotated functions, subunits, similar proteins), (5) cross-references (links to other databases), (6) key words, (7) features, and (8) sequence information. The text in the comments entry provides a function annotation for the protein (e.g., “Vitronectin is a cell adhesion and spreading factor found in serum and tissues. Vitronectins interact with glycosaminoglycans and proteoglycans...”). Cross-references lists the annotations of the protein by other databases, such as GeneCards (Rebhan et al., 1998) and ProDom (Corpet et al., 1999). GeneCards, a database of human genes, shows chromosomal location and the involvement of the protein in certain diseases (if applicable). ProDom contains protein domain families by automated sequence comparisons. Clicking the link to ProDom from SwissProt leads to a nice graphic view for domain parsing, as shown in Figure 2.6.1 for vitronectin.
0
100
200
300
400
500
100
200
300
400
500
VTNC_HUMAN
0
Figure 2.6.1 Domain parsing of human vitronectin by ProDom in SwissProt.
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Various research results are given under features. Some of the features items for VTNC_HUMAN are as follows: SIGNAL CHAIN CHAIN PEPTIDE DOMAIN DOMAIN SITE SITE MOD_RES CARBOHYD DISULFID BINDING CONFLICT
1 20 399 20 150 288 64 398 75 86 293 362 50
19 398 478 63 287 478 66 399 75 86 430 395 50
V65 SUBUNIT. V10 SUBUNIT. SOMATOMEDIN B. HEMOPEXIN-LIKE 1. HEMOPEXIN-LIKE 2. CELL ATTACHMENT SITE. CLEAVAGE. SULFATATION.
HEPARIN. C -> N (IN REF. 5).
where SIGNAL represents the extent of a signal sequence (prepeptide), MOD_RES indicates a post-translationally modified residue (sulfatation, in this case), CARBOHYD shows the glycosylation site, DISULFID means that a disulfide bond exists between the two indicated residues (293 and 430), and CONFLICT shows that different papers report differing sequences. PROTEIN STRUCTURAL DATABASES Searching structure databases is becoming more and more popular in molecular biology. The three-dimensional structures of proteins not only define their biological functions, but also hold a key in rational drug design. Traditionally, protein structures were solved at a low-throughput mode. However, recent advances in new technologies, such as synchrotron radiation sources and high-resolution nuclear magnetic resonance (NMR), accelerate the rate of protein structure determination substantially. There is an overwhelming consensus in the structural biology community that protein structures can be solved en masse (an effort called structural genomics), in a similar fashion as for determining DNA sequences, and that impact of this approach can be compared with that of the Human Genome Project (see Chapter 17). The only international repository for the processing and distribution of protein structures is the PDB (Bernstein et al., 1977). The structures in the PDB were determined experimentally by X-ray crystallography (∼86%) and NMR (∼14%). Theoretical models have been removed from PDB, effective July 2, 2002, based on the new PDB policy. The PDB also contains some structures of chemical ligands and nucleotides. Each PDB entry is represented by a four-character identifier (PDB ID), where the first character is always a number from 0 to 9 (e.g., 1cau, 256b). The PDB can be accessed through the home server (http://www.rcsb.org/pdb/ or http://www.pdb.org in the USA) or through one of many mirror sites from around the world, as listed at the home server. Some of the mirror sites are as follows: Rutgers University, Piscataway, NJ, United States: http://rutgers.rcsb.org/ NIST, Gaithersburg, MD, United States: http://nist.rcsb.org/ Cambridge Crystallographic Data Centre, United Kingdom: http://pdb.ccdc.cam.ac.uk/ National University of Singapore, Singapore: http://pdb.bic.nus.edu.sg/ Osaka University, Japan: http://pdb.protein.osaka-u.ac.jp/ Universidade Federal de Minas Gerais, Brazil: http://www.pdb.ufmg.br/ Max Delbrück Center for Molecular Medicine, Germany: http://www.pdb.mdc-berlin.de/ Protein Databases on the Internet
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The PDB offers three search methods: search by PDB ID, by SearchLite, and by SearchFields. SearchLite is a simple key-word search that uses, for instance, protein name or author’s name. A search using SearchFields, as an advanced search engine, allows a user to specify features of the protein, such as Enzyme Commission (EC) number, name of binding ligand, range of protein size, range of resolution in the X-ray structure, and secondary structure content. The PDB stores structural information in two formats: the PDB file format (Bernstein et al., 1977) and the macromolecular crystallographic information file (mmCIF) format (Bourne et al., 1997). The PDB file format is still the dominant format used in the protein community. It contains three parts: annotations, coordinates, and connectivities. The connectivity part, which shows chemical connectivities between atoms, is optional. It is listed at the end of the PDB file, beginning the line with the key word CONECT. The coordinate part uses each line for a three-dimensional coordinate of an atom, starting from ATOM (for standard amino acids) or HETATM (for nonstandard groups). The following shows an example of the PDB file format: HEADER
OXIDOREDUCTASE
COMPND
GLYCOLATE OXIDASE (E.C.1.1.3.1)
(OXYGEN(A))
14-JUN-89
1GOX
1GOX
3
1GOX
4
... ATOM
232
N
ALA
29
54.035
4.332
19.352
1.00
23.93
1GOX
374
ATOM
233
CA
ALA
29
52.992
65.356 19.569
1.00
24.74
1GOX
375
ATOM
234
C
ALA
29
53.519
66.762 19.309
1.00
25.43
1GOX
376
ATOM
235
O
ALA
29
54.648
67.179 19.655
1.00
25.66
1GOX
377
ATOM
236
C
BALA 29
52.433
65.340 20.993
1.00
24.54
1GOX
378
HETATM
3165
O
HOH 658
62.480
62.480 0.000
0.50
65.79N
1GOX
3170
CONECT
2837 2838
1GOX
3171
... 2854
Each line shows the atom serial number, atom type, residue type, chain identifier (in case of multi-chain structure), residue serial number, orthogonal coordinates (three values), occupancy, temperature factor, and segment identifier. The annotation part of the PDB file format contains dozens of possible record types, including: HEADER (name of protein and release date), COMPND (molecular contents of the entry), SOURCE (biological source), AUTHOR (list of contributors), SSBOND (disulfide bonds), SLTBRG (salt bridges), SITE (groups comprising important sites), HET (nonstandard groups or residues [heterogens]), MODRES (modifications to standard residues), SEQRES (primary sequence of backbone residues), HELIX (helical substructures), SHEET (sheet substructures), and REMARK (other information and comments). The PDB allows a user to view a molecule structure interactively through a Virtual Reality Modeling Language (VRML) viewer, RasMol (Sayle and Milner-White, 1995), Chime, or QuickPDB (a Java applet for viewing sequence and structure) when the browser is configured to support these free rendering tools. The PDB provides related information about the protein, such as secondary structure assignment and geometry. Each PDB entry also links to a wide range of annotations from secondary databases, including (1) summary and display databases such as Graphical Representation and Analysis of Structure Server (GRASS; Nayal et al., 1999), Image Library (Shnel, 1996), Molecular Modelling Database (MMDB; Marchler-Bauer et al., 1999) in Entrez, PDBsum (Laskowski et al., 1997), and Sequence to and within Graphics (STING); (2) domain partition information from 3Dee (Siddiqui and Barton, 1995); (3) the MEDLINE bibliography; (4) structure quality assessment in PDBREPORT from WHAT IF (Vriend, 1990); (5) protein movements recorded in Database of Macromolecular Movement (MolMovDB; Gerstein and Krebs,
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1998); (6) structure families (CATH, CE, FSSP, SCOP, and VAST, as discussed later in this unit); and (7) geometry analyses of the protein, such as CSU Contacts of Structural Units (Sobolev et al., 1999) and castP Identification of Protein Pockets & Cavities (Liang et al., 1998). Several structure databases that are not linked by the PDB can also provide useful information. WPDB (Shindyalov and Bourne, 1995) can be used to visualize and analyze a PDB entry from Microsoft Windows. BioMagResBank (University of Wisconsin, 1999) is a repository for NMR spectroscopy data on proteins, peptides, and nucleic acids. Particularly, it provides partial NMR data (e.g., chemical shifts) before the full structure is solved. PROTEIN FAMILY DATABASES Introduction Proteins can be classified according to their evolutionary, structural, or functional relationships. A protein in the context of its family is much more informative than the single protein itself. For example, residues conserved across the family often indicate special functional roles. Two proteins classified in the same functional family may suggest that they share similar structures, even when their sequences do not have significant similarity. There is no unique way to classify proteins into families. Boundaries between different families may be subjective. The choice of classification system depends in part on the problem; in general, the authors suggest looking into classification systems from different databases and comparing them. Three types of classification methods are widely adopted, based upon the similarity of sequence, structure, or function. Sequence-based methods are applicable to any proteins whose sequences are known, while structure-based methods are limited to the proteins of known structures, and function-based methods depend on the functions of proteins being annotated. Sequence- and structure-based classifications can be automated and are scalable to high-throughput data, whereas function-based classification is typically carried out manually. Structure- and function-based methods are more reliable, while sequence-based methods may result in a false positive result when sequence similarity is weak (i.e., two proteins are classified into one family by chance rather than by any biological significance). In addition, since protein structure and function are better conserved than sequence, two proteins having similar structures or similar functions may not be identified through sequence-based methods. Databases for Sequence-Based Protein Families Sequence-based protein families are classified according to a profile derived from a multiple-sequence alignment. The profile can be shown across a long domain (typically 100 residues or more) or can be revealed in short sequence motifs. Classification methods based on profiles across long domains tend to be more reliable but less sensitive than those based on short sequence motifs.
Protein Databases on the Internet
Several sequence-based methods focus more on profiles across long domains, including Pfam (Bateman et al., 1999), ProDom (Corpet et al., 1999), SBASE (Murvai et al., 1999), and Clusters of Orthologous Group (COG; Tatusov et al., 1997). These methods differ in the techniques used to construct families. Pfam builds multiple-sequence alignments of many common protein domains using hidden Markov models. The ProDom protein domain database consists of homologous domains based on recursive PSI-BLAST searches (UNIT 2.5). SBASE is organized through BLAST neighbors and is grouped by standard protein names that designate various functional and structural domains of protein sequences. COG aims toward finding ancient conserved domains by delineating families of orthologs across a wide phylogenetic range.
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The following shows an example of Pfam for the GRIP domain (accession number PF01465). Pfam lists some useful information for the entry as follows: The GRIP (golgin-97, RanBp2alpha, Imh1p and p230/golgin245) domain is found in many large coiled-coil proteins. It has been shown to be sufficient for targeting to the Golgi. The GRIP domain contains a completely conserved tyrosine residue. The references of the above annotation are also given. In addition, Pfam gives the alignment between the family members: 015045/1511-1558
SAANLEYLKNVLLQFIFLKPG--SERERLLPVINTMLQLSPEEKGKLAAV
O15045
YNP9_CAEEL/633-681
NEKNMEYLKNVFVQFLKPESVP-AERDQLVIVLQRVLHLSPKEVEILKAA
P34562
Q06704/864-909
KNEKIAYIKNVLLGFLEHKE----QRNQLLPVISMLLQLDSTDEKRLVMS
Q06704
Q92805/691-737
REINFEYLKHVVLKFMSCRES---EAFHLIKAVSVLLNFSQEEENMLKET
Q92805
O42657/703-748
MLIDKEYTRNILFQFLEQRD----RRPEIVNLLSILLDLSEEQKQKLLSV
O42657
O70365/1161-1205
EPTEFEYLRKVMFEYMMGR-----ETKTMAKVITTVLKFPDDQAQKILER
070365
Q21071/692-741D
DPAEAEYLRNVLYRYMTNRESLGKESVTLARVIGTVARFDESQMKNVISS
Q21071
Q18013/574-623
STSEIDYLRNIFTQFLHSMGSPNAASKAILKAMGSVLKVPMAEMKIIDKK
Q18013
The alignment shows accession numbers and the range of each sequence. One can identify some features of the family through this pattern (i.e., from particularly conserved residues at specific alignment positions). Some methods are based on “fingerprints” of small conserved motifs in sequences, as with PROSITE (Hofmann et al., 1999), PRINTS (Attwood et al., 1999), and BLOCKS (Heniko et al., 1999). In protein sequence families, some regions have been better conserved than others during evolution. These regions are generally important for the function of a protein or for the maintenance of its three-dimensional structure, and hence are suitable for fingerprinting. The fingerprints can be used to assign a newly sequenced protein to a specific family. Fingerprints are derived from gapped alignments in PROSITE and PRINTS, but are derived from ungapped alignments (corresponding to the highly conserved regions in proteins) in BLOCKS. A fingerprint in PRINTS may contain several motifs from PROSITE, and thus may be more flexible and powerful than a single PROSITE motif. Therefore, PRINTS can provide a useful adjunct to PROSITE. It should be noted that some functionally unrelated proteins may be classified together due to chance matches in short motifs. Other sequence-based protein family databases consist of multiple sources. The ProClass database (Wu et al., 1999) is a nonredundant protein database organized according to family relationships as defined collectively by PROSITE patterns and PIR superfamilies. The MEGACLASS server (States et al., 1993) provides classifications by different methods, including Pfam, BLOCKS, PRINTS, ProDom, and SBASE. The MOTIF search engine at http://motif.genome.ad.jp/ includes PROSITE, BLOCKS, ProDom, and PRINTS. Databases for Structure-Based Protein Families The hierarchical relationship among proteins can be clearly revealed in structures through structure-structure comparison. Structure families often provide more information on the relationship between proteins than what sequence families can offer, particularly when two proteins share a similar structure but no significant sequence identity. Figure 2.6.2 shows an example of a structure-structure alignment between two proteins. Sometimes, sequence similarity between two proteins exists but is not strong enough to produce an unambiguous alignment. In this case, the alignment between two structures can generate Computational Analysis
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Figure 2.6.2 Structure superposition between glycolate oxidase (1gox, in black) and inosine monophosphate dehydrogenase (1ak5, in gray). This figure was made using MOLSCRIPT (Kraulis, 1991).
better alignment in terms of biological significance, and thus may pinpoint the active sites more accurately. Different structure-structure comparison methods yield different structure families. CATH (Class, Architecture, Topology and Homologous superfamily; Orengo et al., 1997) is a hierarchical classification of protein domain structures. CE (Combinatorial Extension of the optimal path; Shindyalov and Bourne, 1998) provides structural neighbors of the PDB entries with structure-structure alignments and three-dimensional superpositions. FSSP (Fold classification based on Structure-Structure alignment of Proteins; Holm and Sander, 1996) features a protein family tree and a domain dictionary, in addition to whole-chain-based classification, sequence neighbors, and multiple structure alignments. SCOP (Structural Classification of Proteins; Murzin et al., 1995) uses augmented manual classification, class, fold, superfamily, and family classification. VAST (Vector Alignment Search Tool; Gibrat et al., 1996) contains representative structure alignments and threedimensional superpositions. Among these five databases, SCOP provides more functionrelated information. However, due to the manual work involved, SCOP is not updated as frequently as the others (as of June, 2003, it was last updated for the PDB release on March 1, 2003), whereas FSSP and CATH follow the PDB updates closely. SCOP is used here as an example to show the features of structure-based families. SCOP can be accessed through its home server in the UK (http://scop.mrc-lmb.cam.ac.uk/scop/). It is also widely mirrored around the world, including: http://pdb.wehi.edu.au/scop/ (Australia) http://mdl.ipc.pku.edu.cn/scop/ (China) http://www.cdfd.org.in:5555/scop/ (India) http://pdb.weizmann.ac.il/scop/ (Israel) http://loki.polito.it/scop/ (Italy) http://scop.protres.ru/ (Russia) http://scop.bic.nus.edu.sg/ (Singapore) http://scop.life.nthu.edu.tw/ (Taiwan) http://scop.berkeley.edu/ (USA) Protein Databases on the Internet
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SCOP describes the hierarchical relationship among proteins through the major levels of (homologous) family, superfamily, and fold. Proteins are clustered together into a (homologous) family if they have significant sequence similarity. Different families that have low sequence similarity but whose structural and functional features suggest a common evolutionary origin are placed together in a superfamily. Different superfamilies are categorized into a fold if they have the same major secondary structures in the same arrangement and with the same topological connections (the peripheral elements of secondary structure and turn regions may differ in size and conformation). Two superfamilies in the same fold may not have a common evolutionary origin. Their structural similarities may arise from the physics and chemistry of proteins favoring certain packing arrangements and chain topologies (Murzin et al., 1995). Figure 2.6.3 shows the SCOP interface using an example of protein 1gox in the PDB. Databases for Function-Based Protein Families There are various protein functional families classified from different perspectives. The ENZYME data bank (Bairoch, 1993) contains the following data for each enzyme: EC number, recommended name, alternative names, catalytic activity, cofactors, pointers to the SwissProt entry, and pointers to any disease associated with a deficiency of the enzyme. PROCAT is a database of three-dimensional enzyme active site templates (Wallace et al., 1996). PDD (Protein Disease Database; Lemkin et al., 1995; Merril et al., 1995) correlates diseases with proteins observable in serum, urine, and other common human body fluids based on biomedical literature. There are also a growing number of databases dedicated to special types of proteins, such as antibodies, G-protein-coupled receptors, HIV proteases, glycoproteins, and RNases, as shown in Table 2.6.1. OTHER DATABASES Protein Binding Databases Protein binding includes protein-substrate docking and protein-protein association. ReLiBase (Hendlich, 1998) is a database system for analyzing receptor-ligand complexes in the PDB. DIP (Database of Interacting Proteins) records protein pairs that are known to bind with each other. The information in DIP may provide information related to signaling pathways, multiple interactions, and complex systems. Protein Energetics Databases There are few databases for protein energetics, due to the low-throughput nature of the data source. One useful energetics database can be found in ProTherm (Thermodynamic Database for Proteins and Mutants; Gromiha et al., 1999). It contains thermodynamic data on mutations, including Gibbs free energy, enthalpy, heat capacity, and transition temperature. These data are important for understanding the structure and stability of proteins. Bibliographic Databases Searching for protein information through traditional bibliographic databases, such as MEDLINE or Grateful Med, can be rewarding. In addition, some bibliographic reference databases dedicated to proteins may provide certain information more directly. For example, SeqAnalRef stores papers dealing with sequence analysis.
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Figure 2.6.3 An example of the SCOP interface when searching the structure of 1gox in the PDB.
COMBINED DATABASES Introduction By integrating different types of protein databases together, a database of databases (or a data warehouse) can be built. Such combined databases not only serve as “one-stop shopping,” but also provide cross-references between entries in different databases. Two combined databases, Entrez and SRS, have been very successful. Entrez Entrez (Schuler et al., 1996) is a combined database consisting of literature, protein sequence and structure, nucleotide sequence, and taxonomy. Different types of information are interconnected through the grouping of sequences/structures and references by computed similarity scores. Entrez can be used through a variety of media, including CD-ROM, a custom Graphical Interface client, a World Wide Web browser, a command line browser (CLEVER), and the National Center for Biotechnology Information’s (NCBI’s) toolkit written in C. SRS SRS (Sequence Retrieval System; Etzold et al., 1996) is the most comprehensive database for molecular biology. The home server at http://srs.ebi.ac.uk supports 264 biological databases (as of March, 2003), including almost all the major protein/genetic databases. As an indexing system, it provides fast access to different databases through searches by sequence or by key words from various data fields. SRS also builds indices using cross-references between databases. An entry from one database can be linked to other databases that contain the entry. However, it should be noted that the contents of SRS may Protein Databases on the Internet
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lag behind the other databases in updating (i.e., some new entries in the original databases may not be included in SRS). SUMMARY This unit reviews several major protein databases on the Internet, and shows what kind of information users can expect from protein databases. Although all technical procedures cannot be described here, most of the protein databases are easy to use and provide detailed on-line manuals so that even users with little computer skill can learn them quickly. Protein databases may not always be easily accessible or usable through the Internet. Sometimes a database server may be down or the Internet connection may be interrupted. Some structures or image files are very large (several megabytes), and the download time may be long. It can be helpful to use a mirror site of the database at a close location in order to accelerate the access speed. For a frequent user, it may be worthwhile to install the database on a local machine. On the other hand, it must be kept in mind that a mirror site or a local copy may contain an older version of the database than the one on the home server. It is important to assess the quality of the data. There are three types of data in protein databases. (1) Experimental data are generally very reliable. However, some entries may contain errors (e.g., some protein sequences) or may be based on low-resolution data (e.g., some protein structures determined by NMR). (2) Annotation data uses computational techniques on experimental data, for example, secondary structure assignment and domain partition in structure. These data depend on the quality of the experimental data and the computational methods used. Different methods may yield different results. (3) Prediction data includes, for example, sequence domain parsing and three-dimensional structure prediction. No matter how good the method, the results are still predictions and should be subjected to experimental verification. In addition, different methods typically give different predictions. In summary, caution is needed when using the data from databases to draw a conclusion. It is worthwhile to check the same type of data from different databases and compare them. It is sometimes necessary to use additional computational tools (e.g., tools to assess the quality of a structure) for further analysis. LITERATURE CITED Altschul, S.F., Madden, T.L., Schaffer, A.A., Zhang, J., Zhang, Z., Miller, W., and Lipman, D.J. 1997. Gapped BLAST and PSI-BLAST: A new generation of protein database search programs. Nucl. Acids Res. 25:3389-3402. Attwood, T.K., Flower, D.R., Lewis, A.P., Mabey, J.E., Morgan, S.R., Scordis, P., Selley, J., and Wright, W. 1999. PRINTS prepares for the new millennium. Nucl. Acids Res. 27:220-225. Bairoch, A. 1993. The ENZYME data bank. Nucl. Acids Res. 21:3155-3156. Bairoch, A. and Apweiler, R. 1999. The SwissProt protein sequence data bank and its supplement TrEMBL in 1999. Nucl. Acids Res. 27:49-54. Barker, W.C., Garavelli, J.S., McGarvey, P.B., Marzec, C.R., Orcutt, B.C., Srinivasarao, G.Y., Yeh, L.L., Ledley, R.S., Mewes, H., Pfeiffer, F., Tsugita, A., and Wu, C. 1999. The PIR-international protein sequence database. Nucl. Acids Res. 27:39-42. Bateman, A., Birney, E., Durbin, R., Eddy, S.R., Finn, F.D., and Sonnhammer, E.L.L. 1999. Pfam 3.1: 1313 multiple alignments match the majority of proteins. Nucl. Acids Res. 27:260-262. Benson, D.A., Boguski, M.S., Lipman, D.J., Ostell, J., Ouellette, B.F., Rapp, B.A., and Wheeler, D.L. 1999. Genbank. Nucl. Acids Res. 27:12-17. Bernstein, F.C., Koetzle, T.F., Williams, G.J.B., Meyer, E.F., Brice, M.D., Rodgers, J.R., Kennard, O., Shimanouchi, T., and Tasumi, M. 1977. The protein data bank: A computer based archival file for macromolecular structures. J. Mol. Biol. 112:535-542. Bourne, P., Berman, H., Watenpaugh, K., Westbrook, J., and Fitzgerald, P. 1997. The macromolecular crystallographic information file (mmCIF). Methods Enzymol. 277:571-590.
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Corpet, F., Gouzy, J., and Kahn, D. 1999. Recent improvements of the ProDom database of protein domain families. Nucl. Acids Res. 27:263-267. Etzold, T., Ulyanov, A., and Argos, P. 1996. SRS: Information retrieval system for molecular biology data banks. Methods Enzymol. 266:114-128. Gerstein, M. and Krebs, W. 1998. A database of macromolecular motions. Nucl. Acids Res. 26:4280-4290. Gibrat, J.F., Madej, T., and Bryant, S.H. 1996. Surprising similarities in structure comparison. Curr. Opinion Struct. Biol. 6:377-385. Gromiha, M.M., An, J., Kono, H., Oobatake, M., Uedaira, H., and Sarai, A. 1999. Protherm: Thermodynamic database for proteins and mutants. Nucl. Acids Res. 27:286-288. Hendlich, M. 1998. Databases for protein-ligand complexes. Acta Crystallogr., Sect. D 1:1178-1182. Heniko, J.G., Heniko, S., and Pietrokovski, S. 1999. New features of the blocks database servers. Nucl. Acids Res. 27:226-228. Hofmann, K., Bucher, P., Falquet, L., and Bairoch, A. 1999. The PROSITE database, its status in 1999. Nucl. Acids Res. 27:215-219. Holm, L. and Sander, C. 1996. Mapping the protein universe. Science 273:595-602. Kraulis, P. 1991. MOLSCRIPT—a program to produce both detailed and schematic plots of protein structures. J. Appl. Crystallogr. 24:946-950. Laskowski, R.A., Hutchinson, E.G., Michie, A.D., Wallace, A.C., Jones, M.L., and Thornton, J.M. 1997. PDBsum: A web-based database of summaries and analyses of all PDB structures. Trends Biochem. Sci. 22:488-490. Lemkin, P.F., Orr, G.A., Goldstein, M.P., Creed, G.J., Myrick, J.E., and Merril, C.R. 1995. The protein disease database of human body fluids: I. Computer methods and data issues. Appl. Theor. Electrophor. 5:55-72. Liang, J., Edelsbrunner, H., and Woodward, C. 1998. Anatomy of protein pockets and cavities: Measurement of binding site geometry and implications for ligand design. Protein Science 7:1884-1897. Marchler-Bauer, A., Addess, K.J., Chappey, C., Geer, L., Madej, T., Matsuo, Y., Wang, Y., and Bryant, S.H. 1999. MMDB: Entrez’s 3D structure database. Nucl. Acids Res. 27:240-243. Merril, C.R., Goldstein, M.P., Myrick, J.E., Creed, G.J., and Lemkin, P.F. 1995. The protein disease database of human body fluids: I. Rationale for the development of this database. Appl. Theor. Electrophor. 5:49-54. Mural, R.J., Parang, M., Shah, M., Snoddy, J., and Uberbacher, E.C. 1999. The Genome Channel: A browser to a uniform first-pass annotation of genomic DNA. Trends Genet. 15:38-39. Murvai, J., Vlahovicek, K., Barta, E., Szepesvari, C., Acatrinei, C., and Pongor, S. 1999. The SBASE protein domain library, release 6.0: A collection of annotated protein sequence segments. Nucl. Acids Res. 27:257-259. Murzin, A.G., Brenner, S.E., Hubbard, T., and Chothia, C. 1995. SCOP: A structural classification of proteins database for the investigation of sequences and structures. J. Mol. Biol. 247:536-540. Nayal, M., Hitz, B.C., and Honig, B. 1999. GRASS: A server for the graphical representation and analysis of structures. Protein Sci. 8:676-679. Ogata, H., Goto, S., Sato, K., Fujibuchi, W., Bono, H., and Kanehisa, M. 1999. KEGG: Kyoto encyclopedia of genes and genomes. Nucl. Acids Res. 27:29-34. Orengo, C.A., Michie, A.D., Jones, D.T., Swindells, M.B., and Thornton, J.M. 1997. CATH—a hierarchic classification of protein domain structures. Structure 5:1093-1108. Rebhan, M., Chalifa-Caspi, V., Prilusky, J., and Lancet, D. 1998. GeneCards: A novel functional genomics compendium with automated data mining and query reformulation support. Bioinformatics 14:656-664. Rodriguez-Tom, P., Stoehr, P.J., Cameron, G.N., and Flores, T.P. 1996. The European Bioinformatics Institute (EBI) databases. Nucl. Acids Res. 24:6-13. Sayle, R.A. and Milner-White, E.J. 1995. RASMOL: Biomolecular graphics for all. Trends Biochem. Sci. 20:374-376. Schuler, G.D., Epstein, J.A., Ohkawa, H., and Kans, J.A. 1996. Entrez: Molecular biology database and retrieval system. Methods Enzymol. 266:141-162. Shindyalov, I.N. and Bourne, P.E. 1995. WPDB: A PC-based tool for analyzing protein structure. J. Appl. Crystallogr. 28:847-852. Shindyalov, I.N. and Bourne, P.E. 1998. Protein structure alignment by incremental combinatorial extension (CE) of the optimal path. Protein Eng. 11:739-747. Shnel, J. 1996. Image library of biological macromolecules. Comput. Appl. Biosci. 12:227-229. Protein Databases on the Internet
Siddiqui, A.S. and Barton, G.J. 1995. Continuous and discontinuous domains: An algorithm for the automatic generation of reliable protein domain definitions. Protein Sci. 4:872-884.
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Sobolev, V., Sorokine, A., Prilusky, J., Abola, E.E., and Edelman, M. 1999. Automated analysis of interatomic contacts in proteins. Bioinformatics. 15:327-332 States, D.J., Harris, N.L., and Hunter, L. 1993. Computationally efficient representation of classes in protein sequence megaclassification. Proc. Intel. Syst. Mol. Biol. 1:387-394. Tatusov, R.L., Koonin, E.V., and Lipman, D.J. 1997. A genomic perspective on protein families. Science 278:631-637. University of Wisconsin. 1999. BioMagResBank. University of Wisconsin, Madison, Wis. Vriend, G. 1990. WHAT IF: A molecular modelling and drug design program. J. Mol. Graphics 8:52-56. Wallace, A.C., Laskowski, R.A., and Thornton, J.M. 1996. Derivation of 3D coordinate templates for searching structural databases: Application to Ser-His-Asp catalytic triads in the serine proteinases and lipases. Protein Sci. 5:1001-1013. Wu, C., Shivakumar, S., and Huang, H. 1999. Proclass protein family database. Nucl. Acids Res. 27:272-274.
INTERNET RESOURCES The Web addresses of the databases mentioned in this unit are listed in Table 2.6.1. Readers can find more protein databases and their related tools in the following Web pages, which collect a large number of useful links. http://compbio.ornl.gov/structure/resource/ Oak Ridge National Laboratory’s resources of protein modeling tools. http://www.public.iastate.edu/~pedro/research_tools.html Pedro’s biomolecular research tools. http://www.expasy.org/alinks.html Amos’ WWW links page.
Contributed by Dong Xu and Ying Xu Oak Ridge National Laboratory Oak Ridge, Tennessee
The authors thank Drs. Edward C. Uberbacher, Michael A. Unseren, Jay Snoddy, and Gwo-liang Chen for helpful discussions. This work is supported by the Office of Biological and Environmental Research, U.S. Department of Energy, under Contract DE-AC05-00OR22725, managed by UT-Battelle, LLC.
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Protein Tertiary Structure Prediction
UNIT 2.7
Predicting the three-dimensional structures of proteins from their amino acid sequences using computational methods is a very important and challenging problem in contemporary molecular biology. The three-dimensional structures of proteins not only define their biological functions, but also provide a key to engineering proteins and to designing drugs that target proteins related to disease. Protein tertiary structure prediction provides timely tools on account of rapid progress being made in genome sequencing, which is proceeding at a rate many times faster than experimental determination of protein structures. In conjunction with supporting experimental data, these tools can provide valuable insights into protein structure and function. In particular, predicted structures often give experimentalists some direction for further studies. Protein tertiary structure prediction is not only important, but also becomes more and more practical. Many nontrivial structure predictions (Nilges and Brünger, 1993; Hu et al., 1995; Madej et al., 1995) obtained prior to experimental structures turned out to be fairly accurate. Most notably, the success of protein structure prediction has been demonstrated in community-wide experiments on the Critical Assessment of Techniques for Protein Structure Prediction (CASP; see CASP, 1995, 1997, 1999). In each of the CASP experiments, crystallographers and NMR spectroscopists were solicited for the names of proteins whose structures were likely to be solved before a given deadline. The sequences of these target proteins were made available, and predictors submitted their models by the deadline. These blind predictions were then assessed through comparison with the experimental structures. It has been shown that many predictions in CASP have good qualities. There are three types of tertiary structure predictions: homology or comparative modeling, fold recognition, and ab initio structure prediction. Homology modeling constructs the coordinates of all the atoms in a query protein based on sequence alignment between the query protein and another protein of known three-dimensional structure. Fold recognition identifies a suitable fold for the query sequence from a structure library and provides an alignment between the query protein and the fold. It consists of two approaches, the sequence profile approach and the sequence-structure comparison (threading) approach. The ab initio structure prediction attempts to predict protein structure based directly on physicochemical principles. Homology modeling has been widely used. It can provide relatively reliable atomic coordinates with a low root mean square deviation (RMSD) between a model and a high-quality experimental structure. Fold recognition, which often generates useful backbone structures, also starts to gain ground. The ab initio prediction is still far from general application, although a few isolated successful predictions have been reported. The boundaries between the different types of predictions are becoming blurred as researchers start to combine them together. For example, one can use the alignment derived from fold recognition in homology modeling, or assemble partial structures predicted by threading in ab initio prediction. Both homology modeling and fold recognition rest on the foundation that the three-dimensional structures of proteins have been better conserved during evolution than their sequences. The relationship between a query protein and its template in a structure database can be classified at different hierarchical levels according to their structural and evolutionary relationship. A widely used classification consists of family, superfamily, and fold (Murzin et al., 1995). A query protein and its template are clustered into a family if they have clear evolutionary relationship with significant sequence identity between them (often 25% or higher). If two proteins have low sequence identity, but their structural Contributed by Dong Xu and Ying Xu Current Protocols in Protein Science (2000) 2.7.1-2.7.17 Copyright © 2000 by John Wiley & Sons, Inc.
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and functional features suggest that a common evolutionary origin is probable, then they are placed in the same superfamily. If two proteins have the same major secondary structures in the same arrangement and with the same topological connections, but they have no significant sequence similarity and may not have a common evolutionary origin, they are defined as having the same fold. It is estimated that the number of folds in all genomes ranges from 650 to 15,000, or less than one thousandth of the number of proteins (Wang, 1998). Different methods of structure predictions are better suited to proteins with different relationship to their templates. When a query protein and its template belong to the same family and they have high sequence identity to produce an unambiguous sequence alignment, homology modeling is the best choice and it typically produces a high-quality model including sidechain atoms. About 17% of identified genes fall into this category (Sanchez and Sali, 1998). The sequence profile approach is suitable for the following two cases: (1) a query protein and its template belong to the same family, but they do not have sufficient sequence identity to produce an unambiguous sequence alignment, or (2) they belong to the same superfamily, and the multiple-sequence alignment of the query protein has a strong pattern. For these two cases, which are also referred to as “remote homologs,” the sequence profile approach typically identifies the structure template with high confidence and good alignment. It extends the genome-wide coverage from 17% to ∼30% to 40% (Gerstein, 1998). The threading approach becomes the major tool for the following two cases: (1) a query protein and its template belong to the same superfamily, but the query protein has few proteins with significant sequence similarities in sequence databases, or the multiple-sequence alignment of the query protein has a weak pattern, or (2) they belong to the same fold, and threading gives a good confidence level for the prediction. This category currently accounts for an additional ∼10% to 20% of identified genes (Jones, 1999). If a query protein and its template belong to the same fold, but threading cannot recognize the template with a good confidence level, or the template does not exist in the structure database (i.e., the query protein is a new fold), it can only be predicted by ab initio methods. In total, structure predictions for about half of the genes do not have to use ab initio prediction and can provide useful information reliably. This number is growing as more and more structures are being solved. This unit provides an overview for homology modeling, fold recognition, and ab initio prediction. It introduces several major protein structure prediction tools. For each tool, it addresses (1) the method, (2) the technical protocol, (3) the expected result, and (4) the confidence level of the prediction. The Web addresses of the major structure prediction tools are listed in Table 2.7.1. The reader can find more detailed information about the tools in their Web pages or manuals. The examples used in this unit are all from the third CASP experiment (CASP-3) held in 1998. This blind test provides an up-to-date benchmark regarding the results that a user can expect from these prediction tools. Among the examples used, the models for t0053, t0067, t0068, and t0074 are from the authors’ own predictions submitted to CASP-3 (Xu et al., 1999). All the results are based on the programs and databases as of August, 1999. HOMOLOGY MODELING
Protein Tertiary Structure Prediction
A homology modeling method typically consists of three steps: (1) identification of protein templates with known three-dimensional structures and production of an alignment between the query sequence and its templates, (2) building a model for the query protein based on its alignment with the template structures, and (3) evaluation of the quality of the model.
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Table 2.7.1
Selected Protein Structure Prediction Tools
Program
Web site
Typea
Sequence alignment: ALIGN
http://www2.igh.cnrs.fr/bin/align-guess.cgi
Server
BLAST FASTA
http://www.ncbi.nlm.nih.gov/BLAST/ http://www.embl-heidelberg.de/cgi/fasta-wrapper-free
Server Server
KESTREL SSEARCH
http://www.cse.ucsc.edu/research/kestrel/ http://vega.igh.cnrs.fr/bin/ssearch-guess.cgi
Server Server
Homology modeling: COMPOSER
http://www-cryst.bioc.cam.ac.uk
SGI executable
CONGEN
http://www.tripos.com/software/composer.html http://www.congenomics.com/congen/congen toc.html
SGI module (GUI) Executables
CPHmodels DRAGON
http://www.cbs.dtu.dk/services/CPHmodels/ http://www.nimr.mrc.ac.uk/∼mathbio/a-aszodi/dragon.html
Server SGI executable
LOOK MODELLER
http://www.mag.com/products/look.html http://guitar.rockefeller.edu/modeller/
Module (GUI) Executables
SwissModel
http://www.msi.com/solutions/products/insight/modules/Modeler.html SGI module (GUI) Server http://www.expasy.ch/swissmod/SWISS-MODEL.html
WHAT IF
http://www.sander.embl-heidelberg.de/whatif/
Executables
http://www.ncbi.nlm.nih.gov/BLAST/ http://www.cse.ucsc.edu/research/compbio/sam.html
Server Server
Sequence profile methods: PSI-BLAST SAM-T99
Singleton-potential threading: 123D http://www-lmmb.ncifcrf.gov/∼nicka/123D.html
Server
SAS TOPITS
http://www.biochem.ucl.ac.uk/bsm/sas/ http://dodo.cpmc.columbia.edu/predictprotein/
Server Server
UCLA-DOE
http://www.doe-mbi.ucla.edu/people/frsvr/frsvr.html
Server
Pairwise-potential threading: NCBI Package PROFIT
http://www.ncbi.nlm.nih.gov/Structure/ http://lore.came.sbg.ac.at
Executables Executables
PROSPECT THREADER
http://compbio.ornl.gov/structure/prospect/ http://globin.bio.warwick.ac.uk/∼jones/threader.html
Executables Executables
ToPLign
http://cartan.gmd.de/ToPLign.html
Server
aExecutables without a specified machine type can be run on multiple platforms. Abbreviations: GUI, Graphic User Interface; SGI, Silicon Graphic,
Inc.
Template Search and Alignment Conventional homology modeling requires that the sequence similarity between a query sequence and its template sequence be significant, so that using a pairwise sequence-sequence alignment can identify the template and produce an unambiguous alignment. The most widely used sequence-sequence alignment tool is BLAST (UNIT 2.5), which is very fast. Using BLAST to search against the Protein Data Bank (PDB; Bernstein et al., 1977) is probably the best starting point for homology modeling. If the identified templates or alignments have any uncertainty, more sensitive but slower methods (e.g., FASTA; Pearson and Lipman, 1988) can be used. An even slower but much more sensitive method
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is the Smith-Waterman alignment (Smith and Waterman, 1981), which has been implemented in SSEARCH and KESTREL (Hughey, 1996). Templates and alignments using fold recognition methods, as described in the following two sections, can also be used for homology modeling. In fact, it has been shown in CASP-3 that profile-based alignment methods and threading methods can produce more accurate alignments than single-sequence alignment methods even when the query protein and its template have some sequence similarity (CASP, 1999). It is sometimes important to find as many templates as possible with a multiple-sequence alignment. Template search and alignment are essential for the correctness and the quality of a homology model. Homology modeling programs always generate a structure for any query sequence using the conformation of the template structures and the alignments between the query protein and its templates. If the templates or the alignments are incorrect, the output model will certainly be a result of “garbage in, garbage out.” Building Atomic Models Different homology modeling methods use different ways to construct a three-dimensional model from given templates and alignments. Some automated homology modeling servers, e.g., SwissModel (UNIT 2.8; Peitsch, 1996) and CPHmodels (Lund et al., 1997), provide interfaces for submitting a sequence and obtaining the model interactively or through e-mail. The use of SwissModel is described in detail in UNIT 2.8. These servers are fast and easy to use. The program WHAT IF (Vriend, 1990) provides the option to construct a crude model quickly or to build a structure using a better, but much slower, method (several hours for a large protein, e.g., >300 amino acids). Another program is COMPOSER (Srinivasan and Blundell, 1993), which has a free academic version and is also integrated into the commercial molecular modeling package SYBYL (Tripos, 1999). It has a specific tool for dealing with loop regions, which contain gaps in the alignment. COMPOSER under SYBYL also provides an interactive Graphic User Interface (GUI) for model building, which allows a user to edit at each step. The most widely used homology modeling program is MODELLER (Sali and Blundell, 1993), which has a free academic version as well as a commercial version integrated into the molecular modeling package INSIGHT (Molecular Simulations, 1998). The model structure starts with an extended strand, and then folds into a compact one by satisfying spatial restraints derived from the alignment between the query sequence and its templates of known structures. In particular, it tries to preserve main chain dihedral angles or hydrogen bonding features from the template structures. Meanwhile, MODELLER uses physical force fields to prevent interatomic clashes. In loop regions with gaps in the alignment, MODELLER uses statistical information derived from the alignment of many proteins of known three-dimensional structure. The final three-dimensional model is obtained by optimization through conjugate gradients and molecular dynamics with simulated annealing. MODELLER provides a wide range of options for structure construction and refinement.
Protein Tertiary Structure Prediction
t0074 (the second EH domain of eps15), which was predicted by the authors, can be used as a CASP-3 example to show some basic features of MODELLER. Constructing a model requires at least three input files: the three-dimensional coordinate file for the template, the alignment file between the query protein and the template, and a script file to run MODELLER. The template found for t0074 is the PDB entry 1ahr (calcium-binding protein), which has three-dimensional coordinates recorded in the file 1ahr.atm (the same file as 1ahr.pdb in the PDB database). The alignment between t0074 and 1ahr is saved in the file 1ahr.ali using the PIR database format as follows:
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>P1;t0074 sequence:t0074: 1 : : 72 : : casp-t0074: : PWAVKPE--DKAKYDAIFDSLSP-VNGFLSGDKVKPVLLNS--KLPVDILGRVWELSDIDHDGML DRDEFAVAMFLV* >P1;1ahr structureX:1ahr: 73 : : 148 : : template ARKMK-D--SEEEIREAFRVFDKDGNGFISAAELRHVMTNLGEKLTDEEVDEMIREADIDGDGQV NYEEFVTMMTSK*
where the first line of each entry specifies the protein code after the “>P1", the second line describes the type, the name, and the residue ranges, and the third line provides the sequence. The following is a sample script file, run.top, for building the model for the query protein t0074: INCLUDE
# Include predefined routines
SET TOPLIB = ’${LIB}/top_allh.lib’ # topology library SET PARLIB = ’${LIB}/par.lib’ # parameters library SET HYDROGEN_IO = ’ON’ SET HETATM_IO = ’ON’
# include hydrogen atoms # include HETATM entries in template
SET ALNFILE = ’1ahr.ali’ SET KNOWNS = ’1ahr’ SET SEQUENCE = ’t0074’
# alignment filename # template name # target sequence name
CALL ROUTINE = ’model’
# do homology modeling
The text following “#” is commentary that will be ignored when the script is run. To run the script, simply type mod run.top
It typically takes several minutes to an hour to run a MODELLER job, depending on the protein sizes and the machine used. After the job mod run.top is finished, a file (t0074.B99990001) containing the three-dimensional coordinates for the atomic model of t0074 is generated, as shown in Fig. 2.7.1A. MODELLER can easily incorporate more sophisticated modeling protocols. For example, several more templates can be used at the same time by changing the SET KNOWNS entry in run.top. For example, to add three templates in addition to “1ahr,” SET KNOWNS would be changed to: SET KNOWNS = ’1ahr’ ’4cln’ ’1cmg’ ’1cdla’
and the alignments of all four templates with the query sequence would be provided in 1ahr.ali. MODELLER can also use specified information (such as NMR data, secondary structure types, and disulfide bonds) as spatial restraints. The following shows an example for building a model with the constraint of disulfide bonds between certain residues: SUBROUTINE ROUTINE = ’special_patches’ # A disulfide between residues 5 and 60: PATCH RESIDUE_TYPE = ’DISU’, RESIDUE_IDS = ’5’ # A disulfide between residues 22 and 38: PATCH RESIDUE_TYPE = ’DISU’, RESIDUE_IDS = ’22’ RETURN END_SUBROUTINE
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A
B
Figure 2.7.1 A comparison between the predicted structure using MODELLER (thick lines) and the experimental structure (thin lines) for CASP-3 targets t0074 (A) and t0068 (B). This figure was made using visual molecular dynamics (VMD; Humphrey et al., 1996).
Model Assessment and Refinement The quality of a model depends primarily on the sequence identity between the query protein and the template. The higher the sequence identity, the more accurate a structure homology modeling can provide. As shown in Figure 2.7.1A, there are some conformational differences between the model and the experimental structure, mainly due to the structural differences between the template and the query protein. In this case, the sequence identity between the query protein and the template is only 15%. For another CASP-3 target, t0068, the sequence identity between the query protein and the template is 25%. As shown in Figure 2.7.1B, the model superimposes the experimental structure better than that of t0074. For high sequence identity, homology modeling often produces models with an all-atom RMSD of <2 Å between the model and the experimental structure (similar to the typical quality of an NMR structure). Another challenge in homology modeling is construction of regions with large alignment gaps. Although loops with short alignment gaps can often be modeled successfully, insertions of ∼8 residues or more in the query sequence usually cannot be modeled reliably.
Protein Tertiary Structure Prediction
It is important to assess models using other computational tools. When the alignment is correct, a homology model often has good stereochemistry and interresidue packing with a smaller backbone RMSD to the experimental structure than the template used. When the alignment has errors or long gaps, it is likely to generate errors, including (1) bad backbone conformations, e.g., artificial cis peptide bonds; (2) poor stereochemistry, e.g., unwanted D-amino residues; or (3) unfavorable interresidue packing. These errors can be detected using programs such as WHAT IF (Vriend, 1990) and PROCHECK (Laskowski et al., 1993). The overall quality of a model can be further assessed by PROVE (PROtein
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Volume Evaluator; Pontius et al., 1996), which checks the departures of the assessed structure from the standard atomic volumes in high-quality experimental structures. If errors are found, the alignment can be adjusted and the model can be rebuilt. Another method is to generate multiple models and find the best model with the least errors. It may be necessary to repeat the process of alignment, model construction, and assessment until a satisfactory model is obtained. SEQUENCE PROFILE METHODS Sequence profile methods detect remote homologs based on a profile in multiple-sequence alignments. Pairwise sequence alignments requires high sequence identity (typically 25% or more) for reliable results. A profile of close homologs can significantly increase the underlying signal while reducing noise, and hence a much lower sequence identity (as low as 15%) is needed for a high-quality alignment. As a huge number of protein sequences are being solved, sequence profile methods become more and more useful for finding a remote homolog with known structure to use as the template for a query protein. There are two successful tools that use sequence profile methods: PSI-BLAST (Altschul et al., 1997; UNIT 2.5) and SAM-T98 (for Sequence Alignment Modeling system; Karplus et al., 1998; Park et al., 1998). The two programs use different methods, as described below. PSI-BLAST is very fast, and the results of each iteration can be returned from the Web server in seconds. It also provides a statistical significance assessment value (EXPECT) for reporting the number of matches that would be expected by chance, given the profile used at the current iteration, according to a stochastic model. PSI-BLAST also allows users to select parameters and proteins for building sequence profiles interactively. On the other hand, SAM-T98 is slower but more sensitive. Compared to PSI-BLAST, SAM-T98 detects more remote homologs and generates fewer false positives at any level of true positives (Park et al., 1998). A statistical significance assessment like the PSIBLAST’s EXPECT value is not available in SAM-T98, but has been added in the newly released SAM-T99. Unlike PSI-BLAST, SAM-T98 and SAM-T99 as e-mail servers do not have the ability to interactively edit parameters and proteins for building sequence profiles during a search process. Users can search using both PSI-BLAST and SAM-T99 and compare the results when any uncertainty exists. PSI-BLAST The Position-Specific Iterated BLAST (PSI-BLAST) program (Altschul et al., 1997) searches the protein database using a position-specific score matrix derived from a multiple-sequence alignment on similar sequences found by BLAST. The search is carried out iteratively until a satisfactory match is found or the search is converged (typically a total of three to four iterations). At each iteration, the position-specific score matrix is updated using the new sequences in addition to the sequences found in previous iterations. One can use PSI-BLAST with a default setting for parameters. However, more remote homologs can be found by trying different settings. CASP-3 target t0081 (methylglyoxal synthase) can be used as an example. PSI-BLAST can be run via the Web server at http://www.ncbi.nlm.nih.gov/cgi-bin/BLAST/ nph_psi_blast. After submitting the sequence for a search using the default setting, the returned result is shown in Figure 2.7.2. If a user continues to use the default in the second iteration, the search will be converged without hitting any new matches with known structure. However, if a user selects the five additional entries whose EXPECT values are <0.01 and then carries out the search using the default setting in subsequent iterations, a
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Figure 2.7.2 The first iteration result of PSI-BLAST for the CASP-3 target t0081. The entries above the “Run PSI-Blast iteration 2" button will be used by the default (with the EXPECT value, listed at the end of each entry, <0.001). The entries below the button are optional to use in the search of the next iteration.
template with known structure will be found at the fourth iteration. The template is carbamoyl phosphate synthetase (chain B of 1jdb in the PDB), whose alignment to t0081 is shown as follows: Score = 74.7 bits (181), Expect = 3e-13 Identities = 23/147 (15%), Positives = 46/147 (30%), Gaps = 9/147 (6%) Query: 2
Sbjct: 990
ELTTRTLPARKHIALVAHDHCKQMLMSWVERHQPLLEQHVLYATGTTGNLISRATGMNVN + T+ L + K+ ++ + L + L AT T ++ A G+N LGSNSTMKKHGRALLSVREGDKERVVDLAAKL--LKQGFELDATHGTAIVLGEA-GINPR AMLSGPMGGDQQVGALISEGKIDVLIFFWDPLNAVPHDPDVKALLRLATVWNIPVATNVA + G + I G+ +I A+ D + + R A + + T + LVNKVHEGRP-HIQDRIKNGEYTYIINTTSGRRAIE---DSRVIRRSALQYKVHYDTTLN
Query: 122
148
Sbjct: 933 Query: 62
TADFIIQSPHFNDAVDILIPDYQRYLA + N + Q A Sbjct: 1046 GG--FATAMALNADATEKVISVQEMHA
Protein Tertiary Structure Prediction
61 989 121 1045
1070
It turned out that the PDB entry 1jdb is the correct template for t0081, and that the above alignment basically agrees with the structure-structure alignment. This example shows that the profile derived from the multiple-sequence alignment can significantly increase the sensitivity and reliability of a sequence search. However, a user should be careful when interpreting the meaning of the EXPECT value (3 × 10−13 in this case). Because EXPECT values are based on the profile used at the current iteration, the validity of using a low EXPECT value as evidence for relationship of a template to the original query protein is contingent upon previous iterations being accurate (Altschul and Koonin, 1998). The significance level for the match between a hit sequence and the query protein is limited to the maximum EXPECT value of all the sequences selected along the search
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path. Hence, changing the default setting to include more proteins to build the profile can be tricky. If improper entries are selected in the iterations, a conflated profile may be built, and some resulting matches could give wrong templates even when their EXPECT values are low. In the example of t0081, the manually added five entries have relatively small EXPECT values; more importantly, they are all synthetases, which are functionally related to the query protein (methylglyoxal synthase). Using these heuristics may help to avoid a conflated profile when adding more entries. Hidden Markov Models Hidden Markov models (HMMs) detect remote homologs with known structures using the following steps (Krogh et al., 1994): (1) a standard sequence-based search to find homologs (not necessarily with known structures) for a query sequence; (2) construction of an HMM based on the alignments between the query sequence and its homologs to describe the position-dependent amino acid (including deletion and insertion) probability distributions; and (3) a search from a library of templates (sequences of proteins in the PDB) to match the constructed HMM model. Similarly, a separate HMM can be constructed for each template, and the query sequence can be scored against each model. Among several computer packages based on HMMs, the most widely used package for finding a structure template of a query protein is SAM-T98/SAM-T99 (Karplus et al., 1998; Park et al., 1998), a particular application of the Sequence Alignment and Modeling system (SAM; Hughey and Krogh, 1996). The SAM-T99 server for identifying templates in the PDB is at http://www. cse.ucsc.edu/research/compbio/sam.html. To run a search, a user needs to enter the returning e-mail address and the query sequence. The result will automatically be sent back via e-mail. The following are the templates that SAM-T99 found for the CASP-3 target t0083 (cyanase): % Sequence ID 1lliA 1lliB 1lrp 1lmb4 1lmb3 1lmdC 1lmdD 1lmdB 1lmdA 1b0nA
Length 92 92 92 92 92 236 236 236 236 111
Simple −17.10 −17.10 −16.51 − 16.51 −16.51 −16.12 −16.12 −16.12 −16.12 −14.99
Reverse −10.38 −10.38 −9.95 −9.95 −9.95 −9.61 −9.61 −9.61 −9.61 −9.21
X count
where simple and reverse are two different scores. Reverse scores are more reliable than simple scores, and the templates are sorted by reverse scores. The first hit, 1lliA, is a good template for t0083, and the SAM-T99 alignment between t0083 and 1lliA is basically correct, as shown in the following: ; 1 user-seq-28992.seqs(1) ; 2 1lliA 1 MIQSQINRNIRLDLADAIL....LSKAKKDLSFAEIADGTGLAEAFVTAALLGQQALPADAARLV 2 STKKKPLTQEQLEDARRLKaiyeKKKNELGLSQESLADKLGMGQSGIGALFNGINALNAYNAALL 1 GAKLDLDEDSILLLQMIPLRGCIDDRIPTDPTMYRFYEMLQVYGTTLKALVHEKFGDGIISAINF 2 AKILKVSVEEFSPSIARE--------------IYEMYEAVS-----------------------1 KLDVKKVADPEGGERAVITLDGKYLPTKPF 2 -----------------------------Quantitation of Protein Interactions
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THREADING Protein threading (Bowie et al., 1991; Godzik et al., 1992; Jones et al., 1992; Sippl and Weitckus, 1992; Bryant and Altschul, 1995; Alexandrov et al., 1996; Fischer et al., 1996; Xu et al., 1998; Crawford, 1999) is the most promising and practical tool for fold recognition as demonstrated in the CASP tests. The basic idea of threading can be summarized as follows. Given a query protein sequence, s, of unknown structure, threading searches all structure templates, T, to find the best fit for s. A threading requires four components (Smith et al., 1997): (1) a library, T, of representative three-dimensional protein structures for use as templates; (2) an energy function to describe the fitness between s and t, where t is a single template in T; (3) a threading algorithm to search for the lowest energy among the possible alignments for a given s-t pair; and (4) a criterion to estimate the confidence level of the predicted structure. The threading approach can be further subdivided into two categories: (1) singleton-potential threading, which only considers the preference of amino acids in the query sequence at single sites of the templates, and (2) pairwise-potential threading that uses the preference on pairs (or even triplets) of amino acids in the query sequence within a contact distance when they are aligned to a given structure. Pairwise-potential threading can also include the singleton energy. Both methods allow gaps in alignments, but the gaps in pairwise-potential threading are often assumed to be in noncore regions (loops and possibly edges of α helices and β sheets) to reduce the search space. Such an assumption is generally true in the structure-structure alignment. In general, singletonpotential threading is faster, whereas pairwise-potential threading is more sensitive in detecting the correct templates. The threading approach generally has a better chance of selecting the right template among the top hits in the ranking than does the sequence profile approach. However, it is difficult to tell whether or not the template of the query protein is in the threading database (i.e., the top hits may be false positive matches). Some templates with similar folds to the query protein may be missed mainly due to the limitation of the energy functions used. A key difficulty for threading is that the structure profile and the residue pairs derived from the template may not describe the corresponding information in the query protein, due to the structure difference between the two proteins, even when they share a common fold. This is a more serious problem in the fold category than in the superfamily category. In CASP-3, almost every protein in the superfamily category was predicted correctly by at least one threading program. However, few proteins in the fold category were predicted correctly by any method. Singleton-Potential Threading Singleton-potential threading constructs a one-dimensional structure profile for each residue position in a template structure using local three-dimensional environmental information such as secondary structure type, degree of environmental polarity, and the fraction of the residue surface accessible to solvent. The energy function is based on the compatibility of the 20 common amino acids for each position in the one-dimensional structure profile. The compatibility is derived from the statistics of the whole template database. Optimal one-dimensional alignments between a query sequence and a template can be determined by dynamic programming. The final template is selected according to the optimal score or its statistical significance. Singleton-potential threading can incorporate secondary structure predictions and position-dependent profiles based on multiple-sequence alignments into the energy function. Protein Tertiary Structure Prediction
Several servers are available for singleton-potential threading, e.g., 123D (Alexandrov et al., 1996), TOPITS (Rost, 1995), SAS (Milburn et al., 1998), and the UCLA-DOE Structure Prediction Server (Fischer and Eisenberg, 1996). The following example uses
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the 123D server, which is located at http://www-lmmb.ncifcrf.gov/∼nicka/run123D.html, and which returns the results interactively from the Web page. An improved version of 123D can be found at http://cartan.gmd.de/ToPLign.html, which sends back results via e-mail. The following output shows the best templates selected by 123D for the CASP-3 target t0079 (the MarA protein): PDB structure raw score 1mseCx 105 a.a. 352 1sra_x 151 a.a. 274 1rcb_x 129 a.a. 58 1yrnBx 78 a.a. 48 1tam_x 120 a.a. 31 1mmoGx 162 a.a. 28 1octCx 131 a.a. 17 1bip_x 122 a.a. 13 ...
Z-score 2.16 1.51 −0.28 −0.37 −0.51 −0.53 −0.62 −0.66
a.a. aligned 99 116 109 73 106 105 100 101
% identities 18 10 17 24 13 24 19 14
Each entry of the output shows the template name, size, raw threading score, Z-score, number of aligned residues, and the sequence identity of the optimal alignment. The Z-score attempts to show the statistical significance of the alignment, but it may not work well in some cases. The optimal alignment between t0079 and 1mseC (c-Myb DNA-binding domain) returned by 123D is as follows: 1mseCx 352 bbb aaaaaaaaaa aaaaaa aaaaaaaaaa MTMSRRNTDAITIHSILDWIEDNLESPLSLEKVSERSGYSKWHLQRMFKKETGHSLGQYI * .. * ::. : . *.: : * .:*.: : *: *. .*. MLIKG------------PWTKEEDQRVIKLVQ---KYGPKRWSV--IAKHLKGR-IGKQC aaaaaaaaaaaaa aa aaaa aaaaaa aaaaa aaaaaaaaaaaa aaaaaaaaa RSRKMTEIAQKLK-----ESNEPILY-LAERYGFESQQTLTRTFKNYFDVPPHKYRMTNM *.* . :.* *... *:* . :*.* :. . :.: :.. * .... ..* RERWHNHLNPEVKKTSWTEEEDRIIYQAHKRLG-NRWAEIAKLLPGRTDNAIKNHWNSTM aaaaa aaaaa aaaaaaaaaaaaaa aaaaaaaa aaaaaaaa QGESRFLHPLNHYNS :.: . RRK-----------V
Among the five lines in the alignment, the second and fourth lines are the query protein sequence and the template sequence, respectively. The first and fifth lines are the predicted secondary structures and the secondary structures of the template, respectively, where a and b are α helix and β sheet, respectively. The third line is the match between the query sequence residues and the template residues, indicating where the two alignment residues are identical (*), similar (:), or related (.). Compared to the experimental structure of t0079, the template 1mseC found by 123D is a correct fold, although its alignment contains apparent errors. Both t0079 and 1mseC are DNA-binding proteins with helixturn-helix motifs. Even though the alignment is not perfect, the prediction may still provide useful information for the query protein. Pairwise-Potential Threading Pairwise-potential threading considers the propensity of two amino acids to be within a specified distance using a score function compiled from a database of structures, typically in addition to the singleton energy. In the recent CASP-3, it was clear that the top performers for fold recognition were among the groups using pairwise-potential threading methods. Several pairwise-potential threading programs are available, including the NCBI Threading Package (Bryant and Lawrence, 1993), PROFIT (Sippl and Weitckus,
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1992), PROSPECT (Xu et al., 1998), and THREADER (Jones et al., 1992). PROSPECT is used below to demonstrate how pairwise-potential threading works. PROSPECT (PROtein Structure Prediction and Evaluation Computer Toolkit) guarantees to find the globally optimal alignments for a given energy function with pairwise interactions. The system runs efficiently when using a distance cutoff. Two sets of threading templates are available: protein chains (defined by an FSSP nonredundant set; Holm and Sander, 1996) and compact domains (defined by the DALI nonredundant domain library; Holm and Sander, 1998). The system allows a user to input experimental knowledge and results from other computational tools into the threading process as restraints. The restraints include disulfide bonds, a match between a residue on the query sequence and a residue on a template, the secondary structure prediction by the neural net program PHD (an automatic e-mail server at http://dodo.cpmc.columbia.edu/predictprotein/; Rost and Sander, 1993), and the position-dependent profile based on the multiple-sequence alignment using SAM (Hughey and Krogh, 1996). PROSPECT also provides a confidence assessment using the Z-score and the compactness of the model. It has an interface to MODELLER for generating atomic structures. PROSPECT can be run in a very simple mode, but includes a wide range of options available to advanced users. The detailed manual of PROSPECT can be found at http://compbio.ornl.gov/structure/prospect/. The CASP-3 target t0057 (the CbiK protein) is used here as an example. To run the program in a default mode, a user only needs to prepare a file, t0053.seq, for the sequence of the protein, and to run the command: prospect t0053.seq
After the program is finished, another file, t0053.sort, will be generated for the ranking of the templates, while a third file, t0053.out, will record alignments and other detailed information. One can find the template with the optimal score (1ak1) and its alignment. To obtain the atomic structure for the query protein based on the threading result, one can simply run: prospect -m t0053.out 1ak1
Advanced options can be used to refine the prediction and yield a better result. For example, the authors refined the alignment between t0053 and the template 1ak1 using the information of an active site. Through a detailed analysis, it was found that 1ak1 has an active site at His-183 (Al-Karadaghi et al., 1997). Searching the BLOCK database (Heniko and Heniko, 1994), His-145 of t0053 was identified as the corresponding active site. PROSPECT was then run using the constraint that His-145 of t0053 is aligned with His-183 of 1ak1. This requires a configuration file, t0053.conf, to include: ANCHOR: anchor: 145:183 EDN
The output gives the detailed energies and assessment: Threading between t0053.seq and 1ak1 ... The raw score of the alignment: −2112.5 (−2113) Mutation = −494.0; Singleton = −872.0; Pairwise = −177.0; InternalPair = 9.5; GapPenalty = 331.0; SSFit = 0.0; AnchorMatch = −910.0 (1-anchor); PairConstrn = 0.0 (0-pair). Protein Tertiary Structure Prediction
Z-score = 11.07 Normalized radius of gyration = 1.83
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where different energy terms are recorded. For example, Pairwise is the pairwise energy between cores, InternalPair is the pairwise energy within cores, SSFit is the fitness energy between the PHD prediction and the secondary structures assigned from the PROSPECT alignment, AnchorMatch is the contribution of matching the predefined alignments between the anchor residues on the query sequence and the positions on the template, and PairConstrn is the contribution of matching the predefined pairs between the residues on the query sequence. The index of normalized radius of gyration gives an estimate for the compactness of the aligned portion in threading. If the value is >3.0, the aligned portion is insufficiently compact, indicating a bad match unless the template is a multidomain protein. The final alignment is as follows, where matches between the two aligned residues are shown to be identical (|), similar (:), or related (.): The full alignment with 264 residues in the target sequence MKKALLVVSFGTSYHDTCEKNIVACERDLAASCPDRD-L--FRA-F--TSGMI-IRKLRQ | |||:..||.|.: . | |: : | : . |. : : . -KMGLLVMAYGTPYKEEDIERYYTHIRR--GRKPEPEMLQDLKDRYEAIGGISPLAQITE RDGIDIDTPLQAL-QKLAAQGYQDV------AIQSLHII--NG-DEYEKIV--------.:: | : :. | . .. . .| | || QQAHNLEQHLNEIQDEITFKAYIGLKHIEPFIEDAVAEMHKDGITEAVSIVLAPHFSTFS --REVQLLRPLFTRL-TLGVPLLSSHNDYVQLMQALRQQMPSLR--------QTEKVVFM : :| | .. . | | : : . .. VQSYNKRAKEEAEKLGGLTITSVESWYDEPKFVTYWVDRVKETYASMPEDERENAMLIVS GH-------GASHHAFA-AYACLDHMMTAQRFP-ARVGAV------ES-Y-PEVDILIDS | . . || :. |:| | AHSLPEKIKEFGDPYPDQLHESAKLIAEGAGVSEYAVGWQSEGNTPDPWLGPDVQDLTRD L-RDEGVTGVHLMP-------L-MLVAGDHAINDMASDDGDSWKMRFNAAGIPATPWLSG | | :| | :| | :. | | |: | .. LFEQKGYQAFVYVPVGFVADHLEVLYDNDYECKVVTDDIGASYY----------RPEMPLGENPAIRAMFVAHLHQALNMAVEEAA | . . | -NAKPEFIDALATVVLKKLG------Identities in the aligned portion = 39/319 (12.2%) Residue range in the template: 5 : : 309 :
The refined alignment basically agrees with the structure-structure alignment, and it is significantly better than the original one without the constraint. The structure using the refined alignment is shown in Figure 2.7.3. The target t0053 and the template 1ak1 belong to the same superfamily. PROSPECT is also able to predict some proteins in the fold category. Figure 2.7.4 shows a prediction for the CASP-3 target t0067 using a template in a different superfamily but the same fold (chain A of 1tup). From both figures, one can get an idea about the quality of the models using threading. It should be noted that even when a perfect alignment is achieved, some conformational differences between the model and the experimental structure are expected on the backbones. The sidechain prediction based on threading alignment is unreliable. Although structure models derived from threading often have lower quality than those from homology modeling with significant sequence similarities, they are typically much better than the models generated from ab initio predictions and can often provide useful information for understanding protein function. Quantitation of Protein Interactions
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A
B
Figure 2.7.3 A comparison between predicted (A) and experimental (B) structures for the CASP-3 target t0053. Cylinders indicate α helices, strands indicate β sheets, dark lines indicate turns, and thin lines indicate loops. This figure was made using the INSIGHT II package (Molecular Simulations, 1998).
A
Protein Tertiary Structure Prediction
B
Figure 2.7.4 A comparison between predicted (A) and experimental (B) structures for the CASP-3 target t0067. Cylinders indicate α helices, strands indicate β sheets, dark lines indicate turns, and thin lines indicate loops. This figure was made using the INSIGHT II package (Molecular Simulations, 1998).
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AB INITIO PREDICTION An ab initio protein structure prediction derives a structure model through the optimization of an energy function that describes the physical properties or statistical preferences of amino acids. After decades of extensive effort, the ab initio tertiary structure prediction from sequence has proven to be extremely difficult. To run an ab initio prediction program requires long computing times, and the prediction results are generally unreliable. However, some recent developments using hierarchical approaches from local structures to the global structure seems to give ab initio predictions some new hope for generating low-resolution structures. When local structures are more or less defined, assembling them significantly reduces the computational search space compared to folding individual residues, and hence increases the chance of predicting a good structure. The optimization process is typically carried out using a genetic algorithm (Unger and Moult, 1992) or Monte Carlo simulations (Skolnick and Kolinski, 1991). Local structures can be built based on empirically derived data of preferred torsion angles in secondary structure elements, as done by the program LINUS (Local Independently Nucleated Units of Structure; Srinivasan and Rose, 1995). The “mini-threading” method (Simons et al., 1997) may be a more efficient way to build local structures. Mini-threading obtains the matches between short structure segments of template and the query sequence for building local structures. Some success of mini-threading has been demonstrated in CASP-3 (CASP, 1999). However, ab initio prediction programs are typically unavailable to the public. Therefore, no technical details about running these programs are described in this unit. COMMENTARY Although computation models often provide valuable information, they are nonetheless predictions no matter how good the methods are. A user should always keep in mind the general quality and the confidence level of the models derived by each method, as described above, when using them to draw any conclusion. It is usually rewarding to use different methods. The consensus and variations in different predictions may provide a clue about the reliability of a prediction. Predictions from other computational tools, e.g., secondary structure prediction, solvent accessibility prediction, and residue contact analysis through correlated mutation in a protein family, can also be used as input or to provide independent assessments in fold recognition and ab initio prediction. Whenever any experimental information for a structure is available, the predictor should incorporate the information in the tools or at least use the information to verify the output model. Last but not least, it is important to visually check the model in a molecular graphics program and manually adjust the model (e.g., change the alignment), if necessary. LITERATURE CITED Alexandrov, N.N., Nussinov, R., and Zimmer, R.M. 1996. Fast protein fold recognition via sequence to structure alignment and contact capacity potentials. In Biocomputing: Proceedings of the 1996 Pacific Symposium (L. Hunter and T. Klein, eds.) pp. 53-72. World Scientific Publishing, Singapore. Al-Karadaghi, S., Hansson, M., Nikonov, S., Jonsson, B., and Hederstedt, L. 1997. Crystal structure of ferrochelatase: The terminal enzyme in heme biosynthesis. Structure 5:1501-1510. Altschul, S.F. and Koonin, E.V. 1998. Iterated profile searches with PSI-BLAST—a tool for discovery in protein databases. Trends Biochem. Sci. 23:444-447. Altschul, S.F., Madden, T.L., Schäffer, A.A., Zhang, J., Zhang, Z., Miller, W., and Lipman, D.J. 1997. Gapped BLAST and PSI-BLAST: A new generation of protein database search programs. Nucl. Acids Res. 25:3389-3402. Bernstein, F.C., Koetzle, T.F., Williams, G.J.B., Meyer, E.F., Brice, M.D., Rodgers, J.R., Kennard, O., Shimanouchi, T., and Tasumi, M. 1977. The protein data bank: A computer based archival file for macromolecular structures. J. Mol. Biol. 112:535-542.
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Bowie, J.U., Luthy, R., and Eisenberg, D. 1991. A method to identify protein sequences that fold into a known three-dimensional structure. Science 253:164-170. Bryant, S.H. and Altschul, S.F. 1995. Statistics of sequence-structure threading. Curr. Opinion Struct. Biol. 5:236-244. Bryant, S.H. and Lawrence, C.E. 1993. An empirical energy function for threading protein sequence through the folding motif. Proteins Struct. Funct. Genet. 16:92-112. CASP (Critical Assessment of Techniques for Protein Structure Prediction). 1995. Protein structure prediction issue. Proteins Struct. Funct. Genet. 23:295-462. CASP. 1997. Protein structure prediction issue. Proteins Struct. Funct. Genet. 29:1-230. CASP. 1999. Protein structure prediction issue. Proteins Struct. Funct. Genet. 37:1-237. Crawford, O.H. 1999. A fast, stochastic algorithm for protein threading. Bioinformatics 15:66-71. Fischer, D. and Eisenberg, D. 1996. Fold recognition using sequence-derived predictions. Protein Sci. 5:947-955. Fischer, D., Rice, D., Bowie, J.U., and Eisenberg, D. 1996. Assigning amino acid sequences to 3-dimensional protein folds. FASEB J. 10:126-136. Gerstein, M. 1998. Patterns of protein-fold usage in eight microbial genomes: A comprehensive structural census. Proteins Struct. Funct. Genet. 33:518-534. Godzik, A., Skolnick, J., and Kolinski, A. 1992. A topology fingerprint approach to the inverse folding problem. J. Mol. Biol. 227:227-238. Heniko, S. and Heniko, J.G. 1994. Protein family classification based on searching a database of blocks. Genomics 19:97-107. Holm, L. and Sander, C. 1996. Mapping the protein universe. Science 273:595-602. Holm, L. and Sander, C. 1998. Dictionary of recurrent domains in protein structures. Proteins Struct. Funct. Genet. 33:88-96. Hu, X., Xu, D., Hamer, K., Schulten, K., Koepke, J., and Michel, H. 1995. Predicting the structure of the light-harvesting complex II of Rhodospirillum molischianum. Protein Sci. 4:1670-1682. Hughey, R. 1996. Parallel hardware for sequence comparison and alignment. CABIOS 12:473-479. Hughey, R. and Krogh, A. 1996. Hidden Markov models for sequence analysis: Extension and analysis of the basic method. CABIOS 12:95-107. Humphrey, W.F., Dalke, A., and Schulten, K. 1996. VMD—visual molecular dynamics. J. Mol. Graphics 14:33-38. Jones, D.T. 1999. GenTHREADER: An efficient and reliable protein fold recognition method for genomic sequences. J. Mol. Biol. 287:797-815. Jones, D.T., Taylor, W.R., and Thornton, J.M. 1992. A new approach to protein fold recognition. Nature 358:86-89. Karplus, K., Barrett, C., and Hughey, R. 1998. Hidden Markov models for detecting remote protein homologies. Bioinformatics 14:846-856. Krogh, A., Brown, M., Mian, I.S., Sjolander, K., and Haussler, D. 1994. Hidden Markov models in computational biology: Applications to protein modeling. J. Mol. Biol. 235:1501-1531. Laskowski, R.A., MacArthur, M.W., Moss, D.S., and Thornton, J.M. 1993. PROCHECK: A program to check the stereochemical quality of protein structures. J. Appl. Crystallogr. 26:283-291. Lund, O., Frimand, K., Gorodkin, J., Bohr, H., Bohr, J., Hansen, J., and Brunak, S. 1997. Protein distance constraints predicted by neural networks and probability density functions. Protein Eng. 10:1241-1248. Madej, T., Gibrat, J.F., and Bryant, S.H. 1995. Threading analysis suggests that the obese gene product may be a helical cytokine. FEBS Lett. 373:13-18. Milburn, D., Laskowski, R.A., and Thornton, J.M. 1998. Sequences annotated by structure: A tool to facilitate the use of structural information in sequence analysis. Protein Eng. 11:855-859. Molecular Simulations. 1998. Insight II (Release 98.0). Molecular Simultions, San Diego, Calif. Murzin, A.G., Brenner, S.E., Hubbard, T., and Chothia, C. 1995. SCOP: A structural classification of proteins database for the investigation of sequences and structures. J. Mol. Biol. 247:536-540. Nilges, M. and Brünger, A. 1993. Successful prediction of the coiled coil geometry of the gcn4 leucine zipper domain by simulated annealing: Comparison to the x-ray structure. Proteins Struct. Funct. Genet. 15:133-146. Protein Tertiary Structure Prediction
Park, J., Karplus, K., Barrett, C., Hughey, R., Haussler, D., Hubbard, T., and Chothia, C. 1998. Sequence comparisons using multiple sequences detect twice as many remote homologues as pairwise methods. J. Mol. Biol. 284:1201-1210.
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Pearson, W.R. and Lipman, D.J. 1988. Improved tools for biological sequence comparison. Proc. Natl. Acad. Sci. U.S.A. 85:2444-2448. Peitsch, M.C. 1996. ProMod and Swiss-Model: Internet-based tools for automated comparative protein modelling. Biochem. Soc. Trans. 24:274-279. Pontius, J., Richelle, J., and Wodak, S.J. 1996. Quality assessment of protein 3D structures using standard atomic volumes. J. Mol. Biol. 264:121-136. Rost, B. 1995. TOPITS: Threading one-dimensional predictions into three-dimensional structures. ISMB 3:314-321. Rost, B. and Sander, C. 1993. Prediction of protein secondary structure at better than 70% accuracy. J. Mol. Biol. 232:584-599. Sali, A. and Blundell, T.L. 1993. Comparative protein modelling by satisfaction of spatial restraints. J. Mol. Biol. 234:779-815. Sanchez, R. and Sali, A. 1998. Large-scale protein structure modeling of the Saccharomyces cerevisiae genome. Proc. Natl. Acad. Sci. U.S.A. 95:13597-13602. Simons, K.T., Kooperberg, C., Huang, E., and Baker, D. 1997. Assembly of protein tertiary structures from fragments with similar local sequences using simulated annealing and Bayesian scoring functions. J. Mol. Biol. 268:209-225. Sippl, M.J. and Weitckus, S. 1992. Detection of native-like models for amino acid sequences of unknown three-dimensional structure in a database of known protein conformations. Proteins Struct. Funct. Genet. 13:258-271. Skolnick, J. and Kolinski, A. 1991. Dynamic monte carlo simulations of a new lattice model of globular protein folding, structure and dynamics. J. Mol. Biol. 221:499-531. Smith, T.F. and Waterman, M.S. 1981. Comparison of biosequences. Adv. Appl. Math. 2:482-489. Smith, T., Conte, L.L., Bienkowska, J., Gaitatzes, C., Rogers, R., and Lathrop, R. 1997. Current limitations to protein threading approaches. J. Comp. Biol. 4:217-225. Srinivasan, B.N. and Blundell, T.L. 1993. An evaluation of the performance of an automated procedure for comparative modelling of protein tertiary structure. Protein Eng. 6:501-512. Srinivasan, R. and Rose, G. 1995. LINUS—a hierarchic procedure to predict the fold of a protein. Proteins Struct. Funct. Genet. 22:81-99. Tripos. 1999. SYBYL 6.5.3. Tripos, Inc., St. Louis. Unger, R. and Moult, J. 1992. Potential of genetic algorithms in protein folding and protein engineering simulations. J. Mol. Biol. 5:637-645. Vriend, G. 1990. WHAT IF: A molecular modelling and drug design program. J. Mol. Graphics 8:52-56. Wang, Z.X. 1998. A re-estimation for the total numbers of protein folds and superfamilies. Protein Eng. 11:621-626. Xu, Y., Xu, D., and Uberbacher, E.C. 1998. An efficient computational method for globally optimal threading. J. Comp. Biol. 5:597-614. Xu, Y., Xu, D., Crawford, O.H., Einstein, J.R., Larimer, F., Uberbacher, E.C., Unseren, M.A., and Zhang, G. 1999. Protein threading by PROSPECT: A prediction experiment in CASP3. Protein Eng. 12:101-109.
Contributed by Dong Xu and Ying Xu Oak Ridge National Laboratory Oak Ridge, Tennessee
The authors thank Drs. Edward C. Uberbacher, J. Ralph Einstein, and Oakley H. Crawford for helpful discussions, and Drs. Stephen Altschul, Kevin Karplus, and Richard Hughey for their constructive comments on this manuscript. This research was sponsored by the Office of Biological and Environmental Research, U.S. Department of Energy, under contract no. DE-AC05-96OR22464 with Lockheed Martin Energy Research Corporation.
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Protein Tertiary Structure Modeling As an experimental science, biology increasingly involves large-scale automated approaches. This data production and gathering activity has profoundly changed the way in which data and information can be handled. The best known manifestation of this paradigm shift is genome sequencing activity, which yields ever increasing numbers of novel genes, for which no biological function is known. It is the proteins that are the “working” molecules responsible for most biological processes. Understanding their modes of action will allow us to unveil their biological role, and how their mutants cause diseases. In the past, proteins and their corresponding genes were generally discovered during functional experiments, which provided the biological context for their role, and which implicitly linked the genes to their function. The genome sequencing approach to biology has completely reversed this paradigm; one now has to discover the function of the large number of newly sequenced genes and their protein products. A widely used means to unravel the function of a protein is to create molecular mutants and observe their deviation from normal behavior. Experience has proven that the efficiency of this experimental approach is considerably enhanced by the insights the 3-D structure of the protein can provide. This can now be achieved by either experimental or, at least in part, by theoretical methods known as protein structure prediction. Experimental protein structure determination methods are often hampered by technical difficulties which, even if they can be overcome, are time and resource intensive. The number of known 3-D structures of proteins thus only represents a small fraction of the known protein sequences. Indeed, over 230,000 protein sequences are logged in the SwissProt/trEMBL database (Bairoch and Apweiler, 1999), while fewer than 10,000 protein sequences have a known 3-D structure and are stored in the Protein Data Bank (http://www.rcsb.org/pdb/). In this context it is not surprising that theoretical approaches have been explored, of which comparative protein modeling is by far the most reliable. Comparative protein modeling follows four steps: It starts with the quest for at least one protein of known structure that shares a significant degree of sequence identity with the se-
quence to be modeled, or target sequence (step 1). These modeling templates serve as the structural basis on which the target model will be built. Then, the target and template sequences are aligned (step 2). This sequence alignment is the key element of the modeling procedure, since it serves as the guide while building the structural “scaffold” of the model. Indeed, each position in the alignment defines the 3-D position of the target sequence’s amino acids. This structural scaffold is then used to build an atomic representation of all amino acids shared by the target and template sequence (step 3), but it is not sufficient to build the portions of the structure that differ from the modeling templates the same way. These deviant sequences are rebuilt using the basic rules that govern protein structures (step 4). This can now be done automatically using the SwissModel World Wide Web server (Guex et al., 1999). This unit will concentrate on delivering sufficient background and guidelines to build protein models.
GLOSSARY For the sake of clarity, a few terms used throughout the unit are defined below: comparative protein modeling often also called “homology modeling” or “knowledgebased modeling”; this is the process by which a model is built. core part of a protein structure that can be considered largely conserved within a protein family (Chothia and Lesk, 1986). ExPDB SwissModel template structure database; contains one entry for each individual protein chain of the PDB. First Approach simplest SwissModel request; initiated by submission of a sequence through the World Wide Web-interface. Alignment between target and template sequences is generated automatically. Optimise Mode more complex SwissModel request; initiated by submitting a project file generated with SPDBV. Allows for manual adjustment of the sequence alignment between target and template sequences. PDB Protein Data Bank; repository of experimentally solved protein structures. P(n) BLAST (Altschul et al., 1990) unlikelihood probabilities (see UNIT 2.5). protein chain a single polypeptide chain; PDB files can contain more than one protein chain.
Contributed by Nicolas Guex, Torsten Schwede, and Manuel C. Peitsch Current Protocols in Protein Science (2000) 2.8.1-2.8.17 Copyright © 2000 by John Wiley & Sons, Inc.
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reference template generally the template sharing the highest degree of sequence similarity with the target. rmsd root mean square deviation between a set of atoms (Å); represents the average displacement of atoms in 3D-space between two comparable structures. SPDBV Swiss-PdbViewer; an interactive software to display, analyze, and modify protein structures. target sequence protein sequence to be modeled. template structure(s) experimentally solved structure(s) used as template(s) for model construction (PDB or ExPDB file). template sequence(s) protein sequence(s) of the template(s) defined above.
ACCESSING SWISSMODEL PROGRAMS AND DOCUMENTATION SwissModel is an Internet-based service for automated comparative modeling of protein tertiary structures. SwissModel requests can be submitted through forms maintained on the ExPASy World Wide Web site, at http://www. expasy.ch/swissmod/SwissModel.html. It is advisable to install the Swiss-PdbViewer (SPDBV) software, which is available for PC, Macintosh, SGI, and i86LINUX platforms (Guex et al., 1999). This software can also b e d ownloaded from ExPASy at http://www.expasy.ch/spdbv/mainpage.htm. SPDBV is an interactive sequence-to-structure workbench, which acts as a client to SwissModel and allows the displaying, analyzing, and modifying of protein structures. By default, SwissModel will return SPDBV project files, which contain not only the coordinates of the model, but also all spatially superposed templates and their corresponding deduced structural alignment. SPDBV can then be used to adjust the alignment between target and template sequences, and to submit the corresponding modeling request.
ExPDB DATABASE
Protein Tertiary Structure Modeling
SwissModel uses ExPDB files as modeling templates. These are derived from the PDB database. Modeling procedures are frequently applied to a protein monomer, and many PDB files contain more than one protein chain. Hence, the ExPDB database has been created, in which each chain from a given PDB file has been split into a different entry. For example, the PDB entry 4MDH contains two protein chains (A and B). This generates the two cor-
responding ExPDB entries 4MDHA and 4MDHB. Some PDB files contain only one chain without a specific chain name (e.g., 1CRN), in this case, an “_” is added to the ExPDB name (1CRN_). SwissModel has been designed to build proteins, thus only amino acids (or modified amino acids) are kept in ExPDB files. This means that all N-terminal blocking groups (e.g., acetyl) and water molecules are removed. However, enzymatic cofactors, phosphorylated residues, and ions are kept, provided they are in close contact with the protein chain. Although they are not yet used during the modeling procedure per se, they are present in the SPDBV project files returned to the user. This allows for a quick inspection of the model in order to see whether these non-proteinaceous molecules could be conserved in the model.
FORMATTING A FIRST APPROACH MODELING REQUEST Mandatory Items As the modeling procedure can take more than 1 hr, depending on the size and complexity of the model to be built, all results are returned by e-mail. Hence one must provide a valid e-mail address, name, and a title for the request. The preferred format for entering the target sequence is the FASTA format (for a description, see UNIT 2.5). Alternatively, if the sequence is already in the SwissProt database, its accession code can be provided (e.g., P02761, used in the example below). Other formats, such as raw sequence, GCG, and SwissProt, are also accepted. Extra spaces and numbers are skipped. This is the minimal information to be submitted for model building. However, a number of options are available for submitting additional information.
Options The automatic template selection is done by comparing the target sequence with ExPDB template sequence. The BLASTP2 P(n) threshold value used to select the modeling templates can be altered. By default a value of 10−5 is used. A lower value (10−10) will select for templates sharing a higher degree of identity with the target sequence, while a higher value (10−4) will allow more remotely similar templates to be used. Up to five templates may be selected by the modeling procedure. However, it may occur that a non-optimal mix of templates is selected. This is the case when several
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distinct structures of the same protein are available in the template database (for example pro-enzyme/mature enzyme). It is then advisable to explicitly provide the name of the ExPDB template(s) to be used.
Finding the Best Templates As discussed above, SwissModel may not choose the optimal template(s), as there is no biological context taken into account by the template-selection procedure. To address this problem, it would be best to have some contextual information about the available templates, as this would allow the user to select the templates based on the experimental conditions under which their structures were elucidated. Indeed, template structures elucidated in the presence or absence of substrates, cofactors, and other variants of the biological unit are not identical. The ExPDB database can be searched with the target sequence for available templates through the World Wide Web server by clicking on the “Search Template” link at, http://www.expasy.ch/swissmod/SwissModel .html. All one needs to do is to provide the target sequence (in FASTA format) in the World Wide Web search form. The resulting report is sorted by BLASTP2 P(n) and contains the name of the proposed ExPDB templates, the method used to elucidate them (X-RAY, NMR), the resolution (for X-RAY), and a short description. A hyperlink to the corresponding PDB entry allows one to reach detailed information about the structure used to generate the ExPDB template (the REMARK fields in PDB files are usually quite instructive). Details of aligned regions between the sequence to be modeled and the template can also be reached through a hyperlink. Up to five templates can be selected for modeling. The first template listed is likely to be the most appropriate one. However, it may happen that several templates have the same P(n) value. Quite often, the server reports multiple entries for the same protein (e.g., multiple chains from a homo-oligomeric protein structure) and it is sufficient to select only one of these. When the same protein structure has been solved twice at different resolutions (e.g., 1CRN 1.5 Å and 1CNR 1.05 Å), only the structure with the best resolution should be selected (in this case 1CNR). Adding other templates with different sequences sometimes helps during side-chain modeling. However, there is no general rule, and it is often worth trying with and without additional templates.
SwissModel will automatically reject templates that cannot be used concurrently with the reference template (first one of the list). This will happen whenever a template cannot be automatically superposed onto the reference one. This limitation can be superseded using the “Optimise” mode.
Using Private Templates Whenever possible, use the ExPDB templates maintained on the server, as they have been processed for optimal use by SwissModel. However, in certain cases one might want to use templates that are not yet available in the ExPDB database. Both “First Approach” and “Optimise” modes allow the use of private templates. To ensure a high probability of success, those templates must comply with these rules: they must (1) be in PDB format, (2) contain only one protein chain with all backbone atoms (no gaps), and (3) be devoid of non-protein atoms (which are marked as “HETATM” in PDB files).
VIEWING SWISSMODEL RESULTS One can specify that the results be delivered in normal mode, in which case only the coordinates of the model, together with a detailed log of the procedure, are sent. The short mode will return just the coordinates of the model, without a log file. The default mode will return a detailed log o f th e m od elin g p ro ce du re an d a SwissPdbViewer project file, which will contain the final model and the template(s), superposed in 3D-space. This allows for the model to be analyzed and compared to its templates. If some regions of the model are dubious due to a bad initial sequence alignment, they can be modified and the altered alignment directly submitted to the SwissModel server as an Optimise mode request (see below). Optionally, a “WhatCheck” report (Hooft et al., 1996) of the model can be sent by the s e r v e r. T h e a n a l y s i s p e r f o r m ed b y WhatCheck includes atom nomenclature, bond-length deviation, side-chain planarity, packing, and many more. The report can be used to assess the accuracy of the model and detect regions that might need fine-tuning. Note that WhatCheck reports are text files that can be opened with SPDBV. Clicking on a report line will automatically center the view on the problem, which allows for quick inspection. Other features can be analyzed directly with SPDBV (for further details refer to the section on Model Analysis).
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By default, the models are sent as e-mail attachments. If your e-mail program does not support attachments, you can specify to receive the results in plain ASCII format. In this case, you will need to copy the content of the mail into a text file that will end with a file extension “.pdb” before dragging it to the SPDBV icon.
EXAMPLES OF SWISSMODEL REQUESTS In this section, various modeling examples are reviewed step by step. They are sorted by level of complexity. As new templates are added to the database, when new proteins are experimentally solved, the results will vary and running the same query again might not deliver exactly the same output.
Protein Tertiary Structure Modeling
First Approach A typical modeling request is shown in Figure 2.8.1. The sequence of rat fatty acid–binding protein (P02761) has been submitted through the “First Approach” mode. The server just acknowledges that the “request submission was successful,” and mentions that all further communications will come via e-mail. As soon as the server starts to build the model, a message containing a unique process identification is sent, as shown in Figure 2.8.2. Further e-mails containing log reports and model coordinates are received up to several hours later depending on the complexity of the model to build, the server load, or the Internet traffic. The report log is divided into two sections. The first section contains the results of the
Figure 2.8.1 Submitting a First Approach modeling request to the SwissModel server.
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>>>>>
Welcome to the SWISS-MODEL Protein Modelling Server
<<<<<
==================================================================== Dear JIM, Title of your Request: Process identification is:
MUP_RAT AAAa002Of
Swiss-Model ‘(SWISS-MODEL Version 3.5)’ started on 18-Aug-1999 The modelling procedure is now in progress, and its results should be sent to you shortly. ==================================================================== If results of this search are reported or published, please mention the publications listed below. (end of message truncated for brevity)
Figure 2.8.2 Message sent when the server starts to build the model. The unique process identification (AAAa002Of) is added in the subject of all subsequent e-mails. This is useful when several modeling requests are treated concurrently.
template identification performed with BLASTP2 (Fig. 2.8.3). The putative template candidates are shown sorted by the percent identity with the target sequence. Templates that share less than 25% identity with the target are automatically discarded, as their automatic alignment is usually not reliable enough to allow for blind model building (see section on Model Accuracy). The regions where the target and template sequences can be aligned are shown graphically. If the suitable templates are clustered in more than one region, the target sequence is split into two or more segments that are modeled separately and labeled Batch.1, Batch.2, and so on up to Batch.n. This occurs e.g., for membrane proteins that have an intra and extracellular domain elucidated by distinct experiments. The second part of the log file contains a summary of the modeling procedure (Fig. 2.8.4). Although some steps will vary from request to request, it typically contains a list of the templates that have been loaded, followed by the result of their 3D-superposition. The first template loaded is considered a reference template, which will guide the modeling procedure. Additional templates that cannot be superposed onto the reference template are discarded. In the case presented here the only template retained is 1MUP_. Recently two experimentally solved structures of rat lipid bind-
ing protein have been deposited in the PDB database (accession codes 2A2G and 2A2U). They may be used to assess the models built in this tutorial based on suboptimal templates. The target sequence is then aligned onto the selected template(s), and the overhangs are discarded (residues that cannot be modeled because they are before or after the first/last residues of the reference template). Once everything is in place, the model is built starting with the core, followed by a loop reconstruction (if the target contains insertions/deletions relative to the reference template). Finally a few energy minimization steps are performed using the GROMOS96 (van Gunsteren et al., 1996) force field as implemented in SPDBV. The final energy is reported at the end of the log file.
First Approach Using Specific Templates The same protein as in the example above has been submitted to SwissModel, but this time modeling templates have been explicitly specified (see Finding the Best Templates for conceptual background, and Optimise Mode for a concrete example). As 1MUP_ is more similar to the target protein than 1A3YB, it is given first to force SwissModel to use it as the reference (Fig. 2.8.5). This is likely to provide a better model than putting 1A3YB first. Specifying the templates will allow SwissModel to skip the template search, and to start
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directly with the pair-wise alignment step. Then the modeling procedure is performed and the reported results (Fig. 2.8.6) are quite similar to those reported in the previous section (Figs. 2.8.3 and 2.8.4). However, in this case the second template (1A3YB) can be superposed onto the reference template (1MUP_). This
adds a step to the procedure during which a structural alignment between the two templates is generated. In addition, information contained in both templates will be used and averaged to build the model. The reference template will be used alone in all regions where additional templates diverge from it too greatly. It may happen
Figure 2.8.3 First part of the modeling log file, containing a summary of the template identification process (see text).
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that the averaging procedure produces small distortions of the backbone (in our case at GLU 50) that will be repaired by the GROMOS96 force field used during the final modeling steps. The remaining steps are essentially the same as for the standard procedure. Depending on the number of templates used, the final energy may differ. In this case the final energy is −4154.777 KJ/mol compared to −6030.488 KJ/mol in the previous example. An all atom comparison between the two models yields 0.98 Å rmsd (for 1279 atoms), while the 157 Cα atoms deviate by 0.35 Å. In
both cases, WhatCheck judges the model satisfactory.
Optimise Mode The “Optimise” mode can be used either to optimize a model generated by the “First Approach” mode, or directly. In both cases this can only be done from SPDBV. In the “Optimise” mode the modeling procedure is uncoupled from the automated template selection, superposition, and target-sequence alignment. Those steps need to be performed manually within SPDBV, which provides a higher level of flexi-
ProModII trace log for Batch.1 ============================================================ ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII:
ProMod version 3.5b3 date Jul 19 1999 17:15 SPDBV version 3.5b3 Loop version 2.60 LoopDB version 2.60 Parameters version 3.5 Topologies version 3.5 Loading Template: 1MUP_.pdb 1MUP: some sidechain atoms are missing --> reconstructing them Loading Template: 1OBPB.pdb Loading Template: 1OBPA.pdb Loading Template: 1PBOA.pdb Loading Template: 1PBOB.pdb Loading Raw Sequence Iterative Template Fitting Protein does not fit well; Removing Layer 1OBPB (if you really want to include this template, use the optimise Iterative Template Fitting Protein does not fit well; Removing Layer 1OBPA (if you really want to include this template, use the optimise Iterative Template Fitting Protein does not fit well; Removing Layer 1PBOA (if you really want to include this template, use the optimise Iterative Template Fitting Protein does not fit well; Removing Layer 1PBOB (if you really want to include this template, use the optimise Aligning Raw Sequence Refining Raw Sequence Alignment N-terminal overhang trimmed for chain ‘ ’. Start at residue: 6 C-terminal overhang trimmed for chain ‘ ’. End at residue: 162 adding blocking groups Averaging Sidechains Adding Missing Sidechains Optimizing Sidechains Dumping Preliminary Model Adding Hydrogens Optimizing loops and OXT (nb = 1) Current Total Energy: -6030.488 KJ/mol Removing Hydrogens Fixing Atom Nomenclature Dumping Sequence Alignment Done.
mode) mode) mode) mode)
Figure 2.8.4 Second part of the modeling log file, containing a summary of action taken during the modeling procedure (see text).
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Figure 2.8.5 Submitting a First Approach modeling request to the SwissModel server with specific EXPDB templates.
Protein Tertiary Structure Modeling
bility, and a real-time visual feedback to help refine the guiding sequence alignment. Hence insertions/deletions can be finely adjusted, and many modeling requests that would otherwise fail in the First Approach mode can be built satisfactorily. The following example describes a typical workflow step by step: 1. Fetch the SwissProt entry Q21735 from the ExPASy site at http://www.expasy.ch/sprot/. 2. Ask for a FASTA-formatted sequence (Fig. 2.8.7). 3. Prepare a text file named Q21735.txt, which will be used later. 4. Paste the FASTA-formatted sequence into the “search template” form on the SwissModel World Wide Web site at http://www. expasy.ch/swissmod/SwissModel.html to find suitable templates. SwissModel returns a list of possible templates (Fig. 2.8.8) aligned onto the sequence to model (Fig. 2.8.9). 5. Download the ExPDB file 1OUNA, the template with the best BLAST score, and save on a local disk as 1OUNA.pdb. Note that other templates give similar scores, but they have been solved at a lower resolution.
The next part of the procedure only outlines the conceptual steps that need to be taken within SPDBV. It is not a full tutorial about the manipulation of SPDBV; this can be found at http://www.expasy.ch/spdbv/. The name of the menu from which a function can be called is shown in [brackets]. 6. Open the ExPDB file 1OUNA.pdb in SPDBV. 7. Use “Load raw sequence to model” [SwissModel] to open the target sequence file Q21735.txt. 8. Open the alignment window [Windows] and click on the name of the target sequence to select it. 9. Use the “Magic Fit” function [Tools]. The alignment should look like the one presented in the Figure 2.8.10. 10. Click “submit a modeling request” [SwissModel]. A SPDBV project file will be saved, containing the target protein, the template, and the alignment. A web-browser window will open with a SwissModel submission form. Note: if it is your first submission, you will be prompted to provide your e-mail address to SPDBV.
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AlignMaster output ============================================================ Length of target sequence: 181 residues Reading user-defined template list Extracting template sequences Running pair-wise alignments with target sequence Sequence identity of templates with target: 1MUP_.pdb: 63.7 % identity 1A3YB.pdb: 34 % identity Looking for template groups Global alignment overview: Target Seq: 1MUP_.pdb 1A3YB.pdb
|============================================================| | ---------------------------------------------------| -----------------------------------------------
AlignMaster found 1 regions to model separately: 1: Using template(s) 1A3YB.pdb 1MUP_.pdb Creating Batch files for ProMod (if any): Batch.1: residues 15 - 181 of submitted sequence. Exiting AlignMaster ProModII trace log for Batch.1 ============================================================ ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII:
ProMod version 3.5b3 date Jul 19 1999 17:15 SPDBV version 3.5b3 Loop version 2.60 LoopDB version 2.60 Parameters version 3.5 Topologies version 3.5 Loading Template: 1MUP_.pdb : some sidechain atoms are missing --> reconstructing them Loading Template: 1A3YB.pdb : some sidechain atoms are missing --> reconstructing them Loading Raw Sequence Iterative Template Fitting Generating Structural Alignment Aligning Raw Sequence Refining Raw Sequence Alignment N-terminal overhang trimmed for chain ‘ ’. Start at residue: 6 C-terminal overhang trimmed for chain ‘ ’. End at residue: 162 adding blocking groups Weighting Backbones Averaging Sidechains Adding Missing Sidechains Small Ligation (C-N < 3.0A) ignored; GROMOS will repair it at residue GLU 50 Optimizing Sidechains Dumping Preliminary Model Adding Hydrogens Optimizing loops and OXT (nb = 1) Current Total Energy: -4154.777 KJ/mol Removing Hydrogens Fixing Atom Nomenclature Dumping Sequence Alignment Done.
Figure 2.8.6 Log file for the optimise mode when modeling templates are specified.
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>Q21735 MSFNPDYESVAKAFIQHYYSKFDVGDGMSRAQGLSDLYDPENSYMTFEGQ QAKGRDGILQKFTTLGFTKIQRAITVIDSQPLYDGSIQVMVLGQLKTDED PINPFSQVFILRPNNQGSYFIGNEIFRLDLHNN Figure 2.8.7 Amino acid sequence of the SwissProt entry Q21735 in FASTA format.
Figure 2.8.8 Results of a template search for the SwissProt entry Q21735.
Protein Tertiary Structure Modeling
11. Select the saved project file by clicking on the [Browse] button in the Web form. Make sure that your name and e-mail address are correct. Optionally you may ask for a WhatCheck report. 12. Submit the request. The results will come by e-mail. The modeling log file (Fig. 2.8.11) will contain the same kind of information as presented earlier (Fig. 2.8.6), with an additional section for each loop that had to be reconstructed (insertions/deletions). In order to build a loop, SwissModel defines two anchor residues—one at each end—that will not move during the procedure. It then attempts to generate a set of acceptable solutions for each
insertion/deletion. If the initially defined anchor residues are not suitable (as no acceptable solution is found), then different anchor residues are selected until at least one acceptable loop is found. For each loop the best solution is reported in the log file, which shows the residues used as anchor points, the number of close contacts (clashes), an energy evaluation with the GROMOS96 force field (as implemented in SPDBV), and a mean-force potential evaluation. This part of the modeling procedure is under constant development, and the reported summary may differ slightly in the future. In general the final force-field energy (FF) should be as low as possible (negative values), while as few clashes as possible should be
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Figure 2.8.9 Alignment between Q21735 and the best template identified.
reported. Furthermore a solution should be found for each insertion/deletion without the need for more than one or two anchor residue reselections. In this example, no loop was found with anchor residues GLY25 and ARG 30, so more flexibility was given to this loop by using ALA31 as a new anchor point. Rebuilding of the next loop (anchor residues ALA31 and LEU34) also failed and needed a second pass, using residues ARG30 and LEU34 as anchor points. In summary, the complete region from GLY25 to LEU34 has been rebuilt. This is most likely an indication that the alignment was not optimal. In order to correct this,
select the residue ASP25 of 1OUNA in the alignment window and move it to the right with the arrow key, until the alignment looks like the one presented in Figure 2.8.12. Then select ASP35 of 1OUNA and move it to the left by one position. Finally select ASP110 and ALA111 and shift them by one position to the right. The new alignment can now be resubmitted to the SwissModel server. The results of this new attempt are shown in Figure 2.8.13. This time, the loop rebuilding procedure seems to find better solutions, and the final global energy is significantly lower than with the default alignment. Figure 2.8.14 shows a comparison of the loop region between and Asn123 derived
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by the First Approach mode (A and B) and the Optimise Mode (C and D). The molecular interactions of the loop based on the automatic alignment (A) are less favorable compared to those found after manually optimizing the placement of insertions in the sequence alignment (C). A ribbon representation of the same loops (B and D) illustrates that the manual adjustment of the insertion into the turn between the two strands reduces the distortions in the rebuilt loop.
It should however be noted that loops are always the least reliable parts of the model. The automated modeling procedure generally returns a reasonable starting point, which can be optimised in only a few steps. The other loops rebuilt in this example are worth considering in more detail, and alternative alignments should be explored. The main reason for SwissModel failures is the misplacement of insertions and deletions in the sequence alignment. Although
A
B Q21735 1OUNA
1 2
MSFNPDYESV AKAFIQHYYS KFDVGDGMSR AQGLSDLYDP ENSYMTFEGQ GDKPIWEQI GSSFIQHYYQ LFDND----R TQ-LGAIYI- DASCLTWEGQ . .* * . . .****** ** * .* *. .* . * .* ***
Q21735 1OUNA
51 45
QAKGRDGILQ KFTTLGFTKI QRAITVIDSQ PLYDGSIQVM VLGQLKTDED QFQGKAAIVE KLSSLPFQKI QHSITAQDHQ PTPDSCIISM VVGQLKADED * *. .*.. *...* * ** *..** * * * *. * * *.****.***
Q21735 1OUNA
101 95
PINPFSQVFI LRPNNQGSYF IGNEIFRLDL HNN PIMGFHQMFL LKNINDA-WV CTNDMFRLAL HNF ** * *.*. *. *.. *..*** * **
Figure 2.8.10 (A) Automatic alignment between Q21735 and its modeling template (1OUNA) as shown within SPDBV. (B) Complete alignment, shown for clarity.
(......) ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: (......) ProModII:
Building CSP loop with anchor residues Number of Ligations found: 19 all loops are bad; continuing CSP with Building CSP loop with anchor residues Number of Ligations found: 244 ACCEPTING loop 113: clash= 1 FF= Building CSP loop with anchor residues Number of Ligations found: 1 all loops are bad; continuing CSP with Building CSP loop with anchor residues Number of Ligations found: 36 ACCEPTING loop 15: clash= 0 FF= Building CSP loop with anchor residues Number of Ligations found: 4 ACCEPTING loop 2: clash= 1 FF= Building CSP loop with anchor residues Number of Ligations found: 5 ACCEPTING loop 4: clash= 0 FF= Current Total Energy:
GLY 25 and ARG 30 larger segment GLY 25 and ALA 31 176.0 PP=-23.72 ALA 31 and LEU 34 larger segment ARG 30 and LEU 34 -185.0 PP=-23.27 ASP 39 and ASN 42 208.2 PP=-23.67 GLY 117 and PHE 120 8.1 PP=-23.38
-5003.404 KJ/mol
Figure 2.8.11 Log file for the optimise mode with default alignment.
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many anchor points for loop building will be tested, it can happen that no acceptable solution can be found. In this case, it is most likely that the guiding alignment is so flawed that model building is completely unrealistic. The log file will have a long succession of consecutive attempts (Fig. 2.8.15) followed by the general error message “Aborting: too many unfruitful attempts to find a compatible spare part”.
Optimise Mode for Modeling a Dimer The functional biological unit of the above example is in fact a homodimer. As a mono-
mer was successfully built, it is worth trying to model the homodimer. As ExPDB files contain only one chain, the templates to build multichain models must be either constructed or obtained elsewhere. The PQS Protein Quaternary Structure Server which is at the EBI (http://pqs.ebi.ac.uk/) may give useful information about the oligomeric state of a protein. Indeed, in this example the original PDB entry (1OUN) contains a properly defined biological unit. It could be used directly in the “optimise mode” as the file contains only amino acids. However, this is not the case for
A
B Q21735 1OUNA
2 2
SFNPDYE SVAKAFIQHY YSKFDVGDGM SRAQGLSDLY DPENSYMTFE GDKPIWE QIGSSFIQHY YQLFDN---- DRTQ-LGAIY ID-ASCLTWE . .* * .. .***** * ** *.* *. .* * .* *
Q21735 1OUNA
49 43
GQQAKGRDGI LQKFTTLGFT KIQRAITVID SQPLYDGSIQ VMVLGQLKTD GQQFQGKAAI VEKLSSLPFQ KIQHSITAQD HQPTPDSCII SMVVGQLKAD *** *. .* ..*...* * ***..** * ** *. * **.****.*
Q21735 1OUNA
99 93
EDPINPFSQV FILRPNNQGS YFIGNEIFRL DLHNN -EDPIMGFHQM FLLKNIN-DA WVCTNDMFRL ALHNF **** * *. *.*. * . *..*** ***
Figure 2.8.12 (A) Manually refined alignment between Q21735 and its modeling template (1OUNA) as shown within SPDBV. (B) Complete alignment, shown for clarity.
(...) ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: ProModII: (...) ProModII:
Building CSP loop with anchor residues Building CSP loop with anchor residues Number of Ligations found: 250 ACCEPTING loop 9: clash= 0 FF= Building CSP loop with anchor residues Number of Ligations found: 1 ACCEPTING loop 0: clash= 0 FF= Building CSP loop with anchor residues Building CSP loop with anchor residues Number of Ligations found: 42 ACCEPTING loop 16: clash= 0 FF= Building CSP loop with anchor residues Building CSP loop with anchor residues Number of Ligations found: 21 ACCEPTING loop 19: clash= 0 FF= Current Total Energy:
GLY 25 and ARG 30 VAL 24 and ARG 30 121.8 PP=-23.78 GLN 32 and SER 35 -98.2 PP=-23.40 PRO 40 and SER 43 ASP 39 and SER 43 -53.6 PP=-22.37 ASN 115 and SER 118 ASN 114 and SER 118 -414.2 PP=-22.62
-5753.147 KJ/mol
Figure 2.8.13 Partial Log file for the optimise mode with manually optimised alignment.
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all PDB files, and as the template file has to follow the rules set forth under “Using private templates” it might be necessary to prepare a compatible file using the steps described below: 1. Open the ExPDB file of the first protein chain (1OUNA) in SPDBV.
2. Select all HETATM [Select; Group Kind]. 3. Remove selected residues [Build]. 4. Open the ExPDB file of the second protein chain (1OUNB). 5. Select all residues in both layers [Select]. Holding down the shift key before and while
Figure 2.8.14 Loop from Ile110 to Ans133 rebuilt with the default alignment. (A) Backbone representation where Cα are emphasized with bigger atoms. Hydrogen bonds are drawn as dotted lines. (B) Ribbon representation of the same loop. (C and D) Same region rebuilt after manual optimisation of the alignment. The H-bonding pattern is more regular, which makes a longer β-sheet.
ProModII: Finding Spare-Part ProModII: Finding Spare-Part ProModII: all loops are bad; (...) ProModII: Finding Spare-Part ProModII: all loops are bad; ProModII: Finding Spare-Part ProModII: Finding Spare-Part ProModII: Finding Spare-Part ProModII: all loops are bad; ProModII: *** Aborting: too compatible spare part
Protein Tertiary Structure Modeling
loop with anchor residues THR 83 and THR 94 loop with anchor residues THR 83 and GLN 95 continuing scan with larger segment loop with anchor residues CYS 73 and continuing scan with larger segment loop with anchor residues CYS 73 and loop with anchor residues CYS 73 and loop with anchor residues CYS 73 and continuing scan with larger segment many unfruitful attempts to find a
GLY 99 PRO 100 ARG 101 SER 102
Figure 2.8.15 Typical example of a Log file for a model that failed because a long loop had to be rebuilt (between THR83 and THR94). As no suitable solution was found after several attempts to gain more flexibility using ever increasing number of residues, the modeling procedure stopped. In most cases, the only way of addressing this problem is to move the gap location in SPDBV, and resubmit an “optimise mode” modeling request.
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>Q21735 MSFNPDYESVAKAFIQHYYSKFDVGDGMSRAQGLSDLYDPENSYMTFEGQ QAKGRDGILQKFTTLGFTKIQRAITVIDSQPLYDGSIQVMVLGQLKTDED PINPFSQVFILRPNNQGSYFIGNEIFRLDLHNN; MSFNPDYESVAKAFIQHYYSKFDVGDGMSRAQGLSDLYDPENSYMTFEGQ QAKGRDGILQKFTTLGFTKIQRAITVIDSQPLYDGSIQVMVLGQLKTDED PINPFSQVFILRPNNQGSYFIGNEIFRLDLHNN
Figure 2.8.16 Twice the amino acid sequence of the SwissProt entry Q21735 separated by a semicolon. This allows for dimer modeling.
selecting the menu item will apply the command on all layers. 6. Create a merged layer from selection [Edit]. This will open a new file containing the biological unit. 7. Close the two original layers (1ONUA and 1ONUB). 8. Save the new modeling template [File]. The rest of the modeling procedure is essentially the same as before: 9. Edit the target sequence in a text editor so that it contains twice the same chain, separated by a semicolon (Fig. 2.8.16). 10. Use “Load raw sequence to model” [SwissModel] to open the edited target sequence file. A different chain name is automatically attributed to each chain. 11. Open the alignment window [Windows] and click on the name of the target sequence to select it. 12. Use the “Magic Fit” function [Tools]. Each target chain is aligned onto its corresponding template chain. 13. Alter the alignment for each monomer, so that gaps are placed at the same position as in Figure 2.8.12. 14. Submit the request following the steps defined previously (see Optimise Mode, steps 10 to 12,).
COMMENTARY Sequence Alignment The most frequent reason for unsuccessful modeling attempts by SwissModel is an incorrect sequence alignment. This can be either generated automatically or provided by the user. Although the automatic mode contains a procedure that will try to optimise the alignment, it is still error prone. As the degree of sequence identity between the target sequence and the reference template decreases, the risk of obtaining no model or worse a severely flawed model increases. For those cases, it is
worth casting a critical eye on the alignment with SPDBV. It might be useful to submit several alternate alignments, and then use the WhatCheck and SPDBV analysis tools to select the best model. As a rule, gaps should be placed at the tip of secondary structure elements, to allow maximum flexibility during loop rebuilding. Also, for soluble proteins, it is worth checking that the polarity of the alignment is correct (i.e., all charged side chains are pointing into the solvent while hydrophobic residues should be mainly pointing into the core of the protein).
Model Accuracy As outlined above, the quality of a model greatly depends on the quality of the alignment used by SwissModel. However, this is not the only reason for structural errors in protein models. For instance, large conformational changes cannot be predicted as the model backbone is constructed by extrapolation of the template backbone (Guex and Peitsch, 1997). In order to assess the value and accuracy of modeling procedures, protein models have been compared with their experimentally elucidated counterpart(s). One calls these structures “experimental control structures.” An evaluation of the performance of SwissModel can be obtained from the World Wide Web page in section “Reliability of models” in the form of a table showing the relationship between sequence identity and backbone rmsd. It is important to realize that the accuracy of a protein model is essentially limited by the structural deviation of the best template relative to the experimental control structure. As a consequence, the Cα atoms of protein models which share 35% to 50% sequence identity with their templates, will generally deviate by 1.0 to 1.5 Å from their experimental counter parts, as do similarly related experimental structures (Chothia and Lesk, 1986). Structural differences between predicted and experimental structures have two sources: (1) the errors in-
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herent to the modeling procedures and (2) the variations caused by the molecular environment and data collection method incorporated into experimentally elucidated structures which will be used as modeling templates. Indeed, crystallographic structures of identical proteins can vary not only because of experimental errors and differences in data collection conditions and refinement (illustrated in Tilton et al., 1992), but also because of different crystal-lattice contacts and the presence or absence of ligands. One of the most interesting examples in which several structures of the same protein, determined by different methods, were compared involves interleukin-4 (IL-4; Harrison et al., 1995). This cytokine consists of a 130-residue 4-helix bundle, and its structure was elucidated by x-ray crystallography as well as by NMR. The backbones of three IL-4 crystal structures (PDB entries 1RCB, 2INT, and 1HIK) show an rmsd of 0.4 to 0.9 Å, while those of three IL-4 NMR forms (PDB entries 1ITM, 1CYL, and 2CYK) give an rmsd of 1.2 to 2.8 Å. These values illustrate the structural differences due to experimental procedures and the molecular environment at the time of data collection. Therefore, “a protein model derived by comparative methods cannot be more accurate than the difference between the NMR and crystallographic structure of the same protein” (Harrison et al., 1995). The accuracy of the model determines the extent to which it can be used. “Low-resolution” models, those derived from templates sharing less than 70% residue identity with the target, are helpful to rationalize site-directed mutagenesis experiments aimed at the identification of residues essential for a given molecular recognition and binding process. On the other hand, models based on more closely related templates may be useful during a compound optimisation process. For example, models of closely related species variants of an enzyme, may allow the optimisation of drug specificity.
structures which were used to build a specific part of the model. This “Model B-factor” is included in the PDB file returned to the user and occupies the B-FACTOR field. For visual inspection, open the project file in SPDBV and color the model by B-factor. Parts that appear in red have been reconstructed from scratch (such as insertions and deletions) and are likely to be where most of the problems reside. The color gradient ranges from green, when only one template was used, to dark blue when five templates were used for a specific region. Third, inspect the Ramachandran plot. In addition to the backbone analysis provided by the WhatCheck report, an interactive graphical representation can be provided by SPDBV. As few residues as possible should lay outside the allowed regions (blue), and most residues should be in the optimal regions (yellow). It should be noted that glycine residues (depicted as squares) are allowed to reside out of the allowed regions as they do not have a side chain. Furthermore, within SPDBV, the protein can be colored by “protein problems,” showing prolines with bad backbone conformation (red), and residues laying out of allowed Ramachandranplot regions (yellow). A cluster of such residues is a good indication that there is something wrong in the area. Buried residues with unsatisfied H-bonding capabilities are displayed in orange. Finally, compute the force-field energy of the protein and color the structure accordingly. Residues with a high energy clearly bear some structural problems. These can be of two types: first stereochemical distortions, meaning that they have out-of-average bond length, bond angles, and torsion angles. Second, proximity problems, residues can be too close to other parts of the structure, meaning that they will bear high nonbonded energies. A residue with a high overall energy usually indicates that a wrong side chain rotamer (combination of torsion angles) was selected during the modeling procedure. This can usually be fixed by selecting another rotamer with the mutate tool of SPDBV.
Model Analysis
Protein Tertiary Structure Modeling
SwissModel returns the model with its template(s) and, when selected, a WhatCheck report. The model quality can be assessed at different levels. First, assess the quality of the sequence alignment. The rules described in the “Sequence Alignment” section should be satisfied. Second, SwissModel computes a confidence factor for each atom in the model structures. This factor gives an indication of the number of
Very Large Scale Protein Modeling There is no doubt that the scrutiny of multiple sequence alignments is more informative than the analysis of an isolated sequence. This allows the identification of conserved and variable residues, and improves the rationalization of site-directed mutagenesis experiments. Likewise, comparative structure analysis involving several members of a protein family, as opposed to single-structure inspection, is ex-
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pected to be invaluable for the understanding of their functional differences. Thus it is increasingly valuable to have models for as many members of a protein family as possible. Following 3DCrunch (Guex et al., 1999), the very large scale comparative protein modeling (Peitsch, 1997) experiment run in 1998 (in collaboration with SGI), a database of protein models has been generated (now containing over 80,000 model structures) that can be accessed at http://www.expasy.ch/ swissmod/ SM_3DCrunch.html. Another database containing models of Saccharomyces cerevisiae, Mycoplasma genitalium, and Methanococcus jannaschii proteins can be reached at http://pipe.rockefeller.edu/modbase/ (Sánchez and Sali, 1998). These databases will evolve over the next year to incorporate new models generated with better algorithms, and will allow cross-searching in order to provide a one-stop shop.
LITERATURE CITED Altschul, S.F., Gish, W., Miller, W., Myers, E.W., and Lipman, D.J. 1990. Basic local alignment search tool. J. Mol. Biol. 215:403-410. Bairoch, A. and Apweiler, R. 1999. The SwissProt protein sequence data bank and its supplement TrEMBL in 1999. Nucl. Acids Res. 27:49-54. Chothia, C. and Lesk, A.M. 1986. The relation between the divergence of sequence and structure in proteins. EMBO J. 5:823-826 Guex, N. and Peitsch, M.C. 1997. SwissModel and SwissPdbViewer: An environment for comparative protein modeling. Electrophoresis 18:27142723.
Guex, N., Diemand, A., and Peitsch, M.C. 1999. Protein modeling for all. Trends Biochem. Sci. 24:364-367. Harrison, R.W., Chatterjee, D., and Weber, I.T. 1995. Analysis of six protein structures predicted by comparative modeling techniques. Proteins Struct. Func. Genet. 23:463-471. Hooft, R.W.W., Vriend, G., Sander, C., and Abola, E.E. 1996. Errors in protein structures. Nature 381:272-272. Peitsch, M.C. 1997. Large scale protein modeling and model repository. In Proceedings of the Fifth International Conference on Intelligent Systems for Molecular Biology, Vol. 5 (T. Gaasterland, P. Karp, K. Karplus, C. Ouzonis, C. Sander, and A. Valencia, eds.) pp. 234-236. AAAI Press, Menlo Park, Calif. Sánchez, R. and Sali, A. 1998. Large-scale protein structure modeling of the Saccharomyces cerevisiae genome. Proc. Natl. Acad. Sci. U.S.A. 95:13597-13602. Tilton, R.F., Dewan, J.C., and Petsko, G.A. 1992. Effects of temperature on protein structure and dynamics: X-ray crystallographic studies of the protein ribonuclease-A at nine different temperatures from 98 to 320 K. Biochemistry 31:2469-2481. van Gunsteren, W.F., Billeter, S.R., Eising, A.A., Hünenberger, P.H., Krüger, P., Mark, A.E., Scott, W.R.P., and Tinoni, I.G. 1996. In Biomolecular Simulation: The GROMOS96 Manual and User Guide. Vdf Hochschulverlag ETH, Zurich (http://igc.ethz.ch/gromos/).
Contributed by Nicolas Guex, Torsten Schwede, and Manuel C. Peitsch Glaxo Wellcome Experimental Research Geneva, Switzerland
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Comparative Protein Structure Prediction
UNIT 2.9
Functional characterization of a protein sequence is one of the most frequent problems in biology. This task is usually facilitated by accurate three-dimensional (3-D) structure of the studied protein. In the absence of an experimentally determined structure, comparative or homology modeling can sometimes provide a useful 3-D model for a protein that is related to at least one known protein structure. Comparative modeling predicts the 3-D structure of a given protein sequence (target) based primarily on its alignment to one or more proteins of known structure (templates). The prediction process consists of fold assignment, target-template alignment, model building, and model evaluation. The number of protein sequences that can be modeled, as well as the accuracy of the predictions, is increasing steadily because of growth in the number of known protein structures and improvements in the modeling software. It is currently possible to model, with useful accuracy, significant parts of approximately one-half of all known protein sequences (Pieper et al., 2002). Despite progress in ab initio protein structure prediction (Baker, 2000; Bonneau and Baker, 2001), comparative modeling remains the only method that can reliably predict the 3-D structure of a protein with an accuracy comparable to a low-resolution experimentally determined structure (Marti-Renom et al., 2000). Even models with errors may be useful, because some aspects of function can be predicted from only coarse structural features of a model (Marti-Renom et al., 2000; Baker and Sali, 2001). There are several computer programs and Web servers that automate the comparative modeling process. The first Web server for automated comparative modeling was the Swiss-Model server (http://www.expasy.ch/swissmod/), followed by CPHModels (http://www.cbs.dtu.dk/services/CPHmodels/), SDSC1 (http://cl.sdsc.edu/hm.html), FAMS (http://physchem.pharm.kitasato-u.ac.jp/FAMS/fams.html), MODWEB (http://guitar.rockefeller.edu/modweb/), and ESyPred3D (http://www.fundp.ac.be/urbm/ bioinfo/esypred/). Several of these servers are being evaluated by EVA-CM (http:// cubic.bioc.columbia.edu/eva), a Web server for assessing protein structure prediction methods in an automated, continuous and large-scale fashion (Eyrich et al., 2001; Marti-Renom et al., 2002). While the Web servers are convenient and useful, the best results in the difficult or unusual modeling cases, such as problematic alignments, modeling of loops, existence of multiple conformational states, and modeling of ligand binding, are still obtained by nonautomated, expert use of the various modeling tools. A number of resources useful in comparative modeling are listed in Table 2.9.1 (an online version of this table is available at http://guitar.rockefeller.edu/bioinformatics_resources.shtml). Next, we describe generic considerations in all four steps of comparative modeling (see Steps in Comparative Modeling, below, and Fig. 2.9.1), typical modeling errors (see Errors in Comparative Models, below, and Fig. 2.9.2), and applications of comparative protein structure models (see Applications of Comparative Modeling, below, and Fig. 2.9.3). Finally, we illustrate these considerations in practice by discussing in detail one application of our program MODELLER (Sali and Blundell, 1993; Sali and Overington, 1994; Fiser et al., 2000).
Computational Analysis Contributed by Marc A. Marti-Renom, Bozidar Yerkovich, and Andrej Sali Current Protocols in Protein Science (2002) 2.9.1-2.9.22 Copyright © 2002 by John Wiley & Sons, Inc.
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Table 2.9.1
Programs and Web Servers Useful in Comparative Modeling
Name
Typea WWW addressb
Databases CATH GenBank GeneCensus MODBASE PDB PRESAGE SCOP TrEMBL
S S S S S S S S
http://www.biochem.ucl.ac.uk/bsm/cath/ http://www.ncbi.nlm.nih.gov/Genbank/ http://bioinfo.mbb.yale.edu/genome/ http://guitar.rockefeller.edu/modbase/ http://www.rcsb.org/pdb/ http://presage.berkeley.edu http://scop.mrc-lmb.cam.ac.uk/scop/ http://srs.ebi.ac.uk
Orengo et al. (2002) Blundell et al. (1987) Gerstein and Levitt (1997) Pieper et al. (2002) Westbrook et al. (2002) Brenner et al. (1999) Lo Conte et al. (2002) Bairoch and Apweiler (2000)
Template search 123D BLAST DALI FastA MATCHMAKER PHD, TOPITS PROFIT THREADER FRSVR
S S S S P S P P S
http://123d.ncifcrf.gov/123D+.html http://www.ncbi.nlm.nih.gov/BLAST/ http://www2.ebi.ac.uk/dali/ http://www.ebi.ac.uk/fasta33/ http://bioinformatics.burnham-inst.org http://cubic.bioc.columbia.edu/predictprotein/ http://www.came.sbg.ac.at http://insulin.brunel.ac.uk/~jones/threader.html http://fold.doe-mbi.ucla.edu
Alexandrov et al. (1996) Altschul et al. (1990) Holm and Sander (1999) Pearson (1990) Godzik et al. (1992) Rost (1995) Flockner et al. (1995) Jones et al. (1992) Fischer and Eisenberg (1996)
http://searchlauncher.bcm.tmc.edu http://www.ncbi.nlm.nih.gov/gorf/bl2.html http://blocks.fhcrc.org/blocks/blockmkr/make_blocks.html
Smith et al. (1996) Altschul et al. (1997) Henikoff et al. (1995)
http://www2.ebi.ac.uk/clustalw/ http://www2.ebi.ac.uk/fasta3/ http://pbil.ibcp.fr
Thompson et al. (1994) Pearson (1990) Corpet (1988)
http://www.tripos.com/software/composer.html http://www.congenomics.com/congen/congen.html http://www.molsoft.com http://www.accelrys.com http://guitar.rockefeller.edu/modeller/modeller.html http://www.accelrys.com http://www.tripos.com http://www.fccc.edu/research/labs/dunbrack/scwrl/ http://www.expasy.org/swissmod/SWISS-MODEL.html http://www.cmbi.kun.nl/whatif/
Sutcliffe et al. (1987) Bruccoleri and Karplus (1990) —c —d Sali and Blundell (1993)e —d —f Bower et al. (1997) Peitsch and Jongeneel (1993) Vriend (1990)
http://www.fundp.ac.be/sciences/biologie/bms/CGI/anolea.html http://urchin.bmrb.wisc.edu/~jurgen/aqua/ http://biotech.embl-heidelberg.de:8400 http://www.doe-mbi.ucla.edu/Services/ERRAT/ http://www.biochem.ucl.ac.uk/~roman/procheck/procheck.html http://www.came.sbg.ac.at http://www.ucmb.ulb.ac.be/UCMB/PROVE http://www.ysbl.york.ac.uk/~oldfield/squid/ http://www.doe-mbi.ucla.edu/Services/Verify_3D/ http://www.sander.embl-heidelberg.de/whatcheck/
Melo and Feytmans (1998) Laskowski et al. (1996) Laskowski et al. (1998) Colovos and Yeates (1993) Laskowski et al. (1998) Sippl (1993) Pontius et al. (1996) Oldfield (1992) Luthy et al. (1992) Hooft et al. (1996a)
Sequence alignment BCM SERVER S BLAST2 S BLOCK S MAKER CLUSTAL S FASTA3 S MULTALIN S Modeling COMPOSER P CONGEN P ICM P InsightII P MODELLER P QUANTA P SYBYL P SCWRL P SWISS-MOD S WHAT IF P Model evaluation ANOLEA S AQUA P S BIOTECHg ERRAT S PROCHECK P ProsaII P PROVE S SQUID P VERIFY3D S WHATCHECK P
Reference
continued
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Table 2.9.1
Name
Programs and Web Servers Useful in Comparative Modeling, continued
Typea WWW addressb
Methods evaluation CASP S CAFASP S EVA S LiveBench S
http://predictioncenter.llnl.gov http://cafasp.bioinfo.pl http://cubic.bioc.columbia.edu/eva/ http://bioinfo.pl/LiveBench/
Reference Moult et al. (2001) Fischer et al. (2001) Eyrich et al. (2001) Bujnicki et al. (2001)
aS, server; P, program. bSome of the sites are mirrored on additional computers. cMolSoft Inc., San Diego, Calif. dAccelrys Inc., San Diego, Calif. eSee Internet Resources for additional information. fTripos Inc., St Louis, Mo. gThe BIOTECH server uses PROCHECK and WHATCHECK for structure evaluation.
STEPS IN COMPARATIVE MODELING Fold Assignment and Template Selection The starting point in comparative modeling is to identify all protein structures related to the target sequence, and then select those structures that will be used as templates. This step is facilitated by numerous protein sequence and structure databases and by the use of database-scanning software available on the Web (Altschul et al., 1994; Barton, 1998; Holm and Sander, 1996; also see Table 2.9.1). Templates can be found using the target sequence as a query for searching structure databases such as the Protein Data Bank (Westbrook et al., 2002), SCOP (Lo Conte et al., 2002), DALI (Holm and Sander, 1999), and CATH (Orengo et al., 2002). The probability of finding a related protein of known structure for a sequence picked randomly from a genome ranges from 20% to 70% (Fischer and Eisenberg, 1997; Huynen et al., 1998; Jones, 1999; Rychlewski et al., 1998; Sanchez and Sali, 1998; Pieper et al., 2002). There are three main classes of protein comparison methods that are useful in fold identification. The first class includes the methods that compare the target sequence with each of the database sequences independently, using pairwise sequence-sequence comparison (Apostolico and Giancarlo, 1998). The performance of these methods in searching for related protein sequences and structures has been evaluated exhaustively. Frequently used programs in this class include FASTA (Pearson and Lipman, 1988; Pearson, 1995) and BLAST (UNIT 2.5; Altschul et al., 1990). The second set of methods relies on multiple sequence comparisons to improve the sensitivity of the search (Altschul et al., 1997; Henikoff and Henikoff, 1994; Gribskov, 1994; Krogh et al., 1994; Rychlewski et al., 1998). A widely used program in this class is PSI-BLAST (Altschul et al., 1997), which iteratively expands the set of homologs of the target sequence. For a given sequence, an initial set of homologs from a sequence database is collected, a weighted multiple alignment is made from the query sequence and its homologs, a position-specific scoring matrix is constructed from the alignment, and the matrix is used to search the database for additional homologs. These steps are repeated until no additional homologs are found. In comparison to BLAST, PSI-BLAST finds homologs of known structure for approximately twice as many sequences (Park et al., 1998; Sternberg et al., 1999).
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start target sequence
template structure
identify related structures
select templates
align target sequence with template structures
alignment target
KLTDSQNFDEYMKALGVGFATRQVGNVTKPTVIISQEGGKVV...
template
KLVSSENFDDYMKEVGVGFATRKVAGMAKPNMIISVNGDLVT...
build a model for the target using information from template structures target model
no
model OK?
yes
end
Pseudo energy
evaluate the model
Residue index
Figure 2.9.1 Steps in comparative protein structure modeling.
Comparative Protein Structure Prediction
The third class of methods are the so-called threading or 3-D template matching methods (Bowie et al., 1991; Jones et al., 1992; Godzik et al., 1992; reviewed in Jones, 1997; Smith et al., 1997; Torda, 1997; Levitt, 1997; David et al., 2000). These methods rely on pairwise comparison of a protein sequence and a protein of known structure. Whether or not a given target sequence adopts any one of the many known 3-D folds is predicted by an optimization of the alignment with respect to a structure-dependent scoring function, independently for each sequence-structure pair—i.e., the target sequence is threaded through a library of 3-D folds. These methods are especially useful when there are no
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sequences clearly related to the modeling target, and thus the search cannot benefit from the increased sensitivity of the sequence profile methods. A useful fold-assignment approach is to accept an uncertain assignment provided by any of the methods, build a full-atom comparative model of the target sequence based on this match, and make the final decision about whether or not the match is real by evaluating the resulting comparative model (Sanchez and Sali, 1997; Guenther et al., 1997; Miwa et al., 1999). Once a list of all related protein structures has been obtained, it is necessary to select those templates that are appropriate for the given modeling problem. Usually, a higher overall sequence similarity between the target and the template sequence yields a better model. In any case, several other factors should be taken into account when selecting the templates:
A
B
D
C
E
Figure 2.9.2 Typical errors in comparative modeling. (A). Errors in side-chain packing. The Trp 109 residue in the crystal structure of mouse cellular retinoic acid binding protein I (thin line) is compared with its model (thick line), and with the template mouse adipocyte lipid-binding protein (broken line). (B) Distortions and shifts in correctly aligned regions. A region in the crystal structure of mouse cellular retinoic acid binding protein I is compared with its model and with the template fatty acid binding protein using the same representation as in panel A. (C) Errors in regions without a template. The Cα trace of the 112–117 loop is shown for the X-ray structure of human eosinophil neurotoxin (thin line), its model (thick line), and the template ribonuclease A structure (residues 111–117; broken line). (D) Errors due to misalignments. The N-terminal region in the crystal structure of human eosinophil neurotoxin (thin line) is compared with its model (thick line). The corresponding region of the alignment with the template ribonuclease A is shown. The black lines show correct equivalences, that is, residues whose Cα atoms are within 5 Å of each other in the optimal least-squares superposition of the two X-ray structures. The “a” characters in the bottom line indicate helical residues. (E) Errors due to an incorrect template. The X-ray structure of α-trichosanthin (thin line) is compared with its model (thick line) which was calculated using indole-3-glycerophosphate synthase as the template.
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syy
[[sty[Å [[syyw
}
STUVWXYZ[\]T]^WTX\ _`\a]YXS_ V`SXZYXYZ[]YV[X_bcdeXYZ ^XZ]YVS Vd\fXYZ[dg[_]\cd_d^`\U^`Sh bc`VX\TXdY[dg[bcdT`XY[b]cTY`cS
uy
eXcTU]^[S\c``YXYZ[]YV Vd\fXYZ[dg[S_]^^[^XZ]YVS
qy
ndV`^[]\\Uc]\W
w[`~U`Y\`[XV`YTXTW
[[stu[Å [[vuw
V`gXYXYZ[]YTXidVW `bXTdb`S
[qtu[Å [xyw
{
_d^`\U^]c[c`b^]\`_`YT[XY jkc]W[\cWST]^^dZc]baW V`SXZYXYZ[\aX_`c]Sh[ST]i^` \cWST]^^Xl]i^`[e]cX]YTS SUbbdcTXYZ[SXT`kVXc`\T`V _UT]Z`Y`SXS c`gXYXYZ[mno[STcU\TUc`S
XYSXZYXgX\]YT[S`~U`Y\`[SX_X^]cXTW
gXTTXYZ[XYTd[^dpkc`Sd^UTXdY `^`\TcdY[V`YSXTW
Comparative Protein Structure Prediction
zty[Å xy[]]
|
gXYVXYZ[gUY\TXdY]^[SXT`S[iW qkr[_dTXg[S`]c\aXYZ STcU\TUc`[gcd_[Sb]cS` `jb`cX_`YT]^[c`STc]XYTS ]YYdT]TXYZ[gUY\TXdY[iW gd^V[]SSXZY_`YT `ST]i^XSaXYZ[`ed^UTXdY]cW c`^]TXdYSaXbS
Figure 2.9.3 Accuracy and application of protein structure models. The vertical axis indicates the different ranges of applicability of comparative protein structure modeling, the corresponding accuracy of protein structure models, and their sample applications. (A) The docosahexaenoic fatty acid ligand (darker strand) was docked into a high-accuracy comparative model of brain lipid-binding protein (right), modeled based on its 62% sequence identity to the crystallographic structure of adipocyte lipid-binding protein (PDB code, 1adl). A number of fatty acids were ranked for their affinity to brain lipid-binding protein consistently with site-directed mutagenesis and affinity chromatography experiments (Xu et al., 1996), even though the ligand-specificity profile of this protein is different from that of the template structure. Typical overall accuracy of a comparative model in this range of sequence similarity is indicated by a comparison of a model for adipocyte fatty acid binding protein with its actual structure (left). (B) A putative proteoglycan binding patch was identified on a medium-accuracy comparative model of mouse mast cell protease 7 (right), modeled based on its 39% sequence identity to the crystallographic structure of bovine pancreatic trypsin (PDB code, 2ptn) that does not bind proteoglycans. The prediction was confirmed by site-directed mutagenesis and heparin-affinity chromatography experiments (Matsumoto et al., 1995). Typical accuracy of a comparative model in this range of sequence similarity is indicated by a comparison of a trypsin model with the actual structure. (C) A molecular model of the whole yeast ribosome (right) was calculated by fitting atomic rRNA and protein models into the electron density of the 80S ribosomal particle, obtained by electron microscopy at 15 Å resolution (Beckmann et al., 2001). Most of the models for 40 out of the 75 ribosomal proteins were based on approximately 30% sequence identity to their template structures. Typical accuracy of a comparative model in this range of sequence similarity is indicated by a comparison of a model for a domain in L2 protein from B. Stearothermophilus with the actual structure (PDB code, 1rl2).
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1. The family of proteins, which includes the target and the templates, can frequently be organized in subfamilies. The construction of a multiple alignment and a phylogenetic tree (Felsenstein, 1985) can help in selecting the template from the subfamily that is closest to the target sequence. 2. The template environment should be compared to the required environment for the model. The term “environment” is used in a broad sense and includes all factors that determine protein structure, except its sequence (e.g., solvent, pH, ligands, and quaternary interactions). 3. The quality of the experimental template structure is another important factor in template selection. The resolution and the R-factor of a crystallographic structure and the number of restraints per residue for an NMR structure are indicative of its accuracy. The priority of the criteria for template selection depends on the purpose of the comparative model. For instance, if a protein-ligand model is to be constructed, the choice of the template that contains a similar ligand is probably more important than the resolution of the template. On the other hand, if the model is to be used to analyze the geometry of the active site of an enzyme, it is preferable to use a high-resolution template. It is not necessary to select only one template. In fact, the use of several templates approximately equidistant from the target sequence generally increases the model accuracy (Srinivasan and Blundell, 1993; Sanchez and Sali, 1997). Target-Template Alignment Most fold-assignment methods produce an alignment between the target sequence and template structures. However, this is often not the optimal target-template alignment for comparative modeling. Searching methods are usually tuned for detection of remote relationships, not for optimal alignments. Therefore, once templates have been selected, a specialized method should be used to align the target sequence with the template structures (Taylor, 1996; Holm and Sander, 1996; Briffeuil et al., 1998; Baxevanis, 1998; Smith, 1999). For closely related protein sequences with identity higher than 40%, the alignment is almost always correct. Regions of low local sequence similarity become common when the overall sequence identity is below 40% (Saqi et al., 1998). The alignment becomes difficult in the “twilight zone” of less than 30% sequence identity (Rost, 1999). As the sequence similarity decreases, alignments contain an increasingly large number of gaps and alignment errors, regardless of whether they are prepared automatically or manually. For example, only 20% of the residues are likely to be correctly aligned when two proteins share 30% sequence identity (Johnson and Overington, 1993). Maximal effort to obtain the most accurate alignment possible is needed, because no current comparative modeling method can recover from an incorrect alignment. There are a great variety of protein sequence alignment methods, many of which are based on dynamic programming techniques (Needleman and Wunsch, 1970; Smith and Waterman, 1981). A frequently used program for multiple sequence alignment is CLUSTAL (Thompson et al., 1994; Higgins et al., 1996), which is also available as a Web server (Table 2.9.1). In the more difficult alignment cases, it is frequently beneficial to rely on multiple structure and sequence information (Barton and Sternberg, 1987; Taylor et al., 1994). First, the alignment of the potential templates is prepared by superposing their structures. Next, the sequences that are clearly related to the templates and are easily aligned with them are added to the alignment. The same is done for the target sequence. Finally, the two profiles are aligned with each other, taking structural information into account as much as possible (Koretke et al., 1998; Thompson et al., 1994; Yang and Honig, 2000; Al Lazikani et al., 2001a).
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Model Building Once an initial target-template alignment has been built, a variety of methods can be used to construct a 3-D model for the target protein. The original and still widely used method is modeling by rigid-body assembly (Browne et al., 1969; Greer, 1990; Blundell et al., 1987). Another family of methods, based on modeling by segment matching, relies on the approximate positions of conserved atoms in the templates (Jones and Thirup, 1986; Unger et al., 1989; Claessens et al., 1989; Levitt, 1992). The third group of methods, modeling by satisfaction of spatial restraints, uses either distance geometry or optimization techniques to satisfy spatial restraints obtained from the alignment (Havel and Snow, 1991; Srinivasan et al., 1993; Sali and Blundell, 1993; Brocklehurst and Perham, 1993; Aszodi and Taylor, 1996; Kolinski et al., 2001). Accuracies of the various model-building methods are relatively similar when used optimally (Marti-Renom et al., 2002). Other factors, such as template selection and alignment accuracy, usually have a larger impact on the model accuracy, especially for models based on less than 40% sequence identity to the templates. There are many reviews of comparative model building methods (Blundell et al., 1987; Sanchez and Sali, 1997; Sali, 1995; Johnson et al., 1994; Bajorath et al., 1993; Marti-Renom et al., 2000; Al Lazikani et al., 2001b). A number of programs and Web servers for comparative modeling are listed in Table 2.9.1. Model Evaluation The quality of the predicted model determines the information that can be extracted from it. Thus, estimating the accuracy of 3-D protein models is essential for interpreting them. The model can be evaluated as a whole, as well as in the individual regions. There are many model-evaluation programs and servers (Laskowski et al., 1998; Wilson et al., 1993; also see Table 2.9.1). The first step in model evaluation is to determine if the model has the correct fold (Sanchez and Sali, 1998). A model will have the correct fold if the correct template is picked and if that template is aligned at least approximately correctly with the target sequence. The confidence in the fold of a model is generally increased by a high sequence similarity with the closest template, a significant energy-based Z-score (Sippl, 1993; Sanchez and Sali, 1998), or conservation of the key functional or structural residues in the target sequence. Once the fold of a model is accepted, a more detailed evaluation of the overall model accuracy can be obtained based on the similarity between the target and template sequences (Sanchez and Sali, 1998). Sequence identity above 30% is a relatively good predictor of the expected accuracy. The reasons are the well-known relationship between structural and sequence similarities of two proteins (Chothia and Lesk, 1986), the “geometrical” nature of modeling that forces the model to be as close to the template as possible (Sali and Blundell, 1993), and the inability of any current modeling procedure to recover from an incorrect alignment (Sanchez and Sali, 1997). The dispersion of the model-target structural overlap increases with the decrease in sequence identity. If the target-template sequence identity falls below 30%, the sequence identity becomes unreliable as a measure of accuracy of a single model. Models that deviate significantly from the average accuracy are frequent. It is in such cases that model-evaluation methods are particularly useful.
Comparative Protein Structure Prediction
In addition to the target-template sequence similarity, the environment can strongly influence the accuracy of a model. For instance, some calcium-binding proteins undergo large conformational changes when bound to calcium. If a calcium-free template is used to model the calcium-bound state of the target, it is likely that the model will be incorrect, irrespective of the target-template similarity or accuracy of the template structure (Pawlowski et al., 1996). This qualification also applies to the experimental determination
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of protein structure; a structure must be determined in the functionally meaningful environment. A basic requirement for a model is to have good stereochemistry. Some useful programs for evaluating stereochemistry (see Table 2.9.1 for URLs) are PROCHECK (Laskowski et al., 1998), PROCHECK-NMR (Laskowski et al., 1996), AQUA (Laskowski et al., 1996), SQUID (Oldfield, 1992), and WHATCHECK (Hooft et al., 1996b). The features of a model that these programs check include bond lengths, bond angles, peptide bond and side-chain ring planarities, chirality, main-chain and side-chain torsion angles, and clashes between nonbonded pairs of atoms. There are also methods for testing 3-D models that implicitly take into account many spatial features compiled from high-resolution protein structures. These methods are based on 3-D profiles and statistical potentials of mean force (Sippl, 1990; Luthy et al., 1992). Programs implementing this approach (see Table 2.9.1 for URLs) include VERIFY3D (Luthy et al., 1992), PROSAII (Sippl, 1993), HARMONY (Topham et al., 1994), and ANOLEA (Melo and Feytmans, 1998). The programs evaluate the environment of each residue in a model with respect to the expected environment as found in the high-resolution X-ray structures. There is a concern about the theoretical validity of the energy profiles for detecting regional errors in models (Fiser et al., 2000). It is likely that the contributions of the individual residues to the overall free energy of folding vary widely, even when normalized by the number of atoms or interactions made. If this expectation is correct, the correlation between the model errors and energy peaks is greatly weakened, resulting in the loss of predictive power of the energy profile. Despite these concerns, error profiles have been useful in some applications (Miwa et al., 1999). ERRORS IN COMPARATIVE MODELS As the similarity between the target and the templates decreases, the errors in the model increase. Errors in comparative models can be divided into five categories (Sanchez and Sali, 1997; also see Fig. 2.9.2): 1. Errors in side-chain packing (Fig. 2.9.2A). As the sequences diverge, the packing of side chains in the protein core changes. Sometimes even the conformation of identical side chains is not conserved, a pitfall for many comparative modeling methods. Side-chain errors are critical if they occur in regions that are involved in protein function, such as active sites and ligand-binding sites. 2. Distortions and shifts in correctly aligned regions (Fig. 2.9.2B). As a consequence of sequence divergence, the main-chain conformation changes, even if the overall fold remains the same. Therefore, it is possible that in some correctly aligned segments of a model the template is locally different (<3 Å) from the target, resulting in errors in that region. The structural differences are sometimes not due to differences in sequence, but are a consequence of artifacts in structure determination or structure determination in different environments (e.g., packing of subunits in a crystal). The simultaneous use of several templates can minimize this kind of an error (Srinivasan and Blundell, 1993; Sanchez and Sali, 1997). 3. Errors in regions without a template (Fig. 2.9.2C). Segments of the target sequence that have no equivalent region in the template structure (i.e., insertions or loops) are the most difficult regions to model. If the insertion is relatively short, less than 9 residues long, some methods can correctly predict the conformation of the backbone (van Vlijmen and Karplus, 1997; Fiser et al., 2000). Conditions for successful prediction are the correct alignment and an accurately modeled environment surrounding the insertion.
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4. Errors due to misalignments (Fig. 2.9.2D). The largest source of errors in comparative modeling are misalignments, especially when the target-template sequence identity decreases below 30%. However, alignment errors can be minimized in two ways. First, it is usually possible to use a large number of sequences to construct a multiple alignment, even if most of these sequences do not have known structures. Multiple alignments are generally more reliable than pairwise alignments (Barton and Sternberg, 1987; Taylor et al., 1994). The second way of improving the alignment is to iteratively modify those regions in the alignment that correspond to predicted errors in the model (Sanchez and Sali, 1997). 5. Incorrect templates (Fig. 2.9.2E). This is a potential problem when distantly related proteins are used as templates (i.e., <25% sequence identity). Distinguishing between a model based on an incorrect template and a model based on an incorrect alignment with a correct template is difficult. In both cases, the evaluation methods will predict an unreliable model. The conservation of the key functional or structural residues in the target sequence increases the confidence in a given fold assignment. An informative way to test protein structure modeling methods is provided by EVA-CM (Eyrich et al., 2001) and LiveBench (Bujnicki et al., 2001) See Table 2.9.1 for the URLs of the EVA and LiveBench servers. APPLICATIONS OF COMPARATIVE MODELING Comparative modeling is an increasingly efficient way to obtain useful information about the proteins of interest. For example, comparative models can be helpful in designing mutants to test hypotheses about a protein’s function (Boissel et al., 1993; Wu et al., 1999), identifying active and binding sites (Sheng et al., 1996), identifying, designing, and improving ligands for a given binding site (Ring et al., 1993), modeling substrate specificity (Xu et al., 1996), predicting antigenic epitopes (Sali et al., 1993), simulating protein-protein docking (Vakser, 1997), inferring function from a calculated electrostatic potential around the protein (Matsumoto et al., 1995), facilitating molecular replacement in X-ray structure determination (Howell et al., 1992), refining models based on NMR constraints (Modi et al., 1996), testing and improving a sequence-structure alignment (Wolf et al., 1998), confirming a remote structural relationship (Guenther et al., 1997; Miwa et al., 1999), and rationalizing known experimental observations. For exhaustive reviews of comparative modeling applications see Johnson et al. (1994), Fiser and Sali (2002), and Baker and Sali (2001). Fortunately, a 3-D model does not have to be absolutely perfect to be helpful in biology, as demonstrated by the applications listed above. However, the type of a question that can be addressed with a particular model does depend on its accuracy (Figure 2.9.3). Three levels of model accuracy and some of the corresponding applications are as follows. 1. At the low end of the spectrum, there are models that are based on less than 30% sequence identity and that sometimes have less than 50% of their atoms within 3.5 Å of their correct positions. Such models still have the correct fold, which is frequently sufficient to predict its approximate biochemical function. More specifically, only nine out of 80 fold families known in 1994 contained proteins (domains) that were not in the same functional class, although 32% of all protein structures belonged to one of the nine superfolds (Orengo et al., 1994). Models in this low range of accuracy, combined with model evaluation, can be used for confirming or rejecting a match between remotely related proteins (Sanchez and Sali, 1997, 1998). Comparative Protein Structure Prediction
2. In the middle of the accuracy spectrum are the models based on ~30% to 50% sequence identity, corresponding to 85% of the residues modeled within 3.5 Å of their correct positions. Fortunately, the active and binding sites are frequently more
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conserved than the rest of the fold, and are thus modeled more accurately (Sanchez and Sali, 1998). In general, medium-resolution models frequently allow refinement of the functional prediction based on sequence alone, because ligand binding is most directly determined by the structure of the binding site rather than by its sequence. It is frequently possible to correctly predict important features of the target protein that do not occur in the template structure. For example, the location of a binding site can be predicted from clusters of charged residues (Matsumoto et al., 1995), and the size of a ligand may be predicted from the volume of the binding site cleft (Xu et al., 1996). Medium-resolution models can also be used to construct site-directed mutants with altered or destroyed binding capacity, which in turn could test hypotheses about the sequence-structure-function relationships. Other problems that can be addressed with medium-resolution comparative models include designing proteins that have compact structures without long tails, loops, and exposed hydrophobic residues, for better crystallization, or with added disulfide bonds for extra stability. 3. The high end of the accuracy spectrum corresponds to models based on more than 50% sequence identity. The average accuracy of these models approaches that of low-resolution X-ray structures (3 Å resolution) or medium-resolution NMR structures (10 distance restraints per residue; Sanchez and Sali, 1997). The alignments on which these models are based generally contain almost no errors. In addition to the already listed applications, high-quality models can be used for docking of small ligands (Ring et al., 1993) or whole proteins onto a given protein (Totrov and Abagyan, 1994; Vakser, 1997). EXAMPLE OF COMPARATIVE MODELING: MODELING LACTATE DEHYDROGENASE FROM TRICHOMONAS VAGINALIS BASED ON A SINGLE TEMPLATE This section contains an example of a typical comparative modeling application. It demonstrates each of the five steps of comparative modeling, using the program MODELLER6. All files described in this section, including the MODELLER program, are available at http://guitar.rockefeller.edu/modeller/tutorials.shtml (also see Internet Resources). A novel gene for lactate dehydrogenase was identified from the genomic sequence of Trichomonas vaginalis (TvLDH). The corresponding protein had a higher similarity to the malate dehydrogenase of the same species (TvMDH) than to any other LDH. The authors hypothesized that TvLDH arose from TvMDH by convergent evolution relatively recently (Wu et al., 1999). Comparative models were constructed for TvLDH and TvMDH to study the sequences in the structural context and to suggest site-directed mutagenesis experiments for elucidating specificity changes in this apparent case of convergent evolution of enzymatic specificity. The native and mutated enzymes were expressed and their activities were compared (Wu et al., 1999). Searching for Structures Related to TvLDH It is necessary to put the target TvLDH sequence into the PIR format readable by MODELLER (see Internet Resources for MODELLER Web site; file TvLDH.ali). The first line of the file (see Fig. 2.9.4) contains the sequence code, in the format >P1;code. The second line, with ten fields separated by colons, generally contains information about the structure file, if applicable. Only two of these fields are used for sequences: sequence (indicating that the file contains a sequence without known structure) and TvLDH (the model file name). The rest of the file contains the sequence of TvLDH, with a “*” marking its end. A search for potentially related sequences of known structure can be performed by the SEQUENCE_SEARCH command of MODELLER. The following
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>P1;TvLDH sequence:TvLDH:::::::0.00: 0.00 MSEAAHVLITGAAGQIGYILSHWIASGELYG-DRQVYLHLLDIPPAMNRLTALTMELEDCAFPHLAGFVATTDPK AAFKDIDCAFLVASMPLKPGQVRADLISSNSVIFKNTGEYLSKWAKPSVKVLVIGNPDNTNCEIAMLHAKNLKPE NFSSLSMLDQNRAYYEVASKLGVDVKDVHDIIVWGNHGESMVADLTQATFTKEGKTQKVVDVLDHDYVFDTFFKK IGHRAWDILEHRGFTSAASPTKAAIQHMKAWLFGTAPGEVLSMGIPVPEGNPYGIKPGVVFSFPCNVDKEGKIHV VEGFKVNDWLREKLDFTEKDLFHEKEIALNHLAQGG*
Figure 2.9.4 File TvLDH.ali. Sequence file in the PIR MODELLER format.
SET SEARCH_RANDOMIZATIONS = 100 SEQUENCE_SEARCH FILE = 'TvLDH.ali', ALIGN_CODES = 'TvLDH', DATA_FILE = ON
Figure 2.9.5 File seqsearch.top. The TOP script file for template research using MODELLER.
script uses the query sequence TvLDH assigned to the variable ALIGN_CODES from the file TvLDH.ali assigned to the variable FILE (file seqsearch.top; Fig. 2.9.5). The SEQUENCE_SEARCH command has many options (see Internet Resources for MODELLER Web site), but in this example only SEARCH_RANDOMIZATIONS and DATA_FILE are set to non-default values. SEARCH_RANDOMIZATIONS specifies the number of times the query sequence is randomized during the calculation of the significance score for each sequence-sequence comparison. The higher the number of randomizations, the more accurate the significance score. DATA_FILE = ON triggers creation of an additional summary output file (seqsearch.dat). Selecting a Template The output of the seqsearch.top script is written to the seqsearch.log file. MODELLER always produces a log file. Errors and warnings in log files can be found by searching for the E> and W> strings, respectively. At the end of the log file, MODELLER lists the hits, sorted by alignment significance. Because the log file is sometimes very long, a separate data file (seqsearch.dat) is created that contains the summary of the search (Fig. 2.9.6). The example in Figure 2.9.6 shows only the top 10 hits from such a file.
Comparative Protein Structure Prediction
The most important columns in the SEQUENCE_SEARCH output are the CODE_2, %ID, and SIGNI columns. The CODE_2 column reports the code of the PDB sequence that was compared with the target sequence. The PDB code in each line is the representative of a group of PDB sequences that share 40% or more sequence identity to each other and have less than 30 residues or 30% sequence length difference. All the members of the group can be found in the MODELLER CHAINS_3.0_40_XN.grp file. The LEN1 and LEN2 are lengths of the protein sequences in the CODE_1 and CODE_2 columns, respectively. NID represents the number of aligned residues. The %ID1 and %ID2 columns report the percentage sequence identities between TvLDH and a PDB sequence, normalized by their lengths, respectively. In general, a %ID value above ∼25% indicates a potential template unless the alignment is short (i.e., less than 100 residues). A better measure of the significance of the alignment is given by the SIGNI column (see Internet Resources for MODELLER Web site). A value above 6.0 is generally significant irrespective of the sequence identity and length. In this example, one protein family represented by 1bdmA shows significant similarity with the target sequence, at more than 40%
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# CODE_1
CODE_2
1 2 3 4 5 6 7 8 9 10
1bdmA 1lldA 1ceqA 2hlpA 1ldnA 1hyhA 2cmd 1db3A 91dtA 1cdb
TvLDH TvLDH TvLDH TvLDH TvLDH TvLDH TvLDH TvLDH TvLDH TvLDH
LEN1 LEN2 NID
335 335 335 335 335 335 335 335 335 335
318 153 313 103 304 95 303 86 316 91 297 88 312 108 335 91 331 95 105 69
% IDI
45.7 30.7 28.4 25.7 27.2 26.3 32.2 27.2 28.4 29.6
%ID2
48.1 32.9 31.3 28.4 28.8 29.6 34.6 27.2 28.7 65.7
SCORE
SIGNI
212557. 183190. 179636. 177791. 180669. 175969. 182079. 181928. 181720. 80141.
28.9 10.1 9.2 8.9 7.4 6.9 6.6 4.9 4.7 3.8
Figure 2.9.6 Top 10 hits from file seqsearch.dat. The summary output file for the SEQUENCE_SEARCH command in MODELLER.
sequence identity. While some other hits are also significant, the differences between 1bdmA and other top scoring hits are so pronounced that we use only the first hit as the template. As expected, 1bdmA is a malate dehydrogenase (from a thermophilic bacterium). Other structures closely related to 1bdmA (and thus not scanned against by SEQUENCE_SEARCH) can be extracted from the CHAINS_3.0_40_XN.grp file: 1b8vA, 1bmdA, 1b8uA, 1b8pA, 1bdmA, 1bdmB, 4mdhA, 5mdhA, 7mdhA, 7mdhB, and 7mdhC. All these proteins are malate dehydrogenases. During the project, all of them, as well as other malate and lactate dehydrogenase structures, were compared and considered as templates (there were 19 structures in total). However, for the sake of illustration, we will investigate only four of the proteins that are sequentially most similar to the target: 1bmdA, 4mdhA, 5mdhA, and 7mdhA. The script in Figure 2.9.7 performs all pairwise comparisons among the selected proteins (file compare.top). The READ_ALIGNMENT command shown in Figure 2.9.7 reads the protein sequences and information about their PDB files. MALIGN calculates their multiple sequence alignment, used as the starting point for the multiple structure alignment. The MALIGN3D command performs an iterative least-squares superposition of the four 3-D structures. The COMPARE command compares the structures according to the alignment constructed by MALIGN3D. It does not make an alignment, but it calculates the RMS and DRMS deviations between atomic positions and distances, differences between the main-chain and side-chain dihedral angles, percentage sequence identities, and several other measures. Finally, the ID_TABLE command writes a file with pairwise sequence distances that can be used directly as the input to the DENDROGRAM command—or the clustering programs in the PHYLIP package (Felsenstein, 1985). DENDROGRAM calculates a clustering tree from the input matrix of pairwise distances, which helps visualizing
READ_ALIGNMENT FILE = ’$(LIB)/CHAINS_all.seq’,; ALIGN_CODES = ’1bmdA’ ’4mdhA’ ’5mdhA’ ’7aidhA’ MALIGN MALIGN3D COMPARE ID_TABLE DENDROGRAM
Figure 2.9.7 TOP script file compare.top.
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>> Least-squares superposition (FIT)
:
T
Atom types for superposition/RMS (FIT_ATOMS) : CA Atom type for position average/variability ( DISTANCE_ATOMS[1]): CA Position comparison (FIT_ATOMS): Cutoff for RMS calculation:
3.5000
Upper = RMS, Lower = numb equiv positions 1bmdA 0.000 310 308 320
1bmdA 4mdhA 5mdhA 7mdhA
4mdhA 1.038 0.000 329 306
5mdhA 0.979 0.504 0.000 307
7mdhA 0.992 1.210 1.173 0.000
>> Sequence comparison: Diag=numb res, Upper=numb equiv res, Lower = % seq ID 1bmdA 4mdhA 5mdhA 7mdhA
1bmdA 327 51 51 48
4mdhA 168 333 98 41
5mdhA 168 328 333 41
7mdhA 158 137 138 351 1bmdA @1.9
49.0000
4mdhA @2.5
2.0000
5mdhA @2.4
55.5000
7mdhA @2.4
+ + + + + + + + + + 57.6400 48.0100 38.3800 28.7500 19.1200 52.8250 43.1950 33.5650 23.9350 14.3050 4.6750
+ + + 9.4900 -0.1400
Figure 2.9.8 Excerpts from the log file compare.log.
differences among the template candidates. Excerpts from the log file are shown in Figure 2.9.8 (file compare.log). The comparison in Figure 2.9.8 shows that 5mdhA and 4mdhA are almost identical, both sequentially and structurally. They were solved at similar resolutions, 2.4 and 2.5 Å, respectively. However, 4mdhA has a better crystallographic R-factor (16.7 versus 20%), eliminating 5mdhA. Inspection of the PDB file for 7mdhA reveals that its crystallographic refinement was based on 1bmdA. In addition, 7mdhA was refined at a lower resolution than 1bmdA (2.4 versus 1.9), eliminating 7mdhA. These observations leave only 1bmdA and 4mdhA as potential templates. Finally, 4mdhA is selected because of the higher overall sequence similarity to the target sequence. Aligning TvLDF with the Template
Comparative Protein Structure Prediction
A good way of aligning the sequence of TvLDH with the structure of 4mdhA is the ALIGN2D command in MODELLER. Although ALIGN2D is based on a dynamic programming algorithm (Needleman and Wunsch, 1970), it is different from standard sequence-sequence alignment methods because it takes into account structural information from the template when constructing an alignment. This task is achieved through a variable gap penalty function that tends to place gaps in solvent-exposed and curved
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READ_MODEL FILE = '4mdh.pdb' SEQUENCE_TO_ALI ALIGN_CODES = '4mdhA', ATOM_FILES = '4mdhA' READ_ALIGNMENT FILE = 'TvLDH.ali', ALIGN_CODES = ALIGN_CODES 'TvLDH', ADD_SEQUENCE = ON ALIGN2D WRITE_ALIGNMENT FILE='TvLDH-4mdhA.ali', ALIGNMENT_FORMAT = 'PIR' WRITE_ALIGNMENT FILE='TvLDH-4mdhA.pap', ALIGNMENT_FORMAT = 'PAP'
Figure 2.9.9 The align2d.top file that contains the MODELLER script executing the ALIGN2D command.
_aln.pos 10 20 30 40 50 60 GSEPIRVLVTGAAGQIAYSLLYSIGNGSVFGKDQPIILVLLDITPMMGVLDGVLMELQDCALPLLKDV 4mdhA MSEAAHVLITGAAGQIGYI LSHWIASGELYG-DRQVYLHLLDIPPA MNRLTALTMELEDCA FPHLAGF TvLDH ** ******* * * * * * * * **** * * * *** *** * * _consrvd **
90 _aln.p 70 80 100 110 120 130 IATDKEEIAFKDLDVAILVGSMPRRDGMERKDLLKANVKIFKCQGAALDKYAKKSVKVIVVGNPANTN 4mdhA VATTDPKAAFKDIDCAFLVASMPLKPGQVRADLISSNSVIFKNTGEYLSKWAKPSVKVLVIGNPDNTN TvLDH **** * * ** *** * * ** * *** * * * ** **** * *** *** _consrvd ** _aln.pos 140 150 170 180 190 200 160 CLTASKSAPSIPKENFSCLTRLDHNRAKAQIALKLGVTSDDVKNVIIWGNHSSTQYPDVNHAKVKLQA 4mdhA CEIAMLHAKNLKPENFSSLSMLDQNRAYYEVASKLGVDVKDVHDIIVWGNHGESMVADLTQATFTKEG TvLDH * **** * ** *** * **** ** * **** * * _consrvd * *
Figure 2.9.10 The alignment file between sequences TvLDH and 4mdhA in the PAP MODELLER format.
regions, outside secondary structure segments, and between two Cα positions that are close in space. As a result, the alignment errors are reduced by approximately one third relative to those that occur with standard sequence alignment techniques. This improvement becomes more important as the similarity between the sequences decreases and the number of gaps increases. In the current example, the template-target similarity is so high that almost any alignment method with reasonable parameters will result in the same alignment. The MODELLER script shown in Figure 2.9.9 aligns the TvLDH sequence in file TvLDH.seq with the 4mdhA structure in the PDB file 4mdh.pdb (file align2d.top). In the first line of the script shown in Figure 2.9.9, MODELLER reads the 4mdhA structure file. The SEQUENCE_TO_ALI command transfers the sequence to the alignment array and assigns it the name 4mdhA (ALIGN_CODES). The third line reads the TvLDH sequence from file TvLDH.ali, assigns it the name TvLDH (ALIGN_CODES), and adds it to the alignment array (ADD_SEQUENCE = ON). The fourth line executes the ALIGN2D command to perform the alignment. Finally, the alignment is written out in two formats, PIR (TvLDH-4mdhA.ali) and PAP (TvLDH-4mdhA.pap). The PIR format is used by MODELLER in the subsequent model-building stage. The PAP alignment format is easier to inspect visually. Due to the high target-template similarity, there are only a few gaps in the alignment. In the PAP format, all identical positions are marked with a “*” (file TvLDH-4mdhA.pap; Fig. 2.9.10).
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INCLUDE SET ALNFILE = ’TvLDH-4mdhA.ali’ SET KNOWNS = ’4mdhA’ SET SEQUENCE = ’TvLDH’ SET STARTING_MODEL = 1 SET ENDING_MODEL = 5 CALL, ROUTINE = ’model’
Figure 2.9.11 TOP script file model-single top.
Model Building Once a target-template alignment is constructed, MODELLER calculates a 3-D model of the target completely automatically. The script in Figure 2.9.11 will generate five models of TvLDH based on the 4mdhA template structure and the alignment in file TvLDH4mdh.ali (file model-single.top). The first line (see Fig. 2.9.11) includes MODELLER variable and routine definitions. The following five lines set parameter values for the “model” routine. ALNFILE names the file that contains the target-template alignment in the PIR format. KNOWNS defines the known template structure(s) in ALNFILE (TvLDH-4mdh.ali). SEQUENCE defines the name of the target sequence in ALNFILE. STARTING_MODEL and ENDING_MODEL define the number of models that are calculated (their indices will run from 1 to 5). The last line in the file calls the “model” routine that actually calculates the models. The most important output files are model-single.log, which reports warnings, errors, and other useful information including the input restraints used for modeling that remain violated in the final model, and TvLDH.B99990001, TvLDH.B9990002, etc., which contain the model coordinates in the PDB format. The models can be viewed by any program that reads the PDB format, such as ModView (http://guitar.rockefeller.edu/modview/; Ilyin et al., 2002). Evaluating a Model If several models are calculated for the same target, the “best” model can be selected by picking the model with the lowest value of the MODELLER objective function, which is reported in the second line of the model PDB file. The value of the objective function in MODELLER is not an absolute measure, in the sense that it can only be used to rank models calculated from the same alignment. Once a final model is selected, there are many ways to assess it. In this example, PROSAII (Sippl, 1993) is used to evaluate the model fold and PROCHECK (Laskowski et al., 1998) is used to check the model’s stereochemistry. Before any external evaluation of the model, one should check the log file from the modeling run for runtime errors (model-single.log) and restraint violations; see the MODELLER manual for details (at the MODELLER Web site listed in Internet Resources for MODELLER Web site). Both PROSAII and PROCHECK confirm that a reasonable model was obtained, with a Z-score comparable to that of the template (-10.53 and -12.69 for the model and the template, respectively). Comparative Protein Structure Prediction
More detailed examples of MODELLER applications can be found in Fiser and Sali (2002).
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CONCLUSION Over the past few years, there has been a gradual increase in both the accuracy of comparative models and the fraction of protein sequences that can be modeled with useful accuracy (Marti-Renom et al., 2000; Baker and Sali, 2001; Pieper et al., 2002). The magnitude of errors in fold assignment, alignment, and the modeling of side-chains, loops, distortions, and rigid-body shifts has decreased measurably. These improvements are a consequence both of better techniques and a larger number of known protein sequences and structures. Nevertheless, all the errors remain significant and demand future methodological improvements. In addition, there is a great need for more accurate detection of errors in a given protein structure model. Error detection is useful both for refinement and interpretation of the models. It is now possible to predict by comparative modeling significant segments of approximately one half of all known protein sequences (Pieper et al., 2002). One half of these models are in the least accurate class, based on less than 30% sequence identity to known protein structures. The remaining 35% and 15% of the models are in the medium- (<50% sequence identity) and high-accuracy classes (>50% identity), respectively. The fraction of protein sequences that can be modeled by comparative modeling is currently increasing by approximately 4% per year (Sanchez et al., 2000). It has been estimated that globular protein domains cluster in only a few thousand fold families, approximately 900 of which have already been structurally defined (Holm and Sander, 1996; Lo Conte et al., 2000). Assuming the current growth rate in the number of known protein structures, the structure of at least one member of most of the globular folds will be determined in less than 10 years (Holm and Sander, 1996). However, there are some classes of proteins, including membrane proteins, that will not be amenable to modeling without improvements in structure determination and modeling techniques. To maximize the number of proteins that can be modeled reliably, a concerted effort towards structure determination of new folds by X-ray crystallography and nuclear magnetic resonance spectroscopy is in order, as envisioned by structural genomics (Terwilliger et al., 1998; Sali, 1998; Montelione and Anderson, 1999; Zarembinski et al., 1998; Burley et al., 1999; Vitkup et al., 2001; Sanchez et al., 2000). It has been estimated that 90% of all globular and membrane proteins can be organized into approximately 16,000 families containing protein domains with more than 30% sequence identity to each other (Vitkup et al., 2001). 3000 of these families are already structurally defined; the others present suitable targets for structural genomics. The full potential of the genome sequencing projects will only be realized once all protein functions are assigned and understood. This aim will be facilitated by integrating genomic sequence information with databases arising from functional and structural genomics. Comparative modeling will play an important bridging role in these efforts. ACKNOWLEDGEMENTS We are grateful to all members of our research group, especially to Dr. András Fiser, for many discussions about comparative structure modeling. Marc A. Marti-Renom was a Burroughs Wellcome Fund Postdoctoral Fellow and is currently a Rockefeller University Presidential Postdoctoral Fellow. Andrej Sali is an Irma T. Hirschl Trust Career Scientist. Support by The Merck Genome Research Institute, Mathers Foundation, and NIH (GM 54762) is also acknowledged. This unit is based on Marti-Renom et al. (2000) and Fiser and Sali (2002).
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lattice modeling for protein structure prediction and refinement. Proteins 44:133-149. Koretke, K.K., Luthey-Schulten, Z., and Wolynes, P.G. 1998. Self-consistently optimized energy functions for protein structure prediction by molecular dynamics. Proc. Natl. Acad. Sci. U.S.A. 95:2932-2937. Krogh, A., Brown, M., Mian, I.S., Sjolander, K., and Haussler, D. 1994. Hidden Markov models in computational biology: Applications to protein modeling. J. Mol. Biol. 235:1501-1531. Laskowski, R.A., Rullmannn, J.A., MacArthur, M.W., Kaptein, R., and Thornton, J.M. 1996. AQUA and PROCHECK-NMR: Programs for checking the quality of protein structures solved by NMR. J. Biomol. NMR. 8:477-486.
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Comparative Protein Structure Prediction
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Computational Analysis
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INTERNET RESOURCES http://guitar.rockefeller.edu/modeller/
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Table 2.9.1 lists a large number of additional Web sites useful in comparative modeling.
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Contributed by Marc A. Marti-Renom, Bozidar Yerkovich, and Andrej Sali The Rockefeller University New York, New York
Comparative Protein Structure Prediction
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Protein Charge Determination
UNIT 2.10
To many researchers the problem of protein charge determination has been solved by the creation of extensive protein data banks and the availability of computer programs for the calculation of net charge from sequence data and a set of pK values for proton dissociation from ionizable groups (Jakubke and Jeschkeit, 1977; Skoog and Wichman, 1986). However, the inaccuracy of that approach was recognized long before the advent of protein data banks and computer programs to facilitate its adoption. Although improvements to these algorithms can be implemented, there is no alternative to experimental measurement for reliable estimation of protein valence. Of the various methods available (Winzor, 2004), capillary zone electrophoresis affords the simplest and most economical approach. This unit also provides a brief discussion of the major pitfalls inherent in the calculation of net charge from protein sequence data.
CALCULATION OF PROTEIN CHARGE FROM SEQUENCE DATA The popularity of this method of charge estimation resides in its availability as an accessible option within the computer software of any protein data bank. Consequently, the reliability of the calculated valence depends upon the adequacy of the quantitative treatment incorporated into the program, which is generally not questioned by its users. On the grounds that the fractional dissociation (αi ) of any ionizable group i is related to pH by the Henderson−Hasselbalch equation:
Equation 2.10.1
αi may be calculated from pKi , the negative logarithm of the effective dissociation constant. In the usual charge determination procedure (Jakubke and Jeschkeit, 1977; Skoog and Wichman, 1986), the pKi for each type of ionizable group is assigned a magnitude. Knowledge of the amino acid sequence (or composition) of a protein is then used to calculate the net charge, zP , as (zP )i , where the protein charge arising from ni ionizable groups of type i, (zP )i , is given by:
Equation 2.10.2
where the ionizable group, i, is anionic (zi is negative) OR
Equation 2.10.3
where the ionizable group, i, is cationic (zi is positive). However, pKi cannot be assigned a fixed magnitude because of a dependence upon the overall charge of the protein; it is more difficult to dissociate a proton from a negatively charged molecule than from one with a net positive charge. This variation of pKi can be taken into account (Compton and O’Grady, 1991) by means of the theoretical expression for proton dissociation that has been in existence for over 80 years (Linderstrøm-Lang, Computational Analysis Contributed by Donald J. Winzor Current Protocols in Protein Science (2005) 2.10.1-2.10.8 C 2005 by John Wiley & Sons, Inc. Copyright
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1924; Linderstrøm-Lang and Nielsen, 1959). Specifically, pKi in Equation 2.10.1 needs to be replaced by the relationship
Equation 2.10.4
where the factor 0.868wzP takes into account the work required to remove the dissociating proton from the charged protein molecule. (pKint )i refers to the intrinsic dissociation constant for ionizable groups i in a molecule with net zero charge. Because of the need to specify the correct magnitude of zP for the calculation of pKi , and hence αi , Equations 2.10.2 to 2.10.4 must be used iteratively to calculate protein valence (Compton and O’Grady, 1991; Winzor, 2003). A problem with this approach is the need to assign a magnitude to the electrostatic interaction factor, w, in Equation 2.10.4. Although Linderstrøm-Lang (1924) provided a theoretical expression for evaluating w from the protein radius and the ionic strength of the medium, the value so obtained invariably exceeds the experimental estimate of its magnitude (Tanford et al., 1955a,b). In addition to errors in zP stemming from that source, there are those that arise from assuming that all charged groups exhibit normal ionization behavior—an assumption that is demonstrably invalid for the tyrosine residues of ribonuclease (Tanford et al., 1955a) and the carboxyl groups of bovine serum albumin (Tanford et al., 1955b). Finally, the neglected consequences of the binding of ions other than protons and hydroxyls preclude any meaningful attempt to calculate protein valence. For example, the calculated isoelectric point (pH associated with net zero charge) of 8.2 for aldolase clearly does not describe the experimental situation (Velick, 1949) in phosphate and acetate buffers with a range of ionic strengths (Fig. 2.10.1)—allowance for the binding of chloride ions also needs to be taken into account. Unfortunately, quantification of chloride binding by proteins (Carr, 1953, 1956; Scatchard et al., 1957) is restricted to measurements at very low ionic strength because of sensitivity limitations in the electrochemical estimation of chloride concentration. Protein valence, therefore, needs to be obtained by experimental measurement under the conditions of interest.
Protein Charge Determination
Figure 2.10.1 Effect of ionic strength (I ) on the isoelectric point of aldolase in acetate and phosphate buffers. The data, taken from Velick (1949), were acquired by interpolating electrophoretic mobility measurements obtained by the moving boundary method.
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ELECTROPHORETIC DETERMINATION OF NET PROTEIN CHARGE The study of proteins by electrophoresis became popular after refinement of the moving boundary technique by Tiselius (1937). Moving boundary electrophoresis afforded an unequivocal means of characterizing proteins on the basis of their migration rate (dx/dt) under the influence of an electrical field (potential gradient), E. The electrophoretic mobility, uP , defined as
Equation 2.10.5
is related to the charge/size ratio of a protein molecule. Most mobility data have been obtained by moving boundary electrophoresis (Tiselius, 1937), but the demise of that technique now necessitates the use of methods such as gel electrophoresis (Chapter 10; Rodbard and Chrambach, 1971) and capillary zone electrophoresis (UNIT 10.13; K´alm´an et al., 1995). These methods have proven more popular than their predecessor because of greater economy in terms of protein requirements. In an electrochromatographic procedure, such as gel electrophoresis, the experimentally determined mobility from a single run necessarily reflects the combined consequences of chromatographic (molecular sieving) and electrophoretic migration. For monophasic buffer systems, this electrochromatographic mobility (uP ) can be obtained via Equation 2.10.5 from the distance migrated by the protein zone during electrophoresis for time t in response to an applied electric field E based on the applied voltage and the length of the gel tube. Because the extent of protein retardation by molecular sieving effects decreases with increasing gel porosity (decreasing gel concentration), it is advantageous to obtain the linear relationship between the logarithm of uP and the gel concentration (Ferguson, 1964; Hedrich and Smith, 1968) to eliminate the chromatographic contribution to uP and hence determine the electrophoretic mobility uP (Rodbard and Chrambach, 1971; Butterman et al., 1988). The need for multiple electrophoresis runs in two series of gels with different porosities (Butterman et al., 1988) is obviously a disadvantage of the electrophoretic mobility determination method. On the other hand, a single experiment suffices for the measurement of mobility by capillary electrophoresis. Because of the experimental design of the apparatus, migration is described by a retention time, tP , the time taken by the protein zone to move a fixed distance (x) from the point of sample application to the detector. For an electric field strength, E, taken as the applied voltage divided by the total length of the capillary across which the voltage is applied, the retention time is related to electrophoretic mobility by the expression:
Equation 2.10.6
Allowance is made for the contribution of electroosmotic flow to protein migration by the inclusion of a retardation time (tM ) for an uncharged marker (mesityl oxide). The reliability of mobility measurement by capillary zone electrophoresis is obviously dependent upon the relative magnitudes of the (x/tP ) and (x/tM ) terms in Equation 2.10.6. In some instances, tP and tM may differ by as little as 2% (Gao et al., 1994), whereupon the magnitude of uP obtained from Equation 2.10.6 is subject to considerable uncertainty. Fortunately, the accuracy of mobility measurement can be improved by coating the inner walls of the capillary tube with a non-crosslinked polyacrylamide layer to decrease the extent of electroosmosis (Hjert´en, 1985; K´alm´an et al., 1995).
Computational Analysis
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Figure 2.10.2 Time course of absorbance (200 nm) at the detection point, located at distance x (20 cm) from the point of sample application, in a zonal capillary electrophoresis experiment on β-lactoglobulin in 0.05 M phosphate buffer, pH 6.7. tP , the time required for the protein zone to migrate the 20 cm under the influence of an electric field strength (E) of 260 V/cm, is the protein characteristic required for determination of electrophoretic mobility (uP ) from Equation 2.10.6. The electrophoretic time-course has been inferred from Figure 3b in Gao et al. (1997).
Capillary zone electrophoresis is currently of limited utility for protein charge estimation because buffers are usually selected on the basis of resolving power rather than physiological considerations. However, the feasibility of conducting capillary zone electrophoresis in buffer with much higher ionic strength is illustrated in Figure 2.10.2, which presents an electropherogram (Gao et al., 1997) for β-lactoglobulin in 0.05 M phosphate buffer, pH 6.7 (I ≈ 0.1 M). Clearly, tP can still be defined sufficiently well for accurate determination of uP via Equation 2.10.6.
RELATIONSHIP BETWEEN MOBILITY AND PROTEIN CHARGE The interpretation of electrophoretic mobility in terms of protein charge has traditionally been considered equivocal because of reliance of the theory upon a microscopic model of electric migration that is known to be imperfect in some respects (Overbeek and Lijklema, 1959). However, the major cause for concern about the validity of protein charge determination from mobility data has been the discrepancies between such estimates and those deduced from pH-titration curves (Longsworth, 1941; Barnett and Bull, 1960). It is now evident that the discrepancy reflected error in the pH titration values rather than in those deduced from electrophoretic mobility (Winzor, 2004). For a uniformly charged spherical molecule, the dependence of electrophoretic mobility upon protein valence is given by the relationship (Abramson et al., 1942):
Equation 2.10.7
Protein Charge Determination
where e denotes electronic charge (1.602 × 10−19 C), κ is the inverse screening length (Debye and H¨uckel, 1923), and f(κRP ) the Henry function (Henry, 1931). The inclusion of Rb , the radius of a buffer ion, makes allowance for the fact that (RP + Rb ) is the distance of closest approach by buffer ion to the center of the protein. η is the dynamic viscosity of the buffer medium (0.00089 N·sec/m2 for water at 25◦ C). Allowance for protein asymmetry entails the incorporation of an additional term ψ (0 < ψ < 1) into the
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Figure 2.10.3 pH dependence of the net charge for ovalbumin (I = 0.1 M) deduced from published electrophoretic mobility measurements (Longsworth, 1941; Creeth and Winzor, 1962). Open symbols denote values calculated by the application of Equation 2.10.7, which assumes spherical geometry for the protein, whereas closed symbols are values of zP based on a modified version of Equation 2.10.7 that takes into account the prolate-ellipsoidal shape of ovalbumin.
numerator of Equation 2.10.7—a modification of the procedure that has been described in detail elsewhere (Winzor, 2004; Winzor et al., 2004). The starting point for the application of Equation √ 2.10.7 is the calculation of the inverse screening length (m−1 ) as κ = 3.27 × 109 I , where I is the molar ionic strength (Abramson et al., 1942). Also required is the assignment of magnitudes to the radius of a buffer ion Rb (0.25 nm) and that of the protein RP , considered to be the Stokes radius. The Henry function may be calculated with sufficient accuracy by the approximate relationship
Equation 2.10.8
with a = 2.8 for κRP < 10, and 2.5 for κRP > 10 (Rodbard and Chrambach, 1971). In the absence of any extensive sets of mobility data obtained by capillary zone electrophoresis, the use of electrophoretic mobility results for the determination of protein valence is illustrated in Figure 2.10.3, which presents the pH dependence of ovalbumin valence that is deduced from electrophoretic mobilities (Longsworth, 1941; Creeth and Winzor, 1962) obtained by the moving boundary technique. Open symbols summarize the interpretation of those data for ovalbumin (I = 0.1 M, 0◦ C) in terms of Equation 2.10.7 with a protein radius (RP ) of 2.9 nm (Creeth and Winzor, 1962; Jeffrey et al., 1977) and a value of 0.00179 N·sec/m2 for the buffer viscosity (η) at 0◦ C (Abramson et al., 1942). However, these estimates of zP refer to a spherical model of ovalbumin, which is more appropriately represented as a prolate ellipsoid with an axial ratio of 1.95 (Harding et al., 1999). Valence estimates for this model (closed symbols, Fig. 2.10.4), which are 32% higher, provide a better indication of the pH dependence of the net charge on ovalbumin (Winzor, 2004). Computational Analysis
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Figure 2.10.4 Estimation of protein valence by capillary zone electrophoresis and the “charge ladder” approach. Published electrophoretic mobilities for Staphylococcal nuclease and mutant variants (Table 4 of Kalm ´ an ´ et al., 1995) are plotted in accordance with Equation 2.10.10, the quantitative basis for valence estimation by the “charge ladder” method. The designated value of zP for each set of conditions is the reciprocal of the slope of the best-fit description of the data in such terms.
VALENCE MEASUREMENT BY THE “CHARGE LADDER” APPROACH A capillary zone electrophoresis procedure has been developed for the determination of protein charge on the basis of mobility differences stemming from alteration of the protein charge either by mutation (K´alm´an et al., 1995) or by chemical modification (Gao et al., 1994). Provided that such modification has no effect on the size and shape of the protein molecule, it follows from Equation 2.10.7 that
Equation 2.10.9
where the P and D subscripts refer to the native protein and its modified derivative, respectively. It has been proposed (K´alm´an et al., 1995) that (zP − zD ) can be deduced from the nature of the mutation in that replacement of an aspartate residue by alanine should change the net charge by +1, whereas substitution of arginine for aspartate should lead to an increase of +2 in net charge. Equation 2.10.9 is therefore rearranged to the form:
Equation 2.10.10 Protein Charge Determination
The valence of the native protein is therefore deduced as the reciprocal of the slope of the dependence of (uP − uD )/uP upon (zP − zD ). From the illustration of this procedure in Figure 2.10.4 for Staphylococcal nuclease (K´alm´an et al., 1995) at pH 6.8 and pH 8.9,
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it is evident that uncertainty surrounds the estimate of zP . Of particular concern is the variation in mobility between mutants bearing the same charge—an observation that invalidates the assumed identity of size and shape for the native protein and all modified forms thereof (Winzor et al., 2004). The lack of any requirement for information about the size and shape of the protein is certainly an attractive feature of this “charge ladder” method for estimating protein valence by capillary zone electrophoresis (Gao et al., 1994; K´alm´an et al., 1995). Unfortunately, valence values determined by this procedure may be prone to considerable error because of failure of the experimental system to comply rigorously with the requirement that chemical modification or mutation has no effect whatsoever on the overall threedimensional characteristics of the protein. A preferred approach would therefore be the interpretation of uP for the native protein in terms of the classical electrophoretic theory described above (Winzor et al., 2004).
CONCLUDING REMARKS This treatment of protein charge determination covers an aspect of protein chemistry with which few biochemists are familiar. Indeed, it refutes the method of charge determination with which protein chemists are most familiar—the calculation of net charge from protein sequence data. Because zP depends not only upon pH but also on the buffer/electrolyte composition of the environment, there is no realistic alternative to the experimental measurement of protein valence. Knowledge of the Stokes radius for a relatively symmetrical (spherical) protein allows the calculation of net charge (via Equation 2.10.7) from the electrophoretic mobility, which is readily measured by capillary zone electrophoresis. In that regard, an improved estimate of zP can be obtained from the same electrophoretic mobility measurements in instances where a prolate ellipsoid provides a more reasonable representation of the three-dimensional protein structure: the only asymmetric model for which the electrophoretic mobility is currently amenable to quantitative interpretation in terms of net protein charge.
Literature Cited Abramson, H.A., Moyer, L.S., and Gorin, M.H. 1942. Electrophoresis of Proteins. Reinhold, New York. Barnett, L.B. and Bull, H.B. 1960. Electrophoresis of ribonuclease and of β-lactoglobulin: Isoelectric points of proteins. Arch. Biochem. Biophys. 89:162-172. Butterman, M., Tietz, D., Orb´an, L., and Chrambach, A. 1988. Ferguson plots based on absolute mobilities in polyacrylamide gel electrophoresis: Dependence of linearity of polymerization conditions and application to the determination of free mobility. Electrophoresis 9:293-298. Carr, C.W. 1953. Studies on the binding of small ions in protein solutions with the use of membrane electrodes. III. The binding of chloride in solutions of various proteins. Arch. Biochem. Biophys. 46:417-423. Carr, C.W. 1956. Studies on the binding of small ions in protein solutions with the use of membrane electrodes. VI. The binding of sodium and potassium ions in solutions of various proteins. Arch. Biochem. Biophys. 62:476-484. Compton, B.J. and O’Grady, E.A. 1991. Role of charge suppression and ionic strength in free zone electrophoresis of proteins. Anal. Chem. 63:2597-2602.
Creeth, J.M. and Winzor, D.J. 1962. Physicochemical studies on ovalbumin 4. Characterization of an iodine-modified derivative by electrophoresis and sedimentation. Biochem. J. 83:566574. Debye, P. and H¨uckel, E. 1923. Theory of electrolytes. I. Lowering of freezing point and related phenomena. Physik. Z. 24:185-206. Ferguson, K.A. 1964. Starch gel electrophoresis: Application to the classification of pituitary proteins and polypeptides. Metabolism 13:9851002. Gao, J., Gomez, F.A., H¨arter, R., and Whitesides, G.M. 1994. Determination of the effective charge of a protein in solution by capillary zone electrophoresis. Proc. Natl. Acad. Sci. U.S.A. 91:12027-12030. Gao, J.Y., Dubin, P.L., and Muhoberac, B.M. 1997. Measurement of the binding of proteins to polyelectrolytes by frontal analysis continuous capillary electrophoresis. Anal. Chem. 69:29452951. Harding, S.E., Horton, J.C., Jones, S., Thornton, J.M., and Winzor, D.J. 1999. COVOL: An interactive program for evaluating second virial coefficients from the triaxial shape or dimensions of rigid macromolecules. Biophys. J. 76:24322438.
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Hedrich, J.L. and Smith, A.J. 1968. Size and charge isomer separation and estimation of molecular weights by disk electrophoresis. Arch. Biochem. Biophys. 126:155-164.
Scatchard, G., Coleman, J.S., and Shan, A.L. 1957. Physical chemistry of protein solutions. VII. The binding of some small ions to bovine serum albumin. J. Am. Chem. Soc. 79:12-20.
Henry, D.C. 1931. The cataphoresis of suspended particles. I. The equation of cataphoresis. Proc. R. Soc. London A. 133:106-129.
Skoog, B. and Wichman, A. 1986. Calculation of the isoelectric points of polypeptides from the amino acid composition. Trends Anal. Chem. 5:82-83.
Hjert´en, S. 1985. High-performance electrophoresis: Elimination of electroendosmosis and solute sorption. J. Chromatogr. 347:191-198. Jakubke, H. and Jeschkeit, H. 1977. Amino Acids, Peptides and Proteins. John Wiley & Sons, New York. Jeffrey, P.D., Nichol, L.W., Turner, D.R., and Winzor, D.J. 1977. The combination of molecular covolume and frictional coefficient to determine the shape and axial ratio of a rigid macromolecule. J. Phys. Chem. 81:566-574. K´alm´an, F., Ma, S., Fox, R.O., and Horv´ath, C. 1995. Capillary electrophoresis of S. nuclease mutants. J. Chromatogr. A 705:135-154. Linderstrøm-Lang, K. 1924. The ionization of proteins. Compt. Rend. Trav. Lab. Carlsberg 15(7):1-29. Linderstrøm-Lang, K. and Nielsen, S.O. 1959. Acid−base equilibria in proteins. In Electrophoresis: Theory, Methods and Applications (M. Bier, ed.) pp. 35-89. Academic Press, New York. Longsworth, L.G. 1941. The influence of pH on the mobility and diffusion of ovalbumin. Ann. N.Y. Acad. Sci. 41:267-285. Overbeek, J.T.G. and Lijklema, J. 1959. Electric potentials in colloidal systems. In Electrophoresis: Theory, Methods and Applications (M. Bier, ed.) pp. 1-33. Academic Press, New York. Rodbard, D. and Chrambach, A. 1971. Estimation of the molecular radius, free mobility and valence using polyacrylamide gel electrophoresis. Anal. Biochem. 40:95-134.
Tanford, C., Hauenstein, J.D., and Rands, D.G. 1955a. Phenolic hydroxyl ionization in proteins. II. Ribonuclease. J. Am. Chem. Soc. 77:64096413. Tanford, C., Swanson, S.A., and Shore, J.S. 1955b. Hydrogen ion equilibria of bovine serum albumin. J. Am. Chem. Soc. 77:6414-6428. Tiselius, A. 1937. A new apparatus for electrophoretic analysis of colloidal mixtures. Trans. Faraday Soc. 33:524-531. Velick, S.F. 1949. The interaction of enzymes with small ions. I. An electrophoretic and equilibrium analysis of aldolase in phosphate and acetate buffers. J. Phys. Colloid Chem. 53:135-149. Winzor, D.J. 2003. Classical approach to interpretation of the charge-dependence of peptide mobilities obtained by capillary zone electrophoresis. J. Chromatogr. A1015:199-204. Winzor, D.J. 2004. Determination of the net charge (valence) of a protein: A fundamental but elusive parameter. Anal. Biochem. 325:1-20. Winzor, D.J., Jones, S., and Harding, S.E. 2004. Determination of protein charge by capillary electrophoresis. Anal. Biochem. 333:225-229.
Contributed by Donald J. Winzor University of Queensland Brisbane, Queensland
Protein Charge Determination
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Basic Protein Sequence Analysis
UNIT 2.11
Protein sequence analysis can be performed from many perspectives and with a vast array of bioinformatics methods. This unit will focus on those analyses that can be performed using only a computer (desktop workstation or laptop), Internet access, and a laboratory notebook to record results. All the resources needed are accessible via publicly available Web servers and databases, and require little or no computational expertise. These tools enable biologists to predict the structure of a protein, the presence of functional motifs or domains, cellular localization (e.g., membrane-bound, secreted, chloroplast, or nuclear), and post-translational modifications. Integration of all these data enables the biologist to make a more informed prediction of the likely molecular functions of proteins of interest. A plethora of tools are available for these tasks, and in choosing preferred methods for each of the separate protocols, the authors have been guided primarily by two criteria: user-friendliness and utility. Some tools are extremely powerful, but inappropriate for the computational novice. Other tools may be easy to use, but relatively ineffectual. We have kept working biologists in mind, and asked ourselves which tools our biologist colleagues would find easiest to use and understand. With these criteria in mind, we selected one or two protocols for each task. Identifying structural domains using different types of servers are presented (Basic Protocol 1 and Alternate Protocols 1 and 2), as are procedures for predicting transmembrane regions and subcellular localization (Basic Protocol 2 and Alternate Protocol 3) and key functional residues and motifs (Basic Protocol 3). Each procedure also includes a brief discussion of some of the Web servers and databases that we feel deserve attention. Support Protocols for each method present a strategy for the integration of the results from the different procedures (Support Protocols 1 to 7).
IDENTIFYING STRUCTURAL AND FUNCTIONAL DOMAINS USING INTEGRATED META-SERVERS
BASIC PROTOCOL 1
Below is a protocol for the use of integrated meta-servers. Meta-servers include prediction results from several independent prediction tools to produce a consensus prediction. This helps a biologist avoid false positive predictions. Our primary protocol for this task involves the use of the SMART (Simple Modular Architecture Research Tool) server provided by the European Molecular Biology Laboratory (EMBL) at Heidelberg (Schultz et al., 1998; Letunic et al., 2004). This server integrates structure and function prediction tools with pre-calculated analyses (including domain structure prediction, intron/exon boundary detection and ortholog identification) for proteins in the Swiss-Prot and TrEMBL databases (Boeckmann et al., 2003) and all proteomes analyzed by the Ensembl group (Hubbard et al., 2002). The extensive results and appealing interface make a biologist’s work much easier, and this is our reason for recommending the use of this server.
Computational Analysis Contributed by Nandini Krishnamurthy and Kimmen V. Sj¨olander Current Protocols in Protein Science (2005) 2.11.1-2.11.24 C 2005 by John Wiley & Sons, Inc. Copyright
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Figure 2.11.1 FASTA format of AVR2 HUMAN protein sequence. The Swiss-Prot accession number is P27037 and the Swiss-Prot ID is AVR2 HUMAN.
Materials Input data The sequence to be analyzed should be in FASTA format, or may alternatively be submitted as raw sequence (the single-letter representation of the protein sequence without the definition line). An example FASTA format is shown in Figure 2.11.1. In cases where the sequence of interest is in the Swiss-Prot or TrEMBL databases, the accession number is normally sufficient to retrieve precalculated results. Otherwise, the protein sequence is required as described in the protocol below. 1. Point the browser at the SMART Web site (http://smart.embl-heidelberg.de/). On the SMART main page that appears, click either SMART MODE: NORMAL or SMART MODE: GENOMIC. In NORMAL SMART, the database contains SwissProt, SP-TrEMBL and stable Ensembl proteomes. In GENOMIC SMART, only the proteomes of completely sequenced genomes are used. We recommend the use of NORMAL SMART. In the page that now appears, paste the sequence of interest in FASTA format in the box provided. If the sequence is from Swiss-Prot, TREMBL, or the EMBL database, one could alternatively submit the Accession number or Entry name in the box marked Sequence ID or ACC (Fig. 2.11.2). 2. Select additional analyses. At this point, one can either click the Sequence SMART button or select additional analyses. In this example (shown in Fig. 2.11.3), we have selected all available analyses. a. Outlier homologs. This option will search sequence databases derived from proteins of solved structures and return the ID and E-value (see Table 2.11.1 for definition) of the top hit and the database searched. The search is done on PDB (Protein Data Bank, a database of solved structures) and position-specific iterated BLAST (PSI-BLAST) profiles derived from SCOP (Structural Classification Of Proteins; Table 2.11.1), a system of classifying proteins based on structural and functional similarity (Murzin et al., 1995).
b. PFAM domains. This option searches for the presence of PFAM domains (Bateman et al., 2002) in the query sequence. Since SMART (as of November, 2004) has 667 hidden Markov models (HMMs), and PFAM (August, 2004) has 7503 HMMs, searching PFAM HMMs increases the likelihood of a match.
c. Signal peptides. This gives information on whether the query sequence contains a signal peptide, using the SigCleave program. Basic Protein Sequence Analysis
d. Internal repeats. This option shows any repeated motifs found in the query.
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Figure 2.11.2 SMART sequence submission form. (A) Submission of sequence in FASTA format. (B) Submission using the Swiss-Prot accession number in the box marked with arrow.
Figure 2.11.3
Sequence submission options in SMART.
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Table 2.11.1 Definitions of Acronyms and Abbreviations Used in this Unit, continued
Acronym/ abbreviation
Full name
Description
3D-PSSM
3-D Position Specific Scoring Matrix
The 3D-PSSM is a protein fold recognition server that uses 1-D and 3-D sequence profiles coupled with secondary structure and solvation potential information
BLAST
Basic Local Alignment Search Tool
BLAST is a sequence comparison algorithm optimized for speed, used to search sequence databases for optimal local alignments to a query. Variants of BLAST enable searches of protein databases using nucleotide queries and vice versa.
CDART
Conserved Domain Architecture Retrieval Tool
For a given protein query sequence, CDART displays the predicted functional/structural domains that make up the protein and lists proteins with similar domain architectures
EMBL
European Molecular Biology Laboratory
EMBL is an international research organization with its main laboratory in Heidelberg, Germany, and four outstations in Hinxton, U.K. (the European Bioinformatics Institute, EBI), Grenoble, France, Hamburg, Germany, and Monterotondo, Italy.
E-value
Expectation value
For a given score S and a database size, the E-value is the number of hits with scores equivalent to or better than S that are expected to occur in a database of that size by chance. The lower the E value, the more significant the score.
HMM
Hidden Markov Model
HMMs for proteins are statistical models (akin to profiles and PSSMs) that represent the amino acid preferences and insertion/deletion likelihoods at each position. They are used to detect remote homologs and to generate alignments of new sequences to a family.
MSA
Multiple Sequence Alignment
A multiple sequence alignment refers to an alignment of more than two sequences; pairwise alignment refers to an alignment of two sequences. Gap characters (typically, dots or dashes) are inserted between amino acids so that all sequences have the same length, and can be arranged in a matrix to display the amino acids conserved across the family. MSAs are used as input to construction of PSSMs and HMMs, to estimate phylogenetic trees, and for numerous other tasks.
NCBI
National Center for Biotechnology Information
The NCBI is the main resource for computational biologists in the U.S., and includes a number of important tools (e.g., BLAST) and databases. Databases include the CDD (Conserved Domain Database), GenBank (a comprehensive sequence database), PubMed (for retrieving papers in biomedical literature), OMIM (Online Mendelian Inheritance in Man), and a host of other important resources. continued
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Table 2.11.1 Definitions of Acronyms and Abbreviations Used in this Unit, continued
Acronym/ abbreviation
Full name
Description
PFAM
Protein Families
PFAM is a large collection of multiple sequence alignments and hidden Markov models representing many protein domains and families. It has a web interface for identification of domains in a given protein, and includes hyperlinks to relevant literature and functional/structural information for many domains.
PSI-BLAST
Position-Specific Iterated BLAST
An iterative search using the BLAST algorithm and profile construction, enabling identification of remote homologs
PSIPRED
PSIPRED is a simple and reliable secondary-structure prediction method, incorporating two feed-forward neural networks, which perform an analysis on output obtained from PSI-BLAST to predict whether a position is likely to be found in an α helix, β strand, or coil.
PSSM
Position-specific scoring matrix
A PSSM is a statistical representation of the amino acids in a multiple sequence alignment, and is used to provide a log-odds score for each amino acid at each position in the alignment. PSSMs, HMMs, and profiles are related.
RasMol
Raster Display of Molecules.
Molecular visualization freeware to visualize 3-D structure of proteins
RPSBLAST
Reverse PSI-BLAST
RPS-BLAST (Reverse PSI-BLAST) searches a query sequence against a database of profiles (i.e., the reverse of PSI-BLAST that searches a profile against a database of sequences).
SAWTED
Structure Assignment With Text Description
SAWTED compares annotations of the query and closely related sequences with annotations associated with database hits having weak scores. This is included to improve the ability to differentiate between true homologs and spurious matches (i.e., SAWTED enables sequences with weak scores but similar annotations to be given more credence in homology search).
SCOP
Structure Classification of Proteins
The SCOP database classifies protein structural domains into a hierarchy (class, fold, superfamily and family), based on functional and structural data. Proteins in the same SCOP superfamily are expected to be related by divergent evolution from a common ancestral protein and to share a similar (albeit potentially high-level) function. Proteins in different SCOP folds are assumed not to be homologous.
SMART
Simple Modular Architecture Research Tool
The SMART Web server allows rapid identification of the domain architecture of a given protein, and includes precomputed results for many sequences
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Figure 2.11.4 SMART results for AVR2 HUMAN. Integrated results from several servers show an N-terminal signal peptide, two PFAM domains, a transmembrane domain, and a low-complexity region. This black and white facsimile of the figure is intended only as a placeholder; for full-color version of figure go to http://www.interscience.wiley.com/c p/colorfigures.htm.
3. Select the Sequence SMART button. This will retrieve precalculated results (when available) or search the sequence against the library of HMMs available. 4. Examine predicted domains and motifs. The features represented in the example (Fig. 2.11.4) are: a. Signal peptide: the small red box at the N-terminus. b. Two PFAM domains: Activin receptor and Pkinase. c. Low-complexity region: pink box at C-terminus. d. Transmembrane domain: Blue vertical box. 5. Place the mouse cursor over predicted domains and unidentified regions to see their start and end amino acid positions. 6. Click on the domains to get a detailed description of the domain. 7. Scroll down to see the E-value of the predicted matches. 8. Analyze results (see Support Protocol 1). SUPPORT PROTOCOL 1
GUIDELINES FOR UNDERSTANDING RESULTS OF ANALYSES FROM INTEGRATED META-SERVERS The domains in SMART (see Basic Protocol 1) are represented by seed alignments (alignments used to build the HMMs). Homologs are gathered by different iterative methods and included in a multiple sequence alignment (MSA) after estimating the statistical significance of sequence similarities and examining experimental biological information, if available. To reduce redundancy in a family of sequences representing a domain, a phylogenetic tree is constructed using the MSA of all the sequences in the family, and, from every branch that is less than a distance of 0.2 on the tree, only a single sequence is chosen. This corresponds to about 80% sequence identity. These sequences are then used in the seed alignment from which the profiles and HMMs representing a domain are constructed (Schulz et al., 1998).
Basic Protein Sequence Analysis
Interpreting the Significance of a Match to a Profile Most similarity search methods employ E-values (Expect values) to report the statistical significance of a match. How an E-value is calculated varies with each method. Generally, the E-value represents the number of matches expected to be found merely by chance. For instance if an E-value is 10, then 10 matches are expected to be found by chance alone. In general, an E-value <0.001 is a good indicator of the credibility of predicted homology, and the likelihood of a spurious result increases as the E-value approaches (or exceeds) 1. A significant E-value implies similarity in structure; the functional specificity may have changed. To confirm functional similarity, it is critical to check for agreement
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at key functional residues (particularly active site residues in the case of an enzyme), and for a high percent identity (as a conservative rule of thumb, we recommend requiring a minimum of 40% identity). If the profile or HMM represents a structural domain, the query should align over a significant fraction of the profile/HMM. As a basic rule of thumb, we recommend requiring a minimum of 75% coverage. Note that the SMART output does not provide this information (i.e., percent identity, coverage of the profile and agreement with key positions); these criteria will need to be confirmed separately.
Factors Affecting E-Value E-values are affected by the length of the profile or HMM, with longer profile/HMMs producing more significant E-values for related sequences and shorter profile/HMMs producing weaker E-values. Weak E-values are particularly common when the profile or HMM is based on very short motifs (e.g., the PFAM Leucine Rich Repeat HMMs, having between 20 and 25 residues). Because a near-exact match to a short motif can occur by chance alone, E-values will be correspondingly high (i.e., approaching or exceeding 1) even for closely related sequences. For this reason, many HMM and profile libraries employ empirically determined score cutoffs instead of E-values (see, e.g., the PFAM gathering threshold); these tend to be more relaxed for shorter profile/HMMs than for longer. Another factor that affects E-values is the size of the database searched. The probability of finding a match by chance increases with the size of the database searched. The upshot of this is that a match between a given query and a hit will have different E-values depending on the database searched. For example, if the E-value of a match between query X and hit Y is 0.01 on searching the National Center for Biotechnology Information (NCBI) nonredundant database (2,182,528 sequences as of November, 2004), then it will be smaller (e.g., 0.001), and hence more significant, on searching Swiss-Prot (159,078 sequences as of November, 2004) using the same search method. The database size becomes an issue especially when E-values are not extremely significant.
IDENTIFYING STRUCTURAL AND FUNCTIONAL DOMAINS USING THE NCBI CD-SEARCH
ALTERNATE PROTOCOL 1
The Conserved Domain Database (CDD; Marchler-Bauer et al., 2003) at NCBI is a database of Position Specific Scoring Matrices (PSSMs) generated from alignments imported from PFAM and SMART and automated alignments from COGs (Cluster of Orthologous Groups; Tatusov, 2001) CD-Search is a tool that uses RPS-BLAST (Reverse PSI-BLAST) to search a sequence against a database of profiles, namely the CDD (Marchler-Bauer et al., 2004). This is the reverse of PSI-BLAST, which searches a profile against a database of sequences.
Materials See Basic Protocol 1. 1. Point the browser at the CDD (http://www.ncbi.nlm.nih.gov/Structure/cdd/cdd.shtml). 2. Input a sequence in FASTA format (or, alternatively, the NCBI accession or GenBank accession number) in the box provided (Fig. 2.11.5). 3. Set the Search Database from the pull-down menu of that name. The default option is CDD, but users can also search SMART, PFAM, and COGs. 4. Set other parameters using the “click here for advanced options” link, if desired. 5. Click Submit.
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Figure 2.11.5
CD-Search sequence submission form.
Figure 2.11.6 Results from CD-Search. (A) The output gives a graphic display and E-values of hits. (B) Pairwise alignment of the query with the top hit. This black and white facsimile of the figure is intended only as a placeholder; for full-color version of figure go to http://www.interscience.wiley.com/c p/colorfigures.htm. Basic Protein Sequence Analysis
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6. The CDD results page returns a graphic display of domains in the query, the Evalues for each hit, and the alignments (Fig. 2.11.6). Analyze results (see Support Protocol 2).
GUIDELINES FOR UNDERSTANDING RESULTS OF ANALYSES FROM THE NCBI CD-SEARCH
SUPPORT PROTOCOL 2
See Alternate Protocol 1 for steps in carrying out this search. 1. The first few lines give details about the version of RPS-BLAST used, the definition line of the query sequence, and the database searched. Because the CDD has regular updates, we recommend noting these data for record-keeping purposes. 2. In the graphic display, the query sequence is represented as a black bar with a ruler above indicating its length. Low-complexity regions are marked in cyan. 3. The domain matches are shown below in different colors depending on the E-value of the hits. The best hits are marked in red, the second best in blue and so on. The CDD contains multiple PSSMs for the same structural domains; these may have differences in length and coverage of the query, and have correspondingly different E-values. Hits to conserved domains that are related by the Conserved Domain Architecture Retrieval Tool (CDART; Geer et al., 2002; also see Table 2.11.1) resource are indicated by the same color. In the example in Figure 2.11.6, the hits to different PSSMs representing kinase domains are all colored red. 4. A jagged edge seen in the hit display indicates incomplete coverage of the domain (a default minimum of 80% coverage is used by the CD search). For example the SPS1 domain in Figure 2.11.6 contains a jagged right edge, indicating that the alignment to the query does not contain this region of the domain. 5. Below the graphical display is a list of hits along with their E-values. A pink dot before the hit indicates the presence of a structure for the predicted hit. Clicking on the dot will display the structure. Prior installation of Cn3D software is required (http://www.ncbi.nlm.nih.gov/Structure/CN3D/cn3d.shtml). 6. Scrolling down the page displays the pairwise alignment between the query and a representative sequence for the PSSM. Identical residues between the query and the hit sequence are colored red, similar residues are colored blue, and masked-out regions are in italics.
Viewing a multiple sequence alignment (MSA) of the query to a conserved domain 7. Clicking on the graphical display of a domain brings up a multiple sequence alignment of the query to selected PSSM training sequences (Fig. 2.11.7). By default, sequences with high similarity to the query protein are shown. This page gives information about the source of the domain alignment, PubMed references, description, and other relevant data. The default MSA viewing option is Hypertext; our guidelines for interpreting results assume this option is used. 8. Interpreting the MSA: aligned positions are in uppercase, unaligned positions are in lowercase, and gaps are indicated by dashes (-). Residues labeled “unaligned” are inserts between positions in the predicted consensus structure. Profiles and hidden Markov models (HMMs) are constructed to represent the conserved structural elements of a protein family or domain. Amino acids corresponding to additional structure (e.g., extended loop regions) will be displayed in lowercase. These lowercase letters are indicative of insertions relative to the consensus structure. Missing parts of the consensus structure are indicated with dash characters (--).
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Figure 2.11.7 Multiple sequence alignment of AVR2 HUMAN kinase domain with a conserved kinase domain hit. The columns marked with # have been identified as critical for kinase function.
9. For domains with a solved structure, one can view the structure using the View 3D Structure option. The coloring scheme of the alignment can be changed to indicate different levels of conservation. For instance using the option Identity from the “color bits” pull-down menu shows identical residues in the alignment in red, aligned residues in blue, and unaligned residues and columns containing many gaps in gray. 10. Features defining a family are highlighted with # above the corresponding columns in the MSA. The evidence for the feature is presented at the bottom of the MSA. This is useful to predict if the query sequence also contains residues important for the family. In our example, clicking on the S-TKc shows a catalytic loop as feature 1 and the 3-D structure as evidence.
Basic Protein Sequence Analysis
Interpreting the significance of results 11. If E-values are weak, it may be helpful to examine the PSSM hits to see if all refer to the same structural or functional class, or if competing hypotheses for a region are presented by the CDD analysis. Checking for significant percent identity and coverage (as described in the SMART discussion earlier in this unit) is important to avoid spurious and misleading predictions.
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12. The Advanced Search mode allows users to change the maximum allowed E-value cutoff. This is generally done if no matches are possible with more restrictive cutoffs. If a permissive threshold (e.g., E-value 1) is used, false positive matches should be expected. 13. Though the domains represented in CD are similar to those in PFAM and SMART, the search and scoring mechanisms are very different. While PFAM uses profile HMM scoring, CD-Search employs RPS-BLAST. Hence, the E-values that are reported from PFAM and CD-Search for the same PFAM domain may not be identical. To interpret the significance of E-values and the effect of profile/HMM lengths on E-values, see the guidelines described in Support Protocol 1.
PREDICTING STRUCTURAL DOMAINS AND SECONDARY STRUCTURE USING 3D-PSSM
ALTERNATE PROTOCOL 2
3D-PSSM is a Web server designed explicitly and exclusively for protein structure prediction, which incorporates a variety of advanced remote homolog detection and threading methods (Kelley et al., 2000). We present 3D-PSSM as a supportive procedure in cases where other servers are unable to provide clues to molecular function or structure for regions of the protein. In these cases, the advanced methods incorporated in the 3DPSSM workflow can often give insights not available via other servers for identifying homologous structures. In contrast to PFAM, the NCBI CDD, and many other servers that model both structural domains and domains having no known structure, any match to a 3D-PSSM domain implies a solved structure for that region. One of the great benefits of 3D-PSSM is the automatic construction of a homology model for any regions in the query matching one or more of the 3D-PSSM models. The 3D-PSSM server is continually updated. As of February, 2005, 3D-PSSM contains a library of 9864 PSSMs representing structural domains.
Materials See Basic Protocol 1. 1. Point the browser at http://www.sbg.bio.ic.ac.uk/servers/3dpssm/. 2. Click on Recognise a Fold (Fig. 2.11.8A) for the sequence submission form. 3. Enter a valid e-mail address. 4. Paste the query sequence in FASTA format. The query sequence must be <800 residues. The maximum number of residues allowed per submission by 3D-PSSM is restricted to 800. If the protein of interest has >800 residues, we suggest that it be submitted in two parts with overlap. For example, for a protein of 1000 residues, submit region 1 to 600 first, and 400 to 1000 next. 5. Click Submit. Results of the run will be returned by e-mail. The e-mail also contains a hyperlink to the Web version of the results. Results are stored only for 5 days in the server (we recommend downloading results from the Web site). 6. Click on the hyperlink in the e-mail to open a Web page with the results (Fig. 2.11.8B). Analyze results (see Support Protocol 3).
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Figure 2.11.8 page.
SUPPORT PROTOCOL 3
Basic Protein Sequence Analysis
Submission form and results page from 3D-PSSM server. (A) The submission form; (B) the results
GUIDELINES FOR UNDERSTANDING RESULTS OF ANALYSES FROM THE 3D-PSSM SERVER We recommend 3D-PSSM (see Alternate Protocol 2) as an alternative for domain identification for those cases where regions of a protein have no significant matches to domains in the CDD or SMART databases. In these cases, the advanced methods used by 3DPSSM may be able to identify a weak but potentially significant match to a protein of solved structure and shed light on the functional role of the protein. Caution must be employed when examining the 3D-PSSM results where the E-value is only marginally significant (e.g., >0.001), as the potential for a false positive match increases as the E-value approaches 1.
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Figure 2.11.9 Sequence alignments produced by 3D-PSSM (A) Alignment of query to its homologs. (B) Pairwise alignment and comparison of secondary structure and solvent accessibility predictions for the query with that of the hit (structural domain). This black and white facsimile of the figure is intended only as a placeholder; for full-color version of figure go to http://www.interscience.wiley.com/c p/colorfigures.htm.
The 3D-PSSM Web server produces a great deal of interesting and informative results, some of which are beyond the scope of this unit. In this protocol, we highlight a selection of the most interesting results. We encourage users to explore further. 1. The first page of the results is separated into two frames. The upper frame contains a summary of results. The lower frame shows the secondary structure of the query sequence predicted by PSIPRED (Jones, 1999; also see Table 2.11.1). We focus here on results in the upper panel. 2. Click on View Multiple Sequence Alignment to examine the alignment of the query and predicted sequence homologs (Fig. 2.11.9A). The 3D-PSSM Web server uses this alignment to construct a PSSM and score proteins of solved structure, as part of a bidirectional search for structural homologies. Each sequence in the alignment has a hyperlink to the corresponding GenBank file at NCBI. The scores and E-value of each hit, and percent ID to query are also available.
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3. Return to the original results page and scroll down to view matches to structural domains. The top 20 structural hits are shown in a tabular format. The table contains the following information. a. Alignment. A link to the sequence alignment of the query to the hit (structural domain) and comparison of the secondary structure and solvent accessibility of residues between query and hit (Fig. 2.11.9B).
b. Fold Library. The ID of the structural hit (d[pdbcode][chain][region]), along with the pairwise ID between query and hit, are shown. Clicking on the ID brings up a page of the fold library containing information about the hit structural domain, including an MSA of the hit with its sequence and structural homologs.
c. Models. A link to the predicted 3-D model of the query protein based on the alignment to the hit which can be viewed by the protein visualization software RasMol (see Table 2.11.1) or Chime.
d. E-value. The E-value of the hit is shown, colored according to level of confidence, red indicating very high confidence.
e. Other information. The other information in the table includes SAWTED (Table 2.11.1) E-value for text matching between query and definitions of hits, SCOP classification of hit, and other data.
If the E-value to a structural hit is >.001, the following analyses are recommended to assess the credibility of the predicted homology 4. Check the sequence alignment. The query and structure should align with few insertions or gap characters, and have significant pairwise identity. As a rule of thumb, a pairwise identity of 25% or more over 80 amino acids can be used to infer homology (assuming very few gaps). As the percent identity drops below this level, the potential for a spurious match by chance alone increases. In these cases, it is critical to confirm that the alignment shows agreement at known critical residues. 5. Compare the agreement between predicted and known secondary structure. Compare the predicted secondary structure of the query with that of hit. The secondary structures are indicated as follows: H for Helix; E for Strand; C for Coil. Agreement between helical and sheet structure is generally more important than in the more variable coil regions. 6. Compare the agreement in the conserved core structure. The hydrophobic core region of the structural domain hit is marked in the MSA page as Core (Fig. 2.11.9B), numerically labeled to indicate how buried the residues are (9 for very buried and making many contacts and 0 for exposed and making few contacts) and color coded to indicate important core residues (red for important core residues). Significant gaps or inserts in the conserved core elements of the structure should be considered evidence against the predicted homology. An important issue to keep in mind is that 3D-PSSM may only identify one structural domain in a multidomain protein. To enable identification of domains in the remaining region(s) of the query, separate submissions of unknown regions are necessary. Basic Protein Sequence Analysis
There are a few other Web servers and databases enabling integrated searches, as shown in Table 2.11.2.
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Table 2.11.2 Web Servers Providing Domain Identification Tools
Web server
Description
URL
SMART
Structural, functional, and localization prediction
http://smart.embl-heidelberg.de/
PFAM
Includes both structural and functional domains
http://www.sanger.ac.uk/Software/Pfam/ http://pfam.wustl.edu
SUPERFAMILY
Based on the Astral PDB90 database of structural domains
http://supfam.mrc-lmb.cam.ac.uk/ SUPERFAMILY/hmm.html
SAM-T02
The UCSC structure prediction HMM library
http://www.cse.ucsc.edu/research/compbio/ HMM-apps/T02-query.html
TigrFAMs
From TIGR, a Web server of HMMs for structure (and function) prediction
http://tigrblast.tigr.org/web-hmm/
PhyloFacts
Includes subfamily HMMs for structural domains.
http://phylogenomics.berkeley.edu/phylofacts/
NCBI Conserved Domain Database
Includes protein domains with solved structures
http://www.ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi
3D-PSSM
Includes a secondary structure prediction to obtain predictions of 3-D fold
http://www.sbg.bio.ic.ac.uk/∼3dpssm/
PREDICTING HELICAL TRANSMEMBRANE REGIONS AND SUBCELLULAR LOCALIZATION
BASIC PROTOCOL 2
The molecular function of a protein is context-dependent; the subcellular localization of a protein plays a large role in determining its partners and pathway interactions. The cellular sorting machinery makes use of sequence signals targeting a protein for secretion, targeting it to different organelles, or targeting it for insertion into the membrane. Bioinformatics methods take advantage of these motifs and patterns as well as other sequence attributes to predict where a protein is localized. These methods are reviewed in this protocol; see UNITS 2.2 & 2.3 for discussion of earlier methods for this type of analysis. Our primary recommended resource for this protocol makes use of a variety of software tools developed by the Center for Biological Sequence Analysis at the Technical University of Denmark (http://www.cbs.dtu.dk/services/). This resource enables the detection of helical transmembrane domains, signal peptides, and other organellar targeting signals. Due to the biological importance of transmembrane proteins, we discuss the detection of these domains first.
Prediction of helical transmembrane regions in protein of interest Transmembrane predictions involve not only the location of probable membrane helices in a protein, but also the full topology of the protein with respect to the membrane (i.e., which regions of the protein span the membrane, which regions are cytoplasmic, and which are extracellular). Among the several Web servers available for transmembrane prediction, we describe the use of the TMHMM server. The TMHMM server uses sophisticated hidden Markov model methodologies to predict transmembrane regions in proteins, incorporating information on hydrophobicity, helix length, charge bias, and topological constraints into its model (Krogh et al., 2001). It has a user-friendly Web interface, produces quick results, and allows batch-mode submissions of up to 4000 sequences. Computational Analysis
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Materials Input data Protein sequence file, in either raw or FASTA format (Fig. 2.11.1). Alternatively, a file containing multiple input sequences in FASTA format can be used. 1. Point the browser at the TMHMM server (http://www.cbs.dtu.dk/services/TMHMM/). 2. Paste in the sequence(s), or upload a file directly from the local computer (recommended when one wants to analyze large numbers of sequences). To upload a file, use the Browse button below the text “Submission of a local file in FASTA format.” Each file can contain at most 4000 sequences. 3. Set display options. If submitting a single sequence, we recommend setting the display as “Extensive, with graphics.” This will provide results in the most informationrich format (see below). The recommended option for multiple sequence search is “One line per protein.” This returns the results in a tab-delimited format that can be saved as a text file to be opened in a program like Excel. 4. Click Submit. After a short time the results will be displayed. 5. Only 10 job (including batch mode) submissions per day are allowed per user. 6. Analyze results (see Support Protocol 4). SUPPORT PROTOCOL 4
GUIDELINES FOR UNDERSTANDING RESULTS OF PREDICTIONS OF HELICAL TRANSMEMBRANE REGIONS AND SUBCELLULAR LOCALIZATION For steps in predicting helical transmembrane regions and subcellular localization, see Basic Protocol 2. 1. Depending on the selection during submission, the results could be either in “long” or “short” format. The long format is shown in Figure 2.11.10. 2. View summary of results containing the following information: a. Length. Length of query protein.
b. Number of predicted TMHs. Number of predicted TM helices in the query.
c. Exp number of AAs in TMHs. Number of amino acids predicted to be in TM helix. If this number is greater than 18, then it is predicted to be a TM.
d. Exp number, first 60 AAs. Number of amino acids expected to be in TM helix in the first 60 residues. A warning POSSIBLE N-term signal sequence is generated if this number exceeds 10.
e. Total prob of N-in. This represents the probability of N-terminal region being on the cytoplasmic side.
Basic Protein Sequence Analysis
3. View plot. The plot summarizes the various signals used by the TM server. The x axis corresponds to the amino acid positions while the y axis corresponds to the likelihood of an individual position being predicted to be in a TM. Note that the y axis continues beyond a value of 1; the region above 1 is used to display the topology. Since the
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Figure 2.11.10 TMHMM ouput for AVR2 HUMAN. Shown are results in the “extensive with graphics” format. A summary of results is followed by a graphic display. The x axis on the graph represents amino acid positions in the query sequence and the y axis represents the probability of a residue to be in a TM. The peaks indicate positions with higher probability of being a TM domain. The predictions of TM and topology are indicated above a y axis value of 1. The region predicted to be inside (cytoplasmic) is represented with a blue line (marked as INSIDE in this figure) and the region on the outside (extracellular) is represented in pink (labeled OUTSIDE in this figure). This black and white facsimile of the figure is intended only as a placeholder; for full-color version of figure go to http://www.interscience.wiley.com/c p/colorfigures.htm.
TMHMM server integrates a variety of signals from the protein, occasionally the predicted number of helices is lower than the number of peaks in the plot. 4. View raw data. Below the plot is a hyperlink to the raw data for the plot. 5. Cautions a. Accuracy of topology predictions. The accuracy of topology prediction (i.e., which parts of the proteins are extracellular and which are cytoplasmic) is known to be one of the most challenging aspects of transmembrane predictions. Even when the location of transmembrane domains is predicted correctly, the topology can be reversed.
b. Potential for false positives and false negatives. All TM prediction servers are known to have problems with false positive and false negative predictions. False positives arise often for hydrophobic stretches and signal peptides. False negatives can arise for any number of reasons, and are particularly common in noncanonical TM stretches containing some polar or charged residues.
c. Incorporating prediction of globular structure. It can be helpful to integrate domain detection, particularly of globular domains, when interpreting TM predictions. If a predicted TM occurs in a region that has a significant match to a protein of solved structure, we recommend examining what is known about that structure. A very limited number of solved structures include
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transmembrane domains. The vast majority of solved structures represent globular (soluble) domains that do not include transmembrane domains. If a predicted TM domain is contained in a region with a significant match to a globular protein, we would discount the likelihood of the TM domain. ALTERNATE PROTOCOL 3
PREDICTING THE SUBCELLULAR LOCALIZATION OF A PROTEIN USING TargetP Cells have evolved a complex machinery for enabling proteins to be targeted to specific sub-cellular locations. Targeted proteins have specific signals recognized by the machinery. For secreted proteins and those targeted to the organelles such as mitochondria or chloroplast, the targeting signals are amino-terminal peptides recognized by the translocation machinery (Schatz and Dobberstein, 1996). The targeting signals for different locations have specific features. For instance, the aminoterminal signal peptide (∼20 amino acids) that targets proteins for secretion starts with a positively charged segment followed by a central hydrophobic region and a polar segment containing the cleavage site recognized by the signal peptidase. Different approaches have been developed to predict protein localization (Emanuelsson and von Heijne, 2001). Most methods specialize in predicting one specific location (e.g., SignalP for predicting signal peptides). In this protocol we review TargetP, an integrated approach that distinguishes proteins targeted to different locations in the cell such as mitochondria, chloroplast, and the secretory pathway.
Materials See Basic Protocol 2. 1. Point the browser at the TargetP server (http://www.cbs.dtu.dk/services/TargetP/). 2. Paste in the sequence(s), or upload a file directly from the local computer. 3. Identify the origin of the protein sequence (i.e., plant or animal). If it is of plant origin, TargetP will search for chloroplast localization signals. 4. Select the option to perform a cleavage site prediction. 5. Use default settings for specificity cutoffs. Click the “Submit sequence/file” button. 6. Analyze results (see Support Protocol 5). SUPPORT PROTOCOL 5
GUIDELINES FOR UNDERSTANDING RESULTS PREDICTING THE SUBCELLULAR LOCALIZATION OF A PROTEIN USING TargetP Results (see Alternate Protocol 3) include scores for different predicted cellular locations (Fig. 2.11.11).
SP : Signal peptide mTP : Mitochondrial target peptide cTP : Chloroplast (if plant sequence is used) Other: For other locations Along with the scores, a “reliability class” (RC) is also calculated. This gives information on the difference between the highest score and the second best.
Basic Protein Sequence Analysis
RC RC RC RC RC
1: 2: 3: 4: 5:
diff > 0.800 0.800 > diff > 0.600 0.600 > diff > 0.400 0.400 > diff > 0.200 0.200 > diff
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Figure 2.11.11
TargetP prediction of the subcellular localization of AVR2 HUMAN.
Table 2.11.3 Cellular Localization Prediction
Web server
Description
URL
TMHMM
The TMHMM transmembrane helix prediction server
http://www.cbs.dtu.dk/services/TMHMM/
DAS
The DAS transmembrane helix prediction server
http://www.sbc.su.se/∼miklos/DAS/
SignalP
The SignalP signal peptide prediction server
http://www.cbs.dtu.dk/services/SignalP/
ChloroP
The ChloroP chloroplast targeting prediction server
http://www.cbs.dtu.dk/services/ChloroP/
PredictNLS
Nuclear localization signal prediction
http://cubic.bioc.columbia.edu/predictNLS/
In interpreting TargetP results, it is important to consider both the predicted location as well as the RC score. An RC of 1 or 2 is a good indicator of reliability. In our example, the score for SP is 0.853 and the RC value is 1, both supporting the prediction of presence of a signal peptide in the protein (Fig. 2.11.11). TargetP is known to have greater specificity in prediction compared to other methods in this class, but this comes at a cost of sensitivity. A null prediction does not imply that the protein is cytoplasmic. By contrast, cleavage-site prediction is less precise. Finally it is critical to indicate whether the input sequence is of plant or animal origin so that the appropriate localization tests can be performed; it is otherwise difficult to distinguish between chloroplast and mitochondrial signals. Other resources for prediction of transmembrane helices or subcellular localization can be found in Table 2.11.3.
PREDICTING KEY FUNCTIONAL RESIDUES AND MOTIFS USING THE PROSITE WEB SERVER
BASIC PROTOCOL 3
Not all residues in a protein are created equal. Residues in active sites, e.g., the catalytic triad of serine proteases, are critical for enzymatic activity. Other positions may be involved in substrate recognition and binding. Still other positions may be conserved for structural reasons. Over the years, biologists have identified a large number of key residues and conserved motifs that are useful in defining and recognizing protein families. Our primary recommended resource for this protocol is PROSITE. The PROSITE Web server enables a biologist to determine whether any experimentally characterized motifs are present in a sequence of interest (Sigrist et al., 2002; Hulo et al., 2004). This resource enables the detection of different protein motifs, post-translational modification sites, and domains. Since domain detection is provided through other servers (presented earlier in this unit), this protocol focuses on the use of the PROSITE Web server for detecting the presence of conserved motifs and functional residues.
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Materials Input data Sequence in raw or FASTA format (Fig. 2.11.1) 1. Point the browser to the PROSITE Web site (http://us.expasy.org/prosite/). 2. Input the sequence in FASTA format in the sequence box provided. 3. Choose the option to exclude patterns with a high probability of occurrence. This will decrease the likelihood of spurious predictions. 4. Click the Quick Scan button to search for conserved motifs in the PROSITE database. 5. Analyze results (see Support Protocol 6).
SUPPORT PROTOCOL 6
GUIDELINES FOR UNDERSTANDING RESULTS OF SEARCHES DONE USING THE PROSITE WEB SERVER The PROSITE Web server (see Basic Protocol 3) includes both pattern and profile search. Profiles are usually constructed to represent domains and protein families, while patterns are confined to the represention of short conserved motifs and critical residues. When a query protein is submitted, both patterns and profiles are searched and the results are returned as “hits by profile” (Fig. 2.11.12A) and “hits by patterns” (Fig. 2.11.12B). For each detected feature, the position and the sequence motif are provided along with a link to detailed information about each feature. Placing the mouse cursor over predicted domains in the graphical display highlights the corresponding region in the sequence. Pattern detection has certain advantages relative to profile search. First, pattern detection is extremely fast. Second, and potentially of greater interest to biologists, the results are easily interpreted; the motifs are typically quite short, and critical residues defining the motif can be examined easily. There are two main disadvantages to pattern detection, relative to profile search. First, a strict agreement with a pattern is required, resulting in lower sensitivity than the more permissive profile searches, which allow substitutions. Second, the results from pattern searches are not given scores; the pattern is either present or it is not. This makes is difficult to evaluate the significance of the match. In addition, some patterns have a high associated false-positive rate (this is more common in very short or highly variable patterns). For example, the post-translational modification site for N-glycosylation is Asn-Xaa-Ser/Thr (where Xaa represents any amino acid). However, the presence of this motif may not be sufficient to ensure glycosylation, as the process is also dependent on the structure of the protein (Sigrist et al., 2002). The PROSITE Web server contains information to help reduce the likelihood of errors. First, each pattern includes information regarding the number of hits obtained while scanning the Swiss-Prot database with that pattern, the number of false positives and false negatives, and results from pattern search using database randomization. These data should be consulted to obtain an understanding of the likely significance of a match to this pattern. In addition, the PROSITE profile searches can provide additional support for a functional classification based on motif and pattern detection. For instance, the presence of a serine/threonine protein kinase signature pattern along with detection of kinase domain for AVR2 HUMAN strengthens the significance of the prediction.
Basic Protein Sequence Analysis
An example of a six-amino-acid PROSITE pattern is shown below. G-{EDRKHPFYW}-x(2)-[STAGCN]-{P}[G is the N-myristoylation site]
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Figure 2.11.12 Prediction of patterns by PROSITE for AVR2 HUMAN. (A) results of profile search; (B) results of pattern search.
Residues in straight brackets ([]) designate residues that are favorable at a position. Residues in curly brackets ({}) designate those amino acids that are not allowed at a position (i.e., in this example, the amino acids EDRKHPFYW are not allowed at position 2). The pattern x(2) indicates any two amino acids. Other resources for similar tasks can be found in Table 2.11.4. Computational Analysis
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Table 2.11.4 Predicting Key Functional Residues and Motifs
Web server
Description
URL
Evolutionary Trace University of Cambridge implementation of Evolutionary Trace algorithm
http://www-cryst.bioc.cam.ac.uk/∼jiye/ evoltrace/evoltrace.html
MEME
The MEME motif discovery software
http://meme.sdsc.edu/meme/Website/intro.html
PROSITE
Detects previously characterized motifs characterizing protein families, as well as post-translational modifications
http://www.expasy.ch/prosite
SUPPORT PROTOCOL 7
HOMOLOG IDENTIFICATION As a support protocol, we encourage biologists to gather homologs for their sequence of interest and to construct and analyze a multiple sequence alignment of these homologs. The BLAST family of methods (Altschul et al., 1990) is by far the most popular means for identifying homologs in database search; we refer readers to UNIT 2.5 in this manual or the BLAST tutorial at the NCBI Website (http://www.ncbi.nlm.nih.gov/ Education/BLASTinfo/tut1.html). Homolog identification is useful for a variety of reasons. First, homologs can give clues to the overall molecular function of a protein. When inferring molecular function by homology, it is critical to confirm that the query and hit have the same overall fold (i.e., are globally alignable), that they are orthologous, and that the database annotation of the hit is based on experiment and not homology (reviewed in Eisen, 1998, and in Sj¨olander, 2004). Gathering homologs is also the first step in construction of a multiple sequence alignment and the (somewhat esoteric) art of alignment gazing. Alignment gazing can help a biologist discriminate between true homologs and spurious inclusions, and assists in the identification of key functional residues. Sequences inserting large numbers of amino acids, that do not align over long regions, or that do not agree at conserved positions (especially longer conserved motifs) should be removed from the alignment. The final set of high-confidence homologs can be used to supplement the structure-prediction efforts (i.e., by parallel analyses using all the servers and approaches described in this unit). In addition, the multiple sequence alignment can be used as input into motif-discovery systems such as MEME. This is particularly helpful when there is limited (or no) success in analysis of the query sequence. Consensus approaches such as these have been shown to dramatically improve prediction accuracy.
COMMENTARY Background Information
Basic Protein Sequence Analysis
Each of the methods presented in this unit has the potential to produce errors of different types. When examining results, it is important to keep in mind the types of errors that are possible with each method, and to attempt to separate those predictions that are highly credible from those that are potentially misleading. Homology-based function prediction is particularly prone to systematic error; domain shuffling, gene duplication, and speciation produce families of proteins sharing some regions of similarity but with different functions and
overall folds. Because of these issues, even a significant E-value between a query and a database hit is not sufficient for inference of molecular function. To avoid these errors, phylogenomic inference of a protein’s molecular function in the context of its related family members is critical. A detailed description of this approach is beyond the scope of this unit; overviews of the issues and methodologies for phylogenomic analysis are presented in Eisen (1998) and Sj¨olander (2004). For these reasons, this unit focuses less on obtaining a prediction of precise molecular
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function and more on a structural annotation of a protein. The protocols presented here have been selected to enable a biologist to label regions in the protein as functional or structural domains, to identify key residues and motifs, and to predict a protein’s cellular localization. Most homolog-detection methods use either profile- or HMM-based strategies. Both profiles and HMMs are generalizations of multiple sequence alignment of homologs. They differ in how they estimate gap parameters; profiles have a fixed cost for insertions and deletions, while HMMs learn them from the training sequences used in the MSA, thus varying the cost for different positions. Both methods are dependent on precision of homolog gathering, alignment accuracy, and estimation of amino acid substitutions. Domain-based approaches to protein annotation have a critical limitation. The profiles (and HMMs and PSSMs) used to detect the presence of these domains are typically optimized for remote homolog detection. This makes them very effective at providing clues to a protein’s fold or function that might otherwise not be achievable. However, because some protein folds are able to provide a multiplicity of distinct functions, a significant score to a profile for a domain may mean only that the query has a similar fold, but may not actually imply a common molecular function. We strongly advise checking for agreement at key functional residues (where this information is available). The use of consensus approaches to predicting a protein’s structure can help avoid errors, provided that the individual methods are orthogonal. For instance, since both SMART and the CDD include PFAM profile HMMs in their structure-prediction process, a consensus structure prediction between the NCBI CDD or SMART and PFAM is much less informative than a consensus of the NCBI CDD and the 3D-PSSM structure prediction Web server (which uses an entirely different approach). Methods for predicting helical transmembrane domains often rely on weak nonspecific signals such as hydrophobicity, and hence have a fairly high frequency of false-positive as well as false-negative predictions (Chen et al., 2002). To boost accuracy in this task, we strongly encourage biologists to examine alternative hypotheses. Does the predicted transmembrane domain occur in a region predicted (with a significant E-value) to be a globular structure? Is it found at the amino-terminus (where it is more likely to be a signal peptide)? In either case, the probability of the
region being a transmembrane domain should be discounted. In summary, our recommendation for enhancing prediction accuracy involves integrating results and information from a variety of different resources, and biological data should be incorporated wherever possible.
Critical Parameters and Troubleshooting On occasion, a Web server will report an error for certain input sequences. In some cases, hidden characters may make identification of the problem difficult. If this happens, try truncating the definition line to the name of the protein, inserting a carriage return after the definition line, and then resubmitting the sequence.
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Recent improvements to the PROSITE database. Nucleic Acids Res. 32:D134-D137. Jones, D.T. 1999. Protein secondary structure prediction based on position-specific scoring matrices. J. Mol. Biol. 292:195-202. Kelley, L.A., MacCallum, R.M., and Sternberg, M.J. 2000. Enhanced genome annotation using structural profiles in the program 3D-PSSM. J. Mol. Biol. 299:499-520. Krogh, A., Larsson, B., von Heijne, G., and Sonnhammer, E.L. 2001. Predicting transmembrane protein topology with a hidden Markov model: Application to complete genomes. J. Mol. Biol. 305:567-580. Letunic, I., Copley, R.R., Schmidt, S., Ciccarelli, F.D., Doerks, T., Schultz, J., Ponting, C.P., and Bork, P. 2004. SMART 4.0: Towards genomic data integration. Nucleic Acids Res. 32:D142D144. Marchler-Bauer, A., Anderson, J.B., DeWeeseScott, C., Fedorova, N.D., Geer, L.Y., He, S., Hurwitz, D.I., Jackson, J.D., Jacobs, A.R., Lanczycki, C.J., Liebert, C.A., Liu, C., Madej, T., Marchler, G.H., Mazumder, R., Nikolskaya, A.N., Panchenko, A.R., Rao, B.S., Shoemaker, B.A., Simonyan, V., Song, J.S., Thiessen, P.A., Vasudevan, S., Wang, Y., Yamashita, R.A., Yin, J.J., and Bryant, S.H. 2003. CDD: A curated Entrez database of conserved domain alignments. Nucleic Acids Res. 31:383-387. Marchler-Bauer, A. and Bryant, S.H. 2004. CDSearch: Protein domain annotations on the fly. Nucleic Acids Res. 32:W327-W331.
Murzin, A.G., Brenner, S.E., Hubbard, T., and Chothia, C. 1995. SCOP: A structural classification of proteins database for the investigation of sequences and structures. J. Mol. Biol. 247:536540. Schatz, G. and Dobberstein, B. 1996. Common principles of protein translocation across membranes. Science 271:1519-1526. Schultz, J., Milpetz, F., Bork, P., and Ponting, C.P. 1998. SMART, a simple modular architecture research tool: Identification of signaling domains. Proc. Natl. Acad. Sci. U.S.A. 95:5857-5864. Sigrist, C.J., Cerutti, L., Hulo, N., Gattiker, A., Falquet, L., Pagni, M., Bairoch, A., and Bucher, P. 2002. PROSITE: A documented database using patterns and profiles as motif descriptors. Brief. Bioinform. 3:265-274. Sj¨olander, K. 2004. Phylogenomic inference of protein molecular function: Advances and challenges. Bioinformatics 20:170-179. Tatusov, R.L., Natale, D.A., Garkavtsev, I.V., Tatusova, T.A., Shankavaram, U.T., Rao, B.S., Kiryutin, B., Galperin, M.Y., Fedorova, N.D., and Koonin, E.V. 2001. The COG database: New developments in phylogenetic classification of proteins from complete genomes. Nucleic Acids Res. 29:22-28.
Contributed by Nandini Krishnamurthy and Kimmen V. Sj¨olander University of California Berkeley, California
Basic Protein Sequence Analysis
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Minimotif Miner: A Computational Tool to Investigate Protein Function, Disease, and Genetic Diversity
UNIT 2.12
Investigating protein function is a very time-consuming, expensive process. Thus, computational tools that aid in prediction of protein function are valuable resources. The Minimotif Miner (MnM) Web site (http://mnm.engr.uconn.edu) has collected information on a large number of short functional motifs, from other databases and primary literature, into a single database. Short motifs are <15 amino acids long, distinguishing them from longer protein domains. The MnM Web tool searches protein queries for the presence of these motifs. Scoring by several approaches allows users to rank and select motifs. By cross-mapping motifs with single nucleotide polymorphisms or disease-associated mutations, users can identify motifs that are altered and generate new hypotheses for mechanisms of genetic diversity and disease. Importantly, MnM is a tool for identifying short functional motifs, not for identifying protein domains, for which numerous resources already exist that are discussed in other units of this chapter (e.g., UNITS 2.3, 2.7, & 2.11). This unit employs Grb2 as an example to describe how MnM can be used to investigate protein function (see Mapping and Ranking Motifs with Minimotif Miner, below). Grb2 is a adaptor protein comprising SH2 and SH3 domains, which links growth-factor receptors to the MAP kinase signal transduction pathways, among other functions (Davis, 1995; Wang and Moran, 1996; Yoon et al., 1997). Analysis of Grb2 with MnM predicts a number of previously known motifs and several motifs that have yet to be studied. Since the definitions of short motifs, by their nature, have lower complexity and higher redundancy than larger protein domains, false-positive predictions are a limitation. Therefore, this unit provides a description of how scores based on increased frequency of occurrence, prediction of motifs on the protein surface, and evolutionary conservation can be used to rank motifs, thereby reducing false-positive predictions. If the protein studied has a known three-dimensional structure, then short motifs predicted by MnM or other short-motif-prediction programs can be mapped onto the surface of the protein (see Mapping Motifs onto Protein Surfaces). A protocol for using PyMOL (a structural display program) to map motifs onto protein surfaces is provided. This approach is much more accurate for identifying motifs on the protein’s surface than the surface prediction score, another approach that MnM uses for ranking motifs (see Interpretation and Selecting Motifs, below). This analysis can also provide insight into the spatial relationship among motifs. The results of MnM can be cross-mapped with known mutations or polymorphisms (see Using Minimotif Miner to Investigate Disease and Genetic Diversity, below). This allows researchers to determine whether mutations identified in patients with a disease perturb an existing motif or introduce a new motif into the protein of interest. This effectively allows investigators to generate new hypotheses for disease pathology. Similarly, if polymorphisms in the coding region of a protein alter the presence or absence of a motif in a protein, this analysis can provide new hypotheses of how such genetic diversity encodes different phenotypes.
Computational Analysis Contributed by Martin R. Schiller Current Protocols in Protein Science (2007) 2.12.1-2.12.14 C 2007 by John Wiley & Sons, Inc. Copyright
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MAPPING AND RANKING MOTIFS WITH MINIMOTIF MINER Input page. The MnM input window accepts protein sequence queries as RefSeq accession numbers or as protein sequences (Fig. 2.12.1; Balla et al., 2006). RefSeq accession numbers for proteins can be readily retrieved using a hyperlink on the input page to the HomoloGene Web site, which can be searched with keywords (Wheeler et al., 2000). Using Grb2 as an example, this keyword retrieves HomoloGene Cluster 1576. This cluster contains RefSeq accession numbers for human Grb2 (NP 002077) and its orthologs from several other species (Pruitt and Maglott, 2001). If using a RefSeq number, the species of the protein sequence should be selected from the pull-down menu on the input page. This is important only for scoring and ranking motifs on the results page. Another pull-down menu allows the user to restrict motifs to the subcellular compartment where they are found. The user then submits the query. Results page. The Results page contains a Protein Sequence window, Motif Table, and Protein Details Table. The protein sequence is initially displayed with all identified motifs highlighted magenta. When human Grb2 is analyzed, 40 different motifs are highlighted. Pop-up balloons for each motif can be visualized by holding the mouse cursor over the motif. The second window on the Results page is a Motif Table that contains information about the motifs highlighted in the sequence window. This table shows the identity of the motif that was found in the protein sequence, the expected and observed number of each motif in the proteome selected from the input page, a ratio of these numbers (Enrichment factor), a short annotation of the motif function, the positions of the first amino acid of the motif in the Grb2 sequence, and a hyperlinked reference to at least one paper that published the consensus motif definition. Selection of a radio button and the View Selected Motif button will highlight only the single selected motif in the Protein Sequence window. The Results page also contains a Protein Details Table that contains database hyperlinks related to the query (Fig. 2.12.2). The content of this table is only displayed if a RefSeq accession number is entered on the input page.
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Figure 2.12.1 MnM Input page, which is used to enter queries. Simplified instructions for entering a query are numbered. Queries can be either a RefSeq accession number (2a) or a protein sequence (2b).
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Figure 2.12.2 Protein Details Table. The accession number for Grb2 was entered on the Input page (see Fig. 2.12.1) and the table shown here was returned, containing relevant information about the query. Database cross-referencing is output only with entry of a RefSeq accession number.
Figure 2.12.3 Protein Sequence window of the MnM results page. Buttons under the window allow mapping of domains (cyan-, green- and yellow-highlighted text) and polymorphisms (capitalized, with blue highlighting). These functions only work with entry of a RefSeq accession number. For the color version of this figure go to http://www.currentprotocols.com.
The Results page has several interactive buttons to aid the user in investigating the identified short motifs. A RefSeq accession number must be used on the input page for these buttons to become active. The View SNPs button will display any known single nucleotide polymorphisms (SNPs) from the dbSNP database with a blue highlight (Fig. 2.12.3; Wheeler et al., 2000). Holding the cursor over the highlighted region reveals a pop-up balloon with more information about the SNP, including its ID number. In the case of human Grb2, SNP-ID # 11552057 is retrieved, which is an allelic variation producing either a Phe or Ile at amino acid 19. The View Domains button will display any protein domains defined by the NCBI Conserved Domain Database (CDD) annotated in the RefSeq database (Marchler-Bauer et al., 2003). These are highlighted yellow, green, or cyan. Again, holding the cursor over the highlighted domain reveals a pop-up balloon
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Table 2.12.1 Scores for ranking short motifs in Grb2a
Motif
Annotation
FS
ECS
SPS
YXXP
SH2 ligand(PLCγ)
2.8
−1.55
0
YXXM
SH2 ligand(PI3K)
8.6
0.56
0
PXXP
SH3 ligand
1.1
−11.1
1
RXX[S,T]V
CamKII substrate
9.8
0.57
1
RXSXSP XP
14-3-3 ligand
172
0.53
1
LXXRR
Erk docking site
13.5
0.56
1
[W,F]W
EH binding motif
7.6
0.56
0
a Abbreviations: FS, frequency score; ECS, evolutionary conservation score; SPS, surface prediction score.
Superscript “P” indicates a phosphorylated residue.
with information about the domain. In the case of Grb2, one SH2 and two SH3 domains are revealed. If there are no SNPs or domains present in the query, then the text in these buttons becomes gray. A View Homologous Proteins button will analyze all orthologs in the same HomoloGene cluster as the query. This analysis adds new information to the Motif Table and sorts the motifs according to degree of evolutionary conservation. The second column of the table now lists the percentage of species that contain the motif and the accession numbers and Taxon IDs of orthologs in the HomoloGene cluster (Wheeler et al., 2000). Taxon IDs are numerical identifiers of species in the Taxonomy database, and these definitions can be retrieved from the MnM help page or from the Taxonomy database (Wheeler et al., 2000). In addition to the frequencies of each motif in the proteome, the expected and observed occurrences and ratio of these numbers for each motif in the query are shown. This is discussed further in the Interpretation and Selecting Motifs section, below. Analysis of Grb2 reveals four Grb2 orthologs from mouse, rat, mosquito, and fly. The 14-3-3 binding motif at the top of the Motif Table is conserved at the same position in Grb2 from all four species, and thus has a conservation of 100%, which is shown in Table 2.12.1.
INTERPRETATION AND SELECTING MOTIFS Three scores are determined for ranking motifs. Above the Motif Table there are buttons for Frequency Score (FS), Surface Prediction Score (SPS), and Evolutionary Conservation Score (ECS). These scores should be used in combination with the known biology of the protein to select motifs for further study.
Frequency Score (FS) FS represents the frequency of occurrence of the motif in the query when compared to all protein sequences in the entire proteome; higher scores are better. An FS >5 is globally more likely to be experimentally confirmed; however motifs with scores <5 are also commonly confirmed and should not be ruled out based on FS alone. Analysis of >2300 experimentally identified motifs that are annotated in the Swiss-Prot database shows that FS is globally larger for motifs that have been experimentally confirmed than for predicted motifs. An example of a prediction with a high FS came from the analysis of EFF-1, a protein that is necessary for fusion of body-wall muscle cells in C. elegans. MnM analysis of EFF-1 revealed a 14-3-3 motif with a high FS score that was subsequently confirmed by biochemical and genetic analyses (Balla et al., 2006). FS has the advantage that it favors motifs rarely found in other proteins, and also motifs Minimotif Miner
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that have multiple copies in a single protein. In many cases, short motifs are found in multiple copies, as exemplified by proline-rich regions with multiple motifs that bind SH3 domains. FS is limited by compartment specificity of motifs, protein surface accessibility of motifs, and random occurrence of motifs, and favors motifs with high complexity and low degeneracy. Therefore, other means of ranking motifs should be used in conjunction with the frequency score. Analysis of human Grb2 yields 40 motifs with FS values ranging from 0.4 to 172. The FS for several motifs in Grb2 are shown in Table 2.12.1. The 14-3-3 motif, which has a score of 172, would be considered more likely to be of biological relevance than the SH3 ligand motif, which has a score of only 1.1 A score of 1.0 would indicate a motif observed at its expected statistical frequency.
Surface Prediction Score (SPS) Given their functions, short motifs should have at least some, if not all, residues on the protein surface. Thus, minimotifs with buried residues are less likely to be functional. The SPS provides a measure of how likely a motif is to be on the protein surface. The scores range between 0 and 1.0, with 1.0 indicating a higher likelihood of the motif being on the surface. Surface prediction was confirmed with two datasets containing nearly 1000 protein structures, and produced an accuracy of 65% to 75% (Balla et al., 2006). For analysis of human Grb2, SPS ranges from 0 to 1.0. A subset of SPS for the 40 identified motifs in Grb2 is shown in Table 2.12.1. It would be expected that the Erk docking site, with a SPS of 1, would be more likely to be on the surface and less likely to be false-positive than the predicted PI3K SH2 ligand, which has an SPS of 0 and is predicted to be buried. Since the three-dimensional structure of Grb2 is known, the motifs shown in Table 2.12.1 were mapped onto backbone and surface plots of the Grb2 crystal structure using MolMol (Fig. 2.12.4; Maignan et al., 1995; Koradi et al., 1996). The two Ser residues in a 14-3-3 binding motif (blue) are surface-exposed, and thus are accessible for Grb2 phosphorylation and binding to 14-3-3. This motif had one of the highest SPS scores. Similarly, residues in the Erk docking site (orange) are exposed on the surface of the Grb2 SH2 domain and had the highest SPS score. The SPS score has limitations. Some motifs may require a specific structure beyond being on the surface, and just because a motif is on the surface does not necessarily mean it is functional. Furthermore, this surface prediction algorithm yields ∼25% to 35% false-positive predictions. Therefore, it is highly recommended that, for proteins with known structures in the PDB, the motifs be mapped onto the surface of the protein. A simplified description on using the PyMOL program to map motifs on protein surfaces is described under Mapping Motifs onto Protein Surfaces, below (also see Delano, 2004). Minimally, SPS should be used in conjunction with other scoring methods.
Evolutionary Conservation Score (ECS) As a quantitative alternative to the View Homologous Proteins function, ECS serves as a quantitative measure of evolutionary conservation of short motifs. The ECS score is generated from a series of BLAST alignments (Balla et al., 2006). This algorithm generates positive or slightly negative scores where a motif is conserved, and strongly penalizes, with relatively large negative scores, if a motif is missing in homologs. The algorithm also takes into account the overall conservation of homologs. ECS, like the FS, has been validated. Examples of ECS scoring for Grb2 are shown in Table 2.12.1. All motifs in Table 2.12.1 have slightly positive or negative ECS values, reflecting conservation of the motif in most
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Figure 2.12.4 Location of conserved minimotifs on the three-dimensional structure of human Grb2. Minimotifs shown in Table 2.12.1 were mapped onto the crystal structure of human Grb2 (1GRI) using MolMol (Koradi et al., 1996). Grb2 contains two SH3 domains (yellow and cyan) and one SH2 domain (light pink). Backbone/stick diagrams (top) and surface plots (bottom) are rotated 180o about the z axis (right). Domains are labeled in the bottom right structure representation. Motifs mapped onto the Grb2 structure are colored: SH2 ligand (PI3K) (red), 14-3-3 ligand (blue), CamKII phosphorylation site (magenta), SH3 ligand (brown), Erk docking site (orange), EF hand (green), and SH2 binding motif(PLCγ) (yellow). For the color version of this figure go to http://www.currentprotocols.com.
of the Grb2 orthologs. An alignment of several Grb2 orthologs is shown in Figure 2.12.5. The PXXP motif has an ECS of −10, reflecting its poor conservation in only 2 of the 6 orthologs. Given the low FS and ECS values for the PXXP motif, it is less likely to be biologically relevant in Grb2. Since evolutionary conservation is considered an important indicator of functional regions in proteins, ECS or View Homologous Proteins can be used to select motifs for further study. However, it must be kept in mind that motifs may be utilized differently among divergent species or that species can have slightly different motif definitions. For example KDEL> is an endoplasmic reticulum retention signal in mammals, while HDEL> is used in yeast (Pelham, 1991; Townsley et al., 1994). In these motif definitions “>” indicates the motif must be on the C-terminus of the protein. The user should be aware that ECS scores are computationally intensive and only currently available through e-mail. In addition to using scoring to select motifs for further study, it is also important to incorporate information about the known biology of the protein. Some of the highscoring motifs in Grb2 participate in functions already known to be associated with the MAP kinase signaling pathway and are consistent with the known biological function of Grb2. For example, the Erk docking site and 14-3-3 binding sites have relatively high FS, ECS, and SPS scores. Erk is one of the kinases in the MAP kinase cascade, and 14-3-3 is already known to bind to Raf, another kinase in the MAP kinase pathway (Davis, 1995; Thorson et al., 1998). It is expected that motifs with functions highly related to the known biology of the protein query are less likely to be false-positive predictions. Minimotif Miner
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Figure 2.12.5 Alignment of Grb2 protein sequences. Alignment of human, mouse, rat, worm, fly, and mosquito Grb2 sequences generated with a multiple sequence alignment tool and Boxshade (http://www.ch.embnet.org/software/BOX doc.html ). SH2 domains are colored green, SH3 domains are colored cyan, and novel minimotifs identified by SMS analysis are colored red. For the color version of this figure go to http://www.currentprotocols.com.
MAPPING MOTIFS ONTO PROTEIN SURFACES If the protein studied has a known three-dimensional structure, then short motifs predicted by MnM or other short-motif prediction programs can be mapped onto the surface of the protein. The SPS score alone has an accuracy of only 65% to 75% for predicting the surface expression of a particular motif. Therefore, if the Protein Data Bank (PDB) contains a structure for the protein being studied, mapping the motifs onto the surface of the protein is a superior approach. Furthermore, by mapping motifs onto the protein surface, additional motif information can be gained to help select other motifs for further consideration. In cases where the structure is known, the spatial relationships of motifs
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can be determined. The structure of a particular motif can be compared to other structures in which the motif ligand is bound to its protein partner in order to further reduce falsepositive predictions. Another consideration pertains to multidomain proteins such as Src, where a motif may be buried between domains and may become accessible only upon other protein-protein interactions (Lerner and Smithgall, 2002; Cowan-Jacob et al., 2005). An example of motif mapping onto the surface of Grb2 is shown in Fig. 2.12.4. Below, a simplified protocol is described for using PyMOL protein structure visualization software to map short motifs onto the surface of proteins with known structure (Delano, 2004). PyMOL is shareware software and instructions for downloading and installing PyMOL are available from the PyMOL Web site (http://pymol.sourceforge.net/). Once the program is installed, follow the instructions below, as done for the Grb2 PDB file 1GRI.pdb similar to Figure 2.12.4 (Maignan et al., 1995). These are simplified instructions; a comprehensive manual is available from the PyMOL Web site (Delano, 2004). MolMol or other structure-plotting programs can also be used to generate surface plots and map motifs (Koradi et al., 1996). 1. Download and save the relevant .pdb file from the Protein Data Bank (1GRI.pdb for Grb2). 2. Download and install PyMOL from http://pymol.sourceforge.net/. 3. Open the PyMOL program. There is a Command Window and Viewer Window.
4. In the Command Window, locate the menu bar and select Open from the File menu, then select the downloaded .pdb file. 5. In the Command Window, choose Display from the menu bar and select Sequence to display the protein sequence in the Viewer Window. 6. In the upper right corner of the Viewer Window, click the “H” button, then select “everything” from the menu that appears. This will hide the molecule.
7. In the upper right corner of the Viewer Window, click the “S” button, then select “ribbon” from the menu that appears. This will show a ribbon plot of the protein molecule.
8. In the Viewer Window, select motif residues in the sequence with the mouse. A new selection (sele) will appear on the right side of window.
9. In the upper right corner of the Viewer Window, to the right of the PDB name, click the “C” button, then choose the color for the backbone. 10. In the upper right corner of the Viewer Window, to the right of “sele,” click the “A” button, choose “rename selection,” type the new motif name, and click Enter, to name the motif. 11. In the upper right corner of the Viewer Window, to the right of the motif name, click the “S” Button, then select “sticks” from the menu that appears to show the motif residues. 12. In the upper right corner of the Viewer Window, to the right of the motif name, click the “C” button, then choose the color for the motif residues. Minimotif Miner
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13. In the upper right corner of the Viewer Window, to the right of the PDB name, click the “S” button, then select “surface” to show the surface. 14. Repeat steps 8 to 12 to color other motifs. 15. Use the mouse to rotate the molecule (by dragging) or zoom the molecule (via the pop-up menu) to view motif(s) on the protein surface. 16. In the upper right corner of the Viewer Window, to the right of the PDB name, click the “H” button, then select “surface” to hide the surface and determine if residues are buried.
USING MINIMOTIF MINER TO INVESTIGATE DISEASE AND GENETIC DIVERSITY If a user is interested in quantitative trait loci, disease-associated mutations, or SNPs that contribute to plant, animal, and human diversity, MnM can be used to investigate the mechanisms by which these substitutions induce a change in phenotype. MnM is useful not only for analyzing missense and nonsense mutations or substitutions, but also for analyzing mutations that affect splicing and use of alternative start/stop codons. Mutations and allelic variations can be identified from numerous resources. Although beyond the scope of this unit, some examples of these databases are provided. The Human Genome Mutation Database (HGMD; http://www.hgmd.cf.ac.uk/ac/index.php) contains over 30,000 mutations that are associated with a variety of diseases (Cooper et al., 1998). The molecular genetics section of the Online Mendelian Inheritance in Man (OMIM; (http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=OMIM) database entries often has a summary of papers that describe allelic variation and diseaseassociated mutations. There are also a fairly large number of smaller databases focused on a specific diseases; these databases contain published and unpublished mutations (Claustres et al., 2002). MnM will map single nucleotide polymorphisms from dbSNP (http://www.ncbi.nlm.nih.gov/projects/SNP/), which contains millions of missense SNPs from several species. There are several additional sources of SNPs. Quantitative trait loci (QTL) are often mapped with SNPs and numerous smaller projects provide a source of additional SNPs (Teufel et al., 2006). A list of many SNP databases can be found at http://hgvbase.cgb.ki.se/databases.htm. Once the protein and mutations/polymorphisms of interest are identified, the amino acid changes resulting from these mutations should be mapped onto the protein sequence. This can be compared to a motif analysis of the protein query by MnM. That comparison will allow the investigator to identify motifs eliminated by mutations in the protein query. To determine whether any motifs are introduced by a mutation, the sequence containing the mutations must be used on the input page. An example analysis of motifs eliminated by mutations in presenilin 1 is shown in Figure 2.12.6. Certain mutations in the presenilin 1 gene are associated with Alzheimer’s disease (Cruts et al., 1996). An alignment of the presenilin 1 proteins from several species is shown in Figure 2.12.6. Mutations found in patients with Alzheimer’s disease are highlighted red. A small subset of the motifs that were identified by analysis with MnM are indicated. Figure 2.12.6 shows that some mutations lie within motifs. For example, mutations have been identified in patients for all three consensus positions of a class II SH3 binding motif. Thus, one arrives at the novel hypothesis that binding of an SH3 domain-containing protein to presenilin 1 is important for its role in disease. Similar analysis can be done for almost all genes implicated to play a role in development or progression of disease. Computational Analysis
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Figure 2.12.6
Legend at right.
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DISCUSSION Predicting protein domains using consensus motifs or positional scoring matrices has proven useful for extrapolating protein function (Letunic et al., 2002; Bateman et al., 2004). This is primarily because there is a very low false-positive domain prediction and a minimal number of false-negative results. Shorter motifs that confer binding, trafficking, or post-translational modification activities to proteins can be equally as important as a functional domain. Because short motifs, by their nature, have low complexity, many false-positive results are expected when trying to correlate motif prediction with function. This is the likely reason why short-motif prediction has not yet developed into as useful an approach as domain prediction. MnM can be used to investigate new functions in proteins as well as functional consequences of mutations or genetic variability. Short motif prediction has several advantages. Because motif prediction is computationally driven, it is fast. For analysis of a typical protein, 50 to 100 motifs are predicted within seconds. Another advantage is that the hypotheses generated with MnM are readily testable. By simply introducing a point mutation that eliminates the short motif and testing for the known function of the motif, it is fairly straightforward to design and perform experiments that test MnM predictions. While there are several Web tools for predicting short motifs, there are two main advantages in using MnM over other motif-searching programs. (1) MnM provides a near-comprehensive view of motifs that are present in other smaller databases. Since most researchers are not aware of several hundred short functional motifs, this Web resource allows investigators to identify new potential functions in proteins. (2) MnM also provides several means for ranking motifs using independent pieces of information. This is important considering that the false-positive rate for short motif prediction is relatively high when compared to protein domain prediction. The most critical consideration in prediction of short motifs is the false-positive rate. There are several other tools that can be used to study motifs. These should be used in conjunction with MnM when ever possible. MnM is the most comprehensive short motif database with >300 nonredundant motifs. Scansite (http://scansite.mit.edu/ motifscan seq.phtml) predicts motifs using position-specific scoring matrices that were built from phage display and random peptide library data (Obenauer et al., 2003). Scansite, while less comprehensive than MnM with 63 motifs, has an advantage over MnM in that experimental data are used to generate the position-specific scoring matrices used for motif prediction. The Eukaryotic Linear Motif (ELM; http://elm.eu.org/) server has ∼80 motifs and several functions for examining motifs that MnM does not have (Puntervoll et al., 2003). “ELM instance” indicates experimentally validated motifs. This server links to pattern-hit initiated BLAST (phi-BLAST; (http://www.ncbi.nlm.nih.gov/blast/), which can be used to search for other proteins that contain the exact sequence of the experimentally identified instance of the motif. The ELM globular domain filter allows users Figure 2.12.6 (at left) Short motifs in presenilin 1 eliminated by mutations in patients with Alzheimer’s disease. Presenilin 1 sequences from human (NP 000012), cow (NP 777146), dog (AAR97724), mouse (NP 032969), rat, (NP 062036), frog (AAH64246), and zebrafish (AAH54639) were aligned and colored using multiple sequence alignment (Smith et al., 1996) and Boxshade (http://www.ch.embnet.org/software/BOX doc.html). Amino acids highlighted black are highly conserved; those highlighted gray are conservative substitutions; those highlighted white are not conserved; and those highlighted red are mutated in human patients with Alzheimer’s disease (http://www.alzforum.org). Amino acids with text colored green are minimotifs that overlap with disease-associated mutations, and those with cyan text are of interest to presenilin function. Motifs are annotated in colored text. For the color version of this figure go to http://www. currentprotocols.com.
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to search only interdomain regions that are thought to be more likely to contain functional motifs (Puntervoll et al., 2003). A few motifs are known to have species-specific consensus definitions, and the ELM server allows species-specific searches. The BLAST Web site has a link with preset options that allows users to search for short, nearly exact matches. In practice, this is useful for motifs of >4 to 5 residues in length (Altschul et al., 1997). The myHITS Web site (http://myhits.isb-sib.ch) is more specialized for this purpose and can be used to search several databases for user-entered motifs that contain degenerate motif definitions using Prosite syntax (Pagni et al., 2001; Falquet et al., 2002). Some motifs such as ubiquitination, O-glycosylation, sulfation, and GPI-anchor attachment are not readily reduced to consensus sequence definitions. Thus, individual servers focused on these motifs can be used to investigate several motifs. A number of these motif predictors are collected at the ExPASy Proteomics Server (Schneider et al., 2004; http://www.expasy.org/). There are also several Web sites that focus on a particular type of motif such as myristoylation, phosphorylation, and proteolysis (Schneider et al., 2004). These sites will generally be superior to MnM, Scansite, and ELM in their ability to predict these specific motifs. The intention of MnM is to inform users of potential functions rather than to flawlessly predict them. Thus, in selecting a motif for experimental analysis, it is recommended that the evolutionary conservation, location of protein surface, increased frequency of occurrence, location in interdomain regions, predictions by other motif programs, and known biology of the protein query all be used to select motifs for study. MnM and other short-motif analysis programs are for prediction, and should be used to generate experimental leads and new hypotheses. All short-motif programs, including MnM, yield a significant number of false-positive predictions, which is the major limitation in motif predictions. MnM, ELM, and Scansite each attempt to limit falsepositive predictions in different ways; thus, using each program will provide additional information in selecting the best motifs for experimental pursuit. It must also be taken into consideration that, for the majority of motifs, every possible amino acid substitution at each position has not been examined; thus, most definitions could possibly be more specific. Furthermore, a number of motifs have been examined using random peptide libraries, or by phage display. While this does assess most, if not all, sequence permutations of the motif, the peptide is out of context of the native protein and may not adopt a structural orientation that is necessary for binding. Thus, motif definitions determined by this approach may also yield false-positive results. Despite these limitations, short-motif analysis will produce many new leads for investigating function. For example, analyzing human dynamin, p53, paxillin, prion, and vAbl with MnM produces 277 motif predictions, of which 18 ± 4% have been previously experimentally confirmed.
ACKNOWLEDGEMENTS This research was funded in part by NIH grant MH65567 to Martin Schiller, grants CCR-9912395 and ITR-0326155 to Sanguthevar Rajasekaran, and ES09847 to Michael Gryk.
LITERATURE CITED Altschul, S.F., Madden, T.L., Schaffer, A.A., Zhang, J., Zhang, Z., Miller, W., and Lipman, D.J. 1997. Gapped BLAST and PSI-BLAST: A new generation of protein database search programs. Nucleic Acids Res. 25:3389-3402. Minimotif Miner
Balla, S., Thapar, V., Luong, T., Faghri, T., Huang, C.H., Rajasekaran, S., del Campo, J.J., Shin,
J.H., Mohler, W.A., Maciejewski, M.W., Gryk, M., Piccirillo, B., Schiller, S.R., and Schiller, M.R. 2006. Minimotif Miner, a tool for investigating protein function. Nat. Methods 3:175177. Bateman, A., Coin, L., Durbin, R., Finn, R.D., Hollich, V., Griffiths-Jones, S., Khanna, A., Marshall, M., Moxon, S., Sonnhammer, E.L.,
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Studholme, D.J., Yeats, C., and Eddy, S.R. 2004. The Pfam protein families database. Nucleic Acids Res. 32:D138-D141. Claustres, M., Horaitis, O., Vanevski, M., and Cotton, R.G.H. 2002. Time for a unified system of mutation description and reporting: A review of locus-specific mutation Databases. Genome Res. 12:680-688. Cooper, D.N., Ball, E.V., and Krawczak, M. 1998. The human gene mutation database. Nucleic Acids Res. 26:285-287. Cowan-Jacob, S.W., Fendrich, G., Manley, P.W., Jahnke, W., Fabbro, D., Liebetanz, J., and Meyer, T. 2005. The crystal structure of a c-Src complex in an active conformation suggests possible steps in c-Src activation. Structure 13:861871. Cruts, M., Hendricks, L., and Broeckhoven, C. 1996. The Presenilin genes: A new gene family involved in Alzheimer’s disease pathology. Hum. Mol. Genet. 5:1449-1455. Davis, R.J. 1995. Transcriptional regulation by MAP kinases. Mol. Reprod. Dev. 42:459467. Delano, W.L. 2004. Use of PYMOL as a communications tool for molecular science. Abstr. Pap. Am. Chem. Soc. 228:U313-U314. Falquet, L., Pagni, M., Bucher, P., Hulo, N., Sigrist, C.J., Hofmann, K., and Bairoch, A. 2002. The PROSITE database: Its status in 2002. Nucleic Acids Res. 30:235-238. Koradi, R., Billeter, M., and Wuthrich, K. 1996. MOLMOL: A program for display and analysis of macromolecular structures. J. Mol. Graph. 14:29-32; 51-55. Lerner, E.C. and Smithgall, T.E. 2002. SH3dependent stimulation of Src-family kinase autophosphorylation without tail release from the SH2 domain in vivo. Nat. Struct. Biol. 9:365369.
cell signaling interactions using short sequence motifs. Nucleic Acids Res. 31:3635-3641. Pagni, M., Iseli, C., Junier, T., Falquet, L., Jongeneel, V., and Bucher, P. 2001. trEST, trGEN and Hits: Access to databases of predicted protein sequences. Nucleic Acids Res. 29:148-151. Pelham, H.R.B. 1991. Recycling of proteins betweeen the Endoplasmic Reticulum and the Golgi Complex. Curr. Opin. Cell Biol. 3:585591. Pruitt, K.D. and Maglott, D.R. 2001. RefSeq and LocusLink: NCBI gene-centered resources. Nucleic Acids Res. 29:137-140. Puntervoll, P., Linding, R., Gemund, C., ChabanisDavidson, S., Mattingsdal, M., Cameron, S., Martin, D.M., Ausiello, G., Brannetti, B., Costantini, A., Ferre, F., Maselli, V., Via, A., Cesareni, G., Diella, F., Superti-Furga, G., Wyrwicz, L., Ramu, C., McGuigan, C., Gudavalli, R., Letunic, I., Bork, P., Rychlewski, L., Kuster, B., Helmer-Citterich, M., Hunter, W.N., Aasland, R., and Gibson, T.J. 2003. ELM server: A new resource for investigating short functional sites in modular eukaryotic proteins. Nucleic Acids Res. 31:3625-3630. Schneider, M., Tognolli, M., and Bairoch, A. 2004. The Swiss-Prot protein knowledgebase and ExPASy: Providing the plant community with high quality proteomic data and tools. Plant Physiol. Biochem. 42:1013-1021. Smith, R.F., Wiese, B.A., Wojzynski, M.K., Davison, D.B., and Worley, K.C. 1996. BCM Search Launcher: An integrated interface to molecular biology data base search and analysis services available on the World Wide Web. Genome Res. 6:454-462. Teufel, A., Krupp, M., Weinmann, A., and Galle, P.R. 2006. Current bioinformatics tools in genomic biomedical research. Int. J. Mol. Med. 17:967-973.
Letunic, I., Goodstadt, L., Dickens, N.J., Doerks, T., Schultz, J., Mott, R., Ciccarelli, F., Copley, R.R., Ponting, C.P., and Bork, P. 2002. Recent improvements to the SMART domain-based sequence annotation resource. Nucleic Acids Res. 30:242-244.
Thorson, J.A., Yu, L.W.K., Hsu, A.L., Shih, N.Y., Graves, P.R., Tanner, J.W., Allen, P.M., PiwnicaWorms, H., and Shaw, A.S. 1998. 14-3-3 proteins are required for maintenance of Raf-1 phosphorylation and kinase activity. Mol. Cell. Biol. 18:5229-5238.
Maignan, S., Guilloteau, J.P., Fromage, N., Arnoux, B., Becquart, J., and Ducruix, A. 1995. Crystal structure of the mammalian Grb2 adaptor. Science 268:291-293.
Townsley, F.M., Frigerio, G., and Pelham, H.R.B. 1994. Retrieval of Hdel proteins is Required for growth of yeast cells. J. Cell Biol. 127:21-28.
Marchler-Bauer, A., Anderson, J.B., DeWeeseScott, C., Fedorova, N.D., Geer, L.Y., He, S., Hurwitz, D.I., Jackson, J.D., Jacobs, A.R., Lanczycki, C.J., Liebert, C.A., Liu, C., Madej, T., Marchler, G.H., Mazumder, R., Nikolskaya, A.N., Panchenko, A.R., Rao, B.S., Shoemaker, B.A., Simonyan, V., Song, J.S., Thiessen, P.A., Vasudevan, S., Wang, Y., Yamashita, R.A., Yin, J.J., and Bryant, S.H. 2003. CDD: A curated Entrez database of conserved domain alignments. Nucleic Acids Res. 31:383-387.
Wheeler, D.L., Chappey, C., Lash, A.E., Leipe, D.D., Madden, T.L., Schuler, G.D., Tatusova, T.A., and Rapp, B.A. 2000. Database resources of the National Center for Biotechnology Information. Nucleic Acids Res. 28:10-14.
Obenauer, J.C., Cantley, L.C., and Yaffe, M.B. 2003. Scansite 2.0: Proteome-wide prediction of
Wang, Z. and Moran, M.F. 1996. Requirement for the adapter protein Grb2 in EGF receptor endocytosis. Science 272:19355-1939.
Yoon, S.Y., Koh, W.S., Lee, M.K., Park, Y.M., and Han, M.Y. 1997. Dynamin II associates with Grb2 SH3 domain in Ras transformed NIH3T3 cells. Biochem. Biophys. Res. Comm. 234:539543.
Computational Analysis
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KEY REFERENCES Balla et al., 2006. See above. Describes the MnM database, algorithms used in scoring, and methods used to validation the scoring approaches.
INTERNET RESOURCES
http://elm.eu.org/ ELM: Online Web site for identifying short functional motifs. http://myhits.isb-sib.ch/cgi-bin/pattern search myHITS (pattern search): Searches several sequence databases for motifs.
http://mnm.engr.uconn.edu Minimotif Miner (MnM):,Online Web site for identifying short functional motifs.
http://www.ncbi.nlm.nih.gov/BLAST/ BLAST: Searches proteins for short, nearly exact matches.
http://pymol.sourceforge.net/ PyMOL: Shareware software for viewing motifs in protein structures.
http://scansite.mit.edu/motifscan seq.phtml SCANSITE: Online Web site for identifying short functional motifs.
http://hugin.ethz.ch/wuthrich/software/molmol/ index.html MolMol: Downloadable software for viewing protein structures.
http://ca.expasy.org/tools/ ExPASy: Contains a collection of tools for prediction of specific motifs.
http://www.rcsb.org/pdb/ Protein Data Bank (PDB): Repository of macromolecular structures. http://www.ncbi.nlm.nih.gov/omim/ McKusick, V.A. 2000. Online Mendelian Inheritance in Man (OMIM). McKusick-Nathans Institute for Genetic Medicine, Johns Hopkins University, (Baltimore, Md. and National Center for Biotechnology Information, National Library of Medicine, Bethesda, Md.
http://www.hgmd.cf.ac.uk/ac/index.php HGMD: The Human Genome Mutations Database, which contains a large collection of mutations known to be associated with human disease.
Contributed by Martin R. Schiller University of Connecticut Health Center Farmington, Connecticut
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CHAPTER 3 Detection and Assay Methods INTRODUCTION
T
he success or failure of protein-centered projects can frequently be traced to the quality of the analytical procedures used to characterize the sample at different stages. Qualitative and quantitative analysis can aid in definition of the sample for the purpose of designing separations. A knowledge of the properties of the desired protein (e.g., whether it has a high aromatic amino acid content) can suggest methods of analysis that will help locate the desired protein in a complex mixture. Establishing the properties of an isolated protein creates benchmarks that future researchers can use to evaluate their protocols and final product. Accurate quantitation of the amount of protein at the beginning, middle, and end of a series of steps is the only valid way to evaluate the overall yield of a procedure. The observation of significant loss of protein following a particular purification procedure, without a substantial increase in the purity of the desired protein, would indicate that the procedure should be omitted or revised.
Several spectroscopic procedures for characterizing protein samples are described in UNIT 3.1. Measuring the absorbance of the aromatic amino acids in a protein at different wavelengths yields a very useful measure of protein concentration. This is nondestructive and requires very little sample or time. A more qualitative, but much more sensitive, evaluation is provided by fluorescence spectroscopy. Another approach is amino acid analysis—qualitative analysis, to determine purity, and quantitative analysis, to provide concentrations—both of which are presented in UNIT 3.2. Procedures are also given for calculating amino acid ratios from primary analytical data. Significant advances that have improved the precision and sensitivity of amino acid analysis have reinvigorated this method, which had for some years been neglected. A common method for enhancing the sensitivity of protein detection is to radiolabel the proteins, as detailed in UNIT 3.3. This approach may also be employed to help quantitate the binding of peptide to other molecules or for radioimmunoassays. Several procedures for iodination (introduction of 125I or 131I) are presented; the choice between them will depend on the sample type, the content of residues sensitive to oxidation, and the amino acids that are critical to function. Procedures for handling radioactive samples, for separating labeled from unlabeled peptides, and for iodination to equimolar or higher levels (using a mixture of radioactive and nonradioactive iodine isotopes) are also provided. When Tyr or His residues are absent, or cannot be iodinated because they are critical for function, alternative methods involving the incorporation of 14C or 3H may be employed instead. The methods described include labeling by acetylation at amino groups on lysine or at the amino terminus using the Bolton-Hunter reagent or acetic anhydride, by reductive alkylation, and by labeling at Lys residues with iodoacetic acid or iodoacetamide. Two synthetic methods, reduction of a peptide containing iodo-Tyr residues and incorporation of labeled amino acids, are also presented. Measurement of the total protein content of a sample is discussed in UNIT 3.4. Three methods based on reduction of Cu2+ to Cu+ followed by chelation of the cuprous ion by various reagents (biuret, Hartree-Lowry, and bicinchinonic acid) are presented. In addition, a quantitative measure of total protein based on protein hydrolysis and the ninhydrin Contributed by Ben M. Dunn Current Protocols in Protein Science (2001) 3.0.1-3.0.2 Copyright © 2001 by John Wiley & Sons, Inc.
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reaction is described. The use of UV absorption, Coomassie dye binding, and weight analysis to determine protein content complete the presentation. Important considerations for performing these assays, such as dealing with interfering substances, are detailed throughout the unit. Some aspects involved in the design of enzyme assays are discussed in UNIT 3.5. Parameters that affect both enzyme and substrate are evaluated and suggestions to optimize the quantitation of enzymatic activity are enumerated. The proper concentration range for conducting initial velocity measurements is presented, and different methods of data presentation and analysis are discussed. Tagging a protein with a group that permits facile detection is an excellent way, for example, to study the movement of a protein through internalization or compartmentalization. A variety of reagents are available for attaching biotin to functional groups or the surface of a protein (UNIT 3.6). Detection and localization can be achieved by the strong and specific binding of avidin or streptavidin. These reactions can also be used to label proteins at cell surfaces. Another highly sensitive approach to the detection of proteins is the incorporation of radiolabeled amino acids into proteins. Integration of a particular radiolabeled amino acid (i.e., [35S]D-methionine or [3H]leucine) can be achieved using the normal cellular machinery (UNIT 3.7). Both pulse-labeling methods that permit study of cellular trafficking and long-term labeling to increase the detection of relatively rare proteins are described. The occurrence of selenium in proteins in the form of selenocysteine is now recognized as essential for several enzymatic functions. Therefore, methods for analysis of selenocysteine in proteins have become more important (UNIT 3.8). A future update will consider selenomethionine incorporation and analysis, as it can be helpful in the determination of three-dimensional structure by X-ray diffraction methods. The availability of genomic sequences for a growing number of organisms has magnified the need for techniques to analyze the protein content of biological fluids, cell and tissue extracts, and other samples. The use of small surfaces with defined properties to capture proteins from complex mixtures with subsequent identification by mass spectrometry is the topic of UNIT 3.9. Analytical assay methods to quantitate total protein content by biological activity will be presented in future updates to this chapter. Ben M. Dunn
Introduction
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Spectrophotometric Determination of Protein Concentration
UNIT 3.1
This unit describes methods for measuring the concentration of a protein in solution using absorbance spectroscopy. The absorbance, A, is a linear function of the molar concentration, c, according to the Beer-Lambert Law: A=ε×l ×c Equation 3.1.1
where ε is the molar absorption coefficient (M−1cm−1) and l is the cell pathlength (cm). It is important to have an accurate value of ε to determine c from A. Basic Protocol 1 shows how to calculate ε for a protein, making use of the average ε values for the three chromophores in proteins (the side chains of tryptophan, tyrosine, and cystine), as determined from an analysis of a large set of proteins with known ε values. Basic Protocol 2 describes how ε can be determined for a folded protein by measuring the absorbance of two solutions of the protein with identical concentrations—one containing just buffer and one containing buffer plus 6 M guanidine⋅HCl. Basic Protocol 3 shows how to measure the concentration of a protein using the Beer-Lambert law after ε has been either calculated or measured. Basic Protocol 4 describes a more sensitive method that is especially useful for proteins that contain few tryptophan or tyrosine residues. Basic Protocol 5 provides an easy way to estimate the total protein concentration in crude protein extracts during the early steps of a purification procedure. CALCULATION OF THE MOLAR ABSORPTION COEFFICIENT (ε) OF A PROTEIN
BASIC PROTOCOL 1
The molar absorption coefficient of a protein at 280 nm, ε280 (in M–1 cm–1 ), is calculated using the following equation (Pace et al., 1995; Pace and Schmid, 1997): ε 280 = (5500 × nTrp ) + (1490 × nTyr ) + (125 × nS-S ) Equation 3.1.2
where the numbers are the molar absorbances for tryptophan (Trp), tyrosine (Tyr), and cystine (i.e., the disulfide bond, S-S), from the first column in Table 3.1.1, and nTrp = number of Trp residues, nTyr = number of Tyr residues, and nS-S = number of disulfide bonds in the protein. This method is simple, fast, and reliable. The amino acid composition of the protein and the number of disulfide bonds are the only information needed. 1. Count the number of Trp residues (nTrp), Tyr residues (nTyr), and disulfide bonds in the protein sequence. 2. Use Equation 3.1.2 and the numbers from step 1 to calculate ε280.
Detection and Assay Methods Contributed by Gerald R. Grimsley and C. Nick Pace Current Protocols in Protein Science (2003) 3.1.1-3.1.9 Copyright © 2003 by John Wiley & Sons, Inc.
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Table 3.1.1
Values of ε for Trp, Tyr, and Disulfide Bonds Used to Determine ε of a Proteina
Average values of ε280 in folded proteinsb (M– 1cm– 1) Trp Tyr S-S
5500 1490 125
Values of ε for model compounds in 6.0 M guanidine⋅HCl at specified wavelengthsc (M– 1cm– 1) 276 nm 5400 1450 145
278 nm 5600 1400 127
279 nm 5660 1345 120
280 nm 5690 1280 120
282 nm 5600 1200 100
aReproduced from Pace and Schmid (1997) with permission from IRL press. bFrom Pace et al. (1995). cData are for N-acetyl-L-tryptophanamide, Gly-L-Tyr-Gly, and for S-S (disulfide bond) in 6.0 M guanidine⋅HCl, 0.02 M phosphate buffer, pH 6.5. Values from Gill and von Hippel (1989); original data from Edelhoch (1967).
BASIC PROTOCOL 2
DETERMINATION OF THE MOLAR ABSORPTION COEFFICIENT (ε) FOR A FOLDED PROTEIN In this protocol, the value of ε for the unfolded protein in 6.0 M guanidine⋅HCl (εU) is first calculated from the number of Trp, Tyr, and disulfide bonds with Equation 3.1.2, but using the ε values for those residues determined in 6 M guanidine⋅HCl, which are given in Table 3.1.1. These ε values are based on experimental studies of appropriate model compounds. This procedure is generally called the Edelhoch method (Edelhoch, 1967). Edelhoch showed that ε values for model compounds determined in 6.0 M guanidine⋅HCl could accurately reproduce the absorbance spectrum of an unfolded protein in 6.0 M guanidine⋅HCl. Values of ε for the reference compounds are given in Table 3.1.1 at five wavelengths between 276 and 282 nm. The εU for the unfolded protein is calculated at the wavelength of the absorbance maximum of the native protein. The ε value for the folded protein (εF) is determined by preparing solutions of folded and unfolded protein with identical concentrations, then measuring their absorbances, AF and AU. Since the concentrations of folded and unfolded protein are equal, εF and εU are related by: AF ε = F AU εU Equation 3.1.3
and εF can be calculated using: A εF = εU × F AU Equation 3.1.4
The steps for determining the molar absorption coefficient of the folded protein are given below. Spectrophotometric Determination of Protein Concentration
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Materials 8 M nitric acid Ethanol 20 mM potassium phosphate buffer, pH 6.5 (APPENDIX 2E) Protein stock solution: ∼1 mg protein in 0.25 ml of 20 mM potassium phosphate buffer, pH 6.5 (see APPENDIX 2E for buffer) 6.6 M guanidine⋅HCl (APPENDIX 3A) in 40 mM potassium phosphate buffer, pH 6.5 (APPENDIX 2E); also see Critical Parameters Double-beam absorbance spectrophotometer Two quartz cuvettes, 10 mm long × 4 mm wide Prepare spectrophotometer and cuvettes 1. Turn on the spectrophotometer and UV and visible lamps. Allow a 30-min warm up period. Thermostatting is not required when the ambient temperature is between 20° and 30°C.
2. Clean cuvettes thoroughly by soaking in an 8 M nitric acid bath and then washing with distilled water. Rinse with ethanol and air dry. Obtain spectrum of native protein 3. Add 910 µl of 20 mM potassium phosphate buffer, pH 6.5 to both the sample and reference cuvettes and scan the absorbance between 350 and 250 nm. This baseline should be flat or deviate by no more than 0.005 AU from the instrument’s baseline with the cells removed.
4. Record the baseline. In addition, individually measure the absorbance values at 276, 278, 279, 280, and 282 nm, as the protein absorbance maximum will typically be in this range. See Critical Parameters for discussion of the linear range of the spectrophotometer.
5. Add 90 µl of the protein stock solution to the sample cuvette and 90 µl of 20 mM potassium phosphate buffer, pH 6.5 to the reference cuvette. Mix the solutions well in the cells. Scan and record the absorbance values at 276, 278, 279, 280, and 282 nm as in steps 3 and 4 above. Repeat the measurement to check reproducibility. 6. Subtract the baseline and the individual absorbance values of the buffer at 276, 278, 279, 280, and 282 nm. Obtain spectrum of unfolded protein 7. Obtain a spectrum of protein as in steps 1 to 6 above, but use 910 µl of 6.6 M guanidine⋅HCl in 40 mM potassium phosphate buffer, pH 6.5 in place of 20 mM potassium phosphate buffer, pH 6.5. 8. Calculate the value of εU for the unfolded protein between 276 nm and 282 nm as described in Basic Protocol 1, using the values of ε for the residues in 6.0 M guanidine⋅HCl (Table 3.1.1).
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Calculate molar absorption coefficient for the folded protein 9. Correct for light scattering, if necessary (see Critical Parameters and Troubleshooting). An absorbance greater than zero at wavelengths above 320 nm generally results from light scattering.
10. Determine the ratios AF/AU at 276 to 282 nm and use Equation 3.1.4 to calculate εF in this wavelength range. BASIC PROTOCOL 3
DETERMINATION OF PROTEIN CONCENTRATION BY ABSORBANCE SPECTROSCOPY USING THE MOLAR ABSORPTION COEFFICIENT This protocol describes how to measure the concentration of a pure protein in solution using the molar absorption coefficient determined in Basic Protocol 1 or 2. Materials Solution of pure protein in either distilled water or buffer Spectrophotometer with UV and visible lamps Quartz cuvette 1. Turn on the spectrophotometer and UV and visible lamps. Allow a 30-min warm up period. 2. Zero the spectrophotometer with a solvent blank (distilled water or buffer) at the wavelength where ε was determined and also at 330 nm. The reading at 330 nm serves to detect light scattering (see Critical Parameters and Troubleshooting).
3. Remove the cuvette, discard the solvent, and thoroughly dry the cuvette. Add the protein solution and replace the cuvette in the spectrophotometer. 4. Record the absorbance of the sample at the appropriate wavelengths. 5. If necessary, correct for light scattering (see Critical Parameters and Troubleshooting). 6. Calculate the protein concentration (c) in mg/ml: c = ( A280 × mol. wt.) /(ε280 × l ) Equation 3.1.5
The path length, l, is usually 1 cm.
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DETERMINATION OF PROTEIN CONCENTRATION BY ABSORPTION SPECTROSCOPY AT 205 nm
BASIC PROTOCOL 4
If the protein of interest contains few or no Trp or Tyr residues, the concentration may be determined by measuring the absorbance at 205 nm (A205). This is generally called the Scopes method (Scopes, 1974). Most of the absorbance at 205 nm is due to peptide bonds, but Scopes described a method for estimating ε at 205 nm (ε205) for proteins that also contain some Trp or Tyr residues. The Scopes method is the method of choice for proteins that do not contain Trp or Tyr residues. It works best for dilute protein solutions (1 to 100 µg/ml) that are in nonabsorbing buffers or solvents. Materials Protein sample Brij 35 solution: 0.01% (v/v) Brij 35 (Sigma) in an aqueous solution appropriate for dissolving the sample protein Spectrophotometer with UV and visible lamps Quartz cuvette 1. Dissolve protein in Brij 35 solution. 2. Turn on the spectrophotometer and UV lamp. Allow a 30-min warm up period. Set the wavelength to 205 nm. 3. Zero the spectrophotometer with the Brij 35 solution without protein. 4. Measure the A205 of the sample protein. 5a. If ε205 is known for the protein, use Equation 3.1.5 to calculate the concentration, substituting A205 for A280 and ε205 for ε280. 5b. If ε205 is not known, estimate the concentration of the sample protein (c) in mg/ml as follows:
c (mg/ml) = A205 /(31 × l )
Equation 3.1.6
SPECTROPHOTOMETRIC DETERMINATION OF TOTAL PROTEIN CONCENTRATION IN CRUDE PROTEIN EXTRACTS
BASIC PROTOCOL 5
A reasonable estimate of the protein concentration in an extract can be obtained by assuming that the absorbance at 280 nm of a 1% solution of the protein (A2801%) equals 13. This value is the average of the measured (A2801%) values for 116 proteins of various molecular weights and amino acid compositions (Pace et al., 1995). This protocol is most useful for monitoring protein concentration in the early stages of a purification procedure. Materials Sample of protein extract Spectrophotometer with UV and visible lamps Quartz cuvette Detection and Assay Methods
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1. Turn on the spectrophotometer and UV and visible lamps. Allow a 30-min warm up period. 2. Zero the spectrophotometer at 280 nm with a solvent blank (distilled water or buffer). 3. Remove the cuvette, discard the solvent, and thoroughly dry the cuvette. Add the protein extract and replace the cuvette in the spectrophotometer. 4. Record the absorbance of the sample at 280 nm. See Critical Parameters for discussion of the linear range of the spectrophotometer.
5. Estimate the protein concentration by assuming that a 1 mg/ml solution of protein will have an absorbance of 1.3. COMMENTARY Background Information
Spectrophotometric Determination of Protein Concentration
There are many methods available for determining protein concentration. Most of these methods rely either on colorimetric assays (e.g., UNIT 3.4) or the use of UV absorbance spectroscopy (Stoscheck, 1990). The choice of method depends upon a variety of factors, including the amount of protein present, the specificity of the method, the presence of interfering substances, and the amino acid composition of the protein. The Lowry method (UNIT 3.4) is a colorimetric method that has been widely used by protein chemists for the past 50 years (Lowry et al., 1951). The fact that Lowry’s publication of the methods is the most cited paper in the history of biochemistry attests to its popularity. In the mid-1970s, the Bradford method (Bradford, 1976; UNIT 3.4), based on binding of a blue dye to the protein, became popular and eventually supplanted the Lowry method. However, colorimetric assays are generally only useful for monitoring protein purification and should not be used when an accurate protein concentration is required. For the accurate determination of protein concentration, UV absorbance spectroscopy is the preferred method. UV absorbance spectroscopy is one of the oldest methods for determining protein concentration (Warburg and Christian, 1942) and has numerous advantages. The protein solution is used directly, without the addition of reagents, and is not modified or inactivated by the process. No incubation period is required, so measurements can be made rapidly and with high reproducibility. Finally, the protein concentration can be determined accurately from absorbance measurements using the Beer-Lambert Law (Equation 3.1.1). An accurate value of ε is essential when using Equation 3.1.1. Determination of ε requires an accurate measurement of A and c. The
measurement of A is straightforward (see Schmid, 1997 for an excellent discussion of how absorbance spectroscopy can be used to study proteins), but the measurement of c is not. The four techniques most often used to measure c are amino acid analysis (UNIT 3.2), Kjelkahl nitrogen determination, the dry weight method, and the Edelhoch method. The Edelhoch method is the simplest and most reliable experimental method to determine c, and consequently ε (Pace et al., 1995). Basic Protocol 1 describes the simplest method of estimating ε. Nevertheless, it is reliable, and in a test with 80 proteins it gave an average deviation from measured values of <4% (Pace et al., 1995). Basic Protocol 2 requires absorbance measurements on two solutions, but it is still simple and straightforward and leads to an experimental determination of ε. This method should be used when an accurate ε value is crucial. Basic Protocol 4 should be used for the rare proteins that contain neither Trp nor Tyr residues.
Critical Parameters and Troubleshooting Chromophores Basic Protocol 1 cannot be used if the protein of interest contains a tightly bound ligand or prosthetic group that absorbs between 250 and 300 nm. By comparing the protein spectrum with the absorbance spectra of Trp and Tyr (Schmid, 1997), one can often detect nonprotein contributions. For example, bound nucleotides lead to a decrease in the ratio A280/A260. If the disulfide bond content is unknown, assume for cytosolic proteins that the number of disulfide bonds (nS-S) is zero and assume for secretory proteins that nS-S = nCys/2 (Pace and Schmid, 1997). Since the molar absorption co-
3.1.6 Supplement 33
Current Protocols in Protein Science
Table 3.1.2
Absorbances of Various Salt and Buffer Components in the Far-UV Rangea
Compoundb
NaClO4 NaF, KF Boric acid NaCl Na2HPO4 NaH2PO4 Sodium acetate Glycine Diethylamine NaOH, pH 12 Boric acid/NaOH, pH 9.1 Tricine, pH 8.5 Tris, pH 8.0 HEPES, pH 7.5 PIPES, pH 7.0 MOPS, pH 7.0 MES, pH 6.0 Cacodylate, pH 6.0
No absorbance Absorbance of a 0.01 M solution in a 0.1-cm cell above 170 nm 170 nm 180 nm 205 nm 210 nm 195 nm 220 nm 220 nm 240 nm 230 nm 200 nm
210 nm 0 0 0 0 0 0 0.03 0.03 0.4 >0.5 0
200 nm 0 0 0 0.02 0.05 0 0.17 0.1 >0.5 >2 0
190 nm 0 0 0 >0.5 0.3 0.01 >0.5 >0.5 >0.5 >2 0.09
180 nm 0 0 0 0.5 >0.5 0.15 >0.5 >0.5 >0.5 >2 0.3
230 nm 220 nm 230 nm 230 nm 230 nm 230 nm 210 nm
0.22 0.02 0.37 0.20 0.10 0.07 0.01
0.44 0.13 0.5 0.49 0.34 0.29 0.20
>0.5 0.24 >0.5 0.29 0.28 0.29 0.22
>0.5 >0.5 >0.5 >0.5 >0.5 >0.5 >0.5
aTable from Schmid (1997) reproduced with permission from IRL press. bBuffers titrated with 1 M NaOH or 0.5 M H SO to the indicated pH values. 2 4
efficient for the disulfide bond is small, errors in the number of disulfide bonds have only a small effect on the calculated absorption coefficient. Basic Protocol 1 works best for globular proteins that contain at least one Trp residue (Pace et al., 1995). Guanidine⋅HCl When performing Basic Protocol 2, use ultrapure guanidine⋅HCl. Since guanidine⋅HCl is quite hygroscopic, it is difficult to prepare stock solutions by weight. Consequently, the molarity of guanidine⋅HCl stock solutions is generally based on refractive index measurements (see Pace and Scholtz, 1997, on how to prepare guanidine⋅HCl stock solutions). For the purposes of this protocol, the exact molarity of the guanidine⋅HCl stock solution is not critical, only that it be near 6 M. This can be approximated by adding 1 g of guanidine⋅HCl per ml of distilled water. A 40 mM buffer is used with the guanidine⋅HCl because in a 6.6 M solution, guanidine⋅HCl occupies about half of the volume of the solution, so the final buffer concentration is actually ∼20 mM. The buffer concentration here is not crucial as long as the correct
pH is maintained. Be sure to filter all solvents to remove dust particles (a 0.45-µm syringe filter works best). Careful pipetting and mixing of all solutions is essential for the reproducibility of this procedure. Light scattering When measuring the absorbance in Basic Protocols 2 and 3, a correction must be made if there is significant light scattering. The absorbance of the protein should be zero above 320 nm. A simple method to correct for light scattering is to multiply the absorbance at 330 nm by 1.929 to get the light scattering contribution at 280 nm or by 1.986 to get the light scattering contribution at 278 nm, then subtract the resulting values from AF and AU as measured at these wavelengths (Pace et al., 1995). This assumes that the light scattering contribution varies as the inverse fourth power of the wavelength and is discussed in detail by Leach and Scheraga (1960). Spectrometer linear range For the most accurate results, the maximum absorbance near 280 should be between 0.2 and
Detection and Assay Methods
3.1.7 Current Protocols in Protein Science
Supplement 33
Table 3.1.3
Concentration Limits of Interfering Reagents for A205 and A280 Protein Assaysa
Reagentsb
A205
A280
Ammonium sulfate Brij 35 DTT EDTA Glycerol KCl 2-ME NaCl NaOH Phosphate buffer SDS Sucrose Tris buffer Triton X-100 TCA Urea
9% (w/v) 1% (v/v) 0.1 mM 0.2 mM 5% (v/v) 50 mM <10 mM 0.6 M 25 mM 50 mM 0.10% (w/v) 0.5 M 40 mM <0.01% (v/v) <1% (w/v) <0.1 M
>50% (w/v) 1% (v/v) 3 mM 30 mM 40% (v/v) 100 mM 10 mM >1 M >1 M 1M 0.10% (w/v) 2M 0.5 M 0.02% (v/v) 10% (w/v) >1 M
aValues from Stoscheck (1990). bAbbreviations: DTT, dithiothreitol; EDTA, ethylenediaminetetraacetic acid; 2-ME.
1.0. When the absorbance is outside this range, make a new protein solution so that A280 is within this range. Finally, be sure that the total volume in the cell is sufficient that the light beam strikes the solution rather than the meniscus. The height of the light beam can be examined visually in the visible range (e.g., near 500 nm) by inserting a strip of white paper into the cell. Buffer effects The Brij 35 solution in Basic Protocol 4 is added to prevent protein adsorption onto vessel surfaces (Stoscheck, 1990), a significant problem when working with very dilute solutions. In addition, at 205 nm, a number of common solvents and buffers may contribute substantially to the total absorbance. Table 3.1.2 provides helpful information for selecting buffers when working in the far-UV region. Table 3.1.3 lists acceptable concentrations of potentially interfering reagents for use at 205 or 280 nm.
Spectrophotometric Determination of Protein Concentration
Nucleic acid interference Basic Protocol 5 is most useful for estimating protein concentration where there is little nucleic acid present. Nucleic acids contribute substantially to A280. To obtain a better estimate of protein concentration in the presence of nucleic acid, measure the absorbance at 260 nm
and 280 nm and calculate the protein concentration, c (in mg/ml), as follows (Stoscheck, 1990):
c = (1.55 × A280 ) − (0.76 × A260 ) Equation 3.1.7
Anticipated Results Accurate protein concentrations can be obtained by measuring the absorbance near 280 nm and then using the Beer-Lambert Law. The accuracy is limited mainly by the accuracy of ε. Equation 3.1.2 can be used to estimate ε quite accurately, but the best approach is to measure rather than predict ε.
Time Considerations All of the protocols described in this unit can be performed in <1 hr for a single sample.
Literature Cited Bradford, M.M. 1976. A rapid and sensitive method for quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72:248-254. Edelhoch, H. 1967. Spectroscopic determination of tryptophan and tyrosine in proteins. Biochemistry 6:1948-1954.
3.1.8 Supplement 33
Current Protocols in Protein Science
Gill, S.C. and von Hippel, P.H. 1989. Calculation of protein extinction coefficients from amino acid sequence data. Anal. Biochem. 182:319-326.
Scopes, R.K. 1974. Measurement of protein by spectrophotometry at 205 nm. Anal. Biochem. 59:277-282.
Leach, S.J. and Scheraga, H.A. 1960. Effect of light scattering on ultraviolet difference spectra. J. Am. Chem. Soc. 82:4790-4792.
Schmid, F.X. 1997. Optical spectroscopy to characterize protein conformation and conformational changes. In Protein Structure: A Practical Approach. (T.E. Creighton, ed.) pp. 261-297. IRL Press, Oxford, U.K.
Lowry, O.H., Rosebrough, J.J., Farr, A.L., and Randall, R.J. 1951. Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193:265275.
Stoscheck, C.M. 1990. Quantitation of protein. Methods Enzymol. 182:50-68.
Pace, C.N. and Schmid, F.X. 1997. How to determine the molar absorbance coefficient of a protein. In Protein Structure: A Practical Approach. (T.E. Creighton, ed.) pp. 253-259. IRL Press, Oxford, U.K.
Warburg, O. and Christian, W. 1942. Isolierung und Kristallisation des Garungsferments Enolase. Biochem. Z. 310:384-421.
Pace, C.N. and Scholtz, J.M. 1997. Measuring the conformational stability of a protein. In Protein Structure: A Practical Approach (T.E. Creighton, ed.) pp. 299-321. IRL Press, Oxford, U.K.
Contributed by Gerald R. Grimsley and C. Nick Pace The Texas A&M University System Health Science Center College Station, Texas
Pace, C.N., Vajdos, F., Fee, L., Grimsley, G., and Gray, T. 1995. How to measure and predict the molar absorption coefficient of a protein. Prot. Sci. 4:2411-2423.
Detection and Assay Methods
3.1.9 Current Protocols in Protein Science
Supplement 33
Quantitative Amino Acid Analysis The use of quantitative amino acid analysis in the characterization of protein samples is often overlooked in many laboratories. This was typically due to the difficulties involved in obtaining reliable amino acid composition data, as well as to the advent of colorimetric dye-binding assays as alternative methods to determine protein concentration. However, many proteins do not bind dyes in the same way as the proteins normally used as standards, making quantitation by those techniques difficult. Fortunately, at many institutions, the creation of protein chemistry core laboratories devoted mainly to protein sequence analysis and peptide synthesis has provided new instrumentation that yields the necessary precision. This unit describes the information that can be derived from quantitative amino acid analysis, presents some details on sample preparation, and gives examples of the calculations that are needed.
INFORMATION OBTAINED FROM AMINO ACID ANALYSIS The benefits of amino acid composition data are two-fold. First, identifying the nature of the amino acids present in a sample can provide an important qualitative measure of the purity of the preparation. Amino acid analysis does not depend on the shape, charge, or function of a protein because the hydrolysis completely destroys structure and reduces the protein to its constituent amino acids. Although large proteins typically contain a mixture of all the commonly occurring amino acids, certain residues are relatively rare—Trp, Met, His, Cys, and occasionally Arg are present at lower than average ratios compared to Gly, Ala, Ser, and Leu. For a pure protein, the ratios of the amino acids should approach the theoretical levels to within ∼10%, with a few caveats. For a sample from a column fraction, for example, the presence of a higher than expected ratio of a certain amino acid is a very good indication that the sample is not pure and that further fractionation is indicated. In the same way, a lower than expected ratio could also indicate impurity— for example, the presence of a second protein that has a lower ratio of some significant amino acid present in higher amounts in the protein of interest. Because the expression of a protein in bacteria or other hosts utilizes a gene of known sequence, the expected amino acid composition
Contributed by Ben Dunn Current Protocols in Protein Science (1995) 3.2.1-3.2.3 Copyright © 2000 by John Wiley & Sons, Inc.
is also known (see Chapters 5 and 6 for expression and purification of recombinant proteins). Second, averaged composition yields an excellent measure of the amount of protein in a sample. On the ion-exchange-based Beckman 6300 amino acid analyzer running with ninhydrin detection chemistry, the minimum amount of the standard (a mixture of amino acids used to derive integration parameters as well as elution times) that is required is typically 1 nmol of each amino acid. Amounts as low as 250 pmol (0.25 nmol) of an amino acid can be measured with confidence from protein hydrolyzates. For a protein of 50 kDa mass, 1 nmol is 50 µg, an amount that can be obtained readily by purification of the protein from a high-expression system followed by purification by standard methods. For isolation of a trace protein from a natural source, this required amount may be too large to be feasible. However, a protein of 50 kDa mass contains ∼400 amino acids, or roughly 20 copies of each amino acid on average. Thus, the actual amount of protein required for detection and semiquantitative analysis could be as low as 0.05 nmol, or 2.5 µg. At that level, rare amino acids cannot be accurately quantitated, but the more common ones can be measured easily. Obviously, the more material available, the more accurate the analysis. Users should check with their local facility for specific advice on the amount of sample that can be processed by existing technologies. The advent of advanced instrumentation, such as the Applied Biosystems 420 amino acid analyzer and the Waters PicoTag system, has provided another increment in sensitivity. These instruments utilize precolumn derivatization followed by separation based on reversed-phase high-performance liquid chromatography (HPLC). With such analyzers, 100 pmol of each amino acid, or 5 pmol of the 50-kDa protein described above, can be quantitated accurately. Another important consideration in the design of a purification scheme that can be accomplished through the use of amino acid quantitation is the compilation of the mass balance—the relationship between how much protein is applied to the chromatography column and how much is recovered in a particular step. Amino acid analysis of a measured aliquot of the sample taken before application (typically ≤1%) along with aliquots of column frac-
UNIT 3.2
Detection and Assay Methods
3.2.1 CPPS
tions or pools, can be used to calculate the amounts of protein loaded and recovered. These calculations can provide essential information to assess the success of a particular purification procedure.
SAMPLE PREPARATION The success or failure of quantitative amino acid analysis is strongly dependent upon the quality of the sample, as well as upon the exact composition of the sample matrix (i.e., the aggregate of all components including salts, detergent, and dirt). The following section gives a few key recommendations for successful sample preparation.
Sample Concentration An ideal sample for amino acid analysis would contain 1 to 10 nmol protein in a volume of 10 to 100 µl water. A sample that has been dialyzed against a low concentration of buffer (if needed to maintain solubility) or pure water is optimal, because the presence of salt, detergent, high concentrations of buffers, or other agents may interfere with either hydrolysis or subsequent analysis. A sample that has been lyophilized to dryness is also usually acceptable, with some important considerations. First, lyophilization should be done in such a way as to concentrate the lyophilized residue at the bottom of a conical tube. Drying a sample in a round-bottom flask may leave the protein sample spread over a large surface area. Such samples are extremely difficult to handle and will not give satisfactory results. Second, lyophilization of a buffered solution will concentrate the salts as well as the protein. Therefore, it is recommended that the total ionic strength of the solution be <50 µM before lyophilization.
Estimate of Total Protein An estimate of the total amount of protein present is very helpful in planning an accurate analysis. This can be based on a colorimetric dye–binding assay, an A280 measurement, or an activity assay in which the results are compared to a standard value. All information about the sample should be communicated to the individual charged with conducting the analysis to ensure the maximum return on the sample. Many facilities provide a form with a series of questions to be answered concerning the sample. Quantitative Amino Acid Analysis
Samples from PVDF Blots With the advent of more sensitive analyzers
based on phenylthiohydantoin (PTH) chemistry, it is now possible to obtain compositional data on samples separated by electrophoresis and blotted onto membranes (UNIT 10.7) such as are used for protein sequence analysis. This is especially helpful in cases where a protein sample fails to yield any sequence during gas-phase Edman degradation. Acid hydrolysis and analysis of sample blot pieces recovered from the sequencer sample compartment yield a rough estimate of the amount of protein present. If the amount is larger than that normally required for sequence analysis (10 pmol), it may then be concluded that the protein sample was N-terminally blocked.
CALCULATION OF THE AVERAGE COMPOSITION Table 3.2.1 presents the results of analysis of a 24-kDa recombinant protein. The initial sample volume was 1.0 ml and was expected to contain ∼4 mg total protein. A 30-µl aliquot was taken for analysis and was estimated to contain 120 µg, or ∼5 nmol. The aliquot was transferred to a glass tube for hydrolysis and dried by application of a vacuum at room temperature. One milliliter of 6 M HCl was added, the tube was cooled in a dry ice bath, then a vacuum was applied and the tube sealed by heating in a gas flame. Hydrolysis was conducted for 24 hr at 124°C, followed by removal of the HCl under vacuum. The sample was taken up in 1.0 ml application buffer, 50 nmol norleucine was added as an internal standard, and a 50-µl aliquot was analyzed by ion-exchange chromatography on a Beckman 6300 system with ninhydrin detection. 1. The total number of nanomoles of amino acids detected is first calculated by summing the second column of Table 3.2.1, yielding 1446 nmol. 2. The total nanomoles is then divided by the total number of expected amino acids (based on protein sequence; in this case, 212) to obtain the average number of nanomoles per residue; 1446 nanomoles/212 residues = 6.82 nmol per residue. If the total number of amino acids is not known, it may be estimated by dividing the mol. wt. of the protein by 110. 3. The number of nanomoles of each amino acid measured is divided by the average number of nanomoles per residue (step 2) to give the ratio1 observed for that amino acid. For Asp, 155 ÷ 6.82 = 22.7. The resulting ratios are given in the fourth column and may be compared to the ratio expected listed in the third column. For most
3.2.2 Current Protocols in Protein Science
Table 3.2.1 Data Used to Calculate Amino Acid Composition for a 24-kDa Protein
Amino acid Asp Thr Ser Glu Pro Gly Ala Val Met Ile Leu Tyr Phe His Lys Arg
Nanomoles measured 155 78 63 153 52 157 90 159 24 88 138 43 56 16 121 53
Ratio expecteda
Ratio1 observed
Ratio2 observed
22.7 11.4 9.2 22.4 7.6 23.0 13.1 23.3 3.5 12.9 20.2 6.3 8.2 2.3 17.7 7.8
21.8 10.9 8.9 21.5 7.3 22.1 12.6 22.3 3.4 12.3 19.4 6.1 7.9 2.3 17.0 7.4
23 13 13 21 7 22 12 23 5 13 19 6 8 3 17 7
aRatio expected based on amino acid sequence of protein.
of the amino acids, the ratio1 observed agrees with the ratio expected. The value of average nanomoles per residue calculated in step 2 is based on the analysis of a 30-µl sample; the total sample of 1 ml contains ∼5.46 mg of protein (average moles/residue × grams/mole protein × 1000/30). This analysis utilized only 50 µl of the 1.0 ml final solution, so a sample size smaller than 3% of the original sample could have been used. Using an excess, however, provides the flexibility to repeat the determination if the first analysis gives peaks that are either too large or too small for accurate integration. The ratio1 observed for Ser, Thr, His, and Met are in this sample are ∼30%, 13%, 24%, and 30% below the expected values, respectively. These results are typical for these four amino acids, but for different reasons. The Ser and Thr results are low because these amino acids undergo partial degradation during acid hydrolysis. Ser and Thr losses of 10% to 40% are normal. The lower-than-expected values for His and Met are an anomaly that is most likely caused by the relatively low copy number of those two amino acids in the protein. Due to these known problems, most investigators choose to delete the values for sensitive residues (Ser, Thr) as well as the values for those amino acids appearing in low amounts (in this case, His, and Met) and calculate the average
number of nanomoles per residue independently of those amino acids. This gives the corrected ratios shown in the last column of Table 3.2.1 (ratio2 observed). In this example, the recalculated values are changed by as much as 0.9 due to the higher average number of nanomoles per residue (7.11). The deviation of ratio1 observed from the expected ratio ranges from 1% to 32% for the various amino acids, with an average error of 8.8%. When the high variation for Ser (32%), Met (32%), and His (24%) are removed from consideration, the average deviation of ratio2 observed from expected ratio drops to 4.1%. By standard criteria, this sample would be judged to be pure; of course, this result needs to be combined with electrophoretic separation (Chapter 10), chromatographic purification (Chapter 8), and perhaps mass spectroscopic analyses before a final assessment of the purity of the sample can be made.
Key Reference Ozols, J. 1990. Amino acid analysis. Methods Enzymol. 182:587-601. Excellent discussion of sample preparation issues and hydrolysis methods.
Contributed by Ben Dunn University of Florida Gainesville, Florida
Detection and Assay Methods
3.2.3 Current Protocols in Protein Science
Supplement 4
In Vitro Radiolabeling of Peptides and Proteins
UNIT 3.3
Radiolabeling of peptides or proteins is often performed to enhance the sensitivity of detection, to quantitate the binding of peptides to other molecules, or for radioimmunoassays. The choice of radioisotope depends in part on the amino acid residue(s) to be modified (Table 3.3.1). The most common and straightforward labeling strategy is radioiodination with 125I because of its relative ease of use, the high resulting specific radioactivities (curies per millimole), and the convenience of detecting γ emissions. Radioiodination may also be performed with 131I, which has a shorter half-life (8 days versus 60 days for 125I), giving higher specific radioactivities but limiting the useful lifetimes of the labeled products; in addition, 131I has higher-energy γ emissions, necessitating more shielding to protect workers. Iodination at tyrosine or histidine residues using Iodo-Beads (Basic Protocol 1) is the most convenient method, but chloramine T or Iodogen (Alternate Protocol 1) may give higher recoveries when working with small quantities of peptide, and lactoperoxidase (Alternate Protocol 2) results in less oxidative damage and lower specific radioactivities, and is commonly used when labeling whole cells. Modifications for achieving equimolar or higher (stoichiometric) rather than trace iodination are presented in Support Protocol 1, and the optional separation of unlabeled peptide from iodinated products by HPLC is described in Support Protocol 2. For peptides that do not contain tyrosine or histidine, or in which labeling of these amino acid residues interferes with biological activity, 125I or 131 I may be added onto primary amino groups (lysine residues and the peptide N-terminus) using Bolton-Hunter reagent (Basic Protocol 2). As an alternative to iodination, peptides or proteins can be labeled with 14C or 3H by acetylation of primary amino groups using anhydride (Basic Protocol 3), by reductive alkylation of primary amino groups using aldehyde and borohydride (Alternate Protocol 3), or by alkylation of cysteine sulfhydryl groups (Alternate Protocol 4). Figure 3.3.1 summarizes these radiolabeling strategies as well as the basic iodination reactions. Two methods are available to label a peptide without altering its structure: 3H labeling by catalytic reduction of (nonradioactive) 127I-labeled peptide (Basic Protocol 4) and de novo peptide synthesis using a radiolabeled form of any amino acid (Basic Protocol 5). However, these methods tend to be more expensive and laborious. Finally, selective Table 3.3.1
Peptide Labeling Strategies
Amino acid residue
Label
Method
Reference
Tyrosine Histidine, tyrosine Tyrosine Lysine, N-terminus Lysine, N-terminus
125I, 131I
Chloramine T, Iodo-Beads Chloramine T, Iodo-Beads Lactoperoxidase Bolton-Hunter Anhydride
Lysine, N-terminus Cysteine Tyrosine
14C, 3H
Any residue Carbohydrate groups
14C, 3H
Greenwood et al. (1963) Wolff and Covelli (1969) Marchalonis (1969) Bolton and Hunter (1973) Fraenkel-Conrat and Colloms (1967) Tack and Wilder (1981) Chersi et al. (1988) Amersham or Du Pont NEN literature Stewart and Young (1984) Gahmberg and Andersson (1977)
125I, 131I 125I, 131I 125I, 131I 14C, 3H
14C, 3H 3H
3H
Aldehyde/borohydride Iodoacetic acid Reduction of 127I Peptide synthesis Periodate/borohydride
Contributed by Ton N.M. Schumacher and Theodore J. Tsomides Current Protocols in Protein Science (1995) 3.3.1-3.3.19 Copyright © 2000 by John Wiley & Sons, Inc.
Detection and Assay Methods
3.3.1 CPPS
I
OH
A
I
OH
OH I
+NH 3
COO –
+NH 3
COO –
+NH 3
COO –
I
B N +NH 3
C
NH
N COO –
I
+NH 3
+
HO
COO–
N COO –
I
COO –
I HO
O N I
NH
+NH 3
O
I
+NH 3
+NH 3
NH
O I
O O
NH
I HO
COO –
NH I
O O
D
+NH 3
+ +NH 3
E
CH3 NH O
O O COO –
CH3 O CH3
COO –
CH3 NH CH3
+NH 3
+NH 3
+
COO –
CH2=O
NH
CH2=N
+
NaCNBH3
COO –
CH2=N
COO –
CH3–NH CH3
CH3 N
CH2=O
(CH3)2N
F SH +NH 3
NaCNBH3
+
I–CH2 –COO –
COO – (or I–CH2 –CONH2)
COO –
S-CH2 –COO – (or S–CH2 –CONH2) +NH 3
COO –
Figure 3.3.1 Common peptide radiolabeling reactions. (A) Iodination on tyrosine. (B) Iodination on histidine. (C) Bolton-Hunter iodination on lysine and N-terminus. (D) Acetylation on lysine and N-terminus. (E) Reductive alkylation on lysine and N-terminus. (F) Alkylation on cysteine.
In Vitro Radiolabeling of Peptides and Proteins
3.3.2 Current Protocols in Protein Science
labeling at the N-terminus of a peptide can be performed during peptide synthesis (Alternate Protocol 5). All peptide-labeling procedures described in this unit are also applicable to larger proteins; however, with larger proteins there is increased potential for idiosyncratic results because of tertiary structure and for undesired side reactions because of sensitive amino acid residues (e.g., tryptophan, methionine, and cysteine in the case of oxidative iodinations). To some extent these problems may be minimized through the use of milder labeling procedures (e.g., lactoperoxidase-catalyzed iodination). NOTE: Various methods for separating radiolabeling agents (e.g., free iodide) from labeled peptide products are described in Basic Protocol 1 and in UNITS 8.2 & 8.3, and any of these methods may be used in conjunction with any of the radiolabeling protocols in this unit. The choice of separation method should depend primarily on the properties of the peptide or protein (see Critical Parameters, discussion of iodination work-up). CAUTION: When working with 125I, 131I, 14C, 3H, or other radiochemicals, always observe appropriate safety precautions, including the use of gloves (consider use of a glove box with a charcoal filter, particularly for >5 mCi iodide), shielding, dosimetry monitoring, and proper disposal of radioactive waste. Consult a radiation safety specialist to ensure that all procedures are safe and in accordance with Nuclear Regulatory Commission guidelines. Oxidation converts iodide to a volatile and hazardous form; peptide-bound iodine is not volatile, but emits γ radiation and remains a potential hazard. Be sure to perform all steps that may give rise to volatile iodine in an externally vented hood. See APPENDIX 2B for additional guidelines. IODINATION AT TYROSINE OR HISTIDINE RESIDUES USING IODO-BEADS
BASIC PROTOCOL 1
Iodo-Beads are nonporous polystyrene beads with an immobilized oxidizing agent (N-chlorobenzenesulfonamide) capable of converting Na125I (or Na131I) to its reactive form (Markwell, 1982). Iodination occurs by electrophilic addition to tyrosine at the two positions ortho to the hydroxyl group, and on histidine at two imidazole carbon positions (Means and Feeney, 1971; Fig. 3.3.1). After reaction of a peptide with oxidized iodide, the mixture is transferred to a solid-phase extraction device (e.g., a Sep-Pak cartridge) to remove free iodide from the labeled peptide. Materials Iodo-Beads (Pierce) 0.1 M sodium phosphate, pH 6.0 or pH 8.5 Carrier-free sodium iodide-125 (Na125I; Amersham, Du Pont NEN, or ICN Biomedicals) or sodium iodide-131 (Na131I) 0.1 M sodium hydroxide Peptide, ideally dissolved in ≤0.4 ml of water or 0.1 M sodium phosphate Methanol HPLC solvent A: 0.1% (v/v) trifluoroacetic acid (TFA) in water HPLC solvent B: 0.1% (v/v) TFA in acetonitrile Filter paper 100-µl syringe with beveled tip, or hot pipettor (i.e., dedicated to radioactive use) Luer-tipped syringes (1-, 5-, and 20-ml) and one 23-G needle Solid-phase extraction device (e.g., Sep-Pak or Sep-Pak Plus C18 cartridge, Waters) Polypropylene or other plastic tubes (12 × 75 mm) Detection and Assay Methods
3.3.3 Current Protocols in Protein Science
1. Wash Iodo-Beads with 1 ml of 0.1 M sodium phosphate, pH 6.0 (for labeling primarily on tyrosine) or pH 8.5 (for labeling on histidine and tyrosine). Dry on filter paper and add to 0.5 ml of same buffer in capped microcentrifuge tube or small reaction vial. For trace labeling use two Iodo-Beads; for stoichiometric labeling more may be needed (see Support Protocol 1). Iodo-Beads are compatible with most salts (including azide), chaotropic agents, and detergents, but not with reducing agents or organic solvents that dissolve polystyrene such as dimethyl sulfoxide (DMSO) and dimethylformamide (DMF). Iodo-Beads must be stored in the presence of desiccant, and have a manufacturer-suggested shelf life of 1 year.
2. Add Na125I (or Na131I) to Iodo-Beads in the reaction tube using a 100-µl syringe with beveled tip or a hot pipettor. Cap tube and wait 5 min at room temperature, agitating the mixture occasionally. Rinse syringe with 0.1 M sodium hydroxide (125I is less volatile at high pH) and then with water before storing. The amount of radioactive iodide depends largely on the desired specific radioactivity, subject to safety and cost limitations; 1 to 5 mCi is typical. Use a fresh batch to minimize loss of radioisotope as volatile molecular iodine. Na125I is supplied in various concentrations of NaOH; the more dilute NaOH solutions are preferable since they are less likely to alter the pH of the reaction.
3. Add peptide to the reaction. Cap tube and wait 30 min, agitating occasionally. The amount of peptide is inversely related to the final specific radioactivity, unless iodinated products are separated from unlabeled peptide (see Support Protocol 2); 0.05 to 2.0 mg is typical. Reaction time can be 45 min or longer if desired, or just 5 to 10 min if oxidationsensitive residues in the peptide (tryptophan, methionine, cysteine) are of concern, and should be optimized empirically if needed.
4. During the iodination reaction (or before if more convenient), prepare a Sep-Pak C18 cartridge by attaching a syringe and passing 5 ml methanol through the cartridge, followed by 5 ml water and 10 to 20 ml HPLC solvent A. Maintain a flow rate of ∼2 ml/min, do not reverse the orientation of the cartridge when changing solvents, and avoid introducing air bubbles or allowing the cartridge to dry out. Although this Sep-Pak method offers the most efficient separation of unbound radioactive iodide from the peptide, anion-exchange (UNIT 8.2) or gel-filtration (UNIT 8.3) chromatography may be used as alternatives, especially for larger peptides at risk of denaturation under the conditions used in steps 4 to 8 (see Alternate Protocol 1, steps 1 and 4). Dialysis is another alternative for large peptides, but takes much longer.
5. Remove the mixture from the reaction tube using a 1-ml syringe fitted with a 23-G needle. Carefully remove the needle and slowly load mixture onto the preconditioned Sep-Pak cartridge, collecting the eluate into a radioactive liquid waste container. 6. Wash Sep-Pak cartridge with 20 ml HPLC solvent A, collecting the eluate as radioactive waste. Neutralize waste with 5 ml of 0.1 M sodium hydroxide. Unbound radioactive iodide will pass through the Sep-Pak cartridge under these conditions, while peptide will remain in the cartridge.
7. Elute two fractions from the Sep-Pak cartridge into 12 × 75–mm plastic tubes, the first using 4 ml of a 1:1 mixture of HPLC solvent A and HPLC solvent B, and the second using 4 ml HPLC solvent B. In Vitro Radiolabeling of Peptides and Proteins
Usually the first tube contains most of the labeled peptide, but a two-step elution improves overall recovery. Because unbound iodide is now removed, the tubes may be transferred out of the glove box or other iodination facility, as long as appropriate shielding is maintained.
3.3.4 Current Protocols in Protein Science
8. Dry the tubes in a Speedvac evaporator. Redissolve in water or desired buffer and pool the two fractions. 9. To determine the specific radioactivity of the labeled peptide, estimate peptide recovery or analyze an aliquot of the sample for peptide concentration (e.g., by a colorimetric assay) and simultaneously determine cpm (in a γ-radiation counter, possibly after serial dilutions in the presence of a carrier protein to reach a level of radioactivity within the linear counting range of the γ counter). Specific radioactivity (in cpm/ìg) = measured radioactivity (in cpm/ìl) ÷ peptide concentration (in ìg/ìl). To convert to millicuries, use 1 mCi = 2.22 × 109 dpm and cpm = dpm × counting efficiency of γ counter (often 50% to 75%, depending on the instrument). Alternatively, if iodinated products are separated from unlabeled peptide by HPLC and the molar ratio of incorporated iodide to peptide is known (or estimated), the specific radioactivity of the peptide may be calculated from that of carrier-free 125I (or 131I; see Support Protocol 2).
IODINATION AT TYROSINE OR HISTIDINE RESIDUES USING CHLORAMINE T OR IODOGEN
ALTERNATE PROTOCOL 1
Instead of using Iodo-Beads, Na125I can be oxidized to its reactive form using soluble chloramine T (N-chloro-4-methylbenzenesulfonamide sodium salt; Greenwood et al., 1963), the solid-phase reagent Iodogen (1,3,4,5-tetrachloro-3α,6α-diphenylglycoluril; Fraker and Speck, 1978), or the enzyme lactoperoxidase (see Alternate Protocol 2). Although slightly less convenient than Iodo-Beads, these methods may give higher recoveries when working with small quantities of peptide (<0.05 mg), which can adsorb to Iodo-Beads. The reagents are inexpensive and stable, and with Iodogen or lactoperoxidase, expose the peptide to relatively mild reaction conditions. Additional Materials (also see Basic Protocol 1) Anion-exchange resin, e.g., Dowex 1-X8, chloride form, 100 to 200 mesh (Bio-Rad) BSA solution: 1.0 mg/ml BSA in PBS, with 0.02% sodium azide (NaN3) 1.0 mg/ml chloramine T in 0.1 M sodium phosphate, prepared immediately before use Iodogen (Pierce), dichloromethane, and glass tubes or vials (optional alternate to chloramine T) Saturated solution of tyrosine in water (∼0.4 mg/ml at 25°C) 2.5 mg/ml sodium metabisulfite (Na2S2O5) in 0.1 M sodium phosphate, prepared immediately before use (optional alternate to tyrosine solution) Pasteur pipet or 1-ml syringe with glass wool plug inserted at bottom Gel-filtration column, e.g., Sephadex G-10 or G-25 (Pharmacia Biotech), or Excellulose Desalting Column (Pierce; optional alternate to anion-exchange column) 1. Suspend Dowex resin in water and pour into plugged Pasteur pipet. Wash with ≥20 ml BSA solution. Alternatively, equilibrate a gel-filtration column with BSA solution. Avoid introducing air bubbles into the column and do not allow column to dry out. Because short peptides (<20 residues) are difficult to separate completely from unbound radioactive iodide by gel-filtration columns, an anion-exchange column may be preferred. Highly negatively charged (e.g., phosphorylated) peptides and very short peptides (<10 residues) may be retained on an anion-exchange column and should therefore be desalted by solid-phase extraction (see Basic Protocol 1). The desalting method should be established in advance.
Detection and Assay Methods
3.3.5 Current Protocols in Protein Science
Supplement 4
2a. Chloramine T procedure: Place peptide in a small tube and buffer with 0.1 M sodium phosphate, pH 6.0 or 8.5 (see Basic Protocol 1, step 1); bring the total volume to 100 to 300 µl. Add Na125I (or Na131I), followed by 10 µl chloramine T (freshly prepared), and mix. Wait 1 min for reaction to occur. 2b. Iodogen procedure:Add sodium iodide to the peptide (as described in step 2a) in a glass tube previously coated with the solid-phase reagent Iodogen by dissolving the solid in dichloromethane and blowing off the solvent under a stream of dry nitrogen. Wait 10 min at room temperature. Iodogen-coated tubes may be stored for many months at room temperature in a desiccator, protected from light. Because the oxiding agent is water insoluble, peptide damage is kept to a minimum.
3. Add 50 µl saturated tyrosine solution to quench the reaction. Alternatively, the reaction may be terminated by adding 10 ìl sodium metabisulfite, although this reducing agent is potentially more damaging to peptides.
4. Load mixture onto a Dowex or gel-filtration column and collect 0.5-ml fractions in 12 × 75–mm plastic tubes, using BSA solution for elution. Monitor fractions with a hand-held γ counter, or count 1-µl aliquots of each fraction in a γ counter to locate the peptide-containing fractions. Radiolabeled peptide should pass through the column; unreacted iodide is retained longer and need not be eluted. Dispose of the entire column as radioactive sharps waste. ALTERNATE PROTOCOL 2
IODINATION AT TYROSINE OR HISTIDINE RESIDUES USING LACTOPEROXIDASE Although the degree of oxidative damage to peptides undergoing iodination is difficult to predict, the lactoperoxidase method results in less oxidative damage than chemical oxidative procedures (Marchalonis, 1969). Iodination is confined to tyrosine. Specific radioactivities are likely to be lower by this method. Because lactoperoxidase (mass 77,500 Da) will be iodinated during the reaction, it should be used only if the labeled peptide or protein of interest can be separated from labeled lactoperoxidase by gel filtration (UNIT 8.3), ion-exchange chromatography (UNIT 8.2), or some other means. Alternatively, it may be possible to use lactoperoxidase coupled to beads, or biotinylated lactoperoxidase and avidin-coated beads to remove the labeled enzyme (Sigma). Because the enzyme does not gain access to the cytoplasm, lactoperoxidase-catalyzed iodination is useful for selectively radiolabeling membrane proteins on whole cells, provided iodide concentrations are kept low and the proportion of dead cells is very small (Marchalonis et al., 1971). Additional Materials (also see Basic Protocol 1) 0.2 mg/ml lactoperoxidase in PBS 0.01% hydrogen peroxide (H2O2) in PBS (diluted from 30% stock immediately before use) 2 U/ml glucose oxidase in PBS and 0.1 M D-glucose in PBS (optional alternate to H2O2) 1. Place peptide in a tube with PBS, ideally at ≥1.0 mg/ml in a small volume. 2. Add 10 µl of 0.2 mg/ml lactoperoxidase.
In Vitro Radiolabeling of Peptides and Proteins
Azide should not be present in the reaction as it inhibits lactoperoxidase.
3.3.6 Supplement 4
Current Protocols in Protein Science
3. Add Na125I (or Na131I), cap tube, and mix. 4. Add 10 µl of H2O2. Wait 10 min at room temperature. The H2O2 concentration is critical and may need fine-tuning for best results. Alternatively, use a coupled glucose/glucose oxidase system to generate H2O2 in situ, i.e., treat with 10 ìl of glucose oxidase and 10 ìl of glucose for 10 min at 4°C (Hubbard and Cohn, 1975).
5. Terminate the reaction and remove unbound radioactive iodide (see Alternate Protocol 1, steps 1, 3, and 4). EQUIMOLAR IODINATION TO GIVE HIGH YIELDS OF PRODUCT Iodination of microgram to milligram amounts of a peptide using low millicurie amounts of carrier-free Na125I (or Na131I), such as those suggested in Basic Protocol 1 and Alternate Protocols 1 and 2, results in trace labeling (iodination of only a small fraction of the peptide molecules). For example, in a reaction between 1 mCi of 125I (∼600 pmol) and 500 µg of a peptide of mass 1000 Da (500 nmol) or a protein of mass 100,000 Da (5 nmol, or ∼200 nmol of tyrosine residues on average), there is at most only enough 125I to label well under 1% of the tyrosines. For proteins, trace labeling is often advantageous as it minimizes the chance that the protein’s activity will be lost (Eisen and Keston, 1949; Hunter and Greenwood, 1962); moreover, a sizable fraction of protein molecules will have one iodine atom on at least one of its tyrosines. However, for short peptides over 99% of the molecules remain unmodified, greatly diluting the specific radioactivities that are possible and complicating the interpretation of experiments in which labeled and unlabeled peptides differ significantly in their activities (see Support Protocol 2).
SUPPORT PROTOCOL 1
To label a higher proportion of a peptide, nonradioactive Na127I (“carrier”) can be added to 125I, giving a higher yield of iodinated product and resulting in a mixture of chemically equivalent 125I- and 127I-labeled peptides. Perform all steps for iodination as described using Iodo-Beads (see Basic Protocol 1) or chloramine T or Iodogen (see Alternate Protocol 1), with two modifications. First, add 0.1 M Na127I in water to the Na125I prior to using Na125I in the indicated step. The amount of 127I should be in moderate molar excess over the peptide (2- to 5-fold) to achieve high yields of iodinated products (70% to 95%). Further increasing the amount of 127I unnecessarily diminishes the products’ specific radioactivities: e.g., at a 4-fold molar excess of 127I, only up to 25% of the 125I added can be incorporated, at a 10-fold molar excess only 10% can be incorporated, etc. Second, a sufficient number of Iodo-Beads (or other oxidizing reagent) must be used to oxidize the total iodide (essentially 127I). Use the manufacturer-determined oxidizing capacity of 0.55 ± 0.05 µmol per Iodo-Bead. Note that Iodo-Beads rapidly develop a yellow-brown color when stoichiometric amounts of iodide are added; failure to observe this color indicates deterioration of the beads. For maximum specific radioactivities, label a small amount of peptide (e.g., 30 µg) with up to an equimolar amount of 125I (e.g., 40 mCi) and separate the unlabeled peptide from iodinated products by HPLC (see Support Protocol 2). It is also possible to iodinate with 127I only, in which case the products will be nonradioactive but chemically equivalent to the corresponding radiolabeled products. This approach is useful for practicing the iodination procedure and for assessing the effects of iodination on a peptide’s activity (Tsomides and Eisen, 1993), as well as for preparing an 127I-labeled peptide to be reductively tritiated (see Basic Protocol 4).
Detection and Assay Methods
3.3.7 Current Protocols in Protein Science
Supplement 3
SUPPORT PROTOCOL 2
SEPARATION OF UNLABELED PEPTIDE FROM IODINATED PRODUCTS BY HPLC Whether trace iodinated with 125I only or stoichiometrically iodinated with a mixture of 125 I and 127I, some peptide will remain unlabeled, and it may be desirable to separate this unlabeled component from the iodinated products. To obtain chemically homogeneous radiolabeled peptides, iodinated products can be resolved from each other and from unlabeled peptide by reversed-phase HPLC. (This optional procedure is distinct from the obligatory separation of peptide from unbound radioactive iodide by solid-phase extraction, anion-exchange chromatography, gel filtration, exhaustive dialysis, or other means as detailed in steps 4 to 8 of Basic Protocol 1.) Because iodine is a bulky, hydrophobic atom relative to hydrogen, the retention times of iodinated peptides will be greater than those of their unlabeled counterparts on reversed-phase HPLC columns (e.g., C18 for short peptides, C4 for longer or more hydrophobic peptides). In most iodinations, more than one product will be generated, even if the peptide has only a single tyrosine (see Fig. 3.3.1). Because the pKa of monoiodotyrosine is lower than that of tyrosine (Edelhoch, 1962), electrophilic addition of the second iodine atom may be accelerated relative to the first; however, this reaction will not go to completion even when iodide is present in molar excess over the peptide. When more than two or three iodinatable residues (tyrosine and histidine) are present (as in a protein), the labeled products may be too complex to allow for individual resolution by HPLC, but unlabeled peptide can still be removed from the radiolabeled mixture. Even peptides as long as 50 to 100 residues can be separated from their iodinated derivatives by reversed-phase HPLC. For peptides with a single iodinatable residue, the two products are usually easily separated by HPLC and can readily be identified by their specific radioactivities, which are in the ratio 1:2 for monoiodinated and diiodinated peptide, respectively. Figure 3.3.2 shows the result of stoichiometrically labeling 2.0 mg of an eight-residue peptide with 20
Figure 3.3.2 Stoichiometric iodination of the eight-residue peptide LSPFPYDL and HPLC separation of the unlabeled starting material (peak 1) from monoiodinated (peak 2) and diiodinated (peak 3) products. Two milligrams of peptide were iodinated with 20 mCi Na125I and 5 µmol Na127I at pH 7.0 using six Iodo-Beads, injected on a Vydac C4 semipreparative column, and eluted with an acetonitrile gradient of 1%/min at a flow rate of 3 ml/min. The specific radioactivities of the three peaks were <105 cpm/µg, 2.1 × 107 cpm/µg, and 4.6 × 107 cpm/µg, respectively.
cpm
1
2
3
A280
In Vitro Radiolabeling of Peptides and Proteins
30
Time (min)
60
3.3.8 Supplement 3
Current Protocols in Protein Science
mCi Na125I mixed with 5 µmol Na127I and separating the unreacted, monoiodinated, and diiodinated products by HPLC. If Edman degradation is available, labeled products may be characterized directly by detecting the monoiodinated and diiodinated derivatives of tyrosine and/or histidine (Tsomides and Eisen, 1993). Note that peptides containing tryptophan, methionine, or cysteine residues may undergo oxidation during iodination. The resulting oxidized products (both radiolabeled and unlabeled) may appear as separate peaks on HPLC, complicating the identification of products. When carrying out stoichiometric iodination with a mixture of 125I and 127I, specific radioactivities of the product mixture or of individual HPLC-purified products can be measured as described for the Iodo-Beads procedure (see Basic Protocol 1, step 9). However, when trace labeling with 125I only and removing unlabeled peptide by HPLC, specific radioactivities may simply be calculated as the specific radioactivity of carrierfree 125I (∼2000 Ci/mmol) multiplied by the molar ratio of 125I to peptide (e.g., 1 for a monoiodinated derivative, 2 for a diiodinated derivative, etc.). When a mixture of 125 I-labeled products is separated from unlabeled peptide by HPLC but not resolved into its individual components, the same calculation may be applied by taking the average molar ratio of 125I to peptide to be the number of iodinatable residues. For example, the specific radioactivity of a peptide of mass 1000 Da with two tyrosine residues that is labeled with 125I and separated from unlabeled peptide by HPLC is ∼5 × 109 cpm/µg, assuming an average of two 125I per labeled peptide. NOTE: Although HPLC fractions can be analyzed for radioactivity by transferring them to a γ counter, it is preferable to use an on-line radioisotope detector connected in series with an ultraviolet absorbance detector (see Fig. 3.3.2). IODINATION AT LYSINE RESIDUES OR N-TERMINUS USING BOLTON-HUNTER REAGENT
BASIC PROTOCOL 2
Many peptides do not contain tyrosine or histidine. Other peptides’ function may be altered if crucial tyrosine or histidine residues are iodinated or if damage to oxidativesensitive residues occurs during iodination. An alternate strategy is to label primary amino groups on lysine residues and/or at the N-terminus of the peptide using Bolton-Hunter reagent 3-(4-hydroxyphenyl)propionic acid N-hydroxysuccinimide ester (Bolton and Hunter, 1973). This reagent introduces a side chain similar to that of tyrosine (see Fig. 3.3.1). Of course, this method carries the potential drawback of altering peptide function if lysine residues (or the N-terminus) are critical for activity; unlike the methods above, peptide charge will be decreased by one unit for each site modified (at pH values below the amino group’s pKa value). Because lysine’s ε-amino group and the N-terminal α-amino group have different pKa values (∼10 vs. 8 to 9), it is possible, in principle, to achieve selective labeling by adjusting the pH (lower pH should favor the N-terminus). However, this is difficult in practice, and a mixture of products is to be expected from lysine-containing peptides unless the N-terminal residue is proline (secondary amine) or a protecting group is present on either lysine or the N-terminus. 125
I-labeled Bolton-Hunter reagent is commercially available in monoiodinated (∼2000 Ci/mmol) or diiodinated (∼4000 Ci/mmol) form. A cost-saving modification is to prepare 125 I-labeled Bolton-Hunter reagent from the less expensive unlabeled reagent by iodination using Iodo-Beads (see Basic Protocol 1) or chloramine T (see Alternate Protocol 1) followed by extraction of the product into anhydrous benzene. (In this case it is crucial to work quickly to minimize hydrolysis, and specific radioactivities will be lower unless unreacted Bolton-Hunter reagent is removed from the labeled product.) Alternatively, the
Detection and Assay Methods
3.3.9 Current Protocols in Protein Science
peptide can first be modified with unlabeled Bolton-Hunter reagent and then radioiodinated using any of the methods previously described. Because 125I-labeled Bolton-Hunter reagent suffers some degradation by competing hydrolysis during its reaction with peptide, this alternate approach gives a higher incorporation of radioactivity; however, it also results in the radiolabeling of any tyrosine or histidine residues present in the peptide, as well as exposure of the peptide to oxidative conditions. Materials 0.1 M sodium borate, pH 8.5 0.1% (w/v) gelatin in 0.1 M sodium borate, pH 8.5 125 I-labeled Bolton-Hunter reagent (Amersham, Du Pont NEN, or ICN Biomedicals); or substitute unlabeled Bolton-Hunter reagent and iodinate later Peptide, ideally ≥1 mg/ml in ≤100 µl of 0.1 M sodium borate, pH 8.5 1 M glycine in 0.1 M sodium borate, pH 8.5 Gel-filtration column, e.g., Sephadex G-10 Dry nitrogen tank and outlet tubing fitted with needle Polypropylene or other plastic tubes (12 × 75 mm) 1. Equilibrate a gel filtration column with 0.1% gelatin/0.1 M sodium borate, pH 8.5, to prevent loss of radiolabeled peptide by nonspecific adsorption. BSA is not recommended as a carrier protein in place of gelatin because albumin binds to Bolton-Hunter reagent. If gelatin is undesired in the final preparation (e.g., iodination is to be performed later), condition the column and then reequilibrate in buffer of choice. Alternatively, use a Sep-Pak cartridge instead of gel filtration to separate by-products and salts from the labeled peptide (see Basic Protocol 1, steps 4 to 8).
2. Immediately prior to use, evaporate 125I-labeled Bolton-Hunter reagent to dryness by inserting a needle through the rubber septum and admitting a gentle stream of dry nitrogen, using the manufacturer-supplied charcoal trap as a vent to contain any released volatile radioiodine. Do not splash solution onto upper parts of the vial. If using unlabeled Bolton-Hunter reagent, prepare a stock solution in anhydrous organic solvent and dilute into borate buffer at the time of reaction to achieve a 3- to 5-fold molar excess over peptide. Unlabeled Bolton-Hunter reagent is also available as a water-soluble (sulfo) derivative (at higher cost), obviating the need for organic solvent and offering an advantage for labeling cell surface proteins because the sulfo derivative is membrane impermeable.
3. Chill peptide in borate buffer to 0°C on ice. Add peptide to vial containing dried 125 I-labeled Bolton-Hunter reagent (or freshly dissolved unlabeled Bolton-Hunter reagent) and wait 15 to 30 min at 0°C, agitating the reaction occasionally. Incorporation is greatest when using high peptide concentrations. Phosphate or bicarbonate buffers, pH 8.5, may be used instead of borate, but avoid amine-containing buffers (ammonium, glycine, Tris) as well as azides and thiols. Being an N-hydroxysuccinimide ester, Bolton-Hunter reagent is sensitive to hydrolysis, with a half-life of a few hours at pH 7.0 and only ∼10 min at pH 8.5; however, its reaction with amino groups is greatly accelerated at the higher pH since more amine is in the deprotonated (active) form.
4. Add 100 µl of 1 M glycine in 0.1 M sodium borate, pH 8.5, to quench the reaction and wait 5 min, or use a corresponding amount of Tris or any other amine for quenching. In Vitro Radiolabeling of Peptides and Proteins
Longer incubations at 0°C (>4 hr) may improve yields and obviate the need for quenching by consuming all of the reagent.
3.3.10 Current Protocols in Protein Science
5. Load mixture onto gel filtration column and collect 0.5-ml fractions in 12 × 75–mm plastic tubes, using borate (or other) buffer for elution. Monitor fractions with a hand-held γ counter, or count aliquots of each fraction in a γ counter to locate the peptide-containing fractions. 6. Resolve products by HPLC if desired and determine specific radioactivities (see Support Protocol 2). 14
C OR 3H LABELING AT LYSINE RESIDUES OR N-TERMINUS BY ACETYLATION USING ANHYDRIDE
BASIC PROTOCOL 3
There are two main advantages of labeling with 14C or 3H instead of 125I. First, for peptides that are unacceptably altered as a result of iodination, different sites may be targeted for modification (i.e., lysine or cysteine), or the peptide may even be labeled without changing its structure by replacing carbon or hydrogen atoms with the corresponding radioisotope (see Basic Protocol 4 and Basic Protocol 5). Second, the half-lives of 14C and 3H are much longer than that of 125I (5760 and 12.26 years, respectively, vs. 60 days), so labeling need not be done often. However, the specific radioactivities of peptides labeled with 14C or 3H are generally correspondingly lower, and the labeling techniques are not as widely applied as radioiodination. Also, counting β emissions involves the use of a liquid scintillation cocktail and is therefore slightly more cumbersome than counting γ emissions. Acetylation with [14C]- or [3H]anhydride is a simple reaction that will modify amino groups (lysine and/or the N-terminus of a peptide; Fig. 3.3.1; Fraenkel-Conrat and Colloms, 1967). The most common side reaction is O-acetylation of tyrosine, which is reversible under mild alkali conditions (Means and Feeney, 1971). CAUTION: All procedures should be carried out in an externally vented hood. Materials Saturated sodium acetate solution Peptide, ideally ≥1 mg/ml in ≤100 µl H2O or sodium acetate [14C]- or [3H]acetic anhydride (Amersham, Du Pont NEN, ICN Biomedicals, or Sigma) Acetic anhydride, unlabeled (optional) 1. Add 1 vol of saturated sodium acetate solution to the peptide (unless already in this buffer) and cool to 0°C on ice. 2. Add [14C]- or [3H]acetic anhydride, with or without 2 molar equivalents of additional unlabeled acetic anhydride, in four equal installments at 10- to 15-min intervals. Because the anhydride is rapidly hydrolyzed, addition over several installments is the preferred method. The use of excess unlabeled anhydride increases the extent of reaction but dilutes the specific radioactivity of the products (analogous to stoichiometric vs. trace iodination; see Support Protocol 1).
3. Remove by-products by gel filtration (UNIT 8.3), dialysis (APPENDIX 3B), or solid-phase extraction (see Basic Protocol 1, steps 4 to 8), and count fractions in an appropriate liquid scintillation cocktail. Resolution of individual products may be possible by HPLC (see Support Protocol 2).
Detection and Assay Methods
3.3.11 Current Protocols in Protein Science
Supplement 2
ALTERNATE PROTOCOL 3
14
C OR 3H LABELING AT LYSINE RESIDUES OR N-TERMINUS BY REDUCTIVE ALKYLATION
An alternative two-step procedure used to label peptides is treatment with an aldehyde followed by reduction of the resulting Schiff base with sodium cyanoborohydride (Fig. 3.3.1; Tack and Wilder, 1981; Jentoft and Dearborn, 1983). Most aldehydes give the monosubstituted alkylamine product, but formaldehyde goes on to give the dimethyl product. 14C can be incorporated via the aldehyde, or 3H can be added via labeled sodium cyanoborohydride. The latter gives higher specific radioactivities, reportedly ≥50 Ci/mmol. One advantage of this approach over acetylation is that the net charge on amino groups is not substantially altered (there is only a small change in pKa value). CAUTION: All procedures should be carried out in an externally vented hood. Materials 0.2 M sodium borate, pH 9.0 Peptide, ideally ≥1 mg/ml in 0.1 to 1.0 ml of 0.2 M sodium borate, pH 9.0 [3H]sodium borohydride (Amersham, Du Pont NEN, or ICN Biomedicals) or [3H]sodium cyanoborohydride (Amersham) 0.01 M NaOH 37% (12.4 M) formaldehyde stock [14C]formaldehyde and unlabeled sodium cyanoborohydride (optional alternative for labeling with 14C instead of 3H) 1. Add an equal volume of 0.2 M sodium borate, pH 9.0, to the peptide (unless already in this buffer) and cool to 0°C on ice. 2. Dilute formaldehyde stock (12.4 M) to 0.1 to 0.2 M and add to peptide on ice to give a 2- to 6-fold molar excess over amino groups (e.g., 10 to 20 µl). More concentrated formaldehyde in this step may lead to peptide cross-linking. For 14C labeling, use [14C]formaldehyde in this step.
3. Immediately dissolve [3H]sodium borohydride (or [3H]sodium cyanoborohydride) in 0.01 M NaOH to a final concentration of 1.0 Ci/ml and add to peptide on ice for 10 min. Use a molar ratio of sodium borohydride to formaldehyde of between 0.25 and 0.40 (e.g., 30 to 50 µl). Sodium cyanoborohydride is a superior reagent for this purpose because it reduces the Schiff base formed initially between the peptide and formaldehyde without reducing disulfides or aldehydes. [3H]sodium borohydride or [3H]sodium cyanoborohydride should be stored frozen in small aliquots rather than frozen and thawed repeatedly. For 14C labeling, use unlabeled sodium cyanoborohydride in this step.
4. Remove by-products by gel filtration (UNIT 8.3), dialysis (APPENDIX 3B), or solid-phase extraction (see Basic Protocol 1, steps 4 to 8), and count fractions in an appropriate liquid scintillation cocktail. Resolution of individual products may be possible by HPLC (see Support Protocol 2).
In Vitro Radiolabeling of Peptides and Proteins
3.3.12 Supplement 2
Current Protocols in Protein Science
14
C OR 3H LABELING AT CYSTEINE RESIDUES USING IODOACETIC ACID OR IODOACETAMIDE
ALTERNATE PROTOCOL 4
To label cysteine residues, a peptide is reduced and allowed to react with either iodoacetic acid (to give the carboxymethyl derivative) or iodoacetamide (to give the carboxamidomethyl derivative, which is uncharged; see Fig. 3.3.1). For large peptides containing disulfide bonds, this procedure is often used to denature the peptide irreversibly. Carboxymethylcysteine content is readily determined by amino acid analysis following acid hydrolysis. [14C]- or [3H]iodoacetic acid and [14C]iodoacetamide are commercially available for radiolabeling. Possible side reactions are alkylation of histidine, methionine, or lysine, each of which can usually be controlled by pH and other reaction conditions (Crestfield et al., 1963; Means and Feeney, 1971). CAUTION: All procedures should be carried out in an externally vented hood. Materials 0.5 M Tris⋅Cl/6 M guanidine⋅HCl/0.002 M disodium EDTA, pH 8.6 Peptide, ideally ≥1 mg/ml in 0.1 to 1.0 ml 0.5 M Tris⋅Cl/6 M guanidine⋅HCl/0.002 M disodium EDTA, pH 8.6 Dithiothreitol 1.0 M NaOH [14C]- or [3H]iodoacetic acid or [14C]iodoacetamide (Amersham, Du Pont NEN, or ICN Biomedicals) Iodoacetic acid or iodoacetamide, unlabeled (optional) 1. Peptides containing disulfide bonds or significant tertiary structure require reduction and/or denaturation prior to alkylation of cysteine residues. For this, prepare a solution of peptide in 0.5 M Tris⋅Cl/6 M guanidine⋅HCl/0.002 M disodium EDTA (pH 8.6), incubate for 1 hr at 37°C, add dithiothreitol (50-fold molar excess over disulfide), and incubate overnight at 37°C. 2. Cool to room temperature and add desired amount of radiolabeled iodoacetic acid (or iodoacetamide) dissolved in 1⁄10 vol of 1.0 M NaOH for 30 min in the dark. Optionally, include additional unlabeled iodoacetic acid (or iodoacetamide) at a 2-fold molar excess over the dithiothreitol added to increase the extent of the reaction (analogous to stoichiometric vs. trace iodination; see Support Protocol 1).
3. Remove by-products by gel filtration (UNIT 8.3), dialysis (APPENDIX 3B), or solid-phase extraction (see Basic Protocol 1, steps 4 to 8), and count fractions in an appropriate liquid scintillation cocktail. Resolution of individual products may be possible by HPLC (see Support Protocol 2). 3
H LABELING AT TYROSINE RESIDUES BY REDUCTION OF 127I-LABELED PEPTIDE
BASIC PROTOCOL 4
It is possible to label one or more tyrosine residues of a peptide stoichiometrically with H to a relatively high specific radioactivity (e.g., 50 Ci/mmol). Since 3H substitutes for hydrogen, the peptide’s structure remains unaltered. However, this approach is more laborious and expensive than the previous methods, requiring the preparation of a 127 I-labeled peptide that is then catalytically reduced in the presence of tritium gas, leading to replacement of each 127I (or other halogen atom) with 3H. 3
The 127I-labeled peptide precursor can be produced in two ways: by stoichiometric iodination with 127I (see Basic Protocol 1, Support Protocol 1, and Support Protocol 2) or by incorporating a suitably protected 127I-labeled tyrosine derivative (either monoiodi-
Detection and Assay Methods
3.3.13 Current Protocols in Protein Science
nated or diiodinated) during de novo peptide synthesis (e.g., N-Boc-O-3-bromobenzyl3,5-diiodo-L-tyrosine, available from Peninsula). The 127I-labeled peptide is then submitted to a specialized facility for reductive tritiation (Amersham and Du Pont NEN offer this service). BASIC PROTOCOL 5
INTRODUCTION OF LABELED AMINO ACID RESIDUES DURING PEPTIDE SYNTHESIS The most specific and least disruptive labeling procedure is incorporation of a suitably protected radioactive amino acid into any desired position of a peptide during its solid-phase synthesis. However, because this requires an entire synthesis rather than use of just an aliquot of peptide, and because the protected derivatives of radioactive amino acids are not generally available and therefore must be prepared, it is not commonly done. Furthermore, introduction of a radioactive amino acid into a peptide synthesizer will be possible only if the operator of the instrument is prepared to work with radioactive samples. However, amino acids containing 14C, 3H, or 35S (for methionine) are available, procedures for attaching various protecting groups to these amino acids are quite well established, and the result may be an abundant supply of a radiolabeled peptide unperturbed in structure and not obtainable by any other means. Specific radioactivities are typically two to three orders of magnitude lower than for radioiodinated peptides. The simplest approach is to select a radioactive amino acid that requires protection during solid-phase peptide synthesis only on its α-amino group and not on its side chain (e.g., 14 C- or 3H-labeled alanine, isoleucine, leucine, valine, or phenylalanine, all available from Amersham, Du Pont NEN, ICN Biomedicals, or Sigma). If classical t-butyloxycarbonyl (t-BOC) chemistry is to be used during peptide synthesis, a variety of procedures are available for attaching t-BOC to α-amino groups (Stewart and Young, 1984); one such procedure is outlined below for leucine. Materials L-Leucine, unlabeled 14 3 L-[ C]- or L-[ H]leucine, 1 to 5 mCi Triethylamine 2-(t-butoxycarbonyloxyimino)-2-phenylacetonitrile (BOC-ON, Aldrich) Dioxane Diethyl ether, anhydrous 1.0 M HCl, 4°C Separatory funnel 1. Add 131 mg L-leucine (1 mmol) and the desired number of millicuries L-[14C]- or 3 L-[ H]leucine (supplied in water) to a 25-ml round-bottom flask. Other amino acids or their side chain protected derivatives can be substituted for leucine. Unlabeled amino acid is used because the amount of 14C- or 3H-labeled material is very small in molar terms (e.g., 2 mCi [3H]leucine is <1 ìmol); safety and cost considerations generally preclude the use of stoichiometric amounts of radiolabeled amino acid.
2. Add 210 µl triethylamine (1.5 mmol). 3. Add 271 mg BOC-ON (1.1 mmol).
In Vitro Radiolabeling of Peptides and Proteins
4. Add a mixture of 600 µl dioxane and enough water to bring the total volume of water to 600 µl; e.g., if the radiolabeled amino acid is supplied in 400 µl water, add 600 µl dioxane and 200 µl water.
3.3.14 Current Protocols in Protein Science
5. Stir with a magnetic stir bar 3 hr at room temperature. The yellow, heterogeneous reaction will become homogeneous. 6. Add 2 ml water and 2 ml diethyl ether and transfer the reaction mixture to a separatory funnel. The aqueous (bottom) layer contains the triethylammonium salt of the t-BOC-amino acid product; the ether (upper) layer contains an oxime by-product from the reaction (bright yellow).
7. Drain the aqueous layer and wash it three times with an equal volume of diethyl ether in the separatory funnel to remove unwanted oxime. 8. Add a small volume of 4°C 1.0 M HCl to bring the aqueous layer to pH 2 and precipitate the t-BOC-amino acid product. 9. Add an equal volume of diethyl ether to the aqueous layer in the separatory funnel, shake, and drain off the aqueous layer. Save the ether layer and repeat ether extraction of the aqueous layer two more times. Pool the combined ether layers into an Erlenmeyer flask. The t-BOC-amino acid product now partitions into the ether layer after being converted by HCl from a salt to the free acid. Some t-BOC-amino acids (serine, threonine) remain soluble in water and are best extracted if the aqueous layer is first saturated with NaCl before adding ether.
10. Allow the product to crystallize for several hours on ice or overnight at 4°C. First blowing off some ether with a gentle stream of dry nitrogen gas may be helpful. Wash the white, crystalline product carefully with cold diethyl ether. The t-BOC-amino acid should be checked for nonreactivity of its amino group by a ninhydrin (Sarin et al., 1981) or other analytical test prior to using it in peptide synthesis.
SELECTIVE LABELING ON N-TERMINUS DURING PEPTIDE SYNTHESIS
ALTERNATE PROTOCOL 5
The N-terminus of a peptide that contains one or more lysine amino groups can be selectively labeled after the peptide has been synthesized by solid-phase methods but before amino acid side chains (including that of lysine) have been deprotected. For example, N-terminal acetylation of a newly synthesized protected peptide can be used to label the peptide with 14C or 3H without affecting lysine residues that might be important for activity (see Basic Protocol 3). Alternatively, addition of a Bolton-Hunter group allows subsequent iodination at this position (see Basic Protocol 2); tyrosine, if present, will also be iodinated unless radiolabeled Bolton-Hunter reagent is used. Various other reagents can be used to modify a peptide at its N-terminus. For example, a dinitrophenyl (DNP) group can be attached specifically to the N-terminus of a protected resin-bound peptide using dinitrofluorobenzene (100 mol equivalents), N,N-diisopropylethylamine (20 mol equivalents), and dichloromethane as the solvent for the coupling reaction (2 hr at room temperature with shaking in the dark; T. Tsomides, unpub. observ.). Specific details of the protocols to be used for N-terminal labeling reactions depend on the particular chemistries involved in the peptide’s synthesis and cleavage. In particular, any label attached selectively to the N-terminus must be stable during the peptide side chain deprotection step (e.g., hydrogen fluoride or trifluoroacetic acid). In the example above, DNP is stable during deprotection with trifluoromethane sulfonic acid. Detection and Assay Methods
3.3.15 Current Protocols in Protein Science
Supplement 4
COMMENTARY Background Information
In Vitro Radiolabeling of Peptides and Proteins
Detection of minute quantities of a peptide or protein is often made possible by radiolabeling procedures. The most common labeling strategy, radioiodination with 125I, is relatively straightforward to perform and low in cost. Thus, a peptide with two tyrosine residues can be radiolabeled with 125I to a specific radioactivity of 5 × 109 cpm/µg (see Support Protocol 2), allowing its detection at subfemtomole levels, or several orders of magnitude lower than absorbance- or fluorescence-based detection methods (UNIT 3.1). The main drawback of radioiodination (other than the need to work safely with a potentially hazardous radioisotope) is that some peptides may be unreactive or may be damaged by the introduction of iodine atoms (each as large as a phenyl ring) or by the oxidative conditions typically employed during the labeling reaction. These problems may be more acute for a large protein with native tertiary structure and more oxidation-sensitive residues (tryptophan, methionine, and cysteine) than for a short peptide; nevertheless, even small peptides can be significantly altered in their biological activities by iodination. In most cases these problems can be obviated by modifying the radiolabeling strategy (see Table 3.3.1). Standard radioiodination methods rely on the oxidation of 125I (in the form of Na125I) to its reactive state (which can be considered “I+”) for electrophilic addition to the phenolic and imidazole side chains of the amino acids tyrosine and histidine, respectively (see Fig. 3.3.1). At a pH between 6 and 7, tyrosine labeling predominates, whereas at a higher pH there is also labeling on histidine (Wolff and Covelli, 1969; Tsomides and Eisen, 1993). Different methods of oxidizing the 125I include IodoBeads (a convenient immobilized form of chloramine T; Basic Protocol 1), soluble chloramine T, and Iodogen (a water-insoluble reagent plated onto tubes prior to use; Alternate Protocol 1). A gentler oxidative procedure uses the enzyme lactoperoxidase along with hydrogen peroxide (Alternate Protocol 2). All four methods can be expected to give reasonable specific radioactivities (lowest for lactoperoxidase), and their relative merits have been assessed for some peptides (Thean, 1990; Rizzino and Kazakoff, 1991). For peptides that lack tyrosine and histidine or that are unacceptably altered by standard
iodination procedures, an alternate strategy is to label free amino groups (lysine side chains and the peptide N-terminus, unless it is proline or blocked) using an iodinated acylating agent (e.g., Bolton-Hunter reagent; see Basic Protocol 2 and Fig. 3.3.1; Bolton and Hunter, 1973). This mild coupling reaction avoids direct contact of the peptide or protein with harsh oxidizing conditions, but the resulting labeled product is likely to have a lower specific radioactivity (because Bolton-Hunter reagent suffers competing hydrolysis during the reaction); a second drawback is the relatively high cost of this reagent. A similar though less commonly tried modification using an iodinated amidinating reagent (Wood et al., 1975) has the potential advantage that the positive charge on lysine (pKa ∼10) is retained in the final labeled product (an amidine with pKa 11.5 to 12.5). Besides labeling with 125I, it is straightforward to radiolabel peptides or proteins with 14C or 3H using various methods (see Table 3.3.1). This may be desirable if all iodination options fail, or if it is critical to obtain a radioactive product devoid of structural alterations. Usually the products of 14C or 3H labeling will have much lower specific radioactivities than the products of 125I labeling, reflecting the different disintegration rates for these radioisotopes (∼35,000 and 75 times lower for 14C and 3H, respectively, than for 125I). Also, counting β emissions requires liquid scintillants and is therefore less convenient than counting γ emissions, which requires no additives to the sample. However, the longer half-lives of 14C and 3H (5760 and 12.26 years, respectively) render the resulting products useful for much longer than 125I-labeled products (t1⁄2 ∼60 days). A selection of protocols for labeling a peptide with 14C or 3H is given in this unit: acetylation on amino groups with acetic anhydride (see Basic Protocol 3), reductive alkylation on amino groups with formaldehyde and sodium cyanoborohydride (see Alternate Protocol 3), and alkylation on cysteine with iodoacetic acid or iodoacetamide (see Alternate Protocol 4). Each of these procedures results in modification of the peptide structure, albeit on different residues than does standard radioiodination (Fig. 3.3.1). To label a peptide without altering its structure, two alternatives are available: 3H labeling by catalytic reduction of (nonradioactive) 127I-labeled tyrosine (see Basic Protocol 4), and de novo peptide synthesis using a radiolabeled (and suitably protected) amino acid
3.3.16 Supplement 4
Current Protocols in Protein Science
derivative (see Basic Protocol 5; Stewart and Young, 1984). These methods are much more laborious and costly, but may be useful when modification of a peptide’s structure is unacceptable.
Critical Parameters and Troubleshooting Amount of radioisotope The amount of radioisotope to be used is often a compromise among the desire for high specific radioactivities (curies per millimole or cpm per microgram), a practical limit to the amount of radioactivity that can be afforded and safely handled, and (particularly for a large protein) any risk of altering tertiary structure by excessive substitution or radiation damage. Commonly, 1 to 5 mCi of 125I (or 131I) is used; however, it is possible to scale up the 125I and increase specific radioactivities proportionately, because iodide is almost always limiting relative to iodinatable sites (e.g., 2 mCi 125I is only ∼1.2 nmol). Conversely, lowering the amount of peptide or protein will also improve specific radioactivities, but lowering the amount of substrate too far will cause unacceptable losses of material by nonspecific adsorption to tubes; a practical lower limit is 5 to 10 µg of peptide. Sensitive amino acid residues During oxidative iodinations, the amino acid residues at greatest risk for damage are tryptophan, methionine, and cysteine. The problem is greatest for tryptophan, as an oxidized indole ring cannot be restored by subsequent treatment with a reducing agent such as dithiothreitol. When these residues are critical for peptide function, the lactoperoxidase or Bolton-Hunter methods may be preferred for 125I labeling. Iodination work-up Several options are available for the removal of unbound radioactive iodide from a peptide once the reaction is complete, including solidphase extraction (e.g., using Sep-Pak cartridges; Basic Protocol 1), anion-exchange chromatography (UNIT 8.2), gel filtration (UNIT 8.3), and dialysis (APPENDIX 3B). The choice of desalting procedure will depend on the peptide, particularly its size and charge. For small peptides the Sep-Pak method works well, because the peptide binds to the Sep-Pak cartridge, allowing extensive washing to remove all traces of free iodide (Tsomides and Eisen, 1993). For
larger peptides and proteins, anion-exchange chromatography (e.g., Dowex column) and gel filtration are popular, as the peptide is eluted immediately while iodide is retained on the column. To ensure that the removal of unbound iodide is complete, many investigators precipitate a radiolabeled protein with trichloroacetic acid (TCA) to determine the proportion of TCA-precipitable counts. Dialysis is a timeconsuming but effective method for removing traces of iodide, provided the peptide is large enough to be retained by the dialysis membrane used. Note that for highly negatively charged peptides, there may be some retention on anion-exchange columns. For peptides of 10 to 20 amino acids, separation of labeled peptide from free iodide may be poor on an anion-exchange or gel-filtration column because the peptide’s molecular mass is so close to the exclusion limit of the column. For such peptides the Sep-Pak method may be advantageous. Once unbound radioactive iodide has been removed, the labeled peptide mixture can be used as is, or the individual products can be separated from each other as well as from the unreacted peptide that is almost always present (see Support Protocol 2). If unlabeled peptide is removed by reversed-phase HPLC, the specific radioactivity of the labeled peptide will be maximized and will equal the specific radioactivity of carrier-free 125I (∼2000 Ci/mmol) multiplied by the number of 125I atoms incorporated per peptide molecule. Poor iodination results Table 3.3.2 gives potential causes and possible remedies for poor iodination results.
Anticipated Results In radioiodinating a purified protein, it is customary to aim for incorporation of up to one 125I atom per protein molecule, i.e., ∼20 µCi/µg for a protein of molecular mass 100,000 Da. For short peptides it is rare to label all the molecules, but specific radioactivities are typically on the same order (e.g., 20 µCi/µg for a peptide of mass 1000 Da, representing only 1% of the peptide labeled and 99% unlabeled). More labeled peptide is obtained when nonradioactive 127I is added during the iodination reaction (see Support Protocol 1). For example, Figure 3.3.2 shows the results of iodinating an eight-residue peptide with 20 mCi 125I and 5 µmol 127I, followed by separating the products from unlabeled peptide by HPLC.
Detection and Assay Methods
3.3.17 Current Protocols in Protein Science
Table 3.3.2
Troubleshooting the Iodination Reaction
Problem
Potential cause
Possible remedy
Poor iodine incorporation
Peptide contains reducing agent, e.g., from synthesis/cleavage Peptide contains detergent or other component that becomes iodinated Tyrosine/histidine residues are unreactive because of tertiary structure Iodo-Beads are unreactive
HPLC-purify or desalt peptide prior to use HPLC-purify peptide
Poor peptide recovery
Solubility problem
Change solvent system or adjust pH
Failed elution from Sep-Pak
Replace with ion exchange, gel filtration, or dialysis Increase amount of peptide or add carrier protein (after iodination reaction) Use chloramine T instead of Iodo-Beads
Nonspecific losses
Labeled peptide unreactive
Tyrosine is crucial for activity
Other peptide damage
Time Considerations Each of the listed protocols is easily completed within a few hours (not including HPLC purification if chosen), and with practice can be done in even less time. The resulting radioactive peptides will, of course, decay according to the half-life of the radioisotope used (or in some cases faster, if there is radiation damage). To calculate specific radioactivity after t days, simply multiply the initial specific radioactivity by 2 ( − t ⁄t ⁄2 ), where t1⁄2 is the half-life of the radioisotope (60 days for 125I). 1
Literature Cited Bolton, A.E. and Hunter, W.M. 1973. The labelling of proteins to high specific radioactivities by conjugation to a 125I-containing acylating agent. Biochem. J. 133:529-539. Chersi, A., Trinca, M.L., and Camera, M. 1988. 14 C-labeling of synthetic peptides. J. Immunol. Methods 110:271-273. In Vitro Radiolabeling of Peptides and Proteins
Treat peptide with denaturing agent Use alternate strategy, e.g., Bolton-Hunter reagent Test for color (see Support Protocol 1); use fresh lot
Crestfield, A.M., Moore, S., and Stein, W.H. 1963. The preparation and enzymatic hydrolysis of reduced and S-carboxymethylated proteins. J. Biol. Chem. 238:622-627.
Iodinate for shorter time and/or use less label Use alternate strategy, e.g., Bolton-Hunter reagent Use alternate strategy
Edelhoch, H. 1962. The properties of thyroglobulin. VIII. The iodination of thyroglobulin. J. Biol. Chem. 237:2778-2787. Eisen, H.N. and Keston, A.S. 1949. The immunologic reactivity of bovine serum albumin labeled with trace amounts of radioactive iodine (131I). J. Immunol. 63:71-80. Fraenkel-Conrat, H. and Colloms, M. 1967. Reactivity of tobacco mosaic virus and its protein toward acetic anhydride. Biochemistry 6:27402745. Fraker, P.J. and Speck, J.C., Jr. 1978. Protein and cell membrane iodinations with a sparingly soluble chloroamide, 1,3,4,6-tetrachloro-3α,6αdiphenylglycoluril. Biochem. Biophys. Res. Commun. 80:849-857. Gahmberg, C.G. and Andersson, L.C. 1977. Selective radioactive labeling of cell surface sialoglycoproteins by periodate-tritiated borohydride. J. Biol. Chem. 252:5888-5894. Greenwood, F.C., Hunter, W.M., and Glover, J.S. 1963. The preparation of 131I-labelled human growth hormone of high specific radioactivity. Biochem. J. 89:114-123.
3.3.18 Current Protocols in Protein Science
Hubbard, A.L. and Cohn, Z.A. 1975. Externally disposed plasma membrane proteins. I. Enzymatic iodination of mouse L cells. J. Cell Biol. 64:438-460.
Thean, E.T. 1990. Comparison of specific radioactivities of human α-lactalbumin iodinated by three different methods. Anal. Biochem. 188:330-334.
Hunter, W.M. and Greenwood, F.C. 1962. Preparation of iodine-131 labelled human growth hormone of high specific activity. Nature 194:495496.
Tsomides, T.J. and Eisen, H.N. 1993. Stoichiometric labeling of peptides by iodination on tyrosyl or histidyl residues. Anal. Biochem. 210:129-135.
Jentoft, N. and Dearborn, D.G. 1983. Protein labeling by reductive alkylation. Methods Enzymol. 91:570-579.
Wood, F.T., Wu, M.M., and Gerhart, J.C. 1975. The radioactive labeling of proteins with an iodinated amidination reagent. Anal. Biochem. 69:339349.
Marchalonis, J.J. 1969. An enzymatic method for the trace iodination of immunoglobulins and other proteins. Biochem. J. 113:299-305.
Wolff, J. and Covelli, I. 1969. Factors in the iodination of histidine in proteins. Eur. J. Biochem. 9:371-377.
Marchalonis, J.J., Cone, R.E., and Santer, V. 1971. Enzymic iodination: A probe for accessible surface proteins of normal and neoplastic lymphocytes. Biochem. J. 124:921-927. Markwell, M.A.K. 1982. A new solid-state reagent to iodinate proteins: Conditions for the efficient labeling of antiserum. Anal. Biochem. 125:427432. Rizzino, A. and Kazakoff, P. 1991. Iodination of peptide growth factors: Platelet-derived growth factor and fibroblast growth factor. Methods Enzymol. 198:467-479. Sarin, V.K., Kent, S.B.H., Tam, J.P., and Merrifield, R.B. 1981. Quantitative monitoring of solidphase peptide synthesis by the ninhydrin reaction. Anal. Biochem. 117:147-157. Stewart, J.M. and Young, J.D. 1984. Solid Phase Peptide Synthesis. Pierce Chemical Co., Rockville, Ill.
Key References Means, G.E. and Feeney, R.E. 1971. Chemical Modification of Proteins. Holden-Day, Inc., San Francisco, Calif. Excellent overview of side chain chemistries, including but not limited to radiolabeling procedures. Tsomides and Eisen, 1993. See above. Describes use of Sep-Pak to remove free iodine, HPLC to separate iodinated products, Edman degradation to identify products, and pH dependence of tyrosine vs. histidine iodination using Iodo-Beads.
Contributed by Ton N.M. Schumacher and Theodore J. Tsomides Massachusetts Institute of Technology Cambridge, Massachusetts
Tack, B.F. and Wilder, R.L. 1981. Tritiation of proteins to high specific activity: Application to radioimmunoassay. Methods Enzymol. 73:138147.
Detection and Assay Methods
3.3.19 Current Protocols in Protein Science
Assays for Determination of Protein Concentration
UNIT 3.4
Accurate measurement of protein concentration is critical since the results are used in other calculations, such as determination of enzyme activity. Errors in protein concentration determination tend to amplify overall errors in these calculations. Furthermore, protein assays are often purchased as kits from commercial suppliers, which result in a poor understanding of the underlying chemistry. This unit presents a survey of the common protein assay methods and highlights their usefulness and limitations. Additionally, the chemistry underlying each assay is explained to aid in troubleshooting and assay selection. There is no single protein assay method that yields absolutely accurate results. Each method has different advantages and limitations. The primary intention of this unit is to inform users how to select the most appropriate protein assay for a specific application. The Kjeldahl method (Ballentine, 1957) and the acid digestion–ninhydrin method (Lovrien and Matulis, 1995) are no longer in general use and are not included here. This unit discusses the following protein assay methods that are commonly used in biochemical laboratories: the Lowry assay (see Basic Protocol 1), the Bradford assay (see Basic Protocol 2 and Alternate Protocol 1), the BCA assay (see Basic Protocol 3 and Alternate Protocol 2), and UV spectroscopy to determine protein concentration (see Basic Protocol 4). Support Protocol 1 discusses standard curves and data processing in detail and is a good place for a novice to start before beginning any experimentation. Also included are two precipitation strategies for dealing with buffer incompatibility. Finally, many applications of SDS-PAGE require equal loading of samples based on total protein. Typically, this requires laborious extraction in a buffer compatible with a protein assay, before adding SDS-PAGE loading buffer. Alternate Protocol 3 details a method for alkylation of excess reducing agent in SDS-PAGE loading buffer for direct analysis in the BCA assay. The protocols in this unit should produce valid results for most applications. There are also kits available for protein determination from Sigma, BioRad, and Pierce. Prior to employing an assay kit or protocol, the user is advised to consult the Strategic Planning section because it describes how to select the most appropriate assay for a particular task. The assays have different strengths and weaknesses, particularly regarding buffer compatibility. Since some investigators may have limited experience with biochemical techniques, the Commentary presents a discussion of the critical parameters for protein assays, focusing on the standard curve and a brief review of spectrophotometry. Finally, a Support Protocol provides suggestions for processing data using a spreadsheet.
STRATEGIC PLANNING Assay choice Two common applications of the protein assay are enzyme activity assays and equal protein loading of SDS-PAGE gels. The strategies and considerations for these applications are considerably different. In enzyme activity calculations, the primary concern is the accuracy of the results, whereas for equal loading of SDS-PAGE gels, precision is more important. There are many other applications for protein assays, e.g., in nutrition and pharmacology, and it is strongly recommend that the literature of the particular field Detection and Assay Methods Contributed by Bradley J.S.C. Olson and John Markwell Current Protocols in Protein Science (2007) 3.4.1-3.4.29 C 2007 by John Wiley & Sons, Inc. Copyright
3.4.1 Supplement 48
be consulted before employing one of the assays presented in this unit to ensure that these assays are applicable in the field. It is particularly important to realize that these assays are estimation procedures unless the individual assay has been standardized to total amino acid analysis (UNIT 3.2). One of the most difficult aspects of assaying protein concentration is the selection of an assay compatible with the sample. All of the assays presented in this unit detect specific properties of a protein, not the entire protein itself; thus protein and buffer composition will be the primary determinants of the appropriate assay. Figure 3.4.1 presents a flow chart of the assay selection process. If selection of a routine estimation procedure for relatively pure protein samples is the goal, highest accuracy will be achieved by standardizing the assay to a total amino acid analysis of the unknown (UNIT 3.2). All of the presented methods have different sensitivities to different proteins such as bovine serum albumin and gamma-globulin (Stoscheck, 1990).
Figure 3.4.1 Flow chart for selecting a protein assay. To use the chart, begin with step A to find the protein assays that are most compatible with the sample. Assays for Determination of Protein Concentration
3.4.2 Supplement 48
Current Protocols in Protein Science
Figure 3.4.1 (continued) Then proceed to step B to obtain the list of assays compatible with the buffer system. Compare the results from each step to find the most compatible assay for the sample. Refer to the text for the assay and Table 3.4.1 to confirm assay compatibility before using a protein assay strategy. Abbreviations: 2-ME, 2-mercaptoethanol; DTT, dithiothreitol; SDS, sodium dodecyl sulfate. Detection and Assay Methods
3.4.3 Current Protocols in Protein Science
Supplement 48
Sample composition Sample composition is critical when choosing a protein assay. For example, a protein rich in arginine residues will produce an artificially high result using a Bradford assay, whereas the same protein will most likely produce a more accurate result using either the Lowry or BCA assay. Conversely, a protein high in cysteine would produce an artificially high result using the BCA assay, but would likely produce better results with the Lowry or Bradford assay. In general, the BCA and Lowry assays perform better with complex protein mixtures. Buffer composition Buffer composition is an important consideration when selecting a protein assay. The Lowry assay is very sensitive for determining protein concentration; however, common buffer components such as ethylenediaminetetraacetic acid (EDTA) interfere with chromophore production. The BCA assay is compatible with a wide range of detergents, including sodium dodecylsulfate (SDS), but does not tolerate reducing agents such as dithiothreitol (DTT). The Bradford assay will not tolerate high concentrations of detergents but will work in the presence of reducing agents, such as DTT or 2-ME. See Table 3.4.1 for a comprehensive table listing interfering compounds and the limiting concentration of each for the different assay methods. If it is not possible to find an assay that is compatible with the buffer system, then the best strategy is to employ a precipitation step (see Support Protocols 2 and 3). After precipitation, the sample is resuspended in a buffer compatible with one of the protein assays. High-throughput adaptations The Bradford and BCA assays are easily adapted for high-throughput analysis using a microtiter plate format, as presented in Alternate Protocols 1 and 2, respectively. These protocols are carried out in flat-bottom 96-well microtiter plates and are the methods of choice when processing a large number of unknowns. If microtiter plate–based assays are performed often, it is recommended to use a multichannel repeating pipettor to increase accuracy and decrease user fatigue as samples are dispensed. A convenient approach is to set up a master plate with pre-diluted standards and unknowns that can then be dispensed onto multiple replicate plates. Another significant advantage of this format is that a larger number of standard curve points can be processed, thus increasing the overall accuracy of the results.
Assays for Determination of Protein Concentration
Selection of microtiter plate and plate reader To adapt protein assays to high-throughput analyses, it is important to select a microtiter plate reader that will accommodate the rigors of analysis. An instrument that has two features: a tungsten and mercury or deuterium lamp as a light source, permitting assays at both UV and visible wavelengths is recommended. The instrument should also utilize interchangeable filter wheels or a diffraction grating to permit tuning of the light source. If a filter wheel is used, it is important to purchase a filter wheel with a fairly narrow bandpass (typically, a bandpass range of 10 to 20 nm; a wider bandpass may result in overlap with unbound Coomassie brilliant blue G-250 dye (CBBG) in the Bradford assay or the copper reagent in the BCA assay). Interference filters are more expensive than simple glass filters, but have a narrower bandpass. For the BCA assay, it is desirable to have a short time between each sample reading. Ideally, all 96 wells in the plate should be read in ≤2 min. Any polystyrene untreated flat-bottomed microplate should be acceptable. Suppliers include BD Falcon, Corning Costar, and Whatman.
3.4.4 Supplement 48
Current Protocols in Protein Science
Table 3.4.1 Concentration Limits of Chemicals in Protein Assaysa
Substanceb
Concentration limits Enhanced copperc
BCAd
Dyee
UVf 280 nm
205 nm
Acids and bases HCl
0.1 M
0.1 M
>1 M
0.5 M
NaOH
0.1 M
0.1 M
>1 M
25 mM
PCA
<1.25%
<1%
10%
1M
TCA
<1.25%
<1%
10%
<1%
Buffers Acetate Ammonium sulfate
>28 mM
Borate
0.2 M
0.6 M
0.1 M
10 mM
20%
1M
>50%
9% >100 mM
10 mM
Citrate
2.5 mM
<1 mM
50 mM
5%
<10 mM
Glycine
2.5 mM
1M
0.1 M
1M
5 mM
HEPES
2.5 µM
100 µM
100 mM
Phosphate
250 mM
250 mM
2M
1M
50 mM
Tris
250 mM
0.1 M
2M
0.5 M
40 mM
1%
1%
10%
<0.1%
0.30%
0.1%
<20 mM
Detergents Brij 35
1%
CHAPS Deoxycholate
1% 625 µg/ml
0.25%
Digitonin
10%
Lubrol PX
1%
10%
Octylglucoside
1%
10%
SDS
1.25%
1%
0.10%
0.10%
0.10%
Triton X-100
0.25%
1%
0.10%
0.02%
<0.01%
>10%
2%
0.30%
0.1%
Triton X-100(R) Tween 20
0.10%
1%
Dithiothreitol
50 µM
<1 mM
1M
3 mM
0.1 mM
2-Mercaptoethanol
1.8 µM
<1%
1M
10 mM
<10 mM
DNA/RNA
0.2 mg
0.1 mg
0.25 mg
1 µg
DMSO
>6.2%
5%
EDTA
125 µM
10 mM
Glycerol
25%
KCl
30 mM
Reductants
Miscellaneous 20%
<10%
0.1 M
30 mM
0.2 mM
10%
100%
40%
5%
<10 mM
1M
100 mM
50 mM continued
Detection and Assay Methods
3.4.5 Current Protocols in Protein Science
Supplement 48
Table 3.4.1 Concentration Limits of Chemicals in Protein Assaysa, continued
Substanceb
Concentration limits Enhanced copperc
BCAd
Dyee
UVf 280 nm
205 nm
NaCl
1.75 M
1M
5M
>1 M
0.6 M
Sucrose
50 mM
40%
1M
2M
0.5 M
Urea
>200 mM
3M
6M
>1 M
<0.1 M
a This table is a general guide. Figures preceded by (<) or (>) symbols indicate that the tolerable limit for the chemical is unknown
but is, respectively, less than or greater than the amount shown. Blank spaces indicate that data were unavailable. Reproduced with permission from Stoscheck (1990). b Abbreviations: PCA, perchloric acid; TCA, trichloroacetic acid; CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]-1propanesulfonate; EDTA, ethylenediamine tetraacetic acid; HEPES, 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid; SDS, sodium dodecyl sulfate R, reduced; DMSO, dimethyl sulfoxide. c Figures indicate the final concentration of the chemical in the assay. d Figures indicate the concentration of the chemical in a 50-µl sample. e Figures indicate the concentration of the chemical in a 25-µl sample. f Figures indicate the concentration of the chemical that does not produce an absorbance of 0.5 over water.
BASIC PROTOCOL 1
Assays for Determination of Protein Concentration
THE LOWRY ASSAY The Lowry method (Lowry et al., 1951) relies on two different reactions. The first reaction is the formation of a copper ion complex with amide bonds, forming reduced copper in alkaline solutions. This is called a Biuret chromophore and is commonly stabilized by the addition of tartrate (Gornall et al., 1949). The second reaction is reduction of the Folin-Ciocalteu reagent (phosphomolybdate and phosphotungstate), primarily by the reduced copper-amide bond complex as well as by tyrosine and tryptophan residues. The reduced Folin-Ciocalteu reagent is blue and thus detectable with a spectrophotometer in the range of 500 to 750 nm. The Biuret reaction itself is not very sensitive. Using the Folin-Ciocalteu reagent to detect reduced copper makes the Lowry assay nearly 100 times more sensitive than the Biuret reaction alone. Several useful modifications of the original Lowry assay have been developed to increase the dynamic range of the assay over a wider protein concentration (Hartree, 1972), to make the assay less sensitive to interference by detergents (Dulley and Grieve, 1975), and to first precipitate the proteins to remove interfering contaminants (Bensadoun and Weinstein, 1976). The Lowry assay is relatively sensitive, but requires more time than other assays and is susceptible to many interfering compounds (Table 3.4.1). The following substances are known to interfere with the Lowry assay: detergents, carbohydrates, glycerol, Tricine, EDTA, Tris, potassium compounds, sulfhydryl compounds, disulfide compounds, most phenols, uric acid, guanine, xanthine, magnesium, and calcium. Many of these interfering substances are commonly used in buffers for preparing proteins or in cell extracts. This is one of the major limitations of the assay. The Lowry assay is also sensitive to variations in the content of tyrosine and tryptophan residues, a trait shared with the ultraviolet assay at 280 nm (see Basic Protool 4). The assay is linear over the range of 1 to 100 µg protein (Fig. 3.4.2). The absorbance can be read in the region of 500 to 750 nm, with 660 nm being the most commonly employed. Other wavelengths can also be used, however, and may reduce the effects of contamination (e.g., chlorophyll in plant samples interferes at 660 nm, but not at 750 nm). Also, if the A660 values are low, sensitivity can be increased by rereading the samples at 750 nm. A typical Lowry assay standard curve is depicted in Figure 3.4.2 and a typical assay spreadsheet is listed in Table 3.4.2.
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Figure 3.4.2 A sample Lowry protein assay standard curve produced using BSA at triplicate points of 0, 10, 20, 30, 40, and 50 µg. The data are fit with a linear regression by the line y = 153.06x + 0.179 with an R2 value of 0.992. The data table used to generate the figure and depiction of a typical Lowry assay is shown in Table 3.4.2.
Materials 1 mg/ml protein standard (e.g., BSA, albumin, or γ-globulin; see Support Protocol 1) Sample Lowry assay mix (see recipe) Freshly prepared diluted Folin-Ciocalteu reagent (see recipe) Test tubes (e.g., 16 × 125–mm) Spectrophotometer warmed up and set to 660 nm (or other appropriate wavelength) and cuvette 1. Prepare standards containing a range of 1 to 100 µg protein (e.g., albumin or γglobulin) to a volume of 1 ml (see Support Protocol 1 for information on standard curve preparation). 2. Add sample to water in a 16 × 125–mm test tube to yield a final volume of 1 ml. Also include two separate tubes containing water for water blanks. Additionally, use buffer blanks if necessary (see Spectrophotometry for more detail). 3. To each tube, add 5 ml of freshly prepared Lowry assay mix and thoroughly vortex. 4. Incubate tubes 10 min at room temperature. 5. Add 0.5 ml of diluted Folin-Ciocalteu reagent to each tube and vortex immediately. 6. Incubate 30 min at room temperature. Detection and Assay Methods
3.4.7 Current Protocols in Protein Science
Supplement 48
Table 3.4.2 Typical Layout of a Lowry Assay and Dataa
Standard 1 mg/ml standard (µl)
H2 O (µl)
Assay mix (ml)
Diluted Folin-Ciocalteu reagent (ml)
BSA (µg)
A660
0
1000
5
0.5
0
0
0
1000
5
0.5
0
0.009
0
1000
5
0.5
0
−0.003
10
990
5
0.5
10
0.062
10
990
5
0.5
10
0.064
10
990
5
0.5
10
0.077
20
980
5
0.5
20
0.125
20
980
5
0.5
20
0.127
20
980
5
0.5
20
0.136
30
970
5
0.5
30
0.172
30
970
5
0.5
30
0.193
30
970
5
0.5
30
0.201
40
960
5
0.5
40
0.25
40
960
5
0.5
40
0.265
40
960
5
0.5
40
0.242
50
950
5
0.5
50
0.344
50
950
5
0.5
50
0.321
50
950
5
0.5
50
0.334
A660
Protein (µg) from Eqn. 1
Equation 1: [µg protein] = (153.06 × A660 ) + 0.179 Unknowns Unknown (µl)
H2 O (µl)
Assay mix (ml)
Diluted Folin-Ciocalteu reagent (ml)
1
999
5
0.5
1
999
5
0.5
5
995
5
0.5
5
995
5
0.5
20
980
5
0.5
20
980
5
0.5
a Typical layout of a Lowry assay and data used to create the sample standard curve shown in Figures 3.4.2 and 3.4.6.
The volumes of unknown used are typical amounts required to be within the dynamic range of the assay.
7. Vortex the tubes, zero the spectrophotometer with the blank, and measure absorbance at 660 nm (or other appropriate wavelength).
Assays for Determination of Protein Concentration
The absorbance will not change significantly if all the samples are read at the same time. The Lowry assay is not an endpoint assay, so samples will change in absorbance if too much time elapses between sample readings. The typical time that elapses during the reading of samples (<10 min) does not usually result in significant changes in absorbance.
8. Refer to Support Protocol 1 for data analysis.
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THE BRADFORD ASSAY The Bradford assay (Bradford et al., 1976) is a popular protein assay because it is simple, rapid, inexpensive, and sensitive. The Bradford assay is based on the direct binding of Coomassie brilliant blue G-250 dye (CBBG) to proteins at arginine, tryptophan, tyrosine, histidine, and phenylalanine residues. The assay primarily responds to arginine residues (eight times as much as the other listed residues), therefore, if the sample is rich in arginines (e.g., histone), it may be necessary to use an arginine-rich standard as well. Anionic CBBG binds to these residues producing an absorbance maximum at 595 nm, whereas the free dye in solution has an absorbance maximum at 470 nm (Fig. 3.4.3). The assay is monitored at 595 nm in a spectrophotometer, and thus measures the CBBG complex with the protein, which results in an absorption peak shift.
BASIC PROTOCOL 2
The absorption spectra of the bound and free dye overlap and cause the Bradford assay to respond nonlinearly to protein concentration, conflicting with a common misconception that the response is linear with respect to protein concentration. However, when a standard curve is performed, a second-order curve will fit the data more accurately than a linear fit (Fig. 3.4.4). Since the data will be analyzed with software that is capable of fitting second-order curves, there is no reason not to use an appropriate curve fit to the standards. One crucial aspect of this assay is the buffer blank (see Spectrophotometry). Since the assay responds nonlinearly to protein concentration, it is important to be certain about the zero point. Because this point is so important to the curve fit, it is highly recommended that at least two buffer blanks be included. If it is determined that interference is not occurring, as is the usual case, proceed to use the water blank. The water and buffer blanks will have a relatively high absorbance, therefore, zeroing the spectrophotometer with air is recommended. This will produce an A595 of ∼0.5 for the blank. However, to zero the spectrophotometer with the blank would force the spectrophotometer to subtract
Figure 3.4.3 Absorbance spectra of the Bradford reagent and the Bradford reagent bound to 20 µg of BSA standard. The free reagent (solid line) has an absorbance peak of 470 nm whereas the Bradford reagent complexed with protein (dashed line) has an absorbance peak near 600 nm. Note that the unbound dye partially overlaps with the bound form of the reagent and thus leads to the nonlinear response of the Bradford assay.
Detection and Assay Methods
3.4.9 Current Protocols in Protein Science
Supplement 48
Figure 3.4.4 A sample micro-Bradford assay standard curve. BSA standard was added in triplicate at 0, 0.25, 1.25, 2.5, 5, 7.5, and 10 µg. Note that the assay does not respond linearly to the concentration of standard. The data is fit with the equation y = 4.5898x2 + 14.424x − 10.694 and has an R2 value of 0.9979. The data table used to generate the figure and depiction of a typical micro-Bradford assay is shown in Table 3.4.3.
a very large amount from every sample read. An absorbance of 0.5 is equivalent to only 31% of the light being transmitted by the reagent blank. This will tend to produce error and imprecision into the sample readings especially at high protein concentration. By zeroing against air, less noise occurs in the readings and A595 values stay below 2.0, a common upper limit for absorbance values (see Spectrophotometry). The choice of standard protein for this assay is important to the success of the assay. Many investigators have noted abnormalities of using various standards with the Bradford assay. Bovine serum albumin (BSA) was the original standard of choice, and is the standard that most likely comes with the assay if purchased as a kit. However, it has been noted that BSA has a significantly higher than “normal” response in the assay (e.g., Stoscheck, 1990) and may not always be suitable. Several researchers, therefore, use immunoglobulin G (IgG) or lysozyme as the preferred standard for the assay, but other standards may be appropriate depending on the particular sample protein. The CBBG dye used in the assay strongly binds to quartz cuvettes. Therefore, glass or plastic cuvettes should be utilized. It is convenient to use disposable plastic cuvettes, making cleanup much easier. Using a single cuvette for all the samples is recommended, as plastic cuvettes are not constructed with the same precision as quartz or glass cuvettes. By using a single microcuvette and maintaining its orientation in the sample holder, this eliminates one potential source error from the system. For the 1-ml micro-Bradford assay, after taking an A595 reading, the contents are aspirated and 0.1 ml of the next sample is used to wash the cuvette. The cuvette is again aspirated and the remaining 0.9 ml of the next sample is added to the cuvette for A595 determination. Assays for Determination of Protein Concentration
There are two major formats of this assay, each with a different detection range. The micro-assay format is designed for protein concentrations between 1 and 20 µg. The macro-assay format is designed for protein concentrations in the range of 20 to 100 µg.
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It is generally more convenient to use the micro-assay format because it uses less of the sample. The micro-assay format also results in less chemical waste and allows the concentrate to be used for more assays. The assay can also be carried out in a microwell plate, which is a very convenient way to process a large number of samples rapidly (see High-Throughput Adaptations). Some proteins, especially hydrophobic, membrane or “sticky” proteins tend to precipitate in the presence of CBBG and will appear as dark-blue clumps after the Bradford reagent has been added. Precipitation of the sample causes light scattering and incorrect reporting of absorbance. If there is formation of precipitate when the dye reagent is added to the sample, add a small amount of sodium hydroxide to help solubilize the protein (see steps 3a and 3b below). It is commonly stated that detergent or surfactant-containing samples are not compatible with the Bradford assay. It is common to lyse bacterial cultures in a proprietary surfactant solution such as BugBuster (Novagen). The authors have found that the amount of surfactant in extracts prepared with 100 µl BugBuster for pelleted cells from a 5-ml culture is sufficiently small that it does not interfere with the Bradford assay. Extracts containing 0.5 to 4 mg/ml protein require <10 µl of sample in a 1-ml total assay volume. A typical standard curve for the micro-Bradford assay is depicted in Figure 3.4.4 and a typical micro-Bradford assay spreadsheet is shown in Table 3.4.3.
Materials 1 mg/ml protein standard (e.g., BSA, albumin, or γ-globulin; see Support Protocol 1) Protein sample 1 M NaOH (optional) Bradford reagent (see recipe) Spectrophotometer warmed up at least 15 min before use and set to 595 nm, and cuvette Test tubes (e.g., 10 × 75–mm for the micro assay or 17 × 100–mm for the macro assay) Micro-Bradford assay 1a. Prepare standards containing a range of 1 to 20 µg protein (e.g., albumin or γglobulin) to a volume of 800 µl (to a volume of 700 µl if adding 1 M NaOH). See Support Protocol 1 for information on standard curve preparation. Table 3.4.3 contains detailed information for preparing a typical Bradford assay with the stand curve results depicted in Figure 3.4.4.
2a. Add sample to water in a final volume of 800 µl (700 µl if adding 1 M NaOH). 3a. Optional: Add 100 µl of 1 M NaOH to each sample and vortex. NaOH is added to ensure that the sample does not precipitate upon addition of Bradford reagent in step 4a. If a sample (not containing NaOH) precipitates upon addition of Bradford reagent then a new sample must be prepared since addition of NaOH after precipitation will not cause it to dissolve. NaOH should thus be added to the new sample prior to addition of Bradford reagent.
4a. Add 200 µl Bradford reagent, vortex, and incubate 5 min at room temperature. The Bradford assay is an endpoint assay, therefore, absorbance readings will not change significantly over time.
5a. Record blank, vortex samples again, and measure the absorbance at 595 nm. 6a. Refer to Support Protocol 1 for information on data processing. Current Protocols in Protein Science
Detection and Assay Methods
3.4.11 Supplement 48
Table 3.4.3 Typical Layout for a Micro-Bradford Assay and Dataa
Standard 1 mg/ml standard (µl)
H2 O (µl)
1 M NaOH (µl)
Bradford reagent (µl)
BSA (µg)
A595
0
700
100
200
0
0.603
0
700
100
200
0
0.603
0
700
100
200
0
0.613
1
699
100
200
1
0.675
1
699
100
200
1
0.664
1
699
100
200
1
0.662
5
695
100
200
5
0.9
5
695
100
200
5
0.878
5
695
100
200
5
0.883
10
690
100
200
10
1.069
10
690
100
200
10
1.062
10
690
100
200
10
1.062
15
685
100
200
15
1.254
15
685
100
200
15
1.257
15
685
100
200
15
1.279
20
680
100
200
20
1.443
20
680
100
200
20
1.444
20
680
100
200
20
1.43
25
675
100
200
25
1.651
25
675
100
200
25
1.631
25
675
100
200
25
1.638
Equation 2: [µg protein] = (4.5898 × A595 ) + (14.424 × A595 ) − 10.694 2
Unknowns Unknown (µl)
H2 O (µl)
1 M NaOH (µl)
Bradford reagent (µl)
1
699
100
200
1
699
100
200
5
695
100
200
5
695
100
200
20
680
100
200
20
680
100
200
A595
Protein (µg) from Eqn. 2
a Typical layout for a micro-Bradford assay and the data used to create the sample standard curve in Figure 3.4.4. The
volumes of unknown used are typical amounts required to be within the dynamic range of the assay.
Assays for Determination of Protein Concentration
Macro-Bradford assay 1b. Prepare standards containing a range of 20 to 200 µg protein (e.g., albumin or γglobulin) in a standard volume (generally ≤1 ml). Refer to Support Protocol 1 for information on standard curves. 2b. Dilute unknowns to an estimated concentration of 20 to 200 µg protein per tube in a same volume as the standards.
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3b. Optional: Add 0.25 ml of 1 M NaOH to each sample and vortex. NaOH is added to ensure that the sample does not precipitate upon addition of Bradford reagent in step 4a. If a sample (not containing NaOH) precipitates upon addition of Bradford reagent then a new sample must be prepared since addition of NaOH after precipitation will not cause it to dissolve. NaOH should thus be added to the new sample prior to addition of Bradford reagent.
4b. Add 5 ml Bradford reagent, vortex, and incubate 5 min at room temperature. 5b. Record blank, vortex samples again, and measure the absorbance at 595 nm. 6b. Refer to Support Protocol 1 for information on data processing.
MICROTITER PLATE BRADFORD For microtiter plate–based Bradford assays, it is recommended that a commercial reagent be considered. The authors find that, due to the small volume, any precipitates that may be in a laboratory prepared reagent will interfere with the determination. A laboratory prepared reagent must be filtered before use. The reagent should be pipetted from the top of the solution to avoid interference from precipitates that may be found at the bottom. The microtiter plate assay version is more prone to protein precipitation due to the higher concentration of dye. Even if the protein sample does not precipitate in the regular Bradford assay, it should again be verifed that the protein is not precipitating in this assay format. If protein sample precipitation is observed, add a small volume of 1 M NaOH (see step 2) to solubilize the protein.
ALTERNATE PROTOCOL 1
Materials Sample 1 mg/ml protein standard (e.g., BSA; see Support Protocol 1) Bradford reagent (see recipe) 1 M NaOH Microtiter plate (see Strategic Planning) Multichannel pipettor or repeating pipettor, optional Microtiter plate reader (see Strategic Planning) 1. Prepare unknowns and standards in a final volume of 160 µl. If protein solubility is an issue in the presence of the Bradford reagent reduce sample volume to 140 µl to accommodate addition of NaOH in step 2.
2. Optional: Add 20 µl of 1 M NaOH. NaOH is added to ensure that the sample does not precipitate upon addition of Bradford reagent in step 4a. If a sample (not containing NaOH) precipitates upon addition of Bradford reagent then a new sample must be prepared since addition of NaOH after precipitation will not cause it to dissolve. NaOH should thus be added to the new sample prior to addition of Bradford reagent.
3. Add 40 µl of Bradford reagent to each well of a microtiter plate and carefully mix. Using a multichannel pipettor facilitates mixing. If one is not available, carefully tap the side of the microtiter plate to avoid cross-contaminating wells or use a minishaker.
4. Incubate 5 min at room temperature. If this is the first time using the microtiter plate format, carefully inspect the microtiter plate wells on a lightbox for signs of precipitate formation. If protein precipitation is observed, add 20 µl of 1 M NaOH and reduce the final sample volume as indicated above.
5. Read samples at an appropriate wavelength (commonly 595 nm). 6. See Support Protocol 1 for information on data processing. Current Protocols in Protein Science
Detection and Assay Methods
3.4.13 Supplement 48
BASIC PROTOCOL 3
BCA ASSAY The BCA assay (Smith et al., 1985) measures the formation of Cu+ from Cu2+ by the Biuret complex in alkaline solutions of protein using bicinchoninic acid (BCA). Originally, the mechanism of the BCA assay was thought to be similar to the Lowry assay, but it is now known that there are two distinct reactions that take place with copper ions unique to the BCA assay (Wiechelman et al., 1988). The first reaction occurs at lower temperatures and is the result of the interaction of copper and BCA with cysteine, cystine, tryptophan, and tyrosine residues in the protein. At elevated temperatures, the peptide bond also is responsible for color development. Hence, performing the assay at 37◦ C or 60◦ C versus room temperature increases the sensitivity and reduces the variation in the response of the assay as a function of protein composition. When possible, the assay should be incubated at 60◦ C since, after the reaction is complete, the absorbance does not increase appreciably, whereas after cooling samples incubated at 37◦ C to room temperature, the blank continues to increase in absorbance at ∼2.3% every 10 min. The BCA reagent replaces the Folin-Ciocalteu reagent in the Lowry assay. The BCA reagent forms a complex with Cu+ , which has a strong absorbance at 562 nm. BCA is advantageous in that it does not interact with as many contaminants and buffer components as the Folin-Ciocalteu reagent, especially detergents. Components that interfere with the BCA assay either lead to reduction of Cu2+ (e.g., DTT) or copper chelators (e.g., EGTA). Generally, these are not critical components of buffers and can be easily removed or omitted prior to the assay. Note that there are two variations of the assay. The micro-based assay requires three reagents, whereas the test tube protocol requires only two reagents. The micro-based assay reagents are at a significantly higher concentration than the macro-based assay reagents. To prevent precipitation of BCA, it is prepared as a separate reagent in the micro-based format. It is important to use the correct reagent mixtures and ratios for the
Assays for Determination of Protein Concentration
Figure 3.4.5 A sample micro-BCA assay standard curve with triplicate BSA standard points of 0, 0.625, 3.125, 6.25, 12.5, and 18.75 µg. Note that the curve is linear across a wide range and is fit with the equation y = 10.571x − 2.0254 and has a R2 value of 0.9893. The data table used to generate the figure and depiction of a typical micro-BCA assay is shown in Table 3.4.4.
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chosen format. In Strategic Planning, a microtiter plate adaptation of the micro-based assay is described. A typical standard curve for the micro-BCA assay is depicted in Figure 3.4.5 and assay setup is listed in Table 3.4.4.
Table 3.4.4 Typical Layout of a Micro-BCA Assay and Dataa
Standard 1 mg/ml standard (µl)
H2 O (µl)
Micro-BCA solution (µl)
BSA total (µg)
A562
0
500
500
0
0.149
0
500
500
0
0.153
0
500
500
0
0.151
0.5
499.5
500
0.5
0.179
0.5
499.5
500
0.5
0.187
0.5
499.5
500
0.5
0.179
2.5
497.5
500
2.5
0.433
2.5
497.5
500
2.5
0.451
2.5
497.5
500
2.5
0.445
5
495
500
5
0.744
5
495
500
5
0.700
5
495
500
5
0.738
10
490
500
10
1.228
10
490
500
10
1.209
10
490
500
10
1.226
15
485
500
15
1.689
15
485
500
15
1.642
15
485
500
15
1.651
20
480
500
20
1.988
20
480
500
20
1.913
20
480
500
20
2.009
A562
Protein (µg) from Eqn. 3
Equation 3: [µg protein] = (10.571 × A562 ) − 2.0254 Unknowns Unknown (µl)
H2 O (µl)
Assay mix (ml)
1
499
500
1
499
500
5
495
500
5
495
500
20
480
500
20
480
500
a Typical layout of a micro-BCA assay and the data used to create the sample standard curve in Figure 3.4.5.
The volumes of unknown used are typical amounts required to be within the dynamic range of the assay.
Detection and Assay Methods
3.4.15 Current Protocols in Protein Science
Supplement 48
Materials Macro-BCA or micro-BCA assay solution (see recipes) Samples 1 mg/ml standard proteins (e.g., BSA, albumin, or γ-globulin; see Support Protocol 1) Test tubes (e.g., 10 × 75–mm for micro-BCA assay or 13 × 100–mm for macro-BCA assay) Spectrophotometer warmed up and set to 562 nm, and cuvette 60◦ C (or 37◦ C) heating block or water bath Micro-BCA assay (1-ml) 1a. Prepare fresh micro-BCA assay solution. Set up test tubes (e.g., 10 × 75–mm) containing samples and known amounts of standard protein in the range of 0.5 to 20 µg with each final sample volume of 500 µl. Refer to Support Protocol 1 for information on preparation of a standard curve. 2a. Add 500 µl of micro-BCA assay solution to each tube, vortex, and incubate the sample 15 min at 60◦ C. Alternatively, incubate 1 hr at 37◦ C, although, in practice, a 60◦ C incubation will yield more consistent results.
3a. Cool samples to room temperature. Note that this is extremely important. The simplest method is to put the test-tube rack into a pan containing room temperature water.
4a. Vortex samples and read absorbance at 562 nm. 5a. Refer to Support Protocol 1 for information on data processing.
Macro-BCA assay (2.1-ml) 1b. Prepare fresh macro-BCA assay solution. 2b. Set up 13 × 100–mm test tubes containing samples and known amounts of standard protein in the range of 0 to 100 µg in a total tube volume of 0.1 ml. 3b. Add 2.0 ml macro-BCA assay solution to each tube and vortex. 4b. Incubate tubes 15 min at 60◦ C. Alternatively, incubate tubes 30 min at 37◦ C.
5b. Cool the tubes to room temperature. 6b. Vortex samples and read the absorbance at 562 nm. 7b. Refer to Support Protocol 1 for information on data processing. ALTERNATE PROTOCOL 2
Assays for Determination of Protein Concentration
MICROTITER PLATE BCA ASSAY The microtiter plate BCA assay is based on the micro-BCA assay (see Basic Protocol 3) and utilizes the same reagents. This protocol results in a final volume of 250 µl and in the authors’ experience, works with most commercially available microtiter plates, but occasionally, some plates cannot accommodate 250 µl without well-to-well contamination. In this case, reducing the volume of reagents may help, but may result in a narrowing of the detection range. For the microtiter plate–based assay, incubating for 15 min at 60◦ C and cooling the plate to room temperature prior to reading the plate is recommended. Some microtiter plate readers can do this entire operation automatically.
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Current Protocols in Protein Science
Materials Micro-BCA working solution (see recipe) 1 mg/ml protein standard (e.g., BSA; see Support Protocol 1) Microtiter plate(s) (BD Falcon 353915; Costar 3370; Whatman 7701-1350) Microtiter plate reader, warmed up and ready to read at 562 nm 60◦ C (or 37◦ C) oven or water bath 1. Prepare sufficient micro-BCA working solution for all assays—prepare ∼15% to 20% extra to account for waste and to avoid running out. 2. Prepare standards and unknowns each in a 125-µl final volume in individual wells of the microtiter plate. 3. Add 125 µl of micro-BCA working solution to each well and gently mix by tapping the side of the microtiter plate or using a minishaker 4. Incubate 15 min at 60◦ C. Alternatively, incubate 30 min at 37◦ C, but the absorbance will continue to increase at 2.3% per 10 min)
5. Cool plate to room temperature. 6. Gently mix by tapping the side of the plate or by using a minishaker and read absorbance at 562 nm.
UV ABSORBANCE TO MEASURE PROTEIN CONCENTRATION The use of ultraviolet absorbance to measure protein concentration is perhaps the most simple and quick method, but also the most likely to produce inaccurate results. Determination of protein concentration by ultraviolet absorption, usually at 280 nm, depends on the presence of aromatic amino acids in proteins. Tyrosine and tryptophan absorb at ∼280 nm, whereas phenylalanine has a peak of absorbance at ∼260 nm. Higher orders of protein structure may also modify the molar absorptivities of tyrosine and tryptophan and thus the UV detection is sensitive to pH and ionic strength.
BASIC PROTOCOL 4
Unless the protein sample is pure and its extinction coefficient is known, UV determination of protein concentration will invariably be significantly in error. This protocol is included here because it is extremely useful for making quick and rough estimates of protein concentration before proceeding to a more accurate assay such as those listed above. A useful equation (Layne, 1957) for approximate concentration determination using UV absorbance is as follows: [protein] (mg/ml) = (1.55 × A280 ) – (0.76 × A260 ) This equation is not sufficiently accurate for routine biochemical calculations. Many other cellular components, particularly nucleic acids, also absorb UV light. Although different proteins will have different amino acid compositions and thus different molar absorptivities, this method can be very accurate when comparing different solutions of the same protein. Glass and polystyrene cuvettes absorb UV light, therefore, quartz cuvettes should be used. Methacrylate plastic cuvettes are also available, which transmit ∼60% of 280-nm light and it is possible to zero spectrophotometers with them and acquire absorbance readings. Methacrylate cuvettes are significantly less expensive than quartz cuvettes and may be a good alternative in environments where cuvette breakage is a concern (e.g., teaching laboratories).
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This method is the least sensitive of the methods presented in this unit. For increased sensitivity, the wavelength can be altered to a range of 210 to 225 nm (Stoscheck, 1990). This measures the amide bonds in proteins and is often the wavelength range used for measuring absorbance of HPLC column fractions. However, it is much more subject to interference from many more biological components and compounds used to make buffer solutions. If using a pure protein with a known extinction coefficient (ε), it is possible to directly determine the concentration from the measured A280 . The extinction coefficients for many proteins have been published (Fasman, 1989). Because of the different mole fractions of tyrosine and tryptophan residues in different proteins, their extinction coefficients can be very different. An example of the difference in extinction coefficient at 280 nm between proteins having different amino acid compositions is as follows: consider the following ε1% values (Fasman, 1989) for horse liver alcohol dehydrogenase, 4.2 to 4.5; bovine serum albumin, 6.3 to 6.8; rabbit IgG, 13.5 to 14.6; and egg white lysozyme, 24.7 to 27.2. Most protein extinction coefficients are not expressed as εmolar (M−1 cm−1 ) as is usual for most biomolecules, but rather as ε1% (%−1 cm−1 ). The ε1% is defined as the A280 value of a 1% solution (1 g/100 ml) of the protein. It is necessary to be aware of this difference when calculating the amount of protein from A280 values. Using the molar absorptivity, A280 /εmolar produces a concentration in molar terms. Using the percent absorptivity, A280 /ε1% produces a concentration in percent. When using the percent absorptivity, it is common to use the equation (A280 /ε1% ) × 10 to yield the concentration in units of mg/ml. If the amino acid sequence of the sample protein is known, it is possible to calculate the theoretical extinction coefficient (Gill and von Hippel, 1989). A procedure for determining the concentration of an unknown using UV absorbance and a standard curve is presented here. In general, the results from this approach will also not be accurate enough to use in downstream calculations.
Materials Sample Protein standard prepared in the same buffer as unknown Sample buffer Spectrophotometer and quartz or UV-transparent cuvettes 1. To perform an ultraviolet reading at 280 nm, warm up the spectrophotometer and set it to 280 nm. 2. Zero the spectrophotometer with a water blank. 3. Construct a standard curve (0 to 3 mg/ml) using a protein standard of choice in the expected concentration range at 280 nm in a quartz or UV-transparent cuvette. 4. Measure the A280 of the buffer used to prepare the sample. 5. Measure the A280 of the sample. If the A280 is >2, dilute an aliquot of the sample in the sample buffer and remeasure. 6. Calculate the protein concentration from the standard curve. 7. Refer to Support Protocol 1 for information on data processing. Assays for Determination of Protein Concentration
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STANDARD CURVES AND DATA PROCESSING Standard Curves and Standard Selection Two details must be understood about standard curves. First, they are called standard “curves” because the response of absorbance to the amount of added protein is not necessarily linear. Second, the range of protein concentrations chosen for the standard curve (e.g., 0 to 25 µg or 0 to 100 µg) will depend on the assay employed. The A280 method (see Basic Protocol 4) is the least sensitive and requires the highest amounts of protein in the standard curve, whereas the Bradford method (see Basic Protocol 2) is sensitive, but not linear over a very wide range, necessitating the use of a relatively narrow range of standard protein.
SUPPORT PROTOCOL 1
Another factor to keep in mind is that the standard curve is based on one protein (usually BSA). The mixture of proteins in the sample may not yield identical results as the standard, in terms of a signal as a function of protein concentration directly corresponding to the standard chosen. While BSA, in general, is a suitable choice as a standard, it does have limitations as a standard (see Standard Selection) and this should be kept in mind when assaying protein samples. One of the most often overlooked assay parameters is the generation of a standard curve with each assay, which is known as an internal standard curve. Internal standard curves are essential for obtaining accurate results, especially for the Lowry (see Basic Protocol 1) and BCA (see Basic Protocol 3 and Alternate Protocol 2) assays, which are nonendpoint reactions. In the case of the Bradford assay, many procedures state that it is not necessary to run internal standard curves with each assay. However, analysis of years of compiled data for various Bradford assays has shown that standard curves can exhibit significant amounts of drift, concluding that neglecting to perform an internal standard curve may yield inaccurate results. Typically, standard curves should be run at least in duplicate.
Sample dilution series In addition to having unknown samples fall within the linear response range of the standard curve, it is good practice to perform a dilution series of the unknown(s). Expect to observe a change in the response of the unknown that follows the slope of the standard curve. For example, if 5 and 10 µl of the sample produce responses equivalent to 7.5 and 15 µg of protein, respectively, confidence may be placed in the results. If the unknowns do not change when diluted, or the response curve does not follow that of the standard curve, assay selection and buffer compatibility may need to re-evaluated. Standard selection Standard selection for protein assays is critical. If the selected standard does not respond to the assay in a manner similar to the unknowns, the magnitude of error in the assay may be significant. The most common standards are BSA or purified-IgG proteins. A review by Stoscheck (1990) describes how differently these two proteins respond in all of the assays presented in this unit. While these have become the de facto standards of choice, there are experiments where these may not be the most appropriate standards. Unfortunately, the only way to ensure total accuracy is by standardizing any assay procedure to quantitative amino acid analysis (UNIT 3.2). Bovine serum albumin (BSA) is the most commonly used standard for protein assays. BSA in no way is the perfect standard, but it has become the arbitrary standard of choice. Serum albumins are “sticky” proteins to the extent that drug adsorption to them is a critical factor in pharmacokinetic calculations (Mandula et al., 2006). This means that BSA tends to bind to other proteins and molecules and, thus, it is very difficult to purchase
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Figure 3.4.6 Data for a sample Lowry protein assay standard curve produced using BSA standard at triplicate points of 0, 10, 20, 30, 40, and 50 µg. The data are identical to those in Figure 3.4.2 except that BSA (µg) is plotted on the x axis and A660 is plotted on the y axis. The data are fit with a linear regression by the line y = 0.0065x + 0.0001 with an R2 value of 0.992. Note that to use the equation to return protein concentration, the equation must be rearranged.
in a 100% pure form. These contaminants can lead to problems with any of the listed assays due to side reactions or blocking interaction sites. Protein standards can be purchased from most chemical suppliers in liquid form or they may be prepared in the laboratory. Usually, a 1 to 5 mg/ml stock solution is prepared in bulk (50 to 100 ml) to avoid variation in the concentration of standards from day to day. When the solution is prepared, do not shake it because this will lead to denaturation of the protein (observed as bubbles on the top of the solution), but rather gently swirl to dissolve the BSA in water. This solution is then dispensed into smaller aliquots (1- to 5-ml) and stored at –20◦ C. As with any solution, it must be mixed upon thawing because melting solutions form an internal concentration gradient. The best strategy for preparing a standard curve dilution series is to prepare the standard as a 1 to 5 mg/ml stock in water and prior to the assay, dilute the stock in the same buffer as the unknowns—including a buffer blank. This strategy will provide the most accurate result, because the effects of the buffer are accounted for in the standard curve. It is also possible to prepare a standard curve dilution series in water and use a buffer blank, however, the buffer used in the unknowns should be fully compatible with the assay.
Assays for Determination of Protein Concentration
Placing absorbance on the x axis during data processing Presented here is a hypothetical example that illustrates why it is more convenient to use the y axis for protein concentration. Consider the data used to generate Figure 3.4.2 and listed in Table 3.4.2. If this data is plotted with the protein (µg) data on the x axis, the standard curve has a curve fit line with the equation y = 0.0065x + 0.0001 (Fig. 3.4.6). The x values are micrograms of protein and the y values are A590 . Thus, the equation can be rewritten as follows to be more intuitive: [A590 ] = 0.0065x[µg protein] + 0.0001
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The measurement taken is the absorbance of the sample, therefore, to calculate the micrograms of protein in the unknown sample, the equation is rearranged as follows: [µg protein] = ([A590 ] – 0.0001)/0.0065 While not a difficult rearrangement, it could introduce errors into the calculation of the amount of protein by an inexperienced laboratory worker. Now consider the same data, rearranged so that the protein (µg) data is on the y axis as in Figure 3.4.2. A linear fit of this data produces the equation y = 153.06x + 0.179. The x values this time are A590 , whereas the y values are micrograms of protein. Thus, the equation becomes [µg protein] = (153.06 × [A590 ]) + 0.179 Now the equation need not be rearranged before use. The advantage of plotting standard curves with the protein amounts on the y axis is especially apparent when using a nonlinear fit as with the Bradford assay (Fig. 3.4.4). In the case where a spreadsheet is being used to calculate standard curve and experimental data, the line-fit equation can be directly copied from the graph and pasted into the spreadsheet formula, thus simplifying the entire process of analyzing data.
Considerations for Processing Data with Spreadsheet Software There are numerous spreadsheet software packages that can be used for processing data from a standard curve. Some general considerations are presented here for preparing the data using these software packages. Standard curve generation Standard curves should be produced using a scatter plot. When possible, plot each data point individually (see examples in Figs. 3.4.2, 3.4.4, and 3.4.5). This will allow the curvefitting algorithm to fit each data point and for the curve fit to reflect the real variation in the data points. As mentioned above, the y axis should be concentration values and the x axis should be absorbance values, as this will allow the curve fit to directly return a function without having to rearrange the equation. Note that many spreadsheet programs allow calculation of a curve fit with a function in the spreadsheet from the set of standard data. For experienced users, this is a convenient shortcut and allows the production of template spreadsheets for data analysis. For inexperienced protein assay users, it is recommended to produce a graphical plot with a curve fit to avoid errors in data analysis and to have a visual check that there has not been an error in entering the data. If performing the Bradford assay, be sure to use a polynomial curve fit as the Bradford assay is a nonlinear assay (see Fig. 3.4.4). The BCA and Lowry assays both perform linearly, therefore, a linear curve fit is acceptable (Figs. 3.4.2 and 3.4.5).
Unknowns within standard curve range A common error made by novices is the utilization of sample data points that do not fit within their standard curves to extrapolate results. To make valid use of a standard curve, data points must be within the linear range of the spectrophotometer (see Spectrophotometry). When unknowns are compared to a standard curve in this way, it is termed interpolation. Avoid extrapolating results in cases where no standard curve data points are in the range of the unknown to support the value that has been determined. Furthermore, the response range of the assay (which is listed with each assay) is considerably narrower than the range of the spectrophotometer. It is generally best to have at least two standard curve data points on either side of the results for the unknowns. This
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may require pilot assays to ensure the addition of the proper amounts or to properly dilute the sample into the correct range. To find the proper range, Basic Protocol 4 describes a crude, but rapid, estimation of the protein concentration by UV spectrophotometry. SUPPORT PROTOCOL 2
ACETONE PRECIPITATION OF PROTEIN Often, the easiest way to remove interfering non-proteinacious substances from protein preparations is to precipitate the protein prior to the assay. In this case, two standard curves should be performed. The first standard curve is prepared from a dilution series of standards that are precipitated under similar conditions as the unknowns, and a second standard curve is prepared without precipitation to allow estimation of the amount of sample lost during the precipitation. It is best to precipitate identical points on the standard curve dilution, however, this is time consuming. Alternatively, a few select points covering the entire range of the standard curve can be precipitated and then compared to the results of the nonprecipitated standard points. For example, in the Lowry assay described in Table 3.4.2 points at 10, 30, and 50 µg of BSA would be precipitated and compared to nonprecipitated standards in a compatible buffer. Acetone precipitation is the easiest method and is the most compatible with downstream assays.
Materials 100% acetone, −20◦ C Protein sample Protein assay–compatible buffer −20◦ C freezer 1.5-ml microcentrifuge tubes (acetone-compatible, not polycarbonate) Microcentrifuge 1. Add cold 100% acetone to the protein sample to a final concentration of 80% (v/v) (e.g., 0.2 ml diluted protein and 0.8 ml acetone) in acetone-compatible microcentrifuge tubes. 2. Vortex the sample vigorously. 3. Incubate 1 hr at −20◦ C in an explosion-proof freezer. 4. Pellet the protein by centrifuging 20 min at 15,000 × g, 4◦ C. CAUTION: Acetone is flammable. Do not scale this precipitation up to volumes >1.5 ml.
5. Carefully decant the supernatant and dispose as organic waste, being careful not to dislodge the pellet. 6. Dry the pellet 20 to 30 min at room temperature by inverting the tube onto a clean paper towel or lint-free wipe. Care should be taken not to dislodge the pellet. Do not over-dry the pellet as this may make is difficult to resuspend. 7. Resuspend the pellet in a protein assay–compatible buffer by vortexing. SUPPORT PROTOCOL 3
Assays for Determination of Protein Concentration
TCA PRECIPITATION OF PROTEIN Acetone precipitation (see Support Protocol 2) is the easiest method and is most compatible with downstream assays, however, a large protein sample may require an impractically large volume to bring the concentration of acetone up to 80%. In these cases, a second protocol utilizing trichloroacetic acid (TCA) precipitation of protein is provided. In TCA precipitation, it is critical that excess TCA be removed before assaying the protein. CAUTION: TCA is caustic and will burn the skin, therefore, proper precautions must be taken when handling this reagent.
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Materials 80% (w/v) TCA Protein sample 80% acetone, −20◦ C Protein assay–compatible buffer −20◦ C freezer 1.5-ml centrifuge tubes (acetone-compatible, not polycarbonate) Microcentrifuge 1. Wearing gloves, add TCA to the protein sample to a final concentration of 4%. 2. Incubate 30 min on ice. 3. Centrifuge 10 min at 15,000 × g, 4◦ C. 4. Decant the supernatant and dispose as organic waste, being careful not to dislodge the pellet. 5. Add an equal volume of 80% cold acetone to the pellet to wash away residual TCA. Vortex vigorously to release the pellet from the bottom of the centrifuge tube. 6. Centrifuge 10 min at 15,000 × g, 4◦ C and repeat 80% acetone wash four additional times. 7. Dry the pellet inverted on a paper towel or lint-free wipe for 30 min being careful not to dislodge or over-dry the pellet. 8. Resuspend the pellet in a protein assay–compatible buffer by vigorous vortexing.
ALKYLATION OF REDUCTANT AND BCA ASSAY OF PROTEIN SAMPLES IN SDS-PAGE LOADING BUFFER
ALTERNATE PROTOCOL 3
One of the most common problems encountered when performing polyacrylamide gel electrophoresis (PAGE; Chapter 10) and immunoblot analysis (UNITS 10.7 & 10.8) of protein extracts is how to load equal amounts of samples for semi-quantitative comparison. All too often researchers load SDS-PAGE gels based on tissue weight before homogenization, but this can lead to significant sample-to-sample variation. Also, some researchers will stain a duplicate gel and compare one or two bands between the gels and state that they are equally loaded. When investigating changes in protein composition between two samples, it is preferable to load based on total protein concentration. Many vendors now offer detergent- and reductant-compatible protein assays to quantitate samples prior to loading of gels for electrophoretic fractionation. These assays are usually more expensive than normal assay kits and either rely on a method similar to what follows, or on protein precipitation prior to an assay such as the Lowry assay (e.g., BioRad RC/DC). One critical factor to this assay is that bromophenol blue should not be added to the SDS-PAGE sample buffer as it may interfere. It is preferable to prepare SDS-PAGE loading buffer as a 6× stock without bromophenol blue and then, if desired, add bromophenol blue from a 10 mg/ml stock to the sample prior to gel loading. Once the amount of sample that is required from each unknown for equal loading is determined, it is critical that all of the samples are diluted to the same volume. If this is not done, band broadening (for high volume samples) and band narrowing (for small volume samples) during the stacking phase of electrophoretic fractionation may be observed. This assay (Hill and Straka, 1988) is a two-step procedure and takes advantage of the fact that the BCA assay is compatible with SDS up to concentrations of 5%. The 6× SDS-PAGE loading buffer described in UNIT 10.1 results in a 1% final concentration of SDS
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in the sample. However, it also contains DTT or 2-mercaptoethanol (2-ME) at greater than millimolar concentrations, which is not compatible with the assay. Thus, prior to the assay, the thiol groups of DTT (or 2-ME) must be alkylated with iodoacetamide to form an S-carboxymethyl derivative of DTT or 2-ME. Iodoacetamide does not interfere with the BCA assay, nor does alkylated DTT or 2-ME, therefore, after the iodoacetamide treatment, samples may be assayed directly in the BCA assay (see Basic Protocol 3 or Alternate Protocol 2). Finally, two standard curves should be performed, especially when starting out with this procedure. Samples for one standard curve will be suspended in SDS-PAGE sample buffer while the others will be in water and will not be treated with iodoacetamide. This will ensure that the addition of the SDS-PAGE loading buffer has not interfered with the BCA assay, and it will also confirm a negligible difference in BCA assay response in the presence of SDS-PAGE loading buffer. In the authors’ experience, little detectable difference has been found between samples prepared in SDS-PAGE sample buffer compared to those in water. It is appropriate to include both water blanks and buffer blanks in this assay. The spectrophotometer should be allowed to warm up and then zeroed with the water blank before recording the A562 of the buffer blank so it can be subtracted from the standard curve containing SDS-PAGE loading buffer.
Materials Stock protein standards 6× SDS-PAGE loading dye without bromophenol blue (UNIT 10.1; see recipe) Unknowns 20 mM iodoacetamide solution (see recipe) Additional reagents and equipment for BCA assay (see Basic Protocol 3) 1. Dilute stock protein standard to 100 µl final volume of water and 100 µl final volume of 1× SDS-PAGE loading buffer (without bromophenol blue) 2. Prepare unknowns in a final volume of 100 µl with 1× SDS-PAGE loading buffer without bromophenol blue. Make sure samples have been homogenized, heated, and centrifuged prior to assaying for total protein.
3. Add 100 µl of 20 mM iodoacetamide solution to the sample. 4. Incubate 20 min at 37◦ C. 5. Proceed to Basic Protocol 3 by adding working reagent directly to the sample. 6. See Support Protocol 1 for data analysis.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Bradford reagent To 50 ml of 95% ethanol, add 100 mg Coomassie Brilliant Blue G-250 (CBBG).
Assays for Determination of Protein Concentration
When the CBBG is partially dissolved, add 100 ml 85% (w/v) phosphoric acid and stir until CBBG is completely dissolved. Let the solution stir overnight to assure maximal dissolution. Dilute to 1 liter with water. Store up to 1 month at room temperature or 3 months at 4◦ C. continued
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Prior to use, filter enough reagent for the day’s work through Whatman no. 1 filter paper. While it is possible to make the reagent in the laboratory, due to the use of phosphoric acid, it more cost effective to purchase a commercial reagent. Commercial reagents also do not need to be filtered prior to use. Note that different brands of commercial reagent (e.g., BioRad or Sigma) have different concentrations and thus different volumes are required.
Bromophenol blue stock solution 10 mg/ml bromophenol blue in water Bromophenol blue is not very soluble in water, therefore, allow the solution to settle prior to use. Store up to 1 year at room temperature.
Folin-Ciocalteu reagent Dilute purchased reagent (e.g., Sigma F9252) with an equal volume of water to prepare the desired volume. Prepare diluted reagent fresh daily. Iodoacetamide solution 20 mM iodoacetamide (2-iodoacetamide) in 100 mM Tris·Cl, pH 8.0 Prepare fresh daily Any buffer compatible with the BCA assay may be used in lieu of Tris·Cl, pH 8.0, as long as the pH is near physiological pH.
Lowry alkaline reagent 2 g NaOH (0.1 M) 10 g Na2 CO3 2.5 g sodium tartrate (use of potassium salt will cause SDS to be insoluble) 12.5 g sodium dodecylsulfate Bring to a final volume of 500 ml with water Store in a plastic bottle up to 3 months at room temperature and check that it is optically clear before using. If the solution is cloudy or particulates are visible, prepare a new batch.
Lowry assay mix 50 ml Lowry alkaline reagent (see recipe) and 0.5 ml Lowry copper reagent (see recipe) Prepare fresh daily Lowry copper reagent 1 g CuSO4 ·5H2 O Bring to a final volume of 100 ml with water Store up to 1 year at room temperature Macro-BCA assay reagent A 1 g sodium bicinchoninate (BCA) 2 g sodium carbonate 0.16 g sodium tartrate 0.4 g NaOH 0.95 g sodium bicarbonate Adjust the pH to 11.25 with 10 M NaOH. Bring to 100 ml with distilled water Store up to 1 year at room temperature Detection and Assay Methods
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Macro-BCA assay reagent B 0.4 g CuSO4 ·5H2 O Bring to 10 ml with distilled water Store for 1 year at room temperature Macro-BCA assay solution 50:1 (v/v) macro-BCA assay reagent A/macro-BCA assay reagent B (see recipes) Make fresh daily Micro-BCA assay reagent A 8 g sodium carbonate monohydrate 1.6 g sodium tartrate Adjust the pH to 11.25 with 10 M NaOH Bring to 100 ml with distilled water Store up to 1 year at room temperature Micro-BCA assay reagent B 4 g BCA in 100 ml distilled water Store up to 1 year at room temperature Micro-BCA assay reagent C 0.4 g CuSO4 ·5 H2 O in 10 ml water. Store up to 1 year at room temperature Micro-BCA assay solution 25:25:1 (v/v/v) micro-assay reagent A/micro-assay reagent B/micro-assay reagent C (see recipes) Prepare fresh daily SDS-PAGE loading buffer without bromophenol blue, 6× 7 ml 0.5 M Tris·Cl, 0.4% SDS, pH 6.8 3.0 ml glycerol 1 g SDS 0.93 g DTT Add water to 10 ml Store in small aliquots at −70◦ C, do not re-freeze Do not add bromophenol blue until after the BCA assay has been performed.
COMMENTARY Critical Parameters and Troubleshooting
Assays for Determination of Protein Concentration
In addition to sample composition (see Strategic Planning) and proper standard curve production (see Support Protocol 1) there are several other elements for a successful protein assay. Be sure to use clean glassware since it is difficult to get accurate results using dirty glassware. For the methods described in this unit, it is recommended to use new glass tubes and discard them after use. High-quality detergents and distilled water should be used to clean glassware. In some cases, such as the Bradford assay, it may be more convenient
to use disposable plastic cuvettes, rather than those made from Pyrex or quartz, because the assay reagents may leave a residue. If a cuvette does not look clean, it should be first washed with soap and water. If the residue remains, a pipe cleaner works well for cleaning cuvettes. Bend the pipe cleaner in the middle and use the bent end to scrub the inside of the cuvette, being careful not to use the other end of the pipe cleaner as the wires will scratch the optical surfaces of the cuvette. If this is not sufficient to remove residues from cuvettes, try soaking in 50% nitric acid overnight. Also, a wash with ethanol or methanol often works well
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to remove contaminating materials. Cuvettes should be thoroughly washed with distilled water between determinations. While carrying out absorbance determinations, hold the filled cuvette up and look though it to determine that the solution is optically clear and without precipitates or floaters. Spectrophotometry Spectrophotometry used in the protein assays presented in this unit either relies on the presence of a metal (e.g., Cu2+ in the BCA and Lowry assays) or involves a system of conjugated double bonds (e.g., CBBG in the Bradford assay) to absorb photons produced by the spectrophotometer. Spectrophotometry is advantageous in that it is a nondestructive method. Because different molecules have different absorption characteristics, a particular compound in a mixture can often be singled out for observation. Most importantly, highly accurate measurements can be made in a short period of time. Lambert-Beer (or Beer-Lambert) equation The Beer-Lambert law of absorption states that the fraction of light absorbed is (1) proportional to the thickness of the absorbing solution, (2) proportional to the concentration of absorbing species, and (3) independent of the light intensity. Mathematically presented as follows: The Beer-Lambert equation: A = –log(I/I0 ) = εcl where ε is the molar extinction coefficient or molar absorptivity and usually has units of M−1 cm−1 , except as noted above (see Basic Protocol 4 introduction) for proteins based on a mass percent. If I0 = intensity of the light entering a colored solution; I = intensity of light leaving the solution; c = concentration; l = length of the light path (pathlength) through the solution; then –log(I/I0 ) is proportional to both the pathlength and the concentration. The term –log(I/I0 ) is called the absorbance (A) of the solution. Extinction coefficients The extinction coefficient is a proportionality constant that relates absorbance to the concentration and pathlength. ε = A/(cl) Absorbance is a dimensionless quantity (essentially a ratio). Pathlength is usually 1 cm. Notice that ε is a value true for only the wavelength at which it is defined. The same
molecule at different wavelengths has different molar absorptivities. Percent transmittance Many spectrophotometers are capable of measuring both absorbance, A, and percent transmittance, %T. Transmittance is defined as I/I0 and is not directly proportional to the absorbance. Absorbance is the –log of the transmittance. Values of transmittance normally vary from 0 to 1.0. Percent transmittance is defined as: %T = (I/I0 ) × 100. Percent transmittance is helpful for zeroing the spectrophotometer prior to use. With a buffer blank, the %T should read 100%, and with the light beam blocked, it should read 0%. There is a practical limit on the absorbance that can be measured in a spectrophotometer. An absorbance of 1 means that the transmittance is 0.10 and that only 10% of the light is reaching the light-measuring photodiode. An absorbance of 2 means that the transmittance is 0.01 and only 1% of the light is passing through the sample. An absorbance of 3 means a transmittance of 0.001 and 0.1% of the light is passing through the sample. Good spectrophotometers can measure absorbance values up to 3. Less sophisticated spectrophotometers that are more commonly found in laboratories can generally only measure absorbance to 2 or 2.5 before there is so little light being measured that the signal-to-noise ratio deteriorates. For the purposes of protein assays, keeping the absorbance range between 0.05 and 2.0 is recommended. Zeroing and blanks The use of the jargon “zero” and “blank” is often confusing and sometimes used interchangeably. Here, “zero” is used as a verb to mean the act of setting the absorbance of a spectrophotometer readout to a value of 0.0. Obviously, one could zero the spectrophotometer without a cuvette in the sample holder; this is not normally done and would be referred to as zeroing with air. It is much more common to zero the spectrophotometer with a cuvette in the sample holder. The cuvette used to zero the spectrophotometer is called a blank. If a cuvette containing only water is used, it is called a water blank. If a cuvette containing water instead of sample, plus the assay reagents, is used to zero the spectrophotometer, it is referred to as a reagent blank.
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If a cuvette containing the buffer components found in the sample to be measured plus the assay reagents is used, this is called a buffer blank. It is important to keep these differences in mind, since they are not equivalent and can lead to misinterpretation and different results.
easily automate both data acquisition and analysis. Typically, the high-throughput methods are the best strategies to use if the assay is compatible with the sample, as they are quick, reliable, and allow acquisition of more data points per sample.
Quantitative amino acid analysis The assays described in this unit have different sensitivities for proteins with different amino acid compositions. This is often not a significant issue for complex protein extracts, but becomes significant for purified proteins. Typically, amino acid analysis is performed by a service facility. An increasing number of companies will perform this service at a reasonable cost. Most service companies will be able to provide the precise protein concentration, and these results can be compared to results from one or all of the assays presented in this unit.
Literature Cited
Anticipated Results Tables 3.4.2 to 3.4.4 demonstrate typical results for the Lowry, Bradford, and BCA assays, respectively. These tables can also be used as assay and spreadsheet templates and are of particular value to new assay users.
Time Considerations
Assays for Determination of Protein Concentration
Basic Protocols 1 and 3 (the Lowry and BCA assays) can easily be completed and data collected in 1 to 1.5 hr. The Bradford assay (see Basic Protocol 2) typically can be completed in <1 hr. UV absorbance (see Basic Protocol 1) is one of the quickest methods and data can be acquired in <30 min. Users unfamiliar with data analysis of protein assay data can anticipate requiring ∼30 min to 1 hr to analyze data. Once familiar with the data analysis process, it is typically completed in 10 to 20 min, especially if using the spreadsheet templates in Tables 3.4.2 to 3.4.4. If sample preparation such as precipitation is required prior to assay, this will typically add 1 hr to analysis time. The high-throughput adaptations can be completed in a similar amount of time as the regular assays for approximately five microtiter plates. Additional microtiter plates only add a few minutes to analysis time. These times can be significantly shortened and the number of samples scaled up by the utilization of multichannel and repeating pipettors. For large-scale operations, these assays will scale nicely with robotics. Many microtiter plate readers also come with software that can
Ballentine, R. 1957. Determination of total nitrogen and ammonia. Methods Enzymol. 3:984-995. Bensadoun, A. and Weinstein, D. 1976. Assay of proteins in the presence of interfering materials. Anal. Biochem. 70:241-250. Bradford, M.M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72:248-254. Dulley, J.R. and Grieve, P.A. 1975. A simple technique for eliminating interference by detergents in the Lowry method of protein determination. Anal. Biochem. 64:136-141. Fasman, G.D. 1989. Practical Handbook of Biochemistry and Molecular Biology. CRC Press, Boca Raton, Florida. Gill, S.C. and von Hippel, P.H. 1989. Calculation of protein extinction coefficients from amino acid sequence data. Anal. Biochem. 182:319-326. Gornall, A.G., Bardawill, C.S., and David, M.M. 1949. Determination of serum proteins by means of the biuret method. J. Biol. Chem. 177:751766. Hartree, E.E. 1972. Determination of protein: A modification of the Lowry method that gives a linear photometric response. Anal. Biochem. 48:422-427. Hill, H.D. and Straka, J.G. 1988. Protein determination using bicinchoninic acid in the presence of sulfhydryl reagents. Anal. Biochem. 170:203208. Layne, E. 1957. Spectrophotometric and turbidimetric methods for measuring proteins. Methods Enzymol. 3:447-455. Lovrien, R. and Matulis, D. 1995. Assays for total protein. Curr. Protoc. Protein Sci. 3.4.4-3.4.24. Lowry, O.H., Rosebrough, N.J., Farr, A.L., and Randall, R.J. 1951. Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193:265-275. Mandula, H., Parepally, J.M., Feng, R., and Smith, O.R. 2006. Role of site-specific binding to plasma albumin in drug availability to brain. J. Pharmacol. Exp. Ther. 317:667-675. Smith, P.K., Krohn, R.I., Hermanson, G.T., Mallia, A.K., Gartner, F.H., Provenzano, M.D., Fujimoto, E.K., Goeke, N.M., Olson, B.J., and Klenk, D.C. 1985. Measurement of protein using bicinchoninic acid. Anal. Biochem. 150:7685. Stoscheck, C.M. 1990. Quantitation of protein. Methods Enzymol. 182:50-68.
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Wiechelman, K.J., Braun, R.D., and Fitzpatrick, J.D. 1988. Investigation of the bicinchoninic acid protein assay: Identification of the groups responsible for color formation. Anal. Biochem. 175:231-237.
Contributed by Bradley J.S.C. Olson Michigan State University East Lansing, Michigan John Markwell University of Nebraska Lincoln, Nebraska
Detection and Assay Methods
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Kinetic Assay Methods The design and use of a particular enzyme assay is dependent on the objectives of the particular investigation. If the focus is on enzyme purification or issues of clinical biochemistry, experimental protocols typically require optimizing conditions for all substrates and effectors—e.g., with respect to pH value, temperature, buffering agent, and ionic strength. However, optimal conditions may not be reasonable for all parameters and factors. For example, substrate instability may preclude studies at the enzyme’s optimal pH or at the physiological temperature. Thus, Silverstein and Sulebele (1969) examined the exchange rates of malate dehydrogenase at 1°C to avoid problems arising from the decomposition of oxaloacetate. Likewise, in clinical studies and enzyme purification, saturating concentrations of substrate(s) are typically required, although this may not be possible or even desirable in many instances, as limitations in substrate solubility or substrate inhibition of the enzyme may discourage the use of elevated concentrations. The focus in clinical studies and enzyme purification is on providing good estimates of the activities of a specific enzyme in a number of physiological samples with a rapid, straightforward, and reproducible assay method. However, for the kineticist the objective is to characterize the initial rates of the enzyme-catalyzed reaction under a wide set of conditions. It is through these detailed studies, often carried out at subsaturating substrate concentrations, that the investigator is able to determine the true optimal parameters to be used in protein purification and clinical studies, as well as to probe the dynamics and mechanism of the reaction and its modulation by other factors. The purpose of this unit is to provide a brief review of issues important in the design of initial-rate assay methods. In most instances, the investigator already has some information about the kinetic properties of the enzyme, obtained from preliminary experiments as well as from the literature. It is necessary to have an approximation of the magnitude of the Michaelis constants of all the substrates in order that detailed initial rate studies can be effectively designed. Certainly, information on the identity of the substrates and products is required, along with an effective procedure for monitoring substrate depletion or product accumulation. The nature of the reaction and the properties of the substrates help Contributed by R. Donald Allison Current Protocols in Protein Science (1996) 3.5.1-3.5.11 Copyright © 2000 by John Wiley & Sons, Inc.
UNIT 3.5 dictate the assay procedure used. Enzyme-assay methods include those based on spectrophotometry, radiometry, pH-stat, turbidometry, polarography, and fluorometry. Spectrophotometric and fluorometric assay protocols rely on the differences in spectral or fluorescent properties, respectively, of the product(s) relative to the substrate(s). With radiometric procedures, effective means for separating labeled substrate(s) from labeled product(s) must be available so that the amount of product formed at a given time can be accurately measured. Those reactions that utilize or generate protons (or hydrons) can be followed by pH-stat methods. If the reaction studied results in a change in solution turbidity (e.g., caused by aggregation of a macromolecular complex), then turbidometric protocols can effectively assay for the protein that causes the change in turbidity. All of these methods, as well as others, can be found in Bergmeyer (1983).
GENERAL ASPECTS OF ASSAY DESIGN An enzyme’s progress curve (i.e., a plot of product formation or substrate depletion as a function of time; see Fig. 3.5.1) consists of four phases that are of interest to the investigator. The duration of the pre-steady-state region is relatively short and is typically followed via rapid-flow or relaxation techniques. Discussion of this region of the progress curve is beyond the scope of this unit, but a review of these techniques has recently appeared (Fierke and Hammes, 1995). The next portion of the curve is the steady-state phase, a longer phase characterized by relatively constant concentrations of each of the individual enzyme forms (e.g., the enzyme-substrate binary complexes). During the third phase of the progress curve— the post-steady-state region—the levels of these enzyme complexes change rapidly, until the last phase—equilibrium—has been reached. Each of these phases contains information of use to the investigator. The steadystate portion of the curve typically exists for a few to several hundred seconds. It is this region of the curve that is most accessible to the majority of investigators. In addition, the relatively straightforward mathematical treatment of steady-state kinetics provides an excellent foundation for the subsequent characterization of the enzyme system. Because enzyme rate expressions can be quite complex, a number of
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Figure 3.5.1 Typical progress curve for an enzyme-catalyzed reaction, where a represents the pre-steady-state region, b the steady-state region, c the post-steady-state region, and d the equilibrium region.
assumptions are made to linearize the equations. The investigator should ensure that each of these factors are addressed when any assay protocol is being designed.
Maintaining Constant Reaction Conditions The status of each experimental parameter—e.g., temperature, ionic strength, and pH—should remain constant over the time course of the assay.
Kinetic Assay Methods
Temperature Prior to initiation of the reaction by addition of a small aliquot of enzyme or substrate, the reaction mixture should be thermally equilibrated with an accurately controlled constanttemperature water bath for several minutes. If the substrate is thermally stable, this incubation should be for ≥10 min or until the desired reaction temperature is reached. If the substrate is unstable at this temperature, it may be necessary either to assay at a different temperature, to correct for substrate decay with time, or to initiate the reaction by addition of a small aliquot of substrate solution that was freshly prepared and maintained in an ice bath. When the reaction is initiated by addition of either the enzyme or the substrate, the volume of the aliquot should be small enough to minimize any alterations in temperature upon addition. Typi-
cally, there is at least a hundred-fold difference in volume between the reaction mixture and the initiating aliquot. If the enzyme and substrate are stable under the assay conditions, then both solutions should be preincubated at this temperature. It should be emphasized that valid initial rate experiments are only obtained when good temperature control is maintained. The temperature-regulated water bath should have a variability of ≤0.1°C. When the reaction is initiated, the solution should be thoroughly mixed. If reaction progress is followed using an instrument such as a spectrophotometer, the sample compartment should be thermally isolated. Another issue of concern is choice of the temperature at which enzyme activity will be measured. In 1964, the International Union of Biochemistry (IUB) recommended a temperature of 30°C. Clinical chemists have often suggested a change to 37°C. However, the final choice of temperature will be dependent on the system being investigated. For example, the activity of glutamate dehydrogenase isolated from the hyperthermophile Pyrococcus furiosus is negligible at both 30°C and 37°C; the enzyme exhibits maximal activity at 97°C (Klump et al., 1992). Clearly, at some point in the investigation, the effect of temperature on the catalytic activity will have to be characterized.
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Buffer selection In setting up the assay protocol, care should also be exercised in selecting an appropriate buffer agent. Unfortunately, many investigators choose a buffer solely on the basis of the desired pH of the reaction medium. Many excellent buffers, however, have a significant oxidationreduction potential that can severely alter an enzyme’s activity. For example, cacodylate (pKa = 6.27) and other organoarsenicals have a potent oxidizing potential under acidic conditions. Borate (pKa = 9.23) and other boratebased buffers form complexes with many diols, polyols, carbohydrates, and ribonucleotides; for example, in the presence of borate ion, both L- and D-serine form a transition complex and strongly inhibit γ-glutamyl transpeptidase (Tate and Meister, 1978). Hence, care should be exercised in selecting a buffer. In determining which buffer is optimal for a given enzyme system, it is best to examine the enzyme’s activity with as many buffers as possible. Buffers in whose presence the enzyme exhibits high activity should then be examined to determine if the buffering agent interacts with the protein. If pH and ionic strength are held constant, the enzyme activity should be independent of the buffer concentration. Another factor of importance in selecting the appropriate buffer is the capacity of the buffering agent to bind metal ions. Ideally, a good buffer will not form any significant complexes with metal-ion cofactors. Metal-ion concentrations The role of metal ions is crucial with many enzymes, particularly with systems that require nucleotides—e.g., phosphotransferases. Care must be exercised in designing initial-rate assays to ensure that the concentration of free metal ions does not vary significantly with substrate concentration. Thus, when varying the concentrations of substrates such as ATP, the concentration of free, uncomplexed Mg2+ should remain relatively constant. One of two methods is commonly utilized to achieve this end: either (1) maintaining the total nucleotide concentration and the metal concentration at a constant stoichiometric ratio (e.g., 10:1) or (2) maintaining a constant excess of metal ion over the total nucleotide concentration (e.g., 1.0 mM free Mg2+). The second method, which has been demonstrated to be the preferable of the two (O’Sullivan and Smithers, 1979), requires knowledge of the stability constant for the metal-ion–nucleotide complex under the conditions of the assay.
Substrate Stability and Purity It is crucial to establish the degree of stability of the substrate as well as that of any enzyme effector under the assay conditions (also see discussion of Maintaining Constant Reaction Conditions). Substrate and effector stabilities can be tested by preparing a solution of the compound at a concentration that will be used experimentally with the other components of the assay mixture, but without the enzyme. Then, samples are removed at different periods of time, usually over a period of several hours, and assayed either chemically or enzymatically (Bergmeyer, 1983). A plot of concentration as a function of time will reveal if the substrate is stable or if there is a decay. When present, this decomposition is usually a first-order process and can be described by the equation [S]t = [S]0 e−kt, where [S]t and [S]0 are the substrate concentrations at time t and time 0, respectively, and k is the first-order rate constant. If the decomposition is greater than first-order or composed of more than one first-order process, then the decay must be evaluated by the appropriate expression, which can be found in any basic kinetics text. An issue related to substrate stability is substrate purity. Impurities in the substrate, effectors, or enzyme preparation can be potential sources of experimental error. Because alternative substrates and competitive inhibitors are often structurally similar to the substrate and frequently have similar chemical and physical properties, it is not surprising to find these compounds as impurities in commercial substrate preparations. Spectral and chromatographic analyses, as well as enzymatic analysis, are crucial in establishing the purity of a particular biochemical substance. If impurities are present in a reaction mixture, a kinetic analysis will give wrong estimates of the kinetic parameters (Allison and Purich, 1979). Heavymetal ions are often found as impurities in commercial substrates. To remove these, it may be necessary to recrystallize the substrates, possibly in the presence of a chelating agent such as EDTA. These issues of purity and stability are also pertinent for any modulators of enzyme activity, including the product(s). The products have to be stable over the time course of the assay. In some instances, the precise nature of the true product of the enzyme-catalyzed reaction may not be known. In these cases, the investigator may follow the reaction via substrate depletion. It is frequently useful to do a kinetic analysis of freshly prepared substrate in the absence and
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presence of enzymatically prepared product. A decent understanding of the possible presence of product inhibition in initial-rate experiments is always useful to the protein chemist.
Enzyme Stability and Purity
Kinetic Assay Methods
Enzymes often undergo a slow inactivation. Usually, the rate of this inactivation is accelerated by dilution of the stock enzyme preparation. Efforts have to be made to eliminate or lessen the loss of enzyme activity with time. Often, multisubstrate enzymes are stabilized by the presence of a substrate. For example, tubulin:tyrosine ligase is stabilized by the presence of its protein substrate (Deans et al., 1992) and the presence of Mg2+–ATP is known to stabilize a number of phosphotransferases. If the enzyme inactivation can be retarded by the presence of a substrate, the investigator should note the amount of that substrate that is present with the enzyme so that proper corrections can be made when the reaction is initiated with the full assay mixture. Other agents—e.g., glycerol, certain salts, high-ionic-strength solutions, and EDTA— have also been used to stabilize proteins. Often, inactivation is a result of surface denaturation or adsorption of the enzyme on glass surfaces following dilution. The presence of stabilizing agents and/or use of plastic containers can lessen these effects significantly. The presence of serum albumin or another protein will frequently minimize the effects of enzyme dilution; the investigator should test several proteins when this technique is used. The stabilizing protein may also bind the substrates or the enzyme’s cofactors; if this is the case, corrections have to be made. If a stabilizing agent cannot be identified, the inactivation should be minimized by using the smallest dilution possible. If practical, the reaction may be initiated by the addition of one of the substrates. Corrections for loss of enzyme activity can also be made by establishing an activity-decay curve. However, it should be verified that the loss of activity is due to total inactivation and not simply to a conformational change to a form having less activity (i.e., a form with different kinetic parameters). This possibility could be tested by kinetic assays at significantly different times following dilution or by two reference assays, one using a saturating substrate level and the other using a subsaturating substrate level. The presence of thiol reagents—e.g., 2-mercaptoethanol, dithiothreitol, or dithioerythritol—is routinely helpful in maintaining activity
for those enzymes that require reduced thiol groups for activity. Care must be exercised that the reducing agent does not produce other effects. For example, γ-glutamylcysteine synthetase is inhibited by the presence of thiol reagents even though cysteine is a substrate (Seelig and Meister, 1985). Enzyme purity is also an issue of concern. It is important in early initial-rate studies to eliminate contaminating activities and to demonstrate that any minor activity does not substantially affect the reaction being studied. For multisubstrate enzymes, this can be readily accomplished by observing the stability of each substrate, effector, and product separately in the presence of the enzyme preparation. Adenylate kinase is a common contaminant in preparations of a wide variety of phosphotransferases and other nucleotide-dependent enzymes. This contaminating enzyme can be inhibited by performing the assay at high ionic strength (Bowen and Kerwin, 1955). Addition of elevated levels of AMP will also inhibit adenylate kinase. However, many phosphotransferases are also inhibited by this mononucleotide or are themselves inhibited by high ionic strength. Purich and Fromm (1972) and Lienhard and Secemski (1973) have shown that the multisubstrate analogs P1,P4-di(adenosine-5′)tetraphosphate and P1,P5-di(adenosine-5′)pentaphosphate are potent and specific inhibitors of this activity. In some instances it may be impossible to completely eliminate side reactions. Enzymes that have branched reaction pathways (i.e., multifunctional enzymes) will exhibit two pathways. Most multifunctional enzymes are transferases having multiple acceptor specificity. Glucose-6-phosphatase catalyzes not only the hydrolysis of glucose-6-phosphate but also its synthesis; thus, it may effect the transfer of phosphoryl groups from pyrophosphate, carbamoyl phosphate, or other glucose-6-phosphate molecules (Nordlie, 1982). This synthetic role of glucose-6-phosphatase is not a minor activity; at a concentration of 100 mM D-glucose, the synthesis and hydrolysis activities are approximately equal. The enzyme γglutamyl transpeptidase catalyzes the hydrolysis of a number of γ-glutamyl-containing molecules (including glutathione and glutathione disulfide) and also exhibits a transferase activity in which the glutamyl moeity is transferred to any of a number of amino acid acceptors, particularly cysteine, glutamine, and methionine (Allison, 1985). The hydrolase and transferase activities of γ-glutamyl transpeptidase
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are approximately equal in the presence of a pool of acceptor substrates at their physiological concentrations (Allison and Meister, 1981). Asparagine synthetase catalyzes the glutamine-dependent synthesis of L-asparagine and also exhibits a significant glutaminase activity (Boehlein et al., 1994). In all of these cases, it is possible to use inactivation studies to observe the parallel loss of both (or all) activities with time.
Enzyme Concentration The duration of the steady-state phase of an enzyme-catalyzed reaction is dependent on a number of factors. Perhaps the most important of these is the relative concentration of the enzyme and substrate. The ratio of the initial substrate concentration to the total enzyme concentration should be ≥100:1. With lower values, the steady-state phase will be of very short duration. It should be pointed out that this relationship holds for enzyme effectors as well. Thus, when investigating the influences of an activator or competitive inhibitor on enzyme activity, the total enzyme concentration should be much less than the concentration of the effector. This may prove difficult if the effector has a very tight affinity with the enzyme. In these cases, the protocols outlined by Williams and Morrison (1979) will prove useful.
Initial Rate and Steady-State Condition In an experiment, the investigator typically measures the rate of an enzyme-catalyzed reaction by calculating the slope of the progress curve. This procedure is only valid if the plot of product concentration as a function of time has been demonstrated to be linear within the time frame of the assay. However, this is not the only criterion that determines initial rate and steady-state kinetics. Linearity has been observed in many non-steady-state systems (Allison and Purich, 1979). The general rule of thumb is that initial rate conditions prevail when the substrate concentration is still within 10% of its starting value. This value is clearly acceptable for reactions that are thermodynamically quite favorable and that do not exhibit any significant product inhibition. However, if the equilibrium constant for the reaction is not large, an estimate of the degree of substrate-to-product conversion should be made. With smaller equilibrium constants, the initial-rate assay protocol necessarily must be more sensitive. In addition, if the product has very tight affinity for the enzyme,
product accumulation can lead to significant errors in initial-rate determinations even in those cases where the degree of conversion is low. A preliminary test to assess this possibility is to look at initial rates in the absence and presence of a known amount of product. Where product inhibition is a problem, product can be depleted if an auxiliary enzyme system for doing this is present. Nevertheless, product inhibition can be an important regulatory scheme for an enzyme. Any kinetic characterization of an enzyme should include accurate measurements of the dissociation constants for all products. Product-inhibition studies can also assist the investigator in establishing mechanistic details of the protein undergoing evaluation (Rudolph, 1979). In addition to establishing that the progress curve is truly linear, demonstrating that the degree of substrate depletion in the initial time course of the assay is low, and assessing the degree of product inhibition that may be present, the investigator should also demonstrate in the early stages of the kinetic studies that the rates observed are linear with respect to the total enzyme concentration ([E]total). If true steadystate conditions are present, a plot of initial rates versus [E]total (in which the enzyme concentration is varied above and below the final value to be used in the initial-rate assays) should be linear.
Substrate Concentration Ranges A wide range of substrate concentrations should be tested in preliminary studies. Such trials will provide an estimate of the Michaelis constant (Km) and an assessment of whether substrate inhibition occurs at elevated substrate levels. These tests will also detect the possible existence of cooperativity. Once an estimate of the Michaelis constant has been made, a more detailed kinetic study can be made, in which the substrate concentration is typically varied from ∼0.2 to 5.0 times the value of the apparent constant. It should be emphasized that reasonable estimates of the Km value can only be made when initial rates have been obtained from substrate concentrations both below and above the true Km value. For a one-substrate enzymecatalyzed reaction: Km = [A](Vmax − v)/v
where [A] is the substrate concentration, Vmax is the maximal velocity of the reaction, and v is the velocity of the reaction at the substrate concentration. A series of substrate concentrations with their respective initial rate velocities
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will provide an estimate of the Michaelis constant. In a standard double-reciprocal plot, the horizontal intercept is numerically equal to −1/Km. For multisubstrate enzymes (see below), estimates of the Michaelis constants will be obtained from slope and intercept replots of the initial rate data. In addition, appropriate statistical treatment of the raw data should be provided (Wilkinson, 1961; Cleland, 1979a) for both Km and Vmax values (or kcat values, equal to Vmax/[E]total, if those are reported). Note that, regardless of whether Vmax or kcat values are reported, the total protein concentration, as well as the specific activity of the enzyme used, should be reported whenever possible. With enzyme systems utilizing two substrates, the concentrations of both substrates must be varied as described above. Typically, three solutions are prepared. One solution contains one of the substrates at its highest level. A second solution contains the second substrate at its highest level. The third solution contains the buffer, all cofactors, metal ions, auxiliary enzymes, and any other effectors, but no substrates. This third solution is used to prepare serial dilutions of the other two stock solutions. Thus, a matrix of initial rate measurements on both substrates can be evaluated from a single experimental trial. Figure 3.5.2 illustrates a typical double-reciprocal plot for a two-sub-
strate enzyme-catalyzed reaction (where the the two substrates are A and B). In this example, Michaelis constants for the two substrates can be obtained by constructing slope and/or intercept replots of the data. For example, plotting the slopes of the five lines as a function of the reciprocal of the concentration of B will yield a straight line with a horizontal intercept numerically equal to −1/KB, where KB is the Michaelis constant for B. For enzymes that utilize three or more substrates, the protocol described above can become unwieldy. Two procedures have been described to address the design of kinetic studies for these proteins. In the first method (Frieden, 1959), one of the three substrates is held constant at a concentration above its respective Michaelis constant that is still subsaturating. The other two are then varied as described above. This same procedure is then followed with each of the remaining substrates. Thus, multisubstrate enzymes are reduced to pseudo-two-substrate systems. One distinct advantage of this procedure is that standard graphical presentations of the data thus produced (e.g., via double-reciprocal plots) are identical to that shown in Figure 3.5.2. However, the concentration value for the third substrate have to be provided. In the second procedure, the concentration of one substrate is varied while the other substrates are held at a
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Figure 3.5.2 Double-reciprocal plot of a two-substrate enzyme-catalyzed reaction, where v = the velocity of the reaction and [A] and [B] represent the concentrations of substrates A and B respectively.
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constant ratio, although the absolute amounts are varied. This is then repeated for all remaining substrates until the concentration of each of the substrates has been varied (Fromm, 1967). One limitation with this approach is the difficulty in obtaining true Michaelis constants. The presence of substrate inhibition can be an annoyance to an investigator who is attempting to determine the kinetic parameters of an enzyme as well as identify the binding pattern. However, the pattern of such inhibition can serve as a tool in understanding the kinetic and possibly the regulatory scheme for the enzyme. Substrate inhibition can occur as a result of a number of different causes—i.e., the formation of a dead-end complex with a particular enzyme form, the appearance of a different binding mechanism at elevated levels of substrate, the presence of an allosteric site for the substrate, or nonspecific inhibition that is due to an elevated ionic strength or another factor. When substrate inhibition occurs, in the case of multisubstrate enzymes, estimates of the Ki value (i.e., the dissociation constant for inhibition) for the substrate can be obtained from slope and intercept replots of the kinetic data. For enzymes having a single substrate, the data can be fit to an enzyme-rate expression containing a term for substrate inhibition (v = Vmax[S]/{Km + [S] + [S]2/Ki}). In addition, estimates of Km, Ki, and Vmax can be obtained from double-reciprocal plots (Cleland, 1979b). Note that in such cases, double-reciprocal plots will be nonlinear, with the degree of curvature dependent on the magnitude of the Ki value. In these cases, it may be necessary to use a computer program to fit the data. Cleland (1979a) has provided a program to address substrate inhibition. In addition, one might consider utilizing a Marmasse plot (Marmasse, 1963). Although rarely used, this plot provides good estimates of Km and Ki. A Marmasse plot is a plot of 1/v versus (α + 1/α ), where α = [A]/[A]m and [A]m represents the concentration of substrate at which the velocity is highest (note that this is not Vmax, as substrate inhibition is present.) This plot, when combined with the observation that [A]m = 1 (Km/Ki) ⁄2, will provide values for the kinetic parameters (Cleland, 1979b).
CONTINUOUS VERSUS STOP-TIME ASSAYS As the best estimate for the initial rate of an enzyme-catalyzed reaction derives from the slope of the steady-state region of the progress curve, continuous assays are greatly preferable to single-point assays. If it is not possible to use
a continuous assay, the investigator must demonstrate that the single-point assay is a true measure of the steady-state velocity of the reaction. In the single-point method, an aliquot of the reaction mixture is removed from the sample at a given time point after the reaction has been initiated (after a “blank” aliquot has been removed from the reaction mixture prior to the initiation step). Aliquots from at least four different time points are removed from the reaction solution; then the reaction is terminated and the amount of product formed (or substrate depleted) is determined. It should be ascertained that the termination procedure truly quenches the reaction. Once the steady-state relationship has been established, replicate kinetic analyses at each substrate concentration will provide sufficient data for a statistical analysis. The number of replicate samples that are assayed should also be reported.
COUPLED ENZYME ASSAYS In many instances, continuous assays are made possible by the presence of auxiliary enzymes. Product formation is measured by having the product react further with another enzyme (or enzymes). Although such assay systems are convenient, precautions must be exercised to ensure that the initial rates observed reflect the enzyme that is undergoing investigation and not the auxiliary system. If the auxiliary system is influencing the initialrate results, the reported values for the kinetic parameters will be in error and the double-reciprocals may exhibit considerable nonlinearity. The product or products formed by the enzyme undergoing study clearly direct the choice of auxiliary enzyme(s). This coupling system should provide turnover in excess over that due to the enzyme being investigated. However, the degree of excess will vary depending upon the assay method, the nature of the coupling system, and the magnitude of the kinetic parameters for each auxiliary enzyme as measured under the same conditions being observed with the target enzyme. Once this information is in hand, the amount of auxiliary enzyme needed can be calculated (McClure, 1969; Storer and Cornish-Bowden, 1974). Test runs should then be undertaken using both the calculated and higher levels of auxiliary enzymes to insure that identical rates are obtained, indicating that a sufficient excess has been achieved. The auxiliary enzyme system should also have minimal effect on cofactors and effectors required by the target enzyme.
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Frequently, a small amount of product may be present in the reaction mixture as a contaminant. For example, commercial preparations of ATP often include a small amount of ADP. The contaminating product can be removed prior to initiation of the assay by incubating the reaction mixture with the auxiliary system. The system should also be checked to insure that no activity is observed in the absence of the auxiliary enzymes. Finally, care should be exercised when using commercial preparations for the auxiliary enzymes. Often these sources of proteins contain contaminating activities that may cause serious difficulties unless they are addressed.
BINDING STUDIES
Kinetic Assay Methods
Measurements of equilibrium ligand binding can frequently add to the characterization of an enzyme’s binding mechanism, and also act as a tool in such procedures as molecularreceptor binding studies and radioimmunoassays. A wide variety of methods have been utilized in characterizing binding phenomena—e.g., gel filtration and equilibrium dialysis. The theory underlying this valuable tool can be found in any physical biochemistry text. Commonly, the binding data are analyzed with Scatchard plots (Scatchard, 1949) in which the number of moles of ligand bound per mole of total protein (ν) are plotted versus the ratio of ν to the free molar concentration of the ligand. These plots are frequently used to obtain values for the ligand dissociation constant, to identify interacting sites, and to estimate the binding capacity of the protein. Care should be exercised in this last use of the binding data, as it is easy to obtain a poor estimate of the stoichiometry of binding from Scatchard plots (Klotz, 1982, 1983). For example, a Scatchard plot of the binding of diazepam to benzodiazepine receptors is reported to provide an incorrect total number of receptor sites. Often, such problems are due to having only a few data points in the Scatchard plot at which the ligand concentration is above the half-maximum binding point. Just as in kinetic data, concentrations of the binding ligand both above and below this half-maximum point should be investigated (Klotz, 1982). Feldman (1983) has recently presented a statistical analysis associated with ligand binding data and Scatchard plots. As this paper points out, estimates of molar binding capacity should always be reported with the appropriate degree of statistical certainty. Without such statistical analyses, reports of the number of binding sites can be meaningless.
PRESENTATION OF INITIAL-RATE DATA The description of the assay protocols used in connection with enzyme purifications, in clinical studies, and in all kinetic investigations should include a clear definition of the unit of enzyme activity. When possible, velocities should be expressed in terms of molarity changes with time, as molarity is an intensive variable. Other units, proportional to molarity, are in common use. In all cases, the conditions of the assay should be fully described, including the volume of the initiating aliquot, the specific activity, and the amount of enzyme used in each experiment. The international unit (U), defined as the conversion of 1 µmol of substrate to product per minute, is in common use. Another unit of catalytic activity, the katal, corresponding to the transformation of one mole of substrate per second, is also in use. It has been suggested that the latter unit be restricted to catalytic activities measured in a clinical context (Dybkaer, 1979). Initial-rate data is frequently best presented via graphical displays. These visual linearizations of kinetic information provide a straightforward evaluation of the binding sequence and mechanism, as well as a ready appraisal of the kinetic parameters, reversible inhibition, cooperativity (both allosteric and hysteretic), and many other steady-state parameters. Whenever possible, slope and/or intercept replots should also be provided in the graphical display, perhaps as insets of the initial rate data. There are three methods frequently employed for depicting kinetic data, with the double-reciprocal plot (i.e., 1/v versus 1/[A]; see Fig. 3.5.2) being the most common. The other common plots are [A]/v versus [A] and v versus v/[A]. Wilkinson (1961) has discussed each of these plots in terms of statistics. When the data are unweighted, the three graphical presentations are not statistically equivalent. However, this problem is considerably mitigated by proper analysis using computer methods and data weighting (Siano et al., 1975). As mentioned earlier, Cleland (1979a) has provided a number of computer programs for the analysis of data. Eisenthal and Cornish-Bowden (1974) have also introduced a novel method for graphical treatment of data that provides a useful tool for evaluating experimental error. An Eisenthal–Cornish-Bowden plot is based on the equation (Vmax/v) − (Km/[A]) = 1. Instead of a particular set of data (i.e., a single initial rate velocity with its respective substrate concentration) being represented by a point in a Car-
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Current Protocols in Protein Science
tesian coordinate system, the velocity (v) is denoted along the vertical axis and the substrate concentration ([A]) is denoted along the horizontal axis. The two points thus plotted are then connected by a line. Thus, whereas in the other graphical procedures the velocity and substrate-concentration data are represented by points, in the Eisenthal–Cornish-Bowden plot the same data are represented by lines. These lines, obtained from a set of initial-rate experiments, will intersect about a point equivalent to v = Vmax and [A] = Km. Because median values of intersection points are used, outlier (i.e., aberrant) data will have minimal effects on estimates of the kinetic parameters. In addition, this plot is very useful in that calculations are not required and data do not have to be weighted. However, it does have limitations (Cornish-Bowden and Endrenyi, 1981, 1986). For example, the plot can become quite unwieldly with multisubstrate enzymes and in inhibition studies. It has recently been suggested (Brand and Johnson, 1992; Johnson and Brand, 1994) that all of the graphical methods described above may be unnecessary now that personal computers and associated new methods of data analysis are available. For example, global analysis of biochemical data is a powerful tool that may prove to be quite useful in the analysis of enzyme kinetics (Beechem, 1992). However, these numerical methods have not yet been commonly applied to studies such as those of multisubstrate systems, equilibrium exchanges, or product inhibition. At this time, with proper statistical weighting of the data, multisubstrate systems are more readily appraised by one of the common graphical procedures. More importantly, these visual depictions of initial-rate data are a clear aid to any individual reviewing the kinetic experiments.
CONCLUDING REMARKS Initial-rate assays are extremely useful tools in the detailed characterization of an enzyme. The behavior, mechanism, and regulatory properties of an enzyme can be readily assessed by straightforward assays. Such assays can frequently be completed within a day, once preliminary runs have identified the magnitude of the kinetic parameters and revealed those factors that affect activity. However, the details of the kinetic mechanism and regulatory properties of the enzyme can easily be obscured by poor assay design. The structural/functional role of an enzyme, as well as its regulation and control, can only be assessed by a reliable
initial-rate assay. In addition, the data assembled from these rigorous studies permit the development of more reliable and sensitive assay protocols for protein purification and identification.
LITERATURE CITED
Allison, R.D. 1985. γ-Glutamyl transpeptidase: Kinetics and mechanism. Methods Enzymol. 113:419-437. Allison, R.D. and Meister, A. 1981. Evidence that transpeptidation is a significant function of γglutamyl transpeptidase. J. Bio l. Ch em. 256:2988-2992. Allison, R.D. and Purich, D.L. 1979. Practical considerations in the design of initial velocity enzyme rate assays. Methods Enzymol. 63:3-22. Beechem, J.M. 1992. Global analysis of biochemical and biophysical data. Methods Enzymol. 210:37-54. Bergmeyer, H.U. (ed.) 1983. Methods of Enzymatic Analysis, 3rd ed., Vols. 1-4. Verlag Chemie, Deerfield Beach, Florida. Boehlein, S.K., Richards, N.G.J., and Schuster, S.M. 1994. Glutamine-dependent nitrogen transfer in Escherichia coli asparagine synthetase B. J. Biol. Chem. 269:7450-7457. Bowen, W.J. and Kerwin, T.P. 1955. The purification of myokinase with an ion exchange resin. Arch. Biochem. Biophys. 57:522-524. Brand, L. and Johnson, M.L. (eds.). 1992. Numerical computer methods. Methods Enzymol. Vol. 210. Cleland, W.W. 1979a. Statistical analysis of enzyme kinetic data. Methods Enzymol. 63:103-138. Cleland, W.W. 1979b. Substrate inhibition. Methods Enzymol. 63:500-513. Cornish-Bowden, A. and Endrenyi, L. 1981. Fitting of enzyme kinetic data without prior knowledge of weights. Biochem. J. 193:1005-1008. Cornish-Bowden, A, and Endrenyi, L. 1986. Robust regression of enzyme kinetic data. Biochem. J. 234:21-29. Deans, N.L., Allison, R.D., and Purich, D.L. 1992. Steady-state kinetic mechanism of bovine brain tubulin:tyrosine ligase. Biochem. J. 286:243251. Dybkaer, R.. 1979. International Union of Pure and Applied Chemistry and International Federation of Clinical Chemistry. IUPAC Section of Clinical Chemistry; Commission on Quantities and Units in Clinical Chemistry and IFCC Committee on Standards, Expert Panel on Quantities and Units; Approved Recommendation (1978) Quantities and units in clinical chemistry. Clin. Chim. Acta 96:157F-183F. Eisenthal, R. and Cornish-Bowden, A. 1974. The direct linear plot. A new graphical procedure for estimating enzyme kinetic parameters. Biochem. J. 139:715-720.
Detection and Assay Methods
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Supplement 5
Feldman, H.A. 1983. Statistical limits in Scatchard analysis. J. Biol. Chem. 258:12865-12867.
hexokinase type III isozyme. Arch. Biochem. Biophys. 170:587-600.
Fierke, C.A. and Hammes, G. 1995. Transient kinetic approaches to enzyme mechanisms. Methods Enzymol. 249:1-32.
Silverstein, E. and Sulebele, G. 1969. Catalytic mechanism of pig heart mitochondrial malate dehydrogenase studied by kinetics at equilibrium. Biochemistry 8:2543-2550.
Frieden, C. 1959. Glutamate dehydrogenase III. The order of substrate addition in the enzymatic reaction. J. Biol. Chem. 234:2391-2396. Fromm, H.J. 1967. The use of competitive inhibitors in studying the mechanism of action of some enzyme systems utilizing three substrates. Biochim. Biophys. Acta 139:221-230. Johnson, M.L. and Brand, L. (eds.). 1994. Numerical computer methods, part B. Methods Enzymol. 240:xi-xiii, 1-36, 51-170, 181-198, 311-322, 781-816. Klotz, I.M. 1982. Numbers of receptor sites from Scatchard graphs: Facts and fantasies. Science 217:1247-1249. Klotz, I.M. 1983. Number of receptor sites from Scatchard and Klotz graphs: A constructive critique. Science 220:981. Klump, H., Di Ruggiero, J., Kessel, M., Park, J.-B., Adams, M.W.W., and Robb, F.T. 1992. Glutamate dehydrogenase from the hyperthermophile Pyrococcus furiosus. J. Biol. Chem. 267:2268122685.
Tate, S.S. and Meister, A. 1978. Serine-borate complex as a transition-state inhibitor of γ-glutamyl transpeptidase. Proc. Natl. Acad. Sci. U.S.A. 75:4806-4809. Wilkinson, G.N. 1961. Statistical estimations in enzyme kinetics. Biochem. J. 80:324-332. Williams, J.W., and Morrison, J.F. 1979. The kinetics of reversible tight-binding inhibition. Methods Enzymol. 63:437-467.
KEY REFERENCES Bergmeyer, 1983. See above. Detailed collection of enzyme assay protocols and methods for a wide variety of enzymes, including descriptions of many special techniques: e.g., fluorometry, turbidimetry, luminometry, and microtechniques.
Lienhard, G.E. and Secemski, I.I. 1973. P1, P5Di(adenosine-5′)pentaphosphate, a potent multisubstrate inhibitor of adenylate kinase. J. Biol. Chem. 248:1121-1123.
Dixon, M. and Webb, E.C. 1979. Enzymes, 3rd ed. Academic Press, New York.
Marmasse, C. 1963. Enzyme inhibition by excess substrate. Biochim. Biophys. Acta 77:530-535.
Fromm, H.J. 1975. Initial Rate Enzyme Kinetics. Springer-Verlag, New York.
McClure, W.R. 1969. A kinetic analysis of coupled enzyme assays. Biochem. 8:2782-2786.
Valuable introductory text with considerable details on experimental design.
Nordlie, R.C. 1982. Kinetic examination of enzyme mechanisms involving branched reaction pathways: A detailed consideration of multifunctional glucose-6-phosphatase. Methods Enzymol. 87:319-353. O’Sullivan, W.J. and Smithers, G.W. 1979. Stability constants for biologically important metal-ligand complexes. Methods Enzymol. 63:294-336.
Classic text on the basics of enzymology; Chapter 4 deals specifically with enzyme kinetics.
Kuby, S.A. 1990. A Study of Enzymes, vol. I. CRC Press, Boca Raton, Florida Detailed text on steady-state kinetics. Contains useful chapters on the effects of metal cofactors and transient phase kinetics. Purich, D.L. (ed.) 1979. Enzyme kinetics and mechanism, part A. Methods Enzymol. Vol. 63.
Purich, D.L. and Fromm, H.J. 1972. Inhibition of rabbit muscle adenylate kinase by the transition state analogue, P1,P4-di(adenosine-5′)tetraphosphate. Biochim. Biophys. Acta 276:563-567.
Describes a variety of initial rate methods and inhibitor studies for characterizing enzyme-catalyzed reactions.
Rudolph, F.B. 1979. Product inhibition and abortive complex formation. Methods Enzymol. 63: 411436.
Purich, D.L. (ed.) 1980. Enzyme kinetics and mechanism, part B. Methods Enzymol. Vol. 64.
Scatchard, G. 1949. The attraction of proteins for small molecules and ions. Ann. N.Y. Acad. Sci. 51:660-673. Seelig, G.F. and Meister, A. 1985. Glutathione biosynthesis: γ-Glutamylcysteine synthetase from rat kidney. Methods Enzymol. 113:379-390. Siano, D.B., Zyskind, J.W., and Fromm, H.J. 1975. A computer program for fitting and statistically analyzing initial rate data applied to bovine Kinetic Assay Methods
Storer, A.C. and Cornish-Bowden, A. 1974. The kinetics of coupled enzyme reactions. Biochem. J. 141:205-209.
Describes a number of isotopic probes for studying enzyme mechanisms and contains valuable chapters on allosterism, hysteresis, immobilized systems, and processivity. Purich, D. L. (ed.) 1982. Enzyme kinetics and mechanism, part C. Methods Enzymol. Vol. 87. Describes methods for characterizing intermediates in enzyme-catalyzed reactions and using stereochemical probes for enzyme mechanisms; includes additional initial-rate and inhibitor methods and discusses further uses of isotopic probes.
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Purich, D.L. (ed.) 1995. Enzyme kinetics and mechanism, part D. Methods in Enzymol. 249:3662. Covers in detail specialized topics in enzyme kinetics including transition-state approaches, kinetic probes with site-directed mutagenesis, partition analysis, positional isotope exchange, interfacial catalysis, and hydrogen tunneling.
Contributed by R. Donald Allison University of Florida Gainesville, Florida
Detection and Assay Methods
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Biotinylation of Proteins in Solution and on Cell Surfaces
UNIT 3.6
A protein is typically labeled with the water-soluble vitamin biotin to tag it for later recovery or for detection with biotin-binding proteins, such as avidin or streptavidin. Applications of this approach include identification and purification of cell surface–exposed proteins, measurement of ligand internalization kinetics, and temporal changes in protein distributions inside cells using light or electron microscopy. Many modified biotins, capable of reacting with protein lysyls, cysteinyls, or carbohydrates, are commercially available (see Table 3.6.1). It is usually possible to find at least one biotin reagent that can derivatize a given protein without affecting the function of interest. For instance, most enzyme activities are independent of protein modification on carbohydrate residues. Also, when used with enzyme-linked or radiolabeled avidin or streptavidin, biotinylation can be an effective technique for visualizing proteins that do not bind well to dyes after separation by SDS–polyacrylamide gel electrophoresis (SDS-PAGE; UNIT 10.1). This unit contains three protocols for biotinylation of isolated proteins: attaching biotin to primary amines (e.g., ε-amino groups of lysyl residues; see Basic Protocol 1); attaching biotin to sulfhydryls (i.e., thiol groups of cysteinyl residues; see Basic Protocol 3); and attaching biotin to carbohydrate residues on proteins (see Basic Protocol 4). As biotinylation of lysyl and cysteinyl residues may alter protein function, modification of protein carbohydrates, which is usually innocuous, may be preferable if intact protein function is required (e.g., for activity assays or affinity purification). Biotinylation of cysteinyl thiols requires that disulfide bonds in isolated proteins be reduced before labeling (see Support Protocol 1). Mild reaction conditions (e.g., in aqueous buffers at pH 7.5 to 8.5) and the small size of the biotin moiety also make biotinylation ideal for labeling surface proteins on living cells (see Basic Protocol 2). Such site-specific labeling enables investigators to monitor and/or effect subcellular purifications. Finally, this unit includes a brief description of methods for detecting biotinylated proteins (see Support Protocol 2). COVALENT ATTACHMENT OF BIOTIN TO LYSINES This protocol is a specialized version of the succinimidyl ester amidation reaction described in Basic Protocol 1 of UNIT 15.2. The reagent used, NHS-LC-biotin (see Fig. 3.6.1), has an N-hydroxysuccinimide (NHS) ester group to increase reactivity with primary amines and a long-chain (LC) spacer arm to extend the biotin from the protein surface, thus improving binding of subsequent reagents (e.g., avidin). The protein of interest is incubated with NHS-LC-biotin. Glycine is added to stop the reaction, and biotinylated proteins are purified by dialysis. This protocol works well over the range of 50 µl to 50 ml (and possibly for much larger volumes), so the volume of the reaction mixture should be chosen to suit the investigator’s needs.
BASIC PROTOCOL 1
If biotinylation of lysines results in loss of protein function, biotinylation of thiols (see Basic Protocol 3) or carbohydrate groups (see Basic Protocol 4) may prove useful. NOTE: NHS-biotin should be stored in the presence of desiccant at −20°C. When the reagent is thawed, it should be warmed thoroughly to room temperature before the container is opened. Enzymatic Manipulation of DNA and RNA Contributed by Elizabeth J. Luna Current Protocols in Protein Science (1996) 3.6.1-3.6.15 Copyright © 1996 by John Wiley & Sons, Inc.
3.6.1 Supplement 6
Table 3.6.1
Commercially Available Biotinylating Reagents
Suppliera
Reagent
Molecular weight
Comments
Specific for reaction with primary amines (see Basic Protocols 1 and 2, Support Protocol 2) NHS-biotin Pierce, Sigma, Vector Labs Soluble in DMF or DMSO; store all NHS 341.4 reagents at −20°C in presence of desiccant Sulfo-NHS-biotin Pierce, Sigma, Vector Labs 443.4 Water soluble NHS-LC-biotin Pierce, Sigma, Vector Labs 454.5 Soluble in DMF or DMSO; spacer arm improves accessibility of protein-bound biotin to affinity matrices NHS-SS-biotin Pierce 606.7 Soluble in DMF; cleavable by disulfide reduction 2-Iminobiotin-NHS Calbiochem, Pierce 421.3 Soluble in DMF; exhibits pH-sensitive binding to avidin Specific for reaction with sulfhydryls (see Basic Protocol 3, Support Protocols 1 and 2) Iodoacetyl-LC-biotin Pierce 510.4 Soluble in DMF or DMSO; reacts with amino groups above pH 7.5 Biotin-LC maleimide Sigma 479.6 Soluble in DMF; reacts with amino groups above pH 7.5 Maleimido-propionyl Molecular Probes, Pierce 523.6 Soluble in DMF; reacts with amino groups biocytin above pH 7.5 Biotin-HPDP Pierce 539.8 Soluble in DMSO; cleavable by disulfide reduction, regenerating initial cysteinyl Specific for reaction with (oxidized) carbohydrates (see Basic Protocol 4, Support Protocol 2) Biotin-LC-hydrazide Calbiochem, Pierce, Vector 258.3 Soluble in water (to ∼5 mM) and DMSO Labs (to ∼50 mM) Biotin-hydrazide Molecular Probes, Pierce, 371.5 Soluble in water (to ∼5 mM) and DMSO Sigma (to ∼50 mM) aSee SUPPLIERS APPENDIX for addresses and phone numbers of manufacturers.
Materials ∼1 mg/ml purified protein Coupling buffer (see recipe), ice cold 2 mg/ml NHS-LC-biotin (Pierce, Vector Labs) in N,N-dimethylformamide (DMF), room temperature, prepared just before use 50 mM glycine in coupling buffer 0.22-µm filter: e.g., Spin-X centrifuge filter (Costar) or syringe filter (Nalgene) Additional reagents and equipment for dialysis (UNIT 4.4 and APPENDIX 3B) 1. If necessary, dialyze the purified protein into coupling buffer to remove traces of interfering buffer components that may have been introduced during purification. Complete exchange of buffer components can be assumed after a 48-hr dialysis against 100 vol of coupling buffer with three to four buffer changes. However, if the initial concentration of the interfering agent is relatively low, a 16- to 24-hr dialysis with two buffer changes is generally sufficient.
Biotinylation of Proteins in Solution and on Cell Surfaces
Azide, glycine, ammonium ions, and Tris are common buffer components that interfere with the biotin labeling reaction. Although these reagents should be avoided, reagents without primary or secondary amino groups can be included to stabilize the protein or to prevent labeling at functional sites. For instance, NaCl, CaCl2, sucrose, glycerol, Triton X-100, and HEPES all can be used in the coupling buffer, if required.
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O HN NaO3S NH2
+
O
NH
O
N O C (CH2)5 NH C (CH2)4
protein
S
NHS-LC-biotin
O HN
O
NH
O
NH C (CH2)5 NH C (CH2)4 protein
S biotinylated protein
+ NaO3S
O N OH O
Figure 3.6.1 Reaction of N-hydroxysuccinimide (NHS)-LC-biotin with protein.
2. Rapidly add the freshly prepared suspension of NHS-LC-biotin in DMF to the protein in coupling buffer, using 1 vol biotin in DMF to 20 vol protein solution. The biotin/protein mixture may become turbid because NHS-LC-biotin forms a fine suspension in water, rather than a true solution. If the protein is sensitive to organic solvents or if the highest affinity binding to avidin afforded by the LC spacer is not necessary, sulfo-NHS-biotin (Pierce, Sigma, or Vector Labs) can be substituted for NHS-LC-biotin. Sulfo-NHS-biotin is very water soluble and can be added at the same final biotin/protein molar ratio (∼10:1) from a freshly prepared aqueous stock solution without precipitate formation.
3. Incubate ∼1 hr on ice. Mix occasionally. Optimal extents of biotinylation must be determined empirically for each protein and are a compromise between high levels of incorporated biotin and retention of protein function. The biotin/protein ratio during labeling and the length of labeling time recommended here have worked well for a number of proteins. However, biotin-to-protein ratios as low as 1:1 and as high as 60:1 have been reported. Similarly, optimal labeling times may vary between 1 and 18 hr.
4. Add 1 vol of 50 mM glycine to bind unreacted biotin. 5. Remove glycine and free biotin by exhaustive dialysis against coupling buffer, or another buffer of choice. Dialysis for ≥36 hr against three or more changes of 500 vol each is recommended. Once the labeling step is complete, any buffer, including Tris⋅HCl, can be used for dialysis (UNIT 4.4 & APPENDIX 3B). Unincorporated sulfo-NHS-biotin can be removed quickly by gel-filtration on an Econo-Pac 10DG desalting column (Bio-Rad) or a Sephadex G-25 column (Pharmacia Biotech; see UNIT 8.3). However, because any suspended micelles of unreacted NHS-LC-biotin will coelute in the void volume with the protein, proteins labeled with NHS-LC-biotin should be dialyzed overnight against one change of buffer before gel filtration.
Enzymatic Manipulation of DNA and RNA
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6. For long-term storage, sterilize biotinylated proteins by passing the solution through a 0.22-µm filter. Spin-X centrifuge filters (Costar) work well for small volumes (<1 ml); syringe filters (Nalgene) are recommended for much larger volumes. Sodium azide also may be added up to 0.02% (w/v) to inhibit bacterial growth, but it is most effective below pH 7.0. Thimerosal (0.1% [w/v]) is more useful in solutions with higher pH values. CAUTION: Sodium azide is toxic; wear gloves when handling.
7. Store biotinylated proteins on ice or at 4°C. Some biotinylated proteins can be frozen and stored in aliquots at −20° or −70°C; others are reported to aggregate or lose function under these conditions. Generally, the biotinylated protein should be stored under the conditions recommended for the unlabeled protein. If no information is available about storage conditions for the protein of interest, it is wise to test small aliquots for relative stability under the following sets of conditions: (1) fast frozen in sterile cryogenic vials (Corning) or in test tubes with O-ring caps (Sarstedt) using a dry ice/ethanol slurry, followed by storage of aliquots (1a) at −20°C, (1b) at −70°C, and (1c) in a liquid nitrogen Dewar flask. Other aliquots should be (2) mixed well with an equal volume of glycerol (to make a 50% glycerol solution) and stored at −20°C, (3) stored on ice, and (4) kept at 4°C. Shortly after the initial storage and at periodic intervals thereafter, aliquots should be removed from each of the storage conditions and tested for biological, binding, or enzymatic activity. Once the most efficient storage condition has been determined, future preparations of the biotinylated protein should be stored accordingly in aliquots sufficient for one, or a few, experiments. Repeated cycles of freezing and thawing should be avoided. CAUTION: Tubes stored under liquid nitrogen can explode on warming if liquid nitrogen has gotten inside the tube. Loosen caps on tubes immediately after taking them out of a liquid nitrogen Dewar flask. The concentration of the biotinylated protein can be determined using any of the assays described in UNIT 3.4, and the concentration of protein-associated biotin can be measured in a variety of solid-phase competition assays with labeled avidin and known amounts of free biotin (Savage et al., 1992). However, because most assays with biotinylated protein are either qualitative or comparative, many investigators merely verify that biotinylation has occurred, i.e., by analysis of electrotransferred protein after SDS-PAGE (UNIT 10.10). BASIC PROTOCOL 2
SIDE-SPECIFIC BIOTINYLATION OF PROTEINS ON CELL SURFACES Biotinylation of proteins at the extracellular surface of living cells is a fast, nonradioactive means of labeling and tracking the plasma membrane during cell fractionation (Ingalls et al., 1986; UNIT 4.2). The biotinylated proteins also are easily isolated after membrane solubilization and affinity chromatography on columns containing bound avidin or streptavidin (Chia and Luna, 1989). Biotinylation of living cells requires the water-soluble biotin derivative sulfo-NHS-biotin. The starting material is intact cells, either freshly isolated or from suspension or adherent cell cultures; control experiments starting with lysed cells or isolated membranes are, however, recommended to verify that cell integrity is maintained during the procedure. Cells are reacted with freshly dissolved sulfo-N-hydroxysuccinimide (NHS)-biotin for 30 min with mixing. Glycine is added to bind unreacted biotin, and cells are washed and lysed to prepare plasma membranes or solubilized proteins.
Biotinylation of Proteins in Solution and on Cell Surfaces
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NOTE: NHS-biotin should be stored in the presence of desiccant at −20°C. When the reagent is thawed, it should be warmed thoroughly to room temperature before the container is opened. Materials 107 to 1010 cells of interest, in suspension (e.g., erythrocytes, leukocytes, or HeLa cells), or cultures of adherent cells Physiological buffer, pH 7.4 (see recipe), ice cold Sulfo-NHS-biotin (Pierce or Vector Labs) 10 mM glycine in physiological buffer Additional reagents and equipment for lysing cells and isolating plasma membranes (UNIT 4.2 and UNIT 4.3) or for solubilizing proteins for SDS gel electrophoresis (UNIT 10.1) NOTE: Carry out all steps in this protocol at 4°C or on ice. 1. Remove serum proteins by washing the cells of interest at least three times with ice-cold physiological buffer, centrifuging cells in suspension for 5 min at ∼500 × g, 4°C, between washes. Adherent cells are most easily washed and labeled while still attached to tissue culture flasks. To avoid labeling cytoplasmic proteins, the biotinylating agent should be thoroughly blocked and removed before dissociating attached cells from the flask (i.e., detach the cells after step 5). The buffer can be varied, depending on the preferences of the cells and the investigator. Dulbecco’s PBS or Earle’s balanced salts (e.g., Sigma) also work well. In contrast, PBS containing only ∼5 mM phosphate (APPENDIX 2E) may not have sufficient buffering capacity. As in Basic Protocol 1, the principal consideration is to omit reagents containing primary amino groups that could compete for binding to the sulfo-NHS-biotin.
2. During the final cell wash, dissolve the sulfo-NHS-biotin in 0.1 vol ice-cold physiological buffer for a final concentration of 2 mg/ml (∼5 mM). For instance, for ∼1010 cells in a volume of 9 ml, dissolve 20 mg sulfo-NHS-biotin in 1 ml physiological buffer. If labeling attached cells, mix the sulfo-NHS-biotin with the entire amount of buffer immediately before adding the mixture to the cells. Minimize the amount of hydrolysis of the succinimide ester by limiting its exposure to water in the absence of cells.
3. Gently suspend the cells by adding ice-cold physiological buffer first. Then add 1⁄10 vol of dissolved sulfo-NHS-biotin. Mix gently without lysing the cells. Although labeling of cell-surface lysines proceeds faster as the pH of the buffer is raised from pH 7.4 to 9.0, so does the hydrolysis of the sulfo-NHS-biotin. In addition, many cells are lysed in higher-pH buffers.
4. Incubate 30 min at 4°C with gentle rotation or swirling. This procedure should label only surface-exposed proteins. Higher temperatures will result in increased biotinylation of membrane proteins, but much of the increase may be due to endocytic membrane turnover.
5. Block unreacted biotinylating reagent for ≥10 min by adding an equal volume of 10 mM glycine in physiological buffer. Wash cells with buffer three times as in step 1. Monitor the cells for lysis during centrifugation, using phase-contrast or differential interference contrast (DIC) light microscopy. If the cells are too fragile to withstand repeated washes, incubate with the glycine at 4°C for 15 to 30 min, then wash only one or two times. Blocking is faster at elevated pH, but pH 7.4 is adequate as long as the cells are not permeabilized during the washes.
Enzymatic Manipulation of DNA and RNA
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6. Lyse the cells and isolate plasma membranes (see UNIT 4.2) or solubilize proteins for SDS-PAGE (see UNIT 10.1, Basic Protocol 1). Store biotinylated membranes in aliquots at −20°C. It is usually a good idea to perform a parallel, control experiment in which cells are biotinylated after lysis or in which isolated membranes are biotinylated. The control procedure should label both surface-exposed and intracellular polypeptides, providing evidence for cellular integrity during the initial labeling procedure. The control can also demonstrate that if a protein of particular interest remains unlabeled under surface-labeling conditions but is labeled in isolated membranes or lysates, it is inherently capable of reacting with the sulfo-NHS-biotin reagent and, thus, is either a cytosolic protein or a membrane protein without accessible external lysines. BASIC PROTOCOL 3
COVALENT ATTACHMENT OF BIOTIN TO THIOLS An alternative to biotinylating lysyl groups (see Basic Protocol 1) is biotinylation of thiols or carbohydrate groups (see Basic Protocol 4). Several modified biotins react specifically with sulfhydryls (see Table 3.6.1). This protocol is described for iodoacetyl-LC-biotin in N,N-dimethylformamide (DMF). However, guidelines for using biotin-maleimide and N-(6-[biotinamido]hexyl)-3′-(2′-pyridyldithio)propionamide (biotin-HPDP) reagents are given in step annotations. Purified protein or membrane fractions are first subjected to disulfide reduction (see Support Protocol 1). The reduced material is mixed with iodoacetyl-LC-biotin. Unreacted components are then separated from labeled proteins by gel filtration or dialysis before the proteins are filter sterilized and stored. Materials 0.5 mg/ml protein or cell fraction of interest, e.g., purified membranes 100 mM sodium phosphate/5 mM EDTA, pH 8.3 (APPENDIX 2E) 4 mM iodoacetyl-LC-biotin (Pierce) in DMF, freshly prepared or thawed from storage at −20°C Additional reagents and equipment for reducing protein disulfides (see Support Protocol 1), dialysis (UNIT 4.4 and APPENDIX 3B), gel-filtration chromatography (UNIT 8.3), or centrifugal filtration (UNIT 15.1, Support Protocol 3) 1. If necessary, reduce protein disulfides in the protein or cell fraction of interest (see Support Protocol 1) and remove the thiol-reducing agent by dialysis (UNIT 4.4 and APPENDIX 3B), gel-filtration chromatography (UNIT 8.3), or centrifugal filtration (UNIT 15.1). Use the sodium phosphate/EDTA buffer to make up the solution of reduced protein. If samples must be dialyzed for prolonged periods, nitrogen gas should be bubbled through the dialysis buffer to help exclude the atmospheric oxygen that will slowly (1 to 7 days) oxidize the reduced disulfides. EDTA chelates free metal ions that can catalyze this oxidation.
2. Add 20 µl of 4 mM iodoacetyl-LC-biotin in DMF for each milliliter of protein or cell fraction in the sodium phosphate/EDTA buffer. For proteins with an “average” number of cysteines, i.e., 1.7 mol % (Table A.1A.1), this provides an equimolar ratio of iodoacetyl-LC-biotin. The cost and relative insolubility of biotinylated alkylating reagents contraindicate their use in a 5- to 10-fold molar excess over total thiols, as described in Basic Protocol 1.
Biotinylation of Proteins in Solution and on Cell Surfaces
Depending on the protein and activity to be studied, another thiol-directed biotinylating reagent may be preferable. At pH 8.3, iodoacetyl-LC-biotin reacts irreversibly and specifically with free sulfhydryls. Maleimide reagents (biotin-LC-maleimide and maleimidopropionyl biocytin) also irreversibly derivatize sulfhydryls but will react with some primary amines unless the pH is lowered to between 6.5 and 7.5. Biotin-
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HPDP reacts rapidly and specifically through disulfide exchange with free protein sulfhydryls; consequently, proteins derivatized with biotin-HPDP should not be exposed to reduced thiol agents unless reversal of the labeling procedure is desired.
3. Incubate ∼2 hr at room temperature or overnight at 0° to 4°C. 4. Remove unreacted reagent by gel-filtration chromatography (UNIT (UNIT 4.4 and APPENDIX 3B).
8.3)
or dialysis
5. Filter the solution through an 0.22-µm filter and store biotinylated proteins at 0° to 4°C or biotinylated membranes in aliquots at −20°C. REDUCTION OF DISULFIDE BONDS Reducing disulfide bonds may be necessary to create the free sulfhydryls on protein cysteines needed for biotinylation with thiol-directed biotin reagents (see Basic Protocol 3). Disulfide reduction can also be used to elute proteins tagged with the amine-directed reagent N-hydroxysuccinimide (NHS)-SS-biotin (see Basic Protocol 1) from avidin affinity columns. This protocol gives an effective method for reducing a thiol-sensitive disulfide bond that is not essential for the quaternary structure, e.g., that in immunoglobulins. The protein of interest is dialyzed into reducing buffer and mixed with reducing agent at elevated temperature (i.e., 37°C). Excess reductant is removed by gel-filtration chromatography (UNIT 8.3). The reduced protein is then ready for immediate biotinylation. In general, disulfide reduction can be assumed to be innocuous, if not advantageous, for proteins that reside in the cell cytoplasm, but it may perturb function in extracellular proteins or in the extracellular domains of transmembrane proteins.
SUPPORT PROTOCOL 1
Materials 1 to 10 mg/ml purified protein Reducing buffer: 100 mM sodium phosphate/5 mM EDTA, pH 6.0 (APPENDIX 2) Reducing agent: 50 mM dithiothreitol (DTT),100 mM 2-mercaptoethanol (2-ME), or 100 mM mercaptoethylamine⋅HCl (Sigma) Econo-Pac 10DG desalting column (Bio-Rad) or Sephadex G-25 column (Pharmacia Biotech) Additional reagents and equipment for dialysis (UNIT 4.4 and APPENDIX 3B) and gel-filtration chromatography (UNIT 8.3) 1. If necessary, dialyze purified protein of interest into reducing buffer. 2. Add 1 vol of 1 to 10 mg/ml purified protein in reducing buffer to an equal volume of reducing agent. Incubate 1 hr at 37°C with occasional mixing. Each mole of DTT contains two thiols, so only half the concentration (50 mM) is required for the same concentration of reducing equivalents used for other reducing agents (100 mM 2-ME or 100 mM mercaptoethanolamine⋅HCl).
3. During the incubation in step 2, equilibrate an Econo-Pac 10DG desalting column or a Sephadex G-25 column with reducing buffer. The column should contain ≥10 vol of gel-filtration matrix (i.e., at least five times more volume than that of the reaction in step 2).
4. Cool the reaction to room temperature. Remove excess reducing agent by gel-filtration chromatography (UNIT 8.3) using the equilibrated column. Collect at least ten fractions, each containing ∼1 vol (i.e., the same volume as the initial volume of protein solution). Alternatively, excess reducing agent may be removed by centrifugal filtration (UNIT 15.1, Support Protocol 3) or dialysis (UNIT 4.4 and APPENDIX 3B). If prolonged dialysis is required,
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reoxidation of protein disulfides can be slowed by including 5 mM EDTA and/or by bubbling nitrogen gas through the dialysis buffer.
5. Read the absorbance of each fraction at 280 nm (A280). Pool fractions that correspond to the first peak of absorbance. 6. As soon as possible after removing excess reducing agent, use the pooled fractions in cysteinyl-directed biotinylations (see Basic Protocol 3). BASIC PROTOCOL 4
COVALENT ATTACHMENT OF BIOTIN TO CARBOHYDRATES Biotinylation of protein carbohydrates requires oxidation of vicinal carbohydrate hydroxyl groups to aldehydes that can bind biotin-LC-hydrazide. Sodium m-periodate is used as the oxidizing agent for glycoproteins in solution, but enzyme-mediated oxidations can be used to oxidize glycoproteins of cells or membrane fractions. Briefly, glycoproteins are oxidized, excess oxidizing agent is removed by gel-filtration, biotin reagent is added, and the biotinylated glycoprotein is purified in a microconcentrator. Materials 0.1 to 0.2 mg/ml glycoprotein Labeling buffer: 100 mM sodium acetate, pH 5.5 (APPENDIX 2E) 30 mM and 3 mM sodium m-periodate (Sigma, mol. wt. 213.9) in water, freshly prepared and stored in the dark 5 mM biotin-LC-hydrazide (Calbiochem, Pierce, or Vector Labs) in labeling buffer (store ≤2 months at 4°C) Stop solution: 100 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) Econo-Pac 10DG desalting column (Bio-Rad) or Sephadex G-25 column (Pharmacia Biotech) Microconcentrator Additional reagents and equipment for dialysis (UNIT 4.4 and APPENDIX 3B) and gel-filtration chromatography (UNIT 8.3) 1. If needed, dialyze the glycoprotein into labeling buffer. 2. Add 1 vol of 30 mM or 3 mM sodium m-periodate to 2 vol of 0.1 to 0.2 mg/ml glycoprotein in labeling buffer for a final periodate concentration of either 10 mM or 1 mM. Incubate ∼30 min at room temperature in the dark. Because proteins differ in sensitivity to periodate, it is a good idea to try at least two different concentrations of the oxidant. For antibodies, O’Shannessy et al. (1984) report excellent results with 10 mM sodium m-periodate. For labeling a transmembrane glycoprotein that requires a detergent for solubilization, avoid the use of octylglucoside, which also reacts with periodate. Cell-surface carbohydrates on erythrocytes and on purified erythrocyte membranes have been biotinylated by labeling cells or purified membranes for 2 hr at room temperature with a solution containing 0.03 U/ml Vibrio cholerae neuraminidase (Boehringer Mannheim), 3 U/ml Dactylium dendroides galactose oxidase, and 0.4 mM biotin-LC-hydrazide in physiological buffer (Bayer et al., 1988). With many cells and cell membranes, however, those conditions will promote cell lysis and/or proteolysis.
3. During the incubation period (step 2), equilibrate an Econo-Pac 10DG desalting column or Sephadex G-25 column with labeling buffer. Biotinylation of Proteins in Solution and on Cell Surfaces
The column should contain ≥15 vol of gel-filtration matrix (i.e., at least five times more volume than that of the reaction in step 2).
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4. Remove unincorporated sodium m-periodate by gel-filtration chromatography (UNIT 8.3) using the equilibrated column. Collect at least ten fractions, each containing ∼1 vol (i.e., the same volume as the initial volume of sodium m-periodate). 5. Read the absorbance of each fraction at 280 nm (A280). Pool fractions that correspond to the first peak of absorbance. 6. Add 1 vol of 5 mM biotin-LC-hydrazide in labeling buffer. Incubate 1 hr at room temperature with occasional mixing. The amount of dilution of protein on the gel-filtration column can vary without significantly affecting the efficiency of the reaction.
7. Terminate the reaction by adding 5 vol stop solution. Tris, glycine, ammonium ions, and other buffer components containing primary amines should be avoided during the oxidation and labeling steps because they form reversible Schiff bonds with the carbohydrate aldehydes. These Schiff bonds will inhibit hydrazone bond formation. Primary amines are, however, good for stopping the labeling reaction. If the biotinylated protein is expected to be less than ∼0.5 mg/ml after concentration, a carrier protein, such as BSA, can be added after the stop solution. Carrier protein helps to eliminate protein loss during concentration but will complicate quantification of the percent recovery of the biotinylated glycoprotein and of the number of biotins incorporated per glycoprotein molecule.
8. Remove unreacted biotin-LC-hydrazide by centrifugation in a microconcentrator with a molecular weight cutoff (MWCO) that is less than half that of the molecular weight of the protein in solution (UNIT 4.4). After concentrating the protein, suspend the sample to the initial volume with labeling buffer and reconcentrate. Repeat until the buffer has been changed a total of three times and the glycoprotein has reached the desired concentration. Any buffer in which the glycoprotein is stable can be substituted for labeling buffer at this step. To estimate the final glycoprotein concentration, assume no more than 50% loss during the entire procedure, as long as the protein is known to be well-behaved in solution. If precision is necessary, the molar ratio of biotin to glycoprotein can be calculated after determining protein concentrations (UNIT 3.4) and biotin concentrations (e.g., by solidphase binding assays with avidin; Savage et al., 1992), after taking into account any added carrier protein. However, the estimate of 50% loss during experimental manipulation generally provides a reasonable starting point for most qualitative experiments (and is seldom off by more than a factor of 2).
9. Store the concentrated glycoprotein in aliquots at −70° or −20°C. For discussion of storage conditions, see Basic Protocol 1 (step 7 annotation).
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SUPPORT PROTOCOL 2
DETECTION OF BIOTINYLATED PROTEINS Once proteins or cells are biotinylated (see Basic Protocols 1 to 4), the next step is often detection (e.g., in the course of performing fluorescence or electron microscopy, assaying enzyme activity, or sizing proteins). Experimental samples and biotinylated standards can be analyzed simultaneously. This protocol gives brief instructions for separating proteins and visualizing them with 125I-labeled avidin or avidin-based chromophores. Avidin-biotin complex (ABC) detection kits are available for isolated proteins (described in this protocol) as well as for proteins on fixed tissue, tissue sections, and microplate wells. Materials Biotinylated experimental protein (see Basic Protocols 1 to 4) Biotinylated protein molecular weight standards (Bio-Rad or Vector Labs) 3% (w/v) BSA in PBS (APPENDIX 2E) Nitrocellulose membrane 0.5 µCi 125I-Bolton-Hunter-labeled avidin (DuPont NEN) in 5 ml avidin buffer (see recipe) or Vectastain ABC kit (Vector Labs) or ImmunoPure ABC kit (Pierce) Additional reagents and equipment for SDS-PAGE (UNIT 10.1), electroblotting (UNIT 10.7), and detecting proteins in immunoblots (UNIT 10.10) 1. Electrophorese biotinylated experimental protein and protein molecular weight standards in separate lanes on a one-dimensional SDS–polyacrylamide gel (UNIT 10.1) 2. Electroblot proteins onto a nitrocellulose membrane (UNIT 10.7) and block ∼1 hr at 22°C with 3% BSA in PBS. Note that the milk-based blocking solution described in UNIT 10.10 should not be used because milk contains free biotin that can lead to unacceptably high backgrounds.
3a. To visualize biotinylated proteins by labeling with radioactive avidin: Incubate the blot in 0.5 µCi of 125I-Bolton-Hunter-labeled avidin (Mock, 1990) in 5 ml avidin buffer for 2 hr at 22°C. Wash the membrane extensively in the same buffer without avidin or BSA, air dry, and expose to film (Goodloe-Holland and Luna, 1987). 3b. To visualize biotinylated proteins with avidin-linked chromophores in the Vectastain ABC kit or ImmunoPure ABC kit: Follow the manufacturer’s instructions. The ABC procedure is essentially identical to that described in UNIT 10.10, except that steps 5 and 6 (incubation with a biotinylated secondary antibody) are omitted.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Avidin buffer 150 mM NaCl 4% (w/v) BSA 0.02% (v/v) Triton X-100 50 mM sodium borate Adjust to pH 10.0 to 10.5 with 1 M NaOH Store ≤2 months at 4°C Biotinylation of Proteins in Solution and on Cell Surfaces
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Coupling buffer Select one of the following buffers (see Critical Parameters for discussion of practical considerations and APPENDIX 2E for recipes): 100 mM NaHCO3, pH 8.5 100 mM Na2CO3, pH 8.5 100 mM sodium phosphate buffer, pH 7.5 100 mM sodium phosphate buffer, pH 8.0 100 mM sodium phosphate buffer/5 mM EDTA, pH 8.3 Physiological buffer, pH 7.4 7 g NaCl (120 mM final) 22 ml 0.2 M KH2PO4 (APPENDIX 2E; 4.4 mM final) 103 ml 0.2 M Na2HPO4 (APPENDIX 2E; 20.6 mM final) Add H2O to 1 liter Check pH (it should be ∼7.4; if not, check concentrations of stock solutions) Store ≤2 months at 4°C or ≤2 weeks at room temperature COMMENTARY Background Information The extraordinarily high affinity of avidin for biotin was first discovered as the explanation for a deficiency disease caused by ingestion of large numbers of raw eggs. In this disease, which is characterized by skin lesions and growth retardation, the essential vitamin biotin is depleted because it is sequestered by avidin in egg whites (György et al., 1941). The dissociation constant at each of the four biotinbinding sites of avidin is ∼10−15 M (Green, 1975), making the avidin-biotin interaction one of the strongest known noncovalent biological interactions. Retention of high-affinity binding after derivatization of the biotin carboxyl group with a wide variety of agents has led to the development of numerous assays involving the recovery or detection of biotintagged moieties with avidin or other high-affinity biotin-binding proteins described below (reviewed by Wilchek and Bayer, 1990 and Savage et al., 1992). Selection of biotinylation reagent There are a number of reagents that can be used for biotinylation (see Table 3.6.1). Selection of reagent depends on the intended uses for the biotinylated product. For instance, most enzyme activities are independent of protein modification on carbohydrate residues, which generally can be assumed to be innocuous; labeling of cysteinyls, however, which generally can be assumed to be innocuous, may disrupt disulfide bonds that are important for structure or activity, so biotinylation of cysteinyls is not recommended for extracellular proteins. Biotinylation of lysyl groups works well
with many proteins, including those on surfaces of intact cells, but it also may lead to loss of protein function. N-Hydroxysuccinimide (NHS) esters of biotin react readily at alkaline pH with any accessible, unprotonated primary amine, such as ε-amino groups of lysyl side chains in proteins (see Basic Protocol 1 and Basic Protocol 2). Because lysine is abundant in proteins, accounting for ∼7% of total amino acid residues in an average protein, NHS-biotin and its derivatives (see Table 3.6.1) are the most commonly used biotinylating agents. Modifications of NHS-biotin include addition of a sulfonate group to improve solubility in water (sulfoNHS-biotin), insertion of a long-chain (LC) spacer arm between the biotin moiety and the reactive ester to increase the availability of the biotin at the protein surface (NHS-LC-biotin), and addition of a cleavable site, such as a disulfide bond (NHS-SS-biotin), to permit release of the protein from biotin using reducing agents such as 2-mercaptoethanol (2-ME) or dithiothreitol (DTT). A cleavable disulfide bond is particularly useful for eluting biotinylated proteins from an affinity matrix containing covalently bound avidin because the exceptionally high affinity (Kd ∼10−15 M) and correspondingly low off-rate of the biotin-avidin interaction complicates elution with chaotropes (but see below). Several commercially available reagents (see Table 3.6.1) permit biotinylation of proteins on cysteinyl residues. At physiological pH (pH 7.5 to 8.0), iodoacetyl-LC-biotin, biotinLC-maleimide, maleimidopropionyl biocytin, biotin-HPDP, and biotin-maleimide all react
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primarily with free sulfhydryl groups on reduced cysteines. For cytoplasmic proteins, in particular, labeling with cysteine-directed reagents may permit retention of functions lost after derivatization of lysines. However, these reagents may not be advisable for labeling extracellular or membrane proteins with conformations stabilized by disulfide bonds because reduction of at least some disulfides with thiol-reducing agents (e.g., 50 mM DTT or 100 mM 2-ME) is generally necessary for efficient labeling (see Support Protocol 1). Biotin hydrazides (see Table 3.6.1), especially those with a six-carbon spacer (biotinLC-hydrazide), can be excellent reagents for biotinylating galactose and N-acetylgalactosamine residues on extracellular and transmembrane proteins (see Basic Protocol 4). The biotin hydrazide reagents covalently bind to aldehydes that have been generated from vicinal carbohydrate hydroxyls (cis-diols) by mild chemical or enzyme oxidation. The resulting hydrazone bonds are stable between pH 2 and 10. Sodium m-periodate provides a fast, effective means for oxidizing cis-diols on many soluble proteins, including immunoglobulins. However, the reagent also can react with serines, threonines, and tyrosines. Consequently, periodate oxidation sometimes affects protein biological activity and often alters migration on SDS–polyacrylamide gels. If these side effects cannot be tolerated (e.g., when identifying cellsurface proteins by biotinylation), then galactose and N-acetylgalactosamine residues should be exposed and oxidized by treating the cells on membranes with a combination of neuraminidase and galactose oxidase.
Biotinylation of Proteins in Solution and on Cell Surfaces
Detection of biotinylated proteins A large number of biotinylated proteins and biotin-detection reagents are commercially available, either as individual compounds or in the form of kits. For instance, biotinylated dipalmitoyl-L-α-phosphatidylethanolamine (Pierce) can be used as a phospholipid tracer in membrane reconstitution studies. Also, biotinylated insulin (Sigma), testosterone (Sigma), and lectins (Pierce or Vector Labs) can be used to identify and/or purify their cellular receptors. Avidin-biotin complex (ABC) detection kits (e.g., Vectastain ABC kit, Vector Labs, and ImmunoPure ABC kit, Pierce; see Support Protocol 2) amplify even weak signals by taking advantage of the multivalency of avidin for biotin and the incorporation of multiple biotins into most biotinylated antibodies and enzymes. Other, similar kits from Vector Labs and Pierce
permit visualization of biotinylated proteins on fixed tissue, thin microscopic sections, and microtiter plate wells, allowing use of the biotinavidin high-affinity interaction in immunofluorescence microscopy, electron microscopy, and enzymatic assays, as well as for protein size determinations. The molecular masses of experimentally obtained biotinylated proteins are best estimated by comparison with the sizes of biotinylated protein molecular weight markers (Bio-Rad or Vector Labs). Because the addition of one or a few of the small biotin molecules does not significantly increase the mass of most proteins, the sizes of biotinylated protein standards are considered identical to those of prestained or unstained markers. Additionally, the markers are visualized with the same detection methods as the proteins of interest, permitting direct size comparisons. Nonspecific, biotin-independent binding to avidin can usually be controlled by employing buffers of pH 10.0 to 10.5, which is near the avidin isoelectric point (pI 10.0). Use of such buffers minimizes electrostatic interactions but causes little or no diminution of the avidin-biotin affinity. When high-pH buffers are undesirable, avidin should be replaced either by a neutralized, deglycosylated form of avidin (Neutralite Avidin from Molecular Probes or Southern Biotechnology, or NeutrAvidin from Pierce) or by streptavidin (Pierce, Sigma, and Vector Labs). Streptavidin is a tetrameric protein from the culture broth of Streptomyces avidinii; it binds four biotin molecules per mole with an affinity similar to that of avidin. Both neutralized avidin (pI 6.3) and streptavidin (pI 5.0) exhibit fewer nonspecific electrostatic interactions at physiological pH and ionic strength. Streptavidin does, however, contain a sequence that causes biotin-independent binding to integrins and related cell-surface fibronectin receptors under certain conditions (Alon et al., 1990). Many commercially available derivatives of streptavidin (Pierce, Sigma, and Vector Labs) and neutralized avidin (Pierce and Southern Biotechnology) may be used in microscopic and enzymatic assays as a replacement for underivatized avidin. Recovery of biotinylated proteins Paradoxically, the high affinity of the biotin-avidin interaction can be a disadvantage during affinity purification of biotinylated proteins. Biotinylated proteins bind tightly to avidin affinity matrices (Molecular Probes, Pierce, Sigma, or Vector Labs) under almost
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any physiological conditions, but the high-affinity binding is reversed only by harsh denaturants, such as 8 M guanidine⋅HCl, pH 1.5, or by boiling with 2% (w/v) SDS. Further, the large positive charge on avidin (pI 10.0) and its carbohydrate residues promote nonspecific binding of many negatively charged proteins and some lectins to avidin affinity columns. For this reason, matrices containing bound NeutrAvidin (Pierce) or streptavidin (Molecular Probes, Pierce, Sigma, and Vector Labs) are often used for the affinity purification of biotinylated proteins, despite the significantly greater expense of these matrices. For preparative purification of biotinylated proteins by affinity chromatography, whether it be on columns containing bound avidin, streptavidin, or neutralized avidin, there are two approaches. The first approach is to accept that the biotin-avidin interaction is essentially irreversible and to attach the biotin label to a dispensable moiety that is left on the column after eluting the protein of interest. For instance, biotinylated monoclonal antibodies can be used to purify individual membrane antigens (Updyke and Nicolson, 1984), and biotinylated insulin and other ligands have been used to purify their specific receptors by avidin or streptavidin affinity chromatography (Kohanski and Lane, 1985/1986; Kraehenbuhl and Bonnard, 1990). Similarly, disulfide bond reduction (see Support Protocol 1) will release proteins that have been biotinylated with NHSSS-biotin (Chia and Luna, 1989) or with biotin-HPDP (see Table 3.6.1). The second approach to purifying native proteins is to moderate the avidin-biotin binding affinity by modifying either the bound avidin or the biotin. For example, “monomeric” avidin columns (Pierce or Sigma), generated by treating immobilized avidin with 0.1 M guanidine⋅HCl, pH 1.5, contain one lower-affinity biotin-binding site (Kd ∼5 × 10−8 M) per bound avidin molecule. After presaturating residual high-affinity sites on the column with excess free biotin, biotin-containing proteins can be bound to the columns and then eluted with either 0.1 M glycine, pH 2.0 to 2.8, or 2 mM free biotin (Henrikson et al., 1979; Kohanski and Lane, 1990). A converse strategy is to derivatize protein lysines with 2-iminobiotin-NHS (see Table 3.6.1), a guanido analog of NHS-biotin that exhibits a strong pH dependence in interactions with avidin and streptavidin (Heney and Orr, 1 98 1) . At p H 9.5, imin ob iotin binds avidin/streptavidin with characteristically high
affinity. However, proteins labeled with iminobiotin can be eluted from avidin columns with either 0.1 M acetic acid or 50 mM ammonium acetate/0.5 M NaCl, pH 4.0. Similarly, columns containing bound 2-iminobiotin (Molecular Probes, Pierce, or Sigma) can be used to purify avidin, streptavidin, and their derivatives. For further information on affinity chromatography, see Chapter 9.
Critical Parameters and Troubleshooting The most critical aspects of protein biotinylation are the storage and solubilization conditions for the biotinylating reagent. This is particularly critical with NHS derivatives, which contain succinimide ester groups that are very reactive with water. The NHS-biotins should be stored in the presence of desiccant at −20°C, as recommended by the suppliers, and the reagent bottles should be warmed completely to room temperature before opening to minimize condensation. Similarly, it is critical to mix the compounds with the protein to be labeled as soon as possible after dissolving the reagents in aqueous solutions. Most other parameters are less critical. Although the rate of protein biotinylation is influenced by temperature, short incubations (e.g., 30 min to 1 hr) at either 0° to 4°C or room temperature are usually satisfactory. The extent of biotinylation is determined by the amounts of reagents mixed and the reaction time. Biotin-to-protein molar ratios from 1:1 to 60:1 and reaction times up to 18 hr have been described. Because biotinylation is directed at selected groups in proteins (e.g., labeling ranges from a single cysteinyl or carbohydrate to some or all lysyls, which make up 7 of 100 residues in a typical protein), the choice of biotinylating reagent is often more critical than the reaction conditions. If an enzyme loses activity during biotinylation and if a change of biotinylating reagent is contraindicated, one strategy is to protect the enzyme’s active site by adding excess substrate during the biotinylation reaction. The medium in which biotinylation takes place must be free of components that will interfere with the reaction chemistry. Coupling buffers for NHS-biotin reagents (see Basic Protocols 1 and 2) must be free of ingredients with primary amines. Excess thiol-reducing agent (see Support Protocol 1) must be removed for efficient use of sulfhydryl-directed reagents (see Basic Protocol 3), and excess cis-diol-oxidizing agent must be eliminated for labeling of protein carbohydrates (see Basic Protocol 4).
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Biotinylation of Proteins in Solution and on Cell Surfaces
Finally, pH is an important factor. Lysyl-directed reagents require unprotonated amines (e.g., pH 7.5 to 8.5), whereas maleimide-based thiol-directed reagents undergo side reactions with amines at pH 7.5 or above and thus need more acid conditions (e.g., pH 6.5 to 7.5) if specificity for cysteines is required. Carbohydrate-directed reagents also require acid conditions (e.g., pH 5.5) for specificity. Reagent solubility may also be a factor. If the protein of interest is sensitive to organic solvents, such as DMF and DMSO, use the water-soluble sulfo-NHS-biotin for labeling lysyl groups. If highest affinity binding is not needed, biotinylating reagents lacking the LC spacer may be more water soluble. When biotinylating a subset of proteins in a complex mixture—for example, when determining which plasma membrane proteins are accessible to a cell-surface biotinylating reagent—it is useful to include two controls in each experiment. First, some cells should always be carried through the labeling and fractionation procedure in the absence of biotinylating reagent, in order to identify endogenously biotinylated proteins, which are involved in many intracellular carboxylation reactions. Second, to help identify proteins that may be exposed by cell breakage during manipulation, cytoplasmically oriented proteins should be labeled with biotinylating reagent either after deliberate mechanical lysis in the presence of 0.2% (v/v) Triton X-100 or after plasma membrane purification. The group of biotinylated cytoplasmically oriented proteins generated with this control should differ dramatically from that observed after biotinylation of unbroken cells. If the two SDS-PAGE profiles appear similar, then the biotinylating procedure should be repeated under conditions that are less conducive to cell lysis (e.g., biotinylate cells that are still attached to tissue culture plastic). Because proteins differ dramatically in the number of surface-accessible reactive groups and how they are affected by labeling with a foreign tag such as biotin, it is hard to predict which biotinylating compound will work best for a particular protein. However, biotinylation chemistries are similar to those used for a variety of other compounds, including fluorescent labels. Thus, if a literature reference exists for fluorescently labeling a protein of choice, the most expeditious strategy is to obtain a biotinylating compound with the same reactive group (i.e., succinimide ester, maleimide, or hydraz-
ide moiety) and to follow the published reaction conditions for labeling the protein. If no information is available about accessible groups or about the functional consequences of a particular derivatization strategy, it is a good idea to try different types of biotinylating reagents at different reagent-to-protein ratios. Reagent/protein molar ratios of 1:1 and 20:1 are usually good starting points. Preliminary characterization of protein functionality should suggest which of the reaction chemistries is superior and whether a higher or lower stoichiometry yields the most efficient labeling.
Anticipated Results The biotinylation protocols in this unit should biotinylate most proteins. The extent of reaction and the degree to which the protein retains its endogenous structure and activity will vary but may be predictable if other investigators have used a similar chemistry to derivatize the protein with a “tag” other than biotin. Soluble proteins biotinylated by these procedures have been used as ligands in binding assays in vitro and as tracers for cellular processes including endocytosis, membrane trafficking, and cytoskeletal remodeling (reviewed in Savage et al., 1992, and Wilchek and Bayer, 1990). Cell surface–biotinylated membrane proteins can be used similarly as markers for plasma membrane proteins in biochemical fractionations and in membrane recycling assays. In addition, biotinylated protein(s) can be purified, either alone or in combination with receptors or other binding partners, on avidin or streptavidin affinity columns.
Time Considerations Labeling reactions involving succinimide esters (NHS-biotin and analogs) and those involving biotin-hydrazides are relatively rapid and usually complete within 1 to 2 hr at 4°C. Reactions of sulfhydryls with maleimides and iodoacetamides are slower, particularly when carried out in lower-pH buffers that optimize reaction selectivity of protein sulfhydryls over amino groups. Although these reactions can be terminated after only 2 to 4 hr, reaction extents are usually increased by longer incubations (e.g., overnight at 4°C). Similarly, depending on the choice of separation technique, removal of unincorporated biotinylating reagent from labeled protein can be either rapid or slow, with gel-filtration chromatography and centrifugal filtration requiring
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no more than 1 to 2 hr, whereas dialysis may take 1 to 3 days. Separation steps also are often dictated more by the properties of the protein being labeled than by investigator preference. Some proteins become irreversibly aggregated if subjected to centrifugal filtration; other proteins are denatured or proteolyzed during prolonged dialysis. Here again, it helps to know what other investigators have done successfully with the particular protein under investigation.
Literature Cited Alon, R., Bayer, E.A., and Wilchek, M. 1990. Streptavidin contains an RYD sequence which mimics the RGD receptor domain of fibronectin. Biochem. Biophys. Res. Commun. 170:1236-1241. Bayer, E.A., Ben-Hur, H., and Wilchek, M. 1988. Biocytin hydrazide—A selective label for sialic acids, galactose, and other sugars in glycoconjugates using avidin-biotin technology. Anal. Biochem. 170:271-281. Chia, C.P. and Luna, E.J. 1989. Phagocytosis in Dictyostelium discoideum is inhibited by antibodies directed primarily against common carbohydrate epitopes of a major cell-surface plasma membrane glycoprotein. Exp. Cell Res. 181:11-26. Goodloe-Holland, C.M. and Luna, E.J. 1987. Purification and characterization of Dictyostelium discoideum plasma membranes. Methods Cell Biol. 28:103-128. Green, N.M. 1975. Avidin. Adv. Protein Chem. 29:85-133. Gyöargy, P., Rose, C.S., Eakin, R.E., Snell, E.E., and Williams, R.J. 1941. Egg-white injury as the result of nonabsorption or inactivation of biotin. Science 93:477-478. Heney, G. and Orr, G.A. 1981. The purification of avidin and its derivatives on 2-iminobiotin-6aminohexyl-Sepharose 4B. Anal. Biochem. 114:92-96.
Henrikson, K.P., Allen, S.H.G., and Maloy, W.L. 1979. An avidin monomer affinity column for the purification of biotin-containing enzymes. Anal. Biochem. 94:366-370. Ingalls, H.M., Goodloe-Holland, C.M., and Luna, E.J. 1986. Junctional plasma membrane domains isolated from aggregating Dictyostelium discoideum amebae. Proc. Natl. Acad. Sci. U.S.A. 83:4779-4783. Kohanski, R.A. and Lane, M.D. 1985/1986. Receptor affinity chromatography. Ann. N.Y. Acad. Sci. 447:373-385. Kohanski, R.A. and Lane, M.D. 1990. Monovalent avidin affinity columns. Methods Enzymol. 184:194-200. Kraehenbuhl, J.-P. and Bonnard, C. 1990. Purification and characterization of membrane proteins. Methods Enzymol. 184:629-641. Mock, D.M. 1990. Sequential solid-phase assay for biotin based on 125I-labeled avidin. Methods Enzymol. 184:224-233. O’Shannessy, D.J., Doberson, M.J., and Quarles, R.H. 1984. A novel procedure for labeling immunoglobulins by conjugation to oligosaccharide moieties. Immunol. Lett. 8:273-277. Savage, M.D., Mattson, G., Desai, S., Nielander, G.W., Morgensen, S., and Conklin, E.J. 1992. Avidin-Biotin Chemistry: A Handbook. Pierce Chemical Co., Rockford, Ill. Updyke, T.V. and Nicolson, G.L. 1984. Immunoaffinity isolation of membrane antigens with biotinylated monoclonal antibodies and immobilized streptavidin matrices. J. Immunol. Methods 73:83-95. Wilchek, M. and Bayer, E.A. (eds.) 1990. Avidinbiotin technology. Methods Enzymol. 184:1-746.
Contributed by Elizabeth J. Luna Worcester Foundation for Biomedical Research Shrewsbury, Massachusetts
Enzymatic Manipulation of DNA and RNA
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Metabolic Labeling with Amino Acids
UNIT 3.7
Metabolic labeling techniques are used to study the biosynthesis, processing, intracellular transport, secretion, degradation and physical-chemical properties of proteins. In this unit, protocols are described for metabolically labeling mammalian cells with radiolabeled amino acids (Table 3.7.1). Cells labeled using these procedures are suitable for analysis by immunoprecipitation, characterization of cellular proteins, analysis of protein trafficking, and one- and two-dimensional gel electrophoresis (UNITS 10.1-10.4). The first protocols describe pulse-labeling (10 to 30 min) of mammalian cells in suspension with [35S]methionine (see Basic Protocol), and necessary modifications for adherent mammalian cells (see Alternate Protocol 1). Alternate protocols are also presented for pulse-chase analysis (see Alternate Protocol 2) and long-term (“steady state”) labeling (6 to 32 hr; see Alternate Protocol 3). This is followed by conditions for metabolic labeling of cells with radiolabeled amino acids other than [35S]methionine (see Alternate Protocol 4). The degree of label incorporation can be determined by precipitation with trichloroacetic acid (TCA; see Support Protocol). NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly. NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4. Table 3.7.1
Radiolabeled Amino Acids Used in Metabolic Labeling of Proteins
Amino acida Leucine Serine Glutamic acid Lysine Alanine Valine Glycine Threonine Arginine Aspartic acid Proline Glutamine Phenylalanine Tyrosine Asparagined Cysteine Isoleucine Histidine Methionine Tryptophan
Frequency (%)b
Radioisotope
10.4 8.1 7.3 7.0 7.0 6.2 5.7 5.6 5.0 4.9 4.9 4.5 4.5 3.6 3.5 3.4 2.9 2.5 1.8 1.3
3H 3H 3H 3H 3H 3H 3H 3H 3H 3H 3H 3H 3H 3H
— 35S 3H 3H 35S 3H
Specific activity (Ci/mmol)
Commentsc
5-190 5-40 15-80 40-110 10-85 10-65 10-60 5-25 30-70 10-50 15-130 20-60 15-140 15-60 — >800 30-140 30-70 >800 20-30
E T, I T, I E T E T, I E E T, I — T, I E — — — E E E E
aAll amino acids in this table are in the L configuration. bFrequency of amino acid residues in proteins, taken from Lathe (1985). cE, essential amino acids; T, amino acids that are modified by transamination (Coligan et al., 1983);
I, amino acids that are converted by cells to other amino acids. See Critical Parameters for more details. dAsparagine is difficult to label (Coligan et al., 1983).
Detection and Assay Methods Contributed by Juan S. Bonifacino Current Protocols in Protein Science (1999) 3.7.1-3.7.10 Copyright © 1999 by John Wiley & Sons, Inc.
3.7.1 Supplement 17
SAFETY PRECAUTIONS FOR WORKING WITH 35S-LABELED COMPOUNDS When working with radioactive materials, take appropriate precautions to avoid contamination of the experimenter and the surroundings. Carry out the experiment and dispose of wastes in an appropriately designated area, following guidelines provided by the local radiation safety officer (also see APPENDIX 2B). Solutions containing 35S-labeled compounds have been found to release volatile radioactive substances (Meisenhelder and Hunter, 1988). In addition to the usual safety practices followed when handling radioactive materials (see APPENDIX 2B), some extra precautions should be taken when using 35S-labeled amino acids: 1. Vials containing 35S-labeled compounds should always be handled in a designated fume hood equipped with an activated charcoal filter. This includes thawing the solution, opening the vial, and adding the radiolabeled amino acid to the labeling medium. Before opening, vials should be vented with a needle attached to a syringe packed with activated charcoal. Tissue culture hoods should not be used for this purpose, as they are likely to become contaminated. 2. Minimize exposure of 35S-containing solutions to the air. 3. Use baths, incubators, and centrifuges designated for work with radioactive materials. Place a tray containing a layer of activated charcoal in the CO2 incubator to reduce the amount of 35S-labeled compounds released to the air during cell labeling. Alternatively, filters impregnated with activated carbon (β-Safe, Schleicher & Schuell) can be attached to the covers of tissue culture dishes. 4. Monitor areas used during labeling by conducting wipe tests. 5. Dispose of solid and liquid 35S waste quickly and with appropriate precautions. BASIC PROTOCOL
Metabolic Labeling with Amino Acids
PULSE-LABELING OF CELLS IN SUSPENSION WITH [35S]METHIONINE Pulse-labeling of proteins is performed by incubating cells for short periods (≤30 min) in culture medium containing a radiolabeled amino acid. The conditions described below are optimized for labeling times of 10 to 30 min, but the same procedure can be used for periods up to 6 hr, although fewer cells and/or more labeling medium should be used for optimal results. Very short pulses (<10 min) may require special conditions (see Critical Parameters). For labeling times >6 hr, see Alternate Protocol 3. [35S]methionine is the radiolabeled amino acid of choice for metabolic labeling of proteins because of its high specific activity (>800 Ci/mmol) and ease of detection. A potential disadvantage of [35S]methionine is its low abundance (∼1.8% of the average amino acid composition; see Table 3.7.1). For proteins that contain little (e.g., only one) or no methionine, other radiolabeled amino acids should be used (see Alternate Protocol 4). Materials [35S]L-Methionine (>800 Ci/mmol) or 35S-labeled protein hydrolyzate (>1000 Ci/mmol) Pulse-labeling medium (see recipe), warmed to 37°C Cell suspension (e.g., Jurkat, RBL, K562, BW5147, or T or B cell hybridomas), grown in a humidified 37°C, 5% CO2 incubator or prepared from tissues (e.g., lymphocytes) PBS (APPENDIX 2E), ice cold Vacuum aspirator with trap for liquid radioactive waste
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Additional reagents and equipment for TCA precipitation (optional; see Support Protocol) 1. Thaw [35S]methionine at room temperature and prepare a 0.1 to 0.2 mCi/ml working solution in prewarmed (37°C) pulse-labeling medium. CAUTION: Volatile 35S-containing compounds can be released during the labeling procedure (refer to Safety Precautions and APPENDIX 2B). Keep [35S]methionine-containing medium in a tightly capped tube in a 37°C water bath until use. Do not let it sit >60 min at 37°C.
2. Harvest 0.5–2 × 107 cells in suspension by centrifuging 5 min at 300 × g, room temperature. 3. Aspirate supernatant and resuspend cells by gently tapping the bottom of the tube. Add 10 ml prewarmed pulse-labeling medium. Mix by inversion of the tube and centrifuge 5 min at 300 × g, room temperature. Repeat this step once. 4. Resuspend cells at 5 × 106 cells/ml in prewarmed pulse-labeling medium and incubate 15 min in a 37°C water bath to deplete intracellular pools of methionine. Invert tube periodically to resuspend cells. 5. Centrifuge cells 5 min at 300 × g, room temperature, and aspirate supernatant. 6. Resuspend cells in 2 ml [35S]methionine working solution (step 1). Cap tube tightly. Incubate cells 10 to 30 min (see Critical Parameters) in a 37°C water bath, resuspending frequently by gentle inversion of the tube. Alternatively, use a rotator placed in a 37°C incubator.
7. Centrifuge cells 5 min at 300 × g, 4°C, and aspirate supernatant. Resuspend with gentle swirling in 10 ml ice-cold PBS and repeat centrifugation. CAUTION: The medium and wash are radioactive—follow applicable safety regulations for disposal. Because of the short times and high concentrations of radiolabeled amino acids employed, pulse-labeling may not result in complete depletion of label from the medium. If this is the case, the labeling mixture can be reused. Collect the labeling medium containing [35S]methionine or 35S-labeled protein hydrolyzate and filter carefully through a 0.45-ìm filter unit. Estimate the percentage of unincorporated radioactivity by scintillation counting aliquots of the labeling mixture before and after labeling the cells. Store the labeling mixture up to 2 months frozen at −20°C.
8. Optional: Determine amount of label incorporation by trichloroacetic acid (TCA) precipitation (see Support Protocol 1). Labeled cells are now ready for the desired processing and analysis. If cell pellets cannot be processed immediately, they can be kept on ice for a few hours or frozen at −80°C for several days. Thaw frozen cell pellets on ice before analysis. In most cases, freezing cells will not affect the biochemical properties of the proteins, but freezing and thawing cell extracts after solubilization with detergents (APPENDIX 1B) can cause dissociation of multisubunit complexes or proteolysis of labeled proteins.
Detection and Assay Methods
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ALTERNATE PROTOCOL 1
PULSE-LABELING OF ADHERENT CELLS WITH [35S]METHIONINE Labeling of adherent cells is essentially the same as described for cells in suspension except that cells are labeled while attached to a dish. As with suspended cells, adherent cells are pulse-labeled 10 to 30 min (see Basic Protocol introduction). Alternatively, some adherent cells can be detached from plates by incubation for 10 min at 37°C with 10 mM EDTA in PBS (APPENDIX 2E) and labeled as a suspension (see Basic Protocol). This is particularly advantageous when labeling cells for pulse-chase experiments, because it simplifies handling of multiple samples and allows more uniform labeling of the cells. Additional Materials (also see Basic Protocol) Adherent cells (e.g., HeLa, NRK, M1, COS-1, CV-1, or fibroblasts or endothelial cells in primary culture) 100-mm tissue culture dishes 1. Grow adherent cells to 80% to 90% confluency in 100-mm tissue culture dishes. Depending on the cell type, a confluent 100-mm dish will contain 0.5–2 × 107 cells.
2. Prepare [35S]methionine working solution (see Basic Protocol, step 1). 3. Aspirate culture medium from the dishes and wash twice by gently swirling with 10 ml prewarmed (37°C) pulse-labeling medium, aspirating medium after each wash. 4. Add 5 ml prewarmed pulse-labeling medium and incubate 15 min in a humidified 37°C, 5% CO2 incubator to deplete intracellular pools of methionine. 5. Remove medium from cells, add 2 ml [35S]methionine working solution (from step 2), and incubate 10 to 30 min in a CO2 incubator. If necessary, as little as 1 ml of labeling medium per plate can be used. It is critical, however, that the plates sit on a perfectly horizontal surface during incubation to avoid drying of the cell monolayer.
6. Remove medium from cells. Wash once with 10 ml ice-cold PBS and remove PBS. CAUTION: The medium and wash are radioactive—follow applicable safety regulations for disposal. If significant detachment of cells occurs during labeling, it may be necessary to scrape the cells carefully in the [35S]methionine-containing medium and transfer the suspension to a 15-ml centrifuge tube before centrifuging and washing with PBS.
7. Add 10 ml ice-cold PBS and scrape cells carefully with either a disposable plastic scraper or a rubber policeman. 8. Transfer the suspension to a 15-ml centrifuge tube, centrifuge 5 min at 300 × g, 4°C, and discard supernatant. 9. Optional: Determine amount of label incorporation by trichloroacetic acid (TCA) precipitation (see Support Protocol 1). Labeled cells are now ready for the desired processing and analysis (see Basic Protocol, step 8 annotation, if cells are not to be used immediately).
Metabolic Labeling with Amino Acids
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PULSE-CHASE LABELING OF CELLS WITH [35S]METHIONINE Pulse-chase protocols are used to analyze time-dependent processes, such as posttranslational modification, transport, secretion, or degradation of newly synthesized proteins. Cells in suspension or adherent cells are pulse-labeled with [35S]methionine (see Basic Protocol and Alternate Protocol 1), after which they are incubated (chased) in complete medium containing excess unlabeled methionine, as described below.
ALTERNATE PROTOCOL 2
Additional Materials (also see Basic Protocol and Alternate Protocol 1) Chase medium (see recipe), 37°C 1. Prepare 0.5–2 × 107 cells per sample per time point, and pulse-label 10 to 30 min with 2 ml of 0.1 to 0.2 mCi/ml [35S]methionine per 0.5–2 × 107 cells (see Basic Protocol, steps 1 to 6, or see Alternate Protocol 1, steps 1 to 5). 2. Remove the [35S]methionine working solution, wash once with 10 ml of 37°C chase medium, and add 10 ml of 37°C chase medium. Rapid termination of the labeling reaction can be achieved by adding two times the volume of chase medium containing excess unlabeled methionine (15 mg/liter) directly to the labeling mixture. Final concentration of cells in suspension should be 2 × 106 cells/ml for chases ≤2 hr, or 0.5 × 106 cells/ml for chases >2 hr. For adherent cells, add 10 ml/100-mm dish.
3. Incubate for the desired time at 37°C. Incubate cell suspensions with rotation in tightly capped tubes. Incubate adherent cells in a CO2 incubator. 4a. For cells in suspension: Collect cells by centrifuging 5 min at 300 × g, 4°C. The supernatant can be discarded or can be collected for analysis of proteins that are secreted or shed into the medium.
4b. For adherent cells: Scrape off adherent cells and transfer to 15-ml centrifuge tubes. Centrifuge 5 min at 300 × g, 4°C, and either save or discard the supernatant as in step 4a. 5. Optional: Determine amount of label incorporation by trichloroacetic acid (TCA) precipitation (see Support Protocol 1). Labeled cells are now ready for the desired processing and analysis (see Basic Protocol, step 8 annotation, if cells are not to be used immediately).
LONG-TERM LABELING OF CELLS WITH [35S]METHIONINE Long-term labeling refers to continuous metabolic labeling of cells for periods of 6 to 32 hr. Long-term labeling is particularly advantageous when studying proteins that are synthesized at low rates. It is also used to accumulate mature labeled proteins, rather than biosynthetic precursors, for characterization of their properties. Steady-state labeling is a form of long-term labeling in which cells are incubated with the radiolabeled amino acid until the rates of synthesis and degradation of the radiolabeled proteins are equal. Steady-state labeling allows calculation of the stoichiometry of subunits within a protein complex, provided that the primary structure of the subunits is known. In these procedures, unlabeled methionine is added to the medium to maintain cell viability and to sustain incorporation of label for the duration of the experiment. The amount of unlabeled methionine added depends on factors such as the length of the labeling period and the cell density. Media containing between 5% and 20% the normal amount of unlabeled methionine are generally used. The conditions described below are suitable for overnight (∼16 hr) labeling of cells in suspension or adherent cells.
ALTERNATE PROTOCOL 3
Detection and Assay Methods
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Additional Materials (also see Basic Protocol and Alternate Protocol 1) Long-term labeling medium (see recipe), warmed to 37°C 75-cm2 tissue culture flask 1. Prepare a 0.02 to 0.1 mCi/ml [35S]methionine working solution in prewarmed (37°C) long-term labeling medium (see Basic Protocol, step 1). For cell suspensions: 2a. Prepare and wash cells once with prewarmed long-term labeling medium (see Basic Protocol, steps 2 and 3). 3a. Resuspend 0.5–2 × 107 cells in 25 ml of 0.02 to 0.1 mCi/ml [35S]methionine working solution and transfer to a 75-cm2 tissue culture flask. For adherent cells: 2b. Grow 0.5–2 × 107 adherent cells in a 75-cm2 tissue culture flask (80% to 90% confluency) in a CO2 incubator. Wash once with prewarmed long-term labeling medium (see Alternate Protocol 1, step 3). 3b. Add 25 ml of 0.02 to 0.1 mCi/ml [35S]methionine working solution to each 75-cm2 flask. 4. Tighten caps to prevent release of volatile 35S-labeled compounds. Incubate 16 hr in a CO2 incubator. 5. Wash cells and determine incorporation (see Basic Protocol, steps 7 and 8, or Alternate Protocol 1, steps 6 to 9). Labeled cells are now ready for the desired processing and analysis (see Basic Protocol, step 8 annotation, if cells are not to be used immediately). ALTERNATE PROTOCOL 4
METABOLIC LABELING WITH OTHER RADIOLABELED AMINO ACIDS In some instances, it may be necessary to label proteins with radiolabeled amino acids other than [35S]methionine—e.g., when the proteins have a low methionine content or have no methionine residues at all. In these cases, the best choices are [35S]cysteine or [3H]leucine. Cysteine residues are more abundant in proteins than methionine residues (3.4% versus 1.8%; Table 3.7.1), although cysteine is less stable. Formulations of [35S]cysteine that have high specific activity (>800 Ci/mmol) can be obtained from several companies. Leucine has the advantage of being the most frequent amino acid in proteins (10.4%; Table 3.7.1) and specific activities of [3H]leucine are the highest among 3H-labeled amino acids (up to 190 Ci/mmol). If proteins are known to be rich in a particular amino acid or if special methods are to be performed (e.g., radiochemical sequencing or multiple labeling), 3H-labeled amino acids other than [3H]leucine can be used. Label cells with these amino acids as described for [35S]methionine in the previous protocols. Substitute [35S]cysteine or 3H-labeled amino acids in the labeling medium. Use labeling medium lacking cysteine or any other respective amino acid. CAUTION: Solutions containing [35S]cysteine release volatile 35S-labeled compounds (refer to Safety Precautions).
Metabolic Labeling with Amino Acids
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TCA PRECIPITATION TO DETERMINE LABEL INCORPORATION In metabolic labeling experiments, it is often useful to monitor the incorporation of radioactivity into total cellular proteins. This can easily be achieved by precipitation with trichloroacetic acid (TCA) using BSA as a carrier protein. This protocol can be used at the end of the labeling procedures in each protocol.
SUPPORT PROTOCOL
Materials Labeled cell suspension (see Basic Protocol or Alternate Protocols 1 to 4) BSA/NaN3: 1 mg/ml BSA containing 0.02% (w/v) sodium azide (NaN3) 10% (w/v) TCA solution (see recipe), ice cold Ethanol Filtration apparatus attached to a vacuum line 2.5-cm glass microfiber filter disks (Whatman GF/C) CAUTION: TCA is extremely caustic. Protect eyes and avoid contact with skin when preparing and handling TCA solutions. 1. Add 10 to 20 µl of a labeled cell suspension to 0.1 ml BSA/NaN3. Place on ice. 2. Add 1 ml ice-cold 10% (w/v) TCA solution. Vortex vigorously and incubate 30 min on ice. 3. Filter the suspension onto 2.5-cm glass microfiber filter disks in a filtration apparatus under vacuum. 4. Wash the disks twice with 5 ml ice-cold 10% (w/v) TCA solution and twice with ethanol. Air dry 30 min. CAUTION: The wash fluids should be handled as mixed chemical/radioactive waste—follow applicable safety regulations for disposal (see APPENDIX 2B).
5. Spot the same volume of the radiolabeled cell suspension used in step 1 (10 to 20 µl) on a glass microfiber disk. Air dry. This disk will be used to measure the total amount of radiolabeled amino acid in the cell suspension.
6. Transfer disks from steps 4 and 5 to 20-ml scintillation vials, add 5 ml scintillation fluid, and measure the radioactivity in a scintillation counter. 7. Calculate the ratio of TCA-precipitable label to total radioactivity (i.e., the ratio of sample radioactivity in step 4 to the total radioactivity in step 5). REAGENTS AND SOLUTIONS Use deionized or distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
NOTE: Use sterile tissue culture technique to prepare these reagents. Chase medium Pulse-labeling medium (see recipe) containing 15 mg/liter unlabeled methionine or equivalent excess amount of other amino acid used for radiolabeling. Store up to 2 weeks at 4°C. Long-term labeling medium (90% methionine-free medium) Mix 9 vol pulse-labeling medium (see recipe), lacking methionine or other appropriate amino acid, with 1 vol chase medium (see recipe) containing the same amino acid. Store up to 2 weeks at 4°C.
Detection and Assay Methods
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Pulse-labeling medium Use supplemented Dulbecco’s modified Eagle medium (DMEM; see APPENDIX 3C but omit nonessential amino acids) or RPMI 1640, each lacking methionine or other specific amino acid, but containing 10% (v/v) FBS (dialyzed overnight against saline solution to remove unlabeled amino acids). Add 25 mM HEPES buffer (with pH adjusted to 7.4 with NaOH). Store up to 2 weeks at 4°C. The amino acid used to label cells is omitted from this medium (e.g., if [35S]methionine is employed, methionine-free DMEM or RPMI 1640 must be used). Specific amino acid–free media can be obtained from several tissue culture medium suppliers, or can be prepared (see APPENDIX 3C) from amino acid–free medium by adding individual amino acid components, omitting the one that will be used to label. A kit for the preparation of such media is available (Select-Amine from Life Technologies).
Trichloroacetic acid (TCA) solution, 10% (w/v) Prepare a 100% (w/v) TCA stock solution by dissolving the entire contents of a newly opened TCA bottle in water (e.g., dissolve the contents of a 500-g bottle of TCA in sufficient water to yield a final volume of 500 ml). Store up to 1 year at 4°C. Prepare 10% (w/v) TCA by dilution and store up to 3 months at 4°C. CAUTION: TCA is extremely caustic. Protect eyes and avoid contact with skin when preparing and handling TCA solutions.
COMMENTARY Background Information Metabolic labeling of cellular proteins is achieved by placing cells in a nutritional medium containing all components necessary for growth of cells in culture, except for one amino acid that is substituted by its radiolabeled form. The radiolabeled amino acids are transported across the plasma membrane by carrier-mediated systems and, once in the cytosol, are loaded onto tRNA molecules before being incorporated into newly synthesized proteins. Because metabolic labeling techniques use the metabolic machinery of the cell to incorporate radiolabeled amino acids, there are limitations on the type of radiolabeled amino acids that can be employed. The list of potential precursors is restricted to L-amino acids normally found in proteins, in which one or more atoms are substituted by a radioisotope (Table 3.7.1). Sulfuric amino acids, such as methionine and cysteine, are conveniently labeled with 35S. Most other amino acids can be obtained labeled with 3H.
Critical Parameters
Metabolic Labeling with Amino Acids
The protocols in this unit should be considered as models from which more specific methods can be designed. The conditions described here have been optimized for radiolabeling proteins that are expressed at low to moderate levels (104 to 105 copies per cell) and assume that the labeled cells will be used for immuno-
precipitation. This level of expression is characteristic of most endogenous membrane proteins, lumenal organellar proteins, signal transduction proteins, and transcription factors. More abundant proteins, such as cytoskeletal proteins or proteins expressed by infection or transfection, can be labeled with less radiolabeled amino acid and/or fewer cells. The choice of a particular labeling protocol and its modification will be aided by careful consideration of a number of parameters that influence the incorporation of radiolabeled amino acids into proteins, as discussed below. Selection of amino acid label Purified [35S]methionine of high specific activity (>800 Ci/mmol) can be obtained from several suppliers of radiolabeled amino acids. Protein hydrolyzates of Escherichia coli grown in the presence of [35SO4]2− (e.g., Tran35S-label from ICN Biomedicals or Expre35S35S from NEN Life Sciences) can be used as substitutes for [35S]methionine in metabolic labeling techniques. These preparations contain 35S distributed among several different compounds. In a typical batch, ∼70% of the radioactivity will be present in the form of [35S]methionine and ∼15% as [35S]cysteine; the remainder is other 35S-labeled compounds. Labeling with these radioactive protein hydrolyzates in methionine-free medium will result in incorporation of label only in methionine residues; use of
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methionine- and cysteine-free medium will result in labeling of both methionine and cysteine residues. Purified [35S]methionine should be used whenever certainty of labeling with only [35S]methionine is required. Another reason for using purified [35S]methionine is that it tends to be more stable and emit less volatile decomposition products than 35S-labeled protein hydrolyzates. In most cases, however, the relatively inexpensive radioactive protein hydroly zates (about one-third the cost of [35S]methionine) can be used instead of purified [35S]methionine. [35S]methionine preparations should be stored frozen at −80°C. Under these conditions, they are stable for at least 1 month; the half-life of 35S is 88 days. For proteins with one or no methionine residues but several cysteine residues, labeling with [35S]cysteine (available with specific activities >800 Ci/mmol) is a good option. 35Slabeled protein hydrolyzates are not good sources of [35S]cysteine for radiolabeling because only ∼15% of the 35S-labeled compounds are [35S]cysteine. Purified [3H]leucine (available with specific activities of up to 190 Ci/mmol) is a good alternative to 35S-labeled amino acids. The halflife of 3H is ∼12 years. The specific activities of other 3H-labeled amino acids range between 5 and 140 Ci/mmol (Table 3.7.1). Several problems can arise when using certain 3H-labeled amino acids due to their participation in metabolic pathways. The following problems must be considered when using tritiated amino acids for metabolic labeling of proteins. Nonessential versus essential amino acids. Cells are able to synthesize nonessential amino acids from other compounds. If the radiolabeled amino acids used are nonessential, their specific activity will be reduced by dilution with the endogenously synthesized amino acids. Therefore, essential amino acids (E in Table 3.7.1) should be favored for metabolic labeling. Transamination. The α-amino groups of many amino acids can be removed in reactions catalyzed by transaminases (T in Table 3.7.1). Deamination of the radiolabeled amino acids can be prevented by addition of the transaminase inhibitor (aminooxy)acetic acid during starvation and labeling of the cells (Coligan et al., 1983). Interconversion. Certain radiolabeled amino acids can be converted by cells into other amino acids (I in Table 3.7.1). This problem is of particular importance when one is attempt-
ing to determine the amino acid composition or sequence of radiolabeled proteins. Labeling time The experimental purpose, the turnover rate of the protein of interest, and the viability of the cells should all be considered when determining the length of time for labeling. If the purpose of the experiment is to identify or characterize a protein precursor, pulse-labeling (10 to 30 min) must be used (see Basic Protocol and Alternate Protocol 1). If the protein of interest has a low turnover rate or if it is necessary to accumulate a labeled mature product, a protocol for long-term labeling (6 to 32 hr) is more appropriate (see Alternate Protocol 3). If the biosynthesis, post-translational modification, intracellular transport, or fate of newly synthesized proteins is being analyzed, a pulsechase protocol should be used (see Alternate Protocol 2). The turnover rate of the protein of interest is an important parameter to be considered when determining the labeling time. Proteins with a high turnover rate are optimally labeled for short times. Longer times will result in increased labeling of other cellular proteins, causing a decrease in the relative abundance of the labeled protein of interest in the cell lysate. This could result in increased detection of nonspecific proteins in immunoprecipitates (see UNIT 3.8). Conversely, proteins that turn over slowly should be labeled for longer times. Incorporation of radiolabeled amino acids into proteins is directly proportional to the labeling time for a certain period, after which it tends to plateau. When all the limiting amino acid is consumed, protein synthesis ceases. The length of the initial phase of linear incorporation will vary with the concentration of the labeling amino acid and the density and metabolic activity of the cells. Concentration and specific activity of the radiolabeled amino acid Because radiolabeled amino acids are used in limiting amounts, their incorporation into proteins is directly proportional to their concentration in the labeling medium. If very short pulses are required, the concentration of labeled amino acid can be increased to compensate for the shorter labeling time. The incorporation of radiolabeled amino acids is also directly proportional to their specific activity. Addition of unlabeled amino acids, as is required in long-term labeling protocols, will result in reduced incorporation over short periods.
Detection and Assay Methods
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Cell density At low cell densities, the amount of labeled protein synthesized is directly proportional to the cell concentration. At high densities, however, incorporation will increase nonlinearly. Very concentrated cell samples (>5 × 107 cells/ml) can only be labeled for short periods (≤5 min) as rapid loss of metabolic activity and cell viability will occur due to acidification of the medium and accumulation of toxic metabolites. Conditions for very short pulses Very short pulses (2 to 10 min) are required to study post-translational modifications that occur shortly after synthesis. These include folding, disulfide bond formation, and early carbohydrate modifications of newly synthesized proteins. Proteins that are rapidly degraded after synthesis are also best studied with very short pulses. The protocol for very short pulses is similar to the Basic Protocol for pulselabeling, with the following modifications: (1) the pulse-labeling medium contains higher concentration of radiolabeled amino acid (up to 1 mCi/ml), (2) labeling is stopped by addition of ice-cold PBS containing 20 mM freshly prepared N-ethylmaleimide (NEM) to prevent oxidation of free sulfhydryls, and (3) if a chase is necessary, 1 mM cycloheximide is added to the chase medium to quickly stop the elongation of nascent polypeptide chains. For additional details on very short pulses, see Braakman et al. (1991). Temperature and pH Unless otherwise required for special conditions (e.g., heat shock, temperature-sensitive mutants), metabolic labeling should be performed at 37°C. For this reason, it is important that the labeling medium be warmed to 37°C before adding it to the cells. A dramatic reduction in incorporation occurs at room temperature. It is also important that the pH of the labeling medium be ∼7.4.
Anticipated Results A typical incorporation of labeled amino acid precursor after a 30-min pulse under the conditions described in this unit (see Basic Protocol and Alternate Protocols 1 and 2) is 5% to 20%. In the case of long-term labeling, the incorporation typically reaches 30% to 60%. If the labeled cells are used for immunoprecipitation (UNIT 3.8), followed by SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11), specific bands should be visible within 2 hr to 2 months of exposure.
Time Considerations It takes 1 to 2 hr to prepare cells and materials for labeling. The actual labeling time depends upon the protocol chosen—pulse-labeling takes 10 to 30 min and long-term labeling takes 6 to 32 hr. Washing and processing cells may take an additional 1.5 hr.
Literature Cited Braakman, I., Hoover-Litty, H., Wagner, K.R., and Helenius, A. 1991. Folding of influenza hemagglutinin in the endoplasmic reticulum. J. Cell Biol. 114:401-411. Coligan, J.E., Gates, F.T. III, Kimball, E.S., and Maloy, W.L. 1983. Radiochemical sequence analysis of metabolically labeled proteins. Methods Enzymol. 91:413-434. Lathe, R. 1985. Synthetic oligonucleotide probes deduced from amino acid sequence data. Theoretical and practical considerations. J. Mol. Biol. 183:1-12. Meisenhelder, J. and Hunter, T. 1988. Radioactive protein labelling techniques. Nature 335:120.
Key Reference Coligan et al., 1983. See above. Contains a detailed description of conditions used to metabolically label proteins with different radiolabeled amino acids.
Contributed by Juan S. Bonifacino National Institute of Child Health and Human Development, NIH Bethesda, Maryland
Metabolic Labeling with Amino Acids
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Current Protocols in Protein Science
Analysis of Selenocysteine-Containing Proteins The presence of selenium in protein as a component in the diet of mammals was not anticipated from its early history. For example, in the 1930s, selenium was recognized as a toxin in livestock that caused severe illness and, if left untreated, death (Franke, 1934a,b). Furthermore, this element was later thought to cause cancer in laboratory animals (reviewed in Combs and Combs, 1986). The overabundance of selenium in soil in certain regions of the United States was therefore considered dangerous, as various “selenium-accumulating” plants that concentrate this element in high levels were being ingested by livestock and other animals. A variety of procedures employing both chemical and biological methods was developed to extract selenium from the soil or to convert it to a form that was not readily available for living organisms. The image of selenium changed completely in the mid-1950s. It was quite surprising when this element was shown to be essential for anaerobic growth of Escherichia coli (Pinset, 1954) and for prevention of liver necrosis in rats (Schwarz and Foltz, 1957). In fact, selenium is now recognized as a micronutrient that is capable of preventing certain forms of heart disease and cancer, decreasing mortality from AIDS in the HIV-infected population, playing a role in sperm maturation, and participating in immunological response and in suppression of viral expression (reviewed in Gladyshev et al., 2000). Perhaps the most recognized role of selenium in human nutrition is its ability to serve as a cancer-preventive agent. Recent doubleblind clinical trials have demonstrated that supplementation of the human diet with a daily dose of 200 µg of selenium resulted in 63%, 53%, and 48% reduction in the incidence of prostate, colon, and lung cancers, respectively (Clark et al., 1996). The majority of these biological and biomedical effects of selenium are most certainly mediated by selenium-containing proteins. Selenium was initially found in protein as a natural and essential component of mammalian glutathione peroxidase (Flohe et al., 1973; Rotruck et al., 1973). This element is incorporated into glutathione peroxidase and most other natural selenoproteins as the amino acid selenocysteine (Sec) in response to specific UGA
codons. Selenocysteine is normally present at the active center of enzymes (Stadtman, 1996), and this critical location of selenium in selenoproteins has generated considerable interest in their characterization. This interest in selenoproteins has resulted in the development of specific methods for their detection and characterization. Moreover, methods have been developed to incorporate selenium into specific sites in proteins, and these artificially derived selenoproteins have been used to obtain unique structural, functional, and mechanistic information.
CHEMICAL FORMS OF SELENIUM IN PROTEINS Selenium may be incorporated into protein as either selenocysteine or selenomethionine, or post-translationally as a cofactor. Both forms may occur nonspecifically when selenium replaces sulfur in sulfur metabolic pathways or when selenium is introduced into a protein by biochemical manipulations. However, both forms may also occur at specific sites in protein if selenium is incorporated by biological processes that are specialized for using this element.
Nonspecific Occurrence of Selenium in Proteins Selenium and sulfur have a similar affinity for sulfur pathways. The ratio of sulfur to selenium in proteins is generally determined by the ratio of sulfur to selenium in the environment or in the biological medium. Such a ratio is normally on the order of 1000:1, so that the proportion of selenium in sulfur-containing compounds and proteins is insignificant. However, when the selenium concentration approaches that of sulfur, the incorporation of selenium may be significant, which may result in selenium toxicity. Selenium toxicity may also result from interactions of low-molecularweight selenium compounds and various cellular metabolic pathways. The ability of selenium to replace sulfur in metabolic pathways has been used extensively to study the role of sulfur in proteins and metabolites (Table 3.8.1). Selenium-containing analogs of sulfur-containing cofactors and metabolites were used to characterize the active center structure and the kinetic properties of several proteins. For example, the nature of the
Contributed by Vadim N. Gladyshev and Dolph L. Hatfield Current Protocols in Protein Science (2000) 3.8.1-3.8.18 Copyright © 2000 by John Wiley & Sons, Inc.
UNIT 3.8
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Table 3.8.1 Overview of Methods to Obtain and Characterize Selenium Analogs of Sulfur-Containing Biological Compounds and Proteins
Method to replace S with Se
Method of analysis
Purpose
Replacement of all Met with Se-Met
X-ray crystallography Kinetic analyses
Replacement of all Cys with Se-Cys
Kinetic analyses Mass spectrometry
Targeted biosynthetic incorporation of Se-Cys in place of any other amino acid
Kinetic analyses Structural studies X-ray absorption spectroscopy EPR spectrometrya
To incorporate heavy atom into protein To solve a phase problem To characterize function of all Met To analyze disulfide bonds To test catalytic roles of all Cys To test redox roles of all Cys To analyze a particular Cys residue (redox and catalytic functions, participation in disulfide bond formation) To identify its direct ligands and to study its coordination to paramagnetic species To test catalytic functions of Ser or Cys To test redox functions of Ser or Cys To analyze coordination spheres of Ser or Cys
Chemical replacement of S (or O) EPR spectrometry with Se NMR spectrometry X-ray absorption spectroscopy Kinetic analyses Chemical synthesis of Kinetic analyses Se-containing compounds and Electrochemical analyses peptides NMR spectrometry X-ray crystallography
To test biological activity of a peptide, compound, cofactor, or biofactor To characterize its catalytic, redox, structural, and functional properties
aEPR spectrometry, electron paramagnetic resonance spectrometry.
Analysis of SelenocysteineContaining Proteins
thiol-binding site within the metal cluster active site of nitrogenase, the iron-molybdenum cofactor, was determined by the Se K-edge X-ray absorption spectroscopy of the selenol-substituted cofactor (Conradson et al., 1994). In addition, a selenium analog of lipoic acid was used to determine the kinetic parameters of selenolipoated proteins in a glycine cleavage system (Fujiwara et al., 1997). The electronic structure of iron-sulfur clusters in ferredoxins was also characterized by nuclear magnetic resonance (NMR) spectroscopy after replacement of sulfur with selenium (Bertini et al., 1993). One of the most advantageous applications of replacing sulfur with selenium has been the use of E. coli methionine auxotrophs grown in media in which selenomethionine replaced methionine. In this system, the methionine residues are quantitatively replaced in overexpressed proteins with selenomethionine residues. The presence of multiple selenium atoms in selenomethionine-containing proteins eliminates the step of obtaining heavy-atom derivatives that are otherwise required to solve a protein crystal structure. The developed multiwavelength anomalous diffraction (MAD) phasing method is now a common technique for determination of X-ray structures of proteins (Hendrickson et al., 1990; Hendrickson, 1991). This
technique, however, cannot be applied to natural selenoproteins, as these proteins usually contain only a single selenium atom. Incorporation of selenomethionine in place of methionine in protein is not very toxic, because methionines rarely occupy positions that are important to protein function. However, similar strategies that attempt to obtain proteins in which all cysteines are replaced with selenocysteines are usually not successful. This may be due to the fact that cysteines are often involved in redox reactions, and the presence of selenium dramatically changes the chemical properties of the protein. Nevertheless, several reports have described the expression of selenocysteine-containing analogs of proteins (Muller et al., 1994; Boschi-Muller et al., 1998). In one report (Muller et al., 1994), thioredoxin was expressed in an E. coli cysteine auxotroph that was grown in the presence of selenocysteine. Selenocysteine was clearly toxic to bacterial cells, and only after carefully adjusting growth conditions did the expression of limited amounts of the selenocysteine-containing protein occur. Interestingly, this selenothioredoxin formed a diselenide bond in place of the normal disulfide bond. However, the diselenide bond was characterized by a decreased redox potential, and thioredoxin reductase failed to reduce selenothioredoxin.
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In addition to the methods that replace either all methionines or all cysteines in proteins with the corresponding selenium-containing amino acids, a technique has been developed for sitespecific incorporation of selenocysteine. Since UGA dictates the incorporation of selenocysteine into selenoproteins (see below), site-directed mutagenesis to generate a TGA codon at a particular site in an open reading frame may be used to signal selenocysteine incorporation. Such a mutant TGA codon must be coupled with an additional signal that is necessary in the resulting mRNA for distinguishing between signals for selenocysteine incorporation and termination of protein synthesis. The latter signal is a stem-loop structure, designated as the selenocysteine insertion sequence (SECIS) element, and it is located in selenoprotein mRNA downstream of the UGA codon. It should be noted that SECIS elements (and systems for selenocysteine incorporation) are different in bacteria and mammals (see below), and therefore SECIS elements that are specific for the host organism must be utilized for selenoprotein expression. An example of an unnatural site-specific incorporation of selenocysteine into proteins is the expression of Methanobacterium formicicum formate dehydrogenase in E. coli (Heider and Bock, 1992). The natural cysteine residue was replaced in the expressed protein with selenocysteine by generating an in-frame TGA codon and a downstream E. coli–type SECIS element. A similar method has also been used to express eukaryotic selenoproteins in E. coli (Arner et al., 1999; Gladyshev et al., 1999b). Another example is the generation of unnatural selenocysteine residues in glutathione peroxidase in various sites in the open reading frame of the gene (Wen et al., 1998). In this case, TGA codons were individually generated in various locations within the open reading frame, but a single eukaryotic SECIS element was present in the 3′ untranslated region (3′-UTR).
Occurrence of Selenium in Natural Selenoproteins In natural selenoproteins, selenium is present either as a dissociable cofactor or as a selenocysteine residue. The dissociable cofactor has only been found in molybdenum-containing enzymes (Dilworth, 1982) in which selenium is coordinated to molybdenum in the enzyme active center (Gladyshev et al., 1994a). For example, in the seleno-molybdoenzyme form of nicotinic acid hydroxylase from Clos-
tridium barkeri, selenium is essential for catalytic activity, as evidenced by comparison of kinetic parameters of enzyme forms isolated from selenium-supplemented and seleniumdeficient cells (Gladyshev et al., 1994a). Other enzymes of this class are clostridial xanthine dehydrogenases and, perhaps, bacterial molybdenum-containing carbon dioxide dehydrogenases (Dobbek et al., 1999). The existence of selenium as a cofactor has not been described in eukaryotic and archaeal proteins. The occurrence of selenium as a selenocysteine residue is by far the most common form of this element in natural proteins. The means by which selenocysteine is incorporated into protein, the known selenoproteins, and the methods to identify and characterize these proteins are discussed below.
COTRANSLATIONAL INCORPORATION OF SELENOCYSTEINE INTO PROTEINS Selenocysteine is the 21st naturally occurring amino acid in protein. It is incorporated into protein cotranslationally in response to the codon UGA (Fig. 3.8.1). In addition to UGA, selenocysteine has its own tRNA, a specific elongation factor that is designated SELB, and the SECIS element that is located in mRNA downstream of UGA. As noted above, the SECIS element is essential for recognizing UGA as a signal for selenocysteine incorporation rather than as a signal for termination.
UGA as a Selenocysteine Codon When the genetic code was deciphered in the mid-1960s, it was thought to be universal (see Gladyshev et al., 2000, and references therein). However, subsequent studies revealed that many changes in the genetic code occurred in evolution. The code therefore has been referred to as the “almost universal genetic code” (Gesteland et al., 1992). In spite of variations in the meaning of codons in different organisms and organelles, only 20 amino acids were thought to be encoded in the gene sequences of organisms until the code was expanded to include selenocysteine in the late 1980s (reviewed in Bock et al., 1991; Hatfield and Diamond, 1993). Addition of selenocysteine marks the 21st amino acid in the genetic code, and is the only addition to the code since it was deciphered. UGA is most often used as a stop signal in nature, but because it also dictates selenocysteine insertion into protein, it has a dual function
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A
initiation of translation
selenocysteine codon
termination of translation SECIS
5′
AUG
UGA
3′ AAAA
UAA UAG UGA
B SECIS 5′
AUG
UGA
UAA UAG UGA
3′
Figure 3.8.1 Structure of selenoprotein mRNA in mammals (A) and E. coli (B). The UGA codon that is present in the open reading frame is decoded as selenocysteine. The stem-loop structure (the SECIS element) is present in the 3′-UTR in mammals and immediately downstream of the UGA selenocysteine codon in E. coli.
(reviewed in Kumaraswamy et al., 1999). AUG also has a dual function in the genetic code, serving as an initiation codon for protein synthesis and as a methionine codon at internal positions of protein. The dual role of AUG, however, has been known since the time the code was deciphered. The presence of UGA in an open reading frame is considered to be the most characteristic feature of selenoprotein genes. However, the occurrence of UGA in an open reading frame does not by itself mean that it codes for selenocysteine, as certain other criteria in gene sequences must also be met (see below).
The SECIS Element
Analysis of SelenocysteineContaining Proteins
The SECIS element is essential for signaling selenocysteine incorporation into protein. Only E. coli and mammalian SECIS elements have been studied in detail, and the stem-loop structures from these organisms clearly differ in terms of conserved sequences and overall structural organization (Low and Berry, 1996). Moreover, the SECIS elements in E. coli occur immediately downstream of the UGA selenocysteine codon, while those in mammals occur further downstream in the 3′-UTR. The distance between the SECIS element and UGA has been determined in mammals and may be as far as 5.4 kb (Buettner et al., 1998) or as close as 51 to 111 bp (Martin et al., 1996). Several archaeal selenoprotein sequences have also been analyzed (Wilting et al., 1997). It
was concluded from these analyses that archaeal SECIS-like structures are different from both mammalian and bacterial SECIS elements, and are likely located in archaea either upstream or downstream of UGA in the 5′- or 3′-UTR (Wilting et al., 1997). The largest number of SECIS elements has been characterized in mammals (GrundnerCulemann et al., 1999). Their structure and function have been probed by site-directed mutagenesis and by chemical and enzymatic methods (Martin et al., 1998; Walczak et al., 1998). These observations have led investigators to draw various conclusions about the conservation of primary and secondary structures of mammalian SECIS elements. A composite structure of the mammalian SECIS element is shown in Figure 3.8.2. It consists of an apical loop and two stems separated by an internal loop. A conserved motif, AUGA (or GTGA), is present at the base of the upper stem. The first nucleotide of this conserved motif is unpaired, while UGA and a following nonconserved nucleotide participate in the quartet of non-Watson-Crick interacting nucleotides. Other conserved nucleotides are an unpaired AA motif in the apical loop and GA in the 3′ portion of the quartet. The distance between the quartet (also designated as the SECIS core) and the conserved AA in the apical loop is 11 or 12 nucleotides. These characteristics of SECIS elements were used to design an algorithm for searching
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Current Protocols in Protein Science
apical loop
AA helix II
N A G U A
• • • •
N G A N
quartet
internal loop
helix I 5′
3′
Figure 3.8.2 Mammalian SECIS element. The conserved nucleotides are indicated by letters and the overall features of the consensus sequence are discussed in the text.
large nucleotide sequence databases and identifying mammalian SECIS elements. This algorithm, designated SECISearch, conducts searches in three steps: (1) an initial primary sequence search for conserved SECIS motifs, (2) a secondary structure search of the motifs of SECIS elements, and (3) determination of the free energy in a candidate SECIS element. Application of SECISearch to analyze the human expressed sequence tag (EST) database resulted in identification of new selenoprotein genes (Kryukov et al., 1999). Two versions of SECISearch are currently available: a database search version and an on-line version. These programs use slightly different parameters and are optimized for different purposes. The former version can be used to search nucleotide databases for the smallest number of sequences that satisfy primary and secondary structure and free energy parameters of the SECISearch algorithm. This version is thus optimized for efficiency of searches, although it may miss SECIS elements that deviate slightly from the consensus SECIS structure. This version was designed to minimize the probability of finding pseudo-SECIS structures. In contrast, the on-line version can be used to analyze a query nucleotide sequence for
the presence of a potential SECIS element. This version is optimized to find all potential SECIS elements in a sequence, which of course means that it may predict a pseudo-SECIS structure. The on-line version was designed to minimize the probability of missing the actual SECIS element. This version should be most helpful to investigators who would like to test a sequence of interest for the presence of a SECIS element and for rapid identification and visualization of potential SECIS elements. The online version is currently available at http:// bio-selenium.unl.edu/SECIS.htm (Kryukov et al., 1999).
Selenocysteine tRNA Selenocysteine, unlike the other 20 amino acids utilized in mammalian protein synthesis, is biosynthesized on its tRNA (Gladyshev et al., 2000). Thus, selenocysteine tRNA has a dual function of serving as both a carrier molecule for the synthesis of its amino acid as well as the molecule that donates selenocysteine to the growing selenopolypeptide in response to specific UGA codons. The tRNA is first aminoacylated with serine, which serves as the backbone for selenocysteine biosynthesis (see below). Therefore, this tRNA has been designated as tRNA[Ser]Sec. Selenocysteine tRNA[Ser]Sec is 90 nucleotides in length and is the longest eukaryotic tRNA sequenced to date (reviewed in Hatfield et al., 1999). The tRNA [Ser]Sec population consists of two major isoacceptors in mammalian cells that differ from each other by a single 2′-O-methylribose in the wobble position (position 34) of the anticodon. One isoacceptor has 5′-methylcarboxymethyluridine (mcmU) in position 34, and the other h a s 5′-methylcarboxymethyluridine-2′-Omethylribose (mcmUm). The degree of methylation of tRNA[Ser]Sec at position 34 is influenced by selenium status (Hatfield et al., 1991; Diamond et al., 1993; Choi et al., 1994; Chittum et al., 1997), and the presence of this methyl group affects tertiary structure (Diamond et al., 1993). However, the biological significance of this methyl group is not known. Biosynthesis of mcmU and mcmUm, as well as the other modified bases in tRNA[Ser]Sec, h av e b ee n e s t ab l i s h e d i n Xenopus oocytes (Choi et al., 1994; Sturchler et al., 1994; Kim et al., unpub. observ.). Selenocysteine tRNA[Ser]Sec is transcribed unlike any other known tRNA in that it begins transcription at the first base within the gene and thus lacks a 5′ leader sequence (Lee et
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al., 1987). The regulatory regions within the tRNA[Ser]Sec gene and the transcription factors and mechanisms governing its expression have been reviewed recently (Hatfield et al., 1999). Subsequent studies involving the regulatory elements of this tRNA have prim ari ly d eal t w ith th e selen ocysteine tRNA[Ser]Sec gene transcription-activating factor (Staf), which specifically enhances transcription of this tRNA gene (Adachi et al., 1998, 1999; Myslinski et al., 1998; Schuster et al., 1998; Schaub et al., 1999).
Selenocysteine Biosynthesis
Analysis of SelenocysteineContaining Proteins
As noted above, the biosynthesis of selenocysteine begins with the attachment of serine to tRNA[Ser]Sec, which is mediated by seryl-tRNA synthetase. The identity elements in tRNA[Ser]Sec for its aminoacylation are therefore governed by seryl-tRNA synthetase and they reside in the discriminator base and the long extra arm (Wu and Gross, 1993; Ohama et al., 1994). The acceptor, T, and D stems also play a role in the identity process (Amberg et al., 1996). The metabolic pathway involving synthesis of selenocysteine on tRNA[Ser]Sec (with serine serving as the backbone; Sunde and Evenson, 1987; Lee et al., 1989; Leinfelder et al., 1989) has been fully characterized in E. coli (see Commans and Bock, 1999, and references therein). In bacteria, a single enzyme, selenocysteine synthase, catalyses the removal of water from serine to form an aminoacrylyltRNA[Ser]Sec intermediate, which in turn catalyzes the acceptance of selenium from the activated form, monoselenophosphate, to yield selenocysteyl-tRNA[Ser]Sec and inorganic phosphate. The synthesis of monoselenophosphate is mediated by selenophosphate synthetase (Glass et al., 1993; Stadtman, 1996). In mammals, further work is required to establish the pathway of selenocysteine synthesis. Selenocysteine tRNA[Ser]Sec was originally discovered as a minor seryl-tRNA that decoded UGA (Hatfield and Portugal, 1970) and formed phosphoseryl-tRNA (Maenpaa and Bernfield, 1970), and was later shown to be selenocysteyltRNA[Ser]Sec (Lee et al., 1989). The role of phosphoseryl-tRNA[Ser]Sec in selenocysteine biosynthesis has not been resolved (Hatfield et al., 1994), but some investigators have proposed that it is “an active storage form” that can be regenerated for selenocysteine biosynthesis after its dephosphorylation (Amberg et al., 1996). It should be noted that the serylated form of tRNA[Ser]Sec has a role of its own in protein
biosynthesis. Seryl-tRNA[Ser]Sec is recognized by the normal elongation factor eEF-1 (Jung et al., 1994) and serves as an in vivo suppressor of UGA stop codons, inserting serine and extending protein synthesis (Chittum et al., 1998).
Elongation Factor SELB and Selenocysteine Incorporation Once selenocysteyl-tRNA[Ser]Sec is synthesized, selenocysteine may be donated to the nascent selenopolypeptide in response to the appropriate UGA codon. In addition to UGA and the SECIS element, another factor that is essential for selenocysteine incorporation into protein is the selenocysteine tRNA[Ser]Sec specific elongation factor SELB. This factor has so far only been described in bacteria (Forchhammer et al., 1990). Bacterial SELB binds to both selenocysteyl-tRNA[Ser]Sec and the SECIS element. The presence of selenocysteine attached to tRNA[Ser]Sec is critical in this interaction, as it avoids recognition of the serylated form (see Commans and Bock, 1999, and references therein). Studies with mammalian systems also suggest the existence of an analogous SELB factor (reviewed in Gladyshev et al., 2000). Several potential SELB candidates have been identified, and one of them appears to specifically recognize the quartet sequence of the SECIS element (Copeland and Driscoll, 1999). This protein also enhanced selenocysteine incorporation into protein from selenocysteyltRNA[Ser]Sec (Copeland et al., 2000). However, this SECIS-binding protein does not appear to bind selenocysteine tRNA and is likely a stimulatory factor. This suggests that the role of bacterial SELB is served by two or more proteins in mammals.
SELENOCYSTEINE-CONTAINING PROTEINS Selenocysteine-containing proteins have been characterized from both prokaryotes and eukaryotes. The number of selenoproteins differs among various organisms. For example, E. coli contains three selenoproteins and all three are formate dehydrogenases (Heider and Bock, 1993). Archaeon M. jannaschii contains up to seven selenoproteins (Wilting et al., 1997). Nematode C. elegans appears to have only a single selenoprotein (Gladyshev et al., 1999b; Buettner et al., 1999), and no selenoproteins are present in S. cerevisiae. To date, the largest number of selenoproteins, sixteen, has been identified in mammals, including humans (Gladyshev and Hatfield, 1999).
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Selenocysteine-containing proteins that have been characterized thus far are shown in Table 3.8.2. Selenocysteine appears to serve different functions in prokaryotic and eukaryotic proteins. In several prokaryotic selenoproteins, selenium is coordinated to a metal such as nickel, molybdenum, or tungsten. The selenium moiety participates in proton transfer in these proteins and also fine tunes the coordination sphere of a metal in the enzyme active center (Boyington et al., 1997). In contrast, selenoproteins in eukaryotes are often associated with disulfide oxidoreductase redox pathways, such as thioredoxin- and glutathione-dependent systems. However, the two common features among all selenoproteins with established function are that these proteins participate in redox processes and that selenocysteine is located at the enzyme active center. Further information on properties and functions of selenium-containing proteins can be found in recent reviews (Low and Berry, 1996; Stadtman, 1996; Gladyshev and Hatfield, 1999; Hatfield et al., 1999; Gladyshev et al., 2000).
SPECIFIC METHODS FOR IDENTIFYING AND CHARACTERIZING SELENOPROTEINS Several methods have been used extensively for the analysis of selenium-containing proteins. These methods have utilized the unique chemical and physical properties of selenium and its isotopes (see Table 3.8.3). For example, 75Se-enriched proteins can be easily detected in protein fractions with γ counters and on polyacrylamide gels by PhosphorImager analyses. In addition, 77Se-enriched proteins can be analyzed by NMR and electron paramagnetic resonance (EPR) spectroscopies. Selenoproteins have also been analyzed with methods such as X-ray absorption spectroscopy, atomic absorption, and mass spectrometry that detect natural selenium species or their derivatives. These procedures are discussed below, and their characteristics, advantages, and limitations are summarized in Table 3.8.3. The use of these methods generally implies that an investigator has sufficient quantities of a homogeneous selenoprotein. However, several methods, most notably metabolic labeling with 75Se, do not require pure protein for analyses. In addition, several techniques, such as NMR, EPR, and X-ray spectroscopies, require sophisticated equipment and extensive expertise, and therefore are often used on a collaborative basis. Finally, identification of new selenoprote-
ins through bioinformatics approaches are discussed.
Methods Utilizing the 75Se Isotope Selenium-containing proteins are most easily detected if they are labeled with 75Se. This γ emitter has a convenient half-life of 121 days and is available from several sources (e.g., University of Missouri Research Reactor Center). Selenoproteins are labeled with 75Se by cotranslational incorporation of selenium when cells are grown in the presence of [75Se]selenite (state of oxidation, +4). The use of selenate (state of oxidation, +6) should be avoided, as this molecule is not easily reduced to biologically active selenide (state of oxidation, −2). To label cells, 75Se, which is normally provided by commercial suppliers in a 35% nitric acid solution, is neutralized with sodium hydroxide and supplied directly to the growth medium (Gladyshev et al., 1999a). Depending on the type of experiment, between 5 and 500 µCi of 75Se are added to 50 ml of a bacterial or mammalian growth medium. To label selenoproteins in animals, the animals are injected peritoneally with freshly neutralized [75Se]selenite (Gladyshev et al., 1998) and then sacrificed at an appropriate time for extraction of proteins from the tissue of interest. It should be noted that [75Se]selenocysteine is not a good source of 75Se for biosynthetic radioactive labeling of selenoproteins. As discussed above, free selenocysteine cannot be incorporated into selenoproteins because this amino acid must be synthesized on tRNA[Ser]Sec from serine and monoselenophosphate. In contrast, [75Se]selenocysteine will enter sulfur pathways and will be incorporated into proteins in place of cysteine, resulting in a higher background during PhosphorImager or γ-counter analyses. Labeling of selenoproteins with 75Se provides an easy means of detecting these proteins in cellular fractions during protein isolation. 75Se may be quantitatively and directly detected in a γ counter. Multidetector counters allow analyses of 100 fractions in only 10 minutes with no loss of material. Therefore, this method is generally superior to spectrophotometric and kinetic assays in terms of sensitivity and time required. Selenoproteins that are labeled with 75Se may also be detected by PhosphorImager analyses on polyacrylamide gels. Under appropriate labeling conditions, this method is more sensitive than immunoblot assays. Growth of mammalian cells (APPENDIX 3C) in the presence
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Table 3.8.2 Selenocysteine-Containing Proteins in Bacteria, Archea, and Eukaryotes
Protein Bacteria
Archaea
Eukaryotes
Formate dehydrogenases Formate dehydrogenase H Formate dehydrogenase N Formate dehydrogenase O Hydrogenases
Organism
Anaerobic bacteria E. coli E. coli E. coli Sulfate-reducing bacteria Glycine reductase protein A Anaerobic bacteria Glycine reductase protein B Anaerobic bacteria Proline reductase protein B Anaerobic bacteria Betaine reductase protein B Anaerobic bacteria Sarcosine reductase protein Anaerobic bacteria B Selenophosphate synthetase Bacteria Formate dehydrogenases Methanogenic archaea Formylmethanofuran Methanogenic archaea dehydrogenase Hydrogenases Methanogenic archaea Selenophosphate synthetase M. jannaschii Cytosolic glutathione Vertebrates peroxidase (GPx1) Gastrointestinal glutathione Vertebrates peroxidase (GPx2) Vertebrates Plasma glutathione peroxidase (GPx3) Vertebrates, S. Phospholipid hydroperoxide glutathione mansoni peroxidase (GPx4) Plasma selenoprotein P Vertebrates (SelP) Selenoprotein W (SelW) Vertebrates Selenoprotein T (SelT) Vertebrates, S. mansoni Selenoprotein R (SelR) Vertebrates 15 kDa selenoprotein Vertebrates Selenophosphate synthetase Vertebrates 2 (SPS2) Thioredoxin reductase 1 Vertebrates, C. elegans (TR1) Thioredoxin reductase 2 Vertebrates (TR2) Thioredoxin reductase 3 Vertebrates (TR3) Thyroid hormone Vertebrates deiodinase 1 (DI1) Thyroid hormone Vertebrates deiodinase 2 (DI2) Thyroid hormone Vertebrates deiodinase 3 (DI3)
Properties; representative reference Contain Mo, FeS; Sawers (1994) 80 kDa; Mo, FeS; Boyington et al. (1997) 110 kDa; Mo, FeS, FAD; Benoit et al. (1998) 110 kDa; Mo, FeS, FAD; Berg et al. (1991) Ni, FeS, FAD; Eidsness et al. (1989) 18 kDa; Kreimer and Andreesen (1995) 47 kDa; Kreimer and Andreesen (1995) 47 kDa; Kabisch et al. (1999) 47 kDa; Andreesen et al. (1999) 47 kDa; Andreesen et al. (1999) ATP dependent; Guimaraes et al. (1996) Mo, FeS, FAD; Wilting et al. (1997) FeS, Mo or W; Vorholt et al. (1997) Ni, FeS, FAD; Wilting et al. (1997) ATP dependent; Guimaraes et al. (1996) Homotetramer of 23-kDa subunits; Chambers et al. (1986) Homotetramer of 23-kDa subunits; Chu et al. (1993) Homotetramer of 25-kDa subunits, glycoprotein; Maser et al. (1994) 20 kDa; Schuckelt et al. (1991)
55 kDa protein with 10-12 Sec; Burk and Hill (1999) 10 kDa protein subunit; Vendeland et al. (1995) 18 kDa protein subunit; Kryukov et al. (1999) 14 kDa protein subunit; Kryukov et al. (1999) 15 kDa protein subunit; Gladyshev et al. (1998) 50 kDa; Guimaraes et al. (1996) Homodimer of 57- kDa subunits; Gladyshev et al. (1996c) Homodimer of 65- kDa subunits; Sun et al. (1999) Homodimer of 57- kDa subunits; Lee et al. (1999) Homodimer of 29- kDa subunits; Berry et al. (1991) 30 kDa; Davey et al. (1995) 31 kDa; Croteau et al. (1995)
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Table 3.8.3
Specific Methods for Analyses of Selenium-Containing Proteins
Methoda
Descriptiona
Radioactive labeling
75Se isotope is commonly used More sensitive than immunoblot assay to detect, isolate, and characterize selenoproteins. Half-life of this γ emitter is 121 days.
NMR spectroscopy
77Se
EPR spectroscopy
Atomic absorption spectroscopy Mass spectrometry
X-ray absorption spectroscopy
Sensitivity
Requires 0.1 to 1 mM protein concentration
may be detected in protein; method may reveal information about coordination of Se and enzyme reaction mechanisms 77Se is used to replace natural Se in a protein by growing cells in presence of [77Se]selenite. If 77Se is in vicinity of a paramagnetic group, method may reveal information about coordination of Se, structure of active center, and reaction mechanism of enzyme Se is directly detected in proteins and biological samples
Advantages Simultaneous detection of many proteins by PhopshorImager analysis; no loss of material in γ-counter assays; detection of selenoproteins in crude extracts Employs 77Se as a nonperturbing probe for protein structure and function
Requires 20 to 200 µM protein Provides unique structural and concentration mechanistic information
nM to µM range depending on sample composition Very sensitive, nM range
ICP-MS: analytical technique for determination of Se in proteins and biological samples; ESI-MS: may detect Se-containing peptides due to a unique isotopic composition of Se Se is directly detected in a protein; EXAFS analysis reveals coordination sphere of Se in a protein and may also determine oxidation state of environmental Se
Not very sensitive; requires ∼1 mM concentration of Se in sample
Reliable method for determination of Se content of a protein Sensitivity; utilizes unique isotopic composition of Se
No interference from contaminants; analysis of intermediates trapped by freezing
aAbbreviations: EPR, electron paramagnetic resonance; ESI-MS, electrospray ionization mass spectrometry; EXAFS, extended X-ray absorption
fine structure; ICP-MS, inductively coupled plasma mass spectrometry; NMR, nuclear magnetic resonance.
of 75Se followed by SDS-PAGE analysis of protein crude extracts (UNIT 10.1) allows simultaneous detection of several major selenoproteins including thioredoxin reductase 1, glutathione peroxidase 1, glutathione peroxidase 4, and the 15 kDa selenoprotein (Gladyshev et al., 1999a). These proteins may be detected simultaneously because they have different mobilities on SDS-PAGE (and nondenaturing) gels (UNITS 10.1 & 10.3). To obtain a selenoprotein signal, gels or immunoblot membranes (UNIT 10.11) are incubated overnight in a Phos-
phorImager cassette. Alternatively, an 75Se signal may be detected by exposing gels or membranes to an X-ray film for two to seven days (UNIT 10.11).
Methods Utilizing the 77Se Isotope Proteins enriched with 77Se (nuclear spin ; 2 natural abundance ∼7.5%) may be analyzed by NMR spectroscopy (for overview, see UNIT 17.5). In addition, if selenium is located in the vicinity of the paramagnetic group, these proteins may be probed with EPR spectroscopy. 1⁄
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77Se-enriched proteins are obtained by growing
cells in the presence of [77Se]selenite. In contrast to labeling of cells with 75Se, which does not require quantitative incorporation of [75Se]selenocysteine into a protein, 77Se must be a major form of selenium in a selenoprotein for subsequent analyses by NMR and EPR spectroscopies. Therefore, this isotope is normally used at concentrations as high as 1 µM to restrict incorporation of other selenium isotopes present in the medium. 77Se-NMR and 77Se-EPR spectroscopies provide information about electronic properties, coordination geometry, and reaction mechanisms of selenium sites in proteins. For example, EPR spectroscopy of 77Se-enriched enzyme preparations has been used to demonstrate the coordination of Se to Mo and to characterize reaction mechanisms of formate dehydrogenase (Gladyshev et al., 1994b; Khangulov et al., 1998) and nicotinic acid hydroxylase (Gladyshev et al., 1994a, 1996b). In addition, coordination of Se to Ni in hydrogenases has been demonstrated by EPR spectroscopy (He et al., 1989; Sorgenfrei et al., 1997). 77Se-NMR spectroscopy has been used in studies involving mammalian glutathione peroxidase (Gettins and Crews, 1991), bacterial selenomethionyl dihydrofolate reductase (Boles et al., 1992), and other enzymes.
Other Methods Atomic absorption spectroscopy This is the most commonly used analytical method for direct quantitation of selenium in biological samples. Conditions have been developed for determination of this element in proteins and biological media. Selenium in proteins and serum may be analyzed with Pd-Mg or Ag-Cu-Mg modifiers by graphite furnace atomic absorption spectroscopy with Zeeman background correction as described by Daher and Van Lente (1994). This method, however, can determine only one chemical element at a time. An additional common technique for quantitative selenium analysis in biological samples is neutron activation analysis (discussed in Chan et al., 1998a).
Analysis of SelenocysteineContaining Proteins
Mass spectrometry The capabilities of another analytical method for determination of selenium in proteins and biological samples, inductively coupled plasma mass spectrometry (ICP-MS), exceeds the slow, single-element analysis by atomic absorption spectroscopy in that it can
be used to simultaneously determine multiple trace elements. This method has been applied to determine selenium concentration in serum, and details of the technique can be found in Chan et al. (1998b). A variation of this technique, electrospray ionization mass spectrometry (ESI-MS), may potentially be used for identification of selenium-containing proteins and peptides due to the unique isotopic composition of selenium. For example, fragmentation of a selenium-containing peptide from human thioredoxin reductase produced a spectrum of masses in which selenium-containing fragments could be detected by a characteristic signature of masses (Fig. 3.2.2; Sun et al., 1999, and figures within). X-ray absorption spectroscopy Selenium in selenoproteins also may be probed with X-ray absorption spectroscopy. Extended X-ray absorption fine structure (EXAFS) may then reveal information about the oxidation state and coordination sphere of selenium. Limitations of this technique are the necessity to use synchrotron radiation and requirements for large amounts of samples. In addition, selenium concentration in the sample should be ≥1 mM, which cannot be reached for many proteins because of protein solubility problems. EXAFS analyses probed selenium ligands in formate dehydrogenase H from E. coli (George et al., 1998), hydrogenase from Desulfovibrio baculatus (Eidsness et al., 1989), selenomethionine-substituted cytochrome ba3 from Thermus thermophilus (Blackburn et al., 1999), selenium-substituted nitrogenase (Conradson et al., 1994), and other proteins.
Cautions to be Taken in Selenoprotein Isolation Selenium-containing proteins are generally very labile due to the high reactivity of selenium in proteins. Selenium may be lost from selenoproteins by hydrolysis in the presence of oxygen, and this process is enhanced in bright light. To obtain selenoproteins of the highest activity, it is recommended that they be isolated as quickly as possible at 4°C in relative darkness. The selenium moiety in natural selenoproteins often serves a redox function, and therefore selenoproteins may be additionally protected during isolation by adding a reducing agent such as dithiothreitol or 2-mercaptoethanol. A number of selenoproteins are quickly inactivated in the presence of oxygen. To isolate such proteins, anaerobic conditions may be required. In isolating most selenoproteins, de-
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oxygenated solutions may be used throughout the isolation procedure. In case of extreme oxygen sensitivity, anaerobic chambers may be necessary (Axley et al., 1990; Gladyshev et al., 1994b). Anaerobic chambers are normally operated at 1 ppm of oxygen or less. Selenoprotein formate dehydrogenase H from E. coli is extremely sensitive to traces of oxygen, but the enzyme can be isolated and crystallized in its active state at <1 ppm of oxygen in an anaerobic chamber (Gladyshev et al., 1996a).
Identification of Selenoprotein Genes in Nucleotide Sequence Databases Until recently, selenoproteins were identified by isolating and characterizing selenocysteine in alkylated protein hydrolysates. This is a labor-intensive method, requiring considerable effort to detect and purify a protein and requiring extensive experience in amino acid analyses. Furthermore, selenoproteins are generally present in low amounts in cells and tissues, and therefore any given selenoprotein may be a limiting factor for analysis. The accumulation of known selenoprotein gene sequences has provided an important tool for making certain generalizations about selenoprotein gene structure that have aided in identifying new selenoproteins. From these generalizations, it initially became apparent that the presence of in-frame TGA is not sufficient information to identify a selenoprotein. Misinterpretations of TGA codons as both selenocysteine and stop signals are known. A more rigorous procedure had to be developed to distinguish between in-frame TGA codons for selenocysteine and those for stop signals, and to identify selenoprotein genes. Three criteria have been defined recently for specifically identifying mammalian selenoprotein genes. The Mammalian Selenoprotein Gene Signature (MSGS) predicts that a protein gene contains a selenocysteine codon if the following criteria are satisfied: 1. Sec is conserved and is flanked by conserved regions in mammalian sequences for this protein. 2. The SECIS element sequence and structure are conserved and are located in the 3′UTRs of mammalian genes for this protein. 3. Genes occur (usually in lower eukaryotes) that contain a Cys codon in place of TGA in the gene for the putative selenoprotein (or distinct homologous genes occur that conserve the TGA codon), and that meet an additional criterion of encoding the sequences with
sufficiently long homologous regions on both sides of the Cys/Sec (or Sec/Sec) pair. MSGS therefore predicts that a selenoprotein will have a conserved Sec and its flanking regions, a conserved SECIS element in 3′-UTR of the corresponding mRNA, and distinct homologs that contain either Sec or Cys in place of selenocysteine in the protein of interest. MSGS has been used to analyze several databases, including the entire genomes of S. cerevisiae, E. coli, and C. elegans, and various EST and nonredundant databases. These searches found all known mammalian selenoprotein genes and, in addition, by satisfying MSGS criteria, they have identified several new selenoproteins (Kryukov et al., 1999; Gladyshev and Kryukov, unpub. data). A similar strategy may be applied to identify selenoprotein genes in bacterial, archaeal, and eukaryotic sequences. In fact, SECISearch and MSGS correctly identified selenoprotein genes in many eukaryotic organisms, such as nematodes, trematodes, flies, amoebae, fish, and birds. On the other hand, the search of the S. cerevsiae genome, the only eukaryotic organism that is known to lack selenoproteins, found no selenoprotein genes. This suggests that MSGS and SECISearch may be used to analyze any eukaryotic genome. However, at this point, it is not known whether deviations from MSGS may occur in eukaryotic genomes that falsely identify selenoproteins, or if sequences selected by MSGS should be further tested before making a final decision.
STRATEGIES FOR IDENTIFYING SELENOCYSTEINE-ENCODING GENES AND SELENOCYSTEINECONTAINING PROTEINS There are several approaches that may be used to determine whether a newly isolated mammalian (or other animal) gene encodes selenocysteine or whether the protein contains a selenocysteine moiety, outlined in Figures 3.8.3 and 3.8.4. The investigator may ask several questions regarding the presence of conserved features observed in known selenoprotein genes that should reside in the newly isolated gene (see Fig. 3.8.3). Does this gene have an in-frame TGA codon? Can a TGA codon that is thought to terminate protein synthesis potentially encode a selenocysteine residue? Does this gene have a putative SECIS element in the 3′-UTR? If the gene has an in-frame TGA codon and a putative SECIS-like structure, are these features conserved in orthologous genes in other mammals? Are there distinct homolo-
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1. Putative Mammalian Selenoprotein Gene Isolated
no
It is unlikely to be a selenoprotein gene
no
It is unlikely to be a selenoprotein gene
4. Do orthologous genes from other mammals conserve the in-frame TGA codon and SECIS element? yes
no
It is unlikely to be a selenoprotein gene
5. Are there distinct homologous proteins that either conserve the in-frame TGA codon or replace it with TGC or TGT (cysteine codons)?
no
It is unlikely to be a selenoprotein gene
2. Does it have an in-frame TGA codon? Is TGA a termination codon?
yes 3. Does it have a putative SECIS element in 3′-untranslated region?
yes
6. Candidate Selenoprotein Gene Identified
7. When this gene is expressed in mammalian cells, is the gene product labeled with 75Se?
8. Optional: Is expression of the protein dependent on levels of Se in cell culture or in the diet? How many equivalents of Se are present in the protein? Does this protein contain Se in the form of Sec?
9. New Selenoprotein Identified
Figure 3.8.3 Strategies to test whether a newly isolated gene encodes a selenoprotein. See text for details.
Analysis of SelenocysteineContaining Proteins
gous proteins that have a cysteine codon (TGC or TGT) in place of the TGA selenocysteine codon? If the answer to any of these questions is no, it is unlikely that the new gene encodes a selenoprotein. It should be noted, however, that some of these questions may not be answered if insufficient information is available for the new gene or its homologs. For example, SECIS elements are often located at the very 3′ ends of mRNAs. If this portion of cDNA is lacking, a SECIS element cannot be easily identified. In addition, variations from conserved features found in known SECIS ele-
ments may be anticipated to occur in at least some new selenoprotein genes. This is because certain point mutations in conserved features of previously characterized SECIS elements do not always correlate with decreased selenocysteine incorporation. Additional difficulties in answering questions in blocks 2 to 5 of Figure 3.8.3 may arise if a new gene does not have known homologs in other species. Therefore, further experimental verification for a new selenoprotein gene may be required. The experimental design in determining if a new gene may encode a selenocysteine may
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Method
1. Putative Selenoprotein Purified
Comments
atomic absorption spectroscopy
2. Does the protein contain Se? How many equivalents of Se are present?
If Se is not present in the protein then it is not a selenoprotein.
γ counter and PhosphorImager analyses
3. When the protein is isolated from cells grown in the presence of 75Se, is it 75Se-labeled?
If 75Se is not present in the protein then it is not a selenoprotein.
native and SDS/PAGE analysis
4. Is Se lost when protein is analyzed by gel electrophoresis in the presence of a reducing agent?
If Se is not present then it is not a Sec-containing protein.
amino acid composition analysis
5. Does amino acid analysis of 75Se-labeled protein show the presence of Sec?
If Sec is not present then it is not a selenoprotein.
amino acid sequencing (Edman degradation)
6. In the isolated 75Se-labeled peptide from a protease digest of an alkylated protein, what is the location of the 75Se-containing amino acid?
Location of Sec is identified by determining a cycle where 75Se is released.
tandem mass spectrometry (MS/MS) analysis
7. In the isolated Se-containing peptide obtained by a protease digest, what is the mass of the alkylated amino acid?
Sec has a unique mass. Se is also detected due to its unique natural isotope abundance.
computational screen (see Fig. 3.8.3)
8. Does the gene for this protein contain an in-frame UGA and/or a SECIS element?
The presence of UGA and SECIS element are expected.
9. New Selenoprotein Identified
Figure 3.8.4 Strategies to test whether a newly isolated protein is a selenoprotein. See text for details.
also include a question of whether the gene product incorporates selenium when it is expressed in mammalian cells. One technique to address this question is to use 75Se labeling of cells transfected with a construct that expresses the new gene. The molecular mass for a new selenoprotein should not correspond to the known masses for major natural selenoproteins expressed in host cells, such as thioredoxin reductase (57 kDa), glutathione peroxidase 1 (25 kDa), glutathione peroxidase 4 (20 kDa), and the 15 kDa protein. This can be achieved
by expressing the new gene as a fusion with a tag protein, such as green fluorescent protein. If all four questions in the computational screen are answered as yes, and if the overexpressed protein is labeled with 75Se, the data may be considered proof for identification of a new selenoprotein gene. Additional (optional) questions may be asked to further resolve the identity of the new putative selenoprotein gene. For example, is the expression of the gene product dependent on levels of selenium in cell culture or in the diet? What is the chemical form of
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Analysis of SelenocysteineContaining Proteins
selenium in the selenoprotein and how many equivalents of selenium are present in the protein? These questions may be addressed using the techniques summarized in Table 3.8.3 and discussed above. In contrast to the experimental design used in identifying new selenoprotein genes, a different set of questions may be asked to determine if an isolated protein is a selenoprotein. The first question to resolve is whether the protein contains selenium. The assay for selenium is straightforward and, if an atomic absorption spectrometer is not available, several companies are available that can determine selenium content in a protein. It is important to determine how many equivalents of selenium are present in a protein. It should be noted that a partial loss of selenium may occur during isolation due to its lability. Therefore, a value of 0.5 equivalents of selenium per protein subunit is not unusual, and a value of 0.9 equivalents is rarely achieved unless specific precautions are taken during isolation (e.g., avoiding direct sunlight, maintaining a minimum time for isolation, using deoxygenated or anaerobic conditions, and using reducing agents and metal chelators). Additional studies may be employed to determine whether the protein is labeled when host cells are grown in the presence of 75Se, and whether 75Se is lost when the protein is analyzed by SDS-PAGE in the presence of a strong reducing agent. Selenium may easily form selenosulfide bonds with reactive protein thiols, and these thioselenides show lower redox potentials compared to corresponding disulfides. Therefore, proteins with reactive cysteines may appear as selenoproteins during isolation, but lose selenium during SDS-PAGE in the presence of excess reducing agent. If possible, the identity of the chemical form of selenium in a protein should be determined. For example, the presence of selenocysteine in a protein can be determined by amino acid analysis, which involves coelution of the isotope from an acid hydrolysate of alkylated 75Se-labeled protein with an alkylated selenocysteine standard. The location of selenocysteine in a protein may also be determined. This can be accomplished, for example, by isolating a 75Se-labeled peptide from a protease digest of the putative selenoprotein and subsequent direct sequencing of the peptide. Although selenocysteine is chemically destroyed during Edman degradation, eluates from each sequencing cycle may be analyzed for the presence of 75Se. Alternatively, a selenium-containing peptide
from a protease digest may be analyzed by tandem mass spectrometry, and the location of the selenocysteine residue determined from the molecular masses of degradation products (Fig. 3.8.4). The location of selenocysteine in a protein may be also deduced by aligning the gene sequence encoding a TGA codon with the expected gene product. The gene must, of course, have been identified as a selenoprotein gene according to the conserved features in Figure 3.8.3.
CONCLUSIONS Selenocysteine has been identified as the 21st amino acid in protein, and it is encoded in genomes of representatives in the three primary life domains: bacteria, archaea, and eukaryotes. As a result, a dramatic increase in research activities related to natural and artificial selenoproteins has occurred in the last several years. The largest number of selenoproteins appears to be present in vertebrate genomes. Methods that are specific for the expression, detection, and characterization of these proteins have been developed and successfully applied. Likewise, significant progress has been made in developing methods for incorporating selenium into a protein in order to obtain useful information on protein structure and properties, particularly on the environment of selenium in selenium-containing proteins. Perhaps the most extensive application of artificial selenoproteins is the use of selenomethionine-enriched proteins in Xray crystallography. Bioinformatics approaches have provided alternative and rapid procedures for identifying and characterizing new selenocysteine-containing proteins. With accelerated discoveries of new natural selenoproteins and the availability of unique methods for selenoprotein analyses, this area of research is expected to be very fruitful in the future.
LITERATURE CITED Adachi, K., Saito, H., Tanaka, T., and Oka, T. 1998. Molecular cloning and characterization of the murine staf cDNA encoding a transcription activating factor for the selenocysteine tRNA gene in mouse mammary gland. J. Biol. Chem. 273:8598-8606. Adachi, K., Saito, H., Tanaka, T., and Oka, T. 1999. Hormonal induction of mouse selenocysteine transfer ribonucleic acid (tRNA) gene transcription-activating factor and its functional importance in the selenocysteine tRNA gene transcription in mouse mammary gland. Endocrinology 140:618623. Amberg, R., Mizutani, T., Wu, X.-Q., and Gross, H.J. 1996. Selenocysteine synthesis in mammalia: An identity switch from tRNA(Ser) to tRNA(Sec). J. Mol. Biol. 263:8-19.
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Andreesen, J.R., Wagner, M., Sonntag, D., Kohlstock, M., Harms, C., Gursinsky, T., Jager, J., Parther, T., Kabisch, U., Grantzdorffer, A., Pich, A., and Sohling, B. 1999. Various functions of selenols and thiols in anaerobic gram-positive, amino acids-utilizing bacteria. Biofactors 10:263-270. Arner, E.S., Sarioglu, H., Lottspeich, F., Holmgren, A., and Bock, A. 1999. High-level expression in Escherichia coli of selenocysteine-containing rat thioredoxin reductase utilizing gene fusions with engineered bacterial-type SECIS elements and coexpression with the selA, selB and selC genes. J. Mol. Biol. 292:1003-1016. Axley, M.J., Grahame, D.A., and Stadtman, T.C. 1990. Escherichia coli formate-hydrogen lyase. Purification and properties of the selenium-dependent formate dehydrogenase component. J. Biol. Chem. 265:18213-18218. Benoit, S., Abaibou, H., and Mandrand-Berthelot, M.A. 1998. Topological analysis of the aerobic membrane-bound formate dehydrogenase of Escherichia coli. J. Bacteriol. 180:6625-6634. Berg, B.L., Li, J., Heider, J., and Stewart, V. 1991. Nitrate-inducible formate dehydrogenase in Escherichia coli K-12. I. Nucleotide sequence of the fdnGHI operon and evidence that opal (UGA) encodes selenocysteine. J. Biol. Chem. 266:2238022385. Berry, M.J., Banu, L., and Larsen, P.R. 1991. Type I iodothyronine deiodinase is a selenocysteine-containing enzyme. Nature 349:438-440. Bertini, I., Ciurli, S., Dikiy, A., Luchinat, C. 1993. The electronic structure of the [4Fe-4Se]3+ clusters in C. vinosum HiPIP and E. halophila HiPIP II through NMR and EPR studies. J. Am. Chem. Soc. 115:12020-12028. Blackburn, N.J., Ralle, M., Gomez, E., Hill, M.G., Pastuszyn, A., Sanders, D., and Fee, J.A. 1999. Selenomethionine-substituted Thermus thermophilus cytochrome ba3: Characterization of the CuA site by Se and Cu K-EXAFS. Biochemistry 38:7075-7084. Bock, A., Forchhammer, K., Heider, J., Leinfelder, W., Sawers, G., Veprek, B., and Zinoni, F. 1991. Selenocysteine: The 21st amino acid. Mol. Microbiol. 5:515-20. Boles, J.O., Tolleson, W.H., Schmidt, J.C., Dunlap, R.B., and Odom, J.D. 1992. Selenomethionyl dihydrofolate reductase from Escherichia coli. Comparative biochemistry and 77Se nuclear magnetic resonance spectroscopy. J. Biol. Chem. 267:2221722223. Boschi-Muller, S., Muller, S., Van Dorsselaer, A., Bock, A., and Branlant, G. 1998. Substituting selenocysteine for active site cysteine 149 of phosphorylating glyceraldehyde 3-phosphate dehydrogenase reveals a peroxidase activity. FEBS Lett. 439:241-245. Boyington, J.C., Gladyshev, V.N., Khangulov, S.V., Stadtman, T.C., and Sun, P.D. 1997. Crystal structure of formate dehydrogenase H: Catalysis involving Mo, molybdopterin, selenocysteine, and an Fe4S4 cluster. Science 275:1305-1308.
Buettner, C., Harney, J.W., and Larsen, P.R. 1998. The 3′-untranslated region of human type 2 iodothyronine deiodinase mRNA contains a functional selenocysteine insertion sequence element. J. Biol. Chem. 273:33374-33378. Buettner, C., Harney, J.W., and Berry, M.J. 1999. The Caenorhabditis elegans homologue of thioredoxin reductase contains a selenocysteine insertion sequence (SECIS) element that differs from mammalian SECIS elements but directs selenocysteine incorporation. J. Biol. Chem. 274:21598-21602. Burk, R.F., and Hill, K.E. 1999. Orphan selenoproteins. Bioessays 21:231-237. Chambers, I., Frampton, J., Goldfarb, P., Affara, N., McBain, W., and Harrison, P.R. 1986. The structure of the mouse glutathione peroxidase gene: The selenocysteine in the active site is encoded by the “termination” codon, TGA. EMBO J. 5:12211227. Chan, S., Gerson, B., and Subramaniam, S. 1998a. The role of copper, molybdenum, selenium, and zinc in nutrition and health. Clin. Lab. Med. 18:673-685. Chan, S., Gerson, B., Reitz, R.E., and Sadjadi, S.A. 1998b. Technical and clinical aspects of spectrometric analysis of trace elements in clinical samples. Clin. Lab. Med. 18:615-629. Chittum, H.S., Hill, K.E., Carlson, B.A., Lee, B.J., Burk, R.F., and Hatfield, D.L. 1997. Replenishment of selenium deficient rats with selenium results in redistribution of the selenocysteine tRNA population in a tissue specific manner. Biochim. Biophys. Acta 1359:25-34. Chittum, H.S., Lane, W.S., Carlson, B.A., Roller, P.P., Lung, F.T., Lee, B.J., and Hatfield, D.L. 1998. Rabbit β-globin is extended beyond its UGA stop codon by multiple suppressions and translational reading gaps. Biochemistry 37:10866-10870. Choi, I.S., Diamond, A.M., Crain, P.F., Kolker, J.D., McCloskey, J.A., and Hatfield, D.L. 1994. Reconstitution of the biosynthetic pathway of selenocysteine tRNAs in Xenopus oocytes. Biochemistry 33:601-605. Chu, F.F., Doroshow, J.H., and Esworthy, R.S. 1993. Expression, characterization, and tissue distribution of a new cellular selenium-dependent glutathione peroxidase, GSHPx-GI. J. Biol. Chem. 268:2571-2576. Clark, L.C., Combs, G.F. Jr., Turnbull, B.W., Slate, E.H., Chalker, D.K., Chow, J., Davis, L.S., Glover, R.A., Graham, G.F., Gross, E.G., Krongrad, A., Lesher, J.L. Jr., Park, H.K., Sanders, B.B. Jr., Smith, C.L., and Taylor, J.R. 1996. Effects of selenium supplementation for cancer prevention in patients with carcinoma of the skin. A randomized controlled trial. Nutritional Prevention of Cancer Study Group. J. Am. Med. Assoc. 276:1957-1963. Combs, G.F. and Combs, S.B. 1986. The Role of Selenium in Nutrition. Academic Press, New York. Commans, S. and Bock, A. 1999. Selenocysteine inserting tRNAs: An overview. FEMS Microbiol. Rev. 23:335. Conradson, S.D., Burgess, B.K., Newton, W.E., Di Cicco, A., Filipponi, A., Wu, Z.Y., Natoli, C.R.,
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Hedman, B., and Hodgson, K.O. 1994. Selenol binds to iron in nitrogenase iron-molybdenum cofactor: An extended x-ray absorption fine structure study. Proc. Natl. Acad. Sci. U.S.A. 91:1290-1293. Copeland, P.R. and Driscoll, D.M. 1999. Purification, redox sensitivity, and RNA binding properties of SECIS-binding protein 2, a protein involved in selenoprotein biosynthesis. J. Biol. Chem. 274:25447-25454. Copeland, P.R., Fletcher, J.E., Carlson, B.A., Hatfield, D.L., and Driscoll, D.M. 2000. A novel RNA binding protein, SBP2, is required for the translation of mammalian selenoprotein mRNAs. EMBO J. 19:306-314. Croteau, W., Whittemore, S.L., Schneider, M.J., and St. Germain, D.L. 1995. Cloning and expression of a cDNA for a mammalian type III iodothyronine deiodinase. J. Biol. Chem. 270:16569-16575. Daher, R. and Van Lente, F. 1994. Concanavalin Abound selenoprotein in human serum analyzed by graphite furnace atomic absorption spectrometry. Clin. Chem. 40:62-70. Davey, J.C., Becker, K.B., Schneider, M.J., St. Germain, D.L., and Galton, V.A. 1995. Cloning of a cDNA for the type II iodothyronine deiodinase. J. Biol. Chem. 270:26786-26789. Diamond, A.M., Choi, I.S., Crain, P.F., Hashizume, T, Pomerantz, S.C., Cruz, R., Steer, C.J., Hill, K.E., Burk, R.F., McCloskey, J.A., and Hatfield, D.L. 1993. Dietary selenium affects methylation of the wobble nucleoside in the anticodon of selenocysteine tRNA[Ser]Sec. J. Biol. Chem. 268:14215-14223. Dilworth, G.L. 1982. Properties of the selenium-containing moiety of nicotinic acid hydroxylase from Clostridium barkeri. Arch. Biochem. Biophys. 219:30-38. Dobbek, H., Gremer, L., Meyer, O., and Huber, R. 1999. Crystal structure and mechanism of CO dehydrogenase, a molybdo iron-sulfur flavoprotein containing S-selanylcysteine. Proc. Natl. Acad. Sci. U.S.A. 96:8884-8889. Eidsness, M.K., Scott, R.A., Prickril, B.C., DerVartanian, D.V., Legall, J., Moura, I., Moura, J.J., and Peck, H.D. Jr. 1989. Evidence for selenocysteine coordination to the active site nickel in the [NiFeSe]hydrogenases from Desulfovibrio baculatus. Proc. Natl. Acad. Sci. U.S.A. 86:147-151. Flohe, L., Gunzler, W.A., and Schock, H.H. 1973. Glutathione peroxidase: A selenoenzyme. FEBS Lett. 32:132-134. Forchhammer, K., Rucknagel, K.P., and Bock, A. 1990. Purification and biochemical characterization of SELB, a translation factor involved in selenoprotein synthesis. J. Biol. Chem. 265:9346-9350. Franke, K.W. 1934a. A new toxicant occurring naturally in certain samples of plant foodstuffs. I. Results obtained in preliminary feeding trials. J. Nutr. 8:597-608. Analysis of SelenocysteineContaining Proteins
Franke, K.W. 1934b. A new toxicant occurring naturally in certain samples of plant foodstuffs. II. The occurrence of the toxican in the protein fraction. J. Nutr. 8:609-613.
Fujiwara, K., Okamura-Ikeda, K., Packer, L., and Motokawa, Y. 1997. Synthesis and characterization of selenolipoylated H-protein of the glycine cleavage system. J. Biol. Chem. 272:19880-19883. George, G.N., Colangelo, C.M., Dong, J., Scott, R.A., Khangulov, S.V., Gladyshev, V.N., and Stadtman, T.C. 1998. X-ray absorption spectroscopy of the molybdenum site of Escherichia coli formate dehydrogenase. J. Am. Chem. Soc. 120:1267-1273. Gesteland, R.F., Weiss, R.B., and Atkins, J.F. 1992. Recoding: Reprogrammed genetic decoding. Science 257:1640-1641. Gettins, P. and Crews, B.C. 1991. 77Se NMR characterization of 77Se-labeled ovine erythrocyte glutathione peroxidase. J. Biol. Chem. 266:4804-4809. Gladyshev, V.N. and Hatfield, D.L. 1999. Selenocysteine-containing proteins in mammals. J. Biomed. Sci. 6:151-160. Gladyshev, V.N., Khangulov, S.V., and Stadtman, T.C. 1994a. Nicotinic acid hydroxylase from Clostridium barkeri: Electron paramagnetic resonance studies show that selenium is coordinated with molybdenum in the catalytically active seleniumdependent enzyme. Proc. Natl. Acad. Sci. U.S.A. 91:232-236. Gladyshev, V.N., Khangulov, S.V., Axley, M.J., and Stadtman, T.C. 1994b. Coordination of selenium to molybdenum in formate dehydrogenase H from Escherichia coli. Proc. Natl. Acad. Sci. U.S.A. 91:7708-7711. Gladyshev, V.N., Boyington, J.C., Khangulov, S.V., Grahame, D.A., Stadtman, T.C., and Sun, P.D. 1996a. Characterization of crystalline formate dehydrogenase H from Escherichia coli. Stabilization, EPR spectroscopy, and preliminary crystallographic analysis. J. Biol. Chem. 271:8095-8100. Gladyshev, V.N., Khangulov, S.V., and Stadtman, T.C. 1996b. Properties of the selenium- and molybdenum-containing nicotinic acid hydroxylase from Clostridium barkeri. Biochemistry 35:212-223. Gladyshev, V.N., Jeang, K.-T., and Stadtman, T.C. 1996c. Selenocysteine, identified as the penultimate C-terminal residue in human T-cell thioredoxin reductase, corresponds to TGA in the human placental gene. Proc. Natl. Acad. Sci. USA 93:61466151. Gladyshev, V.N., Factor, V.M., Housseau, F., and Hatfield, D.L. 1998. Contrasting patterns of regulation of the antioxidant selenoproteins, thioredoxin reductase, and glutathione peroxidase, in cancer cells. Biochem. Biophys. Res. Commun. 251:488-493. Gladyshev, V.N., Stadtman, T.C., Hatfield, D.L., and Jeang, K.T. 1999a. Levels of major selenoproteins in T cells decrease during HIV infection and low molecular mass selenium compounds increase. Proc. Natl. Acad. Sci. U.S.A. 96:835-839. Gladyshev, V.N., Krause, M., Xu, X.M., Korotkov, K.V., Kryukov, G.V., Sun, Q.A., Lee, B.J., Wootton, J.C., and Hatfield, D.L. 1999b. Selenocysteinecontaining thioredoxin reductase in C. elegans. Biochem. Biophys. Res. Commun. 259:244-249. Gladyshev, V.N., Martin-Romero, F.J., Xu, X.-M., Kumaraswamy, E., Carlson, B.A., Hatfield, D.L., and
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Current Protocols in Protein Science
Lee, B.J. 2000. Molecular biology of selenium and its role in cancer, AIDS and other human diseases. Recent Develop. Biochem. In press.
(MAD): A vehicle for direct determination of three-dimensional structure. EMBO J. 9:16651672.
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Jung, J.-E., Karoor, V., Sandbacken, M.G., Lee, B.J., Ohama, T., Gesteland, R.F., Atkins, J.F., Mullenbach, G.T., Hill, K.E., Wahba, A.J., and Hatfield, D.L. 1994. Utilization of selenocysteyltRNA[Ser]Sec and seryl-tRNA[Ser]Sec in protein synthesis. J. Biol. Chem. 269:29739-29745.
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Kabisch, U.C., Grantzdorffer, A., Schierhorn, A., Rucknagel, K.P., Andreesen, J.R., and Pich, A. 1999. Identification of D-proline reductase from Clostridium sticklandii as a selenoenzyme and indications for a catalytically active pyruvoyl group derived from a cysteine residue by cleavage of a proprotein. J. Biol. Chem. 274:8445-8454. Khangulov, S.V., Gladyshev, V.N., Dismukes, G.C., and Stadtman, T.C. 1998. Selenium-containing formate dehydrogenase H from Escherichia coli: A molybdopterin enzyme that catalyzes formate oxidation without oxygen transfer. Biochemistry 37:3518-3528. Kreimer, S. and Andreesen, J.R. 1995. Glycine reductase of Clostridium litorale. Cloning, sequencing, and molecular analysis of the grdAB operon that contains two in-frame TGA codons for selenium incorporation. Eur. J. Biochem. 234:192-199. Kryukov, G.V., Kryukov, V.M., and Gladyshev, V.N. 1999. New mammalian selenocysteine-containing proteins identified with an algorithm that searches for selenocysteine insertion sequence elements. J. Biol. Chem. 274:33888-33897. Kumaraswamy, E., Xu, X.-M., Martin-Romero, F.J., Carlson, B.A., and Hatfield, D.L. 1999. Role of UGA in mammalian protein synthesis: Selenocysteine, stop, suppression and reading gaps. Curr. Top. Biomed. Res. 1:113-123. Lee, B.J., de la Pena, P., Tobian, J.A., Zasloff, M., and Hatfield, D. 1987. Unique pathway of expression of an opal suppressor phosphoserine tRNA. Proc. Natl. Acad. Sci. U.S.A. 84:6384-6388. Lee, B.J., Worland, P.J., Davis, J.N., Stadtman, T.C., and Hatfield, D.L. 1989. Identification of a selenocysteyl-tRNASer in mammalian cells that recognizes the nonsense codon, UGA. J. Biol. Chem. 264:9724-9727. Lee, S.R., Kim, J.R., Kwon, K.S., Yoon, H.W., Levine, R.L., Ginsburg, A., and Rhee, S.G. 1999. Molecular cloning and characterization of a mitochondrial selenocysteine-containing thioredoxin reductase from rat liver. J. Biol. Chem. 274:4722-4734.
Heider, J. and Bock, A. 1993. Selenium metabolism in micro-organisms. Adv. Microb. Physiol. 35:71-109.
Leinfelder, W., Stadtman, T.C., and Bock, A. 1989. Occurrence in vivo of selenocysteyltRNA(SerUCA) in Escherichia coli. Effect of sel mutations. J. Biol. Chem. 264:9720-9723.
Hendrickson, W.A. 1991. Determination of macromolecular structures from anomalous diffraction of synchrotron radiation. Science 254:51-58.
Low, S.C. and Berry, M.J. 1996. Knowing when not to stop: Selenocysteine incorporation in eukaryotes. Trends Biochem. Sci. 21:203-208.
Hendrickson, W.A., Horton, J.R., and LeMaster, D.M. 1990. Selenomethionyl proteins produced for analysis by multiwavelength anomalous diffraction
Maenpaa, P.H. and Bernfield, M.R. 1970. A specific hepatic transfer RNA for phosphoserine. Proc. Natl. Acad. Sci. U.S.A. 67:688-695.
Detection and Assay Methods
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Martin, G.W., Harney, J.W., and Berry, M.J. 1996. Selenocysteine incorporation in eukaryotes: Insights into mechanism and efficiency from sequence, structure, and spacing proximity studies of the type 1 deiodinase SECIS element. RNA 2: 171-182.
Sorgenfrei, O., Duin, E.C., Klein, A., and Albracht, S.P. 1997. Changes in the electronic structure around Ni in oxidized and reduced selenium-containing hydrogenases from Methanococcus voltae. Eur. J. Biochem. 247:681-687. Stadtman, T.C. 1996. Selenocysteine. Annu. Rev. Biochem. 65:83-100.
Martin, G.W. III, Harney, J.W., and Berry, M.J. 1998. Functionality of mutations at conserved nucleotides in eukaryotic SECIS elements is determined by the identity of a single nonconserved nucleotide. RNA 4:65-73.
Sturchler, C., Lescure, A., Keith, G., Carbon, P., and Krol, A. 1994. Base modification pattern at the wobble position of Xenopus selenocysteine tRNA(Sec). Nucl. Acids Res. 22:1354-1358.
Maser, R.L., Magenheimer, B.S., and Calvet, J.P. 1994. Mouse plasma glutathione peroxidase. cDNA sequence analysis and renal proximal tubular expression and secretion. J. Biol. Chem. 269:2706627073.
Sun, Q.A., Wu, Y., Zappacosta, F., Jeang, K.T., Lee, B.J., Hatfield, D.L., and Gladyshev, V.N. 1999. Redox regulation of cell signaling by selenocysteine in mammalian thioredoxin reductases. J. Biol. Chem. 274:24522-24530.
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Wen, W., Weiss, S.L., and Sunde, R.A. 1998. UGA codon position affects the efficiency of selenocysteine incorporation into glutathione peroxidase-1. J. Biol. Chem. 273:28533-28541. Wilting, R., Schorling, S., Persson, B.C., and Bock, A. 1997. Selenoprotein synthesis in archaea: Identification of an mRNA element of Methanococcus jannaschii probably directing selenocysteine insertion. J. Mol. Biol. 266:637-641. Wu, X.G. and Gross, H.J. 1993. The long extra arms of human tRNA((Ser)Sec) and tRNA(Ser) function as major identity elements for serylation in an orientation-dependent, but not sequence-specific manner. Nucl. Acids Res. 21:5589-5594.
Contributed by Vadim N. Gladyshev University of Nebraska Lincoln, Nebraska Dolph L. Hatfield National Cancer Institute Bethesda, Maryland
Schwarz, K. and Foltz, C.M. 1957. Selenium as an integral part of factor 3 against dietary necrotic liver degeneration. J. Am. Chem. Soc. 79:3292-3293. Analysis of SelenocysteineContaining Proteins
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Current Protocols in Protein Science
Solid-Phase Profiling of Proteins
UNIT 3.9
Recent advances in DNA sequencing and nucleic acid microarray technologies have led to the exponential growth of information on genome structure and function. Studies involving the human genome and the publication of almost its entire sequence in the latest special issues of Nature (2001) and Science (HGP, 2001) revealed several puzzling facts. First of all, the number of functional genes in humans is much smaller than was expected. Second, the difference in the number of genes between humans and baker’s yeast is only five-fold. The difference is even less between human and mouse. These facts, in turn, raise fundamental questions—e.g., how is the diversity of the species achieved and what makes human beings human? One of the main answers to this puzzle may lay in the much higher diversity of proteins as compared to the genes encoding them, due to such processes as differential splicing and post-translational processing and modification. At the present moment our knowledge about the protein complement of the genome is quite limited, and progress in this field is directly connected to the development of new approaches and technologies. Major advances in high-throughput technologies, such as protein array technologies (also see UNIT 19.6), are needed to analyze and quantify post-translational and structural protein changes in a large collection of proteins in a relatively short period of time. The eventual objective of the post-genome era is to obtain a comprehensive understanding of the organization and dynamics of the metabolic signaling and regulatory networks of life. Protein array technologies will definitely increase our pace towards this objective. Protein array technologies can be defined as any fabrication and use of arrays containing multiple proteins bound to solid surfaces. The term “protein array” may refer to several different forms of the technology, each differing in the method of protein immobilization and detection, as well as aspects of application. Researchers are encouraged to consult this section in consideration of which type of array technologies best suit their objective of study. Three kinds of arrays are described in this introduction and are noted in Table 3.9.1. The first category of array presented is the ProteinChip (Ciphergen), which is employed to discover new biomarkers or expressed proteins as well as protein-protein interactions (see Basic Protocols 1 and 2). This is retentate chromatography on a chip. Different proteins with different surface characteristics (e.g., charge, hydrophobicity) are retained on the chip surface, which is coated with different chemical groups. The second category of array is employed for large-scale protein-protein interaction studies. Antigens or proteins are immobilized or printed on membrane or modified microscopic slides. Specific interacting proteins or antibodies from a mixture that bind to the printed proteins will then generate a signal, which is detected either by immunoassay, fluorescence imaging, or mass spectroscopy (MS; Chapter 16). These two categories are aimed at screening multiple proteins in a single sample. The third type of chip array, the tissue array, is used to screen multiple samples for changes of a single protein. This requires the researcher to have a prior knowledge of the target protein, such as the availability of specific antibody. It is important to notice (with regret) that most of the technologies described in this unit are in the process of development and are therefore not commercially available at the present time. The only exception is ProteinChip technology (Ciphergen); therefore, we concentrate below on two of the most popular applications of ProteinChip technology, Contributed by Hon-chiu Eastwood Leung, Sau-mei Leung, and Alex Karavanov Current Protocols in Protein Science (2001) 3.9.1-3.9.16 Copyright © 2001 by John Wiley & Sons, Inc.
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Table 3.9.1
Applications and Types of Arrays
Array name and type of solid surface
Applications Biomarker discovery using functional groups
Protein-protein interactions with immobilized proteins
ProteinChip: aluminum chip coated with different chemical groups (retentate chromatography on chips) Membrane immobilized with multiple proteins
Detection method
Website/Reference
Mass spectrometry
http://www.ciphergen.com
ELISA (enzyme linked immunosorbent assay)
Lueking et al. (1999)
Multiple species of proteins printed on glass slides
Screening multiple samples for changes of single biomarker
ELISA, fluorescence Haab et al. (2000), scanning, or radioactivity MacBeath and Schreiber (2000) Rolling circle amplification ELISA or fluorescence Lizardi et al. (1998), technology (RCAT): Microtiter scanning Schweitzer et al. (2000), plates immobilized with Wiltshire et al. (2000) antigen-DNA hybrid molecules ProteinChip Mass spectrometry http://www.ciphergen.com MAGIChips Fluorescence or color http://www.ipd.anl.gov/ reaction biotech/programs/biochip PROfusion technology and Fluorescence or mass http://www.phylos.com HIP chips spectrometry Tissue array: Cylinders of Immunohistochemistry Kononen et al. (1998), tissues on a glass slide or fluorescence Moch et al. (1999), microscopy Sallinen et al. (2000) PROfusion technology and Fluorescence or mass http://www.phylos.com HIP chips spectrometry ProteinChip Mass spectrometry http://www.ciphergen.com
namely, protein profiling (see Basic Protocol 1) and protein-protein interactions (see Basic Protocol 2). Other procedures will be added as the technology becomes more developed (Merchant and Weinberger, 2000; Wright et al., 2000). Chromatography on a Chip: ProteinChip Arrays The ProteinChip arrays (Ciphergen) are aluminum strips that contain 8 or 24 active spots that are 2 mm in diameter. The surface of each spot is coated with a monolayer of chemically active molecules. All spots on the array carry the same kind of active surface, which permits the study of one sample under eight different conditions or eight samples under one particular condition. These molecules can be hydrophobic (C16 aliphatic hydrocarbon chain), ionic (quaternary amine for positively charged surface or carboxylate for negatively charged surface), metal binding (nitrilotriacetate), or hydrophilic material (silicon dioxide), as depicted in Figure 3.9.1. A series of ProteinChip arrays are arranged in parallel. The raw sample solution is deposited on every spot of each array. After binding, the spots on each chip are washed with appropriate buffers of different stringency. Proteins with higher affinity to a particular surface are captured, while proteins with weak affinity are washed away; therefore, a variety of proteins are retained based on the hydrophobicity, surface charge, phosphorylation, glycosylation, or primary sequence.
Solid-Phase Profiling of Proteins
To investigate interactions with particular proteins or DNA segments, subsets of ProteinChip arrays are coated with carbonyldiimidazole or epoxy groups. These chemical groups will attack the primary amine on proteins or nucleic acids, thus forming covalent linkages with the biomolecules. Proteins interacting with these covalently bound bio-
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hydrophobic C16
hydrophilic
SiO2
C16
SiO2
SiO2
metal binding
anionic
metal ions metal metal
– CO2– – CO2–
cationic
– CO2–
–N+R 3 –N+R 3 + –N R 3
Figure 3.9.1 Types of ProteinChip array surfaces. From top to bottom, chromatographic surfaces include hydrophobic surfaces (C16 aliphatic hydrocarbon chain), hydrophilic surfaces (silicon dioxide), metal binding surfaces, anionic (carboxylate) surfaces, and cationic surfaces (quaternary amine).
molecules are pulled down and nonspecific binding proteins are removed with a washing step. Molecular masses of interacting proteins are detected by time-of-flight mass spectrometry. Applications of this technology include: biomarker discovery, protein-protein interactions, on-chip protein modification analysis, epitope mapping, tryptic peptide mapping, and identification of proteins, to name just few (for review see Merchant and Weinberger, 2000). Additional information can be obtained at http://www.ciphergen.com. OTHER PROTEIN ARRAY TECHNOLOGIES Glass surface immobilized with multiple proteins A glass surface is preferred over nitrocellulose or PVDF membrane (see below), as it has greater durability and lower intrinsic fluorescence, producing an increased signal/background ratio; however, choice of technology will depend on a balance of effort, quality, and price. Fluorescence is also more sensitive and gives a more discrete signal than that generated by enzyme-amplified chemiluminescence. Depending on the fabrication process, there are two categories of glass-surface protein microarray. The first category is a glass plate in a 96-well format formed by an enclosing Teflon mask. Each well contains four identical sets of 6 × 6 arrays (144 elements per well). The individual array element has a diameter of ∼275 µm, with a center-to-center spacing of 300 µm. The glass surface
Detection and Assay Methods
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Supplement 26
is activated sequentially by aminopropyltrimethoxysilane and N-hydroxysuccinimide. Protein or antigens are printed onto the activated glass surface by a high-speed printing robot. Specific antibodies conjugated with biotin are added to each well to detect the presence of the target protein. Streptavidin–alkaline phosphatase in the presence of alkaline phosphatase substrate generates a detectable signal. Arrays are quantified using a high-resolution scanning CCD detector. This kind of protein microarray demonstrates simultaneous detection of eight different antigens and a marker protein at concentrations higher than 6.25 µg/ml (Mendoza et al., 1999). Another category of protein microarray is essentially a comparative fluorescence assay on a microscopic slide. Hundreds of specific antibodies or antigens are printed on the surface of derivatized microscopic slides. Protein solutions to be measured are labeled separately by covalent linkage of different fluorescent dyes to the amine group on the proteins. Two differentially labeled protein solutions are mixed together and then incubated with the array, so that the fluorescence ratio at each spot corresponds to the concentration ratio of each protein in the two solutions. Nonoverlapping excitation and emission spectra give the ability to detect and measure interactions simultaneously. Experimentally, 115 pairs of antibody/antigen in an array of approximately 1100 spots were studied, and target protein as low as 1 ng/ml was successfully detected using a fluorescence scanner (Haab et al., 2000; MacBeath and Schreiber, 2000). Membrane surface immobilized with multiple proteins Polyvinylidene difluoride (PVDF) or nitrocellulose filters are loaded with specific antigens (i.e., any protein for which a specific antibody exists) at 250 nmol using an automatic picking/spotting robot. A density of approximately 300 protein samples per square centimeter is spotted onto a membrane filter. The spacing between spots is 250 µm. Filters with immobilized antigens are incubated with monoclonal antibodies and followed by secondary antibodies. Positive reactions are detected as colored spots using a chemiluminescence method. Images are acquired with a cooled charge-coupled device (CCD) detector. Ninety-two different crude lysates of human fetal brain cDNA library are detected with a high level of reproducibility (Lueking et al., 1999). The detection limit is 10 fmol/µl. Rolling circle amplification technology The goal of this technology is to increase the sensitivity of the antigen-antibody assay using in situ amplification of DNA conjugated to the antibody so that a specific antigen or combination of antigens can be detected in complex biological mixtures (e.g., cell extracts, nuclear extracts, bodily fluids). In this technique, antigens are printed on microscopic glass slides or deposited on microtiter plates as described above, and an oligonucleotide primer is covalently attached to an antibody (DNA-antibody conjugate). After the binding of the DNA-antibody conjugate molecules to the specific antigen, amplification of DNA occurs in the presence of circular DNA, DNA polymerase, and nucleotide substrates. Amplification results in a long DNA molecule containing hundreds of copies of the circular DNA sequence that remain attached to the antibody. Detection of the amplified DNA can be carried out in a variety of ways, including direct incorporation of hapten-labeled or fluorescence labeled nucleotides, or by hybridization of fluorescence-labeled or enzymatically labeled complementary oligonucleotide probes (Lizardi et al., 1998; Schweitzer et al., 2000; Wiltshire et al., 2000). Tissue Arrays Solid-Phase Profiling of Proteins
Small tissue core biopsies are punched from selected regions of donor blocks. Tissue cores are transferred into defined array coordinates in the recipient block. Tissue microarrays
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consist of as many as 1000 cylindrical tissue biopsies from individual tumors. Sections of the microarrays provide targets for parallel in situ detection of DNA, RNA, and protein targets in each specimen. Consecutive sections allow the rapid analysis of hundreds of molecular markers in the same set of specimens (Moch et al., 1999). MAGIChips This type of biochip has been developed by Biochip Technology Center at Argonne National Laboratory (http://www.ipd.anl.gov/biotech/programs/biochip) and is based on micro-acrylamide blocks. Protein (or DNA) of interest is immobilized in these blocks, divided by hydrophobic spacers. In this way, each block represents an individual protein and can be manipulated independently of a large number of other individual proteins on the same chip, or the whole array may be manipulated in the same way. The main advantage of this type of array is three-dimensional gel support for individual protein (or DNA) samples, which increases greatly the capacity for immobilized molecules. Two recent publications describe in detail the procedure for preparation of such arrays (Vasiliskov et al., 1999; Arenkov et al., 2000) and present the results of immuno- and enzymatic assays for individual proteins on these chips (Arenkov et al., 2000). This group has developed quantitative fluorescence measurement and color analysis for data collection. MS analysis is under development (Stomakhin et al., 2000). These arrays can be used for protein-protein interaction screening and functional characterization of proteins; however, the disadvantage of this approach at the present stage of development, as well as of some of the other arrays mentioned above, is the need for purified proteins to be immobilized in gel pads. PROfusion technology and HIP chips General information about this very promising technology from Phylos can be found on the company web site (http://www.phylos.com). It is based on RNA-protein fusions where a covalent link is formed between the C-terminus of the protein and the 3′ end of the mRNA that encodes it. Initially mRNA molecules carrying puromycin at the 3′ end are produced. When the ribosome reaches the 3′ end of mRNA during in vitro translation reaction, puromycin enters the peptidyl transferase site and forms a covalent mRNA-protein fusion. A light-induced psoralen cross-linking reaction to attach a puromycin containing nucleotide to the 3′ end of mRNA has been recently described (Kurz et al., 2000). This approach can be used to produce natural and artificial mRNA-protein fusion libraries. At the same time, chemical synthesis and mutagenesis are means to produce such libraries with extremely high complexities (i.e., up to 1 × 1013). Such a library printed on the chip in an addressable manner can be screened for proteins or peptides with the preferred function. The presence of mRNA on one end of the fusion permits amplification and identification of the desired target. Drug target screenings and specific protein-protein interactions are obvious applications for this type of technology. One can envision several other applications for libraries with such enormous diversity. This approach has been extensively reviewed by Roberts (1999). A detailed description of the technology (Liu et al., 2000) has also been published. STRATEGIC PLANNING Success in protein profiling for biomarker discovery requires a consideration of sample components and complexity, the surface chemistry of the ProteinChip arrays, and binding/washing conditions. Different samples may require different preparation techniques before addition to the ProteinChip array. In general, samples can be divided into two major groups: tissue and cells that require protein extraction, pretreatment, and/or fractionation (e.g., bacteria, yeast, laser capture microdissection–procured cells, cultured cells, tissue,
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plant), and fluids, either biological (e.g., serum, urine, cerebral spinal fluid, saliva) or other (e.g., cell culture medium, broth), that require pretreatment and/or fractionation. The choice between protein extraction or prefractionation is defined by the goal of the experiment. Here it should be mentioned that most conventionally used protein extraction methods are compatible with ProteinChip arrays; however, some modifications of the extraction solutions might be needed to improve the detection of proteins in final extracts. Preference should be given to nonionic detergents—e.g., Triton X-100, NP-40, n-octyl β-D-glucopyranoside (OGP)—over ionic detergents (e.g., SDS), and 2-mercaptoethanol is preferred over dithiothreitol (DTT). The use of solutions containing diethylpyrocarbonate should be avoided. A full but very short list of compounds that may interfere with protein detection on ProteinChip arrays can be found in the ProteinChip Users Guide supplied with a system. Choosing the proper control samples is extremely important for correct data interpretation and evaluation of protein profiling results. As far as protein profiling is concerned, controls are especially important for comparisons of normal versus diseased samples. A diseased state is usually accompanied by changes in protein profiles, which might not be specific for the given disease, but rather reflect the general reaction of an organism (e.g., in the case of serum, urine, cerebrospinal fluid), or the tissue microenvironment, to a harmful agent or process (e.g., laser capture microdissected samples, tumor tissue extracts); therefore, control samples should include not only adequate normal extracts, but also extracts from patients with an unrelated disease to discriminate between specific and general disease markers. BASIC PROTOCOL 1
Solid-Phase Profiling of Proteins
SERUM PROTEIN PROFILING USING PROTEINCHIP ARRAY Protein profiling uses different ProteinChip arrays and binding/washing conditions to capture different subsets of proteins from complex biological samples for comparison. Please note that, as described above (see Strategic Planning), it may be important to pretreat or fractionate samples before protein profiling to remove major proteins from samples so the minor ones can be detected (e.g., removing albumin from human serum), to fractionate complex samples into subfractions based on mass or charge in order to detect more peaks (especially high-mass proteins), and to decrease signal suppression. In addition, the proteins must be extracted and solubilized, and the samples must be prepared so that they are compatible with the chip chemistry. Finally, the samples must be diluted into the appropriate binding buffer. Materials 8 M electrophoresis grade urea (Sigma) in 1% CHAPS/1× phosphate buffered saline (PBS), pH 7.2 to 7.4 (APPENDIX 2E): prepare fresh or use aliquot stored at −70°C (do not freeze and thaw the buffer more than once) Serum sample, human Cibacron blue 3GA immobilized on 4% beaded agarose Type 3000 (Sigma) 1× PBS, pH 7.2 (APPENDIX 2E) 1 M urea/0.125% 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS)/1× PBS (APPENDIX 2E): prepare fresh or use aliquot stored at −70°C (do not freeze and thaw the buffer more than once) Buffers: pH 4: 50 mM sodium acetate: adjust pH with acetic acid; store up to 3 months at room temperature pH 6: 50 mM sodium phosphate: adjust pH with 1/7.3 (v/v) 50 mM Na2HPO4/50 mM NaH2PO4; store up to 3 months at room temperature
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pH 8: 50 mM Tris⋅HCl: adjust pH with HCl; store up to 3 months at room temperature pH 10: 100 mM 3-cyclohexamino-1-propanesulfonic acid (CAPS): adjust pH according to manufacturers’ recommendations; store up to 3 months at room temperature Saturated sinapinic acid (SPA) in 0.5% trifluoroacetic acid (TFA)/50% acetonitrile; prepare fresh daily Spin column with 0.45-µm cellulose acetate micro-spin filter (E & K Scientific) SAX2, WCX2 ProteinChip Arrays (Ciphergen Biosystems) 8-well bioprocessor (Ciphergen Biosystems) Adhesive tape Pap pen (i.e., a hydrophobic pen; Zymed) PBSII chip reader (Ciphergen Biosystems) NOTE: HPLC-quality reagents are recommended for preparation of all buffers. Dilute the sample 1. In an appropriate container, add 60 µl 8 M urea buffer to each of two 40-µl human serum samples (samples I and II; see Table 3.9.2) to be compared. 2. Vortex 10 min at 4°C. An automatic vortexer is useful for this purpose.
Prepare filter 3. Wash Cibacron blue 3GA beads (i.e., Cibacron blue 3GA immobilized on 4% beaded agarose Type 3000) five times with 1× PBS, pH 7.2. Adjust to 50% (v/v) beads/buffer with 1× PBS, pH 7.2. The Cibacron blue 3GA beads are shipped and stored in 0.5 M NaCl and 0.02% thimerosal and are used to remove albumin.
4. Add 150 µl of 50% (v/v) bead suspension into a spin column with a 0.45-µm cellulose acetate micro-spin filter. 5. Equilibrate bead suspension by adding 300 µl 1 M urea buffer and microcentrifuging to dryness at 1000 × g for 30 sec. Repeat two more times. Filter sample 6. Add 100 µl diluted serum sample (from step 2) to the filter. 7. Wash the original serum tube with 100 µl 1 M urea/0.125% CHAPS/1× PBS. Add the wash to the micro-spin filter. 8. Put the filter into a fresh collection tube. Vortex at 4°C for 15 min. 9. Microcentrifuge the filter at 1000 × g for 30 sec. Save the filtrate in the collection tube. 10. Add 100 µl of 1 M urea/0.125% CHAPS/1× PBS buffer to the column. 11. Put the filter into a fresh collection tube. Vortex 15 min at 4°C. 12. Put the filter back into the first collection tube and microcentrifuge at 1000 × g for 30 sec. The volume should be ∼300 ìl.
Load samples on the ProteinChip arrays 13. Put SAX2 (strong anionic exchange) and WCX2 (weak cationic exchange) ProteinChip arrays into an 8-well bioprocessor. Attach the top securely. 14. To each well of the arrays, add 125 µl appropriate buffer according to the profiling strategy detailed in Table 3.9.2. Seal wells with adhesive tape. Equilibrate by mixing
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Table 3.9.2 ProteinChip Serum Profiling Strategy
Spot
Samplea
Buffer
A B C D E F G H
I II I II I II I II
pH 4 pH 4 pH 6 pH 6 pH 8 pH 8 pH 10 pH 10
aSamples I and II might be any two samples used for
comparison (e.g., diseased versus normal, drug treated versus control).
at 250 rpm on a platform shaker for 5 min at room temperature. Discard the wash and repeat once with fresh buffer. 15. Dilute serum samples 1:5 with appropriate pH buffer (Table 3.9.2) yielding a final protein concentration of ∼0.5 to 1 mg/ml. For example, for a pH 4 spot, dilute the serum sample with pH 4 buffer.
16. Apply 50 µl diluted serum sample to each appropriate spot. Seal the wells with adhesive tape and mix at 250 rpm on a platform shaker for 20 min at room temperature. 17. Remove the samples from both SAX2 and WCX2 chips by aspiration. Remove unbound protein 18. Wash the chips three times, each time for 5 min, with 125 µl appropriate pH buffer (Table 3.9.2) on a platform shaker at 250 rpm. 19. Wash the chips two times with water by filling and emptying the bioprocessor wells. 20. Remove the chips from the bioprocessor and air-dry. Outline the spots with a Pap Pen. Analyze ProteinChip arrays 21. Add 0.5 µl saturated sinapinic acid (SPA) in 0.5% trifluoroacetic acid (TFA)/50% acetonitrile twice per spot, air drying between additions. 22. Air dry and read the ProteinChip arrays in a PBSII ProteinChip reader (i.e., a time-of-flight mass spectrometer specifically designed to handle ProteinChip arrays). A typical result of serum profiling using a WCX chip with four pH buffer conditions is shown in Fig. 3.9.2. The difference (indicated by arrow) of the two serum samples (with or without spiked known protein) can be clearly seen in pH 6 (spectrum C versus D) and pH 8 (spectrum E versus F) buffers.
Solid-Phase Profiling of Proteins
Other cell lysis and profiling protocols can be found in Wright et al. (2000) and von Eggeling et al. (2000). Figure 3.9.3 is an example of protein profiles from crude rat liver lysate that were obtained using three different ProteinChip arrays and wash conditions (hydrophobic surface H4 chip with 10% acetonitrile, strong anionic exchange SAX chip with pH 8.5 buffer wash, and weak cationic exchange WCX chip with pH 4.5 buffer wash). Each set of conditions was able to generate a different protein profile (e.g., H4 with 10% acetonitrile wash).
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A 75 50 25 0
B
75 50 25
C
0 75 50 25
Intensity (relative units)
D
0 75 50 25
E
0 75 50 25
F
0 75 50 25
G
0 75 50 25
H
0 75 50 25 0 1,000
12,500
15,000
17,500
Mass/charge
Figure 3.9.2 Protein profiles of serum using WCX ProteinChip array and four different pH buffer conditions. Spectrum (A), (C), (E), and (G) are serum spiked with a known protein and spectrum (B), (D), (F), and (H) are serum without the spiked protein. The difference (indicated by the arrow) of the two samples can be clearly seen in pH 6 (spectrum C versus D) and pH 8 (spectrum E versus F) buffer conditions. Detection and Assay Methods
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H4 (10% ACN wash)
20 15 10 5
SAX (pH 8.5 wash)
60 40 20 0 WCX (pH 4.5 wash)
Intensity (relative units)
0
10
5
0 5,000
10,000
15,000
Mass/charge Figure 3.9.3 Protein profiles of crude rat liver lysate in three different ProteinChip arrays and wash conditions. The upper panel depicts hydrophobic surface H4 chip using a 10% acetonitrile wash. The middle panel is a strong anionic exchanger surface (SAX) chip with a pH 8.5 buffer wash. The lower panel is a weak cationic exchange (WCX) chip with a pH 4.5 buffer wash. Arrows indicate the most prominent proteins retained on one surface but not the other.
BASIC PROTOCOL 2
PROTEIN-PROTEIN INTERACTION ANALYSIS USING PROTEINCHIP ARRAYS Protein-protein interaction studies use PS1 or PS2 ProteinChip arrays, both with preactivated surfaces. Both covalently bind any protein of interest to the surface; however, due to the different chemistry used to produce them, they have different properties. The PS-1 surface has a higher capacity, but at the same time, higher nonspecific background binding due to the hydrophobicity of the proteins from cell or nuclear lysates. The PS-2 surface binds less of the protein of interest and, as a consequence, less of the interacting partner; however, nonspecific background binding is greatly diminished as compared to PS1. Most often, the amounts of possible interacting partners in cell lysates are not known; therefore at the beginning of a study one should use both to see which chip gives better results.
Solid-Phase Profiling of Proteins
An absolute requirement for this type of experiment is that one of the interacting partners (i.e., the one to be bound) be purified or at least semipurified by conventional methods. The final preparation should not contain reagents such as Tris, glycine, or sodium azide, as these substances interfere with covalent binding to the chip surface.
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The protocol described below has been used to prove binding specificity of the orphan ligand growth factor pleiotrophin to the orphan receptor anaplastic lymphoma kinase (ALK; Stoica et al., 2001). Materials 1× phosphate buffered saline (PBS), pH 7.2 (APPENDIX 2E) 1 M ethanolamine, pH 8.2 1× PBS, pH 7.2/0.5% and 0.1% Triton X-100 5 M NaCl (APPENDIX 2E) 25% (v/v) ethylene glycol (PS1 array) Cell extract, lysate, or conditioned media (≥1 mg/ml total protein) with or without a molar excess of either the purified protein of interest or a control protein in 1× PBS, pH 7.2 Saturated α-cyano-4-hydroxycinnamic acid (CHCA) or sinapinic acid (SPA) in 0.5% TFA/50% acetonitrile PS1 or PS2 ProteinChip array (Ciphergen Biosystems) PAP pen (PS1 array) Humidified chamber: empty pipet-tip box with damp filter paper fixed on top of the stand 15-ml plastic screw-cap centrifuge tube (Corning) Platform rocker PBSII chip reader (Ciphergen Biosystems) Add probe to the ProteinChip arrays 1. For PS1 ProteinChip array: Outline each spot on a PS1 ProteinChip array with a Pap (hydrophobic) pen and air dry. The PS2 chip does not need this treatment as it has a hydrophobic coating.
2. Prewet each spot with 3 µl of 1× phosphate buffered saline (PBS), pH 7.2, for 5 min in a humidified chamber. 3. Remove PBS from the spots and replace with 1 to 2 µl purified protein of interest or control protein. 4. Place the chip into a humidified chamber and incubate 1 hr at room temperature or overnight at 4°C. The latter is preferable as it ensures complete saturation.
5. Remove the chip from the humidified chamber and remove protein solution from spots by aspiration with a micropipettor. Inactivate surface and remove unbound protein 6. Add 5 µl of 1 M ethanolamine, pH 8.2 per spot, and incubate in a humidified chamber for 30 min at room temperature. 7. Remove ethanolamine solution by aspiration with a micropipettor and wash each spot twice with 1× PBS, pH 7.2/0.5% Triton X-100. IMPORTANT NOTE: Perform this step spot-by-spot without allowing spots to become dry at any time.
8. Place the chip into a 15-ml plastic screw-cap centrifuge tube and add 10 ml 1× PBS, pH 7.2/0.5% Triton X-100. Place the tube on a platform rocker and wash for 10 to 15 min. Repeat the wash one more time. Detection and Assay Methods
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A
2
15868.5+
1
Intensity (relative units)
0
B
2 15874.4+ 1 0
C
2 1 15849.7+ 0
D
2 1
15852.5+
0 25,000
50,000
75,000
100,000
Mass/charge
Figure 3.9.4 (A) Profile of PTN fraction purified on heparin-Sepharose using a normal phase chip (NP-2). (B to D) ALK ECD covalently bound to active spots on preactivated surface (PS1) chip and overlaid with: (B) PTN, (C) PTN+ALK ECD ∼1/1.66 molar ratio, and (D) PTN+TGFβ receptor ∼1/1.7 molar ratio.
9. Transfer the chip into a fresh 15-ml plastic screw-cap centrifuge tube. Add 10 ml 1× PBS, pH 7.2/0.1% Triton X-100 and incubate 5 min. Repeat wash one more time on a platform rocker. Add sample to the ProteinChip arrays 10. Adjust the salt concentration of cell extracts or lysates to 0.15 to 0.2 M NaCl/0.1% Triton X-100 by diluting with the appropriate buffer or adding 5 M NaCl. 11. Remove the chip from the centrifuge tube. To each spot, add 3 to 5 µl cell extract, lysate, or conditioned medium (≥1 mg/ml total protein) with or without a molar excess of either the purified protein of interest or a control protein in 1× PBS, pH 7.2, relative to the protein covalently bound to the surface. 12. Incubate for 1 hr in a humidified chamber. 13. Remove extracts from the spot by aspiration with a micropipettor and wash each spot with 5 µl 1× PBS, pH 7.2/0.5% Triton X-100 for the PS1 chip or 1× PBS, pH 7.2/0.1% Triton X-100 for the PS2 chip.
Solid-Phase Profiling of Proteins
14. Wash the entire array two times with 10 ml 1× PBS, pH 7.2/0.5% Triton X-100 for the PS1 chip or 1× PBS, pH 7.2/0.1% Triton X-100 for the PS2 chip for 10 to 15 min each on a platform rocker.
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15. For PS1 chip only: Wash the array with 10 ml 25% (v/v) ethylene glycol for 5 min. 16. Wash the entire array with 10 ml 1× PBS, pH 7.2 two times for 5 min each. 17. Rinse the array with deionized water two times and air dry. 18. Add 0.5 µl saturated SPA or CHCA in 0.5% TFA/50% acetonitrile twice per spot. Air dry after each addition. Read ProteinChip arrays 19. Read the chips in a PBSII ProteinChip reader. The results of this type of an experiment are shown in Fig. 3.9.4. In this experiment the goal was to prove the specificity of interaction of growth factor pleiotrophin (PTN) with the extracellular domain of ALK receptor protein (ALK-ECD). A commercially available preparation of TGF-β receptor II (Sigma) was used as a control. For Ab capture of protein, the same protocol is applicable. In this case a solution of a specific or control Ab is bound to the chip surface in step 3.
COMMENTARY Background Information Protein array technologies refer to any fabrication and use of arrays containing multiple proteins captured on solid surfaces. This valuable tool is used for high-throughput proteinprotein interaction studies, protein-marker discovery, and other related applications. Although not all of the recently emerged protein array technologies and related detection methods are commercially available, they are noted in this unit. Currently, this unit presents some ProteinChip array (Ciphergen) procedures. Microarray chips are valuable tools for biomarker discovery and protein-protein interaction studies. Protein profiling has been used for finding candidate proteins for soft tissue regeneration (Li et al., 2000) and cancer biomarker discovery, as well as for finding candidate proteins in serum (i.e., “serum profiling”; Wright et al., 2000) or cells procured by laser capture microdissection (LCM; Paweletz et al., 2000; von Eggeling et al., 2000).
Critical Parameters ProteinChip technology and its software is very user-friendly; however, at the least, a very basic knowledge of protein chemistry, chromatography, and biochemistry is required of the user. The protein profiling (i.e., biomarker discovery) application is quite empirical by definition, unless the design of the experiment is dictated by some preliminary data that determine the type of the surface to be used. Usually, success is dependent on the variety of ProteinChip arrays used and the choice of appropriate
control samples. One has to keep in mind that, notwithstanding the enormous advantage conferred by the ability to wash out contaminants from the surface of ProteinChip array, the detection step is still based on mass spectrometry; therefore, top-quality reagents are of the highest priority for all ProteinChip experiments. Reproducibility is highly dependent on the precision of small-volume pipet. Using the special bioprocessor device supplied with the technology, it is possible to apply 50 to 400 µl sample per spot. This range of amounts is highly recommended, especially in cases where samples are very dilute or target components are present at low concentrations (e.g., a conditioned media analysis from cell cultures). In protein-protein interaction experiments, the strength of the interaction and variation of the stringency of washes (detergent and salt concentration) is critical for the success of the experiment. The choice between a PS1 or PS2 ProteinChip array is empirical at the beginning; however, the different chemistries of these two surfaces dictate a preference for the PS2 ProteinChip array when fishing for partners of short peptides. This is due to the longer distance of active chemical groups from the surface provided by this type of array.
Troubleshooting ProteinChip technology is complex, and its performance is influenced by quite a number of factors; therefore, it is almost impossible to give a complete list of all possible factors that may interfere with the results. Table 3.9.3 presents a list of the most common potential prob-
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Table 3.9.3 Troubleshooting Guide for Protein Profiling and Protein-Protein Interaction Studies
Application
Problem
Solution
Apply more sample by performing multiple additions. Incubate sample on spot, remove sample, apply a second addition of fresh sample, and repeat incubation. Perform more additions if necessary. Increase or decrease concentration of sample. Check extraction buffer composition for the presence of ion-suppressive substances. Fractionate sample to resolve more peaks and more high mass peaks. Spin columns for size fractionation or Q-columns for fractionation by charge can be used. Use fresh; most diluted stains are supplemented with protease inhibitors to avoid proteolysis. Micro-dissect cell for as short time as possible (10 to 15 min). Do the protein profiling immediately after the microdissection. Reproducibility Make sure the addition of energy absorbing problems molecule (e.g., sinapinic acid) is consistent from spot to spot. Avoid spreading by drawing PAP pen circle around spots. Check micropipettor calibration.a No specific Modify binding conditions and decrease wash binding stringency. Check antibody specification for the presence of sodium azide or BSA. Sodium azide may be removed by microdialysis. Purify Ab on protein G column. Change the source of Ab.
Protein profiling Small number of peaks are detected
LCM samples profiling
Protein-protein interaction Ab approach
aThe Rainin P-2 and P-10 Pipetman micropipettors have an error of 25% (see manufacturer’s specifications
sheet); therefore, the precision of the micropipettor is of importance, especially when proteins of interest are in low abundance.
lems and their solutions. More information can be found in the ProteinChip Users Guide, supplied with the technology, and in specification sheets supplied with each type of ProteinChip array. As nonspecific background binding of proteins from cell lysates or other biological mixes always accompanies specific binding on ProteinChip arrays, interpretation of the results depends strongly on the type of control experiments used in each particular assay. In general, there are two kinds of binding studies, as described in the following two sections.
Solid-Phase Profiling of Proteins
Direct binding experiments In direct binding experiments, the purified protein of interest is covalently bound to the chip and lysates or extracts are run over the
surface. Two types of control experiments are proper for this approach: (1) a purified protein which does not interact with the ligand, but which has known protein binding properties, is bound to another spot on the same array or (2) a competition experiment where the same protein bound to the chip is spiked into the lysate or extract in molar excess and run over the spot. In the latter case, binding to the protein of interest should be considered specific if the peak(s) on the spot with protein of interest is/are unique or greatly enhanced during the experiment, and if, in control experiments, binding to the protein of interest is reduced by a competitor but not by a control protein.
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Indirect binding experiments In indirect binding experiments, a specific antibody to a protein of interest is covalently bound to the chip and lysates or extracts are run over the spot to fish for the proteins interacting with a protein of interest. In this case, necessary controls should include binding to a nonspecific antibody of the same type (monoclonal or polyclonal) and produced in the same species as the specific antibody.
Anticipated Results Typical results that are possible with this technology are presented in Figures 3.9.2, 3.9.3, and 3.9.4.
Time Considerations Generally, ProteinChip technology is highly time-saving. Most of the experiments do not take more than 2 to 3 hr, not including overnight incubations, which might be needed for protein-protein interaction experiments and especially for antibody capture experiments. For instance, checking the fractions from column chromatography takes no more than 15 to 20 min instead of 6 to 8 hr by SDS-electrophoresis. As some applications include air-drying of the sample on spots of the array, factors such as the humidity in the room might influence the length, but not the quality, of the experiment.
Literature Cited Arenkov, P., Kukhtin, A., Gemmell, A., Voloshchuk, S., Chupeeva, V., and Mirzabekov, A. 2000. Protein microchips: Use for immunoassay and enzymatic reactions. Anal. Biochem. 278:123-131. Haab, B.B., Dunham, M.J., and Brown, P.O. 2000. Protein microarrays for highly parallel detection and quantitation of specific proteins and antibodies in complex solutions. Genome Biology. 2: research0004.1-0004.13 (available online at http://genomebiology.com/2001/2/2/research/ 0004). Human Genome Project (HGP) 2001. The human genome. Science genome map. Science 291:1219-1224. Kononen, J., Bubendorf, L., Kallioniemi, A., Bärlund, M., Schraml, P., Leighton, S., Torhorst, J., Mihatsch, M.J., Sauter, G., and Kallioniemi, O.P. 1998. Tissue microarrays for high-throughput molecular profiling of tumor specimens. Nature Med. 4:844-847. Kurz, M., Gu, K., and Lohse, P.A. 2000. Psoralen photo-crosslinked mRNA-puromycin conjugates: A novel template for the rapid and facile preparation of mRNA–protein fusions. Nucl. Acids Res. 28:e83.
Li, X., Mohan, S., Gu, W., Miyakoshi, N., and Bayling, D.J. 2000. Differential protein profile in the ear punched tissue of regeneration and non-regeneration strains of mice: A novel approach to explore the candidate genes for softtissue regeneration. Biochim. Biophys. Acta 1524:102-109. Liu, R., Barrick, J.E., Szostak, J.W., and Roberts, R.W. 2000. Optimized synthesis of RNA-protein fusions for in vitro protein selection. Methods Enzymol. 318:268-293. Lizardi, P.M., Huang, X., Zhu, Z., Bray-Ward, P., Thomas, D., and Ward, D.C. 1998. Mutation detection and single-molecule counting using isothermal rolling-circle amplification. Nat. Genet. 19:225-232. Lueking, A., Horn, M., Eickhoff, H., Büssow, K., Lehrach, H., and Walter, G. 1999. Protein microarrays for gene expression and antibody screening. Anal. Biochem. 270:103-111. MacBeath, G. and Schreiber, S.L. 2000. Printing proteins as microarrays for high-throughput function determination. Science 289:1760-1763. Mendoza, L.G., McQuary, P., Mongran, A., Gangadharan, R., Brignac, S., and Eggers, M. 1999. High-throughput microarray-based enzymelinked immunosorbent assay (ELISA). Biotechniques 27:778-780, 782-786, 788. Merchant, M. and Weinberger, S. 2000. Recent advancements in surface-enhanced laser desorption/ionization-time of flight-mass spectrometry. Electrophoresis 21:1164-1177. Moch, H., Schraml, P., Bubendorf, L., Mirlacher, M., Kononen, J., Gasser, T., Mihatsch, M.J., Kallioniemi, O.P., and Sauter, G. 1999. Highthroughput tissue microarray analysis to evaluate genes uncovered by cDNA microarray screening in renal cell carcinoma. Am. J. Pathol. 154:981986. Nature 2001. Human genomes public and private. Nature 409:745-964. Editorial. Paweletz, C.P., Gillespie, J.W., Ornstein, D.K., Simone, N.L., Brown, M.R., Cole, K.A., Wang, Q.H., Huang, J., Hu, N., Yip, T.T., Rich, W.E., Kohn, E.C., Linehan,W.M., Weber, T., Taylor, P., Emmert Buck, M.R., Liotta, L.A., and Petricoin, E.F. III. 2000. Rapid protein display profiling of cancer progression directly from human tissue using a protein biochip. Drug Dev. Res. 49:3442. Roberts, R.W. 1999. Totally in vitro protein selection using mRNA-protein fusions and ribosome display. Curr. Opin. Chem. Biol. 3:268-273. Sallinen, S., Sallinen, P.K., Haapasalo, H.K., Helin, H.J., Helén, P.T., Schraml, P., Kallioniemi, O., and Kononen, J. 2000. Identification of differentially expressed genes in human gliomas by DNA microarray and tissue chip techniques. Cancer Res. 60:6617-6622.
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Schweitzer, B., Wiltshire, S., Lambert, J., O’Malley, S., Kukanskis, K., Zhu, Z., Kingsmore, S.F., Lizardi, P.M., and Ward, D.C. 2000. Immunoassays with rolling circle DNA amplification: A versatile platform for ultrasensitive antigen detection. Proc. Natl. Acad. Sci. U.S.A. 97:1011310119. Stoica, G.E., Kuo, A., Aigner, A., Sunitha, I., Soutton, B., Malerczyk, C., Caughey, D.J., Wen, D., Karavanov, A., Riegel, A.T., and Wellstein, A. 2001. Identification of ALK (anaplastic lymphoma kinase) as a receptor for the growth factor pleiotrophin. J. Biol. Chem. In press. Stomakhin, A.A., Vasiliskov, V.A., Timofeev, E., Schulga, D., Cotter, R.J., and Mirzabekov, A.D. 2000. DNA sequence analysis by hybridization with oligonucleotide microchips: MALDI mass spectrometry identification of 5mers contiguously stacked to microchip oligonucleotides. Nucl. Acids Res. 28:1193-1198. Vasiliskov, A.V., Timofeev, E.N., Surzhikov, S.A., Drobyshev, A.L., Shick, V.V., and Mirzabekov, A.D. 1999. Fabrication of microarray of gel-immobilized compounds on a chip by copolymerization. Biotechniques 27:592-594, 596-598, 600 passim.
von Eggeling, F., Davies, H., Lomas, L., Fiedler, W., Junker, K., Claussen, U., and Ernst, G. 2000. Tissue specific microdissection coupled with ProteinChip Array technologies: Applications in cancer research. Biotechniques 29:1066-1070. Wiltshire, S., O’Malley, S., Lambert, J., Kukanskis, K., Edgar, D., Kingsmore, S.F., and Schweitzer, B. 2000. Detection of multiple allergen-specific IgEs on microarrays by immunoassay with rolling circle amplification. Clin. Chem. 46:19901993. Wright, G.L. Jr., Cazares, L.H., Leung, S.M., Nasim, S., Adam, B., Yip, T., Schellhammer, P.F., Gong, L., and Vlahou, A. 2000. ProteinChip surface enhanced laser desorption/ionization (SELDI) mass spectrometry: A novel protein biochip technology for detection of prostate cancer biomarkers in complex protein mixtures. Prostate Cancer Prostatic Diseases 2:264-276.
Contributed by Hon-chiu Eastwood Leung, Sau-mei Leung, and Alex Karavanov Ciphergen Biosystems Inc. Fremont, California
The authors are grateful for helpful comments provided by Christine Yip and to Dr. Huw Davies for providing Figure 3.9.2.
Solid-Phase Profiling of Proteins
3.9.16 Supplement 26
Current Protocols in Protein Science
CHAPTER 4 Extraction, Stabilization, and Concentration INTRODUCTION
R
esearchers can chose from a variety of options when preparing source material to obtain purified proteins. The most practical and productive option is usually to apply recombinant DNA technology, as described in Chapters 5 through 7. There are many advantages of this approach, including the ability to express, in large amounts, proteins that exist only in trace amounts in natural sources. Such proteins may have only been identified by functional assays or analytical detection methods, or at the extreme, as a potential product of a gene’s open reading frame. Proteins prepared by recombinant technology can be used for functional and structural characterization of the protein, as well as to prepare antibodies that can be used to explore a protein’s expression patterns in tissues and cellular organelles. Such information plays a key role in assessing biological functions.
In certain cases, recombinant technology may not be a viable option for protein purification. This is particularly true for cases in which it is desirable to obtain a protein that has its natural post-translational modifications; this is usually not possible with recombinant expression systems. Although a problem of rapidly decreasing proportion, recombinant technology is also not applicable for proteins whose parent gene remains to be identified. For such proteins, tissues and/or cultured cells may serve as a convenient source of starting material. Purification from these sources often begins with cellular fractionation or organelle isolation as a means of enriching for the protein of interest. The first part of this chapter provides methodology for this approach. UNIT 4.1 provides an overview of the principles and procedures of centrifugation that are heavily relied upon for cellular fractionation. UNIT 4.2 provides a range of different procedures that have been used to fractionate tissue homogenates to obtain organelles. Included are criteria for assessing the purity of subcellular fractions and methods for differentiating integral and peripheral membrane proteins. Although the procedures are based on rat liver as starting material, they can be readily modified and optimized for a wide range of tissue and cell types. UNIT 4.3 gives protocols for the immunoisolation of early and late endosomes under conditions where these compartments retain their capacity to support membrane transport in vitro. Criteria for characterizing the purity of endosomal fractions are also provided. For additional purification steps, analyses, storage, and/or examination of structural and functional properties, proteins need to be concentrated and/or the solvent exchanged. Consequently, UNIT 4.4 describes procedures for desalting, concentrating and buffer exchange by dialysis and ultrafiltration. In conjunction with these protocols, UNIT 4.5 provides protocols for the selective precipitation of proteins that can be used for contaminant removal, concentration, and buffer exchange. Finally, UNIT 4.6 deals with the long-term storage of proteins focusing on maintaining structural and functional integrity. The stresses that induce protein aggregation and damage are addressed, including a description of the major pathways of chemical degradation. This is followed by a discussion of suitable long-term storage conditions. John E. Coligan Contributed by John E. Coligan Current Protocols in Protein Science (2002) 4.0.1 Copyright © 2002 by John Wiley & Sons, Inc.
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Overview of Cell Fractionation Cell fractionation has enjoyed widespread use among cell biologists for more than half a century. It continues to be a fruitful, if not essential, approach in the reductionistic efforts to define the composition and functions of the multiple compartments in eukaryotic cells. It also provides the essential ingredients for cell-free assays that are now being used in test-tube reconstructions of complex cellular events involving intercompartmental interactions. Much of the knowledge regarding the composition and function of cell organelles has resulted from fractionation of mammalian tissues, where cells are both abundant and highly differentiated (and thus organelle-rich). However, with widely used techniques in molecular biology and established interest in combining studies of intact cells and functional reconstitution, interest in fractionation has spread to cultured cell lines and genetically tractable lower eukaryotic cells. The goals of cell fractionation often differ depending on the nature of the experiments being conducted. In preparative procedures, in which the intent is to isolate quantities of a particular cell organelle for further study or for subfractionation, the emphasis is on purity and (secondarily) on yield. This is a primary goal in samples now used as starting material for organelle proteomic analyses (e.g., Wu et al., 2000, Bell et al., 2001, Cronshaw et al., 2002, Wasiak et al., 2002). In analytical experiments, in which the intent is not isolation of organelles but evaluation of associations of selected macromolecules with particular organelles, the emphasis is on using one-step procedures that result in different distributions of various organelles (as defined by marker activities) rather than on separating organelles outright. Finally, in preparing organelles for cell-free reconstitution, the goal is to maintain them in a functional state. The investigator generally has developed a specific assay for intercompartmental interaction that does not rely on organelle purity, so the extent of contamination by irrelevant organelles is less important. The separation of distinct organelles during cell fractionation results from their differing physical properties—size (and shape), buoyant density, and surface charge density—which reflect their differing compositions. Particular fractionation techniques capitalize on one or more of these properties. For example, gel fil-
Contributed by J. David Castle Current Protocols in Protein Science (2004) 4.1.1-4.1.9 C 2004 by John Wiley & Sons, Inc. Copyright
tration separates on the basis of size, centrifugation separates on the basis of size and density, and electrophoresis separates on the basis of surface charge density. As knowledge of the specific composition of particular organelles has developed, it has become possible to apply newer techniques such as affinity chromatography and selective density-shift perturbation. In UNIT 4.2, examples are given of fractionation procedures based on each of these approaches. Some are preparative and others are analytical. Although an effort has been made to present procedures that are widely applicable and accessible and that do not require highly specialized and expensive equipment, the procedures may have to be modified to adapt them to individual needs. Even isolation of a particular kind of organelle from different tissue or cell sources may necessitate adjustment of fractionation conditions. This means that in addition to isolating the organelle of interest, one should also plan on confirming the identity of what you have isolated. Centrifugation is the most widely used procedure in cell fractionation, as indicated in UNIT 4.2. It is the only approach commonly used to separate crude tissue homogenates (often having quite large volumes) into subfractions as starting material for more refined purification procedures. Further, the technology available, using rotors with a variety of geometries and diverse media that enable separation according to size, density, or both, now routinely permits refined separations on volumes ranging from submilliliter to several liters. No other technology is this versatile. Gel filtration is limited by the pore sizes of available resins, such that only regularly shaped vesicles with diameters <100 to 200 nm can be purified away from larger or irregularly shaped organelles. Use of electrophoresis for organelle purification, especially at the preparative level, is relatively recent, and, as with gel filtration, its success relies on generating starting material by centrifugation. Although it has shown promise for purifying selected organelles (e.g., endosomes) that otherwise require organellespecific strategies (see UNIT 4.3), surface charge densities on most organelles may not be different enough (or able to be manipulated sufficiently) to make this a versatile procedure. For these reasons, the remaining comments in this overview focus primarily on fractionation
UNIT 4.1
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4.1.1 Supplement 37
of organelles by centrifugation. For more detailed coverage of each of the topics to be discussed, the reader is referred to the excellent book edited by Rickwood (1984).
BASIC PRINCIPLES OF CENTRIFUGATION One of the best ways to understand the rationale for various cell fractionation procedures is to examine the Svedberg equation (Equation 4.1.1), which describes mathematically the sedimentation of a spherical particle in a viscous fluid:
Equation 4.1.1
Overview of Cell Fractionation
where x is the distance from axis of rotation, t is time, ω is the angular velocity, r is the radius of the particle, ρp and ρm are the density of the particle and medium, and η is the viscosity of the medium. This equation states that the sedimentation velocity, dx/dt, per unit centrifugal field ω2 x (which is set by the instrumentation) increases with the square of the particle radius and the difference in density between the particle and the medium and decreases with increasing viscosity. In practical terms, when sedimentation is performed by differential velocity in a medium such as 0.25 M sucrose, which is less dense than all particles and has a low viscosity, the radius of the particle is the dominant factor that determines the sedimentation rate. In contrast, when isopycnic density gradient centrifugation is used, the density of the medium changes from top to bottom in the centrifugation tube, and the range of densities typically includes the buoyant densities of most particles (vesicles). Thus individual types of vesicles will sediment until ρp = ρm , and the vesicles will then stop at their isopycnic densities. Alternatively, in flotation procedures ρp < ρm initially and dx/dt is negative, meaning that particles will move toward the axis of rotation (i.e., toward the top of the tube). Evidently, the equation also indicates that sedimentation or flotation, as the case may be, is slowed in media of higher viscosity. As discussed below, various media used in centrifugation are distinguished by their viscosities (and their osmolarities) at high densities. The Svedberg equation can also be used to discuss three other basic relationships encountered in using centrifugation. First, the velocity of a particle per unit centrifugal field (left
side of the equation) is the sedimentation coefficient. Its dimensions are in seconds, and it is generally reported in units known as Svedbergs (S). 1 S = 10−13 sec, the same order of magnitude as the sedimentation coefficient of several biological macromolecules (e.g., 5S RNA). Second, the centrifugal field, ω2 x is usually normalized to gravity and is expressed in the literature as a relative centrifugal force (RCF = ω2 x/g). Centrifugation conditions are best reported as × g. RCF is usually expressed at the midpoint of the tube (gav ). The centrifugal field term includes the angular velocity in units of radians per second, which is easily converted to revolutions per minute (rpm), the unit most often given in the literature, as follows: RCF = 11.18 r(N/1000)2 , where N is rpm. Third, integration of the Svedberg equation over time yields the following relationship (Equation 4.1.2):
Equation 4.1.2
This is a measure of the clearance time, the time for a particle of given sedimentation coefficient to travel from the top to the bottom of the tube. Included in k are the geometry of the rotor (Xmax and Xmin ) and the angular velocity. kfactors are reported in the printed information that accompanies most rotors. They are useful for estimating the length of centrifugation required to pellet a particular particle and for translating centrifugation conditions between two rotors. For a given particle or vesicle, k/t (rotor 1) = k/t (rotor 2).
INSTRUMENTATION Several pieces of equipment must be available to carry out cell fractionation by centrifugation. These include homogenizers, centrifuges (generally one low-speed and one ultracentrifuge) with compatible rotors, a refractometer for measuring the refractive index of centrifugation media (before centrifugation) and gradient fractions (after centrifugation), a gradient-forming device for generating preformed density gradients, and a gradientcollecting device for unloading the gradients after the run. The refractometer, gradientforming device, and gradient-collection device are very useful, if not essential, for density gradient and rate zonal sedimentation (UNIT 4.2).
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Centrifuges Most organelle purification procedures require that samples be maintained at ≤4◦ C, so the centrifuges used need to be refrigerated. The low-speed centrifuge is used in the early steps of the typical purification procedure for sedimenting large particulates such as nuclei and unbroken cells, frequently from large volumes (e.g., UNIT 4.2). The ultracentrifuge is used for subsequent spins at higher speeds and often with smaller volumes. These instruments have a vacuum system as the rotors are spun in an evacuated chamber to reduce friction (and thus heating).
Rotors Commonly used rotors come in three configurations: fixed-angle, vertical (or near-vertical), and swinging-bucket (Fig. 4.1.1). Continuous-flow and zonal rotors, which are loaded and unloaded during the run, are less widely used. Fixed-angle rotors emphasize speed and capacity. Many of these rotors generate RCFs in excess of 600,000 × g, meaning low k-factors, and can handle a few hundred milliliters of solution at one time. As a result, they are used frequently for pelleting fractions, particularly from large volumes. Most fixed-angle rotors hold tubes in a fixed incline substantially away from the vertical. When the centrifugal force is applied, it is radial rather than along the length of the tube. Thus the path length is across the tilted tube and is reasonably short; particles travel outward until they hit the outer wall and then “slide” down the wall to form a pellet (see Fig. 4.1.1). In vertical rotors, the path is only the width of the tube. As is evident from the figure, in both fixed-angle and vertical rotors the solution in the tubes reorients from the horizontal to the vertical during acceleration and reverses during deceleration. These rotors can be advantageous for density gradients (UNIT 4.2) because equilibrium is reached quite rapidly with the short path, and the separation of bands of different density increases as the contents of the tube reorient during deceleration. With swinging-bucket rotors, the tube and its contents reorient from vertical to horizontal during acceleration and reverse during the end of deceleration. The centrifugal force in this case is along the length of the tube. Note, however, that because the force is directed radially, sedimenting particles tend to concentrate peripherally near the tube walls. Also, the long path length and the generally lower centrifugal forces required for these rotors mean that k-factors are usually much higher than with
fixed-angle rotors. Thus these rotors have been used more for density gradient and rate zonal centrifugations and less for pelleting fractions.
Refractometers A refractometer measures refractive indices of solutions used for organelle separations. Refractive index is linearly related to density, and tables relating these two parameters over a large range of concentrations have been prepared for sucrose (Table 4.1.1) and for other commonly used media—e.g., contact Nycomed Pharma for concentration/refractive index data for metrizamide, Nycodenz, and Optiprep (iodixanol). Alternatively, tables analogous to Table 4.1.1 are readily constructed by weighing aliquots of other accurately prepared gradient solutions. The measurement of refractive index/density is particularly useful for characterizing the position of organelles in gradients after centrifugation and for ensuring reproducibility of gradients from one experiment to another.
Gradient-forming devices These devices consist of two chambers with an interconnecting channel along the bottom and an outlet from one chamber that connects to a peristaltic pump on its way to the centrifuge tube (see Fig. 4.1.2). The channel is gated by a stopcock. For linear gradients (the type most frequently used), the chambers have the same cross-sectional area. Solutions representing the extremes in density of the desired gradient are each placed in one chamber, and a stirring device is placed in the chamber with the outlet going to the tube. As fluid is pumped out of the mixing chamber, fluid also flows through the connecting channel from the reservoir to maintain the same pressure head in the two chambers. Consequently, the density of the solution in the mixing chamber progressively changes. Gradients can be generated either highest density first, if the higher-density solution is placed in the mixing chamber, or lowest density first, if the lower-density solution is placed in the mixing chamber. Note that for highest density first, the delivery point must be located above the level of the solution filling the centrifuge tube, whereas for lowest density first, the delivery point must be at the bottom of the tube. Gradient-collection devices. Gradient collection at the end of centrifugation requires a systematic and reproducible means of emptying the tube. Gradients can be collected from the top or from the bottom. With care, a pipettor (e.g., Pipetman, Rainin) of suitable volume can be used for manual collection from the
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Figure 4.1.1 Rotors used in ultracentrifuges. (A) Fixed-angle rotor. In each profile, the tube on the left shows pelleting from a uniform suspension and the tube on the right shows sedimentation of a band layered at the top above a higher-density fluid. Sedimentation is radial (particles move toward the outer wall of the tube and then down this wall to the pellet). Note how the band in the right tube reorients during the spin and then again at the end of the run. (B) Vertical rotor. The rotor profile shows tubes with sample layered above a higher-density solution before a run. Note that during the run (successive tube profiles shown at right), the layer reorients along the inner wall of the tube and then the bands resolve vertically. During deceleration the bands reorient to a horizontal position. (C) Swinging-bucket rotor. Note the tube on the left in the upper profile has a sample layered at the top. The buckets are mounted vertically on the rotor. During the run, the buckets reorient to the horizontal position, and the bands separate along the length of the tube. Because sedimentation is radial, the particulates in the bands are more concentrated at the walls facing and away from the viewer than in the center of the tube. Toward the end of deceleration, the buckets reorient to the vertical position.
Overview of Cell Fractionation
top, taking care to withdraw the equal-volume fractions from just below the meniscus. Alternatively, Buchler markets a pumping device (Autodensiflow) that continually adjusts to the height of the meniscus and withdraws gradients from the top, and both MSE Scientific and Nyegaard market devices that pump highdensity solution beneath the gradient and pro-
gressively displace the gradient, which is again collected at the top. For collection from the bottom, the centrifuge tube is punctured with a needle and the gradient is collected dropwise.
FRACTIONATION MEDIA Early in the development of centrifugation as a tool for purification of cellular organelles,
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Current Protocols in Protein Science
Table 4.1.1 Density and Refractive Index of Aqueous Sucrose Solutions at 20◦ Ca
Percent (w/w)
Molarity
Density
Refractive index
Percent (w/w)
Molarity
Density
Refractive index
2
0.06
1.006
1.3359
38
1.30
1.166
1.3958
4
0.12
1.014
1.3388
40
1.38
1.176
1.3997
6
0.18
1.022
1.3418
42
1.46
1.187
1.4036
8
0.24
1.030
1.3448
44
1.54
1.197
1.4076
10
0.30
1.038
1.3479
46
1.62
1.208
1.4117
12
0.37
1.046
1.3510
48
1.71
1.219
1.4158
14
0.43
1.055
1.3541
50
1.80
1.230
1.4200
16
0.50
1.064
1.3573
52
1.88
1.241
1.4242
18
0.56
1.072
1.3606
54
1.98
1.252
1.4285
20
0.63
1.081
1.3639
56
2.07
1.263
1.4329
22
0.70
1.090
1.3672
58
2.16
1.275
1.4373
24
0.77
1.099
1.3706
60
2.26
1.286
1.4418
26
0.84
1.108
1.3740
62
2.35
1.298
1.4464
28
0.91
1.118
1.3775
64
2.45
1.310
1.4509
30
0.99
1.127
1.3811
66
2.55
1.322
1.4558
32
1.06
1.137
1.3847
68
2.65
1.335
1.4605
34
1.14
1.146
1.3883
70
2.76
1.347
1.4651
36
1.22
1.156
1.3920
a Modified from Hofmann (1977) by permission of ISCO, Inc.
Figure 4.1.2 Linear gradient-forming device showing the back reservoir connected to the mixing chamber via a channel with a stopcock. A delivery tube leads from the mixing chamber to a peristaltic pump and then to the centrifuge tube. (A) Configuration used when the lower-density solution is delivered first; the lower-density solution is in the mixing chamber and the outlet is at the bottom of the centrifuge tube. (B) Configuration used when the higher-density solution is delivered first; the higher-density solution is in the mixing chamber and the outlet tube is always above the surface of the liquid entering the tube.
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Overview of Cell Fractionation
it was established that nonelectrolyte-based media are preferable to electrolyte-based media for reducing organelle aggregation (Hogeboom et al., 1948). Most media used today are nonelectrolyte-based, although some do have a substantial electrolyte content (e.g., STKM medium, which contains 50 mM Tris·Cl, 25 mM KCl, and 5 mM MgCl2 in addition to 0.25 M sucrose; Adelman et al., 1973). Moreover, media are generally isoosmotic or hyperosmotic as compared to 0.15 M NaCl. Organelles are osmometers, with water moving across their limiting membranes to maintain osmotic equilibrium; thus lysis can occur in hypoosmotic media. The nonelectrolyte solute most commonly used in subcellular fractionation is sucrose. High-quality sucrose (free of RNase) is reasonably inexpensive, very soluble, and can be used readily to prepare solutions that span the range of densities of most biological organelles (Table 4.1.1). In addition, procedures have been developed for purifying most biological organelles in sucrose. Sucrose, however, has limitations; at higher concentrations solutions are quite viscous and hyperosmotic. Isoosmotic sucrose (0.25 M; ∼9% w/v) has a density of only 1.03 g/ml, which is at the lowerdensity end for biological organelles. Thus all isopycnic density gradient centrifugation performed in sucrose constitutes hyperosmotic conditions; intraorganellar water redistributes to the surrounding medium, and the organelles shrink and increase in density. These changes are reversible for some, but not all, organelles. Various sugar alcohols have been used in place of sucrose in some procedures. Glycerol is one, but although its viscosity is lower than that of sucrose, so is its density at a given concentration, and it also permeates some organelles. Mannitol has been used in a few procedures where the use of sucrose is not feasible—e.g., for isolation of epithelial brush border membranes that contain sucrase (Malathi et al., 1979)—and sorbitol has been a favorite for fractionating yeast, which secrete invertase (a form of sucrase; e.g., Walworth and Novick, 1987). Polysaccharides have also been employed as substitutes for sucrose. Ficoll 400 (Pharmacia), a chemical polymer of sucrose with epichlorohydrin that has an average molecular weight of 400,000, has been a popular choice (70,000 is also available, as Ficoll 70). At low concentrations, it has very low osmolarity, but as can be seen in Figure 4.1.3, the osmolarity rises sharply above 30% (w/v; density ∼1.09 g/ml), and it is quite viscous
at all concentrations above 10% (w/v). Ficoll has been used widely for separating different cell populations and is frequently most useful as an additive to media containing other density-modifying agents (see, for example, the procedures in UNIT 4.2). Iodinated nonelectrolytes increase the density of fractionation media through most of the range of biological organelles while maintaining reasonably low viscosity and osmolarity as compared to sucrose. Two closely related compounds are metrizamide and Nycodenz (both marketed by Nycomed Pharma; Accurate Chemical—listed in the SUPPLIERS APPENDIX—is the U.S. outlet). More recently, Nycomed Pharma has synthesized iodixanol (Optiprep)—essentially dimeric Nycodenz— which has become quite popular (Ford et al., 1994). The structures and specific application of these reagents are discussed in more detail, along with the procedure for their use, in UNIT 4.2. As seen from Figure 4.1.3, 40% (w/v) solutions of both metrizamide and Nycodenz have a density >1.2 g/ml (above that of most organelles) and an osmolarity ≤400 mOsm with reasonable viscosity. Strikingly, 60% (w/v) iodixanol has an osmolarity of only 260 mOsm (Fig. 4.1.3), enabling preparation of iso-osmotic solutions over the full range of organelle densities. Further, iodixanol readily can form self-generating gradients during centrifugation, obviating the need to load samples on preformed gradients (see UNIT 4.2). With all these advantageous features, why aren’t these media used more? They are rather expensive, which certainly limits their use for large-scale work. In addition, they absorb in the UV, in the case of metrizamide, and have been reported to inhibit selected enzyme activities (see UNIT 4.2). A different alternative for achieving high densities at low viscosity and osmolarity involves the use of Percoll, colloidal silica coated with polyvinylpyrrolidone to reduce its adsorption to biological organelles. For fractionation of cellular organelles, Percoll is almost always used in combination with isoosmotic sucrose, because pure Percoll is only 10 mOsm. As can be seen from Figure 4.1.3, Percoll-containing media can achieve very high densities with reasonably low viscosities. Because Percoll particles have a significant size (range 30 to 150 Å) and density, they sediment during centrifugation. Thus, concentration gradients self-form fairly rapidly during a spin and the profile of density along the length of the tube continuously changes with time (see Fig 4.2.2). As pointed out in UNIT 4.2, the
4.1.6 Supplement 37
Current Protocols in Protein Science
Figure 4.1.3 Plots of density, viscosity, and osmolarity of gradient media used to fractionate cellular organelles as a function of concentration. Data are shown for sucrose, Ficoll 400, Percoll, metrizamide, Nycodenz, and Iodixanol (optiprep). Modified from Rickwood (1984) by permission of IRL Press.
shallow gradients achieved with Percoll can be very advantageous in separating organelles having densities that differ only slightly; however, for reproducibility, the conditions of centrifugation must be quite carefully set. Percoll also has its limitations: it absorbs in the UV, interferes with protein assays, precipitates in acid and organic solvents [do not attempt to precipitate Percoll-containing fractions with trichloroacetic acid (TCA) or acetone], and is difficult to get rid of entirely. Some strategies for removing Percoll are discussed in UNIT 4.2.
EVALUATION OF FRACTIONATION In setting up a procedure for separating or purifying cellular organelles, it is advisable to evaluate the steps of the protocol to confirm that the outcome is as desired and as stated. This means setting up assays for marker activities and protein and accounting for the totality of both throughout the procedure. This meticulousness enables the investigator to evaluate
the homogenization, yield, fold purification, and extent of contamination by other undesirable organelles, and to account for 100% of the activity at each step. The extent of cell breakage during homogenization can be estimated by determining the fraction of a marker activity of a particular organelle that pellets in the first low-speed centrifugation step (and is likely still to be associated with unbroken cells), while the extent of organelle breakage can be estimated by determining the fraction of the total amount of a soluble intraorganellar protein that is not sedimented by a highspeed spin that pellets all of the organelle. Following marker activity throughout purification enables the investigator to determine the yield of the desired organelle, and, if markers of other organelles are followed, the extent of residual contamination. Finally, if protein concentration is determined in parallel, calculation of the ratio of activity to protein (known as specific activity) at each step provides an estimate of fold purification or enrichment. In
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Table 4.1.2 Preparing Organelle Fractions from Mammalian Tissues and Cells
Organelle
Procedure
Reference
Nucleus
Centrifugation through high-density sucrose
Blobel and Potter (1966)
Endoplasmic reticulum
Discontinuous sucrose gradient
Adelman et al. (1973)
Golgi complex
Sucrose gradient, continuousa and discontinuous
Bergeron et al. (1982)
Endocrine
Metrizamide gradient
Loh et al. (1984)
Exocrine
Discontinuous sucrose gradient
Cameron and Castle (1984)
Synaptic vesicles
Chromatography on controlled-pore glass
Carlson et al. (1978) Huttner et al. (1983)
Plasma membrane
Discontinuous sucrose gradient
Hubbard et al. (1983)
Endosomes
Free flow electrophoresis Density shift with sucrose gradienta
Marsh et al. (1987) Beaumelle et al. (1990)
Lysosomes
Metrizamide gradienta
Wattiaux et al. (1978)
Mitochondria
Velocity sedimentation in sucrose
Schnaitman and Greenawalt (1968)
Peroxisomes
Sucrose gradient, discontinuous and continuous
Leighton et al. (1968)
Secretion granules
a Procedure also presented in detail in UNIT 4.2.
the event that analytical rather than preparative procedures are used, determination of marker activity in every fraction is essential for specifying the distribution of a particular organelle. Finally, it is very important to note that subcellular fractionation is never perfect! The entirety of a particular organelle is never obtained in completely pure form. In fact, the term “fraction” signifies both incomplete yield and incomplete purity. The careful investigator will always be cognizant of the limitations of the procedures and the consequent limitations of the results.
DEFINITIVE PROCEDURES
Overview of Cell Fractionation
A useful starting point in isolating a particular cellular organelle may be to examine a procedure that has been designed for the specific organelle of interest. Table 4.1.2 lists references to procedures for purifying most of the membranous organelles found in eukaryotic cells (several are also presented as protocols in UNIT 4.2). Unfortunately, it is not possible to make this listing comprehensive, but the procedures selected have achieved unusually good purity and in many cases have documented their achievement by bookkeeping of marker activities. Although several of the procedures
are reasonably old, they have been used either as starting points for subfractionation (e.g., nucleus, endoplasmic reticulum, secretion granules, plasma membranes, lysosomes, mitochondria, and peroxisomes) or for functional studies in cell-free assays (e.g., Golgi, endoplasmic reticulum, and mitochondria).
Literature Cited Adelman, M.R., Blobel, G., and Sabatini, D.D. 1973. An improved cell fractionation procedure for the preparation of rat liver membrane-bound ribosomes. J. Cell Biol. 56:191-205. Beaumelle, B.D., Gibson, A., and Hopkins, C.R. 1990. Isolation and preliminary characterization of the major membrane boundaries of the endocytic pathway in lymphocytes. J. Cell Biol. 111:1811-1823. Bell, A.W., Ward, M.A., Blackstock, W.P., Freeman, H.N., Choudhary, J.S., Lewis, A.P., Chotai, D., Fazel, A., Gushue, J.N., Paiement, J., Palcy, S., Chevet, E., Lafreniere-Roula, M., Solari, R., Thomas, D.Y., Rowley, A., and Bergeron, J.J. 2001. Proteomics characterization of abundant Golgi membrane proteins. J. Biol. Chem. 276:5152-5165. Bergeron, J.J.M., Rachubinski, R.A., Sikstrom, R.A., Posner, B.I., and Paiement, J. 1982. Galactose transfer to endogenous acceptors within Golgi fractions of rat liver. J. Cell Biol. 92:139146.
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Blobel, G. and Potter, V.R. 1966. Nuclei from rat liver: Isolation method that combines purity with high yield. Science 154:1662-1665. Cameron, R.S. and Castle, J.D. 1984. Isolation and compositional analysis of secretion granules and their membrane subfraction from the rat parotid gland. J. Memb. Biol. 79:127-144. Carlson, S.S., Wagner, J.A., and Kelly, R.B. 1978. Purification of synaptic vesicles from elasmobranch electric organ and the use of biophysical criteria to demonstrate their purity. Biochemistry 17:1188-1199. Cronshaw, J.M., Krutchinsky, A.N., Zhang, W., Chait, B.T., and Matunis, M.J. 2002. Proteomic analysis of the mammalian nuclear pore complex. J. Cell Biol. 158:915-927. Ford, T. Graham, J., and Rickwood, D. 1994. Iodixanol: A nonionic iso-osmotic centrifugation medium for the formation of self-generated gradients. Anal. Biochem. 220:360-366. Hofmann, G. (ed.) 1977. ISCO Tables, 7th ed. ISCO, Lincoln, Neb. Hogeboom, G.H., Schneider, W.C., and Palade, G.E. 1948. Cytochemical studies of mammalian tissues. I. Isolation of intact mitochondria from rat liver; some biochemical properties of mitochondria and submicroscopic particular material. J. Biol. Chem. 172:619-635. Hubbard, A.L., Wall, D.A., and Ma, A.K. 1983. Isolation of rat hepatocyte plasma membranes: I. Presence of the three major domains. J. Cell Biol. 96:217-229. Huttner, W.B., Schiebler, W., Greengard, P., and DeCamilli, P. 1983. Synapsin I (Protein I), a nerve terminal–specific phosphoprotein: III. Its association with synaptic vesicles studied in a highly purified synaptic vesicle preparation. J. Cell Biol. 96:1374-1388. Leighton, F., Poole, B., Beaufay, H., Baudhuin, P., Coffey, J.W., Fowler, S., and DeDuve, C. 1968. The large-scale separation of peroxisomes, mitochondria, and lysosomes from the livers of rats injected with Triton WR-1339. J. Cell Biol. 37:482-513.
Malathi, P., Preiser, H., Fairclough, P., Mallett, P., and Crane, R.K. 1979. A rapid method for the isolation of kidney brush border membranes. Biochim. Biophys. Acta 554:259-263. Marsh, M., Schmid, S., Kern, H., Harms, E., Male, P., Mellman, I., and Helenius, A. 1987. Rapid analytical and preparative isolation of functional endosomes by free flow electrophoresis. J. Cell Biol. 104:875-886. Rickwood, D. 1984. Centrifugation, A Practical Approach, 2nd ed. IRL Press, Oxford. Schnaitman, C. and Greenawalt, J.W. 1968. Enzymatic properties of the inner and outer membranes of rat liver mitochondria. J. Cell Biol. 38:158-175. Walworth, N.C. and Novick, P.J. 1987. Purification and characterization of constitutive secretory vesicles from yeast. J. Cell Biol. 195:163174. Wasiak, S., Legendre-Guillemin, V., Puertollano, R., Blondeau, F., Girard, M., de Heuvel, E., Boismenu, D., Bell, A.W., Bonifacino, J.S., and McPherson, P.S. 2002. Enthoprotin: A novel clathrin-associated protein identified through subcellular proteomics. J. Cell Biol. 158:855862. Wattiaux, R., Wattiaux-DeConnick, S., RonveauxDupal, M.-F., and Dubois, F. 1978. Isolation of rat liver lysosomes by isopycnic centrifugation in a metrizamide gradient. J. Cell Biol. 78:349368. Wu, C.C., Taylor, R.S., Lane, D.R., Ladinsky, M.S., and Howell, K.E. 2000. GMx33: A novel family of trans-Golgi proteins identified by proteomics. Traffic 1:963-975.
Contributed by J. David Castle University of Virginia Charlottesville, Virginia
Loh, Y.P., Tam, W.W.H., and Russell, J.T. 1984. Measurement of pH and membrane potential in secretory vesicles isolated from bovine pituitary intermediate lobe. J. Biol. Chem. 259:82388245.
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UNIT 4.2
The protocols in this unit illustrate a range of different procedures that have been used to fractionate tissue homogenates. They emphasize different fractionation techniques that have been used for rat liver, an abundant tissue that has been a favorite of many investigators and has served as the source of many organelle preparations of excellent purity. For selected procedures, other examples have been given using other tissue sources (e.g., glandular tissues that maintain protein storage granules for regulated secretion) or, where particularly favorable, cultured cells. The basis (or goal) of each separation and the merits and limitations of each procedure are summarized in Strategic Planning; this section serves as a guide for selecting among the various approaches (see Background Information for more in-depth discussion). In Strategic Planning the methods are described in somewhat general terms, but each corresponding protocol includes specific details relevant to a particular sample cell type (and in many cases a particular organelle).
STRATEGIC PLANNING Because there are so many different techniques available for purifying organelles, the following annotated outline has been prepared as a guide to selecting a purification strategy.
Differential Centrifugation by Velocity (see Basic Protocol 1) Basis/Goal Enrichment of organelles largely according to size by serial spins of increasingly higher force and longer time. Merits 1. This is the nearly universal starting point in fractionation of a homogenate to remove unbroken cells and large debris (as well as nuclei, if not desired). 2. The procedure is the best way to concentrate organelles of a particular size range from large volumes for use as starting material for subsequent procedures. 3. It is generally performed in isoosmotic sucrose medium, thus providing the best chance of organelles retaining function in subsequent cell-free assays. 4. It is useful for sedimenting macromolecular assemblies, e.g., microtubules polymerized from organelle-free postmicrosomal supernatant. 5. The separation is simple to perform. Within limits, successive resuspension and recentrifugation can be used to further enrich a pellet for a given size range of organelle.
Limitations 1. Pelleted fractions are nearly always substantially contaminated with smaller organelles. 2. Aggregation of organelles in pellets can be significant and difficult to reverse, especially if medium has a substantial electrolyte content.
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Differential Centrifugation by Equilibrium (see Basic Protocol 2) Basis/Goal Separation of organelles according their distinct buoyant densities by centrifuging to equilibrium in a density gradient. Merits 1. The technique provides an excellent follow-up to differential centrifugation by velocity (Basic Protocol 1) for resolving similar-sized organelles into subpopulations of distinct densities. 2. Use of this procedure is often advantageous for complex mixtures resolved as multiple bands. 3. It is also useful for analytical experiments in which the distribution of organelles (or co-distribution of proteins with particular organelles) is being examined, and for preparative experiments to obtain purified organelles for further study.
Limitations 1. Extended centrifugation times may be detrimental to labile activities. This limitation can be reduced by selecting an iodinated nonelectrolyte, e.g., iodixanol (Optiprep), in place of sucrose as centrifugation medium as lower viscosities decrease time required to reach equilibrium (UNIT 4.1, see Alternate Protocol 4). 2. Quantities of sample loaded on each gradient must usually be low to maintain high resolution and avoid aggregation.
Rate Zonal Centrifugation Using Sucrose (see Alternate Protocol 1) Basis/Goal Resolution of organelles from a loading layer (most often at the top of the tube) into enriched zones within a density gradient; separation based on size, shape), and density. Merits 1. The procedure provides multiparameter separation within a single run, which is especially advantageous for analytical experiments that rely on different distributions for several organelles. 2. It is widely used for separations of macromolecules and macromolecular assemblies in addition to membrane-bounded organelles: e.g., proteins of different molecular weights (either water-soluble proteins or detergent-solubilized membrane proteins); polysomes of different length.
Limitations 1. Organelle/protein content of the sample loaded must be kept reasonably low for best resolution; aggregation may be a problem with overly large loads. 2. Its main value is in analysis of organelle distribution; use in organelle purification generally requires further steps.
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3. Hyperosmotic high-density sucrose solutions may cause organelle shrinkage due to water efflux, and can thereby reduce density differences between certain types of organelles.
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Gradient Fractionation Using Percoll (see Alternate Protocol 2) Basis/Goal Rapid separation largely by density in an isoosmotic self-forming density gradient. Merits 1. Percoll’s negligible osmolarity but high density and modest viscosity allow its use as an additive to sucrose for achieving a very wide range of densities under isoosmotic conditions. 2. Because Percoll itself sediments appreciably during centrifugation, self-formed density gradients continually change shape during the run. Initially, they are very shallow along most of the length of the tube, allowing excellent separation of organelles that have only slightly different densities and that approach isopycnic density rather rapidly. Large organelles with slightly differing densities can be easily resolved in fixed-angle rotors, whereas small organelles with slightly differing densities can be resolved in vertical rotors (where the short path length during the run reduces the time needed to achieve equilibrium). 3. Equilibrium density separations require relatively short centrifugations, especially compared to those in high-density sucrose. 4. The procedure can be used effectively for large-scale organelle purification, especially where the organelle of interest has either a lower or a higher density than all other organelles in the starting mixture.
Limitations 1. In general, Percoll is not very suitable for generating distinct distributions for several different organelles in the same centrifuge run. With a gradient profile featuring a shallow central segment between steep extremes, low-density organelles tend to concentrate together at the top of the gradient while high-density organelles tend to concentrate together at the bottom of the gradient. 2. Thoroughly removing Percoll from a purified fraction is a major challenge (see Critical Parameters for further discussion). 3. Further limitations may be imposed by the organelles themselves. Certain organelles may have heterogeneous sizes, shapes, and densities; thus, portions of one type of organelle may end up at both extremes of the gradient.
Gradient Fractionation Using D2 O (see Alternate Protocol 3) Basis/Goal Separation by density gradient with heavy (deuterated) water substituted for normal water. Merits 1. There is reduced osmolarity and viscosity of fractionation medium. 2. It is mainly used in analytical procedures.
Limitations 1. The procedure is quite expensive due to the cost of D2 O, limiting its preparative use. 2. Partial exchange of D2 O for H2 O occurs within membrane-bounded organelles, with the extent of exchange differing among organelles. This may either enhance or reduce their separation and requires empirical analysis.
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Gradient Fractionation Using Iodinated Nonelectrolyte Solutes (see Alternate Protocol 4) Basis/Goal Separation by density gradient with iodinated nonelectrolytes substituted for sucrose to achieve high density. Merits Substituting nonionic solutes for sucrose makes it possible to prepare media that span the range of densities of all membrane-bounded organelles, yet have quite low viscosities and only moderate osmolarities. For organelles with a high buoyant density, this significantly reduces efflux of water, which can lead to irreversible changes in some organelles. Limitations 1. The biggest limitation to these very promising media—metrizamide, Nycodenz and iodixanol (Optiprep)—is their expense, which discourages labs from using them for routine large-scale fractionation. 2. Selected enzymes are inhibited by metrizamide but most are not; see Alternate Protocol 4 (explanatory Note included in the protocol). 3. It is important to keep in mind that these media absorb in the UV range.
Gel Filtration to Isolate Secretory Vesicles (see Basic Protocol 3) Basis/Goal Purification of small, homogeneous vesicles from larger and/or irregularly shaped organelles. Merits 1. The procedure offers excellent resolution for purifying some of the smallest membrane-bounded organelles. 2. It is a promising analytical technique for demonstrating specific association of selected proteins with small vesicles. 3. It is very useful for characterizing or purifying protein complexes, e.g., the cytosolic macromolecular assemblies that function in membrane coat formation during vesicular trafficking.
Limitations 1. Separation is limited to organelles that are small enough to enter the pores of the gel filtration resin efficiently. As indicated by examples cited in the protocol, this means vesicles <200 nm diameter. 2. As the separation is based on size alone, gel filtration with the available resins will not distinguish different types of small vesicles having the same size.
Preparative and Analytical Electrophoretic Separation (see Basic Protocol 4 and Alternate Protocol 5) Basis/Goal Separation of small organelles and macromolecular assemblies on the basis of distinct surface charge densities. Purification of Organelles from Mammalian Cells
Merits 1. Although electrophoretic separations have not been used widely in organelle purification, the preparative technique (Basic Protocol 4) shows substantial promise because
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it is reasonably easy, not expensive, and as reported by its originators (L. Rome, pers. comm.) is capable of handling relatively large samples (i.e., 50 mg protein). Furthermore, preliminary findings suggest that larger-porosity agarose gels might be applicable in the separation of organelles up to 300 nm in diameter. 2. The analytical procedure (Alternate Protocol 5) requires a free-flow electrophoresis chamber, either commercially obtained or constructed around a density gradient as described by Tulp et al. (1993) or Lindner (2001); the latter can be done for only a minor fraction of the cost of the commercially available apparatus. The impressive separations that have been reported (Tulp et al., 1994, Lindner, 2001) make this device worthy of serious consideration. Alternatively, electrophoretic analyses of synaptic vesicles (Carlson et al., 1978) have been performed on Ficoll gradients in plastic tissue culture pipets.
Limitations 1. Use of these procedures has been limited to work on coated vesicles or endosomes/lysosomes; therefore, their applicability would need to be tested before they could be applied to other organelles. 2. High-resolution separation requires brief trypsinization of the starting material, limiting the analysis of resulting fractions to components that are refractory to degradation.
Solid-Phase Immunoadsorption (see Basic Protocol 5) Basis/Goal Isolation of vesicles or macromolecular assemblies using specific antibodies attached to solid-phase supports. Merits 1. The technique is very promising and highly selective. Its success is elegantly demonstrated in experiments involving cultured cells in UNIT 4.3. 2. It can be used both analytically and preparatively. Solid phases include fixed S. aureus, protein A–Sepharose, and secondary or primary antibody coupled to several types of beads (e.g., Sepharose, Eupergit, magnetic) or other polymers. 3. Co-localization of components on immunoadsorbed vesicles can be demonstrated secondarily by immunochemical or immunocytochemical procedures (e.g., see Laurie et al., 1993).
Limitations 1. Immunoadsorption requires a specific antibody that can bind to an exposed epitope on the surface of the organelle (or macromolecular assembly). The success of the technique depends on the quality of the antibody (specificity and avidity) and accessibility of its epitope; adsorption of vesicles derived from intracellular compartments requires cytosolically exposed epitopes, whereas adsorption of plasma membrane vesicles (which generally vesiculate with the ectodomain outward) requires epitopes on the extracellular surface. Under circumstances where exogenous proteins are expressed in the cell or tissue of interest, the use of epitope tagging and relevant antibodies may provide a satisfactory alternative for overcoming this type of limitation (see UNIT 4.3). 2. Although sensitivity is generally a positive quality, in immunoadsorption it can be a limitation. Vesicle binding via an avid antibody may require only a few antigenantibody interactions. As many membrane proteins are not confined to single compartments, but instead traffic among compartments, immunoadsorption may not necessarily isolate only a single type of compartment.
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Purification by Lectin Adsorption (see Basic Protocol 6) Basis/Goal Isolation of vesicles derived from plasma membrane by lectin binding to exposed oligosaccharides. Merits 1. The procedure takes advantage of the unique orientation (ectodomain outward) of plasma membrane vesicles. Exposed glycoproteins on these vesicles can bind to immobilized lectins, particularly wheat germ agglutinin (WGA). 2. The procedure is organelle-specific and quite simple as there is limited need for preenrichment steps. 3. N-acetylglucosamine used to elute vesicles from WGA is inexpensive.
Limitations 1. The procedure is largely specific for a single type of membrane. 2. Separation is only as good as the uniformity of orientation of vesiculated membranes. Intracellular vesicles with inverted orientation can potentially bind to WGA, and inverted plasma membrane vesicles will be unable to do so. Respectively, these occurrences will increase contamination and reduce yield. Generally, however, the orientations are fairly uniform, so this does not pose a major problem. 3. Although the purification of plasma membrane is excellent using the protocol presented here, in other procedures it has been observed to be less extensive; possibly adsorbed vesicles or cytoskeletal proteins increase the contamination.
Density Shift Using Digitonin (see Basic Protocol 7) Basis/Goal Analytical procedure in which binding of digitonin to cholesterol increases the buoyant density in proportion to the cholesterol content of the membrane. Density shifts determined following equilibrium centrifugation range from zero (for ER and mitochondria, which have no resident cholesterol) to intermediate (Golgi cisternae) to maximal (postGolgi membranes, including the plasma membrane). Merits An excellent procedure for distinguishing the possible association of a newly identified (or newly synthesized) protein with a particular compartment, using the extent of density shift of compartmental markers for correlation. Limitations Correlation of distributions is a good supporting technique but does not directly demonstrate a particular association. Density Shift Using Colloidal Gold Conjugates (see Basic Protocol 8) Basis/Goal Capitalizes on the high density of colloidal gold and the well-established characteristics of receptor-mediated endocytosis to provide density modification of selected endocytic compartments, thereby enabling their purification. Purification of Organelles from Mammalian Cells
Merits 1. Although this procedure has not been used widely, it appears quite promising for studies aimed at characterizing the various compartments comprising the endocytic network.
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2. The density shifts observed are substantial, making this a good preparative method, and the procedure can be tailored to specific pathways based on the choice of the ligand that is conjugated to colloidal gold.
Limitations 1. Preparation of the gold conjugates is fairly lengthy, but the procedures involved are not difficult. 2. Procedures are limited to cells or tissue where it is possible to introduce endocytic tracers efficiently. 3. The purity of the fractions obtained is limited by the ability to achieve selective compartmental labeling. No endocytic tracers are perfect compartment markers; thus density shifts are likely to spill over into sequential compartments.
Extraction of Extrinsic Proteins from Membranes Using Sodium Carbonate (see Basic Protocol 9) Basic/Goal Stripping of extrinsic (or adsorbed) proteins from membrane vesicles in order to distinguish candidate integral membrane proteins as those that remain membrane bound. Merits 1. Widely used analytical tool to distinguish membrane proteins anchored in the lipid bilayer either by transmembrane segments or by lipid tethers (e.g., prenylation). 2. Frequently a complementary procedure to phase partitioning in the nonionic detergent Triton X-114 (Bordier, 1981).
Limitations 1. The extreme pH may denature proteins of interest; this possibility should be tested on an individual basis. 2. Membrane vesicles develop discontinuities during treatment (see Fujiki et al., 1982), so subsequent procedures should not require intact vesicles.
DIFFERENTIAL CENTRIFUGATION BY VELOCITY This is the classical standard method of cell fractionation using centrifugation, often used as the starting point for a more complex purification procedure. A cell homogenate is subjected to serial centrifugations of increasingly higher force and longer duration, generally in isoosmotic (0.25 to 0.3 M) sucrose medium (see Background Information for discussion of sucrose concentration). Resulting supernatant fractions or pellets (often prepared as a band atop a cushion of high-density medium rather than a bona fide pellet at the bottom of the tube, in an effort to minimize aggregation) are suitable as starting material for further purification or other procedures. This method gives relatively low resolution in that it does not yield fractions of high purity. However, the postnuclear supernatant (PNS) from step 6 and the supernatant from step 8 serve as useful starting materials for procedures that yield high-purity fractions mainly by exploiting differences in organelle density.
BASIC PROTOCOL 1
As an example of the approach, this protocol presents the fractionation of rat liver into various components, including mitochondria, microsomes, and a largely organelle-free “cytosol” fraction. Additional steps for using tissues other than liver are included. Extraction, Stabilization and Concentration
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Materials Rat starved overnight and freshly sacrificed Homogenization medium: 0.25 to 0.3 M sucrose plus optional additives (see recipe; if PMSF is included, add just before use) Motor-driven pestle with high-torque motor (e.g., laboratory stirrer, VWR or Fisher) Cheesecloth Tubes suitable for low-speed centrifugation: 1.5-ml microcentrifuge tubes (for small-scale preparation) or 15- or 50-ml graduated screw-cap tubes (for larger-scale preparations) Low-speed centrifuge capable of producing a 600 × g centrifugal field, 4◦ C Teflon-glass homogenizer (A.H. Thomas or equivalent) of appropriate size for the scale of the experiment: type AA (1 ml) up to type C (40 ml) 200-µm mesh nylon screen (Tetko) Dounce-type glass homogenizer Polycarbonate or polyallomer tubes suitable for ultracentrifugation (e.g., 3-ml size to fit Beckman TLA-100.3 rotor) or polycarbonate bottle assemblies (e.g., 10-ml size to fit Beckman 50Ti rotor or 25-ml size to fit Beckman 60Ti or 70Ti rotor) Ultracentrifuge and fixed-angle rotor appropriate to the scale of the experiment (e.g., Beckman 50Ti, 60Ti, or 70Ti) or tabletop ultracentrifuge and rotor (e.g., Beckman tabletop with TL100 rotor) NOTE: Prepare solutions with Milli-Q-purified water or equivalent. NOTE: All operations after removing and weighing tissue should be conducted at 0◦ C (ice bucket) or 4◦ C (cold room).
Homogenize the sample 1. Remove liver from a freshly sacrificed rat that has been fasted overnight (to deplete hepatic glycogen). Keeping the liver on ice as much as possible, weigh and then mince coarsely with scissors or razor blade in the weighing boat. Stored frozen tissues are not satisfactory for most organelle purification procedures.
2. Homogenize the minced liver at 20% (w/v) in homogenization medium using five up-down strokes at 1700 rpm with a high-torque motor-driven pestle. 3. Filter the homogenate through four layers of cheesecloth.
Perform low-speed centrifugation to remove nuclei and larger particulates 4. Transfer the homogenate to tubes suitable for low-speed centrifugation and centrifuge 10 min at 600 × g (1700 rpm for a 20-cm-radius rotor), 4◦ C. For small-scale procedures, this centrifugation can be performed in 1.5-ml microcentrifuge tubes for 40 sec at 10,000 rpm (6600 × g).
5. For tissues such as pancreas and salivary gland: Resuspend the pellet in the original volume of homogenization medium and rehomogenize with the Teflon-glass pestle, then centrifuge again under the same conditions. Combine the resulting supernatant with the supernatant from the first centrifugation, filter through 200-µm mesh nylon screen, and disperse by 3 to 4 strokes in a Dounce homogenizer. Purification of Organelles from Mammalian Cells
Pancreas and salivary gland (and some other tissues) are more difficult to homogenize than liver. Homogenization is one of the key steps in successful organelle purification procedures, so it is important to employ the two-step homogenization procedure when appropriate.
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6. The pellet is the crude nuclear fraction and the supernatant (or combined supernatants, for optional two-step homogenization) is the postnuclear supernatant (PNS) fraction. Aspirate any lipid that has accumulated at the surface of the PNS, then withdraw the PNS, taking care to leave the pellet undisturbed. Transfer the PNS to polycarbonate tubes that are suitable for ultracentrifugation.
Prepare mitochondrial and microsomal fractions by successive ultracentrifugations 7. Centrifuge the PNS 30 min at 10,000 × g (e.g., 10,000 rpm in Beckman 50Ti, 60Ti, or 70Ti), 4◦ C, to pellet the mitochondrial fraction. 8. Remove the supernatant and transfer to clean polycarbonate ultracentrifuge tubes. 9. Resuspend the mitochondria-rich pellet and transfer to a Dounce homogenizer. Disperse the pellet into a fine suspension by several manual strokes. The pellet also contains lysosomes and peroxisomes (which are much less abundant) and small amounts of rough and smooth microsomes that have happened to pellet.
10. Optionally, further purify the mitochondria away from smaller-sized contaminants by repeating steps 6 and 7. This strategy has been used by Carvalho-Guerra (1974) for preparing liver mitochondrial fractions.
11. Centrifuge the supernatant from step 7 for 2 hr at 100,000 × g, 4◦ C, in fixed-angle rotor. 12. The resulting pellet is the total microsomal fraction, consisting of vesicles derived from rough endoplasmic reticulum and various smooth-membrane-bounded organelles (including fragments from mitochondria and larger-sized organelles that were damaged during homogenization). Remove the postmicrosomal (“high-speed”) supernatant; this is generally used as “cytosol” in various assays. 13. Resuspend the microsomal pellet in an appropriate buffer for assays or additional purification, disperse by Dounce homogenization, and save at 4◦ C until further use.
DIFFERENTIAL CENTRIFUGATION BY EQUILIBRIUM This procedure can be used alone (as presented here) but most often is used as a followup to differential centrifugation by velocity (Basic Protocol 1). Typically, the samples are applied above (or beneath) a sucrose or other nonelectrolyte-based medium having a density that varies—either discontinuously (in steps) or continuously—from top to bottom of the tube. The sample must of course be even lower in density than the lowest density of the gradient (if applied to the top) or higher than the highest density of the gradient (if applied at the bottom). During centrifugation, organelles sediment (or float) until reaching their isopycnic positions, where the buoyant density of the particle equals the density of the medium (UNIT 4.1). In a continuous gradient, organelles separate as bands distributed throughout the gradient; in a discontinuous gradient the organelles concentrate at the interfaces between the particular steps whose densities are lower and higher than that of each organelle. A continuous gradient is preferred for fractionating complex mixtures containing several different organelles.
BASIC PROTOCOL 2
As an example of this type of fractionation, the following one-step procedure (taken from Bergeron et al., 1982) describes separation of fractions from homogenates of rat liver by flotation in a continuous sucrose gradient. Specific information about collecting Golgi fractions (the goal of Bergeron’s work) is described in annotations. Extraction, Stabilization and Concentration
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Additional Materials (also see Basic Protocol 1) Rat starved overnight and freshly sacrificed Sucrose media: 0.25 M, 1.02 M, and 1.8 M sucrose (ultrapure; e.g., ICN Biochemicals) in dilution solution Dilution solution: 50 mM Tris·Cl (pH 7.4)/25 mM KCl/5 mM MgCl2 Ultraclear or polyallomer ultracentrifuge tubes to fit a swinging-bucket rotor (e.g., 38 ml for Beckman SW28 rotor or equivalent; 13 ml for SW41 rotor or equivalent; 5 ml for SW50.1 or SW55 rotor or equivalent) Gradient-forming device (see Fig. 4.1.2) Ultracentrifuge and swinging-bucket rotor (e.g., Beckman SW28, SW41, SW50.1, or SW55) Syringe (with blunt needle), plastic transfer pipet, or siliconized glass Pasteur pipet NOTE: Prepare solutions with Milli-Q-purified water or equivalent. NOTE: All operations after removing and weighing tissue should be conducted at 0◦ C (ice bucket) or 4◦ C (cold room).
Prepare sample by homogenization and low-speed centrifugation 1. Remove liver from a freshly sacrificed rat that has been fasted overnight (to deplete hepatic glycogen). Weigh liver, mince, homogenize, and filter through cheesecloth as for velocity centrifugation (see Basic Protocol 1, steps 1 to 3), except homogenize at 15% (w/v) in 0.25 M sucrose medium. 2. Add 1 vol of 1.8 M sucrose medium to the homogenate to give 1.02 M sucrose final.
Load and run gradients 3. Load the adjusted homogenate into centrifuge tubes to fill ≤ 12 the tube volume. It is important not to overload individual gradients with sample, as overloading promotes aggregation and decreases resolution and purity.
4. Use a gradient-forming device to form a continuous sucrose gradient on top of the loaded samples. To form the gradient, place 0.25 M sucrose solution in the back reservoir of the gradient-forming device and 1.02 M sucrose solution in the mixing chamber. Open the connecting channel and pump the gradient into the tube (high density first), making sure that the outlet remains above the level of the solution in the filling tube (see Fig. 4.1.2). Alternatively, the continuous gradients can be prepared in advance and kept at 4◦ C for a few hours before loading. In this case, use a syringe with a long needle to carefully load the 1.02 M–adjusted homogenate beneath the gradient in each tube.
5. After the gradients are made, insert the tubes into the buckets of the swinging-bucket rotor. Weigh buckets plus tops and balance in pairs by adding a small amount of the lowest-density sucrose medium to the lighter tube of each pair. 6. Centrifuge tubes 3 hr at 100,000 × g (e.g., 28,000 rpm in Beckman SW28), 4◦ C. Allow the rotor to coast to a stop with the brake off.
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If the deceleration is too rapid, the layers of sucrose (e.g., between the load zone and the gradient) rotate relative to one another, which causes vortexing into the overlying sucrose layer(s) and decreases the resolution. With older ultracentrifuges (Beckman L550, L5-65, and older), the rotor coasts to a stop from maximal speed, taking ∼30 min from 28,000 rpm. As most of the disturbance of gradients occurs below the final 1000 rpm of deceleration, newer instruments decelerate to 800 rpm and then coast to a stop, taking
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significantly less time. Run times are given for older instruments, and can be adjusted downward to accommodate the new deceleration pattern.
Unload the gradients and collect fractions 7. Carefully remove the gradients from the rotor buckets, place in a rack, and note the positions of the bands. For the example described here (starved rat liver), a prominent band was found to have floated ∼1/3 of the way into the gradient (i.e., ∼1/3 of the way up) while a second band was observed at the air/0.25 M sucrose interface.
8. Begin collecting the fractions (bands) separately starting from the top, using a syringe (with blunt needle), plastic transfer pipet, or siliconized glass Pasteur pipet. Be sure to collect thoroughly: first sweep the circumference of the tube with the syringe or pipet tip, then pass the tip systematically across the interface while applying suction. Because the direction of centrifugation is radial, bands are more highly concentrated at the walls of the tube than in the center (see UNIT 4.1 for discussion of rotor geometry).
9. If recovery of total activity of the loaded sample is a goal of the experiment, collect aliquots of the solution between bands, as well as aliquots of the original sample and the pellet (after resuspension), as separate samples for assay.
Process individual fractions 10. Dilute each band at least 2- to 3-fold with buffer or a low-concentration sucrose medium. Pellet by velocity sedimentation for ≥100,000 × g (e.g., 40,000 rpm in Beckman SW41 or SW50.1), 4◦ C, then resuspend in the same medium for further use. Process Golgi fraction by adding dilution solution to 0.25 M sucrose final (use a refractometer, if available, to measure the sucrose concentration), then pelleting 1 1 /2 hr at 100,000 × g, 4◦ C. Generally bands obtained by isopycnic centrifugation are not as concentrated as resuspended pellets obtained by differential velocity sedimentation (see Basic Protocol 1). Furthermore, individual bands are suspended in media with differing densities.
11. If desired, subfractionate the samples further—for example, by lysis to separate membranes from soluble content or by high pH (carbonate) treatment, which is frequently used to strip membranes of extrinsic proteins (see Basic Protocol 9). As an alternative to the continuous gradient procedure, it is also possible to generate Golgi fractions by flotation on a discontinuous sucrose gradient. In this case, a homogenate prepared in 0.25 M sucrose/5 mM Tris·Cl (pH 7.4)/25 mM KCl/5mM MgCl2 /4.5 mM CaCl2 (STKCM) is adjusted to 1.15 M sucrose in the same medium. Above this sample in the centrifuge tube are placed layers of 0.95 and 0.4 M sucrose in STKCM. After centrifugation at 200,000 × g for 90 min, the Golgi fraction is collected at the 0.95/0.4 M interface. By using a discontinuous gradient in place of the continuous gradient, it is possible to collect a Golgi fraction with high organelle concentration and thereby avoid the pelleting (step 10 above) after purification. This variation, reported by Dominguez et al. (1999), was used to provide starting material for in vitro studies of membrane fusion. It also served as starting material for organelle proteomic analysis (Bell et al., 2001). Because the liver produces very low density lipoproteins that transiently accumulate in the Golgi before export to the cell surface, Golgi from this source has an unusually low buoyant density and is especially favorable for preparing reasonably purified fractions. Golgi fractions from other sources are more dense, and so far most preparations from nonhepatic sources are not as pure. Extraction, Stabilization and Concentration
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ALTERNATE PROTOCOL 1
RATE ZONAL CENTRIFUGATION USING SUCROSE Rate zonal centrifugation is a nonequilibrium density gradient procedure for fractionating cells, in which sedimentation depends on the size, density, and shape of the particles as well as the viscosity and density of the medium. Typically, samples are loaded atop a continuous sucrose gradient; during subsequent reasonably short centrifugations (length of run and rate often need to be determined empirically), they separate as distinct bands (zones) enriched in particular organelles. This procedure can be adapted to various-sized gradients; available models of gradient-forming devices can be used to generate gradients in tubes ranging from 5 to 38 ml. To prepare gradients in smaller tubes (e.g., for a tabletop ultracentrifuge), investigators often approximate continuous gradients by preparing step gradients having several layers of progressively decreasing density, then letting them stand a few hours before use. Diffusion between the layers makes the approximation quite good. The example described in this protocol is the separation of a postnuclear supernatant (PNS) from exocrine pancreas. It can be adapted to other tissues or cell cultures but this may require adjusting the density limits of the gradient. See, for example, the preparation of Golgi membranes for cell-free assays (Balch et al., 1984). Figure 4.2.1 presents the results of assaying marker activities of several organelles across the gradient. Ficoll 400 (a hydrophilic polymer; see UNIT 4.1) has been added to the sucrose to retard the migration of mitochondria relative to zymogen granules so that these two abundant organelles are resolved as distinct bands.
Materials Rat starved overnight and freshly sacrificed Homogenization medium: 0.3 M sucrose (ultrapure; e.g., ICN Biochemicals) plus 0.2 µg/ml diphenyl-p-phenylenediamine (DPPD, Kodak; from 0.4 mg/ml stock in ethanol) 1 M MES 0.2 M EDTA (APPENDIX 2E) Gradient stock solutions, low- and high-density (see recipe) 0.3 M sucrose/5 mM MES/1 mM EDTA/0.2 µg/ml DPPD, pH 6.5 to 6.7 Cheesecloth Dounce homogenizer Gradient-forming device (see Fig. 4.1.2) suitable for selected gradient size, including compatible magnetic stir bars, tubing, and peristaltic pump Ultraclear or polyallomer centrifuge tubes (of the selected size) suitable for swinging-bucket rotor Plastic tissue culture pipet or polyethylene transfer pipet Ultracentrifuge and appropriate swinging-bucket rotor (e.g., Beckman SW50.1 or SW55 if using 5-ml tubes; Beckman SW41 for 12.5-ml tubes, Beckman SW28 for 38-ml tubes) Automated gradient-collection device (e.g., Buchler Autodensiflow, Nycomed Pharma or MSE Scientific) or micropipettor for manual collection NOTE: Prepare solutions with Milli-Q-purified water or equivalent. NOTE: All operations after removing and weighing tissue should be conducted at 0◦ C (ice bucket) or 4◦ C (cold room). Purification of Organelles from Mammalian Cells
Prepare the pancreatic samples 1. Remove pancreas from freshly sacrificed rat that has been fasted overnight (to maximize the zymogen granule population). Weigh pancreas (usually 0.8 to 1 g per
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Figure 4.2.1 Distribution of marker activities for various organelles after rate zonal centrifugation of a postnuclear supernatant (PNS) from rat pancreas using the conditions specified in Basic Protocol 1. The density profile of the gradient after the run was determined by refractive index and plotted as sucrose (sucrose plus Ficoll) concentration; this is shown in the upper portion of the figure. The density profile of the gradient after the run was determined by refractive index and is shown in the upper portion of the figure. The activities that were assayed and the organelles that they mark are: amylase, a secretory protein concentrated in zymogen granules at the bottom of the gradient but also found in lower amounts in rough endoplasmic reticulum, Golgi, and at the top of the gradient (as a result of organelle lysis during homogenization); GALTRANS, galactosyltransferase, a marker for Golgi; γ-GT, gamma-glutamyl transferase, an integral component of plasma membrane but also of zymogen granule membranes (at lower concentration); MAO, monoamine oxidase, a marker of mitochondria (which is concentrated in outer mitochondrial membranes; also note the small amount near the top of the gradient resulting from organelle breakage); NADH–Cyt.c, NADH–cytochrome c reductase, a marker activity of rough ER and mitochondria. The distinct but overlapping distributions of these markers in this analytical centrifugation is apparent.
animal), mince, homogenize, and filter through cheesecloth (as for liver; see Basic Protocol 1, steps 1 to 3), except use homogenization medium containing 0.2 µg/ml DPPD and homogenize at 15% (w/v). DPPD, an antioxidant, is added because it significantly reduces lysis of zymogen granules.
2. Centrifuge sample, rehomogenize (in the same volume as initially used of homogenization medium containing DPPD), and recentrifuge, then pool supernatants (see Basic Protocol 1, steps 4 and 5). Add MES to 5 mM (from 1 M stock) and EDTA to 1 mM (from 0.2 M stock). Filter and disperse with a Dounce homogenizer (see Basic Protocol 1, step 5). Maintain homogenized sample (PNS) at 0◦ C, preferably briefly, until loaded on the gradients.
Prepare the continuous gradients 3. Set up the gradient-forming device and peristaltic pump as shown in Fig. 4.1.2. 4. Load the gradient-forming solutions. Into the back chamber (reservoir) of the gradient former, pipet the high-density sucrose solution (15 ml for 38-ml tubes; 5 ml for
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12.5-ml tubes; 2.2 ml for 5-ml tubes). Carefully bleed a little of the solution through to the front (mixing) chamber to displace the air in the joining tube, then pipet the low-density solution into the mixing chamber (same volumes as for high-density solution). Be sure that in this configuration that the delivery tube is at the bottom of the centrifuge tube, as the gradient will be generated low-density-first (see Fig. 4.1.2). 5. Start the stirring motor and the peristaltic pump, and immediately open the joining tube between the two chambers. Note the Schlieren patterns in the mixing chamber as high-density sucrose flows in, first as is, then mixed. Set the delivery rate of the peristaltic pump empirically; if it is too fast, the solution level in the mixing chamber will drop below that in the delivery chamber and mixing will appear incomplete. 6. Once delivery of the solutions from the gradient-forming device to the centrifuge tubes is complete, shut off the peristaltic pump before air in the emptying delivery tube reaches the gradient. Slowly withdraw the delivery tube up through the gradient so disturbance to the gradient is minimal. Continuous gradients are quite stable and can be prepared the day before use provided they are stored at 4◦ C without disturbance.
Load the sample on the gradients and centrifuge 7. Using a plastic tissue culture pipet or polyethylene transfer pipet, carefully overlay the gradients with samples of PNS. Maximal volumes of the 7.5% (w/v) PNS that can be loaded without overloading and decreasing resolution are 5.5 ml on a 30-ml gradient, 1.8 ml on a 10-ml gradient, and 0.5 ml on a 4.5-ml gradient. To the inexperienced investigator, use of pipets may sound tricky. In the authors’ experience, nice layering with little mixing can be achieved rather quickly by placing the tip of the pipet on the meniscus of the solution while slightly tilting the tube containing the partly layered gradient, then gradually relaxing the pressure on the finger controlling the flow from the pipet. Some practice may be necessary! As an alternative, polyethylene transfer pipets can be used to load the solutions.
8. Place the loaded gradients into the buckets of the appropriate swinging-bucket rotor, taking care that the tubes are not wet on the outside. Balance opposite buckets to within 0.1 g of each other using homogenization medium. Generally, with accurate pipetting during forming and loading, only minor adjustment is required for balancing.
9. Centrifuge the samples in the ultracentrifuge set at 4◦ C. For rate zonal centrifugation, which is a nonequilibrium procedure, the angular velocity and length of the run may require empirical adjustment. For pancreatic PNS centrifuged in an SW28 rotor, excellent separation conditions are obtained by spinning 110 min at 26,000 rpm (90,000 × g), 4◦ C, then letting the rotor coast to a stop without the brake.
10. At the end of the run, the gradients should show discrete bands or zones that are enriched in specific organelles; these zones are distributed throughout the gradient.
Purification of Organelles from Mammalian Cells
Collect the gradients at 4◦ C 11. Unload the gradients in successive fractions of equal volume using an automated collection device or a micropipettor. For large-volume gradients in 38-ml tubes, a Buchler Autodensiflow device, which collects from the top, works very well. For smallvolume gradients, collection can be satisfactorily performed using a micropipettor, taking care to withdraw the solution slowly and from as close to the surface as possible.
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12. Assay the gradient fractions for the activity of interest. For some assays it may be necessary to reduce the sucrose content and/or concentrate the sample. If so, dilute fractions 2- to 3-fold with 0.3 M sucrose/5 mM MES/1 mM EDTA/0.2 µg/ml DPPD and pellet 1 hr (or shorter if the volume is small) at 100,000 × g, 4◦ C. Figure 4.2.1 shows the results obtained by assaying the gradient fractions obtained by rate zonal centrifugation of the pancreatic PNS for enzymes that mark particular organelles.
GRADIENT FRACTIONATION USING PERCOLL Percoll (polyvinylpyrrolidone-coated colloidal silica) is a useful alternative to highconcentration sucrose gradients, particularly where high densities combined with low viscosity and osmotic activity are desired. Cells are homogenized, mixed with a Percoll solution, and centrifuged, generally using a fixed-angle rotor. The Percoll forms its own density gradient during centrifugation (a self-forming gradient). As it has very low osmotic activity (pure Percoll as purchased is ∼10 mOsm), it is generally added to isoosmotic (or slightly hyperosmotic) sucrose for separating organelles from cell homogenates under isoosmotic (or near isoosmotic) conditions. The density range is controlled by the Percoll concentration, and the slope of the density gradient is determined by the time of centrifugation and the centrifugal field (see Fig. 4.2.2). Generally, the duration of separation procedures is selected so that organelles sediment to near equilibrium density
ALTERNATE PROTOCOL 2
Figure 4.2.2 Percoll density gradient profiles as a function of time of centrifugation at 20,000 × g in a fixed-angle rotor. The starting density in the example shown is 1.07 g/ml. Note that with self-forming density gradients, the slope of the gradient begins as zero; during the run, the shape progressively changes as the Percoll sediments. Early in the run most of the tube contains a shallow gradient that is bordered by Percoll-depleted solution at the top and high-density pelleted Percoll at the bottom. As the run proceeds, the central resolving segment of the gradient becomes steeper and shorter. To produce an isoosmotic sucrose/Percoll solution of a desired density, use the formula: Vx ρ − ρx = o Vo ρi − ρ x where Vx = volume of 2.5 M sucrose, Vo = volume of Percoll, ρo = density of Percoll (1.13 g/ml), ρx = density of 2.5 M sucrose (1.315 g/ml), and ρi = desired density. Figure courtesy of Pharmacia Biotech.
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(isopycnic conditions). Short runs (∼30 min) enable large particles such as 1-µm-diameter secretion granules and mitochondria to approach equilibrium, whereas longer runs (up to 2 hr) may be required for smaller particles. In the example presented in this protocol, Percoll gradients are used to purify secretion granules from exocrine glandular tissue and then to separate the granules into subpopulations according to the extent of their maturity (see Zastrow and Castle, 1987). The initial purification procedure is widely applicable to other types of granules.
Additional Materials (also see Alternate Protocol 1) Percoll gradient solutions (see recipe) Centrifuge and fixed-angle rotor (e.g., Beckman 70Ti or Type 50) Centrifuge tubes (e.g., Beckman thick-walled polycarbonate bottle assemblies; 25-ml for 70Ti rotor or 10-ml for Type 50 rotor) Refractometer NOTE: Prepare solutions with Milli-Q-purified water or equivalent. NOTE: All operations after removing and weighing tissue should be conducted at 0◦ C (ice bucket) or 4◦ C (cold room).
Load and spin the samples 1. Remove parotid salivary gland from freshly sacrificed rats (0.4 to 0.5 g tissue per animal). Weigh gland, mince, homogenize, and filter through cheesecloth (see Basic Protocol 1, steps 1 to 3), except using homogenization medium containing DPPD (as in Alternate Protocol 1) and homogenizing at 12% (w/v). DPPD, an antioxidant, is added because it significantly reduces lysis of zymogen granules.
2. Centrifuge sample, rehomogenize (in homogenization medium with DPPD), and recentrifuge, then pool supernatants (see Basic Protocol 1, steps 4 and 5). Add MES to 5 mM and EDTA to 1 mM. Filter and disperse with a Dounce homogenizer (see Basic Protocol 1, step 5). 3. Add the PNS to 2 vol of 60% Percoll/sucrose and mix. Measure the refractive index using a refractometer. 4. Load into tubes that are appropriate for the fixed-angle rotor that has been selected. Centrifuge the samples under appropriate conditions to separate the organelles of interest. To separate parotid gland secretion granules, appropriate centrifugation conditions are 30 min at 16,400 × g (e.g., 15,000 rpm in 70Ti), 4◦ C. Under these conditions the granules separate as a white band at the bottom of the tube, while the other organelles form a dense band near the top of the tube. A mostly clear zone between the bands corresponds to the shallow portion of the density gradient.
Collect bands and prepare for further purification 5. Collect the bands manually using a Pasteur or plastic transfer pipet. 6. For each fraction, mix by swirling or inverting, then remove a small aliquot and measure its refractive index with a refractometer. Purification of Organelles from Mammalian Cells
7. Adjust the refractive index back to the same value measured for the original diluted PNS by adding homogenization medium.
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Purify fractions further Example: For parotid-gland secretion granule fraction: 8. Load the adjusted granule fraction into centrifuge tubes and recentrifuge under the same conditions as used in the first run. This repeat centrifugation will float much of the residual contaminants away from the granule fraction.
9. Collect the granule fraction manually with a pipet. 10. If desired, process the purified granule fraction as a single organelle population by diluting with 3 vol homogenization medium, then pelleting the granules by centrifuging 45 min at 2000 × g (e.g., 5000 rpm in 70Ti), 4◦ C. Alternatively, proceed to step 11 for further separation of the granule fraction by Percoll centrifugation. The low-speed centrifugation removes most, but not all, of the Percoll and provides an easily resuspended pellet. Removal of Percoll is necessary because it can interfere with later processing of the sample. If necessary, further Percoll removal and purification can be achieved by dilution and recentrifugation under the same conditions, by ultracentrifugation on a step gradient as specified by Zastrow and Castle (1987), or by other methods (see Critical Parameters for further discussion).
11. To separate granule subpopulations: Measure the refractive index of the granule fraction collected in step 9 and adjust to a value of n = 1.3586 using either 0.3 M sucrose or 86% Percoll/sucrose, depending on whether refractive index must be adjusted downward or upward, respectively. The value of n will differ and must be determined empirically for granules from other sources.
12. Place the adjusted granule fraction in a new set of centrifuge tubes and centrifuge 30 min at 45,500 × g (e.g., 25,000 rpm in 70Ti or Type 50 rotor, depending on the volume), 4◦ C. 13. The granule fraction will now be subdivided into a minor upper band, a major lower band, and the clearer (but slightly turbid) zone between the bands. Collect the upper band, the intervening zone, and the lower band as separate fractions. Together these fractions comprise a continuum of increasingly dense granules (and increasing granule maturation), rather than discrete subpopulations. Evidently, the relative amounts of granules in each subfraction depend on the initial density (and thus refractive index) before the spin and on the centrifugation conditions (because Percoll gradients are self-forming and constantly change during the centrifugation).
14. Process the collected bands as in step 10 to remove Percoll. Proceed with analysis.
GRADIENT FRACTIONATION USING D2 O Deuterium oxide fractionation takes advantage of the fact that the density of D2 O is 1.1 g/ml—substantially closer than that of water to the density of biological organelles. The procedure is a form of equilibrium centrifugation in which D2 O (heavy water) is substituted for H2 O. This technique has not been used widely in subcellular fractionation, largely because deuterium oxide is quite expensive, and also because the practical value of the approach has fallen short of expectations. In some cases, however, it can be quite effective, so it should be considered as an option. The procedure presented below (taken from Gumbiner and Kelly, 1981) illustrates use of a combined Ficoll/D2 O density gradient to purify and analyze endocrine secretion granules derived from a pituitary cell line.
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Additional Materials (also see Alternate Protocol 1) Mouse pituitary AtT-20/D16V cells PBS (APPENDIX 2E) containing 4 mM EGTA (from 100 mM EGTA, pH 7.4), 37◦ C Homogenization medium: 0.25 M sucrose (ultrapure; e.g., ICN Biochemicals)/10 mM HEPES (pH 7.4)/2 mM EGTA/1 mM EDTA, 4◦ C Deuterium gradient stock solutions: 0.25 M ultrapure sucrose/10 mM HEPES (pH 7.4)/1 mM EDTA in D2 O (e.g., Aldrich), with and without 20% (w/v) Ficoll 8% (w/v) Ficoll 70 (Pharmacia Biotech) in homogenization medium 75-cm2 tissue culture flasks Dounce glass homogenizer with type B pestle (Kontes or equivalent) Sorvall centrifuge and SS-34 rotor (or equivalent) Gradient-forming device with capacity ≥15 ml/chamber (Hoefer Pharmacia or equivalent) Peristaltic pump to form continuous gradient Ultracentrifuge with swinging-bucket rotor (e.g., Beckman SW27 or SW28) Automated gradient collection apparatus: Autodensiflow device (Buchler) or gradient displacement device (Nycomed Pharma or MSE Scientific) NOTE: Prepare solutions with Milli-Q-purified water or equivalent. NOTE: All operations after harvesting cells should be conducted at 0◦ C (ice bucket) or 4◦ C (cold room).
Homogenize the cells and prepare the crude granule pellet 1. Grow cells almost to confluency in five 75-cm2 tissue culture flasks (∼5 × 106 /flask). Harvest cells at 37◦ C in PBS/4 mM EGTA and pellet at 4◦ C. 2. Resuspend in 15 ml of 4◦ C homogenization medium and homogenize with six strokes in a Dounce homogenizer. A ball-bearing homogenizer designed by Balch and Rothman (1985) is now used widely for homogenizing cultured cells.
3. Pellet large debris and organelles by centrifuging 5 min at 10,000 × g (e.g., 9500 rpm in SS-34 rotor in a Sorvall centrifuge), 4◦ C. 4. Transfer the supernatant to a fresh tube and centrifuge 35 min at 30,000 × g (16,000 rpm), 4◦ C, to pellet the crude granule fraction. 5. Resuspend the crude granule pellet in 1 to 2 ml homogenization medium using a micropipettor until fine particulates are no longer seen. If necessary, homogenize using a small-capacity Dounce homogenizer to achieve the desired uniform suspension. Maintain on ice while preparing the gradient.
Prepare the Ficoll-D2 O gradient 6. Using the gradient stock solutions made in D2 O (with and without 20% Ficoll), prepare layering solutions that are 17%, 14%, 11%, 9%, and 8% Ficoll. 7. Prepare a Ficoll step gradient (in 100% D2 O) at the bottom of each tube by layering the following solutions (from step 6) using either a micropipettor or plastic tissue culture pipet: Purification of Organelles from Mammalian Cells
1 ml 20% Ficoll 2 ml 17% Ficoll 2 ml 14% Ficoll
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2 ml 11% Ficoll 1 ml 9% Ficoll. The volumes specified are for a 38-ml centrifuge tube, which is used in a Beckman SW28 rotor.
8. Mix 8 ml of 20% Ficoll gradient stock solution with 12 ml homogenization medium to give 20 ml of 8% Ficoll in 40% D2 O medium. 9. Place 14.5 ml of the 8% Ficoll/40% D2 O solution in the back reservoir of a gradientforming device. After filling the delivery tube between the chambers, place 14.5 ml of the 8% Ficoll in 100% D2 O gradient stock solution (from step 6) into the mixing chamber. Open the delivery tube, starting stirring the mixing chamber, and start the peristaltic pump to deliver a continuous gradient 40% to 100% D2 O in 8% Ficoll on top of the Ficoll step gradient.
Load the sample and spin 10. Load the resuspended crude pellet on the gradient and centrifuge 12 to 15 hr at 82,500 × g (e.g., 26,000 rpm in SW28 rotor), 4◦ C. The centrifugation conditions listed (including tube size and rotor) are those reported by Gumbiner and Kelly (1981). It is very likely that this procedure can be successfully scaled down for use with smaller samples and smaller-volume gradients.
Collect fractions from the gradient and analyze 11. Collect 1.5-ml gradient fractions using one of the following methods: a. Collect top-first manually using a micropipettor. b. Collect top-first by an automated method: use either a Buchler Autodensiflow device or gradient displacement device (e.g., Nycomed Pharma or MSE Scientific). c. Collect bottom-first by puncturing a hole in the bottom of the tube and collecting drops. The third method was used in the published procedure (Gumbiner and Kelly, 1981). Analyses for the secretion granule marker ACTH and total protein indicate that granules are highly purified, 50-fold on average, and are located within the lower Ficoll gradient/100% D2 O portion of the tube (centered at a density of 1.17 g/ml). Most other organelles are centered within the 8% Ficoll/D2 O gradient portion of the tube as judged by the concentration of protein in this region. Soluble protein remains at the top of the gradient. Lysosomes are a problematic contaminant of endocrine secretion granules (e.g., Roep et al., 1990; Loh et al., 1984) and have not been assayed specifically in this procedure.
GRADIENT FRACTIONATION USING NONIONIC IODINATED SOLUTES Another strategy that is being used to achieve higher-density fractionation media with reduced osmolarity and viscosity involves supplementing or replacing sucrose with iodinated solutes. By far the most successful of these solutes are the nonionic derivatives of triiodobenzoic acid: metrizamide; Nycodenz; and iodixanol, or Optiprep (all available from Nycomed Pharma). Metrizamide [(2-3-acetamido)-5-N-methylacetamido-2,4,6triiodo(benzamido)-2-deoxy-D-glu-cose)], Nycodenz (N,N bis(2,3-dihydroxypropyl)-5N-(2,3-dihydroxypropyl)acetamido-2,4,6-triiodoisophthalamide), and iodixanol (5,5 [(2-hydroxy-1-3-propanediyl)-bis(acetylamino)] bis[N,N bis(2,3-dihydroxypropyl)2,4,6-triiodo-1,3-benzenecarboxamide] are soluble in all aqueous media, stable over the pH range 2 to 12.5 and, unlike related ionic solutes, are not affected by low pH or the presence of divalent cations. Iodixanol is marketed as, Optiprep, a 60% (w/v) solution
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with density 1.320 g/ml, osmolality 260 mOsm, and refractive index 1.4287 without buffer or other additives. These media are advantageous for fractionation under sterile conditions (Nycodenz can be autoclaved, metrizamide must be sterile filtered; Optiprep is sold sterile). The differences in fractionation between metrizamide and Nycodenz are negligible, and for most purposes they are interchangeable. Note, however, that reported buoyant densities of selected organelles can differ slightly between the two (e.g., mitochondria have a density of 1.115 g/ml in metrizamide and 1.132 in Nycodenz). For a more extensive discussion of the properties of these solutes, see the monograph on centrifugation edited by Rickwood (1984). The decreased osmolarity of Optiprep (see Fig. 4.1.3) relative to metrizamide and Nycodenz enables fully iso-osmotic fractionation. This procedure describes purification of lysosomes from rat liver using a metrizamide gradient (Wattiaux et al., 1978); studies isolating other organelles by this method are noted in Background Information. In addition to preparing a high-quality fraction, these investigators show how results obtained with a continuous gradient are used to devise a step-gradient procedure for routine preparations. Subsequently, a procedure for resolving several organelles from mouse liver using iso-osmotic gradients of iodixanol is described (Graham et al, 1994).
Additional Materials (also see Alternate Protocol 1) Rat starved overnight and freshly sacrificed Homogenization medium: 0.25 M sucrose (ultrapure; e.g., ICN Biochemicals), 4◦ C 85.6% metrizamide (from 100% stock, Nycomed Pharma) Gradient stock solutions: for continuous gradients, 19.7% and 50% (w/v) metrizamide, pH 7.4; for step gradients, 19.78%, 24.53%, 26.34%, and 32.82% (w/v) metrizamide, pH 7.4 (adjust pH with 0.01 N NaOH) Teflon-glass homogenizer (type C, 40 ml, A.H. Thomas) Fixed-angle ultracentrifuge rotor appropriate to the size of the experiment (e.g., Beckman 70Ti) Automated gradient collection apparatus: Autodensiflow device (Buchler) or gradient displacement device (Nycomed Pharma or MSE Scientific) NOTE: Prepare solutions with Milli-Q-purified water or equivalent. NOTE: All operations after removing and weighing tissue should be conducted at 0◦ C (ice bucket) or 4◦ C (cold room).
Prepare sample by homogenization and low-speed centrifugation 1. Remove liver from a freshly sacrificed rat that has been fasted overnight (to deplete glycogen stores). Weigh the liver, mince, and homogenize as for velocity centrifugation (see Basic Protocol 1, steps 1 to 2), except homogenize at ∼20% (w/v) using a Teflon-glass homogenizer at 3000 rpm in 4◦ C homogenization medium (0.25 M sucrose). 2. Adjust the homogenate to 15% (w/v), then filter through cheesecloth. Centrifuge 15 min at 2,000 × g, 4◦ C. 3. Save the supernatant on ice and resuspend the pellet, adding homogenization medium to the original homogenate volume. Rehomogenize, then centrifuge again as in step 2. Purification of Organelles from Mammalian Cells
4. Pool the two supernatants and centrifuge 30 min at 8,800 × g in a fixed-angle rotor (e.g., 10,000 rpm in 70Ti). Remove the supernatant, including the fluffy layer
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atop the pellet; resuspend the pellet in homogenization medium and repeat the centrifugation. 5. Resuspend the pellet in homogenization medium and mix with 2 vol of 85.6% metrizamide (to give a final 56.7% metrizamide load for the gradients). Metrizamide, Nycodenz and iodixanol all absorb strongly in the UV range, complicating protein determinations in gradient fractions by OD280 . Also, they interfere with some protein assays (e.g., Lowry). To assay protein in various fractions, the samples can be precipitated with trichloroacetic acid and redissolved in 0.5 N NaOH. Metrizamide inhibits galactosyltransferase activity, so if this Golgi assay is to be performed, the fractions must be diluted, pelleted, and resuspended in a sucrose (or other noninhibitory) medium.
Prepare continuous metrizamide gradient and load sample 6. Using the gradient-forming device (Fig. 4.1.2), place the 50% metrizamide solution (density 1.280 g/ml) in the back reservoir and fill the delivery tube to the mixing chamber. Next place the 19.8% metrizamide solution (density 1.105 g/ml) in the mixing chamber. Set the outlet tube at the bottom of the centrifuge tube. 7. Open the delivery tube between the two chambers, start stirring in the mixing chamber, and pump the gradient (in low-density-first configuration). Wattiaux et al. (1978) treats the continuous gradient as an analytical run and uses 5-ml gradients.
8. After the gradient is delivered, shut off the pump, taking care not to let air pass into the gradient from the outlet tube. Slowly withdraw the outlet tube from the centrifuge, taking care not to disturb the gradient. 9. Layer the sample in 56.7% metrizamide (final) beneath the gradient using either a Pasteur pipet or a syringe with long needle. The reason for loading the gradient at the bottom is that mitochondria exhibit a higher equilibrium density when initially equilibrated in high-density metrizamide than when equilibrated in sucrose homogenization medium. This strategy is critical to achieving separation of mitochondria and lysosomes in the gradient.
10. Centrifuge the gradients 2
1 2
hr at 108,000 × g (e.g., 28,000 rpm in SW28), 4◦ C.
Unload the gradients, measure refractive index, and assay marker activities 11. Working at 4◦ C, collect gradient fractions of equal volume using one of the following methods: a. Collect top-first manually using a micropipettor. b. Collect top-first by an automated method: use either a Buchler Autodensiflow device or gradient displacement device. c. Collect bottom-first by puncturing a hole in the bottom of the tube and collecting drops. 12. Measure the refractive index of each fraction. Using either a published table (available from Nycomed Pharma) or standard curve constructed from a set of precisely made metrizamide stocks, construct a density profile for the gradient. 13. Using each gradient fraction plus an aliquot of the original gradient load, assay marker activities for lysosomes (e.g., acid phosphatase, β-galactosidase), mitochondria (e.g., cytochrome oxidase), peroxisomes (e.g., catalase), and other organelles as desired. Extraction, Stabilization and Concentration
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14. From the activities present in each fraction, calculate the total recoveries of each of the enzymes present in the original gradient load. Plot the frequency of the components for each fraction as activity/(sum of the product of total enzyme activity × the density increment for each fraction). Using these plots, it was found that peroxisomes equilibrate near the bottom of the gradient (median density 1.225 g/ml) with a portion of the catalase remaining in the load; mitochondria equilibrate near the middle (median density 1.162 g/ml); and lysosomes equilibrate in the upper region of the gradient (median density 1.133 g/ml).
Use step gradient to isolate specific organelles Example: Isolate preparative levels of lysosomes 15. Based on the findings made using the continuous analytical metrizamide gradient (step 14), design an appropriate bottom-loaded step gradient. Resuspend the starting sample (light mitochondrial pellet) in homogenization medium, using 1 ml per 3 g original weight of liver, and mix with 2 vol of 85.6% (w/v) metrizamide (to give 56.7% load). Pipet 10 ml into a Beckman SW27 or SW28 tube, then overlay successively with 6 ml of 32.82% metrizamide (density 1.181 g/ml), 6 ml of 26.34% metrizamide (density 1.145 g/ml), 6 ml of 24.53% metrizamide (density 1.135 g/ml), and 9 ml of 19.78% metrizamide (density 1.109). 16. Centrifuge step gradient 2 hr at 95,000 × g (e.g., 27,000 rpm in SW27 or SW28), 4◦ C. 17. Collect bulk fractions from the top of the gradient using a pipet. Fraction 1 extends from the top to about two-thirds of the way through the 19.78% metrizamide layer; fraction 2 includes the 19.78%/24.53% interface (expected from step 14 to be enriched for lysosomes); fraction 3 includes the 24.53%/26.34% interface (expected from step 14 to be enriched for mitochondria); and fraction 4 includes the 32.82%/56.7% (load) interface (expected from step 14 to be enriched for peroxisomes). 18. Assay for marker activities and by electron microscopy to confirm that the fractions are enriched as expected. Notably, Wattiaux et al. (1978) report that fraction 2 contained 10% to 12% of the lysosomal hydrolase activity of the total homogenate (a respectable yield) and had a specific activity for the lysosomal markers that on average was 74-fold higher than that of the homogenate (a very impressive purification).
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A very attractive set of alternative procedures involving use of iso-osmotic iodixanol gradients for preparative purification of nuclei and for resolving ER, Golgi, mitochondria, lysosomes, and peroxisomes from mouse liver in a single analytical run has been developed by Graham et al. (1994). Starting tissue is homogenized, filtered, and subjected to differential velocity centrifugation to prepare crude nuclear and mitochondrial pellets, essentially as described in Basic Protocol 1. Sucrose media (8% w/v) used for homogenization include 20 mM Tricine-NaOH (pH 7.4), 25 mM KCl, and either 5 mM MgCl2 (when purifying nuclei) or 1 mM EDTA (when preparing the crude mitochondrial fraction for subsequent analytical centrifugation). Also, the crude mitochondrial fraction is sedimented 15 min at 17,000 × g instead of 30 min at 10,000 × g. Working solutions (50% iodixanol) are prepared by diluting Optiprep (60% iodixanol) with either 120 mM Tricine-NaOH (pH 7.4)/150 mM KCl/30 mM MgCl2 (for nuclei) or 120 mM Tricine-NaOH (pH 7.4)/6 mM EDTA (for crude mitochondria). For one-step purification of nuclei, the crude nuclear pellet (resuspended in homogenization medium using a Dounce homogenizer) is diluted 1:1 (v/v) with 50% iodixanol solution, and 4 ml is layered over 4 ml each of 30% and 35% iodixanol solution (prepared by mixing homogenization medium and working solution). These volumes are for Beckman SW41 12.5-ml tubes. Centrifugation in a swinging-bucket rotor at 10,000 × g for 20 min yields purified nuclei at the 30%/35% iodixanol interface. For analytical fractionation, the crude mitochondrial pellet resuspended (Dounce homogenizer) in homogenization medium is adjusted to 35% iodixanol with working solution
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and 2.5 ml (per SW41 tube) is layered (using a syringe with long cannula or a long-stem pipet) under a 10% to 30% (w/v) continuous iodixanol gradient (prepared as in Fig 4.1.2 using 5 ml each of 10% and 30% iodixanol solution). Gradients are centrifuged in a swinging-bucket rotor at 52,000 × g for 1.5 hr, and fractions are collected by one of the methods identified in step 11 above. Analysis of organelle marker activities shows distinct distributions for the five organelles mentioned. Notably, the investigators also show that similar resolution can be obtained by using a self-forming iodixanol gradient in which the resuspended mitochondrial pellet is diluted to 17.5% iodixanol and centrifuged at 270,000 × g for 3 hr in a fixed-angle rotor (Graham et al, 1994).
GEL FILTRATION TO ISOLATE SECRETORY VESICLES Gel filtration has not been applied widely in organelle purification; however, for small, homogeneous vesicles that can be included in the matrices of selected inert resins, it has proven to be valuable. As an example of this technique, the following protocol describes the use of Sephacryl S-1000 to fractionate the equivalent of a microsomal pellet obtained from a secretory vesicle–accumulating sec6 mutant of S. cerevisiae (based on Walworth and Novick, 1987).
BASIC PROTOCOL 3
NOTE: During the first run of the column, substantial loss of material may occur, as there appears to be adsorption of lipid-rich vesicles to the column matrix. Adsorption can be substantially reduced by initially running sonicated phospholipid vesicles as a sample (this is also recommended when using controlled-pore glass). Alternatively, glyceryl CPG glass beads, which have a hydrophilic coating (polyethylene glycol or a similar polymer) that reduces surface adsorption, may be used.
Materials Saccharomyces cerevisiae sec6-mutant strain spheroplasted in 1.4 M sorbitol Running buffer: 0.8 M sorbitol/10 mM triethanolamine/1 mM EDTA (pH 7.2; adjust pH with acetic acid) 1.5 × 90–cm to 1.5 × 100–cm column packed with Sephacryl S-1000 superfine resin (Pharmacia Biotech) and equilibrated with running buffer at 4◦ C 1-ml-capacity Dounce homogenizer Centrifuges: low-speed and ultracentrifuge Peristaltic pump capable of producing flow rates of ∼10 ml/hr Fraction collector Additional reagents and equipment for gel-filtration chromatography (UNIT 8.3) NOTE: Prepare solutions with Milli-Q-purified water or equivalent. NOTE: All operations after harvesting cells should be conducted at 0◦ C (ice bucket) or 4◦ C (cold room).
Load and run the column 1. Lyse spheroplasted yeast in running buffer using a Dounce homogenizer. Conditions for spheroplasting can be found in Becker and Lundblad (1994). The running buffer is made with sorbitol instead of sucrose because yeast metabolize sucrose.
2. Centrifuge the lysate 10 min at 10,000 × g, 4◦ C, to remove mitochondria and larger particulates. 3. Remove the supernatant and centrifuge 1 hr at 100,000 × g, 4◦ C, to produce a microsomal pellet.
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4. Resuspend the pellet in 600 µl running buffer and disperse using a Dounce homogenizer. 5. Remove running buffer from above the resin bed of the column and load the sample at 4◦ C. The column size suggested is suitable for most applications; a smaller-dimension column has been used by Barlowe et al. (1994) in fractionating small amounts of radiolabeled material. Additional information on column packing and setup may be found in UNIT 8.3.
6. Once the sample has entered the resin bed, overlay the bed with running buffer and, using a peristaltic pump, elute the column at a flow rate of ∼10 ml/hr, collecting fractions continuously. 80-drop (4-ml) fractions were collected by the investigators who designed this procedure.
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Figure 4.2.3 Distribution of organelle marker activities across eluted fractions from a Sephacryl S-1000 column on which a resuspended microsomal pellet from sec 6-4 mutant (or wild-type) yeast was loaded. Fractions were assayed for invertase (a secretory protein), α-mannosidase (a vacuolar marker), plasma membrane ATPase, and protein. Invertase marks the distribution of secretory vesicles that accumulate in the sec 6-4 mutant (but are absent in wild-type yeast). Part of the plasma membrane ATPase, but <1% of α-mannosidase, codistributes with invertase-containing secretory vesicles in the mutant. The protein indicates that significant protein elutes at the position of the vesicle peak for microsomes derived for mutant but not wild-type yeast. Reproduced from Walworth and Novick (1987) with permission of The Rockefeller University Press.
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Assay the column fractions and process those of interest 7. Vesicles small enough to enter the pores of the resin at least partially run in the included volume of the column; larger vesicles are excluded and appear in the void volume; and soluble proteins elute mostly after the vesicles. Locate the vesicle of interest in the column profile by assaying for a marker activity. A sample elution profile (Walworth and Novick, 1987) is shown in Figure 4.2.3. Invertase is used as a marker in the case of secretory vesicles (biosynthetically labeled secretory protein could also be used, as in Barlowe et al., 1994).
8. Pool the peak fractions and concentrate the vesicles for further study by centrifuging 1 hr at 100,000 × g, 4◦ C.
PREPARATIVE ELECTROPHORETIC SEPARATION Electrophoretic purification can be used to separate complex mixtures of small vesicles that differ only modestly in size and density, because it takes advantage of the differences in their surface charges. This technique can be used preparatively or (as is more common) at the analytical level (Alternate Protocol 5), to separate and analyze organelles that have been selectively labeled. In this protocol, agarose electrophoresis using material derived from rat liver (Kedersha and Rome, 1986) is presented as an example of a preparative electrophoresis procedure. Although the protocol presented here used buffers of pH 6.5, the authors state that other pHs can be used, with resulting changes in the relative migration of different vesicles (as determined empirically).
BASIC PROTOCOL 4
Materials Homogenization medium: 0.25 M sucrose (ultrapure; e.g., ICN Biochemicals)/ 50 mM 4-morpholinoethanesulfonic acid (MES), pH 6.5 6.25% sucrose/6.25% Ficoll 70/50 mM MES, pH 6.5 50 mM MES, pH 6.5 Running buffer: 50 mM MES (pH 6.5) with added sucrose as desired 0.15% to 0.2% Isogel agarose (FMC), or equivalent, in running buffer Centrifuges: low-speed and ultracentrifuge Standard flatbed agarose gel electrophoresis apparatus with 0.06-in. holes drilled in the cover to allow passage of inlet and outlet tubing from sample collecting wells (construct or purchase from Idea Scientific) Standard 1.5-mm-thick plastic (siliconized) or Teflon gel comb 1.5-mm-thick plastic (siliconized) or Teflon eluting combs cut so that teeth are 3 mm wider than loading wells 0.062-in.-o.d. (1.57-mm) polyethylene tubing (Clay-Adams PE-160) Peristaltic pump capable of flow rates of 0.2 to 15.0 ml/hr Fraction collector Additional reagents and equipment for preparing microsomal fractions (see Basic Protocol 1) Prepare crude microsomal fraction 1. Prepare a crude rat liver microsomal fraction according to the procedure for differential centrifugation by velocity (see Basic Protocol 1), except using homogenization medium containing 50 mM MES. At the last step (see Basic Protocol 1, step 13), resuspend the crude microsome pellet in 6.25% sucrose/6.25% Ficoll 70/50 mM MES using a Dounce homogenizer. 2. Centrifuge the homogenate 40 min at 40,000 × g, 4◦ C, to pellet larger membrane vesicles.
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3. Dilute the resulting supernatant with 4 vol of 50 mM MES or homogenization medium and centrifuge 2 hr at 100,000 × g, 4◦ C. Use homogenization medium (containing 0.25 M sucrose) in steps 3 and 4 if possible osmotic damage is a concern.
4. Resuspend the pellet (concentrated microsomes) in ≤0.8 ml of 50 mM MES (pH 6.5) or homogenization medium, and microcentrifuge 3 min at top speed to remove aggregates. Save the sample on ice until it can be applied to the gel (step 9). Samples for preparative electrophoresis can contain as much as 40 to 80 mg/ml protein.
Prepare and load the gel 5. Melt 0.2% Isogel agarose in running buffer using a microwave oven. FMC Isogel agarose is preferred because it has a low content of charged groups; it is usually used at concentrations of 0.15% to 0.2% but may be applied to separation of larger organelles (>200 nm diameter) when used at lower concentrations (<0.15%).
6. Cast the gel, with both the sample loading and eluting combs carefully prepositioned, and allow to solidify ≥2 hr at 4◦ C. Eluting combs are used to facilitate recovery of any particles that have diffused laterally (perpendicular to the field) during electrophoresis. Siliconization of plastic combs is recommended to minimize tearing of low-percentage agarose gels.
7. Remove eluting comb and immediately fill the eluting well(s) with running buffer. Keep the sample comb in place until just before loading. 8. Place the gel in the apparatus, taking care to align the elution wells with the holes in the lid. Electrophoresis is performed at 4◦ C, so all further operations should be conducted at this temperature (in a cold box or cold room).
9. Remove sample comb and load sample (from step 4). 10. Fill buffer reservoirs with running buffer until just level with the upper surface of the gel. 11. Attach the lid of the apparatus. Connect two pieces of 0.062-in.-o.d. polyethylene tubing whose ends have been cut away on one side to a peristaltic pump. Insert tubing through the holes in the lid and fully into the eluting well (see Fig. 4.2.4). The tubing provides buffer inlet and sample collection from the eluting well(s). The cutaway sides on the ends of the tubing should face one another for efficient buffer delivery and sample collection.
Run the gel and continuously collect the eluted fractions 12. Connect the electrodes, with the direction of electrophoresis running toward the (+) electrode (see Fig. 4.2.4), and turn on the power supply and the peristaltic pump. Set the current at 35 mA (∼40 V). Adjust the flow rate on the peristaltic pump empirically. For example, for a 2-cm-wide elution well set 4 cm downfield from the sample well in a 1-cm-thick gel and an electric field of 1 V/cm, a flow rate of 4 ml/hr is reported to give quantitative elution. Purification of Organelles from Mammalian Cells
13. Place a flask of warm water on the apparatus during the run to prevent condensation on the underside of the lid. 14. Carry out the electrophoresis for 16 hr.
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Figure 4.2.4 Schematic showing the horizontal agarose gel system devised by Rome and colleagues. Note the loading well, near (−) electrode, and the eluting well with continuous flow, near (+) electrode. The inset at the lower left shows the configuration of inlet and outlet tubing in the eluting well; note the cutouts from the walls of the tubing within the well. Figure courtesy of L. Rome with permission of Analytical Biochemistry.
15. Collect 3 to 4 ml fractions (generally, 30 to 45 total fractions) and assay initially by analytical SDS-PAGE (UNIT 10.1) or by more specific assays for selected markers (e.g., dot blotting with an antibody). The duration of electrophoresis and the size of fractions collected will generally have to be adjusted empirically to suit the particular fractionation being performed.
16. Concentrate pooled fractions by centrifuging 30 min at 100,000 × g, 4◦ C. Resuspend the pellets in an appropriate buffer for further analysis.
ELECTROPHORETIC SEPARATION ON A DENSITY GRADIENT As indicated in the initial discussion of electrophoretic separations above (see Basic Protocol 4), electrophoresis has proven to be very useful for separating specific membrane populations from among similar-sized vesicles where there are unique surface charge characteristics. In this protocol, the unusually high negative surface charge on endosomes has enabled the identification of a novel population that accumulates MHC class II molecules en route to the cell surface. Its distribution following electrophoresis differs from the distributions of markers of other organelles but corresponds to the distributions of invariant chain processing and antigenic peptide loading (Tulp et al., 1994).
ALTERNATE PROTOCOL 5
Additional Materials (also see Basic Protocol 4) Rat starved overnight and freshly sacrificed Electrophoresis buffer: 0.25 M sucrose (ultrapure; e.g., ICN Biochemicals)/10 mM triethanolamine/10 mM acetic acid/1 mM EDTA, pH 7.4 Trypsin, TPCK-treated (Worthington Biochemical or equivalent) Soybean trypsin inhibitor 10%, 6%, 5%, and 4% (w/v) Ficoll 70, prepared in electrophoresis buffer
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Electrophoresis apparatus: cylindrical electrophoresis chamber with dimensions of 5 cm high and 2.2 cm i.d. with circular palladium and platinum electrodes and cation-permeable membrane (Ionics, Inc.; see Tulp et al., 1993) or Polyprep 2000 apparatus (Buchler) Gradient-forming device capable of forming linear concentration gradients of 10 ml volume (Hoefer Pharmacia) 1-mm-gauge stainless steel mesh Additional reagents and equipment for preparing microsomal fractions (see Basic Protocol 1) NOTE: Prepare solutions with Milli-Q-purified water or equivalent. NOTE: All operations after removing and weighing tissue, including forming and running the gradient, should be conducted at 4◦ C.
Prepare crude microsomal fraction 1. Remove liver from freshly sacrificed rat that has been fasted overnight (to deplete hepatic glycogen). Weigh, mince, homogenize, filter, and centrifuge at low speed as for centrifugation by differential velocity (see Basic Protocol 1, steps 1 to 4), using electrophoresis buffer as the homogenization medium and centrifuging 15 min at 840 × g to remove nuclei and larger debris. 2. Recover the supernatant (PNS), assay its protein content, and add trypsin (25 µg/mg PNS protein). Incubate 5 min at 37◦ C. Trypsin treatment is employed by most investigators who use electrophoresis to fractionate cellular organelles by electrophoresis. Treatment with trypsin enables compartments of the endocytic pathway to be resolved from other microsomal constituents (Marsh et al., 1987). As mentioned under limitations in Strategic Planning, employing a protease restricts subsequent analyses to components that are refractory to digestion.
3. Stop the digestion by adding soybean trypsin inhibitor (100 µg/mg protein) and cooling on ice. Centrifuge 60 min at 100,000 × g, 4◦ C. 4. Resuspend pellet (crude microsomal fraction) in 0.5 ml of 5% Ficoll 70. Maintain on ice until it can be loaded for electrophoresis (step 7).
Set up the electrophoresis apparatus and load with the gradients and sample 5. Fill electrophoresis chamber with 10% Ficoll 70 via the inlet at the bottom. Fill the lower buffer reservoir with electrophoresis buffer. A schematic diagram of an apparatus for electrophoretic separation is shown in Figure 4.2.5. A similar device has been designed by Lindner (2001) and used to resolve Golgi, endosomes, and plasma membrane.
6. Attach the gradient maker to the top inlet of the chamber and introduce a gradient (total volume 6 ml) descending from 10% to 6% Ficoll 70 above the 10% Ficoll. The gradient is formed and introduced into the electrophoresis chamber by displacement of the 10% Ficoll solution, which is allowed to drip slowly out of the lower inlet.
7. Load the crude microsomal sample (from step 4) above the Ficoll gradient, again by displacement of the 10% Ficoll solution, which is allowed to drip out of the lower inlet of the electrophoresis chamber. Purification of Organelles from Mammalian Cells
8. Above the loaded sample, form and load a 9.5-ml continuous gradient of 4% to 0% Ficoll 70 by displacement of the 10% Ficoll at the bottom of the chamber. Upon completion, remove the loading inlet from above the electrophoresis chamber.
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Figure 4.2.5 Apparatus used for electrophoresis on a density gradient. a, separation column (5 cm tall; 2.2 cm wide) containing the density gradient; b, bottom reservoir; c, top reservoir; d, outlet for the top reservoir; e, inlet for the bottom reservoir; f, outlet for the bottom reservoir; i, inlet for the separation column; m, cation-permeable membrane (part 61 AZL-386, Ionics Inc.), which is held in place by a threaded fixture; o, inlet for the upper reservoir; p, circular (3.8-cm2 ) palladium anode; pt, platinum cathode (3.8 cm2 ); t, top cone with flow deflector (the inlet through which the Ficoll gradients are delivered). After removing t, the 1-mm-mesh stainless steel screen is placed atop a while the upper reservoir is filled, and the removed. After removing the screen, pt is installed and the electrophoretic separation is started. Reproduced from Tulp et al., 1993, with permission of VCH Verlagsgesellschaft.
9. Allow 10% Ficoll 70 solution to drip from the inlet at the bottom of the electrophoresis chamber until the meniscus is level with the top of the electrophoresis chamber. Place 1-mm-gauge stainless steel screen over the meniscus (to prevent mixing when the upper reservoir buffer is added) and fill the upper reservoir with electrophoresis buffer. 10. Remove the screen and attach the top (+) electrode. Connect the peristaltic pump to the buffer reservoirs; the upper reservoir buffer is exchanged at 1 ml/min while the lower reservoir is exchanged at 5 ml/min during the run.
Start the electrophoresis 11. Carry out electrophoresis at a constant current of 10 mA (3 V/cm field) for a total of 40 min. At two times during the run, allow 6 ml of 10% Ficoll 70 in electrophoresis buffer to drip slowly (1 ml/min) from the bottom of the electrophoresis chamber, then replace with 6 ml of fresh 10% Ficoll 70. Note that electrophoresis of vesicles with a negative surface charge occurs upward.
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Figure 4.2.6 Results from density gradient electrophoresis of various organelles derived from cultured melanoma cells. (A) Distribution of horseradish peroxidase (HRPO) after endocytosis at 37◦ C. Filled diamonds show distribution of early endosomes (4 min HRPO uptake) and open squares show distribution of late endosomes (4 min HRPO uptake followed by 30 min chase at 37◦ C). (B) Distribution of the lysosomal hydrolase β-hexosaminidase (×) and plasma membranes that were marked in intact cells by surface iodination at 4◦ C using lactoperoxidase (filled circles). (C) Distributions of Golgi membranes (galactosyltransferase activity; open circles); ER (biosynthetically labeled by a 2-min pulse with 35 S methionine/cysteine; filled squares), and total protein (open triangles). The shaded region indicates where the bulk of a novel compartment containing MHC class II molecules are concentrated. Data are from Tulp et al. (1994); reprinted with permission from Nature (copyright 1994, Macmillan Magazines Ltd.).
Replacement of some of the Ficoll 70 is a precaution taken to prevent formation of a low-conductivity layer at the surface of the cation-permeable membrane at the base of the chamber.
Stop the electrophoresis and unload the chamber 12. After ending the run, drain the upper buffer reservoir and reattach the loading inlet above the electrophoresis chamber. 13. Unload the contents of the chamber by upward displacement by pumping 10% Ficoll 70 through the inlet at the bottom of the chamber. Collect the contents in 25- to 30-fractions and use for further analysis. Purification of Organelles from Mammalian Cells
Figure 4.2.6 (from Tulp et al., 1994) illustrates clearly the distinct distributions of different organelles that have been resolved from a crude microsomal preparation of melanoma cells by density gradient electrophoresis.
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SOLID-PHASE IMMUNOADSORPTION Solid-phase immunoadsorption using specific antibodies is a powerful technique for isolating cellular organelles that can be used on both the preparative and analytical levels. The procedure requires a high-quality antibody that reacts to an antigenic epitope exposed on the surface of the organelle of interest; the antigen must be reasonably specific to the organelle of interest. The antibody is adsorbed to a support (e.g., fixed Staphylococcus aureus cells, protein A–Sepharose, or magnetic beads), and the organelles of interest bind to the antibody support.
BASIC PROTOCOL 5
The procedure presented below describes use of antibody-coated magnetic beads, an approach developed for organelles by Gruenberg and Howell (1986). An antibody to the cytoplasmic tail of the polymeric immunoglobulin receptor is bound to the beads coated with secondary antibody; it is then used to isolate relatively large organelles—stacked elements of the Golgi complex—and vesicles that can be induced to bud in vitro from the stacked Golgi elements (Salamero et al., 1990). Also see immunoadsorption procedures presented in UNIT 4.3.
Materials 5 mg/ml BSA in PBS Primary antibody that selectively recognizes an antigenic epitope present on the exposed surface of the vesicle of interest 5 mg/ml BSA in 0.25 M sucrose (ultrapure; e.g., ICN Biochemicals)/0.1 M potassium phosphate (pH 6.65) Rat starved overnight and freshly sacrificed Homogenization medium: 0.5 M sucrose/0.1 M potassium phosphate (pH 6.65)/ 5 mM MgCl2 1.25 M, 1.3 M, and 1.4 M sucrose (ultrapure) in 0.1 M potassium phosphate (pH 6.65)/5 mM MgCl2 4.5-µm-diameter magnetic beads coated with sheep anti-mouse Fc antibody (Dynal) Magnetic collection device (e.g., Dynal) End-over-end rotator capable of slow rotation (2 to 4 rpm; e.g., Pelco rotator, modified) 10-ml and 38-ml centrifuge tubes suitable, respectively, for Beckman Type 50 and SW28 rotors NOTE: Prepare solutions with Milli-Q-purified water or equivalent. NOTE: All operations after removing and weighing tissue should be conducted at 0◦ C (ice bucket) or 4◦ C (cold room).
Coat the magnetic beads with primary antibody 1. Wash an appropriate amount of 4.5-µm-diameter magnetic beads twice with 1 ml of 5 mg/ml BSA in PBS in a microcentrifuge tube. Collect beads with a magnetic collection device between washes. Resuspend to a concentration of 2 mg beads/ml in the same medium. Coating the beads with antibody (steps 1 to 4) is usually done the day before the rest of the experiment. The beads are treated with protein to block nonspecific adsorption. Most procedures call for using bovine serum albumin (5 mg/ml in PBS or in buffered 0.25 M sucrose); one protocol recommends use of fetal bovine serum (3% in PBS) as the best way to reduce nonspecific binding (Saucan and Palade, 1994).
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Magnetic concentration is very quick and requires <1 min per wash. Dynal markets a number of magnetic collection devices, one of which is appropriate for microcentrifuge tubes. Alternatively, a plate magnet (Edmund Scientific) can be used beneath a cell culture dish (Howell et al., 1989). In addition to 4.5-µm beads, Dynal also markets 2.8-µm beads, beads coated with antimouse antibody, and tosyl-activated beads that can be custom-coupled to the antibody of choice by the investigator using a very simple procedure described by the manufacturer.
2. Add primary antibody (in 1 ml of 5 mg/ml BSA in PBS) to 2-fold excess over the amount of bound secondary antibody. Incubate 4 hr to overnight at 4◦ C in a microcentrifuge tube on an end-over-end rotator set at 2 to 4 rpm. Information regarding the amount of secondary antibody bound to the magnetic beads is provided by the manufacturer. A rotator for electron microscopy samples available from Pelco can be easily modified for the slow rotation needed for this protocol.
3. Wash the beads by adding 1 ml of 5 mg/ml BSA in PBS, 4◦ C, then collecting the beads magnetically or by microcentrifugation (30 sec at 10,000 rpm). Repeat twice with 5 mg/ml BSA in 0.25 M sucrose/0.1 M potassium phosphate, pH 6.65. In general, a control for nonspecific binding can be run in parallel using beads that have been coated with an irrelevant primary antibody.
Prepare sample for immunoadsorption Example: Prepare sample enriched in stacked Golgi complexes from rat livers: 4. On the same day the immunoadsorption experiment is to be done, sacrifice a rat that has been starved overnight (to deplete hepatic glycogen) and remove liver. Weigh liver, mince, homogenize, and filter as for velocity differential centrifugation (see Basic Protocol 1), except homogenize at 20% (w/v) in 0.5 M sucrose/0.1 M potassium phosphate/5 mM MgCl2 homogenization medium. This method for preparing a partially purified fraction enriched in stacked Golgi complexes from rat livers is based on the procedure of Leelavathi et al. (1970).
5. Prepare a postnuclear supernatant (PNS) by centrifuging sample as for differential centrifugation by velocity (see Basic Protocol 1, step 4). 6. Place a 4-ml cushion of 1.3 M sucrose/0.1 M potassium phosphate/5 mM MgCl2 into a 10-ml centrifuge tube, and layer 6 ml PNS on top. Centrifuge 60 min at 105,000 × g (in Type 50 rotor), 4◦ C. 7. Collect the band at the surface of the 1.3 M sucrose layer and adjust to 1.1 M sucrose using homogenization medium. 8. Prepare a second set of discontinuous gradients containing 6 ml each of 1.4 M, 1.3 M, and 1.25 M sucrose in 0.5 M potassium phosphate/5 mM MgCl2 . Layer 6 ml of the adjusted sample (step 7) over each gradient, overlay with homogenization medium to fill the tubes, and centrifuge 90 min at 80,000 × g (in SW28 rotor), 4◦ C. 9. Collect the Golgi fraction at the interface between the load and the 1.1 M sucrose layer. Adjust to 0.25 M sucrose and 5 mg/ml BSA. Place on ice. The Golgi fraction is now ready to be used for immunoadsorption (which should be done on the same day). The fraction should be assayed for protein content and maintained on ice until use. Purification of Organelles from Mammalian Cells
Adsorb organelles to antibody-coated beads 10. Add a sample of the organelle fraction (∼0.5 mg protein in 0.3 to 0.4 ml of 0.25 M sucrose/5 mg/ml BSA) to 1 mg antibody-coated magnetic beads in 0.6 to 0.8 ml of
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5 mg/ml BSA in PBS. Incubate 3 hr at 4◦ C with end-over-end rotation to allow binding to occur. Generally, the optimal ratio of input fraction to coated beads is determined in a preliminary binding experiment in which several 1-mg aliquots of beads are incubated with increasing amounts of the organelle fraction to determine the saturation point of binding. Bound and free organelles can be assessed by immunoblotting. For functional studies involving vesicle budding from adsorbed organelles, it is necessary to saturate antigen binding sites so that the vesicles can’t bind once they have budded. Salamero et al. (1990) reached saturation at 0.5 mg (protein) of Golgi fraction per mg of beads.
Wash away unadsorbed organelles 11. Wash the coated beads with 0.25 M sucrose/5 mg/ml BSA. Repeat as many times as needed to remove all unadsorbed organelle, collecting the beads magnetically between washes. The number of washes needed should be determined in the preliminary immunoblotting experiment mentioned above. For adsorbed small vesicles (e.g., endosomes), the concentration device built for microcentrifuge tubes is satisfactory. For large organelles such as the stacked Golgi complexes, it is preferrable to use culture dishes and a plate magnet so that the adsorbed organelles are not damaged or dislodged by mechanical shear.
12. Use the immunoadsorbed fraction for further studies. Examples of further applications include treatment with SDS solubilization buffer to desorb bound protein for SDS-PAGE and immunochemical analysis; suspension in fixative followed by preparation for electron microscopic analysis; incubation with a second primary antibody followed by a colloidal gold–conjugated secondary antibody to test for co-localization with the immunoadsorbing antigen by immunoelectron microscopy (e.g., Laurie et al., 1993); and functional studies of vesicle trafficking (Salamero et al., 1990).
PURIFICATION BY LECTIN ADSORPTION Lectin adsorption has proven to be particularly useful for plasma membrane isolation and to a lesser degree for isolation of endosomes and lysosomes that have been loaded by endocytosis of lectin conjugates at the cell surface. The latter procedures fall into the category of fractionation by density perturbation (Basic Protocol 7). In some procedures, the lectin conjugate is added to intact cells, thereby guaranteeing specific binding to plasma membrane; this approach appears to be applicable to cultured cells but not to tissues. In other procedures that are applicable to tissue samples, immobilized lectins can be used to adsorb selectively to plasma membrane vesicles in cell homogenates or partially purified fractions (e.g., crude microsomes). This strategy is possible because following cell or tissue disruption during homogenization, plasma membranes vesiculate with the ectodomain (glycosylated surface) facing outward, whereas most cytoplasmic organelles retain a cytosolic-surface-outward orientation. Once the fraction is bound to the lectin and washed free of unbound organelles, it can be eluted by competing the interaction between lectin and vesicles using the appropriate hapten sugar.
BASIC PROTOCOL 6
In this protocol, wheat germ agglutinin (WGA) affinity chromatography is used to purify apical plasma membranes from homogenates of pancreatic acinar cells (Paul et al., 1992). WGA is a lectin that binds both N-acetylglucosamine (GlcNAC) and N-acetylneuraminic acid (Neu5Ac) residues in glycoproteins, and interactions with WGA can be competed with free N-acetylglucosamine, which is relatively inexpensive. Additional guidelines for lectin affinity chromatography are presented in UNIT 9.1. Extraction, Stabilization and Concentration
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Materials Freshly sacrificed Sprague-Dawley rat (200 to 275 g), fasted overnight Homogenization medium (see recipe; add 0.1 mM PMSF just before fractionation), 0◦ C 0.2 M KCl/0.2 M NaBr/50 mM Tris·Cl, pH 6.8 50 mM Tris·Cl, pH 6.8 (APPENDIX 2E) Wheat germ agglutinin (WGA) conjugated to Sepharose (Pharmacia Biotech), preequilibrated with 50 mM Tris·Cl, pH 6.8 0.1 M N-acetylglucosamine (e.g., Sigma) in 50 mM Tris·Cl, pH 6.8 Motor-driven Teflon-glass homogenizer Cheesecloth Centrifuge capable of producing a field of 25,000 × g and equipped with a fixed-angle rotor Centrifuge tubes 5-ml chromatography column NOTE: Prepare solutions with Milli-Q-purified water or equivalent. NOTE: All operations after removing and weighing tissue should be conducted at 0◦ C (ice bucket) or 4◦ C (cold room).
Prepare a crude plasma membrane fraction 1. Remove pancreas from rat, weigh (generally 0.6 to 1.0 g tissue per rat), submerge in 0◦ C homogenization medium, and mince coarsely with scissors. Paul et al. (1992) used 6 g of pancreas. Thus, more than one rat may be required.
2. Homogenize at 10% (w/v) in homogenization medium by 10 strokes with a Teflonglass homogenizer. Filter the homogenate through four layers of cheesecloth. Remove a small aliquot (∼1% of total volume) for assay of marker activities. 3. Centrifuge 25 min at 120 × g, 4◦ C, to remove large cellular debris (most nuclei and larger). 4. Collect the supernatant and centrifuge 15 min at 13,000 × g, 4◦ C, to pellet zymogen granules and mitochondria. 5. Collect the supernatant and centrifuge 60 min at 25,000 × g, 4◦ C, to pellet plasma membranes and other vesicles. 6. Discard the supernatant and resuspend the pellet in 1 to 2 ml of 0◦ C homogenization medium. Remove an aliquot (∼1% of total volume) and assay for marker activities. Marker assays include: (1) γ -glutamyl transferase, an apical plasma membrane marker (Orlowsky and Meister, 1963); (2) Na,K-ATPase, a basolateral plasma membrane marker (Baykov et al., 1988); and (3) alkaline phosphatase, a duct cell plasma membrane marker (Emmelot and Bos, 1966). It is reported that 65% of the marker activity of the apical plasma membrane, γ -glutamyl transferase, is present in this fraction (Paul et al., 1992). If activities are being followed during the course of purification, assaying all fractions generated at each step is recommended so that it will possible to account for all activity. Purification of Organelles from Mammalian Cells
7. Dilute the rest of the resuspended membranes 10-fold with 0.2 M KCl/0.2 M NaBr/ 50 mM Tris·Cl, pH 6.8. Resediment by centrifuging 60 min at 25,000 × g, 4◦ C. NaBr is a chaotropic agent that has been used frequently in fractionating pancreas to promote desorption of extrinsic proteins from membrane vesicles.
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8. Resuspend the pellets in 3 ml of 50 mM Tris·Cl, pH 6.8. Mix with 3 ml of WGASepharose preequilibrated in 50 mM Tris·Cl, pH 6.8. Gently mix for 30 min in a closed tube, then transfer the entire mixture to a 5-ml chromatography column. 9. Wash the column after loading with at least 15 ml (5 vol) of 50 mM Tris·Cl, pH 6.8. 10. Elute the bound vesicles using 0.1 M N-acetylglucosamine in 50 mM Tris·Cl, pH 6.8. Concentrate the membrane-containing fractions as necessary by centrifuging 60 min at 25,000 × g, 4◦ C. Most of the membrane vesicles elute within 2 to 4 ml. This can be confirmed by assaying γ -glutamyl transferase of the fractions (a very easy spectrophotometric assay; Orlowsky and Meister, 1963). Measurements of the specific activity of γ -glutamyl transferase (per protein) in the homogenate and the final purified membranes indicates that the purification is 75-fold, which is excellent for plasma membranes.
DENSITY SHIFT USING DIGITONIN Density-shift techniques have found use in both analytical and preparative centrifugation procedures. In this approach, the sample to be separated is subjected to a treatment that differentially alters the densities of the component organelles, which are then separated by centrifugation. The following density-shift technique (based on the work of AmarCostasec et al., 1974) involves the use of digitonin and capitalizes on differences in cholesterol content in the membranes of various cell organelles. Controlled, sublytic concentrations of digitonin are applied to the membranes, leading to the formation of 1:1 molecular complexes with cholesterol, and these complexes result in increased buoyant density during cell fractionation. The magnitude of the effect varies depending on the cholesterol content of different organelle membranes; consequently, different organelles exhibit distinct density shifts as judged from the distribution of marker activities. The example given is for microsomes from rat liver but can be adapted to membranes from other sources.
BASIC PROTOCOL 7
Materials Rat starved overnight to deplete glycogen stores and freshly sacrificed Homogenization medium: 0.25 M sucrose (ultrapure; e.g., ICN Biochemicals)/ 3 mM imidazole (pH 7.4) 20 mg/ml digitonin stock solution (see recipe) 10% (w/v) Triton X-100 Digitonin density shift gradient stock solutions (see recipe) Sample for treatment and analysis Motor-driven Teflon-glass homogenizer (type C, A.H. Thomas) Cheesecloth Low-speed refrigerated centrifuge Ultracentrifuge with fixed-angle (e.g., Beckman 70Ti), and swinging-bucket rotors (e.g., Beckman SW28) Ultracentrifuge tubes to fit rotors Gradient-forming device (e.g., Hoefer Pharmacia) Additional reagents and equipment for microsomal fraction preparation (see Basic Protocol 1) NOTE: Prepare solutions with Milli-Q-purified water or equivalent.
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NOTE: All operations after removing and weighing tissue should be conducted at 0◦ C (ice bucket) or 4◦ C (cold room).
Prepare crude microsomal sample 1. Remove liver from a sacrificed rat (6 to 7 g tissue) and prepare total microsomal preparation as in differential centrifugation by velocity (see Basic Protocol 1, steps 1 to 8 and 11 to 12), except using homogenization medium containing 3 mM imidazole. Imidazole (pH 7.4) was the buffer used when the procedure was devised; MOPS or HEPES can be substituted at the same concentration.
2. Resuspend the microsomal pellet thoroughly in homogenization medium using a Teflon-glass homogenizer and wash by repelleting under the same conditions used to pellet the microsomes initially (see Basic Protocol 1, steps 11 and 12). 3. Resuspend the pellet (total washed microsomes) in homogenization medium, using 1 ml medium per 1 g liver used in step 1 and keep on ice until use.
Determine the digitonin concentration to use for density shift by dose-response assay 4. For liver microsomes, proceed to steps 6 to 8; for membranes from other tissues, determine the digitonin concentration to use for density shift by dose-response assay as follows: Prepare two sets of samples (100 µl/sample) each containing identical amounts of resuspended microsomes. To each set add 270 µl of appropriate digitonin solution to give final concentrations of 0 to 9 mg/ml. To one set, also add Triton X-100 to 0.2% final concentration; to the other set, add equal amounts of water. Incubate 15 min at 0◦ C. Be sure to add the digitonin in the same final volume and as a solution that contains 0.25 M sucrose. Dilute the 20 mg/ml digitonin stock solution to 15 mg/ml using 1 M sucrose; then mix 0.25 M sucrose with 15 mg/ml digitonin in sucrose to give 1.5, 3, 4.5, 6, and 12 mg/ml digitonin, corresponding to final concentrations of 1.1, 2.2, 3.3, 4.4, and 8.8 mg/ml digitonin respectively after addition to samples. The Triton X-100 treatment solubilizes the membrane; this set of samples is used to assay total activity. The set of samples treated with water is used to determine latency of activity—i.e., only limited activity will be detected until digitonin reaches a concentration that causes membrane lysis, at which point nearly the same activity will be detected as in the presence of Triton.
5. Assay the samples for the marker activities of specific organelle(s)—i.e., for the presence of an enzyme activity or other marker that is located in the interior of the vesicle of interest. Determine the highest digitonin concentration that does not cause membrane lysis. In the dose-response performed by Amar-Costasec et al. (1974), 2.7 vol digitonincontaining solution was added to each sample, and the samples were assayed for ER nucleoside diphosphatase and Golgi galactosyltransferase activities. The appropriate sublytic concentration was determined to be 1.7 mg/ml digitonin (Merck), which the authors estimate corresponds to a molar ratio of digitonin/cholesterol of slightly greater than 1:1.
Prepare sucrose density gradients 6. In the bottom of each centrifuge tube, place a cushion of 2.7 M sucrose. For an SW28 tube with 37-ml capacity, this cushion is 3 ml. Purification of Organelles from Mammalian Cells
7. Using the gradient-forming device and working at room temperature, generate a linear sucrose gradient above the cushion extending from 1.9 M sucrose (density 1.25 g/ml) to 0.8 M sucrose (density 1.10 g/ml) as follows: place the 0.8 M sucrose gradient
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Figure 4.2.7 Distribution of marker activities for various organelles after equilibrium centrifugation. Thin lines and open vertical arrows show the distribution and median density, respectively, of control samples; bold lines and filled vertical arrows show the distribution and median density, respectively, of digitonin-treated samples. Activity profiles are determined across the entire gradient (1.10 to 1.27 g/ml); digitonin treatment shifts the activity profile to higher density in proportion to the amount of cholesterol in the membrane. ER (glucose-6-phosphatase; NADH–cytochrome c reductase) and mitochondrial (monoamine oxidase; NADH–cytochrome c reductase) marker activities are practically unshifted, corresponding to the negligible cholesterol content of these organelles. Golgi (galactosyltransferase) and plasma membrane (5 -nucleotidase, alkaline phosphodiesterase, and alkaline phosphatase) marker activities are shifted by 0.015 and 0.030 to 0.036 g/ml (median density), respectively. From Amar-Costasec et al. (1974) by permission of The Rockefeller University Press.
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stock in the back chamber of the gradient maker (15 ml for SW28 tube) and allow it to flow into the junction between the two chambers. Place 15 ml of 1.9 M sucrose gradient stock in the mixing chamber. Open the junction between the chambers of the gradient maker, turn on the stirring device in the mixing chamber, and activate the peristaltic pump to generate the gradient. Once the gradient is generated, chill in a 4◦ C cold room. Because of the high molarity and viscosity of the 1.9 M sucrose, it is placed in the mixing chamber and the gradient is generated high-density-first. Thus the outlet tube must remain above the surface of the gradient as it is formed in the centrifuge tube (see Fig. 4.1.2). The gradients are prepared at room temperature to decrease viscosity.
Treat microsomes and subject to density gradient centrifugation 8. Prepare two samples from the same batch of microsomes used for the dose-response assay (steps 4 and 5)—1 ml each, if an SW28 rotor will be used, or smaller if scaled down—in homogenization medium. Add 2.7 vol of appropriate-concentration icecold digitonin solution to one of the samples. Incubate 15 min on ice. For liver microsomes, Amar-Costasec et al. (1974) used 0.235% (w/v) digitonin in 0.25 M sucrose to give 1.7 mg/ml digitonin.
9. Load the samples on the continuous gradients and centrifuge overnight (12 to 15 hr) at ∼82,500 × g (e.g., 26,000 rpm in SW28) to reach equilibrium.
Collect the gradients and analyze 10. Puncture the tubes with a needle and collect fractions dropwise from the bottom. Alternatively, collect the fractions manually from the top using a micropipettor, taking care to withdraw the viscous solutions slowly so that volumes of fractions remain uniform. 11. Assay the fractions for the marker activities of specific organelles and compare the distributions of activities for digitonin-treated and control samples. Examples of results obtained by Amar-Costasec et al. (1974) are shown in Figure 4.2.7.
12. If relevant, compare the distribution of a protein of interest to those of the marker proteins in the presence and absence of digitonin treatment. BASIC PROTOCOL 8
DENSITY SHIFT USING COLLOIDAL GOLD CONJUGATES Colloidal gold conjugates have not been used very widely as density shift reagents in cell fractionation; however, they offer a number of potential advantages. Conjugates are relatively easy to prepare, can be made with various proteins of interest, and can induce a substantial density shift; the colloidal gold can be produced in a variety of distinct sizes and, because it scatters electrons, can be visualized readily in an electron microscope. In the following procedure, different protein–colloidal gold conjugates have been used to prepare specific organelle fractions from lymphocytes (Beaumelle et al., 1990) and HEp.2 cells (Futter et al., 1995). Selective compartment labeling depends on the trafficking routes normally taken by the protein used in the conjugate and the conditions of incubation of the conjugate with the cells before fractionation.
Materials
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1% (w/v) trisodium citrate dihydrate 1% (w/v) tannic acid (Mallinkrodt) 25 mM K2 CO3 1% (w/v) chloroauric acid (HAuCl4 ; Fisher) in twice-distilled water (store in brown bottle; stable for months at 4◦ C)
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Proteins for conjugation: ricin (galactose-specific lectin; Sigma), anti-mouse transferrin receptor antibody (I.W. Trowbridge), and low-density lipoprotein (LDL; density 1.019 to 1.063 g/ml) 10% (w/v) NaCl 1% (w/v) BSA Phosphate-buffered saline (PBS, APPENDIX 2E) containing 0.5% polyethylene glycol (PEG) 20,000 (Carbowax 20M, Fluka) Cultured cells of an appropriate cell line to be used as a source of membrane fractions: e.g., BW 5147 mouse T cell lymphoma cells (characterized by I.W. Trowbridge, Salk Institute; available from ATCC) or HEp.2 cells (ATCC CCL23) Dulbeccos Modified Eagle Medium (DMEM) containing 100 mg/ml BSA DMEM containing 1% horse serum DMEM containing 5 mg/ml BSA PBS (APPENDIX 2E) Hypoosmotic homogenization medium (see recipe) Hyperosmotic buffer (see recipe) RNase (Sigma) DNase I for treating cell homogenates (Sigma) Pellet resuspension solution: 0.25 M sucrose (ultrapure; e.g., ICN Biochemicals)/ 1 mM EDTA/10 mM Tris acetate (pH 8.0) Continuous density gradient solutions: 30% and 60% (w/v) sucrose (ultrapure) in 1 mM EDTA/10 mM Tris acetate (pH 7.5) 2.5 M sucrose in water Nitrogen bomb and Dounce homogenizer for homogenization Low-speed refrigerated centrifuge Ultracentrifuge with fixed-angle and swinging-bucket rotors (e.g., Beckman ultracentrifuge with Type 50 and SW40 or SW41 rotors) Gradient-forming device Micropipettor NOTE: Prepare solutions with Milli-Q-purified water or equivalent. NOTE: All operations after harvesting cells should be conducted at 0◦ C (ice bucket) or 4◦ C (cold room).
Prepare colloidal gold conjugates 1. Prepare a reducing mixture containing 4 ml of 1% trisodium citrate dihydrate, 0 to 5 ml of 1% tannic acid (depending on the particle size desired), and the same volume of 25 mM K2 CO3 . Add Milli-Q water to 20 ml. As discussed in detail by Slot and Geuze (1985), the amount of tannic acid added determines the size of the particles obtained in the gold sol. Addition of 0.01, 0.1, 0.5, or 5 ml tannic acid yields (after the following steps) gold particles of average diameter 16, 10, 5, and 3 nm, respectively. Potassium carbonate is added to compensate for the acid from the tannic acid so that the pH is maintained between 7.5 and 8. Slot and Geuze used glass-distilled (rather than Milli-Q) water in the original procedure.
2. Mix 1 ml of 1% HAuCl4 with 79 ml Milli-Q water. Heat the gold solution and the reducing mixture separately to 60◦ C, then rapidly mix together with stirring, the solution will turn wine red when the gold sol forms—this takes a few seconds in the presence of maximum tannic acid and up to 1 hr in the absence of tannic acid. After sol formation is finished, continue to heat the samples until boiling. Allow to cool to room temperature and store in clean glass containers until used (up to several months).
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3. To determine the minimum amount of protein required to stabilize the gold sol, set up several 0.25-ml samples of sol and add increasing amounts of the protein to be conjugated (0 to 10 µg using 2 mg/ml stocks in water). Mix, then add 10% (w/v) NaCl to each sample to a final concentration of 1%. If the solution turns blue, then the gold sol has flocculated and the amount of protein is insufficient for stabilization; if the solution stays red, the sol is stable. Slot and Geuze (1985) state that blue color development is slow and subtle in samples that retain excess tannic acid. In this case they recommend adding 0.1% to 0.2% H2 O2 to destroy the tannic acid and facilitate normal color development. Ricin, anti-mouse transferrin receptor antibody, and LDL are the proteins conjugated in the work of Beaumelle et al. (1990), which is being used as an example. Ricin is a galactose-specific lectin (CAUTION: Ricin is a toxin and must be handled with care). Anti-mouse transferrin receptor antibody is discussed in Lesley et al. (1984). Low-density lipoprotein (LDL; density 1.019 to 1.063 g/ml) is purified from fresh human plasma using the procedure of Havel et al. (1955), which involves flotation from NaCl/KBr solutions.
4. Prepare a supply of the stable conjugate by adding more than twice the minimum amount of stabilizing protein with stirring. After 1 min, add BSA to 0.1% final (for added stability). Protein-gold conjugates formed this way are quite uniformly sized. Beaumelle et al. (1990) conjugated 1.9 mg ricin and 1.0 mg anti-transferrin receptor antibody with 100 ml gold.
5. Pellet the protein-gold conjugates in a fixed-angle rotor for 45 min at 50,000 × g (for gold ≥10 nm) or at 125,000 × g (for gold <10 nm), 4◦ C. The pellets usually have a tightly packed portion that adheres to the tube and a larger loose portion that is easily withdrawn with a pipet. Take only the latter. 6. Resuspend the protein-gold conjugate in 3 to 5 ml PBS containing 0.05% PEG 20,000 (as an extra stabilizer). Where protein-gold conjugates are made with serum LDL, the conjugation procedure used is different (Handley et al., 1981), and the conjugate is used immediately. A solution of 1.5 µg LDL in 0.5 ml of 50 mM EDTA (pH 5.5) is added with rapid stirring to 5.0 ml colloidal gold (prepared at the same pH using sodium citrate). The LDL-gold conjugate is sedimented away from excess LDL through a 35% sucrose cushion (centrifuging 20 min at 9000 × g) in a swinging-bucket rotor. The loose pellet is resuspended in cell culture medium and used directly for incubation.
Incubate protein-gold conjugates with cells 7a. For isolation of plasma membranes: Incubate 108 cultured cells 1 hr on ice in 5 ml DMEM containing 100 µg/ml BSA and 200 µl gold-ricin conjugate (from step 6). This step stops all membrane traffic. Beaumelle et al. (1990) report that the lectin concanavalin A does not bind to the plasma membrane sufficiently to cause a density shift during subsequent fractionation.
7b. For isolation of endosomes: Incubate 108 cultured cells 2 hr at 22◦ C in 5 ml DMEM containing 100 µg/ml BSA and 130 µl anti-mouse transferrin receptor antibody–gold conjugate. Incubation at 22◦ C retards both recycling back to the cell surface and trafficking from endosomes to lysosomes. Note that under steady-state conditions, 80% of transferrin receptors in these cells are concentrated in endosomes. Purification of Organelles from Mammalian Cells
7c. For isolation of coated pits: Incubate 108 cultured cells 2 hr at 10◦ C in 5 ml DMEM containing 100 µg/ml BSA and 130 µl anti-mouse transferrin receptor antibody–gold conjugate.
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Incubation at 10◦ C inhibits internalization of coated pits. Note that a large fraction of the transferrin receptor that is present on the cell surface is concentrated in coated pits.
7d. For isolation of lysosomes: Incubate 2 to 5 × 108 cultured cells 18 hr at 37◦ C in DMEM containing 1% horse serum, 5 mg/ml BSA, and 0.8 mg LDL-gold conjugate. Withdraw the medium with a pipet and replace with DMEM containing 5 mg/ml BSA and 150 µg/ml LDL. Incubate an additional 3 hr at 37◦ C. The second incubation (with unconjugated LDL) serves to chase internalized LDL to lysosomes and compete away any surface-associated LDL-gold.
8. After incubation, wash the cells twice with DMEM containing 100 µg/ml BSA and once with PBS (both at the temperature used in step 7a-7d), then chill on ice.
Lyse and homogenize labeled cells and prepare postnuclear supernatant 9. Wash the cells once with hypoosmotic homogenization medium, then scrape (or resuspend, if not attached) into the same buffer, using 1 ml buffer per 5 × 107 cells. 10. Place the cells in a nitrogen bomb for 20 min at 750 lb/in2 and release pressure according to manufacturer’s instructions. Transfer the initial homogenate to a Dounce homogenizer (without altering the composition of the medium) and homogenize with 20 strokes. The initial homogenization using the nitrogen bomb results in ∼30% lysis of the BW 5147 cells (judged by trypan blue exclusion; UNIT 5.5). After Dounce homogenization lysis is ∼90% with no increase in nuclear damage. If a nitrogen bomb is not available, then other homogenization procedures should be tried. Other publications from the same laboratory that developed this procedure report homogenization by passage 15 times through a syringe equipped with a 21-G needle (Futter and Hopkins, 1989) and by repeated passage through a micropipet tip (Beardmore et al., 1987). To a large extent, the homogenization procedure that works best will depend on the type of cells being studied.
11. Add 0.1 vol hyperosmotic buffer and centrifuge 5 min at 800 × g, 4◦ C. 12. Remove the supernatant (PNS). Add 0.1 mg/ml RNase and 0.2 mg/ml DNase I and incubate 3 min at 37◦ C. This step digests any accessible chromatin and RNA and improves the ensuing purification by decreasing aggregation of the total membrane pellet that is prepared in the next step.
Pellet the total membranes of the PNS 13. Pellet organelles in the PNS by centrifuging 30 min at 120,000 × g, 4◦ C, in a fixedangle rotor. These centrifugation conditions will reliably pellet organelles that contain bound colloidal gold.
14. Resuspend the pellets in pellet resuspension solution, using a small-volume Dounce homogenizer if necessary to obtain thorough dispersal.
Prepare and run the continuous sucrose gradients 15. Place sucrose cushions consisting of 0.5 ml of 2.5 M sucrose in the bottom of 13-ml ultracentrifuge tubes. Instructions are given for 13-ml tubes and a SW40 or SW41 rotor, but may be adjusted to different sizes if desired.
16. In each tube, generate a 12-ml continuous (60% to 30%) sucrose gradient above the cushion using the gradient solutions as follows: place 6 ml of 30% sucrose solution in the back chamber of the gradient maker, and fill the junction tube to the mixing chamber. Place 6 ml of 60% sucrose solution in the mixing chamber. Open the
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junction between chambers, start stirring and pumping, and generate gradients (high density first) above the cushion. The delivery tube from the gradient-forming device should be kept above the surface of the fluid in the centrifuge tube.
17. Load 0.5-ml aliquots of the resuspended pellet on top of each gradient and centrifuge 16 hr at 200,000 × g, 4◦ C, in an ultracentrifuge with swinging-bucket rotor (e.g., SW40 or SW41).
Collect the fractions for analysis 18. Unload the gradient from the top in 1-ml fractions. Manual collection with a micropipettor will work fine as long as the high-viscosity sucrose is collected slowly. Otherwise use an automated collection device (Buchler, Nycomed Pharma, or MSE Scientific; see Alternate Protocol 1, step 11). In all cases the organelles that have been purified by density shift are found in the fraction that is just above the pinkish pellet.
19. Analyze the fractions by assaying for marker activities or for protein by twodimensional SDS-PAGE (UNIT 10.4), electron microscopy, and immunoprecipitation or immunoblotting. In the published procedures, the authors also carried out all these procedures using 125 I-radiolabeled protein-gold conjugates and were able to estimate yields of the various organelles by the density shift technique. They obtained 40%, 32%, 10%, and 16% of the associated radiolabel in the high-density fractions of plasma membranes, endosomes, coated pits, and lysosomes, respectively.
BASIC PROTOCOL 9
EXTRACTION OF EXTRINSIC PROTEINS FROM MEMBRANES USING SODIUM CARBONATE The methods for cell fractionation presented elsewhere in this unit focus mostly on purifying specific organelles without further subfractionation into component parts. For many studies that focus on membranes of organelles or on specific membrane-associated proteins, it is desirable to have an easy way to categorize integral and extrinsic polypeptides. Extraction with 0.1 M sodium carbonate (pH 11.5) at 0◦ C (Fujiki et al., 1982) has gained widespread popularity as an easy and efficient method for selectively stripping extrinsic proteins off membranes without affecting the disposition of integral components (including transmembrane and lipid-anchored proteins). The treatment converts vesicles into small membrane sheets, releasing soluble and adsorbed proteins in the process. The membranes are recovered after the treatment by centrifugation. Carbonate extraction has been applied to numerous organelles including endoplasmic reticulum, Golgi, secretory vesicles, plasma membranes, lysosomes, and peroxisomes.
Materials 0.1 M sodium carbonate (Na2 CO3 ), pH 11.5, ice cold 0.3 M sucrose (ultrapure; e.g., ICN Biochemicals), buffered to pH 7 to 7.5 (e.g., with 10 mM Tris·Cl) Cell organelle or membrane fraction to be treated Ultracentrifuge with swinging-bucket rotor Purification of Organelles from Mammalian Cells
NOTE: Prepare solutions with Milli-Q-purified water or equivalent. NOTE: All operations after removing and weighing tissue should be conducted at 0◦ C (ice bucket) or 4◦ C (cold room).
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1. Pellet the organelles or vesicles to be extracted by centrifugation, using conditions described in the appropriate basic or alternate protocol. 2. Resuspend the pellets in a small amount of buffered sucrose sufficient to obtain a well-dispersed suspension. 3. Dilute the sample 50× to 1000× with ice-cold 0.1 M Na2 CO3 . Maintain 30 min on ice. A final protein concentration of 10 to 1000 µg/ml is generally appropriate. Assay protein content if there is concern that this range is exceeded.
4. Place a small cushion of 0.3 M buffered sucrose in a centrifuge tube (∼10% of the volume of the tube), then carefully layer the organelle suspension on top. Pellet the membranes by centrifuging 1 to 2 hr at ∼150,000 × g, 4◦ C. Although not specified in the original procedure, the buffered sucrose cushion prevents formation of a rock-like pellet that is often very difficult to resuspend.
5. Resuspend the pellets and use for assays. Although the high-pH treatment on ice generally does not affect subsequent immunoprecipitations, it can inhibit some enzyme activities (check in a preliminary experiment).
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Digitonin density shift gradient stock solutions Prepare three separate stock solutions: 0.8 M sucrose/3 mM imidazole, pH 7.4 (density 1.10 g/ml) 1.9 M sucrose/3 mM imidazole, pH 7.4 (density 1.25 g/ml) 2.7 M sucrose (density 1.34 g/ml) The 2.7 M sucrose is used for the cushion and contains no buffer. Sucrose should be ultrapure (e.g., ICN Biochemicals). Sucrose stocks can be filtered using 1.2-µm Millipore filter (or equivalent) and stored at least 2 weeks at 4◦ C or indefinitely at −20◦ C. 1 M imidazole (pH adjusted with concentrated HCl) can be filtered using 0.22-µm Millipore filter (or equivalent) and stored indefinitely at 4◦ C.
Digitonin stock solution, 20 mg/ml Dissolve 20 mg/ml digitonin in water, stirring for 15 min at 90◦ to 95◦ C. Filter through 0.22-µm Millipore filter (or equivalent) and store up to several weeks at 4◦ C. Digitonin is available from Merck and Sigma. Neither preparation is pure (they are contaminated mostly by other saponins that bind to cholesterol), but for purposes of the density shift experiment they can be treated as if pure. However, it is preferable not to switch brands without retitration.
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Homogenization medium (lectin adsorption) 0.28 M sucrose (ultrapure; e.g., ICN Biochemicals) 50 mM 2-N-morpholinoethanesulfonic acid (MES), pH 6.0 150 mM NaCl 10 mM MgCl2 0.1 mg/ml soybean trypsin inhibitor (Sigma) 0.1mM 4-(2-aminoethylbenzenesulfonyl fluoride) (AEBSF; Sigma) Pancreatic zymogen granules are labile above pH 7 and the low pH is used to keep them as intact as possible; 150 mM NaCl is included because it is reported to promote plasma membrane vesiculation with ectodomain facing outwards.
Homogenization medium (velocity differential centrifugation) 0.25 to 0.3 M sucrose (ultrapure; e.g., ICN Biochemicals) 50 mM Tris·Cl (pH 7.5; APPENDIX 2E; optional) 25 mM KCl (optional) 5 mM MgCl2 (optional) Proteinase inhibitor cocktail (optional) Although sucrose is the critical component of this medium, other additives are often included as well. Generally the medium is prepared fresh before use. A suitable proteinase inhibitor cocktail includes 1 µg/ml each (final concentration in sucrose homogenization medium) of leupeptin, antipain, pepstatin, and aprotinin (some investigators also include o-phenanthroline; all available from Sigma), along with 0.2 mM AEBSF (Calbiochem) or PMSF (phenylmethanesulfonyl fluoride), in 1 mM EDTA (pH 7). These components are added from 1000× stock solutions prepared ethanol (pepstatin and PMSF) or water and stored at −20◦ C. PMSF has a very short half-life in aqueous solution and should be added just before starting the homogenization.
Hyperosmotic buffer 700 mM KCl 40 mM magnesium acetate 1 mM dithiothreitol (DTT) 10 mM HEPES, pH 7.5 Make fresh before use Store DTT at −20◦ C as a 1 M stock solution in water.
Hypoosmotic homogenization medium (colloidal gold density shift) 15 mM KCl 1.5 mM magnesium acetate 1 mM DTT 10 mM HEPES, pH 7.5 Make fresh before use Store DTT at −20◦ C as a 1 M stock solution in water. Purification of Organelles from Mammalian Cells
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Percoll gradient stock solutions No-Percoll solution (low sucrose) 0.3 M sucrose 5 mM MOPS or MES buffer 1 mM EDTA 0.2 µg/ml DPPD (using 0.4 mg/ml DPPD in ethanol) Adjust pH to 6.5-6.7 using 1 N HCl
No-Percoll solution (high sucrose) 2.15 M sucrose 35 mM MOPS or MES 7 mM EDTA
Prepare solution from the following stock solutions: 2.5 M sucrose (ultrapure; e.g., ICN Biochemicals); 1 M 2-N-morpholinoethanesulfonic acid (MES), pH 6.5; 1 M 3-N-morpholinopropanesulfonic acid (MOPS), pH 6.8; 0.2 M EDTA, pH 7.0; and 0.4 mM DPPD (diphenyl-p-phenylenediamine; Kodak) in ethanol. 2.5 M sucrose can be stored indefinitely at −20◦ C; 1 M MES or MOPS and 0.2 M EDTA should be filtered using 0.22-µm Millipore filter (or equivalent) and can then be stored indefinitely at 4◦ C; and DPPD stock should be made fresh before use. The pKa values of MOPS and MES are 7.2 and 6.1, respectively. 60% Percoll/sucrose: Mix 60 ml Percoll, 14 ml No-Percoll solution (high sucrose), and 26 ml water. Adjust pH as described above and add 0.2 µg/ml DPPD (from 0.4 mg/ml DPPD stock in ethanol). 86% Percoll/sucrose: Mix 86 ml Percoll and 14 ml No-Percoll solution (high sucrose). Adjust pH as described above and add 0.2 µg/ml DPPD (from 0.4 mg/ml DPPD stock in ethanol). Percoll is slightly alkaline, so it is important to check and adjust the pH, especially if the organelles to be purified are labile at alkaline pH.
Rate zonal centrifugation gradient stock solutions Low-density gradient solution 0.5 M sucrose 4% (w/v) Ficoll 400 5 mM MES (pKa = 6.1) 1 mM EDTA 0.2 µg/ml DPPD
High-density gradient solution 1.8 M sucrose 4% (w/v) Ficoll 400 5 mM MES 1 mM EDTA 0.2 µg/ml DPPD
Prepare gradient solutions from the following stock solutions: 2.5 M sucrose (ultrapure; e.g., ICN Biochemicals); 50% Ficoll 400; 1 M 2-N-morpholinoethanesulfonic acid (MES), pH 6.5; 0.2 M EDTA, pH 7.0; and 0.4 mg/ml DPPD (diphenyl-pphenylenediamine; Kodak) in ethanol. Sucrose and Ficoll stocks are stored longterm at −20◦ C; gradient solutions are used within two weeks of preparation; and the DPPD stock is made fresh and used without storage.
Running buffer 0.8 M sorbitol 10 mM triethanolamine 1 mM EDTA, pH adjusted to 7.2 with acetic acid (TEA) Sorbitol is used because yeast metabolize sucrose. The running buffer generally is prepared fresh before use. Extraction, Stabilization and Concentration
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COMMENTARY Background Information
Purification of Organelles from Mammalian Cells
Organelle purification procedures capitalize on the differences in size, density, and (occasionally) surface charge density of individual types of organelles. Most fractionation procedures that are based on centrifugation involve some combination of procedures that distinguish both size and density. Initially, a homogenate is prepared in isoosmotic (or slightly hyperosmotic) sucrose or some other predominantly nonelectrolyte medium. Sucrose solution deviates from ideality; while 0.3 M sucrose and 150 mM NaCl have equivalent particle concentrations, 0.25 M sucrose and 150 mM NaCl have the same osmotic activities. Investigators generally homogenize cells and tissues in solutions that are somewhere in the range of 0.25 to 0.3 M sucrose. The homogenate is filtered through cheese cloth or nylon screen to remove residual large connective tissue debris. Then differential centrifugation by velocity is used at low speed to pellet large particulates—nuclei, erythrocytes, and larger particles (including unbroken and partially disrupted cells). Subsequent treatment of the postnuclear supernatant (PNS) is a function of the mission of the experiment as discussed in the protocols. Differential centrifugation by velocity (Basic Protocol 1) separates organelles within a cell homogenate, largely on the basis of organelle size, through successive spins of increasing centrifugal field and time in a 0.25 to 0.3 M sucrose medium. What does not sediment under the first set of conditions may sediment under one of the later sets of conditions. As all cellular organelles have a greater density than the medium, the separation is largely based on organelle size. The pellets produced during each centrifugation are enriched for organelles of a certain size but are also substantially contaminated with smaller organelles. DeDuve and Berthet (1954) calculated that in a tube where the bottom is twice as far as the surface of the liquid from the axis of rotation, a pellet containing the fastest sedimenting organelle will be contaminated by 40% of an organelle that sediments at one-third the speed and by 13% of an organelle that sediments with one-tenth the speed. Generally, further enrichment is necessary, and purity can be improved by resuspending the selected pellet in medium to its original volume and recentrifuging under the same conditions. Although differential centrifugation by velocity is mainly used to separate and concen-
trate crude organelle fractions, differential centrifugation by equilibrium (Basic Protocol 2) is generally used as a follow-up to further purify organelles from contaminants according to buoyant density differences. Continuous density gradients are widely used to generate unique (but often partially overlapping) distributions along the gradient for various types of organelles within a complex mixture. The resolution is well suited for analytical purposes where the goal is not necessarily preparative purification. In contrast, discontinuous (step) gradients are most frequently used for preparative purifications. Densities of media are selected so that the organelle of interest concentrates at the interface between particular density steps while other organelles either sediment or float to other interfaces. An alternative to differential separation by velocity or equilibrium is rate zonal centrifugation (Alternate Protocol 1), a nonequilibrium method in which organelles separate based on a combination of size, density, and shape into enriched regions within the gradient. This method is particularly suitable for separating several types of organelles into distinct distributions within a relatively short time (1 to 3 hr rather than 8 hr to overnight as in equilibrium centrifugation). However, it should not be viewed as a substitute for differential centrifugation by equilibrium, which is one of the best ways to separate fractions of organelles of similar size but different density (e.g., rough and smooth microsomes). Percoll-gradient fractionation (Alternate Protocol 2) has several advantages. The density gradients are isoosmotic with the homogenization medium, and due to the low viscosity, organelles approach equilibrium density quite rapidly (30 min for 1-µm organelles like granules and mitochondria; 2 hr for small (≤0.2-0.3 µm) vesicles like microsomes and endosomes). Also shallow density gradients are generated with relatively brief centrifugation; this can be advantageous for obtaining good spatial separation of organelle populations that differ only slightly in density from one another. Note, however, that because the shallow region of the gradient is bounded by regions where density changes sharply with distance in the tube (see Fig. 4.2.2), organelles with similar densities but not separated across the shallow region will tend to pile up at the boundaries. Thus Percoll is especially advantageous when the experimental goal can be reduced to separating two components from
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one another (or one component from the rest), and it is unlikely to be the method of choice for resolving complex mixtures into multiple subfractions that are individually enriched in different organelles. Also, it should be kept in mind that if a particular type of organelle has a heterogeneous density (e.g., endosomes), portions of the total population may be distributed on both sides of the shallow resolving segment of the gradient, depending on the concentration of Percoll selected for use. Deuterium oxide fractionation (Alternate Protocol 3) is an equilibrium density procedure in which H2 O is partly replaced with D2 O (heavy water). It is based on the fact that the density of D2 O is 1.1 g/ml—substantially closer than that of water to the density of biological organelles. In principle, this approach should be quite useful as an aid in separating organelles, for two reasons. First, inclusion of D2 O in density gradients can significantly reduce the osmolarity of solutions that are isopycnic with most biological organelles. Second, replacement of H2 O with D2 O can reduce the density difference between organelles and the medium. For two populations of organelles that differ only slightly from one another in density and sediment at only slightly different rates in H2 O-based media, reducing the density difference between the organelles and the medium is one way to magnify the differences in sedimentation rate (this can be verified using the Svedberg equation—see UNIT 4.1). In practice, the value of D2 O has fallen short of expectations, mostly because the densities of several organelles increase in heavy water due to exchange of D2 O for bound water. However, such exchange may be advantageous if it occurs to differing extents between organelles and thereby magnifies their density differences. For example, Beaufay et al. (1959) showed that the buoyant density of liver mitochondria is largely unchanged in H2 O/sucrose vs. D2 O/sucrose, whereas the buoyant densities of both lysosomes and peroxisomes substantially increased in the presence of heavy water. Both D2 O and glycerol (another popular density-modifying agent, e.g., Wagner et al., 1978) readily permeate membranes. In these media, internal and external solvents are identical. For organelles with relatively little internal solute (e.g., synaptic vesicles), sedimentation in D2 O or glycerol gradients has been quite useful in analyzing the physical characteristics of the membranes with minimized contributions from the internal solvent (Wagner et al., 1978).
Excellent fractionation procedures for a variety of cellular organelles have been described that make use of density gradient centrifugation in nonionic iodinated solutes (Alternate Protocol 4). As examples: Barlowe et al. (1994) report use of isopycnic centrifugation in Nycodenz for isolation of biosynthetically labeled, ER-derived transport vesicles from a cell-free assay; Loh et al. (1984) use metrizamide gradients in sucrose for purifying storage granules for peptide hormones from intermediate pituitary, demonstrating reasonably good separation of granules from lysosomes; Wattiaux et al. (1978) use metrizamide gradients to prepare highly purified lysosomes, well resolved from mitochondria and peroxisomes; and Graham et al., (1994) use fully iso-osmotic iodixanol gradients to purify nuclei and to separate ER, Golgi, mitochondria, lysosomes, and peroxisomes. The last three studies thoroughly account for a variety of marker enzyme activities. Gel filtration (Basic Protocol 3) using controlled-pore glass (CPG-3000, Sigma; mean pore diameter 3000 Å) has been employed routinely to prepare highly purified synaptic vesicles from rat brain (Huttner et al., 1983). Sephacryl S-1000 (Pharmacia Biotech), which also excludes particles larger than 3000 to 4000 Å in diameter, has been used to purify secretory vesicles, clathrin-coated vesicles and ER-Golgi/nonclathrin-coated transport vesicles from the yeast S. cerevisiae (Walworth and Novick, 1987; Mueller and Branton, 1984; and Barlowe et al., 1994). For complex mixtures of small vesicles, differences in size and density between individual types of vesicles are frequently insufficient to enable their separation as purified populations. In some of these cases, electrophoretic purification techniques—which capitalize on differences in surface charge density among the organelles—have proved useful at both the preparative and the analytical levels. Organelle purification using preparative electrophoresis (exemplified by Basic Protocol 4) has been performed in large-porosity solid matrices such as agarose—e.g., for clathrin-coated vesicles from bovine brain and liver and from CHO cells (Rubenstein et al., 1981) and the ribonucleoprotein particles known as vaults from rat liver (Kedersha and Rome, 1986). It has also has been carried out in liquid using density gradients (which provide stability but do not contribute to the separation). The latter procedure has been used in the preparative purification of endosomes
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Purification of Organelles from Mammalian Cells
from cultured cells (Marsh et al., 1987) and secretory vesicles from S. cerevisiae mutants (Holcomb et al., 1987, and Basic Protocol 4). In each case, preenrichment of the samples using centrifugation techniques have been used to overcome the limitations in total protein that can be loaded. More commonly, electrophoresis has been used at the analytical level to separate and analyze organelles that are selectively labeled by markers, as exemplified by Alternate Protocol 5. Cases where analytical electrophoresis has been applied include synaptic vesicles isolated from electric organs of marine elasmobranchs (Carlson et al., 1978) and specialized endosomes that accumulate MHC class II molecules (Amigorena et al., 1994; Tulp et al., 1994, Lindner 2001, and Alternate Protocol 5). Both preparative electrophoresis and analytical density-gradient electrophoresis capitalize on equipment that is relatively inexpensive. In the event that a free-flow electrophoresis apparatus (e.g., Bender and Hobein Elphorvap II; cost $100,000) is available, then the procedure for its use presented in detail in Marsh et al. (1987) and subsequently employed by Amigorena et al. (1994) should be seriously considered for either preparative- or analytical-scale isolations. Isolation of cellular organelles by solidphase immunoadsorption using specific antibodies (Basic Protocol 5) has become very popular in the past several years. Clearly, use of these procedures requires availability of a specific antibody capable of reacting with an antigenic epitope exposed on the surface of the organelle of interest, and relies on the quality of that primary antibody and the extent to which the antigen is concentrated in the compartment of interest. These procedures can be conducted at both the preparative and analytical levels. Earlier procedures used either fixed S. aureus or protein A–Sepharose with adsorbed antibody (e.g., Merisko et al., 1981). This approach has continued to be used, especially for preparative-level procedures (e.g., isolation of glucose transporter vesicles; Laurie et al., 1993). As an alternative (as described in Basic Protocol 5), magnetic beads that are coated with secondary antibodies (e.g., sheep anti-mouse Fc or sheep anti-rabbit Fc) provide a new-generation solid support that allows easy isolation and is especially amenable to both functional and morphological analysis of associated organelles. The use of magnetic beads as a solid support for isolating endosomes from cultured cells and subsequently analyzing their
fusion has been elegantly demonstrated by Gruenberg and Howell (1986). The properties of commercially available magnetic beads are discussed in Howell et al. (1989). An elegant procedure for immunoadsorption of endosomes is a major topic in UNIT 4.3. Basic Protocol 6 presents a method that uses affinity chromatography on the lectin wheat germ agglutinin (WGA) to purify apical plasma membranes from homogenates of pancreatic acinar cells (Paul et al., 1992; UNIT 9.2). WGA that binds both N-acetylglucosamine (GlcNAC) and N-acetyl neuraminic acid residues in glycoproteins, and interactions with WGA can be competed with free GlcNAc, which is relatively inexpensive. Other examples of lectin adsorption in subcellular fractionation include use of WGA conjugated to iron dextran for purification of plasma membrane vesicles from CHO cells in a strong electromagnetic field (Warnock et al., 1993) and WGA–colloidal gold conjugates to remove plasma membranes prior to purification of other organelles (Gupta and Tartakoff, 1989). A variety of density-shift techniques exist that are employed for both analytical and preparative purposes. Basic Protocol 6 presents a particularly useful density-shift strategy in which digitonin is applied to a crude organellar fraction; the compound binds to cholesterol, thereby increasing the buoyant density of different organelles to a lesser or greater extent depending on the cholesterol content of the organellar membranes. The protocol is based on the approach of AmarCostasec et al. (1974), who applied it to a crude microsomal fraction from rat liver to demonstrate a substantial shift of marker activities of the plasma membrane (which has a high cholesterol content), a moderate density shift of a marker activity of the Golgi (which has a lower cholesterol content), and negligible shifts of markers of endoplasmic reticulum (rough microsomes) and mitochondria (which both have little or no membrane cholesterol). In turn, the density shifts of these standard markers can serve as references for comparing the behavior of proteins whose localizations are not known. A complementary procedure involving use of EDTA or pyrophosphate to strip ribosomes from the endoplasmic reticulum can be used to achieve a selective density shift of rough microsomes (Amar-Costasec et al., 1974). Both binding to the surfaces of intact cells and endocytosis have also been used to achieve
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density shifts in selected organelles during subsequent fractionation. In one such procedure, receptor-mediated endocytosis of proteins conjugated to horseradish peroxidase (HRPO) has been used to label endocytic compartments which are then shifted to higher buoyant density by carrying out the cytochemical reaction of HRPO using H2 O2 and diaminobenzidene (Courtoy et al., 1984). Unfortunately, the oxidative cytochemical reaction induces wholesale protein cross-linking within the labeled compartment, significantly limiting the potential applicability of this approach (Ajioka and Kaplan, 1987). Another interesting (though rather specialized) application used density shift by an enzyme cytochemical reaction product to distinguish clathrincoated vesicles involved in endocytosis from other types of coated vesicles in liver (Helmy et al., 1986). On the other hand, several procedures using protein–colloidal gold (or irondextran) conjugates appear to offer both wider application and greater possibilities for further analysis of the isolated fractions (e.g., Warnock et al., 1993; Futter and Hopkins, 1989; Beaumelle et al., 1990; Futter et al., 1995). Among these, the use of ricin-gold, LDL-gold, and transferrin receptor antibody– gold conjugates to isolate selected compartments involved in endocytosis (Beaumelle et al., 1990) is presented as Basic Protocol 8. Although Basic Protocol 8 is not an organelle isolation protocol, it is the procedure most widely used to extract extrinsic (or adsorbed) proteins from organelle membranes. Not only has it enabled investigators with an interest in integral membrane proteins to efficiently remove soluble contaminating proteins but it has also been used to distinguish peripheral membrane proteins (those desorbed in the presence of sodium carbonate) from integral membrane proteins (those that remain membrane-associated following treatment). As such, it is complementary to phase partitioning in Triton X-114 in which integral, but not peripheral, proteins bind the detergent and sediment in micelles at the detergent cloud point (Bordier, 1981). Note, however, that lipid-anchored (fatty acylated or prenylated) proteins either partially or fully behave as integral membrane proteins in both procedures.
Critical Parameters and Troubleshooting These procedures can be performed using either tissue from freshly sacrificed animals
(stored frozen tissues are not usually satisfactory) or cultured cells. As compared to cultured cells, however, mammalian tissues offer at least three advantages for cell fractionation and purification of various organelles. First, the cells in mammalian tissues generally are highly differentiated and, depending on the functional specialization of the selected cell type, contain large numbers of organelles per cell. Second, the mass of cells available in tissue taken even from a single rat is much larger than is readily available from reasonably large size cultures. Finally, the connective tissue matrix surrounding cells in tissues contributes to shearing during homogenization and is often an aid in efficiently disrupting cells under reasonably gentle homogenization conditions. Of the procedures discussed in UNIT 4.1, a glass mortar and motor-driven Teflon pestle rotating at ∼1500 to 2000 rpm is used most widely for routine homogenization of tissues. If the investment of connective tissue is extensive (e.g., for parotid salivary gland) making direct homogenization with a Teflon pestle difficult, then a prehomogenization (10 sec at halfmaximal power with a Tekmar Tissumizer (or Polytron) inserted directly in the glass mortar) is used before homogenization with the Teflon pestle. The following requirements are critical for each purification method: Differential centrifugation by velocity The concentration of the homogenate should be standardized somewhere in the 10% to 20% (w/v) range; in especially concentrated homogenates, sedimentation properties change and aggregation is exacerbated. The early steps of differential centrifugation by velocity may be the only chance in a multi-step procedure to remove large organelles. Be sure the centrifugation is hard enough to achieve this task even if it means taking a partial loss of the desired smaller organelles. The loss can be reduced by resuspending the pellet, recentrifuging, and then pooling the second supernatant with the original supernatant. Differential centrifugation by equilibrium Density must be measured accurately— e.g., a refractometer should be used. In addition, reproducible preparation and collection of gradients is very important. Upper limits to gradient loads must be identified so that organelles do not aggregate appreciably and instead continue to sediment according to their individual buoyant densities.
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Deceleration at the end of the centrifugation should be done with the brake in the OFF position (at least for the final 1000 rpm) so that gradients are minimally disturbed by mixing. Following homogenization, vesicles deriving from a particular organelle may be heterogeneous in size and density. Further, the densities of different organelles, especially various types of small vesicles (microsomes, Golgi elements including subcompartments, and endosomes) may not differ much from one another. Thus the investigator should not expect separate, discrete bands. Rather, distributions of marker activities across the gradient must be compared. Rate zonal centrifugation using sucrose Reproducible preparation, loading, running, and unloading of gradients is essential. Conditions are particularly important where distributions of activities/markers are compared between tubes rather than within the same tube. Use of a refractometer to check the concentrations of solutions used to generate the density gradient is recommended. The refractometer is also useful for determining the density of gradient fractions. If a programmable centrifuge is available, use the same acceleration/deceleration program from run to run; otherwise, establish and adhere to a specific manual protocol. The separation between different organelle-enriched bands can be manipulated empirically to some extent by changing the length of centrifugation or the density limits of the gradient.
Purification of Organelles from Mammalian Cells
Gradient fractionation using percoll Because Percoll itself has negligible osmolarity, it must be added to isoomotic media. Almost all membrane-bounded organelles behave as osmometers and will lyse in hypoosmotic media. Although electrolyte-based solutions such as NaCl are generally used in separating populations of cells, organelles tend to aggregate in such media and are generally fractionated in nonelectrolyte-based media. Close attention should be paid in preparing Percoll-sucrose mixtures so that starting densities of the self-forming gradients are reproducible. This is critical in separating organelles with very similar densities. Because the density profile of the Percoll gradient continually changes during the run, centrifugation conditions (time and rpm) should be performed reproducibly. Centrifugation times that are too long can actually elevate the cross-contamination between low- and high-density fractions. After fractionation, it
is necessary to remove the Percoll from purified fractions, as it can interfere with further processing. The simplest method to accomplish this is via additional low-speed centrifugations (as described in Alternate Protocol 2) or through ultracentrifugation on a step gradient (Zastrow and Castle, 1987). If, instead, fractions are concentrated by high-speed centrifugation (as in several of the other protocols in this unit), Percoll will also sediment; it forms a glassy pellet and an overlying high-density gradient at the bottom of the tube. The vesicles of interest loosely congregate above the glassy pellet with the residual Percoll. Concentrating the fraction by centrifuging onto a high-concentration sucrose cushion can help because Percoll will sediment through the cushion. Alternatively, the vesicles of interest can be concentrated by flotation from a medium of higher buoyant density in which Percoll pellets. Finally, for SDS-PAGE of samples that contain residual Percoll, heating 5 to 10 min in sample buffer and then microcentrifuging 10 min at top speed prior to loading on the gel will remove nearly all remaining Percoll. Gradient fractionation using D2 O A main concern in using D2 O for fractionation is maintaining osmotic balance across organelle membranes so that lysis or leakage of contents is avoided. Gumbiner and Kelly (1981) include sucrose in their density gradient solutions to avoid lysis of secretion granules, whereas Wagner et al. (1978) include NaCl to stabilize synaptic vesicles. A similar strategy will be needed if fractionation in D2 O (or glycerol which also permeates membranes) is applied to other organelles. Gradient fractionation using nonionic iodinated solutes Solutions of metrizamide, Nycodenz, and iodixanol should be stored in the dark for stability. As with other density gradient procedures, the concentrations of solutions used to prepare gradients should be standardized using a refractometer. In addition, the densities of gradient fractions collected after centrifugation can be determined by measuring refractive index. Gel filtration The column resin needs to be pretreated to suppress loss of small vesicles due to adsorption. This problem is particularly acute the first few times the column is used. In using controlled-pore glass (CPG-3000), Carlson and Kelly (1978) recommend treating the
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column periodically with polyethylene glycol (PEG; Carbowax) to suppress adsorption. The principle is the same as that used commercially in coating colloidal silica particles with polyvinylpyrrolidone in preparing Percoll. Also, pretreating the resin (either Sephacryl or CPG) by running a sample containing small phospholipid vesicles helps to suppress subsequent adsorptive losses. Satisfactory and reproducible gel filtration of vesicles requires thorough dispersion of the starting sample. Most often this is a microsomal (small vesicle) pellet. After resuspending the pellet, it should be thoroughly dispersed by Dounce homogenization or by passage several times though a small-bore hypodermic needle (>25G). Briefly microcentrifuge the dispersed sample to pellet any residual aggregates and load the supernatant. Preparative and analytical electrophoretic separation To ensure that aggregation of starting material is minimal, after thorough dispersal of microsomal pellets, the samples should be microcentrifuged briefly before loading. Setting the flow rate of the peristaltic pump used to drain the collecting well of the preparative agarose electrophoresis device for efficient clearance of separated organelles is also essential. Solid-phase immunoadsorption The antibody immobilized on the solidphase support must be both stably bound and active. Generally, if an antibody is good for immunoprecipitation, it is a candidate for use in immunoadsorption (contingent on epitope availability). Mouse monoclonal antibodies that have a low affinity for protein A can be adapted for immunoadsorption by using either beads coated with secondary (anti-mouse antibody) or immobilized protein A and a rabbit anti-mouse antibody bridge. Always run a parallel control in which an irrelevant antibody is substituted for the specific antibody. This will indicate the level of nonspecific adsorption in the procedure. Before beginning the immunoadsorption, removal of any organelle aggregates is essential. This is especially critical where centrifugation is used to collect and wash the solid phase. As in the other procedures, thorough dispersal followed by brief centrifugation can be used to eliminate aggregates (which are hopefully only a minor fraction of antigen). For large and/or dense organelles (e.g., stacked Golgi complexes and exocrine secre-
tion granules), it is advisable to use a plate magnet to immobilize the magnetic beads and adsorbed organelles during washing. Repeated packing of the beads by centrifugation or in the commercially available magnetic concentrator used with microcentrifuge tubes can dislodge these organelles. Furthermore, large organelles may sediment independent of their specific association with the immunoadsorbant. Purification by lectin adsorption Identification of a lectin that will bind to the plasma membrane of the cells of interest is essential to the procedure. Efficient elution of bound vesicles is also important; Nacetylglucosamine may not effectively compete for multiple lectin interaction sites per vesicle. An assay for a plasma membrane marker is useful to judge recovery, i.e., the amounts of membrane bound, not bound, and eluted by the competing ligand with respect to the original amount used for adsorption. It is advisable to check marker activities for organelles that are not expected to adhere to the lectin to evaluate potential nonspecific adsorption of contaminating organelles. Density shift using digitonin As digitonin at higher concentrations behaves like a detergent and lyses most membrane-bounded organelles, a preliminary titration experiment is necessary to identify sublytic digitonin concentrations for use in density shift (see Basic Protocol 6, step 4). Density shifts may be fairly subtle, especially where the cholesterol content of the membrane is low. Thus it is important to assay carefully the complete distribution of all markers and the protein (or other macromolecule) of interest across the density gradient. Density shift using colloidal gold conjugates In preparing colloidal gold conjugates, it is essential to identify stabilizing conditions in which the protein coating on the gold particles is sufficient to prevent flocculation. A simple test is described in the protocol. As in most other procedures, minimal aggregation among organelles following homogenization is essential, otherwise, density perturbation extends to organelles that are associated artifactually with the particular densitymodified compartment. Extraction of extrinsic proteins from membranes using sodium carbonate Sodium carbonate treatment is most effective if the content of other electrolytes in the sample is low. This is the reason membrane
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pellets are resuspended in sucrose medium with only a low buffer content and then diluted with a large volume of the high-pH carbonate solution. Carbonate induces discontinuities in the membrane bilayer (see Fujiki et al., 1982) that may be important to effectively removing extrinsic proteins. Higher concentrations of NaCl (and possibly other electrolytes) reduce the formation of discontinuities. Buffers of course decrease the pH elevation. Do not neglect to include the sucrose cushion through which the membranes are spun after carbonate treatment. Membranes pelleted directly in carbonate are rock-like and extremely difficult to resuspend.
Anticipated Results
Purification of Organelles from Mammalian Cells
The key parameters to consider in judging the success of various preparative fractionation procedures are the yield of material and extent of purification. Yield is generally assessed in two ways. First, assays of organelle marker activities are used to determine what percentage of the total activity present in the original tissue homogenate is recovered in the purified fraction. This is a measure of whether the fraction is representative; generally recoveries >10% of total in the fraction of interest are satisfactory, especially as recovery is usually sacrified in the interest of purity. A complementary assessment in definitive studies involves analysis of the fractional contamination by undesirable organelles. The second measure is more practical and involves determining the amount of protein recovered in the fraction from the amount of tissue used. The extent of purification is usually judged by evaluating the enrichment of a marker activity from the homogenate to the final fraction. It is usually expressed as a ratio of activity per mg protein in the final fraction/activity per mg protein in the homogenate. For organelles that constitute large fractions of total protein—e.g., 20% for ER and mitochondria in liver (Wattiaux et al., 1978) and >25% for secretion granules in exocrine cells (Zastrow and Castle, 1987)—enrichment cannot exceed a factor of 4 or 5. However, if an organelle constitutes only 1% to 2% of total cell protein (e.g., Golgi and lysosomes in liver), then theoretical enrichments of 50- to 100-fold are possible. For analytical procedures, the success of separations is mainly judged by the comparative distributions of marker activities for distinct organelles. Examples of anticipated results are given in Figure 4.2.1 (rate zonal
centrifugation), Figure 4.2.6 (analytical electrophoresis on a density gradient), and Figure 4.2.7 (density shift using digitonin). Examples of expected results, particularly for the preparative centrifugation techniques, are given in the following discussion. Good yields but low purity should be expected when using differential centrifugation by velocity (Basic Protocol 1). The microsomal fraction constitutes 13% of homogenate protein (18 to 20 mg protein per gram of liver used) and contains 40% to 50% of ER marker activities (glucose-6-phosphatase and ERassociated esterase), 40% of a Golgi marker (galactosyl transferase), and 25% of a plasma membrane marker activity (5 -nucleotidase; Bergeron et al., 1982). By using differential centrifugation by equilibrium (Basic Protocol 2) subsequent to differential centrifugation by velocity, rough microsomes (derived mainly from rough ER) can be efficiently purified with a yield of 7.5 to 10 mg protein per gram of liver used (Adelman et al., 1973; Bergeron et al., 1982). Mitochondria can be purified by repeated differential centrifugation by velocity (Basic Protocol 1) with a yield of 15 mg protein per gram of rat liver used (CarvalhoGuerra, 1974). Unfortunately the purity was not reported, but it would not be expected to exceed 5-fold for reasons discussed above. Impressive purifications but often lower yields are expected using differential centrifugation by equilibrium (Basic Protocol 2) or its variations (e.g., Alternate Protocol 4). The intact Golgi fraction used as an example in Basic Protocol 2 is more than 100-fold purified from the homogenate (close to the theoretical limit, as Golgi represents close to 1% of liver protein; Wattiaux et al., 1978). The yield of total galactosyl transferase is unusually high (33% of total); however, the absolute amount of material obtained is low (0.24 mg protein per gram liver), as expected (Bergeron et al., 1982). In isolating hepatic lysosomes using gradient centrifugation in iodinated nonionic solutes (Alternate Protocol 4), Wattiaux et al. (1978) obtained a 10% yield of lysosomal marker activities in their purest fraction and the absolute amount of protein was ∼0.25 mg per gram of liver used. However, the purity of the fraction (65- to 80-fold) is extremely impressive, as lysosomes are estimated to be 1% to 1.4% of total liver protein. The procedure of Wattiaux et al. (1978) is a model for careful accounting of recovery of activities and analysis of cross-contamination of fractions by other organelles.
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Yields using Percoll gradient centrifugation (Alternate Protocol 2) generally can be expected to be quite good. For the secretion granules used as an example, about 30% of the secretory protein amylase is recovered in the purified fraction. Generally 20% to 25% of amylase is released from granules that lyse during homogenization, while another 20% fails to be released from cells during homogenization; the latter sediments in the nuclear pellet (see Zastrow and Castle, 1987, and references therein). For secretion granules, which are much more dense than other cytoplasmic organelles, the purity of fractions obtained using Percoll gradients can be expected to be quite good, provided conditions of centrifugation (Percoll density, times of runs) are used reproducibly. Where organelles of interest and contaminants have more similar densities, the possibility of contamination can be significant and needs to be considered. Even though the forte of Percoll is generating very shallow density gradients with good spatial resolution, biological organelles are heterogeneous (either intrinsically or especially after cell homogenization) and distribution of a particular organelle over a range of densities can be expected. Gel filtration to isolate secretory vesicles (Basic Protocol 3) is a procedure that is restricted to small organelles. Losses due to vesicle adsorption to the column matrix can be expected to be significant (possibly as much as 50%) during initial use of the column but should improve significantly thereafter. Even though vesicle purification using the column can be impressive, homogeneity should not be expected. Homogenization generates small vesicles from larger organelles and these will co-fractionate with the secretory (or synaptic) vesicles. Generally it is wise to follow the distribution of marker activities in the fractions eluted from the column (as in Fig. 4.2.3) if co-localization is being evaluated. The use of electrophoretic techniques (e.g., Basic Protocol 4 and Alternate Protocol 5), although quite promising, is still rather limited. The initial efforts suggest that both recovery and purity using the preparative procedure may be quite high for selected organelles (Kedersha and Rome, 1986). However, more widespread use is needed before it is possible to establish expectations for yield and purity. Lectin-based purification of plasma membrane fractions (Alternate Protocol 6) also has not been employed both widely and quantitatively. For the exocrine pancreatic apical plasma membrane fraction used as an example,
yields were not reported. However, the fact that most plasma membrane vesicles are oriented with ectodomain outward (and thus accessible to the lectin) suggests that the yield could potentially be good. The purification (75-fold enrichment over the homogenate) is very good (Paul et al., 1992). Density-shift using colloidal gold conjugates (Basic Protocol 7) appears to be quite promising for obtaining fractions enriched in different segments of the endocytic pathway in both quite good yield and purity (Beaumelle et al., 1990). The yields reported for plasma membranes, endosomes, coated pits, and lysosomes isolated by density modification are 40%, 32%, 10%, and 16%, respectively. Contamination by organelles outside the endocytic pathway appears negligible based on marker assays. Cross-contamination among endocytic compartments also appears to be low but potentially is limited by the extent to which individual compartments can be density-modified selectively in the cells prior to fractionation. Extraction of extrinsic proteins from membranes using sodium carbonate (Basic Protocol 8) generally can be expected to be 90%, while retention of integral proteins in sedimentable membranes is usually quantitative. A result suggesting partial extraction will be difficult to interpret. If established integral membrane proteins are underrecovered the final membrane pellets, then incomplete recovery of membranes following centrifugation needs to be considered. On the other hand, if the loss of the proteins of interest is selective with respect to the markers, then these proteins may be extrinsic but inefficiently extracted. In this case, distribution should be judged following a repeat extraction.
Time Considerations This section provides estimates of the time required for organelle purification by the procedures that are likely to be used most frequently. For the other protocols, total times and convenient breakpoints can be easily estimated from the steps that are listed. In differential centrifugation by velocity (Basic Protocol 1), sample preparation and homogenization takes about 20 min; centrifugation to obtain the PNS takes 10 min; ultracentrifugation to obtain the crude mitochondrial fraction requires 30 min, and further purification takes another 30 min; finally, ultracentrifugation to obtain the microsomal pellet requires 2 hr. For each step requiring resuspension of pellets, include an additional 10 min.
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Purification of Organelles from Mammalian Cells
In differential centrifugation by equilibrium (Basic Protocol 2), preparation of the density gradients requires 30 to 60 min in advance of starting the fractionation. Sample preparation and homogenization takes about 20 min while loading the gradients takes about 15 to 30 min depending on experience. Running the gradient takes 3 hr (with an additional 15 to 30 min to allow the rotor to coast to a stop) in the example given; however, in most equilibrium runs this step goes overnight. Allow ∼30 min to collect the bands from the gradients after the run, 90 min to pellet them by additional centrifugation, and 15 min to resuspend them for use or storage. In many procedures, the gradient step is used in series with differential centrifugation by velocity, so selected times as in Basic Protocol 1 need to be added in. In rate zonal centrifugation (Alternate Protocol 1), preparation of the gradients requires 30 to 60 min in advance of starting the fractionation. Sample preparation and homogenization takes 20 to 30 min (more time is allotted for surgery to remove pancreas, which is more difficult than for liver); low-speed centrifugations to prepare the PNS require 30 min together; loading gradients takes 15 to 30 min; gradient centrifugation requires 2 hr plus a 30-min coast of the rotor to stop; and collection of the fractionations either manually or with a gradientelution device requires 10 to 15 min per centrifuge tube. Any additional centrifugation to pellet fractions generally requires 1 to 1.5 hr, with 15 min to resuspend pellets at the end. For gradient fractionation using Percoll (Alternate Protocol 2), allow 60 min for sample preparation, homogenization, and generation of PNS as above. Any adjustment of density requiring measurement of refractive index takes ∼10 min, while centrifugation of the gradients requires 30 min to 2 hr, depending on the organelle being isolated. Collection of fractions after the run requires 5 or 15 min per centrifuge tube for manual collection or collection with a gradient-elution device, respectively. Subsequent centrifugation to separate most of the Percoll from the fractions requires 45 min in a low-speed centrifuge or 1 to 2 hr if sedimentation in an ultracentrifuge is used. Gradient fractionation using nonionic iodinated solutes (Alternate Protocol 4) requires similar amounts of time as above for sample preparation, homogenization, and centrifugation to generate a PNS. In the example protocol given, the rehomogenization and recentrifugation of the initial nuclear pellet adds another 20 min to give a total time of ∼60 min to
prepare the PNS for loading on gradients. A 10,000-rpm centrifugation to pellet mitochondria requires an additional 30 min. Continuous gradients can be made in advance of starting the experiment whereas discontinuous gradients are usually made right before use during the low-speed centrifugation steps; allow 10 min per tube. Ultracentrifugation of the gradients requires 2.5 to 3 hr (including coast of the rotor to a stop). Unloading gradients takes 10 to 15 min per tube. Any additional centrifugation to pellet the isolated fractions requires 1.5 hr plus 5 min per tube to resuspend the final pellets for assays or storage. For gel filtration (Basic Protocol 3), sample preparation and centrifugation to prepare the column load approximately follows the timecourse for differential centrifugation by velocity. Where yeast are used as a source of sample, allow 60 min to spheroplast and harvest the yeast by centrifugation before starting the homogenization. The gel-filtration column is run overnight; fractions are maintained at 4◦ C during assay, and then an additional 1 to 1.5 hr is needed for pooling the selected fractions and pelleting them by centrifugation. For electrophoretic separations (Basic Protocol 4 and Alternate Protocol 5), about 3 to 4 hr is required for preparing and homogenizing tissue and sedimenting the microsomal starting material for the electrophoretic separation. Centrifugation times are similar to those in Basic Protocol 1, and additional time is included to allow for assay of total protein and brief trypsinization (Alternate Protocol 5). The agarose gel for preparative electrophoresis (Basic Protocol 4) can be prepared in advance of the experiment and requires at least 2 hr to solidify before use. The density gradient for analytical electrophoresis (Alternate Protocol 5) requires ∼30 min and is set up just before loading. Preparative electrophoresis in agarose is conducted for 16 hr, while analytical electrophoresis on the density gradient will take an hour, including the solution changes during the run. Fractions for assay and further processing are collected continuously during electrophoresis on the agarose gel whereas fractions are collected at the end of the run on the analytical density gradient, requiring 30 min. Solid-phase immunoadsorption using antibody-coated magnetic beads (Basic Protocol 5) requires 4 hr overnight incubation in advance of the experiment to coat the beads with antibody and subsequently wash them. During the experiment, preparation of the PNS
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requires about 30 min (as in Basic Protocol 1). Subsequent centrifugation to prepare membranes to load on the gradient requires 1.5 hr (including loading and unloading the tubes) in the example given. Preparation and loading of discontinuous gradients, centrifugation, and collection of fractions requires a total of 2 to 2.5 hr. Immunoadsorption of organelles requires 3 hr and subsequent washing needs another 30 min before samples are harvested or used in functional assays. Extraction of extrinsic proteins from membranes using sodium carbonate (Basic Protocol 8) can be performed either immediately after preparation by any of the protocols listed or after storage of samples that have been resuspended in buffered sucrose and frozen. Carbonate treatment requires 30 min; loading of treated samples for pelleting through a sucrose cushion requires 5 min per centrifuge tube; and centrifugation requires 1 to 2 hr. Collecting the final supernatants and resuspended pellets requires 5 min per tube.
Literature Cited Adelman, M.R., Blobel, G., and Sabatini, D.D. 1973. An improved cell fractionation procedure for the preparation of rat liver membrane-bound ribosomes. J. Cell Biol. 56:191-205. Ajioka, R.S. and Kaplan, J. 1987. Characterization of endocytic compartments using the horseradish peroxidase/diaminobenzidene density shift technique. J. Cell Biol. 104:77-85. Amar-Costasec, A., Wibo, M., Thines-Sempoux, Beaufay, H., and Berthet, J. 1974. Analytical study of microsomes and isolated subcellular membranes from rat liver. J. Cell Biol. 62:717745. Amigorena, S., Drake, J.R., Webster, P., and Mellman, I. 1994. Transient accumulation of new class II MHC molecules in a novel endocytic compartment in B lymphocytes. Nature 369:113-120. Balch, W.E. and Rothman, J.E. 1985. Characterization of protein transport between successive compartments of the Golgi apparatus: Asymmetric properties of donor and acceptor activities in a cell-free system. Arch. Biochem. Biophys. 240:413-425. Balch, W.E., Dunphy, W.G., Braell, N.A., and Rothman, J.E. 1984. Reconstitution of the transport of protein between successive compartments of the Golgi measured by the coupled incorporation of N-acetylglucosamine. Cell 39:405-416. Barlowe, C., Orci, L., Yeung, T., Hosobuchi, M., Hamamoto, S., Salama, N., Rexach, M.F., Ravazzola, M., Amherdt, M., and Schekman, R. 1994. Cop II: A membrane coat formed by sec proteins that drive vesicle budding from the endoplasmic reticulum. Cell 77:895-907.
Baykov, A.A., Evtushenko, O.A., and Avoeva, S.M. 1988. A malachite green procedure for orthophosphate determination and its use in alkaline phosphatase–based enzyme immunoassay. Anal. Biochem. 171:266-270. Beardmore, J., Howell, K.E., Miller, K., and Hopkins, C.R. 1987. Isolation of an endocytic compartment from A431 cells using a density modification procedure employing a receptor-specific monoclonal antibody complexed with colloidal gold. J. Cell Sci. 87:495-506. Beaufay, H., Bendall, D.S., Baudhuin, P., Wattiaux, R., and DeDuve, C. 1959. Tissue fractionation studies: 13. Analysis of mitochondrial fractions from rat liver by density gradient centrifuging. Biochem. J. 73:628-637. Beaumelle, B.D., Gibson, A., and Hopkins, C.R. 1990. Isolation and preliminary characterization of the major membrane boundaries of the endocytic pathway in lymphocytes. J. Cell Biol. 111:1811-1823. Becker, D. M. and Lundblad, V. 1994. Introduction of DNA into yeast cells. In Current Protocols in Molecular Biology (F.A. Ausubel,R. Brent,R.E. Kingston,D.D. Moore,J.G. Seidman,J.A. Smith,K. Struhl, eds.) pp.13.7.113.7.10. John Wiley and Sons, New York. Bell, A.W., Ward, M.A., Blackstock, W.P., Freeman, H.N., Choudhary, J.S., Lewis, A.P., Chotai, D., Fazel, A., Gushue, J.N., Paiement, J., Palcy, S., Chevet, E., Lafreniere-Roula, M., Solari, R., Thomas, D.Y., Rowley, A., and Bergeron, J.J. 2001. Proteomics characterization of abundant Golgi membrane proteins. J. Biol. Chem. 276:5152-5165. Bergeron, J.J.M., Rachubinski, R.A., Sikstrom, R.A., Posner, B.I., and Paiement, J. 1982. Galactose transfer to endogenous acceptors within Golgi fractions of rat liver. J. Cell Biol. 92:139146. Bordier, C. 1981. Phase separation of integral membrane proteins in Triton X-114 solution. J. Biol. Chem. 256:1604-1607. Carlson, S., Wagner, J.A., and Kelly, R.B. 1978. Purification of synaptic vesicles from elasmobranch electric organ and the use of biophysical criteria to demonstrate purity. Biochemistry 17:1188-1199. Carvalho-Guerra, F. 1974. Rapid isolation techniques for mitochondria: Technique for rat liver mitochondria. Methods Enzymol. 31:299-305. Courtoy, P.J., Quintart, J., and Baudhuin, P. 1984. Shift of equilibrium density induced by 3,3 diaminobenzidene cytochemistry: A new procedure for the analysis and purification of peroxidase-containing organelles. J. Cell Biol. 98:870-876. DeDuve, C. and Berthet, J. 1954. The use of differential centrifugation in the study of tissue enzymes. Int. Rev. Cytol. 3:225-275. Dominguez, M., Fazel, A., Dahan, S., Lovell, J., Hermo, L., Claude, A., Melancon, P., and Bergeron, J.J.M. 1999. Fusogenic domains of Golgi membranes are sequestered into specialized
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regions of the stack that can be released by mechanical disruption. J. Cell Biol. 145:673-688. Emmelot, P. and Bos, C.J. 1966. Studies on plasma membranes. II. K+-dependent p-nitrophenyl phosphatase activity of plasma membranes isolated from rat liver. Biochim. Biophys. Acta 121:375-385. Fujiki, Y., Hubbard, A.L., Fowler, S., and Lazarow, P.B. 1982. Isolation of intracellular membranes by means of sodium carbonate treatment: Application to endoplasmic reticulum. J. Cell Biol. 93:97-102. Futter, C. and Hopkins, C.R. 1989. Subfractionation of the endocytic pathway: Isolation of compartments involved in the processing of internalized epidermal growth factor-receptor complexes. J. Cell Sci. 94:685-694. Futter, C.E., Connolly, C.N., Cutler, D.E., and Hopkins, C.R. 1995. Newly synthesized transferrin receptors can be detected in the endosome before they appear on the cell surface. J. Biol. Chem. 270:10999-11003. Graham, J., Ford, T., and Rickwood, D. 1994. The preparation of subcellular organelles from mouse liver in self-generated gradients of iodixanol. Anal. Biochem. 220:367-373.
Kedersha, N.L. and Rome, L.H. 1986. Preparative agarose gel electrophoresis for the purification of small organelles and particles. Anal. Biochem. 156:161-170. Laurie, S.M., Cain, C.C., Lienhard, G.E., and Castle, J.D. 1993. The glucose transporter GluT4 and secretory carrier membrane proteins (SCAMPs) colocalize in rat adipocytes and partially segregate during insulin stimulation. J. Biol. Chem. 268:19110-19117. Leelavathi, D.E., Estes, L.W., Feingold, D.S., and Lombardi, B. 1970. Isolation of a Golgi-rich fraction from rat liver. Biochim. Biophys. Acta 211:124-138. Lesley, J., Domingo, D.L., Schulte, R., and Trowbridge, I.S. 1984. Effect of an anti-murine transferrin receptor–ricin A conjugate on bone marrow stem and progenitor cells treated in vitro. Exp. Cell Res. 150:400-407.
Gruenberg, J.E. and Howell, K.E. 1986. Reconstitution of vesicle fusions occurring in endocytosis with a cell-free system. EMBO J. 5:3091-3101.
Lindner, R. 2001. One-step separation of endocytic organelles, Golgi/trans-Golgi network, and plasma membrane by density gradient electrophoresis. Electrophoresis 22:386-393.
Gumbiner, B. and Kelly, R.B. 1981. Secretory granules of an anterior pituitary cell line, AtT20, contain only mature forms of corticotropin and β-lipotropin. Proc. Natl. Acad. Sci. U.S.A. 78:318-322.
Loh, Y.P., Tam, W.W.H., and Russell, J.T. 1984. Measurement of pH and membrane potential in secretory vesicles isolated from bovine pituitary intermediate lobe. J. Biol. Chem. 259:82388245.
Gupta, D. and Tartakoff, A.M. 1989. Lectin– colloidal gold–induced density perturbation of membranes: Application to affinity elimination of the plasma membrane. Methods Cell Biol. 31:247-263.
Marsh, M., Schmid, S., Kern, H., Harms, E., Male, P., Mellman, I., and Helenius, A. 1987. Rapid analytical and preparative isolation of functional endosomes by free flow electrophoresis. J. Cell Biol. 104:875-886.
Handley, D.A., Arbeeny, C.M., Witte, L.D., and Chen, S. 1981. Colloidal gold–low density lipoprotein conjugates as membrane receptor probes. Proc. Natl. Acad. Sci. U.S.A. 78:368371.
Merisko, E., Farquhar, M.G., and Palade, G.E. 1981. Coated vesicle isolation by immunoadsorption on Staphylococcus aureus cells. J. Cell Biol. 92:846-857.
Havel, R.J., Eder, H.A., and Bragdon, J.H. 1955. The distribution and chemical composition of ultracentrifugally separated lipoproteins in human serum. J. Clin. Invest. 34:1345-1353. Helmy, S., Porter-Jordan, K., Dawidowicz, E.A., Pilch, P., Schwartz, A.L., and Fine, R.E. 1986. Separation of endocytic from exocytic coated vesicles using a novel cholinesterase-mediated density shift technique. Cell 44:497-506. Holcomb, C., Etcheverry, T., and Schekman, R. 1987. Isolation of secretory vesicles from Saccharomyces cerevisiae. Anal. Biochem. 166:328-334.
Purification of Organelles from Mammalian Cells
Huttner, W.B., Schlieber, W., Greengard, P., and DeCamilli, P. 1983. Synapsin I (Protein I), a nerve terminal-specific phosphoprotein: III. Its association with synaptic vesicles studied in a highly purified synaptic vesicle preparation. J. Cell Biol. 96:1374-1388.
Howell, K.E., Schmid, R., Ugelstad, J., and Gruenberg, J. 1989. Immunoisolation using magnetic solid supports: Subcellular fractionation for cell-free functional studies. Methods Cell Biol. 31:265-292.
Mueller, S.C. and Branton, D. 1984. Identification of coated vesicles in Saccharomyces cerevisiae. J. Cell Biol. 98:341-346. Orlowsky, M. and Meister, A. 1963. γ-glutamylp-nitroanilide: A new convenient substrate for the determination and study of L- and D-γglutamyltranspeptidase activities. Biochim. Biophys. Acta 73:679-681. Paul, E., Hurtubise, Y., and LeBel, D. 1992. Purification and characterization of the apical plasma membrane of the rat pancreatic acinar cell. J. Memb. Biol. 127:129-137. Rickwood, D. 1984. Centrifugation, a Practical Approach (2nd ed). IRL Press, Oxford. Roep, B.O., Arden, S.D., deVries, R.R.P., and Hutton, J.C. 1990. T-cell clones from a type 1 diabetes patient respond to insulin secretory granule proteins. Nature 345:632-634.
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Rubenstein, J.L.R., Fine, R.E., Luskey, B.D., and Rothman, J.E. 1981. Purification of coated vesicles by agarose gel electrophoresis. J. Cell Biol. 89:357-361.
Walworth, N.C. and Novick, P.J. 1987. Purification and characterization of constitutive secretory vesicles from yeast. J. Cell Biol. 105:163174.
Salamero, J., Sztul, E.S., and Howell, K.E. 1990. Exocytic transport vesicles generated in vitro from the trans-Golgi network carry secretory and plasma membrane proteins. Proc. Natl. Acad. Sci. U.S.A. 87:7717-7721.
Warnock, D.E., Roberts, C., Lutz, M.S., Blackburn, W.A., Young, W.W., and Baenziger, J.U. 1993. Determination of plasma membrane lipid mass and composition in cultured Chinese hamster ovary cells using high gradient magnetic affinity chromatography. J. Biol. Chem. 268:1014510153.
Saucan, L. and Palade, G.E. 1994. Membrane and secretory proteins are transported from the Golgi complex to the sinusoidal plasmalemma of hepatocytes by distinct vesicular carriers. J. Cell Biol. 125:733-741. Slot, J.W. and Geuze, H.J. 1985. A new method of preparing gold probes for multiple-labeling cytochemistry. Eur. J. Cell Biol. 38:87-93. Tulp, A., Verwoerd, D., and Pieters, J. 1993. Application of an improved density gradient electrophoresis apparatus to the separation of proteins, cells and subcellular organelles. Electrophoresis 14:1295-1301. Tulp, A., Verwoerd, D., Dobberstein, B., Ploegh, H.L., and Pieters, J. 1994. Isolation and characterization of the intracellular MHC class II compartment. Nature 369:120-126.
Wattiaux, R., Wattiaux-DeConnick, S., RonveauxDupal, M.-F., and Dubois, F. 1978. Isolation of rat liver lysosomes by isopycnic centrifugation in a metrizamide gradient. J. Cell Biol. 78:349368. Zastrow, M.V. and Castle, J.D. 1987. Protein sorting among two distinct export pathways occurs from the content of maturing exocrine storage granules. J. Cell Biol. 105:2675-2684.
Contributed by J. David Castle University of Virginia Charlottesville, Virginia
Wagner, J.A., Carlson, S.S., and Kelly, R.B. 1978. Chemical and physical characterization of cholinergic synaptic vesicles. Biochem. 17:1199-1206.
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Subcellular Fractionation of Tissue Culture Cells
UNIT 4.3
The development of cell fractionation techniques over the last few decades has provided the means to analyze the composition and properties of purified cellular elements. In particular, subcellular fractionation is essential for the development of cell-free assays that reconstitute complicated cellular processes. These assays have provided new and important tools to understand the molecular mechanisms of complex cellular functions, permitting these functions to be studied in the test tube as a series of biochemical reactions. General methods for fractionating mammalian tissues are presented in UNIT 4.2. The protocols in this unit describe fractionation of tissue culture cells to immunoisolate early and late endosomes under conditions where these compartments retain their capacity to support membrane transport in vitro. To prepare different populations of endosomes, monolayers of cells are exposed to a fluid-phase marker for endosomal compartments (horseradish peroxidase; HRP) either briefly to label early endosomes or with a chase period to label endosomal carrier vesicles or late endosomes (see Basic Protocol 1). For subsequent immunoisolation, many antibodies are available against cytoplasmically exposed epitopes present on early or late endosome proteins (Gruenberg, 2001). Antibody-antigen couples do not, however, always work efficiently in immunoisolation experiments. An attractive alternative is to use a foreign antigen, against which appropriate antibodies are already available (e.g., an epitope added to an ectopically expressed protein). This protocol was used for the immunoisolation of recycling endosomes from MDCK cells expressing a myc-tagged version of the transferrin receptor (Gagescu et al., 2000). Alternatively, vesicular stomatitis virus (VSV) can be fused with the cells to add VSV G protein to the cell membranes (see Basic Protocol 2), and the cells are then exposed to HRP. Labeled cells from either protocol are then homogenized (see Basic Protocol 3), and the postnuclear supernatant (PNS) is fractionated by a flotation gradient (see Basic Protocol 4). Specific endosomal compartments are isolated using magnetic or polyacrylamide beads coupled to an antibody specific for VSV G protein (see Basic Protocol 5). The Support Protocol describes growth conditions for preparing BHK-21 cells for isolation of endosomes. The Commentary discusses preparation of a “balance sheet” for the isolation procedure (see Background Information) and problems that may be encountered in subcellular fractionation of tissue culture cells (see Critical Parameters as well as the Commentary in UNIT 4.2). INTERNALIZATION OF FLUID-PHASE MARKER Several proteins, including Rab GTPases and receptors (Table 4.3.1), can be used to follow early or late endosomes. In the absence of appropriate reagents, markers endocytosed in the fluid phase can allow convenient discrimination between the compartments (Table 4.3.1). A word of caution is warranted, however, because solutes can efficiently be internalized by other routes in some cell types, in particular by macropinocytosis in cells with high ruffling activity. Horseradish peroxidase (HRP) is widely used as a marker for fluid-phase endocytosis. HRP internalized at a concentration of 0.5 to 10 mg/ml distributes uniformly within the lumen of endosomal compartments and is easily measured using a colorimetric reaction. Internalization is carried out directly in the tissue culture dish for cells growing attached to the substratum. For suspension cells, the method is essentially identical, except that all changes of buffers and solutions must be accomplished by centrifugation. Once labeled, the cells must be washed thoroughly to remove nonspecifically adsorbed marker. Washed cells may be incubated further in the absence of marker to label endosomal carrier vesicles and late endosomes.
BASIC PROTOCOL 1
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Contributed by Fernando Aniento and Jean Gruenberg Current Protocols in Protein Science (2003) 4.3.1-4.3.21
4.3.1
Copyright © 2003 by John Wiley & Sons, Inc.
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Table 4.3.1
Labeling Conditions for Characterization of the Different Endosomal Compartmentsa,b
Duration of incubation Endosomal compartment at 37°C
Markers
References
EE
Rab5, Rab4
Chavrier et al. (1990); Gorvel et al. (1991)
Annexin II Receptors recycling with the plasma membrane (e.g., transferrin-R)
Emans et al. (1993) Gruenberg and Howell (1989); Emans et al. (1991)
ECV
LE
5 min
5-min pulse + 45-min chase after depolymerization of microtubules 5-min pulse + 45-min chase
Gruenberg et al. (1989); Bomsel et al. (1990); Aniento et al. (1993) Rab7, Rab9
Chavrier et al. (1990); Gorvel et al. (1991)
CI-MPR lgps
Griffiths et al. (1988); Aniento et al. (1993a) Geuze et al. (1983); Griffiths et al. (1988); Gorvel et al. (1991)
aSeveral proteins listed here exhibit a restricted distribution within endosomes, but can also be found in other compartments (e.g., MPR in TGN and
late endosomes; Rab5 in plasma membrane and early endosomes). Analysis of purified fractions must thus combine different markers to identify endosomes unambiguously. bAbbreviations: CI-MPR, cation-independent mannose-6-phosphate receptor; EE, early endosomes; ECV, endosomal carrier vesicles; LE, late
endosomes; lgps, lysosomal glycoproteins; TGN, trans Golgi network; transferrin-R, transferrin receptor. The 4a1 and 2a5 monoclonal antibodies recognize two lgps that are abundant in late endosomes (Aniento et al., 1993).
Materials Monolayer cultures of BHK-21 cells in 10-cm tissue culture plates, 14 to 18 hr after plating (see Support Protocol 3) PBS (APPENDIX 2E), ice cold 2 to 10 mg/ml horseradish peroxidase (HRP) in internalization medium (see recipe), 37°C PBS/BSA: PBS containing 5 mg/ml BSA, ice cold and 37°C IM/BSA: Internalization medium (see recipe) containing 2 mg/ml BSA, 37°C Cold plate: wet metal plate lying flat on top of ice in an ice bucket Rocking platform Warm plate: 37°C water bath with a flat metal plate slightly below the water level or 37°C incubator 1. Place tissue culture dishes containing monolayer cultures of BHK-21 cells on a cold plate. Aspirate the medium and add 5 to 10 ml ice-cold PBS to each dish. Incubate 2 to 3 min on a rocking platform. Repeat this step twice to thoroughly wash the cells. Use two to three dishes of cells per time point. Keep the dishes level on the plate in the ice bucket.
2. Aspirate the PBS and prewarm each dish 1 to 2 sec on a warm plate. Add 2.5 ml prewarmed HRP to the dish and incubate 5 min on the warm plate. In most cases, HRP at a concentration of 2 mg/ml provides sufficient labeling. If necessary, higher concentrations (up to 10 mg/ml) can be used to increase the signal (for example, when fractions are further purified by immunoisolation). Subcellular Fractionation of Tissue Culture Cells
To reduce costs, small volumes of HRP (e.g., 2.5 ml) can be used for incubation at 37°C. The culture dish should be placed on a metal plate mounted on a rocking platform in a 37°C incubator to ensure that the cells are adequately covered with liquid during the incubation.
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After a 5-min incubation at 37°C, membrane proteins and fluid-phase tracers distribute predominantly within early endosomes (Gruenberg et al., 1989; Gorvel et al., 1991; Emans et al., 1993). After longer internalization times, the markers appear first in endosomal carrier vesicles and then sequentially in late endosomes and lysosomes (Bomsel et al., 1990; Aniento et al., 1993a; Lebrand et al., 2002).
3. Return dishes to the cold plate. Aspirate HRP and wash each dish three times, 5 to 10 min each, with 5 ml PBS/BSA. Aspirate PBS/BSA and wash each dish twice with 5 ml PBS. To label endosomal carrier vesicles or late endosomes, proceed to steps 4 and 5. Otherwise, proceed to homogenization (see Basic Protocol 3). 4. Optional: If the experiment requires fluid-phase labeling of endosomal carrier vesicles or late endosomes, immediately after the internalization step rinse each dish two or three times with 5 ml of 37°C PBS/BSA. Add 10 ml IM/BSA and incubate 45 min at 37°C. These pulse-chase conditions are preferred to continuous internalization of HRP because under these conditions the amount of HRP in earlier compartments is reduced.
5. Return dishes to the cold plate. Then, as in step 3 above, aspirate medium and wash each dish twice, 5 min each, with 5 ml PBS/BSA. Aspirate PBS/BSA and rinse twice with PBS. Markers can also be allowed to accumulate in the intermediate endosomal carrier vesicles by preincubating the cells 1 to 2 hr at 37°C in medium containing 10 ìM nocodazole to depolymerize microtubules. The conditions for fluid-phase labeling and marker internalization are the same, except that all wash solutions and media used, including internalization and chase media, should contain 10 ìM nocodazole. Alternatively, endosomal carrier vesicles can be labeled by incubating the cells 2 hr at 20°C without nocodazole. However, internalization rates and labeling efficiency will be higher at 37°C than at 20°C.
IMPLANTATION AND INTERNALIZATION OF VSV G PROTEIN The transmembrane glycoprotein G of vesicular stomatitis virus (VSV), or another transmembrane endosomal antigen for which there is an antibody, can be used to immunoisolate endosomal fractions (Fig. 4.3.1). Virus is first bound to the plasma membrane by wheat germ agglutinin– mediated adhesion, and then fused with the plasma membrane by a brief exposure to low-pH medium, which inhibits endocytosis (Davoust et al., 1987). The cells are then washed and incubated with HRP at 37°C. Under these conditions, the G protein, a membrane marker, is co-internalized with HRP, a fluid-phase marker. The G protein cytoplasmic domain becomes exposed on the surface of endosomal elements and is accessible to antibodies after homogenization.
BASIC PROTOCOL 2
Materials Monolayer cultures of BHK-21 cells in 10-cm tissue culture plates, 14 to 18 hr after plating (see Support Protocol 3) 100 µg/ml wheat germ agglutinin (WGA) from Triticum vulgaris (Sigma) in 7.4 M medium (see recipe), ice cold 20 µg/ml vesicular stomatitis virus (VSV; see Support Protocol 1) in 7.4 M medium (see recipe), ice cold Fusion medium: MEM containing 20 mM succinate, pH 4.9, 37°C 60 mM N-acetylglucosamine (GlcNAc) in 7.4 M medium (see recipe), ice cold Vacuum aspirator connected to a trap containing disinfectant (e.g., chloramine T) Additional reagents and equipment for fluid-phase internalization (see Basic Protocol 1) Extraction, Stabilization, and Concentration
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0 min
perform membrane labeling (Basic Protocol 2, steps 1 to 10)
internalize label (Basic Protocol 2, step 11)
45 min without nocodazole
45 min with nocodazole
5 min perform fluid-phase labeling with HRP (Basic Protocol 1; optional)
homogenize cells (Basic Protocol 3)
ECV + LE EE
ECV
fractionate the homogenate (Basic Protocol 4)
LE
immunoisolate desired endosome fraction (Basic Protocol 5)
ECV + LE EE
EE
Figure 4.3.1 Outline of the complete procedure followed for the immunoisolation of early endosomes (EE), endosomal carrier vesicles (ECV), and late endosomes (LE). The G protein of the VSV virus is implanted into the plasma membrane by low pH–mediated fusion of the virus with the plasma membrane. Then, G protein molecules are cointernalized with horseradish peroxidase (HRP) to introduce the G protein into EE, ECV, or LE, respectively. The cells are then homogenized and the postnuclear supernatant (PNS) is prepared. The PNS is fractionated on a flotation gradient. The EE fraction or the ECV/LE fractions are used as input for immunoisolation using beads coupled to an antibody against the G protein cytoplasmic domain. Depending on the internalization conditions for the G protein, ECV can be separated from LE.
CAUTION: VSV is a rodent pathogen. Consult with the local hazardous materials authority for proper handling and disposal procedures. All plastic, glassware, and solutions can be disinfected with chloramine T. Wash cells 1. Chill monolayer cultures of BHK-21 cells on a cold plate. Place the cold plate on a rocking platform. The dishes must lie perfectly flat on the plate. A rocking platform should be used for all incubations, particularly those with small volumes (e.g., <5 ml).
2. Rapidly aspirate the medium and rinse twice with 5 ml ice-cold PBS to ensure proper chilling of cells and removal of cellular debris. Bind WGA to cells 3. Aspirate the PBS and add 2.5 ml ice-cold 100 µg/ml WGA. Incubate 1 hr on the cold plate. Subcellular Fractionation of Tissue Culture Cells
4. Aspirate the medium and wash twice with 5 ml PBS.
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Bind VSV to cells 5. Aspirate the PBS and add 2.5 ml ice-cold 20 µg/ml VSV. Incubate 1 hr on the cold plate. Addition of 50 ìg VSV protein to a 10-cm dish of confluent BHK-21 cells results in the fusion of 8.5 ìg VSV protein with the plasma membrane; this corresponds to ∼80 molecules of VSV G protein/ìm2 membrane surface area (Gruenberg and Howell, 1987).
6. Aspirate virus solution and wash cells twice with 5 ml PBS. 7. Remove dish from the cold plate, aspirate wash solution, and add 20 ml of 37°C fusion medium. Incubate 30 sec at 37°C. Low pH–mediated fusion at 37 °C results in the incorporation of G protein molecules in the correct transmembrane orientation. Because all steps except the fusion are done on ice and because endocytosis is arrested at pH 4.9, G protein is restricted to the plasma membrane.
8. Aspirate fusion medium and quickly place the dish back on the cold plate. Rinse twice with 5 ml ice-cold PBS. Remove unfused VSV 9. Remove unfused virions by aspirating the PBS, and then adding 5 ml ice-cold 60 mM GlcNAc. Incubate 1 hr on the cold plate. 10. Aspirate the GlcNAc and wash twice with PBS. Internalize G protein 11. Add 10 ml IM/BSA and incubate 5 min or 45 min at 37°C. After 5 min, G protein is found predominantly in early endosomes; after 45 min, it is found primarily in late endosomes. Endosomal carrier vesicles contain the G protein after 45 min if microtubules are depolymerized by treatment with nocodazole (see Basic Protocol 1).
12. Optional: If a general marker for endosomes is required (e.g., to establish a balance sheet for fractionation), co-internalize HRP with the G protein (see Basic Protocol 1). PREPARATION OF VESICULAR STOMATITIS VIRUS STOCKS For this immunoisolation experiment, the antigen is provided by the cytoplasmic domain of the transmembrane glycoprotein G of the enveloped virus, vesicular stomatitis virus (VSV). VSV stocks are produced and harvested as described in Gruenberg et al. (1989). Briefly, subconfluent 2-day cultures of BHK cells are infected with 0.2 pfu VSV/cell and incubated 19 to 20 hr at 37°C. Medium containing virus is removed from the cells and centrifuged 30 min at 7000 × gavg in an HS4 rotor (Sorvall) to remove cell debris. The supernatant is layered on top of a two-step gradient of 10% and 55% (w/v) sucrose in 50 mM Tris⋅Cl/100 mM NaCl/0.5 mM EDTA (pH 7.4) and centrifuged 90 min at 90,000 × gavg. Virions are collected at the 10%/55% interface. Aliquots of 50 and 200 µg are frozen in liquid nitrogen and stored at −70°C.
SUPPORT PROTOCOL 1
HOMOGENIZATION OF TISSUE CULTURE CELLS Gentle homogenization conditions should be used to limit possible damage to endosomal elements, particularly when using fluid-phase markers. Clearly, the markers should remain entrapped in vesicles (latent) after homogenization. In addition, harsh conditions should always be avoided to limit the breakage of lysosomes and consequent proteolysis by released hydrolases. Cells grown for 14 to 16 hr are easily homogenized, so each step of the homogenization process should be monitored using a phase-contrast microscope. A balance sheet should be used to follow the distribution of protein and markers during homogenization, flotation, and immunoisolation (see Table 4.3.2 for an example). La-
BASIC PROTOCOL 3
Extraction, Stabilization, and Concentration
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beled cells are scraped from the dish, homogenized, and centrifuged to prepare a postnuclear supernatant (PNS). Materials Monolayer cultures of BHK-21 cells labeled with VSV G protein and/or fluid-phase marker (see Basic Protocols 1 and 2) PBS (APPENDIX 2E), 4°C Homogenization buffer (HB; see recipe) Protease inhibitor cocktail (see recipe) 15-ml plastic conical centrifuge tube Refrigerated centrifuge and rotor appropriate for cell sedimentation 22-G needle attached to 1-ml syringe Phase-contrast microscope Beckman Airfuge or equivalent ultracentrifuge Additional reagents and equipment for assay of horseradish peroxidase activity (see Support Protocol 2) Collect cells 1. Remove cells from the dish by scraping with a rubber policeman in 2.5 ml of 4°C PBS and transfer to a 15-ml plastic conical centrifuge tube. Centrifuge 5 min at 700 × g, 4°C. Typically, the contents of three to five dishes are pooled into a 15-ml tube.
2. Remove and save the supernatant. Resuspend cell pellet very gently in 5 ml HB using a wide-bore pipet. Centrifuge 10 min at 1500 × g, 4°C. 3. Analyze supernatant for the presence of markers (e.g., HRP, alkaline phosphatase) due to cell breakage during removal from the plate. Cell breakage during removal should not exceed 5% to 10%. Support Protocol 2 presents a method for detecting HRP. UNIT 10.10 describes a detection method for alkaline phosphatase, another common marker.
Table 4.3.2
Methods of Subcellular Fractionation of Endosomes
Separation method
References
Density modification DAB shift procedure
Courtoy et al. (1984); Ajioka and Kaplan (1986); Stoorgovel et al. (1987)
Colloidal gold method
Beardmore et al. (1987); Beaumelle et al. (1990)
Free-flow electrophoresis
Marsh et al. (1987); Schmid et al. (1988)
Immunoisolation
Howell et al. (1989b); Thomas et al. (1992); Aniento et al. (1993a); Emans et al. (1993)
Gradients
Subcellular Fractionation of Tissue Culture Cells
Percoll
Pertoft et al. (1978); Sahagian and Neufeld (1983); Bergeron et al. (1985)
Nycodenz
Kindberg et al. (1984); Mullock et al. (1989)
Sucrose
Khan et al. (1986); Mueller and Hubbard (1986); Gorvel et al. (1991); Aniento et al. (1993a); Emans et al. (1993)
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Homogenize cells 4. Add 0.5 ml protease inhibitor cocktail in HB to the cell pellet and resuspend using a 1000-µl pipettor with a blue (1000-µl) tip. Homogenization is easier at a relatively high cell density (e.g., 20% to 30% [v/v]). Cells that are grown in suspension or that are released from the dish by trypsin treatment may require harsher homogenization conditions. The use of a tight-fitting glass-glass Dounce homogenizer (15 to 20 passages of the pestle) or a cell cracker may be required to achieve good cell breakage. A cell cracker is a device used to pass cells through a precision clearance between the wall of a metal chamber and a metal ball bearing (Balch and Rothman, 1985).
5. Draw the homogenate three to ten times through a 22-G needle attached to a 1-ml syringe. Monitor homogenization using a phase-contrast microscope. Be careful not to over-homogenize. Nuclei should be intact and free of cellular materials. Damage to endosomes and lysosomes can be quantified by measuring the latency of content markers, lysosomal enzymes, or other integral markers. (Latency corresponds to the percentage of a marker that remains entrapped within vesicles, such as endosomes, after homogenization.) Under optimal conditions, latency should be >75% after homogenization.
6. Centrifuge homogenate 15 min at 1500 × g, 4°C and collect the postnuclear supernatant (PNS) and the nuclear pellet (NP). Resuspend NP in 1 ml PBS. Save a 50-µl aliquot of the PNS and the NP for analysis. The PNS is ready for fractionation (see Basic Protocol 4).
Determine latency of markers 7. Dilute a 20-µl aliquot of the PNS with HB to a total volume of 0.2 to 1 ml, depending on the amount of PNS necessary to detect the marker. 8. Centrifuge 15 min at 200,000 × g (e.g., 20 psi in a Beckman Airfuge), 4°C. 9. Collect the supernatant. Resuspend the pellet in the same volume of HB as used in step 7. 10 Determine the amount of HRP or other marker in the original PNS, in the pellet, and in the supernatant (see Support Protocol 2). 11. Calculate the amount of marker in the pellet as a percentage of the total amount present in the PNS (yield). DETERMINATION OF HORSERADISH PEROXIDASE ACTIVITY Internalized HRP is measured using o-dianisidine and peroxide as substrates; the browncolored product is quantified by measuring the absorbance at 455 nm with a spectrophotometer.
SUPPORT PROTOCOL 2
Materials Subcellular fraction containing internalized HRP (see Basic Protocol 3) HRP substrate solution (see recipe) HRP standards: 10 to 100 ng HRP/ml in the same buffer used to prepare or dilute sample 10 µM potassium cyanide (KCN; optional) Extraction, Stabilization, and Concentration
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1. Quickly mix 0.1 ml sample with 0.9 ml substrate solution. Mix 0.1 ml of each HRP standard with 0.9 ml substrate solution. Incubate at room temperature in the dark. Time the reaction. If necessary, sample may be diluted with PBS (APPENDIX 2E) or other buffer before analyzing a 0.1-ml aliquot.
2. When a brown color develops, note the time and measure the absorbance of sample and standards at 455 nm. With small amounts of HRP, the reaction can be developed for up to 2 hr in the dark. It is not necessary to stop the reaction, but the heme group of HRP can be blocked at the desired time by adding a respiratory inhibitor such as 10 ìM KCN.
3. Use the values obtained for HRP standards to quantify the amount of HRP in the sample. BASIC PROTOCOL 4
FLOTATION-GRADIENT FRACTIONATION OF TISSUE CULTURE HOMOGENATES Early and late endosomal fractions can easily be prepared from postnuclear supernatants (PNS) by flotation in a step gradient of 40.6% to 8.6% (w/w) sucrose/D2O. Early endosomes containing the transferrin receptor, the small GTPase Rab5 protein, and annexin II can be separated from late endosomes containing the Rab7 protein, lysosomal glycoproteins (lgps), and the cation-independent mannose-6-phosphate receptor (Table 4.3.1). Fractions from this gradient can be used to determine the general protein composition of the fractions in high-resolution two-dimensional gels (UNIT 10.4) and to reconstitute the different endosomal membrane fusion events (Fig. 4.3.2). Materials Postnuclear supernatant (PNS; see Basic Protocol 3) Sucrose solutions (see recipes) Homogenization buffer (HB; see recipe) Refractometer SW60 centrifuge tubes Ultracentrifuge and SW60 rotor Peristaltic pump connected to 50-µl capillary tube 1. Dilute a PNS sample 1:1 (v/v) with 62% sucrose solution in H2O to give a solution of 40.6% sucrose/0.5 mM EDTA and check the final sucrose concentration with a refractometer.
1
EE
EE
2 ECV
ECV
3 Subcellular Fractionation of Tissue Culture Cells
LE
4
LE
Figure 4.3.2 Outline of the different endosome fusion events reconstituted in an in vitro fusion assay. Early endosomes (EE) undergo homotypic fusion with each other (1; Gruenberg and Howell, 1989). Several components regulating this process have been identified (Gruenberg and Clague, 1992). Endosomal carrier vesicles (ECV) form from EE (2) in a process that requires vacuolar ATPase activity (Clague et al., 1994). ECV fuse with late endosomes (LE); (3) in a process facilitated by microtubules and cytoplasmic dynein (Aniento et al., 1993a). LE also undergo homotypic fusion with each other (4). All other combinations (EE-ECV, ECV-ECV, and EE-LE) are refractory to fusion (Aniento et al., 1993a).
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2. Load 1 ml diluted PNS in the bottom of an SW60 centrifuge tube and overlay sequentially with 1.5 ml of 16% sucrose solution in D2O, then 1 ml of 10% sucrose solution in D2O. Add HB to fill the tube. Early and late endosomes can also be isolated on a gradient with 35% and 25% sucrose solutions in H2O. Also, the low-density sucrose step (10% in D2O or 25% in H2O) can be omitted to yield fractions containing mixed populations of early and late endosomes.
3. Mount the tubes in an SW60 rotor and centrifuge 1 hr at 14,000 × gavg (35,000 rpm), 4°C. 4. In a 4°C cold room, use a peristaltic pump connected to a 50-µl capillary tube to collect the gradient. The early endosome fraction containing the transferrin receptor, the small GTPase Rab5, and annexin II is collected from the 16%/10% interface (or 35%/25% interface). The late endosome fraction containing the Rab7 protein, lysosomal glycoproteins (lgps), and the cation-independent mannose-6-phosphate receptor (CI-MPR) is collected from the upper portion of the 10% cushion (or 25% cushion in H2O).
5. Divide early and late endosome fractions into 50-µl aliquots, and then freeze and store them in liquid nitrogen. Aliquots should be placed in a liquid nitrogen bath for rapid freezing; similarly, they should be thawed as rapidly as possible prior to use or analysis.
IMMUNOISOLATION OF ENDOSOMAL FRACTIONS Immunoisolation can be used to purify any subcellular fraction as long as a specific antibody is available. The specific antibody is coupled to magnetic or polyacrylamide beads with a covalently coupled species-specific linker antibody, and then the beads are used to precipitate the desired organelle. Since this protocol was introduced (Howell et al., 1989a), a similar strategy has been used to immunoisolate a long list of organelles and vesicles from different cell types with different antigen-antibody couples. The procedure for immunoisolating endosomes using VSV G protein as the antigen is described here.
BASIC PROTOCOL 5
Materials Rabbit anti-mouse immunoglobulin covalently coupled to polyacrylamide or magnetic beads (e.g., Immunobeads; Bio-Rad) PBS/BSA: PBS (APPENDIX 2E) containing 2 mg/ml BSA, ice cold Mouse antibody against the cytoplasmic domain of VSV G protein (P5D4; Kreis, 1986) Endosomal fractions (see Basic Protocol 4) Microcentrifuge, refrigerated Low-speed rotating wheel (2 to 20 rpm) 1. Resuspend 1 to 10 mg rabbit anti-mouse immunoglobulin covalently coupled to polyacrylamide or magnetic beads in 1 ml PBS/BSA and microcentrifuge 2 to 3 min at 3000 × g, 4°C. Immunobeads are also available with antibodies to immunoglobulins from other species.
2. Wash the beads four times with PBS/BSA by centrifugation as in step 1. 3. Add 5 to 10 µg mouse anti– G protein antibody/mg bead in a total volume of 1 ml PBS/BSA for 1 to 10 mg of beads. Rotate end over end overnight at 4°C.
Extraction, Stabilization, and Concentration
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4. Remove excess antibody by washing three times with PBS/BSA. Resuspend beads at 10 mg/ml in PBS/BSA. 5. Dilute the endosomal fraction 1:3 with ice-cold PBS/BSA in a 1.5-ml microcentrifuge tube to a final volume of ∼1 ml. 6. Add 1 mg beads with bound antibodies to the cellular fraction. Rotate end over end 2 hr at 4°C. For either polyacrylamide or magnetic beads, 1 mg solid support (a pellet of ∼20 ìl) is used for immunoisolation from 300 to 500 ìg PNS protein or 50 ìg protein of a fraction collected from the gradient (the equivalent of cells from 10 to 20 cm2 of tissue culture surface area).
7. Centrifuge the beads and fraction mixture 5 min at 3000 × g in a refrigerated centrifuge. Collect supernatant for analysis. Gently resuspend pellet in 1 ml PBS/BSA to prevent vesicularization and repeat centrifugation. If the sample is to be analyzed by gel electrophoresis, the last wash should be in PBS without BSA. The amount of immunoisolated protein cannot be quantified directly because of the added BSA and antibodies. It is possible, however, to obtain an estimate by using cells metabolically labeled with [35S]methionine.
8. Resuspend the beads plus bound vesicles in 100 µl medium or buffer. The choice of medium or buffer depends on how the sample will be analyzed; for example, use homogenization buffer (see recipe) for a cell-free assay of endosome fusion (Gorvel et al., 1991; Aniento et al., 1993a) or lysis buffer for two-dimensional gel electrophoresis (UNIT 10.4). SUPPORT PROTOCOL 3
PREPARATION OF TISSUE CULTURE CELLS FOR SUBCELLULAR FRACTIONATION Cells from a baby hamster kidney line (BHK-21) are grown as monolayers in G-MEM supplemented with 5% (v/v) fetal calf serum (FCS), 10% (v/v) tryptose phosphate broth, and 2 mM glutamine in a humidified 5% CO2, 37°C incubator. The cells are seeded at 4 × 104 cells/cm2 in 10-cm tissue culture dishes (∼3.2 × 106 cells per dish) to form an even, confluent monolayer by 14 to 16 hr after seeding. These conditions yield ∼1.3 × 107 cells and ∼2.5 to 3.0 mg total protein per dish at the time of the experiment. This protocol has the additional advantage that cells from 14- to 16-hr cultures can easily be homogenized. Typically, two to three 10-cm dishes are used per experimental condition, and fifteen to twenty 10-cm dishes are used for preparing stocks. To isolate endosomal carrier vesicles, microtubules can be depolymerized prior to labeling and homogenization to inhibit transport of the marker from early endosomes to late endosomes and lysosomes. The conditions required to depolymerize microtubules may, however, vary between cell types. In BHK-21 cells, preincubating the cells 1 to 2 hr at 37°C in the presence of 10 µM nocodazole is sufficient (Gruenberg et al., 1989). No apparent depolymerization of BHK microtubules is observed after prolonged incubation at 4°C. In contrast, the depolymerization of MDCK microtubules requires both a cold treatment and an incubation at 37°C in the presence of nocodazole (Bomsel et al., 1990).
Subcellular Fractionation of Tissue Culture Cells
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REAGENTS AND SOLUTIONS 7.4 M medium MEM containing: 10 mM TES [N-tris(hydroxymethyl)methyl-2-aminoethanesulfonic acid] 10 mM MOPS [3-(N-morpholino)propanesulfonic acid] 15 mM HEPES (N-2-hydroxyethylpiperzazine-N′-2-ethanesulfonic acid) 2 mM NaH2PO4 0.42 mM NaHCO3 Adjust pH to 7.4 Store sterile medium 4 to 8 weeks at 4°C Homogenization buffer (HB) 250 mM sucrose 3 mM imidazole, pH 7.4 10 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) or 10 mM HEPES (N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid) Store sterile buffer 4 to 8 weeks at 4°C HRP substrate solution 0.05 M sodium phosphate buffer, pH 5.0 (APPENDIX 2E) 0.3% (v/v) Triton X-100 0.342 mM o-dianisidine 0.003% (v/v) H2O2 Store at 4°C protected from light Prepare this solution in very clean glassware to avoid contamination. Add o-dianisidine first; after it has been solubilized, add H2O2. Discard if a straw yellow color develops.
Internalization medium MEM containing: 5 mM D-glucose 10 mM HEPES (N-2-hydroxyethanepiperazine-N′-2-ethanesulfornic acid) Adjust pH to 7.4 Store sterile medium 4 to 8 weeks at 4°C Protease inhibitor cocktail Stock solution 10 mM leupeptin 10 mg/ml aprotinin 1 mM pepstatin Store several months at −20°C Working solution: Dilute stock solution 1:1000 in HB (see recipe). Sucrose solutions 10% (w/w) sucrose solution in D2O 0.303 M sucrose 3 mM imidazole, pH 7.4 Store sterile solution 4 to 8 weeks at 4°C The refractive index of this solution should be 1.3478 at 20°C.
16% (w/w) sucrose solution in D2O 0.497 M sucrose 3 mM imidazole, pH 7.4 Store sterile solution 4 to 8 weeks at 4°C Extraction, Stabilization, and Concentration
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The refractive index of this solution should be 1.3573 at 20°C.
25% (w/w) sucrose solution in H2O 0.806 M sucrose 3 mM imidazole, pH 7.4 Store sterile solution 4 to 8 weeks at 4°C The refractive index of this solution should be 1.3723 at 20°C.
35% (w/w) sucrose solution in H2O 1.177 M sucrose 3 mM imidazole, pH 7.4 Store sterile solution 4 to 8 weeks at 4°C The refractive index of this solution should be 1.3904 at 20°C.
62% (w/w) sucrose solution in H2O 2.351 M sucrose 3 mM imidazole, pH 7.4 1 mM EDTA Store sterile solution 4 to 8 weeks at 4°C The refractive index of this solution should be 1.4463 at 20°C.
COMMENTARY Background Information
Subcellular Fractionation of Tissue Culture Cells
Subcellular fractionation was initially developed with rat liver and, in most cases, involves separation of organelles based on physical properties such as density. However, two major problems have interfered with the widespread use of subcellular fractionation in cell and molecular biology. First, studies of membrane transport and cellular organization have revealed that several subcellular compartments share similar physical properties and hence cofractionate, at least to some extent, in conventional gradients (e.g., Golgi and endosomes). In addition, the use of tissue culture cells is now widespread because cultures of cells can be manipulated in a manner impossible to achieve in animals or separate tissues. Tissue culture cells are, however, more difficult to fractionate than most tissues, presumably because of differences in organization of the cytoskeleton (Howell et al., 1989a). Subcellular fractionation consists of two major steps: (1) disruption of the cellular organization (homogenization) and (2) fractionation of the homogenate to separate the different organelles. Since the early attempts to quantitatively fractionate liver tissue (Claude, 1946), methods have been developed to reproducibly disaggregate the tissue into a suspension of subcellular components and organelles called tissue homogenate (UNIT 4.2; DeDuve et al., 1955, DeDuve, 1971; Fleischer and Kervina, 1974; Beaufay and Amar-Cortesec, 1976). This
homogenate can then be resolved by differential centrifugation into several fractions: (1) nuclei, heavy mitochondria, and plasma membrane; (2) light mitochondria, lysosomes, and peroxisomes; (3) Golgi, endosomes, and microsomes; and (4) cytosol. From these main fractions, organelles of interest can be further separated according to their size and density by additional purification steps. It is essential to point out that achievement of a complete purification is a barely accessible goal. Each type of organelle is composed of populations of elements that vary in size, density, and other properties on which separation relies. The complexity of tissues and cells, together with the structural alterations induced at all stages of fractionation, increases the physical heterogeneity. Additionally, the relative abundance of the organelle of interest in the cell will affect the yield in fractionation. Clearly, it is easier to purify major subcellular components (e.g., endoplasmic reticulum and mitochondria) than, for instance, transport vesicles, which are minor cellular elements. In the latter case, a low yield is often the price to be paid to achieve purification. Subcellular fractionation Traditional fractionation procedures are based on differences in the size and density of organelles or organelle-derived vesicles as measured in density gradients. Although sucrose is most widely used to form gradients,
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there are many other alternatives, such as metrizamide, Ficoll (Amersham Biosciences), Percoll (Amersham Biosciences), Nycodenz (Progen Biotechnik), or Optiprep (Progen Biotechnik). However, many of the smooth-membrane vesicles derived from the compartments involved in membrane traffic share common densities and are difficult to isolate by these procedures, so alternative methods have been developed. Table 4.3.2 lists a number of procedures that have been used to isolate endosomal fractions (UNIT 4.2). Gradients are extremely useful to separate populations of endosomes and lysosomes. Gradients are often best combined, however, with one of the techniques described below to achieve higher levels of purification. This unit describes the use of a flotation gradient combined with a method to immunologically isolate endosomal fractions using the vesicular stomatitis virus (VSV) G protein. Density-shift methods rely on altering the density of a specific compartment by the introduction of a perturbant. These methods have proven extremely useful for isolation of fractions from the endocytic or lysosomal pathway because of the ease with which density perturbants can be loaded into these compartments from the extracellular space. This experimental approach was pioneered by the DeDuve group, who used the detergent Triton WR-1339 to isolate lysosomes from rat liver (Leighton et al., 1968). Ligands coupled to horseradish peroxidase (HRP; Courtoy et al., 1984; Ajioka and Kaplan, 1986; Stoorgovel et al., 1987) or gold (Beardmore et al., 1987) have also been used to isolate endosomes. Golgi-derived clathrincoated vesicles can be separated using a newly synthesized cholinesterase activity present in the vesicles (Fishman and Fine, 1987). Free-flow electrophoresis separates subcellular compartments on the basis of charge and was first used to purify lysosomes from rat liver (Stahn et al., 1970). This method has been particularly useful for the separation of endosomes and lysosomes, which are deflected towards the anode in the electric field. A twostep procedure in which endosomes and lysosomes are first separated from all other organelles by free-flow electrophoresis and then from each other by Percoll-gradient centrifugation has been carried out using both CHO and BHK-21 cells (Marsh et al., 1987). This work has been extended to separate two distinct populations corresponding to early and late endosomes (Schmid et al., 1988). Immunoisolation relies on the unique localization of an antigen exposed on the outer
surface of the compartment of interest, rather than on a physical property of the organelle (Howell et al., 1989b). Isolation is carried out using a specific antibody bound to a solid support (e.g., magnetic beads, cellulose fibers, acrylamide beads, or Staphylococcus aureus cells). The use of immunoisolation for separating different populations of endosomes is described in detail in Basic Protocol 5 and illustrated in Figure 4.3.1. This technique has been used to isolate endosomes using the cytoplasmic domain of the G protein as antigen (Howell et al., 1989a,b). Different solid supports are available: heterodisperse polyacrylamide beads (Bio-Rad), Staphylococcus aureus expressing protein A (Pansorbin, Calbiochem), monodisperse magnetic beads (Dynal; Howell et al., 1989b), Eupergit particles (Rohm GmbH), and cellulose fibers (Richardson and Luzio, 1986; Luzio et al., 1988). The efficiency of immunoisolation for one particular antigen-antibody pair may vary with different immunoadsorbents depending on the abundance and/or accessibility of the antigen on the membrane of interest. Clearly, the actual immunoisolation experiment provides the only appropriate trial for selecting an immunoadsorbent. In the method described in this unit, the G protein of VSV is present at an estimated density of 80 G molecules/µm2 membrane surface area (Gruenberg and Howell, 1987). The cytoplasmic domain of the G protein is relatively short (consisting of the 22 carboxyl-terminal amino acids) and the antibody used was raised against a synthetic carboxyl-terminal peptide (Kreis, 1986). To reconstitute fusion in the cell-free assay, magnetic beads (Dynal) with a coupled linker antibody raised in sheep against the Fc domain of mouse IgG (Howell et al., 1989b; Gagescu et al., 2000) or anti-mouse polyacrylamide beads (Bio-Rad) were used. In this system, >70% of fluid-phase markers internalized for 5 to 45 min could be immunoisolated by adding ≤300 µg of postnuclear supernatant (PNS) protein to the beads; enrichment was 10- to 15-fold (Gruenberg and Howell, 1987) based on the ratio of markers immunoisolated on specific beads to markers bound nonspecifically to control beads. The immunoisolated fractions immobilized on solid supports retain their fusion activity (Gruenberg and Howell, 1986, 1987), with only a minimal decrease in latency of fluid-phase markers (Gruenberg et al., 1989). A significant advantage is that the bound fraction can be introduced into and then retrieved from a reaction mixture at the end of the reaction while
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bound to the solid support (Gruenberg and Howell, 1986, 1987, 1989; Gruenberg et al., 1989). Retrieval is simple, and separation can be repeated sequentially with the same fraction through a series of experimental conditions. Separation is easily achieved with magnetic beads because they can be sedimented within minutes using a small magnet. A magnet should, however, only be used to retrieve the beads after the immunoisolation step itself to limit magnetic aggregation of the beads. By combining a flotation gradient with immunoisolation, it has been possible to purify early endosomes, endosomal carrier vesicles, and late endosomes (Fig. 4.3.1). In the first case, the G protein was internalized for 5 min so it distributed in early endosomes. Then cells were homogenized and the PNS was fractionated on a flotation gradient. The early endosome band (interface between 16% and 10% sucrose in D2O) was then used as input for immunoisolation. To isolate endosomal carrier vesicles (ECV) and late endosomes, the G protein was internalized for 45 min, either in the absence of microtubules to introduce the G protein into ECV or in the presence of microtubules to introduce the G protein into late endosomes (Table 4.3.1; Fig. 4.3.1). After homogenization, the PNS was fractionated on the flotation gradient. In this case, the ECV and late endosome band (interface between 10% sucrose in D2O and homogenization buffer) was used as input for immunoisolation. Thus, depending on the internalization conditions, either ECV or late endosomes can be immunoisolated. Analysis of subcellular fractions The functional specialization of subcellular organelles is reflected by their morphology, protein and lipid composition, and enzymatic activities, all of which can be used to characterize and analyze purified cell fractions.
Subcellular Fractionation of Tissue Culture Cells
Markers Traditionally, enzymes that are localized, ideally in one cell organelle (e.g., sugar transferases in the Golgi), are used as marker enzymes for the corresponding compartments. Purification of an organelle can then be quantified by following the activity of the marker enzyme, if known, during the fractionation procedure (see example in Table 4.3.2). More importantly, many proteins that are preferentially found in a given organelle have been characterized, and they provide useful markers during fractionation.
Biochemical and morphological characterization can easily be achieved using specific antibodies against proteins that localize to specific subcellular compartments. If antibodies are not available, such proteins can be tagged with an epitope and expressed ectopically. Green fluorescent fusion proteins are now routinely used to study the proteins of interest by light microscopy (Lippincott-Schwartz et al., 1997)—studies that can easily be combined with subcellular fractionation using antibodies against the green fluorescent protein. Finally, the large family of small GTPases, each of which has a restricted distribution, is an essential tool for monitoring the composition of subcellular fractions (Zerial and McBride, 2001). Exogenous markers can be endocytosed and thus serve as markers for endocytic organelles. The markers can be in the fluid phase or added to the plasma membrane. Protein profiles Identification and characterization of proteins present in organelles is becoming a much easier task than in the past, in particular because of major progress in protein analysis by mass spectrometry, combined with the fact that several genomes have been sequenced—and that a first draft of the human genome has been completed (Bock et al., 2001; Mann et al., 2001). Moreover, the proteome of several organelles has already been—or is being—determined (Rout et al., 2000; Wu et al., 2000; Garin et al., 2001). Morphological analysis Progress in light microscopy, combined with the use of green fluorescent fusion proteins has tremendously improved the analysis of protein distribution not only in fixed cells but also in living cells (Lippincott-Schwartz et al., 1997). Such in vivo analyses provide fundamental information on organelle dynamics and protein trafficking. Electron microscopy is, however, the method of choice for ultrastructural studies and the detailed analysis of protein distribution. Organelles retain many of their structural features after isolation, and thus electron microscopy is also an important method for analyzing fractions. Because many compartments can have the same appearance—tubular or vesicular—electron microscopy is best combined with immunocytochemistry or immunogold labeling of frozen sections.
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Balance sheet Typically, the distribution of the desired component, for example a new protein or the mutagenized form of a known protein, is compared with the distribution of known markers. Analysis of the purified fraction and assessment of the fractionation procedures require a complete balance sheet of the values measured for the different markers that are being used, including a general marker (e.g., a protein), specific markers, and contaminants, if any. These values must be expressed so as to indicate the yield and enrichment of specific markers, as well as contaminants. The following parameters must be calculated for the different steps of the fractionation. The specific activity measures the amount of the desired component in the fraction normalized to the total amount of protein. Traditionally, the specific activity refers to the activity of a marker enzyme per amount of total protein, and is measured in units per milligram protein. Clearly, the amounts of the desired component can also be quantified by other techniques, for example by immunoprecipitation with specific antibodies followed by SDSPAGE, autoradiography, and scanning of the autoradiogram. The yield, the activity of the marker enzyme in the fraction per total activity of the marker enzyme or protein in the starting material (i.e., homogenate), which is expressed as a percentage, is a measure of the amount of the material recovered in that fraction. The relative specific activity (RSA) is the absolute specific activity of the fraction per absolute specific activity of the starting material. This value provides an adequate estimate for the degree of purification achieved in each fraction. Table 4.3.3
An example illustrating the use of these parameters after fractionation of rat liver lysosomes is presented in Table 4.3.3. Fractionation is followed using a spectrophotometric assay to quantify the enzymatic activity of a lysosomal enzyme, β-N-acetyl-D-glucosaminidase, as described (Aniento et al., 1993b). Livers excised from rats are homogenized, and then the homogenate is centrifuged 10 min at 3000 × g, 4°C, to prepare a nuclear and mitochondrial pellet (NM fraction) and a postmitochondrial supernatant (PMS fraction). Fraction L corresponds to the pellet obtained after two sequential centrifugations of the PMS for 10 min at 25,000 × g, 4°C. The pellet is then resuspended in homogenization buffer and fractionated in a metrizamide gradient (Wattiaux et al., 1978; Aniento et al., 1993b). At the end of the centrifugation, fractions are collected and analyzed. Fractions 1 to 3 are pooled together and contain mainly lysosomes. Fractions 4 to 5 contain mainly mitochondria but also some lysosomes. Functional characterization The use of in vitro assays has become an essential tool to study complex cellular processes in the test tube, where they can easily be manipulated and analyzed. Most steps of membrane traffic have now been reconstituted in vitro, leading to the identification of the key components regulating membrane transport (Rothman, 1994; Chen and Scheller, 2001; Zerial and McBride, 2001). Using an in vitro assay to monitor endosome fusion, early endosomes have been observed to exhibit a high homotypic fusion activity (Gorvel et al., 1991; Aniento et al., 1993a). Endosome fusion assays have led to the characterization of proteins that regulate early endosome dynamics, including
Balance Sheet for Isolation of Rat Liver Lysosomesa
β-N-acetyl-D-glucosaminidase Protein (g)
Total act. (units)
Spec. act. (U/mg)
Yield (%)
5.000 2.175 2.825 0.110 0.011 0.084
65.0 34.4 30.6 10.7 6.9 3.5
0.013 0.016 0.011 0.097 0.623 0.042
100.0 52.9 47.1 16.5 10.6 5.4
Homogenate NM fraction PMS fraction L fraction Fractions 1-3 Fractions 4-5
RSA 1.0 1.2 0.8 7.5 143.9 8.1
aAbbreviations: β-NAGase, β-N-acetylglucosaminidase; total act., β-NAGase total activity; Spec. act., β-NAGase specific activity; RSA, β-NAGase relative specific activity.
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Subcellular Fractionation of Tissue Culture Cells
the small GTPase Rab5 and its numerous effectors (Gorvel et al., 1991; Zerial and McBride, 2001). In addition to functionally different domains that are involved in protein recycling and degradation, early endosomes also exhibit a complex ultrastructural architecture. It is now becoming clear that this mosaic of morphological and functional domains reflects differences in the distribution of protein complexes associated with specific lipids and protein-lipid domains within membranes (Gruenberg, 2001; Zerial and McBride, 2001). Early endosomes do not, however, undergo direct fusion with late endosomes in vitro (Fig. 4.3.2). Intermediate endosomal carrier vesicles, which exhibit a characteristic multivesicular appearance, mediate transport from early to late endosomes in vivo and undergo fusion with late endosomes in vitro. Recent studies highlight the role of phosphoinositol-3phosphate (PI3P) signaling and the endosomal sorting complex required for transport (ESCRT) protein complexes in sorting a subset of proteins destined for degradation within the internal membranes of these multivesicular endosomes (Gruenberg, 2001; Conibear, 2002). The mechanisms that regulate the biogenesis of these vesicles are still unclear. Candidate regulators include proteins that interact with PI3P, components of the ESCRT complex, as well as an endosomal coat protein (COP) complex and the small GTPase ARF1 (Gruenberg, 2001). These (multivesicular) carrier vesicles do not exhibit homotypic fusion properties and do not undergo fusion with early endosomes, as expected. The interactions of carrier vesicles with late endosomes is facilitated by the presence of polymerized microtubules and motor proteins (Bomsel et al., 1990; Aniento et al., 1993a), whereas docking and fusion are presumably regulated by molecules that belong to the conventional docking and fusion machinery (Chen and Scheller, 2001; Zerial and McBride, 2001). Late endosomes also exhibit a high homotypic fusion activity in vitro (Aniento et al., 1993a) and form highly dynamic networks in vivo (Lebrand et al., 2002) that contain distinct morphologically visible regions (Gruenberg, 2001) and membrane domains selectively enriched in Rab7 or Rab9 (Barbero et al., 2002). In addition, major lipid remodeling occurs in late endosomes. Their multivesicular or multilamellar elements accumulate large amounts of the poorly degradable phospholipid lysobisphosphatidic acid (LBPA), which is involved in protein and lipid transport through this compartment (Kobayashi et al., 1998,
1999). This, together with the finding that different types of membrane can be isolated from late endosomes (Fivaz et al., 2002; Kobayashi et al., 2002), supports the notion that late endosomes, much like early endosomes, are formed from a mosaic of regions and membrane domains that differ both structurally and functionally.
Critical Parameters The major impediment to successful fractionation of tissue culture cells is the difficulty encountered when trying to produce an ideal homogenate, that is, the release of various subcellular components (i.e., organelles) as a free suspension of intact, individual elements. Often, it is necessary to compromise between complete homogenization and damage to cellular compartments. After homogenization, cytoplasmic aggregates that contain cytoskeletal elements as well as various organelles can often be observed. To some extent, these aggregates can form after homogenization when conditions are too extreme because of the presence, for instance, of DNA released by breakage of the nucleus. Aggregates can, however, also reflect some cellular organization, particularly due to the cytoskeleton, that remains after homogenization. Some organelles also remain associated with cytoskeletal elements surrounding the nucleus and become entrapped in “clumps” of cytoplasm that readily sediment. As a result, as much as 50% of the components of the homogenate may be pelleted along with the nucleus during the initial centrifugation step. Because the cytoplasmic organization of different tissue culture cell lines varies enormously, homogenization conditions must frequently be optimized for each cell line. Homogenization of tissue culture cells Under ideal homogenization conditions, particulate organelles such as nuclei, mitochondria, lysosomes, and peroxisomes remain intact. Golgi complexes, plasma membranes, and reticular organelles will fragment and can, at least to some extent, vesiculate. These fragments will, however, retain many of the in vivo activities of the corresponding organelles (e.g., protein translocation across the endoplasmic reticulum membrane). Following homogenization, nuclei are totally and exclusively removed by centrifuging 10 min at 500 to 1000 × g to form a nuclear pellet (NP). The PNS contains cytosol and other organelles in free suspension. It is not possible to outline a general protocol suitable for the production of a reasonable ho-
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mogenate for all tissue culture cells, because different conditions are required for different cell types and experimental purposes. However, four parameters influence the efficiency of homogenization of cultured cells and may provide general guidelines. Growth and experimental conditions of the cells To achieve comparable homogenization over extended periods of time, it is essential to use cells that are grown from the same original parent culture (frozen in most cases immediately after cloning). The growth conditions of the cells must also be standardized and optimized for homogenization to remain reproducible. Cells that have been growing in a confluent state for several days are often more difficult to homogenize, possibly because they have a more complex network of cytoplasmic filaments (Kreis and Vale, 1993). In many cases, better homogenization is achieved with cells that are 12 to 36 hr postpassage, depending upon the density at which cells are seeded. Finally, the experimental conditions to which cells are subjected (e.g., prolonged cold treatment) may also influence homogenization conditions. Cells growing in suspension (or cells normally grown in monolayer but brought into suspension, such as with trypsin) can require significantly harsher homogenization conditions than monolayers of cells that are mechanically scraped from the dish. Homogenization medium Upon homogenization, the cell components are unavoidably transferred from a physiological environment to an entirely different medium. The homogenization medium should fulfill a number of basic requirements. It should protect the organelles against osmotic burst and possible damage due to proteases, preserve their enzymatic activities and biochemical functions, prevent any kind of agglutination, minimize leakage of constituents from organelles, and interfere as little as possible with biochemical analysis and functional assays. It is obvious that it is not always possible to fulfill all of these requirements, and the most appropriate medium will largely depend on the purpose of the experiment. To maintain the structural integrity of particulate organelles, isotonic (250 to 320 mosM) conditions at neutral pH are preferable. These conditions are also essential to retain the content within the lumen of vesicles derived from the organelles of the secretory and endocytic
pathways. Some buffers appear to be more useful for the production of a monodisperse homogenate in isotonic sucrose solutions. Among these are imidazole (Beaufay and Am ar- Cortesec, 1976) and triethanolamine/acetic acid (the latter was first introduced for and routinely used in free-flow electrophoresis experiments; Stahn et al., 1970). Some cells are very difficult to break in isotonic buffers. The forces required also disrupt the nuclear membrane releasing DNA, which causes the cytoplasm to clump and prevents further separation. In these cases, hypotonic conditions (e.g., 3 mM imidazole or 10 mM Tris⋅Cl) may become necessary to achieve reasonable homogenization, but isotonic conditions should be restored as soon as possible. It may be important to include a cocktail of proteolytic inhibitors in the homogenization medium; the choice of these is largely empirical (Gruenberg and Howell, 1986; Howell et al., 1989a; Gruenberg and Gorvel, 1992). The use of disruptive agents to depolymerize filaments of the cytoskeletal network—microtubules, microfilaments, and intermediate filaments (Kreis and Vale, 1993)—may improve the quality of the homogenate obtained from tissue culture cells. Many interactions between the cytoskeletal network and organelles are functionally significant in vivo, but must be disrupted to allow separation of individual vesicles in the homogenate. Depolymerization of the microtubules (e.g., with nocodazole) and actin filaments (e.g., with cytochalasin) does not, however, result in significantly better homogenization in many cases, presumably because of the presence of intermediate filaments, which are extremely difficult to depolymerize. The addition of salts may or may not be recommended. High salt conditions (≤1 M KCl) have been used to reduce vesicular aggregation and to prevent redistribution of luminal proteins released during homogenization of guinea pig pancreas (Scheele et al., 1978). In spleen homogenates, agglutination is avoided by the addition of 0.2 M KCl (Bowers et al., 1967). Salts can, however, promote aggregation in rat liver and FAO cells (E. Devaney and K.E. Howell, unpub. observ.). High salt has also been reported to promote the condensation of intermediate filaments, a condition that is likely to increase aggregation of organelles (Steinert et al., 1982). Finally, and most importantly, high salt concentrations cause the release of peripheral membrane proteins, which can be an
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advantage or a disadvantage, depending on the goal of the experiment. Homogenization devices When the conditions of the cells and the homogenization medium are optimized, equally efficient homogenization can be obtained by many methods: by glass-glass (Dounce) or glass-Teflon (Potter-Elvehjem) homogenization vessels and pestles; by repeated pipetting with a pipet or the plastic tip of an automatic pipettor (Harms et al., 1980); or by passage through a small-gauge needle with a syringe (Gruenberg et al., 1989). According to the authors’ experience, cells grown in suspension are better homogenized in a cell cracker, a device used to homogenize cells by passage through a precision clearance (5 to 20 µm ± 1 µm) between the wall of a metal chamber and a metal ball bearing (Balch and Rothman, 1985). The choice of homogenization device often depends on the volume of the original sample. Evaluation of the homogenate The results of homogenization should be assessed by morphological and biochemical means. Using a phase-contrast microscope, it is possible to assess the extent of cellular disruption, the appearance of intact and clean nuclei (a simple method to evaluate a good homogenization), and the absence of cytoplasmic clumps, which often appear upon overhomogenization when the nuclei start to break and the DNA is released into the cytoplasm. The quality of the homogenate is mainly revealed by its biochemical properties. Traditionally, a marker for each of the major organelles is assayed to determine its distribution in the NP and PNS (Table 4.3.1). In addition, the structural integrity of the organelles may be monitored by measuring the structure-linked latency of marker proteins or enzymes. With endocytic compartments, the latency of a marker endocytosed in the fluid phase can easily be quantified.
Anticipated Results
Subcellular Fractionation of Tissue Culture Cells
After subcellular fractionation, the purified cellular components are characterized according to several criteria, including the composition of the fraction with respect to known markers, general protein pattern, and morphological appearance. Early endosomes, endosomal carrier vesicles, and late endosomes from BHK cells have different characteristics as summarized in Table 4.3.1.
Early endosomes can be labeled with a fluidphase marker (i.e., horseradish peroxidase; HRP), which is internalized from the medium during a 5-min incubation at 37°C. A number of early endosomal markers can be used, including the small GTPase Rab5 and its effector EEA1 (Gruenberg, 2001; Zerial and McBride, 2001). Endosomal carrier vesicles can be labeled after microtubule depolymerization using HRP internalized for 5 min and then chased for 45 min at 37°C (Aniento et al., 1993a). Late endosomes are labeled with HRP internalized under the same conditions in the presence of polymerized microtubules. In addition, several proteins can be used to identify late endosomes, including the small GTPases Rab7 and Rab9, LAMP1, or the late endosomal lipid LBPA (Gruenberg, 2001; Zerial and McBride, 2001). Although the proteome of endosomes is not yet available, the three endosomal compartments have been characterized with respect to their general protein composition by analyzing the immunoisolated fractions in high-resolution bidimensional gels (Aniento et al., 1993a; Emans et al., 1993). Early endosomes contain 50 major proteins, many of which cannot be detected at later stages of the pathway (e.g., proteins recycling with the plasma membrane). Late endosomes also exhibit a complex composition with ∼50 major proteins, including unique LAMP1 and other so-called lysosomal glycoproteins. In contrast, ECV do not appear to contain unique proteins; >90% of the proteins they contain are also detected in late endosomes, consistent with the view that ECV are transient elements of the pathway and function as carrier vesicles.
Time Considerations Cells for fractionation have to be seeded 14 to 16 hr before the experiment. Fluid-phase internalization takes 60 to 90 min depending on the internalization conditions (Table 4.3.1), including washes before and after the internalization step. When microtubules have to be depolymerized, an additional 60 to 90 min are required. For endosomes that will be immunoisolated using the VSV G protein, ∼3 to 4 hr are required for viral fusion before internalization and an overnight incubation to bind antibodies to the beads. Once the markers have been internalized (if required), cells have to be scraped from the dishes and homogenized. It takes ∼1 hr to obtain the PNS. The flotation gradient is centrifuged for 1 hr. Fractions collected from the flotation gradient can be analyzed for protein and endocytosed marker and frozen in
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liquid nitrogen. It is convenient to divide the fractions into small volumes (i.e., 50 to 100 µl) that must be frozen quickly to limit damage to the organelles. These aliquots must also be thawed very quickly and shortly before use.
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Fleischer, S. and Kervina, M. 1974. Subcellular fractionation of rat liver. Methods Enzymol. 31:6-40. Gagescu, R., Demaurex, N., Parton, R.G., Hunziker, W., Huber, L., and Gruenberg, J. 2000. The recycling endosome of MDCK cells is a mildly acidic compartment rich in raft components. Mol. Biol. Cell 11:2775-2791. Garin, J., Diez, R., Kieffer, S., Dermine, J.F., Duclos, S., Gagnon, E., Sadoul, R., Rondeau, C., and Desjardins, M. 2001. The phagosome proteome: Insight into phagosome functions. J. Cell Biol. 152:165-180. Gorvel, J.-P., Chavrier, P., Zerial, M., and Gruenberg, J. 1991. Rab 5 controls early endosome fusion in vitro. Cell 64:915-925. Gruenberg, J. 2001. The endocytic pathway: A mosaic of domains. Nat. Rev. Mol. Cell Biol. 2:721730. Gruenberg, J. and Clague, M. 1992. Regulation of intracellular membrane transport. Curr. Opin. Cell Biol. 4:593-599. Gruenberg, J. and Gorvel, J.-P. 1992. In vitro reconstitution of endocytic vesicle fusion. In Protein Targetting: A Practical Approach (A.I. Magee and T. Wileman, eds.) pp. 187-216. Oxford University Press, Oxford.
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Gruenberg, J.E. and Howell, K.E. 1986. Reconstitution of vesicle fusions occurring in endocytosis with a cell-free system. EMBO J. 5:3091-3101. Gruenberg, J. and Howell, K.E. 1987. An internalized transmembrane protein resides in a fusioncompetent endosome for less than 5 min. Proc. Natl. Acad. Sci. U.S.A. 84:5758-5762 Gruenberg, J. and Howell, K.E. 1989. Membrane traffic in endocytosis: Insights from cell-free assays. Annu. Rev. Cell Biol. 5:453-481. Gruenberg, J., Griffiths, G., and Howell, K.E. 1989. Characterisation of the early endosome and putative endocytic carrier vesicles in vivo and with an assay of vesicle fusion in vitro. J. Cell Biol. 108:1301-1316. Harms, E., Kern, H., and Schneider, J.A. 1980. Human lysosomes can be purified from diploid skin fibroblasts by free-flow electrophoresis. Proc. Natl. Acad. Sci. U.S.A. 77:6139-6143. Howell, K.E., Devaney, E., and Gruenberg, J. 1989a. Subcellular fractionation of tissue culture cells. Trends Biochem. Sci. 14:44-47. Howell, K.E., Schmid, R., Ugelstad, J., and Gruenberg, J. 1989b. Immunoisolation using magnetic solid supports: Subcellular fractionation for cellfree functional studies. Methods Cell Biol. 31A:264-292. Khan, M.N., Savoie, S., Bergeron, J.J.M., and Posner, B.I. 1986. Characterization of rat liver endosomal fractions: In vitro activation of insulinstimulable kinase in these structures. J. Biol. Chem. 261:8462-8472.
Leighton, F., Poole, B., Beaufay, H., Baudhuin, P., Coffey, J.W., Fowler, S., and DeDuve, C. 1968. The large-scale separation of peroxisomes, mitochondria, and lysosomes from the livers of rats injected with Triton WR-1339. J. Cell Biol. 37:482-513. Lippincott-Schwartz, J. and Smith, C.L. 1997. Insights into secretory and endocytic membrane traffic using green fluorescent protein chimeras. Curr. Opin. Neurobiol. 7:631-639. Luzio, J.P., Mullock, B.M., Branch, W.J., and Richardson, P.J. 1988. Immunoaffinity techniques for the purification and functional assessment of subcellular organelles. Prog. Clin. Biol. Res. 270:91-100. Mann, M., Hendrickson, R.C., and Pandey, A. 2001. Analysis of proteins and proteomes by mass spectrometry. Annu. Rev. Biochem. 70:437-473. Marsh, M., Schmidt, S.L., Kern, H., Harms, E., Male, P., Mellman, I., and Helenius, A. 1987. Rapid, analytical, and preparative isolation of functional endosomes by free-flow electrophoresis. J. Cell Biol. 104:875-886. Mueller, S.C. and Hubbard, A.L. 1986. Receptormediated endocytosis of asialoglycoproteins by rat hepatocytes: Receptor-positive and receptornegative endosomes. J. Cell Biol. 102:932-942.
Kindberg, G.M., Ford, T., Blomhoff, R., Rickwood, D., and Berg, T. 1984. Separation of endocytic vesicles in Nycodenz gradients. Anal. Biochem. 142:455-462.
Mullock, B.M., Branch, W.J., vanSchaik, M., Gilbert, L.K., and Luzio, J.P. 1989. Reconstitution of an endosome-lysosome interaction in a cellfree system. J. Cell Biol. 108:2093-2099.
Kobayashi, T., Stang, E., Fang, K.S., de Moerloose, P., Parton, R.G., and Gruenberg, J. 1998. A lipid associated with the antiphospholipid syndrome regulates endosome structure/function. Nature 392:193-197.
Pertoft, H., Warmegard, B., and Hook, M. 1978. Heterogeneity of lysosomes originating from rat liver parenchymal cells. Metabolic relationship of subpopulations separated by density-gradient centrifugation. Biochem. J. 174:309-317.
Kobayashi, T., Beuchat, M.-H., Lindsay, M., Frias, S., Palmiter, R.D., Sakuraba, H., Parton, R., and Gruenberg, J. 1999. Late endosomal membranes rich in lysobisphosphatidic acid regulate cholesterol transport. Nat. Cell Biol. 1:113-118.
Richardson, P.J. and Luzio, J.P. 1986. Immunoaffinity purification of subcellular particles and organelles. Appl. Biochem. Biotechnol. 13:133145.
Kobayashi, T., Beuchat, M.H., Chevallier, J., Makino, A., Mayran, N., Escola, J.M., Lebrand, C., Cosson, P., and Gruenberg, J. 2002. Separation and characterization of late endosomal membrane domains. J. Biol. Chem. 277:3215732164.
Subcellular Fractionation of Tissue Culture Cells
Lebrand, C., Corti, M., Goodson, H., Cosson, P., Cavalli, V., Mayran, N., Fauré, J., and Gruenberg, J. 2002. Late endosome motility depends on lipids via the small GTPase Rab7. EMBO J. 21:1289-1300.
Rothman, J.E. 1994. Mechanisms of intracellular protein transport. Nature 376:55-63. Rout, M.P., Aitchison, J.D., Suprapto, A., Hjertaas, K., Zhao, Y., and Chait, B.T. 2000. The yeast nuclear pore complex. Composition, architecture, and transport mechanism. J. Cell Biol. 148:635-652.
Kreis, T.E. 1986. Micro-injected antibodies against the cytoplasmic domain of vesicular stomatitis virus glycoprotein block its transport to the cell surface. EMBO J. 5:931-941.
Sahagian, G.G. and Neufeld, E.F. 1983. Biosynthesis and turnover of the mannose 6-phosphate receptor in cultured Chinese hamster ovary cells. J. Biol. Chem. 258:7121-7128.
Kreis, T.E. and Vale, R.D. 1993. Guidebook to the Cytoskeletal and Motor Proteins. Oxford University Press, Oxford.
Scheele, G.A., Palade, G.E., and Tartakoff, A.M. 1978. Cell fractionation studies on the guinea pig pancreas. Redistribution of exocrine proteins during tissue homogenization. J. Cell Biol. 78:110-130.
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Schmid, S.L., Fuchs, R., Male, P., and Melman, I. 1988. Two distinct populations of endosomes involved in membrane recycling and transport to lysosomes. Cell 52:73-83. Stahn, R., Maier, K.P., and Hannig, K. 1970. A new method for the preparation of rat liver lysosomes. Separation of cell organelles of rat liver by carrier-free continuous electrophoresis. J. Cell Biol. 46:576-591. Steinert, P.M., Zachroff, R.V., Anyardi-Whitman, M., and Goldman, R.D. 1982. Isolation and characterization of intermediate filaments. Methods Cell Biol.. 24:399-419. Stoorgovel, W., Geuze, H.J., and Strous, G.J. 1987. Sorting of endocytosed transferrin and asialoglycoprotein occurs immediately after internalization in HepG2 cells. J. Cell Biol. 104:1261-1268. Wattiaux, R., Wattiaux-De Coninck, S., RonveauxDupal, M.F., and Dubois, F. 1978. Isolation of rat
liver lysosomes by isopycnic centrifugation in a metrizamide gradient. J. Cell Biol. 78:349-368. Wu, C.C., Yates, J.R.R., Neville, M.C., and Howell, K.E. 2000. Proteomic analysis of two functional states of the Golgi complex in mammary epithelial cells. Traffic 1:769-782. Zerial, M. and McBride, H. 2001. Rab proteins as membrane organizers. Nat. Rev. Mol. Cell Biol. 2:107-117.
Contributed by Fernando Aniento University of Valencia Valencia, Spain Jean Gruenberg University of Geneva Sciences II Geneva, Switzerland
Extraction, Stabilization, and Concentration
4.3.21 Current Protocols in Protein Science
Supplement 32
Desalting, Concentration, and Buffer Exchange by Dialysis and Ultrafiltration
UNIT 4.4
Various steps in protein purification result in concentrations of low-molecular-weight materials, such as (NH4)2SO4, that exceed levels appropriate for subsequent procedures. In cases where simple dilution cannot sufficiently reduce the concentration of these impurities, dialysis or diafiltration—dialysis by ultrafiltration at constant volume— should be considered. Ultrafiltration is also convenient for concentrating protein samples without concomitantly concentrating low-molecular-weight impurities. This is achieved by forcing water and dissolved low-molecular-weight substances through a semipermeable membrane under pressure; the high-molecular-weight protein components are retained by the membrane. Alternatively, ultrafiltration can be carried out via a rapidly recirculating flow of solution over a semipermeable membrane (tangential-flow ultrafiltration). Dialysis and ultrafiltration are also effective methods for buffer exchange—i.e., the replacement of a buffer used for a previous procedure with a different buffer appropriate for a subsequent one. Gel filtration (UNIT 8.3) can also be used for desalting, but because this purification technique normally employs a column containing a differentially porous gel matrix, rather than a semipermeable membrane, it is less readily adapted to large sample volumes. However, despite the availability of membranes with a wide range of porosities, dialysis and ultrafiltration are generally less efficient than gel filtration for separation of proteins on the basis of size difference, except where the difference in size is quite large (i.e., by a factor of ≥10; see Background Information). This unit includes a variety of dialysis and ultrafiltration techniques that can be used for desalting, concentration, or buffer exchange. Basic Protocol 1 describes standard dialysis by diffusion across cellulose tubing, a technique used for desalting or buffer exchange. Basic Protocol 2 describes ultrafiltration under pressure, which can be used either for concentrating protein or, where the sample volume is replenished with a desired buffer, for desalting/buffer exchange (i.e., diafiltration). Alternate Protocol 1 describes ultrafiltration by tangential flow, which serves the same purposes as ultrafiltration under pressure but is better suited to handling large volumes of solution. Alternate Protocol 2 is an ultrafiltration technique employing centrifugal microconcentrators, which allow concentration of very small volumes of solution. Additional information on standard dialysis using membrane diffusion is provided in APPENDIX 3B. DESALTING AND BUFFER EXCHANGE BY DIALYSIS THROUGH REGENERATED CELLULOSE TUBING
BASIC PROTOCOL 1
When sample volume is <200 ml, dialysis can be accomplished by placing the sample in standard dialysis tubing immersed in a buffer of the desired ionic strength and continuously stirring the buffer. For larger volumes, multiple dialysis bags may be used, but desalting or buffer exchange on a large scale is more rapidly achieved by tangential-flow diafiltration (see Alternate Protocol 1). Because the exposure time of the sample to the buffer is relatively long, the composition, pH, and ionic strength of the buffer should be dictated by considerations of protein stability; when large volumes are involved, buffer price may be a factor as well. Extraction, Stabilization, and Concentration Contributed by Allen T. Phillips Current Protocols in Protein Science (1995) 4.4.1-4.4.13 Copyright © 2000 by John Wiley & Sons, Inc.
4.4.1 Supplement 2
Materials Dialysis tubing: regenerated cellulose, MWCO 12,000 (e.g., Spectra/Por Type 4, Spectrum) for general protein use or MWCO 3500 (e.g., Spectra/Por Type 3) for small proteins, stored at 4°C in plastic bag 50% (v/v) ethanol in large wash bottle 10 mM EDTA, pH 8.0 0.05 M NaHCO3 Protein solution (clarified by centrifugation if necessary) Appropriate dialysis buffer 1-liter beaker Tubing closures, two (one weighted and one not weighted) per dialysis bag (e.g., Spectra/Por closures and weighted closures, Spectrum) Small plastic funnel Conductivity meter and probe (optional) Prepare the dialysis tubing 1. Select tubing of appropriate MWCO that has a width such that the sample can be contained in sections ≤25 cm in length (see Table 4.4.1). Cut the dry tubing to the proper length, allowing an additional 10 cm for closing off the ends. Handle tubing only with powder-free gloves to avoid getting skin oils on membrane.
2. Immerse the sections of tubing in distilled water and rub each between two fingers until the layers separate. Using a large wash bottle containing 50% ethanol, rinse the inside and outside of each section thoroughly. The ethanol rinse serves to remove glycerol (used as a humectant) and residual sulfides (from the manufacturing process).
3. Mix 400 ml of 10 mM EDTA (pH 8.0) and 400 ml of 0.05 NaHCO3 in a 1-liter beaker. Transfer the wetted and washed tubing to this beaker and stir 30 min using a magnetic stirrer. 4. Decant the EDTA/NaHCO3 solution and replace with 800 ml distilled water. Stir 10 min on the magnetic stirrer, decant the water, and repeat the water wash. Transfer the tubing to fresh distilled water. If not to be used promptly, the tubing should be stored covered at 4°C in water containing 0.1% sodium azide. Once wetted, the membranes must not be allowed to dry out. Boiling tubing is not recommended prior to use in protein work. For more vigorous removal of glycerol and sulfide in situations where these impurities could be troublesome, the tubing may be immersed in 1 liter of 50% ethanol and heated 1 hr at 50°C,
Table 4.4.1 Approximate Capacities for Dialysis Tubing of Various Diameters
Desalting, Concentration, and Buffer Exchange by Dialysis and Ultrafiltration
Flat width (mm)
Inflated diameter (mm)
Volume/length (ml/cm)
10 18 25 45 54 75
6.4 11.5 16 29 34 48
0.3 1.0 2.0 6.4 9.3 18
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instead of being rinsed as in step 2. Regenerated cellulose tubing may be autoclaved, but this alters the MWCO. Individually packaged 14-cm lengths of radiation-sterilized tubing containing no glycerol, sulfides, or heavy metals are available at several MWCO values between 8,000 and 25,000 (Spectra/Por Biotech Sterile Membranes, Spectrum). These are suitable for small (2- to 20-ml) samples.
Dialyze the sample 5. Close one end of a washed section of tubing firmly with a weighted tubing closure. Squeeze the tubing between two fingers to remove excess water from inside. Alternatively, the tubing section may be closed by tying two simple overhand knots near one end.
6. Slip the open end of the tubing section over the narrow end of a small plastic funnel, then add the protein sample through the funnel. When sample addition is complete, remove the funnel and work the sample down the tubing by squeezing with the fingers to eliminate air space. Clamp the end shut with a non-weighted tubing closure (or knot the tubing carefully twice; see annotation to step 5) and trim off excess tubing. Check for leakage at both ends by applying moderate pressure to the sample area. Excessive air space will lead to dilution of the sample by buffer that enters and displaces air during dialysis; however, extra space should be allowed if the sample is very high in salt concentration, as it may be necessary to permit an increase in volume to prevent rupture of the bag. If the sample is very viscous, it should be diluted with the buffer against which it will be dialyzed, to facilitate outward diffusion.
7. Place the filled dialysis bag in a beaker containing a volume at least ten times the sample volume of dialysis buffer at the desired temperature, making sure that the bag is completely immersed. Stir the dialysis buffer continuously with a magnetic stirrer. Continual stirring serves to maximize the difference in salt concentration between the interior and immediate exterior of the dialysis bag. A larger initial buffer-to-sample ratio (≥100:1) will speed attainment of the desired final ionic strength with fewer changes of dialysis buffer. Although diffusion rate is somewhat reduced in the cold, almost all protein dialysis is conducted at 4°C to enhance protein stability.
8. Follow the progress of the dialysis by measuring the conductivity of the dialysis buffer (if this is practical). Replace the used dialysis buffer with an equal volume of fresh buffer when the increase in conductivity slows down (indicating approach to equilibrium and a drop in the efficiency of the dialysis). Continue replacing the dialysis buffer in this way until the conductivity of the buffer remains essentially unchanged after stirring for 1 to 2 hr. If a conductivity meter is not available, the buffer should be changed at least twice, allowing 4 to 6 hr between changes to permit the system to approach equilibrium. Conductivity measurements are particularly useful when large amounts of salt are to be removed and when the buffer-to-sample ratio is <100:1. In such cases, several replacements of dialysis buffer will be necessary to achieve the desired final ionic strength; conductivity measurements permit dialysis to be completed in minimum time by eliminating guesswork regarding when the system is approaching equilibrium. When a conductivity meter is not used and large samples (or tubing diameters >20 mm) are involved, it may be necessary to extend the time between buffer changes to 8 to 12 hr to insure that equilibrium is attained.
9. Remove the dialysis bag from the buffer and rinse the exterior with distilled water. While holding the bag just below the upper closure (or knot), remove that closure (or cut off the knot). Carefully tip the bag and pour the dialyzed sample into an appropriate container. Confirm that the desired salt concentration was achieved by comparing conductivity measurements of the dialyzed sample and unused dialysis buffer.
Extraction, Stabilization, and Concentration
4.4.3 Current Protocols in Protein Science
Supplement 2
If the bag is extremely taut as a result of buffer infiltration, do not cut the upper knot; instead put one end of the tubing into a beaker large enough to hold the entire sample, then puncture the tubing near the lower knot with a long needle and allow the sample to drain directly into the beaker. BASIC PROTOCOL 2
CONCENTRATION OR DIALYSIS BY ULTRAFILTRATION THROUGH ASYMMETRIC MEMBRANE DISKS Protein samples at an initial concentration ≤20 mg/ml can easily be reduced in volume using an ultrafiltration system and membrane, as in the protocol described below. The stirred-cell system used in this protocol—in which nitrogen gas pressure forces molecules smaller than the MWCO through the pores—is available with a variety of chamber sizes and membrane properties. Because most salts pass through the membrane along with water, the sample volume is reduced without a significant change in its surrounding buffer composition, an obvious advantage over lyophilization. Ultrafiltration can also be used to remove or exchange salts without changing the protein concentration. This process, called diafiltration, resembles dialysis but is driven by the rate of ultrafiltration (see Critical Parameters and Troubleshooting), not by a concentration gradient. Diafiltration is carried out as in the concentration protocol here, except that continuous or periodic additions of buffer of the desired final pH, concentration, and composition are made to compensate for changes in protein concentration that would otherwise occur as a result of volume reduction. Materials Protein solution (clarified by centrifugation if necessary) N2 gas cylinder and two-stage pressure regulator capable of controlling delivery up to 100 psi 5% ethanol Ultrafiltration membrane (YM10, Amicon; PLGC, Millipore; or Molecular/Por Type C, Spectrum) with MWCO >10,000 and diameter to fit stirred cell Ultrafiltration unit, stirred-cell type (Series 8000, Amicon, or Molecular/Por, Spectrum) with: Tubing for connection to N2 source Glass funnel with stem diameter to fit through cell fill hole (optional; used in certain cell models) Ultrafiltrate collection container (Griffin beaker with graduations) Pressurized reservoir (optional; for use when sample size is large and protein is dilute, or when diafiltration is performed with constant readdition of new buffer): e.g., Spectra/Chrom Pressure Reservoir (Spectrum) or RG5 Fiberglass Reservoir (Amicon) 1. Rinse a new ultrafiltration membrane with distilled water to wet it thoroughly. Handle membrane only with powder-free gloves and hold by the edge to avoid scratching the smooth, thin “skin” surface.
Desalting, Concentration, and Buffer Exchange by Dialysis and Ultrafiltration
Some ultrafiltration membranes can be used more than once. The Millipore PLGC (MWCO 10,000) or PLTK (MWCO 30,000) cellulose membranes are intended mainly for single use. Many other suppliers indicate that their membranes can be reused until clogged or damaged. Such membranes may be more economical, but only if each user makes sure that the previously used membrane is undamaged and properly cleaned. Amicon YM series or Spectrum Molecular/Por Type C membranes can be cleaned by soaking in 0.01 M SDS solution or 1% Terg-A-Zyme (e.g., Baxter or VWR) for several
4.4.4 Supplement 2
Current Protocols in Protein Science
hours; the membranes are then thoroughly rinsed with distilled water before reuse. Used, cleaned membranes should be stored covered with 5% ethanol in a closed container at 4°C. For membranes other than the ones mentioned, the manufacturer’s instructions must be consulted for a suitable cleaning procedure.
2. Load the membrane into the stirred cell with the “skin” or glossy side up. Complete assembly of the ultrafiltration system, taking special care to ensure that O ring seals (if used) are not nicked and are properly placed on the membrane disk to prevent leakage around the membrane when pressure is applied. The size and thus the membrane diameter of the ultrafiltration cell should be as large as practical for the amount of sample to be concentrated; a larger cell reduces the necessity to refill the unit and provides the largest filtration area, speeding the concentration process. Cells are commonly available in sizes ranging from 10 to 400 ml.
3. Fill stirred cell with sample to the maximum operating volume, either by pouring the sample directly through the fill hole or by using the glass funnel provided with the cell in systems where the fill hole is small. Replace the fill-hole cover and check that the pressure-release valve (often an integral part of the fill-hole cover) is in the closed position. If a funnel is used, contact between the funnel tip and membrane surface should be avoided.
4. Place the unit on a magnetic stirrer and stir at the maximum rate that does not generate foam or a sizeable vortex at the surface of the solution. Direct the outlet tubing to the ultrafiltrate collection container. Because of protein stability requirements, most ultrafiltrations are conducted in the cold. It is advisable to check the surface temperature of the magnetic stirrer periodically; if it is warm, a styrofoam sheet 0.5-in. thick should be placed between the stirrer surface and the cell bottom.
5. Connect the cell to the nitrogen gas regulator and pressurize to 50 psi. Flow of liquid into the outlet tubing should quickly become evident. The system should be examined for appearance of liquid at the base of the cell where the cylinder body contacts the O ring; this indicates a leak around the membrane, in which case the unit should be depressurized and its assembly examined. Lower or higher pressures may be needed depending on the membrane type and MWCO value, but pressure should not exceed 70 psi in most stirred cells.
6. Monitor the flow rate and ultrafiltrate volume until the desired rate of concentration has been achieved. If the total sample volume exceeds the cell capacity, the cell should be refilled when the volume has decreased by one-third. Sample additions are then continued until all of the sample has been added to the cell. A pressurized sample reservoir is recommended when the sample volume is many times the cell capacity. At a constant temperature and pressure, the flow rate is a function of the filter area and the degree to which concentration polarization can be avoided (see Critical Parameters). Vigorous stirring is needed to minimize concentration polarization of protein at the membrane surface. Buildup of protein on the surface will result in slow filtration, even when the protein concentration of the sample is relatively low. Filtration rates at 4°C are often only one-half those seen at 25°C because of the influence of viscosity. Do not concentrate the sample to the point where it is no longer in contact with the stir bar, as lack of stirring will lead to severe concentration polarization in all but the most dilute samples. Overconcentration makes sample recovery difficult and may require readdition of buffer to wash the membrane, thereby adding to the volume.
7. Turn off the stirrer and the nitrogen flow to the cell. Slowly release the internal cell pressure, then turn the stirrer back on and allow stirring to continue for 2 min to
Extraction, Stabilization, and Concentration
4.4.5 Current Protocols in Protein Science
Supplement 2
resuspend protein in the gel layer. Finally, turn the stirrer off and remove the concentrated sample by pouring or pipetting through the fill hole. If a pipet is used and the membrane is of the reusable type, care should be taken not to touch the pipet to the membrane surface. An alternative means of removing the sample is to unclamp the cell and pour out the sample, after checking that the body of the unit is firmly screwed into the base.
8. If the membrane is of the reusable type, hold it by the edges and rinse it well with distilled water. Proceed as soon as possible with the manufacturer’s suggested cleaning procedure, keeping the membrane wet at all times, then store the cleaned filter in 5% ethanol in a closed container at 4°C. ALTERNATE PROTOCOL 1
DIAFILTRATION OR CONCENTRATION BY TANGENTIAL-FLOW ULTRAFILTRATION The most rapid method for dialyzing large samples is the use of ultrafiltration membranes fabricated into hollow fibers or spiral-wound layers, or, as in this protocol, in the form of stacked plates. The sample is allowed to flow tangentially over these membranes with recirculation. The semipermeable nature of the membrane then permits water and materials with molecular weights that are substantially below the MWCO value to cross through the membrane surface relatively unimpeded; retained materials, including the proteins of interest, are returned to the sample reservoir. To use this technique for diafiltration (as opposed to concentration), fresh dialysis buffer of the desired composition should be added to the sample reservoir in a volume equal to that of the ultrafiltrate collected, so that the concentration of macromolecular species remains unchanged. Removal of microsolutes to which the membrane is completely permeable can be estimated from the volume of ultrafiltrate (or diafiltrate, in this case) collected. Collection of an amount of diafiltrate equal to 3 vol of original sample will remove 95% of the starting solute to which the membrane is permeable, while a diafiltrate of 5 vol will correspond to a 99% separation. The same process is used to concentrate samples, except that the retentate volume is allowed to decrease until the desired degree of concentration is reached. Maintaining a rapid flow over the membrane with a moderate transmembrane pressure minimizes concentration polarization and keeps the filtration rate high even as protein concentration in the sample increases. The procedure described here is based on the Millipore Minitan acrylic ultrafiltration system, which employs stacked filter plates and a peristaltic pump. It is designed for sample sizes from 200 ml to 2 liters, but will work with larger samples if they are dilute. Equivalent units include the Spectrum MP-3 system and Amicon CH2PRS concentrator, both of which employ spiral-wound membrane cartridges. Additional Materials (also see Basic Protocol 1) 1% (w/v) Terg-A-Zyme detergent (e.g., Baxter or VWR), 40°C 0.05% (w/v) sodium azide
Desalting, Concentration, and Buffer Exchange by Dialysis and Ultrafiltration
Minitan membrane filter plates (Millipore) made of cellulose (PLGC, MWCO 10,000 or PLTK, MWCO 30,000) or polysulfone (PTTK, MWCO 30,000) Minitan acrylic ultrafiltration system with pump assembly and tubing (Millipore) Hose clamps (adjustable with screws) Container capable of holding entire sample
4.4.6 Supplement 2
Current Protocols in Protein Science
Prepare the system 1. Select a filter plate of appropriate MWCO. Filtration rates are higher with MWCO 30,000 membranes, but the large volumes to be filtered in diafiltration may lead to some loss of proteins with molecular weight <50,000 using such membranes. Membranes of lower MWCO are also available (MWCO 1000, 3000, and 5000) but these filter more slowly and can reject some of the buffer ions, thus requiring more filtrate volume to provide complete buffer exchange.
2. Disassemble the Minitan cell to expose the lower acrylic manifold. Place the first silicone retentate separator on the acrylic surface. Carefully slip a filter plate down on the alignment pins until it is seated against the silicone separator. Follow this with the second silicone retentate separator, then another filter plate, until the desired number of filter plates have been stacked—varying the left-to-right orientation of plates at each level to permit tangential flow across each plate. After the last plate and separator have been inserted, replace the upper acrylic manifold, then the top steel plate. Complete the assembly by adding the washers and nuts, tightening them against the plate with a torque wrench to the recommended pressure. Layers of filter plates alternating with silicone separators can be added up to a total of 10, each with an area of 60 cm2. Use multiple filter plates if available, especially when initial volumes exceed 2 liters. Additional filter plates increase the filtration rate in a linear fashion by providing more cross-sectional area.
3. Attach tubing to the retentate inlet on the lower manifold and run it through the pump to the sample reservoir. Likewise, attach tubing to the retentate outlet on the upper manifold and bring it to the sample reservoir. Attach a hose clamp on this section of tubing to permit the transmembrane pressure to be increased if necessary. Attach tubing from both filtrate outlets to a collection vessel; provide each section of outlet tubing with a hose clamp. A Y-shaped tube connector can be used to direct the output of both filtrate exit ports to a single piece of tubing.
4. Place a small amount of dialysis buffer in the sample reservoir and slowly bring the pump to a mid-range operating speed. Run the pump until the dialysis buffer has circulated through and purged the filter plates and all tubing lines. Stop the pump and discard the contents of the sample reservoir and collection vessel. Perform ultrafiltration 5. Fill the sample reservoir with the protein sample to be diafiltered or concentrated. Restart the pump and adjust the flow to 100% recirculation by clamping off the filtrate outlets. Measure the flow and readjust the pump speed until the maximum cross-flow that provides an inlet pressure of 25 psi is achieved. 6. Open the filtrate outlets and begin collection of the ultrafiltrate. If filtrate flow is inadequate, some increase can be achieved by partially restricting the retentate outlet line by adjusting the screw on the hose clamp, thereby increasing the transmembrane pressure. The inlet pressure should not be allowed to exceed the recommended level for the tubing being used (usually 25 psi for silicone tubing with 4 mm i.d.). However, reduced tangential flow can lead to greater concentration polarization and ultimately a reduced dialysis rate. A recirculation flow/filtrate flow ratio of 20:1 may be satisfactory for dilute samples, but higher ratios are more successful at preventing concentration polarization. Because shear forces are normally not high during recirculation, protein stability is seldom reduced as a consequence of increasing this ratio.
Extraction, Stabilization, and Concentration
4.4.7 Current Protocols in Protein Science
Supplement 2
7. As filtration proceeds, frequently refill the sample reservoir with fresh dialysis buffer to restore the original volume. Continue doing this until the volume of ultrafiltrate is three to five times the volume of the original sample, indicating that removal of diffusible material is 95% to 99% complete. If concentration is desired, allow the sample volume to decrease after this point until the desired volume is achieved. 8. Clamp off the filtrate line and slow the pump to terminate filtration. Remove the retentate inlet line from the sample reservoir and place it in a solution of fresh dialysis buffer, then allow the pump to push sample into the reservoir until a volume of buffer equal to the internal volume of the system has been collected (usually ∼25 ml). Finally, stop the pump and remove the contents of the sample reservoir. Clean the system 9. Clean the unit and membrane before disassembly by first pumping water through all lines to flush out remaining sample and buffer salts, then circulating 1% Terg-AZyme, 40°C, through the system for 30 min. Collect output from filtrate and retentate line in the same reservoir so that all solution is reused. Flush the system with 5 to 10 liters of water, discarding all output from the filtrate and retentate lines and making sure that all lines are flushed. 10. Disassemble the unit and store the filter plate(s) in 0.05% sodium azide in a covered plastic container at 4°C. ALTERNATE PROTOCOL 2
MICROSCALE CONCENTRATION AND DESALTING WITH CENTRIFUGAL ULTRAFILTERS Sample preparation for analytical purposes and small-scale purification may require concentration and/or desalting of volumes <1 ml. Stirred-cell assemblies (see Basic Protocol 2), although able to deal with amounts only slightly greater than that, often have unacceptably large internal volumes making sample recovery difficult. Moreover, as concentration proceeds the risk of excessive volume reduction (even to dryness) increases, because filtration with stirred cells is not self-limiting. The advent of commercially available, single-use ultrafiltration microconcentrators powered by centrifugal force has eliminated many of these problems. These are available with a range of MWCO values in sizes suitable for handling volumes of 50 µl to 2 ml (e.g., Microcon units from Amicon). The microconcentrators can often be combined with an additional microporous membrane of 0.22-µm pore size to remove large particulates— e.g., polyacrylamide—at the same time the protein sample is being concentrated or desalted (e.g., Micropure Separator inserts from Amicon, used with the Microcon units). Centrifugal ultrafilters are also produced in sizes that will accommodate samples of ≥10 to 25 ml, but their limited membrane area and minimal remixing can result in low rates of concentration as compared with stirred cells, except when samples are dilute. Some recent designs appear to have greatly reduced concentration polarization (e.g., Centriprep from Amicon); these mostly involve improved methods of producing movement of liquid through the membrane via centrifugal force.
Desalting, Concentration, and Buffer Exchange by Dialysis and Ultrafiltration
Additional Materials (also see Basic Protocol 1) Centrifuge microconcentrator units: e.g., Microcon-10, -30, or -50 (MWCO 10,000, 30,000, and 50,000 respectively; Amicon); Ultrafree-MC with MWCO 5,000, 10,000, or 30,000 (Millipore); or Centri/Por with MWCO 10,000, 25,000, or 50,000 (Spectrum) 1. Pipet ≤500 ml of protein solution into microconcentrator and seal with the cap supplied by the manufacturer. When using a single concentrator or when multiple ones are not
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Current Protocols in Protein Science
matched in weight, prepare a balance tube (if necessary) by pipetting a volume of water equal to that of the sample into a previously used microconcentrator. 2. Place the microconcentrator opposite its balance tube in a microcentrifuge in a 4°C refrigerator or cold room. Microcentrifuge 20 to 40 min at the speed specified for the filter unit by the manufacturer. A nonrefrigerated microcentrifuge may develop temperatures detrimental to protein samples when operated for extended periods; therefore it is usually best to have the microcentrifuge in a refrigerator or cold room for this operation, even though the filtration rate is reduced by the cold. Microcon units can withstand forces of 13,000 × g, whereas the Ultrafree units are limited to 5,000 × g. If the sample is not sufficiently concentrated after 20 to 40 min, centrifugation should be continued an additional 20 min to see if further volume reduction occurs. Most centrifugal microconcentrators are designed to stop at a volume of 5 to 15 ìl. Typical centrifugation times to concentrate 500 ìl of 0.1% bovine serum albumin to 10 ìl at 4°C are 25 min for the Microcon-30 and 70 min for the Microcon-10. These times are twice those observed at 25°C; recovery is ∼95% in either instance. If desalting without concentration is required, remove the microconcentrator from the microcentrifuge at the end of the centrifugation period, unscrew the sample reservoir from the filtrate tube, then pour off the filtrate from the filtrate tube. Replace the reservoir on the filtrate tube and add a sufficient amount of suitable buffer to restore the original sample volume. Replace the cap and invert to mix. If the sample does not show the desired final ionic strength as determined by conductivity measurement, repeat the centrifugation and readdition of buffer.
3. Once the desired degree of concentration has been achieved, remove the cap from the unit and place the filter portion upside down into the retentate tube supplied by the manufacturer. Place the reversed unit back in the microcentrifuge and spin 15 sec to drive the concentrate down to the tip of the tube. Remove the unit from microcentrifuge and recover the sample from the retentate tube. With some microconcentrators, adsorption of protein to the walls of the unit as well as to the filter itself can be significant when the sample is very dilute (≤25 ìg/ml). Passivation of the surfaces by soaking the entire unit overnight in 5% Tween-20, followed by thorough washing with distilled water, may help alleviate this problem. Suelter and Deluca (1983) suggest that modification of the protein solvent by addition of glycerol or Triton X-100 (below the critical micelle concentration) is superior to treatment of the surfaces of the microconcentrator.
COMMENTARY Background Information Used primarily for removal of excess salts or organic solvents (e.g., acetone or ethanol) employed as precipitants in protein purification or analysis, molecular separations by means of differentially permeable membranes have been an integral part of protein biochemistry since the earliest purifications. Membranes of natural origin have long been supplanted for use in dialysis by regenerated cellulose tubing. Relatively recent developments in membrane technology have provided the asymmetric designs now found in ultrafiltration membranes. This asymmetry refers to the “sidedness” built into ultrafiltration membranes, wherein a dense
layer or skin ∼0.1 to 0.5 µm in thickness with a carefully controlled pore size is bonded to one side of a thicker structural layer (composed of the same or different material) that has a large open-celled structure highly permeable to most molecules. A variety of materials have been used to fabricate these semipermeable membranes, ranging from cellulose and cellulose esters to polyethersulfone or polyvinylidene difluoride (PVDF). Because of proprietary interests, specific details concerning construction of these filters—particularly in regard to the more complex materials used in tangentialflow ultrafiltration—are not widely disseminated by their manufacturers. However, several
Extraction, Stabilization, and Concentration
4.4.9 Current Protocols in Protein Science
Supplement 2
Desalting, Concentration, and Buffer Exchange by Dialysis and Ultrafiltration
companies (Millipore, Amicon, and Spectrum) make available extensive information on their product lines and offer considerable technical advice and literature pertaining to their use. All membranes, whether used for dialysis or ultrafiltration, are characterized by their molecular-weight cutoff (MWCO) value. This is usually defined as the molecular weight of a solute that is 90% prevented from penetrating the membrane under a chosen set of conditions. For some membranes the MWCO value is based on calibration using a mixture of dextrans, while for others it is based on calibration with specific globular proteins. How readily a particular protein is rejected by the membrane is a function of the shape, hydration state, and charge of the protein molecule. Hence, it is possible for two proteins of similar molecular weight to have decidedly different retention properties when a membrane with a stated MWCO that is near their molecular weight is used to separate them. Moreover, MWCO values are not sharp; rather, there is a gradual increase in retention as the size of solute molecules approaches and exceeds the average membrane pore size. Only at the point where all pores are smaller than a particular solute molecule is that molecule completely excluded (Saltonstall, 1992a). In standard dialysis procedures, the requirement is for a large amount of a diffusible material to migrate through the membrane while high-molecular-weight components are retained. The rate of diffusion is influenced primarily by four factors: the concentration gradient of the diffusible material, the diffusion constant of that material, the area of the membrane, and the temperature. An increase in any or all of these variables produces more rapid dialysis; however the two that can most readily be varied are the gradient and the membrane area. The gradient should be kept large by rapid stirring of the dialysis buffer, which keeps the concentration of diffusible solutes as low as possible at the membrane exterior surface, as well as by frequent changes of the buffer solution. It is also desirable to stir the contents of the dialysis bag—which speeds up internal diffusion and minimizes concentration polarization—but this is not easily done without special equipment. The primary reason that dialysis is commonly used is that it is technically uncomplicated; even though the technique is moderately slow, arrangements intended to increase efficiency (e.g., rocking dialyzers) are rarely used (McPhie, 1971).
The rate of movement for permeable solutes in dialysis is difficult to predict, but the end state that can be achieved can be readily estimated. Ignoring possible inequality between the concentrations of low-molecular-weight ions on the two sides of a membrane that may result from a Donnan effect, the equilibrium salt concentration is predictable from simple dilution considerations. This is illustrated by the following example. If a 20-ml protein sample in 2 M ammonium sulfate/50 mM potassium phosphate is dialyzed against 1000 ml of a dialysis buffer containing 50 mM potassium phosphate without ammonium sulfate, the dilution factor will be 1,020⁄20; thus, at equilibrium the (NH4)2SO4 concentration will be 0.039 M. If, after equilibration, this buffer is replaced with 1000 ml of fresh 50 mM ammonium sulfate, upon equilibration the ammonium sulfate concentration will be <0.001 M. If only 200 ml of 50 mM potassium phosphate is used for each of the two buffer changes, however, the ammonium sulfate concentration in the sample will approach 0.016 M. It is thus highly desirable to change the dialysis buffer several times or to use a large ratio of dialysis buffer to sample. Both of these strategies serve to maximize the concentration differential that drives the process of dialysis towards the solute concentration present in the original dialysis buffer. Separation by means of dialysis or ultrafiltration of two protein species that differ in their molecular weights (and R values; see Critical Parameters) is possible under some conditions, but generally will be incomplete unless the size difference is large. Saltonstall (1992b) has concluded that an efficient separation from an initial 1:1 ratio to a final 10:1 ratio would require a ratio of retention values of 10:1 (e.g., 0.9 versus 0.09), corresponding to a molecularweight ratio of 23:1 for several different types of membrane.
Critical Parameters and Troubleshooting Membrane impurities Dialysis membranes made from regenerated cellulose generally contain appreciable amounts of heavy metals, with iron, zinc, and lead present at levels in excess of 1 to 10 ppm. They also contain ∼0.1% sulfur from the manufacturing process. Pretreatment may be necessary to remove these materials when their presence is detrimental. Cellulose ester mem-
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branes do not contain these contaminants and thus can be used directly; however they are less tolerant of organic solvents and high pH than regenerated cellulose membranes. Ultrafiltration The advantage of desalting processes based on ultrafiltration over those based on simple dialysis is that the rate of low-molecular-weight solute removal is not determined by a concentration differential, but rather by the flow rate of solvent and the rejection of the solute by the ultrafiltration membrane employed. The rejection (R) of a solute is defined by the relationship R = 1 − (CF/CI), where CF is the final concentration of the solute (in the filtrate) and CI is the initial concentration of the solute (in the solution prior to filtration). In desalting by diafiltration, where the filtrate is replaced by fresh dialysis buffer at the same rate at which it is removed, a membrane is usually selected whose MWCO is much larger than the molecular weight of the solute to be removed. This produces an R equal to zero for the permeable solute, and, optimally, an R essentially equal to 1 (i.e., total rejection) for the protein(s) of interest—a situation in which loss of protein is minimized. When the diafiltration involves removal of a solute with an R equal to zero, the degree of washout of the solute can be calculated from the volume of filtrate collected relative to the original sample volume according to the equation: ln (CI/CR) = (VF/VI), where CR is the final concentration of the solute in the retentate, VF is the volume of ultrafiltrate collected (also equal to the volume of dialysis buffer added back to the sample), and VI is the initial sample volume (which remains constant in diafiltration because of readdition of buffer). Using this equation, the volume of filtrate that must be collected relative to sample size (VF/VI) can be estimated for any degree of desalting required. Hence, to reduce the concentration of solute in the sample 10-fold (i.e., CI/CR = 10), the volume of filtrate collected must be 2.3 × the original sample volume; for a 100-fold reduction, this volume ratio must be 4.6. Membranes for ultrafiltration are generally selected on the basis of the MWCO needed to retain the protein of interest but allow the maximum amount of other materials to pass through. It is usually best to choose an MWCO value that is roughly one-half the molecular weight of the species to be retained. This provides a reasonable margin of retention whereby almost none of the protein of interest should be lost, but at the same time provides the largest differ-
ence between the MWCO value and the molecular weight of the salts to be removed, thereby maximizing filtration rate. Table 4.4.2 lists flow rates for several types of membrane disks. Concentration polarization The term concentration polarization describes the concentration gradient of solutes (mainly those to which the membrane is not readily permeable) that forms immediately above the membrane on the side under pressure. This condition exists to some degree even at low solute levels. Concentration polarization can be reduced by efficient remixing of the solution—either by stirring action (in stirredcell systems) or tangential movement of solution over the membrane (in tangential-flow systems). However, if the concentration of material within the gel layer at the membrane surface reaches a value that is sufficiently high, the filtration rate becomes pressure-independent and may fall to near zero. One of the chief advantages of tangentialflow ultrafiltration systems is that they allow better control of concentration polarization than stirred-cell systems. In tangential-flow systems, concentration polarization is controlled by varying the recirculation rate of the retentate and adjusting the transmembrane pressure. Flux of filtrate through the membrane is a linear function of transmembrane pressure, until gel formation on the surface becomes the limiting factor. When this happens, increasing the velocity across the membrane surface without further increasing the transmembrane pressure will improve flux. Adsorption of protein to the membrane In regard to the degree of nonspecific adsorption of protein to membranes, some important differences exist among the various materials used in membrane fabrication. Most suppliers indicate that their cellulose membranes exhibit the least nonspecific adsorption of protein; these are therefore recommended for protein work even though they may have slower filtration rates than membranes fabricated from other materials. The actual amount of protein bound depends on the quantity present, its charge characteristics, and the membrane area. Even with the best membranes, losses of 1% to 5% are not uncommon when dealing with total quantities of protein in the range of 1 to 10 mg using a filter with a 43-mm diameter. The nature of the buffer can also affect adsorption of protein; some membranes exhibit altered flow
Extraction, Stabilization, and Concentration
4.4.11 Current Protocols in Protein Science
Supplement 2
Table 4.4.2
Typical Flow Rates for Disk-Type Ultrafiltration Membranes
Membranea
MWCO (kD)
Approximate flow rate at 25°C (ml/min/cm2) For water
For protein solution
Amicon YM3 YM10 YM30 PM10 PM30
3 10 30 10 30
0.07b 0.2b 0.9b 2.5b 4.0b
0.07c 0.15c 0.2c 0.2c 0.3c
Spectrum Molecular/Por Type C Molecular/Por Type C
3 10
0.1d 0.2d
— —
Millipore Ultrafree Ultrafree
10 30
0.05e 0.4e
0.05f 0.1f
aAddresses and telephone numbers of suppliers are provided in the SUPPLIERS APPENDIX. bDetermined at 55 psi. cDetermined at 55 psi using 0.1% albumin. dDetermined at 50 psi. eDetermined at 2000 × g. fDetermined at 2000 × g using 0.1% IgG.
properties when high levels of ions are present. In this regard, phosphate buffers seem to present more of a problem than Tris buffers. Precipitation of protein Desalting sometimes results in precipitation of a portion of the protein in the sample. Although the insoluble material can be removed by centrifugation, it should not be discarded until assays establish that the desired protein has remained in solution. If the precipitated material includes the protein of interest, it can often be redissolved in a solution of higher ionic strength.
Anticipated Results
Desalting, Concentration, and Buffer Exchange by Dialysis and Ultrafiltration
When carried out correctly, dialysis and diafiltration should produce a nearly complete recovery of the protein of interest in a buffer solution of the desired type and ionic strength. In the case of dialysis, this requires that the procedure be carried out for a sufficient time period to equilibrate the salt concentrations inside and outside the dialysis bag. In the case of diafiltration with an asymmetric membrane, this requires that a volume of filtrate be collected that is equal to approximately five times the initial sample volume.
The degree of concentration to be achieved by ultrafiltration should be that required for subsequent work. However, it should be kept in mind that once a protein concentration >30 mg/ml has been reached, further concentration is attained with increasing difficulty and production of very concentrated solutions can therefore be slow. Recovery of sample following concentration is generally >95%; failure to achieve this value usually indicates leakage into the filtrate or nonspecific binding to the membrane and/or concentration apparatus itself.
Time Considerations Dialysis through regenerated cellulose tubing for desalting or buffer exchange usually requires 6 to 12 hr for volumes ≤10 ml and 12 to 24 hr for volumes from 10 to 200 ml. Dialysis time can be reduced by more frequent buffer changes, by increasing the ratio of dialysis buffer to sample, by increasing the surface area of the bag, and by monitoring the conductivity of the dialysis buffer to determine when the desired ionic strength has been reached. The rate of ultrafiltration, whether performed for concentration or diafiltration, varies considerably as a function of the MWCO and surface area of the membrane and the initial
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sample concentration. A 10-fold reduction in sample volume (or 10-fold reduction in the concentration of a low-molecular-weight impurity by diafiltration) can usually be accomplished in 1 to 4 hr.
Key References Pohl, T. 1990. Concentration of proteins and removal of solutes. Methods Enzymol. 182:68-83. A recent review of the topics covered here.
Literature Cited
Scopes, R.K. 1994. Protein Purification: Principles and Practice, 3rd ed. pp. 14-21. Springer-Verlag, New York.
McPhie, P. 1971. Dialysis. Methods Enzymol. 22:23-32.
Covers concentration and dialysis in the context of protein purification techniques.
Saltonstall, C.W. 1992a. Separations with dialysis and ultrafiltration: Theoretical and practical considerations. Am. Biotechnol. Lab. 10:32-34. Saltonstall, C.W. 1992b. Dialysis and ultrafiltration: Estimating quantitative separations. Am. Biotechnol. Lab. 10:18-20.
Contributed by Allen T. Phillips Pennsylvania State University University Park, Pennsylvania
Suelter, C.H. and Deluca, M. 1983. How to prevent losses of protein by adsorption to glass and plastic. Anal. Biochem. 135:112-119.
Extraction, Stabilization, and Concentration
4.4.13 Current Protocols in Protein Science
Supplement 2
Selective Precipitation of Proteins
UNIT 4.5
Selective precipitation of proteins (Rothstein, 1994) can be used as a bulk method to recover the majority of proteins from a crude lysate, as a selective method to fractionate a subset of proteins from a protein solution, or as a very specific method to recover a single protein of interest from a purification step. Except for antibody-mediated precipitation (see UNIT 10.8), selective precipitation methods are usually not protein-specific; the process depends on the physical and/or chemical interaction of the precipitating agent with one or more proteins that possess certain characteristics. It may be necessary to use a combination of selective precipitation techniques to isolate the desired protein. The success of precipitation is monitored by measuring total protein content (UNIT 3.4), by identifying proteins in the fractions using SDS-PAGE (UNIT 10.1), and by performing a suitable bioassay to identify the fraction containing the desired protein. Developing a selective precipitation technique for a particular protein requires identification of the appropriate precipitating agent and optimization of the precipitation procedure using that agent. Often, more than one reagent will successfully precipitate a particular protein, so selecting a reagent is a matter of identifying the one that provides the desired protein in the most optimal state. Some reagents cause denaturation or adversely affect bioactivity, and some form complexes that bind the protein tightly. Still others lack the ability to selectively enrich for the desired protein. This unit describes a number of methods suitable for selective precipitation (see Table 4.5.1). In each of the protocols that are outlined, the physical or chemical basis of the precipitation process, the parameters that can be varied for optimization, and the basic steps for developing an optimized precipitation are described.
Table 4.5.1
Protein Precipitative Techniques
Technique
Protocol
Salting out
Basic Protocol 1, Alternate Protocol 1
Isoionic precipitation
Basic Protocol 2, Alternate Protocol 2
Two-carbon (C2) organic cosolvent precipitation of proteins
Basic Protocol 3
C4 and C5 organic cosolvent precipitation, phase partitioning, and extraction of proteins
Basic Protocol 4
Protein exclusion and crowding agents (neutral polymers) and osmolytes
Basic Protocol 5
Synthetic and semisynthetic polyelectrolyte precipitation
Basic Protocol 6
Metallic and polyphenolic heteropolyanion precipitation
Basic Protocol 7
Hydrophobic ion pairing (HIP) entanglement ligands
Basic Protocol 8
Matrix-stacking-ligand coprecipitation
Basic Protocol 9
Di- and trivalent metal cation precipitation
Basic Protocol 10 Extraction, Stabilization, and Concentration
Contributed by Rex E. Lovrien and Daumantas Matulis Current Protocols in Protein Science (1997) 4.5.1-4.5.36 Copyright © 1997 by John Wiley & Sons, Inc.
4.5.1 Supplement 7
STRATEGIC PLANNING Clarification of Samples If raw samples are laden with particulate matter that cannot be centrifuged (at 5 to 50 × g) or filtered out by ordinary means, sometimes suspensions can be clarified by filtration on a 2- to 4-mm layer of “filter-aid” material—e.g., Filter-Aid (Schleicher and Schuell)—on Whatman no. 1 filter paper. Several kinds of filter aid materials for particulate and gelatinous samples are available—e.g., cellulosic fibers (Filter-Aid from Schleicher and Schuell or Hyflo Supercel from Fisher), diatomaceous minerals (e.g., Celatom from Aldrich, kieselguhr from Baxter, or diatomaceous earths from Sigma), silicates (e.g., Celite from Aldrich), specialized clay absorbents, calcium minerals, bentonite, or Fuller’s earth. These should be experimented with—e.g., by adding 0.05 to 0.2 g of the material to 1 to 2 ml of the sample to find out which filter aid adsorbs gelatinous or particulate material without excessive adsorption of the protein. The layer or pad of filter aid for use with the main sample is prepared by adding a few grams to the filter paper and distributing it evenly by puddling it with water while pulling moderate suction on a Buchner funnel. Frequently, a means for freeing proteins in solution from particles and haze by centrifugation or simple filtration can be quickly worked out by adjusting the pH or salt content (or both). The pH may be adjusted up or down by one to three pH units; also, salt may be added to make particulates and colloidal material easier to centrifuge or filter out. Bioactivity assays should be carried out even at the clarification stage to ensure that pH changes and salt additions did not destroy the activity or functions of the proteins to be isolated. Figure 4.5.1 summarizes the steps in clarification of protien solutions. If the conservative techniques for clarification (filter aids and pH/salt variation) do not work, more aggressive techniques may be used that employ some of the rather powerful particle and colloid agglomerating agents such as synthetic polyelectrolytes (described below; see Basic Protocol 6), or metal-ion flocculating agents such as aluminum ions. These need to be added in quite limited amounts (in micrograms to a few milligrams per ml of sample), cautiously and slowly, to coagulate unwanted particles without precipitating sought-for proteins at this preliminary clarification stage. An advantage of metal ions as particulate coagulants or filter aids is that any excess of metal ions can be conveniently removed by conventional cation-exchange resins in K+, Na+, or NH4+ form (see Basic Protocol 2). Unwanted particulates like whole cells, cell membranes, and haze may also be removed by centrifugation in some samples. However, for very large volumes (and where centrifugation is not successful) filtration using filter-aid compounds is frequently useful. Pressure membrane ultrafiltration (UNIT 4.4) with a large MWCO (>30,000) may be tried if such membranes pass the sought-for proteins and hold back the haze particles. Concentration of Samples
Selective Precipitation of Proteins
Very dilute samples beyond the reach of normal precipitating agents for proteins may need to be concentrated before adding precipitating agents. Lyophilization (freeze-drying) is sometimes successful for concentrating certain proteins that are able to withstand removal of virtually all water. However, a majority of proteins, including clotting-cascade and many blood plasma and cytosol proteins, cannot withstand lyophilization. It is more generally preferred, therefore, to use pressure membrane ultrafitration (UNIT 4.4) with a membrane of appropriate MWCO to concentrate sought-for proteins while keeping them cautiously in solution.
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Current Protocols in Protein Science
crude sample, particulate/haze laden
haze-removal steps: clarify by precipitation:
pH shift, 2 to 4 pH units up or down
salt (NaCl) addition plus pH shift
precipitating agents, limited amount, to precipitate particulates but not proteins in solution
clarify by centrifugation (500 –1000 × g) or filtration over filter aid on Whatman no. 1 paper; use pressure membrane (high MWCO) filtration if necessary
particle-free protein in solution: assay protein content and bioactivity; adjust total protein concentration (by pressure membrane ultrafiltration using low to medium MWCO to remove water if necessary) to bring it into range of 0.1% to 2% total protein (depending on precipitative technique to be used); see Table 4.5.1
perform pilot experiments for optimizing one of the precipitative techniques in Table 4.5.1
Figure 4.5.1 First steps and options for clearing particulates—e.g., haze and cell walls—from crudes before precipitating proteins. Limited amounts of precipitating agents may be effective in removal of haze from proteins. However precipitating agents for haze removal should not reach lower threshold concentrations for starting precipitation of sought-for proteins. Abbreviation: MWCO, molecular-weight cutoff.
Often it is best not to overconcentrate—i.e., not to attempt to remove nearly all of the water but to get the sought-for proteins to >1% to 2% weight concentration—before precipitating. Unwanted materials such as cell debris, polysaccharides, glycoproteins, and lipoproteins, may precipitate out early, coprecipitate desired proteins, or otherwise become difficult to deal with when they are overconcentrated. In some instances it may be the best choice to dilute samples with a buffer that can also shift pH, then carry out the concentration step. A general starting range of desired protein concentration (clear of haze) is 0.01% to 2%. Pilot Experiments After the protein sample has been clarified and concentrated (if needed), aliquots of haze-free sample should be placed in small test tubes or 2-ml plastic centrifuge tubes. A series (Fig. 4.5.3) or a 3 × 3 or a 4 × 4 grid of tubes (Fig. 4.5.11) are used. At first, buffers should be added to the aliquots of sample in rather large steps, at intervals of 1 or even 2 pH units. As the procedure is developed, much narrower pH intervals will be used to zero in on the optimum pH, depending on where bioactivity and protein-specific activity assays locate sought-for proteins in each tube. Likewise, in the first pilot experiments one may make rather large, well-spaced strides in concentrations of precipitating agents added— perhaps 5- to 10-fold differences between pilot samples to locate the neighborhood of precipitation versus nonprecipitation for chosen agents. Then, in a second series, more narrowly spaced conditions should be used to get smoother control over precipitation.
Extraction, Stabilization, and Concentration
4.5.3 Current Protocols in Protein Science
Supplement 7
Protein precipitation and coprecipitation frequently are quite cooperative (in the equilibrium sense of the term) like other types of phase changes—i.e., protein precipitations may tend to be all-or-none processes. Therefore good control sometimes can be obtained only within small ranges of conditions, such as in variation of precipitating agents. However, the neighborhood or range of parameters where that can be achieved may best be located by the first, wide-ranging set of experiments. Pilot experiments should make it possible to decide whether the process shall be one of first precipitating out desired proteins, as opposed to first taking out unwanted proteins and perhaps other materials and then bringing down desired proteins in a second stage. Investigation of optimum temperatures is usually useful and even mandatory. Pilot samples may be kept in refrigerators, at room temperature, and at still higher (heat-shock) temperatures, to find the best temperatures for both protecting and precipitating proteins. Protein precipitation, even when optimized, is sometimes nearly as dependent on rates (kinetics) as on equilibria (thermodynamics). In practice, therefore, attention must be paid to controlling the time of incubation of pilot samples under the various conditions. Strong precipitating agents (synthetic polyelectrolytes are leading examples) may form coprecipitates within a few seconds, but be irreversible. Intermediate and even quite weaker precipitants may take hours to develop protein coprecipitates but afford more controllability and reversibility. Choice of Procedure Table 4.5.1 lists ten possible techniques for protein precipitation. Choice of which to use likely depends on how easy or difficult it may be to reverse precipitation to recover the protein(s) and discard excess precipitating agent. For most uses to which precipitated proteins are put—e.g., nutrition, therapy, or simply for assay—the proteins must be freed of precipitating agents. The quite powerful agents, such as synthetic polyelectrolytes, can be difficult or slow (or both) to remove and release from desired proteins. Consequently, synthetic polyelectrolyte coprecipitation may best be used for first bringing down unwanted proteins (i.e., where those precipitates are to be discarded). Remaining proteins in the supernatant may then be captured using a less aggressive but more controllable, reversible precipitating agent from the list in Table 4.5.1 (e.g., Basic Protocols 1 to 5 or 8 to 10). Removal of Nucleic Acids
Selective Precipitation of Proteins
Crude proteins from cell cytosols, especially from bacteria and yeasts, may be laden with nucleic acids (RNA and DNA). Nucleic acids may interfere with protein isolation and contribute to haze and colloidal character. It is useful to remove nucleic acids in the early stages of crude workup. Techniques for removal of nucleic acids also help remove some kinds of particulates when the particulates have much anionic character—e.g., many cellular and subcellular membranes. There are two general methods for removing nucleic acids—i.e., precipitation using polyamines (UNIT 1.3) and using positively charged anionexchanger resins (UNIT 6.1). Nucleic acids are polyanions that are charged negatively and rather densely with phosphate groups. Naturally occurring “protamines” (Felix, 1960; Budavari, 1989), which are proteins themselves, but very cationic proteins, thoroughly coprecipitate most nucleic acids and ribosomes. However, overshooting the endpoint— i.e., adding more than enough protamine proteins to remove the nucleic acids—may leave excess protamines in samples, contaminating the sought-for proteins and perhaps interfering with their isolation downstream. Accordingly, it may be necessary to determine the endpoint for stopping protamine addition using a suitable spectroscopic technique for nucleic acid content (see UNIT 7.2). Synthetic polyethyleneimine (PEI; Sigma) and syn-
4.5.4 Supplement 7
Current Protocols in Protein Science
thetic polyamino butane compounds (spermine; Sigma) also precipitate nucleic acids, and are useful for large samples (milliliter to liter in volume). A detailed review of PEI (also called Polymin by some vendors) for nucleic acid precipitation is found in Jendrisak (1987). Between 20 and 80 µl of 5% PEI, added per ml of crudes containing 4% (dry weight) nucleic acids, nearly completely removes nucleic acids. This procedure should be carried out at a pH of ∼7.5 to 8, where PEI amine groups are fully cationic and nucleic acids are fully anionic. However, PEI is a very able precipitant for proteins with isoionic points below pH 7. If the sought-for protein coprecipitates with both PEI and nucleic acids, as some enzymes do, pH variation and salt extraction (e.g., with NaCl; Jendrisak, 1987) then becomes necessary to free such enzymes from the PEI agglomerate. Anion exchangers may also be used to remove nucleic acids and have an advantage and a disadvantage compared to polyamines used in solution. The advantage is that an excess of solid exchanger may adsorb less of the sought-for proteins than an excess of such reagents as PEI in solution. The disadvantage is that—because solid resins can only adsorb very large macromolecules like nucleic acids at the resin surface (not in its interior)— rather large quantities of resin may be required. The Amberlite IRA-400 series of resins (Sigma) and Dowex-1 anion exchangers (Sigma) are adsorbants for nucleic acids in neutral pH ranges. The DEAE-celluloses (Sigma), which are quaternized amines and therefore cationic, also are capable of adsorbing nucleic acids to remove them from crudes preliminary to isolating the proteins from the crudes. See Basic Protocol 2 for directions on washing and conditioning such resins prior to use. SELECTIVE PRECIPITATION BY SALTING OUT Ammonium sulfate is the best, first-choice salt for initial development of a salting-out program to precipitate sought-for proteins (see Fig. 4.5.2). Other salts may be chosen, and frequently they yield successful results. However because of sulfate’s kosmotropic and protein-molecule exclusionary powers (see Commentary), ammonium sulfate is most preferred and best understood. Sulfate salting out may be both thermodynamically limited and rate-limited. If precipitation of proteins does not occur quickly after addition of estimated amounts of sulfate salt, it may be best to wait for a period, perhaps a few hours, to allow time for precipitates to form. This becomes necessary for dilute protein solutions—less than ~1% in overall concentration. For several mammalian enzymes, the threshold for protein precipitation and solubility—i.e., the visibly seen development of precipitation versus proteins remaining dissolved (apparent equilibrium)—occurs at ~2 to 5 mg/ml protein in 2 to 3 M ammonium sulfate, at neutral pH (Scopes, 1987).
BASIC PROTOCOL 1
The pH and temperature also are likely to be important. If the isoelectric pH of the sought-for protein is known, it may be used as the center point around which to vary pH, in increments of ~0.5 to 1 pH unit, in initial searches for optimum conditions. A number of proteins, peptide antibiotics, and drugs are best salted out as coprecipitates with SO42– as a bound counterion. Hence their optimal pH for salting out may be acidic with respect to their isoelectric point. Figure 4.5.2 illustrates the main steps in salting out, starting with mixtures of proteins in solution without particulate matter. In both this protocol and Alternate Protocol 1, the sample is first carried through all steps on a small scale for a pilot experiment to determine optimal conditions. The procedure is then scaled up to purify the protein sample, using those optimal conditions. Figure 4.5.3 illustrates the procedures for such a pilot experiment. Extraction, Stabilization, and Concentration
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Materials Crude protein solution of interest, particle-free (see discussion of Clarification in Strategic Planning) Appropriate pH buffer Ammonium sulfate Additional reagents and equipment for dialysis (UNIT 4.4 & APPENDIX 3B) 1. Dialyze protein samples against a pH buffer or a pH buffer/ammonium sulfate mixture having a sulfate concentration below that needed to start precipitation (see UNIT 4.4 & APPENDIX 3B). By carrying out dialysis, three objectives may be achieved at the expense of an added step. First, it is possible to dialyze out unwanted lower-molecular-weight materials—e.g.,
simplest procedure; one-stage
more elaborate procedure
adjust crude protein sample to a single, preset pH
adjust crude protein sample to a total protein concentration between 0.05% and 2%; adjust pH to optimum value as determined by pilot experiment
add (NH4)2 SO4 in various concentrations as aliquots from a 100% saturated stock or as crystalline salt
time to develop visible copious precipitate: 2.5 to 4 hours, depending on temperature
harvest precipitate; dialyze to remove excess salt; assay activity; measure total protein
Selective Precipitation of Proteins
add salt to precipitate 20% to 60% of the total protein
assay precipitate and supernatant for protein activity
active protein in precipitate: discard supernatant
active protein in supernatant: discard precipitate
redissolve precipitate in buffer to shift pH as needed for optimal salting out
increase salt concentration; shift pH as needed
increase salt to precipitate out the majority of the proteins; dialyze away salt; measure total protein and total activity
harvest precipitate; redissolve in the operating buffer or in the assay running buffer; assay
Figure 4.5.2 Two-salting out protocols, single-stage and two-stage, starting from a particle-free aqueous crude preparation (e.g., cell or tissue extract or centrifuged fermentation broth).
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sugars, amino acids, and components of fermentation broth or cell extract. Second, dialysis makes it possible to bring the protein sample smoothly to the prescribed pH. Finally, dialysis allows the investigator to adjust the protein sample to a known beginning salt concentration—that of the dialyzing buffer. Such dialysis may be carried out in dialysis bags (cellulose acetate tubing), or by pressure membrane concentrators with membranes having nominal molecular-weight cutoff points below that of the sought-for protein.
2. Set up a pilot experiment (see discussion of Pilot Experiments in Strategic Planning) to determine the optimal protein concentration, pH, salt concentration, temperature, and incubation time. See Critical Parameters and Troubleshooting for guidelines on how to vary these parameters. See Figure 4.5.3 for illustration of a small-scale pilot experiment. pH adjustments generally are performed by addition of acid (HCl), base (NaOH, KOH, or ammonia), or a buffer. At the same time, it is usually necessary to avoid adding too much overall volume, so the buffering capacity of the sample itself must be taken into account. If feasible, it is generally best to adjust pH of proteins with ~0.1 or 0.2 M HCl, or comparable concentrations of aqueous ammonia. Trying to move the pH appreciably with buffers, if dilute buffers must be used, increases overall volumes and decreases salt concentration. That, in turn, may require the use of considerably more salt to reach a specified concentration, in contrast to operations where pH has been adjusted by use of HCl or NaOH. CAUTION: The protein solution should be stirred well during addition of strong acid or strong base. Beware of changes in temperature.
3. Add ammonium sulfate to the optimal concentration (see Critical Parameters and Troubleshooting). Incubate (for the optimal period of time) until a precipitate forms. The salt may generally be added as a saturated solution. However, there is an advantage to adding the salt in solid form, as increases in volume are thereby lessened. There are few indications that addition of solid ammonium sulfate does harm to proteins already in solution.
increasing ammonium sulfate concentration
saturated or subsaturated ammonium sulfate
mix
starting volume of sample plus pH-adjusting buffer completion visible protein precipitate
Figure 4.5.3 Salting out—small-scale investigation. Maximum ammonium sulfate concentration ~4 M. Development of precipitate depends on the protein and unwanted materials which sometimes precipitate out in samples or fractions, apart from the sought-for protein. Other adjustable parameters are pH, temperature, and concentration of starting sample.
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4. Collect the precipitate by simple decantation (pouring off the supernatant). Ammonium sulfate precipitates are usually dense and settle out readily. However, if simple settling and decantation will not work, a slow-speed centrifugation (10 to 100 × g) should easily suffice.
5. Redissolve the precipitate in a buffer suitable for the next step (e.g., running buffer for electrophoresis or assay buffer). High concentrations of K+ and other salts are not compatible with SDS-PAGE and isoelectric focusing. K+ salts of SDS are poorly soluble. Large amounts of ammonium ion grossly interfere with some total protein assays (UNIT 3.4). Therefore, excess salt should be dialyzed away by simple cellulose-tubing dialysis (UNIT 4.4 & APPENDIX 3B) or by pressure ultrafiltration (UNIT 4.4).
6. Perform a bioassay for the protein of interest. If the sample is to be diluted considerably for the bioassay, the redissolved ammonium sulfate may not need to be dialyzed beforehand. However, if appreciable salt interferes (as in SDS-PAGE and total protein assays), the assay may require a dialyzed sample input. ALTERNATE PROTOCOL 1
SELECTIVE PRECIPITATION BY STEPWISE SALTING OUT A stepwise salting-out procedure is used to fractionate a crude mixture into two portions, only one of which retains the desired protein. The result is a partial purification of the desired protein from a crude mixture of proteins; this may improve the extent of recovery in subsequent purifications. In this protocol, as in Basic Protocol 1, the sample is first carried through all steps on a small scale for a pilot experiment to determine optimal conditions. The procedure is then scaled up to purify the protein sample, using those optimal conditions. Figure 4.5.3 illustrates the procedures for such a pilot experiment. 1. Adjust the concentration and pH of the crude protein solution to the optimal values as determined by a pilot experiment (see discussion of Pilot Experiments in Strategic Planning). See Critical Parameters and Troubleshooting for guidelines on how to vary these parameters. See Figure 4.5.3 for illustration of a small-scale pilot experiment.
2. Add ammonium sulfate at an appropriate concentration. Incubate until precipitate forms. 3. Collect the precipitate and supernatant and assay for the appropriate bioactivity in each. 4a. If the active fraction is the precipitate: Redissolve the material in a buffer suitable for assay or the next general procedure. If the next steps are not to be performed immediately, the precipitate need not be redissolved. It should be stored with its salt because salts, especially ammonium sulfate, are protective and stabilizing.
4b. If the active fraction (or part of it) remains in the supernatant: Increase salt concentrations by 5% to 10% and perhaps change pH by ~0.5 to 1 unit, in renewed pilot experiments and in larger-scale precipitation. 5. Collect the precipitate and supernatant and assay for the appropriate bioactivity in each. Selective Precipitation of Proteins
6. Repeat steps 4b and 5 until the protein of interest is obtained in as clean a preparation as possible.
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SELECTIVE PRECIPITATION BY ISOIONIC PRECIPITATION: COLUMN METHOD
BASIC PROTOCOL 2
Proteins frequently are least soluble and most precipitable when they are isoionic. In the isoionic, salt-free state, protein molecules are in their most compact, least hydrated conformation—a phenomenon that is closely related to the condition of proteins at their isoelectric point. The distinction between isoionic and isoelectric properties is drawn in detail by Tanford (1961). Deionization using a column (this protocol) or dialysis (see Alternate Protocol 2) aim at rendering proteins isoionic to precipitate them. Two important parameters determine solubility of many proteins: solution pH with respect to each protein’s isoionic point (pI) and the low salt concentration (zero to 0.1 to 0.2 M salt). A number of proteins—e.g., β-lactoglobulin—are sharply dependent on these parameters. Accordingly, if these parameters are well controlled and carefully adjusted, the solubility/precipitability behavior of a protein will be a practical basis for scaleable, selective isolation of the desired protein. The column method used in this protocol is appropriate only for proteins that remain soluble at their isoionic point. In addition to adjusting proteins to their isoionic pH, the general method described here, using mixed-bed resin deionization, is able to strip away all salts from proteins. Salts, even in small concentrations (<0.05 M in many cases) often have large effects on protein solubilities and therefore on precipitability. Inorganic salts tend to “salt in” many proteins, thus enhancing their solubilities. Accordingly, sharp control and thorough removal of salts as desired is an important part of the general technique of isoionic precipitation. Figure 4.5.4 shows a Dintzis deionization column with a lower, main mixed bed of anion-cation exchange resin beads in their respective OH− and H+ form. Two categories
guard band: anion exchanger resin in acetate form, IRA-400Ac– (4 to 5 cm) guard band: cation exchanger resin in ammonium form, IR-120NH4+ (3 to 4 cm) cation exchanger in its H + form (e.g., Amberlite IR-120 resin beads)
main mixed-bed exchanger anion exchanger in its OH– form (e.g., Amberlite IRA-400 resin beads)
Figure 4.5.4 Deionization column, Dintzis design, described in Edsall and Wyman (1958). Passage through this column renders the protein salt-free and automatically adjusted to the protein’s isoionic pH. This column is used for proteins that remain soluble at their isoionic point.
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of salt ions are removed by the mixed bed: (1) the supporting electrolyte—i.e., the free salt in the solution; and (2) salt ions that are counterions, which are in effect bound to the protein if the protein is not isoionic. If the pH of the starting solution is below the isoionic point, the protein is a cation with anion counterions, commonly Cl−. If the pH is greater than the isoionic point, the protein is a net macroanion with cationic counterions, commonly K+ or Na+. In the mixed bed, cationic and anionic salt ions are exchanged for H+ and OH−, respectively, going on to simply form water. Protein molecules deprived of all mobile counterions are quantitatively forced to the single pH at which they can exist without mobile counterions or protein-bound ions—i.e., the isoionic pH. Therefore the effluent of the column is isoionic protein at the isoionic pH, in plain water. The net salt exchange and entrapment processes are similar to conventional desalting by resin ion exchangers, except the effluent is buffered by the isoionic protein and adjusted to its pH. In the original Dintzis column, illustrated in Figure 4.5.4, guard exchanger bands may be positioned at the top of the column above the main bed. They exchange incoming salt cations and anions (or protein counterions) for NH4+ ion and acetate ion from the resins. This is intended to protect incoming proteins from direct mixing with the equivalent of strong acid and strong base, respectively, from the mixed-bed strong exchange resins in their respective H+ and OH− form. The NH4+ and acetate ions are exchanged and removed in the main bed, and the isoionic protein sample flows through. Materials Cation exchanger: Amberlite IR-120H+ resin (Sigma or Aldrich) Anion exchanger: Amberlite IRA-400Cl– resin (Sigma or Aldrich) ~0.5 and 1 M NaOH or KOH 0.5 to 1 M HCl 0.2 M acetic acid 0.2 M ammonium hydroxide Crude protein solution of interest, particle-free (see discussion of Clarification in Strategic Planning) Plastic beakers Large Buchner funnel Whatman no. 1 filter paper Chromatography column of appropriate size 1. Determine exchange capacity required. Necessary capacity for deionizing and trapping salts depends on the number of counterions and therefore on pH of the protein solution, concentration of the supporting electrolyte or buffer, and overall volume. The following is an example of the calculation method used. Assume that 50 ml of 5% serum albumin in 0.2 M acetate buffer, pH 4, is to be deionized. On the basis of the titration curve (with a pI of 5.0 for albumin; see UNIT 8.2 for discussion of titration curves), ~35 counterions (e.g., Cl−) are present per protein molecule (mol.wt. 67,000), amounting to ~1.3 meq of ions. The buffer contains ~10 meq of total acetate, along with 10 meq of Na+, which were counterions for the buffer’s acetate. Hence, there are a total of 22 meq of ions to be exchanged. The mixed bed is calculated to hold 1 meq/ml of settled mixed-bed resin, so ~12 ml of resin, minimally, is required. Thus a 2 × 30–cm bed including 80 to 90 cm3 of mixed-bed resin is in good excess (by a factor of ~4) of the minimum required to strip out all inorganic ions, buffer, and protein counterions. See Critical Parameters for additional discussion.
Selective Precipitation of Proteins
2. Place IR-120H+ cation-exchange resin in a plastic beaker, cover with an excess of ~0.5 M NaOH or KOH, and incubate ~20 min to convert resin to the Na+ or K+ form. Transfer to a large Buchner funnel containing Whatman no. 1 filter paper, wash with water, then transfer to a beaker and cover with an excess of 0.5 to 1 M HCl. Incubate
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20 min, then transfer to a Buchner funnel containing filter paper and wash with water again. Repeat this exchange cycle at least one more time before mixing the exchange resins. This full-cycle treatment removes undesirable low-molecular-weight hydrolysis fragments from resin polymers, which otherwise would bind strongly to many proteins and elute with them when proteins are passed through the column. Each step requires ~20 min, to allow penetration, diffusion, and exchange of ions with the interiors of the resin beads. More detailed specifications are given in full treatises describing exchangers, such as Kressman (1957) and in the manufacturers’ literature.
3. Place IRA-400Cl– resin in a plastic beaker, cover with an excess of ~1 M NaOH or KOH, and incubate ~20 min to convert resin to the OH– form. Transfer to a large Buchner funnel containing Whatman no. 1 filter paper and wash with water to remove all excess OH– ion. The resin is now IRA-400OH−. It should be kept cold and should not be exposed to air during long periods of storage because the CO2 in air slowly neutralizes the IRA-400OH− to a bicarbonate form. The resin should be taken through this full exchange cycle within a few days prior to use. Anion exchangers, both old and new, which are made of quaternary amine polymers, self-hydrolyze and slough off organic amines, especially when stored in their hydroxyl form (e.g., IRA-400OH−). Accordingly, IRA-400 in whatever starting form (Cl− or OH−), needs be taken through a full exchange cycle. Each step requires ~20 min, to allow penetration, diffusion, and exchange of ions with the interiors of the resin beads. More detailed specifications are given in full treatises describing exchangers, such as Kressman (1957) and in the manufacturers’ literature.
4. Prepare resin for preexchange guard bands as follows. React IRA-400OH− (from step 3) with 0.2 M acetic acid, then wash with water to prepare anion-exchanger IRA-400Ac−. React IR-120H+ (from step 2) with 0.2 M ammonium hydroxide, then wash with water to prepare IR-120NH4+. 5. Mix and load ion-exchange resins into columns (also see Fig. 4.5.4). See Critical Parameters for guidelines on preparing the mixed-bed resin. Some precautions need be taken to prevent unmixing—i.e., formation of separated bands of cation and anion exchangers in their respective H+ and OH− forms—when filling the column. The density of IR-120H+ is appreciably larger than that of IRA-400OH−. The resins used for the guard bands have a similar disparity in density. If large volumes of premixed resins are dropped through several centimeters of water, they unmix on the way down. Hence, mixed-bed resins should be dropped in small increments, with forward flow of water, to prevent unmixing. The same general considerations pertaining to procedures such as packing and loading chromatographic columns, dead-volume minimization, and prevention of channeling that are described in UNIT 8.4 apply to this column technique.
6. Determine flow rate required. If there is adequate capacity in the mixed bed to trap all salts, a 2 × 30–cm column deionizes 50 to 100 ml of a 1% to 5% protein solution in ~3 hr. Suitable flow rates average 0.5 to 1 ml/min.
7. Perform column separation on crude protein solution. The first cut, ~20 to 30 ml of eluate, will be quite dilute as the protein solution displaces water in the column, and may be set aside.
8. Perform bioassay for the protein of interest. 9. Recycle used resins. Mixed-bed resins previously used for deionization—e.g., of serum proteins—are good general binding agents and can be reused. After being used for deionizing a protein, the
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resin should be flushed with 0.2 M HCl to remove any protein and to prevent bacterial growth. Mixed-bed resin pairs of the IRA-400 and IR-120 series, as well many of the Dowex resin pairs, separate well from one another in saturated KCl. Crystalline KCl can be added to a slurry of used mixed-bed resin in water. As KCl saturation approaches, the anion exchanger that is least dense—i.e., the Cl− form—floats upward. The cation exchanger in K+ form sinks. The anion-exchange resin slurry is decanted off the top, away from the cation exchanger, isolating both resins. These may then be individually recycled back to their respective OH− and H+ forms, whereupon they may again be mixed as needed (see steps 2 and 3). ALTERNATE PROTOCOL 2
SELECTIVE PRECIPITATION BY ISOIONIC PRECIPITATION: DIALYSIS METHOD One of the older means of rendering proteins salt-free or nearly salt-free (i.e., isoionic) is dialysis (also see UNIT 4.4 & APPENDIX 3B). However, two problems frequently arise with conventional dialysis: (1) when appreciable amounts of protein are present, osmotic effects result in swelling of dialysis bags as salt diffuses outward; and (2) often it is quite uncertain where the isoionic point is, even if it is feasible to deionize by dialysis against a buffer. The resin deionization method (see Basic Protocol 2), sometimes called Dintzis’ method (Edsall and Wyman, 1958), automatically adjusts a protein precisely to its isoionic pH without prior knowledge of it. Proteins that precipitate immediately when deionized are troublesome in the column method described in Basic Protocol 2 and Figure 4.5.4. Precipitates plug the column and pose the problem of separating precipitated protein from resin beads. β-lactoglobulin and a number of other proteins behave in this way; they have solubilities very sensitive to low salt conditions near the isoionic pH. This protocol (also see Fig. 4.5.5) describes a means of overcoming this problem. Proteins to be deionized and precipitated in isoionic form are confined in dialysis tubing with a molecular weight cutoff that will retain the protein of interest and the mixed-bed resin is placed outside as a slurry of 40 to 60 g of mixed-bed resin (50 to 80 ml of wet resin; see Basic Protocol 2) per 200 to 400 ml of dialyzing solvent. Salt ions and protein counterions exchange through the membrane and are trapped outside in the exchanger resins. After exchange is completed, precipitated protein in the dialysis tubing is recovered by centrifugation of the bag’s contents. This general technique requires several hours. Salting out and diffusion slow as the protein concentration inside the dialysis bag decreases; hence this method is slower than the flow-through column method (see Basic Protocol 2).
Slurry of mixed-bed exchanger resins
rocking
precipitated protein
Selective Precipitation of Proteins
dialysis bag containing protein solution
Figure 4.5.5 Deionization by dialysis. When proteins are insoluble and precipitate at their isoionic point, deionization is accomplished by placing dialysis tubing containing the protein sample in a slurry of mixed-bed resin exchangers (i.e., a mixture of IRA-400OH− and IR-120H+) and incubating with rocking. Proteins insoluble at their isoionic point precipitate inside the bag. The mixed-bed exchange resins remove free salts, forcing the protein to its isoionic pH. This method is considerably slower than the column method.
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SELECTIVE PRECIPITATION USING A TWO-CARBON (C2) ORGANIC COSOLVENT
BASIC PROTOCOL 3
Protein precipitation using very cold ethanol or acetone is a method that was developed over a century ago. Both of these cosolvents are still frequently used. Ethanol surpassed acetone for precipitating proteins at the time E. J. Cohn, and co-workers at Harvard perfected the Cohn fractionation (Cohn et al., 1946) for large-scale preparation of plasma proteins—albumin, fibrinogen, prothrombin, and γ-globulins. The Cohn procedures are still employed, although modified. One of the advantages ethanol has over acetone pertains to fire hazard. Ethanol is less volatile than acetone and generates less flammable vapor under otherwise similar conditions. Both cosolvents, when handled in in >0.1-liter amounts, should be used in ventilated rooms. Materials Crude protein solution of interest, particle-free (see discussion of Clarification in Strategic Planning) Buffering system Ethanol Acetone Centrifuge 1. Adjust the crude protein solution of interest to the selected pH. If the operating buffer (used as the solvent in the separation) or pH-calibration buffers (used to check pH-meter calibration) are to be selected for low-temperature operation, the tables by Alner et al. (1967) and Perrin and Dempsey (1974) give formulations for several buffers together with their pH values at temperatures in the 0°C range.
2. Arrange for good stirring and good metering to control mixing of organic cosolvents with the aqueous proteins. It is necessary to suppress local high concentrations of C2 cosolvents. It is also necessary to suppress the heat generated by mixing of cosolvents, especially of ethanol with water, to avoid undue denaturation and temperature increases greater than ~2°C above the set temperature. Dilution of organic cosolvents with water or with aqueous running buffer before they are added to the protein solution may repress generation of heat during mixing and may also repress formation of large local concentrations of cosolvents at the point at which they are mixed with aqueous protein. However excessive dilution of organic cosolvents may add to the final volumes necessary to reach precipitation thresholds. A 3% to 8% (v/v) volume of water in organic cosolvent is a suitable dilution for achieving protein precipitation before overall volumes become excessive.
3. Cool the two components—i.e., protein in aqueous buffer and cosolvent system to be used—to the running temperature. Do not allow temperatures to rise more than 2°C above the running temperature. Lower temperatures can be obtained by using an auxiliary refrigeration unit pumping coolant through an exchanger coil or by incubating C2 cosolvents in the deep freeze. Salt/ice baths are capable of reaching approximately −10°C.
4. Slowly mix the the cosolvent system into the aqueous protein solution while monitoring the mixture with a thermometer. If the temperature rises much during cosolvent (or cosolvent/buffer) addition, slow the rate of solvent addition down to let the cooling system remove excess heat. Predilution of organic cosolvents (see annotation to step 2) may be helpful here.
5. Collect precipitate by centrifugation or allow precipitate to settle out. Remove supernatant and redissolve precipitate by adding water or suitable aqueous buffer.
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Organic cosolvents admixed with aqueous systems usually decrease the density of the solvent mixture, which may aid settling or centrifugation of precipitated protein. However, the centrifuge should also be cooled down to the precipitation running temperature. Some centrifuges become surprisingly warm, depending on the vintage and specific features. A test of the actual centrifugation temperature is advisable; this is done simply by immersion of a thermometer in centrifuge vessels after a run.
6. Perform a bioassay for the protein of interest. BASIC PROTOCOL 4
SELECTIVE PRECIPITATION USING C4 AND C5 ORGANIC COSOLVENTS Figure 4.5.6 outlines the technique for three-phase partitioning (TPP) using t-butanol in conjunction with ammonium sulfate (Lovrien, et al., 1987). Some C5 and C6 cosolvents— e.g., alcohols such as pentanol—may be useful in some cases. However C5 and larger cosolvents tend to be so insoluble in water as to be of limited use. The cosolvent used here, t-butanol, is infinitely soluble in water alone, but in the presence of large concentrations of ammonium sulfate it forms a second phase. Simultaneously, proteins are precipitated, forming a third phase intermediate between the lower (aqueous) and upper (organic) liquid phases (see Fig. 4.5.6). The various phases are mutually saturated with respect to one another. The protein precipitate is usually well developed and easily retrieved by low-speed centrifugation. TPP behavior depends on protein molecular charge (precipitated products are often sulfate salts of protein). Therefore, the pH of the aqueous phase is an important parameter for modulating protein precipitation, partitioning, and extraction of unwanted compounds. When proteins have been extracted from their original source using detergents, TPP or related methods such as Morton’s n-butanol extraction technique (Morton, 1950) are frequently effective in removing detergents, lipids, and pigments from the sought-for protein.
t-butanol layer third phase: precipitated enzymes add t-butanol, mix, let settle
aqueous crude enzymes. Adjust to suitable pH and salt content.
Selective Precipitation of Proteins
aqueous phase
perform semimicro-scale centrifugation to collect protein
Figure 4.5.6 Procedure for precipitation of proteins from aqueous solutions using t-butanol. This figure applies to both large-scale precipitation and small-scale pilot experiments. The t-butanol layer develops when adequate salt is present in the aqueous solution. Both the t-butanol layer and the aqueous lower phase act as extraction solvents as well as partitioning systems.
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Materials Ammonium sulfate Buffering system Crude protein solution of interest, particle-free (see discussion of Clarification in Strategic Planning) C4 organic cosolvent (e.g., t-butanol, analytical grade) 1. Add ammonium sulfate and buffer to adjust the salt concentration and pH of the crude protein solution of interest appropriately. The amount of ammonium sulfate used is determined by pilot experiments and is in general less than that used for simple salting out (20% to 100% saturation); i.e., the amount added is less than sufficient to induce precipitation.
2. Add ~0.5 to 1 ml C4 organic cosolvent per ml crude protein solution, mix, and incubate until three phases form. If only technical grade t-butanol is available it may be either recrystallized (the melting point of pure t-butanol = 25.5°C) at room temperature, or distilled (the boiling point = 82°C).
3. Transfer the middle phase containing the precipitated protein to a centrifuge tube of volume approximately twice that of the protein precipitate. Centrifuge at 10 × g, 4° to 25°C. See Critical Parameters for discussion of the separation of phases.
4. Redissolve the precipitate in an appropriate buffer and perform suitable bioassay. SELECTIVE PRECIPITATION USING PROTEIN EXCLUSION AND CROWDING AGENTS AND OSMOLYTES Protein exclusion and crowding agents and osmolytes are neutral, very water-soluble compounds—such as sugars, organic diols, and polymers—used in concentrations of 2% to 20% to precipitate proteins. They operate primarily by entropically driven “crowding” (Herzfeld, 1996), by preferential hydration (Timasheff, 1992), or a combination of both mechanisms. Such agents push proteins out of solutions in the mechanical/physical sense and in the thermodynamic sense. At concentrations greater than ~5% by weight, exclusion agents and osmolytes occupy so much volume on the molecular scale that they leave less room in solution available for protein molecules.
BASIC PROTOCOL 5
Crowding and molecular-exclusion polymers that have been used for protein precipitation include polyethylene glycol (PEG), polypropylene glycol (PPG), polyvinyl alcohol (PVA), methylcellulose, dextran, and hydroxypropyldextran. Osmolytes and zwitterions used for protein precipitation include sucrose, 2-methyl-2,4-pentanediol (MPD), raffinose, maltose, sarcosine, and betaine. These compounds are readily available from suppliers such as Sigma and are able to protect and precipitate a number of proteins. Development of each protein-precipitation program requires attention to pH, temperature, and other factors that modulate parameters governing protein solubility and precipitability. The literature is scant concerning rate limitation of protein precipitation by these parameters. It is likely that in lower concentration ranges of proteins, neutral polymers, and osmolytes (<1% to 2%), precipitation rates may be slowed. Accordingly the general method should be treated as a rates (kinetics) problem when precipitates are not quickly formed. The sample is first carried through all steps on a small scale for a pilot experiment to determine optimal conditions. The procedure is then scaled up to purify the protein sample, using those optimal conditions. Figure 4.5.3 illustrates the setup for such a pilot experiment.
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Materials Crude protein solution of interest, particle-free (see discussion of Clarification in Strategic Planning) Appropriate pH buffer Precipitating agent: neutral polymer or osmolyte 1. Set up a pilot experiment (see discussion of Pilot Experiments in Strategic Planning) to determine the optimal concentrations of protein and precipitating agent as well as the optimal pH, temperature, and incubation time. In preliminary searches, use a series of test tubes as in Figure 4.5.3. On addition of components to each other, mixing or strong vortexing may be needed because some neutral polymers are highly viscous.
2. Prepare protein solution and solution of precipitating agent. Neutral polymers should be dissolved in water before adding them to aqueous proteins. Moderate heating may speed dissolving. It is poor practice to attempt addition of dry polymers directly to dissolved proteins as the polymers tend to form gummy masses that dissolve very slowly. Mechanisms here depend on molecular crowding; hence concentrations of proteins and dissolved polymers to be added must be maximized. These components dilute each other when mixed. If protein samples need pH adjustment before the mixing step, use buffers in fairly concentrated form—or even 0.2 to 1 M acid or base if feasible—to keep volumes minimized and component concentrations maximized.
3. Mix protein solution and solution of precipitating agent. Incubate (for the optimal period of time) until a precipitate forms. If mixtures of proteins are very crude and only one or two proteins are sought after, it may be preferable not to drive the precipitation too hard. In such cases it may be best to accept more slowly developed precipitates to maximize the amount of unwanted protein and excess precipitant left behind in the supernatant and the amount of sought-after protein captured in the precipitate.
4. Collect the precipitate. Trapped and coprecipitated neutral polymers in precipitates may need be “reversed” or split away later to release proteins free of polymers. How this can be done depends on the chemistry of the systems, and perhaps also on the chromatographic properties of the precipitated proteins. In the case of dextrans, modest amounts can be degraded by dextranase enzyme. Coprecipitated proteins may be captured on ion-exchange matrices, whereupon neutral polymer may be eluted and the protein displaced from the exchanger using a pH shift and/or salt (see UNIT 8.2).
5. Perform bioassay for the protein of interest. BASIC PROTOCOL 6
Selective Precipitation of Proteins
SELECTIVE PRECIPITATION USING SYNTHETIC AND SEMISYNTHETIC POLYELECTROLYTES Four categories of polyelectrolytes can precipitate proteins. First, there are the synthetic polyelectrolytes—e.g., vinyl polymers such as polyacrylate (PAA) and polymethacrylate (PMA); acid salts (polyanions); and polyethyleneimine (PEI, a polycation). Next, there are the semisynthetic polyelectrolytes—carboxymethylcellulose (CMC), sulfated cellulose, and sulfated dextrans. The third category includes the protamines (very basic proteins), and the fourth includes naturally occurring sulfate polysaccharides—e.g., heparin and chondroitin sulfate. Heparin and related biochemical polyelectrolytes are not generally used on a large scale because of their cost; they are used in research for precipitating narrow classes of proteins such as β-lipoproteins. For a recent, short review
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Table 4.5.2
Synthetic Polyelectrolyte Coprecipitating Agents for Proteins
Polyelectrolyte
Proteins preciptated
References
Polyacrylate Polyacrylate Polyacrylate
Lysozyme, hemoglobin Eight industrial enzymes Ferrihemoglobin, catalase, bovine serum albumin Cellulase enzymes Serum albumin, hemoglobin Recombinant (RecA) protein RNA polymerase II β-galactosidase fusion protein Restriction endonucleases β-lactoglobulin, α-lactalbumin Fibrinogen Protein A Trypsin, RNase A, lysozyme Serum albumin
Sternberg and Hershberger (1974) Sternberg (1976) Berdick and Morawetz (1954)
Polymethacrylate Polymethacrylate Polyethyleneimine Polyethyleneimine Polyethyleneimine Polyethyleneimine Carboxymethylcellulose (CMC) Polylysine Copolymer Eudragit Tri-block polyampholytes Polydimethyldiallylammonium chloride
Whitaker (1953) Morawetz and Hughes (1952) Shibata et al. (1981) Jendrisak and Burgess (1975) Niederauer et al. (1994) Bickle et al. (1977) Hidalgo and Hansen (1971) Sela and Katchalski (1959) Mattiasson and Kaul (1994) Patrickios et al. (1994) Dubin et al. (1987)
of polyelectrolytes for protein precipitation see Singh (1995). Table 4.5.2 lists a number of polyelectrolytes in each category and cites references regarding their use. Figure 4.5.7 outlines the main steps for polyelectrolyte-mediated protein precipitation. Frequently, the main practical problem with this technique is in reversing coprecipitation—i.e., dissolving the coprecipitate in a way that releases proteins and splits away the polyelectrolyte. In the case of polyacrylate and polymethacrylate, these polymers bind calcium and barium ions rather strongly. Thus protein/polyacrylate coprecipitates may first be shifted upward in pH, which renders proteins and polyelectrolytes negatively charged and helps them redissolve. After this, Ca2+ or Ba2+ is introduced to precipitate away the polyelectrolyte and release the protein. Several papers describing examples of the use of this technique are referred to in the Commentary (see Critical Parameters and Table 4.5.2). The general range of starting protein concentrations from which proteins may be coprecipitated in fair to good yield (at optimum pH) is ~0.05% to 3% (w/v) protein. The range of quantities of polyelectrolyte required, given a suitable kind of polyelectrolyte, tends to be less than the amount of protein. By weight, ~2% to 20% polyelectrolyte with respect to the protein is generally required. The variation between systems is large, partly because polyelectrolytes have large variations in charge density—e.g., completely ionized polyacrylic acid carries one anionic charge per 56 g polymer; carboxymethylcellulose, substituted on each glucose unit, carries one anion per 240 g polymer if completely ionized. Materials Water-soluble polyelectrolytes (Aldrich or Sigma) Crude protein solution of interest Test tubes Glass or plastic cuvettes Spectrophotometer
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1. Choose a polyelectrolyte. For proteins not previously investigated, first experiment with from two to four easily available, water-soluble polyelectrolytes. Polyacids (e.g., polyacrylic acids), and polybases (e.g., polyethyleneimine), can be titrated with NaOH, or HCl respectively, to two or three pH units, respectively, above or below the target protein’s isoelectric point (if known). Such polyelectrolytes are buffers themselves, as are proteins, over the pH range of acid/base titration. Hence, it is unnecessary to use additional conventional inorganic buffers to control pH. If polyelectrolytes and proteins are mismatched in pH before mixing, the mixture may shift in pH; the pH should be measured, if necessary, to determine H+ transfer on mixing the main components.
2. Prepare stock polyelectrolyte solutions. Initial stock polyelectrolyte solutions should be prepared in water in concentrations three to ten times the final coprecipitating concentrations. Dilutions will be performed in the succeeding steps.
polyelectrolyte: dissolve in water at a concentration, 5× to 10× larger than precipitating concentration
adjust pH, salt content
protein sample: free of particulate matter (see Strategic Planning) adjust pH, salt content
add polyelectrolyte to protein with metering and good mixing
observe turbidity by visual observation or spectrophotometry using near-UV or visible wavelengths
stop polyelectrolyte addition when coprecipitation ceases (end point)
centrifuge coprecipitate down
supernatant: analyze for total protein, activity
precipitate: attempt to reverse
analyze separated protein; recover polyelectrolyte
Selective Precipitation of Proteins
Figure 4.5.7 Principal steps in protein coprecipitation with polyelectrolytes. “Metered” addition is slow addition (with good stirring) of aliquots of polyelectrolyte stock to the protein sample near the threshold of precipitate formation as seen by eye or spectrophotometric turbidity determination.
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3. Render protein sample clear and nonparticulate (see discussion of Clarification in Strategic Planning). 4. Set up several test tubes on a small scale, adding ~2 ml sample and 0.05 to 1 ml of each polyelectrolyte stock solution at three or four different pH values to construct a matrix of polyelectrolyte/protein mixtures. Observe the point at which turbidity appears. The intent is to roughly locate (by visual comparison to detect turbidity) the coprecipitation starting point, the coprecipitation end point, the relative concentrations of polyelectrolyte and protein to be used, and the optimum pH. This enables construction of a rough phase diagram (i.e., a plot of turbidity versus variables such as relative amount of polyelectrolyte). Also see discussion of Pilot Experiments in Strategic Planning.
5. Set up a more finely grained pilot experiment as in the previous step, but carry out turbidity measurements in glass or plastic cuvettes. For scale-up, use controlled rates of polymer addition (i.e., adding polymer to the protein sample slowly in reproducible aliquots of stock solution near the threshold of precipitate formation), temperature control (water bath), and mixing using a magnetic stir-bar. Measure spectrophotometric absorbance in the 400- to 500-nm range and determine where overt coprecipitation begins. Sharply climbing absorbances mark the ranges where turbidity forms as a result of particulate coprecipitates. Coprecipitation is in large part a rate process as well as an equilibrium process. If the aim is to achieve selection of particular proteins, rather than precipitation of all proteins, the best strategy may to bring down some protein(s) in a first-stage separation of the coprecipitate, followed by a second stage in which polyelectrolyte is again added.
6. Carry out full-scale separation. Centrifuge to collect coprecipitate; retain the supernatant and analyze for total protein activity. 7. Reverse coprecipitation to free captured protein from synthetic polymers. This step should start with resuspension of particles in water or in buffers to shift pH to a position where polyelectrolyte and protein have similar charges—e.g., both anionic or both cationic. Separating out polyelectrolytes is often achieved by addition of metal ions (Ca2+ or Ba2+ in the form of chlorides) to carboxylate polyelectrolytes. An attempt may also be made to adsorb polyelectrolytes on bead resins of opposite charge—e.g, on DEAE cellulose for carboxylated polyelectrolytes. In developing a procedure for polyelectrolyte removal and release of sought-for proteins, measurement of supernatant protein concentration and/or activity if the protein is an enzyme should facilitate monitoring of protein release. Most of the polyelectrolytes in Table 4.5.2 have low A280 absorbances (and small molar extinction coefficients). Hence simple measurement of A280 should locate the conditions at which proteins are released into solution, provided there are no particulates present. However, such particulates cause very strong scattering and turbidity in the UV range, far beyond the absorbance of equivalent amounts of truly dissolved protein.
8. Perform bioassay for protein of interest and recover polyelectrolyte.
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BASIC PROTOCOL 7
SELECTIVE PRECIPITATION USING METALLIC AND POLYPHENOLIC HETEROPOLYANIONS Under strongly acid conditions, inorganic and some organic strong anions—known as heteropolyanions—ably precipitate proteins. Inorganic anions used under acid conditions for this purpose include perchlorate, tungstate, phosphotungstate, molybdate, phosphomolybdate, tungstosilicate, and ferrocyanide. Inorganic ions (used under acid to neutral conditions) include sulfosalicylate, picrate, and diverse plant polyphenols and tannates (also see Critical Parameters). Prior to overt precipitation, protein molecules under strongly acidic conditions are acid-expanded and remain soluble. When the heteropolyanions are introduced and allowed to bind, the protein molecules are forced back to a compact, poorly hydrated conformation. Molecular expansion and contraction of the protein in solution, which leads to precipitate formation, may be followed by biophysical tools (Fink, 1995). Driven further with additional heteropolyanions, such as perchlorate and tungstate in ~0.2% to 2% or 3% concentration, proteins coprecipitate with these anions in dense aggregates that easily settle or centrifuge down. In some procedures for precipitating proteins, perchloric acid (HClO4) is the precipitating agent of choice because it is ultraviolet-transparent above 250 nm. Perchloric acid precipitates whole protein molecules, but not their low-molecular-weight fragments— amino acids and oligopeptides. This is the basis for analysis with many proteolytic enzymes—at the end of the proteolytic cleavage assay, whole or intact proteins are precipitated out by a few percent HClO4. Proteolytic fragments are left in the supernatant for subsequent A280 measurement, giving a rather direct measure of the amount of hydrolysis that occurred before adding the HClO4.
BASIC PROTOCOL 8
SELECTIVE PRECIPITATION USING HYDROPHOBIC ION PAIRING (HIP) ENTANGLEMENT LIGANDS Hydrophobic ion pairing (HIP) is coprecipitation of proteins using flexible hydrocarbon “tails” of alkane anions (detergents) that have their anion head groups bound in the target proteins. Figure 4.5.8 illustrates the composition of such a coprecipitate. Organic anions, especially sulfonates and sulfates, often bind strongly to proteins bearing cationic sites.
SO – + 3
+ SO3–
SO3–
+
Selective Precipitation of Proteins
+ –O S 3 + –O S 3 –O S 3
+
+ – SO3 + SO3–
SO3– +
+ –O S 3 + + SO3– –O S 3 protein cation sites, ZH + ≅ ϑ ligand anions –O S SO3– +3 +
+ SO3– –O S + 3
+
+ SO3–
SO3– +
–O S 3
+
–O S 3 –O S 3
+
Figure 4.5.8 HIP coprecipitate. Strong anions (sulfate or sulfonate-bearing C10–C14 alkane tails, e.g., dodecyl sulfate) bind to protein cationic sites forming ligand-protein complexes. Complexes draw together via tail-tail hydrophobic entanglement, aggregate, and coprecipitate. At maximum coprecipitating efficiency with ~10−5 M protein, the stoichiometric ratio of bound ligand anions (ν) is to cation side chains (ZH+) in the protein molecule is close to 1:1.
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The organic alkane groups entangle and bind to each other in a manner similar to tail/tail association in micelles. Entanglement draws protein/ligand complexes together as shown in Figure 4.5.8. Such complexes go on to aggregate, then coprecipitate. Ordinary detergents such as dodecyl sulfate, (DDS−) in correct protein/detergent ratios are able protein precipitants. Such ligands, especially detergent anions, need to be carefully regulated with regard to concentration. Suitable overall concentrations of precipitating detergents are usually 10−4 to 10−5 M when used with overall protein concentrations ranging from 0.01% to 0.001% (w/w) or 10−4 to 10−6 M protein. If suitable ligand concentrations are exceeded, the ligands go on to form micelles and solubilize proteins rather than precipitate them. Inside their optimum coprecipitation range with respect to total protein concentration, these ligands are rather generally effective towards many proteins. Protein coprecipitation by HIP is frequently easy to reverse. The precipitate is redissolved in solvent with a pH ~2 or 3 units more alkaline than the coprecipitation pH. Detergent anions may be quantitatively stripped out with strong ion exchange resins, usually in the chloride form—e.g., Amberlite IRA-400Cl− or Dowex-1Cl−, or equivalent resins. Released proteins—e.g., enzymes in the supernatant—are then ready for assay. SELECTIVE PRECIPITATION BY MATRIX-STACKING LIGAND COPRECIPITATION
BASIC PROTOCOL 9
Matrix-stacking ligands function (see Fig. 4.5.9) to draw protein molecules together in dense complexes that aggregate and coprecipitate protein. Ligand/ligand association forms a matrix host; protein molecules trapped in the matrix are guests. Occasionally, these ligands cocrystallize proteins and peptides (Conroy and Lovrien, 1992). Ligand organic tails bear rather rigid coplanar groups that stack and bind to one another π-face–π-face fashion, in a manner similar to stacking of aromatic bases in nucleic acid double helices. Tail-to-tail matrix association is reinforced by short alkane groups on the periphery of the tails—i.e., methyl, ethyl, and t-butyl substituents. The alkyl substituents lend hydrophobicity and promote water displacement, which helps drive tail-to-tail association. The structures of a few anionic ligands, are shown in Figure 4.5.10. As with flexible-entanglement ligands (see Basic Protocol 8), the more rigid matrix-stacking ligands achieve maximal coprecipitation efficiency when ν ligands are bound per protein molecule with close to a 1:1 ratio of ν to ZH+ —i.e., ν = ~ZH+, where ZH+ = protein net cationic charge from H+ titration of side chains. Hence coprecipitation depends on pH and how pH governs the protein charge, ZH+. Stacking and entanglement ligands in their most useful overall concentrations of 10−4 to 10−5 M often protect proteins as well as precipitate them. Such ligands tighten protein conformation, sometimes profoundly (Matulis and Lovrien, 1996). Protection often starts in solution before overt coprecipitation. The capacity to protect proteins against several kinds of degradation is an advantage of matrix-stacking ligand coprecipitation. Two variables around which to design early protein coprecipitate experiments are illustrated in Figure 4.5.11. The following steps may be used for developing a matrix-ligand coprecipitation protocol. Materials Crude protein solution of interest, particle-free (see discussion of Clarification in Strategic Planning) Potential ligands (Fig. 4.5.10; also see Aldrich catalog) Appropriate buffers Dowex-1Cl– resin Small test tubes or 2- to 3-ml plastic microcentrifuge tubes
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HO
R1 N
N
SO–3
R2 pH R2 R1
H
O
N
+ SO–3
N
+
–O S 3
N N O
H R1
R2
Figure 4.5.9 Molecular structure-function basis for matrix-stacking ligand coprecipitation of proteins. Protein molecules initially soluble in water and bearing a positive net charge attract strong anion (sulfonate) ligands. Such ligands associate via organic tail/organic tail stacking interaction, reinforced by alkane R-groups. Ligand/protein complexes are thus drawn together and coprecipitate out of solution.
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1. Set up grids of small test tubes or 2- to 3-ml plastic microcentrifuge tubes as in Figure 4.5.11. In each grid, vary two parameters versus one another to optimize coprecipitation—e.g., pH versus ligand concentration (with a fixed total protein concentration), pH versus kind of ligand, and other pairs of variables as described in Figure 4.5.11. 2. If possible, determine protein concentrations in samples to be precipitated (see UNIT 3.4). Usually if stacking and entanglement ligands are to function, proteins need be adjusted in pH in such a way that proteins of 20- to 60-kDa molecular weight carry a ZH+ of ~15 to 40 net cationic charges. If protein molecular weights are unknown, assume ~40,000 Da, or assume the molecular weights typical of the kinds of proteins being worked with (e.g., common microbial and digestive proteases range from 20 to 35 kDa, and coagulation proteases from 50 to 75 kDa).
3. Determine isoionic point for the protein and optimal pH for precipitation. Expect to adjust the pH of some samples to between 2.5 and 5 if information from titration curves or amino acid analysis is not available for predicting ZH+. Preferably this is done
–O S 3
H O N N
CH3
C Little Rock Orange
–O S 3
CH3
CH3 Orange II
H O N N
H O N N
CH2
CH2
Gopher Maroon
SO3–
CH3
Jenelle's Orange
H O N N
H O N N
–O S 3
Crocein Orange G
CH3
SO3–
H O –O S 3
N N
SO3–
Orange ROF
Figure 4.5.10 Anionic ligands synthesized with reinforcing alkane groups (Little Rock Orange and Jenelle’s Orange) are very strong protein coprecipitants. Commercially available dye anions—Orange II, Crocein Orange G, and Orange ROF (available from Aldrich)—frequently are also efficient coprecipitating ligands for cationic proteins in pH ranges 2 to 4 units below the isoionic pH of the protein.
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by direct acidification with ~0.01 M HCl accompanied by pH monitoring. Alternatively, buffers—e.g., glycine or formate buffers—are recommended. Do not adjust protein pH into acid regions before adding the ligands unless there is a need for protection by ligands in control experiments. If sought-for proteins are fragile at acid pH, ligands should be added before titrating the proteins to the lowered pH. Control (contrast) experiments may use the opposite sequence—i.e., acidify first and add ligand second. If there turns out to be much difference in the results obtained via the two sequences, protection of proteins from acid-pH denaturation is indicated.
4. Experiment with two or three ligands from Figure 4.5.10 or choose other potential ligands from among the azoaromatic sulfonate dyes listed in the Aldrich catalog. Prepare ligand stock solutions. Ligands are preferably dissolved in water or in dilute buffer at the running pH. Stay below the limit of ligand solubility, which can vary considerably, in making stock solutions. Upper solubility limits for a number of such ligands are 0.01 to 1 mM.
5. Add ligands from ligand stock solutions to protein sample in various volume ratios—e.g., ~1 vol stock solution to 2 vol protein solution and vice versa. Briefly vortex the mixtures. Extreme ratios—e.g., 10 vol of ligand solution to 1 vol protein samples—should be avoided as these will dilute the protein too much. If possible, arrange for modest mutual dilutions of ligand and proteins on mixing.
6. Optimize temperature, carrying out initial investigations at room temperature and proceeding to higher or lower temperatures as needed.
variable y
variable pH
Selective Precipitation of Proteins
Figure 4.5.11 Sets of small test tubes or plastic microcentrifuge tubes, 2 to 3 ml in volume, may be used to to explore optimum conditions for matrix-ligand/protein mutual coprecipitation by varying pairs of important parameters. Five principal variables determining coprecipitation are: kind of ligand; ligand-protein ratio (y; often between 2 and 20); initial pH; temperature; and concentrations of auxiliary coprecipitating agents such as Zn2+ (10−3 to 10−4 M).
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7. Determine rate of precipitation giving the optimal yield. Matrix-ligand coprecipitation is sometimes rate-limited when protein concentrations start from <0.1% (w/v). If this is the case, incubate ligand/protein mixtures for a few hours. Rather often, coprecipitation occurs rapidly; however, to avoid including unwanted compounds in coprecipitates, do not drive the coprecipitation reaction too rapidly (if possible).
8. Collect coprecipitates by centrifugation at 5 to 50 × g. Perform bioassay of the coprecipitates and, if necessary, of the supernatant for the protein of interest. In some systems, e.g., peroxidase isolations from crudes, myriad unwanted proteins precipitate out first and the peroxidase enzyme is left in the supernatant.
9. Redissolve coprecipitates by shifting pH upward with 0.1 M acetate, imidazole, or phosphate buffers, pH 5 to 7. Add ~100 mg of Dowex-1Cl− resin and incubate 5 to 15 min with gentle shaking to trap the anionic ligands. Monitor the completeness of trapping via the color of the matrix ligand. Approximately 100 mg of resin per milliliter of the higher-pH redissolving buffer provides a good excess of trapping capacity. See Basic Protocol 2 for discussion of the capacities of anion exchange resins. Dowex-1 resins have ~1 meq of capacity per ml (wet volume) of resin beads. Complete trapping is easily followed from matrix ligand’s color. Released proteins in solution should be colorless.
10. Measure total protein (and total released activity if an enzymatic or other bioassay is available) to determine specific activity, total recovered activity, and recovered protein for comparison to samples before coprecipitation. SELECTIVE PRECIPITATION USING DI- AND TRIVALENT METAL CATION PRECIPITANTS
BASIC PROTOCOL 10
Di- and trivalent metal cations—e.g., Zn2+, Mn2+, Ca2+, and Al3+ (general symbol M2+ or M3+)—provide two means for precipitating proteins: (1) they may act directly, or (2) frequently they are useful as auxiliary agents for other precipitation methods such as the crowding method using neutral polymers (see Basic Protocol 5) or matrix-stacking ligand coprecipitation (see Basic Protocol 9). This technique is reviewed by Rothstein (1994) and Gurd and Wilcox (1956). Zinc ions are the ions most commonly used for direct precipitation and as auxiliary agents, especially when working with plasma and coagulation proteins (Cohn et al., 1950). Zinc ions are required for insulin crystallization, including large-scale industrial production of this polypeptide. The discussion below represents a small sector of the landscape of chemical behavior in regard to metal cations. Metal cations provide a diverse, largely undeveloped means of protein precipitation. The intrinsic solution behavior of metal ions (without proteins), especially their chelation-coordination reactions, must be taken into account for application of such ions to the development of protein precipitation methods. Materials Analytical Reagent (AR)-grade metal salts (generally as chloride or nitrate) Crude protein solution of interest, particle-free (see discussion of Clarification in Strategic Planning) Appropriate buffer(s) 1. Determine optimal buffering system. The precipitation system may be buffered in two respects: (1) toward H+ ions and pH in the conventional way and (2) also with respect to metal ions, using metal ion buffers which are coordinating and chelating agents (Perrin and Dempsey, 1974). A number of com-
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pounds—e.g., EDTA, citrate, and several amino acids—can serve both buffering purposes. Evidence that compounds serve as one or both of these types of buffer is that pH dependency and/or metal ion concentration dependency of protein precipitation is rather sharply dependent on the kind of buffering compound. Zwitterionic buffers such as HEPES may be useful for controlling pH because they have much less tendency to bind or coordinate with metal ions than conventional Sorenson buffers (e.g., phosphate). Because proteins are also buffers, their self-buffering action may suffice to control pH if present in concentrations ≥0.3%.
2. Select an appropriate metal cation and precipitate protein. Di- and trivalent metal cations (M2+ and M3+) are vigorously precipitated by salts and inorganic buffer anions, which are present in protein extracts and samples at various stages of isolation. Sulfate (from salting out; see Basic Protocol 1 and Alternate Protocol) powerfully precipitates Ca2+ as CaSO4. Solubilities of <100 mg/ml or even <10 mg/100 ml are seen with Zn(NH4)PO4, Mn3(PO4)2⋅7H2O, MnCO3, Mg(NH4)PO4⋅6H2O, MgCO3, CaCO3, and a number of calcium phosphates. Sulfates and carbonates of Ba2+ are extremely insoluble (Hogness and Johnson, 1954). Before committing a metal ion as a precipitating agent: (1) review general behavior with respect to precipitability by buffer and salt anions in the protein solvent; (2) carry out control experiments with protein-free samples to find if the solvent(s) are likely to produce massive precipitates of simple inorganic compounds with the metal ion; and (3) arrange to remove interfering anions if the anions are in concentrations ≥0.05 M or if concentrations of ions such as SO4−2 and CO3−2 re <0.05 M. Trivalent metal ions such as Al3+ and Fe3+ form a complicated series of oxo-, hydrate, and hydroxylated species in water. They go on to form insoluble hydroxides (solubility products 10−30 to 10−40; Hogness and Johnson, 1954), at neutral and even somewhat acidic pH. Metal cations of this kind are interesting protein precipitants (Gurd and Wilcox, 1956). However, instead of complexing with protein molecules, they may form very insoluble hydroxyl gels and flocculate proteins by adsorption on the gels at pH ≥5. Hence development of protein precipitation with such cations may need be to confined to rather acidic pH regions. It is not necessarily a disadvantage if low-molecular-weight salts or other solvent components precipitate M2+ first, before proteins are precipitated. Inorganic precipitates of sulfate, phosphate, and carbonate may form first. These can then be centrifuged away, whereupon more metal ions are added to bring down the sought-for protein, in a second stage of precipitation. Total protein measurements (UNIT 3.4) are used to locate the starting point for protein precipitation by the metal ions.
3. Analyze precipitate. Analysis of metal content of coprecipitates and their supernatants is useful for characterizing products and following how variations in procedures determine product composition. The most generally accessible methods are spectrophotometric, using color-developing reagents capable of quantitating metal cations at 10−4 to 10−5 M concentrations (Sandell and Onishi, 1978). Figure 4.5.12 shows a calibration curve for Zn2+ analysis using pyridylazoresorcinol (PAR; Aldrich). The optimal pH for use of PAR is ~7 (Cheng et al., 1992) in HEPES or other nonchelating buffer. 0.1 mM PAR is prepared in 0.05 M HEPES buffer, pH 7.0, then dilutions of Zn2+ (as zinc acetate) are prepared in the same buffer at various concentrations ranging from 5 to 30 ìM. The PAR reagent may be made up as a 1 mM stock solution and diluted 10-fold to provide the working reagent at 0.1 mM. Spectrophotometric samples are made by mixing 1.0 ml of 0.1 mM PAR with 1.0 ml of each of the Zn2+ dilutions. A495 is then measured versus a reference of equal PAR concentration without zinc. The slope of the plot in Figure 4.5.12 is the molar extinction coefficient, ε495 = 4.5 × 104 M−1 cm−1. An example of Zn2+ release from a zinc enzyme monitored by PAR color development is given in Hunt et al. (1985). Zinc analysis is a criterion for the identification and purity of insulin. Selective Precipitation of Proteins
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0.8
A495
0.6
0.4
0.2
0 0
4
8
12
16
[Zn+2] in final assay volume (µM)
Figure 4.5.12 Calibration plot for zinc ion spectrophotometric analysis using pyridylazoresorcinol reagent.
4. Remove metal ions from precipitate. Metal ions may be removed from protein precipitates via cation-exchange resin trapping, or by treating precipitates with a buffer at a pH at which it is expected to dissolve. It is preferable to mix the precipitate with a release buffer first to see if in fact it does dissolve, then test the resin-bead exchanger afterwards. Trapping resins in the Na+ or K+ form may be tested; conventional cation resins such as Amberlite IR-120Na+ usually give adequate results. A specialty chelating resin—Chelex 100 exchange beads (Bio-Rad)—may also be used. The exchange capacities of such resins are 1.8 and 0.4 meq (milliequivalents) per milliliter of wet resin. The resins should not be used in great excess (i.e., >5× resin equivalents per estimated equivalent of metal to be removed) because such resins may also adsorb proteins. Exchange of metal ions out of precipitates may be accelerated by conventional soluble chelating agents (e.g., 10−4 M EDTA) or by amino acids such as aspartic acid in the case of Zn2+ (Fiabane and Williams, 1977).
5. Perform bioassay for protein of interest. COMMENTARY Background Information In practice, most protein-precipitating agents and methods yield a variety of different products depending upon conditions such as pH, metal ions present in the solution, and/or presence of cosolvents. Many “precipitates” are actually coprecipitates in which molecules of the precipitating agent remain bound to the proteins. How much precipitating agent or ligand becomes bound and thus coprecipitated usually remains unclear until the coprecipitate is analyzed. Precipitation and coprecipitation
are most important in “upstream” stages of protein isolation—for crude, dilute, solutions in large volumes up to several liters. However, precipitation is sometimes useful downstream also, as it reduces volume—e.g., excess solvent and buffers may be removed from chromatographic eluates via this technique. Sometimes it is an efficient strategy to precipitate unwanted proteins in the first stages of purification and then capture the sought-for protein in a second precipitation step later. Some precipitation methods are old, and
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some have been developed more recently. Some of the old methods, particularly salting out with ammonium sulfate (which is more than 130 years old), remain as useful as ever. Ammonium sulfate offers the ability to protect sensitive proteins and may be scaled up at a reasonable cost. Some precipitating and coprecipitating agents yield precipitates that readily reverse. Reversibly acting agents can be stripped away later—e.g., ammonium sulfate salting out is reversed by dialysis, pressure filtration, or resin exchange to remove sulfate. Other agents, despite their great powers in precipitating, may be difficult to remove completely, or may catalyze damage to proteins (e.g., sulfhydryl oxidation). Consequently, precipitating agents must be evaluated in relation to later steps that may have to be used to compensate for the disadvantages that some strong precipitating agents carry with them. Sometimes the optimal precipitating agent is a weaker rather than a very strong agent, if the weaker agent is more easily dissociated, trapped out, or dialyzed away to release the sought-for protein. Besides the agents and individual methods listed in Table 4.5.1, combinations of methods are frequently advantageous. For example, sometimes it is helpful to add modest concentrations of Zn2+ to proteins that one is trying to precipitate by another method. Another precipitation technique, TPP (see Basic Protocol 4), is also a “hybrid”—a combination of organic cosolvent precipitation and sulfate kosmotropic precipitation. Precipitation and coprecipitation are often quite protective of proteins, especially in salting-out techniques. However, whether or not protection is achieved frequently depends on the sequence of steps used in each protocol. For example, if salting out or coprecipitation with a particular ligand needs to be carried out at a pH very different from the starting point, a choice must be made between shifting pH first and then adding salt, or alternatively adding salt or ligand first and then readjusting pH. The latter is usually best choice in terms of protecting the protein.
Critical Parameters and Troubleshooting
Selective Precipitation of Proteins
Salting out In the simplest version of salting out, desired proteins are precipitated relatively early and the supernatant is set aside or discarded. In this case, unwanted proteins are often left in the supernatant. The supernatant may, however, be readjusted in pH, increased in salt concentra-
tion, or both, to bring down yet other proteins, perhaps the sought-for protein, in a second stage or “second cut” of salting out. Capture of sought-for protein activities in second and even third stages or cuts, with intervening pH shifts, are discussed in various sources (e.g., Scopes, 1987). In each step, or cut, the sequence of salt addition, pH shifting, and changes in other parameters (such as temperature) must be considered. The salt (ammonium sulfate) may be added before shifting other parameters, especially the pH. Alternatively, the opposite sequence may be used: adjust the pH first to some desired set point, then add the salt. These two sequence options can have quite different effects for several reasons, stemming from the fact that salts, particularly ammonium sulfate in concentrations greater than ~0.5 M, often serve as protein-protective agents. The proteinprotective effects of ammonium sulfate include conformation tightening, which guards against pH-induced unfolding. Sometimes it is advantageous to impose quite acidic or alkaline pH extremes or a thermal shock, to first precipitate unwanted, denatured proteins, and leave sought-for proteins in the supernatant. However, the amount of salt and when to add it are variables; these need attention in weaving the optimum path for salting out. Several treatises and textbooks present tables, formulas, and nomograms for relating percent saturation, molar concentration, weight of ammonium sulfate to be added, and dilution factors that are useful for increasing or decreasing amounts of ammonium sulfate so as to arrive at prescribed concentrations of the salt. The numerical relationships are mostly cited for 0° to 30°C, although the solubility of ammonium sulfate in water changes little. Saturated ammonium sulfate in water is close to 4.1 M. Two relationships cited by Scopes (1987) give the number of grams (g) to be added to a liter of solution at 20°C for shifting molarity (M) and percent saturation (S). The first of these equations is for shifting to a molarity M2 starting at molarity M1: 553( M2 − M1 ) g= 4.05 − 0.3 M2 The second equation is for shifting to a percent saturation S2 starting at percent saturation S1. 553( S2 − S1 ) g= 100 − 0.3S2 These relationships pertain to solutions of
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ammonium sulfate in water. Proteins that are already in diverse buffers and that contain other salts, in unknown concentrations, throw off the accuracy of nomograms and formulas for shifting saturation percentages and molarities. Nevertheless, these are useful for estimating added salt concentrations to be used and for reporting. Neutral salts other than ammonium sulfate, such as sodium or potassium chloride, may be tested in attempting to precipitate the soughtfor protein (and later in attempting to split excess salt away—i.e., reverse the process). However, ammonium sulfate is usually the best primary choice, not because of tradition but because the sulfate anion, SO4−2, is one of the strongest yet most benign Hofmeister kosmotropes (UNIT 8.4) over nearly the whole “biological” pH range, approximately from pH 2 to 10 (Collins and Washabaugh, 1985). The orthophosphate dianion, HPO4−2 is also a strong, inexpensive kosmotrope and salting-out agent. However the pK of H2PO4− is 7.2; accordingly, below pH ~6.5 the H2PO4− monoanion predominates even though it is a considerably weaker kosmotrope. Other anions such as fluoride, F−, are good kosmotropes but are very toxic. For a combination of desirable properties, including least expense, the sulfate systems remain the most convenient, reliable, and most tested. One disadvantage of sulfate arises in the case of proteins requiring calcium ion, Ca2+. Because calcium sulfate and calcium phosphate are extremely insoluble, both SO4–2 and HPO4–2 precipitate Ca2+. Hence, for calcium-requiring proteins, salting out may have to be developed with salts other than sulfates— e.g., acetates are a possibility. Isoionic precipitation Amberlite resins IR-120 (a cation exchanger) and IRA-400 (an anion exchanger), in the form of 6% or 8% cross-linked beads, are the basis of the original Dintzis mixed-bed exchangers. The equivalent Dowex exchangers, Dowex 1 and Dowex 50 series, in the form of 6% to 8% cross-linked resins, serve equally well but are not superior. Anion and cation exchangers have differing exchange capacities per settled milliliter of wet resin. IR-120H+ has ~1.8 meq/ml of capacity, and IRA-400OH− has ~1.2 meq/ml. Accordingly each 100 ml of mixed exchanger may be made by mixing 40 ml of the cation exchanger in the H+ form with 60 ml of anion exchanger in the OH− form, a volume ratio inversely proportional to their intrinsic exchange capacity ratio. Mixed-bed exchangers may be stored in
the cold (do not freeze them) for long periods without self-neutralization. Mixed-bed exchangers, such as the MB-1 Amberlite series, may be purchased premixed. However it is recommended that the exchanger be mixed in the laboratory to ensure that resins have been “exercised” (see Basic Protocol 2, steps 2 and 3) and leached of any polymer hydrolysis products. Anion exchangers based on various quaternary amines are especially susceptible to self-hydrolysis (detectable by an amine odor) upon storage in the OH− form. Commercial ready-mixed bed resins usually incorporate dye chromogens (acid-base indicator molecules), which show (by a blue to amber color change) when their capacity has been titrated. Such resins may be usable with some protein molecules, but if there is any leaching out of resin materials, the eluants will be strongly colored, indicating contamination of proteins. Two-carbon organic cosolvent precipitation In Basic Protocol 3, no general criterion can be put forth for choosing between acetone and ethanol, for deciding how pH should be adjusted versus the isoionic point of the protein, or for determining the exact salt concentration to be used. There exists a large body of literature describing empirical, experimental means for C2 cosolvent–based isolation of dozens of proteins. A few guidelines may be useful. Ethanol was the C2 cosolvent used for the isolation of most blood-plasma proteins. Many respiratory enzymes have been isolated with a step involving an acetone cosolvent. In systems where lipids and triglycerides are troublesome, many workers have used acetone cosolvents, sometimes at nearly 100% acetone, because acetone and acetone-water mixtures extract lipid materials. Acetone in such applications functions as an extractant as well as a protein precipitating agent. Proteins and whole-tissue agglomerates that have been rendered nearly lipid-free by acetone, then lyophilized, are referred to as “acetone powders.” A review with many references was recently written by Rothstein (1994), which includes discussion of the seminal papers by Cohn and coworkers. With a number of proteins, a complex relationship exists between salts, the C2 organic cosolvents, the proteins, (which bind salts to some extent), and the pH. These parameters operate to make organic cosolvent precipitation quite empirical. Methanol and the two-carbon organic cosolvents appear to be able to penetrate protein
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molecules and compete with water for sites inside. However C4 and larger cosolvents are largely excluded from the inside of proteins if the proteins are kept compact and folded. Accordingly, it is unlikely that the larger cosolvent molecules, such as C4 alcohols, behave congruently with the C1 and C2 cosolvents for most proteins.
Selective Precipitation of Proteins
C4 and C5 organic cosolvent precipitation Some C4 organic cosolvents, particularly t-butanol and n-butanol, are able precipitating and extracting agents for proteins. They are useful in “upstream” stages of isolation, starting with crude solutions. They are variously used in combination with osmolytes and salting-out agents such as ammonium sulfate. Conditions are adjusted in such a way that separated layers of cosolvent are obtained, along with an aqueous layer and a layer of precipitated protein. The separated liquid layers act as extraction and partitioning systems as well as protein precipitation systems. Precipitated proteins usually appear at the interface between liquid layers. Because some of the C4 and C5 cosolvents act either as kosmotropes or in parallel to kosmotropes like the sulfate anion, these cosolvents frequently are protective of aqueous proteins even at normal temperatures, 20° to 30°C. They induce protein-molecule conformation tightening, from which the protein-molecule protection partly originates. The similarity between sulfate interaction with proteins and C4/C5 cosolvent interaction with proteins does not, however, have the same origin. Sulfate exerts electrostatic forces toward many proteins (Chakrabarti, 1993), whereas t-butanol exerts much less electrostatic-based interaction, perhaps none at all. A rough model of the effect of the t-butanol cosolvent is that it behaves like a neutral osmolyte in water. It is not always strictly necessary to choose t-butanol as the cosolvent. However, t-butanol has some advantages. First, it is relatively inexpensive. Only ~0.3 to 0.5 ml of t-butanol is required per milliliter of starting aqueous protein solution, after the optimal amount of salt has been added. When t-butanol is used, smaller amounts of ammonium sulfate are usually required for a given precipitation, relative to the amounts needed in conventional salting out. Second, t-butanol is much less volatile than C2 cosolvents, especially in water-dominated systems. Hence it poses less of a fire hazard and less hazard from inhalation. Third, t-butanol is, in general, far less denaturing at 20° to 30°C temperatures than C2 cosolvents. Fourth, t-bu-
tanol behaves similarly to kosmotropes and osmolytes, protecting proteins. Finally, the extractive and partitioning capacities of threephase partitioning (TPP) using C4 and C5 organic cosolvents (see Fig. 4.5.6) tend to remove pigments, lipids, and amino acids from proteins. C4 and C5 organic cosolvents in the aqueous phase produced by techniques like TPP exert themselves as osmolytes and crowding agents, up to a point, when they reach concentrations of several percent. However, the main differences between the C4/C5 organic compounds (of the types discussed here—e.g., butanols and pentanols) and water-soluble osmolytes such as sugars are: (1) osmolytes like sugars and amino acids tend not to form a separate organic phase that goes on to act as an extraction phase for lipids, pigments, and other nonprotein material, whereas the butanols and pentanols do; and (2) hydrocarbonaceous C4 and C5 compounds bind to some extent to proteins that have a high degree of nonpolar, hydrophobic character. This latter property causes such proteins to position themselves as illustrated in Figure 4.5.6, floating above the aqueous layer. When the protein precipitate (i.e., coprecipitate) is redissolved in buffer with low salt, then shifted away from the precipitative pH (usually upward ~2 to 3 pH units), in most instances the C4/C5 compounds dissociate. In contrast, conventional osmolytes, especially sugar, tend not to bind to proteins. In some versions of TPP, the system is operated in two stages. In the first stage, the precipitate containing unwanted proteins that forms at the initial pH is discarded. The aqueous phase is then shifted in pH, and/or more salt is added, giving a second precipitate containing the sought-for protein. In the example described in Lovrien et al. (1987), the enzyme α-amylase is isolated in this way. Many sorts of variations can be made in pH, salt concentration, and other parameters. Protein-exclusion and crowding agents Exclusion agents and osmolytes have high affinities for water and compete with proteins in solution for water. The crowding action, aided by any degree of affinity of protein molecules for one another (protein-to-protein association) promotes protein precipitation. Thermodynamic pushing occurs because hostile environments imposed on proteins in osmolytes and and in crowded solution raises protein chemical potential in a positive direction. This is then relieved by protein precipitation, which
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enables protein molecules to exist with lowered chemical potential. Crowding and exclusion agents frequently act upon conformationally motile (loose), water-penetrated proteins before overt precipitation. Conformationally loose protein molecules are also “squeezed” on by these agents, promoting protein molecule tightening and sometimes promoting an ordered protein conformation. Proteins thus “squeezed on” often become protected, sometimes dramatically. Moreover if their conformation is forced into good order, protein molecules may crystallize instead of simply being precipitated in amorphous form. Thus, the osmolyte MPD (2methyl-2,4-pentanediol) is often used with success for protein crystallization. MPD crowds, excludes, and squeezes to push proteins into a narrow or even single conformation necessary for arriving at the crystalline state. As a general rule, osmolytes and crowding agents do not bind strongly to proteins. Hence, ideally, precipitated proteins are free of such agents. However in fact, some agents bind rather weakly, and appear in the precipitate (which is in fact a coprecipitate). Accordingly, selection of the agent may depend on the ease or difficulty of removing trapped agent from protein precipitates if in fact the desired redissolved protein needs be stripped of precipitating agent. Neutral polymers such as polyethylene glycol (PEG) and dextran, which do not affect electric charge, might be expected to act toward proteins independently of electrostatic parameters such as protein net charge and salt concentration. However, the opposite often occurs: i.e., buffer, salt, and pH-dependent protein charge frequently sharpen and either greatly enhance or sometimes depress protein precipitation via PEG. Salts in concentrations quite below normal kosmotropic concentrations— i.e., 0.01 to 0.5 M salts—have a large influence, decreasing chymotrypsin solubility and increasing its precipitation by PEG (Miekka and Ingham, 1978). In other examples, chaotropic anions and “Hofmeister-neutral” anions (e.g., chloride) increase protein solubility and depress precipitation. Readers may profit by consulting some of the papers referred to here, particularly to Miekka and Ingham (1978) to review how large and variable these “electrostatic” effects may be. Neutral polymers, acting as steric-exclusion precipitants, and the monomers of such polymers, acting as osmolytes, are far from equivalent in protein-precipitating power when used
in equal concentrations. The polymers appear to be considerably most effective. For example, Laurent (1963) investigated dextran polymer and its monomer glucose as precipitants for albumin and fibrinogen. Dextran polymer considerably outstripped glucose in precipitation power when each reagent was used at a concentration of ~0% to 10% (w/v). On the other hand, concentrations of proteins for achieving protein precipitation using neutral polymers usually must be from 1% to 5% (w/v). Hence the neutral polymers are considerably less sensitive and less effective at low protein concentrations than some of the other methods discussed in this unit—e.g., the matrix-stacked ligand technique (see Basic Protocol 9 and discussion below). Neutral polymers are equivalent to neutral polymers used in aqueous two-phase partitioning (Walter and Johansson, 1994) and as “precipitants” in protein crystallization (McPherson, 1982). PEG is the dominant polymer used in two-phase partitioning and as a precipitant for protein crystallization. PEG polymers in several molecular-weight ranges—low, medium, and high—are available from Sigma and other suppliers. Choice of PEG molecular weights must be balanced between two considerations. Higher-molecular-weight polymers tend to be more effective precipitants for some proteins (i.e., able to precipitate albumins out of solution in ~5% to 15% PEG concentration ranges; Atha and Ingham, 1981). Lower-molecular-weight PEG polymers of molecular weight <3000 to 4000 generally must be used in 20% to 50% concentrations to effectively precipitate a number of proteins that have been studied. However, the practical problem of ridding precipitated proteins of trapped PEG polymers is eased with low-molecular-weight PEGs because they dialyze away relatively easily. Proteins generally are more sensitive to precipitation by high-molecular-weight PEGs than low molecular-weight-PEGs, frequently precipitating, if indeed they do so, in 3% to 20% concentrations of large PEGs (molecular weight greater than ~4000; Atha and Ingham, 1981). The downside of high-molecularweight PEGs is that, if trapped, these PEGs become difficult to separate from coprecipitated proteins by simple dialysis. Also, highmolecular-weight PEG solutions tend to be quite viscous, whereas oligomeric low-molecular-weight PEGs are easier to handle and mix. Alkane ether polymers (trade name Ucon; Union Carbide; see SUPPLIERS APPENDIX), which are copolymers of ethylene and propylene ox-
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ide related to PEGs, go into and out of solution with marked temperature dependency (i.e., they “phase-separate”; Albertsson and Tjerneld, 1994). Accordingly, with the use of such polymers, manipulation of temperatures in initial precipitation and in redissolving protein precipitates may afford means for balancing these two problems.
Selective Precipitation of Proteins
Synthetic and semisynthetic polyelectrolytes Polyelectrolytes have been synthesized as copolymers with nonionic groups to help alleviate problems that arise with homopolymers such as polyacrylic and polymethacrylic acid. The latter homopolyelectrolytes, when ionized, are so strongly coprecipitating that they are often poorly selective. Hence, copolymers of acrylic acid and methacrylic acid with a nonionic methylmethacrylate ester side function as agents with softened coprecipitating power but increased selectivity. A series of copolymers of this type, with varying ratios of ionic and nonionic groups, are named “Eudragit” agents, and are available commercially from Rohm GmbH (see SUPPLIERS APPENDIX and Mattiasson and Kaul, 1994). Copolymerization of two and even three separate vinyl monomers provide polyampholytes of myriad kinds, differing in ratios of vinyl monomers. Synthetic polyampholytes take on net positive, net negative, or zero charge, depending on the pH (like proteins, which are also polyampholytes). The advantage of such compounds is that after coprecipitating proteins, the synthetic polyampholyte can be split away from the protein by adjusting the pH of the mixture to the isoionic pH of the polyampholyte, which will precipitate it separately (Patrickios et al., 1994). Many of the polymers that have been discussed so far—e.g., carboxymethylcellulose and polymethacrylic acid anion—are negatively charged above the pK of their side-chain carboxyl groups, ~pH 5. Such polymers should primarily be used for precipitating proteins that are cationic—i.e., in pH ranges below the target protein’s isoelectric point. However, it may be necessary to experiment with synthetic polyelectrolytes which are cations and which are therefore primarily aimed at precipitating anionic proteins in pH ranges impractical for synthetic polyanions. The most common, readily available neutralcationic synthetic polyelectrolyte of this type is polyethyleneimine (PEI). The H+-titratable nitrogen groups of PEI are integral parts of its backbone, not relegated to side chains. PEI is very basic and exists as a polycation over a very wide range of pH. PEI is also a powerful copre-
cipitating agent for proteins. Table 4.5.2 lists a small number of polyelectrolytes, several of which are sold by Aldrich and Sigma. References in Table 4.5.2 should enable the reader to survey conditions used for coprecipitation of various enzymes. Metallic and polyphenolic heteropolyanions There is considerable variability but lack of much quantitative information concerning recovery and bioactivity measurements of most proteins that have been precipitated with heteropolyanions, in cases where their activities must be recovered. The great acidity and also the oxidizing power of some of these agents— e.g., HClO4—tends to seriously denature many proteins. Deamidation of proteins sometimes follows strong acidification, as well as unwanted hydrolysis and conformational unfolding. However, weaker acids such as tannic acid (a mixture of several phenolic products) are less denaturing, and yield intact active enzymes and other proteins after tannate precipitation at neutral pH. Accordingly, choosing among heteropolyanion precipitation agents depends on the use to which the protein precipitates are to be put. For very rapid, complete precipitation of unwanted proteins down to ~50 µg/ml (Bensadoun and Weinstein, 1976), in cases where destruction of proteins is acceptable, HClO4 in 1% to 5% concentration is used. Picric acid is almost equally effective, but it is highly colored and hence interferes in analytic procedures requiring spectrophotometric measurement. As with polyelectrolyte precipitation, which agent to use may depend on the ease versus difficulty of removing such agents from proteins after coprecipitation. Examples of precipitation of diverse proteins by heteropoly anions at mildly acidic pH (i.e., pH 4 to 6) are described in two papers. Sternberg (1970) coprecipitated ovalbumin, hemoglobin, pepsin, and amyloglucosidase from protein crudes using four heteropoly anions. Sternberg removed the inorganic anions by dialysis, ultrafiltration, and ion exchange, recovering fair to good activities in some cases. Astrup et al. (1954) worked with six heteropolyanions in staged purification of serum proteins including α- and γ-globulins. Tannic acids are polyphenolic compounds (sometimes conjugated with one or more saccharides) from plant extracts. They are quite diverse in structure, depending on the plant source. Sigma sells a crude tannic acid that is convenient to start with. An advantage to using tannates as precipitants is that they function in
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neutral pH regions, pH ~4 to 8. Hence, severe acid and hydrolytic conditions, often necessary with the metallic heteropolyanions, may be avoided with tannates, which are all-organic anions. Little detail is known concerning how tannate precipitation works. Tannates are sometimes effective in precipitating proteins on a broad front, and inclusively, but in some systems fail to precipitate proteins to which they are bound (Hagerman and Butler, 1991). Amounts of tannic acid and of tannate needed for precipitation of diverse proteins range from approximately half as much by weight as the sought-for protein, down to quite smaller amounts of tannin; these quantities are also quite dependent on pH. Hagerman and Butler (1978, 1991) survey principal parameters for controlling precipitation of individual proteins such as albumin and lysozyme. They recommend quebracho tannin and commercial tannic acids as initial agents for attempting precipitation of mixtures of proteins. Although there are gaps in the precipitating power of tannate compounds for proteins, they provide alternatives for scaleable, reversible procedures under conditions where the metallic heteropolyanions (strong acid) are too severe. The strong spectrophotometric absorption spectra of tannins when reacted with Folin phenol reagent or with ferric chloride, both of which generate characteristic colors for phenols, afford convenient means for quantitating tannates in coprecipitates and in soluble remnants (Hagerman and Butler, 1978). Hydrophobic ion-pairing entanglement ions It may seem surprising that ligands such as those used in Basic Protocol 8—especially detergents—are efficient precipitating agents; these reagents are normally used in solubilizing and denaturing proteins. Detergent anions (i.e., dodecyl sulfate) acting as protein coprecipitants were first studied at length by Putnam and Neurath (1944) for seven proteins. Since then, the detergent/protein and subsequent complex/complex association reactions have sometimes been called hydrophobic ion-pairing reactions. A rather strict detergent-anion to protein-cation association occurs in 1:1 stoichiometry to form ion pairs between charged groups; simultaneously an alkane/alkane association occurs between the hydrophobic tails. The two sets of forces strongly pull the complexes together and out of the solution to agglomerate and form the visible coprecipitate. Protein-molecule to protein-molecule association may also occur, but there are rather few
detailed studies of this. The ion/ion association, which is the electrostatic component of HIP, is dependent on and adjusted by the pH, because the pH and the number of protein H+–titratable side chains determine the number of net cationic charges on the global protein molecule— i.e., ZH+. This, in turn, determines the number of ligand anions expected to bind when coprecipitates are formed. Because strong anions from detergents usually bind strongly, there is seldom a need to add extra ligand (detergent) to provide for large amounts of free (unbound) ligand. Between 60% and 95% of the total detergent anions added under the aforementioned conditions are likely to be bound for 10-carbon, 12-carbon, and larger detergent sulfates. However, short-chain detergent anions— e.g., hexyl and octyl sulfates—are usually poor HIP precipitants. Matrix-stacking ligand coprecipitation In organic anion–matrix ligands, the rigid azoaromatic sulfonates are able to stack their tail groups to force association and coprecipitation. These reagents have several similarities to flexible detergent coprecipitating anions (HIP agents; see discussion above). As in the case of the flexible detergents, matrix-ligand coprecipitation is sharply dependent on protein electrostatic charge, thence on pH, on protein side-chain composition, and on the isoionic point. In contrast to the detergent anions, whose coprecipitating powers are mainly dependent on alkane-chain length, matrix-stacking ligands are dependent on organic structural detail and placement of groups, not simply on size. The placement of short alkane “reinforcement” substituents and the location of sulfonate anions on aromatic rings, as well as the location of tautomerizing hydroxyl groups, make stacking-ligand structural character more complicated and unpredictable than that of flexibledetergent ligands. In practice, one may need to experiment with a variety of matrix ligands to find an optimal kind for selectively coprecipitating each protein. It appears that matrix-stacking ligands need their hydrophobic tail groups to extend some distance away (greater than ~10 to 15 Å) from the sulfonated head group (Matulis and Lovrien, 1996). This requirement presumably maximizes tail-tail stacking and hydrophobic association, distancing them so as not to interfere with sulfonate group–protein cation association. In any case, functions of matrix ligands are sensitive to placement of substituents such as alkane groups on the azoaromatic rings, as
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well as the number of such substituents. In practice, one expects to initially experiment with ~3 to 5 such ligands to find a ligand suitable for investigation in more detail. Because matrix ligands generally are quite protective of proteins with which they coprecipitate, it may not be necessary to always develop the coprecipitation stage of protein isolation at cold temperatures. Indeed, the method often interfaces well with a heat-shock step to clear out unwanted proteins susceptible to heat shocking. Pilot experiments therefore can generally be carried out at ordinary room temperatures. Matrix ligands strongly coprecipitate proteins. However, overall isolation processes for most proteins usually need an exit. The protein and ligand should be released from one another by resin-exchange trapping as described in Basic Protocol 2. Hence, it may occur that somewhat more weakly coprecipitating, but more easily released ligands may be optimal in the procedures, so as to allow proteins to dissolve, ready for assay or the next isolation steps.
Selective Precipitation of Proteins
Di- and trivalent metal cation precipitation Metal cations exert themselves in three ways to precipitate proteins. First, their charges add to protein molecule net charge, enhancing binding of coprecipitating ligand ions such as matrix or detergent sulfate anions. Second, metal ions draw in protein side chains and lysine amines, as well as carboxylate and histidyl side chains, to fill metal ion orbitals with their electron pairs. Protein molecules thus act as large chelating agents in a manner analogous to the behavior of EDTA and other multidentate chelators and become tightened and sometimes protected because of the chelation to M2+. Finally, M2+ and M3+ ions can become complexed with buffer ions and other electrolytes chosen by the researcher. When that happens, actual net charge and remaining metal ion ligand positions available to the protein may be adjusted to modulate protein precipitation. Metal ion complexes of M2+ and M3+ may actually be made neutral, or even anionic, depending on presence of chloride, acetate, or citrate, and their concentrations (in the 10−3 to 10−1 M range); this phenomenon is also dependent on pH. Cotton and Wilkinson (1988) cite Zn(H2O)6+2, ZnCl(H2O)5+1, ZnCl2(H2O)4, ZnCl4−2, and ZnCl4(H2O)4–2 as examples. Cobalt and a number of other transition metal cations complex with ammonia and water molecules to make a series of coordinate cations of considerably larger size than the parent Co2+ but which still carry the central metal ion’s
charge. On the other hand, addition of CN− or SCN− generates a series of complexes such as Fe(CN)6−3 which are strong, very stable anions. A number of metal cations—e.g., Ni+2 and Zn+2—are capable of switching configuration from square planar to tetrahedral to octahedral, depending on coordinate ligands, chelating agent, and solvent conditions. Some metal cations useful in protein precipitation are amphoteric, switching water molecules and OH− ligands into and out of metal orbital positions depending on pH. Metal cations such as Al3+ strongly bind oxo-ligands, including water molecules and hydroxyl (OH–) to form metallic gels that settle or centrifuge down and capture proteins by adsorption (Collingwood et al., 1988). This mode of precipitation is often called flocculation. Flocculated products tend to have uncertain composition, but have the advantage of being dense and separable by low-speed centrifugation or simply by settling. If the choice of metal ion is otherwise not obvious, the first choice as protein coprecipitant is the zinc ion. Zn2+ is of intermediate strength in chelation-coordination reactions with respect to the kinds of ligands offered by amine, carboxylate, and imidazole side chains. Therefore it can be “reversed”—i.e., removed from coprecipitates. Zn2+ rather rapidly exchanges ligands that coordinate with it (Berg and Shi, 1996). More importantly, zinc ions do not promote electron transfer and oxidation-reduction reactions, whereas many of the transition metals—e.g., iron, copper, and nickel ions—indeed do so. Hence Zn2+ is fairly safe to use for precipitating proteins with sensitive sulfhydryl (–SH) groups. Other metal ions– e.g., Cu2+ and Ni2+ promote oxidation, usually irreversibly, and therefore can be rather destructive. Zn2+ in the form, for example, of zinc chloride, is also the probable first choice to use as an “assist” agent in conjunction with other precipitants. Atha and Ingham (1981) show excellent examples of the ability of ZnCl2 to decisively decrease the concentration of PEG crowding agent necessary to precipitate out human serum albumin from a 0.5% solution. Zn2+ appears to tighten the albumin molecule’s conformation, easing PEG’s task in precipitating the protein.
Literature Cited Albertsson, P.A. and Tjerneld, F. 1994. Phase diagrams. Methods Enzymol. 228:3-13. Alner, D.J., Creczek, J.J., and Smeeth, A.G. 1967. Precise pH standards from 0° to 60°. J. Chem. Soc. A 1205-1211.
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Astrup, T., Birch-Andersen, A., and Schilling, K. 1954. Fractional precipitation of serum proteins by means of specific anions. Acta Chem. Scand. 8:901-908.
Edsall, J.T. and Wyman, J. 1958. Biophysical Chemistry, pp. 591-662. Academic Press, New York.
Atha, D.H. and Ingham, K.C. 1981. Mechanism of precipitation of proteins by polyethylene glycols. J. Biol. Chem. 256:12108-12117.
Fiabane, A.M. and Williams, D.R. 1977. The Principles of Bio-Inorganic Chemistry. In Chemical Society Publication no. 31, pp. 32-48. The Chemical Society, London.
Berg, J.M. and Shi, Y. 1996. The galvanization of biology: A growing appreciation for the roles of zinc. Science 271:1081-1085. Bensadoun, A. and Weinstein, D. 1976. Assay of proteins in the presence of interfering materials. Anal. Biochem. 70:241-250. Berdick, M. and Morawetz, H. 1954. The interaction of catalase with synthetic polyelectrolytes. J. Biol. Chem. 206:959-971. Bickle, T.A., Pirotta, V., and Imber, R. 1977. A single, general procedure for purifying restriction endonucleases. Nucl. Acids Res. 4:25612572. Budavari, S. (ed.) 1989. The Merck Index, 11th ed. Merck & Co., Rahway, N.J. Chakrabarti, P. 1993. Anion binding sites in protein structures. J. Mol. Biol. 234:463-482. Cheng, K.L., Ueno, K., and Imamura, T. (eds.) 1992. CRC Handbook of Organic Analytical Reagents, pp. 219-226. CRC Press, Boca Raton, Fla. Cohn, E.J., Strong, L.E., Hughes, W.L., Mulford, D.J., Ashworth, J.N., Melin, M., and Taylor, H.L. 1946. Preparation and properties of serum and plasma proteins. IV. A system for separation into fractions of the protein and lipoprotein components of biological tissue and fluids. J. Am. Chem. Soc. 68:459-475. Cohn, E.J., Gurd, F.R., Surgenor, D.M., Barnes, B.A., Brown, R.K., Derouaux, G., Gillespie, J.M., Kahnt, F.W., Lever, W.F., Liu, C.H., Mittelman, D., Mouton, R.F., Schmid, K., and Uroma, E. 1950. A system for separation of the components of human blood: Quantitative procedures for separation of the protein components of human plasma. J. Am. Chem. Soc. 72:465-474. Collingwood, T.N., Daniel, R.M., and Langdon, A.G. 1988. An M(III) facilitated flocculation technique for enzyme recovery and concentration. J. Biochem. Biophys. Methods 17:303-310. Collins, K.D. and Washabaugh, M.W. 1985. The Hofmeister effect and the behavior of water at interfaces. Q. Rev. Biophys. 18:323-422. Conroy, M.J. and Lovrien, R.E. 1992. Matrix coprecipitating and cocrystallizing ligands (MCC ligands) for bioseparations. J. Crystal Growth 122:213-222. Cotton, F.A. and Wilkinson, G. 1988. Advanced Inorganic Chemistry, 5th ed., pp. 606-608. Wiley-Interscience, New York. Dubin, P., Ross, T.D., Sharma, I., and Yegerlehner, B. 1987. Coacervation polyelectrolyte-protein complexes. In ACS Symposia Series 342 (W. Hinze and D. Armstrong, eds.), pp. 162-169. American Chemical Society, Washington, D.C.
Felix, K. 1960. Proamines. Adv. Protein Chem. 15:120.
Fink, A.L. 1995. Compact intermediate states in protein folding. Annu. Rev. Biophys. Biomol. Struct. 24:495-522. Gurd, F.R.N. and Wilcox, P.E. 1956. Complex formation between metallic cations and proteins, peptides and amino acids. Adv. Protein Chem. 11:311-427. Hagerman, A.E. and Butler, L.G. 1978. Protein precipitation method for the quantitative determination of tannins. Agric. Food Chem. 26:809-812. Hagerman, A.E. and Butler, L.G. 1991. Tannins and lignins. In Herbivores: Their Interactions with Secondary Plant Metabolites, Vol. 1 (G. Rosenthal, ed.) pp. 355-388. Academic Press, New York. Herzfeld, J. 1996. Entropically driven order in crowded solutions: From liquid crystals to cell biology. Acc. Chem. Res. 29:31-37. Hidalgo, J. and Hansen, P.M.T. 1971. Selective precipitation of whey proteins with carboxymethylcellulose. J. Dairy Sci. 54:1270-1274. Hogness, T.R. and Johnson, W.C. 1954. Qualitative Analysis and Chemical Equilibrium, 4th ed., pp. 567-598. Henry Holt Co., New York. Hunt, J.B., Neece, S.H., and Ginsburg, A. 1985. The use of 4-(2-pyridylazo)resorcinol in studies of zinc release from E. coli aspartate transcarbamylase. Anal. Biochem. 146:150-157. Jendrisak, J. 1987. Use of polyethyleneimine in protein purification. In Protein Purification: Micro to Macro (R. Burgess, ed.) pp. 75-97. Alan R. Liss, New York. Jendrisak, J.J. and Burgess, R.R. 1975. A new method for the large scale purification of wheat germ DNA-dependent RNA polymerase II. Biochemistry 14:4639-4645. Kressman, T.R. 1957. Commercial materials. In Ion Exchangers in Organic and Biochemistry (C. Calmon and T. Kressman, eds.) pp. 107-109. Interscience Publishers, New York. Laurent, T.C. 1963. The interaction between polysaccharides and other macromolecules. Biochem. J. 89: 253-257. Lovrien, R., Goldensoph, C., Anderson, P.C., and Odegaard, B. 1987. Three phase partitioning (TPP) via t-butanol: Enzyme separations from crudes. In Protein Purification: Micro to Macro (R. Burgess, ed.), pp. 131-148. Alan R. Liss, New York. Mattiasson, B. and Kaul, R. 1994. One-pot purification by process integration. Biotechnology 12:1088-1089.
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Matulis, D. and Lovrien, R. 1996. Coprecipitation of proteins with matrix ligands: Scaleable protein isolation. J. Mol. Recognit. In press.
Scopes, R.K. 1987. Protein Purification: Principles and Practice, 2nd ed., pp. 45-54. Springer-Verlag, New York.
McPherson, A. 1982. Preparation and Analysis of Protein Crystals, pp. 102-108. John Wiley & Sons, New York.
Sela, M. and Katchalski, E. 1959. Biological properties of poly-α-amino acids. Adv. Protein Chem. 14:391-478.
Miekka, S.I. and Ingham, K.C. 1978. Influence of self-association proteins on their precipitation by polyethylene glycol. Arch. Biochem. Biophys. 191:525-536.
Shibata, T., Cunningham, R.P., and Radding, C.M. 1981. Homologous pairing in genetic recombination. J. Biol. Chem. 256:7557-7564.
Morawetz, H. and Hughes, W.L., Jr. 1952. The interaction of proteins with synthetic polyelectrolytes. I. Complexing of serum albumin. J. Phys. Chem. 56:64-69. Morton, R.K. 1950. Separation and purification of enzymes associated with insoluble particles. Nature 166:1092-1095. Niederauer, M.Q., Suominen, I., Rougvie, M.A., Ford, C.F., and Glatz, C.E. 1994. Characterization and polyelectrolyte precipitation of βgalactosidase containing genetic fusions of charged polypeptides. Biotechnol. Prog. 10:237245. Patrickios, C.S., Hertler, W.R., and Hatton, T.A. 1994. Protein complexation with acrylic polyampholytes. Biotechnol. Bioeng. 44:10311039. Perrin, D.D. and Dempsey, B. 1974. Metal-ion buffers. In Buffers for pH and Metal Ion Control, pp. 94-102. Chapman & Hall, London. Putnam, F.W. and Neurath, H. 1944. The precipitation of proteins by synthetic detergents. J. Am. Chem. Soc. 66:692-697. Rothstein, F. 1994. Differential precipitation of proteins. In Protein Purification Process Engineering (R.G. Harrison, ed.) pp. 115-208. Marcel Dekker, New York. Sandell, E.B. and Onishi, H. (eds.) 1978. Photometric Determination of Traces of Metals, 4th ed. John Wiley & Sons, New York.
Singh, R.K. 1995. Precipitation of biomolecules. In Bioseparation Processes in Foods (R.K. Singh and S.S. Rizvi, eds.) pp. 139-173. Marcel Dekker, New York. Sternberg, M.Z. 1970. The separation of proteins with heteropolyacids. Biotechnol. Bioeng. 12:117. Sternberg, M. 1976. Purification of industrial enzymes with polyacrylic acids. Process Biochem. 11:11-12. Sternberg, M. and Hershberger, D. 1974. Separation of proteins with polyacrylic acids. Biochim. Biophys. Acta 342:195-206. Tanford, C. 1961. Multiple equilibria. In Physical Chemistry of Macromolecules, pp. 561-564. John Wiley & Sons, New York. Timasheff, S.N. 1992. Water as ligand: Preferential binding and exclusion of denaturants in protein unfolding. Biochemistry 31:9857-9864. Walter, H., and Johansson, G. 1994. Aqueous twophase systems. Methods Enzymol. 228: 1-17. Whitaker, D.R. 1953. Purification of M. verrcaria cellulase. Arch. Biochem. Biophys. 43: 253-267.
Contributed by Rex E. Lovrien and Daumantas Matulis University of Minnesota St. Paul, Minnesota
Selective Precipitation of Proteins
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Long-Term Storage of Proteins After a protein has been purified, there is often a need to store it for weeks or months before further study or use. For basic science investigations, it is critical that the protein’s physicochemical properties are not altered during storage. For example, oxidation of a methionine residue at the active site of an enzyme may render the protein inactive. Alternatively, a substantial fraction of the protein may precipitate and thus, no longer be usable for studies of protein structure and function. The means by which proteins degrade and the optimal methods to prevent damage during storage have been most extensively studied with recombinant proteins that are used as therapeutics. For these proteins to be approved for human use, each type of degradation product must be characterized and must be sufficiently inhibited in the final formulation such that the product is still safe and effective after 18 to 24 months of storage. A major problem is the formation of nonnative protein aggregates, which even at relatively low levels (e.g., 1% to 2% of total dose) may cause adverse responses in patients. Conversely, with some proteins, a given type of chemical degradation (e.g., deamidation) may not alter activity or safety. Because of the stringent stability requirements for therapeutic proteins, great insight into how to minimize protein degradation has been obtained from the study of these systems. The purpose of this overview is to provide a summary of some of this information as it relates to the issues that researchers face when attempting to store purified proteins. First, the stresses that induce protein aggregation and the pathway or pathways for this route of damage will be treated briefly. Then, the major chemical degradation routes for proteins will be highlighted. Finally, the use of various storage strategies to increase the long-term stability of proteins will be explained. When appropriate as part of this discussion, the authors will point out critical mistakes to avoid.
PROTEIN AGGREGATION In many academic labs, the first indication that a protein has stability problems is the appearance of precipitates, which usually contain nonnative protein aggregates. Such aggregation is usually considered irreversible, because the native protein conformation typically
cannot be restored unless the aggregates are dissolved in highly concentrated chaotrope solutions and the protein molecules refolded by removal of the chaotrope. Aggregation and precipitation of proteins can be induced by numerous different acute stresses including exposure to interfaces (e.g., air-water interface during mixing or agitation or liquid-solid interface during filtration), reduction in pH, freeze-thawing, exposure to subdenaturing concentrations of chaotropes, and exposure to elevated temperature. These stresses usually cause perturbation of the protein’s native tertiary structure and the formation of aggregation-competent, partially unfolded molecules. However, it is important to realize that aggregation can occur on reasonable time scales (e.g., days to weeks) under “physiological conditions” (e.g., pH 7 at 37°C), under which the native protein conformation is highly favored thermodynamically. Recent research has documented that aggregates can form from protein molecules that are only slightly perturbed from the most compact native conformation (e.g., Kendrick et al., 1998; Kim et al., 2000; Webb et al., 2001). Thus, during long-term storage in aqueous solutions, aggregation of proteins is a major problem, even if steps are taken to avoid exposure of the protein to obvious acute stresses. Nonnative protein aggregates, which are the precursors of visible precipitates, can remain in solution if they are sufficiently small (e.g., dimers, tetramers, octamers). Soluble aggregates can be detected readily with size exclusion chromatography and/or light-scattering techniques. It is important to know their levels because they may have altered structure and/or activity compared to the native protein. More importantly, soluble aggregates, even at relatively low levels (e.g., only 2% to 3%), can assemble into nuclei that promote rapid aggregation and precipitation of the protein. Such processes explain why a protein solution that appears to be “stable” for weeks or months, with only slight loss of activity, can suddenly form massive amounts of precipitate. It is thought that precipitation does not occur until soluble aggregates reach a threshold level that triggers nucleation. Alternatively, exposure of a solution containing soluble aggregates to an acute stress such agitation or filtration can rapidly promote nucleation and precipitation.
Contributed by John F. Carpenter, Mark C. Manning, and Theodore W. Randolph Current Protocols in Protein Science (2002) 4.6.1-4.6.6 Copyright © 2002 by John Wiley & Sons, Inc.
UNIT 4.6
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CHEMICAL DEGRADATION
Long-Term Storage of Proteins
Although the multiple functionalities of amino acid side chains could result in a wide range of chemical modifications, oxidation and hydrolysis reactions are the most prevalent (Manning et al., 1989; Volkin et al., 1997; Jaenicke, 2000). Oxidation occurs most readily at methionine and cysteine residues as well as those that possess aromatic side chains (i.e., tryptophan, histidine, tyrosine). Hydrolysis can occur both at amino acid side chains, as in the case of deamidation reactions at asparagine and glutamine residues, or along the peptide backbone, resulting in cleavage of a peptide group. Oxidation can arise readily from either the presence of molecular oxygen or from oxidative contaminants in the bulk drug, excipients, or containers (Cleland et al., 1993). For example, buffers can possess trace quantities of redox active metal ions that can foster oxidation. In addition, stoppers are known to contain residual free radical impurities that can lead to oxidative damage. Oxidation occurs most readily at residues that are highly solvent exposed, but even interior residues can oxidize during long-term storage in aqueous solution. The rate of oxidation of partially exposed residues can be retarded by compaction of the native state via addition of certain stabilizing additives such as sucrose (DePaz et al., 2000; Hovorka et al., 2001). Deamidation of asparagine residues is highly dependent on neighboring residues, where glycine in the following position displays the fastest rate of degradation (Patel and Borchardt, 1990a). Local flexibility of the polypeptide chain (Kossiakoff, 1988; Kosky et al., 1999) can also influence deamidation. Most importantly, extrinsic factors, such as pH and buffer species, can accelerate the reaction, with the rates being especially rapid at alkaline pH and in the presence of high concentrations of certain buffers, such as phosphate (Patel and Borchardt, 1990b). Nonenzymatic hydrolysis of the peptide backbone (termed proteolysis) is most likely to occur at asparagine residues, especially when flanked by proline (Manning et al., 1989). In addition, the reaction is both acid- and basecatalyzed. Other less common chemical reactions that have been observed during long-term storage of proteins includes disulfide exchange (Costantino et al., 1994), racemization (Senderoff et al., 1998), and beta-elimination (Manning et al., 1989; Volkin et al., 1997).
STORAGE IN NONFROZEN AQUEOUS SOLUTIONS The most convenient way to store proteins is in nonfrozen aqueous solutions in the refrigerator (i.e., at 4° to 8°C). For short-term storage or for exceptionally stable proteins this may be a viable approach; however, it must be realized that even if the protein does not aggregate, there may be progressive chemical degradation (especially oxidation). Aggregation can be minimized with solution conditions that optimize thermodynamic stability of the native state, because the protein population is shifted away from partially folded, aggregation-competent species (e.g., Kim et al., 2000). For example, most proteins have a relatively narrow pH range over which the free energy of unfolding is maximum. Also, specific ligands that bind avidly to the native state (e.g., calcium or co-factors) can greatly increase native state stability. Nonspecific stabilizing additives, such as sucrose, are also beneficial, but usually these compounds must be employed at concentrations of at least 0.5 M to be effective. These compounds are preferentially excluded from the surface of protein molecules with a concomitant increase in protein chemical potential (Timasheff, 1998). The magnitude of exclusion and chemical potential increase are directly proportional to the surface area of the protein molecule and, thus, are greater for partially or fully unfolded states than for the native conformation. As a result, the free energy barrier between native and unfolded states is increased. Numerous additives work through the preferential exclusion mechanism including salting-out salts (e.g., ammonium sulfate), amino acids (e.g., glycine), polyols (e.g., glycerol) and sugars (e.g., sucrose). One compound that appears to be particularly effective at conferring thermodynamic stability is citrate. The mechanism for its action has not been studied, but it most likely works by preferential exclusion. The rate of protein aggregation can also be reduced by storage of a protein at reduced concentration. Aggregation is usually a second- or higher-order kinetic process, and thus its rate is very sensitive to protein concentration. However, protein concentration should not be reduced too low (e.g., < ∼0.1 mg/ml) because of the risk of losing a substantial fraction of the protein by adsorption to the storage vessel walls. For many proteins, storage in solutions with high ionic strength (e.g., phosphate-buffered saline solution with ∼150 mM NaCl) favors the formation of nonnative aggregates. This is be-
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cause charge shielding by the ions in solution reduces the charge-charge repulsion between protein molecules. However, the actual effect of ionic strength on physical stability of a given protein must be determined empirically.
STORAGE AS SALTED-OUT PRECIPITATES A common means by which commercially available enzymes are shipped and stored is as salted-out precipitates. Salting-out (UNIT 4.5) is also routinely used to fractionate proteins during purification and as a storage method for proteins purified in research laboratories. Most often, ammonium sulfate at relatively high concentrations (e.g., 70% saturated) is used to precipitate the protein (APPENDIX 3F). The mechanism for salting-out is the same as that for increased thermodynamic stability by preferential exclusion of solutes. In the case of salting out, sufficient solute is added such that the protein’s chemical potential is high enough to greatly reduce its solubility. The structure of protein molecules remains native in salted-out precipitates. Upon dilution (e.g., dialysis) of the salting-out agent, the protein molecules dissolve and should be native and functional, without the need for refolding. Storage stability is greatly increased in salted-out precipitates relative to that for an aqueous solution of the same proteins. Often, with storage at 4°C, a salted-out protein preparation can remain stable for several months. Formation of nonnative aggregates is inhibited because the concentration of free protein molecules in solution is extremely low. Also, the salting-out solutes will greatly increase the thermodynamic stability of the native state of protein molecules. These two effects virtually eliminate partially folded, aggregation-competent protein molecules from the solution phase. Similarly, the conformational dynamics of salted-out protein molecules should be substantially reduced, which should serve to minimize the rate of chemical degradation, at least of interior residues. Although salted-out protein preparations are usually stored in the refrigerator, storage stability may be further enhanced by freezing the suspension. The salting-out compounds are potent cryoprotectants for proteins (see below) and freezing-induced damage should be minimized in a precipitate of native protein molecules. By reducing the temperature, the rate of chemical degradation can be slowed substantially.
STORAGE AS FROZEN SOLUTIONS Freezing of soluble protein preparations as a storage method is one of the most commonly used approaches. It is simple and convenient to place a test tube of protein solution into a mechanical freezer. At subzero temperatures, chemical degradation rates are usually reduced, and assuming that freezing itself did not denature the protein, the sample should be stable for weeks or months, especially if it is stored at −80°C or below. However, freezing can denature proteins because of several stresses including low temperature, concentration of solutes, formation of ice-liquid interfaces, and potentially drastic reductions in pH (see below). The risk of denaturation can be reduced by cryoprotectants in the solution. The mechanisms of nonspecific cryoprotection are the same as that for increasing thermodynamic stability in aqueous solution by additives (Heller et al., 1996), and the same compounds are effective in both instances. Examples of good cryoprotectants for proteins include glycerol, salting-out salts (see above), and disaccharides. The degree of protection against acute freeze-thawing stress usually correlates directly with the initial concentration of the solute. Often concentrations exceeding 0.3 M are needed for optimal protection. Another class of compounds that afford protection during freeze-thawing are surfactants such as polysorbates and polyethylene glycols, which are often optimally effective at concentrations as low 0.1% (w/v). It is thought that surfactants inhibit protein denaturation by competing with protein molecules for ice-liquid interfaces (Chang et al., 1996). Other proteins (e.g., BSA) and synthetic polymers can also inhibit freezing-induced protein unfolding at relatively low concentrations of approximately a few weight percent. Although these compounds are preferentially excluded from the surface of the protein, the molal concentration is not sufficiently high to alter protein chemical potential significantly by this mechanism. Rather, it appears that polymers inhibit protein denaturation during freezing by sterically hindering unfolding due to macromolecular crowding. The resistance of a protein preparation to damage during freeze-thawing can be increased by increasing the initial protein concentration. If a fraction of the protein damage noted in a given frozen-thawed sample is due to protein adsorption to denaturing ice-liquid interfaces, this damage is finite for a given
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sample volume. Thus, the percentage of protein molecules denatured will be less as the initial protein concentration is increased. Also, protection by the excluded volume effects discussed above can apply to individual protein molecules in a solution of the same protein. To test if a given solution is appropriate for frozen storage, small aliquots can be frozen and then thawed the next day. In general, if the protein survives the acute stresses of freezethawing, it will have relatively good storage stability when stored frozen in the same solution. Storage at −80°C is preferred over −20°C. First, the lower temperature will slow degradative reactions to a much greater degree. Second, the freezer compartment of refrigerators is often not −20°C in many areas, especially inside the door. Furthermore, in a frost-free freezer the sample may be subjected to repeated freeze-thawing. Finally, −20°C is near the eutectic temperature of certain salts, and cycling of temperature above and below this temperature can cause melting and reformation of the eutectic crystals, which might denature the protein. Repeated freeze-thawing can progressively increase protein denaturation in a sample and should be avoided. Instead, the protein solution should be divided into numerous small aliquots that can be removed as needed, thawed, and then used. The most common mistake that is made when researchers freeze proteins for storage is to use sodium phosphate buffer. During freezing, the dibasic salt crystallizes, while the acidic salt remains soluble. Thus, in frozen solutions the pH of sodium phosphate can drop very low (e.g., pH 4), which can cause denaturation of many proteins (e.g., Anchordoquy and Carpenter, 1996). Buffers that are less likely to undergo pH changes during freezing include citrate, Tris, and histidine.
STORAGE AS FREEZE-DRIED SOLIDS
Long-Term Storage of Proteins
A properly formulated and freeze-dried protein preparation can remain stable for years, even at room temperature (Carpenter and Chang, 1996); however, development of optimal solution compositions and processing conditions for a freeze-dried formulation is well beyond the scope of the typical academic lab. One main difficulty is that the type of lyophilizers available in most labs, which usually have no capacity to control sample temperature, cannot achieve the low level of residual water (e.g., <1% to 2% by mass) that is required for long-
term storage stability of freeze-dried products. To dry samples sufficiently, their temperature must be increased to greater than room temperature during the terminal stages of the freeze-drying process. Also, without temperature control during drying the sample can collapse into a clump, which makes it even more difficult to remove water. Even if appropriate processing conditions can be developed with a given lyophilizer, it is critical to use the correct additives to prevent protein denaturation during both freezing and drying (reviewed in Carpenter and Chang, 1996; Carpenter et al., 1997). The disaccharides sucrose and trehalose are very effective at protecting proteins during these stresses and during storage in the dried solid. These sugars are nonreducing. Reducing sugars such as glucose and maltose should not be used since they degrade proteins via the Maillard reaction. The degree of protection during freezing depends on the initial bulk concentration of sugar present, whereas that during drying depends on the sugar:protein mass ratio (Carpenter and Chang, 1996). As noted above, freezing protection is due to preferential exclusion of the sugar from the protein’s surface, which increases the free energy of unfolding. During drying, protein molecules unfold as the water hydrating their surface is removed. Sugars prevent this damage by hydrogen bonding to the dried protein in place of the lost water. The storage stability of a dried formulation depends on a low residual water content (e.g., <1% to 2% by mass), retention of the native protein conformation during freezing and drying (which can be accomplished with protective additives) and storage of the samples below the glass transition temperature of the amorphous phase, which contains the protein and stabilizing sugars (Carpenter and Chang, 1996). Measuring these physical parameters is often beyond the capabilities of most academic labs. Thus, for most labs freeze-drying should be considered only when all other methods for enhancing storage stability have been found to be inadequate. Assuming that suboptimal processing will be necessary (i.e., there is no way to control sample temperature in the freezedrier) and that critical physical parameters cannot be measured, is there any chance of obtaining a stable dried formulation? The answer is “maybe.” The practical approach is to prepare the protein at a relatively high concentration (which increases resistance to freezing and reduces volume to be dried) with sufficient trehalose or sucrose to inhibit unfolding during
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freezing and drying. If the protein preparation is resistant to freezing, usually a sugar:protein mass ratio in the range of 1 to 2 is adequate to prevent protein unfolding during drying. If protection is required during freezing, often an initial sugar concentration of 200 to 300 mM is required for optimal protection. It may also be beneficial to include a carbohydrate polymer (e.g., dextran, Ficoll, hydroxyethyl starch), which can provide sample bulk and increase the glass transition temperature of the amorphous phase. Sometimes adding a bulking agent is needed to prevent protein from escaping from the container during drying. Also, these polymeric amorphous bulking agents make it easier to dry the samples without collapse. An initial polymer concentration of 2% to 3% (w/v) usually provides adequate bulking. The samples are usually frozen by either placing them into a mechanical freezer or by immersing the sample containers into liquid nitrogen. It is preferred to aliquot the preparation into several small test tubes (e.g., 1.5-ml polypropylene microcentrifuge tubes), rather than trying to dry a relatively large volume in a few containers. The frozen samples should be placed into a container (e.g., desiccator or flask), which is attached to the lyophilizer, and immediately exposed to the system’s vacuum. Sublimation cooling should keep the samples frozen during the initial phase of the process when ice is removed. After ice is removed the sample temperature will increase to room temperature, during which time water is partially desorbed from the remaining non-ice phases. The samples should be kept under vacuum for sufficient time to “complete” the drying process. Drying is never actually complete. Rather the water content eventually reaches a minimal level, after which more time under vacuum at room temperature will not facilitate further drying. The water content resulting after drying at room temperature in sucrose or trehalose formulations is usually >4% to 5% by mass. The actual duration of drying needed must be determined empirically, but this can be a problem if the required instrumentation is not available. To be safe, for typical samples with a volume of 0.5 ml for example, it is advisable to keep samples under vacuum for at least 1 to 3 days. Also, to minimize degradation of the protein, it is recommended that the dried samples be stored in the freezer at −20°C or, even better, at −80°C. Again, all of this effort seems to be excessive compared to simply freezing the solutions, es-
pecially since subzero storage temperatures probably will be required for suboptimally freeze-dried samples.
SELECTING AN APPROPRIATE STORAGE METHOD The choice of storage method depends on several criteria, including the intrinsic stability of the protein, the duration of storage required, and how stringent the need for preventing damage is. There is tremendous variability in these criteria. However, the following general guidelines should apply in most instances. If the goal is simply to keep a small batch of purified protein around for a few days, then often storage in unfrozen solution at 4°C is sufficient. If greater duration of storage is required or if there is obviously protein degradation (e.g., visible precipitates or unacceptable loss of function), then the next option is to choose either frozen storage or storage as a salted-out precipitate (UNIT 4.5; APPENDIX 3F). Both methods are relatively straightforward and, when conducted properly, should afford several weeks or months of storage stability. Prior to attempting to design a salting out strategy, preliminary freeze-thawing experiments should be conducted to determine a given protein preparation’s resistance to this stress. If the protein is stable during this treatment, then long-term storage in the frozen state will probably provide adequate stability. However, if preliminary experiments document that the protein is sensitive to acute freeze-thawing stress, it will be necessary to test the appropriate cryoprotectants or to opt to explore the utility of salting out the protein. The latter is probably somewhat easier, but it does not take a large effort to optimize freeze-thawing resistance. At this point, the choice of method to use depends on prior experience with the protein, as well as the history of how a given lab has successfully dealt with protein storage stability issues. Finally, only if all other methods fail or if there is a need for very long-term storage (e.g., 18 to 24 months) at ambient temperatures should freeze-drying be explored.
INHIBITION OF PROTEASES This overview has been written with the assumption that purification of the protein was good enough that detrimental levels of proteases were removed. Of course, even with highly refined purification protocols that result in preparations of >99% homogeneity, there still may be sufficient protease activity that the “purified” protein is rapidly hydrolyzed. Hy-
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drolysis is usually easily recognized with SDS polyacrylamide electrophoresis (UNIT 10.1), which documents the presence of protein fragments. Enzymatic fragmentation proceeds relatively rapidly (e.g., on a time scale of hours or days) compared to nonenzymatic hydrolysis. If proteolysis occurs then the protein will have to be purified further or protease inhibitors will have to be added to the protein preparation. A comprehensive guide to selection and use of protease inhibitors can be found at the Roche Molecular Biochemicals web site. The URL for specific descriptions of proteases and their inhibitors is: http://biochem.roche.com/prodinfo_fst.htm/ proteaseinhibitor/prod07.htm.
Kendrick, B.S., Carpenter, J.F., Cleland, J.L., and Randolph, T.W. 1998. A transient expansion of the native state precedes aggregation of recombinant human interferon-gamma. Proc. Natl. Acad. Sci. U.S.A. 95:14142-14146.
LITERATURE CITED
Manning, M.C., Patel, K., and Borchardt, R.T. 1989. Stability of protein pharmaceuticals. Pharm. Res. 6:903-918.
Anchordoquy, T.J. and Carpenter, J.F. 1996. Polymers protect lactate dehydrogenase during freeze-drying by inhibiting dissociation in the frozen state. Arch. Biochem. Biophys. 332:231-238. Carpenter, J.F. and Chang, B.S. 1996. Lyophilization of protein pharmaceuticals. In Biotechnology and Biopharmaceutical Manufacturing, Processing and Preservation (K. Avis and V. Wu, eds.) pp. 199-263. Intepharm Press, Buffalo Grove, Ill. Carpenter, J.F., Pikal, M.J., Chang, B.S., and Randolph, T.W. 1997. Rational design of stable lyophilized protein formulations: Some practical advice. Pharm. Res. 14:969-975. Chang, B.S., Kendrick, B.S., and Carpenter, J.F. 1996. Surface-induced denaturation of proteins during freezing and its inhibition by surfactants. J. Pharm. Sci. 85:1325-1330. Cleland, J.L., Powell, M.F., and Shire, S.J. 1993. The development of stable protein formulations: A close look at protein aggregation, deamidation, and oxidation. Crit. Rev. Ther. Drug Carrier Syst. 10:307-377. Costantino, H.R., Langer, R., and Klibanov, A.M. 1994. Solid-phase aggregation of proteins under pharmaceutically relevant conditions. J. Pharm. Sci. 83:1662-1669. DePaz, R.A., Barnett, C.C., Dale, D.A., Carpenter, J.F., Gaertner, A.L., and Randolph, T.W. 2000. The excluding effects of sucrose on a protein chemical degradation pathway: Methionine oxidation in subtilisin. Arch. Biochem. Biophys. 384:123-132. Heller, M., Carpenter, J.F., and Randolph, T.W. 1996. Effect of phase separating systems on lyophilized hemoglobin. J. Pharm. Sci. 85:1358-1362. Hovorka, S.W., Hong, J., Cleland, J.L., and Schoneich, C. 2001. Metal-catalyzed oxidation of human growth hormone: Modulation by solvent-induced changes of protein conformation. J. Pharm. Sci. 90:58-69. Long-Term Storage of Proteins
Jaenicke, R. 2000. Stability and stabilization of globular proteins in solution. J. Biotech. 79:193-203.
Kim, Y.-S., Wall, J.S., Meyer, J., Murphy, C., Randolph, T.W., Manning, M.C., Solomon, A., and Carpenter, J.F. 2000. Thermodynamic modulation of light chain amyloid fibril formation. J. Biol. Chem. 275:1570-1574. Kosky, A.A., Razzaq, U.O., Treuheit, M.J., and Brems, D.N. 1999. The effects of alpha-helix on the stability of Asn residues: Deamidation rates in peptides of varying helicity. Protein Sci. 8:2519-2523. Kossiakoff, A.A. 1988. Tertiary structure is a principal determinant to protein deamidation. Science 240:191-194.
Patel, K. and Borchardt, R.T. 1990a. Chemical pathways of peptide degradation. III. Effect of primary sequence on the pathways of deamidation of asparaginyl residues in hexapeptides. Pharm. Res. 7:787-793. Patel, K. and Borchardt, R.T. 1990b. Chemical pathways of peptide degradation. II. Kinetics of deamidation of an asparaginyl residue in a model hexapeptide. Pharm. Res. 7:703-711. Senderoff, R.I., Kontor, K.M., Kreilgaard, L., Chang, J.J., Patel, S., Krakover, J., Heffernan, J.K., Snell, L.B., and Rosenberg, G.B. 1998. Consideration of conformational transitions and racemization during process development of recombinant glucagon-like peptide-1. J. Pharm. Sci. 87:183-189. Timasheff, S.N. 1998. Control of protein stability and reactions by weakly interacting cosolvents: The simplicity of the complicated. Adv. Prot. Chem. 51:355-432. Volkin, D.B., Mach, H., and Middaugh, C.R. 1997. Degradative covalent reactions important to protein stability. Mol. Biotechnol. 8:105-122. Webb, J.N., Webb, S.D., Cleland, J.L., Carpenter, J.F., and Randolph, T.W. 2001. Partial molar volume, surface area, and hydration changes for equilibrium unfolding and formation of aggregation transition state: High-pressure and cosolute studies on recombinant human IFN-γ. Proc. Natl. Acad. Sci. U.S.A. 98:7259-7264.
Contributed by John F. Carpenter and Mark C. Manning University of Colorado Health Sciences Center Denver, Colorado Theodore W. Randolph University of Colorado Boulder, Colorado
4.6.6 Supplement 27
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Extraction of Proteins from Plant Tissues Proteins are extracted from plant tissues for a wide range of reasons, including to assay an enzyme in a crude extract for physiological studies, to purify a protein in order to identify the gene that encodes it, and to resolve plant proteins by SDS-PAGE. Plants differ from animals in that they have a rigid cellulose cell wall, which may be further strengthened by lignification. In addition, they often have a large vacuole that contains segregated secondary plant products (especially phenolics and polyphenolics), organic acids, and proteinases. These vacuoles are broken upon grinding, releasing contents that may modify, inactivate, precipitate, or degrade proteins. Consequently, special techniques are required to disrupt the cell walls and to protect proteins from damaging components released on breakage (see Critical Parameters).
UNIT 4.7 BASIC PROTOCOL
Materials Frozen or fresh plant material Liquid nitrogen Extraction buffer (see recipe) Mortar and pestle or tissue grinder (Freezer/mill from SPEX CertiPrep or equivalent) 20- to 70-µm pore size nylon mesh (screen printing mesh) or Calbiochem Miracloth for filtration Small paintbrush Refrigerated centrifuge and centrifuge tubes NOTE: All operations at should be carried out at 0◦ to 4◦ C (i.e., in a cold room, or on ice) unless there are reasons to do otherwise. The extraction procedure should be performed rapidly to minimize exposure of proteins of interest to potentially damaging compounds and enzymes released upon cell breakage.
Disrupt tissue 1. Grind frozen plant material to a fine powder using a mortar and pestle prechilled to −186◦ C in liquid nitrogen, or in a tissue grinder designed to work with liquid nitrogen. It is easiest to grind without having the liquid nitrogen come into contact with the tissue. To ensure that the sample stays frozen, the mortar may be placed in a shallow pool of liquid nitrogen (e.g., in the lid of a standard polystyrene box). Fresh material may be ground directly in ice-cold buffer. A tissue homogenizer is especially effective for grinding fresh plant material. Other options include a blender or a mortar and pestle plus acid-washed sand. Foaming can be a problem when blending, and some measure of control may be achieved by adding a few drops of n-octanol. CAUTION: Wear gloves and safety glasses to avoid burns while working with liquid nitrogen.
2. Transfer the frozen sample powder to ice-cold extraction buffer, mix quickly, and grind further with a chilled mortar and pestle or a tissue homogenizer. In most cases, use a tissue/buffer ratio equal to 1:5 (w/v) (see Critical Parameters). Do not add buffer to the −186◦ C mortar and pestle, as the sample will then take a very long time to thaw. Instead, add the frozen powder to the extraction buffer in an ice-cold container and mix the buffer into the powder quickly, before thawing of the powder occurs. Use a paintbrush to aid the transfer. Extraction, Stabilization, and Concentration Contributed by William Laing and John Christeller Current Protocols in Protein Science (2004) 4.7.1-4.7.7 C 2004 by John Wiley & Sons, Inc. Copyright
4.7.1 Supplement 38
Purify extract 3. Filter the extract immediately through a 20- to 70-µm nylon mesh (as used in screen printing) or Miracloth, squeezing by hand to remove cell walls and other debris. For particularly large volumes, a press, such as a hand-pumped cider press, can be used.
4. To remove insoluble material from the filtered sample, centrifuge large-volume extracts >10 min at 30,000 × g, 4◦ C; microcentrifuge small-volume extracts 10 min at maximum speed, 4◦ C. 5. Decant the supernatant carefully, watching out for an unstable pellet. Chlorophyll will be solubilized if there is a detergent in the extraction buffer (see Reagents and Solutions); if no detergent is present, most chlorophyll will be in the pellet. A white precipitate may be present, consisting of starch and/or insoluble polyvinylpyrrolidone.
6. If quantitative recovery is important, extract the pellet again with more buffer, filter, and centrifuge again. Combine the supernatants. 7. Depending on the purpose of the experiment, perform additional standard protein purification steps (see Chapter 4) and/or direct analysis by SDS-PAGE (UNIT 10.1) on the supernatant. Alternatively, if the protein is being extracted for resolution by SDS-PAGE (Sch¨agger and von Jagow, 1987), the tissue can be ground until finely powdered (as described in step 1), a small amount (e.g., 50 mg) quickly weighed and placed into a tube, and a suitable volume of hot SDS sample buffer added. This is mixed and immediately heated to denature proteins, centrifuged, and applied to the gel.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Extraction buffer 0.2 M 3-(N-morpholino)propanesulfonic acid (MOPS), pH 7.0 5% (w/w) polyvinyl polypyrrolidone (PVPP; weigh out, add to mixture, and allow to hydrate 1 hr before use) 1% (v/v) Triton X-100 (optional; see Critical Parameters) 10% (v/v) glycerol (optional; see Critical Parameters) Store buffer with above components up to several weeks at 4◦ C On day of use, add the following: 2 mM dithiothreitol (DTT) Suitable plant proteinase inhibitor cocktail (Sigma, Roche, or equivalent; add according to manufacturer’s instructions, or 30 min before tissue extraction; see Critical Parameters) COMMENTARY Background Information
Extraction of Proteins from Plant Tissues
There are two main purposes for extracting proteins from plant tissues: to assay protein levels or activity in a crude extract, or to purify relatively large quantities of an enzyme or protein. The methods chosen thus depend on purpose, tissue type, and enzyme or protein to be extracted. In this unit, various options are discussed for making appropriate modifications to cope with a particular
situation. The more that is known about the properties of the protein of interest, the easier it is to make choices regarding extraction methods. Generally, the better the health of the plant tissue, the greater the activity that will be extracted (unless the target protein is a stress- or disease-related protein). In addition, younger plants and unstressed tissues often have fewer interfering compounds.
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Critical Parameters Extraction buffer Buffer is required to maintain stability of enzymes from both a pH and an ionicstrength standpoint. Because plant cytoplasm is around pH 7 and chloroplasts are at a higher pH, it is generally optimal to maintain extracted enzymes at a similar, neutral pH. However, extraction at pH > 7 is not recommended (see below). Alternative buffers to MOPS include N-2-hydroxyethylpiperazineN -2-ethanesulfonic acid (HEPES), phosphate, or 2-(N-morpholino)-ethanesulfonic acid (MES), depending on the goals of the experiment and the need to avoid particular buffers. As the contents of plant cell vacuoles are often quite acidic, sufficient buffering capacity should be used to ensure the pH does not drop below the acceptable range for the protein of interest. The volume of extraction buffer used per unit weight of tissue depends on the tissue and the purpose of the extraction: e.g., the need for complete protein extraction versus the convenience of small volumes. Better extraction is obtained with a higher ratio of buffer volume to fresh tissue weight (e.g., 10 to 1). For largescale extractions, however, this often produces too large a volume to work with in subsequent steps unless the sample is first concentrated. An insufficient volume ratio of buffer risks changes in the pH of the extract and modification of the proteins in the tissues. A buffer containing 10% to 20% glycerol (v/v) may be necessary to ensure the stability of particularly unstable proteins. Some proteins (e.g., cell wall proteins and some seed storage proteins) are only solubilized in the presence of high salt concentrations, in the range of 0.1 to 1 M (Mika and L¨uthje, 2003). Antioxidants Antioxidants in the extraction buffer are essential to maintain the reduced state of free sulfhydryl groups in enzymes and to reduce oxidation of other components, such as phenolics. Options include DTT (2 mM or more), β-mercaptoethanol (5 to 10 mM), or ascorbic acid (5 to 10 mM), and in some circumstances, stronger reducing agents such as sodium dithionite (Melville and Ryan, 1972). More than one reducing agent can be used (e.g., DTT and β-mercaptoethanol). Antioxidants also reduce the activity of polyphenol oxidases that, in the presence of oxygen, rapidly synthesize polyphenols from phenolics upon tissue disruption. Polyphenol oxidase (PPO)
inhibitors, such as sodium diethyldithiocarbamate (1 to 5 mM) are effective against copper-containing PPOs. As plant tissues are often naturally high in ascorbic acid, it should be noted that ascorbic acid can be oxidized in the presence of metals, producing hydrogen peroxide, which can oxidize cysteine (Schmalhausen et al., 2003). The iron chelator deferoxamine is effective at stopping this oxidation, whereas EDTA actually promotes it (Schmalhausen et al., 2003). Detergents Detergents are used to disrupt membranes and are therefore essential for solubilizing membrane proteins (Mika and L¨uthje, 2003). However, they may also be useful for maximizing the yield of soluble proteins in some circumstances, especially when low ratios of extraction buffer to tissue are used. A series of detergents or other chaotropes can be used to carry out differential extractions. Examples include Tween 80 (0.1% to 1%), Triton X-100 (0.1% to 1%), or sometimes 3-[(3-cholamidopropyl)dimethylammonio]2-hydroxy-1-propanesulfo-nate (CHAPS) or octyl glycosides. Generally, initial empirical testing is needed to determine optimal combinations. Protective agents Protective agents are required to prevent polyphenols such as tannins (water-soluble polyphenols of varying molecular weight that are the most abundant polyphenols; Spencer et al., 1988; Singh et al., 2003) from binding to and inactivating proteins. The insoluble PVPP or soluble polyvinyl pyrrolidone (PVP) are useful in most applications (Loomis, 1974). These bind polyphenols best at pH values below 7 (Makkar et al., 1995). PVP is useful when making organelles or otherwise separating samples from the medium (by centrifugation or chromatography) early in the procedure. PVPP has the advantage that it is removed from the medium at the filtration and centrifugation steps. PVPP (0.5 to 5%, w/v) must be hydrated for ∼1 hr in the extraction buffer before use. Polyethylene glycol (e.g., PEG 6000) is also very useful for binding polyphenols. Proteinase inhibitors Proteinase inhibitors (PIs) are essential for controlling degradation of proteins following maceration. Cysteine proteinases are most widespread in plant tissues but metallo-, serine, and aspartic proteinases have been reported in different tissues. Furthermore, a
Extraction, Stabilization, and Concentration
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buffer pH of around 7 is conducive to cysteine proteinase activity, as is the presence of antioxidants. Some proteinase inhibitors are irreversible and chemically modify their targets (e.g., phenylmethylsulfonyl fluoride, PMSF), whereas others are tight binding (e.g., bovine pancreatic trypsin inhibitor, BPTI, also known as aprotinin) and some proteinase activity may be recovered in later steps if inhibitor is no longer present. This may cause problems if PIs are omitted in later purification steps. PMSF (1 mM), BPTI (1 µM), and 4(2-aminoethyl)-benzenesulfonyl-fluoride hydrochloride (AEBSF or PEFA-BLOC, 1 mM) are useful against serine proteinases. PMSF is toxic and rapidly hydrolyzed, and should be added to the extraction buffer just before tissue disruption. EDTA, 1,10-phenanthroline, or 2, 2 bipyridyl (both at 1 to 10 mM) are used against metalloproteinases, and trans-epoxysuccinylL-leucyl-amido-(4-guanidino)butane, (E64, 10 µM) is effective against cysteine proteinases. Antipain and leupeptin (both at 10 µM) have activity against all serine and cysteine classes. Pepstatin (10 µM) is effective against aspartic acid proteinases, but may not be necessary at pH values near 7, as aspartic proteinases generally function at low pH values. Commercial proteinase inhibitors (e.g., those provided in the form of tablets by Roche and as solutions by Sigma) containing a cocktail of these inhibitors are available, tailored for plants. These are a convenient way to add a range of these inhibitors. In cases where it is not desirable to add chelating agents, PIs without EDTA can be used. Note that PI tablets are slow to dissolve, and so should be added to buffer about 30 min before extracting the tissue. Exercise caution, as some proteinase inhibitors are toxic.
Extraction of Proteins from Plant Tissues
Other special considerations Filtration through Miracloth is effective, but this material rips easily and is rather expensive. Nylon mesh is extremely strong, can be washed and reused, and is readily available from screen printing supply companies. Whereas most plant proteins are not particularly heat- or acid-stable, some proteins can be extracted successfully at low pH and/or at high temperatures. For example, some small proteinase inhibitors are stable at pH 3 (Ryan et al., 2003), others at both pH 3 and 80◦ C for 6 min (Melville and Ryan, 1982). Tissues with high lipid levels, such as seeds, may need to be defatted before further purifica-
tion (Pastorello and Tranbaioli, 2001). Where the protein of interest can withstand high levels of organic solvent (e.g., as with proteinase inhibitors), the ground seeds can be defatted in acetone, the residue powder dried, and the sample then extracted in aqueous buffer. Enzyme inactivation is reduced by slowly adding very cold acetone during extraction. Rapid desalting of extracts Large-scale extracts greater than 0.2 liters (e.g., for protein purification) take longer to process because centrifugation, filtering, and other processes are slower. Consequently, there can be a significant problem with polyphenolics binding to proteins. By applying the centrifuged filtrate immediately to a large (e.g., 40 × 10 cm) G-75 Superdex or G-25 Sephadex column at a flow rate of least 5 ml/min, various low-molecular-weight polyphenols can be separated from proteins. The column should be equilibrated with a compatible buffer containing at least 100 mM buffer to ensure that proteins do not bind to the column. For the preparation of small tissue samples for crude assays, the extract can be desalted using small gel-filtration columns (e.g., PD-10 or NAP columns, Amersham Biosciences) equilibrated with assay buffer or another compatible buffer. All steps can be done in microcentrifuge tubes in a microcentrifuge placed in a cold room. Often, there is a requirement to both protect the enzyme from modification in the extraction media and to rapidly concentrate the protein for further work. The protein may be concentrated by standard salt or organic solvent precipitation (Brovko and Zagranichnaya, 1998; UNIT 4.5). Alternatively, the extract can be concentrated by dialysis against a volatile buffer (e.g., ammonium carbonate), by using a filter-based concentrator (UNIT 4.4), or by ion-exchange chromatography (UNIT 8.2). However, concentration still may be slow and may expose the protein to polyphenols and proteinases during these processes. Tissue types The types of tissue being extracted will affect the choice of method used and the yield of protein. Leaves and roots have a fresh weight of about 80% to 90% water, and can vary from those that have few “problem” compounds except proteinases (e.g., in spinach) to those requiring the full spectrum of protectants (e.g., pine needles). Whereas protein concentration in leaves can be high (1% to 4% of fresh
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weight), the concentration in fruit and stem tissue can be up to an order of magnitude lower, and large extraction buffer ratios will lead to very dilute protein concentrations in the extracts. Seeds, which are up to 90% dry weight with correspondingly high protein concentrations, often contain high amounts of lipids, which are difficult to separate from the aqueous extract. Fruits are often acidic, with high levels of polyphenolics, and need high buffer concentrations to maintain pH. The protein in this sort of tissue is often very dilute and needs concentration. Some fruits (e.g., kiwifruit, papaya) are also very high in cysteine proteinases. Tissue culture cells and single-celled algae can be difficult to break, and sonication, use of a French press, or a cell wall–digesting enzyme preparation (e.g., macerozyme) may be necessary. Vascular tissue needs to be separated from surrounding material, although cambium peels are sometimes possible. This can be done by peeling the bark away from the stem and scraping soft tissue from the inside of the bark (“phloem”) or the outside of the stem (“xylem”). Sometimes, specialized extracts can be obtained; for example, it is very easy to obtain large volumes of phloem sap from cucurbits (Murray and Christeller, 1995). It is more difficult to extract proteins from tissues with secondary thickening of their cell walls, such as woody tissues. Tissues can be subfractionated to enrich for the protein of interest (e.g., chloroplast or mitochondrial proteins) by density gradient fractionation of these organelles after tissue disruption. However, yields are usually low because short and gentle disruption methods are needed to break cell walls without damaging the membranes of the organelles. Membranes can be isolated by differential centrifugation (Rontein et al., 2003; Mika and L¨uthje, 2003)
before proteins are solubilized. Vacuoles, with their single membrane, are particularly fragile and difficult to isolate. A common method is to isolate vacuolar membranes (Yuasa and Maeshima, 2000; Heyen et al., 2002). These specialized methods are outside the scope of this unit. Each species may have unique problems. Spinach is regarded as benign and easy to disrupt and is often used to make delicate active chloroplasts, whereas apples, tobacco, and conifers are high in polyphenolics that crossreact with proteins. Arabidopsis is a model plant species and is quite easy to work with and extract proteins from (Che et al., 2003; Bonin et al., 1997), as are potato (Geigenberger et al., 1997) and tomato (Delmas et al., 2003).
Troubleshooting See Table 4.7.1 for a listing of common problems that may arise during extraction of protein from plant tissues and suggestions for how to overcome or avoid these problems; these suggestions are elaborated upon in Critical Parameters.
Anticipated Results Depending on the stability of the protein to be extracted or purified, active protein should be obtained easily from a wide variety of plant materials including leaves of tree species, woody tissue, and seeds (Table 4.7.2). Of course, there are exceptions, and totally different approaches may need to be taken. For example, the most difficult tissue the authors have encountered is the leaf of the New Zealand beech tree (Nothofagus species), where the best buffer was 50 mM borate, pH 7. For some reason, this buffer complexed compounds in the leaves that otherwise bound to proteins and inactivated them (W. Laing and D. Stevenson, unpub. observ.).
Table 4.7.1 Extraction Problems Common to Plants
Problem with plant tissue
Solutiona
Acidity (large acidic vacuoles)
Use high buffer amounts/weight tissue
Polyphenols
Use protective reagents
Oxidants
Use reducing reagents
Proteinases
Use proteinase inhibitors
Cell walls
Use efficient grinding techniques, liquid nitrogen
High lipid content (e.g., seeds) a See Critical Parameters for details.
Defat using organic solvents
Extraction, Stabilization, and Concentration
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Table 4.7.2 Examples of Proteins Extracted from Different Plants
Proteins assayed or purified
Points illustrated by assay
Reference
Cell suspension Arabidopsis thaliana cultures
GDP-mannose 3 ,5 -epimerase
Extraction and partial purification of an enzyme from cell culture
Wolucka et al. (2001)
Fruit cortex
Acid phosphatases
Illustrates some of the Turner and complications of a “difficult” tissue Plaxton (2001)
Fruit cortex and Actinidia deliciosa seeds (kiwifruit)
Phytocystatins
Extraction of robust proteinase inhibitors, capable of withstanding reversed-phase HPLC
Rassam and Laing (2004)
Fruit phloem exudate
Cucurbita maxima (squash)
Aspartic proteinase inhibitor
Extraction of a less robust proteinase inhibitor
Christeller et al. (1998)
Fruit tissues
Apple, kiwifruit, avocado
Proteins for two-dimensional electrophoresis
A general method for analytic separation of proteins from “difficult” tissues
Barraclough et al. (2004)
Leaf chloroplasts
Spinacia oleracea (spinach)
Glutamyl proteinase
Extraction involving isolation of intact chloroplasts as the first step in protein purification
Laing and Christeller (1997)
Leaves
Oryza sativa (rice)
Sucrose-1-phosphate phosphohydolase
Extraction and purification of an enzyme to homogeneity from a monocot
Lunn et al. (2000)
Seeds
Malus domestica (apple)
Trypsin and papain proteinase inhibitors
Extraction of robust proteinase Ryan et al. inhibitors, proteins capable of (2003) withstanding reversed-phase HPLC
Whole seedlings
Arabidopsis thaliana
26S proteasome
Purification of a complex protein from a model plant
Yang et al. (2004)
Wood/xylem
Picea sitchensis (Sitka spruce)
Laccase
Extraction of proteins from woody tissue
McDougall (2000)
Tissue
Plant
Musca cavendishii (banana)
Time Considerations In the authors’ laboratory, to purify proteins in large extracts, sequentially grinding two 100 g lots of fresh tissue, extracting the protein in extraction buffer, centrifuging, and passing the extract through a large G-75 column can all be accomplished within a day, with loading of the fractions containing the desired activity onto a Hitrap Q FF ion exchange column the same evening. Alternatively, it is reasonable to expect to be able to grind small amounts of tissue (100 mg fresh weight), extract the frozen powder in buffer, centrifuge, and apply to and elute from a NAP or PD-10 desalting column within 30 min. Where there is a series of samples to extract, ten samples can be extracted by two people within an hour.
Literature Cited Extraction of Proteins from Plant Tissues
Barraclough, D., Obenland, D., Laing, W., and Carroll, T. 2004. A method for quick and easy two-dimensional electrophoresis of plant samples. Postharvest Biol. Technol. 32:175-181.
Bonin, C.P., Potter, I., Vanzin, G.F., and Reiter, W.D. 1997. The MUR1 gene of Arabidopsis thaliana encodes an isoform of GDP-D-mannose-4,6dehydratase, catalyzing the first step in the de novo synthesis of GDP-L-fucose. Proc. Natl. Acad. Sci. U.S.A. 94:2085-2090. Brovko, F.A. and Zagranichnaya, T.K. 1998. Separation of proteins from phenols in cereal leaf extract by hydrophobic interaction: Ammonium sulfate fractionation. Plant Physiol. Bioch. 36:773-777. Che, P., Weaver, L.M., Wurtele, E.S., and Nikolau, B.J. 2003. The role of biotin in regulating 3-methylcrotonyl-coenzyme A carboxylase expression in Arabidopsis. Plant Physiol. 131:1479-1486. Christeller, J.T., Laing, W.A., Ramsay, R.J., Cutfield, J., Cutfield, S., and Sullivan, P. 1998. Purification and characterization of an aspartic acid proteinase inhibitor from squash phloem exudate. Euro. J. Biochem. 254:160-167. Delmas, F., Petit, J., Joub`es, J., S´eveno, M., Paccalet, T., Hernould, M., Lerouge, P. Mouras, A., and Chevalier, C. 2003. The gene expression and enzyme activity of plant 3-deoxy-D-manno2-octulosonic acid-8-phosphate synthase are
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preferentially associated with cell division in a cell cycle-dependent manner. Plant Physiol. 133:348-360. Geigenberger, P., Reimholz, R., Geiger, M., Merlo, L., Canale, V., and Stitt, M. 1997. Regulation of sucrose and starch metabolism in potato tubers in response to short-term water deficit. Planta 201:502-518. Heyen, B., Alsheikh, M.K., Smith, E.A., Torvik, C.F., Seals, D.F., and Randall, S.K. 2002. The calcium-binding activity of a vacuoleassociated, dehydrin-like protein is regulated by phosphorylation. Plant Physiol. 130:675-687. Laing, W.A. and Christeller, J.T. 1997. A plant chloroplast glutamyl proteinase. Plant Physiol. 114:715-722. Loomis, W.D. 1974. Overcoming problems of phenolics and quinones in the isolation of plant enzymes and organelles. Methods Enzymol. 31:528-544. Lunn, J.E., Ashton, A.R., Hatch, M.D., and Heldt, H.W. 2000. Purification, molecular cloning, and sequence analysis of sucrose-6F-phosphate phosphohydrolase from plants. Proc. Natl. Acad. Sci. U.S.A. 97:12914-12919. Makkar, H.P.S., Bl¨ummel, M., and Becker, K. 1995. Formation of complexes between polyvinyl pyrrolidones or polyethylene glycols with tannins, and their implications in gas production and true digestibility in in vitro techniques. Br. J. Nutr. 73:897-913. McDougall, G.J. 2000. A comparison of proteins from the developing xylem of compression and non-compression wood of branches of sitka spruce (Picea sitchensis) reveals a differentially expressed laccase. J. Exp. Bot. 349:1395-401. Melville, J.C. and Ryan, C.A. 1972. Chymotrypsin inhibitor I from potatoes: Large scale preparation and characterization of its subunit components. J. Biol. Chem. 247:3445-53. Mika, A. and L¨uthje, S. 2003. Properties of guaiacol peroxidase activities isolated from corn root plasma membranes. Plant Physiol. 132:14891498. Murray, C. and Christeller, J.T. 1995. Purification of a trypsin inhibitor (PFTI) from pumpkin fruit phloem exudate and isolation of putative trypsin and chymotrypsin inhibitor cDNA clones. Biol. Chem. Hoppe Seyler 376:281-287. Pastorello, E.A. and Trambaioli, C. 2001. Isolation of food allergens. J. Chromatogr. B. Biomed. Sci. Appl. 756:71-84. Rassam, M. and Laing, W. 2003. Purification and characterization of phytocystatins from kiwifruit cortex and seeds. Phytochemistry 65:19-30. Rontein, D., Wu, W.I., Voelker, D.R., and Hanson, A.D. 2003. Mitochondrial phosphatidylserine decarboxylase from higher plants: Functional complementation in yeast, localization in plants, and overexpression in Arabidopsis. Plant Physiol. 132:1678-1687. Ryan, S.N., McManus, M.T., and Laing, W.A. 2003. Identification and characterisation of proteinase
inhibitors and their genes from seeds of apple (Malus domestica). J. Biochem. 134:31-42. Sch¨agger, H., and von Jagow, G, 1987. Tricinesodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa. Anal. Biochem. 166:368-379. Schmalhausen, E.V., Pleten, A.P., and Muronetz, V.I. 2003. Ascorbate-induced oxidation of glyceraldehyde-3-phosphate dehydrogenase. Biochem. Biophys. Res. Comm. 308:492-496. Singh, B., Bhat, T.K., and Singh, B. 2003. Potential therapeutic applications of some antinutritional plant secondary metabolites. J. Agric. Food Chem. 51:5579-5597. Spencer, C.M., Cai, Y., Martin, R., Gaffney, S.H., Goulding, P.N., Magnolato, D., Lilley, T.H., and Haslam, E. 1988. Polyphenol complexation— some thoughts and observations. Phytochemistry 27:2397-2409. Turner, W.L. and Plaxton, W.C. 2001. Purification and characterization of banana fruit acid phosphatase. Planta 214:243-249. Wolucka, B.A., Persiau, G., Van Doorsselaere, J., Davey, M.W., Demol, H., Vandekerckhove, J., Van Montagu, M., Zabeau, M., and Boerjan, W. 2001. Partial purification and identification of GDP-mannose 3 ,5 -epimerase of Arabidopsis thaliana, a key enzyme of the plant vitamin C pathway. Proc. Natl. Acad. Sci. U.S.A. 9826:14843-14848. Yang, P., Fu, H., Walker, J., Papa, C.M., Smalle, J., Ju, Y.M., and Vierstra, R.D. 2004. Purification of the Arabidopsis 26S proteasome: Biochemical and molecular analyses revealed the presence of multiple isoforms. J. Biol. Chem. 279:64016413. Yuasa, K. and Maeshima, M. 2000. Purification, properties, and molecular cloning of a novel Ca2+ -binding protein in radish vacuoles. Plant Physiol. 124:1069-1078.
Key References Loomis, 1974. See above. Describes in detail procedures to avoid the problems of phenolics. Barraclough et al., 2004. See above. Describes a robust method to process plant tissue for two-dimensional electrophoresis.
Contributed by William Laing The Horticultural and Food Research Institute of New Zealand Auckland, New Zealand John Christeller The Horticultural and Food Research Institute of New Zealand Palmerston North, New Zealand
Extraction, Stabilization, and Concentration
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CHAPTER 5 Production of Recombinant Proteins INTRODUCTION
T
his chapter deals with the production of recombinant proteins using the expression systems in common use. Methods describing the actual construction of the expression vectors or plasmids containing the genes coding the proteins of interest are described in detail elsewhere (Sambrook et al., 1989; Ausubel et al., 1997).
The most widely used and most important expression systems are those using E. coli as the host cell. An overview of the production of proteins in E. coli is provided in UNIT 5.1. General strategies for gene expression described include direct intracellular expression, secretion of the protein into the periplasmic space, and the expression of fusion proteins. Some of these strategies (e.g., secretion procedures) are used to overcome the common problem of inclusion body formation (see UNIT 6.1 for additional discussion); others (e.g., fusions involving affinity tags) are used simply to assist in protein purification. Many of the expression vectors discussed are commercially available and suppliers are indicated throughout the text. Protocols describing protein expression in transformed E. coli cells are provided in UNITS 5.2 & 5.3. First, protein expression in E. coli requires that a suitable host strain (tabulated in UNIT 5.2) be transformed with an expression vector. Electroporation, a quick and simple method for transforming E. coli host strains, is presented in UNIT 5.2. Second, the growth conditions and methods used to induce gene expression are optimized for the particular host strain and controllable transcriptional promoter used. Standard procedures for small-scale expression in shaker flasks using several of the most common expression vectors are also presented, along with protocols for determining plasmid stability and storing transformed strains at low temperature. Finally, methods for establishing the level of protein expression as well as solubility and location (intracellular or periplasmic) are described. Once suitable host/vector combinations have been selected on the basis of the small-scale experiments described in UNIT 5.2, large-scale protein production can be carried out in fermentors with careful monitoring of cell growth. UNIT 5.3 describes fermentation in batch mode using a fixed amount of medium. For higher-density fermentations, the fed-batch process is described: in this procedure the growth carbon source is continuously fed into the culture. Modifications to the standard fed-batch process are used to introduce the heavy atom derivative selenomethionine into recombinant proteins to facilitate X-ray structure determination and to uniformly label proteins with combinations of the stable isotopes 2H, 13C, and 15N. Isotopically labeled proteins are required for high-resolution structural studies using nuclear magnetic resonance (NMR) spectroscopy (see UNIT 7.1 for additional details). Protocols for growth monitoring and sterility checking are also included. There are several eukaryotic cell–based expression systems available for overproducing recombinant proteins (consult Ausubel et al., 1997, Chapter 16), and most of these will eventually be covered in this chapter. One system, which uses insect cells infected with recombinant baculovirus, has become widely used. An overview of protein expression using the baculovirus system is given in UNIT 5.4. References covering the molecular Contributed by Paul T. Wingfield Current Protocols in Protein Science (2002) 5.0.1-5.0.3 Copyright © 2002 by John Wiley & Sons, Inc.
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biology required to construct the recombinant baculovirus vector that expresses the gene of interest are provided in the overview. Furthermore, references to protocols describing small-scale expression experiments are supplied in the introduction to UNIT 5.5. The large-scale growth of insect cells infected with baculovirus is carried out in bioreactors, requiring large (liter) amounts of viral stock (UNIT 5.5). These stocks must have known titers and are analyzed in small-scale (1- to 3-liter) cultures in order to evaluate and optimize protein expression. For developing protein purification methods and for isolating moderate amounts of recombinant protein, small-scale cultures in shaker or spinner culture flasks may provide adequate starting material. Large-scale insect cell culture is carried out with more sophisticated equipment using bioreactors in either batch mode or perfusion mode. Cell harvesting is carried out by centrifugation or tangential-flow microfiltration. The cells (for intracellular expressed proteins) or the culture medium (for secreted proteins) are retained for downstream processing. Other protocols in this unit provide methods for preparing samples for SDS-PAGE to check protein expression levels and for monitoring nutrient levels in culture media. Yeast is another important eukaryotic host used for protein expression. The molecular biology of Saccharomyces cerevisiae (baker’s yeast) is described in Chapter 13 of Ausubel et al. (1997). An overview of protein expression in S. cerevisiae is given in UNIT 5.6, which details, as a case study, some of the machinations required to achieve high-level production of the human glycoprotein antithrombin 111 secreted into the extracellular medium. Other strains of yeast apart from S. cerevisiae have also been used for heterologous protein expression, including the methylotropic yeast Pichia pastoris. There may be some advantages to using this host system over the more common S. cerevisiae, as expression levels, especially for intracellular proteins, can be higher. Protein expression using the P. pastoris system is reviewed in UNIT 5.7. The commonly used expression vectors are described, together with many examples of proteins successfully expressed with the system. Protocols describing the culture of the S. cerevisiae and P. pastoris strains with various expression vectors for heterologous protein expression are provided in UNIT 5.8. An overview of protein expression using mammalian cells is presented in UNIT 5.9. The unit reviews the stages involved in protein production using the stable expression approach. As post-transcriptional modification and intracellular protein transportation are integral components of mammalian protein expression, descriptions of these mechanisms are included. Protocols for the production of recombinant proteins in mammalian cells using various culture configurations are described in UNIT 5.10. These are supplemented with a number of support protocols that include the first steps in downstream processing. The transient expression of proteins in mammalian cells using a viral vector system is described in UNITS 5.11-5.15. Methods for constructing recombinant vaccinia virus are presented in UNITS 5.11-5.13 & 5.15. The characterization of the recombinant virus is described in UNIT 5.14. Proteins produced using this expression system are subject to normal mammalian post-translational processing.
Introduction
The choice of protein expression system does depend on the intended use and characteristics of the protein. If a protein can be expressed in E. coli in a soluble state either directly into the cytoplasm or by secretion into the periplasm, then it may be possible to isolate large amounts of correctly folded and active protein (e.g., see UNIT 6.2). On the other hand, if the protein must be post-translationally modified (i.e., glycosylated or phosphorylated), a eukaryotic expression system must be used. Furthermore, if the protein is expressed in an insoluble state in E. coli, one way to circumvent this problem is to try expression in an eukaryotic system.
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Current Protocols in Protein Science
The baculovirus system complements E. coli expression systems and for many investigators is the system of choice, especially for expressing post-translationally modified proteins. Yeast has the advantage that it is cheap to grow (analogous to E. coli) but can produce hyperglycosylated proteins; moreover, occasionally protein produced at high levels is insoluble (again analogous to E. coli). If a glycoprotein is required for clinical studies, it must usually be produced by mammalian cell expression, as discussed in detail in UNITS 5.9 & 5.10. However, no expression system is without its drawbacks; some of the problems reported using the baculovirus and yeast systems are mentioned in UNITS 5.4 & 5.6-5.8 and elsewhere (Bradley, 1990). In addition, a brief summary of the advantages and disadvantages of all the commonly used host systems is given in the booklet The Recombinant Protein Handbook (Pharmacia Biotech, 1994). A more detailed discussion of the choices for cellular protein expression is given in UNIT 5.16. Rather than commit to any one expression system for the expression a particular protein, the Gateway universal cloning technology (Invitrogen) allows for the rapid screening of many different vectors and host combinations. In UNIT 5.17 the use of the Gateway system for protein expression in multiple hosts is described and examples of traditional and high-throughput protein expression are provided. LITERATURE CITED Ausubel, F.M., Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G., Smith, J.A., and Struhl, K. (eds). 1997. In Current Protocols in Molecular Biology. John Wiley & Sons, New York. Bradley, M.K. 1990. Overexpression of proteins in eukaryotes. Methods Enzymol. 182:112-143. Pharmacia Biotech. 1994. The Recombinant Protein Handbook (18-1105-02). Pharmacia Biotech, Piscataway, N.J. Sambrook, J., Fritsch, E.F. and Maniatis, T. 1989. Molecular Cloning. A Laboratory Manual (2nd ed.). Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York.
Paul T. Wingfield
Production of Recombinant Proteins
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Production of Recombinant Proteins in Escherichia coli The genetics and biochemistry of Escherichia coli are probably the best understood of any known organism. The knowledge gained in the study of E. coli biology has been applied to the development of many of today’s molecular cloning techniques. Most cloning vectors and methods utilize E. coli or its phages as a preferred host, primarily because of the ease with which the bacterium can be grown and genetically manipulated. These same characteristics made E. coli an attractive early choice as a host for the production of large quantities of protein encoded by cloned genes. Aside from its well-studied biology, E. coli is suitable as the basis of an expression system because of its rapid doubling time and its ability to grow in inexpensive media. Years of study devoted to gene expression in E. coli have provided numerous choices for transcriptional and translational control elements that can be applied to the expression of foreign genes (UNITS 5.2 & 5.3). As a result, E. coli has been and continues to be the expression system of choice and a substantial body of literature has accumulated on the successful expression of foreign genes in this host. Several problems with protein expression in E. coli have been encountered, and many have been ultimately solved. This unit describes methods that have been developed for production of recombinant proteins in E. coli and potential pitfalls that may be encountered.
GENERAL STRATEGIES FOR GENE EXPRESSION IN E. COLI The basic approach used to express foreign genes in E. coli begins with insertion of the gene into an expression vector, usually a plasmid (obtained from a commercial supplier or from the author of a published study). The next step involves transforming a suitable E. coli host strain with the plasmid, for example, by electroporation (UNIT 5.2). Transformed cells are then subjected to evaluation of plasmid stability and foreign protein expression following induction of the controllable transcriptional promoter present in the expression vector (UNIT 5.2). Once small-scale shaker flask experiments have identified successful expression systems (UNIT 5.2), the transformed E. coli strain can be used in large-scale fermentation systems (UNIT 5.3), which can be designed to produce recombinant
Contributed by Edward R. LaVallie Current Protocols in Protein Science (1995) 5.1.1-5.1.8 Copyright © 2000 by John Wiley & Sons, Inc.
UNIT 5.1
proteins with altered amino acids (e.g., selenomethionine in place of methionine) or stable isotope tags to facilitate structural studies. Production is followed by protein purification (Chapter 6) and characterization (Chapter 7). If the gene to be expressed in E. coli is of eukaryotic origin, it must be a cDNA copy, as E. coli will not recognize and splice out introns from transcripts of genomic copies of eukaryotic genes. Typical E. coli expression vectors contain the following elements.
Selectable Marker Expression plasmids contain sequences encoding a selectable marker to ensure maintenance of the vector in the host cell. Commonly used selectable markers in E. coli include bla (which encodes β-lactamase and confers resistance to ampicillin and other β-lactam antibiotics), cat (which encodes chloramphenicol acetyltransferase and confers resistance to chloramphenicol), and tet (which encodes a membrane protein that confers resistance to tetracycline).
Origin of Replication Replication of a plasmid as an independent extrachromosomal element is controlled by its origin of replication. Some plasmids are under so-called stringent control, wherein the plasmid’s replication is coupled to replication of the host chromosome. As such, only one or at most a few copies of the plasmid are maintained in each cell. More commonly, E. coli vectors used for expression of cloned genes utilize relaxed control such as the ColE1 replicon found in pBR322 and derivatives. These plasmids generally have copy numbers of 10 to 200 plasmids per cell. The presence of multiple copies of the gene and its associated control elements can provide advantages in mRNA production and, as a result, may be reflected in an increase in the total amount of protein that accumulates. Nevertheless, the reduced copy number of stringent replicons may sometimes be advantageous: it allows for modulation of gene dosage, which can sometimes be helpful for altering the kinetics of expression or reducing the basal transcriptional levels of the plasmid-encoded promoter. Production of Recombinant Proteins
5.1.1 CPPS
Promoters
Production of Recombinant Proteins in Escherichia coli
An essential component of expression vectors is a controllable transcriptional promoter which, when induced, can direct the production of large amounts of mRNA from the cloned gene. There are a variety of controllable promoters that are routinely used. The lac promoter system utilizes the transcriptional control elements from the E. coli β-galactosidase gene (Miller and Reznikoff, 1978). The lac promoter (like the tac and trc promoters, which have optimized lac promoter RNA polymerase recognition sequences) is controlled by binding of lac repressor (the lacI gene product) to an operator sequence in the promoter region. The repressor can be expressed either from the host genome (single copy, in which case an overexpressing repressor allele called lacIq should be used) or from the expression vector (multiple copies, with resultant tighter transcriptional control). Induction of the promoter is generally accomplished by the addition of isopropyl-βD-thiogalactopyranoside (IPTG), a lactose analog that binds to the lac repressor and prohibits its binding to the lac operator. An example of a commercially available lac promoter vector is pBluescript (Stratagene). The pKK223-3 vector from Pharmacia Biotech is a source of the tac promoter. The trc promoter is used in the vectors pSE280, pSE380, and pSE420 from Invitrogen. Another commonly used promoter system utilizes the major leftward promoter of bacteriophage lambda, pL, and its control elements (Shimatake and Rosenberg, 1981). The pL promoter contained on an expression vector can be controlled by the phage-encoded cI repressor, which is typically expressed from an integrated copy of the phage in the host genome. A temperature-sensitive cI repressor (called cI857) is usually used; this encodes a repressor that is functional at lower temperatures but denatures at temperatures above 37.5°C. Thus, pL-mediated protein synthesis can be induced by a simple temperature shift. Alternatively, temperature-independent pL promoter systems have been developed that utilize a cI repressor gene under the control of a separate inducible promoter (Mieschendahl et al., 1986; LaVallie et al., 1993a). A commercial source for the pL vector is the pPL-Lambda vector from Pharmacia Biotech. The T7 RNA polymerase promoter is another popular transcriptional control element for heterologous expression. The RNA polymerase from bacteriophage T7 is highly selective for specific T7 phage promoter sequences that
are uncommon in other DNAs (Studier et al., 1990). A gene of interest placed under the control of a T7 promoter can be selectively expressed by induction of host-encoded T7 polymerase synthesis, which itself is under the control of an inducible promoter such as lac. Induction of T7 polymerase synthesis causes almost exclusive expression of the gene under the control of the T7 promoter. In some instances, the T7 transcription can outcompete the host RNA polymerase, resulting in accumulation of large amounts of the gene product of interest. There are many commercially available expression vectors that utilize the T7 polymerase/promoter system, such as the pGEMEX vectors from Promega, the pRSET vectors from Invitrogen, and the pET vectors from Novagen.
Translation Initiation Sequence Initiation of translation on mRNAs requires the presence of a so-called Shine and Dalgarno sequence or ribosome binding site (RBS) in close proximity to an initiator methionine (Shine and Dalgarno, 1974). The RBS consists of a purine-rich stretch of nucleotides complementary to the 3′ end of 16S RNA, located 5 to 13 bases 5′ to an initiator ATG. RBS elements typically used in expression vectors derive from well-translated E. coli or bacteriophage genes. For instance, the pTrcHis and pRSET vectors (Invitrogen) and the pGEMEX (Promega) vectors use the T7 gene 10 RBS.
SPECIFIC EXPRESSION STRATEGIES Direct Intracellular Expression Direct expression refers to the fusing of the coding sequence of interest to transcriptional and translational control sequences on an expression vector, with an initiator methionine codon preceding the open reading frame. This approach can be used to produce cytoplasmic proteins, and it can also be used for the intracellular expression of normally secreted proteins. In the latter case, the DNA sequence encoding the signal peptide is replaced by the initiator methionine codon. Success with the direct approach is often variable. First, translation initiation is inconsistent due to the fact that sequences 3′ to the initiator methionine can influence the efficiency of ribosome binding (Looman et al., 1987; Bucheler et al., 1990). For reasons that are not fully understood, maximizing the A+T content of the 5′ end of the coding sequence (taking advantage of the degeneracy of the
5.1.2 Current Protocols in Protein Science
genetic code) can sometimes improve the efficiency of translation initiation (De Lamarter et al., 1985; Devlin et al., 1988). Second, recombinant proteins produced in the cytoplasm often form dense, insoluble aggregates of protein called inclusion bodies (Schein, 1989). In some ways, this can be viewed as an advantage because inclusion bodies are easily purified from the soluble proteins (UNIT 6.3). In addition, proteins in inclusion bodies are usually resistant to proteolytic degradation. In cases where activity of the expressed protein is unnecessary (e.g., in production of protein to be used as an antigen to produce antibodies), inclusion body formation and resultant insolubility of the protein may actually be preferred. If active protein is desired, however, recovery of properly folded protein from inclusion bodies requires the total denaturation of the protein using reagents such as urea or guanidine, followed by subsequent refolding using protocols that must be determined empirically for each protein (UNIT 6.5). The success of refolding protocols is variable, depending on the particular protein, and may yield little or no correctly folded material. If protein insolubility is encountered and is undesired, production of soluble protein can sometimes be enhanced by simply lowering the growth temperature during protein synthesis (Bishai et al., 1987; Schein and Noteborn, 1988). If the protein is normally secreted, then a secretory construct may produce more soluble protein (see section on Secretion). Alternatively, production of the protein as a fusion to a highly soluble partner may alleviate the problem (see section on Fusion Proteins). If the gene is expressed poorly, translation efficiency may be a problem. If this is the case, it is often advantageous to maximize the A+T content of the 5′ coding sequence and/or replace the RBS with a sequence that initiates translation more efficiently (Olins and Rangwala, 1989). Another potential cause of poor translation is a preponderance of “rare” codons in the coding sequence, that is, codons that are underutilized by E. coli (Robinson et al., 1984). Stretches of contiguous rare codons may contribute to lower expression levels and should be changed to codons that are frequently used by E. coli. Finally, poor expression levels may be caused by product instability. In this instance, use of host strains and induction methods that minimize proteolytic degradation should be attempted (UNIT 5.2). In instances where the gene is expressed well and soluble protein can be produced, the protein must then be purified from an abundant
and diverse mixture of other E. coli cytoplasmic proteins. This can be a difficult and time-consuming process, and the purification scheme will be different for each protein based on its physical and biochemical characteristics (Chapters 6 and 8).
Secretion Secretion of proteins in E. coli is mediated by the presence of an N-terminal signal sequence that is cleaved after translocation of the protein. Expression of cloned gene products as secreted proteins in E. coli has been utilized as an alternative to cytoplasmic expression for proteins that are normally secreted. In E. coli the protein is secreted to the periplasmic space between the cytoplasmic and outer membranes, in contrast to extracellular secretion that occurs in gram-positive bacteria and eukaryotic cells. The result is that in E. coli the secreted protein remains cell-associated, although in a “compartment” separated from the cytoplasmic proteins that make up the vast majority of the total cellular protein. This can be advantageous in terms of protein purification if techniques are used that release only periplasmic contents while leaving the cytoplasmic membrane intact (Neu and Heppel, 1965). Secretion of heterologous gene products has been successfully employed for various proteins that are difficult to produce in the cytoplasm of E. coli as soluble and active proteins, including various growth factors (Cheah et al., 1994), receptors (Fuh et al., 1990), and recombinant Fab fragments (Skerra, 1994). Although eukaryotic signal peptides have been reported to function in E. coli, most secretion vectors utilize signal peptides derived from prokaryotic genes such as the ompA (Takahara et al., 1988; Cheah et al., 1994), pelB (Power et al., 1992), phoA (Oka et al., 1985), or hisJ (Vasquez et al., 1989) signal peptides. Although secretion provides an alternative to cytoplasmic expression that can sometimes result in the production of properly folded and active protein, the yield of desired protein is often low. Also, overexpressed gene products have been reported to form inclusion bodies even in the periplasm (Bowden and Georgiou, 1990); so secretion is not a panacea for the problem of insolubility.
Fusion Proteins The problems that have plagued successful overexpression of cloned gene products in E. coli—namely, inconsistent expression levels, protein insolubility, and difficult purification of the gene product from E. coli contaminants—
Production of Recombinant Proteins
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Production of Recombinant Proteins in Escherichia coli
have been most successfully addressed by the use of fusion proteins. Fusion proteins are created via a translational fusion of the coding sequence for the protein of interest to a gene for a highly expressed protein partner (or “carrier” protein). Typically, the gene encoding the protein of interest is inserted in-frame 3′ to the coding sequence for the carrier protein, in place of the usual termination codon. This allows uniform translational initiation of the carrier protein regardless of the coding sequence fused to its 3′ end, which helps to ensure consistent expression levels. Carrier proteins are usually chosen based on specific attributes that make them suitable in this role. The most successful fusion systems employ the maltose-binding protein (MBP; Maina et al., 1988), glutathione S-transferase (GST; Smith and Johnson, 1988), or thioredoxin (TRX; LaVallie et al., 1993a). The genes for these proteins are well expressed, and the proteins are highly soluble and provide specific physical characteristics or affinities to aid purification. These qualities also extend to the protein sequences fused to them, thereby enhancing the solubility and ease of purification of the entire fusion protein. Maltose-binding protein is a 43-kDa secreted protein from E. coli. As its name implies, it binds specifically to maltose or amylose, a property that can be exploited in purification schemes (Guan et al., 1988). MBP fusions can be secreted into the periplasm, or they can be expressed without the MBP signal peptide, which results in accumulation of the fusion protein in the cytoplasm. MBP fusions are usually well expressed, and a high proportion of MBP fusion proteins are soluble. MBP expression plasmids and reagents can be purchased from New England Biolabs. The MBP expression vectors utilize the tac promoter and a plasmid-encoded lacIq gene for transcriptional control. These vectors contain a recognition sequence for the site-specific protease factor Xa to allow removal of the carrier protein following purification (see section on Fusion Protein Cleavage Methods). Glutathione S-transferase is a 26-kDa cytoplasmic protein from Schistosoma japonicum. Carboxy-terminal protein fusions to GST are usually soluble and well-expressed (Smith and Johnson, 1988). GST binds specifically to glutathione, and GST fusion proteins can be purified in a single step from crude bacterial lysates by affinity chromatography on immobilized glutathione. The GST expression plasmids are
available from Pharmacia Biotech, and utilize the tac promoter and plasmid-encoded lacIq for inducible transcriptional control. A variety of vectors are available which contain either the factor Xa or the thrombin recognition sequence to allow cleavage and removal of the GST from the protein of interest following affinity purification (see section on Fusion Protein Cleavage Methods). Thioredoxin is a 12-kDa intracellular E. coli protein. It is very soluble, can be highly overexpressed from plasmid vectors (Lunn et al., 1984), and has been shown to be a very successful fusion partner. A wide variety of gene products can be produced abundantly in soluble fashion when fused to TRX (LaVallie et al., 1993a). Two different characteristics of TRX can be exploited to allow specific purification of some TRX fusion proteins. TRX accumulates at specific sites along the inner surface of the cytoplasmic membrane, called Bayer’s patches or adhesion zones. These sites constitute an osmotically sensitive compartment, and TRX (and some TRX fusions) can be selectively released from the cytoplasm and separated from the bulk of E. coli proteins by osmotic shock or freeze/thaw procedures. In addition, TRX is thermostable, and some TRX fusions can be purified by selective thermal denaturation of contaminants. In instances where osmotic shock or heat treatments do not provide adequate purification, an altered TRX protein has been developed (E.R.L., manuscript in preparation) that allows specific purification by metal-chelate affinity chromatography. The TRX fusion vector (available from Invitrogen) contains an enterokinase recognition sequence positioned at the junction between TRX and the C-terminal fusion partner. In addition to the carrier proteins listed above, there are many other proteins that have been used to produce fusions for the purpose of generating large amounts of protein in E. coli. Among them are E. coli β-galactosidase (Ruther and Muller-Hill, 1983) and TrpE (Yansura, 1990), Staphylococcus aureus protein A (Nilsson et al., 1985; vector available from Pharmacia Biotech), chloramphenicol acetyltransferase (Knott et al., 1988), bacteriophage lambda cII protein (Nagai and Thøgersen, 1987), and various carbohydrate-binding proteins (Taylor and Drickamer, 1991; Helman and Mantsala, 1992). Table 5.1.1 lists many fusion partners that have been described in the literature.
5.1.4 Current Protocols in Protein Science
Common Fusion Proteins
Table 5.1.1
Fusion protein
Specific purification methoda
Maltose-binding protein Glutathione S-transferase Thioredoxin Protein A β-Galactosidase Chloramphenicol acetyltransferase lac repressor Galactose-binding protein Cyclomaltodextrin glucanotransferase Lambda cII protein TrpE protein
Amylose binding Glutathione binding Selective release, heat stability, MCAC IgG binding APTG or anti-β-galactosidase antibody binding Chloramphenicol binding lac operator binding (Lundeberg et al., 1990) Galactose binding Cyclodextrin binding None None
aMCAC, metal-chelate affinity chromatography; IgG, immunoglobulin G; APTG, p-amino-β-D-thiogalactoside.
Table 5.1.2
Fusion Tags
Amino acid tag
Ligand
Reference/supplier
Polyhistidine Polyaspartic acid Polyarginine Polyphenylalanine Polycysteine In vivo biotinylated peptide Flag peptide
Metal-chelate affinity resin Anion-exchange resin Cation-exchange resin HIC resina Thiol Avidin or streptavidin Anti-Flag antibody
Qiagen Dalbøge et al. (1987) Brewer and Sassenfeld (1985) Persson et al. (1988) Persson et al. (1988) Schatz (1993) International Biotechnologies (IBI)
aHIC, hydrophobic interaction chromatography (UNIT 8.4).
Fusion Tags Fusion tags are small stretches of amino acids added to the N-terminal or C-terminal end of a protein. Although they do not usually help to increase expression levels or protein solubility, they can be advantageous in protein purification and detection. The tags are generally chosen either because they encode an epitope that can be detected and purified using an antibody that binds to it, or because the tag amino acids provide a physical characteristic that can be exploited for easy and specific purification. A popular example is the polyhistidine tag, usually a stretch of six consecutive histidine residues added to either the N or C terminus of a protein, that provides specific binding to metal chelate resins. There are a number of polyhistidine vectors on the market, such as the pTrcHis vector from Invitrogen. Other examples, such as polyarginine (Brewer and Sassenfeld, 1985) or polyaspartic acid tags (Dalbøge et al., 1987), can be used to alter the binding
behavior of a protein on ion-exchange resins. A noteworthy fusion tag is a small stretch of amino acids recognized and biotinylated in vivo by the biotin-protein ligase in E. coli (Schatz, 1993). This allows specific capture of the biotinylated protein on immobilized avidin or streptavidin. Table 5.1.2 lists a number of the more popular fusion tags.
Fusion Protein Cleavage Methods Whereas fusion proteins and fusion tags have proved useful for the reasons outlined above, the end result is that the protein of interest will carry additional sequences that may hamper its functional activity. There are several methods for removal of the carrier protein, which can be divided into chemical cleavage methods and enzymatic (proteolytic) cleavage methods. Chemical cleavage of fusion proteins can be accomplished with reagents such as cyanogen bromide (Met↓; Itakura et al., 1977; Villa et al., 1989), 2-(2-nitrophenyl-
Production of Recombinant Proteins
5.1.5 Current Protocols in Protein Science
Production of Recombinant Proteins in Escherichia coli
sulfenyl) - 3 - methyl - 3′ - bromoindolinine or BNPS-skatole (Trp↓; Dykes et al., 1988), hydroxylamine (Asn↓Gly; Bornstein and Balian, 1977), or low pH (Asp↓Pro; Szoka et al., 1986). The use of these reagents, as described in UNIT 11.4, can be applied to the site-specific cleavage of fusion proteins provided that the fusion is designed so that the chemically labile bond is positioned at the desired point of scission. Chemical cleavage reagents tend to be inexpensive and efficient, and many of the reactions can be performed under denaturing conditions so that even insoluble proteins can be cleaved. However, one disadvantage of using chemical cleavage reagents for the site-specific cleavage of fusion proteins is that the reactions are generally performed under extremes of pH and/or temperature that can result in unwanted amino acid side-chain modifications. Chemical cleavage methods also have the disadvantage of low specificity, so it is more likely that the protein of interest will contain an internal cleavage site. Enzymatic digestion is usually the method of choice for soluble fusion protein cleavage as the reactions are carried out under relatively mild conditions. Proteases such as trypsin (Lys↓ or Arg↓), endoproteinase Asp-N (↓Asp), and Staph V8 S. aureus V8 protease (Glu↓ or Asp↓) can be used in this role. High-quality trypsin, S. aureusV8 protease (endoproteinase Glu-C), and endoproteinase Asp-N can all be obtained from Boehringer Mannheim. These proteases, like chemical cleaving reagents, are limited by their low degree of specificity (recognizing and cleaving at single amino acids). Alternatively, thrombin (Leu-Val-Pro-Arg↓Gly-Ser; Gearing et al., 1989), factor Xa [Ile-Glu(or Asp)-Gly-Arg↓; Nagai and Thøgersen, 1984, 1987; Gardella et al., 1990], renin (Pro-Phe-His-Leu↓Leu-Val-Tyr; Haffey et al., 1987), collagenase (Pro-X↓Gly-Pro-Y↓; Germino and Bastia, 1984), and enterokinase (Asp-Asp-Asp-Asp-Lys↓; Dykes et al., 1988; LaVallie et al., 1993b) are much more suitable in this role. All of these enzymes have extended substrate recognition sequences (up to seven amino acids in the case of renin), which greatly reduces the likelihood of unwanted cleavages elsewhere in the protein. Factor Xa and enterokinase are most useful for cleaving off C-terminal fusion partners because they cleave on the carboxyl-terminal side of their respective recognition sequences. This allows the release of fusion partners containing their authentic amino terminus. In addition, the catalytic subunit of bovine enterokinase has been cloned
and expressed (LaVallie et al., 1993b), and the recombinant enzyme has been shown to be approximately 100-fold more efficient than the native intestinally derived enzyme in fusion protein cleavage reactions (Racie et al., 1995). The recombinant enterokinase is available from New England Biolabs.
SUMMARY Methods for the overexpression of cloned gene products in E. coli have improved significantly since it was first attempted. Common problems such as variable expression levels, inclusion body formation, and purification difficulties have been successfully addressed by advancements in expression technology. Probably the most significant of these advancements has been the development of fusion proteins and fusion tag expression and purification techniques. These methods have resulted in more consistent production of soluble and active protein, and have allowed for simple and efficient purification of the proteins from bacterial lysates. Although the production of soluble, properly folded, and active recombinant proteins in E. coli is still not guaranteed, the likelihood of success is far greater than it was just a few years ago. This progress should help ensure that E. coli will continue to be the host organism of choice for recombinant protein production.
LITERATURE CITED Bishai, W.R., Rappuoli, R., and Murphy, J.R. 1987. High-level expression of a proteolytically sensitive diphtheria toxin fragment in Escherichia coli. J. Bacteriol. 169:5140-5151. Bornstein, P. and Balian, G. 1977. Cleavage at AsnGly bonds with hydroxylamine. Methods Enzymol. 47:132-145. Bowden, G.A. and Georgiou, G. 1990. Folding and aggregation of β-lactamase in the periplasmic space of Escherichia coli. J. Biol. Chem. 265:16760-16766. Brewer, S.J. and Sassenfeld, H.M. 1985. The purification of recombinant proteins using C-terminal poly-arginine fusions. Trends Biotechnol. 3:119122. Bucheler, U.S., Werner, D., and Schirmer, R.H. 1990. Random silent mutagenesis in the initial triplets of the coding region: A technique for adapting human glutathione reductase-encoding cDNA to expression in Escherichia coli. Gene 96:271-276. Cheah, K.C., Harrison, S., King, R., Crocker, L., Well, J.R., and Robins, A. 1994. Secretion of eukaryotic growth hormones in Escherichia coli is influenced by the sequence of the mature proteins. Gene 138:9-15.
5.1.6 Current Protocols in Protein Science
Dalbøge, H., Dahl, H.H.M., Pedersen, J., Hansen, J.W., and Christensen, T. 1987. A novel enzymatic method for production of authentic hGH from an Escherichia coli-produced hGH precursor. Bio/Technology 5:161-164.
Knott, J.A., Sullivan, C.A., and Weston, A. 1988. The isolation and characterisation of human atrial natriuretic factor produced as a fusion protein in Escherichia coli. Eur. J. Biochem. 174:405-410.
De Lamarter, J.F., Mermod, J.J., Liang, C.M., Eliason, J.F., and Thatcher, D.R. 1985. Recombinant murine GM-CSF from E. coli has biological activity and is neutralized by a specific antiserum. EMBO J. 4:2575-2581.
LaVallie, E.R., DiBlasio, E.A., Kovacic, S., Grant, K.L., Schendel, P.F., and McCoy, J.M. 1993a. A thioredoxin gene fusion system that circumvents inclusion body formation in the E. coli cytoplasm. Bio/Technology 11:187-193.
Devlin, P.E., Drummond, R.J., Toy, P., Mark, D.F., Watt, K.W.K., and Devlin, J.J. 1988. Alteration of the amino-terminal codons of human granulocyte colony-stimulating factor increases expression levels and allows efficient processing by methionine aminopeptidase in Escherichia coli. Gene 65:13-22.
LaVallie, E.R., Rehemtulla, A., Racie, L.A., DiBlasio, E.A., Ferenz, C., Grant, K.L., Light, A., and McCoy, J.M. 1993b. Cloning and functional expression of a cDNA encoding the catalytic subunit of bovine enterokinase. J. Biol. Chem. 268:23311-23317.
Dykes, C.W., Bookless, A.B., Coomber, B.A., Noble, S.A., Humber, D.C., and Hobden, A.N. 1988. Expression of atrial natriuretic factor as a cleavable fusion protein with chloramphenicol acetyltransferase in Escherichia coli. Eur. J. Biochem. 174:411-416. Fuh, G., Mulkerrin, M.G., Bass, S., McFarland, N., Brochier, M., Bourell, J.H., Light, D.R., and Wells, J.A. 1990. The human growth hormone receptor. Secretion from Escherichia coli and disulfide bonding pattern of the extracellular binding domain. J. Biol. Chem. 265:3111-3115. Gardella, T.J., Rubin, D., Abou-Samra, A.-B., Keutmann, H.T., Potts, J.T. Jr., Kronenberg, H.M., and Nussbaum, S.R. 1990. Expression of human parathyroid hormone (1-84) in Escherichia coli as a factor X-cleavable fusion protein. J. Biol. Chem. 265:15854-15859. Gearing, D.P., Nicola, N.A., Metcalf, D., Foote, S., Willson, T.A., Gough, N.M., and Williams, R.L. 1989. Production of leukemia factor in Escherichia coli by a novel procedure and its use in maintaining embryonic stem cells in culture. Bio/Technology 7:1157-1161. Germino, J. and Bastia, D. 1984. Rapid purification of a gene product by genetic fusion and site specific proteolysis. Proc. Natl. Acad. Sci. U.S.A. 81:4692-4696. Guan, C., Li, P., Riggs, P.D., and Inouye, H. 1988. Vectors that facilitate the expression and purification of foreign peptides in Escherichia coli by fusion to maltose-binding protein. Gene 67:2130. Haffey, M.L., Lehman, D., and Boger, J. 1987. Sitespecific cleavage of a fusion protein by renin. DNA 6:565-571. Hellman, J. and Mantsala, P. 1992. Construction of an E. coli export-affinity vector for expression and purification of foreign proteins by fusion to cyclomaltodextrin glucanotransferase. J. Biotechnol. 23:19-34. Itakura, K., Hirose, T., Crea, R., Riggs, A.D., Heyneker, H.L., Bolivar, F., and Boyer, H.W. 1977. Expression in Escherichia coli of a chemically synthesized gene for the hormone somatostatin. Science 198:1056-1063.
Looman, A.C., Bodlaender, J., Comstock, L.J., Eaton, D., Jhurani, P., de Boer, H.A., and van Knippenberg, P.H. 1987. Influence of the codon following the AUG initiation codon on the expression of a modified lacZ gene in Escherichia coli. EMBO J. 6:2489-2492. Lundeberg, J., Wahlberg, J., and Uhlen, M. 1990. Affinity purification of specific DNA fragments using a lac repressor fusion protein. Genet. Anal. Techn. Appl. 7:47-52. Lunn, C.A., Kathju, S., Wallace, B.J., Kushner, S.R., and Pigiet, V. 1984. Amplification and purification of plasmid-encoded thioredoxin from Escherichia coli K12. J. Biol. Chem. 259:1046910474. Maina, C.V., Riggs, P.D., Grandea, A.G., Slatko, B .E., Mora n, L.S., Ta gliam onte, J.A., McReynolds, L.A., and Guan, C. 1988. An Escherichia coli vector to express and purify foreign proteins by fusion to and separation from maltose-binding protein. Gene 74:365-373. Mieschendahl, M., Petri, T., and Hanggi, U. 1986. A novel prophage independent trp regulated lambda pL expression system. Bio/Technology 4:802-808. Miller, J.H. and Reznikoff, W.S. (eds.) 1978. The Operon. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Nagai, K. and Thøgersen, H. C. 1984. Generation of β-globin by sequence-specific proteolysis of a hybrid protein in Escherichia coli. Nature 309:810-812. Nagai, K. and Thøgersen, H.C. 1987. Synthesis and sequence-specific proteolysis of hybrid proteins produced in Escherichia coli. Methods Enzymol. 153:461-481. Neu, H.C. and Heppel, L.A. 1965. Release of enzymes from Escherichia coli by osmotic shock during the formation of spheroplasts. J. Biol. Chem. 240:3685-3692. Nilsson, B., Abrahmsen, L., and Uhlen, M. 1985. Immobilization and purification of enzymes with staphylococcal protein A gene fusion vectors. EMBO J. 4:1075-1080. Oka, T., Sakamoto, S., Miyoshi, K., Fuwa, T., Yoda, K., Yamasaki, M., Tamura, G., and Miyake, T. 1985. Synthesis and secretion of human epider-
Production of Recombinant Proteins
5.1.7 Current Protocols in Protein Science
mal growth factor by Escherichia coli. Proc. Natl. Acad. Sci. U.S.A. 82:7212-7216. Olins, P.O. and Rangwala, S.H. 1989. A novel sequence element derived from bacteriophage T7 mRNA acts as an enhancer of translation of the lacZ gene in Escherichia coli. J. Biol. Chem. 264:16973-16976. Persson, M., Bergstrand, M.G., Bulow, L., and Mosbach, K. 1988. Enzyme purification by genetically attached polycysteine and polyphenylalanine tags. Anal. Biochem. 172:330-337. Power, B.E., Ivancic, N., Harley, V.R., Webster, R.G., Kortt, A.A., Irving, R.A., and Hudson, P.J. 1992. High-level temperature-induced synthesis of an antibody VH-domain in Escherichia coli using the PelB secretion signal. Gene 113:95-99. Racie, L.A., McColgan, J.M., Grant, K.L., DiBlasio-Smith, E.A., McCoy, J.M., and LaVallie, E.R. 1995. Production of recombinant bovine enterokinase catalytic subunit in Escherichia coli using the novel secretory fusion partner DsbA. Bio/Technology. In press. Robinson, M., Lilley, R., Little, S., Emtage, J.S., Yarranton, G., Stephens, P., Millican, A., Eaton, M., and Humphreys, G. 1984. Codon usage can affect efficiency of translation of genes in Escherichia coli. Nucl. Acids Res. 12:6663-6671. Ruther, U. and Muller-Hill, B. 1983. Easy identification of cDNA clones. EMBO J. 2:1791-1794. Schatz, P.J. 1993. Use of peptide libraries to map the substrate specificity of a peptide-modifying enzyme: A 13 residue consensus peptide specifies biotinylation in Escherichia coli. Bio/Technology 11:1138-1143. Schein, C.H. 1989. Production of soluble recombinant proteins in bacteria. Bio/Technology 7:1141-1149. Schein, C.S. and Noteborn, M.H.M. 1988. Formation of soluble recombinant proteins in Escherichia coli is favored by lower growth temperature. Bio/Technology 6:291-294. Shimatake, H. and Rosenberg, M. 1981. Purified λ regulatory protein cII positively activates promoters for lysogenic development. Nature 292:128-132.
Shine, J. and Dalgarno, L. 1974. The 3′-terminal sequence of Escherichia coli 16S ribosomal RNA: Complementarity to nonsense triplets and ribosome binding sites. Proc. Natl. Acad. Sci. U.S.A. 71:1342-1346. Skerra, A. 1994. A general vector, pASK84, for cloning, bacterial production, and single-step purification of antibody Fab fragments. Gene 141:79-84. Smith, D.B. and Johnson, K.S. 1988. Single-step purification of polypeptides expressed in Escherichia coli as fusions with glutathione Stransferase. Gene 67:31-40. Studier, F.W., Rosenberg, A.H., Dunn, J.J., and Dubendorff, J.W. 1990. Use of T7 RNA polymerase to direct expression of cloned genes. Methods Enzymol. 185:60-89. Szoka, P.R., Schreiber, A.B., Chan, H., and Murthy, J. 1986. A general method for retrieving components of a genetically engineered fusion protein. DNA 5:11-20. Takahara, M., Sagai, H., Inouye, S., and Inouye, M. 1988. Secretion of human superoxide dismutase in Escherichia coli. Bio/Technology 6:195-198. Taylor, M.E. and Drickamer, K. 1991. Carbohydrate-recognition domains as tools for rapid purification of recombinant eukaryotic proteins. Biochem. J. 274:575-580. Vasquez, J.R., Evnin, L.B., Higaki, J.N., and Craik, C.S. 1989. An expression system for trypsin. J. Cell. Biochem. 39:265-276. Villa, S., DeFazio, G., and Canosi, U. 1989. Cyanogen bromide cleavage at methionine residues of polypeptides containing disulfide bonds. Anal. Biochem. 177:161-164. Yansura, D.G. 1990. Expression as trpE fusion. Methods Enzymol. 165:161-166.
Contributed by Edward R. LaVallie Genetics Institute, Inc. Cambridge, Massachusetts
Production of Recombinant Proteins in Escherichia coli
5.1.8 Current Protocols in Protein Science
Selection of Escherichia coli Expression Systems
UNIT 5.2
Use of Escherichia coli as a protein factory capable of being retooled to meet the needs of the investigator is a well-established alternative to purification of proteins from natural sources. Moreover, expression—the directed synthesis of a foreign gene—is often the next step for researchers who have isolated a gene and want to study the protein it encodes. Table 5.2.1 lists the most useful expression strains for fermentation processes. A more comprehensive table of E. coli strains can be found in Raleigh et al. (1989) and Sambrook et al. (1989). A summary of four of the most widely used expression systems is presented in Table 5.2.2. The available expression systems have been reviewed (e.g., Goeddel, 1990). A number of systems use thermoinducible promoters that predictably induce the E. coli heat-shock response. We therefore frequently use strains partially or totally defective in this response (e.g., Coli B, HB101, or DH5α) since some of the induced heat-shock proteins are proteases.
Table 5.2.1
Escherichia coli Fermentation Strains
Strain
Genotype
Source
Comment
B
Wild type
Commonly used
B834
F− ompT gal met rBmB (DE3)a
ATCC 23227/nonK12 lon deficient Novagen
F− ompT rBmB (DE3)a F− gyrA96 recA1 relA1 endA1 thi-1 hsdR17 supE44 λ− mcrA mcrB nalR DH5α F− gyrA96 recA1 relA1 endA1 thi-1 hsdR17 supE44 δlacU169 (φ80lacZδM15) λ− mcrA mcrB nalR HMS174 F−recA r−K12 m+K12 RifR HB101 F− leuB6 thi-1 hsdS20 lacY1 proA2 ara-14 galK2 xyl-5 mtl-1 recA13 rpsL20 supE44 λ− mcrA strR JM101 supE thi-1 δ(lac-proAB) F′[traD36 proAB+ lacIq lacZδM15] MM294 F− endA1 thi-1 hsdR17 supE44 λ− mcrA mcrB RV308 su− ∆lacX74 galISII::OP308 strA SG936 F− lac(am) trp(am) pho(am) supC(ts) rpsL mal(am) htpR(am) tsx::Tn10 lonR9 W3110 F− λ− mcrA mcrB BL21 DH1
Novagen ATCC 33849
Host for selenomethionine labeling with T7 expression Host for T7 expression General host
GIBCO/BRL
General host
Novagen ATCC 33694
Host for T7 expression Inhibited at 42°C
ATCC 33876
Host for lac and tac expression
ATCC 33625
General host
ATCC 31608
Host for secretion
ATCC 39624
Defective in heat-shock response
ATCC 27325
General host
aλDE3 lysogen
Production of Recombinant Proteins
Contributed by Alain Bernard and Mark Payton
5.2.1
Current Protocols in Protein Science (1995) 5.2.1-5.2.18 Copyright © 2000 by John Wiley & Sons, Inc.
Table 5.2.2
Summary of Expression Systems
Promoter
Repressor Induction Host
Selection
Reference
pL lac, tac trp T7
cI857 lacIQ Trp lacIQ
Tet Amp Tet Amp/Cm
Remaut et al., 1987 Stahl et al., 1989 Rosenberg et al., 1984 Studier et al., 1986
30°-42°C IPTG IAAa IPTG
E. coli B E. coli JM101 E. coli B E. coli BL21 (DE3) E. coli HMS174 (DE3)
aThe trp promoter is naturally induced by tryptophan starvation; addition of IAA is optional.
This unit describes standard procedures for several expression systems, namely, temperature induction via the pL promoter (Basic Protocol 1) and chemical induction via the trp promoter (Basic Protocol 2), lac or tac promoters (Alternate Protocol 1), and the T7 promoter (Alternate Protocol 2). These protocols require that the gene encoding the protein of interest has been identified and cloned into an appropriate expression vector using standard molecular biology techniques. Transformation of a suitable host strain (e.g., by electroporation; Support Protocol 2) is also a prerequisite. Once an appropriate expression strain has been selected, the stability of the plasmid can be analyzed (Support Protocol 2) and the strain can be stored (Support Protocol 3). Other support protocols in this unit describe how to prepare samples for electrophoresis (Support Protocol 4), how to analyze the solubility of the expressed proteins (Support Protocol 5), and how to make samples of periplasmic extracts (Support Protocol 6) and extracellular media (using TCA precipitation, Support Protocol 7). Many of the support protocols are small-scale analysis procedures that are used to guide subsequent purification strategies and determine the suitability of the expression system for further development and scale-up. BASIC PROTOCOL 1
EXPRESSION UNDER CONTROL OF pL PROMOTER: TEMPERATURE INDUCTION Protein expression from vectors containing the pL promoter from bacteriophage lambda can be induced by raising the temperature. The E. coli host strains used with such vectors must harbor a temperature-sensitive repressor protein encoded by cI857 either integrated in the host chromosome as a dormant phage, or built into the same plasmid as the one carrying the heterologous gene. At high temperatures, expression of the cI857 gene is inactive, releasing the repression and allowing transcription of genes downstream of the pL promoter. In this protocol, cells are first grown at 30°C to the desired density in rich medium containing antibiotic to maintain the plasmid; synthesis of the protein is then induced by a temperature shift to 42°C. The proper antibiotic is determined by the expression vector. Materials Host E. coli strain (e.g., Coli B, W3110) transformed with plasmid containing gene for the protein of interest under control of pL promoter (see Support Protocol 1) LB medium (see recipe) Antibiotic stock solution (see recipe)
Selection of Escherichia coli Expression Systems
20-ml sterile culture tube 42°C shaking water bath 150-ml shaker flask, sterile
5.2.2 Current Protocols in Protein Science
1. Prepare a small liquid culture in stationary phase of the host E. coli strain transformed with plasmid containing the gene for the protein of interest under control of the pL promoter by inoculating a 20-ml tube containing 2 ml LB medium (supplemented with appropriate antibiotic stock solution) with a single colony from a fresh agar plate of the same medium. Incubate the tube overnight on a rotary shaker at 30°C, 200 rpm. Do not allow the temperature to exceed 30°C. Temperatures above 30°C may induce expression of the target protein and interfere with growth.
2. The next morning, read the OD600 against water as a blank to verify that cultures have reached stationary phase. In a 1-cm light path cuvette, the OD should be in the range of 1 to 2. (An OD of 1 is equal to 0.5-1 × 109 cells/ml.) The OD is read on an aliquot of the culture diluted 10-fold in PBS buffer against water as a blank. Readings higher than 0.4 are not correlated to cell density. Absolute absorbance values will vary among spectrophotometers depending on the cuvettes, light path, and construction of the apparatus.
3. Place a 150-ml shaker flask containing 50 ml LB medium with antibiotic in a 42°C shaking water bath to prewarm the medium. Again, make sure that the temperature is accurate; too low a temperature will lead to incomplete repressor inactivation and poor induction. To measure temperature, insert a thermometer into a control flask (containing water only) also placed in the water bath.
4. Transfer the 2-ml overnight culture into the flask containing prewarmed medium. Take a 1-ml sample and process it as described in step 5. Return the 50-ml culture to the 42°C shaking water bath and incubate 3 hr with vigorous agitation (200 rpm). At the end of the incubation, take another 1-ml sample. Inoculation and sample removal should be done quickly to keep the temperature stable.
5. Microcentrifuge each sample 2 min at maximum speed, room temperature. Discard the supernatant and prepare for analysis by SDS-PAGE (see Support Protocol 4). Pellets may be stored at −20°C at this point before SDS-PAGE. They may be stored frozen indefinitely (for years).
6. Use the remainder of the 50-ml culture to analyze plasmid stability (Support Protocol 2), determine solubility of the sample (see Support Protocol 5), and prepare periplasmic extracts or extracellular medium samples (see Support Protocol 5 and Support Protocol 6) to obtain recombinant protein. Store leftover culture, if desired (see Support Protocol 3). EXPRESSION UNDER CONTROL OF trp PROMOTER: CHEMICAL INDUCTION This protocol describes induction of synthesis of the protein of interest under control of the E. coli trp promoter. Cells are grown to exponential phase in minimal medium to achieve tryptophan starvation, and residual trp repressor is inactivated by the tryptophan analog indole-3-acrylic acid. Incubation of cultures is conducted at 37°C throughout, and the choice of antibiotic is determined by the plasmid construction. Materials Host E. coli strain transformed with plasmid containing trp promoter and gene for the protein of interest (see Support Protocol 1) M9 medium (see recipe) with 0.5% (w/v) Casamino Acids (Difco) Antibiotic stock solution (see recipe) Indole-3-acrylic acid (IAA; see recipe)
BASIC PROTOCOL 2
Production of Recombinant Proteins
5.2.3 Current Protocols in Protein Science
20-ml sterile culture tube 150-ml shaker flask 37°C orbital shaker 1. Use a single colony of the host E. coli strain transformed with plasmid containing trp promoter and gene for the protein of interest, from a fresh agar plate, to inoculate a 20-ml tube containing 2 ml of M9 medium with 0.5% Casamino Acids and appropriate antibiotic. Incubate the tube overnight at 37°C on a rotary shaker (200 rpm). 2. The next morning, read OD600 against water as a blank. 3. Transfer the overnight culture to a 150-ml shaker flask containing 50 ml of M9/Casamino Acids supplemented with appropriate antibiotic. 4. Take an initial 1-ml sample as a control for uninduced cells and either immediately treat as described in step 7 or refrigerate at −20°C until ready to perform step 7. 5. Incubate the culture for 1 to 2 hr in a 37°C orbital shaker with vigorous agitation (200 rpm). Check OD600. The cells should be in full exponential growth phase (OD600 of 0.2 to 0.5).
6. Induce protein expression by adding IAA (2.5 ml of the 400× stock per liter) to a final concentration of 25 µg/ml. Incubate for another 3 hr and take a second (final) 1-ml sample. 7. For each 1-ml sample, microcentrifuge 2 min at maximum speed, room temperature. Discard supernatant and prepare the sample for SDS-PAGE analysis (see Support Protocol 4). If it is not convenient to electrophorese samples right away, pellets can be stored at −20°C.
8. Use remaining culture for analysis of protein solubility (see Support Protocol 5), and localization of proteins in the periplasmic space (see Support Protocol 6) or culture medium (see Support Protocol 7), or analysis of plasmid stability (if needed; see Support Protocol 2). If desired, store leftover cells (see Support Protocol 3). ALTERNATE PROTOCOL 1
EXPRESSION UNDER CONTROL OF lac/tac PROMOTERS For protein expression under control of the lac promoter or tac promoter (a hybrid between lac and trp promoters), induction via release of the lac repressor is achieved by adding isopropyl-β-thiogalactopyranoside (IPTG) to cultures, which binds and inactivates the repressor. The host strain must harbor the lacI repressor gene in the chromosome or in a plasmid (either the same plasmid as the target gene or a compatible one). Additional Materials (also see Basic Protocol 2) Host E. coli strain containing lacI repressor gene and transformed with plasmid containing lac or tac promoter and gene for the protein of interest Isopropyl-β-thiogalactopyranoside (IPTG; see recipe) For this protocol, follow steps 1 to 5 of Basic Protocol 2 (trp promoter), substituting the appropriate host strain as the starting material. Replace step 6 with the following and then return to steps 7 and 8 of Basic Protocol 2.
Selection of Escherichia coli Expression Systems
6a. Induce protein expression by adding IPTG to a final concentration of 0.4 to 1.0 mM. Incubate for a further 3 hr and take a second (final) 1-ml sample.
5.2.4 Current Protocols in Protein Science
T7 RNA POLYMERASE/PROMOTER EXPRESSION SYSTEM The T7 RNA polymerase/promoter expression system utilized in this protocol involves induction, via the lacUV5 promoter, of the highly active and efficient T7 RNA polymerase, which binds to the T7 promoter (pT7) and transcribes genes under its control. An appropriate E. coli strain—in which the T7 polymerase gene under control of lacUV5 has been integrated into host DNA and which has been transformed with plasmid DNA containing an ampicillin-resistance gene and the gene for the protein of interest under control of pT7—is induced with IPTG. In a variation of the system, T7 lysozyme, which binds the polymerase and thus eliminates transcription under noninduced conditions, can be introduced on a second plasmid (e.g., pLysE or pLysS, which also encode chloramphenicol resistance) and maintained under selection with a second antibiotic.
ALTERNATE PROTOCOL 2
Additional Materials (also see Basic Protocol 2) Host E. coli strain transformed with DNA encoding T7 RNA polymerase under control of lacUV5 promoter, plasmid containing pT7 and gene for the protein of interest, and (optional) plasmid containing gene for T7 lysozyme Isopropyl-β-thiogalactopyranoside (IPTG; see recipe) For this protocol, follow steps 1 to 5 of Basic Protocol 2, except begin with the appropriate transformed E. coli host strain, use ampicillin selection, and include a second antibiotic if the optional T7 lysozyme plasmid strategy is used (e.g., chloramphenicol at 30 mg/liter for pLysE or pLysL transformation). Replace step 6 with the following and return to steps 7 and 8 of Basic Protocol 2. 6b. Induce protein expression by adding IPTG to the culture to achieve a final concentration of 0.5 to 1.0 mM (10 to 20 ml of 100× stock per liter). Incubate for another 3 hr before taking a second (final) 1-ml sample. TRANSFORMATION BY ELECTROPORATION Electroporation is a quick, simple, and efficient method for introducing foreign DNA into E. coli (i.e., transformation). Cultures of the appropriate host strain are grown to exponential phase in rich medium to produce competent cells (i.e., cells able to take up foreign DNA). Competent cells can be stored frozen at −80°C until use. Plasmid DNA is introduced by subjecting cells to an electrical pulse, using a commercially available electroporation apparatus, and cells are allowed to recover in rich medium supplemented with glucose. After the transformation, strains can be stored (see Support Protocol 3) and/or used for protein expression (see Basic Protocol 1, Basic Protocol 2, Alternate Protocol 1, and Alternate Protocol 2). Plasmid stability analysis (see Support Protocol 2) is not absolutely required and should be carried out only in cases where expression with a given strain is not reproducible. Several different host strains can be transformed in the same experiment to allow investigation of the influence of the host on protein expression efficiency. Materials Host E. coli strain LB medium (see recipe) Water, ice-cold 10% (w/v) glycerol, ice-cold Plasmid DNA SOC medium (see recipe) LB plates with appropriate antibiotic (see recipe)
SUPPORT PROTOCOL 1
Production of Recombinant Proteins
5.2.5 Current Protocols in Protein Science
1-liter and 50-ml centrifuge bottles, chilled Centrifuge (e.g., Sorvall RC3B equipped with H6000A rotor, or equivalent), 4°C 1.5-ml sterile microcentrifuge tubes, chilled Electroporation apparatus (e.g., E. coli Pulser Apparatus; Bio-Rad) Electroporation cuvettes, 0.2-cm electrode gap, sterile and prechilled Prepare the competent cells 1. Prepare an overnight culture with the E. coli host strain to be transformed by inoculating a 100-ml flask containing 20 ml LB medium with a single colony from an LB agar plate. As the host strain is plasmid free, do not add any antibiotic; with shaking, grow at 37°C. The next morning, verify that cultures have reached stationary growth phase (OD600 of 1 to 2). In the case of T7 expression and when using the additional pLys plasmids (optional; see Alternate Protocol 2), the host already contains a plasmid and should be kept under selective pressure (30 mg/liter chloramphenicol).
2. Transfer the overnight culture to 800 ml LB medium in a 3-liter Erlenmeyer flask. Incubate cells 1 to 2 hr in a 37°C orbital shaker with vigorous agitation (200 rpm). 3. Verify that the cells are in full exponential growth phase (OD600 of 0.5 to 0.6). Transfer cells to a 1-liter chilled centrifuge bottle. 4. Centrifuge 15 min at 4000 × g, 4°C. Discard supernatant and resuspend the pellet in 5 to 10 ml ice-cold water. Bring to a final volume of 800 ml with ice-cold water. 5. Repeat step 4 twice. 6a. If cells are to be transformed the same day: Resuspend the pellet in ice-cold water to a final volume of 50 ml, and pellet again in a chilled 50-ml centrifuge bottle. Discard supernatant, estimate the volume of the pellet, and add an equal volume of ice-cold water. Vortex, distribute 100-µl aliquots into prechilled 1.5-ml sterile microcentrifuge tubes, and keep on ice. Proceed to step 7. 6b. If competent cells are to be stored frozen: Resuspend the pellet in ice-cold 10% glycerol to a final volume of 50 ml, and pellet again in a 50-ml centrifuge bottle. Discard supernatant, estimate the volume of the pellet, and add an equal volume of ice-cold 10% glycerol. Vortex, place 100-µl aliquots into prechilled 1.5-ml microcentrifuge tubes, and store in a −80°C freezer. To use frozen competent cells for transformation, thaw microcentrifuge tubes on ice and proceed to step 7.
Transform the competent cells 7. Add 5 pg to 0.5 µg plasmid DNA (in 1 to 5 µl) to 100 µl competent cells. Mix by inverting the tube several times by hand. Keep on ice. Quantities of plasmid DNA can be estimated from average yields of a plasmid prep, or measured by UV absorbance. Some host cells (such as E. coli B) require more DNA for efficient transformation, depending on their ability to restrict foreign DNA.
8. Set the electroporation apparatus to 2.5 kV, 25 µF, 200 Ω. Selection of Escherichia coli Expression Systems
Some variation of the resistance (100 to 400 Ω) may be necessary to obtain optimal results (see Seidman et al., 1989).
9. Transfer the DNA and cells to a prechilled electroporation cuvette, wipe away ice and
5.2.6 Current Protocols in Protein Science
water from the cuvette outer walls, place the cuvette into the electroporation chamber, and apply the electrical pulse. Include a control with no DNA to ensure that transformants arise only by introduction of foreign DNA into the competent cells.
10. Remove the cuvette, immediately add 1 ml SOC medium and transfer the contents of the cuvette to a 20-ml sterile culture tube. Incubate 60 min with moderate shaking. The incubation temperature depends on the expression vector. Use 30°C for pL-based expression plasmids and 37°C for others.
11. Spread aliquots of the transformation culture (10 to 100 µl) on LB plates with appropriate antibiotic. Check for appearance of individual colonies after 24 to 48 hr of incubation. 12. Individually streak and label 5 to 10 colonies from transformation plates onto fresh plates with appropriate antibiotic to ensure they are pure, viable, and antibiotic resistant. 13. Store an aliquot of each strain (see Support Protocol 3). Proceed to the appropriate expression protocol (see Basic Protocol 1, Basic Protocol 2, Alternate Protocol 1, and Alternate Protocol 2). PLASMID STABILITY ANALYSIS Analysis of plasmid stability is important for assessing the quality of recombinant strains. This check is not absolutely required and is really only needed in cases where expression with a given strain is not reproducible. An aliquot of the frozen stock is thawed and serial dilutions of the strain are prepared and spread on plates containing rich medium with or without the antibiotic used to maintain the plasmid. Plated cultures are grown overnight, colonies counted, and the percentage of plasmid-carrying cells in the population determined.
SUPPORT PROTOCOL 2
Materials PBS (APPENDIX 2E) Transformed E. coli host strain (Basic Protocols 1 and 2, Alternate Protocols 1 and 2) LB plates with and without antibiotic (see recipe) 20-ml sterile culture tubes Sterile triangular glass rod NOTE: Cells should be kept in an ice bath throughout the procedure to minimize alterations to the samples. 1. Set up a rack of 20-ml sterile culture tubes and place 9 ml PBS in each tube. Pipet a 1-ml sample of the transformed E. coli host strain to be analyzed into the first tube. Vortex for 5 sec. An OD600 reading should be done before diluting the culture to facilitate steps 2-3.
2. With a new sterile pipet, transfer 1 ml from the previous tube into the next tube containing PBS. 3. Repeat step 2 until the final desired dilution is reached. The dilution range depends on the initial density of the culture. For an OD600 of 1, the final dilution factor should be 10−6.
Production of Recombinant Proteins
5.2.7 Current Protocols in Protein Science
4. Spread 100 µl of the final three dilutions onto triplicate sets of LB plates with and without antibiotic. Use a sterile triangular glass rod to achieve an even distribution of the cells on the plate. Start with the highest dilution (i.e., with the fewest cells) to minimize effects of carryover from one plate to the next. As 0.1 ml is spread on the plate and as a culture at an OD600 of 1 contains ∼109 cells/ml, diluting the culture to a maximum of 10−6 should give ∼100 colonies per plate.
5. Incubate plates overnight at the appropriate temperature. A temperature of 30°C should be used for pL-based expression strains and 37°C for all others. Incubate the plates upside down to collect water condensation in the plate cover.
6. The next morning, count the colonies on each plate. Calculate the number of cells present in the original sample that grew on LB and the number that grew on LB plus antibiotic. Determine the ratio of the two numbers to obtain the percentage of plasmid-carrying cells in the population. Serial dilutions can be made by a different combination of volumes leading eventually to a different dilution factor (0.1 ml into tubes containing 0.9 ml, 0.01 ml in 0.99 ml, etc.) For best accuracy, count plates with >30 colonies. Strains showing more than 50% plasmid retention before induction of protein expression are acceptable. SUPPORT PROTOCOL 3
STRAIN STORAGE For long-term storage, recombinant E. coli strains are kept at low temperature, frozen with a cryopreservative agent. A temperature of −80°C is sufficient for most strains. Multiple strains can be prepared in parallel and frozen in a single operation. It is good practice to keep at least one frozen copy that is not thawed and frozen repeatedly. The frozen stocks can be retrieved from storage and used for protein expression (Basic Protocols 1 and 2, and Alternate Protocols 1 and 2). Materials Transformed E. coli host strain (see Support Protocol 1) LB medium (see recipe) Antibiotic stock solution (see recipe) 20% (w/v) glycerol, dispersed in sterile cryovials (see recipe) LB plate (see recipe) 20-ml culture tube 30° or 37°C orbital shaker 1. Grow an overnight culture of transformed E. coli host strain by inoculating 2 ml of LB medium (supplemented with appropriate antibiotic, if necessary) in a 20-ml culture tube with a single colony from a fresh agar plate. Incubate the tube overnight on an orbital shaker. The next morning, verify that cells are in stationary phase (OD600 of 1 to 2). The incubation temperature is dependent on the expression vector. Use 30°C for pL-based systems and 37°C for others.
2. Inoculate a 20-ml tube (containing 2 ml LB plus appropriate antibiotic) with 50 µl from the overnight culture. Incubate for 2 to 3 hr. Selection of Escherichia coli Expression Systems
5.2.8 Current Protocols in Protein Science
3. Add 0.9 ml culture to each cryovial, label, vortex well, and freeze immediately. Store frozen strains at −80°C. 4. To start a new culture from the frozen stock, scrape the surface of the vial with a sterile loop and either streak on an LB plate or dip into liquid medium. Return the cryovial to −80°C for storage. Several hundred strains can be stored in a single freezer, but it is advisable to have alarm and safety devices against mechanical or electrical failure.
PREPARATION OF SAMPLES FOR ANALYSIS BY SDS-PAGE Electrophoretic separation of proteins in a cell extract (total lysate or fraction of the cell) by SDS-PAGE is described in detail in UNIT 10.1. This protocol describes only the preparation of the samples and applies to material derived from Basic Protocol 1, Basic Protocol 2, Alternate Protocol 1, and Alternate Protocol 2. The objective is to bring all proteins of the sample into solution in a denatured state so the mixture can be resolved by one-dimensional electrophoresis in SDS gels. Solubilization is achieved by sonicating samples in a buffer containing detergent and reducing agents, followed by brief heating. A large excess of sample is prepared so that it can be reloaded on several gels, if necessary. Once solubilized in SDS sample buffer, samples can be stored for years at −20°C.
SUPPORT PROTOCOL 4
Materials Sample of interest 2× and 1× SDS sample buffer (UNIT 10.1) Toothpicks, sterile Sonicator (Branson, with microtip) 1a. For soluble samples (e.g., soluble cellular proteins, periplasmic extracts, extracellular medium): Add the sample of interest to an equal volume of 2× SDS sample buffer in a 1.5-ml microcentrifuge tube and proceed directly to step 4. 1b. For pellet samples (total cells, insoluble fraction, trichloroacetic acid precipitate): Transfer 5 to 10 mg pellet material from the sample of interest with a sterile toothpick to a preweighed 1.5-ml microcentrifuge tube. 2. Add 1× SDS sample buffer, using 400 µl per 10 mg pellet. 3. Sonicate 10 sec with a sonicator (with power setting at 30 to 50 W) to resuspend the pellet. Be careful to avoid foaming by keeping the sonicator probe well below the liquid surface at all times.
4. Heat to 90°C for 5 min in a heating block. After cooling, samples are ready to be applied to gel wells for electrophoresis (UNIT 10.1). Loading amounts are based on gel and well dimensions, as well as the detection system to be used (e.g., silver staining is 100 times more sensitive than Coomassie blue staining; UNIT 10.5). Recommended amounts are as follows: 0.5 to 1 ìl for microgels (e.g., 4.3 × 5 cm; 0.45 mm thick), 5 to 10 ìl for minigels (e.g., 7.3 × 8.3 cm; 1 mm thick), and up to 20 ìl for full-size gels (e.g., 14 × 14 cm; 1.5 mm thick).
Production of Recombinant Proteins
5.2.9 Current Protocols in Protein Science
SUPPORT PROTOCOL 5
SOLUBILITY ANALYSIS Analysis of the solubility of recombinant proteins is needed to determine the subsequent purification strategy (UNIT 6.1). The protocol is based on mechanical disruption of cells followed by high-speed centrifugation to separate the soluble fraction from the insoluble fraction. Samples are then prepared for SDS-PAGE analysis (see Support Protocol 4). Materials Transformed E. coli host cells expressing the protein of interest 25 mM HEPES (pH 7.6; autoclave and store up to 1 yr at 4°C) 50- and 5-ml centrifuge tubes Centrifuge (e.g., Sorvall RC5B equipped with SS34 rotor), 4°C Sonicator NOTE: Cells should be kept in an ice bath throughout the procedure to minimize alterations to the samples. 1. Harvest 25 ml transformed E. coli host cells expressing the protein of interest by centrifuging 15 min at 4000 × g, 4°C, in a 50-ml centrifuge tube. 2. Discard the supernatant and resuspend the pellet in 1 to 1.5 ml 25 mM HEPES (pH 7.6) to obtain an OD600 of ∼40. 3. Set the tube in an ice bath and sonicate three times for 1 min each, allowing 30-sec intervals between pulses. 4. Place a 100-µl aliquot of the cell lysate (total protein) in a 1.5-ml microcentrifuge tube and keep at −20°C for later analysis (step 7). The 30-sec intervals allow the suspension to cool down between sonications. Be careful to avoid overheating at all stages.
5. Transfer the suspension to a 5-ml centrifuge tube and centrifuge 30 min at 30,000 × g (16,000 rpm in an SS-34 rotor), 4°C. 6. Pour off supernatant (soluble fraction) into another 5-ml tube to separate it from the pellet (insoluble fraction). 7. Prepare the three fractions (total, soluble, and insoluble proteins) for analysis by SDS-PAGE (see Support Protocol 4). The proportion of soluble/insoluble proteins can be quantified after SDS-PAGE separation (UNIT 10.1), staining of gels (UNIT 10.5), and scanning densitometry of the gels.
Selection of Escherichia coli Expression Systems
5.2.10 Current Protocols in Protein Science
PREPARATION OF PERIPLASMIC EXTRACTS This protocol describes a small-scale procedure to confirm the expression of proteins that are designed to be localized to the periplasmic space of E. coli. It is based on subjecting cells to an osmotic shock in a concentrated sucrose solution, which mechanically forces the contents of the periplasm through the outer membrane and cell wall.
SUPPORT PROTOCOL 6
Materials Transformed E. coli cells expressing the protein of interest PBS (APPENDIX 2E) 20% (w/v) sucrose/10 mM Tris⋅Cl (pH 7.5), ice-cold 0.5 M EDTA (pH 8.0) Ice-cold water Microcentrifuge, 4°C NOTE: Cells should be kept in an ice bath throughout the procedure to minimize alterations to the samples. Release the periplasmic proteins by osmotic shock 1. Place 1 ml transformed E. coli cells expressing the protein of interest in a 1.5-ml microcentrifuge tube and microcentrifuge 2 min at maximum speed, 4°C. Depending on the culture density, adjust the volume of the sample so that it contains ∼2 × 109 cells. (An OD600 of 1 is equivalent to 109 cells/ml).
2. Discard supernatant and resuspend the pellet in 1 ml PBS. Vortex tube well. 3. Pellet the cells again in the microcentrifuge for 2 min. 4. Discard supernatant and resuspend the pellet in 150 µl cold 20% sucrose/10 mM Tris⋅Cl (pH 7.5). Vortex well and add 5 µl of 0.5 M EDTA (pH 8.0). 5. Save a 50-µl aliquot representing the whole-cell fraction and store at −20°C. Leave remainder of the sample on ice for 10 min. The short incubation allows time for the EDTA to loosen the cell wall structure.
Separate the sample into periplasmic and cytoplasmic/membrane fractions 6. Pellet the cells for 5 min in the microcentrifuge. 7. Discard supernatant and resuspend the pellet by vigorous vortexing in 100 µl ice-cold water. 8. Leave on ice for 10 min and then repeat step 6. 9. Carefully collect and save the supernatant, which contains the periplasmic fraction in a 1.5-ml microcentrifuge tube and store at −20°C. 10. Eliminate remaining fluid from the pellet with a cotton-tipped swab and resuspend the sample in 100 µl ice-cold water. Save this solution as the cytoplasmic and membrane fraction. 11. Prepare the three fractions (whole-cell, periplasm, and cytoplasm/membrane fractions) for SDS-PAGE analysis (see Support Protocol 4).
Production of Recombinant Proteins
5.2.11 Current Protocols in Protein Science
SUPPORT PROTOCOL 7
PREPARATION OF EXTRACELLULAR MEDIUM SAMPLES Expression systems that direct extracellular secretion of the desired protein can be checked by analyzing samples of the culture medium. This protocol describes treatment of culture media with trichloroacetic acid (TCA) to precipitate extracellular proteins and thus concentrate proteins for subsequent analysis. Addition of bovine serum albumin (BSA) may be helpful if protein concentrations in the medium are low. Materials Transformed E. coli cells expressing the protein of interest 2 g/liter BSA (optional; sterilize through a 0.22-µm filter and store ≤1 month at 4°C) 200 g/liter trichloroacetic acid (TCA; sterilize through a 0.22-µm filter and store ≤1 month at 4°C) 70% (v/v) ethanol 0.22-µm filters suitable for aqueous solutions Microcentrifuge, 4°C Ice bath 1. Place a 1-ml culture of E. coli cells expressing the protein of interest in a 1.5-ml microcentrifuge tube and microcentrifuge 2 min at maximum speed, 4°C. 2. Collect the supernatant and pass it through a 0.22-µm filter into a 1.5-ml microcentrifuge tube. Discard pellet. 3. Add 100 µl of 2 g/liter BSA, if necessary. BSA acts as a carrier and nucleus for precipitation. It can be omitted if the medium already contains >100 mg/l of protein.
4. Add 1 ml of 200 g/liter TCA, vortex well, and leave the tube on ice for 15 to 30 min to allow proteins to precipitate. 5. Microcentrifuge for 10 min at maximum speed, 4°C. 6. Discard supernatant and wash the pellet at least twice with 2 ml of 70% ethanol. The washings remove the TCA, which would otherwise disturb subsequent analysis by SDS-PAGE.
7. Prepare the pellet for analysis by SDS-PAGE (see Support Protocol 4).
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Selection of Escherichia coli Expression Systems
Antibiotic stock solutions (1000× concentrated) Ampicillin: Prepare a 40 g/liter stock in water. Store ≤2 weeks at 4°C. Chloramphenicol: Prepare a 30 g/liter stock in ethanol. Store ≤1 year at −20°C. Kanamycin: Prepare a 40 g/liter stock in water. Store ≤1 year at 4°C. Tetracycline: Prepare a 5 g/liter stock in 70% ethanol. Store ≤1 year in the dark at −20°C. Sterilize all stock solutions by filtration through a 0.22-µm filter before use. Aliquot in 5 to 10 portions, depending on quantities needed.
5.2.12 Current Protocols in Protein Science
Glycerol Dispense 1 ml of 20% (v/v) glycerol into 2.5-ml cryovials. Autoclave with the caps loose. After sterilization, tighten the caps. Store up to 1 year at 4°C. An entire rack of cryovials can be prepared at once, as they can be stored for a long time.
Indole-3-acrylic acid (IAA) Store IAA as a powder at 4°C. Prepare solution fresh as a 400× concentrated solution in ethanol (10 g/liter). Isopropyl-β-thiogalactopyranoside (IPTG) Store IPTG as a powder at −20°C. Prepare solution fresh as a 100× concentrated solution in water (50 mM, i.e., 11.9 mg/liter). Sterilize by filtration before use. LB medium Per liter: 10 g Bacto-tryptone 5 g yeast extract 5 g NaCl 10 ml 1 N NaOH (to pH 7.0) H2O to 1 liter Sterilize 30 min at 121°C Dispense into 20-ml culture tubes (2 or 5 ml/tube) or shaker flasks (see Commentary). Plug opening with rigid aluminum cap or with hydrophobic cotton wool covered with aluminum foil. Antibiotics should be added (if appropriate) just before use. The original recipe for LB medium (sometimes referred to Luria or Lenox broth) does not contain NaOH. There are many different recipes for LB that differ only in the amount of NaOH added. We use this formula in our own work. Even though the pH is adjusted to near 7.0 with NaOH, the medium is not highly buffered, and the pH of a culture growing in it drops as it nears saturation.
M9 minimal medium Per liter: 6 g Na2HPO4 3 g KH2PO4 1 g NH4Cl 0.5 g NaCl 3 mg CaCl2 H2O to 1 liter Sterilize 30 min at 121°C Dispense into 20-ml culture tubes (2 or 5 ml/tube) or shaker flasks (see Commentary). Plug opening with rigid aluminium cap or with hydrophobic cotton wool covered with aluminum foil. Antibiotics should be added (if appropriate) just before use.
Production of Recombinant Proteins
5.2.13 Current Protocols in Protein Science
SOC medium Per liter: 5 g yeast extract 20 g Bacto-tryptone 580 mg NaCl (10 mM) 186 mg KCl (2.5 mM) 940 mg MgCl2 (10 mM) 1.2 g MgSO4 (10 mM) 3.6 g glucose (20 mM) H2O to 1 liter Sterilize 30 min at 121°C Dispense into 20-ml culture tubes (2 or 5 ml/tube) or shaker flasks (see Commentary). Plug opening with rigid aluminium cap or with hydrophobic cotton wool covered with aluminum foil. Antibiotics should be added (if appropriate) just before use. The amounts given in the recipe refer to the anhydrous salts.
Plates Use liquid medium formulation (see recipes for LB, SOC, and M9 media) and add 20 g/liter agar. For LB or SOC plates, autoclave the agar together with the other ingredients. For M9 plates, autoclave 20 g agar in 800 ml water for 15 min, then add sterile concentrated M9 medium (i.e., make up components in 200 rather than 1000 ml water). Cool the agar solutions to about 50°C before adding desired supplements (i.e., antibiotics; see recipe). The media will stay liquid at 50°C but will rapidly solidify if the temperature drops below 45°C. Pour 32 to 40 ml medium into each sterile disposable petri dish to get 25 to 30 plates per liter. Allow the agar to solidify. Dry the plates by leaving them out at room temperature for 2 or 3 days, or by leaving them with the lids off for 30 min in a 37°C incubator or in a laminar-flow hood. Store dry plates at 4°C, wrapped in the original bags used to package the empty plates. Antibiotic-containing plates have a limited shelf-life: 2 weeks for ampicillin; 6 weeks for chloramphenicol, kanamycin, and tetracycline. COMMENTARY Background Information
Selection of Escherichia coli Expression Systems
As described in UNIT 5.1, the choice of an expression system and corresponding host strain for the production of a recombinant protein in E. coli is dictated by the ultimate goal of the study. If the protein is needed for structural studies requiring tens to hundreds of milligrams of a given protein, correctly folded, then either secretion of the protein or high-level production inside the cell (followed by protein refolding if the protein is in aggregated form; UNIT 6.5) may be the methods of choice. If an antigen is needed for raising antibodies, gene fusion strategies are often the method of choice. The toxicity of some recombinant proteins will also drive a requirement for very tightly repressed systems such as T7 RNA polymerase, which have particular host requirements (see Alternate Protocol 2). In other words, the type of
protein and its solubility dictate the vector construction and, thus, the E. coli host strain used. The pL promoter from lambda phage initiates high levels of transcription that can be repressed by the cI protein (Shatzman et al., 1990). A temperature-sensitive mutant of the repressor, cI857, which is inactive at high temperature, allows induction by a temperature shift from 30° to 42°C (Remaut et al., 1981). The repressor gene can be carried on the host chromosome or on the same plasmid as pL. In general, induction of pL is most efficient when the cells are in full exponential growth phase in a rich medium (see Basic Protocol 1). Tac is a hybrid between lac and trp promoters and is regulated by the same lacI repressor as lac. These promoters are inducible by the addition of a nonmetabolizable galactose analog, isopropyl-β-thiogalactopyranoside
5.2.14 Current Protocols in Protein Science
(IPTG), which binds and inactivates the repressor. The lacI repressor gene can be integrated in the host chromosome, on the same plasmid as the target gene, or on another compatible one. This expression system (see Alternate Protocol 1) is extensively used for recombinant protein expression (Harris, 1983). T7-driven expression (see Alternate Protocol 2) is based on the efficiency and specificity of the phage T7 RNA polymerase for initiating transcription from T7 promoters (Tabor, 1990; Studier and Moffat, 1986). Two elements are introduced into the host cell: the T7 polymerase gene, under control of an inducible promoter (lacUV5), which is integrated into the host DNA as a λDE3 lysogen; and the target gene, which is introduced on a plasmid under control of the T7 promoter. This plasmid also carries an antibiotic (ampicillin or kanamycin) resistance gene for selection. Two hosts are routinely used: E. coli BL21 and HMS174 (Table 5.2.1). For toxic target gene products and because basal levels of the T7 polymerase can promote some transcription of the target gene, a third optional element—T7 lysozyme—is sometimes added. T7 lysozyme binds the T7 polymerase, thereby inactivating it and eliminating transcription of the target gene under noninduced conditions. The T7 lysozyme gene is carried by either of two plasmids (pLysE or pLysS) and expressed constitutively. Both plasmids also carry a selection marker (chloramphenicol-resistance gene). The pET system is a series of vectors for cloning into the T7 promoter system (Novagen). Some vectors carry an ampicillin-resistance gene; others carry kanamycin resistance. Novagen distributes a complete set of tools associated with the system, including host strains, competent cells, and pLysS and pLysE vectors. The promoter for the tryptophan operon (trp) is strong and well-repressed under basal conditions (Somerville, 1988). Tryptophan present in the culture medium acts as a corepressor by binding to the trp repressor. This complex binds to the operator and prevents transcription. As the cells consume tryptophan for growth, its concentration decreases to a point where the operator is unbound and transcription of genes downstream (see Basic Protocol 2) is initiated. Addition of indole-3acrylic acid (IAA), a tryptophan structural analog, inactivates any residual repressor. High-copy-number plasmids are desirable for high-level expression but also often result in plasmid loss. An analysis of plasmid stability (see Support Protocol 2) allows evaluation of
plasmid loss and control of the quality and integrity of recombinant strains. The relative proportions of soluble and insoluble recombinant protein (also referred to as inclusion bodies) are affected by many variables such as protein primary sequence, expression system, and host strain. Environmental factors such as temperature and pH of the culture are also key determinants as there is a consensus that lower temperatures (20° to 25°C) lead to a higher proportion of soluble protein. The soluble/insoluble ratio dictates the subsequent purification strategy (see Chapter 6). It is therefore essential to analyze the recombinant protein solubility (see Support Protocol 5), which, in some cases, can be considered more important than the total protein expression level. Secretion to the periplasmic space should be evaluated, because this compartment provides a more oxidative environment than the cytoplasm and favors disulfide bridge formation. With an appropriate expression system, it is possible to direct secretion of the recombinant protein to the periplasmic space of E. coli (Hsiung et al., 1986). Analysis of expression and confirmation of the proper localization thus require the preparation of a periplasmic extract. The osmotic shock method (see Support Protocol 6) to prepare periplasmic extracts was originally described by Koshland (1980). Fusion to periplasmic (e.g., MalE, PhoA) or outer membrane (e.g., OmpA, LamB) protein signal peptides has been extensively used to direct recombinant proteins to the periplasm (Hsuing et al., 1986; Hewinson and Russell, 1993; Karim et al., 1993). This approach was successfully applied to antibody fragments (Carter et al., 1992; Plückthun, 1991). Complete secretion to the extracellular medium has also been achieved by fusing the gene of interest at the N terminus to protein A signal peptide from S. aureus or at the C terminal domain of hemolysin A (reviewed in Blight and Holland, 1994). Several expression systems are now available that allow efficient secretion of the recombinant protein into the extracellular medium (Blight and Holland, 1994). However, the secreted protein is usually quite dilute and concentration of the protein is necessary for analyzing and confirming expression by SDSPAGE. Ultrafiltration or precipitation can be used for this purpose. This protocol describes precipitation by TCA, which is quick, reliable, and inexpensive. Several samples can be handled in the same batch.
Production of Recombinant Proteins
5.2.15 Current Protocols in Protein Science
Critical Parameters and Troubleshooting
Selection of Escherichia coli Expression Systems
The temperature shift necessary to induce pL will also induce a heat-shock response, and in particular turn on some protease genes. Also, some host strains (such as E. coli HB101) do not tolerate 42°C, and growth is drastically reduced at this temperature. Tac and lac are weaker than pL or trp, but still drive high-level expression of some proteins. They are not recommended for toxic gene products because repression is often incomplete. This occurs because, in the lac operon, normal repression is mediated in part by an operator site internal to the lacZ gene, which is not usually found in the target gene. It is also important to note that repression is more efficient with the mutated repressor gene lacIQ. The expression protocols can be scaled to any size shaker flask, as long as the culture is kept well aerated (maximum ratio of liquid/flask volume of 0.3). Care should also be taken to ensure that incubation temperatures are accurate. It takes ∼95 min for an 800-ml culture in a 2-liter flask to heat from 20° to 42°C in an air shaker at 200 rpm. Pre-warming of culture media can be done adequately in an air shaker if enough time is allowed for the warmup, but it is probably better if a water bath is used for temperature induction steps. Attempts to increase solubility of proteins formed in aggregates or inclusion bodies have included coexpression with chaperones, manipulation of temperature and other environmental factors, and secretion of the desired protein to the periplasmic space and/or the extracellular medium. These strategies have been reviewed (Hockney, 1994). Incomplete repression is quite frequent with genes under control of a lac-type promoter. In fact, for some cases, one may find that induction does not increase the constitutive level of expression. This is due in part to the mechanism of repression by the LacI protein. It has been shown that the repressor binds the natural lac operator in a DNA region that also contains part of the 5′ coding sequence for LacZ. If this region is exchanged for that of a foreign gene, then the affinity of the repressor for the operator is decreased and transcription can occur at a reduced level. The optional pLysE or pLysS plasmids for the T7 system should be used with considerable caution. The presence of pLysE leads to a high level of lysozyme in the cell, which can dampen expression and eventually weaken the cell wall. It also, however, brings an additional advantage
for purification: a simple freeze-thaw cycle usually effects cell lysis efficiently. In the case of no detectable expression, we advise (1) trying another promoter system; (2) testing several different host strains; (3) fusing the gene of interest to well-expressed DNA sequences such as LacZ, glutathione transferase (GST), thioredoxin, or protein A, to aid expression and/or recovery (UNIT 5.1). Optimization of codon usage is a topic outside the scope of this unit (Robinson et al., 1984). Overexpression of rare tRNAs and mutagenesis of rare codons have been used for enhancing expression of heterologous proteins in E. coli (De Boer and Hui, 1990). DNA used for electroporation should be as free of salts as possible, as they contribute to decreasing the resistance of the sample and can lead to arcing. It is quite useful to make large batches of competent cells (e.g., 800-ml cultures) and freeze them in aliquots. Several host strains can be transformed quickly (in 2 hr) tested in parallel for expression. It should also be stressed that the goal of the transformation is only to obtain a few colonies of E. coli cells carrying the expression vector. Efficiency is not a major concern. Usually a few hundred transformant colonies are generated; only a small proportion is tested for the ability to express the recombinant protein of interest. SDS-PAGE analysis of total E. coli cell proteins is straightforward and reproducible. One minor difficulty may arise from pellet sample preparation—if DNA has not been broken down well enough at the sonication step, the sample will be highly viscous and hard to load on the gel. In this instance, sonication should be repeated. At low levels of expression, it is possible that Coomassie staining will not be sensitive enough. It is not recommended that more sensitive staining methods (e.g., silver staining) be tried, as this will only increase the background. Rather, immunoblotting should be used to analyze the proteins. It is also good practice to include negative controls, such as noninduced cell extract or extract of induced cells carrying an empty vector, as well as positive controls, such as samples from previous experiments or partially pure protein where available.
Anticipated Results Expression levels of the desired proteins may range from <1% to >40% of total cell protein. In shaker-flask cultures this is equivalent to hundreds of milligrams per liter, which can be increased to grams-per-liter level in
5.2.16 Current Protocols in Protein Science
fermenters (where greater cell densities are achieved; see UNIT 5.3). The expressed product can usually be visualized by running appropriate extracts of cells on SDS-polyacrylamide gels and staining with Coomassie blue. The majority of the proteins produced will be insoluble, but the material can often be solubilized and renatured to an active state (UNITS 6.3, 6.4, & 6.5).
Hsiung, H.M., Mayne, N.G., and Becker, G.W. 1986. High level expression, efficient secretion and folding of human growth hormone in E. coli. Bio/Technology 4:991-995.
Time Considerations
Plückthun, A. 1991. Antibody engineering: Advances from the use of Escherichia coli expression systems. Bio/Technology 9:545-551.
Transformation of cells requires about 1 day. Cell growth, induction, and harvesting take 6 to 8 hr depending on the expression system and strain of E. coli. Analyzing plasmid stability if necessary extends into the following day. It is often convenient to conduct analysis of samples by SDS-PAGE the next day, as gel preparation, running, staining, and destaining will require several hours. The requirement to prepare overnight cultures to be induced or analyzed must also be taken into account. Gel preparation can be avoided by using commercially available gels (e.g., Phast-gels from Pharmacia Biotech or minigels from Novex). Overall, 2 to 3 days should be required for full testing of a given expression vector in different host strains.
Literature Cited Blight, M.A. and Holland, B. 1994. Heterologous protein secretion and the versatile E. coli haemolysin translocator. Trends Biotechnol. 12:450-455. Carter, P., Kelley, R.F., Rodrigues, M.L., Snedecor, B., Covarrubias, M., Velligan, M.D., Wong, W.L.T., Rowland, A.M., Carver, M.E., Yang, M., Bourell, J.H., Shephard, H.M., and Henner, D. 1992. High level Escherichia coli expression and production of a bivalent humanized antibody fragment. Bio/Technology 10:163-167. De Boer, H.A. and Hui, A.S. 1990. Sequences within ribosome binding site affecting messenger RNA translatability and method to direct ribosomes to single messenger RNA species. Methods Enzymol. 185:103-114. Goeddel, D.V. (ed.) 1990. Gene expression technology. Methods Enzymol. 185. Harris, T.J.R. 1983. Expression of eukaryotic genes in E. coli. Genet. Eng. 4:127-185. Hewinson, R.G. and Russell, W.P. 1993. Processing and secretion by Escherichia coli of a recombinant form of the immunogenic protein MPB70 of Mycobacterium bovis. J. Gen. Microbiol. 139:1253-1259. Hockney, R.C. 1994. Recent developments in heterologous protein production in E. coli. Trends Biotechnol. 12:456-463.
Karim, A., Kaderbhai, N., Evans, A., Harding, V., and Kaderbhai, M.A. 1993. Efficient bacterial export of a eukaryotic cytoplasmid cytochrome. Bio/Technology 11:612-618. Koshland, D. and Botstein, D. 1980. Secretion of beta-lactamase requires the carboxy end of the protein. Cell 20:749-760.
Raleigh, E.A., Lech, K., and Brent, R. 1989. Selected topics from classical bacterial genetics. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 1.4.1-1.4.14. John Wiley & Sons, New York. Remaut, E., Stanssens, P., and Fiers, W. 1981. Plasmid vectors for high-efficiency expression controlled by the pL promoter of coliphage lambda. Gene 15:81-93. Remaut, E., Marmenout, A., Simons, G., and Fiers, W. 1987. Expression of heterologous unfused proteins in Escherichia coli. Methods Enzymol. 153:416-431. Robinson, M., Lilley, R., Little, S., Emtage, J.S., Yarranton, G., Stephens, P., Millican, A., Eaton, M., and Humphreys, G. 1984. Codon usage can affect efficiency of translation of genes in Escherichia coli. Nucl. Acids Res. 12:6663-6671. Rosenberg, S.A., Grimm, E.A., McCrogan, M., Doyle, M., Kawasaki, E., and Koths, K. 1984. Biological activity of recombinant human interleukin-2 produced in E. coli. Science 223:14121415. Sambrook, J., Fritsch, E.F., and Maniatis, T. 1989. In Molecular Cloning, A Laboratory Manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Seidman, C.E., Struhl, K., and Sheen, J. 1989. Introduction of plasmid DNA into cells. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 1.8.11.8.8. John Wiley & Sons, New York. Shatzman, A.R., Gross, M.S., and Rosenberg, M. 1990. Expression using vectors with phage λ regulatory sequences. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 16.3.1-16.3.11. John Wiley & Sons, New York. Somerville, R.L. 1988. The trp promoter of E. coli and its exploitation in the design of efficient protein production systems. Biotechnol. Genet. Eng. Rev. 6:1-41. Stahl, S.J. and Murray, K., 1989. Immunogenicity of peptide fusions to hepatitis B virus core antigen. Proc. Natl. Acad. Sci. U.S.A. 86:6283-6287.
Production of Recombinant Proteins
5.2.17 Current Protocols in Protein Science
Studier, F.W. and Moffat, B.A. 1986. Use of bacteriophage T7 RNA polymerase to direct selective high level expression of cloned genes. J. Mol. Biol. 189:113-130. Tabor, S. 1990. Expression using the T7 RNA polymerase/promoter system. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 16.2.1-16.2.11. John Wiley & Sons, New York.
Key References Goeddel, 1990. See above. Reviews most common expression systems in E. coli and provides a complete source of background information on the expression vectors as well as some hints on strategies for optimization.
Hockney, 1994. See above. Comprehensive review on advances in protein expression in E. coli. Provides useful information regarding solubility, secretion, and fusions. Also discusses the difficult problem of expressing complex proteins such as multi-transmembrane proteins in E. coli.
Contributed by Alain Bernard and Mark Payton Glaxo Institute for Molecular Biology Geneva, Switzerland
Selection of Escherichia coli Expression Systems
5.2.18 Current Protocols in Protein Science
Fermentation and Growth of Escherichia coli for Optimal Protein Production
UNIT 5.3
Large-scale production of recombinant proteins in Escherichia coli requires growth of cells in fermentors. Fermentation involves the transformation of simple organic compounds by the enzymatic machinery of E. coli, in this case to produce proteins. There are many choices of bacterial strain, vector, expression system, and protein localization and purification strategy that must be set before proceeding to batch fermentation. UNIT 5.1 gives an overview of protein expression in E. coli, and UNIT 5.2 outlines procedures for transforming cells and inducing synthesis of foreign proteins in various expression systems on a small scale in shaker flasks. Table 5.2.1 lists E. coli strains appropriate for use in fermentors. The Commentary of this unit discusses important characteristics of fermentation equipment. Starting with a strain of E. coli that has been transformed to produce the desired protein (UNIT 5.2) this unit focuses on production of recombinant proteins in batch fermentations (Basic Protocol 1). Alternate protocols introduce variations of fermentation systems that enable continuous growth and protein production in high-cell-density, fed-batch cultures (Alternate Protocol 1) and that permit labeling of recombinant proteins with heavy atom derivatives such as selenomethionine (Alternate Protocol 2) or with stable isotopes such as 2H, 13C, and 15N (Alternate Protocol 3). Production of labeled proteins facilitates structural studies by X-ray crystallography and nuclear magnetic resonance (NMR) spectroscopy. The protocols in this unit are designed for expression systems directing intracellular or periplasmic localization of recombinant proteins; however, in the case of extracellular secretion of the desired protein, the culture medium itself, rather than pelleted cells, would be saved, concentrated, and subjected to purification processes. Fermentation experiments require careful monitoring of cell growth and assurance of preinoculation sterility, which are described in Support Protocols 1 and 2, respectively. It must be kept in mind, however, that the outcome of fermentation experiments, even if derived from successful small-scale shaker flask cultures (UNIT 5.2), may be negatively affected by unexpected toxicity of the target protein in fermentation systems. PRODUCTION OF RECOMBINANT PROTEINS IN BATCH FERMENTATIONS
BASIC PROTOCOL 1
This protocol describes the basic sequence of operations, their timing, and the key issues associated with batch fermentations. Simple batch fermentations involve growth of fermenting organisms in a fixed amount of medium, without additional nutrients. The fermentor is set up, filled with fermentation medium (but not including heat-sensitive ingredients), and sterilized. Filter-sterilized heat-sensitive ingredients are then added, and environmental parameters are set to the desired values (using acid and base to adjust pH) before final preinoculation samples are taken to check sterility (Support Protocol 2). An inoculum prepared from a fresh overnight culture of the transformed E. coli strain is introduced into the fermentor and the cells allowed to grow. Synthesis of the recombinant protein following induction of cells is monitored, and cells are harvested by centrifugation using a batch pot or continuous feed centrifuge, depending on the volume of medium processed. Special fermentation equipment is required. Several commercial manufacturers produce fermentation equipment (Table 5.3.1) for laboratory use (1 to 20 liters) or small pilot-scale experiments (50 to 200 liters). For recombinant E. coli processes, users should pay particular attention to heat and oxygen transfer, process monitoring and control (particuContributed by Alain Bernard and Mark Payton Current Protocols in Protein Science (1995) 5.3.1-5.3.18 Copyright © 2000 by John Wiley & Sons, Inc.
Production of Recombinant Proteins
5.3.1 CPPS
Table 5.3.1
Fermentation Equipment Suppliers
Suppliera
Vessel size (liters)
B. Braun
2 to 50,000
Bioengineering
2.4 to 50000
Inceltech
2.5 to 800
New Brunswick Scientific Co. LSL Biolafitte New MBR
Control
Comments
DDCb, Computer supervision Analog, DDC, Computer supervision
Microprocessor-based instrumentation Special vessel designs (airlifts, fluidized beds, membrane dialysis) Complete plants Very high oxygen transfer
Analog, DDC, Computer supervision 1 to 15,000 Analog, DDC, Computer supervision 1 to 50,000 DDC, Computer supervision 3.5 to 100,000 Analog, DDC, Computer supervision
Very high oxygen transfer Windows-based control package Microprocessor-based instrumentation Complete plants
aFor complete listing of suppliers’ addresses, see SUPPLIERS APPENDIX. bDDC, direct digital control
larly if high-cell-density processes are needed; Alternate Protocol 1), and ease of maintenance and servicing. Utilities such as air, water, steam, and electricity are needed in defined qualities and quantities. Consult the fermentor manufacturer for specific requirements. Materials Transformed E. coli host strain expressing the protein of interest (UNIT 5.2) LB medium with antibiotic (UNIT 5.2) Antifoam: polypropylene glycol (PPG) 2000 (pure; store indefinitely at room temperature) Nutrient solution (optional, for fed-batch methods; see Alternate Protocol 1) ECPM1 medium (see recipe) 85% (w/v) H3PO4 28% (w/v) NH4OH 10 N NaOH Fermentor (Fig. 5.3.1) 5-ml and 50-ml sterile syringes and 1 × 40–mm needles Centrifuge, with capacity up to 6 liters per run (for 1- to 10-liter scale; e.g., Sorvall RC3B, equipped with H6000A rotor) or continuous feed centrifuge (for scale >10 liters; Fig. 5.3.2; e.g., Sharples AS16) Heat-sealable plastic bags and heat-sealing apparatus Additional reagents and equipment for inducing protein expression and for preparing samples for polyacrylamide gel analysis (UNIT 5.2)
Fermentation and Growth of E. coli for Optimal Protein Production
Prepare the inoculum and fermentor 1. Prepare an overnight culture of the transformed E. coli host strain expressing the protein of interest by inoculating a suitable amount of LB medium, supplemented with the appropriate antibiotic to maintain plasmid-carrying cells, with a single colony picked from an LB/antibiotic plate or retrieved from a vial of frozen cells (see UNIT 5.2). The next morning, read the OD600 against water as a blank to verify that the
5.3.2 Current Protocols in Protein Science
exhaust air
exhaust air filter
reflux cooler
flow meter air
drain cold water inoculum antifoam base acid
air inlet filter top
sparger reactor vessel
sample harvest
jacket
cold water stirrer agitator
drain steam drain mechanical seal
motor electrically controlled valves
manual valve
circulating pump for water jacket
Figure 5.3.1 Schematic diagram of a bacterial fermentor, depicting major functions. Variations in the design can be made to suit specific requirements.
culture has reached stationary phase (i.e., OD>1.5, see Support Protocol 1). Use the overnight culture to inoculate the fermentor. The size of the inoculum can be varied between 1% and 10% of the intended fermentation volume. For a 10-liter fermentor, the required inoculum of 100 to 1000 ml can be conveniently prepared in 500-ml to 5-liter shaker flasks (a maximum ratio of liquid/flask volume of 0.3 ensures that the culture is well aerated). Fermentors of sizes above 10 liters can be inoculated with several shaker flasks, representing 1% of the fermentor volume. The contents of all the inoculum flasks are pooled into a single large sterile container, fitted with the appropriate transfer line. The choice of antibiotic is dictated by the transformation vector, and the incubation temperature depends on the expression system (use 30°C for pL-based systems and 37°C for others). The inoculum is needed in step 15 after the fermentor has been loaded with fermentation medium and sterilized.
Production of Recombinant Proteins
5.3.3 Current Protocols in Protein Science
motor
cooling water out
supernatant
cooling water in
feed
Figure 5.3.2 Model AS16 continuous feed tubular bowl centrifuge (Alfa Laval). Other models (disc stack, with pellet ejection) are suitable for pilot-scale operations. Broth is fed by pressure at the bottom of the bowl, and cells sediment against a liner inside the bowl. Cell-free supernatant flows from the top of the bowl. The capacity is ∼6 liters. A cooling jacket is provided to avoid high temperatures.
2. Empty the reactor vessel of the fermentor and inspect it for residual biological material on the inner wall, agitator shaft baffles, or other parts. Between fermentation processes, the fermentor is usually kept partially filled with water.
3. Calibrate sensors: pH and pO2 probes, load cells, and pressure gauges according to manufacturer’s instructions. Although calibration is not necessary for all sensors and every run, it should be performed on a regular basis (every 10 runs or so). The pH and pO2 probes, however, should be recalibrated before each run because of the possible drift of the signal with time or fouling of the probe membrane. The calibration of the pO2 probe done here is a rough determination of whether the probe is responding to changes of dissolved O2. The precise calibration for pO2 is performed at step 10, after the probe is in the sterilized fermentation medium, in an environment closest to the one of the process.
4. Prepare accessory lines, transfer vessels, and ports, including septums and addition lines for inoculum, acid, base, antifoam, nutrients, and inducers. Sterilize by autoclaving. Also sterilize antifoam and, if applicable, nutrient solution, if possible while linked to the respective lines. Some nutrient solutions (e.g., for fed-batch processes) contain heat-sensitive components and should be sterilized according to the recipe (see Reagents and Solutions).
Fermentation and Growth of E. coli for Optimal Protein Production
5. Prepare ECPM1 medium by mixing all heat-sterilizable ingredients with water to ∼80% of the final volume. Pour into fermentor. For fermentation volumes exceeding 5 liters, dissolve all ingredients as a concentrate (at 100× the final required concentration) and pour into the fermentor.
5.3.4 Current Protocols in Protein Science
6. Adjust presterilization volume by adding water directly to the reactor vessel. The presterilization volume should be calculated from the final culture volume (which should not exceed 3⁄4 of the total fermentor volume) by deducting volumes of inoculum, heat-sensitive ingredients and antibiotic to be added and water that evaporates during sterilization. The addition of water resulting from steam condensation during sterilization must also be taken into account. The amounts of liquid loss or gain should not exceed 10% of the final culture volume and is best empirically determined by measuring accurately the liquid level pre- and post-sterilization in a mock trial sterilization with water. Culture volume does vary during the process, due to mass cell accumulation, sampling, additions of inducers and for pH and foam correction, but these phenomena usually result in a net volume variation that is negligible (except for fed-batch processes).
Sterilize the fermentor and medium 7. Start sterilization cycle. For autoclavable glass transfer vessels (usually 1 to 5 liters), disconnect all cables and tubings from the fermentor; allow the inside of the chamber to be ventilated through one or two lines (protected with air filters) that communicate with the reactor headspace. For transfer vessels that can be sterilized in situ, engage the automatic sequence controller. It is best to insert the temperature sensor of the autoclave into the temperature port of the reactor, if possible. This ensures that the autoclave control system will sense the exact temperature of the broth inside the reactor. For in situ sterilizable vessels, most fermentors are fitted with an automatic sequence controller that initiates and controls the whole sterilization cycle with predetermined parameters.
8. Sterilize at 121°C for 30 min. If cooling proceeds too quickly, an internal vacuum may form that will open the way to contamination. A cooling rate of ≤40°C/min is acceptable—if it is higher than this, restrict the cooling water flux.
9. As soon as the temperature reaches the range of 102° to 108°C, apply headspace pressure to the reactor with sterile air (0.05 to 0.1 bar). Start water flow through the reflux cooler to condense exhaust gases. Applying headspace pressure is important to avoid an internal vacuum and to create a barrier to any potential external contaminant. It is also good practice, once pressure has stabilized, to set a small air flow through the headspace (0.2 liters air/culture/min) to dry the exhaust gas filter.
Conduct final preparations for inoculation 10. For pH correction, pour 85% H3PO4 into one transfer vessel (acid) and 28% NH4OH into the other (base) under a biological hood. Connect acid, base, and antifoam lines aseptically to the fermentor. Once the temperature is ≤40°C, proceed to step 11. Aseptic connections are easily made by using a sterile needle to puncture a sterile septum port. Use flame sterilization and/or a 70% ethanol spray to ensure sterility. Some connectors can be sterilized again by steam after the connection is completed.
11. Add heat-sensitive ECPM1 medium ingredients (the 1 M MgCl2⋅6H2O previously sterilized by 0.22-µm filtration; see Reagents and Solutions) with a sterile syringe by puncturing a septum port with the needle. 12. Calibrate pO2 sensor by sparging air at 0.5 vol gas/vol culture/min with vigorous agitation (≥500 rpm) and adjusting the 100% saturation point. The zero point of the sensor can be calibrated by sparging N2 gas instead of air, or with a sensor simulator, but this point should not need frequent recalibration—after every 10 sterilization cycles should suffice.
Production of Recombinant Proteins
5.3.5 Current Protocols in Protein Science
13. Adjust all environmental parameters to the desired set points. Typical parameter settings are: temperature, 30° or 37°C; pH, 7.0; agitation, 300 rpm; air flow, 1 vol/vol culture/min; pO2, 100% air saturation.
14. Withdraw a small sample aseptically and check for sterility (see Support Protocol 2). A sterile fermentor can be prepared 2 to 3 days in advance. If this is done, it is best to recheck the sterility of a fresh sample taken just before inoculation. Because the results of this sterility check are not known for another 12 to 16 hr, the procedure is initiated and continued. In case of a contamination revealed after inoculation, the batch should be considered suspect and/or discarded. A massive contamination (which could occur if the fermentor has been prepared more than 24 hr in advance) will be obvious from the evolution of the pH and/or pO2 and turbidity of the sample. If any contamination is detected before inoculation, the entire run should be cancelled and the fermentor cleaned (see step 24).
Inoculate the fermentor, growth cells, and induce protein expression 15. Aseptically connect inoculum line to fermentor septum port. Inoculate fermentor by transferring the inoculum (step 1) aseptically into the reactor vessel. Disconnect inoculum line. A short spray of 70% ethanol on the septum and needle improves sterility. Inoculation is achieved either via an in-line peristaltic pump or by pressurizing the inoculum vessel with sterile air.
16. Monitor growth by measuring OD600 of samples taken at 1-hr intervals (see Support Protocol 1). When the target cell concentration is reached, proceed to induction. Target cell concentration depends on the strain and expression system. As a general rule, and in the absence of any previous experience with a given strain, a target of OD 5 to 10 is applicable. It is possible, however, that the same process can also be run with induction at a later stage (OD 15 to 20), but this needs empirical determination.
17. Induce protein expression via temperature or chemical induction using one of the following methods, as appropriate. a. For the pL system: Perform a fast temperature shift (within a few minutes) from 30° to 42°C and incubate 5 hr. b. For the trp promoter system: Add IAA to a final concentration of 25 mg/liter (see UNIT 5.2). Incubate to stationary phase (usually overnight). c. For the lac and tac promoter and T7 systems: Add IPTG to a final concentration of 0.5 to 1.0 mM (UNIT 5.2) and incubate 5 hr. 18. At 1-hr intervals, withdraw samples of the culture, monitor growth by OD600 measurements, and prepare them for polyacrylamide gel analysis, as described in UNIT 5.2, to monitor recombinant protein expression. A band of the expected molecular weight should appear in lanes of induced samples, but not in those of uninduced or early-time-of-induction samples. Results of this analysis cannot be known before the projected harvesting time, so for the first experiment, the process should be run with the time lines indicated above. The best combination of high expression (judged by band intensity on the gel) and high cell density should dictate optimal harvesting time—these conditions can then be applied to repeat experiments. It it also important to monitor growth closely during the induction phase because some strains rapidly lyse. In case of cell lysis, proceed to harvesting. Fermentation and Growth of E. coli for Optimal Protein Production
5.3.6 Current Protocols in Protein Science
Harvest the cells For small fermentors (1 to 10 liters): 19a. To harvest cells from small fermentors, pressurize the fermentor and transfer the reactor contents into an intermediate container. 20a. Distribute liquid culture, preferably under a biological hood, into centrifuge bottles. Record final culture volume in order to estimate final cell yield. Aerosols should be minimized at all times. Use closed containers fitted with sterile filter vents.
21a. Centrifuge 15 min at 5000 × g (4000 rpm in an H6000A rotor), 4°C. 22a. Collect pellets from each centrifuge bottle and place together in a heat-sealable plastic bag. Proceed to step 23. If the protein is secreted to the extracellular medium, collect the supernatant and concentrate by tangential flow ultrafiltration.
For large fermentors (>10 liters): 19b. For larger fermentors, prepare for cell harvest by removing all flexible tubing connections on the reactor, stopping pH and pO2 control loops, and setting the temperature set point at minimum. Set pressure set point to 0.8 bars and direct gas flow to the headspace of the reactor. In Figure 5.3.2, the broth is transferred from the fermentor to the bottom of a continuous feed centrifuge by pressurizing the reactor through the headspace. Consult the manufacturer’s instructions for specific operating conditions. It is important to cool the culture liquid quickly to minimize proteolytic degradation.
20b. Start the continuous feed centrifuge and accelerate to an effective centrifugation force of ≥10,000 × g. When bowl speed is stable, open connecting valves from the fermentor to the centrifuge. 21b. Set inlet flow rate to the centrifuge at ∼1 to 2 liters/min. Centrifuge at 4°C until all the medium has been processed. 22b. Stop centrifuge, dismount the bowl, and collect the pelleted cells into a heat-sealable plastic bag. If the purification process cannot handle the entire pellet volume, divide the material into smaller aliquots. If the protein is secreted to the extracellular medium, collect the supernatant and concentrate by tangential flow ultrafiltration.
Store the harvested cells and clean the fermentor 23. Retain a small amount of the pellet (50 to 100 mg) in a separate microcentrifuge tube for polyacrylamide gel analysis (step 18) to verify that centrifugation has not altered the target protein expression level. Close pellet-containing bags by heat sealing. Record total pellet weight and freeze at −20°C. Most recombinant proteins in the pelleted cells remain stable for months when stored at −20°C. However, some membrane receptors (e.g., neurokinin, adrenergic, transmembrane receptors in general) are best preserved by snap freezing in dry ice/isopropanol and storing at −70°C.
24. To clean the fermentor, fill it with water and adjust to pH 11.0 to 12.0 by adding 10 N NaOH. Set temperature to 50°C and leave with vigorous agitation overnight. Local biosafety rules may require sterilization of the reactor before cleaning. For sterili-
Production of Recombinant Proteins
5.3.7 Current Protocols in Protein Science
zation, fill the reactor with water and proceed to steps 7 and 8. If the fermentor is still not clean after this treatment, alternate alkali and acid (H3PO4) treatments.
25. Empty the reactor, rinse extensively with water, and sterilize as described in steps 7 and 8. Prepare the fermentor for another run, if desired. Electrochemical sensors (pH, pO2, pCO2) should be removed and stored according to manufacturer’s instructions. The fermentor should then be filled with water, sterilized, and stored sterile until the next run. ALTERNATE PROTOCOL 1
PROTEIN PRODUCTION IN HIGH-CELL-DENSITY PROCESSES: FED-BATCH METHODS Fed-batch fermentations take advantage of the high-protein-synthesizing capacity of an exponentially growing culture while minimizing the buildup of toxic by-products by keeping growth carbon source–limited. This protocol describes a simple glycerol-supplementation method that is applicable to a wide range of culture conditions. Cells are first grown in a batch fermentation in a medium containing 10 g/liter glycerol. After the initial glycerol is consumed, the culture is supplemented with an increasing feed rate of concentrated medium containing 900 g/liter glycerol. Except for cell density, induction of protein synthesis and harvest are the same as for batch fermentation (see Basic Protocol 1). Additional Materials (also see Basic Protocol 1) HCDF1 medium (see recipe) HCDM1 medium (see recipe), containing 10 g/liter glycerol O2 gas, pure 1. Prepare an inoculum culture of the transformed E. coli host strain expressing the protein of interest, then set up the fermentor as for batch fermentation (see Basic Protocol 1, steps 1 to 4). 2. Prepare HCDF1 feed medium in a feed tank (transfer vessel) and set on a balance or a load cell to keep track of feed consumption. 3. Proceed to preparing the fermentation medium, sterilizing the medium and fermentor, and conducting the final preparations for inoculation for batch fermentation procedure (see Basic Protocol 1, steps 5 to 14), substituting HCDM1 batch medium containing 10 g/liter glycerol as the fermentation medium. 4. Inoculate the fermentor and grow cells (see Basic Protocol 1, steps 15 and 16). After the initial glycerol is consumed, start the feed of HCDF1 medium. A sharp rise in pO2 is indicative of reduced oxygen consumption and thus of glycerol becoming limiting. Monitor growth during this phase to estimate maximum growth rate (see Sinclair and Cantero, 1990, for methodology).
5. Increase rate of addition of HCDF1 medium gradually and withdraw samples to follow the increase in OD600. Adjust rate of HCDF1 medium addition to the target feed rate. Monitor feed rates by measuring the mass of the feed tank over time. The target feed rate can be estimated as F = ì × Vf × OD/(Y × 0.4 × 0.9) Fermentation and Growth of E. coli for Optimal Protein Production
where F is the feed rate (ml/hr), ì is the target growth rate (hr−1), Vf is the fermentor volume (liters), OD is the optical density at 600 nm, Y is the yield of cell mass per unit carbon source (g/g), 0.4 represents the conversion factor from OD to cell mass (OD/g/liter), and 0.9 is the glycerol content in the feed solution (g/ml). An estimate of
5.3.8 Current Protocols in Protein Science
80% of the maximum growth rate (step 4) is a good starting value. An estimate of 0.3 is a good value for the yield, Y. When monitoring feed rates, make sure to take into account the density of the feed and the increase in fermentation volume with feeding. The target growth rate is dependent on each strain and each process. To avoid accumulation of toxic by-products (organic acids), the target growth rate should be set below 80% of the maximum growth rate observed during the batch phase.
6. Blend pure O2 gas into the air that is sparged into the reactor (either by solenoid valve pulses or by a motor-driven proportional valve) to make sure that pO2 is properly controlled (set point between 20% and 50%). 7. Once the target high biomass density is obtained (OD600 above 50), induce expression of the protein as in batch fermentation procedure (see Basic Protocol 1, step 17). Final cell densities can be as high as 100 to 150 OD units. Induction by a chemical inducer can also be achieved by simply adding the inducer to the feed solution (gradual induction).
8. Proceed to monitoring expression and harvesting (see Basic Protocol 1, steps 18 to 23) while continuing to match the rate of HCDF1 supplementation with growth. IN VIVO LABELING OF RECOMBINANT PROTEINS WITH HEAVY METAL DERIVATIVES
ALTERNATE PROTOCOL 2
To maximize incorporation of selenomethionine in place of methionine in recombinant proteins to facilitate structural studies, it is preferable to use a methionine auxotroph host strain. The protocol is best suited for small fermentors (1 to 10 liters) because the medium change step requires sterile harvesting via a batch pot centrifuge. Additional Materials (also see Basic Protocol 1) Methionine auxotroph host strain: E. coli DL41 or B834 (Table 5.2.1) Expression vector containing gene for the protein of interest DLM medium (see recipe) Complete rich medium (e.g., LB medium; UNIT 5.2) Additional reagents and materials for electroporation and testing expression of transformants (UNIT 5.2) Construct the expression strain 1. Transform methionine auxotroph host strain with expression vector containing the gene for the protein of interest (see UNIT 5.2). For T7 expression systems, use E. coli B834 (with or without pLys plasmid) as a host (also see UNIT 5.2 Commentary and see Table 5.2.1).
2. Test expression of transformants in a shaker flask induction test (see UNIT 5.2). Set up the fermentation and induce protein expression 3. Prepare DLM medium (same volume as fermentor) and keep at room temperature. 4. Prepare fermentor with complete rich medium and inoculate with the recombinant strain (see Basic Protocol 1, steps 1 to 16). 5. Grow overnight in rich medium at 30°C for pL strains, 37°C for others. Production of Recombinant Proteins
5.3.9 Current Protocols in Protein Science
6. Withdraw sample and measure OD600. Harvest cells under sterile conditions by batch pot centrifugation (see Basic Protocol 1, steps 19a to 22a). Stop all control loops on the fermentor. OD values should be in the range of 20 to 30 after overnight growth. Harvest only part of the culture. Calculate the volume to be harvested so that, once the cells are resuspended in the same volume of DLM labeling medium, the OD600 will be between 10 and 15.
7. Resuspend the pellet in DLM medium. Transfer to a sterile container and then back to the fermentor. Resume all control loops. In practice, there will be several pellets, distributed in several centrifuge bottles. Pellets are individually resuspended, then suspensions are pooled into a single vessel to avoid having to transfer each separate bottle back into the fermentor. Remember that sterility is a must here, and handling one container is less risky than dealing with several.
8. Wait for 30 min until all environmental parameters are back to their set points and are stable before inducing expression (see Basic Protocol 1, step 17). Proceed to monitoring expression and harvesting cells (see Basic Protocol 1, steps 18 to 23). ALTERNATE PROTOCOL 3
STABLE ISOTOPIC LABELING OF RECOMBINANT PROTEINS Uniform incorporation of stable isotopes in recombinant proteins is an alternative to selective labeling of a given amino acid (Alternate Protocol 2) to facilitate structural studies. The method requires the use of a semi-defined medium containing salts, trace elements, and yeast extract. Cells are supplied with [13C]glucose, 15NH4Cl, and/or D2O, and induced to synthesize recombinant protein labeled with 13C, 15N, and/or 2H. The three labels can be used individually or in combination. This protocol makes use of the fed-batch method (Alternate Protocol 1) but substitutes (labeled) glucose for glycerol as the carbon source and is run at much lower cell densities. Additional Materials (also see Basic Protocol 1) Transformed E. coli host strain expressing the protein of interest (UNIT 5.2) HCDM1 medium (see recipe) 50% (w/v) [13C]glucose 100 g/liter 15NH4Cl stock solution D2O Base for pH correction: 2 N NaOH (for 15N label) 1. Grow an inoculum culture of the transformed E. coli host strain expressing the protein of interest, then prepare fermentor and start the batch fermentation with HCDM1 medium (see Basic Protocol 1, steps 2 to 14), setting up 50% [13C] glucose as the nutrient solution (place feed tank on a balance or load cell). In this protocol, the carbon source is labeled glucose instead of glycerol. For 2H labeling, use D2O instead of H2O in making up HCDM1 medium.
2. After medium sterilization add 10 g/l [13C]glucose and the first 5-ml portion of 100 g/liter 15NH4Cl stock solution per liter fermentation volume (using a sterile syringe; see Basic Protocol 1, step 12).
Fermentation and Growth of E. coli for Optimal Protein Production
Glucose is added after sterilization to avoid partial destruction when heat sterilized with nitrogen-containing compounds present in the medium. To achieve the target 10 g/liter concentration, add 40 ml of the feed solution per liter of culture.
5.3.10 Current Protocols in Protein Science
15
NH4Cl is added in three separate batches to avoid inhibition by growth by excess NH4Cl in the medium.
3. Inoculate fermentor (see Basic Protocol 1, step 15). Allow cells to grow. When the initial [13C]glucose is consumed, start feed with [13C]glucose. A sharp rise in pO2 is indicative of glucose becoming limiting and thereby reducing oxygen consumption. To set initial and subsequent feed rates, use the equation in Alternate Protocol 1, step 5, substituting 0.5 for 0.9 as the glucose feed content (g/ml), using a target growth rate of 0.3/hr and an estimate of 0.4 for the yield Y.
4. Monitor growth by withdrawing samples periodically and measuring OD600. At an OD600 ∼5, add second batch of 15NH4Cl and induce protein expression (see Basic Protocol 1, step 17). In this protocol a high cell density is not desirable, and therefore no pure O2 should be necessary for pO2 control.
5. At 1 hr postinduction add the last batch of 15NH4Cl. 6. Proceed to monitoring protein expression and harvesting (see Basic Protocol 1, steps 18 to 23) while continuing to match the rate of glucose supplementation with growth. GROWTH MONITORING Bacterial growth can be monitored off-line by several different methods (Lech and Brent, 1994a). For fermentation process control, spectrophotometry is fast, reliable, and simple. Microscope counting is an alternative but is more cumbersome and time-consuming. On-line sensors also exist and have been used successfully in some cases (Schügerl et al., 1993). Most sensors, however, are sensitive to particulate matter, gas bubbles, fouling, and calibration drifts.
SUPPORT PROTOCOL 1
Materials Test sample (e.g., E. coli inoculum or fermentation medium) Culture medium (optional, for diluting visibly turbid cultures) Spectrophotometry cuvettes (e.g., 1-cm light path) Fixed-wavelength spectrophotometer (e.g., 600 or 660 nm) Count slide or hemacytometer Phase-contrast microscope (400×) For spectrophotometry: 1a. Place test sample in spectrophotometry cuvette. Fill reference cuvette with water. 2a. Transfer cuvettes to a fixed-wavelength spectrophotometer and measure the optical density at 600 nm. The level of absorbance at 600 nm will depend on the distance between the cuvette and the detector and will vary among instruments, often by a factor of 2. It is thus wise to calibrate each instrument by recording the OD600 of a culture that contains a known number of cells determined by some other method, such as observation on a count slide or titering for viable colonies (Lech and Brent, 1994b). For a culture grown in rich medium, a good rule of thumb is that each 1 OD unit is roughly equivalent to 109 cells/ml. OD readings above 0.4 cannot be accurately correlated with cell numbers and the sample should be diluted 10- or 100-fold with PBS to bring reading back into the 0.4 range. Production of Recombinant Proteins
5.3.11 Current Protocols in Protein Science
For cell counting: 1b. Take a clean count slide (or hemacytometer) and cover it with a clean coverslip. Dip a 0.1- or 1-ml pipet into the test sample, allow a small drop of liquid to form on the end of the pipet, and touch it lightly to the surface of the slide at the periphery of the coverslip. 2b. Put the slide on the stage of a phase-contrast microscope set to 100×, and focus on the cells. Calculate cell densities. Each cell in a small square of a count slide is equivalent to 2 × 107 cells/ml. SUPPORT PROTOCOL 2
STERILITY CHECKING Although results are known only after an overnight incubation, checking the sterility of the fermentation medium prior to inoculation is essential to ensure reproducible results and aids in troubleshooting potential problems. To absolutely eliminate contamination during the test itself, it is preferable to perform this protocol under a microbiological hood. A quick check of sterility requires a sample size of 100 µl, whereas more sensitive detection requires up to 100 ml. Materials Test sample (e.g., fermentation medium) LB plates with and without antibiotic (UNIT 5.2) 0.45-µm membrane (Millipore, type HA or HC) Sterile filter holder For a quick check: Spread 100 µl test sample onto one LB plate and one LB/antibiotic plate. Incubate at 30°C overnight. Inspect for appearance of any colonies. For a more sensitive detection: Filter up to 100 ml test sample by vacuum aspiration through a 0.45-µm membrane set in a sterile filter holder. Disassemble filter holder and transfer membrane to an LB agar plate, face up. Incubate plate 24 to 48 hr at 30°C. Inspect membrane for appearance of any colonies. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Fermentation and Growth of E. coli for Optimal Protein Production
DLM medium 2.5 g adenine⋅HCl 2.2 g L-glycine 1.3 g guanosine 1 g L-alanine 1 g L-arginine 0.8 g L-aspartic acid 0.06 g L-cystine 1.3 g L-glutamic acid 0.66 g L-glutamine 0.12 g L-histidine 0.5 g L-isoleucine 0.5 g L-leucine 0.7 g L-lysine 0.2 g L-methionine continued
5.3.12 Current Protocols in Protein Science
0.26 g L-phenylalanine 0.2 g L-proline 0.2 g seleno-L-methionine 4.2 g L-serine 0.46 g L-threonine 0.34 g L-tyrosine 0.46 g L-valine 0.34 g thymine 1 g uracil 20 g glucose 0.75 g NH4Cl 1.2 g K2HPO4 0.4 g MgCl2⋅6H2O 10 ml trace element solution 1 (see recipe) H2O to 1 liter CAUTION: Seleno-L-methionine, which replaces methionine, is highly toxic. Dissolve each ingredient one after the other in water, except for L-aspartate, L-glutamate, L-tyrosine, guanosine, thymine, and uracil, which need to be dissolved first in a small amount of 0.1 N NaOH. Sterilize by filtration through a membrane with 0.22-µm pores. Store at room temperature. ECPM1 medium 4 g K2HPO4 1 g KH2PO4 1 g NH4Cl 2.4 g K2SO4 132 mg CaCl2⋅2H2O 10 ml trace element solution 1 (see recipe) 20 g Casamino Acids (enzymatic hydrolysate; Difco) 3 g yeast extract (Difco) 40 g glycerol H2O to 800 ml Mix ingredients, pour into fermentor, and sterilize by heat (see Basic Protocol 1, steps 5 to 8) adding balance of water to 1 liter in step 6. Add 2 ml of 1 M MgCl2⋅6H2O (sterilized by filtration through a membrane with 0.22-µm pores) as directed in step 12 of Basic Protocol 1. Once sterilized, the medium can be stored up to 1 week at 4°C. HCDF1 medium 900 g glycerol H2O to 910 ml Sterilize by heat Add 20 ml 1 M MgCl2⋅6H2O (sterilized by filtration through 0.22-µm membrane) Add 70 ml trace element solution 2 (see recipe)
Production of Recombinant Proteins
5.3.13 Current Protocols in Protein Science
Supplement 6
HCDM1 medium 10 g K2HPO4 10 g KH2PO4 9 g Na2HPO4⋅2H2O 1.1 g NH4Cl 9 g K2SO4 10 ml trace element solution 2 (see recipe) 2 g yeast extract H2O to 800 ml Sterilize by heat (see Basic Protocol 1, steps 5 to 8), adding balance of water to 1 liter in step 6. Add 2 ml 1 M MgCl2⋅6H2O (sterilized by filtration through 0.22-µm membrane; see Basic Protocol 1, step 12). Supplement with carbon source (glycerol or glucose) as indicated in Alternate Protocol 1 and Alternate Protocol 3. Trace element solution 1 5 g EDTA 0.5 g FeCl3⋅6H2O 0.05 g ZnO 0.01 g CuCl2⋅2H2O 0.01 g Co(NO3)2⋅6H2O 0.01 g (NH4)6Mo7O24⋅4H2O Dissolve EDTA in 700 ml water. Dissolve each element separately by adding just enough 10 N HCl to bring it into solution. Add each element in solution one after the other to the EDTA solution. Add water to 1 liter. Adjust pH to 7.0 with concentrated NaOH. Autoclave. Store in the dark at 4°C. Trace element solution 2 5 g EDTA 6 g CaCl2⋅2H2O 6 g FeSO4⋅7H2O 1.15 g MnCl2⋅4H2O 0.8 g CoCl2⋅6H2O 0.7 g ZnSO4⋅7H2O 0.3 g CuCl2⋅2H2O 0.02 g H3BO3 0.25 g (NH4)6Mo7O24⋅4H2O Dissolve EDTA in 700 ml water. Dissolve remaining ingredients separately in just enough 10 N HCl to achieve solution and add one after the other to the EDTA solution. Add water to 1 liter. Adjust to pH 7.0 with concentrated NaOH. Autoclave. Store in the dark at 4°C.
COMMENTARY Background Information
Fermentation and Growth of E. coli for Optimal Protein Production
The choice of a strategy for fermentation is based primarily on the type of expression system (see UNIT 5.1). For example, chemical induction (e.g., via trp, lac, or tac promoters) may be more feasible than temperature induction (e.g., via the lambda promoter) in larger-volume fermentors. The strategy also depends on the final intended use for the protein: immunization and generation of antibodies, structure-function in-
vestigations, structural studies, or in vivo assays. Because the fermentation process (i.e., the media, temperature, pH, and pO2 used) can affect the solubility and localization of the protein, it should be designed with the purification process in mind (Chapter 6). Localization of the protein in the E. coli cell (cytoplasmic, periplasmic, or secreted in the extracellular medium) is also a prime factor in
5.3.14 Supplement 6
Current Protocols in Protein Science
the design of the process. Finally and most importantly, the toxicity of the target protein for E. coli is a key determinant in the success of the production. For example, cell lysis has been observed repeatedly when over-expressing proteins of the apolipoprotein family. Fermentation of recombinant E. coli has been developed by many different research groups, and several well-documented processes have been published (Knorre et al., 1991; Strandberg et al., 1991; Yang, 1992; Gram et al., 1994). In general, accurate control of environmental parameters (e.g., temperature, pH, and pO2) is prerequisite for any production. This will also ensure reproducibility among different batches. Table 5.3.1 summarizes vendors of fermentation equipment that we have used successfully. The list is not meant to be exhaustive, and local supply may vary. Although most recombinant protein production protocols described in the literature are based on a simple batch type of operation (see Basic Protocol 1), fed-batch methods (see Alternate Protocol 1) allow a more productive use of a given reactor by allowing growth of the culture to a high cell density prior to induction. The principle is to supply increasing amounts of nutrients so as to match exponential growth of the culture while minimizing accumulation of toxic by-products. Growth is made carbon source–limited in order to keep the concentration of residual carbon source low and to maximize yield of recombinant proteins. Fermentations may also be designed to produce labeled products. The replacement of methionine residues by selenomethionine (SeMet) in proteins (see Alternate Protocol 2) was first suggested as a new strategy to tackle a major hurdle in protein X-ray crystallography, namely, determination of the phases (Yang et al., 1990). The approach consists of using crystals of the SeMet-substituted protein in multiwavelength anomalous diffraction (MAD) experiments, which require access to a synchrotron. The SeMet protein can also be used, however, in fixed-wavelength analysis to provide landmarks (i.e., the heavy atoms of selenium) that facilitate data analysis and resolution of the structure. Nuclear magnetic resonance (NMR) spectroscopy is used increasingly for determining the three-dimensional structure of proteins (up to 40 kDa) in solution. In conventional NMR techniques, however, proteins, because of their large size, generate spectra with overlapping signals, in particular in the region of 1H resonances. This led several teams to develop a
general method for labeling proteins with different stable isotopes such as 2H, 13C, and/or 15N (see Alternate Protocol 3). Isotopic labeling allows NMR specialists to study heteronuclear coupling and to use powerful two-, three-, and four-dimensional editing of spectra. These techniques greatly facilitate assignment of resonances (for a review, see Clore and Gronenborn, 1994).
Critical Parameters and Troubleshooting Equipment. For optimal operation, several fermentor characteristics are essential. Heat transfer is best achieved through a double jacket around a stainless steel base. Water flow should be guided in loops around the vessel wall to avoid any dead zones. Steam, if available, is the most efficient means of heating. Heating efficiency is critical for temperature shift inductions, where the broth should be heated from 30° to 42°C in a few minutes. In fact, for fermentors with a working volume >10 liters, there should be provision for direct steam injection to obtain a very quick temperature shift. Cooling requires consumption of cold water, which, in some instances can be recycled. Recycling is most useful for installations where supercold water (−4° to 4°C) is used. In addition to the essential sensors (i.e., temperature, agitation, dissolved O2, pH, and foam level), sensors such as gas flow, pressure, load, redox potential, culture fluorescence, and mass spectrometer on exhaust gas (Todd, 1989) can be mounted to monitor the environment. Halling (1990) gives detailed information on basic principles and calibration of the essential monitors and Dusseljee and Feijen (1990) review sensors and process control information. Dissolved oxygen and O2 transfer can be controlled by agitation, air flow, gas composition, and headspace pressure. In practice, using high-agitation-rate motors with Rushton turbines and sparging O2-enriched air allow maximal transfers, but oxygen transfer can still become limiting at high cell densities. The average specific oxygen uptake rate for E. coli strains growing at a rate of 0.3 hr−1 in ECPM1 medium at 30°C is around 0.4 g O2/g cells/hr. Sterilization is a critical operation that can be hazardous if not properly handled. It should always be carried out according to the instructions of the fermentor supplier. An automatic controller is helpful and considerably limits the risks; it also ensures reproducibility of the operation from one batch to another. Sterility is, of course, a prerequisite for any fermentation
Production of Recombinant Proteins
5.3.15 Current Protocols in Protein Science
Fermentation and Growth of E. coli for Optimal Protein Production
experiment. Fermentor designs should be checked carefully to ensure that all parts in contact with the culture are properly sterilized by heat. The bottom region of the reactor vessel where the agitator shaft enters the fermentor is critical. Double mechanical seals with lubrication by steam condensates provide an efficient sterility barrier. Magnetically coupled agitation systems are even safer because all seals are static. Computer control is not absolutely required for most routine purposes. It is helpful, however, for fed-batch processes and for those requiring model-based feedback control from off-line analysis. It also allows extensive data logging and analysis both from the fermentor itself and from peripheral instrumentation (e.g., off-gas analysis, flow injection analyzers). High-cell-density methods. Fed -b atch methods can be classified into two types: (i) growth-linked feeding using process parameters such as pH (Bauer and White, 1976) or pO2 (Mori et al., 1979) to control the feed of nutrients, and (ii) model-dependent feeding, controlled by a mathematical model of growth (Yee and Blanch, 1992). The first class requires little modification to a batch operation, whereas the second class necessitates on-line computer control. In the latter case, one needs to estimate growth from off-gas measurement (Reiling et al., 1985) or by real-time measurement of growth by spectrophotometry. The feed rate can be adjusted via computer control to match the increase in growth monitored by OD600. As high cell densities are obtained, limitations eventually arise from broth viscosity, oxygen transfer, or the cooling capacity of the fermentor. Figure 5.3.3 illustrates a typical fed-batch process run according to Alternate Protocol 1, with a recombinant E. coli strain and without induction of expression. Isotopic labeling. As an alternative to the defined medium described in Alternate Protocol 3, a commercial medium (Celtone) is available from Martek with labeled amino acids derived from unicellular algae. Celtone gives higher cell densities and is inexpensive and easy to make. To use this medium, follow Basic Protocol 1 for growth and induction, use 5 N NaOH for pH control when labeling with 15N, and add another 1× concentration of Celtone medium (100 ml of 10× stock solution per liter fermentation volume) at the start of induction. Harvesting. Harvesting can be achieved by tangential flow microfiltration as an alternative to continuous centrifugation (Bailey et al., 1990). Tangential flow microfiltration allows the operation to be more contained, but requires a
final centrifugation to obtain a cell cake. However, it can also be coupled to cell wash, diafiltration, buffer exchange, and cell breakage by continuous-flow high-pressure homogenizers. The most common problem found in scaling up from shaker flask experiments is the decrease in expression level. In this case, one can vary the time of induction, check plasmid stability (UNIT 5.2), and test other host/vector combinations. Changing the composition of the medium may also solve the problem but is lengthy and labor-intensive. The absence of cell lysis should be confirmed by frequent microscopic observations, and cells must be quickly harvested as soon as lysis begins. Sterility check. Because the results of this test are not known for 12 to 16 hr, the procedure is initiated and continued. In case of a contamination revealed after inoculation, the batch should be considered suspect and/or discarded. A massive contamination (which could occur if the fermentor has been prepared more than 24 hr in advance) will be obvious from the evolution of the pH and/or pO2 and turbidity of the sample. If any contamination is detected before inoculation, the entire run should be cancelled and the fermentor cleaned (see Basic Protocol 1, step 24).
Anticipated Results With an average level of expression (e.g., 10% to 15% of total cellular proteins), a normal batch process will yield ∼100 g wet cell pellet per liter, containing ∼500 mg of the recombinant protein. This shows that even small-scale fermentations (1 to 10 liters) are quite effective in providing the starting material for purification developments. For actual production, a fed-batch process is more typically used—this maximizes fermentor output, but also requires more manpower to operate. In this case, overall yields of 1.5 to 2 g recombinant protein per liter culture, in the crude extract, are expected.
Time Considerations Preparation of a fermentor and its ancillaries can be done in 4 to 5 hr. Preparation of the inoculum can be started after that, allowing an overnight test for reactor sterility. The following morning inoculation can proceed, and the fermentation should be completed within 24 to 36 hr. Harvesting takes 1 to 2 hr. Analysis requires an additional 3 hr. While running the analysis, the fermentor can be cleaned, sterilized with water, and made ready for another run. Overall, a complete cycle takes 2.5 to 3 days.
5.3.16 Current Protocols in Protein Science
OD600
1000
100 90 80 70 60 50 40 30 20 10 0
100
10
1
B
• Acetate (g/liter)
A
90 Feed rate (g/liter/hr)
80 70 60 50 40 30 20 10 0 Cumulated CO2 production (mM/liter)
0
C
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Figure 5.3.3 High-cell-density growth. (A) Kinetics of growth (OD600) and acetate production. (B) Feed rate profile. Note that the feed rate was adjusted according to growth rate as described in Alternate Protocol 1. (C) Kinetics of cumulated CO2 production. Mass spectrometry of the exhaust gas was performed as described by Todd (1989). Acetate formation was measured as described by Reiling et al. (1985).
Production of Recombinant Proteins
5.3.17 Current Protocols in Protein Science
Bailey, F.J., Warf, R.T., and Maigetter, R.Z. 1990. Harvesting recombinant microbial cells using crossflow filtration. Enzyme Microb. Technol. 12:647-652.
Reiling, H.E., Laurila, H., and Fiechter, A. 1985. Mass culture of Escherichia coli: Medium development for low and high density cultivation of Escherichia coli B/r in minimal and complex media. J. Biotech. 2:191-206.
Bauer, S. and White, M.D. 1976. Pilot scale exponential growth of Escherichia coli to high cell concentration with temperature variation. Biotech. Bioeng. 18:839-846.
Sinclair, C.G. and Cantero, D. 1990. Fermentation modelling. In Fermentation, A Practical Approach. (B. McNeil and L.M. Harvey, eds) pp. 65-85. IRL press, Oxford.
Clore, G.M. and Gronenborn, A.M. 1994. Multidimensional heteronuclear nuclear magnetic resonance of proteins. Methods Enzymol. 239:349363.
Schügerl, K., Brandes, L., Wu, X., Bode, J., Ree, J.I., Brandt, J., and Hitzmann, B. 1993. Monitoring and control of recombinant protein production. Anal. Chim. Acta 279:3-16.
Dusseljee, P.J.B. and Feijen J. 1990. Instrumentation and control. In Fermentation, A Practical Approach. (B. McNeil and L.M. Harvey, eds.) pp. 149-171. IRL press, Oxford.
Strandberg, L., Koehler, K., and Enfors, S.O. 1991. Large scale fermentation and purification of a recombinant protein from Escherichia coli. Process Biochem. 26:225-234.
Gram, H., Ramage, P., Memmert, K., Gamse, R., and Kocher, H.P. 1994. A novel approach for high level production of a recombinant human parathyroid hormone fragment in Escherichia coli. Bio/Technology 12:1017-1023.
Todd, D. 1989. Mass spectrometry of fermentation emissions. BioTechnology 7:1182-1183.
Halling, P.J. 1990. pH, dissolved oxygen and related sensors. In Fermentation, A Practical Approach. (B. McNeil and L.M. Harvey, eds.) pp. 131-147. IRL press, Oxford.
Yang, W., Hendrickson, W.A., Kalman, E.T., and Crouch, R.J. 1990. Expression, purification and characterization of natural and selenomethionyl recombinant ribonuclease H from Escherichia coli. J. Biol. Chem. 265:13553-13559.
Literature Cited
Knorre, W.A., Deckwer, W.D., Korz D., Pohl, H.D., Riesenberg, D., and Ross, A. 1991. High cell density fermentation of recombinant Escherichia coli with computer controlled optimal growth rate. Ann. N.Y. Acad. Sci. 646:300-306. Lech, K. and Brent, R. 1994a. Growth in liquid media. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 1.2.1-1.2.2. John Wiley & Sons, New York. Lech, K. and Brent, R. 1994b. Growth on solid media. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 1.3.1-1.3.5. John Wiley & Sons, New York. Mori, H., Yano, T., Kobayashi, T., and Shimizu, S. 1979. High density cultivation of biomass in fed-batch system with DO-stat. J. Chem. Eng. 12:313-319.
Yang, X.M. 1992. Optimization of a cultivation process for recombinant protein production by Escherichia coli. J. Biotechnol. 23:271-289.
Yee, L. and Blanch, H.W. 1992. Recombinant protein expression in high cell density fed-batch cultures of Escherichia coli. Bio/Technology 10:1550-1556.
Key Reference Fermentation, A Practical Approach. 1990. (B. McNeil and L.M. Harvey, eds.) IRL press, Oxford. Covers the complete fermentation process cycle, illustrating many diagrams for equipment and giving essential background theoretical information.
Contributed by Alain Bernard and Mark Payton Glaxo Institute for Molecular Biology Geneva, Switzerland
Fermentation and Growth of E. coli for Optimal Protein Production
5.3.18 Current Protocols in Protein Science
Overview of the Baculovirus Expression System Baculoviruses have emerged as a popular system for overproducing recombinant proteins in eukaryotic cells (Luckow and Summers, 1988; Miller, 1988; Miller et al., 1986; Luckow, 1991). Several factors have contributed to this popularity. First, unlike bacterial expression systems (UNITS 5.1-5.3), the baculovirus-based system is a eukaryotic expression system and thus uses many of the protein modification, processing, and transport systems present in higher eukaryotic cells. In addition, the baculovirus expression system uses a helperindependent virus that can be propagated to high titers in insect cells adapted for growth in suspension cultures, making it possible to obtain large amounts of recombinant protein with relative ease. The majority of this overproduced protein remains soluble in insect cells, in contrast to the insoluble proteins often obtained from bacteria. Furthermore, the viral genome is large (130 kbp) and thus can accommodate large segments of foreign DNA. Finally, baculoviruses are noninfectious to vertebrates, and their promoters have been shown to be inactive in mammalian cells (Carbonell et al., 1985), which gives them a possible advantage over other systems when expressing oncogenes or potentially toxic proteins. This unit describes vectors and protocols for using the baculovirus expression system. A comprehensive guide outlining most of the protocols described here (as well as others) can be found in O’Reilly et al. (1992). Procedures related to large-scale production of recombinant proteins using the baculovirus expression system are covered in UNIT 5.5.
BACULOVIRUS LIFE CYCLE Currently, the most widely used baculovirus expression system utilizes a lytic virus known as Autographa californica nuclear polyhedrosis virus (AcMNPV; hereafter called baculovirus). This virus is the prototype of the family Baculoviridae. It is a large, enveloped, double-stranded DNA virus that infects arthropods. The baculovirus expression system takes advantage of some unique features of the viral life cycle (Fig. 5.4.1). See Doerfler and Bohm (1986) for a comprehensive review. As with mammalian DNA viruses, the baculovirus life cycle is divided temporally into immediate early, early, late, and very late
phases. Viruses enter the cell by adsorptive endocytosis and move to the nucleus where their DNA is released. DNA replication begins ∼6 hr after infection. Replication is followed by viral assembly in the nucleus of the infected cell. Two types of viral progeny are produced during the life cycle of the virus: extracellular virus particles (nonoccluded viruses) during the late phase and polyhedra-derived virus particles (occluded viruses) during the very late phase of infection. Extracellular virus is released from the cell by budding, beginning at ∼12 hr postinfection, and is produced at a logarithmic rate until 20 hr postinfection, after which production drops off. Polyhedra-derived virus, on the other hand, appears in the nucleus at ∼18 hr postinfection and continues to accumulate as late as 72 hr postinfection, or until the cells lyse. Occluded viral particles are embedded in proteinaceous viral occlusions called polyhedra within the nucleus of infected cells. The polyhedrin protein (29 kDa) is the major protein component of the occlusion bodies. The polyhedrin protein serves an important function for the survival and propagation of the virus in nature. Because baculoviruses are lytic, they quickly kill their insect host after infection. The polyhedrin protein serves to sequester, and thereby protect, hundreds of virus particles from proteolytic inactivation by the decomposing host tissue. The virus is transmitted when occlusion bodies are ingested by a new host as it feeds on a contaminated food source. The polyhedrin protein dissolves in the alkaline environment of the new host’s gut and the occluded virus is released. This virus infects the gut epithelial cells and virus replication takes place. Nonoccluded virus is then produced and budded from the infected gut cells. At this point, the virus spreads throughout the tissues of its new host. Although the polyhedrin protein is essential for survival of the virus in nature, it is dispensable for virus survival and propagation in tissue culture cells.
BACULOVIRUS EXPRESSION SYSTEM The baculovirus expression system takes advantage of several facts about polyhedrin protein: (1) that it is expressed at very high levels in infected cells, constituting more than
Contributed by Cheryl Isaac Murphy and Helen Piwnica-Worms Current Protocols in Protein Science (1995) 5.4.1-5.4.4 Copyright © 2000 by John Wiley & Sons, Inc.
UNIT 5.4
Production of Recombinant Proteins
5.4.1 CPPS
replication
viral occlusion
uncoating
fusion (midgut cells)
primary infection
budding
secondary infection of cells and tissues
solubilization in gut ingestion
secondary infection of cells and tissues
infection of host insect
Figure 5.4.1 Baculovirus life cycle. Viruses enter cells by adsorptive endocytosis and move to the nucleus where their DNA is released. Both DNA replication and viral assembly take place in the nuclei of infected cells to generate two types of viral progeny. These include extracellular (nonoccluded) virus particles and polyhedra-derived (occluded) virus particles. Extracellular virus is released from the cell by budding, starting at ∼12 hr postinfection and ending ∼36 hr postinfection. Polyhedra-derived virus appears later (∼18 hr postinfection) and accumulates in the nuclei of infected cells ≤72 hr postinfection or until cellular lysis. Polyhedra-derived virus is embedded in proteinaceous viral occlusions, the major protein component of which is the viral polyhedrin protein. Secondary infection of cells and tissues occurs by two pathways. In the first, the extracellular virus, once budded from the site of primary infection, is free to infect neighboring cells by the pathway just described. Alternatively, polyhedra-derived virus is released from occlusion bodies after an infected food source is ingested by a new host. Reproduced from Summers and Smith (1987) with permission from the Texas Agricultural Experiment Station.
Overview of the Baculovirus Expression System
half of the total cellular protein late in the infectious cycle; (2) that it is nonessential for infection or replication of the virus, meaning that the recombinant virus does not require any helper function; and (3) that viruses lacking the polyhedrin gene have a plaque morphology which is distinct from that of viruses containing the gene. Recombinant baculoviruses are generated by replacing the polyhedrin gene with a foreign gene through homologous recombination. In this system, the distinctive plaque morphology provides a simple visual screen for identifying the recombinants. To produce a recombinant virus that expresses the gene of interest, the gene is first
cloned into a transfer vector (described below). Most baculovirus transfer vectors contain the polyhedrin promoter followed by one or more restriction enzyme recognition sites for foreign gene insertion. Once cloned into the expression vector, the gene is flanked both 5′ and 3′ by viral-specific sequences. Next, the recombinant vector is transfected along with wild-type viral DNA into insect cells. In a homologous recombination event, the foreign gene is inserted into the viral genome and the polyhedrin gene is excised. Recombinant viruses lack the polyhedrin gene and in its place contain the inserted gene, whose expression is under the control of the polyhedrin promoter.
5.4.2 Current Protocols in Protein Science
Homologous recombination between circular wild-type DNA and the recombinant plasmid DNA occurs at a low frequency (typically 0.2% to 5%). However, linearization of wildtype baculovirus DNA before cotransfection with plasmid DNA increases the proportion of recombinant virus to ∼30% (Kitts et al., 1990). If the DNA is linearized such that an essential portion of the 1629 open reading frame (ORF) downstream from the polyhedrin gene is deleted, 85% to 99% of the viruses obtained by cotransfection with a plasmid vector that complements the deletion express the heterologous gene (Kitts and Possee, 1993). Two companies (Pharmingen and Clontech) market linear AcMNPV DNA containing such a deletion. Once a virus stock is obtained after cotransfection, it is necessary to purify recombinant virus by plaque assay so the recombinant virus can be identified. Limiting dilution can also be used (see O’Reilly et al., 1992). One of the beauties of this expression system is a visual screen allowing recombinant viruses to be distinguished. As mentioned above, the polyhedrin protein is produced at very high levels in the nuclei of infected cells at late times after viral infection. Accumulated polyhedrin protein forms occlusion bodies that also contain embedded virus particles. These occlusion bodies, up to 15 µm in size, are highly refractile— i.e., they have a bright, shiny appearance that is readily visualized under the light microscope. Cells infected with recombinant viruses lack occlusion bodies. Thus, when the virus is plaqued onto Sf 9 (Spodoptera frugiperda) cells, plaques can be screened for the presence (indicative of wild-type virus) or absence (indicative of recombinant virus) of occlusion bodies. Recombinant viruses can also be identified by DNA hybridization and polymerase chain reaction (PCR) amplification.
POST-TRANSLATIONAL MODIFICATION OF PROTEINS IN INSECT CELLS Because baculoviruses infect invertebrate cells, it is possible that the processing of proteins produced by them is different from the processing of proteins produced by vertebrate cells. Although this seems to be the case for some post-translational modifications, it is not the case for others. For example, two of the three post-translational modifications of the tyrosine protein kinase, pp60c-src, that occur in higher eukaryotic cells (myristylation of the N-terminal glycine residue and phosphorylation of serine 17) also take place in insect cells.
However, another modification of pp60c-src observed in vertebrate cells, phosphorylation of tyrosine 527, is almost undetectable in insect cells (Piwnica-Worms et al., 1990). In addition to myristylation, palmitylation has been shown to take place in insect cells. However, it has not been determined whether all or merely a subfraction of the total recombinant protein contains these modifications. Cleavage of signal sequences, removal of hormonal prosequences, and polyprotein cleavages have also been reported, although cleavage varies in its efficiency. Internal proteolytic cleavages at arginine- or lysine-rich sequences have been reported to be highly inefficient, and alpha-amidation, although it does not occur in cell culture, has been reported in larvae and pupae (Hellers et al., 1991). In most of these cases a cell- or species-specific protease may be necessary for cleavage. Protein targeting seems conserved between insect and vertebrate cells. Thus, proteins can be secreted and localized faithfully to either the nucleus, cytoplasm, or plasma membrane. Although much remains to be learned about the nature of protein glycosylation in insect cells, proteins that are N-glycosylated in vertebrate cells will also generally be glycosylated in insect cells. However, with few exceptions the N-linked oligosaccharides in insect cell-derived glycoproteins are only high-mannose type and are not processed to complex-type oligosaccharides containing fucose, galactose, and sialic acid. O-linked glycosylations have been even less well characterized in Sf 9 cells, but have been shown to occur. For further information on protein processing in insect cells, see Jarvis and Summers (1992) and O’Reilly et al. (1992).
LITERATURE CITED Carbonell, L.F., Klowden, M.J., and Miller, L.K. 1985. Baculovirus-mediated expression of bacterial genes in dipteran and mammalian cells. J. Virol. 56:153-160. Doerfler, W. and Bohm, P. 1986. The Molecular Biology of Baculoviruses. Springer-Verlag, New York. Hellers, M., Gunne, H., and Steiner, H. 1991. Expression and post-translational processing of preprocecropin-A using a baculovirus vector. Eur. J. Biochem. 199:435-439. Jarvis, D.L. and Summers, M.D. 1992. Baculovirus expression vectors. In Recombinant DNA Vaccines: Rationale and Strategies (R.E. Isaacson, ed.) pp. 265-291. Marcel Dekker, New York. Kitts, P.A. and Possee, R.D. 1993. A method for producing recombinant baculovirus expression vectors at high frequency. BioTechniques 14:810-817.
Production of Recombinant Proteins
5.4.3 Current Protocols in Protein Science
Kitts, P.A., Ayres, M.D., and Possee, R.D. 1990. Linearization of baculovirus DNA enhances the recovery of recombinant virus expression vectors. Nucl. Acids Res. 18:5667-5672. Luckow, V.A. 1991. Cloning and expression of heterologous genes in insect cells with baculovirus vectors. In Recombinant DNA Technology and Applications (A. Prokop, R.K. Bajpai, and C. Ho, eds.) pp. 97-152. McGraw-Hill, New York. Luckow, V.A. and Summers, M.D. 1988. Trends in the development of baculoviral expression vectors. Bio/Technology 6:47-55.
and its association with polyoma virus middle T antigen in insect cells. J. Virol. 64:61-68. Summers, M.D. and Smith, G.E. 1987. A manual of methods for baculovirus vectors and insect cell culture procedures. Texas Agricultural Experiment Station Bulletin No. 1555. College Station, Texas.
Key Reference O’Reilly et al. 1992. See above. A guide assembled to aid researchers using the baculoviral expression system, containing detailed protocols for using this system effectively.
Miller, L.K. 1988. Baculoviruses as gene expression vectors. Annu. Rev. Microbiol. 42:177-199. Miller, D.W., Safer, P., and Miller, L.K. 1986. An insect baculovirus host vector for high-level expression of foreign genes. In Genetic Engineering, Vol. 8 (J.K. Setlow and A. Hollaender, eds.) pp. 277-298. Plenum, New York. O’Reilly, D.R., Miller, L.K., and Luckow, V.A. 1992. Baculovirus Expression Vectors. W.H. Freeman and Company, New York.
Contributed by Cheryl Isaac Murphy Cambridge Biotech Corporation Worcester, Massachusetts Helen Piwnica-Worms Washington University School of Medicine St. Louis, Missouri
Piwnica-Worms, H., Williams, N.G., Cheng, S.H., and Roberts, T.M. 1990. Regulation of pp60c-src
Overview of the Baculovirus Expression System
5.4.4 Current Protocols in Protein Science
Protein Expression in the Baculovirus System
UNIT 5.5
Insect cell–recombinant baculovirus co-cultures offer a protein production system that complements microbial systems (UNITS 5.1-5.3) by providing recombinant proteins in soluble form and with most post-translational modifications. Moreover, the large size of the viral genome (130 kbp) enables cloning of large segments of DNA and consequent expression of complex protein aggregates. This unit describes methods associated with the large-scale production of recombinant proteins in the baculovirus expression system. UNIT 5.4 gives an overview of this system. Performing the protocols in this unit requires that the desired protein has already been cloned into an appropriate vector for expression in cultured insect cells under the control of a strong baculoviral promoter; it is also assumed that both infection of cells by the recombinant virus and production of recombinant proteins have been analyzed in preliminary experiments. Detailed protocols for the construction of a recombinant baculovirus, maintenance of insect cells in culture, and analysis in small-scale expression experiments can be found in several manuals (Murphy and Piwnica-Worms, 1994a,b; Summers and Smith, 1987). Production of proteins in large amounts requires correspondingly large quantities of starting materials, most importantly viral stocks of high titer. Basic Protocol 1 gives a method for large-scale production of viral stocks. Quality control is absolutely vital, and methods for titration of virus are provided (Support Protocols 1 and 2 describe the plaque assay and the end-point assay, respectively). Moreover, once viral stocks have been prepared and titered, the virus is tested in small-scale cultures to determine the kinetics of expression (Basic Protocol 2), which allows evaluation of various cell culture and infection conditions aimed at developing optimal levels of protein production (e.g., comparisons of different host cell lines, media, and environmental parameters). Support Protocol 4 provides instructions for preparing culture samples for protein analysis by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE; UNIT 10.1), and Support Protocol 5 discusses analytical methods for monitoring nutrient levels in cell culture fluids. Once optimal process parameters are identified, the target protein can be produced on a large scale in fermentors. Basic Protocol 3 describes regular batch production in bioreactors, whereas Alternate Protocol 1 covers a variant fed-batch procedure of production in perfusion cultures. Techniques for harvesting cultures from bioreactors are provided in Support Protocol 3. These protocols produce cell pellets or culture supernatants containing the protein of interest, which must then be subjected to purification and analysis as described elsewhere in this manual (Chapters 6 and 7). Despite being based on the use of a virus, the expression of recombinant protein with the baculovirus system is not hazardous to humans, as the virus cannot infect cells of noninsect origin. Nonetheless, handling of virus stocks should be restricted to well-defined equipment, materials, and rooms, kept separate from those where the host cell lines are maintained. This helps prevent any uncontrolled latent viral infection of the host cell line stocks. NOTE: All protocols (except Support Protocol 4 and Support Protocol 5) should be carried out using strict sterile technique, in the absence of antibiotics.
Production of Recombinant Proteins Contributed by Alain Bernard, Mark Payton, and Kathryn R. Radford Current Protocols in Protein Science (1995) 5.5.1-5.5.18 Copyright © 2000 by John Wiley & Sons, Inc.
5.5.1 CPPS
BASIC PROTOCOL 1
LARGE-SCALE PRODUCTION OF VIRAL STOCK The first phase of large-scale production of recombinant proteins is the generation of a large volume (1 to 5 liters) of highly infective recombinant virus. A culture of insect cells is prepared, grown to mid-exponential phase, and infected with recombinant virus. On day 4 or 5 postinfection, the culture is harvested by centrifugation and then stored. A small sample of the culture is reserved for titering by the plaque assay or by the end-point dilution method (see Support Protocols 1 and 2). High-titer stocks are used in the analysis of expression kinetics (see Basic Protocol 2) and in the infection of reactor cultures (see Basic Protocol 3 and Alternate Protocol 1). Materials Spodoptera frugiperda (Sf 9) cells Insect cell medium: TC100 (GIBCO/BRL) or IPL41 (GIBCO/BRL), with 5% to 10% (w/v) fetal calf serum (FCS); or Sf-900 II (GIBCO/BRL), serum free Pure recombinant virus (at 107 pfu/ml) 1-liter to 10-liter capacity shaker flasks or spinner culture flasks 27°C shaker incubators, humidity controlled Low-speed centrifuge (Sorvall RC3B or equivalent) 1-liter glass conical-bottom centrifuge bottles (Corning or Bellco), sterile Additional reagents and equipment for monitoring cell growth by cell counting and determining cell viability with trypan blue (see Support Protocol 6), monitoring nutrient levels in culture media (see Support Protocol 5), and titration of virus (see Support Protocols 1 and 2) 1. Prepare a 500-ml culture of Sf 9 cells in a shaker flask or spinner culture flask containing prewarmed insect cell medium (with or without serum) in a 27°C shaker incubator under controlled humidity. Always use a low liquid/total volume ratio (<0.4) to ensure good aeration of the culture while minimizing mechanical shear effects. Pluronic F-68 (BASF) should be included at 2 g/liter in media that lack a shear protectant. Set agitation to 50 to 100 rpm to ensure good mixing and aeration. To avoid uncontrolled infection, noninfected cultures should be handled separately and kept physically apart from infected cultures (use different incubators and different microbiological hoods). The Sf 9 cell line should be maintained and regularly passaged two to three times per week in either monolayer cultures or small shaker cultures in the same medium as the one chosen for the protocol. Maintainance of the cell line is described in Murphy and Piwnica-Worms (1994a).
2. Withdraw a sample of culture and perform appropriate tests to monitor cell growth, viability, and nutrient levels to ensure an adequate growth environment.
Protein Expression in the Baculovirus System
Growth should be monitored by cell counting using a hemocytometer and microscope. The culture should be initiated with a minimum of 2-3 × 105 cells/ml, at a viability higher than 98%. Healthy cells of high viability are characterized under the microscope by regular spherical shapes, smooth membranes, and refractile nuclei. Viable cells exclude trypan blue dye, whereas dead cells don’t (see Support Protocol 6). Elongated cell shapes or rough cytoplasmic membranes may be an indicator of oxygen limitation (this can be relieved by increased agitation or reduced culture volume/vessel volume ratios). Nutrient levels are measured using an assortment of specific assays (see Support Protocol 5). Typically, glucose and glutamine levels should not decrease below 0.2 g/liter and 0.2 mM, respectively.
3. When the culture reaches mid-exponential phase (∼ 1 to 5 × 106 cells/ml depending on the cell line and medium employed), infect with an aliquot of the pure recombinant
5.5.2 Current Protocols in Protein Science
virus to achieve a low multiplicity of infection (0.01 to 0.1). Transfer the spinner or shaker flask to a biological hood reserved for handling virus-infected cultures. Carefully open the flask and using a sterile pipet, add the appropriate volume of virus suspension to the culture. Mix the contents of the flask by gentle swirling, close the flask, and return it to the incubator. For a 500-ml culture at 1 × 106 cells/ml, 5 ml of a virus suspension at 10 appropriate.
7
pfu/ml is
The multiplicity of infection (MOI) is the ratio of infective virus particles per cell. The volume of virus solution (Vv, in ml) to add to a culture volume (Vc, in ml) is calculated as Vv = MOI × N × Vc/T, where N is the cell density in the culture (cells/ml) and T is the titer of the virus solution (pfu/ml).
4. Incubate culture 4 to 5 days in a 27°C shaker incubator under controlled humidity to allow virus production. Monitor cell growth once per day. At 4 to 5 days postinfection a large proportion of the cells (>50%) display hallmarks of infection: large cell bodies, condensed nuclei, rough membranes, and, for some cell lines, sausage shapes. At this stage, viability should be <20%.
5. Harvest the culture by centrifugation in sterile conical-bottom bottles for 30 min at 200 × g, 4°C, to remove cell debris. Retain the supernatant as the viral stock. Before discarding cell debris, inactivate all viruses in the pellet by soaking 30 min in 1 N NaOH.
6. Withdraw a small sample (e.g., 5 ml) for viral titration and store remaining stock at 4°C, protected from light, in 175-cm2 tissue culture flasks. Viral stocks can be stored in the dark for several months at 4°C without significant loss of infectivity. Low-titer viral stocks (<5 × 107 pfu/ml) may be more unstable, and titration of existing viral stocks should be performed immediately before use. Exposure to light has been shown to reduce virus infectivity during long-term storage (Jarvis et al., 1994). Two to three aliquots of 1 ml each can be dispensed in sterile cryovials under the biological hood (see UNIT 5.2, Support Protocol 3), tightly closed, and stored in liquid nitrogen as a back-up source for the virus. Freezing of the virus suspension, however, may lead to loss of virus titer. This loss, combined with the small volume that is frozen, would not eliminate the need to go back to step 1 of this protocol to generate a high-titer virus stock.
7. Estimate the titer of the reserved stock sample either by the plaque assay or by the end-point dilution method (see Support Protocols 1 and 2). Stocks of <107 pfu/ml are useless and should be discarded. 1-5 × 107 pfu/ml are considered to be low titer. The expected average titer is 108 pfu/ml.
DETERMINATION OF EXPRESSION KINETICS Once a viral stock is made with a titer above 5 × 10 pfu/ml, the kinetics and the level of protein expression are characterized in small-scale cultures in terms of the optimal cell culture and infection parameters. The following protocol can be used to compare host cell lines, media, virus constructs, and usual process parameters (e.g., cell density, pH, and agitation). It involves growing small-volume cultures of insect cells, infecting with virus stock, and periodically withdrawing culture samples that are prepared for protein analysis (see Support Protocol 4). The results are used to guide the timing of harvesting in large-scale production systems (see Basic Protocol 3 and Alternate Protocol 1). 7
BASIC PROTOCOL 2
Production of Recombinant Proteins
5.5.3 Current Protocols in Protein Science
Materials Cell line: Spodoptera frugiperda (Sf 9, Sf 21; ATCC, Pharminogen, Invitrogen) or Trichoplusia ni (Tn5 line; High Five cell line, Invitrogen) Insect cell medium: TC100 (GIBCO/BRL) or IPL41 (GIBCO/BRL), serum free or with 5% to 10% (w/v) fetal calf serum (FCS); or Ex-Cell 401 or Ex-Cell 405 (JRH Biosciences) or Sf-900 II (GIBCO/BRL), serum free Recombinant viral stock (see Basic Protocol 1) 1.5- to 3-liter shaker flasks or spinner culture flasks 27°C incubator, humidity controlled Additional reagents and equipment for monitoring cell growth by cell counting and determining cell viability with trypan blue (see Support Protocol 6), monitoring nutrient levels in culture media (see Support Protocol 5), and preparing samples for protein analysis (see Support Protocol 4) 1. Prepare a culture of cells in a 1.5- to 3-liter shaker flask or spinner culture flask containing 0.5 to 1 liter insect cell medium (with or without serum). Place flask in a 27°C incubator and allow cells to grow. Monitor cell growth, cell viability, and nutrient levels. Growth should be monitored by cell counting using a hemacytometer and microscope, and cell viability is determined by the trypan blue exclusion method (see Support Protocol 6). Nutrient levels are measured using an assortment of specific assays (see Support Protocol 5). In serum-containing media, the cell density should not exceed 1 × 106 cells/ml at time of infection. In serum-free media, infection can be started at up to 5 × 106 cells/ml. Prepare separate test cultures for each cell culture and infection variable to be analyzed (e.g., MOI, cell line, medium, and cell density).
2. Infect culture at high MOI (5 to 10) by sterile addition of an aliquot of recombinant viral stock. Return culture to 27°C incubator and incubate under controlled humidity. To calculate the amount of virus stock to add, use the calculations given in Basic Protocol 1, step 3. For example, for a 500-ml culture at 106 cells/ml, adding 5 ml of a virus stock (at 108 pfu/ml) would give an MOI of 1; 50 ml of the same stock would result in an MOI of 50. In some cases, it may be advantageous to centrifuge the cells and suspend them in fresh medium just prior to infection. This avoids possible nutrient limitation and toxic by-product inhibition during infection.
3. Withdraw 10-ml culture samples, measure nutrient levels (see Support Protocol 5), and prepare aliquots for protein analysis (see Support Protocol 4). Monitor nutrient levels and cell viability using trypan blue on a regular basis (twice per day) until 4 to 5 days postinfection. Once infected, cells are not necessarily completely permeable to trypan blue, and intermediate levels of staining can be observed. Also, some cell lines still exclude trypan blue quite effectively even when fully infected. An increase in cell size is usually a better indicator of infection.
4. Determine the optimal harvest time from the kinetics of target protein expression. SDS-PAGE (UNIT 10.1), immunoblotting, and biological assay, if available; may be used to analyze protein levels. For some proteins, it may be beneficial to harvest before the actual peak of expression because of progressive proteolytic degradation. Such degradation may be apparent as undersized immunoreactive bands in blots. Protein Expression in the Baculovirus System
Because the infection has been carried well beyond the optimal harvesting time, and unless the desired protein is expressed at very high levels and is very stable, the test cultures cannot be used further. They should be neutralized for 30 min in 1 N NaOH.
5.5.4 Current Protocols in Protein Science
PRODUCTION IN BIOREACTORS This protocol describes large-scale production of recombinant proteins in insect cell cultures conducted in bioreactors. Equipment preparation involves inspecting the reactor for cleanliness, calibrating sensors, and sterilizing. Culture medium is introduced through a sterilizing filter and then inoculated with exponentially growing, uninfected insect cells prepared in shaker or spinner flasks (see Basic Protocol 1, steps 1 and 2). Once cells reach half-maximal density, viral stock is introduced. The culture is terminated at the appropriate time, as determined in small-scale experiments (see Basic Protocol 2), and harvested (see Support Protocol 3). Because cultures are performed for days or weeks in the absence of antibiotics, maintaining the sterility of all operations is essential. This method may be adapted to perfusion cultures (see Alternate Protocol 1).
BASIC PROTOCOL 3
Materials Insect cell medium: TC100 (GIBCO/BRL) or IPL41 (GIBCO/BRL), serum free or with 5% to 10% (w/v) fetal calf serum (FCS); or Ex-Cell 401 (JRH Biosciences) or Sf-900 II (GIBCO/BRL), serum free Inoculum (15% to 20% bioreactor culture volume): spinner or shaker flask culture of uninfected insect cells and in full exponential growth phase Recombinant viral stock (see Basic Protocol 1) Bioreactor fitted with temperature, pH, agitation, air flow, and pO2 control (see Table 5.3.1 and Fig. 5.3.1) Millipak filter, sterile (Millipore) Viral stock and inoculum transfer vessel fitted with transfer line (see Fig. 5.3.2) Peristaltic pump (optional) Additional reagents and equipment for checking sterility and monitoring cell growth and determining cell viability with trypan blue (see Support Protocol 6) 1. Prepare bioreactor by inspecting for the absence of residual biological material on the reactor vessel walls, agitator shaft baffles, or other parts; calibrating sensors for temperature, pH, pO2 and pressure; and sterilizing accessory lines and transfer vessels by autoclaving. Fill reactor vessel with water such that all probe tips are immersed and sterilize. Empty reactor. If the bioreactor is autoclaved, it is important to ensure proper monitoring of the temperature inside the reactor (UNIT 5.3).
2. Connect medium addition line and add medium through a sterilizing filter. Disconnect medium addition line. The volume of the medium added is 75% to 80% of the final culture volume desired. This allows for cell inoculum and viral stock additions.
3. Set all parameters to the respective set points. Among strategies available to control dissolved oxygen levels in the culture (e.g., flow rate of gas, agitation, control through headspace), pure O2 pulsing through the sparger is the easiest method to monitor. A permanent airflow through the reactor headspace (at 1 liter air/liter culture/min) also damps possible oscillations around the set point. Generally, a set point of 30% to 50% air saturation should be optimal. If possible, leave the bioreactor in an uninoculated state for 24 to 48 hr. Check sterility to detect contaminants and to avoid loss of valuable inoculum.
4. Inoculate bioreactor with inoculum consisting of a spinner or shaker flask culture of uninfected insect cells at high viability and in full exponential growth phase so as to obtain a starting density of 2 to 3 × 105 cells/ml. Pool cultures if necessary and transfer
Production of Recombinant Proteins
5.5.5 Current Protocols in Protein Science
into a sterile inoculation transfer vessel under a biological hood. Connect transfer vessel as described in UNIT 5.3. Use a septum port that has not been previously used for emptying or filling the reactor. The growth characteristics of the inoculum should have exhibited predictable and reproducible growth rates during previous subculture.
5. Transfer the inoculum aseptically into the reactor either via a peristaltic pump or by pressurizing the inoculum vessel with sterile air. Foaming of the culture should be avoided at all times during the transfer.
6. Monitor cell growth until cell density reaches half-maximal levels, as estimated from inoculum growth curves. 7. Infect culture at high MOI (5 to 10) by aseptically adding recombinant viral stock as described for inoculum addition in step 5. Allow virus infection, virus replication and protein expression to proceed for 4 to 5 days, withdrawing samples twice daily and determining cell viability and nutrient levels (see Support Protocol 5). In addition, prepare aliquots of the sample twice daily for protein analysis (see Support Protocol 4). Terminate the process by harvesting at the optimal time point (see Basic Protocol 2 regarding determination of this time point). The multiplicity of infection (MOI) is the ratio of infective virus particles per cell. The volume of virus solution (Vv in ml) to add to a culture volume (Vc in ml) is calculated as Vv = MOI × N × Vc /T, where N is the cell density in the culture (cells/ml) and T is the titer of the virus solution (pfu/ml).
8. Harvest by centrifugation or tangential flow microfiltration (see Support Protocol 3). Depending on the target protein localization, keep the cell pellet or the supernatant fraction. Representative samples should be collected before and after any of the following downstream processes (see Support Protocol 3 and Support Protocol 4) to monitor changes in target protein stability or conformation. ALTERNATE PROTOCOL 1
Protein Expression in the Baculovirus System
PRODUCTION IN PERFUSION CULTURES Perfusion cultures permit high cell densities and production of greater protein amounts than simple batch methods (see Basic Protocol 3), particularly if proteins are designed for secretion. Equipment setup is as in Basic Protocol 3 except that a tank containing extra medium is set on a load cell and connected to the reactor, and a cell retention device is included along with a holding tank for spent medium. After the reactor is inoculated with insect cells, fresh medium is added to the culture at increasing rates to match the increase in cell density. Spent medium (free of cells) is withdrawn at the same rate to keep reactor volume constant. Because of the large holding tanks required, this protocol can be easily implemented at reactor scales of 1 to 5 liters but requires an industrial production environment for larger scales. Additional Materials (also see Basic Protocol 3) Cell retention device: spin filter, internal membrane filter, external membrane filter, or external centrifuge Fermentor level controller (contact probe) Medium feed and filtrate polypropylene holding tanks (10 to 15 times reactor volume; Nalgene) Load cell or electronic balance for feed tank (Mettler or equivalent) Peristaltic pumps (Watston-Marlow or equivalent) Additional reagents and equipment for determining cell viability with trypan blue
5.5.6 Current Protocols in Protein Science
(see Support Protocol 6), and preparing samples for protein analysis (see Support Protocol 4) 1. Prepare the bioreactor and start the culture (see Basic Protocol 3, steps 1 to 6). Be sure to equip the bioreactor with a cell retention device and a fermentor level controller. Set up the level controller so that it activates addition of fresh medium to the reactor by a peristaltic pump as culture volume decreases. The decrease in culture volume is caused by continuous withdrawal of cell-free filtrate by another pump. The speed of this second pump determines the perfusion rate (0.5 to 4 culture volumes/day). The equipment used here is available from most fermentor manufacturers and can be adapted to standard batch-oriented designs (see Table 5.3.1). For a thorough review of perfusion hardware, see Tokashiki and Takamatsu (1993).
2. Prepare a large volume of medium in a feed tank. Set tank on load cell or electronic balance. Feed and filtrate holding tanks should be of at least 10× the reactor capacity. This explains why it is more difficult to run large reactors (more than 5 to 6 liters) in perfusion in a common laboratory environment.
3. Connect feed tank to the reactor and connect filtrate holding tank to the filtrate port of the cell retention device. Set up peristaltic pumps for medium feed and filtrate removal. 4. Monitor cell growth, and, once cell density has reached half the maximal density for a batch operation, start perfusion with insect cell medium at a rate of 0.5 reactor volume/day. Monitor feed rates by recording changes in the weight of the feed tank per unit time (be sure to take into account the density of the feed medium).
5. Gradually increase perfusion rate in proportion to the increase in cell density. Allow 3 to 5 days of perfusion to reach the target cell density. The target cell density for infection is 1 to 3 × 107 cells/ml at more than 95% viability. Ideally, a perfusion process should be run without infection first to determine the maximal cell density achievable with a given set of hardware, cell line, and medium. However, most reports available in the literature seem to agree with a maximal cell density of 5-6 × 107 cells/ml.
6. Infect the culture by adding recombinant viral stock to the reactor (see Basic Protocol 3). Withdraw a 5-ml sample for analysis of protein expression. Halt perfusion for a short time after infection (1 to 2 hr) to allow virus attachment and/or penetration inside the cells. This sample will serve as a negative control in protein expression analysis (see Support Protocol 4). In this process, because the cell density is unusually high, it may be difficult to achieve a high MOI. However, it has been shown that infection at very low MOI (0.01) is still as productive as a batch operation with a high MOI. Perfusion is temporarily halted to prevent virus escape through the cell retention device. To avoid nutrient limitation during the arrest of perfusion, the feed rate can be increased 2- to 3-fold during the hour preceding the infection. Halting perfusion can simply be done by turning off the filtrate withdrawal pump. The rest of the cell retention hardware (filters, pump, centrifuge) should be left running without interruption to avoid any heterogeneity of the whole system.
Production of Recombinant Proteins
5.5.7 Current Protocols in Protein Science
7. Set perfusion rate to maximum (3 to 4 reactor volumes/day). 8. Let infection proceed for at least 4 days. Monitor cell viability and protein expression. 9. To avoid dilution of the product in the filtrate (and if the protein is secreted), remove the accumulated filtrate every 24 hr and concentrate by ultrafiltration (see Support Protocol 3). Prepare sample for analysis (see Support Protocol 4). If expression of the secreted protein is under control of a late promoter, the accumulated filtrate for the first 24 hr postinfection usually contains little product and can be discarded. SUPPORT PROTOCOL 1
TITRATING VIRAL STOCKS WITH THE PLAQUE ASSAY It is important to know the titer of a viral stock when preparing new stocks or when infecting insect cells to produce recombinant proteins. Two methods are used to determine viral titers in stock preparations, namely, the plaque assay and the end-point titration assay (see Support Protocol 2). Both methods are biological assays employing cell culture and quantify only the extracellular infectious forms of baculovirus. The plaque assay involves infecting exponentially growing Sf 9 cell monolayers with serial dilutions of viral stock (see Basic Protocol 1). After removing the viral supernatant following a period of viral adsorption, an agarose overlay is added and culture is continued for 5 to 7 days before plaques are counted in the culture treated with the highest dilution of virus that elicits an infection. Materials Sf 9 cells (30 ml, 8 × 105 cells/ml) TC100 medium (GIBCO/BRL), with and without 5% (w/v) fetal calf serum (FCS) Test sample: viral stock or culture supernatant 3% (w/v) low gelling/melting temperature agarose, autoclaved 2× TC100 medium/10% (w/v) FCS 0.3% (w/v) neutral red, diluted 1:20 in 1× phosphate-buffered saline (PBS, APPENDIX 2E; filter sterilize and store in the dark) 6-well cluster dishes, sterile Pipet tips, sterile Tubes, sterile Humid box 1. Seed three sterile 6-well cluster dishes by placing 1.5 ml of exponentially growing (8 × 105 cell/ml) Sf 9 cells in TC100 medium containing 5% FCS in each well. Place dishes in incubator at 27°C. 2. After monolayer formation (after 30 min to 1 hr incubation), wash cells with TC100 medium (no serum). Serum-free medium facilitates the adsorption of the virus to the cells.
3. Using sterile pipet tips and tubes, dilute test sample to 10−6, 10−7, and 10−8 in serum-free (TC100 medium). Add 300 µl of each dilution to each well of the host cell monolayers. Six repeats of one dilution per plate should be performed. Hold the pipet tip against the wall of the well while dispensing and carefully tilt the plate by hand to ensure an even distribution of the inoculum. Make six replicates of each dilution per plate. Protein Expression in the Baculovirus System
4. Return plates to incubator and let the inoculum virus adsorb 2 hr at 27°C. During incubation, prepare the agarose overlay by melting autoclaved 3% low gelling/melt-
5.5.8 Current Protocols in Protein Science
ing temperature agarose in a microwave, cooling to 37°C, and mixing 1:1 with 2× TC100 medium/10% FCS at 37°C. 5. Remove the virus by vacuum pipetting. Again, hold the pipet tip against the wall of the well, plate tilted.
6. Rapidly cover with 2 ml agarose overlay per well, and let the agarose harden for 3 to 4 hr at 27°C. Overlay with 1 ml TC100 medium/5% FCS. The agarose prevents diffusion of virus released from infected cells over the course of the assay. Thus, only cells immediately adjacent to the cell of primary infection are subsequently infected by progeny virus. When enough cells in the immediate vicinity of the primary infection are lysed, a plaque (or hole in the cell monolayer) results.
7. Incubate plates 5 to 7 days in a humid box in the 27°C incubator. 8. Stain plates for 2 hr with 0.3% neutral red. Invert the plate on absorbent paper to decant excess liquid and store 5 hr in the dark at room temperature to allow plaques to become visible. Count plaques and calculate average of all wells showing clear, separate plaques. Express titers as pfu/ml original test sample. Staining of cells is not essential but facilitates plaque observation. Uninfected cells stain red, and areas of cell lysis remain clear due to plaque formation.
TITRATING VIRAL STOCKS WITH THE END-POINT ASSAY Titrating virus by the end-point assay is similar to the plaque assay (see Support Protocol 1), but here a single virus can replicate and spread throughout the culture well and ultimately infect all cells present in the well, leading to the cytopathic effect. The titer is first given in TCID50 units/ml, which can be transformed to an approximate result in pfu/ml. Infection of cells is assayed by detecting either the recombinant protein or the cytopathic effect.
SUPPORT PROTOCOL 2
Materials Sf 9 cells (1 × 106 cells/ml), grown in Sf-900 II medium (GIBCO/BRL) Test sample: viral stock or culture supernatant Chromogenic substrate for detecting recombinant protein (optional, if available; e.g., Xgal for β-galactose) 96-well microtiter plate, sterile Multitip pipettor Pipet tips, sterile Humid box 1. Add 225 µl exponentially growing Sf 9 cells (1 × 106 cells/ml, in Sf-900 II medium) to every well of a sterile 96-well microtiter plate. Do not allow the plates to stand.
2. Immediately add 25-µl undiluted test sample of virus to the first well. Make three replicate dilutions of each sample.
3. Make 10-fold serial dilutions across the plate, directly into the insect cell suspensions, using a multitip pipettor and changing pipet tips between each dilution to prevent carryover of virus on the outside of the tip to the low dilutions. Alternatively, dilutions can be made in sterile 1.5-ml microcentrifuge tubes.
Production of Recombinant Proteins
5.5.9 Current Protocols in Protein Science
4. Transfer 10 µl of each of the 10−6 to 10−8 dilutions to a microtiter plate. Up to 60 replica wells per dilution can be made.
5. Incubate microtiter plates in a humid box 6 days at 27°C. It is important that no evaporation be allowed to occur due to the small volume in each well. Wrapping parafilm around the plate also avoids desiccation.
6. If available, add chromogenic substrate for detecting recombinant protein. Score wells such that evidence of viral infection is taken as a positive result. Evidence of viral infection may be simple in the case of virus coding for recombinant protein that can be easily detected (e.g., via the use of chromogenic substrates). Otherwise a positive well is scored on the basis of the presence of the cytopathic effect.
7. Count the number of positive (infected) and negative (uninfected) wells and calculate the theoretical dilution that would yield 50% positives, following the method described in Summers and Smith (1987). To convert TCID 50 into pfu, multiply by ln(2) (i.e., 0.69). SUPPORT PROTOCOL 3
HARVESTING Harvesting of cell cultures can be done by centrifugation or tangential flow microfiltration to separate cell bodies from culture medium. Centrifugation can be done in a batch mode by dividing the material into large centrifuge bottles and processing conventionally. Continuous feed centrifugation requires a continuous feed centrifuge and is effective for volumes of 5 to 20 liters. Larger volumes must be harvested by tangential flow microfiltration, which provides a retentate that must then be transferred to centrifuge bottles and subjected to batch centrifugation. Depending on the target protein localization, either the cell pellet or the supernatant fraction is kept for further purification. Protease inhibitors should be added in all cases and temperature should be kept at ∼4°C to limit possible degradation. In practice, however, it becomes expensive to add them before the material is concentrated to a small volume. Materials Protease inhibitor mix (see recipe) 1-liter conical-bottom centrifuge bottles (Bellco), sterile Low-speed centrifuge (Sorvall RC3B or equivalent), continuous feed centrifuge (Heareus Contifuge or equivalent), or tangential flow microfiltration system (Microgon KrosFloII or equivalent; 0.04 m2/liter culture) To harvest by batch centrifugation (≤5 liters) 1a. Transfer the reactor contents into 1-liter conical-bottom glass centrifugation bottles by pressurizing the headspace into the bottles. 2a. Centrifuge 30 min at 200 × g. Concentrate the cell-free supernatant by tangential flow ultrafiltration. To harvest by continuous feed centrifugation (5 to 20 liters): 1b. Connect the inlet line of the centrifuge to the reactor harvest valve, then connect the filtrate outlet line from the centrifuge to a harvesting tank.
Protein Expression in the Baculovirus System
2b. Start centrifuge at 200 × g, 250 ml/min. Check that supernatant is clear and free of cells. Concentrate the cell-free supernatant by tangential flow ultrafiltration.
5.5.10 Current Protocols in Protein Science
To harvest by tangential flow microfiltration (>20 liters) 1c. Connect the inlet of the filtration system to the reactor harvest valve. Connect the retentate line to a return port at the bottom of the reactor. It is important that the return port is immersed in the culture to avoid foaming. Alternatively, the reactor contents can be transferred to a harvesting tank, then filtered as described.
2c. Start the circulation pump with the filtrate line closed. Operate without filtration for 3 to 5 min, then open the filtrate line. When the reactor is almost empty, transfer the retentate to centrifuge bottles and centrifuge as described in step 2a. Concentrate the cell-free supernatant by tangential flow ultrafiltration. The circulation rate and transmembrane pressure should be set so that the average shear rate is <2500 sec−1, and transmembrane pressure should not exceed 500 mbar to obtain optimal filtration and avoid membrane clogging (Maiorella et al., 1991).
PREPARING SAMPLES FOR PROTEIN ANALYSIS The preparation of samples for protein analysis depends on the protein that is expressed. For a cell-associated protein, a simple lysis in a detergent-containing buffer usually releases most intracellular and membrane proteins. Alternatively, direct resuspension in the SDS sample buffer also solubilizes all proteins. For a secreted protein, dilute the culture supernatant (after concentration if necessary) with an equal volume of sample buffer. Once prepared by this protocol, samples can be loaded directly onto polyacrylamide gels (UNIT 10.1).
SUPPORT PROTOCOL 4
Materials Test sample SDS sample buffer (UNIT 10.1) or detergent lysis buffer (see recipe) Low-speed centrifuge 1. Pellet cells 5 min at 300 × g, 4°C. 2. Suspend in SDS sample buffer or lysis buffer (50 to 100 µl per 106 cells). If the cell pellet is resuspended in lysis buffer, an aliquot of the lysate should then be diluted 1:1 with sample buffer.
3. Store at −20°C or −80°C until analyzed. Samples can be loaded on SDS-PAGE gels (UNIT 10.1) and analyzed by direct detection (e.g., by Coomassie blue or silver staining; UNIT 10.5) of a band at the expected molecular weight with maximum intensity at optimal harvest time, or by immunodetection after blotting the gel on a membrane (UNIT 10.7). It is good practice to include negative controls (e.g., time 0 of infection, cultures infected with wild-type virus, or noninfected culture samples) as well as positive controls (e.g., partially purified recombinant or natural protein when available, or samples from other expression experiments).
Production of Recombinant Proteins
5.5.11 Current Protocols in Protein Science
SUPPORT PROTOCOL 5
MONITORING NUTRIENT LEVELS IN CULTURE MEDIA Expression of recombinant proteins by cultured cells depends directly on levels of nutrients supplied by the culture medium. This protocol describes analytical techniques for glucose, lactate, glutamine, and ammonia. Monitoring nutrient levels is not an absolute requirement for successful exploitation of the baculovirus expression technology. It is useful, however, for troubleshooting in cases of problems with cell growth, virus stock amplification, and protein production. It allows quality control checks of media and cultures and points to eventual nutrient limitations (glucose, glutamine, and other amino acids) or inhibitory product accumulation (lactate, ammonia). A more exhaustive compendium of biochemical analytical techniques for cell culture can be found in Doyle et al., 1994. The test sample of 5 to 10 ml is withdrawn from the culture (see Basic Protocols 1 and 2), the cells removed by brief centrifugation or filtration, and the culture fluid subjected to analysis. Glucose and lactate. Glucose can be analyzed in an automated enzyme assay (Yellow Spring Instruments). Glucose oxidase, immobilized on a membrane, oxidizes glucose in the sample to Glucono-δ-lactone, producing H2O2. A platinum electrode, held at an anodic potential, in turn oxidizes H2O2 to produce electrons. The electron flow is measured and is directly proportional to the H2O2 and therefore also proportional to the glucose concentration. The same machine can analyze lactate on the same sample. Another similar membrane electrode with immobilized L-lactate oxidase at its surface gives a parallel amperometric signal proportional to the lactate concentration in the sample. Glutamine. Glutamine is a critical nutrient for insect cell metabolism. Because glutamine degrades in liquid media, it is wise to check whether it becomes limiting for growth. Glutamine can be measured by high-performance liquid chromatography (HPLC) or by an enzyme assay in which it is first converted to glutamate and ammonia by glutaminase (Ozturk et al., 1989). Ammonia. Ammonia can be measured by an enzymatic assay kit (Sigma) or by an ion-specific electrode (Orion 9512BN). For analysis with the ammonia electrode, the solution to be analyzed is first made alkaline by the addition of an equal volume of 0.2 N NaOH. Ammonia gas is generated which diffuses through the semipermeable membrane of the electrode and leads to a voltage potential difference. The assay is linear from 0.2 to 12 mM ammonia (log scale; Ozturk et al., 1989).
SUPPORT PROTOCOL 6
DETERMINATION OF CELL GROWTH AND VIABILITY This protocol describes growth monitoring by cell counts and viability estimation by the trypan blue exclusion method. The culture sample is first stained with trypan blue, and an aliquot is applied to the chamber of a hemocytometer and examined under an inverted microscope. At least 200 cells should be counted to guarantee accuracy. Materials Fresh cell culture sample 0.2% (w/v) trypan blue solution (Sigma) Ethanol
Protein Expression in the Baculovirus System
Hemacytometer Inverted microscope (×10 objective, phase contrast) 1. Mix 100 µl of the fresh cell culture sample with 100 µl trypan blue solution.
5.5.12 Current Protocols in Protein Science
2. Clean the hemocytometer plate and coverslip with ethanol. Dry and fix the coverslip. 3. Fill the hemocytometer chamber with the stained sample. 4. Examine under the inverted microscope and count live (bright) and dead (blue) cells on at least two areas of 25 small squares. Average each count (live and dead) and multiply by 2 × 104 to obtain cell density/ml. If there are too many cells to count, dilute the original sample by 5- or 10-fold in PBS and start at step 1 again, taking dilution factor into account for the final result.
5. Calculate viability by the proportion of live cells in the total number (live + dead). Healthy, noninfected suspension cultures should give a viability above 95% at all times.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Detergent lysis buffer 1% (v/v) Nonidet P-40 (NP-40) 150 mM NaCl (APPENDIX 2E) 50 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) Protease inhibitor mix 1 mM phenylmethylsulfonyl fluoride (PMSF) 5 mM iodoacetamide 0.2 mM Nα-p-tosyl-L-lysine chloromethyl ketone (TLCK; prepared fresh) 2 mM benzamidine⋅HCl CAUTION: Prepare stock solutions in a chemical fume hood.
COMMENTARY Background Information The baculovirus expression system has become a popular technology to produce recombinant proteins. It is based on the infection of cultured insect cells by a recombinant virus vector in which foreign DNA is integrated under control of the strong viral promoter for polyhedrin protein (polh; UNIT 5.4). It is used for obtaining recombinant proteins for various purposes; such as biochemical studies, analysis of structure-function relationships, in vivo animal studies, and structural studies. The system recognizes vertebrate protein targeting signals and thus allows expression of different classes of proteins: cytoplasmic, nuclear, membrane-spanning, and secreted. The host insect cell performs most post-translational modifications, including phosphorylation, acylation, glycosylation, precursor processing, subunit assembly, secretion, and nuclear translocation. The glycosylation patterns reported in the recombinant proteins, however, have been restricted to the high-mannose type in almost all cases. The baculovirus expression system is also extensively used for the construc-
tion of complex biological aggregates (e.g., ion channels, viral capsids, and multisubunit enzymes). It is biologically safe. Instructions for constructing recombinant baculovirus and small-scale expression experiments are available in other manuals (Murphy and PiwnicaWorms, 1994a,b; Summers and Smith, 1987). The polh promoter is the most widely used promoter to date and many of the available transfer vectors use it. However, p10, another very late, very strong promoter is also being used (McCutchen et al., 1991; Stewart et al, 1991; Belyaev and Roy, 1993; Wang et al., 1993; Bonning et al., 1994; Deramoudt et al., 1994; van-Lier et al., 1994). In addition, the use of earlier promoters (e.g., basic protein promoter, also called pCor) is also exploited. With this promoter, expression of the recombinant protein starts at about 20 hr postinfection and peaks at 30 to 36 hr postinfection (Sridhar and Hasnain, 1993; Sridhar et al., 1993; Rankl et al., 1994; Chazenbalk and Rapoport, 1995). At these times, the host functions involved in protein processing are much less affected by viral infection.
Production of Recombinant Proteins
5.5.13 Current Protocols in Protein Science
Before large-scale production of recombinant proteins with the baculovirus system can be undertaken, it is necessary to prepare largevolume stocks of recombinant virus (see Basic Protocol 1). The stocks must have a known titer (see Support Protocols 1 and 2) and must be analyzed in small-scale cultures (see Basic Protocol 2) to determine optimal process parameters. Analysis includes preparing samples for electrophoresis (see Support Protocol 4) and monitoring nutrient levels in culture media (see Support Protocol 5). The results of analysis of the kinetics of recombinant protein expression are used to determine the optimal harvest time, which is a balance between maximal production and minimal degradation. Once optimal process parameters (e.g., cell line, media, MOI, time of harvest) are identified, the target protein can be produced on a large scale in bioreactors (see Basic Protocol 3). The design of an animal cell bioreactor is similar to that of a microbial fermentor (Fig. 5.3.1) with modifications of the agitation (e.g., lower range of motor speeds, 20 to 200 rpm; marine impeller instead of Rushton turbine) and aeration (lower range of gas flow rates, headspace aeration) systems (for a review, see Hu and Wang, 1986). Equipment preparation is identical to that of a microbial reactor (UNIT 5.3), and harvesting is by batch centrifugation, continuous feed centrifugation, or tangential flow microfiltration (see Support Protocol 3). Because cell culture processes are run for days or weeks, without any antibiotics, special attention should be paid to the sterility of all operations. Perfusion of insect cell cultures (see Alternate Protocol 1) is an effective technique to increase cell density and volumetric productivity of a reactor. It is particularly well suited to processes for the production of secreted proteins. The basis of the technique is to retain the cells inside the reactor by an internal spin filter, an external filter, or an external centrifuge (for a review, see Tokashiki and Takamatsu, 1993) while adding fresh medium and withdrawing spent medium at equal rates to maintain a constant reactor volume.
Critical Parameters and Troubleshooting
Protein Expression in the Baculovirus System
Insect cell line. Recombinant product expression levels are cell line–dependent (Hink et al., 1991), and it may therefore be advantageous to screen a number of cell lines for relative productive capacities. The robustness, ease of handling in suspension and in large-
scale reactors, and nutrient requirements are also key parameters to consider when choosing a production cell line. Generally, Sf 9 cells (Spodoptera frugiperda clone 9, derived from fall armyworm ovaries) are superior for production of virus, whereas Tn5 cells are superior for secreted proteins. From a practical point of view, it is possible to generate virus stocks in one cell line to be used for recombinant protein production in another. Maintenance of cell lines in a predictable manner (i.e., same cell count and viability profiles over time) is extremely important to ensure reproducible and optimal recombinant product yields. As cell lines grow to different maximum densities under different medium conditions, they should be subcultured at different initial cell inoculum densities. Ideally, cell lines should be subcultured by 10-fold dilution with fresh medium and allowed to grow to densities that are ≤40% of the maximum achievable cell density (for Sf 9 in TC100/5% FCS medium, maximum density is 2 × 106 cells/ml). Above this critical density, the culture deviates from the maximum specific growth rate and may exhibit inefficient utilization of nutrients and subsequent variability in maximal production. Viral stocks. Two major rules are associated with stock virus preparation. (1) Keep the number of times a virus is serially passaged to a minimum (<10). Otherwise, mutant virus can become the predominant species (Kool et al., 1991), leading to a subsequent decrease in recombinant product formation in infected insect cell cultures (Wickham et al., 1991). (2) Cultures serially infected at a high MOI (5 to 10) are more likely to have higher proportions of defective interfering particles than those infected at low MOI. A high cell viability (>95%) is essential for good results. A low MOI (<0.1) allows some degree of postinfection cell growth, which leads to higher numbers of infected cells and the production of high-titer viral stocks. High-titer viral stocks are necessary to prevent the need to add large quantities of viral stock inoculum to large-scale cultures employed for recombinant protein expression. Titration of virus. The relative advantages of the plaque assay and end-point assay may be dependent on the availability of an easy marker for infection (some viruses carry β-galactosidase or luciferase as a reporter gene). Both titration methods take at least 6 days to obtain an estimate with a large error range (50% to 100%). The end-point assay, however, is technically less difficult than the plaque assay,
5.5.14 Current Protocols in Protein Science
which allows many more replicates to be made. This can reduce the error range significantly (down to 15% to 20% for 100 replicates per dilution). In this case initial dilutions can be performed in sterile 1.5-ml microcentrifuge tubes. When, in contrast to β-gal expressing viruses, the recombinant virus does not express a protein that can be quickly and easily detected in the 96-well microtiter plates, the end-point method is more difficult to interpret, and it is advisable to use the plaque assay even at the sacrifice of accuracy. The major hurdles in virus quantification are associated with the inherent nature of the bioassay, which depends on the condition and density of the cells. Cells from poorly growing cultures with low viability cannot be used. Even for healthy cultures, however, if the cell density used is too low, underestimation of the virus titer occurs. If the cell density is too high, nutrients essential for recombinant product expression may be consumed to a large degree by cell growth, which would also lead to the underestimation of virus titers. Expression kinetics. Evaluation of expression kinetics needs to be performed for every recombinant virus. In particular, the optimal time of harvest is a parameter that is specific to each protein. It varies largely with the virus construct (specifically, the temporal class of the viral promoter used for driving gene expression), the type of protein (i.e., cytoplasmic versus secreted), and the extent of post-translational modifications. Initially, infection at a higher MOI (1 to 5) is desirable to establish the optimal times of infection and harvest, especially if product instability is an issue. The MOI cannot directly increase the productive capacity of individual insect cells, but a low MOI (0.01 to 0.0001) can indirectly increase the final volumetric yield due to increased postinfection cell growth. This may be useful for the large-scale expression of stable recombinant proteins and when virus stocks are dilute. A low MOI may lead to variability of product expression due to the inherent inaccuracy of the methods employed for virus stock quantification. Slight underestimation of the true low MOI leads to excessive cell growth and increases the risk of substrate limitation. Conversely, slight overestimation of the true low MOI leads to suboptimal postinfection cell growth, and therefore decreased predicted volumetric yield. Optimal, less variable product yields can be achieved without instability concerns by optimizing the cell density at the time of infection,
employing a high MOI. The cell density at the time of infection can significantly affect production yields of proteins as well as virus. For highest yields of recombinant proteins, it is necessary to infect cultures at the highest possible cell density that supports infection and maximum specific productivites. This density usually occurs immediately prior to the midexponential growth phase and is dependent on the medium and cell line employed. Maintenance of separate virus and cell banks. Physical separation of all virus-associated material and equipment from cell-associated material and equipment is essential. This includes incubators, fume hoods and associated apparatus, bioreactors, probes, cell culture reagents, and media. Accidental contamination of stock cell cultures with virus is potentially disastrous and difficult to detect. Small quantities of contaminating virus may involve a latent infection that is continuously diluted during normal subculture. If a virally contaminated culture is left to overgrow, it may be difficult to detect the infection because the culture may stop growing due to exhaustion of nutrients before a full-scale viral infection can be detected. Production in bioreactors. Expression decreases in batch cultures infected beyond the mid-exponential phase of cell growth. Dissolved oxygen (pO2) is the major additional parameter that can be easily controlled with bioreactors compared to spinner flasks. As a general rule, a set point of 30% to 50% dissolved oxygen tension (of oxygen contained in air) should be optimal and has been shown to be fairly independent of the target protein. Because the culture is sparged either continuously or at intervals with gas, the addition of a shear protectant such as pluronic F-68 is required to avoid mechanical damage to the cells (Murhammer and Goochee, 1988). Basic Protocol 3 describes a procedure for stirred tank reactors. Minor modifications, in particular for pO2 control strategy, are necessary for airlift type reactors. Rice et al. (1993) compared the two reactor types and showed that they were equivalent in terms of protein expression levels. Finally, viability should be monitored closely in large-scale systems, even when the optimal time of harvest has already been established at a small scale, to avoid excessive contamination of the supernatant with cellular proteins and to facilitate subsequent downstream processing. Media. Recombinant product expression is nutrient dependent and therefore medium de-
Production of Recombinant Proteins
5.5.15 Current Protocols in Protein Science
Protein Expression in the Baculovirus System
pendent. Serum-free medium generally supports higher expression levels due to the incorporation of greatly increased quantities of nutrients, including carbohydrates and amino acids. For instance, Sf-900 II contains five times the amount of glucose and twice the amount of glutamine as serum-containing IPL-41 medium and is able to sustain higher maximum cell densities and recombinant product yields. Absence of serum is not directly responsible for this difference, however, as serum-supplemented IPL-41 medium is more enriched and sustains higher cell growth and protein production than serum-supplemented TC100 medium. If not included in the commercial medium, the addition of pluronic F-68 (≤4 g/liter) is essential to protect the cells against the shear stress imposed by a combined sparging and mechanical agitation in bioreactors. Fetal calf serum is available from many vendors. Different lots of serum from a number of suppliers should be obtained and tested. The lot that promotes the best growth rate and cell viability should be purchased in bulk. Antibiotics. The routine use of antibiotics to allow contaminant-free cell culture is not recommended for two reasons. Antibiotics may affect the growth and production potential of the culture. Additionally they may lead to latent bacterial infection in the stock cell cultures with possible resistance that cannot be detected until extended cell growth or during valuable production runs. Harvesting. Establishment of the degree of sensitivity of the target protein to degradation due to cell separation and concentration protocols should be established at small-scale prior to large-scale harvest, as it is considered to be protein specific. In the case of a secreted product, concentration by ultrafiltration should directly follow cell separation. For products that are particularly sensitive to proteolysis, we find it useful to add protease inhibitors (1 mM benzamidine, 1 mM PMSF, 5 mg/liter leupeptin, 5 mg/liter SBTI, and 5 mg/liter pepstatin A) immediately after cell separation. Degradation of cytoplasmic and secreted proteins can happen very quickly at late times of infection and make the protein difficult to purify (McGlynn et al., 1992). For a secreted protein, perfusion alleviates this problem by reducing the residence time of the protein in the bioreactor (Jäger et al., 1992). Tangential flow microfiltration can be performed by attaching the filter system directly to the reactor (see Support Protocol 3) or, alternatively, by transferring the contents of the
reactor to a harvest tank and filtering the tank contents. The filter system should have a minimum of 0.04 m2 per liter of reactor volume so as to allow complete separation in <1 hr. Microfiltration should avoid cell breakage and release of cellular proteases (see Maiorella et al., 1991). This can be difficult considering the fragility of infected cells. It should be carried out quickly (<1 hr for large volumes).
Anticipated Results Generally, reported levels of foreign gene expression range from about 1 to 500 mg recombinant product per 2 × 109 infected cells (Luckow and Summers, 1988; Miller, 1988). Reported expression levels of 25% to 70% total cellular protein produced in infected cultures often represent viral structural proteins such as BlueTongue virus proteins (Roy, 1990) or rabies nucleoprotein (Prehaud et al., 1990). It should be emphasized that these are examples of the potential levels, and 10 to 100 mg per 109 infected cells is the usual expression level range for most heterologous proteins. Comparison with different eukaryotic expression systems such as those employing mammalian cell lines indicates higher production by the baculovirus expression vector system.
Time Considerations The baculovirus life cycle dictates that 4 to 5 days after cell infection be allowed for viral reproduction. Moreover, up to 1.5 to 2 weeks may be required to achieve target cell densities of host cells before infection. Viral stock production takes ∼4 or 5 days postinfection, and, although titration experiments can be set up in 1 day, it takes at least 6 days for titers to be determined. The time required for analysis of expression in small-scale cultures, which depends on how quickly target cell densities are achieved, ranges from 7 to 10 days. Preparing samples for electrophoresis can be accomplished in ∼30 min, but SDS-PAGE or other analysis procedures require 1 to 3 days. Measuring nutrient levels in culture media can range from as short as ∼15 min for simple enzyme assays employing kits or electrodes to as long as 1 day if HPLC is required. Production in bioreactors entails similar time considerations. Preparing a reactor and loading it with medium can take half a day, and it is best to observe the system for 1 or 2 days prior to inoculation to ensure sterility. Perfusion cultures can theoretically be maintained indefinitely. The time required for harvesting batch cultures depends on the volume to be
5.5.16 Current Protocols in Protein Science
processed, ranging from 30 min for batch centrifugation to 45 min for continuous feed centrifugation to 1 hr for tangential flow microfiltration. The last procedure is followed by batch centrifugation, leading to a total time of 1.5 hr.
Literature Cited Belyaev, A.S. and Roy, P. 1993. Development of baculo virus triple and quadruple expression vectors: Co-expression of three or four bluetongue virus proteins and the synthesis of bluetongue virus-like particles in insect cells. Nucl. Acids Res. 21:1219-1223. Bonning, B.C., Roelvink, P.W., Vlak, J.M., Possee, R.D., and Hammock, B.D. 1994. Superior expression of juvenile-hormone-esterase and betagalactosidase from the basic protein promoter of Autographa californica nuclear-polyhedrosis virus compared to the p10 protein and polyhedrin promoters J. Gen. Virol. 73:1551-1556. Chazenbalk, G.D. and Rapoport, B. 1995. Expression of the extracellular domain of the thyrotropin receptor in the baculovirus system using a promoter active earlier than the polyhedrin promoter. Implications for the expression of functional highly glycosylated proteins. J. Biol. Chem. 270:1539-1543. Deramoudt, F.X., Monnet, S., Rabaud, J.N., Quiot, J.M., Cerutti, M., Devauchelle, G., and Kaczorek, M. 1994. Production of a recombinant protein in a high density insect cell cytoflow reactor. In Animal Cell Technology: Products of Today, Prospects for Tomorrow. (R.E. Spier, J.B. Griffiths, and W. Berthold, ed.) pp 222-226. Butterworth. Doyle, A., Griffiths, J.B., and Newell, D.G. 1994. Biochemistry of cells in culture. In Cell & Tissue Culture: Laboratory Procedures. John Wiley & Sons, Chichester, UK. Hink, W.F., Thomsen, D.R., Davidson, D.J., Meyer, A.L., and Castellino, F.J. 1991. Expression of three recombinant proteins using baculovirus vectors in 23 insect cell lines. Biotechnol. Prog. 7:9-14. Hu, W.S. and Wang, D.I.C. 1986. Mammalian cell culture technology: A review from an engineering perspective. In Mammalian Cell Technology (W.G. Thilly, ed.) pp. 167-199. Butterworths, London. Jäger V., Grabenhorst, E., Kobold, A., Deutschmann, S.M., and Conradt, H.S. 1992. High density perfusion culture of insect cells for the production of recombinant glycoproteins. In Baculovirus and Recombinant Protein Production Processes (J.M. Vlak, E.J. Schlager, and A.R. Bernard, eds.) pp. 274-284. Roche Editions, Basel. Jarvis, D.L. and Aurelio, G., Jr. 1994. Long-term stability of baculoviruses stored under various conditions. BioTechniques 16:508-513.
Kool, M., Voncken, J.W., van Lier, F.L.J., Tramper, J., and Vlak, J.M. 1991. Detection and analysis of Autographa californica nuclear polyhedrosis virus mutants with defective interfering properties. Virology 183:739-746. Luckow, V.A. and Summers, M.D. 1988. Trends in the development of baculovirus expression vectors. Bio/Technology 6:47-55. Maiorella, B., Dorin, G., Carion, A., and Harano, D. 1991. Crossflow microfiltration of animal cells. Biotechnol. Bioeng. 37:121-126. McCutchen, B.F., Choudary, P.V., Crenshaw, R., Maddox D., Hammock, B.D., and Maeda S. 1991. Development of a recombinant baculo virus expressing an insect-selective neurotoxin: Potential for pest control. Bio/Technology 9:848852. McGlynn, E., Becker, M., Mett, H., Reutener, S., Cozens, R., and Lydon, N.B. 1992. Large-scale purification and characterisation of a recombinant epidermal growth-factor receptor proteintyrosine kinase. Modulation of activity by multiple factors. Eur. J. Biochem. 207:265-275. Miller, L.K. 1988. Baculoviruses as gene expression vectors. Annu. Rev. Microbiol. 42:177-199. Murhammer, D.W. and Goochee, C.F. 1988. Scaleup of insect cell cultures: Protective effects of Pluronic F68. Bio/Technology 6:1411-1418. Murphy, C.I. and Piwnica-Worms, H. 1994a. Preparation of insect cell cultures and baculovirus stocks. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 16.10.1-16.10.8. John Wiley & Sons, New York. Murphy, C.I. and Piwnica-Worms, H. 1994b. Generation of recombinant baculoviruses and analysis of recombinant protein expression. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 16.11.116.11.19. John Wiley & Sons, New York. Ozturk, S.S., Meyerhoff, M.E., and Palsson, B.O. 1989. Measurement of ammonia and glutamine in cell culture media by gas sensing electrodes. Biotechnol. Tech. 3:217-222. Prehaud, C. Harris, R.D., Fulop, V., Koh, C.L., Wong, J., Flamand, A., and Bishop, D.H. 1990. Expression, characterization, and purification of a phosphorylated rabies nucleoprotein synthesized in insect cells by baculovirus vectors. Virology 178:486-497. Rankl, N.B., Rice, J.W., Gurganus, T.M., Barbee, J.L., and Burns, D.J. 1994. The production of an active protein kinase C-delta in insect cells is greatly enhanced by the use of the basic protein promoter. Protein Expression Purif. 5:346-356. Rice, J.W., Rankl, N.B., Gurganus, T.M., Marr, C.M., Barna, J.B., Waters, M.M., and Bruns, D.J. 1993. A comparison of large-scale Sf 9 insect cell growth and protein production: Stirred vessel vs. airlift. BioTechniques 15:1052-1059.
Production of Recombinant Proteins
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Roy, P. 1990. Use of baculovirus expression vectors: Development of diagnostic reagents, vaccines and morphological counterparts of bluetongue virus. FEMS Microbiol. Immunol. 2:223-234. Sridhar, P. and Hasnain, S.E. 1993. Differential secretion and glycosylation of recombinant human chorionic gonadotropin (beta-HCG) synthesized using different promoters in the baculo virus expression vector system. Gene 131:261-264. Sridhar, P., Panda, A.K., Pal, R., Talwar, G.P., and Hasnain, S.E. 1993. Temporal nature of the promoter and not relative strength determines the expression of an extensively processed protein in a baculovirus system. FEBS Lett. 315:282-286. Stewart, L.M.D., Hirst, M., Lopez-Ferber, M., Merryweather, A.T., Cayley, P.J., and Possee, R.D. 1991. Construction of an improved baculo virus insecticide containing an insect-specific toxin gene. Nature 352:85-88. Summers, M.D. and Smith, G.E. 1987. A manual of methods for baculovirus vectors and insect cell culture procedures. Texas Agricultural Experimental Station Bulletin No. 1555. College Station, Texas. Tokashiki, M. and Takamatsu, H. 1993. Perfusion culture apparatus for suspended mammalian cells. Cytotechnology 13:149-159. van-Lier, F.L.J., van-Duijnhoven, G.C.F., de-Vaan, M.M.J.A.C.M., Vlak, J.M., and Tramper, J. 1994. Continuous beta-galactosidase production in insect cells with a p10 gene based baculo virus vector in a two-stage bioreactor system. Biotechnol. Prog. 10:60-64.
Wang, M.Y., Kwong, S., and Bentley, W.E. 1993. Effects of oxygen/glucose/glutamine feeding on insect cell baculo virus protein expression: A study on epoxide-hydrolase production. Biotechnol. Prog. 9:355-361. Wickham, T.J., Davis, T., Granados, R.R., Hammer, D.A., Shuler, M.L., and Wood, H.A. 1991. Baculovirus defective interfering particles are responsible for variations in recombinant protein production as a function of multiplicity of infection. Biotechnol. Lett. 13:483-488.
Key References Summers and Smith, 1987. See above. O’Reilly, D., Miller, L.K., and Luckow, V.A. 1994. Baculovirus expression vectors: A laboratory manual. Oxford University Press, New York. King, L.A. and Possee, R.D. 1992. The baculovirus expression system. A laboratory guide. Chapman & Hall, London. These manuals give excellent background information on the insect cell and baculovirus biology, expression vectors and strategies for optimization. Includes very practical aspects of many elementary but specialized procedures.
Contributed by Alain Bernard, Mark Payton, and Kathryn R. Radford Glaxo Institute for Molecular Biology Geneva, Switzerland
Protein Expression in the Baculovirus System
5.5.18 Current Protocols in Protein Science
Overview of Protein Expression in Saccharomyces cerevisiae For more than a decade, the yeast Saccharomyces cerevisiae has been extensively utilized for the production of foreign proteins. Numerous characteristics of this system account for its popularity, among them (1) the existence of well-developed tools for manipulation of yeast DNA, including expression vectors and host strains; (2) a vast knowledge base regarding the genetics and biochemistry of the organism; (3) a eukaryotic secretory pathway; (4) eukaryotic post-translational modification pathways, such as those for N-linked and O-linked glycosylation; and (5) an extensive track record as a safe organism. Foreign gene expression in S. cerevisiae has been reported for a wide variety of proteins derived from fungal and mammalian species, and many laboratories have contributed to a substantial base of vectors and host strains. Romanos et al. (1992) provide an extensive overview of strategies and progress toward the expression of foreign genes in S. cerevisiae as well as other yeasts. However, although S. cerevisiae has a well-developed eukaryotic secretory pathway, it is not the most efficient yeast for high-level export of proteins to the extracellular medium. There have been notable successes with respect to secretion of some foreign proteins in S. cerevisiae (Innis et al., 1985; Van Arsdell et al., 1987; Penttila et al., 1988; Hitze-
UNIT 5.6
man et al., 1990), but often transport to the extracellular medium is inefficient, especially for complex mammalian proteins of high molecular weight. Nevertheless, the application of advanced genetic and molecular biology tools should bring continued improvements in secretory capabilities of engineered S. cerevisiae. As with all heterologous expression systems, the production of foreign proteins in S. cerevisiae poses various challenges, some of which have not yet been resolved. In general, the goals for expression of foreign proteins in S. cerevisiae are (1) attainment of desired protein yield; (2) production of protein with appropriate post-translational modifications, desired conformation, and biological function; and (3) secretion to the extracellular medium. Although these criteria do not apply to all expression projects, many proteins intended for human therapeutic use are normally secreted, and complex post-translational processing is required for biological function. It is especially for these cases that further engineering of S. cerevisiae vectors and host strains is required to improve both the product yield and product quality—particularly with regard to post-translational glycosylation, for which there are several important differences between S. cerevisiae and mammalian cells (Ballou et al., 1980; Kukuruzinska et al., 1987).
Table 5.6.1
Representative Vectors for Foreign Gene Expression in S. cerevisiaea
Vector
Promoter
Signal
Selectable marker
Replication platform
pTK46
PGK1
MF-α1
URA3
pTE432 pMA230
CUP1 PGK1 PHO5/GPD1 GPD1
Integrated (TY-δ) 2µm 2µm 2µm Integrated (rDNA) 2µm 2µm
pMIRY 2
pYBDT-10 PHO5 PGK1 pYEULS1
PGK1
— — — — tPA (human) levanase (B. subtilis) killer toxin (K. lactis)
TRP1 LEU2 LEU2 LEU2-d TRP1 LEU2
LEU2-d 2µm and URA3
Reference Sakai et al. (1991) Etcheverry (1990) Tuite et al. (1982) Hinnen et al. (1989) Lopes et al. (1989) Lemontt et al. (1985) Wanker et al. (1995) Castelli et al. (1994)
aAbbreviations: CUP1, copper-binding metallothionein; GPD1, glyceraldehyde-3-phosphate dehydrogenase; MF-α1,
α-factor; PGK1, 2-phosphoglycerate kinase; PHO5, acid phosphatase; rDNA, ribosomal DNA; tPA, tissue plasminogen activator; Ty-δ, δ sequences of Ty transposable elements.
Contributed by Robert L. Strausberg and Susan L. Strausberg Current Protocols in Protein Science (1995) 5.6.1-5.6.7 Copyright © 2000 by John Wiley & Sons, Inc.
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VECTORS AND HOST STRAINS Through the efforts of a diverse and talented community, there are now many vectors and host strains available to direct gene expression in S. cerevisiae (Romanos et al., 1992). A variety of choices is available with respect to specific elements used to direct expression and secretion—e.g., the promoter to direct expression, the signal sequence for secretion, the expression cassette copy number and mechanism for replication, and the selectable marker to establish and/or maintain transformants. Some representative vectors that have been successfully used to direct foreign gene expression in S. cerevisiae are listed in Table 5.6.1. These examples illustrate the diversity of choices available to effect production of foreign proteins in this yeast.
Promoters
Overview of Protein Expession in Saccharomyces cerevisiae
Many different promoters have been used to successfully direct expression of foreign genes in S. cerevisiae. In some cases, highly efficient promoters from the glycolytic pathway (glyceraldehyde-3-phosphate dehydrogenase, alcohol dehydrogenase, and phosphoglycerate kinase) have been used (Shuster, 1989). Because these promoters are among the strongest in S. cerevisiae, they are reasonable choices to effect high-level expression of foreign genes. However, these promoters should be used with care because expression is up-regulated by glucose, a common carbon source in yeast media. Many tightly regulated, relatively strong yeast promoters are available to direct expression in S. cerevisiae, including those from GAL1, PHO5, and CUP1. In these cases, regulated expression can be achieved through simple manipulation of the growth medium by addition of metabolites or ions (Kramer et al., 1984)—galactose (GAL1), phosphate (PHO5), and copper (CUP1)—in a manner that will allow the transition from shaker flasks to largescale fermentors. Through the use of these promoters, it is relatively simple to regulate the timing of expression, such as toward the end of a large-scale fermentation. In employing these and other expression systems, it is especially important to carefully consider the choice of host strain. For example, many host strains carry the gal2 mutation, making the cells refractive to galactose import. Also, host strains vary considerably in their resistance to copper ions. In addition to the naturally occurring regulated promoters, investigators have creatively utilized host strains carrying genetic alterations
to achieve regulated expression of strong yeast promoters. For example, the MF-α1 (α-factor) promoter is constitutively expressed in wildtype cells of the α mating type. Brake et al. (1984) employed a MATα strain carrying a mutation in the sir3 gene (which regulates mating type–specific gene expression) to achieve temperature-regulated expression of epidermal growth factor driven by the MF-α1 promoter.
Vector Maintenance and Copy Number In general, assuming the criterion of regulated expression can be achieved, high copy number of the expression cassette is considered desirable. High copy number can sometimes enhance expression levels and can result in greater stability of transformed yeast strains. To effect autonomous regulation of the expression cassette, many investigators choose the 2µm circle replication system. Copy number of vectors based on this system can vary depending on choice of selectable marker (see Selectable Markers) and range from ∼10 to 200 copies per cell. An alternative approach to achieve both high copy number and stable replication is to integrate the expression cassette at repeated sites within the yeast genome. Lopes et al. (1989) developed an integrating plasmid, pMIRY2, that carries a portion of the rDNA of S. cerevisiae and can integrate at high copy number (greater than 100 copies) within this repetitive genomic region. Sakai et al. (1991) employed a strategy that led to integration of multiple copies of expression cassettes between δ sequences of Ty transposable elements on yeast chromosomes.
Transcription Terminators Terminators from many yeast genes have been employed to efficiently terminate expression of heterologous mRNAs. It is difficult to discern clear differences in expression efficiencies based on the particular terminator employed, but it is important to note that inclusion of a transcription terminator in the expression cassette is desirable for assuring maximal effectiveness for the cassette.
Selectable Markers An important consideration in developing expression vectors is the choice of selectable marker. This component can influence the ease of yeast transformation, copy number and stability of the expression cassette, and adaptability of the transformed strains for large-scale
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fermentation. The most widely used selectable markers are yeast genes that complement auxotrophic mutations in host strains. These include, for example, the LEU2 and URA3 genes used in strains that are leu2 or ura3, respectively. Host strains have been assembled that carry very stable forms of these mutant loci, including strains that carry two lesions within the LEU2 gene (leu2-3 and leu2-112). In addition to maintenance based on auxotrophic complementation, positive selection is also available based on resistance to drugs such as G418 (Webster and Dickson, 1983) or to ions such as copper (Henderson et al. 1985). Positive selection is particularly useful for transformation of strains lacking auxotrophic mutations, especially industrial yeasts. Selectable markers should be chosen carefully because the marker may have unexpected effects, e.g., on expression cassette copy number and ease of transformation of host strains. For example, two forms of the LEU2 selectable marker are available, one of which (LEU2-d) carries only a portion of the LEU2 promoter (Beggs, 1978; Erhart and Hollenberg, 1983). As a result, vectors carrying this marker are maintained at very high copy number, but transformation efficiencies are much reduced.
CHALLENGES TO THE EXPRESSION OF FOREIGN PROTEINS Although the differences among expression vectors can be very subtle, effects on expression of foreign proteins can be quite profound. To illustrate this, results are presented from a series of studies on the expression of a human glycoprotein, antithrombin III (At-III; Strausberg and Strausberg, 1991). This example was chosen because it amply illustrates many of the challenges that can be encountered in attempting to attain high-level expression, post-translational modification, and secretion to the extracellular medium. At-III is a human plasma protein with three disulfide bonds, three sites for N-linked glycosylation, and a nonglycosylated molecular weight of 43,000. A number of important goals were set for expression of this protein in S. cerevisiae, including (1) expression levels of at least several hundred milligrams per liter of culture; (2) proper signal processing to produce an amino acid sequence identical to that of the protein expressed in humans; (3) proper conformation, including the formation of the three disulfide bonds; (4) post-translational amino acid modification, especially N-linked glyco-
sylation of three asparagine residues; and (5) secretion to the extracellular medium. The potential of using S. cerevisiae for the production of glycosylated, biologically active At-III was initially tested using an expression vector carrying sequences known to efficiently direct secretion. A vector carrying the promoter and signal sequence from the well-characterized MF-α1 gene was chosen because this gene encodes an efficiently secreted yeast peptide, α-factor (see Fig. 5.6.1). In this case, secretion is directed by both pre and pro sequences; proteolytic processing of these pre and pro sequences is well understood both genetically and biochemically. Other important yeast elements included in the initial expression vectors for At-III production were a replication origin from the yeast 2µm circle (chosen because vectors with this origin replicate in a stable manner and can be maintained at medium or high copy number), a transcription terminator from the highly expressed glyceraldehyde-3-phosphate dehydrogenase (GPD1) gene for efficient termination, and the yeast LEU2-d gene as a selectable marker. (LEU2-d is a truncated form of the LEU2 gene lacking a portion of the promoter region. An order-ofmagnitude increase in vector copy number is associated with use of this selectable marker, compared to that of the full-length LEU2 gene in leu2 host strains.) Yeast strains of MATa and MATα mating types were transformed with this vector and several very striking results were obtained. First, it was clear from immunoblot analysis and biological activity assays that yeast can produce glycosylated At-III with biological activity. The At-III protein detected through the immunoblot appeared as a very heterogeneous band, indicating the addition of N-linked core glycosylation in the endoplasmic reticulum and outer chains of variable length in the Golgi complex. This pattern is very typical of yeast glycosylated proteins such as invertase (Kukuruzinska et al., 1987). Second, examination of the At-III expressed in yeast and treated with endoglycosidase H to remove carbohydrate (UNIT 12.4) revealed that the α-factor prepro sequence was retained in a substantial portion of the protein population. Third, overall expression was estimated at less than one percent of total cell protein. Interestingly, transformation with the high-copy-number vector (YpGX235) was only successful in cells of the MATα mating type, where activity of the MF-α1 promoter was expected to be very low. This result suggested that high-level expression of At-III, ex-
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UAS
GAL1
TIS
signal
cDNA
MF –α1
SUC2
terminator
PHO5
GPD1
Figure 5.6.1 Genetic constructions for expression of At-III. Abbreviations: GAL1, galactose; GDP1, glyceraldehyde-3-phosphate dehydrogenase; MF-α1, α-factor; PHO5, acid phosphatase; SUC2, invertase; TIS, transcription initiation site; UAS, upstream activation site.
pected in MATα cells, might be toxic to S. cerevisiae. Even at this low level of expression, most of the At-III remained cell-associated and segregated with insoluble cellular material. This result was consistent with retention of the α-factor prepro sequence.
Attainment of Desired Production Yield
Overview of Protein Expession in Saccharomyces cerevisiae
To achieve regulated, high-level expression, a hybrid promoter was assembled from the yeast GAL1 and MF-α1 genes. This promoter design was chosen because regulation of yeast GAL1 is well understood and permits a variety of regulation schemes suitable for large-scale fermentation. The hybrid promoter consisted of the upstream activation site (UAS) from GAL1 positioned upstream from the MF-α1 transcription initiation site. Other important elements in the vector include a combination of the 2µm origin of replication and LEU2-d (for high copy number) and a GPD1 transcription terminator. Initial expression results in yeast cells transformed with this vector were only somewhat encouraging. Although regulated expression of At-III protein was observed, the actual production levels were still quite low, possibly due to the limited availability of regulatory proteins associated with expression from the GAL1 promoter. In this system, inducible expression is
dependent upon a trans-acting regulatory protein, GAL4, that interacts with the UAS (Johnston, 1987). In the absence of galactose, GAL4 complexes with another protein, GAL80, and is unavailable for interaction with the UAS. Upon induction with galactose, GAL4 protein dissociates from GAL80 and binds to the UAS, thereby stimulating protein induction. It was known that the GAL4 protein is available in very limited quantities in wildtype yeast cells, so it seemed reasonable that most of the copies of the hybrid GAL1/MF-α1 promoter might remain dormant. Therefore, an additional modification was made so that the GAL4 gene was positioned within the highcopy-number expression vector. The results were again only moderately encouraging because, although expression was increased several fold, the overall expression levels were no more than a few percent of total soluble protein. Immunoblot analysis revealed that ∼90% of the At-III remained associated within the insoluble cell fraction. A very striking impact on expression of At-III resulted from the use of different signal peptide coding sequences upstream from the start of mature At-III. The main goal in assessing the use of different sequences was to assess the impact on secretion, but the most noteworthy result was observed with respect to effect on expression level. Vectors were assembled
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Table 5.6.2
Effect of Signal Sequence on Production of Soluble At-III Protein
Vectora
Signal
Promoter
YpGX243GAL4 YpGX254GAL4 YpGX262GAL4
MF-α1 prepro SUC2 PHO5
GAL1/MF-α1 GAL1/MF-α1 GAL1/MF-α1
Soluble At-III (mg/liter) producedb 2.7 3.6 121.0
aYeast strain D8 (MATα leu2-3 leu2-112) was transformed with each vector, and cells were grown
and induced as described in Strausberg and Strausberg (1991). bDetermined by radioimmunoassay.
with signal sequences derived from various secreted proteins including invertase (SUC2), α-factor (MF-α1), and acid phosphatase (PHO5; see Fig. 5.6.1). For extensive discussions of signal sequences used to direct secretion in yeast see Kingsman et al. (1985) and Romanos et al. (1992). Each of the signal peptide coding sequences was assembled through insertional directed mutagenesis so that no extraneous codons were present between the signal peptide and mature protein coding sequences. For this experiment, the PHO5 signal peptide was designed predominantly from preferred yeast codons, the codons most often utilized in highly expressed yeast genes. Expression levels were monitored by radioimmunoassay and immunoblotting. These studies were very enlightening with regard to parameters that can influence expression levels for foreign proteins in S. cerevisiae. First, upon induction, expression levels of soluble At-III (most of the At-III is insoluble) differed by more than an order of magnitude among the different vectors (Table 5.6.2). This result was most striking in the case of YpGX262GAL4, in which secretion is directed by the PHO5 signal. In this case, At-III protein accumulated to ∼20 percent of total cell protein and microscopic examination of the yeast cells revealed particulate bodies reminiscent of the inclusion bodies associated with high-level protein expression in Escherichia coli. Although the mechanism for the very high level of At-III expression associated with this vector was not determined, a reasonable interpretation is that the use of preferred codons in the PHO5 signal peptide coding sequence enhanced translation initiation.
Production of Protein with Appropriate Post-translational Modifications, Conformation, and Function One of the most difficult issues concerning production in yeast of glycoproteins intended
for human therapy is the difference in glycosylation patterns for both N- and O-linked glycosylations. Although N-linked glycosylation in the endoplasmic reticulum in yeast is quite similar to that in mammals, substantial differences are associated with processing in the Golgi complex. In the endoplasmic reticulum, both systems add core units composed of mannose, N-acetylglucosamine, and glucose residues, and processing of certain glucose and mannose residues follows. However, in the Golgi complex, mammalian cells continue to process mannose residues and additional residues such as galactose and sialic acid. In contrast, S. cerevisiae performs neither of these functions, but instead adds extensive outer chains with many branches that can be heterogeneous in size. Genetic approaches have been employed to facilitate the production in yeast of glycoproteins more similar to their human counterparts. Ballou and colleagues (Ballou et al., 1980) provided a basis for this approach based on careful genetic dissection of the glycosylation pathway. For example, yeast strains with the mnn9 mutation do not add further mannose units in the Golgi complex. In the case of At-III, use of strains carrying this mutation resulted in production of a homogeneous protein with only core glycosylation. Further utilization of the mnn family of mutations may permit engineering of yeast-produced glycoproteins with improved properties. Expression levels can markedly affect posttranslational modification of foreign proteins. Thus, in the At-III example, expression at relatively low levels resulted in protein that carried full glycosylation, including inner and outer chains. However, as expression was enhanced, glycosylation was reduced, and at high-level expression directed by YpGX262, most of the At-III was unglycosylated. This results, of course, from the important interplay that occurs between the expression level and the ability of
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yeast cells to process the molecules during their transport through the secretory pathway.
Secretion to the Extracellular Medium
Overview of Protein Expession in Saccharomyces cerevisiae
Although some success has been obtained in secreting complex mammalian proteins extracellularly from S. cerevisiae, the greatest success has only been achieved with smaller peptides or proteins normally secreted by fungal sources. Indeed, in the At-III example, no extracellular protein was detected. Other yeasts appear to secrete mammalian proteins more effectively, posing a major challenge to the future of S. cerevisiae as the system of choice for secreted proteins. However, increased understanding of yeast genetics and biochemistry may be used to substantially improve secretion of foreign proteins in S. cerevisiae. Transport of proteins through a eukaryotic secretory pathway is a complex process involving the participation of many cofactors. For instance, signal peptidase activity is required for removal of signal sequences from mature proteins. In the case of α-factor, additional proteases encoded by the KEX2 and STE13 genes are required for cleavage of the pro sequence from the mature peptide. Thus, it is possible that with overexpression of secreted proteins, these cofactors become limiting. This is supported by the observation of HaguenauerTsapis and Hinnen (1984) that upon overexpression of yeast acid phosphatase, precursor protein accumulates. In some cases, overexpression of limited processing enzymes has a beneficial effect. Barr et al. (1987) noted that multiple copies of KEX2 enhanced the ability of S. cerevisiae to process and secrete a foreign protein, transforming growth factor, linked to the α-factor prepro sequence. Therefore, careful examination of processing defects can identify limited processing enzymes within the cell and their increased expression can, at least in some cases, improve processing and secretion. In addition to protease activities, many other protein functions are required for efficient secretion. The activity of several accessory proteins is required for proper conformational folding within the endoplasmic reticulum— e.g., molecular chaperones as well as proteins specifically associated with disulfide bond formation. Robinson et al. (1994) reported that overexpression of protein disulfide isomerase, an enzyme that catalyzes disulfide exchange within the endoplasmic reticulum, resulted in improved secretion of both human platelet-derived growth factor B homodimer and Schizo-
saccharomyces pombe acid phosphatase from S. cerevisiae. Furthermore, the ease of S. cerevisiae genetic manipulation offers the potential for improved secretion even in cases where specific defects have not been identified. Toward that end, several groups have applied mutagenesis strategies in conjunction with rapid screening methods to identify yeast strains with enhanced secretion capability. Smith et al. (1985) applied this approach to identify yeast strains that carried mutations leading to enhanced secretion of chymosin. Importantly, improved secretion of additional, but not all, foreign proteins was associated with one of these mutations.
TIME FRAME AND FUTURE PROSPECTS As summarized in this commentary, tools for the expression of foreign genes in S. cerevisiae are very well developed, and allow the individual researcher to establish within a few months the relative capabilities of S. cerevisiae as a host for expression of new target proteins. However, in many cases the challenges are substantial and long-term dedicated efforts are required for ultimate success. Prospects for further engineering of S. cerevisiae cells for the expression of foreign proteins seem quite bright, in part because a cooperative international effort is likely to soon complete the entire sequence of the S. cerevisiae genome. The availability of a complete repertoire of genetic information will provide a very firm basis for better understanding of complex eukaryotic pathways, and will likely suggest new avenues for engineering this organism for enhanced expression of complex mammalian proteins.
LITERATURE CITED Ballou, L., Cohen, R., and Ballou, C. 1980. Saccharomyces cerevisiae mutants that make mannoproteins with a truncated carbohydrate outer chain. J. Biol. Chem. 255:5986-5991. Barr, P.J., Steimer, K.S., Sabin, E.A., Parkes, D., George-Nascimento, C., Stephans, J.C., Powers, M.A., Gyenes, A., Nan Nest, G.A., Miller, E.T., Higgins, K.W., and Luciw, P.A. 1987. Antigenicity and immunogenicity of domains of the human immunodeficiency virus (HIV) envelope protein expressed in the yeast Saccharomyces cerevisiae. Vaccine 5:90-101. Beggs, J.D. 1978. Transformation of yeast by a replicating hybrid plasmid. Nature 275:104-109. Brake, A.J., Merryweather, J.P., Coit, D.G., Heberlein, U.A., Masiarz, F.R., Mullenbach, G.T., Urdea, M.S., Valenzuela, P., and Barr, P.J. 1984. α-factor-directed synthesis and secretion of mature foreign proteins in Saccharomyces cere-
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Current Protocols in Protein Science
visiae. Proc. Natl. Acad. Sci. U.S.A. 81:46424646. Castelli, L.A., Mardon, C.J., Strike, P.M., Azad, A.A., and Macreadie, I.G. 1994. High-level secretion of correctly processed beta-lactamase from Saccharomyces cerevisiae using a highcopy-number secretion vector. Gene 142:113117. Erhart, E. and Hollenberg, C.P. 1983. The presence of a defective LEU2 gene in 2µm DNA recombinant plasmids is responsible for curing and high copy number. J. Bacteriol. 156: 625-635. Etcheverry, T. 1990. Induced expression using yeast copper metallothionein promoter. Methods Enzymol. 185:319-329. Haguenauer-Tsapis, R. and Hinnen, A. 1984. A deletion that includes the signal peptidase cleavage site impairs processing, glycosylation, and secretion of cell-surface yeast acid phosphatase. Mol. Cell. Biol. 4:2668-2675. Henderson, R.C.A., Cox, B.S., and Tubb, R. 1985. The transformation of brewing yeasts with a plasmid containing the gene for copper resistance. Curr. Genet. 9:133-138. Hinnen, A., Meyhack, B., and Heim, J. 1989. Heterologous gene expression in yeast. In Yeast Genetic Engineering (P.J. Barr, A.J. Brake, and P. Valenzuela, eds.) pp. 193-213. Butterworths, London. Hitzeman, R.A., Chen, C.Y., Dowbenko, D.J., Renz, M.E., Lui, C., Pai, R., Simpson, N.J., Kohr, W.J., Singh, A., Chisholm, V., Hamilton, R., and Chang, C.N. 1990. Use of heterologous and homologous signal sequences for secretion of heterologous proteins from yeast. Methods Enzymol. 185:421-440. Innis, M.A., Holland, M.J., McCabe, P.C., Cole, G.E., Wittman, V.P., Tal, R., Watt, K.W.K., Gelfand, D.H., Holland, J.P., and Meade, J.H. 1985. Expression, glycosylation, and secretion of an Aspergillus glucoamylase by Saccharomyces cerevisiae. Science 228:21-26. Johnston, M. 1987. A model fungal gene regulatory mechanism: The GAL genes of Saccharomyces cerevisiae. Microbiol. Rev. 51:458-47. Kingsman, S.M., Kingsman, A.J., Dobson, M.J., Mellor, J., and Roberts, N.A. 1985. Heterologous gene expression in Saccharomyces cerevisiae. Biotechnol. Genet. Eng. Revs. 3:377-416. Kramer, R.A., DeChiara, T.M., Schaber, M.D., and Hilliker, S. 1984. Regulated expression of a human interferon gene in yeast: Control by phosphate concentration or temperature. Proc. Natl. Acad. Sci. U.S.A. 81:367-370.
Lopes, T.S., Klootwijk, J., Veenstra, A.E., van der Aar, P.C., van Heerikhuizen, H., Raue, H.A., and Planta, R.J. 1989. High-copy-number integration into the ribosomal DNA of Saccharomyces cerevisiae: A new vector for high-level expression. Gene 79:199-206. Penttila, M.E., André, L., Lehtovaara, P., Bailey, M., Teeri, T.T., and Knowles, J.K.C. 1988. Efficient secretion of two fungal cellobiohydrolases by Saccharomyces cerevisiae. Gene 63:103-112. Robinson, A.S., Hines, V., and Wittrup, K.D. 1994. Protein disulfide isomerase overexpression increases secretion of foreign proteins in Saccharomyces cerevisiae. Bio/Technology 12:381-384. Romanos, M.A., Scorer, C.A., and Clare, J.J. 1992. Foreign gene expression in yeast: A review. Yeast 8:423-488. Sakai, A., Ozawa, F., Higashizaki, T., Shimizu, Y., and Hishinuma, F. 1991. Enhanced secretion of human nerve growth factor from Saccharomyces cerevisiae using an advanced δ-integration system. Bio/Technology 9:1382-1385. Shuster, J.R. 1989. Regulated transcriptional systems for the production of proteins in yeast: Regulation by carbon source. In Yeast Genetic Engineering (P.J. Barr, A.J. Brake, and P. Valenzuela, eds.) pp. 83-108. Butterworths, London. Smith, R.A., Duncan, M.J., and Moir, D.T. 1985. Heterologous protein secretion from yeast. Science 230:1219-1224. Strausberg, R.L. and Strausberg, S.L. 1991. Composite yeast vectors. U. S. Patent 5,013,652. Tuite, M.F., Dobson, M.J., Roberts, N.A., King, R.M., Burke, D.C., Kingsman, S.M., and Kingsman, A.J. 1982. Regulated high-efficiency expression of human interferon-α in Saccharomyces cerevisiae. EMBO J. 1:603-608. Van Arsdell, J.N., Kwok, S., Schweikart, V.L., Ladner, M.B., Gelfand, D.H., and Innis, M.A. 1987. Cloning, characterization, and expression in Saccharomyces cerevisiae of endoglucanase I from Trichoderma reesei. Bio/Technology 5:6064. Wanker, E., Klingsbichel, E., and Schwab, H. 1995. Efficient secretion of Bacillus subtilis levanase by Saccharomyces cerevisiae. Gene 161:45-49. Webster, T.D. and Dickson, R.C. 1983. Direct selection of Saccharomyces cerevisiae resistant to the antibiotic G418 following transformation with a DNA vector carrying the kanamycin resistance gene of Tn 903. Gene 26:243-252.
Kukuruzinska, M.A., Bergh, M.L.E., and Jackson, B.L. 1987. Protein glycosylation in yeast. Annu. Rev. Biochem. 56:915-944.
Contributed by Robert L. Strausberg National Institutes of Health Bethesda, Maryland
Lemontt, J.F., Wei, C.M., and Dackowski, W.R. 1985. Expression of active human tissue plasminogen activator in yeast. DNA 4: 419-428.
Susan L. Strausberg University of Maryland Rockville, Maryland Production of Recombinant Proteins
5.6.7 Current Protocols in Protein Science
Supplement 2
Overview of Protein Expression in Pichia pastoris During the early 1970s, the methylotrophic yeast Pichia pastoris was developed as a biological tool to convert methanol—a cheap and readily available carbon source that was being discarded by the oil industry as a waste product—into high-quality protein that could be used in the livestock industry. A combination of the oil crisis that began in the mid-1970s, the fact that Pichia as a component of livestock feed was never able to compete in cost with soybeans, and the emergence of the biotechnology industry led to new attempts to find potential uses for the high protein-production capacity of Pichia. By the early 1980s the focus had shifted to the use of Pichia as a eukaryotic microbial system to produce large quantities of heterologous proteins of interest to biotechnologists, including those in the pharmaceutical industry (Ratner, 1989; Wegner, 1990; Buckholz et al., 1991; Cregg et al., 1993). The successful and rapid introduction of P. pastoris into the general scientific community as a heterologous expression system is a result of several important factors, including the goal of developing P. pastoris into a single-cell protein (SCP) production system, its utility as a model eukaryotic organism for basic molecular and cellular biology studies (Gould et al., 1992), and its similarity—both biologically and technically—to one of the most well-characterized systems in modern biology, the baker’s yeast Saccharomyces cerevisiae (UNIT 5.6). The exploitation of Pichia as a tool to produce useful quantities of biologically relevant proteins has moved into high gear (see Table 5.7.1). Pichia as an expression system is now readily available to the research community and can be used to generate proteins destined for academic scrutiny in addition to proteins produced in Pichia for commercialization. The number of heterologous proteins successfully being expressed in Pichia is doubling almost monthly.
GENERAL CHARACTERISTICS OF PICHIA PASTORIS As a yeast, Pichia pastoris is a microbial eukaryote and is as easy to manipulate as Escherichia coli. It has many of the advantages of eukaryotic expression (e.g., protein processing, folding, and post-translational modifications), and it is faster, easier, and cheaper to use than Contributed by David R. Higgins Current Protocols in Protein Science (1995) 5.7.1-5.7.18 Copyright © 2000 by John Wiley & Sons, Inc.
UNIT 5.7
other eukaryotic expression systems, such as baculovirus (UNITS 5.4 & 5.4) or mammalian tissue culture. It also generally gives higher expression levels. P. pastoris is completely amenable to the genetic, biochemical, and molecular biological techniques that have been developed over the past several decades for S. cerevisiae with little or no modification. In particular, methods for transformation by complementation, gene disruption, and gene replacement developed for S. cerevisiae work equally well for P. pastoris (Cregg et al., 1987, 1989; Guthrie and Fink, 1991). In addition, heterologous protein expression levels are often 10- to 100-fold higher in Pichia. The genetic nomenclature adopted for P. pastoris mirrors that used for S. cerevisiae. For example, the gene from S. cerevisiae that encodes the enzyme histidinol dehydrogenase is called the HIS4 gene; the homologous gene from P. pastoris is called the P. pastoris HIS4 gene, and so on. There is also a very high degree of cross-functionality between P. pastoris and S. cerevisiae. For instance, many S. cerevisiae genes have been shown to genetically complement the comparable mutants in P. pastoris, and vice versa—e.g., the P. pastoris HIS4 gene functionally complements S. cerevisiae his4 mutants, and the S. cerevisiae HIS4 gene functionally complements P. pastoris his4 mutants. Other cross-complementing genes that have been identified include LEU2, ARG4, TRP1, and URA3.
P. pastoris as a Methylotrophic Yeast P. pastoris, one of four different genera of methylotrophic yeasts (the others being Candida, Hansenula, and Torulopsis), is capable of metabolizing methanol as its sole carbon source. The first step in this process is the oxidation of methanol to formaldehyde by the enzyme alcohol oxidase, generating hydrogen peroxide in the process. To avoid hydrogen peroxide toxicity, methanol metabolism takes place within a specialized cell organelle, the peroxisome, which sequesters toxic intermediates from the rest of the cell. Alcohol oxidase is sequestered in the peroxisomes and functions as a homo-octomer, with each subunit containing one noncovalently bound flavin adenine dinucleotide (FAD) cofactor. Alcohol oxidase
Production of Recombinant Proteins
5.7.1 Supplement 2
Table 5.7.1
Examples of Heterologous Protein Expression in Pichia pastoris
Expression level (g/liter)
Where/how expresseda
Reference
I/MutS I/Mut+
Cregg et al. (1987) Tschopp et al. (1987a)
Invertase
0.4 20,000 (U/mg total protein) 2.3
S/Mut+
Tschopp et al. (1987b)
Tumor necrosis factor (TNF) Bovine lysozyme c2
10.0 0.55
I/MutS S/Mut+
Sreekrishna et al. (1989) Digan et al. (1989)
Streptokinase (active) Tetanus toxin fragment C
0.08 12.0
Hagenson et al. (1989) Clare et al. (1991a)
3.0
I/n.r. I/Mut+ and MutS I/MutS
0.45
S/MutS
Clare et al. (1991b)
0.8
S/Mut+/MutS
Vedvick et al. (1991)
1.0 4.0
S/Mut+
Wagner et al. (1992) Barr et al. (1992)
Protein Hepatitis B surface antigen β-Galactosidase
Pertussis antigen P69 Mouse epidermal growth factor (EGF) Aprotinin Kunitz protease inhibitor Human serum albumin
S/Mut+
and
Romanos et al. (1991)
MutS
1.7
I and S/Mut+ S/Mut+ and MutS S/Mut+
Scorer et al. (1993) Despreaux and Manning (1993) Larouche et al. (1994)
Porcine leukocyte lipoxygenase (72 kD) Human proteinase cathepsin E (homodimer of 42 kD protein)
n.r.
I/MutS
Reddy et al. (1994)
n.r.
S/MutS
Yamada et al. (1994)
Bm86 (tick gut glycoprotein) Alpha amylase
1.5 2.5
S/MutS S/MutS
Rodriguez et al. (1994) Paifer et al. (1994)
Kunitz protease inhibitor (APLP-2)
1.0
S/Mut+
Van Nostrand et al. (1994)
S/MutS S/MutS
White et al. (1994) Ridder et al. (1995)
HIV-1 gp120 Carboxypeptidase B
1.25 0.8
TAP (tick anticoagulant protein)
Thrombomodulin Rabbit scFv (30 kD)
0.01-0.03 >0.1
CP15 (Cryptosporidium parvum) Rat Pit-1
n.r. n.r.
n.r. n.r.
Jenkins and Fayer (1995) Howard and Maurer (1995)
Human N-terminal transferrin Amyloid precursor protein
0.27 0.001-0.004
S/MutS S/MutS
Steinlein et al. (1995) Ohsawa et al. (1995)
S/MutS I/MutS
Fryxell et al. (1995) Weiss et al. (1995)
S/Mut+ I/MutS
Halaas et al. (1995) Shiba et al. (1995)
S/Mut+
Vozza et al., in press
Human CD38 Hamster prion protein PrPc Mouse OB gene (leptin) Human alanyl-tRNA synthetase
∼0.16 <0.001
0.01 3% (total protein) Recombinant enterokinase (26 kD) 0.006 Overview of Protein Expression in Pichia pastoris
aAbbreviations: I, intracellular; Mut+, methanol utilization wild type (genotype AOX1 AOX2); MutS, methanol utilization
slow (genotype aox1 AOX2); n.r., not reported or not available; S, secreted.
5.7.2 Supplement 2
Current Protocols in Protein Science
has a poor affinity for O2 and P. pastoris compensates for this by generating large amounts of the enzyme. There are two genes in P. pastoris that code for alcohol oxidase—AOX1 and AOX2—but the AOX1 gene is responsible for the vast majority of alcohol oxidase activity in the cell. Expression of the AOX1 gene is tightly regulated and is induced by methanol to very high levels, typically ≥30% of the total soluble protein in cells grown with methanol as the carbon source. The AOX1 gene has been isolated and a plasmid-borne version of its promoter is used to drive expression of the gene of interest encoding a desired heterologous protein (Ellis et al., 1985; Tschopp et al., 1987a; Koutz et al., 1989). Expression of the AOX1 gene is controlled at the level of transcription. In methanolgrown cells ∼5% of the polyA+ RNA is from the AOX1 gene. Regulation of the AOX1 gene is similar to regulation of the GAL1 gene (and others) of S. cerevisiae in that control involves a two-step process: a repression/derepression mechanism plus an induction mechanism. However, unlike the situation in S. cerevisiae, derepression alone of the AOX1 gene (i.e., absence of a repressing carbon source such as glucose) is not sufficient to generate even minute levels of expression from the AOX1 gene. The inducer, methanol, is also necessary to obtain even minimally detectable levels of AOX1 expression (Ellis et al., 1985; Tschopp et al., 1987a; Koutz et al., 1989). Loss of the AOX1 gene (see Transformation by Integration), and thus a loss in most of the cell’s alcohol oxidase activity, results in a strain that is phenotypically MutS (methanol utilization slow). This phenotype is caused by the loss of the alcohol oxidase activity encoded by the AOX1 gene, which results in the cell’s inability to metabolize methanol. Thus, a poor growth
Table 5.7.2 Expression
phenotype on methanol media is observed. (Mut+ strains are wild type and capable of metabolizing methanol as their sole carbon source.)
HETEROLOGOUS PROTEIN EXPRESSION P. pastoris has been used successfully to express a wide range of heterologous proteins as either intracellular or secreted proteins (see Table 5.7.1). Secretion requires the presence of a signal sequence on the expressed protein to target it to the secretory pathway. Although several different secretion signal sequences have been used successfully, including the native secretion signal present on some heterologous proteins, success has been variable. The secretion signal sequence from the S. cerevisiae MATα factor prepro peptide (MF-α1) has been used with the most success (Cregg et al., 1993; Scorer et al., 1993). The major advantage of expressing heterologous proteins as secreted proteins is that P. pastoris secretes very low levels of native proteins. That, combined with the very low amount of protein in the minimal Pichia growth medium, means that the secreted heterologous protein comprises the vast majority of the total protein in the medium, and this serves as a first step in purification of the expressed protein (Barr et al., 1992).
Strains for Expression The P. pastoris host strains have defects in either the histidinol dehydrogenase activity encoded by the HIS4 gene (strains GS115, KM71, SMD1163, SMD1165, and SMD1168), or the arginosuccinate lyase activity encoded by the ARG4 gene (strain GS190), or both (strain PPF1; see Table 5.7.2). All of these strains will grow on complex media such as YPD (yeast
Some Strains of Pichia pastoris Used for Heterologous Protein
Strain
Genotype
Feature
GS115 GS190 KM71 PPF1
his4 arg4 his4 arg4 aox1::ARG4 his4 arg4
SMD1163 SMD1165 SMD1168
his4 pep4 prB1 his4 prB1 his4 pep4
Wild-type strain Wild-type strain MutS Two selectable auxotrophic markers Double protease mutant Protease mutant Protease mutant
Vector selectable marker required HIS4 ARG4 HIS4 HIS4 or ARG4 HIS4 HIS4 HIS4
Production of Recombinant Proteins
5.7.3 Current Protocols in Protein Science
Supplement 2
Table 5.7.3
Media for Pichia pastoris
Mediaa
YNBb
YPD RDBe MDf MMf MGY MM BMGYg
— 0.34% 0.34% 0.34% 0.34% 0.34% 0.34%
— 1% 1% 1% 1% 1% 1%
2% — — — — — 2%
BMMY
0.34%
1%
2%
(NH4)2SO4
Peptone
Carbon sourcec
Phosphate bufferd
Other
Use
1% — — — — — 1%
2% glu 2% glu 2% glu 0.5% meth 1% gly 0.5% meth 1% gly
— — — — — — 100 mM
— 1 M sorbitol — — — — —
1%
0.5% meth
100 mM
Rich medium Transformation Mut+/MutS Mut+/MutS Expression Expression Alternate expression Alternate expression
Yeast extract
—
aAll of these media can be made either as liquid or, by adding agarose to 2% (w/v), as plates. bYeast nitrogen base without amino acids and without (NH ) SO . 42 4 cAbbreviations: glu, glucose (same as dextrose); gly, glycerol; meth, methanol. dStock: 1 M potassium phosphate buffer, adjusted to desired pH between 5 and 8.
eNo amino acid supplements; these media are normally used with transformed strains that are His+ (or Arg+) prototrophs.
fMD and MM media can be used to identify MutS clones. Growth of both Mut+ and MutS clones will be wild type on MD medium. Growth of Mut+ will be wild type on MM medium, but growth of MutS on MM will be severely retarded to completely absent after 2 days because of the inability of MutS clones to efficiently metabolize methanol as a carbon source. gThe addition of yeast extract and peptone to expression media may improve the results of heterologous protein expression in some cases. The mode of action of this improvement is not known, but it may be that yeast extract and peptone serve as competitive inhibitors of proteases and/or act as weak medium buffering agents.
Overview of Protein Expression in Pichia pastoris
extract, peptone, dextrose) and on minimal media, such as MD (minimal dextrose), MM (minimal methanol), and MGY (minimal glycerol), that have been supplemented with histidine and/or arginine (Table 5.7.3). Until transformed with the appropriate vectors, these strains will not grow on minimal medium lacking histidine (those strains with genotype including his4) or arginine (those strains with genotype including arg4). Selection of His+ or Arg+ prototrophs among his4/arg4 mutant strains transformed with a plasmid carrying the wild-type gene is used as the basis for selecting recombinant strains containing the expression plasmid. Most expression experiments begin with wild-type strain GS115 (his4) in which vectors carrying the HIS4 gene can be selected, but there are several alternative strains, mostly derivatives of GS115, that can be employed to achieve success empirically or when a specific need can be predicted. Both Mut+ and MutS as well as single- and multicopy vector recombinant versions of GS115 can be generated. KM71 (his4 arg4 aox1::ARG4) is a MutS strain because the chromosomal AOX1 gene has been replaced with the ARG4 gene. It can be used when only MutS recombinant strains are desired (see Table 5.7.2).
SMD1163 (his4 pep4 prB1), SMD1165 (his4 prB1), and SMD1168 (his4 pep4) are a series of protease-deficient strains that may provide a more suitable environment for certain proteins that are being expressed as heterologous proteins. The PEP4 gene codes for proteinase A, a vacuolar aspartyl protease required for activation of other vacuolar proteases such as carboxypeptidase Y and proteinase B. Prior to processing and activation by proteinase A, proteinase B has about half of its protease activity. The PRB1 gene codes for proteinase B itself. Therefore, pep4 mutants display a substantial decrease in or elimination of proteinase A and carboxypeptidase Y activity, and partial reduction in proteinase B activity. prB1 mutants show a substantial decrease in or elimination of proteinase B activity only, and pep4 prB1 double mutants show a substantial reduction in or elimination of all three of these protease activities (see Table 5.7.4).
Expression Plasmids Plasmid vectors designed for heterologous protein expression in P. pastoris all have general features in common. These are illustrated in the generic plasmid depicted in Figure 5.7.1. (1) The 5′PAOX1 is the promoter from the Pichia alcohol oxidase (AOX1) gene; it is used to drive
5.7.4 Supplement 2
Current Protocols in Protein Science
Sig MCS TT NotI or BgI II 5′ AOX1
generic plasmid
f1 ori
HIS4
Ap r 3′ AOX1
NotI or BgI II
Kanr
Figure 5.7.1 Generic plasmid for heterologous protein expression in Pichia pastoris. Elements present in all expression plasmids are shown in dark boxes, optional elements present in some plasmids are shown in gray boxes. Abbreviation: TT, transcriptional termination sequence.
methanol-inducible expression of the gene of interest. (2) The MCS is a multiple cloning site with unique restriction endonuclease sites; it is used to insert the gene of interest into the plasmid. (3) Transcription termination sequences (TT) are derived from the native Pichia AOX1 gene; they are used to promote efficient mRNA processing and polyadenylation. (4) HIS4, the wild-type gene for histidinol dehydrogenase, is a selectable marker; it is used to positively select for recombinant Pichia strains that have acquired the vector, which complements the auxotrophic his4 mutation. (5) The 3′AOX1 sequence is derived from a region of the native gene that lies 3′ to the transcription termination sequences; it is required for integration of vector sequence by gene replacement or gene insertion 3′ to the chromosomal AOX1 gene. (6) The ColE1 origin of replication and ampicillin resistance gene (Apr) are required for replication and maintenance in bacteria. (7) Unique restriction endonuclease sites are included that allow for the generation of linear plasmids that can integrate at the AOX1 locus by gene replacement (NotI and BglII). Other unique sites at 5′PAOX1 (SacI and BstXI) and HIS4 (SalI and StuI) are included to allow
generation of a linear plasmid that can integrate into the genome at these loci by gene insertion. There are several additional features that are included in some expression vectors; these serve as tools for specialized functions (see Fig. 5.7.1 and Table 5.7.4). Sig is a DNA sequence juxtaposed between 5′PAOX1 and the MCS region; it encodes a protein secretion signal that is expressed as an N-terminal fusion to the protein of interest. This sequence directs the passage of the protein through the secretory pathway and targets proteins that carry it out of the cell. Sig sequences that have been used for expression in Pichia include the S. cerevisiae mating type α prepro sequence (MF-1α prepro; Scorer et al., 1993, 1994), the Pichia acid phosphatase signal sequence (PHO), the S. cerevisiae invertase signal sequence (Tschopp et al., 1987b), and several hybrid sequences (Larouche et al., 1994). Alternatively, the gene of interest may contain a sequence that encodes its own native secretion signal; that signal sequence may be functional in Pichia (Digan et al., 1989; Barr et al., 1992; Despreaux et al., 1993). Kanr is a bacterial kanamycin resistance gene; it may be included to confer resistance to the drug G418. Recombinant Pichia strains that contain multiple copies of Kanr are resistant to
Production of Recombinant Proteins
5.7.5 Current Protocols in Protein Science
Supplement 2
Table 5.7.4
Vectors for Heterologous Protein Expression in Pichia pastoris
Vector
Size (bp)
Selectable Secretion signal marker
pHIL-D2 pHIL-S1 pPIC3 pPIC3K pPIC9 pPIC9K pAO815
8157 8207 7768 9009 8034 9272 7837
HIS4 HIS4 HIS4 HIS4 HIS4 HIS4 HIS4
None Pichia PHO1 None None S. cerevisiae α factor S. cerevisiae α factor None
Multicopy feature
Type of fusion
Transcriptiona Translationb Transcriptiona Transcriptiona Translationb Kan genec (G418r) Translationb In vitro multimers Transcriptiona
None None None Kan genea (G418r)
aTranscription fusion: Vector does not supply an ATG initiation codon for translation of the open reading frame contained
within the expressed insert; insert must bring an in-frame ATG in a favorable eukaryotic context (i.e., Kozak consensus sequence); gene of interest carried on the DNA fragment inserted at the MCS of the vector is fused within the 5′ untranslated region of the mRNA expressed from the AOX1 promoter. bTranslation fusion: Vector supplies an ATG initiation codon to initiate translation of the secretion signal peptide destined to be expressed as an N-terminal fusion to the heterologous protein of interest; open reading frame contained within insert may or may not have an in-frame ATG; gene of interest carried on the DNA fragment inserted at the MCS of the vector is fused within the open reading frame region of the mRNA expressed from the AOX1 promoter. cKan gene: Bacterial Kan gene confers resistance to the drug G418; recombinant Pichia strains that contain multiple copies of the Kan gene can be resistant to higher concentrations of G418, with this resistance being dependent on Kan gene copy number.
higher concentrations of G418 in a manner that correlates with the Kanr copy number. The f1 ori sequence is a bacterial single-strand origin of replication; this sequence can be included to generate single-stranded DNA in Escherichia coli to facilitate plasmid mutagenesis.
Transformation by Integration
Overview of Protein Expression in Pichia pastoris
As in S. cerevisiae, linear DNA can generate stable transformants of P. pastoris via homologous recombination between the transforming DNA and regions of homology within the genome (Cregg et al., 1985, 1989). Such integrants show extreme stability in the absence of selective pressure, even when present in multiple copies. The most commonly used expression vectors carry the HIS4 gene for selection and are designed to be linearized with a restriction enzyme such that His+ recombinants can be generated by vector integration at the his4 or AOX1 locus (see Fig. 5.7.2). Integration events at the AOX1 locus can occur as either gene insertions or gene replacements. A gene replacement (omega insertion) event arises from a double cross-over between the AOX1 promoter and 3′AOX1 regions of the vector and genome; this results in the complete removal of the AOX1 coding region (i.e., gene replacement; Fig. 5.7.2). The resulting phenotype is His+ MutS. His+ transformants can be
readily and easily screened for their Mut phenotype, with MutS serving as a phenotypic indicator of integration via gene replacement at the AOX1 locus. Gene insertion events at the AOX1 locus arise from a single cross-over event between the AOX1 locus in the chromosome and any of the three AOX1 regions on the vector (AOX1 promoter, AOX1 TT, or 3′ AOX1); these events result in the insertion of one or more copies of the vector upstream or downstream of an otherwise intact and functional native AOX1 gene (see Fig. 5.7.3). The phenotype of such a transformant is His+ Mut+ (Mut+ is wild-type for methanol utilization); the His+ transformant is still able to utilize methanol because the AOX1 gene remains intact and functional (see Fig. 5.7.3). Gene insertion events at the his4 locus arise from a single cross-over event between the his4 locus in the chromosome and the HIS4 gene on the vector, and result in the insertion of one or more copies of the vector at the his4 locus (see Fig. 5.7.4). Because the genomic AOX1 locus is not involved in this recombination event, the phenotype of such a His+ transformant remains Mut+. Multiple gene insertion events at a single locus in a cell do occur spontaneously, with a low but detectable frequency—between 1% and 10% of all selected His+ transformants (see Fig. 5.7.5). Multicopy events can occur as gene
5.7.6 Supplement 2
Current Protocols in Protein Science
HIS4
TT gene of interest
linearized plasmid
5′ PAOX1
3′ AOX1
AOX1 gene Pichia genome 5′
ORF
3′
TT
gene of interest 5′ PAOX1
plasmid integrated into genome 3′ AOX1
HIS4
TT
Figure 5.7.2 Gene replacement. A gene replacement event at the AOX1 locus by a plasmid fragment carrying an “expression cassette” consisting of an AOX1 promoter–driven gene of interest and the HIS4 selectable marker allows for selection of His+ prototrophs using a his4 Pichia strain. The event represents a double cross-over between two linear DNA molecules. Double-stranded ends of DNA molecules act to target and stimulate homologous recombination. The result is a loss of the AOX1 locus (the strain becomes MutS) and the gain of PAOX1-gene of interest-HIS4 (gene of interest+ His+). Abbreviations: ORF, open reading frame; TT, transcriptional termination sequence.
gene of interest
TT
HIS4 5′ PAOX1 3′ AOX1
AOX1 gene Pichia genome 5′
AOX1 gene 5′
ORF
TT 3′
TT
ORF
3′
gene of interest 5′ PAOX1
TT
HIS4
3′ AOX1
expression cassette
Figure 5.7.3 Gene insertion. A gene insertion event 3′ of AOX1 in the genome by the plasmid carrying an expression cassette allows for selection of His+ prototrophs using a his4 Pichia strain. The event represents a single cross-over event between circular and linear DNA molecules. Recombination takes place within regions of homology between the plasmid and the genome. The result is an insertion of the plasmid 3′ to the intact AOX1 locus (the strain remains Mut+) and the gain of PAOX1-gene of interest-HIS4 (gene of interest+ His+). This event also could have taken place between the 5′ AOX1 regions on the plasmid and genome with the resulting insertion positioned 5′ to an intact AOX1 locus. Abbreviations: ORF, open reading frame; TT, transcriptional termination sequence.
Production of Recombinant Proteins
5.7.7 Current Protocols in Protein Science
Supplement 2
gene of interest
5′ AOX1 3′ AOX1
TT
HIS4
Pichia genome with his4 mutation
his4
gene of interest
HIS4
TT
5′ PAOX1
His+
3′ AOX1
his4 His–
Figure 5.7.4 Gene insertion in his4. A gene insertion event within the his4 locus of the genome by the plasmid carrying an expression cassette allows for selection of His+ prototrophs using a his4 Pichia strain. The event consists of a single cross-over between circular and linear DNA molecules, in which recombination takes place between the HIS4 locus on the plasmid and the his4 locus in the genome. The result is an insertion of the plasmid between duplicated copies of the HIS4/his4 genes, one still mutant and the other wild-type (the strain remains Mut+, and becomes gene of interest+ His+). Abbreviations: TT, transcriptional termination sequence.
Overview of Protein Expression in Pichia pastoris
insertions at either the AOX1 or his4 locus and can be detected by PCR analysis, Southern blot analysis, differential hybridization, or phenotypic identification based on the resulting hyperresistance to G418 (see Figs. 5.7.5 and 5.7.6). Three specific expression plasmids listed in Table 5.7.4 have been developed to facilitate the generation of multicopy expression cassette recombinant strains, pPIC3K, pPIC9K, and pAO815 (Cregg et al., 1993; Scorer et al., 1993, 1994). Expression plasmids that contain the bacterial Kanr gene act as a marker that can be used to identify phenotypically His+ recombinant strains which carry multiple copies of the gene of interest (see Fig. 5.7.6). Each expression cassette introduces a copy of the bacterial Kanr gene. This gene confers resistance to G418, and multiple copies of the gene make recombinant strains hyperresistant to the drug G418. Hyperresistance to G418 allows for phenotypic identification of strains carrying multiple copies of the Kanr gene and, by linkage, multiple copies of the expression cassette (PAOX1-gene of interest and HIS4; Scorer et al., 1993, 1994). Another plasmid system designed to gener-
ate recombinant strains that contain multiple copies of expression cassettes is pAO815 (Table 5.7.4). Multiple copies of PAOX1-gene of interest are ligated together in a plasmid in vitro—forced into a directional configuration by use of BamHI-BglII ligations—and linked to a single HIS4 gene in an expression cassette. Transformation of P. pastoris his4 strains with these in vitro–formed multimers is used as a tool to increase the frequency with which multiple copy expression cassette–recombinant strains are generated (Fig. 5.7.7; Thill et al., 1990; Cregg et al., 1993). The his4 mutant allele found in the strains listed in Table 5.7.2 does not contain a deletion mutation. Spontaneous reversion of the his4 mutant allele to His+ prototrophy is less than 10−8. However, apparent “hit-and-run” gene conversion events of his4 strains transformed with a HIS4 plasmid—that is, nonintegrative recombination between the his4 gene on the chromosome and the HIS4 gene on the plasmid that results in conversion of the chromosomal his4 gene to His+—occurs with a frequency of 10% to 50% of all His+ transformants. The His+ gene conversion frequency apparently can vary
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gene of interest
TT HIS4 5′ PAOX1
3′ AOX1 gene of interest
AOX1 gene 5′
ORF
second insertion event
AOX1 gene 5′
ORF
TT HIS4 Kanr 3′ AOX1 expression cassette
5′ PAOX1
TT 3′
TT
3′
gene of interest 5′ PAOX1
TT HIS4 3′ AOX1 expression cassette 1 gene of interest 5′ PAOX1
TT
HIS4 3′ AOX1
expression cassette 2 third and subsequent insertion events
Figure 5.7.5 Multiple insertion events. Multiple insertions of an expression plasmid 3′ to an otherwise intact AOX1 locus can result from multimer formation prior to a single insertion event or from multiple sequential insertion events at the same locus. These events can also take place between the 5′ AOX1 regions on the plasmid and genome, with the resulting insertions positioned 5′ to an intact AOX1 locus. Multiple events (two or more plasmids integrated) occur spontaneously, although infrequently, accounting for 1% to 10% of all His+ transformants. Abbreviations: ORF, open reading frame; TT, transcriptional termination sequence.
depending on the transformation method, with electroporation yielding a higher frequency of His+ gene conversion than spheroplast transformation (D.R.H., unpub. observ.). These His+ convertants that do not contain a copy of the gene to be expressed can be identified by PCR or Southern blot analysis and eliminated from subsequent protein-expression testing. Figure 5.7.8 outlines the steps required to generate and analyze recombinant strains of P. pastoris.
Post-Translational Modifications P. pastoris utilizes most of the post-translational modification pathways typically associ-
ated with higher eukaryotes. In comparison to S. cerevisiae, Pichia may have an advantage in the glycosylation of secreted proteins because it may have less of a tendency to hyperglycosylate. Although both S. cerevisiae and P. pastoris have a majority of N-linked glycosylation of the high-mannose type, the length of the oligosaccharide chains added post-translationally to proteins is much shorter in Pichia than in S. cerevisiae (an average 8 to 14 mannose residues per side chain in P. pastoris compared to 50 to 150 in S. cerevisiae; Tschopp et al., 1987b; Grinna and Tschopp, 1989). There is also a difference between S. cerevisiae and P. pastoris in the structure of the oligosaccharides added.
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gene of interest
TT
HIS4
Kanr
5′ PAOX1 3′ AOX1
5′
3′
ORF TT AOX1 gene
first insertion of vector
gene of interest 5′
TT 3′
ORF
5′ PAOX1
TT
AOX1 gene
HIS4
Kanr 3′ AOX1
single copy Kanr gene results in resistance to 0.25 mg/ml G418
gene of interest
TT
HIS4
second insertion event Kanr
5′ PAOX1
3′ AOX1
5′
3′
ORF TT AOX1 gene
gene of interest 5′ PAOX1
TT
HIS4
Kanr 3′ AOX1
expression cassette 1
gene of interest 5′ PAOX1
HIS4
TT
gene of interest
TT HIS4
Kanr
3′ AOX1
expression cassette 2
third and subsequent insertions of vector Kanr
5′ PAOX1
3′ AOX1
Overview of Protein Expression in Pichia pastoris
>10 copies of Kanr gene result in resistance to >4 mg/ml G418
Figure 5.7.6 Multiple insertions with a Kanr-containing plasmid. Multiple insertions of an expression plasmid 3′ to an otherwise intact AOX1 locus can result from multimer formation prior to a single insertion event or from multiple sequential insertion events at the same locus. These events can also occur between the 5′ AOX1 regions on the plasmid and genome, with the resulting insertions 5′ to an intact AOX1 locus. Each expression cassette introduces a copy of the bacterial Kanr gene, which confers resistance to G418; multiple copies make recombinant strains hyperresistant to G418, which phenotypically identifies strains carrying multiple copies of the expression cassette. Abbreviations: ORF, open reading frame; TT, transcriptional termination sequence.
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Bam HI TT
gene of interest
HIS4 5′ PAOX1 BgI II
3′ AOX1
BgI II
Bam HI
BgI II
5′ PAOX1 gene of interest
TT
Bam HI TT
gene of interest
HIS4 5′ PAOX1 BgI II 3′ AOX1
BgI II BgI II + BamHI digestion
BgI II
gene of interest
5′ PAOX1
BamHI/BgI II
TT
5′ PAOX1
gene of interest
BamHI
TT
expression cassettes
Figure 5.7.7 In vitro multimer formation. By this scheme, multiple copies of PAOX1-gene of interest are ligated together in a plasmid in vitro, and linked to a single HIS4 gene in an expression cassette. Transformation of Pichia with these in vitro–formed multimers is used as a tool to increase the frequency with which multiple-copy expression cassette recombinant strains are generated. Abbreviations: OEF, open reading frame; TT, transcriptional termination sequence.
S. cerevisiae core oligosaccharides have terminal α1-3 glycan linkages, whereas P. pastoris does not. Furthermore, it is believed that the α1-3 glycan linkages of S. cerevisiae are primarily responsible for the hyperantigenic nature of glycosylated proteins from S. cerevisiae
that makes them particularly unsuitable for therapeutic use. Although not yet proven, this is predicted to be less of a problem for glycoproteins generated in P. pastoris, because their glycoprotein structure may resemble that of higher eukaryotes (Cregg et al., 1993).
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generate recombinant strain: select expression vector construct gene of interest in vector select strain transform strain with construct
identify His+ recombinants
identify His+ recombinants that are hyperresistant to G418
confirm integration of gene of interest
generate biomass
perform Mut+ induction: pellet cells and resuspend in methonal medium sample at 0, 6, 12, 24, 36, and 48 hr
perform MutS induction: pellet cells and resuspend in methonal medium sample at 0, 24, 48, 72, 96, and 144 hr
analyze protein expression in medium and/or cell lysate using: PAGE with Coomassie staining (UNIT 10.1) immunoblotting ELISA immunoprecipitation
Figure 5.7.8 Flow chart for the generation of recombinant strains of Pichia pastoris.
Expression Examples
Overview of Protein Expression in Pichia pastoris
There are several parameters that can be varied to optimize protein expression levels using the Pichia system, including the strain (e.g., protease-deficient strains, Mut+ or MutS recombinant strains), intracellular or secreted expression, and flask or fermenter format. Table 5.7.1 lists published examples of heterologous protein expression in Pichia. Unpublished examples of proteins expressed in Pichia include MHC class II, shark cytochrome p450, muscle kinase, human serum proteins, maize heat shock protein, α-glycosidase, diphenol oxidase, porcine cytokines, antibody heavy and light chains, lysoxygenase, cytochrome c reductase, spinach glycolate oxidase, oncostatin M, lactoferrin, bovine rhodopsin, and trans-
membrane transferrin. Although expression— or very-high-level expression—of heterologous proteins in Pichia has not been successful in every case, there are sufficient examples of success to warrant inclusion of Pichia in the standard expression repertoire. In addition to the well-publicized examples of high-level expression (Tschopp et al., 1987a; Clare et al., 1991a; Barr et al., 1992), there are several examples that demonstrate Pichia’s behavior as a eukaryote capable of correct post-translational processing, including expression of correctly folded heterologous proteins that contain many disulfide bonds (White et al., 1994), proteins that are active when expressed in Pichia but not when expressed in E. coli (Yamada et al., 1994; Jenkins and Fayer, 1995; Steinlein
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et al., 1995; Vozza et al., 1995), and proteins for which Pichia properly recognized the native higher eukaryote secretion signal (Barr et al., 1992; Ridder et al., 1995). Levels of expression vary widely, as do the optimal conditions for expression.
Legal Issues Heterologous protein expression in Pichia pastoris is covered by issued patents that are owned and administered by Research Corporation Technologies (RCT). Invitrogen has the exclusive rights for the distribution of Pichia expression technology. The Pichia expression system (strains, vectors, and reagents) is available to all researchers simply by purchasing the Pichia Expression Kit from Invitrogen. Protein expression in Pichia, even for research purposes only, requires a “research license” that is automatically granted with the purchase of the kit from Invitrogen. A “commercial license” is required for expression intended for commercial purposes. The commercial license must be obtained from RCT after the kit is purchased. Questions regarding commercial licenses or the definition of commercial application should be addressed to RCT.
COMMENTARY Critical Parameters Successful expression—loosely defined as achieving expression levels that meet the users’ needs—can be viewed as a two-step process. The first step is to achieve expression at some level, even if suboptimal. Often successful expression is fully achieved in the first step. If the levels of expression obtained initially are not adequate, however, the second step is optimization of expression. Several parameters can be manipulated to optimize expression levels, including (1) expression as an intracellular protein versus a protein targeted for secretion; (2) expression in a Mut+ versus a MutS recombinant strain; (3) expression in a wild-type versus a protease-deficient mutant; (4) expression in recombinant strains with multiple copies of expression cassettes; and (5) optimization of medium conditions, including specific components (i.e., MM versus BMMY), pH, and expression scale (i.e., flasks versus fermentation). Tools have been developed that allow the researcher to manipulate these parameters and optimize expression. It is essential to keep optimization steps in mind when planning the initial expression experiments. Other critical tools required for success in-
clude good, well-defined methods to unambiguously detect heterologous protein expression. Simple visualization via polyacrylamide gel electrophoresis (PAGE; UNIT 10.1) with Coomassie or silver staining (UNIT 10.5) is ideal, but other, more sensitive tools—such as antibodies for immunoblots, ELISAs, or immunoprecipitations, or assays for an activity associated with the protein being expressed—are also very useful.
Troubleshooting Although the Pichia pastoris system is easy to use, it requires a number of media that may include components and procedures that are new to some labs. The proper recipes and components should be obtained and the media should be prepared well in advance of beginning the experiments (see UNIT 5.8). All the media are very stable and have long shelf lives, so it is feasible to prepare them in advance. There are several methods to transform P. pastoris (Ausubel et al., 1995, Chapter 13), although the spheroplast and electroporation methods are probably the most commonly used. Spheroplast transformation is a relatively straightforward procedure, but it may be the most difficult aspect for first-time users of the system to master. Gentle handling of the cells and meticulous attention to the protocol are required for success. Most other problems that are likely to arise in expression using P. pastoris involve protein expression levels, and are outlined in Table 5.7.5.
Anticipated Results As the number of expressed proteins in P. pastoris increases, more information is being generated to help define routes to successful expression. Based on available data, there is an ∼50% to 75% chance of expressing a protein of interest in P. pastoris at reasonable levels. The biggest hurdle seems to be generating initial success—that is, expression of the protein at any level. After success at this stage, there are many well-defined parameters that can be manipulated to achieve optimization of expression, and it is often at this stage that acceptable levels of expression are achieved. Although there have been relatively few instances of expression of ≥10 g/liter, there have been many cases of expression in the ≥1 g/liter range, ranking the P. pastoris expression system as one of the most productive eukaryotic expression systems available. Likewise, there are several proteins that have been successfully expressed
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Table 5.7.5
Troubleshooting Guide for Failure to Obtain Heterologous Protein
Possible cause of failure
Solution
Expression cassette not present
Demonstrate that recombinant strain contains the gene of interest by PCR or Southern blot hybridization
No transcription
Check for mRNA by RT-PCR or northern blot hybridization; check for AT-rich regions that may be functioning as transcription termination sites within coding region of expressed gene (see Scorer et al., 1993)
No translation of mRNA
Check sequence for possible secondary structure in 5′ untranslated region of mRNA Check codon usage and initiation context for gene of interest
Protein instability
Use protease-deficient strain(s) to prevent proteolysis Change medium or pH medium to optimal for protein
Secretion problem: incomplete secretion, retained in secretory pathway, secreted protein improperly processed, improper modification
Check cell pellet for protein Determine apparent MW of protein
Expression too low
Increase copy number of gene of interest by using pPIC3K, pPIC9K, or pAO815; or scale up fermentation Examine at earlier time points
Expressed protein is deleterious to cells
in P. pastoris following a complete lack of success in baculovirus or S. cerevisiae, making the P. pastoris system an important alternative in the protein expression toolbox.
Time Considerations
Overview of Protein Expression in Pichia pastoris
Numerous media are needed for recombinant strain generation and protein expression (Table 5.7.3); medium preparation takes about 1⁄ day and should be completed at least 1 day 2 prior to starting the transformation. The transformation procedure takes ∼4 hr with an overnight culture inoculated the day before. It takes 4 to 6 days for prototrophic transformants (e.g., His+) to form colonies. Screening of transformants to distinguish the His+ Mut+ from the His+ MutS takes an additional 3 days. Genomic DNA preps to generate template DNA for PCR analysis and PCR analysis itself can be completed in 1 day. Purification of mRNA for RTPCR or northern analysis takes ∼1 day. For protein expression experiments in flasks, growth of cultures in glycerol medium to generate biomass takes 1 to 2 days. Methanol induction and a time-course study of protein expression goes from 6 hr to 2 days postinduction for Mut+ recombinant strains and 1 to 6 days for MutS recombinant strains. Analysis of
expression by PAGE (UNIT 10.1), immunoblotting, ELISA, or specific assay usually requires one additional day. The total elapsed time from the first transformation to the first protein analysis is ∼3 to 4 weeks, with fairly continuous attention required. Commonly, the next phase involves optimization and scale-up once acceptable protein expression levels have been demonstrated. It is reasonable to expect that it will take 6 months from the first transformation to achieving medium- or large-scale production of acceptable levels of heterologous protein expression.
Literature Cited Ausubel, F.M., Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G., Smith, J.A., and Struhl, K. (eds.) 1995. Current Protocols in Molecular Biology. John Wiley & Sons, New York. Barr, K.A., Hopkins, S.A., and Sreekrishna, K. 1992. Protocol for efficient secretion of HSA developed from Pichia pastoris. Pharm. Eng. 12:48-51. Buckholz, R.G. and Gleeson, M.A.G. 1991. Yeast systems for the commercial production of heterologous proteins. Bio/Technology 9:10671072.
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Clare, J.J., Rayment, F.B., Ballantine, S.P., Sreekrishna, K., and Romanos, M.A. 1991a. Highlevel expression of tetanus toxin fragment C in Pichia pastoris strains containing multiple tandem integrations of the gene. Bio/Technology 9:455-460. Clare, J.J., Romanos, M.A., Rayment, F.B., Rowedder, J.E., Smith, M.A., Payne, M.M., Sreekrishna, K., and Henwood, C.A. 1991b. Production of epidermal growth factor in yeast: Highlevel secretion using Pichia pastoris strains containing multiple gene copies. Gene 105:205212. Cregg, J.M. and Madden, K.R. 1989. Use of sitespecific recombination to regenerate selectable markers. Mol. Gen. Genet. 219:320-323. Cregg, J.M., Barringer, K.J., Hessler, A.Y., and Madden, K.R. 1985. Pichia pastoris as a host system for transformations. Mol. Cell. Biol. 5:3376-3385. Cregg, J.M., Tschopp, J.F., Stillman, C., Siegel, R., Akong, M., Craig, W.S., Buckholz, R.G., Madden, K.R., Kellaris, P.A., Davis, G.R., Smiley, B.L., Cruze, J., Torregrossa, R., Velicelebi, G., and Thill, G.P. 1987. High-level expression and efficient assembly of hepatitis b surface antigen in the methylotrophic yeast Pichia pastoris. Bio/Technology 5:479-485. Cregg, J.M., Vedvick, T.S., and Raschke, W.C. 1993. Recent advances in the expression of foreign genes in Pichia pastoris. Bio/Technology 11:905-910. Despreaux, C.W. and Manning, R.F. 1993. The dacA gene of Bacillus stearothermophilus coding for D-alanine carboxypeptidase: Cloning, structure, and expression in Escherichia coli and Pichia pastoris. Gene 131:35-41. Digan, M.E., Lair, S.V., Brierley, R.A., Siegel, R.S., Williams, M.E., Ellis, S.B., Kellaris, P.A., Provow, S.A., Craig, W.S., Velicelebi, G., Harpold, M.M., and Thill, G.P. 1989. Continuous production of a novel lysozyme via secretion from the yeast Pichia pastoris. Bio/Technology 7:160164. Ellis, S.B., Brust, P.F., Koutz, P.J., Waters, A.F., Harpold, M.M., and Gingeras, T.R. 1985. Isolation of alcohol oxidase and two other methanol regulatable genes from the yeast Pichia pastoris. Mol. Cell. Biol. 5:1111-1121. Fryxell, K.B., O’Donoghue, K., Graeff, R.M., Lee, H.C., and Branton, W.D. 1995. Fucntional expression of soluble forms of human CD38 in Escherichia coli and Pichia pastoris. Protein Express. Purif. 6:329-336. Gould, S.J., McCollum, D., Spong, A.P., Heyman, J.A., and Subramani, S. 1992. Development of the yeast Pichia pastoris as a model organism for a genetic and molecular analysis of peroxisome assembly. Yeast 8:613-628. Grinna, L.S. and Tschopp, J.F. 1989. Size distribution and general structural features of N-linked oligosaccharides from the methylotrophic yeast Pichia pastoris. Yeast 5:107-115.
Guthrie, C. and Fink, G.R., eds. 1991. Guide to Yeast Genetics and Molecular Biology. Methods Enzymol.:194.. Hagenson, M.J., Holden, K.A., Parker, K.A., Wood, P.J., Cruze, J.A., Fuke, M., Hopkins, T.R., and Stroman, D.W. 1989. Expression of streptokinase in Pichia pastoris yeast. Enzyme Microbiol. Technol. 11:650-656. Halaas, J.L., Gajiwala, K.S., Maffei, M., Cohen, S.L., Chait, B.T., Rabinowitz, D., Lallone, R.L., Burley, S.K., and Friedman, J.M. 1995. Weightreducing effects of the plasma protein encoded by the obese gene. Science 269:543-546. Howard, P.W. and Maurer, R.A. 1995. A composite Ets/Pit-1 binding site in the prolactin gene can mediate transcriptional responses to multiple signal transduction pathways. J. Biol. Chem. 270:20930-20936. Jenkins, M.C. and Fayer, R. 1995. Cloning and expression of a cDNA encoding an antigenic Cryptosporidium parvum protein. Mol. Biochem. Parasitol. 71:149-152. Koutz, P.J., Davis, G.R., Stillman, C., Barringer, K., Cregg, J.M., and Thill, G. 1989. Structural comparison of the Pichia pastoris alcohol oxidase genes. Yeast 5:167-177. Larouche, Y., Storme, V., De Meutter, J., Messens, J., and Lauwereys, M. 1994. High-level secretion and very efficient isotopic labeling of tick anticoagulant peptide (TAP) expressed in the methylotrophic yeast, Pichia pastoris. Bio/Technology 12:1119-1124. Ohsawa, I., Hirose, Y., Ishiguro, M., Imai, Y., Ishiura, S., and Kohsaka, S. 1995. Expression, purification, and neurotrophic activity of amyloid precursor protein–secreted forms produced by yeast. Biochem. Biophys. Res. Commun. 213:5258. Paifer, E., Margolies, E. Cremata, J., Montesino, R., Herrera, L, and Delgado, J.M. 1994. Efficient expression and secretion of recombinant alpha amylase in Pichia pastoris using two different signal sequences. Yeast 10:1415-1419. Ratner, M. 1989. Protein expression in yeast. Bio/Technology 7:1129-1133. Reddy., R.G., Yoshimoto, T. Yamamoto, S., and Marnett, L. 1994. Expression, purification, and characterization of porcine leukocyte 12 lipoxygenase produced in methylotrophic yeast, Pichia pastoris. Biochem. Biophys. Res. Commun. 205:381-388. Reddy, R., Schmitz, R., Legay, F., and Gram, H. 1995. Generation of rabbit monoclonal antibody fragments from a combinatorial phage display library and their production in the yeast Pichia pastoris. Bio/Technology 13:255-260. Ridder, R., Schmitz, R., Legay, F., and Gram, H. 1995. Generation of rabbit monoclonal antibody fragments from a combinatorial phage display library and their production in the yeast Pichia pastoris. Bio/Technology 13:255-260. Rodriguez, M., Rubiera, R., Penichet, M., Montesinos, R., Cremata, J., Falcon, V., Sanchez, G.,
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Bringas, R., Cordoves, C., Valdes, M., Lieonart, R., Herrera, L. and de la Fuente, J. 1994. Higlevel expression of the b. microplus Bm86 antigen in the yeast Pichia pastoris forming highly immunogenic particles for cattle. J. Biotechnol. 33:135-146. Romanos, M.A., Clare, J.J., Beesley, K.M., Rayment, F.B., Ballantine, S.P., Makoff, A.J., Dougan, G., Fairweather, N.F., and Charles, I.G. 1991. Recombinant Bordetella pertussis pertactin (P69) from the yeast Pichia pastoris: High-level production and immunological properties. Vaccine 9:901-906. Scorer, C.A., Buckholz, R.G., Clare, J.J., and Romanos, M.A. 1993. The intracellular production and secretion of HIV-1 envelope protein in the methylotrophic yeast Pichia pastoris. Gene 136:111-119. Scorer, C.A., Clare, J.J., McCombie, W.R., Romanos, M.A., and Sreekrishna, K. 1994. Rapid selection using G418 of high-copy-number transformants of Pichia pastoris for high-level foreign gene expression. Bio/Technology 12:181-184. Shiba, K., Ripmaster, T., Suzuki, N., Nichols, R., Plotz, P., Noda, T. and Schimmel, P. 1995. Human alanyl-tRNA synthetase: Conservation in evolution of catalytic core and microhelix recognition. Biochemistry 34:10340-10349. Sreekrishna, K., Nelles, L., Potenz, R., Cruze, J., Mazzaferro, P., Fish, W., Fuke, M., Holden, K., Phelps, D., Wood, P., and Parker, K. 1989. Highlevel expression purification and characterization of recombinant human tumor necrosis factor synthesized in the methylotrophic yeast Pichia pastoris. Biochemistry 28:4117-4125. Steinlein, L.M., Graf, T.N., and Ikeda, R.A. 1995. Production and purification of N-terminal halftransferrin in Pichia pastoris. Protein Express. Purif. 6-619-624. Thill, G.P., David, G.R., Stillman, C., Holtz, G., Brierly, R., Engel, M., Buckholz, R., Kenney, J., Provow, S., Vedvick, T., and Siegel, R.S. 1990. Positive and negative effects of multi-copy integrated expression vectors on protein expression in Pichia pastoris. In Proceedings of the 6th International Symposium on Genetics of Microorganisms, Vol. II (H. Heslot, J. Davies, J. Florent, L. Bobichon, G. Durand, and L. Penasse, eds.) pp. 477-490. Société Française de Microbiologie, Paris. Tschopp, J.F., Brust, P.F., Cregg, J.M., Stillman, C., and Gingeras, T.R. 1987a. Expression of the lacZ gene from two methanol-regulated promoters in Pichia pastoris. Nucl. Acids Res. 15:3859-3876. Tschopp, J.F., Sverlow, G., Kosson, R., Craig, W., and Grinna, L. 1987b. High-level secretion of glycosylated invertase in the methylotrophic yeast Pichia pastoris. Bio/Technology 5:13051308. Overview of Protein Expression in Pichia pastoris
Van Nostrand, W.E., Schmaier, A.H., Neiditch, B.R., Siegel, R.S., Raschke, W.C., Sisodia, S.S., and Wagner, S.L. 1994. Expression, purification, and characterization of the Kunitz-type proteinase
inhibitor domain of the amyloid β-protein precursor-like protein-2. Biochim. Biophys. Acta 1209:165-170. Vedvick, T., Buckholz, R.G., Engel, M., Urcan, M., Kinney, J., Provow, S., Siegel, R.S., and Thill, G.P. 1991. High-level secretion of biologically active aprotonin from the yeast Pichia pastoris. J. Ind. Microbiol. 7:197-201. Vozza, L., Wittwer, L., Higgins, D.R., Purcell, T.J., Bergseid, M., Collins-Racie, L.A., LaVallie, E.R., and Hoeffler, J.P. 1995. Production of a recombinant bovine enterokinase catalytic subunit in the methylotrophic yeast Pichia pastoris. Bio/Technology. Wagner, S.L., Siegel, R.S., Vedvick, T.S., Raschke, W.C., and Van Nostrand, W.E. 1992. High-level expression purification and characterization of the Kunitz-type protease inhibitor domain of protease nixin-2/amyloid β-protein precursor. Biochem. Biophys. Res. Commun. 186:11381145. Wegner, G.H. 1990. Emerging applications of the methylotrophic yeasts. FEMS Microbiol. Rev. 87:279-283. Weiss, S., Famulok, M., Edenhofer, F., Wang, Y.H., Jones, I.M., Groschup, M., and Winnacker, E.L. 1995. Overexpression of active Syrian golden hamster prion protein PrPc as a glutathione-Stransferase fusion in heterologous systems. J. Virol. 69:4776-4783. White, C.E., Kempi, N.M., and Komives, E.A. 1994. Expression of higly disulfide-bonded proteins in Pichia pastoris. Structure 2:1003-1005. Yamada, M., Azuma, T., Matsuba, T., Iida, H., Suzuki, H., Yamamoto, K., Kohli, Y., and Hori, H. 1994. Secretion of human intracellular aspartic proteinase cathepsin E expressed in the methylotrophic yeast Pichia pastoris and characterization of produced recombinant cathepsin E. Biochim. Biophys. Acta 1206:279-285.
Key References Cregg et al., 1993. See above. Good overview of Pichia pastoris as an expression system with a discussion of glycosylation of expressed proteins. Scorer et al., 1993. See above. Describes in detail the advantages of intracellular versus secreted expression, the effect of multiplecopy expression cassettes on expression levels, and an overall approach to optimizing expression levels. Scorer et al., 1994. See above. Details the generation of multiple-copy expression cassette recombinant strains using the bacterial Kanr gene and screening for hyperresistance to G418.
Contributed by David R. Higgins Invitrogen Corporation San Diego, California
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Culture of Yeast for the Production of Heterologous Proteins
UNIT 5.8
This unit describes culture of the yeast strains Saccharomyces cerevisiae and Pichia pastoris for the production of foreign proteins. The theoretical background to foreign gene expression in yeast is discussed in the previous units (UNITS 5.6 & 5.7). General techniques for handling yeast are described in Ausubel et al. (1995), Chapter 13, and will not be covered here. S. cerevisiae expression systems are generally based on the yeast 2µm plasmid, but they differ in choice of promoter and selectable marker, which affect the culture medium and induction conditions that must be used. Vectors with auxotrophic selection markers must be used in appropriate strains (e.g., LEU2 plasmids in leu2 strains), and growth must be in appropriate selective medium, i.e., minimal medium containing all the necessary supplements. Vectors with dominant selectable markers (e.g., G418r) can be used in any host strain and selected in rich medium, giving higher growth rate and cell density. The protocols listed here are for three widely used types of promoter: galactose-regulated (GAL1, GAL7, GAL10; Basic Protocol 1), glucose-repressible (e.g., ADH2; Alternate Protocol 1), and constitutive glycolytic (e.g., PGK or GAPDH; Alternate Protocol 2). Minor variations to each can be made depending on the selection system used. The P. pastoris expression system uses integrating vectors with the methanol-regulated AOX1 promoter and HIS4 selection marker; although transformants are stable, they are generally grown in minimal selective medium. In most cases, initial analysis is carried out with protein produced in small-scale (5-ml to 1-liter) cultures, and this may be sufficient in cases where the yield is high. Where larger amounts of protein are required, cultures can be grown to high cell density using fermenters. Methods are described for small-scale S. cerevisiae (Basic Protocol 1) and P. pastoris cultures (Basic Protocol 2) and also for high-density fermentations with these yeasts (Basic Protocols 3 and 4). High-density (>100 g/liter dry weight) growth of S. cerevisiae requires computer-controlled carbon source feeding based on continual monitoring of growth parameters (Wang et al., 1979; Fieschko et al., 1987). The protocol given here applies a much simpler feeding strategy based on calculated feed rates (Basic Protocol 3) and yields cell densities of 10 to 30 g/liter (Aiba et al., 1976; Hsieh et al., 1988; Bock Gu et al., 1989). In contrast, with P. pastoris, it is simple to obtain extremely high-density cultures (e.g., 130 g/liter) using basic fermenter equipment (Basic Protocol 4). Finally, Support Protocol 1 describes small-scale preparation of protein extracts. NOTE: All solutions and equipment coming into contact with cells must be sterile, and proper sterile technique should be used accordingly. SMALL-SCALE EXPRESSION USING S. CEREVISIAE GALACTOSE-REGULATED VECTORS Galactose-regulated (GAL) promoters are induced >1000-fold by addition of galactose but are strongly repressed by glucose, so that in glucose-grown cultures maximal induction can be achieved only following depletion of glucose. For rapid induction, cells can be grown using a nonrepressing carbon source (e.g., raffinose), then induced by direct addition of galactose. First, saturated overnight starter cultures are prepared, and these are used to inoculate fresh medium. These secondary cultures are grown for two doublings (4 to 5 hr including growth lag) to late exponential pase, then induced by direct addition of galactose. Cells are then harvested by centrifugation and may be stored at −70°C as dry pellets or disrupted immediately, using glass beads, to prepare a total protein extract. Contributed by Michael A. Romanos and Jeffrey J. Clare Current Protocols in Protein Science (1995) 5.8.1-5.8.17 Copyright © 1997 by John Wiley & Sons, Inc.
BASIC PROTOCOL 1
Production of Recombinant Proteins
5.8.1 Supplement 2
With either very stable vectors, or vectors having dominant selectable markers, growth and induction can be carried out in rich medium to maximize cell density. Where the plasmid requires auxotrophic selection, minimal selective medium must be used at least for the starter culture, although the induction itself can still usually be carried out in rich medium. The protocol described below is applicable to culture volumes of 5 to 1000 ml. Materials YP broth (see recipe) containing 2% (w/v) raffinose or YNB medium (see recipe) for auxotrophic selection 50 mg/ml G418 (100×) or supplements for auxotrophic selection, as required Saccharomyces cerevisiae transformed with a galactose-regulated expression vector and plated on selective medium (see UNIT 5.6 and Ausubel et al., 1995, Chapter 13) 20% (w/v) galactose (filter sterilize and store at room temperature) H2O, sterile Plastic 25-ml universal containers, sterile (e.g., Nunc) Heraeus omnifuge centrifuge and 2250 rotor (or equivalent) 15-ml polypropylene snap-top tubes Centricon concentrator (Amicon; also see UNIT 4.4) 1. Place 10 ml YP broth containing 2% raffinose in a sterile 25-ml universal container and inoculate with cells of a single colony of transformed S. cerevisiae. Incubate overnight at 30°C with shaking. As expression levels may vary between individual transformants, it may be advisable to test several when looking for optimal yield. For vectors with G418-resistance markers, add G418 (0.5 mg/ml final) to the raffinose preinduction medium. For unstable vectors with auxotrophic markers, use YNB medium (see recipe) with 2% raffinose and the appropriate supplements (cell density will be up to 5-fold lower). Glucose can be used in place of raffinose in the preinduction growth stages, especially where it is necessary to maintain tight repression of expression, e.g., in expressing toxic proteins. However, excess glucose must be removed by centrifuging and washing the cells in sterile saline prior to resuspending them in induction medium. For glucose-grown cultures, induction will have a several-hour lag period, during which the glucose is exhausted.
2. Measure the A600. The A600 of an overnight culture should be ∼5.0.
3. Dilute the overnight culture into 10 ml YP broth containing 2% raffinose to an A600 of 0.25 (typically a 20-fold dilution). Incubate at 30°C with shaking until the culture density reaches an A600 of ∼1.0 (4 to 5 hr). 4. Add 1 ml of 20% galactose to induce the culture. Incubate 8 to 24 hr (e.g., overnight) with shaking at 30°C. 5. Measure the A600 of the culture. The A600 of the induced cultures should be 10 to 15.
Culture of Yeast for the Production of Heterologous Proteins
6a. For intracellular protein expression: Harvest the cells by centrifuging 5 min at 2000 × g, 4°C. Resuspend the cell pellet in 10 ml water, transfer to a 15-ml snap-top tube, and centrifuge again as before. Remove excess water from the pellet. Store at −70°C. 6b. For secreted protein expression: Harvest the cells by centrifuging 5 min at 2000 × g, 4°C. Collect the medium (supernatant) and concentrate proteins in the medium by
5.8.2 Supplement 2
Current Protocols in Protein Science
ultrafiltration using a Centricon concentrator (Amicon). Store the protein concentrate at −70°C. Retain the cell pellet for analysis. Several secreted proteins have been found to accumulate to lower levels in minimal medium than in rich medium. Yields in minimal medium have been improved by adding Casamino acids (see recipe) or by buffering to pH 6.0 using potassium phosphate (unbuffered media will reach pH ∼4). To concentrate secreted proteins further, concentrate the cells 20-fold by centrifugation prior to induction.
EXPRESSION USING GLUCOSE-REPRESSIBLE ADH2 VECTORS The ADH2 promoter is repressed 100-fold by glucose. To maintain repression, cells must always be grown in excess glucose—e.g., 8% (w/v)—because normal concentrations (e.g., 2%) can become depleted. It is also preferable to store transformants on plates containing 8% glucose. Inductions can be carried out by first growing cells in medium containing excess glucose, then inducing by changing to fresh medium containing a nonfermentable carbon source such as raffinose.
ALTERNATE PROTOCOL 1
Additional Materials (also see Basic Protocol 1) YP broth (see recipe) containing 8% (w/v) glucose Saccharomyces cerevisiae transformed with a glucose-repressible ADH2 expression vector and plated on selective medium (see UNIT 5.6 and Ausubel et al., 1995, Chapter 13) YP broth (see recipe) containing either 2% (w/v) raffinose and 2% (v/v) ethanol or 1% (w/v) glucose 100-ml Erlenmeyer flask, sterile Diastix glucose test strips (Bayer Diagnostics; optional) 1. Prepare a 10-ml exponential phase culture of the S. cerevisiae transformant in YP broth containing 8% glucose (see Basic Protocol 1, steps 1 to 3). Depending on the vector characteristics, selective medium may need to be used, as in Basic Protocol 1.
2. Harvest the cells by centrifuging 5 min at 2000 × g, 20°C. Resuspend the pellet in 10 ml sterile water. 3. Centrifuge again as in step 3 and resuspend the pellet in 10 ml YP broth containing 2% raffinose/2% ethanol. 4. Incubate 8 to 24 hr (e.g., overnight) at 30°C under conditions of maximum aeration (e.g., by incubating in a 100-ml Erlenmeyer flask with vigorous agitation). Induction starts after a lag of several hours. As an alternative to steps 3 and 4, use the overnight starter culture in 8% glucose to inoculate YP broth containing 1% glucose to a starting A600 of 0.05 (typically a 100-fold dilution). Incubate 18 to 24 hr at 30°C with vigorous aeration. Monitor glucose depletion using Diastix glucose test strips if possible.
5. Measure the A600. 6. Harvest the cells by centrifuging 5 min at 2000 × g, 4°C. 7. Analyze the cell pellet for proteins expressed intracellularly and analyze the cell pellet and the medium (supernatant) for secreted proteins.
Production of Recombinant Proteins
5.8.3 Current Protocols in Protein Science
Supplement 8
ALTERNATE PROTOCOL 2
EXPRESSION USING VECTORS WITH GLYCOLYTIC GENE PROMOTERS A number of commonly used expression vectors have constitutive promoters from glycolytic genes (e.g., PGK, GAPDH, and ADH1). Because instability problems are more frequent with constitutive vectors, it is recommended that selective media be used throughout. Materials (also see Basic Protocol 1) YNB medium (see recipe) containing 2% (w/v) glucose and the required supplements Saccharomyces cerevisiae transformed with an expression vector containing a glycolytic gene promotor and plated on selective medium (see UNIT 5.6 and Ausubel et al., 1995, Chapter 13) 1. Prepare an overnight starter culture of the S. cerevisiae transformant in YNB medium containing 2% glucose and required supplements (see Basic Protocol, step 1). Doubling time for the transformed strain will usually be longer than for the untransformed strain. Constitutive high-level expression using vectors that contain glycolytic gene promotors causes a metabolic burden on the cell that usually results in reduced growth rates. The degree of the effect will depend on the gene expressed, but doubling times may increase to 3 or 4 hr, or greater in extreme cases. Very long doubling times are diagnostic of toxicity, and indicate that a regulated vector should be used.
2. Measure the A600. The A600 of the overnight culture should be 5.
3. Dilute the overnight culture into 10 ml fresh supplemented YNB medium to an A600 of 0.0025. Using medium with a higher glucose concentration—e.g., 8% (w/v)—may improve expression levels, because glycolytic promoters are partly glucose-inducible.
4. Incubate at 30°C with shaking until the A600 reaches 0.25 to 0.5—18 to 24 hr, depending on the foreign protein. 5. Harvest the cells for analysis by centrifuging 5 min at 2000 × g, 4°C. 6. Analyze the cell pellet for proteins expressed intracellularly and analyze the cell pellet and the medium (supernatant) for secreted proteins. BASIC PROTOCOL 2
SMALL-SCALE EXPRESSION IN PICHIA PASTORIS Small-scale cultures of Pichia pastoris are used for the initial analysis of expressed proteins and for monitoring established expression strains. As different transformants may vary widely in product yield, it may be necessary to screen a significant number (see Critical Parameters and UNIT 5.7). Aeration is a key consideration because the efficiency of induction is reduced when oxygen is limited. To avoid this, incubate with vigorous shaking, keep the culture volume to a minimum (e.g., 5 to 10 ml) and use large flasks (e.g., >100 ml, preferably baffled) with loose or permeable covers. Some evaporation may occur, and during extended inductions (>1 day) it may be necessary to add further aliquots of methanol. The optimal induction period may vary for different proteins, and it is recommended that a time course be carried out to maximize yields.
Culture of Yeast for the Production of Heterologous Proteins
5.8.4 Supplement 8
Current Protocols in Protein Science
Materials YNB medium (see recipe) containing 2% (w/v) glycerol Pichia pastoris transformed with an expression vector and plated on selective medium (see UNIT 5.7 and Ausubel et al., 1995, Chapter 13) 0.002 mg/ml biotin (500×; filter sterilize and store <1 year at 4°C) H2O, sterile YNB medium (see recipe) containing 1% (v/v) methanol Plastic 25-ml universal containers, sterile Heraeus omnifuge centrifuge and 2250 rotor (or equivalent) 100-ml Erlenmeyer flasks, sterile 15-ml polypropylene snap-top tubes 1. Place 10 ml YNB broth containing 2% glycerol in a sterile 25-ml universal container and inoculate with cells of a single colony of transformed P. pastoris. Incubate overnight at 30°C with shaking. 2. Measure the A600. The A600 of the overnight culture should be ∼5.
3. Dilute the overnight culture into 10 ml YNB medium containing 2% glycerol to an A600 of 0.25. Incubate 6 to 8 hr at 30°C with vigorous shaking to obtain an exponentially growing culture. 4. Collect the cells by centrifuging 5 min at 2000 × g, 4°C. Resuspend the cell pellet in 10 ml sterile water. 5. Pellet the cells by centrifuging 5 min at 2000 × g, 4°C. Resuspend the cell pellet in 10 ml YNB medium containing 1% (v/v) methanol. Incubate 1 to 5 days at 30°C with maximum aeration (e.g., incubate culture in a 100-ml Erlenmeyer flask with vigorous shaking). If inducing >1 day, add an additional 0.5% methanol on the second day. The final density of the culture should be A600 = 5 to 10. The concentration of a secreted product in the medium is generally proportional to the density of the culture. Thus, to increase the concentration of a secreted protein, grow a larger culture (e.g., 100 ml) and concentrate the cells prior to the induction by resuspending in 5 ml of the methanol-containing induction medium. In addition, the yield of some secreted proteins may be reduced by extracellular proteases. This can sometimes be ameliorated by altering the composition of the medium, for example by buffering the pH or by adding amino acid–rich supplements such as Casamino acids (see recipe) or peptone (Clare et al., 1991b; Cregg et al., 1993).
6. Collect the cells by centrifuging 5 min at 2000 × g, 4°C. Resuspend the cell pellet in 10 ml water. If the expressed protein is secreted, retain the supernatant, concentrating the protein by ultrafiltration (see Basic Protocol 1, step 6b). 7. Transfer the cell suspension to a 15-ml polypropylene snap-top tube. Centrifuge 5 min at 2000 × g, 4°C. Remove excess water from the pellet and store the washed cell pellet at −70°C. 8. Analyze the cell pellet for intracellularly expressed protein and analyze the cell pellet and the supernatant for secreted protein.
Production of Recombinant Proteins
5.8.5 Current Protocols in Protein Science
Supplement 2
BASIC PROTOCOL 3
LARGE-SCALE EXPRESSION USING S. CEREVISIAE GALACTOSE-REGULATED VECTORS High-density fermentation of S. cerevisiae is problematic because the yeast has a limited capacity to metabolize glucose oxidatively; ethanol accumulates, which eventually inhibits growth to a specific growth rate of 0.2 to 0.25 hr−1 (Sonnlietner and Kappeli, 1986). Additionally, glucose concentrations above ∼0.1% repress respiration. To prevent the conversion of glucose to ethanol and to maximize biomass, S. cerevisiae is grown using a fed-batch process: glucose is slowly fed to the fermenter, the feed rate increasing as the cell density increases, so as to obtain an almost constant growth rate (µ). The growth rate above which ethanol is formed (µcrit), and hence the critical feed rate, are determined by both the strain and the growth medium. Figure 5.8.1 shows the relationship between the growth rate and biomass yield, and the effect of exceeding the µcrit (Pirt, 1985). The following simplified protocol uses calculated feed rates. Materials YNB medium (see recipe) containing 1% (w/v) glucose and appropriate supplements Saccharomyces cerevisiae transformed with a galactose-regulated expression vector and plated on selective medium (see UNIT 5.6 and Ausubel et al., 1995, Chapter 13) BSSM fermenter medium (see recipe) 20% (w/v) glucose 20% (w/v) galactose 3 M phosphoric acid 3 M sodium hydroxide Antifoam (Dow Corning M30 antifoam) 250-ml and 2-liter Erlenmeyer flasks, sterile Benchtop fermenter, 5-liter total volume, with data logging and MENTOR control system (Life Science Laboratories) Electronic balance (Mettler) with RS232 communications Sorvall RC-3B centrifuge and H6000A rotor (or equivalent)
Yield (g protein/g glucose)
0.5
0.4
0.3
0.2
0.1
0
Culture of Yeast for the Production of Heterologous Proteins
0
0.1
0.2
0.3
0.4
0.5
Growth rate (liter/hr)
Figure 5.8.1 Effect of growth rate on yield.
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Prepare the cultures 1. Place 50 ml YNB medium containing 1% glucose and supplements in a sterile 250-ml Erlenmeyer flask and inoculate with cells from a single colony of transformed S. cerevisiae. Incubate overnight at 30°C with shaking. 2. Measure the A600. The A600 of the overnight culture should be ∼4.0.
3. Dilute 10 ml of the overnight culture into 250 ml YNB medium containing 1% glucose in a 2-liter shaker flask. Incubate 18 to 24 hr at 30°C with shaking (150 rpm). 4. Measure the A600. The A600 should be ∼4.0.
Calculate the feed rate 5. Determine the feed rate for the fermenter using the following equation (see Table 5.8.1 for definitions of terms) for a 20% glucose solution, where C1 = (X0 × V × ρ)/(Y × Sf). cumulative weight of feed = (C1 × eµt) − C1
Based on the approximate values for a 5-liter fermentation protocol given in Table 5.8.1, C1 = 2.7, and the feeding algorithm reduces to: cumulative weight of feed = (2.7 × e0.16t) − 2.7
6. Determine the feed profile by calculating the cumulative weight of feed for t = 1, 2,…, n hr. Enter these values into the MENTOR computer system. Express the protein 7. Place the fermenter reservoir containing 20% glucose feed on an electronic balance linked to MENTOR, which will control the feed rate via a dosing pump. The fermentation consists of two phases: a glucose fed-batch phase followed by batch addition of galactose to induce expression.
Table 5.8.1
Variable
Variables for Calculating S. cerevisiae Feeding Algorithm
Definition
Value for 5-liter protocol
Xo
Biomass in fermenter at start of feeding (g/liter)
0.07
V
Working volume of fermenter (liter)
3.5
Y
Yield coefficient (g cells per g glucose)
0.45
Sf µ
Glucose concentration in feedstock (g/liter) Target growth rate (hr−1)
ρ
Feed density (g/liter)
200 0.16 1000
Comment Derived by multiplying the starting A600 in the fermenter by 0.22 (assume shaker flask inoculum has A600 = 4) Maximum working volume Theoretical value is 0.5, but a more conservative value of 0.45 is recommended — Suggested value (see text for optimization) Approximation is sufficient
Production of Recombinant Proteins
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8. Add 2.5 liters BSSM fermentation medium to the fermenter. Then add the culture from step 2 to the fermenter and initiate feeding with 20% glucose. Feed for 28 hr. Monitor the A600 closely during feeding; if the growth rate drops below the target (ì = 0.16 hr−1, stop glucose feeding early. The growth rate selected in this protocol serves as a useful starting point for any strain. To optimize the protocol, determine the maximum growth rate (ìmax) for the strain in shaker-flask cultures, then calculate the feed rate, setting the desired growth rate at ì = 0.5ìmax. This should be safely below ìcrit; if necessary, increase ì from this value. If the feed rate is too high, glucose or ethanol will be detected in the fermentation and biomass yield will be reduced. The effect of altering ì on the feeding profile is shown in Figure 5.8.2.
9. Induce the culture by adding 250 ml of 20% galactose. Incubate the culture 24 hr. If glucose has accumulated, then induction will be delayed for several hours. Higher expression levels have been reported when 1% (w/v) galactose is pulsed in during the glucose feeding stage, and the glucose feed is then replaced with a galactose feed (Fieschko et al. 1987). The duration of induction should be optimized by determining the kinetics of product formation. Harvesting shortly before the culture stops growing may help minimize proteolytic degradation of the product. Cells in stationary phase are more resistant to mechanical breakage.
10. Harvest the culture after induction by centrifuging 10 min at 7000 × g in a Sorvall RC-3B centrifuge with H6000A rotor. 11. Analyze the cell pellet for intracellularly expressed protein and analyze the cell pellet and the supernatant for secreted protein. LARGE-SCALE EXPRESSION USING PICHIA PASTORIS Optimal expression in P. pastoris requires the use of fermenters, which increases both the efficiency of induction and the culture density. The method is simple, requiring only a basic fermenter with monitors and controls for pH, dissolved O2, stirring speed, temperature, and air flow. Cultures are grown in a defined medium containing glycerol. The pH is maintained by the addition of ammonium hydroxide, which also serves as the nitrogen source. Dissolved O2 must be maintained at >20% for optimal growth. Fermentations are carried out in three stages: an initial batch growth phase, a glycerol-limited phase during which the culture reaches high cell density, and, finally, a methanol-induction phase of
Cumulative weight of feed added (g)
BASIC PROTOCOL 4
Culture of Yeast for the Production of Heterologous Proteins
1600 0.22 0.20 0.18 0.16 0.13 0.11 0.09
1400 1200 1000 800 600 400 200
0 0
5
10
15
20
25
30
Time (hr)
Figure 5.8.2 Effect of growth rate on feed profile.
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several days during which product accumulates. The protocol given below is for a 1-liter “bench-top” fermenter, but very similar conditions have been used for cultures up to 240 liters (Cregg et al., 1987). Materials YNB medium (see recipe) containing 2% (v/v) glycerol Pichia pastoris transformed with an expression vector and plated on selective medium (see UNIT 5.7 and Ausubel et al., 1995, Chapter 13) 5× basal salts solution (see recipe) PTM1 solution (see recipe) Glycerol/PTM1: 12 ml PTM1 solution/liter 50% (v/v) glycerol 50% (v/v) ammonium hydroxide Methanol/PTM1: 12 ml PTM1 solution/liter 100% methanol Fermenter equipped with monitors or controls for pH, temperature, dissolved O2, airflow, and stirring speed Erlenmeyer flask, sterile 1. Place 10 to 50 ml YNB medium containing 2% glycerol in an Erlenmeyer flask and inoculate with cells of a single colony of transformed P. pastoris. Incubate overnight at 30°C with shaking. 2. Add 1 liter of 5× basal salt solution, 4 ml PTM1 solution, and 5% (v/v) glycerol to the fermenter and mix well. 3. Add the overnight culture to the fermenter. Incubate at 30°C until the glycerol is exhausted (∼24 to 30 hr). Adjust the aeration and agitation to maintain dissolved O2 at >20%, and maintain the pH at 5.0 with 50% ammonium hydroxide. The dissolved O2 level will begin to rise sharply when the glycerol becomes exhausted.
4. Initiate a limited glycerol/PTM1 feed at 12 ml/hr and continue for 4 to 24 hr, maintaining pH, temperature, and dissolved O2. During this period the culture should grow to a density of 30 to 100 g dry weight/liter. Cell density can be monitored by measuring A600; 1 A600 unit = ∼0.33 g/liter, although different spectrophotometers will vary and each should be calibrated individually. To achieve optimal yield, the duration of this glycerol-limited phase, and thus the cell density at induction, can be varied depending on the type of product and the Mut phenotype of the host (see UNIT 5.7). Mut+ strains can be induced at low cell density because they continue to grow during induction. MutS strains should be allowed to achieve high density before induction. The yield of secreted proteins may be improved by early induction, allowing secretion to occur during a period of active growth.
5. Induce the culture by replacing the glycerol feed with a methanol/PTM1 feed at the rate of 1 ml/hr. Gradually increase the methanol feed rate over a period of 6 hr to 6 to 10 ml/hr and continue fermentation for an additional 46 to 92 hr. For efficient induction of Mut+ strains, the aim is to achieve maximal methanol feed rates (after an initial period to adapt the culture to methanol utilization) while maintaining sufficient aeration. If dissolved oxygen levels cannot be maintained >20%, the methanol feed rate should be reduced. This will also prevent accumulation of toxic levels of methanol. To avoid this problem with MutS strains, the methanol feed rate should be reduced to 1 ml/hr after 48 hr of induction.
Production of Recombinant Proteins
5.8.9 Current Protocols in Protein Science
Supplement 2
SUPPORT PROTOCOL
SMALL-SCALE PREPARATION OF PROTEIN EXTRACTS Following induction using one of the previous basic or alternate protocols, protein extracts may be made from the harvested yeast cells either by mechanical disruption or by spheroplasting followed by lysis in hypotonic buffer. A simple mechanical disruption method is described here, in which cells are vortexed in the presence of glass beads. This method can be used for both analytical- (e.g., 10-ml) or preparative-scale cultures; for the latter a Bead Beater (BioSpec Products) may be used. Cells from 10-ml inductions can be disrupted rapidly using glass beads on a vortex mixer with tube rack. For Pichia pastoris high-density fermentations of >2 liters, a larger bead mill with efficient cooling is required (Dyno Mill). Although proteases are less of a problem than with Escherichia coli, cell breakage should be carried out in the cold, ideally in the presence of protease inhibitors. A typical extraction buffer is given here, but the particular buffer used will depend on the protein being studied and the subsequent analysis or purification to be performed. If the aim is simply to detect a polypeptide by SDS-polyacrylamide gel electrophoresis (UNIT 10.1), the composition of the buffer is less important, because the total extract is denatured before analysis. Materials Cell pellet from an induced culture of transformed yeast cells Protein extraction buffer (see recipe), ice-cold 5× protease inhibitor cocktail (see recipe) 15-ml polypropylene snap-top tubes 0.45-mm glass beads, acid washed (see recipe) Platform vortex mixer (e.g., IKA-Vibrax-VXR with test tube rack) 1. Resuspend the cell pellet from an induced culture of transformed yeast cells in 0.5 ml ice-cold extraction buffer plus 125 µl of 5× protease inhibitor cocktail in a 15-ml polypropylene snap-cap tube. 2. Add glass beads to two-thirds the height of the meniscus. Place the tubes on a platform vortex mixer in a cold room and vortex 10 min at full speed. Samples can be vortexed by hand using a vortex mixer in six 30-sec bursts punctuated by 30-sec intervals on ice. Breakage can be checked using a phase-contrast light microscope: intact cells appear bright and broken cells are phase-dark “ghosts.” Breakage is most efficient in wide round-bottom tubes. For larger volumes use 30-ml centrifuge tubes or a Bead Beater.
3. Recover the cell extract from the glass beads using a fine-tipped plastic Pasteur pipette. To recover as much of the extract as possible, reextract the beads with 0.5 ml ice-cold extraction buffer. Alternatively, centrifuge the cell extract and beads 2 min at 1000 × g, 4°C, through a plastic disposable filter (e.g., Poly-Prep chromatography columns, Bio-Rad). Check that the protein of interest is not retained by the filter. For large-scale purification of cytosolic proteins, the preparation may be centrifuged 1 hr at 100,000 × g, 4°C, to pellet membranes.
4. Determine the total protein concentration of the extract using a standard assay (e.g., Bio-Rad protein assay; also see UNIT 3.4). The yield of total protein from a 10-ml culture grown to A600 = 10 is ∼10 mg. Culture of Yeast for the Production of Heterologous Proteins
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5. If desired, separate insoluble protein by microcentrifuging 15 min at 12,000 × g, 4°C. Transfer the soluble protein to a new tube and determine the protein concentration; retain this and the pellet containing insoluble proteins for analysis. To determine the proportion of a protein that is soluble, resuspend the insoluble protein pellet in the same volume of buffer as the soluble protein preparation, and analyze equal volumes of total, soluble, and insoluble protein by SDS-PAGE (see UNIT 10.1).
6. Store protein extracts at −20° or −70°C. Upon thawing, a precipitate, largely consisting of cell membranes, may appear; this should be dispersed prior to SDS-PAGE (e.g., by pipetting up and down).
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Basal salts solution, 5× Per liter: 42 ml phosphoric acid (0.62 M final) 1.8 g calcium sulfate⋅2H2O (10 mM final) 28.6 g potassium sulfate (0.16 M final) 23.4 g magnesium sulfate⋅7H2O (10 mM final) 6.5 g potassium hydroxide (0.12 M final) Autoclave Store indefinitely at room temperature BSSM fermenter medium 3.35 g monobasic potassium phosphate (1 mM final) 1.8 g magnesium sulfate⋅7H2O (3 mM final) 26.25 g ammonium sulfate (0.08 M final) 2.5 liters distilled water 0.6 ml 0.07 M manganese sulfate⋅4H2O (20 µM final) 0.6 ml 0.08 M copper sulfate⋅5H2O (20 µM final) 0.6 ml 0.6 mM zinc sulfate⋅7H2O (4 µM final) 0.1 ml 33 mM potassium iodide (1 µM final) 0.1 ml 0.39 M sodium molybdate⋅2H2O (20 µM final) 0.1 ml 0.08 M ferric chloride⋅6H2O (3 µM final) Place in 5-liter fermenter vessel and autoclave Add 50 ml vitamin solution (see recipe) Casamino acids, 10× Prepare a 20% (w/v) solution of Casamino acids (low-salt, Difco). Autoclave. Store up to 1 year at 4°C. Glass beads, acid washed Wash 0.45-mm-diameter glass beads (B. Braun) for 16 hr in concentrated HCl. Rinse thoroughly in distilled water. Bake 16 hr at >150°C. Store indefinitely at room temperature.
Production of Recombinant Proteins
5.8.11 Current Protocols in Protein Science
Supplement 2
Protease inhibitor cocktail, 5× 20 mM EDTA 20 mM EGTA 20 mM PMSF 10 mg/ml pepstatin 10 mg/ml leupeptin 10 mg/ml chymostatin 10 mg/ml antipain Store in 1-ml aliquots up to 1 year at −20°C Protein extraction buffer 50 mM Tris⋅Cl, pH 7.4 100 mM NaCl 2 mM EDTA Autoclave Store indefinitely at room temperature Chill on ice before use PTM1 salt solution Per liter: 6 g cupric sulfate⋅5H2O (24 mM final) 0.08 g potassium iodide (0.5 mM final) 3 g manganese sulfate⋅H2O (18 mM final) 0.2 g sodium molybdate⋅2H2O (0.8 mM final) 0.02 g boric acid (0.3 mM final) 0.5 g cobalt chloride⋅6H2O (2 mM final) 20 g zinc chloride (0.15 M final) 65 g ferrous sulfate⋅7H2O (0.23 M final) 0.2 g biotin (0.8 mM final) 5 ml sulfuric acid (0.9 M final) Autoclave Store indefinitely at room temperature Vitamin solution Per 50 ml: 10 mg biotin 65 mg pantothenic acid 100 mg meso-inositol 5 mg 4-aminobenzoic acid 65 mg pyridoxine⋅HCl 50 mg thiamine 50 mg nicotinic acid Filter sterilize Store up to 1 year at 4°C The vitamin solution can also be prepared as a 10× stock.
YNB stock, 10× Dissolve 6.7 g yeast nitrogen base without amino acids (Difco) in 100 ml distilled water. Filter sterilize with 0.2-µm filter. Store up to 1 year at 4°C. Culture of Yeast for the Production of Heterologous Proteins
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YNB medium Add 10× YNB stock (see recipe) and 500× biotin stock to sterile distilled water to give 1× final concentration for basic YNB medium. Add a carbon source (e.g., glucose or raffinose) and nutritional supplements (see Table 5.8.2) according to vector selection requirements. Store up to 1 year at 4°C. Table 5.8.2 Nutritional Supplements for YNB Medium
Nutrienta Adenine sulphate L-Arginine⋅HCl L-Aspartic acid L-Glutamic acid L-Histidine⋅HCl L-Isoleucine L-Leucine L-Lysine⋅HCl L-Methionine L-Phenylalanine L-Serine L-Threonine L-Tryptophan L-Tyrosine
Uracil Valine
Final concentrationb (mg/liter) 20 20 100 100 20 30 30 30 20 50 375 200 20 30 20 150
aPrepare as 100× stock solutions and filter sterilize. Store at 4°C. bAdd to filter-sterilized medium.
YP broth 10 g yeast extract 20 g Bacto Peptone 1 liter H2O Autoclave Store up to 1 year at room temperature COMMENTARY Background Information Yeast expression systems have advantages over other commonly used systems for the production of proteins in certain situations (see UNITS 5.6 & 5.7 and Romanos et al., 1992). Although typical percentage yields of proteins in Saccharomyces cerevisiae (1% to 5% of cell protein) are less than those obtained with Escherichia coli or baculovirus, the volumetric yield with S. cerevisiae may be higher than with the baculovirus system due to the much higher cell density. Depending on the productivity of
the strain, shaker-flask inductions, scaled up to 1 liter if necessary, may be sufficient to provide the amount of protein needed. Where a significant amount of protein is required, e.g., >100 mg, fermenters may be used to culture yeast to extremely high density, e.g., 100 g dry weight/liter; this is particularly valuable with secretion, where the concentration of protein in the medium can increase almost proportionately. Because of the technical complexity involved in achieving such densities with S. cerevisiae, this unit includes a much simplified
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Table 5.8.3 Comparison of Yields for Heterologous Expression of Two Proteins
Expression system
Tetanus toxin fragment C (% of total protein)
Pertactin (% of total protein)
E. coli Baculovirus S. cerevisiae GAL7 P. pastoris
24 10a 2-3 28b
30 >40a 0.1 10c
aEstimate; variable yield greatly reduced on scale-up. bScale-up without loss of yield to >12 g/liter. cScale-up without loss of yield to >4 g/liter.
protocol using calculated feeding rates, which gives lower cell densities. However, the authors recommend the use of Pichia pastoris for highdensity fermentation wherever possible. P. pastoris is an industrial yeast, initially selected for its high-density growth in simple defined media; this organism has also been developed as an expression system (Romanos et al., 1992; Cregg et al., 1993). There are numerous cases in which much higher percentage yields of intracellular proteins have been obtained using P. pastoris than using S. cerevisiae, often to levels equivalent to those in E. coli and baculovirus—e.g., 5% to 40% of cell protein (Sreekrishna et al., 1989; Clare et al., 1991a; Romanos et al., 1991). Table 5.8.3 shows comparative data for two proteins expressed in E. coli, baculovirus, S. cerevisiae, and P. pastoris. P. pastoris is also generally more efficient in secreting foreign proteins (Clare et al., 1991b). Moreover, scale-up of P. pastoris to high-density fermentations is very simple and can give enormous yields on a volumetric basis. The tight regulation of the AOX1 promoter means that the growth phase is essentially identical for different protein products, and scale-up to large fermenters does not affect biomass or product yield. Additionally, contrary to the experience with S. cerevisiae, an immediate improvement in percentage yield of foreign protein is generally seen on going from shaker flasks to fermenters. This appears to be due at least in part to the high O2 demand of the organism, which is not satisfied in shaker flasks. For further general information on use of the P. pastoris system, consult the instruction manual available from Invitrogen and UNIT 5.7. Culture of Yeast for the Production of Heterologous Proteins
Critical Parameters S. cerevisiae shaker-flask inductions are relatively straightforward, but improvements can be made by testing different host strains,
including protease-deficient strains. In S. cerevisiae fed-batch fermentations, achieving the correct feed rate is critical; overfeeding, especially in the first few hours, will result in lower cell yield. Because the feed rate in the protocol given here is calculated, it is essential that the density of the inoculum reaches the required level. Another critical factor is dissolved oxygen: levels <30% of saturation affect cell growth, while <10% may affect expression. Increasing the agitation or air-flow rate can increase dissolved oxygen. The most important factors affecting expression in P. pastoris have been discussed in reviews (e.g., Romanos et al., 1992; Cregg et al., 1993; UNIT 5.7). When using P. pastoris, selection of the transformant and growth conditions are highly critical. Selection of the transformant is important because P. pastoris transformations, especially BglII transplacements, result in a highly heterogeneous population of transformants. The variables are Mut phenotype (determined by whether or not AOX1 is disrupted), site of integration, and copy number. For intracellular proteins this usually results in extreme clonal variation in yield between different transformants, dependent mainly on the number of copies of the vector integrated into the chromosome (Sreekrishna et al., 1989; Clare et al., 1991a; Romanos et al., 1991; Scorer et al., 1994). A number of approaches have been used to maximize vector copy number—e.g., screening transformants by DNA dot blot (Romanos et al., 1991) and G418 selection (Scorer et al., 1994). Although it appears that the highest copy numbers result from transplacements using the spheroplast technique, for rapid isolation of nearoptimal transformants the authors recommend the much simpler method of G418 selection following electroporation (Scorer et al., 1994). With secretion of foreign proteins, by contrast, the presence of many copies is sometimes
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suboptimal, and the Mut phenotype may have a greater influence on yields; consequently, a significant number of transformants with different copy numbers may need to be tested for maximum yield (Thill et al., 1990; Clare et al., 1991b). An alternative approach is to preselect efficient secretors empirically by micro-scale inductions in 96-well plates (Laroche et al., 1994). A critical factor during P. pastoris growth is its tendency to become oxygen-limited in small-scale inductions, leading to low and variable yields. This effect can be minimized by taking steps to ensure good aeration. However, it is nevertheless a major reason for switching to fermenters as soon as the most efficiently expressing transformants have been identified from small-scale inductions. The change from small-scale to fermenter is usually accompanied by an increase in percentage yield. A factor that affects P. pastoris fermentations is the Mut phenotype, which determines whether or not there is continued growth during the methanol induction phase. Mut+ strains have a greater tendency to become oxygen-limited during induction and may give more variable yields in small-scale inductions. The methanol feeding rate will be different for Mut+ and MutS strains because the latter can be poisoned by excess methanol feeding.
Troubleshooting Problems in expression can arise at numerous stages from vector stability through to protein turnover. The following problems are among the most common: (1) genetic instability of the transformed yeast strain, particularly on scale-up; (2) inability to produce some proteins because of their high toxicity; (3) failure to express AT-rich genes due to premature transcriptional termination; (4) inefficient secretion of some larger (>30 kDa) proteins; (5) proteolysis of secreted proteins; and (6) inappropriate glycosylation of mammalian glycoproteins. These problems have been discussed in previous reviews (e.g., Romanos et al., 1992; UNITS 5.6 & 5.7), and this section will concentrate on those problems related purely to strain stability and growth conditions. In S. cerevisiae, low yields of foreign proteins may arise because of reduced copy number, rearrangement, or mutation of the vector, most frequently during the constitutive expression of toxic proteins. The solution to most of these problems is simply to use a tightly regulated promoter—e.g., a GAL promoter or the P.
pastoris AOX1 promoter—to reduce selection pressure during the growth phase of the culture. It is also good practice to always prepare banks of −70°C glycerol stocks of selected transformants before the start of a project, so that the starting point of each culture is the same. However, extreme product toxicity can be an intractable problem, and it may then be more profitable to try a transient expression system, such as baculovirus. In P. pastoris, a common problem is proteolysis of secreted proteins (reviewed in Cregg et al., 1993). The problem may occur in shaker flasks but is exacerbated in high-density fermenter cultures, possibly due to the higher concentration of proteases and the different medium used. A number of strategies have been used successfully to overcome proteolysis: adding peptide supplements (peptone or Casamino acids), buffering the medium to pH 3 or 6, or using a pep4 protease-deficient strain.
Anticipated Results Yields of intracellular proteins from S. cerevisiae are usually in the range of 0.5% to 5% of total cell protein, or 1.5 to 15 mg/liter/A600 unit (15 to 150 mg/liter for typical culture densities in YP medium). Many small nonglycosylated polypeptides are efficiently secreted and may reach levels of 10 mg/liter/A600 unit in the culture supernatant. Larger proteins (>25 kDa) are secreted far less predictably, some efficiently and some poorly. The cell density at the end of the fed-batch phase of S. cerevisiae fermentations should reach at least A600 = 20 (4.0 g/liter) and A600 = 40 (8 g/liter) at harvest. Using this protocol with minor modifications, such as the use of complex medium, cell densities in excess of 30 g/liter have been reported (Hsieh et al., 1988). Yields are highly strainand protein-specific. Typical yields with this protocol are ∼0.3 g/liter for intracellular products and ∼100 mg/liter for extracellular products. P. pastoris frequently yields intracellular products at 5% to 30% of cell protein. Cell density reaches 130 g dry weight/liter in the fermenter, of which >30% is protein; this is equivalent to at least 2 to 12 g/liter of product. As with S. cerevisiae, yields of secreted proteins are less predictable but frequently reach several hundred mg/liter at high cell density, and may constitute the only major protein in the culture medium. It is recommended that positive control strains—e.g., a strain expressing β-galactosidase—be used to ensure that the
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Supplement 2
induction conditions are optimal; control strains for P. pastoris can be obtained from Invitrogen.
Time Considerations Small-scale S. cerevisiae inductions take a total of 2 days from inoculation of the starter culture to harvesting of the induced cells. Several inductions (e.g., 24) can be carried out at a time. P. pastoris inductions are generally carried out for 2 to 5 days; therefore, the procedure takes 3 to 6 days from start to finish, depending on the optimal length of induction for the particular protein. S. cerevisiae fermentations typically take 4 days from start to finish: 2 days for inoculum development, 1 day for biomass accumulation on glucose, and ∼1 day for galactose induction. P. pastoris 1-liter fermentations typically take 5 days from start to finish: 1 day to produce a starter inoculum, 2 days for biomass accumulation on glycerol, and up to 2 days for methanol induction. Some workers have reported that MutS inductions require a longer period, but this has not been the authors’ experience. Half a day is required after the fermenter run to harvest and wash cells and to clean the fermenter.
Literature Cited Aiba, S., Nagai, S., and Nishizawa, Y. 1976. Fedbatch culture of Saccharomyces cerevisiae: A perspective of computer control to enhance the productivity of Baker’s yeast cultivation. Biotechnol. Bioeng. 18:1001-1016. Ausubel, F.M., Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G., Smith, J.A., and Struhl, K. (eds.) 1995. Current Protocols in Molecular Biology. John Wiley & Sons, New York.
Culture of Yeast for the Production of Heterologous Proteins
B.L., Cruze, J., Torregrossa, R., Velicelebi, G., and Thill, G.P. 1987. High-level expression and efficient assembly of hepatitis B surface antigen in the methylotrophic yeast Pichia pastoris. Bio/Technology 5:479-485. Cregg, J.M., Vedvick, T.S., and Raschke, W.C. 1993. Recent advances in the expression of foreign genes in Pichia pastoris. Bio/Technology 11:905-910. Fieschko, J.C., Egan, K.M., Ritch, T., Koski, R.A., Jones, M., and Bitter, G.A. 1987. Controlled expression and purification of human immune interferon from high-cell-density fermentations o f Saccharomyces cerevisiae. Biotechnol. Bioeng. 29:113-1121. Hsieh, J.-H., Shih, K.-Y., Kung, H.-F., Shiang, M., Lee, L.-Y., Meng, M.-H., Chang, C.-C., Lin, H.-M.., Shih, S.-C., Lee, S.-Y., Chow, T.-Y., Feng, T.-Y., Kuo, T.-T., and Choo, K.-B. 1988. Controlled fed-batch fermentation of recombinant Saccharomyces cerevisiae to produce hepatitis B surface antigen. Biotechnol. Bioeng. 32:334-340. Laroche, Y., Storme, V., De Meutter, J., Messens, J., and Lauwereys, M. 1994. High-level secretion and very efficient isotopic labeling of tick anticoagulant peptide (TAP) expressed in the methylotrophic yeast, Pichia pastoris. Bio/Technology 12:1119-1124. Pirt, S.J. 1985. Batch cultures with substrate feeds. In Principles of Microbe and Cell Cultivation, pp. 211-222. Blackwell Scientific, Oxford. Romanos, M.A., Clare, J.J., Beesley, K.M., Rayment, F.B., Ballantine, S.P., Makoff, A.J., Dougan, G., Fairweather, N.F., and Charles, I.G. 1991. Recombinant Bordetella pertussis pertactin (P69) from the yeast Pichia pastoris: High-level production and immunological properties. Vaccine 9:901-906. Romanos, M.A., Scorer, C.A., and Clare, J.J. 1992. Foreign gene expression in yeast: A review. Yeast 8:423-488.
Bock Gu, M., Jung, K.-H, Park, M.N., Shin, K.S., and Kim, K.H. 1989. Production of HbsAg by growth-rate control with recombinant Saccharomyces cerevisiae in fed-batch. Biotech. Lett. 11:1-4.
Scorer, C.A., Clare, J.J., McCombie, W.R., Romanos, M.A., and Sreekrishna, K. 1994. Rapid selection using G418 of high-copy-number transformants of Pichia pastoris for high-level foreign gene expression. Bio/Technology 12:181-184.
Clare, J.J., Rayment, F.B., Ballantine, S.P., Sreekrishna, K., and Romanos, M.A. 1991a. Highlevel expression of tetanus toxin fragment C in Pichia pastoris strains containing multiple tandem integrations of the gene. Bio/Technology 9:455-460.
Sonnlietner, B. and Kappeli, O. 1986. Growth of Saccharomyces cerevisiae is controlled by its limited respiratory capacity: Formation and verification of a hypothesis. Biotechnol. Bioeng. 28:927-937.
Clare, J.J., Romanos, M.A., Rayment, F.B., Rowedder, J.E., Smith, M.A., Payne, M.M., Sreekrishna, K., and Henwood, C.A. 1991b. Production of mouse epidermal growth factor in yeast: High-level secretion using Pichia pastoris strains containing multiple gene copies. Gene 105:205-212. Cregg, J.M., Tschopp, J.F., Stillman, C., Siegel, R., Akong, M., Craig, W.S., Buckholz, R.G., Madden, K.R., Kellaris, P.A., Davies, G.R., Smiley,
Sreekrishna, K., Nelles, L., Potenz, R., Cruze, J., Mazzaferro, P., Fish, W., Motohiro, F., Holden, K., Phelps, D., Wood, P., and Parker, K. 1989. High-level expression, purification, and characterization of recombinant human tumour necrosis factor synthesized in the methylotrophic yeast Pichia pastoris. Biochemistry 28:4117-4125. Thill, G.P., Davis, G.R., Stillman, C., Holtz, G., Brierley, R., Engel, M., Buckholtz, R., Kinney, J., Provow, S., Vedvick, T., and Seigel, R.S. 1990. Positive and negative effects of multi-copy
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integrated expression vectors on protein expression in Pichia pastoris. In Proceedings of the 6th International Symposium on Genetics of Microorganisms, Vol. II (H. Heslot, J. Davies, J. Florent, L. Bobichon, G. Durand, and L. Penasse, eds.) pp. 477-498. Société Française de Microbiologie, Paris. Wang, H.Y., Cooney, C.L., and Wang, D.I. 1979. Computer control of bakers’ yeast production. Biotechnol. Bioeng. 21:975-995.
Key References Cregg et al., 1993. See above. An excellent review of P. pastoris expression technology. McNeil, B. and Harvey, L.M. eds. 1990. Fermentation: A practical approach. Oxford University Press, Oxford. A detailed laboratory manual covering all aspects fermentation, especially useful to those new to fermentation technology. Romanos et al., 1992. See above.
foreign gene expression, secretion, alternative yeasts (including P. pastoris), and fermenter scaleup. Romanos, M.A., Scorer, C.A., and Clare, J.J. 1995. Expression of cloned genes in yeast. In DNA Cloning 2, Expression Systems: A Practical Approach (D.M. Glover and B.D. Hames, eds.) pp. 123-167. IRL Press, Oxford. A detailed practical review of the basic strategies and techniques for high-level expression of genes in yeast including approaches for diagnosing and solving problems.
Contributed by Michael A. Romanos and Jeffrey J. Clare Wellcome Research Laboratories Beckenham, United Kingdom Crawford Brown British Bio-Technology Ltd. Cowley, United Kingdom
A detailed review covering all aspects of yeast expression, including vector systems, factors affecting
Production of Recombinant Proteins
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Overview of Protein Expression by Mammalian Cells In recent years, mammalian cells have been used in the production of recombinant proteins, antibodies, viruses, viral-subunit proteins, and gene-therapy vectors. In addition to being used in commercial biotechnology, mammalian cell systems have served as a means for examining fundamental aspects of gene replication, transcription, translation, and post-translational protein processing. The availability of transformable cell lines, along with viral and plasmid-based mammalian-cell vector systems, has provided tools through which important aspects of mammalian gene function can be investigated. The following are typical uses for mammalian expression systems: 1. verification of a cloned gene product; 2. analysis of the effects of protein expression on cell physiology; 3. production and isolation of genes from cDNA libraries; 4. production of correctly folded and glycosylated proteins for assessment of biological activity in both in vitro and in vivo systems; 5. production of suitable quantities of proteins and glycoproteins for structural characterization of protein and carbohydrate moieties; 6. production of important clinically active viral surface antigens—e.g., prehepatitis B virus surface antigen (preS2 HBVsAg)—as well as therapeutic proteins—e.g., β-interferon, tissue plasminogen activator (tPA), erythropoietin (EPO), and Factor VIII; and 7. production of monoclonal antibodies. Important features of mammalian cells include their ability to perform post-translational modifications and to secrete glycoproteins that are correctly folded and contain complex antennary oligosaccharides with terminal sialic acid. These covalent modifications may modulate the clinical efficacy of the protein (e.g., circulatory half-life and biospecificity) or result in properties that are of interest for biochemical characterization—e.g., with respect to structural stabilization, functional groups, and biological role. Mammalian-produced proteins are quality-controlled through a process whereby the progress of incompletely folded, misassembled, and unassembled proteins into the secretory pathway is selectively inhibited (Hurtley and Helenius, 1989). The correctly processed material progresses and is generally Contributed by David Gray Current Protocols in Protein Science (1997) 5.9.1-5.9.18 Copyright © 1997 by John Wiley & Sons, Inc.
UNIT 5.9
secreted as fully active protein. Another important feature of mammalian cell expression has been the ability to provide proteins bearing sialyl glycoforms closely resembling glycoproteins produced by humans. In contrast, E. coli does not possess any glycosylation machinery and yeast tends to produce hypermannose-type glycosylation. The yeast-produced glycoproteins are rapidly cleared from the circulation by the liver and also may induce adverse immunogenic responses in humans. The post-translational modifications performed by mammalian cells include the proteolytic processing of the propeptide (Neurath, 1989) as well as glycosylation and carbohydrate trimming (Hirschberg and Snider, 1987). Other modifications may include γ carboxylation of glutamic acid residues (Suttie, 1985; Kaufman et al., 1986a); hydroxylation of aspartic acid and asparagine residues (Stenflo, 1989); sulfation of tyrosine residues (Baeuerle and Huttner, 1987); phosphorylation of proteins through cell receptor–protein interaction (Myers et al., 1994); fatty acid acylation (McIlhinney, 1990); and correct assembly of multimeric proteins. In contrast to what happens in mammalian cells, the production of intracellular recombinant proteins in E. coli leads to accumulation of the protein in reduced form, often as insoluble refractile bodies requiring subsequent denaturation and refolding. This unit will briefly review the stages involved in protein production in mammalian cells using the stable-expression approach as outlined in Figure 5.9.1. Discussion will include choice of cell type, transfection of the host cells, methods for selection and amplification of transformants, and growth of cells at appropriate scale for protein production. Since post-transcriptional modification and intracellular protein transportation are important features of recombinant-protein production in mammalian cells, some description of these mechanisms is included.
CHOICE OF MAMMALIAN CELL HOST Although a variety of mammalian cell hosts are available for protein production, only a small number have emerged as systems of choice for production of proteins to be used clinically. The most common of these cell hosts
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are summarized in Table 5.9.1. The narrowing down of choices is largely due to the need for cell lines that: (1) are capable of continuous growth; (2) can be grown in suspension (in bioreactors); (3) have low risk of adventitious infection by potentially pathogenic viruses; (4) have genetic stability; and (5) can be readily characterized with respect to karyology, morphology, isoenzymes, and gene copy number. The existence of a variety of host-cell systems, the availability of viral or cDNA-based vectors, and the possibility of either stable or transient
construction of vector(s)
expression requires that the prospective user define an expression strategy based on ultimate goals. When the researcher’s objectives require <1 mg of protein, transient expression in COS-7 cells is the relevant route. Transient expression in the COS-cell and vaccinia systems has been recently reviewed (Moss and Earl, 1991; Aruffo, 1997), and detailed protocols for construction of suitable vectors and protein expression by these systems can be found in those articles. In transient expression a burst of production occurs in the host cell and is usually accompa-
selection of host cell
transfection of host cell (with protein gene and marker function gene)
cloning (dilution to single colony)
initial growth and selection for production of protein (in presence of inhibitor)
expansion of clones (growth)
amplification of gene (stepwise increases in inhibitor concentration)
additional round of selection (selecting higher producers)
adaptation (to medium compatible with production)
cell banking (choice of best cell line and freezing of aliquots)
Overview of Protein Expression by Mammalian Cells
growth of cells in stirred reactors
Figure 5.9.1 Flow chart of activities in the development of a mammalian-cell expression system.
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Table 5.9.1
Common Mammalian Cell Hosts
Description
Growth
Utility
Sourcea
Namalwa
Burkitt’s lymphoma–transformed lymphoblastoid cell
Large-scale suspension
Production of α interferon (e.g., by Burroughs Wellcome)
ATCC #CRL-1432
HeLa
Aneuploid cervical carcinoma cell
Small-scale suspension
Production of small quantities of research material (few mg)
ATCC #CCL-2
293
Transformed kidney cell
Small-scale suspension
Production of small quantities of research material (few mg)
ATCC #CCL-1573
WI-38
Human diploid normal embryonic lung cell
Attachment only
Host for virus production
ATCC #CCL-75
MRC-5
Human diploid normal embryonic lung cell
Attachment only
Hardier host for virus production—e.g., hepatitis A
ATCC #CCL-171
HepG2
Liver carcinoma transformed cell
Attachment
Small-scale evaluation of expression—e.g., HBVsAg
ATCC #HB-8065
3T3
Swiss mouse embryo fibroblast
Attachment
Used in testing transforming agents, expression evaluation
ATCC #CCL-92
L-929
Normal connective tissue fibroblasts
Attachment
Small-scale evaluations of expression
ATCC #CCL-1
Myeloma (e.g., NS/O)
Many types
Large-scale suspension
Monoclonal antibody production
ATCC (and commercial sources)
BHK-21
Baby hamster kidney cell
Large-scale suspension
Host for virus production or for stable gene integration
ATCC #CCL-10
CHO-K1
Chinese hamster ovary cell
Large-scale suspension
Used with glutamine synthetase system
ATCC #CCL-61
CHO DG44
Chinese hamster ovary cell
Large-scale suspension
Host for DHFR coamplification
L. Chasinb
CHO DXB11
Chinese hamster ovary cell
Large-scale suspension
Preferred host for DHFR coamplification
L. Chasinb
COS-7
Transformed African green monkey kidney cell
Small-scale attachment
Transient expression host
ATCC #CCL-1651
Vero
Normal African green monkey kidney cell
Large-scale attachment
Production of viruses
ATCC #CCL-81
Cell line Human
Rodent
Monkey
aAbbreviation: ATCC, American bE-mail:
Type Culture Collection (see SUPPLIERS APPENDIX).
[email protected]
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Overview of Protein Expression by Mammalian Cells
nied by death and rapid lysis of the cell. This presents the purification scientist with the challenge of fishing out the protein of interest, which may be present at ∼5 µg/ml, from a soup of lysed cellular protein, nucleic acids, and viral particles. The yield of product during purification may be low as a result of the low titer and starting purity; however, when only small quantities of protein are required, transient expression in COS cells or the vaccinia system is a quick and suitable system to employ. For production of larger quantities of protein, stable expression must be used because of the difficulty in scaling up transient expression into a bioreactor system. Stable expression in Chinese hamster ovary (CHO) cells is described in UNIT 5.10. Some cell lines have successfully been used as hosts in production of viruses and useful proteins. These include the human embryonic lung cell line MRC-5 and the normal embryonic cell line WI-38. These cells are attachment growth–dependent and have a finite lifespan of ∼50 generations, after which time the cells enter a senescent phase and begin to die. Cell lines that have undergone transformation by virus or that have experienced alteration of chromosomes can exhibit a capacity for infinite growth as well as an ability to grow in suspension. Examples of such cell lines are HeLa cells from a human cervical cancer and Namalwa cells from a human lymphoma. There has been some reluctance to use these cell lines in production of clinical agents because of the possibility of transfer of tumorigenic agents to the product. For production of larger amounts of recombinant protein, stable expression is required and has been primarily performed in CHO cells, baby hamster kidney (BHK-21) cells, or myeloma cells (e.g., NS/O). These cell lines are capable of indefinite growth on a large scale and are suitable hosts for stable integration of heterologous DNA. Stable expression results from integration into the host-cell genome of the gene for the expression of the heterologous protein. The integrated gene is transcribed efficiently and the protein is expressed persistently over many generations by the host cell. A stable producercell line typically provides ∼1 to 10 mg of secreted protein per 109 cells per day (specific productivity). For monoclonal antibody production, productivity levels of 35 to 100 mg per 109 viable cells per day in myeloma cells (Bebbington et al., 1992; Shitara et al., 1994) and 15 to 110 mg per 109 viable cells per day in CHO cells (Page and Sydenham, 1991) are
obtainable. Such levels may allow secreted-antibody titers of 1 to 1.5 mg/liter to be achieved in optimized large-scale systems; hence these systems have been popular in biotechnology. Where protein is accumulated intracellularly, it may reach little more than 0.1% to 0.5% of total cell protein. Thus, the focus has tended to be on secretion of protein where significant amounts of product are obtainable. The obvious drawback associated with stable expression in mammalian cells is the time required to obtain stable productive cell banks (months) versus that required to obtain large amounts of non-post-translationally modified proteins in E. coli or large quantities of highmannose-containing glycoprotein using yeast and insect cells (2 to 3 weeks). The advantages of obtaining the more complex proteins from mammalian cells include the ability to understand how these molecules interact in biological models both in vitro and in vivo. Using mammalian systems, much knowledge has been gained in the understanding of post-translational modification, protein quality control, and protein translocation. This knowledge ultimately facilitates improved expression in mammalian hosts. Stable expression requires the transfer of the foreign DNA, along with the relevant DNAbased signals for transcription by RNA polymerase II, into the chromosomal DNA of the host cell (e.g., CHO). A suitable plasmid vector is required to carry the functions through to the host DNA. Vectors are commercially available from a variety of vendors—e.g., Invitrogen, Clontech, and Promega (see SUPPLIERS APPENDIX). A large variety of vectors are described in Pouwels et al. (1988). Suitable replicons contain 5 to 10 kb of DNA and usually are designed to contain phenotypic markers for selection in E. coli (e.g., Apr) and CHO cells (DHFR+). Other features of appropriate plasmids include a promoter-distal cloning site along with other cloning sites; a replication origin for E. coli; a polyadenylation sequence from SV40 or bovine growth hormone; a eukaryotic origin of replication (e.g., SV40 or oriP); the gene for expression of the protein in a site proximal to the promoter; and a promoter and associated elements. Commonly used constitutive promoters for CHO cell applications are the SV40 early promoter (Moreau et al., 1981; Neuhaus et al., 1984), the adenovirus late promoter (McKnight and Tjian, 1986), and the cytomegalovirus (CMV) early promoter (Boshart et al., 1985). Transcriptional enhancers such as the human CMV enhancer (Boshart et al.,
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Figure 5.9.2 Representation of a mammalian expression vector based on the vector pRSC, published by Tsang et al. (1997); adapted with permission. The plasmid has two multiple cloning sites (MCSs) and three promoters. One promoter drives the neomycin-resistance protein while the remaining two MCSs have specific promoters and can be used to incorporate an amplification marker (DHFR) and a gene for the protein of interest.
1985) can enhance transcriptional activity by 10- to 100-fold and may be incorporated into the vector. These features are illustrated in Figure 5.9.2. CHO cells have been used extensively as a host for stable expression of proteins. A number of CHO mutants have been developed that provide the user with tools to examine the synthesis of DNA, RNA, and protein as well as protein secretion, protein glycosylation, and intermediary metabolism. The most popular CHO sublines (also see Urlaub and Chasin, 1980) are CHO DXB11 (dhfr+/dhfr−) and CHO DG44 (dhfr−/dhfr−). The DXB11 cell was derived from CHO-K1 in 1978 by Lawrence Chasin. The CHO-K1 cells were originally derived in the 1950s, and the cells that were used in Chasin’s laboratory for the development of the DXB11 line were obtained from Ted Puck and Fa-ten Kao at the Eleanor Roosevelt Cancer Institute in Denver in 1970. The DG44 cell line contains a double mutation in the dhfr genes and is not capable of natural reversion to the dhfr+ phenotype. The DG44 cell line was derived from CHO pro3− cells by Chasin in 1982. The CHO pro3− was derived from the cell line
established in the 1950s and is sometimes referred to as CHO Toronto, because it was used extensively in that city by Louis Siminivitch and colleagues. Both these cell lines can be obtained from Dr. Lawrence Chasin of Columbia University (e-mail: [email protected]). Other mutants have been produced, and a brief list is included in Wirth and Hauser (1993).
TRANSFECTION, SELECTION, AND AMPLIFICATION For introduction of genetic material into mammalian cells, three types of vectors are available—viruses, plasmid vectors with a replicon for animal cells, and plasmid vectors without a replicon for animal cells. Most viruses that replicate in mammalian cells ultimately cause death and lysis of the host cell, so expression of virus-borne genes is transient and can only be obtained over a short period prior to lysis. Retroviruses are capable of integrating genetic material into the chromosome of host cells—i.e., stable transformation. As with viruses, the use of plasmid vectors can result in either stable or transient expression of a gene product; the plasmid vectors without a mam-
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Overview of Protein Expression by Mammalian Cells
malian cell replicon would require stable integration for expression of gene product. Stable production of foreign proteins by mammalian cells requires use of a host cell line that is capable of incorporating the heterologous-gene DNA into a chromosome. In addition to the need for successful stable integration, a selection method is needed to fish out the stable transformants from the large population of cells that may contain unstable, intracellular heterologous gene copies. This necessitates cotransfection of a gene conferring some unique phenotypic property. Resistance to cytotoxic drugs is the most frequently applied method for selection of stable transformants. Normally, a gene for recessive drug resistance would be cotransfected into a host cell deficient in that particular activity. The most common approach is the use of dihydrofolate reductase (DHFR), which can be used as a selectable marker in DHFR-deficient CHO cells (Kaufman and Sharp, 1982). Other common transfectable selectable amplification markers that can be used with CHO cells are adenosine deaminase (in presence of inhibitors of ADA or cytotoxic levels of adenine or its analogs) in CHO DXB11 (DHFR−; Yeung et al., 1983; Kaufman et al., 1986b); carbamyl phosphate synthetase–aspartate transcarbamylase–dihydroorotase in CHO UrdA mutant cells (de Saint Vincent et al., 1981); asparagine synthetase in CHO N3 (AS−) cells (Cartier and Stanners, 1990); glutamine synthetase in any CHO cell (Bebbington and Hentschel, 1987); ornithine decarboxylase in CHO C55.7 (ODC−; Chiang and McConlogue, 1988); multiple drug resistance in any CHO cell (Kane et al., 1989); and a mutant DHFR (Leu 22 changed to Arg 22) which can be used in any CHO cell host (Simonsen and Levinson, 1983). See Mortensen et al. (1997) for additional discussion of the use of selectable markers with mammalian cells. It has been reported that a highly variable but typically large fraction (20% to 100%) of CHO clones established through calcium phosphate– or lipofection-mediated cotransfections (of dhfr and heterologous protein genes) coexpressed the dual-transfected expression vectors (Wurm and Petropoulos, 1994). It is likely that integration occurs at random sites. Given that the mammalian-cell genome is large (>109 bp), has 20% to 30% of the DNA as single-copy, and has only 0.1% of the genomic DNA containing coding sequences, it is critical to exert selective pressure to screen out the large number of nonproductive clones. Further amplification in the presence of selective pressure is
necessary to obtain clones in which multiple copies of the heterologous gene are obtained through coamplification with the dhfr or other selectable marker gene. In this respect it is wise to screen for high producers at moderate levels of methotrexate (<1 µM) to prevent enrichment of high-copy-number but unstable integrands. Stability is assessed through determination of productivity after extended growth or passaging in the absence of selective pressure in complete medium. Following transfection of cells using calcium phosphate precipitation or lipofection (Kaufman, 1990a), ∼5% to 50% of the cells in the population acquire DNA and transiently express the encoded protein over a period of days before unstable, nonintegrated DNA is lost from the cells. A smaller portion of the cells will integrate the DNA chromosomally, resulting in stable protein expression. Following selection of these stable-expressing cells by application of selective pressure, less than or equal to approximately 0.1% of the original population will express the foreign protein. It is thus imperative to use selective pressure in the production of the early stable-expressing clones to prevent overgrowth with nonproducing cells. For CHO dhfr− host–based production, the cells are grown in medium deficient in nucleosides (e.g., α-MEM medium from Life Technologies), containing serum that has been dialyzed (to remove low-molecular-weight nucleosides such as hypoxanthine), and supplemented with 0.1 to 1.0 µM methotrexate. In deficient medium, the cell’s requirement for purine and pyrimidine nucleosides is satisfied by the action of heterologous dihydrofolate reductase, which converts folate from the medium to tetrahydrofolate (FH4). FH4 and serine are then used by cellular serine hydroxymethyl transferase to provide methylene-FH4 which can go on to donate C1 units in the reactions required for inosinic acid and thymidine production. Cells that acquire the capacity to express DHFR can be selected by medium deficient in nucleosides. Stepwise increases in the methotrexate concentration with increased passaging in deficient medium will select for cells containing high copy number of DHFR (Kaufman, 1990b).
PROTEIN TRANSLATION, QUALITY CONTROL, AND COVALENT MODIFICATION Mammalian cells are complex factories with large numbers of compartments, each with specialized functions. Synthesis of proteins begins
5.9.6 Supplement 10
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Table 5.9.2
Protein Translocation and Post-Translational Modification Signals
Protein-based signal sequence
Related cell function
Example
Reference
KDELCOOH
Retention in ER
BiP, protein disulfide isomerase, prolyl isomerase
Pelham and Bienz (1982)
Transmembrane sequences Mannose-6-phosphate
Golgi retention
Coronavirus matrix protein Lysosomal hydrolases
SKLCOOH
Added in Golgi for transport to lysosome Peroxisomes
Mayer et al. (1988); Machamer and Rose (1987) Reitman and Kornberg (1981) Keller et al. (1987); Miyazawa et al. (1989)
Leader peptide
Mitochondria
RRNRRRRW (other specific signals exist, e.g., for p53 and SV40 late T antigen) Asn-X-Ser(Thr)
Nucleus
Luciferase acyl coenzyme A oxidase CoXIV (cytochrome c oxidase subunit IV) Rev
Hurt et al. (1984)
Many normal secreted and recombinant proteins
Hirschberg and Snider (1987)
Malim et al. (1989)
Cys (ER) Cys (Golgi)
N-linked glycosylation in ER (dolichol precursor addition) Golgi-based Oglycosylation Palmitoylation Myristoylation
Includes some recombinant Hart et al. (1988); Conradt proteins—e.g., IL-2, tPA et al. (1990) Membrane anchoring—e.g., HBV McIlhinney (1990) preS1
Tyr (Golgi)
Sulfation
Biological function
Rosenquist and Nicholas (1993); Han and Martinage (1992)
Signal transduction and some viral proteins—e.g., HBV core
O-Ser/Thr
Tyr (cell-surface kinases) Phosphorylation
Asn, Asp (Golgi)
Hydroxylation
Biological function
Myers et al. (1994); Albin and Robinson (1980); Aitken (1996) Stenflo et al. (1989)
Glu (Golgi)
γ-carboxyglutamate
Calcium binding—e.g., bloodcascade factors
Suttie (1985); Kaufman et al. (1986a)
in the nucleus with transcription and progresses to the cytoplasm, where translation and translocation into the endoplasmic reticulum (ER) occurs. The protein can contain sequences (motifs) that will order specific post-translational modifications as the protein progresses through the ER into the Golgi and before reaching the final destination either as a secreted protein or as a specialized resident cellular protein (see Table 5.9.2). In general, initiation of gene-transcription is the rate-limiting step. The rate of transcription elongation is constant until the movement of RNA polymerase II is stopped by termination signals. It follows that a highly expressed gene would be distinguished by the presence of a high density of polymerase molecules, resulting from high-frequency initiation. For recombinant proteins that are destined
for secretion, the translocation of expressed protein follows the so-called constitutive route (Walter and Lingappa, 1986), which is shown in part in Figure 5.9.3. A signal-recognition particle (SRP) binds to the signal sequence of the growing peptide when it protrudes into the cytoplasm, thereby halting further synthesis of the protein until the ribosome-SRP complex has bound to the docking protein (SRP receptor) at the ER membrane. Protein synthesis then continues and the peptide chain progresses into the lumen of the ER. At this point the aminoterminal signal sequence is cleaved off. Signal peptides encompass 13 to 30 amino acids. They usually have a net positive charge at the N terminus, a core of nine or more hydrophobic amino acids, and a cleavage site (Von Heijne, 1983, 1984). The tissue plasminogen activator
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in the ER. The ER provides an environment optimized for protein folding and is distinguished by a relatively high concentration of calcium (Baumann et al., 1991). The folding and stability of some proteins is calcium-dependent (Lodish and Kong, 1990). Another significant feature of the ER is that its redox potential is sufficiently high to enable disulfidebond formation (Hwang et al., 1992). It appears
(tPA) leader peptide has been used for efficient translocation (Burke et al., 1986). The protein is folded and glycosylation is initiated in the ER. Detailed discussion of the intracellular factors involved in protein translocation are given in a number of excellent reviews (Gething and Sambrook, 1992; Rothman and Orci, 1992; Wirth and Hauser, 1993; Rothman, 1994). The protein begins to fold cotranslationally
spliced mRNA
N
SRP-ribosome docking
translation (cytoplasm)
hsp70 N binding
N-linked glycoform relative to passage through ER and Golgi
mitochondria Dol-P-P
nucleus CNX
PDI PI
ER
ATP BiP sugar BiP refolding and assembly ER misfolding BiP KDEL signal salvage pathway
translocation to Golgi
transport vesicle v-SNARE
degradation pathway
coated cis Golgi bud (CGN) ARF COPs
ER
target t-SNARE ATP GTP acyl-CoA
COP-coated vesicle t-SNARE
ER
cis
hybrid type (endo H resistant)
5
medial
6
medial 7
trans 8
9
complex type endosomes/ lysosomes
Overview of Protein Expression by Mammalian Cells
2
4
medial Golgi
trans Golgi (TGN)
ER
high mannose ER (endo H sensitive) cis Golgi 3
NSF, SNAPs, GTP bp ATP, GTP, acyl-CoA, other proteins
further vesicular transport
1
cell surface
secretory/ storage vesicles
Figure 5.9.3 Intracellular protein translation and cisternal transport. Cytoplasmic translation results in a nascent polypeptide that is chaperoned by the hsp70 protein, which may direct the ribosome-peptide complex to the mitochondrion or the ER upon N-terminal peptide signaling. A signal-recognition particle (SRP) binds to the signal sequence, halting translation until it has docked at the SRP receptor. The hsp70 protein maintains the protein in a translocation-competent state, enabling membrane protrusion. Inside the ER the protein undergoes refolding assisted by protein disulfide isomerase (PDI), BiP, and calnexin (CXN). Initial N-glycosylation takes place cotranslationally in the ER. Misfolded protein, associated with BiP, is retained and eventually degraded. Transport of folded, assembled protein between the ER and Golgi cisternae occurs through a series consisting of budding, vesicle formation, and specific binding to target cisternae surfaces, through to the trans Golgi (TGN), where distribution to particular destinations occurs. Abbreviations: ARF, ADP-ribosylation factor; COPs, a family of coat proteins; Dol, dolichol; endo H, endoglycosidase H; GTP bp, GTP binding protein; NSF, NEM-sensitive cytosolic factor (identical to yeast sec 18p); SNAPs, three soluble NSF attachment proteins; v-SNARE and t-SNARE, SNAP receptors. Symbols: solid circles, N-acetyl glucosamine; open circles, mannose; solid diamonds, glucose; open squares, galactose; solid squares, sialic acid; solid triangles, fucose; circled 1, α-glucosidase I; circled 2, α-glucosidase II; circled 3, α-1,2-mannosidase; circled 4, GlcNAc transferase I; circled 5, α-mannosidase II; circled 6, GlcNAc transferase II; circled 7, α-1,6-fucosyl transferase; circled 8, β-1,4-galactose transferase; and circled 9, α-2,3-sialyl transferase.
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that glutathione is the redox buffer and that the redox potential of the ER is 20 to 100 times more oxidizing than that of the cytosol as indicated by the GSH/GSSG ratios (between 1:1 and 1:3 in the ER as compared to between 30:1 and 100:1 in the cytosol; Hwang et al., 1992). The ER contains soluble folding enzymes such as protein disulfide isomerase (PDI) and prolyl isomerase, which serve to aid the refolding process. Another group of proteins known as chaperones (Ellis and Hemmingsen, 1989) are present in the ER and appear to assist in protein folding and to ensure that incorrectly folded protein is not released from the ER, thus playing a quality-assurance role for newly synthesized protein in the cell. The best-studied chaperone is the glucose response protein BiP (or GRP 78), a member of the hsp70 family of proteins (Munro and Pelham, 1987; Hendershot et al., 1988). An ER integral membrane protein called calnexin (Bergeron et al., 1994) also appears to function in quality control of protein folding in the ER and affects the retention of incorrectly folded protein. Calnexin is unrelated to the heat-shock chaperone Hsp60, Hsp70, and Hsp90 families. As the protein is folding in the ER, N-linked oligosaccharides are added to the nascent chain via dolichol phosphate. Intramolecular disulfide bonds are formed with the assistance of protein disulfide isomerase (PDI), and the molecular chaperone BiP/GRP78 associates with the refolding peptide chain. Multiple cycles of BiP binding and release occur in a process that requires ATP. Protein folding occurs with high efficiency and with a half-time on the order of 1 min. Pulse-chase studies of hepatitis B surface antigen (HBVsAg) secretion in mouse fibroblast cells (Huovila et al., 1992) showed that the formation of HBVsAg dimers in the ER occurred rapidly, within 2 min, while formation of higher-order oligomers occurred with a halftime of 2 hr. The study also provided evidence that the rate-limiting step was likely to be passage from the ER into the Golgi, as mature intracellular glycosylated HBV was not detected. The higher-order assembly of subunits or assembly of multimeric proteins begins in the ER and is likely completed in the so-called 15° compartment (named because of accumulation of molecules there at low temperature). This appeared to be the case for HBVsAg (Huovila et al., 1992). When a monomeric subunit is extensively folded, the BiP dissociates, and for multimeric proteins such as viral membrane proteins, subunit-subunit recognition and assembly occurs prior to transport into the Golgi
complex (Doms et al., 1993). Only correctly folded protein progresses from the ER to the Golgi apparatus. Protein may be misfolded as a result of inefficient folding, mutations, heat shock, energy depletion, or addition of exogenous reductants such as dithiothreitol. These misfolded proteins usually form aggregates and typically display permanent and stable association with BiP. Aggregates are eventually degraded unless they are salvaged through a pathway that facilitates refolding from the misfolded protein species under specific conditions—e.g., altered temperature for temperature-sensitive folding mutants or where the ER environment is experimentally altered. Proteins that have successfully folded in the ER are destined for secretion to the plasma membrane or the intracellular compartments, including the ER itself. Proteins destined to be retained in the ER possess the C-terminal signal sequence KDEL (Munro and Pelham, 1987). Proteins containing the KDEL signal that escape the ER are efficiently salvaged from the Golgi through a divalent cation–dependent receptor-based salvage pathway (Rothman and Orci, 1992). Proteins destined to be integrated into the ER membrane have a signal consisting of two lysine residues separated by three and four or five amino acids apart from the C terminus (Jackson et al., 1990). Signals for Golgi retention appear to involve histidine and cysteine within the membrane-spanning domain (Aoki et al., 1992). N-linked oligosaccharide assembly begins in the ER compartment (Kornfeld and Kornfeld, 1985). In brief, N-linked glycosylation begins with synthesis of a lipid-linked oligosaccharide moiety—Glc3Man9GlcNAc2-P-Pdolichol—which is transferred to the nascent peptide chain at the relevant Asn-X-Ser/Thr sequon site (i.e., peptide sequence recognition site). Truncations of the initial oligosaccharide structure follow this initial step in the ER. The folded and primary-glycosylated protein is transported to the Golgi, where maturation of the carbohydrate chains takes place. O-glycosylation is thought to occur after the N-glycosylation in the ER or the early part of the Golgi. Differences in glycosylation of a given protein are a result of divergence in the Golgibased processing of the oligosaccharides in different cell types (Jenkins et al., 1996). The human lymphoblastoid Namalwa cell line performs O- and N-linked glycosylation of recombinant tPA efficiently to produce human-type glycosylation characteristics (Okamoto et al., 1991; Khan et al., 1995). Most CHO cell lines
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Overview of Protein Expression by Mammalian Cells
used in production of recombinant proteins do not possess active α1,3-galactosyltransferase, although the gene is present in these cells, which make low levels of galactosamine neuraminic acid. CHO and BHK cells also lack a functional α2,6-sialyltransferase enzyme and synthesize exclusively α2,3–linked terminal sialic acids via α2,3-sialyltransferase, in contrast to human cell lines, which contain both enzymes (Lee et al., 1989). In CHO cells the dominant sialic acid found is N-acetyl neuraminic acid. Transport of the protein through the secretory and endocytic pathways is mediated by small vesicles that bud off from one compartment and fuse to the next compartment (Rothman and Orci, 1992). The cis Golgi stack (CGN) and trans Golgi stack (TGN) are defined areas of the entire Golgi network. The CGN and TGN constitute the entry and exit faces, respectively, and are involved in sorting and distribution of the protein. The CGN includes the KDEL salvage pathway for ER-based proteins. The TGN is where proteins with differing final destinations—e.g., lysosomes, secretory vesicles, and plasma membrane—diverge. The Golgi consists of stacks of cisternae where carbohydrate-chain trimming and elongation as well as other covalent modifications occur. The mechanism of flow through the Golgi cisternae appears to involve assembly of coated vesicles, which then transfer to the next compartment with concomitant loss of the coat. The loss of coat is linked to GTP-dependent hydrolysis, while subsequent fusion at the destination cisternae is ATP-dependent and also involves an N-ethylmaleimide (NEM)-sensitive enzyme, Sec18p. The specificity of vesicle targeting involves a docking mechanism based on binding of soluble NEM-sensitive attachment protein (SNAP) to a receptor designated as SNARE. Heterogeneity in the SNARE family allows for specialization and the ability to direct a given protein in the appropriate direction by selective fusion to progressive cisternae and ultimately the destination membrane. Although it is usually desirable to obtain secretion of protein out of the cell prior to purification, in some cases the goal may be to express a protein receptor that would be transported to the cell-surface membrane. This was described for the production of the erythropoietin EPO receptor (EPO-R) in transfected hematopoietic cells (Hilton et al., 1995). The polypeptide was mutated to improve EPO-R refolding in the ER and increase cell-surface expression. The protein was obtained as a crude
extract from the cell culture medium following the completion of the cell culture process. The glycoprotein may then be purified for further physicochemical analysis or testing in biological assays and animal studies. Some proteins are not normally secreted by mammalian cells. This is the case for ER-resident proteins such as BiP, prolyl isomerase, and protein disulfide isomerase, which contain the C-terminal ER-retention peptide sequence KDEL. Also, a number of proteins including receptors and signal-transduction proteins are expressed and retained in the cell or on the cell surface. In cases where specific inhibitors of glycosylation enzymes are added to the culture medium, proteins that are normally secreted may be poorly secreted and thus accumulate in the ER or Golgi of the cell (Hickman and Kornfeld, 1978). Release of these intracellular or surface proteins requires disruption of the cell using a method capable of breaking the cell membrane (homogenization); in cases where membrane association is expected, nonionic detergents at 0.1% to 0.5% (v/v) are used in the lysis buffer. Covalent modifications of amino acids by mammalian cells are numerous and to some extent may be a feature of a particular cell type. For example the γ carboxylation of factor VIII, which is essential for clotting-cascade activity, can be accomplished in the BHK and human 293 cell lines. This microsome-based, vitamin K–dependent modification was nonfunctional in CHO cells that had been transformed with complementing genes for carboxylation (Wirth and Hauser, 1993). Other processing events occur during passage of the protein from the ER to the Golgi. These involve removal of certain residues from the carbohydrate chains—e.g., glucose and mannose trimming. Other sugars are added to the carbohydrate chains as the protein passes through the medial and trans Golgi—e.g., addition of N-acetylglucosamine, galactose-N-acetate, fucose, galactose, and sialic acid. O-linked glycosylation is important for stabilization of some proteins— e.g., low-density-lipoprotein (LDL) receptor (Kingsley et al., 1986).
GROWTH OF MAMMALIAN CELLS FOR PROTEIN EXPRESSION Mammalian cell culture media are complex in comparison to the simple defined media used for propagation of bacteria and yeast. This complexity stems from the need to satisfy the many functions required for normal growth and
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metabolism of mammalian cells (Bettger and McKeehan, 1986). Nutrients must satisfy the requirements for growth, maintenance of the cell, and expression of products. Additional specific growth factors such as insulin or insulin-like growth factor (IGF) are usually required. Minimal nutritional requirements were first defined in the 1950s (Eagle, 1955). These early studies showed that thirteen amino acids (Arg, Cys, Gln, His, Ile, Leu, Lys, Met, Phe, Thr, Trp, Tyr, and Val), eight vitamins (biotin, folic acid, niacinamide, pantothenic acid, pyridoxine, riboflavin, and thiamine), and various salts and minerals (Na, K, Ca, Mg, Cl, PO4, Fe, Zn, Se, and possibly Cu, Mn, Mo, and V, which are usually present as trace contaminants) are required. The commonly used CHO derivatives DXB11 and DG44 have an absolute requirement for proline that results from a genetic deficiency in the conversion of glutamic acid to glutamic γ-semialdehyde. Inorganic ions may function as catalysts or exert physiological effects. Glucose (5 to 20 mM) and glutamine (0.5 to 5 mM) are commonly used to provide the energy source for the cells. These are consumed at a rate proportional to the viable-cell concentration. Cultured mammalian cells do not use the complete tricarboxcylic acid (TCA) cycle, and the glucose-consumption rate is lower than the glutamine consumption rate. Consumption of glucose yields lactate and consumption of glutamine yields ammonia as the metabolic byproduct. Other nutrients are consumed at a rate proportional to the rate of increase in cell density. Yield-based uptake rates for amino acids by CHO DG 44 cells range from 0.1 mM tryptophan/109 cells to 0.8 mM arginine/109 cells at a specific growth rate (the rate of growth divided by the cell density) of 0.25 to 0.30 days−1 (D. Gray, unpub. observ.), reflecting the relative incorporation frequency of the individual amino acids into general protein. CHO cells, like most mammalian cells, produce alanine as a result of glutaminolysis. Accumulation of ammonia, as a result of glutamine degradation in the medium and consumption by cells, may become inhibitory to cell growth and ultimately limit the density of the culture (Butler and Spier, 1984). In addition, physiological parameters such as pH, osmolality, and redox potential must be kept within acceptable limits. The use of serum is often advocated and has been regarded as a source of important components such as hormones, lipids, and growth factors. These serum-borne ingredients provide some consumable nutritional components as well as mito-
gens or growth factors that overall are not consumed by the cells (Klinman and McKearn, 1981). A further level of complication in mammalian-cell medium design occurs through the need to maintain balanced relationships between nutrients while maintaining the osmolality of the medium within the physiological constraints (290 to 380 mOsmol/kg). Recent medium development has focused on reducing the need for high levels of serum (Jayme, 1991). Fetal bovine serum (FBS) is the most frequently used version of serum and can be obtained from HyClone (see SUPPLIERS APPENDIX). FBS costs ∼$500 to $600 per liter and is a factor contributing to the high cost of producing commercial clinical lots of protein. There may be batch-to-batch variation in serum which necessitates screening by small T-flask or shake-flask growth studies with the chosen mammalian cell line. Serum contains ∼35 mg/ml of protein, of which ∼60% is albumin. Thus if 10% serum were added to medium, the level of initial exogenous protein in the media would be 3 to 5 mg/ml. Growth of cells in low serum (<0.5% v/v) has been accomplished through step-wise adaptation of cells to growth in medium with low or zero serum. The important functions of serum are replaced by the addition of the lipid precursors choline, inositol, and ethanolamine, the polyamine putrescine, and recombinant insulin (Nucellin-Zn from Eli Lilly) at 1 to 5 mg/liter (Jenkins, 1990) along with FeCl3/citrate (Keenan and Clynes, 1996). A number of serum-free media have been developed and are now commercially available from, e.g., BioWhittaker. Another approach to implementing protein-free media has been the genetic modification of CHO cells through incorporation of the heterologous genes coding for transferrin and IGF-1 (Pak et al., 1996). As a result of cellular expression of the IGF-1 and transferrin, the cell is then able to grow, without addition of exogenous proteins, in a modified DMEM/F12 medium supplemented with sodium selenite. A major concern in producing recombinant protein in protein-free medium is the potential for proteolysis and glycosidase activity. Such activity results in processing of the polypeptide chain and degradation of the carbohydrate moiety (Gramer and Goochee, 1993; Teige et al., 1994). Serum contains endogenous protease inhibitors—e.g., α2-macroglubulin (Barrett, 1981)—and for this reason may be a valuable reagent in its own right in cell culture media
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Overview of Protein Expression by Mammalian Cells
when the produced protein is susceptible to proteolysis. Sialidases retain significant activity at pH 7.0 and are active in cell culture supernatants. Caution should be taken to minimize cell lysis during the culture process if the goal is to minimize desialylation of the expressed protein. This may be achieved by harvesting the supernatants during the exponential-growth phase prior to the beginning of the decreasing-viability phase, chilling it, and adding sialidase inhibitors such as 2,3-dehydro-2deoxy-N-acetylneuramic acid to the supernatant (Gramer et al., 1995). In batch culture in serum-free medium, cell viability may decrease rapidly at the end of culture, thereby releasing intracellular proteases and glycosidases into the culture medium, which may in turn cause rapid proteolysis and carbohydrate degradation of the product. Production of sialylated forms of glycoproteins is important in clinical applications to avoid rapid clearance by asialo- and asialogalactoprotein receptors in the liver. The genes for some of the key enzymes in the glycosylation pathway have been transfected into CHO cells (Lee et al., 1989) in attempts to tailor the glycoforms. A number of CHO-glycosylation mutants have been made available that exhibit a defect in one or more steps in the pathway (Stanley, 1989). Erythropoietin (EPO) expressed in the CHO mutant Lec 3.2.8.1. exhibited less heterogeneity than EPO produced in wild-type CHO cells (Stanley, 1991). Studies with CHO-320 cells in glucose-limited chemostat culture (using RPMI-1640 medium) at a low dilution rate of 0.008 day−1 (i.e., volume of fresh medium added to the reactor per day divided by the reactor medium volume) showed a deviation of the specific growth rate from the dilution rate and an increase in specific cell-death rate (i.e., rate of cell death per liter per day divided by the number of viable cells per liter), suggesting that a minimum specific growth rate of 0.011 day−1 existed (Hayter et al., 1993). In the same study, the specific rate of γ interferon (γ-IFN) production increased with the specific growth rate (µ), indicating growth-related product production. The pattern of γ-IFN glycosylation was similar at all growth rates except the lowest, where an increase in nonglycosylated γ-IFN was observed. This was further addressed using batch culture (Leelavatcharamas et al., 1994), where it was concluded that the growth-associated production was a reflection of a cell cycle–related phenomenon whereby lower growth rate resulted in a prolongation of the cell-cycle time with an
increase in the relative frequency of the G1 cells in the culture population. It was suggested that the G1 phase is the least productive phase in the cell cycle of CHO cells. Similarly S phase–specific synthesis of dihydrofolate reductase in nontransfected CHO-K1 cells was observed in which the maximum peak of DHFR activity coincided with the maximum rate of DNA synthesis in late S phase (Mariani et al., 1981). The expression of recombinant β-galactosidase by the CMV late promoter in CHO βG72-16 was shown to be S-phase related (Gu et al., 1993). It is likely that CHO cells will generally exhibit an S-phase dependency for expression of foreign genes using the typical promoter elements such as the CMV late promoter, and that a direct correlation of specific heterologous protein production rate with specific growth rate will be observed (Bock et al., 1993; Gu et al., 1996). As growth rate increases, the ratio of S/(G1 + G2/M) increases (Hooker et al., 1995) and the frequency of cells in productive S phase in the culture increases. A given glycoprotein produced in cell culture by mammalian cells will present a population of molecules exhibiting differences in glycosylation. The existence of families of structurally related yet distinct carbohydrate chains makes structural analysis of a distinct species complicated, since it is difficult to separate and purify the structural variants. Variation may result from incomplete occupancy of a directed site on the protein and from carbohydrate-structure variations imparted to the cells via cell culture–based conditions. Furthermore the action of glycosidases in the extracellular environment, particularly during batch culture, may degrade the carbohydrate structure of secreted glycoprotein, leading to a wider range of carbohydrate structures in the population. Changes in the oligosaccharides of the glycoprotein may lead to diminished biological efficacy in vivo—e.g., through increased clearance—or may enhance unwanted antigenicity of the molecule in a clinical application (Goochee et al., 1991). Antibodies produced in CHO cells in serumcontaining medium exhibited more galactosylation compared to antibodies produced by cells grown in serum-free medium (Lifely et al., 1995). The culture-medium glucose concentration affected the degree of glycosylation of γ-IFN produced in CHO cells in continuous culture (Hayter et al., 1992). Lipid supplements, with or without lipid carriers such as lipoprotein, appeared to increase N-glycosylation-site occupancy in γ-IFN (Jenkins et al.,
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Table 5.9.3
Estimated Quantity of Foreign Protein Produced by a Given Production Systema
Vol. of medium in system
System
Total cells in system (vol. × conc.)
Total product in systemb
Batch T-flask (T-175): 75 ml
1.8 × 107
0.5-1 mg
100 ml
2 × 108
2-4 mg
1000 ml
1 × 109
10-20 mg
Batch 5-liter spinner flask fitted with O2 frit spargerc
1000 ml
2×
20-40 mg
Batch 15-liter bioreactor with pH and O2 controlc
10 liters
2 × 1010
200-400 mg
5-20 liter perfusion per day
1 × 1011
100-400 mg per day
175 cm2 attached growth in DMEM/F12/10% serumc Batch shake flask (250 ml), 0.51% serumc Batch 5-liter spinner flaskc
Continuous-mode 15-liter perfusion reactor with pH and O2 control and cell retention aQuantities
assume a specific productivity of 1 to 4 mg/109 viable cells per day.
bProduction cBatch
109
of recombinant intracellular protein may achieve ∼0.1% of total cell protein.
culture would be performed for 4 days after inoculation of 1 × 105 viable cells/ml.
1994). In perfusion culture of BHK-21 cells producing a recombinant interleukin 2 (IL-2) mutant, nutrient limitations (i.e., glucose, amino acids, and dissolved O2) led to shortterm changes in the general level of glycosylation, but the specific type of sugar incorporated was largely unchanged (Gawlitzek et al., 1995). Direct intervention in cell physiological processes has an impact on glycosylation. A decreased protein-synthesis rate following addition of cyclohexamide improved the glycosylation-site occupancy of recombinant prolactin produced by C127 cells (Shelikoff et al., 1994). Low concentrations of dithiothreitol, which prevent cotranslational disulfide-bond formation in the ER, led to complete glycosylation of tPA at a tripeptide sequon that normally exhibited variable occupancy (Allen et al., 1995). This suggests that refolding may in itself limit the accessibility of glycosylation sites. Sodium butyrate is sometimes used to improve protein synthesis, but it can change glycosylation by inducing the GlcNAc-transferase involved in O-glycosylation (Datti and Dennis, 1993) and by increasing sialyltransferase activity in recombinant CHO cells (Chotigeat et al., 1994; Gebert and Gray, 1995).
SCALE OF OPERATION Batch growth in shake flasks or spinner flasks may provide titers in stable producer cell lines of 10 to 50 mg/liter, thus yielding up to
30 to 150 mg of crude secreted product which may then be purified from the culture supernatant. Following purification, the researcher may recover ∼10% to 30% of the protein. Thus, small laboratory-scale cell culture may provide 3 to 50 mg of pure protein (>90% purity). Larger quantities of material require use of bioreactors in which pH, dissolved O2, temperature, and stirring may be controlled. Should the user wish to control the cell growth rate during the production of the product, a continuous reactor may be used at a fixed dilution rate. In cases where high cell density is desired, the mode of operation would be continuous medium perfusion with cell retention in the bioreactor. Tables 5.9.3 and 5.9.4 provide some guidance for the likely scale requirements for a given objective. The bioreactor environment may influence the product quality. Mild hypoxia may decrease the specific activity of sialyl transferase and hence the sialylation of the oligosaccharide chain, as was observed with recombinant human follicle-stimulating hormone produced in CHO cells at 10% dissolved oxygen saturation (Chotigeat et al., 1994). It appears that pH values within the range 6.9 to 8.2 have no dramatic effect on glycosylation, but clearly optimal growth would require control of pH within a narrower range, typically 7.0 to 7.4. Metabolic byproducts may be the limiting factors in achieving high cell density in perfu-
Production of Recombinant Proteins
5.9.13 Current Protocols in Protein Science
Supplement 10
Table 5.9.4
Typical Protein Requirements for Various Analyses
Assay ELISA
<5 µg
Identification of activity possible with crude extract
Amino acid analysis and sequencing (including development of method)
20 µg
Pure protein required; thus starting requirement would be >100 µg in order to account for purification losses
Carbohydrate analysis Extinction-coefficient determination
20 µg 500 µg
Pure protein required Pure protein required; thus starting requirement would be 3-5 mg
50 mg
Must obtain suitable purity of product to prevent collateral effects on physiology due to contaminants
Biological testing in animal-based model Human clinical trials
Overview of Protein Expression by Mammalian Cells
Quantity of protein Comments required
1-100 g
sion culture. Increases in ammonium ion concentration to 10 mM in the culture medium resulted in substantial growth inhibition in hybridoma cells (McQueen and Bailey, 1990) and appeared to inhibit growth in most cell lines. The likely cause of this is the transport of ammonium ion by membrane-bound ion pumps and ultimate acidification of the cytoplasm (Martinelle et al., 1996). Lower levels of ammonia (∼2 mM) may exert influences on glycosylation (Andersen et al., 1994). One approach to eliminating this potential inhibition of cell growth is through adaptation to a nonammoniagenic medium. This medium would contain glutamate or 2-oxoglutarate in place of glutamine (Hassel and Butler, 1990). Lactate generated from glucose can also reach growth inhibitory levels at >15 mM. Optimization of medium and perfusion rate may enable reduction in the levels of lactate and ammonia, allowing higher cell density. Lowering the glucose level may reduce the yield of lactate, but concomitant limitation of glycosylation may occur. CO2 may accumulate in high-density cultures, thereby exerting an inhibitory effect on the culture (Gray et al., 1996). Limitations that are due to CO2 accumulation may be overcome by choosing an optimal sparging strategy. Finally mammalian cells are weaker and larger than bacterial cells and limits to the degree of agitation are important to prevent cell damage and death. Where agitation is used in reactors or in sample handling, cell damage at the gas-liquid interface as well as bubble damage can be reduced by adding low levels—
>95% purity; requires extensive testing to ensure safety
<0.1% (w/v)—of the surfactant pluronic polyol (Murhammer and Goochee, 1990).
SUMMARY Both yeast and E. coli heterologous protein expression are very well defined. The components of these expression systems are easily obtained through many commercial vendors. Use of these systems enables the researcher to obtain large amounts of pure material in a short period of time. In contrast, mammalian expression systems are not comprehensively defined or understood and require a significant investment of time before heterologous protein is obtained. At the present time the mammalian expression systems are not available as turnkey, catalog-based systems. The user must define the components for a particular application. Although this complexity exists with the mammalian cell–based systems, they are uniquely important for the production of glycoproteins that possess oligosaccharides close to or identical to those of human origin. Proteins secreted by mammalian cells possess correct conformations and exhibit full biological activity. The ability to transfer foreign DNA into these cells enables a greater understanding of the mammalian cell transcriptional, translational, and post-translational machinery. The ability to incorporate foreign gene sequences into a mammalian-cell chromosome and the ability to grow mammalian cells in suspension in large-scale bioreactors has enabled the production of large quantities of pure clinically important proteins.
5.9.14 Supplement 10
Current Protocols in Protein Science
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Production of Recombinant Proteins
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Overview of Protein Expression by Mammalian Cells
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Current Protocols in Protein Science
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Production of Recombinant Proteins
5.9.17 Current Protocols in Protein Science
Supplement 10
Overview of Protein Expression by Mammalian Cells
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Contributed by David Gray Chiron Corporation Emeryville, California
5.9.18 Supplement 10
Current Protocols in Protein Science
Production of Recombinant Proteins in Mammalian Cells
UNIT 5.10
Mammalian cells are the relevant hosts for expression of properly folded glycoproteins that possess post-translational modifications and that have biological activities similar to those of proteins derived from human cells (UNIT 5.9). Increasing interest in the production of glycoproteins (Chapter 12) for clinical application and as reagents for the study of biological mechanisms has led to greater use of mammalian cell protein expression systems. In response to this, developments in molecular biology and process science have provided mammalian cell expression vectors, cell lines, and growth media—and have established methods for high-density suspension culture in compact bioreactors. When DNA is transferred into a mammalian host cell, the genes may be expressed without being integrated into the chromosomal DNA (transient expression). A small percentage of cells incorporate the DNA into a transcriptionally active chromosomal locus and may express the gene on a permanent basis from generation to generation (stable expression). In UNIT 5.9 it was stated that the strategy for expression is dependent on the ultimate goal of the project. When one-time production of a few hundred micrograms of purified protein is sufficient, then rapid, transient (“burst production”) expression of the gene in COS-7 cells (Aruffo, 1997), the vaccinia system (Earl and Moss, 1998; Earl et al., 1998a,b; Elroy-Stein and Moss, 1998; Moss and Earl, 1998), Sindbis virus (Invitrogen), or Semliki Forest virus (Liljestrom and Garoff, 1995) are appropriate. The best strategy for consistent production of larger quantities of pure protein is stable expression. Popular hosts for stable expression are Chinese hamster ovary (CHO) cells, baby hamster kidney (BHK-21) cells, myeloma cells, and the transformed kidney cell line 293. Protocols for stable production in CHO cells are described in this unit. Typical methods for transfection using commercially available plasmid expression vectors (see Table 5.10.1) are described, along with methods to select for stable expression and methods for amplifying the expression level in the transfected cell. Following this, procedures are presented for efficient cell growth to obtain significant amounts of protein product. A general flow scheme of the overall process is provided in Figure 5.9.1. Stable expression is obtained when the gene of interest and associated expression elements (cis elements) are incorporated into a transcriptionally active region of the host cell chromosome. Production of the plasmid vector for use in transfection is usually performed in E. coli cells and the plasmid is purified using standard column chromatography methods (see Basic Protocol 1). DNA is transferred into the cell by lipofection (see Basic Protocol 2) or electroporation (see Alternate Protocol 1). Derivation of stable transfectants requires selection of clones producing the recombinant protein (see Basic Protocol 3, Support Protocol 1, and Alternate Protocol 2) and single-cell cloning (see Support Protocol 2), and as such is a more laborious process than the repeated transientexpression approach. Significant effort and time is needed for several rounds of gene amplification and selection because cell cloning and selection is required at each round. Although the additional work is a possible deterrent to use of this approach, it is possible to produce larger quantities of protein in this way. The other simple advantage of stable transfection is that cell banks can be produced, which enable reproducible production of the product. These advantages will more than provide a dividend after the initial investment of time. To obtain high-level expression, the gene amplification and selection processes can be combined (see Basic Protocol 4). Growth of the production cells in spinner flasks and/or bioreactors is necessary to obtain sufficient total protein for further purification and study. Procedures for adapting cells to Contributed by Su Chen, David Gray, Johnny Ma, and Shyamsundar Subramanian Current Protocols in Protein Science (1998) 5.10.1-5.10.41 Copyright © 1998 by John Wiley & Sons, Inc.
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Table 5.10.1
Useful Commercially Available Vectors for Transfection of CHO Cellsa
Poly(A)
E. coli Plasmids and Selectable selectable suppliers marker maker
Dual-plasmid SV40, hCMV vector cotransfection
SV40
Neo, Pac
Amp
pSI, pCI (Promega); pPUR, pSV2neo (Clontech); pSVK3 (Pharmacia),
Single-plasmid vector (regular cassette)
hCMV, RSV, SV40, MMTV
SV40, BGH
Neo, Zeo
Amp, Zeo
pcDNA3.1, pZeoSV2, pRc/CMV2 , pRc/RSV (Invitrogen); pMAMneo (Clontech); pCI-neo (Promega),
hCMV
SV40
GS
Amp
pEE14 (CellTech)
Single plasmid vector (dicistronic cassette)
hCMV
BGH
Neo, Hyg
Amp
pIRES1neo, pIRES1hyg Uses encephalomyo(Clontech) carditis virus IRES sequence
Special plasmid vectors
pCMV
BGH
Zeo
Amp
pSecTag (Invitrogen)
Secretion of proteins lacking signal peptides
pHook (Invitrogen)
Isolation of positive clones through magnetic affinity cell sorting (MACS)
pTracer-CMV (Invitrogen)
Isolation of positive clones using GFP and FACS
Strategy
Promoter
Notes
Amplification using glutamine synthetase with MSX
aAbbreviations: hCMV, human cytomegalovirus (CMV) major immediate early promoter; Hyg, hygromycin phosphotransferase B gene; MSX, methionine sulfoximine gene; Neo, aminoglycoside phosphotransferase II gene; Pac, puromycin resistance gene; RSV, Rous sarcoma virus long terminal repeat; Zeo, Zeocin resistance gene.
suspension conditions and low-serum media are included (see Basic Protocol 5) as well as procedures for growing cells in various vessels and in batch culture (see Basic Protocols 6 and 7), and in continuous and perfusion cultures (see Alternate Protocol 3). Most often, the product is secreted into the medium and must be separated from the cells prior to subsequent storage or purification (see Basic Protocol 8). In some cases, the protein of interest may be intracellular or membrane-associated, and thus the cells become the product and must be harvested from the culture (see Basic Protocol 9). Support protocols describe freezing of cells (see Support Protocol 3), determination of growth rates (see Support Protocols 4 and 5), determination of specific productivity of cells (see Support Protocols 6 and 7), preparing samples for assay (see Support Protocol 8), and setting up a 10-day shaker-flask growth curve (see Support Protocol 9). NOTE: Milli-Q purified water or equivalent is used for all aqueous buffer reagents. NOTE: All reagents and equipment coming into contact with live cells must be sterile, and proper sterile technique should be used accordingly. Production of Recombinant Proteins in Mammalian Cells
NOTE: All culture incubations are performed in a humidified 37°C, 5% CO2 incubator unless otherwise noted.
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PLASMID PURIFICATION BY ALKALINE-LYSIS/ANION-EXCHANGE CAPTURE
BASIC PROTOCOL 1
The gene of interest (in the form of cDNA) is inserted into the multiple cloning region of the plasmid (which has been constructed with a number of restriction sites) after any required modifications in the sequence. Commercially available plasmids provide various choices of elements; a subset are shown in Figure 5.10.1. Sufficient purified plasmid is produced in competent E. coli cells (e.g., strain DH5α) using the preparation approach discussed here. The alkaline-lysis/anion-exchange capture method is commonly used to prepare large amounts of DNA for transfection of host cells. A number of commercially available kits are available, as mentioned below, making this method quick and easy to use. Following DNA preparation, transfection of the CHO cells is performed using lipofection (see Basic Protocol 2) or electroporation (see Alternate Protocol 1).
A dual-plasmid vector cotransfection gene of interest
poly (A)
selectable marker
poly (A)
B single plasmid vector dicistronic cassette gene of interest
intron
IRES
selectable marker
poly (A)
multiple cassette gene of interest
poly (A) promoter
selectable marker
poly (A)
C active-site directed vectors weak
strong gene of interest
splice-site intron
selection marker sequence 1 selectable marker
splice-donor site
poly (A)
gene of interest
selection marker sequence 2 poly (A)
splice accepter site
Figure 5.10.1 Expression vector elements for common transfection strategies. (A) Dual- and (B) single-plasmid vectors are shown for commercial vectors types described in Table 5.10.1. The active site–directed vectors (C) are reported throughout the literature (Lucas, 1996; Wurm et al., 1996); although not currently available as commercial catalog items, they may provide advantages in obtaining high expression levels. Abbreviations: IRES, internal ribosomal entry site.
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Materials cDNA containing gene of interest Plasmid with appropriate cis-acting elements (Fig. 5.10.1 and UNIT 5.2) Competent E. coli cells (e.g., strain DH5-α; Life Technologies) LB medium (UNIT 5.2) containing antibiotics for selection (e.g, 100 µg/ml ampicillin, 40 µg/ml kanamycin, or 25 µg/ml tetracycline, depending on plasmid kit; all antibiotics available from Sigma) Resuspension buffer (see recipe) Lysis solution: 0.2 M NaOH/1% (w/v) SDS Neutralization solution: 1.3 to 3.0 M potassium acetate, pH 4.8 to 5.5 Isopropanol TE buffer (APPENDIX 2E) 80% ethanol Sterile glass test tubes for bacterial culture Environmental shaker (New Brunswick Scientific) 3-liter Erlenmeyer flask Beckman J2-21 centrifuge with JA-10 rotor or Sorvall RC-5B centrifuge with GS-3 rotor (or equivalent) and appropriate polycarbonate centrifuge bottles UV/visible light spectrophotometer (e.g., Perkin-Elmer) Additional reagents and equipment for ligation of linkers to cDNA (Klickstein and Neve, 1991), plasmid selection and insertion of DNA into plasmid vectors (UNIT 5.2), transformation of E. coli (UNIT 5.2), and purification of DNA by anion-exchange chromatography (Budelier and Schorr, 1998) NOTE: Several DNA plasmid preparation kits are available commercially—e.g., Plasmid Maxi Kit (Qiagen), Wizard Maxipreps (Promega), and FlexiPrep Kit (Pharmacia Biotech). Kits include resuspension buffer, lysis solution, and neutralization solution as described in the protocol. Prepare transformed cells 1. Ligate linkers to cDNA containing gene of interest. Insert the cDNA into an appropriate plasmid vector and transform E. coli host cells with the vector (UNIT 5.2). Ligation of linkers to cDNA is described in Klickstein and Neve (1991).
2. Prepare a starter culture by transferring a single colony from a selective agar plate containing transformants (UNIT 5.2) into a sterile glass test tube containing 5 ml LB medium with the appropriate selective antibiotic. 3. Incubate 6 to 8 hr at 37°C with vigorous shaking (on environmental shaker set to ∼300 rpm) until a culture with an optical density of 2 to 4 at 600 nm (OD600) is obtained. 4. Add 1 ml of this starter culture to a 3-liter Ehrlenmeyer flask containing 1 liter LB medium with the same selective antibiotic used in step 2. Incubate culture 12 to 16 hr at 37°C with vigorous shaking (∼300 rpm). 5. Transfer cell suspension to a polycarbonate centrifuge bottle. Harvest cells by centrifuging 15 min at 6,000 × g (6,000 rpm in JA-10 or GS-3 rotor), 4°C. Decant supernatant and resuspend pellet in 30 ml resuspension buffer.
Production of Recombinant Proteins in Mammalian Cells
Isolate plasmid DNA 6. Add 30 ml of lysis solution to resuspended cells. Mix thoroughly by inverting (not vortexing) and incubate ∼10 min at room temperature. Cell lysis is complete when solution becomes relatively clear and viscous.
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7. Add 30 ml of neutralization solution to the lysed cells and mix immediately by inverting. 8. Centrifuge 20 min at 10,000 to14,000 × g, (7,500 to 9,000 rpm in JA-10 or GS-3 rotor), 4° to 8°C. This step separates the cell debris from the lysate. The plasmid will be in the lysate.
9. Precipitate crude DNA from the clear lysate by adding 0.5 vol isopropanol, then centrifuge 30 min at 15,000 × g (9,200 rpm in JA-10 or GS-3 rotor), 4° to 8°C. Mark the region in the centrifuge tube where the pellet is expected to stick, as it may not be visible after isopropanol precipitation.
10. Dissolve DNA pellet in 2 ml TE buffer. Purify plasmid DNA 11. Apply resuspended DNA solution to an anion-exchange resin (e.g., as provided with Qiagen Plasmid Maxi Kit; also see Budelier and Schorr, 1998). The plasmid DNA will bind to the resin. Budelier and Schorr (1998) describes the anion-exchange procedure for DNA.
12. Wash resin with 80% ethanol or with the wash buffer provided with the kit (Budelier and Schorr, 1998). 13. Elute DNA from resin with TE buffer or the elution buffer provided with the kit. Determine DNA concentration by measuring OD260 (50 µg/ml = 1.0 OD260) using a UV/visible light spectrophotometer. The eluted DNA can be used directly in transfection of host cells (see Basic Protocol 2 and see Alternate Protocol 1) or stored at −20°C.
TRANSFECTION OF CELLS BY LIPOFECTION A good method for transfecting CHO cells and adherent cells in general is lipofection. This technique can typically achieve a 50% transfection efficiency (Subramanian and Srienc, 1996), although the value depends on the sensitivity of detection. In the process of lipofection, plasmid DNA is complexed with cationic lipids; the lipid then interacts with the cell membrane and delivers the DNA to the cell. In the protocol below, the lipid reagent used is Lipofectamine, which has been very effective with CHO cells. There are several other lipids available commercially from Life Technologies, Boehringer Mannheim, and Invitrogen, which may be more suitable for other cell types.
BASIC PROTOCOL 2
Materials Chinese hamster ovary (CHO) cells (Table 5.9.1) αMEM medium with and without 10% FBS (see recipe) Lipofectamine (Life Technologies; store up to 6 months at 4°C) Plasmid DNA to be transfected (see Basic Protocol 1) 100-mm tissue culture dishes or T-75 flasks Polystyrene tubes Additional reagents and equipment for growing mammalian cells in culture (APPENDIX 3C) 1. Plate CHO cells, at a density of 1.5–3.0 × 106 viable cells per 100-mm tissue culture dish or T-75 flask, using αMEM medium/10% FBS. Incubate ∼24 hr to achieve 50% 80% confluency (APPENDIX 3C).
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2. Dilute 48 µl of Lipofectamine in 0.8 ml serum-free αMEM in a polystyrene tube. Dilute 8 µg of DNA to be transfected in 0.8 ml serum-free αMEM in a separate polystyrene tube. 3. Mix lipid and DNA solutions and incubate 45 min at room temperature to form complexes. Dilute the complexes in 6.4 ml serum-free αMEM. 4. Wash cells with 10 to 20 ml serum-free αMEM per dish, remove the medium, then add the diluted lipid-DNA complexes to cells. Alternatively, the lipid-DNA complexes can be directly diluted in the tissue culture dish.
5. Incubate cells with the complexes for 5 to 6 hr under normal culture conditions. Replace complexes with αMEM/10% FBS, then incubate 24 to 48 hr. Note that the recipe for áMEM (see Reagents and Solutions) includes no antibiotics. Cells are ready for either transient analysis or selection (see Basic Protocol 3) 24 to 48 hr post-transfection. ALTERNATE PROTOCOL 1
TRANSFECTION OF CELLS BY ELECTROPORATION Electroporation allows delivery of DNA into cells by creation of transient pores in the cell membrane. In many cell types that grow in suspension, the lipofection method (see Basic Protocol 2) does not result in high transfection efficiencies. In these situations electroporation is an appropriate choice, as it has been shown to work with numerous cell types (both suspension and adherent). Electroporation may also be the preferred method when single-copy integration is desired, since this technique delivers lower amounts of DNA to the cell than lipofection. Both methods work well in the case of CHO cells. The authors have found that the Bio-Rad Gene Pulser II, equipped with a capacitance extender, is suitable for electroporation of mammalian cells. This instrument delivers a decaying exponential pulse. Other instruments—e.g., from BTX and Invitrogen—can also be used. Additional Materials (also see Basic Protocol 2) Electroporator and electroporation cuvettes with 0.4-cm electrode gap (Bio-Rad) Additional reagents and equipment for growing mammalian cells in culture, trypsinizing cells, and counting viable cells by trypan blue exclusion (APPENDIX 3C) 1. Harvest exponentially growing (subconfluent) cultures by trypsinization and count cells (APPENDIX 3C). Centrifuge cells 10 min at 200 to 500 × g, room temperature. Resuspend pellet in αMEM/10% FBS medium at 1–2 × 107 viable cells/ml. Dispense 0.5-ml aliquots into electroporation cuvettes. 2. Add 10 to 20 µg of DNA to be transfected and incubate 10 min on ice. 3. Resuspend cells by flicking, then pulse at high capacitance (∼1000 µF) at 200 to 400 V (which should result in a time delay in 5 to 20 msec range). Use 10- to 50-V steps. 4. Incubate electroporated cells 10 min at 37°C. Transfer cells into a T-75 flask or 100-mm dish with 20 ml of medium and incubate 24 to 48 hr under normal culture conditions (APPENDIX 3C). The cuvette may need to be rinsed for complete removal of cells.
Production of Recombinant Proteins in Mammalian Cells
Cells are ready for either transient analysis or selection (see Basic Protocol 3) 24 to 48 hr post-transfection.
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SELECTION FOR NEOMYCIN PHOSPHOTRANSFERASE (NPTII) WITH GENETICIN (G418)
BASIC PROTOCOL 3
Following the transfection, cells are subjected to the selection process, in which they are exposed to a drug that allows cells expressing the resistance marker to survive while killing the rest. The commonly used dominant selection markers include neomycin phosphotransferase (NPTII, aminoglycoside phosphotransferase II) which enables growth in the presence of G418 (geneticin). Other selection approaches in common use are the glutamine synthetase (GS) system, which uses growth in glutamine-free medium for selection (Kingston et al., 1993). Another alternative and common approach is the use of a recessive marker such as dihydrofolate reductase (DHFR) with initial selection in nucleoside-free medium (see Basic Protocol 4). Selection, as described in this protocol, and screening (see Alternate Protocol 2), are the two different methods that help isolate cells that express the marker gene. The marker gene is an indirect indication that the protein of interest is also being expressed. In selection, the drug kills cells that do not express the marker. This protocol describes selection with the commonly used dominant selection marker neomycin phosphotransferase II. Materials Culture of transformed cells at 50% to 70% confluency, in T-25 or T-75 flasks (24 to 48 hr post-transfection; see Basic Protocol 2 or Alternate Protocol 1) Selective medium: αMEM medium (see recipe) containing 5% to 10% FBS and 0.1 to 1.0 mg/ml G418 (Geneticin, Life Technologies; add from 50 mg/ml stock prepared in 100 mM HEPES, pH 7.0 to 7.2; see Support Protocol 1 for optimization of concentration) 96-well flat-bottom tissue culture plates Inverted microscope T-75 tissue culture flasks Additional reagents and equipment for growing mammalian cells in culture, trypsinizing cells, and counting viable cells by trypan blue exclusion (APPENDIX 3C) 1. At 24 to 48 hr post-transfection, harvest cells by trypsinization and count viable cells by trypan blue exclusion (APPENDIX 3C). 2. Plate 100 to 1000 cells in each well of a 96-well tissue culture plate using 200 µl selective medium per well. Begin incubation. 3. Replace selective medium once every 3 to 4 days and observe cells using an inverted microscope to monitor cell death and detachment. Large colonies of live cells (containing 100 to 200 cells) should appear within 3 weeks.
4. As the colonies grow and become confluent (indicated by change in color of the phenol red indicator in the medium from dark red to orange as the pH decreases), screen individual clones for the protein of interest using different methods depending on the targeting of the gene product. The supernatant is assayed for secreted proteins using an ELISA. For cytoplasmic proteins, the cells are screened through the detection of surrogate markers such as green fluorescent protein (GFP; Kahana and Silver, 1996). Membrane proteins may be amenable to immunofluorescent staining followed by flow cytometric sorting. An alternative approach is to pick 10 to 20 clones and screen cell lysates with ELISAs or immunoblots (UNIT 10.8). If the protein is localized to the membrane protein, immunofluorescent staining with specific antibodies (UNIT 10.10) can be directly applied.
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5. Trypsinize viable productive colonies and transfer to wells of a 24-well plate, each containing 2 ml selective medium. After cells reach confluency, trypsinize and transfer to a T-75 flask with 20-ml medium (APPENDIX 3C). When the attachment culture in a tissue culture flask grows to 80% to 90% confluency, it should be trypsinized and subcultured into a new flask with a starting cell density of 1 × 104/cm2 for continuation of cell growth. SUPPORT PROTOCOL 1
DETERMINATION OF BACKGROUND SENSITIVITY TO G418 The drug concentration and the plating cell density are very important for selection to be effective. In the case of selection, the optimal drug concentration is determined as described below. This concentration is set to the minimum required to kill nontransfected cells in 2 to 3 weeks. The selection becomes less stringent when cells are confluent, especially with G418 selection, probably because of lowered metabolic activity. It is important to choose a concentration of the toxic drug that will kill cells that are not expressing the resistance gene but that will allow growth. Additional Materials (also see Basic Protocol 3) Selective media containing various concentrations (100 to 1000 µg/ml in 100-µg/ml increments) of G418 (add from 50 mg/ml stock prepared in 100 mM HEPES, pH 7.0 to 7.2) 100-mm tissue culture dishes 1. Plate cells at a density of 0.6–1.2 × 106 per 100-mm dish in selective media containing G418 at various concentrations. Begin incubation. 2. Replace medium every 3 to 4 days and observe cells under inverted microscope. 3. Determine the minimum concentration of G418 required for complete killing of cells after 2 weeks.
ALTERNATE PROTOCOL 2
SCREENING OF GREEN FLUORESCENT PROTEIN BY FLOW CYTOMETRY An alternative to selection (see Basic Protocol 3) is the process of screening, where cells expressing the protein of interest are detected in the population using a noninvasive method and then isolated using flow cytometry or other techniques such as affinity chromatography and magnetic-affinity cell sorting. These methods have not been developed extensively because few noninvasive detection methods have become available. In the protocol below, the green fluorescent protein (GFP; Kahana and Silver, 1996) has been used as a screening marker and provides a flow cytometric method for measuring single-cell fluorescence, which may be combined with sorting based on the signal (Subramanian and Srienc, 1996). Any membrane protein against which specific antibodies can be directed may be used instead of GFP. An additional staining step is required when membrane proteins are used.
Production of Recombinant Proteins in Mammalian Cells
Materials Cells transformed with plasmid (see Basic Protocol 2 or Alternate Protocol 1) containing GFP reporter gene (pIRES1-EGFP from Clontech or pTracer-CMV from Invitrogen) at 50% to 70% confluency in T-25 or T-75 flasks 1% (w/v) bovine serum albumin (BSA; Sigma) in PBS (APPENDIX 2E) 10× (0.05 mg/ml) propidium iodide stock (PI; Molecular Probes; optional) Appropriate culture medium
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Flow cytometer with cell sorting capability (FACS Vantage and FACSort from Becton Dickinson Immunocytometry Systems or EPICS Elite ESP cell sorter from Coulter Electronics or equivalent) T-25 tissue culture flasks Additional reagents and equipment for trypsinizing cells, counting viable cells by trypan blue exclusion, and growing mammalian cells in culture (APPENDIX 3C) and flow cytometry (Robinson et al., 1998) 1. At 24 to 48 hr post-transfection, harvest cells by trypsinization and count viable cells by trypan blue exclusion (APPENDIX 3C). Make sure cells are in single-cell suspension and not in clumps.
2. Resuspend cells at a concentration of ∼1 × 106 cells/ml in 1% BSA/PBS. If desired, immediately before analysis, counterstain by adding 0.05 mg/ml (10×) PI to a final concentration of 5 µg/ml. 3. Run cells through flow cytometer at a flow rate of 200 to 1000 cells/sec. Tune laser wavelength to 488 nm and collect the following parameters: Forward-angle light scatter (FALS; linear) Right-angle light scatter (RALS; linear) Green fluorescence: linear and logarithmic using 510- to 540-nm band-pass (FITC) filter Red fluorescence (if counterstained with PI): linear or logarithmic using 670 or 690 long-pass filter. Gate on viable cells from bivariate plot of FALS versus RALS or FALS versus PI. Set gate to distinguish nonfluorescent from fluorescent cells, then select top 1% to 10% (or all) of the fluorescent cells for sorting (see Robinson et al., 1998, for detailed protocols in flow cytometry and cell sorting). 4. Sort 1 × 103 to 1 × 105 viable cells within the defined fluorescence threshold into a tube containing culture medium. Inoculate cells into T-25 flask with 5 to 10 ml culture medium. Begin incubation. 5. Sort cells again after 2 to 3 weeks, directly into 96-well plates containing 200 µl medium per well, at 1 cell/well. Continue incubation. Allow enough time for cells to establish a stable phenotype. Stringent criteria on GFP may be used at this step—i.e., selecting the top 1% or less of fluorescent cells (0.01% to 0.1% of cells can be sorted). After the clones grow out they can be screened for the production of the protein of interest.
SINGLE-CELL CLONING After cells expressing the gene product have been selected, single-cell cloning is usually applied to obtained monoclonal cultures. This is usually done by diluting a cell suspension and plating the cells in 96-well plates so that only single cells result in colonies, which may then be isolated. The cell suspension should consist mainly of single cells, with no clumps, and the viable cell count should be accurate. Cell clumps after trypsinization should be carefully broken up by repeated pipetting. Since hemacytometer-based counts have a 20% standard deviation (even with two hemacytometers) it is recommended that cells be counted using a Coulter Counter or an Elzone particle counter and that the count for viability (determined by trypan blue exclusion and a hemacytometer; APPENDIX 3C be corrected on the basis of the automated cell count to determine the viable cell density).
SUPPORT PROTOCOL 2
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Materials Cells expressing gene product of interest (see Basic Protocol 3 or Alternate Protocol 1) growing in tissue culture flasks or dishes at 70% to 90% confluency Appropriate culture medium Coulter Counter or equivalent 96-well tissue culture plates Additional reagents and equipment for growing mammalian cells in culture, trypsinizing cells, and counting viable cells by trypan blue exclusion (APPENDIX 3C) 1. Trypsinize cells (APPENDIX 3C). Break up any cell clumps by pipetting up and down repeatedly, then count cells using a Coulter Counter and determine number of viable cells by trypan blue exclusion. 2a. Cloning method 1: Resuspend cells in fresh medium at a concentration such that there will be 0.3 to 1.0 cell per well of a 96-well plate, in 200 to 250 µl of medium per well. Pipet a 200- to 250-µl aliquot of this suspension into each well. 2b. Cloning method 2 (visual check): Resuspend in medium at a concentration of 1 cell per 5 to 10 µl medium. Place a 5- to 10-µl drop (a Pasteur pipet can be used for this purpose) in center of flat-bottom 96-well tissue culture plate. Observe ∼5 wells at time under an inverted microscope at 100× to 200× magnification, then add 200 to 250 µl of medium to wells with only a single cell. 3. Incubate, replacing medium once every 1 to 2 weeks. Large colonies with at least 2000 cells take ∼2 to 4 weeks to form. BASIC PROTOCOL 4
AMPLIFICATION OF DIHYDROFOLATE REDUCTASE (DHFR) WITH METHOTREXATE (MTX) The level of expression after selection is usually not sufficient when the goal is to create a cell line for the production of the protein on a large scale. Amplification to increase gene dosage is a method for creating high producers. Cells are allowed to adapt to growth in the presence of a drug that inhibits the activity of the selection marker. This adaptation usually results from an increase in gene copy number to overcome the inhibition. Stepwise increases in drug concentration and adaptation of the cells amplify the cell line. In this protocol, the selection marker used is dihydrofolate reductase and the drug that stoichiometrically binds and inhibits DHFR activity is methotrexate (MTX). The glutamine synthetase marker and its corresponding inhibiting drug, methionine sulfoximine, is another amplification system that can be used with CHO cells. Materials Culture of transformed cells at 50% to 70% confluency, in T-25 or T-75 flasks (24 to 48 hr post-transfection; see Basic Protocol 2 or Alternate Protocol 1) Nucleoside-free αMEM medium (see recipe but omit nucleosides) containing 10% FBS that has been dialyzed to remove nucleosides (HyClone) 10 mM methotrexate (MTX) stock solution (Sigma; store up to 1 year at −20°C; thaw just before use)
Production of Recombinant Proteins in Mammalian Cells
96-well flat-bottom tissue culture plates 24-well tissue culture plates T-25 tissue culture flasks 100-mm tissue culture dishes
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Additional reagents and equipment for growing mammalian cells in culture, trypsinizing cells, and counting viable cells by trypan blue exclusion (APPENDIX 3C) Perform selection 1. At 24 to 48 hr post-transfection, harvest cells by trypsinization and determine number of viable cells by trypan blue exclusion (APPENDIX 3C). 2. Plate 100 to 1000 cells in each well of a 96-well tissue culture plate using 200-µl ml nucleoside-free αMEM/10% dialyzed FBS per well. Begin incubation. 3. As colonies grow and become confluent, screen clones (see Basic Protocol 3, step 4). 4. Trypsinize viable productive colonies and transfer to wells of a 24-well plate, each containing 2 ml nucleoside-free αMEM/10% dialyzed FBS. After cells reach confluency, trypsinize and transfer half to inoculate T-25 flasks containing 5 to 10 ml nucleoside-free αMEM/10% dialyzed FBS. Combine the remaining cells from all clones into one single pool and use in step 5. The clones transferred to the T-25 flasks can undergo selection. Screening is recommended at this stage before starting the amplification process.
Perform first round of amplification 5. Plate pooled cells at a density of 0.6–1.2 × 106 cells/ml in nucleoside-free αMEM/10% dialysed FBS (selective medium) containing 0.02, 0.05, 0.1, and 0.2 µm MTX (added from 10 mM stock) in a series of 100-mm dishes. 6. Replace selective medium in each dish with selective medium containing the same MTX concentration every 3 to 4 days and observe cells using an inverted microscope to monitor cell death and detachment. Large colonies of live cells (containing 100 to 200 cells) should appear within 3 weeks.
7. Trypsinize cells and count number of viable cells by trypan blue exclusion (APPENDIX 3C). 8. Plate cells from each of the methotrexate concentrations in wells of the same 96-well tissue culture plate at 0.3 to 1.0 viable cells per well in 200 µl selective medium with same MTX concentration as used before. Incubate until confluent, then transfer to 24-well plates and screen clones for productivity (see Basic Protocol 3, step 4). Perform additional rounds of amplification 9. Plate remaining cells from step 7 at a density of 0.6–1.2 × 106 cells per 100-mm dish in selective medium containing an MTX concentration 2- to 5-fold higher than that used before (0.5, 1, 2, 5, and 10 µM MTX). 10. After cells are adapted to the next level of methotrexate carry on with steps 7 to 9.
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BASIC PROTOCOL 5
ADAPTATION OF SUSPENSION CELLS TO PRODUCTION MEDIUM THROUGH MULTIPLE PASSAGING Adaptation of transfected CHO cell lines to suspension growth is recommended to enable growth in spinner bottles or reactors. Suspension growth in low-serum medium may improve the initial specific purity of the product by providing the opportunity for higher cell density and product titer than can be obtained by surface-attachment growth, as well as by reducing serum proteins in the medium. Growth of cells in suspension, in medium containing adequate levels of nutrients, enables use of compact production systems in which high cell density can be achieved. Such compact systems are economical choices in laboratory environments. Materials Mid–log phase culture of stable-transfected cells in T-75 flask (or thawed 1-ml aliquot of cryopreserved cells; see Support Protocol 2), produced by DHFR selection/MTX amplification (see Basic Protocol 4) or other selection/amplification procedure Nucleoside-free αMEM medium (see recipe but omit nucleosides) containing 10% FBS that has been dialyzed to remove nucleosides (for DHFR selection/MTX amplification cells) or regular αMEM medium (see recipe) containing 10% FBS (prepare as appropriate for cells produced by other selection/amplification procedures) 10 mM methotrexate (MTX) stock solution (Sigma; store up to 1 year at −20°C; thaw just before use) Dispase solution (Boehringer Mannheim) DMEM/F12 custom-modified medium (see recipe) with 10%, 1%, and 0.5% dialyzed FBS CD CHO medium (for DHFR systems; Life Technologies), Super CHO (for non-DHFR systems; Bio-Whittaker), or CHO-S-SFM (for non-DHFR systems; Life Technologies)—optional, for adaptation to serum-free medium 1 g/liter Nucellin-Zn salt (recombinant insulin; Eli Lilly)—optional, for adaptation to serum-free medium T-175 flasks Sterile 45-ml polycarbonate screw-cap tubes Inverted phase-contrast microscope 250-ml shaker flasks Platform shaker (New Brunswick Scientific) Additional reagents and equipment for growing mammalian cells in culture, trypsinizing cells, and counting viable cells by trypan blue exclusion (APPENDIX 3C), determining specific growth rate of cells (see Support Protocols 4, 5, and 9), determining specific productivity of cells producing heterologous protein (see Support Protocols 6 and 7), and freezing cells (see Support Protocol 2) Establish cultures in adaptive medium 1. Preincubate a T-175 flask and add 50 ml of prewarmed fresh αMEM/10% FBS (nucleoside-free and containing MTX concentration used for amplification if produced via Basic Protocol 4; prepare as appropriate for other selection/amplification schemes). Keep flask in incubator until the inoculum is prepared.
Production of Recombinant Proteins in Mammalian Cells
2. Remove the medium from a mid–log phase (50% to 90% confluent) T-75 flask of stable-transfected cells using a sterile 25-ml pipet. Trypsinize and harvest the cells into a sterile 45-ml polycarbonate tube (APPENDIX 3C).
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When the attachment culture in a tissue culture flask grows to 80% to 90% confluency, it should be trypsinized and subcultured into a new flask with a starting cell density of 1 × 104/cm2 for continuation of cell growth. If the culture is transferred from a tissue culture flask into a shaker flask to begin suspension adaptation, the starting cell density for the new culture should be ∼1 × 105 viable cells/ml. The procedures for trypsinizing and passaging cells are described in APPENDIX 3C.
3. Determine the concentration of viable cells by trypan blue exclusion (APPENDIX 3C), or, if clumping is evident, by adding 0.3 to 2.4 U/ml dispase, incubating 15 min at 37°C, then counting the number of viable cells by trypan blue exclusion. From the viable cell concentration determine the volume of suspension to inoculate into a T-175 flask to obtain a concentration of 1 × 104 viable cells/cm2. Add the required volume of inoculum to the T-175 flask and incubate. CHO cells have a tendency to clump when grown at high cell density in the higher-serum medium. Clumping may be minimized by adapting cells to growth in lower serum (≤0.5% FBS). Reduced-calcium media such as DMEM/F12 also may produce less cell clumping. In cultures that contain clumping cells the only secure counting method is through use of dispase before hemacytometer/trypan blue counting.
4. Periodically observe cells in the tissue culture flask using an inverted phase-contrast microscope at 125× magnification. When the cells reach a confluency of 50%, remove the spent medium and replace with the new medium, DMEM/F12 supplemented with 10% dialyzed FBS and the concentration of MTX (or other selective agent) used previously. Continue incubation, periodically observing growth using the inverted microscope. 5. When cells reach 80% to 95% confluency, trypsinize and harvest the cells. Determine the viable cell density (APPENDIX 3C) and calculate the volume of inoculum required to give a final concentration of 1 × 105 viable cells/ml in 100 ml. Simultaneously, prepare a 250-ml shaker flask with 100 ml of fresh DMEM/F12/10% dialyzed FBS/MTX and preincubate to 37°C. 6. Remove the volume of medium calculated for the inoculum from the shaker flask and transfer the corresponding volume of inoculum. Mix by gentle swirling, then remove a 0.25-ml sample and determine post-inoculation viable cell density (APPENDIX 3C). Loosely cap the flask, and begin incubation with agitation at 75 to 125 rpm on a platform shaker. 7. Continue incubation and calculate specific growth rate from the growth curve (see Support Protocols 4, 5, and 9) and specific productivity (see Support Protocols 6 and 7). Determine the viable cell density (APPENDIX 3C) daily. At 4 days post inoculation, or when the cell density has reached 4 × 105 viable cells/ml, transfer sufficient cells to inoculate a second passage flask as in step 6. 8. Passage the cells in changes of the same medium until the passage-specific growth rate reaches a constant maximum value. At this passage, reduce the serum concentration in the medium to 1% dialyzed FBS. Also ascertain that the specific productivity remains within reasonable bounds with increased passage number, as an indication of expression stability.
9. When the passage-specific growth rate shows no further increase, reduce the serum to 0.5% and repeat adaptation passaging. When growth rate shows no further gain, the cells have adapted to the low level of serum.
Confirm adaptation to production medium 10. Confirm that cells have adapted to the low level of serum by splitting the cells into two flasks: one with medium plus 10% FBS and another with medium plus 0.5% FBS. Determine growth rate, which show no large difference for correctly adapted cells.
Production of Recombinant Proteins
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11. If serum-free growth is desired: Perform a further adaptation from 0.5% to 0% serum by repeating the adaptation steps through a final round using specific-protein-free medium such as CD CHO, Super CHO or CHO-S-SFM (or equivalent). Supplement with 5 mg/liter of Nucellin (added from 1 g/liter stock) and reduce to 1 mg/liter gradually 12. Freeze adapted cells (see Support Protocol 2) or proceed to growth in batch mode (see Basic Protocol 6). BASIC PROTOCOL 6
GROWTH OF CELLS IN BATCH MODE IN LARGE-SCALE SPINNER CULTURE VESSELS Suspension-adapted cells may be grown in spinner flasks accommodating liquid volumes of up to 16 liters. Flasks are placed in a humidified temperature-controlled environment, which contains 5% CO2 gas. The CO2 is used to provide dissolved carbon dioxide and bicarbonate, which provide nutrient substrate and buffer components in the culture medium. The flasks are set on a rotating platform to agitate the liquid surface and provide adequate oxygen transfer to support cell oxygen demand. The procedure may be initiated from a vial of the cells or from ongoing shaker flasks that are being passaged (see Basic Protocol 5). The ultimate scale of production depends on the expression level, the purification yield, and the quantity of protein required by the researcher. Flasks can accommodate up to 100 ml of culture while small glass spinner jars can be used to produce up to 3 liters of culture. When larger volumes of liquid culture are required, the spinner flasks may become more complicated in design, including dissolved oxygen (dO2) controllers and pH controllers. Volumes greater than 5 liters of culture can be obtained by using 20-liter spinner jars. The increased volume and subsequent apparatus size becomes cumbersome and is worthy of arranging in a fixed location. In order to supply adequate oxygen to support the cultures, it is necessary to set up such a spinner jar in a warm room or with an external wraparound band heater (Watlow). The spinner jar is usually fitted with a top-entry mechanical stirrer (Bellco Biotechnology). The supply of oxygen to the culture is achieved through sparging with O2/air mixtures through a sparge tube fitted with a frit (Mott Cups) and by adding a CO2/O2/air mixture into the headspace. Similarly the control of pH is best achieved through use of an in-vessel pH probe with adjustment by addition of 0.5 M NaOH using automatic pH controllers or manually performing daily pH checks. A typical large volume (20-liter) spinner set up is shown in Figure 5.10.2. Here continued use of MTX is desired, use nucleoside-free medium and dialyzed FBS. In most cases where stable integration and production has been acceptable at levels <1 µM MTX, the continued use of methotrexate should not be required. Materials 1-ml cryovial or mid–log phase culture of suspension-adapted cells (see Basic Protocol 5) Medium to which cells have been adapted (see Basic Protocol 5), supplemented with 4 mM L-glutamine and 4.5 to 5.5 g/liter D-glucose Medical-grade, sterile-filtered 5% CO2/air mixture Medical-grade, sterile-filtered 100% O2 (if necessary)
Production of Recombinant Proteins in Mammalian Cells
250-ml screw-cap shaker flasks Platform shaker (New Brunswick Scientific) 1-, 5-, and 20-liter borosilicate glass spinner jars (Bellco Biotechnology) Flat-plate stirrer platform (Bellco Biotechnology)
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Peristaltic pump (head size 15, 16, 24; with 0 to 600 rpm capability; Watson-Marlow or Cole-Parmer) Sterile silicone tubing (Pt-cured; size 15, 16, 24; Masterflex from Cole-Parmer) Sterile 50-ml syringes Dissolved oxygen probe (Ingold) pH probe (Ingold or Broadley James) Additional reagents and equipment for growing mammalian cells in culture and counting viable cells by trypan blue exclusion (APPENDIX 3C), determining specific growth rate of cells (see Support Protocols 4, 5, and 9), and determining specific productivity of cells producing heterologous protein (see Support Protocols 6 and 7) Establish cultures 1. Thaw a 1-ml cryovial of the culture in a 37 ± 1°C water bath for ∼2 min with gentle shaking. 2. To a 250-ml screw-cap shaker flask, add a volume of cells from the vial sufficient to inoculate 100 ml of the medium to which the cells have been adapted (supplemented with L-glutamine and D-glucose and prewarmed to 37°C), at 1 × 105 viable cells/ml.
motor filter (0.45 µm)
filter air gas out
top air inoculum line
sparge sample
oxygen line clamps
line clamps attach sampling device
maximum liquid level
filter
base inoculation vessel (5 liters) stirrer
harvest/inoculum line
frit sparger
Figure 5.10.2 Typical arrangement of 20-liter spinner jar for batch cell culture.
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Incubate flask in a humidified 37°C ± 0.5°C, 5% CO2 incubator with rotation at 100 rpm on a platform shaker. Measure the viable cell density daily (APPENDIX 3C). When the culture reaches mid–log phase (∼3 to 4 days) the cells are suitable for inoculation of a 1-liter spinner jar containing 400 ml medium.
Inoculate spinner jar cultures 3. Add 400 ml of 37°C medium to a preincubated (5% CO2, 37°C) 1-liter glass spinner jar. Add 100 ml of the mid–log phase shaker flask culture through the side arm. Determine the post-inoculation viable cell density, then place the spinner jar in the incubator on a flat-plate stirrer platform and set the stirrer speed to 60 to 80 rpm. Incubate, measuring the viable cell density daily. When the culture reaches mid–log phase (∼3 to 4 days) the cells are suitable for inoculation of a 5-liter spinner jar containing 1600 ml of medium. The culture may be continued in the 400-ml spinner culture by adding glucose to a final concentration of 2 g/liter and glutamine to a final concentration of 2 mM on days 4, 6, and 8, along with daily pH monitoring and adjustment to 7.0 to 7.4 using 0.5 N NaOH.
4. Add 1600 ml of 37°C medium to a preincubated 5-liter glass spinner jar. Add the 400 ml of the mid–log phase culture. Determine the post-inoculation viable cell density and incubate with stirring as in step 3. Measure the viable cell density daily and monitor growth rate (see Support Protocols 4, 5, and 9) and specific productivity (see Support Protocols 6 and 7). When the culture reaches mid–log phase (∼3 to 4 days) the cells are suitable for inoculation of a 20-liter spinner jar containing 4500 ml of medium. Otherwise the culture may be continued in the 5-liter jar by adding nutrients as described above. Knowledge of the growth curve will enable the researcher to choose the point at which the cells may be harvested with adequate viability and cell density. Where still larger volumes up to 16 liters are required, continue with step 5 onwards.
Increase production 5. Preincubate a 20-liter spinner jar at 37°C overnight in a warm room and flush the jar with filtered sterile 5% CO2/air mixture through the sparger. The 20-liter spinner jar is fitted with a sterilizable dissolved oxygen probe and pH probe for measurement of dO2 and pH.
6. Add 4.5 liter fresh prewarmed medium to the empty preincubated jar. 7. Transfer 1.5 liter of mid–log phase inoculum to the 20-liter spinner using a peristaltic pump with size 16 silicone tubing or by gravity feeding using a size 16 silicone tube between the inoculum bottle and the inoculum port on the spinner jar. Determine post-inoculation viable cell density. After inoculation, if the cell density is in the range 0.5–1.5 × 105 viable cells/ml, surface aeration is sufficient to supply the oxygen to the cells.
8. Set the flow rate of air/5% CO2 mixture in the headspace to 100 ml/min. 9. Sample the culture daily through a side-arm port using sterile tubing and presterilized 50-ml syringes.
Production of Recombinant Proteins in Mammalian Cells
10. As the culture grows, and the dissolved oxygen begins to fall to 50% air saturation (measured by dO2 probe), set the flow rate of air/5% CO2 mixture in the sparger to ∼25 ml/min. Increase this rate, as oxygen demand increases, up to 50 ml/min. If the demand for oxygen is not met (i.e., if the dO2 remains <50% of saturation) using the air/5% CO2 mixture, add 100% oxygen to the mixture. Maintain the dissolved oxygen
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between 20% and 100% air saturation. Maintain pH in the range 7.2 to 7.4 using automatic pH control or daily manual addition of 0.5 N NaOH. 11. When the culture reaches mid–log phase (around day 4) add 6 liters of fresh prewarmed (37°C) medium using a peristaltic pump and size 16 silicone tubing (see step 7). 12. When the 12-liter culture reaches mid–log phase (around day 6) add 4 liters of fresh prewarmed (37°C) medium. The 16-liter culture may be grown up to the desired point for harvesting or may be operated in a feed-and-bleed mode. In feed-and-bleed mode, typically 4 to 10 liters of medium can be harvested from the late log-phase culture and replaced with the equivalent volume of fresh medium.
13. Harvest the protein product (see Basic Protocols 8 and 9). GROWTH OF CELLS IN LARGE-SCALE BATCH REACTORS Stirred-tank bioreactors may range in scale from 10 liters to 200 liters in a laboratory/pilot plant location. Bioreactors are distinguished from glass jar spinners and similar devices by their robust stainless-steel design and automatic control systems for temperature, dissolved oxygen, and pH, as well as instrumentation for monitoring important parameters such as medium flow rate, gas flow rate, pressure, weight, and liquid level. Bioreactors should possess ports around the base of the vessel for insertion of pH and dissolved oxygen probes. Since bioreactors may contain large volumes of culture and may operate at high cell density, the total oxygen demand is larger than can be supplied by agitation alone. In such reactors the addition of oxygen is achieved through sparging of the culture. This involves adding the gas through a frit-type device at the base of the reactor, or through a drilled-hole sparger. The authors recommend the frit design for reactors up to 500 liters. Bioreactor systems are supplied by specialized manufacturers of process equipment and may require lead time of 3 to 6 months from order to delivery.
BASIC PROTOCOL 7
Materials Inoculum of mid–log phase cells in spinner jar (see Basic Protocol 6) Ultrafiltered deionized water Medium to which cells have been adapted (see Basic Protocol 5), supplemented with 8 mM L-glutamine and 4.5 to 5.5 g/liter D-glucose Medical-grade, sterile-filtered 95% N2/5% CO2 and 95% air/5% CO2 mixtures and 100% oxygen 10- to 100-liter bioreactor (New Brunswick Scientific, Bioengineering, Braun, New MBR, Applikon) pH probe (Broadley James or Ingold), calibrated Dissolved oxygen probe (Ingold), pretested Autoclave large enough to accommodate bioreactor vessel (Finn Aqua) Inoculum transfer bottle with bottom side port attached to sterile size 16 tubing (Bellco Biotechnology) Sterile flexible disposable medium and harvest bags (Stedim Laboratories) Additional reagents and equipment for counting viable cells by trypan blue exclusion (APPENDIX 3C), determining specific growth rate of cells (see Support Protocols 4, 5 and 9) and determining specific productivity of cells producing heterologous protein (see Support Protocols 6 and 7) Production of Recombinant Proteins
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Prepare the bioreactor assembly 1. Assemble bioreactor components—e.g., agitator, sparger lines, outlet gas line, top headspace gas inlet, base addition lines, medium addition lines, inoculum transfer lines, harvest lines, and sample port lines. Recommended equipment includes sterile tubing (Pharmed) and connection tubing welder SCD IIB (Terumo Medical).
2. Prepare an inoculum for inoculation of the bioreactor. This would be between 0.1 and 0.15 liter of mid–log phase culture per liter of bioreactor operating volume. For a 100-liter bioreactor, an inoculum volume of 10 to 15 liters is required and would be produced by the 20-liter spinner method (see Basic Protocol 6).
3. Calibrate the pH probe and insert into reactor through the side port. Insert the pretested dissolved oxygen probe. Fill the vessel with ultrafiltered deionized (UFDI) water to cover the probes if sterilization is not immediately scheduled. Prior to sterilization, remove water. 4. Add ∼10 ml ultrafiltered deionized water per liter of vessel volume and sterilize the vessel in an autoclave for 1 hr at 121°C, using slow exhaust for cooling phase. 5. Pump the culture medium into the vessel to cover the probes. Equilibrate system by sparging the medium and headspace at ∼0.005 vessel liquid volumes per minute with a 95% N2/5% CO2 mixture, and allow the medium to reach the 37°C operation temperature. After equilibration (∼5 hr required) is complete (as judged by an x-axis asymptotic dissolved oxygen trace), set the dissolved oxygen probe to 0% air saturation, then sparge the vessel with 95% air/5% CO2 mixture. After equilibration, set the dissolved oxygen probe span to 95% air saturation. Optional recommended equipment includes a YSI 2700 Select glucose, glutamine, and lactate analyzer (YSI Instruments) and an IBI Biolyzer for ammonia and lactate (Kodak Lab Research Products).
Inoculate the bioreactor 6. Adjust the medium volume to 40% of the target final harvest volume (e.g., 4 liters in a 10-liter-operating-volume vessel). Equilibrate to 37°C, then inoculate to a starting density of 1 to 2 × 105 viable cells/ml using the inoculum transfer bottle. 7. Set the sparge air-flow rate to ∼0.005 vessel volumes of air per vessel liquid volume and set the dissolved oxygen controller to add pure (100%) oxygen to the sparger on demand. Purge the headspace 0.0025 vessel liquid volumes of air. Agitate the culture at an rpm producing a tip speed of 1000 to 1500 cm/sec. 8. Continue incubation, determining the post-inoculation viable cell density daily (APPENDIX 3C). Monitor growth rate of cells 9. Feed medium beginning on day 2 and adjust the rate daily to give a rate of 2 liters per day per 1 × 109 total viable cells until the target volume is reached. At that point, stop feeding fresh medium. 10. During the initial-feed batch and final batch phases maintain the pH in the range 7.2 ± 0.1, the temperature at 37°C ± 0.5°C, and the dissolved oxygen in the range of 20% to 100% air saturation. Production of Recombinant Proteins in Mammalian Cells
The feed rates can be set through use of precalibrated pumps and checked using load cells that measure the medium supply container weights.
11. Harvest the protein product (see Basic Protocols 8 and 9) using a sterile flexible bag.
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GROWTH OF CELLS IN CONTINUOUS CULTURE IN BIOREACTORS Continuous culture refers to the operation of a bioreactor where the cells are removed at a particular rate and replaced with fresh medium at an equal rate. The ratio of the medium-addition rate to the culture volume is known as the dilution rate. Continuous culture may be desirable where large quantities of protein product are required for further studies, and is also of interest to the researcher for examining the potential effects of growth rate on the product. In this situation, a continuous reactor is set up with no cell retention devices. Where the researcher wishes to maximize the productivity of the given system, a continuous perfusion (cell retention) system would be used. Medium is removed at a rate corresponding to the addition rate of fresh medium but a large proportion of cells are retained in the reactor through use of suitable retention devices. The retention devices are usually either internal spin filters (supplied with bioreactors), external tangential cross-flow microporous filtration devices (Millipore, A/G Tech, Sartorius, Pall-Filtron) with cell recycle, external centrifugal devices (Centritech from Sorvall) with cell retention, or inclined settling surfaces (custom designed), which may be internal or external. Recently acoustic settlers have become available (Sonosep) and these are possible candidates for bioreactor scales around 10 to 30 liters.
ALTERNATE PROTOCOL 3
The reactor assembly would incorporate the required peripherals for continuous operation (Fig. 5.10.3). In the case of perfusion, the settling device would be incorporated and sterilized along with the bioreactor. The basic setup is the same as described previously (see Basic Protocol 7).
peristaltic pump
cells
conditioned medium out peristaltic pump
fresh medium supply cell bleed
0.5 N NaOH peristaltic pump
conditioned medium
solenoid
95% air/ 5% CO2
sparger
sample line
100% O2
pH base addition
95% air/ 5% CO2
motor s
O2 suppl y fresh medium
process controller
temp temperature dissolved oxygen heating belt
spin filter heating blanket pH
liquid volume
vessel weight (load cell)
Figure 5.10.3 Compact-scale perfusion cell culture system. Shows typical connection of medium and product tanks to compact (100-liter) reactor. Simple pH, dissolved oxygen, and vessel weight control loops are illustrated. Medium and product vessels (for conditioned medium and/or cells) should be set up in a cold-box environment.
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Additional Materials (also see Basic Protocol 7) Bioreactor with accessories for continuous operation (consult New Brunswick Scientific, New MBR, Bioengineering AG, Braun, or other fermenter manufacturer; for perfusion operation where retention is required, use reactors with spin filter design, e.g., New MBR or Braun) Sterile bags (Stedim Laboratories) or sterilized stainless steel containers for collection of medium 1. Set up and inoculate bioreactor (see Basic Protocol 7, steps 1 to 7). For the simple continuous mode and perfusion operation, feed medium at a rate of 2 liters per day per 1 × 109 total viable cells, up to the total operating volume of the reactor. 2. When the cells have reached mid to late log phase, begin cell removal at the target rate corresponding to the desired dilution rate (the volume of medium added to the vessel per day per reactor operating volume) The reactor operating volume is constant with respect to time. Fresh medium is fed to the reactor to maintain constant volume. In the case of the perfusion reactor, the retention device results in cells being returned to the bioreactor medium, and the cell density increases until it becomes limited by a nutrient or by toxic waste products such as ammonia and lactate. At that point a steady-state condition will be reached and cell density will remain constant.
3. Collect the conditioned medium as it removed from the bioreactor in sterile bags or sterilized stainless steel containers. Cool the stream to 4° to 8°C to minimize product degradation. 4. Maintain the pH in the range 7.2 ± 0.1, the temperature at 37°C ± 0.5°C, and the dissolved oxygen in the range of 20% to 100% air saturation. The feed rates can be set through use of precalibrated pumps and checked using load cells.
5. Harvest the protein product (see Basic Protocol 8 and 9). HARVESTING OF PROTEIN PRODUCT FROM BATCH MODE SPINNER CULTURES AND LARGE-SCALE BATCH REACTORS Most often, protein production in mammalian cells leads to secretion of the product across the cell wall into the culture medium where it accumulates with time in a batch mode (see Basic Protocol 8). In some cases the researcher may wish to harvest the cells in order to obtain an intracellular or membrane-associated protein (see Basic Protocol 9). In general, all strategies will necessitate a solid/liquid separation step and most frequently will employ batch centrifugation on a small scale or tangential cross-flow filtration (TFF) at larger scales, e.g., ≥20 liters. For continuous-mode bioreactors, harvesting typically involves accumulation of conditioned medium, which is processed using depth filters and ultrfiltration concentration. Cell-associated product must be obtained by disrupting the cell wall and again separating the solid and liquid phase. Nonionic detergents may be used to weaken the membrane and aid the release of non–integral-membrane–associated material. Where the product is an integral membrane protein, the disrupted cell membrane should be recovered and a typical membrane protein extraction and purification protocol pursued (UNIT 4.2). Production of Recombinant Proteins in Mammalian Cells
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Harvesting a Secreted Product
BASIC PROTOCOL 8
Materials Cells in batch culture (see Basic Protocols 6 or 7 or Alternate Protocol 3) Phosphate-buffered saline (PBS; APPENDIX 2E) TE buffer or PBS (APPENDIX 2E) Sterile bags (Stedim Laboratories) Refrigerated centrifuge: Sorvall RC-5B or RC-3B, Beckman J2-21 or S600, or equivalent Microfiltration system with membranes for cell concentration and debris/product separation (Sartorius, Pall-Filtron, or Millipore) Depth filtration membranes for large-scale debris separation prior to sterile filtration (Cuno Bio-Cap 90 SP or equivalent products from Millipore or Sartorius) Filter-sterilizing capsules (0.2-µm membrane; Sartorius, Pall, or Millipore) Silicone tubing, autoclaved Sterile polycarbonate containers Ultrafiltration system with membranes for protein concentration (MWCO 10-kDa; Sartorius, Pall-Filtron, or Millipore) 1. Following completion of the batch process in spinner jar or a bioreactor remove medium from the reactor harvest port using sterile silicone tubing and a peristaltic pump. Collect into a sterile container such as a presterilized glass jar or sterile disposable plastic flexible bag. Chill collected material to 4° to 10°C 2a. For volumes ≤6 liters: Centrifuge 10 min at 1000 to 4000 × g, 4° to 10°C, in a Sorvall RC-3B centrifuge with standard rotors. For up to 20-liter volumes it may be feasible to use a larger-capacity laboratory centrifuge such as the Sorvall RC-5B fitted with the swinging bucket rotor accommodating six 1-liter buckets; using this setup the cells should be centrifuged 15 min at 4000 × g, 4° to 10°C.
2b. For volumes >20 liters: Use a wide-channel nonscreened microporous cross-filtration unit fitted with ∼1 ft2 (0.2 µm or 0.45 µm pore rating) per 2-liter culture. Recirculate the cells through the retentate loop to give an inlet gauge pressure ≤10 psi. Limit the transmembrane gauge pressure to <4 psi using a peristaltic pump on the permeate line. Concentrate the cells in the retained filter stream by reducing the volume 10-fold, then permeate a further 3 vol PBS to wash the cells and recover the remaining product. Collect and chill the permeate in sterile plastic bags or sterile containers. 3. Filter the centrifuge supernatant, perfusion conditioned medium, or permeates using a rinsed, prewet liquid depth filter followed by a prewet or rinsed 0.2-µm rated filter-sterilizing capsule. Use autoclaved silicone tubing on the sterile side of the filter capsule and recover product in sterile polycarbonate containers or sterile plastic bags. Rinse the capsules with a minimum amount (50 to 100 ml) of TE buffer or PBS to recover the product. IMPORTANT NOTE: It is very important to prewet or rinse capsule filters as they hold significant amounts of liquid, which can result in a loss of product.
4. Optional: Further concentrate filtrate by tangential ultrafiltration using a 10-KDa MWCO ultrafiltration membrane sized to 1 ft2 of screened membrane per 20 liters. Recirculate at an inlet gauge pressure of 20 to 30 psi and a transmembrane gauge pressure of 10 to 15 psi. A minimum recirculation loop volume of 100 ml is feasible for small-scale systems (Millipore, Sartorius, or Filtron Slice units) and will define the maximum physical concentration factor.
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5. Rinse the filtration loop and filters with 50 to 100 ml of TE buffer or PBS to recover the product. The product may be purified directly or stored frozen (see Support Protocol 3) until required. BASIC PROTOCOL 9
Harvesting a Cell-Associated Product Materials Cells in batch culture (see Basic Protocols 6 or 7 or Alternate Protocol 3) Phosphate-buffered saline (PBS; APPENDIX 2E) Homogenization buffer (see recipe), 4°C Cell homogenizer: e.g., Microfluidizer (Microfluidics), Tissumizer (Tekmar-Dohrmann), Dounce homogenizer (Bellco Biotechnology), or Manton-Gaulin-APV homogenizer (APV-Gaulin) Additional reagents and equipment for centrifugation or cross-flow microfiltration of cell suspension (see Basic Protocol 8) 1. Subject cell suspension from batch culture to centrifugation or cross-flow microfiltration (see Basic Protocol 8, steps 2a or 2b), but retain centrifuge pellets or cross-flow filtration–concentrated slurry. Keep cells at 4° to 10°C. 2. Resuspend cells in 3 to 5 vol PBS (or other isotonic buffer) and centrifuge 10 min at 1000 to 4000 × g, 4° to 10°C. The cell pellet may be stored frozen (<−35°C) or processed directly.
3. Resuspend pellet in 2 to 3 vol of 4°C homogenization buffer. Lyse cells using a tissue homogenizer at 4°C. For larger volumes, it is possible to recirculate the suspension through a gear-type pump at high flow rate (obtain 10-20 psig pump outlet pressure in a 0.25-0.5 in.-diameter tube recirculation loop).
4a. For volumes <20 liters: Centrifuge the homogenate 30 min at 5000 × g, 4°C to remove the bulk of the debris. Filter sterilize the supernatant, which will contain the released soluble protein, prior to storage or further processing. 4b. For large volumes: Remove the debris using microfiltration/tangential-flow filtration (MF-TFF) at an inlet gauge pressure of 20 psi and limit the transmembrane gauge pressure to <4 psi using a peristaltic pump on the permeate line. Wash the retained concentrated debris with 3 to 5 vol buffer. Filter-sterilize the permeate, which will contain the released soluble protein, prior to storage or further processing. Cell debris containing integral membrane protein may be recovered as the retentate concentrated slurry. The permeate stream may be concentrated using cross-flow ultrafiltration, as described for secreted protein (see Basic Protocol 8).
Production of Recombinant Proteins in Mammalian Cells
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FREEZING CELLS FOR PRODUCTION OF CELL BANKS In order to be able to produce recombinant proteins consistently from batch to batch, it is advisable to produce a number of equivalent storage vials of the cell line—often referred to as a parent or master cell bank. Before attempting large-scale culture with a new cell bank the researcher is strongly advised to ensure that the cell bank is not contaminated by microbes, including mycoplasma, and that it grows normally (i.e., compared to the growth curve obtained during preparation). A growth curve test of the vial would detect contamination by bacteria, yeast, and fungi, while mycoplasma may be determined using commercial kits such as the Fluorescent DNA stain (Hoechst). A primary cell bank, or parent or master cell bank (MCB), can be used as the source to create subsequent working cell banks (MWCB) when necessary, depending on the scale of production. General principles and procedures for cryopreservation of attachment-dependent cells are provided in APPENDIX 3C. The following protocol is applicable to cells that have been adapted for suspension growth.
SUPPORT PROTOCOL 3
Materials High-viability (>95%) log-phase culture of cells Freezing medium: complete medium used for growing cells supplemented with 10% to 20% (v/v) FBS and 5% to 10% (v/v) DMSO (Sigma), 4°C 45-ml sterile screw-cap tubes polycarbonate centrifuge tubes (Falcon from Becton Dickinson) Sorvall T-6000B centrifuge (DuPont) or equivalent 1.5-ml Nalgene cryotubes (Nunc) Additional reagents and equipment for counting viable cells by trypan blue exclusion (APPENDIX 3C) 1. Take an aliquot of ∼1 ml from a high-viability log-phase culture and determine the viable cell density (APPENDIX 3C). Determine the number of vials to be produced, then, based on the viable cell density and the total number of frozen vials to be prepared for the cell bank, calculate the volume of suspension culture needed. Each vial should usually contain 0.5 or 1 × 107 viable cells with a density of 1 × 107 cells/ml.
2. Transfer the desired volume of cell suspension to one or more sterile 45-ml polycarbonate centrifuge tubes and centrifuge 10 min at 300 to 350 × g in a T-6000B or equivalent centrifuge, room temperature. 3. Decant supernatant from the centrifuge tube(s) and resuspend pellet in required volume of 4°C freezing solution for a final concentration of 1 × 107 viable cells/ml. 4. Transfer 0.6- or 1.1-ml aliquots of the cell suspension into labeled cryotubes. 5. Cool and freeze cells at a rate of −1°C/min. Three general methods can provide this freezing profile: (1) the vials can be placed in a styrofoam box with a wall thickness of ∼15 mm, which is then placed in a −70°C freezer; (2) the vials can be placed in a special cell-preservation container supplied by Nalgene which is filled with isopropanol and placed at −70°C; (3) an expensive programmable cooler can be used that can sense the temperature of the vial and feed in liquid N2 accordingly. Placing the vials directly into liquid N2 can result in the formation of ice crystals in the cells and poor viability when thawed.
6. When vials reach −70°C (or after 2 days) transfer to liquid N2 for long-term storage.
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SUPPORT PROTOCOL 4
NUMERICAL DETERMINATION OF SPECIFIC GROWTH RATE OF CELLS Specific growth rate of cells any given time is defined as the rate of total biomass production divided by viable cell density at that given time: µ(t ) =
d (V ⋅ X ) 1 V (t ) ⋅ X (t ) dt
where µ(t) is the specific growth rate at time t, V(t) is the volume at time t, and X(t) is the cell density at time t. Specific growth rate is a function of cell type as well as the depletion of growth-limiting nutrients in the medium of choice and/or the accumulation of growth-inhibitory waste products in the medium. Both graphical (see Support Protocol 5) and numerical methods have been used to determine the specific growth rate of a culture. The growth rate is determined numerically by the following procedure. 1. Take a 1-ml sample from the culture and count viable cells (APPENDIX 3C) at time point 1. 2. After 4 to 24 hr (depending on how fast cells are growing) take another 1-ml sample from the culture to count viable cell density at time point 2. 3. Calculate the specific growth rate µ of cells as: ln µ time 2 =
SUPPORT PROTOCOL 5
viable cell density at time 2 viable cell density at time 1 time 2 − time 1
GRAPHICAL DETERMINATION OF SPECIFIC GROWTH RATE OF CELLS When complete growth curve data are obtained, the data may be input to a spreadsheet/graphics-type program capable of curve fitting and defining a polynomial best fit. The information can be used to determine the derived parameter of specific growth rate. For definition of specific growth rate, see Support Protocol 4. The growth curve is determined graphically as follows. Materials Spreadsheet software with graphical output (e.g., Microsoft Excel or Lotus 123) 1. Take a 1-ml sample from the culture for viable cell count (APPENDIX 3C) every 4 to 24 hr, depending on how fast the cells are growing, until late log or stationary phase. 2. Plot the viable cell density versus time on an x-y chart using spreadsheet software capable of graphical output. 3. Curve-fit the chart to a best 3- or 4-order polynomial function. The logarithmic growth phase (days 0 to 5, typically) can also be fitted with a log function or linear fit on a log scale.
4. Calculate the specific cell growth rate at time = T hr as the first-order derivative of the function divided by the viable cell density at time = T hr. Production of Recombinant Proteins in Mammalian Cells
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NUMERICAL DETERMINATION OF SPECIFIC PRODUCTIVITY OF CELLS PRODUCING HETEROLOGOUS PROTEIN
SUPPORT PROTOCOL 6
The specific productivity of a protein at any given time is defined as the rate of increase in the protein mass divided by total viable cell mass at that given time: specific productivity (t ) =
dP(t ) 1 X (t ) dt
where X(t) is the cell concentration at time t and P(t) is the product concentration at time t. Specific productivity is calculated as follows. 1. Take a 1-ml sample from the culture, count viable cells, and determine the product protein concentration by ELISA at time point 1. 2. After 4 to 24 hr (depending on how fast cells are growing), take another 1-ml sample from the culture, count viable cells, and determine product protein concentration by ELISA at time point 2. 3. Calculate the specific productivity (qp) by cells as: qp ( time 1 → time 2) =
protein concentration at time 1 − protein concentration at time 2 ( time 2 − time 1) × Xavg ( time 1 → time 2 )
where Xavg ( time 1 → time 2) =
viable cell density time 2 − viable cell density time 1 ln ( viable cell density time 2 − viable cell density time 1 )
GRAPHICAL DETERMINATION OF SPECIFIC PRODUCTIVITY OF CELLS PRODUCING HETEROLOGOUS PROTEIN
SUPPORT PROTOCOL 7
For definition of specific productivity, see Support Protocol 6. Materials Spreadsheet software with graphical output (e.g., Microsoft Excel or Lotus 123) 1. Sample cultures at time points 1 and 2, count viable cells, and determine protein concentrations (see Support Protocol 6, steps 1 and 2). 2. Plot the viable cell densities versus time and plot the protein concentrations versus time on x-y formatted charts using spreadsheet software capable of graphical output. 3. Curve-fit the charts with the best 3- or 4-order polynomial functions. 4. Calculate the specific productivity as the first-order derivative of the protein concentration curve-fit function divided by the viable cell density at any given time point.
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SUPPORT PROTOCOL 8
PREPARATION OF SAMPLES FOR PRODUCT ASSAY Samples taken from the culture need special preparation for protein assay and storage. The handling procedures are different for cultures that make intracellular product and those that make secreted product. Freezing whole-cell cultures and subsequent thawing may affect the value and consistency of the assay in some cases. If samples must be stored frozen as whole-cell cultures prior to assay, the effects of freeze/thawing should be evaluated, but this is not recommended as a consistent method. In the procedures below, the culture is centrifuged or filtered to separate the cells (which are lysed to produce a cell extract if the product is intracellular) from the supernatant (which is handled separately when the product is secreted). Materials Sample of cell culture (see Basic Protocols 6 or 7 or Alternate Protocol 3) 0.9% NaCl, filter-sterilized, 4°C Refrigerated centrifuge Ultrasonic homogenizer (Misonix) 1. Centrifuge 1 to 5 ml of sample 10 min at 300 to 350 × g, 4°C. Decant supernatant; retain for assay or store frozen at ≤30°C for future analysis if protein product is secreted. If protein product is intracellular, proceed with step 2 and subsequent steps. 2. Resuspend the cell pellet in 1 ml of cold 0.9% NaCl solution. 3. Centrifuge again for 10 min at 300 to 350 × g, 4°C. 4. Decant supernatant and resuspend pellet in 1 ml of 4°C 0.9% NaCl. 5. Keeping the sample tube in an ice bucket, place the sample tube under the ultrasonic homogenizer and immerse the microtip probe in the cell suspension. Operate the sonicator for a 10-sec cycle with continuous power-on time of 0.5 sec and continuous power-off time of 1.5 sec, with the power set at low to medium level. After the completion of a 10-sec cycle, load a drop of cell suspension onto a hemacytometer slide and observe the lysate under 125× magnification using phase contrast. If insufficient disruption is observed, perform further 10-sec sonication cycles until cells lysed.
6. Centrifuge the suspension of disrupted cells 10 min at 800 to 1000 × g, 4°C. 7. Retain supernatant for assay or store frozen at ≤30°C for future analysis. SUPPORT PROTOCOL 9
DETERMINATION OF SHAKER FLASK GROWTH CURVE A complete 10-day growth curve is useful for determining the expected cell yield in batch culture, for optimization of time of harvest, and for information making it possible to compare culture performance at different scales. Additional Materials (also see Basic Protocol 6) Thawed vial of cryopreserved cells (0.5 ml to 1 ml at 107 viable cells/ml) or aliquot of healthy cells in exponential growth phase (107 cells total in ∼25 ml) Additional reagents and equipment for determination of specific growth rate (see Support Protocols 4 and 5) and specific protein productivity of cells (see Support Protocols 6 and 7)
Production of Recombinant Proteins in Mammalian Cells
1. Preincubate a 250-ml shaker flask, containing sufficient medium such that the volume after adding the aliquot of cells will be 50 or 100 ml total, in a humidified 37°C, 5% CO2 incubator. Add an aliquot of cells that will give a starting density of 0.5–1.0 × 105 viable cells/ml.
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2. Remove a 1.0-ml sample to determine post-inoculation viable cell density (APPENDIX 3C) and protein product titer (in supernatant or cells). Place the flask containing the rest of the cells on a rotating platform in the incubator and incubate with agitation at 80 to 100 rpm. 3. Sample 1.0 ml from the flask daily and determine viable cell density and protein product titer. 4. Feed cells by adding glutamine to a final concentration of 2 mM and glucose to a final concentration of 2 mg/ml on days 4, 6, and 8. Check the pH on a daily basis from day 4 onwards, and correct to pH 7.2 ± 0.1 if necessary using 0.5 N NaOH. 5. Keep a tally of sample volumes removed and the glucose, glutamine, and NaOH volumes added to enable calculation of the cell density dilution that has occurred as a result of additions and sampling. 6. Determine the specific growth rate (see Support Protocols 4 and 5) and productivity (see Support Protocols 6 and 7). REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
αMEM medium Purchase Minimal Essential Medium Alpha (αMEM; Irvine Scientific, Life Technologies, Sigma, JRH Biosciences, or HyClone) without L-glutamine and supplemented with 2.2 g/liter sodium bicarbonate. For growth medium, purchase formulation containing ribonucleosides and deoxyribonucleosides; for DHFR selection medium, purchase nucleoside-free formulation. Supplement with Pluronic F-68 (BASF) to a final concentration of 0.05% (w/v). Add 10 ml of 200 mM L-glutamine stock solution (HyClone; store at –20°C) per liter of medium immediately prior to use. Supplement as required by protocol with γ-irradiated fetal bovine serum (FBS; HyClone; store at –20°C prior to addition), heat-inactivated 30 min at 56°C. For DHFR selection, use dialyzed FBS (HyClone) to ensure absence of nucleosides. Store prepared medium (without the serum) up to 1 month at 4° to 8°C. DMEM/F12 custom-modified medium DMEM/F12 medium (1:1 DMEM/F12; e.g., Life Technologies, JRH Biosciences, HyClone, or Irvine Scientific), without thymidine, hypoxanthine, or L-glutamine, containing the following (final concentrations): 25 mM HEPES 5 g/liter D-glucose 0.11665 g/liter CaCl2 (anhydrous) 0.04884 g/liter MgSO4 (anhydrous) 0.061 g/liter MgCl2⋅6H2O 2.5 g/liter sodium bicarbonate 0.05% (w/v) Pluronic F-68 (BASF) Osmolality 280 to 320 mOsm/kg, adjusted with NaCl Store up to 1 month at 4° to 8°C Add L-glutamine to 4 mM final concentration just prior to use Supplement as required by protocol with γ-irradiated, dialyzed fetal bovine serum (FBS), heat-inactivated 30 min at 56°C. This formulation (minus the L-glutamine and serum) may be prepared to order by Life Technologies, JRH Biosciences, HyClone, or Irvine Scientific.
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Homogenization buffer Appropriate buffer (e.g., 50 mM Tris⋅Cl, pH 7.5; APPENDIX 2E) containing: 1 mM EDTA 0.1 to 1.0% Tween 80 (omit for extraction of soluble intracellular proteins) Store up to 2 weeks at 4° to 8°C Resuspension buffer 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 10 mM EDTA 100 µg/ml RNase A Store up to 1 week at 4° to 8°C COMMENTARY Background Information
Production of Recombinant Proteins in Mammalian Cells
Mechanisms of expression The level of expression of genes introduced into mammalian cells will depend on the gene copy number, the site of chromosomal integration, the transcriptional control elements, and mRNA stability. Availability of translational components—e.g., aminoacyl-tRNA, ATP, GTP, and other nutrients—is not likely to be limiting in mammalian cells growing under cell culture conditions. Expression vectors are modular in design, with various elements such as promoter-enhancer sequences, transcription termination/poly(A) sequences, multiple cloning sites, selection marker sequences with promoters and poly(A) sequences, and bacterial elements for replication and maintenance. Constitutive expression systems are most frequently used for stable expression. These include the strong promoters such as the cytomegalovirus (CMV) promoter, the β actin promoter, the SR α promoter, and the Rous Sarcoma Virus (RSV) promoter, which are capable of high levels of transcription. Following transcription in the nucleus, the primary transcript is capped and processed to remove any introns and the poly(A) tail is added before the mRNA is transported to the cytoplasm. Since most recombinant proteins are isolated as cDNAs, they do not contain introns. The poly(A) signal is required at the 3′ end of the foreign protein gene sequence for the transport and stability of the mature transcript. Commonly used poly(A) sequences are BGH, SV40, and HSV-tk; these are found in commercially available vectors. Although many commercially available plasmids contain introns, these offer no significant advantage in tissue culture cells (Brinster et al., 1988).
Translation of the message in the cytoplasm involves scanning of the message by the 30S ribosome complex from the cap in the 5′ to 3′ direction until it reaches an AUG codon in a favorable context, which then initiates translation. Thus, alteration of the original context of translation initiation to the Kozak’s consensus sequence—GCCA/GCCAUGG—in the DNA molecule, may improve expression through increased efficiency of translation initiation (Kozak, 1986). In the consensus sequence, the −3 position and the +4 position are most critical. Another essential requirement is a stop codon at the end of the foreign-gene cDNA for termination of translation. For efficient translation, the DNA sequence should be altered to incorporate preferred codons of the expression system of choice where there is great disparity in codon usage. The cDNAs should also contain the appropriate signal sequences in frame, either on the 5′ or 3′ end, depending on the targeted organelle (see Aitken, 1997). The selection marker is the other most crucial part of the vector apart from the cis elements that regulate expression of the protein. The cis elements required in the transcription unit of the selection marker are similar to those used for the protein. When the goal is to obtain high-level expression, it is important for the selection to be stringent and for the expression of the gene of interest to be linked to the expression of the marker gene. Methods for obtaining stringent selection involve weakening the expression of the marker protein through altering the context of translation initiation (Kozak, 1986), using a weak promoter such as SV40, or decreasing the mRNA stability (Miloux and Lupker, 1994). Mutation of the protein sequence to decrease activity has been used as a means to increase the stringency of selection, for example in the neomycin phosphotransferase gene (Yenofsky et al., 1990). In
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many earlier transfections, the marker gene and the protein gene were on separate plasmids. Current vectors carry both genes on the same plasmid, which provides better probability of cointegration in stable transfection. The singlevector strategy does not essentially guarantee that the marker and protein expression are correlated in cases of very low expression of the marker gene (Kaufman et al., 1985). This motivated the incorporation of internal ribosomal entry site (IRES) sequences from encephalomyocarditis (EMCV) virus (Kaufman et al., 1991) to allow linking of expression at the RNA level. Although the fusion-protein approach (UNITS 5.1, 6.6 & 6.7) provides the best linking of genes it is seldom applied to produce proteins for structural or biological characterization since it will pose issues such as the effect of the fusion on the structure and function of the fusion protein. Linking reduces the chance of vector rearrangement, which could result in nonexpression of protein of interest with continued expression of surrogate marker. The use of multiple elements that can be selected or screened for, such as neomycin and GFP together or neomycin and DHFR together, reduces the chance that clones with expression of two surrogate markers will end up with no production of the protein of interest. Thus far the methods discussed assume random integration of the heterologous DNA into the genome. The random-integration approach results in a very low frequency of integration into a transcriptionally active locus. This frequency can be increased through targeted integration using retroviral flanking sequences (Wurm et al., 1996), or by utilizing DNA sequences that flank the gene of interest in highproducer cell lines (Fukushige and Sauer, 1992; Morris et al., 1997). Selection and screening of transfected cells Selection and screening are the two different methods that help isolate cells expressing the marker gene. The marker gene is an indirect indication that the protein of interest is also being expressed. In selection, cells that do not express the marker are killed by the drug, and in screening, the cells that express the marker are isolated through flow cytometric or other techniques. Marker genes for selection are most often drug-resistance genes or easily detectable protein markers such as green fluorescent protein (GFP). In addition, a membrane-targeted protein that may be detected at the cell surface by a fluorescent-labeled specific antibody may act as a useful marker. Examples of drug-resis-
tance markers include hygromycin phosphotransferase, which confers resistance to hygromycin; neomycin phosphotransferase, which confers resistance to geneticin (G418); thymidine kinase, which confers resistance to aminopterin; xanthine-guanine phosphoribosyltransferase, which confers resistance to mycophenolic acid; adenosine deaminase, which confers resistance to 9-β-D-xylofuranosyl adenine; and a mutant dihydrofolate reductase, which confers resistance to high levels of methotrexate. Although several marker systems are available (Mortensen et al., 1997; Kriegler, 1990; Kaufman, 1990), the neomycin phosphotransferase system is most commonly used and is available in a number of commercially available vectors (Table 5.10.1). Amplification provides the means to go beyond the selection stage and enables higher gene dosage to be obtained. This usually leads to increased specific production of the gene of interest. Amplifiable elements are provided by recessive markers, which may be transferred into host cells lacking the related cellular function. Common examples are genes for dihydrofolate reductase (DHFR), which allows the transfected cells to grow in methotrexate (MTX), and for glutamine synthetase (GS), which enables growth in the presence of methionine sulfoximine (MSX). It is interesting to note that these drugs inhibit activity of marker enzyme through tight stoichiometric binding, whereas most drugs used in selection are the substrate for the marker enzyme and are inactivated by it. Other less frequently used recessive markers are adenosine deaminase (ADA), asparagine synthetase (AS), and ornithine decarboxylase (ODC; Mortensen et al., 1997; Kaufman, 1990). In the case of the recessive marker gene DHFR in CHO (dhfr−) cells, the methotrexate concentration is increased (when cells are growing adequately) by stepwise increases up to 10µM. After each step increase in the methotrexate concentration, the cells adapt and grow through increased dhfr gene expression. The higher-producing clones are screened and sorted for further rounds of amplification. The GS recessive marker has been used in CHO cells, and high titers (>100 mg/L) of foreign proteins have been obtained after two rounds of amplification using MSX (Cockett et al., 1990). DHFR can serve as a selection/amplification marker in DHFR+ cells, such as CHOK1, through the use of strong promoters to drive its expression, or with the low MTX affinity– mutated mouse dhfr gene (Simonsen and
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Levinson, 1983), or multiple selection markers (e.g., neoR; Kaufman, 1990). In these instances, selection for stable integration of dhfr requires moderately high (200 nM) MTX level to overwhelm the endogeneous DHFR and therefore the amplification potential is limited. Amplification provides a means to enhance the specific productivity of a cell, particularly where the level of expression is gene-dosage limited. Limits to the potential increased productivity exist and may be due to saturation of the translational capacity of the cells (Pendse et al., 1992) or to instability of gene copy number (Wurm, 1992). Thus, there is likely a point where the level of message is not limiting the overall expression of protein. Although high synthesis rates have been described, the overall expression rate may be limited by the secretory capacity of the cell (Cockett et al., 1990). Coexpression of a protein with higher levels of endoplasmic reticulum resident proteins (BiP, PDI, and calnexin) has been performed as a means to facilitate higher secretion rates, but there has been no general approach that guarantees higher secretion rate. Where cells are amplified to high gene copy number there is no absolute assurance that the amplicons are stable in the absence of the MTX. Following amplification, it is useful to subpassage the cells without MTX and determine the stability of production by detection of the protein of interest over several passages. Another method for evaluating the heterogeneity of gene copy number and stability of the foreign genes in the cell population is the use of fluorochrome-labeled MTX (MTX-F) and flow cytometry (Kaufman, 1978). Although there will always be significant time investment in amplification of expression, time savings may be possible by accelerating the selection and cloning procedure by use of flow cytometric sorting of producer cells. Use of these methods requires marker that can be noninvasively detected through fluorescencebased methods. A unique reporter that satisfies these criteria is the green fluorescent protein (GFP) from jellyfish (Chalfie et al., 1994; Kahana and Silver, 1996). Cells that express this protein can easily be detected and sorted using flow cytometric techniques (Subramanian and Srienc, 1996). GFP can be substituted with membrane-targeted proteins that are also amenable to flow cytometric detection and sorting. For example, the pHOOK system, commercially available from Invitrogen (pHOOK-2 vector; Capture-Tec 2 kit) can be used (Mortensen et al., 1997; Chesnut et al., 1996). In this
system, a single-chain antibody (sFv) against a hapten is expressed in addition to the transcription unit for the gene of interest. There are also two immunofluorescence tags, c-myc and HA, fused to the sFv. Antibodies against the sFv or the tags could be stained through direct immunofluorescence staining (primary antibody is conjugated with a fluorescent label, i.e., FITC or PE) or through indirect immunofluorescence staining (the primary antibody has no label but a secondary antibody that recognizes the primary is labeled). Alternatively, cells expressing the sFv can be separated using magnetic affinity cell sorting (MACS) where micron-size paramagnetic beads coated with the hapten are mixed with transfected cells and the cells to which the beads are bound are separated using a magnet. One disadvantage with the MACS is that it can only separate positive from negative but cannot differentiate between cells with various amount of surface proteins (density). This situation may eventually not be limiting, with the introduction of continuous magnetic separation devices that can fractionate on the basis of surfacemarker density (Chalmers et al., 1998). The use of surrogate markers can be completely avoided if the protein of interest can be directly quantitated and high-producer clones can be isolated. Such an assay, which measures product secreted from single cells encapsulated in agarose beads using an immobilized matrix for the capture and detection of the protein of interest, has been applied to the isolation of hybridomas with high antibody productivity (Gray et al., 1995). Mechanisms of secretion Secretion of protein from the cell into the medium is the most commonly used expression approach. In the case of glycoproteins, the routing of the protein through the cell is dependent on a leader sequence in front of the protein sequence for endoplasmic reticulum docking and uptake, and lack of the C-terminal endoplasmic reticulum–retention sequences. The protein will be transported from the endoplasmic reticulum (with addition of a lipid-linked oligosaccharide dolichol phosphate precursor and trimming of mannose and glucose) through to the Golgi (with trimming of mannose and addition of N-acetyl glucosamine, glucose, galactose, fucose, and sialic acid residues) and to the cell surface for secretion. In the absence of a leader sequence, the protein will remain in the cytoplasm where other motifs and chaperones may channel the protein to the nucleus, to the mitochondria, or into membranes.
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It may also simply accumulate in the cytoplasm as unglycosylated protein. For more information on the various protein motifs, see Aitken (1997) and UNIT 17.1. Proteins that are channeled through the ER and Golgi of the cell are presented to the cell’s glycosylation machinery. The compartmentalization and translocation mechanisms are briefly reviewed in UNIT 5.9. Choice of cell lines Chinese hamster ovary cells have become a widely used host for stable expression of heterologous genes. The CHO cell lines DG44 and DXB11 are DHFR− and commonly used to enable methotrexate selection and amplification of chromosomal integrated genes (see UNIT 5.9). Both these cell lines may be obtained from Dr. Lawrence Chasin of Columbia University (e-mail [email protected]). Selection approaches, other than methotrexate, use CHOK1 cells (ATCC #CCL-61). A number of CHO cell mutants exist that enable the production of glycoforms with removal or addition of linkages or sugar types. The mutants were isolated by their ability to grow in the presence of specific toxic lectins. The resistant cells do not bind the toxic lectins on their cell surface. This is a consequence of the mutant cell line lacking a specific glycotransferase enzyme and the resulting lack of the related lectin-binding carbohydrate structure on the cell surface. A number of these mutant CHO cells have been isolated and described and can be grown in culture (Stanley and Ioffe, 1995), providing interesting potential options for tailoring the carbohydrate structures. A mutant designated as Lec3.2.8.1 can provide glycoproteins with all the N-linked carbohydrate as Man5-7 and the O-linked truncated to a single GalNAc (Stanley, 1989). Other approaches to in vitro modulation of the glycoprotein carbohydrate structures involve use of specific inhibitors such as tunicamycin, castanospermine, deoxynojirimycin, deoxymannojirimicin, and swainsonine, which inhibit specific steps in the endoplasmic reticulum such as the addition of the lipid-oligosaccharide dolichol phosphate precursor to asparagine, or the trimming of glucose and mannose residues (UNIT 12.3). If the inhibitors are toxic to cells at their functional level, then gradual adaptation of the cells to the compounds may be required. Analysis of the detailed carbohydrate sequence and distribution of structures for purified glycoproteins is still challenging, although the advent of capillary zone electrophoresis and electrospray mass spectrometry have been use-
ful for fractionation and analysis of glycoforms present in recombinant proteins (Parekh and Patel, 1992; Yim, 1991). Although there are no preparative-scale separation methods to resolve larger quantities of glycoforms to homogeneity, immobilized lectin columns are available that enable purification of high-mannose glycoprotein from heterogeneous mixtures (UNIT 9.1). Use of purified specific exoglycosidases may be useful for sequential and specific digestion of the carbohydrate structures to a desired species (Brady et al., 1994). Choice of medium CHO cells readily adapt to suspension growth in medium containing lower Ca2+ and Mg2+, in the range of 0.2 to 0.6 mM respectively. Spinner MEM medium, with reduced calcium and increased phosphate, was first used for anchorage-independent growth in 1959. A commonly used medium is DMEM/F12 (Life Technologies, JRH Biosciences, or HyClone) supplemented with fetal calf serum (Life Technologies, HyClone). Following adaptation to suspension, cells may be subjected to stepwise changes in medium components—e.g., decreasing levels of serum. Adapting the cells to growth in medium containing low serum will generally make the cells grow with less clumping. Another benefit of growing in lowered serum is reduction of contaminant protein level, which makes the subsequent purification easier. Serum contains bovine transferrin and mitogenic growth factors. When the concentration of serum is <1% (v/v), addition of insulin or IGF to the medium is needed. Mechanisms of cell growth Cells may be grown in shaker flasks, spinners, or larger bioreactors. The choice of approach is dictated by the quantity of final purified product required (Table 5.9.3 and Table 5.9.4). Bioreactors may be set up in glass spinner vessels fitted for pH and dO2 control (Griffiths, 1992). Larger-scale operation requires use of stainless steel vessels, which are available from 10 to 10,000 liters. Scale-up to 100 liters is adequate for intermediate-scale production. Such reactors may be operated in batch, fed batch, or continuous mode. Batch operation is the least complicated approach and the one that is least vulnerable to contamination, since fewer manipulations are required. Highest titer levels are achievable in fed batch because of the high cell density and high specific growth rate. Perfusion reactors enable high product titer and
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productivity, which result from the higher cell density achievable, although the lower target growth rate may decrease the specific productivity. Cell-specific productivity is usually higher in fast-growing cells because of an increase in the frequency of S-phase cells in culture, and the use of S-phase-active promoters (Gu et al., 1993). High density is obtained by operation in continuous mode with cell retention in the bioreactor. Cell-retention devices include internal or external inclined settlers, spin filters, or external microporous filtration devices. Recently, an acoustic cell settler (Sonosep) and a cell-recycling external centrifuge were developed (Centritech AB, DuPont) and have been made available for compact operation up to 40 liters. Cell-retention devices should provide a cell retention efficiency of >75% to support a reactor steady-state cell density at a minimum specific growth rate of 0.3 per day. A target growth rate should be stipulated in continuous-mode strategies. Perfusion reactors require an initial fed-batch phase to establish high cell density. As the specific growth rate declines, the removal rate of cells is set to the target value (typically 0.25 to 0.4 per day). Following completion of batch growth, or during continuous-mode operation of cell culture bioreactors, it is necessary to recover the product. The primary recovery steps may be designed to harvest the cells when the product is intracellular, or most commonly to harvest the conditioned medium containing the secreted product. The types of methods used are not very different from those typically used in bacterial or yeast bioprocessing. A summary of typical recovery operations used at small and large scale is given in Figure 5.10.4. Mammalian cells are ∼10 to 20 times larger in dimension than bacteria and possess a lipid bilayer cell wall, which renders the mammalian cell much weaker than the more complex cell walls of bacteria and yeast. The larger size provides for a sedimentation rate of 10 to 20 cm/min at 1 g, in the order of 100 times larger than that of bacteria; hence, low-speed centrifugation is able to separate the cells without compression and cell damage. Thus, recovery of the product from the culture may be readily accomplished at laboratory and compact bioreactor scale. Batch centrifuges capable of spinning at 2000 to 5000 × g can separate cells from the conditioned medium. For larger volumes (>20 liters), tangential-flow microporous membranes (0.2 µm pore size) may be used to obtain a permeate containing extracellular pro-
tein and a retentate containing the concentrated cells. It is important to minimize cell damage and product contamination and loss, which could result from exposure to shear forces or bursting bubbles. Cross-filtration devices are available for mammalian cell operations and feature open-channel design to minimize shear damage. These operate at low transmembrane gauge pressure (<5 psi) to avoid cell deformation and fouling at the membrane surface. Since most mid-scale-up to pilot-scale continuous centrifuges contain acceleration zones which can damage cells, they are rarely used in primary recovery of cells or conditioned medium. In dealing with large volumes of conditioned medium from perfusion reactors, it may be possible to allow the cells in the conditioned medium to settle overnight. The top 80% of the liquid could then be clarified by tangential flow filtration (TFF) or by depth-filtration membranes (Seitz, Cuno, FPI) and then concentrated using tangential cross-flow ultrafiltration. If intracellular product is desired, it is necessary to disrupt the cell membrane to release the product. Typically, the first step in such a scheme would be to separate the cells from the culture medium to eliminate the serum contaminants. This is achieved by centrifugation or gravity separation of the cells overnight in a cold room, followed by decanting of the clear top liquid phase. The cells can be resuspended in a simple buffer containing a low concentration of a suitable nonionic detergent to weaken the cell membrane. Exposure of the cells to high laminar shear using a homogenizer (APVGaulin), or recirculation at high rates using a rotary displacement pump, will rupture the cells further and release the product. After cell disruption, the difficult step of separating the cell debris from the released product must be accomplished. For this step, high-speed centrifugation could be applied or TFF could be used at low transmembrane pressure and high recirculation rate employing screened membranes. Throughout the recovery operations the product may be exposed to physical conditions, or degradative enzymes released by cell lysis, which may modify the protein. Typically, serine proteases are active at neutral pH and lysosomal-based proteins are active at low pH (4 to 6). Resistance to proteolysis may result from inherent structural properties of the proteins such as buried protease sites or the conditions of processing such as inclusion of protease inhibitors or speedy processing at <10°C. The presence of sialidase, as a result of cell lysis,
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bioreactor
perfusion conditioned media
depth filtration
bioreactor cell broth
intracellular product
secreted product
laboratory scale (< 20 liter(
pilot scale (> 20 liter(
laboratory scale (< 20 liter(
pilot scale (> 20 liter(
centrifugation
concentration (MF-TIFF)
centrifugation
MF-TFF
resuspend pellet in PBS buffer resuspend pellet in homogenization buffer
reduce volume 5-10-fold
filter supernatant
diafiltration (wash concentrate with 3+0 5 vol. PBS)
cell lysis
concentration (UF)
filtrate
add homogenization buffer centrifugation
cell lysis sterile filtration
recover supernatant concentration (UF-1 TIFF) sterile filtration
depth filtration or MF-TFF filtrate or permeate
purification (or storage)
purification (or storage)
Figure 5.10.4 Cell culture product recovery flow schemes for laboratory and pilot scale processes. Options for intracellular and secreted proteins are shown. For details, see Basic Protocol 8. Abbreviations: MF, microfiltration; TFF, tangential-flow filtration; UF, ultrafiltration.
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may also modify the glycoprotein and decrease the sialic acid content. Processing quickly and keeping the product at 4° to 10°C is the basis for a recovery operation with minimal physical product damage. Where specific problems of aggregation, proteolysis, and degradation are encountered, the researcher may have to consider use of chemical inhibitors.
Critical Parameters and Troubleshooting Expression vector construction No molecular biology protocols have been provided for constructing an expression vector, since such information is widely available (e.g., Ausubel et al., 1998, and Sambrook et al., 1989). However, the following guidelines should be considered no matter which approach is used. Construction of the vector with most commercial vectors is done by subcloning the insert cDNA into the multiple cloning sites by restriction digestion of vector and insert and maybe blunt ending with T4 DNA polymerase or Klenow polymerase, after which the fragments are ligated with T4 DNA ligase and transformed and amplified in E. coli. PCR is usually employed if modifications need to be made in the cDNA such as alteration of context of translation initiation or in-frame fusion to a signal sequence or another protein. Following purification of the PCR product it should be sequenced before being used for expression. The preparation of DNA is usually straightforward, but resuspension of the E. coli after the initial pelleting is important for efficient lysis. Centrifugation at higher speed or longer duration will make the pellet compact and difficult to resuspend. Tapping the tube on the bench aids the suspension process. One other critical step is lysis, where going beyond the recommended time is to be avoided. The quality of plasmid DNA, and especially the presence of endotoxins, has been shown to affect the transfection efficiency. If it is determined that this is an issue, a new plasmid preparation kit for endotoxin removal is available from Qiagen. The E. coli host chosen for plasmid preparation should be recombination deficient (recA1 or recA13) and endonuclease deficient (endA1) to avoid rearrangement and degradation of the plasmid sequence. Production of Recombinant Proteins in Mammalian Cells
Transfection Before embarking on the laborious and lengthy process of transfection, selection, and amplification, it is advisable to do transient
expression of the protein of interest. At 24 to 48 hr post transfection (which may be extended to 72 to 96 hr for secreted proteins), the cell or supernatant fraction is harvested and assayed for the protein of interest using ELISAs and immunoblots (UNIT 10.8). The transfection protocol may also be initially tested using reporter genes—e.g., GFP, β-galactosidase, or luciferase—in a vector construction that is identical to that of the protein of interest (Kain and Ganguly, 1996). Lipofection (see Basic Protocol 2) and electroporation (see Alternate Protocol 1) are popular methods of gene transfer since a broad range of cell types can be transfected. The efficiency of transfection may be expressed as the total amount of gene product obtained at peak expression (24 to 48 hr), and this can be by directly assayed from cell lysates or supernatants. In cases where single-cell expression is measured using flow cytometry, as with GFP or through immunofluorescent staining, the transfection efficiency is the product of the percentage of cells positive for gene expression multiplied the average level of gene product per cell. In CHO cells, the highest transfection efficiency was obtained when 6 µL of lipid and 1 µg of DNA were used for transfecting cells in a 6-well plate (Subramanian and Srienc, 1996). The inoculation cell density and the amounts of lipid and DNA recommended in Basic Protocol 2 were obtained by scaling up in proportion to the surface area available for growth in 100-mm dishes or T-75 flasks. The most common problems encountered in transient-expression experiments are the lack of expression and cytotoxicity. Most often, the optimal conditions are a compromise between transfection efficiency and post-pulse viability. Troubleshooting lack of expression or lowlevel expression can be accomplished by optimization of the conditions of transfection. In lipofection, the optimization parameters include: lipid (24 to 72 µl); DNA (4 to 16 µg); cell density; medium (αMEM, DMEM, or Opti-MEM); and incubation time with complexes (3 to 9 hr). In electroporation, the voltage is the most critical variable. In addition, the amount of DNA can be increased to 40 µg, but this increase will usually decrease the viability later. Similarly, the volume in the cuvette can be reduced by 50% or the medium could be changed to increase the conductivity. This will also increase the cytotoxicity. Cell death is more of concern in electroporation, giving viabilities after pulsing of up to 50%. This may be acceptable and may be close to optimal in
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many cell types. In lipofection, cytotoxicity is less of a concern; here it can be reduced by decreasing the amounts of lipid and DNA or the time of incubation with complexes. The use of linearized DNA for stable transfection is recommended with the electroporation method. Circular DNA is essential for transient expression with either method. The use of necessary positive and negative controls is a must for any expression experiment. It is recommended that sensitive reporters, such as the E. coli β-galactosidase or the firefly luciferase, be included in transfection experiments as positive controls. Negative controls could be included through the use of no DNA or through using the vector without any protein insert. The negative control will be used to determine background (baseline) and any nonspecificity of assay. Selection and screening In the amplification process, after selecting the initial clones, those clones or a subset of those that represent high producers may be individually amplified. This means no further cloning is required and the danger of losing clones by outgrowth of other clones is removed. However, much more work may result, depending on the number of clones to be taken along. The amplification of individual clones is a risky enterprise since they may not amplify well. This approach may work when there are two selection markers in the same vector such as neo and DHFR (as a fusion protein or in separate transcription units) and the initial selection may be made in nucleoside-free medium containing G418. Following this initial selection of clones a few high producers may be carried on through the amplification procedure separately. The number of clones at the end of selection that need to be analyzed and the method of analysis (serum level, suspension versus adherent, production conditions, and cell growth) are very important if the requirement is to obtain a high producer. In selection and amplification, if growth is not impeded (especially in amplification) with the drug level chosen, either drug concentration should be increased or plate cell density should be decreased. In the selection procedure, if the number of cells per well exceeds what would give rise to 1 clone per well, then multiple clones will exist in the same well, which is not effective if single clones are required. The number of cells to be plated depends on the marker expression and the drug concentration. Even if the frequency of a stable transfectant is 1 in 105 and
the number of cells plated is 100, there is always a chance that multiple clones will get into the same well, so recloning is recommended after selection/amplification procedures so as to obtain monoclonal cultures. Limiting dilution through visual inspection is the best method, since the usual plating of 0.3 cells per well to minimize number of wells with >1 cell according to the Poisson distribution again does not assure monoclonality. Assuming Poisson distribution of viable cells, plating at a density of 0.3 cells per well would result in 74.08% wells being empty, 22.22% containing 1 cell, and ∼3.69% containing >1 cell. The cloning efficiency (number of observed growth-positive wells divided by the number of expected growth-positive wells) will likely be less than expected since not all single cells form colonies as a result of stress from evaporation (higher osmolarity) or their inability to grow as single cells. Diluting the medium can be useful if evaporation and the resulting increase in osmolarity become a problem. Low serum level may reduce cloning efficiency, and it is recommended that 10% serum be included in the medium at this stage. Even at this level of stringency, it is quite possible that cultures obtained may not be monoclonal because of the growth advantage conferred upon wells with multiple cells. Visual inspection provides the only confirmation, and should be incorporated after dilution and plating to eliminate the nonsingle wells. Almost invariably, as cells are amplified, there is no improvement in productivity after ∼1 µM MTX, either because of saturation of some component in the cellular machinery or as a result of mechanisms other than amplification of DHFR for gaining resistance to methotrexate, such as a reduced influx of the drug. Amplification usually results in a high gene copy number in a nonchromosomal location, such as double minute chromosomes, which are highly unstable and which may be lost during cell division. This can be monitored as a drop in productivity following culture of cells in MTX-free medium. Low-level amplification, at or below 1 µM, is recommended to reduce instability-related issues. Maintaining cells in culture It is critical for any experiment that the host cells be healthy, which is made possible by maintaining them in the exponential phase of growth in medium containing 10% FBS. It is recommended that the growth of the host be checked in 10% FBS in complete medium, that
Production of Recombinant Proteins
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Production of Recombinant Proteins in Mammalian Cells
the cells be split at appropriate times, and that confluent cells not be exposed to prolonged nutrient limitation. A doubling time of 24 ± 8 hr is typical for most continuous cell lines. It is also advisable to start work with a frozen vial from a cell bank, which the investigator can go back to. This is important since cells that have been in culture for fewer generations (1 to 2 months after thawing) are generally easier to transfect. Some of the host cells used require special medium additives—e.g., proline for all CHO cells, as well as hypoxanthine (or adenine and deoxyadenosine), thymidine, and glycine for DHFR− CHO cells (DG44 and DXB11). CHO cells have a tendency to clump when grown at high cell density in serum-containing medium. Clumping may be minimized by adapting cells to growth in lower serum (≤0.5% FBS). Reduced-calcium medium such as DMEM/F12 also may produce less clumping of cells. In cultures that contain clumping cells, the only secure counting method is through use of dispase and hemacytometer/trypan blue counting (APPENDIX 3C). When culturing cells, it is critical to keep the temperature at 37°C ± 0.5°C. Cells will generally not tolerate rapid increases in temperature to levels above 38.5°C and will die following such shock. Exposure to lower temperature has less drastic consequences, but should be minimized so as to prevent metabolic perturbations. Optimal pH for growth is ∼7.2, but cells will grow between pH 6.8 and 7.5. The optimal pH for protein expression may differ from the optimal growth temperature, and it may be worthwhile to investigate growth of cells at pH 7.0, 7.2, and 7.4 in shaker flasks to determine if there is a useful advantage to be gained in production at varied pH. All mammalian cells require Lglutamine as an energy-providing carbon source in addition to glucose. Glutaminolysis leads to the accumulation of ammonia in the medium, while glucose consumption leads to the accumulation of lactate in the medium. Both these catabolites can reach levels that may inhibit growth of cells. Typically ammonia reaches toxic levels at ∼15 to 20 mM and lactate may be inhibitory at 20 mM. In cases where high cell density is desired, it is most probable that the ammonia and lactate level may limit the culture. Glutamine may also undergo spontaneous degradation to pyrrolidonecarboxylic acid and ammonia. The rate of this degradative process increases with temperature. At 37°C, ∼20% of the glutamine is converted after 2 days, whereas at 4°C, <20% loss is experienced after 65 days. At small scale, medium can be pur-
chased without glutamine and the user can add the reagent to give a concentration of 4 to 8 mM glutamine just prior to using the medium. Clearly, glutamine stock solutions (200 mM) should be stored frozen prior to use. Ammonia and lactate can be determined rapidly using convenient metabolite analyzers such as the Kodak biolyser. In the case of large-scale operation where larger volumes of medium are needed, the user should order and use fresh complete medium with glutamine added to 4 to 8 mM. Hyperbaric levels of oxygen, which are possible where oxygen is used to supplement sparge gas, may produce peroxide and oxygen radicals which can result in poor growth and cell death. Careful control of dissolved oxygen is recommended for largescale reactors. In the case of shaker flasks or small spinners, air is usually sufficient to support cells up to typical levels of density. In spinners or shaker flasks, pH control is done manually, and care should be taken to add the base slowly to a swirling liquid bulk. Use of 0.5 N NaOH for pH control is recommended to avoid local high-pH spots. In reactors, use of a 6 mm internal diameter tube with a short U-bend at its end below the surface and close to the agitator is adequate. Buffering of medium using sodium bicarbonate/CO2 is not unproblematic, and alkaline pH excursions are common when adding base or where flasks are left outside on a bench in the absence of CO2. Such excursions may cause injure cells or affect nutrient balances. The authors recommend use of medium containing 50 to 75 mM HEPES for shaker flasks or spinner culture where manual pH adjustment is required and 25 mM in reactors where automatic pH control is available. Bicarbonate is required in the medium and is present at ∼2.5 g/liter. Preincubation of spinners and flasks is important to avoid the formation of hydroxyl ions and CO2 from the bicarbonate. Always include 5% CO2 in the sparge and headspace gas when operating at low cell density in bioreactors. As the cell density is higher, in particular in high-density perfusion culture, the cells produce enough CO2; therefore supplementation of sparge gas is not required. Mammalian cells are sensitive to turbulent shear, especially as may be experienced around the liquid surface where sparge gas bubbles may be bursting. Cell death can be very significant in such cases, but addition of pluronic polyol F68 (BASF) has proven to minimize bubble–cell interaction. All medium used should contain 0.05% (w/v) of Pluronic Polyol F68. Certain batches of Pluronic may be
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toxic (which is related to the polyoxypropylene group) and preliminary screening prior to purchase is a secure way to avoid any serious problems. Cell growth and productivity The specific productivity of the cell (see Support Protocols 6 and 7) is a function of the particular protein being expressed. A shaker flask growth curve (see Support Protocol 9) and titer profile will provide information on the specific productivity during the initial exponential phase and will enable calculation of the effect of lower growth rate on specific productivity as the culture progresses through to day 10. Generally the specific productivity is highest in cells growing at their maximum specific growth rate and decreases as specific growth rate is decreased. Thus, the highest specific productivity is seen in batch reactors, and high productivity can be obtained in feed-and-bleed mode. Perfusion reactors operate at lower specific growth rates with associated lower specific productivity, but provide the benefit of high cell density and higher cellular yield in the given medium. In cases where there is evidence of proteolysis or glycosidase degradation of the product, it is advisable to operate a perfusion system with high viability. In this type of system the product residence time at 37°C and cell lysis are minimized. Further minimization of proteolysis may be obtained by addition of specific protease inhibitors such as aprotinin (for inhibition of serine proteases). Addition of sialidase inhibitors such as 2,3-dehydro-2-deoxy-N-acetylneuramic acid (Gramer et al., 1995) has been described. Many of the low-molecular-weight inhibitors are toxic and undesirable in production of material destined for clinical applications. In such cases, rapid processing at low temperature (<10°C) is the best option. Where proteins are exquisitely sensitive to proteolysis, it is advisable to operate with 0.5% to 1.0% FBS in the cell culture medium as a reagent to prevent proteolysis (Tiege et al., 1994), as serum contains the potent protease inhibitor α2-macroglobulin. Many cell culture–dictated conditions may influence the microheterogeneity of the glycoprotein produced. In order to reproduce equivalent product from cell culture batches, it is important to standardize the cell culture protocol. Spin filters are a commonplace option for cell retention in bioreactors; however, fouling of the screen with cells is usual after 2 to 3 weeks of operation. The passive settler devices
are low-maintenance and predictable for cells of a given size and clumpiness. At present, settlers are custom-designed, although several approaches have been described in the literature (Batt et al., 1990; Brown et al., 1993). Harvesting cultures When harvesting cultures, it is important to maintain the cells intact to prevent contamination of the product in the extracellular compartment with intracellular constituents or to prevent loss of product if it is intracellular. Use of peristaltic pumps with size 16 or 24 silicone tubing or use of gravity transfer is recommended. Static laboratory centrifugation is a gentle approach and useful for clarifying relatively small volumes of culture liquid. The largest rotor capacities are ∼5 liters, and the researcher can gauge the time investment needed to process the material with such an approach. It would be feasible to process up to 30 liters of culture broth in ∼2.5 hr using such an approach. Where cross-flow filtration is employed to separate cells and medium, it is important to minimize cell lysis, which could result in a decrease in yield, purity, or performance of the membrane system. Operation at cold-room temperature (4° to 12°C) is advisable as in all of the downstream recovery operations. Avoidance of average wall shear rates greater than 3000 sec−1, using membrane pore sizes of 0.2 µm and minimization of pump shear rates will minimize the potential for cell damage (Maiorella et al., 1991). If small membrane systems of 1 ft2 are used, then peristaltic pumps (Watson Marlow or Cole-Parmer) are adequate, while for larger systems the larger-gap rotary positive- displacement pumps are employed (Flowtech, Waukesha). Cross-linked cellulosic membranes are available (Sartorius Hydrosart) which are useful in minimizing surface fouling and for facilitating high protein flux.
Anticipated Results For plasmid preparation, 1 liter of E. coli culture grown to an OD650 of 4.0 can yield up to 5 mg of high-copy-number plasmids. The DNA may be stored frozen without loss of quality. Confluent growth on a surface corresponds to a cell-surface density of 1–1.5 × 105 cells/cm2, so one T-75 flask can provide ∼1 × 107 total viable cells and one T-175 flask can provide ∼2–3 × 107 total viable cells, enough for the transfection process. Approximately 50% of the cells will have taken up the plasmid
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Table 5.10.2
Specific Productivities for a Selection of Proteins Expressed in Chinese Hamster Ovary Cells
Protein
Cell Line
Promoters Amplification GOI Heterologous proteins TIMP CHO-K1
hCMV-TIMP
TIMP
hCMV-TIMP
CHO DXB11 CHO DXB11 CHO DXB11 CHO DXB11 CHO DXB11
htPA IFN-β IFN-γ h-AT III
Monoclonal antibodies Campath-1H CHO DXB11
Subunit vaccines HBsAg CHO DHFR− aAbbreviations:
SM
SV40 late-GS 20 µM MSX
SV40 earlydhfr Ad2 MLP-tPA Ad2 MLPdhfr SV40 early-IFN SV40 earlydhfr SV40 early-IFN SV40 earlydhfr SV40 earlyMMTV-LTRATIII dhfr
1 µM MTX
Productivity (µg/10−6 cells/day)
110 17.5
0.5 µM MTX
66
1 µM MTX
0.5
0.3 µM MTX
10
10 µM MTX
22
β-actin–light weak SV40dhfr chain β-actin–heavy MMT-neo chain
1 µM MTX
100
SV40 earlyHBV
10 µM MTX
1-2 (stable)
MMTV-LR dhfr
Notes
Reference
copy number ∼30 copy number∼100, dual plasmid cotransfection dual plasmid cotransfection dicistronic vector
Cockett (1990)
Kaufman (1985) Chernajovsky (1984) Mory (1966) Zettlmeissl (1987)
Dual plasmid transfection
Page (1991)
copy number Michel (1985) ∼400 (T-flask)
GOI, gene of interest; MTX, methotrexate; SM, selection marker.
Production of Recombinant Proteins in Mammalian Cells
vector following lipofection versus approximately 10% in electroporation. The proportion of cells that chromosomally integrate the foreign DNA is low, in the range of 0.001% to 0.01%. Of these cells, the frequency of stable high producers ∼0.1% in general, but may reach 1% in cases where targeted integration methods are used. From these numbers, it is apparent that an initial total of 1 × 107 cells will give ∼10 to 100 productive stable clones with perhaps 1 to 10 clones being high producers. The specific productivity of the transfected CHO cells will be heterogeneous within the cell population. The level and range of productivity is largely a function of the protein being expressed as well as the particular culture conditions. Typical specific productivity levels are ∼5 to 20 µg/106 cells/day, although it may be higher, with examples indicating recombinant antibody production at >100 µg/106 cells/day. Lower values, ∼1 to 5 µg/106 cells/day, are typically found for recombinant viral-antigen proteins. Amplification may increase the specific productivity 10- to 100-fold, depending on the protein and the clone selected. Examples of
productivities following amplification that have been reported in the literature are provided in Table 5.10.2. Where a high producer is initially selected, the amplification process may provide little gain in productivity. Productivity may be limited by secretion rates in the case of large glycoproteins with numerous disulfides. CHO cells adapt quickly to growth in suspension and can be adapted to growth in low (0.5%) serum in ∼15 to 20 passages. High specific growth rate is usually reestablished in 3 to 5 passages following a step change in a medium concentration. Complete elimination of serum may take a further 5 to 10 passages before cells become truly serum-independent. Once cells are adapted to growth in medium such as DMEM/F12, it is possible to obtain a cell density of 2–4 × 106 per ml in batch shaker flasks or spinners. In the case of bioreactors, with precise pH and dissolved oxygen control, cell densities may reach 4 × 106 cells/ml for batch operation and as high as 10 × 106 cells/ml for bioreactors operating in perfusion mode. Product titers of 5 to 20 mg/liter are possible in batch-mode flasks, spinners, and bioreactors. Fed batch reactors may provide higher titers,
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since the promoters that have been recommended are generally dominant in the S phase of the CHO cell cycle and faster specific growth will result in a higher specific productivity than slower growing cells. A cell that has a specific productivity of 5 pg/cell/day at a specific growth rate of 0.3 per days is capable of providing 75 to 100 mg/day/liter culture in a perfusion reactor operating with a cell density of 15 × 106 cells/ml, a dilution rate of 1.0, and 70% overall cell retention. The purity of the product, with a titer of 30 mg/liter, is estimated at ∼5% for cells grown in bioreactors that achieve 10 × 106 cells/ml with a viability of 90%, using 1% FBS. In serum-free medium under the same conditions, the purity may be 10%. Intracellular-produced proteins rarely exceed 1% of total intracellular protein. Recovery yield of the product from the reactor should be in the region of 75% to 90%. At large scale, the recovery yield for intracellular protein may be lower (50% to 60%) if large volumes of high-debris-containing material result in membrane fouling. Use of cellulose-based membranes and/or depth filtration may improve such low yields.
Time Considerations Preparation of DNA and the process of transfection can be accomplished in <1 week. Following transfection, it takes 3 weeks to obtain colonies on plates. Expansion of the colonies up to the tissue culture flask stage requires 2 further weeks. Thus it will take a total of ∼6 weeks to obtain stable transfected cells that grow in T-75 flasks in the presence of the selection drug. Subjecting the cells to amplification, such as an increased level of methotrexate, will take up 3 weeks to produce a pool of cells. Each further round of amplification will require 3 weeks. Based on these time lines, a cell that has been subjected to two rounds of amplification would be available 12 weeks after initiating the full process. Following amplification, the process of single-cell cloning requires 3 weeks to obtain colonies and a further 2 weeks to scale up to the T-75 flask stage. At this point the higherproducing clones can be adapted from tissue culture flask growth to shaker flask growth in 1 week, and can be further adapted to 1% serum in a 2 weeks, 0.5% serum in 5 weeks, and 0% serum in 8 weeks. Laying down a cell bank of the adapted cells takes ∼1.5 weeks. Based on a cell-growth doubling rate of 1 to 2 days, a batch culture requires 5 to 7 days to complete, and depending on the scale would
require an inoculum buildup process. For a 100-liter-scale process it may take 10 to 15 days to scale up to a suitable inoculum volume. Recovery of product is usually performed as a batch process and should require <8 to 10 hr.
Literature Cited Aitken, A. 1997. A library of concensus sequences. Methods Mol. Biol. 64:357-367. Aruffo, A. 1997. Transient expression of proteins in COS cells. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 16.13.1-16.13.7. John Wiley & Sons, New York. Ausubel, F.A., Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G., Smith, J.A., and Struhl, K., (eds.) 1998. Current Protocols in Molecular Biology. John Wiley & Sons, New York. Batt, B.C., Davis, R.H., and Kompala, D.S. 1990. Inclined sedimentation for selective retention of viable hybridomas in a continuous suspension bioreactor. Biotechnol. Prog. 6:458-464. Brady, R.O., Murray, G.J., and Barton, N.W. 1994. Modifying exogenous glucocerebrosidase for effective replacement therapy in Gaucher Disease. J. Inherited Metab. Dis. 17:510-519. Brinster, R.L., Allen, J.M., Behringer, R.R., Gelinas, R.E., and Palmiter, R.D. 1988. Introns increase transcriptional efficiency in transgenic mice. Proc. Natl. Acad. Sci. U.S.A. 85:836-840. Brown, P.C., Wininger, M., Chow, R., and Cox, J. 1993. Perfusion culture of CHO cells in suspension facilitated by inclined sedimentation chambers for cell recycle. Presented at the 205th National Meeting of the ACS, Division of Biotechnology, March 31, 1993. Budelier, K. and Schorr, J. 1998. Purification of DNA by anion-exchange chromatography. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 2.1.11-2.1.18. John Wiley & Sons, New York. Chalfie, M., Tu, Y., Euskirchen, G., Ward, W.W., and Prasher, D.C. 1994. Green fluorescent protein as a marker of gene expression. Science 263:802805. Chalmers, J.J., Zborowski, M., Sun, L., and Moore, L. 1998. Flow- through immunomagnetic cell separation. Biotechnol. Prog. 14:141-148. Chernajovsky, Y., Mory, Y., Chen, L., Marks, Z., Novick, D., Rubinstein, M., and Revel, M. 1984. Efficient constitutive production of himan fibroblast interferon by hamster cells transformed with the IFN-β1 gene fused to an SV40 early promoter. DNA 3:297-308. Chesnut, J.D., Baytan, A.R., Russel, M., Chang, M-P., Bernard, A., Maxwell, I.H., and Hoeffler, J.P. 1996. Selective isolation of transiently transfected cells from a mammalian cell population with vectors expressing a membrane anchored
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single-chain antibody. J. Immunol. Methods 193:17-27.
K. Struhl, eds.) pp. 9.7.22-9.7.28. John Wiley & Sons, New York.
Cockett, M.I., Bebbington, C.R., and Yarranton, G.T. 1990. High level expression of tissue inhibitor of metaloproteinase in Chinese hamster ovary cells using glutamine synthetase gene amplification. Biotechnol. 8:662-667.
Kain, S. and Ganguly, S. 1996. Overview of genetic reporter systems. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 9.6.3-9.6.12. John Wiley & Sons, New York.
Earl, P. and Moss, B. 1998. Characterization of recombinant vaccinia viruses and their products. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 16.18.1-16.18.10. John Wiley & Sons, New York. Earl, P., Wyatt, L., Carroll, M., and Moss, B. 1998a. Generation of recombinant vaccinia viruses. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 16.17.1-16.17.16. John Wiley & Sons, New York. Earl, P.L., Cooper, N., Wyatt, L., Carroll, M., and Moss, B. 1998b. Preparation of cell cultures and vaccinia virus stocks. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 16.16.1-16.16.7. John Wiley & Sons, New York. Elroy-Stein, O. and Moss, B. 1998. Gene expression using the vaccinia virus/T7 RNA polymerase hybrid system. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 16.19.1-16.19.9. John Wiley & Sons, New York. Fukushige, S. and Sauer, B. 1992. Genomic targeting with a positive-selection lox integration vector allows highly reproducible gene expression in mammalian cells. Proc. Natl. Acad. Sci. U.S.A. 89:7905-7909. Gramer, M.J., Goochee, C.F., Chock, V.Y., Brousseau, D.T., and Sliwkowski, M.B. 1995. Removal of sialic acid from a glycoprotein in CHO cell culture supernatant by action of an extracellular CHO cell sialidase. Bio/Technology 13:692-298. Gray, F., Kenney, J.S., and Dunne, J.F. 1995. Secretion capture and report web: Use of affinity derivatized agarose microdroplets for the selection of hybridoma cells. J. Immunol. Methods 182:155-163. Griffiths, B. 1992. Scaling-up of animal cell cultures. In Animal Cell Culture, 2nd ed. (R.I. Freshney, ed.), pp. 47-93. IRL Press, Oxford. Gu, M.B., Todd, P., and Kompala, D.S. 1993. Foreign gene expression (β-galactosidase) during the cell cycle phases in recombinant CHO cells. Biotechnol. Bioeng. 42:1113-1123.
Production of Recombinant Proteins in Mammalian Cells
Kahana, J. and Silver, P. 1996. Use of the A. victoria green fluorescent protein to study protein dynamics in vivo. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and
Kaufman, R.J. 1978. Quantitation of dihydrofolate reductase in individual parental and methotrexate-resistant murine cells. J. Biol. Chem. 16:5852-5860. Kaufman, R.J. 1990. Selection and coamplification of heterologous genes in mammalian cells. Methods Enymol. 185:537-566. Kaufman, R.J., Wasley, L.C., Spiliotes, A.J., Gossels, S.D., Latt, S.A., Larsen, G.R., and Kay, R.M. 1985. Coamplificaton and coexpression of human tissue-type plasminogen activator and murine dihydrofolate reductase sequences in Chinese hamster ovary cells. Mol. Cell. Biol. 5(7):1750-1759. Kaufman, R.J., Davies, M.V., Wasley, L.C., and Michnick, D. 1991. Improved vectors for stable expression of foreign genes in mammalian cells by use of the untranslated leader sequence from EMC virus. Nucl. Acids Res. 19(16):4485-4490. Kingston, R.E., Kaufman, R.J., Bebbington, C.R., and Rolfe, M.R. 1993. Amplification using CHO cell expression vectors. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.), pp. 16.14.116.14.13. John Wiley & Sons, New York. Klickstein, L.B. and Neve, R.L. 1991. Ligation of linkers or adapters to double-stranded cDNA. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.), pp. 5.6.1-5.5.10. John Wiley & Sons, New York. Kozak, M. 1986. Point mutations define a sequence flanking the AUG initiator codon that modulates translation by eukaryotic ribosomes. Cell 44:283-292. Kriegler M. 1990. Gene Transfer and Expression, A Laboratory Manual. Stockton Press, New York. Liljestrom, P. and Garoff, H. 1995. Expression of proteins using Semliki Forest virus vectors. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 16.20.1-16.20.16. John Wiley & Sons, New York. Lucas, B.K., Giere, L.M., DeMarco, R.A., Shen, A., Chisholm, V., and Crowley, C.W. 1996. High level expression of recombinant proteins in CHO cells using adicistronic DHFR intron expression vector. Nucl. Acids Res. 24(9):1774-1779. Maiorella, B.M., Dorin, G., Carion, A., and Harano, D. 1991. Crossflow microfiltration of animal cells. Biotechnol. Bioeng. 37:121-126.
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Michel, M-L., Sobczak, E., Malpiece, Y., Tiollais, P., and Streeck, R.E. 1985. Expression of amplified hepatitis B surface antigen in Chinese hamster ovary cells. Bio/Technology 3:561-566.
Stanley, P. 1989. Chinese hamster ovary cell mutants with multiple glycosylation defects for production of glycoproteins with minimal carbohydrate heterogeneity. Mol. Cell Biol. 9:377-383.
Miloux, B. and Lupker, J.H. 1994. Rapid isolation of highly productive recombinant Chinese hamster ovary cell lines. Gene 149:341-344.
Stanley, P. and Ioffe, E. 1995. Glycosyltransferase mutants: Key to new insights in glycobiology. FASEB J. 9:1436-1444.
Morris, A.E., Lee, C-C., Hodges, K., Aldrich, T.L., Krantz, C., Smidt, P.S., and Thomas, J.N. 1997. Expression augmenting sequence element (EASE) isolated from Chinese hamster ovary cells. In Animal Cell Technology (M.J.T. Carrondo, ed.) pp. 529-534. Kluwer Academic Publishers, Boston.
Subramanian, S. and Srienc, F. 1996. Quantitative analysis of transient gene expression in mammalian cells using green fluorescent protein. J. Biotechnol. 49:137-151.
Mortensen, R., Chesnut, J.D., Hoeffler, J.P., and Kingston, R.E. 1997. Selection of transfected mammalian cells. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D.Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 16.5.1-16.5.19. John Wiley & Sons, New York. Mory, Y., Ben-Barak, J., Segev, D., Cohen, D., Novick, D., Fischer, D.G., Rubinstein, M., Kargman, S., Zilberstein, M., and Revel, M. 1986. Efficient constitutive production of human IFN-γ in Chinese hamster ovary cells. DNA 5:181-193. Moss, B. and Earl, P.L. 1998. Overview of the vaccinia virus expression system. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 16.15.11615.5. John Wiley & Sons, New York. Page, M.J. and Sydenham, M.A. 1991. High level expression of the humanized monoclonal antibody Campath-1H in Chinese hamster ovary cells. Bio/Technology 9:64-68. Parekh, R.B. and Patel, T.P. 1992. Comparing the glycosylation patterns of recombinant glycoproteins. Trends Biotechnol. 10(8):276-280. Pendse, G.J., Karkare, S., and Bailey, J.E. 1992. Effect of cloned gene dosage on cell growth and Hepatitis B surface antigen synthesis and secretion in recombinant CHO cells. Biotechnol. Bioeng. 40:119-129. Robinson, J.P., Darzynkiewicz, Z., Dean, P.N., Orfao, A., Rabinovitch, P.S., Stewart, C.C., Tanke, H.J., and Wheeless, L.L. (eds.) 1998. Current Protocols in Cytometry. John Wiley & Sons, New York. Sambrook, J., Fritsch, E.F., and Maniatis, T. 1989. Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Simonsen, C. and Levinson, A. 1983. Isolation and expression of an altered mouse dihydrofolate reductase cDNA. Proc. Natl. Acad. Sci. U.S.A. 80:2495-2499.
Tiege, M., Weidemann, and R., Kretzmer, G. 1994. Problems with serum-free production of antithrombin III regarding proteolytic activity and product quality. J. Biotechnol. 34:101-105. Wurm, F.M., Pallavicini, M.G., and Arathoon, R. 1992. Integration and stability of CHO amplicons containing plasmid sequences. Dev. Biol. Stand. 76:69-82. Wurm, F.M., Johnson, A., Ryll, T., Kohne, C., Scherthan, H., Glaab, F., Lie, Y.S., Petropoulos, C.J., and Arathoon, W.R. 1996. Gene transfer and amplification in CHO cells: Efficient methods for maximizing specific productivity and assessment of genetic consequences. Ann. N.Y. Acad. Sci. 782:70-78. Yenofsky, R.L., Fine, M., and Pellow, J.W. 1990. A mutant phosphotransferase II gene reduces the resistance of transformants to antibiotic selection pressure. Proc. Natl. Acad. Sci. U.S.A. 87:3435-3439. Yim, K.W. 1991. Fractionation of human recombinant tissue plasminogen activator (rtPA) glycoforms by high-performance capillary zone electrophoresis and capillary isoelectric focusing. J. Chromatogr. 559:401-410. Zettlmeissel, G., Ragg, H., and Karges, H.E. 1987. Expression of biologically active Antithrombin III in Chinese hamster ovary cells. Bio/Technology 5:720-725.
Key References Butler, M. (ed.). 1991. Mammalian Cell Biotechnology, IRL Press, Oxford. Excellent introductory background with protocol and tips. Freshney R.I. (ed.). 1992. Animal Cell Culture. (2nd ed.), IRL Press, Oxford. Practical guide to cell culture with useful protocols and tips. Kriegler, 1990. See above Comprehensive manual for molecular biology.
Contributed by Su Chen, David Gray, Johnny Ma and Shyamsundar Subramanian Chiron Corporation Emeryville, California Production of Recombinant Proteins
5.10.41 Current Protocols in Protein Science
Supplement 12
Overview of the Vaccinia Virus Expression System Vaccinia virus was introduced in 1982 as a vector for transient expression of genes in mammalian cells (Mackett et al., 1982; Panicali and Paoletti, 1982). This expression system differs from others in that transcription occurs in the cytoplasm of the cell rather than in the nucleus. As a vector, vaccinia virus has a number of useful characteristics, including a capacity that permits cloning large fragments of foreign DNA (>20 kbp) with retention of infectivity, a wide host range, a relatively high level of protein synthesis, and “appropriate” transport, secretion, processing, and posttranslational modifications as dictated by the primary structure of the expressed protein and the cell type used. For example, N- and O-glycosylation, phosphorylation, myristylation, and cleavage, as well as assembly of expressed proteins, occur in an apparently faithful manner. Laboratory applications of vaccinia virus vectors include production of biologically active proteins in tissue culture, analysis of mutant forms of proteins, and determination of transport and processing signals. In addition, recombinant vaccinia viruses have been important for immunological studies (Bennink and Yewdell, 1990). Infected cells can serve as targets to analyze the antigenic specificity of cytotoxic T cells. Recombinant viruses can be used to infect animals in order to determine cell-mediated and humoral responses to specific proteins, and are being evaluated as candidate vaccines for human and veterinary uses. Several variations of the vaccinia vector system have been developed and either standard or more attenuated, host-restricted strains may be used. After obtaining the virus stock (UNIT 5.12), the gene of interest is placed under control of a vaccinia virus promoter and integrated into the genome of vaccinia so as to retain infectivity (UNIT 5.13). Alternatively, expression can be achieved by transfecting a plasmid containing the vaccinia promoter–controlled gene into a cell that has been infected with vaccinia virus. These recombinant viruses are then characterized using various methods (UNIT 5.14). In still another variation, the bacteriophage T7 RNA polymerase gene can be integrated into the genome of vaccinia so that a gene controlled by a T7 promoter, either in a
UNIT 5.11
transfected plasmid or a recombinant vaccinia virus, will be expressed.
VACCINIA REPLICATION CYCLE Vaccinia is the prototypic member of the Orthopoxvirus genus of the Poxviridae family. Poxviruses differ from other eukaryotic DNA viruses in that they replicate in the cytoplasm rather than in the nucleus. Vaccinia virus has a linear, double-stranded DNA genome of nearly 200,000 bp that encodes most of the proteins needed for replication and transcription in the cytoplasm. The replication cycle of poxviruses is represented in Figure 5.11.1. The virus particle or virion consists of a complex core structure surrounded by a lipoprotein envelope. Remarkably, all proteins necessary for transcription of the early class of genes are packaged with the genome in the core. These include the following virus-encoded proteins: a multisubunit, DNA-dependent RNA polymerase, an early transcription factor, capping and methylating enzymes, and a poly(A) polymerase. The transcription system is activated upon infection, and early mRNAs and proteins can be detected within the first hour. The early mRNAs closely resemble their eukaryotic counterparts—they are capped, methylated, polyadenylated, and of discrete size. Termination of transcription occurs ∼50 bases after the sequence TTTTTNT (where N can be any nucleotide; the termination signal is actually recognized in the nascent RNA as UUUUUNU). There is no evidence for splicing or other kinds of processing involving RNA cleavage. DNA replication begins within a few hours postinfection and leads successively to the intermediate and late phases of gene expression. The promoters associated with intermediate and late genes differ in sequence from early promoters and have different transcription factor requirements. The intermediate factors are synthesized early in order to facilitate transcription of newly replicated viral DNA. Some of the late factors are products of intermediate genes and hence are made and used after viral DNA replication. The mRNAs transcribed from intermediate and late genes differ from typical early mRNAs Production of Recombinant Proteins
Contributed by Bernard Moss and Patricia L. Earl Current Protocols in Protein Science (1998) 5.11.1-5.11.5 Copyright © 1998 by John Wiley & Sons, Inc.
5.11.1 Supplement 13
uncoating
intermediate mRNA late transactivators
e tor as tiva er c sa early mRNA lym ran t po te A dia DN rme e int early enzymes
er resoluti
envelope RNA polymerase transcription factor poly(A) polymerase capping enzyme methylating enzymes
DNA replication (2-12 hr)
on
core
growth factor
nucleus
mature virion
morphogenesis (4-20 hr)
concatem
DNA
late mRNA late enzymes early transcription factor structural proteins cleavage glycosylation acylation phosphorylation
cell Figure 5.11.1 Replication cycle of vaccinia virus. After entry of vaccinia virus into the cells, early genes are expressed, leading to secretion of several proteins (including a growth factor), uncoating of the virus core, and the synthesis of DNA polymerase (and other replication proteins), RNA polymerase subunits, and transcriptional transactivators of the intermediate class of genes. After DNA replication, intermediate mRNAs are made, some of which encode late transcriptional transactivators. This in turn leads to expression of late genes, which encode structural proteins, enzymes, and early transcription factors that are packaged in the assembling virus particles. Some of the mature virions are wrapped in Golgi-derived membranes and are released from the cell. The bold arrows indicate products that exit the cell. Reprinted with permission from Raven Press (Moss, 1996a).
Overview of the Vaccinia Virus Expression System
as follows. First, the termination signal UUUUUNU is not recognized; consequently the mRNAs are long and heterogeneous in length, making procedures like northern blotting virtually useless. Second, the 5′ ends of intermediate and late mRNAs contain a capped poly(A) leader of ∼35 nucleotides that is probably the result of an RNA polymerase slippage mechanism within the conserved AAA at the initiation site. Many early proteins are not synthesized beyond ∼6 hr postinfection (unless an inhibitor of DNA replication such as cytosine arabinoside is added) because of the cessation of early gene transcription and the relatively short half-life of all mRNAs at late times. Some genes, however, have tandem early and late promoters so that they are expressed throughout the growth cycle. Intermediate genes are expressed for a relatively short period after DNA replication. Either because of intrinsic promoter strength, DNA copy number, or prolonged expression (>20 hr), much more protein is made from the
strongest late promoters than from early or intermediate promoters.
EFFECTS OF VACCINIA INFECTION Standard vaccinia virus strains can productively infect most mammalian and avian cell lines, with a few exceptions such as Chinese hamster ovary (CHO) cells, but may not complete the replication cycle in primary lymphocytes or macrophages. The host-restricted modified vaccinia virus Ankara (MVA) strain has been shown to replicate efficiently only in chick embryo fibroblasts and BHK-21 cells, but viral and recombinant protein expression occurs in all cell lines infectable by standard strains. Vaccinia infection generally results in rapid inhibition of host nucleic acid and protein synthesis. Inhibition of host protein synthesis is dramatic and probably results from several factors whose identities are uncertain; the relative contribution of each factor may depend on the virus multiplicity, cell type, and time of
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Current Protocols in Protein Science
analysis. At the time of maximal late gene expression, host protein synthesis has been largely suppressed, facilitating the identification of viral or recombinant proteins by pulselabeling with radioactive amino acids (UNIT 3.3). In fibroblasts, the initial cytopathic effect is cell rounding, and is obvious by several hours postinfection. Nevertheless, the majority of cells remain intact for ≥48 hr. Some virus strains may cause less cytopathic effects than others, but mutants that efficiently express recombinant proteins but do not inhibit host gene expression have not been identified. Approximately 100 to 200 plaque-forming units (pfu), equivalent to ∼2500 to 5000 particles, are made per cell within a 20- to 40-hr period. With the commonly used vaccinia virus Western Reserve (WR) strain, >95% of the infectious virus remains cell-associated. With some other vaccinia virus strains, notably IHD-J, larger amounts of extracellular virus are produced.
VACCINIA VECTOR EXPRESSION SYSTEM Genes or cDNAs containing open reading frames derived from prokaryotic, eukaryotic, or viral sources have been expressed using vaccinia virus vectors. The gene of interest is usually placed next to a vaccinia promoter and this expression cassette is then inserted into the virus genome by homologous recombination or direct ligation (UNIT 5.13). Use of poxvirus promoters is essential because cellular and other viral promoters are not recognized by the vaccinia transcriptional apparatus. Strong late promoters are preferable when high levels of expression are desired. An early promoter, however, may be of use if it is desirable to express proteins prior to the occurrence of major cytopathic effects, or when the purpose is to make cells that express antigens in association with major histocompatibility class I molecules so they form cytotoxic T cell targets or prime animals for a cytotoxic T cell response. (The ability of vaccinia viral vectors to direct this type of antigen presentation seems to diminish late in infection.) The most versatile and widely used promoters contain early and late promoter elements. Transcripts originating early will terminate after the sequence TTTTTNT; thus, any cryptic TTTTTNT termination motifs within the coding sequence of the gene should be altered by mutagenesis if an early poxvirus promoter is used (Earl et al., 1990). To mimic vaccinia virus mRNAs, untranslated leader and 3′-terminal sequences are usually kept short.
A number of plasmids have been designed with restriction endonuclease sites for insertion of foreign genes downstream of vaccinia promoters (UNIT 5.13). The expression cassette is flanked by vaccinia DNA to permit homologous recombination when the plasmid is transfected into cells that have previously been infected with a vaccinia virus. The flanking vaccinia virus DNA is chosen so that recombination will not interrupt an essential viral gene. Without selection, the ratio of recombinant to parental vaccinia virus is usually ∼1:1000. Although this frequency is high enough to permit the use of plaque hybridization or immunoscreening to pick recombinant viruses, a variety of methods have been employed to facilitate identification of recombinant viruses. Some widely used selection or screening techniques are described in UNIT 5.13. Commonly, the expression cassette is flanked by segments of the vaccinia thymidine kinase (TK) gene so that recombination results in inactivation of TK. Virus with a TK− phenotype can then be distinguished from those with a TK+ phenotype by infecting a TK− cell line in the presence of 5-bromodeoxyuridine (BrdU), which must be phosphorylated by TK to be lethally incorporated into the virus genome. Alternatively, recombinant viruses can be selected by the coexpression of a bacterial antibiotic resistance gene such as guanine phosphoribosyltransferase (gpt). Co-expression of the Escherichia coli lacZ gene or other color markers allows rapid screening of recombinant virus plaques. Complementation of host-range and plaqueforming defects are also useful for isolation of recombinant viruses (UNIT 5.13).
STEPS FOR EXPRESSION OF GENES USING VACCINIA VECTORS The expression of genes using the vaccinia expression system is presented in detail in UNITS 5.12-5.15 and is outlined in the flowchart in Figure 5.11.2. A brief overview is presented below. 1. Prepare a stock of standard, host-rangerestricted, or bacteriophage T7 RNA polymerase–expressing vaccinia virus. At the same time, subclone the gene of interest into a plasmid transfer vector (UNITS 5.12 & 5.13). 2. Infect cells with vaccinia virus and transfect with the recombinant plasmid (UNIT 5.13). 3. Lyse the cells and plaque the virus under suitable selection or screening conditions (UNIT 5.13).
Production of Recombinant Proteins
5.11.3 Current Protocols in Protein Science
Supplement 13
culture cell lines (UNIT 5.12, Basic Protocols 1 & 2)
infect cell line and prepare wild-type vaccinia stock (UNIT 5.12, Basic Protocol 3)
purify vaccinia virus (UNIT 5.13, Support Protocol 1)
isolate vaccinia virus DNA (UNIT 5.13, Support Protocol 2)
titer vaccinia virus stock using plaque assay (UNIT 5.12, Support Protocol 1)
infect cells with vaccinia virus and transfect with vector (UNIT 5.13, Basic Protocol 1)
amplify a plaque (UNIT 5.13, B asic Protocol 3)
titer the amplified plaque (UNIT 5.12, Support Protocol 1)
prepare recombinant virus stock (UNIT 5.12, Basic Protocol 3)
express gene and analyze protein by • polyacrylamide gel electrophoresis • immunoblotting • immunoprecipitation
insert gene into transfer vector (UNIT 5.13, Basic Protocol 1)
select and screen recombinant virus plaques (UNIT 5.13, Basic Protocol 2)
analyze recombinant virus by • polymerase chain reaction (UNIT 5.14, Basic Protocol 1) • Southern blot hybridization (UNIT 5.14, Basic Protocol 2) • DNA dot-blot hybridization (UNIT 5.14, Basic Protocol 3) • immunoblotting (UNIT 5.14, Alternate Protocol) • immunostaining (UNIT 5.12 )
Figure 5.11.2 Flow chart showing protocols for gene expression using the recombinant vaccinia virus system.
Overview of the Vaccinia Virus Expression System
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4. Pick plaques and confirm the presence and/or expression of the foreign gene (UNIT 5.14). 5. Amplify the plaque and prepare recombinant virus stock (UNIT 5.13). 6. Infect cells and analyze proteins synthesized (UNIT 5.14).
SAFETY PRECAUTIONS FOR USING VACCINIA Vaccinia virus is not to be confused either with variola virus, another member of the Orthopoxvirus genus that caused smallpox prior to its eradication, or with varicella virus, a herpes virus that causes chicken pox. Until 1972, vaccinia virus was routinely used in the United States as a live vaccine to prevent smallpox, and a residual scar, commonly on the upper arm, is evidence of that vaccination. To prevent laboratory infections, the Centers for Disease Control (CDC) and the National Institutes of Health (NIH) recommend that individuals who come into contact with vaccinia virus receive vaccinations at 10-year intervals (Richmond and McKinney, 1993). The CDC has supplied vaccine for such purposes when requested by qualified health workers. Eczema or an immunodeficiency disorder in the laboratory worker or a close contact, however, may be a contraindication to vaccination, which should only be given under medical supervision. The benefits of routine vaccination for healthy investigators have also been questioned (Baxby, 1989; Wenzel and Nettelman, 1989). Vaccinia virus is very stable and parenteral inoculation, ingestion, and droplet or aerosol exposure of mucous membranes are the primary hazards to laboratory or animal care personnel. Standard biosafety level 2 (BL-2) practices and class I or II biological safety cabinets should be employed (Richmond and McKinney, 1993). The NIH intramural program has lowered the safety requirements for highly attenuated vaccinia virus strains such as modified vaccinia virus Ankara (MVA; UNIT 5.12) and NYVAC (Tartaglia et al., 1992). However, local institutional biosafety offices should be contacted to determine current policy regarding vaccination and physical containment. Additional precautions may be necessary for expression of certain genes such as toxins or large segments of other viral genomes, and guidelines for recombinant DNA work should be consulted. Approval of local biosafety committees may be necessary.
LITERATURE CITED Bennink, J.R. and Yewdell, J.W. 1990. Recombinant vaccinia viruses as vectors for studying T lymphocyte specificity and function. Curr. Topics Microbiol. Immunol. 163:153-184. Baxby, D. 1989. Smallpox vaccination for investigators. Lancet 2:919. Earl, P.L., Hügin, A.W., and Moss, B. 1990. Removal of cryptic poxvirus transcription termination signals from the human immunodeficiency virus type 1 envelope gene enhances expression and immunogenicity of a recombinant vaccinia virus. J. Virol. 64:2448-2451. Mackett, M., Smith, G.L., and Moss, B. 1982. Vaccinia virus: A selectable eukaryotic cloning and expression vector. Proc. Natl. Acad. Sci. U.S.A. 79:7415-7419. Moss, B. 1996a. Poxviridae: The viruses and their replication. In Virology (B.N. Fields, D.M. Knipe, and P. M. Howley, eds.) pp. 2637-2671. Raven Press, New York. Panicali, D. and Paoletti, E. 1982. Construction of poxviruses as cloning vectors: Insertion of the thymidine kinase gene from herpes simplex virus into the DNA of infectious vaccinia virus. Proc. Natl. Acad. Sci. U.S.A. 79:4927-4931. Richmond, J.Y. and McKinney, R.W. 1993. Biosafety in Microbiological and Biomedical Laboratories, 3rd ed. U.S. Government Printing Office, Washington, D.C. Tartaglia, J., Perkus, M.E., Taylor, J., Norton, E.K., Audonnet, J.C., Cox, W.I., Davis, S.W., Vanderhoeven, J., Meignier, B., Riviere, M., Languet, B., and Paoletti, E. 1992. NYVAC–A highly attenuated strain of vaccinia virus. Virology 188:217-232. Wenzel, R.P. and Nettelman, M.D. 1989. Smallpox vaccination for investigators using vaccinia recombinants. Lancet 2:630-631.
KEY REFERENCES Johnson, G.P., Goebel, S.J., and Paoletti, E. 1993. An update on the vaccinia virus genome. Virology 196: 381-401. Moss, B. 1996b. Genetically engineered poxviruses for recombinant gene expression, vaccination, and safety. Proc. Natl. Acad. Sci. U.S.A. 93:11341-11348.
Contributed by Bernard Moss and Patricia L. Earl National Institute of Allergy and Infectious Diseases Bethesda, Maryland
Production of Recombinant Proteins
5.11.5 Current Protocols in Protein Science
Supplement 13
Preparation of Cell Cultures and Vaccinia Virus Stocks
UNIT 5.12
This unit describes the maintenance of cell lines used with vaccinia virus, both in monolayer cultures (see Basic Protocol 1) and in suspension (see Basic Protocol 2). The suspended cell culture is then used in the preparation of vaccinia virus stocks (see Basic Protocol 3). The preparation of chick embryo fibroblasts (CEF) is also presented (see Basic Protocol 4), for use in the production of the highly attenuated and host range–restricted modified vaccinia virus Ankara (MVA) strain of vaccinia virus (see Basic Protocol 5). Additionally, support protocols are presented for the titration of standard and MVA vaccinia virus stocks (see Support Protocols 1 and 2, respectively). Because standard vaccinia virus strains have a broad host range, there is considerable latitude in the selection of cell lines; those described below (see Basic Protocols 1 and 2) have been found to give good results. BS-C-1 cells give the best results for a plaque assay, whereas HeLa cells are preferred for preparation of virus stocks. CV-1 cells can be used for both procedures, but they are generally used for transfection (UNIT 5.13). Human thymidine kinase–negative (TK−) 143B cells are used when TK selection is employed (UNIT 5.13), but they can be used for transfection as well as for a plaque assay. Either CEF or BHK-21 cells are used to propagate MVA and prepare recombinant MVA. Table 5.12.1 presents a summary of the uses for specific cell lines. NOTE: Carry out all procedures in this unit using sterile technique, preferably in a biosafety cabinet.
Table 5.12.1
Cell Lines Used in Specific Vaccinia Protocols
Cell line
Usea
Procedure
HeLa S3
Virus stock preparation Virus purification Plaque amplification
UNIT 5.12 Basic
Plaque assay Transfection (optional) XGPRT selection Plaque amplification (optional)
UNIT 5.12
CV-1
Transfection Virus stock preparation (optional) Plaque assay (optional)
UNIT 5.13 Basic
HuTK− 143B
TK selection Plaque assay (optional) Transfection (optional)
UNIT 5.13 Basic
MVA procedures
UNIT 5.12 Basic Protocol 5, Support Protocol 2; UNIT 5.13 Basic Protocol
4
UNIT 5.12 Basic Protocol 5, Support Protocol 2; UNIT 5.13 Basic Protocol
4
BS-C-1
CEF BHK-21
MVA procedures
Protocol 3 Support Protocol 1 UNIT 5.13 Basic Protocol 3 UNIT 5.13
Support Protocol 1 Protocol 1 UNIT 5.13 Basic Protocol 2 UNIT 5.13 Basic Protocol 3 UNIT 5.13 Basic
Protocol 1 UNIT 5.12 Basic Protocol 3 UNIT 5.12 Support Protocol 1 Protocol 2 Support Protocol 1 UNIT 5.13 Basic Protocol 1 UNIT 5.12
aThe preferred use(s) for each cell line is listed first; if optional is indicated, the cell line can be used for the indicated
procedure, but the results may not be as good as those from the preferred cell line.
Production of Recombinant Proteins Contributed by Patricia L. Earl, Norman Cooper, Linda S. Wyatt, Bernard Moss, and Miles W. Carroll
5.12.1
Current Protocols in Protein Science (1998) 5.12.1-5.12.12 Copyright © 1998 by John Wiley & Sons, Inc.
Supplement 13
BASIC PROTOCOL 1
CULTURE OF MONOLAYER CELLS Frozen cells are thawed and grown in appropriate complete medium containing twice the maintenance amount of serum (see below). When the cells are confluent, they are treated with trypsin/EDTA, diluted, and maintained in appropriate complete medium containing 10% FBS (see Table 5.12.2). Table 5.12.2
Media Used for Growth and Maintenance of Cell Linesa
Cell line
Maintenance medium
Start-up mediuma
BHK-21 BS-C-1 CEF CV-1 HeLa S3 HuTK− 143B
Complete MEM-10 Complete MEM-10 Complete MEM-10 Complete DMEM-10 Complete spinner medium-5 Complete MEM-10/BrdU
Complete MEM-20 Complete MEM-20 Complete MEM-10 Complete DMEM-20 Complete MEM-10 Complete MEM-20/BrdU
aSee Reagents and Solutions for recipes.
Materials Frozen ampule of cells (Table 5.12.2): BS-C-1 (ATCC #CCL26), CV-1 (ATCC #CCL70), HuTK− 143B (ATCC #CRL8303), or BHK-21 (ATCC #CCL10) cells 70% ethanol Start-up medium (Table 5.12.2): complete MEM-20, complete MEM-20/BrdU (see recipes), or complete DMEM-20 (APPENDIX 3C) 37°C Maintenance medium (Table 5.12.2): complete MEM-10, complete MEM-10/BrdU (see recipes), or complete DMEM-10 (APPENDIX 3C) 37°C PBS (optional; APPENDIX 2E) Trypsin/EDTA: 0.25% (w/v) trypsin/0.02% (w/v) EDTA, 37°C 25-cm2 and 150-cm2 tissue culture flasks Humidified 37°C, 5% CO2 incubator Begin the culture 1. Thaw a frozen ampule of cells in a 37°C water bath. 2. Sterilize the ampule tip with 70% ethanol, break the neck, and transfer the cells with a pipet into a 25-cm2 tissue culture flask containing 5 ml start-up medium. Rotate the flask to evenly distribute the cells and place overnight in a humidified 5% CO2 incubator at 37°C. 3. Aspirate the start-up medium and replace with appropriate maintenance medium. Return cells to the CO2 incubator at 37°C and check daily for confluency. Cells should be passaged when they become confluent. Generally, if cells are split 1:20, they reach confluence in 1 week and need not be counted.
Maintain the culture 4. When the cells are a confluent monolayer, aspirate medium. 5. Wash cells once with PBS or trypsin/EDTA to remove remaining serum from the cells by covering cells with the solution and pipetting it off. Preparation of Cell Cultures and Vaccinia Virus Stocks
6. Overlay cells with 37°C trypsin/EDTA using a volume that is just enough to cover the monolayer (e.g., 0.3 ml for a 25-cm2 flask). Allow to sit 30 to 40 sec (cells should become detached) and shake the flask to completely detach cells.
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7. Add 1.4 ml appropriate maintenance medium. Pipet the cell suspension up and down several times to disrupt clumps (these cells are ready for passage). 8. Remove 0.5 ml cell suspension and add it to a new 150-cm2 tissue culture flask containing 30 ml maintenance medium. Rotate the flask to evenly distribute the cells and place in a CO2 incubator at 37°C until the cells are confluent (∼1 week). Maintain the cells by splitting ∼1:20 in maintenance medium at approximately weekly intervals. Cells can be maintained in smaller flasks if desired. If so, volumes should be adjusted proportionately.
CULTURE OF CELLS IN SUSPENSION HeLa S3 cells are maintained in complete spinner medium-5.
BASIC PROTOCOL 2
Materials Frozen ampule of HeLa S3 cells (ATCC #CCL2.2) 70% ethanol Complete MEM-10 (see recipe), 37°C Trypsin/EDTA: 0.25% (w/v) trypsin/0.02% (w/v) EDTA, 37°C Complete spinner medium-5 (see recipe), 37°C 25-cm2 tissue culture flask Humidified 37°C, 5% CO2 incubator Sorvall H-6000A rotor (or equivalent) 100- or 200-ml vented spinner bottles and caps with filters (Bellco) Additional reagents and equipment for counting cells with a hemacytometer (APPENDIX 3C) Begin the culture 1. Thaw a frozen ampule of HeLa S3 cells in a 37°C water bath. 2. Sterilize the ampule tip with 70% ethanol, break the neck, and transfer the cells with a pipet to a 25-cm2 tissue culture flask containing 5 ml complete MEM-10. Rotate the flask to evenly distribute the cells and place overnight in a humidified 5% CO2 incubator at 37°C. 3. Aspirate medium. Overlay cells with 0.5 ml of 37°C trypsin/EDTA and let sit 30 to 40 sec. Since these cells do not attach firmly to the flask, they should not be washed prior to trypsinization.
4. Add 10 ml complete spinner medium-5 and transfer cells to a 50-ml centrifuge tube. Centrifuge 5 min in a Sorvall H-6000A rotor at 2500 rpm (1800 × g), room temperature, and discard supernatant. 5. Suspend cell pellet in 5 ml complete spinner medium-5 by pipetting up and down to disrupt clumps. 6. Add 50 ml complete spinner medium-5 to a 100- or 200-ml vented spinner bottle and transfer the cell suspension to this bottle. 7. Remove 1 ml cell suspension and count the cells using a hemacytometer (APPEN5 DIX 3C). Add complete spinner medium-5 to adjust the cell density to 3–4 × 10 cells/ml. Place cells in a 37°C incubator without CO2 and stir continuously. The initial high density is used because some cells are not viable.
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8. Grow cells for two successive days, counting cells daily and adding complete spinner medium-5 as necessary to maintain a concentration of 3–4 × 105 cells/ml. Maintain the culture 9. Remove 1 ml cell suspension and monitor the cells using a hemacytometer. 10. When the density is 4–5 × 105 cells/ml, dilute the cells to 1.5 or 2.5 × 105 cells/ml with complete spinner medium-5 for alternate day or daily feeding, respectively. 11. Place a 100- or 200-ml vented spinner bottle containing 50 ml or 100 ml cells, respectively, in 37°C incubator without CO2 and stir continuously. Passage every 1 to 2 days. HeLa S3 cells are grown and maintained in complete spinner medium-5 in vented spinner bottles at 37°C without CO2. Cells are diluted with fresh medium at 1- to 2-day intervals to keep the cell density between 1.5 × 105 and 5 × 105 cells/ml. BASIC PROTOCOL 3
PREPARATION OF A VACCINIA VIRUS STOCK To prepare a vaccinia virus stock, HeLa S3 cells from a spinner culture (see Basic Protocol 2) are plated the day before infection and allowed to attach. They are then infected with trypsinized virus. After several days, the infected cells are harvested and lysed during repeated freeze-thaw cycles. The virus stock is then aliquoted and stored at −70°C. To titer this stock, see Support Protocol 1. The protocol can be modified for monolayer cultures. Materials HeLa S3 cells from suspension culture (see Basic Protocol 2) Complete MEM-10 and -2.5 (see recipe), 37°C Vaccinia virus (ATCC #VR1354 or equivalent) 0.25 mg/ml trypsin (2× crystallized and salt-free; Worthington; filter sterilize and store at −20°C) Sorvall H-6000A rotor (or equivalent) 150-cm2 tissue culture flask Humidified 37°C, 5% CO2 incubator Additional reagents and equipment for counting cells with a hemacytometer (APPENDIX 3C) Prepare cells 1. Count HeLa S3 cells from a suspension culture using a hemacytometer (APPENDIX 3C). 2. Centrifuge 5 × 107 cells 5 min in a Sorvall H-6000A rotor at 2500 rpm (1800 × g), room temperature, and discard supernatant. 3. Resuspend cells in 25 ml of 37°C complete MEM-10, dispense in a 150-cm2 tissue culture flask, and place overnight in a humidified, 5% CO2 incubator at 37°C. Increase the number of HeLa cells proportionately if more than one 150-cm2 flask is to be infected. As an alternative to HeLa suspension cells, prepare 150-cm2 flasks of monolayer cells as described (see Basic Protocol 1) and continue with step 4 below.
Preparation of Cell Cultures and Vaccinia Virus Stocks
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Trypsinize virus 4. Just prior to use, mix an equal volume of vaccinia virus stock and 0.25 mg/ml trypsin, and vortex vigorously. Incubate 30 min in a 37°C water bath, vortexing at 5- to 10-min intervals throughout the incubation. Virus stocks are usually at a titer of ∼2 × 109 pfu/ml, but may be significantly lower depending on the source. Vortexing usually breaks up any clumps of cells. However, if there are still visible clumps, chill to 0°C and sonicate 30 sec on ice (UNIT 5.13). Sonication can be repeated several times but the sample should be allowed to cool on ice between sonications.
Infect cells 5. Dilute trypsinized virus in complete MEM-2.5 at 2.5–7.5 × 107 pfu/ml. Decant or aspirate medium from the 150-cm2 flask of cells and add 2 ml diluted, trypsinized virus. Place 2 hr in a CO2 incubator at 37°C, rocking flask by hand at 30-min intervals. The optimal multiplicity of infection (MOI) is 1 to 3 pfu/cell. Multiplicities of 0.1 pfu/cell may be necessary if the titer of the initial virus stock is low. The trypsinized virus must be diluted ≥10-fold to avoid detaching the cells.
6. Overlay cells with 25 ml complete MEM-2.5 and place 3 days in a CO2 incubator at 37°C. 7. Detach the infected cells from the flask by shaking and pour or pipet into a sterile plastic screw-cap tube. Centrifuge 5 min at 1800 × g, 5° to 10°C, and discard supernatant. 8. Resuspend cells in 2 ml complete MEM-2.5 (per initial 150-cm2 flask) by gently pipetting or vortexing. Harvest virus stock 9. Lyse the cell suspension by freeze-thaw cycling as follows: freeze in dry ice/ethanol, thaw in a 37°C water bath, and vortex. Carry out the freeze-thaw cycling a total of three times. 10. Keep the virus stock on ice and divide it into 0.5- to 2-ml aliquots. Store the aliquots indefinitely at −70°C. TITRATION OF VACCINIA VIRUS STOCKS BY PLAQUE ASSAY Serial dilutions of the trypsinized virus stock (see Basic Protocol 3) are used to infect the appropriate cell line. After several days growth, the medium is removed and the cells are stained with crystal violet. Plaques appear as 1- to 2-mm-diameter areas of diminished staining due to the retraction, rounding, and detachment of infected cells.
SUPPORT PROTOCOL 1
Additional Materials (also see Basic Protocols 1 and 3) BS-C-1 cells from confluent monolayer culture (see Basic Protocol 1) Virus stock (see Basic Protocol 3) 0.1% (w/v) crystal violet (Sigma) in 20% ethanol (store indefinitely at room temperature) 6-well, 35-mm2 tissue culture dishes Additional reagents and equipment for counting cells with a hemacytometer (APPENDIX 3C) Production of Recombinant Proteins
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Prepare cell and virus stocks 1. Trypsinize confluent monolayer of BS-C-1 cells as described (see Basic Protocol 1, steps 4 to 7). 2. Count the cells using a hemacytometer (APPENDIX 3C). 3. Seed wells of a 6-well, 35-mm2 tissue culture dish with 5 × 105 cells per well BS-C-1 cells in 2 ml complete MEM-10. Place overnight in a humidified 5% CO2 incubator at 37°C to reach confluency. 4. Trypsinize virus stock (see Basic Protocol 3, step 4). 5. Make nine 10-fold serial dilutions of the trypsinized virus in complete MEM-2.5, using a fresh pipet for each dilution. Perform assay 6. Remove medium from BS-C-1 cells and infect cells in duplicate wells with 0.5 ml of the 10−7, 10−8, and 10−9 trypsinized virus dilutions. Place 1 to 2 hr in a CO2 incubator at 37°C, rocking dish at 15- to 30-min intervals to spread virus uniformly and keep cells moist. 7. Overlay cells in each well with 2 ml complete MEM-2.5 and place 2 days in a CO2 incubator at 37°C. 8. Remove medium and add 0.5 ml of 0.1% crystal violet to each well. Incubate 5 min at room temperature. 9. Aspirate crystal violet and allow wells to dry. 10. Determine the titer by counting plaques within the wells and multiplying by the dilution factor. Most accurate results are obtained from wells with 20 to 80 plaques. In determining the titer, take into account the 1:1 dilution of the virus stock with trypsin. BASIC PROTOCOL 4
PREPARATION OF CHICKEN EMBRYO FIBROBLASTS The attenuated, replication-deficient modified vaccinia virus Ankara (MVA) can be used as an alternative to standard strains of vaccinia virus, for use as a vector to express foreign genes. MVA was derived from the vaccinia virus strain Ankara by multiple (>570) passages in chick embryo fibroblasts (CEF). Although MVA expresses recombinant proteins efficiently, the assembly of infectious particles is interrupted in human and most other mammalian cells lines, providing an added degree of safety to laboratory personnel. The NIH Intramural Biosafety Committee has determined that individuals do not need to be vaccinated to work with MVA, and can do so under biosafety level 1 (BL-1) conditions if no other vaccinia virus strains are being manipulated at the same location by other investigators. Recombinant MVA expressing bacteriophage T7 RNA polymerase, which can be used for high level transient expression, has also been constructed (UNIT 5.15) CEF or Syrian hamster kidney cells (BHK-21; see Basic Protocol 1) are permissive for growth of MVA. MVA is titered by immunostaining (see Support Protocol 2), because it does not form distinct plaques for accurate quantitation.
Preparation of Cell Cultures and Vaccinia Virus Stocks
In this protocol, nine-day-old chicken embryos are trypsinized and plated in appropriate medium. After the cells form a confluent monolayer, they are transferred directly to a 31°C incubator where they can be held for 2 to 3 weeks before making secondary CEF for virus infection. With each passage, CEF require longer to become confluent; therefore use of the second passage is recommended.
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Materials Nine-day-old embryonated eggs (Specific Pathogen Free Eggs, SPAFAS) 70% ethanol MEM with no additives, 37°C Trypsin/EDTA: 0.25% (w/v) trypsin/0.02% (w/v) EDTA, 37°C Complete MEM-10 (Table 5.12.2; see recipe), 37°C Sterile dissecting scissors and forceps 100-cm2 sterile petri dishes 10-ml syringes Sterile trypsinization flask with magnetic stir bar Humidified 37° and 31°C, 5% CO2 incubators 500-ml beakers with two layers of gauze taped over tops Sorvall RC-3B centrifuge and 250-ml centrifuge bottles (or equivalent) 150-cm2 tissue culture flasks Prepare tissue 1. Position ten 9-day-old embryonated eggs with air space (blunt end) up and spray with 70% ethanol. 2. Crack the top of an egg with sterile dissecting scissors, and cut off shell to just above the membrane while keeping the latter intact. Remove membrane with sterile forceps. Repeat with remaining eggs. NOTE: Healthy eggs have well-formed blood vessels.
3. Remove the embryo from each egg and combine in a 100-cm2 sterile petri dish. 4. Remove head and feet from each embryo and place the rest of the body in a 100-cm2 sterile petri dish containing 10 ml MEM with no additives. Mince embryos by squeezing through 10-cc syringes (∼5 embryos/syringe) into a sterile trypsinization flask. Dissociate cells 5. Add 100 ml of 37°C trypsin/EDTA and incubate 5 min in a humidified 5% CO2 incubator at 37°C, with stirring. 6. Decant fluid from the trypsinization flask into a 500-ml beaker covered with gauze. Transfer filtrate to a 250-ml centrifuge bottle. 7. Add 100 ml fresh trypsin/EDTA to the remaining tissue in the trypsinization flask, and incubate 5 min at 37°C. 8. Pour digest into a second 500-ml beaker covered with gauze, and add filtrate to the first filtrate in the 250-ml centrifuge bottle. 9. Centrifuge 10 min at 1200 × g, 4°C, in a Sorvall RC-3B centrifuge. 10. Aspirate and discard supernatant from pellet, add 10 ml complete MEM-10, resuspend by pipetting ∼10 to 15 times, and adjust volume to 100 ml. 11. Transfer to a 250-ml centrifuge bottle and centrifuge 10 min at 1200 × g, 4°C. 12. Resuspend pellet in 5 ml complete MEM-10 and adjust volume to 30 ml. Prepare confluent culture 13. Add 1 ml cell suspension to each of thirty 150-cm2 tissue culture flasks containing 30 ml complete MEM-10.
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14. Incubate at 37°C for several days until confluent, and move flasks to a humidified, 31°C, 5% CO2 incubator for storage. The CEF cell cultures can be held for 2 to 3 weeks at 31°C without further attention. Primary CEF can also be used directly for virus growth, or the trypsinized stock of CEF can be frozen in liquid nitrogen for future use. BASIC PROTOCOL 5
PREPARATION OF AN MVA STOCK To prepare an MVA stock, CEF (see Basic Protocol 4) or BHK-21 (see Basic Protocol 1) cells are plated several days before infection and allowed to approach confluency. They are then infected with MVA. After several days, the infected cells are harvested, lysed by repeated freezing and thawing, sonicated, dispensed in small aliquots, and stored at −70°C. NOTE: BHK-21 cells should be acquired from ATCC, as cells from alternative sources may support lower levels of MVA replication. Materials 150-cm2 tissue culture flasks of nearly confluent CEF (see Basic Protocol 4) or BHK-21 cells (see Basic Protocol 1) Complete MEM-10 and -2.5 (see recipe), 37°C Modified vaccinia virus Ankara (MVA; A. Mayr, Institut für Med. Mikrobiologie, Munich, Germany, or B. Moss, email [email protected]) 150-cm2 tissue culture flasks Humidified 37°C, 5% CO2 incubator Sorvall RC-3B centrifuge and sterile 250-ml centrifuge bottles (or equivalent) Additional reagents and equipment for trypsinizing cells (see Basic Protocol 1) 1. Trypsinize seven 150-cm2 tissue culture flasks of nearly confluent CEF or two of BHK-21 cells as described (see Basic Protocol 1, steps 4 to 7), using 1.5 ml trypsin/EDTA and 8.5 ml maintenance medium for the larger flask. Distribute to twenty 150-cm2 flasks. Incubate at 37°C in a humidified 5% CO2 incubator until nearly confluent monolayers have formed (usually 2 days). CEF and BHK-21 cells are split 1:3 and 1:10, respectively. It is advisable to use monolayers of BHK-21 cells at 90% confluency, as completely confluent cultures degrade within 48 hr.
2. Remove medium and add 30 ml complete MEM-2.5 to each 150-cm2 flask. 3. Thaw MVA virus and sonicate 30 sec on ice to break up clumps. Trypsinizaton of virus is avoided with MVA because it may lower the titer.
4. Add 1 to 3 infectious units of virus/cell to the medium and place cells in a CO2 incubator for 3 days at 37°C. A monolayer of CEF or BHK-21 cells in a 150 cm2 flask contains ∼1–2 × 107 cells. If there is insufficient virus, then use fewer or smaller flasks of confluent cells.
5. Detach the cells with a cell scraper or, if possible, by shaking. Transfer cells and medium to a sterile 250-ml centrifuge bottle and centrifuge 10 min at 1200 × g, 4°C, in a Sorvall RC-3B centrifuge. Discard supernatant. 6. Resuspend cells in 1 ml complete MEM-2.5 (per 150-cm2 flask) by pipetting or vortexing. Preparation of Cell Cultures and Vaccinia Virus Stocks
7. Lyse cells by freeze-thaw cycling: freeze in dry ice/ethanol, thaw in a 37°C water bath, and vortex. Carry out the freeze-thaw cycling a total of three times.
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8. Sonicate stock on ice for 1 min, pause for 1 min, and repeat sonication. Dispense in 0.5-ml aliquots and store indefinitely at −70°C. TITRATION OF MVA STOCKS BY IMMUNOSTAINING The titer of MVA is routinely determined by immunostaining, because MVA does not form distinct plaques in CEF or BHK-21 cells. To determine the MVA titer, serial dilutions of virus stock are prepared and used to infect CEF or BHK-21 cells. At 24 hr after infection, the cells are fixed with acetone/methanol and immunostained using a polyclonal vaccinia virus antiserum and a secondary peroxidase-conjugated antibody. A focus of infected cells appears as a 0.3- to 0.4-mm reddish brown–stained area. If the cells are left for two days before immunostaining, secondary foci may form by virus spread giving inaccurate titers.
SUPPORT PROTOCOL 2
Materials CEF (see Basic Protocol 4) or BHK-21 cells (see Basic Protocol 1) in a 150-cm2 tissue culture flask Complete MEM-10 and -2.5 (see recipe), 37°C MVA stock (see Basic Protocol 5) 1:1 (v/v) acetone/methanol PBS (APPENDIX 2E), with and without 3% FBS Rabbit anti-vaccinia antibody (e.g., Access BioMedica; or see Linscott, 1998) Horseradish peroxidase–conjugated whole anti–rabbit Ig antibody (HRP-anti-rabbit; Amersham) Dianisidine, or premade peroxidase substrate kit (Sigma) PBS/H2O2 (add 10 µl 30% H2O2 to 10 ml PBS immediately before use) 6-well, 35-mm2 tissue culture dishes Humidified 37°C, 5% CO2 incubator Additional reagents and equipment for trypsinizing cells (see Basic Protocol 1) Infect and fix cells 1. Trypsinize a 150-cm2 tissue culture flask of CEF or BHK-21 cells as described (see Basic Protocol 1, steps 4 to 6), using 1.5 ml trypsin/EDTA for the larger flask. Resuspend cells by repeated pipetting in 5 ml complete MEM-10, and add an additional 60 ml complete MEM-10. 2. Add 2 ml cell suspension to each well of a 6-well, 35-mm2 tissue culture dish. Incubate overnight in a humidified 5% CO2 incubator at 37°C to reach near confluency. 3. Remove medium and replace with 2 ml complete MEM-2.5. 4. Thaw MVA stock and sonicate 30 sec on ice. 5. Make eight 10-fold serial dilutions of the virus in complete MEM-2.5, using a fresh pipet for each dilution. 6. Add 0.1 ml of each of the 10−6, 10−7, and 10−8 dilutions to cells in duplicate wells. Swirl gently to mix, and incubate 24 hr in a CO2 incubator at 37°C. 7. Remove fluid and fix cells 2 min with 1 ml of 1:1 acetone/methanol. Remove fixative and add 2 ml PBS to each well. At this point, the plates can be immunostained immediately or stored at 4°C for several weeks.
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Perform immunostaining 8. Dilute rabbit anti-vaccinia antibody in PBS containing 3% FBS and add 1 ml/well. Incubate 1 hr at room temperature, with gentle rocking, if desired. The optimal dilution for each antibody must be determined empirically, but a 1:500 or 1:1000 dilution is a good place to start.
9. Wash twice with 2 ml PBS. 10. Dilute HRP-anti-rabbit secondary antibody in PBS containing 3% FBS and add 1 ml/well. Incubate for 30 to 45 min at room temperature, with gentle rocking, if desired. The optimal dilution for each antibody must be determined, but a 1:500 or 1:1000 dilution is a good place to start. If another anti-vaccinia antibody is used in step 8, the appropriate HRP-conjugated anti-species secondary antibody should be substituted here.
11. Wash twice with 2 ml PBS. 12. Make a saturated solution of dianisidine in 0.5 ml ethanol, vortex, incubate 5 min at 37°C, and clarify by microcentrifugation at maximum speed for 1 min. Add 0.2 ml dianisidine solution to 10 ml PBS/H2O2. Alternatively, use premade substrate tablets according to manufacturer’s instructions. CAUTION: Dianisidine is carcinogenic and the powder should be manipulated with gloves in a fume hood. 30% H2O2 is caustic to skin and should be handled with gloves.
13. Add 0. 5 ml dianisidine substrate solution to each well. Rotate dish gently and let stand ∼10 min. Check microscopically for foci of infected cells. When development is complete, wash dishes with water and overlay with 1 ml water to preserve stain. Weaker antibody may take longer to develop.
14. Count the number of stained foci and multiply by the dilution factor to express titer as infectious units/ml. Most accurate results are obtained from wells with 20 to 100 plaques. In determining the titer as infectious units/ml, be sure to mulitply by 10 to take into account the 0.1 ml addition of virus to the wells.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Complete MEM-2.5, -10, or -20 Minimum essential medium (MEM) containing: 2.5%, 10%, or 20% FBS 0.03% glutamine 100 U/ml penicillin 100 µg/ml streptomycin sulfate Store up to several months at 4°C Add supplements from stock solutions prepared in water at the following initial concentrations: 3% glutamine (100×), 20,000 U/ml penicillin (200×), and 20 mg/ml streptomycin (200×). Filter sterilize stock solutions. Store 100× glutamine 4 months at 4°C, and 200× penicillin/streptomycin 4 months at −20°C. Preparation of Cell Cultures and Vaccinia Virus Stocks
Fetal bovine serum is added at 10% to complete medium for maintenance of cells and at 20% to encourage initial growth upon thawing.
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Complete MEM-10/BrdU or -20/BrdU Prepare as for complete MEM-10 or -20 (see recipe) and add a 5 mg/ml solution of 5-bromodeoxyuridine (BrdU) to 25 µg/ml final. Store up to several months at 4°C. Prepare the 5 mg/ml BrdU stock solution (200×) in water and filter sterilize. Store in the dark at −20°C. After thawing 5 mg/ml BrdU, vortex to be sure it is in solution before adding to MEM.
Complete spinner medium-5 MEM spinner medium 5% horse serum Store up to several months at 4°C Supplements are in MEM spinner medium; no addition is necessary. Horse serum is used because it is cheaper than FBS and may give less cell clumping.
COMMENTARY Background Information An overview of the vaccinia life cycle and expression system is presented in UNIT 5.11. Because HeLa cells consistently give high yields of virus, they are routinely used for preparation of virus stocks. HeLa S3 cells, as obtained from the ATCC, grow well in monolayer culture but can also be put into suspension culture. After repeated passages in suspension, they do not adhere well to flasks and grow poorly in monolayer cultures. The authors prefer HeLa cells adapted to suspension culture because large numbers can be grown in a single bottle. However, suspension cultures require more maintenance than monolayer cultures and the latter may be more convenient. Suspension cells are allowed to form a monolayer before infection to increase the chances of cell-to-cell spread if not all cells are initially infected with the viral inoculum. Thus, good yields of virus may be obtained even if the inoculum is <1 pfu/cell. HeLa cells—even those adapted to monolayer culture—are fairly round to begin with and therefore show little visible evidence of infection. BS-C-1 cells, by contrast, are long and spindle-shaped but round up dramatically a few hours after infection. This property accounts for the highly visible plaques obtained with BS-C-1 cells, and their preference for titration by plaque assay. The Western Reserve (WR) strain of vaccinia virus (ATCC # VR1354) is widely used for laboratory studies. It gives high yields of cell-associated virus, discrete plaques, and is well adapted to mice and other laboratory animals. Other strains of vaccinia virus are available from the ATCC and private sources. Horse serum is used for growth of HeLa cells in suspension because it is cheaper than FBS and may give less cell clumping. Horse serum
is used in spinner medium for growth of HeLa cells in suspension. All monolayer cells are grown in medium with FBS. Modified vaccinia virus Ankara (MVA; Mayr et al., 1975) was one of several highly attenuated strains of vaccinia virus that were developed but were not extensively used for smallpox vaccination, in part because of the successful eradication of the disease with the conventional vaccine strains. Restriction enzyme analysis demonstrated that MVA had suffered multiple deletions (Meyer et al., 1991) during its long passage history in CEF that may account for its severe host restriction, including an inability to replicate efficiently in human and most other mammalian cells (Carroll and Moss, 1997). It was somewhat surprising to find that the replication defect occurred at a late stage of virion assembly and consequently that viral or recombinant gene expression was unimpaired (Sutter and Moss, 1992). Therefore, MVA can be viewed as an efficient single-round expression vector with little potential to spread. Although MVA causes less cytopathic effects than standard vaccinia virus in some cell lines, in others the effect is quite severe (Carroll and Moss, 1997). Because of its host-range restriction, recombinant MVA were initially made and propagated exclusively in primary or secondary CEF. A recent survey demonstrated that a stable Syrian hamster kidney cell line, BHK-21, was also permissive for MVA growth and could be used for making recombinant viruses (Carroll and Moss, 1997). Although many laboratories may prefer to use a stable cell line, the authors continue to employ CEF because they give slightly higher MVA titers, can be stored in a dormant state for several weeks (whereas BHK-21 cells must be passaged every 5 days),
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and are better suited to immunostaining because of greater adherence than BHK-21 cells to plastic dishes.
titer for many years when stored as indicated above (see Critical Parameters).
Time Considerations Critical Parameters Proper maintenance of actively growing cell lines is important in order to achieve efficient infections and high yields of virus. This is especially true for HeLa S3 cell suspension cultures, which should be maintained at 1.5–5 × 105 cells/ml (requiring passaging every 1 to 2 days). HeLa cells can achieve a density of 8–10 × 105 cells/ml, but exhibit a lag period upon dilution before optimal growth is resumed; prolonged maintenance at high density will lead to cell death. HeLa S3 cells adapted to grow in suspension cultures do not grow well in monolayer cultures. However, the cells will adhere to flasks, which allows the cells to be infected on the following day (see Basic Protocol 3). Since most progeny viruses remain cell-associated, infected cells must be disrupted by freeze-thaw cycling and trypsinization. These procedures are important for releasing virus from the host cells. An entire stock of virus can be subjected to freeze-thaw cycling, but only the portion to be used should be trypsinized. Sonication also helps in disaggregating virus but is usually unnecessary if the virus stock has been trypsinized. Unlike standard replicationcompetent vaccinia protocols, trypsin is not used as a pretreatment to enhance the infectivity of MVA. Pretreatment of MVA with trypsin can decrease virus titer. Although vaccinia virus is relatively stable, stocks should be kept on ice while in use and should be stored at −70°C. A vaccinia virus stock should have a stable titer for many years when stored at this temperature. Although the stock can be frozen and thawed several times without loss of infectivity, storing in aliquots is recommended. The preparation of a seed stock of virus that is sufficient for future experiments is strongly recommended. A common mistake is to continually passage the virus.
Anticipated Results
Preparation of Cell Cultures and Vaccinia Virus Stocks
A vaccinia stock should have a titer of ≥1–2 × 109 pfu/ml. MVA stocks can approach a titer of 1 × 1010 infectious units/ml when prepared as described (see Basic Protocol 5), and recombinant virus can reach 1–3 × 109 infectious units/ml. The viral stock should have a stable
Actively growing cells should be prepared in advance, as it will take ≥1 week to revive frozen cells. For preparation of a vaccinia virus stock, infected cells should be incubated for 3 days. During this period, the BS-C-1 cells for plaque titration can be prepared. For a small volume of stock (i.e., from one 150-cm2 flask), harvesting and freeze-thaw cycling can be done in <1 hr. Determination of the titer of a virus stock requires 2 days of incubation to allow for development of plaques. Preparation of CEF cells from embryonated eggs requires ∼2 hr. Thereafter, they can be stored for several weeks with no maintenance. Starting BHK-21 cells from frozen cultures can take >1 week and requires passaging every 5 days. For preparation of MVA stocks, 3 days are required for virus growth, 2 hr for harvesting, freeze-thaw cycling, and dispensing, and 1 day of incubation for determination of the virus titer by immunostaining. The immunostaining procedure takes ∼3 hr.
Literature Cited Carroll, M.W. and Moss, B. 1997. Host range and cytopathogenicity of the highly attenuated MVA strain of vaccinia virus: Propagation and generation of recombinant viruses in a nonhuman mammalian cell line. Virology 238: 198-211. Linscott, W.D. 1998. Linscott’s Directory of Immunological and Biological Reagents, 10th ed. W.D. Linscott, Santa Rosa, CA. Mayr, A., Hochstein-Mintzel, V., and Stickl, H. 1975. Abstammung, eigenschaften and verwendung des attenuierten vaccinia-stammes MVA. Infection 3:6-14. Meyer, H., Sutter, G., and Mayer, A. 1991. Mapping of deletions in the genome of the highly attenuated vaccinia virus MVA and their influence on virulence. J. Gen. Virol. 72:1031-1038. Sutter, G. and Moss, B. 1992. Nonreplicating vaccinia virus efficiently expresses recombinant genes. Proc. Natl. Acad. Sci. U.S.A. 89:1084710851.
Contributed by Patricia L. Earl, Norman Cooper, Linda S. Wyatt, and Bernard Moss National Institutes of Allergy and Infectious Diseases Bethesda, Maryland Miles W. Carroll Oxford BioMedica Oxford, United Kingdom
5.12.12 Supplement 13
Current Protocols in Protein Science
Generation of Recombinant Vaccinia Viruses
UNIT 5.13
This unit first describes how to infect cells with vaccinia virus and then transfect them with a plasmid-transfer vector to generate a recombinant virus (see Basic Protocol 1). Methods are also presented for purifying vaccinia virus (see Support Protocol 1) and for isolating viral DNA (see Support Protocol 2), which can be used during transfection. Also presented are selection and screening methods used to isolate recombinant viruses (see Basic Protocol 2) and a method for the amplification of recombinant viruses (see Basic Protocol 3). Finally, a method for live immunostaining that has been used primarily for detection of recombinant modified vaccinia virus Ankara (MVA) is presented (see Basic Protocol 4). HeLa S3 cells are used for large-scale growth of vaccinia virus. However, several other cell lines may be required for plaque purification and amplification. For thymidine kinase (TK) selection, HuTK− 143B cells are used; for xanthine-guanine phosphoribosyltransferase (XGPRT) selection, BS-C-1 cells are used. CV-1 or BS-C-1 cells are used for transfection, and either can be used for determination of virus titer (UNIT 5.12). With MVA, all steps are carried out in CEF or BHK-21 cells (see Basic Protocol 4). CAUTION: Proceed carefully and follow biosafety level 2 (BL-2) practices when working with standard vaccinia virus (see UNIT 5.11 for safety precautions). NOTE: Carry out all procedures for preparation of virus in a biosafety cabinet. TRANSFECTION OF INFECTED CELLS WITH A VACCINIA VECTOR The foreign gene of interest is subcloned into a plasmid transfer vector (Fig. 5.13.1 and Fig. 5.13.2) so that it is flanked by DNA from a nonessential region of the vaccinia genome. This recombinant plasmid is then transfected into cells that have been infected with vaccinia virus. Homologous recombination occurs between the vaccinia virus genome and homologous sequences within the plasmid (Fig. 5.13.1). The recombinant virus is obtained in a cell lysate, which is then subjected to several rounds of plaque purification using appropriate selection and/or screening protocols (see Basic Protocol 2). For modified vaccinia virus Ankara (MVA), the same transfection procedure is used, except that the virus is not trypsinized and either CEF (UNIT 5.12) or BHK-21 cells are substituted for CV-1 or BS-C-1 cells. For live immunostaining to detect recombinant MVA or standard virus, see Basic Protocol 4.
BASIC PROTOCOL 1
Materials pSC11, pRB21, pSC65, pLW9, or other suitable vector (Table 5.13.1) CV-1, BS-C-1, BHK-21, or CEF cells (UNIT 5.12) Complete MEM-10 and -2.5 media (UNIT 5.12) Vaccinia virus stock (UNIT 5.12) 0.25 mg/ml trypsin (2× crystallized and salt-free; Worthington; filter sterilize and store at −20°C) Transfection buffer (see recipe) 2.5 M CaCl2 20 mM HEPES, pH 7.4 DOTAP liposomal transfection reagent (Boehringer Mannheim) OptiMEM medium (Life Technologies) Dry ice/ethanol bath 25-cm2 tissue culture flask Contributed by Patricia L. Earl, Bernard Moss, Linda S. Wyatt, and Miles W. Carroll Current Protocols in Protein Science (1998) 5.13.1-5.13.19 Copyright © 1998 by John Wiley & Sons, Inc.
Production of Recombinant Proteins
5.13.1 Supplement 13
12 × 75–mm polystyrene tubes Disposable scraper or rubber policeman, sterile 15-ml conical centrifuge tubes Sorvall centrifuge with H-6000A rotor (or equivalent) Additional reagents and equipment for subcloning and isolation of plasmid (APPENDIX 4C), and tissue culture (APPENDIX 3C) NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly.
TKL
TKR
IacZ p11
L
p 7.5
foreign gene
TK
R double cross-over event results in integration of lacZ and foreign gene segment of plasmid into TK gene of vaccinia virus
TKL
Generation of Recombinant Vaccinia Viruses
lacZ
p11 p 7.5
foreign gene
TKR
Figure 5.13.1 Homologous recombination between a transfected plasmid and the vaccinia virus genome. TKL and TKR are vaccinia virus DNA sequences flanking the foreign gene. p11 and p7.5 are promoters.
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Prepare recombinant plasmid DNA 1. Subclone the gene of interest into the multiple cloning site (MCS) in pSC11, pRB21, pSC65, pLW9, or other suitable vector and isolate plasmid (APPENDIX 4C). Prepare vaccinia-infected cells 2. Seed a 25-cm2 flask with 1 × 106 CV-1, BS-C-1, BHK-21, or CEF cells in complete MEM-10 medium. Incubate to near confluency (usually overnight). UNIT 5.12
details the culture of these cells.
3. Just prior to use, mix equal volumes of vaccinia virus stock and 0.25 mg/ml trypsin, and vortex vigorously. Incubate 30 min in a 37°C water bath, vortexing at 5- to 10-min intervals during the incubation. For MVA, substitute 30 sec sonication on ice for trypsinization to break up clumps (also see UNIT 5.12). Virus stocks are usually at a titer of ∼2 × 109 pfu/ml, but may be significantly lower depending on the source. Vortexing usually breaks up any clumps of cells. However, if there are still visible clumps, chill to 0°C and sonicate 30 sec on ice. Sonication can be repeated several times but the sample should be allowed to cool on ice between sonications. Trypsinizaton of MVA is avoided because it may lower the titer.
Figure 5.13.2 Plasmid transfer vector pSC11.
Production of Recombinant Proteins
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Supplement 13
4. Dilute trypsinized or sonicated virus in complete MEM-2.5 to 1.5 × 105 pfu/ml. Aspirate medium from confluent monolayer of cells and infect with 1 ml diluted vaccinia virus (0.05 pfu/cell). Incubate 2 hr with rocking at ∼15-min intervals. Approximately 30 min before the end of the infection period, prepare DNA according to the CaCl2 or DOTAP method as described below.
Prepare DNA For CaCl2 method 5a. Place 1 ml transfection buffer into a 12 × 75–mm polystyrene tube and add 5 to 10 µg (in <50 µl) of the recombinant plasmid containing the gene of interest (from step 1). Inclusion of vaccinia DNA (see Support Protocol 2) in the transfection is not required but results in a higher efficiency of recombination. If it is added, combine 1 ìg vaccinia DNA Table 5.13.1 Vaccinia Virus Transfer Vectors
Insertion sitesd
Selection/ screening
Vectora
Promoterb
Cloning sitesc
pGS20 pSC11
p7.5 (E/L) p7.5 (E/L)
BamHI; SmaI TK TK SmaI; MCS
TK− TK−, β-gal
pMJ601, pMJ602 pRB21 pMC02 pSC59 pSC65 pJS4
psyn (L)
MCS
TK
TK−, β-gal
Mackett et al., 1984 Chakrabarti et al., 1985; Earl et al., 1990; Bacik et al., 1994 Davison and Moss, 1990
MCS MCS MCS MCS MCS
F12L/F13L TK TK TK TK
Plaque TK−, GUS TK− TK−, β-gal TK−
Blasco and Moss, 1995 Carroll and Moss, 1995 Chakrabarti et al., 1997 Chakrabarti et al., 1997 Chakrabarti et al., 1997
MCS
TK
TK−, gpt
Chakrabarti et al., 1997
Reference
psyn (E/L) psyn (E/L) psyn (E/L) psyn (E/L) psyn (E/L) ×2 pJS5 psyn (E/L) ×2 pG06 psyn (E/L) ×2 pLW-7 psyn (E/L) pMC03 psyn (E/L) pLW-9 pH5 (E/L) pLW-17 pH5 (E/L)
MCS
Del III
Transient gpte Sutter et al., 1994
MCS MCS MCS MCS
Del III Del III Del III Del II
Transient gpte GUS Transient gpte None
pLW-21 psyn (E/L)
MCS
Del II
None
pLW-22 psyn (E/L)
MCS
Del II
β-gal
pLW-24 p7.5 (E/L)
MCS
Del II
None
Wyatt et al., 1996 Carroll and Moss, 1995 Wyatt et al., 1996 L. Wyatt and B. Moss, unpub. observ. L. Wyatt and B. Moss, unpub. observ. L. Wyatt and B. Moss, unpub. observ. L. Wyatt and B. Moss, unpub. observ.
apRB21 was specifically designed for use with vaccinia virus vRB12, which has a deletion in the F13L gene. The plasmids
pG06, pLW-7, pMC03, pLW-9, pLW-17, pLW-21, pLW-22, and pLW-24 were designed for MVA. bAbbreviations: E, early; L, late; E/L, early and late. The designation “×2" refers to two oppositely oriented promoters
Generation of Recombinant Vaccinia Viruses
that can be used for expression of two genes. cSmaI digestion gives a blunt end for cloning any fragment that has been blunt-ended. MCS signifies multiple cloning sites. dAbbreviations: TK, thymidine kinase locus; F12L/F13L, between F12L and F13L open reading frames; Del III, site of natural deletion in MVA. eTransient selection in which XGPRT gene is deleted from recombinant vaccinia virus during recombination; see Background Information.
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with 5 to 10 ìg recombinant plasmid DNA and vortex vigorously to shear the high-molecular-weight DNA before adding CaCl2 or DOTAP. A discussion of transfection can be found in UNIT 5.10; also see Critical Parameters.
6a. Slowly add 50 µl of 2.5 M CaCl2 and mix gently. Leave 20 to 30 min at room temperature. Gentle mixing is essential; see Critical Parameters. A fine precipitate should appear.
For DOTAP method 5b. Add 70 µl of 20 mM HEPES, pH 7.4, and 30 µl DOTAP to a 12 × 75–mm polystyrene tube. In a second tube, add 5 µg recombinant plasmid containing the gene of interest (from step 1) to 20 mM HEPES for a final volume of 100 µl. See step 5a regarding optional addition of vaccinia DNA.
6b. Add the DNA solution to the DOTAP solution and leave 15 min at room temperature, then add 1 ml OptiMEM medium. Transfect cells 7. Aspirate virus inoculum from monolayer of cells (step 4). 8a. For CaCl2 method: Add the precipitated DNA suspension (step 6a) to the cells and leave 30 min at room temperature, then add 9 ml complete MEM-10 and incubate 3 to 4 hr at 37°C. 8b. For DOTAP method: Add the DNA/DOTAP solution (step 6b) to the cells and incubate 4 hr at 37°C. 9. Aspirate medium, replace with 5 ml complete MEM-10, and incubate 2 days at 37°C. 10. Dislodge cells with a disposable scraper or sterile rubber policeman and transfer to a 15-ml conical centrifuge tube. Centrifuge 5 min at 1800 × g (2500 rpm in a Sorvall H-6000A rotor), 5° to 10°C, then aspirate and discard medium. 11. Resuspend cells in 0.5 ml complete MEM-2.5. Lyse the cell suspension by performing three freeze-thaw cycles, each time by freezing in a dry ice/ethanol bath, thawing in a 37°C water bath, and vortexing. 12. Store the cell lysate at −70°C until needed in the selection and screening procedure (see Basic Protocol 2). PURIFICATION OF VACCINIA VIRUS Vaccinia is usually purified by zonal sucrose gradient centrifugation. Purified virus is useful for preparation of vaccinia DNA (see Support Protocol 2), in studies in which contaminating infected-cell proteins are undesirable, and as a very high titer stock. For large-scale purification (as in this protocol, which is for 1-liter cultures or multiples thereof) it is preferable to use HeLa cell suspensions for infection rather than monolayer cultures. If monolayer cells are used, follow the alternative procedure for monolayers (see UNIT 5.12, Basic Protocol 3) then continue with step 9 of this protocol. For many purposes, virus that is partially purified (through step 14 of this protocol) may suffice. For purification of MVA, virus is not trypsinized and either CEF or BHK-21 cells are used instead of HeLa.
SUPPORT PROTOCOL 1
Production of Recombinant Proteins
5.13.5 Current Protocols in Protein Science
Supplement 13
Materials Vaccinia virus stock (UNIT 5.12) 0.25 mg/ml trypsin (2× crystallized and salt-free; Worthington; filter sterilize and store at −20°C) HeLa S3 cells growing in suspension culture (UNIT 5.12) Complete spinner medium–5 (UNIT 5.12) 10 mM and 1 mM Tris⋅Cl, pH 9.0 (APPENDIX 2E) 36% (w/v) sucrose solution in 10 mM Tris⋅Cl, pH 9.0 40%, 36%, 32%, 28%, and 24% (w/v) sucrose solutions in 1 mM Tris⋅Cl, pH 9.0 Sorvall centrifuge with H-6000A rotor (or equivalent) 2-liter vented spinner flasks (microcarrier type; Bellco) Dounce homogenizer, glass, with tight pestle Cup sonicator (e.g., Ultrasonic Processor VC-600 from Sonics and Materials) Ultracentrifuge with Beckman SW 27 or SW 28 rotor (or equivalent) and sterile centrifuge tubes Spectrophotometer Additional reagents and equipment for tissue culture and counting cells (APPENDIX 3C), and titering virus (UNIT 5.12) NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly. Infect the cells 1. Just prior to use, mix equal volumes of vaccinia virus stock and 0.25 mg/ml trypsin and vortex vigorously. Incubate 30 min in a 37°C water bath, vortexing at 5- to 10-min intervals during the incubation. Virus stocks are usually at a titer of ∼2 × 109 pfu/ml, but may be significantly lower depending on the source. Vortexing usually breaks up any clumps of cells. However, if there are still visible clumps, chill to 0°C and sonicate 30 sec on ice. Sonication can be repeated several times but the sample should be allowed to cool on ice between sonications.
2. Count HeLa S3 cells using a hemacytometer (APPENDIX 3C). 3. Transfer 5 × 108 HeLa S3 cells into a centrifuge tube, then centrifuge 10 min at 1800 × g (2500 rpm in an H-6000A rotor), room temperature, and discard supernatant. 4. Resuspend cells in complete spinner medium-5 at 2 × 107 cells/ml. 5. Add trypsinized virus (from step 1) at an MOI of 5 to 8 pfu/cell and incubate 30 min with stirring. 6. Transfer cells to a vented spinner flask containing 1 liter complete spinner medium-5 and incubate 2 to 3 days with stirring. 7. Centrifuge cells 5 min at 1800 × g, 5° to 10°C, and discard supernatant. 8. Resuspend cells in 14 ml of 10 mM Tris⋅Cl, pH 9.0. Keep samples on ice for the remainder of the protocol. Generation of Recombinant Vaccinia Viruses
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Current Protocols in Protein Science
Lyse the cells 9. Homogenize cell suspension with 30 to 40 strokes in a glass Dounce homogenizer with tight pestle. Check for cell breakage by light microscopy. 10. Centrifuge 5 min at 300 × g (900 rpm in H-6000A rotor), 5° to 10°C, to remove nuclei. Save the supernatant. 11. Resuspend cell pellet in 3 ml of 10 mM Tris⋅Cl, pH 9.0. Centrifuge 5 min at 300 × g, 5° to 10°C. Save supernatant and pool with supernatant from step 10. 12. Sonicate pooled supernatants (lysate), keeping the lysate cold the entire time, using a cup sonicator as follows. a. Split the sample into 3-ml aliquots to be sonicated separately. b. Fill the cup with ice-water (∼50% ice). Place the tube containing the lysate in the ice-water and sonicate 1 min at full power. c. Repeat this three to four times, placing the lysate on ice for ≥30 sec between sonications. Because sonication melts the ice, it is necessary to replenish the ice in the cup.
Obtain purified virus 13. Layer the sonicated lysate onto a cushion of 17 ml of 36% sucrose in a sterile SW 27 (or SW 28) centrifuge tube. Centrifuge 80 min at 32,900 × g (13,500 rpm in an SW 27 rotor), 4°C. Aspirate and discard the supernatant. 14. Resuspend the viral pellet in 1 ml of 1 mM Tris⋅Cl, pH 9.0. At this stage, the virus may be sufficiently pure for some purposes—e.g., isolation of DNA.
15. Sonicate once for 1 min as in step 12. 16. Prepare a sterile 24% to 40% continuous sucrose gradient in a sterile SW 27 centrifuge tube the day before it is needed by carefully layering 6.8 ml each of 40%, 36%, 32%, 28%, and 24% sucrose. Let sit overnight in refrigerator. 17. Overlay the sucrose gradient with 1 ml of sonicated viral pellet from step 15. Centrifuge 50 min at 26,000 × g (12,000 rpm in an SW 27 rotor), 4°C. 18. Observe the virus as a milky band near the middle of the tube. Aspirate the sucrose above the band and discard. Carefully collect the virus band (∼10 ml) with a sterile pipet, place in a sterile tube, and save. 19. Collect aggregated virus from the pellet at the bottom of the sucrose gradient after aspirating the remaining sucrose from the tube. Resuspend the viral pellet by pipetting up and down in 1 ml of 1 mM Tris⋅Cl, pH 9.0. 20. Sonicate resuspended pellet once for 1 min as in step 12. 21. Reband the virus from the pellet as in steps 16 to 18 and pool band with band from step 18. Add 2 vol of 1 mM Tris⋅Cl, pH 9.0, and mix. Transfer to sterile SW 27 centrifuge tubes. The total volume should be ∼60 ml, which is enough to fill two SW 27 centrifuge tubes. If less volume is obtained, fill the tubes with 1 mM Tris⋅Cl, pH 9.0.
22. Centrifuge 60 min at 32,900 × g, 4°C, then aspirate and discard supernatant. 23. Resuspend the virus pellets in 1 ml of 1 mM Tris⋅Cl, pH 9.0. Sonicate as in step 12 and divide into 200- to 250-µl aliquots. Save one aliquot for step 24 and freeze remainder at −70°C.
Production of Recombinant Proteins
5.13.7 Current Protocols in Protein Science
Supplement 13
Quantitate virus 24. On the unfrozen aliquot, estimate the amount of virus spectrophotometrically at 260 nm. Use this to determine the amount of virus needed in the protocol for isolation of vaccinia virus DNA (see Support Protocol 2). One optical density unit is ∼1.2 × 1010 virus particles, which is ∼2.5–5 × 108 pfu. This value is due to light scattering, not absorbance, and therefore may vary slightly with different spectrophotometers.
25. Titer the virus by sonicating 20 to 30 sec on ice (see step 12), preparing 10-fold serial dilutions down to 10−10, and infecting, in duplicate, confluent BS-C-1 cell monolayers in 6-well tissue culture plates using the 10−8, 10−9, and 10−10 dilutions (UNIT 5.12). SUPPORT PROTOCOL 2
ISOLATION OF VACCINIA VIRUS DNA If vaccinia DNA is used in the transfection protocol (see Basic Protocol 1, step 5a annotation), it is isolated after a proteinase K digestion and phenol extraction. The extracted DNA is precipitated and then dissolved in water. The DNA concentration is determined by measuring A260. Materials Purified vaccinia virus (see Support Protocol 1) 50 mM and 1 M Tris⋅Cl, pH 7.8 (APPENDIX 2E) 10% (w/v) sodium dodecyl sulfate (SDS; APPENDIX 2E) 60% (w/v) sucrose 10 mg/ml proteinase K Buffered phenol: phenol equilibrated with 50 mM Tris⋅Cl, pH 7.8 (APPENDIX 2E) 1:1 (v/v) phenol/chloroform 1 M sodium acetate, pH 7.0 100% and 95% ethanol Spectrophotometer Sorvall centrifuge with H-6000A rotor (or equivalent) Additional reagents and equipment for phenol extraction of DNA (APPENDIX 4E) and measuring DNA concentration (APPENDIX 4K) NOTE: Avoid vortexing the DNA throughout this protocol. 1. Measure the optical density of purified vaccinia virus at 260 nm and bring 20 optical density units of virus (also see Support Protocol 1, step 24) to 1.2 ml final volume with 50 mM Tris⋅Cl, pH 7.8. Because the vaccinia virus genome is large, care must be taken to avoid shearing (i.e., by not vortexing) if full-length DNA (such as for restriction digestion analysis) is desired.
2. Add the following to the vaccinia suspension (for 2-ml final volume): 0.1 ml 1 M Tris⋅Cl, pH 7.8 0.1 ml 10% SDS 0.2 ml 60% sucrose 0.4 ml 10 mg/ml proteinase K. Incubate 4 hr at 37°C.
Generation of Recombinant Vaccinia Viruses
3. Extract twice with phenol, each time by adding an equal volume of buffered phenol, mixing gently by rocking the tube, centrifuging 10 min at 300 × g (900 rpm in an H-6000A rotor), room temperature, then aspirating and saving the aqueous phase using a pipet with the tip cut off.
5.13.8 Supplement 13
Current Protocols in Protein Science
4. Using the technique described in step 3, extract once with 1:1 phenol/chloroform. 5. Add 1⁄10 vol of 1 M sodium acetate, pH 7.0, and 2.5 vol of 100% ethanol. Mix gently and chill several hours at −20°C. 6. Microcentrifuge 10 min at maximum speed, 4°C. Aspirate and discard supernatant. 7. Wash pellet twice with 95% ethanol, each time by adding the ethanol, microcentrifuging at maximum speed, and aspirating the supernatant. Air dry and dissolve in 100 µl water. 8. Prepare dilutions (using only a small amount of the total material) and measure A260 to determine DNA concentration (APPENDIX 4K). SELECTION AND SCREENING OF RECOMBINANT VIRUS PLAQUES For standard vaccinia virus strains, procedures are described involving xanthine-guanine phosphoribosyltransferase (XGPRT; Falkner and Moss, 1988) or thymidine kinase (TK; Mackett et al., 1984) for selecting virus plaques that contain recombinant DNA. A newer and simpler drug-free method (plaque selection) is based on repair of a mutation that caused the parental virus to form pin-point plaques (Blasco and Moss, 1995; see Background Information regarding choice of procedure). In addition, β-galactosidase screening (Chakrabarti et al., 1985) or β-glucuronidase (GUS) screening (Carroll and Moss, 1995) can be used alone or in conjunction with TK selection to discriminate TK− recombinants from spontaneous TK− mutants. For each method, recombinant virus (obtained in the transfection; see Basic Protocol 1) is used to infect a monolayer culture of cells—for XGPRT selection, BS-C-1 cells are used because large plaques are obtained; for TK selection, it is necessary to use a cell line such as HuTK− 143B that is deficient in thymidine kinase. Molten agarose with medium containing the appropriate selective drugs is then pipetted onto the infected cell monolayer. Because of cell-to-cell spread of virus, each productively infected cell gives rise to a plaque. After two days, a second agarose overlay containing neutral red is placed on top of the first agarose overlay, whereupon neutral red is taken up by viable cells. In infected areas of the monolayer, the cells are rounded and dead, and thus appear as colorless plaques after neutral red staining. If β-galactosidase (or GUS) screening is used, the substrate Xgal (or Xgluc) is included in the second agarose overlay. Plaques containing infected cells that have expressed β-galactosidase or GUS turn blue; thus, blue (recombinant) plaques can be distinguished from clear (parental) plaques. A Pasteur pipet is used to aspirate infected cells from the plaques and the virus is released by freeze-thaw cycling and sonication. Several rounds of plaque purification are used to ensure the absence of residual non-recombinant virus.
BASIC PROTOCOL 2
For MVA, the virus is not trypsinized and the XGPRT, β-galactosidase, and GUS methods are used with CEF or BHK-21 cells. TK selection has not been used with MVA. Basic Protocol 4 describes live immunostaining of recombinant MVA. Materials BS-C-1, HuTK− 143B, BHK-21, or CEF confluent monolayer cells (UNIT 5.12) and appropriate complete medium Complete MEM-2.5 medium (UNIT 5.12) Selective agents (for XGPRT selection; filter sterilize, and store at –20°C): 10 mg/ml (400×) mycophenolic acid (MPA; Calbiochem) in 0.1 N NaOH 10 mg/ml (40×) xanthine in 0.1 N NaOH 10 mg/ml (670×) hypoxanthine in 0.1 N NaOH Transfected cell lysate (see Basic Protocol 1)
Production of Recombinant Proteins
5.13.9 Current Protocols in Protein Science
Supplement 23
2% low-melting point (LMP) agarose (Life Technologies) in H2O, sterilized by autoclaving Complete 2× plaque medium-5 (see recipe) 5 mg/ml 5-bromodeoxyuridine (BrdU) in H2O (for TK selection; filter sterilize and store at −20°C) 10 mg/ml neutral red in H2O 4% Xgal in dimethylformamide (optional, for β-galactosidase screening) 2% Xgluc in dimethylformamide (optional, for GUS screening) Dry ice/ethanol bath 6-well, 35-mm tissue culture plates Cup sonicator (e.g., Ultrasonic Processor VC-600 from Sonics and Materials) 45°C water bath Cotton-plugged Pasteur pipets, sterile Additional reagents and equipment for tissue culture and counting cells (APPENDIX 3C) NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly. Prepare the cells 1. Trypsinize confluent monolayer culture and resuspend in appropriate complete medium as in UNIT 5.12, steps 4 to 7 of Basic Protocol 1. a. For XGPRT selection, plaque selection, or color screening, use BS-C-1 cells. b. For TK selection, use HuTK− 143B cells. c. For MVA, use BHK-21 or CEF cells. 2. Count cells using a hemacytometer (APPENDIX 3C). 3. Plate 5 × 105 cells/well in a 6-well tissue culture plate (2 ml/well final). Incubate until confluent (this should take <24 hr). 4. Prepare cells as follows. a. For XGPRT selection, preincubate monolayer for 12 to 24 hr in filter-sterilized complete MEM-2.5 containing 1⁄400 vol of 10 mg/ml MPA, 1⁄40 vol of 10 mg/ml xanthine, and 1⁄670 vol of 10 mg/ml hypoxanthine. b. For plaque or TK selection or color screening methods, do not preincubate. Prepare the lysate and infect cells 5. Just prior to use, mix 100 µl of transfected cell lysate and 100 µl of 0.25 mg/ml trypsin, and vortex vigorously. Incubate 30 min in a 37°C water bath, vortexing at 5- to 10-min intervals during the incubation, then sonicate 20 to 30 sec on ice. For MVA, omit trypsinization but sonicate 20 to 30 sec on ice to break up clumps. 6. Make four 10-fold serial dilutions (ranging from 10−1 to 10−4) of the trypsinized and/or sonicated cell lysate in complete MEM-2.5 as follows. a. For XGPRT selection, add MPA, xanthine, and hypoxanthine at the concentrations indicated in step 4, substep a, above. Generation of Recombinant Vaccinia Viruses
b. For plaque or TK selection or color screening methods, make no additions.
5.13.10 Supplement 23
Current Protocols in Protein Science
7. Aspirate medium from the cell monolayers (from step 3) and infect with 1.0 ml diluted lysate per well. Incubate 2 hr, rocking at 30-min intervals. Use dilutions between 10−2 and 10−4.
Perform agarose overlays 8. Before the 2-hr infection is finished, melt 2% LMP agarose (1.5 ml × number of wells) and place in a 45°C water bath to cool (be sure it cools to 45°C before using it to overlay cells). Prepare and warm to 45°C the necessary amount of selective plaque medium (1.5 ml × number of wells) by making the following additions to complete 2× plaque medium-5. a. For XGPRT selection, include MPA, xanthine, and hypoxanthine at twice the concentrations indicated in step 4, substep a. Twice the concentration is necessary because the 2× plaque medium will be mixed 1:1 with agarose.
b. For TK selection, include 1⁄100 vol of 5 mg/ml BrdU. c. For plaque selection or color screening, make no additions. 9. Prepare appropriate selective agarose by mixing equal volumes of 2% LMP agarose and selective plaque medium from step 8, substep a or b. 10. Aspirate the viral inoculum from the infected cells (from step 7). Overlay each well with 3 ml appropriate selective agarose and allow to solidify at room temperature or 4°C. Incubate 2 days. 11. Prepare the second agarose overlay by mixing equal volumes of 2% LMP agarose (1 ml × number of wells, melted and cooled to 45°C as in step 8) and 2× plaque medium-5 (1 ml × number of wells, warmed to 45°C) with 1⁄100 vol of 10 mg/ml neutral red. If β-galactosidase screening is to be used, add 1⁄120 vol of 4% Xgal to the agarose/plaque medium. If GUS screening is to used, add 1⁄100 vol of 2% Xgluc. Overlay each well with 2 ml of this second agarose overlay, allow to solidify, and incubate overnight. There is no need to add any FBS, glutamine, penicillin/streptomycin, or any selection drugs to this overlay medium.
Obtain the plaques 12. Add 0.5 ml complete MEM-2.5 to sterile microcentrifuge tubes. When incubation period is complete (step 11), pick well-separated plaques by squeezing the rubber bulb on a sterile, cotton-plugged Pasteur pipet and inserting the tip through the agarose to the plaque. Scrape the cell monolayer and aspirate the agarose plug into the pipet. Transfer to a tube containing 0.5 ml complete MEM-2.5. Repeat for six to twelve plaques with separate pipets, placing each in a separate tube. 13. Vortex each virus-containing tube, then perform three freeze-thaw cycles, each time by freezing in a dry ice/ethanol bath, thawing in a 37°C water bath, and vortexing. 14. Place tube containing virus into a cup sonicator containing ice-water and sonicate 20 to 30 sec at full power. If TK selection only has been used, plaque isolates should be tested by PCR, DNA dot-blot hybridization, or immunostaining (all procedures described in UNIT 5.14), because some plaques will contain spontaneous TK− mutations and not recombinant virus.
Carry out several rounds of plaque purification 15. Prepare monolayers of the appropriate cell line as described in steps 1 to 4. One 6-well plate is needed for each plaque isolate.
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16. Make three 10-fold serial dilutions at 10−1, 10−2, and 10−3, of each of several plaque isolates as described in step 6. If XGPRT selection is used, cells must be preincubated with selective drugs and serial dilutions of the viral isolates must also contain selective drugs (step 4, substep a).
17. Aspirate medium from cell monolayers and infect two wells with 1.0 ml of each dilution of virus. Incubate 2 hr, rocking by hand at 30-min intervals. 18. Repeat steps 8 to 14 for three or more rounds of plaque purification to ensure a clonally pure recombinant virus. BASIC PROTOCOL 3
AMPLIFICATION OF A PLAQUE A recombinant plaque isolate (obtained after the selection and screening protocol; see Basic Protocol 2) is amplified by infection of successively larger numbers of cells. Medium containing drugs for XGPRT or TK selection is usually used, up to and including the infection of cells in a 25-cm2 flask. Freeze-thaw cycling is carried out to release the recombinant virus from the cells. The titer of the virus stock is determined as described in UNIT 5.12. For amplification of MVA, see Basic Protocol 4. Materials Resuspended recombinant plaque (see Basic Protocol 2) Confluent monolayer cultures of appropriate cells in both a 12-well, 22-mm tissue culture plate and a 25-cm2 tissue culture flask (UNIT 5.12) Complete MEM-2.5 and -10 media (UNIT 5.12) Selective agents (for XGPRT selection; filter sterilize, and store at –20°C): 10 mg/ml (400×) mycophenolic acid (MPA; Calbiochem) in 0.1 N NaOH 10 mg/ml (40×) xanthine in 0.1 N NaOH 10 mg/ml (670×) hypoxanthine in 0.1 N NaOH 5 mg/ml 5-bromodeoxyuridine (BrdU) in H2O (for TK selection; filter sterilize and store at −20°C) Dry ice/ethanol bath Spinner culture of HeLa S3 cells (UNIT 5.12) Cup sonicator (e.g., Ultrasonic Processor VC-600 from Sonics and Materials) 15-ml conical centrifuge tubes Sorvall centrifuge with H-6000A rotor (or equivalent) 150-cm2 tissue culture flask Additional reagents and equipment for tissue culture and counting cells (APPENDIX 3C) NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly. Infect a monolayer culture of cells with a plaque 1. Place tube containing resuspended recombinant plaque into a cup sonicator containing ice-water and sonicate 20 to 30 sec at full power. 2. Infect appropriate confluent monolayer culture in 12-well plate with 250 µl (1⁄2) of each plaque isolate. Incubate 1 to 2 hr, rocking at ∼15-min intervals.
Generation of Recombinant Vaccinia Viruses
If XGPRT selection is used, the monolayer culture is preincubated for 12 to 24 hr in complete MEM-2.5 containing MPA, xanthine, and hypoxanthine (see Basic Protocol 2, step 4, substep a). Infection should also be done in the presence of these drugs.
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3. Overlay with 1 ml complete MEM-2.5 medium containing the appropriate selective agents. a. For XGPRT selection, include 1⁄400 vol of 10 mg/ml MPA, 1⁄40 vol of 10 mg/ml xanthine, and 1⁄670 vol of 10 mg/ml hypoxanthine. b. For TK selection, include 1⁄200 vol of 5 mg/ml BrdU. Incubate 2 days or until cytopathic effect (cell rounding) is obvious. 4. Scrape cells, transfer to microcentrifuge tube, and microcentrifuge 30 sec at maximum speed. Aspirate and discard medium. 5. Resuspend cells in 0.5 ml complete MEM-2.5 then perform three freeze-thaw cycles, each time by freezing in a dry ice/ethanol bath, thawing in a 37°C water bath, and vortexing. 6. Place tube containing cell suspension into a cup sonicator and sonicate 20 to 30 sec at full power. Scale up the culture 7. Dilute 0.25 ml of lysate from step 6 with 0.75 ml complete MEM-2.5 containing selective agents (see steps 2 and 3). Infect appropriate confluent monolayer culture in a 25-cm2 flask and incubate 30 min. 8. Overlay with 4 ml complete MEM-2.5 containing the appropriate selective agents (step 3). Incubate 2 days or until cytopathic effect is obvious. 9. Scrape cells, transfer to 15-ml conical centrifuge tube, and centrifuge 5 min at 1800 × g (2500 rpm in H-6000A rotor), 5° to 10°C. Resuspend cells in 0.5 ml complete MEM-2.5 and repeat freeze-thaw cycling and sonication as described in steps 5 and 6. 10. Count HeLa S3 cells from a spinner culture using a hemacytometer (APPENDIX 3C). Alternatively, prepare monolayer cells as described in UNIT 5.12 and proceed to step 13, below.
11. Place 5 × 107 HeLa S3 cells in a centrifuge tube. Centrifuge 5 min at 1800 × g, room temperature, and discard supernatant. 12. Resuspend cells in 25 ml complete MEM-10, dispense in one 150-cm2 flask, and incubate overnight (for infection the following day). 13. Remove medium from cells and replace with a mixture of 0.25 ml lysate (from step 9) and 1.75 ml complete MEM-2.5. Incubate 1 hr, rocking the flask at 15- to 30-min intervals. 14. Overlay with 25 ml complete MEM-2.5 (selection is not required at this step) and incubate 3 days. 15. Detach cells from the flask by shaking or scraping if necessary. Transfer to centrifuge tube by pipetting, then centrifuge 5 min at 1800 × g, 5° to 10°C. Aspirate and discard supernatant. 16. Resuspend cells in 2 ml complete MEM-2.5 and carry out freeze-thaw cycling three times as described in step 5. 17. Determine titer of the viral stock as described in UNIT 5.12. Freeze viral stock at −70°C. Production of Recombinant Proteins
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BASIC PROTOCOL 4
LIVE IMMUNOSTAINING OF MVA RECOMBINANTS Live immunostaining was developed for modified vaccinia virus Ankara (MVA) because this strain does not form discrete, easily recognizable plaques and because this technique helps to avoid the incorporation of selectable or screening marker genes. Immunostaining can be used for recombinant proteins that are expressed on the cell surface or in the cytoplasm. These protocols are also applicable to standard strains of vaccinia virus. The strong adherence of chicken embryo fibroblasts (CEF) to concanavalin A–coated plastic dishes make them superior to BHK-21 or other cell lines that the authors have tried. Materials 150-cm2 flask of confluent CEF (UNIT 5.12) Complete MEM-2, -2.5, and -10 media (UNIT 5.12) Transfected cell lysate (see Basic Protocol 1) Dry ice/ethanol bath Primary antibody to protein product of foreign gene Horseradish peroxidase–conjugated secondary antibody (to species of primary antibody) Concanavalin A–coated 6-well tissue culture plates (see Support Protocol 3) Cup sonicator (e.g., Ultrasonic Processor VC-600 from Sonics and Materials) Inverted microscope Sterile toothpicks Cell scraper or plunger of 1-ml syringe 75- and 150-cm2 tissue culture flasks Additional reagents and equipment for culture, trypsinization, and immunostaining of CEF cells, titering of MVA, and preparation of MVA stocks (UNIT 5.12) NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly. Prepare, infect, and immunostain CEF cells 1. Trypsinize confluent CEF monolayer cultures, seed cells in concanavalin A–coated 6-well tissue culture plates, and incubate until nearly confluent (UNIT 5.12). 2. Thaw transfected cell lysate and sonicate in a cup sonicator for 30 sec at full power, on ice. Add 100, 10, 1, or 0.1 µl of lysate to duplicate wells of CEF containing 2 ml of complete MEM-2.5 medium. Gently swirl to mix. Incubate 2 days. 3. Immunostain the unfixed cells using antibody to the protein product of the foreign gene (see UNIT 5.12, Support Protocol 2, steps 8 to 13). If the protein of interest is expressed intracellularly, remove the fluid from the wells and place plate at −70°C for 1 hr. Allow to thaw and begin staining as described in UNIT 5.12. This procedure ruptures the cells in situ and allows the antibody to penetrate.
Generation of Recombinant Vaccinia Viruses
Isolate recombinant virus 4. Aspirate the medium and examine the cell monolayer using an inverted microscope. Touch immunostaining foci with sterile toothpicks. Place toothpicks individually in small vials containing 0.5 ml of complete MEM-2.5, and break off each toothpick so that the sterile part is inside the tube.
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5. Perform three freeze/thaw cycles on each tube, each time by freezing in a dry ice/ethanol bath, thawing in a 37°C water bath, and vortexing. Place tube into a cup sonicator and sonicate 30 sec at full power. 6. Replate onto new CEF monolayers and plaque purify the MVA recombinant a second time as in steps 2 to 5. Plaque purify an additional three times. In this protocol, plaque-purification is done under a liquid overlay instead of agarose to allow immunostaining. To check stability and purity, an extra plate can be held for 3 days and then fixed and immunostained as in UNIT 5.12, Support Protocol 2. The presence of nonstaining foci, which may be detected at this time because of cytopathic effects, are due to wild-type virus or an unstable recombinant.
Amplify plaque-purified virus 7. Infect 1 well of a concanavalin A–coated 6-well plate of CEF or BHK-21 cells with 0.25 ml of plaque-purified MVA recombinant in a total volume of 2 ml MEM-2.5. Incubate 2 days. 8. Remove and discard 1 ml of medium covering the cell monolayer. Dislodge cells into the 1 ml of remaining medium with a cell scraper (or plunger of a 1 ml syringe) and transfer to a vial. Carry out three freeze/thaw cycles as in step 5 to produce a lysate. 9. Amplify the virus by inoculating 0.5 ml of lysate into 75-cm2 flask of CEF or BHK-21 cells containing 15 ml MEM-2.5. After 2 days harvest and lyse cells as in step 8. 10. Inoculate a 150-cm2 flask of CEF or BHK-21 cells containing 30 ml MEM-2.5 with 0.5 ml of lysate from step 9. 11. Determine titer and prepare large stock of recombinant MVA (UNIT 5.12, Basic Protocol 5). Freeze viral stock in aliquots at −70°C. COATING PLATES WITH CONCANAVALIN A This protocol describes the preparation of concanavalin A–coated plates to be used growing CEF monolayers for live immunostaining as in Basic Protocol 4.
SUPPORT PROTOCOL 3
Materials Concanavalin A (Sigma) Phosphate-buffered saline (PBS; APPENDIX 2E) 6-well, 35-mm tissue culture dishes Plastic bags for storage NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly. 1. Add 12 mg concanavalin A to 120 ml sterile PBS for a final concentration of 100 µg/ml. 2. Add 1 ml of the PBS/concanavalin A solution to each well of twenty 6-well plates. Incubate 1 hr at room temperature. 3. Aspirate the liquid from each well and rinse with 2 ml PBS. 4. Remove the fluid from each well and let plates dry by storing open in a biological safety hood (to keep sterile). 5. Store plates in a plastic bag (to preserve sterility) at room temperature (will keep for months).
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Complete 2× plaque medium-5 2× plaque medium (Life Technologies) containing: 5% fetal bovine serum (FBS; APPENDIX 3C) 0.03% (w/v) glutamine 100 U/ml penicillin 100 µg/ml streptomycin Store up to 3 months at 4°C Transfection buffer 0.14 M NaCl 5 mM KCl 1 mM Na2HPO4⋅2H2O 0.1% (w/v) dextrose 20 mM HEPES Adjust to pH 7.05 with 0.5 M NaOH and filter sterilize Store indefinitely at −20°C The pH of this buffer is critical and should be between 7.0 and 7.1.
COMMENTARY Background Information
Generation of Recombinant Vaccinia Viruses
Plasmid-transfer vectors have three essential components: an expression cassette consisting of a natural or synthetic vaccinia promoter, restriction endonuclease sites for insertion of foreign genes, and flanking vaccinia virus DNA that determines the site of homologous recombination. An additional component may provide antibiotic selection or color screening. Various transfer vectors are listed in Table 5.13.1 and one is presented in detail in Figure 5.13.2. Homologous recombination (Fig. 5.13.1) is the usual way of generating recombinant vaccinia viruses. Initially, recombination may result from a single cross-over event, resulting in integration of the circular plasmid and the creation of a tandem duplication; however, this intermediate is highly unstable. A second recombination event then occurs between the tandem repeats, resolving the structure into a small circular DNA molecule and either a wildtype or recombinant viral genome. An alternative method of direct ligation using an unique restriction endonuclese site in the vaccinia genome has also been described (Pfleiderer et al., 1995; Merchlinsky et al., 1997). This method avoids an E. coli cloning step and can potentially be used for direct cloning of cDNA libraries in vaccinia virus. Considerable attention has been devoted to promoters because they affect both the time and
level of expression (UNIT 5.11). The most widely used promoter, p7.5, actually contains tandem late and early promoters (Cochran et al., 1985) and provides a moderate level of expression throughout infection. The modified pH5 promoter provides higher levels of both early and late expression than p7.5 (Wyatt et al., 1996). Strong natural late promoters include p11 (Bertholet et al., 1985) and pCAE (Patel et al., 1988). About a two-fold increase in expression may be achieved with the synthetic late promoter in pMJ601 and pMJ602 (Davison and Moss, 1990) or the synthetic early/late compound promoter in pSC59 and pSC65 (Chakrabarti et al., 1997; Table 5.13.1). The ability to synthesize many different kinds of proteins, including those with transmembrane domains, is one advantage of the vaccinia virus expression system. Nevertheless, very high expression of certain genes might adversely affect virus replication. If difficulty is experienced in obtaining recombinant vaccinia with strong promoters, the weaker p7.5 or pH5 promoter or the inducible vaccinia virus/bacteriophage T7 hybrid system should be tried (UNIT 5.15). Several different methods for selecting recombinant viruses are available. The widely used TK selection method is based on the insertional inactivation of the thymidine kinase gene (UNIT 5.10). In the presence of active TK, added BrdU is phosphorylated and incorpo-
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rated into viral DNA, where it causes lethal mutations. If TK− cells are used, then TK− virus will replicate normally in the presence of BrdU, whereas TK+ virus will not. Because the product of a single cross-over event is still TK+, only double cross-over events are selected. However, not all TK− viral plaques will be recombinants because spontaneous TK− mutants arise at a frequency of 1:10,000. Depending on the transfection efficiency, recombinants may comprise 10% to >90% of the TK− plaques. β-galactosidase or β-glucuronidase (GUS) screening is based on the coinsertion of the E. coli lacZ or GUS gene, under the control of a vaccinia virus promoter, into the vaccinia virus genome along with the gene of interest. If the TK gene is insertionally inactivated by such an event, recombinant viruses will be TK− and will make blue plaques on medium containing Xgal or Xgluc. This combination of color screening and TK selection will discriminate TK− recombinants from spontaneous TK− mutants. An example of an insertion vector that provides both TK− and β-galactosidase screening is shown in Figure 5.13.2. XGPRT selection uses mycophenolic acid (MPA), an inhibitor of purine metabolism (see UNIT 5.10). Because MPA blocks the pathway for GMP synthesis, it interferes with the replication of vaccinia and severely reduces the size of vaccinia plaques. This effect can be overcome, however, by expression of the E. coli gpt gene in the presence of xanthine and hypoxanthine (i.e., XGPRT can use either of these as substrate for synthesis of GMP). Thus, coexpression of XGPRT provides a useful selection system. Usually the XGPRT gene is placed adjacent to the gene of interest and within the vaccinia virus DNA flanking sequences. Unlike the TK− situation, the viral products arising from both single and double cross-over events will be selected. Therefore, it is important to do successive plaque isolations so that the single cross-over events will be resolved. A reverseselection procedure can be used to delete the XGPRT gene from a recombinant vaccinia virus or replace it with another gene (Isaacs et al., 1990). Another procedure, transient XGPRT selection, is performed using a transfer vector with the XGPRT gene outside of the vaccinia virus DNA flanking sequences (Falkner and Moss, 1990). Under these conditions, only the single cross-over recombinant virus will express XGPRT, providing substantial enrichment over the parental virus. However, when MPA selection is removed, the desired double cross-over recombinant virus without the
XGPRT gene will have to be differentiated from the parental virus by PCR, immunostaining, or other methods. Plaque selection (Blasco and Moss, 1995) provides an extremely simple method of selecting recombinant vaccinia viruses that involves neither drugs nor reporter genes. The parental virus (vRB12) contains a deletion of most of the F13L gene, which prevents vaccinia virus from making normal-size plaques. The transfer vector pRB21 contains a segment of the F13L gene adjacent to the gene of interest, so that recombinant viruses will have a functional F13L gene and will make normal-size plaques that are easily distinguished from the pin-point plaques of vRB12. Therefore, one merely needs to pick large plaques to isolate recombinant viruses. However, it is important to check that the virus is a double cross-over recombinant. To avoid further impairment of MVA, foreign genes have been targeted to existing deletion sites. Deletion II encompasses parts of the HindIII N-K fragments; deletion III lies within the HindIII A fragment (Meyer et al., 1991). The TK gene has been successfully used as an insertion site for MVA (Carroll and Moss, 1997; T. Blanchard, pers. comm.), although there have been reports of some difficulties in isolating stable TK− MVA (Scheiflinger et al., 1996). Since there are no MVA-permissive TK− cell lines, the main attraction for using the TK site is the large number of available transfer vectors that target this site. When some foreign genes were placed under the control of the very strong synthetic early/late promoter, the resulting recombinant MVA were difficult to isolate or unstable (Wyatt et al., 1996). To date, the examples have been membrane proteins. In most cases, stable recombinants could be made by using a promoter of lower strength, e.g., pH5 or p7.5.
Critical Parameters During purification of vaccinia virus (see Support Protocol 1) it is important to check the extract after Dounce homogenization to be sure that cells are broken; if they are not, repeat with more force for ten more strokes and check again. In addition, be sure to perform all steps on ice (this is particularly important when sonicating). Use sterile technique throughout. Obtaining a very fine precipitate of calcium phosphate and DNA is critical for efficient transfection. Several factors can influence the quality of the precipitate. First, the pH of the transfection buffer should be between 7.0 and 7.1. Second, if wild-type vaccinia DNA is used,
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it should be vortexed vigorously to shear the high-molecular-weight DNA prior to precipitation. Although coprecipitation of viral DNA with the insertion vector increases the efficiency of recombination, it is not required if a selection procedure is used. Third, the CaCl2 should be added slowly and mixed gently (not vortexed), only until full mixing is achieved. The tube should then be left undisturbed until the solution is layered on the cells. Cationic lipids are simpler to use and provide similar or better transfection efficiencies. Although the DOTAP procedure was described in this unit, other cationic lipids are also effective. During plaque purification and amplification of a new recombinant virus, it is important to maintain the appropriate selective pressure to prevent growth of any contaminating wildtype virus. After isolation, selection is not required. Check the purity of a recombinant virus by PCR or Southern blot analysis of a HindIII digest of the virus; the fragment into which the foreign gene has been cloned (the 5.1-kb HindIII J fragment for TK selection) should have increased by the size of the insert. Freezing vaccinia stocks causes clumping of virus particles; thus, stocks should be trypsinized and/or sonicated after thawing. This is particularly important when plaque-purifying a virus, as each plaque should have arisen from a single virus. Trypsinization has been avoided with MVA because preliminary experiments suggested a loss in titer. Some lots of agarose contain contaminants that are toxic to cells (the authors have used low-meltingpoint agarose from Life Technologies successfully). It is prudent to save half of each stock at each step in case of contamination or failure at a succeeding step. The authors strongly recommend preparation of a seed stock of the final recombinant virus that will be adequate for future needs. A common mistake is to continually passage the virus. If immunostaining is used instead of selection or color screening, the antibody must give a good signal to initially pick out the few small foci of cells infected with recombinant virus among a great excess of cells infected with the parental nonstaining virus. Before using an antibody for isolation of recombinant MVA, it can be tested by staining the MVA-infected cells after the first transfection step. See UNIT 5.11 for safety precautions that need to be taken when working with vaccinia virus.
Anticipated Results
Approximately 1–5 × 1010 pfu of purified virus should be obtained per liter of 5 × 108 HeLa cells. Depending on the efficiency of the transfection, single, well-isolated plaques should be visible in cells infected with one of the recommended virus dilutions. With TK selection, from 10% to 90% of the plaques will contain recombinant virus. If β-galactosidase or GUS screening is also used, only blue recombinant virus plaques should be picked. With XGPRT selection, all plaques picked should contain recombinant virus. If the titer of recombinant virus is low, amplification can be achieved by a round of growth in the presence of MPA prior to plaquing (use the procedure described in amplification of a plaque). Cytopathic effects should be clearly visible at each step of amplification except with final infection (infected HeLa cells do not exhibit clear cytopathic effects). The titer of the final crude stock should be 1–2 × 109 pfu/ml.
Time Considerations During purification of vaccinia virus, the viral amplification takes 2 to 3 days. After harvesting the infected cells, the entire purification can be done in 1 day. However, the protocol can be stopped after resuspension at step 12 or after collecting the band at step 16. The virus can be stored overnight at 4°C or at −70°C for longer periods. The infection/transfection procedure for generation of recombinant virus takes ∼6 hr and cells are harvested after 2 days. Each round of selection and plaque purification takes 3 days, although little working time is required. Plaques can be picked and reinfections performed on the same day. Amplification of a single plaque isolate to a small high-titer crude stock will take ∼7 days. Large stocks should then be prepared as described in UNIT 5.13. Transfection of cells with the plasmid transfer vector requires ∼2 hr to complete, and the cells are harvested after 2 days. Each round of isolation and plaque purification requires about 30 min for infection, 2 days for incubation, 3 hr for color or immunostaining, and 2 hr for isolating new recombinants and freeze-thaw cycling. Recombinant isolates can be picked and reinfections performed on the same day. From completion of plaque purification to the preparation and titering of a virus stock takes another 10 days.
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Literature Cited Bacik, I., Cox, J.H., Anderson, R., Yewdell, J.W., and Bennink, J.R. 1994. TAP (transporter associated with antigen processing)–independent presentation of endogenously synthesized peptides is enhanced by endoplasmic reticulum insertion sequences located at the amino- but not carboxyl-terminus of the peptide. J. Immunol. 152:381-387. Bertholet, C., Drillien, R., and Wittek, R. 1985. One hundred base pairs of 5′ flanking sequence of a vaccinia virus late gene are sufficient to temporally regulate late transcription. Proc. Natl. Acad. Sci. U.S.A. 82:2096-2100. Blasco, R. and Moss, B. 1995. Selection of recombinant vaccinia viruses on the basis of plaque formation. Gene 158:157-162. Carroll, M.W. and Moss, B. 1995. E. coli β-glucuronidase (GUS) as a marker for recombinant vaccinia viruses. BioTechniques 19:352-355. Carroll, M.W. and Moss, B. 1997. Host range and cytopathogenicity of the highly attenuated MVA strain of vaccinia virus: Propagation and generation of recombinant viruses in a nonhuman mammalian cell line. Virology 238:198-211. Chakrabarti, S., Brechling, K., and Moss, B. 1985. Vaccinia virus expression vector: Coexpression of beta-galatosidase provides visual screening of recombinant virus plaques. Mol. Cell. Biol. 5:3403-3409. Chakrabarti, S., Sisler, J.R., and Moss, B. 1997. Compact, synthetic, vaccinia virus early/late promoter for protein expression. BioTechniques 23:1094-1097. Cochran, M.A., Puckett, C., and Moss, B. 1985. In vitro mutagenesis of the promoter region for a vaccinia virus gene: Evidence for tandem early and late regulatory signals. J. Virol. 54:30-37. Davison, A.J. and Moss, B. 1990. New vaccinia virus recombination plasmids incorporating a synthetic late promoter for high level expression of foreign proteins. Nucl. Acids Res. 18:42854286. Earl, P., Koenig, S., and Moss, B. 1990. Biological and immunological properties of human immunodeficiency virus type 1 envelope glycoprotein: Analysis of proteins with truncations and deletions expressed by recombinant vaccinia viruses. J. Virol. 65:31-41. Falkner, F.G. and Moss, B. 1988. Escherichia coli gpt gene provides dominant selection for vaccinia virus open reading frame expression vectors. J. Virol. 62:1849-1854. Falkner, F.G. and Moss, B. 1990. Transient dominant selection of recombinant vaccinia viruses. J. Virol. 64:3108-3111.
Mackett, M., Smith, G.L., and Moss, B. 1984. General method for production and selection of infectious vaccinia virus recombinants expressing foreign genes. J. Virol. 49:857-864. Meyer, H., Sutter, G., and Mayr, A. 1991. Mapping of deletions in the genome of the highly attenuated vaccinia virus MVA and their influence on virulence. J. Gen. Virol. 72:1031-1038. Merchlinsky, M., Eckert, D., Smith, E., and Zauderer, M. 1997. Construction and characterization of vaccinia direct ligation vectors. Virology 238:444-451. Patel, D.D., Ray, C.A., Drucker, R.P., and Pickup, D.J. 1988. A poxvirus-derived vector that directs high levels of expression of cloned genes in mammalian cells. Proc. Natl. Acad. Sci. U.S.A. 85:9431-9435. Pfleiderer, M., Falkner, F.G., and Dorner, F. 1995. A novel vaccinia virus expression system allowing construction of recombinants without the need for selection markers, plasmids and bacterial hosts. J. Gen. Virol. 76:2957-2962. Scheiflinger, F., Falkner, F.G., and Dorner, F. 1996. Evaluation of the thymidine kinase (tk) locus as an insertion site in the highly attenuated vaccinia MVA strain. Arch. Virol. 141:663-669. Sutter, G., Wyatt, L.S., Foley, P.L., Bennink, J.R., and Moss, B. 1994. A recombinant vector derived from the host range-restricted and highly attenuated MVA strain of vaccinia virus stimulates protective immunity in mice to influenza virus. Vaccine 12:1032-1040. Wyatt, L.S., Shors, S.T., Murphy, B.R., and Moss, B. 1996. Development of a replication-deficient recombinant vaccinia virus vaccine effective against parainfluenza virus 3 infection in an animal model. Vaccine 14:1451-1458.
Key References Mackett et al., 1984. See above. Piccini, A., Perkus, M.E., and Paoletti, E. 1987. Vaccinia virus as an expression vector. Methods Enzymol. 153:545-563.
Contributed by Patricia L. Earl, Bernard Moss, and Linda S. Wyatt National Institute of Allergy and Infectious Diseases Bethesda, Maryland Miles W. Carroll Oxford BioMedica Oxford, United Kingdom
Isaacs, S.N., Kotwal, G.J., and Moss, B. 1990. Reverse guanine phosphoribosyltransferase selection of recombinant vaccinia viruses. Virology 178:626-630. Production of Recombinant Proteins
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Characterization of Recombinant Vaccinia Viruses and Their Products
UNIT 5.14
After a recombinant vaccinia virus is made (UNIT 5.13), its DNA and protein products can be analyzed in several ways. Identification of the recombinant virus can be carried out by PCR (see Basic Protocol 1), with verification of correct insertion of the DNA by Southern blotting (see Basic Protocol 2). Vaccinia DNA can also be detected by dot-blot hybridization (see Basic Protocol 3). Finally, when antibodies are available, protein expression can be analyzed by immunological methods such as dot blotting with an antibody (see Alternate Protocol) or immunoblotting (see Basic Protocol 4) and/or immunoprecipitation (see Basic Protocol 5). In addition, immunostaining can be used for identification of recombinant plaques as well as for determination of the purity of a recombinant virus stock. For these purposes, the support protocol for immunostaining in UNIT 5.12 should be used; however an antibody to the foreign protein should be substituted for anti-vaccinia antibody. All of the protocols in this unit can be used for characterization of modified vaccinia virus Ankara (MVA) recombinant viruses (UNIT 5.13). CAUTION: Proceed carefully and follow biosafety level 2 (BL-2) practices when working with vaccinia virus (see UNIT 5.11 for safety guidelines). NOTE: Carry out all procedures for growth of vaccinia virus using sterile technique in a biosafety cabinet. DETECTION OF VACCINIA DNA USING PCR The presence of recombinant virus in isolated plaques can be determined by infecting cells with the virus, extracting the DNA, and performing PCR analysis utilizing oligonucleotide primers in the flanking vaccinia DNA. Although PCR analysis can be done directly on DNA in picked virus plaques (Zhang and Moss, 1991), individual researchers sometimes have problems with this shorter procedure. PCR is also useful for determining the purity of a viral stock, as both wild-type and recombinant DNAs are amplified using this protocol. Here a method is described for isolation of DNA from infected cells. An alternative to this protocol is to use a commercial kit such as Qiagen QIAamp Blood Kit for extraction of DNA. Materials Confluent BS-C-1 or HuTK− 143B cell monolayer (UNIT 5.12) Phosphate-buffered saline (PBS; APPENDIX 2E) Trypsin/EDTA (0.25%:0.02%), 37°C Complete MEM-10, -2.5, and -5 media (UNIT 5.12) Complete MEM-10/BrdU medium (UNIT 5.12) Recombinant virus plaques, picked and resuspended in tubes (UNIT 5.13) Selective agents (filter sterilize and store at −20°C; also see UNIT 5.13): 10 mg/ml (400×) mycophenolic acid (MPA; Calbiochem) in 0.1 N NaOH (for XGPRT selection) 10 mg/ml (40×) xanthine in 0.1 N NaOH (for XGPRT selection) 10 mg/ml (670×) hypoxanthine in 0.1 N NaOH (for XGPRT selection) 5 mg/ml (200×) 5-bromodeoxyuridine (BrdU) in water (for TK selection) Dry ice/ethanol bath DNA extraction buffer (see recipe) 3 M sodium acetate, pH 6.0 (APPENDIX 2E) Contributed by Patricia L. Earl and Bernard Moss Current Protocols in Protein Science (1998) 5.14.1-5.14.11 Copyright © 1998 by John Wiley & Sons, Inc.
BASIC PROTOCOL 1
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95% and 70% ethanol, ice cold 10× MgCl2-free PCR amplification buffer (APPENDIX 4J) 10 mM 4dNTP mix (APPENDIX 4J) 100 µM primer (sequence depends on where foreign gene is inserted) 15 mM MgCl2 24-well tissue culture plates Cup sonicator Additional reagents and equipment for tissue culture and counting cells (APPENDIX 3C), phenol/chloroform extraction of DNA (APPENDIX 4E), PCR amplification (APPENDIX 4J), and agarose gel electrophoresis (APPENDIX 4F) NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly. Set up cells for infection 1. Aspirate medium from a confluent cell monolayer. a. For XGPRT selection, use BS-C-1 cells. b. For TK selection, use HuTK− 143B cells. 2. Wash cells once with 37°C PBS or trypsin/EDTA to remove the remaining serum. 3. Overlay cells with 37°C trypsin/EDTA using a volume that is just enough to cover the monolayer (1.5 ml for a 150-cm2 flask). Allow to sit ∼30 sec (the cells should become detached). Shake to detach cells completely. 4. Add 8.5 ml of the appropriate medium. a. For BS-C-1 cells, use complete MEM-10. b. For HuTK− 143B cells, use complete MEM-10/BrdU. Pipet cell suspension up and down several times to disrupt clumps. 5. Count cells using a hemacytometer (APPENDIX 3C). 6. Plate 1.25 × 105 cells/well in 24-well tissue culture dishes (0.5 ml/well final). Incubate until confluent (this should take <24 hr). Infect cells with virus 7. Place tube containing virus into a cup sonicator containing an ice/water mixture and sonicate 20 to 30 sec at full power. 8. Aspirate medium from each confluent well and infect each well with one recombinant virus plaque, using half the volume in which each plaque is suspended (i.e., 0.25 ml where 0.5 ml is used to resuspend the plaque). Incubate 1 to 2 hr, rocking at 15- to 30-min intervals. Perform selection 9. Overlay with 1 ml complete MEM-2.5 to which appropriate selection drugs have been added. Incubate until cytopathic effect (cell rounding) is evident (usually 24 to 48 hr). Characterization of Recombinant Vaccinia Viruses and Their Products
a. For XGPRT selection, add 1⁄400 vol of 10 mg/ml MPA, 1⁄40 vol of 10 mg/ml xanthine, and 1⁄670 vol of 10 mg/ml hypoxanthine. b. For TK selection, add 1⁄200 vol of 5 mg/ml BrdU.
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10. Scrape cells and transfer to a microcentrifuge tube. Microcentrifuge 30 sec at maximum speed. Aspirate medium, add 1 ml PBS, vortex to resuspend, then microcentrifuge again, 30 sec at maximum speed. Aspirate PBS, then resuspend pellet in 0.5 ml PBS. Cells in a 24-well plate can be scraped using the plunger of a 1-ml syringe.
Prepare DNA for PCR amplification 11. Lyse the cell suspension by carrying out three freeze-thaw cycles, each time by freezing in a dry ice/ethanol bath, thawing in a 37°C water bath, and vortexing. 12. Place tube containing lysate into a cup sonicator containing an ice/water mixture and sonicate 20 to 30 sec at full power. 13. Place 50 to 100 µl of the lysate into a microcentrifuge tube. Add 1⁄10 vol DNA extraction buffer. Mix by gently inverting the tube several times. Incubate 2 hr to overnight at 37°C. 14. Extract once with an equal volume of buffered phenol and once with an equal volume of chloroform (APPENDIX 4E). 15. Add 1⁄10 vol of 3 M sodium acetate, pH 6.0 (0.3 M final) and 2.5 vol of 95% ethanol. Place at −20°C for 20 min. 16. Microcentrifuge 15 min at maximum speed, 4°C. Aspirate and discard supernatant. Wash pellet with ice-cold 70% ethanol, then air dry. 17. Dissolve pellet in 20 µl deionized water. Prepare the following reaction mix on ice: 10 µl redissolved (template) DNA pellet 5 µl 10× MgCl2-free PCR amplification buffer 5 µl 10 mM 4dNTP mix 0.5 µl 100 µM primer 5 µl 15 mM MgCl2 24.5 µl H2O. Overlay with mineral oil. A PCR kit such as the Expand High Fidelity PCR System (Boehringer Mannheim) is appropriate for this step.
18. Carry out PCR (APPENDIX 4J) using the following thermal cycling parameters: 35 cycles:
1 cycle: Final step:
20 sec 20 sec 1 to 2 min 5 min indefinitely
95°C 55°to 60°C 72°C 72°C 4°C
(denaturation) (annealing) (extension) (extension) (hold).
These conditions are for a 1-kb fragment.
19. Analyze PCR product on agarose gel (APPENDIX 4F).
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BASIC PROTOCOL 2
DETECTION OF VACCINIA DNA USING SOUTHERN BLOT HYBRIDIZATION Historically, vaccinia virus DNA has been characterized by its HindIII restriction endonuclease digestion pattern. The TK gene, into which most foreign genes are inserted in recombinant viruses, is located in the 5.1-kbp HindIII J fragment. Thus in a recombinant virus, if the inserted gene has no HindIII sites, this fragment should be increased in size. The 5.1-kbp HindIII J fragment should be absent, indicating lack of contamination with wild-type vaccinia virus. Southern blot analysis can be done with DNA from purified virus (UNIT 5.13) or, more easily, from a crude lysate of infected cells as described below. Materials Confluent BS-C-1 cell monolayer (UNIT 5.12) Complete complete MEM-10 and -5 media (UNIT 5.12) Recombinant vaccinia virus stock (UNIT 5.12) 0.25 mg/ml trypsin (2× crystallized and salt-free;Worthington; filter sterilize and store at −20°C) Phosphate-buffered saline (PBS; APPENDIX 2E) Low-salt buffer (see recipe) 20 mg/ml proteinase K Buffered phenol (APPENDIX 2E) 1:1 phenol/chloroform 3 M sodium acetate, pH 6.0 (APPENDIX 2E) 95% and 70% ethanol, ice-cold TE buffer, pH 7.8 (APPENDIX 2E) HindIII restriction endonuclease (APPENDIX 4I) and appropriate buffer 12-well tissue culture plates Additional reagents and equipment for preparing BS-C-1 cells for vaccinia infection (see Basic Protocol 1), tissue culture and counting cells (APPENDIX 3C), phenol/chloroform extraction of DNA (APPENDIX 4E), restriction endonuclease digestion of DNA (APPENDIX 4I), Southern blotting (APPENDIX 4G), and hybridization (APPENDIX 4H) NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly. Prepare cell cultures and infect cells 1. Set up cell cultures for infection (see Basic Protocol 1, steps 1 to 5) starting with a confluent BS-C-1 cell monolayer. 2. Plate 2.5 × 105 cells/well in 12-well tissue culture plates (1 ml/well final) using complete MEM-10 medium. Incubate until confluent (this should take <24 hr). 3. Just prior to use, mix equal volumes of recombinant vaccinia virus stock and 0.25 mg/ml trypsin and vortex vigorously. Incubate 30 min in a 37°C water bath, vortexing at 5- to 10-min intervals during the incubation. Virus stocks are usually at a titer of ∼2 × 109 pfu/ml but may be significantly lower depending on the source.
Characterization of Recombinant Vaccinia Viruses and Their Products
Vortexing usually breaks up any clumps. However, if clumps still remain, chill to 0°C and sonicate 30 sec on ice. Sonication can be repeated several times but the sample should be allowed to cool on ice between sonications.
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4. Aspirate medium and infect monolayer culture at a multiplicity of infection (MOI) of 10 to 20 pfu/cell in 250 µl complete MEM-5. Incubate 1 to 2 hr, rocking at ∼30-min intervals. 5. Overlay with 1 ml complete MEM-5 and incubate ∼24 hr. Harvest cells 6. Scrape cells and transfer to a microcentrifuge tube. Microcentrifuge 30 sec at maximum speed, room temperature, then aspirate and discard supernatant. The plunger of a 1-ml syringe can be used to scrape cells.
7. Resuspend pellet in 50 µl PBS and vortex. 8. Add 300 µl of low-salt buffer and 10 µl of 20 mg/ml proteinase K to a microcentrifuge tube. Add cell suspension to this same tube. Vortex, then incubate 5 hr to overnight at 37°C. Extract DNA and analyze by Southern blotting 9. Extract once with an equal volume of buffered phenol, then once with an equal volume of 1:1 phenol/chloroform (APPENDIX 4E). If the solution is viscous after phenol extraction, pass it through a 25-G needle to shear DNA and reduce viscosity.
10. Add 1⁄10 vol 3 M sodium acetate, pH 6.0 (0.3 M final) and 2.5 vol 95% ethanol. Place 30 min on dry ice or overnight at −20°C. 11. Microcentrifuge 10 min at maximum speed, 4°C. Aspirate and discard supernatant. 12. Wash pellet with ice-cold 70% ethanol, then air dry. 13. Dissolve pellet in 100 µl TE buffer, pH 7.8. 14. Digest 15 µl of DNA solution with HindIII (APPENDIX 4I) and analyze by Southern blotting and hybridization (APPENDIX 4H). DETECTION OF VACCINIA DNA USING DOT-BLOT HYBRIDIZATION In addition to PCR and Southern blotting, DNA dot-blot hybridization, as described in this protocol, may be used for identifying recombinant virus in isolated plaques.
BASIC PROTOCOL 3
Materials 0.5 N NaOH 1 M Tris⋅Cl, pH 7.5 (APPENDIX 2E) 2× SSC (APPENDIX 2E) Nitrocellulose membrane Whatman 3MM filter paper 80°C vacuum oven Additional reagents and equipment for preparing cells, infecting with vaccinia virus, and performing selection (see Basic Protocol 1), dot blotting (APPENDIX 4G), and hybridization (APPENDIX 4H) NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly.
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Prepare and infect cells and harvest cell suspension 1. Infect cells with virus and perform selection (see Basic Protocol 1, steps 1 to 9). 2. Scrape cells and transfer to a microcentrifuge tube. Microcentrifuge 30 sec at maximum speed, then aspirate and discard medium. Resuspend cell pellet in 200 µl PBS. Cells in a 24-well plate can be scraped using the plunger of a 1-ml syringe.
3. Freeze thaw and sonicate cell suspension (see Basic Protocol 1, steps 11 and 12). Perform blotting procedure 4. Cut nitrocellulose membrane to the appropriate size to fit in the dot-blot apparatus. Wet the membrane by immersing it in water and place in the apparatus. Add 20 to 100 µl cell lysate (from step 3) to each well. Draw the liquid through the membrane by applying suction, then remove the nitrocellulose from the apparatus. 5. Cut six pieces of Whatman 3MM filter paper so they are larger than the nitrocellulose membrane. Soak one piece of 3MM paper with 0.5 N NaOH. Lay the nitrocellulose membrane on top of the wet 3MM paper and let sit for 1 min. Remove membrane and place it on a separate piece of dry 3MM paper for 1 min. Repeat this procedure two times using the same piece of 3MM paper. Forceps should be used to transfer the membrane.
6. Repeat step 5 (three times per membrane) with Whatman 3MM paper soaked in 1 M Tris⋅Cl, pH 7.5. 7. Repeat step 5 (three times per membrane) with 3MM Whatman paper soaked in 2× SSC. 8. Bake membrane 2 hr at 80°C under vacuum. Baking fixes the DNA to the filter.
9. Proceed with DNA hybridization as described in APPENDIX 4H. ALTERNATE PROTOCOL
DETECTION OF EXPRESSED PROTEIN BY A DOT-BLOT PROCEDURE This protocol is an alternative to DNA dot-blot hybridization for checking plaques to identify recombinant virus. Infected cell lysates are spotted onto a nitrocellulose membrane as in DNA dot-blotting (see Basic Protocol 3). The membrane is then incubated with an antibody that recognizes the expressed foreign gene product. 125I-labeled protein A is used to detect the bound antibody. Alternatively, a chemiluminescent detection system such as Amersham ECL can be used. The membrane is then exposed to X-ray film. Note that infected cells comprising a plaque can be stained in situ by following the protocol in UNIT 5.12 (Basic Protocol 4, titration of MVA by immunostaining) except that antibody to the recombinant protein should be substituted for antibody to vaccinia virus.
Characterization of Recombinant Vaccinia Viruses and Their Products
Additional Materials (also see Basic Protocol 3) PBS/Tween (see recipe) with and without 4% (w/v) BSA Antibody to foreign protein 100 µCi/ml [125I]-labeled protein A (30 mCi/mg; Amersham) Additional reagents and equipment for autoradiography (UNIT 10.11) 1. Prepare cells and blot cell lysate (see Basic Protocol 3, steps 1 to 4).
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2. Soak membrane ∼30 min in PBS/Tween containing 4% BSA. Wash once with PBS/Tween without BSA. Washing is done by pouring liquid off and replacing it.
3. Dilute antibody to foreign protein 1:50 to 1:5000 in a minimal volume of PBS/Tween. Incubate membrane in this solution ≥1 hr at room temperature. 4. Wash approximately five times with an excess volume of PBS/Tween. 5. Incubate membrane for 30 min in a minimal volume of PBS/Tween containing 1 µCi of [125I]-labeled protein A. Alternatively, a chemiluminescent detection kit (e.g., Amersham ECL) may be used to detect the antibody
6. Wash membrane approximately five times with an excess volume of PBS/Tween. 7. Wrap membrane in plastic wrap and autoradiograph by exposing to X-ray film. DETECTION OF EXPRESSED PROTEIN USING IMMUNOBLOTTING After a recombinant virus has been plaque purified and amplified, immunoblotting can be used to determine if the recombinant protein is of the expected size and whether or not it is secreted from infected cells. The steps below are for nonsecreted (intracellular) proteins, but annotations describe procedures when working with cells in which the recombinant protein is secreted into the medium.
BASIC PROTOCOL 4
Materials Confluent BS-C-1 cell monolayer (UNIT 5.12) Complete MEM-5 medium (UNIT 5.12) Recombinant vaccinia virus stock (UNIT 5.12) 0.25 mg/ml trypsin (2× crystallized and salt-free;Worthington; filter sterilize and store at −20°C) Cell lysis buffer (see recipe) 5× SDS sample buffer (UNIT 10.1) 6-well tissue culture plates Sorvall centrifuge with H-6000A rotor (or equivalent) 95°C water bath Additional reagents and equipment for preparing cell cultures for infection (see Basic Protocol 1) and immunoblotting (UNIT 10.10) NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly. 1. Set up cell cultures for infection (see Basic Protocol 1, steps 1 to 5) starting with a confluent BS-C-1 cell monolayer. 2. Plate 5 × 105 cells/well in a 6-well tissue culture plate (2 ml/well final) using MEM-5 medium. Incubate until confluent (this should take <24 hr). 3. Just prior to use, mix equal volumes of recombinant vaccinia virus stock and 0.25 mg/ml trypsin and vortex vigorously. Incubate 30 min in a 37°C water bath, vortexing at 5- to 10-min intervals during the incubation. Virus stocks are usually at a titer of ∼2 × 109 pfu/ml but may be significantly lower depending on the source.
Production of Recombinant Proteins
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Vortexing usually breaks up any clumps. However, if clumps still remain, chill to 0°C and sonicate 30 sec on ice. Sonication can be repeated several times but the sample should be allowed to cool on ice between sonications.
4. Aspirate medium and infect monolayer culture at a multiplicity of infection (MOI) of 10 to 30 pfu/cell in 1 ml (final volume) complete MEM-5. Incubate 1 to 2 hr, rocking at ∼30-min intervals. If purified virus is used, sonicate 30 sec on ice prior to use (see Basic Protocol 1, step 7). If analyzing proteins that are secreted into the medium, it is important to infect and overlay the cells with medium that does not require the addition of serum or to use only 1% serum, since a high concentration of serum proteins interferes with gel analysis.
5. Overlay with 2 ml complete MEM-5 and incubate 24 to 48 hr. 6. Scrape cells and transfer to a centrifuge tube. Centrifuge 5 min at 1800 × g (2500 rpm in a H-6000A rotor), 5° to 10°C. Aspirate and discard supernatant. For secretory proteins, concentrate the medium by spinning in a Centricon 10 microconcentrator (Amicon) until the volume is reduced ∼5-fold. Alternatively, proteins can be precipitated from the medium by trichloroacetic acid (TCA) precipitation as follows: (1) Add Triton X-100 to the sample to 0.03% (v/v) final; (2) add an equal volume of 20% TCA; (3) place 15 min on ice; (4) microcentrifuge 15 min at maximum speed, 4°C; (5) aspirate supernatant and wash the pellet with 70% ethanol, (6) air dry the pellet, add 5 ìl 1 M Tris base, and dissolve in 1× SDS/sample buffer.
7. Resuspend cell pellet in 200 µl cell lysis buffer and transfer to a microcentrifuge tube. Vortex and let sit 10 min on ice. Cell lysis buffer contains nonionic detergent only. This may not be sufficient for extraction of some proteins. In such cases, SDS-containing buffer may be necessary.
8. Microcentrifuge 10 min at maximum speed, 4°C. Separate supernatant (cytoplasm) and pellet (nuclei). 9. To 20 µl of the supernatant, add 5 µl of 5× SDS/sample buffer and heat 5 min at 95°C. Dissolve pellet in 200 µl of 1× SDS/sample buffer and heat 5 min at 95°C. 10. Proceed with immunoblotting (UNIT 10.10). BASIC PROTOCOL 5
DETECTION OF EXPRESSED PROTEIN USING IMMUNOPRECIPITATION When strong late promoters are used, the expressed recombinant protein can be labeled with radioactive amino acids and detected by polyacrylamide gel electrophoresis. Immunoprecipitation of the labeled proteins can be used to increase sensitivity and specificity when early or late promoters are used. Materials Complete MEM-5 medium (UNIT 5.12) Complete methionine- or cysteine-free MEM-5 medium (see recipe in UNIT 5.12 but use methionine- or cysteine-free MEM, e.g., from Life Technologies) prepared with dialyzed FBS (e.g., HyClone) [35S]methionine or [35S]cysteine (depending on which amino acid was omitted from the medium; see Critical Parameters) Phosphate-buffered saline (PBS; APPENDIX 2E) Cell lysis buffer (see recipe)
Characterization of Recombinant Vaccinia Viruses and Their Products
Additional reagents and equipment for preparing cells and infecting with vaccinia virus (see Basic Protocol 4) and immunoprecipitation (UNIT 10.10) NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified.
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NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly. 1. Set up cell cultures and infect cells (see Basic Protocol 4, steps 1 to 4). 2. Overlay with 2 ml complete MEM-5 medium and continue incubation for the appropriate period of time. The time after infection at which metabolic labeling is performed will depend upon which promoter is used to express the gene of interest. If an early promoter only is used, labeling should be done between 1 and 6 hr post infection. If a late promoter is used, labeling should be done between 4 and 20 hr post infection. With a compound early/late promoter, either time may be used.
3. At the appropriate time after infection, aspirate medium and replace with 0.5 ml complete methionine- or cysteine-free MEM-5 containing 25 to 50 µCi of [35S]methionine or [35S]cysteine and 35 µl complete MEM-5. Incubate 2 to 3 hr to overnight. 4. Add 0.15 ml complete MEM-5 as chase. Continue incubation for 1 hr. 5. Remove medium and save if the expressed protein is secreted. If the medium is to be analyzed, remove any free cells by microcentrifuging 3 min.
6. Overlay cells with 1 ml PBS, scrape, and transfer to a microcentrifuge tube. Microcentrifuge 3 min at maximum speed, room temperature. Aspirate and discard supernatant. 7. Resuspend cells in 200 µl cell lysis buffer. Vortex and place 10 min on ice. 8. Microcentrifuge 10 min at maximum speed, 4°C. Remove supernatant and proceed with immunoprecipitation (UNIT 10.10). REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Cell lysis buffer 100 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 100 mM NaCl 0.5% (v/v) Triton X-100 or NP-40 Store indefinitely at room temperature Add PMSF (or other protease inhibitors; optional) from a 100× stock to 0.2 mM final just before use. Store 100× inhibitor stock at −20°C.
DNA extraction buffer 20 mM EDTA 20 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 0.5% (w/v) SDS 0.5 to 1.0 mg/ml proteinase K (add immediately before use) Store components without proteinase K indefinitely at room temperature Low-salt buffer 20 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 10 mM EDTA 0.75% (w/v) sodium dodecyl sulfate (SDS) Store indefinitely at room temperature
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PBS/Tween Add 0.5% (v/v) Tween-20 to PBS (APPENDIX 2E). Store indefinitely at room temperature. COMMENTARY Background Information Several different methods of characterizing recombinant vaccinia viruses are available. Each is suitable at a different time during isolation and characterization of a recombinant virus. During plaque purification of a new virus, PCR, DNA dot-blot hybridization, or immunoblotting of individual plaque isolates are quick and easy ways of determining which plaque contains the foreign gene. This is particularly important if TK selection only is used, since not all plaques will contain recombinant virus. After an isolate has been plaque purified several times and amplified, analysis of the DNA by PCR or Southern blotting can be used to determine whether or not a pure isolate has been obtained. Restriction endonuclease digestion of vaccinia DNA with HindIII gives a specific pattern. The HindIII fragment into which the foreign gene has been inserted should be increased in size by the size of the foreign gene (unless the latter contains a HindIII site). The TK gene, which is the most commonly used region for insertion of a foreign gene, is located in the 5.1-kbp HindIII J fragment. Western blotting or radioimmunoprecipitation using an antibody specific for the gene of interest will determine whether the correct gene product is made. Analysis of the extracellular medium will show whether the protein is secreted. If a nuclear localization signal is present, the protein may be translocated into the nucleus. In this case, the nuclei should also be analyzed (see Basic Protocol 4, step 8). For large-scale production of protein, HeLa cell suspension cultures may be infected as described in the support protocol for purification of vaccinia virus in UNIT 5.13, and harvested after 48 to 72 hr. Alternatively, microcarrier cultures of Vero cells may be used (Barrett et al., 1989).
Critical Parameters
Characterization of Recombinant Vaccinia Viruses and Their Products
In all protocols for identifying plaques containing recombinant virus, it is important to include positive and negative controls. For PCR or DNA dot-blot hybridization, the plasmid used for transfection is an appropriate positive control, while cells infected with wild-type
vaccinia virus can serve as a negative control. For immunoblotting, an extract of cells known to express the foreign gene can be used (e.g., influenza virus–infected cells could be used as a positive control for analysis of a recombinant vaccinia virus expressing an influenza gene). Cells infected with wild-type vaccinia virus can be used as a negative control. In planning an immunoprecipitation, check the amino acid composition of the expressed protein for numbers of methionine and cysteine residues and then label with the appropriate amino acid(s). If lysates are frozen prior to immunoprecipitation, it is important to microcentrifuge 5 min at maximum speed to pellet subcellular debris before mixing a sample of the supernatant with antibody.
Anticipated Results The Southern blotting protocol in this unit is a quicker, easier method than the one in APPENDIX 4G. Since the most common region used for insertion of foreign genes into vaccinia virus is the TK locus within the 5.1-kbp HindIII J fragment, the size of this fragment should be larger in a recombinant virus (if there is no HindIII site within the inserted DNA). Hybridization of a second blot or rehybridization of the original blot with 32P-labeled vaccinia virus DNA will show the overall HindIII restriction digestion pattern. The 5.1-kbp HindIII J fragment should be absent, indicating that the virus isolate does not contain any contaminating wild-type recombinant virus. A control of wildtype HindIII-digested vaccinia DNA is helpful for a comparison to the original restriction endonuclease pattern. Visualization of protein bands from an immunoblotting procedure is usually accomplished with an X-ray film exposure of 1 to 24 hr. Additional time may be required if the antibody recognition is poor; if this is the case, cells can be lysed in a smaller volume, allowing more lysate to be analyzed in one lane. Additional incubation time or a lower dilution of antibody may also help to intensify the signal. A western blot that has been exposed to X-ray film but has not been completely dried can be reincubated with the same antibody (if the original antibody
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concentration was too low) or with a different antibody. Negative and positive controls should be included to help identify specific protein bands. For an immunoprecipitation, the gel exposure time will depend on several factors, including the amount of label incorporated into the protein and the affinity of the antibody for the protein. It may be necessary to optimize results by adjusting the amount of labeled amino acid used, the amount of antibody used, or the time of incubation of the lysate with the antibody.
blot and immunoblotting protocols varies depending upon the hybridization probe or antibody used.
Literature Cited Barrett, N., Mitterer, A., Mundt, W., Eibl, J., Eibl, M., Gallo, R.C., Moss, B., and Dorner, F. 1989. Large-scale production and purification of a vaccinia recombinant-derived HIV-1 gp 160 and analysis of its immunogenicity. AIDS Res. Hum. Retroviruses 5:159-171. Zhang, Y. and Moss, B. 1991. Inducer-dependent conditional-lethal mutant animal viruses. Proc. Natl. Acad. Sci. U.S.A. 88:1511-1515.
Time Considerations The PCR, DNA dot-blot hybridization, and immunoblotting protocols require 2 days for amplification of the virus. An additional day is necessary for PCR amplification and agarose gel analysis or for hybridization and washing. Time of exposure to X-ray film for DNA dot-
Contributed by Patricia L. Earl and Bernard Moss National Institute of Allergy and Infectious Diseases Bethesda, Maryland
Production of Recombinant Proteins
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Gene Expression Using the Vaccinia Virus/ T7 RNA Polymerase Hybrid System
UNIT 5.15
This unit describes a transient cytoplasmic expression system that relies on the synthesis of the bacteriophage T7 RNA polymerase in the cytoplasm of mammalian cells. To begin, a gene of interest is inserted into a plasmid such that it comes under the control of the T7 RNA polymerase promoter (pT7). Using liposome-mediated transfection, this recombinant plasmid is introduced into the cytoplasm of cells infected with vTF7-3, a recombinant vaccinia virus encoding bacteriophage T7 RNA polymerase (see Basic Protocol 1). During incubation, the gene of interest is transcribed with high efficiency by T7 RNA polymerase. This transfection protocol is adequate for analytical purposes and is simple because no new recombinant viruses need to be made. For large-scale work, the pT7-regulated gene can be inserted into a second recombinant vaccinia virus by homologous recombination (UNIT 5.13) and used in combination with vTF7-3 to coinfect cells grown in suspension (see Basic Protocol 2) or used to infect OST7-1 cells (a stable cell line that constitutively expresses the T7 RNA polymerase; see Basic Protocol 3). Expressed protein is then analyzed by pulse-labeling (see Support Protocol) or by the methods detailed in UNIT 5.14, and purified as described in Chapter 6. There have been several innovations to this vaccinia virus/T7 RNA polymerase hybrid expression system (see Background Information). One new development is the VOTE inducible expression system, which eliminates the need to use two recombinant viruses or a special cell line (see Basic Protocol 4). NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly. LIPOSOME-MEDIATED TRANSFECTION FOLLOWING RECOMBINANT VACCINIA VIRUS (vTF7-3) INFECTION
BASIC PROTOCOL 1
This protocol begins with a plasmid vector containing the gene of interest at a site between pT7 and the T7 terminator (Figs. 5.15.1 & 5.15.2; also see Critical Parameters). This recombinant plasmid is then introduced via liposome-mediated transfection (UNIT 5.10) into cells already infected with vaccinia virus vTF7-3 (a recombinant virus expressing the T7 RNA polymerase gene). After harvesting, the cells are analyzed for expression of the gene product as described in UNIT 5.14 or by pulse-labeling (see Support Protocol). Materials Recombinant plasmid: gene of interest subcloned into pTF7-5 or pTM1 vectors (available from B. Moss, e-mail [email protected]; Figs. 5.15.1 and 5.15.2) or other plasmid containing the T7 promoter (e.g., pBluescript, Stratagene) Confluent CV-1 cell monolayer (ATCC #CCL70; UNIT 5.12) Complete DMEM-10 (APPENDIX 3C) vTF7-3 vaccinia virus stock (ATCC #VR-2153) Opti-MEM I reduced serum medium (Life Technologies) Liposome suspension (Lipofectin or TransfectAce; Life Technologies) 6-well tissue culture dishes with 35-mm-diameter wells Cup sonicator (385-W) Contributed by Orna Elroy-Stein and Bernard Moss Current Protocols in Protein Science (1998) 5.15.1-5.15.11 Copyright © 1998 by John Wiley & Sons, Inc.
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5.15.1 Supplement 14
MCS 807
pT7
Clal 737
T-T7 TKL Xbal 139 Hin dlll 1
TKR Xma lll 1589 pTF7-5 4600 bp
Scal 2870 pUC
multiple cloning site: pTF7-5: GGATCC BamHl
pEB2:
GATCGAATTCAGGCCTAATTAATTAAGTCGAC
EcoRl
Stul
stop
Sall
Figure 5.15.1 pTF7-5 (Fuerst et al., 1986) contains pT7 and a terminator, between which is a unique BamH1 site for insertion of a gene. The derivative, pEB2 (Berger et al., 1988) contains unique EcoRI and StuI sites followed by translation stop codons in all three reading frames. The expression cassette is flanked by segments of the vaccinia virus TK gene; thus, TK− selection can be used for isolation of recombinant plaques (UNIT 5.13).
12 × 75–mm polystyrene tubes (Falcon) Additional reagents and equipment for purification of plasmid (APPENDIX 4C) 1. Purify recombinant plasmid for transfection by CsCl/ethidium bromide centrifugation or PEG precipitation (APPENDIX 4C). Transfection will require 5 ìg DNA/well of cells. When using the pTM1 plasmid (Fig. 5.15.2), make sure that the initiator ATG of the gene of interest is the ATG of the pTM1 NcoI site (see Critical Parameters). Determine that the correct construction has been obtained by restriction mapping and DNA sequencing.
2. The day before infection, seed wells of a 6-well tissue culture dish with 5 × 105 CV-1 cells/well in complete DMEM-10. Incubate until cells are near confluency (usually overnight). Cells should be counted with a hemacytometer (APPENDIX 3C). Gene Expression Using the Vaccinia Virus/T7 RNA Polymerase Hybrid System
3. Vortex stock of vTF7-3 to disperse clumps. Sonicate the lysate using a cup sonicator as follows. a. Fill the cup with ice water (∼50% ice).
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Cla l 737
p T7
Hin dlII 1075 MCS 1423 -1487
EMC
TK L Xbal 139
T-T7 Hind lII 1
* pTM1 5357 bp TKR
Xma lII 2245
Scal 3628
pUC multiple cloning site: pTM1: CCATGGGAATTCCCCGGGGAGCTCACTAGTGGATCCCTGCAGCTCGAGAGGCCTAATTAATTAAGTCGAC Sal I BamHI NcoI Eco RI SmaI SacI SpeI PstI XhoI StuI stop Acc I Hin cII pTM3 same as pTM1 except it includes p 7.5 and gpt gene at* Figure 5.15.2 pTM1 contains the encephalomyocarditis virus untranslated leader region (EMCV-UTR) downstream of pT7. The NcoI site contains the translation initiation codon and should be used for insertion of the 5′ end of the protein-coding DNA segment. The 3′ end of the DNA can be inserted into any downstream sites preceding the T7 terminator. The expression cassette is flanked by segments of the vaccinia virus TK gene; thus, TK− selection can be used for isolation of recombinant virus (UNIT 5.13). pTM3 is the same as pTM1 except it includes the guanine phosphoribosyltransferase (gpt) gene to permit mycophenolic acid selection for recombinant virus (Moss et al., 1990; UNIT 5.10).
b. Place tube containing vTF7-3 stock in the ice water and sonicate at full power, 30 sec. c. Place lysate on ice ≥30 sec, then repeat sonication. Because sonication melts the ice, it is necessary to replenish the ice in the cup.
4. Dilute an aliquot of the virus to 2 × 107 pfu/ml in Opti-MEM I. 5. Infect subconfluent monolayer of CV-1 cells from step 2 with 0.5 ml/well diluted virus stock (i.e., 10 pfu/cell). Incubate 30 min, rocking every 5 to 10 min. 6. Approximately 5 min before the end of the infection, prepare liposome suspension. Place 1 ml Opti-MEM I in separate 12 × 75–mm polystyrene tubes (one for each well to be tested). Vortex liposome suspension and add 15 µl to the medium; vortex briefly. Add 5 µg recombinant plasmid (from step 1) and mix gently. Vortex the liposome suspension before use to suspend the liposomes, which gradually settle during storage. Do not use a polypropylene tube, as the DNA/liposome complex may adhere to the surface.
Production of Recombinant Proteins
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7. Aspirate virus inoculum from CV-1 cells and add the DNA/liposome complex directly to the cells. Incubate 5 to 24 hr, then analyze expressed protein by pulse labeling (see Support Protocol) or by methods described in UNIT 5.14. The time needed for incubation depends on the analysis that will be done. Although expression is detectable within a few hours after transfection, pulse-labeling at late times is preferred because host protein synthesis is inhibited and synthesis of the recombinant protein is increased (UNIT 5.11). BASIC PROTOCOL 2
COINFECTION WITH TWO RECOMBINANT VACCINIA VIRUSES The recombinant plasmid containing the gene of interest under control of pT7 (Basic Protocol 1) is incorporated into vaccinia virus by homologous recombination (UNIT 5.13) and a stock is made (UNIT 5.12). This stock is used together with vTF7-3 to coinfect cells in monolayer or suspension cultures. During incubation, the gene of interest is efficiently transcribed by T7 RNA polymerase and translated in the cytoplasm of the infected cells. The protein is then analyzed (UNIT 5.14) or purified (Chapter 6). This coinfection protocol is recommended for mass production of proteins. For this purpose, cells in suspension (i.e., Vero or HeLa S3 cells) are used. Materials vTF7-3 vaccinia virus stock (ATCC #VR-2153) Stock of recombinant vaccinia virus encoding the gene of interest under control of pT7 (UNITS 5.12 & 5.13) Confluent monolayer culture of CV-1 cells (UNIT 5.12) or HeLa S3 cells from a spinner culture (UNIT 5.12) 0.25 mg/ml trypsin (Worthington 2× crystalline and salt-free; filter sterilize and stored at −20°C) Complete DMEM-10, MEM spinner-5, or MEM-2.5 (depends on cell type; see below and UNIT 5.12) Sorvall H-6000 rotor or equivalent 6-well tissue culture dishes with 35-mm-diameter wells or larger tissue culture flasks for scaled-up analysis Additional reagents and equipment for counting cells with a hemacytometer (APPENDIX 3C), vaccinia virus analysis (UNIT 5.14), and protein purification (Chapter 6) Prepare the cells 1a. For infection of a CV-1 monolayer: The day before infection, seed wells of a 6-well tissue culture dish with 5 × 105 cells in complete DMEM-10. Incubate until cells are near confluency (usually overnight). Increase volumes proportionally if large-scale production is desired.
1b. For infection of HeLa S3 suspension cells: Just prior to infection, count cells using a hemacytometer (APPENDIX 3C) and centrifuge the desired amount 5 min at 1800 × g (2500 rpm in Sorvall H-6000 rotor), room temperature. Discard supernatant, resuspend cells in MEM spinner-5 medium at 2 × 107 cells/ml, and transfer to a smaller sterile tissue culture flask or Erlenmeyer flask. Gene Expression Using the Vaccinia Virus/T7 RNA Polymerase Hybrid System
2. Typsinize virus stocks. Place an aliquot of each stock in a separate tube and add an equal volume of 0.25 mg/ml trypsin. Vortex vigorously. Incubate 30 min, vortexing at 5- to 10-min intervals. The highest level of expression using this system is obtained with an MOI of 10 pfu/cell for each virus.
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Vortexing usually breaks up any clumps of cells. However, if any visible clumps remain, chill to 0°C and sonicate 30 sec on ice. Sonication can be repeated several times but the sample should cool on ice between sonications (see Basic Protocol 1).
3. Combine the two trypsinized virus stocks. Infect the cells For cells in monolayer culture 4a. Dilute trypsinized virus mixture with complete MEM-2.5 to 4 × 107 pfu/ml. This represents 2 × 107 pfu/ml from each virus stock.
5a. Aspirate medium from cells and infect wells of a 6-well dish with 0.5 ml virus mixture per well. Incubate 1 to 2 hr, rocking dish at 15- to 30-min intervals. 6a. Overlay cells with 2 ml DMEM-10 and incubate. Analyze gene expression 5 to 24 hr after infection, depending on the method used. Alternatively, harvest 2 to 3 days after infection for protein purification. Although expression is detectable within a few hours after infection, analysis by immunoprecipitation, immunoblotting, or northern blotting is preferred at late times because the T7-regulated gene product accumulates to a higher level. Earlier analysis may be preferred for assaying the biological activity of the expressed protein.
For suspension cells 4b. Add trypsinized virus mixture to the concentrated cells. Stir with a sterile magnetic stir bar 30 min at 37°C. 5b. Transfer infected cells to a larger vented spinner flask and dilute with MEM spinner-5 to bring the cell density to 5 × 105 cells/ml. Stir 1 to 3 days at 37°C. 6b. Harvest cells and analyze or purify the expressed protein (see annotation to step 6a above). INFECTION OF OST7-1 CELLS WITH A SINGLE VIRUS OST7-1 cells are derived from mouse L929 cells and constitutively express the bacteriophage T7 RNA polymerase in the cytoplasm. Thus, using this cell line eliminates the need for infection with vTF7-3. This protocol describes the infection of OST7-1 cells with a single recombinant virus containing the gene of interest.
BASIC PROTOCOL 3
Materials Confluent monolayer of OST7-1 cells (available from B. Moss, e-mail [email protected]) Complete DMEM-10 (APPENDIX 3C) containing 400 potent µg/ml Geneticin (G418, UNIT 5.10; Life Technologies; prepared from 80 mg/ml stock in PBS, filter sterilized and stored at −20°C) Stock of recombinant vaccinia virus containing the gene of interest under the control of pT7 promoter (UNITS 5.12 & 5.13) 0.25 mg/ml trypsin (Worthington 2× crystalline and salt-free; filter sterilize and store at −20°C) Complete MEM-2.5 (UNIT 5.12) 6-well tissue culture dish with 35-mm-diameter wells 1. Seed wells of a 6-well tissue culture dish with 1 × 106 OST7-1 cells/well in complete DMEM-10 containing 400 potent µg/ml Geneticin. Incubate until cells reach confluency (usually overnight). Complete DMEM-10 containing Geneticin is used to maintain the culture, but Geneticin is not required for the expression experiment itself.
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2. Mix an equal volume of the virus stock and 0.25 mg/ml trypsin by vortexing vigorously. Incubate 30 min at 37°C, vortexing at 5- to 10-min intervals. Use an MOI of 10 pfu/cell.
3. Dilute the trypsinized virus to 2 × 107 pfu/ml with complete MEM-2.5. 4. Infect confluent monolayer of OST7-1 cells with 0.5 ml virus mixture/well. Incubate 1 to 2 hr, rocking at 15- to 30-min intervals. 5. Add 1.5 ml/well complete MEM-2.5 and continue to incubate. 6. Analyze the cells 3 to 24 hr after infection (see Basic Protocol 2, annotation to step 6a). BASIC PROTOCOL 4
GENE EXPRESSION USING THE VOTE SYSTEM In this more recently developed method for vaccinia virus/T7 RNA polymerase hybrid expression, the gene of interest is cloned into the plasmid pVOTE.1 or pVOTE.2 and then incorporated into the genome of vaccinia virus vT7lacOI by homologous recombination (UNIT 5.13) and a stock is made (UNIT 5.12). Cells are then infected with the recombinant vaccinia virus in the presence of isopropyl-1-thio-β-D-galactoside (IPTG) and the gene of interest is efficiently transcribed by T7 RNA polymerase and translated in the cytoplasm of infected cells. Figure 5.15.3 illustrates this system. Materials Vaccinia virus vT7lacOI (available from B. Moss, e-mail [email protected]) DNA containing gene of interest pVOTE.1 or pVOTE.2 (available from B. Moss) CV-1 or BS-C-1 confluent monolayer (UNIT 5.12) Cells for protein expression (e.g., HeLa) Isopropyl-1-thio-β-D-galactoside (IPTG) Additional reagents and equipment for PCR (APPENDIX 4J), restriction enzyme digestion (APPENDIX 4I), preparation of vaccinia virus stock (UNIT 5.12), infection of cells with vaccinia virus, transfection of infected cells with a vaccinia vector, harvest of vaccinia virus, selection of recombinant vaccinia virus plaques, and plaque purification of virus (UNIT 5.13), and determination of protein expression in the vaccinia system (UNIT 5.14; also see Support Protocol in this unit) Insert gene into pVOTE.1 or pVOTE.2 1. Amplify the open reading frame from the gene of interest by PCR (APPENDIX 4J) so that the translation-initiation codon is part of an NcoI (CCATGG) site for cloning into pVOTE.1 or a NdeI (CATATG) site for cloning into pVOTE.2. The other end of the PCR product should be compatible with a unique restriction endonuclease site present in the MCS (XhoI, SacI, SmaI, EcoRI, PstI, SalI, or BamHI). For high expression, the translation start codon must be adjacent to the encephalomyocarditis virus ribosome entry site.
2. Digest the PCR product and either pVOTE.1 or pVOTE.2 with the chosen restriction endonucleases (APPENDIX 4I) and ligate the PCR product and plasmid together. Gene Expression Using the Vaccinia Virus/T7 RNA Polymerase Hybrid System
Transfer gene to vT7lacOI 3. Prepare a stock of vT7lacOI (UNIT 5.12) and use the stock to infect CV-1 or another cell line (UNIT 5.13).
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4. Transfect recombinant plasmid derived from pVOTE.1 or pVOTE.2 (from step 1) into the cells infected with vT7lacOI (from step 3). Harvest the virus. All of these procedures are described in UNIT 5.13.
5. Select recombinant virus for expression of XGPRT and plaque purify virus (UNIT 5.13). Because the gene is inserted into the hemagglutinin site of vT7lacOI, the cells infected with recombinant virus can be recognized with the light microscope by their formation of syncytia.
6. Prepare a stock of the plaque-purified virus (UNIT 5.12). Express the gene by infecting cells in the presence of IPTG 7. Infect monolayer or suspension cultures of HeLa or other cells with 10 pfu/cell of the recombinant virus and add 5 µM to 2 mM IPTG to the medium at the start of infection. 8. Determine expressed proteins by standard methods (UNIT Protocol in this unit).
5.14;
also see Support
Ð inducer EMC
lacO T7gene1
p L p E/L lacl
SLO p T7 p E/L gpt
TT target
active repressor
+ inducer
EMC
lacO T7gene1
pL pE/L lacl
TT target
SLO pT7 pE/L
gpt
target gene expression inactive repressor
Figure 5.15.3 Schematic representation of the VOTE expression system. Portions of a recombinant vaccinia virus genome containing regulatory elements are diagrammed. Transcription of the lacI gene under control of a vaccinia virus early/late promoter (pE/L) results in continuous synthesis of repressor monomers (represented by solid circles) which assemble into tetramers and bind to the lacO adjacent to a vaccinia virus late (pL) promoter, preventing transcription of the bacteriophage T7 RNA polymerase gene (T7gene1), and to the modified lacO (SLO) next to the T7 promoter (pT7), preventing transcription of the target gene should leaky synthesis of the T7 RNA polymerase occur. Upon induction, the repressor is inactivated, vaccinia virus RNA polymerase transcribes T7 gene 1, and T7 RNA polymerase is made. The latter binds to the T7 promoter and transcribes the target gene from the SLO to the triple terminator (TT). Translation of the target gene mRNA is enhanced by the EMC leader. From Ward et al., 1995; reprinted with permission of the National Academy of Sciences.
Production of Recombinant Proteins
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SUPPORT PROTOCOL
DETECTION OF EXPRESSED PROTEIN USING PULSE LABELING Pulse labeling infected cells is a quick and sensitive way to monitor expression of the T7-regulated gene relative to expression of vaccinia late proteins. When performed 24 hr post-infection under optimal conditions, up to 80% of the [35S]methionine is incorporated into the product of the T7-regulated gene. This protein is detected using SDS-PAGE. If the EMC untranslated leader is present (e.g., if the pTM1 plasmid is used), hypertonic conditions may be used to selectively enhance cap-independent translation. Additional Materials Infected cells expressing the desired T7-regulated gene of interest (see Basic Protocols 1, 2, 3, or 4) in a 6-well tissue culture dish Methionine- or cysteine-free, serum-free MEM (Life Technologies, Select-Amine Kit) 10 mCi/ml [35S]methionine (1175 Ci/mmol; Amersham) or 15 mCi/ml [35S]cysteine (600 Ci/mmol; Amersham) Phosphate-buffered saline (PBS; APPENDIX 2E), ice-cold Cell lysis buffer (UNIT 5.14) 6× SDS sample buffer (UNIT 10.1) Fixing solution: 50% (v/v) methanol, 10% (v/v) acetic acid, 40% H2O Fluorographic solution (EN3HANCE from Du Pont NEN; or Amplify from Amersham) Cell scraper 95°C water bath Additional reagents and equipment for denaturing (SDS) gel electrophoresis (UNIT 10.1) and autoradiography (UNIT 10.11) Label the cells 1. Obtain infected cells expressing the desired T7-regulated gene of interest 24 hr after infection. Aspirate medium and replace it with 0.3 ml methionine- or cysteine-free MEM per well. Incubate 20 min. Hypertonic conditions selectively enhance cap-independent translation. Therefore, if the EMC leader is present in the plasmid used in the infection (e.g., pTM1; Fig. 5.15.2), add 4.8 ìl of 5 M NaCl to each well (190 mM final).
2. Add 30 µCi of [35S]methionine or [35S]cysteine to each well. Incubate 30 min. Lyse the cells 3. Aspirate medium and add 1 ml ice-cold PBS to each well. 4. Scrape cells with a disposable cell scraper and transfer to a 1.5-ml microcentrifuge tube. Microcentrifuge 1 min at high speed. Aspirate and discard supernatant. 5. Resuspend cell pellet in 100 µl cell lysis buffer. Vortex briefly and incubate 5 min on ice. 6. Microcentrifuge 5 min at maximum speed and transfer supernatant to a clean microcentrifuge tube. Analyze proteins 7. Remove 10 µl lysate and place in another microcentrifuge tube with 2 µl of 6× SDS sample buffer. Heat 5 min at 95°C. Gene Expression Using the Vaccinia Virus/T7 RNA Polymerase Hybrid System
8. Load sample on single well of denaturing SDS-polyacrylamide gel and perform electrophoresis (UNIT 10.1). A 10% polyacrylamide gel is recommended for separation of vaccinia late proteins that will appear as background bands on the autoradiogram.
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It is important to use a control of infected cells that do not express the gene of interest.
9. Fix gel 20 min in fixing solution. 10. Incubate gel in fluorographic solution according to manufacturer’s instructions. 11. Dry gel and autoradiograph by exposing to X-ray film. If the pTM1 vector is used, the gene product is usually at a high enough level to appear as a major band among the vaccinia virus late gene products. If the T7-regulated gene product co-migrates with a vaccinia gene product, it will be necessary to analyze the expression using other methods as described in UNIT 5.14.
COMMENTARY Background Information The most efficient procedures for expression of genes in the cytoplasm of mammalian cells utilize a recombinant vaccinia virus encoding the bacteriophage T7 RNA polymerase. Many of the properties of T7 RNA polymerase that contribute to its successful use in prokaryotic gene expression (UNIT 5.2) are also relevant to its application for expression in eukaryotic cells. These properties include (1) the single subunit structure of T7 RNA polymerase, (2) the processive transcriptase activity of the enzyme, and (3) stringent promoter specificity in the eukaryotic cytoplasm. In one version of the hybrid expression system described in this unit, plasmid DNA containing a gene of interest under the control of the pT7 promoter is transfected into cells infected with the recombinant vaccinia virus, after which the gene is transcribed efficiently by T7 RNA polymerase (Fuerst et al., 1986). Alternatively, DNA can be transfected into a cell line that constitutively expresses T7 RNA polymerase in the cytoplasm. However, for reasons that remain unclear, high-level expression in this cell line still requires infection with wild-type vaccinia virus (even when the capindependent EMC virus untranslated region is used; Elroy-Stein and Moss, 1990). Therefore, the T7 RNA polymerase–cell line does not offer a clear advantage over the T7 RNA polymerase–vaccinia virus recombinant for transfection experiments. In the vaccinia/T7 system, mRNA derived from the transfected gene can be as much as 10% to 30% of total cytoplasmic RNA. However, although mRNA levels of the T7-regulated gene are often quite high, only moderate amounts of protein are made. This limited translation is presumably due to the low efficiency of capping the transcripts made by T7 RNA polymerase (Fuerst and Moss, 1989). This difficulty was addressed by incorporating
a feature of picornavirus mRNAs—a long untranslated leader region (UTR or “ribosomal landing pad”) that facilitates cap-independent ribosome binding—just downstream of the T7 promoter in certain vectors. The use of such vectors (e.g., pTM1; Fig. 5.15.2) in the vaccinia/T7 system enhances expression 5- to 10fold (Elroy-Stein et al., 1989). Transfection of DNA can be mediated by calcium phosphate or cationic liposomes. The latter method has reproducibly allowed 80% to 90% of the cells to be transfected (Felgner et al., 1987; Elroy-Stein and Moss, 1990; Rose et al., 1991). There have been several more recent innovations in the vaccinia virus/T7 RNA polymerase hybrid expression system. One is the use of the highly attenuated MVA strain of vaccinia virus to express the T7 RNA polymerase gene. Overall, the levels of expression of transfected plasmids are similar using vTF7-3 and MVA/T7pol, but one may be higher than the other in certain cells (Wyatt et al., 1995). The main advantages of MVA/T7pol over vTF7-3 are lower biosafety requirements and the expression of genes in the absence of virus replication in nonpermissive cells (UNIT 5.12). Stocks of MVA/T7pol are prepared as described for MVA in UNIT 5.12. Liposome-mediated transfection is carried out as described in Basic Protocol 1 of this unit. Recombinant vaccinia viruses that express SP6 RNA polymerase have been made, and are used in conjunction with plasmid vectors that use SP6 promoters for gene regulation (Usdin et al., 1993). Another innovation is the development of the VOTE inducible expression system (see Basic Protocol 4 and Ward et al., 1995). This system should supersede the two-virus system and the need for cell lines expressing T7 RNA polymerase (UNIT 5.15). The VOTE system is tightly regulated, because there is an E. coli
Production of Recombinant Proteins
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Gene Expression Using the Vaccinia Virus/T7 RNA Polymerase Hybrid System
lacO regulating the T7 RNA polymerase gene and another one regulating the T7 promoter adjacent to the gene of interest. The level of expression can be titrated with IPTG. Another version of the VOTE system has a thermolabile E. coli lac repressor, so that induction occurs by temperature elevation (Ward et al., 1995).
of infected cells (when the gene of interest is incorporated in a vaccinia virus) are observed to express the desired protein. If protein is being detected by pulse labeling, an overnight exposure should result in the protein product appearing as a dominant band over the background of vaccinia proteins.
Critical Parameters
Time Considerations
To maximize protein expression, the gene of interest is cloned into a plasmid vector (e.g., pTF7-5 or pTM1) that contains both pT7 and the T7 terminator (Fig. 5.15.1 & Fig. 5.15.2). Better expression (5- to 10-fold) is obtained by using the encephalomyocarditis virus (EMCV)-UTR downstream of pT7 in pTM1. Alternatively, any plasmid that carries the T7 promoter (e.g., pBluescript; see UNIT 5.1) can be used but may give lower expression levels. When the “ribosomal landing pad” within the EMCV-UTR is used, translation initiation occurs at the ATG, which is in the NcoI site of pTM-1. If native protein is to be expressed, the initiator ATG codon must be in the NcoI site. If necessary, the NcoI site may be mutated but the position of the initiator ATG should not be changed. If a fusion protein is desired, the coding sequence is inserted into one of the multiple cloning sites, in frame with the ATG of the NcoI site. For some methods of detection (e.g., immunofluoresence; Earl et al., 1990), it is preferable to harvest the cells ≤24 hr after transfection. If the incubation continues much beyond this point, the cells may detach from the monolayer and be lost during washing or staining. For analytical purposes, expression can be achieved by transfecting the recombinant plasmid into cells that have been infected with a recombinant vaccinia virus, such as vTF7-3, which expresses the T7 RNA polymerase gene. For mass production of proteins—e.g., 8 µg CAT/106 cells (Elroy-Stein et al., 1989) or 11 mg HIV envelope protein/liter (Barret et al., 1989)—a recombinant vaccinia virus that contains the pT7-regulated target gene should be generated by homologous recombination (UNIT 5.13). It should be introduced into cells constitutively expressing the T7 RNA polymerase, or used with vTF7-3 to coinfect any mammalian or avian cell line. Alternatively, the VOTE system eliminates the need for special cell lines or vTF7-3.
Starting with a plasmid containing the gene of interest cloned in the appropriate vector, the entire procedure from infection or infection/transfection to harvesting and analyzing the expressed product requires 2 to 3 days.
Anticipated Results In most cases, 60% to 90% of transfected cells (following infection with vTF7-3) or 99%
Literature Cited Barret, N., Mitterer, A., Mundt, W., Eibl, J., Eibl, M., Gallo, R.C., Moss, B. and Dorner, F. 1989. Large-scale production and purification of a vaccinia recombinant-derived HIV-1 gp160 and analysis of its immunogenicity. AIDS Res. Hum. Retroviruses 5:159-171. Berger, E.A., Fuerst, T.R., and Moss, B. 1988. A soluble recombinant polypeptide comprising the amino-terminal half of the extracellular region of the CD4 molecule contains an active binding site for human immunodeficiency virus. Proc. Natl. Acad. Sci. U.S.A. 85:2357-2361. Earl, P.L., Koenig, S. and Moss, B. 1990. Biological and immunological properties of human immunodeficiency virus type 1 envelope glycoprotein: Analysis of proteins with truncations and deletions expressed by recombinant vaccinia viruses. J. Virol. 65:31-41. Elroy-Stein, O. and Moss, B. 1990. Cytoplasmic expression system based on constitutive synthesis of bacteriophage T7 RNA polymerase in mammalian cells. Proc. Natl. Acad. Sci. U.S.A. 87:6743-6747. Elroy-Stein, O., Fuerst, T.R., and Moss, B. 1989. Cap-independent translation of mRNA conferred by encephalomyocarditis virus 5′ sequence improves the performance of the vaccinia virus/bacteriophage T7 hybrid expression system. Proc. Natl. Acad. Sci. U.S.A. 86:6126-6130. Felgner, P.L., Gadek, T.R., Holm, H., Roman, R., Chan, H.W., Wenz, M., Northrop, J.P., Ringold, G.A., and Danielsen, G.A. 1987. Lipofection: A highly efficient, lipid-mediated DNA transfection procedure. Proc. Natl. Acad. Sci. U.S.A. 84:7413-7417. Fuerst, T.R. and Moss, B. 1989. Structure and stability of mRNA synthesized by vaccinia virusencoded bacteriophage T7 RNA polymerase in mammalian cells. Importance of the 5′ untranslated leader. J. Mol. Biol. 206:333-348. Fuerst, T.R., Niles, E.G., Studier, F.W., and Moss, B. 1986. Eukaryotic transient expression system based on recombinant vaccinia virus that synthesizes bacteriophage T7 RNA polymerase. Proc. Natl. Acad. Sci. U.S.A. 83:8122-8126.
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Moss, B., Elroy-Stein, O., Mizukami, T., Alexander, W.A., and Fuerst, T.R. 1990. New mammalian expression vectors. Nature (Lond.) 348:91-92. Rose, J.K., Buonocore, L., and Whitt, M.A. 1991. A new cationic liposome reagent mediating nearly quantitive transfection of animal cells. Biotechniques 10:520-525. Usdin, T.B., Brownstein, M.J., Moss, B., and Isaacs, S.N. 1993. SP6 RNA polymerase containing vaccinia virus for rapid expression of cloned genes in tissue culture. BioTechniques 14:222224. Ward, G.A., Stover, C.K., Moss, B., and Fuerst, T.R. 1995. Stringent chemical and thermal regulation of recombinant gene expression by vaccinia virus vectors in mammalian cells. Proc. Natl. Acad. Sci. U.S.A. 92:6773-6777.
Wyatt, L.S., Moss, B., and Rozenblatt, S. 1995. Replication-deficient vaccinia virus encoding bacteriophage T7 RNA polymerase for transient gene expression in mammalian cells. Virology 210:202-205.
Contributed by Orna Elroy-Stein Tel Aviv University Tel Aviv, Israel Bernard Moss National Institute of Allergy and Infectious Diseases Bethesda, Maryland
Production of Recombinant Proteins
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Choice of Cellular Protein Expression System Over the last twenty years or so recombinant protein expression has become a routine laboratory tool, no longer an art form practiced by start-up biotechnology companies. Today, given the plethora of choices for producing proteins, there is even more need for the investigator to wisely consider the alternative expression and production strategies. Although it is very encouraging to see an intense Coomassie blue band by SDS-PAGE that corresponds to an expressed product of interest, that may be as far as the project progresses if the product cannot be extracted and purified from the cells. On the other, a band only visible by immunoblotting may represent a better product— being soluble, correctly folded, and easily purified by affinity chromatography. Widespread protein expression using Escherichia coli, Bacillus subtilis, yeast, and mammalian systems has led to the formulation of a simple generalized approach (for review, see Goeddel, 1990). It was suggested that, for expression purposes, heterologous proteins could be assigned to one of four broad categories. The first class covered small peptides (<80 amino acids), best expressed as fusion proteins in E. coli. The second class included polypeptides that are normally secreted proteins (80 to 500 amino acids) and that could be expressed in all systems, but preferably as a secreted product in yeast or mammalian cells. The third class consisted of very large (>500 amino acids) secreted and cell-surface proteins best expressed in mammalian systems. The fourth class was nonsecreted proteins (>80 amino acids) for which the choice of expression system depends heavily on the nature of the protein. A large number of proteins fall into this fourth class, and so, within this category, further considerations should be given to choosing a host system. In the late 1980s insect cell culture was also noted as a useful approach for obtaining small amounts of proteins possessing eukaryotic post-translational modifications. Another review (Marino, 1989) was based on making choices for production of proteins for commercial biotechnology application and raised points to be considered in selection of expression systems. First and foremost, the user should establish the complexity and quantity of protein required. Secondary criteria included in this 1989 review involved consideration of eco-
nomic and regulatory compliance issues, more of interest for commercial biotechnology applications. Later, a simple guide was offered for production of glycoproteins in Chinese hamster ovary (CHO) cells, insect cells, and yeasts cells (Hodgson, 1993). Since then, many more proteins have been expressed using optimized expression cassettes in various hosts. The focus of the choice has shifted from optimization of expression towards obtaining proteins with suitable post-transcriptional modification, relevant purity, and adequate yield. The desired aim of this unit is to provide a current guide for selecting the appropriate cellular-based expression system for obtaining proteins of suitable quality and quantity. Many systems are available, each with peculiar advantages and disadvantages that may be pertinent to the user’s project goal. In the majority of cases, the popular well-described systems will be suitable choices—i.e., E. coli (UNITS 5.1-5.3); baculovirus in insect cells (UNITS 5.4 & 5.5); yeast systems in Saccharomyces cerevisiae (UNITS 5.6 & 5.8) and Pichia pastoris (UNITS 5.7 & 5.8); mammalian CHO cells (UNITS 5.9 & 5.10); and transient expression in vaccinia (UNITS 5.11-5.15). Special considerations may be required in a few cases and are also briefly described.
BACKGROUND As a starting point in the choice of expression systems, the strategic parameters of greatest importance are the desired quality and quantity of the protein (Table 5.16.1). Quality encompasses not only protein purity, but also the need for and type of post-translational protein modifications. Quantity is a measure of the mass of purified protein required to meet the project goal.
Quality With respect to quality, the most important issue is the inclusion or exclusion of post-translational modifications of the protein. The most frequent issue to address is inclusion of glycosylation during expression of typical glycoproteins. The requirement for glycosylation is of major importance, since carbohydrate modification of proteins can influence important physicochemical properties such as solubility, stability, and protein folding. The various oligosaccharide additions also modulate key biological parameters such as specific activity,
Contributed by David Gray and Shyam Subramanian Current Protocols in Protein Science (2000) 5.16.1-5.16.34 Copyright © 2000 by John Wiley & Sons, Inc.
UNIT 5.16
Production of Recombinant Proteins
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antigenicity/immunogenicity, targeting, pharmacokinetics, and clearance (Goochee et al., 1991; Varki, 1993; Bhatia and Mukhopadhyay, 1998). Given the huge diversity of glycoforms found in natural proteins, it is at first difficult to ascribe the most appropriate expression host. Knowledge of post-translational modifications performed in the various eukaryotic hosts will
Table 5.16.1
Protein Quality and Quantity Considerations
Feature
Parameters or considerations
Glycosylation, low stringency
Chemical properties: Increased heterogeneity Increased diversity Physical properties: Resistance to proteolysis Resistance to denaturation Solution properties: Increased viscosity Increased solubility Protein folding: Minimizes aggregation Interacts with chaperones (quality control) Nucleates β turns In vitro activity: Altered protein-protein and protein-saccharide interaction In vivo activity: Changes in targeting Altered clearance Various modifications include phosphorylation, acylation, carboxylation, and hydroxylation. Covalent modifications are specific and modify in vivo activity and influence in vitro properties such as stability, solubility, and activity. Macroheterogeneity: >90% protein purity typically required Presence or absence of glycan at putative site (occupancy) Microheterogeneity (expression): Glycan variation N-terminal methionine incompletely removed (cytosolic) Amino acid misincorporation (norleucine) Incomplete proteolytic processing (eukaryotic) Microheterogeneity (processing): Met oxidation Cys oxidation Asp-Pro cleavage at low pH Deamidation Proteolysis Mixed disulfides
Glycosylation, high stringency
Other post-translational modifications (eukaryotic hosts)
Purity considerations
Choice of Cellular Protein Expression System
be useful in guiding the user to the appropriate system for the task at hand. In this unit, glycosylation is discussed within the context of host system from among those typical well-studied expression systems (see Post-Translational Modifications and Expression System Choice). The current major choices of systems for preparation of glycosylated proteins are yeasts, insect cells, and mammalian cells. Transgenic plants
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and animals can be used for the expression of recombinant glycoproteins, but these systems are not discussed, as they require significantly greater time investment and are not unicellular expression hosts, the subject of this discussion. Production of glycoproteins excludes E. coli, as it does not contain the cellular machinery for elaborate post-translational modification. Of course, the glycobiologist is aware and interested in the various prokaryotic oligosaccharides found in bacterial membranes, but these are not considered here.
Quantity The second important primary criterion is the amount of protein required (see UNIT 5.9, Table 5.9.4). This parameter dictates both the strategy and the scale of operation. A calculation of the overall yield should include projected purification yield as well as putative cell mass yield and product expression level. Quantity requirements will influence whether transient expression systems are appropriate, and will dictate the choice of scale of operation and the potential need to invest time for optimizing the bioreactor and purification steps. Expression in E. coli often results in accumulation of the foreign protein as cytosolic refractile inclusion bodies. The strategy for purification is discussed elsewhere (UNITS 6.1 & 6.3). A folding yield of 10% may be acceptable for laboratory-scale requirements and is commonly achieved without optimization for simple proteins. Poor overall protein recovery consequent to poor folding yield can be compensated for by increasing the scale of operation or by increasing the E. coli fermentation productivity (high cell density, increased expression level). S. cerevisiae has been successfully used to produce large quantities of recombinant proteins (UNIT 5.6; for additional review, see Barr, 1992). In addition to secretion of small proteins, yeast allow expression and secretion of multidomain proteins (>50 kDa) that are often poorly expressed in E. coli (e.g., Epstein-Barr virus major envelope glycoprotein; Schulz et al., 1987). Recently, higher levels of productivity, which are relevant to economical production of large quantities of protein, have been provided through use of the yeasts Pichia pastoris (UNIT 5.8) and Hansenula polymorpha (Faber et al., 1995). When very small quantities of proteins bearing post-translational modifications are needed (<50 µg quantity), it is quicker to use a transient expression system approach. Transient expres-
sion systems have been described for the Vaccinia virus expression system (UNIT 5.11), COS cells (Aruffo, 1998), and other transient mammalian expression hosts (UNITS 5.9 & 5.10). The insect cell–recombinant baculovirus expression method can be used to obtain moderate quantities of protein (500 mg) in stirred bioreactors in a relatively short time (UNIT 5.5). When larger quantities of protein are needed, stable expression in E. coli, yeast, or mammalian cells is required. Standard expression systems are the best choice for expression of the vast majority of heterologous proteins. Production of glycoprotein mandates a secretion expression strategy. Mammalian cell systems are required for producing glycoproteins with structures resembling the native counterparts. If complex glycosylation is unimportant to the application, secretion in yeast is simpler and quicker for obtaining glycoprotein. Alternatively, intracellular cytosolic expression in E. coli or yeast is often used for the production of proteins that do not require glycosylation.
EXPRESSION SYSTEMS The expression strategy used has an impact on the protein recovery and purification approach. In intracellular expression, protein can accumulate as either insoluble or soluble product. Often proteins with multiple native disulfides are slowly refolded during cytosolic expression, and will probably accumulate as refractile bodies in E. coli or as insoluble aggregates in yeast. When in vitro folding is inefficient, it may be expedient to take advantage of the secretory pathway offered in mammalian cell expression. The expression systems discussed here are the common systems that may be applied in a typical laboratory equipped with fundamental molecular biology tools, cell growth capabilities (shaken flask or fermentation systems), and protein recovery and purification equipment. These systems have been widely applied in heterologous protein expression and have a predictable range of performance based on historical perspective. This information provides the user with initial estimates of the potential titer and amounts of protein that can be obtained (Table 5.16.2).
Escherichia coli Despite the complexity involved in purifying proteins from inclusion bodies and the presence of endogenous cell membrane endotoxin, the use of the E. coli expression system is usually the most popular choice where post-
Production of Recombinant Proteins
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translational modifications are not required for biological specificity, activity, or other project goals. E. coli has been widely used for the expression of foreign proteins and is usually the best choice for expressing proteins with the following characteristics: single domain; Table 5.16.2
nonglycosylated; size <50 kDa; expressed as either soluble cytosolic folded protein or insoluble protein that can be extracted and folded with adequate yield. Some proteins require disulfide bonds for stable native conformation. Since the cyto-
Application of General Expression Systems
General host Strategy
Productivity (g/liter)
Expression (% TCP)a
Comments
E. coli
0.5-5
5%-25%
Common strategy
0.5-5
5%-25%
S. cerevisiae
P. pastorisb
Direct intracellular: inclusion body Direct intracellular: soluble Secretion
<0.5
Fusion
1-3
Membrane
<1
Secretion
<0.25
Direct intracellular: soluble
1-4
Direct intracellular: inclusion body Secretion
1-4 0.2-4
Direct intracellular
0.2-12
H. Secretion polymorphab
Mammalian cells
Insect cells (Sf9)
0.2-3
Direct intracellular Secretion
0.2-8 0.001-0.1
Intracellular
0.001
Baculovirus (AcMNPVc)
0.001-0.5
Minority of proteins accumulate as soluble cytoplasmic product 0.3%-4% (80% Periplasmic accumulation as soluble protein; purity) high purity in periplasm; some insoluble aggregates 5%-15% Increased expression stability; recent approaches promote formation of soluble cytoplasmic product; requires cleavage of fusion protein from product <5% Recently useful for production of some functional eukaryotic membrane proteins <1% (30%-60% Many examples of normally secreted proteins, purity in although potentially hypermannosylated medium) 5%-20% Proteins expressed as insoluble aggregates in E. coli may be expressed as soluble cytoplasmic products in yeast (FV111a) 5%-20% Aggregated inclusion body–like production occurs in many cases (HSA) High volumetric productivity resulting from 0.5%-10% high cell density (300-500 g wet cell/liter) (30%-80% purity in medium) 1%-30% High expression and cell density provide high productivity 0.5%-10% Similar to P. pastoris (30%-80% purity in medium) 1%-20% Slightly less than with P. pastoris <0.5% (1%-5% Low productivity, but complex in medium) post-translational modifications possible <0.1% Rare strategy, but may be applied for endoplasmic reticulum and membrane proteins <30% Provides quick method for obtaining a few hundred mg protein; post-translational modifications may be limited
a% TCP, total cell protein. bCommercial systems: Pichia from Salk Institute Biotechnology/Industrial Associates (SIBIA) and Philips Petroleum Company; Hansenula from Rhein
Biotech. cAcMNPV, Autographica californica nuclear polyhedrosis virus (most commonly used virus for baculovirus-based systems).
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plasm of E. coli is reducing, due to the presence of reduced glutathione (Hwang et al., 1995), these disulfide bonds do not form, and the incompletely folded protein tends to be less soluble and accumulates as inclusion bodies. Completion of folding in such proteins requires extraction from the cells, denaturation, and reduction followed by folding in vitro (Marston, 1986; Weir and Sparks, 1987; Ruldolph and Lilie, 1996). Direct intracellular expression in E. coli yields a protein sequence possessing an N-terminal methionine that may also bear an N-formyl group. Native proteins are normally processed by endogenous E. coli N-terminal methionine amino peptidase (MAP). In many cases, the N-terminal processing rate for recombinant proteins is limiting and N-terminal heterogeneity is observed. This has been discussed in the case of interleukin β (IL-β) expressed as a soluble cytosolic protein (UNIT 6.2). The presence of an N-terminal methionyl residue has been reported for recombinant proteins such as granulocyte/macrophage colonystimulating factor (GM-CSF; Burgess et al., 1987), IL-2 (Itri, 1987), and human growth hormone (hGH; Kaplan et al., 1986). Although the presence of the N-terminal methionine generally does not affect the activity of proteins (Hampsey et al., 1986), potential undesirable properties may occur as in the case of recombinant hGH, where a higher incidence of antibody response to methionyl-hGH compared to pituitary-derived hGH was reported (Kaplan et al., 1986). Heterologous proteins secreted into the periplasm have resulted in production of proteins without the N-terminal heterogeneity, as well as translocation of the protein to an oxidizing compartment that enables more rapid folding/oxidation of disulfide proteins (Missiakas et al., 1993). However, there are examples such as insulin-like growth factor 1 (IGF-I; Hart et al., 1994) where inclusion bodies form in the periplasm. Recently, various fusion-protein approaches have re-emerged in E. coli. These are aptly described in UNIT 5.1 and listed in Tables 5.1.1 and 5.1.2. The fusion or tagged approach enables rapid purification and may be a suitable route to obtain sufficient amounts of protein for characterization. To produce larger amounts by this approach for in vivo testing, the need for cleavage of the product from the fusion moiety should not result in deleterious protein modification. Thioredoxin-lymphokine fusions result in the expression of soluble intracellular protein (UNIT 6.7; La Vallie et al., 1993). Another current
fusion technique used in E. coli involves using pGEX vectors. Here the fusion expression of glutathione-S-transferase (GST) enables the rapid purification of large amounts of soluble protein using glutathione–Sepharose 4B as an affinity matrix (UNIT 6.6). The fusion approach has been useful for intracellular expression of small proteins that are rapidly proteolyzed. This was one of the earliest biotechnology strategies for expression of hormones and viral antigens (Kleid, 1983). E. coli can be useful for obtaining heterologous membrane proteins. Successful expression of functionally active membrane-based proteins, using endogenous short N-terminal fusions, has been reported for β-adrenergic receptors (Marullo et al., 1988) and transmembrane helix receptors (for review, see Schertler, 1992).
Yeast S. cerevisiae provides an expression host for both secretion and intracellular expression. The host is also advantageous for production of clinical protein products because no endotoxins are produced. The importance of yeast as a host system for production of recombinant proteins was established in 1982, when it was demonstrated that hepatitis B surface antigen (HBsAg) was expressed and recovered as antigenic particles (Valenzuela et al., 1982). S. cerevisiae naturally exports only a few proteins. They are α-factor, a-factor, killer factor, and Barrier activity protein. These proteins are generally small and nonglycosylated, except in the case of Barrier activity protein, which is a large (140 kD) complex glycoprotein. Typically, yeast has been regarded as a host for the production of small non-disulfide-containing proteins that are expressed in the cytoplasm, or for nonglycosylated disulfide-containing growth factors in the secretion pathway. In general, large proteins such as tissue plasminogen activator (tPA), urokinase-type plasminogen activator (uPA), and coagulation factors have been poorly secreted from yeast (for review, see Mackay et al., 1990) and are hypermannosylated. Proteins such as factor VII, factor IX, and protein C require post-translational modifications such as γ-carboxylation and βhydroxylation that have not been detected in yeast (Foster et al., 1987; Thim et al., 1988). Attempts to increase the secretory capacity of yeast by coexpression of the endoplasmic reticulum folding chaperones immunoglobulin-binding protein (BiP) and protein disulfide isomerase (PDI) appear to increase the secre-
Production of Recombinant Proteins
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Choice of Cellular Protein Expression System
tion titers of single-chain antibody fragments by 2 to 8 fold (Shusta et al., 1998). Secretion of bovine prochymosin was increased with BiP levels (Harmsen et al., 1996). The method does not seem to work for all secretion projects and, in fact, the secretion of factor VIII and Von Willebrand factor by animal cells was reduced at higher BiP levels (Dorner and Kaufman, 1994). Direct intracellular expression has been used for HBsAg (Valenzuela et al., 1982), functional antibodies (Wood et al., 1985), human superoxide dismutase (Hallewell et al., 1987), fibroblast growth factors (Barr et al., 1988), HIV-1 env and gag (Barr et al., 1987a; Bathurst et al., 1989), hemoglobin (Wagenbach et al., 1991), surface antigens of malaria parasites (Barr et al., 1987b), and α-antitrypsin (α1-PI; Travis et al., 1985). Intracellular expression of proteins as ubiquitin fusions in yeast is useful for the production of proteins with authentic N termini (Barr et al., 1991). This approach enables highlevel production of intracellular protein, whereas the secretion of proteins with processed N-termini from Saccharomyces may reach saturation levels, thus limiting production of authentic protein. Nevertheless, protein secretion in Saccharomyces has enabled production of fully active proteins that were poorly expressed in E. coli—e.g., epidermal growth factor (EGF), in which the three disulfide bonds were correctly formed (George-Nascimento et al., 1988). Native yeast proteins are, on the average, larger than those in E. coli and include native multidomain proteins. Thus, yeast would be expected to be a good choice of host for cytosolic production of large proteins (>45 kDa). S. cerevisiae may not be the optimal yeast host for large-scale operation or high-level protein expression. One drawback has been the lack of strong and regulatable promoter systems compatible with fermentation to high cell density. The achievement of high cell density usually requires carefully fed batch approaches to supply substrates for growth and maximal induction of the promoter (e.g., alcohol dehydrogenase/glyceraldehyde phosphate dehydrogenase or ADH/GAPDH; Suster, 1989). Large amounts of proteins can usually only be obtained after significant investments in fermentation development and host strain selection. The methylotroph P. pastoris system uses the tightly regulated alcohol oxidase promoter (AOX1). Addition of methanol as the carbon source induces expression using the AOX1 pro-
moter, enabling high levels of protein expression. P. pastoris has no native plasmids, thus the foreign gene and expression cassette must be integrated into the chromosome. The transformation, selection, and optimization of integrated gene copy number are time consuming. Similarly, Hansenula, using the methanol oxidase promoter (MOX), provides a useful system for obtaining high volumetric productivity. Hansenula provides an excellent host for secretion of large amounts of protein (Gellissen et al., 1992; Gellissen, 1994). The various yeast secretion systems are capable of processing propolypeptides exhibiting a KEX2 protease recognition site. For secretion of the desired protein into the growth medium, the DNA encoding the protein of interest is fused to DNA encoding a secretory signal sequence such as that derived from the yeast α-factor mating pheromone. Alternatively, the protein DNA is linked to a yeast signal peptide sequence or, if applicable, is expressed as a preproprotein form with its own signal peptide (for review, see Barr, 1992). The various expression strategies related to yeast expression are illustrated and discussed in Figure 5.16.1.
Mammalian Cells Mammalian cell production of complex multidomain proteins through the secretion pathway is the best method at present to produce biologically active protein. The initial and substantial time investment in such a project may be repaid by avoiding the need to deal with designing a folding step for such proteins. The main reasons to avoid the use of mammalian cells have been the low volumetric productivity of protein (typically, 1 to 100 mg/liter for recombinant proteins or 0.2 to 1.0 g/liter for recombinant antibodies), the high cost of media, and the large time investment. To overcome the lower productivity, largescale operation of bioreactors for suspension cultures is possible (UNIT 5.10). Attachment-dependent cells require microcarrier support media in order to obtain suitable volumetric productivity. Alternatively, dense cultures of attachment-dependent cells can be attempted in fixed-bed reactors to obtain a high product titer. In addition, plasmid vectors containing strong promoters, such as the cytomegalovirus immediate early (CMV IE) promoter, and dominant and recessive selection markers (neo and dihydrofolate reductase or DHFR, respectively) are available (UNIT 5.10, Table 5.10.1), and can enable higher productivity. Selection and amplification techniques have been developed, and
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A. Secretion promoter
signal pre-peptide (~ 20 amino acids)
Met
pro-sequence
heterologous gene
KEX2
signal peptidase
B. Secretion promoter
signal/leader peptide (~ 20 amino acids)
heterologous gene
signal peptidase
Met
C. Direct promoter
heterologous gene
Met
D. Fusion promoter
fusion partner
heterologous gene
Met
Figure 5.16.1 General representation of gene constructions for secretion or intracellular expression in yeast. Protein secretion (A, B) is the most common strategy used in rDNA expression using yeast. The most commonly used signal sequence is the yeast MFα-1 prepro region with an introduced XhoI cloning site towards the end of the pro region. The PHO5 and SUC2 signal sequences have also been useful. A few heterologous signal sequences have been successful in only a few cases. For heterologous gene expression, constructs are designed so that release of the mature polypeptide requires only the KEX2 cleavage event (as shown in A). In direct expression (C), the ubiquitin fusion approach enables enhanced expression stability and gives rise to mature authentic N-terminal heterologous protein through proteolytic cleavage by an endogenous yeast hydrolase (see Barr, 1992). In cases where the N-terminal amino acid is Pro, the cleavage is inefficient. Direct intracellular expression of a fusion protein (D) can be used to enhance levels (e.g., hSOD, α-IFN fusions).
most commonly the DHFR or the glutamine synthetase (GS) system is used (UNITS 5.9 & 5.10). Obtaining a high-producing line requires either laborious screening of many clones or stringent selection for marker expression (UNIT 5.10). The likelihood of obtaining a clone that is a high producer can be improved if the “position” effects are removed—i.e., the effects on transcription from the region of the chromo-
some where the expression cassette is integrated. Targeted integration of the expression cassette into “hot spots” (regions of high transcription), through homologous recombination using the Cre-lox system (Fukushige and Sauer, 1992) or matrix scaffold regions (Morris et al., 1997), may result in more predictable and higher levels of gene expression. Additionally, with respect to the positional effects of DNA
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integration, the productivity is usually not limited at the transcriptional or translational level if strong viral promoters are used, many cDNA copies are integrated (or few in a hot spot), and the mRNA is stable. This, of course, would have to be verified on a case-by-case basis to determine the cause of the bottleneck. In cases where mRNA levels are limiting, improved expression can be obtained by sodium butyrate induction or retransfection of cDNA (Fann et al., 1999). In most glycoproteins, the limitation is usually post-translational. Rate-limiting stages of protein production include folding, glycosylation, or other post-translational modification; subunit assembly; transport in the endoplasmic reticulum (ER) and Golgi; and secretion (Schroder and Friedl, 1997; Fann et al., 1999). Approaches to alleviate these problems, e.g., by manipulating levels of foldases such as PDI (Kitchin and Flickinger, 1995) and the chaperones BiP/GRP78 (Dorner and Kaufman, 1994), have not been successful to date. Today, technology exists for tailoring glycosylation of a protein through manipulation of a cell line. A custom continuous cell line can be created by transforming primary cells with desired glycosylation characteristics through specific knockouts or overexpression of genes involved in the regulation of the cell cycle. However, there are only a few such reports (e.g., PER.C6 for adenovirus production; Fallaux et al., 1998). This is probably due to a lack of understanding of the transformation process and its impact on cell characteristics, a reluctance to use uncharacterized cell lines, and time considerations. The currently available cell lines could also be further modified through expression of heterologous glycosyltransferases (Youakim and Shur, 1994) or isolation of mutants by using lectins (Stanley, 1992).
Insect Cells
Choice of Cellular Protein Expression System
Baculovirus-mediated expression has become a common approach for the rapid production of high levels of heterologous proteins in insect cells (UNIT 5.4). The main options for virus are the Autographica californica nuclear polyhedrosis virus (AcMNPV) and the Bombyx mori nuclear polyhedrosis virus (BmNPV). For cell culture applications, AcMNPV is the preferred choice due to the diversity of available transfer plasmids and the protein yield and growth properties of cells hosting AcMNPV. Insect larvae have been used for protein expression using BmNPV-based vectors. Larval production is more tedious than cell culture, but may be advantageous when higher efficiency
of post-translational modification is a prime requisite. The commonly used insect cell hosts are the Spodoptera frugiperda (fall armyworm) lines Sf9 and Sf21. These cells grow well (doubling times 18 to 22 hours) in both monolayer and suspension culture, and support the replicaton of AcMNPV. Other cell lines that support AcMNPV replication, such as the Trichoplusia ni–derived BTI-Tn-5B1-4 (“High5”), do not grow well in suspension culture, but may be better secretors of the recombinant protein than Sf9 cells. The insect cell approach falls into the category of transient expression. As such, it is capable of providing milligram quantities of recombinant proteins with desirable post-transcriptional modifications. Insect cells express high levels of heterologous proteins bearing complex post-translational modifications. These modifications include: correct proteolytic processing (signal-peptide cleavage and internal proteolytic cleavage); N-terminal blocking; phosphorylation; glycosylation; and oligomerization and assembly. As in any eukaryotic system, it is important to be aware that differences in post-translational modifications between a particular cell and the natural cell are likely. In insect cells, the high expression level and polh and p10 promoter activity in the very late phase of infection can lead to incomplete post-transcriptional modification. This is evident for phosphorylated proteins, where serine and threonine sites are incompletely phoshorylated as the efficiency of the processing becomes limited due to high expression level and very late-phase expression. Tyrosine phosphorylation may occur, but in some instances can require specific phosphotransferase enzymes that may not be present in the insect cell. Notwithstanding the potential limitations, insect cell culture has proven to be particularly useful for the expression of some phosphorylated proteins (Ramer et al., 1991; Sicheri et al., 1997; Xu et al., 1997). Methods for preparation of recombinant virus and identification of recombinant plaques are quite straightforward (UNIT 5.5; O’Reilly et al., 1994). The parameters for optimizing infection—multiplicity of infection (MOI), cell density, time—and the kinetics of infection for maximal protein production have been well studied. The growth medium can influence the output of this system, but the availability of commercial serum-free media enables serumfree production in any laboratory. Most posttranslational modifications are carried out by
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Table 5.16.3
Post-Translational Modifications of Proteins in Different Host Cellsa
Expression system Post-translational modification
N-Acetylation Amidation γ-Carboxylation of glutamate N-Glycosylation O-Glycosylation Heterodimer Hydroxylation Myristoylation Palmitoylation Phosphorylation Protein proteolytic processing Sulfation N-Terminal methionine removal
Insect cells Mammalian (baculovirus) cells
E. coli
Yeast
Yes No No No No Yes No Yes (cloned) No Yes E coli membrane protein signal sequence removal No Yes (and partial)
Yes Yes No Yes Yes Yes Yes Yes Yes Yes Yes
Yes Yes No Yes Yes Yes Yes Yes Yes Yes Yes
Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes
No Yes (and partial)
Yes Yes
Yes Yes
aAdapted from Yarranton (1992) and UNIT 5.9.
insect cells, and the quality of proteins in this regard can be considered second only to mammalian cells. Also, mammalian signal peptides are recognized. The overloading of the cellular protein-processing capacity of the insect cell is a common problem as the expression phase takes place late in infection. Overexpression of glycosyltransferases, foldases, and chaperones has been used fairly successfully in alleviating some of these problems (Jarvis et al., 1998; Ailor and Betenbaugh, 1999). Choice of the host insect cell may also be important in the selection process, as some cells, such as Estigmineae acrea (Ea), have a greater capacity for complex glycosylation (Ogonah et al., 1996). Transient expression in baculovirus is not the only available insect cell method. Stable expression of proteins is also becoming common in Sf9 cells as well as in whole larvae (Hollister et al., 1998; Farrell and Keshishian, 1999).
POST-TRANSLATIONAL MODIFICATIONS AND EXPRESSION SYSTEM CHOICE Native eukaryotic proteins contain various post-translational modifications of amino acid side chains that are required for correct in vivo biological function. Modifications to the amino acid side chains of proteins provide an additional level of diversity to protein function.
Given that the genetic code provides for only 20 amino acids, it is intriguing to consider that the numerous post-translational modifications add diversity by providing ∼200 distinct amino acid units (for review, see Krishna and Wold, 1998; also see Chapter 14). The first important decision in choosing an expression system is whether or not protein post-translational modification is required. This section covers the various types of posttranslational modifications obtained in various systems and the implications of such modifications on potential use of expressed proteins. The various choices of host cells available for heterologous expression may provide the necessary modifications (Table 5.16.3).
Glycosylation The most common issue is the need to obtain glycosylated proteins. The majority of proteins secreted by mammalian and yeast cells are glycoproteins. Carbohydrates are attached to proteins at asparagine residues in the case of N-linked glycosylation, or to serine/threonine residues in the case of O-linked glycosylation (see Chapter 12 for further information). The inherent complexity of glycosylation leads to the existence of diversity in structure within given proteins. Glycostructures produced in different cell types exhibit major gen-
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eral differences. Glycosylation diversity appears to be related to species-dependent and cell-to-cell variations in Golgi-based processing. For example, an N-glycosylation sequon of human interleukin 1β (hIL-1β) was occupied following expression in yeast, but unoccupied when expressed in human macrophage cells (Fleer et al., 1991). In this case, the glycosylation of hIL-1β led to a 95% decrease in biological activity. Yeast, insect cells, and mammalian cells are all capable of adding carbohydrate to protein; however, it is important to decide whether the project goals require stringent use of a specific host. The highest stringency will be required when a specific in vivo effect is to be pursued and will most often require use of a mammalian host system. When the emphasis is on in vitro protein stability, folding, or related physicochemical properties, the use of yeast or insect cell hosts will generally suffice.
Choice of Cellular Protein Expression System
N-Linked glycosylation The initial steps involved in the addition of N-linked oligosaccharides to proteins occur in the ER. Although the structure of the N-linked oligosaccharide (Glc3Man9GlcNAc2) and the initial removal of glucose (Glc) and mannose (Man) residues in the ER is the same in all eukaryotes, the structures subsequently added in the Golgi are very diverse (see UNIT 5.9 and Kornfield and Kornfield, 1985, for a more complete description of the N-linked glycosylation pathway). Based on the kind of Golgi processing, the glycosylation at any N-linked site can be classified as high mannose, complex, or hybrid. Complex glycosylation implies that residues such as galactose (Gal), N-acetyl glucosamine (GlcNAc), fucose (Fuc), or sialic acid (Sia) are added to Man5GlcNAc2 after the mannosetrimming step in the Golgi. In the high-mannose form, no new sugars are added after mannose trimming in the ER and Golgi (Man58GlcNAc2). Hybrid glycosylation is when one branch of the core is complex and the other is high mannose. Various commonly encountered glycan structures are illustrated in Figure 5.16.2. Mammalian systems. Mammalian cells would seem to be the safest choice for expressing a human therapeutic protein in order to maintain the correct glycosylation, folding, and other post-translational modifications. However, mammalian cell hosts are not all the same with respect to glycosylation capabilities, and it would be difficult to find a host capable of exactly simulating a human protein glycosyla-
tion pattern. To achieve this, human cell lines are the best choice, but not many such lines are available for large-scale expression. Time-dependent cellular substrate and enzyme variations occur in cells and result in normal variation of glycosylation within various cell types in the human body. To date, glycosylation has not been studied sufficiently in the available human cell lines (HEK 293, HT1080, Namalwa, human hybridoma; also see UNIT 5.9). Also, transformed human cell lines may have glycosylation features that are different from usual human glycoproteins (Laidler and Litynska, 1997). Most of our knowledge about glycosylation comes from analysis of recombinant proteins expressed in commonly used laboratory cell lines from other genera—e.g., CHO, NS0, baby hamster kidney (BHK-21), or C127—or from the analysis of proteins obtained from natural sources such as serum or urine. Host-based differences in the glycosylation of given proteins would be expected due to species and host variety of glycosyltransferases, each possessing defined specificities and present at differing levels in the various hosts. Further heterogeneity may be derived from protein substrate concentration. In many host cells, certain glycosyltransferases are mutated or not present, thereby eliminating certain sugars/linkages. The endogenous level of glycosyltransferases is not the only variable. For example, the expression level of various nucleotide transporters and the cytosolic concentration of substrates can also influence glycosylation patterns (Abeijon et al., 1997). Another difference between mammalian cell hosts is the type of sialic acid linkages. Humans contain both α26- and α2-3-sialyltransferase, whereas CHO and BHK-21 cells contain only the α2-3-sialyltransferase, and C127 cells only the α2-6 enzyme (Goochee et al., 1991). It is not clear whether these two types of linkages have a significant effect on protein properties. In fact, erythropoietin (EPO) produced in modified CHO cells, engineered to express α2-6-sialyltransferase, contains α2-6-linked sialic acids but shows lower bioactivity (Zhang et al., 1998). Several reports have shown that it is possible to tailor glycosylation in CHO, BHK21, and other mammalian cells by heterologous expression of glycosyltransferases, and to thereby create desired glycoforms that were not previously available (Youakim and Shur, 1994; Grabenhorst et al., 1995; Minch et al., 1995). NS0 myeloma cells (mouse cells) add Nglycolylneuraminic acid (NeuGc) in addition
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to the usual N-acetylneuraminic acid (NeuAc) to recombinant proteins. NeuGc is an oncofetal antigen that is not present in human adults. An increased level of NeuGc can result in an immunogenic response and rapid clearance (Jenkins et al., 1996). A low level of NeuGc is detected in CHO- and BHK-21-derived glycoproteins (Jenkins et al., 1996). In addition to the above-mentioned host cell lines, there are spontaneous mutants of CHO cells (screened using lectins) that have lost or gained some glycosylation function and can be used to synthesize high-mannose-type or other novel glycan structures (Stanley, 1992; Stanley and Ioffe, 1995). The Lec 3.2.8.1 mutant enables the production of proteins with high-mannose glycans at all glycosylated N-linked sites (Man5GlcNAc2) and with unsubstituted N-acetylgalactosamine (GalNAc) at O-linked sites (Stanley, 1992). Yeast systems. In yeast (UNITS 5.6 & 5.7), hypermannosylation causes concern for proteins destined for in vivo use (Herscovics and Orlean, 1993; Dean, 1999; Gemmill and Trimble, 1999). Trimming of one mannose from Man9GlcNAc2 occurs in the yeast ER. Yeast cells do not trim mannose residues in the Golgi, but they add further mannose units to form an inner core of Man9-15GlcNAc2. For secreted glycoproteins, the outer chain is formed by addition of mannose through α1-6 linkages to the lower arm. Branching of the outer chain occurs by the addition of α1-2-linked mannose residues. These α1-2 linkages are also decorated with terminal α1-3-linked mannose residues. The total number of mannose residues in outer chains can vary between 50 and 200, and can cause heterogeneity in the product. In contrast to animal cells, yeast do not add any sialic acid to the ends of their glycans, but may use phosphate (analogous to lysosome-targeted enzymes in mammalian cells), glucuronic acid, or pyruvate to add negative charge (Gemmill and Trimble, 1999). S. cerevisiae can add up to 200 mannose residues in the outer chain. Other yeasts such as P. pastoris have produced proteins without hypermannosylation (Miele et al., 1997; Montesino et al., 1998), but hypermannosylation may still occur for certain proteins (Martinet et al., 1997; Kang et al., 1998). The glycosylation mechanisms in H. polymorpha are largely similar to those of P. pastoris (Gemmill and Trimble, 1999). In addition, these systems do not add the immunogenic terminal α1-3-linked mannose residues. Several S. cerivisiae mnn and other mutants are available that produce proteins that are not hypermanno-
sylated, do not contain immunogenic mannose residues, and have homogeneous glycosylation (Ballou, 1990; Lehle et al., 1995). These mutants have weaker membranes and exhibit poorer growth compared to wild-type cells (Ballou et al., 1980). Genetic engineering may also make yeasts capable of producing therapeutic proteins. For example, galactosylation of glycoproteins, not normally observed in S. cerevisiae, has been made possible through coexpression of a galactosyltransferase and a nucleotide sugar transporter protein (Kainuma et al., 1999). A large number of eukaryotic glycosyltransferases have been cloned and can be found listed in a recent review (Sears and Wang, 1998). Insect cell systems. N-Linked glycosylation in lepidopteran insect cells results in mostly a trimannosyl core with or without fucosylation of the protein-proximal GlcNAc (Hooker et al., 1999; Kulakosky et al., 1998). Only a few cases of complex glycosylation have been observed (Davidson et al., 1990; Davidson and Castellino, 1991b; Wagner et al., 1996a; Hsu et al., 1997). Sialylation has been observed only with plasminogen and limited to the late phase of infection (Davidson and Castellino, 1991a). Complex glycosylation may also be species dependent. Insect cells such as Ea4 may have a greater ability for complex glycosylation than others (Ogonah et al., 1996). There is evidence that the apparatus for complex glycosylation does exist in insect cells (Velardo et al., 1993; Altmann et al., 1995). The glycosylation pattern of the protein depends on the interplay between glycosidase and glycosyltransferase activities, and thus their relative levels determine the outcome (van Die et al., 1996; Wagner et al., 1996a). The commonly observed trimannosyl core in glycoproteins expressed in insect cells is believed to be formed as follows (Jarvis et al., 1998). After α-mannosidase I action, the Man5GlcNAc2 substrate is acted upon by Nacetylglucosaminyltransferase, which adds a GlcNAc residue. This residue is apparently removed by a membrane-bound N-acetylglucosaminidase. The resulting structure is further trimmed by α-mannosidase II, resulting in the Man3GlcNAc2 structure The outcome of this pathway may depend on whether the protein is a bad substrate for the glycosidase or a good substrate for the glycosyltransferase under the conditions for expression (Jarvis et al., 1998). The use of immediate early (IE) baculovirus vectors instead of p10 and polyhedrin promoter-linked expression may yield less hetero-
Production of Recombinant Proteins
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A.
N-linked glycosylation precursor Manα1-2Manα1-6
Manα1-6
Manα1-2Manα1-3
Manβ1-4GlcNAcβ1-4GlcNAc-Asn
Glcα1-2Glcα1-3Glcα1-3Manα1-2Manα1-2Manα1-3
B.
Mammalian N-linked high- mannose and Lec 1 mutant Manα 1-2 Manα1-6 Manα1-6 Manα1-3
Manβ1-4GlcNAcβ1-4GlcNAc-Asn
Manα 1-2 Manα 1-2Manα1-3
C.
Mammalian N-linked hybrid
Manα 1-6
Manα1-6
Manα1-3
Manβ1-4GlcNAcβ1-4GlcNAc-Asn
NeuAcα 2-3 Galβ1-4GlcNAcβ1-2Manα1-3
D.
Mammalian N-linked complex biantennary NeuAcα 2-3 Galβ1-4GlcNAcβ1-2Manα1-6
Fucα1-6 Manβ1-4GlcNAcβ1-4GlcNAc
Asn
NeuAcα 2-3 Galβ1-4GlcNAcβ1-2Manα1-3
E.
Mammalian N-linked complex tetraantennary NeuAcα 2-3Galβ1-4GlcNAcβ1-2 Manα1-6 NeuAcα 2-3Galβ 1-4GlcNAcβ 1-6 NeuAcα 2-3Galβ 1-4GlcNAcβ 1-4
Fucα1-6 Manβ1-4GlcNAcβ1-4GlcNAc
Asn
Manα1-3
NeuAcα 2-3Galβ1-4GlcNAcβ1-2
Figure 5.16.2 (above and at right) Typical glycan structures obtained from various eukaryoytic hosts expressing heterologous proteins. In all eukaryotic cells, N-linked glycosylation is identical in the addition of the precursor, Glc3Man9GlcNAc2 to appropriate asparagine residues and the initial processing in the ER. The process diverges in the Golgi, resulting in various glycoforms. In mammalian cells, high-mannose, complex, or hybrid glycosylation is observed and is dependent upon the host cell, the environment, and the polypeptide itself. Hybrid and complex glycosylation are nonexistent in insect and yeast cells. Apart from mannose trimming, there is little processing in the Golgi in insect cells. In yeast cells, Golgi processing involves addition of mannose residues through various linkages resulting in hypermannosylation. Many mutants that have certain pathways added or removed from the glycosylation machinery yield different useful glycoforms. O-Linked glycosylation involves the addition of individual sugars to Ser/Thr residues and is initiated in the late ER or Golgi. It is specific and different from N-linked glycosylation in many ways. Many kinds of sugars present in complex glycoforms (with one notable exception being GalNAc) are added to the protein backbone in mammalian cells. Only mannose residues are added in yeast cells. Limited data is available for insect cells, but it is believed to be similar to mammalian cells, except for that there is no addition of sialic acid in insect cells. Key: Normal text indicates residues that are normally present; italic text indicates residues that may or may not be present. In F, X ≈ 10 for wildtype. In F and G, * indicates potential phosphorylation sites. References: (A) Kornfield and Kornfield (1985); (B-E,H,I) Goochee et al. (1991); (F,G,J) Ballou (1990).
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F.
Yeast N-linked wildtype
Manα1-2 —
Manα1-6* —
Manα1-3 —Manα1-6
outer chain
Manβ1-4G1cNAcβ1-4 GlcNAc—Asn
Manα1-6*—Manα1-6 —Manα1-6 —Manα1-6 —Manα1-6
Manα1-2
Manα1-2
Manα1-2
Manα1-2 —
Manα1-3
— —
Manα1-2 ManPO4α1-6 —
—
Manα1-3
Manα1-2*
—
—
Manα1-2
Manα1-2
— —
Manα1-2
— —
Manα1-2
Manα1-6 —Manα1-3 —
Manα1-2
— —
—
Manα1-2
—
—
—
Manα1-2
X
Manα1-3
Manα1-3
core
G.
Yeast N-linked mnn1 mnn9 mutant Manα1-2
—
Manα1-6* — Manα1-3 — Manα1-6
Man β1-4GlcNAcβ1-4GlcNAc — Asn
Manα1-6 — Manα1-3 —
—
Manα1-2
Manα1-2*
—
Manα1-2
H.
Insect N-linked
Fucα1-6
Manα1-6 Manβ1-4GlcNAcβ1-4GlcNAc
Asn
Manα1-3
I.
Mammalian O-linked
NeuAcα 2-3 Galβ1-3 GalNAcα1 – Ser/Thr
NeuAcα 2-6
J.
Yeast O-linked
Manα 1-3 Manα 1-3 Manα 1-2 Manα 1-2 Manα – Ser/Thr
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geneous glycosylation and probably allows complete glycosylation when the enzymes are present in low quantities. Another use of such vectors is to increase the levels of endogenous or heterologous processing enzymes that may not be expressed to high levels or may be a burden late in infection, as this could increase the efficiency of glycosylation or at least alter it (Jarvis and Finn, 1996). The use of promoters active in the late phase of infection overloads the glycosylation machinery due to the high rate of protein transit through the secretory pathway. This may be true for any post-translational modification in the insect cell system.
Choice of Cellular Protein Expression System
O-Linked glycosylation It is noteworthy that the expression of the mammalian glycosyltransferases, or the addition of substrates, can effect the glycosylation profile (Wagner et al., 1996b; Donaldson et al., 1999; Hollister et al., 1998). Sialylation in insect cells may be difficult to engineer, as both the substrate and the enzymes required are not normally present in insect cells (Hooker et al., 1999). O-Linked glycosylation in insect cells is similar to mammalian cells except that no sialic acid is added. As the potential for complex glycosylation does appear to exist in insect cells, it may be possible to engineer a more useful host by heterologous expression of glycosyltransferases and nucleotide sugar transporters and by supplying deficient substrates (Jarvis et al., 1998). O-Linked glycosylation is initiated in the late ER or Golgi with the transfer of GalNAc from the nucleotide sugar UDP-GalNAc to appropriate Ser/Thr amino acids (Van den Steen et al., 1998). No consensus sequence similar to N-glycosylation has been found. Sugars such as Sia, GlcNAc, GalNAc, Gal, and Fuc extend the chain. The glycosyltransferases in the Oglycosylation pathway are specific and different from the enzymes in the N-linked pathway. The O-linked glycosylation processing in yeasts is different from that of mammalian cells. The process is initiated in the ER through a lipid-linked Man donor and addition of up to four Man residues to form a linear chain. Occupancy of N-linked and O-linked glycosylation sites may not be similar between hosts (Van den Steen et al., 1998). Yeasts and mammalian cells may not O-glycosylate the same Ser/Thr sites. Thr-29 in IGF-1 is not usually O-glycosylated in human serum, but it is when produced in S. cerevisiae (Goochee et al., 1991).
Implications of Glycosylation Patterns on Protein Use In general, carbohydrate modification seems to make glycoproteins more soluble and resistant to proteases and increases the stability towards temperature and denaturants (Goochee et al., 1991). Invertase in S. cerevisiae is a good example, as unglycosylated and glycosylated forms are produced by the same organism. Resistance to thermal and denaturant inactivation was observed in the form with only the added core glycosylation (Kern et al., 1992). Protein folding, assembly, and secretion The correct folding, assembly, and secretion of proteins may also be dependent on glycosylation. Glycosylation can stabilize native-like protein conformation (Imperiali and Rickert, 1995). In invertase, the rate of folding was similar for deglycosylated and glycosylated protein, but competing processes such as intermediate aggregation greatly reduced the recovery of folded aglycosyl protein (Kern at al., 1992). Chaperones are involved in the prevention of aggregation of nascent proteins in the ER and allow correct folding to proceed. Calnexin and calreticulin are two chaperones that enable quality control of protein folding by binding to monoglucosylated proteins obtained after two glucose residues are trimmed by glucosidases. A glucosyltransferase acts as a sensor of folding and monoglucosylates the protein if the last glucose residue has been trimmed by α-glucosidase II and if the protein has not completely folded (Helenius et al., 1997; Trombetta and Helenius, 1998). Calnexin-mediated folding is required for some proteins such as factor VIII, HIV gp120, and hepatitis B virus middle (M) antigen (Kaufman, 1998; Trombetta and Helenius, 1998), but not for others such as tPA (Allen and Bulleid, 1997). The folding of the protein can in turn affect glycosylation. The N-linked site Asn-184 in tPA usually shows variable occupancy and becomes completely glycosylated when disulfide bond formation is inhibited using dithiothreitol (Allen et al., 1995). In general, core glycosylation seems to be sufficient for solubility, stability, and folding of glycoproteins. Specific activity of proteins The effect of glycosylation on the specific activity of proteins is very difficult to generalize and is dependent on the definition of activity. For stable proteins, in vitro activity is usually comparable or better in the absence of glycosylation. In vivo activity is difficult to assess
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per se, as the protein clearance rate differs with glycosylation pattern. In pituitary and placental glycoprotein hormones, the removal of glycans reduced effector activity toward target cells, although receptor binding was still apparent (Thotakura and Blithe, 1995). For EPO, GM-CSF, and tPA, in vitro activity increased with the removal of sugars (Moonen et al., 1987; Wittwer et al., 1989; Wittwer and Howard, 1990; Howard et al., 1991; Delorme et al., 1992; Higuchi et al., 1992). In EPO, the level of sialic acid was found to be inversely related to in vitro activity, and its removal abolished in vivo activity, probably due to hepatic clearance of asialoerythropoietin (Imai et al., 1990). Glycosylation regulates the activity of tPA through variable sequon occupancy at Asn-184. Native tPA is a single-chain molecule (sc-tPA) that can be cleaved by plasmin into a two-chain form (tc-tPa). Native sc-tPA also occurs as two glycoforms: type I tPA is glycosylated at Asn-117, Asn-184, and Asn-448, while type II tPA is glycosylated only at Asn-117 and Asn-448. Type I tc-tPA has lower activity than type II tc-tPA, and sc-tPA has a lower activity and susceptibility to inhibition than tc-tPA. Interestingly, the rate of conversion of type I sc-tPA into type I tc-tPA is half the rate of conversion of type II sc-tPA into type II tc-tPA. Therefore, the presence of glycosylation at Asn-184 hinders the conversion of sc-tPA to tc-tPA, thereby regulating activity (Wittwer and Howard, 1990). Antibodies have recently emerged as candidates for therapy in cancer indications. The Fc (CH2 domain) of IgG is usually glycosylated at Asn-237 with complex biantennary glycans. These glycans disrupt protomer-protomer interactions in their vicinity. This domain is responsible for antibody effector functions such as complement activation, antibody-dependent cell-mediated cytotoxicity (ADCC), and complement-mediated lysis (CML). Modification or removal of glycans does not affect antibody binding functions (Fab), but it reduces or eliminates effector functions associated with monocytes and macrophages of the ADCC and CML systems (Jefferis and Lund, 1997; Wright and Morrison, 1997). Recently, the creation of a bisected glycoform on an IgG1 through overexpression of GlcNAcT III in CHO cells resulted in increased ADCC (Umana et al., 1999). From such results it is evident that the modulation of antibody glycosylation affects in vivo function.
Effect on in vivo properties Certain features of glycoproteins may affect the persistence of a glycoprotein in vivo. The Galα1-3Gal moiety is not present in humans but is present in many glycosylated proteins in nonprimate cells (e.g., rodents). Between 0.5% and 1% of circulating IgG in humans can recognize this epitope (Galili, 1993). Mouse epithelial cells (C127 and mouse myeloma NS0) contain the epitope, while CHO and BHK-21 cells do not. No difference has been observed in the clearance of recombinant proteins such as factor VIII and tPA, which may contain this epitope (Tanigawara et al., 1990; Kaufman, 1998). Interestingly, a retroviral vector that does contain this moiety is inactivated through the classical complement pathway (Takeuchi et al., 1996). Terminal sugars influence in vivo properties of the glycoprotein, as these are surface exposed and thus can be recognized by cell receptors. There are a variety of carbohydrate receptors in hepatic and other cells. Clearance of glycoproteins from the circulation is brought about by receptor-based retrieval systems. Two major in vivo clearance mechanisms exist. One mechanism, present in the liver, specifically sequesters asialoglycoproteins. The second mechanism, present in liver endothelial cells and macrophages of the lung, liver, and spleen (Kupffer cells), involves the retrieval of mannose- and fucose-terminated glycoproteins. This receptor also has low affinity for GlcNAc, Glc, and Gal moieties. EPO is an example where sialic acid residues prevent clearance of proteins by the asialoglycoprotein receptor (Fukuda et al., 1989). Proteins secreted in S. cerevisiae are mannose-terminated and, therefore, are cleared rapidly. Even the presence of high-mannose type glycosylation in tPA (Asn117) from CHO cells results in enhanced clearance through the mannose receptor (Hotchkiss et al., 1988). In addition, terminal mannose (α1-3) residues render the protein immunogenic due to the presence of circulating antibodies against this moiety (Herscovics and Orlean, 1993). Glycoproteins produced in other yeast hosts, such as P. pastoris, and insect cells face the same problem with low in vivo halflives, but they lack the immunogenic Manα13Man moiety. Humans with Gaucher disease lack the enzyme glucocerebrosidase. Enzyme replacement therapy uses placental/recombinant glucocerebrosidase that is treated with exoglycosidases to expose Man/GlcNAc residues for targeting and increased uptake by macrophages
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(Brady et al., 1994). Thus, mannose-terminated proteins are capable of providing activity suitable for human therapy and disease control. Other physical clearance mechanisms, not directly based on carbohydrate structure, are also present in humans. Proteins of <70 kDa are normally filtered in the kidney. The cutoff is not absolute and is dependent on structure, charge, and molecular weight. Sialic acid can add charge, and sugars can significantly increase the hydrodynamic radius of the protein and thereby prevent renal clearance (Goochee et al., 1991). Covalent modification of proteins, such as PEGylation, can also increase size and prevent clearance (Knauf et al., 1988). Effect on protein antigenicity The effect of carbohydrates on protein antigenicity is better understood than is its effect on immunogenicity (for review, see Goochee et al., 1991). Circulating antibodies exist against blood group antigens, Manα1-3Man, and Galα1-3Gal. The presence of these epitopes on glycoproteins may elicit an antibody response or facilitate antibody-mediated clearance. Little data is available concerning the possible immunogenicity of a glycoprotein with a native amino acid sequence but non-native oligosaccharide structure. In the case of recombinant CHO-derived EPO, no antibodies were detected when administered in humans (Canaud et al., 1990). The well-studied example of EPO suggests the recombinant glycans are not necessarily immunogenic and are likely representative of variations that exist in vivo. It is not known exactly how glycosylation influences immunogenicity. Aglycosyl proteins can aggregate and be immunogenic (Moore and Leppert, 1980). Interestingly, PEGylation of IL-2 reportedly resulted in increased solubility and a decrease in nondesirable immunogenicity (Katre, 1990).
Choice of Cellular Protein Expression System
Effect of O-linked glycosylation The effect of O-linked glycosylation on protein properties is not well established but may provide functions similar to N-linked sugars. The importance of this modification for proteins such as proteoglycans and mucins, which show extensive O-glycosylation, is well known (Varki, 1993; Van den Steen et al., 1998). As O-linked sugars tend to be clustered, they serve as multivalent ligands for processes such as antibody binding, cell adhesion, and microbe binding (Hounsell et al., 1996). For EPO and factor VIII, O-linked glycosylation does not play a role in secretion and in vitro/in vivo
activity (Imai et al., 1990; Delorme et al., 1992; Higuchi et al., 1992; Kaufman, 1998). In the case of GM-CSF, the removal of the single O-linked sugar results in polymerization with subsequent loss of activity (Oh-eda et al., 1990). Effect of production system environment Analysis of the glycan structures of natural glycoproteins reveals micro- and macroheterogeneity (Rudd et al., 1994). This is to be expected due to the complexity of the glycosylation patterns in a given protein. Microheterogeneity, i.e., the variation in composition and structure of sugars at a particular glycosylation site, results through competition between various glycosylation pathways in the cell. Macroheterogeneity, i.e., the degree of occupancy of N-linked glycosylation sites, is caused by the interaction of the protein and the oligosaccharyltransferase complex during the folding process. For a particular polypeptide, heterogeneity in glycosylation is dependent on the host cell and the bioreactor environment. The effect of cell culture environmental parameters on the glycosylation profile is documented for the most part in mammalian cells (Goochee and Monica, 1990), but these results may apply to other expression systems as well. In any expression system, the depletion of precursors for glycosytransferase substrates (glucose and lipids for mammalian cells) will alter the glycosylation pattern (Jenkins et al., 1996). Induction or repression of glycosyltransferases and glycosidases through genetic or physiological means (hormones) can also affect the glycosylation pattern (Goochee and Monica, 1990). By-products of mammalian cell metabolism such as ammonia can also induce changes in the glycosylation pattern. High ammonia levels result in reduced sialylation and increased antennarrity of glycoproteins. Elevated ammonia levels in the cell culture medium can lead to increased local pH in the Golgi, which may lower the activity of sialyltransferases. Increased N-acetylhexosamine (UDP-GNAc) levels, which reduce substrate (CMP-NeuC) availability, can influence the glycosylation when high ammonia is present in the culture (Grammatikos et al., 1998; Zanghi et al., 1998). The serum level in media can also influence the glycosylation pattern of proteins (Jenkins et al., 1996). Glycosidase action in supernatants and cell lysates can result in the degradation of the glycans on the product. Various glycosidases have been detected for many cell types such as CHO, 293, NS0, and hybridoma cells
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(Gramer and Goochee, 1993, 1994). In CHO cell supernatants, the presence of a stable sialidase is a major problem, as the enzyme is active at neutral pH (Gramer et al., 1995). Prevention of glycoprotein desialylation through the use of competitive inhibitors such as sialic acid analogs has been successful (Gramer and Goochee, 1993). Recently it has been shown that feeding of sialic acid precursors like N-acetylmannosamine can improve sialylation (Gu and Wang, 1998). The maintenance of high cell viability in the bioreactor can reduce the release of glycosidases, thereby alleviating the problem. Given that it is improbable that a homogeneous glycosylation profile will be obtained, it is desirable that sequential preparations be consistent. Therefore, the glycosylation should be characterized and controlled by regulating the environment. In other words, reproducible production of glycoprotein requires control of the cell culture process (UNIT 5.10). Summary From the previous discussion of comparative glycoprotein production, it is clear that glycobiology is a complex field that makes choosing a host a daunting task. It is also difficult to predict the effect of particular glycosylation patterns on a protein’s physical and biological characteristics. However, there are known cases that can help in choosing a host. In general, it is advantageous to have the native protein (human-like) as a reference to enable comparison of glycosylation structures to that of the recombinant protein. For most applications, the desired protein properties are high solubility, stability, and activity. For therapeutics, additional characteristics such as antigenicity/immunogenicity and pharmacokinetics are important.
Other Post-Translational Modifications In addition to glycosylation, specific modifications of amino acids and addition of lipids may be required for authentic protein production. These modifications include removal of N-terminal methionine, acylation, phosphorylation, sulfation, γ-carboxylation, β-hydroxylation, and protein subunit assembly (Table 5.16.3). Removal of N-terminal methionine Methionine amino peptidase (MAP) is present in all cell types. Specificity for removal of the initiator methionine is substrate defined through the residue adjacent to the N-terminal
methionine. The amino acids with the smallest radii of gyration are substrates for MAP, and the thirteen amino acids bearing large side chains are not (Tsunasawa et al., 1985). High-level cytoplasmic production of heterologous proteins may overburden the cellular MAP, so that a portion of the protein will bear N-terminal methionine. In prokaryotes, the deformylating enzyme rate may also become limiting (Sugino et al., 1980). The removal of the N-terminal methionine in eukaryotes or the formyl or formyl-methionine in prokaryotes does not appear to have any major effect on activity of proteins (Hampsey et al., 1986). N-Terminal heterogeneity may be circumvented by secretion or by the fusion protein approach. The fusion approach for intracellular production of N-terminus-processed proteins is achieved in yeast using the ubiquitin fusion protein strategy (for review, see Barr et al., 1991). Using this approach, high intracellular levels of ubiquitin fusion proteins can be produced and, as the initiating methionine is located at the N terminus of the ubiquitin, proteins with authentic N termini are produced. Ubiquitin-specific hydrolases act in vivo to release the desired protein. For yeast-secreted proteins, the normal processing of the secretion leader generally results in production of correctly N- terminally processed proteins. This is accomplished through the action of an endogenous subtilisinlike protease produced by the KEX2 gene. The protein KEX2 is a transmembrane calcium-dependent serine protease of the subtilisin-like proprotein convertase (SPC) family, with specificity for cleavage after paired basic amino acids. It is mainly located in the late Golgi. Growth hormone–releasing factor (GRF) and transforming growth factor α (TGF-α) were inefficiently processed by KEX2, giving rise to α-factor leader/peptide fusions observed in growth media. This problem was circumvented by coexpression of the KEX2 gene and recombinant protein gene (Barr et al., 1991). In mammalian cells it is believed that a similar endoproteinase, Furin, performs the cleavage of signal/leader sequences. Acylation Acylation involves addition of the acetyl or formyl alkyl groups or of fatty acids (myristate, palmitate) to the N-terminal amino group. NTerminal acylation of amino acid amine groups occurs in eukaryotes and prokaryotes. In prokaryotes, the formyl group is added posttranslationally through Met-tRNAfMet. The for-
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Choice of Cellular Protein Expression System
myl group is found almost exclusively in prokaryotes. Acetyl groups are found in eukaryotes and are added to the N-terminal amino group from acetyl CoA, catalyzed by acyltransferase. This transfer occurs onto the nascent polypeptide, and the reaction follows complicated rules involving other enzymes and is also influenced by the N-terminal amino acid itself. Greater than 95% of N-terminal acetylation examples feature Ala, Ser, Gly, Thr, or Met as the N-terminal amino acid. Some N-terminal modifications, such as N-terminal α-keto acids, are found and result from spontaneous chemical cleavage (van Poelje and Snell, 1990). Other non-acyl N-terminal amino modifications have been found. The N-terminal lysine amino groups may become glycosylated in the presence of high levels of reducing sugars through a Schiff’s base reaction followed by a stable Amadori rearrangement. This would be unusual in common expression strategies, but may be an important consideration when attempting stable formulation of a protein. The C terminus does not typically exhibit general significant post-translational modifications. A few specialized proteins undergo Omethylation (Clarke, 1992; Inglese et al., 1992) or O-ribosylation (Riquelme et al., 1979). Amidation of the carboxyl group occurs in small active peptide toxins and hormones. Acylation is not limited to the N-terminal amino acid. Lysines in the region of the N terminus can also be acylated. Addition of the acetyl group is through acyltransferase and removal of acetyl groups is achieved through the action of a deacetylase enzyme (for review, see Kuo and Allis, 1998). Protein acylation (UNIT 14.2) is naturally performed by yeast, insect cells, and mammalian cells. This modification occurs in some eukaryotic natural proteins, where the N-terminal acetylation of nuclear matrix proteins and histones are important regulatory processes. Acetylation at the N-terminal tails of core histones alters nucleosome and higher-order chromatin structure, aiding transcriptional elongation and facilitating the binding of transcription factors to the nucleosomes associated with regulatory DNA sequences (for review, see Davie, 1997). Acetylation is an energy-dependent process and is catalyzed by the enzyme N-acetyltransferase. This modification has been demonstrated in E. coli in expressing recombinant proteins, although it has been shown that these adventitious post-translational modifications are mostly undesirable. A recombinant human
basic fibroblast growth factor mutein, expressed in E. coli, exhibited ε-N-acetylation of three lysines, although there is no apparent functionality associated with this modification (Suenaga et al., 1996). In epithelial cells, keratin undergoes a number of post-transcriptional modifications, including acetylation of the Nterminal serine (Omary et al., 1998). In yeast, recombinant human superoxide dismutase underwent N-terminal acetylation and indicated that yeast would provide this modification for proteins normally acetylated in mammalian cells (Hallewell et al., 1987). Addition of hydrophobic moieties to proteins is important for protein-membrane interactions. Myristoylation (14:0) occurs cotranslationally at N-terminal glycine through an amide linkage, and palmitoylation occurs post-translationally at Cys, Ser, or Thr, and requires prior myristoylation. Myristate is exclusively linked through an amide bond to an N-terminal glycine and palmitate is bound by a thioester linkage predominantly at a cysteine residue. Acylation provides a lipophilic handle for attachment of protein to specific membranes. This is important in picono and retrovirus assembly, and in a few cases may be important in modulation of protein activity in the cytosol (e.g., p60src myristoylation; Kamps et al., 1985). E. coli does not naturally perform acylation, but N-myristoylation of a mammalian protein was reconstituted in E. coli by coexpression of the yeast N-myristoyltransferase (NMT1) gene and a murine cDNA encoding the catalytic (C) subunit of cAMP-dependent protein kinase (Duronio et al., 1990). This is an early example of a prokaryotic host with an engineered post-translational system. Phosphorylation Phosphorylation is a ubiquitous post-translational protein modification. It is usually reversible and involves the action of phosphorylases and phosphatases present in the host cell. It is important to many cellular functions (UNIT 13.1). In essence, phosphorylation controls biological function by bringing about structural changes that may stabilize protein-substrate complexes. The protein phosphate esters occur on the hydroxyamino acids Ser, Thr, and Tyr. The baculovirus system is particularly useful for production of phosphorylated proteins that may be candidates for structural analysis (Ramer et al., 1991; Sicheri et al., 1997; Xu et al., 1997).
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Sulfation Sulfation of Tyr residues occurs in the lumen of the ER and Golgi apparatus. In general, this modification occurs in cells of multicellular and not unicellular organisms. Since the mechanism is based in the secretory pathway, sulfate is found in many naturally secreted mammalian cell proteins. Sulfation and sulfate conjugate hydrolysis play an important role in metabolism and in the stabilization/destabilization of proteins (Niehrs et al., 1994), and are catalyzed by members of the sulfotransferase and sulfatase enzyme superfamilies. In vivo sulfation is an important deactivating, detoxification pathway. γ-Carboxylation A number of coagulation factor proteins that are synthesized in the liver require γ-carboxyglutamic acid residues within the N terminus of the mature protein. The attachment of the carboxyl group to glutamate is achieved through the activity of a vitamin K–dependent enzyme. Expression of recombinant factor IX in CHO cells resulted in high levels of extracellular product but with negligible specific activity. Addition of vitamin K to the conditioned medium resulted in detection of active factor IX. Purification of the active fraction revealed that the activity was associated with the population of γ-carboxyglutamate-modified protein (Kaufman et al., 1986). Whether or not a recombinant protein is expressed bearing all the desirable post-transcriptional modifications is an important issue. The faithfulness in execution of post-translational reactions is difficult to predict for glycoproteins. Studies of γ-interferon glycostructures from CHO, transgenic mice, and insect cells showed extensive differences (James et al., 1995). Faithful expression of a liver glycoprotein in yeast would expectedly be problematic, since the complex glycans of liver would not be added by the yeast, which will by nature add hypermannose. This ostensible host/protein incompatibility may be eliminated in the future by effective coexpression of processing enzymes along with the protein. As mentioned, myristoylation of a recombinant protein was achieved in E. coli by this strategy. The human kidney 293 cell produced protein C bearing all the required post-translational requirements, which included formation of γ-carboxylation at the first nine Glc residues in the sequence; β-hydroxylation of Asp-71; N-linked glycosylation at four glycosylation sites; disulfide bond formation; peptide bond cleavage to remove the
signal sequence, propeptide, and an internal dipeptide; and cleavages that yield the active serine protease. In other mammalian cell hosts, this was not accomplished or expression was very poor (Yan et al., 1990). The 293-produced product had higher anticoagulant activity than the plasma-derived protein C. Interestingly, the sialic acid content of the recombinant protein was lower than that of the native protein, and the fucose content was approximately five times higher. The recombinant protein also contained a unique nonreducing terminal trisaccharide, GalNAcβ1-4Fucα1-3GlcNAcβ1R. Heterologous expression of mature proteins may avoid the need to consider whether cellular proteolytic enzymes will correctly process a precursor or zymogen protein. These proteolytic maturation events are highly specific and necessitate the use of cell lines close to the primary natural cell line. Direct expression of the mature sequence is a first choice for proteins normally produced by cellular complex proteolytic maturation processes. The processing of signal sequences that yield a mature protein from a preproprotein is a general process in cells capable of protein secretion (yeast and mammalian cells). A number of important human proteins are derived from precursors that are cleaved at dibasic amino acid processing sites. Some well-studied examples exist for proalbumin (Rhodes et al., 1989), proinsulin (Davidson et al., 1988), and proopiomelanocortin (POMC), which is a highly complex polyprotein that, upon processing, gives rise to seven neuroendocrine hormones (reviewed in Thomas et al., 1988). The yeast KEX2 protein was capable of processing POMC to the mature proteins in vitro. It is also important to consider whether artificial post-translational modifications occur in overproducing recombinant proteins. Such undesirable modifications involve peptide chain extensions from termination readthrough in prokaryotes, truncations, unique reactions, and faulty amino acid incorporations. Interesting but unexpected post-translational derivatives have been reported that are unique to recombinant expression. A peculiar example is a trisulfide with an extra sulfur between the Cys-182 and Cys-189 residues in recombinant hGH produced in E. coli (Jespersen et al., 1994). Codon usage is an important consideration. In E. coli, the Arg codon AGA led to incorporation of Lys in IGF, whereas the CGT codon led only to incorporation of Arg. Incorporation of norleucine instead of methionine was reported in recombinant bGH when E. coli
Production of Recombinant Proteins
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were grown under conditions where high levels of norleucine were made and competed with methionine for incorporation (Bogosian et al., 1989). Elimination of the leucine operon or increasing methionine concentration prevented misincorporation.
PRODUCTION OF SOLUBLE PROTEINS BY HOST CELLS The secondary decisions for choosing a host system are centered on the desire to obtain the protein in soluble form when expressed intracellularly. When soluble protein is produced, the user is able to extract the active protein from the cell and obtain purified material through standard purification procedures. It is now well accepted that foreign proteins often accumulate as refractile bodies in E. coli and must be solubilized and refolded to generate activity. The folding step may be difficult to optimize for highly complex disulfide-rich proteins, and the folding yield of native structure can be very low (<10%). In such cases, it may be necessary to use larger quantities of cells to obtain sufficient protein through a “brute force” approach. Prior knowledge of protein folding provides information allowing the user to assess the ability to obtain suitable quantities of protein through a refractile body–based process. Knowledge of the folding yield, along with standard yield for extraction and purification steps, will enable the user to determine the overall yield of protein, which in turn dictates the scale of operation.
In Vitro Folding
Choice of Cellular Protein Expression System
Optimization of in vitro protein folding is required to obtain high yield of correctly refolded protein. Aggregation of folding intermediates occurs and competes with the folding pathway. Aggregation may become a significant limitation in attaining adequate yield of correctly refolded protein. Proteins that form multimers through strong subunit interactions, and small proteins that are naturally glycosylated and contain a high area of hydrophobic nonpolar surface, tend to have a greater occurrence of aggregation during folding. These proteins are likely to accumulate as insoluble aggregates when expressed in E. coli, and thus require that a folding step be developed. Development of an efficient folding step for a complex protein requires a significant investment of time. For complex proteins, a single set of folding conditions will likely be nonoptimal, given the sequential nature of the overall fold-
ing process. Also, in batch oxidative folding processes, where protein folding is initiated with dilution of the denaturant, the concentration of the components in the folding mixture will change with time and may become suboptimal during the overall process. Low yield results when the rate of aggregation of incompletely refolded intermediates is greater than the sequential folding rate. The in vitro rate of aggregation may be slowed through folding at low protein concentration, or by using a fed batch reactor to minimize intermediate concentration (Katoh et al., 1999).
In Vivo Folding In E. coli, in vivo protein folding depends on cell-based chaperones and chaperonins. In vitro folding of protein at low concentration can reduce overall aggregation; however, in vivo folding must proceed at cellular concentrations at which aggregation would be expected to dominate. During in vivo folding, the unfolded polypeptides expose hydrophobic side chains that may associate and result in the formation of aggregates. More advanced intermediates that precede the rate-limiting final stages of folding may give rise to aggregation (Mitraki et al., 1991). The existence of inclusion bodies consisting of aggregates of near-native protein may be explained through the aggregation of folding intermediates that accumulated in the later stages of intracellular folding (Fig. 5.16.3). Interferon-γ refractile bodies, examined by density and size, appeared to be formed of regular structure in comparison to the amorphous prochymosin inclusion bodies (Taylor et al., 1986). Also, cytoplasmic β-lactamase inclusion bodies observed by electron microscopy showed a regular substructure array (Bowden et al., 1991). The concentration of nascent, ribosome-associated polypeptide chains in E. coli is believed to be as high as 35 µM, assuming a uniform distribution (Ellis and Hartl, 1996). Their effective concentration would be higher as a result of molecular crowding and because the ribosomes are usually organized into polysomes. Molecular crowding occurs because macromolecules in the cell can have total concentrations of up to 340 g/liter, thus excluding other macromolecules (Zimmerman and Trach, 1991). This is predicted to cause an increase in molecular association constants by several orders of magnitude. Thus, the high success of in vivo folding is dependent on cellular-based systems that de-emphasize the aggregation problem.
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cleavage of leader sequence KEX2 endoplasmic reticulum mannose addition N
DNA
RNA
nascent polypeptide U
U
Golgi -mannose trimming -addition of complex carbohydrate
transport
secretion
folded polypeptide N
refolding intermediates X1 X2 Xn l N-terminal MAP association
cytosolic accumulation
aggregation
refractile bodies
ribosome
Figure 5.16.3 Schematic of in vivo protein folding and transport. Simplified representation of intracellular protein folding. Proteins destined for export (yeast and mammalian hosts) usually fold efficiently in the oxidizing ER with assistance from ER-resident chaperones (UNIT 5.9). Protein folding in the cytoplasm must proceed in a reducing environment, at a pH close to neutral. In E. coli, resident proteins may undergo chaperone-assisted folding, but heterologous proteins follow a physical folding pathway. Folding proceeds through a series of intermediates (X1 through Xn), which may associate reversibly with oligomeric intermediates. This is a physical process, as was described for in vitro folding (UNIT 6.4). Late-stage intermediates may associate and accumulate as aggregates that are not in reversible equilibrium. These aggregates form inclusion bodies. Some proteins exhibit rapid and stable folding, which leads to a lower steady-state concentration of folding intermediates. As a result of the lower concentration of intermediates, the rate of the side reactions (involving association and aggregation of intermediates) is much lower than the rate-limiting step in the folding pathway, and the folded protein accumulates as soluble protein in the cytoplasm. In yeast and mammalian cells, the rate of mRNA translation is 10- to 50-fold lower than that of E. coli, resulting in a lower steady-state concentration of nascent polypeptide and folding intermediates. A consequence of this is an increased probability that a cytoplasmic protein, insoluble in E. coli, will be soluble in a eukaryotic cytoplasmic expression system. See text for further discussion.
Complete protein folding requires the presence of a full protein domain (∼100 to 200 amino acid residues in length). Therefore, the nascent chain can only fold after the C terminus is disengaged from the ribosome. In a growing cell, a large population of emerging, aggregation-prone, nascent peptide chains must be maintained in a folding-competent conformation. This is achieved by the cotranslational binding of molecular chaperones to the nascent chain (Beckmann et al., 1990; Hendrick et al., 1993; Nelson et al., 1993; Frydman et al., 1994; Kudlicki et al., 1995). In E. coli, cytosolic folding occurs in a reducing environment at pH 7.4 to 7.8, which is suboptimal for folding disulfide-containing proteins. Thus, it would be expected that the overall folding rates would be limited by disulfide bond formation.
Several heat shock proteins of E. coli serve as chaperonins. These include DnaJ (Hsp70), Dnak (the main Hsp70 of E. coli), and the cofactor GrpE. These folding chaperones of E. coli have been recently reviewed (Hartl, 1996; Netzer and Hartl, 1998), and an abbreviated form relevant to this purpose is provided here. In E. coli, the chaperone system involves a number of stages: (1) binding of unfolded polypeptide to DnaJ, followed by binding of DnaK-ATP; (2) release of DnaJ with hydrolysis of ATP to ADP; (3) binding of GrpE to the unfolded polypeptide and release of ADP; and (4) ATP-dependent release of the polypeptide, which may then complete folding or undergo a further cycle of chaperone binding. A further mechanism, the GroEL-GroES system, acts to prevent aggregation of polypeptides of <55
Production of Recombinant Proteins
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Choice of Cellular Protein Expression System
kDa. The GroEL cylinder provides a cage that protects a single polypeptide chain and facilitates folding of single chains, thereby preventing aggregation (for review, see Netzer and Hartl, 1998). The eukaryotic cytosol contains the chaperonin TriC, which does not utilize a GroES-like function. Given that these chaperone systems exist, why is the formation of inclusion bodies so common for heterologous expression in E. coli? One explanation is that only a small fraction of E. coli proteins utilize the GroEL system in vivo. In fact, it is estimated that the chaperonin is responsible for only 2% to 7% of the protein synthesis flux (Ellis and Hartl, 1996; Lorimer, 1996). Further, only 10% to 15% of all cytosolic proteins of E. coli interact with GroEL, although this may increase to 30% after a brief heat shock (Netzer and Hartl, 1998). Hence, the majority of E. coli cytosolic proteins are folded independently of chaperonin intervention. The proteins that interact with GroEL have a distinct size of <55 kDa, resulting from spatial limitations within the chaperonin cage (Ewalt et al., 1997). The chaperonin cycle is ∼15 sec, which is the time required to synthesize an average E. coli protein. Pulse-chase experiments revealed that the vast majority of the rapidly released proteins are in the size range of 25 kDa or smaller. The larger substrates of between 25 and 50 kDa remain associated with GroEL for >100 sec on average, and must undergo several chaperonin cycles. Thus, the fraction of E. coli proteins using GroEL is limited by the low concentration of GroEL (3 to 4 µM) and the recycling required by larger polypeptides. Cytosolic E. coli protein synthesis can be categorized based on protein interactions with chaperonins (Ewalt et al., 1997). Class I proteins consist of small, rapidly folding proteins that fold independently of chaperones. Class II proteins, the majority of E. coli cytosolic proteins, exhibit an intermediate dependence on GroEL, although they may still fold rapidly and independently. Class III proteins are dependent on GroEL, fold slowly, and are predominantly larger proteins (up to 55 kDa). In E. coli, the majority of the larger proteins are membrane proteins, and only a small proportion of the endogenous proteins are rated as class III. The average size of E. coli proteins is 35 kDa, with only 13% exceeding 55 kDa. Approximately 60% of the 4300 proteins of E. coli are cytosolic. In contrast, eukaryotes have a larger proportion of large multidomain proteins that are >55 kDa. The average yeast protein is 53 kDa,
with 38% of the total number of proteins being >55 kDa (Netzer and Hartl, 1998). Large multidomain proteins are most often difficult to fold in vitro (Garel, 1992). Eukaryotic cytosolic chaperones (TriC) are present at low levels, appear to be specialized for folding of actin and tubulin, and have little importance in folding of heterologous proteins (Kubota et al., 1995; Lewis et al., 1996). However, in stress conditions the cytosolic chaperones may assist folding of a wider range of proteins. The high efficiency of eukaryotic protein folding may be a result of cotranslational “domain-wise” folding. Eukaryotic translation rates are 2 to 4 amino acids per second, which is only 10% of the E. coli rate. The protein “domain-folding” rate must be in synchrony with the translation rate for minimal accumulation of folding intermediates. With the lower translation rate in eukaryotic cells, it may be that aggregation of hydrophobic domains is minimized as a result of the lower steady-state concentration of potential aggregating intermediates. The slow rate of folding of these proteins coupled with the faster translation rate may account for the fact that larger proteins accumulate as insoluble inclusion bodies in E. coli. This is illustrated to an extent in Figure 5.16.1. For secreted protein, the folding process takes place in the ER. This environment is more optimal for protein folding and disulfide formation. PDI and prolyl isomerase assist in the folding of protein, and several chaperones ensure complete folding as the protein is translocated through the secretory pathway (UNIT 5.9). This method of producing recombinant protein is generally useful for producing active glycoproteins and has also provided a means for producing active recombinant human serum albumin (hSA), which was otherwise aggregated in cytosolic expression (Quirk et al., 1989).
PROTEIN-BASED FEATURES As not all proteins form inclusion bodies when expressed at high levels in E. coli, it is of interest to ask whether there are features of the host and/or protein that especially contribute to this phenomenon. Several reviews have dealt with this issue (Schein, 1989; Wilkinson and Harrison, 1991), but no clear picture arises, except that large (>50 kDa) multidomain proteins are usually insoluble, and this is exacerbated when the protein contains many unpaired cysteines. The latter is often true regardless of the protein size. It is probably safe to say that one cannot predict expression solubility based
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on primary sequence analysis, just as one cannot predict water solubility based on knowledge of a protein’s three-dimensional structure. However, if the authentic candidate protein has been well characterized in terms of its in vitro folding behavior, it might be possible to make some predictions. Also, as mentioned earlier, if a normally glycosylated protein is expressed in E. coli, the lack of carbohydrate can effect folding, especially by reducing the solubility of folding intermediates. The size issue requires a little more elaboration. Large multisubunit complexes with masses >1 × 106 Da can quite happily accumulate in E. coli; one of the best examples is the hepatitis core antigen (Wingfield et al., 1995). Many other more moderately sized oligomers, for example, tumor necrosis factor (TNF, a trimer), have been produced (Wingfield et al., 1987). The common feature here is that the monomeric unit is produced in a folded soluble state and that the monomers self-associate to form higher oligomers. An analogous scenario for a single-chain polypeptide is that separate folding domains form (as previously mentioned, ∼50 to 200 residues), followed by their intramolecular association into the parent molecule. All these processes can be derailed by aggregation at various steps in the folding/association pathways. Given that one can neither predict nor prevent the aggregation of most proteins in the host, the real question to ask is whether it is possible to fold the protein into its native conformation. If the full-length native protein has been expressed and there have been no unexpected post-translational modifications (proteolytic processing, deamidation, oxidation), the chances are high that protein from inclusion bodies can be folded. The yield of product tends to decrease with increasing size and the number of disulfide bonds that must be formed. If one is attempting to produce portions of the protein that typically correspond to defined domains, then insoluble products may be produced, which may not be capable of folding into native structures, although there are many examples where this has been done successfully. Another problem to be aware of is proteins that are produced as precursors, which must undergo specific proteolytic processing to form the mature product.
SPECIAL CONSIDERATIONS Expression A number of specialized systems in various host cells have been described. The various
special applications are summarized below in Table 5.16.5. In addition to these specialized expression systems, there are a number of process-based parameters that may bear influence on the strategy to be used. Growth of recombinant E. coli at suboptimal temperature (<28°C) may increase the proportion of soluble folded protein (Schein, 1989). This approach is worth attempting but may not be generally applicable, as the outcome is dependent on many interacting mechanisms including transcription rate, translation rate, inherent in vivo folding rate, and competing in vivo aggregation rates. When protein is required for structural studies and glycosylation is unimportant, fusion expression can be used to obtain soluble protein. This is described in UNITS 6.6 & 6.7 for GST and thioredoxin fusions. When glycosylation is the subject of study, the choice of host requires careful consideration. For stringent biological activity applications, it may be necessary to use a specialized host that is capable of providing complex glycosylation. In this category, the coexpression of particular glycosyltransferases (see review by Sears and Wang, 1998) may be necessary and will necessitate a large investment of effort and time. When it is important to obtain humanlike glycans, mammalian cell systems (generally CHO or 293 cells) must be used. Also, since CHO and BHK cells lack a functional copy of the α2-6-sialyltransferase, it may be worth engineering the host cell by co-cloning the gene for the enzyme (Minch et al., 1995). Similar approaches are capable of influencing the quality of glycosylation in insect cells (Ailor and Betenbaugh, 1999). Other related considerations include use of glycosyltransferase mutants (lec mutants), described by Stanley and Ioffe (1995), with added cloned enzymes. The lec mutants have proven useful in ascribing function relationships for various glycostructures such as the Lewis antigen (Lex) and the selectin (SLex) determinants found in specific natural N-linked glycans (Nishihara et al., 1994). For large-scale transient expression, the user may chose to use cell lines that are very similar to the normal source of the eukaryotic protein of interest. The dominant cells for this application are human embryonic kidney 293 cells (HEK 293), COS cells, and BHK cells. Largescale transient expression in mammalian cells for recombinant protein production has been succinctly reviewed recently (Wurm and Bernard, 1999).
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requires postyes translational modification
determine amount of pure protein needed
< 0.05 mg
complex sialyl >0.05 mg
requires authentic N terminus
transient expression
mammalian cell secretion
mannose
insect cell secretion
mannose
yeast cell secretion
yeast cytosolic ubiquitin fusion
yes
E. coli secretion (periplasmic)
no
E. coli direct cytosolic (soluble)
no express as soluble cytosolic protein
very low
express as insoluble cytosolic protein
E. coli fusion protein (soluble) >45k Da >45k Da
yeast direct cytosolic
<45k Da
E. coli direct cytosolic (refractile body)
prior or predictable acceptable protein folding yield
low molecular weight (<10 kDa) unstable
E. coli fusion protein (insoluble)
Figure 5.16.4 Decision flow chart for initial expression strategy choices. The main choices are E. coli, yeast, insect cell, and mammalian cell expression. The first major decision is whether a protein will require post-translational modification features such as glycosylation, phosphorylation, carboxylation, or acylation. When this is required, a eukaryotic host should be chosen. The need for complex glycans dictates a mammalian host or requires significant further investment for producing an engineered insect cell host. Where stringency of glycan structure is less important but the user wishes to examine general effects of glycan on protein function, a yeast or insect cell approach will be expedient. The second most important decision is the amount of protein required, and this dictates use of stable expression in systems capable of high productivity when very large amounts are needed. Determination of the amount of protein required includes the process yield calculation, which should include folding yield for inclusion body–based processes in E. coli. Other decision points are whether N-terminal methionine removal is of high importance; whether soluble expressed protein is desired; whether a fusion-affinity strategy is worthwhile; whether the protein can be folded efficiently; and whether the protein is >45 kDa and more likely to be expressed as active protein in a eukaryotic host. In some cases, specialized systems may be pertinent and are discussed in the text.
Choice of Cellular Protein Expression System
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Downstream Considerations When proteins are poorly expressed, either as intracellular or secreted products, it is often necessary to use additional purification steps to obtain highly purified protein with concomitant poor recoveries. For an intracellular product, the fusion expression approach may alleviate this problem through potential enhanced solubility and higher ease of recovery through ligand-affinity chromatography. For proteins secreted from mammalian cells, the initial purity may be increased by reduction of the serum concentration in the growth medium. This requires investing time to adapt the cells to growth and production at low serum levels (<1% v/v). For example, if a protein is secreted at 1 mg/liter in 10% serum (3 to 5 mg/ml protein), it will be present at <0.05% purity. Decreasing the serum concentration to 0.5% (v/v) would increase the purity 20-fold, assuming there is no cell lysis. In low-serum cell culture, the user should investigate the stability of the protein. Serum contains protease inhibitors, which will also be at lower levels as the serum is decreased. For the stringent production of glycoproteins, the activity of glycosidases in cell culture supernatants can result in product heterogeneity. This may be minimized by designing the cell culture process for high cell viability, thereby minimizing cell lysis. Glycosidase inhibitors can be added as a reagent to minimize glycoprotein degradation (Gramer et al., 1995).
Table 5.16.4
The ability to recover acceptable levels of activity from refractile bodies following denaturation and folding may be further hampered by the presence of a protease activity that cosediments with the inclusion bodies in centrifugation, and that is activated during the folding process (Babbit et al., 1990). Washing inclusion bodies with neutral detergents (Triton, Tween) may remove protease, thus minimizing the problem. When producing products for human clinical applications, several biosafety issues are important. In light of potential bovine spongiform encephalitis (BSE) contamination, process reagents should be sourced from non-animal organ origin. Governmental information on this important issue can be found at the U.S. Food and Drug Administration (FDA) Web site http://www.fda.gov, and at http://www.eudra.org for Europe. For applications requiring large quantities of pure protein (>100 g), the user should consider high-volumetric-productivity bioreactor strategies. Where precise glycan detail is not important, E. coli should be used. If some glycosylation is desired but strict human-like structure is not required, the yeast P. pastoris or H. polymorpha host expression systems have proven to be capable of >3 g/liter recombinant protein at high cell density.
CONCLUSIONS The early categorization of expression systems by Goeddel (1990) was a good basis for
Commonly Used Cellular Expression System Strategies
Host
Strategy
Comments
E. coli (UNITS 5.1 & 5.2)
Intracellular fusion
Strategy for obtaining small proteins that are unstable when expressed directly In few cases leads to soluble active protein; most often leads to inclusion body formation Performs most post-transcriptional modifications; small proteins and larger proteins secreted with hypermannosylation High expression level possible Can perform human-like post-translational modifications; can produce complex glycoprotein Production of proteins with glycosylation and other post-translational modifications; high level of expression; N-linked glycans are Man-trimmed but not galactosylated or sialylated Production of small amounts of protein; vaccinia can be used to transfect many mammalian cell hosts (not CHO)
Direct intracellular S. cerevisiae (UNIT 5.6)
Secretion
Intracellular direct Mammalian cells (CHO; UNIT 5.10) Secretion Insect cells (Sf9, Sf21, Tn5; UNIT 5.5)
Baculovirus vectors
Transient expression (UNITS 5.12 &
Vaccinia-HeLa
5.13)
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deciding an expression route. Some additional categorization is possible following later advances in the process sciences. The most important first decisions are whether complex and specific post-translational modifications are required and the amount of pure protein that is required for the project. A decision flow path is shown in Figure 5.16.4. In most cases, standard commonly used expression approaches (Table 5.16.4) can be applied. When stringent complex glycosylation is not important, small proteins (<8 kDa) can be produced by the secretion pathway in yeast to provide active protein or can be suitable candidates for the fusion
protein route in E. coli. Recent fusions in E. coli may also lead to improved purification by affinity methods and possible intracellular soluble expression. Large, typically secreted proteins are still best expressed in mammalian cells or in yeast. For typically nonsecreted proteins, the approach is still case dependent. Given that E. coli naturally produces cytosolic proteins that are <55 kDa, nonsecreted proteins that are >55 kDa are probably best expressed in eukaryotes. For production of gram quantities of nonglycosylated proteins, use of E. coli is the best and easiest strategy. For complex proteins
Table 5.16.5
Specialized Expression Strategies
Host cell
Strategy
Strains
Vectors
Comments
E. coli
Intracellular fusion
General (UNIT 5.2, Table 5.2.1) General (UNIT 5.2, Table 5.2.1)
See UNIT 5.1, Table 5.1.1 See UNIT 5.1, Table 5.1.2
Periplasmic (secretion)
BL21(DE3), RV308 (UNIT 5.2, Table 5.2.1)
pET26b (Novagen)
Membrane localization
pop 6510 (thr leu tonB thi lacY1 recA dex5 metA supE)
pAC1 (Boulain et al., 1986; Chapot et al., 1990)
Small unstable proteins stabilized in inclusion bodies Improved purification using ligand-affinity approach; often produces soluble cytosolic product Provides oxidizing environment that facilitates disulfide formation and folding; results in removal of N-terminal Met to give authentic sequence Examples for functional membrane protein production
Fusion tag
Low temperature (20°-25°C) S. cerevisiae
Cytosolic ubiquitin fusion
Various
P. pastoris
Secretion
UNIT 5.7, Table 5.7.2
Cytosolic H. polymorpha
Secretion
Cytosolic Transient expression
Medium
Various
Integration vectors (UNIT 5.7, Table 5.7.4); requires secretion signal UNIT 5.7, Table 5.7.2 Integration vectors (UNIT 5.7, Table 5.7.4) RB11 Integration vectors pRBMOXApro MF α-gene RB11 Integration vectors pRBMOXApro gene HEK 293, COS, Plasmids (mammalian BHK cell expression); viral vectors (Sindbis, Semliki Forest virus, adenovirus, vaccinia)
Can lead to increased solubility in cytosolic expression (Schein, 1989) Can achieve high expresson level with natural processing to remove N-terminal Met (Barr et al., 1991) High productivity
High productivity High productivity
High productivity Large-scale transient expression can yield 1-20 mg/liter in stirred bioreactors at 1- to 10-liter scale
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with poor folding yield, low overall recovery will necessitate scaling up the volume of the E. coli fermentation process beyond the typical 10-liter scale. When folding yield is less than satisfactory (<5%) and scaling up is not possible, an alternative strategy has to be pursued. For example, the use of yeast intracellular expression and yeast or mammalian secretion routes will directly provide active protein. Although some limited success has been reported for promotion of soluble expression in E. coli through manipulation of fermentation temperature, it is likely that changes in growth conditions or strains will not satisfactorily improve solubility for complicated multi-disulfide-containing proteins. In such cases, yeast or mammalian expression should be the first choice. In large-scale manufacturing, the advantages of the E. coli and yeast systems are yield and economy of production. Although E. coli is a rapid and high-productivity option, high levels of intracellular expression are attainable using the alternative yeast hosts P. pastoris and H. polymorpha. They offer the ability to obtain high cell density coupled to protein expression from strong tightly regulated promoters. Once fundamental molecular biology details are optimized (e.g., codon optimization, promoter, copy number), intracellular expression in these hosts can lead to high expression levels. Secondary criteria may, however, influence the decision of host choice as shown in Figure 5.16.4. These secondary criteria may lead the user to specialized expression systems (Table 5.16.5). One drawback in E. coli expression is a propensity for incomplete N-terminal methionine removal by MAP. Potential solutions to this problem are secretion into the periplasm in E. coli, direct fusion expression and cleavage, or the ubiquitin fusion intracellular expression approach in yeast. Secretion leads to correct N-terminal processing in most cases, although some yeast-secreted proteins were heterogeneous, due to rate limitations in the S. cerevisiae KEX2 protease processing. Increased stringency for very pure protein bearing complex glycosylation structures has led to a greater use of mammalian cell expression systems. This is an area where structure/function relationships are not easily defined. The carbohydrate moieties mediate molecular recognition in several instances, for example, antibody function, cell adhesion, virus infection, and cell receptor function. It is not possible to define a general motif for each of these interactions. If the exact effect or role of the sugar structure is unknown, the choice
of host system is empirical and has to be dealt with on a case-by-case basis. There are many examples of glycoproteins that have been produced in CHO cells. The murine glycan structures obtained appear to provide product with useful properties (increased circulation time in vivo, resistance to proteolysis, thermal stability). It is also clearly apparent that purified glycoproteins exhibit glycan heterogeneity that stems from variation in the cellular glycosylation apparatus and may be further diversified through extracellular environmental enzymatic activities. Attempts to obtain more human-like specific glycan structures of medical importance currently necessitate the use of human host cell lines or CHO glycosylation mutant cell lines (see Stanley and Ioffe, 1995). When glycoprotein with close resemblance to the natural form is required, the use of a host cell similar to the natural source is best. This may limit the amount of protein obtainable using a laboratory-scale procedure if such cell lines are hard to grow and expression levels are poor. Future developments will likely result in the production of host cell lines engineered for production of proteins with more authentic human-like glycoforms.
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Choice of Cellular Protein Expression System
Boulain, J.C., Charbit, A., and Hofnung, M. 1986. Mutagenesis by random linker insertion into the lamB gene of Escherichia coli K12. Mol. Gen. Genet. 205:339-348.
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Fukuda, M.N., Sasaki, H., Lopez, L., and Fukuda, M. 1989. Survival of recombinant erythropoietin in the circulation: The role of carbohydrates. Blood 73:84-89. Fukushige, S. and Sauer, B. 1992. Genomic targeting with a positive-selection lox integration vector allows highly reproducible gene expression in mammalian cells. Proc. Natl. Acad. Sci. U.S.A. 89:7905-7909. Galili, U. 1993. Evolution and pathophysiology of the human natural anti-α-galactosyl IgG (antiGal) antibody. Springer Semin. Immunopathol. 15:155-171. Garel, J.-R. 1992. Folding of large proteins: Multidomain and multisubunit proteins. In Protein Folding (T.E. Creighton, ed.) pp. 405-454. W.H. Freeman, New York. Gellissen, G. 1994. Heterologous gene expression in C1 compound-utilizing yeasts. Bioprocess Technol. 19:787-796. Gellissen, G., Weydemann, U., Strasser, A.W., Piontek, M., Janowicz, Z.A., and Hollenberg, C.P. 1992. Progress in developing methylotrophic yeasts as expression systems. Trends Biotechnol. 10:413-417. Gemmill, T.R. and Trimble, R.B. 1999. Overview of N- and O-linked oligosaccharide structures found in various yeast species. Biochim. Biophys. Acta 1426:227-237. George-Nascimento, C., Gyenes, A., Halloran, M., Merryweather, J., Valenzuela, P., Steimer, K.S., Masiarz, F.R., and Randolph, A. 1988. Characterization of recombinant human epidermal growth factor produced in yeast. Biochemistry 27:797-802. Goeddel, D.V. 1990. Systems for heterologous gene expression. Methods Enzymol. 185:3-7. Goochee, C.F. and Monica, T. 1990. Environmental effects on protein glycosylation. Bio/Technology 8:421-427. Goochee, C.F., Gramer, M.J., Andersen, D.C., Bahr, J.B., and Rasmussen, J.R. 1991. The oligosaccharides of glycoproteins: Factors affecting their synthesis and their influence on glycoprotein properties. In Frontiers in Bioprocessing II (S.K. Sidkar, M. Bier, and P. Todd, eds.) pp. 199-240. American Chemical Society, Washington, D.C. Grabenhorst, E., Hoffmann, A., Nimtz, M., Zettlmeissl, G., and Conradt, H.S. 1995. Construction of stable BHK-21 cells coexpressing human secretory glycoproteins and human Gal(β1 -4)GlcNAc-R α2,6-sialyltransferase. α2,6-linked NeuAc is preferentially attached to the Gal(β1-4)GlcNAc(β1-2)Man(α1-3)-branch of diantennary oligosaccharides from secreted recombinant β-trace protein. Eur. J. Biochem. 232:718-725. Gramer, M.J. and Goochee, C.F. 1993. Glycosidase activities in Chinese hamster ovary cell lysate and cell culture supernatant. Biotechnol. Prog. 9:366373. Gramer, M.J. and Goochee, C.F. 1994. Glycosidase activities of the 293 and NS0 cell lines, and of an
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Choice of Cellular Protein Expression System
Hooker, A.D., Green, N.H., Baines, A.J., Bull, A.T., Jenkins, N., Strange, P.G., and James, D.C. 1999. Constraints on the transport and glycosylation of recombinant IFN-γ in Chinese hamster ovary cells and insect cells. Biotechnol. Bioeng. 63:559572. Hotchkiss, A., Refino, C.J., Leonard, C.K., O’Connor, J.V., Crowley, C., McCabe, J., Tate, K., Nakamura, G., Powers, D., Levinson, A., et al. 1988. The influence of carbohydrate structure on the clearance of recombinant tissue-type plasminogen activator. Thromb. Haemost. 60:255-261. Hounsell, E.F., Davies, M.J., and Renouf, D.V. 1996. O-linked protein glycosylation structure and function. Glycoconjugate J. 13:19-26. Howard, S.C., Wittwer, A.J., and Welply, J.K. 1991. Oligosaccharides at each glycosylation site make structure-dependent contributions to biological properties of human tissue plasminogen activator. Glycobiology 1:411-418. Hsu, T.A., Takahashi, N., Tsukamoto, Y., Kato, K., Shimada, I., Masuda, K., Whiteley, E.M., Fan, J.Q., Lee, Y.C., and Betenbaugh, M.J. 1997. Differential N-glycan patterns of secreted and intracellular IgG produced in Trichoplusia ni cells. J. Biol. Chem. 272:9062-9070. Hwang, C., Lodish, H.F., and Sinskey, A.J. 1995. Measurement of glutathione redox state in cytosol and secretory pathway of cultured cells. Methods Enzymol. 251:212-221. Imai, N., Higuchi, M., Kawamura, A., Tomonoh, K., Oh-eda, M., Fujiwara, M., Shimonaka, Y., and Ochi, N. 1990. Physicochemical and biological characterization of asialoerythropoietin. Suppressive effects of sialic acid in the expression of biological activity of human erythropoietin in vitro. Eur. J. Biochem. 194:457-462. Imperiali, B. and Rickert, K.W. 1995. Conformational implications of asparagine-linked glycosylation. Proc. Natl. Acad. Sci. U.S.A. 92:97-101. Inglese, J., Glickman, J.F., Lorenz, W., Caron, M.G., and Lefkowitz, R.J. 1992. Isoprenylation of a protein kinase. Requirement of farnesylation/αcarboxyl methylation for full enzymatic activity of rhodopsin kinase. J. Biol. Chem. 267:14221425. Itri, L.M. 1987. Report of Roche recombinant human interleukin-2 clinical trials in cancer patients. J. Biol. Respir. Med. 6:220-221. James, D.C., Freedman, R.B., Hoare, M., Ogonah, O.W., Rooney, B.C., Larionov, O.A., Dobrovolsky, V.N., Lagutin, O.V., and Jenkins, N. 1995. N-glycosylation of recombinant human interferon-γ produced in different animal expression systems. Bio/Technology 13:592-596.
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Jarvis, D.L. and Finn, E.E. 1996. Modifying the insect cell N-glycosylation pathway with immediate early baculovirus expression vectors. Nature Biotechnol. 14:1288-1292.
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Jefferis, R. and Lund, J. 1997. Glycosylation of antibody molecules: Structural and functional significance. Chem. Immunol. 65:111-128. Jenkins, N., Parekh, R.B., and James, D.C. 1996. Getting the glycosylation right: Implications for the biotechnology industry. Nature Biotechnol. 14:975-981. Jespersen, A.M., Christensen, T., Klausen, N.K., Nielsen, F., and Sorensen, H.H. 1994. Characterization of a trisulphide derivative of biosynthetic human growth hormone produced in Escherichia coli. Eur. J. Biochem. 219:365-373. Kainuma, M., Ishida, N., Yoko-o, T., Yoshioka, S., Takeuchi, M., Kawakita, M., and Jigami, Y. 1999. Coexpression of α 1,2 galactosyltransferase and UDP-galactose transporter efficiently galactosylates N- and O-glycans in Saccharomyces cerevisiae. Glycobiology 9:133-141. Kamps, M.P., Buss, J.E., and Sefton, B.M. 1985. Mutation of NH2-terminal glycine of p60src prevents both myristoylation and morphological transformation. Proc. Natl. Acad. Sci. U.S.A. 82:4625-4628. Kang, H.A., Sohn, J.H., Choi, E.S., Chung, B.H., Yu, M.H., and Rhee, S.K. 1998. Glycosylation of human alpha 1-antitrypsin in Saccharomyces cerevisiae and methylotrophic yeasts. Yeast 14:371-381. Kaplan, S.L., Underwood, L.E., August, G.P., Bell, J.J., Blethen, S.L., Blizzard, R.M., Brown, D.R., Foley, T.P., Hintz, R.L., Hopwood, N.J., et al. 1986. Clinical studies with recombinant-DNAderived methionyl human growth hormone in growth hormone–deficient children. Lancet 1:697-700. Katoh, S., Sezai, Y., Yamaguchi, T., Katoh, Y., Yagi, H., and Hohara, D. 1999. Refolding of enzymes in a fed-batch operation. Proc. Biochem. 35:297300. Katre, N.V. 1990. Immunogenicity of recombinant IL-2 modified by covalent attachment of polyethylene glycol. J. Immunol. 144:209-213. Kaufman, R.J. 1998. Posttranslational modifications required for coagulation factor secretion and function. Thromb. Haemost. 79:1068-1079. Kaufman, R.J., Wasley, L.C., Furie, B.C., Furie, B., and Schoemaker, C. 1986. Expression, purification and characterization of recombinant γ-carboxylated factor IX synthesised in Chinese hamster ovary cells. J. Biol. Chem. 261:9622-9628. Kern, G., Schulke, N., Schmid, F.X., and Jaenicke, R. 1992. Stability, quaternary structure, and folding of internal, external, and core-glycosylated invertase from yeast. Protein Sci. 1:120-131. Kitchin, K. and Flickinger, M.C. 1995. Alteration of hybridoma viability and antibody secretion in transfectomas with inducible overexpression of protein disulfide isomerase. Biotechnol. Prog. 11:565-574. Kleid, D.G. 1983. Using genetically engineered bacteria for vaccine production. Ann. N.Y. Acad. Sci. 413:23-30.
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Marston, F.A.O. 1986. The purification of eukaryotic proteins synthesized in Escherichia coli. Biochem. J. 240:1-12. Martinet, W., Saelens, X., Deroo, T., Neirynck, S., Contreras, R., Min Jou, W., and Fiers, W. 1997. Protection of mice against a lethal influenza challenge by immunization with yeast-derived recombinant influenza neuraminidase. Eur. J. Biochem. 247:332-338. Marullo, S., Delavier-Klutchko, C., Eshdat, Y., Strosberg, A.D., and Emorine, L. 1988. Human β2-adrenergic receptors expressed in Escherichia coli membranes retain their pharmacological properties. Proc. Natl. Acad. Sci. U.S.A. 85:7551-7555. Miele, R.G., Nilsen, S.L., Brito, T., Bretthauer, R.K., and Castellino, F.J. 1997. Glycosylation properties of the Pichia pastoris–expressed recombinant kringle 2 domain of tissue-type plasminogen activator. Biotechnol. Appl. Biochem. 25:151157. Minch, S.L., Kallio, P.T., and Bailey, J.E. 1995. Tissue plasminogen activator coexpressed in Chinese hamster ovary cells with α(2,6)-sialyltransferase contains NeuAcα(2,6)Galβ(1,4)GlcNAcR linkages. Biotechnol. Prog. 11:348-351. Missiakas, D., Georgopoulos, C., and Raina, S. 1993. Identification and characterization of the Escherichia coli gene dsbB, whose product is involved in the formation of disulfide bonds in vivo. Proc. Natl. Acad. Sci. U.S.A. 90:7084-7088. Mitraki, A., Fane, B., Haase-Pettingell, C., Sturtevant, J., and King, J. 1991. Global suppression of protein folding defects and inclusion body formation. Science 253:54-58. Montesino, R., Garcia, R., Quintero, O., and Cremata, J.A. 1998. Variation in N-linked oligosaccharide structures on heterologous proteins secreted by the methylotrophic yeast Pichia pastoris. Protein Express. Purif. 14:197-207. Moonen, P., Mermod, J.J., Ernst, J.F., Hirschi, M., and DeLamarter, J.F. 1987. Increased biological activity of deglycosylated recombinant human granulocyte/macrophage colony-stimulating factor produced by yeast or animal cells. Proc. Natl. Acad. Sci. U.S.A. 84:4428-4431. Moore, W.V. and Leppert, P. 1980. Role of aggregated human growth hormone (hGH) in development of antibodies to hGH. J. Clin. Endocrinol. Metab. 51:691-697. Morris, A.E., Lee, C.-C., Hodges, K., Aldrich, T.L., Krantz, C., Smidt, P.S., and Thomas, J.N. 1997. Expression augmenting sequence element (EASE) isolated from Chinese hamster ovary cells. In Animal Cell Technology (M.J.T. Carrondo, ed.) pp. 529-534. Kluwer Academic Publishers, Boston. Nelson, R.J., Ziegelhoffer, T., Nicolet, C., WernerWashburne, M., and Craig, E.A. 1993. The translation machinery and 70 kd heat shock protein cooperate in protein synthesis. Cell 71:97-105. Choice of Cellular Protein Expression System
Netzer, W.J. and Hartl, F.U. 1998. Protein folding in the cytosol: Chaperonin-dependent and -inde-
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Production of Recombinant Proteins
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Characterization and novel purification of recombinant human protein C from three mammalian cell lines. Biotechnology 8:655-661. Yarranton, G.T. 1992. High-level expression in Escherichia coli. In Transgenesis: Applications of Gene Transfer (J.A.H. Murray, ed.) pp. 1-29. John Wiley & Sons, New York. Youakim, A. and Shur, B.D. 1994. Alteration of oligosaccharide biosynthesis by genetic manipulation of glycosyltransferases. Ann. N.Y. Acad. Sci. 745:331-335. Zanghi, J.A., Mendoza, T.P., Schmelzer, A.E., Knop, R.H., and Miller, W.M. 1998. Role of nucleotide sugar pools in the inhibition of NCAM polysialylation by ammonia. Biotechnol. Prog. 14:834844. Zhang, X., Lok, S.H., and Kon, O.L. 1998. Stable expression of human α-2,6-sialyltransferase in Chinese hamster ovary cells: Functional consequences for human erythropoietin expression and bioactivity. Biochim. Biophys. Acta 1425:441452. Zimmerman, S.B. and Trach, S.O. 1991. Estimaton of macromolecule concentrations and excluded volume effects for the cytoplasm of Escherichia coli. J. Mol. Biol. 222:599-620.
INTERNET RESOURCES http://www.fda.gov U.S. Food and Drug Administration Web site. Provides governmental information on replacing animal-derived products with non-animal products when using expressed protein for human clinical applications. http://www.eudra.org European site providing similar information.
Contributed by David Gray Chiron Corporation Emeryville, California Shyam Subramanian Merck Research Laboratories West Point, Pennsylvania
Choice of Cellular Protein Expression System
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Use of the Gateway System for Protein Expression in Multiple Hosts Current biology depends to a large extent on cloned DNA molecules. Hundreds of plasmid and viral vectors have been described that have facilitated many advances in biomedical science. A large fraction of these vectors are designed for expressing a gene in some new context—for example in a heterologous host, or at levels that enable purification. An obstacle to the use of this resource is the task of transferring a gene of interest into new vectors using the traditional tools of restriction enzymes and ligases, which is often cumbersome and time consuming. The Gateway cloning method (Invitrogen) allows a gene to be cloned and subsequently transferred into any vector by in vitro site-specific recombination. It does not necessarily follow, however, that a gene can be cloned once and expressed in all the available host-vector combinations with uniformly satisfactory results. This is because different organisms have different mechanisms of translating mRNA into protein, and also because choices always have to be made when designing an expression construct (for example, the presence or absence of a stop codon). This unit first provides an overview of Gateway cloning, then reviews aspects of protein expression that limit the universality of the use of one clone in many vectors and hosts, and finally discusses how some of the conflicts between the structure of a Gateway clone of a
B1
gene
UNIT 5.17
gene and the rules of protein expression can be minimized or resolved. Examples of how Gateway cloning has been used in traditional and high-throughput applications are also described.
A BRIEF OVERVIEW OF GATEWAY Gateway cloning uses in vitro site-specific recombination, instead of restriction enzymes and ligase, to move a gene from one vector to another (Hartley et al., 2000; Fig. 5.17.1). Sitespecific recombination can be envisioned as a combination of restriction endonuclease cleavage and ligation, but with an important difference: the DNA termini produced by the cleavage reactions of a site-specific recombinase are not allowed to roam about and find a ligation partner by random collisions. Instead, the ligation partner is part of the reaction complex, and the transfer of DNA ends from one partner to another is programmed and controlled. One of the principal attractions of Gateway cloning is that potentially (and, up to this point, in reality) all Gateway clones and vectors are compatible with each other. By convention, all Gateway expression vectors have the same reading frame (in the case of fusion vectors) and the same orientation of att recombination sites, and all genes are cloned into the Gateway vector in the compatible reading frame. Converting existing expression vectors for Gateway vector compatibility is not complicated,
B2 C-tag
B1
gene
B2
native protein
C-terminal fusion
L1
gene L2
entry clone N-tag B1 gene
B2 C-tag
N-, C-terminal fusion
N-tag B1 gene
B2
N-terminal fusion
Figure 5.17.1 Gateway cloning for protein expression.
Production of Recombinant Proteins
Contributed by James L. Hartley
5.17.1
Current Protocols in Protein Science (2002) 5.17.1-5.17.10 Copyright © 2002 by John Wiley & Sons, Inc.
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------------------ at t B1 --------------------------------T
S
. . . a ca agt
L
Y
K
K
A
G FLSYCW
t tg
tac aaa aaa gca ggc tnn
nnn nnn . . .
ORF . . .
------------------ at t B2 --------------------------------------YHND P ...
nnn nnn
nac
A
cca gct
F
L
Y
K
V
V
ttc
ttg
tac
aaa
g tg
g tn . . . . . .
Figure 5.17.2 attB1 and attB2 sites in the conventional reading frame and orientation in a clone expressing a protein. The amino acids shown are made part of the desired protein only in the case of amino fusions (attB1) or carboxy fusions (attB2). Bases believed to be most important for att site function are shown in bold.
Use of the Gateway System for Protein Expression in Multiple Hosts
requiring ligation of a 1.7-kb cassette into any cloning site. Since reading frame and orientation are fixed, and subcloning with Gateway is trivial, vectors and genes can be exchanged and immediately used among different laboratories. Gateway is also useful in dealing with large genes (≥10 kb) or vectors (>100 kb), because att recombination sites should be extremely rare, and because the concerted mechanism of site-specific recombination brings both DNA partners into the reaction complex (Weisberg and Landy, 1983). While Gateway cloning and subcloning uses all four of the phage lambda recombination sites (attB, attP, attL, and attR), only the attB sites are of concern here, because only attB sites are present in the expression clones that produce recombinant proteins. There are two attB sites—attB1 on the amino side of the gene and attB2 on the carboxy side. If an open reading frame is cloned between attB1 and attB2 in the standard reading frame, and the attB1-ORFattB2 region is translated (i.e., fusions are made on both the amino and carboxy termini), the resulting protein will contain the sequence shown in Figure 5.17.2. Attempts to induce antibodies to attB1 and attB2 peptides have thus far been unsuccessful (G. Temple, pers. comm.). It should be noted that shorter, more efficient attB sites are now known, and there is a great deal of sequence flexibility possible (Cheo et al., 2001), so that, for example, translation signals could be made to be contained within a new version of attB1. However, using such alternative att sites might require reconstructing the other plasmids of the Gateway system,
and compatibility with existing Gateway vectors and gene libraries might be lost. As shown in Figure 5.17.1, the starting clone of an open reading frame (ORF) in the Gateway system is called an entry clone. An ORF in an entry clone cannot be usefully expressed because of the large size (100 bp) of the flanking attL sites. In fact, the vector backbones of entry clones typically contain a transcription terminator upstream of attL1 to decrease any inadvertent expression of the ORF and possible toxicity that might result. Entry clones are often obtained by in vitro recombination of attB-containing PCR products (Hartley et al., 2000). Two other technologies that allow DNA segments to be transferred from one vector to another by in vitro site-specific recombination are commercially available. The Echo system (Invitrogen) uses Cre-lox recombination to fuse a plasmid containing a gene of interest to a plasmid containing a promoter and, when desired, an amino-terminal fusion domain. The Creator technology (BD BioSciences) is also based on Cre-lox recombination, and transfers a cassette containing both the gene of interest and a drug resistance marker (positioned 3′ to the gene) to a new vector. Disadvantages for the Cre-lox-based methods are: (1) lox sites (34 bp) are larger than attB sites (21 to 25 bp); (2) single-stranded loxP sites (as in mRNA) form cruciforms that inhibit expression when they are in 5∼ UTRs (Liu et al., 1998); (3) carboxyterminal fusions are complicated (requiring intron splicing in eukaryotes) or impossible (must be included in the starting ORF for E. coli expression, cannot be changed); and (4) genes from expression clones cannot be recovered by recombination. Advantages are the relative
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simplicity of the systems, and the lower cost of Cre recombinase. Comparing the selections used in traditional subcloning (REaL, i.e., restriction enzymes and ligase) with Gateway, Echo, and Creator is instructive. For REaL, the combination of linearizing the new vector and inhibiting the circularization of the vector unless it has received the gene of interest (e.g., phosphatase treatment, incompatible ends) is fundamental. Thus, complete digestion of the recipient vector with restriction enzyme(s) is critical, as are the terminal bases of both the insert and the vector. For the recombinational cloning schemes, in contrast, the recipient vector is often circular, and the new covalent bonds that form the desired cloning product do not involve 5′ and 3′ ends at all. Instead, genetic selections are used to avoid the unreacted recipient and donor vectors, and the recombination reactions occur within the recombination sites. While the genetic selections impose some costs (special E. coli strains and/or growth media), the great advantage is the ease and universality of vector preparation. The cost of using recombination sites instead of restriction sites is larger size. What is gained from this cost is great specificity, with the important consequences that all reactions can be done with essentially the same reagents and conditions (some modifications may be required to compensate for DNA concentration or size), and that the precise structures of the cloning products (i.e., orientation and reading frame) are completely specified by the starting reagents.
PROTEIN EXPRESSION: A GENE IN A VECTOR IN A HOST Expression of a recombinant protein requires three components: a gene, a vector that supplies replication and expression functions for the gene, and milieu supplying the expression machinery (e.g., RNA polymerase, NTPs, ribosomes, and aminoacyl tRNAs). The gene is the information acted upon by the expression machinery in the context of the various cis elements present in the expression vector (e.g., transcription, translation, replication functions, and fusion tags). Usually, for protein expression, the gene consists of an initiator methionine (an ATG codon), an ORF of widely varying size (encoding from a few to thousands of amino acids), and a stop codon (TAG, TGA, or TAA). Historically, protein overexpression was achieved by simply cloning the native gene so as to increase the copy number of the gene, and
thus increase the number of transcripts and the amount of protein. With the advent of PCR, genes can be precisely tailored to produce native or fusion proteins in the hosts of choice. Introns are usually omitted by recovering the gene from a cDNA clone, to permit expression in prokaryotic hosts (usually E. coli), and to decrease the size and complexity of the gene. The Mammalian Gene Collection of the Natio nal In stitutes of Health (http://mgc.nci.nih.gov/) is a valuable source of PCR templates from which expression clones can be constructed. For expression, the gene must be transcribed and translated, and motifs that facilitate these processes must be provided by the DNA molecule containing the gene. The RNA polymerase recognition motif (promoter) may be a simple sequence (18 base pairs in the case of bacteriophage T7 RNA polymerase), or it may comprise hundreds of base pairs (as for the cytomegalovirus enhancer/promoter). Importantly, the promoter may be positioned tens or hundreds of base pairs upstream of the gene, and usually produces an RNA transcript that contains untranslated regions (UTRs) on both ends of the protein-coding region. The 5′ UTR of the mRNA often contains translation start signals (see below), while the 3′ UTR may contain RNA processing motifs, e.g., for polyadenylation. Once the mRNA containing the gene is synthesized, ribosomes must assemble on the 5′untranslated region (UTR) and find the initiating ATG codon. This process proceeds very differently in prokaryotes and eukaryotes. The DNA sequences involved in translation initiation and termination must briefly be discussed, because their arrangement is a significant limitation on the use of the same genetic construct in a variety of expression hosts and vectors. In E. coli, where the mRNA does not have a 5′ modification (cap), ribosomes apparently scan the message for ATGs with appropriate sequence context and spacing. The sequence context was originally identified by Shine and Dalgarno (1974) as a purine-rich region complementary to bases within the 16S RNA of the E. coli ribosome. The Shine-Dalgarno (SD) sequence must be positioned near and upstream of the initiating ATG, but the size and degree of the homology and its spacing relative to the ATG are quite variable. It is also important that the SD site be contained within a longer region of the 5′ UTR that has minimal secondary structure (deSmit and VanDuin, 1990). An SD motif within a larger (preferably unstructured)
Production of Recombinant Proteins
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region on amino side of gene supplied by expression vector
promoter
region that moves with ORF from an entry clone to new expression vectors
attB1
protease site? amino fusion tags (must have ATG and translation start)?
ORF
ATG? translation initiation? protease site? amino fusion tags?
region on carboxy side of gene supplied by expression vector
att B2
stop codon? protease site? carboxy fusion tags?
stop codon? protease site? carboxy fusion tags? transcription stop?
Figure 5.17.3 Motifs required for Gateway cloning (attB sites) and protein expression.
Use of the Gateway System for Protein Expression in Multiple Hosts
sequence comprises an E. coli ribosome binding site (RBS). Since bacteriophages are efficient at producing large amounts of capsid protein, a widely used group of E. coli expression vectors, the pET series (Rosenberg et al., 1987), incorporates a portion of the phage T7 capsid protein RBS (Olins and Rangwala, 1989). No equivalent to the E. coli ribosome binding site exists in eukaryotes. Instead, ribosomes appear to load onto the 5′ end of the mRNA as directed by the 5′ cap structure, and scan down the message to an ATG start codon (Kozak, 1999). Insect cells (Chang et al., 1999) and yeast cells (McCarthy, 1998; Yun et al., 1996) appear to follow a “first ATG” rule, i.e., the 5′-most ATG is highly preferred for translation initiation, with sequence context playing a minor role. In mammalian cells, on the other hand, there are well documented cases of ATGs within the 5′ UTR being bypassed in favor of others located further downstream. Comparison of sequences around these ATGs has resulted in the “Kozak” consensus for efficient translation initiation: GCCRCCaugG (Kozak, 1987). When the –3 base is A and the +4 base is G, other parts of the context are not very important (Kozak, 1999). Purines at –3 and +4 are found in highly expressed genes in other eukaryotes, e.g., plants (Joshi et al., 1997; Lukaszewicz et al., 2000), green algae (Ikeda and Miyasaka, 1998), and marine invertebrates (Mankad et al., 1998). Thus, eukaryotes follow relatively simple context rules for translation initiation (either “first ATG” or one or two “Kozak” bases near the ATG) for starting translation (from capped messages), whereas the predominant prokary-
otic expression host, E. coli, requires a longer motif containing more specific sequence to achieve the same result (with uncapped mRNAs).
COMBINING GATEWAY CLONING WITH PROTEIN EXPRESSION To what extent can the situation depicted in Figure 5.17.1 be realized—i.e., to what extent can a single gene construct, cloned in Gateway, be transferred to the appropriate expression vector to yield both native and fusion proteins in useful forms? Also, to what extent can these proteins be produced in E. coli and insect and mammalian and yeast cells, from a single entry clone? When an ORF is flanked by motifs necessary for one mode of expression (for instance, an RBS and a stop codon for native expression in E. coli), what is the effect of those motifs on another desired mode of expression, e.g., an amino fusion in mammalian cells? There are relatively few considerations to keep in mind when constructing a fusion protein at the carboxy end of the ORF (Fig. 5.17.3); if the entry clone has a stop codon, carboxy-terminal fusions are not possible. Carboxy fusion domains, e.g., an epitope tag or His6, must either be included upstream of the stop codon (and thus always remain with the ORF), or the stop codon must be omitted (in which case the amino acids encoded by attB2 will always be added to the carboxy terminus of the protein of interest). Partial relief from this situation may someday be possible through the use of suppression mechanisms (i.e., with suppressor tRNAs; J. Hartley, unpub. observ.), but the necessary vectors and/or hosts for this are not presently available. At least in E. coli, TAG
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Table 5.17.1
Effects of Translation Motifs and Protease Sites on Modes of ORF Expressiona
Example Motifs between att sites no.
Good for
Bad for
1
attB1-RBS-ATGORF-stop-attB2
Native protein in E. coli Carboxy fusions; amino fusions; native protein in some mammalian cells
2
attB1-Kz-ATGORF-stop-attB2
3
attB1-Kz-ATGORF- attB2
4
attB1-RBS-KzATG-ORF-attB2
Native protein in eukaryotes; all amino fusion proteins Carboxy fusions in eukaryotes; native amino end in eukaryotes; all amino fusions Carboxy fusions in E. coli and eukaryotes
5
attB1-TEV-Kz-ATG- All amino fusions; ORF-stop-attB2 native protein in eukaryotes attB1-ORF-attB2 All amino and carboxy fusions
Native protein in E. coli
attB1-Kz-ATG-ORF- All amino fusions; His6-stop-attB2 near-native protein in eukaryotes; affinity purification
Carboxy fusions; native protein in E. coli
6
7
Carboxy fusions; native protein in E. coli Carboxy fusions in E. coli (no RBS); no native protein at C-terminus, attB2 amino acids always present No native protein at C-terminus; amino fusions in E. coli
No native protein, at least a short amino fusion required for translation start
Notes Amino fusions will translate the RBS; native protein will coexpress with amino fusions in E. coli No internal starts in either E. coli or eukaryotes No expression in E. coli without RBS from vector, therefore no carboxy fusion without amino fusion Amino fusions have extra amino acids from RBS; in E. coli, amino fusions contaminated with native protein Near-native protein in E. coli after protease cleavage of amino fusions No ATG, no stop; this is the arrangement of the 19000-gene C. elegans ORFeome project (Reboul et al., 2001) His6 not removable; short C-terminal tails have little effect on activity
aAbbreviations: RBS, E. coli ribosome binding site (contains Shine-Dalgarno sequence); Kz, eukaryotic “Kozak” translation initiation sequence;
ORF, open reading frame; TEV, tobacco etch virus protease site.
(amber) stop codons are highly suppressible but are also efficient translation terminators (Mottagui-Tabar, 1998), so using TAG instead of TGA (weaker stopping of translation) or TAA (strongest terminator, but not as suppressible) as stop codons may allow C-terminal fusions to be made in the future. The options for making fusions on the amino side of the gene are more complex (Fig. 5.17.3). To produce protein in eukaryotic cells that is native at the amino end, a purine, preferably an A, at −3 from the ATG (dispensable in insect cells and yeast), significantly enhances translation initiation. An entry clone with such an arrangement can be used to make native protein in eukaryotic cells (when the expression vector supplies a promoter) or an amino fusion protein (when the expression vector supplies both a promoter and a translation start upstream of attB1). The codon containing the −3 purine
adds only one amino acid to the fusion protein. Since fusions through attB1 already add about 9 amino acids, this is a relatively minor consideration. However, adding an E. coli ribosome binding site to an entry clone is more significant. The RBS (which contains the SD motif) comprises from 4 (for modest levels of expression) to more than 10 amino acids (as noted above). In addition to adding more amino acids to an amino-terminal fusion, having an E. coli RBS in the entry clone presents another complication: any amino fusion expressed in E. coli will be contaminated with native protein, and the yield of the amino fusion will be reduced, due to internal translation initiation (in the RBS from the entry clone) as well as initiation at the amino fusion domain’s ATG. The addition of protease sites to entry clones can relieve some of the difficulties that can
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Table 5.17.2
Examples of Gateway Cloning for Protein Expression of One or a Few Genesa
Expression host(s)
Proteins expressed
Reference
Human cells, yeast
MASK kinase and mutants, expressed for activity and reporter assays, immunoprecipitations, two-hybrid Neuropeptide Y receptor expressed, ligand binding investigated Active glycosyltransferase secreted and purified from insect cells DCC, membrane protein implicated in colon cancer, expressed as fusions for coimmunoprecipitations, apoptosis DHFR expressed as TEV fusion, activity increased after GST tag was removed Klotho homolog, expressed, purified, and assayed as GST fusion Dicer localization determined by fusion to GFP Conditional expression vector used to express toxic Drosophila cyclin A Atrogin1 expressed as GST fusions for coimmunoprecipitations Two E. coli trehalose synthase genes fused, expressed in human cells
Dan et al. (2001)
Human Insect Human
E. coli
E. coli Human Yeast Human Human
Holmberg et al. (2002) Iwai et al. (2002) Liu et al. (2002)
Wang et al. (2001)
Arking et al. (2002) Billy et al. (2001) Finley et al. (2002) Gomes et al. (2001) Lao et al. (2001)
aAbbreviations: DCC, [protein] deleted in colorectal cancer; DHFR, dihydrofolate reductase; GFP, green fluorescent
protein; GST, glutathione-S-transferase; MASK, multiple ankyrin repeats single KH domain; TEV, tobacco etch virus protease site.
Use of the Gateway System for Protein Expression in Multiple Hosts
result from trying to express an ORF in both fusion and native forms, because the protease can be used to remove a fusion domain during purification. TEV (tobacco etch virus) protease (Carrington et al., 1989; Parks et al., 1994; Lucast et al., 2001) appears to be more specific than alternatives such as enterokinase, thrombin, and factor Xa. The canonical recognition sequence of TEV, ENLYFQ/G, has recently been shown to be very forgiving at the last amino acid, allowing proteins to be engineered so that TEV cleavage can leave any amino terminus except proline (Kapust et al., 2002). Thus, a TEV site upstream of the ATG can remove an amino-terminal tag and yield “native” protein (i.e., amino-terminal methionine). A TEV site adjacent to the last amino acid of an ORF can be used to remove a carboxy-terminal tag, leaving a 6-amino-acid tail. It should be noted that not all TEV sites designed into a construct are accessible to the protease. Additional amino acids between the TEV site and the fusion partner, e.g., 6 glycines, will sometimes help the efficiency of TEV cleavage (Evdokimov et al., 2001). TEV protease is 50% active in 1.5 M urea (D. Esposito, pers. comm.),
which may allow removal of a fusion tag under mildly denaturing conditions. A number of possible arrangements of ORFs, attB sites, and protein expression motifs in entry clones are shown in Table 5.17.1. The choice of most suitable expression construct must be made in advance, along with a list of acceptable compromises. For instance, the RBS needed to express native or carboxy fusions in E. coli (examples 1 and 4 in Table 5.17.1) is a liability for amino fusions in E. coli (a second RBS will be present for the amino fusion tag, so both the native protein and the fusion protein will be expressed), and the RBS adds extra amino acids when expressing amino fusions in eukaryotic hosts. Adding a codon upstream of the ATG that contains the Kozak A at –3, as in example 2 of Table 5.17.1, will allow native protein to be made in all eukaryotic hosts, but expression in E. coli will be weak except as amino fusions. Since a surprising number of these fusions display biological activity, perhaps this is not such a bad choice. However, if purification from E. coli is to be followed by structural studies, e.g., crystallization, the absence of an RBS (to make native
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Table 5.17.3 Examples of Gateway Cloning for High-Throughput Protein Expressiona
Expression host(s)
Proteins expressed
Reference
E. coli
Each of 27 small human proteins expressed as fusion with seven amino tags, tested for solubility Each of 32 human proteins expressed as fusions with 4 amino tags to test expression and solubility; 336 human proteins expressed as His6 fusions to test purification 107 human proteins expressed as N-terminal or C-terminal fusions with fluorescent proteins for intracellular localization Components of three multiprotein complexes expressed in retroviral vectors for identification of interacting proteins 60 yeast vectors converted to Gateway, various promoters and tags, tested with GFP 11 Agrobacterium-based Gateway vectors 30 proteasome proteins tested against C. elegans ORFeome by yeast two-hybrid 75 C. elegans DNA damage repair genes expressed as 2-hybrid fusions in yeast; dsRNA of same genes expressed in E. coli for RNAi in C. elegans
Hammarström et al. (2002)
E. coli
Monkey cells
Human cells
Yeast
Plants Yeast Yeast, RNAi in E. coli
Braun et al. (2002)
Simpson et al. (2000)
Gavin et al. (2002)
Funk et al. (2002)
Karimi et al. (2002) Davy et al. (2001) Boulton et al. (2002)
aAbbreviations: dsRNA, double-stranded RNA; GFP, green fluorescent protein; RNAi, RNA interference.
protein) or a protease site (to remove a fusion tag) on the amino side is a significant limitation. Including a His6 tag on the carboxy end of an ORF just upstream of the stop codon (example 7 in the table) will always allow affinity purification. C-terminal polyhistidine tags (up to His12) are widely used to purify proteins for structural studies, and accessibility of such tags has been cited as evidence of the native configuration of recombinant protein produced in E. coli (Pedelacq et al., 2002).
RESEARCH THAT COMBINES GATEWAY WITH PROTEIN EXPRESSION As the preceding sections indicate, modes of protein expression (native and fusion proteins, in various hosts) cannot be seamlessly combined. In addition, the attB sites required for moving genes among various expression vectors with the Gateway system may impose costs of extra nucleotides and amino acids, depending on the particular context. In light of
these considerations, how have investigators used Gateway to make proteins? No single user of Gateway has reported expressing the same cloned gene in all the forms shown in Figure 5.17.1 (native, amino fusion, carboxy fusion, double fusion), and this is not surprising given the difficulties discussed above (e.g., truly native expression requires a stop codon, making carboxy fusions impossible without additional maneuvering). Most of those expressing one or a few genes have used Gateway to either make multiple fusion proteins (e.g., His6, GST, or fluorescent proteins), or to make their cloning simpler or more successful. Some examples of these uses of Gateway are shown in Table 5.17.2. The report of Dan et al. (2001) exemplifies the new possibilities for research that are created when transferring a gene into a new expression context is made easier. In this report, proteins expressed from Gateway plasmids were used for immunoprecipitations, two-hybrid assays, investigating domains responsible for activity, and activating reporters in vivo,
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among other purposes. Holmberg et al. (2002) noted that their gene of interest, a G protein– coupled receptor, was “exceedingly difficult to clone,” and Gateway was the only method that worked. They expressed protein in mammalian cells that bound the physiological ligand. The study of Iwai et al. (2002) is notable because they purified an active protein that was secreted from insect cells that contained a FLAG tag and the attB1 coding sequence. They used a Gateway expression vector of their own construction that contained the human immunoglobulin κ signal peptide to facilitate secretion. Liu et al. (2002) expressed a protein implicated in colorectal cancer as His6-FLAG, His6, and GST amino fusions, and used deletion derivatives to map the regions of the protein that contribute to apoptosis. Wang et al. (2001) cloned the rat dihydrofolate reductase gene as a GST fusion via Gateway, and included a TEV protease site just upstream of the ORF; removal of the GST tag increased the enzymatic activity 30-fold. As would be expected, attention to compatibility of cloned ORFs with multiple modes of protein expression becomes more important as the number of genes becomes larger. Thus, Simpson et al. (2000) opted to include the ATG of each of 107 genes and omit its stop codon. In this way, they were able to make an amino fusion of each ORF with one fluorescent protein, and a carboxy fusion to the same ORF with a different fluorescent protein. They proposed that if each expressed protein localized to the same cellular compartment, the native gene product was likely to be found there also. The C. elegans ORFeome project (Reboul et al., 2001) has accepted the presence of the amino acids encoded by the attB1 and attB2 sites, and cloned 19000 C. elegans ORFs without either the ATG or stop codons (as in Table 5.17.1, example 6). This collection of clones is useful for yeast two-hybrid and RNAi applications (Walhout et al., 2000; Davy et al., 2001; Boulton et al., 2002). However, structural biologists have not found this context complete satisfactory (Chance et al. 2002). This has not prevented a large-scale effort to solve the structures of hundreds of the genes represented in this ORF collection (http://sgce.cbse.uab.edu/index.htm). Table 5.17.3 contains other examples of high-throughput applications of Gateway.
Use of the Gateway System for Protein Expression in Multiple Hosts
SUMMARY OF GATEWAY STRENGTHS AND WEAKNESSES Gateway’s weaknesses include (1) the requirement for long PCR primers; (2) addition of 25 base pairs or 9 amino acids to UTRs or
fusion proteins, respectively; (3) the expense of recombination proteins; and (4) licensing issues that may hinder adoption by commercial organizations. Gateway’s strengths include (1) all entry clones are compatible with all Gateway expression vectors; (2) reading frame and orientation are always preserved; (3) amino, carboxy, and double fusions are possible; (4) attB sites have no reported effect on expression level, proteolysis, or solubility; (5) amino acids encoded by attB sites are weakly or nonimmunogenic; (6) cloning (of attB PCR products) and subcloning (of genes into new vectors) are highly reliable, specific, simple, automatable, and efficient; (7) existing vectors are easily converted to compatibility with Gateway; (8) all components except the recombination proteins can be propagated by the investigator; and (9) vector and gene size are relatively unimportant. There is no single perfect cloning method for protein expression. Reactions that join two different DNA molecules into one can be highly specific (good) by using relatively large (bad) nucleic acid motifs (as in Gateway), or they can be less specific (bad) by using smaller (good) sequences (restriction enzymes and ligase). Different organisms require different transcription and translation motifs, thus requiring different expression vectors and different forms of the gene to be expressed. While it has its faults, Gateway offers the possibility of making hundreds of vectors constructed by clever biologists more accessible to the general research community.
LITERATURE CITED Arking, D.E., Krebsova, A., Macek, M., Sr, Arking, A., Mian, I.S., Fried, L., Hamosh, A., Dey, S., McIntosh, I., and Dietz, H.C. 2002. Association of human aging with a functional variant of klotho. Proc. Natl. Acad. Sci. U.S.A. 99:856-861. Billy, E., Brondani, V., Zhang, H., Muller, U., and Filipowicz, W. 2001. Specific interference with gene expression induced by long, doublestranded RNA in mouse embryonal teratocarcinoma cell lines. Proc. Natl. Acad. Sci. U.S.A. 98:14428-14433. Boulton, S.J., Gartner, A., Reboul, J., Vaglio, P., Dyson, N., Hill, D.E., and Vidal, M. 2002. Combined functional genomic maps of the C. elegans DNA damage response. Science 295:127-131. Braun, P., Hu, Y., Shen, B., Halleck, A., Koundinya, M., Harlow, E., and LaBaer, J. 2002. Proteomescale purification of human proteins from bacteria. Proc. Natl. Acad. Sci. U.S.A. 99:2654-2659. Carrington, J.C., Cary, S.M., Parks, T.D., and Dougherty, W.G. 1989. A second proteinase en-
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Gomes, M.D., Lecker, S.H., Jagoe, R.T., Navon, A., and Goldberg, A.L. 2001. Atrogin-1, a musclespecific F-box protein highly expressed during muscle atrophy. Proc. Natl. Acad. Sci. U.S.A. 98:14440-14445.
Chang, M.J., Kuzio, J., and Blissard, G.W. 1999. Modulation of translational efficiency by contextual nucleotides flanking abaculovirus initiator AUG codon. Virology 259:369-383. Cheo, D., Brasch, M.A., Temple, G.F., Hartley, J.L., and Byrd, D. 2001. Use of multiple recombination sites with unique specificity in recombinational cloning. International Patent Application Number WO 01/42509 A1. Dan, I., Ong, S.E., Watanabe, N.M., Blagoev, B., Nielsen, M.M., Kajikawa, E., Kristiansen, T.Z., Mann, M., and Pandey, A. 2001. Cloning of MASK, a novel member of mammalian germinal center kinase-III subfamily, with apoptosis-inducing properties. J. Biol. Chem. 276:3211532121. Davy, A., Bello, P., Thierry-Mieg, N., Vaglio, P., Hitti, J., Doucette-Stamm, L., Thierry-Mieg, D., Reboul, J., Boulton, S., Walhout, A.J., Coux, O., and Vidal, M. 2001. A protein-protein interaction map of the Caenorhabditis elegans 26S proteasome. EMBO Rep. 2:821-828. de Smit, M.H. and van Duin, J. 1990. Control of prokaryotic translational initiation by mRNA secondary structure. Prog. Nucleic Acid Res. Mol. Biol. 38:1-35. Evdokimov, A.G., Tropea, J.E., Routzahn, K.M., Copeland, T.D., and Waugh, D.S. 2001. Structure of the N-terminal domain of Yersinia pestis YopH at 2.0 A resolution. Acta Crystallogr. D Biol. Crystallogr. 57:793-799. Finley, R.L., Zhang, H., Zhong, J., and Stanyon, C. A. 2002. Regulated expression of proteins in yeast using the MAL61–62 promoter and a mating scheme to increase dynamic range. Gene 285:49-57. Funk, M., Niedenthal, R., Mumberg, D., Brunkmann, K., Ronicke, V., and Henkel, T. 2002. Vector systems for the heterologous expression of proteins in Saccharomyces cerevisiae. Methods Enzymol. 350:248-257. Gavin, A.C., Bosche, M., Krause, R., Grandi, P., Marzioch, M., Bauer, A., Schultz, J., Rick, J.M., Michon, A.M., Cruciat, C.M., Remor, M., Hofert, C., Schelder, M., Brajenovic, M., Ruffner, H., Merino, A., Klein, K., Hudak,M., Dickson, D., Rudi, T., Gnau, V., Bauch, A., Bastuck, S., Huhse, B., Leutwein, C., Heurtier, M.A., Copley, R.R., Edelmann, A., Querfurth, E., Rybin, V., Drewes, G., Raida, M., Bouwmeester, T., Bork, P., Seraphin, B., Kuster, B., Neubauer, G., and Superti-Furga, G. 2002. Functional organization of the yeast proteome by
Hammarström, M., Hellgren, N., van den Berg, S., Berglund, H., and Härd, T. 2002. Rapid screening for improved solubility of small human proteins produced as fusion proteins in Escherichia coli. Prot. Sci. 11:313-321. Hartley, J.L., Temple, G.F., and Brasch, M.A. 2000. DNA cloning using in vitro site-specific recombination. Genome Res. 10:1788-1795. Holmberg, S.K., Mikko, S., Boswell, T., Zoorob, R., and Larhammar, D. 2002. Pharmacological characterization of cloned chicken neuropeptide Y receptors Y1 and Y5. J. Neurochem. 81:462-471. Ikeda, K. and Miyasaka, H. 1998. Compilation of mRNA sequences surrounding the AUG translation initiation codon in the green alga Chlamydomonas reinhardtii. Biosci. Biotechnol. Biochem. 62:2457-2459. Iwai, T., Inaba, N., Naundorf, A., Zhang, Y., Gotoh, M., Iwasaki, H., Kudo, T., Togayachi, A., Ishizuka, Y., Nakanishi, H., and Narimatsu, H. 2002. Molecular cloning and characterization of a novel UDP-GlcNAc:GalNAc-peptide beta1,3N-acetylglucosaminyltransferase (beta 3GnT6), an enzyme synthesizing the core 3 structure of O-glycans. J. Biol. Chem. 277:12802-12809. Joshi, C.P., Zhou, H., Huang, X., and Chiang, V.L. 1997. Context sequences of translation initiation codon in plants. Plant Mol. Biol. 35:993-1001. Kapust, R.B., Tozser, J., Copeland, T.D., and Waugh, D.S. 2002. The P1′ specificity of tobacco etch virus protease. Biochem. Biophys. Res. Commun. 294:949-955. Karimi, M., Inze, D., and Depicker A. 2002. GATEWAY vectors for Agrobacterium-mediated plant transformation. Trends Plant Sci. 7:193-195. Kozak, M. 1987. An analysis of 5′-noncoding sequences from 699 vertebrate messenger RNAs. Nucleic Acids Res. 15:8125-8148. Kozak, M. 1999. Initiation of translation in prokaryotes and eukaryotes. Gene 234:187-208. Liu, Q., Li, M.Z., Leibham, D., Cortez, D., and Elledge, S.J. 1998. The univector plasmid-fusion system, a method for rapid construction of recombinant DNA without restriction enzymes. Curr. Biol. 8:1300-1309. Liu, J., Yao, F., Wu, R., Morgan, M., Thorburn, A., Finley, R.L. Jr., and Chen, Y.Q. 2002. Mediation of the DCC apoptotic signal by DIP13 alpha. J. Biol. Chem. 19:26281-26285. Lao, G., Polayes, D., Xia, J.L., Bloom, F.R., Levine, F., and Mansbridge, J. 2001. Overexpression of trehalose synthase and accumulation of intracellular trehalose in 293H and 293FTetR:Hyg cells. Cryobiology 43:106-113.
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Lucast, L.J., Batey, R.T., and Doudna, J.A. 2001. Large-scale purification of a stable form of recombinant tobacco etch virus protease. Biotechniques 30:544-550.
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Mankad, R.V., Gimelbrant, A.A., and McClintock, T.S. 1998. Consensus translational initiation sites of marine invertebrate phyla. Biol Bull. 195:251-254. McCarthy, J.E. 1998. Posttranscriptional control of gene expression in yeast. Microbiol Mol. Biol. Rev. 62:1492-1553. Mottagui-Tabar, S. 1998. Quantitative analysis of in vivo ribosomal events at UGA and UAG stop codons. Nucl. Acids Res. 26:2789-2796. Olins, P.O. and Rangwala, S.H. 1989. A novel sequence element derived from bacteriophage T7 mRNA acts as an enhancer of translation of the lacZ gene in Escherichia coli. J. Biol. Chem. 264:16973-16976.
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Pedelacq, J.D., Piltch, E., Liong, E.C., Berendzen, J., Kim, C.Y., Rho, B.S., Park, M.S., Terwilliger, T.C., and Waldo, G.S. 2002. Engineering soluble proteins for structural genomics. Nat. Biotechnol. 20:927-932.
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Reboul, J., Vaglio, P., Tzellas, N., Thierry-Mieg, N., Moore, T., Jackson, C., Shin-i, T., Kohara, Y., Thierry-Mieg, D., Thierry-Mieg, J., Lee, H., Hitti, J., Doucette-Stamm, L., Hartley, J.L., Temple, G.F., Brasch, M.A., Vandenhaute, J., Lamesch, P.E., Hill, D.E., and Vidal, M. 2001. Open-reading-frame sequence tags (OSTs) support the existence of at least 17,300 genes in C. elegans. Nat. Genet. 27:332-336.
Contributed by James L. Hartley Science Applications International Corporation (SAIC)/National Cancer Institute Frederick, Maryland
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Wheat Germ Cell-Free Expression System for Protein Production
UNIT 5.18
The wheat germ cell-free pipeline protocol developed at the Center for Eukaryotic Structural Genomics (CESG) for NMR structure determinations is presented as a workflow diagram in Figure 5.18.1. The approach consists of four steps: (1) creation of a plasmid for in vitro transcription, (2) small-scale (50-µl) screening to assay the level of protein production and solubility, (3) larger-scale (4- to 12-ml) production of uniformly 15 N-labeled protein to evaluate whether solution conditions can be found that render the target suitable for NMR structure determination (soluble, monodisperse, folded, and stable), and (4) production of sufficient uniformly 13 C,15 N-labeled protein for multidimensional, multinuclear magnetic resonance data collection. A well-defined series of cloning steps is used to create a DNA plasmid containing the target gene and 5 and 3 extensions that promote efficient transcription and translation (see Vinarov et al., 2004). A construct resulting in a His-tagged or GST-tagged fusion protein will facilitate subsequent protein purification. Small-scale protein expression and purification trials are generally carried out in a 96-well format (Basic Protocols 1 and 2). Successful candidates from these screens (i.e., those estimated to yield ≥0.5 mg/ml target protein under larger-scale production conditions, with solubility >50% as determined by SDS-PAGE) are then selected for larger-scale protein expression using 15 N-labeled amino acids (Basic Protocols 3 and 4). Purified, uniformly 15 N-labeled proteins are then assayed by 1 H-15 N correlation spectroscopy for their suitability as structural candidates; they must be folded, monodisperse, and stable at room temperature for at least 14 days. Targets that pass these tests are then prepared as uniformly 13 C,15 N-labeled protein samples. The purification workflow diagram for methods described in Support Protocols 1, 2 and 3 is shown in Figure 5.18.2. One of the features of cell-free protein production is that only the protein of interest is labeled, so contaminating proteins do not show up in normal multinuclear spectra. Dilution of the added labeled amino acids by endogenous natural abundance amino acids is not a problem; the level of label incorporation is >95% as determined by mass spectrometry. The approaches described in this unit are rapid and amenable to automation and form the basis for a high-throughput “pipeline protocol” for producing labeled proteins in the CESG (see Basic Protocol 5). With the use of the CellFree Sciences GeneDecoder1000 robotic system it is possible to carry out as many as 1052 small-scale (50-µl) screening reactions per week, and with use of the CellFree Sciences Protemist10 robotic system it is possible to carry out 24 of the production-scale (4-ml) reactions per week. These protocols can also be used to produce proteins labeled with selenomethionine for crystallization trials (D. Vinarov, unpub. observ.). Wheat germ extract is available from CellFree Sciences as well as from several other manufacturers (e.g., Promega, Invitrogen, or Novagen). Extracts can also be made inhouse, but the process is laborious and expensive and is not recommended. The advantages of wheat germ extract over other cell-free systems are discussed in the Commentary section.
Production of Recombinant Proteins Contributed by Dmitriy A. Vinarov, Carrie L. Loushin Newman, Ejan M. Tyler, John L. Markley, and Mark N. Shahan
5.18.1
Current Protocols in Protein Science (2006) 5.18.1-5.18.18 C 2006 by John Wiley & Sons, Inc. Copyright
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Figure 5.18.1 Workflow for production of isotopically enriched NMR samples based on the wheat germ cell-free expression system.
noncleavable N-(His)6 tag
cell free reaction
cleavable N-GST tag
1st GSTrap column HisTrap HP chelating column
concentrate
concentrate
PreScission protease cleavage
2nd GSTrap column
Superdex 75 in NMR Buffer
concentrate
NMR sample
Figure 5.18.2 Workflow for purification of isotopically enriched NMR samples produced using the wheat germ cell-free expression system.
BASIC PROTOCOL 1
MANUAL SMALL-SCALE (50-µ l) TRANSCRIPTION REACTION AND ETHANOL PRECIPITATION OF RESULTANT mRNA Small-scale screening offers the most economical way of testing whether a target will proceed through the critical stages leading to structure determination. In vitro protein expression requires transcription and translation reactions, which could be carried out in either coupled or decoupled mode. Performing transcription as a separate step, as described here, offers better control over quantity and quality of mRNA that is used in the translation reaction.
Wheat Germ Cell-Free Expression System
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Materials Template DNA construct (purified with Qiagen miniprep kit) 5× transcription buffer (see recipe) 25 mM NTP mixture (see recipe) Current Protocols in Protein Science
80 U/µ RNasin RNase inhibitor solution (Promega) 80 U/µl Sp6 RNA polymerase solution (Promega) Milli-Q-purified H2 O 6 M ammonium acetate (sterilize by passing through a 0.2-µm filter) 70% and 100% ethanol 1.7-ml microcentrifuge tubes, sterile Benchtop refrigerated centrifuge −80◦ C freezer 1. Add 2 to 5 µg template DNA from Qiagen miniprep to a sterile 1.7-ml microcentrifuge tube. 2. To the same tube, add the following components:
10 µl 5× transcription buffer 5 µl 25 mM NTP mixture 0.19 µl 80 U/µl Sp6 RNA polymerase solution 0.16 µl 80 U/µl RNasin RNase inhibitor solution Milli-Q-purified H2 O to a final volume of 50 µl Preparation a master mix of the first four components is recommended to provide sufficient mix for at least four 50-µl transcription reactions to ensure accurate pipetting.
3. Gently mix the reaction and incubate 3 hr at 37◦ C. The reaction should become cloudy due to the formation of pyrophosphate.
4. After incubation, centrifuge the sample 2 min at 11,000 × g, 4◦ C, transfer the supernatant to a sterile 1.7-ml microcentrifuge tube, and add 10 µl 6 M ammonium acetate and 150 µl 100% ethanol. 5. Invert the tube several times until a white precipitate forms. 6. Incubate 20 min on ice. 7. Centrifuge 20 min at 11,000 × g, 4◦ C and carefully pipet off the supernatant so as not to disturb the mRNA pellet. 8. Wash with 600 µl 70% ethanol and centrifuge 3 min at 11,000 × g, 4◦ C, again carefully pipetting off the supernatant. Let the pellet air dry ∼10 min. Use the purified mRNA immediately (e.g., small-scale translation in Basic Protocol 2) or store up to 3 months at –80◦ C as a 100% ethanol suspension.
MANUAL SMALL-SCALE (50-µ l) TRANSLATION REACTION Small-scale translation reactions performed separately from the transcription reaction allows for better control over the input parameters such as mRNA quantity and quality.
BASIC PROTOCOL 2
Materials Dialysis buffer with 0.3 mM amino acid mixture, unlabeled (see recipe) mRNA pellet (Basic Protocol 1) Wheat germ cell-free extract OD600 200 (CellFree Sciences, http://www.cfsciences.com) 20 mg/ml creatine kinase (Roche) solution in H2 O 80 U/µ RNasin RNase inhibitor solution (Promega) 8 mM amino acid mixture (see recipe) 24-well plate or dialysis cup holder Dialysis cups (12-kDa MWCO; Biotech International)
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Plastic wrap 26◦ C incubator Additional reagents and equipment for SDS-PAGE (UNIT 10.1) 1. Prepare the dialysis cups and check for leaks by setting them in a sterile 24-well plate containing 2.5 ml of Milli-Q-purified water and adding 500 µl water to each cup. The level of water in each cup should stay constant as long as the membrane is intact.
2. Prepare sufficient dialysis buffer with 0.3 mM amino acid mixture (unlabeled) for the number of samples, taking into account the dialysis cup holder volume. For example, if using a 24-well plate, each sample will require 2.5 ml of buffer; if using a dialysis cup holder, each sample will require 700 µl of dialysis buffer. This protocol assumes the use of a 24-well plate, so each sample will require 2.5 ml of dialysis buffer plus 31.215 µl for the actual 50-µl translation reaction (step 3). Preparing a slight excess of buffer to account for pipetting and other losses is recommended.
3. To the 1.7-ml tube containing the clean, dry mRNA pellet (from Basic Protocol 1) add the following components:
15 µl wheat germ cell-free extract OD600 200 (final OD600 = 60) 1.75 µl 20 mg/ml creatine kinase solution 0.16 µl RNasin RNase inhibitor solution 1.875 µl 8 mM amino acid mixture 31.215 µl dialysis buffer with 0.3 mM amino acids 4. Allow the pellet to dissolve ∼5 min and gently mix to homogeneity with a pipet. Make sure the mRNA pellet is completely dissolved, and be careful not to form any air bubbles while mixing.
5. Shake out the water from the prepared dialysis cups and add the 50-µl reaction mixture. 6. Add 2.5 ml dialysis buffer with 0.3 mM amino acid mixture (unlabeled) to a clean, sterile well in the 24-well plate, and set the dialysis cup in the well, making sure there are no bubbles in the buffer/dialysis cup interface. Wrap the plate with plastic wrap and incubate 16 hr at 26◦ C. 7. Analyze using SDS-PAGE (UNIT 10.1). Since creatine kinase is added to the reaction mix at 700 µg/ml, it is clearly visible on the SDS-PAGE gel at 42 kDa and provides a good internal control for the level of expression of the target proteins. Typically, targets expressed at ∼500 µg/ml with at least a 75% soluble fraction should be selected for large-scale protein production. BASIC PROTOCOL 3
MANUAL LARGE-SCALE (6-ml) TRANSCRIPTION REACTION AND ETHANOL PRECIPITATION OF RESULTANT mRNA This large-scale transcription reaction is used to provide starting material for both manual and automated large-scale protein translation reactions.
Materials
Wheat Germ Cell-Free Expression System
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Template DNA construct (purified from plasmid DNA with a Marligen maxiprep kit; http://www.marligen.com) 5× transcription buffer (see recipe) 25 mM NTP mixture 80 U/µ RNasin RNase inhibitor solution (Promega) 80 U/µl Sp6 RNA polymerase solution (Promega) Milli-Q-purified H2 O Current Protocols in Protein Science
6 M ammonium acetate (sterile filtered through 0.2 µm filter) 70% and 100% ethanol 50-ml conical polypropylene centrifuge tube 37◦ C incubator or water bath Benchtop refrigerated centrifuge −80◦ C freezer (optional) 1. Using a 50-ml conical polypropylene centrifuge tube add the following components:
120 µg template DNA construct 1.2 ml 5× transcription buffer 720 µl 25 mM NTP mixture 18.75 µl 80 U/µl RNasin RNase inhibitor solution 22.5 µl 80 U/µl Sp6 RNA polymerase solution Milli-Q-purified H2 O to a final volume of 6 ml The amount of water added will vary because of different plasmid-DNA concentrations.
2. Gently pipet the reaction to fully mix the added components. 3. Incubate at 37◦ C for 3 to 5 hr. The reaction should turn cloudy due to the presence of pyrophosphate.
4. Centrifuge the sample 5 min at 10,000 × g, 4◦ C, and decant the supernatant into a clean 50-ml conical tube. To the tube, add 1.2 ml 6 M ammonium acetate and 18.0 ml 100% ethanol. 5. Invert the tube several times until a white precipitate forms, and incubate 20 min on ice. 6. Centrifuge 20 min at 10,000 × g, 4◦ C and decant the supernatant, being careful not to disturb the pellet. 7. Add 40 ml 70% ethanol and centrifuge 5 min at 10,000 × g, 4◦ C. Carefully decant the supernatant without disturbing the pellet, and let the pellet air dry for at least 1 hr. Pipet off any liquid that collects at the bottom of the tube. Use the purified mRNA immediately (e.g., large-scale translation in Basic Protocol 4) or store at −80◦ C for up to 3 months as a 100% ethanol suspension.
MANUAL LARGE-SCALE (4-ml) TRANSLATION REACTION AND SETUP FOR AUTOMATION
BASIC PROTOCOL 4
The large-scale translation reactions performed separately from the transcription reaction allow for better control over the input parameters such as mRNA quantity and quality, while automation allows for greater control over reaction parameters such as temperature and centrifugation speeds and times.
Materials Dried mRNA pellet (Basic Protocol 3) Wheat germ cell-free extract OD600 200 (CellFree Sciences, http://www.cfsciences.com) 16.8 mg/ml creatine kinase (Roche) in 1× dialysis buffer with 0.3 mM amino acids (see recipe) 80 U/µ RNasin RNase inhibitor solution (Promega) Dialysis buffer with 0.3 mM amino acids (see recipe) 15-ml Amicon centrifugal filtration unit (MWCO 10 kDa; Millipore) Protemist10 (CellFree Sciences, http://www.cfsciences.com)
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26◦ C incubator Benchtop refrigerated centrifuge capable of handling 50-ml conical tubes 50-ml conical centrifuge tubes Additional reagents and equipment for SDS-PAGE (UNIT 10.1) NOTE: Steps for preparation of the RNA are for both the manual synthesis reaction in a benchtop centrifuge and the automated Protemist10 reaction. 1. To the dried mRNA pellet (from Basic Protocol 3) add the following components:
1.2 ml wheat germ cell-free extract OD600 200 (final OD600 = 60). 167 µl of 16.8 mg/ml creatine kinase in 1× dialysis buffer with 0.3 mM amino acids 12.5 µl RNasin RNase inhibitor solution 2.62 ml dialysis buffer containing 0.3 mM amino acids. 2. Allow the pellet to dissolve ∼10 min at room temperature (22◦ C to 25◦ C), and then gently agitate the reaction mixture with a pipet until the pellet is completely dissolved.
For manual synthesis 3a. Load the 4-ml reaction mixture into a 15-ml Amicon centrifugal filter unit (MWCO 10 kDa), and incubate 30 min at 26◦ C. Add 2.5 ml dialysis buffer with 0.3 mM amino acids and gently mix with a pipet. 4a. Centrifuge 7 to 12 min at 4,000 × g, 4◦ C (until the flow-through volume is 2.5 ml). Discard the flow-through (or save it in a different conical tube until the reaction is complete). Add 2.5 ml dialysis buffer with 0.3 mM amino acids, gently mix with a pipet, and incubate 30 min at 26◦ C. Repeat this step for 18 cycles.
For automated synthesis 3b. Place the 4-ml reaction mix and 50 ml dialysis buffer containing 0.3 mM amino acids into the Protemist10. 4b. Start the program (∼14 to16 hr for one to four samples and 20 hr for five to eight samples). Robots are preprogrammed with several available programs for various numbers of reactions and reaction volumes. Although it is possible to change the program parameters, it is not recommended without a significant level of training.
5. Analyze reaction products by SDS-PAGE (UNIT 10.1). BASIC PROTOCOL 5
SMALL-SCALE AUTOMATED TRANSCRIPTION AND TRANSLATION REACTIONS ON THE GeneDecoder1000 The GeneDecoder1000 operates in two modes: (1) screening mode, which uses four 96well plates, 2.5 to 5.0 ml of wheat germ extract per plate, and an overnight run to produce 2 to 10 µg of protein per well overnight run; and (2) small-scale protein production mode, which uses two 96-well plates, 5.0 to 10.0 ml of wheat germ extract per plate and an overnight run to produce 10 to 50 µg of protein per well. The Protemist10 is capable of carrying out eight 4-ml reactions using 3 ml of wheat germ extract per reaction in an overnight run to produce 1 to 3 mg of protein per reaction.
Wheat Germ Cell-Free Expression System
This protocol uses one, two, or four 96-well plates to screen for protein expression on a micro-scale in a bilayer reaction. It is necessary to be familiar with the GeneDecoder1000 programs and software for this protocol. This protocol is written for one 96-well plate; the values can be doubled or quadrupled for two or four plates, respectively.
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Materials ∼5 µg/ml template DNA constructs (purified from plasmid DNA with a Qiagen miniprep kit) 5× transcription buffer (see recipe) 80 U/µl RNasin RNase inhibitor solution (Promega) 80 U/µl Sp6 RNA polymerase solution (Promega) 25 mM NTP mixture (see recipe) Milli-Q-purified H2 O 6 M ammonium acetate (sterilize by passing through a 0.2-µm filter) 70% and 100% ethanol Dialysis buffer with 0.3 mM amino acids (see recipe) 8 mM amino acid mixture (see recipe) Wheat germ cell-free extract OD600 200 (CellFree Sciences; http://www.cfsciences.com) 20 mg/ml creatine kinase (Roche) in H2 O 96-well PCR plates with lids (Greiner Bio-One 652280) GeneDecoder1000 and corresponding sterile troughs (CellFree Sciences, http://www.cfsciences.com) MTP plates with lids (Greiner Bio-One 650201) 20-µl BioRobotix tips (Molecular BioProducts 918-261; http://www.mbpinc.com) 300-µl MBP tips (Molecular BioProducts 3771; http://www.mbpinc.com) Ethanol towels (e.g., Wypall L-40) Additional reagents and equipment for SDS-PAGE (UNIT 10.1) 1. Dispense the ∼5µg/ml template DNA constructs into a 96-well PCR plate. The robot normally adds 10 µl of DNA to each transcription reaction. If the DNA concentration is >5 µg/ml, the volume of DNA preparation can be adjusted along with the corresponding volume of Milli-Q-purified water (see step 2).
2. Prepare the transcription mixture (sufficient for 110 × 25-µl reactions) by combining the following:
550 µl 5× transcription buffer (see recipe) 8.8 µl 80 U/µl RNasin RNase inihibitor solution 10.45 µl 80 U/µl Sp6 RNA polymerase solution 274 µl 25 mM NTP mixure 805.2 µl Milli-Q-purified H2 O and add to a small trough in the reagent incubator of the GeneDecoder1000. 3. Combine 1.125 ml 6 M ammonium acetate and 18.875 ml 100% ethanol, and add the mixture to a large trough in the reagent incubator of the GeneDecoder1000. 4. Prepare 19 ml dialysis buffer with 0.3 mM amino acids and dispense into a large sterile trough. The next step will use 1.873 ml dialysis buffer with 0.3 mM amino acids. The rest will be used to overlay each reaction well with 125 µl.
5. Prepare the translation mixture (sufficient for 120 × 25-µl reactions) by combining:
900 µl wheat germ cell-free extract OD600 200 105 µl creatine kinase 112.5 µl 8 mM amino acid mixture 9.6 µl RNasin RNase inhibitor solution 1.873 ml dialysis buffer with 0.3 mM amino acids and add to a small trough in the reagent incubator of the GeneDecoder1000. Current Protocols in Protein Science
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6. Set up the MTP plates with lids, 20-µl BioRobotix tips, 96-well PCR plates with lids, and ethanol towels in the GeneDecoder1000 according to the manufacturer’s directions. Select the correct preprogramed conditions in the software and start the program. Be sure to add a balance MTP plate with 150 µl of Milli-Q-purified water in each well to the centrifuge “dummy” position when screening only 1 plate.
7. Analyze reaction products by SDS-PAGE (UNIT 10.1). SUPPORT PROTOCOL 1
IMMOBILIZED METAL AFFINITY CHROMATOGRAPHY (IMAC) PURIFICATION OF His6 -TAGGED FUSION PROTEINS AND PREPARATION OF SAMPLE FOR NMR ANALYSIS Protein production is typically followed by protein purification. His-tagged target proteins produced by wheat germ cell-free protein expression can be purified efficiently and effectively by immobilized metal affinity chromatography (IMAC). HisTrap columns tolerate DTT concentrations up to 10 mM, and their use eliminates the need for dialysis to lower the levels of DTT in the reactions mixtures containing the protein product. Typical protein purity of approximately 75% is attained using IMAC.
Materials Reaction mixture (Basic Protocol 4) Protease inhibitor cocktail V (Sigma-Aldrich) Ni elution buffer (see recipe) Ni binding buffer (see recipe) 1-ml HisTrap HP chelating column (Amersham 17-5247-01) ¨ Pump: automated purifier (e.g., AKTA; GE Healthcare, http://www.gehealthcare. com) or 1-ml, 3-ml, or 10-ml syringes with a 0.7-mm blunt end needle, or peristaltic pump Amicon Ultra-4 concentrators (5- or 10-kDa MWCO; Millipore) Refrigerated centrifuge(s) capable of handling 1.5-ml microcentrifuge tubes and 15- and 50-ml conical tubes Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and NMR analysis (see UNIT 17.5) Prepare samples 1. Treat reaction mixture with protease inhibitor cocktail V (1:100 v/v) and incubate at least 30 min at 4◦ C. 2. Centrifuge samples 15 min at 9,300 × g, 4◦ C to clear. Save supernatant on ice to load on the HisTrap HP chelating column.
Prepare and run columns 3. Wash the HisTrap HP chelating column with 2 to 3 column volumes of Ni elution buffer followed by 5 column volumes of Ni binding buffer. ¨ This protocol can be performed automatically using an automated purifier (e.g., AKTA) or manually using 1-ml, 3-ml, or 10-ml syringes with a 0.7-mm blunt end needle, or a peristaltic pump
4. Load the sample at a rate of 1.0 ml per min at room temperature. Wheat Germ Cell-Free Expression System
5. Elute with 5 column volumes of Ni elution buffer and collect in single column volume fractions. Elutions can be performed in either a gradient or one-step profile. Gradient elution can produce a 90% pure sample; however, this is not typical. The advantage of the one-step
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elution procedure is primarily in the smaller final volume, which significantly reduces the time needed for the concentration step. The major contaminants that typically copurify by this method can easily be separated from monomeric proteins by gel filtration (UNIT 8.3). Additionally, the contaminating bands do not interfere with prescreening by NMR, because they are unlabeled. Therefore samples can be prescreened by NMR after the initial IMAC purification. ¨ Time to purify is ∼35 min when an AKTA purifier is used. Concentration and buffer exchange times vary greatly between proteins, but on average it takes 30 to 45 min to reduce the volume from 4 ml to 300 µl. The column can be stripped and regenerated according to manufacturer’s instructions.
6. Concentrate the sample to a final volume of <300 µl by centrifuging the samples 5 min at maximum speed, 4◦ C in an Amicon Ultra-4 concentrator. Proceed to Support Protocol 2 for gel filtration purification of sample. Use 5-kDa MWCO units for proteins <15 kDa and 10 kDa MWCO units for proteins >15 kDa.
GEL-FILTRATION PURIFICATION AND PREPARATION OF SAMPLE FOR NMR ANALYSIS
SUPPORT PROTOCOL 2
Purification by gel filtration allows target proteins to be obtained at >95% purity.
Materials NMR screening buffer (see recipe) Glycerol Concentrated sample (Support Protocol 1) NMR screening buffer with 7% D2 O (see recipe) Protease inhibitor cocktail V (Sigma-Aldrich) Superdex 75 column (GE Healthcare, http://www.gehealthcare.com) ¨ Pump: automated purifier (e.g., AKTA; GE Healthcare, http://www.gehealthcare.com) or 1-ml, 3-ml, or 10-ml syringes with a 0.7-mm blunt end needle, or peristaltic pump Refrigerated centrifuge Amicon Ultra-4 concentrators (5- or 10 kDa MWCO) 1.5-ml microcentrifuge tubes Additional reagents and equipment for SDS-PAGE (UNIT 10.1), protein quantitation (UNIT 3.4), and NMR analysis (see UNIT 17.5) Prepare and run columns 1. Equilibrate the Superdex 75 column by washing with 2.5 column volumes of appropriate NMR screening buffer. 2. Mix 10 µl glycerol and 90 µl NMR screening buffer to use as a band sharpener during the loading procedure. 3. Load 300 µl concentrated sample on column followed by 50 µl of the band sharpener. 4. Run 2 additional column volumes of buffer through the column at 0.5 ml per min and collect 0.5 ml fractions after void volume. Pool selected fractions. ¨ This protocol can be performed automatically using an automated purifier (e.g., AKTA) or manually using 1-ml, 3-ml, or 10-ml syringes with a 0.7-mm blunt end needle, or a peristaltic pump. ¨ The computer that runs the AKTA automated purifier displays the chromatogram absorbance units versus time. This information can be used to select fractions.
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Prepare NMR sample 5. Centrifuge pooled fractions in an appropriate Amicon Ultra-4 concentrator 5 min at 7,000 × g, 4◦ C to concentrate the protein to a 400-µl volume. 6. Add NMR screening buffer with 7% D2 O to a total volume of 4 ml. Concentrate to final volume of 300 µl as in the previous step. 7. Centrifuge the samples 5 min at full speed, 4◦ C. Remove the supernatant to new tube for NMR. 8. Save the pellet and 0.5 µl of the supernatant for gel analysis and 7.5 µl of the supernatant for BCA protein assay (UNIT 3.4). 9. Add 3 µl protease inhibitor cocktail V to sample, label, and analyze by NMR (see UNIT 17.5). SUPPORT PROTOCOL 3
PURIFICATION OF GST-FUSION PROTEIN AND PREPARATION OF SAMPLE FOR NMR ANALYSIS Wheat germ cell-free protein expression allows for the production of GST-tagged proteins. Purification and subsequent tag removal described in this protocol offer effective and efficient means of obtaining purified target protein.
Materials Phosphate-buffered saline (PBS; diluted from 10× PBS; see recipe) with 10 mM dithiothreitol (DTT): PBS+DTT 10× PBS (see recipe) GST-fusion construct (Basic Protocol 4) 10 mM reduced glutathione (Sigma, G-6529) in PBS+DTT NMR screening buffer (see recipe) 10× protease buffer, pH 7 (see recipe) PreScission protease solution (GE Healthcare) NMR screening buffer with 7% D2 O (see recipe) BCA protein assay reagents (Pierce) 1-ml GSTrap HP column (Amersham 17-5281-01 or 17-5282-02) or GST Microspin purification kit (Amersham, 27-4570-03) ¨ Pump: automated purifier (e.g., AKTA; GE Healthcare, http://www.gehealthcare.com) or 1-ml, 3-ml, or 10-ml syringes with a 0.7-mm blunt end needle, or peristaltic pump Refrigerated centrifuge Amicon Ultra-4 concentrator (5 or 10 kDa; Millipore) 1.5-ml microcentrifuge tubes Nutator mixer (Clay Adams) Additional reagents and equipment for SDS-PAGE (UNIT 10.1), NMR analysis (see UNIT 17.5), and protein quantitation (UNIT 3.4) Purify GST-fusion construct 1. Prepare GSTrap column by washing with 5 column volumes of PBS+DTT. The GST Microspin purification kit may be used to purify small-scale reaction product.
2. Add 10× PBS to sample to final concentration of 1×. Wheat Germ Cell-Free Expression System
3. Load GST-fusion construct on to the GSTrap column at a rate of 0.5 ml per min. 4. Wash with 5 column volumes PBS+DTT.
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5. Elute with 10 column volumes of 10 mM reduced glutathione in PBS+DTT, and collect 0.7-ml fractions. Save column for second purification step.
6. Select fractions to pool for protease digestion. Centrifuge pooled fractions in an appropriate Amicon Ultra-4 concentrator 5 min at 7,000 × g, 4◦ C to to concentrate protein in a 400 µl volume. Protein usually elutes within the first 3 ml. ¨ This protocol can be performed automatically using an automated purifier (e.g., AKTA) or manually using 1-ml, 3-ml, or 10-ml syringes with a 0.7-mm blunt end needle, or a peristaltic pump. ¨ The computer that runs the AKTA automated purifier displays the chromatogram absorbance units versus time. This information can be used to select fractions.
7. Add PBS and concentrate the protein as in the previous step. Repeat several times to exchange the buffer 100-fold (∼8 ml total). Concentrate to a final volume of ∼600 µl. Remove 1 µl for analysis by SDS-PAGE (UNIT 10.1). The glutathione concentration must be reduced to <1 mM to facilitate binding of the protease enzyme and GST tag in the second purification step.
Remove the GST tag 8. Add 70 µl 10× protease buffer and 15 µl PreScission protease solution to the protein concentrate. Incubate overnight at 4◦ C on the Nutator mixer. Agitation of the sample increases the efficiency of cleavage.
9. Centrifuge the samples 5 min at 15,000 × g, 4◦ C. Remove the supernatant to a new tube. 10. Save the pellet and 0.5 µl of the supernatant for analysis by SDS-PAGE (UNIT 10.1).
Purify target protein 11. Wash the GSTrap column with 5 column volumes 1× PBS+DTT. 12. Load the sample (supernatant from step 9) onto the column used in the first purification. 13. Wash sample with 3 to 5 column volumes PBS+DTT. Collect flow-through, which contains the cleaved protein. Contaminants may be removed from the column at this time by washing the column with 10 mM glutathione PBS+DTT. This procedure cleans the column for repeated use.
14. Centrifuge selected fractions in an appropriate Amicon Ultra-4 concentrator 5 min at 7,000 × g, 4◦ C to concentrate protein to a volume of 400 µl. 15. Add NMR screening buffer with 7% D2 O and concentrate the protein as in the step 6. Repeat several times to exchange the buffer 100-fold (∼8 ml total). Concentrate to a final volume of ∼300 µl. 16. Remove supernatant to new 1.5-ml microcentrifuge tube and analyze by NMR (see UNIT 17.5). 17. Save the pellet and 0.5 µl of the supernatant for analysis by SDS-PAGE (UNIT 10.1) and 7.5 µl of the supernatant for protein quantitation by BCA protein assay (UNIT 3.4). Production of Recombinant Proteins
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX. Use only certified RNA/DNase-free buffers and components. Have on hand sterile, baked, 1-liter and 250-ml graduated cylinders and magnetic stir bars.
Amino acid mixture, 8 mM Combine the 20 amino acids to a final concentration of 8 mM each in a 50-ml conical tube. Add 0.01 N HCl to 50 ml, and stir with a stir bar for 2 hr. Store up to 1 year at −20◦ C. According to the stage of protein production (see Fig. 5.18.1 and the unit introduction) the amino acids may be unlabeled or labeled with 15 N, 13 C and 15 N (Cambridge Isotope Laboratories) or selenomethionine (Sigma-Aldrich).
Dialysis buffer with 0.3 mM amino acids Add 1/5 the volume necessary of the 5× dialysis buffer (see recipe). Add enough of the 8 mM amino acid mix to make the final concentration 0.3 mM. Bring solution to the final volume with H2 O. Sonicate 5 to 10 min, and sterilize by passing through a 0.2-µm filter. Prepare fresh on day of use. Dialysis buffer, 5× 45 ml 2 M HEPES stock, pH 7.8 (see recipe; 150 mM) 200 ml H2 O 28.4 g potassium acetate (500 mM) 1.74 g magnesium acetate (13.5 mM) 1.16 g dithiothreitol (DTT; 12.5 mM) 2.18 g ATP (6 mM) 426 mg GTP (1.25 mM) 15.7 g creatine phosphate (80 mM) 150 mg sodium azide (0.25%) 183.2 µl of 6.54M spermidine (Fluka; 2 mM)). Bring volume to 600 ml with H2 O Add a stir bar to the buffer in a 1-liter graduated cylinder, and mix until all components are competely dissolved. Sterilize by passing through a 0.2-µm filter using a vacuum filter unit. Dispense aliquots into 12 × 50–ml conical tubes, and store up to 6 months at −80◦ C. Dilute if necessary with H2 O. Final concentrations of components in the 5× stock are given in parentheses.
Glutathione 10 mM reduced glutathione (Sigma-Aldrich) 1× PBS:dilute 10× PBS (see recipe) with H2 O 10 mM DTT Prepare fresh on day of use
Wheat Germ Cell-Free Expression System
HEPES stock, pH 7.5 Dissolve 95.32 g HEPES in just under 200 ml water in a 250 ml graduated cylinder using a stir bar on a magnetic plate. Adjust the pH to 7.5 using solid potassium hydroxide. Remove the stir bar and add H2 O to a final volume of 200 ml. Check the pH again and adjust if necessary. Sterilize by passing through a 0.2-µm filter using a vacuum filter unit. Store up to 6 months at room temperature. HEPES buffer stock, pH 7.8
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HEPES stock, pH 7.8 Dissolve 95.32 g HEPES in just under 200 ml H2 O in a 250 ml graduated cylinder using a stir bar on a magnetic plate. Adjust the pH to 7.8 using solid potassium hydroxide. Remove the stir bar, and add H2 O to a final volume of 200 ml. Check the pH again, and adjust if necessary. Sterilize by passing through a 0.2-µm filter using a vacuum filter unit. Store up to 6 months at room temperature. Ni binding buffer, pH 8.00 50 mM NaH2 PO4 300 mM NaCl 50 mM imidazole Store up to 6 months at room temperature Add DTT to a final concentration of 2 mM just prior to use
Ni elution buffer, pH 8.00 50 mM NaH2 PO4 300 mM NaCl 250 mM imidazole Store up to 6 months at room temperature Add DTT to a final concentration of 2 mM just prior to use
NMR screening buffer with and without 7% (v/v) D2 O Choose any of the following buffers depending on the studied protein’s isoelectric point: 10 mM sodium acetate, pH 5.0/100 mM NaCl/0.02% NaN3 (pI between 6 and 7 or greater than 7) 10 mM Bis-Tris, pH 6.0/100 mM NaCl/0.02% NaN3 (pI less than 5 or greater than 7) 10 mM Bis-Tris, pH 7.0/100 mM NaCl/0.02% NaN3 (pI less than 6 or greater than 8) When required, add D2 O (Sigma-Aldrich) to a final concentration of 7% (v/v). Store up to 6 months at room temperature. Add DTT to a final concentration of 5 mM just before use. It is important to add DTT when proteins contain cysteine. The composition of the NMR screening buffer must be determined by the user, depending on the characteristics of the protein being isolated. The three recipes given are suggestions.
NTP mixture, 25 mM In a 50 ml conical vial, add the following components: 708 mg GTP 756 mg ATP 688 mg UTP 658 mg CTP Add H2 O to a final volume of 50 ml. Mix until dissolved and sterilize by passing through a 0.2-µm filter into five 15-ml conical tubes (10 ml each). Store up to 1 year at −20◦ C.
Phosphate buffered saline (PBS), 10×, pH 7.30 100 mM Na2 HPO4 1.50 M NaCl 1.4 M NaCl
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27 mM KCl Adjust pH with 1 N HCl Store up to 6 months at room temperature Dilute with H2 O to desired concentration Protease buffer, 10×, pH 7.00 500 mM Tris·Cl 10 mM KH2 PO4 10 mM EDTA 10 mM DTT Adjust the pH with 1 N HCl Store up to 6 months at −20◦ C in 100-µl aliquots Before use, dilute to 1× and add 2 M DTT fresh to a final concentration of 2 mM. Transcription buffer, 5× To a 250-ml graduated cylinder, add the following components: 40 ml HEPES stock, pH 7.5 (see recipe; 400 mM) 3.43 g magnesium acetate (80 mM) 306 µl spermidine (6.54 M solution; Fluka) 1.54 g DTT (50 mM) Add H2 O to a final volume of 200 ml Mix the buffer with a stir bar until all components are dissolved. Sterilize by passing through a 0.2-µm filter using a vacuum filter unit. Dispense aliquots into 50-ml conical tubes. Store up to 6 months at −20◦ C.
COMMENTARY Background Information
Wheat Germ Cell-Free Expression System
Cell-free methods for protein synthesis with extracts from prokaryotic (Kramer et al., 1999) or eukaryotic (Clemens and Prujin, 1999) sources offer an alternative to the E. coli cell-based platforms that are the mainstay of most structural proteomics centers. Stableisotope or selenomethionine labeling is easier with cell-free systems than with the simple eukaryote, D. discoideum (Cubeddu et al., 2000), or insect cell systems (Bruggert et al., 2003; Strauss et al., 2003). Cell-free systems permit the production of proteins that undergo proteolysis in cells (Goff and Goldberg, 1987; Maurizi, 1987) or aggregate in inclusion bodies (Chrunyk et al., 1993). Cell-free systems also support selective labeling strategies that cannot be achieved in bacterial whole-cell systems (Kigawa et al., 1995; Yabuki et al., 1998; Torizawa et al., 2002; Shi et al., 2004). They have been used for the production of various kinds of proteins, including membrane proteins (Klammt et al., 2004) and proteins that are toxic to cells (Henrich et al., 1982; Chrunyk et al., 1993). It is possible to collect NMR spectra of 15 N-labeled proteins prior to isolation from the cell-free protein synthesis mixture (Guignard et al., 2002). Successful implementation of cell-free protein expression
obviates the need for cell harvesting, cell lysis, and some lengthy steps (e.g., large-volume protein concentration) in protein purification. Protein purification is usually simpler because the protein of interest starts out more concentrated and is isolated from a smaller set of contaminants. The RIKEN Structural Genomics Center, in collaboration with Roche, has pioneered the use of cell-free protein production through a coupled transcription-translation system employing E. coli extracts (Kim et al., 1996; Kigawa et al., 1999; Yokoyama et al., 2000; Kigawa and Yokoyama, 2002; Yokoyama, 2003). In the experience of another laboratory that has investigated this approach (Masatsune Kainosho, personal communication), most of the proteins that are produced well in E. coli cell-free systems are the same ones that are produced successfully from E. coli cells. Thus, despite other potential advantages, this approach may not greatly expand the range of proteins that can be produced in a soluble, folded state, although it may be possible to overcome this limitation by redesigning the gene sequence (see below), adding chaperones or other factors (Kim and Swartz, 2000; Yin and Swartz, 2004), or reengineering ribosomal proteins (Chumpolkulwong et al., 2004).
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The Center for Eukaryotic Structural Genomics (CESG), in cooperation with Ehime University, Japan, and the CellFree Sciences Co. (Yokohama, Japan), is investigating the potential of wheat germ cell-free protein production (Madin et al., 2000; Sawasaki et al., 2002a,b; Kawasaki et al., 2003; Morita et al., 2003) as an enabling technology for NMR-based structural proteomics. CESG has developed the protein-production platform and has found that it offers potential advantages over E. coli cell and E. coli cell-free approaches. A larger proportion of targets are produced in folded, soluble form by the wheat germ cell-free approach than by the E. coli cell approach. In a comparison of 96 randomly chosen Arabidopsis thaliana targets carried through CESG’s wheat germ cell-free and E. coli cell pipelines to 15 N-labeled proteins, 8 proteins from the cell-free pipeline vs. 5 proteins from the cell-based pipeline were found suitable for NMR structural analysis on the basis of 1 H-15 N correlation NMR spectra (Tyler et al., 2005). The platform utilizes a single construct for all targets without any redesign of the DNA or RNA; thus it offers advantages over commercial cell-free methods utilizing E. coli extracts (Betton, 2003) that require multiple constructs or redesign of the ORF (e.g., Roche Protein Expression Group, http://www.rpeg.com/rpeg/content/custom protein.html#custom; ProteoExpert RTS E. coli HY, http://www.roche-applied-science .com/sis/proteoexpert). The wheat germ cellfree protocol employs no additives such as polyethylene glycol, which has been found to improve yields in E. coli S30 cell-free synthesis (Kim and Swartz, 2000). Because of the small volume requirements for screening (25 to 50 µl) and protein production for structural studies (4 to 12 ml), the method is amenable to automation. The high success rate of the approach in providing folded protein samples, coupled with the efficient level of incorporation of labeled amino acids makes the costs of the wheat germ cell-free method comparable to those of E. coli cell-base methods for protein production for NMR structure determinations. One of the major bottlenecks in highthroughput protein structure determination is the production of soluble, folded protein. Proteomics projects require the expression and purification of thousands of proteins and/or protein fragments. However, many individual proteins cannot be expressed in soluble form in bacteria. Insolubility arises either from an intrinsic property of a protein (e.g., ag-
gregation due to a very hydrophobic patch on the surface) or because a protein is not susceptible to folding mechanisms in the expression host, in which case there is an aggregation of folding intermediates. These insoluble proteins include one-third to one-half of all prokaryote proteins. This proportion is likely to be higher for eukaryotic proteins, particularly those that comprise multiple domains, require cofactors or protein partners for proper folding, or require extensive post-translational modification (see Protein Structure Initiative Target Database: http://www.rcsb.org/pdb/strucgen.html). For this reason, the development of improved platforms for the high-throughput production of soluble proteins or protein domains is an important goal. Optimal platforms for NMR spectroscopy must support the production of proteins labeled with stable isotopes (15 N, 13 C, and/or 2 H), and those for X-ray crystallography must enable incorporation of selenomethionine or other appropriately labeled amino acids. CESG, in cooperation with Ehime University, Japan and CellFree Sciences, Ltd., Japan, is investigating the potential of wheat germ cell-free protein translation as an enabling technology for structural genomics. The potential advantages of cell-free protein synthesis over other approaches are as follows: expression trials carried out without subcloning; improved yields of folded, soluble protein; amenability to automation since only small volumes are required; simplified protein purification (fewer contaminants, proteases); efficient incorporation of labeled amino acids without label scrambling; expression of proteins toxic to cells or degraded in cells; and the possibility of post-translation modifications and S-S bond formation. Two important applications of this technology are (1) facile assessment of whether a particular sequence can be produced in soluble form in adequate yields for proteomics investigations and (2) production of labeled proteins required for critical aspects of protein structure determination. The successful implementation of cellfree protein expression may minimize problems inherent in cell harvesting, cell lysis, and manipulations prior to using columns. Moreover, this approach may simplify purification, because the protein of interest is isolated from a smaller set of contaminants. Cell-free systems may also permit labeling strategies that cannot be achieved in whole cell systems, while potentially providing a substantial economy in the labeled material required to
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produce a target protein. This promise warrants serious consideration of how cell-free translation can contribute to high-throughput structural biology.
Critical Parameters and Troubleshooting
Wheat Germ Cell-Free Expression System
Most structural genomics centers are reporting overall success rates from target selection through structure determination and refinement in the range between 1% and 10%, with success rates lower for eukaryotic proteins. In the interest of efficiency and cost savings, is important to analyze where failures occur and to devise strategies to minimize them. The most effective routes for improvement involve a combination of bioinformatics and small-scale screening. Bioinformatics relies on prior information and mathematical models for correlating success rates with gene sequences. Small-scale screening offers the most economical way of testing whether a target will proceed through the critical stages leading to a structure: whether it has been cloned, supports the production of sufficient soluble protein, crystallizes for X-ray analysis, or is suitable for solution-state NMR analysis. The protocols described in this unit should allow one to take advantage of the efficiency and effectiveness of the wheat germ cell-free protein expression technology to rapidly and cost-effectively prescreen targets for expression and solubility (with and without the Nterminal tag) on the small scale, and foldedness, stability and suitability as a structural candidate on a large scale. In our experience, the cell-free approach is much less labor intensive than an E. coli cell-based pipeline, but it requires more expensive reagents and supplies. Current limitations of the method stem from the restricted availability and high cost of highly active wheat germ extract. These problems should ease as the wheat germ cell-free approach becomes more widespread and as demands for extract stimulate improvements in production technology. Currently the CESG purchases wheat germ extract from CellFree Sciences under a collaborative agreement. Current average supplies costs are: $87/target for cloning and expression-solubility testing, $276/mg for producing 15 N-labeled protein, and $436/mg for producing 13 C,15 N-labeled protein. The costs for labeled protein could be lowered by employing less expensive mixtures of labeled amino acids, tailoring the mixture of amino acids to match the amino acid composition of the target protein, or recovering and recycling
the unused amino acids. If amino acid mixtures are used, missing amino acids or those present at low abundance relative to the amino acid composition of the target protein must be added as supplements. CESG recently carried out a detailed cost analysis of its protein production pipelines, which considered the costs (labor and supplies) of individual steps (cloning, expression testing, and preparation of uniformly 15 Nlabeled and uniformly 13 C,15 N-labeled proteins) and took into account the yields at each step. The overall costs determined from this analysis were similar for the E. coli cell and wheat germ cell-free approaches to preparing samples for NMR structural investigations. The major advantage of the wheat germ cell-free method is that it supports the production of a larger fraction of targets as folded, soluble protein; the major advantage of the E. coli approach, when successful, is that larger quantities of protein are produced at a given cost.
Anticipated Results The authors’ overall success rate for producing highly soluble protein with an Nterminal (His)6 tag is 40% for Arabidopsis thaliana; 37% for human and 36% for mouse targets. “Highly soluble” indicates that the target protein is at least 75% soluble. In the case of targets from Arabidopsis GST-tagged proteins, 95% of target proteins remained >90% soluble after tag cleavage. The overall success rate for producing highly soluble protein with an N-terminal GST tag after the tag cleavage has been 49% for targets from Arabidopsis. Bioinformatics plays a key role in target selection and the resulting success rates.
Time Considerations One of the main advantages of the wheat germ cell-free protein expression is that the overall process is much less time and effort consuming when compared to the cellbased methods. Not including the cloning steps it generally takes 48 (using GeneDecoder1000TM) to 72 (manually) hr to screen 96 targets for expression and solubility on the small scale. The purification protocols are also less time and effort consuming owing to smaller volumes. Using the latest in GE Healthcare HisTrap purification technology, IMAC purification of His-tagged proteins requires 2 hr of processing time and results in 75% to 85% pure protein samples. Gel filtration adds an additional 3 hr and can increase the purity to >95% for proteins
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<15 kDa and to 90% for proteins <20 kDa. GST purification results in >95% purity regardless of size, however, minimal time to process the sample is greater than 10 hr.
Kigawa, T. and Yokoyama, S. 2002. Highthroughput cell-free protein expression system for structural genomics and proteomics studies. Tanpakushitsu Kakusan Koso 47:10141019.
Acknowledgments
Kigawa, T., Muto, Y., and Yokoyama, S. 1995. Cellfree synthesis and amino acid-selective stable isotope labeling of proteins for NMR analysis. J. Biomol. NMR 6:129-134.
Research carried out at the CESG was supported by the NIH Protein Structure Initiative through grant 1 P50 GM64598 to John L. Markley. The authors thank Professor Yaeta Endo of Ehime University (Matsuyama, Japan) and CellFree Sciences Co., Ltd. (Yokohama, Japan) for encouragement in using the wheat germ cell-free system.
Literature Cited Betton, J.M. 2003. Rapid translation system (RTS): A promising alternative for recombinant protein production. Curr. Protein Pept. Sci. 4:73-80. Bruggert, M., Rehm, T., Shanker, S., Georgescu, J., and Holak, T.A. 2003. A novel medium for expression of proteins selectively labeled with 15 N-amino acids in Spodoptera frugiperda (Sf9) insect cells. J. Biomol. NMR 25:335-348. Chrunyk, B.A., Evans, J., Lillquist, J., Young, P., and Wetzel, R. 1993. Inclusion-body formation and protein stability in sequence variants of interleukin-1-beta. J. Biol. Chem. 268:1805318061. Chumpolkulwong, N., Hori-Takemoto, C., Hosaka, T., Inaoka, T., Kigawa, T., Shirouzu, M., Ochi, K., and Yokoyama, S. 2004. Effects of Escherichia coli ribosomal protein S12 mutations on cell-free protein synthesis. Eur. J. Biochem. 271:1127-1134. Clemens, M.M. and Prujin, G.J. 1999. In Protein Expression. A practical approach. (S.J. Higgens. and B.D. Hames, eds.) pp. 129-165. Oxford University Press, Oxford. Cubeddu, L., Moss, C.X., Swarbrick, J.D., Gooley, A.A., Williams, K.L., Curmi, P.M., Slade, M.B., and Mabbutt, B.C. 2000. Dictyostelium discoideum as expression host: Isotopic labeling of a recombinant glycoprotein for NMR studies. Protein Expr. Purif. 19:335-342. Goff, S.A. and Goldberg, A.L. 1987. An increased content of protease LA, the lon gene-product, increases protein-degradation and blocks growth in Escherichia coli. J. Biol. Chem. 262:45084515. Guignard, L., Ozawa, K., Pursglove, S.E., Otting, G., and Dixon, N.E. 2002. NMR analysis of in vitro-synthesized proteins without purification: A high-throughput approach. FEBS Lett. 524:159-162. Henrich, B., Lubitz, W., and Plapp, R. 1982. Lysis of Escherichia coli by induction of cloned PhiX174 genes. Mol. Gen. Genet. 185:493-497. Kawasaki, T., Gouda, M.D., Sawasaki, T., Takai, K., and Endo, Y. 2003. Efficient synthesis of a disulfide-containing protein through a batch cell-free system from wheat germ. Eur. J. Biochem. 270:4780-4786.
Kigawa, T., Yabuki, T., Yoshida, Y., Tsutsui, M., Ito, Y., Shibata, T., and Yokoyama, S. 1999. Cell-free production and stable-isotope labeling of milligram quantities of proteins. FEBS Lett. 442:15-19. Kim, D.M. and Swartz, J.R. 2000. Prolonging cellfree protein synthesis by selective reagent additions. Biotechnol. Prog. 16:385-390. Kim, D.M., Kigawa, T., Choi, C.Y., and Yokoyama, S. 1996. A highly efficient cell-free protein synthesis system from Escherichia coli. Eur. J. Biochem. 239:881-886. Klammt, C., Lohr, F., Schafer, B., Haase, W., Dotsch, V., Ruterjans, H., Glaubitz, C., and Bernhard, F. 2004. High level cell-free expression and specific labeling of integral membrane proteins. Eur. J. Biochem. 271:568-580. Kramer, G., Kudlicki, W., and Hardesty, B. 1999. Cell-free coupled transcription-translation systems from Escherichia coli. In Protein Expression: A Practical Approach (S.J. Higgens and B.D. Hames, eds.) pp. 201-223. Oxford University Press, Oxford. Madin, K., Sawasaki, T., Ogasawara, T., and Endo, Y. 2000. A highly efficient and robust cell-free protein synthesis system prepared from wheat embryos: Plants apparently contain a suicide system directed at ribosomes. Proc. Natl. Acad. Sci. U.S.A. 97:559-564. Maurizi, M.R. 1987. Degradation in vitro of bacteriophage lambda N protein by Lon protease from Escherichia coli. J. Biol. Chem. 262:2696-2703. Morita, E.H., Sawasaki, T., Tanaka, R., Endo, Y., and Kohno, T. 2003. A wheat germ cell-free system is a novel way to screen protein folding and function. Protein Sci. 12:1216-1221. Sawasaki, T., Hasegawa, Y., Tsuchimochi, M., Kamura, N., Ogasawara, T., Kuroita, T., and Endo, Y. 2002a. A bilayer cell-free protein synthesis system for high-throughput screening of gene products. FEBS Lett. 514:102-105. Sawasaki, T., Ogasawara, T., Morishita, R., and Endo, Y. 2002b. A cell-free protein synthesis system for high-throughput proteomics. Proc. Natl. Acad. Sci. U.S.A. 99:14652-14657. Shi, J., Pelton, J.G., Cho, H.S., and Wemmer, D.E. 2004. Protein signal assignments using specific labeling and cell-free synthesis. J. Biomol. NMR 28:235-247. Strauss, A., Bitsch, F., Cutting, B., Fendrich, G., Graff, P., Liebetanz, J., Zurini, M., and Jahnke, W. 2003. Amino-acid-type selective isotope labeling of proteins expressed in Baculovirusinfected insect cells useful for NMR studies. J. Biomol. NMR 26:367-372.
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Torizawa, T., Terauchi, T., and Kainosho, M. 2002. Recent developments in NMR methods for structural biology. Seikagaku 74:1279-1284. Tyler, R.C., Aceti, D.J., Bingman, C.A., Cornilescu, C.C., Fox, B.G., Frederick, R.O., Jeon, W.B., Lee, M.S., Newman, C.S., Peterson, F.C., Phillips, G.N., Shahan, M.N., Singh, S., Song, J., Sreeenath, H.K., Tyler, E.M., Ulrich, E.L., Vinarov, D.A., Vojtik, F.C., Volkman, B.F., Wrobel, R.L., Zhao, Q., and Markley, J.L. 2005. Comparison of cell-based and cell-free protocols for producing target proteins from the Arabidopsis thaliana genome for structural studies. Proteins 59:633-643. Vinarov, D.A., Lytle, L.B., Peterson, F.C., Tyler, E.M., Volkman, B.F., and Markley, J.L. 2004. Eukaryotic cell-free structural genomics: NMR structure of a beta-grasp fold protein from Arabidopsis thaliana. Nat. Methods 1:149-153. Yabuki, T., Kigawa, T., Dohmae, N., Takio, K., Terada, T., Ito, Y., Laue, E.D., Cooper, J.A., Kainosho, M., and Yokoyama, S. 1998. Dual amino acid-selective and site-directed stableisotope labeling of the human c-Ha-Ras protein by cell-free synthesis. J. Biomol. NMR 11:295306. Yin, G. and Swartz, J.R. 2004. Enhancing multiple disulfide bonded protein folding in a cell-free system. Biotechnol. Bioeng. 86:188-195. Yokoyama, S. 2003. Protein expression systems for structural genomics and proteomics. Curr. Opin. Chem Biol. 7:39-43. Yokoyama, S., Hirota, H., Kigawa, T., Yabuki, T., Shirouzu, M., Terada, T., Ito, Y., Matsuo, Y., Kuroda, Y., Nishimura, Y., Kyogoku, Y., Miki, K., Masui, R., and Kuramitsu, S. 2000. Structural genomics projects in Japan. Nat. Struct. Biol. 7:943-945.
Klammt et al., 2004. See above. The authors demonstrate high level expression of integral membrane proteins (IMPs) in a cell-free coupled transcription/translation system using a modified Escherichia coli S30 extract preparation. Madin et al., 2000. See above. One of the original papers by Endo and colleagues describing the development and evolution of the wheat germ cell-free protein production. Tyler et al., 2005. See above. The authors describe a comparative study of protein production from 96 Arabidopsis thaliana open reading frames (ORFs) by cell-based and wheat germ cell-free protocols. Vinarov et al., 2004. See above. Describes a wheat germ cell-free platform for protein production that supports efficient NMR structural studies of eukaryotic proteins and offers advantages over cell-based methods. Yokoyama, 2003. See above. Discusses application of the E. coli cell-free protein synthesis system for large-scale production of protein samples for NMR (stable-isotope labeling) and X-ray crystallography (selenomethionine substitution).
Contributed by Dmitriy A. Vinarov, Carrie L. Loushin Newman, Ejan M. Tyler, and John L. Markley University of Wisconsin-Madison Madison, Wisconsin Mark N. Shahan Quintessence Biosciences, Inc. Madison, Wisconsin
Key References Kigawa et al., 1995. See above. Discusses the applicability and advantages of E. coli cell-free protein synthesis for selective stable isotope labeling of proteins for the application of multidimensional NMR spectroscopy to larger proteins.
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CHAPTER 6 Purification of Recombinant Proteins INTRODUCTION
T
he principle aim of protein overexpression using the various systems described in Chapter 5 and elsewhere (e.g., Chapter 16 of Ausubel et al., 1997) is to obtain purified protein, and that is the subject matter of this chapter. The purification of any protein, recombinant or nonrecombinant, is critically dependent on the starting material (or source). Some natural proteins—for example, cellular receptors and growth factors—may be present in biological source material only in trace amounts. In these cases, complex purification schemes are required to obtain micro-amounts of partially or completely pure protein (Simpson and Nice, 1989, and references therein). With recombinant sources, the overproduced protein is usually (though not always) abundant, and hence the amount available is not generally a critical factor. The gene expressing the recombinant protein must of course have been previously cloned, and in fact may have been done so using oligonucleotide probes derived from the amino acid sequence data, in turn derived from the authentic protein isolated by (to use Scope’s expression) “heroic efforts” (UNIT 1.1). What is at issue with overproduced proteins made by the various host/vector systems is their authenticity, in terms of both noncovalent (i.e., conformational—is the protein correctly folded or aggregated?) and covalent structure (i.e., is the correct disulfide bond pattern present and are post-translational modifications such as glycosylation required?). In UNIT 6.1, an overview of protein purification is presented; although purification from E. coli is emphasized, other host systems are also referred to where appropriate. Purification of recombinant protein can be aided by the fact that the investigator knows the primary sequence of the recombinant protein and can thus estimate its isoelectric point. Furthermore, the purified protein can thus be checked against this theoretical sequence for artificial modifications arising from the expression system and purification method using some of the methods summarized in Chapter 7. This unit has been recently updated and expanded (2002) and forthcoming revisions will include more detailed discussions of purification characteristics from hosts other than E. coli. If the recombinant source contains the protein to be purified in a soluble state, then the protein can be isolated using standard methods, often involving column chromatography. In UNIT 6.2, the purification of interleukin 1β (IL-1β) is used to illustrate the basic approach for purifying an abundant and soluble protein expressed in E. coli. Using the basic fractionation and chromatographic approach described, it is often possible to purify other soluble recombinant proteins. For situations where the starting material is less abundant, albeit soluble (e.g., proteins secreted into the medium, or, in the case of E. coli, the periplasmic space), higher-resolution techniques such as biospecific affinity chromatography may be required. These are discussed in UNIT 6.1 and Chapter 9. The expression of proteins genetically engineered to contain various affinity tags or carrier proteins that aid purification are reviewed in UNIT 5.1 as well as in UNITS 6.1, 6.5, 6.6 & 6.7 of this chapter. IL-1β is somewhat of an ideal protein from the viewpoint of both expression and purification. The authentic protein is not post-translationally modified and although it contains two sulfhydryl groups, it contains no disulfide bonds. For this protein, the authentic and recombinant forms have the same covalent structure, as is discussed in detail in UNIT 6.2 (Background Information). Contributed by Paul T. Wingfield Current Protocols in Protein Science (2002) 6.0.1-6.0.4 Copyright © 2002 by John Wiley & Sons, Inc.
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In many cases high-level expression of protein in the E. coli host leads to the formation of highly aggregated proteins, called inclusion bodies. The determination of whether a protein is soluble or not is made using cell lysates and simple centrifugation schemes as outlined in UNIT 6.1 (see also UNIT 5.3). Proteins in inclusion bodies usually have the correct primary sequences, identical to those of their native and authentic counterparts, but are aggregated and inactive due to noncovalent or conformational differences. In UNIT 6.3, the preparation of insoluble inclusion body protein from E. coli lysates by a simple washing procedure is described. Before insoluble proteins in inclusion bodies can be purified, they must be extracted and solubilized; in this respect their purification is analogous to the extraction of intrinsic membrane proteins from membranes. The solubilization of inclusion body protein, however, requires denaturation with reagents such as concentrated solutions of urea or guanidine⋅HCl (see APPENDIX 3A), whereas the extraction of membrane proteins is normally carried out under nondenaturing conditions using nonionic or zwitterionic detergents (von Jagow et al., 1994). As the solubilized recombinant protein is denatured or unfolded, it must be folded into the correct tertiary and quaternary structures in order for the protein to acquire its functional and biological properties. Many nonrecombinant proteins that have been isolated under native conditions can be reversibly denatured and renatured using well-established methods. A concise account of protein folding is given in UNIT 6.4 (see also Pain, 1994) to provide some of the theoretical background required to interpret protein folding experiments. The preparation of folded and active recombinant proteins from inclusion bodies involves the process known as “preparative protein folding.” In traditional protein folding experiments, pure native proteins such as ribonuclease were used to study the reversible denaturation/renaturation process to gain information about folding pathways (Pain, 1994). These experiments are usually carried out using low protein concentrations (in the micrograms per milliliter range) to minimize nonspecific inter- and intramolecular interactions that lead to aggregation. In preparative protein folding, the aim is to obtain folded protein in as high a yield as possible using relatively high starting concentrations (in the milligrams per milliliter range) and without using very large reaction volumes. The various strategies usually used to achieve this are illustrated in UNIT 6.5. In preparative protein folding, apart from the need to avoid or minimize aggregation, formation of the correct disulfide bonds is required. After extraction of the recombinant protein from inclusion bodies using a protein denaturant and reducing agent (UNIT 6.3), the solubilized protein is unfolded and sulfhydryl groups, if present, are reduced. To form the correctly folded protein, disulfide bond formation and protein folding must both occur, and usually in a concomitant manner. Folding pure or partially purified protein may be advantageous, leading to higher yields of folded protein. In UNIT 6.3, gel filtration in the presence of guanidine⋅HCl is described as a simple and rapid method for partially purifying extracted and denatured proteins prior to folding. A recent example of this approach is the production of the ectodomain of HIV-1 and SIV gp41 (Wingfield et al., 1997). Other methods that can be used to purify proteins in high concentrations of urea or guanidine⋅HCl are discussed in UNIT 6.1.
Introduction
Three examples of the folding and purification of insoluble proteins expressed in E. coli are described in UNIT 6.5. Bovine growth hormone (somatotropin) is extracted from inclusion bodies using guanidine⋅HCl; the protein is then folded and then purified by conventional chromatographic methods. Aggregation during the folding process is minimized by using a solvent additive cosolvent, in this case urea in a limiting (i.e., nonde-
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naturing) concentration. Oxidation of the two disulfide bonds is enhanced by inclusion of an oxido-shuffling system based on oxidized and reduced glutathione (UNITS 6.1 & 6.4). In the second example, purification of human interleukin-2 (T cell growth factor), a different approach is used. The protein is extracted from inclusion bodies using acetic acid, then fractionated by gel filtration in the same acid. The partially purified protein is folded and air-oxidized by the simple approach of dialysis against water. Because interleukin-2 contains three sulfhydryl groups, only two of which are involved in an intramolecular disulfide bond, it is possible for incorrect disulfide bonds to form during folding. An HPLC-based support protocol is included that allows rapid determination of the disulfide bonding pattern. Finally, the folded protein is purified to homogeneity using preparative reversed-phase HPLC. The acid stability of the protein is clearly an advantage and necessary for the success of the overall process. The third example describes the purification of the catalytic domain of the immunodeficiency virus (HIV-1) integrase. This protein is essential for the HIV-1 life cycle and is therefore a potential drug target. The integrase catalytic domain is expressed as a fusion protein containing an additional 20 amino acids at the N-terminus. This “fusion domain” includes a tract of six histidine residues and is commonly called a His tag. The expression of proteins with His tags is fairly common practice, as it allows proteins to be purified by metal-chelate chromatography (MCAC) under both native and denaturing conditions (see UNIT 5.1 for additional discussion). The HIV-1 integrase, extracted with denaturant and partially purified by gel filtration and MCAC, is folded into native protein using a slow and continuous addition of denatured protein into a renaturation solution. This approach was originally described by Ruldolph and his colleagues (see Ruldolph, 1989, and references therein) and is generally applicable for avoiding and/or minimizing aggregation during folding. The His tag is removed from the folded fusion protein using specific protease digestion (in this example, the protease used is thrombin, and methods for its handling and specific removal are also described). A widely used method for producing fusion proteins in E. coli is the glutathione S-transferase (GST) system (UNIT 5.1). Analogous to His tag fusions, there is a simple affinity chromatographic method for purification of GST fusions using immobilized glutathione as an affinity matrix. The affinity purification of GST fusions is probably more specific than the metal chelate chromatography used for His tags, but it has the limitation that it cannot be performed under denaturing conditions. Fusion proteins usually contain a linker region with a specific proteolytic cleavage site, and often this site is specific for thrombin. Thus, once the GST fusion protein has been purified, the protein moiety can be readily released by proteolysis. Further purification may include one more round of affinity chromatography (to remove GST) and a gel filtration step to remove excess thrombin and other minor contaminates. Purification may be possible by performing only the gel filtration step, depending on the size of various protein products (GST is a dimer of 51 kDa and thrombin is ∼36 kDa). In UNIT 6.6 the expression and purification of GST fusion proteins is described. Protocols for protein expression using shaker cultures and purification methods of soluble GST fusion proteins are included. If the GST fusion protein is insoluble and forms inclusion bodies, the protein must be solubilized and folded into a native-like conformation prior to affinity chromatography. An extraction and folding protocol using the protein denaturant urea is described. (It should be noted that the GST moiety can be readily folded from either urea or guanidine⋅HCl denatured samples.) Protocols for proteolytic cleavage of the fusion protein and further purification are also described. Structural biologists may note with interest the recent paper by Kuge et al. (1997) in which the crystallization of a GST fusion protein was achieved.
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Another popular gene fusion system is based on the E. coli intracellular enzyme thioredoxin. This 12-kDa protein is very soluble, is monomeric, and has been thoroughly characterized (UNIT 5.1). The target protein can be positioned either N-terminal (proteinthioredoxin) or C-terminal (thioredoxin-protein) to the fusion partner. This dual orientation, also possible with Hist-tag fusions, represents a difference from GST fusions, as the GST moiety is normally only N-terminal. It is worth noting that as thioredoxin is a monomeric protein, it may be a more suitable fusion partner for oligomeric proteins than, for example, dimeric GST (Hurd and Hornby, 1996). In UNIT 6.7 the construction and expression of thioredoxin fusion proteins are discussed. Support protocols deal with the release of the fusion proteins by mechanical lysis and osmotic stress. E. coli–derived thioredoxin exhibits thermal stability, and a heat treatment of cell lysates at 80°C may be a useful initial purification step. This approach is described in the third support protocol. The commentary section includes a brief discussion of more specific purification methods. However, if ease of purification is of critical importance, the GST fusion system should be considered as the glutathione-based affinity method is probably more efficient than the corresponding affinity-based method used for thioredoxin. It should be emphasized that although most of the protocols described in this chapter are applied to specific examples, they are fairly representative of the methods and approaches used in general for laboratory-scale preparation of recombinant proteins from E. coli. The individual characteristics of a given protein (e.g., solubility, isoelectric point, pH stability, and number of sulfhydryl residues) will dictate the conditions required for any particular purification step or folding process. Method optimization is usually empirical unless characterization of the purified protein provides useful guidelines. LITERATURE CITED Ausubel, F.M., Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G., Smith, J.A., and Struhl, K.S. (eds.). 1997. Current Protocols in Molecular Biology. John Wiley & Sons, New York. Hurd, P.J. and Hornby, D.P. 1996. Expression systems and fusion proteins. In Proteins LabFax (N.C. Price, ed.) pp. 109-117. BIOS Scientific Publishers, Oxford, UK. Kuge, M., Fujii, Y., Shimizu, T., Hirose, F., Matsukage, A., and Hakoshima, T. 1997. Use of a fusion protein to obtain crystals suitable for X-ray analysis: Crystallization of a GST-fusion protein containing the DNA-binding domain of DNA replication-related element-binding factor, DREF. Protein Sci. 6:17831786. Pain, R.H. 1994. Mechanisms of Protein Folding. IRL Press, Oxford, UK. Ruldolph, R. 1989. Renaturation of recombinant, disulfide-bonded proteins from “inclusion bodies.” In Modern Methods in Protein and Nucleic Acid Research: Review Articles (H. Tschesche, ed.) pp. 149-171. Walter de Gruyter, Berlin. Simpson, R.J. and Nice, E.C. 1989. Strategies for the purification of subnanomole amounts of proteins and polypeptides for microsequence analysis. In The Use of HPLC in Receptor Biochemistry (A.R. Kerlavage, ed.) pp. 210-244. Alan R. Liss, New York. von Jagow, G., Link, T.A., and Schägger, H. 1994. Purification strategies for membrane proteins. In A Practical Guide to Membrane Protein Purification (G. von Jagow and H. Schägger, eds.) pp. 3-21. Academic Press, San Diego. Wingfield, P.T., Stahl, S.J., Kaufman, J., Zlotnick, A., Hyde, C.C., Gronenborn, A.M., and Clore, G.M. 1997. The extracellular domain of immunodeficiency virus gp41 protein: Expression in Escherichia coli, purification, and crystallization. Protein Sci. 6:1653-1660.
Paul T. Wingfield
Introduction
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Overview of the Purification of Recombinant Proteins Produced in Escherichia coli
UNIT 6.1
The expression of recombinant proteins, especially using bacterial vectors and hosts, is a mature technology. With the appropriate cDNA and PCR methods, expression plasmids can be rapidly produced. Following sequence determination of the constructs, plasmids are transformed into expression hosts, single colonies picked, and fermentation performed. With E. coli, a 2-liter fermentation using complex media will generate ∼50 to 80 g (wet weight of cells). Assuming modest protein expression (2% to 5% of the total cellular protein), between 100 and 300 mg of recombinant protein is available in the cells. The problem is, of course, how to isolate it in an active form. Soluble proteins can be recovered with good yields (>50%), and insoluble proteins, which must undergo a denaturation and folding cycle, can be recovered with more modest yields (5% to 20%). Hence, using small-scale fermentations and laboratory-scale processing equipment, proteins (or subdomains thereof) can usually be produced in sufficient quantities (10 to 100 mg) to initiate most studies including detailed structural determinations. Some strategies for achieving high-level expression of genes in E. coli have been reviewed by Makrides (1996) and Baneyx (1999). Some of the above characteristics also hold true for the production of proteins using yeast and baculovirus eukaryotic expression systems, although more effort and expertise is required to construct the vectors and, with the baculovirus system, produce cells for processing. A yeast expression system may be a wise choice for proteins that form insoluble inclusions in bacteria, and for the production of membrane-associated proteins (Cereghino and Clegg, 1999; UNITS 5.6-5.8). The baculovirus system has proven very useful for producing phosphorylated proteins and glycoproteins (Kost, 1999; UNITS 5.4-5.5) and for the co-expression of interacting proteins. The construction of stable mammalian protein expression vectors requires considerably more time and effort but may be the only approach for producing complex multidomain proteins (UNITS 5.9-5.10). Cells growing to cell densities of 1-5 × 109 cells/ml can be expected to typically secrete >10 mg/liter of product. Alternatively, transient gene expression systems using various viral vectors (e.g., vaccinia virus; UNITS 5.12-5.15), can be used to produce lesser amounts of protein, which is useful for feasibility studies. It is of interest to note that the large-scale transient expression systems in mammalian cells are being actively developed by biotechnology companies (Wurum and Bernard, 1999). The initial choice of host system for the production of recombinant proteins for many investigators is Escherichia coli. This is due to such factors as ease of genetic manipulation, availability of optimized expression plasmids, and ease of growth. This unit presents an overview of recombinant protein purification with special emphasis on proteins expressed in E. coli. Practical aspects and strategies are stressed throughout, and wherever possible, the discussion is cross-referenced to the example protocols described in the rest of Chapter 6. The first section deals with information pertinent to protein purification that can be derived from translation of the cDNA sequence. This is followed by a brief discussion of some of the common problems associated with bacterial protein expression (see also UNIT 5.1). Planning a protein purification strategy requires that the solubility of the expression product be determined; it is also useful to establish the location of the protein in the cell—e.g., cytoplasm or periplasm. This unit includes flow charts that summarize approaches for establishing solubility and localization of bacterially produced proteins (see also UNIT 5.2). Contributed by Paul T. Wingfield Current Protocols in Protein Science (2002) 6.1.1-6.1.37 Copyright © 2002 by John Wiley & Sons, Inc.
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Purification strategies for both soluble and insoluble proteins are reviewed and summarized in flow charts (see also Chapter 1). Many of the individual purification steps, especially those involving chromatography, are covered in detail in Chapters 8 and 9, and elsewhere (Scopes, 1994; Janson and Ryden, 1998). The methodologies and approaches described here are essentially suitable for laboratory-scale operations. Large-scale methodologies have been previously reviewed (Asenjo and Patrick 1990; Thatcher, 1996; Sofer and Hagel, 1997). A section on glycoproteins produced in bacteria in the nonglycosylated state is included to emphasize that, although they may not be useful for in vivo studies, such proteins are well suited for structural studies. The final sections deal with protein handling, scale and aims of purification, and specialized equipment needed for recombinant protein purification and characterization. PROTEIN SEQUENCE AND COMPOSITIONAL ANALYSIS Analyzing the Protein Sequence The protein sequence translated from the DNA coding sequence is usually available, and before attempting any laboratory work, it is useful to carry out a literature survey and basic computer analyses (see Chapter 2). First, if the natural protein has been isolated and characterized, reviewing the physicochemical properties of the protein and the established purification techniques used may aid in planning a strategy for isolation from the recombinant host. Recombinant proteins that accumulate as insoluble aggregates or inclusion bodies, require folding into native-like conformations (Lilie et al., 1998; De Bernardez Clark et al., 1999; UNIT 6.4). Information on the conformational properties, including denaturation/folding curves, can help rationalize the development of preparative folding processes. Second, for uncharacterized proteins, analyses of related proteins with sequence similarities or known motifs may provide useful clues for selecting purification steps (UNIT 2.1; see also the PROSITE database of protein families and domains at the ExPASy Molecular Biology Server at http://ca.expasy.org/prosite). For example, if the protein contains the well-known kringle domain, lysine affinity chromatography might be a successful purification technique (Cleary et al., 1989). On the other hand, if the protein contains no recognizable motifs and has no similarity to other proteins, yet contains many cysteine residues, other strategies and precautions would be warranted as described in UNITS 6.3-6.5. The amino acid sequence can be used to direct the synthesis of peptides corresponding to potential epitopes (e.g., 10 to 20 residues; UNIT 2.2). Polyclonal antibodies raised against the peptides may be suitable for detecting the protein of interest by immunoblotting. This approach may be especially valuable for monitoring proteins expressed at low levels— e.g., when E. coli secretion vectors are used. The antibodies may also be useful for immunoaffinity chromatography. Analyzing the Amino Acid Composition
Purification of Recombinant E. coli Proteins
The amino acid composition (UNIT 3.2) of the protein will also allow calculation of some basic physicochemical parameters. Using average pKa values for ionizable side chains in proteins (Matthew et al., 1978), the isoelectric point (pI) can be estimated by applying the well-known Henderson-Hasselbach relationship. The calculations can be performed using an electronic spreadsheet such as Excel or via the internet using one of the many molecular biology servers, e.g., ExPASy (http://www.expasy.ch/tools/pi_tool.html). The values obtained, although only approximate, are useful for guiding the initial selection of
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ion-exchange resins and the pH of column buffers. When eukaryotic hosts are selected for protein expression, it should be noted that post-translational modifications such as phosphorylation and glycosylation will affect the pI. Another parameter that can be estimated from the amino acid composition is the extinction coefficient (ε), usually at 280 nm (Pace et al., 1995). Although this information will be more useful when the protein has been purified, as most columns are monitored by UV absorption, proteins with an unusually low ε (no tryptophan and little or no tyrosine) may be difficult to detect during the early stages of purification. Other physicochemical parameters that can be calculated include hydrodynamic parameters such as molecular radii and sedimentation coefficients, the program SEDNTERP is especially useful (http://www.jphilo.mailway.com/download.htm). These parameters may help in interpreting results of gel-filtration and centrifugational separations. CHARACTERISTICS OF THE HOST-VECTOR SYSTEM Choosing an Expression System Popular protein expression systems include E. coli, yeast, baculovirus-infected insect cells, and cultured mammalian cell lines (see Chapter 5). If the requirement is to obtain a protein post-translationally modified via glycosylation (see Chapter 12) or phosphorylation (see Chapter 13), then a eukaryotic expression system must be used. Stable mammalian expression systems are the most time-consuming to establish and require the most expertise; however, they may be the only successful system for certain requirements including, e.g., proteins with authentic glycosylation patterns; large multidomain and multisubunit proteins, and especially proteins that are insoluble in E. coli. Post-translational modifications may aid purification (e.g., lectin affinity chromatography can be used for glycoproteins; UNIT 9.1). On the other hand, these modifications may introduce charge heterogeneity—as is commonly observed with glycosylation due to loss of sialic acid— which may then complicate purification, especially with methods such as ion-exchange chromatography (UNIT 8.2). Specific modification of proteins expressed in E. coli can be achieved by the co-expression of modifying enzymes, such as phosphorylation of tryrosyl residues by tyrosine kinase (Ren and Schaefer, 2001; http://www.stratagene.com/manuals/200124.pdf). However, most of the post-translational modifications observed in E. coli are nonspecific, such as deamidation (Wingfield et al., 1987a) and proteolytic clipping (Nagata et al., 1986). Other less common sources of protein heterogeneity arising from E. coli expression are: (1) internal starts in translation (Dale et al., 1994); (2) partial readthroughs of the termination codon (Danley et al., 1991), and (3) translation errors (Lu et al., 1993). The initial choice for protein expression is often E. coli but if direct expression of a protein of interest fails or yields an insoluble product, there are many other options available including generating fusion proteins and many other approaches discussed elsewhere in this overview. If other expression hosts are to be screened, there are universal cloning systems commercially available (e.g., Gateway cloning system at http://www.invitrogen.com) that allow the rapid transfer of the gene of interest into multivector systems including yeast, baculovirus, and mammalian cells. Minimizing Proteolysis If the protein is expressed in the cytoplasm in a soluble state, the purification can be carried out directly after cell lysis. Soluble recombinant proteins are, however, susceptible to proteolysis, which can occur before or after extraction from the cell (Maurizi, 1992). Choosing protease-deficient E. coli host strains (Goff and Goldberg, 1985), manipulating growth conditions, especially the time of induction for inducible promoters (Allet et al.,
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1988), and using exogenous protease inhibitors can minimize this problem. Nevertheless, more extreme steps may be required, such as inducing the expressed protein to form insoluble inclusion bodies, using a secretion vector to locate the protein to the periplasm or medium, and changing to a eukaryotic expression system. In addition, there is a protein engineering approach that requires knowledge of the proteolytic cleavage site(s) to stabilize the protein. It requires alteration of one or both of the residues forming the scissile bond by site-directed mutagenesis (Mildner et al., 1994). For more detailed discussions on strategies to minimize proteolytic degradation, see reviews by Murby et al. (1996) and Makrides (1996). Removing the Amino-Terminal Methionine Another common problem with proteins expressed directly in E. coli is retention of the N-terminal methionine derived from the initiating N-formylmethionine (the formyl group is almost always removed). The N-terminal methionine is generally removed when the second amino acid is alanine, glycine, proline, serine, threonine, or valine (cleavable residues), but not when it is arginine, asparagine, aspartic acid, glutamic acid, glutamine, isoleucine, leucine, lysine, or methionine (noncleavable residues; Sherman et al., 1985). When recombinant proteins are expressed at very high levels, the N-terminal methionine can be retained regardless of the nature of the second amino acid, presumably due to saturation of the processing enzymes or depletion of required metal cofactors. Removal of cleavable N-terminal methionine can be carried out in vitro by digestion with purified methionine aminopeptidase (Miller et al., 1987) or by co-expression of the processing enzyme (Ben-Bassat et al., 1987; Hwang, et al., 1999). The need to remove noncleavable methionines can be circumvented by incorporating an N-terminal secretion leader sequence that localizes the protein, minus the leader, to the periplasmic space (Holland et al., 1990). Other approaches utilize the incorporation of N-terminal fusions with, e.g., ubiquitin, which can be cleaved in vitro or in vivo with a processing enzyme (ubiquitin hydrolase). This approach involves co-expression of the hydrolase (Miller et al., 1989). Finally, it should be noted that it is sometimes possible to resolve proteins containing N-terminal Met from those lacking it by chromatographic methods (Wingfield et al., 1987b). Dealing with Inclusion Bodies The expression of eukaryotic proteins in E. coli often leads to the accumulation of insoluble protein called inclusion bodies (UNITS 6.3 & 6.5). Inclusion bodies can be easily observed by phase-contrast microscopy as dense bodies, usually located at the polar extremities of the cell and they can be isolated by centrifugation (Georgiou and Valax, 1999).
Purification of Recombinant E. coli Proteins
The rate of protein biosynthesis in prokaryotes is about ten times faster than in eukaryotes. Comparison of the rates of in vitro refolding of orthologous prokaryotic and eukaryotic proteins indicates that the former refolds six times faster. This suggests that the rate of folding correlates with the rate of elongation of polypeptide chains. Hence, part of the problem in expressing eukaryotic proteins in bacteria might be due to combination of fast synthesis and slow folding, which favors aggregation (Widmann and Christen, 2000). Proteins in the unfolded state at high concentration, even small rapidly folding proteins, are prone to aggregation due to exposure of hydrophobic surfaces that are normally buried in the native state (see Fersht, 1999, for further discussion). Some proteins are helped to fold in vivo by binding to accessory proteins called chaperones (see Protein-Assisted Folding and Oxidation in the discussion of Performing Protein Folding).
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The formation of inclusion bodies can occasionally protect proteins against proteolysis and can also allow accumulation of proteins normally toxic to the cell; some examples include proteases (HIV-1 protease; Cheng et al., 1990) and membrane-spanning domains (Jones et al., 2000). The formation of inclusion bodies also simplifies purification of the protein, albeit in a denatured/aggregated state (see below). The main disadvantage is that the protein must be extracted with protein denaturants and then folded into a native-like conformation. For small (10- to 20-kDa) single-domain proteins, this is usually not problematic, although the overall recoveries may only be 5% to 20% of those of similar or identical proteins expressed in a soluble state. For large (40- to 70-kDa), multiple-domain proteins, recoveries may be negligible, although there have been a number of successful cases, such as the 69-kDa tissue plasminogen activator (Grunfeld et al., 1992). The formation of inclusion bodies can sometimes be prevented by changing the promoter, host strain, and combinations thereof; controlling the growth conditions (especially the pH of the culture); adding nonmetabolizable sugars such as sucrose and sorbitol to the fermentation medium; and changing the temperature of induction, usually by lowering it (for reviews, see Schein, 1989; Wetzel, 1992; Baneyx, 1999). The recombinant protein may be located in both the insoluble and soluble fractions of the cell (mixed-phase expression), and in these cases, better yields may be obtained by processing the soluble material (discarding the insoluble) even though it might constitute only a minor portion of the total expressed protein (Thatcher and Panayotatos, 1986; Wingfield et al., 1987c). Soluble protein purified from mixed-phase expressions should be carefully analyzed to check its authenticity (e.g., by mass spectrometry) as the solubility may have resulted from minor modifications such as deamidation or proteolysis of a few residues from either the N or C terminus (P.T. Wingfield unpub. observ.). A successful approach for avoiding inclusion body formation is the use of an appropriate secretion vector (Guisez et al., 1998; Cornelius, 2001) The N-terminal secretion signal directs protein to the periplasmic space (see Localizing Protein), and translocation across the plasma membrane results in cleavage of the secretion leader sequence. The periplasm contains enzymes that accelerate folding and formation of disulfide bonds (for reviews, see Missiakas and Raina 1997). Purification is also simplified as the protein content of the periplasmic space constitutes only 4% of the total E. coli protein (Beacham, 1979). Fusion Proteins Apart from direct expression, there are many examples of fusion protein expression. Fusion proteins consist of the protein of interest partnered or tagged with proteins or protein domains appended to either the N- or C-terminus (or both) (UNIT 5.1; Uhlen et al., 1992; also see Table 3 in Makrides, 1996). The appended moieties are commonly called “tags” and are often linked to the host protein by a short linker sequence containing a specific chemical (e.g., Met or Asp-Pro) or protease cleavage site (e.g., thrombin). One of the main purposes of constructing fusion proteins is to facilitate the recovery and purification of the recombinant protein. The most popular fusion partners are the polyhistidine tag (His-tag) and the glutathione-S-transferase (GST-tag), these are discussed in more detail in UNITS 6.5 & 6.6. A tag may help maintain the solubility of a protein that is normally expressed in an insoluble form (LaVallie et al., 1993; Zhang et al., 1998). Alternatively, the tags may promote insolubility, especially useful for protecting short, partially structured polypeptides, and for expressing proteins that are normally toxic to E. coli (e.g., proteins with membrane-associating or -spanning regions). The Gateway universal cloning system, previously mentioned, has been used to screen for improved solubility by comparing the effects of six different N-terminal fusion proteins and the
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Table 6.1.1
Protein Expression in Escherichia coli
Protein expressiona Native sequence
Native–fusion
Native–secretion
Locationb
Soluble
C
Advantages
Disadvantages
Yes
High-level expression Direct purification with good recovery
N-terminal Met may be retained Susceptible to proteolysis
C
No
High-level expression May protect against proteolysis Toxicity effects of protein to cell may be avoided Easy partial purification (washed pellets—see Fig 6.1.1)
Protein folding must be carried out Recovery of purified native protein can be low to zero N-terminal Met may be retained
C
Yes
High-level expression Purification aided with affinity-tagged protein Solubility and stability of expressed protein (or peptide) can be enhanced by fusion partners Authentic N terminus after site-specific cleavage (not always true)
To obtain native sequence site-specific cleavage of fusion protein required Overall yield of native protein may be low
C
No
See comments for native insoluble Purification of denatured protein aided (e.g., His-tag) Protein folding may be enhanced by binding to affinity matrix (Sinha et al., 1994; UNIT 9.4)
See comments for native insoluble Protein folding required
C
No
P/M
Yes
Ease of purification Correct N terminus Protein folded and oxidized
Expression level and recovery may be low
P
No
Correct N terminus Expression level may be high May be protected against proteolysis
Protein folding must be carried out
Secretion leader unprocessed, purification usually not attempted
aNative, native protein sequence, including any site-specific mutations or deletions; Native–fusion native sequence with N- or C-terminal extension sequence (e.g., polyhistidine tag); Native—secretion, native sequence plus N-terminal leader sequence coding for an E. coli secretory signal (e.g., OmpA). bC, cytoplasm; P, periplasmic space; M, medium.
His-tag (Hammarstrom et al., 2002). This type of study using conventional cloning and expression would represent a major undertaking.
Purification of Recombinant E. coli Proteins
The expression of fusion proteins with affinity handles, such as those containing stretches of polyhistidine (His-tagged; see also UNIT 6.5), has become extremely popular due to the ease of protein purification under both nondenaturing and denaturing solvent conditions (for more details, see discussion on Purifying Denatured Proteins). The soluble fusion proteins often have native-like conformations and are biologically active. It cannot be assumed, however, that a tag will have no effect on the protein’s function or activity. From
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cells break cells mechanical: French press enzymatic: lysozyme
extract
centrifuge 30 min at 10,000 x g
low-speed supernatant: polymers, soluble proteins
low-speed pellet: inclusion bodies, cell wall components
centrifuge 90 min at >60,000 x g
extract (Triton/EDTA/urea) centrifuge 30 min at 10,000 x g repeat two times
high-speed pellet: membrane vesicles, ribosomal particles
high-speed supernatant: cytoplasmic proteins, periplasmic proteins
purify
low-speed extract supernatant: cell wall components
low-speed washed pellet: inclusion bodies
solubilize
denatured protein
native protein
fold
purify
Figure 6.1.1 Differential centrifugation of E. coli cell lysates. Cells are broken with a French press or by lysozyme treatment. Insoluble (inclusion body) proteins, from either the cytoplasm or periplasm, are located in the low-speed pellet, which is subjected to preextraction to remove outer membrane and peptidoglycan material. Inclusion bodies are extracted from washed pellets with strong protein denaturants such as guanidine⋅HCl. The solubilized protein, which is denatured and reduced (free sulfhydryl residues), is either directly folded and oxidized (disulfide bonds formed) or purified before folding. Soluble proteins (from the periplasm and cytoplasm) are located in the low-speed and high-speed supernatants. The latter can be used directly for chromatography, whereas the former requires clarification by other techniques such as ammonium sulfate fractionation or membrane filtration.
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fermentation broth
centrifuge 30 min at 10,000 x g
supernatant: secreted protein
pellet: cells
S1
wash in isotonic medium with Mg 2+ centrifuge 30 min at 10,000 x g
pellet: washed cells
supernatant: wash
digest with lysozyme (200 µg/ ml) dilute 1:2 with water centrifuge 30 min at 10,000 x g
suspend in hypertonic medium (20% sucrose/EDTA/Tris•CI) centrifuge 30 min at 10,000 x g
supernatant: wash
pellet: plasmolyzed cells
P1
S2
supernatant: periplasmic proteins
lyse cells treat with detergent, sonicate, or subject to hypotonic shock
suspend in hypotonic medium (10 mM Tris•CI) centrifuge 90 min at >60,000 x g
supernatant: periplasmic proteins
P2
pellet: spheroplasts
cell lysate
pellet
P3
Figure 6.1.2 Localization of secreted and periplasmic proteins in E. coli. Periplasmic protein produced via a secretion vector can leak into the medium and be recovered by centrifugation (supernatant, S1) or filtration. Washing cells with an isotonic solution such as lightly buffered 0.15 M NaCl or 0.25 M sucrose can also release protein (S2). The compartmentalized periplasmic proteins are released by isotonic shock treatment by directly suspending normal cell paste or plasmolyzed cell paste into hypotonic medium. Plasmolyzed cell paste is derived by suspending cells in hypertonic medium and then pelleting. (In hypertonic medium the cell contracts, separating the inner membrane from the cell wall, and is said to be osmotically sensitized.) The hypertonic wash often releases protein (P1). The supernatant from shocked cells (P2) will contain constitutive E. coli proteins and the recombinant product. Osmotically sensitized cells can also be treated with lysozyme to fragment the outer membrane, thus releasing periplasmic proteins (P3). The pellet from the lysozyme treatment contains spheroplasts (cells with fragmented outer membranes), which are easily disrupted by detergents, sonication, or hypotonic shock to release cytoplasmic proteins. Purification of Recombinant E. coli Proteins
a protein purification viewpoint, the main advantage of affinity-tagged proteins is realized when they are combined with a secretion vector (Skerra et al., 1991); in such a case, the
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cell lysate (e.g., low-speed supernatant of Fig. 6.1.1)
clarify lysate centrifuge (90 min at >60,000 x g) filter or salt fractionate and exchange buffer
clarified lysate
conduct ion exchange (1) DEAE-Sepharose
conduct ion exchange (2) weak cationic (CM) strong cationic (S) strong anionic (Q) weak anionic (DEAE) phosphocellulose
preference
exchange buffer
order of
perform affinity methods
perform other chromatography methods (3) dye matrix hydrophobic hydroxylapatite chromatofocusing
concentrate perform gel filtration
sterile-filter
purified protein
Figure 6.1.3 Purification of soluble proteins from E. coli lysates. Abbreviations for ion-exchange resins are as follows: CM, carboxymethyl; DEAE, diethylaminoethyl; Q, quaternary ammonium; S, methyl sulfonate. The order of preference for the stages of ion-exchange (2) and other methods (3) is based on the author’s opinion and does not necessarily represent a consensus view. On the other hand, the use of a DEAE-based matrix at an early stage (1) is common practice. Affinity methods (see text and Chapter 9) can be performed at any stage following clarification of the lysate.
protein will be translocated to the periplasm or the medium, though often at low concentrations, and the tagged protein can be readily purified from the culture medium after osmotically shocking the cells. Table 6.1.1 briefly summarizes some of the major advantages and disadvantages of the various expression scenarios using E. coli. If attempts to express the protein in E. coli or
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to fold inclusion body proteins fail, then a eukaryotic system must be considered. The decision of which system to use is often dictated by the expertise available to the laboratory. Alternatively, many companies offer custom expression services using various protein expression systems—this can be an expedient, although expensive, solution. SOLUBILITY AND LOCATION OF THE PROTEIN Determining Solubility Figure 6.1.1 shows a simple centrifugation scheme that indicates how to determine the solubility of a protein expressed in E. coli (see also UNIT 5.2). The recombinant protein in the various fractions is assayed by SDS-PAGE (UNIT 10.1); if more sensitive methods are required, immunoblotting or biological assays may be used. Cell breakage carried out with a French press (UNIT 6.2) will disrupt both the outer and inner membranes. The peptidoglycan layer, which lies underneath the outer membrane in Gram-negative bacteria such as E. coli, will be fragmented into sheets. Low-speed centrifugation (30 min at 10,000 × g) separates unbroken cells, bacterial outer membrane, and peptidoglycan components, and highly aggregated inclusion body proteins (pellet fraction) from soluble bacterial proteins, soluble recombinant proteins, and polymeric materials, including ribosomal protein complexes and inner membrane vesicles (supernatant fraction). High-speed centrifugation (90 min at 100,000 × g) of the low-speed supernatant will pellet polymers. Soluble proteins, derived mainly from the cytoplasm and periplasmic space, can then be recovered from the clarified supernatant. Soluble proteins in the low-speed or high-speed supernatant are purified directly using conventional methods (UNIT 6.2). It should be noted that a working definition of solubility is the presence of protein in the supernatant after centrifugation for 100 min at 100,000 × g. This definition applies to solvents of viscosity or density close to that of water. Occasionally, recombinant protein will be found in both the pellet and supernatant fractions after low-speed centrifugation due to the accumulation of both soluble and inclusion body proteins. If partitioning is observed only following high-speed centrifugation, then specific self-association or nonspecific association involving E. coli proteins and nucleic acid may be suspected. Recombinant proteins that normally bind RNA or DNA often bind nonspecifically to bacterial nucleic acid (Sherman and Fyfe, 1990; Wingfield et al., 1990). Lindwall et al. (2000) have developed a sparse screen approach to optimizing the buffer composition for extracting and solubilizing folded (non-aggregated) proteins. Inclusion body proteins, which are located in the low-speed pellet fraction, can be partially purified by extracting with a mixture of detergent [usually 1% to 5% (v/v) Triton X-100] and denaturant, either urea or guanidine⋅HCl. The concentration of denaturant used for pellet washing is determined empirically and should be below the concentration required for solubilization of the recombinant protein; the usual ranges are ∼1 to 4 M urea and ∼0.5 to 1.5 M guanidine⋅HCl. The cloudy extract will consist of complex carbohydrate from the fragmented peptidoglycan layer, lipopolysaccharide, and outer membrane proteins. The inclusion body proteins in the washed pellets are then extracted with solvents that disrupt protein-protein interactions (e.g., 6 to 8 M urea or guanidine⋅HCl) and processed as described below. Purification of Recombinant E. coli Proteins
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Localizing Protein When proteins incorporating a secretion vector are expressed in E. coli, advantage can be taken of the fact that the recombinant proteins will be located in the periplasmic space and/or the culture medium. Secretion into the medium is due to “leakage” from the periplasm and appears to depend on the level of accumulation and the fermentation conditions. Figure 6.1.2 summarizes approaches used to recover proteins selectively from the periplasmic space or the medium (see also UNIT 5.2). High-level secretion into the periplasm sometimes results in the formation of aggregates, analogous to cytoplasmic inclusion bodies (Bowden et al., 1991). Periplasmic inclusion bodies can be extracted from the low-speed pellet fraction following normal cell breakage (see Fig. 6.1.1). Proteins in the medium can be recovered by subjecting the culture medium to centrifugation or filtration, steps that remove intact cells and large debris. The clarified protein is usually dilute and is often concentrated prior to purification by affinity or conventional chromatography. Periplasmic proteins can be selectively released by osmotic shock (preferred method) or by selective disruption of the outer membrane and peptidoglycan layer using lysozyme. Apart from its use in dissecting the bacterial compartments, lysozyme is often employed to prepare complete cell lysates, especially in laboratories that do not have access to a French press. Cells treated with lysozyme can be disrupted with detergents or by brief sonication (UNIT 6.5). Useful microscale (<1 ml) E. coli cell fractionation schemes have been based on osmotic shock treatment (Yarranton and Mountain, 1992) or repeated freezing and thawing of cells (Johnson and Hecht, 1994). UNIT 5.2 describes small-scale (1- to 25-ml) procedures for preparing samples of periplasmic extracts and extracellular media for analysis by SDSPAGE (UNIT 10.1). STRATEGIES FOR ISOLATION OF SOLUBLE PROTEINS There are no set formulas for isolating soluble recombinant or nonrecombinant proteins; there are, however, some basic strategies and precautions that can be followed. A flow chart summarizing some of the methods commonly used for E. coli is shown in Figure 6.1.3 (see also Chapter 1). In Figure 6.1.3, the step, Perform Affinity Methods, refers not only to conventional affinity purification methods (see Chapter 9), but also to affinity methods based on the use of fusion proteins. A specific protocol detailing the purification of the soluble protein interleukin-1β (IL-1β) is presented in UNIT 6.2 and two more recent examples are discussed below. Comments on the various stages are given in order of their application. Determining the Isoelectric Point The section on Analyzing the Amino Acid Composition mentions how the isoelectric point of a protein can be estimated from the pKa values for ionizable side-chain groups. The pI can also be determined experimentally by subjecting the soluble protein extract to 1-D isoelectric focusing (UNIT 10.2) or 2-D titration curve analysis (Watanabe et al., 1994; UNIT 7.3). If the recombinant protein is not a major component in the cell extract, specific detection on the 2-D gel by immunoblotting will be required. The calculated pI can be used to optimize the buffer pH in subsequent ion-exchange steps. Purification of Recombinant Proteins
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Breaking Cells Cells are efficiently broken by high-pressure homogenization using a continuous-fill French press, which is suitable for processing volumes of 40 to 250 ml (reviewed by Hopkins, 1991; see UNIT 6.2). (Yeast cells can also be conveniently broken with the French press, although two passes are required). For volumes exceeding 500 ml, the MantonGaulin-APV homogenizer (APV Gaulin) should be used. Sonication is also useful for breaking cells but is best suited for volumes <100 ml. Alternatively, the outer cell wall can be enzymatically digested with lysozyme (200 µg/ml) and the cells broken by detergents, sonication, or both (Kaback, 1971; Burgess and Jendrisak, 1975; UNIT 6.5). Proteins that are secreted into the periplasmic space can be selectively released by hypotonic (osmotic) shock (Heppel, 1967). The viscosity of the cell lysate may be high due to released nucleic acid. Before centrifugation, the viscosity must be reduced either by sonicating or by adding DNase (25 to 50 µg/ml plus 5 to 10 mM Mg2+) and RNase (50 µg/ml; no Mg2+ requirement). A standard protease inhibitor mixture should be included in the buffer—containing, for example, 2 to 5 mM EDTA, 0.5 to 1.0 mM phenylmethylsulfonyl fluoride (PMSF) or 5 mM benzamidine, and 1 µM pepstatin A. The serine protease inhibitor 4-(2-aminoethyl)benzenesulfonyl fluoride hydrochloride (AEBSF) is a water-soluble substitute for PMSF with a much longer half-life in aqueous solution and is used at ∼50 µM. (Roche Applied Science: http://www.roche-applied-science.com. Go to the Biochemistry section to download the booklet: The complete guide for protease inhibition that lists properties for most commercially available reagents). The addition of α2-macroglobulin (1 µg/mg recombinant protein) before the final purification step(s) can protect protease-sensitive proteins (Ultsch et al., 1991; see also section on MAP30 purification below). The crude extracts should be kept cold and the recombinant protein taken rapidly to a stage of the purification process where it is stable against contaminating proteases (e.g., as an ammonium sulfate precipitate). Clarifying Cell Extract by Centrifugation or Selective Precipitation The lysate is subjected first to low-speed centrifugation to remove unbroken cells and large cellular debris, then to high-speed centrifugation to remove ribosomal material and other particulates (see Fig. 6.1.1). If an ultracentrifuge is not available, the extract can be clarified by the following techniques: ammonium sulfate or polyethylene glycol fractionation (reviewed by Scopes, 1994), phase partitioning (reviewed by Walter and Johansson, 1986), and membrane filtration (van Reis and Zydney, 2001; useful guides on filtration technology are available from Millipore at http://www.millipore.com). A fairly recent technology is expanded bed adsorption where crude extracts can be directly applied to adsorbents, for example ion exchangers, without initial clarification (see Amersham Biosciences for literature at http://www.apbiotech.com). Proteins that bind tightly to nucleic acid can be selectively precipitated with polyethyleneimine and resolubilized by salt extraction (Burgess and Jendrisak, 1975). In practice, particular properties of the protein can be exploited at this stage; for example, the protein of interest may be soluble under conditions where most E. coli proteins are insoluble, such as acidic pH or high temperature. Applying Clarified Extract to a Weak Anion Exchanger
Purification of Recombinant E. coli Proteins
Fractionating the extract with an anion-exchange resin is a useful first step as it removes host E. coli proteins, many of which have pI values in the range 5.0 to 7.0 and will thus bind to a column equilibrated in 50 to 100 mM Tris⋅Cl, pH 7.5 to 8.0. The positively charged matrix will also tightly bind nonproteinaceous materials such as nucleic acids
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low-speed washed pellet (e.g., Fig. 6.1.1): inclusion body protein extract 6-8 M Gu•HCI + DTT, 8 M urea + DTT, or 10-20% acetic acid purify protein with metal chelate chromatography (His-tagged proteins) using urea or Gu•HCI as solvent
denatured protein: monomeric and reduced protein
purify protein with ion-exchange chromatography using urea or nonionic detergents as solvent
purify protein gel filtration using urea or Gu•HCI as solvent HPLC using TFA/acetonitrile as solvent
denatured protein: purified protein
fold protein
fold and oxidize proteins (1) remove denaturant by dilution or dialysis (2) include cosolvents (e.g. ,1-4 M urea or PEG or arginine or nonionic detergents) (3) include redox system (e.g., 10:1 GSH/GSSG) folded protein: purified and oxidized protein
purify protein
conduct gel filtration chromatography to remove aggregates and misfolded protein
product
Figure 6.1.4 Folding and purification of inclusion body proteins from E. coli. The protein is extracted with protein denaturants such as guanidine⋅HCl (Gu⋅HCl), urea, or an organic acid. The reductant dithiothreitol (DTT) is included to prevent artificial disulfide bond formation (especially intermolecular bonds). The denatured protein can be purified by various methods and then folded, or it can be directly folded. Typically, some purification (e.g., gel filtration in Gu⋅HCl) prior to folding is recommended as it often results in higher folding yields. Protein folding and oxidation are carried out concurrently. Disulfide bond formation is catalyzed by low-molecular-weight thiol/disulfide pairs such as reduced (GSH) and oxidized (GSSG) glutathione. GSH/GSSG ratios of 5:1 to 10:1 are normally used, which are similar to those found in vivo in the endoplasmic reticulum (Hwang et al., 1992). A cosolvent is included to maintain solubility during folding. Folded protein is purified if necessary (purification is usually needed if the protein is directly folded). Gel filtration is a useful final step for removing aggregated and or misfolded protein.
and other polyanionic species (e.g., lipopolysaccharide derived from the bacterial outer membrane). A useful cleanup of the protein will take place whether or not the protein of interest binds to the column (see UNITS 6.2 & 6.5). The following column sizes are recommended for processing extracts: for 5 g cells, 2.5-cm diameter packed to a height of 10 to 15 cm; for 50 g cells, 5.0-cm diameter packed to a height of 20 cm.
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cells
break lysozymetreated cells with French press
break untreated cells with French press
centrifuge 30 min at 10,000 x g
centrifuge 30 min at 10,000 x g
low-speed centrifugation
A
B s
s
lp ib c
lp ib c
washing steps
C
ib c
Figure 6.1.5 Preparation of washed pellets using lysozyme and the French press. Cells are broken with the French press with or without prior treatment with lysozyme. After low-speed centrifugation using a fixed-angle rotor, the contents of the centrifuge tubes have the characteristics shown. The contents of tubes A and B are labeled: s, supernatant; lp, loose pellet; ib, inclusion body protein; and c, unbroken cells and large cellular debris. The loose pellet material is derived from the outer cell wall and outer membrane (see text for further details). After washing the insoluble material (UNIT 6.3), the pellet should consist mainly of the inclusion body layer (tube C), and the supernatant should be fairly clear.
Preparing for Repeat Ion-Exchange Step
Purification of Recombinant E. coli Proteins
Before repeating ion exchange, the solvent pH and ionic strength usually need adjustment. This can be carried out by dialysis (UNIT 6.2) or by gel filtration on a desalting column using, for example, Sephadex G-25 or G-50 (UNIT 8.3). In preparation for cation-exchange chromatography, dialysis against slightly acidic buffers (pH 5.0 to 6.0) will result in the helpful precipitation of some E. coli proteins. It may also be advisable to include a relatively low concentration of urea (0.5 to 2 M) or a nonionic or zwitterionic detergent in the dialysis buffer to minimize coprecipitation with contaminants (for an extensive listing of detergents and properties, see http://psyche.uthct.edu/shaun/SBlack/detergnt.html). Basic proteins (pI >9.0), which do not bind to the DEAE column, can be
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applied after dilution to a cation exchanger equilibrated at pH 7.0 to 7.5 without careful buffer exchange (Allet et al., 1988). Repeating Ion-Exchange Chromatography For a second round of ion-exchange chromatography, one of the ion-exchange resins indicated in Figure 6.1.3 should be used. Selection kits are available for rapidly screening and selecting the most suitable ion-exchanger (Amersham Biosciences). For cation-exchange chromatography, phosphate buffer (10 to 50 mM) between pH 5.0 and 7.5 should be tried first. Cellulose Phosphate (a bifunctional cation exchanger manufactured by Whatman: http://www.whatman.com) is effective for nucleic acid–binding proteins (Kelley and Stump, 1979). Protein is usually eluted from cellulose phosphate columns using phosphate gradients. After two stages of ion exchange, many proteins will be pure enough for the final gel-filtration step (see Performing Gel Filtration). However, if the sample contains contaminants close in size to the protein of interest, then further purification is required. Some of the frequently used methods are listed in Figure 6.1.3. Hydrophobic-interaction chromatography (UNIT 8.4) is especially useful following ammonium sulfate fractionation or salt elution from an ion-exchange resin. Screening kits are also available for rapidly checking protein binding on several different agarose-dye matrices (Sigma at http://www.sigmaaldrich.com). Performing Gel Filtration The final purification step of gel filtration (using a column 1.5 to 5.0 cm in diameter and 60 to 100 cm in length) will provide good separation of the recombinant protein from higher- and lower-molecular-weight E. coli protein contaminants. Gel filtration will also separate aggregated or highly associated recombinant protein from the physically stable form of the protein (e.g., monomer or dimer). Finally, gel filtration chromatography allows for easy exchange of the buffer. The protein solution is usually concentrated before being applied to the column. After chromatography, the protein will be diluted three- to five-fold (or more) and may therefore require repeat concentration. Other Methods In addition to the generalized approach described, affinity methods can be applied at any stage following clarification of the extract. Biospecific affinity can be exploited with immobilized natural ligands such as antibodies, substrates, and receptor ligands. Affinity chromatography, which selects for particular classes of proteins, is carried out with immobilized lectins (for glycoproteins), dyes (for nucleotide-binding proteins), and nucleic acids or heparin (for RNA- and DNA-binding proteins). Commercially available antibodies against post-translationally modified residues (e.g., phosphotyrosine) are also useful. The application of affinity tags or fusions has been previously described. Affinity methods are most useful when high degrees of purification are required—e.g., for proteins secreted into the medium, for small-scale isolations, or for rapid purification requirements. The most commonly used affinity method is immunoaffinity chromatography. The ideal reagent is a monoclonal antibody that has been specifically selected to have a moderateto-low affinity for the ligand in question, thus allowing elution under nondenaturing conditions. Antibodies raised against peptides often have lower affinities for the native protein than antibodies raised against the intact protein. Elution from peptide-antibody immunoaffinity columns can be achieved using the competing immunizing peptide (reviewed by Sutcliffe et al., 1983). Directed immobilization of the antibody, where only
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the Fc domain is bound to the column matrix and the antigen binding site (Fab domain) is thus oriented away from the matrix, results in higher binding efficiencies. An oriented antibody matrix can be made by binding antibody to immobilized protein A-Sepharose (or protein G-Sepharose) and fixing it in position with a covalent cross-linking reagent (Schneider et al., 1982; commercial kits can be obtained from Pierce at http://www.piercenet.com/). Compilations of standard chromatographic fractionation media are available (Table 8.2.2; Patel, 1993). However, the best source of information is often the literature from the various manufacturers. STRATEGIES FOR ISOLATION OF INSOLUBLE PROTEINS Recombinant proteins expressed in E. coli that are located in the low-speed pellet fraction (see Fig. 6.1.1) following cell lysis are highly aggregated (i.e., inclusion bodies). Inclusion bodies are normally derived from protein aggregation in the cytoplasm, or in the periplasm if a secretion vector was used. As mentioned above, protein can also be located in either the low- or high-speed pellet fractions because of interaction with bacterial nucleic acids. Furthermore, if the protein is known to undergo polymerization in vitro (e.g., viral nucleocapsid subunits), expression in E. coli can also be expected to lead to polymerization in vivo to varying degrees, and such proteins will be partitioned in both the supernatant and pellet fractions (Wingfield et al., 1995). There are also examples of membrane proteins that, when expressed in E. coli, associate with the inner cytoplasmic membrane and can be extracted with nondenaturing detergents (Bibi and Beja, 1994, and references cited therein). When apparent insolubility is due to interactions involving folded protein as described above, extraction under nondenaturing conditions should be attempted, for example, using various pH buffers containing salt (e.g., 0.25 to 1.0 M NaCl) and nondenaturing detergents (e.g., 10 mM CHAPS or 2% Triton X-100). Insolubility due to classic inclusion body formation requires extraction with denaturing solvents, and the remainder of this section deals with this subject. The flow chart in Figure 6.1.4 illustrates some of the approaches possible for processing protein extracted from inclusion bodies. Breaking Cells Cells can be broken by mechanical means (UNIT 6.2), enzymatically with lysozyme (UNIT 6.5), or by a combination of methods (UNIT 6.5). It is advantageous to break cells as completely as possible, as any unbroken cells will be located in the low-speed pellet fraction from which the recombinant, insoluble protein will be extracted. Preparing Washed Pellets The object of the initial low-speed centrifugation and pellet “washing” is to extract as many E. coli contaminants as possible without solubilizing the recombinant protein. This is usually carried out as described in the section on Determining Solubility (see also Fig. 6.1.1).
Purification of Recombinant E. coli Proteins
When a fixed-angle rotor is used, pellets from the low-speed centrifugation consist of at least two light-colored layers and a darker, hard-packed pellet at the bottom of the tube (Fig. 6.1.5). The hard-packed material is probably a small amount of unbroken cells. The next layer is inclusion body protein, and the top layer (least dense and lightest in color) is outer membrane and peptidoglycan fragments. Analysis of the top layer by SDS-PAGE (after heating proteins in SDS sample buffer at >80°C) will reveal two strong bands at ∼35 and 38 kDa representing OmpA and the matrix proteins OmpC and OmpF, respec-
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tively, from the outer membrane (DiRienzo et al., 1978; see also Fig. 6.3.1). The outer membrane/peptidoglycan layer can be partially removed by resuspending and centrifuging at reduced speed (or time) or by diluting the suspension. Alternatively, the cells can be pretreated with lysozyme prior to the French-press cell breakage as described in UNIT 6.5. The lysozyme treatment reduces the size of the loosely pelleted outer membrane/peptidoglycan material so it locates predominately in the low-speed supernatant (Fig. 6.1.5). The recombinant protein in a well-prepared washed pellet will typically be >60% pure when analyzed by SDS-PAGE (UNIT 6.3). Extracting Protein The washed pellets are extracted with high concentrations of protein denaturants such as 6 to 8 M guanidine⋅HCl or urea. It should be noted that some proteins are resistant to denaturation with high concentrations of these reagents, especially urea. Some washed pellets extracted with 8 M guanidine⋅HCl can be viscous and unsuitable for direct chromatography. In these cases, pre-extraction of the washed pellets with a limiting concentration (0.5 to 2.0 M) of guanidine⋅HCl can often overcome this problem. Solubilization with the anionic detergent N-lauroylsarcosine (Nguyen et al., 1993; Burgess and Knuth, 1996) and with 10% to 20% acetic acid has also been useful (UNIT 6.5); other denaturants for extracting inclusion bodies are described elsewhere (UNIT 6.3; Marston and Hartley, 1990). For background information on the mode of action of protein denaturants, readers should consult the reviews of Tanford (1968) and Creighton (1993). If the protein contains cysteine residues, it is essential to include a reducing agent, preferably 5 to 10 mM dithiothreitol (DTT). Even in the presence of strong protein denaturants, it may be necessary to sonicate or heat samples briefly to completely disperse and solubilize the protein. The extraction process should completely disaggregate and denature the protein into unfolded monomers. Urea is not recommended for the initial extraction. For example, even if it is known that a native version of protein can be unfolded with 4 M urea, the same protein in an E. coli inclusion body will almost certainly not be completely extracted as unfolded monomers with that same concentration of urea (or in most cases, even with 8 M urea). Initial extraction trials should be carried out with guanidine⋅HCl, which is more effective than urea. Most proteins will be extracted with 6 to 8 M guanidine⋅HCl. There should be adequate reductant present to maintain sulfhydryl groups in the reduced state, and thus prevent artificial disulfide bond formation. The presence of EDTA and a slightly acidic pH of 6.0 to 6.5 will help minimize cysteine oxidation. The extract may require clarification by filtration or centrifugation. Choosing Purification or Folding The extracted protein can be further purified, or it can be directly folded and then purified. Protein folding appears to be unaffected by the protein background in bacterial extracts (London et al., 1974), however, removal of nonproteinaceous material prior to folding has been reported to be beneficial (Darby and Creighton, 1990). Based on recent work, it is worth noting that high concentrations of background bacterial protein may promote aggregation of the unfolded recombinant protein by macromolecular crowding effects (Ellis, 2001). If purification of protein in the denatured state is possible, use the purified material to develop a folding protocol. Then use this protocol with clarified protein extracts, or better still with protein partially purified by DEAE-Sepharose, to observe if the presence of contaminants has any effect on the yield of folded protein. Finally, there may be specific reasons for purifying proteins in the denatured state. For example, some proteolytic enzymes, such as HIV-1 protease, self-digest (undergo auto-
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proteolysis) in the uninhibited state (Mildner et al., 1994, and references cited therein) but can be purified intact in the denatured (inactive) state, then refolded when required. Other proteins once folded may have low solubilities and be especially susceptible to aggregation, resulting in poor behavior on column matrices (see VP26 purification below). However, in general, unfolded proteins are more susceptible to chemical and proteolytic modifications. Purifying Denatured Proteins If the protein is extracted with guanidine⋅HCl, gel filtration is a useful first purification method; often protein >80% pure can be obtained (UNIT 6.3; Wingfield et al., 1997). The proteins exist as random coils in the denaturant and their elution from the column should be in order of their molecular weight and not be influenced by shape. If the protein is located in several peaks there may have been incomplete solubilization during the extraction. In this case, 8 M guanidine⋅HCl should be used for the extraction and the protein dispersed by sonication or by heating if necessary. Another possibility is intermolecular disulfide bond formation, in which case the DTT concentration in the sample and column buffers should be increased. It is worth noting that the column can often be equilibrated and eluted with lower guanidine⋅HCl concentrations (e.g., 4 M) than those used for the actual extraction process. Only monomeric protein should be selected for further processing. The protein at this stage can be stored frozen, ideally at −80°C. The partially purified protein in guanidine⋅HCl can be directly folded (see Performing Protein Folding), or the denaturant can be exchanged by dialysis or gel filtration for 1% to 5% (v/v) acetic or formic acids (acetonitrile at 5% to 10% v/v can also be included) and then lyophilized. Alternatively, the protein can be acidified with trifluoroacetic acid (TFA; ≤0.1% v/v) and further purified by reversed-phase chromatography (Wingfield, 1997; Wingfield et al., 1999). Useful high-flow matrices (Source 15RPC from Amersham Biosciences) can be purchased as bulk media. These matrices may not have the resolution of traditional prepackaged silica-based reversed-phase columns, but they have high capacity, can be eluted at higher flow rates, and are stable over a wide range of pH. Proteins eluted with acetonitrile/TFA are also suitable for lyophilization. Proteins tagged with histidine residues can be purified in guanidine⋅HCl-, urea-, or even SDS-containing buffers, using metal chelate chromatography (UNIT 6.5). There are many reports of “on-column protein folding” by binding the unfolded protein in guanidine⋅HCl or urea and then accomplishing folding using a reverse urea gradient (e.g., Gulnik et al., 2001). Proteins in urea and non-ionic or zwitterionic detergents (e.g., CHAPS) can be purified by ion-exchange chromatography (e.g., Wingfield et al., 1990). For ion-exchange chromatography, better results have been reported using protein that has been first extracted with guanidine⋅HCl, and then exchanged into urea (Shire et al., 1984). If urea is used either for extraction or for maintaining solubility during refolding, a cyanate scavenger such as a glycine- or Tris-based buffer should be included to prevent carbamylation of the protein (Stark et al., 1960). For critical work, urea can be deionized with a mixed bed ion-exchange resin (see discussion of Protein Folding Reagents in APPENDIX 3A). Performing Protein Folding
Purification of Recombinant E. coli Proteins
Protocols for folding proteins basically involve controlled removal of the denaturant under conditions that minimize aggregation and allow correct formation of disulfide bonds. For overviews of the practical aspects of protein folding, see UNIT 6.4; Wetzel (1992); Thatcher
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et al. (1996); Rudolph et al. (1997); Lilie et al. (1998); De Bernardez Clark et al. (1999); and De Bernardez Clark (2001). To minimize nonproductive aggregation, folding is normally carried out at low protein concentrations (e.g., 0.01 to 0.10 mg/ml); for small, single-domain proteins, higher concentrations (e.g., 0.1 to 1.0 mg/ml) can often be tolerated. Dilution and dialysis are the most common methods for removing the denaturant. Solubility during folding can be maintained with co-solvents such as nondenaturing concentrations of urea (1 to 4 M; London et al., 1974; UNIT 6.5) or guanidine⋅HCl (0.1 to 1.5 M; Orsini and Goldberg, 1978), arginine (0.4 to 0.8 M; De Bernardez Clark et al., 1999), nonionic detergents and lipids (Zardeneta and Horowitz, 1994), cationic detergents (Puri et al., 1992), and polyethylene glycol (PEG; Cleland et al., 1992). These various additives function by minimizing intermolecular associations between “sticky” hydrophobic surfaces present in folding intermediates. For further discussion of aggregation versus folding, see Goldberg et al. (1991) and Kiefhaber et al. (1991). Additives such as ammonium sulfate, glycerol, sucrose, enzyme substrates or inhibitors, and ligands have also been used to improve protein folding (see Table 1 in De Bernardez Clark et al., 1999, for a useful list of additives used in folding). Protein expressed in the cytoplasm of E. coli is in the reduced state; this is true for both soluble and insoluble proteins. Once insoluble protein is solubilized, it needs to be maintained in a reduced state by the presence of reductant until protein folding is initiated. The oxidative formation of disulfide bonds (one of the rate-limiting steps in protein folding) can be catalyzed by low-molecular-weight thiol and disulfide pairs such as reduced and oxidized glutathione (GSH/GSSG). Redox buffers facilitate oxidation through thiol/disulfide exchange reactions (reviewed by Wetlaufer, 1984; Creighton, 1984; Gilbert, 1995). Normally GSH/GSSG ratios of 5 to 10 are used with a total glutathione concentration of 1 to 5 mM (Wetlaufer, 1984). To reduce the rate of GSH loss due to air oxidation, 1 mM EDTA should be included in the buffer (Wetlaufer et al., 1987). The optimal concentrations and ratios of reagents must be established in an empirical manner. Folding and oxidation are normally carried out concurrently (for further details, see Rudolph et al., 1997). Analogous to the approach commonly used to optimize conditions for protein crystallization, various screens have been developed to establish initial conditions for protein renaturation and oxidation (Hofmann et al., 1995; Armstrong et al., 1999) and kits are commercially available (FoldIt Screen from Hampton Research at http://www.hamptonresearch.com). For examples of preparative protein folding, see UNIT 6.5. In addition, some recent examples from the author’s laboratory are given below. The refolding of Fab fragments expressed in E. coli (Buchner and Rudolph, 1992) is illustrative of the systematic and empirical approach used to optimize folding conditions. Other examples of interest are described by Kohno et al. (1990) and Grunfeld et al. (1992). Protein-assisted folding and oxidation Protein folding in vivo is assisted in both eukaryotes and prokaryotes by two classes of accessory proteins: folding catalysts (for a review, see Schiene and Fischer, 2000) and molecular chaperones (Eisenberg, 1999; Feldman and Frydman, 2000). Folding catalysts accelerate rate-limiting steps in protein folding such as disulfide bond formation (protein disulfide isomerases) and the rotation of X-Pro bonds (peptidyl prolyl cis-trans isomerase) during protein folding. Chaperones bind denatured or unfolded proteins thus preventing misfolding and aggregation. The cytoplasm of E. coli is maintained in the reduced state by thioredoxin and the glutathione/glutaredoxin pathways. In hosts where the reduction of thioredoxin and glutathione is impaired by mutations to the thioredoxin
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reductases and glutathione reductase genes, the resultant oxidizing conditions allow the formation of disulfide bonds in expressed proteins located in bacterial cytoplasm (Bessesste et al., 1999; cells and expression kits are commercially available from Novagen). The periplasm of E. coli also contains protein disulfide isomerases, the Dsb enzymes, which have thioredoxin-like folds and act as strong thiol:disulfide oxidants (Missiakkas and Raina, 1997; Braun et al., 1999). Secretion of proteins into the periplasmic space has been the traditional approach for producing oxidized proteins in vivo, but with the aforementioned advances in cytoplasmic oxidations, this approach is probably best suited for proteins that are toxic to the cell when expressed in the cytoplasm (Cornelis, 2000). As mentioned, molecular chaperones prevent aggregation by interacting transiently with hydrophobic patches on unfolded proteins and suppressing aggregation and promoting folding (UNIT 6.4; reviewed by Jaenicke, 1993; Ellis and Hart, 1999; Feldman and Frydman, 2000). There are now many examples of chaperone-assisted protein expression in which the endogenous levels of the bacterial chaperones GroES and GroEL (∼1%) are increased up to ten-fold by co-expression with a target protein (Cole, 1996; Goenka and Rao, 2001). Often, increases in soluble protein expression are observed, but this is not always the case. Chaperones have also been used in vitro as protein folding reagents and some examples of folding in the presence of protein disulfide isomerase, peptidyl prolyl cis-trans isomerase and GroES/GroEL are given in Rudolph et al. (1997). Protocols for the high-level expression and rapid purification of E. coli GroEL and GroES are described by Kamireddi et al. (1997). Purifying Folded Protein Once the protein has been folded, any of the purification methods discussed in Chapters 8 and 9 can be used. The number of purification steps required should be fewer than those for a protein expressed in a soluble state because of the purification factor obtained by preparation of washed inclusion bodies (UNIT 6.3). One of the purification methods that should be included is gel filtration, which may be the only one required. A correctly selected matrix should remove any remaining E. coli proteins and separate aggregated and misfolded protein from the native folded protein. Misfolded protein may be expected to have a larger molecular radius (higher apparent mass) than the corresponding native protein. Monitoring Protein Folding The restoration of function (e.g., enzymatic or biological activity) is perhaps the best criterion for detecting successful folding. However, it is not always practical to use activity measurements to monitor folding. It is also worth mentioning that an unfolded protein may become activated following the dilution required for many activity measurements. Conversely, native proteins can be denatured or inactivated during prolonged incubation at 37°C or by adsorption to microtiter plates. The use of antibodies to monitor protein folding is briefly reviewed by Goldberg (1991), and reviews of common spectroscopic methods, such as circular dichroism and fluorescence, are provided in Chapter 7 and by Schmid (1997). BACTERIAL EXPRESSION OF PROTEINS NORMALLY GLYCOSYLATED
Purification of Recombinant E. coli Proteins
Because E. coli lacks glycosylation machinery, expression of glycoproteins in E. coli systems results in the synthesis of nonglycosylated variants. Glycoproteins expressed in E. coli are often, but not always, insoluble. In vitro folding studies with glycosylated and nonglycosylated forms of proteins indicate that the carbohydrate can stabilize folding
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intermediates, and thus enhance folding, while not necessarily affecting the stability of the native state (Kern et al., 1993, and references cited therein). In eukaryotic cells, interference with protein glycosylation can lead to the formation of misfolded, aggregated, and degraded protein. This indicates that in vivo glycosylation (N-linked) may also prevent the aggregation of folding intermediates (reviewed by Helenius, 1994). Detailed NMR studies on glycoproteins have clearly shown that carbohydrates stabilize folded proteins and even prevent marginally stable proteins from unfolding (for a review, see Wyss and Wagner, 1996). Despite potential pitfalls, many nonglycosylated protein variants have been successfully folded from E. coli inclusion bodies. Examples include cytokines of biomedical importance such as granulocyte/macrophage colony-stimulating factor (GM-CSF; Diederichs et al., 1991) and interleukin 5 (IL-5; Milburn et al., 1993). Inclusion body formation was avoided in some studies by using secretion vectors; examples include GM-CSF (Walter et al., 1992) and the extracellular domain of the human growth hormone receptor (deVos et al., 1992). The aforementioned proteins have been crystallized and their structures determined by X-ray crystallography, supporting the view that the structural integrity and conformation of the proteins were not affected by the lack of glycosylation and their respective preparative histories. If a glycoprotein of interest is available from a eukaryotic recombinant expression system or if the natural protein is available, then before investing time with E. coli expression, it may be worthwhile to determine whether the protein can be denatured and refolded in vitro. Pilot experiments can be carried out on intact protein and on protein enzymatically deglycosylated with glycosidases and, if disulfides are present, with and without reduction. Of course, if the protein can be secreted to the periplasm, aggregation and the necessity for in vitro folding may be avoided. The production of deglycosylated proteins in E. coli expression systems for in vitro biochemical and structural studies is obviously of great value; however, the proteins may not always be suitable for in vivo studies due to low biological activity. Compared to authentic proteins, nonglycosylated variants can have a reduced circulatory lifetime and can exhibit increased immunogenicity and protease sensitivity (Rasmussen, 1992). SOME EXAMPLES OF PROTEIN EXPRESSION AND PURIFICATION Examples of protein expression and purification can be found in most biochemical journals, two which may be especially useful: Protein Expression and Purification (http://www.academicpress.com/pep), which covers advances in the expression and purification of recombinant proteins mainly from E. coli although other expression systems are often included; and Current Opinion in Biotechnology, which regularly provides updates on various aspects of recombinant protein production as well as useful reference lists. Detailed protocols are also given in the units of this Chapter and a few recent examples of protein expression and purification are discussed below to illustrate some of the general approaches used to deal with soluble and insoluble E. coli protein expression. Soluble Proteins HIV Nef Nef is a 205-residue myristolylated protein expressed at high levels in the early stages of HIV infection. The protein is important for the induction of AIDS and is being actively researched as a potential drug target. Unlike most HIV-1 and related proteins expressed in bacteria, Nef is recovered from the soluble fraction of E. coli extracts. The purification
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protocol adopted following cell breakage and low-speed centrifugation is fairly straightforward comprising two stages of ion-exchange chromatography using DEAE-Sepharose (weak exchanger) followed by Q Sepharose (strong exchanger) and finally gel filtration using Superdex 75. Characterization of the purified protein yielded the following information. 1. Nef has a maximum solubility of ∼0.5 to 0.6 mM (∼10 mg/ml) in low-ionic strength buffers at pH 7.5 to 8.0, (e.g., 5 mM Tris⋅Cl). The solubility can be increased by the inclusion of nondenaturing concentrations (2 M) of urea, as established by titration studies monitored by far-UV circular dichroism. Acetonitrile (5% to 10%) also increases the solubility of protein. 2. The protein contains three cysteines (positions 54, 141, and 205), none of which are involved in native disulfide bond formation. The cysteines at positions 54 and 205 are solvent-exposed. 3. Digestion of the purified protein with proteases indicated rapid digestion of the N-terminal region (residues 1-38). For example, digestion was complete with a few minutes using relatively low concentration of trypsin (1% w/w). The above information was exploited to increase the robustness of the purification protocol. Low solubility was a major issue during purification and this was improved by including 4 M urea in the extraction buffer and 2 M urea in the two anion exchange column buffers. For the final gel filtration step, 10% acetonitrile was included to help maintain both the solubility of Nef and fortuitously cause aggregation of some E. coli contaminants that eluted in the void volume. Neither the urea nor the acetonitrile at the concentrations used resulted in Nef denaturation. The problem of cysteine oxidations was circumvented by mutating cysteines 54 and 205 to alanines. Mutation of cysteine 205 alone and including 5 mM DTT in all the column buffers was also a satisfactory solution. The high susceptibly of the N-terminal region to proteolytic processing indicates that it is solventaccessible and likely to be unstructured. In the case of Nef, this region can be deleted without affecting the folding of the protein and removes the potential for heterogeneity due to partial processing by E. coli proteases. The NMR structure of HIV Nef was determined with protein prepared as described above (Grzesiek et al., 1997). MAP30 MAP30 is a plant protein obtained from bitter melon that has anti-HIV and anti-tumor activities. The 30-kDa protein is well expressed in E. coli as a soluble protein and is purified by two stages of exchange chromatography followed by gel filtration. The clarified extract is first applied to a DEAE-Sepharose column at pH 8.0; the majority of MAP30 does not bind or weakly binds the exchange resin. The column flow-through and early eluting fractions are dialyzed against pH 6.5 buffer then fractionated using SPSepharose (strong cation exchanger). The final step is gel filtration using a Superdex 200 column at pH 8.0.
Purification of Recombinant E. coli Proteins
There are clear similarities between the MAP30 purification scheme and the one developed for the Nef protein; both utilize an initial clean-up step using DEAE-Sepharose followed by a second more discriminating ion-exchange step and finally a “polishing” step using gel filtration. For Nef, the second ion-exchange step employs an anion-exchange resin while the MAP30 method uses a cation-exchange resin. The choice of resin for the second step reflects the difference in the isoelectric points of these proteins. Nef has a calculated pI of ∼5.95 and is positively charged at pH values greater than this. MAP30 has a slightly basic pI of 9.00 and is negatively charged at pH values below this. Thus, Nef binds to DEAE-Sepharose and Q-Sepharose at pH 7.4 and 8.0, respectively.
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On the other hand, MAP30 does not bind to DEAE-Sepharose at pH 7.4 but binds strongly to a cation exchanger at pH 6.5. Apart from purification, there is also another similarity between Nef and MAP30, namely susceptibility to proteolytic processing during purification. As previously mentioned, the N-terminal region (residues 1-38) of Nef is at risk for proteolysis, and to maintain the structural integrity, especially during cell breakage and the initial processing, protease inhibitor cocktails must be included in the buffers. MAP30 also has a region susceptible to processing, namely, the ∼20 residues at the C-terminal end of the protein. Again, this is due to the fact that this region is largely unstructured in an otherwise folded and stable molecule (Wang et al., 1999). When purifying MAP30, standard protease inhibitors are included during the early stages of purification and, in addition, α-macroglobulin (15 to 2.0 µg/mg protein) is added to the protein prior to the gel-filtration step. The macroglobulin inhibits a wide range of proteases by a trapping mechanism (Sottrup-Jensen, 1989). If proteins are to be used for structural studies, deletion mutants can eliminate unstructured regions at the N- and C- terminal regions. Deletions of such regions from either Nef or MAP30 do not significantly change the pI of either protein, so the same purification procedures can be applied to the deletion mutants. Although incremental structural determination is an important strategy in structural biology, one should always be aware that regions deleted, even those that appear unstructured, may have important functional roles. There are many examples of disordered proteins and protein domains that adopt folded structures upon binding to their biological targets (for a review, see Dyson and Wright, 2001), and in the case of Nef, it appears that the apparently unstructured N-terminal region (residues 1-57) mediates binding to the tumor suppressor protein p53, possibly enhancing HIV-1 replication (Greenway et al., 2002). A dual vector co-expression system for producing heteromeric complexes in E. coli (Johnson et al., 2000) may be particularly useful for producing proteins requiring binding partners for folding and stability. Insoluble proteins HIV-1 gp41 ectodomain The membrane-associated glycoproteins of HIV-1 include gp120 and gp41, the latter mediating membrane fusion with the host cell. These viral envelope proteins have been the subject of intense structural analysis over the last several years as inhibition of membrane fusion, hence viral entry, is a potential drug target in the development of therapeutics for AIDS. A basic strategy in tackling membrane-associated proteins is to remove the membrane-spanning region by expressing the non-membrane-associated region or ectodomain. The gp41 ectodomain is a 150-residue protein that is recombinantly expressed in E. coli as an insoluble protein. The protein can be extracted from inclusion bodies with 8 M guanidine⋅HCl and purified by one step of gel filtration in the presence of 4 M guanidine⋅HCl. The guanidine is removed by preparative reversed-phase HPLC and the protein folded upon dialysis against 50 mM sodium formate at pH 3.0. The yield of folded protein is >90%. Characterization of the protein indicates that its solubility decreases dramatically below pH 4.0. Between pH 3.0 and 4.0, the protein has an all α-helical secondary structure with a trimeric subunit structure. The protein was demonstrated to have folded by determining its full structure at pH 3.5 using multidimensional NMR (Caffrey et al., 1998). The protein was also crystallized from a buffer at pH 3.5 and its structure determined by X-ray crystallography (Yang et al, 1999). Purification of Recombinant Proteins
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Other insoluble proteins expressed in E. coli that exhibit acid stability similar to the gp41 ectodomain can be processed and folded using a similar scheme as described above. For example, the HIV protease can be purified and folded with this method. The HIV protease, after folding at pH 3.5, exhibits fair solubility up to pH 5.0, with solubility decreasing at higher pH values. Other proteins may only be partially folded or unfolded at acidic pH values; in these cases, the reversed-phase HPLC step could be used to simply remove the denaturant, then the protein can be freeze dried from TFA-acetonitrile solvent and used for folding trials. The gp41 ectodomain contains two cysteine residues in a loop region connecting N- and C-terminal helical domains. These cysteines do not form intramolecular disulfides and can be substituted by alanine residues. This is a common theme. If a protein contains free solvent-accessible cysteines that play no structural or functional role, it is often a good idea to substitute them (usually with Ala), especially if structural studies are planned. Human Tissue Inhibitor of Metalloprotease-2 (TIMP-2) and hepatocyte growth factor isoforms (NK1 and NK2) The TIMP families of proteins are inhibitors of the matrix metalloproteases and are critical effectors of extracellular matrix turnover. The hepatocyte growth factor (HGF) is a multifunctional protein stimulating a wide range of cellular targets. The HGF gene codes for three distinct proteins: the full-length form and two truncated isoforms that include an N-terminal domain (N) and one-kringle (NK1) or two-kringle domains (NK2). TIMP-2 (21 kDa), NK1 (21 kDa), and NK2 (30 kDa) contain multiple disulfides that stabilize the folded conformations. For example, TIMP-2, apart from having 12 cysteines that form 6 disulfides, contains a cysteine as the N-terminal residue. All three proteins were expressed in E. coli as insoluble proteins, extracted with guanidine⋅HCl and reductant, and the unfolded protein separated by gel filtration in a similar manner to that previously discussed. The partially purified proteins can be conveniently stored frozen in guanidine⋅HCl at −80°C for several years without deleterious effects on folding or recovery of active protein. The folding and oxidation of the proteins are detailed in the respective publications, Stahl et al. (1997) and Wingfield et al. (1999), but briefly, the protocols involve equilibrium dialysis incorporating urea as a co-solvent to maintain solubility during folding, and a glutathione-based oxido-shuffling system (redox buffer) to promote formation of disulfide bonds (this approach is also detailed in Basic Protocol 1 in UNIT 6.5). The final stage of the purification process is gel filtration of the folded proteins, which, apart from removing host contaminates, separates folded monomers from any misfolded and aggregated protein.
Purification of Recombinant E. coli Proteins
When recombinant expressed proteins are insoluble in E. coli, the purification scheme can be very simple as illustrated above where one or two steps of gel filtration may be all that is required; the challenge is determining a method to fold and oxidize the protein. In all three examples discussed above, the key to efficient folding is maintaining solubility, whether it be by taking advantage of the acid stability of the protein and working at pH 3.5, or by including the co-solvent urea. As mentioned above, TIMP-2 has an N-terminal cysteine residue. When this protein was originally expressed, an alanine was appended to the N-terminus since it had been observed that partial N-terminal processing occurred when cysteine was the terminal residue. The alanine residue was added in an effort to produce homogeneous protein for structural studies. The purified Ala+ TIMP-2 appeared monomeric and folded, yet was devoid of its normal inhibitory activity (Wingfield et al., 1999). It was determined that the coordination of a zinc atom by the N-terminal cysteine stabilized substrate binding and required a free amino terminal group. This was demonstrated by exopeptidase digestion (using aminopeptidase 1) of Ala+ TIMP-2, which
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removed the N-terminal alanine making cysteine the N-terminal residue and, thus, restoring biological activity. A GST fusion protein The protein VP26 is a 12-kDa capsid protein of the herpes simplex virus and initial attempts to directly express this protein in E. coli failed. It was possible, however, to produce this protein at fairly high levels in E. coli as a GST fusion (Wingfield et al., 1997a). The insoluble protein was treated in the usual manner: solubilized with guanidine⋅HCl and partially purified by gel filtration also in guanidine⋅HCl. The usual purification for GST fusion proteins is affinity chromatography using immobilized glutathione, which requires that the GST moiety be folded (UNIT 6.6). Due to the low solubility of VP26 and its high propensity for aggregation, the following approach was used. First, the VP26-GST fusion was folded from the guanidine⋅HCl solution by equilibrium dialysis against buffer containing 2.5 M urea, 10 mM CHAPS, and 0.25 M NaCl, and then against the same buffer lacking the urea. The buffer additives were included to maintain protein solubility (solubility is improved with >0.25 M NaCl, but the cleavage of the GST moiety by thrombin is inhibited by high salt concentrations). Following cleavage of GST and VP26, the proteins were denatured again with guanidine⋅HCl, separated by gel filtration and the purified VP26 refolded from urea and CHAPS as described above. As an aside, the GST moiety is readily refolded from guanidine⋅HCl and does not require high salt or CHAPS to maintain solubility during the dialysis steps. The purification approach used here may appear inelegant, but the fusion system was used not to facilitate purification, but to facilitate expression of the protein. PROTEIN HANDLING Storing Purified Proteins Purified protein should be filter-sterilized prior to storage. Millex-GV 0.22-µm filters (Millipore) employ hydrophilic membranes with low binding capacities and are recommended for most proteins. Proteins are best stored at −80°C or may be stored on ice; freezing at −20°C is not recommended. Rapid freezing in small aliquots using dry ice/ethanol mixtures is preferred to slow freezing at −20°C . The addition of sucrose or glycerol often increases protein stability during storage and during freezing and thawing cycles (Arakawa and Timasheff, 1985; Timasheff and Arakawa, 1997). Lyophilization is best for long-term storage; however, care should be taken in choosing the protein solvent (Franks, 1993). Promoting Protein Solubility and Stability If the recombinant protein contains reactive unpaired sulfhydryl groups in the native conformation, 1 to 5 mM DTT should be included in the column buffers during purification. However, reductant should not be used gratuitously, as the native protein may contain intra- or intermolecular disulfide bonds, disruption of which can reduce the stability and solubility of the protein. Reductants should be included, for example, during gel filtration if dimers or higher aggregates need to be converted to active monomeric protein. The presence of intermolecular (and occasionally intramolecular) disulfide bonds can be determined analytically by SDS-PAGE under nonreducing conditions (UNIT 6.5) by pretreating proteins sequentially with iodoacetamide (to prevent artificial disulfide exchange) and then with SDS in the absence of reductant. The use of reductants can best be rationalized once the native protein has been characterized. Purification of Recombinant Proteins
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EDTA (1 to 5 mM) is often included in buffers to remove heavy metals that can catalyze oxidative processes and inhibit certain proteases. It should be noted that EDTA will bind to anion exchange resins (Scopes, 1994). Other components often added to buffers to promote protein solubility during purification include nonionic or zwitterionic detergents, low concentrations of urea (1 to 2 M), and salt (0.5 to 1 M NaCl). These additives are compatible with ion-exchange chromatography, except for high-salt concentrations, which are compatible with hydrophobic-interaction chromatography (UNIT 8.4), affinity chromatography (Chapter 9), and gel-filtration chromatography (UNIT 8.3). Solvent pH is one of the most important variables for maintaining protein solubility; in general, proteins are least soluble at or near their isoelectric points. Preventing Contamination Precautions to prevent contamination of the protein of interest are as follows: 1. To avoid cross-contamination, especially from other recombinant proteins, dedicate one set of chromatography resins for the purification of each protein. If this is not possible, or if expensive prepackaged matrices are used, be sure to clean resins thoroughly after each use. Check the manufacturer’s recommendations and be aware of the chemical stability of the resin, especially for extremes of pH. 2. Store resins with preservatives (e.g., 1 mM sodium azide) and avoid storage in phosphate buffers, which provide a good medium for bacterial growth. 3. To generate reproducible protocols using ion-exchange methods, monitor the pH and conductivity of all buffers and column effluents (the latter ideally in-line). 4. Avoid protein cross-contamination in concentration equipment such as stirred ultrafiltration cells with ultrafiltration membranes. 5. Keep pH and conductivity probes scrupulously clean, especially when used with solutions containing proteases. Likewise, use care when using cuvettes for UV measurements. 6. Avoid vigorous stirring of protein solutions to prevent shear denaturation, and handle soft agarose-based column matrices carefully to prevent bead fragmentation. Removing Pyrogens Recombinant proteins used for in vivo studies should be free of endotoxins (pyrogenic lipopolysaccharide derived from the bacterial outer membrane of Gram-negative bacteria). Yeast and mammalian cell hosts do not contain endotoxins; however, exogenous contamination from water and others must be avoided. Pyrogens can be detected using the sensitive Limulus amoebocyte lysate (LAL) assay kits available from Sigma and other suppliers. As endotoxins are negatively charged, they will be removed by anion-exchange chromatography. Other methods are reviewed in detail by Petch and Anspach (2000). SCALE OF OPERATIONS AND AIMS OF PURIFICATION Determining Scale Purification of Recombinant E. coli Proteins
The amount of protein required and the level of purity will vary dramatically from laboratory to laboratory and study to study. The following guidelines will help in planning a strategy.
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If a Coomassie blue–stained band corresponding to the expressed protein is observed on one-dimensional SDS-PAGE analysis of a whole-cell extract, then the protein constitutes at least 0.5% to 1% of the total protein. Wet E. coli cell paste contains ∼10% to 15% protein by weight (reviewed by Neidhardt, 1987). If the level of expression is low to average (e.g., 5%), then 1 g wet weight of cells will contain ∼5 mg recombinant protein. Hence, a cell paste of 20 to 50 g (a typical yield from a 1- to 2-liter benchtop fermentation) will contain 100 to 250 mg recombinant protein, and often two- to five-fold more. Shaker-flask fermentations of equivalent volumes might yield 5% or 10% of these amounts. Thus, for soluble proteins, or insoluble ones that can be refolded (with ≥5% yield), significant amounts of protein can be obtained from relatively small fermentations. For proteins secreted into the periplasm or medium, fermentations on larger scales may be required, as expression levels are usually considerably lower than that for direct expression. Deciding the Aims of the Purification There are many reasons, both scientific and commercial, for producing purified recombinant proteins. The development of laboratory-scale purification schemes that produce pure protein (a single band on SDS-PAGE) should be relatively straightforward given the relatively high abundance of recombinant proteins in cell extracts. Protein present at 1% of the total cell extract requires only a 100-fold purification compared to the several thousand-fold sometimes required for the purification of nonrecombinant proteins (reviewed by Stein, 1991). The widely used method of affinity tagging proteins allows the nonspecialist to rapidly purify protein for biochemical and activity studies without investment in some of the specialized equipment mentioned below. However, far more time and expertise is required to develop protocols that produce purified recombinant proteins having the physical and chemical homogeneity required for clinical use and for structural determinations. Furthermore, only after detailed characterization of the isolated protein will chemical and physical heterogeneities be revealed in enough detail for steps to be taken to either prevent their occurrence or rationalize fractionation of modified species. Therapeutic proteins Mammalian cells are the production host for many current protein therapeutics, however, E. coli, is also used to produce major biotechnological products including insulin and bovine growth hormone. Some advances in E. coli production of therapeutic proteins and methods used to fold solubilized protein for industrial processes have been recently reviewed (De Bernardez Clark, 2001; Swartz, 2001). Proteins used for clinical studies must be manufactured according to applicable FDA guidelines that include Good Manufacturing Procedures (GMP). Sofer and Hagel (1997) provide practical coverage of modern process development, including process chromatography and its scale-up. The physiochemical characterization of protein pharmaceuticals can be especially challenging and many of the methods and approaches used rely on mass spectrometry (see Chapter 16). Structure Determination For many investigators, a primary goal is to correlate the structure of a protein with its function (and vice versa). Many proteins produced by recombinant DNA technology are present only in trace amounts in nature (e.g., interferons and other cytokines; Ealick et al., 1991), and authentic material is not available for detailed molecular characterization. Knowledge of the 3-D structure allows a rational approach to protein engineering and the design of drugs that modulate the biological activity of the protein. The substitution,
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deletion, and insertion of residues allow a structure-function hypothesis to be tested and new, sometimes improved protein variants (or mutants) to be produced. NMR Spectroscopy It is a major challenge to produce proteins suitable for structural determination, not only in terms of quality, but in terms of the quantity which may be required, especially for NMR (UNIT 17.5). Many proteins, although they have native-like structure and biological activity, are not suitable for structural determination due to, e.g., limited solubility, conformational flexibility (floppy regions/domains), and heterogeneity of posttranslational modifications (especially carbohydrate). Often, these problems can be resolved by a combination of protein biochemistry and protein engineering approaches and requires a close collaboration between the structural biologist and the molecular/protein chemist. The NMR determination of the HIV-1 Nef structure is an example of this integrated approach (see above). Structural determination in solution by multidimensional NMR is presently limited to proteins 30 to 40 kDa (reviewed by Clore and Gronenborn, 1994). One of the largest proteins solved to date is the 44-kDa trimeric SIV gp41 ectodomain (Caffrey et al., 1998). Larger proteins can be studied incrementally (using the dissection approach) if information on the domain boundaries is known (Campbell and Downing, 1998). The sample demands can be as high as several hundred milligrams, and larger proteins (>10 kDa) must be uniformly labeled with various combinations of 2H, 13C, and 15N. Some of the labeling scenarios required to solve the HIV Nef structure are presented in Table 1 of Grzesiek et al. (1997). New developments in isotope labeling strategies are reviewed by Goto and Kay (2000). Labeling in E. coli is achieved by growing the bacteria in minimal medium containing one or more of the following stable (nonradioactive) isotopes: 15 NH4Cl (sole nitrogen source), [13C] glucose (sole carbon source), and 2H2O (UNIT 5.3). The 15N and 13C labeling of the HIV protease using a 2-liter fermentor is detailed by Yamazaki et al. (1996). Label incorporation is conveniently monitored by mass spectrometry of the purified protein. Over the lifetime of the structural study (4 to 12 months), because of the multisample requirements, a reliable and robust purification method is essential. It should also be noted that the labeling requirement usually dictates that the protein be produced in bacteria, although labeled proteins have been produced in yeast and insect cells (Goto and Kay, 2000). For NMR purposes, the recombinant protein must be homogeneous and soluble at 1 to 3 mM concentrations, preferably with solvents below pH 7.0 and at temperatures >30°C. As measurements take many hours to complete, the presence of trace amounts of proteases can ruin the experiment. In addition, particular attention must be paid to maintaining solvent-accessible and reactive cysteines (unpaired) in the reduced state (usually by including DTT or TCEP) and often cysteines are mutated to alanine residues (Wingfield et al., 1997). Protein crystallization and X-ray crystallography
Purification of Recombinant E. coli Proteins
The rate-limiting step in structure determination using X-ray crystallography is production of crystals that diffract to high resolution (UNIT 17.4). The scientists involved in the production and characterization of the protein are often best situated to crystallize the protein. Furthermore, once crystallization conditions have been optimized, it can be quite easy to interest structural groups in collaboration.
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Determining optimal crystallization conditions may require as little as a few milligrams or as much as 100 mg of pure protein. The protein itself must usually be physically and chemically homogeneous; small amounts of protein impurities may significantly interfere with crystallization. In general, physical homogeneity is more critical than chemical homogeneity. Some of the methods used to establish physical and chemical homogeneity are discussed elsewhere (Chapter 7; Jones et al., 1994). Many investigators use the sparse matrix sampling technology to screen for initial crystallization conditions and commercial kits are available for this purpose (e.g., Hampton Research: http://www.hamptonresearch.com/index.html. The company site also has useful tips and protocols). The phase problem in crystallographic analysis has traditionally been solved by isomorphous replacement with heavy atoms, but over the last 10 years, multiwavelengh anomalous diffraction (MAD) has gained popularity (Hendrickson et al., 1990). For the latter approach, selenium is incorporated into recombinant proteins via selenomethionine (seleno-L-methionine; available from Sigma and others) using a methionine-requiring auxotroph. In studies of the gp41 protein (mentioned above), the T7 expression system (Novagen, http://www.novagen.com) and the host strain B834/DE3 (Novagen) were used. Briefly, transformed cells were grown overnight in a 0.5-liter shaker flask containing minimal media plus 1 mM methionine. Cells were collected and resuspended in 0.5 liters of media minus methionine. The cells were grown at 37°C in a small fermentor and fed 5 ml of 10 mg/ml selenomethionine. The cells were induced for 3 hr with IPTG and fed an additional 50 mg of selenomethionine (total feed: 100 mg). Cells were collected (∼7.0 g wet weight) and 130 mg of pure gp41 ectodomain was isolated as described above. Mass spectrometry indicated that the single methionine was >98% labeled. The protein was then crystallized as previously described (Wingfield et al., 1997). For more details on labeling using E. coli, see Chapter 5 (UNIT 5.3). Selenomethionine incorporation into eukaryotic systems is not as successful as in E. coli; incorporation can be as high as 90% in baculovirus, but only ∼60% in yeast. The production of well-defined protein complexes for structural studies can be straightforward. For example, monomeric proteins can be expressed in bacteria which self-associate into stable complexes ranging from simple dimers (e.g., γ-IFN) and trimers (e.g., α-TNF) to complex structures such as viral nucleocapsids (e.g., Hepatitis B Virus core antigen, 180-mer). These stable (tightly associated) homopolymers are well suited for structural studies. Heteroprotein complexes can be made by either co-expression of protein subunits (Johnson et al., 2000; Kholod and Mustelin, 2001) or by in vitro assembly of individual components. The former approach may be required in the case where individual subunits are unstable (Nash et al., 1987). In contrast to stable complexes, there are many biologically significant complexes characterized by weak association. Many protein-protein interactions of interest, e.g., signal transduction pathways, may be somewhat transitory and involve weak interactions. In these complexes, the dissociation constants (Kd) between proteins are <10−6 M, and therefore, tend to exhibit concentrationdependent reversible self-association. This behavior results in physical heterogeneity, thus, complicating crystallization attempts. NMR has been used to examine weakly associating systems, e.g., the binding of the CD4 determinant to HIV-1 Nef (Grzesiek et al., 1996). To study protein-protein complexes characterized by low Kd, it may be necessary to use protein engineering and other approaches to generate more stable and tighter interactions. It has been mentioned that one of the reasons for fusion tagging proteins is to increase their potential for stable expression and accumulation. The enhancement of a protein’s physical properties, especially solubility, by appending protein (e.g., GST) or peptide (e.g., FLAG) tags makes these fusion proteins good candidates for crystallization trials.
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Some investigators have reported problems with His-tagged proteins and it is generally recommended to either remove this tag for structural work or use another tag. Biophysical Studies Low-resolution structural studies using various biophysical methodologies (Jones et al., 1994) can be made with less material (<1 to 10 mg). Proteins for spectroscopic studies should be >95% pure and previously fractionated on a gel-filtration column to remove aggregated and possibly misfolded variants. The removal of aggregates is especially important for spectroscopic studies including UV/vis, fluorescence and circular dichroism where excessive light scattering must be avoided (see Chapter 7; Colon, 1999). Various labeling and tagging strategies can be used to aid both structural and functional studies. The most common approach is to append affinity tags that can then be used to immobilize the protein in a directed manner (Nilsson et al., 1997). This approach is especially useful for studying protein interactions. Also, analogous to the in vivo protein labeling scenarios as described above for selenomethionine, specific residues can be modified. For example, tryptophan in recombinant proteins can be replaced by 5-hydroxytryptophan by using an E. coli Trp auxotroph. Protein thus labeled has a strong absorbance at 310 nm that can be exploited in structure-function studies (Laue et al., 1993). SPECIALIZED EQUIPMENT Breaking and Fractionating Cells For small- to medium-scale work on a regular basis, a French press (Thermo Spectronic, http://www.thermo.com) with a continuous-fill cell is recommended (UNIT 6.2). It is also useful for breaking yeast cells. For large-scale work (>500 ml), the Manton-Gaulin-APV homogenizer (APV Gaulin) is recommended. For further processing of cells and cell lysates (e.g., UNITS 6.2 & 6.3), an ultrasonic homogenizer is required. An instrument with a 400-W (or higher) capacity is recommended (Branson, http://www.bransonultrasonics.com). After low-speed centrifugation using standard preparative centrifuges (Beckman Coulter Preparative Centrifuge, Avanti Series can be found at http://www.beckman.com), highspeed centrifugation is a convenient and rapid cleanup step before column chromatography (Fig. 6.1.3). With Beckman ultracentrifuges, the 45 Ti rotor is recommended. This six-place rotor has a maximum speed of 235,000 × g; with thick-walled polycarbonate tubes, its capacity is ∼400 ml. Chromatographing Proteins
Purification of Recombinant E. coli Proteins
Most chromatography is carried out at 4°C either in a cold room or, more conveniently, in a cold cabinet in the laboratory. The basic components of a chromatography system are as follows: column, column matrix, pumps, a gradient-making device, UV/visible or other detection system, and a fraction collector. These components can be bought as units such as the AKTA Explorer or FPLC chromatograph systems (Amersham Bioscience, http://www.apbiotech.com), which can be used for laboratory-scale to large-scale work. Systems can also be custom assembled from individual components from Amersham and other vendors. Column matrices can be purchased prepacked or as bulk media that are packed in columns by the user. Ion-exchange separations, using standard low- to mediumpressure resins (agarose/dextran/cellulose-based), require at least one narrow (2.5-cm) and one wide (5.0-cm) column with adjustable flow adapters so that the resin height can be varied between 5 and 30 cm. Gel filtration requires columns with diameters of 1.25
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and 2.5 cm (5 cm for larger-scale work) and lengths of 60 to 100 cm. Simple gradient makers with capacities of 150 ml to 2 liters are generally available. Concentrating Proteins Stirred ultrafiltration cells are recommended for laboratory-scale work. The cells range in size from 3 ml to 2 liters and are used in conjunction with variable molecular weight cutoff membranes (Millipore, http://www.millipore.com). For larger volumes, Millipore also sells various systems. For smaller volumes (0.5 to 15 ml), centrifugational concentrators are available (Millipore and others). For a review of the equipment used for protein concentration, see Harris (1989). Making Analytical Measurements A protein purification laboratory should have a dependable scanning UV/visible spectrophotometer, ideally an instrument with computerized data collection and analysis. Hewlett Packard (Agilent) instruments with diode array detectors are recommended for most routine work (http://www.chem.agilent.com). For laboratories specializing in purifying recombinant proteins from E. coli, access to a spectropolarimeter (e.g., Jasco J-810, http://www.jascoinc.com) will be helpful for monitoring and developing folding protocols. For rapid chemical characterization and identity check of proteins, access to a mass spectrometer is also desired (Chapter 16). Most of the companies mentioned above have excellent Web sites where technical information is posted. The series of handbooks on chromatographic separations published by Amersham Biosciences can be conveniently downloaded as pdf files. Literature Cited Allet, B., Payton, M., Mattaliano, R.J., Gronenborn, A.M., Clore, G.M., and Wingfield, P.T. 1988. Purification and characterization of the DNA-binding protein Ner of bacteriophage Mu. Gene 65:259-268. Arakawa, T. and Timasheff, S.N. 1985. Theory of protein solubility. Methods Enzymol. 114:49-77. Armstrong, N., De Lencastre, A., and Gouaux, E. 1999. A new protein folding screen: Application to the ligand binding domain of a glutamate and kainite receptor and to a lysozyme and carbonic anhydrase. Protein Sci. 8:1475-1483. Asenjo, J.A. and Patrick, I. 1990. Large-scale protein purification. In Protein Purification Applications: A Practical Approach (E.L.V. Harris and S. Angal, eds.) pp. 1-27. IRL Press, Oxford. Baneyx, F. 1999. Recombinant protein expression in Escherichia coli. Curr. Opin. Biotechnol. 10:411-421. Beacham, I.R. 1979. Periplasmic enzymes in Gram-negative bacteria. Int. Biochem. 10:877-883. Ben-Bassat, A., Bauer, K., Chang, S.-Y., Myambo, K., Boosman, A., and Chang, S. 1987. Processing of the initiation methionine from proteins: Properties of the E. coli methionine aminopeptidase and its gene structure. J. Bacteriol. 169:751-757. Bessette, P.H., Aslund, F., Beckwick, J., and Georgiou, G. 1999. Efficient folding of proteins with multiple disulfide bonds in the Escherichia coli cytoplasm. Proc. Natl. Acad. Sci. U.S.A. 96:13703-13708. Bibi, E. and Beja, O. 1994. Membrane topology of multidrug resistant protein expressed in E. coli. J. Biol. Chem. 31:19910-19915. Bowden, G.A., Paredes, A.M., and Georgiou, G. 1991. Structure and morphology of protein inclusion bodies in E. coli. Biotechnology 9:725-730. Braun, P., Gerritse, G., van Dijl, J.-M., and Quax, W.J. 1999. Improving protein secretion by engineering components of the bacterial translocation machinery. Curr. Opin. Biotechnol. 10:376-381. Buchner, J. and Rudolph, R. 1992. Renaturation and characterization of recombinant Fab fragments produced in Escherichia coli. Biotechnology 9:157-162. Burgess, R.R. and Jendrisak, J.J. 1975. A procedure for the rapid, large-scale purification of E. coli DNA-dependent RNA polymerase involving polymin P precipitation and DNA-cellulose chromatography. J. Biol. Chem. 14:4634-4638. Purification of Recombinant Proteins
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Burgess, R.R. and Knuth, M.W. 1996. Purification of a recombinant protein overproduced in Escherichia coli. In Strategies for Protein Purification and Characterization: A Laboratory Course Manual (D.R. Marshak, J.T. Kadonaga, R.R. Burgess, M.W. Knuth, W.A. Brennan, and S.-H. Lin, eds.) pp. 205-217 and 245-262. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Caffrey, M., Cai, M., Kaufman, J., Stahl, S., Wingfield, P.T., Covell, D.G., Gronenborn, A.M., and Clore, G.M. 1998. Three dimensional solution structure of the 44 kDa ectodomain of SIV gp41. EMBO J. 17:4572-4584. Campbell, I. and Downing, A.K. 1998. NMR of modular proteins. Nat. Struct. Biol. 5:496-499. Cereghino, G.P.L. and Clegg, J.M. 1999. Applications of yeast in biotechnology: Protein production and genetic analysis. Curr. Opin. Biotechnol. 10:422-427. Cheng, Y.-S.E., McGowan, M.H., Kettner, C.A., Schloss, J.V., Erickson-Viitanen, S., and Yin, F.H. 1990. High synthesis of recombinant HIV-1 protease and the recovery of active enzyme from inclusion bodies. Gene 87:243-248. Cleary, S., Mulkerrin, M.G., and Kelley, R.F. 1989. Purification and characterization of tissue plasminogen activator kringle-2 domain expressed in E. coli. J. Biol. Chem. 28:1884-1891. Cleland, J.L., Builder, S.E., Swartz, J.R., Winkler, M., Chang, J.Y., and Wang, D.I.C. 1992. Polyethylene glycol enhanced protein refolding. Biotechnology 10:1013-1019. Clore, G.M. and Gronenborn, A.M. 1994. Multidimensional heteronuclear nuclear magnetic resonance of proteins. Methods Enzymol. 239:249-363. Cole, P.A. 1996. Chaperone-assisted protein expression. Structure 4:239-242. Colon, W. 1999. Analysis of protein structure by solution optical spectroscopy. Methods Enzymol. 309:605632. Cornelis, P. 2000. Expressing genes in different Escherichia coli compartments. Curr. Opin. Biotechnol. 11:450-454. Creighton, T.E. 1984. Disulfide bond formation in proteins. Methods Enzymol. 107:305-329. Creighton, T.E. 1993. Proteins: Structures and Molecular Properties, 2nd ed. pp. 292-296. Freeman, New York. Dale, G.E., Broger, C., Langen, H., D’Arcy, A., and Struber, D. 1994. Improving protein stability through rationally designed amino acid replacements: Solubilization of the trimethoprim-resistant type S1 dihydrofolate reductase. Protein Eng. 7:933-939. Danley, D.E., Strick, C.A., James, L.C., Lanzetti, A.J., Otterness, I.G., Grenett, H.E., and Fuller, G.M. 1991. Identification and characterization of a C-terminally extended form of recombinant murine IL-6. FEBS Lett. 283:135-139. Darby, N.J. and Creighton, T.E. 1990. Folding proteins. Nature 344:715-716. De Bernardez Clark, E. 2001 Protein folding for industrial processes. Curr. Opin. Biotechnol. 12:202-207. De Bernardez Clark, E., Schwartz, E., and Rudolph, R. 1999. Inhibition of aggregation side reactions during in-vitro protein folding. Methods Enzymol. 309:217-236. deVos, A.M., Ultsch, M., and Kossiakoff, A.A. 1992. Human growth hormone and extracellular domain of its receptor: Crystal structure of the complex. Science 255:306-312. Diederichs, K., Boone, T., and Karplus, A. 1991. Novel fold and putative receptor binding site of granulocyte-macrophage colony stimulating factor. Science 254:1779-1782. DiRienzo, J.M., Nakamura, K., and Inouye, M. 1978. The outer membrane proteins of Gram-negative bacteria: Biosynthesis, assembly and function. Annu. Rev. Biochem. 47:481-532. Dyson, H.J. and Wright, P.E. 2001. Coupling of folding and binding for unstructured proteins. Curr. Opin. Struct. Biol. 12:54-60. Ealick, S.E., Cook, W.J., Vijay-Kumar, S., Carson, M., Nagabhushan, T.L., Trotta, P.P., and Bugg, C.E. 1991. Three-dimensional structure of recombinant human interferon-gamma. Science 252:698-702. Eisenberg, D. 1999. How chaperones protect virgin proteins. Science 285:1021-1022. Ellis, R.J. 1994. Role of chaperones in protein folding. Curr. Opin. Struct. Biol. 4:117-122. Ellis, R.J. 2001. Macromolecular crowding: An important but neglected aspect of the intracellular environment. Curr. Opin. Struct. Biol. 11:114-119. Ellis, R.J. and Hart, F.U. 1999. Principles of protein folding in the cellular environment. Curr. Opin. Struct. Biol. 9:102-110. Purification of Recombinant E. coli Proteins
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Janson, J.-C. and Ryden, L. 1989. Protein Purification: Principles, High Resolution Methods, and Applications. VCH Publishers, New York. Johnson, B.H. and Hecht, M.H. 1994. Recombinant proteins can be isolated from E. coli by repeated cycles of freezing and thawing. Biotechnology 12:1357-1360. Johnson, K., Clements, A., Venkataramani, R.N., Trievel, R.C., and Marmorstein, R. 2000. Coexpression of proteins in bacteria using a T7-based expression plasmid: Expression of heteromeric cell cycle and transcriptional regulatory complexes. Protein Expr. Purif. 20:435-443. Jones, C., Mulloy, B., and Thomas, A.H. 1994. Microscopy, optical spectroscopy, and macroscopic techniques. Methods Mol. Biol. 22:1-245. Jones, D.H., Ball, E.H., Sharpe, S., Barber, K.R., and Grant, C.W.M. 2000. Expression and membrane assembly of a transmembrane region from Neu. Biochemistry 39:1878-1878. Kaback, H.R. 1971. Bacterial membranes. Methods Enzymol. 22:99-120. Kamireddi, M., Eisenstein, E., and Reddy, P. 1997. Stable expression and rapid purification of Escherichia coli GroEL and GroES chaperones. Protein Expr. Purif. 11:47-52. Kelley, W.S. and Stump, K.H. 1979. A rapid procedure for isolation of large quantities of E. coli DNA polymerase 1 utilizing a λ polA transducing phage. J. Biol. Chem. 254:3206-3210. Kern, G., Kern, D., Jaenicke, R., and Seckler, R.L. 1993. Kinetics of folding and association of differentially glycosylated variants of invertase from Saccharomyces cerevisiae. Protein Sci. 2:1862-1868. Kholod, N. and Mustelin, T. 2001. Novel vectors for co-expression of two proteins in E.coli. Biotechniques 31:322-328. Kiefhaber, T., Rudolph, R., Kohler, H.-H., and Buchner, J. 1991. Protein aggregation in vitro and in vivo: A quantitative model of the kinetic competition between folding and aggregation. Biotechnology 9:825-829. Kohno, T., Carmichael, D.F., Sommer, A., and Thompson, R.C. 1990. Refolding of recombinant proteins. Methods Enzymol. 185:187-195. Kost, T.A. and Condreay, J.P. 1999. Recombinant baculovirus as expression vectors for insect and mammalian cells. Curr. Opin. Biotechnol. 10:428-433. Laue, T.M., Senear, D.F., Eaton, S., and Ross, J.B.A. 1993. 5-Hydroxytryptophan as a new intrinsic probe for investigating protein-DNA interactions by analytical ultracentrifugation. Study of the effect of DNA on the self-assembly of the bacteriophage λ cI repressor. J. Biol. Chem. 32:2469-2472. LaVallie, E.R., DiBlasio, E.A., Kovacic, S., Grant, K.L., Schendel, P.F., and McCoy, J.M. 1993. A thioredoxin gene fusion expression system that circumvents inclusion body formation in the E. coli cytoplasm. Biotechnology 11:187-193. Lilie, H., Schwartz, E. and Rudolph, R. 1998 Advances in refolding of proteins produced in E.coli. Curr. Opin. Biotechnol. 9:497-501. Lindwall, G., Chau, M.-F., Gardner, S.R., and Kohlstaedt, L.A. 2000. A sparse matrix approach to the solubilization of overexpression proteins. Protein Eng. 13:67-71. London, J., Skrzynia, C., and Goldberg, M.E. 1974. Renaturation of Escherichia coli tryptophanase after exposure to 8 M urea. Eur. J. Biochem. 47:409-415. Lu, H.S., Fausset, P.R., Sotos, L.S., Clogston, C.L., Rohde, M.F., Stoney, K.S., and Herman, A.C. 1993. Isolation and characterization of three recombinant human granulocyte colony stimulating factor His to Gln isoforms produced in E. coli. Protein Expr. Purif. 4:465-472. Markrides, S. 1996. Strategies for achieving high-level expression of genes in Escherichia coli. Microbiol.l Rev. 60:512-538. Marston, F.A.O. and Hartley, D.L. 1990. Solubilization of protein aggregates. Methods Enzymol. 182:264276. Matthew, J.B., Friend, S.H., Botelho, L.D., Lehman, L.D., Hanania, G.I., and Gurd, F.R.H. 1978. Discrete charge calculations of potentiometric titrations for globular proteins. Biochem. Biophys. Res. Commun. 81:416-421. Maurizi, M.R. 1992. Proteases and protein degradation in Escherichia coli. Experientia 48:178-201. Milburn, M.V., Hassel, A.M., Lambert, M.H., Jordon, S.R., Proudfoot, A.E.I., Graber, P., and Wells, T.N.C. 1993. A novel dimer configuration revealed by the crystal structure at 2.4 angstrom resolution of human interleukin-5. Nature 363:172-176. Mildner, A.M., Rothrock, D.J., Leone, J.W., Bannow, C.A., Lull, J.M., Reardon, I.M., Sarcich, J.L., Howe, W.J., Tomich, C.-S.C., Smith, C.W., Heinrikson, R.L., and Tomasselli, A.G. 1994. The HIV-1 protease as enzyme and substrate: Mutagenesis of autolysis sites and generation of a stable mutant with retained kinetic properties. Biochemistry 33:9405-9413. Purification of Recombinant E. coli Proteins
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Sottrup-Jensen, L. 1989. Alpha-macroglobulins: Structure, shape, and mechanism of proteinase complex formation. J. Biol. Chem. 264:11539-11542. Stahl, S.J., Wingfield, P.T., Kaufman, J.D., Pannell, L.K., Cioce, V., Sakata, H., Taylor, W.G., Rubin, J.S., and Bottaro, D.P. 1997. Functional and biophysical characterization of recombinant human growth factor isoforms produced in Escherichia coli. Biochem. J. 326:763-772. Stark, G.R., Stein, W.H., and Moore, S. 1960. Reactions of cyanate present in aqueous urea with amino acids and proteins. J. Biol. Chem. 235:3177-3181. Stein, S. 1991. Isolation of natural proteins. In Fundamentals of Protein Biotechnology (S. Stein, ed.) pp. 137-160. Marcel Dekker, New York. Sutcliffe, J.G., Shinnick, T.M., Green, N., and Lerner, R.A. 1983. Antibodies that react with predetermined sites on proteins. Science 219:660-665. Swartz, J.P. 2001. Advances in Escherichia coli production of therapeutic proteins. Curr. Opin. Biotechnol. 12:195-201. Tanford, C. 1968. Protein denaturation. Adv. Protein Chem. 23:122-275. Thatcher, D.R. 1996. Industrial scale purification of proteins. In Proteins LabFax (N.C. Price, ed.) pp. 131-137. BIOS Scientific Publishers, Oxford. Thatcher, D.R. and Panayotatos, N. 1986. Purification of recombinant IFN-2. Methods Enzymol. 119:166177. Thatcher, D.R., Wilks, P., and Chaudhuri, J. 1996. Inclusion bodies and refolding. In Proteins LabFax (N.C. Price, ed.) pp. 119-130. BIOS Scientific Publishers, Oxford. Timasheff, S.N. and Arakawa, T. 1997. Stabilization of protein structure by solvents. In Protein Structure: A Practical Approach. Second Edition (T.E. Creighton, ed.) pp. 349 -363. IRL Press, Oxford. Uhlen, M., Forberg, G., Moks, T., Hartmanis, M., and Nilsson, B. 1992. Fusion proteins in biotechnology. Curr. Opin. Biotechnol. 3:363-369. Ultsch, M., deVos, A.M., and Kossiakoff, A.A. 1991. Crystals of the complex between human growth hormone and the extracellular domain of its receptor. J. Mol. Biol. 222:865-868. Van Reis, R. and Zydney, A. 2001. Membrane separations in biotechnology. Curr. Opin. Biotechnol. 12:208-211. Walter, H. and Johansson, G. 1986. Partitioning in aqueous two-phase systems: An overview. Anal. Biochem. 155:215-242. Walter, M.R., Cook, W.J., Ealick, S.E., Nagabhushan, T.L., Trotta, P.P., and Bugg, C.E. 1992. Three-dimensional structure of human recombinant granulocyte-macrophage colony stimulating factor. J. Mol. Biol. 224:1075-1085. Wang, Y.-X., Neamati N, Jacob, J., Palmer, I., Stahl, S.J., Kaufman, J.D., Huang, P.L., Huang, P.L., Winslow, H.E., Pommier, Y., Wingfield, P.T., Lee-Huang, S., Bax, A., and Torchia, D.A. 1999. Solution structure of anti-HIV-1 and anti-tumor protein MAP30: Structural insights into its multiple functions. Cell 99:433-442. Watanabe, E., Tsoka, S., and Asenjo, J.A. 1994. Selection of chromatographic protein purification operations based on physicochemical properties. Ann. N.Y. Acad. Sci. 721:348-364. Wetlaufer, D.B. 1984. Nonenzymatic formation and isomerization of protein disulfides. Methods Enzymol. 107:301-304. Wetlaufer, D.B., Branca, P.A., and Chen, G.-X. 1987. The oxidative folding of proteins by disulfides plus thiol does not correlate with redox potential. Protein Eng. 1:141-146. Wetzel, R. 1992. Principles of protein stability. Part 2—Enhanced folding and stabilization of proteins by suppression of aggregation in vitro and in vivo. In Protein Engineering: A Practical Approach (A.R. Rees, M.J.E. Sternberg, and R. Wetzel, eds.) pp. 191-216. IRL Press, Oxford. Widmann, M. and Christen P. 2000. Comparison of folding rates of homologous prokaryotic and eukaryotic proteins. J. Biol. Chem. 275:18619-18622. Wingfield, P.T., Mattaliano, R.J., MacDonald, H.R., Craig, S., Clore, G.M., Gronenborn, A.M., and Schmeissner, U. 1987a. Recombinant-derived interleukin 1α stabilized against specific deamidation. Protein Eng. 1:413-417. Wingfield, P.T., Graber, P., Rose, K., Simona, M.G., and Hughes, G.J. 1987b. Chromatofocusing of N-terminally processed forms of proteins: Isolation and characterization of two forms of interleukin 1β and bovine growth hormone. J. Chromatogr. 387:291-300. Wingfield, P.T., Payton, M., Graber, P., Rose, K., Dayer, J.-M., Shaw, A.R., and Schmeissner, U. 1987c. Purification and characterization of human interleukin 1α produced in Escherichia coli. Eur. J. Biochem. 165:537-541. Purification of Recombinant E. coli Proteins
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Current Protocols in Protein Science
Wingfield, P.T., Stahl, S.J., Payton, M.A., Venkatesan, S., Misra, M., and Steven, A. 1990. HIV-1 Rev expressed in recombinant Escherichia coli: Purification polymerization and conformational properties. Biochemistry 30:7527-7534. Wingfield, P.T., Stahl, S.J., Williams, R.W., and Steven, A.C. 1995. Hepatitis core antigen produced in E. coli: Conformational analysis, and in vitro assembly. Biochemistry 34:4919-4932. Wingfield, P.T., Stahl, S.J., Kaufman, J., Zlotnick, A., Hyde, C.C., Gronenborn, A.M., and Clore G.M. 1997. The extracellular domain of immunodeficiency virus gp41 protein: Expression in Escherichia coli, purification and crystallization. Protein Sci. 6:1653-1660. Wingfield, P.T., Stahl, S.J., Thomsen, D.R., Homa, F.L., Booy, F.P., Trus, B.L., and Steven, A.C. 1997a. Hexon-only binding of VP26 reflects differences between the hexon and penton conformations of the VP5, the major capsid protein of Herpes Simplex Virus. J. Virology 71:8955-8961. Wingfield, P.T., Stahl, S.J., Kaufman, J., Palmer, I., Chung, V., Sax, J.K., Kleiner, D.E., and Stetler-Stevenson, G.W. 1999. Functional and biophysical characterization of full length, recombinant human TIMP-2 produced in Escherichia coli: Comparison of wild type and N-terminal alanine substituted variant. J. Biol. Chem. 274:21362-21368. Wurm, F. and Bernard, A. 1999. Large–scale transient expression in mammalian cells for recombinant protein production. Curr. Opin. Biotechnol. 10:156-159. Wyss, D.F. and Wagner, G. 1996. The structure of sugars in glycoproteins. Curr. Opin. Struct. Biol. 7:409-416. Yamazaki, T., Hinck, A.P., Wang, Y-X., Nicholson, L.K., Torchia, D.A., Wingfield, P.T., Stahl, S.J., Kaufman, J.D., Chang, C.-H., Domaille, P.J., and Lam, P.Y.S. 1996. Three dimensional solution structure of the HIV-1 protease complexed with DMP 323, a novel cyclic urea-type inhibitor, determined by nuclear magnetic resonance spectroscopy. Protein Sci. 5:495-506. Yang, Z.-N., Mueser, T.C., Kaufman, J., Stahl, S.J., Wingfield, P.T., and Hyde, C. 1999. The structure of the SIV gp41 ectodomain at 1.47 A. J. Struct. Biol. 126:133-144. Yarranton, G.T. and Mountain, A. 1992. Expression of proteins in prokaryotic systems—Principles and case studies. In Protein Engineering: A Practical Approach (A.R. Rees, M.J.E. Sternberg, and R. Wetzel, eds.) pp. 303-324. IRL Press, Oxford. Zardeneta, G. and Horowitz, P.M. 1994. Detergent, liposome, and micelle-assisted protein refolding. Anal. Biochem. 223:1-6. Zhang, Y., Olsen, D.R., Nguyen, K.B., Olson, P.S., Rhodes, E.T., and Mascarenhas, D. 1998. Expressoin of eukaryotic proteins in soluble form in Escherichia coli. Protein Expr. Purif. 12:159-165.
Contributed by Paul T. Wingfield National Institutes of Health Bethesda, Maryland
Purification of Recombinant Proteins
6.1.37 Current Protocols in Protein Science
Supplement 30
Preparation of Soluble Proteins from Escherichia coli
UNIT 6.2
Once a suitable protein expression system involving Escherichia coli is developed and optimized (UNITS 5.1 & 5.2), large-scale production of recombinant proteins (UNIT 5.3) generates large quantities of culture material from which the protein of interest must be purified. Harvesting (UNIT 5.3) produces cell concentrate or culture medium, depending on the subcellular localization of the protein. Cell paste is the starting material for purification of proteins expressed in soluble form inside cells, such as interleukin 1β (IL-1β). Human IL-1β is a 153-residue (17.4-kDa) protein cytokine of biomedical importance that plays a central role in immune and inflammatory responses. Purification of human IL-1β is used as an example of the preparation of soluble proteins from E. coli. Bacteria containing IL-1β are lysed, and the resulting supernatant is clarified to remove ribosomes and other particulate matter. The sample is then applied to an anion-exchange column to separate recombinant IL-1β from cellular contaminants, such as E. coli proteins, nucleic acids, and lipopolysaccharides. The sample is further purified through salt precipitation and cation-exchange chromatography, then concentrated. Finally, the IL-1β protein is applied to a gel-filtration column to separate it from remaining higherand lower-molecular-weight contaminants, the purified protein is stored frozen or is lyophilized. The purification protocol described is typical for a protein that is expressed in fairly high abundance (i.e., >5% total protein) and accumulates in a soluble state. With these expression levels, only about a 20-fold overall purification is required to obtain pure protein. Therefore, conventional chromatographic methods can be used, and normally only three or four purification stages are required (see Table 6.2.1 for an outline of the procedure as applied to IL-1β, including time considerations). The process can be shortened somewhat through the use of the Pharmacia Biotech BioPilot or FPLC systems (see Time Considerations). SDS-PAGE (UNIT 10.1) is used to monitor column fractions for the presence of IL-1β, which is detected as a stained band (UNIT 10.5) at the expected 17.4-kDa location. This is common practice in recombinant protein purification, as the specific assays for many proteins, including IL-1β, are relatively complex. The original expression and fermentation experiments involving IL-1β (Wingfield et al., 1986) followed procedures similar to those given in UNIT 5.3. CAUTION: Avoid direct contact with IL-1β-containing solutions. Trace material from aerosols or from hand contact can cause severe inflammation of the eyes. Safety glasses and gloves should be worn. PURIFICATION OF A PROTEIN EXPRESSED IN ESCHERICHIA COLI IN A SOLUBLE STATE: INTERLEUKIN 1β Materials DEAE Sepharose CL-4B resin (Pharmacia Biotech) Anion-exchange buffer (see recipe) 0.26% (w/v) sodium hypochlorite/70% ethanol or 5% (v/v) bleach (e.g., Clorox)/70% ethanol E. coli cells (∼50 g wet weight) from fermentation (UNIT 5.3) containing IL-1β Lysis buffer (see recipe) Contributed by Paul T. Wingfield Current Protocols in Protein Science (1995) 6.2.1-6.2.15 Copyright © 1995 by John Wiley & Sons, Inc.
BASIC PROTOCOL
Purification of Recombinant Proteins
6.2.1 CPPS
Bovine pancreas DNase I and RNase A (Worthington; optional, for reducing solution viscosity) 2 N sodium hydroxide Ammonium sulfate, ground with mortar and pestle Cation-exchange buffer (see recipe) CM Sepharose CL-4B (Pharmacia Biotech) Cation-exchange buffer/250 mM NaCl (see recipe) Tris base Gel-filtration buffer (see recipe) Ultrogel AcA54 gel-permeation resin (BioSepra) Lyophilization buffer (see recipe; optional) 2- or 3-liter sintered glass funnel with fritted disc (coarse porosity) and 5-liter filter flask Chromatography columns (preferably glass) with adjustable flow adapters: one (or optionally two) 5 × 50 cm and one 2.5 × 100 cm (Pharmacia Biotech, Amicon, or equivalent) RK50 packing reservoir (Pharmacia Biotech) Peristaltic pump, UV monitor, and fraction collector (Pharmacia Biotech or equivalent) 16 × 150–mm culture tubes 40-ml French pressure cell and rapid-fill kit (SLM-AMINCO) Aminco laboratory press (SLM-AMINCO) 1-liter Waring commercial blender
Table 6.2.1
Day
Outline of Interleukin β Purificationa
Steps (low pressure)
1
Preparation of DEAE Sepharose column As for low pressure (steps 1-17) (steps 1-3) Cell breakage (steps 4-8) Clarification of lysate (steps 9 and 10) DEAE Sepharose chromatography (step 11) SDS-PAGE of DEAE Sepharose fractions (step 12) (NH4)2SO4 fractionation (steps 13-15) Dialysis (steps 16 and 17)
2
CM Sepharose chromatography (steps 18-22) SDS-PAGE of CM Sepharose fractions (step 22)
Fast Flow (substitute for CM Sepharose; steps 18-21) SDS-PAGE of Fast Flow fractions (steps 22) Concentration (step 23a) Superdex 75 gel filtration (substitute for Ultrogel; steps 24-26)
3
Concentration (step 23a or 23b)
SDS-PAGE of Superdex 75 fractions (step 26) Concentration (step 27)
Ultrogel gel filtration (steps 24-26) 4 Preparation of Soluble E. coli Proteins
Steps (BioPilot or FPLC)
SDS-PAGE of gel-filtration fractions (step 26) Concentration (step 27)
aStep numbers in parentheses refer to the Basic Protocol. Chromatography materials as well as the BioPilot and FPLC
systems are from Pharmacia Biotech.
6.2.2 Current Protocols in Protein Science
250, 500, and 1000-ml stainless steel beakers Ice bucket, ∼4 liter Tissue-grinder homogenizer (Polytron Model PT 10/35, Brinkmann) Ultrasonic homogenizer, ≥400 W, with sound enclosure (Branson or equivalent) Preparative centrifuge: Beckman J2-21M Rotors for preparative centrifuge: Beckman JA-14 (capacity 6 × 250 ml) or JA-20 (capacity 8 × 50 ml) Ultracentrifuge: Beckman Optima XL-90 Rotors for ultracentrifuge: Beckman 45Ti (capacity 6 × 100 ml) or 35Ti (capacity 6 × 94 ml) Conductivity meter (Radiometer America) Spectra/Por 1 dialysis tubing (Spectrum) Gradient maker: Model 2000 (working volume 0.65 to 2 liters; Life Technologies) 200- or 400-ml stirred ultrafiltration cell and Diaflo ultrafilter PM10 or YM3 membranes (Amicon; optional) Millex-GV 0.22-µm-pore-size filter units (Millipore) 10- or 20-ml syringe Additional materials and equipment for SDS-PAGE (UNIT 10.1) and dialysis (APPENDIX 3B) NOTE: All protocol steps are carried at 4°C unless otherwise stated. Forces for centrifugation steps refer to the maximum × g (i.e., centrifugal force at the bottom of the tubes). Prepare anion-exchange column 1. Pour 400 to 500 ml DEAE Sepharose CL-4B ion-exchange resin into a sintered-glass funnel and wash with several liters water followed by 1 liter anion-exchange buffer (pH 8.5). Measure the conductivity of the starting buffer and eluted buffer to make sure they are the same before proceeding to the next step. The resin is supplied in 500-ml bottles as a slurry in 20% ethanol. When washing the resin, do not allow it to run dry on the filter funnel. Laboratory vacuum (e.g., water aspirator) is adequate for filtering.
2. Suspend the washed resin in anion-exchange buffer to 75% settled gel/25% buffer by volume, per manufacturer’s recommendations. Degas in a filter flask and pour into a 5 × 50–cm chromatography column fitted with a filling reservoir. After settling, the height of the resin should be ∼20 to 25 cm (390 to 490 ml packed resin). For details on packing columns, see UNIT 8.4. Because the solubility of gases decreases with increases in temperature, it is usual practice to pack the column at room temperature and then run it in a cold room or cold box.
3. Elute column with anion-exchange buffer at 100 to 150 ml/hr using a peristaltic pump. Make sure there is no compression of column contents. Monitor the absorbance of the effluent at 260 or 280 nm with a UV detector. Collect ∼15-ml fractions in 16 × 50–mm culture tubes using a fraction collector. Check that the pH and conductivity of the column effluent are the same as the for anion-exchange buffer applied to the column (this indicates that the column matrix is correctly equilibrated). The bed height should not change significantly once the column is packed. Compression indicates that the pressure applied to the column is too high (see manufacturer’s recommendations for maximum flow rates).
Break cells with a French press 4. Clean bench areas that may come in direct contact with cells with 0.26% sodium hypochlorite/70% ethanol.
Purification of Recombinant Proteins
6.2.3 Current Protocols in Protein Science
5. Assemble the French pressure cell and and chill to ∼4°C either by incubation in ice or by refrigeration. Install the cell (first dried with paper towels if necessary) in the Aminco laboratory press. It is important to cool the equipment because pressurizing will generate heat. The 20K rapid-fill French pressure cell (1-in.-diameter piston) has a capacity of 40 ml and can be continuously filled while installed on the press. Before using the pressure cell, replace the nylon ball at the end of the flow valve assembly or, at the very least, check it for distortion. For small-scale work, a miniature French pressure cell (3/8-in.-diameter piston) with a 3.7-ml capacity is available.
6. Suspend thawed E. coli cells (∼50 g wet weight) with 150 ml lysis buffer using a Waring blender. Place the suspension in a stainless steel beaker and homogenize with the Polytron tissue-grinder homogenizer until clumps are no longer detected. IMPORTANT NOTE: Wear disposable gloves and safety glasses while working with E. coli. The high-pressure homogenization may generate aerosols. The E. coli cells are stored frozen at −80°C as a flattened paste in heat-sealable plastic bags (UNIT 5.3). The cells are thawed at room temperature. Complete suspension of the cells with the blender is important, as any visible clumps of bacteria will block the French pressure cell. A clogged cell may have to be disassembled to clear the blockage.
7. Lyse the cells with two passes through the French press operated at 16,000 to 18,000 lb/in2 (with the high-ratio setting, pressure gauge readings between 1011 and 1135). Chill the cell suspension to 4°C after each pass through the pressure cell by incubation on ice. When filling the pressure cell, avoid drawing air into the cylinder to prevent foaming. If a French press is not available, the cells can be broken by including 200 ìg/ml lysozyme (Worthington) and 0.05% (w/v) sodium deoxycholate (Calbiochem) in the lysis buffer and incubating cells ∼20 min at 20° to 25°C with intermittent homogenization using the tissue grinder (Burgess and Jendrisak, 1975). Cell breakage by lysozyme treatment and sonication is described in Basic Protocol 3 of UNIT 6.5.
8. Place the suspension (contained in a steel beaker) on an ice bath and, using an ultrasonic homogenizer, sonicate 5 min at full power with 50% duty cycle (on for 0.5 sec then off for 0.5 sec). While sonicating, stir the suspension using a magnetic stirrer. IMPORTANT NOTE: Wear sound-protection earmuffs to protect ears from ultrasonic noise. Because sonication will generate some aerosol, use the sonicator in a microbiological hood if possible. High viscosity reduces the rate of sedimentation of the various contaminating cellular material and thus longer (sometimes much longer) centrifugation times are required. Sonication reduces the viscosity of the suspension prior to centrifugation by shearing the released DNA and RNA. The viscosity can also be reduced by digesting the lysate 15 to 30 min at 4° to 10°C with bovine pancreas DNase I (25 to 50 ìg/ml) and RNase A (50 ìg/ml). If the nucleases are used, EDTA in the lysis buffer should be replaced by 5 mM MgCl2 as DNase requires Mg2+.
Clarify the lysate 9. Transfer sample from the beaker to centrifuge bottles and centrifuge the cell lysate 40 min at 22,000 × g (e.g., in a Beckman J2-21M preparative centrifuge at 12,000 rpm using JA-14 rotor or at 13,500 rpm using JA-20 rotor), 4°C. Decant the supernatants, pool, and recentrifuge 90 min at ∼100,000 × g (30,000 rpm in Beckman Optima XL-90 ultracentrifuge using Ti45 rotor), 4°C. Preparation of Soluble E. coli Proteins
Low-speed centrifugation removes unbroken cells and large cellular debris. Highspeed centrifugation removes smaller particles such as ribosomes and membrane
6.2.4 Current Protocols in Protein Science
vesicles; the Beckman 70Ti rotor (capacity 8 × 39 ml) can be used in the ultracentrifuge for smaller-scale work. Clarification of the lysate can also be carried out by salt fractionation (see Background Information, section on determining solubility, for further details). Pellets are usually discarded immediately; but see Critical Parameters and Troubleshooting, section on protein purification for further comments.
10. Dilute the supernatant from step 6 (∼160 ml) 1:2 (three-fold) with anion-exchange buffer and adjust to pH 8.5 (if necessary) with 2 N NaOH. Using a conductivity meter, measure the conductivity of the diluted supernatant. If it is higher than 5.0 to 5.3 mS/cm, reduce by dilution with water. The conductivity of the protein solution is carefully adjusted to ensure that the proteins (in this case contaminants) are bound to the matrix. Too high an ionic strength will reduce or prevent binding. The DEAE Sepharose column is normally prepared (steps 1 to 3) before lysis of the cells is initiated. If there are any delays in applying sample to column, store the clarified lysate at 0° to 4°C (for example, in a covered beaker or flask embedded in an ice bucket). The same applies to the other chromatographic stages.
Chromatograph cleared lysate on anion-exchange resin 11. Apply the clarified lysate (480 to 500 ml) to the DEAE Sepharose column (step 3) at 150 ml/hr and elute the column with anion-exchange buffer. Continue the elution, collecting 15-ml fractions, until the effluent absorbance is close to the baseline value. In solutions above ∼pH 7.0, IL-1β (pI 6.8) is negatively charged. Therefore, the protein should, in principle, bind to an anion exchanger (positively charged matrix) buffered at pH 8.5. In fact, under the conditions described, IL-1β is only weakly bound to the matrix, which allows for partial resolution from proteins that do not bind to the matrix. Much of the unbound protein is of high molecular weight (or highly aggregated) and is partially separated from IL-1β by the gel filtration effect of the matrix. IL-1β is therefore usually located in the latter two-thirds (∼500 ml) of the column flowthrough volume.
12. Assay every second or third column fraction by SDS-PAGE (UNIT 10.1). See Figure 6.2.1A for an example of results from SDS-PAGE. For rapid analysis, use precast gels or the Hoefer Pharmacia Phast system. Alternatively, if speed of purification is important, the entire flowthrough can be used, eliminating the need to analyze separate fractions. Most protein contaminants bound to the column can be removed by step elution with 1 M NaCl in column buffer. After use, the resin should be unpacked from the column and washed on a sintered-glass funnel with 1 liter of 2 M NaCl/0.5% (w/v) Triton X-100, followed by 10 liters water. If the resin is to be stored, suspend it in 5% ethanol or 5 mM sodium azide and store at 4°C. In order to avoid potential cross-contamination, dedicate the used resin for repeat purifications of IL-1β only.
Fractionate sample with ammonium sulfate 13. Pool IL-1β-containing fractions from the DEAE Sepharose column, record the volume (∼500 ml), and transfer the solution to a 1-liter beaker (preferably stainless steel). Add 30.2 g (NH4)2SO4 per 100 ml solution at 0°C (51.3% saturation or 2 M final concentration): add powdered (NH4)2SO4 slowly over 30 min, mixing gently with a magnetic stirrer, then allow a further 30 min of mixing. Methods for calculating the percent saturation of ammonium sulfate solutions, which specifically refer to 0°C, have been described by Wood (1976). When using solid ammonium sulfate, calculations must include volume increases on addition of the solid to fixed volumes (as carried out above). Purification of Recombinant Proteins
6.2.5 Current Protocols in Protein Science
A
B
a
1.0
b
c
d
e
g
f
h
1.0 .75 .50
0.75
V0
Vi
200 300 400 Elution volume (ml)
500
A280
.25
0 100
0.50
V0
Vi
0.25
P 0 100
200
300
400
500
Elution volume (ml)
Figure 6.2.1 Purification of IL-1β. (A) SDS-PAGE analysis of samples at various stages. Analysis was conducted on a gel of dimensions 12 cm × 16 cm × 1.5 mm. Lane a, purified protein (100 µg loaded); lane b, purified protein (10 µg loaded); lane d, CM Sepharose pool (80%); lane e, DEAE Sepharose pool after ammonium sulfate fractionation (56%); lane f, high-speed supernatant (starting material for DEAE Sepharose column; 13.5%); lane g, cell lysate (12.0%). The percentages refer to specific IL-1β contents of the fractions determined by densitomeric scanning of the Coomassie blue–stained gel lanes. Lanes c and h contain the following protein standards (low-range standards supplied by Bio-Rad) in order of increasing migration distance: phosphorylase b (97.4 kDa), bovine serum albumin (66.2 kDa), hen egg white ovalbumin (45 kDa), bovine carbonic anhydrase (31 kDa), soybean trypsin inhibitor (21.5 kDa), and hen white lysozyme (14.4 kDa). (B) Analysis of results from gel filtration on Ultrogel AcA54. The excluded volume (V0) and the fully included volume (Vi) are indicated. Inset, analytical rechromatography of the protein from the pooled fractions (indicated P in larger chromatogram).
Preparation of Soluble E. coli Proteins
6.2.6 Current Protocols in Protein Science
14. Centrifuge the slightly cloudy solution 30 min at 22,000 × g (12,000 rpm in JA-14), 4°C. Decant the supernatant into a beaker and add an additional 17 g (NH4)2SO4 per 100 ml solution (77% saturation or 3 M final concentration). Equilibrate with stirring and centrifuge 30 min at 22,000 × g, 4°C. For the addition of (NH4)SO4 follow the same method as described in step 13.
15. Decant the supernatant and drain the pellets by inverting the tubes on a paper towel. Save the pellets. Dialyze the fractionated sample 16. Suspend the pellets in ∼300 ml cation-exchange buffer and dialyze, using Spectra/Por 1 dialysis tubing, against 5 liters cation-exchange buffer. Change the dialysis buffer at least once. The dialysis step is conveniently performed overnight; the CM Sepharose column used in step 18 can be prepared during this period. The dialysis tubing is prepared by heating 30 to 60 min at 90° to 95°C in 5 mM EDTA. The tubing is then washed well with water and stored in 10% ethanol at 4°C prior to use. A suitable length of tubing is filled to about one-half to three-quarters capacity with solution (to allow for expansion) and sealed with two knots at each end. Use gloves when handling the tubing, check for leaks before use, and make sure the magnetic stir-bar does not rub against the tubing. See APPENDIX 3B for further information concerning dialysis.
17. After dialysis, remove the slightly cloudy solution from the tubing and centrifuge 30 min at 22,000 × g (12,000 rpm in JA-14), 4°C. Save the supernatant. Chromatograph dialyzed sample on cation-exchange resin 18. Prepare ∼200 to 225 ml CM Sepharose CL-4B resin by washing on a sintered-glass funnel first with water, then with cation-exchange buffer (pH 5.7; wash as in step 1 except using different buffer). Pack the degassed resin into a 5 × 50–cm column as in step 2. The packed column will have a bed height of ∼11 to 12 cm. The comments made in the annotation to step 3 also apply here.
19. Elute the column using cation-exchange buffer at 100 to 150 ml/hr with a peristaltic pump. Monitor the column effluent at 280 or 260 nm using a suitable UV detector. Check that the pH and conductivity of the column effluent are the same as for the buffer applied to the column. 20. Check the pH and conductivity of the dialysate supernatant from step 17 and, if necessary, dilute with water so that the conductivity is in the range 1.0 to 1.2 mS/cm (at 4° to 6°C). Apply the clear solution to the CM Sepharose column at a flow rate of 150 ml/hr. When the UV absorbance of the column effluent approaches baseline, proceed to the next step. 21. Prepare a 0 to 250 mM NaCl gradient in cation-exchange buffer by adding 500 ml buffer to the inner chamber of the gradient maker and 500 ml buffer/250 mM NaCl to the outer chamber. Apply the gradient to the column at 150 ml/hr. Collect 15-ml fractions. The total volume of the gradient is 1 liter (∼4.5 column volumes). At pH 5.7, IL-1β (pI 6.8) is positively charged and binds to the negatively charged cation-exchange resin. IL-1β is eluted from the column with ∼100 mM NaCl, and it will be located in the major absorbance peak (see Wingfield et al., 1986, for figure of typical elution profile).
Purification of Recombinant Proteins
6.2.7 Current Protocols in Protein Science
22. Monitor the progress of the gradient using an in-line conductivity meter positioned after the absorbance flow cell. Assay column fractions for IL-1β by SDS-PAGE (UNIT 10.1) and pool fractions containing IL-1β. Deciding what fractions to pool is dictated by the fact that the remaining purification stage is gel filtration, a method that will not remove contaminants with sizes close to that of IL-1β (17.4 kDa). As IL-1β is a well-expressed protein (>5% total protein), one can afford to be conservative and pool for purity rather than yield. The used CM-Sepharose matrix can be cleaned up and stored as described in the annotation to step 12. CM Sepharose Fast Flow or SP Sepharose FF (a strong cation exchanger; both resins are also from Pharmacia Biotech) can be used instead of CM Sepharose CL-4B. Similar results are obtained with either matrix, with the advantage of faster flow rates.
Concentrate the proteins 23a. To concentrate proteins by ultrafiltration: Adjust the CM Sepharose pool (150 to 250 ml) to pH 7.5 with Tris base. Assemble a 200- or 400-ml stirred ultrafiltration cell containing a washed ultrafilter membrane, either YM3 (3-kDa cutoff) or PM10 (10-kDa cutoff). Pressurize the cell with nitrogen according to manufacturer’s recommendations. Collect the effluent from the cell into a measuring cylinder and occasionally record the absorbance at 280 nm to check that the membrane is not leaking. When the volume remaining in the cell is ∼15 ml, depressurize the cell and carefully remove the solution using a Pasteur pipet with a small length of polyethylene tubing attached to the end so as not to scratch the membrane. Use ∼3 ml gel-filtration buffer to wash the membrane, adding washings to the main concentrate. Exposure to solutions below pH 4.5 and above pH 10.5 to 11.0 often causes protein denaturation. Hence, when adjusting the pH of protein solutions, there is less chance of overshooting the required pH by using concentrated buffer components instead of pure acid or base. For example, Tris base is used rather than dilute NaOH. Select the ultrafiltration membrane pore size based on the size of the protein. The membrane can be reused; store in 5% ethanol at 4°C. It is good practice to reserve a membrane for use with a particular protein (compare comment on resin usage) and to wash the concentration cell carefully after use.
23b. To concentrate proteins by salt precipitation: Adjust the CM Sepharose pool to pH 7.5 with Tris base and slowly add 53.9 g (NH4)2SO4 per 100 ml solution at 0°C (82.2% saturation or 3.2 M final concentration). Follow the basic salt precipitation method (steps 13 to 15) and suspend the pellets in 18 to 20 ml gel-filtration buffer. Conduct gel-filtration chromatography 24. Prepare ∼480 ml Ultrogel AcA54 resin by washing on a sintered-glass funnel with water, then gel-filtration buffer (as in step 1); suspend washed resin in gel-filtration buffer to 75% settled gel/25% buffer by volume and degas (step 2). Pour a slurry of degassed resin into a 2.5 × 100–cm chromatography column fitted with a filling reservoir. Pack the column at ∼35 ml/hr. UNITS 6.3 & 8.3 should be consulted for further details on the preparation and elution of gel-filtration columns. The column should be prepared in advance so that the protein samples can be applied as soon as the concentration step is completed.
Preparation of Soluble E. coli Proteins
The resin should be free-flowing yet concentrated enough to produce a packed bed with one pouring. A freshly packed gel-filtration column can be checked for packing irregularities by prerunning the column with a few colored markers. Blue dextran will be excluded from the gel matrix and will elute at the void volume (V0, which equals 30% to 35% of the total column volume); cytochrome c (red; 12.4 kDa) will elute close
6.2.8 Current Protocols in Protein Science
to the expected position of IL-1β; and potassium dichromate (yellow) will be fully included in the gel matrix and will elute at the included volume (Vi, ∼480 ml). Superdex 75 gel-filtration matrix (Pharmacia Biotech) can be substituted for Ultrogel AcA54. The former allows higher flow rates and thus is more compatible with the FPLC and BioPilot systems. Both Superdex and the cation-exchange Fast Flow resins mentioned above can be purchased in various prepacked columns that are useful for method development.
25. Filter the concentrated protein using a Millex-GV 0.22-µm filter unit attached to a 10- or 20-ml syringe and apply to the gel-filtration column. The sample can be applied either directly to the top of the column using a Pasteur pipet (care is required to prevent breaking the tip and contaminating the column) or via a three-way valve and syringe without removing the top flow adapter. The volume of sample applied to the column (20 ml) represents ∼4% of the total column volume. For columns with different dimensions, apply the same proportionate volume of sample. The column should be ≥60 cm long. For analytical separations, the sample volume should not exceed 2.5% of the column volume.
26. Elute the column at 35 ml/hr with gel-filtration buffer and collect 10-ml fractions. The major eluting peak contains the IL-1β; monitor the fractions by SDS-PAGE (UNIT 10.1) and pool fractions that contain pure protein. Save side fractions that contain small amounts of contaminants; this material can be rechromatographed after concentration as described in step 23a or 23b. See Figure 6.2.1A for an example of results from SDS-PAGE. Once the sample has been run into the column, it can be eluted with any buffer or even with a suitable column storage solvent such as 5 mM sodium azide in water.
Concentrate and store purified protein 27. Concentrate the purified protein, if required, by ultrafiltration (see step 23a). Determine the protein concentration of purified IL-1β by measuring absorbance at 280 nm. A molar absorbance coefficient (ε) of 10.61 mM−1 cm−1 is used. This corresponds to an absorbance of 0.63 for a 1 mg/ml solution using a 1-cm-path length cell.
28. For short-term storage (≤12 months), filter the protein with a Millex-GV 0.22-µm filter unit, divide the solution into sterile plastic vials, and freeze aliquots rapidly with dry ice/ethanol. Store at −80°C. For long-term storage of IL-1β (>12 months), lyophilize the protein. Dialyze the sample using a volatile buffer such as 50 mM ammonium bicarbonate or a nonvolatile buffer such as lyophilization buffer. To circumvent the dialysis step, the phosphate-based lyophilization buffer can be used for gel filtration instead of the Tris⋅Cl gel-filtration buffer.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX. Buffer pH and conductivities are for solutions at 4° to 6°C. Units of conductance are given in siemens (S = Ω−1).
Anion-exchange buffer (50 mM Tris⋅Cl, pH 8.5) Dilute 1 M Tris⋅Cl, pH 8.0 20-fold with water and adjust to pH 8.5 with NaOH. Make immediately before use. Conductivity of the solution is 1.57 mS/cm.
Purification of Recombinant Proteins
6.2.9 Current Protocols in Protein Science
Cation-exchange buffer, 10× (15 mM sodium phosphate, pH 5.7/1 mM sodium azide) 19.4 g NaH2PO4⋅H2O 1.4 g Na2HPO4⋅2H2O 0.65 g sodium azide H2O to 1 liter Store up to 1 month at 4°C Dilute 10-fold immediately prior to use Conductivity of the solution is 0.9 mS/cm. Sodium azide is an antibacterial agent.
Cation-exchange buffer (1×)/250 mM NaCl 14.61g NaCl 100 ml 10× cation-exchange buffer (see recipe) H2O to 1 liter Gel-filtration buffer (100 mM Tris⋅Cl, pH 7.5/1 mM sodium azide) Dilute 1 M Tris⋅Cl, pH 8.0 10-fold with water and adjust to pH 7.5 with HCl. Add sodium azide from a 1 M stock solution (65 g/liter). Make immediately before use. Lyophilization buffer (25 mM sodium phosphate, pH 7.5/0.5 mM sodium azide) 2.2 g NaH2PO4⋅H2O 11.9 g Na2HPO4⋅2H2O 1.3 g sodium azide H2O to 4 liters Make fresh and use immediately Usually, 2 liters is required for dialysis, with one buffer change (4 liters total). Gel filtration requires smaller volumes.
Lysis buffer 100 mM Tris⋅Cl, pH 8.0 2 mM EDTA, pH 8.0 5 mM benzamidine⋅HCl (780 mg/liter) Make immediately prior to use; alternatively, make ahead of time and store up to several days at 4°C. Conductivity of the solution is 1.57 mS/cm. It should be noted that a 1°C decrease in temperature increases the pH of the Tris buffer by ∼0.03 pH units. Both Tris⋅Cl and EDTA stock solutions are commercially available (e.g., Life Technologies). Benzamidine⋅HCl is a water-soluble serine protease inhibitor. An alternative is 50 ìM 4-(2-aminoethyl)benzenesulfonyl fluoride hydrochloride (AEBSF; Perfabloc SC, Boehringer Mannheim), a water-soluble inhibitor with the same spectrum of activity as phenylmethylsulfonyl fluoride (PMSF).
COMMENTARY Background Information
Preparation of Soluble E. coli Proteins
Expression of soluble proteins The basic stages involved in preparing soluble proteins from E. coli are achieving protein expression, harvesting and breaking cells (or collecting culture medium, in the case of secreted proteins), purifying proteins, and characterizing the purified protein. Choices that must be made at each stage are briefly reviewed.
Optimizing the expression system The basic requirements for protein expression in E. coli are discussed in UNIT 5.1. Experience has shown that it is usually possible to express any given protein in E. coli, but whether the protein will be expressed in a soluble and unmodified state is unpredictable. Tinkering with the various elements of the expression plasmid, changing the E. coli host strains, and altering the fermentation conditions are carried out to optimize the expression
6.2.10 Current Protocols in Protein Science
of unmodified soluble protein (UNIT 5.1). Use of secretion vectors and fusion protein constructs to prevent aggregation has been discussed elsewhere (UNITS 5.1 & 6.1). Even if efforts to express soluble protein fail and inclusion bodies are produced, there is often a good chance that the protein can be extracted and folded into native protein (see UNITS 6.4 & 6.5). There are situations where the intrinsic properties of a protein can preclude its expression in a soluble and stable form. Examples include the following: (1) individual subunits of a heterodimer, which may be unstable and may degrade or aggregate when expressed separately (coexpression of the subunits may solve the problem; Nash et al., 1987); (2) proteins derived from activated precursors (UNIT 6.4; Hlodan and Hartl, 1994); and (3) proteins in which the native (authentic) sequences have been modified by site-directed mutagenesis (Chrunyk et al., 1993). It is worth noting that the solubility of proteins that are otherwise insoluble or poorly soluble can, conversely, sometimes be improved by site-directed mutagenesis (Dyda et al., 1994). Once conditions have been found that result in the expression of soluble protein, the next hurdle is to recover the protein in an unmodified form. Avoiding proteolysis and chemical modification during the isolation process is crucial. Breaking cells Methods for breaking cells or selectively extracting proteins are reviewed in UNIT 6.1. In the protocol for purifying human IL-1β, cells are disrupted with a French press. This method is the most efficient way to break E. coli cells and is the method of choice. The equipment, however, requires dedicated laboratory space and may be expensive for some laboratories. Cell breakage with lysozyme (UNIT 6.5) and/or sonication is also frequently used; such methods are especially suited to small-scale work (reviewed by Hopkins, 1991). It is important to consider whether the cell breakage method influences the final solubility and stability of the recombinant protein in the cell lysates. A soluble recombinant protein constituting ∼12% of the total bacterial protein will be present in the cytoplasmic space at a concentration of at least 30 mg/ml, assuming a total protein and RNA concentration in the cytoplasm of ∼340 mg/ml in nontransformed cells (Zimmerman and Trach, 1991) and a 2.7:1 weight ratio of protein to RNA (Neidhart, 1987). Cell breakage can be expected to in-
crease the solubility of the recombinant protein by a dilution effect (discussed by Zimmerman and Trach, 1991). Cell lysis procedures, however, especially when mechanical shear is used (e.g., French press and sonication), can produce local heating that may have a denaturing effect and lead to aggregation. Mechanical shearing will also be expected to decrease the size of nucleic acids and release polyanionic lipopolysaccharides; both classes of coumpounds can bind nonspecifically to proteins, especially basic proteins, thereby reducing their solubility. Including detergents in the lysozyme treatment may also release potent membrane-bound proteases. Despite these potential pitfalls, the recovery of soluble protein is usually not dramatically influenced by the cell breakage method, but there is always the potential that it might be. Protein recovery as a function of cell breakage methodology is rarely systematically studied or reported. Johnson and Hecht (1994) reported that freezing and thawing of cells selectively releases recombinant proteins located in the cytoplasm. The freeze-thaw procedure, in principle, should be the best method for releasing soluble proteins in an unmodified state, but its general applicability remains to be seen. Determining solubility The solubility of recombinant protein in the cell lysate or extract is usually established by differential centrifugation (UNITS 6.1 & 5.3). Alternatively, the extract can be clarified by filtering and applied to a small gel-filtration column. An HPLC or FPLC gel-filtration separation in conjunction with a rapid screening method for assaying column fractions (e.g., SDS-PAGE using PhastSystem from Hoefer Pharmacia) can produce a determination of both solubility and size distribution in ∼2 hr or less. Purifying protein There are many approaches to purifying proteins, such as expressing fusion proteins that contain engineered affinity handles or tags (see UNITS 5.1 & 6.1 for further details). Most soluble proteins that are well behaved—i.e., do not contain excessive charge heterogeneity—are non-self-associating and can often be purified by two stages of ion-exchange chromatography and one or more polishing steps, one of which should be gel filtration. The basic approach is outlined in Figure 6.1.3. The purification of human IL-1β is a typical example. Specific accounts of the various chromatographic meth-
Purification of Recombinant Proteins
6.2.11 Current Protocols in Protein Science
ods are found in Chapters 8 and 9; Scopes (1993) particularly emphasizes first principles. Characterizing protein Once a protein has been purified, it is usually characterized (Chapter 7) in order to establish chemical homogeneity (primary sequence) and physical homogeneity (size and conformation). Not only is this information required for a fundamental description of the protein, but it will also give insights on how best to rationalize the purification process. Moreover, the detection of certain heterogeneities may give clues on how to avoid their occurrence. For example, chemical modifications such as deamidation may be prevented by changing one or more purification steps that expose the protein to extremes of pH. Other modifications that arise from construction of the expression plasmid itself, once recognized, can often be fixed.
Preparation of Soluble E. coli Proteins
Characteristics of recombinant and authentic interleukin 1β The cytokine IL-1β and the closely related IL-1α are synthesized mainly by monocytes and mononuclear cells as 31-kDa cytoplasmic precursor proteins (reviewed by Dinarello, 1989). A unique cytoplasmic cysteine protease (IL-1β-converting enzyme, or ICE) cleaves the IL-1β precursor in half, generating the mature form of the protein comprising the C-terminal 153 residues. The biological activities of IL-β (and IL-1α) are initiated by interaction with either type I or type II cellular receptors (McMahan et al., 1991). Both receptors have three immunoglobulin-like extracellular ligand-binding domains and single membranespanning segments. Another form of IL-1, the interleukin 1 receptor antagonist (IL-1ra), is a natural competitive antagonist of IL-1β. Authentic IL-1β been isolated from human monocyte cell cultures by HPLC using anionexchange and gel-filtration (or reversed-phase) matrices. A 3000-fold purification was required to obtain 16 µg protein from 5 liters of cell culture extract (Gery and Schmidt, 1985). The authentic protein was established to be a nonglycosylated monomer (∼18 kDa), with a pI of 6.8 and containing a single N-terminal sequence (AlaProValArg) as predicted by the DNA coding sequence. The recombinant protein produced in E. coli has the same properties as the natural product except that N-terminal heterogeneity is often observed. The recombinant protein usually contains a mixture of forms with Met (20%), Ala (67%), or Pro (13%) as the N terminus. The
Met form is derived by incomplete processing of the initiating Met, and the Pro form is due to cleavage of the authentic N-terminal Ala by E. coli protease. Despite the slight differences among the recombinant variants, partial or complete separations have been achieved (Wingfield et al., 1987; Yem et al., 1988). Both the crystal structure of IL-1β (Priestle et al., 1988) and its nuclear magnetic resonance (NMR) structure in solution (Clore et al., 1991) have been determined using material purified by the protocol described here. For some NMR studies it was necessary to produce material with a single N terminus, and this was fortuitously achieved by replacing the N-terminal Ala residue with a Cys residue. Purified protein from cells expressing the mutant (N-terminal CysProValArg) contain only N-terminal Pro. It should be noted that deletion of the first four residues of IL-1β has no effect on activity; however, the presence of the unprocessed Met reduces receptor binding 10-fold. Interleukin 1β does not contain a disulfide— the cysteines are either buried (Cys-71) or partially buried (Cys-8)—so there is no need to include reductants in column buffers. As mentioned earlier, the authentic protein is nonglycosylated even though there is a potential Nglycosylation site (-Asn7-Cys8-Thr9-) near the N terminus. Interestingly, when IL-1β is expressed in yeast, equal amounts of N-glycosylated (21 kDa) and nonglycosylated protein (17 kDa) are produced (Livi et al., 1991). Other approaches to purifying recombinant interleukin 1β Because of the biomedical importance of IL-1β, several reports have described expression and purification of the protein. Some examples are discussed. Kronheim et al. (1986) acidified E. coli cell lysates to pH 4 and thus precipitated ∼70% of the total protein while most of the IL-1β remained soluble. IL-1β was further purified by cation-exchange, anion-exchange, and dyematrix chromatographies (UNITS 8.2 & 9.2); ∼200 mg protein was recovered from 2.5 liters of culture. The noteworthy feature of this method is the acid precipitation stage. Extracts of E. coli contain predominately negatively charged (anionic) proteins: 60% have pI values between 5.0 and 6.0 and 80% between 4.5 and 6.7 (Sherwood, 1992). Adjustments of cell extracts to pH 5.0 and below thus result in isoelectric precipitation of substantial amounts of E. coli protein. This is a useful purification step as long
6.2.12 Current Protocols in Protein Science
as the recombinant protein remains soluble (or does not coprecipitate) under the conditions. In another approach, Meyers et al. (1987) carried out ammonium sulfate fractionation directly on the E. coli extract. The IL-1β protein was recovered in the fraction corresponding to 50% to 80% saturation. After desalting, the protein was further purified by ion-exchange (cation and anion) and gel-filtration chromatographies; ∼400 mg protein was recovered from 250 g wet weight cells. The high-speed centrifugation step used to remove particulate material (see Basic Protocol, step 6) can often be replaced by salt fractionation. For IL-1β, advantage can be taken of the fact that a relatively high (NH4)2SO4 concentration is required to precipitate the protein. The disadvantage of using this approach early in the purification is that the protein must be desalted before being applied to an ion-exchange column. Finally, it is of interest to note that IL-1β can be released from E. coli cells by osmotic shock treatment (Joseph-Liauzun et al., 1990). It is well known that certain E. coli cytoplasmic proteins, including thioredoxin, can be released by osmotic shock. Scale of procedure The Basic Protocol described starts with 50 g cell paste and yields ∼400 mg pure IL-1β. For smaller (1 to 5 g) or larger (100 g) cell quantities, the method can be adapted by simply reducing or increasing, respectively, the crosssectional area of the chromatography columns. For example, for 5 g cells, the diameters of the DEAE and CM Sepharose CL-4B columns are reduced to 2.5 cm, with column lengths of 20 and 10 cm, respectively. This results in 1:4 reductions in the resin volumes compared to those for the protocol utilizing 50 g cells. The original gel-filtration column can be used (2.5 × 100 cm); however, the volume of ∼1:4 sample applied should be reduced to 8 to 10 ml. Alternately, a column of about the same length (60 to 100 cm) but smaller diameter (e.g., 1.25 cm) should be used. For discussion on scaling up column chromatography procedures, see UNIT 8.3 (Strategic Planning and Scopes, 1993).
Critical Parameters and Troubleshooting High-level protein expression with good cell yield (UNITS 5.1-5.3) is required to obtain reasonable amounts of pure protein. This is obviously true no matter how efficient (or inefficient) the purification protocol. Once the host/vector sys-
tem and the fermentation conditions have been optimized and, equally important, standardized, the level and solubility of recombinant protein in the starting material should be consistent. However, it is sensible to check both the expression level and solubility before starting each protein purification. The expression level can usually be monitored by SDS-PAGE; UNIT 5.2 describes how to prepare samples for analysis by SDS-PAGE, and UNIT 10.1 gives electrophoresis procedures. Solubility can be easily monitored by breaking a small amount of cells (≤1.0 g) by sonication (UNIT 5.3) or by treating cells with lysozyme (UNIT 6.5) and microcentrifuging the resulting cell lysate to judge whether the protein is highly aggregated. There are many reasons why protein expression may not be as high as anticipated and, worse, why the expressed protein may switch from being soluble to insoluble. For example, faulty pH control during the fermentation may cause insolubility. Whatever the situation, one should be in a position to rationally decide whether to continue with the purification or repeat the initial fermentation. If the protein expression level is normal (>5%) and the protein has accumulated in a soluble state, implementing the Basic Protocol, which uses only standard purification methods, should be trouble-free. Common mistakes involve buffer preparation and other trivial errors such as pooling the wrong column fractions. As discussed later, do not discard column fractions until they have been completely analyzed. A classic purification table (see for example, Table III.1 in Dixon and Webb, 1979), has not been presented, as the biological activities of IL-1β could not be measured reliably in crude E. coli cell extracts by assays available to the author when the method was developed (Wingfield et al., 1986). As is often the case with proteins requiring complex biological assays, protein size analysis by SDS-PAGE is used to follow purification. During the initial stages, it is of course necessary to confirm the identity of the particular band one is following, and this can be accomplished by eluting or blotting the protein from SDS-polyacrylamide gels and performing N-terminal amino acid sequence analysis. Immunoblotting with specific antibody can be used to monitor protein purification when the recombinant protein is in low abundance (e.g., for secreted proteins in the cell medium). Finally, it should be noted that several manufacturers (e.g., R&D Systems and Genzyme) now supply assay kits for many cytokines, including IL-1β.
Purification of Recombinant Proteins
6.2.13 Current Protocols in Protein Science
Cell breakage Efficient cell breakage should be troublefree as long as the French press is operated in accordance with the manufacturer’s instructions. It takes a little practice to operate the flow valve. The aim is to generate as high a flow rate as possible while maintaining a pressure gauge reading of ∼1000. If the flow rate is too fast, the pressure reading will drop and unbroken cells will pass into the flow stream. Toward the end of the run, the flow rate should be reduced, as it becomes difficult to control the pressure. After use the French pressure cell should be cleaned and dried, and the flow valve ball should be replaced. Store the cell at 4°C. Protein purification In general, troubleshooting a purification method will be much easier if fractions are not discarded until the appropriate monitoring of the purification steps is complete. When using an established method, fractions from a chromatographic run are frequently pooled on the basis of absorbance only and the remainder quickly discarded. Ammonium sulfate supernatants or pellets are often discarded on the basis of previous fractionation behavior or pilot-scale work. As stated, do not discard any fractions until they have been checked, usually by SDS-PAGE. If there has been a problem, it can usually be easily sorted out if all the fractions from the various stages are still available. If in doubt about conditions for storing fractions, freeze selected fractions at as low a temperature as possible (ideally −80°C) and discard when appropriate. When freezing material, it is worthwhile to take the extra effort to dispense small samples (<1 ml) into microcentrifuge tubes amenable for rapid analysis, if necessary, to avoid having to thaw large volumes merely to take 10 µl for SDS-PAGE. For reproducible results, careful recording of the pH and conductivity of starting buffers, protein solutions, and column eluents is necessary. For example, if the protein does not bind to the cation exchanger (step 19), the ionic strength of the sample or column buffer may be too high. Alternatively, the buffer pH may be too high. Errors of this kind are avoided by simply measuring the conductivity and pH of all solutions at each stage. Other precautions and critical steps relating to chromatography procedures are mentioned in Chapters 8 and 9. Preparation of Soluble E. coli Proteins
Anticipated Results
Purification results in recovery of ∼8 mg purified protein per gram wet weight cells. That
result is based on a 10% expression level; IL-1β expression levels of 10% to 15% are routinely obtained with the expression system previously described (Wingfield et al., 1986). With a 1.5liter fermenter, at least 50 g wet weight cells/liter can be obtained using a standard complex medium (UNIT 5.3). Thus, ∼400 mg protein is obtained from 1 liter of culture, and optimal conditions can lead to nearly twice that amount. The yield corresponds to an overall recovery of ∼60% of the IL-1β originally in the cells. Figure 6.2.1 illustrates the protein content at various purification stages. The starting material for the final stage of gel filtration (Fig. 6.2.1A, lane d) is mainly contaminated with E. coli proteins >44 kDa in size, and these are easily separated from the IL-1β protein (17.4 kDa) by gel filtration. Analytical rechromatography of the protein from the pooled fractions (indicated P in Fig. 6.2.1B) reveals a single symmetrical peak indicative of purity and physical homogeneity (Fig. 6.2.1B inset). A typical cation-exchange chromatogram for the purification appears elsewhere (Wingfield et al., 1986).
Time Considerations Because IL-1β appears to be stable against proteolytic degradation and other chemical modications during purification, the speed of purification was not critical in this case. The low-pressure chromatographic method described in the Basic Protocol requires ∼4 days, which can be shortened to ∼3 days using the Pharmacia Biotech BioPilot or FPLC systems in conjunction with matrices that allow faster flow rates. The times required for purification whether using low-pressure chromatography (as described in the Basic Protocol) or mediumpressure chromatography in the FPLC or BioPilot systems are summarized in Table 6.2.1.
Literature Cited Burgess, R.R. and Jendrisak, J.J. 1975. A procedure for the rapid, large-scale purification of E. coli DNA-dependent RNA polymerase involving polymin P precipitation and DNA-cellulose chromatography. J. Biol. Chem. 14:4634-4638. Chrunyk, B.A., Evans, J., Lillquist, J., Young, P., and Wetzel, R. 1993. Inclusion body formation and protein stability in sequence variants of Interleukin-1β. J. Biol. Chem. 268:18053-18061. Clore, G.M., Wingfield, P.T., and Gronenborn, A.M. 1991. High-resolution structure of interleukin 1β in solution by three- and four-dimensional nuclear magnetic resonance spectroscopy. Biochemistry 30:2315-2323.
6.2.14 Current Protocols in Protein Science
Dinarello, C.A. 1989. Interleukin-1 and its biologically related cytokines. Adv. Immunol. 44:153205. Dixon, M. and Webb, E. 1979. Enzyme isolation. In Enzymes (3rd ed.) pp. 23-46. Academic Press, New York. Dyda, F., Hickman, A.B., Jenkins, T.M., Engelman, A., Craigie, R., and Davies, D.R. 1994. Crystal structure of the catalytic domain of HIV-1 integrase: Similarity to other polynucleotidyltransferases. Science 266:1981-1986. Gery, I. and Schmidt, J.A. 1985. Human interleukin 1. Methods Enzymol. 116:456-467. Hlodan, R. and Hartl, F.U. 1994. How the protein folds in the cell. In Mechanisms of Protein Folding (R.H. Pain, ed.) pp. 194-228. IRL Press, Oxford. Hopkins, T.R. 1991. Physical and chemical cell disruption for the recovery of intracellular proteins. In Purification and Analysis of Recombinant Proteins (R. Seetharam and S.K. Sharma, eds.) pp. 57-83. Marcel Dekker, New York. Johnson, B.H. and Hecht, M.H. (1994) Recombinant proteins can be isolated from E. coli by repeated cycles of freezing and thawing. Bio/Technology 12:1357-1360. Joseph-Liauzun, E., Legoux, R., Guerveno, V., Marchese, E., and Ferra, P. 1990. Human recombinant interleukin-1β isolated from E. coli by simple osmotic shock. Gene 86:291-295. Kronheim, S.R., Cantrell, M.A., Deeley, M.C., March, C.J., Glackin, P.J., Anderson, D.M., Hemenway, T., Merriam, J.E., Cosman, D., and Hopp, T.P. 1986. Purification and characterization of human interleukin-1 expressed in Escherichia coli. Bio/Technology 4:1078-1082. Livi, G.P., Lillquist, J.S., Ferrara, A., Sathe, G.M., Simon, P.L., Meyers, C.A., Gorman, J.A., and Young, P.R. 1991. Secretion of N-glycosylated interleukin-1β in Saccharomyces cerevisiae using a leader peptide from Candida albicans. Effect of N-linked glycosylation on biological activity. J. Biol. Chem. 266:15348-15348. McMahan, C.J., Slack, J.L., Mosley, B., Cosman, D., Lupton, S.D., Brunton, L.L., Grubin, C.E., Wignall, J.M., Jenkins, N.A., Brannan, C.I., Copeland, N.G., Huebner, K., Croce, C.M., Cannizzarro, L.A., Benjamin, D., Dower, S.K., Spriggs, M.K., and Sims, J.E. 1991. A novel IL-1 receptor, cloned from B cells by mammalian expression, is expressed in many cell types. EMBO J. 10:2821-2832. Meyers, C.A., Johanson, K.O., Miles, L.M., McDevitt, P.J., Simon, P.L., Webb, R.L., Chen, M.-J., Holskin, B.P., Lillquist, J.S., and Young, P.R. 1987. Purification and characterization of human recombinant interleukin-1β. J. Biol. Chem. 262:11176-11181.
Nash, H.A., Robertson, C.A., Flamm, E., Weisberg, R.A., and Miller, H. 1987. Overproduction of Escherichia coli integration host factor, a protein with nonidentical subunits. J. Bacteriol. 169:4124-4127. Neidhardt, F.C. 1987. Chemical composition of Escherichia coli. In Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology (F.C. Neidhardt, J.L. Ingraham, K.B. Low, B. Magasanik, M. Schaechter, and H.E. Umbarger, eds.) pp. 3-6. American Society for Microbiology, Washington, D.C. Priestle, J.P., Schar, H.-P., and Grutter, M.G. 1988. Crystal structure of the cytokine interleukin 1β. EMBO J. 7:339-343. Scopes, R.K. 1994. Protein Purification: Principles and Practice, 3rd ed. Springer-Verlag, New York and Heidelberg. Sherwood, R.F. 1992. Making bacterial extracts suitable for chromatography. Meth. Mol. Biol. 11:287-305. Wingfield, P., Payton, M., Tavernier, J., Barnes, M., Shaw, A., Rose, K., Simona, M.G., Demczuk, S., Williamson, K., and Dayer, J.M. 1986. Purification and characterization of human interleukin1β expressed in recombinant Escherichia coli. Eur. J. Biochem. 160:491-497. Wingfield, P.T., Graber, P., Rose, K., Simona, M.G., and Hughes, G.J. 1987. Chromatofocusing on N-terminally processed forms of proteins. J. Chromatogr. 387:291-300. Wood, W.I. 1976. Tables for the preparation of ammonium sulfate solutions. Anal. Biochem. 73:250-257. Yem, A.W., Richard, K.A., Staite, N.D., and Deibel, M.R. 1988. Resolution and biological properties of three N-terminal analogues of recombinant human interleukin-1β. Lymphokine Res. 7:8592. Zimmerman, S.B. and Trach, S. 1991. Estimation of macromolecular concentrations and excluded volume effects in the cytoplasm of Escherichia coli. J. Mol. Biol. 222:599-620.
Key Reference Wingfield et al., 1986. See above. The original publication on which Basic Protocol 1 is based.
Contributed by Paul T. Wingfield National Institutes of Health Bethesda, Maryland
Purification of Recombinant Proteins
6.2.15 Current Protocols in Protein Science
Preparation and Extraction of Insoluble (Inclusion-Body) Proteins from Escherichia coli
UNIT 6.3
High-level expression of many recombinant proteins in Escherichia coli leads to the formation of highly aggregated protein commonly referred to as inclusion bodies (UNITS 5.1 & 6.1). Inclusion bodies are normally formed in the cytoplasm; alternatively, if a secretion vector is used, they can form in the periplasmic space. Inclusion bodies recovered from cell lysates by low-speed centrifugation are heavily contaminated with E. coli cell wall and outer membrane components. The latter are largely removed by selective extraction with detergents and low concentrations of either urea or guanidine⋅HCl to produce so-called washed pellets. These basic steps result in a significant purification of the recombinant protein, which usually makes up ∼60% of the washed pellet protein. The challenge, therefore, is not to purify the recombinant-derived protein, but to solubilize it and then fold it into native and biologically active protein. Basic Protocol 1 describes preparation of washed pellets and solubilization of the protein using guanidine⋅HCl. The extracted protein, which is unfolded, is either directly folded as described in UNIT 6.5 or further purified by gel filtration in the presence of guanidine⋅HCl as in Basic Protocol 2. A support protocol describes the removal of guanidine⋅HCl from column fractions so they can be monitored by SDS-PAGE (UNIT 10.1). PREPARATION AND EXTRACTION OF INSOLUBLE (INCLUSION-BODY) PROTEINS FROM ESCHERICHIA COLI
BASIC PROTOCOL 1
Bacterial cells are lysed using a French press, and inclusion bodies in the cell lysate are pelleted by low-speed centrifugation. The pellet fraction is washed (preextracted) with urea and Triton X-100 to remove E. coli membrane and cell wall material. Guanidine⋅HCl (8 M) and dithiothreitol (DTT) are used to solubilize the washed pellet protein. Extraction with the denaturant simultaneously dissociates protein-protein interactions and unfolds the protein. As a result, the extracted protein consists (ideally) of unfolded monomers, with sulfhydryl groups (if present) in the reduced state. Materials E. coli cells from fermentation (UNIT 5.3) containing the protein of interest Lysis buffer (see recipe) Wash buffer (see recipe), with and without urea and Triton X-100 Extraction buffer (see recipe) 250- and 500-ml stainless steel beakers 0.22-µm syringe filters (e.g., Millex from Millipore) 20-ml disposable syringe Additional equipment for breaking cells, homogenizing cells and pellets and centrifuging at low and high speeds (UNIT 6.2) Break cells and prepare clarified lysate 1. Place thawed E. coli cells in a stainless steel beaker. Add 4 ml lysis buffer per gram wet weight of cells. Keep bacterial cells cool by placing the beaker on ice in an ice bucket. The cells can be pretreated with lysozyme prior to lysis in the French press. Lysozyme treatment involves incubating cells ∼20 min at 20° to 25°C in lysis buffer supplemented Contributed by Ira Palmer and Paul T. Wingfield Current Protocols in Protein Science (1995) 6.3.1-6.3.15 Copyright © 2000 by John Wiley & Sons, Inc.
Purification of Recombinant Proteins
6.3.1 CPPS
with 200 ìg/ml lysozyme, with intermittent homogenization using a tissue grinder. It should be emphasized that this optional step is carried out before French press breakage and is not simply an alternative method of cell breakage (compare the comments made in the annotation to step 4 of UNIT 6.2). Its purpose is to aid removal of the peptidoglycan and outer membrane protein contaminants during the washing steps (steps 6 to 9; for further details see UNIT 6.1 and Fig. 6.1.5). An example of this approach is given in Basic Protocol 1 of UNIT 6.5.
2. Suspend cells using a Waring blender and homogenize using the Polytron tissuegrinder homogenizer until all clumps are disrupted, as described in UNIT 6.2, step 3. 3. Lyse cells with two passes through the French pressure cell operated at 16,000 to 18,000 lb/in2 (with the high-ratio setting, pressure gauge readings between 1011 and 1135), chilling the cell suspension to 4°C after each pass, as described in UNIT 6.2, steps 2 and 4. 4. Reduce the viscosity of the suspension by sonicating 5 min at full power with 50% duty cycle (on for 5 sec, off for 5 sec) using an ultrasonic homogenizer, as described in UNIT 6.2, step 5. 5. Clarify the lysed cell suspension by centrifuging 1 hr at 22,000 × g (12,000 rpm in a JA-14 rotor in a Beckman J2-21M centrifuge), 4°C. Unbroken cells, large cellular debris, and the inclusion body protein will be pelleted. The JA-14 rotor uses 250-ml centrifuge bottles. For processing smaller volumes the Beckman JA-20 rotor (or equivalent) with 50-ml tubes can be used, at 13,500 rpm (22,000 × g). The procedure for dealing with insoluble inclusion-body proteins now diverges from that for purifying soluble proteins (UNIT 6.2).
Prepare washed pellets 6. Carefully pour off the supernatant from the pellet. Using a tissue homogenizer, suspend the pellet with 4 to 6 ml wash buffer per gram wet weight cells. Complete homogenization of the pellet is important to wash out soluble proteins and cellular components. Removal of cell wall and outer membrane material can be improved by increasing the amount of wash solution to 10 ml per gram cells. The concentration of urea and Triton X-100 in the wash buffer can be varied. The urea concentration is usually between 1 and 4 M; higher concentrations may result in partial solubilization of the recombinant proteins. The usual detergent concentration is 0.5% to 5%. Triton X-100 will not solubilize inclusion body proteins; it is included to help extract lipid and membrane-associated proteins.
7. Centrifuge the suspension 30 min at 22,000 × g (12,000 rpm in JA-14), 4°C. Discard supernatant and, using the tissue homogenizer, suspend the pellet in 4 to 6 ml wash buffer per gram wet weight of cells. 8. Repeat step 7 two more times. If the supernatant is still cloudy or colored, continue washing the pellet until the supernatant is clear.
9. Suspend the pellet with wash buffer minus the Triton X-100 and urea, using 4 to 6 ml buffer per gram wet cells. Centrifuge 30 min at 22,000 × g (12,000 rpm in JA-14), 4°C. Preparation and Extraction of Inclusion Bodies
The final wash removes excess Triton X-100 from the pellet.
6.3.2 Current Protocols in Protein Science
If necessary the washed pellets can be stored at −80°C. It is better to store material at this stage rather than after the extraction stage (see comments to step 13).
Extract recombinant protein from washed pellets with guanidine⋅HCl 10. Using the tissue homogenizer, suspend the pellet with guanidine⋅HCl-containing extraction buffer. Use 0.5 to 1.0 ml buffer per gram wet weight of original cells if the extract will be subjected to gel filtration, and 2 to 4 ml buffer if the extract will be used in protein folding procedures. Perform this step at room temperature. To estimate the amount of recombinant protein in the washed pellets, use the following guidelines. (1) An expression level of 1% corresponds to ∼1 mg recombinant protein per 1 g wet cells. (2) The recovery of highly aggregated recombinant protein in the washed pellets is ∼75% that originally present in the cells. (3) About 60% of the total washed pellet protein is recombinant-derived. Thus, if 50 g cells is processed and the expression level is 5%, the washed pellets contain ∼200 mg recombinant protein. The total amount of recombinant-derived protein in washed pellets can be directly determined by measuring the total protein concentration or by analyzing the washed pellets via SDS-PAGE (see Support Protocol and UNIT 10.1) to determine the proportions of the protein constituents. For gel-filtration purposes, the pellets from 50 g wet weight E. coli cells are solubilized with 40 to 50 ml extraction buffer (see Basic Protocol 2); the concentration of recombinant protein in the extract will be 4 to 5 mg/ml. For direct protein folding (UNIT 6.5), the pellets are extracted with 100 to 200 ml buffer, and the concentration of recombinant protein 1 to 2 mg/ml. If the washed pellet is heavily contaminated with outer cell wall and peptidoglycan material, the extract must be diluted further with extraction buffer (usually 1:1 to 1:3) to reduce the viscosity before it can be used for chromatography.
11. Centrifuge the suspension 1 hr at 100,000 × g (30,000 rpm in Ti45 rotor in a Beckman Optima XL-90 ultracentrifuge), 4°C. For volumes <250 ml the Beckman 70Ti rotor (capacity 6 × 39 ml) can be used at 32,000 rpm (∼100,000 × g).
12. Carefully pour off the supernatant from the pellet. Filter the supernatant through a 0.22-µm syringe filter attached to a 20-ml disposable syringe. The filter removes unpelleted large cell wall debris that will clog most chromatography columns.
13. Use the clarified inclusion body extract for preparing folded protein (UNIT 6.5) or purify further by gel filtration (see Basic Protocol 2). The extract can be stored at −80°C until required. Freeze in plastic (or polyethylene) containers rather than glass. Divide sample into 10- to 20-ml aliquots instead of freezing in one large lot and fill containers to only 50% to 75% capacity.
MEDIUM-PRESSURE GEL-FILTRATION CHROMATOGRAPHY IN THE PRESENCE OF GUANIDINE HYDROCHLORIDE
BASIC PROTOCOL 2
Washed, extracted pellets (see Basic Protocol 1) contain >50% recombinant protein and are used as the starting material for purification of the protein of interest by gel-filtration chromatography. Superdex 200 gel-filtration medium, which allows high flow rates, is washed and packed into a column. The column is equilibrated at 4°C and the sample is applied. Assay of column fractions by gel electrophoresis in the presence of SDS is complicated by the fact that guanidine⋅HCl forms a precipitate with SDS. Therefore, preparing samples for gel analysis involves selective precipitation of protein from guanidine⋅HCl prior to
Purification of Recombinant Proteins
6.3.3 Current Protocols in Protein Science
SDS-PAGE (see Support Protocol). The purified (or partially) purified protein is used as the starting material for procedures (e.g., UNIT 6.5) in which the denatured protein is folded into a native and biologically active structure. Materials Gel-filtration medium: Superdex 200 PG (preparative grade; Pharmacia Biotech) 5% (v/v) ethanol Gel-filtration buffer (see recipe) Guanidine⋅HCl extract of E. coli cells containing the protein of interest (see Basic Protocol 1) 4- to 6-liter plastic beaker Chromatography column: Pharmacia Biotech XK 16/100, 26/100, or 50/100 Packing reservoir: Pharmacia Biotech RK 16/26 (for 16- and 26-mm-i.d. columns) and RK 50 (for 50-mm-i.d. column) Chromatography pump: Pharmacia Biotech P-6000 or P-500 Injection valve (to select between sample loop and pump) UV monitor and fraction collector Sample loop (volume determined by size of column) NOTE: The various components of the chromatography system (pumps, valves, monitors, and sample loops) listed separately above are supplied as components of the BioPilot chromatography system (Pharmacia Biotech), which is used to run the XK 50/100 column. The smaller XK columns (2.6 and 2.5 cm i.d.) are run using the FPLC chromatography system (also from Pharmacia Biotech), which is designed for small- to mediumscale work. For further details on this equipment see the manufacturer’s literature (e.g., Process Products, Pharmacia Biotech). NOTE: Perform steps 1 to 11 at room temperature. After the column is packed, equilibrate and elute at 4°C. Pack the column 1. Wash the gel-filtration medium in a large plastic beaker with 5% ethanol. Let the medium settle and adjust the volume of liquid to give a gel slurry concentration of 65% to 75%. The XK 16/100, 26/100, and 50/100 columns are 100 cm long and have inner diameters of 16, 26, and 50 mm, respectively. Hence, for an XK 50/100 column, column volume = radius (2.5 cm)2 × 3.1416 × bed height (97 cm) ≅ 1900 ml, and ∼2 liters preparative-grade Superdex 200 is required. To pack this column, the gel medium is suspended in 5% ethanol to give a total volume of 3 liters which corresponds to ∼70% gel slurry (it should be noted that the RK 50 reservoir has a capacity of 1 liter, so the 3 liters of gel slurry can be poured in a single operation).
2. Fix the chromatography column in an upright position, using a level to adjust the position. Attach the packing reservoir. 3. Add sufficient 5% ethanol to displace the air from a few centimeters of the bottom of the column. Clamp off the bottom of the column. 4. Gently mix the gel-filtration medium in the plastic beaker to an even slurry of 70% medium suspended in 5% ethanol. 5. Degas the suspension 5 to 10 min using a vacuum flask and laboratory vacuum. Preparation and Extraction of Inclusion Bodies
The ethanol is included to reduce the surface tension and density of the solvent, thus allowing air bubbles that form to rise to the surface more quickly.
6.3.4 Current Protocols in Protein Science
6. Carefully pour the slurry of medium into the column, introducing material along the side of the column to avoid creating air bubbles. 7. Let the column stand 5 min and then unclamp the bottom of the column. 8. Attach the chromatography pump to the packing reservoir and pump 5% ethanol (degassed) into the column at an appropriate flow rate (based on manufacturer’s instructions). Pack the column at a pressure greater than the pressure at which the column will be run (up to twice as high), but not greater than the maximum pressure rating of the column. The XK 50/100 column (rated to 0.5 MPa) is packed at ∼20 to 30 ml/hr and ∼0.4 MPa.
9. After the medium has settled, turn off the pump and close the bottom of the column. Pipet fluid from the reservoir and remove the reservoir. Once the column has been packed, be careful to prevent air from entering the column bed. Air will disturb the bed and reduce the column separation resolution.
10. Attach the column top adapter to the column. Place the top of the adapter onto the top of the packed medium and gently compress the medium. 11. Reattach the pump to the column and wash the column with water at a flow rate that will generate the maximum pressure to be used. If the medium continues to settle, readjust the top adapter to maintain a firm fit against the gel. From this point onward, perform all steps at 4°C. Equilibrate the column 12. Equilibrate the column with at least 1 column volume of gel-filtration buffer. Although the proteins were extracted with buffer containing 8 M guanidine⋅HCl (see Basic Protocol 1), the gel-filtration buffer contains only 4 M guanidine⋅HCl. The concentration is reduced to allow faster flow rates and for reasons of economy. Most proteins remain unfolded at the lower guanidine⋅HCl concentration. If, however, the protein elutes in an anomalous manner (e.g., in more than one peak or at an elution position not consistent with its size), and assuming there is adequate reducing agent present, then try increasing the guanidine⋅HCl concentration in the gel-filtration buffer.
13. Measure the actual flow rate while running the column at a flow rate that generates a back pressure about one-half of that generated when packing the column (step 8). For an XK 50/100 column packed using Superdex 200 at 0.4 MPa, a running pressure of ∼0.2 MPa is used, which generates flow rates of 5 to 10 ml/min that are equivalent to linear flow rates of 15.3 to 30.6 cm/hr. The linear flow rate equals the flow rate (ml/hr)/cross-sectional area (cm2). At these flow rates it takes between 3 and 6 hr to complete the chromatography.
14. Connect tubing from the end of the column to the UV monitor and the fraction collector. Apply the sample 15. Load the sample loop with the guanidine⋅HCl extract to be separated. Avoid loading a sample volume >5% of the total column volume; the optimum sample size is 2% (∼40 ml for the XK 50/100 column). The sample consists of washed pellets extracted with guanidine⋅HCl (see Basic Protocol 1). A sample size of 40 to 50 ml is usually derived from ∼50 g wet weight cells. With smaller sample sizes, use columns with proportionally smaller diameters (e.g., XK 16/100 or 26/100 columns). If purchase of only one column is possible, a 2.5 × 100–cm size is a good compromise for variable sample loading.
Purification of Recombinant Proteins
6.3.5 Current Protocols in Protein Science
16. Monitor column effluent with the UV monitor and collect fractions with the fraction collector. For an XK 50/100 column, collect 15- to 20-ml fractions in 16 × 20–mm culture tubes. The eluent from the column is usually monitored at 280 nm or, if the protein has a particularly low extinction coefficient, at 230 nm (guanidine⋅HCl strongly absorbs below 225 nm). For an XK 50/100 column, fractions need only be collected after ∼500 ml of elution. The excluded volume (void volume) is ∼570 ml. Run one column volume (1900 ml) to ensure all of the load material is eluted from the column.
17. Prepare the fractions to be assayed for SDS-PAGE (see Support Protocol and UNIT 10.1). SUPPORT PROTOCOL
PREPARATION OF SAMPLES CONTAINING GUANIDINE HYDROCHLORIDE FOR SDS-PAGE Because guanidine⋅HCl forms a precipitate with SDS, it is necessary to remove the former before carrying out SDS-PAGE. Protein in column fractions is separated from guanidine⋅HCl by precipitation using 90% ethanol (Pepinsky, 1991). Materials Sample containing the protein of interest 100% ethanol, 0° to 4°C 1× SDS sample buffer (UNIT 10.1) Gilson Pipetman (Rainin Instrument) Additional reagents and equipment for gel electrophoresis (UNIT 10.1) 1. Pipet 25 µl sample containing the protein of interest into a 1.5-ml microcentrifuge tube. 2. Add 225 µl cold (0° to 4°C) ethanol to the sample in the tube. The final ethanol concentration is 90% by volume.
3. Mix the sample and ethanol well. Chill 5 to 10 min at −20°C or colder (e.g., −80°C). 4. Microcentrifuge the sample 5 min at maximum speed (∼15,000 × g), 4°C. Carefully withdraw the supernatant and retain the pellet. The pellet may be difficult to see. Be careful not to draw the pellet out of the microcentrifuge tube with the supernatant.
5. Suspend the pellet with 250 µl cold 90% (v/v) ethanol. Mix thoroughly using a vortex mixer. The 90% ethanol is made by mixing 225 ìl ethanol and 25 ìl H2O.
6. Microcentrifuge the sample 5 min at maximum speed, 4°C. Carefully pipet off the supernatant and suspend the pellet in 25 µl of 1× SDS sample buffer. Some proteins are more difficult than others to suspend from an ethanol precipitate. Electrophoresis sample buffer containing 8 M urea is helpful for such proteins (UNIT 10.1). Sonication with a microtip probe can also be used to disperse the sample. A volume of sample buffer >25 ìl may be required in this case (e.g., 50 ìl), and great care must be taken to prevent foaming of the sample caused by excessive sonication power. Preparation and Extraction of Inclusion Bodies
6.3.6 Current Protocols in Protein Science
7. Heat the sample 3 to 5 min at 90° to 100°C. Load on an SDS-polyacrylamide gel (UNIT 10.1). REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Extraction buffer 50 mM Tris⋅Cl, pH 7.0 5 mM EDTA 8 M guanidine⋅HCl (764 g/liter) 5 mM DTT (770 mg/liter) If the buffer is cloudy, filter through a 0.45- to 0.5-µm filter (the solution should be clear if high-quality guanidine⋅HCl—e.g., ultrapure grade, ICN Biomedicals—is used; see APPENDIX 3A). Buffer can be stored minus DTT at least 1 month at 4°C. Gel-filtration buffer 50 mM Tris⋅Cl, pH 7.5 4 M guanidine⋅HCl (382 g/liter; ultrapure, ICN Biomedicals) 5 mM DTT (770 mg/liter) Buffer can be stored minus DTT at least 1 month at 4°C. Filter (as for extraction buffer; see recipe) and degas before use. Higher concentrations of guanidine⋅HCl (up to 8 M) may be required for some proteins (see comment at step 12).
Lysis buffer 100 mM Tris⋅Cl, pH 7.0 5 mM EDTA 5 mM DTT (770 mg/liter) 5 mM benzamidine⋅HCl (780 mg/liter) Prepare immediately before use The Tris⋅Cl and EDTA are diluted from concentrated stock solutions. The other components are added to the diluted buffer before use.
Wash buffer 100 mM Tris⋅Cl, pH 7.0 5 mM EDTA 5 mM DTT (770 mg/liter) 2 M urea (120 g/liter; ultrapure, ICN Biomedicals) 2% (w/v) Triton X-100 (20 g/liter; Calbiochem-Novabiochem) Add DTT, urea, and Triton X-100 to the other components directly before use. Prepare this buffer in two forms: one with and one without the urea and Triton X-100 (the latter for use in Basic Protocol 1, step 9).
Purification of Recombinant Proteins
6.3.7 Current Protocols in Protein Science
COMMENTARY Background Information The decision of whether to work with insoluble recombinant protein or to put more effort into generating soluble protein (e.g., by modifying the expression vector or changing the host strain and fermentation conditions) can be dictated by the nature of the protein. A small protein (10 to 17 kDa) with only one or two cysteine residues might be expected to fold in reasonable yield from extracted inclusion bodies. Larger proteins (>25 kDa) with many cysteine residues may be more problematical, and lower folding yields can normally be expected. In the latter case, if only small amounts of material are needed then yield is not such an important issue. It should be emphasized that, unless proved otherwise, a protein folded from insoluble inclusion bodies can be expected to have the same structural and conformational integrity as the same protein directly purified from soluble extracts (also see UNIT 6.1). It is similarly true that a purified soluble protein can be denatured and renatured (reversible denaturation) without structural or conformational modifications (reviewed by Anfinson, 1973; Ghelis and Yon, 1982).
Preparation and Extraction of Inclusion Bodies
How inclusion bodies are formed Inclusion bodies are noncrystalline, amorphous structures; however, there is some evidence that the constituent densely packed proteins may have nativelike secondary structures (Oberg et al., 1994). This suggests that the aggregates are formed by the association of partially folded protein or so-called folding intermediates. Furthermore, the aggregation appears to take place late, rather than early, in the folding pathway. Although inclusion body formation and its prevention are of great academic and commercial interest, the pragmatic situation is clearly summarized by Seckler and Jaenicke (1992), who state: “That inclusion bodies are aggregates of otherwise intact polypeptides in nonnativelike conformations has been proven repeatedly by the successful refolding of active proteins after dissociation of the aggregates by chemical dissociation.” It must be noted that the work of Oberg et al. (1994) does not invalidate this statement. It is simply a matter of terminology: the protein in aggregates may have nativelike secondary structures, but compared with soluble folded protein it must still be considered nonnative.
The propensity of a protein to form inclusion bodies is not related to the presence of sulfhydryl residues, as proteins without sulfhydryls still form such aggregates. Furthermore, where carefully studied, it has been found that aggregates derived from proteins that in their native state contain disulfides consist mainly of reduced protein (Langley et al., 1987). This also supports the view that inclusion bodies are not aggregates of completely unfolded protein because, if so, random disulfide bond formation would occur despite the reducing environment of the E. coli cytoplasm. Regardless of the sulfhydryl state in the aggregates, once the protein has been solubilized, reducing agents must be included to prevent the formation of nonnative disulfide bonds. Extracting inclusion body protein Proteins are extracted from inclusion bodies using strong protein denaturants. Protein denaturation can be induced by the following solvent conditions or reagents. 1. pH. Protein denaturation occurs because of the ionization of side chains. Generally, proteins retain less residual structure (are more denatured) when exposed to high pH (e.g., >10.5) compared to low pH (<4.5). Acidic or basic pH may be used in conjunction with urea or inorganic salts. An example of acidic pH extraction is given in UNIT 6.5. 2. Organic solvents. Examples of organic solvents include ethanol and propanol. Generally, proteins are not completely unfolded in these solvents. Organic solvents are infrequently used for the primary extraction of inclusion bodies but can be useful cosolvents to enhance protein folding (Chang and Swartz, 1993). 3. Organic solutes. Examples of organic solutes include guanidine⋅HCl (used at 6 to 8 M; effectiveness independent of solvent pH but temperature-dependent) and urea (used at 6 to 9 M; effectiveness modulated by pH, ionic strength, and temperature). Organic solutes are the most versatile and most commonly used denaturants for solubilizing inclusion bodies. The denaturing power of guanidinium salts increases in the order Cl− < Br− < I− < SCN−. 4. Detergents. The most common examples of protein-denaturing detergents are sodium dodecyl sulfate (SDS, an anionic detergent) and cetyltrimethylammonium bromide (CTAB, a cationic detergent). Both are effective protein denaturants, but they are not nor-
6.3.8 Current Protocols in Protein Science
mally used to solubilize inclusion bodies as they interfere with recovering folded protein in reasonable yields (e.g., see van Kimmenade et al., 1988). 5. Inorganic salts. Salts at high concentration (>1 M) often denature proteins. The denaturing power increases in the following order for anions: SO42− < CH3COO− < Cl− < Br− < ClO4− < SCN−; for cations, the order is (CH3)4N+, NH4+, K+, Na+ < Li+ < Ca2+ < Gdn+ (reviewed by von Hippel and Schleich, 1969). The organic guanidinium ion (Gdn) is included for comparative purposes. Salts have not been widely used for solubilizing inclusion bodies. On the other hand, they are frequently used for selective extraction of extrinsic membrane proteins and in principle should be useful for preextracting inclusion bodies. Denaturing salts such as KSCN or LiBr are sometimes referred to as chaotropes (tending to disorder). 6. Temperature. Thermally induced extraction is rarely used as it often results in irreversible protein denaturation (Zale and Klibanov, 1986). However, denaturants such as guanidine⋅HCl and urea are more effective at elevated temperatures (e.g., 37° to 60°C). Care must be taken when heating urea solutions because of the increased rate of cyanate formation, which will covalently modify amino groups on the protein, especially at pH >6. A comparison of some protein denaturants and their relative effectiveness is described by Pace and Marshall (1980, and references cited therein). Monera et al. (1994) provide an explanation of why guanidine⋅HCl is usually two to three times per mole (2- to 3-fold) more effective than urea at unfolding proteins. The mechanism of protein denaturation is reagent-specific; this topic has been reviewed by Tanford (1968), Ghelis and Yon (1982), and Creighton (1993). The extraction conditions can be empirically determined on a small scale (<1 ml) by taking aliquots of washed pellet suspension and microcentrifuging. If preweighed microcentrifuge tubes are used, the extraction conditions can be related to a given wet weight of pellet. A fixed volume of extraction buffer is added plus a reducing agent (5 to 20 mM DTT). Initially, guanidine⋅HCl (4 to 8 M) or urea (6 to 9 M) should be tried. The pellets are resuspended by vortexing, then incubated for various times at different temperatures. (Suspension of the pellet is aided by brief sonication or by using a tissue homogenizer.) After equilibration, solutions are clarified either by filtration or by
centrifugation. For the latter, a benchtop ultracentrifuge such as the Beckman XL-100 or Airfuge is ideal. The comparative effectiveness of extractants can determined by measuring the amount of protein solubilized from the washed pellets using standard protein estimation methods and/or SDS-PAGE (UNIT 10.1). It should be noted that solubilized protein must at some stage be folded into the native conformation; hence, the most effective solubilizing conditions might not necessarily be the best, especially if they cause irreversible denaturation. Irreversible denaturation appears to result from chemical modification of the protein and is induced by such factors as high temperature, extremes of pH, and tight binding of denaturants such as SDS. The extraction process (Basic Protocol 1) should result in protein that is both monomeric (assumed to be unfolded) and contains cysteine residues in the reduced state. This provides a defined starting point from which to develop a reproducible folding protocol or from which to further purify the protein by, for example, gel filtration under denaturing conditions (Basic Protocol 2). Extraction of inclusion bodies with some denaturants, such as urea, may not completely convert the protein to monomers, resulting in physically heterogeneous mixtures. As pointed out in UNIT 6.1, extraction can be accomplished with strong denaturants (e.g., guanidine⋅HCl) that can then be exchanged by dialysis or gel filtration for weaker ones (e.g., urea). This particular solvent exchange often results in much better yields of folded protein compared to that obtained by the direct removal of guanidine⋅HCl (e.g., UNIT 6.5). Gel-filtration chromatography (Basic Protocol 2) is not commonly considered a high-resolution separation technique. However, as the recombinant-derived protein content of wellprepared washed pellets will be >50% of the total (see Fig. 6.3.1, lanes h and i), only a 2-fold purification is required to obtain pure protein. Protein extracted with guanidine⋅HCl in the presence of reductant will ideally be in a random coil conformation with all sulfhydryl residues in the reduced state. Under such conditions, the order in which proteins elute from a gel-filtration matrix in guanidine⋅HCl can be directly correlated with molecular size (Mann and Fish, 1972). Selection of the proper chromatography resin is critical for success (for detailed discussion, see Critical Parameters and Troubleshooting). The main disadvantage of gel filtration in
Purification of Recombinant Proteins
6.3.9 Current Protocols in Protein Science
guanidine⋅HCl, especially when using some of the (soft) agarose-based resins (e.g., Sepharoses and Bio-Gels), is that flow rates can be very slow due to the high viscosity of the gel-filtration buffer. Basic Protocol 2 uses a Superdex matrix that permits fast flow rates in the presence of high guanidine⋅HCl (or urea). A column size of 5 × 100 cm allows ∼40 to 50 ml guanidine⋅HCl-containing extract to be processed. For smaller sample sizes, columns with proportionally smaller diameters are used. Because the proteins separated by gel filtration in the presence of guanidine⋅HCl are unfolded, they have no biological activity, so the column fractions are usually assayed by SDSPAGE. Direct addition of SDS to samples containing guanidine⋅HCl results in the formation of the insoluble guanidinium salt of dodecyl sulfate. The guanidine⋅HCl must therefore be removed before addition of the sample buffer used for SDS-PAGE analysis (UNIT 10.1). The approach used in Support Protocol 1 takes advantage of the fact that guanidine⋅HCl is soluble in 90% ethanol whereas the protein is not. An alternative is to simply remove the guanidine⋅HCl by dialysis. Small samples (10 to 100 µl) can be dialyzed using one of various microdialyzing systems commercially available (e.g., the Microdialyzer System 100 from Pierce, which allows simultaneous dialysis of up to 12 samples) or a simple homemade device that uses modified microcentrifuge tubes (Falson, 1992).
Critical Parameters and Troubleshooting
Preparation and Extraction of Inclusion Bodies
Breaking cells and preextracting inclusion bodies Once it has been established that the recombinant protein is insoluble (UNIT 6.1), extra care should be taken to ensure complete cell lysis. If unbroken cells are present in the low-speed pellets, they will leach contaminants when the pellets are extracted with strong protein denaturants. Proper operation of the French press is described in UNIT 6.2. Other major sources of contamination are outer membrane and cell wall material, most of which can be preextracted with detergents such as Triton X-100 (0.5% to 5%) and urea (1 to 4 M). The proper concentration of urea in the wash buffer is determined empirically in small-scale extractions. It is probably safe to use 2 M urea for most aggregates. The effectiveness of the wash process at removing the cell wall and outer membrane material can be improved by
increasing the volume of wash buffer (see Basic Protocol 1, step 6: use 10 ml instead of 6 ml buffer per gram cells). Treatment of the cells with lysozyme prior to homogenization with the French press will also help remove this material (UNIT 6.5, Basic Protocols 1 and 2). Although insoluble proteins are less susceptible than native proteins to proteolysis, inclusion of EDTA and a serine protease inhibitor such as benzamidine⋅HCl or AEBSF is recommended. The choice of cell lysis buffer is not critical; however, the buffer pH should be >6.5, as many soluble E. coli proteins precipitate at slightly acidic pH values. After preextraction, wash the aggregates with buffer alone to remove excess detergent. When attempting to develop a reproducible folding protocol, note that varying amounts of detergent carryover may influence the outcome. In fact, nonionic detergents can be used as cosolvents to aid folding (UNIT 6.1); even so, it is important to know how much detergent is present. Extracting inclusion bodies with guanidine⋅HCl The washed pellets containing the insoluble inclusion body proteins are extracted with guanidine⋅HCl (see Basic Protocol 1, step 10). To obtain complete dissolution of the pellets, some form of mechanical dispersion is often required. Homogenization at room temperature with a tissue grinder (as described) is often adequate; however, sonication should be used if the pellet is especially recalcitrant. Heating the solution will also aid protein solubilization; 10 to 15 min at 50° to 60°C is usually a good starting point. Excess heating, whether direct or caused by sonication without adequate cooling, should be avoided. It is worth remembering that if extraction involves extremes of heat and or pH, such conditions will favor deamidation of Asn and may also promote cleavage at the labile Asp-Pro bond and other chemical modifications of the protein (Zale and Klibanov, 1986). It is best to use the extract directly after preparation rather than storing it frozen. In general, an unfolded protein in solution is much more susceptible to proteolytic degradation and chemical modification than its native (folded) counterpart. Selecting medium and column for gel filtration in guanidine⋅HCl Selecting the proper gel-filtration resin is one of the most critical steps. First of all, the
6.3.10 Current Protocols in Protein Science
Table 6.3.1 Gel-Filtration Matrices Suitable for Use with Solutions Containing Guanidine Hydrochloride
Mass range (kDa)
Matrixa
Native proteins Sepharose CL-6B Bio-Gel A-5m Sepharose CL-4B Sephacryl S-100 HR Sephacryl S-200 HR Sephacryl S-300 HR Sephacryl S-400 HR Superdex 75 Superdex 200
Unfolded proteinsb
10-4,000 10-5,000 60-20,000 1-100 5-250 10-1,500 20-8,000 3-70 10-600
1-80 1-80 10-300 <1-30c 1-50 1-100c 1->100c <1-25 1-80
Reference Mann and Fish (1972) Mann and Fish (1972) Mann and Fish (1972) — Belew et al. (1978) — — I.P. and P.T.W. (unpub. observ.) I.P. and P.T.W. (unpub. observ.)
aAll resins are from Pharmacia Biotech except Bio-Gel A-5m, which is from Bio-Rad. The Sepharose and Bio-Gel matrices
are normally run under low pressure; all other resins can be run under low or medium pressure. Medium pressure is achieved using one of the chromatography pumps indicated in Basic Protocol 2; the pumps are normally included in the Pharmacia Biotech FPLC or BioPilot systems. bData on the fractionation range in the unfolded state refer to proteins unfolded with guanidine⋅HCl; however, the guidelines also apply to proteins unfolded and eluted with urea (assuming they are random coils). cEstimates based on fractionation range for native proteins.
chromatography medium must have high chemical stability that permits prolonged exposure to high concentrations of denaturing solvents such as guanidine⋅HCl and urea. It must also have high physical strength that allows high flow rates with viscous solvents. Suitable media are listed in Table 6.3.1. The Sepharoses and Bio-Gels are cross-linked agaroses, Sephacryl is dextran cross-linked with bisacrylamide, and Superdex is cross-linked agarose with bonded dextran. All the matrices listed are stable in high concentrations of urea and guanidine⋅HCl. Details on the properties of various media can be obtained from the manufacturers. Second, selecting a medium with the proper pore size is important for good separation. Manufacturers provide information about the separation ranges of their products. These ranges are determined for proteins in an aqueous buffer. In guanidine⋅HCl, denatured proteins (assuming they are random coils) will have a radius 3- to 4-fold greater than an average folded globular protein of the same mass. Thus, the mass range values for unfolded proteins in Table 6.3.1 should be used rather than the native protein values. Third, the pressure rating of the column must be adequate. Because 4 M guanidine⋅HCl has a higher viscosity than most aqueous buff-
ers, higher pressures are needed to achieve the same flow rates. The column should be capable of safely withstanding moderate pressures of around 0.5 MPa (5 bar) or ∼75 lb/in2 (the XK columns are rated to 0.5 MPa). The proteins separate as they pass through the column, so column length is important. A column 60 to 100 cm in length will generally be suitable. Some manufacturers offer prepacked columns with a choice of media. If column packing is to be done in the laboratory, then selecting a column with a packing reservoir is helpful. Prepacked columns (diameters 1.6, 2.6, 3.5, and 6.0 cm and length 60 cm) prepacked with Sephacryl resin (Sephacryl S-100, S-200, or S-300) or Superdex resin (Superdex 200 and 75) are commercially available from Pharmacia Biotech. Superdex 200 and 75 PG (preparative grade) 60/600 columns (6.0 cm i.d. × 60 cm length) are used with the BioPilot system (Pharmacia Biotech) either individually or sequentially to obtain, in many cases, pure protein (for example, see Fig. 6.3.1). Performing gel filtration in guanidine⋅HCl The general guidelines and critical parameters that apply to gel-filtration chromatography in general and are detailed elsewhere (UNIT 8.3) also apply here. The main distinction from typical gel filtration is the use of a highly
Purification of Recombinant Proteins
6.3.11 Current Protocols in Protein Science
Preparation and Extraction of Inclusion Bodies
viscous solvent, which requires robust matrices to obtain reasonable flow rates. On the other hand, it should be noted that the presence of high concentrations of guanidine⋅HCl gives higher-resolution separations than those obtained with normal solvents because the high viscosity of the solvent reduces diffusion, which otherwise tends to broaden elution peaks. It is important to ensure that all connections and tubing are leak-free. Concentrated guanidine⋅HCl is highly corrosive to metal and electronic components. The sample derived from extraction of highly aggregated protein must be clarified before being applied to the column. Furthermore, to prevent the sample from precipitating near or at the top of the column, it should be equilibrated to the same temperature as the column matrix and buffer. This is especially important if chromatography is carried out at 4°C and the sample was heated during extraction. The column must be equilibrated with freshly made buffer before use. The column buffer contains the sulfhydryl reagent DTT, which will oxidize over time, thus lessening its effectiveness and increasing the background UV absorbance of the buffer. If the recombinant protein has been effectively solubilized, it will be physically homogeneous and its elution position will be proportional to its mass. If the protein elutes anomalously (i.e., is located in more than one peak), there are several possible reasons. 1. Incomplete protein solubilization. Try more drastic extraction conditions—in particular, increase the denaturant concentration and temperature during extraction. Use a stronger chemical denaturant, especially if urea was originally used. 2. Aggregation/precipitation of protein(s) in the column. Ensure that the concentration of guanidine⋅HCl in the column buffer will maintain the solubility of all proteins during chromatography. The column can be run at room temperature (as opposed to 4°C), and reasonable flow rates can still be obtained with buffers containing 6 to 7 M guanidine⋅HCl (4 M is used in Basic Protocol 2). 3. Conformational heterogeneity. Related to the above two situations is the case where the protein, although monomeric, contains both unfolded and partially folded species. The folded protein will elute later (appear smaller) than the unfolded protein. Conformational heterogeneity is usually not a major problem in high (>4 M) guanidine⋅HCl concentrations.
4. Oxidation of sulfhydryl residues. The formation of intermolecular disulfide bonds is indicated by the appearance of protein eluting earlier than expected and corresponds to the formation of multimers (dimers, trimers, etc.). Disulfide interchange occurs more readily at high pH; thus, use of a slightly acidic buffer (pH 6.0 to 5.0) and inclusion of fresh reductant in both the sample and column buffer are recommended. For analytical separations, cysteine residues are normally capped by alkylation with iodoacetamide; cysteines can also be reversibility modified by sulfitolysis (Glazer et al., 1975). 5. Proteolysis. (a) The sample contains partially proteolyzed protein, with proteolysis by E. coli proteases occurring before or after cell breakage: In fully denatured protein, it should be possible to separate clipped protein from unmodified protein if there is sufficient mass difference. Once the protein has been separated by gel filtration, it should be checked by mass spectrometry to confirm its integrity. If the protein is not fully denatured, clipped protein may have similar elution properties as unmodified protein; however, the processing will be revealed when the protein is boiled in SDS and analyzed by gel electrophoresis. (b) Where the recombinant protein is itself a protease: As pointed out by Fish et al. (1969), “Since denaturation is a kinetic process, the dissolution of proteases in guanidine⋅HCl might result in a condition where the fraction of the protein not denatured at a given instance may digest the unfolded protein leading to low molecular weight estimate.” The converse of this situation is also true: if proteases are extracted under conditions where they are fully denatured and, thus, inactive (e.g., 8 M guanidine⋅HCl), changes in the solvent conditions (e.g., reducing the guanidine⋅HCl concentration or exchanging for urea) might lead to a fraction of the protein folding into an active conformation. If autolytic processing is suspected, include an appropriate inhibitor or choose a solvent pH where the enzyme has minimal activity.
Ancipitated Results Cell lysis and preparation of washed pellets Pelleted aggregates after washing contain ∼30% dry weight, of which 90% is protein. SDS-PAGE of a typical washed pellet preparation (Fig. 6.3.1, lanes h and i) indicates that recombinant bovine growth hormone (21 kDa) makes up >60% of the total protein. The
6.3.12 Current Protocols in Protein Science
a
b
c
d
e
f
g
h
i
Figure 6.3.1 Analysis by SDS-PAGE of fractions from low-speed centrifugation of E. coli cell lysates containing aggregated bovine growth hormone. A 12.5% acrylamide gel of dimensions 12 cm × 16 cm × 1.5 mm was used with the Laemmli buffer system (UNIT 10.1). Lanes a and g contain standard proteins (low-range standards, Bio-Rad) in order of increasing migration distance: phosphorylase b (97.4 kDa), bovine serum albumin (66.2 kDa), hen egg white ovalbumin (45 kDa), bovine carbonic anhydrase (31 kDa), soybean trypsin inhibitor (21.5 kDa), and hen egg white lysozyme (14.4 kDa). After low-speed centrifugation of the clarified lysate and of the washed pellet homogenate (see Basic Protocol 1, steps 5 and 7), the supernatants will be cloudy (lane f) and the pellets usually consist of two layers (see Fig. 6.1.5). The bottom layer is inclusion body protein plus unbroken cells (lanes b and c) and the top layer consists of outer membrane and peptidoglycan fragments (lanes d and e). The outer membrane proteins OmpA (35 kDa) and OmpF/C (38 kDa) are indicated by ω and ο, respectively. After the washing steps, the growth hormone (marked β, 21 kDa) is the major constituent (lanes h and i) together, in this example, with another plasmid-encoded protein, namely kanamycin phosphotransferase (marked α, 30.8 kDa), the product of the gene conferring resistance to the antibiotic kanamycin.
washed pellets analyzed in Figure 6.3.1 are typical starting materials for the protein folding and purification described in UNIT 6.5 (Basic Protocol 1). The multiple bands in the background are either derived from unbroken cells (most likely) or are E. coli cytoplasmic proteins coprecipitated or trapped during aggregate formation. The gel analysis indicates the presence of another plasmid-encoded protein, namely, kanamycin phosphotransferase (30.8 kDa), a product of the gene conferring resistance to the antibiotic kanamycin (Kane and Hartley, 1991). The washing procedure described only partially extracts the phosphotransferase but removes most of the outer membrane proteins. For further details, see the legend to Figure 6.3.1.
Gel filtration in guanidine⋅HCl Fractionation of human immunodeficiency virus type 1 (HIV-1) protease illustrates expected results from gel-filtration chromatography. The protein is expressed in E. coli as a 17-kDa precursor that at some stage undergoes autolytic processing to form the mature-sized protease (10.7 kDa). The protease and undigested precursor accumulate as insoluble inclusions in the E. coli cytoplasm. Washed pellets prepared and extracted with 6 M guanidine⋅HCl as described (see Basic Protocol 1) were applied to a Superdex 200 column. The protease eluted in a single peak (Fig. 6.3.2, fractions 66 to 72), well separated from unprocessed protein (e.g., fraction 60) and larger molecular mass proteins (e.g., fractions 50 to 55). The fractions
Purification of Recombinant Proteins
6.3.13 Current Protocols in Protein Science
2.00
A280
1.50
S 50 55 60 65 70 75 1.00
0.50
P 0.00 0
20
40
60
80
100
Fraction number
Figure 6.3.2 Gel filtration using Superdex 200 in 4 M guanidine⋅HCl. Column dimensions, 6 × 60 cm; buffer, 50 mM Tris⋅Cl (pH 7.5)/4 mM guanidine⋅Cl/2 mM DTT; flow rate, 5 ml/min (300 ml/hr).The sample was an extract containing HIV-1 protease, which has a mass of 10 kDa. Protein fractions 66 to 72 (pool P) was further purified under the same conditions using a Superdex 75 matrix. The inset shows SDS-PAGE analysis of selected fractions. The protein markers (lane S) correspond to standards with mass values of 66.2, 45, 30, 21.5, and 14.4 kDa, respectively (migration order top to bottom).
Preparation and Extraction of Inclusion Bodies
indicated in Figure 6.3.2 by P were pooled and the protein further purified by repeat chromatography on a Superdex 75 column under the same conditions. The protein at this stage is >95% pure and after solvent exchange is folded into active protein. (As it happens, the native protease is a 20-kDA homodimer that is very susceptible to autolytic digestion; hence, purification in the denatured, inactive state is highly advantageous). In the inset of Figure 6.3.2, analysis by SDS-PAGE of fraction 60 illustrates incomplete dissociation of the sample, a problem commonly observed when electrophoresing samples precipitated with ethanol and trichloroacetic acid (TCA). If necessary, repeat the analysis using SDS sample buffer (UNIT 10.1; see Support Protocol). The fractionation of HIV-1 protease is somewhat of a best-case scenario due to the small size of the protein. Usually it is not possible to purify the recombinant protein completely, especially for proteins close in size to the main
E. coli contaminants (30 to 45 kDa). In general, small proteins (<25 kDa) are best suited for the method; however, any protein will be purified to some extent, and further purification can be attempted using some of the methods discussed in UNIT 6.1. Moreover, partial purification may be all that is required to enhance protein folding significantly. Recovery of recombinant protein from gelfiltration chromatography is close to quantitative, and the purified (or partially purified) protein can be folded or can be stored in aliquots at −80°C. On thawing the protein, add fresh DTT (an additional 1 or 2 mM) and warm the solution at 37° to 40°C for several minutes before use. The protein concentration can be conveniently estimated by UV spectroscopy.
Time Considerations Cell lysis and preparation of washed pellets French press lysis of ∼150 to 200 ml cell suspension will take ∼30 to 35 min. Preparation
6.3.14 Current Protocols in Protein Science
and extraction of the washed pellet take about half a day. Gel filtration in guanidine⋅HCl Using Superdex resins in conjunction with an FPLC or BioPilot system, gel-filtration chromatography takes 3 to 6 hr and the gel analysis 1.5 to 2 hr. With low-pressure resins, elution takes much longer. For example, a Sephacryl S-300 column (2.5 × 98 cm) eluted with 6 M guanidine⋅HCl runs at ∼25 to 30 ml/hr. An average-sized protein (15 to 30 kDa) is eluted after 10 to 12 hr. A column can be loaded in late afternoon and run overnight.
Literature Cited Anfinson, C.B. 1973. Principles that govern the folding of protein chains. Science 181:223-230. Belew, M., Fohlman, J., and Janson, J.-C. 1978. Gel filtration on Sephacryl S-200 superfine in 6 M guanidine⋅HCl. FEBS Lett. 91:302-304. Chang, J.Y. and Swartz, J.R. 1993. Single-step solubilization and folding of IGF-1 aggregates from Escherichia coli. In Protein Folding: In Vivo and in Vitro (J.L. Cleland, ed.) pp. 178-188 (ACS Symposium Series No. 526). American Chemical Society, Washington, D.C. Creighton, T.E. 1993. Proteins: Structures and Molecular Properties, 2nd ed., pp. 293-296. W.H. Freeman, New York. Falson, P. 1992. An efficient procedure to dialyze volumes in the range of 10-200 µl. BioTechniques 13:20. Fish, W.W., Mann, K.G., and Tanford, C. 1969. The estimation of polypeptide chain molecular weights by gel filtration in 6 M guanidine hydrochloride. J. Biol. Chem. 244:4989-4994. Ghelis, C. and Yon, Y. 1982. Simulation of protein folding: Studies of in-vitro denaturation-renaturation. In Protein Folding, pp. 225-243. Academic Press, San Diego. Glazer, A.N., Delange, R.J., and Sigman, D.S. 1975. Modifications of sulfhydryl and disulfide groups. In Chemical Modification of Proteins, pp. 101-120 (Laboratory Techniques in Biochemistry and Molecular Biology Series). North-Holland Publishing Company, New York. Kane, J.K. and Hartley, D.L. 1991. Properties of recombinant protein-containing inclusion bodies in E. coli. In Purification and Analysis of Recombinant Proteins (R. Seetharam and S.K. Sharma, eds.) pp. 121-145. Marcel Dekker, New York.
Langley, K.E., Berg, T.F., Strickland, T.W., Fenton, D.M., Boone, T.C., and Wypych, J. 1987. Recombinant-DNA-derived growth hormone from Escherichia coli: Demonstration that the hormone is expressed in the reduced form, and isolation of the hormone in the oxidized, native form. Eur. J. Biochem. 163:313-321. Mann, K.G. and Fish, W.W. 1972. Protein polypeptide chain molecular weights by gel chromatography in guanidinium chloride. Methods Enzymol. 26:28-42. Monera, O.D., Kay, C.M., and Hodges, R.S. 1994. Protein denaturation with guanidine hydrochloride provides a different estimate of stability depending on the contributions of electrostatic interactions. Protein Sci. 3:1984-1991. Oberg, K., Chrunyk, B.A., Wetzel, R., and Fink, A.L. 1994. Nativelike secondary structure in interleukin-1β inclusion bodies by attenuated total reflectance FTIR. Biochemistry 33:2628-2634. Pace, C.N. and Marshall, H.F. 1980. A comparison of protein denaturants for β-lactoglobulin and ribonuclease. Arch. Biochem. Biophys. 199:270276. Pepinsky, B.R. 1991. Selective precipitation of proteins from guanidine hydrochloride-containing solutions with ethanol. Anal. Biochem. 195:177181. Seckler, R. and Jaenicke, R. 1992. Protein folding and protein refolding. FASEB J. 6:2545-2552. Tanford, C. 1968. Protein denaturation. Adv. Prot. Chem. 23:122-275. van Kimmenade, A., Bond, M.W., Schumacher, J.H., Laquoi, C., and Kastelein, R.A., 1988, Expression, renaturation and purification of recombinant human interleukin 4. Eur. J. Biochem. 173:109-114. von Hippel, P.H. and Schleich, T. 1969. Ion effects on the solution structure of biological macromolecules. Acc. Chem. Res. 2:257-265. Zale, S.E. and Klibanov, M. 1986. Why does ribonuclease irreversibly inactivate at high temperature? J. Biol. Chem. 25:5432-5444.
Contributed by Ira Palmer and Paul T. Wingfield National Institutes of Health Bethesda, Maryland
Purification of Recombinant Proteins
6.3.15 Current Protocols in Protein Science
Overview of Protein Folding HOW PROTEINS FOLD Where proteins are expressed as insoluble aggregates—i.e., bacterial inclusion bodies— the obligatory first step in the recovery of functional protein is to disperse the inclusion bodies in a denaturant such as 6 M guanidine⋅HCl (guanidinium chloride; GdmCl), which will yield a solution of unfolded and essentially noninteracting polypeptide chains. The next problem is the one that faced Anfinsen in his pioneering work on the mechanism of protein folding—i.e., how to recover the protein in its unique, functional, three-dimensional conformation. The main factors determining the folding of proteins, and hence their three-dimensional conformation, are summarized in the following three sections.
All the Information is in the Sequence Anfinsen (1973) showed that proteins fold up solely as a result of thermodynamic drive, which leads to the creation of the specific noncovalent interactions and disulfide bonds characteristic of the functional state. In other words, a protein folds because, under folding conditions, its native state is more stable than the unfolded state–the information determining the relative stabilities of the two states resides solely in the primary sequence. No additional energy or information, for example, in the form of a template, is required. Other factors that may increase the yield and rate of folding such as chaperone proteins, protein disulfide isomerase (PDI), and peptidyl prolyl cis-trans isomerase (PPI), all depend on this thermodynamic drive for the intrinsic ability of the intact polypeptide chain to fold.
Proteins Fold Along a Limited Number of Pathways Like a large river, proteins in the process of folding start from a large number of sources or conformations—i.e., the unfolded state(s). These rapidly condense, reducing the number of paths (intermediate states) available to the flow, until the ocean (the functional, folded state) is finally reached through something like a single pathway. As the protein folds, the number of possible intermediate states becomes increasingly limited, and further folding is thus confined to pathways by which the vast majority of conformational states potentially accessible to the polypeptide chain are precluded. In this way, proteins can fold in vivo at Contributed by Roger H. Pain Current Protocols in Protein Science (1995) 6.4.1-6.4.7 Copyright © 2000 by John Wiley & Sons, Inc.
a rate that can keep pace with the rate of synthesis; in vitro, protein molecules can fold with half-times varying from tens to thousands of milliseconds. Consider first proteins that do not contain disulfide bonds in the native state. Very early in the folding pathway, in less than a few milliseconds, the chain collapses to intermediates in which substantial proportions of the secondary structure and hydrophobic core are assembled. The stabilization of secondary structure elements and the packing of side chains continues until a compact intermediate termed the “molten globule” is formed. This contains essentially all the native secondary structure and, in many proteins, has been shown to accumulate to a significant extent (Ptitsyn et al., 1990). Although the topology of the molten globule is similar to that of the protein in the native state, the specific and persistent tertiary interactions characteristic of the native state are still lacking. This accounts for its ability to bind the hydrophobic probe 8-anilino-1-naphthalene sulfonic acid (ANS) and presumably also for its observed tendency to aggregate. The molten globule finally folds to the native state in a reaction that usually constitutes the rate-limiting step of folding. Although several proteins have been shown to exhibit multiple kinetic phases and hence small energy barriers during folding, the early stages are fast (t1⁄2 = 1 to 25 msec). Many of the slower steps observed in folding are rate-limited by the slow cis-trans isomerization of peptide bonds joining X-Pro residues (Nall, 1994). This isomerization, with half-times on the order of a minute, occurs at later stages of folding, and therefore within rather collapsed forms of the protein. The variable accessibility of the bonds involved in cis-trans isomerization, resulting from the collapsed state of the polypeptide chain, explains the somewhat variable success of peptidyl prolyl cis-trans isomerase (PPI) in catalyzing the reaction in vitro. In comparison to these generally fast folding processes, disulfide bond–containing proteins fold somewhat more slowly in vitro. Where the noncovalent interactions are relatively strong and constitute the main factors driving the folding (at least up to a molten globule state), the stages of folding prior to disulfide formation will be similar to those described above. In some disulfide-containing proteins, however, the noncovalent interactions are weaker; thus
UNIT 6.4
Purification of Recombinant Proteins
6.4.1 CPPS
quently through large nonpolar surfaces—it is not surprising to find that the occurrence of aggregation during folding is a much greater problem. The dependence of yield on aggregation is illustrated in Figure 6.4.1.
the drive to shift the equilibrium toward the compact folded state must come from the formation of the native disulfide bonds. In such cases the intermediate that accumulates during folding will be relatively expanded. Protein disulfide isomerase (PDI) is located in the lumen of the endoplasmic reticulum and is important in catalyzing formation of disulfide bridges in proteins as they fold in vivo. The isolated enzyme can, in many cases, be used to increase the rate of folding in vitro. This application of PDI has had variable success, which is accounted for (as in the case of PPI, discussed above) by the variable accessibility of the protein thiols. As PDI is a low-efficiency enzyme, large amounts are required to achieve catalysis.
HOW TO FOLD PROTEINS The three principles outlined in the discussion of How Proteins Fold lead to three sets of conditions necessary for the successful recovery of functional proteins after denaturation: (1) the folded protein should be the most stable state (see All the Information is in the Sequence, above), (2) kinetic barriers that slow down or block the folding pathway should be minimized (see Proteins Fold Along a Limited Number of Pathways), and (3) intermolecular aggregation between folding intermediates should be reduced to acceptable limits (see Aggregation is the Main Competitor to Folding). These three goals will be considered as they bear on the practical problems involved in folding recombinant proteins. They will thus provide a rationale for the protocols in UNIT 6.5.
Aggregation is the Main Competitor to Folding Most proteins tend to exhibit some tendency toward association, reflecting the high proportion (usually ∼50%) of their surface that is nonpolar. In the case of folding intermediates, which have somewhat higher proportions of nonpolar groups exposed to the solvent, it is not surprising that aggregation is a frequently experienced hazard. Aggregation during folding in vivo has been well-documented in the cases of phage tail spike protein mutants (Haase-Pettingell and King, 1988) and a-1-antitrypsin (Lomas et al., 1993). It has also been established that a major role of the families of chaperone proteins is to prevent aggregation from competing with folding in the cell (Hlodan and Hartl, 1994). In the case of multimeric proteins, whose subunit chains have evolved to interact strongly with each other—fre-
U
X1
Stabilizing the Native Functional State The “simple” case Some proteins—e.g., phosphoglycerate kinase—are composed of a single intact polypeptide chain, constituting the simple case in which all the information required to define the folding and stability of the three-dimensional conformation is present in the sequence. In such a case, the native state will represent the most readily accessible stable state. Where disulfide
X2 .... Xn
As
Overview of Protein Folding
I
N
Al
Figure 6.4.1 Illustration of the dependence of yield on aggregation. The unfolded protein (U) folds through the intermediate forms (X1 to Xn), then through the molten globule (I) to the native state (N). Some or all of the intermediates will be capable of associating, reversibly at first, to form small aggregates (As). Subsequently, however, these will associate irreversibly to form larger insoluble aggregates (Al). The rate of aggregation (ra) will depend on the concentrations of the intermediates (and hence on the rate constants for the individual steps in the folding pathway) and also on the rate constant (ki,aggr) for aggregation of each intermediate (Xi) according to the equation ra = ki,aggr [Xi]. Aggregation will therefore depend on the overall protein concentration as well as the solubility properties of each intermediate.
6.4.2 Current Protocols in Protein Science
110 100
N
90 80
% Native circular dichroism
70 60 50 40 30 20 10 0
U
–10 0
1
2
3
4
Guanidine-HCl (mol/liter)
Figure 6.4.2 Graphical illustration of reversible unfolding transition of recombinant interleukin 1β in guanidine⋅HCl, monitored by circular dichroism (CD). Near-ultraviolet CD, which indicates changes in tertiary structure, was measured at 260 nm (triangles), 287 nm (circles), and 295 nm (squares). The plateau regions are marked for the native (top; N) and unfolded (bottom; U) states respectively, and represent regions of denaturant concentration within which there is no change in protein conformation detectable by CD. CD measurements in the far-ultraviolet range, which reflect changes in secondary structure, resulted in a curve that was coincident with this one, indicating that at equilibrium, this transition is largely two-state—i.e., only the native and unfolded states are significantly populated (Craig et al., 1987).
bonds are involved, they supplement the noncovalent interactions in contributing to the stability and specificity of the native three-dimensional conformation and the folding intermediates (see discussion of Disulfide-bonded proteins, below). Covalently bonded ligands (e.g., heme in cytochrome c) may also contribute to the stability of both the native and intermediate forms and therefore play an important role in folding (Roder and Elöve, 1994). When such proteins are subjected to dena-
turing agent, such as guanidine⋅HCl, a reversible unfolding transition is normally observed (Fig. 6.4.2). For the majority of proteins, the equilibrium transition region reflects the varying proportions of the native and unfolded states only, because the concentrations of folding intermediates are rather low at equilibrium. To fold a protein from inclusion bodies solubilized with 6 M guanidine⋅HCl, for example, the concentration of denaturant must be reduced to a value within the native plateau re-
Purification of Recombinant Proteins
6.4.3 Current Protocols in Protein Science
Overview of Protein Folding
gion. It is useful therefore to have access to an experimentally determined transition curve such as that for interleukin 1β illustrated in Figure 6.4.2. Effects of denaturants. Within the plateau region of the folding transition curve (Fig. 6.4.2), the stability of the native protein decreases with increasing concentration of denaturant. The possible disadvantage of limiting the concentration of denaturant in the interest of protein stability may be balanced, however, by the fact that the tendency to associate will be decreased. This may apply also to the transient intermediates formed during folding. Guanidine⋅HCl is not only a more powerful denaturant than urea (∼2.5-fold more effective on a molar basis), but its solubilizing effect on different chemical groups in proteins is different from that of urea. In particular, the ratio of the solubilizing power for the peptide bond relative to that for the average nonpolar side chain is greater for guanidine⋅HCl than for urea (Lapanje et al., 1978; Mitchinson and Pain, 1985). Thus, for a folding intermediate, where secondary structure (in which peptide bonds are shielded from the solvent) has formed but where nonpolar groups are more exposed, urea may be more effective than guanidine⋅HCl in stabilizing the intermediate against aggregation (assuming denaturant concentrations at which both urea and guanidine-HCl have the same effect on overall protein stability). Hence, urea is occasionally used as an intermediate solvent during renaturation. Effects of cosolvents. Cosolvents such as glycerol and ethylene glycol, glucose and sucrose, certain anions such as phosphate and sulfate, and certain cations such as 2-(N-morpholino)ethanesulfonic acid (MES) can stabilize proteins markedly against denaturation by urea or guanidine⋅HCl. They exhibit little or no effect on the folding rate constant, but dramatically decrease the unfolding rate constant. In certain favorable cases, the unfolded protein can be refolded simply by adding sulfate to the solution containing denaturant (Mitchinson and Pain, 1985). However these cosolvents, which act by stabilizing hydrophobic interactions, also stabilize aggregates; hence their usefulness is likely to be limited, probably to proteins with a relatively high proportion of polar residues.
of a number of chains. Functional proteins that are derived from precursors activated in this manner pose a more complex problem for folding in vitro. Some of the information required to define the conformation, originally resident in the intact precursor protein, has been lost during activation and the conformation is stabilized by the disulfide bonds. For example, reduction of the disulfide bonds in insulin results in unfolding and subsequent aggregation even in the absence of denaturant (Anfinsen, 1967). Penicillin acylase, which contains no disulfide bonds, is more difficult to fold in the absence of the linking peptide present in the proenzyme (Lindsay and Pain, 1991). In both of these cases, at least part of the function of the linking peptide is thought to be to increase the frequency of productive interactions between the two elements of structure (Hlodan et al., 1991). Denaturation of such proteins is not a simple thermodynamically reversible process. In cases where it is not possible to produce and activate the preprotein to obtain the functional protein, each chain may have to be produced separately and alternative folding and assembly strategies may have to be explored. For example, one chain that aggregates when folded by itself may be stabilized in its folded state if folding is carried out by diluting it into a solution of the already folded second chain (Lindsay and Pain, 1991). Stabilizing cosolvents may induce native-like conformation in the isolated chains, encouraging assembly of the chains into a functional molecule.
Proteolytically processed proteins Other proteins—e.g., chymotrypsin—are derived from precursors that have been processed by proteolytic cleavage and are composed
Figure 6.4.3 Equilibrium for noncovalent interactions during protein folding, where n is the number of disulfide bonds in the native, functional protein.
Disulfide-bonded proteins Disulfide bonds contribute to the stability of the native conformation to an extent that varies from one protein to another. The overall stability may be expressed thermodynamically in terms of the equilibria for the sidechain and backbone noncovalent interactions (Fig. 6.4.3) and the covalent interaction (Fig. 6.4.4) in which two thiol groups produce a disulfide bond.
U(–SH)2n
N′(–SH)2n
6.4.4 Current Protocols in Protein Science
X – SH + HS –Y
X – S – S –Y
Figure 6.4.4 Equilibrium for the covalent interactions (formation of disulfide bonds) during the folding of a protein, X and Y are cysteine residues in the amino acid sequence of the folding protein that interact specifically in the folded structure.
At one extreme—where the conditions of folding (e.g., denaturant concentration and pH) lead to noncovalent interactions that are strong enough to stabilize the protein in a native-like form, with the native pairs of thiol groups forced close to one another (i.e., the equilibrium illustrated in Fig. 6.4.3 lies to the right)—very little oxidizing pressure will be needed to form the disulfide bonds. If, on the other hand, the noncovalent interactions are relatively weak (i.e., the equilibrium illustrated in Fig. 6.4.3 lies well over to the left and hence toward the expanded state of the protein), more energy—a stronger oxidation potential—will be required to drive the equilibrium to the native state (Gilbert, 1994). To fold proteins containing disulfide bonds, it is necessary to pay attention to the chemistry of the covalent bonds involved in cysteine thiol oxidation. Two principal folding routes are possible—one utilizing air oxidation in the presence of trace heavy metals (Fig. 6.4.5) and the
SH
S
S– + R–S–S–R
+ 2e – + 2H+
P
SH
P
Figure 6.4.5 Formation of disulfide bond by air oxidation of thiol groups in the presence of trace heavy metals.
S heavy metals
P
other involving formation of mixed-disulfide intermediates (Fig. 6.4.6). The former route (Fig. 6.4.5), although cheaper when using large volumes, is less easy to optimize and control. The latter route (Fig. 6.4.6) utilizes a low-molecular-weight dithiol (R-S-S-R), usually glutathione. In practice, a redox pair of reduced glutathione (GSH) and oxidized glutathione (GSSG) is needed to create the appropriate oxidizing “pressure” that will allow the disulfides in the folding intermediates to make and break, so that the noncovalent interactions can shuffle the protein to its lowest free energy state. Because of the energy balances discussed above, the lowest free energy state will be expected to vary from protein to protein; hence the optimal redox conditions will also differ. However, a GSH:GSSG molar ratio of 10:1 (at a concentration of 2 to 5 mM GSH) gives maximal rates of folding to the native structure for a number of proteins. Because each step in the oxidation involves nucleophilic attack by an S− group (see Fig. 6.4.6), the rate of disulfide formation is expected to increase within the range of pH 6 to 10. However, pH may also affect the stability of folding intermediates in an unpredictable way. In cases where the noncovalent interactions alone do not lead to a condensed intermediate (i.e., where the equilibrium illustrated in Fig. 6.4.3 lies to the left; see discussion above), addition of a stabilizing cosolvent such as phosphate or sulfate can drive the equilibrium in the direction of condensation, thereby offering the possibility of increasing the rate of disulfide formation
S
S–S–R + R–S–
P
S–
S–
+ R–S–
P S
Figure 6.4.6 Formation of disulfide bond through mixeddisulfide intermediates.
Purification of Recombinant Proteins
6.4.5 Current Protocols in Protein Science
and interchange. Other factors (e.g., ionic strength and temperature) that can affect the stability of intermediates will also influence the kinetics and yield of folding.
Achieving an Acceptable Rate of Folding Within the plateau region of the unfolding transition (see Fig. 6.4.2), the rate of folding will generally be in inverse proportion to the concentration of denaturant. The rate of folding will increase with temperature, up to that of thermal denaturation. For certain proteins whose folding is rate-limited by proline cistrans isomerization (see Proteins Fold Along a Limited Number of Pathways, above), the rate can be increased by the addition of peptidyl prolyl cis-trans isomerase (Nall, 1994). However, when working with proteins lacking disulfide bonds, rate is not as important for practical purposes as yield. The rate of folding for disulfide-bonded proteins in vitro can be very slow, with half-times of the order of minutes or hours, and conditions must be arranged to reduce the time required to realize an acceptable yield. The linkage between noncovalent and disulfide stability (see discussion of disulfide-bonded proteins under Stabilizing the Native Functional State) has important consequences in defining conditions for folding disulfide-bonded proteins. Certain intermediates in some folding pathways have been shown to possess nonnative disulfides (Gilbert, 1994). The energy barriers involved in making and breaking both covalent disulfide links and noncovalent interactions must therefore not be too high if such a pathway is to be followed. If, as a result of the redox conditions, the nonnative disulfides in the intermediates are so strong that they cannot be broken to allow them to shuffle, the native noncovalent interactions will not be able to form. It is necessary therefore to create a redox potential such that the equilibrium illustrated in Figure 6.4.4 is appropriately balanced. The converse of this requirement is that the equilibrium in Figure 6.4.3 must be appropriately balanced to allow the noncovalent interactions to shuffle and thereby allow the native disulfide pairs to form. This can be achieved by adjusting the concentrations of denaturant and cosolvent.
Limiting Aggregation Aggregates of unfolded proteins are intrinsically more stable than the folded, monomeric
state, as is shown by the formation of inclusion bodies. A major factor in stabilizing aggregates appears to be hydrophobic interaction; therefore factors that can decrease the strength of such interaction or increase the solubility of other parts of the protein chain will help relieve problems of low yield stemming from aggregation. The characteristic inverse dependence of stability on temperature in hydrophobic interactions suggests that lower temperatures should lessen aggregation. Nondenaturing concentrations of urea or guanidine⋅HCl have been also used with considerable success to lessen aggregation. These concentrations have usually been arrived at empirically, by dialyzing denaturant out of the denaturation mixture. Increasing the net charge by moving away from the isoelectric point and/or lowering the ionic strength may also lessen aggregation, other factors being equal. Chaperone proteins, notably GroEL, have in certain cases been shown to dramatically increase the yield of refolding proteins in vitro (Goloubinoff et al., 1989). This multimeric protein, cylindrical in shape with a cylindrical core, will bind partially folded protein monomers, effectively removing them from free solution, and thereby protecting them against aggregation. The protein is then released with the cooperation of a second chaperone protein, GroES, under the influence of ATP. The obvious factor governing aggregation is protein concentration (Hlodan et al., 1991; Thatcher and Hitchcock, 1994). Diluting the unfolded protein into larger volumes of buffer should increase the proportion of molecules that fold, although problems of recovery will also increase. Dropwise addition of the solution of unfolded protein to a volume of buffer over a longer period of time can reduce the concentration of “sticky” intermediates at any given time. Removal of denaturant from the protein while the protein is isolated in reverse micelles or a gel-exclusion medium, or while it is in equilibrium with unfolded protein bound to an ion-exchange column, are possible approaches to refolding at higher concentrations of protein (Thatcher and Hitchcock, 1994). It should be emphasized that the problem of aggregation is a kinetic one depending on the ratio of rates (see Fig. 6.4.1) under the particular working conditions. Although general principles can provide a rationale for possible methods, success will come from largely empirical application of the principles.
Overview of Protein Folding
6.4.6 Current Protocols in Protein Science
LITERATURE CITED Anfinsen, C.B. 1967. The formation of the tertiary structure of proteins. Harvey Lect. 61:95-116. Anfinsen, C.B. 1973. Principles that govern the folding of protein chains. Science 181:223-230. Craig, S., Schmeissner, U., Wingfield, P., and Pain R.H. 1987. Conformation, stability and folding of interleukin-1β. Biochemistry 26:3570-3576. Gilbert, H.F. 1994. The formation of native disulfide bonds. In Mechanisms of Protein Folding (R.H. Pain, ed.) pp. 109-111. Oxford University Press, Oxford. Goloubinoff, P., Christeller, J.T., Gatenby, A.A., and Lorimer, G.H. 1989. Reconstitution of active dimeric ribulose bisphosphate carboxylase from an unfolded state depends on two chaperone proteins and ATP. Nature 342:884-889. Haase-Pettingell, C.A. and King, J. 1988. Formation of aggregates from a thermolabile in vivo folding intermediate in P22 tail spike maturation. J. Biol. Chem. 263:4977-4983. Hlodan, R., Craig, S., and Pain, R.H. 1991. Protein folding and its implications for the production of recombinant proteins. Biotechnol. & Genet. Eng. Rev. 9:47-88. Hlodan, R. and Hartl, F.U. 1994. How the protein folds in the cell. In Mechanisms of Protein Folding (R.H. Pain, ed.) pp. 194-228. Oxford University Press, Oxford. Lapanje, S., Skerjane, J., Glavnik, S., and Zibret, S. 1978. Thermodynamic studies of the interactions of guanidinium chloride and urea with some oligoglycines and oligolysines. J. Chem. Thermodynam. 10:425-433.
assembly of penicillin acylase, an enzyme composed of two polypeptide chains that result from proteolytic activation. Biochemistry 30:90349040. Lomas, D.A., Evans, D.Ll., Stone, S.R., Chang, W.-S.W., and Carrell, R.W. 1993. Effect of the Z mutation on the physical and inhibitory properties of α1-antitrypsin. Biochemistry 32:500-508. Mitchinson, C. and Pain, R.H. 1985. Effects of sulfate and urea on the stability and reversible unfolding of β-lactamase from Staphylococcus aureus. J. Mol. Biol. 184:331-342. Nall, B.T. 1994. Proline isomerization as a rate-limiting step. In Mechanisms of Protein Folding (R.H. Pain, ed.) pp. 80-103. Oxford University Press, Oxford. Ptitsyn, O.B., Pain, R.H., Semisotnov, G.V., Zerovnik, E., and Razgulaev, D.I. 1990. Evidence for a molten globule state as a general intermediate in protein folding. FEBS (Fed. Eur. Biochem. Soc.) Lett. 262:20-24. Roder, H. and Elöve, G.A. 1994. Early stages of protein folding. In Mechanisms of Protein Folding (R.H. Pain, ed.) pp. 37-40. Oxford University Press, Oxford. Thatcher, D. and Hitchcock, A. 1994. Protein folding in biotechnology. In Mechanisms of Protein Folding (R.H. Pain, ed.) pp. 242-250. Oxford University Press, Oxford.
Contributed by Roger H. Pain Jozef Stefan Institute Ljubljana, Slovenia
Lindsay, C.D. and Pain, R.H. 1991. Refolding and
Purification of Recombinant Proteins
6.4.7 Current Protocols in Protein Science
Folding and Purification of Insoluble (Inclusion Body) Proteins from Escherichia coli
UNIT 6.5
Heterologous expression of recombinant proteins in E. coli often results in the formation of insoluble and inactive protein aggregates, commonly referred to as inclusion bodies. To obtain the native (i.e., correctly folded) and hence active form of the protein from such aggregates, four steps are usually followed. In the first step, the bacterial cells are lysed and the aggregates collected by low-speed centrifugation (UNITS 6.2 & 6.3). In the second step (which is usually but not always included) the aggregates are preextracted to remove cell wall and outer membrane components, producing washed pellets (UNIT 6.3). In the third step, the aggregates are solubilized (or extracted) with strong protein denaturants— e.g., guanidine⋅HCl, 8 M urea, or organic acids at high concentrations (UNIT 6.3). In the fourth step, the solubilized, denatured proteins are folded with concomitant oxidation of reduced cysteine residues into the correct disulfide bonds to obtain the native protein. Basic Protocols 1, 2, and 3 in this unit feature three different approaches to the final step of protein folding and purification. Basic Protocol 1 describes the extraction of bovine growth hormone (BGH) using guanidine⋅HCl, after which the solubilized protein is folded (before purification) in an “oxido-shuffling” buffer system to increase the rate of protein oxidation. A nondenaturing concentration of urea (a cosolvent; see UNIT 6.4) is added to the buffer used in the folding step to increase the yield of folded protein by minimizing the formation of aggregated, misfolded protein. The folded protein is then purified by chromatography. Basic Protocol 2 describes folding and purification of the cytokine interleukin 2 (IL-2). This procedure differs from Basic Protocol 1 in that (1) acetic acid is used to solubilize (i.e., extract) the protein, (2) the acid-soluble protein is partially purified by gel filtration before folding, and (3) protein folding and oxidation are carried out with no additives; the protein (in acid solution) is simply dialyzed against water, thereby allowing the pH to slowly increase and air oxidation of the protein to occur. The folded protein is further purified by reversed-phase high-performance liquid chromatography (RP-HPLC). A support protocol is included for rapidly determining the amount of folded protein that contains the correct disulfide linkage pattern. Basic Protocol 3 describes the folding and purification of a fusion protein containing the catalytic domain of the human immunodeficiency virus 1 (HIV-1) integrase. The fusion partner is a 20-amino-acid peptide containing a track of six histidine residues (a His tag; see UNIT 5.1). The protein is extracted with guanidine⋅HCl (as with BGH), then purified by gel filtration in guanidine⋅HCl (UNIT 6.3). The protein is further purified by metal-chelate affinity chromatography, which takes advantage of the His tag. Next, the purified protein is folded by very slow dilution into a folding buffer containing additives required to maintain the solubility of the folded protein. The His tag is removed by digestion with the protease thrombin, the thrombin is removed by affinity chromatography using an immobilized substrate, and finally the cleaved protein is run through a gel-filtration column to remove any aggregated protein.
Purification of Recombinant Proteins Contributed by Paul T. Wingfield, Ira Palmer, and Shu-Mei Liang Current Protocols in Protein Science (1995) 6.5.1-6.5.27 Copyright © 2000 by John Wiley & Sons, Inc.
6.5.1 CPPS
BASIC PROTOCOL 1
FOLDING AND PURIFICATION OF BOVINE GROWTH HORMONE The high-level expression of bovine growth hormone (BGH) in E. coli results in the formation of highly aggregated protein (inclusion bodies). To restore the BGH to its native, functional state, the E. coli cells are first lysed with the French press and the inclusion bodies are partially purified by selective extraction of soluble and particulate E. coli contaminants. The inclusion bodies are solubilized with guanidine⋅HCl to give protein that is unfolded and reduced (i.e., with four free sulfhydryl residues). The protein is then simultaneously folded and oxidized by the slow removal of protein denaturant, resulting in the formation of two disulfide bonds from the sulfhydryl residues. The solubility of the protein during folding is maintained using a cosolvent (urea) and the oxidation of cysteine residues is catalyzed by a mixture of reduced and oxidized glutathione (GSH/GSSG; see also APPENDIX 3A). The folded, oxidized protein is finally purified to homogeneity by anion exchange and gel-filtration chromatography. Materials E. coli cells expressing BGH: ∼50 g wet weight from a 1.5-liter fermentation (UNIT 5.3), and stored as a flattened paste in a sealed polyethylene bag at −80°C BGH break buffer A (see recipe), 4°C BGH break buffer B (see recipe), 4°C BGH wash buffer A (see recipe), 4°C BGH wash buffer B (see recipe), 4°C BGH extraction buffer (see recipe), 4°C BGH folding buffer A (see recipe), 4°C BGH folding buffer B (see recipe), 4°C 2 M HCl BGH column buffer A (see recipe), 4°C DEAE Sepharose CL-4B ion-exchange resin (Pharmacia Biotech) Sephadex G-100 gel filtration resin (Pharmacia Biotech) BGH column buffer B (see recipe), 4°C Blender (e.g., Waring; ≥1 liter capacity) Tissue homogenizer (e.g., Polytron, Brinkmann) Beckman J2-21M centrifuge and JA-20 rotor (or equivalent) Sonicator, ≥400 W, with sound enclosure (Branson or equivalent) 0.7-µm glass-microfiber filters, 4.7-cm diameter (Whatman GF/F) with vacuum filtration apparatus (optional) Spectra/Por 1 dialysis tubing, 40-mm diameter (MWCO 6000 to 8000; Spectrum) 5 × 50–cm and 5 × 100–cm glass chromatography columns with adjustable flow adaptors (Pharmacia Biotech) Stirred cell with Diaflo PM 10 ultrafiltration membrane (Amicon) Millex-GV 0.22-µm filter units (Millipore) Additional reagents and equipment for cell breakage using a French press (UNIT 6.2), dialysis (APPENDIX 3B), ion-exchange chromatography (UNIT 8.2), gel-filtration chromatography (UNIT 8.3), and SDS-PAGE (UNIT 10.1) NOTE: All steps are carried at 4°C unless otherwise stated. Break cells 1. Resuspend ∼ 50 g (wet weight) of E. coli cells expressing BGH in 250 ml BGH break buffer A using a blender.
Folding and Purification of Insoluble Proteins from E. coli
The lysozyme in BGH break buffer A aids in removal of the peptidoglycan and outer membrane protein contaminants (see UNIT 6.1).
6.5.2 Current Protocols in Protein Science
2. Stir cell suspension 30 min at 20° to 25°C using a heavy-duty magnetic stirrer. To reduce viscosity of the suspension, homogenize with tissue homogenizer until no clumps are detected, then centrifuge 35 min at 20,000 × g (13,500 rpm in JA-20 rotor), 4°C. 3. Resuspend pellet in 150 ml BGH break buffer B using tissue homogenizer. Break cells using French press as described in UNIT 6.2, Basic Protocol 1, steps 1 to 5. The viscosity can also be reduced by digestion with DNase and RNase as described in UNIT 6.2, in which case the EDTA in BGH break buffer B should be replaced with 5 mM MgCl2.
Prepare and extract washed pellet 4. Centrifuge suspension 40 min at 20,000 × g. Resuspend pellet in 250 ml BGH wash buffer A using the tissue homogenizer, then centrifuge 30 min at 20,000 × g. Repeat centrifugation and resuspension once using BGH wash buffer A and again using BGH wash buffer B. Discard the cloudy supernatants and save the washed pellet. If not to be used immediately, the washed pellet may be stored at −80°C.
5. Resuspend washed pellet in 150 ml BGH extraction buffer. Sonicate briefly to completely disperse protein, then clarify solution by centrifuging 30 min at 20,000 × g or by filtering through a 0.7-µm glass-microfiber filter under vacuum. Generally, when protein is extracted with guanidine⋅HCl, the reductant dithiothreitol is included in the buffer system to ensure that the protein is fully reduced prior to folding. In this procedure it was established that >80% of the protein in the inclusion bodies was already in the reduced state (P.T.W., unpub. observ.). In the extraction buffer used here, therefore, only the relatively weak reductant glutathione is included. This compound serves to maintain thiols in the reduced state, and is a component of the redox buffer that will be used later to oxidize the protein. It cannot, however, be assumed that other inclusion-body proteins contain only free thiols.
Fold protein 6. Dilute the clear amber-colored solution with an equal volume of BGH folding buffer A and pour into prewashed dialysis tubing. Use two or three pieces of tubing and fill dialysis bags only three-quarters of the way to allow for any volume increase during dialysis. The BGH concentration from step 5 should be 2 to 4 mg/ml, and after dilution should not exceed 1.0 to 2.0 mg/ml. If higher concentrations are found, the solution must be diluted further with folding buffer A. BGH concentration can be estimated by diluting the sample with 4 M guanidine⋅HCl in water and measuring the absorbance at 280 nm and 260 nm in a cell with a 1-cm path length. The total protein concentration (mg/ml) is estimated as 1.55 A280 − 0.76 A260 nm (Stoscheck, 1990). The BGH content may either be assumed to be 60% of the total protein or, more accurately, be estimated by performing SDS-PAGE and densitometry using the washed pellet from step 4. Urea is included in folding buffer A as a cosolvent to maintain solubility of the protein during refolding. Removal of guanidine⋅HCl by dialysis or dilution results in precipitation if no cosolvent is used. The urea concentration chosen (in this case 4 M) should be low enough to allow the native structure to form (see UNIT 6.4). Urea unfolding/folding profiles for BGH (i.e., equilibrium-denaturation curves in which protein conformation is measured as a function of denaturant concentration; see UNIT 6.4) were available in the literature prior to development of this method (Edelhoch and Burger, 1966). The urea concentration that induces protein unfolding can be determined rapidly by urea gradient electrophoresis (Goldenberg, 1989).
7. Dialyze solution 12 to 16 hr (or overnight) against 5 liters of BGH folding buffer A, then dialyze an additional 6 to 8 hr against 5 liters of BGH folding buffer B. The second dialysis can be continued overnight if necessary.
Purification of Recombinant Proteins
6.5.3 Current Protocols in Protein Science
a
b
c
d
e
f
g
Figure 6.5.1 SDS-PAGE of bovine growth hormones on 12.5% polyacrylamide gel. Lane A, BGH-expressing E. coli cells minus the expression vector; lanes B and C, BGH-expressing E. coli cells with ∆A-4 and A-9 BGH expression vectors, respectively; lane D, purified recombinant A-9 BGH; lane E, BGH purified from pituitary (supplied by A.F. Parlow, UCLA); lane F, purified recombinant A-9 BGH with no reductant in sample buffer; lane G, BGH purified from pituitary with no reductant in sample buffer. In lane E (pituitary BGH), the two bands correspond to full-length protein and protein truncated at the N-terminus by 4 residues. It can be seen that the bottom band has the same mobility as E. coli extracts containing the ∆A-4 BGH construct. In lane G, it it may be noted that the two bands are not resolved under nonreducing conditions.
Oxidation of the protein during dialysis can be monitored by SDS-PAGE (UNIT 10.1). SDS-denatured oxidized BGH is more compact than SDS-denatured reduced BGH and thus migrates faster as a result of the lower apparent molecular weight (i.e., 18 kDa for the oxidized form versus 22 kDa for the reduced form). SDS-PAGE band patterns of oxidized and reduced BGH are shown in Figure 6.5.1. Any free thiol groups in the sample are quenched by addition of 20 mM iodoacetamide. SDS sample buffer (UNIT 10.1) minus reductant is then added. The pH of the SDS-treated sample may have to be readjusted with dilute alkali. The proportion of oxidized versus reduced protein is finally determined by densitometry of the Coomassie blue–stained gel. It should be noted that this approach does not prove whether or not the correct disulfide bond(s) have been formed. The shift in mobility upon reduction occurs because of the formation of the disulfide bond linking Cys-51 to Cys-163. BGH in which the second disulfide bond (linking Cys-180 to Cys-188) has been selectively reduced (Graf et al., 1975) still exhibits the gel shift. Despite these potential pitfalls, the gel method is useful and correlates with other approaches—e.g., direct monitoring of disulfide formation using 2-nitro-5-thiosulfobenzoate in the presence of sodium sulfite after the free sulfhydryls have been quenched and the buffer components removed (Thannhauser et al., 1984). Sulfhydryl groups can be assayed using Ellmans reagent (Riddles et al., 1979).
8. Centrifuge the slightly cloudy solution 30 min at 20,000 × g. Discard pellet and adjust supernatant to pH 9.0 with 2 M HCl. The volume of the supernatant should be 300 to 320 ml.
Folding and Purification of Insoluble Proteins from E. coli
Purify protein by ion-exchange and gel filtration chromatography 9. Pack a 5 × 50–cm column with DEAE Sepharose CL-4B to give a bed height of ∼10 cm and equilibrate with BGH column buffer A. Apply the solution from step 8 at a flow rate of 60 ml/hr and begin collecting 15-ml fractions. See UNIT 8.2 for further information on preparing an ion-exchange column.
6.5.4 Current Protocols in Protein Science
10. Elute with BGH column buffer A at the same flow rate and continue fraction collection until A280 or A260 of eluant approaches a baseline value. Assay fractions by SDS-PAGE and pool fractions containing BGH. Under the ion-exchange conditions used, BGH does not bind to the matrix and is located in the flowthrough fractions. The more acidic E. coli contaminants bind tightly to the top portion of the column, which turns brown. Some of the earlier flowthrough fractions may be slightly contaminated with aggregated protein that separates from soluble protein as a result of the gel-filtration effect of the matrix. A BGH concentration of 0.2 to 0.3 mg/ml in ∼400 to 450 ml of pooled eluant should be obtained. This should exhibit a single band on the SDS-PAGE gel. Pooled fractions can be stored 1 to 2 days at 4°C and for longer periods at −80°C.
11. Concentrate pooled BGH-containing fractions to 50 to 60 ml (i.e., to ∼2 to 3 mg/ml protein concentration) using a stirred cell with Diaflo PM 10 ultrafiltration membrane. Centrifuge filtrate 15 min at 20,000 × g to remove any precipitated protein. 12. Pack a 5 × 100–cm column with Sephadex G-100 to a bed height of 95 cm and equilibrate with BGH column buffer B. Apply the filtrate from step 11 at a flow rate of 40 ml/hr and begin collecting 15-ml fractions. See UNIT 8.3 for additional information on preparing gel-filtration columns. If a 5 × 100–cm column is not available, a 2.5 × 100–cm column can be used. Depending on the amount of sample, the smaller-diameter column may be run several times, loading 18 to 20 ml of sample per run.
13. Elute with BGH column buffer B at the same flow rate and continue fraction collection until A280 or A260 of eluant approaches a baseline value. Assay fractions by SDS-PAGE and pool fractions containing BGH. The protein elutes in a single but slightly asymmetrical peak. The gel-filtration elution peak has a sharp front edge, whereas the descending portion is more diffuse and trailing. This elution behavior results from the fact that BGH is a rapidly associating/dissociating monomer/dimer system (with a Kd of ∼0.8 to 1.0 × 10−5 M; see Ackers, 1970, for discussion of theory). The apparent molecular weight estimated by gel filtration is 30 to 35 kDa (with a monomer mass of ∼22 kDa). The slightly alkaline pH of the column buffer and folding buffers is required to maintain solubility of the protein.
14. Repeat step 11, then filter sterilize filtrate using a Millex-GV 0.22-µm filter unit. Store purified BGH in aliquots at −80°C. For long-term storage the protein can be lyophilized. If this is to be done, the pooled fractions from step 10 should be directly dialyzed (APPENDIX 3B) against 0.1 M ammonium bicarbonate (pH 9.2 to 9.4, adjusted with ammonium hydroxide), and the dialysate filter-sterilized and freeze-dried. If an essentially salt-free protein is required, two cycles of freeze-drying should be performed and the protein reconstituted with water alone after the first drying cycle. The concentration of the purified BGH can be conveniently determined by UV absorbance measurement—i.e., 1 mg/ml native BGH has an A280 of 0.7 in a 1-cm quartz cuvette. Protein concentration may also be estimated in crude extracts using the Bio-Rad Protein Assay Kit (based on the Bradford method) and in partially purified fractions as described in the annotation to step 6. Biological activity of the protein is measured as described in Wingfield et al. (1987a). Purification of Recombinant Proteins
6.5.5 Current Protocols in Protein Science
BASIC PROTOCOL 2
FOLDING AND PURIFICATION OF HUMAN INTERLEUKIN 2 Human interleukin 2 (hIL-2) is a hydrophobic, acid-stable polypeptide with a molecular weight of 15 kDa. Recombinant hIL-2 expressed in E. coli is deposited in inclusion bodies. Extraction of the inclusion bodies with denaturants followed by folding in aqueous solution often results in formation of insoluble hIL-2 aggregates. In this protocol, inclusion bodies containing recombinant hIL-2 are first extracted with acetic acid and the soluble monomeric hIL-2 is separated from other hIL-2 aggregates by gel-filtration chromatography. The soluble protein is then folded and oxidized by dialysis against water. Correctly folded hIL-2 is finally separated from unfolded hIL-2 and other E. coli contaminants by reversed-phase high-performance liquid chromatography (RP-HPLC). Materials hIL-2 break buffer (see recipe), 4°C E. coli cells expressing hIL-2: ∼20 g wet weight from a 3-liter fermentation (UNIT 5.3), and stored as a flattened paste in a sealed polyethylene bag at −80°C Sucrose Lysozyme (Worthington) hIL-2 wash buffer: 0.75 M guanidine⋅HCl/1% (w/v) Tween 20 (prepare immediately before use), 4°C PBS (APPENDIX 2E), 4°C 10% and 20% (v/v) acetic acid, 4°C (prepare fresh from glacial acetic acid) Sephadex G-100 gel-filtration resin (Pharmacia Biotech) Acetonitrile (HPLC grade) Trifluoroacetic acid (TFA; HPLC grade) 7 × 250–mm 300-Å octyl Aquapore RP-300 semiprep column (Brownlee column; Thomson Instrument) RP-HPLC solvent A (see recipe), room temperature RP-HPLC solvent B (see recipe), room temperature 25 mM acetic acid, 4°C Tissue homogenizer (e.g., Polytron, Brinkmann) 30°C water bath Sorvall RC-5C centrifuge with SS-34 rotor (or equivalent) 2.6 × 100–cm glass chromatography column Spectra/Por 3 dialysis tubing, 11.5- and 45-mm diameters (MWCO 3500; Spectrum) Sterivex-GS 0.22-µm filter units (Millipore) HPLC system with pumps, UV detector, and fraction collector (Waters) Additional reagents and equipment for cell breakage using a French press (UNIT 6.2), gel-filtration chromatography (UNIT 8.3), and dialysis (APPENDIX 3B) Break cells 1. Add 60 ml hIL-2 break buffer to 20 g (wet weight) of E. coli cells expressing hIL-2 in a 250-ml glass beaker. Mix well using tissue homogenizer. UNIT 6.2
describes use of a tissue homogenizer to resuspend E. coli cells.
2. Add 21 g sucrose and mix well with cell paste using tissue homogenizer. Add 34 mg lysozyme and mix again using tissue homogenizer. Incubate 30 min in a 30°C water bath, then dilute with 100 ml hIL-2 break buffer and cool on ice. Folding and Purification of Insoluble Proteins from E. coli
3. Break cells by passing suspension through a French press twice as described in UNIT 6.2. Centrifuge 30 min at 13,000 × g (10,400 rpm in SS-34 rotor), 4°C. Save pellet. The wet weight of the pellet should be ∼1.5 g.
6.5.6 Current Protocols in Protein Science
4. Wash pellet by resuspending in 30 ml hIL-2 wash buffer and centrifuging 30 min at 13,000 × g. Repeat wash once with hIL-2 wash buffer and again using 30 ml PBS in place of the wash buffer. Washed pellets should be used immediately or stored at −70°C until required.
Extract protein with acid and prepare gel-filtration column 5. Add 12.5 ml of 20% acetic acid to the washed pellet and mix using tissue homogenizer. Centrifuge 30 min at 13,000 × g, 4°C, and save the supernatant. 6. Pack a 2.6 × 100–cm column with Sephadex G-100 to give a bed height of 95 cm and equilibrate with 10% acetic acid. For convenience the column may be packed the day before use.
7. Repeat step 5 and pool the two clear supernatants (total volume should be ∼25 ml). Isolate hIL-2 monomer from solubilized protein by gel filtration 8. Immediately apply pooled supernatants to column prepared in step 6. Perform gel filtration chromatography at 4°C as described in UNIT 8.3 and construct a chromatogram, pooling the fractions that make up the third peak. The solution should be applied to the column immediately after extraction to prevent any covalent modification of the protein. The column eluate usually shows three major peaks in A280. The first peak (in the void volume) contains aggregates, the second peak contains dimeric hIL-2, and the third peak contains monomeric hIL-2 (S.M.L., unpub. observ.). A T cell proliferation assay shows that the third peak contains the highest biological activity of hIL-2 (Liang et al., 1986; Bottomly et al., 1991).
9. Using 45-mm diameter Spectra/Por 3 dialysis tubing (MWCO 3500), dialyze the pooled fractions (∼60 ml) making up the third peak at 4°C overnight against 5 liters of Milli-Q water, then for an additional 3 to 4 hr against 5 liters of fresh Milli-Q water. The pooled eluate fractions should be extensively dialyzed against water to remove acetic acid. The dialysate should have a volume of ∼60 ml with an hIL-2 concentration of 0.2 mg/ml. After dialysis, the hIL-2 monomer can be stored at −20°C until required. As the acetic acid is removed; sample pH slowly increases, thereby allowing hIL-2 thiols to form disulfide bonds.
Purify protein by RP-HPLC 10. Filter the dialysate through a 0.22-µm filter unit, then add 30 ml acetonitrile (33% v/v final) and 0.09 ml TFA (0.1% v/v final). 11. Prepare and test the HPLC system according to manufacturer’s instructions. Place a 300-Å octyl Aquapore RP-300 reversed-phase chromatography column in line. Other reversed-phase columns; e.g., the 2.2 × 15–cm TMS-250 (TosoHaas) can also be used (Kato, 1985).
12. Fill the reservoirs of the HPLC system gradient maker with RP-HPLC solvent A and RP-HPLC solvent B. Carry out a blank run (at room temperature) with the following gradient program: 0 min: 0% solvent B/100% solvent A 10 min: 0% solvent B/100% solvent A 60 min: 50% solvent B/50% solvent A 90 min: 50% solvent B/50% solvent A 150 min: 70% solvent B/30% solvent A
Purification of Recombinant Proteins
6.5.7 Current Protocols in Protein Science
160 min: 70% solvent B/30% solvent A 170 min: 100% solvent B/0% solvent A 180 min: 100% solvent B/0% solvent A 190 min: 0% solvent B/100% solvent A 210 min: 0% solvent B/100% solvent A. Repeat blank run as necessary to ensure that the baseline profile has stabilized.
13. Pump sample from step 10 into column at a flow rate of 4 ml/min. Run the gradient program described in step 12 at a flow rate of 1 ml/min at room temperature. Collect 8-ml fractions at a rate of 1 ml/min and pool the fractions making up the correctly folded hIL-2 peak, which elutes at ∼70 min. Care should be taken to avoid introducing air bubbles while pumping the sample into the column, as these will generate false peaks in the chromatogram. If the peaks in the HPLC profile are too broad, the gradient program should be varied to improve the separation (see Chapter 8).
14. Using 11.5-mm diameter Spectra/Por 3 dialysis tubing (MWCO 3500), dialyze the pooled hIL-2 fractions against Milli-Q water or 25 mM acetic acid at 4°C. Filter the purified protein solution using a 0.22-µm filter unit and store at −20°C or below. SUPPORT PROTOCOL
RESOLUTION OF NATIVE AND MISFOLDED FORMS OF hIL-2 BY RP-HPLC Human interleukin 2 (hIL-2) contains three cysteine residues—at positions 58, 105, and 125. In the correctly folded (native) protein, the cysteines at positions 58 and 105 form a disulfide bond that is important for biological activity of the protein (Robb et al., 1984; Wang et al., 1984; Liang et al., 1985). As hIL-2 contains three cysteines, three possible intramolecular disulfide–linked forms can exist, only one of which is the native form. RP-HPLC can separate these isomers, and is used to verify that the purified IL-2 contains the native disulfide. Additional Materials (also see Basic Protocol 2) 0.46 × 10–cm SynChropak RP-P C18 column (Thomson Instrument) RP-HPLC solvent C (see recipe) 25 mM acetic acid Purified folded hIL-2 solution (see Basic Protocol 2) 1. Prepare and test HPLC system according to manufacturer’s instructions. Place a 0.46 × 10–cm SynChropak RP-P C18 column in line. 2. Fill the reservoirs of the HPLC system gradient maker with RP-HPLC solvent A and RP-HPLC solvent C. Apply 20 µl of 25 mM acetic acid as a blank sample. Perform a blank run at room temperature using a gradient program starting with 100% solvent A for 5 min, followed by a linear gradient starting at 100% solvent A/0% solvent C and ending at 0% solvent A/100% solvent C, at a flow rate of 1 ml/min. Repeat as necessary to check that the baseline profile is reproducible. 3. Prepare a solution of ∼5 mg/ml purified folded hIL-2 solution in 25 mM acetic acid and filter through a 0.22-µm filter.
Folding and Purification of Insoluble Proteins from E. coli
4. Apply 20 µl of purified folded hIL-2 solution in 25 mM acetic acid to the column and carry out RP-HPLC at room temperature using the gradient program described in step 2. Collect the fractions containing the properly folded hIL-2.
6.5.8 Current Protocols in Protein Science
A
B
A214
0.3 0.2 0.1 0.0 C
D 1
0.3 0.2 0.1 0.0
2
0
40
80
120
0
40
80
120
Time (min)
Figure 6.5.2 Chromatograms illustrating peaks produced by (A) correctly folded hIL-2 (in absence of denaturant); (B) correctly folded hIL-2 (in presence of denaturant but at pH 3.5, which is too low for disulfide-bond exchange); (C) scrambled hIL-2 isomers (resulting from denaturant treatment at pH 8.5); (D) unfolded hIL-2 (resulting from denaturant/reductant treatment).
Properly folded hIL-2 usually elutes at ∼54% acetonitrile and unfolded hIL-2, which has no intramolecular disulfide bond, elutes at 57% acetonitrile. In the presence of denaturants such as guanidine⋅HCl, hIL-2 rapidly scrambles into a mixture of three disulfide-linked isomers. The incorrectly folded hIL-2 elutes at a lower acetonitrile concentration (<54%) as shown in Figure 6.5.2. For each separation in Figure 6.5.2, two Vydac C4 columns were connected in series and run at 40°C with a 1 ml/min flow rate. The sample was eluted with an acetonitrile gradient that increased from 0% to 44% in 15 min, and then to 64% in 120 min. In the separation represented by the chromatogram in panel A, 100 ìg hIL-2 in 40 ìl of 50 mM acetic acid was dissolved in 400 ìl of 20% formic acid and chromatographed. The elution peak (at ∼57.3% acetonitrile) represents native hIL-2. In panel B, hIL-2 was added to 300 ìl of 6 M guanidine⋅HCl/0.2 M acetic acid, pH 3.5, and incubated 10 min at room temperature. 200 ìl of 20% formic acid was added and the sample was chromatographed. Note that the chromatogram has not changed, as the pH is below the point where isomerization (disulfide-bond exchange) occurs. In panel C, hIL-2 was added to 300 ìl of 6 M guanidine⋅HCl/0.05 M Tris⋅Cl, pH 8.5, incubated 5 min at room temperature, then quenched with formic acid as in the separation represented by panel B. Peak 1 represents the scrambled isomer with a disulfide bond between Cys-105 and Cys-125, whereas peak 2 is probably the scrambled isomer with a disulfide bond between Cys-58 and Cys-125. Finally, in panel D, hIL-2 was added to 6 M guanidine⋅Cl as in panel C. 50 mM DTT was then added and the sample was incubated 40 min at room temperature, then quenched with formic acid as in panel B. The peak (at ∼59.4% acetonitrile) represents completely reduced (unfolded) hIL-2. (see also Browning et al., 1986). IMPORTANT NOTE: If the pressure in the HPLC column exceeds 4000 psi, the column should be cleaned.
Purification of Recombinant Proteins
6.5.9 Current Protocols in Protein Science
BASIC PROTOCOL 3
FOLDING AND PURIFICATION OF A HISTIDINE-TAGGED PROTEIN: HIV-1 INTEGRASE This protocol describes the folding and purification of the central core domain of the HIV-1 integrase, which consists of residues 50 to 212 (IN50-212). The protein is expressed in E. coli with an N-terminal extension of 20 residues that includes a track of six histidines (i.e., a His tag; see UNIT 5.1). Cell are first broken using lysozyme and sonication. The protein is then extracted from inclusion bodies using 8 M guanidine⋅HCl and partially purified by gel filtration in the presence of 4 M guanidine⋅HCl. Further purification by metal-chelate affinity chromatography (MCAC; also see UNIT 9.4) takes advantage of the His tag. The purified protein, which at this stage is unfolded, is then folded by removal of guanidine⋅HCl through dilution into a buffer containing the zwitterionic detergent 3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate (CHAPS). The His tag is removed from the folded protein by digestion with the protease thrombin, and the thrombin is removed by affinity chromatography using immobilized p-aminobenzamidine. A final cleanup of the protein is carried out by gel filtration. Materials E. coli cells expressing HIV-1 integrase: 100 g wet weight from a 3.5-liter fermentation (UNIT 5.3), stored as a flattened paste in a sealed polyethylene bag at −80°C IN break buffer (see recipe), 4°C Suspension buffer: IN break buffer (see recipe) prepared without lysozyme, 4°C IN extraction buffer (see recipe), 4°C 6 × 60–cm Superdex 200 prep grade prepacked gel-filtration column (Pharmacia Biotech) IN column buffer A (see recipe), 4°C Ni-NTA-Sepharose CL-6B MCAC resin (Qiagen) IN column buffer B (see recipe), 4°C IN column buffer C (see recipe), 4°C IN column buffer D (see recipe), 4°C 6 × 60–cm Superdex 75 prep grade prepacked gel-filtration column (Pharmacia Biotech) IN column buffer E (see recipe), 4°C 4 M guanidine⋅HCl/5 mM DTT (4°C) IN folding buffer (see recipe), 4°C 50 mM Tris⋅Cl (pH 7.5 at 4°C)/10 mM CHAPS, 4°C 2000 NIH unit/ml thrombin (see recipe) IN column buffer F (see recipe), 4°C 9 M urea, 4°C IN column buffer G (see recipe), 4°C p-aminobenzamidine immobilized on Sepharose 6B (Pierce, Pharmacia Biotech, or Sigma) 0.1 mM 4-(2-aminoethyl)benzenesulfonyl fluoride (AEBSF; Boehringer Mannheim or ICN Biomedicals)
Folding and Purification of Insoluble Proteins from E. coli
Blender (e.g., Waring; ≥1 liter capacity) 1-liter steel beaker Sonicator, ≥400 W, with sound enclosure (Branson or equivalent) Centrifuge with Beckman J-14 rotor (or equivalent) Tissue homogenizer (e.g., Polytron, Brinkmann) Ultracentrifuge with Beckman 45Ti rotor (or eqivalent) 5 × 50–cm chromatography column with adjustable flow adaptors
6.5.10 Current Protocols in Protein Science
Stirred cells (400-ml and 2-liter capacities) with Diaflo PM 10 ultrafiltration membranes (Amicon) Peristaltic pump Millex-GV 0.22-µm pore size filter units (Millipore) 28°C water bath 1 × 10– to 1 × 20–cm chromatography column Additional reagents and equipment for gel-filtration chromatography (UNIT 8.3) and SDS-PAGE (UNIT 10.1) NOTE: All steps are carried at 4°C unless otherwise stated. Break cells and prepare washed pellets 1. Resuspend 100 g (wet weight) of E. coli cells expressing HIV-1 integrase in 400 ml IN break buffer using a blender. Stir with a magnetic stirrer at room temperature (20° to 25°C) for 30 min. 2. Pour the viscous suspension into a 1-liter steel beaker in an ice bath. Sonicate 20 min at full power with a 60% duty cycle (i.e., on for 0.6 sec, then off for 0.4 sec), with stirring. Be sure that the temperature does not rise above 20°C.
3. Centrifuge 1 hr at 30,000 × g (14,000 rpm in J-14 rotor), 4°C. Resuspend pellet with 500 ml suspension buffer and centrifuge 1 hr at 30,000 × g. Extract washed pellet and purify by gel filtration 4. Dissolve pellet in 40 ml IN extraction buffer using tissue homogenizer. Centrifuge 45 min at 100,000 × g (35,000 rpm in 45Ti rotor), 4°C. 5. Apply the clear supernatant to a 6 × 60–cm Superdex 200 gel-filtration column (UNIT 8.3) equilibrated with IN column buffer A. Elute at 300 ml/hr using IN column buffer A, collecting 15-ml fractions. Assay fractions by SDS-PAGE (UNIT 10.1) and pool those containing IN50-212. IN50-212 should be the major band visible on SDS-PAGE after gel filtration, representing 20% to 50% of the total protein.
Purify IN50-212 by MCAC 6. Apply the IN50-212 sample to a 5 × 50–cm column that has been packed with Ni-NTA-Sepharose CL-6B MCAC resin to a bed height of 5 cm and equilibrated with IN column buffer B at a flow rate of 300 ml/hr. Continue washing column with buffer until absorbance approaches the baseline. 7. Continue washing the column with 5 to 10 column volumes (490 to 980 ml) of IN column buffer C. The absorbance of the column eluate should be close to baseline before the sample is applied. Histidine is bound to the Ni-NTA resin by coordination bonding of the nonprotonated nitrogen atom of the imidazole side chain to Ni2+ ions. Because the pH of the IN column buffer C (6.4) is close to the average pKa of histidine (6.0 to 7.0), only proteins that contain a track of six histidines (i.e., the His tag) are bound strongly enough to be retained. The sample applied to the column should not contain EDTA, and if reductant is required 1 mM 2-mercaptoethanol should be used rather than dithiothreitol. The Ni-NTA matrix is reported to have a binding capacity of ∼5 to 10 mg protein per ml resin.
Purification of Recombinant Proteins
6.5.11 Current Protocols in Protein Science
8. Elute bound protein with a linear pH gradient in a total volume of 980 ml (10 column volumes) beginning at pH 6.4 and ending at pH 4.5: make the gradient by placing 490 ml IN column buffer C (pH 6.4) and 490 ml of IN column buffer D (pH 4.5) in the appropriate reservoirs of the gradient maker. Elute column at ∼300 ml/hr, collect 10-ml fractions and pool those making up the major peak (containing IN50-212; eluted at ∼pH 5.5). After elution, the column can be cleaned by washing with 0.2 M acetic acid/6 M guanidine⋅HCl, then reequilibrated with IN column buffer B. Under the conditions described, the column can be used two or three times before it turns from light blue-green to brownishgray, indicating that regeneration is required. The column can be regenerated by passing 2 column volumes of 100 mM EDTA to strip Ni from the resin, washing the column sequentially with 2 column volumes each of 0.1 M NaOH and 6 M guanidine⋅HCl, and finally passing 2 column volumes of 10 mM NiSO4 through the column to charge the resin with Ni. The column is then washed with water and equilibrated with IN column buffer B. Further details on use of the Ni-NTA resin are given in the manufacturer’s literature (Qiagen, 1992) and in UNIT 9.2.
9. Concentrate pooled IN50-212-containing fractions to ∼40 ml (i.e., ∼8 to 10 mg/ml protein) using a stirred cell with Diaflo PM 10 ultrafiltration membrane. 10. Apply concentrated sample to a 6 × 60–cm Superdex 75 gel-filtration column equilibrated with IN column buffer E. Pool the peak fractions. Reductants (either dithiothreitol or 2-mercaptoethanol) are included in the buffers to maintain the IN50-212 cysteine thiol groups in the reduced state. It should be noted that IN50-212 contains three cysteine residues that do not participate in disulfide bond formation. Pooled fractions may be stored in 10 to 20 ml aliquots at −80°C until ready to use. This gel-filtration step removes minor protein contaminants and metal ions leached from the MCAC column. It also exchanges the protein into fresh buffer containing DTT. Alternatively, the pooled fractions from step 8 can be concentrated by ultrafiltration to ≤1 mg/ml protein, then dialyzed (APPENDIX 3B) against IN column buffer E.
Fold protein by dilution 11. Dilute pooled fractions from step 10 to ∼1 mg/ml protein using 4 M guanidine⋅HCl/5 mM DTT. Pump 30 ml (or other volume as required) of the diluted solution into 1 liter IN folding buffer using a peristaltic pump at a flow rate of ∼3 ml/hr while stirring gently with a magnetic stirrer (see Fig. 6.5.3).
~3 ml/hr flow rate
folding buffer stir bar
HIV-1 integrase
peristaltic pump
Folding and Purification of Insoluble Proteins from E. coli
magnetic stirrer
Figure 6.5.3 Setup for folding of HIV-1 integrase by dilution into buffer.
6.5.12 Current Protocols in Protein Science
After dilution, the final protein concentration will be ∼30 ìg/ml. Protein concentration can be estimated using A280 = 1.46 for 1 mg/ml folded IN50-212 (still containing the His tag) measured in a cuvette with a 1-cm path length.. If A280 is >2.0, dilute with 4 M guanidine⋅HCl in water. The scale of operation at this step is limited by the large dilution of the protein. This step can be scaled up if equipment suitable for concentration of large volumes is available (see annotation to step 12). Otherwise, the step can be repeated if more protein is needed. Protein folding is usually carried out at a low concentration to avoid aggregation, which is thought to arise from partially folded protein in the folding pathway (i.e., folding intermediates; Goldberg et al., 1991; Jaenicke, 1991; also see UNIT 6.4). Because aggregation is an intermolecular event, it is concentration-dependent. In this step the protein concentration is increased very slowly, allowing the protein already in solution to fold before additional unfolded protein is added. It is assumed that folded protein does not participate in aggregation.
12. Concentrate solution to ∼1 mg/ml protein using a stirred cell with Diaflo PM 10 ultrafiltration membrane. Filter concentrated solution (25 to 30 ml volume) through a Millex-GV 0.22-µm filter. If it is not be be used immediately, the concentrated solution can be stored at −80°C for several months. The largest stirred cell produced by Amicon has a 2-liter maximum capacity (i.e., the Model 2000). This cell can be used to concentrate the solution to ∼100 ml, and this must be further concentrated using a smaller cell (e.g., with a 200- or 400-ml capacity). For volumes >2 liters, or for a more rapid process, the Minitan tangential-flow membrane system (Millipore) can be used. This system allows a concentration rate of 0.5 to 1.0 liter/hr.
Cleave histidine tag with thrombin 13. Dilute folded protein from step 12 1:1 with 50 mM Tris⋅Cl (pH 7.5)/10 mM CHAPS (∼0.5 mg/ml final protein concentration). This dilution is carried out because thrombin digestion (in step 14) works best at NaCl concentrations <200 mM.
14. Add 300 µl of 2000 NIH unit/ml thrombin (600 NIH units) to 60 ml of diluted protein from step 13 (30 mg IN50-212). Incubate 30 min at 28°C with occasional mixing. Add an additional 300 µl of 2000 NIH unit/ml thrombin and incubate another 30 min. Cool solution on ice to 4°C and proceed immediately to step 15. Digestion conditions are optimized by conducting small-scale digestions (using <1.0 ml of protein solution from step 13). See Critical Parameters and Troubleshooting for details. The substrate/enzyme ratio used here is ∼0.02% (w/w). Thrombin digestions are more reproducible when carried out on a relatively small scale— e.g., ∼20 to 30 mg protein per batch. If more cleaved protein is required, several batches can be processed concurrently.
Remove excess thrombin by affinity chromatography 15. Pack a 1 × 10–cm column with p-aminobenzamidine immobilized on Sepharose 6B. Equilibrate with IN column buffer F. 16. Add 7.5 ml of 9 M urea (final concentration 1 M) to the 60-ml thrombin digest prepared in step 14. Purification of Recombinant Proteins
6.5.13 Current Protocols in Protein Science
17. Apply digest to equilibrated column from step 15 at a flow rate of 2 ml/min. Collect the column flowthrough, then wash with IN column buffer F until A280 is close to the baseline (this usually requires 1 to 2 column volumes). IN50-212 cannot be separated from thrombin by gel filtration because the IN50-212 (mol. wt. 18,200) undegoes self-association (Hickman et al., 1994), and therefore elutes with a molecular weight close to that of thrombin (36,000). Affinity chromatography using the thrombin inhibitor p-aminobenzamidine is therefore employed. The 1 M urea is added to the sample and column buffers to prevent nonspecific binding of IN50-212 to the column. The affinity column may be regenerated by washing with several column volumes of 6 M guanidine⋅HCl in water to remove the bound thrombin; alternatively, the thrombin can be eluted under native conditions with 50 mM Tris⋅Cl (pH 7.8)/10 mM benzamidine⋅HCl.
18. Pool flowthrough and wash fractions, add 0.1 mM AEBSF, then concentrate to ∼ 2 mg/ml (in ∼10 ml) using a stirred cell with Diaflo PM 10 ultrafiltration membrane. Peform final purification and assay protein 19. Purify protein by gel filtration as in step 10, using IN column buffer G in place of IN column buffer E for equilibration and elution. The column size can be reduced to 2.6 × 60 cm for fractionation of smaller (∼30 mg) quantities of protein; the sample volume should be 5 to 10 ml.
20. Pool IN50-212-containing fractions and filter sterilize with Millex-GV 0.22-µm filter. The final protein solution can be concentrated if required. However, it should be noted that when CHAPS-containing solutions are concentrated by ultrafiltration, the detergent will also be concentrated. The excess detergent can be slowly removed by dialysis (APPENDIX 3B; the critical micellar concentration of CHAPS is 8 mM and its micellar size is ∼8 kDa). Unfortunately, radiolabeled CHAPS is not available for monitoring the removal of that detergent; however, an enzymatic assay is available (Talalay, 1960; Coleman et al., 1979).
21. Measure A280 of protein solution in a cuvette with a 1-cm path length. Native IN50-212 (after the His tag has been cleaved) at a concentration of 1 mg/ml will have an A280 = 1.54.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
BGH break buffer A 100 mM Tris⋅Cl, pH 7.8 (pH as determined at 4°C) 20 mM EDTA (from 0.5 M stock) 10% (w/v) sucrose (100 g/liter) 200 µg/ml lysozyme (200 mg/liter) Prepare immediately before use
Folding and Purification of Insoluble Proteins from E. coli
BGH break buffer B 100 mM Tris⋅Cl, pH 7.8 (pH as determined at 4°C) 10 mM EDTA (from 0.5 M stock) 0.5 mM phenylmethylsulfonyl fluoride (PMSF) 5 mM benzamidine⋅HCl (780 mg/liter) Prepare immediately before use PMSF can be added as 2.5 ml/liter of 0.2 M PMSF stock solution.
6.5.14 Current Protocols in Protein Science
BGH column buffer A 60 mM ethanolamine⋅HCl, pH 9.0 5% (w/v) sucrose (50 g/liter) Prepare immediately before use Ethanolamine (molecular weight 61.08; density 1.02) is a strong base (pKa = 9.4). The pH 9.6 buffer can made by dissolving 3.7 ml liquid ethanolamine free base or 5.9 g ethanolamine⋅HCl salt (molecular weight 97.5) per liter, and adjusting the pH to 9.6 with HCl or NaOH.
BGH column buffer B Solution A: 0.5 M (62 g/liter) anhydrous Na2HCO3 (mol. wt. 84) Solution B: 0.5 M (53 g/liter) anhydrous Na2CO3 (mol. wt. 106) Mix 52 ml solution A and 148 ml solution B and dilute to 1 liter with water. The solution may also be made directly by dissolving 7.8 g NaHCO3 and 2.2 g Na2CO3. The 0.5 M stock solutions may be stored at least 1 month at 4°C.
BGH extraction buffer To 420 ml H2O add (in the order indicated): 17.2 ml 150 g/liter (2 M) glycine stock (50 mM final; store stock at 4°C) 7.8 ml 2 M NaOH 7.7 g reduced glutathione (GSH) or 50 ml 0.1 M GSH stock (see recipe; 5 mM final) 764 g guanidine⋅HCl (8 M final) H2O to 1 liter Prepare immediately before use Because of the volume increase on addition of guanidine⋅HCl (i.e., 1 g increases the volume 0.76 ml), buffer components must be added to 420 ml water and total volume adjusted to 1 liter at the end. The buffer should be pH 9.6 as determined at 4°C; if it is not, it can be adjusted with 2 M NaOH or 2 M HCl. If solid glutathione (which is acidic) is added, pH will require adjustment. If high-quality guanidine⋅HCl is used, the solution will be colorless and clear (see APPENDIX 3A).
BGH folding buffer A To ∼500 ml H2O add (in the order indicated): 17.2 ml of 150 g/liter (2 M) glycine stock (50 mM glycine final; store stock at 4°C) 7.8 ml 2 M NaOH 100 g sucrose (10% w/v final) 2 ml 0.5 M EDTA (1 mM final) 310 mg reduced glutathione (GSH) or 10 ml of 0.1 M GSH stock (see recipe; 1 mM final) 60 mg oxidized glutathione (GSSG) or 1 ml of 0.1 M GSSG stock (see recipe; 0.1 mM final) 240 g urea (4 M final) Add 4°C H2O to 1 liter Prepare immediately before use The buffer should be pH 9.6 as determined at 4°C; if it is not it can be adjusted with either 2 M NaOH or 2 M HCl. Because of the volume increase on addition of urea (i.e., 1 g increases the volume 0.76 ml), buffer components must be added to ∼500 ml and total volume adjusted to 1 liter at the end. There is no need to deionize urea if ultrapure-grade material is used.
Purification of Recombinant Proteins
6.5.15 Current Protocols in Protein Science
BGH folding buffer B 60 mM ethanolamine⋅HCl, pH 9.6 (pH as determined at 4°C) 10% (w/v) sucrose (100 g/liter) 1 mM EDTA (from 0.5 M stock) 0.1 mM reduced glutathione (GSH) 0.01 mM oxidized glutathione (GSSG) Prepare immediately before use GSH and GSSG can be added from stock solutions (1 ml of 0.1 M GSH and 0.1 ml of 0.1 M GSSG per liter) or as solids. If solid glutathione (which is acidic) is added, pH may require adjustment. Ethanolamine (molecular weight 61.08; density 1.02) is a strong base (pKa = 9.4). The pH 9.6 buffer can made by dissolving 3.7 ml liquid ethanolamine free base per liter or 5.9 g ethanolamine⋅HCl salt (molecular weight 97.5) per liter, and adjusting the pH to 9.6 with HCl or NaOH. It can be stored several days at 4°C.
BGH wash buffer A 100 mM Tris⋅Cl, pH 7.8 (pH as determined at 4°C) 2% (w/v) Triton X-100 (20 g/liter) 4 M urea (240 g/liter) 5 mM EDTA (from 0.5 M stock) 0.5 M DTT (77 mg/liter) Prepare immediately before use When preparing solution, allow for volume increase on addition of urea; 1 g urea will increase solution volume by 0.76 ml. There is no need to deionize urea if ultrapure-grade material is used. Triton X-100 is a viscous liquid at 20° to 25°C; for the use intended here, there is no need for highly purified Triton.
BGH wash buffer B 100 mM Tris⋅Cl, pH 7.8 (pH as determined at 4°C) 0.5 mM DTT (77 mg/liter) Prepare immediately before use It is preferable to add DTT as a solid, but it may also be added as 0.2 M stock in water (30.8 g/liter; store at or below −20°C; stable ≥1 month).
Glutathione stock solutions, 0.1 M Reduced glutathione: Dissolve 30.73 mg/ml reduced glutathione (GSH) in water and adjust pH to 7 with NaOH. Store at or below −20°C (stable ≥1 month). Oxidized glutathione: Dissolve 61.3 mg/ml oxidized glutathione (GSSG) in water and adjust pH to 7 with NaOH. Store at or below −20°C (stable ≥1 month). hIL-2 break buffer 0.1 M Tris⋅Cl, pH 7.5 (pH as determined at 4°C) 50 mM EDTA (from 0.5 M stock; APPENDIX 2E) Prepare immediately before use
Folding and Purification of Insoluble Proteins from E. coli
IN break buffer 50 mM Tris⋅Cl, pH 7.5 (pH as determined at 4°C) 5 mM EDTA (from 0.5 M stock) 5 mM benzamidine⋅HCl (780 mg/liter) 5 mM DTT (770 mg/liter) 0.1 mg/ml lysozyme (100 mg/liter) Prepare immediately before use It is preferable that DTT be added as a solid, but it may also be added as 0.2 M stock in water (30.8 g/liter; store at or below −20°C; stable ≥1 month).
6.5.16 Current Protocols in Protein Science
IN column buffer A 10 mM Tris⋅Cl, pH 7.5 (pH as determined at 4°C) 4 M guanidine⋅HCl (382 g/liter) 1 mM 2-mercaptoethanol (2-ME; 0.070 ml/liter) Because of the volume increase on addition of guanidine⋅HCl (i.e., 1 g increases the volume 0.76 ml), buffer components must be added to 420 ml water and total volume adjusted to 1 liter at the end. Concentrated 2-ME (14.3 M) should be stored at 4°C and opened bottles replaced every 2 or 3 months.
IN column buffer B To 500 ml H2O add: 10 ml 1 M Tris⋅Cl, pH 8.0 (pH as determined at 4°C; 10 mM final) 0.73 g NaH2PO4⋅H2O (monobasic; mol. wt. 137.99) 13.44 g Na2HPO4 (dibasic; mol. wt. 141.96) 0.07 ml 2-mercaptoethanol (2-ME; 1 mM final) After salts have dissolved add: 573.2 g guanidine⋅HCl (6 M final) Bring temperature to 4°C and add 4°C H2O to 1 liter Adjust pH to 8.0 (as determined at 4°C) if necessary with 2 M NaOH or 2 M HCl Prepare immediately before use Because of the volume increase on addition of guanidine⋅HCl (i.e., 1 g increases the volume 0.76 ml), buffer components must be added to ∼500 ml and total volume adjusted to 1 liter at the end. Concentrated 2-ME (14.3 M) should be stored at 4°C and opened bottles replaced every 2 or 3 months.
IN column buffer C Titrate IN column buffer B (see recipe) to pH 6.4 (as determined at 4°C) with HCl. Prepare immediately before use. IN column buffer D Prepare 100 mM (13.8 g/liter) NaH2PO4⋅H2O. If necessary adjust to pH 4.5 with 2 M NaOH or 2 M HCl. IN column buffer E Immediately before use, prepare IN column buffer B (see recipe), replacing the 2-ME with 5 mM DTT. It is preferable that DTT be added as a solid, but it may also be added as 0.2 M stock in water (30.8 g/liter; store at or below −20°C; stable ≥1 month).
IN column buffer F 50 mM Tris⋅Cl, pH 7.5 (pH as determined at 4°C) 0.25 M NaCl (14.6 g/liter) 10 mM 3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate (CHAPS; mol. wt. 614.9; 6.2 g/liter) 1 mM DTT (154 mg/liter) 1 mM EDTA (prepare from 0.5 M stock) 1 M urea (60 g/liter) Prepare immediately before use It is preferable that DTT be added as a solid, but it may also be added as 0.2 M stock in water (30.8 g/liter; store at or below −20°C; stable ≥1 month).
IN column buffer G Prepare IN column buffer F (see recipe), but omit urea.
Purification of Recombinant Proteins
6.5.17 Current Protocols in Protein Science
IN extraction buffer 10 mM Tris⋅Cl, pH 7.5 (pH as determined at 4°C) 8 M guanidine⋅HCl (764 g/liter) 5 mM DTT (770 mg/liter) Prepare immediately before use Because of the volume increase on addition of guanidine⋅HCl (i.e., 1 g increases the volume 0.76 ml), buffer components must be added to 420 ml and total volume adjusted to 1 liter at the end. If high-quality guanidine⋅HCl is used, the solution will be colorless and clear (see APPENDIX 3A). It is preferable that DTT be added as a solid, but it may also be added as 0.2 M stock in water (30.8 g/liter; store at or below −20°C; stable ≥1 month).
IN folding buffer 50 mM Tris⋅Cl. pH 7.5 (pH as determined at 4°C) 0.5 M NaCl (29.2 g/liter) 10 mM 3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate (CHAPS; mol. wt. 614.9; 6.2 g/liter) 2 mM DTT (308 mg/liter) Prepare immediately before use It is preferable that DTT be added as a solid, but it may also be added as 0.2 M stock in water (30.8 g/liter; store at or below −20°C; stable ≥1 month).
RP-HPLC solvents A, B, and C Solvent A: Prepare 0.1% (v/v) acetonitrile in water (both HPLC grade; e.g., Sigma). Solvent B: Prepare 0.1% (v/v) TFA in acetonitrile (both HPLC grade) Solvent C: Mix 800 ml acteonitrile and 1 ml TFA (both HPLC grade). Dilute with HPLC-grade water to 1 liter. Prepare all solutions fresh; filter and degas before use. Thrombin, 2000 NIH units/ml Immediately before use reconstitute contents of a 1030 NIH-unit vial (2100 NIH units/mg) of thrombin from human plasma (Sigma) with 0.5 ml water to give a solution containing 50 mM sodium citrate, pH 6.5, 0.15 M NaCl, and 2000 NIH units/ml thrombin. The A280 of 1 mg/ml thrombin is 1.83 in a cuvette with a 1-cm path length. Store the lyophized powder at or below −20°C.
COMMENTARY Background Information
Folding and Purification of Insoluble Proteins from E. coli
Bovine growth hormone (BGH) There has been much commercial interest over the past fourteen years in producing large quantities of recombinant growth hormones. Human growth hormone is used as a pharmaceutical agent for treating dwarfism. Nonhuman growth hormones, including bovine and porcine growth hormones, are used in agriculture for stimulating growth and lactation in animals. These hormones were among the first group of proteins pursued by the biotechnology industry; they had been investigated decades before by protein chemists.
Bovine growth hormone (BGH; also known as bovine somatotropin) is a pituitary hormone, and the mature protein contains 191 residues (mol. wt. 21,876) with two disulfide bonds— one between residues 51 and 163 and another between residues 180 and 188. The closely related porcine growth hormone (pGH) was the first growth hormone for which the three-dimensional structure was determined (AbdelMeguid et al., 1987). Since then, the structure of the human growth hormone (hGH) as a complex with its receptor has also been determined (deVos et al., 1992). It should be noted that hGH and pGH share 68% and 91% sequence similarity, respectively, with BGH.
6.5.18 Current Protocols in Protein Science
In Basic Protocol 1, conditions are chosen that allow the unfolded, reduced protein to be folded and oxidized at relatively high protein concentrations (∼1 to 2 mg/ml). The procedure is based on the original work of Goldberg (Orsini and Goldberg, 1978; see also references therein), who pioneered the use of cosolvents (e.g., urea in Basic Protocol 1; also see UNIT 6.4 and Maeda et al., 1994) to help maintain solubility during refolding. The use of redox buffer systems containing low-molecular-weight thiol-disulfide pairs (e.g., reduced and oxidized glutathione) to enhance the rate of disulfidebond formation during the folding of reduced protein is based on Wetlaufer’s work (Wetlaufer, 1984; see also references therein). Although the methodology of Basic Protocol 1 was developed in 1982-1983, it remains a fairly typical example of a technique for preparative protein folding. Details of BGH expression in E. coli can be found in Wingfield et al. (1987a). BGH folding and thiol oxidation The structure of both porcine and human growth hormone consists of four antiparallel α-helices (Abdel-Meguid et al., 1987). The topology of BGH is almost certainly similar to that of pGH, due to the high degree of sequence and functional similarity. In pGH, one disulfide bond is formed between Cys-53 (found in the middle of the region connecting helices 1 and 2) and Cys-164 (located in helix 4). The other disulfide bond links the C-terminal region (at Cys-189) to helix 4 (at Cys-181). Examination of the solution conditions for folding and oxidation of the reduced pituitaryderived BGH has shown that reduced and unfolded protein—in 6 M guanidine⋅HCl diluted into 4.5 M urea—can be air-oxidized with a high (90%) yield (Holzman et al., 1986). These conditions are similar to those used in Basic Protocol 1, except that in the latter, the protein concentration is >5-fold higher (>1 mg ml) and a redox buffer is used. For further information on in vitro folding of BGH and the closely related pGH, consult Brems and Havel (1989) and Bastiras and Wallace (1992). Havel et al. (1989) present a concise and useful review of the some of the spectroscopic methods used to monitor protein conformation and structure (e.g., UV absorption, circular dichroism, and fluorescence) with examples of studies on BGH.
Comparison of authentic and recombinant BGH Natural BGH is purified from bovine pituitaries, and examples of purifications leading to crystallizable protein are described by Bell et al. (1985) and Spitsberg (1987). The method that appears to give the most homogeneous product (Wood et al., 1989) involves extraction of pituitaries with 4.5 M urea (pH 8.8) followed by purifications employing anion- and cationexchange chromatography in the presence of urea. The urea maintains solubility of the protein during purification; at the concentration used (4.5 M) it does not perturb or unfold the protein. The urea is finally removed from the purified protein by dialysis at pH 10. A single bovine pituitary yields ∼16 mg of pure protein by this method. Preparations of the natural protein often show varying degrees of N-terminal heterogeneity, depending on the method of purification. This may involve truncations of either one (∆A- 1) or four (A-4) residues—e.g., (Ala)PheProAlaMetSerLeuSerGlyLeuCys AlaPhe…—where the residue in parentheses is lost in a ∆ A-1 truncation and the four boldface residues are lost in a A-4 truncation. An example of a A-4 truncation is shown in Figure 6.5.1, lane E, where the two closely migrating bands at 23 and 22 kDa correspond to full-length and N-terminal-deleted (A-4) protein, respectively (for further details see Wingfield et al., 1987b, and references therein; also see Langley et al., 1987a). The two forms can be resolved under native conditions by chromatofocusing, which takes advantage of the small difference in their isoelectric points; full-length and A-4 BGH have pI values of 8.2 and 8.0, respectively (Wingfield et al., 1987b). Protein purified by the method of Wood et al. (1989) has a single and correct N-terminus. Recombinant hormones have been expressed that contain the full BGH sequence (e.g., Langley et al., 1987a), as well as others with the ∆A-1 deletion (e.g., Wingfield et al., 1987a; Bogosian et al., 1989) and the A-9 deletion (Wingfield et al., 1987a). Purified A-1 BGH retains the initiating Met at the N-terminus (MetPhePro...), but when the initiating Met is placed in front of the Ala, the Met is removed to give the authentic sequence (AlaPhePro...). Deletion of at least the first nine N-terminal residues does not appear to effect the biological activity of BGH. Another source of chemical heterogeneity in both authentic and recombinant growth hor-
Purification of Recombinant Proteins
6.5.19 Current Protocols in Protein Science
mones is the deamidation of specific Asn and Gln residues (Secchi et al., 1986). This is partly responsible for the multiple band patterns observed in isoelectric focusing gels (Wingfield et al., 1987a; 1987a). In typical authentic and recombinant BGH preparations, ∼20% of the protein is deamidated. However, because these modified forms have lower pIs than unmodified protein, they can be separated by chromatofocusing (Wingfield et al., 1987a). The cationexchange chromatography step (at pH 6.9 in 4.5 M urea) used by Wood et al. (1989) for purification of pituitary BGH also appears to separate deamidated from unmodified protein. Deamidated protein is more positively charged than unmodified protein, and will therefore bind more tightly to a negatively charged cation-exchange matrix. Authentic and recombinant BGH both exhibit self-association with the characteristics of a rapid monomer-dimer equilibrium. The dissociation constant (Kd) for authentic BGH is 6.6 × 10−6 M (Fernandez and Delfino, 1983), which is similar to that determined for both the A-1 and A-9 recombinant forms of BGH (P.T.W., unpub. observ.). At concentrations of 1.0 and 0.1 mg/ml, ∼76% and 42% of the protein will be dimeric, respectively. Under physiological conditions, the monomer appears to be the active species. Monomeric protein binds to two molecules of the extracellular domain of the cell-surface receptor, forming a 1:2 molar complex analogous to the human growth hormone (Staten et al., 1993).
Folding and Purification of Insoluble Proteins from E. coli
Other approaches used to extract and fold recombinant BGH Langley et al. (1987a) extracted protein from inclusion bodies using 6 M guanidine⋅HCl (pH 8.0) as the denaturant. The extract was allowed to oxidize in air >72 hr at room temperature in the presence of the denaturant. Oxidized protein was fractionated by gel filtration in guanidine⋅HCl, the fractions making up the monomer peak pooled, and the protein folded by removal of denaturant using dialysis. The rationale for this method is that, in a protein with four cysteine residues that form two disulfide bonds (such as BGH), there are three possible intramolecular disulfide linkage combinations. Hence, even if the oxidation proceeds in a random manner as expected for unfolded protein, at least 33% of the protein produced will have the correct pattern. In practice, the recovery of correctly folded oxidized protein was significantly higher than 33%—suggest-
ing that even in 6 M guanidine⋅HCl, BGH may contain some elements of native-like structure. This method is geared toward large-scale protein production and gives good protein recovery. However, the long incubation period in guanidine⋅HCl appears to result in ∼50% of the protein being deamidated (Langley et al., 1987b). Bogosian et al. (1989) used 4.5 M urea at pH 10.7 to extract BGH from inclusion bodies. The extract was then stirred for 48 hr, allowing the protein to oxidize in air. This method is mentioned because it results in formation of a novel concatenated dimer, formed by the interlocking of disulfide loops (Violand et al., 1989). It is most likely that under the conditions used for extraction, the protein folding and oxidation is initiated from partially folded associated protein. Ovine growth hormone (which is closely related to BGH) was folded and purified with an overall recovery of 30% by a procedure similar in approach to Basic Protocol 1 (Wallis and Wallis, 1989). This method, however, used 2 M guanidine⋅HCl instead of urea as the cosolvent to maintain solubility during folding. It also employed air oxidation for 24 hr instead of the redox buffer system. Interleukin 2 Interleukin 2 (IL-2) is a member of the cytokine family. It is composed of a compact core bundle of four antiparallel α-helices (Bazan, 1992; McKay, 1992). IL-2 plays important roles in the proliferation and differentiation of T lymphocytes as well as in regulation of the immune system. The biological effects of IL-2 are mediated via binding to IL-2 receptors (Minami et al., 1993). The human IL-2 (hIL-2) gene has been cloned and expressed in E. coli as insoluble inclusion bodies (Devos et al., 1983). The purification of recombinant hIL-2 has been described and the protein thoroughly characterized (Liang et al., 1985; Kato et al., 1985; Weir and Sparks, 1987). Basic Protocol 2 details the solubilization, refolding, and purification of recombinant hIL-2. This procedure can be scaled up to yield hIL-2 on the scale of hundreds of milligrams. hIL-2 has three cysteines; hence, three possible disulfide-linked forms can exist. The native isomer consists primarily of one form— i.e., the one that contains a disulfide linkage between Cys-58 and Cys-105 (Robb et al., 1984; Wang et al., 1984; Liang et al., 1985).
6.5.20 Current Protocols in Protein Science
Reversed-phase-HPLC has been used to separate these isomers (Browning et al., 1987; also see Fig. 6.5.3). HIV-1 integrase An early part of the life cycle of all retroviruses (including the human immunodeficiency virus, HIV) is the integration of a DNA copy of the viral genome into the host chromosome. This step is essential for viral replication. Retroviral DNA integration is carried out by a defined set of DNA cutting and joining reactions, catalyzed by integrase protein (Katz and Skalka, 1994). The HIV-1 integrase is a 288-residue protein (mol. wt. 32,200) which has been expressed in E. coli (Sherman and Fyfe, 1990). The protein is located in the pellet obtained by low-speed centrifugation following cell breakage, but is apparently not highly aggregated into inclusion bodies, as it can be extracted into high-ionicstrength solution (e.g., 1 M NaCl) without a denaturant. The insolubility of the protein is probably a result of nonspecific binding to E. coli nucleic acid. The salt-extraction procedure was originally used by Terry et al. (1988) for extraction of avian sarcoma-leukosis virus integrase expressed in E. coli. Dissection of the HIV-1 integrase by preparation of a series of deletion mutants (Bushman et al., 1993; see also references therein) demonstrated that a central core region of residues 50 to 212 was enzymatically active, carrying out a subset of the reactions catalyzed by the full-length enzyme. This deletion mutant is interesting for structural studies as it has better solubility than the full-length enzyme, which is notoriously difficult to handle as a result of limited solubility in the usual aqueous solvents. The HIV-1 integrase deletion mutant (IN50-212) is expressed in E. coli as a fusion protein with the N-terminal extension sequence GlySerSerGlyHisHisHisHisHisHisSerSer GlyLeuValProArgGlySerHisMet. This sequence (a His tag) contains a six-residue histidine repeat which is responsible for the selective high-affinity binding of the fusion protein to a nickel-chelate column. The boldface portion of the His tag sequence indicates the location of the specific thrombin-cleavage site between Arg-16 and Gly-17, which is exploited in removal of the tag using thrombin. In Basic Protocol 3 the His-tagged IN50-212 is expressed as an insoluble (i.e., inclusionbody-type) protein, and is purified under denaturing conditions, taking advantage of the fact that metal-chelate affinity chromatography
(MCAC) can be carried out in the presence of protein denaturants such as guanidine⋅HCl. Following purification, the protein is folded and the His tag removed by exploiting the specific thrombin-cleavage site of the tag sequence (underlined above). The cleaved protein has the N-terminal sequence GlySerHisMetHisGlyGln.... The Met at position 4 corresponds to Met 50 of the wild-type integrase and residues 1 to 3 (GlySerHis) are derived from the tag. Excess thrombin is removed by affinity chromatography using an immobilized inhibitor (p-aminobenzamidine). In the last step of the purification, the IN50-212 is subjected to gel-filtration to remove any aggregated multimers. The protein produced is 166 residues long, the molecular weight estimated from the DNA coding sequence is 18,200, and the calculated isoelectric point is ∼7. Although the folded deletion mutant has enhanced solubility as compared with the wildtype enzyme, the zwitterionic detergent CHAPS is included in buffers to maintain solubility, especially during protein folding. Following protein purification the detergent can be removed or exchanged for other buffer additives. Substitution of Phe-185 for a Lys residue appears to increase the solubility of IN50-212. This mutant protein, expressed in E. coli with the N-terminal His tag, was extracted with 1 M salt and purified by MCAC. The extraction with high-ionic-strength salt solution in the absence of denaturant is analogous to the procedure used with the full-length protein, discussed above. The crystal structure of this protein has been determined at 2.5-Å resolution. The overall topology consists of five β-sheets flanked by helical regions, showing that the integrase domain belongs to a superfamily of polynucleotidyl transferases that includes ribonuclease H (Dyda et al., 1994).
Critical Parameters Bovine growth hormone (BGH) Protein extraction. The protein should be extracted from the inclusion bodies in a monomeric and fully reduced state. This provides a defined starting point from which to develop a reproducible folding protocol. Based on pilot-scale experiments in which washed pellets (from step 4 of Basic Protocol 1) were extracted with various protein denaturants (P.T.W., unpub. observ.; also see UNIT 6.3), 8 M guanidine⋅HCl was chosen for solubilization of aggregated BGH. The most effective ones were
Purification of Recombinant Proteins
6.5.21 Current Protocols in Protein Science
Supplement 4
Folding and Purification of Insoluble Proteins from E. coli
urea and guanidine⋅HCl at 6 M to 8 M concentrations, each of which solubilized >80% of the protein. The extracts were further analyzed by gel filtration on Sephacryl S-300 in the presence of the same concentration of the particular denaturant used for the initial extraction. The results indicated that guanidine⋅HCl at concentrations >6 M yielded solubilized protein of which >80% was monomeric. Although 8 M urea also solubilized >80% of the protein, only ∼20% of this protein was monomeric, the remainder consisting of dimers and higher-order aggregates. Protein folding and oxidation. Basic Protocol 1 describes an empirical approach based on some of the principles discussed in UNITS 6.1 & 6.4. The determination of basic physicochemical/conformational properties for a particular protein will take some of the guesswork out of developing a suitable folding protocol. For BGH, purified authentic protein from bovine pituitary glands was available, and had been fairly well characterized before recombinant proteins were produced. This situation is not common, as the natural counterparts of many recombinant proteins are rare and may never have been purified. The optimal ratio of reduced to oxidized glutathione in the folding buffers used in Basic Protocol 1 was empirically determined. However, it is commonly observed that the highest folding yields are obtained when the ratio of GSH/GSSG is between 5 and 10 and the total reduced and oxidized glutathione concentration is in the range of 1 to 10 mM. Deamidation. As mentioned above, BGH is susceptible to deamidation. Although deamidated forms (charge isomers) can be resolved by ion-exchange methods (Wingfield et al., 1987a), their formation should be minimized. This is achieved by limiting the amount of time the protein spends in the fully unfolded state, especially at pH 9.5. For example, the guanidine⋅HCl extract should be processed immediately and diluted as described in step 6 of Basic Protocol 1. The dialysis steps (at pH 9.5) should be performed for the times indicated, not longer. The folded protein is stable and does not appear to deamidate spontaneously on storage. Scaleup. For larger-scale production, a Manton-Gaulin-APV homogenizer (UNIT 6.1) is used to break cells instead of the French press. In addition, a larger Sephadex G-100 column (e.g., 5.0 × 90–cm) is used in step 12 of Basic Protocol 1.
Interleukin-2 Protein purification. Recombinant hIL-2 should be extracted as a monomer (see Basic Protocol 2, step 5). The extraction has been tested with various concentrations of acetic to determine the optimal condition; 20% acetic acid has been found to extract the maximum amount of monomeric hIL-2 from inclusion bodies. However, in the gel-filtration step (see Basic Protocol 2, step 8), 20% acetic acid is too corrosive for the metal parts of the chromatography system (e.g., the stands, pump, and fraction collector); thus 10% acetic acid is recommended. Protein folding. The hIL-2 monomer that elutes from the Sephadex G-100 column (see Basic Protocol 2, step 8) yields two peaks upon analytical reversed-phased chromatography (see Support Protocol). These two peaks represent the refolded, oxidized hIL-2 and unfolded, reduced hIL-2. The proportion of the oxidized form is increased by the dialysis step (see Basic Protocol 2, step 9). The time and volume recommended in the dialysis step have been determined empirically to produce the best yield of oxidized hIL-2. Longer periods of dialysis (e.g. 48 hr) causes do not increase the yield of oxidized hIL-2. Scaleup. The parameters for larger-scale production are the same as those for BGH (see above). HIV-1 integrase MCAC. It is important to use low concentrations of 2-mercaptoethanol in the sample and column buffer (1 mM is safe) and to completely avoid dithiothreitol, as either reductant will strip Ni2+ from the MCAC matrix (see UNIT 9.4). Protein folding. Because HIV-1 integrase is not a very soluble protein and has a high tendency toward self-association and aggregation (Hickman et al., 1994), the solubility and stability of the folded protein must be maintained by including a relatively high salt concentration as well as the detergent CHAPS in the folding buffer and in all subsequent column buffers. The buffer additives indicated for HIV-1 integrase are usually not required for folding of the average protein. Thrombin cleavage. It is necessary to perform pilot-scale experiments to optimize the conditions for removing the His tag using thrombin. The usual parameters to vary are enzyme/substrate ratios and incubation time and temperature. Thrombin is a serine protease, and can be irreversibly inhibited with either PMSF or AEBSF. Removal of the tag can be
6.5.22 Supplement 4
Current Protocols in Protein Science
monitored by SDS-PAGE (UNIT 10.1), in which case the thrombin should be quenched with 1 mM PMSF or AEBSF prior to addition of SDS and heating. There is an ∼2-kDa mass difference between tagged and nontagged protein (see Fig. 6.5.2). The digestion should be as complete as possible at the specific cleavage site (see Background Information); overdigestion may result in nonspecific cleavage at other sites. It is also important to avoid denaturation of the (thrombin) (e.g., by vigorous stirring or overheating), as partially denatured proteases often exhibit some loss of specificity, cleaving at unpredictable sites.
Troubleshooting Bovine growth hormone (BGH) Most precautions and guidelines for folding and oxidation are given in annotations to the individual steps of Basic Protocol 1. It should be cautioned that, when using a mixture of oxidized and reduced glutathione for oxidizing proteins that contain both free and disulfidelinked cysteines in the native state (e.g., IL-2), the potential exists for glutathionylation, in which the unpaired cysteine(s) form a mixed disulfide with glutathione. This modification will cause charge heterogeneity, and can be readily detected by electrospray ionization mass spectrometry (ESI-MS), which will indicate a mass increase of 305.3 for each GSH moiety incorporated. Interleukin-2 Low-molecular-weight impurities extracted together with monomeric hIL-2 in acetic acid (see Basic Protocol 2, step 5) are usually eluted earlier than the oxidized form of hIL-2 in RPHPLC (see Basic Protocol 2, step 12) and can thus be removed. If the RP-HPLC peak containing the oxidized form of hIL-2 is contaminated by low-molecular-weight impurities, SDSPAGE analysis of each fraction composing the peak is recommended to avoid pooling fractions containing impurities. Additional information regarding troubleshooting is given in annotations to individual steps of Basic Protocol 2. HIV-1 integrase MCAC. Protein will not bind to the column if the His tag has been nonspecifically degraded or removed by E. coli proteases. SDS-PAGE (UNIT 10.1) of the cell extracts and fractions during the purification should indicate if the tag is present. The His tag adds ∼2 kDa to the
apparent mass of the protein. The denaturant concentration should be high enough to maintain solubility of the protein during chromatography. Insoluble or aggregated protein will not bind to the column and will be located in the column flowthrough or will clog the top of the matrix. If protein aggregates on the column, a higher concentration of denaturant should be tried, although aggregation should not happen using 6 M guanidine⋅HCl in column buffer A. Further details on troubleshooting the use of the Ni-NTA resin are given in the manufacturer’s literature (Qiagen, 1992). Thrombin cleavage. If undigested (Histagged) protein still remains after digestion, the standard procedure is to reapply the mixture to an MCAC column, whereupon the matrix will specifically bind the unprocessed protein containing the His tag. The processed (cleaved) protein is not bound and elutes in the column flowthrough. This approach, although satisfactory for monomeric proteins, is complicated with multimeric proteins and those that exhibit reversible self-association—e.g., HIV-1 integrase. For example, in a partially digested dimeric protein, the subunits can be arranged in three possible configurations—TT, TM, and MM—where T is still tagged and M has had the tag cleaved. It may be possible to separate TT and TM by gradient elution from the MCAC column, as the former might be expected to bind more tightly than the latter. A simpler approach is to use solvent conditions that dissociate the protein subunits without denaturing them. For HIV-1-integrase, this can be achieved with 2 to 3 M urea.
Anticipated Results BGH The purification of ∆ A-9 BGH is summarized in Table 6.5.1. About 90 to 100 mg of protein are obtained from 50 g (wet weight) of cells with an overall yield in the range 15% to 25%. For larger and/or multidomain proteins, much lower yields (1% to 5%) may be more typical. Gel analysis of the BGH in cell extracts and in purified protein is shown in Figure 6.5.1. Interleukin-2 If the expression of hIL-2 in E. coli is ∼10%, 20 g of cell pellet will yield 4 to 5 mg of purified protein, with a specific activity of 5 × 106 U/mg. The purity of the hIL-2 should be >98% as indicated by analytical C18 column chromatography (see Support Protocol).
Purification of Recombinant Proteins
6.5.23 Current Protocols in Protein Science
Table 6.5.1
Purification of a Recombinant Bovine Growth Hormone Analoga
Total Proteinc (mg)
Stage of Purificationb Cells Washed pellet (step 4) Dialysis supernatant (step 8) DEAE-Sepharose pool (step 11) Sephadex G-100 pool (step 14)
5000 375 200 131 95
Specific BGH contentd (%) 7.5 60 75 95 99
Total BGH (mg) 375 225 150 125 94
Yield (%) 100 60 40 33 25
aThe summary refers specifically to a biologically active analog of BGH (δ-9 BGH) in which the full-length sequence is
truncated at the N-terminus by eight residues and serine is substituted for glycine at the first position. Similar relative yields were obtained with a δ-1 analog in which the N-terminal Ala of the native sequence was replaced by Met, but the amount of this protein expressed in E. coli was several-fold lower than that obtained for δ-9 BGH (Wingfield et al., 1987a) bThe numbers in parentheses refer to Basic Protocol 1 steps. cDeterminations were made using the Bio-Rad Protein Assay Kit, except those for the DEAE and Sephadex pools, which
were made by UV absorbance measurements. dPercentage of total protein that is BGH. Estimates made by densitometric scanning of Coomassie blue stained
SDS-polyacrylamide gels.
a
b
c
d
In a scaled-up process (see Critical Parameters), 25 to 30 mg of purified hIL-2 may be obtained from 200 g of E. coli cell pellet.
Folding and Purification of Insoluble Proteins from E. coli
HIV-1 integrase Figure 6.5.4 shows results of SDS-PAGE of the purified IN50-212 before (lane B) and after (lane A) removal of the His tag by thrombin digestion. The molecular weight difference of ∼2000 kDa corresponds to the removal of the
Figure 6.5.4 Results of SDS-PAGE of H1V-1 integrase50-212 on a 12.5% polyacrylamide gel stained with Coomassie blue. Lane A, recombinant HIV-1 integrase50-212; lane B, recombinant HIV-1 integrase50-212 with N-terminal His tag; lane C, extract of HIV-1 integrase–expressing E. coli cells used for purification.
16 amino acid residues comprising the N-terminal His tag. The overall recovery of purified protein is ∼30%. For example, from 1 g (wet weight) of cells (containing ∼5% IN50-212) ∼1.5 mg of purified protein are obtained (see Figure 6.5.4, lane C, for SDS-PAGE analysis of whole IN50212–expressing E. coli). Most of the steps, including protein folding, have high recoveries (80% to 95%). The yield of thrombin-digested
6.5.24 Current Protocols in Protein Science
protein from purified tagged protein is between 60% to 70%. The enzymatic and biophysical properties of the protein are detailed in Hickman et al. (1994).
Time Considerations BGH Basic Protocol 1 for the purification of BGH takes 5 days. Protein can be stored at −80°C at the end of step 4 (as washed pellets) or step 10 (as pooled fractions from ion-exchange). Day 1: Cell breakage, preparation of washed pellets and extraction with guanidine⋅HCl (steps 1 to 5) requires 1⁄2 day of work. The protein extract is then dialyzed overnight (step 5). Day 2: After changing the dialysis buffer (step 7), dialysis is continued at least 6 hr. At this step the dialysis can be left overnight or directly processed by ion-exchange chromatography (steps 8 to 10), which takes 6 to 8 hr and can be run overnight. Day 3: Ion-exchange chromatography is run, or if ion-exchange was performed on day 2, the pooled fractions are concentrated for several hours (step 11), then applied to the gel-filtration column and chromatographed for several hours (steps 12 and 13). The gel-filtration column is run is overnight if a low-pressure matrix (e.g., Sephadex G-100) is used or on the same day if a medium-pressure matrix (e.g., Superdex 200) is used. Day 4: The protein is analyzed (e.g., by SDS-PAGE or isoelectric focusing), concentrated if required (step 14), and frozen or prepared for lypohilization. Interleukin-2 In Basic Protocol 2, preparation and lysis of the cells (steps 1 and 2) will take 1 hr, cell breakage and preparation of washed pellets (steps 3 and 4) will take 3 hr, acid extraction (steps 5 and 6) will take 1 hr, Sephadex G-100 column chromatography (step 7) will take 12 hr, dialysis of the pooled fractions will be carried out overnight, and RP-HPLC (steps 9 to 12) will require 7.5 hr. The final dialysis is run overnight. In the Support Protocol for resolution of native and misfolded forms of hIL-2, it will take ∼1 hr to run a sample after the blank runs to establish the baseline profile have been completed.
HIV-1 integrase Basic Protocol 3 is usually carried in two stages. In stage 1, purification of the unfolded protein that still contains the His tag (steps 1 to 8) is usually carried out on a relatively large scale (using 100 g of cells). If the Superdex 200 column used in step 5 and the MCAC column used in step 6 are run using the Pharmacia Biotech Biopilot system, this stage will take 3 to 4 days to complete. It will take longer if low-pressure columns are used. In stage 2, protein folding (steps 11 and 12), removal of the His tag by thrombin cleavage (steps 13 and 14), affinity chromatography (steps 15 to 17), and gel filtration (step 19) are performed. This stage is carried out repeatedly with relatively small amounts of protein (i.e., 30 mg; this represents <10% of the protein produced in the Stage 1). Using the Pharmacia Biotech FPLC system for the chromatography steps, stage 2 takes 2 days.
Literature Cited Abdel-Meguid, S.S., Shieh, H.-S., Smith, W. W., Dayringer, H.E., Violand, B.N., and Bentle, L.A. 1987. Three-dimensional structure of a genetically engineered variant of porcine growth hormone. Proc. Natl. Sci. U.S.A. 84:6434-6437. Ackers, G.K. 1970. Analytical gel chromatography of proteins. Adv. Protein Chem. 24:343-446. Bastiras, S. and Wallace, J. C. 1992. Equilibrium denaturation of recombinant porcine growth hormone. Biochemistry 31:9304-9309. Bazan, J.F. 1992. Unraveling the structure of IL-2. Science 257:410-413. Bell, J.A., Moffat, K., Voderhaar, B.K., and Golde, D.W. 1985. Crystallization and preliminary Xray characterization of bovine growth hormone. J. Biol. Chem. 260:8520-8525. Bogosian, G., Violand, B.N., Dorward-King, E.J., Workman, W.E., Jung, P.E., and Kane, J.F. 1989. Biosynthesis and incorporation into protein of norleucine by Escherichia coli. J. Biol. Chem. 264:531-539. Bottomly, K., Davis, C.S., and Lipsky, P.E. 1991. Measurement of human and murine interleukin2 and interleukin-4. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeek, D.H. Marguiles, E.M. Shevach, and W. Strober, eds.) pp. 6.3.1-6.3.12. John Wiley & Sons, New York. Brems, D.N. and Havel, H.A. 1989. Folding of bovine growth hormone is consistent with the molten globule hypothesis. Proteins Struct. Funct. Genet. 5:93-95. Browning, J.L., Mattaliano, R.J., Chow, E.P., Liang, S.-M., Allet, B., Rosa, J., and Smart, J.E. 1986. Disulfide scrambling of interleukin-2: HPLC resolution of the three possible isomers. Anal. Biochem. 155:123-128.
Purification of Recombinant Proteins
6.5.25 Current Protocols in Protein Science
Bushman, F.D., Engelman, A., Palmer, I., Wingfield, P., and Craigie, R. 1993. Domains of the integrase protein of human immunodeficiency virus type 1 responsible for the polynucleotidyl transfer and zinc binding. Proc. Natl. Acad. Sci. U.S.A. 90:3428-3432. Coleman, R., Iqbal, S., Godfrey, P.P., and Billington, D. 1979. Membranes and bile formation. Composition of several mammalian biles and their membrane-damaging properties. Biochem. J. 178:201-208 deVos, A.M., Uttsch, M., and Kossiakoff, A.A. 1992. Human growth hormone and extracellular domain of its receptor: Crystal structure of the complex. Science 255:306-312. Devos, R., Plaetinc, G., Cheroutr, H., Simons, G., Degrave, W., Tavernie, J., Remaut, E., and Fiers, W. 1983. Molecular cloning of human interleukin-2 carrier DNA and its expression in Escherichia coli. Nucl. Acids Res. 11:4307-4323. Dyda, F., Hickman, A.B., Jenkins, T.M., Engelman, A., Craigie, R., and Davies, D.R. 1994. Crystal structure of the catalytic domain of HIV-1 integrase: Similarity to other polynucleotidyl transferases. Science 266:1981-1986. Edelhoch, H. and Burger, H.G. 1966. The properties of bovine growth hormone. II. Effects of urea. J. Biol Chem. 241:458-463. Fernandez, H.N. and Delfino, J.M. Covalent crosslinking of the bovine somatotropin dimer. Biochem. J. 209:107-115. Goldberg, M.E., Ruldolph, R., and Jaenicke, R. 1991. A kinetic study of the competition between renaturation and aggregation during the refolding of denatured-reduced egg white lysozyme. Biochemistry 30:2790-2797. Goldenberg, D.P. 1989. Analysis of protein conformation by gel electrophoresis. In Protein Structure: A Practical Approach (T.E. Creighton, ed.) pp. 225-250. IRL Press, Oxford. Graf, L., Li, C.H., and Bewley, T. 1975. Selective reduction and alkylation of the COOH-terminal disulfide bridge in bovine growth hormone. Int. J. Pep. Prot. Res. 7:467-473. Havel, H.A., Chao, R.S., Haskell, R.J., and Thamann, T.J. 1989. Investigations of protein structure with optical spectroscopy: Bovine growth hormone. Anal. Chem. 61:642-650. Hickman, A., Palmer, I., Engelman, A., Craigie, R., and Wingfield, P. 1994. Biophysical and enzymatic properties of the catalytic domain of HIV-1 integrase. J. Biol. Chem. 269:29279-29287. Holzman, T.F., Brems, D.N., and Dougherty, J.J. 1986. Reoxidation of reduced bovine growth hormone from a stable secondary structure. Biochemistry 25:6707-6917. Jaenicke, R. 1991. Protein folding: Local structures, domains, subunits, and assemblies. Biochemistry 30:3147-3161. Folding and Purification of Insoluble Proteins from E. coli
Kato, K., Yamada, T., Kawahara, K., Onda, H., Asano, T., Sugino, H., and Kakinuma, A., 1985. Purification and characterization of recombinant human interleukin-2 produced in Escherichia
coli. Biochem. Biophys. Res. Commun. 130:692699. Katz, R.A. and Skalka, A.M. 1994. The retroviral enzymes. Annu. Rev. Biochem. 63:133-173. Langley, K.E., Berg, T.F., Strickland, T.W., Fenton, D.M., Boone, T.C., and Wypych, J. 1987a. Recombinant-DNA-derived bovine growth hormone from E. coli. I. Eur. J. Biochem. 163:313321. Langley, K.E., Lai, P.-H., Wypych, J., Everett, R.R., Berg, T.F., Krabill, L.F., and Souza, L.M. 1987b. Recombinant-DNA-derived bovine growth hormone from E. coli. II. Eur. J. Biochem. 323-330. Liang, S.-M., Allet, B., Rose, K., Hirschi, M., Liang, C.-M., and Thatcher, D.R. 1985. Characterization of human interleukin 2 derived from Escherichia coli. Biochem. J. 229:429-439. Liang, S.-M., Thatcher, D., Liang, C.-M., and Allet, B. 1986. Studies of structure activity relationships of human interleukin-2. J. Biol. Chem. 261:334-337. Maeda, Y., Ueda, T., Yamada, H., and Imoto, T. 1994. The role of net charge on the renaturation of reduced lysozyme by sulfhydryl-disulfide interchange reaction. Prot. Engineering 7:12491254. McKay, D.B. 1992. Unraveling the structure of IL2-response. Science 257:412-413. Minami, Y., Kono, T., Miyazaki, T., and Taniguchi, T. 1993. The IL-2 receptor complex: Its structure, function, and target genes. Annu. Rev. Immunol. 11:245-267. Orsini, G. and Goldberg, M.E. 1978. The renaturation of reduced chymotrypsin A in guanidine HCl. J. Biol. Chem. 253:3453-3458. Qiagen. 1992. The QIAexpressionist, 2nd ed. Qiagen, Chatsworth, Calif. Riddles, P. W., Blakeley, R.L., and Zerner, B. 1979. Ellman’s reagent: 5,5′-dithiobis(2-nitrobenzoic acid)—a reexamination. Anal Biochem. 94:7581. Robb, R.J., Kutney, R.M., Panico, M., Morris, H.R., and Chowdhry, V. 1984. Amino acid sequence and post-translational modification of human interleukin-2. Proc. Natl. Acad. Sci. U.S.A. 81:6486-6490. Secchi, C., Biondi, P. A., Negri, A., Borrini, R., and Ronchi, S. 1986. Detection of desamido forms of purified bovine growth hormone. Int. J. Pept. Prot. Res. 28:298-306. Sherman, P.A. and Fyfe, J.A. 1990. Human immunodeficiency virus integration protein expressed in Escherichia coli possesses selective DNA cleavage activity. Proc. Natl. Acad. Sci. U.S.A. 87:5119-5123. Spitsberg, V.L. 1987. A selective extraction of growth hormone from bovine pituitary gland and its further purification and crystallization. Anal. Biochem. 160:489-495. Staten, N.R., Byatt, J.C., and Krivi, G.G. 1993. Ligand-specific dimerization of the extracellular
6.5.26 Current Protocols in Protein Science
domain of the bovine growth hormone receptor. J. Biol. Chem. 268:18467-18473.
isomerization of protein disulfides. Methods Enzymol. 107:301-304.
Stoscheck, C.M. 1990. Quantitation of protein. Methods Enzymol. 182:50-67.
Wingfield, P.T., Graber, P., Buell, G., Rose, K., Simona, M., and Burleigh, B.D. 1987a. Preparation and characterization of bovine growth hormones produced in recombinant Escherichia coli. Biochem. J. 243:929-839.
Talalay, P. 1960. Enzymic analysis of steroid hormones. Biochem. Anal. 8:119-142. Terry, R., Soltis, D.A., Katzman, M., Cobrinik, D., Leis, J., and Skalka, A.M. 1988. Properties of avian sarcoma-leukosis virus pp32-related polendonucleases produced in Escherichia coli. J. Virol. 62:2358-2365. Thannhauser, T.W., Konishi, Y., and Scheraga, H.A. 1984. Sensitive quanititative analysis of disulfides in polypeptides and proteins. Anal. Biochem. 138:181-188. Violand, B.N., Takano, M., Curran, D.F., and Bentle, L.A. 1989. A novel concatenated dimer of recombinant bovine somatotropin. J. Prot. Chem. 8:619-628. Wallis, O.C. and Wallis, M. 1989. Purification and properties of recombinant DNA–derived ovine growth hormone analogue (oGH1) expressed in Escherichia coli. J. Mol. Endocrinology 4:61-69. Wang, A., Lu, S.-D., and Mark, D. 1984. Studies of structure-activity relationships of human interleukin-2. Science 224:1431-1433. Weir, M.P. and Sparks, J., 1987. Purification and renaturation of recombinant human interleukin2. Biochem. J. 245:85-91. Wetlaufer, D.B. 1984. Nonenzymatic formation and
Wingfield, P.T., Graber, P., Rose, K., Simona, M.G., and Hughes, G.J. 1987b. Chromatofocusing of N-terminally processed forms of proteins: Isolation and characterization of two forms of interleukin-1β and bovine growth hormone. J. Chromatogr. 387:291-300. Wood, D.C., Salsgiver, W.J., Kasser, T.R., Lange, G.W., Rowold, E., Violand, B.N., Johnson, A., Leimgruber, R.M., Parr, G.R., Siegel, N.R., Kimack, N.M., Smith, C.E., Zobel, J.F., Ganguli, S.M., Garbow, J.R., Bild, G., and Krivi, G.G. 1989. Purification and characterization of pituitary bovine somatotropin. J. Biol. Chem. 264:14741-14747.
Contributed by Paul T. Wingfield (BGH and HIV-1 integrase) and Ira Palmer (HIV-1 integrase) National Institutes of Health Bethesda, Maryland Shu-Mei Liang (interleukin-2) North American Vaccine Corp. Beltsville, Maryland
Purification of Recombinant Proteins
6.5.27 Current Protocols in Protein Science
Expression and Purification of GST Fusion Proteins
UNIT 6.6
This unit describes how pGEX vectors (available from Pharmacia Biotech) can be used for high-level inducible intracellular expression of polypeptides as fusions with glutathione-S-transferase (GST) in Escherichia coli. GST fusion proteins are easily purified under nondenaturing conditions by affinity chromatography (Chapter 9) using a glutathione–Sepharose 4B conjugate. The amino-terminal GST moiety can then be cleaved from the protein of interest using a specific protease cleavage site located between the GST moiety and the recombinant polypeptide. Finally, the GST moiety can be removed by rechromatographing the sample on the glutathione-Sepharose column. Potential applications of the system include the expression and purification of large quantities of individual polypeptides for use in structural determinations using either nuclear magnetic resonance (NMR) or crystallography, immunological studies, vaccine production, and structure-function studies involving protein-protein and DNA-protein interactions. The basic protocols are chromatography-based adaptations of the manufacturer’s recommendations and include batch and column purification methods. The experience of this laboratory is that, although batch purifications may be slightly faster, column-based purification steps usually provide higher protein yields and higher purity of the final product. GST fusion proteins can be expressed at high levels in E. coli grown using an environmental shaker (see Basic Protocol 1). The soluble fusion protein is purified using a single-step glutathione–Sepharose 4B affinity column (see Basic Protocol 2). Fusion proteins that are found in inclusion bodies can be extracted using denaturants and then refolded before affinity chromatography using a glutathione–Sepharose 4B column (see Alternate Protocol 1). Soluble fusion proteins can also be purified batchwise using glutathione–Sepharose 4B affinity matrix (see Alternate Protocol 2). The fusion protein can be cleaved with either thrombin or factor Xa in solution to separate the protein of interest from glutathione (see Basic Protocol 3). The protein of interest can also be released by protease digestion of the fusion protein immobilized on a glutathione– Sepharose 4B column (see Alternate Protocol 3), or by batchwise protease cleavage and separation from the resin (see Alternate Protocol 4). Affinity purification using glutathione–Sepharose 4B can be used to remove the GST moiety after enzymatic cleavage (see Support Protocol 1). HPLC gel filtration is used as a final purification step to obtain >98% pure protein (see Support Protocol 2). The relationships between the protocols are shown in Figure 6.6.1. STRATEGIC PLANNING A variety of pGEX expression vectors are commercially available (Pharmacia Biotech) that contain a tac promoter for chemically inducible, high-level protein expression. The available pGEX vectors have an open reading frame encoding glutathione-S-transferase (GST) followed by multiple cloning sites. These are followed by termination codons in each reading frame (Figs. 6.6.2 and 6.6.3). A fragment of DNA containing the genetic sequence for the polypeptide of interest is ligated into an appropriate pGEX vector and transformed into E. coli. It should be noted that although expression in E. coli is efficient, there is no post-translational modification machinery. Successful expression of GST
Contributed by Sandra Harper and David W. Speicher Current Protocols in Protein Science (1997) 6.6.1-6.6.21 Copyright © 1997 by John Wiley & Sons, Inc.
Purification of Recombinant Proteins
6.6.1 Supplement 9
expression of GST fusion protein (Basic Protocol 1)
on-column affinity purification (Basic Protocol 2 or Alternate Protocol 1)
cleavage on column (Alternate Protocol 3)
elution
batchwise purification with affinity resin (Alternate Protocol 2)
elution
cleavage in solution (Basic Protocol 3)
elution
cleavage bound to resin (Alternate Protocol 4)
cleavage in solution (Basic Protocol 3)
protein purification by affinity chromatography (Support Protocol 1) and/or HPLC (Support Protocol 2)
Figure 6.6.1 Flow chart showing the relationships between the various protocols in this unit.
fusion proteins using baculovirus systems (Davies et al., 1993) and yeast (Mitchell et al., 1993) have also been reported. One factor that influences which pGEX vector to choose is whether or not the GST moiety will ultimately be cleaved away from the protein of interest. The pGEX-2T and pGEX-4T series of vectors contain a protease cleavage site for thrombin, and the pGEX-3X and pGEX-5X series of vectors contain protease cleavage sites for factor Xa. A more recently developed expression vector is the pGEX-6P series, which contains a cleavage site for PreScission protease (Pharmacia Biotech). PreScission protease has the advantage that it is effective at low temperatures (5°C). It is also a GST fusion protein, a feature that facilitates removal of the protease from the target protein after cleavage. Fusion proteins with a thrombin recognition site have the advantage that relatively small amounts of thrombin and short digestion times at 37°C will often cleave the fusion protein with high efficiency. Thrombin digestions are often the most cost effective on a per milligram of cleaved target polypeptide basis. Factor Xa is more expensive and typically requires use of much higher enzyme-to-substrate ratios for efficient cleavage. Solutions of factor Xa also have a more limited shelf life since freezing and thawing inactivates this enzyme.
Expression and Purification of GST Fusion Proteins
For preparation of cDNA inserts encoding the desired polypeptide, see Ausubel et al. (1994) or Sambrook et al. (1989). Briefly, a set of oligonucleotides is designed for polymerase chain reaction (PCR) amplification of the region of interest of a pertinent cDNA. These oligonucleotides should also contain appropriate restriction sites adjacent to the desired coding region that are compatible with a restriction site in the cloning site of the selected pGEX vector (see Figs. 6.6.2 and 6.6.3). PCR amplification is performed,
6.6.2 Supplement 9
Current Protocols in Protein Science
pGEX-1λT thrombin Leu Val Pro Arg ↓ Gly Ser Pro Glu Phe Ile Val Thr Asp CTG GTT CCG CGT GGA TCC CCG GAA TTC ATC GTG ACT GAC TGA CGA
BamHI
stop codons
EcoRI Tth111I Aat II
BalI glutathione-S- transferase Ptac
r
pSj10∆Bam7Stop7
Ap
BspMI
PstI
pGEX ~4950 bp
la
NarI EcoRV BssHII ApaI Bst EII MluI
Alw NI
p4.5
q
cl
pBR322 ori
Figure 6.6.2 pGEX vectors are plasmid expression vectors that express a cloned gene as a fusion protein with glutathione-S-transferase (GST). The lac repressor gene binds to the lac promoter (ptac) and represses expression of the GST fusion protein until induction with isopropyl-1-thio-β-D-galactopyranoside (IPTG). The polypeptide of interest can be inserted immediately after the GST gene using the polylinker site shown in brackets (pGEX-1λT, shown here, is the most common; see Fig. 6.6.3 for other PGEX polylinkers). Protease cleavage sites (brackets above the polylinker sequences) are located between the GST carrier protein and the protein of interest so that the GST moiety can be removed. Restriction endonuclease sites are indicated below the sequence of the polylinker and on the plasmid. An important consideration in selecting a vector and appropriate cloning sites is to minimize the number of extraneous residues introduced into the N-terminal of the target polypeptide. Vector map courtesy of Pharmacia Biotech.
followed by digestion of the PCR product and the pGEX vector with the appropriate restriction enzymes. The PCR product is then ligated into the pGEX vector and transfected into a suitable E. coli host. Several transformants should be grown in minicultures and induced with isopropyl-1-thio-β-D-galactopyranoside (IPTG) to check for expression of the desired fusion protein. Fusion protein expression can be monitored by SDS-PAGE (UNIT 10.1) or by Western blot (UNIT 10.10) detection of the GST fusion protein using an antibody specific for either the target protein or the GST moiety. Once successful expression is achieved, the integrity of the DNA should be verified by sequencing to ensure that no errors were introduced during PCR. Before conducting a large-scale purification, it is worthwhile to perform a small pilot purification (∼10-fold less than protocol descriptions) to determine optimal conditions. The purification can then easily be scaled up. All stages of purification should be monitored using SDS-PAGE (UNIT 10.1). In most cases, GST fusion protein expression is very high and a major band at the expected molecular weight (the GST moiety contributes 26 kDa to the molecular weight) is obvious when uninduced and induced cells are compared on SDS gels (Fig. 6.6.4). This band can then be monitored at each step of the purification. However, as noted above, if the level of protein expression obtained is low or band identification is ambiguous, the fusion protein can be monitored by Western blot
Purification of Recombinant Proteins
6.6.3 Current Protocols in Protein Science
Supplement 9
pGEX-2T thrombin Leu Val Pro Arg ↓ Gly Ser Pro Gly Ile His Arg Asp CTG GTT CCG CGT GGA TCC CCG GGA ATT CAT CGT GAC TGACTG ACG
stop codons
BamHI SmaI EcoRI pGEX-2TK kinase
thrombin
Leu Val Pro Arg ↓ Gly Ser Arg Arg Ala Ser Val CTG GTT CCG CGT GGA TCT CGT CGT GCA TCT GTT GGA TCC CCG GGAATT CATCGT GAC TGA
stop codon
BamHI SmaI EcoRI pGEX-4T-1
thrombin
Leu Val Pro Arg ↓ Gly Ser Pro Glu Phe Pro Gly Arg Leu Glu Arg Pro His Arg Asp CTG GTT CCG CGT GGA TCCCCG GAATTC CCG GGT CGA CTC GAG CGGCCG CAT CGT GAC TGA
BamHI
SalI
EcoRI SmaI
stop codons
NotI
XhoI
pGEX-4T-2
thrombin
Leu Val Pro Arg ↓Gly Ser Pro Gly Ile Pro Gly Ser Thr Arg Ala Ala Ala Ser CTG GTTCCG CGT GGA TCC CCA GGA ATT CCC GGG TCGACT CGA GCG GCC GCA TCG TGA
BamHI
EcoRI SmaI
SalI
stop codons
NotI
XhoI
pGEX-4T-3
thrombin
Leu Val Pro Arg ↓ Gly Ser Pro Asn Ser Arg Val Asp Ser Ser Gly Arg Ile Val Thr Asp CTG GTT CCG CGT GGA TCC CCG AAT TCC CGG GTC GAC TCG AGC GGC CGC ATC GTG ACT GAC TGA
BamHI
factor Xa
EcoRI SmaI
SalI
stop codons
NotI
XhoI
pGEX-3X
Ile Glu Gly Arg ↓Gly Ile Pro Gly Asn Ser Ser ATC GAA GGT CGT GGG ATC CCC GGG AAT TCA TCG TGA CTG ACT GAC
stop codons
BamHI SmaI EcoRl pGEX-5X-1
factor Xa
Ile Glu Gly Arg ↓ Gly Ile Pro Glu Phe Pro Gly Arg Leu Glu Arg Pro His Arg Asp ATC GAA GGT CGT GGG ATC CCC GAATTC CCG GGTCGA CTC GAG CGG CCG CAT CGT GAC TGA
BamHI
EcoRl SmaI
Sall
XhoI
Not l
stop codons
pGEX-5X-2
factor Xa
Ile Glu Gly Arg ↓ Gly Ile Pro Gly Ile Pro Gly Ser Thr Arg Ala Ala Ala Ser ATC GAA GGT CGT GGG ATC CCC GGA ATTCCC GGG TCG ACT CGA GCGGCC GCATCG TGA
BamHI
EcoRl SmaI
Sall
XhoI
Not l
stop codons
pGEX-5X-3
factor Xa
Ile Glu Gly Arg ↓ Gly Ile Pro Arg Asn Ser Arg Val Asp Ser Ser Gly Arg lle Val Thr Asp ATC GAA GGT CGT GGG ATC CCC AGG AAT TCC CGGGTC GAC TCG AGC GGC CGC ATC GTG ACT GAC TGA
BamHI
EcoRI SmaI
SalI
XhoI
NotI
Stop codons
Figure 6.6.3 Variations on the polylinker site shown in Figure 6.6.2 (courtesy of Pharmacia Biotech). Expression and Purification of GST Fusion Proteins
6.6.4 Supplement 9
Current Protocols in Protein Science
95 66 43 36
66
25 17
29
6 18
1
2
L
S 30 C
Figure 6.6.4 SDS gel stained with Coomassie brilliant blue showing the expression of a glutathione-S-transferase (GST) fusion protein. Lane 1, total cell lysate before induction with isopropyl1-thio-β-D-galactopyranoside (IPTG). Lane 2, total cell lysate after 3-hr induction with IPTG. The position of the fusion protein is indicated by the arrow.
P
L
S 25 C
P
L
S
P
18 C
Figure 6.6.5 SDS gel stained with Coomassie brilliant blue showing the effects of different growth temperatures on solubility of an expressed glutathione-S-transferase (GST) fusion protein. E. coli transfected with a recombinant pGEX-2T vector were grown at different temperatures as indicated. Proportional aliquots of whole-cell lysate after sonication (L), the supernatant after centrifuging the lysate (S), and the remaining pellet (P) are shown. This recombinant was primarily in inclusion bodies (P) at 37°C (not shown), 30°C, and 25°C. At 18°C, however, nearly all of the expressed protein was in a soluble native form.
analysis (UNIT 10.10) using a GST-specific antibody (Pharmacia Biotech). It is recommended that the lysed cell extract, extracted pellet, and all other collected fractions from the purification be saved on ice until after careful analysis of the entire purification by SDS-PAGE and/or immunoblotting to ensure that fractions containing the fusion protein are not mistakenly discarded. Basic Protocol 1 describes protein production in cells grown at 37°C; however, at this temperature some fusion proteins may be found in inclusion bodies in a denatured form. As an alternative to attempting to renature the protein after extraction from inclusion bodies (UNIT 6.1), expression at lower temperatures, such as 30°, 25°, 20°, or 15°C, can be evaluated to determine whether the protein can be obtained in good yield in the soluble fraction (Fig. 6.6.5). When expressing fusion proteins at lower temperatures, the initial overnight culture can be grown at 37°C followed by growth at a lower temperature prior to induction. EXPRESSION OF GLUTATHIONE-S-TRANSFERASE FUSION PROTEIN Transformed E. coli cells expressing the glutathione-S-tranferase (GST) fusion protein of interest are grown in culture in the presence of isopropyl-1-thio-β-D-galactopyranoside (IPTG) at the desired preparative scale. Since the expression level of GST fusion proteins is usually very high, adequate amounts of protein can usually be conveniently obtained by preparing a few liters of cells grown in shaker cultures. This protocol describes the preparation of 1.8 liters of transfected E. coli in three 600-ml units using 2-liter flasks in a shaker incubator. Moderate further scale-up is feasible by using more or larger flasks.
BASIC PROTOCOL 1
Purification of Recombinant Proteins
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Further scale-up can be accomplished using a fermentor (see UNITS 5.3 & 5.4). This protocol describes protein production in cells grown at 37°C. At this temperature, however, some fusion proteins may be recovered from inclusion bodies in a denatured form, and culture conditions may need to be modified to improve protein yield in the soluble fraction (see Strategic Planning). Culture growth can be monitored by reading the optical density at 550 nm (OD550) as well as by analysis of the bacterial culture using SDS-PAGE. Cells should not be allowed to grow for extended periods of time after induction as cell lysis can occur; this releases proteases that may degrade the fusion protein. Visual inspection of the cells using a microscope is a useful method for identifying cell breakage (see UNITS 5.1-5.3 & 6.1-6.5 for additional details concerning recombinant protein expression in E. coli). Materials Luria broth (LB medium; UNIT 5.2; pH adjusted to 7.2) 5 mg/ml ampicillin (see recipe) Glycerol culture of transformed E. coli cells expressing GST fusion protein of interest in a pGEX vector 100 mM isopropyl-1-thio-β-D-galactopyranoside (IPTG; see recipe) 2-liter culture flasks 500-ml culture flasks Large centrifuge bottles (e.g., 1-liter capacity) Low-speed refrigerated centrifuge (e.g., Beckman J6-B and JS-4.2 rotor or equivalent), 4°C Additional reagents and equipment for SDS-polyacrylamide gel electrophoresis (SDS-PAGE; UNIT 10.1) Grow bacterial cells 1. Prepare LB medium and add 600 ml to each of three 2-liter flasks and 100 ml to each of two 500-ml flasks. Autoclave 20 to 30 min at a slow exhaust (liquid) setting. Flasks should be filled to only 20% to 30% of their capacity to ensure adequate aeration of the medium during cell growth. Autoclave LB medium immediately after preparing it to prevent any incidental bacterial growth from occurring. It can then be stored up to 1 month at room temperature under sterile conditions.
2. Allow medium to cool to room temperature. Add 1 ml of 5 mg/ml ampicillin to one flask containing 100 ml LB medium. It is important to allow medium to cool before proceeding, because ampicillin is inactivated at temperatures >50°C. Flame the opening of all bottles and flasks to reduce the risk of contamination.
3. Using an inoculating loop, transfer some of the glycerol culture containing the transfected E. coli expressing the GST fusion protein of interest to the flask containing 100 ml LB medium with ampicillin. Sterile flame the inoculating loop as well as the opening of all bottles. Allow the loop to cool before transferring the inoculating culture. If the loop temperature is too high, all the cells could be killed during the transfer.
4. Incubate the inoculated culture on an environmental shaker set at 250 to 300 rpm overnight at 37°C. Expression and Purification of GST Fusion Proteins
5. The next morning, remove the culture from the shaker and read the optical density at 550 nm (OD550) using a UV/visible light spectrophotometer. The OD550 of the overnight culture should be ∼1.0. Use medium from the second 500-ml flask that did not receive ampicillin as a reference to zero the spectrophotometer.
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6. Using sterile technique, add 6 ml of 5 mg/ml ampicillin to each 2-liter flask containing 600 ml LB medium (0.1 mM final concentration). 7. Dilute the overnight culture 1:20 by adding 30 ml to each of the three 2-liter flasks containing 600 ml LB medium. 8. Incubate the 600-ml cultures on a shaker at 37°C at 250 to 300 rpm until the OD550 is 0.5 to 0.7. It should take ∼2 hr for the culture to reach this early log stage of growth at 37°C. If cells are grown at lower temperatures to shift the expressed protein from inclusion bodies into the soluble fraction, this time must be increased, since the cells will grow more slowly at lower temperatures.
Induce expression of fusion protein 9. Remove a 1-ml sample from each flask and save for gel analysis. Induce cells by adding 6 ml of 100 mM IPTG per flask (1.0 mM final). Incubate at 37°C for an additional 2.5 to 3 hr. Optimal growth conditions (OD550 at time of induction, growth temperature, and growth time after induction) for each recombinant protein should be empirically determined. The conditions given here will generally be suitable for growth at either 37° or 30°C. At lower temperatures (≤25°C), longer growth times will definitely be needed, and the best protein yields will usually be obtained if the OD550 prior to induction is 0.65 to 0.85.
10. Remove cultures from the shaker 2.5 to 3 hr after induction. Remove 1 ml from each culture and save for gel analysis. Check final OD550. Culture growth can be monitored at OD550. When the cells reach saturation, they will stop dividing. A typical SDS-PAGE gel of an uninduced control culture and an induced culture after 3 hr growth is shown in Figure 6.6.4.
Recover bacterial cells 11. Pour each culture into a large centrifuge bottle and centrifuge 20 min at 4000 × g, 4°C. Medium from the second 500-ml flask without ampicillin can be used to balance the centrifuge bottles.
12. Carefully decant the supernatant, leaving 15 to 50 ml in the centrifuge bottle. Resuspend the pelleted cells in the remaining supernatant. Transfer the cell suspension to a 50-ml centrifuge tube. 13. Centrifuge 20 min at 4000 × g, 4°C. Decant the supernatant. 14. Freeze cell pellet by placing in a −80°C freezer. 15. Analyze saved 1-ml samples from before and after induction using SDS-PAGE (UNIT 10.1) to check for protein expression. As noted above, a 1-ml aliquot of culture is removed prior to induction and a second 1-ml sample is removed after the 3-hr induction. These aliquots can be centrifuged for 2 min and the supernatants carefully removed with a pipet. These samples can be stored at 0° to 4°C overnight prior to running a gel; for longer-term storage prior to gel analysis, store frozen (−20°C) to prevent proteolysis. The cell pellet can be directly resuspended in SDS sample buffer (200 ìl) and heated 3 to 5 min at 90°C. Typical results are shown in Figure 6.6.4. Purification of Recombinant Proteins
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BASIC PROTOCOL 2
AFFINITY CHROMATOGRAPHY PURIFICATION OF A SOLUBLE GST FUSION PROTEIN Soluble glutathione-S-transferase (GST) fusion proteins can easily be purified from cell lysate supernatants by affinity chromatography on glutathione–Sepharose 4B using either batch or column loading of the sample. Protease inhibitors should be added to the lysis buffer to minimize potential proteolysis. As an aid for cell lysis, 1% (v/v) Triton X-100 may be added to the lysis buffer. After cells are lysed, centrifugation is used to pellet any unlysed cells and inclusion bodies. Location of the fusion protein can usually easily be determined by analyzing a small aliquot of both the supernatant and the pellet by SDS-PAGE (UNIT 10.1). The glutathione–Sepharose 4B can be regenerated and reused for subsequent purifications. To avoid potential cross-contamination of different proteins or mutant forms of a single protein, however, it is recommended that a given column or batch of resin be reserved for use with a single protein. Preequilibration of the glutathione column (steps 1 to 4) can be performed either prior to or in parallel with cell lysis. All steps except SDS-PAGE (room temperature) should be performed in a cold room at 4°C, unless otherwise noted. Materials Glutathione–Sepharose 4B resin (Pharmacia Biotech) PBS (APPENDIX 2E) Glutathione buffer (see recipe) PBS/EDTA/PMSF buffer (see recipe) Pelleted E. coli culture expressing fusion protein (see Basic Protocol 1, step 14) Lysis buffer (see recipe), ice cold Wash buffer (see recipe), ice cold PBS/EDTA (see recipe) 2.5 × 8–cm glutathione–Sepharose 4B column (e.g., Bio-Rad Econo) Peristaltic pump Sonicator equipped with microtip probe (e.g., Branson) Dounce homogenizer 60-ml centrifuge bottles (capable of handling force of 48,000 × g) High-speed refrigerated centrifuge (e.g., Beckman JZ-21M centrifuge and JA-18 rotor or equivalent), 4°C Additional reagents and equipment for pouring chromatographic columns (UNIT 8.3) and for SDS-PAGE (UNIT 10.1) Preequilibrate glutathione column 1. Pour 20 ml glutathione–Sepharose 4B resin into 2.5 × 8–cm column (glutathione column; see UNIT 8.3 for details of pouring procedure). This amount of resin should be adequate for purification of fusion protein from three 600-ml cultures that contain ∼20 to 40 mg fusion protein per culture. Although glutathione– Sepharose 4B has an advertised minimum binding capacity of 8 mg/ml resin, the actual capacity may be substantially different and should be empirically determined. The actual amount of resin used and column size can be increased or decreased depending on the amount of fusion protein to be purified.
2. Wash the glutathione column with 5 to 10 bed volumes PBS at a flow rate of 1.5 ml/min to remove the ethanol storage solution. Expression and Purification of GST Fusion Proteins
A bed volume is one-half the amount of glutathione–Sepharose 4B that was added to the column as a 50% slurry.
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New columns should be prepared as described in steps 1 to 4. Previously used columns should be preequilibrated using steps 3 to 4 only, to ensure that the column is completely reduced and has maximal binding capacity. Use of a peristaltic pump is recommended for convenient control of flow rates. Compression of the resin bed indicates that the column pressure is too high and that the flow rate should be lowered.
3. Wash the glutathione column with 3 to 5 bed volumes glutathione buffer at 1.5 ml/min. Previously used columns may become partially oxidized on storage and should be preequilibrated (steps 3 to 4) within 24 hr before use.
4. Wash the glutathione column with 10 bed volumes PBS/EDTA/PMSF at 1.5 ml/min. Lyse cells 5. Resuspend each pelleted 600-ml culture in 15 ml ice-cold lysis buffer. Pellets should be resuspended in 25 to 50 ìl buffer per milliliter of culture.
6. Sonicate the suspension using a probe-tip sonicator ten times for 10 sec each, with 1-min rests between sonications to lyse the cells. Save a sample (∼100 µl) of the lysate for gel analysis and transfer remainder to a 60-ml centrifuge tube. The cells are usually adequately lysed at the point when the suspension turns a slightly darker color and becomes clearer. To minimize proteolysis in the sample, it is essential to keep the cells on ice throughout the sonication procedure, and sonication should be performed in short bursts to minimize sample heating. Avoid excessive sonication, as this can lead to co-purification of E. coli host proteins along with the fusion protein of interest. Avoid frothing during sonication, which can denature the fusion protein.
7. Centrifuge the lysate 20 min at 48,000 × g, 4°C. Unbroken cells, large cellular debris, and inclusion-body protein will be pelleted.
8. Decant the supernatant containing the soluble fusion protein into a clean 50-ml centrifuge tube. 9. Add a volume of ice-cold wash buffer equal to the volume of lysis buffer used in step 5 to the pellet. Use a dounce homogenizer to resuspend the pellet. Pellets should be resuspended in 25 to 50 ìl buffer per milliliter of culture.
10. Analyze the lysate, supernatant, and resuspended pellet using SDS-PAGE (UNIT 10.1) to verify that the fusion protein is in the supernatant. If the fusion protein is in the supernatant, proceed to the next step. If the protein is in the pellet, it is necessary to purify the GST fusion proteins from inclusion bodies (see Alternate Protocol 2) or to start over, shifting the protein into the supernatant by growing the cultures at a lower temperature (see Troubleshooting and Fig. 6.6.5).
Load the column 11. Load the supernatant onto a preequilibrated glutathione column (see step 4). Collect fractions and analyze multiple fractions across the unbound peak by SDS-PAGE (UNIT 10.1) to verify that the fusion protein has bound and that column capacity was not exceeded. For a 2.5-cm diameter column, a sample-loading flow rate of ≤0.1 ml/min is recommended to permit complete binding of the fusion protein to the resin. Faster flow rates may decrease the yield of bound fusion protein due to slow kinetics of association at 4°C, unless a large excess of resin is used. This step is most conveniently performed overnight; the flow rate
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can be adjusted so that most of the supernatant will be loaded by the next morning. Do not allow the column to run dry. SDS-PAGE analysis of fractions collected during sample loading will reveal whether the fusion protein is bound to the column or is present in the unbound fractions. Absence of fusion protein in early unbound fractions combined with its appearance in late unbound fractions indicates that column capacity has been exceeded. If this condition is observed, reduce the protein load or increase the column size.
12. Wash the column with 5 to 10 bed volumes PBS/EDTA/PMSF at 1.5 ml/min. 13. Wash the column with 10 bed volumes PBS/EDTA at 1.5 ml/min to remove the PMSF. If samples are to be cleaved with thrombin or factor Xa, any serine protease inhibitor (e.g., PMSF) must be removed from the sample before cleavage.
Elute the fusion protein 14. Elute the fusion protein from the column by washing the column with 5 bed volumes glutathione buffer. A flow rate of 0.3 ml/min for a 2.5-cm column is recommended to elute the fusion protein in a minimal volume. Elution of the fusion protein can be monitored at A280 either with an online UV monitor or by reading the absorption of individual fractions.
15. Analyze the fractions by SDS-PAGE (UNIT 10.1) and pool fractions containing the GST fusion protein. Store at 0° to 4°C. Fusion protein should typically be >90% pure at this point. ALTERNATE PROTOCOL 1
AFFINITY CHROMATOGRAPHY PURIFICATION OF GST FUSION PROTEIN FROM INCLUSION BODIES In some cases, fusion proteins are entirely or primarily located in a denatured, aggregated form in inclusion bodies. Glutathione-S-transferase (GST) fusion proteins can often be purified from inclusion bodies after solubilization in urea or another denaturant followed by renaturation by dialysis (UNIT 6.3). Other methods of solubilization include addition of detergents such as Sarkosyl (N-laurylsarosine; Grieco et al., 1992; Frangioni, 1992). After denaturation and renaturation, it is important to ensure that the protein has regained its native conformation and function (UNITS 6.4 & 6.5). Although denatured GST will not bind to the glutathione column and hence will not be recovered by this method, the possibility that the GST moiety has refolded when the fusion partner has not folded properly should also be considered. As an alternative to purifying proteins from inclusion bodies, growing the cells at a lower temperature will often shift the fusion protein into the supernatant while still producing ≥10 mg of fusion protein per liter of culture when pGEX vectors are used (see Strategic Planning and see Troubleshooting). All steps should be performed in a cold room at 4°C unless otherwise noted. Additional Materials (also see Basic Protocol 2) U buffer (see recipe) Triton X-100 PBS/glycerol buffer (see recipe) Low-speed refrigerated centrifuge (e.g., Beckman J6-B and JS-4.2 rotor or equivalent), 4°C Additional reagents and equipment for dialysis (APPENDIX 3B)
Expression and Purification of GST Fusion Proteins
1. Preequilibrate the glutathione column, lyse the cells, and separate the lysate pellet, which includes the inclusion bodies, and supernatant (see Basic Protocol 2, steps 1 to 9).
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2. Centrifuge washed pellet (see Basic Protocol 2, step 9) 20 min at 48,000 × g, 4°C. Decant the supernatant and resuspend the pellet in 12 ml freshly prepared U buffer per 600 ml original culture. Incubate 2 hr on ice. Pellets should be resuspended in 20 ìl U buffer per ml of culture.
3. Centrifuge 20 min at 48,000 × g, 4°C. Carefully transfer the supernatant to a clean 50-ml centrifuge tube. The extracted fusion protein should now be in the supernatant.
4. Add Triton X-100 to the supernatant to give a final concentration of 1% (v/v). 5. Dialyze the sample (APPENDIX 3B) 2 to 3 hr in PBS/glycerol buffer. Dialysis buffer volume should be 20 times the sample volume.
6. Dialyze sample overnight in PBS/EDTA/PMSF. Dialysis buffer volume should be >100 times the sample volume.
7. Remove the sample from the dialysis bag and centrifuge 20 min at 4000 × g, 4°C. 8. Column purify the fusion protein (see Basic Protocol 2, steps 11 to 15). BATCH PURIFICATION OF GST FUSION PROTEIN Soluble glutathione-S-transferase (GST) fusion proteins in cell lysate supernatants or renatured proteins extracted from inclusion bodies can be batch purified on glutathione– Sepharose 4B as an alternative to column-based purification (see Basic Protocol 2 and see Alternate Protocol 1). Batch purification requires less equipment and is relatively quick and easy, but resulting protein yield and sample purity are lower than in a chromatographic separation. In addition, the room temperature incubations recommended by the resin manufacturer (Pharmacia Biotech), especially the batch incubation of E. coli lysate supernatant or inclusion body extract with glutathione-Sepharose, increase the risk of proteolytic degradation of the fusion protein.
ALTERNATE PROTOCOL 2
Prepare fusion protein 1. Extract soluble fusion proteins (see Basic Protocol 2, steps 5 to 10). If the fusion protein is in inclusion bodies, lyse the cells and collect the pellet (Basic Protocol 2, steps 5 to 9, followed by Alternate Protocol 1, steps 2 to 7).
Prepare resin slurry 2. Prepare a 50% slurry of glutathione–Sepharose 4B in PBS: For each milliliter of bed volume, centrifuge 1.33 ml of a 75% slurry of glutathione–Sepharose 4B for 5 min at 500 × g, room temperature. Discard the supernatant. Wash with 10 bed volumes PBS. Invert tube containing resin several times to mix, then centrifuge 5 min at 500 × g, room temperature, and remove supernatant. Add 1 ml PBS for each 1.33 ml of original slurry. Mix well before using. Bind fusion protein to resin 3. Add 2 ml of 50% slurry of equilibrated glutathione-Sepharose to each 100 ml bacterial lysate supernatant. Incubate 30 min at room temperature with gentle agitation on a platform or orbital shaker. 4. Centrifuge the suspension 5 min at 500 × g, room temperature. Remove supernatant and save at 0° to 4°C for later analysis by SDS-PAGE (UNIT 10.1) to determine the efficiency of binding of the fusion protein to the resin.
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5. Wash the pellet with 10 bed vol PBS. A bed volume is one-half the volume of the 50% slurry that was added to the column.
6. Centrifuge the suspension 5 min at 500 × g, room temperature. Discard the supernatant. 7. Repeat the wash and centrifugation steps for a total of three washes with 10 bed volumes each of PBS. Elute fusion protein 8. Elute the bound fusion protein by gently resuspending the sedimented resin in 1.0 ml glutathione buffer per milliliter resin bed volume. Incubate 10 min at room temperature with gentle agitation. 9. Centrifuge the suspension 5 min at 500 × g, room temperature. Transfer supernatant to a separate tube. 10. Repeat the elution and centrifugation (steps 8 to 9) a total of three times. Store at 0° and 4°C. The supernatants may be pooled into one tube or analyzed separately by SDS-PAGE (UNIT 10.1) to monitor for fusion protein content. The yield of fusion protein can be monitored by measuring the absorbance at 280 nm (A280). The extinction coefficient will partially depend on the absorbance characteristics of the experimental component of the fusion protein. For the GST moiety alone, the concentration can be estimated using 1 A280 = ∼0.6 mg/ml protein. As noted above, batch purification at room temperature increases the risk of proteolytic digestion of the target protein. To minimize such degradation, this procedure can alternatively be performed in a cold room at 4°C with incubation times in steps 3 and 8 increased 2- to 4-fold. BASIC PROTOCOL 3
PROTEASE CLEAVAGE OF FUSION PROTEIN IN SOLUTION TO REMOVE GST AFFINITY TAG The glutathione-S-transferase (GST) affinity tag is removed by cleaving with thrombin (pGEX-T vectors) or factor Xa (pGEX-X vectors). Conditions for optimal cleavage of each recombinant fusion protein must be empirically determined. Some of the parameters that can be varied include temperature, enzyme-to-substrate ratio, length of incubation, and buffer conditions. Proteolysis can usually be performed in the glutathione buffer used to elute the fusion protein from the column, either with or without addition of NaCl or Ca2+. After proteolysis, the glutathione must be removed by dialysis prior to rechromatography on the glutathione column to separate the cleaved target protein from the GST moiety and uncleaved fusion protein. Materials Solution of affinity-purified fusion protein (see Basic Protocol 2, step 15, or Alternate Protocol 2, step 10) Thrombin (Sigma) reconstituted in water at 0.5 U/µl or 1 µg/µl factor Xa (Boehringer Mannheim) 0.15 M PMSF in isopropanol PBS/EDTA/PMSF buffer (see recipe) Beckman J6-B centrifuge and JS-4.2 rotor (or equivalent), 4°C
Expression and Purification of GST Fusion Proteins
Additional reagents and equipment for dialysis (APPENDIX 3B) and SDS-PAGE (UNIT 10.1)
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1. To the solution of affinity-purified fusion protein, add an appropriate amount of thrombin (when using pGEX-T vectors) or factor Xa (when using pGEX-X vectors) per microgram of purified fusion protein and digest protein 2 to 8 hr in a shaking water bath at 37°C (thrombin) or 25°C (factor Xa). The appropriate amount of enzyme must first be empirically determined for each fusion protein by surveying a range of conditions in a pilot analytical proteolysis experiment. A convenient method is to digest 100 ìg fusion protein per condition over a range of enzyme-to-substrate ratios and for differing incubation times. Typical digestion times range from 2 to 8 hr and typical enzyme-to-substrate-ratios are 1:100, 1:350, 1:1000, and 1:3000 (in units of enzyme per microgram fusion protein) at 37°C for bovine plasma thrombin, or 1:10, 1:25, 1:50, 1:100, 1:300 (microgram enzyme per microgram fusion protein) at 25°C for bovine plasma factor Xa. At the desired times, proteolysis is stopped by adding an aliquot of the sample to boiling SDS sample buffer, and samples are analyzed on an SDS minigel (UNIT 10.1; 2 ìg/lane) to determine the best digestion conditions. If digestion is incomplete at the highest enzyme-to-substrate ratio tested with digestion times of 6 to 8 hr, consider reengineering the protease cleavage site by introducing a linker between the GST moiety and target polypeptide to decrease steric hindrance. In some cases, >10-fold improvement in cleavage efficiency can be achieved by adding as few as two glycines next to the thrombin site. The SDS gel in Figure 6.6.6 shows a typical thrombin digestion optimization experiment where the amount of enzyme has been varied.
2. Stop preparative digestion by adding a 1:500 dilution of 0.15 M PMSF stock solution. Incubate sample an additional 15 min at 37°C for thrombin or 30 min at 25°C for factor Xa to covalently inhibit the enzyme with the PMSF. 3. Dialyze the sample (APPENDIX 3B) twice versus 2 liters PBS/EDTA/PMSF for a minimum of 4 hr per buffer change at 4°C. Complete removal of glutathione is important if samples are to be rechromatographed on glutathione-Sepharose to remove the GST moiety and uncleaved fusion protein. Larger volumes of dialysate may be necessary depending on sample volume: e.g., if the sample volume is >40 ml, increase the dialysis buffer volume to 4 liters per change or use three changes of buffer. In addition, since glutathione equilibrates slowly during dialysis, when
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Figure 6.6.6 SDS gel stained with Coomassie brilliant blue showing pilot thrombin digestions of two recombinant glutathione-S-transferase (GST) fusion proteins. Samples were digested 3 hr at 37°C in buffer with varying enzyme-to-substrate ratios. Lane 1, a 78-kDa fusion protein. Lanes 2 to 5, thrombin digestion of the 78-kDa fusion protein using enzyme-to-substrate ratios of 1:3000, 1:1000, 1:350 and 1:100, respectively. Lane 6, a 45-kDa fusion protein. Lanes 7 to 10, thrombin digestion of the 45-kDa fusion protein using substrate ratios of 1:3000, 1:1000, 1:350, and 1:100, respectively. In each case, an enzyme-to-substrate ratio of 1:1000 was chosen as the optimal digestion condition
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dialysis tubing with a MWCO <12,000 is used, the dialysis conditions should be increased to at least three changes with 2 liters dialysis buffer per change.
4. Centrifuge the dialyzed sample 20 min at 4000 × g, 4°C, to remove any precipitated material. Transfer the supernatant to a clean tube at 0° to 4°C and analyze by SDS-PAGE (UNIT 10.1). The cleaved protein of interest can be separated from the GST moiety and uncleaved fusion protein by rechromatographing it on the glutathione column (see Support Protocol 1). ALTERNATE PROTOCOL 3
PROTEASE CLEAVAGE OF FUSION PROTEIN BOUND TO A GLUTATHIONE-SEPHAROSE COLUMN The target polypeptide can be cleaved from the glutathione-S-transferase (GST) moiety while the fusion protein is immobilized on the affinity matrix as described by Gearing et al. (1989). The fusion protein is loaded onto the glutathione column after cell lysis in the normal manner. Rather than elute the fusion protein with glutathione buffer, a solution of thrombin (or factor Xa) is loaded onto the column and allowed to incubate at room temperature for several hours. The cleaved protein is then eluted from the column by washing with PBS. This method is much faster and can yield higher purity products than batch protease digestions and in some cases, secondary cleavages can be avoided or minimized using the on-column method. Disadvantages of this method include much less effective protease cleavage, much lower final polypeptide yields, decreased control of digestion conditions, and inability to easily monitor the extent of cleavage using on column digestion. Additional Materials (also see Basic Protocol 3) PBS (APPENDIX 2E) Glutathione–Sepharose 4B column containing bound fusion protein (see Basic Protocol 2, step 13) Glutathione buffer (see recipe) 1. For each milliliter of glutathione–Sepharose 4B column containing bound fusion protein, dilute desired amount of reconstituted thrombin to 1.0 ml using PBS. Enzyme concentrations must be empirically determined for each fusion protein (see Basic Protocol 3, step 1 annotation).
2. Load the thrombin reaction mixture onto the column. When the reaction mixture has been added, replace the bottom cap and allow the column to stand at room temperature for 2 to 16 hr. Incubation times must be empirically determined for each fusion protein.
3. Elute the protein of interest by washing the column with PBS and immediately add an aliquot of 0.15 M PMSF stock solution to give a final PMSF concentration of 0.3 mM (e.g., 6 µl PMSF stock solution per 3.0-ml fraction). The protein of interest will be in the flowthrough and the GST moiety and undigested fusion protein will remain bound to the glutathione-Sepharose.
4. Remove the GST moiety and uncleaved fusion protein from the column by washing with 5 bed volumes glutathione buffer at 0.3 ml/min. 5. Immediately after collecting fractions, store all fractions at 0° to 4°C. Expression and Purification of GST Fusion Proteins
6. Evaluate unbound and bound peaks using SDS-PAGE (UNIT 10.1). A large proportion of the fusion protein is usually not cleaved by this method.
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BATCH PROTEASE CLEAVAGE AND SEPARATION OF FUSION PROTEIN ON GLUTATHIONE RESIN
ALTERNATE PROTOCOL 4
As a subsequent step to batch purification of soluble glutathione-S-transferase (GST) fusion proteins (see Alternate Protocol 2), the GST affinity tag can be cleaved from the protein of interest using thrombin (pGEX-2T) or factor Xa (pGEX-3X) before elution from the glutathione–Sepharose 4B resin in a manner analogous to the on-column cleavage described in Alternate Protocol 3. In this protocol, the fusion protein on the glutathione-Sepharose resin is incubated in the presence of protease, and the cleaved protein is released into the supernatant and collected by centrifugation. Additional Materials (also see Basic Protocol 3) Glutathione–Sepharose 4B resin with bound fusion protein (see Alternate Protocol 2, step 7) PBS (APPENDIX 2E) 1. For each milliliter of glutathione–Sepharose 4B resin with bound fusion protein, dilute the desired amount of reconstituted thrombin to 1.0 ml with PBS. Enzyme concentrations must be empirically determined for each fusion protein (see Basic Protocol 3, step 1 annotation).
2. Add the reconstituted thrombin reaction mixture to the pelleted glutathioneSepharose resin containing the bound fusion protein (from Alternate Protocol 2, step 7). 3. Gently resuspend the glutathione-Sepharose and agitate on an orbital shaker 2 to 16 hr at room temperature. Incubation times must be empirically determined for each fusion protein. Cleavage may be monitored by removing an aliquot of the slurry from the incubation mixture at different time points, centrifuging the slurry to separate the resin and supernatant, and analyzing the fractions by SDS-PAGE (UNIT 10.1).
4. Centrifuge the slurry 5 min at 500 × g, room temperature. Transfer supernatant to a separate tube. The supernatant will contain the protein of interest, while the GST moiety and uncleaved fusion protein will remain bound to the resin.
5. Inhibit the reaction by adding a 1:500 dilution of 0.15 M PMSF stock solution (0.3 mM PMSF final). Incubate the sample an additional 15 min at 37°C for thrombin or an additional 30 min at 25°C for factor Xa to covalently inhibit the enzyme with the PMSF. Store the samples at 0° to 4°C. 6. Analyze the final fractions by SDS-PAGE (UNIT 10.1). AFFINITY CHROMATOGRAPHY PURIFICATION OF POLYPEPTIDES AFTER ENZYMATIC CLEAVAGE
SUPPORT PROTOCOL 1
The target polypeptide is affinity purified by rechromatography on a glutathione– Sepharose 4B column after enzymatic cleavage of the fusion protein and dialysis to remove glutathione present in the buffer from the initial column eluate. Uncleaved fusion protein and the glutathione-S-transferase (GST) moiety both bind to the column, and the protein of interest is in the column flowthrough. All steps should be performed in a cold room at 4°C unless otherwise indicated. Purification of Recombinant Proteins
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Materials Solution of fusion protein that has been cleaved with thrombin or factor Xa and dialyzed into PBS/EDTA/PMSF buffer (see Basic Protocol 3, step 4) Glutathione buffer (see recipe) PBS/EDTA/PMSF buffer (see recipe) 2.5 × 8–cm glutathione–Sepharose 4B column (e.g., Bio-Rad Econo) Additional reagents and equipment for SDS-PAGE (UNIT 10.1) 1. Wash the glutathione–Sepharose 4B column with >3 bed volumes glutathione buffer at 1.5 ml/min. A bed volume is one-half the amount of glutathione-Sepharose that was added to the column. The same 2.5 × 8–cm column used for initial purification of a given fusion protein can be used for repurification of the same target polypeptide after protease cleavage. GlutathioneSepharose must be fully reduced in order for the GST moiety to bind efficiently. If the column was washed with glutathione buffer <48 hr before, this step may be skipped.
2. Wash the glutathione column with 10 bed vol PBS/EDTA/PMSF at 1.5 ml/min. 3. Load the solution of fusion protein that has been cleaved onto the column using a flow rate of 0.1 ml/min. Collect fractions for later analysis by SDS-PAGE (UNIT 10.1) and store at 0° to 4° C. The protein of interest will be in the unbound or column flowthrough peak, and the GST moiety will bind to the glutathione-Sepharose. A low flow rate for sample application, such as 0.1 ml/min, should be used to ensure complete binding of GST to the column. Faster flow rates are likely to result in elution of excessive amounts of the GST moiety in the unbound fraction.
4. Wash the column with 2 to 3 bed volumes PBS/EDTA/PMSF at 1.5 ml/min. 5. Elute the bound GST moiety from the resin by washing with 5 bed volumes glutathione buffer at 0.3 ml/min. 6. Analyze bound and unbound peaks by SDS-PAGE (UNIT 10.1), pool the fractions containing the cleaved target polypeptide, and store at 0° to 4°C. SUPPORT PROTOCOL 2
Expression and Purification of GST Fusion Proteins
FURTHER PURIFICATION OF CLEAVED RECOMBINANT PROTEIN USING HPLC GEL FILTRATION Cleaved proteins after rechromatography on glutathione columns (see Support Protocol 1) or eluted after on-column protease cleavage (see Alternate Protocol 3) contain the inactivated protease used to cleave the protein. These samples also typically contain small amounts of aggregates, a small amount of glutathione-S-transferase (GST) moiety that did not rebind to the column, and possibly other minor contaminants, including proteolytic cleavage products or host cell proteins. Although the protein is typically ∼90% pure at this stage, a further purification step may be desirable for many applications. HPLC gel filtration is a fairly rapid high-resolution purification step that usually separates properly folded polypeptides from aggregates and incorrectly folded chains. Selection of appropriate matrices and column dimensions for particular applications is discussed in UNIT 8.3. Alternatively, ion-exchange chromatography (UNIT 8.2) can be used as a further purification strategy, either instead of or in addition to HPLC gel filtration. This protocol describes the HPLC gel filtration purification of cleaved polypeptides after rechromatography. All steps, including HPLC, should be performed at 4°C unless otherwise indicated.
6.6.16 Supplement 9
Current Protocols in Protein Science
Materials Cleaved fusion protein (see Support Protocol 1, step 6, or Alternate Protocol 3, step 5) PBS (APPENDIX 2E) Centriprep concentrator with appropriate MWCO (Amicon) Low-speed refrigerated centrifuge (e.g., Beckman J6-B and JS-4.2 rotor or equivalent, 4°C), or 0.22-µm low-protein-binding filter (Costar) Gel-filtration columns (see UNIT 8.3) Additional reagents and equipment for SDS-PAGE (UNIT 10.1) 1. Concentrate the unbound cleaved fusion protein using a Centriprep concentrator. Samples should be concentrated to a volume of no greater than 0.5% to 1% of the column volume to be used for the separation.
2. Centrifuge the concentrated sample 20 min at 4000 × g, 4°C. Remove the supernatant, being careful not to disturb any pellet that may have formed (precipitated material formed during concentration). Alternatively, a 0.22-ìm low-protein-binding filter may be used to remove particulates. It is especially important to remove particulates to avoid clogging of the HPLC column end frits.
3. Inject the concentrated sample onto gel filtration column(s) equilibrated in PBS. 4. Monitor the absorbance at 280 nm (A280) using an online HPLC detector and collect fractions. 5. Analyze the fractions by SDS-PAGE (UNIT 10.1) to determine fractions of interest and to evaluate the purity of the target polypeptide, which should be in the major peak. 6. Pool the desired fractions and store at 0° to 4°C. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or its equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Ampicillin, 5 mg/ml 500 mg ampicillin 100 ml H2O Filter sterilize using 0.22-µm filter Store up to several months at 4°C Do not autoclave the solution; ampicillin is inactivated above 50°C.
Glutathione buffer 50 mM Tris base 10 mM reduced glutathione (Sigma) Adjust pH to 8.0 with 6 M HCl Bring to final volume with cold H2O Prepare fresh daily and store at 4°C until needed Isopropyl-1-thio-â-D-galactopyranoside (IPTG), 100 mM 1 g IPTG 42 ml sterile H2O Divide into 6-ml aliquots in sterile tubes Store up to 1 year at −20°C
Purification of Recombinant Proteins
6.6.17 Current Protocols in Protein Science
Supplement 9
Lysis buffer 50 mM NaCl 50 mM Tris base 5 mM EDTA 1 µg/ml leupeptin 1 µg/ml pepstatin 0.15 mM phenylmethylsulfonyl fluoride (PMSF) in isopropanol 1 mM diisopropyl fluorophosphate (DFP) Adjust pH to 8.0 with 6 M HCl Bring to final volume with cold H2O Prepare fresh daily CAUTION: DFP is a dangerous neurotoxin. Handle the neat reagent with double gloves in a chemical fume hood only. Carefully follow all precautions supplied by the manufacturer for this chemical.
PBS/EDTA 1× PBS (APPENDIX 2E) 5 mM EDTA Adjust pH to 7.4 with 1 M NaOH Bring to final volume with cold H2O Store up to 1 month at 4°C PBS/EDTA/PMSF buffer 1× PBS (APPENDIX 2E) 5 mM EDTA 0.15 mM phenylmethylsulfonyl fluoride (PMSF) in isopropanol Adjust pH to 7.4 with 1 M NaOH Bring to final volume with cold H2O Store up to 1 week at 4°C PBS/glycerol buffer 1× PBS (APPENDIX 2E) 20% (v/v) glycerol 1% (v/v) Triton X-100 5 mM 2-mercaptoethanol (2-ME) 5 mM EDTA 0.1 µg/ml leupeptin 0.1 µg/ml pepstatin 0.15 mM phenylmethylsulfonyl fluoride (PMSF) in isopropanol 0.1 mM diisopropyl fluorophosphate (DFP) Adjust pH to 7.4 with 1 M NaOH Bring to final volume with cold H2O Prepare fresh daily CAUTION: DFP is a dangerous neurotoxin. Handle the neat reagent with double gloves in a chemical fume hood only. Carefully follow all precautions supplied by the manufacturer for this chemical.
Expression and Purification of GST Fusion Proteins
U buffer 5 M urea 50 mM Tris base 5 mM EDTA 5 mM 2-mercaptoethanol (2-ME) 1 µg/ml leupeptin 1 µg/ml pepstatin continued
6.6.18 Supplement 9
Current Protocols in Protein Science
0.15 mM phenylmethylsulfonyl fluoride (PMSF) in isopropanol 1 mM diisopropyl fluorophosphate (DFP) Adjust pH to 8.0 with 6 M HCl Bring to final volume with cold H2O Prepare fresh daily CAUTION: DFP is a dangerous neurotoxin. Handle the neat reagent with double gloves in a chemical fume hood only. Carefully follow all precautions supplied by the manufacturer for this chemical.
Wash buffer 50 mM Tris base 5 mM EDTA 1 µg/ml leupeptin 1 µg/ml pepstatin 0.15 mM phenylmethylsulfonyl fluoride (PMSF) in isopropanol Adjust pH to 8.0 with 6 M HCl Bring to final volume with cold H2O Prepare fresh daily COMMENTARY Background Information The pGEX vectors were designed for highlevel, inducible, intracellular expression of glutathione-S-transferase (GST) fusion proteins produced in Escherichia coli. GST is a common 26-kDa protein of eukaryotes. The GST gene used in the development of the pGEX vectors was originally cloned from the parasitic helminth Schistosoma japonicum (Smith and Johnson, 1988). Purification of fusion proteins from a wholecell lysate is readily achieved through the strong affinity of the GST moiety for glutathione, which is immobilized on Sepharose beads. The fusion protein can be displaced under mild conditions from the glutathioneSepharose beads using neutral-pH buffers containing free reduced glutathione. The main advantages of this system are the very high level of fusion protein expression (often >10 to 50 mg/liter of E. coli culture grown on an environmental shaker) and the facile purification methods for both initial isolation and subsequent separation of cleaved polypeptide and GST moiety. Since nondenaturing purification conditions are employed, polypeptides that do not normally contain posttranslational modifications usually retain their functional and antigenic properties. Other advantages of this system include availability of several alternative protease cleavage sites and the large number of bacterial hosts that can be used. Purification of most soluble GST fusion proteins is straightforward, and success in the
purification of insoluble products is determined largely by the ability to refold the fusion protein after extracting it from inclusion bodies. Since the GST moiety refolds fairly readily after urea solubilization, the presence of this moiety may facilitate refolding of the adjacent polypeptide. However, the high protein expression level of the vector, even at lower growth temperatures means that the preferred strategy for solubilizing proteins that are initially expressed in inclusion bodies is to attempt to grow the transfected cells at lower temperatures, where the protein often remains in the soluble fraction (see Fig. 6.6.5 and Troubleshooting). Difficulties with vector construction, protein expression, and protease cleavage may be greater when the molecular weight of the desired polypeptide is >100 kDa. The GST moiety can usually be easily removed from the protein of interest by cleavage with thrombin or factor Xa. For some studies, such as some immunological or functional assays, removal of the GST moiety may not be required. An important consideration is that the GST moiety is a dimer; hence, steric hindrance effects and nonequivalence of functional sites on the associated fusion partner within this dimeric structure must be considered.
Troubleshooting Contamination of the glutathione-S-transferase (GST) fusion protein after affinity purification with E. coli host cell proteins is usually an indication that sonication has been too severe. Other contaminants may represent de-
Purification of Recombinant Proteins
6.6.19 Current Protocols in Protein Science
Supplement 9
Expression and Purification of GST Fusion Proteins
graded fragments of the fusion protein that might be eliminated by the addition of protease inhibitors and/or carefully keeping all samples, buffers, and sample tubes as cold as possible (i.e., keeping the materials at 0°C, on ice, often results in noticeably lower proteolysis than occurs where samples are kept at 4°C in a refrigerator). Degradation of fusion proteins during expression can sometimes be minimized by adding isopropyl-1-thio-β-D-galactopyranoside (IPTG) later in the bacterial culture growth with cells at higher OD at time of induction and by decreasing the duration of the induction period. If the fusion protein does not completely bind to the glutathione-Sepharose resin, several options may be tried: increasing the quantity of glutathione-Sepharose, decreasing the protein loading flow rate, ensuring that the column is completely reduced by pretreating it with a freshly prepared glutathione buffer within 24 hr prior to using the column for purification, and ensuring that any denaturants or reducing reagents used to treat the sample have been completely removed through exhaustive dialysis. Expression of GST fusion protein in inclusion bodies can be addressed in several ways. The preferred method is to lower the growth temperature from 37° to 30°, 25°, 20°, or 15°C. The optimal growth temperature is that temperature where >50% of the expressed protein is in the supernatant. This strategy is efficient even if the expressed fusion protein yield decreases somewhat at the lower growth temperatures. It is usually more time and cost effective to grow a larger quantity of cells than to attempt to renature fusion protein extracted from inclusion bodies since the latter method nearly doubles the time required to purify the fusion protein. In one study, when the authors’ laboratory expressed eleven GST fusion proteins at 37°C, four proteins were primarily in inclusion bodies and the remaining seven proteins were partially in the supernatant, although a substantial amount of fusion protein was also in inclusion bodies. When these eleven fusion proteins were expressed at 30°C, ten were primarily (>80%) in the soluble fraction. The remaining recombinant protein was primarily in inclusion bodies at either 30° or 25°C, but could be switched to the soluble fraction and produced in high yield by growing the cells at 18°C, as shown in Figure 6.6.5. If lowering the growth temperature does not result in expression of a soluble protein, renaturing the protein after extracting inclusion bodies with a chaotropic reagent such as urea
can be attempted (see Alternate Protocol 1). Finally, even when most of a protein is primarily in inclusion bodies, there may be enough protein in the soluble fraction that it can be purified in low milligram yields per liter of culture. In some cases, using high enzyme-to-substrate ratios and prolonged incubation times is not sufficient to cleave the fusion protein in good yield. To minimize steric hindrance at the thrombin cleavage site, modified pGEX vectors that contain a glycine-rich, nine-amino-acid “kinker” region immediately before or after the thrombin cleavage site (Guan and Dixon, 1991; Hakes and Dixon, 1991) can be tried. Sometimes introduction of as few as one or two extra glycines immediately after to the thrombin site may substantially improve the rate of protease cleavage. The probability that extraneous residues on the N-termini of cleaved polypeptides could significantly affect structural and functional properties of expressed polypeptides is very likely to increase as the length of the extraneous sequence increases. Hence, the number of extra residues remaining on the target polypeptide after protease cleavage should be minimized. Although thrombin cleavage is usually more cost effective than factor Xa cleavage, a minimum of two extra residues from the thrombin recognition site (GS) are left on the cleaved peptide. Although no residues from the factor Xa recognition site are left on the cleaved polypeptide, at least several extraneous amino acids are introduced onto the N-terminal of all pGEX-X vectors from the sequence remaining at the cloning site (see Figs. 6.6.2 and 6.6.3). Depending upon the pGEX vector and cloning site utilized, as few as two and as many as thirteen extra amino acids may remain on the cleaved N-terminal end of the target protein. The yield of GST fusion proteins is typically quite high, i.e., 10 to 50 mg of fusion protein per liter of culture when cells are grown on an environmental shaker. When low protein expression does occur, it can often be improved by optimizing the bacterial culture growth conditions: for example, growing the cells to a somewhat higher OD550 at the time of induction, increasing the length of the induction period, using another strain of E. coli such as a protease-negative strain, or growing the cells at higher temperatures (although proteins are more likely to be in inclusion bodies at higher temperatures; see Background Information). Caution should be taken when using higher cell ODs and longer growth times after induction,
6.6.20 Supplement 9
Current Protocols in Protein Science
since the cells may start to lyse under these conditions. For additional information concerning GST fusion protein expression, Pharmacia Biotech publishes a reference list, bulletin number 181117-27, that classifies related publications into several useful categories to assist in troubleshooting a variety of parameters.
Anticipated Results Yields of fusion protein can vary widely. Typical yields are 10 to 50 mg/liter, but can occasionally be much lower, especially if the fusion protein is toxic to the cells or is unstable. In some cases, >50 mg/liter can be obtained when expression conditions have been well optimized. A single-step affinity purification should yield fusion protein that is >90% pure in most cases. The relationship between yield of fusion protein and yield of cleaved, repurified recombinant target polypeptide is in part due to the mass of the target polypeptide. A good final yield of a cleaved protein after repurification on a glutathione column followed by HPLC gel filtration might be ∼2 mg/liter for a 10-kDa protein and ∼10 mg/liter for a 50-kDa protein.
Time Considerations Protein expression takes 1.5 days of intermittent work requiring approximately 3 to 4 hr of operator time. Longer induction periods may be required at temperatures lower than 30°C, but total operator time remains the same. A small-scale batch purification can be completed in 1 day. A large-scale column purification, cleavage of fusion protein, and repurification of the cleaved peptide by affinity and gel filtration chromatography will take ∼5 to 8 days of intermittent work requiring several hours of operator time per day. The purification should be completed in as short a time as practical to minimize proteolysis, aggregation, and precipitation of impure fractions.
Literature Cited Ausubel, F.M., Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G., Smith, J.A., and Struhl, K. (eds.). 1994. Current Protocols in Molecular Biology. John Wiley & Sons, New York. Davies, A.H., Jowett, J.B.M., Jones, I.M. 1993. Recombinant baculovirus vectors expressing glutathione-S-transferase fusion proteins. Bio/Technology 11:933-936. Frangioni, J.V. 1992. Solubilization and purification of enzymatically active glutathione-S-transferase (pGEX) fusion proteins. Anal. Biochem. 210:179-187. Gearing, D.P., Nicola, N.A., Metcalf, D., Foote, S., Willson, T.A., Gough, N.M., and Williams, R.L. 1989. Production of leukemia inhibitory factor in Escherichia coli by a novel procedure and its use in maintaining embryonic stem cells in culture. Bio/Technology 7:1157-1161. Grieco, F., Hull, J., and Hull, R. 1992. An improved procedure for the purification of protein fused with glutathione-S-transferase. Biotechniques 13:856-857. Guan, K.L. and Dixon, J.E. 1991. Eukaryotic proteins expressed in Escherichia coli: An improved thrombin cleavage and purification procedure of fusion proteins with glutathione-S-transferase. Anal. Biochem. 192:262-267. Hakes, D.J. and Dixon, J.E. 1991. New vectors for high level expression of recombinant proteins in bacteria. Anal. Biochem. 202:293-298. Mitchell, D.A., Marshall, T.K., and Deschenes, R.J. 1993. Vectors for the overexpression of glutathione-S-transferase fusion proteins in yeast. Yeast 9:715-722. Sambrook, J., Fritsch, E.F., and Maniatis, T. 1989. Molecular Cloning: A Laboratory Manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. Smith, D.B. and Johnson, K.S. 1988. Single-step purification of polypeptides expressed in Escherichia coli as fusions with glutathione-Stransferase. Gene 67:31-40.
Key Reference Smith and Johnson, 1988. See above. Original description of the pGEX system.
Contributed by Sandra Harper and David W. Speicher The Wistar Institute Philadelphia, Pennsylvania
Purification of Recombinant Proteins
6.6.21 Current Protocols in Protein Science
Supplement 9
Expression and Purification of Thioredoxin Fusion Proteins
UNIT 6.7
This unit describes a gene fusion expression system that uses thioredoxin, the product of the Escherichia coli trxA gene, as the fusion partner. The system is particularly useful for high-level production of soluble fusion proteins in the E. coli cytoplasm; in many cases heterologous proteins produced as thioredoxin fusion proteins are correctly folded and display full biological activity. Although the thioredoxin gene fusion system is routinely used for protein production, high-level production of peptides—i.e., for use as antigens— is also possible because the prominent thioredoxin active-site loop is a very permissive site for the introduction of short amino acid sequences (10 to 30 residues in length). The inherent thermal stability of thioredoxin and its susceptibility to quantitative release from the E. coli cytoplasm by osmotic shock can also be exploited as useful tools for thioredoxin fusion protein purification. In addition, a more generic method for purification of any soluble thioredoxin fusion employs a modified form of thioredoxin (called “His-patch Trx”), which has been designed to bind to metal chelate resins. Protein fusions to His-patch Trx can usually be purified in a single step from cell lysates (see Strategic Planning). The first step is construction of a fusion of trxA to any desired gene and expression of the fusion protein in an appropriate host strain at 37°C (see Basic Protocol). Additional protocols describe E. coli cell lysis using a French pressure cell and fractionation (see Support Protocol 1), osmotic release of thioredoxin fusion proteins from the E. coli cytoplasm (see Support Protocol 2), and heat treatment to purify some thioredoxin fusion proteins (see Support Protocol 3). STRATEGIC PLANNING The thioredoxin gene fusion expression vectors pTRXFUS and hpTRXFUS, both of which carry the E. coli trxA gene (Fig. 6.7.1), are used for high-level production of C-terminal fusions to thioredoxin. The vector hpTRXFUS differs from pTRXFUS in that it contains a modified E. coli trxA gene which produces a mutant protein (“His-patch” thioredoxin) that can specifically bind to metal chelate matrices charged with nickel or cobalt, otherwise known as native metal-chelate affinity chromatography (MCAC; UNIT 9.4). The trxA translation-termination codon has been replaced in both vectors by DNA encoding a ten-residue peptide linker sequence that includes an enterokinase (enteropeptidase; LaVallie et al., 1993a) cleavage site. This highly specific site can be cleaved with enterokinase following purification of the fusion protein to release the protein of interest from its thioredoxin fusion partner. Immediately downstream of the DNA encoding the enterokinase site in pTRXFUS and hpTRXFUS lies a DNA polylinker sequence containing a number of unique restriction endonuclease sites that can be used for forming in-frame translational fusions of any desired gene to trxA. Downstream of the DNA polylinker lies the E. coli aspA transcription terminator. Replication of these vectors is controlled by a modified colE1 replication origin similar to that found in pUC vectors (Norrander et al., 1983). Plasmid selection and maintenance is ensured by the presence of the β-lactamase gene on the vector. The vector pALtrxA-781 (Fig. 6.7.1) is very similar to pTRXFUS. However in this plasmid the trxA gene is followed by a translation termination codon, and the sequences encoding the enterokinase-site peptide linker are absent. A unique RsrII site, present in both pALtrxA-781 and pTRXFUS, allows for the easy insertion of short peptide-encoding DNA sequences into trxA within the region that encodes the active-site loop. Contributed by John McCoy and Edward LaVallie Current Protocols in Protein Science (1997) 6.7.1-6.7.14 Copyright © 1997 by John Wiley & Sons, Inc.
Purification of Recombinant Proteins
6.7.1 Supplement 10
BLA
ori
pALtrxA–781 pTRXFUS p L hpTRXFUS
trxA aspA Rsr II
TGGTGCGGTCCGTGCAAA W C G33 P34 C K
Sfi I
pALtrxA–781:
Xba I
Sal I
Pst I
AACCTGGCCTAGCTGGCCATCTAGAGTCGACCTGCAG N L A *
aspA terminator
thioredoxin
Kpn I Bam HI Xba I
pTRXFUS:
Sal I
Pst I
AACCTGGCCGGTTCTGGTTCTGGTGATGACGATGACAAGGTACCCGGGGATCCTCTAGAGTCGACCTGCAG
N
L
thioredoxin
A
G
S
G
S
G
D
D
D
D
K
fusion point
aspA terminator
linker peptide enterokinase site
Figure 6.7.1 Thioredoxin gene fusion expression vectors pTRXFUS, hpTRXFUS, and pALtrxA-781. pALtrxA-781 contains a polylinker sequence at the 3′ end of the trxA gene. pTRXFUS and hpTRXFUS contain a linker region encoding a peptide that includes the enterokinase cleavage site between the trxA gene and the polylinker. The sequence surrounding the active site loop of thioredoxin has a single RsrII site that can be used to insert peptide coding sequence. The asterisk indicates a translational stop codon. Abbreviations: trxA, E. coli thioredoxin gene; BLA, β-lactamase gene; ori, colE1 replication origin; pL, bacteriophage λ major leftward promoter; aspA terminator, E. coli aspartate amino-transferase transcription terminator.
Expression and Purification of Thioredoxin Fusion Proteins
pTRXFUS, hpTRXFUS, and pALtrxA-781 carry the strong bacteriophage λ promoter pL (Shimatake and Rosenberg, 1981) positioned upstream of the trxA gene. Transcription initiation at the pL promoter is controlled by the intracellular concentration of λ repressor protein (cI). cI857-containing strains (Shatzman et al., 1990) can be used for heat inductions of pL at 42°C; alternatively, in the strains carrying the wild-type repressor, pL can be induced by a prior induction of the E. coli SOS stress response. However, it is often desirable to express heterologous genes in E. coli at temperatures considerably lower than 42°C, or under conditions where cells are not undergoing a physiological stress. Strains GI698, GI724 and GI723 were designed to allow the growth and induction of pL expression vectors, including pTRXFUS, hpTRXFUS, and pALtrxA-781, under mild conditions over a wide range of temperatures (see Table 6.7.1; Mieschendahl et al., 1986). Each of these strains carries a wild-type allele of cI stably integrated into the E. coli chromosome at the nonessential ampC locus. A synthetic trp promoter integrated into ampC upstream of the cI gene in each strain directs the synthesis of cI repressor only when intracellular tryptophan levels are low. When tryptophan levels are high, synthesis of cI is switched off; therefore, the presence of tryptophan in the growth medium of GI698, GI723, or GI724 will block expression of λ repressor and thus will turn on pL. Because the three strains carry ribosome-binding sequences of different strengths at the 5′-end of
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Current Protocols in Protein Science
Table 6.7.1 E. coli Strains for Production of Thioredoxin Fusion Proteins at Varying Temperatures
Strain
Desired production temperature (°C)
Pre-induction growth temperature (°C)
Induction period (hr)
GI698
15
25
20
GI698 GI698
20 25
25 25
18 10
GI724 GI724
30 37
30 30
6 4
GI723
37
37
5
their respective cI genes, they maintain intracellular concentrations of λ repressor that increase in the order GI698 < GI724 < GI723. The choice of which strain to use for a particular application is dependent on the desired culture conditions as described below. Although some thioredoxin fusion proteins produced at 37°C are insoluble, expression at lower temperatures can often result in the fusion protein being produced in a soluble form. Each of the three pL host strains GI698, GI723, and GI724 is suitable for the production of thioredoxin fusion proteins over a particular temperature range. Table 6.7.1 indicates the correct strain for expression of thioredoxin fusion proteins at any temperature between 15°C and 37°C. The induction protocol at any of these temperatures is the same as that described for induction of GI724 at 37°C (see Basic Protocol), except the preinduction growth temperature and the length of the induction period vary according to the strain used and the temperature chosen. Cultures should be grown at the indicated preinduction growth temperature until they reach a density of 0.4 to 0.6 OD550/ml. They should then be moved to the desired induction temperature and induced by the addition of 100 µg/ml tryptophan. Low-temperature inductions are best performed in strain GI698. However, this strain makes only enough cI repressor protein to maintain the vectors in an uninduced state at temperatures below 25°C. GI698 should therefore never be grown above 25°C when it carries a pL plasmid. A nonrefrigerated water bath can be maintained below room temperature by placing it in a 4°C room and setting the thermostat to the desired temperature. It is often a good idea to collect time points during the course of a long induction period and to fractionate cells from these time points (see Support Protocol 1, steps 9 to 13). Although a particular fusion protein may be soluble during the early part of an induction, during the later phases of induction it may become unstable or its concentration inside the cell may exceed a critical threshold above which it will precipitate and appear in the insoluble fraction.
Purification of Recombinant Proteins
6.7.3 Current Protocols in Protein Science
Supplement 10
BASIC PROTOCOL
CONSTRUCTION AND EXPRESSION OF A THIOREDOXIN FUSION PROTEIN This protocol describes construction and subsequent expression of a gene fusion between trxA (encoding thioredoxin) and a gene encoding a particular protein or peptide. After a clone carrying the correct fusion sequence is constructed, analyzed, and isolated, cultures are grown and expression is induced. The protocol is described in terms of the E. coli host strain GI724 with expression at 30°C; it may also be applied to strains GI698 and GI723 (also available from Genetics Institute) for expression at other temperatures by using the parameters specified in Table 6.7.1 (see Strategic Planning). Materials DNA fragment encoding desired sequence Thioredoxin expression vectors (Fig. 6.7.1): pTRXFUS or pALtrxA-781 (Genetics Institute or Invitrogen) or hpTRXFUS (Genetics Institute) E. coli strain GI724 (Genetics Institute or Invitrogen), grown in LB medium and made competent LB medium (UNIT 5.2) IMC plates (see recipe) containing 100 µg/ml ampicillin CAA/glycerol/ampicillin 100 medium (see recipe) IMC medium (see recipe) containing 100 µg/ml ampicillin 10 mg/ml tryptophan (see recipe) SDS-PAGE sample buffer (see recipe) 30°C convection incubator 18 × 50–mm culture tubes Roller drum (New Brunswick Scientific) 250-ml culture flask 70°C water bath Microcentrifuge, 4°C Additional reagents and equipment for SDS-PAGE (UNIT 10.1), and Coomassie brilliant blue staining (UNIT 10.5) Construct the trxA gene fusion 1. Use DNA fragment encoding the desired sequence to construct either an in-frame fusion to the 3′-end of the trxA gene in pTRXFUS or hpTRXFUS, or a short peptide insertion into the unique RsrII site of pALtrxA-781. A precise fusion of the desired gene to the enterokinase linker sequence in pTRXFUS or hpTRXFUS can be made by using the unique KpnI site trimmed to a blunt end with the Klenow fragment of E. coli DNA polymerase. The desired gene can usually be adapted to this blunt-end construct by using a synthetic oligonucleotide duplex ligated between it and any convenient downstream restriction site close to the 5′ end of the gene. When designing the fusion junction, note that enterokinase is able to cleave —DDDDK↓X—, where X is any amino acid residue except proline. Synthetic oligonucleotides encoding short peptides for insertion into the thioredoxin active-site loop at the RsrII site will insert only in the desired orientation, because the RsrII sticky end consists of three bases.
2. Transform the ligation mixture containing the new thioredoxin fusion plasmid into competent GI724 cells. Plate transformed cells onto IMC plates containing 100 µg/ml ampicillin to select transformants. Incubate plates in a 30°C convection incubator until colonies appear. Expression and Purification of Thioredoxin Fusion Proteins
Strains GI698, GI723, and GI724 are all healthy prototrophs that can grow under a wide variety of growth conditions, including rich and minimal media and a broad range of growth temperatures (see Table 6.7.1). These strains can be prepared for transformation with pL-containing vectors by growing them in LB medium at 37°C. LB medium may also
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Current Protocols in Protein Science
be used for these strains during the short period of outgrowth immediately following transformation. This growth period of 30 min to 1 hr is often used to express drug-resistance phenotypes before plating out plasmid transformations onto solid medium. Subsequently, however, these strains should be grown only on minimal or tryptophan-free rich media, such as IMC medium containing 100 ìg/ml ampicillin (for expression of the fusion protein) or CAA/glycerol/ampicillin 100 medium (for plasmid DNA preparations). Except during transformation, LB medium should never be used with these three strains when they carry pL plasmids because LB contains tryptophan. The pL promoter is extremely strong and should be maintained in an uninduced state until needed so that expression of the protein will not lead to selection of mutant or variant cells with lower expression due to undesirable genetic selections or rearrangements in the expression strain.
3. Grow candidate colonies in 5 ml CAA/glycerol/ampicillin 100 medium overnight at 30°C. Prepare minipreps of plasmid DNA and check for correct gene insertion into pTRXFUS by restriction mapping. 4. Sequence plasmid DNA of candidate clones to verify the junction region between thioredoxin and the gene or sequence of interest. Induce expression 5. Streak out frozen stock culture of GI724 containing thioredoxin expression plasmid to single colonies on IMC plates containing 100 µg/ml ampicillin. Grow 20 hr at 30°C. Occasionally there is induction of pL plasmids grown in GI698 and GI724 at 37°C, even in medium containing no tryptophan. Such induction appears to be a temperature-dependent phenomenon. If growth at 37°C prior to pL induction is essential, then GI723 should be used as the host strain because GI723 produces higher levels of cI repressor than both GI698 and to GI724. Otherwise, plasmid-containing GI698 should be grown at 25°C and plasmid-containing GI724 should be grown at 30°C prior to induction (see Table 6.7.1).
6. Pick a single fresh, well-isolated, colony from the plate and use it to inoculate 5 ml IMC medium containing 100 mg/ml ampicillin in an 18 × 150–mm culture tube. Incubate overnight at 30°C on a roller drum. 7. Add 0.5 ml overnight culture to 50 ml fresh IMC medium containing 100 µg/ml ampicillin in a 250-ml culture flask (1:100 dilution). Grow at 30°C with vigorous aeration until absorbance at 550 nm reaches 0.4 to 0.6 OD/ml (∼3.5 hr). 8. Remove a 1-ml aliquot of the culture (uninduced cells). Measure the optical density at 550 nm and harvest the cells by microcentrifuging 1 min at maximum speed, room temperature. Carefully remove all the spent medium with a pipet and store the cell pellet at −80°C. 9. Induce pL by adding 0.5 ml of 10 mg/ml tryptophan (100 µg/ml final) to remaining cells immediately. 10. Incubate 4 hr at 37°C. At hourly intervals during this incubation, remove 1-ml aliquots of the culture and harvest cells as in step 8. 11. Harvest the remaining cells from the culture 4 hr post-induction by centrifuging 10 min at 3000 rpm (e.g., in a Beckman J6 rotor), 4°C. Store the cell pellet at −80°C. Procedures for further analysis of these cells are outlined in the support protocols.
Verify induction 12. Resuspend the pellets from the induction intervals (steps 8 and 10) in 200 µl of SDS-PAGE sample buffer/OD550 cells. Heat 5 min at 70°C to completely lyse the cells and denature the proteins. Run the equivalent of 0.15 OD550 cells per lane (30 µl) on an SDS-polyacrylamide gel (UNIT 10.1).
Purification of Recombinant Proteins
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13. Stain the gel 1 hr with Coomassie brilliant blue (UNIT 10.5). Destain the gel and check for expression. Most thioredoxin fusion proteins are produced at levels that vary from 5% to 20% of the total cell protein. The desired fusion protein should exhibit the following characteristics: it should run on the gel at the mobility expected for its molecular weight; it should be absent prior to induction; and it should gradually accumulate during induction, with maximum accumulation usually occurring 3 hr post-induction at 37°C. SUPPORT PROTOCOL 1
E. COLI LYSIS USING A FRENCH PRESSURE CELL A small, 3.5-ml French pressure cell can be used as a convenient way to lyse E. coli cells. The whole-cell lysate can be fractionated into soluble and insoluble fractions by microcentrifugation. Other lysis procedures may be used—for example, sonication (UNIT 6.6) or treatment with lysozyme-EDTA (UNIT 6.5). For use of the larger 40-ml French pressure cell, see UNIT 6.2. Materials Cell pellet from 4-hr post-induction culture (see Basic Protocol) 20 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E), 4°C Lysis buffer: 20 mM Tris⋅Cl (pH 8.0) with protease inhibitors (optional)—0.5 mM phenylmethylsulfonyl fluoride (PMSF), 1 mM p-aminobenzamidine (PABA), and 5 mM EDTA SDS-PAGE sample buffer (see recipe) French press and 3.5-ml mini-cell (SLM Instruments), 4°C Additional reagents and equipment for SDS-PAGE (UNIT 10.1) Lyse the cells 1. Resuspend cell pellet from 4-hr post-induction culture in 20 mM Tris⋅Cl, pH 8.0, to a concentration of 5 OD550/ml. Protease inhibitors can be included in the resuspension if desired. Cells can also be resuspended at densities of 100 OD550/ml or greater; however, at high densities cell lysis may be less efficient.
2. Place 1.5 ml resuspended cell pellet in the 3.5-ml French pressure cell. Hold the cell upside down with the base removed, the piston fully extended downwards, and the outlet valve handle that holds the nylon ball seal in the open position (loose). Before filling the pressure cell, check that the nylon ball, which seals the outlet port and sits on the end of the outlet valve handle, is not deformed. If it is, replace it with a new one. Both the condition of the nylon ball and its seat in the pressure-cell body are critical for the success of the procedure.
3. Bring the liquid in the pressure cell to the level of the outlet port by raising the piston slowly to expel excess air from the cell. With the outlet valve open and at the same time maintaining the piston in position, install the pressure-cell base. Gently close the outlet valve. CAUTION: Do not over-tighten the valve as this will deform the nylon ball and may irreparably damage its seat on the pressure-cell body.
4. Turn the sealed cell right-side-up and place it in the hydraulic press. Expression and Purification of Thioredoxin Fusion Proteins
5. Turn the pressure regulator on the press fully counter-clockwise to reset it to zero pressure. Set the ratio selector to medium. Turn on the press. CAUTION: The larger (50-ml) pressure cell is usually used with the selector set on high. The small (3.5-ml) cell is only used on medium ratio.
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6. Slowly turn the pressure regulator clockwise until the press just begins to move. Allow the press to compress the piston. The press will stop moving after a few seconds.
7. Position a collection tube under the pressure-cell outlet. Slowly increase the pressure in the cell by turning the pressure regulator clockwise. Monitor the reading on the gauge and increase the pressure to 1000 on the dial, corresponding to an internal cell pressure of 20,000 lb/in2. 8. While continuously monitoring the gauge, very slowly open the outlet valve until lysate begins to trickle from the outlet. The lysate should flow slowly and smoothly, and the cell pressure should not drop more than 100 divisions on the dial. At 20,000 lb/in2 and 5 OD550/ml, cell lysis will be complete after one passage through the press. Lower pressures and/or higher cell densities may require a second passage.
Fractionate the lysate 9. Remove a 100-µl aliquot of the lysate and freeze at −80°C (whole-cell lysate). 10. Fractionate the remainder of the lysate by microcentrifuging 10 min at maximum speed, 4°C. 11. Remove a 100-µl aliquot of the supernatant and freeze at −80°C (soluble fraction). Discard the remainder of the supernatant. Because this is a pilot experiment, it would not produce enough material to warrant saving any remaining supernatant.
12. Resuspend the pellet in an equivalent volume of lysis buffer. Remove a 100-µl aliquot and freeze at −80°C (insoluble fraction). 13. Lyophilize the 100-µl aliquots to dryness in a Speedvac evaporator. Solubilize in 100 µl SDS-PAGE sample buffer. Analyze 30-µl samples by SDS-PAGE (UNIT 10.1). This crude fractionation provides a fairly reliable indication of whether a protein has folded correctly. Usually proteins in the soluble fraction have adopted a correct conformation and proteins in the insoluble fraction have not. However, occasionally proteins found in the soluble fraction are not truly soluble; instead they form aggregates that do not pellet in the microcentrifuge. Conversely, sometimes a protein found in the insoluble fraction may be there because it has an affinity for cell-wall components and cell membranes, and it may not be intrinsically insoluble. Occasionally proteins can be recovered from these insoluble fractions by extracting with agents such as mild detergents.
OSMOTIC RELEASE OF THIOREDOXIN FUSION PROTEINS Thioredoxin and some thioredoxin fusion proteins can be released with good yield from the E. coli cytoplasm by a simple osmotic shock procedure.
SUPPORT PROTOCOL 2
Materials Cell pellet from 4-hr post-induction cultures (see Basic Protocol) 20 mM Tris⋅Cl (pH 8.0)/2.5 mM EDTA/20% (w/v) sucrose, ice-cold 20 mM Tris⋅Cl (pH 8.0)/2.5 mM EDTA, ice-cold Additional reagents and equipment for SDS-PAGE (UNIT 10.1) 1. Resuspend cell pellet from 4-hr post-induction cultures at a concentration of 5 OD550/ml in ice-cold 20 mM Tris⋅Cl (pH 8.0)/2.5 mM EDTA/20% sucrose. Incubate 10 min on ice. 2. Microcentrifuge 30 sec at maximum speed, 4°C, to pellet the cells.
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3. Discard the supernatant and gently resuspend the cells in an equivalent volume of ice-cold 20 mM Tris⋅Cl (pH 8.0)/2.5 mM EDTA. Incubate 10 min on ice and mix occasionally by inverting the tube. Osmotic release from the cytoplasm occurs at this stage.
4. Microcentrifuge 30 sec at maximum speed, 4°C. Save the supernatant (osmotic shockate). Resuspend the cell pellet in an equivalent volume 20 mM Tris⋅Cl (pH 8.0)/2.5 mM EDTA (retentate). 5. Lyophilize 100-µl aliquots of osmotic shockate and retentate to dryness in a Speedvac evaporator. 6. Solubilize each in 100 µl SDS-PAGE sample buffer. Analyze 30-µl aliquots by SDS-PAGE (UNIT 10.1). The osmotic-shock procedure provides a substantial purification step for some thioredoxin fusion proteins. This procedure will remove most of the contaminating cytoplasmic proteins as well as almost all of the nucleic acids. However the shockate will contain as contaminants about half of the cellular elongation factor-Tu (EF-Tu) and most of the E. coli periplasmic proteins. SUPPORT PROTOCOL 3
PURIFICATION OF THIOREDOXIN FUSION PROTEINS BY HEAT TREATMENT Wild-type thioredoxin is resistant to prolonged incubations at 80°C. A subset of thioredoxin fusion proteins also exhibit corresponding thermal stability, and heat treatment at 80°C can sometimes be used as an initial purification step. Under these conditions the majority of contaminating E. coli proteins are denatured and precipitated. Materials Cell pellet from 4-hr post-induction cultures (See Basic Protocol) 20 mM Tris⋅Cl (pH 8.0)/2.5 mM EDTA SDS-PAGE sample buffer (see recipe) 80°C water bath 10-ml glass-walled tube Additional reagents and equipment for lysis using a French pressure cell (Support Protocol 1) and SDS-PAGE (UNIT 10.1) 1. Resuspend cell pellet from 4-hr post-induction cultures at a concentration of 100 OD550/ml in 20 mM Tris⋅Cl (pH 8.0)/2.5 mM EDTA. It is important to start off with a high protein concentration in the lysate to ensure efficient precipitation of denatured proteins.
2. Lyse the cells at 20,000 lb/in2 in a French pressure cell (see Support Protocol 1, steps 2 to 8). Collect whole-cell lysate in a 10-ml glass-walled tube. 3. Incubate whole-cell lysate 10 min at 80°C. Remove 100-µl aliquots after 30 sec, 1 min, 2 min and 5 min and plunge immediately into ice. At 10 min, plunge the remaining heated lysate into ice.
Expression and Purification of Thioredoxin Fusion Proteins
A glass-walled tube (not plastic) provides good thermal conductivity to provide a rapid rise in temperature to 80°C and then a rapid drop in temperature to 4°C. A suitable volume to use in a 10-ml glass tube is 1.5 ml lysate. For large-scale work, a glass-walled vessel should be used and the lysate should be mixed well during both heat treatment and cooling.
4. Microcentrifuge the aliquots 10 min at maximum speed, 4°C to pellet heat-denatured, precipitated proteins.
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5. Remove 2-µl aliquots of the supernatants and add 28 µl SDS-PAGE sample buffer. Analyze the samples by SDS-PAGE (UNIT 10.1) to determine the heat stability of the fusion protein and the minimum time of heat treatment required to obtain a good purification. REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Casamino Acids (CAA), 2% (w/v) 20 g Casamino Acids (Difco-certified) H2O to 1 liter Autoclave or filter sterilize through a 0.45-µm filter Store ≤2 months at room temperature Do not use technical-grade Casamino Acids because it has a higher NaCl content.
CAA/glycerol/ampicillin 100 medium 800 ml 2% (w/v) Casamino Acids (see recipe; 1.6% final) 100 ml 10× M9 salts (see recipe; 1× final) 100 ml 10% (v/v) glycerol (sterile; 1% final) 1 ml 1 M MgSO4 (sterile; 1 mM final) 0.1 ml 1 M CaCl2 (sterile; 0.1 mM final) 1 ml 2% (w/v) vitamin B1 (sterile; 0.002% final) 10 ml 10 mg/ml ampicillin (sterile; 100 µg/ml final) Prepare fresh IMC medium 200 ml 2% (w/v) Casamino Acids (see recipe; 0.4% final) 100 ml 10× M9 salts (see recipe; 1× final) 40 ml 20% (w/v) glucose (sterile; 0.5% final) 1 ml 1 M MgSO4 (sterile; 1 mM final) 0.1 ml 1 M CaCl2 (sterile; 0.1 mM final) 1 ml 2% (w/v) vitamin B1 (sterile; 0.002% final) 658 ml glass-distilled H2O (sterile) 10 ml 10 mg/ml ampicillin (sterile; optional; 100 µg/ml final) Use fresh IMC plates 15 g agar [Difco; 1.5% (w/v)] 4 g Casamino Acids [Difco-certified; 0.4% (w/v)] 858 ml glass-distilled H2O (sterile) Autoclave 30 min Cool in a 50°C water bath 100 ml 10× M9 salts (see recipe; 1× final) 40 ml 20% (w/v) glucose (sterile; 0.5% final) 1 ml 1 M MgSO4 (sterile; 1 mM final) 0.1 ml 1 M CaCl2 (sterile; 0.1 mM final) 1 ml 2% (w/v) vitamin B1 (sterile; 0.002% final) 10 ml 10 mg/ml ampicillin (sterile; optional; 100 µg/ml final) Mix well and pour into petri plates Store ≤1 month at 4°C Purification of Recombinant Proteins
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M9 salts, 10× 60 g Na2HPO4 (0.42 M) 30 g KH2PO4 (0.24 M) 5 g NaCl (0.09 M) 10 g NH4Cl (0.19 M) H2O to 1 liter Adjust pH to 7.4 with NaOH Autoclave or filter sterilize through a 0.45-µm filter Store ≤6 months at room temperature SDS-PAGE sample buffer 15% (v/v) glycerol 0.125 M Tris⋅Cl, pH 6.8 (APPENDIX 2E) 5 mM Na2EDTA 2% (w/v) SDS 0.1% (w/v) bromphenol blue 1% (v/v) 2-mercaptoethanol (2-ME; add immediately before use) Store indefinitely at room temperature Tryptophan, 10 mg/ml Heat 500 ml glass-distilled H2O to 80°C. Stir in 5 g L-tryptophan until dissolved. Filter sterilize the solution through a 0.45 µm filter and store ≤6 months in the dark at 4°C. COMMENTARY Background Information
Expression and Purification of Thioredoxin Fusion Proteins
Two significant problems plague researchers who hope to express heterologous proteins in Escherichia coli: inefficient initiation of translation of many eukaryotic mRNA sequences on bacterial ribosomes (Stormo et al., 1982), and proteins that often form insoluble aggregates, called inclusion bodies, that are composed of misfolded or denatured proteins (Mitraki and King, 1989). Although successful protocols for refolding eukaryotic proteins from inclusion bodies can be developed, the process is always uncertain and usually time-consuming; in most instances it is preferable to prevent inclusion-body formation in the first place. The use of trxA fusions provides a solution to both problems. Inefficient initiation of translation of eukaryotic messages in E. coli can often be improved by modifying sequences at the 5′ end of the gene. A more reliable technique that avoids the problem entirely is to use a gene-fusion strategy in which the gene of interest is linked in-frame to the 3′ end of a highly translated partner gene. In this case protein synthesis always initiates on the same efficiently translated fusion-partner mRNA, thus ensuring high-level expression. Some earlier gene-fusion expression systems, for example the trpE and lacZ systems, offer very reliable ways of
producing large quantities of any desired eukaryotic protein. However, these gene-fusion systems still suffer from the pervasive inclusion-body problem. They are thus mainly useful for the production of antigens, rather than correctly folded, biologically active proteins. More recently the maltose binding protein (MBP; Riggs, 1994; UNIT 5.1) and glutathioneS-transferase (GST) gene fusion expression systems (see UNIT 6.6) have proven more successful in producing soluble fusion proteins; these systems retain the translation advantage of the earlier fusion systems. Apart from the obvious advantages in making a correctly folded product, the synthesis of soluble fusion proteins also allows for the development of generic purification schemes based on some unique property of the fusion partner. Why would any particular eukaryotic protein produced in the E. coli cytoplasm be more soluble when it is linked to a fusion partner than it would be by itself? It is likely that physical properties of the fusion-partner protein are important, with efficient self-folding and high solubility being useful in this role. It is possible that some good fusion partners (proteins that fold efficiently and are highly soluble), by virtue of their desirable physical qualities, are able to keep folding intermediates of linked heterologous proteins in solution long enough for
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them to adopt their correct final conformations. In this respect the fusion partner may serve as a covalently joined chaperone protein, in many ways fulfilling the role of authentic chaperone proteins (McCoy, 1992), analogous to the covalent chaperone role proposed for the N-terminal pro regions of a number of protein precursors (Silen et al., 1989; Shinde et al., 1993). Many of the known properties of E. coli thioredoxin (Holmgren, 1985) suggested that it would make a particularly effective fusion partner in an expression system. First, thioredoxin, when overproduced from plasmid vectors, can accumulate to 40% of the total cellular protein, yet even at these expression levels all of the protein remains soluble. Second, the molecule is small (11,675 Mr) and would contribute a relatively modest amount to the total mass of any fusion protein, in contrast to other systems such as the lacZ system. Third, the tertiary structure of thioredoxin (Katti et al., 1990) reveals that both the N- and C-termini of the molecule are accessible on the surface and in good position to link to other proteins. The structure also shows that the molecule has a very tight fold, with >90% of its primary sequence involved in strong elements of secondary structure. This provides an explanation for thioredoxin’s observed high thermal stability (Tm = 85°C), and suggests that the molecule might possess the robust folding characteristics that could make it a good fusion-partner protein. In support of this view, complete thioredoxin domains are found in a number of naturally occurring multidomain proteins, including E. coli DsbA (Bardwell et al., 1991), the mammalian endoplasmic reticulum proteins ERp72 (Mazzarella et al., 1990), and protein disulfide isomerase (PDI; Edman et al., 1985). These proteins can all be considered as natural precedents for thioredoxin fusion proteins. The synthesis of small peptides in E. coli is often difficult, with the products frequently being extensively degraded or insoluble. The thioredoxin tertiary structure revealed that the characteristic active site, —CGPC—, protrudes from the body of the protein as a surface loop, with few interactions with the rest of the molecule. The loop does not seem to contribute to the overall stability of thioredoxin, so the production of peptides as insertions at this site was an attractive possibility. In this location they would be protected from host-cell aminoand carboxypeptidases, and thioredoxin’s high solubility should help keep them in solution. In addition, the conformation of peptides inserted at this position would be constrained, which
could be an advantage for applications in which it is desirable for the peptide to adopt a particular form. Thioredoxin has indeed proven to be an excellent partner for the production of soluble fusion proteins in the E. coli cytoplasm (LaVallie et al., 1993b). Figure 6.7.2 demonstrates the production of soluble fusion proteins between thioredoxin and eleven human and murine cytokines and growth factors using the trxA vectors. All of these mammalian proteins had been previously produced in E. coli only as insoluble inclusion bodies. As thioredoxin fusions, the growth factors are not only made in a soluble form, but in most cases they are also biologically active in in vitro assays. Experience gained while working with these and a number of other trxA fusion proteins shows that two further characteristics of thioredoxin can be exploited as purification tools. The first is the inherent thermal stability of the molecule, a property that is retained by some thioredoxin fusion proteins. This enables heat treatment to be used as an effective purification step. The second additional property relates to thioredoxin’s cellular location. Although E. coli thioredoxin is a cytoplasmic protein, it has been shown to occupy a special position within the cell—it is primarily located on the cytoplasmic face of the adhesion zones that exist between the inner and outer membranes of the E. coli cell envelope (Lunn and Pigiet, 1982). From this location thioredoxin is quantitatively released to the exterior of the cell by simple osmotic shock or freeze/thaw treatments, a remarkable property that is retained by some thioredoxin fusion proteins, thus providing a simple purification step. A more generic method for purification of any soluble thioredoxin fusion employs a modified form of thioredoxin (called “Hispatch Trx”), which has been designed to bind to metal chelate resins (Lu et al., 1996; UNIT 9.4). If the fusion protein is soluble after lysis or osmotic release (see Support Protocols 1 and 2), then any of the conventional purification methods described in Chapter 8 can be used. Alternatively, an affinity-based purification method can be used where an immobilized arsenical compound forms an adduct with the redox-sensitive vicinal dithiols present at the active site of thioredoxin. This affinity matrix was originally prepared by covalently linking 4-aminophenylarsenine oxide to cyanogen bromide–activated Sepharose 4B (Hannestad et al., 1982). Hoffman and Lane (1992) coupled the same ligand to Affi-Gel 10 (Bio-Rad); the
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1
2
3
4
5
6
7
8
9
10
11
12
MW(kDa) 97.4 66.2
45.0
31.0
21.5
14.4
Figure 6.7.2 Expression of thioredoxin gene fusions. The gel shows proteins found in the soluble fractions derived from E. coli cells expressing eleven different thioredoxin gene fusions. Lane 1, host E. coli strain GI724 (negative control, 37°C); lane 2, murine interleukin-2 (IL-2; 15°C); lane 3, human IL-3 (15°C); lane 4, murine IL-4 (15°C); lane 5, murine IL-5 (15°C); lane 6, human IL-6 (25°C); lane 7, human MIP-1a (37°C); lane 8, human IL-11 (37°C); lane 9, human macrophage colony-stimulating factor (M-CSF; 37°C); lane 10, murine leukemia inhibitory factor (LIF; 25°C); lane 11, murine steel factor (SF; 37°C); and lane 12, human bone morphogenetic protein-2 (BMP-2; 25°C). Temperatures in parentheses are the production temperature chosen for expressing each fusion. This is a 10% SDS-polyacrylamide gel, stained with Coomassie brilliant blue.
Expression and Purification of Thioredoxin Fusion Proteins
resultant affinity matrix has a 10-atom spacer which makes it more useful for affinity chromatography. The resin is commercially available from Invitrogen (ThioBond) and Sigma. Protocols detailing the affinity-based purification approach can be obtained on-line from the I nvitr og en web site (http://www.invitrogen.com/manuals.html), and a recent example from the literature is the expression of human glutamate decarboxylase (Papouchado et al., 1997). Other specific purification approaches include immunoaffinity chromatography using antibodies against either the target protein or against the thioredoxin moiety (monoclonal antibodies against thioredoxin are commercially available). Thioredoxin fusions with additional histidine tags or thioredoxin fusions based on mutant thioredoxins with metalchelating affinity (Lu et al., 1996) can be purified using metal-chelate chromatography (see UNIT 9.4). Finally, if the fusion protein is insoluble, then attempts to solubilize and fold the fusion protein can be made prior to purification,
in a technique analogous to that used with insoluble GST fusion proteins (see UNIT 6.6). Thioredoxin from E. coli is a very soluble protein that can be readily denatured and refolded (Kelley et al., 1987).
Critical Parameters Lack of protein solubility leading to inclusion-body formation in E. coli is a complex phenomenon with many contributing factors: simple insolubility as a result of high-level expression, insolubility of protein-folding intermediates, lack of appropriate bacterial chaperone proteins, and lack of glycosylation mechanisms in the bacterial cytoplasm. Fusion of heterologous proteins to thioredoxin or to other fusion partners can help address most of these solubility issues. However, another important factor contributing to inclusion body formation is the inability to form essential disulfide bonds in the reducing environment of the bacterial cytoplasm, which leads to incorrect folding. Thermal lability of even correctly
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folded heterologous proteins in the absence of these stabilizing disulfide cross-links is a significant problem, so the expression of fusion genes should be attempted over a wide range of temperatures, even as low as 15°C (the limit for E. coli growth is ∼8°C). Thermal denaturation is a time-dependent process, so it is also prudent to monitor the solubility of the expressed fusion protein over the time course of induction. A great many proteins contain distinct structural domains. For example, hormone receptor proteins usually have an extracellular ligandbinding domain, a transmembrane region, and an intracellular effector domain. Sometimes, expressing these domains individually as fusion proteins can yield better results than expressing the entire protein. The exact positions chosen for boundaries of the domains to be expressed in the fusion protein are important and can be determined from a knowledge of the tertiary structure of the protein of interest, by homology comparisons with similar proteins, by limited proteolysis or other domain-mapping experiments, or empirically by generating multiple fusions that test different boundary positions. It is important to be consistent in treating samples for loading on gels. For example, using different heating conditions from one experiment to the next can result in a mobility shift for the protein of interest.
Anticipated Results Thioredoxin fusion protein yields are usually in the range of 5% to 20% of total cell protein. At these expression levels, a 1-liter induction culture in a shaker flask will yield ∼3 g (wet weight) of cells, 300 mg total protein, and 15 to 60 mg of thioredoxin fusion protein. The final recovered yield will depend on factors such as solubility of the fusion protein and the efficiency of downstream purification procedures.
Time Considerations From a single colony on a plate, the basic induction protocol requires an overnight growth to prepare a liquid inoculum and a 3.5-hr preinduction growth at 30°C the next day, followed by a 4-hr 37°C induction period. These times are significantly longer if lower induction temperatures are required (see Table 6.7.1). Lysis of a sample in the French pressure cell should require ≤ 5 min, and both the heattreatment and osmotic-shock procedures require <1 hr each. SDS-PAGE takes 2.5 hr.
Literature Cited Bardwell, J.C.A., McGovern, K., and Beckwith, J. 1991. Identification of a protein required for disulfide bond formation in vivo. Cell 67:581589. Edman, J.C., Ellis, L., Blacher, R.W., Roth, R.A., and Rutter, W.J. 1985. Sequence of protein disulphide isomerase and implications of its relationship to thioredoxin. Nature 317:267-270. Hannestad, U., Lundqvist, P., and Sorbo, B. 1982. An agarose derivative containing an arsenical for affinity chromatography of thiol compounds. Anal. Biochem. 126:200-204. Hoffman, R.D. and Lane, M.D. 1992. Isodophenylarsine oxide and arsenical affinity chromatography: New probes for dithiol proteins. J. Biol. Chem. 267:14005-14011. Holmgren, A. 1985. Thioredoxin. Ann. Rev. Biochem. 54:237-271. Katti, S.K., LeMaster, D.M., and Eklund, H. 1990. Crystal structure of thioredoxin from Escherichia coli at 1.68 angstroms resolution. J. Mol. Biol. 212:167-184. Kelley, R.F., Shalongo, M., Jagannadham, M.V., and Stellwagen, E. 1987. Equilibrium and kinetic measurements of the conformational transition of reduced thioredoxin. Biochemistry 26:14061411. LaVallie, E.R., Rehemtulla, A., Racie, L.A., DiBlasio, E.A., Ferenz, C., Grant, K.L., Light, A., and McCoy, J.M. 1993a. Cloning and functional expression of a cDNA encoding the catalytic subunit of bovine enterokinase. J. Biol. Chem. 268:23311-23317. LaVallie, E.R., DiBlasio, E.A., Kovacic, S., Grant, K.L., Schendel, P.F., and McCoy, J.M. 1993b. A thioredoxin gene fusion expression system that circumvents inclusion body formation in the E. coli cytoplasm. Bio/Technology 11:187-193. Lu, Z., DiBlasio-Smith, E.A., Grant, K.L., Warne, N.W., LaVallie, E.R., Collins-Racie, L.A., Follettie, M.T., Williamson, M.J., and McCoy, J.M. 1996. Histidine patch thioredoxins. Mutant forms of thioredoxin with metal chelating affinity that provide for convenient purifications of thioredoxin fusion proteins. J. Biol. Chem. 271:5059-5065. Lunn, C.A. and Pigiet, V.P. 1982. Localization of thioredoxin from Escherichia coli in an osmotically sensitive compartment. J. Biol. Chem. 257:11424-11430. Mazzarella, R.A., Srinivasan, M., Haugejorden, S.M., and Green, M. 1990. ERp72, an abundant luminal endoplasmic reticulum protein, contains three copies of the active site sequences of protein disulfide isomerase. J. Biol. Chem. 265:1094-1101. McCoy, J.M. 1992. Heat-shock proteins and their potential uses for pharmaceutical protein production in microorganisms. In Stability of Protein Pharmaceuticals, Part B. (T. Ahern and M. Manning, eds.) pp 287-316. Plenum Press, New York.
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Mieschendahl, M., Petri, T., and Hanggi, U. 1986. A novel prophage independent trp regulated lambda pL expression system. Bio/Technology 4:802-808.
Shinde, U., Chatterjee, S., and Inouye, M. 1993. Folding pathway mediated by an intramolecular chaperone. Proc. Natl. Acad. Sci. U.S.A. 90:6924-6928.
Mitraki, A. and King, J. 1989. Protein folding intermediates and inclusion body formation. Bio/Technology 7:690-697.
Silen, J.L., Frank, D., Fujishige, A., Bone, R., and Agard, D.A. 1989. Analysis of prepro-α-lytic protease expression in Escherichia coli reveals that the pro region is required for activity. J. Bacteriol. 171:1320-1325.
Norrander, J., Kempe, T., and Messing, J. 1983. Construction of improved M13 vectors using oligonucleotide-directed mutagenesis. Gene 26:101-106.
Stormo, G.D., Schneider, T.D., and Gold, L. 1982. Characterization of translation initiation sites in E. coli. Nucl. Acids Res. 10:2971-2996.
Papouchado, M.L., Valdez, S.N., Ghiringhelli, D., Poskus, E., and Ermacora, M.R. 1997. Expression of properly folded human glutamate decarboxylase 65 as a fusion protein in Escherichia coli. Eur. J. Biochem. 246:350-359.
Key Reference
Riggs, P. 1994. Expression and purification of maltose-binding protein fusions. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.F. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp.16.6.116.6.14. John Wiley & Sons, New York.
A source of protocols for cloning and analyzing DNA.
Shatzman, A.R., Gross, M.S., and Rosenberg, M. 1990. Expression using vectors with phage λ regulatory sequences. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.F. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 16.3.1-16.3.11. John Wiley & Sons, New York. Shimatake, H. and Rosenberg, M. 1981. Purified λ regulatory protein cII positively activates promoters for lysogenic development. Nature 292:128-132.
Ausubel, F.M., Brent, R., Kingston, R.F., Moore, D.D., Seidman, J.G., Smith, J.A., and Struhl, K. (eds.) 1997. Current Protocols in Molecular Biology. John Wiley & Sons, New York.
Internet Resources http://www.invitrogen.com/manuals.html Source for protocols on affinity-based purification.
Contributed by John McCoy and Edward LaVallie Genetics Institute Cambridge, Massachusetts
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Automated Large-Scale Purification of a Recombinant G-Protein-Coupled Neurotensin Receptor
UNIT 6.8
Structure determination of integral membrane proteins requires milligram amounts of purified, functional protein on a regular basis. As an example, this unit describes a method for purifying a G-protein-coupled neurotensin receptor fusion protein at the 10-milligram level, using immobilized metal affinity chromatography and a neurotensin affinity column in a fully automated mode. The method is designed to be simple and robust. Integral membrane proteins reside in lipid membranes. The hydrophobic character of an integral membrane protein makes it necessary to use detergents to extract receptors from the membrane environment and to keep them in solution. Then the same purification procedures as those used for water-soluble proteins can be applied to detergent-solubilized membrane proteins. The purification of integral membrane proteins can be conceptually divided into two steps: extraction from membranes with detergents (solubilization), followed by purification procedures. Basic Protocol 1 describes the solubilization step, and Basic Protocol 2 describes an affinity purification procedure. Many integral membrane proteins are stable in a membrane environment but show reduced stability in detergent solution; therefore, the purification procedures must be performed rapidly to minimize protein denaturation. A timeline guide is provided to suggest the most time-efficient approach for the affinity purification procedure. Basic Protocol 2 describes the purification of a histidine-tagged neurotensin receptor generated as a recombinant protein as described in Support Protocol 1. The purification is based on two steps: the first, generic step, exploits binding of the receptor to a metal affinity column through its C-terminal histidine affinity tag. The first purification step selects for tagged receptors irrespective of their conformations. However, not all detergent-solubilized receptors can be assumed to be fully functional. The second purification step is receptor-specific and is based on binding of a high-affinity ligand to the receptor. It separates functional from nonfunctional receptors. The generation of a neurotensin affinity column is described in Support Protocol 2. The progress of receptor purification can be monitored by radioligand binding experiments (Support Protocol 3) and protein concentration determination (Support Protocol 4). Protein concentration establishes the total amount of protein, i.e., purified receptor (whether functional or not) and contaminants. Radioligand binding detects correctly folded receptors, which bind ligand. The availability of both parameters allows estimation of the quality of the final receptor preparation. The two-step purification procedure described in this unit allows for production of large (10 mg) quantities of pure, fully functional neurotensin receptor fusion protein.
SOLUBILIZATION OF MEMBRANE-BOUND RECEPTORS FROM TOTAL CELL LYSATE The solubilization and purification of receptors, including all preparations, requires four days to complete. Day 1 is used to make buffers and to start the equilibration of the columns. The solubilization of receptors (described in this protocol) and initiation of purification (Basic Protocol 2) take place on day 2. This solubilization protocol should
Contributed by Jim F. White and Reinhard Grisshammer Current Protocols in Protein Science (2007) 6.8.1-6.8.31 C 2007 by John Wiley & Sons, Inc. Copyright
BASIC PROTOCOL 1
Purification of Recombinant Proteins
6.8.1 Supplement 47
be started on day 2 after the Ni-NTA and NT columns have been equilibrated as described in Basic Protocol 2. The purifier runs through day 3, and purified receptors are available on day 4. Solubilization involves the suspension of cells in buffer, addition of detergent solutions to extract the receptors from the membranes, mild sonication to enhance solubilization efficiency, and ultracentrifugation to remove nonsolubilized membranes and cell debris. The supernatant is then used for purification of receptors (Basic Protocol 2).
Materials 250 g frozen E. coli cells expressing the NTS1 fusion protein NTS1-624 (Support Protocol 1) 2× solubilization buffer (100 mM Tris·Cl, pH 7.4/60% v/v glycerol/400 mM NaCl) 1 M MgCl2 10 mg/ml DNase I (e.g., Sigma) Deionized water, 4◦ C CHAPS/CHS stock solution: 6% (w/v) 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS; Anatrace)/1.2% (w/v) cholesteryl hemisuccinate Tris salt (CHS; Anatrace) detergent stock solution; dissolve CHS in CHAPS solution by stirring for several hours; prepare fresh and cool to 4◦ C LM stock solution: 10% (w/v) n-dodecyl-β-D-maltoside (LM; Anatrace) detergent stock solution in H2 O; prepare fresh and cool to 4◦ C Protease inhibitor stock solutions 70 mg/ml phenylmethylsulfonyl fluoride (PMSF) in ethanol 1 mg/ml leupeptin (MP Biomedicals) in H2 O 1.4 mg/ml pepstatin A (MP Biomedicals) in methanol 2 M imidazole (e.g., Fluka Ultra; Sigma-Aldrich) stock solution: adjust to pH 7.4 with 12 N HCl Waring blender 2-liter beaker Magnetic stir bar and stir plate Misonix sonicator 3000 with 1/2-inch flat tip 70-ml polycarbonate ultracentrifuge bottles 45Ti ultracentrifugation rotor (Beckman-Coulter; or equivalent) and ultracentrifuge 0.22-µm filter (Stericup; Millipore, or equivalent) Suspend frozen cells 1. Wrap 250 g of frozen cell paste between plastic sheets (e.g., Saran Wrap) and gently crush with a hammer to obtain smaller pieces. Place cells into a Waring blender (which is at room temperature). 2. Add 500 ml of cold 2× solubilization buffer to the cells. 3. Operate the Waring blender for 30 sec on low speed, followed by ∼3 min on high speed, until all cells are suspended. Incomplete cell suspension will reduce the solubilization efficiency, but prolonged operation of the Waring blender will lead to warming of the material, which must be avoided.
4. Transfer the cell suspension into a 2-liter beaker with a magnetic stir bar, and place on a stir plate in a cold room.
Automated Large-Scale Production of a Recombinant GPCR
5. While stirring, add 1 ml of each of the protease inhibitor stocks, 5 ml of 1 M MgCl2 solution (final concentration 5 mM), 0.6 ml DNaseI solution, and 50 ml cold water. The volume of the suspension should now be close to 800 ml.
6.8.2 Supplement 47
Current Protocols in Protein Science
Solubilize membrane-bound receptors 6. While stirring, add dropwise 100 ml CHAPS/CHS stock solution. Slow addition of the detergent is recommended. Fast addition leads to high local detergent concentrations, and possible denaturation of receptors.
7. While stirring, add dropwise 100 ml LM stock solution. 8. Continue to stir in the cold room for 15 min. 9. Sonicate 33 min (8 sec/g of cells), 1 sec on, 2 sec off, level 4 (for the Misonix sonicator 3000 with a 1/2-in. flat tip). Keep the sample in an ice/water bath, and stir continuously during sonication to avoid local heating. 10. Add an additional 1 ml of each protease inhibitor stock. 11. Determine the volume of the preparation. 12. Add cold water dropwise with stirring to a final volume of 1000 ml. The amount of water needed at this step is ∼20 ml, which takes ∼2 min to add.
13. Stir sample in the cold room for an additional 30 min.
Clarify solubilized receptor preparation 14. Transfer sample to 70-ml polycarbonate ultracentrifuge bottles and ultracentrifuge 1 hr at 235,000 × g (45,000 rpm at rmax in a Beckman 45Ti rotor), 4◦ C. A volume of ∼1 liter requires three Beckman 45Ti rotors (or equivalent), using polycarbonate bottles with a capacity of 70 ml.
15. Decant the supernatant (containing solubilized proteins) and determine volume. 16. Add the cold 2 M imidazole stock solution dropwise while stirring to attain a final concentration of 50 mM. The presence of imidazole eliminates the binding of most E. coli contaminants to the Ni-NTA resin.
17. Pass sample through 0.22-µm filter. Keep the sample cold. The filtered sample is now ready for loading onto the Ni-NTA column (Basic Protocol 2). Storage of the sample at this point is not recommended.
AUTOMATED PURIFICATION OF THE NTS1 FUSION PROTEIN USING Ni-NTA CHROMATOGRAPHY AND NEUROTENSIN (NT) AFFINITY COLUMNS The step-by-step protocol for the automated purification of the NTS1 fusion protein is given in Table 6.8.1. The routines ensure that each buffer line, pump head, and column are filled with the correct buffer at any given time. Solubilized proteins are passed through line S1 by the external sample pump P950 onto the Ni-NTA column. An air sensor (air2) detects the end of loading. After extensive washing of the Ni-NTA column using system pump A, bound receptor fusion protein is eluted in the presence of 200 mM imidazole (system pump B), and collected (line F3) into a vessel which contains 1 ml of buffer NT200, into which line B22 is submersed. The Ni-NTA column eluate is diluted with NT0 buffer (see Background Information) using the system pumps, and passed onto the NT column (see Support Protocol 2 for preparation of the NT column). Again, an air sensor (air1 in line B22) detects the end of loading. After washing of the NT column, the NTS1 fusion protein is eluted and collected. The columns are then washed into storage buffer, and the system pumps are cleaned with water and 20% v/v ethanol. Starting with solubilization, a 10-milligram preparation is completed after 55 hr.
BASIC PROTOCOL 2
Purification of Recombinant Proteins
6.8.3 Current Protocols in Protein Science
Supplement 47
Table 6.8.1 Purification Scheme Using a 100-ml Ni-NTA Superflow Column and a 20-ml NT Column
Flow rate system pumps (ml/min)
Buffer
Buffer line
Sample
S1 air sensor 2.0 (sample pump)
NiA
S8
NiA
Volume Injection (ml) valve
Column position
Outlet valve
Gradient Comment (% B)
∼ 960
Inject
4
F4 NiFT
NA
Load Ni-NTA column
2.0 (sample pump)
10
Inject
4
F4 NiFT
NA
Rinse sample pump
A2
2.0
140
Load
4
F4 NiFT
0
Wash Ni-NTA column
NiA
A2
2.0
1360
Load
4
F1 waste
0
Wash Ni-NTA column
NiB
B1
5.0
20
Load
Bypass
F1 waste
100
Rinse line
NiB
B1
2.0
72
Load
4
F1 waste
100
Elute Ni-NTA column
NiB
B1
2.0
122
Load
4
F3 peak collection
100
Elute Ni-NTA column
NiB
B1
2.0
506
Load
4
F1 waste
100
Wash Ni-NTA column
NT200
B28
5.0
25
Load
Bypass
F1 waste
100
Rinse line
NT0
A12
5.0
25
Load
Bypass
F1 waste
0
Rinse line
NT0/NT200
A12/B28
5.0
50
Load
Bypass
F1 waste
35
Rinse lines
NT0/NiNTA eluate
A12/B22 air sensor
0.4
351
Load
5
F5 NTFT
35
Air sensor load Ni-NTA eluate onto NT column
NT0/NT200
A12/B28
0.4
50
Load
5
F5 NTFT
35
Rinse lines
NT0/NT200
A12/B28
0.7
160
Load
5
F1 waste
35
Wash NT column
NT1K
A13
5.0
50
Load
Bypass
F1 waste
0
Rinse line
NT1K
A13
0.5
75
Load
5
F2 1-ml fractions
0
Elute NT column
NT1K
A13
0.7
75
Load
5
F1 waste
0
Elute NT column
H2 O
A11
5.0
20
Load
Bypass
F1 waste
0
Rinse line
H2 O
A11
3.0
500
Load
4
F1 waste
0
Wash Ni-NTA column
B3
A16
5.0
20
Load
Bypass
F1 waste
0
Rinse line
A3
A17
5.0
20
Load
Bypass
F1 waste
0
Rinse line
A3
A17
0.7
150
Load
5
F1 waste
0
Wash NT column
B3
A16
0.7
150
Load
5
F1 waste
0
Wash NT column
H2 O/ H2 O
A11/B21
5.0
100
Load
Bypass
F1 waste
50
Wash system pumps with H2 O
20% (v/v) A18/B27 ethanol/20% (v/v) ethanol
2.5
300
Load
Bypass
F1 waste
50
Wash system pumps with 20% (v/v) ethanol
a Abbreviations: NA, not applicable; NiFT, flowthrough of Ni-NTA column; NTFT, flowthrough of NT column.
6.8.4 Supplement 47
Current Protocols in Protein Science
Figure 6.8.1 Schematic representation of the chromatography system used for automated purification of an NTS1 fusion protein. Circles denote valves (1, injection valve; 2, column valves; 3, outlet valve; 4, sample valve; 5, buffer valve A1; 6, buffer valve B2). The system pumps are indicated as A and B. Buffer lines F3 and B22 are used to collect and to load the Ni-NTA column eluate (NiE) onto the NT column. The NT column flowthrough is collected by line F5. The NT column eluate (NTE) is collected by the fraction collector (frac). Labels in other shaded boxes indicate names of buffers. For clarity, not all buffers and buffer lines are indicated. S1, S2, and S8 represent sample inlet lines. The NiE collected in vessel F3 is diluted with NT0 buffer to reduce the NaCl concentration from 200 mM to 70 mM before passing onto the NT column (syntax setting “35% B” in the UNICORN 4.11 program for purification). Adapted and reprinted by permission of the Federation of the European Biochemical Societies from White et al., 2004.
¨ This protocol describes the use of an Akta Purifier P100 system (Amersham Biosciences, GE Healthcare; see Fig. 6.8.1). However, any other chromatography system capable of ¨ executing the operations described below is adequate. The Akta Purifier is equipped with a sample valve (buffer line S1 with air sensor Air-912, S3 and S8), sample pump P950, injection valve (with the sample pump outlet connected to port 2, and port 3 connected to port 6), column valves, outlet valve, fraction collector Frac950, buffer valve A1 (with buffer lines A11, A12, A13, A16, A17, A18) connected to buffer line A1, buffer line A2, buffer line B1, and buffer valve B2 (with buffer lines B21, B22 with air sensor Air-912, B27, B28) connected to buffer line B2. Teflon tubing (i.d. 1.6 mm) is used for the buffer inlet lines, whereas Tefzel tubing (i.d. 0.75 mm) is used for the air sensor inlet lines. The 100-ml Ni-NTA Superflow (Qiagen) column (XK50, Amersham Biosciences) is connected to port 4 of the column valves, and the 20-ml NT column is connected to port 5 of the column valves (White et al., 2004).
Materials Milli-Q-purified water (or equivalent) NiA buffer: 50 mM Tris·Cl, pH 7.4/30% (v/v) glycerol/50 mM imidazole/200 mM NaCl/0.5% (w/v) 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS; Anatrace)/0.1% (w/v) cholesteryl hemisuccinate Tris salt (CHS; Anatrace)/0.1% (w/v) n-dodecyl-β-D-maltoside (LM; Anatrace)
Purification of Recombinant Proteins
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NiB buffer: NiA buffer with 200 mM imidazole NT0 buffer: 50 mM Tris·Cl, pH 7.4/30% v/v glycerol/1 mM EDTA/0.5% (w/v) CHAPS/0.1% (w/v) CHS/0.1% (w/v) LM NT200 buffer: NT0 buffer with 200 mM NaCl NT1K buffer: NT0 buffer with 1 M NaCl A3 buffer: 50 mM Tris·Cl, pH 7.4/1 mM EDTA/1 M NaCl B3 buffer: 50 mM Tris·Cl, pH 7.4/1 mM EDTA/3 mM NaN3 XK50 column (GE Healthcare) packed with 100 ml of Ni-NTA resin (Qiagen) NT column (Support Protocol 2) 95% (v/v) ethanol Clarified preparation containing solubilized proteins (Basic Protocol 1) 0.22-µm filter (Stericup; Millipore, or equivalent) ¨ Akta Purifier P100 chromatography system and operating software, e.g., UNICORN 4.11 (GE Healthcare) 50-ml Falcon test tube 75 borosilicate glass fraction collector tubes (12 × 75 mm) 500- and 1000-ml beakers 250-ml Pyrex bottle NOTE: All steps are performed at 4◦ C or on ice unless otherwise indicated. The chromatography equipment should be located in a cold (4◦ C) room.
Day 1: Prepare buffers and purifier 1. Using Milli-Q-purified water (or equivalent), prepare all buffers in the materials list in the volumes indicated in Table 6.8.2. Pass through a 0.22-µm filter. Table 6.8.2 Buffer Requirements for Receptor Solubilization, Column Equilibration, and Subse¨ quent Purification Using the Akta Purifier with a 100-ml Ni-NTA column and a 20-ml NT column
Stock solutions
Amount needed (ml)
10% (w/v) LM
155
–
6% (w/v) CHAPS/1.2% (w/v) CHSa
525
–
2× solubilization buffer
500
–
Buffer
Automated Large-Scale Production of a Recombinant GPCR
6.8.6 Supplement 47
Amount needed (ml)
Buffer line
H2 O
1000
A11–B21b
NT0
800
A12
NT1K
320
A13
B3
300
A16
A3
300
A17
20% (v/v) ethanol
1500
A18–B27b
NiA
2600
A2
NiB
1000
B1
NT200
1
NT200
300
Solubilized sample NiA
Variable 35
F3–vessel–B22–air1c B28 S1–air2–P950 S8
a CHS is not water soluble on its own and needs to be dissolved in CHAPS. b Buffer lines share one container. c Outlet F3 leads into a vessel that contains 1 ml of NT200 buffer, into which line B22 is submersed.
Current Protocols in Protein Science
¨ 2. Pack a 100-ml Ni-NTA column and connect to the Akta purifier at the column valve position 4. Wash the Ni-NTA column in manual mode using Milli-Q-purified water. The Ni-NTA resin can be used for ∼5 purifications.
¨ 3. Connect the NT column to the Akta purifier at the column valve position 5. The 20-ml NT column can be used many times.
Purge sample pump 4. Fill all purifier buffer inlet lines and pump heads with Milli-Q-purified water, using the manual mode. 5. Place line S3 into a container of 95% ethanol (minimum 60 ml), line S8 into water (minimum 60 ml), and purge the sample pump (see the Appendix at the end of this unit for the UNICORN 4.11 program for purging the sample pump P950). ¨ The software the authors use to operate the Akta purifier is UNICORN version 4.11. Consult the manufacturer’s instructions for syntax and details.
6. After completion of the purge program, place the purifier buffer inlet lines into the respective buffers (see Table 6.8.2).
Equilibrate columns 7. Place lines S1 and S8 into a 50-ml Falcon tube containing 35 ml of NiA buffer. After equilibration, line S1 will be transferred into the supernatant, obtained after ultracentrifugation on day 2 (see step 14 and Basic Protocol 1).
8. Place the exit lines F3-F5 into a waste container (these lines need to be rearranged after the equilibration is completed). 9. Start equilibration method (see Table 6.8.3 for buffers and purifier line designations; see the Appendix at the end of this unit for the UNICORN 4.11 method for programming the purifier for equilibration of the Ni-NTA and NT columns). The equilibration of both columns is started on Day 1 and continues overnight in an automated manner. The equilibration of both columns is completed after 12 hr.
Day 2: Purify receptor 10. Prepare the fraction collector with 75 test tubes. 11. Place line F4 into an empty 1-liter beaker to collect the Ni-NTA column flowthrough. 12. Place line F5 into an empty 500-ml beaker to collect the NT column flowthrough. 13. Place line F3 into a 250-ml Pyrex bottle containing 1 ml of buffer NT200, into which line B22 (to air sensor 1) is submersed. 14. Place line S1 into the supernatant obtained after ultracentrifugation, imidazole addition, and filtration (Basic Protocol 1). 15. Execute the method for the purification of the NTS1 fusion protein (see the Appendix at the end of this unit for details about the UNICORN 4.11 method for purifying the NTS1 fusion protein). The purifier runs through day 3.
Day 4 16. Collect NT column eluate in 1-ml fractions. 17. Analyze [3 H]NT binding (Support Protocol 3) and determine protein concentration (Support Protocol 4).
Purification of Recombinant Proteins
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Supplement 47
Table 6.8.3 Equilibration of the 100-ml Ni-NTA Superflow Column and the 20-ml NT Column
Buffer
Buffer line
Flow rate Volume Injection system pumps (ml) valve (ml/min)
Column Outlet valve position
NT1K
A13
2.0
10
Load
Bypass
NT200
B22 air sensor
2.0
5
Load
NT200
B28
2.0
10
NT0
A12
2.0
NT0/NT200
A12/B28
NT0/NT200
Gradient (% B)
Comment
F1 waste
0
Rinse line
Bypass
F1 waste
100
Rinse line
Load
Bypass
F1 waste
100
Rinse line
10
Load
Bypass
F1 waste
0
Rinse line
2.0
25
Load
Bypass
F1 waste
35
Rinse lines
A12/B28
0.7
175
Load
5
F1 waste
35
Equilibrate NT column
NT0/NT200
A12/B28
0.7
25
Load
5
F5 waste
35
Equilibrate NT column
NiA
S1 air sensor
2.0 (sample pump)
5
Inject
Bypass
F1 waste
NA
Rinse line
NiA
S8
2.0 (sample pump)
10
Inject
Bypass
F1 waste
NA
Rinse line
NiB
B1
2.0
20
Load
Bypass
F3 waste
100
Rinse line
NiA
A2
2.0
20
Load
Bypass
F4 waste
0
Rinse line
NiA
A2
2.0
800
Load
4
F1 waste
0
Equilibrate Ni-NTA column
Day 5 18. Clean the purifier with Milli-Q-purified water and wash all lines and pumps with 20% (v/v) ethanol to prevent bacterial growth, using the purifier’s manual mode. It is essential to maintain the purification equipment in good condition to obtain repeatedly optimal results. SUPPORT PROTOCOL 1
Automated Large-Scale Production of a Recombinant GPCR
EXPRESSION OF NEUROTENSIN RECEPTOR IN E. col i A maltose-binding protein (MBP) fusion approach is used in this protocol for the expression of functional, membrane-inserted neurotensin receptor. This approach promotes correct folding of receptors. The expression plasmid encodes MBP with its own signal peptide, followed by the receptor. After translocation across the cytoplasmic membrane, the signal peptide is cleaved by the leader peptidase. Therefore, the NTS1 fusion protein MBP-rT43NTR-TrxA-H10 (NTS1-624) used for purification starts with (1) the mature E. coli maltose-binding protein (MBP, Lys1 to Thr366 ), followed by (2) Gly-Ser, (3) the N-terminally truncated rat neurotensin type I receptor (NTS1; rT43NTR, Thr43 to Tyr424 ; Tanaka et al., 1990), (4) three Ala residues, (5) the E. coli thioredoxin (TrxA, Ser2 to Ala109 ; the presence of TrxA results in increased expression; see Tucker and Grisshammer, 1996), (6) Gly-Thr, and (7) a decahistidine tag (H10) (Grisshammer and Tucker, 1997). The production of functional receptors necessitates that the E. coli translocon (the probable site of membrane insertion for MBP-NTR fusion proteins) is not overloaded by the nascent receptor chain. This can be achieved by using a low-copy number plasmid with a weak promoter (see Tucker and Grisshammer, 1996) and a low temperature (22◦ C) during production. Fewer protein molecules are produced at any given moment, but accumulation of correctly folded receptors is observed over time.
6.8.8 Supplement 47
Current Protocols in Protein Science
Purification of 10 milligrams of a NTS1 fusion protein requires 250 g of cells, which is equivalent to 50 liters of cell culture. Expression at the 200-liter scale, as described below, provides material for four large-scale purifications. Fermentation can be scaled down to 50 liters if a larger fermentor is not available. Use of shake flasks would make this step rate limiting.
Materials 2× TY medium (APPENDIX 4A; 200 liters) 100 mg/ml ampicillin stock solution (140 ml) 40% (w/v) glucose stock solution (1 liter) 200-ml overnight 37◦ C culture of E. coli DH5α containing the plasmid pRG/III-hs-MBP-rT43NTR-TrxA-H10 (plasmid # 624; see Grisshammer and Tucker, 1997) 1 M isopropyl-β-D-thiogalactopyranoside (IPTG; 50 ml) 300-liter working volume fermentor (New Brunswick Scientific) Continuous flow centrifuge (Sharples As 16P) 1. In a 300-liter working volume fermentor, prepare 200 liters 2× TY medium and add 140 ml of 100 mg/ml ampicillin stock solution and 1 liter of 40% (w/v) glucose stock solution. 2. Inoculate with 200 ml of an overnight culture of E. coli DH5α containing the expression plasmid. 3. Grow cells at 37◦ C to an OD600 of 0.5 with constant stirring (200 rpm) and air supply of 1 vvm (volume per volume per minute). It takes 3 to 4 hr to reach the induction point.
4. Induce expression of the NTS1 fusion protein with 0.25 mM IPTG (by adding 50 ml of 1 M IPTG solution) and decrease the temperature to 22◦ C. 5. Harvest the cells ∼40 hr later using a continuous flow centrifuge at a flow rate of 100 liters per hr. At harvest, the cell suspension has an OD600 of ∼3.
6. Freeze the cells in liquid nitrogen, and store up to 1 year at −85◦ C. Cells can be stored for at least 1 year without compromising receptor quality.
PREPARATION OF THE NEUROTENSIN AFFINITY COLUMN Neurotensin (NT) has subnanomolar affinity for its receptor in the presence of detergents. This property can be exploited for an efficient affinity purification step (Tucker and Grisshammer, 1996; Grisshammer et al., 1999). The NT matrix described below is based on N-terminally biotinylated NT, bound to a tetrameric avidin resin. Modification of NT at its N-terminus does not interfere with receptor binding. Avidin has a high isoelectric point and needs to be succinylated to reduce nonspecific binding of E. coli proteins to the affinity matrix (Tucker and Grisshammer, 1996). Biotinylated NT is easy to obtain by solid-phase peptide synthesis.
SUPPORT PROTOCOL 2
Materials Tetrameric avidin resin suspension (Sigma) 50 mM borate buffer, pH 9.0 2 mg/ml succinic anhydride in 50 mM borate buffer, pH 9.0 (dissolves upon gentle warming) Buffer I (10 mM sodium phosphate pH 7.0) Buffer II (50 mM Tris·Cl, pH 7.4/1 mM EDTA)
Purification of Recombinant Proteins
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Supplement 47
Biotinylated NT (biotin-βAla-βAla-Gln-Leu-Tyr-Glu-Asn-Lys-Pro-Arg-Arg-ProTyr-Ile-Leu-OH), custom synthesis (a crude preparation may contain some nonbiotinylated NT) Buffer II containing 5 mM biotin Buffer II containing 500 mM NaCl Buffer II containing 3 mM azide Suitable column (e.g., 2.5 × 10 cm EconoColumn; Bio-Rad) NOTE: All steps are performed at room temperature. 1. Wash resin in 2 vol 50 mM borate buffer, pH 9.0 by gentle rocking for 10 min. 2. Resuspend resin in 2 mg/ml succinic anhydride in borate buffer (1 mg succinic anhydride per ml resin). Incubate 1.5 hr (see Dimroth, 1986). 3. Allow the resin to settle and remove the supernatant. 4. Resuspend resin in 1 vol buffer I. 5. Transfer resin into a column and allow to settle. 6. Wash resin with 10 vol buffer I. 7. Wash resin with 15 vol buffer II. 8. Incubate resin with 2-fold molar excess of biotinylated NT over resin-bound avidin groups in 2 volumes of buffer II for 1 hr. Calculate the amount of peptide needed from the biotin binding capacity of the resin and the proportion of biotinylated to nonbiotinylated peptide.
9. Wash the resin with 15 vol water. 10. Block unoccupied biotin binding sites with buffer II containing 5 mM biotin. 11. Wash the NT resin extensively with 20 vol buffer II containing 500 mM NaCl, then with 10 vol water. 12. Store NT resin at 4◦ C in buffer II containing 3 mM azide. The NT resin can be used for many years. SUPPORT PROTOCOL 3
Automated Large-Scale Production of a Recombinant GPCR
LIGAND-BINDING ASSAY USING SOLUBILIZED RECEPTORS Ligand binding with radiolabeled ligands provides a sensitive measure of the amount of functional receptors in a given preparation. Tritium-labeled neurotensin ([3 H]NT), an agonist for the neurotensin receptor, is very hydrophilic and not “sticky”, resulting in low background binding. The detergent-solubilized, purified NTR fusion protein (Basic Protocol 2) binds NT with high affinity (KD of 0.073 nM; Grisshammer et al., 1999). The dissociation rate is sufficiently slow to allow separation of the ligand-receptor complex from free ligand by centrifugation-assisted gel filtration, without losing bound agonist during this procedure. The size difference between [3 H]NT and the detergent-receptorligand complex is large enough to avoid leakage of free [3 H]NT. For routine analyses of purified receptors, the authors determine [3 H]NT binding at a concentration of 2 nM, which corresponds to ∼96.5% occupancy. In simple terms and with some assumptions, the law of mass action predicts the fractional receptor occupancy at equilibrium as a function of the ligand concentration: fractional occupancy = [ligand]/{[ligand] + KD }. A theoretical value for specific binding (Bmax ) of 10.4 nmol/mg is calculated for the NTS1-624 fusion protein (molecular mass of 96.5 kDa), assuming one ligand-binding site per receptor molecule.
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Current Protocols in Protein Science
Materials Receptor dilution buffer: 50 mM Tris·Cl, pH 7.4/1 mM EDTA/0.2% (w/v) 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS; Anatrace)/0.04% (w/v) cholesteryl hemisuccinate Tris salt (CHS, Anatrace)/0.1% (w/v) n-dodecyl-β-D-maltoside (LM; Anatrace) Solubilized NT receptor (Basic Protocol 2) 4 nM [3 H]NT (Perkin-Elmer; specific activity ∼100 Ci/mmol or 222,000 dpm/pmol) in TEBB buffer CHAPS/CHS stock solution: 6% (w/v) 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS; Anatrace)/1.2% (w/v) cholesteryl hemisuccinate Tris salt (CHS; Anatrace) detergent stock solution; dissolve CHS in CHAPS solution by stirring for several hours; prepare fresh and cool to 4◦ C LM stock solution: 10% (w/v) n-dodecyl-β-D-maltoside (LM; Anatrace) detergent stock solution in H2 O; prepare fresh and cool to 4◦ C TEBB buffer: 50 mM Tris·Cl, pH 7.4/1 mM EDTA /40 µg/ml bacitracin /0.1% (w/v) BSA 40 µM unlabeled neurotensin (NT) solution in TEBB buffer Scintillant (e.g., Ecoscint A; National Diagnostics) Bio-Spin 30 Tris columns (Bio-Rad) Scintillation counter vials (e.g., SnapCap Bio-vials; Beckman), Centrifuge with swinging-bucket rotor Liquid scintillation counter 1. Set up the required number of 1.5-ml microcentrifuge tubes as specified in Table 6.8.4. Each sample is assayed in duplicate (two tubes for total binding, two tubes for nonspecific binding, i.e., a total of four tubes per sample).
2. Incubate samples for 1 hr on ice. Equilibrium of ligand binding is reached after 1 hr on ice.
3. During incubation, prepare spin columns as follows: a. Drain columns into 2-ml microcentrifuge tubes without lids (provided by the spin column supplier). b. Apply 1 ml receptor dilution buffer. c. Allow the buffer to drip by gravity. Table 6.8.4 Pipetting Scheme for Setting Up One 1.5-ml Microcentrifuge Tube for Ligand Binding Analysis
Material/buffer
Volume (µl)
Final concentrationsa
20
≤0.4 nM
[ H]NT
75
2 nM
CHAPS/CHS stock
4.3
0.2%/0.04%
LM stock
1.3
0.1%
TEBB buffer
41.9
Unlabeled NT solution or TEBB buffer
7.5
Solubilized NT receptor 3
2 µM unlabeled NT
a The receptor preparation must be diluted with receptor dilution buffer such that the total [3 H]NT binding (specific
and nonspecific) does not exceed 15,000 dpm in the 150 µl setup, i.e., the total [3 H]NT binding should not be more than 20% of the total input of radioligand (0.3 pmoles/150 µl = 2 nM).
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d. Add another 1 ml receptor dilution buffer. e. Again allow the buffer to drip by gravity. f. Place columns into refrigerator until needed. 4. Just before use empty the 2-ml microcentrifuge tubes and place the spin columns back into tubes. 5. Place the 2-ml microcentrifuge tubes, containing the spin columns, into empty scintillation counter vials. Centrifuge the vials with the spin columns 2 min at 1000 × g, room temperature, in a swinging bucket rotor. Higher g-values will likely crush the resin and therefore lower the return of the receptor/ligand complex. The purpose of this step is to empty the columns by prespinning according to the manufacturer’s instructions.
6. Load 100 µl sample (from step 2) per spin column. 7. Place spin columns into scintillation counter vials equipped with an adapter (e.g., the upper part of a 1.5-ml microcentrifuge tube with bottom cut off and without lid), so the spin columns fit snugly into the scintillation counter vials. 8. Centrifuge columns 4 min at 1000 × g, room temperature. 9. Retrieve spin columns from centrifuge. 10. Add scintillant to the scintillation counter tubes containing receptor and bound ligand. Discard spin columns, which contain the unbound radioligand, into radioactive waste. 11. Quantify radioactivity in the samples using a liquid scintillation counter. 12. Calculate specific binding: dpm (total binding) − dpm (nonspecific binding) = dpm (specific binding). 13. Calculate pmol/ml value of the original receptor sample. Take into account the receptor dilution. 14. Calculate fractional receptor occupancy: [3 H]NT/([3 H]NT+ KD ) where the concentration of [3 H]NT in the assay is in nM and the KD for solubilized NTR is 0.073 nM (Grisshammer et al., 1999). 15. Correct for percent occupancy by dividing the pmol/ml value from step 13 by the fractional receptor occupancy from step 14. 16. Calculate final pmol/mg protein value as follows: [(pmoles/ml)/(mg protein/ml)]. See Support Protocol 4 for determination of protein concentration in mg/ml. SUPPORT PROTOCOL 4
Automated Large-Scale Production of a Recombinant GPCR
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AMIDO BLACK ASSAY FOR PROTEIN CONCENTRATION DETERMINATION Many methods for the determination of protein content are inaccurate in the presence of detergents and lipids. The Amido Black assay (Schaffner and Weissmann, 1973) includes a precipitation step to ensure that there is no interference by detergents, lipids or other buffer components. Not all proteins bind dyes such as Coomassie Blue or Amido Black with equal affinity and protein concentration determinations may be biased if dye binding is different for the protein under investigation compared to the protein used as reference (e.g., BSA). Therefore, amino acid analysis (UNIT 3.2) of the purified target protein is recommended to calibrate the Amido Black data to the actual concentration of the target membrane protein (Weiss and Grisshammer, 2002; Butler et al., 2004). Current Protocols in Protein Science
Materials 2 mg/ml BSA (Pierce Perbio, albumin standard, fraction V, no. 23209) stock solution Samples Tris-SDS solution: 1 M Tris·Cl, pH 7.5/2% (w/v) SDS 6% and 90% (w/w) trichloroacetic acid (TCA) 0.1% (w/v) Amido Black (Sigma; also called Naphthol Blue Black, Amido Black 10B, Buffalo Black NBR) solution in 45:10:45 (v/v/v) methanol/acetic acid/H2 O Wash solution: 90:2:8 (v/v/v) methanol /acetic acid/ H2 O Elution solution: 25 mM NaOH, 0.05 mM EDTA in 50% (v/v) ethanol Custom-made cylindrical Teflon block (one piece with the base 57 mm in diameter and 5 mm high; the upper part 35 mm in diameter and 27 mm high) with 12 evenly spaced holes of 3.5 mm in diameter, to fit on the filtration base 0.45-µm Millipore HA filters (HAWP 047 00) Millipore glass filtration base with stopper (XX10 047 02) for 47-mm disc filters and anodized aluminum spring clamp (XX10 047 03) Spectrophotometer with 630-nm capability 1.5-ml disposable plastic cuvettes 1. Dilute BSA stock solution to a working concentration of 200 µg/ml. 2. Set up BSA standard curve as described in Table 6.8.5. 3. Prepare samples (0.5 to 10 µg of protein) in a total volume of 225 µl. 4. Add 30 µl of Tris-SDS solution to each sample, mix. 5. Add 50 µl of 90% TCA, vortex. Leave 2 min at room temperature. 6. Assemble Teflon filter block onto filtration base with an HA filter and connect to a vacuum source. Wet the filters with water during the assembly step. 7. Filter samples onto HA filter. A maximum of 12 samples can be filtered onto one HA filter.
8. Rinse sample tubes with 200 µl of 6% TCA solution each, and filter this wash. 9. Wash filter gently in 6% TCA. 10. Stain filter for 3 min with Amido Black. 11. Rinse filter for 30 sec in water to remove excess Amido Black. 12. Rinse filter 3 times, 1 min each time with the wash solution. 13. Rinse filter for 3 min with water. 14. Blot filter dry, cut out blue spots containing stained protein. Table 6.8.5 Pipetting Scheme for Amido Black Standard Curve
µg BSA
µl BSA (200 µg/ml)
µl H2 O
0
0
225
2
10
215
4
20
205
8
40
185
12
60
165
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15. Elute Amido Black dye in 2-ml tubes with 1 ml of elution solution, vortex periodically over a period of about 10 min. 16. Measure absorbance of samples at 630 nm in 1.5-ml disposable plastic cuvettes. 17. Calculate protein concentration (in mg/ml) by comparison to the standard curve and linear regression. The protein concentration determination by the Amido Black method may be biased if dye-binding is different for the protein under investigation compared to BSA used as reference. Therefore, correct the Amido Black data to the actual concentration obtained by amino acid analysis, if necessary. No correction is required for the NTS1-624 fusion protein because the values obtained by the Amido Black assay have been confirmed by amino acid analysis.
COMMENTARY Background Information
Automated Large-Scale Production of a Recombinant GPCR
Finding a rational approach for the highlevel expression of recombinant integral membrane proteins is still a challenge. Mechanistic details are still sparse as to why some membrane proteins are produced at high levels in membrane-inserted, functional form but closely related targets are not. This lack of rationalization often results in a trial-and-error approach to obtain recombinant membrane protein in quantities sufficient for purification attempts. Two recent reports warrant emphasis: First, elegant work combining theory with biochemical approaches provides insight into how transmembrane helices are recognized by the endoplasmic reticulum translocon (biological hydrophobicity scale; Hessa et al., 2005a,b). Second, successful recombinant overexpression of some membrane proteins seems to be linked to avoidance of stress responses in host cells (Griffith et al., 2003) and can be related to the differential expression of genes involved in membrane protein secretion and cellular physiology (Bonander et al., 2005). Simple approaches for increasing recombinant membrane protein yields may therefore be unsuccessful. It may be worth considering a whole-cell approach for developing general strategies to obtain high levels of functional membrane proteins, especially for eukaryotic targets. Heterologous expression and purification of membrane proteins has been covered by a number of reviews (Grisshammer and Tate, 1995; Tate, 2001; Sarramegna et al., 2003; Tate and Grisshammer, 2003). In these the reader will find information on both prokaryotic and eukaryotic membrane proteins. Until recently, all structures of eukaryotic membrane proteins (e.g., the visual pigment rhodopsin; Palczewski et al., 2000; Li et al., 2004) had been solved with material from natural sources, re-
flecting the difficulty of producing functional eukaryotic membrane proteins in heterologous expression systems. Therefore, the recently published structure of a recombinant mammalian voltage-gated potassium channel, produced in the yeast Pichia pastoris, is good news (Long et al., 2005). Crystallization and other applications (e.g., nuclear magnetic resonance; see Reeves et al., 1999; Inooka et al., 2001; Williamson et al., 2002; Luca et al., 2003) require milligram quantities of functional receptors. Only a few receptors have been purified in such quantities (Hulme and Curtis, 1998; Ohtaki et al., 1998; Klaassen et al., 1999; Reeves et al., 2002; Weiss and Grisshammer, 2002; Warne et al., 2003). Given the current expression levels of the NTR fusion protein (see Support Protocol 1), the authors have optimized the protocol for immobilized metal affinity chromatography by using receptor fusion proteins with C-terminal tails of ten (rather than six) histidine residues (see Grisshammer and Tucker, 1997) in combination with Ni-NTA resin. The tight binding to Ni-NTA resin of decahistidine-tagged receptors allows stringent washing steps using 50 mM imidazole. Imidazole at this concentration eliminates binding of most of the E. coli contaminants to the Ni-NTA resin, but it does not lead to elution of the fusion protein. This strategy not only results in efficient purification of receptors from crude membranes, but works particularly well for purification from total cell lysate (Grisshammer and Tucker, 1997). Using whole cells as starting material is of advantage, since the preparation of crude membranes on a large scale is time consuming. This unit describes the use of Ni-NTA resin for the purification of the NTS1 fusion protein. However, other metal ions and resin types may equally be suitable for such a first purification step and have, in fact, been explored for
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other receptors (e.g., see Hulme and Curtis, 1998; Warne et al., 2003). The apparent capacity of the Ni-NTA resin in the buffer system described in this unit is low (<0.5 mg of NTS1 fusion protein/ml resin); hence, it is necessary to use large columns and large (and expensive) amounts of detergent-containing buffers. There is no obvious explanation for this observation; however, if binding of the affinity tag to the resin is too slow, then batch loading may be considered, which often results in much higher receptor yields (e.g., see Weiss and Grisshammer, 2002). However, batch loading cannot be automated. The apparent affinity of NT for NTS1 is reduced in the presence of the high concentrations of sodium ions and imidazole in the NiB buffer (Grisshammer et al., 1999) precluding binding of functional receptors contained in the Ni-NTA column eluate to the NT column. The concentration of NaCl and imidazole is therefore reduced from 200 mM to 70 mM by dilution with buffer (see Fig. 6.8.1 and Table 6.8.1), to allow binding of functional NTS1 to the NT column. The NT column flowthrough contains a considerable amount of receptors that do not bind ligand. This receptor population may be slightly misfolded, but it is still soluble in detergent solution (as opposed to heavily aggregated material which cannot be maintained in detergent-containing buffer). These misfolded receptors may arise from inefficient insertion and folding in the E. coli cytoplasmic membrane. Therefore, the availability of a high-affinity ligand resin, specific for functional, correctly folded receptors proves very powerful in isolating receptors of high quality. The fusion protein NTS1-624 described above does not allow removal of the MBP and TrxA-H10 moieties. To obtain receptor without fusion partners, one must use different expression constructs where rT43NTR is surrounded by tobacco etch virus (Tev) protease recognition sites (White et al., 2004). Incubation of such a purified receptor fusion protein with Tev protease generates rT43NTR, MBP, and TrxA-H10. The receptor can be isolated from this mixture by size-exclusion chromatography (White et al., 2004).
Critical Parameters and Troubleshooting Proper handling of solubilized membrane proteins is mandatory for good results. Many integral membrane proteins are quite stable in a membrane environment, but become destabilized in the presence of detergents. Therefore
it is of advantage to keep the target membrane protein cold during all stages of the solubilization and purification. Avoid the generation of foam when handling membrane proteins in detergent solutions; receptors at the air-liquid interface will denature. Equally important is the source of detergents. Only high-quality (high-purity) detergents should be considered for work with membrane proteins. CHS Tris salt and the detergents LM and CHAPS are powders and should be stored at −20◦ C in a box containing desiccant. Before use, bring the bottles to room temperature to avoid condensation of water inside the detergent bottle.
Anticipated Results Typical results for the solubilization and purification of the NTS1-624 fusion protein are given in Table 6.8.6 and Figure 6.8.2. For Support Protocol 1 (Expression of neurotensin receptor in E. coli), radioligand binding analyses with intact cells typically indicates ∼1000 receptors/cell. The cell density at harvest is between 2.5 and 3 OD600 . One 200liter run yields about 1.1 kg of wet bacterial cell paste containing 90 to 100 mg of functional receptor fusion protein (White et al., 2004).
Time Considerations The purification of 10 mg of an NTS1 fusion protein (see Table 6.8.6) requires 4 days for completion. The preparation of detergent stocks and buffers requires most of day 1. The equilibration of the columns is started at the end of day 1 and proceeds overnight. Day 2 is dedicated to cell resuspension and receptor solubilization, which takes 6 to 7 hr. The loading of the Ni-NTA column is started as soon as the material is available. Solubilized receptors are then analyzed by ligand binding and protein assay, completing day 2. No hands-on time is required on day 3 and most of day 4 while the purification process is underway. The purified NTS1 fusion protein becomes available for analysis in the afternoon of day 4. The washing of the Ni-NTA and NT columns completes automatically overnight. On day 5, the purifier is cleaned with water, and all lines and pumps are rinsed into 20% (v/v) ethanol to prevent bacterial growth. This cleaning requires less than 1 hr.
Acknowledgements The authors’ research is supported by the Intramural Research Program of the NIH, National Institute of Diabetes and Digestive and Kidney Diseases.
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Table 6.8.6 Automated Large-scale (10-mg) Purification of the Fusion Protein NTS1-624a, b
Volume (ml)
Functional receptor (nmol)
Protein (mg)
Supernatant
929
198.2
15025
0.011
1
100
d
1060
21.3
ND
ND
NA
11
122
145.5
30
5.2
473
73 (100)
NTFT
411
4.3
14
ND
NA
2 (3)
NTE
14
94.3
9.9
9.6
873
48 (65)
NiFT e
NiE
e
Specific Purification binding (-fold) (nmol/mg)c
Recovery (%)f
a Data for the 10-mg preparation are the average of five purifications. b Abbreviations: NA, not applicable; ND, not determined; NiE, Ni-NTA column eluate; NiFT, flowthrough of Ni-
NTA column; NTE, NT column eluate; NTFT, flowthrough of NT column. c A theoretical value for specific binding of 10.4 nmol/mg is calculated for NTS1-624 (molecular mass of 96.5 kDa)
assuming one ligand binding site per receptor molecule. d NiFT contains the actual flowthrough and buffer used to wash the sample pump. e The volume of the NTFT is larger than that of the NiE because of the dilution applied to reduce the NaCl and imidazole concentrations. f Recovery data listed in parentheses refer to the NT column.
Figure 6.8.2 SDS-PAGE of NTS1 fusion protein purification fractions. A 10-mg preparation of the fusion protein NTS1-624 was purified on a 100-ml Ni-NTA column followed by a 20-ml NT column, starting from 250 g of cells. The progress of purification was monitored by SDS-PAGE (NuPAGE 4% to 12% Bis-Tris gel, Invitrogen; 1× MES buffer) and Coomassie R-250 staining. Lane M: Novagen Perfect Protein Marker (15 to 150 kDa); lane 1: 10 µg supernatant; lane 2: 10 µg Ni-NTA column flowthrough; lane 3: 5 µg Ni-NTA column eluate; lane 4: 10 µg NT column flowthrough; lane 5: 5 µg NT column eluate. Reprinted by permission of the Federation of the European Biochemical Societies from White et al., 2004. Automated Large-Scale Production of a Recombinant GPCR
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Current Protocols in Protein Science
Literature Cited Bonander, N., Hedfalk, K., Larsson, C., Mostad, P., Chang, C., Gustafsson, L., and Bill, R.M. 2005. Design of improved membrane protein production experiments: Quantitation of the host response. Protein Sci. 14:1729-1740. Butler, P.J., Ubarretxena-Belandia, I., Warne, T., and Tate, C.G. 2004. The Escherichia coli multidrug transporter EmrE is a dimer in the detergent-solubilised state. J. Mol. Biol. 340:797-808. Dimroth, P. 1986. Preparation, characterization, and reconstitution of oxaloacetate decarboxylase from Klebsiella aerogenes, a sodium pump. Methods in Enzymology 125:530-540. Griffith, D.A., Delipala, C., Leadsham, J., Jarvis, S.M., and Oesterhelt, D. 2003. A novel yeast expression system for the overproduction of quality-controlled membrane proteins. FEBS Lett. 553:45-50. Grisshammer, R. and Tate, C.G. 1995. Overexpression of integral membrane proteins for structural studies. Q. Rev. Biophys. 28:315-422. Grisshammer, R. and Tucker, J. 1997. Quantitative evaluation of neurotensin receptor purification by immobilized metal affinity chromatography. Protein Expression and Purification 11:53-60. Grisshammer, R., Averbeck, P., and Sohal, A.K. 1999. Improved purification of a rat neurotensin receptor expressed in Escherichia coli. Biochemical Society Transactions 27:899-903. Hessa, T., Kim, H., Bihlmaier, K., Lundin, C., Boekel, J., Andersson, H., Nilsson, I., White, S.H., and von Heijne, G. 2005a. Recognition of transmembrane helices by the endoplasmic reticulum translocon. Nature 433:377-381. Hessa, T., White, S.H., and von Heijne, G. 2005b. Membrane insertion of a potassium-channel voltage sensor. Science 307:1427. Hulme, E.C. and Curtis, C.A.M. 1998. Purification of recombinant M1 muscarinic acetylcholine receptor. Biochem. Soc. Trans. 26:S361. Inooka, H., Ohtaki, T., Kitahara, O., Ikegami, T., Endo, S., Kitada, C., Ogi, K., Onda, H., Fujino, M., and Shirakawa, M. 2001. Conformation of a peptide ligand bound to its G-protein coupled receptor. Nature Structural Biology 8:161-165. Klaassen, C.H., Bovee-Geurts, P.H., Decaluwe, G.L., and DeGrip, W.J. 1999. Large-scale production and purification of functional recombinant bovine rhodopsin with the use of the baculovirus expression system. Biochem. J. 342:293-300. Li, J., Edwards, P.C., Burghammer, M., Villa, C., and Schertler, G.F.X. 2004. Structure of bovine rhodopsin in a trigonal crystal form. J. Mol. Biol. 343:1409-1438. Long, S.B., Campbell, E.B., and MacKinnon, R. 2005. Crystal structure of a mammalian voltagedependent Shaker family K+ channel. Science 309:897-903.
Luca, S., White, J.F., Sohal, A.K., Filippov, D.V., van Boom, J.H., Grisshammer, R., and Baldus, M. 2003. The conformation of neurotensin bound to its G protein-coupled receptor. Proc. Natl. Acad. Sci. U.S.A. 100:10706-10711. Ohtaki, T., Ogi, K., Masuda, Y., Mitsuoka, K., Fujiyoshi, Y., Kitada, C., Sawada, H., Onda, H., and Fujino, M. 1998. Expression, purification, and reconstitution of receptor for pituitary adenylate cyclase-activating polypeptide: Large-scale purification of a functionally active G protein-coupled receptor produced in Sf9 insect cells. J. Biol. Chem. 273:15464-15473. Palczewski, K., Kumasaka, T., Hori, T., Behnke, C.A., Motoshima, H., Fox, B.A., Le Trong, I., Teller, D.C., Okada, T., Stenkamp, R.E., Yamamoto, M., and Miyano, M. 2000. Crystal structure of rhodopsin: A G protein-coupled receptor. Science 289:739-745. Reeves, P.J., Klein-Seetharaman, J., Getmanova, E.V., Eilers, M., Loewen, M.C., Smith, S.O., and Khorana, H.G. 1999. Expression and purification of rhodopsin and its mutants from stable mammalian cell lines: application to NMR studies. Biochem. Soc. Trans. 27:950-955. Reeves, P.J., Kim, J.-M., and Khorana, H.G. 2002. Structure and function in rhodopsin: A tetracycline-inducible system in stable mammalian cell lines for high-level expression of opsin mutants. Proc. Natl. Acad. Sci. 99:1341313418. Sarramegna, V., Talmont, F., Demange, P., and Milon, A. 2003. Heterologous expression of G-protein-coupled receptors: Comparison of expression systems from the standpoint of largescale production and purification. Cell. Mol. Life Sci. 60:1529-1546. Schaffner, W. and Weissmann, C. 1973. A rapid, sensitive, and specific method for the determination of protein in dilute solution. Anal. Biochem. 56:502-514. Tanaka, K., Masu, M., and Nakanishi, S. 1990. Structure and functional expression of the cloned rat neurotensin receptor. Neuron 4:847854. Tate, C.G. 2001. Overexpression of mammalian integral membrane proteins for structural studies. FEBS Lett. 504:94-98. Tate, C.G. and Grisshammer, R. (eds.) 2003. Special Issue on Overexpression of Integral Membrane Proteins. Biochim. Biophys. ActaBiomembranes 1610:1-153. Tucker, J. and Grisshammer, R. 1996. Purification of a rat neurotensin receptor expressed in Escherichia coli. Biochem. J. 317:891-899. Warne, T., Chirnside, J., and Schertler, G.F.X. 2003. Expression and purification of truncated, non-glycosylated turkey beta-adrenergic receptors for crystallization. Biochim. Biophys. Acta 1610:133-140. Weiss, H.M. and Grisshammer, R. 2002. Purification and characterization of the human
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adenosine A2a receptor functionally expressed in Escherichia coli. Eur. J. Biochem. 269:8292. White, J.F., Trinh, L.B., Shiloach, J., and Grisshammer, R. 2004. Automated large-scale purification of a G-protein-coupled receptor for neurotensin. FEBS Letters 564:289-293. Williamson, P.T.F., Bains, S., Chung, C., Cooke, R., and Watts, A. 2002. Probing the environment of neurotensin whilst bound to the neurotensin receptor by solid-state NMR. FEBS Letters 518:111-115.
Contributed by Jim F. White and Reinhard Grisshammer NIDDK, NIH Bethesda, Maryland
Automated Large-Scale Production of a Recombinant GPCR
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¨ APPENDIX: UNICORN 4.11 PROGRAMS FOR OPERATING THE AKTA PURIFIER ¨ The software the authors use to operate the Akta purifier is the UNICORN version 4.11 (see the manufacturer’s instructions for details). Newer UNICORN versions are available, and may require a slightly different syntax than that for UNICORN 4.11. UNICORN 4.11 Program for Purging Sample Pump P950 Main method: Main 0.00 Base Volume 0.00 ColumnPosition Position1Bypass 0.00 InjectionValve Waste 0.00 Flow 0.10 {ml/min} 0.00 Gradient 0.00 {%B} 0.00 {base} 0.00 BufferValveA1 A11 0.00 Block water wash P950 S8 water wash P950 S8 0.00 Base SameAsMain 0.00 Flow 0.00 {ml/min} 0.00 InjectionValve Waste 0.00 SampleValve S8 0.00 DirectLoad 5.00 {ml/min} 10.00 {ml} No 0.10 {ml/min} 0.00 End block 0.00 Block ethanol wash P950 S3 ethanol wash P950 S3 0.00 Base SameAsMain 0.00 Flow 0.00 {ml/min} 0.00 InjectionValve Waste 0.00 SampleValve S3 0.00 DirectLoad 5.00 {ml/min} 50.00 {ml} No 0.10 {ml/min} 0.00 End block Purification of Recombinant Proteins
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0.00 Block water wash P950 S8 again water wash P950 S8 again 0.00 Base SameAsMain 0.00 Flow 0.00 {ml/min} 0.00 InjectionValve Waste 0.00 SampleValve S8 0.00 DirectLoad 1.00 {ml/min} 5.00 {ml} No 0.10 {ml/min} 0.00 End block 0.00 Block water wash P950 S8 again again water wash P950 S8 again again 0.00 Base SameAsMain 0.00 Flow 0.00 {ml/min} 0.00 InjectionValve Waste 0.00 SampleValve S8 0.00 DirectLoad 10.00 {ml/min} 30.00 {ml} No 0.10 {ml/min} 0.00 End block 0.00 End method
UNICORN 4.11 Program for Equilibrating the Ni-NTA and NT Columns Main method: Main 0.00 Base Volume 0.00 Block Line washes for NT column Line washes for NT column 0.00 Base SameAsMain 0.00 Flow 2.00 {ml/min} 0.00 Gradient 0.00 {%B} 0.00 {base} 0.00 PumpAInlet A1 0.00 BufferValveA1 A13 Automated Large-Scale Production of a Recombinant GPCR
0.00 InjectionValve Load 0.00 ColumnPosition Position1Bypass
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0.00 OutletValve WasteF1 10.00 Gradient 100 {%B} 0.00 {base} 10.00 PumpBInlet B2 10.00 Valve8 2 15.00 Gradient 100.00 {%B} 0.00 {base} 15.00 PumpBInlet B2 15.00 Valve8 8 25.00 Gradient 0.00 {%B} 0.00 {base} 25.00 BufferValveA1 A12 25.00 PumpAInlet A1 35.00 Gradient 35.00 {%B} 0.00 {base} 35.00 BufferValveA1 A12 35.00 PumpAInlet A1 35.00 PumpBInlet B2 35.00 Valve8 8 60.00 End block 0.00 Block Equilibrate NT column Equilibrate NT column 0.00 Base SameAsMain 0.00 Flow 0.70 {ml/min} 0.00 Gradient 35.00 {%B} 0.00 {base} 0.00 ColumnPosition Position5 0.00 BufferValveA1 A12 0.00 PumpAInlet A1 0.00 PumpBInlet B2 0.00 Valve8 8 0.00 InjectionValve Load 0.00 OutletValve WasteF1 175.00 OutletValve F5 200.00 End block Purification of Recombinant Proteins
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0.00 Block Line wash S1 for NiNTA column Line wash S1 for NiNTA column 0.00 Base SameAsMain 0.00 Flow 0.00 {ml/min} 0.00 InjectionValve Inject 0.00 ColumnPosition Position1Bypass 0.00 OutletValve WasteF1 0.00 SampleValve S1 0.00 DirectLoad 2.00 {ml/min} 5.00 {ml} No 0.10 {ml/min} 0.00 End block 0.00 Block Line wash S8 for NiNTA column Line wash S8 for NiNTA column 0.00 Base SameAsMain 0.00 Flow 0.00 {ml/min} 0.00 InjectionValve Inject 0.00 ColumnPosition Position1Bypass 0.00 OutletValve WasteF1 0.00 SampleValve S8 0.00 DirectLoad 2.00 {ml/min} 10.00 {ml} No 0.10 {ml/min} 0.00 End block 0.00 Block Line washes for NiNTA column Line washes for NiNTA column 0.00 Base SameAsMain 0.00 DirectLoad 0.00 {ml/min} 0.00 {ml} No 0.10 {ml/min} 0.00 PumpBInlet B1 0.00 Flow 2.00 {ml/min} 0.00 Gradient 100.00 {%B} 0.00 {base} 0.00 InjectionValve Load Automated Large-Scale Production of a Recombinant GPCR
0.00 ColumnPosition Position1Bypass 0.00 OutletValve F3
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20.00 Flow 2.00 {ml/min} 20.00 PumpAInlet A2 20.00 Gradient 0.00 {%B} 0.00 {base} 20.00 InjectionValve Load 20.00 ColumnPosition Position1Bypass 20.00 OutletValve F4 40.00 End block 0.00 Block Equilibrate NiNTA column Equilibrate NiNTA column 0.00 Base SameAsMain 0.00 DirectLoad 0.00 {ml/min} 0.00 {ml} No 0.10 {ml/min} 0.00 Flow 2.00 {ml/min} 0.00 Gradient 0.00 {%B} 0.00 {base} 0.00 InjectionValve Load 0.00 ColumnPosition Position4 0.00 OutletValve WasteF1 0.00 PumpAInlet A2 800.00 End block 0.00 End method
UNICORN 4.11 Program for Purifying the NTS1 Fusion Protein Main method: Main 0.00 Base CV 100.00 {ml} Any 0.00 Flow 1.00 {ml/min} 0.00 AveragingTimeUV 2.56 {sec} 0.00 Alarm Pressure Enabled (10.00){\#}Alarm pressure {MPa} 0.00 {MPa} 0.00 Block Direct load sample Direct load sample 0.00 Base Volume 0.00 AutoZeroUV
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0.00 OutletValve F4 0.00 Flow 0.00 {ml/min} 0.00 PumpAInlet A2 0.00 Gradient 0.00 {%B} 0.00 {base} 0.00 ColumnPosition Position4 0.00 InjectionValve Inject 0.00 MethodBase SamplePump 0.00 SampleValve S1 0.00 SampleFlow 2.00 {ml/min} 5.00 Watch AirSensor2 Equal 1 END BLOCK 2000.00 SampleFlow 0.00 {ml/min} 2000.00 End block 0.00 Block Rinse sample line Rinse sample line 0.00 Base Volume 0.00 MethodBase SamplePump 0.00 Watch off AirSensor2 0.00 SampleValve S8 0.00 OutletValve F4 0.00 Flow 0.00 {ml/min} 0.00 Gradient 0.00 {%B} 0.00 {base} 0.00 PumpAInlet A2 0.00 InjectionValve Inject 0.00 ColumnPosition Position4 0.00 SampleFlow 2.00 {ml/min} 10.00 MethodBase SystemPump 10.00 SampleFlow 0.00 {ml/min} 10.00 End block 0.00 Block Wash Ni NTA Automated Large-Scale Production of a Recombinant GPCR
Wash Ni NTA 0.00 Base SameAsMain 0.00 MethodBase SystemPump
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0.00 SampleFlow 0.00 {ml/min} 0.00 OutletValve F4 0.00 Gradient 0.00 {%B} 0.00 {base} 0.00 PumpAInlet A2 0.00 Flow 2.00 {ml/min} 0.00 ColumnPosition Position4 0.00 InjectionValve Load 1.40 OutletValve WasteF1 15.00 End Block 0.00 Block Prep system for elution Ni NTA Prep system for elution Ni NTA 0.00 Base Volume 0.00 Gradient 100 {%B} 0.00 {base} 0.00 PumpBInlet B1 0.00 InjectionValve Load 0.00 OutletValve WasteF1 0.00 ColumnPosition Position1Bypass 0.00 Alarm Pressure Enabled 10.00 {MPa} 0.00 {MPa} 0.00 Flow 5.00 {ml/min} 20.00 End block 0.00 Block Elution Ni NTA Elution Ni NTA 0.00 Base SameAsMain 0.00 Flow 2.00 {ml/min} 0.00 Gradient 100 {%B} 0.00 {base} 0.00 InjectionValve Load 0.00 ColumnPosition Position4 0.00 PumpBInlet B1 0.00 OutletValve WasteF1 0.00 Alarm Pressure Enabled 10.00 {MPa} 0.00 {MPa} 0.72 OutletValve F3 1.94 OutletValve WasteF1
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1.94 Flow 2.00 {ml/min} 7.00 End Block 0.00 Block Wash B28 with NT200 buffer Wash B28 with NT200 buffer 0.00 Base Volume 0.00 Flow 5.00 {ml/min} 0.00 Gradient 100.00 {%B} 0.00 {base} 0.00 InjectionValve Load 0.00 ColumnPosition Position1Bypass 0.00 OutletValve WasteF1 0.00 PumpBInlet B2 0.00 Valve8 8 25.00 End block 0.00 Block Wash A12 with NT0 buffer Wash A12 with NT0 buffer 0.00 Base Volume 0.00 Flow 5.00 {ml/min} 0.00 Gradient 0.00 {%B} 0.00 {base} 0.00 InjectionValve Load 0.00 ColumnPosition Position1Bypass 0.00 OutletValve WasteF1 0.00 BufferValveA1 A12 0.00 PumpAInlet A1 25.00 End block 0.00 Block Prep system for NT column Prep system for NT column 0.00 Base Volume 0.00 Flow 5.00 {ml/min} 0.00 Gradient 35.00 {%B} 0.00 {base} Automated Large-Scale Production of a Recombinant GPCR
0.00 InjectionValve Load 0.00 ColumnPosition Position1Bypass 0.00 OutletValve WasteF1
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0.00 BufferValveA1 A12 0.00 PumpAInlet A1 0.00 PumpBInlet B2 0.00 Valve8 8 50.00 End block 0.00 Block Load dilute NiE onto NT column Load dilute NiE onto NT column 0.00 Base Volume 0.00 Flow 0.40 {ml/min} 0.00 Gradient 35.00 {%B} 0.00 {base} 0.00 PumpBInlet B2 0.00 Valve8 2 0.00 BufferValveA1 A12 0.00 PumpAInlet A1 0.00 InjectionValve Load 0.00 ColumnPosition Position5 0.00 AutoZeroUV 0.00 OutletValve F5 0.00 Alarm Pressure Enabled 10.00 {MPa} 0.00 {MPa} 5.00 Watch AirSensor1 Equal 1 END BLOCK 700.00 End Block 0.00 Block Rinse B2 line wash NT column Rinse B2 line wash NT column 0.00 Base Volume 0.00 Watch off AirSensor1 0.00 Flow 0.40 {ml/min} 0.00 Gradient 35.00 {%B} 0.00 {base} 0.00 BufferValveA1 A12 0.00 PumpAInlet A1 0.00 PumpBInlet B2 0.00 Valve8 8 0.00 InjectionValve Load
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0.00 ColumnPosition Position5 0.00 OutletValve F5 50.00 End Block 0.00 Block Wash NT column Wash NT column 0.00 Base Volume 0.00 Flow 0.70 {ml/min} 0.00 Gradient 35.00 {%B} 0.00 {base} 0.00 BufferValveA1 A12 0.00 PumpAInlet A1 0.00 PumpBInlet B2 0.00 Valve8 8 0.00 InjectionValve Load 0.00 ColumnPosition Position5 0.00 OutletValve WasteF1 160.00 End Block 0.00 Block Prep NT column for elution Prep NT column for elution 0.00 Base Volume 0.00 Flow 5.00 {ml/min} 0.00 Gradient 0.00 {%B} 0.00 {base} 0.00 PumpAInlet A1 0.00 BufferValveA1 A13 0.00 InjectionValve Load 0.00 ColumnPosition Position1Bypass 0.00 OutletValve WasteF1 50.00 End block 0.00 Block Elution NT column Elution NT column Automated Large-Scale Production of a Recombinant GPCR
0.00 Base Volume 0.00 Flow 0.50 {ml/min} 0.00 Gradient 0.00 {%B} 0.00 {base}
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0.00 InjectionValve Load 0.00 PumpAInlet A1 0.00 BufferValveA1 A13 0.00 ColumnPosition Position5 0.00 Fractionation 12mm 1.00 {ml} FirstTube Volume 75.00 Flow 0.70 {ml/min} 75.00 FractionationStop 150.00 End Block 0.00 Block Wash NiNTA column Wash NiNTA column 0.00 Base Volume 0.00 Flow 5.00 {ml/min} 0.00 Gradient 0.00 {%B} 0.00 {base} 0.00 InjectionValve Load 0.00 ColumnPosition Position1Bypass 0.00 OutletValve WasteF1 0.00 BufferValveA1 A11 0.00 PumpAInlet A1 20.00 ColumnPosition Position4 20.00 Flow 3.00 {ml/min} 520.00 End Block 0.00 Block Prep for NT column wash Prep for NT column wash 0.00 Base Volume 0.00 Flow 5.00 {ml/min} 0.00 Gradient 0.00 {%B} 0.00 {base} 0.00 InjectionValve Load 0.00 ColumnPosition Position1Bypass 0.00 OutletValve WasteF1 0.00 BufferValveA1 A16 0.00 PumpAInlet A1 20.00 BufferValveA1 A17
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40.00 End Block 0.00 Block Wash NT column with A3 B3 Wash NT column with A3 B3 0.00 Base Volume 0.00 Flow 0.70 {ml/min} 0.00 Gradient 0.00 {%B} 0.00 {base} 0.00 InjectionValve Load 0.00 ColumnPosition Position5 0.00 OutletValve WasteF1 0.00 PumpAInlet A1 0.00 BufferValveA1 A17 150.00 BufferValveA1 A16 300.00 End Block 0.00 Block Wash pumps A and B with water Wash pumps A and B with water 0.00 Base Volume 0.00 Flow 5.00 {ml/min} 0.00 Gradient 50.00 {%B} 0.00 {base} 0.00 InjectionValve Load 0.00 ColumnPosition Position1Bypass 0.00 OutletValve WasteF1 0.00 PumpAInlet A1 0.00 BufferValveA1 A11 0.00 PumpBInlet B2 0.00 Valve8 1 100.00 End Block 0.00 Block Wash pumps A and B with EtOH Wash pumps A and B with EtOH 0.00 Base Volume Automated Large-Scale Production of a Recombinant GPCR
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CHAPTER 7 Characterization of Recombinant Proteins INTRODUCTION
O
nce a protein has been purified, it is necessary to characterize the protein preparation. The initial and most fundamental analyses performed are checks of purity and of biological or enzymatic activity. Indeed, to purify a protein in the first place a specific assay for the protein must be available, and limited purity determinations are usually made en route to the finished product by SDS-PAGE monitoring of the sample at different purification stages. The first unit of this chapter (UNIT 7.1) provides an overview of the approaches and methods used for more detailed characterization of recombinant proteins. It should be noted that none of the techniques or methods mentioned are used uniquely for recombinant proteins, and protocols dealing with some of the methods are included elsewhere in this book. However, the emphasis of the discussion in UNIT 7.1 (and of the protocols which will appear in future updates) is on addressing the detection and determination of the most pertinent sources of contamination and heterogeneity commonly associated with proteins isolated from recombinant sources. The first section of the overview describes common sources of contamination, both proteinaceous and nonproteinaceous. Because these contaminants are derived from the host cell, their specific characteristics will depend on the particular expression system used (also see Chapter 5). Once the purity of the protein has been established, the primary structure (amino acid sequence) is checked by standard protein-chemical methodologies. Throughout the overview, the use of HPLC and mass spectrometry is emphasized; these are important tools for checking the primary structure and especially for detecting both nonspecific and specific post-translational modifications. Because many proteins expressed in E. coli form insoluble aggregates known as inclusion bodies, before purification they must be extracted under conditions that unfold the protein (see UNITS 6.1, 6.3 & 6.5). After folding and purification of the protein, there is always the possibility that varying amounts of misfolded conformers may be present. Misfolded protein can be considered as a contaminant. A number of spectroscopic and other methods used to study protein conformation and structure are tabulated in the overview. Protocols describing the most commonly used of these methods are included in this chapter (see UNITS 7.2, 7.4, 7.6, 7.7 & 7.9). Proteins expressed in E. coli can retain N-terminal methionine derived from the initiating N-formylmethionine. The retention or nonretention of N-terminal methionine is a potential source of heterogeneity, and is discussed in the last section of the overview (also see UNIT 6.1). In UNIT 7.2 the analysis of proteins using ultraviolet (UV) absorption spectroscopy is described. The first part deals with direct (or batch-mode) spectroscopic measurement, in which the purified protein is simply placed in a quartz cuvette and analyzed. Data collection and interpretation protocols allow assessment of the identity and, to a lesser degree, the purity of the protein. The second part of UNIT 7.2 describes the analysis of proteins simultaneously separated by HPLC chromatography. This can be considered to be a two-dimensional method in which the first dimension is chromatography; the peaks thus separated are spectroscopically analyzed using diode array detection in the second Contributed by Paul T. Wingfield and Roger H. Pain Current Protocols in Protein Science (2003) 7.0.1-7.0.4 Copyright © 2003 by John Wiley & Sons, Inc.
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dimension. Both batch-mode and column-based spectroscopic analysis complement purity analysis using SDS-PAGE (Chapter 10) in that the former methods can detect the presence of nonproteinaceous contaminants, especially nucleic acid. The primary sequence of a recombinant protein is known from the cDNA coding sequence. However, the sequence of an isolated recombinant-derived protein may differ from the predicted sequence due to, for example, post-translational modifications; it is therefore important to confirm its primary sequence. The amount of analysis required to do this will depend on the intended use of the protein. For many investigators the analysis may consist of limited N-terminal and C-terminal sequencing, peptide mapping, and either amino acid analysis (UNIT 3.2) or molecular weight determination using mass spectrometry (Chapter 16). On the other hand, a full structural determination is usually required for proteins intended for pharmaceutical use. describes some techniques that give information on the primary structure. The method most commonly used to review the primary sequence of a protein is peptide mapping. Peptides derived from protein chemically or enzymatically fragmented are separated by HPLC or electrophoresis. The interpretation of the resultant chromatograms, or band patterns, is based on the specificity of the cleavage method (usually very selective) and the expected sequence of the protein. Often comparisons are made with well-characterized standards. The most common post-translation modification is the oxidation of paired sulfhydryl residues to form a disulfide bond. The technique of peptide mapping is usually used to determine which sulfhydryl residues are paired (the linkage pattern). Specific details of HPLC methodology (Chapter 11) and gel electrophoresis (Chapter 10) are detailed elsewhere and peptide analysis using mass spectrometry will be described in Chapter 16. The electrophoretic technique of two-dimensional titration curve analysis is also included in UNIT 7.3, as it can give information on the charged residue composition of the protein in addition to guiding the selection of ion-exchange resins used for protein purification.
UNIT 7.3
The topic of protein conformation is addressed in UNIT 7.4, in which the electrophoretic method known as transverse urea-gradient gel electrophoresis (UGGE) is described. The unfolding of a protein exposed to a linear gradient of urea (usually 0 to 8 M) is monitored simply by observation of the electrophoretic mobility. Gel patterns can be analyzed to estimate conformational stability and help identify covalent or conformational heterogeneity. Spectroscopic methods such as circular dichroism (UNIT 7.6) and fluorescence (UNIT 7.7), and to a lesser degree, UV-spectroscopy (UNIT 7.2), are among the standard methods used to analyze protein conformation, and descriptions of these methods are planned for this chapter. However, for investigators not wishing or not able to invest in expensive and dedicated equipment, the gel electrophoretic approach described herein may be a useful alternative, although not as flexible or rigorous (potentially) as spectroscopic and calorimeter-based methods. For a purified protein, determination of the molecular weight and physical homogeneity in solution are of fundamental importance. With recombinant proteins the molecular weight can be predicted from the cDNA coding sequence and rapidly confirmed by mass spectroscopy (Chapter 16). Determination of the molecular weight of the native protein in solution is not so straightforward, but is still fairly routine using the modern analytical ultracentrifuge. UNIT 7.5 is an overview of analytical ultracentrifugation that includes a discussion of methods for determining both the molecular weight and hydrodynamic properties of proteins.
Introduction
Light-scattering methods can also provide information about the native molecular weight, oligomeric composition, and overall conformation. Techniques using size-exclusion
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chromatography with on-line light scattering detectors are especially useful for characterizing proteins and glycoproteins (for review see Wen et al., 1996). UNIT 7.8 provides an overview of the various light-scattering methods used to characterize proteins and macromolecular complexes. It should be noted that this method, like analytical centrifugation, provides direct molecular weight information without the use of calibration standards or models. Circular dichroism and fluorescence spectroscopy are popular methods for determining protein conformation. Basic protocols for determining the circular dichroic spectra of proteins are described in UNIT 7.6 and for determining the fluorescence spectra in UNIT 7.7. Circular dichroism and fluorescence spectroscopy are similar to most other spectroscopic methods in that careful interpretation of the data is required together with an understanding of the limitations and pitfalls of the method. These points are emphasized throughout these units. Differential scanning calorimetry (DSC), described in UNIT 7.9, is an instrumental method used to measure heat energy uptake (heat capacity) in proteins undergoing thermal denaturation. This is accomplished by varying the temperature of a dissolved protein sample and measuring the difference in heat generated by the protein compared to a reference cell that contains solvent alone. This heat difference is related to the conformational energy of the protein molecule. In practice, when proteins undergo thermal denaturation, typically at 40° to 80°C, there is usually heat generated (i.e., it is an exothermic process); this heat can be very accurately measured by modern DSC instruments. The unit discusses the theoretical background of the method and details methods for actually performing DSC. For background on basic thermodynamic principles consult, for example, Tinco et al. (1995), and for overview of DSC applications in biotechnology see Chowdhry and Cole (1989). The Internet sites of the various instrument manufacturers are also useful sources of information. Gel filtration is a technically simple method that is nonetheless one of the most powerful available for characterizing recombinant proteins. Working in an analytical mode using modern automated equipment, information on protein purity, conformation, and physical homogeneity can be obtained within a very short time. When the analysis is combined with mass spectroscopy, the gel filtration chromatogram can usually be fully interpreted to determine the chemical identity of all the peaks. UNIT 7.10 details the complementary use of HPLC gel filtration and mass spectroscopy to analyze recombinant proteins. The determination of protein structures by multidimensional NMR (UNIT 17.5) requires soluble and folded protein usually produced by expression in E. coli. In UNIT 7.11 the rapid screening by heteronuclear NMR of E. coli extracts containing 15N-labeled recombinant proteins is described. This approach allows a rapid assessment of the suitability of the protein for more detailed structural analysis without the need for purification. CONFORMATIONAL ANALYSIS OF RECOMBINANT-PRODUCED PROTEINS The assays for many protein functions, for example in the case of cytokines, can be time consuming and of relatively low precision. As proteins depend for their function on the correct 3D fold of the polypeptide chain, demonstration that the conformation of the recombinant protein is the same as that of the authentic protein should provide a more convenient means of characterizing the product. The methods described in UNITS 7.4, 7.6 & 7.7, although providing average structural information only, as compared with X-ray crystallography or NMR, are sensitive indicators of conformation and conformational change and, as such, can provide a highly specific “fingerprint” of a protein. Such
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conformational analysis will follow assurance of chemical authenticity and homogeneity, as described in UNITS 7.1-7.3 (and elsewhere in the book). Urea gradient gel electrophoresis (UNIT 7.4) is a hydrodynamic technique that monitors the degree of compactness of the protein. It allows unfolding and refolding to be studied, leading to characterization of the protein in terms of its stability and kinetics. Near-UV circular dichroism (CD) results in spectra that are characteristic of the polar asymmetry of the environments of the aromatic residues, mainly tryptophan and tyrosine. Far-UV CD monitors the polypeptide backbone conformation, and hence, the secondary structure. Fluorescence spectra are determined chiefly by the polarity of the environment of the tryptophan and tyrosine residues and their interactions.The four techniques thus provide different, but complementary, fingerprints of the protein conformation that provide a high degree of assurance of nativeness within a few hours. For quality assurance of the native conformation and for general characterization, it is important to employ as broad a range of the available techniques as possible. Besides confirming the nativeness of the folded conformation of a sample of a protein that has already been characterized, either as the previously produced recombinant protein or as the authentic protein, the techniques described here offer a rapid means of obtaining structural and dynamic information, together with estimates of the stability of the protein. LITERATURE CITED Chowdhry, B.Z. and Cole, S.C. 1989. Differential scanning calorimetry: Applications in biotechnology. Trends. Biotechnol. 7:11-18. Tinco, I., Sauer, K., and Wang, J.C. 1995. Physical Chemistry: Principles and Applications in Biological Sciences (3rd ed.). Prentice Hall, Upper Saddle River, N.J. Wen, J., Arakawa, T. and Philo, J.S. 1996. Size-exclusion chromatography with on-line light scattering, absorbance, and refractive index detectors for studying proteins and their interactions. Anal. Biochem. 240:155-166.
Paul T. Wingfield and Roger H. Pain
Introduction
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Overview of the Characterization of Recombinant Proteins With the advent of recombinant DNA technology and the availability of numerous expression systems, scientists are now able to prepare large amounts of proteins in relatively short periods of time. Many of the proteins produced by this technology are new and previously uncharacterized. Current developments in protein purification techniques, especially the use of affinity matrices, can eliminate a number of less-specific purification steps, accelerating this process. It is still challenging, however, to purify a protein to complete homogeneity, removing all proteinaceous and nonproteinaceous contaminants. How clean a protein product needs to be largely depends on its final use. Some applications require exhaustive testing to ensure that no foreign material accompanies the protein. This is extremely important, for example, when the protein is to be used therapeutically, because even a minor contaminant can lead to side reactions that may be fatal. For such applications, it is therefore critical to evaluate the recombinant protein not only for its physical impurities, but also for its potential to cause immunogenic or allergic responses (Bristow, 1989). Strict guidelines for validating the purity of proteins for therapeutic use are in place at the U.S. Food and Drug Administration (Geisow, 1991; Garnick et al., 1991). Such careful characterization may not be required for proteins used in research, where a slight impurity may not affect the activity of a protein. It is known, however, that failure of crystallization is often due to microheterogeneity of the protein. The most troublesome impurities result from deamidation of Asn and Gln, Cys oxidation, proteolysis, sequence polymorphisms, and variability in post-translational modifications (Wood, 1989). A number of different expression systems are available for producing recombinant proteins, including bacteria (most commonly E. coli), yeast, insect cells, and mammalian cells infected with retroviruses (Chapter 5). Bacterial expression systems are generally the easiest to handle, and the first systems tried. Some proteins, however, require specific post-translational modifications (e.g., phosphorylation, glycosylation, and/or fatty acid acylation) for optimal activity, thus forcing the use of the other (eukaryotic) expression systems. Each expression system has its own set of advantages
and disadvantages (Chapter 5), and each has its own set of likely contaminants. A comprehensive set of analytical techniques is available to ascertain the purity of the protein products.
COMMON SOURCES OF CONTAMINATION Contaminating Proteins It is critical to check that the sample is free of other contaminating proteins. The hardest to detect and eliminate are those of similar molecular weight, pI, and hydrophobicity. The best way to analyze for such impurities is by HPLC, using at least two complementary chromatographic separations (e.g., reversed-phase chromatography on a C4 column and ion-exchange chromatography, or the use of two types of ion-pairing agents in reversed phase) with the aid of a diode-array detector. Another powerful method is the use of high-resolution peptide mapping by HPLC, alone or in combination with electrospray mass spectrometry (Scoble and Martin, 1990; Carr et al., 1991) or matrix-assisted laser-desorption ionization time-of-flight (MALDI TOF) mass spectrometry (see UNIT 11.6). If authentic pure protein is available for comparison, proteolytic digestion of both the authentic and the recombinant protein under the same conditions can be used to assess purity of the recombinant protein. Both proteins should generate the same fragmentation pattern; differences may be interpreted as being due to contaminants or as indicating chemical heterogeneity. Electrospray and MALDI TOF mass spectrometry eliminate the requirement for pure authentic protein, because the mass assignments for each proteolytic fragment of the recombinant protein are accurate enough on their own to indicate if the fragments are all derived from the parent protein. Any fragments that do not fit may belong to a contaminant or indicate incorrect primary sequence.
Proteolytically Clipped Proteins Many proteins become proteolytically cleaved during the purification process, even in the presence of protease inhibitor cocktails, due to the ubiquitous distribution of proteolytic agents in cell lysates. In some circumstances it may be difficult to separate the fragments from the product of interest, especially when the protein is nibbled from one end or the other by
Contributed by Nancy D. Denslow, Paul T. Wingfield, and Keith Rose Current Protocols in Protein Science (1995) 7.1.1-7.1.13 Copyright © 2000 by John Wiley & Sons, Inc.
UNIT 7.1
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7.1.1 CPPS
exoproteases, as these products will be almost the same size and have nearly identical physical properties. This task becomes especially difficult when the level of contamination is below normal detection levels. For proteins used therapeutically, this problem may have major ramifications, because it is well known that degraded proteins, or those that are not folded correctly, may be more antigenic than their intact counterparts, causing the patient to produce antibodies to the protein itself (Walsh and Headon, 1994). Determination of the N- and C-terminal sequences may be required to ensure purity from this type of contamination.
Nucleic Acid Host DNA may accompany the purified protein. The main concern with this contaminant is the possibility that an oncogene from the host may be present in the sample. DNA is generally efficiently removed from samples by chromatography through anion exchangers such as DEAE. Spectroscopic measurements at 260 nm will indicate the presence of nucleic acids.
Pyrogens Bacterial endotoxins and lipopolysaccharides can often accompany a protein through its purification (Bristow, 1989). These agents, if present even in small amounts, can produce fever in patients. Thus, for proteins used therapeutically, it is important to prove that such contaminants are not present in the sample. Apart from chromatography by HPLC, sensitive immunological methods also exist to test for the most common of these contaminants. For example, bacterial proteins can be detected by using antibodies against total E. coli cell lysates in immunoblots of the recombinant protein of interest. If the presence of lipopolysaccharides is suspected, they usually can be removed by gel filtration or by using membranes of the appropriate molecular weight cutoff, as these contaminants are very hydrophobic and tend to aggregate. For further details on the detection and analysis of nonproteinaceous contaminants, see Walsh and Headon (1994).
CHEMICAL IDENTITY
Overview of the Characterization of Recombinant Proteins
Checking Primary Structure of Recombinant Proteins Once the purity of the recombinant protein has been ascertained, it is imperative to prove that the isolated protein indeed has the correct
amino acid sequence. There are a number of analytical techniques that can be used to characterize the protein, each providing different but overlapping answers. Some of the techniques are more exacting than others. They all, however, generally require expensive instrumentation, such as HPLC machines or mass spectrometers, that may not be available in all laboratories. Other techniques are less precise in their measurements, but provide reasonable data if authentic protein is available as a control. The first test is to determine the mass of the recombinant protein, especially as it compares to the authentic protein. This can be accomplished in most laboratories by SDS-PAGE. Although the mass determined in this manner is not very accurate, it can be compared to the control to determine whether the protein is close to the right size. A more precise measure of the mass can be obtained by mass spectrometry, either electrospray or MALDI TOF. These measurements are generally within 0.01% to 0.05% of the correct mass. Next, it is useful to determine the isoelectric point (pI) of the recombinant protein. This can be accomplished by isoelectric focusing (IEF) gel electrophoresis (UNIT 10.2) using flat-bed agarose or acrylamide gels or slab gels. The pI is determined from a standard curve constructed using IEF standards (proteins of known pI). It is good practice to include the native protein, if available, in an adjacent lane. Differences in pI from the expected value are diagnostic for differences in charged amino acids. A small sample of the protein (1 to 5 µg) should be analyzed for its amino acid composition. This can usually be done in a protein chemistry core facility, available at most universities, for a small fee. This analysis should provide confirmation that all the expected amino acids are present in the protein. A second sample (25 to 50 pmol) should be submitted for N-terminal amino acid sequencing. This sample can be provided on a PVDF membrane (electroblotted from an SDS gel), or in solution (provided the sample is free of salts). This analysis should verify that the protein starts with the expected sequence, an important consideration if the N-terminal Met is cleaved after translation. A blocked N-terminus, such as occurs when the terminal amino acid is post-translationally modified, requires special procedures to be identified (LeGendre et al., 1993). High-resolution peptide maps (fingerprinting) provide information on internal sequences
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by comparing digests of the recombinant and the authentic protein, both treated under the same conditions. The best diagnostic fingerprints are analyzed by HPLC, where the retention times of over a hundred different fragments can often be compared. A difference in amino acid sequence between the recombinant protein and the authentic protein is highly likely to result in a change in the mobility of at least one fragment. If an HPLC unit is not available, one can get relatively good information from analysis of digested proteins by high-resolution SDS-PAGE. In that case one should use multiple partial digests to compare fragments that are ≥2 kDa in size. The Tris-tricine gel system (UNIT 10.1) is recommended for this type of analysis. New, powerful methods are being developed that combine peptide mapping with mass spectrometry (Pappin et al., 1993). In these procedures, the mass of each fragment is obtained with high accuracy, allowing direct comparison with the predicted sequence from the cDNA. Thus, it is not necessary to include the authentic protein in the analysis, unless post-translational modifications are expected.
Characterizing Post-Translational Modifications It is important to determine the nature and location of all post-translational modifications of recombinant proteins. Among the most problematic are unwanted, nonspecific modifications, including oxidation of Met residues and chance deamidation of Asn or Gln. These modifications can easily be detected as retention time shifts when HPLC profiles of peptide maps of the recombinant protein are compared to those of the authentic protein. The presence of reducing agents, such as 2-mercaptoethanol or dithiothreitol, in the isolation buffer often prevents oxidation of Met. There are situations in which one wants to determine if the correct post-translational modification has been made in the recombinant protein, especially if the modification is important for the activity of the protein. Usually, the expression system has been carefully selected to provide the appropriate modification. Among the most frequent post-translational modifications are phosphorylation, N-methylation, acetylation, glycosylation, farnesylation, and sialic acid capping of oligosaccharides. Pinpointing these modifications can be time-consuming, requiring specific methods for each modification if traditional approaches are to be used (see Chapters 12 and 13 for
protocols for determining sites of glycosylation and phosphorylation). The usual method is to digest the recombinant protein, separate the fragments by HPLC, then pick fragments suspected of being modified for further analysis by N-terminal sequencing using Edman chemistry. A major disadvantage to this approach is that some of the modifications are not stable when exposed to the chemicals used in sequencing, and will therefore be missed. For example, phosphorylated Ser and Thr residues are not easily detected because they undergo β-elimination of the phosphoryl group (Mercier et al., 1971). Instead, a system using radioactive tracers has been developed in which the assignment of the phosphorylation sites is based on the release of [32P]Pi during Edman degradation (Wettenhall and Cohen, 1982). Newer approaches are being developed that rely on mass spectrometry to pinpoint the location of the modifications (Wang et al., 1994). These approaches offer direct procedures for identifying the modified residues and should cut the time required for analysis.
PHYSICAL AND CONFORMATIONAL CHARACTERIZATION Once the chemical identity of a protein has been established, its physical and conformational properties are probed using a variety of biophysical techniques. Unlike chemical analysis, most of these techniques require that the protein be pure, as otherwise the interpretation of the data can be difficult or impossible. Some of the basic questions that are addressed are: (1) Is the protein monodisperse? (2) What is the quaternary (subunit) structure? (3) Is the protein folded? (4) What are the secondary and tertiary structures of the protein? The commonly used methods for studying protein conformation and structure are listed in Table 7.1.1 (nonspectroscopic) and Table 7.1.2 (spectroscopic). References have been included that emphasize the practical aspects of the various techniques. Because of the complexity of the subject matter, several outstanding textbooks that cover most of the theoretical background have also been cited (see Key References).
Physical Appearance and Absorbance It is always useful to visually inspect purified protein placed in a glass or quartz cuvette; it may be obvious on holding the cuvette up to the light that the protein is colored, cloudy, or perhaps slightly opalescent. For recombinant
Characterization of RecombinantProduced Proteins
7.1.3 Current Protocols in Protein Science
Nonspectroscopic Methods for Studying Protein Structure and Conformation
Table 7.1.1
Method
Information obtained
Notes
Urea gradient electrophoresis
Size, shape, and unfolding transitions
Amount of material required depends on size of Goldenberg (1989) gel and detection system. If antibodies are used, crude samples can be analyzed.
Gel filtration
Size, shape, and molecular interactions
Most commonly used method for determining molecular radii. Mr can be determined if sedimentation coefficient is known. Conformation changes can be followed.
Ultracentrifugation
Reference(s)
Ackers (1970); Ptitsyn et al. (1990)
Various sedimentation velocity measurements used to gain information on shape/conformation.
Sedimentation velocity
Shape
Useful check of homogeneity. Determination of Van Holde (1975); sedimentation coefficient (S). S is sensitive to Harding (1994); protein conformation. Small differences in S Ralston (1993) between samples may be difficult to detect.
Difference sedimentation
Shape
The measurement of very small differences in Van Holde (1975) sedimentation coefficient (∼0.01 S). Sensitive to very small conformation differences between samples.
Sedimentation equilibrium
Molecular weight, quaternary structure, and molecular interactions
The method of choice for direct determination of the molecular weight (precision ±3%). Also widely used for studying reversible self-association of proteins and ligand binding.
Van Holde (1975); Harding (1994); Ralston (1993); McRorie and Voelker (1993)
Conformation
Protease action on folded proteins depends on both the cleavage-site specificity and its accessibility. In unfolded proteins only the former is important.
Price and Johnson (1989)
Similar in principal to above except using chemical reagents specific for various amino acid side chains (e.g., sulfhydryl reagents).
Ballery et al. (1993)
Accessibility assays Protease sensitivity
Chemical reactivity Conformation
2H
and 3H-exchange Conformation/dynamics HPLC and NMR used to measure exchange reactions.
Englander and Kallenbach (1984)
Calorimetry Differential scanning Conformation; thermal stability
Sturtevant (1987); Measurement of heat uptake (heat capacity) during change in temperature. Used for study of Cooper and thermal unfolding of proteins and nucleic acids. Johnson (1994) In modern instruments sample requirements are modest: 1-2 ml at 0.5-2.0 mg/ml per scan. Proteins often irreversibly denature at neutral pH; this is sometimes circumvented by analysis at acid pH or by addition of low concentrations of chaotropes.
Isothermal titration
Direct measurement of the energetics of bimolecular interactions (including binding constants). Larger samples needed than for differential scanning.
Overview of the Characterization of Recombinant Proteins
Molecular interactions
Cooper and Johnson (1994)
continued
7.1.4 Current Protocols in Protein Science
Table 7.1.1
Nonspectroscopic Methods for Studying Protein Structure and Conformation, continued
Bimolecular interaction analysis
Molecular interactions
Bimolecular interaction analysis using the Pharmacia BIAcore instrument. Uses surface plasmon resonance to detect bimolecular interactions. Examples: protein ligand binding to receptor, protein binding to nucleic acid, and antibody-antigen interactions. Wet chemistry required to immobilize one of the reaction partners on a matrix.
Chaiken et al (1993); Cunningham and Wells (1993)
Immunological assays
Conformation
Affinity of an antibody for its antigen depends on the structural complementarity between the molecules. Changes in the antigen conformation result in changes in the binding affinity for appropriate antibodies. Monoclonal antibodies are usually used.
Figuet et al. (1989)
proteins isolated from E. coli, all of these observations are usually bad news. Running an absorption spectrum from 240 nm (in the nearUV) to 600 nm (in the visible region) will enable one to decide if the protein is partially aggregated or is contaminated with colored chromophores. The absorbance of a ∼1 mg/ml protein solution (using a 1-cm path length cell) in the region of 400 and 320 nm should be <0.05; this is a spectral region where pure protein does not absorb, and high absorbance (>0.1) is usually due to light scattering from aggregated protein. It should be noted that light scattering from macroscopic particles increases with decreasing wavelength; hence, the detection of scattering artifacts is aided by measuring an absorption spectrum rather than single wavelengths. Filtration with a 0.2-µm-poresize filter, or low-speed centrifugation, often removes the material responsible for light scattering. A clearer picture of what proportion of the sample is aggregated requires further analysis by gel-filtration (see discussion of Molecular Size and Subunit Structure) or analytical ultracentrifugation. The main absorption peak should be at ∼280 nm and the A280/A260 ratio should be ∼1.5 to 2.0. If there is high absorbance at 260 nm, it may be due to nucleic acid or other chemical contaminants.
Molecular Size and Subunit Structure The next step is to apply the sample to a gel-filtration column to see if the protein elutes in a single symmetrical peak as expected for a monodisperse protein (UNIT 8.3). The appearance of material eluting at the void volume of the column indicates highly aggregated protein. If the column is calibrated with molecular size
markers, an approximate idea of the size can be obtained. It should be remembered that to get an accurate molecular weight, it is necessary to combine the gel-filtration data with a knowledge of the sedimentation coefficient, which must be obtained either by analytical ultracentrifugation or density gradient analysis (UNIT 4.2). If the molecular weight of the native protein is known, the subunit composition can be determined, as the monomeric mass is known from the amino acid sequence translated from the DNA coding sequence. The only commonly used methods capable of directly determining molecular weight are sedimentation equilibrium and light scattering (osmotic pressure measurement also gives this information but is rarely used nowadays). Determination of the molecular weight of recombinant glycoproteins by gel filtration is especially unreliable. If the amount of carbohydrate associated with the protein has been determined (e.g., by mass spectrometry), the molecular weight can be directly determined by sedimentation equilibrium. With ultracentrifugation, the association state of a glycoprotein (e.g., whether it is a monomer or dimer) can be determined even when the carbohydrate composition is unknown (Pennica et al., 1992; Shire, 1992). Apart from some of the methods referred to above, a more modest approach, equipmentwise, to molecular weight determination is the use of native gel electrophoresis (UNIT 10.3).
Conformation and Structure It is important to assess if the purified recombinant protein has been folded correctly,
Characterization of RecombinantProduced Proteins
7.1.5 Current Protocols in Protein Science
Table 7.1.2
Spectroscopic Methods for Studying Protein Structure and Conformation
Method
Information obtained
Notes
Reference(s)
Method available to most laboratories. Modern instruments have diode array detectors and are computer controlled.
Wetlaufer (1962)
Ultraviolet absorption Standard spectroscopy
Secondary and tertiary structure
Difference spectroscopy
Conformational change Spectral differences between perturbed (e.g., using pH, temperature, and denaturants) and nonperturbed samples give information on relative conformation.
Fluorescence
Donovan (1969, 1973)
Very sensitive; small samples needed compared to other spectroscopic methods.
Steady-state
Secondary and tertiary structure
Intrinsic fluorescence mainly due to tryptophan. Proteins can be labeled with fluorophores (extrinsic fluorescence). Steady-state quenching studies give information on exposure of chromophores.
Time-resolved
Conformational change Trp fluorescence lifetimes (decay time) lie in the nanosecond region. Molecular dynamics can be studied by time-resolved quenching and depolarization.
Circular dichroism
Conformational change One of the most commonly used techniques to compare overall (or gross) conformational properties of proteins.
Chen et al. (1969); Haugland (1994)
Eftink and Ghiron (1981); Varley (1994)
Far-UV
Secondary structure
Measurements in the range 240-180 nm. α-helix most accurately determined. Computer-aided analysis of spectra by various methods to estimate secondary structure contents.
Yang et al. (1986); Johnson (1990)
Near-UV
Tertiary structure
Measurements in the range 340-240 nm. Depends on environment of aromatic residues. Conformational fingerprint. Interpretation not easy, but useful for comparative purposes.
Strickland (1974)
Secondary structure
Raman scattering (and infrared absorption) known as vibrational spectroscopy. Useful for large macromolecular assemblies, precipitates, and crystals. Secondary structure estimates using Raman Amide I and III spectra.
Bussian and Sander (1989); Williams (1986)
Infrared absorption Secondary structure
Modern computerized Fourier transform (FT-IR) instrumentation used; allow measurements in H2O (particulate samples can also be measured). Secondary structure from Amide I spectra. Better than circular dichroism for β-structure estimation.
Haris and Chapman (1992, 1994); Surewicz et al. (1993)
Light scattering
Direct Mr determination (cf. sedimentation equilibrium). Especially useful for large macromolecular assemblies.
Raman scattering
continued Overview of the Characterization of Recombinant Proteins
7.1.6 Current Protocols in Protein Science
Table 7.1.2
Spectroscopic Methods for Studying Protein Structure and Conformation, continued
Static light scattering
Molecular weight and shape
Classical light scattering. Modern instruments use lasers. Samples must be dust-free. Determination of Mr requires that the refractive index increment (dn/dc) be known. Mr distributions and conformational information can be obtained by combining gel filtration with an in-line multi-angle light scattering detector.
Harding (1994)
Dynamic quasi-elastic
Shape and molecular weight
Also known as photon correlation spectroscopy. The translation diffusion coefficient (D) is measured. Knowledge of the sedimentation coefficient (S) is required to determine Mr.
Harding (1994); Phillies (1990)
Nuclear magnetic resonance 1H
Secondary structure; Large sample requirements. Proteins must be 3-D structure of small very pure and stable. proteins and polypeptides (<10 kDa)
Clore and Gronenborn (1992)
Multidimensional
Secondary structure; 3-D structure of proteins 10-30 kDa
Clore and Gronenborn (1994)
Proteins uniformly labeled with combinations of 15N, 13C, and 2H by expression in E. coli using minimal medium (see UNIT 5.3). 2-D 1H-15N correlation spectra are very useful as protein conformational fingerprints.
X-Ray diffraction
3-D structure at atomic resolution
Scanning Molecular weight transmission electron microscopy
especially if it was prepared from inclusion bodies (insoluble aggregates of the overexpressed protein), which are often found in bacterial expression systems (UNITS 6.3-6.5). Spectroscopic methods and calorimetry are the usual techniques used to monitor or measure protein conformation. Urea-gradient electrophoresis is also used and has the advantage that pure protein is not required. It is advantageous—as in mapping the chemical identity of the protein— to have some type of standard available for comparative purposes. The power of the spectroscopic methods, especially circular dichroism (CD) and fluorescence, lies in their ability to detect differences rather than to make absolute determinations. The standard may be the natural or authentic protein or may be the same recombinant protein separately prepared and analyzed by independent methodologies.
Proteins must be crystallized. Commercially available Crystal screen kit widely used.
McRee (1993); Ducruix and Giege (1992); Hampton Research
Scanning transmission electron microscopy; mass determination of individual molecules. Useful for large particles beyond the mass range of sedimentation equilibrium.
Thomas et al. (1994)
Apart from the use of protein standards, model compounds are often used as conformation standards (especially for fluorescence and UV spectroscopy). These may be the free amino acids tryptophan and tyrosine: the model protein is constructed using the appropriate molar ratio of tryptophan and tyrosine and represents a fully denatured protein in which the aromatic residues are fully exposed to the solvent. Alternatively, the recombinant protein can be denatured with 4 to 8 M guanidine⋅HCl (see APPENDIX 3A). Comparison of either simulated or denatured protein with the test protein will give some indication of whether the protein is folded. The most commonly used spectroscopic methods are CD and fluorescence. Nuclear magnetic resonance (NMR) spectroscopy measurements are especially useful when the re-
Characterization of RecombinantProduced Proteins
7.1.7 Current Protocols in Protein Science
Overview of the Characterization of Recombinant Proteins
combinant protein is uniformly labeled with 15N; for example, 1H-15N correlation spectra are excellent protein conformational fingerprints. (A conformational fingerprint is related to the overall tertiary structure as a classical two-dimensional peptide map is related to the primary sequence.) CD spectra in the far-UV region (240 to 180 nm) can be used to estimate secondary structure, and spectra in the near-UV region (350 to 240 nm) give information on the tertiary structure. When interpreting CD spectra, it should be borne in mind that a protein can have an apparently normal far-UV CD spectrum, yet have a near-UV spectrum characteristic of an unfolded protein. This is characteristic of partially folded protein or so-called molten globules (UNIT 6.4). Apart from the analysis of recombinant proteins that contain the native sequence, mutant proteins are produced for various structurefunction studies. It is almost mandatory for some conformational analyses to be made on these proteins so that functional differences can be related to specific chemical differences (i.e., different amino acid side chains) rather than merely being explained as due to conformational perturbation. For an example of a spectral-based analysis of site-directed mutants, see Mulkerrin and Cunningham (1993). The folding question is also important for proteins prepared by other expression systems (e.g., yeast and baculovirus). Although these systems offer the advantage of providing proteins with the correct post-translational modifications and thus by supposition the correct structure, it is critical to demonstrate that the recombinant product has the same secondary and tertiary structures as the authentic protein. Incorrect disulfide bonding is sometimes encountered in these systems and will introduce conformational heterogeneity. When analyzing protein conformation, it is a mistake to depend on any one physicochemical technique, especially the spectroscopic methods. The more corroborative data that is accumulated using different techniques and approaches, the more reliance can be placed on any conclusions. For spectroscopic analysis, the ideal sample is protein freshly purified by all titration. The peak fractions of a symmetrically shaped elution peak are tested using the column elution buffer as reference material. If rapid buffer exchange is required, for example to remove excess salt prior to far-UV CD measurements, the sample (0.1 to 1.0 ml) can be applied to a prepacked column (1.5-cm diameter × 5 cm) of
Sephadex G-25 (PD-10 columns from Pharmacia Biotech) and eluted in <5 min.
High-Resolution Structure The ultimate test of a protein’s structural integrity and conformational homogeneity is determination of its three-dimensional structure. For X-ray structure determination, crystals must be obtained that diffract to high resolution (<3 Å). The fact that the protein can be crystallized is in itself a fairly good indication that the preparation is both physically and chemically homogeneous. For reviews of the methodologies involved in protein crystallization and X-ray diffraction, see McRee (1993) and Ducruix and Giege (1992). Structural determination of proteins in solution can be achieved by NMR spectroscopy. For small proteins (<10 kDa), conventional homonuclear two-dimensional methods have been used. For larger proteins, three- and fourdimensional heteronuclear methods are required to resolve the spectral complexity due to the larger numbers of protons. For this purpose, proteins expressed in E. coli are uniformly labeled with 15N and/or 13C as mentioned in UNIT 5.3. Structures of medium-sized proteins (∼15 to 20 kDa) can be determined at high resolution using the current methodologies. For further details see references listed in Table 7.1.1 and Table 7.1.2.
Biological Activity and Immunogenicity The biological activity of the isolated protein should be carefully studied. When possible, the activity should be compared to that of the authentic protein. For an apparently pure protein, a lower activity is often a clue that the protein has not folded correctly, is partially aggregated, or has been improperly post-translationally modified. A low specific activity may also indicate the presence of other proteinaceous contaminants that have been undetected by the analytical methods used. It is of course essential to accurately determine protein concentrations (Chapter 3) in order to make realistic comparisons between samples. The main problem with all types of activity measurement is to know what proportion of the preparation is active. For an enzyme, active-site titration can give this information (Fersht, 1985). However, a small proportion (<5%) of any preparation may be inactive due to denaturation upon storage and standing, and this would not be detected by most of the spectroscopic methods referred to above. The presence
7.1.8 Current Protocols in Protein Science
of small amounts of misfolded protein may normally be of no consequence, the main requirement being that the protein have similar biological activity to the standard. (The standard, especially if it is an authentic protein, will probably be even less well characterized than the recombinant protein). If, on the other hand, the protein is to be used for in vivo studies, misfolded protein may be immunogenic and, thus, must be considered as a contaminant even though it may be chemically indistinguishable from the correctly folded protein. This is of special concern for therapeutic proteins, especially those repeatedly administered (for review, see Walsh and Headon, 1994). The ability of antibodies to detect subtle conformational change has long been known (Goldberg, 1991). Conformational changes in local regions of the protein can be measured by determining changes in affinity. Furthermore, conformational differences between samples can be detected where spectroscopic and, more importantly, activity measurements, do not show a difference (Hattori et al., 1993).
BASIC PLAN OF ACTION The task tree presented in Figure 7.1.1 is a suggested path for fully characterizing recombinant proteins. Some of the analytical techniques overlap in the type of information they provide. In order to avoid running multiple tests, we recommend using the high-technology methods, such as HPLC, mass spectrometry, circular dichroism, NMR, and fluorescence spectroscopy. However, if these are not available, good information can still be extracted from the more traditional approaches, keeping in mind that they are less resolving and therefore require complementary analytical strategies.
ANALYSIS OF N-TERMINAL HETEROGENEITY OF RECOMBINANT PROTEINS PRODUCED IN E. COLI When a purified recombinant protein presents as a single band on an SDS-PAGE gel, gives a satisfactory amino acid analysis, gives the expected N-terminal amino acid sequence upon Edman degradation, shows the correct
PROTEIN Determination of Purity HPLC analysis with diode array detector
SDS-PAGE
isoelectric focusing
peptide mapping mass spectrometry
Chemical Characterization N-terminal amino acid sequencing analysis
peptide mapping
internal sequencing
characterization of posttranslational modifications
Conformational Analysis circular nuclear magnetic dichroism resonance spectroscopy
fluorescence spectroscopy
calorimetry
Analysis of Quaternary Structure
gel filtration
native gel electrophoresis
analytical centrifugation
Activity Assays
Figure 7.1.1 Task tree graphically representing the five steps (branches) required to fully characterize a recombinant protein. The first step is to determine the purity of the protein, which can be done by a variety of methods; the choice will depend on the availability of instrumentation and the degree of purity desired. This is followed by chemical characterization of the protein. Specific methods for the third and fourth steps, determination of protein conformation (folding) and structure, are listed in in Table 7.1.1 and Table 7.1.2. The last step, determining the activity of the protein, will require unique assays developed specifically for the protein of interest.
Characterization of RecombinantProduced Proteins
7.1.9 Current Protocols in Protein Science
disulfide bond linkages, and is biologically active, most people are quite satisfied. However, insufficient resolution on SDS-PAGE, imprecise amino acid analysis (4% at best) and bioassays, and nonquantitative yield (particularly on the first cycle) during Edman degradation may all combine to mask protein heterogeneity. High-resolution, charge-sensitive techniques, such as two-dimensional gel analysis involving isoelectric focusing (UNIT 10.4) and capillary electrophoresis, may reveal quite extensive heterogeneity in proteins that seem pure by the criteria described above. This heterogeneity may be due to deamidation of Asn or Gln residues—some of which, because of their position in the polypeptide chain, may be particularly susceptible to hydrolysis of their side chain amide—but other explanations are possible. For example, the carboxy terminus may be “ragged” (i.e., longer or shorter polypeptide chains may be present, with not all molecules terminating at the same residue). A ragged N-terminus is normally detected by Edman degradation, but other forms of post-translational modification at the N-terminus can go unnoticed because the amino group is blocked and so not accessible to the Edman chemistry. Various kinds of post-translational modification can create heterogeneity (when they do not affect all the protein molecules in question), and some can affect all of the molecules in such a way that the product is homogenous but is not the expected protein product.
Importance of Mass Spectrometry
Overview of the Characterization of Recombinant Proteins
Mass spectrometry is a very useful technique for identifying all post-translational modifications, not merely those occurring at the N-terminus (Wang and Chait, 1994; Bradshaw and Stewart, 1994). The molecular weight of the recombinant protein can be obtained with high accuracy and precision (better than 0.01%) and minor impurities identified. This performance is normally quite sufficient to detect modifications such as missing residues or additional groups (see discussion of Two Instructive Examples). However, it is not usually sufficient to identify deamidation (1 mass unit change) of a protein larger than 10 kDa. Particular sites of deamidation are best located by peptide mapping, and Staphylococcus aureus V8 endoprotease Glu-C and Pseudomonas fragi endoprotease Asp-N are useful in this regard because they can cleave at the sites of Glu and Asp residues, respectively (UNIT 11.1). If deamidation has occurred to a small extent at many sites, the
problem of locating such sites becomes much more difficult. Ideally, a protein should be purified to a single species prior to characterization, but this is not always possible. Once it is known that a portion of the sample has different charge (by IEF) or mass (by mass spectrometry), attempts should made to identify the site and nature of the modification by peptide mapping (UNIT 11.6). As is evident from the two examples that follow, and from UNIT 11.6, mass spectrometry has an essential role to play in the characterization of recombinant proteins.
Two Instructive Examples A simple but instructive example is that provided by interleukin 5 (IL-5). Recombinant human IL-5 expressed in E. coli is a single band on SDS-PAGE but shows several bands on IEF (Rose et al., 1992a). Electrospray ionization mass spectrometry showed a major component with the expected molecular weight (observed, 13282.5 ± 2; calculated, 13280.4), but also showed evidence of the presence of species of slightly greater molecular weight. Peptide mapping led to isolation of four forms of the N-terminal peptide—the expected (methionyl) form, a formyl-methionyl form, a methionyl-sulfoxide form, and a carbamoyl-methionyl form— all of which were characterized by mass spectrometry. The dimeric nature of the native protein added to the complexity of the IEF results. The formyl and carbamoyl forms were not accessible to Edman degradation, and the yield on the first cycle (∼50%) was in the acceptable range for the technique. Amino acid analysis did not help to detect the presence of these two forms, as both modifying groups (formyl and carbamoyl) are hydrolyzed off during hydrolysis. A second instructive example concerns the recombinant DNA-binding protein Ner expressed in E. coli (Rose et al., 1992b). This small protein was purified and characterized by conventional techniques (Allet et al., 1988). Every residue was identified by Edman degradation except the N-terminal Cys residue (Cys was not derivatized, so a blank degradation cycle was not unexpected). Nevertheless, the expected one Cys residue was detected by amino acid analysis after oxidation with performic acid. It was only when mass spectrometry showed that the protein was too heavy by 70 mass units that a problem was recognized. After much effort, involving peptide mapping, mass spectrometry, microchemistry, and twodimensional NMR spectroscopy, it turned out
7.1.10 Current Protocols in Protein Science
that the N-terminal residue was modified with pyruvate, but in a way not involving the carboxyl group of pyruvate (Rose et al., 1992b). This modification was unstable in mild acid and in mild base (although stable at physiological pH), which explained why it had been missed when the protein was analyzed by Edman degradation and amino acid analysis. Thus, although the protein was blocked (N-terminally modified), the modifying group became detached during the first coupling step of the Edman chemistry and so was invisible to the technique.
LITERATURE CITED Ackers, G.K. 1970. Analytical gel chromatography of proteins. Adv. Protein Chem. 24:342-443. Allet, B., Payton, M., Mattaliano, R.J., Gronenborn, A.M., Clore, G.M., and Wingfield. P.T. 1988. Purification and characterization of the DNAbinding protein ner of bacteriophage mu. Gene 65:259-268. Ballery, N., Desmadril, M., Minard, P., and Yon, J.M. 1993. Characterization of an intermediate in the folding pathway of phosphoglycerate kinase: Chemical reactivity of genetically introduced cysteinyl residues during the folding process. Biochemistry 32:708-714. Bradshaw, R.A. and Stewart, A.E. 1994. Analysis of proteins modifications: Recent advances in detection, characterization and mapping. Measurement as a tool for studying proteins. Curr. Opin. Biotechnol. 5:85-93. Bristow, A.F. 1989. Purification of proteins for therapeutic use. In Protein Purification Applications: A Practical Approach (E.L.V. Harris and S. Angal, eds.) pp. 29-44. IRL Press, Oxford. Bussian, B.M. and Sander, C. 1989. How to determine protein secondary structure in solution by Raman spectroscopy: Practical guide and test case DNase I. Biochemistry 28:4271-4277. Carr, S.A., Hemling, M.E., Bean, M.F., and Roberts, G.D. 1991. Integration of mass spectrometry in analytical biotechnology. Anal. Chem. 63:28022824. Chaiken, I., Rose, S., and Karlsson, R. 1992. Analysis of macromolecular interactions using immobilized ligands. Anal. Biochem. 201:197-210. Chen, R.F., Endelhoch, H., and Steiner, R.F. 1969. Fluorescence of proteins. In Physical Principles and Techniques of Protein Chemistry (S.J. Leach, ed.) pp. 171-244. Academic Press, New York. Clore, G.M. and Gronenborn, A.M. 1992. Methods of structural analysis of proteins. Part 2—Nuclear magnetic resonance. In Protein Engineering: A Practical Approach (A.R. Rees, M.J.E. Sternberg, and R. Wetzel, eds.) pp. 33-56. IRL Press, Oxford.
Clore, G.M. and Gronenborn, A.M. 1994. Multidimensional heteronuclear nuclear magnetic resonance of proteins. Methods Enzymol. 239:349363. Cooper, A. and Johnson, C.M. 1994. Microcalorimetry; differential scanning calorimetry; and isothermal titration calorimetry. Methods Mol. Biol. 22:109-150. Cunningham, B.C. and Wells, J.A. 1993. Comparison of a structural and functional epitope. J. Mol. Biol. 234:554-563. Donovan, J.W. 1969. Ultraviolet absorption. In Physical Principals and Techniques of Protein Chemistry, Part A (S.J. Leach, ed.) pp. 101-170. Academic Press, New York. Donovan, J.W. 1973. Ultraviolet difference spectroscopy—new techniques and applications. Methods Enzymol. 27:497-548. Ducruix, A. and Giege, R. (eds.). 1992. Crystallization of Nucleic Acids and Proteins: A Practical Approach. IRL Press, Oxford. Eftink, M.R. and Ghiron, C.A. 1981. Fluorescence quenching studies with proteins. Anal. Biochem. 114:199-227. Englander, S.W. and Kallenbach, N.R. 1984. Hydrogen exchange and structural dynamics of proteins and nucleic acids. Q. Rev. Biophys. 16:521-655. FDA (Food and Drug Administration). FDA Handbook. Points to Consider in the Production and Testing of New Drugs and Biologicals Produced by Recombinant DNA Technology. Office of Biologics Research and Review, Center for Drugs and Biologics, Food and Drug Administration, Bethesda, Md. Fersht, A. 1985. Enzymes: Structure and Mechanism, 2nd Edition. Freeman, New York. Figuet, B., Djavadi-Ohaniance, L., and Goldberg, M.E. 1989. Immunological analysis of protein conformation. In Protein Structure: A Practical Approach (T.E. Creighton, ed.) pp. 287-310. IRL Press, Oxford. Garnick, R.L., Rose, M.J., and Baffi, R.A. 1991. Characterization of proteins from recombinant DNA manufacture. In Drug Biotechnology Regulation: Scientific Basis and Practices. (Y.-Y. H. Chiu and J.L. Gueriguian, eds.) pp. 263-313. Marcel Decker, New York. Geisow, M.J. 1991. Characterizing recombinant proteins. Bio/Technology 9:921-924. Goldberg, M.E. 1991. Investigating protein conformation, dynamics and folding with monoclonal antibodies. Trends Biochem. Sci. 16:358-362. Goldenberg, D.P. 1989. Analysis of protein conformation by gel electrophoresis. In Protein Structure: A Practical Approach (T.E. Creighton, ed.) pp. 225-250. IRL Press, Oxford. Hampton Research Macromolecular Crystallization Reagent Kits: Crystal Screen I and II. Hampton Research, Calif. Harding, S.E. 1994. Sedimentation velocity; sedimentation equilibrium. Methods Mol. Biol. 22:61-83.
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7.1.11 Current Protocols in Protein Science
Harding, S.E. 1994. Classical light scattering and dynamic light scattering. Methods Mol. Biol. 22:61-83. Haris, P.I. and Chapman, D. 1992. Does Fouriertransform infrared spectroscopy provide useful information on protein structures? Trends Biochem. Sci. 17:328-333. Haris, P.I. and Chapman, D. 1994. Analysis of polypeptide and protein structures using Fouriertransform infrared spectroscopy. Methods Mol. Biol. 22:183-202. Hattori, M., Ametani, A., Katakura, Y., Shimizu, M., and Kaminogawa, S. 1993. Unfolding/refolding studies on bovine β-lactoglobulin with monoclonal antibodies as probes. J. Biol. Chem. 268:22414-22419.
Ralston, G. 1993. Introduction to analytical ultracentrifugation. Vol. 1, Analytical Ultracentrifugation Series. Beckman Instruments, Fullerton, Calif. Rose, K., Anderegg, R., Wells, T.N., and Proudfoot, A.E. 1992a. Human interleukin-5 expressed in E. coli has N-terminal modifications. Biochem. J. 286:825-828.
Haugland, R.P. 1994. Handbook of Fluorescent Probes and Research Chemicals, 5th Ed. Molecular Probes, Eugene, Oreg.
Rose, K., Simona, M. G., Savoy, L.-A., Green, B. M., Gronenborn, A.M., Clore, G.M., and Wingfield, P.T. 1992b. Pyruvic acid is attached to the amino terminus of the recombinant-DNA derived DNA-binding protein ner of bacteriophage mu. J. Biol Chem. 267:19101-19106.
Johnson, W.C. 1990. Protein secondary structure and circular dichroism: A practical guide. Proteins Struct. Funct. Genet. 7:205-214.
Scoble, H.A. and Martin, S.A. 1990. Characterization of recombinant proteins. Methods Enzymol. 193:519-536.
LeGendre, N., Mansfield, M. Weiss, A., and Matsudaira, P. 1993. Purification of proteins and peptides by SDS-PAGE. In A Practical Guide to Protein and Peptide Purification for Microsequencing (P. Matsudaira, ed.) pp. 74-101. Academic Press, San Diego.
Shire, S.J. 1992. Determination of the molecular weight of glycoproteins by analytical ultracentrifugation. Beckman Technical Information, Publication DS-837. Spinco Business Unit, Palo Alto, Calif.
McRee, D.E. 1993. Practical Protein Crystallography. Academic Press, San Diego. McRorie, D.K. and Voelker, P.J. 1993. Self-associating systems in the analytical ultracentrifuge. Vol. II, Analytical Ultracentrifugation Series. Beckman Instruments, Fullerton, Calif. Mercier, J.-C., Grosclaude, F., and Ribadeau-Dumas, B. 1971. Primary structure of bovine S1 casein. Complete sequence. Eur. J. Biochem. 23:41-51. Mulkerrin, M.G. and Cunningham, B.C. 1993. Spectral analysis of site-directed mutants of human growth hormone. In Protein Folding: In Vivo and In Vitro (J.L. Cleland, ed.) pp. 240-253. ACS Symposium Series 526. American Chemical Society, Washington, DC. Pappin, D.J.C., Hojrup, P., and Bleasby, A.J. 1993. Rapid identification of proteins by peptide-mass fingerprinting. Curr. Biol. 3:327-333. Pennica, D., Kohr, W.J., Fendly, B.M., Shire, S.J., Raab, H.E., Borchardt, P.E., Lewis, M., and Goeddel, D.V. 1992. Characterization of a recombinant extracellular domain of the type I tumor necrosis factor receptor. Biochemistry 31:1134-1141. Phillies, G.D. 1990. Quasielastic light scattering. Anal. Chem. 62:1049-1057.
Overview of the Characterization of Recombinant Proteins
Ptitsyn, O.B., Pain, R.H., Semisotnov, G.V., Zerovnik, E., and Razgulyaev, O.I. 1990. Evidence for a molten globule state as a general intermediate in protein folding. FEBS Lett. 262:20-24.
Price, N.A. and Johnson, C.M. 1989. Proteinases as probes of conformation of soluble proteins. In Proteolytic Enzymes: A Practical Approach (R.J. Beynon and J.S. Bond, eds.) pp. 163-179. IRL Press, Oxford.
Strickland, E.H. 1974. Aromatic contributions to circular dichroism spectra of proteins. Crit. Rev. Biochem. 2:113-175. Sturtevant, J.M. 1987. Biochemical applications of differential scanning calorimetry. Ann. Rev. Phys. Chem. 38:463-488. Surewicz, W.K., Mantsch, H.H., and Chapman, D. 1993. Determination of protein secondary structure by Fourier transform infrared spectroscopy: A critical assessment. Biochemistry 32:389-393. Thomas, D., Schultz, P., Steven, A.C., and Wall, J.S. 1994. Mass analysis of biological macromolecular complexes by STEM. Biol. Cell 80:181-192. Van Holde, K.E. 1975. Sedimentation analysis of proteins. In The Proteins, 3rd Edition, Vol. 1 (H. Neurath, ed.) pp. 225-291. Academic Press, N.Y. Varley, P.G. 1994. Fluorescence spectroscopy. Methods Mol. Biol. 22:203-218. Walsh, G. and Headon, D.R. 1994. Therapeutic proteins: Special considerations. In Protein Biotechnology. pp. 119-162. John Wiley & Sons, Chichester, U.K. Wang, R. and Chait, B.T. 1994. High-accuracy mass measurement as a tool for studying proteins. Curr. Opin. Biotechnol. 5:77-84. Wang, R., Chait, B.T., and Kent, S.B.H. 1994. Protein ladder sequencing towards automation in techniques. In Protein Chemistry V (J.W. Crabb, ed.) pp. 19-26. Academic Press, New York. Wetlaufer, D.B. 1962. Ultraviolet spectra of proteins and amino acids. Adv. Protein Chem. 17:303390.
7.1.12 Current Protocols in Protein Science
Wettenhall, R.E. and Cohen, P. 1982. Isolation and characterization of cyclic AMP–dependent phosphorylation sites from rat liver ribosomal protein S6. FEBS Lett. 140:263-269. Williams, R.W. 1986. Protein secondary structure analysis using Raman amide I and III spectra. Methods Enzymol. 130:311-331. Wood, S.P. 1989. Purification for crystallography. In Protein Purification Applications. A Practical Approach (E.L.V. Harris and S. Angal, eds.) pp. 45-58. IRL Press, Oxford. Yang, J.T., Wu, C.-S., and Martinez, H. 1986. Calculation of protein conformation from circular dichroism. Methods Enzymol. 130:208-297.
Kyte, J. 1995. Structure in Protein Chemistry. Garland Publishing, N.Y. Structural biology emphasis, with a clear and concise explanation of X-ray diffraction analysis. Excellent illustrations. Campbell, I.D. and Dwek, R.A. 1984. Biological Spectroscopy. Benjamin/Cummings Company, Menlo Park, Calif. Very good explanations of the theoretical principles, along with many practical applications.
KEY REFERENCES
Contributed by Nancy D. Denslow University of Florida Gainesville, Florida
Cantor, C.R. and Schimmel, P.R. 1980. Biophysical Chemistry. Part II: Techniques for the Study of Biological Structure and Function. Freeman, San Francisco.
Paul T. Wingfield National Institutes of Health Bethesda, Maryland
Fairly advanced text that includes descriptions of the spectroscopic and ultracentrifugation methods. Freifelder, D. 1982. Physical Biochemistry: Applications to Biochemistry and Molecular Biology. Freeman, N.Y.
Keith Rose University Medical Centre (CMU) Geneva, Switzerland
Excellent introductory text with especially strong sections on spectroscopy and centrifugation. Read this book first.
Characterization of RecombinantProduced Proteins
7.1.13 Current Protocols in Protein Science
Determining the Identity and Purity of Recombinant Proteins by UV Absorption Spectroscopy
UNIT 7.2
Ultraviolet absorption spectrophotometry can be used to confirm the identity and, to a lesser extent, assess the purity of recombinant proteins and their peptide fragments. The near-UV (250 to 350 nm) absorbance spectrum of a protein is almost entirely a function of its aromatic amino acids—tryptophan, tyrosine, and phenylalanine (Fig. 7.2.1). Because each protein (gene product) has a unique amino acid sequence, the particular aromatic amino acid content of each protein results in a unique spectrum in the near-UV region. The highly specific microenvironment experienced by each aromatic residue in the three-dimensional protein matrix results in fine shifts in a protein’s spectrum, which can be detected (Basic Protocol 1) and analyzed (Support Protocol). Thus, quality UV spectra serve as indicators of the structural integrity of proteins. In contrast to electrophoretic analyses (Chapter 10), UV spectroscopy allows the detection of nucleic acids, certain prosthetic groups, and metal ions which are frequently associated with proteins. The unique UV spectral properties of proteins can in turn be used to assess their purity. This application is inherent in the use of a diode array detector to monitor the effluent
+200
Trp
Trp
5540
– 200
Tyr
Absorbance
1480
d2(absorbance)/dλ2
0 +50
Tyr
– 50
0 Phe
Phe
+10
195
– 10 0 240 250 260 270 280 290 300 310 Wavelength (nm)
0 240 250 260 270 280 290 300 310 Wavelength (nm)
Figure 7.2.1 Normal (zero-order) and second-derivative spectra of aromatic model compounds: N-acetyl-L-tryptophanamide, N-acetyl-L-tyrosineamide, and N-acetylphenyl ethyl ester (solid lines) and “average” aromatic amino acid residues as determined using a set of globular, water-soluble proteins (dotted lines). Absorbance and second-derivative values correspond to 1 M solutions, 1-cm path length. The band positions in denatured proteins usually have intermediate positions.
Characteristics of Recombinant Proteins
Contributed by Henryk Mach, C. Russell Middaugh, and Nancy D. Denslow
7.2.1
Current Protocols in Protein Science (1995) 7.2.1-7.2.21 Copyright © 2000 by John Wiley & Sons, Inc.
Supplement 1
from a high-performance liquid chromatography (HPLC) column. A protocol using this technique to assess the purity of recombinant proteins is presented in Basic Protocol 2. BASIC PROTOCOL 1
ANALYSIS OF PROTEINS USING NEAR-UV SPECTROPHOTOMETRY This procedure identifies proteins using near-UV spectrophotometry. Protein solution is placed in a UV-transparent quartz cuvette, absorption of light is measured as a function of wavelength, and the resulting spectrum is transferred either to a displaying device (e.g., CRT or printer) for inspection or to a computer that can employ a spreadsheet or other program for derivative calculations and/or quantitative analyses (see Support Protocol). Materials Detergent solution Methanol 0.1 N hydrochloric acid Acid/ethanol cleaning solution (see recipe) 10 N NaOH 4% holmium oxide (aqueous solution in 10% perchloric acid; see recipe) Buffer identical to that containing the protein (reference standard) Sample for analysis in solution at an appropriate concentration (e.g., 0.05 to 1 mg/ml; see below) Vacuum-powered cuvette washer (NSG Precision Cells, Kontes Glass, or equivalent) Two factory-matched synthetic quartz (Suprasil or equivalent) cuvettes (Hellma, NSG Precision Cells, or equivalent; for double-beam spectrophotometers) or a single cuvette (for single-beam use) Gel-loading plastic pipet tips (Marsh Biomedical Products; Rainin Instruments) for a pipettor (Pipetman, Eppendorf, or equivalent) CAUTION: Employ appropriate safety precautions when using concentrated acids and bases. Ready the spectrophotometer 1. Turn on the spectrophotometer and the UV lamp; allow 20 min for warm-up. Lamp stability can be monitored by recording a baseline as a function of time. Only the UV (deuterium) lamp need be turned on; it should provide sufficient light intensity up to ∼400 nm. Higher wavelengths are needed only for proteins with visible-light-absorbing chromophores, such as heme proteins and metalloproteins. The instrumental wavelength cross-over point (the wavelength at which the light source is switched from the deuterium to the tungsten lamp) must be set ≥ to 360 nm.
Determining the Identity and Purity of Recombinant Proteins by UV Absorption Spectroscopy
Prepare the cuvette 2. Use the vacuum-powered cuvette washer to wash the cuvettes. Apply a detergent solution briefly and wash three or four times with water, waiting each time until all the water is exhausted from the cuvette. Finally wash with methanol to remove water-insoluble impurities and speed up drying. Wipe the outside of the cuvette with a tissue (Kimwipe) or, better, with lens cleaning paper and inspect visually. If not clean, repeat the cleaning procedure starting with 0.1 N HCl instead of detergent. If the cuvette is still not clean, soak it in acid/ethanol solution for several hours. As a last resort, soak briefly (20 min) in 10 N NaOH. Cuvettes should always be cleaned immediately after use. Denatured proteins may often be removed by soaking overnight in a mild pepsin solution. Prolonged contact with the NaOH may partially dissolve quartz faces of the cuvette.
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Current Protocols in Protein Science
Set up appropriate instrument parameters 3. Identify the cross-sectional area of the light beam (from the instrument’s instruction manual) and determine the correct height to which the cuvettes should be fitted. The light beam may be as large as 1 × 2 cm. The sample volume must be large enough so that the meniscus is completely above the beam. The light must not impinge on the air-solution interface. If the beam is wider than the portion of the cuvette containing the solution, the remaining portions of the cuvette walls should be masked by a light-absorbing coat; special blackened cuvettes are available for this purpose. In some instruments the height of the beam can also be reduced; do so if sample availability is a problem. The cuvettes must be positioned vertically; any deviations may alter the effective path length and reduce the precision of measurements.
4. Adjust the wavelength (scanning) range to 240- to 360-nm. For diode-array spectrophotometers, use 10-sec measurement times. For conventional scanning spectrophotometers use bandwidths of 1 or 2 nm, time constants (response times) of 2 sec (medium) or less, and scanning speeds of 60 nm/min or less. Set the data point interval to 1 nm (scanning instruments only). Use the same settings if spectra are to be compared to previously collected ones.
Measure the spectrum of a reference standard 5. Measure the baseline (background) spectrum with no cuvettes in the sample holders. Insert a sealed cuvette containing 4% holmium oxide (aqueous solution in perchloric acid; Weidner et al., 1985) into the sample holder. Measure the spectrum and save it on a disc or other permanent form of storage. Collect a second spectrum using a scanning speed approximately one-half of that initially used. 6. Determine the exact peak positions by calculating the first-derivative spectrum in the 280- to 290-nm range. If the spectrophotometer has a built-in derivative calculation ability, choose derivative order 1, polynomial degree 2, and a window of five data points. If not, transfer the spectra in ASCII (text) format (or type it) into a spreadsheet (Excel, LOTUS-123, or equivalent) and perform the calculation using Equation 7.2.1 (Savitzky and Golay, 1964; Steiner et al., 1972), where FD(λ) is the first-derivative value at the integral wavelength λ, A(λ ± n) is the absorbance values at wavelength λ + n (where n = −2, −1, 1, and 2), and k is the data point interval (in nanometers). FD(λ) =
−2[A(λ − 2)] − A(λ − 1) + A(λ + 1) + 2[A(λ + 2)] 10 × k Equation 7.2.1
The values of n double if a 2-nm data point spacing is employed. The derivative calculation using an Excel spreadsheet is performed as follows: (1) copy the absorbance values of the spectrum into the first column (typically A1:A100 range); (2) fill second column (B1:B100) with the corresponding wavelength values, entering the first wavelength and using Edit/Fill/Series commands; (3) enter the formula in the cell of the third column, which corresponds to A(λ) absorbance value (B3)—e.g., for Equation 7.2.1 and k=1, cell C3 contains =(−2×A1−A2+A4+2×A5)/10 (derivative values cannot be calculated for first two and last two data points because a five-data point window is used); (4) copy the content of that cell (the formula) down onto the entire column (B3:B98) using Edit/Copy and Edit/Paste commands; and (5) use the Chart Wizard option to plot the resulting spectrum.
7. Note where the spectra cross the zero line (change sign from positive to negative) and calculate the exact positions of these intercepts using Equation 7.2.2, where λ is the
Characteristics of Recombinant Proteins
7.2.3 Current Protocols in Protein Science
Supplement 1
wavelength prior to the intersection point, k is the data point interval (in nanometers), and |FD(λ)| and |FD(λ + 1)| are the absolute values of the first derivative at the integral wavelengths λ (before the intersection point) and λ + 1 (after the intersection point). This equation arises from simple geometric interpolation, illustrated for the second-depeak position (nm) = λ + k
|FD (λ)| |FD (λ)| + |FD (λ + 1)|
Equation 7.2.2
rivative spectra in the inset of Figure 7.2.2B. Note that the resulting intersection point is the position of the peak near 287 nm and serves as a reference point for further determination of exact peak positions of the protein spectra.
8. Calculate the intersection point for the second spectrum collected using a slower scanning speed (scanning instruments only); if the position is different, decrease the time constant (response time) or use a slower scanning speed until the positions of the intersections become independent of these parameters. Test for the proper response time only with the first spectra acquired. Once established, the settings can then be used with confidence. The 287-nm holmium oxide peak is found at 287.18 nm (for 1-nm bandwidths) and at 287.47 nm (for 2- and 3-nm bandwidth) on an instrument extensively verified by measurements of mercury and deuterium emission lines (Weidner et al., 1985). Commercial instruments will deviate somewhat from this value. Correct the wavelength results for proteins for the difference observed. Diode-array instruments (see Basic Protocol 2) will typically require such calibration once a year; scanning instruments should be calibrated frequently depending on use. If a holmium oxide standard is not readily available, Trp (monomeric amino acid) or a related compound can be used for reference purposes.
Measure the protein spectrum 9. Identify the number of light beams by looking at the number of sample holders and their positions in the sample compartment. In a double-beam spectrophotometer, use two matched cuvettes and perform an initial background (baseline) measurement with just buffer (identical to that in which the protein sample is dissolved) in both cuvettes. Then substitute protein solution for the buffer in the sample position cuvette, leaving the reference buffer in the other cuvette. Acquire the sample spectrum and subtract the previously measured background spectrum if necessary. In a single beam spectrophotometer, use a single cuvette. Measure the background with buffer in the cuvette. Then remove the buffer, measure the spectrum of the unfiltered protein solution using the same cuvette in the same orientation, and subtract the background spectrum (baseline). Sample can be removed from the cuvette using a pipettor with gel-loading plastic pipet tips. Do not scratch the inner optical surface of the cuvette. Many proteins, especially when partially unfolded, strongly adhere to quartz and are not removed by brief detergent or organic solvent washes. If UV absorption peaks at 270- to 280-nm are detected in the baseline, clean the cuvette again (step 2).
Determining the Identity and Purity of Recombinant Proteins by UV Absorption Spectroscopy
10. Note the absorbance at the protein peak centered between 275 and 282 nm. If the absorbance value is >1.0 at the maximum, check the range of linearity of the instrument in use (from the instruction manual). If the absorbance value observed is above this range, dilute the sample or decrease the path length. If a fluorescence-type 2-5 × 10–mm cell is used, turn the cuvette sideways to reduce the path length. A new baseline should be obtained. If the absorbance falls below 0.2, carefully observe the
7.2.4 Supplement 1
Current Protocols in Protein Science
noise levels and use longer acquisition times (both for the baseline and the actual spectrum). 11. Save the spectrum in a proper format for the particular instrument employed or transfer the spectrum in an ASCII (text) format to a disc for retrieval by a spreadsheet for further analysis. If this is not feasible, print or manually record absorbance values as a function of wavelength. Most protein solutions containing 1 mg/ml protein have an absorbance near 1.1 ± 0.5 in the 280-nm region. If an estimate of the concentration of the solution is available, the sample can be diluted or concentrated to obtain an absorbance in the 0.2 to 1.0 range. Alternately, make a preliminary measurement at a single wavelength (e.g., 280 nm) to obtain this information. Note that with modern spectrophotometers very low values of absorbance (<0.01) can be accurately measured; however, care must be taken to avoid adsorption artifacts, as a substantial portion of the protein may now reside on the interior surface of the cuvette. This can be checked by obtaining a spectrum of the empty cuvette after carefully removing the protein solution.
INTERPRETING PROTEIN NEAR-UV SPECTRA Normal (zero-order) spectra are spectra of absorbance versus wavelength as measured by a spectrophotometer without derivative calculation. Different regions of the spectrum contain information about various aspects of the structure of the protein. The 320- to 350-nm region may contain information about aggregation state (due to light scattering) or the presence of UV-absorbing prosthetic groups; the 250- to 300-nm region provides information about the aromatic amino acids and their environment (Fig. 7.2.1 and Table 7.2.1). Numerical calculation of the second derivative of the spectrum minimizes the effect of broad spectral components (such as light scattering or anion absorption), exaggerating fine structure originating from absorption by aromatic rings. This method is particularly useful for precise determination of tertiary structure–dependent band positions as well as for quantitative analysis.
SUPPORT PROTOCOL
Correct for light scattering 1. Examine the spectral region between 320 and 350 nm (Fig. 7.2.2 contains an example for a typical protein). Significant “apparent absorbance” of gradually, almost linearly decreasing intensity with increasing wavelength indicates that a portion of the light beam did not reach the detector because it was scattered. Absorbance that falls to zero at 350 nm or does not gradually decline (Fig. 7.2.2) is probably true absorbance arising from prosthetic groups or contaminants.
2. Use Equation 7.2.3 to calculate optical density (OD) due to light scattering, where a and b are constants. log OD = a log λ + b Equation 7.2.3
If the instrument’s software cannot make such a correction, use a spreadsheet (Microsoft Excel, LOTUS-123, or equivalent; see Basic Protocol 1, step 6 annotation, for an example) to generate the light-scattering curve using optical density values at 320 nm and 350 nm. Calculate the light-scattering correction from Equation 7.2.4, where m = 64.32 − 25.67 log λ. Ac = 10[(m + 1) × log OD320 − m × log OD350] Equation 7.2.4
Characteristics of Recombinant Proteins
7.2.5 Current Protocols in Protein Science
Supplement 2
If this is not feasible, calculate the light-scattering OD at 260 nm (m = 2.31) and 280 nm (m = 1.50) with a calculator. If significant light scattering is detected, remeasure the sample after filtration through a 0.1 to 0.45-micron diameter cut-off filter. The difference of protein concentration before and after filtration will indicate the fraction of aggregated protein. Compare the second-derivative spectra to determine whether the initial sample was homogenous. Note that optical density (OD) refers to the total extinction of light produced by all processes (including light scattering) whereas absorbance (A) refers only to the actual absorption process. The coefficient a varies from −4 (the ideal case of scattering from a small particle, not larger than one-tenth the wavelength of the light; this situation is known as Rayleigh scattering) to +2.2 for submicron-sized aggregates (Mie theory, in which the scattering intensity becomes a complex function of molecular size; Timasheff, 1966). Because the light-scattering contribution to the optical density is almost linear in the 300- to 350-nm region, a simple linear fit using a spreadsheet or even a crude graphic estimate can serve as a rudimentary measure of the correction necessary to obtain the correct absorbance in the 250- to 300-nm range.
Visually recognize the presence of the individual types of aromatic residues 3. Inspect the spectrum in the 250- to 300-nm range. A small inflection point on the declining shoulder of the main peak around 290 nm (Fig. 7.2.2) originates from Trp residues; the sharper the inflection point, the more homogeneous and apolar the average microenvironment of these side chains. The distinctive spectral features of Tyr residues are usually masked and less pronounced as a consequence of the less-sharp fine structure of these residues, their broadening by the medium, and their greater heterogeneity from their usually greater frequency in proteins. The presence of the Tyr residues can be deduced, however, from the deviation of the overall spectral shape from that expected for pure Trp. The presence of Phe residues is usually manifested as a series of weak but distinctive bumps superimposed on the rising 250- to 270-nm region, which is dominated by more strongly absorbing Trp and Tyr side chains.
Table 7.2.1 Spectroscopic Properties of UV-Absorbing Amino Acids, Natural Ligands (Prosthetic Groups), and Nucleic Acids
Chromophore
Determining the Identity and Purity of Recombinant Proteins by UV Absorption Spectroscopy
Tryptophan (in native proteins) Tyrosine (in native proteins) Disulfide bond (glutathione) N-Acetyl-L-tryptophanamide N-Acetyl-L-tyrosinamide N-Acetyl-L-tyrosinamide N-Acetyl-L-phenylalanine ethyl ester Cu2+ (azurin) FAD (pyruvic dehydrogenase) Fe3+-heme (cytochrome c, reduced) FMN (amino acid oxidase) Retinal-Lys (rhodopsin) DNA (native, per base) RNA (per base)
λ (nm)a
ε (M−1 cm−1)b
280 280 280 280 275 280 257 781 460 550 455 498 258 258
5540 1480 134 5390 1390 1185 195 320 1270 2770 1270 4200 6600 7400
aWavelengths listed indicate the peak maxima, except for tyrosine, N-Acetyl-L-tyrosinamide, and L-cystine at 280 nm. bMolar absorptivity values of tryptophan and tyrosine in native proteins as well as oxidized glutathione (disulfide bond) at 280 nm are used for the calculation of molar extinction coefficients of proteins.
7.2.6 Supplement 2
Current Protocols in Protein Science
A
1.2 Total Trp Tyr LS S-S Phe
Absorbance
1 0.8 0.6 0.4 0.2 0
B
0.015 0.01
d2(absorbance)/dλ2
0.005 0 –0.005 –0.01
band position
–0.015
0
λ+1 λ
–0.02 –0.025 250
260
270
280
290
300
310
320
330
340
350
Wavelength (nm)
Figure 7.2.2 (A) Normal (zero-order) spectrum of a 100 µM solution of a hypothetical protein containing 1 Trp, 2 Tyr, 3 Phe, and 4 Cys (S-S) residues. The constituent spectral components and a typical light scattering (LS) contribution are also shown. (B) Calculated second-derivative spectra of A. The second-derivative spectra of cystine and light-scattering components are essentially zero. Inset: Enlargement of the marked segment illustrating the interpolation between adjacent secondderivative points that yields the precise position of the intersect with the wavelength axis.
Estimate the nucleic acid content 4. Calculate the ratio R of absorbances at 260 nm and 280 nm, A260,c to A280,c after correction for light scattering (Equation 7.2.4). Use Table 7.2.2 to estimate % weight of DNA (%N) in the sample (Glasel, 1995). Estimate protein concentration in the sample 5. If a significant amount of DNA was detected (R>0.70), calculate the protein absorbance A280,c,p using % weight of protein and DNA found in Table 7.2.2, where E0.1% is the extinction coefficient of the protein (per 1 mg/ml). If the amino acid composition is known, calculate E0.1% using Equation 7.2.7 and divide the result (ε) by the
Characteristics of Recombinant Proteins
7.2.7 Current Protocols in Protein Science
Supplement 1
Table 7.2.2 Theoretical R(260/280) Values vs %P and %N in Mixtures of Nucleic Acids and Proteinsa
%Pb
%Nb
100 99 98 97 96 95 90 80 60 40 20 0
R(260/280)
0 1 2 3 4 5 10 20 40 60 80 100
0.57 0.70 0.81 0.90 0.99 1.06 1.32 1.59 1.81 1.91 1.97 2.00
aAmount of DNA as small as 1% can be detected. This is due to the
approximately tenfold higher values of the DNA extinction coefficient (i.e., E0.1% of 10 and 20 at 280 nm and 260 nm, respectively). Average value of R for 12 globular proteins was 0.57 +/− 0.06. The empirical equation for this table is: %N=(11.16×R−6.32)/(2.16−R) (Glasel, 1995; see also Manchester, 1995, for further comments). bAbbreviations: %N, percent weight of DNA in the sample; %P, percent weight of protein in the sample.
molecular weight of the protein. For a crude estimation, use the average value of E0.1% = 1.1. A280,c,p =
% P× E0.1% % P × E0.1% + % N × 10
Equation 7.2.5
6. Use the spectral intensity values at 280, 320, and 350 nm to assess the concentration of protein based on the known amino acid composition of the protein using Equation 7.2.6 and Equation 7.2.7 (Mach et al., 1992b), where C is the protein concentration (molar), ε is the molar extinction coefficient (M−1 cm−1) that would be obtained from a solution of 1 M concentration and a 1-cm path length (L), and NTrp, NTyr, and NS-S are the numbers of Trp, Tyr, and disulfide bonds (cystine residues) in the protein. A280,c is the optical density at 280 nm (OD280) corrected for light scattering (see Equation 7.2.4). When sufficient amount of protein is available, the extinction coefficient ε is usually determined by quantitative amino acid analysis (UNIT 3.2). C =
A280,c εL
Equation 7.2.6 Determining the Identity and Purity of Recombinant Proteins by UV Absorption Spectroscopy
ε = 5540NTrp + 1480NTyr + 134NS−S Equation 7.2.7
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Based on studies of a series of proteins (Equation 7.2.7), the above relationships can be used to determine the extinction coefficients of proteins and, consequently, their aqueous concentration to an accuracy generally within 2%.
Calculate and visually inspect the second-derivative spectra 7. Calculate the second-derivative spectrum. If the instrument’s software has derivative computation capability, choose derivative order 2, polynomial order 2, and a window of five data points. Otherwise use Equation 7.2.8 (Savitzky and Golay, 1964; Steiner et al., 1972), where SD(λ) is the second-derivative value at integral wavelength λ, A(λ ± n) is the absorbance value at λ + n wavelength (where n = −2,−1, 0, 1 and 2), and k is the data point interval (in nanometers). SD(λ) =
2[A (λ − 2)] − A (λ − 1) − 2[A (λ) − A (λ + 1) + 2[A (λ + 2)] 7×k Equation 7.2.8
See Basic Protocol 1, step 6 annotation, for an example. Note that the values of n double for 2-nm spectral bandwidth.
8. Inspect the features of the resultant second-derivative spectrum. The negative peak centered near 290 nm (Fig. 7.2.2) originates from the Trp inflection point observed in the zero-order spectrum. Although its position is partially influenced by the overlapping Tyr spectrum, the position of the negative 290-nm peak reflects the average polarity of Trp microenvironments: the more hydrophobic the environment, the higher the wavelength (i.e., an apolar environment produces a “red shift” in this peak; see Fig. 7.2.1). The magnitude of this negative peak is diminished by a Tyr band of opposite (positive) sign. In contrast, Trp and Tyr bands of the same sign (both negative) amplify the composite second-derivative peak near 280 nm. The positive peak centered at ∼285 nm is composed mostly of the Trp component, as the second-derivative spectrum of Tyr crosses the zero axis in this region. In some proteins individual Tyr residues have microenvironments different enough to produce 2- to 3-nm shifts and band overlap sufficient to obviate individual peaks in this region. The alternating positive and negative peaks between 250 and 270 nm are the result of the wave-like fine structure from Phe side chains seen in the zero-order spectrum and are affected only marginally by Trp and Tyr counterparts. The magnitudes of the second-derivative peaks are proportional to the concentration of aromatic amino acids, revealing both the relative content of each type of aromatic residue and the protein concentration. The relative heights of the second-derivative peaks constitute a characteristic “fingerprint” by which purity and structural integrity of a protein can be recognized. Some instruments (e.g., Hewlett-Packard 8452) have software capable of comparing two spectra using linear regression and providing a quantitative “match factor.” Unlike that for holmium oxide, the first derivative cannot be used for protein solutions because of residual nonzero values originating from light scattering, cystine, and other amino acid absorbance below 270 nm.
9. Calculate the positions of the intersections of the second derivative spectrum with the wavelength axis in the 285- to 295-nm region. Use Equation 7.2.9, where λ is the wavelength prior to the intersection point, k is the data point interval (in nanometers), and |SD(λ)| and |SD(λ + 1)| are the absolute values of the second derivative at the integral wavelengths λ (before the intersection point) and λ + 1 (after the intersection point).
peak position (nm) = λ + k
| SD(λ)| | SD(λ)| + | SD(λ + 1)|
Equation 7.2.9
Characteristics of Recombinant Proteins
7.2.9 Current Protocols in Protein Science
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These values can be used as probes of protein structural integrity. In future manipulations or purification of the same protein, identical values of these parameters will indicate that the structure of the protein did not significantly change. The values of intersection points are typically reproducible within a 0.05-nm error margin using diode-array spectrophotometers. Scanning spectrophotometers require frequent wavelength calibration using standardized spectra to obtain this precision. The intersections of the second-derivative spectra of the holmium oxide standard determined using Equation 7.2.9 provide suitable reference points with which to compare results obtained at different times or using different instruments. BASIC PROTOCOL 2
ANALYSIS OF PROTEINS USING AN HPLC WITH A DIODE-ARRAY DETECTOR Purifying recombinant proteins relies on large-scale chromatographic procedures that generally lack the resolution of the analytical methods on which they are based. A common problem in the large-scale process is that a second protein (or some other compound, e.g., a detergent) may coelute with the protein of interest, contaminating the sample. Highperformance liquid chromatography (HPLC) with a diode-array detector is a good method to determine the purity of a sample. By exquisitely regulating the pumps, this chromatography system minimizes flow problems that can otherwise interfere with peak detection, yielding highly reproducible gradients. Thus, HPLC provides the best chromatographic resolution of a protein. A diode-array detector can be coupled to virtually any type of chromatographic separation, including reversed-phase, ion exchange (UNIT 8.2), and gel filtration (UNIT 8.3). The method assesses the purity of proteins or peptides equally well. In this protocol, the purity of a protein is determined using a C4 reversed-phase column. The same protocol can be adapted to peptides by substituting a C18 reversed-phase column. Materials Buffer A: 0.1% (v/v) trifluoroacetic acid (TFA; reagent grade) in water (Milli-Q or equivalent purity) Buffer B: 0.09% (v/v) TFA in 80% acetonitrile (HPLC grade) C4 reversed phase column and guard: (e.g., Vydac C4, 250 × 4.6-mm-i.d. column, 5-µm particle size, 300-Å pore size for samples in the 0.5 to 5 nmol range or a 250 × 2.1-mm-i.d. column for samples in the 100 to 500 pmol range) HPLC with a diode-array detector (e.g., Hewlett-Packard, Waters, Beckman) 1. Set scan limits on the diode array detector to cover the 200- to 300-nm region. Peptide bonds have strong absorption in the 200- to 230-nm region (wavelength maximum = 188 nm) and aromatic amino acids have specific maxima in the 240- to 300-nm region.
2. Set the primary sample wavelength at 215 nm, with secondary tracings being made at 254 nm and 280 nm. Set bandwidth at 4 nm. The sample wavelength is the wavelength at which absorbance is measured as a function of time for the whole run. The bandwidth can be set from 4- to 20-nm for a sample. A bandwidth of 4 nm for the primary wavelength of 215 nm means that the absorbance measurement will extend between 213 and 217 nm.
Determining the Identity and Purity of Recombinant Proteins by UV Absorption Spectroscopy
It is important to set the primary wavelength to measure peptide bonds. The peak absorbance (188 nm) cannot be used, however, because many of the HPLC reagents also absorb strongly in this region and obscure the signal from the protein. At 215 nm, the solvents do not interfere with the protein. The secondary tracings cover the absorption of aromatic amino acids. Although having
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the whole chromatogram at these secondary wavelengths is not essential for this method, it can be useful (see Anticipated Results).
3. Set reference wavelength at 550 nm with a bandwidth of 100 nm. The reference wavelength is generally set outside the range of protein absorption. Because proteins do not absorb at this wavelength, a wide bandwidth is recommended to avoid spurious readings. The absorbance of the reference wavelength is subtracted from the absorbance of the sample wavelength to give the absorbance of the signal for the protein.
4. Determine the points at which diode-array spectra will be taken and saved. Set acquisition and storage of spectrum to “peak controlled.” This means that diode-array spectra will be taken at a point on the upslope, at the apex, and at a point on the downslope of a peak. The information obtained will be used to assess peak purity. If a peak has shoulders or is asymmetrical, spectra will automatically be obtained at additional points using the peak purity program.
5. Set threshold to define height of signal expected for the smallest peak from 0.1 to 999 milliabsorbance units (mAU). A threshold of 5 mAU is suggested for a sample of 50 pmol protein. This parameter is relevant to the size of the record that will be made and saved during the run. Diode-array spectra will be saved only for peaks that are higher than the threshold height defined, not for peaks that are lower.
6. Set peakwidth to equal the half-height width expected for the narrowest peak, with limits from 0.007 to 10 min. A suggested starting value for this parameter is 0.15 min. Together with the threshold, the peakwidth determines the size of the record made for the run. Diode-array spectra will be saved only for peaks displaying a wider half-height width than the value chosen, not for peaks narrower than this width. This eliminates saving spectra for peaks that are due to noise.
7. Set stop time for acquisition of spectra. This value controls the diode-array detector only; it can be set to stop independently from the run, earlier than the run, or automatically with the run.
8. Set a gradient rate of 1% buffer B/min and a flow rate of 1 ml/min for the 4.6-mm-i.d. column, 200 µl/min for the 2.1-mm-i.d. column. Set the amount of buffer A plus the amount of buffer B to equal 100%. HPLC pumps are controlled by the B pump, which can vary from 0% to 100%. The amount of buffer A plus the amount of buffer B must equal 100%. Thus 0% buffer B means that the B pump is off and only the A pump is working (e.g., 100% A), and 100% buffer B means that the A pump is off and only the B pump is working. To achieve the same chromatographic separation on a 2.1-mm-i.d. column as obtained with a 4.6-mm-i.d. column, adjust the flow rate according to the ratio of the cross-sectional areas of the two columns. A 4.6-mm-i.d. column is roughly five times the cross-sectional area (πr2) of a 2.1-mm-i.d. column. Gradient rate defines the change in the gradient over time. The gradient should not be started until the sample has had time to flow onto the column. For a sample 100 ìl in volume and a flow rate of 1 ml/min, it will take 0.1 min to load the sample onto the column. The following table illustrates typical gradient conditions for a 100-ìl sample and a flow rate of 1 ml/min. This information would be entered into a liquid chromatography (LC) time table. The time required for the run should be determined and entered at the appropriate place in the program. This time limit should be long enough to allow for all of the events specified in the LC time table. Characteristics of Recombinant Proteins
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Time (min)
%B
0 1 51 56 66
10 10 60 80 10
After a trial run, if peaks of interest are too close to each other, the gradient rate can be made shallower (e.g., 0.5% buffer B/min) to spread out the gradient or steeper (e.g., 1.5% buffer B/min) to sharpen well-resolved peaks.
9. Equilibrate the column with 10% buffer B by running a volume equivalent to at least 3 column volumes. 10. Prepare experimental and control samples in 10% buffer B, keeping the volume to a minimum and making all samples the same volume. A good control sample will contain everything present in the experimental sample except the protein. The following table illustrates a sample preparation:
Protein sample Sample buffer HPLC buffer B
Control
Experimental
— 90 µl 10 µl
90 µl — 10 µl
Quantities of protein in the sample can range from 0.5 to 5 nmol for a 4.6-mm-i.d. column and from 100 to 500 pmol for a 2.1-mm-i.d. column. For reversed phase chromatography, sample volumes of 20 to 100 ìl will work.
11. Filter samples through a low-protein-absorption 0.21-µm pore size filter or microcentrifuge 5 min at maximum speed to remove particulates. 12. Choose an appropriate label for the control run. Inject the control sample into the HPLC apparatus. The control sample is used to control for spectral impurities in buffers. The control should be run before the experimental sample to permit careful inspection of the entire gradient. The HPLC starts acquiring data on injection. If a rheodyne injector is used, leave the sample loop on line long enough to allow the whole sample to be applied to the column.
13. Choose an appropriate label for the experimental run. Inject the experimental sample into the HPLC. As above, the injection of the sample starts the acquisition of data. The data is stored in a computer file under the given label.
Data analysis 14. Subtract the UV spectrum of the control from that for the protein to take into account impurities found in the sample buffer. 15. Run the peak purity program, which compares spectra for points at the apex and on the upslope and downslope of each peak. 16. Compare normalized overlaid spectra. Determining the Identity and Purity of Recombinant Proteins by UV Absorption Spectroscopy
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Acid/ethanol cleaning solution (3 N HCl/50% ethanol) Add 50 ml of 12 N HCl to 50 ml deionized water. Add the resulting solution to 100 ml ethanol. Store several months at room temperature. CAUTION: Never pour water or ethanol into the concentrated acid. Always pour the acid into water or ethanol. Wear appropriate personal protection.
Holmium oxide reference solution 4% holmium oxide in aqueous solution of 10% (v/v) perchloric acid sealed in a nonfluorescent fused silica cuvette of optical quality (National Institute of Standards and Technology, Standard Reference Materials catalog; see SUPPLIERS APPENDIX). Store at room temperature for several months. COMMENTARY Background Information Ultraviolet absorption spectroscopy played a key role in early investigations of protein structural transitions (Wetlaufer, 1962). The explosive growth of molecular biology and expanded use of more powerful spectroscopic techniques such as nuclear magnetic resonance (NMR), X-ray crystallography, circular dichroism, and fluorescence relegated this technique to use in simple assays and concentration determinations. During this period, however, electronic and optical systems improved considerably, allowing extraction of much more detailed information from simple near-UV spectra. It was soon shown that the magnitude of the near-UV second derivative peaks can be used for quantitative determination of the content of aromatic amino acids in proteins (Ichikawa and Terada, 1977; Servillo et al., 1982, Levine and Federici, 1982; Nozaki, 1990). These methods employ either peak-topeak distances or least-squares multicomponent analysis and are capable of resolving all three components simultaneously. In fact, the spectrum of a mixture of three proteins can be easily resolved by this method if the spectra of the individual proteins are known (Mach et al., 1989). The composition of simple protein mixtures and complexes can then be monitored without actual chromatographic separation. Generally, the spectra of the individual aromatic amino acids become red-shifted (move to higher wavelengths) upon incorporation into protein structures (see Fig. 7.2.1). These shifts can be as large as 5 nm for Tyr, 4 nm for Trp, and 2 nm for Phe residues (Mach and Middaugh, 1994). Of course, many side chains naturally exist in quite polar microenviron-
ments in native structures, so these shifts are not always so large. Unfolding of a protein usually results in some increased solvent exposure of aromatic side chains and, consequently, in blue shifts (toward lower wavelengths). If the spectral signal near 295 nm (Fig. 7.2.2) shifts left even a fraction of a nanometer, the absorbance value at any given wavelength in this vicinity may change significantly because of the steepness of the slope. This phenomenon can be effectively utilized to follow the unfolding of a protein by measuring the spectral intensity at this wavelength as a function of a structure-perturbing variable (e.g., temperature, pH, protein concentration, or urea concentration) or by measuring the difference spectra between two proteins of identical concentration, one of which is subject to structural perturbation. The resulting values, however, are relative. Use of the intersection of a portion of the second-derivative spectrum with the wavelength axis is therefore recommended. Application of more complex fitting routines and use of systematically shifted standard spectra allow the individual spectra of Trp, Tyr, and Phe to be resolved from the composite protein spectrum (Mach and Middaugh, 1994). This allows the fractional exposure of aromatic residues to be assessed using additions of apolar compounds as perturbants. The observed shifts are compared to the shifts in model, solvent-exposed compounds (e.g., acetylated and amidated aromatic amino acid derivatives), and the solvent exposure is estimated from a comparison of these values. In addition to proteins, DNA and RNA possess strongly absorbing chromophores (purine and pyrimidine bases) in the near-UV (Table 7.2.1), resulting
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Determining the Identity and Purity of Recombinant Proteins by UV Absorption Spectroscopy
in the well-known featureless spectrum of nucleic acids with a peak near 260 nm. The molar absorptivity (extinction coefficient) of polynucleotides is strongly structure dependent and this is often utilized in monitoring thermal structural transitions (i.e., in melting experiments). In addition, DNA and RNA have welldefined, albeit weak, second-derivative spectra that can be employed to determine the amount of protein in the presence of DNA (Mach et al., 1992a). Very strong absorbance is also produced by all proteins in the far-UV range (180 to 250 nm). This is primarily the result of the intense absorption by the peptide bond in the far-UV. Because of amide absorption near 188 to 190 nm, a peak is actually present, but the intense absorption by oxygen as well as most solutes in this region makes it difficult to resolve experimentally. In some spectrophotometers, this peak can be resolved by flushing the instrument with N2 and by the use of UV-transparent anions (e.g., phosphate or perchlorate), but this is rarely done today. Although the structure and position of this peak contain secondary-structure information as well as side-chain contributions, the peak is rarely used experimentally because of the superiority of circular dichroism measurements in the same region. It is occasionally employed, however, for concentration measurements or simple protein detection during chromatography by monitoring wavelengths on the slope of the peak (e.g., 210 and 215 nm). If the protein or peptide studied lacks aromatic amino acids, or the concentration of the sample does not produce sufficient signal, its concentration can be calculated by employing a set of average extinction coefficient values in the far-UV range that were obtained based on the analysis of twelve proteins (Scopes, 1974): (1) E(1 mg/ml, 1 cm) at 205 nm = 31 with stated error of 5% (2) E(1 mg/ml, 1 cm) at 205 nm = 27.0 + 120 A280/A205 with stated error of 2%. Extreme care should be taken when correcting for the many buffer components absorbing in this region. In principle, methods such as SDSpolyacrylamide gel electrophoresis (SDSPAGE; UNITS 10.1 & 10.3) are more effective than UV spectroscopy in assessing the purity of a protein sample, based on detection of the presence and absence of additional bands. Such methods, however, may fail to ascertain contaminating proteins of similar molecular weight, unless isoelectric focusing (IEF) gels
are used (UNITS 10.2 & 10.8). In contrast to electrophoretic (Chapter 10) or chemical (Chapter 11) analyses, the UV absorption technique is both nondestructive and rapid. Moreover, only a modest amount of sample is required. This unit describes recombinant protein analyses using the two general types of UV spectrophotometers (Fig. 7.2.3): conventional scanning instruments, in which light absorption is measured by mechanically moving a monochromator allowing data to be accumulated temporally as a function of wavelength (Basic Protocol 1 and Support Protocol) and diode-array spectrophotometers (Basic Protocol 2), in which a light beam containing all frequencies is simultaneously split onto an array of detectors (diodes) after passing through the sample. Diode-array spectrophotometers (HewlettPackard 8450 to 8453 series or Beckman DU 7000) are simpler and consequently have excellent wavelength reproducibility. A single spectrum is typically acquired in 0.1 sec; longer acquisition times provide an average of a number of measurements. Spectral bandwidth is fixed at 1 or 2 nm. The small cross-sectional area of the light beam (∼1×5 mm for the Hewlett-Packard diode-array instrument) allows small sample volumes and free choice of cuvette format. Such instruments can be easily used on-line to collect and analyze spectra of protein fractions leaving a chromatographic column—e.g., from gel filtration (UNIT 8.3), ionexchange (UNIT 8.2), and reversed-phase media. In contrast, conventional scanning spectrophotometers require longer acquisition times, since at any given time only one wavelength is read.
Critical Parameters The spectrophotometers contain moving internal parts that require frequent wavelength calibration. Of great importance when using scanning spectrophotometers is the setting of proper detector response time relative to scanning speed. A decrease in absorbance values, broadening of bands, and shifts are often observed in such cases. Although the intrinsic absorbance of the aromatic amino acids varies little with temperature, it is advisable in many cases to control the temperature of the sample. Many processes destructive to proteins (e.g., oxidation, deamidation, unfolding, and aggregation) are accelerated at elevated temperature (Volkin and Middaugh, 1992). Under most conditions of humidity, however, a temperature of 15°C should be regarded as minimal. Decreasing the
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A
b I0
I db
light source
sample
detector array
holographic grating
B
b I0
I db
light source
grating monochromator
sample
detector
Figure 7.2.3 Basic scheme for spectrophotometric measurements: lamp, sample and detector; b (path) and db are shown in the sample and I and I0 are denoted (see below). (A) Diode-array and (B) conventional scanning spectrophotometers. The relative decrease of the intensity of the light beam after passing through the sample is proportional to the distance (db) and the concentration of the absorbing component: −db/b = kc × db, where k is an absorption coefficient. Upon integration, the result is ln(I/I0) = − kcl. The fraction of light transmitted through the sample is known as the transmittance T = I/I0. Absorbance is defined as A = log I0/I = ε cb, where ε is a molar absorptivity (extinction coefficient), c is the molar concentration, and b is the path length (in centimeters). Alternatively, weight extinction coefficients E1 cm, 0.1% expressed per 1 mg/ml (0.1%) concentration are used.
temperature below this level may result in water condensing on the optical surface of the cuvette. The power of deuterium lamps used in UV spectrophotometers is usually not sufficient to cause photooxidation of aromatic residues. However, if the fluorescence spectra were acquired using the same sample, some of the tryptophan residues may have been oxidized to N-formylkynurenine and kynurenine by light from more powerful xenon lamps. This condition can be recognized by the appearance of a broad spectral band extending over 350 nm (similar to light scattering) and can be identified by comparing spectra as a function of irradiation time in a fluorometer or by direct observation of photoproduct fluorescence bands at ∼400 nm (with excitation at 320 nm). Solvent-accessible tryptophan side chains are particularly prone to photooxidation, with half-lives as short as several minutes (Mach et al., 1995). An aqueous buffer in which the protein of interest is stable should always be used. For example, phosphate-buffered saline, pH 7.4
(PBS; APPENDIX 2E), or an equivalent buffer containing 10 mM sodium phosphate and 120 to 150 mM sodium chloride, pH 7.0 to 7.4, is often employed. Choice of pH is important, because the spectral contribution of absorbing residues may be directly or indirectly (effect on protein conformation) pH dependent. Whereas tyrosine ionization (pK=10) results in an entirely different spectrum (λmax = 294 nm), ionization of nonabsorbing or weakly absorbing side chains (e.g., lysine, pK∼10; aspartic and glutamic acids, pK∼4.5; histidine, pK∼6.5; or cysteine, pK∼8.5) may produce subtle shifts if the microenvironments of UV-absorbing chromophores or the overall conformation are altered. The pH should be sufficiently different from the isoelectric point to ensure good solubility. Note that many agents used to improve protein solubility (e.g., surfactants, denaturing agents, and sugars; Middaugh and Volkin, 1992) may affect spectral band positions. For HPLC analysis, it is important that HPLC-grade solvents, free of UV-absorbing contaminants, be used. Similarly, the sample
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should be prepared carefully, with the highestgrade components, to avoid inadvertently introducing contaminants. All samples should be either filtered or centrifuged to remove particulate matter. In addition, HPLC tracings should be examined for shoulders and unusual peak broadening, which may indicate impurities (Zavitsanos and Goetz, 1991). Diode-array spectra will be obtained automatically at additional points if the computer detects an asymmetrical peak. This information is invaluable in determining peak purity. Finally, HPLC column performance should be routinely assessed by injecting a set of standard proteins or peptides (Mant and Hodges, 1991), as poor resolution may broaden peaks. Resolution deteriorates on columns that have been used excessively. One of the major advantages of using a diode-array detector to determine purity is that it is relatively unaffected by gradients and requires no prior knowledge of the sample. Absence of a strong absorption band between 200 and 230 nm indicates that the analyzed peak is not a protein but rather a nonproteinaceous contaminant, such as a detergent or a chromophore, picked up during the purification procedure.
Troubleshooting
Determining the Identity and Purity of Recombinant Proteins by UV Absorption Spectroscopy
A few recurring problems are encountered in the analysis of proteins by UV spectrophotometry. A major, often unrecognized difficulty is the phenomenon known as absorption flattening. Imagine a large number of protein molecules aggregating so that some incident light is able to pass “in between” them without encountering an absorbing entity. However, because of the high concentration of chromophores in the aggregated particle, photons that happen to encounter the aggregate are unlikely to emerge on the other side. As a result, the intensity of such spectra are significantly reduced and often distorted. If this problem is suspected, the length of the effective light path should be changed. If absolute values of protein concentration are found to be pathway dependent, this phenomenon may be present. The presence of anomalies such as significant curvature, negative values, or “bumps” in the 300- to 350-nm spectral region indicates either technical problems or the presence of unknown absorbing components, perhaps due to contaminating solutes. The spectra of any suspected absorbing impurities can be measured and compared to the experimental spectra to help determine if impurities are present.
Detergents are often found to produce such extraneous absorbance, and their potential contribution should be evaluated if they are employed at any stage in the purification process. If the second-derivative spectra do not resemble the examples shown in Figures 7.2.1 and 7.2.2, the most probable reason is too wide a bandwidth. Typically, a 2-nm bandwidth is the largest value permitting accurate derivative calculation. If the derivative spectrum is calculated by the instrument’s software, probably more than five data points (e.g., 7 to 21) are used for calculation of each value or the polynomial order is different from 2. If the calculated second-derivative spectra resemble random noise, the signal-to-noise ratio of the normal (zero-order) spectrum is probably too low to allow this type of analysis. This problem may be overcome by increasing spectral acquisition times or averaging a number of spectra. Problems in obtaining a stable baseline may arise from a number of factors. If the incorrect lamp was turned on—the tungsten-halogen (or “visible”) instead of the deuterium (or “UV”) lamp—spectra will not be obtainable. Failure of the UV lamp will usually manifest itself in an inability to obtain zero absorbance values during a “blank” measurement after collecting the baseline with both cuvettes containing the buffer. Many instruments contain an indicator of this condition. Care should be taken not to directly inspect the deuterium lamp for ignition, as UV irradiation can seriously damage the retina of the eye. The cuvette itself should also be reexamined. Only synthetic quartz provides sufficient transparency over the UV region. The inadvertent use of a glass cuvette will block passage of light through the system. If spectra cannot be obtained, the cuvette may have been positioned incorrectly, with opaque or black walls perpendicular to the light beam. Instability of the absorbance signal can occur for a number of other reasons, such as liquid smears, water vapor condensation or fingerprints on the optical surfaces, or air bubbles or other suspended or sedimenting particles in the test solution. As indicated previously, the meniscus should be above the upper edge of the beam and no light should be able to bypass the sample. In some cases, the problem of insufficient volume of sample available can be solved by inserting some space-filling material beneath the cuvette. In diode-array detection schemes, it is important to run a control sample consisting of the solution in which the protein is dissolved, to detect possible contaminants. Threshold and
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peak width values should be set correctly to ensure that all peaks of interest are seen by the diode-array detector. Otherwise, small contaminating peaks may be skipped.
Anticipated Results Figure 7.2.2A illustrates a typical example of a near-UV protein spectrum. Despite differences in aromatic amino acid compositions in proteins, spectra are usually qualitatively similar, with a maximum falling between 276 and 282 nm and only marginal absorbance above 300 nm. Small soluble monomeric proteins do not usually produce any light scattering. The second-derivative pattern should have the distinct features shown in Figure 7.2.2B characteristic of tryptophan, tyrosine, and phenylalanine residues, if these are present in the sequence. The position of the intersect with the wavelength axis of the declining second-derivative region near 290 nm is usually found at 289.41 ±0.61 nm using 1-nm data point spacing (based on the average from 14 tryptophan-containing proteins). This value should be ∼0.5 nm lower when the 2-nm data point interval is used. The reproducibility should be within 0.05 nm for pure and structurally intact proteins. The diode-array detector obtains spectra of very high quality and precision, enabling the operator to check the purity of a protein by comparing normalized spectra from different points of the eluted peak. It uses information within the protein (i.e., its amino acid composition) to check purity. Proteins of differing composition will have different spectral absorptions, particularly if they vary widely in aromatic amino acid content. Thus, if the retention times of two proteins are close enough, the proteins appear to be in the same peak. Based on their absorption characteristics the diode-array detector can distinguish them by comparing normalized spectra at the front end of the peak on the upward slope, at the back end of the peak on the downward slope, and at the apex. If the protein is pure, the spectra collected from these three points will superimpose exactly. If the protein is impure, the spectra collected from the three points will deviate, aside from the rare case in which the impurity and the sample exactly coelute from the column. The general rule is that small-to-moderate levels of impurity can be detected in a contaminated sample, provided the spectral properties of the contaminant vary from the recombinant protein. The examples in Figures 7.2.4 and 7.2.5
illustrate pure and impure samples. Figures 7.2.4A and 7.2.5A show the main chromatography tracings at 215 nm. The first visible peak in each chromatogram is the injection peak and the second is the protein of interest. Figures 7.2.4B and 7.2.5B show the stored diode-array spectra for points collected on the upswing, the apex, and the downswing of the protein peak of interest. A pure protein is characterized by identical spectra at these points, whereas an impure protein will have different spectra at each point. In Figure 7.2.4B, the spectra are close to identical, indicating that the protein is fairly pure. In Figure 7.2.5B, on the other hand, the spectra vary widely. Diode-array spectra for points 1 and 2 were collected early in the peak. These spectra have little absorbance in the region from 200 to 220 nm, indicating that a nonproteinaceous component is eluting at the front end of the BSA peak. Peaks 3, 4, and 5 correspond to the upswing, the apex, and the downswing, respectively, of the main peak. The diode-array spectra of these points are different enough to indicate contamination. Figures 7.2.4C and 7.2.5C show the superposition of tracings measured at the primary, secondary, and tertiary wavelengths. Note that these closely superimpose for the pure sample (Fig. 7.2.4C) but not for the impure sample (Fig. 7.2.5C).
Time Considerations The entire procedure for measurement and analysis of protein UV spectra can usually be completed within 30 min, of which 20 min is needed for lamp warm-up and sample preparation. This time should be shorter if the instrument is already turned on. Diode-array spectrophotometers interfaced to computers equipped with data processing programs are especially time efficient when a larger number of samples is analyzed. Characteristically, even the most complex analysis routines, when implemented within the existing software, usually require only seconds for completion. In the case of conventional scanning instruments, calibration (measurements of standards) combined with slower scanning rates may require some additional time. An HPLC run takes ∼2 hr, including the column equilibration with buffer A. Examination of the diode-array detector information should take at most 1 additional hour for the novice and 30 min for the experienced operator. Characteristics of Recombinant Proteins
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40 30 20
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Determining the Identity and Purity of Recombinant Proteins by UV Absorption Spectroscopy
75
76 77 Time (min)
78
79
Figure 7.2.4 Spectra of a pure protein sample (carbonic anhydrase). (A) Main chromatography tracings at 215 nm. Arrow indicates retention time of the component analyzed. (B) Stored diode-array spectra for points collected on the upswing, the apex, and the downswing of the protein peak. (C) Superposition of tracings measured at the primary, secondary, and tertiary wavelengths.
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70 60
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69 70 71 Time (min)
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Figure 7.2.5 Spectra of an impure protein sample (bovine serum albumin). (A) Main chromatography tracings at 215 nm. Arrow indicates retention time of the component analyzed. (B) Stored diode-array spectra for points collected on the upswing, the apex, and the downswing of the protein peak. (C) Superposition of tracings measured at the primary, secondary, and tertiary wavelengths.
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Literature Cited Glasel, J.A. 1995. Validity of nucleic acid purities monitored by 260 nm/280 nm absorbance ratios. BioTechniques 18:62-63. Ichikawa, T. and Terada, H. 1977. Second derivative spectrophotometry as an effective tool for examining phenylalanine residues in proteins. Biochim. Biophys. Acta 494:267-270. Levine, R.L. and Federici, M.M. 1982. Quantitation of aromatic residues in proteins: Model compounds for second-derivative spectroscopy. Biochemistry 21:2600-2606. Mach, H. and Middaugh, C.R. 1994. Simultaneous monitoring of the environment of tryptophan, tyrosine and phenylalanine residues in proteins by near-UV second derivative spectroscopy. Anal. Biochem. 222:323-331. Mach, H., Thomson, J.A., and Middaugh, C.R. 1989. Quantitative analysis of protein mixtures by second derivative absorption spectroscopy. Anal. Biochem. 181:79-85. Mach, H., Middaugh, C.R., and Lewis, R.V. 1992a. Detection of proteins and phenol in DNA samples with second derivative absorption spectroscopy. Anal. Biochem. 200:20-26. Mach, H., Middaugh, C.R., and Lewis, R.V. 1992b. Statistical determination of the average values of the extinction coeficients of tryptophan and tyrosine in native proteins. Anal. Biochem. 200:7480. Mach, H., Burke, C.J., Sanyal, G., Tsai, P.-K., Volkin, D.B., and Middaugh, C.R. 1995. Origin of the photosensitivity of a monoclonal immunoglobulin G. In Formulation and Delivery of Proteins and Peptides (J.L. Cleland and R. Langer, eds.) pp. 72-84. American Chemical Society, Washington, D.C. Manchester, K.L. 1995. Value of A260/A280 ratios of measurement of purity of nucleic acids. BioTechniques 19:208-210. Mant, C.T. and Hodges, R.S. 1991. Requirement for peptide standards to monitor column performance and the effect of column dimensions, organic modifiers, and temperature in reversedphase chromatography. In High-Performance Liquid Chromatography of Peptides and Proteins: Separation, Analysis, and Conformation (C.T. Mant and R.S. Hodges, eds.) pp. 289-295. CRC Press, Boca Raton, Fla. Middaugh, C.R. and Volkin, D.B. 1992. Protein solubility. In Stability of Protein Pharmaceuticals, Part A: Chemical and Physical Pathways of Protein Degradation (T.J. Ahern and M.C. Manning, eds.) pp. 109-134. Plenum Press, New York.
Determining the Identity and Purity of Recombinant Proteins by UV Absorption Spectroscopy
Nozaki, Y. 1990. Determination of tryptophan, tyrosine and phenylalanine by second derivative spectrophotometry. Arch. Biochem. Biophys. 277:324-333.
Savitzky, A. and Golay, M.J.E. 1964. Smoothing and differentiation of data by simplified least squares procedures. Anal. Chem. 36:1627-1639. Scopes, R.K. 1974. Measurement of protein by spectrophotometry at 205 nm. Anal. Biochem. 59:277-282. Servillo, L., Colonna, G., Balestrieri, C., Ragone, R., and Irace, G. 1982. Simultaneous determination of tyrosine and tryptophan residues in proteins by second-derivative spectroscopy. Anal. Biochem. 126:251-257. Steiner, J., Termonia, Y., and Deltour, J. 1972. Comments on smoothing and differentiation of data by simplified least squares procedure. Anal. Chem. 44:1906-1909. Timasheff, S.N. 1966. Turbidity as a criterion of coagulation. J. Colloid Interface Sci. 21:489497. Volkin, D.B. and Middaugh, C.R. 1992. The effect of temperature on protein structure. In Stability of Protein Pharmaceuticals, Part A: Chemical and Physical Pathways of Protein Degradation (T.J. Ahern and M.C. Manning, eds.) pp. 215247. Plenum Press, New York. Weidner, V.R., Mavrodineanu, R., Mielenz, K.D., Velapoldi, R.A., Eckerle, K.L., and Adams, B. 1985. Spectral transmittance characteristics of holmium oxide in perchloric acid solution. J. Res. Natl. Bur. Stand. 90:115-125. Wetlaufer, D.B. 1962. Ultraviolet spectra of proteins and amino acids. Adv. Protein Chem. 17:303390. Zavitsanos, P. and Goetz, H. 1991. The practical application of diode array UV-visible detectors to high-performance liquid chromatography analysis of peptides and proteins. In High-Performance Liquid Chromatography of Peptides and Proteins: Separation, Analysis, and Conformation (C.T. Mant and R.S. Hodges, eds.) pp. 553-562. CRC Press, Boca Raton, Fla.
Key References Cantor, C.R. and Schimmel, P.R. 1980. Absorption spectroscopy. In Biophysical Chemistry, Part II: Techniques for the Study of Biological Structure and Function, pp. 349-408. W.H. Freeman, New York. Provides theoretical background and practical considerations in absorption spectroscopy. Wetlaufer, D.B. 1962. See above. Provides comprehensive discussion of factors involved in analysis of proteins using UV spectroscopy. Savitzky, A. and Golay, M.J.E. 1964. See above. Provides formulas for smoothing and differentiation of data.
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Steiner, J. et al. 1974. See above. Provides formulas for smoothing and differentiation of data. Levine, R.L. and Federici, M.M. 1982. See above. Describes an application of second-derivative spectroscopy to analyze aromatic amino acid content employing a diode-array spectrophotometer. Mach, H., Middaugh, C.R., and Lewis, R.V. 1992b. See above.
Contributed by Henryk Mach and C. Russell Middaugh Merck Research Laboratories West Point, Pennsylvania Nancy Denslow University of Florida Gainesville, Florida
Describes calculation of protein extinction coefficients based on sequence and provides a formula for light-scattering correction at 280 nm.
Characteristics of Recombinant Proteins
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Determining the Identity and Structure of Recombinant Proteins
UNIT 7.3
After isolating a recombinant-derived protein and establishing that it is pure, it is necessary to confirm that it has the same primary sequence as predicted from the cDNA coding sequence. There are several reasons why this may not be the case: (1) errors in sequencing the gene (e.g., single-base errors resulting in single amino acid substitutions) and typographical errors in translating the sequence; (2) translation errors leading to the misincorporation of various amino acids (e.g., Harris et al., 1993; Randhawa et al., 1994); (3) unexpected translation start and finish points (see UNIT 6.1 for sample references); (4) retention or nonretention of N-terminal methionine when expressing proteins in E. coli (UNIT 6.1 and UNIT 7.1); (5) proteolytic processing by host cell proteases or, if the expressed protein is a protease, autolysis; and (6) chemical modification of specific amino acids: e.g., deamidation of amides (Asn and Gln), oxidation of cysteine and methionine, and, if urea was used for protein purification or folding, carbamylation of lysine residues (UNIT 6.1). Although this list is not exhaustive, it is enough to indicate that there are sufficient reasons to warrant a careful analysis of the primary sequence of the isolated protein to confirm that it corresponds to that expected from the cDNA sequence. The standard methods for confirming the primary sequence of a recombinant protein are combinations of (1) accurate amino acid analysis (UNIT 3.2) and mass spectrometry (Stults, 1995); (2) N- and C-terminal amino acid sequence determination by Edman degradation; and (3) peptide mapping in conjunction with the first two methodologies. In this unit peptide mapping protocols with separation of the constituent peptides by high-performance liquid chromatography (HPLC) analysis (Basic Protocol 1) and by high-resolution SDS-polyacrylamide gel electrophoresis (Alternate Protocol) are presented. The latter approach is included for the benefit of laboratories that do not have access to the more expensive HPLC equipment. Other peptide mapping techniques are described in UNIT 11.6. When peptide mapping a recombinant protein, the peptide bonds are first cleaved using specific proteases or chemical reagents to produce a mixture of peptides. The composition of the mixture is a function of both the primary sequence and the specificity of the protease or chemical reagent. In the protocols described, proteases with strict specificity are used, and as the translated protein sequence is known, it is possible to predict the number and size of the peptides. The composition of the peptides can be determined by mass spectrometry and/or amino acid analysis (UNIT 3.2). Peptide mapping is ideally suited for comparative purposes—for example, combined analysis of the recombinant protein and its natural counterpart (or some other well-characterized standard). The method also has the advantage of reviewing the whole sequence. The most common post-translational modification, regardless of whether the protein is derived from a prokaryotic or eukaryotic host, is the oxidation of paired cysteine residues to form disulfide bonds. Proteins can contain one or more intramolecular disulfide bonds (for a general review, see Thornton, 1981). For dimeric proteins, intermolecular disulfide bonds can occur between the subunits, as in interleukin-5 (Proudfoot et al., 1991) and macrophage colony-stimulating factor (Glocker, et al., 1993). Some dimeric proteins contain both intra- and intermolecular disulfide bonds; the best known example is insulin (Sanger, 1959). In this unit the general strategy used to determine the linkage pattern of a monomeric recombinant protein containing two intramolecular disulfide bonds is described (Basic
Characteristics of Recombinant Proteins
Contributed by Nancy D. Denslow, Keith Rose, and Pier Giorgio Righetti
7.3.1
Current Protocols in Protein Science (1996) 7.3.1-7.3.26 Copyright © 2000 by John Wiley & Sons, Inc.
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Protocol 2). The approach used is an extension of peptide mapping, where the aim is to isolate and characterize peptides containing only a single disulfide bond. In addition to the standard protein chemical approaches, isoelectric focusing in polyacrylamide gels can indicate charge heterogeneity arising from specific and nonspecific post-translational modifications. The usefulness of isoelectric focusing is increased when it is combined with electrophoresis. The resultant two-dimensional electrophoretic method, in which the protein isoelectric point is displayed as a function of pH, is described as electrophoretic titration curve analysis (Basic Protocol 3). This method is especially useful for checking for deamidation (e.g., of Asn to Asp) in which additional negative charge is introduced into the modified protein (Wingfield et al., 1987). Other modifications that can introduce charge heterogeneity include oxidation of cysteine residues, carbamylation of lysine side chains resulting from use of urea, and internal proteolytic clipping in which additional N- and C-termini are introduced (i.e., within a disulfide loop; Niekamp et al., 1969). Proteins expressed in eukaryotic systems can be subject to specific post-translational modification such as glycosylation and phosphorylation (Chapters 12 and 13, respectively), which can also introduce charge heterogeneity. Proteins produced in E. coli can also be subject to certain post-translational modifications such as acetylation of the N terminus and of the ε-side chain of lysine. These modifications will also result in charge heterogeneity (Violand et al., 1994). Electrophoretic titration curve analysis is also extremely useful for predicting suitable conditions for ion-exchange chromatography (see UNIT 6.1 for further details). BASIC PROTOCOL 1
PEPTIDE MAPPING BY HPLC The most important way to characterize a recombinant protein is to show that its primary amino acid sequence matches the predicted translation of the DNA coding sequence. In theory, one can simply digest the protein with an enzyme or chemical agent that is highly specific and compare the resulting fragments with those expected. There are computer programs that predict fragmentation patterns for proteins with all of the commonly used enzymes or chemical agents. In practice, however, the comparison can be a little tricky, because often digestions do not go to completion or peptide fragments behave differently than predicted due to slight variations in technique. This section covers protocols for two different strategies for peptide mapping. The first, based on high-performance liquid chromatography (HPLC), provides the best resolution of the digested protein and can be used in tandem with other analytical techniques to identify each fragment if this is necessary. The second method is based on high-resolution SDS-polyacrylamide gel electrophoresis (SDS-PAGE). Though less powerful than HPLC, in the sense of providing lower resolution of fragments, SDS-PAGE offers the advantage of allowing comparison to minute amounts of native protein, by using either very sensitive stains or radiolabeled protein. SDS-PAGE is also more feasible than HPLC in some laboratories.
Determining the Identity and Structure of Recombinant Proteins
Peptide mapping by HPLC is the method of choice if native material is not available for comparison. In this event, HPLC fractions must be collected for further analyses, including amino acid compositional analysis and N-terminal sequencing. New methods, recently developed, that couple peptide mapping by HPLC with mass spectrometry, are accurate enough to determine to within 0.01% the masses of the components of a digest of a recombinant protein. Tandem MS methods can even provide extensive sequence information. However, mass spectrometers may not be easily accessible and are not absolutely required for this analysis.
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Of course, the quickest way to determine the primary structure of a recombinant protein is to compare it to the native material. In this case, both the native material and the recombinant protein are digested under similar conditions and the resulting chromatograms (or SDS gels) are simply compared. Often identical chromatograms (or patterns in a gel) suffice to identify the recombinant protein in a preliminary way. However, native material is not always available, and when it is, it may be available only in minute quantities. For all of these procedures, a recombinant protein is first denatured and alkylated and is then digested with a specific protease or chemical agent (in these protocols endoproteinase Lys-C, which specifically cleaves at Lys residues, is used as an example). The resulting fragments are analyzed by HPLC (Basic Protocol 1) or by high-resolution SDS-PAGE (Alternate Protocol). Another option presented that may be convenient in some cases is to perform the protein digestion or chemical cleavage in an SDS-polyacrylamide gel slice (see Support Protocol). Of course, both methods can be readily adapted to a number of other specific proteases or chemical cleaving agents. These are discussed in the Commentary section; specific protocols are given in UNITS 11.1 & 11.4. Materials Samples of recombinant and, if possible, native protein Acetone/1 M HCl, −20°C (acidified acetone; optional) 8 M urea/50 mM Tris⋅Cl, pH 7.5 45 mM dithiothreitol (DTT) 100 mM iodoacetamide, prepared fresh and kept wrapped in foil Sequencing-grade protease (UNIT 11.1) or chemical cleavage agent (UNIT 11.4): e.g., endoproteinase Lys-C (Wako Chemicals) Digestion buffer (see recipe) HPLC solvents A and B for peptide mapping (see recipes) 50°C water bath or heating block 15- to 25-cm, 2.5-mm-i.d. C18 HPLC column (e.g., Vydac C18, 300 Å, 5-µm bead) Analytical high-performance liquid chromatography system Additional reagents and equipment for proteolytic digestion (UNIT 11.1) and/or chemical cleavage (UNIT 11.4) of proteins in solution and for digestion in SDS-polyacrylamide gel (optional; see Support Protocol) Prepare samples and controls 1. Prepare a 50- to 300-pmol protein sample and, if available, an equal amount of the native protein (as control) by lyophilizing or by precipitating from solution with 9 vol of −20°C acidified acetone. In experiments in which both the recombinant protein and the native protein are used, the amounts of both should be similar; refer to UNITS 11.1 & 11.4 for appropriate protocols. Proteolytic digests can also be done directly in a gel slice for proteins purified by gel electrophoresis; in this case steps 1 to 6 of this protocol should be replaced by gel purification and digestion in the gel slice (see Support Protocol and UNIT 11.3). In the event that the protein cannot be precipitated or lyophilized, the conditions can be adjusted by adding solid urea directly to the protein sample, then adding DTT to the appropriate concentration and incubating (see step 3), before continuing with alkylation (step 4). In this case, expect that solid urea added to 8 M concentration will double the volume of the sample.
2. Label three microcentrifuge tubes: one for the recombinant protein, one for a control consisting of enzyme (or chemical cleavage agent) alone, and one for the native protein, if available.
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Denature and alkylate protein 3. Place each sample in the appropriately labeled microcentrifuge tube, add 50 µl of 8 M urea/50 mM Tris⋅Cl, pH 7.5, and dissolve. Add 5 µl of 45 mM dithiothreitol. Incubate 15 min at 50°C. In the presence of 8 M urea and dithiothreitol, the protein sample becomes completely denatured and all disulfide bonds are reduced.
4. Add 5 µl of 100 mM iodoacetamide to alkylate cysteine residues. Incubate 15 min at room temperature in the dark. Iodoacetamide solution should be made fresh and kept protected from light. Wrapping the tube in foil works well. If a larger volume is used in step 3, the volume of iodoacetamide should be adjusted proportionately. The iodoacetamide concentration should always be at least twice the concentration of sulfhydryls, including those present in the protein and in the reducing agent, DTT, added to reduce disulfides.
Digest or chemically cleave the sample 5a. For enzymatic digestion: Add an appropriate quantity of the selected protease in digestion buffer: e.g., 140 µl digestion buffer containing 0.003 U endoproteinase Lys-C per µg protein, or ∼1:25 (w/w) enzyme/protein. Cap tube, wrap a strip of Parafilm tightly around the cap to prevent evaporation, and allow to digest overnight (∼16 hr) at 37°C. Refer to UNIT 11.1 for protocols for different proteolytic enzymes, including appropriate digestion buffers. It is important to reduce the urea concentration to ≤2 M for this step. Proteolytic enzymes are not active in higher urea concentrations because they become denatured. Most proteolytic agents will tolerate up to 10% acetonitrile as well; several published procedures include 10% acetonitrile in the digestion buffer (Fernandez et al., 1992).
5b. For chemical cleavage: Precipitate the reduced and alkylated protein with 9 vol ice-cold (−20°C) acidified acetone for 2 hr in a dry ice/ethanol bath and microcentrifuge 15 min at maximum speed, 4°C, to collect the precipitate. Solubilize the sample in the solvent appropriate for the cleavage. Specific conditions for use of CNBr, which cleaves at Met groups; BNPS-skatole, which cleaves at Trp groups; or hydroxylamine, which cleaves at Asn-Gly bonds, can be found in UNIT 11.4.
6. Microcentrifuge 15 min at maximum speed, room temperature, to pellet any particulate material that might have formed during the digestion or cleavage. The sample is now suitable for direct injection onto the HPLC column.
Perform HPLC analysis 7. Equilibrate a C18 column with HPLC solvent A, setting flow rate at 150 µl/min for a 2.1-mm-i.d. column. If a column with a wider i.d. is used (e.g., 4.6 mm), the flow rate should be adjusted in proportion to the increase in the cross-sectional area of the column. A 4.6-mm-i.d. column has roughly five times the cross-sectional area (πr2) of a 2.1-mm-i.d. column.
8. Set gradient conditions on the chromatograph. A recommended gradient protocol for peptide mapping is as follows: Determining the Identity and Structure of Recombinant Proteins
0-5 min 5-65 min 65-70 min 70-80 min
10% B 10%-60% B 60%-90% B 90% B.
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Gradient rate defines the change in the gradient over time. HPLC gradients are controlled by the pump dispensing buffer A, which can vary from 0% to 100% of the total flow. The amount of buffer A plus the amount of buffer B must equal 100%. Thus 0% B means that the B pump is off and only the A pump is dispensing buffer, i.e., 100% A; and 100% B means that only the B pump is running.
9. Set detector settings at 215 nm primary trace and 280 nm secondary trace. A single-channel HPLC apparatus is perfectly acceptable for this procedure; monitoring should be at 215 nm.
10. Begin the HPLC analysis. Perform a blank run with buffers alone, a second run with the enzyme-only (or chemical cleavage agent–only) control, and finally the run with the protein sample (with a separate run for the native-protein control, if available). Inject 100 µl of each sample to start run and do not start gradient until the sample has had time to flow onto the column. Equilibrate with HPLC buffer A between runs. A 100-ìl sample will take 0.66 min to load at a flow rate of 150 ìl/min. The injection of the sample initiates data acquisition.
11. Compare traces at all wavelengths. See Anticipated Results for a discussion of data analysis.
PEPTIDE MAPPING BY HIGH-RESOLUTION SDS-PAGE This method has less resolving power than HPLC, but allows a comparison of the recombinant product with very small amounts (in the 10 to 100 ng range) of purified native protein using sensitive stains (such as silver or gold staining) or radiolabeling as a detection method. In addition, gel electrophoresis equipment is more commonly available in laboratories.
ALTERNATE PROTOCOL
Additional Materials (also see Basic Protocol 1) 2× SDS sample buffer (UNIT 10.1) 16 × 16–cm × 1-mm or 16 × 20–cm × 1-mm Tris-tricine slab gel (UNIT 10.1) 85°C water bath or heating block XAR X-ray film or equivalent (optional; for detection of radiolabeled proteins, if used) Additional reagents and equipment for radiolabeling of proteins (UNITS 3.3; optional), proteolytic digestion (UNIT 11.1) and/or chemical cleavage (UNIT 11.4) of proteins in solution, digestion in SDS-polyacrylamide gel (optional; see Support Protocol), one-dimensional SDS-PAGE (UNIT 10.2), and detection of proteins in gels (UNIT 10.5) or on membranes (UNIT 10.8) Prepare protein sample 1. Prepare the sample as for peptide mapping by HPLC by digesting either in solution (see Basic Protocol 1, steps 1 to 5) or in SDS-polyacrylamide gel (see Support Protocol, steps 1 to 9). It is important to prepare the recombinant and native proteins in the same way. Always include an enzyme-only control. Start with 25 to 50 pmol protein if silver staining is to be used as the detection method. For Coomassie staining, start with 150 to 300 pmol protein. For more sensitive detection, proteins in the 1 to 5 pmol range can be visualized by gold staining (UNIT 10.8) or radiolabeled before digestion with 125I by the iodogen method (Markwell and Fox, 1978; UNIT 3.3).
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2. Add 1 vol of 2× SDS sample buffer to each sample. Heat 5 min at 85°C and then load into wells formed in a high-resolution Tris-tricine slab gel. For this method to work, it is important to use Tris-tricine gels, as regular Laemmli gels do not provide the required resolution of small fragments. If the sample was prepared by digestion in a gel slice (Support Protocol) and already contains some SDS, then simply calculate how much 2× SDS sample buffer needs to be added to provide a final concentration of 2% SDS. If the sample was precipitated with acetone, redissolve it in 10 ìl of 2× SDS sample buffer and add 10 ìl water.
3. Perform one-dimensional electrophoresis as indicated in UNIT 10.1. Bromphenol blue accompanies fragments in the 3-kDa range. It is wise to stop the run when the bromophenol blue band is at least 1 cm from the bottom of the gel.
4. Prepare gel for staining as recommended in UNIT 10.5 for the particular stain (silver or Coomassie blue) that is to be used. With silver staining, best results are obtained if the gel is washed with at least four changes of 50% methanol over 2 hr or equilibrated with 50% methanol overnight. Small peptides do not stain as intensely as larger peptides and may leak out of the gel. Fixing with glutaraldehyde (UNIT 10.5) may prevent this from happening and will enhance the staining obtained with silver.
5a. Detect proteins by standard methods, such as silver or Coomassie blue staining (UNIT 10.5). 5b. If proteins have previously been radiolabeled, fix and dry the gel, then detect proteins directly using a phosphoimager or by autoradiography (for 32P or 125I) or fluorography (for 3H, 14C, or 35S) with XAR X-ray film. Methods for autoradiography and fluorography can be found in Laskey and Mills (1975). SUPPORT PROTOCOL
DIGESTION OF PROTEIN IN AN SDS-POLYACRYLAMIDE GEL SLICE On occasion, it is desirable to compare a recombinant protein with authentic material after purification by SDS-PAGE. For this protocol, ∼50 pmol protein is required. The samples are subjected to SDS-PAGE in the normal manner (UNITS 10.1 & 10.4) and stained with Coomassie blue (UNIT 10.5). Bands of interest are simply cut from the gel and treated as described below; the digested fragments can then be analyzed by HPLC (Basic Protocol 1) or by high-resolution gel electrophoresis (Alternate Protocol). Additional Materials (also see Basic Protocol 1) 10 mM Tris⋅Cl (pH 7.5)/0.1% (w/v) SDS 1.5-ml Kontes microcentrifuge tube and pestle, or equivalent 1.5-ml low-protein-binding microfilterfuge tube (0.2-µm cellulose acetate; Rainin or Costar) Dry ice/ethanol bath (optional) Additional reagents and equipment for one- or two-dimensional SDS-PAGE (UNITS 10.1 & 10.4), staining with Coomassie blue (UNIT 10.5), and proteolytic digestion of proteins in gels (UNIT 11.3)
Determining the Identity and Structure of Recombinant Proteins
Gel purify the protein of interest 1. Load a sample containing ≥50 pmol protein onto an SDS-polyacrylamide gel and perform one- or two-dimensional SDS-PAGE (UNITS 10.1 & 10.4), then stain with Coomassie blue and destain in standard fashion (UNIT 10.5). The following list of equivalences is helpful for determining the required amount of protein: for a 100-kDa protein, 100 ìg = 1 nmol; for a 50-kDa protein, 50 ìg = 1 nmol;
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and for 25-kDa protein, 25 ug = 1 nmol. Because proteins are reduced and denatured in SDS sample buffer, it is not generally necessary to further treat the protein with reducing or alkylating agents. However, if there is concern about disulfides, the protein can be reduced and alkylated prior to electrophoresis.
2. Wash the gel slice in water, using four changes in 1 hr with constant shaking, to remove as much of the destaining solution as possible. 3. Dry the gel slice in air for 15 min. Digest or chemically cleave the sample 4. Rehydrate the gel slice with digestion buffer containing an appropriate amount of the selected enzyme or chemical cleavage agent: e.g., 140 µl digestion buffer containing 0.003 U endoproteinase Lys-C per µg protein, or ∼1:25 (w/w) enzyme to protein. Refer to UNITS 11.1 & 11.3 for protocols for different proteolytic enzymes, including appropriate digestion buffers. This methodology can also be used to cleave proteins using chemical agents such as CNBr, which cleaves at Met groups; BNPS-skatole, which cleaves at Trp groups; or hydroxylamine, which cleaves at Asn-Gly bonds. Procedures for using these agents can be found in UNIT 11.4.
5. Macerate the gel with a tight-fitting pestle (a Kontes microcentrifuge tube and pestle are ideal). Carefully rinse the pestle with an additional 50 µl digestion buffer. 6. Cap the tube, wrap a strip of Parafilm tightly around the cap to prevent evaporation, and allow to digest overnight (∼16 hr) at 37°C. 7. After digestion, separate the protein fragments from the gel pieces by centrifuging 5 min at ∼2000 to 4000 × g (see manufacturers’ recommendations) through a 1.5-ml low-protein-binding microfilterfuge tube. An efficient way to transfer the gel mixture to a microfilterfuge tube is to simply cut the bottom off the microcentrifuge tube containing the digest with a microcentrifuge cutter (Fisher) and assemble onto a microfilterfuge tube. The whole unit (cut microcentrifuge tube and microfilterfuge tube) can be stabilized by assembling it inside a 10-ml round plastic disposable tube. The unit is then centrifuged 5 min at low to medium speed (e.g., ∼2000 to 4000 × g; see manufacturers’ recommendations) in a tabletop centrifuge to allow all gel pieces to flow into the top compartment of the microfilterfuge. The use of a micropipet to transfer the mashed gel pieces to the microfilterfuge is not recommended. The use of a micropipet is not recommended because gel pieces will adhere to the inside of the pipet tip, reducing fragment recovery. Extracting the protein fragments from the gel is critical for good resolution of fragments by high-resolution gel electrophoresis.
8. Reextract the gel pieces with 100 µl of 10 mM Tris⋅Cl (pH 7.5)/0.1% SDS for 20 min at room temperature and then recentrifuge through the microfilterfuge tube. 9. Combine the extracts (containing the peptide fragments). If samples are to be analyzed by high-resolution SDS-PAGE, these extracts can be applied directly to the gel and steps 10 to 12 of the present protocol should be omitted; instead, proceed directly to electrophoresis (see Alternate Protocol, step 2).
Acetone precipitate samples that are to be analyzed by HPLC 10. Add 9 vol of −20°C acidified acetone to sample. Allow to precipitate overnight at −20°C or 2 hr in a dry ice/ethanol bath. 11. Microcentrifuge sample 15 min at maximum speed, 4°C. Carefully decant supernatant from the pellet.
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12. Dry the pellet in a Speedvac to remove all traces of acetone. Resuspend pellet in HPLC buffer A and proceed to separation by HPLC (see Basic Protocol 1, step 7). Acetone precipitation is necessary to remove excess SDS present when digestion is performed in a gel slice. SDS levels should be reduced below 0.05% before the sample is applied to the HPLC column. BASIC PROTOCOL 2
DETERMINING THE DISULFIDE LINKAGE PATTERN OF A RECOMBINANT PROTEIN Correct formation of the disulfide linkages of a recombinant protein is at least as important for protein function as correct amino acid sequence, but can sometimes be more difficult to establish. First, the sample is treated to alkylate any free thiol groups so as to block their ability to catalyze disulfide interchange (“scrambling”); this is done using denaturing but nonreducing conditions to expose any buried thiols. In the next, key step the protein is digested by chemical or enzymatic means in such a way as to cleave the polypeptide chain at least once between successive Cys residues (except where two consecutive Cys residues share a disulfide bond, when this is not necessary). Resulting fragments containing only a single disulfide bond are then isolated by reversed-phase high-performance liquid chromatography (RP-HPLC) and identified by a combination of mass spectrometry and Edman analysis; each bond will either be holding two small peptide chains together, or holding a single polypeptide chain in a loop. This apparently simple task is often frustrated by the presence of closely spaced Cys residues (because it is necessary to cleave between them) and by the difficulty in ensuring that disulfide scrambling does not occur. Given the huge variety of possible protein properties, it is impossible to give a single recipe for determining disulfide linkage patterns that can be blindly followed in all cases. The protocol that follows takes protein idiosyncracies into account as much as possible in the annotations which follow (or sometimes precede) critical steps; the proteins granulocyte/macrophage colony-stimulating factor (GM-CSF) and macrophage colonystimulating factor (M-CSF) are discussed as examples. NOTE: All reagents used should be of analytical grade or better. Solutions should be made fresh except where otherwise stated (see Reagents and Solutions). Special precautions are necessary when working with iodoacetic acid, concentrated formic acid, cyanogen bromide, and trifluoroacetic acid (TFA): these precautions are described at the points where use of these chemicals is described. Materials Sample of recombinant protein Alkylation buffer (see recipe) Nitrogen or argon, oxygen-free Guanidine hydrochloride 50 mM and 2 M ammonium bicarbonate 2 mg/ml cyanogen bromide in 70% formic acid (see recipe) HPLC solvents A and B for disulfide linkage analysis (see recipe) 1% formic acid Proteases: pepsin, Staphylococcus aureus V8 protease (endoproteinase Glu-C), and/or trypsin (porcine variety, Sigma) Glacial acetic acid 1 M dithiothreitol (DTT)
Determining the Identity and Structure of Recombinant Proteins
Pretreated dialysis tubing with appropriate MWCO (see recipe) Apparatus for reversed-phase high-pressure liquid chromatography (RP-HPLC), preferably with automatic sample injector
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Appropriate analytical HPLC column: e.g., 25 cm × 4 mm i.d. filled with Nucleosil 5 µm, 300 Å C8 particles (Macherey-Nagel) Appropriate preparative HPLC column: e.g., 25 cm × 4 cm i.d. filled with Nucleosil 5 µm (as above) for purifying up to ∼0.2 mg protein, or 25 cm × 10 mm i.d. with same packing for up to ∼1.5 mg protein (see Critical Parameters for further guidance) Vacuum centrifuge: e.g., Speedvac (Savant) Equipment for mass spectrometry of peptides Additional reagents and equipment for dialysis (UNIT 4.4; APPENDIX 3B) Block free thiols 1. Dissolve the protein to ∼0.1 mM in alkylation buffer and incubate 30 min at room temperature (∼22°C) in the dark under nitrogen or argon. Use 0.2 to 2 mg protein (depending on availability) for steps 1 to 3 in the first instance, and repeat on a larger scale if necessary. This step serves to alkylate any easily accessible thiol groups. Radiolabeled alkylating reagent (e.g., [14C]iodoacetic acid) may be used to facilitate the later location of modified peptide.
2. Add solid guanidine hydrochloride to a final concentration of 6 M and continue incubation for a further 60 min. At this stage any previously buried thiols should become alkylated. Do not add thiol at the end of the incubation to consume excess alkylating agent in the normal fashion, as disulfides would then be at risk. It may be necessary to add the guanidine earlier on in order to solubilize the protein, in which case the 30-min preincubation should be omitted.
3. Transfer the sample to a prepared dialysis bag. Dialyze the sample (UNIT 4.4; APPENDIX 3B) in the dark at 4°C against 50 mM ammonium bicarbonate, using a bath volume ≥2000 times greater than the sample volume and stirring the bath magnetically. Change the solution after 1 hr, 2 hr, and 4 hr, and continue dialysis overnight (∼18 hr total). Recover the protein by lyophilization. IMPORTANT NOTE: Take great care not to damage the dialysis bag. A plastic automatic pipet tip is preferred for the transfer, but if a glass Pasteur pipet is used, remove sharp edges by heating it in a flame prior to taking up sample; avoid damage due to sharp fingernails, especially when knotting the bag. Proteins known not to precipitate on removal of chaotrope can be recovered more quickly by gel filtration than by dialysis. Furthermore, steps 1 to 3 are usually omitted when it is known from attempted reaction with thiol-specific reagents or radiolabeled reagents that there are no unpaired Cys residues. Whenever there is reason to suspect the presence of an unpaired Cys—e.g., an odd number of Cys residues in the sequence, reaction with thiol-specific reagents (Ellman, 1959; Thannhauser et al., 1984) or radiolabeled reagents —these blocking steps must precede the fragmentation procedures. For example, blocking was necessary for the protein G-CSF (Wingfield et al., 1988) but not for GM-CSF (Schrimsher et al., 1987).
Fragment the polypeptide chains 4. Depending on the known or expected amino acid sequence, select digestion conditions (cyanogen bromide, one or more proteolytic enzymes, or a combination of these in succession) that will cleave the polypeptide chain at least once between successive Cys residues (see Critical Parameters for guidance). Perform an appropriate combination of the treatments described in steps 5 to 8. If the protein is known to contain a few well-spaced Met residues, cyanogen bromide is often useful as a first fragmentation step.
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For cyanogen bromide treatment 5. Dissolve the recombinant protein at 1 mg/ml in 2 mg/ml cyanogen bromide/70% formic acid. Seal the tube and incubate 24 hr in the dark at room temperature. CAUTION: Cyanogen bromide is toxic and volatile; concentrated formic acid is very corrosive (see recipes for detailed precautions).
6a. If the complexity of the analysis is high (i.e., a large protein and/or many disulfide bridges are involved) and the expected sequence suggests the problem can be simplified by working individually with the cyanogen bromide fragments, dilute the cyanogen bromide digest at least three-fold with HPLC solvent A and separate by RP-HPLC (steps 9 and 10), then proceed to subdigest the large fragments enzymatically. 6b. Otherwise, simply remove solvent and reagent under high vacuum in a vacuum centrifuge (avoid exposing other samples in the vacuum centrifuge to reagent vapors), then proceed to enzymatic digestion. Here and throughout the protocol, use the vacuum centrifuge with the heater switched off to lessen the risk of such reactions as deamidation, oxidation, and N-terminal cyclization.
For protease treatment 7a. Using pepsin: Dissolve the protein or protein fragments at ∼1 mg/ml in 1% (v/v) formic acid, add pepsin to obtain a 1/100 (w/w) enzyme/substrate ratio, and incubate 16 to 24 hr at room temperature. Pepsin was useful for G-CSF, although trypsin or V8 protease were more useful for GM-CSF (Schrimsher et al., 1987).
7b. Using trypsin or V8 protease: Dissolve the protein or protein fragment at ∼1 mg/ml in 0.1 M phosphate buffer, pH 7.0. Add enzyme to obtain a 1/100 (w/w) enzyme/substrate ratio, and incubate 4 hr at 37°C. When using trypsin, the porcine variety (Sigma) is preferred over the bovine enzyme as it retains its activity at lower pH.
8. If trypsin or V8 protease is used for digestion at neutral or slightly alkaline pH, acidify the digest to pH 3 with acetic acid just prior to HPLC (step 9) to stop the digestion and prepare the sample to be placed in the automatic injector. A peptic digest is already acidic and is injected directly; if it sits in the automatic injector, the digestion will continue.
Analyze digest by RP-HPLC 9. Subject the cyanogen bromide or enzyme digest to analytical RP-HPLC—e.g., using the trifluoroacetic acid/acetonitrile RP-HPLC system (UNIT 11.6), which is very convenient because both solvents are completely volatile. Use a gradient of 2% HPLC solvent B/min up to 100% B. Monitor at low wavelength (e.g., 214 nm), as many fractions may not contain aromatic residues.
Determining the Identity and Structure of Recombinant Proteins
Perform preparative HPLC 10. Perform preparative HPLC separations on digests that showed a suitable number of well-separated fragments in step 9. For best results, use a much shallower gradient (e.g., 0.2% solvent B/min) over the percentages of interest deduced from the screening run. For further discussion of the design of the preparative gradient, see, for example, Rose et al. (1992).
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Particularly for preparative HPLC runs, it is preferable to acidify the digest (if not already acid) to pH 3 with acetic acid prior to injection (check by spotting onto pH paper), as otherwise the early part of the chromatogram may be compromised. Ideally, choose column dimensions and flow rate compatible with the quantity of digested protein (see Critical Parameters). If several milligram of protein are available, it makes life much easier to work at the 1.5-mg scale, but remember that it may sometimes be necessary to return to step 7 and continue the analysis with a different digest.
11. Recover peptides by drying the collected fractions in the vacuum centrifuge, then redissolve each in 200 µl HPLC solvent A. Divide each fraction, placing two analytical-sized aliquots (10 µl each if a major peak and >0.5 mg digest were used; 30 µl if a minor peak and <0.5 mg digest were used) directly into the automatic injector sample tubes, and retaining the remainder separately. The original chromatogram obtained during collection of the corresponding preparative fraction should be used to select an appropriate sample volume and attenuation level for the analytical runs that will follow.
Locate fractions containing disulfide bonds 12. To one sample add 2 M ammonium bicarbonate to 50 mM final and 1 M DTT to 50 mM final. To the second sample (control) add ammonium bicarbonate but no DTT. Close tubes and incubate 2 hr at 37°C. Steps 12 and 13 constitute the simplest of several possible ways of locating fragments containing a disulfide bond: attempted reduction of disulfide bonds in each HPLC fraction with DTT followed by reinjection on a shallow gradient. To perform this assay on a dry sample, the peptide can simply be dissolved in 50 mM ammonium bicarbonate/50 mM DTT.
13. Acidify the samples with a few microliters of acetic acid (test a blank with pH paper), agitate to liberate the carbon dioxide bubbles, then analyze the samples in pairs (reduced and nonreduced) by RP-HPLC, using the analytical column but with the shallow gradient used for the preparative run (step 10). As mentioned in step 11, the original chromatogram obtained during collection of the corresponding preparative fraction, and the proportion taken for analysis, permit an appropriate choice of attenuation to be made for these analytical runs.
14. Interpret the pairs of chromatograms (attempted reduction and control). When all has gone well, if no disulfide bond is present there should be essentially no difference between attempted reduction and control. If a disulfide bond is present, then either this bond is holding two small peptide chains together, or it is holding a single polypeptide chain in a loop. In the former case, two peaks should be visible on reduction; in the latter case all that will be observed is a small shift to later retention time (opening of the loop exposes more of the peptide chain to the stationary phase). The reason for performing a control incubation in the absence of reductant is that some fractions may shift retention time or develop structure due to changes such as opening of a homoserine lactone, cyclization of an N-terminal Gln, or oxidation of methionine to its sulfoxide, and this change might otherwise be mistaken for reduction. Other sources of difficulty in interpreting results include the following: One of the components liberated by reduction of a disulfide bond may elute early and be difficult to detect because of the presence of salt and DTT peaks. One component may be short and devoid of aromatic residues and so be much less visible on the chromatogram. Finally, because only one HPLC dimension was used for the initial collection, there may be a nonreducible component present in addition to a reducible one; this need not be a problem when both Edman and mass spectrometric techniques are used for characterization.
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Characterize Cys-containing peptides 15. For each fraction identified as containing a disulfide bond, repeat the reduction (step 12) with an appropriate portion of the remaining nonreduced fraction (set aside in step 11), repeat the HPLC separation, and collect the reduced components. For an example of where this repeated reduction and separation strategy was used, see Schrimsher et al. (1987). To determine sample size, the mass spectroscopist who will perform that analysis should be consulted regarding how much protein will be required.
16. Divide the fractions and submit these reduced components, together with some of the unreduced fraction, for examination both by mass spectrometry and by Edman degradation. The combination of mass spectrometry and Edman degradation provides the quickest and surest approach to characterizing the fractions containing disulfide bonds. Mass spectrometry provides the molecular weight of the peptide component (or components) present, while Edman analysis should give their N-terminal sequences. The expense and sophistication of mass spectrometric equipment is such that usually, where it exists, a competent person is present to give advice on its use. Electrospray ionization mass spectrometry (ESI-MS) is most suited to this application, although good results may also be obtained by fast-atom-bombardment (liquid secondary-ion) mass spectrometry if a high-performance machine is available. Matrix-assisted laser-desorption ionization time-of-flight (MALDI-TOF) mass spectrometry may also be used (see UNIT 11.6); mass precision is generally lower than with fast-atom-bombardment or ESI spectrometry, but can be adequate if the time-of-flight analyzer is operated in reflectron mode and carefully calibrated (using an internal standard), and is excellent if the MALDI source is connected to a Fourier-transform ion cyclotron resonance analyzer. Some peptide components may not be susceptible to Edman degradation. A peptide may be N-terminally blocked (e.g., by acetyl for the N-terminal peptide of the protein, or by a residue of pyroglutamic acid if a Gln residue liberated during proteolysis has cyclized). This problem is discussed in Critical Parameters and Troubleshooting.
17. Align the peptides identified in step 16 with the known sequence of the recombinant protein. Interpret the molecular weights determined quite precisely by mass spectrometry. Even if the Edman degradation did not provide sequence information up to the C terminus of a given peptide, the N-terminal starting point should be unambiguous: the C terminus can then be deduced from the mass of the peptide. Taken together, the Edman and mass spectrometry data generally allow the disulfide bonding pattern of the component peptides to be deduced, and hence the bonding pattern of the recombinant protein. Even where this is not the case, sufficient information is gained to greatly facilitate a second attempt. In some cases, the fragments identified indicate neither a single disulfide bond in a loop nor two short chains held together by one disulfide bond. In such cases (e.g., where two chains are both linked to a third by disulfide bonds; Rose et al., 1992), further cycles of fragmentation, isolation, and identification must be performed in order to cleave between the Cys residues. Scintillation counting of fractions permits rapid location of any Cys residues radioalkylated in Step 1. Characterization of any such fractions by Edman degradation and mass spectrometry completes linkage assignment by proving that the Cys residues so alkylated could not have been involved in disulfide bonds.
Determining the Identity and Structure of Recombinant Proteins
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Current Protocols in Protein Science
DETERMINING CHARGE HETEROGENEITY BY TWO-DIMENSIONAL TITRATION CURVE ANALYSIS
BASIC PROTOCOL 3
Unfolding of proteins can be monitored by running the sample across a urea gradient—the resulting denaturation curve, often sigmoidal in shape, represents the transition from the folded to the unfolded state. In general, pH/mobility curves of proteins are generated in a polyacrylamide IEF slab gel (see Fig. 7.3.1 for an outline of the procedure). Materials Acrylamide/bisacrylamide stock solution (see recipe) Carrier ampholytes: e.g., Pharmalyte (Pharmacia Biotech), Bio-Lyte (Bio-Rad), or Servalyte (Serva) 10% (w/v) ammonium persulfate stock solution (see recipe) N,N,N′,N′-tetramethylethylene diamine (TEMED) Kerosene Electrode solutions (see recipe) Sample of recombinant protein in aqueous solution containing ≤10 mM Tris⋅acetate (or other salt or buffer consisting of weak acids and bases) Coomassie blue staining solution: 0.1% (w/v) Coomassie brilliant blue R-250/25% (v/v) ethanol/8% (v/v) acetic acid Destaining solution: 25% (v/v) ethanol/8% (v/v) acetic acid Gel Bond PAG sheets Electrophoresis unit for horizontal slab gel operation: e.g., Multiphor (Pharmacia Biotech) Levelling table IEF electrode filter paper strips (Pharmacia Biotech)
A
B –
+
C
D
–
+ 9 7 pH 5 3
Figure 7.3.1 Experimental procedure for generating titration curves by isoelectric focusing. First the pH gradient is formed by focusing the carrier ampholyte mixture without sample (first-dimension run; A); the electrode strips and the gel layer underneath are then removed (B); the sample is applied in the trench now perpendicular to the pH gradient (C); the second-dimension run is performed perpendicular to the first dimension axis (D). From Righetti et al. (1979).
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Constant-power supply (up to 3000 V, 200 mA, 20 W) Thermostatic water bath: e.g., MultiTemp (Pharmacia Biotech) Titration curve kit (Pharmacia Biotech) Additional reagents and equipment for denaturing IEF in slab gels (UNIT 10.2) Cast the gel 1. Cut a Gel Bond PAG sheet in half (to 12.5 × 12.5 cm) to form a gel-supporting foil. Roll the supporting foil on the glass plate of the electrophoresis unit, putting a few drops of water on the back to secure adhesion. The hydrophilic surface of the foil should face the operator; this is the surface designed for adhesion of the gelling layer.
2. Complete the cassette by placing the glass plate–foil sandwich between the two marks on top of the slot former and fastening them together using four clamps. Lay out the completed cassette horizontally, preferably on a levelling table. 3. Prepare 28 ml of an appropriate gel solution—e.g., containing 5%T and 4%C monomers and 2% ampholytes—from acrylamide/bisacrylamide stock solution and carrier ampholytes (see UNIT 10.2 for details of gel casting). Degas for 5 to 10 min. Preparing 28 ml of solution provides a slight excess, as 25 ml will be needed to fill up the gel cassette.
4. Add 4 µl ammonium persulfate stock solution and 1 µl pure TEMED per milliliter of gelling solution; mix carefully. Immediately inject the gel solution between the slot-former and the upper glass plate (the solution will fill the cassette by capillary action). Let stand while polymerization occurs (this will be complete after 1 hr). IMPORTANT NOTE: When titration curves are run under denaturing conditions (in 8 M urea), the gel solution will polymerize very quickly. High temperatures will also greatly accelerate polymerization. If polymerization occurs too quickly, there may not even be sufficient time to pour out the liquid from the flask. If this occurs, repeat the procedure and this time precool the gelling liquid to ∼10°C.
Perform first-dimension electrophoresis 5. Remove the clamps and separate the glass plate from the Gel Bond PAG sheet with a gentle twist of a spatula. Holding the cassette upside down, use the spatula to introduce air bubbles between the gel and slot former. 6. Starting from a corner, remove the gel from the plastic cover containing the slot former. Spread a few drops of kerosene onto the cooling block of the Multiphor chamber (set at 10°C) to provide good adhesion, and then gently lower the gel onto the cooling surface. Any trapped air bubbles should be eliminated, as their presence will lead to the formation of hot spots that will distort the pH gradient.
7. Prepare IEF electrode filter paper strips by cutting them to the length of the gel. Wet in electrode solutions and blot excess solution with a paper tissue. Apply strips to gel surface. For good performance, the electrode strips and platinum wires should all be parallel to one another and the sample trench should be perpendicular. Refer to Figure 7.3.1 for this and the following operation. Determining the Identity and Structure of Recombinant Proteins
8. Perform first-dimension electrophoresis at 10 W constant power with voltage and current set to a maximum (typically, at steady state, the voltage should be ∼1000 V). This focusing step is usually terminated in 50 min.
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Perform second-dimension electrophoresis 9. Remove the electrode strips and use a long blade to cut away the strips of gel underlying the electrode strips (see Fig. 7.3.1B). This step is important, as otherwise considerable heating will occur in these regions due to high conductivity.
10. Turn the gel around 90° on the cooling plate. Prepare and apply new electrode strips, using the same electrode solutions as for the first dimension. The sample trench is now parallel to the electrode strips (see Fig. 7.3.1). 11. Apply the sample by filling the trench with ∼100 µl of protein solution (typically 100 to 400 µg protein, depending on sample heterogeneity, at 1 to 4 mg/ml; see Fig. 7.3.1C). 12. Perform second-dimension electrophoresis at 600 V for 5 to 20 min (based on sample mobility). High levels of salts, as well as salts and buffers comprising strong acids and bases, must be strictly avoided in any focusing process. For best results, the sample should not contain more than 10 mM salt or buffer, and these should consist of weak acids and bases—e.g., Tris-acetate, but not Tris⋅Cl or sodium acetate.
Stain and destain gel 13. Place the gel in Coomassie blue staining solution and allow to stain 1 hr with shaking. Staining solution can be reused at least 10 times. Store at room temperature protected from light.
14. Replace the staining solution with destaining solution. Destain with shaking, changing solution several times until a clear background is seen. Destaining solution can also be stored at room temperature and reused several times if filtered through a charcoal cartridge after each use. Beware of excessive evaporation of both ethanol and acetic acid, which will weaken the destaining process. If the gel is over-destained, the titration curve zones will also fade away. In this case, it can be restained and destained. When too little sample is available to be visualized in this fashion, any variant of silver staining may be used (Rabilloud, 1990; UNIT 10.5). In this last case, the amount of sample can be 20 times smaller. For best results, however, the gel slab should be thinner (e.g., 0.5 mm thick).
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Acrylamide/bisacrylamide stock solution Dissolve 28.7 g acrylamide and 1.3 g N,N′-methylene bisacrylamide in distilled water to a final volume of 100 ml (30%T, 4%C final). Store up to 3 weeks in the dark at 4°C. Alkylation buffer, pH 8.0 100 mM Tris⋅Cl adjusted to pH 8.0 with 6 M HCl or glacial acetic acid 125 mM NaCl 5 mM EDTA 4 mM iodoacetic acid Prepare fresh just before use Concentrated stock solutions of EDTA (sodium salt; APPENDIX 2E) and iodoacetic acid in water may be made up and kept at −20°C (stable at least 1 month). Iodoacetic acid should continued
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be bought in small amounts and should take the form of white crystals. The solid and its solutions are light-sensitive and should therefore be shielded with aluminum foil and kept protected from direct sunlight. CAUTION: Solid iodoacetic acid and its concentrated solutions are corrosive—wear gloves when handling these materials.
Ammonium persulfate, 10% (w/v) Dissolve 50 mg ammonium persulfate in 0.5 ml distilled water. Prepare fresh each time. Cyanogen bromide, 2 mg/ml in 70% formic acid Weigh an empty open small glass tube or vial together with its cap, and then move this container to a chemical fume hood. Add a few small crystals of cyanogen bromide, replace the cap, and then reweigh the container to calculate the amount of cyanogen bromide added. Dilute with sufficient 70% formic acid to obtain a 2 mg/ml solution. (This procedure avoids the need to use a balance inside the chemical fume hood.) Prepare fresh. CAUTION: Cyanogen bromide, though a solid at room temperature, is quite volatile and sufficiently toxic to have been used as a chemical warfare agent (albeit a rather ineffective one). It should be manipulated in a chemical fume hood and gloves should be worn when handling. Buy good-quality reagent, keep solid protected from light at 4°C to minimize degradation, and make up fresh solution as required, as otherwise contaminating free bromine will tend to oxidize Met and Trp residues, which can result in cleavage at Trp and failure to cleave at Met.
Dialysis tubing, pretreated Obtain dialysis tubing with appropriate MWCO (molecular weight cut-off) for the protein under study—e.g., Visking 18/32, which has a cut-off after boiling of <5000—and prepare an appropriate length of tubing fresh before use by treating it as follows to remove glycerol and residues from manufacture. Bring 1% sodium bicarbonate solution to a simmer in a beaker which has no chips or cracks (as these could pierce the tubing). Place the tubing in the solution and simmer 10 min; do not boil vigorously or the tubing will protrude from the solution and become brittle. Cool by adding distilled water, then thoroughly rinse the tubing inside and out with distilled water. Excess prepared tubing can be stored in water or buffer for a few days at 4°C if precautions are taken to avoid microbial growth (e.g., add sodium azide to 0.005%). IMPORTANT NOTE: Throughout tubing preparation, beware of sharp edges, including fingernails and especially pipets, which can easily damage the tube and lead to complete loss of sample during the experiment.
Digestion buffer 10 mM Tris⋅Cl, pH 8.0 0.1 mM EDTA 0.02 M NH4HCO3 0.1% SDS (include only if protein is to be digested in a gel slice) Prepare fresh Different proteolytic enzymes and chemical cleavage agents require different digestion buffers; refer to UNITS 11.1 & 11.4 for a discussion of these requirements. Determining the Identity and Structure of Recombinant Proteins
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Electrode solutions Use 50 mM solutions of either weak acids (e.g., acetic acid) or weak bases (e.g., Tris) as anolyte and catholyte, respectively (strong acids and bases should be avoided). Alternatively, use 50 mM solutions of isoionic zwitterions (e.g., Asp, pI 2.77, or Glu, pI 3.22) as anolyte and 50 mM Lys (pI 9.74) or 50 mM Arg (pI 10.76) as catholyte. Store in cold room to avoid bacterial growth. Keep basic solutions tightly capped and check the pH before use. If the pH decreases ≥0.5 pH units (due to CO2 absorption), discard the solution. HPLC solvents for disulfide linkage analysis RP-HPLC solvent A: Add 1 g HPLC-grade trifluoroacetic acid (TFA; Pierce or ABI) to 1 liter of HPLC-grade water (e.g., purified using a Milli-Q system, Millipore) and mix. Degas either by vacuum filtration in an all-glass device (Millipore) or by sparging with helium. The solution is good for at least 3 months when stored at room temperature, protected from dust, and degassed prior to each use. HPLC solvent B: Add 1 g TFA to 100 ml HPLC-grade water and mix. Dilute to 1 liter with HPLC-gradient-grade acetonitrile. Degas as for solvent A except, if using vacuum filtration, employ a solvent-resistant filter membrane (e.g., Millipore). The solution is good for at least 3 months when stored at room temperature, protected from dust, and degassed prior to each use. CAUTION: Undiluted TFA is very corrosive: wear eye protection (preferably a face shield) and gloves. Once diluted to 1 g/liter, it presents no danger.
HPLC solvents for peptide mapping HPLC solvent A: 0.1% (v/v) TFA in water HPLC solvent B: 0.07% (v/v) TFA/80% acetonitrile Filter and degas both solvents. For best results, balance absorption of solvents A and B at 215 nm by adding TFA until absorption values are equal. Store up to 1 week at room temperature. COMMENTARY Background Information Peptide mapping The number of different digests required to characterize a recombinant protein depends on the analytical method used and on the quality of information produced. Analysis by high-performance liquid chromatography (HPLC; Basic Protocol 1) allows the use of proteolytic agents that cut proteins generally into small fragments (such as trypsin, which cuts at Arg and Lys residues; endoproteinase Lys-C, specific for Lys residues; or endoproteinase Asp-N, specific for Asp residues). Analysis by SDSpolyacrylamide gel electrophoresis (SDSPAGE; Alternate Protocol), on the other hand, requires larger fragments (≥2 kDa in size) and thus is limited to agents that cut the protein of interest at amino acid residues that are less frequent (such as CNBr or BNPS-skatole, which cut at Met and Trp residues, respectively) or to partial digests with proteolytic enzymes (such as endoproteinases Lys-C or Asp-N,
which cut at Lys and Asp residues, respectively). These proteolytic agents have very strict specificities and easily predicted patterns of digestion. Other proteases have rather broad specificities (e.g., chymotrypsin is specific for Tyr, Phe, and Trp and to a lesser extent for Leu, Met, Ala, Asp, and Glu) and therefore produce a large number of fragments. These proteases are ideal for comparing a recombinant protein to the native protein because many more sites can be probed with only one digestion. Still other proteases, such as endoproteinase Glu-C (otherwise known as Staphylococcus aureus V8 protease), recognize one or two sites (in this case Glu and Asp) depending on the buffer conditions used for the digestion. For a useful discussion of these enzymes, see Allen (1989) and UNIT 11.1. For recombinant proteins, the investigator has the advantage of knowing the predicted amino acid sequence and can design a specific strategy based on the distribution of key amino acids that define chemical and protease cleav-
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age sites. Several computer programs predict cut sites, HPLC elution patterns, and molecular weights of fragments. The Peptide Sort program from Genetics Computer Group (GCG; University of Wisconsin, Madison, Wis.) is one such program. Because many cleavage agents give rise to a combination of complete and partial digests, it is important to treat the recombinant protein and the native isolated protein (if available) under the same conditions. If it is impossible to obtain enough native material for comparison by the methods described above, there are a number of analytical techniques that can be used to prove the recombinant protein has the correct structure. The easiest and most cost-effective method is to analyze individual peaks from an HPLC run for their amino acid compositions; this can be compared to the predicted composition for each fragment. This method is most powerful when combined with N-terminal amino acid sequencing, which can identify each fragment collected by HPLC. The intact protein can also be submitted for N-terminal sequencing. With new instruments, it should be possible to identify as many as 50 to 100 residues directly from the N-terminus. On several occasions it has been possible to sequence up to 50% of a protein from a single digest (N. Denslow, unpub. observ.). Newer analytical methods rely on mass spectrometry to analyze the digest of recombinant protein (Langen et al., 1993; Patterson, 1994). The mass accuracy of mass spectrometers is sufficient to identify a fragment directly. The preferred technique relies on the use of electrospray ionization (ESI)–quadrupole mass spectrometers, especially if the mass spectrometer is in-line with the HPLC. ESI-MS has a mass accuracy of 0.01%. Accurate data, however, can also be obtained from a matrix-assisted laser-desorption ionization time-offlight (MALDI-TOF) mass spectrometer, which typically has a mass accuracy of 0.03%. In this case, HPLC fractions must be collected and mixed with a matrix for analysis.
Determining the Identity and Structure of Recombinant Proteins
Disulfide linkage analysis Correct formation of the disulfide linkages of a recombinant protein is at least as important for function as correct amino acid sequence, but (as can be appreciated from the complexity of Basic Protocol 2) it is sometimes more difficult to establish. The key to success generally involves digestion of the protein by chemical or enzymatic means in such a way as to cleave the polypeptide chain at least once between succes-
sive Cys residues—except where two consecutive Cys residues share a disulfide bond, when this is not necessary. This apparently simple task is often frustrated by the presence of closely spaced Cys residues, because it is not always easy to cleave between these. Some instructive examples showing the difficulties encountered when two Cys residues are very close or juxtaposed are discussed by Morris and Pucci (1985); Hojrup and Magnusson (1987); and Rose et al. (1992). Cleavage techniques are somewhat limited by the need to prevent disulfide scrambling (Hojrup and Magnusson, 1987; Rose et al., 1992). Scrambling can occur in the presence of strong mineral acid, and also in neutral to basic medium, which is why mildly acidic conditions are preferred where possible. Scrambling is catalyzed by the presence of free thiol; for this reason any unpaired Cys residues (which may be present in a small fraction of improperly refolded material) must be blocked in the first step. It must be emphasized that people working with a recombinant protein usually have two enormous advantages over those working with a rare polypeptide isolated from natural sources—they have much more material, and they know most, if not all, of the amino acid sequence. These advantages must be exploited to the full: it is advisable to use trial amounts, change tactics if solubility problems are insurmountable, and design the fragmentation strategy around the known sequence. The method described is designed to give results as reliably and rapidly as possible with recombinant proteins, and would need extensive modification to be suitable for dealing with trace amounts of proteins derived from natural sources. Titration curve analysis Protein titration curves (in reality, pH/mobility curves) became suddenly popular in the late seventies after an original report by Rosengren et al. (1977), who demonstrated a simple method to force a protein to migrate across a quasi-stationary pH gradient in such a way as to trace a sigmoidal path resembling a classical potentiometric titration curve. These pH/mobility curves were found to be quite useful in a number of applications: (1) for screening of genetic mutants (Righetti et al., 1978a; Righetti et al., 1979); (2) for studying macromoleculeligand interactions (Krishnamoorthy et al., 1978; Constans et al., 1980); (3) for demonstrating macromolecule-macromolecule interactions (Righetti et al., 1978b; Lostanlen et al., 1980; Gianazza et al., 1980); (4) for direct pK
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determination of simple cations and anions and of uni-uni-valent amphoteric molecules (Righetti et al., 1979; Valentini et al., 1980); (5) for determining dissociation constants (Kd) of ligands and proteins and their pH dependence (Ek et al., 1980; Ek and Righetti, 1980); (6) for detecting protein and peptide microheterogeneity (Arnaud et al., 1980; Henner and Sitrin, 1984; Picard et al., 1987; Van Den Oetelaar and Hoenders, 1987); and (7) for choosing optimal pH conditions in ion-exchange chromatography (Fägerstam et al., 1983). The times must have been ripe for such an idea, because at just about the same time, Creighton (1979) reported a method for studying the unfolding of proteins by running them across a urea gradient: these denaturation curves, often sigmoidal in shape, represent the transition from the folded to the unfolded state. In an analogous manner, it is possible to study the melting profiles of DNA (from double helix to single strand) by running them across a thermal gradient: these curves too are sigmoidal in shape (Riesner et al., 1991). In general, pH/mobility curves of proteins are generated in a polyacrylamide slab gel. However, a number of modifications have been reported, including the use of: (1) cellulose acetate as support (Vickers and Robinson, 1983); (2) agarose thin layers (Troungos et al., 1982); (3) highly cross-linked (highly porous) polyacrylamide matrices (Bianchi-Bosisio et al., 1980); (4) thin-layer silica gels (Stahl and Muller, 1981)—in this last instance, however, no electric field is used and the analysis performed is a standard ascending thin-layer chromatography against a pH gradient created by buffer diffusion; (5) paper sheets as supports (Tate, 1981)—here too the pH gradient is obtained by bathing the paper sheet in a number of buffers titrated at different pH values; and (6) paper sheets in water-alcohol solvents (Jokl et al., 1979)—again, the pH gradient is obtained as in Tate (1981). The typical gel slab size normally employed consists of a polyacrylamide plaque 12.5 × 12.5 cm and 1.6 mm in thickness. Because this is the typical plate commercially available, that is the only method described extensively in Basic Protocol 3. However, it must be emphasized that by using the Pharmacia Biotech PhastSystem, titration curves can be run in a small format (typically 4 cm × 4 cm × 0.5 mm) and the entire operation (including staining and destaining) can be performed in only 50 min. Additionally, Rüchel and Trost (1981) have described a
minichamber only 35 × 15 × 0.5 mm vertically inserted into two cylindrical buffer reservoirs. The gel slab is polymerized by using a two-vessel gradient mixer with two limiting solutions titrated to different pH values; thus, in this case too an artificial pH gradient (utilizing diffusion of nonamphoteric buffers) is created instead of a natural one (created by a current with the aid of a multitude of amphoteric buffers).
Critical Parameters and Troubleshooting Peptide mapping Excess SDS (>0.01%) is a problem for peptide mapping by HPLC (Basic Protocol 1). For samples digested in a gel slice in the presence of 0.1% SDS, the detergent must be removed prior to applying the sample to the HPLC. Otherwise, it will interfere with the resolution of fragments and the retention time on the column, and numerous artifactual peaks will be observed. SDS can be removed to a certain extent by precipitation in acidified acetone. Total removal of SDS from a protein solution requires either binding to an ion exchanger or electrophoresis. Ion-exchange precolumns that can be put in-line with the reverse phase column to remove SDS from a contaminated sample are available commercially (e.g., from The Nest Group). Excess of either SDS or salt causes problems for high-resolution SDS-polyacrylamide gels. These factors should not be a problem if the method described in the Alternate Protocol is used. In the event that a protocol containing high salt or excess SDS is used to cleave a protein, levels of both can be reduced by precipitation with acidified acetone. Generally, proteins that are precipitated by acidified acetone in the presence of SDS are readily solubilized after precipitation, probably because a small amount of SDS adheres to the protein. It is important to use the correct digestion buffer for each protease (Allen, 1989). If the wrong conditions are used, the enzyme is likely to lose its specificity, if it works at all. Some proteases can withstand up to 0.1% SDS or 10% acetonitrile without loosing activity. Others are sensitive to 0.01% SDS, and in these cases the conditions described here may not be suitable. Refer to UNITS 11.1 & 11.4 for specific protocols. For best results, proteases should be of sequencing grade. For high-resolution SDS-PAGE of proteolytic digests done in gel slices, it is crucial to
Characteristics of Recombinant Proteins
7.3.19 Current Protocols in Protein Science
Supplement 3
separate the polypeptide fragments from the gel pieces prior to running the sample. As indicated in the protocol, this can be done easily by centrifuging the sample through a microfilterfuge. Recoveries are in the 80% range for this step. Failure to remove the gel pieces will widen each peptide band, giving rise to poor resolution in the gel. Gels that will be silver stained should be washed with 50% methanol (four changes in 2 hr, or equilibration overnight) to remove substances that interfere with the staining process.
Determining the Identity and Structure of Recombinant Proteins
Disulfide linkage analysis Most steps are critical when working with a given protein for the first time, as one can lose it by precipitation or adsorption to columns, or in various other ways! Usually, however, many properties of a protein (e.g., solubility and stability) are discovered during its isolation and purification: it is important to find out all that is already known about the protein before attempting to determine the disulfide bridge positions. Unless the protein is available in great abundance, it is advisable not to commit >0.1 to 0.2 mg to any untried step. Blocking steps 1 to 3 must be performed whenever there is reason to suspect the presence of an unpaired Cys (e.g., an odd number of Cys residues in the sequence or reaction with thiol-specific reagents: Ellman, 1959; Thannhauser et al., 1984). These blocking steps must precede the fragmentation procedures to prevent scrambling (see Background Information). Good HPLC technique is critical to success: the appearance of spurious peaks on chromatograms is often due to poor technique. The analytical column should be run at a flow rate of 0.6 ml/min. This column may also be used for preparative purposes with digests of up to ∼0.2 mg protein: use flow rates of 0.6 ml/min when working with up to 0.1 mg digest and 1 ml/min when working with 0.1 to 0.2 mg digest. If working with larger amounts (up to ∼1.5 mg digested protein), use a 25-cm × 10-mm-i.d. column filled with the same packing and operate at 3 ml/min (for 0.5 to 1 mg digest) or 4 ml/min (for 1 to 1.5 mg digest). When no longer required, columns must first be flushed with five column volumes of pure acetonitrile; flow may then be stopped. If the flow is stopped with HPLC solvent A or B present, the column will rapidly deteriorate and silica or stationary phase may block the detector when flow is restarted. It is important to prevent contamination of shared stock reagents or solutions and
to avoid storing them in an inappropriate vessels—e.g., acetonitrile, trifluoroacetic acid, concentrated acetic, and formic acids may leach UV-absorbing materials from certain plastic tubes or stoppers. Use glass or polypropylene where possible, and beware of black rubber disks (even if faced with PTFE) in solvent bottle stoppers. Always start a series of runs with a blank injection of HPLC solvent A; run some standards (e.g., glucagon and insulin, obtainable from Sigma) if peaks are broad or fail to elute (expect analytical peaks that are not overloaded to have a width at half height of ∼30 sec with a gradient of 2% B/min). Ask a chromatography expert for advice if problems continue. Correct fragmentation (in general, cleavage between Cys residues, without scrambling) is crucial to success. Depending on the known or expected amino acid sequence, digestion conditions are chosen that will cleave the polypeptide chain at least once between successive Cys residues. Acidic conditions (for cyanogen bromide, see Basic Protocol 2 and UNIT 11.4; for pepsin, see Basic Protocol 2) help to avoid disulfide scrambling. If neutral or basic conditions must be used (see UNIT 11.1), it is important to keep the pH as low as possible while preserving adequate enzyme activity; keep digestion times short by using a high protein concentration and enzyme/substrate ratio; and, if possible, operate in the presence of a thiol-specific reagent to trap any transient thiols. For recombinant CD23, for example, the tryptic digestion was performed at pH 7.0 in the presence of 10 mM iodoacetic acid (Rose et al., 1992). Note that commercially available asparaginyl-endopeptidase (Pierce), in contrast with Asp-N protease (UNIT 11.1), requires reducing conditions and so cannot generally be used for disulfide bond assignments (Rose et al., 1992). If saving time is more important than the amount of material consumed, perform several trial fragmentations in parallel and examine them analytically by reversed-phase HPLC. Sometimes an isolated peptide component may not be susceptible to Edman degradation: it may be N-terminally blocked (e.g., by formyl or acetyl if it is the N-terminal peptide of the protein, or by a residue of pyroglutamic acid if a Gln residue liberated during proteolysis has cyclized). This problem is easily recognized by the fact that more components are found by mass spectrometry than by Edman degradation. It can usually be solved more easily by amino acid analysis of the isolated (reduced) component than by digestion with acyl amino acid
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Current Protocols in Protein Science
hydrolase or pyroglutamyl amino peptidase to expose an amino group for a further attempt at Edman degradation. Alternatively, sophisticated mass spectrometric sequencing techniques may be employed; however, these are not available in the average biochemical laboratory. Titration curve analysis Avoid overfilling the sample trench with protein solution! Sample running freely on the gel surface will produce smears and curtains. If some sample overflows outside the trench accidentally, blot it immediately with filter paper. If condensation is seen on the glass lid supporting the electrodes, it means that too much joule heat has been generated. This could be due to too high a power setting (i.e., 10 W); if this is the case, lower the setting. Most often, however, condensation comes from electrode filter paper strips being too wet with electrode solution; this results in solution flooding the gel surface in the first-dimension run. The remedy, at this point, would be to excise a wider area of gel in front of the electrode wicks when they are removed in the passage from the first gel to the second dimension (see Fig. 7.3.1B for this operation). Occasionally, the thermostatic water bath might not have been switched on, the coolant might not be recirculating, or the water bath might be jammed (in the Pharmacia Biotech instrument, press the button release to restart the thermostat). If, upon staining, the protein titration curves appear too wavy or excessively broad, there are three things to check. First, inspect the contact between the platinum wire and the electrode strips. The platinum wire might have been damaged or broken, or might be dislodged from its groove. Integrity, good contact, and good alignment of the platinum wire are essential for good technique. Second, check the amount of salt in the protein (if enough liquid volume is available, one can use a conductivity meter). If too much salt is present, the high sample conductivity might generate too much heat in the trench and provoke convective flows or even partial protein denaturation due to heat shock or generation of strongly acidic and/or strongly basic ion boundaries. Remember that use of salts made of strong acids or bases is an anathema to all focusing techniques. Finally, check the protein concentration. If too much protein has been applied, the titration curves will be rather broad, since diffusion is proportional to the initial sample concentration.
Anticipated Results Peptide mapping Note that partial digestion products are usually present in varying amounts in both chemical and enzymatic cleavage procedures. For this reason, if the recombinant protein will be compared with the native protein, it is important to treat both samples concurrently so that they can be compared directly. The large number of fragments that result are easily resolved by HPLC, giving many points of comparison between the recombinant and native proteins. In the event that native protein is not available (the most likely circumstance), then the peptide fragments of the recombinant protein must be analyzed by additional techniques, including amino acid analysis, N-terminal sequence analysis, and/or mass spectrometry. Analyzing peptide maps by high-resolution gel electrophoresis, on the other hand, may fail to resolve all of the fragments. While Tris-tricine gels are ideal for resolving peptides in fairly tight bands down to 2 kDa (UNIT 10.1), the error in mass measurement can be as high as 5%. Fragments close in size may not be resolved from each other by this method. In general, several digests may be required to get a good set of data to compare with the native protein. In the event that a native protein is not available for comparison, fragments can be transferred to PVDF membranes and analyzed by amino acid analysis or N-terminal sequence analysis. Disulfide linkage analysis Interpretion of the pairs of chromatograms (attempted reduction and control) is usually straightforward. If no disulfide bond is present, there should be essentially no difference in retention time between the peak present in the attempted reduction and in the control. If a disulfide bond is present, then either this bond is holding two small peptide chains together or it is holding a single polypeptide chain in a loop. In the former case, two peaks should be visible on reduction, whereas in the latter case all that is observed is a small shift to later retention time (opening of the loop exposes more of the peptide chain to the stationary phase). If the results of the analytical reduction of a fraction are very clear-cut (small shift to later retention time, or splitting into two components), it may be sufficient to submit a portion of that fraction for Edman degradation, a portion for mass spectrometry, and a portion for in
Characteristics of Recombinant Proteins
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+ Lys
NAA
Arg
NAA
Lys
His
NAA
Glu, Asp
NAA
Mobility
0
– + Glu
Arg
His
0
– 11
8
5
11
8
5
11
8
5
pH
Figure 7.3.2 Theoretical titration curves of a wild-type protein and its genetic mutants in the presence of a given amino acid substitution. NAA: neutral amino acid; broken lines represent the zero-mobility plane (sample application trench); arrows indicate the direction of amino acid mutation. From Righetti et al. (1979).
situ reduction during mass spectrometry, thus avoiding the preparative reduction and HPLC step. Ask if the mass spectroscopist can perform in situ reduction: if this service is not available, then do preparative reduction, isolation, and analysis as described in Basic Protocol 2.
Determining the Identity and Structure of Recombinant Proteins
Titration curve analysis Some examples of applications are presented in the figures. Figure 7.3.2 offers an example of differential titration curves obtained by mixing the presumptive mutant with the wild-type species. According to the various types of amino acid substitutions, different relative shapes will be detected, as shown. Note that only in the case of a replacement of arginine with a neutral amino acid (NAA) will the two curves run parallel all along the titration interval (typically not exceeding the pH 3 to 10 range). Figure 7.3.3 gives a good example of affinity-titration curves obtained by incorporating a ligand into the gel matrix. In this case, a lectin from Ricinus communis was run in presence of increasing amounts of allyl-α-D-galactose grafted onto the gel matrix. This technique gives a unique depth of field, because not only does it permit measurement of the dissociation constant (Kd), but it also makes it possible to
evaluate the variation in Kd with respect to pH (see Fig. 7.3.4).
Time Considerations Peptide mapping. A peptide mapping experiment can be completed in ∼2 days. The sample is generally digested overnight; analysis by either HPLC or high-resolution SDSPAGE is then done on the second day. HPLC runs generally take ∼2 hr each, including the preequilibration step for the column. Tris-tricine gels can be prepared ahead of time and stored overnight in a cold-box. Large gels generally require ∼5 hr to run. Staining requires ∼2 hr for Coomassie blue, whereas gels that will be silver stained should be washed with 50% methanol (four changes in 2 hr, or equilibration overnight) to remove substances that can interfere with the staining process. The silver staining process requires ∼1.5 hr. Disulfide linkage analysis. Each step (including dialysis) has been described in such a way as to make it possible to estimate how much time it will take. The time necessary for vacuum centrifugation and lyophilization steps depends somewhat on the scale, but samples containing ammonium bicarbonate must remain on the lyophilizer overnight or the salt
7.3.22 Supplement 3
Current Protocols in Protein Science
A
B
C
E
F
_
EL
+
_
IEF
+
D
9
8
7
6
5
4
9
8
7
6
5
4
9
8
7
6
5
4
pH
Figure 7.3.3 Affinity-titration curves of lectin from Ricinus communis seeds. The gel contained 6%T, 4%C, 2% Ampholine (pH 3.5 to 10), and 2 mM each Glu, Asp, Lys, and Arg. The amounts of allyl-α-D-galactose copolymerized in the gel matrix were: (A) 0 (no-ligand control); (B) 1 × 10−5 M; (C) 4 × 10−5 M; (D) 5 × 10−5 M; (E) 7 × 10−5 M; and (F) 10 × 10−5 M. The amount of protein loaded was 200 µg in 100 µl volume in all cases. The first- and second-dimension runs were 50 min at 10 W constant power and 20 min at 700 V constant voltage, respectively. The two arrows in (A) and (D) indicate the sample application trench (zero-mobility plane); the two double arrows with positive and negative symbols in (D) represent the direction and polarity of isoelectric focusing (IEF) and electrophoresis (El). From Ek et al. (1980).
50
40 Lens culinaris
Kd x
105
30
20
10
Ricinus communis
0 4
5
6
7
8
9
pH
Figure 7.3.4 Variation of the dissociation constants (Kd) as a function of pH for the lectins Ricinus communis and Lens culinaris. These values were calculated from the retarded mobilities observed at different pH values in affinity-titration curves such as the one reported in Figure 7.3.3. The two vertical arrows indicate the pI of each lectin. From Ek et al. (1980).
Characteristics of Recombinant Proteins
7.3.23 Current Protocols in Protein Science
Supplement 3
will not vaporize fully and some will remain with the protein. If blocking (steps 1 to 3), cyanogen bromide treatment, and enzymatic digestion must all be performed, the bonding pattern should be established within 2 weeks provided that an automatic injector is available for the HPLC. Titration curve analysis. The titration curve analysis can be done in one day. Casting the gel requires ∼1 hr. The first dimension electrophoresis takes ∼1 hr, and the second dimension run requires ∼30 min. Staining and destaining the gel can take ∼1 to 2 hr.
Literature Cited Allen, G. 1989. Specific cleavage of the protein in sequencing of proteins and peptides. In Laboratory Techniques in Biochemistry and Molecular Biology, 2nd ed. (R.H. Burdon and P.H. van Knippenberg, eds.) pp. 73-104. Elsevier Science Publishing, New York. Arnaud, P., Gianazza, E., Righetti, P.G., and Fudenberg, H.H. 1980. The role of sialic acid in the microheterogeneity of α1 acidic glycoprotein: Study by isoelectric focusing and titration curves. In Electrophoresis ’79 (Radola, B.J., ed.) pp. 151-163. Walter de Gruyter, Berlin. Bianchi-Bosisio, A., Loherlein, D., Snyder, R., and Righetti, P.G. 1980. Protein titration curves by isoelectric focusing-electrophoresis in highly porous (highly cross-linked) polyacrylamide gels. J. Chromatogr. 189:317-330.
Fernandez, J., DeMott, M., Atherton, D., and Mische, S.M. 1992. Internal protein sequence analysis: Enzymatic digestion for less than 10 µg of protein bound to polyvinylidene difluoride or nitrocellulose membranes. Anal. Biochem. 201:255-264. Gianazza, E., Gelfi, C., and Righetti, P.G. 1980. Isoelectric focusing followed by electrophoresis of proteins for visualizing their titration curves by zymogram and immunofixation. J. Biochem. Biophys. Methods 3:65-75. Glocker, M.O., Arbogast, B., Schreurs, J., and Deinzer, M.L. 1993. Assignment of the inter- and intra-molecular disulfide linkages in recombinant human macrophage colony-stimulating factor using fast atom bombardment mass spectroscopy. Biochemistry 32:482-488. Harris, R.J., Murnane, A.A., Utter, S.L., Wagner, K.L., Cox, E.T., Polastri, G.D., Helder, J.C., and Sliwkowski, M.B. 1993. Assessing genetic heterogeneity in production cell lines: Detection by peptide mapping of a low level Tyr to Gln sequence variant in a recombinant antibody. Bio/Technology 11:1293-1297. Henner, J. and Sitrin, R.D. 1984. Isoelectric focusing and electrophoretic titration of antibiotics using bioautographic detection. J. Antibiot. 37:1475-1478. Hojrup, P. and Magnusson, S. 1987. Disulphide bridges of bovine Factor X. Biochem. J. 245:887892.
Constans, J., Viau, M., Gouaillard, C., Bouisson, C., and Clerc, A. 1980. Binding affinities between VDBP and vitamin D3,25-(OH)-D3, 24-25(OH)2-D3 and I-25-(OH)2-D3 studied by electrophoretic methods (PAGE-IEF combined IEFelectrophoresis) and print-immunofixation. In Electrophoresis ’79 (Radola, B.J., ed.) pp. 701710. Walter de Gruyter, Berlin.
Krishnamoorthy, R., Bianchi-Bosisio, A., Labie, D., and Righetti, P.G. 1978. Titration curves of liganded hemoglobins by combined isoelectric focusing electrophoresis. FEBS Lett. 94:319323.
Creighton, T.E. 1979. Electrophoretic analysis of the unfolding of proteins by urea. J. Mol. Biol. 129:235-264.
Langen, H., Sander, B., Vilbois, F., and Lahm, H.-W. 1993. Characterization of the proteins c-kit ligand and DHFR by electrospray mass spectrometry. In Techniques in Protein Chemistry IV (R.H. Angeletti, ed.) pp. 47-54. Academic Press, San Diego.
Ek, K. and Righetti, P.G. 1980. Determination of protein ligand dissociation constants and of their pH-dependence by combined isoelectric focusing electrophoresis (titration curves). Binding of phosphorylases A and B to glycogen. Electrophoresis 1:137-140. Ek, K., Gianazza, E., and Righetti, P.G. 1980. Affino-titration curves: Determination of dissociation constants of lectin-sugar complexes and of their pH-dependence by isoelectric focusing electrophoresis. Biochim. Biophys. Acta 626:356-365. Ellman, G.L. 1959. Tissue sulfhydryl groups. Arch. Biochem. Biophys. 82:70-77. Determining the Identity and Structure of Recombinant Proteins
monobeads ion exchangers for the separation of biopolymers. Prot. Biol. Fluids 30:621-628.
Fägerstam, L., Söderberg, L., Wahlström, L., Fredriksson, U.B., Plith, K., and Walldèn, E. 1983. Basic principles used in the selection of
Jokl, V., Dolejsovà, J., and Matusovà, M. 1979. Zone electrophoresis of organic acids and bases in organic solvents. J. Chromatogr. 172:239-248.
Laskey, R.A. and Mills, A.D. 1975. Quantitative film detection of 3H and 14C in polyacrylamide gels by fluorography. Eur. J. Biochem. 56:335341. Lostanlen, D., Gacon, G., and Kaplan, J.C. 1980. Direct enzyme titration curve of NADH:cytochrome b5 reductase by combined isoelectric focusing/electrophoresis. Interactions between enzyme and cytochrome b5. Eur. J. Biochem. 112:179-183. Markwell, M.A.K. and Fox, C.F. 1978. Surface-specific iodination of membrane protein of viruses and eucaryotic cells using 1,3,4,6-tetrachloro3α,6α-diphe ny lglyco lu ril. Biochemistry 17:4807-4817.
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Current Protocols in Protein Science
Morris, H.R. and Pucci, P. 1985. A new method for rapid assignment of S-S bridges in proteins. Biochem. Biophys. Res. Commun. 126:1122-1128. Niekamp, C.W., Hixson, H.F., and Laskowski, M. 1969. Peptide-bond hydrolysis equilibria in native proteins. Conversion of virgin into modified soybean trypsin inhibitor. Biochemistry 8:16-22. Patterson, S.D. 1994. From electrophoretically separated protein to identification: Strategies for sequence and mass analysis. Anal. Biochem. 221:1-15. Picard, B., Goullet, P., and Krishnamoorthy, R. 1987. A novel approach to study of the structural basis of enzyme polymorphism. Biochem. J. 241:877-881. Proudfoot, A.I., Davies, G.D., Turcatti, G., and Wingfield, P.T. 1991. Assignment of the disulfide bridges of unglycosylated recombinant interleukin-5 expressed in recombinant Escherichia coli using proteolysis at low pH. FEBS Lett. 238:61-64. Rabilloud, T. 1990. The mechanism of protein silver staining in polyacrylamide gels: A 10-year synthesis. Electrophoresis 11:785-794. Randhawa, Z.I., Witkowska, H.E., Cone, J., Wilkins, J.A., Hughes, P., Yamanishi, K., Yasada, S., Masui, Y., Arthur, P., Kletke, C., Bitsch, F., and Shackleton, C.H.L. 1994. Incorporation of norleucine at methionine positions in recombinant human macrophage stimulating factor (M-CSF, 4-153) expressed in Escherichia coli: Structural analysis. Biochemistry 33:4352-4362. Riesner, D., Henco, K., and Steger, G. 1991. Temperature gradient gel electrophoresis. In Advances in Electrophoresis (Chrambach, A., Dunn, M.J., and Radola, B.J., eds.) vol. 4, pp. 169-250. VCH Publishers, Weinheim and New York. Righetti, P.G., Krishnamoorthy, R., Gianazza, E., and Labie, D. 1978a. Protein titration curves by combined isoelectric focusing-electrophoresis with hemoglobin mutants as models. J. Chromatogr. 166:455-460. Righetti, P.G., Gacon, G., Gianazza, E., Lostanlen, D., and Kaplan, J.C. 1978b. Titration curves of interacting cytochrome b5 and hemoglobin by isoelectric focusing electrophoresis. Biochem. Biophys. Res. Commun. 85:1575-1581. Righetti, P.G., Menozzi, M., Gianazza, E., and Valentini, L. 1979. Protolytic equilibria of doxorubicin as determined by isoelectric focusing and electrophroetic titration curves. FEBS Lett. 101:51-55. Rose, K., Turcatti, G., Graber, P., Pochon, S., Regamey, P.-O., Jansen, K.U., Magnenat, E., Aubonney, N., and Bonnefoy, J.-Y. 1992. Partial characterization of natural and recombinant human soluble CD23. Biochem. J. 286:819-824.
Rosengren, A., Bjellqvist, B., and Gasparic, V. 1977. A simple method of choosing optimum pH conditions for electrophoresis. In Electrofocusing and Isotachophoresis (Radola, B.J. and Graesslin, D., eds.) pp. 165-172. Walter de Gruyter, Berlin. Rüchel, R. and Trost, M. 1981. A study of the structural conversions of two carboxyl proteinases employing electrophoresis across a pH gradient. In Electrophoresis ’81 (Allen, R.C. and Arnaud, P., eds.) pp. 667-676. Walter de Gruyter, Berlin. Sanger, F. 1959. The chemistry of insulin. Science 129:1340-1344. Schrimsher, J.L., Rose, K., Simona, M.G., and Wingfield, P.T. 1987. Characterization of human and mouse granulocyte-macrophage-colonystimulating factors derived from Escherichia coli. Biochem. J. 247:195-199. Stahl, E. and Muller, J. 1981. pH-gradient-dunnschicht-chromatographie von benzodiazepinen. J. Chromatogr. 209:484-488. Stults, J.T. 1995. Matrix-assisted laser desorption/ionization mass spectroscopy (MALDIMS). Curr. Opin. Structur. Biol. 5:691-698. Tate, M.E. 1981. Determination of ionization constants by paper electrophoresis. Biochem. J. 195:419-426. Thannhauser, T.W., Konishi, Y., and Scheraga, H.A. 1984. Sensitive quantitative analysis of disulfide bonds in polypeptides and proteins. Anal. Biochem. 138:181-188. Thornton, J.M. 1981. Disulfide bonds in globular proteins. J. Mol. Biol. 151:261-287. Troungos, C., Krishnamoorthy, R., Elion, J., and Labie, D. 1982. Titration curves by combined isoelectric focusing-electrophoresis on a thin layer of agarose gel. 1982. J. Chromatogr. 250:73-79. Valentini, L., Gianazza, E., and Righetti, P.G. 1980. pK determinations via pH-mobility curves obtained by isoelectric focusing electrophoresis. Theory and experimental verification. J. Biochem. Biophys. Methods 3:323-338. Van Den Oetelaar, P.J. and Hoenders, H.J. 1987. Electrophoretic titration curve in 6 M urea of the bovine eye lens protein α-crystallin. J. Chromatogr. 411:507-509. Vickers, M.F. and Robinson, P.A. 1983. Protein titration curves using modified cellulose acetate membranes. J. Chromatogr. 275:428-431. Violand, B.N., Schlitter, M.R., Lawson, C.Q., Kane, J.K., Siegal, N.R., Smith, C.E., Kolodziej, E.W., and Duffin, E.L. 1994. Isolation of Escherichia coli synthesized eukaryotic proteins that contain ε-N-acetyllysine. Protein Sci. 3:1089-1097.
Characteristics of Recombinant Proteins
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Wingfield, P., Mattaliano, R., MacDonald, R.H., Craig, S., Clore, G.M., Gronenborn, A.M., and Schmeissner, U. 1987. Recombinant-derived interleukin-1a stabilized against specific deamidation. Protein Eng. 1:413-417. Wingfield, P., Benedict, R., Turcatti, G., Allet, B., Mermod, J.-J., DeLamarter, J., Simona, M.G., and Rose, K. 1988. Characterization of recombinant-derived granulocyte-colony stimulating factor (G-CSF). Biochem. J. 256:213-218.
Key Reference Allen, G. 1989. See above. Contains excellent information about common proteolytic agents.
Contributed by Nancy D. Denslow (peptide mapping) University of Florida Gainesville, Floria Keith Rose (disulfide linkage analysis) University Medical Center Geneva, Switzerland Pier Giorgio Righetti (two-dimensional titration curve analysis) University of Milano Milan, Italy
Determining the Identity and Structure of Recombinant Proteins
7.3.26 Supplement 3
Current Protocols in Protein Science
Transverse Urea-Gradient Gel Electrophoresis Monitoring the cooperative unfolding transition induced when a protein is exposed to elevated temperature or a chemical denaturant is an important strategy for characterizing the conformational properties of a globular protein. This transition may be analyzed quantitatively by a variety of spectroscopic techniques, but a simpler alternative is urea-gradient gel electrophoresis. The pattern produced in the resulting gel can be used to estimate both the free energy change for unfolding and the rate of the unfolding transition. In addition, the technique can help identify either covalent or conformational heterogeneity in a protein sample. Because urea-gradient gel patterns are sensitive to several parameters, including hydrodynamic volume, net charge, and conformational stability, the technique can be particularly useful for comparing two forms of a protein, e.g., a natural form and the product of recombinant bacteria.
UNIT 7.4 BASIC PROTOCOL
Transverse urea-gradient gels are slab gels prepared with a gradient of urea concentration perpendicular to the direction of electrophoresis (Creighton, 1979). A single sample is applied to the top of the gel, and as the protein is electrophoresed, molecules at different positions across the gel are exposed to different urea concentrations. Unfolding is detected by a decrease in electrophoretic mobility, reflecting the increased hydrodynamic volume of the unfolded protein. The gels are prepared with the glass plates rotated 90° from the orientation used for electrophoresis, which requires minor modifications of most standard electrophoresis equipment. The major requirements are spacers to fit the sides of the glass plates as oriented for casting and some arrangement for holding the glass plates in this orientation. An arrangement devised for preparing gels for the Bio-Rad Mini-Protean II apparatus is illustrated in Figure 7.4.1. The spacers and a casting box for preparing multiple gels simultaneously are available from Aquebogue. The gels are prepared using a photoacti-
90
casting
electrophoresis
Figure 7.4.1 Spacers for preparing urea-gradient gels for the Bio-Rad Mini-Protean II apparatus. (A) A gel “sandwich” assembled and oriented for casting. The spacers are designed fit along the sides of the gel, as oriented for casting, and to keep the glass plates from sliding when placed in a casting box. (B) After the gel has polymerized, the gel is rotated 90°, the casting spacers are removed, and short spacers are placed on the sides of the gel to form a single sample well. The spacers are made to prepare gels 1 mm thick and can be constructed by machining single pieces of plastic or by glueing layers together. The standard glass plates for the Mini-Protean II apparatus are used. In the figure, the thicknesses of the spacers and glass plates have been exaggerated for clarity.
Characteristics of Recombinant Proteins
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vating polymerization catalyst to allow sufficient time to prepare the gradient. After the gels have polymerized, the spacers are removed to expose the top and bottom of the gel, as oriented for electrophoresis. The gel is then preelectrophoresed and the sample is loaded. After electrophoresis, the gel is stained and analyzed. The following instructions assume that the total volume of the gels is 40 ml for five 10-cm-wide gels in the Aquebogue casting box. The gels are prepared so that 2 cm on the left and right side (as oriented for electrophoresis) contain 0 and 8 M urea, respectively. Volumes should be adjusted appropriately for gels of other dimensions. Materials Gel overlay solution: 20% (v/v) ethanol/0.002% (w/v) bromophenol blue Urea/acrylamide gel solutions with photopolymerization catalysts (see recipes) 10× electrophoresis buffer (see recipe) Protein samples Acidic or basic protein sample buffer (see recipes) Coomassie blue R-250 stain solution (see recipe) Gel rinse solution: 50% (v/v) methanol/7.5% (v/v) acetic acid Gel destain solution: 5% (v/v) methanol/7.5% (v/v) acetic acid Gel electrophoresis tank and glass plates (e.g., Bio-Rad) Spacers for casting gels (Aquebogue) Gel casting box (Aquebogue) Three-channel peristaltic pump (e.g., ISCO Tris pump) Light source to initiate photopolymerization of gels (e.g., two 15-W blue fluorescent lamps) Air-displacement pipettor Gel-loading tip Plastic boxes Set up apparatus for casting gradient gels 1. Assemble glass plates and spacers into sandwiches and place them in the casting box. 2. Connect casting box and peristaltic pump as illustrated in Figure 7.4.2 and set up the reservoir and mixing chamber. Position the pump so that it is at a slightly higher elevation than the mixing chamber and slightly below the inlet to the casting box to minimize mixing of the gradient as the solution of increasing density passes from the mixing chamber to the casting box.
3. Degas the gel overlay solution 5 min with a water aspirator. 4. Fill the bottom 2 to 4 cm of the casting box with overlay solution. Use a 20-ml syringe to draw the solution through the tubing between the mixing chamber and the casing box. Clamp the tubing between the peristaltic pump and the casting box. Make sure that the level of the solution is still above the bottoms of the glass plates. Prepare the gel solutions 5. Working in subdued lighting, prepare 30 ml each of 0 M and 8 M urea/acrylamide gel solutions containing photopolymerization catalyst. Do not add TEMED (for riboflavin polymerization) or methylene blue (for methylene blue polymerization).
Transverse Urea-Gradient Gel Electrophoresis
Because of the time required to prepare the gradients, it is convenient to use a photoactivated polymerization catalyst, such as riboflavin or methylene blue. The gels can be cast under subdued lighting (complete darkness is not necessary), and then exposed to direct light to initiate polymerization. Appropriate pH and acrylamide concentrations of gel solutions must be chosen for each protein (see Critical Parameters).
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gel sandwiches in casting box
resembles a label
mixing chamber
0M urea
peristaltic pump
8M urea magnetic stirrer
reservoir
Figure 7.4.2 Preparation of urea-gradient gels using a three-channel peristaltic pump. The gel sandwiches are held in a clear plastic casting box with an inlet for the gel solution placed below the bottoms of the glass plates. Gel solutions containing 0 and 8 M urea are placed in the mixing chamber and reservoir respectively. As solution from the mixing chamber is pumped into the casting box using two channels of the pump, the 8 M urea solution is pumped into the mixing chamber using a single channel, leading to a linear increase in urea concentration in the mixing chamber. Before the gel solution is pumped into the casting box, the bottom of the box and the tubing between it and the mixing chamber are filled with gel overlay solution. The flow rate from the pump to the casting box should be ∼2 ml/min.
Urea should be dissolved just prior to preparing the gels, because urea in solution breaks down to produce cyanate that can react with protein amino groups.
6. Degas the gel solutions 5 min with a water aspirator. 7. For photopolymerization, add 36 µl TEMED (for riboflavin) or 1.5 ml 2 mM methylene blue. Cast the urea-gradient gels 8. Place 12 ml of 8 M urea gel solution in the reservoir. Use a 20-ml syringe to fill the tubing connecting the reservoir to the mixing chamber. Clamp the tubing in the peristaltic pump and temporarily place the outlet in the 8 M urea reservoir. 9. Place 8 ml of 0 M urea solution in the mixing chamber. Remove the clamp between the pump and the casting box. Turn on the peristaltic pump to begin pumping 0 M urea solution into the casting box at a rate of ∼2 ml/min (1 ml/min per channel). When the last of the 0 M urea solution has just entered the tubing, stop the pump. Be careful not to allow any air into the tubing.
10. Place 12 ml of 0 M urea solution in the mixing chamber and position the outlet from the 8 M urea solution in the mixing chamber. Turn on the magnetic stirrer and the peristaltic pump. The concentration of urea in the flask will now increase linearly as the solution is pumped into the casting box.
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11. When the last of the solution has entered the tubing, stop the pump. 12. Place the remaining 8 M urea solution in the mixing chamber and turn on the peristaltic pump. When the boundary between the ethanol overlay solution and the 0 M urea reaches the top of the gel plates, turn off the pump. 13. Clamp off the inlet to the casting box and disconnect it from the peristaltic pump. 14. Place the gels ∼10 cm from the light source to initiate polymerization, which should take ∼30 min. 15. Promptly rinse the gel solution out of the tubing before it begins polymerizing. Set up the gel 16. After the gels have polymerized, remove them from the casting box, remove the spacers, and mount the gel in the electrophoresis chamber. Urea-gradient gels can be stored 1 to 2 days at 4°C without significant dissipation of the gradient. Before storing, wrap the gels in plastic wrap to prevent drying.
17. Dilute electophoresis buffer to 1× and fill the electrophoresis tank. 18. Preelectrophorese the gel 30 min at 100 V in the same direction as will be used for electrophoresis. For negatively charged proteins, electrophorese with the positive electrode at the bottom of the gel; for positively charged proteins, electrophorese with the negative electrode at the bottom of the gel. These conditions are usually adequate to remove reactive compounds resulting from the breakdown of urea or the polymerization process in a gel that is 1 mm thick and 10 cm wide.
Prepare and electrophorese samples 19. Prepare the sample for electrophoresis by diluting ∼50 µg of protein to a total volume of 60 µl. Add 15 µl sample buffer appropriate for acidic or basic protein. Small samples of unfolded proteins in 8 M urea can be prepared by dissolving 48 mg solid urea in 63 ìl aqueous solution (including the protein) to yield a total of 100 ìl. It is not necessary to add glycerol to samples that contain 8 M urea.
20. Apply the sample evenly across the top of the gel using an air-displacement pipettor with a gel-loading tip. 21. Electrophorese the sample. The voltage and duration of electrophoresis appropriate for a given protein and set of electrophoresis conditions must be determined empirically, but 50 to 100 V for 3 to 4 hr is a good starting point.
22. At the end of the run, turn off the power supply and disconnect the electrodes. Remove the gel from the apparatus and remove the plates. Mark the orientation of the gel by cutting off one corner. Stain and destain the gel 23. Place the gel in a plastic box containing ∼50 ml Coomassie blue R-250 stain solution. Agitate gently ≥2 hr at room temperature. Transverse Urea-Gradient Gel Electrophoresis
The high acid concentration of the recommended stain makes this solution particularly effective for fixing smaller proteins that are not easily precipitated by milder staining conditions. For some applications, staining solutions commonly used for SDS gel
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electrophoresis—e.g., 0.1% (w/v) Coomassie blue R-250/50% (v/v) methanol/7.5% (v/v) acetic acid—may also be suitable.
24. Transfer the gel to a clean plastic box containing gel rinse solution and rinse ≤5 min. 25. Transfer the gel to a plastic box containing gel destain solution. Agitate gently and change the gel destain solution as necessary to obtain a nearly clear background. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocols steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Acidic protein sample buffer 50% (w/v) glycerol 0.01% (w/v) bromophenol blue Store at room temperature Acrylamide/bisacrylamide stock solution 30 g acrylamide (30% final) 0.8 g N,N′-methylene bisacrylamide (0.8% final) H2O to 100 ml Filter through paper or an 0.45-µm filter Store at 4°C in the dark CAUTION: Acrylamide and bisacrylamide are toxic. The powders should be handled in a fume hood, and both the powder and solutions should be handled wearing gloves.
Basic protein sample buffer 50% (w/v) glycerol 0.2% (w/v) methyl green Store at room temperature Coomassie blue R-250 stain solution 0.5 g Coomassie blue R-250 dissolved in ∼400 ml H2O (1 mg/ml final) 50 g trichloroacetic acid (0.61 M final) 50 g 5-sulfosalicylic acid (0.39 M final) H2O to 500 ml Store at room temperature CAUTION: Trichloroacetic acid and 5-sulfosalicylic acid are very caustic. Wear gloves when handling the solids or stain solution.
Electrophoresis buffer, 10× Buffer solutions that have been used successfully for urea-gradient gels include: 0.5 M acetate-Tris, pH 4.0: 0.5 M acetic acid adjusted to pH 4.0 with Tris base 0.5 M imidazole/0.5 M MOPS, pH ∼7.0 0.5 M Tris-acetate, pH 8.0: 0.5 M Tris base adjusted to pH 8.0 with acetic acid 0.5 M Tris/0.5 M Bicine, pH ∼8.4 Store all buffer solutions at room temperature Urea/acrylamide gel solutions containing photopolymerization catalysts 0 M urea/15% acrylamide gel solution for riboflavin-polymerized gels 15 ml acrylamide/bisacrylamide stock solution (see recipe) 3 ml 10× electrophoresis buffer (see recipe; 1× final) 3.75 ml 0.04 mg/ml riboflavin (5 µg/ml final) 8.25 ml H2O Immediately before casting gels, degas gel solution, then add 36 µl TEMED. continued
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8 M urea/11% acrylamide gel solution for riboflavin-polymerized gels 14.4 g solid urea 11 ml acrylamide/bisacrylamide stock solution (see recipe) 3 ml 10× electrophoresis buffer (see recipe; 1× final) 3.75 ml 0.04 mg/ml riboflavin (5 µg/ml final) 1.2 ml H2O Immediately before casting gels, degas gel solutio, then add 36 µl TEMED. 0 M urea/11% acrylamide gel solution for methylene blue–polymerized gels 11 ml acrylamide/bisacrylamide stock solution (see recipe) 3 ml 10× electrophoresis buffer (see recipe; 1× final) 1.5 ml 20 mM sodium toluene sulfinate (1 mM final) 1.5 ml 1 mM diphenyliodonium chloride (50 µM final) 11.5 ml H2O Immediately before casting gels, degas solution and add 1.5 ml 2 mM methylene blue (100 µM final). 8 M urea/7% acrylamide gel solution for methylene blue–polymerized gels 14.4 g solid urea 7 ml acrylamide/bisacrylamide stock solution (see recipe) 3 ml 10× electrophoresis buffer (see recipe; 1× final) 1.5 ml 20 mM sodium toluene sulfinate (1 mM final) 1.5 ml 1 mM diphenyliodonium chloride (50 µM final) 4.4 ml H2O Immediately before casting gels, degas solution and add 1.5 ml 2 mM methylene blue (100 µM final). COMMENTARY Background Information
Transverse Urea-Gradient Gel Electrophoresis
One of the most general properties of folded globular proteins is the tendency to undergo a cooperative unfolding transition when exposed to elevated temperature or to chaotropic agents such as urea or guanidinium chloride (Tanford, 1968). The unfolding transition for a particular protein under defined conditions is characterized by several thermodynamic properties, including the free energy change for unfolding, the enthalpy change, the entropy change, and, even more fundamentally, whether or not the transition is sufficiently cooperative that only fully folded and fully unfolded molecules are present at equilibrium. The free energy change for unfolding, ∆Gu, is a particularly important parameter that represents the net stability of the folded conformation relative to the unfolded state and is widely used for comparing the stabilities of related proteins. Unfolding-refolding transitions are also characterized by their kinetic properties, including their rates and the accumulation of any transient intermediates. The thermodynamics and kinetics of protein unfolding and refolding reactions have been
studied extensively using a variety of different biophysical methods, including spectroscopic and calorimetric techniques. These studies have led to important advances in understanding the physical interactions that stabilize folded proteins and the mechanisms by which these structures form (Schellman, 1987; Privalov, 1989; Kim and Baldwin, 1990; Creighton, 1992; Matthews, 1993). Although spectroscopic techniques and calorimetry have been mastered in a growing number of laboratories specializing in protein stability and folding, these measurements are quite technically demanding and require equipment and expertise that is not always found in typical protein biochemistry laboratories. Urea-gradient gel electrophoresis was developed by T.E. Creighton (1979) as a simple alternative to spectroscopic methods for studying protein folding and unfolding. In this procedure, a single sample is applied across the top of a polyacrylamide slab gel containing a horizontal gradient of urea concentration. As the sample is electrophoresed through the gel, molecules at different positions across the gel are exposed to increasing urea concentrations.
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Those molecules that unfold have a larger hydrodynamic volume than the folded protein molecules and therefore migrate more slowly. After electrophoresis, the stained gel produces a graphic representation of the unfolding transition. As discussed later (see Anticipated Results), the pattern generated can provide information about whether or not the protein undergoes a urea-induced unfolding transition, information about the presence of significantly populated intermediate states in this transition, and semiquantitative information about the net stability of the folded protein and the rates of interconversion between the native and unfolded states. Although the data obtained from urea-gradient gels are not generally as quantitative as those derived from spectroscopic or calorimetric measurements, the electrophoretic method offers a number of advantages for many applications. In addition to being relatively simple to implement, urea-gradient gel electrophoresis requires only small amounts of protein—a 50µg sample is sufficient for detection with Coomassie blue, and even less can be used with more sensitive detection methods (UNIT 10.5 & 10.6). Furthermore, heterogeneous samples may often be used, because different components are likely to be separated during electrophoresis (Hollecker and Creighton, 1982). Indeed, ureagradient electrophoresis is a particularly sensitive method for detecting heterogeneity, because molecules that differ in their electrophoretic mobilities in either the native or unfolded form can be distinguished, as can molecules that differ only with respect to the urea concentration at which they unfold. In addition to being convenient, urea-gradient gels can often provide information about protein-folding transitions that is complementary to the information obtained from more conventional biophysical methods. Most spectroscopic techniques—e.g., UV absorbance (UNIT 7.1), fluorescence, and circular dichroism—measure only the averaged properties of a sample. As a consequence, it is often difficult to determine if, for instance, the signal measured at an intermediate urea concentration arises exclusively from a mixture of fully unfolded and fully native molecules or if a partially folded species may be present. Separation techniques such as gel electrophoresis can sometimes help resolve such ambiguities, particularly if different conformational states interconvert relatively slowly on the time scale of the separation.
The most useful applications of urea-gradient gels are often those where a qualitative comparison of different proteins is required, as in comparing the protein produced in recombinant bacteria with that obtained from a natural source. If the recombinant-produced protein has the same covalent structure and has folded to the same conformation as that of the natural protein, the two samples should display identical electrophoresis patterns. If, on the other hand, the two proteins differ in their covalent structure, this might be reflected in differences in the mobilities of either the native or unfolded forms, particularly if there are differences in net charge. If the conformations of the folded proteins are different, this may lead to differences in the mobilities of the native form or in the urea concentration at which the proteins unfold. Other applications include screening protein variants for differences in folding thermodynamics and kinetics (Klemm et al., 1991; Creighton and Shortle, 1994) and carrying out preliminary experiments to define conditions for more detailed studies using spectroscopic methods.
Critical Parameters Because the behavior of a protein during urea-gradient gel electrophoresis depends on several properties of the particular protein, including its hydrodynamic volume, net charge, and conformational stability, there are more parameters that must be considered than for SDS gel electrophoresis, for instance. These parameters include pH, direction of electrophoresis, gel composition, and duration of electrophoresis. pH Because electrophoresis depends on the intrinsic net charge of the protein, the pH must be sufficiently far from the isoelectric point to maintain a significant net charge, usually 5 to 10 charge units (the charge of a proton or electron). For proteins with extremely high or low isoelectric points, it will probably be necessary to work below or above the isoelectric point, respectively, but proteins with isoelectric points near neutrality can be electrophoresed under either acidic or basic conditions. Although it may be desirable, in general, to use a pH close to physiological, in some cases it may be necessary to use more extreme conditions. For instance, it may be necessary to use a higher or lower pH to maintain a large enough net charge on the protein for electrophoresis or to
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keep the protein soluble, because many proteins are least soluble close to their isoelectric points. In addition, pH can be used as a means to manipulate the stability of the folded protein. Many proteins are too stable to be unfolded in 8 M urea at neutral pH, but can be unfolded by urea under alkaline or acidic conditions. Several different buffers have been used successfully for urea-gradient gels (see Reagents and Solutions). Ideally, the ionic strength of the buffer solution should be as low as possible to minimize the heat generated during electrophoresis. Meeting this condition can be facilitated by using buffers in which both the anionic and cationic component provide buffering capacity; several buffer systems of this type have been described by McLellan (1982). The direction of electrophoresis is determined by the pH chosen. If the pH is above the isoelectric point, the protein will have a net negative charge and will migrate towards the anode in the electrophoresis tank, as in SDS–polyacrylamide gel electrophoresis. If, on the other hand, a pH below the isoelectric point is used, the orientation of the electrodes must be reversed.
Transverse Urea-Gradient Gel Electrophoresis
Gel composition As in other forms of electrophoresis, the gel composition must be chosen to match the hydrodynamic volume of the protein of interest, with larger proteins requiring lower acrylamide concentrations. The sieving properties of the gel and, therefore, the choice of acrylamide concentration also depend on the presence of urea in the gel and the method of polymerization used. For reasons that have not been fully characterized, urea in a polyacrylamide gel decreases the electrophoretic mobility, so that even in the absence of an unfolding transition there will be a progressive reduction of mobility across a urea-gradient gel. This can be largely compensated for by incorporating an acrylamide gradient so that there is a lower acrylamide concentration at the high-urea side of the gel. In his original protocol, Creighton (1979) used a 15% to 11% (w/v) acrylamide gradient (photopolymerized with riboflavin) superimposed on the 0 to 8 M urea gradient, and these conditions have worked well for a variety of monomeric proteins, including bovine serum albumin, with Mr = 66,000. Larger proteins, however, are likely to require lower acrylamide concentrations. Gel sieving is also greatly influenced by the conditions used for polymerization. The two catalysts that have been most widely used to
polymerize acrylamide gels are ammonium persulfate and riboflavin, the latter used for photoactivation. Both are used in conjunction with N,N,N′,N′-tetramethylethylenediamine (TEMED). Photopolymerization is particularly convenient for preparing gradient gels, because the timing of polymerization can be easily controlled. A major drawback to the use of riboflavin, however, is that polymerization is not as complete as when ammonium persulfate is used. Although this is not a serious difficulty for gels prepared at slightly acidic pH, at neutral or alkaline pH the gels may be very soft or the solution may not gel at all. The efficiency of photopolymerization can be increased by adding a small amount of ammonium persulfate, but it can be very tricky to find a concentration that will stimulate photopolymerization without also causing early polymerization. More recently, a procedure for photopolymerizing gels using methylene blue has been described, and it appears that this catalyst leads to much more efficient polymerization than riboflavin does (Lyubimova et al., 1993). Urea-gradient gels containing 11% to 7% (w/v) acrylamide and polymerized with methylene blue have been reported to have sieving properties similar to those of 15% to 11% (w/v) acrylamide gels polymerized with riboflavin (Creighton and Shortle, 1994). Sample size For urea-gradient gels ∼10 cm wide, a sample volume of 75 µl is appropriate. This is large enough to apply evenly across the top of the gel and small enough to give reasonably narrow bands. It should be possible, in principle, to use discontinuous buffer systems with a stacking gel, but this does not appear to be necessary. Even in the absence of a discontinuous buffer system, there is usually some stacking effect when a macromolecular sample enters a gel, because the first molecules to enter the gel are retarded and accumulate near the top of the gel while the rest of the molecules enter the gel. The difference between the mobilities of molecules above and within the gel can be enhanced by making the ionic strength of the sample lower than that of the electrophoresis buffer. Duration of electrophoresis The time required for electrophoretic separation depends on the voltage applied to the gel, and this parameter can be manipulated according to the type of information desired. As discussed later (see Anticipated Results), the elec-
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Troubleshooting The difficulties most likely to arise in using urea-gradient gel electrophoresis are the formation of protein aggregates or covalent modifications either before or during the electrophoresis. Aggregation is likely to lead to smearing of the protein bands or a complete failure of the protein to enter the gel. It may be possible to minimize aggregation by choosing a different pH or ionic strength for electrophoresis or by decreasing the concentration of protein applied to the gel. Covalent modifications are likely to cause multiple protein bands, particularly if they lead to changes in net charge. Such modifications may arise from residual free radicals from the polymerization reaction or from cyanates generated by hydrolysis of urea. To minimize these reactions, the urea solutions should be prepared immediately before casting the gels, and the gels should be preelectrophoresed before applying the sample.
Anticipated Results The simplest patterns generated by ureagradient gel electrophoresis arise when the native (N) and unfolded (U) forms of the protein are in rapid equilibrium and there are no significant populations of partially folded intermediates. In these cases, a simple sigmoidal band is produced (see Fig. 7.4.3). If the two forms are in rapid equilibrium on the time scale of the electrophoresis, the band should remain sharp through the transition region, and the same pattern should be obtained whether native or urea-unfolded protein is applied to the gel. Provided that only the native
Electrophoresis
trophoresis pattern generated depends to a large degree on whether the interconversion between native and unfolded states is rapid on the time scale of the electrophoresis. In particular, if the two forms interconvert many times during the electrophoresis, then a smooth, continuous band of protein should be generated, and the position of the observed transition can be used to estimate the thermodynamic stability of the protein. On the other hand, if the two forms interconvert slowly, a discontinuous or blurred band will be generated, and under favorable conditions it is possible to estimate the rate of the transition from the appearance of the bands and the duration of electrophoresis. Significant separations have been observed after as little as 8 min of electrophoresis (Creighton, 1980), but times as long as 10 hr have been used to generate smooth patterns for relatively slow transitions (Klemm et al., 1991). An additional parameter that can be manipulated to advantage is the presence of other molecules or ions in the gel that influence the unfolding transition by binding selectively to either the native or unfolded forms. For instance, the influence of a divalent cation can be examined by comparing the patterns observed when the ion is present in the electrophoresis buffers with the patterns obtained when EDTA is added to the buffers. This approach is limited, of course, to compounds that do not interfere with either gel polymerization or electrophoresis. Thiol reagents, for instance, react with free radicals generated during polymerization, and high salt concentrations lead to very high temperatures during electrophoresis.
unfolded protein
native protein 0 M
8 M
Urea gradient Figure 7.4.3 Diagram of an unfolding curve produced by urea-gradient gel electrophoresis of a protein undergoing rapid interconversion between the folded and unfolded states.
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studies of protein unfolding transitions have shown that the dependence of ∆Gu on denaturant concentration can usually be well described by a simple linear relationship,
and unfolded forms are significantly populated, the observed mobility at the intermediate urea concentrations will reflect the relative concentrations of N and U ([N], [U]), and the pattern can be interpreted as a plot of the fraction of protein unfolded versus urea concentration. Although urea-gradient gels are used primarily for qualitative analysis of protein unfolding transitions, they may also be used to obtain an estimate of the conformational stability of a protein, provided that the conditions described above are met. The free energy change for unfolding is given by ∆Gu = −RT ln
∆Gu = m ⋅ (C − Cm)
where Cm is the urea concentration at the transition midpoint (i.e., where ∆Gu = 0, and the average mobility is midway between that of the native and unfolded forms), C is any other urea concentration and m is an empirical parameter (Schellman, 1978; Pace et al., 1989). Hollecker and Creighton (1982) have shown that m is proportional to the derivative of the fraction unfolded (fu) with respect to urea concentration evaluated at the midpoint according to the relationship:
[U] [N]
where R is the gas constant and T is the absolute temperature. The value of ∆Gu will depend on the urea concentration, becoming more negative at higher concentrations. Because ∆Gu can only be measured at urea concentrations where N and U are both detectable, i.e., in the transition zone, it is usually necessary to extrapolate values measured near the middle of the transition to 0 M urea in order to estimate the stability under physiological conditions. Numerous
m = −4RT
dfu (Cm) dC
This derivative can be estimated from urea-gradient gel patterns by drawing a line tangent to the gel band at the midpoint and measuring the change in urea concentration, ∆C, associated with the extrapolated change in fu from 0 to 1. This procedure is illustrated in Figure 7.4.4.
fu = 1
m = –4RT ∆C
fu = 0 ∆C
0
2
4
6
8
Urea concentration (M) Transverse Urea-Gradient Gel Electrophoresis
Figure 7.4.4 Estimating the parameter m from a urea-gradient gel. See text for details.
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Electrophoresis
Cm
0 M
8 M
Urea gradient
Figure 7.4.5 Urea-gradient gel pattern produced by a protein that slowly interconverts between the native and unfolded states when native protein is applied to the gel.
The derivative is simply 1/∆C, and m is calculated according to: m=
−4RT ∆C
From the value of m and Cm, the value of ∆Gu can be calculated for any other urea concentration, including 0 M, where ∆Gu = −mCm (Hollecker and Creighton, 1982; Klemm et al., 1991). Quantitating urea-gradient gel results in this manner requires knowing the urea concentration at different positions across the gel, which can be estimated from the dimensions of the gel and from the known volumes of the linear gradient and the two bordering segments containing 0 and 8 M urea. Although values of Cm, m, and ∆Gu estimated from urea-gradient gels are certainly not expected to be as accurate as those obtained by spectroscopic methods, they have been found to be surprisingly consistent with more rigorous measurements (Hollecker and Creighton, 1982; Pace et al., 1989). If the native and unfolded forms do not interconvert rapidly during electrophoresis, a more diffuse band or a discontinuity will be generated in the transition zone (see Fig. 7.4.5). In addition, different patterns are expected when the native and unfolded forms are applied to the gel, with the transition appearing slightly above the equilibrium midpoint (Cm) when native protein is applied and below Cm when unfolded protein is applied. These patterns arise because the rate constants for unfolding and refolding, as well as the equilibrium constant, vary with urea concentration, with unfolding rates increasing and folding rates decreasing at higher urea concentrations. When native pro-
tein is applied to the gel, it remains in its initial state at low urea concentrations both because this state is thermodynamically favored and because unfolding is very slow. At urea concentrations near the equilibrium midpoint, the native conformation does not predominate thermodynamically, but at least a fraction of the molecules remain in this state simply because they do not have time to unfold during the time of electrophoresis. At urea concentrations somewhat above the midpoint, the unfolding rate is fast enough that most or all of the molecules unfold during the electrophoresis. When unfolded protein is applied to the gel, the opposite effect is seen, with a fraction of the molecules remaining unfolded even at urea concentrations below the equilibrium midpoint. When the unfolding and refolding rates are not fast compared to the time of electrophoresis, the patterns cannot be used to estimate thermodynamic parameters, but it may be possible to obtain some information about the kinetics of the transition. If the half-time for unfolding (or folding) at the midpoint is approximately equal to the time of electrophoresis, a smeared band will be generated in the transition zone, and slower transitions will give rise to a discontinuity, as described above. The rate constants can be estimated by comparing the observed patterns with those predicted by numerical simulations, as discussed in greater detail elsewhere (Creighton, 1979; Goldenberg and Creighton, 1984; Goldenberg, 1989). Much more complicated gel patterns can arise if there are additional species present either at equilibrium or accumulating as transient kinetic intermediates. If there is a partially
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Electrophoresis
A
unfolded protein
native protein 0 M
8 M Urea gradient
Electrophoresis
B
unfolded protein
native protein 0 M
8 M Urea gradient
Figure 7.4.6 Urea-gradient gel patterns produced by proteins with additional species. (A) A protein with a partially unfolded species that predominates at intermediate urea concentrations. (B) A protein with two unfolded species, one of which refolds very slowly.
Transverse Urea-Gradient Gel Electrophoresis
unfolded species that predominates at intermediate urea concentrations, a two-step transition may be observed (see Fig. 7.4.6A). A frequently observed type of conformational heterogeneity is one in which there are multiple unfolded forms that refold at different rates. Multiple unfolded forms can arise from isomerization of prolyl peptide bonds (Brandts et al., 1975) and can give rise to the pattern depicted in Figure 7.4.6B when unfolded protein is applied to the gel (Creighton, 1980). This pattern is generated because some of the unfolded molecules can rapidly refold and establish an equilibrium with the native protein, but others do not have time to refold during the electrophoresis and remain unfolded even at urea concentrations at which the native protein is thermodynamically preferred. At the lowest urea concentrations, some of the slowly refolding form may have time to fold to the native form, giving rise to a blurred or discontinuous
transition. Also, the slowly folding form may equilibrate with partially folded forms, giving rise to a downward curvature in the upper band. When native protein is applied to the gel, only the lower, sigmoidal, band is expected, because the slowly refolding form is generated relatively slowly after the protein unfolds. Other complexities in the patterns may arise from multiple folded forms, intermediates that convert slowly with the native or unfolded protein, or oligomeric proteins that undergo dissociation either prior to or in concert with unfolding.
Time Considerations Most urea-gradient gel electrophoresis experiments can easily be completed in 1 day. A few hours should be allocated for preparing the gels, and electrophoresis may take as little as 20 min or as long as several hours. Once prepared, the gels can be kept at least 2 days at 4°C without apparent changes in their properties.
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Literature Cited Brandts, J.F., Halvorson, H.R., and Brennan, M. 1975. Consideration of the possibility that the slow step in protein denaturation reactions is due to cis-trans isomerism of proline residues. Biochemistry 14:4953-4963. Creighton, T.E. 1979. Electrophoretic analysis of the unfolding of proteins by urea. J. Mol. Biol. 129:235-264. Creighton, T.E. 1980. Kinetic study of protein unfolding and refolding using urea-gradient electrophoresis. J. Mol. Biol. 137:61-80. Creighton, T.E. 1992. Protein Folding. W.H. Freeman, New York. Creighton, T.E. and Shortle, D. 1994. Electrophoretic characterization of the denatured states of staphylococcal nuclease. J. Mol. Biol. 242:670-682.
Matthews, C.R. 1993. Pathways of protein folding. Annu. Rev. Biochem. 62:653-683. McLellan, T. 1982. Electrophoresis buffers for polyacrylamide gels at various pH. Anal. Biochem. 126:94-99. Pace, C.N., Shirley, B.A., and Thomson, J.A. 1989. Measuring the conformational stability of a protein. In Protein Structure: A Practical Approach. (T. E. Creighton, ed.) pp. 311-330. IRL Press, Oxford. Privalov, P.L. 1989. Thermodynamic problems of protein structure. Annu. Rev. Biophys. Chem. 18:47-69. Schellman, J.A. 1978. Solvent denaturation. Biopolymers 17:1305-1322. Schellman, J.A. 1987. The thermodynamic stability of proteins. Annu. Rev. Biophys. Chem. 16:115137.
Goldenberg, D.P. 1989. Analysis of protein conformation by gel electrophoresis. In Protein Structure: A Practical Approach (T.E. Creighton, ed.) pp. 225-250. IRL Press, Oxford.
Tanford, C. 1968. Protein denaturation. Adv. Protein Chem. 23:121-282.
Goldenberg, D.P. and Creighton, T.E. 1984. Gel electrophoresis in studies of protein conformation and folding. Anal. Biochem. 138:1-18.
Creighton, 1979. See above.
Hollecker, M. and Creighton, T.E. 1982. Effect on protein stability of reversing the charge on amino groups. Biochim. Biophys. Acta 701:395-404. Kim, P.S. and Baldwin, R.L. 1990. Intermediates in the folding reactions of small proteins. Annu. Rev. Biochem. 59:631-660. Klemm, J.D., Wozniak, J.A., Alber, T., and Goldenberg, D.P. 1991. Correlation between mutational destabilization of phage T4 lysozyme and increased unfolding rates. Biochemistry 30:589594. Lyubimova, T., Caglio, S., Gelfi, C., Righetti, P.G., and Rabilloud, T. 1993. Photopolymerization of polyacrylamide gels with methylene blue. Electrophoresis 14:40-50.
Key References The original description of urea-gradient gels; includes extensive discussion of most of the important parameters and considerations and detailed analysis of the relationships between transition rates and electrophoretic patterns. Goldenberg and Creighton, 1984. See above. General review of gel electrophoresis and applications to studies of protein folding, including urea gradients.
Contributed by David P. Goldenberg University of Utah Salt Lake City, Utah
Characteristics of Recombinant Proteins
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Supplement 3
Analytical Ultracentrifugation Analytical ultracentrifugation is the grandfather of biochemical methods, with a 70-year history of development and application (Svedberg and Pederson, 1940; Schachman, 1959). Recently, the attention of protein scientists and molecular biologists has been drawn to macromolecular interactions, an area where analytical ultracentrifugation excels, resulting in a renewed interest in this technology (Harding et al., 1992; Ralston, 1993; McRorie and Voelker, 1993; Schuster and Laue, 1994). As this unit will describe, analytical ultracentrifugation is one of the most powerful, though as yet underexploited, techniques available to molecular biology and biochemistry. Table 7.5.1
UNIT 7.5
Two different but complementary methods of sample analysis are possible using an analytical ultracentrifuge: sedimentation velocity and sedimentation equilibrium (Table 7.5.1). Each encompasses a range of techniques suitable for characterizing everything from crude mixtures to highly purified proteins. Sedimentation velocity provides such hydrodynamic information as size, shape, and, when combined with diffusion measurements, molecular weight. Sedimentation equilibrium provides thermodynamic information about molecular weight, stoichiometry, association energy, and nonideality. Both sedimentation methods are understood from first principles. Thus, no standards are
Methods of Analytical Ultracentrifugation
Method
Category
Velocity
Direct fitting to transport equations
Modern way to obtain s (and the diffusion coefficient) for a single solute. When combined with molecular weight, s yields information concerning the size and shape of the protein (Philo, 1994). g(s)–particle size Powerful method for analyzing complicated distribution analysis mixtures of molecules. Holds promise for analyzing associating systems (Stafford, 1992; Stafford, 1994). By increasing the gravitational field over the course of an experiment, fractionation of incredibly complicated mixtures may be achieved (Mächtle, 1988). Differential Sensitive means of detecting small (0.005%) sedimentation differences in sedimentation due to conformational changes (Richards and Schachman, 1957). Active enzyme Means of characterizing an impure enzyme (Cohen sedimentation et al., 1971) using a chromophoric assay and monitoring the movement of color development as the enzyme sediments. Van Holde and Graphical means of removing the effects of Weischet extrapolation diffusion, thus improving resolution of a paucidisperse solution (Van Holde and Weischet, 1978).
Equilibrium
Short-column analysis (750 µm, 15 µl samples) Longer-column analysis (3 mm, 110 µl samples)
Useful for quick surveys of molecular weight, association properties, and nonideality (Yphantis, 1960; Laue, 1992). Provides higher precision than short-column; useful for low-molecular-weight solutes and analysis of heterogeneity (Yphantis, 1964).
Miscellaneous
Extinction coefficient
Combines absorbance and refractive optical reading for precise measurement of extinction coeffiecient. Sensitive test for sample homogeneity. Very sensitive and selective for characterizing interactions between purified proteins. Obtained at low speed.
Tracer sedimentation Diffusion coefficient
Information obtained and notes
Contributed by Thomas M. Laue Current Protocols in Protein Science (1996) 7.5.1-7.5.9 Copyright © 2000 by John Wiley & Sons, Inc.
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Analytical Ultracentrifugation
required to interpret data and the same values should be obtained for parameters (e.g., molecular weight and s20,w) from preparation to preparation and from laboratory to laboratory. Of the two methods, sedimentation velocity provides a greater degree of sample fractionation, and it is uniquely suited to particle size distribution analysis (see Anticipated Results for definitions and further discussions). The standardized sedimentation coefficient (s020,w; measured in Svedberg units, S) is a constant for a given molecule; it therefore is often used as a species descriptor (e.g., 30S subunit, 5S RNA). The reasons behind the revival of analytical ultracentrifugation reveal many of its virtues. Fundamental information concerning sample purity, native molecular weight, association energies, and molecular size and shape are all readily obtainable using this technique. The methods are not fussy with regard to solvent conditions, and can be applied to any sort of protein (fibrous or globular, large or small) over an enormous range of molecular weights (<100 to >10,000,000 g/mol). Relatively small quantities of materials are required for analysis and, because the techniques are nondestructive, samples can be recovered afterwards. Moreover, great experimental precision can be anticipated, even by a novice experimenter. The disadvantages of sedimentation analysis are different for the velocity and equilibrium methods. Sedimentation velocity analysis requires relatively large sample volumes (450 µl) and, being a broad-zone method, exhibits poor sample fractionation when compared with various chromatographic and electrophoretic methods (Table 7.5.2). Subtle changes readily apparent upon electrophoresis may not be revealed by sedimentation. Another consequence of the technique’s broad-zone nature is that only the slowest-sedimenting species can be examined in a pure state; all other boundaries will be composed of mixtures of components. The interpretation of sedimentation equilibrium data requires that highly purified proteins be used. For the analytical ultracentrifuge, relatively high concentrations of proteins (0.1 to 1.0 mg/ml) are required for detection (Table 7.5.3). Even if these criteria are met, complicated association schemes (e.g., highly nonideal monomer:dimer:tetramer:octamer associations) are difficult to analyze in great detail. Finally, neither sedimentation velocity nor sedimentation equilibrium is suitable for kinetic analysis.
Even with these disadvantages, sedimentation analysis compares favorably with alternative methods (Table 7.5.2). There are three classes of methods that can be compared with sedimentation: (1) other primary methods (light scattering and osmotic pressure), (2) spectroscopic analysis for obtaining thermodynamic information, and (3) secondary methods built around electrophoresis and column chromatography. Both light scattering and osmotic pressure are well-grounded in thermodynamics and provide accurate estimates of molecular weight. However, neither affords any fractionation of the solution and neither provides hydrodynamic information equivalent to that from sedimentation velocity. Both are subject to problems with contamination (osmotic pressure from small particles and light scattering from large ones), though this sensitivity can be useful for detecting such contaminants. Finally, both require larger quantities and higher concentrations of sample than does sedimentation. Spectroscopic methods—e.g., fluorescence, absorbance, circular dichroism (CD), nuclear magnetic resonance (NMR), and electron spin resonance (ESR)—are often useful for examining sample purity and for conducting binding studies. For sample purity, these methods are usually applied to detect the presence of specific contaminants (e.g., detecting protein in a DNA sample by absorbance). The concentrations needed to conduct a study range from quite low (fluorescence) to rather high (NMR). In addition to sample purity, spectroscopy often is used phenomenologically to monitor associations, including macromolecular interactions. Because of their respective sensitivities, the concentration range accessible to these methods is quite variable. All spectroscopic methods suffer from certain common problems. First, because they are phenomenological, it is not guaranteed that binding will lead to the production of an appropriate signal. Hence, it is possible for an interaction to occur that gives rise to no measurable spectroscopic change. Moreover, if the stoichiometry of binding is >1, it is possible for different binding events to give rise to proportionately different signals. In these cases, the stoichiometry of binding will be nearly impossible to determine and estimates of the strength of binding may be seriously in error. There are several chromatographic and electrophoretic methods available for making measurements similar to those from sedimentation. These fractionate the sample and therefore are extremely useful for detecting contam-
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Current Protocols in Protein Science
Table 7.5.2
Alternative Analytical Methods
Class
Method
Notes
Primary methods No standards required; data interpretation grounded in physical first principles; no sample fractionation.
Scattering
Light and low-angle X-ray can yield molecular weight and radius of gyration. For mixtures, weightaverage molecular weights and z-average radii are obtained. (Laue, 1995; Yphantis, 1964). Sensitive to large contaminants. Small cuvettes reduce sample volume, but sensitivity is limited to relatively high concentrations (>1 mg/ml). Particle scattering (e.g., neutrons) is very expensive but yields details about particular aspects of structure.
Osmotic pressure (other colligative methods)
Useful for determining molecular weights of small molecules. For mixtures, number-average molecular weights are obtained. Sensitive to low-molecularweight contaminants. Relatively high concentrations (>1 mg/ml) are required. Vapor pressure osmometers require small sample volumes.
Absorbance
Useful for detecting particular contaminants. Simple to use. Sensitivity to contaminants depends on their extinction coefficient at the wavelength of interest. Changes in absorbance with binding can be used to monitor macromolecular interactions with varying success.
Fluorescence
Very sensitive. Both luminescence and quenching of fluorescence useful for detecting contaminants. Changes in fluorescence can be used to monitor macromolecular interactions with varying success. Not generally as useful as other spectroscopic methods for detecting contaminants. Can be used to monitor macromolecular interactions with varying success. Instrumentation expensive.
Spectroscopic methods Primarily useful phenomenologically; sample size and sensitivity vary from method to method; no sample fractionation.
Circular dichroism
Secondary methods Require standards for interpretation; can be particularly useful for detecting contaminants; most are used as “thin-zone” methods, which limits their usefulness in characterizing interactions.
Nuclear magnetic resonance, electron spin resonance
Not generally as useful as other spectroscopic methods for detecting contaminants. Can be used to monitor macromolecular interactions with varying success. Requires high concentrations of material and instrumentation expensive.
Column chromatography
Depending on column matrix, can be sensitive to size, charge, or specific binding. In studying binding, useful qualitatively but less so for quantitative work. Fractionation can be excellent. Ability to measure contaminants depends on detector used to monitor effluent, but can be exquisitely sensitive. A modified detector in which the matrix is attached to the detector is at the heart of the Pharmacia Biotech Biacore and related devices. Several methods based on electrophoretic mobility exist (e.g., gel electrophoresis, capillary zonal electrophoresis, and isoelectric focusing). Fractionation is usually superb; sensitivity depends on detector. Because of the high degree of sample fractionation, only limited, qualitative information about macromolecular interactions is obtainable.
Electrophoresis
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Methods of Solute Detection
Table 7.5.3
Method of detection
Usea
Pb
Ac
Sd
De
Notes
Absorbance
A, P
G
F
G
F
Refractive
A
E
E
G
P
Fluorescence
P
F
F
E
E
Enzyme assay
P, A
F
F
E
E
Radiolabel
P
G
F
E
E
Other
P
—
—
—
—
Sensitivity depends on solute’s extinction coefficient. Discrimination depends on separation between absorbances. Accuracy and precision cannot be matched by other techniques. Method of choice for nonabsorbing solutes or buffers that absorb in the UV. Specific labels can provide excellent sensitivity and superb discrimination, but the linearity of the concentration determination is subject to several types of error. Excellent discrimination and sensitivity. Can be used to determine which form of an enzyme is active. Precision and accuracy can be quite good or rather poor, depending on the assay. A band sedimentation method for the analytical ultracentrifuge works quite well, but requires that a suitable colorimetric assay be available. Superb discrimination and sensitivity can be attained. Accuracy of the concentration determination can range from very good to very poor depending on the particulars of the radiolabeling. Any assay (RIA, ELISA, bioassay) that yields a signal proportional to the concentration may be used for determining molecular weight and s20,w. Of particular interest are methods in which the sample is fractionated (e.g., by gels or HPLC) subsequent to sedimentation, which allows detection of interactions between components in extremely complicated mixtures. Typically, the accuracy of these methods is not sufficiently high to allow further analysis (i.e., determination of association constants).
aDesignates the instrument that may be used: A, analytical ultracentrifuge; P, preparative centrifuge with subsequent microfractionation. Where both types of instruments may be used, the better of the two is listed first. In general, the analytical ultracentrifuge offers improved accuracy in the determination of radial position and concentration, which is necessary for determining accurate association constants and stoichiometries. The preparative centrifuge offers lower cost and, in general, greater solute sensitivity and discrimination, but lower precision and accuracy. bPrecision of detection. Usually given as a standard error of replicates, the precision of a detection scheme may be very high, yet the accuracy may be poor if systematic error is present. E, excellent; G, good; F, fair; P, poor. cAccuracy of detection. This depends on the lack of systematic errors (e.g., nonlinearity of the response of the detector with concentration). dSensitivity to solute. eDiscrimination between solutes.
Analytical Ultracentrifugation
inants. Most of these methods are now highly refined, are easy to use, and can examine a wide range of sensitivities. However, with regard to sample characterization, they are less rigorous than sedimentation. This stems from the fact that all are secondary methods, requiring standards for calibration and comparison. For example, gel permeation chromatography is often used to estimate molecular weights, radii, and stoichiometries. However, if a protein does not conform to the properties of the standards (e.g., if the unknown is asymmetric and the standards are all spherical), erroneous conclusions may be reached with this approach. A second, more fundamental, problem arises when these methods are used to examine macromolecular interactions. Both chromatography and electropho-
resis usually entail the monitoring of thin zones of material. Because the concentration in the zone varies both spatially and with time, it is impossible to relate the zone shape or position to any association parameters in a rigorous fashion.
PLANNING AN EXPERIMENT The principle measurement obtained in any sedimentation experiment is the concentration as a function of radial position. Knowledge of the concentration distribution, along with the rotor speed and temperature, is needed to obtain a molecular weight or an s20,w. There are several means of determining the concentration distribution (Table 7.5.3). Analyses may be conducted using either a preparative or an analyt-
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ical ultracentrifuge. It is worth taking a moment to consider which instrument will be most useful for elucidating a particular problem. To do this, it is necessary to have a clear notion of what information is being sought from the centrifugation experiment. If all that is needed is a rough estimate of the molecular weight to confirm a stoichiometry, then a preparative centrifuge will be sufficient. On the other hand, if association constants or accurate sedimentation coefficients are needed, an analytical ultracentrifuge will be required. The biggest difference between an analytical and a preparative ultracentrifuge is the analytical instrument’s ability to monitor the concentration distribution as a sample is spinning. To do this, a special cell and rotor are used that permit light to pass through the sample. Light sources and detectors are arranged to monitor the cell contents throughout an experiment. Data acquisition is automated, and a large number (500 to 2000) of data points are available across the cell (1.5 cm), yielding the high radial resolution needed for the analysis of interacting systems. At present, absorbance and refractive detection are available for the XLA analytical ultracentrifuge (Beckman), and prototype fluorescence optics are under development. Data acquisition requires 5 to 30 sec per cell, and up to four cells may be used. The ability to acquire data rapidly and with high radial resolution has led to improvements in both sedimentation equilibrium and velocity analysis. For sedimentation equilibrium, there has been a resurgence of interest in short-column methods (Yphantis, 1960; Laue, 1992), whose small sample volumes (15 µl), rapid equilibrium (typically ∼1 hr), and large throughput (up to 16 samples simultaneously) are well-suited to addressing biochemical problems. The development of time-derivative sedimentation velocity has improved the sensitivity of the refractive optical systems nearly 100fold, and promises to revolutionize the range of systems suitable for sedimentation analysis. Experimental protocols in which the rotor speed is accelerated over the course of an experiment (i.e., gravitational sweep sedimentation) have been described (Stafford, 1992; Machtle, 1988) for analyzing and characterizing polydisperse samples. The fractionation range and accuracy of this method far exceed those of any other technique presently available. A good preparative ultracentrifuge (either a floor or tabletop model) may also be used for analytical ultracentrifugation, as confirmed by
the long-time success of sucrose gradient analysis. In many cases, these methods offer great advantages because the sample is analyzed after centrifugation by taking fractions (i.e., radial slices) from the centrifuge tube. Because of this, any detection scheme may be used to determine the concentration distribution. This means that unparalleled sensitivity and selectivity are afforded by schemes that use the preparative ultracentrifuge. Moreover, each fraction may be further fractionated—e.g., by high-performance liquid chromatography (HPLC) or gel electrophoresis—yielding good analytical sedimentation data for complicated mixtures. Detailed protocols for sedimentation velocity and sedimentation equilibrium in a preparative ultracentrifuge are available, and a superb microfractionator is available (from Brandel) specifically for conducting sedimentation analyses (Attri and Minton, 1986). However, it must be recognized that the information obtained from a preparative centrifuge is a snapshot of the cell at the time of deacceleration (assuming no mixing occurs during deacceleration). This precludes the use of the elegant time-derivative methods that have been developed for the analytical machine. Moreover, the time delay between stopping the run and sampling the cell allows diffusion to distort the concentration gradient, leading to inaccuracy in its determination. This problem becomes severe when steep gradients are involved. In general, the radial resolution for the preparative techniques is not as high as with the analytical ultracentrifuge. The concentration of protein needed for an experiment depends on what question is being addressed. If one merely seeks the molecular weight or sedimentation coefficient of a solute, then any convenient concentration may be used. However, if one is interested in knowing the association state of a sample under particular conditions (e.g., at concentrations found in vivo, or in an assay), the choice of detection scheme may become critical. In this case, one must work out what concentration is needed and what detection schemes are suitable for the concentration range, and match this to an entry in Table 7.5.3. It is essential to realize that any experimental protocol must examine the sedimentation behavior over a range of concentrations to take account of thermodynamic nonideality (sedimentation equilibrium) and hydrodynamic effects (sedimentation velocity; Laue et al., 1992). The stability and accuracy of the temperature is a critical experimental parameter. The
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Analytical Ultracentrifugation
XLA ultracentrifuge can be operated at temperatures between 0° and 40°C. Although this is convenient for most biochemical work, there are some studies (e.g., thermal denaturation) where a wider range is desired. In this case, one is limited to using an appropriately configured preparative centrifuge. Regardless of the temperature used, temperature stability is important in preventing convection. The accuracy of the temperature measurement is also important, particularly for sedimentation velocity analysis, where adjustment of a measured sedimentation coefficient (s) to standard conditions (20°C and water; the s20,w) must be made. Methods for making these adjustments are well established and have been automated (Laue et al., 1992). The rotor speed should be chosen carefully, as the gravitational field (g) increases as its square (g = ω2r, where ω = π × rpm/30 and r is the rotor radius). Different criteria are used to choose the rotor speed for sedimentation velocity and for sedimentation equilibrium analyses. For a sedimentation velocity experiment, two options can be considered: (1) gravitational sweep, in which the rotor speed is increased periodically throughout the experiment, and (2) s, where a single rotor speed is chosen for the analysis. Gravitational sweep is particularly useful for the initial characterization of a system where it is useful to get a general idea of how broad the particle size distribution is. In this method, the rotor is accelerated to a low speed (e.g., 3000 rpm) and several (six to ten) concentration distributions are collected to determine if there are any fast-moving boundaries. Next, the rotor speed is accelerated twofold, and data collection again undertaken. This process is continued until the maximum rotor speed (60,000 rpm) is attained, following which data are acquired until all boundaries have sedimented at least three-quarters of the way down the cell. Using this scheme, a sample can be “scanned” for the presence of an extraordinarily broad range of particle sizes (from greater than 1000S to ∼1S). The second method of analysis acquires data at a fixed rotor speed. This is useful for samples that have one or only a few types of proteins with similar sizes (within an order of magnitude). If one is trying to resolve two proteins of similar size, the highest rotor speed that allows a sufficient quantity of data to be obtained should be used (i.e., such that the fastest boundary takes ∼30 min to travel the length of the cell). The time-derivative data analysis routines work best with large numbers of data sets
closely spaced in time, so data should be acquired frequently throughout a run. Multiple rotor speeds should be used for sedimentation equilibrium analysis. Choosing appropriate rotor speeds is relatively simple if the molecular weight of the largest species is known (rpm ≈ 2 × 106 √ M−1 ), and a practical scheme for choosing an initial rotor speed is available for other cases. Low speeds are useful for detecting aggregates and high speeds are useful for detecting proteolytic fragments. In general, rotor speeds should be used that keep the reduced molecular weight, σ, between 2 and 12. An estimate of σ can be calculated from the molecular weight: _ 2 σ = M(1 − νρ) ω ⁄RT
_ where ν is the partial specific volume (∼0.72 ml/g is a good rough estimate), ρ is the solvent density (1 g/ml for the dilute aqueous buffers usually used in biochemistry), ω is the rotor angular velocity (see above), R is the gas constant (8.31 × 107 erg/mol o), and T is the absolute temperature. Even a crude estimate of σ will suffice. The range of rotor speeds used in an analysis should vary σ over a 4-fold or greater range, if possible. The experimental protocol should involve incrementing rather than decrementing the rotor speed, as (because of diffusion) it takes much longer for a rotor to reach equilibrium when approaching it from a higher speed. There are many other practical tips for choosing the rotor speed, estimating σ, and designing an appropriate experimental protocol (Ralston, 1993; Laue, 1992). In general, conducting an experiment with an analytical ultracentrifuge is very simple. The usual cause of difficulty is the sample, either in terms of its purity (for experiments trying to extract thermodynamic details) or its behavior (for previously uncharacterized biochemical systems). Problems often encountered for either sedimentation velocity or sedimentation equilibrium, along with suggestions for resolving them, are discussed in Table 7.5.4.
ANTICIPATED RESULTS Different, yet complementary, information is obtained from sedimentation velocity and sedimentation equilibrium experiments. For sedimentation velocity, the rate of movement of the boundary yields the sedimentation coefficient, s. Straightforward adjustment of this measured value provides s20,w, the expected sedimentation coefficient for the molecule if it were sedimented in pure water at 20°C (Laue et al., 1992). A second adjustment is made by
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Table 7.5.4
Troubleshooting Guide for Analytical Ultracentrifugation
Problem
Possible cause
Velocity method No boundary or very broad bound- Convection ary Sample heterogeneity (aggregates or fragments) Loss of sample to cell walls
Too few data points; boundary reaches base in <30 min Boundary that moves too slowly
Interacting system (concentrationdependent; may show pressure dependence) Rotor speed too high Rotor speed too low Presence of proteolytic fragments Improper sample dialysis
Solution May be caused by thermal instability or mechanical instability (rotor vibration) Clean sample by gel filtration Use centerpiece made of different material. Analyze by sedimentation equilibrium Shake cell and rerun at lower rotor speed Shake cell and rerun at higher rotor speed Fractionate sample by gel filtration Buffer component not equilibrated in sample and reference channels
Equilibrium method Decrease in molecular weight with increased concentration
Nonideality; little dependence of rotor Use a nonideal model in nonlinear least speed on molecular weight squares analysis; extrapolate to zero protein concentration. Information about molecular size and charge available. Fractionate sample Sample heterogeneity; molecular weight also decreases with increased rotor speed Characterize by nonlinear least-squares Increase in molecular weight with Mass-action association analysis increased concentration, but little change as rotor speed is varied Use sedimentation velocity analysis to Slow aggregation; flocculant Somewhat higher molecular weight than expected; slow loss of formation (data are often noisier than detect this as a fast boundary or sloping usual; inconsistent molecular weights plateau concentration. Only cure is to material find buffer conditions that stabilize are obtained from one sample to the protein solubility. For intracellular next) proteins, be sure a reducing agent (2ME or DTT) is present. Somewhat lower molecular Proteolysis (e.g., molecular weight Use protease inhibitors in sample; weight than expected drops with time) refractionate by gel filtration. Improper dialysis of sample prior to Be sure sample and reference buffers are analysis at dialysis equilibrium prior to run. Use centrifugal gel filtration for rapid equilibration. measuring the sedimentation coefficient of the boundary at several different protein concentrations, then extrapolating the values of s20,w to zero protein concentration to get s020,w. For a nonassociating protein, s020,w decreases with increasing concentration; this decrease occurs gradually if the protein is spherical (by ∼1% per mg/ml) and more strongly if it is asymmetric. In addition to gaining insight into the shape of a protein, there are three reasons for adjusting to standard conditions. First, s020,w is a practical, descriptive constant for a protein that can be used to compare preparations made under different conditions or in different laboratories.
Second, determination of s020,w provides a firstprinciple means of detecting changes in the hydrodynamics of the protein (see below). Third, an increase in s020,w with increasing concentration is a useful diagnostic for mass action association of a protein. Sedimentation velocity is also useful for characterizing more complicated mixtures of molecules (Stafford, 1992). In these cases sedimentation coefficient profiles are obtained. For samples containing only a few boundaries (paucidisperse samples), appropriate extrapolation of the data to infinite time provides higher precision and improved resolution of the
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boundaries (Van Holde and Weischet, 1978; Stafford, 1994). In addition to s020,w, the Stokes radius, RS, of a particle may be obtained from sedimentation analysis (Laue et al., 1992). Models can be applied in which RS is analyzed in terms of either the extent of hydration or the degree of asymmetry (Laue et al., 1992). Such analyses often provide the first insight into the shape of a protein. Equilibrium sedimentation provides thermodynamic information about a sample. Analyzed in the simplest way, the data can be used to obtain the molecular weight. For a mixture of molecules, various average molecular weights (number, weight, and z) may be calculated (Yphantis, 1964; Laue, 1995). By measuring the molecular weight in a denaturing solvent (e.g., 6 M guanidine⋅HCl; APPENDIX 3A) and in a p hysiological solvent, the stoichiometry of the native molecule can readily be determined (Laue, 1992). More detailed analysis of the concentration distributions can provide energies of assembly and nonideality coefficients (McRorie and Voelker, 1993).
TIME CONSIDERATIONS
Analytical Ultracentrifugation
For a typical run with the XLA ultracentrifuge involving three cells per rotor, sample preparation and cell loading requires ∼1 hr. For sedimentation velocity analysis, a run requires ∼2 hr (once the rotor is at the desired temperature). Gravitational sweep analysis requires somewhat longer. The time required to conduct a sedimentation equilibrium analysis is discussed below. In any case, operation of the centrifuge, including data acquisition, is completely automated. After the run, sample recovery and cell cleaning require ∼1 hr. For maximum efficiency, it is worthwhile to have two complete sets of cells so that one may be cleaned and loaded while the other is being run. Data analysis for sedimentation velocity experiments is computerized and rather straightforward, and usually requires <1 hr. The time required to conduct a sedimentation equilibrium experiment depends a great deal on the particulars of the experimental setup. The number of samples that can be analyzed simultaneously depends on the choice of centerpieces, so that up to sixteen samples may be analyzed simultaneously. Cell loading still only requires ∼1 hr. The time needed to reach sedimentation equilibrium depends on the solvent viscosity, the diffusion coefficient of the protein, and the distance between the meniscus and the base of the solution column. In general,
the time increases linearly with the viscosity, as the cube root of the molecular weight, and as the square of the column height of the solution. Shorter solution columns can be accommodated in any of the centerpieces. For a typical aqueous buffer and a 50,000-molecularweight protein, the time needed to reach equilibrium is ∼9 hr for a 3-mm solution column (∼110 µl of sample) and ∼1 hr using a short column centerpiece (∼15 µl of sample). Because a typical experimental protocol requires equilibrium at three or four rotor speeds, a full 1 or 2 days are needed to complete data acquisition. Computer programs have automated equilibrium data analysis. Even so, the time needed to analyze equilibrium data depends on the level of detail of the scrutiny. If only an average molecular weight is required, this can be accomplished in a few minutes. On the other hand, determination of both the stoichiometry and energetics of a self-associating protein may require several days of analysis if no previous characterization has been done. Both the level of detail required to answer a biological question and the experience of the researcher will affect the time needed to complete an analysis. For either velocity or equilibrium sedimentation, there are no time-critical steps once the centrifuge run has started. Zealots often begin data analysis during the centrifuge run. Although this is not strictly necessary, it is particularly useful when examining a previously uncharacterized system as it allows adjustment of the experimental conditions to maximize the quality of the data. Methods using a preparative centrifuge can handle a larger number of samples in a run (depending on which rotor is used). Likewise, the time required for preparation will vary, though it will be on the same order of magnitude as for the XLA. Unlike with the XLA, data acquisition is not fully automated in that the radius of each fraction must be calculated on the basis of its volume and the sample-tube geometry (Attri and Minton, 1986). Likewise, assaying the fractions may or may not require considerable time.
HELP FOR THE NOVICE It is widely recognized that undertaking sedimentation analysis for the first time can be a daunting task. Fortunately, the most difficult parts—including data acquisition, data analysis, and data interpretation—have been automated. Commercial computer programs for these tasks are available from Beckman Instru-
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ments. In addition, there is an Internet bulletin board ([email protected]) where analytical sedimentation is discussed and a wide variety of computer programs are available free of charge. Often, though, it is not the process of conducting analytical ultracentrifugation that is a problem for biochemists, but rather their uncertainty about the solution physical chemistry of proteins. Fortunately, the cadre of researchers working with the ultracentrifuge welcome the opportunity to help newcomers to the field. For further information contact the author by e-mail at [email protected].
Richards, E.G. and Schachman, H.K. 1957. A differential ultracentrifuge technique for measuring small changes in sedimentation coefficient. J. Am. Chem. 79:5324-5325.
LITERATURE CITED
Stafford, W.F. III. 1994. Sedimentation boundary analysis of interacting systems. In Modern Analytical Ultracentrifugation: Acquisition and Interpretation of Data for Biological and Synthetic Polymer Systems (T.M. Schuster and T.M. Laue, eds.) pp. 119-137. Birkhauser, Boston.
Attri, A.K. and Minton, A.P. 1986. Technique and apparatus for automated fractionation of the contents of small centrifuge tubes: Application to analytical ultracentrifugation. Anal. Biochem. 152:319-328. Cohen, R. and Mire, M. 1971. Analytical-band centrifugation of an active enzyme-substrate complex. 1. Principle and practice of the centrifugation. Eur. J. Biochem. 23:267-275. Harding, S.E., Rowe, A.J., and Horton, J.C. 1992. Analytical Ultracentrifugation in Biochemistry and Polymer Science. Royal Society of Chemistry, Cambridge. Laue, T.M., Shah, B., Ridgeway, T.M., and Pelletier, S.L. 1992. Computer-aided interpretation of sedimentation data for proteins. In Analytical Ultracentrifugation in Biochemistry and Polymer Science (S.E. Harding, A.J. Rowe, and J.C. Horton, eds.) pp. 90-125. Royal Society of Chemistry, Cambridge. Laue, T.M. 1992. Short Column Sedimentation Equilibrium Analysis for Rapid Characterization of Macromolecules in Solution. Technical Information DS-835. Beckman Instruments, Palo Alto, Calif. Laue, T.M. 1995. Sedimentation equilibrium as a thermo dy namic to ol. Methods Enzymol. 259:427-452. Mächtle, W. 1988. Coupling particle size technique. A new ultracentrifuge technique for determination of the particle size distribution of extremely broad distributed dispersions. Die Angew. Makromol. Chem. 162:35-52. McRorie, D.K. and Voelker, P.J. 1993. Self Associating Systems in the Analytical Ultracentrifuge. Beckman Instruments, Fullerton, Calif. Philo, J.S. 1994. Measuring sedimentation, diffusion and molecular weights of small molecules by direct fitting of sedimentation velocity concentration profiles. In Modern Analytical Ultracentrifugation: Acquisition and Interpretation of Data for Biological and Synthetic Polymer Systems (T.M. Schuster and T.M. Laue, eds.) pp. 156-170. Birkhauser, Boston. Ralston, G. 1993. Introduction to Analytical Ultracentrifugation. Beckman Instruments, Fullerton, Calif.
Schachman, H.K. 1959. Ultracentrifugation in Biochemistry. Academic Press, New York. Schuster, T.M. and Laue, T.M. 1994. Modern Analytical Ultracentrifugation: Acquisition and Interpretation of Data for Biological and Synthetic Polymer Systems. Birkhauser, Boston. Stafford, W.F. III. 1992. Boundary analysis in sedimentation transport experiments: A procedure for obtaining sedimentation coefficient distributions using the time derivative of the concentration profile. Anal. Biochem. 161:70-79.
Svedberg, T. and Pederson, K.O. 1940. The Ultracentrifuge. Clarendon Press, Oxford. Van Holde, K.E. and Weischet, W.O. 1978. Boundary analysis of sedimentation-velocity experiments with monodisperse and paucidisperse solutes. Biopolymers 17:1387-1403. Yphantis, D.A. 1960. Rapid determination of molecular weights of peptides and proteins. Ann. N.Y. Acad. Sci. 88:586-601. Yphantis, D.A. 1964. Equilibrium ultracentrifugation of dilute solutions. Biochemistry 3:297-317.
KEY REFERENCES Harding et al., 1992. See above. Excellent book containing descriptions of many of the modern methods of analytical ultracentrifugation. Schachman, 1959. See above. Classic work describing many useful methods still used today. Schuster and Laue, 1994. See above. Up-to-date collection of methods for analytical ultracentrifugation. Svedberg and Pederson, 1940. See above. A classic work that is still relevant in that it provides some of the clearest descriptions available of analytical centrifugation.
INTERNET RESOURCES www.bbri.harvard.edu/rasmb/rasmb.html Web site for most recent programs and discussion group on analytical ultracentrifugation.
Contributed by Thomas M. Laue University of New Hampshire Durham, New Hampshire
Characteristics of Recombinant Proteins
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Determining the CD Spectrum of a Protein
UNIT 7.6
There are two requirements for a molecule or group of atoms in a molecule to exhibit a circular dichroism (CD) spectrum. The first is the presence of a chromophore—i.e., a group that can absorb radiation by virtue of the electronic configuration of its resting or ground state at room temperature. The energy absorbed results in a transition to a higher-energy or excited state, which has a different distribution of electrons around the nucleus. It can therefore interact with its environment in a way that differs from the ground state. In proteins, tryptophan, tyrosine, and phenylalanine are the main chromophores in the near-UV (240- to 320-nm) region; the peptide bond is the main chromophore in the far-UV (180- to 240-nm) region. Disulfide bonds and histidine residues are two other chromophores whose contribution to CD are, in general, less marked. Most chromophores exhibit more than one transition (e.g., see Fig. 7.6.1, Fig. 7.6.2, and Table 7.6.2) and their spectra are a composite of several absorption bands. It is an important feature of CD that, unlike optical rotation, the wavelength region within which spectra are observed is limited strictly to the wavelength region of the individual absorption bands. This confers greater specificity on spectra and allows a certain degree of assignment to particular chromophores. The second requirement for CD is that the chromophore be in, or closely associated with, an optically asymmetric environment. The chromophores in proteins are themselves not chiral and exhibit no optical activity. The phenolic group of tyrosine, for example, exhibits a CD spectrum only because it is connected to an optically asymmetric carbon atom. However, when the same group is packed in a folded protein in an environment that is asymmetric with respect to polarity, or is interacting through the phenolic hydroxyl, it exhibits a different CD spectrum that is specific to the particular environmental influences acting on the chromophore. In practice, the latter spectrum is usually much more intense than that for free tyrosine and may also differ in shape. When the peptide-bond chromophore is part of a regularly folded structure, with particular backbone angles and hydrogen-bond interactions as in an α-helix or β-sheet, it becomes closely associated with a conformationally asymmetric structure—i.e., one that cannot be superimposed on its mirror image. A CD spectrum for the protein will be generated that is a composite of the individual spectra corresponding to each of the peptide-bond absorption transitions (Fig. 7.6.1). The intensity of the spectrum may be further affected by interaction between neighboring chromophores (Bayley, 1980). Plane-polarized radiation comprises two circularly polarized vectors of equal intensity, one right-handed and the other left-handed (Fig. 7.6.3A), which are separately measured in the CD spectrometer by means of a photoelastic modulator. A chromophore situated in an optically symmetrical environment will normally absorb the two components equally so that, when recombined after passing through a solution of the chromophore, they result once again in radiation oscillating in a single plane. A chromophore situated in an optically asymmetric environment, however, will absorb each of the two components to a different extent, the difference being ∆A. When recombined, the resultant vector describes an ellipse (Fig. 7.6.3B), the ratio of whose major and minor axes determines the ellipticity. The value of ∆A, and hence that of ellipticity, can be positive or negative depending on the nature of the asymmetric environment. This is exemplified by the contribution of the tyrosines in IL-1β (Fig. 7.6.4A) and of the tyrosines and tryptophans in GM-CSF (Fig. 7.6.5). It is a general consequence of the above principles that CD spectra of molecules in solution are located in the same wavelength region as their absorption bands. For proteins this means the far-UV and near-UV regions, as well as regions extending into the visible and near infrared. These regions have their origin in and provide information about the Contributed by Roger Pain Current Protocols in Protein Science (1996) 7.6.1-7.6.23 Copyright © 2000 by John Wiley & Sons, Inc.
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1
A perpendicularly polarized π0 π – band Absorbance
Figure 7.6.1 Relation between absorbance and circular dichroism for a polypeptide chain in the helical conformation. (A) The absorbance spectrum of poly-γ-methyl-L-glutamate in the α-helical form, with an assignment of its three constituent transitions. (B) The far-UV CD spectrum of the same compound, deconvoluted on the assumption that the shapes of the absorbance and dichroic bands are directly related (broken lines). The solid line is the observed spectrum and the filled circles the sum of the three bands. In general, the sign of a CD band may be either positive or negative, and its intensity does not necessarily follow that of the absorbance band. (From Holzwarth and Doty, 1965.)
parallel polarized π – band π0
n1
π – band
0 10
B
8
[θ]mrw x 10–4 (deg cm2 dmol–1)
6
4
2
0
–2
–4
180
200
220 240 Wavelength (nm)
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0.4 272 278 282 289.5
Absorbance
0.3
1L a
0.2
0.1 1L b
0 230
240
250
260 270 280 Wavelength (nm)
290
300
310
Figure 7.6.2 The two near-UV absorbance transitions for tryptophan. The near-UV absorption spectrum for N-stearyl tryptophan n-hexyl ester dissolved in methylcyclohexane at 24°C (solid line) has been resolved into the 1La and 1Lb bands. (Drawn from Strickland, 1974.)
polypeptide backbone and its conformation (far-UV), the aromatic amino acid residues and their environments (near-UV), and bound ligands such as heme and cofactors (visible and near infrared). Experimentally, the near-UV and the visible and near-infrared regions can be treated together and the protocols in this unit will therefore refer explicitly only to the collection of far- and near-UV spectra. In both the far- and near-UV regions, CD spectra can be used empirically as “fingerprints” of a particular protein, with the spectrum resulting from the aromatic residues being rather more specific and hence diagnostic. The far-UV spectra, however, can provide information about the protein conformation in terms of its secondary structure. As for fluorescence spectroscopy or any spectroscopic method, the sample needs to be chemically pure and homogeneous. The output of CD spectrometers is in two alternative but related types of unit. The difference in absorbance with respect to left- and right-handed circularly polarized radiation (AL − AR = ∆A) is measured in absorbance units; ellipticity (θ) is measured in millidegrees (mdeg). These units are related by the expression θ = 33,000 × ∆A (see Fig. 7.6.3A). Ellipticity is usually used for far-UV measurements. Because essentially every residue in a protein is associated with a peptide bond, spectra in this region can be directly compared for different proteins by calculating the mean residue ellipticity, [θ]mrw =
θ ×Mmrw 10 ×c ×1
where c is the protein concentration in mg/ml, l is the cell path length in cm, and Μmrw is the mean residue molecular weight—i.e., protein formula weight/number of residues. The dimensions of [θ]mrw are deg cm2/dmol, and the quantity is independent of molecular weight. Μmrw is in the range 110 to 115 for most proteins.
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A
B I
L′ L
I′
R
O
O
R′
Figure 7.6.3 The relation of ellipticity to the differential absorption of circularly polarized radiation. The oscillating radiation sine wave, OI, is proceeding out of the plane of the paper towards the viewer. (A) Plane-polarized radiation is made up of left- and right-handed circularly polarized components, OL and OR, respectively. Absorption by a chromophore in a nonchiral environment results in an equal reduction in intensity of each component, whose resultant is a vector oscillating only in the vertical plane—i.e., plane-polarized radiation. (B) Interaction of the radiation with a chiral chromophore leads to unequal absorption, so that combination of the emerging vectors, OL’ and OR’, leads to a resultant that describes an elliptical path as it progresses out of the plane of the paper. The ratio of the major and minor axes of the ellipse is expressed by tan θ, thus defining ellipticity. The major axis of the ellipse makes an angle (φ) with the original plane, which defines the optical rotation. This figure thus demonstrates the close relation between optical rotation and circular dichroism.
For example, for serum albumin (see Fig. 7.6.6), the ellipticity measured at 209 nm using a 1-mm path-length cell for a solution containing 0.361 mg/ml is −61.7 mdeg. Taking a mean residue molecular weight of 110, [θ]mrw = −18.8 × 103 deg cm2/dmol. For small molecules, such as aromatic amino acids and model compounds, the molar difference absorption coefficient is frequently used. By analogy with absorbance spectroscopy, this quantity is given by ∆ε = ∆A/(c × l), where c is the concentration in mol/liter, l the path length in cm and ∆ε has the units liters/mol/cm (or cm2/mmol). However, in the case of proteins, near-UV CD spectra are made up of contributions from tryptophan, tyrosine, and phenylalanine residues—the content of all of which vary from protein to protein. It is therefore confusing—and achieves no useful end—to present results in terms of mean residue ellipticity. Instead, dichroism should be presented as values of ∆A. For a given protein, different samples may be compared by calculating ∆A/(c × l), with c in mg/ml. Owing to the variety of units found in the literature and in software—e.g., mdeg and deg for ellipticity, and mm, cm or dm for path length—it is necessary to take care that experimental parameters be entered in the units requested by the machine software.
Determining the CD Spectrum of a Protein
The Strategic Planning section deals with considerations regarding instrumentation and reagents for CD spectrometry. The Basic Protocol outlines the steps in recording a CD spectrum. The two support protocols explain the interpretation of CD spectra—Support Protocol 1 deals with near-UV spectra and Support Protocol 2 with far-UV spectra.
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0.5
A
Tyr 121
0 Tyr 24 ∆) x 10 "
Figure 7.6.4 The contributions of different aromatic residues in a folded protein to circular dichroism. Interleukin 1β contains one tryptophan and four tyrosine residues. Their individual contributions have been resolved using site-directed mutagenesis. (A) The near-UV CD spectra of the tyrosine residues show that, although Tyr 68 makes a significant contribution to the negative ellipticity, indicating its buried, asymmetric environment, the contributions resulting from residues 24 and 90 are little different from that of free tyrosine. The derived spectrum of Tyr 121 is interesting in that it clearly contains a contribution arising from tryptophan (Fig. 7.6.2), indicating its close interaction with Trp 120. (B) The contribution from the tryptophan is seen to dominate, reflecting its nonpolar environment. The intensity of the protein spectrum is less intense than it might otherwise be, because of the opposing signs of the individual spectra. Spectra were measured using a 1-nm bandwidth (Craig et al., 1989).
Tyr 90 –0.5
Tyr 68
–1.0 3
B Trp 120
∆) x 10 "
2
1
0 Tyr 68
–1 250 260 270
280 290
300 310 320
Wavelength (nm)
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0
A
U –1 R N ∆ A x 10 4
Figure 7.6.5 The near-UV CD spectra of granulocyte/macrophage-colony stimulating factor (GM-CSF). (A) Comparison of the spectrum of the native (N) murine GM-CSF (with one tryptophan residue, which is exposed, and four tyrosine residues, 2.5 of which are exposed) with those for the protein unfolded in 3.25 M guanidine⋅HCl (U) and refolded from this solution (R) by dialysis against 3 M urea. The intense negative spectrum of the folded protein, with a maximum at 278 nm, is characteristic of tyrosine, with contributions from phenylalanine. The intensity is probably a result of the interaction of Tyr 81 and Tyr 82. The absence of a band at 292 nm shows that the Trp 14 makes little contribution, as expected from its exposed location. Refolding restores the detailed tertiary structure of the native protein. (B) Reduction (RED) of native human GM-CSF (with two tryphtophan residues, one of which is exposed, and two tyrosine residues, which are buried) reduces the ellipticity of the native protein (N) to that of the fully unfolded protein (U). The secondary structure is however, unaffected (see inset for far-UV CD spectra). Notice that, in contrast to the murine protein, human GMCSF exhibits a less intense, but positive, near-UV CD band at 290 nm with little fine structure, implying a contribution from a 1La transition of Trp 123 and indicating the buried nature of this residue. Spectra were measured using 0.01-mm path-length cells and 2-nm bandwidth in the far-UV, and 10-mm path-length cells with 1-mm bandwidth in the near-UV (Wingfield et al., 1988).
–2
–3
–4
B 1
N
∆ A x 10 4
U 0
[θ] mr w x (10 – 3/deg cm 2 dmol –1)
RED
–1
–2 250
0
RED N
200
260 270 280 290 300 310 Wavelength (NM)
250 320
330
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40
[θ]mrw x 10 –3 (deg cm2 dmol –1)
30
20
10
0
–10
–20 180
190
200
210
220
230
240
250
Wavelength (nm)
Figure 7.6.6 Characteristic far-UV CD spectra of proteins containing high contents of helix (dotted line) and β-sheet (solid line) structures. Protein concentrations were 0.36 mg/ml human serum albumin and 0.03 mg/ml immunoglobulin G. Spectra were determined using a 1-mm path-length cell. Each spectrum is the smoothed average of four repeat scans, measured with 1-nm bandwidth, reading ellipticity every 1 nm with an averaging time of 5 sec—i.e., at a scan speed of 12 nm/min.
STRATEGIC PLANNING CD Spectrometer CD spectra are measured using a CD spectrometer, of which there are three different makes on the market (AVIV Associates, Instruments SA, and Jasco; see SUPPLIERS APPENDIX). These instruments are fitted for control, data collection, and data handling by computer. The principles of how they function are generally outlined in the manufacturers’ manuals and a good general account is given in Bayley (1980). A supply of high-purity nitrogen is essential to displace oxygen, in order to avoid degradation of the mirrors by ozone generated by the high-power xenon sources as well as to reduce absorbance from oxygen bands below 200 nm (see discussion of Nitrogen Supply, below). A satisfactory means of thermostatting cells is essential for work with proteins (see discussion of Cells, below), using a water-circulation system or the more convenient Peltier thermostat. The CD spectrometer is usually required to work near the limits of sensitivity—e.g., reading ∆A values of ≤10−4 at a total absorbance of 1. Thus, particular care needs to be taken with cleanliness and orientation of cells and with settings of scan rate, time constant, and bandwidth. It is also important, especially when recording far-UV spectra, that the lamp is not old and that the mirrors are not clouded from radiation and traces of ozone. Because the spectrometer is a single-beam instrument, it is essential always to watch carefully for evidence of instrumental drift during measurements of sample and baseline. Characterization of Recombinant Proteins
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[θ] x 10 –3 (deg cm2 dmol –1)
8
0
–8
–16 150
200
250 Wavelength (nm)
300
350
Figure 7.6.7 CD spectrum of D-(+)-10-camphorsulfonic acid (CSA) in water. The vertical bars represent variations of ±1.5% and ± 5%. The broken line represents the extrapolation of a gaussian band. Commercial CSA was twice recrystallized. (From Chen and Yang, 1977.)
Calibration of the Spectrometer Depending on the make of the spectrometer, wavelength calibration may require the use of a mercury lamp and the services of an engineer, or the use of a holmium oxide solution with a simple computer-operated adjustment. It is unlikely to vary significantly during the lifetime of a xenon lamp. Calibration of ellipticity, on the other hand, must be carried out regularly—at least monthly, if the machine is in regular use—using a purified compound of known absorbance and ellipticity. D-(+)-10-camphorsulfonic acid (CSA) is the substance of choice (Fig. 7.6.7), but care must be taken to purify it from optically active impurities. Crystallization from ethyl acetate is recommended. CSA is hygroscopic; hence, although a stable monohydrate can be prepared (Yang et al., 1986), the concentration of the aqueous solution (∼1 mg/ml) should be determined from A2851 mg/ml = 0.149 using a high-quality spectrophotometer. Calibration of the spectrometer is carried out according to the maker’s instructions, using θ = 33.5 mdeg at 290.5 nm for a solution of 1.00 mg/ml anhydrous CSA in a 1-mm path-length cell. CSA exhibits a second peak at 192.5 nm where θ ≈ −67.0 mdeg for the same solution. If the value of the ratio of ellipticity at the two wavelengths falls below 2.0, this indicates a loss of performance of the spectrometer. Epiandrosterone, at 304 nm, and D-(−)-pantoyllactone, at 219 nm, have also been used for calibration (Schmid, 1989). The peaks of the spectra scanned for calibration are sufficiently sharp to provide a check on the wavelength calibration. Cells
Determining the CD Spectrum of a Protein
Choice of cells Quartz windows can exhibit dichroism, which is due to strain either remaining after the annealing process during manufacture or induced by distortion caused by heating or pressure. It is important for measurement of CD that the optical faces of the quartz QS cuvettes have low strain and exhibit low dichroism. In the near-UV region—where path lengths ≥5 mm are required—rectangular cells, in their standard, semimicro, or micro forms, are normally used. These are not only more economical with regard to protein than the cylindrical type, but allow mixing to be carried out in the cell. Testing a number of
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rectangular cells in the CD spectrometer usually yields one or two with low dichroism; it is also possible to purchase cells with certificates of low dichroism. Careful observation of the height of the light beam at the cell allows the minimum volume of solution to be used—usually <1 ml in a 10 × 10–mm cell. Because standard quartz (Suprasil) cells are constructed from two different types of glass, differential expansion of the faces with temperature leads to a significant change in cell dichroism. At least for temperature-dependence experiments, it is recommended to use cells with all four faces made from fused quartz. Semimicro cuvettes with 1-cm path length are available; these require ∼300 µl of solution. For far-UV CD spectra, short-path-length cells—from 1 mm down to 0.01 mm—are necessary because of the higher absorbance and ellipticity in this region. Demountable cells, cylindrical or rectangular, of this size are easily cleaned, but require practice to fill without allowing evaporation and may be subject to solvent loss over long periods of time. Such leakage can be minimized by the use of Parafilm on the edges of the cells. Quartz-jacketed cylindrical cells with two filling holes, available from Hellma (see SUPPLIERS APPENDIX), provide the best economy of protein, temperature control, and reproducibility. The path length of cells should be checked by absorbance measurements. A solution of 300 mg/liter of potassium dichromate in 0.05 M KOH gives an A280 of 0.705 in a 1-mm cell. Preparation and handling of cells Cells must be reproducibly free from external and internal contamination. Not only will contamination make a contribution to the measured ellipticity, but dirty surfaces cause bubbles to be trapped in short-path-length cells, leading to additional artifacts. Cleaning should always be repeated if there are any signs of “tailing.” The short-path-length cells, being inaccessible to mechanical cleaning, have to rely on effective soaking and rinsing. Cuvettes must be thoroughly and reproducibly cleaned by soaking in a proprietary cuvette cleaning fluid such as Hellmanex II (Hellma), an alkaline liquid concentrate, or RBS-35 Detergent Concentrate (Pierce), following the makers’ instructions. The makers of Hellma cuvettes caution against the use of ultrasonics as a means of boosting cleaning efficiency. 50% nitric acid is also effective, but requires care and a fume hood. Detergents frequently contain fluorescent material and, if used, should be removed from cuvettes with acid (2M HCl) followed by water. After the final rinse with copious amounts of high-quality distilled water, the cells are best dried using suction through a Pasteur pipet protected with a small length of fine plastic tubing. This is safer than using acetone or alcohol, which can leach out fats from fingers and deposit them on the cuvette. The faces of cuvettes must not be touched after cleaning. Filling, emptying, and rinsing between measurements of spectra must be done using pipets, to avoid spillage onto the outer faces. A Pasteur pipet protected with fine plastic tubing is satisfactory. The baseline scans, with buffer in the cell, must be examined as a constant check on the reproducibility of the state of the cell. Buffer Solutions Buffers must be optically inactive and have the lowest possible absorbance in the desired spectral region. Most buffers in common use are essentially transparent above 230 nm, which is fine for near-UV CD. For far-UV CD using a 1-mm pathlength cell, 0.01 M sodium phosphate buffer (APPENDIX 2E), is generally acceptable as a solvent for most purposes, and will still have a reasonably low absorbance between 190 and 185 nm—the lower the pH, the more transparent the buffer. NaCl should be avoided in the far-UV range. Solutions of 0.01 M NaClO4, NaF, KF, or boric acid are all transparent down to the present range of commercial instruments, but are not frequently used except for vacuum UV measurements. The absorbance of 20 mM Tris⋅Cl is acceptable down to 190 nm. Buffers
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in common use, together with their absorbances, are listed by Schmid (1989). Urea or guanidine⋅HCl solutions (high-purity spectroscopic grades must be used) absorb very strongly below 220 nm; even in the shortest-path-length cells, they are opaque below 205 nm. Bubbles form readily in the cell and can distort the CD. It is important therefore to degas the buffer before filling the cell by applying vacuum in a suitable container, such as a Buchner funnel, and swirling. Clarification of Solutions Protein solutions must be freed from the turbidity that results from dust and aggregated protein. Turbidity will reduce the sensitivity of the measurements and can also cause artifacts in both the intensity and the shape of the CD spectrum, particularly in the far-UV (Swords and Wallace, 1993). Many proteins can be filtered, usually as a stock solution, using a 0.22-µm filter (ideally of the grade special for proteins) fitted to a syringe. If the protein is adsorbed by filters, the solution may be clarified by centrifugation; 10 min in a refrigerated microcentrifuge is convenient and usually effective. Note that the protein concentration must be measured after clarification. Buffer solutions are filtered on a larger scale, using a 0.22-µm filter. It is essential to monitor clarification if artifacts are to be avoided. Indicators of adequate clarification that should be adopted for spectroscopic characterization are available from the procedures basic to the measurement of CD. First, determination of protein concentration requires an absorbance spectrum to be recorded on a good quality spectrophotometer from 240 to 350 nm. Aromatic amino acid residues do not absorb above 320 nm, so the spectrum between 320 and 350 nm should be only marginally above baseline. The presence of turbidity will result in a sloping spectrum in this region. From the absorbance spectrum obtained in this manner, the ratio Amax/Amin is calculated, where Amin is the absorbance in the trough in the region of 240 to 250 nm. This ratio will be specific for a given protein and should be measured for a sample that has been carefully and reproducibly clarified. Because of the strong wavelength dependence of scattering, the ratio is a very sensitive indicator, decreasing sharply with turbidity, and can be used as a semiquantitative check on the clarity of subsequent solutions of the protein. For membrane proteins in membrane fragments, the intrinsic turbidity offers special problems, which should be addressed by instrumental modifications. A useful discussion of the problems involved can be found in Swords and Wallace (1993). Nitrogen Supply Oxygen-free nitrogen can be obtained from cylinders, but must be filtered to remove oil contaminants. Liquid nitrogen provides a better supply of gas, but impurities accumulate at the bottom of the Dewar flask, so that the bottom 10% should not be used (Johnson, 1990). Accessories to monitor and purify nitrogen gas are available commercially. BASIC PROTOCOL
Determining the CD Spectrum of a Protein
RECORDING A CD SPECTRUM Simple comparison of the CD spectra of a protein in buffer and in 6 M guanidine⋅HCl will immediately tell whether the protein has achieved some degree of folding. However, for the purpose of determining whether a sample of recombinant protein is correctly folded, the CD spectrum can provide a complex fingerprint that can be compared with that of a specimen that has been well authenticated as being native and functional. This constitutes an empirical, but powerful means of characterization, with the near-UV spectrum usually being more sensitive than its far-UV counterpart. There are, for example,
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many well-documented cases where partially unfolded proteins exhibit a major change in the aromatic region accompanied by relatively minor changes in the far-UV CD (e.g., see Fig. 7.6.5)—the classical molten globule being an important example (Ptitsyn, 1992). For these reasons, it is important to obtain good quality near-UV spectra, paying particular attention to details of any fine structure that may exist. Materials CD spectrometer (calibrated), nitrogen supply, cells, buffer solution, and clarified protein solution for analysis (see Strategic Planning) 1. Set up the CD spectrometer by purging the optics with nitrogen in the manner designated in the manual, turning on the cooling water, and finally switching on the lamp. Allow to warm up for the recommended period of time, usually 30 min. Regulate the thermostat system to the desired temperature. Particular attention must be paid to the procedure in the manual for switching on and warming up the spectrometer. Failure to purge with nitrogen can lead to rapid and irreversible degradation of the optics. CD spectra, particularly in the near-UV (see Support Protocol 1) reflect the dynamics of the chromophore and may therefore show dependence on temperature. It is important to stabilize the temperature reproducibly. Accurate temperature control is particularly important in denaturation experiments to determine protein stability.
2. Enter the settings required for the scan. Careful optimization of the settings is necessary to achieve the lowest-noise spectra consistent with total time spent in data collection. To check that optimal settings have been achieved, a small sample region of the spectrum can be scanned. Table 7.6.1 is a rough guide for scanning a protein solution with A280 = ∼1. Settings will differ according to the sample and the spectrometer.
a. Set wavelength. For near-UV spectra, the scan should be from 240 to 340 nm, in order to cover contributions from tryptophan, tyrosine and phenylalanine. Although little or no dichroism will be found above 320 nm, it is advisable to extend the scan to 340 nm to ensure that both sample and baseline scans coincide at zero ellipticity in the region where there is no dichroism. This provides assurance that the cell and buffer are optically identical for the two scans (see Fig. 7.6.8). For far-UV spectra, it is usual to scan between 250 nm—where the ellipticities of the baseline and sample effectively coincide, given the scale of ellipticity being used—and as
Table 7.6.1 A280 = ∼1
Rough Guide for Scanning a Protein Solution with
Cell path length (cm) Wavelength (nm) Bandwidth (nm) Averaging time (sec) or Time constant (sec) Repeat scans Step size (nm) Resulting time per scan (min)
Far-UV
Near-UV
0.01–0.05 250–180 0.5–1 1
1 340–250 0.2–0.5 1–5
2–8 1–2 0.5–1 70–140
8 2–4 0.2–0.5 3–37
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15 corrected spectrum
–15
buffer
6 protein
–30
protein
∆A x 10 5
∆A x 10 5
0
0 280 –45 250
260
270
280
290
285 300
290 310
295 320
Wavelength (nm)
Figure 7.6.8 Obtaining the corrected near-UV CD spectrum for hen egg white lysozyme. The protein and baseline spectra were collected using a 10-mm cylindrical cell and 0.5 mg/ml protein in 0.067 M phosphate buffer, pH 6.0. Instrument settings were 1-nm bandwidth, 0.2-nm step size, scan speed 2 nm/min, time constant 8 sec (scan speed × time constant = 0.27 nm). Protein solution and buffer were scanned once each. The spectra were smoothed, a sample of the fit being shown in the inset. Reproducibility of the instrument and of the state of the cell are demonstrated by the coincidence of the ellipticity above 300 nm. The corrected spectrum was obtained by subtraction, using the instrument software.
near to 180 nm as the absorbance of the solution will permit. The lower limit may be dictated by the absorbance of the sample or buffer, as indicated by the value of the dynode voltage (see step 2e).
b. Set bandwidth. The radiation passing through the sample comprises a spread of intensities distributed around the selected wavelength (Fig. 7.6.9). The bandwidth (in nm) of this spread is determined by the spectrometer slit width (in mm) and is the width of the distribution at half the maximum intensity. For proteins exhibiting broad CD bands, including most far-UV and many near-UV spectra, a wide setting (2 nm) will have little effect on the shape or intensity of the spectrum. For bands with fine structure, including some near-UV spectra, a more narrow bandwidth is necessary to avoid smoothing of narrow peaks and reduction in peak intensity. The largest bandwidth that causes no distortion of the spectrum should be used, in order to maximize the signal-to-noise ratio. To optimize the slit-width setting, preliminary scans should be made at a series of slit widths over a critical region of the spectrum.
c. Set averaging time or time constant, scan speed, and number of accumulations. The time constant determines the time that the spectrometer takes to respond to changes in signal. The reason for setting it is to obtain the best signal-to-noise ratio consistent with the time spent scanning. When the signal is strong, the time constant need only be small. The appropriate time constant for a signal of 500 mdeg is 0.25, that for a signal of 100 mdeg is 0.25 to 0.5, that for a signal of 10 mdeg is 0.5 to 0.8, and that for a signal of 2 mdeg is 2 to 30. Determining the CD Spectrum of a Protein
The faster the scan speed (with economies in operator time), the smaller the time constant must be in order to allow the instrument to respond fully to changes in ellipticity. However, less time is then available for collecting data at a given wave-
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Relative intensity
1
0.5
b
0 Wavelength
Figure 7.6.9 The distribution of intensity of radiation passing through the sample cell. Radiation emerging from a monochromator set at a wavelength corresponding to the peak maximum will contain components at other wavelengths, with intensities described by the curve. The bandwidth (b) is the width of the distribution curve at half the peak height. Its magnitude will depend on the exit slit width. The setting of the slit width is usually programmed to change during scanning, so as to give a constant bandwidth and hence constant spectral resolution.
length, so the noise is greater. Scan speed and time constant are thus interdependent, and their product should not exceed 0.33 nm. For example, a time constant of 2 sec is recommended for a scan speed of 10 nm/min. Some instruments, in the wavelength scan mode, specify an averaging time, which takes the place of a time constant setting; this is simply the time spent collecting data at each wavelength. The signal-to-noise ratio can be enhanced either by increasing the time constant or averaging time or by accumulation of a number of scans. The latter technique is preferable to using long scan times because the effect of instrument drift will be to distort the spectrum from a single long scan, whereas the average of quicker, repeat scans will lead to a more uniformly shifted spectrum. The latter can then be corrected using the average of buffer baselines measured before and after the sample scans. The problem of optimizing signal-to-noise ratio is greater in the case of samples possessing a low ellipticity. An all-β protein, for example, will demand more repeat scans than an all-helix protein at the same concentration. There are realistic time limits to the degree of enhancement of signal-to-noise ratio, as the latter is proportional to the square root of both the time constant and the number of scans (see Yang et al., 1986).
d. Set step size. The step size (in nm) between successive data-collection wavelengths, together with the scan speed, defines the number of data points collected per nm. Spectra with more fine structure require higher resolution. For far-UV spectra, which are smooth and relatively featureless, a step size of 0.5 nm is adequate. For some, near-UV spectra, but not all, a resolution of 0.2 or 0.1 nm is advisable. For a given scan speed, the greater resolution gained by using a small step size will result in more noise, so it is more economical of time to use as large a step size as possible.
e. Note dynode voltage. Dynode voltage cannot be preset, but its value can be controlled within limits by adjusting the protein concentration and/or cell pathlength, so it is appropriate to consider it here.
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The photomultiplier detector operates with a high voltage in the region of 1000 V across the anode and cathode. When light falls on the photosensitive cathode, current flows and the potential difference is reduced. Readings of the dynode voltage thus give a measure of the intensity of light passing through the sample. Higher values indicate low levels of light (high sample absorbance) with consequent decrease in signal-to noise-ratio; very high values indicate incorrect readings. Values of dynode voltage should always be noted during a scan in order to assess the reliability of the ellipticity measurements. Most instruments record this parameter as a matter of course. Special care must be exercised in the far-UV region, where sample and buffer absorbance can rise sharply with decreasing wavelength.
3. Fill the cuvette and place it in the cell holder, making sure that the orientation is reproducible both longitudinally and rotationally. Mark the cell to allow reproducible orientation in the light beam. Techniques for filling the cell will depend on the type and path length of cell being used. It is essential in all cases to avoid the presence of small bubbles, which can easily be trapped at the optical face, especially with shorter-path-length cells. Even slight contamination of the faces will make this difficult, necessitating frequent cleaning of the cell after contact with certain proteins and with buffer components such as guanidine⋅HCl. For rectangular-type cuvettes, the volumes of solution required are approximately 2.5 ml (for 1-cm standard), 1 ml (for 1-cm semimicro), 300 ìl (for 0.1-cm standard) and 25 ìl (for 0.01-cm demountable). Volumes for cylindrical-type cells are somewhat larger, depending on the type. Choice of path length will depend on the available concentration of the protein solution. If the absorbance of the buffer is unavoidably high, it will be advantageous to use a shorter path length and a higher protein concentration. As a general guide, for a protein solution with A280 = 1, path lengths of 0.01 to 0.05 cm for the far-UV and 1 cm for the near-UV make suitable starting points. Rectangular cells have the advantages over cylindrical cells of having smaller sample volume and better temperature control with available cell holders. Good temperature control with cylindrical cells requires the use of jacketed cells. For shorter (<0.1-cm) path lengths, where rectangular cells are currently demountable, the rigid cylindrical cells have the advantage of avoiding leakage over prolonged periods of time.
4. Scan the baseline using the same cell and buffer as required for the sample, as well as the same instrument settings. Preliminary scans of the samples should be carried out to assess whether valid measurements can be made in the required wavelength region or whether it is necessary to adjust the sample concentration or the cell path length. The wavelength is lowered, noting the point at which the photomultiplier voltage exceeds the value corresponding to low light intensity—i.e., to an absorbance of 1 to 1.5. Experience will indicate the value above which noise becomes excessive. It is helpful, however, to get a more objective assessment by noting the voltage at a series of low wavelengths and then measuring the actual absorbance of the same cell using a quality spectrophotometer. When recording spectra for a series of samples, buffer scans should be recorded at least twice (at the beginning and end of the series) in order to check for instrument drift, especially when the lamp is beginning to age. A convenient procedure when scanning a number of samples is to scan air blanks—i.e., with no cell in the light path—at appropriate intervals between samples. This saves additional cell filling, washing, and drying operations. A single scan (S) of the cell containing buffer accompanied by a scan of the air blank then allows correction of all the protein spectra for solvent baseline, according to the expression:
{Sprotein − Sair} − {Ssolvent − Sair} = {Sprotein − Ssolvent}. Determining the CD Spectrum of a Protein
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Ideally, the buffer scan should be run before the protein sample, as this requires less rigorous washing of the cell between the two runs and hence better assurance that the optical surfaces will be identical for both. This however requires prior knowledge of the settings appropriate to the particular sample. It is tempting to use water in place of buffer on the grounds that both are optically inactive and, for water, the cell needs only to be dried between scans. This is unwise, however, particularly for buffers with higher concentrations of solutes and hence higher refractive index. Because in most instruments the light beam is not parallel when passing through the cell, the area and position of the photomultiplier surface illuminated is likely to vary with the refractive index of the solution, which may give slightly different voltage readings. This may result in significant errors when the signal is low, as in the case of near-UV CD.
5. Remove the buffer solution, rinse the cell with water and dry, taking care not to contaminate or touch the outer surfaces, refill the cell with the clarified sample solution, and scan the sample using the same instrument settings as for the buffer. Because washing, drying, and filling the cell takes time, it is advisable to work with two cells alternately (both with solvent blanks properly determined) when processing a number of samples. The protein concentration should be as high as possible, consistent with the absorbance in the cell not exceeding 1.0 to 1.5, and must be accurately determined by UV absorbance. Values of mean residue ellipticity and secondary structure content results are both affected by uncertainties in protein concentration. In spectral regions such as the far-UV, where absorbance varies markedly with wavelength, the spectrum may have to be recorded in overlapping regions, at appropriate concentrations or cell path lengths. A good test for the absence of artifacts that can arise at higher absorbances is to run a blank with a non–optically active molecule, such as 3-methylindole or a DL–amino acid, at the same absorbance as the test samples. The scan should be indistinguishable from the solvent blank.
6. Subtract the baseline from the sample scan. The ellipticity of the spectrum corrected for baseline should be close to zero above 320 nm for near-UV and above 250 nm for far-UV spectra (see Fig. 7.6.6 and Fig. 7.6.8). Significant differences usually indicate problems with reproducibility of the buffer or cell and must be remedied for quantitative characterization of samples. Depending on the level of noise, spectra and baselines can be smoothed using the instrument software. Smoothed and raw spectra should be compared directly by overlaying on the computer screen in order to avoid the distortion that can arise from overenthusiastic smoothing.
INTERPRETATION OF NEAR-UV CD SPECTRA CD spectra reflect, in different ways, the detailed conformational environment of individual chromophores in a protein. To the extent that the transitions for the different types of chromophore differ in wavelength, the spectra can be assigned to particular types of residue and, in the case of the peptide bond, to different conformations or types of secondary structure. A brief account of the interpretation of near- and far-UV spectra is therefore pertinent. Near-UV CD spectra provide evidence of tertiary structure and dynamics. The CD of chromophores absorbing in the near-UV region is associated with the π−π* electronic transitions. Relevant transitions for the aromatic amino acids are listed in Table 7.6.2. Tryptophan, tyrosine and phenylalanine possess partial or complete planes of symmetry, so that their dichroism arises almost entirely as a result of interaction of the transitions with neighboring groups. Dipole-dipole interactions, especially with other aromatic rings
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Table 7.6.2
Transitions of Aromatic Amino Acid Side Chainsa
∆εmax (M−1 cm−1)b
Electronic transitions
Peak wavelengths (nm)
Phenylalanine
1L b
262, 268
±0.3
Tyrosine
1L a 1L b
far UV 277, 283 268, 289 (shoulders)
±2.0
1L a
266 ∼295-303c
1L b
285, 292, 305d
Amino acid
Tryptophan
±3 ±2.5
aData from Strickland (1974). bEstimated values for an isolated side chain. cMay be seen as a low intensity shoulder or peak from a Trp in a nonpolar environment. dUnusual; observed with, e.g., lysozyme.
and with the π−π* transition of peptide bonds, make a major contribution to CD and are relatively long-range, being effective up to ∼1 nm. Electric-magnetic coupling also has an influence on the CD, but is very short-range and makes a relatively minor contribution. In contrast to the aromatic residues, the disulfide bond has no plane of symmetry and is, as a result, intrinsically optically active, particularly around a dihedral angle of 90°. These interactions of the environment with transitions of aromatic residues result in dichroic bands that are either positive or negative so that statistically the more such residues there are, the less intense the resultant ellipticity is likely to be. An intense signal frequently indicates the presence of one particularly strong interaction that overrides other minor contributions—such as the strong band at 292 nm seen in aspartic proteinases (Fig. 7.6.10), and the contributions of the one tryptophan and four tyrosine residues to the CD spectrum of interleukin 1β (Fig. 7.6.4). The 1Lb transition for phenylalanine resembles that for benzene (Strickland, 1974), but the CD signal is of relatively low intensity, so that it is seen only as shoulders or minor peaks to the low-wavelength side of tyrosine and tryptophan bands (Fig. 7.6.10). As a result of the high degree of symmetry in the side chain, the presence of these peaks is an indication of a highly specific and well-packed environment and can act as a marker for native folding. The 1Lb transition for tyrosine (the 1La transition lies in the far-UV) is more intense than that for phenylalanine and overlaps the two transitions for tryptophan (Strickland, 1974). It can result in a significant contribution to the near-UV CD (e.g., see Fig. 7.6.4 and Fig. 7.6.5) but is not always easy to assign in the presence of tryptophan. Hydrogen bonds to peptide carbonyl, carboxylate (C=O), and imidazole (–N=) groups contribute significantly to its CD, as do the presence of neighboring polar groups. The direction of the optical transition is in the plane of the ring and perpendicular to the axis of symmetry (Fig. 7.6.11) so that interaction with neighboring groups makes the CD very sensitive to rotation of the residue and to the dynamics of the protein in general.
Determining the CD Spectrum of a Protein
The two transitions for tryptophan exhibit distinct features, which lead to quite different CD bands. The 1La transition is broad, relatively featureless, and intense (Fig. 7.6.2). The 1 Lb transition is weaker but exhibits fine structure (Fig. 7.6.2) similar to that of tyrosine. It can be masked by or superimposed on the 1La transition. Interactions similar to those listed for tyrosine strongly affect the peak wavelengths and intensities. The presence of
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∆ A x 10 4
2
1
240
260 280 Wavelength (nm)
300
320
Figure 7.6.10 The near-UV CD spectra of cathepsin D. The solid line represents the porcine enzyme and the dashed line represents the bovine enzyme, each cleaved into two chains which remain associated; the dotted line represents the bovine intact enzyme. The spectra were recorded using 10-mm- and 20-mm-path-length cells, with enzyme concentrations of 0.8 to 0.4 g/liter. The far-UV CD and fluorescence spectra of the cleaved and intact enzymes are not significantly different (Pain et al., 1985).
OH
Figure 7.6.11 The near-UV absorbance transition dipole moment of the tyrosine side chain. The 1Lb transition is seen to be perpendicular to the axis of rotation of the phenolic ring. Interactions of the transition with the environment, expressed as an enhanced CD band, will be sensitive to rotation of the side chain around this axis.
1L
b
C
β
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bands above 285 nm is diagnostic for tryptophan in a specific environment; interactions that cause the 1Lb transition to be shifted to the red (or high-wavelength) end of the spectrum result in bands as high as 310 nm which are highly conformation-specific. The two transitions may be affected quite independently by interactions, and their combination can thereby result in four distinct types of spectrum (Strickland, 1974). Disulfide bonds may exhibit a broad band of ellipticity in the range of 240 to 350 nm. This can be confused with the band associated with the 1La transition of tryptophan but, in cases where a disulfide makes a significant contribution to the CD, it can be recognized by its ellipticity above 320 nm. It can augment or diminish the apparent intensity of tryptophan and tyrosine contributions without being recognized as such and without fine structure necessarily being lost. Some of the above features can be seen in the spectra of cathepsin D (Fig. 7.6.10), where the intact single-chain bovine enzyme is compared with the same material cleaved at an exposed loop but without dissociation. The latter has 50% of the specific activity of the intact molecule. Phenylalanine residues can be seen to be present in specific environments in both forms. The fine structure of the 1Lb transition of tryptophan is superimposed on the broad peak of the 1La transition, which is apparently more intense in the intact enzyme. Alternatively, there could be a greater contribution from disulfide bonds, but the absence of ellipticity above 320 nm favors the former assignment and the CD is therefore consistent with a limited increase in dynamics of the molecule as a result of the chain cleavage. On the other hand, the persistence of the pronounced peak at 292 nm, where the contribution of the 1La transition is low, indicates that the interaction responsible for this band is unchanged by cleavage. Coupled with the fact that both far UV CD and fluorescence spectra are identical for the two forms, it is concluded that the overall conformations are structurally similar, differing mainly in their packing and dynamics. Comparison with the spectrum for porcine cathepsin D demonstrates the conformational similarity expected from sequence homology, as evidenced in particular by the persistence of the interaction responsible for the appearance of the 1Lb band with its intense peak at 292 nm. Differences in the relative intensities of the minor bands provide a sensitive fingerprint for distinguishing two proteins with strong sequence homology. SUPPORT PROTOCOL 2
Determining the CD Spectrum of a Protein
INTERPRETATION OF FAR-UV CD SPECTRA The peptide bond exhibits intense circular dichroism bands when it is located in regular, rigid conformations of the protein backbone chain. It is not surprising therefore to find that the different secondary structures—helix, pleated sheets, and β-turns—each give rise to characteristic spectra (see Fig. 7.6.6). In principle it should be possible, knowing the spectra for each type, to deconvolute the far-UV spectrum for a particular protein to give the proportions of each form of secondary structure in the folded conformation. Early work involved the use of the spectra of polyamino acids as models for α-helix, β-pleated sheet, and random coil conformations. These turned out to be inadequate models for the relatively short helices, distorted intramolecular β-sheets, and aperiodic—but far from statistically random—coil conformations found in the average globular protein. The alternative approach, which forms the basis of most estimations and avoids the need to define reference spectra, involves the use of a database of CD spectra (see Yang et al., 1986, for listings) and corresponding secondary structure contents derived from X ray structure determination. Linear combination of these spectra is carried out to give a resultant that best fits the experimental spectrum for the test protein. The two algorithms that have been most commonly used are those of Provencher and Glöckner (1981) and of Johnson and coworkers (Johnson, 1990). The former used a bank of 16 proteins with spectra measured between 240 and 190 nm. Using a different statistical approach, Johnson and coworkers provided an algorithm that uses 15 reference spectra measured between
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260 and 178 nm. The latter, together with a more recent development of the method, has been reviewed and compared with the Provencher and Glöckner algorithm (Johnson, 1990) and the underlying assumptions have been critically discussed (Manning, 1989). Greenfield (1996) has provided a useful and comprehensive review of all the main algorithms for determining secondary structure. Commercial CD spectrometers include at least one algorithm in their software, although it is worth installing the two original programs independently so that results from each can be compared. Copies are available from the authors of the programs and from Dr. Greenfield. Attempts have been made to assess the success of different algorithms in deriving secondary structure content from CD spectra (see Yang et al., 1986). The general conclusion is that the helix content is usually estimated well, regardless of the structural type of protein. This is not surprising given the high intensity of the helix spectrum. β-sheet content is estimated well in the case of “all-β” proteins, but in mixed α and β proteins there is greater uncertainty. Some proteins, such as subtilisin, perform badly regardless of the method used. The estimation of the content of β-turns is similarly subject to greater uncertainty. Reasons for these problems are various. First, estimations are dependent on the correct values of mean residue ellipticity, which depend critically on determination of the protein concentration. Second, it is reasonably claimed that the further down in wavelength spectra are recorded, the better the estimate of structure (Johnson, 1990). It is not possible with all proteins to obtain reliable spectra below 185 nm with commercial instruments— inability to use the appropriate buffer and intrinsic scattering from larger proteins are limiting factors, although loss of optimal instrument performance as a result of insufficient maintenance is frequently the main cause of the problem. Third, there is uncertainty about the degree of contribution of aromatic and cystine residues in the far-UV (see Chaffotte et al., 1992, for references and discussion). Aromatic residues are probably significant when the residues are located in clusters, and cystine residues when the dihedral angle is of the order of 90°. Fourth, criteria vary for defining the secondary structure content from the three-dimensional coordinates of structures. In conclusion, the analysis of spectra properly recorded to 185 nm, or lower where possible, can give useful estimates of secondary structure content, but the content of turns and of β-structure should be interpreted with caution. In principle, Fourier transform infrared spectroscopy should provide better estimates of the latter. Secondly, when using the results of far-UV CD determination to characterize reproducibility of folding for different samples, it is important first to compare the spectra visually and to look for possible trends or factors that may explain small differences, rather than to rely solely on comparison of derived secondary structure contents. COMMENTARY Background Information CD spectra in the near-UV are of much lower intensity than those in the far UV, in part as a result of the much lower concentration of the chromophore, but can be measured reproducibly with due care. They are very important in characterizing the degree and correctness of folding of recombinant proteins and their mutants. Near-UV CD depends critically on the atomic environment and closeness of packing of the aromatic residues, including their acces-
sibility to solvent. Although the structural information is limited strictly to the immediate neighborhood of the relatively few aromatic residues, the generally cooperative nature of protein conformation means that changes in aromatic CD frequently reflect more widespread changes in conformation and dynamics. A slight loosening of a protein structure may not result in any change in solvent accessibility to aromatic residues and therefore will not change the maximum fluorescence wavelength of tryptophan. Such a change may also be
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outside the resolution of X-ray crystallographic analysis. The resulting change in dynamics of the residue, and hence in the asymmetry and detailed interactions with the immediate protein environment, will however affect the nearUV CD spectrum, usually quite markedly (see Fig. 7.6.5 and Fig. 7.6.12). Changes in the far-UV CD spectrum resulting from slight changes in conformation or
[θ]mrw x 10–3 (deg cm2 dmol–1)
A
partial unfolding will not in general be so marked as in the near-UV region, so that the value of the former for the detailed characterization of correct folding is not as great as that of the latter. It is, however, applicable to proteins with little or no aromatic content. The fact that the far-UV CD arises from the transitions of a single type of chromophore—i.e., the peptide bond—and that the backbone conforma-
5
0
–5
–10
–15
∆) x 104
B
200
210
220 230 Wavelength (nm)
240
250
250
260
270 280 Wavelength (nm)
290
300
0
–1
–2
Determining the CD Spectrum of a Protein
Figure 7.6.12 Effect of mutations detected by CD. The far-UV CD spectra (A) show that the secondary structure of β-lactamase PC1 (solid line) from Staphylococcus aureus is essentially unaffected by point mutations P2 (Thr 140→Ile; dashed line) and P54 (Asp 146→Asn; dotted line). The crystallographic structure of P54 (Herzberg et al., 1991) confirms that, apart from a loop region, the main body of the molecule that contains the thirteen tyrosine residues is very closely similar to that in the wild-type enzyme. The intensity of the tyrosine ellipticity (B) is, however, markedly decreased in each of the mutants, the lower thermodynamic stabilities of which support the interpretation of increased dynamics (Craig et al., 1985).
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tion constitutes the main influence on its dichroism, means that this region is good for identifying family relationships between proteins. CD in general is a useful tool for examining proteins made by site-directed mutagenesis for structure-function studies, where it is important to know to what extent the conformation has been affected by the mutation. The far-UV spectrum will tell whether the mutant has folded with a backbone conformation similar to that of the wild-type protein, whereas the near-UV spectrum will detect many minor changes in conformation or dynamics (Fig. 7.6.12). CD complements fluorescence as a technique for characterizing the folded conformation in solution and for providing, in a single spectrum, a highly specific fingerprint for the native state. It requires only modest amounts of protein and measurements are relatively quick to make. Like fluorescence, its use as a comparative method requires a control spectrum from a carefully characterized sample of the natural or recombinant protein, recorded under defined and optimal conditions, using a spectrometer in good condition.
Critical Parameters Many of the critical experimental parameters for CD spectrometry are the same as those for measuring fluorescence. Spectra will only be as reliable as the protein solution, the machine, and the operator. Cells must be scrupulously clean, solutions must be well-clarified, and buffers must have a low absorbance. In addition, not only must the protein concentration be known precisely but, together with the path length, must be chosen to optimize the absorbance in the region of interest. This will sometimes require recording a spectrum in overlapping sections. Preliminary experiments to optimize the instrumental parameters are important. Lastly, it is essential that the spectrometer be well-maintained and up to specification. The useful life for a lamp is on the order of 800 to 1000 hr, but performance in the far-UV may start to fall off before this.
Troubleshooting The problem most frequently met (though a casual scan of the literature shows that it is not so frequently recognized) is the loss of light intensity at low wavelengths. Under such conditions, many spectrometers will produce traces that are smooth, giving the appearance of being valid spectra, but which are in fact
artifacts. The example of such an artifact shown in Figure 7.6.13 provides a cautionary tale. In this case, the glutamine absorbs light in the far-UV on account of its side-chain amide group. The increasing concentration of glutamine results in a progressively longer wavelength at which light is cut off, to which this particular spectrometer responded by returning to zero dichroism, giving the impression of a peak with a maximum at increasing wavelengths. To avoid such artifacts, it is essential to monitor the photomultiplier high voltage during a scan. Similar problems can arise from using too high a protein concentration or too long a light path, from too much absorbance by the buffer, from turbidity in the protein solution, from a dirty cell, or—below 200 nm— from absorption by oxygen that results from incomplete purging with nitrogen. A further problem that is sometimes encountered is that the spectrum obtained after baseline subtraction does not return to zero above 320 nm (in near-UV CD) or 250 nm (in far-UV CD). If, above these wavelengths, the spectrum runs parallel to the zero base, a provisional spectrum may be derived by subtracting the difference in ellipticity, as a constant, from the whole spectrum. For quantitative assessment of a sample, however, the spectrum should be redetermined, paying attention to the correct buffer blank, cell reproducibility, and instrumental drift (see Strategic Planning). Care must be taken to rectify all of the above mentioned problems if meaningful spectra are to be obtained for reliable characterization.
Anticipated Results Several examples of near- and far-UV CD spectra are given in the figures included in this unit. With current instruments the final spectra exhibit low noise and—provided that the instrument and sample parameters have been optimized—should be true and reproducible within the limits of protein-in-buffer absorbance of <1. The most frequent source of nonreproducibility of intensity in spectra is the difficulty of determining the protein concentration, particularly of larger proteins that scatter more and of those with a greater tendency to aggregate. What constitutes a significant difference between two spectra? When the differences are small, the answer depends on sample preparation and sample stability as well as accuracy of concentration determination, identification of and compensation for drift in the spectrometer, correct baseline correction, absence of bubbles
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3.0 f 2.5
g
e 2.0
∆) x 103
h
d
1.5 c
1.0 b
0.5 0.0 –0.5
a
–1 190
200
210
220
230
240
250
Wavelength (nm)
Figure 7.6.13 A convincing artifact. In an attempt to study the conformational consequences of adding an acceptor, D-glutamine, to a DD-peptidase, far-UV CD spectra were recorded for the enzyme in the presence of increasing concentrations of glutamine: a = 0 mM, b = 3 mM , c = 7 mM, d = 10 mM, e = 20 mM, f = 30 mM, g = 50 mM, and h = 95 mM. The enzyme concentration was 0.1 mg/ml, equivalent to ∼10−3 M peptide bond, in 10 mM sodium phosphate pH 7.2. A 2-mm cell path length was used.
in the sample, reproducible cleanliness of the cuvette, and the level of general handling procedures. Ultimately, an assessment of significance depends on the experience, competence, and confidence of the operator. As CD is very sensitive to small changes in conformation and dynamics, wild-type and mutant proteins will frequently give spectra that differ slightly in intensity rather than shape. If the dihedral angle for a disulfide bond is such as to yield an intense contribution, small conformational perturbations in a mutant protein may further affect the spectrum. Hence, there is the need to examine proteins by a number of different spectroscopic, hydrodynamic, and functional probes in order to assess correct folding of mutant proteins.
Time Considerations
Determining the CD Spectrum of a Protein
A single spectrum, with associated baseline, will require ∼1 hr, including the time needed to fill and clean the cuvette. Up to an additional hour may be needed for optimizing instrument parameters if the sample is unfamiliar. For a number of samples, economies of time can be effected (see Basic Protocol).
Literature Cited Bayley, P. M. 1980. Circular dichroism and optical rotation. In An Introduction to Spectroscopy for Biochemists (S.B.Brown, ed.) pp. 148-235. Academic Press, London. Chaffotte, A.F., Guillou, Y., and Goldberg, M.E. 1992. Kinetic resolution of peptide bond and side chain far-UV circular dichroism during the folding of hen egg white lysozyme. Biochemistry 31:9694-9702. Chen, G.C. and Yang, J.T. 1977. Two-point calibration of circular dichrometer with d-10-camphorsulfonic acid. Anal. Lett. 10:1195-1207. Craig, S., Hollecker, M., Creighton, T.E., and Pain, R.H. 1985. Single amino acid mutations block a late step in the folding of β-lactamase from Staphylococcus aureus. J. Mol. Biol. 185:681687. Craig, S., Pain, R.H., Schmeissner, U., Virden, R., and Wingfield, P.T. 1989. Determination of the contributions of individual aromatic residues to the CD spectrum of IL-1β using site directed mutagenesis. Int. J. Peptide Protein Res. 33: 256-262. Greenfield, N.J. 1996. Methods to estimate the conformation of proteins and polypeptides from circular dichroism data. Anal. Biochem. 235:1-10. Herzberg, O., Kapadia, G., Blanco, B., Smith, T.S., and Coulson, A. 1991. Structural basis for the inactivation of the P54 mutant of β-lactamase
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from Staphylococcus aureus PC1. Biochemistry 30:9503-9509. Holzwarth, G. and Doty, P. 1965. The ultraviolet circular dichroism of proteins. J. Am. Chem. Soc. 87:218-228.
Yang, J.T., Wu, C.-S. and Martinez, H.M. 1986. Calculation of protein conformation from circular dichroism. Methods Enzymol.130:208-269.
Key References
Johnson, W.C. 1990. Protein secondary structure and circular dichroism: a practical guide. Proteins Struct. Funct. and Genet. 7:205-214.
Bayley, 1980. See above
Manning, M.C. 1989. Underlying assumptions in the estimation of secondary structure content in proteins by circular dichroism spectroscopy - a critical review. J. Pharmacol. Biomed. Anal. 7:1103-1119.
Johnson, 1990. See above.
Pain, R.H., Lah, T., and Turk, V. 1985. Conformation and processing of cathepsin D. Biosci. Rep. 5:957-967.
A good introduction to the principles and practice of circular dichroism.
A good account of the practice of far-UV CD spectroscopy and a description and assessment of the determination of protein secondary structure content.
Provencher, S. W. and Glöckner, J. 1981. Estimation of globular protein secondary structure from circular dichroism. Biochemistry 20:33-37.
Sears, D.W. and Beychok, S. 1973. Circular dichroism. In Physical Principles and Techniques of Protein Chemistry, Part C (S.J. Leach, ed.) pp. 445-593. Academic Press, New York and London.
Ptitsyn, O.B. 1992. The molten globule state. In Protein Folding (T.E. Creighton, ed.) pp. 243300. W.H. Freeman, New York.
A comprehensive review of the theory of CD, together with a useful discussion of near-UV spectra in particular for selected proteins.
Schmid, F.X. 1989. Spectral methods of characterizing protein conformation and conformational changes. In Protein Structure (T.E. Creighton, ed.) pp. 251-285. IRL Press, Oxford.
Strickland, 1974. See above
Strickland, E.H. 1974. Aromatic contributions to circular dichroism spectra of proteins. C.R.C. Crit. Rev. Biochem. 2:113-175. Swords, N.A. and Wallace, B.A 1993. Circular dichroism analyses of membrane proteins: Examination of environmental effects on bacteriorhodopsin spectra. Biochem. J. 289:215-219. Wingfield, P., Graber, P., Moonen, P., Craig, S., and Pain, R.H. 1988. The conformation and stability of recombinant-derived granulocyte-macrophage colony stimulating factors. Eur. J. Biochem. 173:65-72.
An older but still excellent account of the practice, spectroscopic basis, and interpretation of the near UV CD spectroscopy of proteins and model compounds. Yang et al. 1986. See above. A good account of the practice of far-UV CD spectroscopy and a description and assessment of the determination of protein secondary structure content.
Contributed by Roger Pain JoÓef Stefan Institute Ljubljana, Slovenia
Characterization of Recombinant Proteins
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Determining the Fluorescence Spectrum of a Protein
UNIT 7.7
Fluorescence spectra provide a sensitive means of characterizing proteins and their conformation. The spectrum is determined chiefly by the polarity of the environment of the tryptophan and tyrosine residues and by their specific interactions. The diminution of intensity of the fluorescence by added “quenchers” provides a measure of the degree to which these residues are buried in the protein structure or exposed to solvent. One of the main values of fluorescence as a technique for probing protein conformation is that it is highly sensitive and very economical of material. This means, however, that small traces of fluorescent impurities in the solvent or on the cell are readily detected and, if care is not taken, can lead to misinterpretation of the spectra. The essential aim in using this technique, therefore, must be to obtain a fluorescence emission spectrum for the recombinant protein that is guaranteed free from all too easily generated artifacts. STRATEGIC PLANNING The following points are essential to obtaining reproducible spectra for reliable comparison between samples. Fluorescence Spectrometer Fluorescence spectra are measured with a fluorescence spectrometer (spectrofluorimeter), of which there are a variety of models on the market. For protein characterization, a xenon light source and optics that ensure low levels of stray light are essential. Spectrometers with dual holographic gratings in the excitation monochromator are preferred. There are two main types of fluorescence detection available: analog and photon counting. The latter counts individual photons, which enables high sensitivity and stability, but limits the range of intensities over which a linear response is obtained. Analog spectrometers provide high-precision measurements over a wide range of intensities. Stability is generally high, but is improved by the presence of a reference photomultiplier. In general, the photon-counting spectrometer is for more specialized use and the analog type of instrument is more widely used for the kind of work described in this unit. A thermostatted cell holder—either one connected to a good circulating thermostat bath or one of the Peltier controlled type—is essential. The latter controls the temperature by electrical means, the advantages being that no water needs to be brought into the spectrometer and, secondly, that the temperature can be raised and lowered very quickly. The spectrometer should incorporate shutters that protect the sample and the photomultiplier from radiation when scanning is not in progress. A magnetic stirrer incorporated into the cell holder is useful to facilitate mixing and to minimize artifacts due to photochemical degradation. Careful attention must be paid to the manufacturer’s instructions in order to achieve the optimum settings for measurement. Calibration of Spectrometer In order to ensure reproducibility, the wavelength setting should be correct within ±0.5 nm. This will generally not be a problem, but it is just as well to check, most readily with a mercury lamp inserted into the cell compartment. The procedure described by Lakowicz (1983) is recommended. Calibration of intensity can be more difficult, and it is always more reliable to scan samples in parallel with a solution of a known, stable compound, be it the protein in question or a model compound such as N-acetyl-L-tryptophanamide. Contributed by Roger H. Pain Current Protocols in Protein Science (1996) 7.7.1-7.7.19 Copyright © 1996 by John Wiley & Sons, Inc.
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Cells Choice of cells The fluorescence cuvettes best used for reliable characterization of proteins are either the standard 10 × 10–mm fluorescence quartz cuvettes (with all four faces polished), taking 2 to 3 ml of solution, or semimicro cuvettes, taking ~0.7 ml. In order to ensure reproducible positioning of the cuvette, it should be placed in the cell holder with the inscription facing the incident light beam. Preparation and handling of cells Cuvettes must be thoroughly and reproducibly cleaned by soaking in a proprietary cuvette-cleaning fluid such as Hellmanex II (Hellma), an alkaline liquid concentrate, or RBS-35 Detergent Concentrate (Pierce), following the manufacturer’s instructions. The manufacturers of Hellma cuvettes caution against the use of ultrasonics as a means of boosting cleaning efficiency. Fifty percent nitric acid is also effective, but requires greater care and a fume hood. Detergents frequently contain fluorescent material and, if used, should be removed from cuvettes with 2 M HCl followed by water. After the final rinse with copious amounts of high-quality distilled water, the cells are best dried using suction through a Pasteur pipet protected with a small length of plastic tubing. This is safer than using acetone or alcohol, which can leach out fats from fingers and deposit them on the cuvette. The faces of cuvettes must not be touched after cleaning. Filling, emptying, and rinsing between measurements of spectra must be done using pipets, to avoid spillage onto the outer faces. A Pasteur pipet protected with fine plastic tubing is satisfactory. The baseline scans, with buffer in the cell, must be examined as a constant check on the reproducibility of the state of the cell. The cleanliness of the cuvette can be checked by filling with distilled water and scanning the appropriate wavelength region of the spectrum. If there is no contamination by fluorescent material, the spectrum should be flat, save for the Raman band (see Fig. 7.7.1) on the high-wavelength side of the excitation wavelength, λmax. Buffer Solutions Buffers should have low absorbance and low fluorescence in the regions of excitation and emission. Absorbance will usually be expected to be <0.1 and fluorescence close to zero. This will normally be the case for standard buffers made from analytical-grade reagents or materials of equivalent purity but they should, nevertheless, be routinely checked for fluorescence. If a fluorescent component is added to the solution—e.g., as a ligand—it should be checked that the fluorescence observed arises solely from that component. The actual buffer solution used to dissolve or to dialyze the protein should be used for the fluorescence blank. Plastic containers (and stirring bars) may contribute fluorescent agents; if these are used, appropriate blanks should be carefully monitored for fluorescence. Clarification of Solutions Protein solutions must be freed from the turbidity resulting from dust and aggregated protein. Many proteins can be filtered (preferably as a stock solution) using a 0.22-µm filter—ideally of the grade special for proteins such as Durapore (Millipore)—fitted to a syringe. Alternatively, if the protein is adsorbed by filters, the solution can be clarified by centrifugation. A microcentrifuge operated at maximum speed for 10 min at 4°C provides reliable clarification for the majority of proteins that do not sediment appreciably under these conditions. Buffer solutions are filtered on a larger scale, using a 0.22-µm filter. Determining the Fluorescence Spectrum of a Protein
It is essential to monitor clarification if artifacts are to be avoided. There are three indicators of the completeness of clarification, each available from the standard procedures that should be adopted for spectroscopic characterization.
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Wavelength (nm) Figure 7.7.1 Fluorescence spectrum of hen egg white lysozyme (EWL). The Rayleigh and Raman bands are seen in the scans for the solvent baseline and protein solutions (solid lines). Circles represent the spectrum of EWL with baseline subtracted. Parameters: EWL A280 = 0.05; λex = 280 nm; excitation and emission bandwidths, 2.5 nm; scan rate, 100 nm/min; five scans accumulated. Spectrum was measured using a Perkin Elmer LS50B fluorescence spectrometer.
1. Determination of protein concentration (UNIT 7.2) requires an absorbance spectrum to be recorded on a good quality spectrophotometer from 240 to 350 nm. Aromatic amino acid residues do not absorb above 320 nm, so the spectrum between 320 and 350 nm should be only marginally above baseline. The presence of turbidity will result in finite attenuance in this region that increases toward lower wavelengths. 2. From the above absorbance spectrum, the ratio Amax/Amin is measured, where Amin is the absorbance at the trough in the region 240 to 250 nm. This ratio will be specific for a given protein and should be measured for a sample that has been carefully and reproducibly clarified. Due to the strong wavelength dependence of scattering, the ratio is very sensitive, decreasing sharply with turbidity, and can be used as a semiquantitative check on the clarity of further solutions of the protein. 3. The scan of a fluorescence emission spectrum should routinely include the region of the Rayleigh peak (see Fig. 7.7.1). This provides a built-in indicator of turbidity of the buffer or protein solutions. The height of the peak should not differ greatly from that obtained for the solvent from the equivalent buffer blank scan; it will increase markedly in height if there is dust or aggregate in the solution. Characterization of Recombinant Proteins
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Figure 7.7.2 Temperature dependence of fluorescence. (A) Tyrosine (6 µM) at 303 nm (λex = 274 nm). (B) Tryptophan (1 µM) at 355 nm (λex = 278 nm). Both chromophores are in a polar environment, 0.01 M potassium phosphate, pH 7.0. Reprinted from Schmid (1989) with permission of Oxford University Press.
BASIC PROTOCOL 1
RECORDING A FLUORESCENCE EMISSION SPECTRUM A fluorescence emission spectrum is the plot of fluorescence intensity as a function of wavelength. Fluorescence emission takes place from the wavelength of excitation (λex) upwards. The range of the spectrum recorded is described below (step 5). Materials Calibrated fluorescence spectrometer, buffer solution, and cleaned cuvettes (see Strategic Planning) Clarified protein solution of known concentration (see Strategic Planning for clarification methods and UNIT 7.2 for concentration determination) with absorbance usually in the range of 0.05 to 0.1 at the wavelength to be used for excitation Prepare spectrometer 1. Switch on the spectrometer and allow it to warm up for the time specified in the manufacturer’s instructions (usually ∼30 min). Regulate the thermostat control to the desired temperature (normally 25°C unless the stability of the protein dictates otherwise). The intensity of fluorescence is highly temperature-dependent (see, e.g., Fig. 7.7.2) and it is important that the temperature be accurately controlled and measured in the cuvette with a calibrated electric thermometer. It is useful for future reference to note the time taken for the sample to reach the operating value.
2. Set the excitation wavelength (λex). Setting λex to the λmax of the absorbance spectrum (usually 280 nm) will excite both tyrosine and tryptophan residues. Alternatively, λex may be set to 295 nm in order to excite only tryptophan (see annotation to step 3). Determining the Fluorescence Spectrum of a Protein
3. Set the excitation bandwidth. The term bandwidth is described and defined in UNIT 7.6, Basic Protocol step 2b. The excitation bandwidth (in nm) is determined by the excitation slit width (in mm). It will
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Figure 7.7.3 Fluorescence spectrum of ribonuclease (RNase), a “tyrosine only” protein. Parameters: Rnase A275 = 0.05; λex = 275 nm; one scan, smoothed; other settings as for Figure 7.7.1. Baseline was measured with the same settings and subtracted. Measurements made using a Perkin-Elmer LS50B.
determine the intensity of the excitation and the amount of scattered light from the Rayleigh peak that will be “seen” in the emission spectrum. It is important in certain cases in determining which chromophore is excited. It is particularly important, for example, to limit the bandwidth when measuring the contribution of tyrosine in the presence of tryptophan. The emission maximum of tyrosine at 303 nm (see Fig. 7.7.3) is close to the required excitation wavelength of 295 nm. The envelope of the Rayleigh peak must not, in this case, significantly overlap the emission band of tyrosine. Settings of 2.5 to 5 nm for λex ≈280 nm and ≤2.5 nm for λex ≈295 nm are usual.
4. Set the emission bandwidth. Since protein emission spectra are generally rather broad, larger emission bandwidths can usually be tolerated. Only where it is important to resolve tyrosine fluorescence from tryptophan fluorescence and the Rayleigh scattering peak is it necessary to minimize the bandwidth. In cases where only small amounts of material are available, it is necessary to strike a balance between resolution and light intensity in order to obtain the best possible signal-to-noise ratio. Settings of 2.5 to 10 nm are normal.
5. Set the range of emission wavelengths to be scanned for the emission spectrum. The lower wavelength should be ∼10 nm below the excitation wavelength in order to record the Rayleigh peak (see discussion of Clarification of Solutions in Strategic Planning). The higher end of the range should be chosen to include as much of the spectrum as possible, so as to allow integration of the peak if necessary. Values of 400 and 340 nm are suitable for proteins with and without tryptophan, respectively. Hence, for example, a typical emission wavelength range for a Trp-containing protein excited at 295 nm would be 285 to 400 nm. Characterization of Recombinant Proteins
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Figure 7.7.4 The Raman band measured under different scanning conditions. The appropriate section of the baseline in Figure 7.7.1 is shown for: a single scan at 1500 nm/min (dotted line); ten scans at 1500 nm/min (dashed line); and ten scans at 100 nm/min (solid line). The times required for scanning the complete baselines for these three measurements were 0.087 min, 0.87 min, and 13 min, respectively. Measurements made using a Perkin-Elmer LS50B.
6. Set the scan rate. The scanning rate for the emission wavelength has to be slow enough to allow the instrument to respond to changes in intensity with wavelength. Too fast a scan, even with a number of accumulations, results in an apparent spectrum that can differ markedly from the true spectrum (Fig. 7.7.4). When, as is usually the case with proteins, the emission band is rather broad (Fig. 7.7.1), a scan rate setting in the region of 100 nm/min will usually be appropriate.
Scan buffer blank and protein sample 7. Fill a cleaned cuvette with a filtered sample from the actual batch of buffer used to dissolve or dialyze the protein, then place the cuvette in the cell holder of the spectrometer in a reproducible orientation (see discussion of Cells in Strategic Planning). Scan this buffer blank using the same instrument settings as are appropriate for the sample, store the spectrum, and check for any unexpected fluorescence bands. If the settings appropriate for the protein sample cannot be estimated in advance, the sample may be run before the buffer blank, although special care will have to be taken to remove any protein adsorbed to the inner faces of the cuvette. Temperature equilibration is not usually important for the buffer, except in cases where it contains fluorescent components. Cell holders are susceptible to corrosion by salt solutions. Cells should therefore be filled outside the spectrometer. Determining the Fluorescence Spectrum of a Protein
Apart from the Raman band, a buffer solution should give an almost flat baseline of low intensity (see Fig. 7.7.1). At the usual working concentrations of protein (A280 ≥0.05), buffer fluorescence constitutes only a minor contribution to the fluorescence spectrum of the protein and is accounted for by the appropriate baseline subtraction.
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It is advisable to repeat baseline scans at intervals during a series of experiments to check on contamination of the outer faces of the cuvette, and of the inner faces due to protein adsorption.
8. Remove the cuvette from the spectrometer, empty with a protected Pasteur pipet (see discussion of Cells in Strategic Planning) and refill in the same way with the clarified protein solution of known concentration with an absorbance of 0.05 to 0.1 at the wavelength used for excitation. Allow to come to temperature in the cell holder, then scan and store spectrum. If the protein concentration has to be low, and/or the intensity of fluorescence is low, it may be necessary to accumulate a larger number of scans. Proteins differ widely in their sensitivity to photodegradation, but exposure to UV radiation may lead to damage to those molecules in the light path, with subsequent reduction in fluorescence intensity. Excitation slit widths should, therefore, be as narrow as is consistent with good signal-to-noise ratio. Stirring the solution in the cuvette will bring undamaged molecules into the light path (Fig. 7.7.8) and further reduce significant artifacts due to radiation damage.
Calculate results 9. Subtract the baseline from the sample spectrum using the instrument software (Fig. 7.7.1). Fluorescence has no units. Hence, if the fluorescence intensities of different samples are to be compared, careful attention must be paid to reproducible concentrations and instrumental conditions. Instrument response may vary with time, so must be controlled by the use of stable standards.
10. Correct the apparent fluorescence intensities for the inner filter effect. A protein solution will absorb a certain proportion of the radiation before it reaches the volume of solution whose fluorescence is viewed by the optics of the instrument (see Fig 7.7.5). For a solution of absorbance 0.1, the intensity at the center of the cell is about 10% less than that of the incident radiation, with a corresponding decrease in the observed fluorescence. Similarly, the fluorescence intensity may be reduced by absorption in the exit path. This is known as the inner filter effect. In practice, it means that the observed fluorescence intensity is directly proportional to protein concentration only at low concentrations, when absorbance is <0.1 at λex. The apparent fluorescence intensity (Fapp) can be corrected by the following approximation: Fcorr = Fapp antilog{(Aex + Aem)/2} where Aex and Aem are the solution absorbances at the excitation and emission wavelengths, respectively. The effect is much more marked if a third component that absorbs in the wavelength range of the protein absorption or fluorescence is added to the solution (see Basic Protocol 2).
11. Determine the value of the fluorescence intensity at the wavelength of maximum intensity (Imax at λmax), using, where available, the instrument software. It is possible to obtain corrected spectra, which take into account the variation of intensity of the light source as well as variation of the detector response with wavelength. The procedures for calculating these are not simple (see Lakowicz, 1983), and for the purposes of characterization described here the uncorrected spectra are normally used. Because of the above variations, however, it is important when comparing spectra to do so with spectra obtained using the same instrumental settings and, preferably, the same instrument. In order to compare complete spectra from different samples of a protein, they should be normalized by dividing the value of fluorescence intensity by the protein concentration (i.e., Iem/c). Characterization of Recombinant Proteins
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1 I/I0 0.9 0.8
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Figure 7.7.5 The inner filter effect. A cuvette (10 × 10–mm) is represented in plan view, with the collimated incident beam from the monochromator having intensity I0. As a result of absorption by the protein solution, the intensity through the cuvette will decrease steadily, emerging with intensity I. The values are illustrated for a solution having absorbance at the excitation wavelength of 0.1. The optics of the fluorescence detector are focused so that only the fluorescence originating from the volume depicted by the heavily shaded square is “seen” by the photomultiplier. Thus the observed fluorescence intensity will be less than that expected from the fluorescence of the protein at infinite dilution. The fluorescence passes through the protein solution on its way to the detector and will be further decreased in intensity if the solution absorbs at the wavelengths of the emitted radiation.
BASIC PROTOCOL 2
DETERMINATION OF FLUORESCENCE QUENCHING Fluorescence quenching provides a means of probing the accessibility of tryptophan residues to small molecules and thus gives information about their structural environment. The technique involves quantifying the decrease in protein fluorescence intensity in the presence of increasing concentrations of quencher, followed by analysis of the data to give details of the interaction of the quencher with the tryptophan residue. Materials Protein solution in buffer, A280 = 0.05 to 0.1 Quenchers: these are most conveniently made up as concentrated stock solutions; examples of frequently used quenchers include: 5 M NaI or 2.5 M KI solution containing 1 mM Na2S2O3 to prevent I3− formation 5 M CsCl (optical grade, Aldrich) 8 M acrylamide (Electran grade from BDH; ε295 = 0.236 liter/mol/cm) 2.5 M succinimide (recrystallized from ethanol with activated charcoal treatment; ε295 = 0.03 liter/mol/cm)
Determining the Fluorescence Spectrum of a Protein
Additional reagents and equipment for recording a fluorescence spectrum (see Basic Protocol 1)
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1. Scan the fluorescence spectrum of a measured volume of the protein in the required buffer (see Basic Protocol 1). 2. Add a known small volume of a stock solution of quencher, mix thoroughly and again scan the spectrum. Alternatively, for economy of time, measure the intensity of fluorescence at just two wavelengths—315 and 350 nm—allowing a sufficient time of accumulation to achieve a reproducible average reading. The actual amount of quencher added is a matter of judgment, depending on the degree of quenching found. It is important to obtain at least ten measurements of fluorescence intensity spread evenly between zero and full quenching (see Fig. 7.7.6). Quenchers should be carefully purified. When iodide is used, it should be in the presence of 1 mM Na2S2O3 to avoid the formation of I3−, which can react with tyrosine. The tryptophan residues in a given protein are likely to be in different environments and, therefore, to exhibit different values of λmax. Measurement of the degrees of quenching at the red and blue ends of the emission spectrum will determine whether this is so and will also add further detail to the overall fingerprint of the protein (see discussion of Quenching in Support Protocol).
3. Correct each reading of the fluorescence intensity for dilution and for the absorbance of the quencher (see Basic Protocol 1). See also Eftink and Ghiron (1981).
4. Plot the resulting fluorescence intensities as a function of concentration of quencher according to the Stern-Volmer equation (see Fig. 7.7.6A): F0/F = 1 + KD[Q]
where F0 and F are the fluorescence intensities in the absence and presence, respectively, of a given concentration of quencher [Q], and KD is the Stern-Volmer quenching constant. KD is determined by the bimolecular quenching constant (kq) and by the fluorescence lifetime of the fluorophore (τ0), in the absence of quencher (KD = kqτ0). The slope of a plot of F0/F against [Q] gives KD directly, in units of liters/mol, with an intercept of unity. This plot is expected to be linear for a single fluorophore in a single environment. For a more complex system the plot will be non-linear (see Fig. 7.4.6A) and the data should be plotted according to a modified Stern-Volmer equation: F0/∆F = 1/faKD[Q] + 1/fa where fa is the fraction of the initial fluorescence accessible to the quencher, fa = F0a/(F0a + F0b) and KD is the Stern-Volmer quenching constant for that fraction of the fluorescence. A plot of F0/∆F against 1/[Q] gives an intercept of 1/fa and a slope of 1/ faKD (see Fig. 7.7.6B).
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Determining the Fluorescence Spectrum of a Protein
Figure 7.7.6 Quenching of fluorescence of 3-phosphoglycerate kinase (PGK). (A) Fluorescence emission intensities, expressed as a fraction of the unquenched fluorescence, are plotted according to the Stern-Volmer equation (see Basic Protocol 2, step 4) at 316 nm (filled symbols) and 350 nm (open symbols). Quenchers used were succinimide (circles; recrystallized from ethanol and treated with activated charcoal, 2.5 M stock solution) and potassium iodide (squares; 2.5 M stock solution with 1 mM Na2S2O3). Fluorescence intensities were adjusted for dilution and for quencher absorbance using a molar extinction coefficient ε295 = 0.03 liter/mol/cm for succinimide. The greater level of quenching at 350 nm reflects the fact that the more exposed of the two tryptophan residues has a more red-shifted fluorescence maximum. (B) Modified Stern-Volmer plot for the quenching of PGK fluorescence by succinimide. The data in panel A for quenching at 350 nm is plotted according to the modified Stern-Volmer equation (see Basic Protocol 2, step 4 annotation). The intercept on the y-axis is 2.36 ± 0.2. The results in panels A and B are characteristic of a protein where part of the fluorescence is inaccessible to a given quencher, in this case, iodide and succinimide. Measurements made using a Perkin-Elmer MPF3-L. Reprinted from Varley (1991) with permission of Elsevier Science.
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BASIC THEORY AND INTERPRETATION OF FLUORESCENCE SPECTRA When radiation is incident on a solution, most of it is scattered without change in wavelength. Thus, exciting radiation at 280 nm will be scattered and a fraction of it will be detected by the spectrofluorimeter as a peak at the same wavelength. The width of this Rayleigh scattering peak (Fig. 7.7.1) will depend on instrumental settings, mainly those of the excitation and emission slit widths, which determine the range of wavelengths observed. The bandwidth is the width, in nm, of the distribution of intensity measured at half the height of the maximum intensity.
SUPPORT PROTOCOL
A further low-intensity peak, known as the Raman band, is also observed in spectra (see Fig. 7.7.1). This is a low-intensity band of scattered radiation whose distance from the excitation band is a measure of the vibrational energy of the H–O bond in solvent water. At λex = 280 nm, the Raman band for water occurs at 311 nm; in general, it occurs at 1/{(1/λex) − (3.6 × 10−4)}, where λex is the excitation wavelength in nm. The intensity and resolution of the Raman band provides a useful empirical check on the performance of the spectrofluorimeter. A decrease in the signal-to-noise ratio usually indicates a deterioration of the lamp. Chromophores, such as the sidechains of tryptophan and tyrosine, will absorb some of the radiation in ∼10−15 sec and become excited from the ground electronic state (S0) to higher vibrational levels in a higher energy electronic state (S1; see Fig. 7.7.7). The energy in the vibrational levels is rapidly (∼10−12 sec) lost to the surroundings as thermal energy, so that the excited chromophore is very largely in its ground vibrational level. Return to the ground electronic state can result in the emission of radiation of lower energy and hence longer wavelength (Figs. 7.7.1 and 7.7.3), or the energy can be dissipated in a number of ways that are nonradiative (Fig. 7.7.7), including quenching (see discussion
S1
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Figure 7.7.7 The processes of absorption and emission of radiation by a chromophore. The electronic structure of a chromophore is in the ground state (S0) under normal conditions. Absorption of radiation, a rapid process, leads to excitation into an excited state (S1). Within each electronic state are a number of vibrational states, represented by horizontal lines. Under normal thermal conditions, only the lower one or two vibrational levels are occupied. Thus, although excitation takes place into a range of levels in the excited electronic state, emission, in the form of fluorescence, takes place from the lowest vibrational level. The lengths of the vertical lines representing absorption and emission illustrate the relative energies of the radiation in each process and illustrate why fluorescence occurs almost always at longer wavelengths than absorption. The vertical broken line represents the dissipation of absorbed energy by means other than radiation.
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of Quenching, below). The ratio of the rate constants for these two alternative processes defines the quantum yield or fluorescence efficiency, which is almost always less than unity. The chromophores tyrosine and tryptophan absorb radiation in the 280 nm region, re-emitting a proportion of it as fluorescence. The quantum yields for tryptophan, tyrosine and phenylalanine are 0.2, 0.1, and 0.04, respectively. Coupled with their relative absorption coefficients (UNIT 7.2), it can be seen why most protein fluorescence spectra are dominated by tryptophan. Quantitation of the fluorescent properties of a protein can provide a number of rather specific parameters that allow assessment of the correct folding, besides giving additional structural information. Tyrosine Fluorescence Is Frequently Not Seen in the Presence of Tryptophan The fluorescence intensity of free tyrosine is about one-fifth of that of tryptophan, but in proteins it is usually much weaker still. This is due to a combination of interactions—such as hydrogen bonding to peptide carbonyl groups, the presence of neighboring charged groups, and nonradiative energy transfer to tryptophan or disulfide bonds—all of which can result in tyrosine fluorescence being diminished or quenched. The degree of quenching varies from protein to protein and can be observed by paying careful attention to bandwidths (see Basic Protocol 1). In Figure 7.7.8, where the peak intensity at 335 nm of
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Determining the Fluorescence Spectrum of a Protein
Figure 7.7.8 Quenching of the tyrosine fluorescence of chymotrypsinogen A. The solid line represents the emission spectrum excited at 280 nm. The circles represent the corresponding spectrum obtained with λex = 295 nm. Because the latter spectrum is considerably less intense than the former (as a result of the differences in absorbance at 280 nm and 295 nm), the spectra were normalized using the ratio of their fluorescence intensities at 335 nm. Parameters: chymotrypsinogen A280 = 0.1; excitation and emission bandwidths, 2.5 nm; scan rate, 100 nm/min, two scans accumulated for λex = 280 nm and four scans for λex = 295 nm. Fluorescence was measured using a Perkin Elmer LS50B.
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fluorescence excited at 295 nm (i.e., tyrosine is not excited) has been normalized to that excited at 280 nm (where both tryptophan and tyrosine are excited), it is evident that there is no separate contribution of tyrosine to the fluorescence excited at the latter wavelength. Essentially all the tyrosine fluorescence is quenched (see Lakowicz, 1983). The unfolded protein shows a separate shoulder resulting from unquenched tyrosine fluorescence.
λmax and Imax Reflect the Detailed Environment of Aromatic Residues The precise wavelength of emission is, in the case of tryptophan, dependent on the polarity of the immediate environment of the chromophore in the protein, and maximum values have been observed over a broad range between 308 nm (highly nonpolar) and 352 nm (fully exposed to solvent). In tumor necrosis factor (TNF), the maximum emission wavelength is 320 nm, compared to 350 nm for the fully unfolded protein (Fig. 7.7.9). The value of 336.5 nm for λmax in acid solution indicates partial unfolding. By comparison, tryptophan residues in native chymotrypsinogen A, where λmax = 332 nm (Fig. 7.7.10), are in a slightly less nonpolar environment than in TNF. The value of λmax therefore provides both a characteristic parameter for a particular protein and structural information concerning the average degree of burial of the tryptophans. The wavelength of tyrosine fluorescence, on the other hand, does not vary with polarity of environment and occurs with a maximum at 303 nm. Phenylalanine does not absorb above 275 nm, so that its weak fluorescence is not normally observed. The intensity of fluorescence is, for both tyrosine and tryptophan, dependent upon interactions with neighboring groups that may quench the fluorescence emission from the native conformation. Tryptophan and tyrosine fluorescence is quenched, relative to the emission intensity in water, in solutions of denaturant. Depending on the degree of quenching of these chromophores in the folded protein, therefore, denaturation can result in an increase or a decrease in intensity, as well as a shift in λmax to longer wavelengths (350 to 355 nm). In the case of TNF (Fig. 7.7.9), the large reductions in fluorescence intensity observed on unfolding the protein in guanidine or acid show that the tryptophan residues in the native protein are relatively unquenched. By contrast, tryptophan fluorescence in native chymotrypsinogen A is seen to be considerably quenched (Fig. 7.7.10). Although specific assignments of these interactions cannot be derived from the spectra, the intensity of fluorescence provides a further empirical characteristic of a protein. Note that caution should observed when comparing recorded values of λmax and Imax (see Basic Protocol 1). Quenching Fluorescence quenching provides an indicator of the solvent accessibility of tryptophans and of the dynamics of protein conformation. The presence of certain extrinsic groups in physical contact with an excited chromophore can lead to sharing or transfer of the excitation energy, with consequent reduction in the quantum yield and diminution of fluorescence intensity. The efficiency of this quenching depends on the electronic structure of the extrinsic molecule or ion. The fact that it depends also on its accessibility to the chromophore means that a study of quenching can provide a further and independent probe of tryptophan environment. Iodide ion (I−) cesium ion (Cs+), and succinimide are useful quenchers for determining accessibility (a fuller list of quenchers for proteins is provided by Eftink and Ghiron, 1981). The ionic character of I− and Cs+ and the relatively large size of succinimide mean that these quenchers cannot readily penetrate the nonpolar interior of the protein, so they can be used to differentiate between buried and solventaccessible chromophores. Their accessibility to an aromatic residue in the surface of the protein will depend, in addition, on the stereochemistry, chemical nature, and, in particular, charge of the surrounding groups. Quenching can be readily quantitated using the
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1.0 0.9 0.8
Fluorescence
0.7 0.6 0.5 0.4 0.3 0.2 1.0 0 300
350
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Figure 7.7.9 Quenching of fluorescence of tumor necrosis factor (TNF) upon denaturation. Fluorescence emission spectra are shown for native TNF (solid line), guanidine-unfolded TNF (dotted line), and acid-denatured TNF (dashed line). Parameters: TNF concentration, 30 µg/ml; λex = 280 nm; bandwidths ranging from 16 to 24 nm. Measurements made using Perkin-Elmer MPF3 spectrofluorimeter. Reprinted from Hlodan and Pain (1994) with permission of Elsevier Science.
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300
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Determining the Fluorescence Spectrum of a Protein
Figure 7.7.10 Dequenching of chymotrypsinogen A on unfolding. Native chymotrypsinogen A (N) was scanned as in Figure 7.7.8. Unfolded chymotrypsinogen A (U) was scanned in 6 M guanidinium chloride using the following parameters: A280 = 0.05; λex = 280 nm; excitation and emission bandwidths, 2.5 nm; scan rate 100 nm/min; single scan, smoothed. The tryptophan fluorescence is seen to be considerably quenched in the folded protein. Measurements made using a Perkin-Elmer LS50B.
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Stern-Volmer equations to give the quenching constant (KSV) and the fraction of the initial fluorescence that is accessible to the quencher (see Basic Protocol 1). The quenching of fluorescence of phosphoglycerate kinase (PGK) is shown plotted according to the Stern-Volmer equation (see Fig. 7.7.6A). The quenching levels off above 0.9 M succinimide and 0.6 M iodide, respectively, indicating that part of the fluorescence is unquenched by these quenchers. The different degrees of quenching by iodide and succinimide is due to the nature of the neighboring groups affecting access to the tryptophan. The plot of some of the data according to the modified Stern-Volmer equation (Fig. 7.7.6B) shows the fluorescence quenched by succinimide to be 42% of the total. The different degrees of quenching at 316 and 350 nm are due to the two tryptophans having substantially different values of λmax, and hence different degrees of accessibility to solvent. Thus, taking into account the crystal structure of PGK, one of the two tryptophans, W333, is seen to be buried and inaccessible to quencher while the other, W308, is quenched with a quenching constant of KSV = 0.39 liters/mol. Another quencher, acrylamide, can quench the fluorescence not only of exposed but also of buried tryptophan residues based on its ability to diffuse through the protein structure. The efficiency of quenching is, therefore, a reflection of the tightness of packing and the dynamics of the protein. Figure 7.7.11 shows the results of acrylamide quenching of the
2.00
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Quencher concentration (mol/liter) Figure 7.7.11 The quenching of buried tryptophan fluorescence in 3-phosphoglycerate kinase (PGK). In the presence of 1.12 M succinimide (shown to quench the exposed tryptophan fluorescence; see Fig. 7.7.6), fluorescence is further quenched by acrylamide, which is able to gain access to the buried tryptophan. Fluorescence was measured at 316 nm (filled circles) and at 350 nm (open circles) and plotted according to the Stern-Volmer equation (see Basic Protocol 2, step 4). Fluorescence intensities were corrected for dilution and for quencher absorbance using ε295 = 0.236 for acrylamide. The linear plot indicates the quenching of a single residue. Measurements made using a Perkin-Elmer MPF3-L. Reprinted from Varley (1991) with permission of Elsevier Science.
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buried W333 of PGK (in this experiment the more exposed W308 is quenched by succinimide). The linearity of the plot, combined with the coincidence of the quenching measured at 316 and 350 nm (in contrast to Fig. 7.7.6), shows that a single tryptophan only is being quenched by the acrylamide. A quenching constant KSV = 0.94 liters/mol is obtained from the slope. In general, bimolecular rate constants for quenching, which reflect more directly the dynamics of the protein, can also be calculated if fluorescence lifetime measurements are available (see discussion of Stern-Volmer equation in Basic Protocol 2; see also Eftink and Ghiron, 1981). Fluorescence Depolarization If a polarizer is placed in the excitation beam, the radiation passing through the sample solution will be plane-polarized. This beam will now be absorbed only by those chromophores whose transition dipoles are in the same plane (see Fig. 7.7.12). If the chromophore does not rotate within 10−8 sec (see Fig. 7.7.7), the fluorescence will be emitted with the same degree of polarization and will pass through a second polarizer (or analyzer) oriented parallel to the first polarizer. If, on the other hand, the excited chromophores rotate before fluorescence takes place, emission will take place from a smaller number of molecules reoriented with transition dipoles parallel and perpendicular to the exciting radiation
S
no rotation P
D
A
rotation
Determining the Fluorescence Spectrum of a Protein
Figure 7.7.12 Depolarization of fluorescence indicates rotation of the chromophore. Monochromatic radiation from the source (S) has all but the vertically polarized electric vector removed by the polarizer (P). This is absorbed only by those molecules (see Fig. 7.7.5) in which the transition dipole of the chromophore is aligned vertically. In the case where these molecules do not rotate appreciably before they fluoresce (“no rotation”), the same molecules will fluoresce (indicated by shading) and their emitted radiation will be polarized parallel to the incident radiation. The intensity of radiation falling on the detector (D) will be zero when the analyzer (A) is oriented perpendicular to the polarizer. In the case where the molecules rotate significantly before fluorescence takes place, some of the excited chromophores will emit radiation with a horizontal polarization (“rotation”) and some with a vertical polarization. Finite intensities will be measured with both parallel and perpendicular orientations of the analyzer. The fluorescence from the remainder of the excited molecules will not be detected. The heavy arrows on the left of the diagram illustrate the case where there is rotation.
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(Fig. 7.7.12). The emitted radiation is now said to be depolarized. The polarization (P) can be quantitated as P = (Ipar − Iperp)/(Ipar + Iperp) where Ipar and Iperp, respectively, are the intensities measured parallel and perpendicular to the original plane of polarization, usually vertical. For chromophores that are part of small molecules, or that are located flexibly on large molecules, the depolarization is complete—i.e., P = 0. A protein of, say, Mr = 25 kDa however, has a rotational diffusion coefficient such that only limited rotation occurs before emission of fluorescence and only partial depolarization occurs, measured as 1 > P > 0. The depolarization can therefore provide access to the rotational diffusion coefficient and hence the asymmetry and/or degree of expansion of the protein molecule, its state of association, and its major conformational changes. This holds provided that the chromophore is firmly bound within the protein and not able to rotate independently. Chromophores can be intrinsic tryptophan or extrinsic, covalently bound fluorophores—e.g., the dansyl (5-dimethylamino-1-naphthalenesulfonyl) group. More detailed information can be obtained from time-resolved measurements of depolarization, in which the kinetics of rotation, rather than the average degree of rotation, are measured. For further details, see Lakowicz (1983) and Campbell and Dwek (1984). Although potentially powerful, the interpretation of fluorescence depolarization results requires extreme caution. Table 7.7.1 provides a summary guide for interpreting results obtained using the techniques described in this unit. In conclusion, given a protein in which tryptophan is partially or wholly buried, quantitative estimates of quenching by different quenchers provide a further fingerprint that is specific to the particular conformation of a recombinant protein and which, combined with other fluorescence, circular dichroism, and gel electrophoresis data, forms the basis for specific identification and quality assessment. COMMENTARY Background Information Fluorescence depends on the atomic environment of the tryptophan and tyrosine residues and on their accessibility to solvent. Although the structural information is limited strictly to the immediate neighborhood of the relatively few aromatic residues, the cooperative nature of protein conformations means that changes in aromatic fluorescence frequently reflect more widespread changes in conformation. Similarly, measures of solvent accessibility obtained from fluorescence quenching reflect changes in the folding and packing of nonaromatic residues in a very sensitive way. Along with its sensitivity to small changes in conformation, fluorescence offers the additional advantage of requiring very small amounts of protein, in favorable cases down to tens of micrograms. The method is less powerful in characterizing proteins lacking tryptophan, for rea-
sons mentioned earlier (see Support Protocol), and does not apply at all to the rare proteins that lack both fluorophores. Near-UV circular dichroism measurements (UNIT 7.6) can still be of considerable value for proteins containing tyrosine and, in both cases, both far-UV circular dichroism and urea-gradient gel electrophoresis (UNIT 7.4) provide powerful contributions to characterization. Extrinsic fluorescent probes can provide additional useful information about the integrity of protein conformation. 8-{2-[(iodoacetyl)ethyl]amino} naphthalene sulfonic acid (ANS) fluoresces only weakly in free solution but strongly when bound to a nonpolar surface. Originally used as a competitive probe to monitor fatty-acid binding to albumin, it has been widely used in studies of protein conformation—in partially denatured states at equilibrium as well as in transient intermediates that accumulate during protein folding (reviewed
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by Christensen and Pain, 1994). Fully folded native proteins in general bind only limited amounts of ANS, although these are sufficient to characterize the surface hydrophobicity of a number of proteins (Cardamone and Puri, 1992). Fully unfolded proteins do not bind
Table 7.7.1
ANS. Partially folded or partially unfolded proteins will, however, bind ANS—indicating the presence of associated nonpolar groups that constitute a surface site, or the existence of relatively loosely packed hydrophobic cores into which ANS can intercalate. This and sim-
Summary of the Interpretation of Fluorescence Spectra of Proteinsa,b
1. All fluorescence of a protein is due to tryptophan, tyrosine, and phenylalanine, unless the protein contains another fluorescent component. 2. Maximum fluorescence of tryptophan occurs at lower wavelengths (<350 nm) and has greater intensity the lower the polarity of its environment. a. If λmax is located at lower wavelengths when the protein is in a polar solvent—e.g., water—the tryptophan must be buried in a nonpolar environment. b. If λmax is shifted to shorter wavelengths when the protein is transferred to a nonpolar medium, either the tryptophan is on the surface of the protein or the solvent induces a conformational change that causes it to be exposed to the solvent. 3. If a substance known to be a quencher quenches tyrosine and/or tryptophan fluorescence in a protein, these residues must be on the surface. If it fails to do so, there are several possible reasons: a. The residue may be buried in the interior of the protein. b. The residue may be in a crevice whose dimensions are too small for the quencher to enter. c. The residue may be in highly charged surroundings whose charge may repel a quencher with the same charge. In such a case, a quencher of the opposite charge or no charge would be expected to quench the fluorescence. 4. If a substance that does not affect the quantum yield of the free amino acid affects the fluorescence of a protein, it probably does so by inducing a conformational change in the protein. 5. If tryptophan or tyrosine are in a polar environment, their quantum yield decreases with increasing temperature, whereas in a nonpolar environment there is little change. Deviation from a monotonic decrease with increasing temperature signifies a change in polarity of the tryptophan environment, the sign of the change indicating the direction of change in polarity. Either indicates a conformational change in the protein. 6. Tryptophan fluorescence is quenched by protonated acidic groups, either in the neighboring environment or covalently attached as an α-carboxyl group. If the fluorescence changes with a particular pK, then a residue with that pK must be close to the tryptophan. Such an interpretation only holds if evidence is provided that no pH-induced conformational change taking place. 7. If the binding of a substance is accompanied by quenching of tryptophan fluorescence, either there is a resulting conformational change or tryptophan is in or very near the binding site. Furthermore, because a decrease in polarity causes a shift to shorter wavelengths, a decrease in λmax on binding indicates that water is excluded in the complex. 8. If the absorbance spectrum of a small molecule overlaps the emission spectrum of tryptophan and if the distance between them is small, quenching will be observed. If binding of such a molecule to a protein quenches tryptophan fluorescence, tryptophan must be in or near (<2 nm away from) the binding site. aAdapted from Freifelder (1982). Reproduced with permission of W.H. Freeman, New York. bIn making these interpretations, two important facts must be borne in mind. First, fluorescence is so sensitive to
Determining the Fluorescence Spectrum of a Protein
environmental factors that alternative interpretations must always be considered. Second, if a protein contains more than one tryptophan, each may have a different quantum yield. For this reason the absolute magnitude of changes cannot be used to determine the fraction in a given environment.
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ilar probes are mainly used as empirical indicators of conformational change, as the detailed mechanisms of interaction are not well defined. Fluorescence energy transfer, which can be observed when the absorbance spectrum of a fluorescent probe such as the dansyl group overlaps the emission spectrum of tryptophan, provides a means of estimating distances between groups, and hence of conformation and conformational change. An example where this approach is used alongside ANS fluorescence to follow protein folding is reported by Agashe et al. (1995). In the present context, fluorescence is an important tool for the rapid comparison of a sample of recombinant wild-type or mutant protein with a standard sample of the recombinant or the authentic protein, as part of the process of characterizing and authenticating a recombinant protein. Coupled with measurements of the depolarization of fluorescence, the method can provide information on the dynamics of a protein and on the change in dynamics in response to, for example, binding of ligands.
Critical Parameters Recording a spectrum is basically very simple, but the critical element is to avoid the production of artifacts. For this reason, it has to be emphasized that scrupulous attention to the cleanliness of cells and of the cell compartment of the spectrofluorimeter is essential, as is attention to the clarification of solutions and buffers (see Strategic Planning). It is important also to make intelligent choices in the selection of instrumental parameters to optimize resolution and signal-to-noise ratio (see Basic Protocol 1).
solubility of many proteins. For this reason, it is important to pay strict attention to monitoring aggregation (see discussion of Clarification of Solutions in Strategic Planning).
Literature Cited Agashe, V.R., Shashtry, M.C.R., and Udgaonkar, J.B. 1995. Initial hydrophobic collapse in the folding of barstar. Nature 377:754-757. Cardamone, M. and Puri, K. 1992. Spectrofluorimetric assessment of the surface hydrophobicity of proteins. Biochem. J. 282:589-593. Campbell, I.D. and Dwek, R.A. 1984. Biological Spectroscopy. Benjamin/Cummings, Menlo Park, Calif. Christensen, H. and Pain, R.H. 1994. The contribution of the molten globule model. In Mechanisms of Protein Folding (R.H. Pain, ed.) pp. 55-79. Oxford University Press, Oxford. Eftink, M.R. and Ghiron, C. 1981. Fluorescence quenching studies with proteins. Anal. Biochem. 114: 199-227. Hlodan, R. and Pain, R.H. 1994. Tumour necrosis factor is in equilibrium with a trimeric molten globule at low pH. FEBS Lett. 343:256-260. Lakowicz, J.R. 1983. Principles of Fluorescence Spectroscopy. Plenum Press, New York. Schmid, F.X. 1989. Spectral methods of characterizing protein conformation and conformational changes. In Protein Structure (T.E. Creighton, ed.) pp. 251-285. IRL Press, Oxford. Varley, P.G., Dryden, D.T., and Pain, R.H. 1991. Resolution of the fluorescence of the buried tryptophan in yeast 3-phosphoglycerate kinase using succinimide. Biochim. Biophys. Acta 1077:1924.
Key Reference Lakowicz, J.R. 1983. See above.
Troubleshooting
An essential text, providing a thorough, but clear and readable description of the practice and interpretation of fluorescence spectroscopy, with particular reference to proteins.
One of the main causes of artifacts in the spectroscopy of proteins is the frequent tendency of proteins to aggregate, either during the folding step in recovery from inclusion bodies or as a result of the delicate balance of
Contributed by Roger H. Pain JoÓef Stefan Institute Ljubljana, Slovenia
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Light Scattering
UNIT 7.8
Light scattering methods can provide information about the native molecular weight, oligomeric composition, and gross conformation of a protein in solution. These methods are particularly well suited for studying large oligomeric systems or glycoproteins and can be used to characterize much larger structures involving protein such as viruses and even bacterial spores (Harding, 1997). Light scattering techniques are not so well suited for characterization of smaller protein systems (where the molecular weight, M, is <30,000 Da); in these cases other methods such as analytical ultracentrifugation (UNIT 7.5) and solution X-ray scattering are more suitable. All light scattering measurements on solutions of proteins and protein assemblies are based on the principle of analyzing the intensity of light scattered by the solution (Fig. 7.8.1), either in terms of the time-averaged intensity (“classical” or “static” light scattering) or intensity fluctuations with time (“dynamic” or “quasielastic” light scattering) at a given angle or series of angles. There are three types of “static” light scattering experiment:
1. Turbidimetry, which is simple but gives only crude molecular-weight estimates for large assemblies; 2. Low-angle light scattering, which is also simple and gives molecular-weight and molecular-weight-distribution information; 3. Multiangle light scattering, which gives more reliable molecular-weight and molecularweight-distribution information and, for proteins of molecular weight at least ∼50,000 Da, solution conformation information. There are also three types of “dynamic” light scattering measurements: 1. Fixed-angle (90°-angle) measurements, which are simple and give an estimate for the translational diffusion coefficient and an idea of sample polydispersity for approximately globular macromolecules; 2. Variable-angle measurements, which are less simple, give more reliable estimates for translational diffusion coefficient and sample polydispersity, and can in some circumstances yield an estimate for rotational diffusion coefficients;
sample cell monochromatic laser radiation
I0
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Figure 7.8.1 Light scattering studies on biomolecular solutions involve consideration of the relationship between the time-averaged scattered light intensity, I(θ), with the incident intensity, I0, and angle of detection, θ (“static light scattering”) or of the rapid fluctuations of the scattered intensity, I(θ), with time, τ (“dynamic light scattering”). Turbidimetry involves static light scattering measurements on the relation between I0 and transmitted light intensity, I (at zero angle) alone. Contributed by Stephen E. Harding and Kornelia Jumel Current Protocols in Protein Science (1998) 7.8.1-7.8.14 Copyright © 1998 by John Wiley & Sons, Inc.
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3. Electrophoretic light scattering measurements (“ELS”), popularly used for studying colloid solubility.
STATIC LIGHT SCATTERING ANALYSIS OF PROTEIN SOLUTIONS Basic Theory The basic equation for the angular dependence of light scattered from a solution of proteins or protein assemblies is the Debye-Zimm relation (Zimm, 1948), shown in Equation 7.8.1,
1 Kc (1 + 2 A2 c +) = Rθ MP(θ) Equation 7.8.1
In this equation, A2 (ml mol/g2) is the thermodynamic (2nd virial) nonideality coefficient. Rθ is the Rayleigh excess ratio (the ratio of the intensity, Iθ, of excess light scattered compared to pure solvent) at an angle θ to that of the incident light intensity, I0 (a correction term is necessary if unpolarized light is used but not necessary if lasers are used). K is an experimental constant dependent on the square of the solvent refractive index, the square of the refractive index increment (dn/dc in ml/g), and the inverse fourth power of the incident wavelength, λ (cm). M is the molecular weight in Da, c is the solute concentration (g/ml), and P(θ) is the form factor. Equation 7.8.1 is valid if the proteins/protein assemblies satisfy the Rayleigh-Gans-Debye criteria illustrated in Equation 7.8.2:
n − 1 << 1 n0 and 4 πn0 n − 1 << 1 d λ 0 n0 Equation 7.8.2
where n is the refractive index of the solution, n0 is the refractive index of the solvent, λ0 is the incident wavelength (in vacuo), and d is the maximum dimension of the particle. The form factor P (θ) can also be given to a good approximation by Equation 7.8.3, Light Scattering
1 + 16 πRθ2 2 θ 1 1 ≈ sin ( )( + 2 A2 c) P(θ) 3λ2 2 M Equation 7.8.3
Rg is extensively referred to as the “radius of gyration” of the macromolecule and c is the solute concentration (g/ml). If the solute is heterogeneous, M (g/mol = Da) will be a weight average, MW, and Rg a z-average. Equation 7.8.1 is generally a good representation for particles whose maximum dimension is between λ/20 and λ. Since λ is typically ∼600 nm, this covers all proteins and protein assemblies up to the sizes of filamentous viruses. For larger particles, much more complex representations are necessary. For particles of dimension <λ/20 (M < 50,000 Da) the angular term in Equation 7.8.1 is small. No angular-dependence measurements are necessary to obtain M (although Rg cannot be obtained; if this is needed then X-ray or neutron solution scattering measurements need to be employed instead). For particles of dimension >λ/20, a double extrapolation to zero angle and zero concentration is necessary. This is usually performed on a grid-like plot referred to as a “Zimm plot” or via measurement at a single angle assumed small enough so that sin2(θ/2) ∼ 0. Other methods of representing the data have been in terms of “disymmetry”: z(θ) (defined as the ratio of the scattering intensity at an angle θ, typically 45°, to that at 180° − θ) versus θ plots. Both z(θ) and Rg provide useful guides to the conformation of a macromolecule, with z(θ) being more popular with linear DNA molecules and Rg being more suited for descriptions of protein conformation. Other useful representations for nonprotein systems are also available (see Burchard, 1992). For fairly rigid protein systems, Rg can be used directly to model gross conformation— either as an additional parameter to the diffusion coefficient (see Dynamic Light Scattering Analysis) and other hydrodynamic parameters for representing the structure in solution of complex protein systems in terms of bead modeling (Garcia de la Torre et al., 1997)—or as a parameter, after combination with the second virial coefficient A2 and a parameter from solution viscometry known as the intrinsic viscosity, for representing the triaxial structure of a protein (Harding et al., 1997). The principal useful parameters to be derived from static light scattering measurements
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0. 0 0. 010 0 0. 014 00 17
0 C
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0 0
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Figure 7.8.2 Zimm biaxial plot for a (diptheria) toxin-antibody aggregate. The arbitrary scaling constant, k, equals 200 ml/g. The molecular weight (Mw) as calculated from the (reciprocal) common intercept of the c = 0 and θ = lines is equal to ∼78 × 106 Da. Data from Johnson and Ottewill (1954) and Johnson (1993).
are thus M, Rg, and to a lesser extent A2. M can be obtained to a reasonable accuracy (usually to within 5%). The extraction of Rg is more difficult, requiring considerable care in the form of the angular extrapolation, and becomes even more difficult as the lower limit of λ/20 is approached. The use of light scattering photometers incorporating a flow cell that can be linked directly on-line to size-exclusion chromatography columns is becoming increasingly popular, particularly for the characterization of polydisperse systems (the hallmark, for example, of many glycoproteins), and as an on-line sample clarification system.
Turbidimetry Turbidimetry involves the measurement of the total loss of intensity by a solution through scattering, summed over the entire angular intensity envelope and compared with the intensity of the incident radiation. It can be used to measure the molecular weights of protein assemblies of M >105 Da (Bahls and Bloomfield, 1977). It is a type of measurement that can be performed on a good-quality spectrophotometer (i.e., one whose detector does not accept appreciable amounts of scattered light). Measurements have to be made at wavelengths away from the influence of absorption maxima.
Low-Angle Light Scattering (LALLS)
As θ approaches 0, Equation 7.8.1 becomes Equation 7.8.4:
Kc 1 = + 2 A2 c Rθ M Equation 7.8.4
Low-angle light scattering photometers permit the measurement of Kc/Rθ at one fixed, small scattering angle θ (usually <8°) at which Equation 7.8.4 is taken to be valid (see Jumel et al., 1992). Although the angle used is assumed low enough that no angular correction of the scattering data is required, an extrapolation to zero concentration of Kc/Rθ may be necessary. The method can thus provide values for M and A2 of a system, but not for Rg since no record is made of the angular dependence of Kc/Rθ . At low concentration, and especially when the light scattering detector is on-line to size-exclusion chromatography columns, the further approximation that 2A2c ∼ 0 can be made, and hence M is simply ∼Rθ/Kc. Since lasers are now routinely used as the light source, this technique takes the popular acronym of “LALLS” (low-angle laser light scattering).
Multiangle Light Scattering (MALLS) Multiangle light scattering measurements are based on the full Debye-Zimm equation (Equation 7.8.1). Performing measurements at multiple angles permits extrapolation of the ratio Kc/Rθ to zero sin2(θ/2), which, together with an extrapolation to zero concentration,
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forms the basis of the Zimm plot (Fig. 7.8.2), a method that can yield M, A2 and Rg. Values for A2 returned are typically between 10−5 ml mol/g2 (for globular proteins) and 10−3 ml mol/g2 (for large glycoconjugates). A warning is in order here—a common misconception is to ignore these values as small numbers. In actuality, it is the product 2 × A2 × M × c that manifests the influence of nonideality. The expression 1/(1 + 2A2Mc) represents the factor by which an “apparent” molecular weight measured at a finite concentration c underestimates the true or “ideal” value (Tanford, 1961). An opposing effect to that of thermodynamic nonideality is that of reversible interaction phenomena (as represented for example by a molar dissociation constant, Kd), and in some cases (“pseudoideality”) the two effects approximately cancel. As with LALLS, lasers are now routinely used as the light source, and hence the acronym “MALLS” (multiangle laser light scattering) has now been adopted. Besides the potential for extracting Rg, a more important advantage of MALLS over LALLS is that the angular extrapolation permits identification and avoidance of any spurious results at the lowest angles. Plots of Kc/Rθ are only linear over a limited range of angles. For globular proteins of M < 50,000 Da (corresponding to a maximum dimension ∼λ/20), the angular dependence of Kc/Rθ will be negligible, permitting a more accurate determination of M but of course excluding the possibility of measuring Rg. As with LALLS, at low concentration, the further approximation that 2A2c ∼ 0 can be made; hence Equation 7.8.1 reduces to
Kc 1 = Rθ MP(θ) Equation 7.8.5
and where there is no angular dependence, this reduces further to M ∼ Rθ/Kc, as noted above.
Limitations of Static Light Scattering Methods
Light Scattering
The principal limitation for all these “static” light scattering measurements is the need for sample clarification to avoid dust and supramolecular aggregates (see Clarification of Solutions and Scattering Cells). This is especially serious if solutions of proteins of M <50,000 Da are being studied, and also if LALLS is used, since large-particle contamination effects become disproportionally larger at low angles and
the LALLS instruments provide no angular check for this. Another limitation is that a separate precise measurement of the refractive index increment dn/dc is required, preferably at the same wavelength used in the light scattering photometer. With MALLS, Rg cannot be measured as noted above for particles of molecular weight <50,000 Da. For very large macromolecular assemblies (maximum dimension >λ/2), the reliability of the theory on which Equation 7.8.1 is based (known as RayleighGans-Debye theory) becomes doubtful.
SEC-LALLS and SEC-MALLS A revolutionary development has been the coupling of LALLS or MALLS photometers to size-exclusion chromatography (SEC) systems (Jumel et al., 1992; Wyatt, 1992). This is made possible by replacing the standard light scattering cuvette with a flow cell that can be coupled downstream from an HPLC pump and size-exclusion chromatography column(s), and upstream from a (UV-absorption or refractive index) concentration detector (Fig. 7.8.3). This has the double advantage of providing an online clarification system to remove supramolecular contamination and allowing fractionation of polydisperse materials prior to light scattering analysis. However, care must still be taken, particularly against the spurious shedding of column material. The coupled SEC systems are referred to as “SEC-LALLS” or “SEC-MALLS.”
Samples for Analysis in Static Light Scattering Solutions should be dialyzed against an appropriate buffer of defined pH and ionic strength, I. Normally an ionic strength of at least 50 mM is needed to suppress the contribution of molecular charge on the protein to the nonideality coefficient (A2 in Equation 7.8.1) although, of course, the I chosen will obviously depend on its effect on the protein. The concentration of protein required depends on: (1) its molecular weight, since the scattering signal is approximately proportional to the product of concentration and molecular weight; (2) the output of the laser; and (3) the clarity of the solutions, since substantial material may be lost on filtering or from the column in SEC-LALLS or SEC-MALLS. For scrupulously clean solutions, a 5-mW laser for a loading concentration of ∼3-mg/ml is sufficient for proteins of molecular weight ∼40,000 Da. For smaller proteins, a proportionally higher con-
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eluent injector degasser
columns
pump
light-scattering detector
concentration detector
data transfer
waste
computer
Figure 7.8.3 Experimental setup for SEC-MALLS (see Wyatt, 1992).
centration and/or higher laser power is required. With regard to volume requirements, if a flow cell arrangement is used (linked, e.g., to an SEC system), loading volumes can be as low as 0.1 ml. For use with scintillation vials (which are nowadays used instead of fluorimeter-type cuvettes), ≥3 ml of solution is required. Square cuvettes, of course, prevent measurements in the angular part of the scattering envelope near the corners of the cuvette; if cylindrical cuvettes are employed, then small diameters (<2 cm) are to be avoided because of extraneous scattering or reflections from the glass walls, although large-diameter cuvettes can be expensive in terms of quantity of solution required. Flow cells are now preferred, and a typical experimental set up would have MALLS coupled to an SEC, but with a separate injection port (via a filter or guard column) if the SEC separation is not needed.
Clarification of Solutions and Scattering Cells For molecular weights <200,000 Da, the most serious experimental problem has been that of clarification. All traces of dust and supramolecular aggregates have to be removed
since experiments on incompletely purified material are “not useful” (Johnson, 1993), and clarification can lead to a significant loss of material. Contaminating particles can be removed by ultracentrifugation and ultrafiltration. If scintillation vials are used, these too must be scrupulously clean. They should be cleaned and rinsed in double-distilled water, then dried and covered with aluminum foil to prevent any dust from entering the vials. It is also extremely important that there are no scratches or fingerprints or other marks on the outside of the vial. Sample preparation for batch work should, if possible, be carried out in a flow cabinet to prevent any dust from entering the solutions. If a flow-cell device is used for clarification (Sanders and Cannell, 1980), this too will frequently need to be removed and cleaned using detergent, acetic acid, and a final necessary rinse with ultrafiltered water. An acetone reflux may also be occasionally necessary. When attached to an SEC column, prior to solution injection and after all the solution has been eluted, the solvent eluant must be monitored for any shedding from the column material. When the solution is injected, this needs to be done via a Millipore filter of appropriate size.
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Calibration (LALLS and MALLS) The quantities directly measured by a light scattering photometer are voltages and not light scattering intensities. For this reason, the instrument has to be “calibrated”—usually with a strong Rayleigh scatterer with a known Rayleigh ratio such as toluene (Stacey, 1956; Johnson and McKenzie, 1977). Measurement of the scattered and incident light intensities (Iθ and I0, respectively) then facilitates calculation of the calibration constant, which is then used to calculate Rayleigh ratios for sample solutions from the output of the instrument. Calibration does not mean that the light scattering measurements have to be made relative to protein standards as in calibrated gel chromatographic methods. A typical calibration procedure would include the following steps. 1. Switch on the light scattering detector and laser at least 1 hr before measurements are made. 2. Make sure that the sample cell is scrupulously clean and the calibrating solvent (in most cases toluene as it is the strongest Rayleigh scatterer with a known Rθ of 1.406 × 10−5cm−1 at a wavelength of 633 nm) is HPLC grade. 3. Inject the solvent into the cell via a membrane filter (preferably ≤0.2 µm pore size). 4. Measure the scattered light intensity and calculate the instrument calibration constant by means of the dedicated software (e.g., ASTRA, Wyatt Technology), using the known Rayleigh ratio of the solvent.
Normalization (MALLS)
Light Scattering
For simultaneous multiangle detection in MALLS photometers of the type now commonly available, the detectors have to be “normalized” to allow for the different scattering volumes as a function of angle and the differing responses of the detectors. This is normally achieved using a solution of molecules that scatter light isotropically (i.e., with equal intensity in all directions)—as is the case when the diameter of the sample molecule is less than ∼λ/20 of the incident wavelength. The best molecules of this kind for use in a chromatography system are either a polystyrene standard in toluene or tetrahydrofuran (THF) with molecular weight of ∼30,000 Da or a pullulan or dextran standard of 20,000 to 30,000 Da in aqueous solution; the Rg of these molecules is ∼5 nm. In the batch mode—i.e., with the sample in a vial–10 mg/ml solutions of 4000-Da polystyrene in toluene or THF, or 5000 Da pullulan in water work well; these standards have Rg’s of ∼2 nm.
The procedures described below are typical normalizations for the MALLS instruments distributed by Wyatt Technology. Normalization for chromatography Inject a known amount of sample into the system and collect data using the ASTRA software. After the run has finished, set baselines and peak limits. For normalization, it is necessary to set very narrow, symmetrical limits at the top of the peak. Enter the Rg value for the standard used in the Normalize menu, click “normalize,” and the normalization coefficients will be displayed on the screen. This procedure only needs to be repeated if the solvent is changed or the flow cell has been cleaned. Normalization for batch collection Remove the flow cell from the read head and assemble alignment rings and flow-to-batch conversion plate in read head as described in the manual. First, place a scintillation vial containing the filtered solvent into the aperture and observe the scattering intensities. Once these have stabilized, begin data collection, which will stop after a preset volume. Save the data. Next, insert the vial containing the sample for normalization measurement and again wait for signals to stabilize. Append the file containing the values for the solvent. After collection has stopped, measure the scattering for the solvent again. Set a baseline and peak limits that cover most of the plateau corresponding to the normalization standard. Go to the Normalization menu and enter the concentration and Rg of the normalization standard. Click “normalize” and the normalization coefficients will appear on the screen.
Evaluation of Molecular Weight (LALLS) After measurement of Rθ, the molecular weight, M, can be calculated from this value, the concentration, the experimental constant K, and a subsequent extrapolation to c = 0 (Equation 7.8.4) if the nonideality term A2c is significant. The concentration, c (from, e.g., UV absorbance or refractometry) should be known as accurately as possible.
Evaluation of Molecular Weight and Radius of Gyration (MALLS). If the nonideality term A2c (and of course any higher-order terms) is insignificant (i.e., if the measurements can be performed at low enough concentration), then only an extrapola-
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2.00 × 107
Rθ /Kc
1.60 × 107 1.20 × 107 8.00 × 106 4.00 × 106 0.00
0.0
0.2
0.4
0.6
0.8
1.0
sin2(θ /2)
Figure 7.8.4 Debye plot (angles from 21.7° to 58.3°) for a heavily glycosylated protein system (pig gastric mucin glycoprotein). A first-order (linear) fit is shown. This figure illustrates the need for caution when interpreting static light-scattering data: the upward curvature at the lower angles for a simple protein system (or downward curvature in a Kc/Rθ versus sin2(θ/2) plot) would normally be ascribed to supramolecular contamination, and the data points would be ignored or given low weighting. However, for a genuinely polydisperse system, the lower-angle measurements contain proportionally higher information about the higher-molecular-weight species in a distribution, and removal of the low-angle data and/or choice of a first-order fit could bias the distribution towards the low-molecular-weight end.
tion to zero angle and not zero concentration is necessary (see Equation 7.8.5). Sometimes the inverse form of Equation 7.8.4 is used, known as a “Debye plot” of Rθ/Kc versus sin2(θ/2) (Wyatt, 1992). Although this invokes a further approximation, the ordinate axis does correspond to an apparent molecular-weight axis (Fig. 7.8.4). It must be noted that Figure 7.8.4 is a rather extreme example for a highly glycosylated and polydisperse system. Figure 7.8.4 also shows that care must be taken in choosing the appropriate fit, and that there is a proportionally larger effect at low angles of supramolecular contaminants. The order of the fit chosen (usually linear or quadratic) has a more dramatic effect on the (limiting) slope of such plots and hence on the value for Rg returned. This feature needs to be borne in mind when conclusions on macromolecular conformation based on the relationship between M and Rg are being drawn, particularly using the SECMALLS devices considered below. If the nonideality term is significant, and an assumed value for A2 cannot be taken as zero (which can in fact be predicted from the triaxial shape of the protein; Harding et al., 1997)—and charge effects are not significant, then an additional extrapolation to zero concentration is necessary. Both angular and concentration extrapolations are usually done on the same plot, known as a Zimm plot (Fig. 7.8.2), which can yield M from the reciprocal of the common
intercept, and estimates for A2 (from the θ = 0, c extrapolation line) or Rg [from the c = 0, sin2 (θ/2) extrapolation line]. However it cannot be overstressed that the Rg returned can be very sensitive to the order of extrapolation used. An arbitrary (positive or negative) constant k is used on the abscissa to scale the data points better.
Evaluation of Molecular Weight Distribution (SEC-LALLS) and (SEC-MALLS) Absolute molecular weight distributions (Fig. 7.8.5) of mixed protein systems or glycosylated protein systems such as mucins, glycoproteins, or glycosaminoglycans may be obtained with either of the above techniques.The sample is injected via the injection valve and separated by the column system. Effluent from the column(s) is monitored by light scattering and concentration detectors. Once the peak area from the chromatogram has been chosen, the concentration and Rθ at each data point within this area are known from light scattering and concentration detectors, respectively, and the molecular weight (M) at each of these points is calculated using Equation 7.8.4 (for SEC/LALLS) or Equation 7.8.5 (for SEC/MALLS). Molecular weight averages for the whole of the peak area may then be calculated. In addition, a so-called “calibration plot” of molecular weight versus elution vol-
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5.0 (b) 4.0
(c)
g(M )
3.0
(a)
2.0
1.0
0.0 10 5
10 6
107
10 8
Molecular weight (g/mol)
Figure 7.8.5 SEC-MALLS molecular-weight distribution for colonic mucin glycoprotein (peak a) and its thiol-reduced (peak b) and papain digested (peak c) forms (from Jumel et al., 1997). The quantity g(M) is in arbitrary units.
ume can be constructed, thus enabling the elution volume versus concentration plot to be converted to a molecular weight distribution. Such molecular weight distributions thus found will be absolute in the sense that, unlike with conventional SEC, calibration standards of known M are not required. This is particularly useful not only for protein mixtures but for heavily glycosylated systems because of their polydispersity and the difficulty in obtaining standards of the appropriate (often uncertain) conformation. For a more detailed account of the theory the reader is referred to Yau et al. (1979).
DYNAMIC LIGHT SCATTERING ANALYSIS OF PROTEIN SOLUTIONS Basic Theory
Light Scattering
Whereas static light scattering is concerned with the time-averaged scattering intensity properties, dynamic light scattering is concerned with time fluctuations in intensity caused by motions of macromolecules and macromolecular assemblies. Whereas lasers are highly desirable for static light scattering because of their high intensity, collimation, and monochromaticity—with dynamic light scattering they are mandatory because of the requirement for spatial and time (“temporal”)
coherence—light has to be emitted from the source as a continuous wave rather than as short bursts. The physics underlying dynamic light scattering is rather complicated (Brown, 1993), although the basic principle and experimental setup is relatively simple (Fig. 7.8.1 and Fig. 7.8.6). Laser light is directed onto a thermostatted protein solution, and the intensity is recorded at either a single angle or multiple angles using a photomultiplier/photodetector. The moving biomolecules will “Doppler broaden” the otherwise monochromatic incident radiation. The scattered intensity recorded as the number of photons received by a detector (photomultiplier) will fluctuate because of “beating interference” of scattered waves of different but similar wavelength. The situation is analogous to the fluctuations in intensity of a radio channel caused by interference from another radio channel of very close wavelength. This “broadening” of the otherwise monochromatic incident radiation is why the technique is often referred to as “quasielastic light scattering” or QLS. The detector sends the intensity signal to a special computer called an autocorrelator, which compares or correlates the intensity at different times (hence the other name often used—photon correlation spectroscopy or PCS).
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bath cell laser
θ
photomultiplier computer storage
correlator
amplifier-discriminator
Figure 7.8.6 Schematic dynamic light scattering setup for multiangle measurement (see Johnson, 1993).
How rapidly the intensity fluctuates over short time periods or “delay times,” τ (in nsec to msec depending on how mobile the scattering biomolecules/assemblies are), is represented by how a parameter known as the normalized intensity autocorrelation function— g(2)(τ)—decays as a function of τ. The superscript “(2)” is used to indicate that it is an intensity as opposed to an electric field or “(1)” autocorrelation function, and many data sets of g(2)(τ) as a function of τ are accumulated and averaged. The amount of averaging necessary depends on the incident laser intensity and the size and concentration of the scattering biomolecule (at a given concentration, larger molecules scatter more). For globular proteins, sufficient data can usually be acquired over the time scale of one to several minutes. The calculation of the correlation function from the intensity fluctuations is performed after the signal has passed through an amplifier-discriminator via the autocorrelator. Calculation of the diffusion coefficient from the decay of g(2)(τ) with τ is performed on a computer. Some of the most modern instrumentation (particularly fixed-angle photometers) have all of the units depicted in Figure 7.8.6 built into one instrument. Analysis of how the normalized intensity autocorrelation function g(2)(τ) decays as a function of τ can be used to evaluate the trans-
lational diffusion coefficient, D. For dilute systems of spherical or near-spherical (i.e., globular) biomolecules and assemblies, the variation of g(2)(τ) with τ can be represented by the simple logarithmic equation
[
]
ln g( 2 ) ( τ − 1) = −2 Dg( 2 ) τ Equation 7.8.6
where q is known as the Bragg wave vector whose magnitude is defined by q = {4πn/λ}sin(θ/2)—n being the refractive index of the medium, θ being the scattering angle, and λ being the wavelength of the incident light. Thus D can be found from a plot of ln[g(2)(τ) − 1] versus τ, and Figure 7.8.7 shows an example for the motility protein dynein. D can then be converted to standard conditions (i.e., the viscosity and temperature of water at 20.0°C) to give D20,w as noted above, and then extrapolated to zero concentration to give D020,w. An additional extrapolation is necessary if the biomolecule is not globular: this is because at finite angles θ there will be an extra term on the right-hand side of Equation 7.8.6 deriving from rotational diffusional phenomena. This term approaches 0 as θ approaches 0; therefore true D can be measured according to Equation 7.8.6 so long as measurements of the apparent D are made at a
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x x x
–1.0
x
x
x
x
x x
–1.5
x xx x x
x xx x
x
–2.0
x xx xx xx x x x
In[g (2)(τ) – 1]
x
x
x
x
–0.5
x
0.0
–2.5 –3.0 0
10
20
30
40
50
60
Channel number
Figure 7.8.7 Plot of ln[g(2)(τ) – 1] versus channel number for the motility protein dynein. Channel number = delay time/sample time (Wells et al., 1990).
number of angles and at an additional extrapolation to zero angle (or Bragg vector q) is performed. The extrapolations to zero concentration and zero angle (or q) can be performed simultaneously on a biaxial extrapolation plot known as a dynamic Zimm plot (Burchard, 1992). If an angular extrapolation is not necessary, a scattering angle of 90° is usually chosen, and fixed-angle instruments are usually set at this angle (see Claes et al., 1992). At lower angles the problems due to supramolecular contamination are accentuated. If a system is heterogeneous it is possible, at least in principle, to obtain a distribution of diffusion coefficients after various assumptions and mathematical manipulations of the autocorrelation data. The various methods of manipulation have been reviewed by Johnsen and Brown (1992) and several commercially available computer routines are available for performing these. A more simple way of representing heterogeneity is the polydispersity factor (PF) which is obtained by comparing linear with quadratic or quadratic with cubic-order fits of the normalized autocorrelation function decay data (Pusey, 1974).
Translational Diffusion Coefficient
Light Scattering
The translational diffusion coefficient, D20,w or D020,w, obtained by dynamic light scattering or boundary spreading in the ultracentrifuge, can be used to provide a number of useful characteristics about a biomolecular system.
Equivalent hydrodynamic radius (rH) The simplest deduction one can make from D020,w is the size of the biomolecule as represented by the equivalent hydrodynamic radius (also known as the Stokes radius)—rH.What this means is, although the biomolecular system may not be a sphere at all, its diffusive behavior can be represented by an equivalent spherical particle of radius rH. The value of rH can be easily obtained from D020,w via the Stokes-Einstein relation: rH =
k BT 6 πη20,w D0 20,w
Equation 7.8.7
where kB, the Boltzmann constant, is equal to 1.379 × 10−16 erg/°K, T = 293.15°K and η20,w (the viscosity of water at 20.0°C) is equal to 0.01 poise. If D values are not corrected to standard conditions then the appropriate values for T and η have to be used. Frictional coefficient (f) The frictional coefficient provides a handle on molecular weight and shape as discussed below. Like rH it can also be calculated simply from D020,w according to Equation 7.8.8. f=
RT N A D0 20,w
Equation 7.8.8
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where T = 293.15°K and R = 8.314 × 10−7 erg/mol °K and NA is Avogadro’s number (6.022137 × 1023 mol). Molecular weight A more absolute description of biomolecular size than rH is the molecular weight, M. Calculation of M from dynamic light scattering is less direct than from static measurements. However, combination of the D020,w value with the analogous parameter from sedimentation velocity experiments (UNIT 7.5) in the analytical ultracentrifuge—s020,w (in sec)—provides a popular route for obtaining the molecular weight of a biomolecule. The analogous equation to Equation 7.8.8 for the sedimentation coefficient is Equation 7.8.9. f=
(
M 1-vρ20, w NAS
0
)
20, w
Equation 7.8.9
Elimination of f between Equation 7.8.8 and Equation 7.8.9 yields the Svedberg (1927) equation (Equation 7.8.10):
for biomolecules of known D and M (Claes et al., 1992), M of an unknown can be found from its measured D. A conformation has of course to be assumed; the Equation 7.8.11 method for M evaluation is thus not as reliable as the method shown in Equation 7.8.10. Equation 7.8.11 can be taken even further in converting a distribution of diffusion coefficients into a molecular-weight distribution, although such distribution information is not as reliable (because of the approximations made) as that obtained with static light scattering methods coupled directly on-line to a gel-filtration column (Wyatt, 1992). Conformation The translational frictional coefficient (f) can be used directly to provide information on conformation. More convenient, however, is to use the corresponding dimensionless ratio called the frictional ratio (f/f0), where f0 is the frictional coefficient of a spherical particle of the same anhydrous mass and density as the biomolecule. The quantity f/f0 can be calculated from D°20,w by the relation in Equation 7.8.12. 1
s0 RT M = 020 ,w D 20 ,w 1− vρ20 ,w
k T 4 πN A 3 1 f / f0 = B 0 6 πη20,w 3vM D 20,w
Equation 7.8.10
Equation 7.8.12
_ where again T = 293.15°K. The variable v is the partial specific volume of the biomolecule (which can be measured from densimetry or from the composition of the biomolecule) and ρ20,w is the density of water at 20.0°C (0.9982 g/ml). It is possible in principle to measure both D020,w and s020,w simultaneously from analysis of the boundary shape in sedimentation velocity analytical ultracentrifugation, although in practical terms this needs data of very high quality and the absence of any effects of sample heterogeneity. Equation 7.8.10 also of course provides a route for measurement of D020,w if s020,w and M are known. A simpler method for M measurement is possible if the conformation of the biomolecule is assumed and use is made of a power law relation, known as a Mark-Houwink relation (Equation 7.8.11): D = KM −ε Equation 7.8.11
where ε = 0.333, 0.85, or 0.5 to 0.6 for sphere-, rod-, and coil-shaped molecules, respectively. From a calibration plot of log D versus log M
The frictional ratio depends intrinsically on the conformation, flexibility, and degree of solvent association (water plus other salt ions and any other solvent molecules) of the biomolecule. This degree of water association is termed the “hydration” of the biomolecule, δ, and is defined as the mass (in g) of associated solvent per g of anhydrous biomolecule. This associated solvent includes both chemically bound solvent and solvent physically entrained in the interstices of the molecule. The value of δ is typically between 0.2 and 0.5 g/g for proteins, although it is a notoriously difficult parameter to pin down with any accuracy. The function defining the shape and flexibility of the biomolecule is the Perrin translational frictional function, P, illustrated by Equation 7.8.13:
δ P = ( f / f0 )1 + v ρ0
−
1 3
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where ρ0 is the density (g/ml) of the bound solvent. For a molecule that is fairly rigid on a time-averaged basis, the gross conformation can be specified using P in terms of the axial ratio of the equivalent hydrodynamic ellipsoid or in terms of sophisticated arrangements of spheres called hydrodynamic bead models. Computer programs are available for both types of modeling strategy (Harding et al., 1997; Garcia de la Torre et al., 1997)—although particularly with the latter, the diffusion coefficient should be used in conjunction with other hydrodynamic measurements to minimize uniqueness problems of the determined solution conformation.
Limitations of Dynamic Light Scattering Methods As with static light scattering, the main limitations here relate to sample clarification. The technique is most suited to larger proteins and protein assemblies (M >100,000 Da) where problems due to contamination are less, and, as with static light scattering, the problems become disproportionally higher at the lower scattering angles. If the protein systems are approximately globular, then to a good approximation these low angles can be avoided and measurements at a single higher angle (usually 90°) can suffice. For nonspheroidal systems, angular extrapolations are necessary to eliminate rotational diffusion contributions. The user should also be wary of interpreting data from fixed-angle photometers, which do not permit such angular measurements. The other limitations concern direct molecular-weight evaluation via Equation 7.8.11 and interpretation of the autocorrelation data in terms of polydispersity or size distribution. The user should be aware that such distributions are obtained purely by mathematical manipulation of the autocorrelation data, instead of involving a physical separation of different sizes (as with SEC-LALLS or SEC-MALLS), and the user is advised to be extremely cautious when interpreting such information produced by computer packages.
Samples for Analysis in Dynamic Light Scattering Aqueous solvents should be of sufficient ionic strength to suppress charge effects. The loading concentrations required—which, as with static light scattering, should be measured after clarification—will depend principally on the size of the scatterer and the output from the
laser. For example, if a 25-mW He-Ne laser is used, a loading concentration of at least ∼1 mg/ml (and a volume of 2 to 3 ml) is required for a large protein assembly whose molecular weight (M) is ∼5 × 106 Da. For proteins of molecular weight down to a lower limit of ∼10,000 Da, more powerful lasers (∼100 mW) and/or higher concentrations and/or longer experimental duration times are generally necessary to obtain meaningful results.
Temperature Control D is very sensitive to temperature, mainly because of the dependence of diffusion on the viscosity of the solvent. Temperature needs to be controlled or, at the very least, monitored accurately during the measurement, and a water bath is highly desirable.
Clarification of Solutions and Scattering Cells As with static light scattering, the scattering signal in dynamic light scattering is very sensitive to the presence of trace amounts of dust or supramolecular aggregation products; solutions and scattering vessels (called scattering cells or cuvettes) need to be scrupulously clean and free of particulates. Appropriate filtration, centrifugation of solutions, and washing of vessels with solvent is necessary. Special in-house cell-filling devices can be used along the lines described by Sanders and Cannell (1980); these can also be used for static light scattering.
Choice of Scattering Cuvette/Cell The same criteria apply as with static light scattering (see Samples for Analysis in Static Light Scattering). The cell design depends on whether the photometer is a fixed-angle or multi-angle design.
Procedure for Making Scattering Measurements Using a Fixed-Angle Instrument with a Flow Cell The following steps describe the injection procedure for a fixed (90°) angle photometer with a 20-mW infrared (780 nm) semiconductor laser (Claes et al., 1992). 1. The user should be satisfied that the protein/protein assembly is approximately globular. If not, then a multiangle instrument should be used (see Procedure for Making Scattering Measurements Using a Multiangle Instrument with a Conventional Cell). 2. Switch on the instrument and allow the laser to warm up 5 to 10 min.
Light Scattering
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3. Inject pure ultrafiltered water or buffered solvent via a 0.1-µm filter to get the clean water count rate. 4. If the count rate is below the manufacturer’s threshold, check the instrument alignment or increase the protein concentration. 5. Inject the protein solution in the same way as the water or buffer, using the appropriate filter, and check that the count rate meets the criteria specified by the manufacturer. 6. The choice of sample time and experimental duration time is normally done automatically. 7. Use the instrument’s software to obtain the diffusion coefficient, and, where appropriate, the in-built calibration to directly obtain the approximate molecular weight. 8. Rinse and dry the flow cell of the photometer.
rinsing with detergent and mild acetic acid), dry with ultrafiltered air, and dry the cuvette.
Procedure for Making Scattering Measurements Using a Multiangle Instrument with a Conventional Cell
The mobility, U = V/E, can thus be defined and hence the zeta potential, ζ:
1. Choose the appropriate cuvette or cell, using the same criteria (square versus cylindrical) as discussed above for static light scattering (see Samples for Analysis in Static Light Scattering). 2. Ensure that the cell is clear and free from dust and perform a check with pure ultrafiltered water, which should give only a negligibly small number of photon counts. Visually check for any “sparkling” in the laser beam passing through the water. 3. Remove the water from the cuvette, dry with ultrafiltered air, and inject the sample solution using the appropriate filter. 4. Set the goniometer for a scattering angle of 90°. 5. The choice of sample time and experimental duration time is normally done automatically. 6. Use the instrument’s software to obtain the apparent diffusion coefficient, Dapp. Inspect the autocorrelation decay plots (see Fig. 7.8.7). Nonlinearity can be due to sample polydispersity, particle asymmetry (effect of rotational diffusion phenomena), or particle settling (usually only observed for cellular systems, not for macromolecules). If it is not reasonable to assume an approximately globular system, then measure Dapp at a series of angles and extrapolate to zero angle, but be aware of enhanced supramolecular contamination problems at the lower angles. 7. Remove the solution, rinse the cell with ultrafiltered pure water (after, if necessary, prior
Electrophoretic Light Scattering Electrophoretic light scattering (ELS) uses dynamic light scattering to measure the velocity of migration, V, of a protein or other macromolecular system under the influence of an electric field, E (Langley, 1992; McNeil-Watson and Parker, 1992) The velocity V is related to the Doppler broadening, ∆ν, of the frequency of the incident laser radiation due to the velocity of the macromolecule. ∆v is related to V by Equation 7.8.14.
2 nV θ ∆ν = sin λ0 2 Equation 7.8.14
U = εζ / η Equation 7.8.15
This procedure is however used more for the investigation of the stability of colloids rather than protein structure and so will not be discussed further here.
LITERATURE CITED Bahls, D.M. and Bloomfield, V.A. 1977. Turbidimetric determination of bacteriophage molecular weights. Biopolymers 16:2797-2799. Brown, W. (ed.) 1993. Dynamic Light Scattering: The Method and Some Applications. Clarendon Press, Oxford. Burchard, W. 1992. Static and dynamic light scattering approaches to structure determination of biopolymers. In Laser Light Scattering in Biochemistry (S.E. Harding, D.B. Sattelle, and V.A. Bloomfield, eds.) pp. 3-22. Royal Society of Chemistry, London. Claes, P., Dunford, M., Kenney, A., and Vardy, P. 1992. An on-line dynamic light scattering instrument for macromolecular characterisation. In Laser Light Scattering in Biochemistry (S.E. Harding, D.B. Sattelle, and V.A. Bloomfield, eds.) pp. 66-76. Royal Society of Chemistry, London. Garcia de la Torre, J., Carrasco, B., and Harding, S.E. 1997. SOLPRO: Theory and computer program for the prediction of SOLution PROperties of rigid macromolecules and bioparticles. Evr. Biophys. J. 25:361-372. Harding, S.E. 1997. Microbial laser light scattering. Biotechnol. Genet. Eng. Rev. 14:145-164.
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Harding, S.E., Horton, J.C., and Colfen, H. 1997. The ELLIPS suite of macromolecular conformation algorithms. Eur. Biophys. J. 25:347-359. Johnsen, R.M. and Brown, W. 1992. An overview of current methods of analysing QLS data. In Laser Light Scattering in Biochemistry (S.E. Harding, D.B. Sattelle, and V.A. Bloomfield, eds.) pp 7791. Royal Society of Chemistry, London. Johnson, P. 1993. Light scattering in the study of colloidal and macromolecular systems. Int. Rev. Phys. Chem. 12:61-87. Johnson, P. and McKenzie, G.H. 1977. A laser light scattering study of haemoglobin systems. Proc. R. Soc. Lond. B. 199:263-278. Johnson, P. and Ottewill, R.H. 1954. A light scattering study of diptheria toxin-antitoxin interaction. Disc. Faraday Soc. 18:327-337. Jumel, K., Browne, P., and Kennedy, J.F. 1992. The use of low angle laser light scattering with gel permeation chromatography for the molecular weight determination of biomolecules. In Laser Light Scattering in Biochemistry (S.E. Harding, D.B. Sattelle, and V.A. Bloomfield, eds.) pp. 23-34. Royal Society of Chemistry, London. Jumel, K., Fogg, F.J.J., Hutton, D.A., Pearson, J.P., Allen, A., and Harding, S.E. 1997. A polydisperse linear random coil model for the quaternary structure of pig colonic mucin. Eur. Biophys. J. 25:477-480. Langley, K.H. 1992. Developments in electrophoretic laser light scattering and some biochemical applications. In Laser Light Scattering in Biochemistry (S.E. Harding, D.B. Sattelle, and V.A. Bloomfield, eds.) pp. 151-160. Royal Society of Chemistry, London. McNeil-Watson, F.K. and Parker, A. 1992. New tools for biochemists: Combined laser Doppler microelectrophoresis and photon correlation spectroscopy. In Laser Light Scattering in Biochemistry (S.E. Harding, D.B. Sattelle, and V.A. Bloomfield, eds.), pp. 59-65. Royal Society of Chemistry, London.
Sanders, A.H. and Cannell, D.S. 1980. Techniques for light scattering from hemoglobin. In Light Scattering in Liquids and Macromolecular Solutions (V. Degiorgio, M. Corti, and M. Giglio, eds.) pp. 173-182. Plenum, New York. Stacey, K.A. 1956. Light Scattering in Physical Chemistry. Butterworths, London. Svedberg, T. 1927. Nobelvortrag gehalten zu Stockholm am 19. Mai 1927. Kolloid-chem. Beih. 26:230-244. Tanford, C. 1961. Physical Chemistry of Macromolecules. Chapters 4 and 5. John Wiley & Sons, New York. Wells, C., Molina-Garcia, A.D., Harding, S.E., and Rowe, A.J. 1990. Self-interaction of dynein from Tetrahyamena cilia. J. Muscle Res. Cell Motil. 11:344-350. Wyatt, P.J. 1992. Combined differential light scattering with various liquid chromatography separation techniques. In Laser Light Scattering in Biochemistry (S.E. Harding, D.B. Sattelle, V.A. Bloomfield, V.A. eds.) pp. 35-58. Royal Society of Chemistry, London. Yau, W.W., Kirkland, J.J., and Bly, D.D. 1979. Modern Size Exclusion Chromatography. John Wiley & Sons, New York. Zimm, B.H. 1948. J. Chem. Phys. 16:1093-1099.
KEY REFERENCES Harding S.E., Sattelle, D.B. and Bloomfield, V.A. (eds.) 1992. Laser Light Scattering in Biochemistry. Royal Society of Chemistry, London. Huglin, M.B. (ed.) 1972. Light Scattering from Polymer Solutions. Academic Press, London.
Contributed by Stephen E. Harding and Kornelia Jumel University of Nottingham School of Biology Sutton Bonington, United Kingdom.
Pusey, P.N. 1974. Correlation and light beating spectroscopy In Photon Correlation and Light-Beating Spectroscopy (H.Z. Cummings and E.R. Pike, eds.) p. 387. Plenum, New York.
Light Scattering
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Current Protocols in Protein Science
Measuring Protein Thermostability by Differential Scanning Calorimetry
UNIT 7.9
Every chemical reaction is accompanied by heat effects that, at constant pressure, are described by the enthalpy of the process. Protein folding/unfolding, being an isomerization reaction between different macroscopic states, is no exception (Sturtevant, 1972; Privalov and Khechinashvili, 1974; Krishinan and Brandts, 1978). Temperature-induced changes in the enthalpy of the macroscopic states of a protein are of particular significance. Temperature and enthalpy are coupled extensive and intensive thermodynamic parameters. Thus, the functional relationship between enthalpy and temperature, f = H(T), includes all thermodynamic information about the macroscopic states in the system. This information can be extracted using the formalism of equilibrium thermodynamics. The H(T) function of a system can be determined experimentally from calorimetric measurements of its heat capacity at constant pressure Cp(T), over a broad temperature range: C p (T ) =
I
∂H and thus H (T ) = T Cp (T )dT + H (T0 ) ∂T p T 0
Equation 7.9.1
The heat capacity function, Cp(T), of a system can be directly measured by differential scanning calorimetry (DSC), making DSC the method of choice to study the conformational stability of biological macromolecules and proteins in particular. This unit presents step-by-step protocols for DSC, including sample preparation and interpretation of the results in the simplest cases (see Basic Protocol) as well as calibration of the apparatus (see Support Protocols 1, 2, and 3) and maintenance of DSC cells (see Support Protocol 4). Various general experimental considerations, including possible pitfalls and errors, are discussed in the Commentary (see Critical Parameters; see Troubleshooting). STRATEGIC PLANNING To study biological macromolecules using DSC instrumentation, a number of requirements must be satisfied: the measurements must be performed (1) in a physiological milieu, i.e., aqueous solution; (2) on protein samples that are dilute—i.e., at concentrations as low as 1 mg/ml (0.1%)—to minimize intramolecular interactions; and (3) in a small volume, since isolating and purifying protein samples is costly (in most cases only a few milligrams of protein are available, which at 1 mg/ml is only a few milliliters). Under such conditions the partial heat capacity of a protein in aqueous solution is on the order of 0.03% to 0.5%, and this is measured on the background of >99.5% heat capacity of solvent. Such measurements require extreme sensitivity, reproducibility, and signal stability from the DSC instrumentation. This implies that particular care should be taken with the experimental procedures for sample preparation and routine DSC operation. Currently two companies manufacture DSC instruments capable of working with solutions of biological macromolecules (Calorimetric Science and Microcal; see SUPPLIERS APPENDIX). These instruments are fully automated for control and for data collection, handling, and analysis using PC computers. The general principles of DSC instrument operation are outlined by Privalov et al. (1995) and Plotnikov et al. (1997).
Contributed by George I. Makhatadze Current Protocols in Protein Science (1998) 7.9.1-7.9.14 Copyright © 1998 by John Wiley & Sons, Inc.
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In performing a DSC experiment, it is important to keep in mind that DSC operates in differential mode. This means that the heat capacity of the protein in aqueous solution is measured relative to the heat capacity of the surrounding buffer. In an ideal situation, when the heat capacities and volumes of both sample and reference cells are precisely the same, a single protein/buffer scan will suffice. However, in reality, sample and reference cells are not identical and this difference has to be taken into consideration. Experimentally this requires the use of an internal instrument standard—the buffer/buffer scan, or baseline of the instrument. Before starting the protein scan it is very important to establish the stability of the baseline, i.e., its relative position and shape. Most contemporary calorimeters have very high baseline stability and reproducibility. Before starting to prepare the protein it is always a good idea to take several overnight buffer/buffer baselines and compare these to make sure that they are reproducible in accordance with previous observations. If this criterion is satisfied, one may start preparing the protein sample (see Critical Parameters for crucial steps in sample preparation). When setting the DSC control program, one of the most important parameters is the heating rate. Several considerations have to be taken into account. First, the higher the scan rate, the higher the sensitivity of the instrument. The increase in sensitivity is linear with respect to the heating rate: i.e., sensitivity at a heating rate of 2°/min is twice as high as at 1°/min. One needs to keep in mind that the increase in sensitivity does not mean an increase in the signal-to-noise ratio. Second, if the expected transition is very sharp—i.e., occurs within a few degrees—a high heating rate will affect the shape of the heat absorption profile and lead to an error in the determination of all thermodynamic parameters for this transition. This error will be particularly large for the transition temperature. Third, the higher the heating rate, the less time the system has to equilibrate. So for slow unfolding/refolding processes it is advisable to use low heating rates. In practice, for small globular proteins exhibiting reversible temperature-induced unfolding, the maximal heating rate (on most DSC instruments) of 2°/min is acceptable. For large proteins lower heating rates (0.5° to 1°/min) are more appropriate. In the case of fibrillar proteins, which usually exhibit very narrow transitions, a heating rate of 0.1° to 0.25°/min is most suitable. BASIC PROTOCOL
DIFFERENTIAL SCANNING CALORIMETRY Before beginning calorimetry, an instrument baseline is established by a buffer/buffer scan. If there are changes in the shape and/or relative position of the buffer/buffer scan, the cells are probably dirty (see Support Protocol 4 for cleaning procedure). Materials Protein solution, dialyzed (UNIT 4.4 & APPENDIX 3.B) and in dialysis bag (see Critical Parameters) Equilibration buffer (buffer solution used for the last dialysis; see Critical Parameters for discussion of buffer selection)
Measuring Protein Thermostability by Differential Scanning Calorimetry
Pasteur pipet or automatic pipettor Spectrophotometer and quartz cuvettes of appropriate optical path length (depending on the expected protein concentration) 2-ml all-glass syringe (e.g., Becton Dickinson 2-cc Yale syringe) Precut needle supplied with the DSC instrument Differential scanning calorimeter (DSC), calibrated (see Support Protocols 1, 2, and 3)
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1. Turn on the spectrophotometer. This step is required only if a spectrophotometric method of determining protein concentration is used. See Critical Parameters for discussion of appropriate methods for determining protein concentration, which is crucial to obtaining accurate DSC data.
2. Transfer the protein solution from the dialysis bag into a 1.5-ml microcentrifuge tube using a clean Pasteur pipet or an automatic pipettor, and centrifuge 15 to 20 min at ∼12,000 to 13,000 × g, 4°C. This step removes insoluble material.
3. While the protein is centrifuging, fill two quartz spectrophotometer cuvettes with equilibration buffer and take a buffer/buffer spectrum. This step establishes the buffer-buffer baseline of the spectrophotometer, which is particularly important with buffers containing absorbing components (e.g., dithiothreitol).
4. Empty both calorimetric cells, and rinse an all-glass syringe and attached needle (precut needle supplied with the DSC instrument) at least five to seven times with equilibration buffer. 5. Rinse one calorimetric cell (the reference cell) as follows: fill up the syringe, pump out all air bubbles, and insert the needle into the cell, letting the plunger slide down under its own weight (needles are precut to a specific length so that the tip of the needle will be barely above the bottom of the cell). Once the buffer appears in the overflow reservoir, start abruptly pumping buffer in and out of the cell, for a total of at least eleven to thirteen syringe volumes. Then fill the cell with more buffer. The abrupt pumping will force trapped air bubbles out of the cell. Calorimetric cells can be refilled using disposable plastic syringes as well (e.g., in the case of Ca2+-binding proteins, where it is desirable to avoid glass syringes since they will “leak” calcium). In this case the operator should slowly inject solution into the cell until the solution appears in the overflow reservoir, and then briskly pump the solution in and out of the cell. During the last step, the solvent level should never get below the level of the overflow reservoir.
6. Repeat rinsing procedure with the second calorimetric cell (sample cell) and leave it empty. Remove all buffer leftovers from the overflow reservoir. 7. Stop the centrifuge. Using the syringe, transfer the supernatant of the protein solution into a spectrophotometric cuvette and record the protein spectrum. It is advisable not to try and remove all of the supernatant, but to leave last 50 to 70 ìl in the microcentrifuge tube.
8. Transfer the protein solution from the spectrophotometric cuvette into the sample calorimetric cell. Follow the same technique as for filling the reference cell (see step 5).
9. Apply pressure, maintaining at least 1.5 atm (∼22 psi) in the cell. All calorimetric experiments should be performed under excess pressure (see Critical Parameters).
10. Select scan parameters and start the DSC control program. See Strategic Planning for discussion of control parameters; see Anticipated Results for information on data analysis procedures. Characterization of Recombinant Proteins
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CALIBRATION OF THE DSC INSTRUMENT DSC instruments are precalibrated by their manufacturers for temperature scale and heat capacity. It is unlikely that the characteristics of an instrument will change, but it is nonetheless advisable to check them once or twice a year. Two other calibrations should be done in the lab: electrical calibration (y axis of the instrument readout) using the system built into the instrument (see Support Protocol 1), and temperature calibration (x axis) using either hydrocarbon capsules obtained from the manufacturer (see Support Protocol 2) or lipid suspensions that can be prepared in the laboratory (see Support Protocol 3). SUPPORT PROTOCOL 1
Electrical Calibration
SUPPORT PROTOCOL 2
Temperature Calibration Using Hydrocarbon Capsules
SUPPORT PROTOCOL 3
Temperature Calibration Using Lipid Suspensions
All DSC instruments have a built-in capability to perform heat capacity calibration using an electrical pulse. Electrical calibration is performed using a special built-in circuit with a precisely measured resistance. A precise electrical current applied to this circuit generates a defined amount of power, δW, supplied to one of the cells. Exactly the same amount of power will be supplied to the other cell by the cell heaters in order to compensate for the inequality of temperature between the sample and reference cells. The power generated by the heaters of the instrument is related to the apparent heat capacity, δCp; the y-axis value generated by the instrument is δCp = δW/(dT/dt), where dT/dt is the heating rate. Follow the manufacturer’s suggested procedures for electrical calibration.
Temperature calibration is performed by melting pure hydrocarbons of known melting temperatures. The instrument manufacturers supply capsules containing hydrocarbons that can be inserted into a DSC cell filled with water (follow the manufacturer’s suggested procedure). Alternatively, aqueous suspensions of certain lipids with known melting temperatures can be used (see Support Protocol 3).
The following procedure for preparing lipid suspensions was suggested by Mabrey and Sturtevant (1976). Additional Materials (also see Basic Protocol) Lipid: dipalmitoylphosphatidylcholine (DPPC) or distearoylphosphatidylcholine (DSPC) 10 mM sodium phosphate buffer, pH 7.0 (APPENDIX 2E) Vortex mixer Water bath, 60°C 1. Add the lipid (DPPC or DSPC) to 10 mM sodium phosphate buffer, pH 7.0, to a 0.5 to 1.0 mg/ml final lipid concentration. 2. Heat the suspension at 60°C for several minutes, then vortex vigorously. 3. Fill the reference calorimetric cell with 10 mM sodium phosphate buffer; fill the sample cell with the lipid suspension. 4. Set the heating rate of the calorimeter at 0.1° to 0.25°C/min.
Measuring Protein Thermostability by Differential Scanning Calorimetry
If it is possible to set the response time of the instrument, use the fastest available.
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5. Start the scan. The DPPC suspension has two transitions, a low-temperature, broad transition at 35.3°C and a high-temperature, sharp transition at 41.4°C. For the DSPC suspension the low-temperature transition is at 51.5°C and the high-temperature transition is at 54.9°C.
MAINTENANCE AND CLEANING OF DSC CELLS DSC cells that become clogged by protein aggregates must be cleaned carefully in order to provide a reproducible baseline and thus accurate results. Proper use of calorimeters (i.e., avoidance of aggregating systems) minimizes the need for this sometimes very tedious procedure. Depending on the composition of the calorimetric cells (gold, platinum, titanium, tungsten), the chemical resistance of the material and the degree of contamination, a number of different procedures can be recommended.
SUPPORT PROTOCOL 4
Additional Materials (also see Basic Protocol) 1 M HCl or 1 M NaOH Distilled water Concentrated formic acid 95% (190-proof) ethanol 2% (w/v) SDS, or other detergent Concentrated nitric acid 1. Rinse calorimetric sample cell with one syringe volume of 1 M HCl or 1 M NaOH. Rinse the cells well with water (∼20 to 30 syringes-full; see Basic Protocol, step 5). Fill both cells and record a water-water scan. If the baseline does not return to its original position, proceed with step 2. The choice between acid and base depends on the properties of the protein aggregates formed. This can be checked by titrating the aggregated protein sample removed from the calorimeter with acid or base (half and half). IMPORTANT NOTE: Do not use NaOH with cells made out of tungsten or titanium, as they will corrode.
2. Fill the sample cell with formic acid and let it sit for ∼10 to 15 min at room temperature. Rinse the cells well with water (∼20 to 30 syringes-full), fill both cells, and record a water-water scan. If the baseline does not return to its original position, proceed with step 3. 3. Fill both cells with formic acid and scan several times up to 70°C. Rinse the cells well with water (∼20 to 30 syringes-full), fill both cells, and record a water-water scan. If the baseline does not return to its original position, proceed with step 4. 4. Fill both cells with 95% ethanol and scan several times to 70°C. Rinse the cells well (∼20 to 30 syringes-full of water), fill both cells, and record a water-water scan. If the baseline does not return to its original position, proceed with step 5. 5. Fill both cells with detergent and scan several times to the maximal temperature allowed by the instrument. Rinse very well (it is quite difficult to rid cells of detergent, but ∼40 to 50 syringes-full of water per cell usually suffice). Fill both cells and record a water-water scan. If the baseline does not return to its original position, proceed with step 6. 6. Fill both cells with nitric acid and scan to 70°C several times. Rinse well (∼20 to 30 syringes-full), fill both cells with water, and record a water-water scan. Particular care should be taken to prevent exposing the pan or pressure sensor (in some designs) to the nitric acid fumes, which will cause rusting. Put wet cotton wool or a Kimwipe on top of the cell outlets and pressure-sensor hole.
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If none of these treatments helps, a possible alternative reason for changes in the shape and position of the baseline is changes in the intrinsic thermal properties of the cells that can occur over time. Contact the instrument manufacturer for advice.
COMMENTARY Background Information The major advantage of using DSC to study protein stability compared to other methods for monitoring temperature-induced unfolding of proteins is that it provides direct information about the modes of unfolding. This technique is unique for establishing whether the unfolding process is a two-state or a multi-state process. It also provides direct measurement of the energetics of the unfolding transition. Another important parameter directly measured by DSC is the heat capacity change upon unfolding, which can be used to compute the temperature dependencies of enthalpy, entropy, and Gibbs energy functions. Until recently, the major drawback to DSC was the requirement for relatively large amounts of a protein sample. Contemporary DSC instruments have overcome this and allow measurements at spectroscopic concentrations (0.1 mg/ml).
Critical Parameters Protein purity The purity of the sample should be maintained very scrupulously. Traditional electrophoresis (native, UNIT 10.1, and SDS-PAGE, UNIT 10.3) as well as emerging, more sensitive methods (MALDI-TOF or ion-spray mass spectroscopies, UNIT 16.2) should be used to check sample purity. Impurities can affect both the shape of the calorimetric profiles and the estimation of the thermodynamic parameters of unfolding through errors in determination of the sample concentration (see below).
Measuring Protein Thermostability by Differential Scanning Calorimetry
Buffer solutions Careful attention should be paid to the buffer composition. Two factors are particularly important. 1. Since calorimetric experiments are performed over a broad temperature range, the pH of the buffer should have a low temperature dependence. For this reason the very popular “physiological” buffer Tris is not the best choice. Glycine, sodium or potassium acetate, sodium or potassium phosphate, sodium cacodylate, and sodium citrate are all suitable buffers. See Izatt and Christensen (1976) for tabulated data on the physicochemical properties of various buffers.
2. Protein unfolding is usually accompanied by changes in the ionization state of certain moieties within the protein (i.e., amino acid functional groups). The heat of ionization can be very large (e.g., 30 to 44 kJ/mol for histidine, depending on pH) and can obscure the “true” conformational enthalpy of unfolding. In some cases this problem can be avoided by selecting buffers with heats of ionization close to the heats of ionization of the protein groups. In this case the heat of proton release/absorption for the protein will be compensated by the heat of proton absorption/release for the buffer. This can be achieved in the acidic pH ranges of 2.0 to 3.2 (with the glycine/HCl buffer system) and 3.5 to 5.0 (with the sodium acetate/acetic acid buffer system). In this overall pH range (2.0 to 5.0), the carboxyl groups of aspartic and glutamic acid are titrated, and their heat of ionization will be counterbalanced by the heat of ionization of the carboxyl groups of the buffer. In neutral and alkaline pH ranges, such compensation is not possible and corrections for the effects of functional group ionization should be made during data analysis (see Anticipated Results). Equilibration of the protein solution with the solvent Equilibration of the protein with buffer is a vital part of sample preparation for calorimetric experiments. One should keep in mind that what the DSC instrument measures is the difference in heat capacity between buffer and protein solutions. This means that any small variations in buffer composition will result in an incorrect absolute value and an incorrect slope of the heat capacity profile. Direct dissolution of a protein sample in the corresponding buffer gives an equimolar concentration of the low-molecular-weight component (the buffer). In the case of ionic compounds, this will lead to nonidentical buffer compositions, since the ionic solute—i.e., protein—will directly interact with the buffer components, thereby decreasing its bulk concentration. The best way to achieve equilibrium between buffer and protein solution is dialysis. The protein sample should be dialyzed against at least two changes of buffer (see UNIT 4.4). The appropriate dialysis time will vary depending
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on the molecular weight cutoff (MWCO) of the dialysis membrane: the lower the MWCO, the longer the dialysis should be. For membranes with MWCO of 3500, 6 hr per change are enough. Higher-MWCO membranes equilibrate somewhat faster. It is advisable to filter the buffer solutions prior to dialysis using a 0.45-µm filter to eliminate all insoluble material. Clarification of the solution After dialysis, all insoluble particles should be carefully removed from the protein solution. This can be achieved either by passing the protein solution through a 0.22- to 0.45-µm filter or by microcentrifuging the solution for 15 to 20 min at ∼12,000 × g. Centrifugation is preferable for several reasons: (1) quite often there is nonspecific adsorption of protein to the filtration membrane; (2) the filtration membrane can contain compounds that will be washed off into the protein solution, thus changing the buffer composition of the protein solution compared to the equilibration buffer; and (3) filtration membranes are costly. Concentration of the protein solution Knowledge of the exact protein concentration after dialysis against the corresponding buffer is another important prerequisite to DSC. Among all available methods for determining protein concentration, the spectrophotometric method seems to be most accurate and rapid. This method requires (1) a standard doublebeam UV/visible spectrophotometer with resolution of at least 0.002 absorption units, and (2) knowledge of the extinction coefficient of the protein at a given wavelength. The extinction coefficient can be determined experimentally by determination of the nitrogen content (Jaenicke, 1974) or by quantitative amino acid analysis (UNIT 3.2). Alternatively, if the protein of interest contains at least one tyrosine or tryptophan residue or disulfide bond, the extinction coefficient can be calculated from the number of aromatic residues and disulfide bonds using an empirical equation (Gill and von Hippel, 1989): 0.1%,1 cm
ε 280 nm
=
5690 NTrp + 1280 NTyr + 120 NSS MW Equation 7.9.2
where NTrp, NTyr, and NSS are the number of tryptophans, tyrosines, and disulfide bonds, respectively, and MW is the molecular weight of
the protein in daltons. A more elaborate procedure, described by Pace et al. (1995), improves the accuracy of this calculation. For proteins that do not contain any aromatic residues, the method of Scopes (1974), which also uses spectrophotometry, gives fast and accurate results (see UNIT 3.1). Light scattering in spectrophotometric measurements of protein concentration can be a significant problem. This can be taken into account using a procedure suggested by Winder and Gent (1971). Pressure in the calorimetry cell All calorimetric experiments are performed under excess pressure. There are two main reasons for this. (1) The solubility of gases in aqueous solution is temperature dependent— thus, the lower the temperature, the higher the solubility. Excess pressure prevents air bubble formation in the cell at elevated temperatures. In some cases this can also be achieved by degassing protein solutions by exposing them to vacuum. However, degassing inevitably leads to a change in the solvent composition due to evaporation, which will in turn introduce a distortion in the calorimetric profile. (2) Excess pressure keeps sample evaporation from the calorimetric cell to a minimum. It is important to obtain heat capacity measurements for as broad temperature range as possible; for some proteins, the unfolding transition is not complete until the temperature is above the boiling point of water. Excess pressure will elevate the boiling temperature, and allow measurements at temperatures above the boiling point of water (100°C) in these cases. It is recommended that the pressure in the cell be kept to at least 1.5 atm (∼22 psi) in all experiments.
Troubleshooting The most frequent problem in DSC experiments is irreversibility of the unfolding transition. This applies particularly to large proteins that tend to aggregate after unfolding. The sensitivity of contemporary DSC instruments provides reliable heat capacity profiles at concentrations as low as 0.1 mg/ml for proteins 6 to 17 kDa in size, and the concentration can be even lower for larger proteins. Low protein concentration decreases the probability of aggregation. However, even at low protein concentrations, finding the proper solvent conditions to prevent aggregation is not a simple task. A rule of thumb is that at a pH value close to the isoelectric point of the protein, aggregation
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after unfolding is highly probable. High concentrations of neutral salts also promote aggregation. In many cases, aggregation can be avoided by keeping the ionic strength of the solvent very low (10 to 50 mM) and by using pH values a couple of units away from (usually below) the isoelectric point.
Anticipated Results Calculation of the partial heat capacity profile The first step in calculating the partial heat capacity profile is subtraction of the scan rate– normalized baseline (buffer/buffer scan) from the scan rate–normalized protein scan (protein/buffer) to obtain ∆Capp p (T), the heat capacity difference between sample and reference cells at temperature T. Knowledge of ∆Capp p (T) allows calculation of the partial heat capacity of the protein at temperature T, Cexp p,pr (T), as: C pexp , pr ( T ) = C p , buf ( T ) ⋅
Vpr ( T ) Vbuf ( T )
−
∆C app p (T ) m pr ( T )
Equation 7.9.3
where Cp,buf(T) _is _ the heat capacity of buffer at temperature T, Vpr (T) is the partial __ volume of the protein at temperature T, Vbuf (T) is the partial volume of buffer at temperature T, and mpr(T) is the mass of the protein in the calorimetric cell at temperature T. The mass of the protein in the cell is calculated as mpr(T) = vcell(T)⋅Wpr(T), where vcell(T) is the volume of the cell at temperature T and wpr(T) is the concentration of the protein at temperature T. In practice, however, for aqueous buffers C p,buf Vbuf
≅
C p, H O 2 VH O 2
Equation 7.9.4
and mpr(T) ≅ constant, because ∂vcell (T ) ∂T
≅
∂wpr (T ) ∂T
Equation 7.9.5
so that Equation 7.9.3 is simplified to: Measuring Protein Thermostability by Differential Scanning Calorimetry
C pexp , pr ( T )
C p, H O ∆C papp (T ) 2 = ⋅ Vpr − VH 2 O mpr Equation 7.9.6
where only one term, ∆Capp p (T), is temperature dependent. In any case, both equations require knowledge of the partial volume of the protein, _ _ Vpr. This parameter can be calculated from the amino acid composition of the protein using an additivity scheme described by Makhatadze et al. (1990). Calculations can also be performed via the World Wide Web at the URL http:// www.ttu.edu/~chem/makhatadze/Vpartial.html. Interpretation of DSC profiles Two important pieces of information should be established prior to the analysis of the calorimetric profiles. These will help to decide whether the formalism of equilibrium thermodynamics is applicable to the protein being studied. 1. Reversibility of the unfolding transition should be examined under several conditions by rescanning the sample. Reversibility depends on the upper temperature of heating. The unfolding transition can be considered reversible if rescanning after the sample has been heated to the temperature just past the completion of the transition leads to the recovery of at least 85% to 90% of the initial endotherm. If no endotherm is observed after rescanning, the unfolding reaction is irreversible. Thermodynamic analysis for irreversible transitions is possible only in special cases (Sanchez-Ruiz, 1992). 2. The equilibrium conditions of reversible unfolding should be examined to find the optimal scan rate that permits establishment of equilibrium between native and unfolded states at all temperatures—i.e., the scan rate should not be faster than the folding/unfolding rates. This can be accomplished by comparing the heat capacity profiles obtained on the same exact sample at two different heating rates, usually 2°/min and 0.5°/min. If the difference in the transition temperature exceeds 0.5° to 0.7°C, additional experiments should be performed at a lower heating rate (i.e., 0.1°/min) and compared to the 0.5°/min experiment. If there is still a difference in the transition temperature, all experiments at a given solvent composition should be performed at several different heating rates and the transition temperatures should be extrapolated to a zero heating rate (for more details see Yu et al., 1994). NOTE: Sensitivity of DSC is reversibly proportional to the heating rate, so these experiments should be performed at protein concentrations high enough to obtain the transition temperature with a relatively high accuracy.
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30 Heat capacity (kJ K – 1 mol – 1)
Tmax > Tm 25 Cp,U > (T)
20 ∆Cp (T )
C exp
15
p, pr
< Cp (T ) > prg Cp,N (T ) Tm
10 < Cp (Tm ) > max
5
∆Hexp (Tm )
< Cp (T ) > exc 0 10
20
30
40
50
60
70
80
90
Temperature (oC)
Figure 7.9.1 The relation of the partial molar heat capacity, Cpexp ,pr (T), to the excess heat capacity, exc, progress heat capacity, Cprg p (T), and partial heat capacities of the native, Cp,N(T), and unfolded, Cp,U(T), states.
By visual inspection of the calorimetric profiles one can establish whether the unfolding transition represents a single endotherm (as shown in Fig. 7.9.1) or is more complex. These two cases are analyzed differently. If the DSC profile is a single endotherm Heat capacity functions for the native and unfolded states. The partial specific heat capacity of the native state, Cp,N, ranges from 1.25 to 1.80 J/(K⋅g) at 25°C depending on the protein (Makhatadze, 1998). Cp,N has a linear dependence on temperature with a slope from 0.005 to 0.008 J/(K−2 g), also depending on the protein (Makhatadze, 1998). The partial specific heat capacity of the unfolded state, Cp,U, in the temperature range from 5° to 100°C is always higher than the heat capacity of the native state. At 25°C, Cp,U values for different proteins range from 1.85 to 2.2 J/(K⋅g), whereas at 100°C these values are higher, from 2.1 to 2.4 J/(K⋅g) (Makhatadze, 1998). The partial heat capacity of the unfolded state has a nonlinear dependence on temperature (e.g., Wintrode et al., 1994). It increases gradually (with a slope comparable to that for the native state) and approaches a constant value at 60° to 75°C. At temperatures above 60° to 75°C, Cp,U does not depend on temperature. As a result of the different temperature dependencies of the heat capacities of the native and unfolded states, the
heat capacity change upon protein unfolding, ∆Cp = Cp,U − Cp,N, appears to be a temperaturedependent function. Excess heat capacity profile. The so-called excess heat capacity function exc is the heat capacity caused exclusively by the heat absorbed due to the unfolding reaction. exc thus is expressed as a difference between the experimental, Cexp p (T), and progress, Cprg heat capacity functions of a T), ( p protein: < C p ( T ) > exc = C pexp ( T ) − C pprg ( T ) Equation 7.9.7
where the progress heat capacity is defined by the intrinsic (partial) heat capacity of the protein in the entire temperature range:
Cpprg (T ) = FN (T ) ⋅ Cp,N (T ) + FU (T ) ⋅ Cp,U (T ) Equation 7.9.8
where Cp,N(T) and Cp,N(T) are the temperature dependencies of the partial heat capacities of the native and unfolded states, respectively. Because of the difference in the partial heat capacities of the native and unfolded states, in the temperature range where transition occurs,
Characterization of Recombinant Proteins
7.9.9 Current Protocols in Protein Science
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progress heat capacity is not experimentally accessible. However, it can be calculated as a population-weighted sum of the partial heat capacities of the native and unfolded states. FN(T) and FU(T) can thus be defined as fractions of the protein in the initial (native) and final (unfolded) states, respectively, and consequently FN(T) + FU(T) = 1. Fractions of protein in the native or unfolded states are in turn calculated from the progress of the reaction as: FU (T ) = 1 − FN (T ) =
I I
Qi (T )
< C p (T ) > exc dT
0 ∞
< C p (T ) >
exc
dT
Equation 7.9.9
where Qtot is the total heat absorbed upon unfolding and Qi(T) is the heat absorbed at temperature T. Transition temperature, Tm. The transition temperature, Tm, is defined as the temperature at which the areas under the excess heat capacity profile on both sides of Tm are equal, i.e.:
I 0
I
∞
< C p ( T ) > exc dT =
< C p ( T ) > exc dT
Tm
Figure 7.9.10
Tm always differs from the temperature of maximum partial heat capacity, Tmax, due to the difference in the heat capacities between the native and unfolded states, ∆Cp = Cp,U − Cp,N. Tmax − Tm decreases with an increase in the relative ratio of the area under the excess heat capacity profile to ∆Cp. Calorimetric enthalpy, ∆Hexp (Tm). Calorimetric enthalpy is defined as the area under the excess heat capacity function, i.e.,
I
∞
∆Hexp (Tm ) = Qtot = < C p (T ) > exc dT 0
Equation 7.9.11 Measuring Protein Thermostability by Differential Scanning Calorimetry
Equation 7.9.12
The enthalpy of ionization arises from two different processes, ionization of protein groups and ionization of buffer; i.e.: ion ion ∆H ion ( Tm ) = ∆H pr ( Tm ) + ∆v ⋅ ∆H buf ( Tm )
Equation 7.9.13
0
Tm
∆Hexp ( Tm ) = ∆H cnf ( Tm ) + ∆H ion ( Tm )
Qtot
T
=
release protons that will be absorbed or released by the buffer. This process will be accompanied by heat effects and will be included in the experimentally measured calorimetric enthalpy, ∆Hcal(Tm). Thus ∆Hcal(Tm) consists of two parts, the conformational enthalpy of protein unfolding, ∆Hcnf(Tm), and the ionization effects, ∆Hion (Tm):
and is referred to as the enthalpy at the transition temperature. Enthalpy of the conformational transition, ∆Hcnf(Tm). When a protein unfolds, internal groups buried in the native state can bind or
where Hion pr (Tm) is the enthalpy of ionization of protein, Hion buf (Tm) is the enthalpy of ionization of buffer, and ν is the change in the protonation state of the protein (see below). When the enthalpies of ionization of protein groups and of the buffer are close, the calorimetric and conformational enthalpies are also close to each other, i.e, ∆Hcal (Tm) ≅ ∆Hcnf(Tm). Such a situation occurs in glycine or acetate buffer systems with an acidic pH. In the acidic pH range, the most probable titratable groups of the protein are the carboxylic acids (aspartic and glutamic acid). Their heats of ionization are very close to the heats of ionization for carboxylic groups of glycine or acetic acid. However, at neutral or alkaline pH, titratable groups (histidine, lysine, arginine, tyrosine, and cysteine) have heats of ionization very different from those of the buffer systems used (e.g., phosphate and cacodylate). In these cases, differences between calorimetric and conformational enthalpies can be significant (see, for example, Yu et al., 1994). Changes in the protonation state of the protein upon unfolding, ∆ν. Under certain solvent conditions protein unfolding is accompanied by a change in the protonation state of the protein, ∆ν. This difference in the number of protons bound to the unfolded, νU, and native, νN, states can be determined from the dependence of the transition temperature on the pH values of the solution dTm/dpH as: ∆ν = νU − ν N =
∆H exp ( Tm ) 2.303 ⋅ R ⋅ Tm2
⋅
dTm dpH
Equation 7.9.14
7.9.10 Supplement 12
Current Protocols in Protein Science
dTm =0 dpH
60 55
T m (oC)
50
dTm >0 dpH
dTm >0 dpH
45 40 35 30 25
2
3
4
5
6
7
8
9
10
pH
Figure 7.9.2 Dependence of the transition temperature on solution pH for a hypothetical protein.
Typically, the curve representing the dependence of the transition temperature on the pH of the solution has a bell shape (Fig. 7.9.2). On the left shoulder, dTm/dpH > 0 and thus ∆ν < 0—i.e., the protein releases protons upon unfolding. On the right shoulder of Tm dependence on pH, dTm/dpH < 0 and thus the protein absorbs protons upon unfolding (∆ν > 0). At the maximum of the Tm(pH) function, dTm/dpH = 0 and thus the protonation states of the native and unfolded states of a protein are identical (∆ν = 0). Van’t Hoff enthalpy, ∆HνH (Tm), and modes of unfolding transition. The so-called van’t Hoff enthalpy for a two-state unfolding process is determined by the temperature dependence of the equilibrium constant, Keq: ∆H vH (Tm ) = − R ⋅
d ln K eq d (1 / T )
= R⋅T2 ⋅
Equation 7.9.15
where Keq is defined as K eq ( T ) =
FN ( T ) FN ( T ) + FU ( T )
Equation 7.9.16
d ln K eq dT
From this definition, it follows that van’t Hoff enthalpy can be estimated from Equation 7.9.8 or Equation 7.9.9. However, for the calorimetric curves ∆HνH (Tm) can be estimated as: ∆H vH ( Tm ) =
4 ⋅ R ⋅ Tm2 ⋅ < C p ( Tm ) > max ∆H exp ( Tm )
Equation 7.9.17
where max is the maximum value of the excess heat capacity function. A direct estimate of two enthalpies—experimental calorimetric enthalpy, ∆Hexp (Tm), and van’t Hoff enthalpy, ∆HνH(Tm)—from a single DSC experiment provides a unique way to determine the modes of protein unfolding (Biltonen and Freire, 1978; Privalov and Potekhin, 1986; Kidokoro and Wada, 1987; Marky and Breslauer, 1987; Ghosaini et al., 1988). When the ratio β = ∆HvH(Tm)/∆Hcal(Tm) equals 1 (in practice, ∼0.95 to 1.05), this indicates that the observed transition is two-state, proceeding from the native to the unfolded state without a significant population of intermediates: i.e., the unfolding reaction can be represented by N ↔ U. Deviations from unity indicate that the transition is more complicated. There are two possible explanations of β deviating from unity for
Characterization of Recombinant Proteins
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Measuring Protein Thermostability by Differential Scanning Calorimetry
the reversible unfolding of the protein showing a single-isotherm DSC profile. 1. The reaction is not monomolecular. Information on whether or not the reaction is monomolecular can be obtained from the dependence of the transition temperature for the reversible reaction on the protein concentration. The possibility of oligomerization of the protein in the native or/and unfolded states should be examined by performing experiments under exactly the same solvent conditions using protein concentrations that differ by at least a factor of 5. The dependence of the transition temperature, Tm, on protein concentration, w, will be an indication of the mode of unfolding reaction:
of the heat capacity functions of the unfolded and native states at the transition temperature: ∆Cp = Cp,U(Tm) − Cp,N(Tm). An alternative, indirect method for the determination of ∆Cp uses a basic thermodynamic equation, ∆Cp = (∂∆H/∂T)p, which shows that the slope of the dependence of the conformational enthalpy of unfolding, ∆Hcnf(Tm), on the transition temperature, Tm, is equal to ∆Cp. Heat capacity change upon unfolding, ∆Cp, defines the temperature dependencies of the enthalpy
a. ∂Tm/∂w = 0: N ↔ U, unfolding reaction is monomolecular; b. ∂Tm/∂w > 0: N ↔ nU, unfolding reaction is of nth order and the native protein exists in an oligomeric state; c. ∂Tm/∂w < 0: N ↔ Un, unfolding reaction is of nth order and the unfolded protein exists in an oligomeric state.
Equation 7.9.18
Situations (a) and (b) are very common and the thermodynamic formalism for their analysis is well developed. Situation (c) is rare for reversible reactions, but does occur for irreversible reactions when irreversibility is caused by aggregation (usually nonspecific) in the unfolded state. 2. Intermediate states. An alternative explanation for β being >1 is the existence of intermediate states during equilibrium reversible unfolding: i.e., N ↔ I1 ↔ I2, ... In ↔ U. Consider the most common example of such an unfolding reaction, which is the unfolding of a two-domain protein. Both domains have similar intrinsic stability, ∆G1(T) ≈ ∆G2(T), and their interactions with one another are described by Gibbs free energy ∆G12(T). The ratio of calorimetric to van’t Hoff enthalpy, β, will depend on the strength of the interdomain interactions. If ∆G12(T) = 0, i.e., there is no interaction between domains, each domain will unfold independently of the other, and β = 2. If there is a strong interaction between domains, i.e., neither domain can exist in a folded state if the other domain is unfolded, unfolding will be very cooperative and β = 1. Heat capacity change upon unfolding, ∆Cp, and thermodynamic functions for protein unfolding: enthalpy, ∆H(T), entropy, ∆S(T), and Gibbs free energy, ∆G(T). One way of estimating the heat capacity change upon protein unfolding is directly from the difference
I
T
∆H (T ) = ∆H (T0 ) +
∆C p ⋅ dT
T0
and entropy functions,
I
T
∆S (T ) = ∆S(T0 ) +
∆C p ⋅ d ln T
T0
Equation 7.9.19
and thus the Gibbs free energy function: ∆G(T ) = − R ⋅ T ⋅ ln Keq = ∆H (T ) − T ⋅ ∆S( T ) Equation 7.9.20
The integration constants ∆H(To) and ∆H(To) in Equation 7.9.19 and Equation 7.9.20 have to be determined experimentally. The integration constant the enthalpy function can be set as ∆H(To) = ∆Hexp(Tm). The integration constant for the entropy function (Equation 7.9.19) for a two-state transition is determined as follows. If the transition is two-state—i.e., ∆Hexp(Tm) ≈ ∆HνH(Tm)—then at the transition temperature (Tm) the population of native and unfolded states is equal. This means that at Tm the equilibrium constant of the reaction Keq = FN/FU is equal to 1, and thus the Gibbs free energy difference between the native and unfolded states at Tm is equal to zero and ∆G(Tm) = ∆H(Tm) − Tm⋅∆S(Tm) = 0. Thus the entropy change upon protein unfolding at the transition temperature, ∆S(Tm), is equal to: ∆S ( Tm ) =
∆H ( Tm ) Tm
Equation 7.9.21
7.9.12 Supplement 12
Current Protocols in Protein Science
This equation provides a constant of integration for Equation 7.9.20, ∆S(Tm) = ∆S(To). A complete thermodynamic description of the system at any other temperature can then be calculated as follows:
I
T
∆H (T ) = ∆H (Tm ) +
∆C p ⋅ dT
Tm
I I
T
∆Cp T
Tm
=
∆H(Tm ) Tm
⋅ dT
T
+
∆Cp ⋅ d ln T
Tm
Equation 7.9.23
∆G(T ) = ∆H (Tm ) − T ⋅
I
T
+
I
∆H (Tm ) Tm
T
∆C p ⋅ dT − T ⋅
Tm
Gill, S.C. and von Hippel, P.H. 1989. Calculation of protein extinction coefficients from amino acid sequence data. Anal. Biochem. 182:319-326. Izatt, R.M. and Christensen, J.J. 1976. Heats of proton ionization, pK, and related thermodynamic quantities. In CRC Handbook of Biochemistry and Molecular Biology, Vol. 1: Physical and Chemical Data (G.D. Fasman, ed.) pp. 151-269. CRC Press, Cleveland.
Equation 7.9.22
∆S(T ) = ∆S(Tm ) +
Ghosaini, L.R., Brown, A.M., and Sturtevant, J.M. 1988. Scanning calorimetric study of the thermal unfolding of catabolite activator protein from Escherichia coli in the absence and presence of cyclic mononucleotides. Biochemistry 27:52575261.
∆C p ⋅ d ln T
Tm
Equation 7.9.24
where the heat capacity change upon protein unfolding, ∆Cp, is a function of temperature itself. In practice, however, if no extrapolations over a wide temperature range are necessary, as a first approximation a temperature-independent ∆Cp is a reasonable assumption.
Time Considerations Sample preparation (of which dialysis is the most time-consuming step) usually requires 12 to 16 hr. Consult UNIT 4.4 for possible ways of reducing dialysis time. Baseline data is usually collected in overnight experiments; a calorimetric experiment with a protein sample requires several hours, depending on the heating rate. However, optimization of experimental conditions (reversibility and equilibrium) when first working with an unfamiliar sample may take several days. Time requirements for data analysis vary in individual cases.
Literature Cited Biltonen, R.L. and Freire, E. 1978. Thermodynamic characterization of conformational states of biological macromolecules using differential scanning calorimetry. CRC Crit. Rev. Biochem. 5:85124.
Jaenicke, L. 1974. A rapid micromethod for the determination of nitrogen and phosphate in biological material. Anal. Biochem. 61:623-627. Kidokoro, S. and Wada, A. 1987. Determination of thermodynamic functions from scanning calorimetry data. Biopolymers 26:213-229. Krishinan, K.S. and Brandts, J.F. 1978. Scanning calorimetry. Methods Enzymol. 49:3-14. Mabrey, S. and Sturtevant, J.M. 1976. Investigation of phase transitions of lipids and lipid mixtures by sensitivity differential scanning calorimetry. Proc. Natl. Acad. Sci. U.S.A. 73:3862-3866. Makhatadze, G.I. 1998. Heat capacities of amino acids, peptides and proteins. Biophys. Chem. 71:1-26. Makhatadze, G.I., Medvedkin, V.N., and Privalov, P.L. 1990. Partial molar volumes of polypeptides and their constituent groups in aqueous solution over a broad temperature range. Biopolymers 30:1001-1010. Marky, L.A. and Breslauer, K.J. 1987. Calculating thermodynamic data for transitions of any molecularity from equilibrium melting curves. Biopolymers 26:1601-1620. Pace, C.N., Vajdos, F., Fee, L., Grimsley, G., and Gray, T. 1995. How to measure and predict the molar absorption coefficient of a protein. Protein Sci. 4:2411-2423. Plotnikov, V.V., Brandts, J.M., Lin, L.N., and Brandts, J.F. 1997. A new ultrasensitive scanning calorimeter. Anal. Biochem. 250:237-244. Privalov, G., Kavina, V., Freire, E., and Privalov, P.L. 1995. Precise scanning calorimeter for studying thermal properties of biological macromolecules in dilute solution. Anal. Biochem. 232:79-85. Privalov, P.L. and Potekhin, S.A. 1986. Scanning microcalorimetry in studying temperature-induced changes in proteins. Methods Enzymol. 131:4-51. Privalov, P.L. and Khechinashvili, N.N. 1974. A thermodynamic approach to the problem of stabilization of globular protein structure: A calorimetric study. J. Mol. Biol. 86:665-684. Sanchez-Ruiz, J.M. 1992. Theoretical analysis of Lumry-Eyring models in differential scanning calorimetry. Biophys. J. 61:921-935.
Characterization of Recombinant Proteins
7.9.13 Current Protocols in Protein Science
Supplement 12
Scopes, R.K. 1974. Measurement of protein by spectrophotometry at 205 nm. Anal. Biochem. 59:277-282. Sturtevant, J.M. 1972. Calorimetry. Methods Enzymol. 26:227-253. Winder, A.F. and Gent, W.L. 1971. Correction of light-scattering errors in spectrophotometric protein determinations. Biopolymers 10:12431251. Wintrode, P.L., Makhatadze, G.I., and Privalov, P.L. 1994. Thermodynamics of ubiquitin unfolding. Proteins 18:246-253. Yu, Y., Makhatadze, G.I., Pace, C.N., and Privalov, P.L. 1994. Energetics of ribonuclease T1 structure. Biochemistry 33:3312-3319.
Key References
Pace et al., 1995. See above. Detailed discussion on the calculation of protein extinction coefficients from amino acid compositions. Privalov, P.L. 1979. Stability of proteins: Small globular proteins. Adv. Protein Chem. 33:167241. Privalov, P.L. 1982. Stability of proteins: Proteins which do not present a single cooperative system. Adv. Protein. Chem. 35:1-104. Makhatadze, G.I. and Privalov, P.L. 1995. Energetics of protein structure. Adv. Protein. Chem. 47:307-425. Three reviews describing and assessing the determination and interpretation of protein stability by DSC.
Biltonen and Freire, 1978. See above.
Sanchez-Ruiz, 1992. See above.
Privalov and Potekhin, 1986. See above.
Detailed discussion of the analysis of irreversible denaturation processes.
Kidokoro and Wada, 1987. See above. Three references that discuss in detail the theoretical background for deconvolution of the heat capacity profile, each describing a separate approach to data analysis. Izatt and Christensen, 1976. See above.
Contributed by George I. Makhatadze Texas Tech University Lubbock, Texas
Basic reference containing tables of heats of ionizations and other thermodynamic information for different buffer systems.
Measuring Protein Thermostability by Differential Scanning Calorimetry
7.9.14 Supplement 12
Current Protocols in Protein Science
Characterizing Recombinant Proteins Using HPLC Gel Filtration and Mass Spectrometry
UNIT 7.10
It is essential to determine the identity, sequence integrity, purity, oligomeric state, and conformational integrity of a protein before structural or functional studies are carried out. This is true for proteins isolated from any source, but is particularly important for recombinant proteins, which are subject to several problems associated with their artificial overexpression in a heterologous host. These problems include expression in a denatured state, aggregation, mutations, incorrect translation (e.g., stop codon readthrough), incorrect proteolytic processing, and unwanted post-translational modifications. Often these problems cannot be detected using SDS-PAGE. Fortunately, most sources of heterogeneity in recombinant proteins can be identified by a preliminary analysis of the sample using a combination of analytical high-performance liquid chromatography/fast protein liquid chromatography (HPLC/FPLC) and matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS). Analytical gel filtration is used to detect and quantify contaminants in the sample (including protein aggregates), and can give an approximate measure of the protein size in solution (Stokes radius). Mass spectrometry is then used to determine the molecular weight of the protein to within a few daltons, which enables the detection and identification of incorrect sequence length and most chemical modifications of amino acid side chains. Apart from detecting unwanted modifications to the protein of interest, HPLC and mass spectrometry can be used to help characterize post-translational modifications such as glycosylation (see Chapter 12). In this unit, analytical HPLC gel filtration is used to characterize a purified recombinant protein sample in the Basic Protocol. A method for determining the Stokes radius of a protein using HPLC is described in Support Protocol 1. MALDI-MS is used to characterize a protein sample in Support Protocol 2. NOTE: Milli-Q-purified or equivalent quality water should be used for all buffers. STRATEGIC PLANNING Selecting Column Type and Size The first step in planning a size-exclusion fractionation is selecting the type of column to be used. Several factors need to be considered. Most importantly, the molecular weight of the recombinant proteins to be analyzed should fall within the fractionation range of the column, which is determined by the pore size of the particles. The fractionation range should be large enough to encompass the molecular weights of all species to be separated, and the protein of interest should be near the midpoint of the fractionation range. The fractionation range may be increased by placing two columns with different pore sizes in series. Further selection of the column depends upon several related factors: the required resolution, the sample volume, and the desired run time. For analytical gel filtration, the aim is to maximize resolution while minimizing run time. Resolution is affected by the particle size and size distribution, as well as the column length and the flow rate (for a detailed discussion, see UNITS 8.1 & 8.3, or Neue, 1997). Columns with small, rigid, well-defined particles (<15 µm diameter) show the best resolution while allowing fast flow rates and therefore short run times (12 to 60 min). Table 7.10.1 lists several common commercial prepacked columns that feature small particle sizes and good resolution. The disadvantage of these columns is that the small particle sizes lead to high back pressures, requiring an isocratic chromatography system that can operate at moderate pressure (FPLC; 200 to 600 psi) or preferably higher pressure (HPLC; >500 psi). Resolution may Contributed by Gillian E. Begg, Sandra L. Harper, and David W. Speicher Current Protocols in Protein Science (1999) 7.10.1-7.10.15 Copyright © 1999 by John Wiley & Sons, Inc.
Characterization of Recombinant Proteins
7.10.1 Supplement 16
Table 7.10.1
Common Prepacked Columns for HPLC/FPLC Size-Exclusion Chromatographya
Fractionation range (kDa)
Resinb
Particle size (µm)
Dimensions available (i.d. × length, mm)
Supplierc
5-150 10-500
S S
4 4
4.6 × 300 4.6 × 300
TH TH
5-150 10-500 20-10,000 5-100
S S S S
5 5 8 10
TH TH TH TH
TSK G3000SW
10-500
S
10
TSK G4000SW
20-7000
S
13
Bio-Sil SEC 125 Bio-Sil SEC 250 Bio-Sil SEC 400 Protein-Pak 60 Protein-Pak 125 Protein-Pak 200SW Protein-Pak 300SW Superose 12 HR 10/30 Superdex Peptide Superdex 75 HR Superdex 200 HR Superose 6 HR
5-100 10-300 20-1000 1-20 2-80 1-60 10-300 1-3000 0.1-7 3-70 10-600 5-5000
S S S S S S S CL A CL A/D CL A/D CL A/D CL A
5 5 5 10 10 10 10 10 13 13 13 13
7.8 × 300 7.8 × 300 7.8 × 300 7.5 × 300 7.5 × 600 7.5 × 300 7.5 × 600 7.5 × 300 7.5 × 600 7.8 × 300 7.8 × 300 7.8 × 300 7.8 × 300 7.8 × 300 8.0 × 300 8.0 × 300 10.0 × 300 10.0 × 300 10.0 × 300 10.0 × 300 10.0 × 300
Preparative TSK G2000SW
5-100
S
13
TSK G3000SW
10-500
S
13
TSK G4000SW
20-7000
S
17
TSK G2000SW
5-100
S
20
TSK G3000SW
10-500
S
20
Superdex 30 HiLoad
1-10
CL A/D
34
Superdex 75 HiLoad
3-70
CL A/D
34
Superdex 200 HiLoad
10-600
CL A/D
34
Column type Microanalytical TSK Super SW2000 TSK Super SW3000 Analytical TSK G2000SWXL TSK G3000SWXL TSK G4000SWXL TSK G2000SW
21.5 × 300 21.5 × 600 21.5 × 300 21.5 × 600 21.5 × 300 21.5 × 600 55.0 × 300 55.0 × 600 55.0 × 300 55.0 × 600 16.0 × 600 26.0 × 600 16.0 × 600 26.0 × 600 16.0 × 600 26.0 × 600
TH TH BR BR BR WA WA WA WA APB APB APB APB APB TH TH TH TH TH TH TH TH
aData reported by manufacturers. bAbbreviations: CL A, cross-linked agarose; CL A/D, cross-linked agarose/dextran; S, coated silica. cAbbreviations: APB, Amersham Pharmacia Biotech; BR, Bio-Rad; TH, TosoHaas; WA, Waters. Addresses and phone
numbers of suppliers are provided in the SUPPLIERS APPENDIX.
Characterizing Recombinant Proteins Using HPLC and MS
7.10.2 Supplement 16
Current Protocols in Protein Science
be further enhanced by increasing the length of the column; if a longer column is not available, two columns may be placed in series to achieve the same effect. Low-pressure gel filtration columns with softer, larger particles are generally less expensive and allow larger protein loads, but require lower flow rates, which markedly increases the run time for the separation while providing reduced resolution even at low flow rates. If it is necessary to scale up an analytical separation, the sample volume will determine the size of the column required—the larger the sample volume and amount of protein to be separated, the greater the column volume needed. One option is to increase the length of the column; for example, a 60-cm column has twice the sample volume capacity of a 30-cm column, with little loss of resolution. Increasing the diameter of the column will also increase the maximum load in proportion to the square of the increase in diameter. Table 7.10.1 lists prepacked columns for both preparative work (i.d. = 21.5 or 55 mm) and analytical work (i.d. ≤ 10 mm). Another factor to consider when selecting a column is the potential for interactions between the sample and the column. For example, silica-based columns are available in the smallest particle sizes and can be operated at higher pressures compared to agarose gels; however, some adsorption can occur due to electrostatic interactions with unmodified silanol groups. Usually these effects can be minimized by maintaining at least a physiological ionic strength. Stainless steel columns can withstand high back pressures, but also have drawbacks. For example, some samples might interact with the metal surfaces (especially stainless steel end frits) and common buffer components (e.g., halogens such as chloride) can react with incompletely passivated stainless steel surfaces such as the compression screws. For samples or buffer systems that are incompatible with stainless steel, glass columns should be used. Selecting Instrument Type An isocratic (one pump, one buffer) HPLC or FPLC system is required for the columns described in this unit. Selection of an appropriate chromatography system involves two major decisions: operating pressure range and whether or not biocompatibility is needed. FPLC, HPLC, and biocompatible systems are appropriate for different types of applications. An FPLC system is used to run columns that require moderate back pressure (typically 50 to 600 psi). An HPLC system is used to run columns with higher back pressures (typically >500 psi). Biocompatible systems are available for operation in a variety of pressure ranges and are used in cases where the protein of interest may be adversely affected by trace metal ions that could exist with a system containing stainless steel. Biocompatibility is a heterogeneously applied term, generally assumed to mean a system with few or no stainless steel components. The importance of biocompatibility for most protein applications has not been clearly established. Many companies sell integrated computer-controlled systems that consist of reliable pumps, an injector, a UV detector, a fraction collector, and a computer output for data collection and manipulation. Of course, a dual-pump HPLC or FPLC system may be used by utilizing only one of the pumps. It is most critical to have a pumping system that can deliver a constant flow rate in the desired range, which is typically 0.1 to 1.5 ml/min for analytical columns and up to 5 ml/min for preparative columns. Because of the small number of samples typically analyzed, an autosampler is not necessary. However, it is important to have a computer integrated with the injector to ensure precise starting points for sample injection. While most binary HPLC and FPLC systems are relatively expensive, there are less expensive alternatives, such as integrated isocratic systems that cover the required pressure range (e.g., 0 to 2500 psi) and cost under $10,000 (e.g., D-Star Instruments Integrated Isocratic HPLC system, available through Thomas Scientific).
Characterization of Recombinant Proteins
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Selecting Appropriate Buffers The buffer selected for gel filtration should be optimized for protein stability. Highly hydrophobic or membrane proteins, for example, require a nondenaturing detergent in the buffer (see next section). Silica-based resins are not stable above pH 7.5, and stainless steel columns should typically not be used with halogens. Buffer type and strength may affect resolution and mass recovery. The ionic strength of the buffer is also important. In high–ionic strength buffers (>1 M), hydrophobic interactions between the protein and the resin may occur, while at low ionic strength (<0.1 M), ionic interactions may occur. Analysis of Membrane Proteins in Detergents The separation of membrane proteins in the presence of detergents (typically a nonionic or zwitterionic detergent) is the same in principle as separation in aqueous solutions. However, in the presence of detergent, the apparent size of the protein will probably increase due to detergent binding to the protein. For this reason, the fractionation range of gel filtration columns specified for water-soluble proteins will not apply to detergentsolubilized proteins (Schagger, 1994). Accurate size determination is not feasible in the presence of detergents, since membrane proteins will bind detergent molecules and/or interact with detergent micelles to differing degrees. The procedure for gel filtration in the presence of detergent is essentially as described in the Basic Protocol, with a few additional factors to take into account. First, the protein sample should be equilibrated with the detergent prior to the run. Detergents with a low critical micelle concentration (CMC) do not dialyze well, so the detergent can be added directly to the sample, or a desalting column can be used for buffer exchange. The choice of detergent is another important consideration. If the protein elution profile is to be detected via UV absorbance, for example, the detergent needs to be transparent at the wavelength selected (for example, detergents containing aromatic ring systems, e.g., Triton, absorb strongly at 280 nm), unless the concentration of detergent can be kept low enough to keep the absorbance signal within the linear range of the UV detector. However, the gel filtration buffer should contain detergent at a concentration well above the CMC (i.e., ≥4×) to prevent association of membrane proteins (Schagger, 1994). The micelle size is also important, as large micelles will change the elution behavior of proteins more than small micelles, and may be harder to separate from the protein of interest. A common problem in gel filtration of membrane proteins is adsorption of protein to the resin or the column. The extent of this problem depends upon the detergent type and concentration, as well as the resin type and column hardware. Unfortunately, predicting the column and detergent that will minimize adsorptive losses is difficult, and a trial-anderror approach may be needed to find the optimum combination. BASIC PROTOCOL
Characterizing Recombinant Proteins Using HPLC and MS
HPLC ANALYSIS OF RECOMBINANT PROTEINS This protocol describes analytical HPLC gel filtration of soluble recombinant proteins (refer to UNITS 8.1 & 8.3 for a complete description of gel filtration chromatography). Analytical HPLC is used to evaluate sample heterogeneity and quantify sample components. For example, HPLC may be used to monitor the proteolytic cleavage of a fusion protein (Fig. 7.10.1), to separate two or more oligomers, or to determine the amount of aggregate or other contaminates in a sample. Although the protocol is written for HPLC systems, FPLC and HPLC are largely interchangeable (see Strategic Planning), and either can be used. As analytical HPLC is often used to compare different preparations of a recombinant protein, to monitor the homogeneity of a sample over time, or to compare a recombinant
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protein with standards, accurate and reproducible determination of retention time is critical. For this reason it is important to use high-precision chromatography pumps that provide a constant, accurate flow rate. It is also important that the column packing be in good condition and maintained at a constant temperature; typically 4°C is used to minimize potential proteolysis of the protein. This can be accomplished by placing either the column or the entire chromatography system in a cold cabinet. Materials Gel filtration buffer (see recipe) Protein standard (e.g., Bio-Rad gel filtration standard) Protein sample 0.05% (w/v) sodium azide or 20% (v/v) ethanol
0.25
*
GST
Absorbance (280 nm)
0.20
0.15
*
0.10
*
0.05
0.00
0
5
10
15
20
25
30
Time (min)
Figure 7.10.1 Comparison of related fusion proteins analyzed immediately after protease cleavage using analytical HPLC gel filtration to evaluate potential self-aggregation. The top panel shows 100 µg of the wild-type 52-kDa recombinant protein, and the lower panels show two different mutants. The asterisk indicates the retention time of the monomeric proteins. The elution position of the glutathione-S-transferase (GST) moiety is also indicated. Peaks eluting earlier than the monomeric form of the protein are self-aggregates. These results show that the mutants, but not the wild type, rapidly aggregate during protease digestion. When monomeric forms of the mutants were isolated by preparative HPLC gel filtration and reanalyzed by analytical gel filtration, a substantial increase in aggregation occurred in a concentration- and time-dependent manner (data not shown).
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Isocratic liquid chromatography system (HPLC or FPLC) Prepacked HPLC or FPLC column (Table 7.10.1) 0.2-µm filters 1. Select an appropriate column for the type of proteins to be analyzed. Refer to Strategic Planning and Table 7.10.1 for a listing of prepacked analytical HPLC columns. 2. Prepare an isocratic liquid chromatography system as in Basic Protocol 1 of UNIT 8.3, steps 14 to 16 (also see Strategic Planning of this unit). It is desirable to have an HPLC UV detector in line with outputs to both a strip chart recorder and to a computer for peak quantification.
3. Filter gel filtration buffer that is compatible with both the column and the sample, using a 0.2-µm filter to degas and remove any small particles that may clog the column frit. Refer to the manufacturer’s recommendations for column-compatible buffers. The column buffer does not need to be identical to the buffer of the injected sample, as gel filtration can also be used to exchange buffers; however, the sample should be tested with the column buffer to ensure that no precipitate occurs when the sample enters the column. If bacteriostatic agents are not used, buffer should be filtered daily.
4. Equilibrate the column with three column volumes of gel filtration buffer. Do not exceed the manufacturer’s recommended flow rates or pressure limits. If operating at low temperatures, it may be necessary to decrease the flow rate as the viscosity of the buffer may increase, generating higher back pressures.
5. Pass a protein standard through a protein-compatible 0.2-µm filter or centrifuge at 10,000 × g for 10 min to remove particulate matter. 6. Rinse the sample application loop with gel filtration buffer and evaluate the performance of the column by injecting the protein standard. Figure 7.10.2 shows chromatograms obtained with columns showing optimal (upper panel) and poor performance (lower panel). Note the reduced peak heights, increased peak widths, and decreased separation between peaks in the lower chromatogram.
7. Pass a protein sample through a 0.2-µm filter or centrifuge at 10,000 × g for 10 min to remove particles. 8. Rinse the sample application loop again and inject an aliquot of the recombinant protein to be analyzed. The lower limit of protein concentration is determined by the sensitivity of the detector used, while the upper limit is influenced by multiple factors including sample viscosity, potential nonideal behavior of the sample at high concentrations (e.g., aggregation), and degradation of resolution between adjacent peaks as protein loads increase. For optimum resolution, sample volumes should be ≤1% of the total column volume (column volume [ml] = πr2l, where radius [r] and length [l] are in cm). For initial analytical separations, a sample volume of ∼0.5% of the column volume can generally be used, with a total protein concentration of ∼0.5 to 1.0 mg/ml. Figure 7.10.3 shows a typical analysis of a 55-kDa recombinant protein by SDS-PAGE, HPLC, and MALDI-MS.
9. Optional: Collect fractions of the eluting proteins.
Characterizing Recombinant Proteins Using HPLC and MS
Collection of fractions during a gel filtration separation allows subsequent identification of individual peaks with other methods such as MALDI-MS (see Support Protocol 2 and Fig. 7.10.5C), SDS-PAGE (UNIT 10.1 and Fig. 7.10.5B), immunoblotting (UNIT 10.10), or N-terminal sequencing (UNIT 11.10). However, it should be noted that because the amount of protein typically loaded on an analytical HPLC column is small, and proteins are diluted
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0.40
Absorbance (280 nm)
0.30
0.20
0.10
0.00 0
5
10
15
20
25
Time (min)
Figure 7.10.2 Standard protein mix to evaluate column performance. A comparison between new (upper panel) and old (lower panel) TSK G3000SWXL (TosoHaas) columns. In this case, the decreased resolution of the old column was primarily attributed to a void that had formed at the top of the column (see Troubleshooting).
during the separation, the protein yield and/or concentration may not be high enough for some forms of analysis. In some cases the fractions may be pooled and/or concentrated prior to analysis, otherwise the separation may have to be scaled up (see Strategic Planning). If collecting fractions, it is a good idea to collect through the entire run time of the separation, using fraction volumes that will provide at least four to six fractions across a typical peak (this can be estimated from the average peak widths of standard proteins).
10. Store column in a solution containing a bacteriostat such as 0.05% sodium azide or 20% ethanol. Refer to the manufacturer’s recommendations regarding column storage. DETERMINATION OF STOKES RADIUS BY HPLC GEL FILTRATION This protocol describes the use of HPLC gel filtration to rapidly determine the Stokes radius of proteins (also see UNIT 8.3, Basic Protocol 3). Protein standards may be ordered individually or as part of a kit. Table 8.3.8 provides a list of commercially available standards. It is important to note that a Stokes radius is a measure of molecular size and not molecular mass; effectively, it is the radius of an equivalent hydrated sphere (Cantor and Schimmel, 1980). Thus, an approximate molecular mass may be obtained only when the protein of interest and the protein standards that are used in calibration have the same shape. As most standards are roughly spherical, the protein of interest should also be spherical. A better estimate of molecular mass (independent of shape) may be calculated from the Stokes radius combined with a sedimentation coefficient, which can be obtained using analytical ultracentrifugation or sucrose gradients (Seigel and Monty, 1966; UNIT 7.5). 1. Select a column of appropriate fractionation range for samples of interest and equilibrate the column in a compatible gel filtration buffer (see Basic Protocol, steps 1 to 4).
SUPPORT PROTOCOL 1
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A
B
0.20
Absorbance (280 nm)
0.15 66 46 0.10 36 V0
Vt 25
0.05
0.00 0
5
10
15
20
25
30
Time (min)
66,431
C
2,000
55,861.3
33,217
27,897
Counts
4,000
0 20,000
30,000
40,000
50,000
60,000
70,000
Mass (m/z)
Figure 7.10.3 Analysis of a purified recombinant protein by HPLC gel filtration, SDS-PAGE, and mass spectrometry. (A) HPLC chromatograph of an α-actinin recombinant (25 µg injected). (B) SDS-PAGE Coomassie blue–stained gel showing the α-actinin recombinant. (C) Mass spectra of α-actinin recombinant protein. The apparent mass from SDS-PAGE is close to the expected mass. The HPLC chromatograph shows a single species, which migrates near the midpoint of the fractionation range, indicated by the void volume (V0) and the total column volume (Vt). However, when mass analysis was performed, the actual mass observed ([M + H]+ = 55,861.3 or [M] = 55,860.3) was 738 Da larger then the sequence molecular mass, corresponding to a readthrough of the insert stop codon. Instead, the sequence terminated at the in-frame stop codon in the cloning vector. The peak at 66,431 is the single protonated form of BSA, an internal standard. The peaks at 27,897 and 33,217 correspond to the double-protonated species of the recombinant and standard, respectively. Characterizing Recombinant Proteins Using HPLC and MS
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Stokes radius (Å)
1000
100
10 0
10
20
30
40
50
Ve (ml)
Figure 7.10.4 Stokes radius calibration. Two Superdex 200 columns (Amersham Pharmacia Biotech) were connected in series and run at a flow rate of 0.5 ml/min in PBS, pH 7.4, containing EDTA, phenylmethylsulfonyl fluoride, and sodium azide. Individual protein standards from the Amersham Pharmacia Biotech high- and low-molecular-weight gel filtration kits were prepared at a concentration of 1 to 5 mg/ml in gel filtration buffer. A total of 50 µg of each standard was injected. Samples included: ribonuclease A (16.4 Å), chymotrypsinogen A (20.9 Å), ovalbumin (30.5 Å), albumin (35.5 Å), aldolase (48.1 Å), catalase (52.2 Å), ferritin (61.0 Å), and thyroglobulin (85.0 Å). The retention time was multiplied by the flow rate (measured as 0.45 ml/min) to determine the elution volume (Ve), which was plotted verses the log of the known Stokes radii (Rs). The equation for the fitted line is logRs= −0.04Ve+ 2.65. This equation can then be used to determine the Stokes radius of an unknown, by injecting a small amount of the sample, determining Ve, and solving the equation for Rs.
2. Dissolve individual protein standards to a concentration of 1 to 5 mg/ml in gel filtration buffer. Pass through a protein-compatible 0.2-µm filter or centrifuge at 10,000 × g for 10 min to remove particulate matter. Protein standards can be mixed together to save time and reduce the number of injections; however, some proteins may interact and affect the elution of other proteins in the mix.
3. Inject each protein standard or the protein standard mixture onto the column. Rinse the sample application loop with buffer prior to each injection. For accurate comparison of the standard and sample retention times, the volume of the protein standard injected onto the column should be the same as that of the sample to be tested.
4. Determine the elution volume (Ve) of each standard by multiplying the retention time by the flow rate. Because pump performance will vary on most systems as piston seals wear, it is a good idea to check the actual flow rate by measuring the precise time required for 10 ml of eluant to be collected in a 10-ml graduated cylinder or volumetric flask.
5. Construct a graph plotting the log of the known Stokes radius (Rs) versus elution volume for each standard. Figure 7.10.4 shows a graph of logRs versus Ve for a series of protein standards run on a Superdex 200 column.
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6. Fit the points with a straight line. Determine the slope (a) and y offset (b) of the line. Most graphing or spreadsheet programs may be used to perform linear regression analysis on the data points, giving the slope and offset of the fitted line.
7. Inject protein samples with unknown Stokes radii onto the same column. Calculate Ve and use the equation logRs = aVe + b to determine the Stokes radius of each unknown (see Fig. 7.10.4). SUPPORT PROTOCOL 2
MALDI-MS ANALYSIS OF RECOMBINANT PROTEINS The following protocol describes a basic method for MALDI-MS analysis of proteins carried out on a Voyager RP spectrometer (PerSeptive BioSystems). See UNITS 16.1-16.3 or Beavis and Chait (1996) for a more detailed discussion of MALDI-MS theory and methods. Two matrices are commonly used for recombinant proteins: α-cyano-4-hydroxycinnamic acid (for proteins <20 kDa) and sinapinic acid (for proteins >20 kDa), each in 33% acetonitrile/0.1% trifluoroacetic acid. The optimum matrix may have to be determined empirically for each sample. Materials 1 to 10 pmol/µl HPLC-purified protein sample of interest (see Basic Protocol) MALDI-MS-compatible buffer (e.g., 10 mM ammonium bicarbonate, pH 8) Molecular weight standards Saturated matrix solution (see recipe) Sample target (e.g., gold or stainless steel plate) Matrix-assisted laser desorption/ionization (MALDI) mass spectrometer (Voyager RP, PerSeptive BioSystems) 1. Optional: Dialyze an HPLC-purified protein sample into a MALDI-MS-compatible buffer. This step removes buffer components that may interfere with ionization or crystal formation. See Table 16.2.1 for tolerance limits of buffer components. Sodium azide, which is typically used in HPLC gel filtration buffers as a bacteriostatic reagent, is particularly problematic if not removed by dialysis, as it causes strong signal suppression. Very small samples (<300 ìl) can be dialyzed in a microcentrifuge tube (Overall, 1987).
2. Dilute sample to a concentration of 1 to 5 pmol/µl with water or MALDI-MS buffer. 3. Select and prepare molecular weight standards. Two mass peaks are typically used to calibrate the mass spectrometer; these can be the singly and doubly charged species of a single standard, such as protein A, or two different protein standards. Ideally, the molecular weight of the standards should bracket the expected masses of the sample. See Table 7.10.2 for a list of typical molecular weight standards. The most accurate calibration is obtained when the standards are included in the sample of interest (internal calibration). However, the standard may suppress ionization of the recombinant protein, in which case the ratios of standards and sample must be adjusted empirically or an external calibration must be used.
4. Apply 0.5 µl saturated matrix solution to a sample target and allow to air dry. Apply 0.5 µl protein sample onto the dried matrix and allow to dry. Characterizing Recombinant Proteins Using HPLC and MS
Alternatively, mix 2 ìl matrix with 2 ìl sample in a microcentrifuge tube, apply 1 ìl to the sample target, and allow to air dry.
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Table 7.10.2 of Proteins
Common Molecular Weight Standards for MALDI-MS
Standard Bradykinin Insulin B Insulin Ubiquitin Cytochrome c α-Lactalbumin Trypsin inhibitor A α-Chymotrypsin Ovalbumin Protein A BSA
Molecular weight (Da)
Concentration (pmol/µl)
1061.2 3496.9 5734.6 8565.9 12231.9 14071.1 20091.7 25234.7 44401.0 44614.0 66341.0
1 1 2-4 1-2 1-2 1-2 1-2 1-2 3-5 3-5 3-5
Matrix HCCAa HCCA HCCA HCCA HCCA HCCA Either Either Sinapinic acid Sinapinic acid Sinapinic acid
aHCCA, α-cyano-4-hydroxycinnamic acid.
5. Insert sample target into a MALDI mass spectrometer and follow the manufacturer’s procedure for obtaining a spectrum. 6. Compare the mass obtained with that expected from the amino acid sequence of the recombinant protein. The masses should agree to within ∼0.1% when an external standard is used, and ∼0.03% using an internal standard. If this is not the case, or if additional peaks are present, there may be some modification of the protein. Table 7.10.3 lists some common modifications seen with recombinant proteins. Figure 7.10.5 shows typical MALDI-MS results.
Table 7.10.3
Interpreting Mass Spectra of Recombinant Proteins
Problem/change
Comment
Typical mass change
Stop codon readthrough (Fig. 7.10.3)
Stop codon in the construct is missed; translation continues until the next stop codon, often in the vector
+50 to 1000
Disulfide adduct
Disulfide bond formed between a reducing agent and a highly reactive cysteine in the recombinant protein
+76 (2-mercaptoethanol) +119 (dithiothreitol) +305 (glutathione)
Disulfide dimer
Formed between two reactive cysteines
2× expected mass
Protease degradation
For example, due to contaminating proteases, or secondary cleavage sites for proteases used to cleave fusion protein
Variable size reduction
Incomplete removal of tag sequences
For example, His tag (∼6 His) or Flag sequence
Increased mass should equal calculated tag sequence
Post-translational modifications Only in eukaryotic expression systems; (e.g., glycosylation; Fig. 7.10.5C) glycosylation usually gives broad or multiple peaks Removal of initiation Met residue
Variable (+ ≥300) −131
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0.80
S 1 2 3
C 3,500
3,000
36 25
0.60
12
0.40 0.20 1
2
29,314.9
Absorbance (280 nm)
3
1.00
200 95 68
28,262.5
B
1.20
Counts
A
2,500
2,000
0.00 5
10 15 20 25 30
1,500
Time (min) 1,000 25,000 30,000 35,000 Mass (m/z )
Figure 7.10.5 Identifying post-translational modifications using MALDI-MS. (A) HPLC chromatograph of a recombinant extracellular domain expressed in Sf9 insect cells. The recombinant protein (peak 3) is effectively separated from high-molecular-weight contaminants (peaks 1 and 2). (B) SDS-PAGE of the sample before gel filtration (lane S), and fractions taken from HPLC peaks 1 to 3 (lanes 1 to 3). (C) MALDI mass spectrum of the purified extracellular domain from peak 3. Two broad peaks are detected, with molecular masses 0.6 to 1.7 kDa greater than the molecular mass calculated from the sequence (27,531 Da), indicating heterogeneous glycosylation of the protein. The sequence mass was calculated after determining the site of signal peptide cleavage using N-terminal sequence analysis of the intact recombinant.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Gel filtration buffer PBS (APPENDIX 2E) or other buffer compatible with both the column and sample. Protease inhibitors (e.g., 1 mM EDTA, 0.15 mM phenylmethylsulfonyl fluoride) as well as a bacteriostat (0.05 % sodium azide) may also be added to the buffer. A reducing agent (2-mercaptoethanol, dithiothreitol, or tris(2-carboxyethyl)phosphine hydrochloride [TCEP⋅Cl]) may also be included to prevent proteins containing reactive cysteines from forming disulfide linkages. Buffers should be degassed and made fresh each week unless antimicrobial agents have been added, in which case it may be used for up to one month.
Saturated matrix solution Prepare 10 mg/ml matrix (α-cyano-4-hydroxycinnamic acid or sinapinic acid) in 33% (v/v) acetonitrile/0.1% (v/v) trifluoroacetic acid. Centrifuge briefly to sediment undissolved matrix. Use the saturated solution to prepare samples. Stored for up to 2 weeks at 4°C, protected from light. Characterizing Recombinant Proteins Using HPLC and MS
For optimum sensitivity, the α-cyano-4-hydroxycinnamic acid matrix supplied by the manufacturer can be repurified (see UNIT 16.3, Support Protocol).
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COMMENTARY Background Information Analytical HPLC gel filtration and MALDIMS are relatively rapid and convenient methods that can be used to detect (and in some cases help identify) many protein modifications and potential self-aggregation seen after expression of recombinant proteins. Gel filtration chromatography is one of the oldest methods used to characterize proteins, and its resolution and speed are substantially improved by modern small-particle high-performance resins. The general aim of analytical HPLC gel filtration is to separate and quantify the components of a protein mixture based on their solution size. In the case of recombinant proteins, the method is used primarily to quantify contaminants, especially self-aggregates (Fig. 7.10.1), and to obtain an approximate Stokes radius of each of the sample components. One of the major advantages of gel filtration over other techniques (e.g., SDS-PAGE) is that it is nondenaturing, enabling the separation, quantification, and approximate size determination of oligomers and aggregates of the protein of interest as they occur in solution. Furthermore, gel filtration can be performed in a variety of solution conditions, so the effects of ionic strength, pH, temperature, or reducing agents on the sample may be tested. Other advantages of the technique are that the sample is recoverable—allowing further analyses (e.g., SDS-PAGE or mass spectrometry) to be performed on the separated components— and that the analysis time is relatively short (typically ~30 to 60 min). There are some limitations of gel filtration, and additional techniques should be used to fully characterize recombinant proteins. Depending on the sensitivity and detector wavelength used, HPLC analysis may fail to detect some contaminants that might be seen by SDSPAGE, such as proteins without tyrosine or tryptophan residues, which will not have substantial absorbance at 280 nm. For size determination, HPLC gel filtration is fast, requires very little protein, and gives excellent fractionation, but has inherent drawbacks, such as the potential for retardation of protein elution caused by electrostatic or hydrophobic interactions with the column matrix. For comparison, velocity sedimentation (UNIT 7.5) requires no calibration standards and there is no solid phase to interact with the protein, but the fractionation of complex mixtures is not as great. It is important to note that, due to the large variations in protein shape and degree of hydration, a Stokes
radius is often not a good indicator of oligomeric state. While HPLC is a rapid first analysis method, the oligomeric state of a protein should be definitively determined using equilibrium sedimentation in an analytical ultracentrifuge (UNIT 7.5). MALDI-MS is an extremely valuable tool for the protein chemist, allowing determination of protein molecular mass within a few daltons, regardless of the size, shape, or hydration of the protein. Other advantages of this technique are that it can be applied to protein mixtures, only ∼10 pmol of protein is needed, and the analysis can be completed in a few minutes. The drawbacks of this technique are that some proteins are difficult to ionize and that it is not quantitative, as different proteins can yield highly variable signal intensities. Thus, the utility of MALDI-MS is the accurate mass determination of the protein of interest, which can detect most post-translational and artifactual modifications of that protein. It should also be noted that while obtaining the mass predicted by the sequence of the protein of interest is a good indication that the sequence is correct, confirmation of sequence identity may also require N-terminal sequence analysis of the intact protein and HPLC/MS mapping of tryptic peptides (UNIT 7.3).
Critical Parameters The maximum sample volume that is applied to the column should not exceed 0.5% to 1% of the total column volume for optimal resolution of the sample components. Sample capacity may be increased by using a larger-diameter column. Another way of achieving maximum resolution of components is to place two or more columns in series, effectively increasing the length of the column. Using columns of smaller particle size can increase the resolution of closely eluting components as well as reducing band broadening. Using a lower flow rate than the maximum recommended by the manufacturer may also lead to better resolution of closely eluting components. Proper care of the columns and HPLC system will ensure the maximum lifetime of the system. After use, the columns and HPLC system should always be stored in a bacteriostat as recommended by the manufacturer. Worn pistons and seals should be replaced to maintain flow rate, and hence retention time reproducibility. Factors that can reduce the lifetime of a column include deposition of aggregated ma-
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terial on the column frit, introduction of air into the column, running the columns at flow rates and pressures exceeding the manufacturer’s recommendations, and exposing silica-based columns to pHs >7.5.
Troubleshooting
Characterizing Recombinant Proteins Using HPLC and MS
The most common problems encountered in HPLC gel filtration are decreased sample recovery, decreased resolution of sample components, increased back pressure, and changes in retention time. The first step in troubleshooting is tracing the problem to its source: the column, sample, buffer, or instrumentation. Use a process of elimination to find the source by changing or removing one component of the system at a time. Decreased sample recovery may be due to adsorption of the sample to the column matrix. It has been the authors’ observation that some proteins tend to adhere to silica-based matrices. In particular, hydrophobic or membrane proteins may not elute from the column except in the presence of a detergent. A decrease in resolution (due to peak tailing or broadening) or a change in retention time is often due to the formation of a void at the top of the column bed or a dead volume elsewhere in the system—e.g., a bad tubing connection between the column(s) and the detector, which creates a small mixing chamber. A void at the top of the column can be due to bed compression, introduction of air onto the column, or dissolution of silica at pHs >7.5. Bed compression can occur if the flow rate or back pressure is too high. To prevent particulate matter from clogging the frit or column matrix, buffers should be filtered and samples should be filtered or centrifuged prior to injection. Another method is to install a guard column in front of the fractionating column, which will trap any particles or precipitating proteins that are accidentally introduced into the system. Air bubbles can occur if the buffer has not been degassed properly, or they may be introduced into the system during sample injection if the sample loop is not completely filled with buffer and sample. A void may be filled using some of the same column matrix; however, care should be taken not to disturb the existing column bed. An increase in back pressure may be due to a clogged frit. The best remedies for this are to reverse flush the column, clean the column according to the manufacturer’s recommendations, or replace the end fitting. When cleaning the column, make sure the cleaning buffers are compatible with all other components of the
system. The column should be run at a lower flow rate and the detector should be taken out of line. The column frit should also be changed if a visible space develops between the adaptor and the filter. Resolution of the columns should be checked with a standard protein mix rather than with an experimental sample. An experimental sample may contain either a leading or trailing edge due to other components or impurities, such as aggregated or proteolyzed material. Routine column tests using a well-characterized standard mixture also facilitate comparative monitoring of a column’s performance over the lifetime of the column. Problems with mass spectrometry usually involve failure of the protein sample to ionize (no signal), or signal suppression due to internal standards or buffer components such as high salt or detergent. Ionization of a sample may be improved by experimenting with different matrices, sample concentrations, sample/matrix ratios, and laser power. Signal suppression due to buffer components can be remedied by dialyzing the sample into an MS-compatible buffer such as ammonium bicarbonate.
Anticipated Results If the recombinant protein is pure and exists in a single oligomeric state with no aggregation or degradation, it is anticipated that the HPLC chromatograph will show a single, symmetrical peak. Additional peaks that elute before the protein of interest may be oligomers or aggregates of the protein, or contaminants. Peaks eluting after the protein of interest may be proteolysis products of the protein or contaminants. To identify the components, collect fractions throughout the separation procedure and analyze by another technique such as SDSPAGE (UNIT 10.1), MALDI-MS (see Support Protocol 2), immunoblotting (UNIT 10.10), or Nterminal sequencing (UNIT 11.10). MALDI-MS analysis of a pure sample with the correct sequence will show a single peak with a mass that should be within 0.1% of that expected from the amino acid sequence. Final confirmation of identity can be obtained by N-terminal sequence analysis of the intact protein and/or tryptic peptide mapping (UNIT 7.3). Additional peaks in the spectrum may sometimes be identified by their mass (Table 7.10.3).
Time Considerations Equilibration of the gel filtration column typically takes several hours. In some cases, equilibration may have to be performed over-
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night. An analytical HPLC gel filtration run can typically be performed in ∼30 min depending upon flow rate and column length.
Literature Cited Beavis, R.C. and Chait, B.T. 1996. Matrix-assisted laser desorption ionization mass-spectrometry of proteins. Methods Enzymol. 270:519-551.
Seigel, L.M. and Monty, K.J. 1966. Determination of molecular weights and frictional ratios of proteins in impure systems by use of gel filtration and density gradient centrifugation. Application to crude preparations of sulfite and hydroxylamine reductases. Biochim. Biophys. Acta 112:346-362.
Key References
Cantor, C.R. and Schimmel, P.R. 1980. Biophysical Chemistry, Part II: Techniques for the Study of Biological Structure and Function. W.H. Freeman, New York.
Beavis and Chait, 1996. See above.
Neue, U.D. 1997. HPLC Columns: Theory, Technology, and Practice. John Wiley & Sons, New York.
Neue, 1997. See above.
Overall, C.M. 1987. A microtechnique for dialysis of small volume solutions with quantitative recoveries. Anal. Biochem. 165:208-214. Schagger, H. 1994. Chromatographic techniques and basic operations in membrane protein purification. In A Practical Guide to Membrane Protein Purification (G.V. Jagow and H. Schagger, eds.) pp. 23-57. Academic Press, San Diego.
A practical guide to MALDI-MS of proteins, including details of sample preparation and data analysis.
A theoretical and practical guide to HPLC, including properties and selection of columns, methods development, and troubleshooting.
Contributed by Gillian E. Begg, Sandra L. Harper, and David W. Speicher The Wistar Institute Philadelphia, Pennsylvania
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Rapid Screening of E. coli Extracts by Heteronuclear NMR
UNIT 7.11
Assessing whether a protein or protein complex is amenable to structural analysis is an important component in the structural genomics effort. In particular, if complete sets of structures for entire genomes are to be obtained within a reasonable time frame, high throughput methodologies for all steps along the way have to be developed. These days, cloning and expression systems are highly optimized and a variety of commercially available vectors can be used. However, heterologous proteins or protein domains expressed in bacteria may not be soluble or correctly folded, necessitating intricate solubilization and refolding schemes prior to structural or functional studies. NMR spectroscopy is an important tool for assessing the solubility, stability, and structural integrity of a gene product. It allows efficient evaluation of many variations of polypeptide length and sequence (without time- and labor-intensive purification/refolding procedures) using 1H-15N-HSQC spectroscopy of samples of 15N-labeled proteins directly from crude E. coli extracts (Clore and Gronenborn, 1996). In addition to screening for particular properties of the expressed protein alone, it is also possible to map intermolecular interactions such as ligand binding (Huth et al., 1997). In this unit, the basic methodology for bacterial growth, isotope labeling, and spectroscopic evaluation of the protein structure is provided. STRATEGIC PLANNING Choice of Expression System A variety of efficient expression systems are available (see Chapter 5). Important for the present purpose is a tightly controlled transcription and translation system. As an example, the T7 promoter-driven system originally developed by Studier (Studier et al., 1990) has enjoyed widespread use, although other systems (λ pL, tac, trp, lac, araBAD promoters) can be employed as well. In addition, the decision regarding whether expression as an untagged protein or fusion protein is preferable should take into account any previous knowledge about solubility and stability of the protein. For novel and uncharacterized proteins or domains, it is generally advisable to express these as fusions, increasing the probability of producing large amounts of protein in a folded form. For most N-terminal fusion proteins, such as those containing domains from glutathione S-transferase, maltose binding protein, protein A, thioredoxin, or the immunoglobulin-binding domain of streptococcal protein G (GB1), yields are high and there is generally no need to optimize expression conditions. However, careful attention should be paid to the selection of host strains in each particular expression vector–host system. It must be established that growth of the bacterial host in a modified minimal medium is not impaired since labeling is achieved using 15NH4Cl and glucose as sole nitrogen and carbon sources, respectively. Unfortunately, the published genotypes of host strains may not list important amino acid requirements. A comprehensive description of methods for biosynthetic enrichment of proteins with 15N is provided in Muchmore et al. (1989). NMR Instrumentation Any high-field NMR spectrometer (11.75 to 21.15 Tesla, 500 to 900 MHz 1H frequency) equipped with a triple-resonance gradient probe can be used to obtain 1H-15N HSQC spectra. Higher field instruments obviously produce spectra with higher resolution and sensitivity, although excellent performance for screening purposes can be obtained with 500-MHz instruments and cryo-probes. For high-throughput structural genomics appliContributed by Angela M. Gronenborn Current Protocols in Protein Science (2003) 7.11.1-7.11.8 Copyright © 2003 by John Wiley & Sons, Inc.
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cations, an automated sample changer may be advantageous. For an introduction to NMR of proteins, see UNIT 17.5. BASIC PROTOCOL
15
N PROTEIN LABELING AND NMR CHARACTERIZATION
This protocol describes the preparation and screening of crude extracts for a GB1-fusion system. Variations of this Basic Protocol are necessary for different expression systems. The key to success is tight repression prior to induction and robust expression after induction. In general, 50 to 250 ml of bacterial culture should suffice for several NMR samples. Materials Bacteria (E. coli strains BL21(DE3) or HMS174) containing the expression plasmid (GEV1 or GEV2) carrying the coding sequence for the protein of interest Minimal medium (see recipe) 1 M IPTG PBS, pH 7.4 20 mM sodium phosphate, pH 5.4 D2O Shaker incubator (New Brunswick Scientific) French pressure cell (UNIT 6.7); alternatively the bacteria can be ruptured in a microfluidizer Concentrators (e.g., Centriprep-3, Amicon) NMR spectrometer Grow bacteria, express protein, and prepare extract 1. Transform (UNIT 5.2) or obtain a 50-ml culture of bacteria (E. coli strains BL21(DE3) or HMS174) containing an expression plasmid (GEV1 or GEV2) carrying the coding sequence for the protein of interest. It is best to start with fresh transformants or a previously tested glycerol stock.
2. At an OD600 of ∼1⁄3 of the OD600 reached at stationary phase without induction, harvest the cells by centrifuging 10 min at 1200 × g, 25°C. UNIT 5.3
describes growth monitoring of E. coli.
It is important not to form a hard cell pellet since it needs to be resuspended again for further growth.
3. Resuspend the cells in pre-warmed (37°C) minimal medium containing 15NH4Cl as sole nitrogen source. Culture for an additional 30 min. This minimal media works well for growth in shake flasks as well as for small fermentations.
4. Induce protein production by adding 1 to 2 ml of 1 M IPTG per liter of culture and incubate an additional 3 to 5 hr. 5. Harvest cells by centrifuging 10 min at 4000 × g, 4°C, decant the supernatant and resuspend the cell pellet in 20 ml PBS, pH 7.4. Lyse the cells by passing them through a French pressure cell two times (UNIT 6.7). 6. Remove the cell debris by high-speed centrifuging 20 min at 20,000 × g, 4°C. Rapid Screening of E. coli Extracts by Heteronuclear NMR
7. Concentrate the supernatant in a Centriprep-3 device to a volume of ∼1 ml. At the same time change the buffer to 20 mM sodium phosphate, pH 5.4 by adding ∼10 ml of buffer to the 1-ml concentrate and concentrating this sample again two to three
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HSQC
15H
1H
Figure 7.11.1 Schematic outline of cloning, expression, extract preparation, and NMR screening.
times. Transfer 0.3 to 0.5 ml of this concentrated extract to an NMR tube and add D2O to yield a final concentration of 5% (v/v). The general outline of this approach is illustrated in Figure 7.11.1.
Perform NMR spectroscopy 8. Record 1H-15N HSQC spectra using pulse sequences previously described (Bax et al., 1990; Piotto et al., 1992) in the experimental libraries of the instrument manufacturer. In general, 128 × 512 complex points in the indirect (15N) and acquisition (1H) dimensions are recorded with total acquisition times of 64 msec in both dimensions and 16 scans per T1 increment. Figure 7.11.2 presents 1H-15N HSQC spectra of crude E. coli extracts containing 15N-labeled GB1 domain (56 amino acids) or Il-1β (153 amino acids).
ENRICHMENT OF 15N-LABELED PROTEIN For cases in which a fusion construct is employed for expression, partial purification or enrichment of the 15N-labeled fusion protein can be carried out in one step by passing the extract over an affinity column. For highly expressing constructs (>0.1 mg/ml of culture), this is not necessary, but it could be useful for poorer expressers. In the case of the GB1 fusion protein vector (Huth et al., 1997), a His-tag resides at the C-terminus in addition to the GB1 domain at the N-terminus and both domains can be used for affinity purification (e.g., employing a Ni2+ affinity column to bind the His tag; UNIT 9.4).
ALTERNATE PROTOCOL
Additional Materials (also see Basic Protocol) 50 mM Tris⋅Cl, pH 7.5/5 mM EDTA/5 mM benzamidine solution 20-ml IgG Sepharose fast-flow column 50 mM Tris⋅Cl, pH 7.5/150 mM NaCl 5 mM sodium acetate buffer, pH 5.0 5 M sodium acetate buffer, pH 3.5 10 to 20 mM sodium phosphate, pH 5 to 6 Additional reagents and equipment for SDS-PAGE (UNIT 10.1) 1. Grow cells and induce protein expression as described in Basic Protocol, steps 1 to 4.
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IL-1b
GB-1
Figure 7.11.2 SDS/PAGE of crude extract of IL-1β and GB1 and the corresponding 1H-15N HSQC spectra. Samples prepared as described in the Basic Protocol were separated on SDS/PAGE and Coomassie stained. The identical samples were used for NMR spectroscopy. Crosspeaks are labeled with the residue name and number.
2. Harvest cells by centrifuging 10 min at 4000 × g, 4°C. Decant the supernatant and resuspend the pelleted cells in 20 ml 50 mM Tris, pH 7.5/5 mM EDTA/5 mM benzamidine solution. Disrupt the cells using a French pressure cell as described in Basic Protocol, step 5. 3. Remove the cell debris by high-speed centrifuging 20 min at 20,000 × g, 4°C, and apply the supernatant containing the fusion protein onto a 20-ml IgG Sepharose fast-flow column equilibrated in 50 mM Tris⋅Cl, pH 7.5/150 mM NaCl either at room temperature or 4°C depending on stability of protein. 4. Wash the column with one to two column volumes of 5 mM sodium acetate buffer, pH 5.0 and then elute the GB1 fusion protein with 5 M sodium acetate buffer, pH 3.5. Collect 1-ml column fractions and check for protein by SDS-PAGE (UNIT 10.1). 5. Concentrate the protein-containing fractions in a Centriprep-3 device to a volume of ∼1 ml. At the same time, change the buffer to 10 to 20 mM sodium phosphate, pH 5 to 6 (see Basic Protocol, step 7). The final buffer is selected based on known properties of the expressed protein. If not information is available, a low molarity phosphate buffer, slightly on the acidic side, is recommended.
6. Transfer 0.3 to 0.5 ml of this concentrated solution to an NMR tube, and add D2O to a final concentration of 5% (v/v). 7. Acquire NMR spectra as described in Basic Protocol, step 8. Figure 7.11.3 presents the 1H-15N HSQC spectra of two different GB1- fusion proteins after one-step purification on IgG Sepharose. The spectrum in panel A illustrates the appearance of a fusion partner that is basically a random coil (resonances reside between 7.8 and 8.4 ppm for 1H) while the resonances in panel B are dispersed over the entire spectral widths, indicative of a folded fusion partner. Rapid Screening of E. coli Extracts by Heteronuclear NMR
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A
B
Figure 7.11.3 1H-15N HSQC spectra of GB1-fusion proteins. (A) The fusion partner exhibits a random coil structure (GB1-GCN4) and (B) a folded fusion partner (GB1-Fas) is present. Resonances arising from the GB1 portion are labeled by residue name and number. For GB1-GCN4 (A) all additional cross-peaks are located in the narrow region of 7.8-8.4 ppm, indicative of a random coil conformation for the GCN-4 portion. For GB1-Fas (B), cross-peaks are distributed over the entire region of the spectrum, indicative of folded Fas protein.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or its equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Minimal medium 2 ml 20% (w/v) glucose 500 µl trace elements (see recipe) 500 µl 1 M MgCl2⋅6 H2O 300 µl 5 mg/ml thiamine-vitamin B1 100 µl 10% yeast extract (optional) 50 µl 50 mg/ml antibiotic Add sterile water to 50 ml and mix Add to 950 ml of minimal salts (see recipe) Make fresh for use within 1 or 2 days Filter sterilize all stock solutions through a 0.22-µm filter Minimal salts 13 g/liter KH2PO4 (anhydrous) 10 g/liter K2HPO4 (anhydrous) 9 g/liter Na2HPO4 (anhydrous) 2.4 g/liter K2SO4 1.1 g/liter NH4Cl (either 14N or 15N) Dissolve in 950 ml H2O and autoclave Store up to several months at room temperature
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Trace elements 6.0 g CaCl2⋅2H2O 6.0 g FeSO4⋅7H2O 1.15 g MnCl2⋅4H2O 0.8 g CoCl2⋅6H2O 0.7 g ZnSO4⋅7H2O 0.3 g CuCl2⋅2H2O 0.02 g H3BO3 0.25 g (NH4)6Mo7O24⋅4H2O 5.0 g EDTA Add reagents one at a time to 1 liter of water with vigorous stirring. Wait 5 to 10 min between each addition. Each reagent must be completely dissolved before adding the next one. After addition of EDTA, stir until the solution is golden-brown. This may take 2 to 3 days, including a transition from green to a yellowish-brown color. When the color is golden yellow, sterilize by filtering through a 0.22-µm filter. Store up to several months in the dark at room temperature. COMMENTARY Background Information
Rapid Screening of E. coli Extracts by Heteronuclear NMR
A prerequisite for any structural investigation by NMR is the availability of a “well-behaved” sample. For proteins, it is necessary to have an over-producing organism, fast and effective purification methods, and the knowledge of conditions that allow the sample to survive days, or sometimes weeks, at or above room temperature. Thus, it is imperative to establish conditions (buffer, salt, pH) under which the protein is biochemically stable, folded, and amenable to spectroscopy. Usually, these conditions are determined by trial and error, using large amounts of protein, costly purification procedures, and considerable time and manpower. As such, they are ill-suited for any high-throughput strategies. For many proteins, sub-cloning of truncated versions or domains as well as point mutants is a common approach and frequently a large number of protein variants is evaluated before one with the desired biochemical and biophysical properties emerges. The methodology outlined in this unit addresses these issues and is aimed at overcoming the shortcomings traditionally encountered. Some specific advantages of this system are as follows: (1) The coupling between the highly efficient T7 polymerase expression system with the N-terminal GB1 domain in the GEV vectors ensures a high level of protein production. Depending on the particular case, protein expression can reach 25% of total cellular protein. (2) The addition of the highly soluble GB1 domain enhances the solubility of the fusion
partner. Indeed it has been shown that using this approach, the structure determination of a protein-protein complex that was insufficiently soluble and stable became possible (Zhou et al., 2001). (3) Sample conditioning can be carried out without having to cleave the fusion protein. This entails evaluating different buffer compositions and pH values for obtaining the best spectral resolution. The particular advantage of GB1 compared to other fusion proteins is its small size (6.2 kD). Most other fusion partners are fairly large (GST, 27.5 kD, and MBP, 38 kD), and as a result, their properties can easily mask those of the protein of interest. For NMR spectroscopy, this implies that a larger number of signals will arise from the protein tag, obscuring the observation of resonances from the interesting partner. For the GB1 fusions, this implies that all screening and even a structural characterization can be carried out on the fused protein (Huth et al., 1997). (4) There appears to be no interaction between the GB1 and its fusion partner for all cases studied to date (Huth et al., 1997; Zhou et al., 2001). Thus, the likelihood of inducing an artificial conformational change in the protein of interest by coupling to the GB1 domain is minimal. (5) The use of the GB1 fusion vector in combination with screening of E. coli extracts by 1H-15N HSQC spectroscopy is highly amenable to automation. In particular, the NMR screening part can be carried out in highthroughput mode using spectrometers with automated sample changers.
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(6) Using 1H-15N HSQC spectra for screening purposes exploits the intrinsically higher sensitivity of this experiment compared to other 2-D experiments, overcoming the inherent low sensitivity of NMR. Since chemical shifts are extremely sensitive to the local environment, a qualitative assessment of structural identity and similarity can be carried out by NMR, even without resonance assignments. The pattern of cross-peaks in a 1H-15N HSQC spectrum of uniformly 15N-labeled protein can be used as a fingerprint of the structure. Thus, closely similar resonance frequencies of amide groups provide a reliable measure for similarities in overall structure for related proteins or mutants and this property can be exploited for fast and efficient screening of mutant libraries by NMR (Gronenborn et al., 1996). The high quality of the 1H-15N HSQC spectra is remarkable and a direct result of the high expression levels reached in the cell. When combined with the sensitivity of a cryoprobe, these spectra can be recorded directly without further purification or concentration on <100 µM samples.
Critical Parameters With the current state of NMR methodology and instrumentation, the most critical parameter for success in this approach is the level of expression of the protein of interest. In bacterial systems, such as E. coli, high expression levels of up to 30% of total protein have been obtained. In order to restrict 15N labeling to the protein of choice and ensure >90% isotopic enrichment, the target gene has to be completely repressed before induction and this repression has to be completely lifted upon induction. Not all E. coli expression systems exhibit these properties. In other organisms, such as the yeast Pichia pastoris (UNIT 5.7) and the insect cell/baculovirus system (UNITS 5.4 & 5.5), large amounts of protein (25% to 50%) can be produced and media for labeling over-expressed proteins with NMR-active isotopes are available (Creemers et al., 1999; de Lamotte et al., 2001). Methodologies for these systems are, however, not as advanced as those for protein production in E. coli. More developmental work is needed in alternate systems and the associated costs need to come down substantially to allow for routine use. Complete media for labeling in E. coli are commercially available from Cambridge Isotopes (www.isotope.com), Isotec (www.isotec.com), Spectra Stable Isotopes (www.spectrastableisotopes.com) and Silantes GmbH (www.silan-
tes.com); media for labeling in pichia or SF9 cells are in development. Higher-field NMR instruments will obviously have higher sensitivity and therefore allow screening at lower concentrations (expression levels). The availability of cryoprobes for 500- and 600-MHz spectrometers that provide significant improvement in signal-to-noise ratios for protein samples and allow much shorter data collection times has changed the requirement for high field instruments. It is possible to record 1H-15N HSQC spectra on 15N-labeled proteins on a 600-MHz spectrometer equipped with a cryoprobe on 10 µM or 100 µM samples in 5 to 10 hr or 30 min, respectively.
Troubleshooting A frequent, but rarely reported problem with expression systems is their leakiness. If repression of the desired protein is not efficient before introduction of isotopically labeled media, substantial amounts of unlabeled material may accumulate and dilute the concentration of the labeled material. If a new expression system is investigated for purposes of screening with the methodology described in this unit, it is advised to monitor production of the expressed protein along the growth curve in medium containing 14NH Cl by SDS/PAGE prior to growth in 4 15NH Cl-containing medium. 4 Slow, insufficient growth can be caused by lesions in amino acid metabolism genes in the host strains. Commonly used host strains that grow well in the minimal medium described in this unit are BL21, DH5α, and HMS174. Strains that grow poorly or not at all are TOP10 and HB101. In addition, optimization of expression for the system at hand should be done before setting up a screening program for a large number of samples. For the GB1 fusions discussed in this unit, high yields are invariably observed, but this may not translate to other systems to the same extent.
Anticipated Results Several examples of 1H-15N HSQC spectra recorded on crude extracts and enriched samples are provided in Figures 7.11.2 and 7.11.3. With current instrumentation, excellent signalto-noise ratios can be achieved, provided the parameters for sample preparation are optimized (using >20 µM labeled protein) and the parameters for the NMR experiment are set correctly. The most likely reason for low signal intensity is aggregation of the proteins, caused by instability or incorrect folding of the ex-
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pressed protein. Adding small amounts of detergent (CHAPS) or salt may alleviate problems associated with aggregation. Figure 7.11.2 displays 1H-15N HSQC spectra of GB1 and IL-1β recorded on crude extracts and Figure 7.11.3 those of the partially enriched GB1 fusion proteins of GB1-HMG-I with and without the DNA ligand.
Time Considerations After optimization of the system, all steps, from expression to the completed NMR spectrum can be completed in <24 hr. This opens the door to high-throughput screening by NMR spectroscopy, with a drastically reduced number of sample handling steps. Employing parallel sample preparation renders this methodology amenable to easy automation.
Literature Cited Bax, A., Ikura, M., Kay, L.E., Torchia, D.A., and Tschudin, R. 1990. Comparison of different modes of two-dimensional reverse correlation NMR for the study of proteins. J. Magn. Reson. 86:304-318. Creemers, A.F., Klaassen, C.H., Bovee-Geurts, P.H., Kelle, R., Kragl, U., Raap, J., de Grip, W.J., Lugtenburg, J., and de Groot, H.J. 1999. Solid state 15N NMR evidence for a complex Schiff base counterion in the visual G-protein-coupled receptor rhodopsin. Biochemistry 38:71957199. de Lamotte, F., Boze, H., Blanchard, C., Klein, C., Moulin, G., Gautier, M.F., and Delsuc, M.A. 2001. NMR monitoring of accumulation and folding of 15N-labeled protein overexpressed in Pichia pastoris. Protein Expr. Purif. 22:318-324.
Gronenborn, A.M. and Clore, G.M. 1996. Rapid screening for structural integrity of expressed proteins by heteronuclear NMR spectroscopy. Protein Sci. 5:174-177. Gronenborn, A.M., Frank, M.K., and Clore, G.M. 1996. Core mutants of the immunoglobulin binding domain of Streptococcal protein G: Stability and structural integrity. FEBS Lett. 398:312-316. Huth, J.R., Bewley, C.A., Jackson, B.M., Hinnebusch, A.G., Clore, G.M., and Gronenborn, A.M. 1997. Design of an expression system for detecting folded protein domains and mapping macromolecular interactions by NMR. Protein Sci. 6:2359-2364. Muchmore, D.C., McIntosh, L.P., Russell, C.B., Anderson, D.E., and Dahlquist, F.W. 1989. Expression and nitrogen-15 labeling of proteins for proton and nitrogen-15 nuclear magnetic resonance. Methods Enzymol. 177:44-73. Piotto, M., Saudek, V., and Sklenar, V. 1992. Gradient-tailored excitation for single quantum NMR spectroscopy of aqueous solutions. J. Biomol. NMR 2:661-665. Studier, F.W., Rosenberg, A.H., Dunn, J.J., and Dubendorf, J.W. 1990. Use of T7 RNA polymerase to direct expression of cloned genes. Methods Enzymol. 185:60-89. Zhou, P., Lugovskoy, A.A., and Wagner, G. 2001. A solubility-enhancement tag (SET) for NMR studies of poorly behaving proteins. J. Biomol. NMR 20:11-14.
Contributed by Angela M. Gronenborn National Institute of Diabetes and Digestive and Kidney Diseases National Institutes of Health Bethesda, Maryland
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CHAPTER 8 Conventional Chromatographic Separations INTRODUCTION
I
t can be argued with conviction that the purification of proteins has been a key step in many of the advances in our understanding of biological structure and function. The careful dissection of enzyme mechanisms would have been impossible without isolated catalytic species. Before the advent of gene sequencing, determination of protein structure absolutely required clean protein samples. Precise determination of the biological effects of cytokines and growth factors necessitates the preparation of pure polypeptides. Impressive advances from X-ray structure determination must begin with homogeneous, concentrated protein samples. The utility of site-specific mutagenesis, which is crucial to asking precise questions about structure-function relationships, is limited if the recombinant protein cannot be separated from contaminating cellular proteins. Finally, the revolution created by the availability of restriction enzymes is founded upon the preparation of pure samples of the reagents.
This chapter presents a rational, systematic approach to the discovery and optimization of protein separations by chromatographic methods. The first unit provides a description of how to plan an overall strategy to attempt the isolation of a new protein (UNIT 8.1). Important initial considerations include the desired quality level of the final product and the properties of the source as well as the anticipated properties of the target protein. The choice of a strategic approach is described, beginning with consideration of the needs of the final sample with respect to concentration, buffer composition, and pH. The competing factors of selectivity, efficiency, and capacity are discussed with reference to how each method can influence the choice of the next step. The second unit provides a series of steps to identify conditions for optimizing a separation strategy (UNIT 8.2). Alternative methods for sample processing and elution are described, thus providing numerous choices for the fractionation of a variety of proteins. In addition, critical parameters for the long-term care of a chromatography system are detailed. The relationship of the ion-exchange method to other procedures is also discussed. A single procedure, no matter how carefully performed, is unlikely to totally resolve a mixture of proteins. Accordingly, use of additional procedures that provide separation based on different physical properties are essential to complete the isolation procedure. Protein fractionation by gel-filtration chromatography can provide a separation based on molecular size (UNIT 8.3). Gel filtration can also be used to remove salts from a protein at various stages of isolation; this may be valuable before ion-exchange chromatography, for example. When performed using standard proteins for calibration, gel filtration can also provide information about the size of a new protein. Another orthogonal separation method that has proven to be very valuable, particularly for protein isolation, is hydrophobic interaction chromatography (UNIT 8.4). In this method the protein interacts with a matrix through surface-exposed hydrophobic groups. Guidelines for beginning and optimizing a separation based on hydrophobic interaction chromatography are provided.
Contributed by Ben M. Dunn Current Protocols in Protein Science (2001) 8.0.1-8.0.2 Copyright © 2001 by John Wiley & Sons, Inc.
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Chromatofocusing (UNIT 8.5) is a technique that separates proteins on the basis of their isoelectric points. High resolution is achieved, making the procedure perfect for intermediate polishing steps. Selection of proper media and buffers is essential to the technique, as described in the unit. Binding of the charged groups of proteins (positive as well as negative) to the calcium phosphate matrix of hydroxylapatite provides another unique separation mechanism to resolve protein mixtures (UNIT 8.6). The unit also includes recommendations for alternative methods to derive the optimal advantages of this strategy. With the advent of more sensitive methods for the analysis of protein sequence (UNITS 11.8 & 11.10), it became clear that high resolution protein and peptide separation methods were needed. Fortunately, high performance liquid chromatography was developed at the right time to fill this need. By utilizing smaller bead sizes, the resolving power of many types of chromatography could be enhanced. UNIT 8.7 provides a thorough discussion of the methodology for optimizing a separation using HPLC procedures. General instructions for the care of the equipment are presented along with standard procedures for running a separation with high-pressure pumps. This is followed by specific directions for the different types of separation mechanisms that may be achieved by HPLC methods. Each unit defines chromatographic parameters and describes methodology to provide quantitative evaluation of column efficiency. In addition, each unit presents summaries of the properties of chromatographic media available from commercial sources, which have undergone significant improvements over the past twenty years. The advantages of some features such as pore size and bead diameter are discussed in the context of the competing needs of resolution, capacity, and speed. Although all the procedures are presented so that separations can be done manually, the advantages of automation for some steps are described. As separation technology is rapidly changing, future updates will expand the methodology described in this chapter. Ben M. Dunn
Introduction
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Overview of Conventional Chromatography The key to success in purification of any protein is developing the right separation strategy. This involves selecting and correctly applying a combination of separation techniques, chosen on the basis of information about the target protein and contaminants. There is no single separation technique and no single purification scheme that will allow successful purification of all types of proteins. However, there is a single approach to developing a successful purification strategy. Three basic steps must be completed before beginning to develop a purification strategy. First, the amount and purity of the target protein (the protein to be purified) required for its intended use must be determined. Second, all available physical and chemical information concerning potential source materials, the nature of the target protein itself, and major contaminants must be gathered. Third, the best source material from which to purify the target protein must be determined. These three steps are mutually interdependent—what is learned in one will have an impact on the others. The more that is known before beginning development of a purification strategy, the more successful that strategy will be.
YIELD AND PURITY OF TARGET PROTEIN How much protein must be purified? How pure must the protein be? These two questions establish the criteria for selecting specific chromatographic techniques and for judging the efficacy of any individual purification step or overall purification procedure. The answers to these simple questions depend upon the nature of the source material from which the target protein will be purified, the intended use for the purified protein, and the methods available to assess the purity of the target protein.
Amount of Protein Required The amount of protein required from a purification process depends primarily on the intended use for the target protein. However, the amount of protein that will be used during the purification process to develop methods and assess purity must also be considered. It is necessary only to estimate whether analytical (≤1 mg), preparative (≤1 g), pilot (≤ 1 Kg), or process (>1 Kg) amounts of target protein will be needed. This is often referred to as the scale of operation. When the source material is very Contributed by Alan Williams Current Protocols in Protein Science (1995) 8.1.1-8.1.9 Copyright © 2000 by John Wiley & Sons, Inc.
UNIT 8.1
expensive, unstable, or difficult to obtain, it is necessary to develop and optimize the purification strategy on a small scale first, then scale up. Conversely, if multiple samples are to be processed, as for analytical applications, it may be desirable to develop the purification process at a larger scale, then scale down. The amount of target protein to be purified may also be determined by limited availability of source material.
Source Material Once the scale of operation has been determined, the amount of source material needed to begin the procedure must be clearly defined. This obviously depends on the concentration of target protein in the source material, which can vary greatly. Certain proteins and peptides are present in natural sources only in very minute quantities. Purification of even a microgram of pure material may require many kilograms of tissue, or many liters of fermentation broth if microorganisms are the source. On the other hand, some recombinant proteins are expressed at such high levels in a host that the protein may be present in milligram per milliliter or higher concentrations. Source materials containing very low or very high concentrations of target protein each present unique problems that must be addressed in selecting a purification strategy.
Yield of Purification Procedure The next major consideration in deciding how much protein to purify is what yield is necessary at each step of the purification procedure to ensure that there will be enough pure target protein at the end, taking into account the availability of source material. If a 75% yield of target protein is obtained at each of four steps in a purification procedure, less than one-third of the desired protein will remain at the end of the procedure (Fig. 8.1.1). It is obviously advantageous to use as few steps as possible in a purification process while maintaining as high a yield as possible at each step. The yield from any purification step may be reported as total protein, percentage of total activity, or change in specific activity. The yield information from progressive purification steps is used to construct a purification table (e.g., see Table 8.1.1). In Table 8.1.1, the yield is reported as percent of total activity remaining from the crude
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8.1.1 CPPS
95%/step 90%/step
85%/step 80%/step 75%/step
Target protein recovered (%)
100 90 80 70 60 50 40 30 20 10 0 0
1
2
3
4
5
6
7
8
9
10
Number of steps in procedure
Figure 8.1.1 Theoretical yields from multistep protein purifications. Each curve represents a multistep process with a given percentage yield per step. The percentage of the original quantity of target protein remaining at the end of purification is plotted against the number of purification steps.
homogenate; the step yield is the percent activity relative to the previous step; and the purification factor reflects the change in specific activity relative to the crude homogenate. It is not unusual to have yields >100% based on activity (e.g., the yield after anion-exchange chromatography in Table 8.1.1). This is most often due to the removal of inhibitors or contaminants that interfere with the activity assay.
Assessment of Protein Purity
Overview of Conventional Chromatography
Assessing protein purity is often the most time-consuming and labor-intensive step in the purification process. It is generally desirable to limit the number of steps in a purification process simply to reduce the number of assays that must be performed to assess purity at each step. Protein purity is judged from a structural and/or functional perspective, depending on the nature of the target protein and its intended use. If the intention is to examine the function of the target molecule, the activity of the target protein must be maintained during the purification process. A functional assessment of the purity of the target protein will thus be required at each step. If the intention is to examine the structure of the target molecule, structure must be maintained, but function may or may not have to be preserved. A structural assessment of purity will, however, be required in this case. If struc-
tural/functional relationships involving the target molecule are to be examined, assessment of both structural and functional purity will be required. Methods for assessing protein purity must quantitate the amount of target protein relative to the amount of contaminant(s) in the sample. This requires two separate analytical methods: one for the target protein and one for the total protein including contaminants. For example, with enzymatic proteins, assessment of functional purity typically relies on kinetic assays in which the amount of substrate consumed or product produced per unit time is proportional to the amount of enzyme present. The total activity of the sample is calculated and compared to the total amount of protein to give the specific activity of the sample. Methods for determining total protein are discussed in Chapter 3. An increase in specific activity at a purification step reflects a loss of contaminant proteins. Functional purity of structural proteins, denatured proteins (i.e., proteins that have lost their function during the purification procedure), and proteins for which a functional assay does not exist must usually be assessed by measuring structural attribute(s)—e.g., molecular weight, pI, presence of a metal ion or cofactor, or presence of an antibody binding
8.1.2 Current Protocols in Protein Science
Table 8.1.1
Purification Table for an Arbitrary Enzymea
Purification step
Total activity (U)
Crude homogenate 12,000 × g supernatant (NH4)2SO4 fraction (20-50%) HIC AEX Gel filtration
200 180 150 125 208 148
Total protein Specific Yield (% total Step Purification (mg) activity (U/mg) activity) yieldb factorc 50,000 35,000 25,000 2500 29.3 5.9
0.004 0.005 0.006 0.05 7.1 25.1
100 90 75
— 90 83
1.00 1.25 1.50
62.5 104 74
83 166 71
12.50 1775 6275
aAbbreviations: HIC, hydrophobic-interaction chromatography; AEX, anion-exchange chromatography. bPercentage activity relative to previous step. cIncrease in specific activity.
site. As in the case of functional purity, an increase in the amount of target protein relative to total protein reflects an increase in purity.
STEPS IN A PURIFICATION STRATEGY A purification strategy for any protein at any scale of operation can be broadly divided into three sequential stages—capture, intermediate purification, and polishing. Each stage represents a set of specific problems that may be encountered during a purification process. The nature of the sample and the scale of operation will dictate what equipment and methodology are appropriate to solve the problem.
Capture Stage The capture stage is the initial purification of the target protein from the source material. The goal of capture is to concentrate the target protein while removing as much of the major contaminant(s) as possible; to this end, an adsorptive chromatographic technique should be employed. Ion-exchange chromatography (UNIT 8.2) and hydrophobic-interaction chromatography (HIC; UNIT 8.4) are generally the best chromatographic techniques for capture at any scale of operation. Certain affinity techniques are also effective, including lectin affinity chromatography (UNIT 9.1), dye affinity chromatography (UNIT 9.2), immunoaffinity chromatography, and metal-chelate affinity chromatography (MCAC). Affinity chromatography is very useful for capture on a small scale (e.g., <1 g). Larger-scale capture using affinity-based methods is generally avoided because of the cost of the chromatographic media and reagents (especially for immunoaffinity chromatography).
At the capture stage the sample is very crude. It may be turbid from the presence of cellular debris or viscous from the presence of DNA, and will typically represent the largest volume to be processed during the purification procedure. Water is often the major contaminant, especially in samples from cell-culture fluids or fermentation broth. The sample may also contain pigments, particulates, and lipophilic materials that can easily clog chromatography columns or bind nonspecifically to certain media. Once such contaminants are introduced into a chromatography system, they can be very difficult to remove. The sample may also contain proteinases or other agents that can potentially destroy or denature the target protein. Ion-exchange or HIC media suitable for capture must allow high throughput of dilute, turbid, or viscous samples. This is achieved by using bead diameters of 40 to 300 µm. Columns packed with such relatively large beads can usually be operated at lower pressures and higher flow rates, and are less prone to fouling or clogging from particulates in the sample. Higher flow rates reduce the time required to handle large volumes, which in turn reduces the potential for product loss from degradation or denaturation. When the volume of crude sample is very large (or larger than can be reasonably handled with available pumps or columns), batch techniques (UNIT 8.2) can be used at the capture stage. In batch adsorption of the target protein by ion exchange, HIC, or affinity chromatography, the chromatography medium is mixed directly with the sample in a suitable vessel. When the target protein is adsorbed, the medium is separated from the sample by filtration, centrifuga-
Conventional Chromatographic Separations
8.1.3 Current Protocols in Protein Science
tion, or settling and decantation. The target protein is then eluted from the medium and collected. For both column and batch techniques, elution of the target protein at the capture stage is often achieved using simple step gradients, but continuous gradients may be used to achieve higher resolution with column techniques. Specific medium requirements and methodologies for use at the capture stage are presented in UNIT 8.2 for ion-exchange chromatography and in UNIT 8.4 for HIC.
Intermediate Purification Stage
Overview of Conventional Chromatography
The next stage in the purification process is intermediate purification. Once the solution containing the target protein has been clarified and concentrated, only small amounts of other types of biomolecules (e.g., lipids and nucleic acids) should be present. The target protein and remaining protein contaminants have a functional or structural attribute in common, whose nature depends on the technique that was used at the capture stage. If ion exchange was used, the contaminants and target protein will be similar in charge characteristics; if HIC was used, similar in hydrophobicity; if affinity methods were used, similar in a ligand- or antibody-binding site, and if ultrafiltration or gel filtration was used, similar in size. Because the contaminants have a physicochemical similarity to the target protein, it is best to exploit a different attribute of the target protein or contaminant during the intermediate purification stage than was used during the capture stage. Use of the same chromatographic technique for both capture and intermediate purification is not recommended. The combination of ion exchange and HIC is very effective in most purification strategies. Proteins are eluted from HIC media at low ionic strength, which is ideal for direct application of the sample to an ion-exchange column, where binding occurs at low ionic strength. Conversely, proteins are eluted from ion-exchange media with increasing ionic strength, which is ideal for subsequent direct application of the sample to a HIC column, where binding occurs at high ionic strength. Cation and anion exchange in combination can also be very effective. The reduced sample volume in the intermediate purification stage also allows use of negative adsorption chromatography, which is designed to bind contaminants to the column, allowing the target protein to pass through. Chromatography techniques used in intermediate purification should resolve the target protein from contaminants on the basis of small
differences in a single physicochemical attribute. High resolution and high yield are the primary goals of intermediate purification. To achieve these goals, a chromatography medium of smaller bead size than that used at the capture stage should be selected. The advantage of smaller bead diameters is increased resolution resulting from decreased peak width. However, smaller bead diameters typically require higher operating pressures. To further enhance resolution during intermediate purification, elution of the target protein is often done using a linear gradient. Specific medium requirements and methodologies for use at the intermediate purification stage are presented in UNIT 8.2 for ion-exchange chromatography and in UNIT 8.4 for HIC.
Polishing Stage Polishing is the final stage in the purification process. The goal of polishing is to remove structural and functional variants of the target protein, as well as any trace contaminants. After polishing, the target protein should be in a suitable form for its intended use. The contaminants and structural variants that must be removed during polishing share very similar physicochemical attributes with the target protein. Chromatographic techniques employed in the polishing stage should exploit a different physicochemical attribute of the target protein than those used for capture and intermediate purification. The similarity of contaminants to target protein heightens the need for high-resolution techniques during this stage. Chromatofocusing, gel filtration (UNIT 8.3), reversed-phase chromatography (RPC; Corran, 1989), and affinity-based techniques (Chapter 9) are all useful in this stage. RPC is very effective for resolving structural variants of a target protein, but generally leads to loss of activity of the target protein because of the organic solvents employed. Adsorptive techniques such as chromatofocusing and affinity chromatography often require special eluants to recover the target protein, which may have to be removed before the purified target protein can be used. Gel filtration does not generally require any special eluants, and is often the best technique for polishing.
Optimizing Purification Throughout the three stages of the purification process, the concentration of target protein is increasing with respect to the concentration of contaminants. In addition, the contaminants to be removed at each stage are increasingly
8.1.4 Current Protocols in Protein Science
peak 1
peak 2
VR2 – VR1 1 (W b2 + Wb1)
A280
Rs =
2
VR1
VR2
Wb1
Wb2 Elution volume
Figure 8.1.2 Hypothetical chromatogram illustrating calculation of elution volumes and peak widths for two peaks. Total elution volume is plotted against absorbance at 280 nm. VR1 and VR2 are the elution volumes (at the peak maxima) of peaks 1 and 2, respectively; Wb1 and Wb2 are the peak widths (at base) of peaks 1 and 2, respectively. Peak widths at base represent the segment of the baseline intercepted by the tangents drawn to the inflection points of the peak. Resolution (Rs) is defined as the distance between the elution volumes at peak maxima divided by the average of the peak widths, according to Equation 8.1.1 (see discussion of Resolution).
similar to the target protein. This increases the need for higher resolution and better yields at each progressive step. It is desirable to examine the efficacy of several techniques at each stage of the procedure before deciding on a specific one to use. Once a specific chromatographic technique has been chosen, the methodology for that step should be optimized: first to achieve maximum resolution, then maximum capacity and yield, and finally maximum speed. This approach will result in a fast, efficient, economical, and robust purification strategy.
PARAMETERS AFFECTING CHROMATOGRAPHIC RESOLUTION Resolution Resolution is a measure of the relative separation achieved between two chromatographically distinct materials, and maximum resolution is the primary goal of any purification step. This discussion covers the main theoretical parameters that affect ability to resolve components of a protein sample into chromatographically distinct forms, thereby providing a basis on which to judge chromatographic results. The main parameters affecting resolution are selec-
tivity, efficiency, and capacity. More detailed discussions are found in Giddings and Keller (1965) and Janson and Ryden (1989). The result of any chromatographic separation is often expressed as the resolution between the zones or peaks containing the chromatographically distinct sample components. The resolution (Rs) of two peaks obtained in a chromatography step may be determined from the chromatogram shown in Figure 8.1.2 according to Equation 8.1.1, where VR2 is the elution volume of peak 2, VR1 is the elution volume of peak 1, Wb1 is the width of peak 1, and Wb2 is the width of peak 2. In cases where a constant eluant flow is maintained and fractions are collected at regular time intervals, VR1, VR2, Wb2, and Wb1 may be expressed in units of time rather than volume. It is necessary that the same units (time or volume) be used to express all terms in the equation to give a dimensionless value to Rs. Rs =
VR2 − VR1 1 ( Wb2 + Wb1 ) 2
Equation 8.1.1
Conventional Chromatographic Separations
8.1.5 Current Protocols in Protein Science
Resolution values can be used to determine if further optimization of a chromatographic procedure is required, as they provide a numerical indication of whether two peaks have been completely resolved. Figure 8.1.3 compares two separation results: one with completely resolved peaks and the other with incompletely resolved
peaks. It should be kept in mind that a completely resolved peak is not necessarily a pure substance. A single peak frequently represents multiple substances that are not resolved by the particular chromatographic technique used. The resolution achievable in any chromatographic system is proportional to the capacity,
A peak 1
peak 2
A280
RS = 1.0
98% peak 1
98% peak 2
2% peak 2 2% peak 1 Elution volume
B peak 1
peak 2
A280
RS = 1.5
~100% peak 1
~100% peak 2
Elution volume
Overview of Conventional Chromatography
Figure 8.1.3 Separation results with different resolutions. (A) When Rs = 1.0, 98% purity has been achieved—i.e., 98% of the fraction collected at peak 1 consists of a single protein with the remaining 2% being contamination from the protein collected at peak 2; conversely, 98% of the fraction collected at peak 2 consists of a single protein with the remaining 2% being contamination from peak 1. (B) Complete (baseline) resolution requires that Rs >1.5. At this value, the peaks are completely separated from each other (i.e., the purity of each peak is >99.9%).
8.1.6 Current Protocols in Protein Science
efficiency, and selectivity of the system. Each of these factors must be considered and controlled to achieve success. The theoretical expression for resolution is Equation 8.1.2, where k is the average capacity factor for the two peaks, N is the efficiency factor for the system, and α is the selectivity factor of the medium. Rs =
k=
VR2 − Vm Vm
Equation 8.1.3
In Equation 8.1.2 for Rs, presented in the discussion of Resolution, k is the average of k1 (capacity factor for peak 1) and k2 (capacity factor for peak 2). Adsorption techniques such as ion exchange, HIC, chromatofocusing, RPC, and affinity chromatography can have high capacity factors because experimental conditions can be manipulated so that the elution volume for a peak can exceed the total bed volume (Vm as is the case with peaks 2 and 3 in Fig. 8.1.4.) However, in gel filtration, which is a nonadsorptive technique, all peaks must elute within the volume Vm−Vo as is the case with peak 1 in Fig. 8.1.4. The impact of experimental conditions on capacity at various stages of protein purification are discussed for different purification techniques in subsequent units of Chapter 8.
α−1 k 1 ) ( N−2 ) ( ) ( 4 1+k α Equation 8.1.2
The impact of various experimental factors on resolution in various stages of protein purification is discussed for specific chromatographic techniques in subsequent units of Chapter 8.
Capacity The capacity or retention factor (k) is a measure of retention of a sample component. It should not be confused with the loading capacity of a column, which is expressed as milligrams of sample bound per milliliter of gel and represented by the area under a peak. The capacity factor may be calculated for any individual peak in a chromatogram. For example, the capacity factor for peak 2 in Figure 8.1.4 is derived from Equation 8.1.3, where VR2 is the elution volume of peak 2 and Vm is the volume of the mobile phase (i.e., the total bed volume).
Efficiency The efficiency factor (N) is a measure of zone broadening (peak width) occurring on a column. It can be calculated for any given peak from Equation 8.1.4, where VR is the elution volume (i.e., the total volume of eluant that has
gel filtration
absorptive techniques 2
3
A280
1
V0
Wb1 VR1 VM VR2 VR3 Elution volume
Figure 8.1.4 Hypothetical chromatogram. Vo = void volume; VR1 = elution volume for peak 1; VR2 = elution volume for peak 2; VR3 = elution volume for peak 3; VM = volume of mobile phase; Wb1 = peak width for peak 1; Wb2 = peak width for peak 2.
Conventional Chromatographic Separations
8.1.7 Current Protocols in Protein Science
A
A280
high efficiency
low efficiency
B
A280
high efficiency
low efficiency
Elution volume
Figure 8.1.5 Effect of selectivity and efficiency on resolution. (A) Chromatogram obtained using experimental technique that provides good selectivity. Where conditions are chosen to provide high efficiency, two narrow and distinct peaks are obtained (high resolution). When the conditions provide only low efficiency, the peaks are broadened and overlap more (lower resolution). (B) Chromatogram obtained using experimental technique that provides bad selectivity. Even when high-efficiency conditions are used, there is lower resolution than in (A), and the resolution deteriorates further with low-efficiency conditions.
passed through the column at peak maximum) and Wh is the peak width at half the peak height. VR N = 5.54 W h
2
Equation 8.1.4
Overview of Conventional Chromatography
Efficiency (N) may be expressed as the number of theoretical plates for the column under specific experimental conditions (see UNIT 8.3).
Efficiency is also frequently defined as the number of plates per meter of chromatographic bed, or in terms of H, the height equivalent to a theoretical plate. H is simply the column length (L) divided by the efficiency factor (N); i.e., H = L/N. The main cause of zone broadening (i.e., loss of efficiency) in a chromatographic bed is diffusion. Diffusion perpendicular to the flow is restricted by the walls of the column. Therefore, longitudinal diffusion is the primary fac-
8.1.8 Current Protocols in Protein Science
tor contributing to zone broadening. While a protein is adsorbed to the medium, little or no diffusion takes place, but once the protein is unbound, diffusion begins. The amount of diffusion that occurs is proportional to the time required for the material to emerge from the system. Loss of efficiency resulting from diffusion is minimized if the distances available for diffusion in the mobile phase, gel beads, and system are minimized. In practice, efficiency increases with increasing uniformity in particle size and with decreasing bead size. Good experimental technique is required for high efficiency. Unevenly packed chromatography beds and trapped air will lead to channeling, zone broadening, and consequent loss of resolution. Loss of efficiency also stems from system effects such as dead volumes in the system, poor mixing during gradient formation, and pulsations in flow.
Selectivity is more important than efficiency (N) in determining resolution because Rs is directly proportional to selectivity, but is proportional only to the square root of efficiency (see Fig. 8.1.5 along with Equation 8.1.1 and Equation 8.1.2 for Rs in the discussion of Resolution). Hence, a four-fold increase in efficiency is required to double resolution, as compared with a two-fold increase in selectivity. In practice, selectivity depends partly on the chromatographic technique employed but can usually be controlled by manipulating experimental conditions, such as the pH and ionic strength of the mobile phase. Because this can be done easily and predictably, selectivity is the factor that is exploited to achieve maximum resolution in column chromatography rather than efficiency, which is fixed by the particle size and uniformity of the medium selected.
LITERATURE CITED Selectivity Selectivity of a chromatographic medium defines the ability of that medium to separate peaks (i.e., it is a measure of the distance between two peaks in a chromatogram; see Fig. 8.1.5). The selectivity factor (α) can be calculated from a chromatogram using Equation 8.1.5, where k2 is the capacity factor for the second peak (see discussion of Capacity), k1 is the capacity factor for the first peak, VR2 is the elution volume of the second peak, VR1 is the elution volume of the first peak (see Fig. 8.1.2), and Vm is the volume of the mobile phase.
α=
k2 VR2 − Vm VR2 = ≈ k1 VR1 − Vm VR1
Corran, P.H. 1989. Reversed-phase chromatography of proteins. In HPLC of Macromolecules: A Practical Approach (R.W.A. Oliver, ed.) pp. 127156. IRL Press, Oxford. Giddings, J.C. and Keller, R.A. (eds.) 1965. Dynamics of Chromatography, Part 1: Principles and Theory. Marcel Dekker, New York. Janson, J.-C. and Ryden, L. (eds.) 1989. Protein Purification: Principles, High Resolution Methods and Applications. VCH Publishers, New York.
Contributed by Alan Williams Pharmacia Biotech Piscataway, New Jersey
Equation 8.1.5
Conventional Chromatographic Separations
8.1.9 Current Protocols in Protein Science
Ion-Exchange Chromatography
UNIT 8.2
Ion-exchange chromatography separates biomolecules on the basis of charge characteristics. Charged groups on the surface of a protein interact with oppositely charged groups immobilized on the ion-exchange medium. As illustrated in Figure 8.2.1, the charge of a protein depends on the pH of its environment (the operating pH). The pH at which the net charge of a protein is zero (i.e., where the number of positive charges equals the number of negative charges) is known as the isoelectric point (pI). When the operating pH is greater than the pI, the protein will have a net negative charge, and should bind to anion-exchange media, which are positively charged. When the operating pH is less than the pI, the protein will have a net positive charge, and should bind to cation-exchange media, which are negatively charged. The Strategic Planning section outlines the basic steps in planning and carrying out ion-exchange chromatography to separate proteins. Basic Protocol 1 describes batch adsorption of protein to an ion-exchange medium followed by elution using a step gradient of increasing salt concentration. This technique accommodates a wide range of sample volumes and is most often used in the initial capture stage of protein purification (UNIT 8.1). Batch techniques have minimal system requirements. The Alternate Protocol describes use of a buffer of a different pH to elute via step gradient. Basic Protocol 2 describes adsorption of protein to an ion-exchange medium in a column, followed by elution with a linear gradient. Basic Protocol 2 provides a higher resolution than Basic Protocol 1, and is therefore used in the intermediate purification and final polishing stages of protein separation (UNIT 8.1). Support Protocol 1 describes a pilot experiment to determine initial conditions for batch or column chromatography (i.e., pH required for binding, change in pH or salt concentration required for elution, and available capacity of a medium). Support Protocol 2 describes a means of calculating the dynamic capacity (UNIT 8.1) of an ion-exchange column, Support Protocol 3 describes methods for producing continuous gradients of pH and salt concentration to elute proteins from ion-exchange
+ Net positive charge of protein isoelectric point (pl)
Will bind to cation exchanger
select system pH above pI for anion exchange
0
system pH 4
6
8
10
select system pH below pI for cation exchange Will bind to anion exchanger
– Net negative charge of protein
Figure 8.2.1 Net charge of a protein as a function of pH, showing the pH ranges in which protein is bound to anion or cation exchangers. The pH range over which the protein is stable may be only a small fraction of the binding range; this must also be taken into consideration when choosing an ion-exchange medium. Contributed by Alan Williams and Verna Frasca Current Protocols in Protein Science (1999) 8.2.1-8.2.30 Copyright © 1999 by John Wiley & Sons, Inc.
Conventional Chromatographic Separations
8.2.1 Supplement 15
columns, Support Protocol 4 describes regeneration of used ion-exchange media, and Support Protocol 5 details storage of ion-exchange media. STRATEGIC PLANNING There are a number of basic steps in developing an ion-exchange method for protein purification. First, an appropriate ion-exchange medium must be selected, as well as the optimal operating pH and buffer system for the medium and sample (see following discussions on Selecting an Ion-Exchange Medium and Selecting a Buffer System). It is then necessary to decide whether batch or column chromatography is appropriate given the purity, protein concentration, and physical characteristics of the sample, the sample volume to be used, and the availability of suitable equipment (see following discussion on Selecting Batch Versus Column Purification). Next, pilot experiments are conducted to determine conditions for binding and eluting the protein (see Support Protocol 1). It is also necessary to make sure that the capacity of the medium is sufficient to isolate the desired quantity of protein (see Support Protocol 1 and Support Protocol 2). When these initial conditions have been determined, the medium and sample are prepared, the sample is bound to the medium, unbound sample components are washed away, and bound sample components are selectively eluted and collected (see Basic Protocol 1, Basic Protocol 2, or Alternate Protocol). The medium may then be cleaned and regenerated (see Support Protocol 5). Finally, results are evaluated and conditions optimized if necessary (see Critical Parameters). Selecting an Ion-Exchange Medium A wide variety of ion-exchange media are commercially available, but no miracle medium exists that is best for every protein purification. Criteria for selecting an ion-exchange medium include the specific requirements of the application, the pI and molecular size of the sample components (i.e., target protein and contaminants), and the available equipment (e.g., pumps and columns). It is necessary to begin by selecting either anion or cation exchange. This requires knowledge of the pI and pH stability of the target protein. If the pI of the target protein is known, an anion-exchange medium with an operating pH above the pI of the target protein or a cation-exchange medium with an operating pH below the pI of the target protein should be selected. If the pI of the target protein is unknown, it is desirable to determine it before beginning. The optimal operating pH can be determined empirically (see Support Protocol 1). Because the pI for most proteins is below pH 7 (Gianazza and Righetti, 1980), it is reasonable to select an anion-exchange medium and an operating pH of 8.5 to start, then evaluate the results and optimize conditions as necessary. It is also useful to know the pI and binding characteristics of the contaminants present in the protein solution. For example, if the pH of the binding buffer is higher than the pI of a major contaminant, that contaminant will not bind to an anion-exchange medium. If the starting material is crude cell lysate, DNA will be present and generally needs to be removed from the protein. Since DNA is anionic, it binds tightly to anion-exchange media and typically is not eluted with the salt and pH conditions used during protein purification. Selecting a Buffer System A buffer system must be selected for the desired pH range. As with selection of an ion-exchange medium, there are several factors that must be considered in selecting a buffer system, including the type of ion exchange to be performed, the pH stability of the sample and the pH range to be used, the required buffering capacity, and, finally, the cost. Ion-Exchange Chromatography
8.2.2 Supplement 15
Current Protocols in Protein Science
Table 8.2.1
Buffers for Ion-Exchange Chromatographya
pKa (25°C)
pH range
Bufferb
Anion exchange 4.75 5.68 5.96 6.46 6.80 7.76 8.06 8.52 8.88 8.64 9.50 9.73 10.47 11.12
4.5-5.0 5.0-6.0 5.5-6.0 5.8-6.4 6.4-7.3 7.3-7.7 7.6-8.5 8.0-8.5 8.4-8.8 8.5-9.0 9.0-9.5 9.5-9.8 9.8-10.3 10.6-11.6
N-Methylpiperazine Piperazine L-Histidine bis-Tris bis-Tris propane Triethanolamine Tris⋅Cl N-Methyldiethanolamine Diethanolamine 1,3-Diaminopropane Ethanolamine Piperazine 1,3-Diaminopropane Piperidine
Cation exchange 2.00 2.88 3.13 3.81 3.75 4.21 4.76 5.68 7.20 7.55 8.35
1.5-2.5 2.4-3.4 2.6-3.6 3.6-4.3 3.8-4.3 4.3-4.8 4.8-5.2 5.0-6.0 6.7-7.6 7.6-8.2 8.2-8.7
Maleic acid Malonic acid Citric acid Lactic acid Formic acid Butanedioic acid Acetic acid Malonic acid Phosphate HEPES BICINE
Working Temperature concentration (mM) factorc 20 20 20 20 20 20 20 50 20 (pH 8.4) 20 20 20 20 20 20 20 20 50 50 50 50 50 50 50 50
−0.015 −0.015 −0.017 −0.02 −0.028 −0.028 −0.025 −0.031 −0.029 −0.026 −0.026 −0.031
−0.0024 +0.0002 −0.0018 +0.0002 −0.0028 −0.0140 −0.0180
aInformation from Pharmacia Biotech (1995). bAbbreviations: BICINE, N,N-bis[2-hydroxyethylglycine; bis-Tris, bis[2-hydroxyethyl]iminotris[hydroxymethyl]methane; bis-Tris propane, 1,3-bis[tris(hydroxymethyl)methylamino]-propane; HEPES, N-[2-hydroxyethyl]piperazine-N′-[2ethanesulfonic acid]. cChange in pK per °C (i.e., ∂pK /∂T). a a
Anionic buffers (e.g., acetate and phosphate) are preferred for cation exchange and cationic buffers (e.g., Tris⋅Cl, ethanolamine, and piperazine) are preferred for anion exchange. It is important to ensure that the buffering ion will have the same charge as the ion exchanger, and hence will not be bound. A constant buffering capacity and pH will thus be maintained during the ion-exchange experiment. Table 8.2.1 lists a variety of buffers useful for anion and cation exchange. Additives to be included in the buffers (e.g., detergents or protease inhibitors) should also carry the same charge as the ion-exchange medium, to preclude binding. Selecting Batch Versus Column Purification If the sample volume is large in relation to the size of the pumps and columns available in the laboratory, batch adsorption techniques (see Basic Protocol 1) are appropriate for the capture stage of purification. Batch methods, in which the sample and medium are directly mixed without use of a column, are employed to reduce the sample volume and
Conventional Chromatographic Separations
8.2.3 Current Protocols in Protein Science
Supplement 15
Table 8.2.2
Ion-Exchange Media for Batch Adsorption
Medium
Type
Suppliera
Dextran bead matrix QAE Sephadex A-25 QAE Sephadex A-50 SP Sephadex C-25 SP Sephadex C-50
Strong anion Strong anion Strong cation Strong cation
Amersham Pharmacia Biotech Amersham Pharmacia Biotech Amersham Pharmacia Biotech Amersham Pharmacia Biotech
Microgranular cellulose matrix QA-52 Strong anion QA-53 Strong anion SE-52 Strong cation SE-53 Strong cation
Whatman Whatman Whatman Whatman
Polymer-coated silica matrix Accell Plus QMA Strong anion Accell Plus CM Weak cation
Waters Waters
Agarose bead matrix DEAE Bio-Gel A CM Bio-Gel A
Bio-Rad Bio-Rad
Weak anion Weak cation
aFor addresses and telephone numbers of suppliers, see SUPPLIERS APPENDIX.
total protein content prior to column chromatography. Batch techniques are also very useful for capture of the target protein from extremely crude samples (e.g., cell lysates and samples containing particulates and/or aggregates). In batch methods the flow and packing characteristics of the medium are of minor importance, but economy and capacity are of primary importance. Ion-exchange media suitable for batch purification at the capture stage of protein purification are presented in Table 8.2.2. High resolution is required in the intermediate purification and polishing stages of protein purification, and this can only be achieved using column chromatography. However, higher-than-necessary resolution is frequently traded off for greater throughput (i.e., amount of material processed in a defined time). The packing and flow characteristics of the medium are of primary importance when using column chromatography. Columns packed with smaller beads usually require higher operating pressures than columns packed with larger beads. However, smaller beads offer higher resolution, resulting from the increased efficiency that comes with decreasing bead size. Larger beads are desirable for applications requiring higher throughput. A medium with the smallest bead size that will provide the necessary throughput (and whose operating pressure can be accommodated by available equipment) should be selected. Table 8.2.3 lists a variety of media suitable for column purification at various stages in the purification process. See SUPPLIERS APPENDIX for contact information of suppliers of these media, all of whom provide technical support for selection and use of specific media. Capacity of Ion-Exchange Media
Ion-Exchange Chromatography
The amount of ion-exchange medium required for a particular application will be determined by the protein capacity of the medium (i.e., the amount of protein that can be bound to the matrix) at the chosen pH. The amount of medium required will in turn determine the size of column required. The capacity of an ion-exchange medium is dependent on a variety of factors, including charge and molecular size of the components in the sample and experimental conditions employed.
8.2.4 Supplement 15
Current Protocols in Protein Science
Conventional Chromatographic Separations
8.2.5
Current Protocols in Protein Science
Supplement 15
34 90 90 200 200
PS/DVB
Agarose Agarose Agarose Agarose
Agarose
Q Sepharose High Performance SP Sepharose High Performance Q Sepharose Fast Flow
SP Sepharose Fast Flow
SP Sepharose Big Beads Agarose Agarose
SOURCE 15S
STREAMLINE DEAE
STREAMLINE SP
Hi-Trap Qc HiTrap SPd
34
PS/DVB
SOURCE 15Q
200
15
15
10
PS/DVB
Mono S
Avg. bead diameter (µm)
10
Matrix
B/C
B/C
C
C/I
C/I
C/I/P
C/I/P
I/P
I/P
I/P
I/P
Use
BU
BU
BU
PP/BU
PP/BU
PP/BU
PP/BU
PP/BU
PP/BU
PP
PP
Form available
Ion-Exchange Media for Column Purificationa
Amersham Pharmacia Biotech Mono Q PS/DVB
Mediumb
Table 8.2.3
170
205
210
215
215
170
170
ND
ND
160
320
Ionic capacity (meq/ml)
—
—
—
—
—
—
—
—
—
—
—
Available capacity (mg/ml)
70
55
80
50
120
90
108
85
54
75
65
Dynamic capacity (mg/ml)
continued
4.5 mg/ml BSA; 50 mM Tris⋅Cl, pH 8.0; 100 cm/hr; 0.5 × 5–cm column Human IgG; 100 mM sodium acetate, pH 5.0; 300 cm/hr; 0.5 × 5–cm column 4.5 mg/ml BSA; 50 mM Tris⋅Cl, pH 8.0; 100 cm/hr; 0.5 × 5–cm column 55 mg/ml lysozyme; 20 mM phosphate, pH 6.8; 1 ml/min; 0.63 × 3–cm column 4.5 mg/ml BSA; 5 mM Tris⋅Cl, pH 8.0; 100 cm/hr; 0.5 × 5–cm column RNase; 100 mM sodium acetate, pH 5.0; 150 cm/hr; 0.5 × 5–cm column Human serum albumin; 50 mM Tris⋅Cl, pH 8.3 Human IgG; 100 mM sodium acetate, pH 5.0; 300 cm/hr; 0.5 × 5–cm column 2 mg/ml BSA; 100 mM sodium acetate, pH 5.0; 300 cm/hr BSA; 50 mM Tris⋅Cl, pH 7.5; 300 cm/hr; 5-cm-i.d. expanded-bed column Lysozyme; 50 mM phosphate, pH 7.5, 300 cm/hr; 5-cm-i.d. expanded-bed column
Conditions for determining dynamic capacity
Ion-Exchange Chromatography
8.2.6
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PM
PM PM
Macro-Prep high S
Macro-Prep high Q
Macro-Prep high S
65 35 100 65 35
PM PM PM PM PM PM PM PM PM PM PM
Waters Protein-Pak Q HR Protein-Pak SP HR Protein-Pak Q HR Protein-Pak SP HR Protein-Pak Q HR Protein-Pak SP HR 8 8 15 15 40 40
100 100 100
PM PM PM
50
50
10
10
Avg. bead diameter (µm)
TosoHaas Toyopearl QAE-550C Toyopearl SP-550C Toyopearl SUPER Q-650C Toyopearl SUPER Q-650M Toyopearl SUPER Q-650S Toyopearl SP-650C Toyopearl SP-650M Toyopearl SP-650S
Econo-Pac high Qe Econo-Pac high Sf
PM
Matrix
P P I/P I/P B/C/I B/C/I
C/I C/I/P I/P
I/P
C/I/P
C/I C/I C/I
B/C/I
B/C/I
I/P
I/P
Use
PP PP PP/BU PP/BU BU BU
PP/BU PP/BU PP/BU
PP/BU
PP/BU
PP/BU PP/BU PP/BU
P/B
PP/BU
PP/BU
PP/BU
Form available
Ion-Exchange Media for Column Purificationa, continued
Bio-Rad Macro-Prep high Q
Mediumb
Table 8.2.3
200 225 200 225 200 225
150 150 150
240
240
330 160 280
160
400
127
115
Ionic capacity (meq/ml)
ND ND ND ND
126g 45h 50h 50h
65 40 75 40 60 20
ND
143g
— — — — — —
ND ND ND
60
40
50
20
Dynamic capacity (mg/ml)
70g 110h 129g
—
—
—
—
Available capacity (mg/ml)
continued
BSA; 20 mM Tris⋅Cl, pH 8.2 Cytochrome c; 25 mM MES, pH 5.0 BSA; 20 mM Tris⋅Cl, pH 8.2 Cytochrome c; 25 mM MES, pH 5.0 BSA; 20 mM Tris⋅Cl, pH 8.2 Cytochrome c; 25 mM MES, pH 5.0
— — —
—
—
— — —
5 mg/ml BSA; 50 mM Tris⋅Cl, pH 8.3; 100 cm/hr; 1.5 × 10–cm column 5 mg/ml human IgG; 20 mM sodium acetate, pH 5.0; 100 cm/hr; 1.5 × 10–cm column 5 mg/ml BSA; 50 mM Tris⋅Cl, pH 8.3; 100 cm/hr; 1.5 × 10–cm column 5 mg/ml human IgG; 20 mM sodium acetate, pH 5.0; 100 cm/hr; 1.5 × 10–cm column
Conditions for determining dynamic capacity
Conventional Chromatographic Separations
8.2.7
Current Protocols in Protein Science
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CP
CP
CP
CP
CP
CP
CP
CP
BioSepra Q-Hyper D
S-Hyper D
Q-Hyper D
S-Hyper D
Q-Hyper D
S-Hyper D
Q-Hyper D
S-Hyper D
60
60
35
35
20
20
10
10
Avg. bead diameter (µm)
C/I
C/I
I/P
I/P
I/P
I/P
P
P
Use
BU
BU
PP/BU
PP/BU
PP/BU
PP/BU
PP
PP
Form available
100
200
100
250
100
250
110
250
Ionic capacity (meq/ml)
—
—
—
—
—
—
—
—
Available capacity (mg/ml)
100
100
100
40
105
90
105
90
Dynamic capacity (mg/ml)
5 mg/ml BSA; 50 mM Tris⋅Cl, pH 8.6; 150 cm/hr; determined at 10% breakthrough 5 mg/ml lysozyme; 50 mM acetate, pH 4.5; 150 cm/hr; determined at 10% breakthrough 5 mg/ml BSA; 50 mM Tris⋅Cl, pH 8.6; 150 cm/hr; determined at 10% breakthrough 5 mg/ml lysozyme; 50 mM acetate, pH 4.5; 150 cm/hr; determined at 10% breakthrough 5 mg/ml BSA; 50 mM Tris⋅Cl, pH 8.6; 150 cm/hr; determined at 10% breakthrough 5 mg/ml lysozyme; 50 mM acetate, pH 4.5; 150 cm/hr; determined at 10% breakthrough 5 mg/ml BSA; 50 mM Tris⋅Cl, pH 8.6; 150 cm/hr; determined at 10% breakthrough 5 mg/ml lysozyme; 50 mM acetate, pH 4.5; 150 cm/hr; determined at 10% breakthrough
Conditions for determining dynamic capacity
hConditions for determining available capacity: 5 mg/ml lysozyme; 20 mM phosphate, pH 6.0.
gConditions for determining available capacity: 5 mg/ml BSA; 50 mM Tris⋅Cl, pH 8.7.
f Prepacked column suitable for use with syringe. For medium specification, see Macro-Prep high S.
e Prepacked column suitable for use with syringe. For medium specification, see Macro-Prep high Q.
d Prepacked column suitable for use with syringe. For medium specification, see SP Sepharose High Performance.
c Prepacked column suitable for use with syringe. For medium specification, see Q Sepharose High Performance.
bFor addresses and telephone numbers of suppliers, see SUPPLIERS APPENDIX.
aAbbreviations: B, batch; BSA, bovine serum albumin; BU, bulk; C, capture; CP, composite polymer, I, intermediate purification; i.d., inner diameter; MES, 2-[N-morpholino]ethanesulfonic acid; ND, not determined; P, polishing; PM, polymethacrylate; PP, prepacked; PS/DVB, polystyrene/divinylbenzene.
Matrix
Ion-Exchange Media for Column Purificationa, continued
Mediumb
Table 8.2.3
The number of charged groups on a specified amount of matrix that can participate in ion exchange is termed ionic or total capacity. However, ion-exchange media are porous and the ion-exchange groups are usually distributed throughout the bead. The amount of protein that will bind is dependent on the ability of the sample components to diffuse into the bead, and this ability is related to the molecular weight and shape of the components and the pore structure of the matrix (see UNIT 8.3). In addition, large molecules such as proteins and nucleic acids can bind to multiple ion-exchange groups (or sterically hinder other molecules from binding) depending on the shape and surface-charge distribution of the protein, as well as the spatial orientation of the exchange ligands on the matrix. These factors are taken into account in the available and dynamic capacity of a matrix or column. Available capacity is defined as the amount of a specific protein that will bind to an ion-exchange medium at a defined pH, salt concentration, and sample concentration in a batch purification process. When medium is packed in a column, the amount of a specific protein that will bind is dependent on many of these factors, as well as on the column dimensions and flow rate. The capacity of a column operating under defined conditions is termed the dynamic capacity. The dynamic capacity is typically less than the available capacity for the same sample under the same operating conditions. Ion-exchange media in which the charge state of the ligand is sensitive to the operating pH are termed weak ion exchangers. The usable pH range of weak ion exchangers is narrower than that of strong ion-exchange media, in which the charge on the ligand used for exchange is constant (i.e., insensitive to pH). Ion-exchange media with DEAE (diethylaminoethyl) or CM (carboxymethyl) groups as the ligand for exchange are considered weak ion exchangers. Because the ionic capacity of weak ion-exchange media is variable, strong ion-exchange media are recommended for pilot experiments (see Support Protocol 1). When selecting ion-exchange media for batch or column purifications, the amount of medium should be estimated from the manufacturer-supplied capacity data for a protein similar in size to the target protein to be purified. Be aware of what type of capacity (i.e., ionic, available, or dynamic) is being reported and the experimental conditions under which the reported capacity was determined (see Table 8.2.3). A general recommendation is to begin pilot experiments at ∼10% of the published capacity data, then determine the capacity for your system. Preparing Solutions All aqueous solutions for use in ion-exchange chromatography should be prepared with distilled or deionized water (Milli-Q purified or equivalent). Solutions and samples for column chromatography should be filtered, degassed, and equilibrated to the appropriate temperature prior to use. Special attention must be given to any additives to be included in the buffer to be sure that they will not bind to the column (see preceding discussion on Selecting a Buffer System). If the chosen additives increase the viscosity of the mobile phase, the flow rate should be decreased. Chromatography Systems
Ion-Exchange Chromatography
Column techniques typically result in higher resolution than batch techniques, and should be used during the intermediate purification and polishing stages (UNIT 8.1) of a purification procedure. However, to achieve high resolution, an appropriate chromatography system is required. A chromatography system may be described simply by the functional operations required to perform a chromatographic separation. The minimum functions required (Fig. 8.2.2) include a buffer delivery system capable of gradient formation, a mechanism for introducing sample into the system, the column itself (properly packed with the appro-
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Table 8.2.4 Scale-Up Calculations for Ion-Exchange Column Chromatography
Scale
1×
10×
100×
Sample Sample concentration (mg/ml) Sample load (mg) Sample volume (ml)
1 10 10
1 100 100
1 1000 1000
Column Column diameter (cm) Cross-sectional area (cm2) Bed height (cm) Bed volume (ml)
0.5 0.2 5 1
1.6 2 5 10
5 20 5 100
Flow rate Volumetric flow (ml/min)a Linear velocity (cm/hr)
2 300
20 300
200 300
Gradient Gradient volume (ml)
20
200
2000
aLinear flow (cm/hr) multipled by the cross-sectional area of the column (cm2) equals the volumetric flow rate (ml/hr); cm/hr × cm2 = cm3/hr = ml/hr.
priate ion exchanger), an in-line UV monitor suitable for use at 280 nm, a chart recorder, and a fraction collector to preserve the separation. Additional unit operations may include system controllers for automated operation as well as data-acquisition systems. The equipment required to perform each of these unit operations is ultimately determined by the chromatography medium and column selected. Specifically, the operating pressure and flow characteristics of the packed column will be the most important factors for selecting equipment to fit the column, or for selecting columns to fit existing equipment. It is highly recommended that prepacked columns be used and that the manufacturer be consulted prior to purchase as to equipment requirements for proper operation. This will save time and money and increase the probability of success. If columns are to be packed in the laboratory, the manufacturer’s packing instructions for the medium should be followed and column efficiency and function should be tested with representative standards prior to use with the actual sample. There is no single packing methodology that is optimal for all types of media. Table 8.2.3 lists a variety of ion-exchange media available in prepacked columns and/or in bulk for packing in the laboratory. Several manufacturers offer inexpensive disposable ion-exchange columns, suitable for many applications, that can be operated using only a syringe. Scale-Up Conditions Ion-exchange separations are typically developed and optimized on a small scale before the entire sample is committed. It is thus important to choose an ion exchanger that will allow simple and convenient scale-up, so that methods established on a small column can be applied more or less directly to a larger one. Scaling up in ion-exchange chromatography can be approached in two different ways— either the same ion-exchange medium can be used in a larger-diameter column with the same bed height, or a different medium having the same charged group immobilized on a matrix with a higher throughput can be substituted for the medium used in the small-scale separation.
Conventional Chromatographic Separations
8.2.9 Current Protocols in Protein Science
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The former approach, using the same medium throughout, is the simplest. Scaling up a separation with the same medium and bed height used in the small-scale optimization is achieved by employing simple scale factors to adapt the flow, gradient volume, and sample load to the increased volume of a larger-diameter column. The scale factor for the required increase in flow rate is the ratio of the cross-sectional area of the larger column to that of the smaller column (i.e., the factor by which the total flow of solution through the column must be increased so that the same linear velocity used in the small-scale experiment is maintained). The increase in gradient volume and sample loading are directly proportional to the increase in column volume. Table 8.2.4 shows an example of changes that must be made to various parameters for a 10- and 100-fold increase in scale. BASIC PROTOCOL 1
BATCH ADSORPTION AND STEP-GRADIENT ELUTION WITH INCREASING SALT CONCENTRATION This protocol describes adsorption of a protein to an ion-exchange medium by direct mixing of the protein-containing sample and the medium (in this case, an anion-exchange gel). This batch purification is in contrast to column purification (see Basic Protocol 2), in which the gel is packed in a column and the sample is passed through it. Batch adsorption is particularly useful where large volumes of sample are involved, which is usually the case in the capture stage of protein purification (UNIT 8.1). The equipment used here is sufficient for sample volumes up to 1.5 liter. Larger samples may be divided and processed with parallel apparatuses, or processed sequentially on a single apparatus. Different-sized funnels and side-arm flasks may be used when appropriate. Once the starting conditions have been determined (see Support Protocol 1), the range of pH and salt concentration where binding and elution occur is well defined. Conditions for batch adsorption and elution used in this procedure are based on the results, presented in Figure 8.2.3, of a test tube pilot experiment performed as described in Support Protocol 1. Buffer volumes and equipment were chosen on the basis of a sample volume of 1 liter containing 50 to 1000 mg of total protein. Elution of sample components adsorbed to an ion-exchange medium can be achieved either by increasing the salt concentration of the mobile phase, as in this protocol, or by changing its pH (see Alternate Protocol). In batch techniques, discontinuous (step) gradients are used for elution. Column chromatography (see Basic Protocol 2) allows the use of either step or continuous gradients. Formation of continuous gradients requires special equipment (see Basic Protocol 2 and Support Protocol 3). The number of incremental steps to include in a step gradient must be judged empirically based on the resolution achieved. Ideally (as in the procedure described here), three steps should be used in which all weakly bound materials are desorbed and eluted in the first gradient step using wash buffer, the target protein is eluted in the second gradient step using elution buffer, and all strongly adsorbed materials are eluted in the third gradient step using regeneration buffer. A minimum increment of 50 mM NaCl is recommended for experiments using step-gradient elution with increasing salt concentration. Steps in the gradient may be added, omitted, or modified as determined empirically.
Ion-Exchange Chromatography
Materials QAE Sephadex A-25 (Amersham Pharmacia Biotech) or equivalent anion-exchange gel Binding buffer: 20 mM Tris⋅Cl, pH 7.5 (or other buffer as determined empirically; see Support Protocol 1) Protein sample to be purified Wash buffer: 20 mM Tris⋅Cl (pH 7.5)/100 mM NaCl (or other buffer/salt solution as determined empirically; see Support Protocol 1)
8.2.10 Supplement 15
Current Protocols in Protein Science
Elution buffer: 20 mM Tris⋅Cl (pH 7.5)/350 mM NaCl (or other buffer/salt solution as determined empirically; see Support Protocol 1) Regeneration buffer: 20 mM Tris⋅Cl (pH 7.5)/2 M NaCl (also see Support Protocol 5) Boiling water bath (optional) 500-ml sintered-glass filter funnel, medium porosity Three 2000-ml side-arm flasks Conductivity meter Swell and equilibrate ion-exchange gel 1. Add 10 g QAE Sephadex A-25 to 1 liter binding buffer and swell 2 days at room temperature or 2 hr in a boiling water bath. The volume of swollen gel will be ∼70 ml.
2. Mount 500-ml sintered-glass filter funnel on 2000-ml side-arm flask and apply suction. Centrifugation (∼5000 × g, 1 min) may be used instead of filtration to collect the gel, and may be preferred for processing small sample volumes. It is also possible to simply allow the gel to settle under gravity, then decant or aspirate the supernatant; this may be preferable for very large-scale operations.
3. Gently swirl swollen gel to resuspend. Pour gel slurry into funnel as fast as the fluid level in funnel permits, allowing buffer to collect in flask. Continue until all gel has been collected. Release suction after all buffer has been removed. IMPORTANT NOTE: Never use a magnetic stir-bar to resuspend chromatography medium. This can damage beads by grinding them against the bottom of the vessel, resulting in fine particles that can slow the filtration process.
4. Add 200 ml binding buffer to funnel and resuspend gel using a stirring rod. Allow resuspended gel to stand for 5 min, then apply suction to remove buffer. 5. Repeat step 4 at least three times to ensure equilibration. The gel is considered equilibrated when the pH and salt concentration of the eluant are the same as those of the binding buffer. If the procedures here are followed, there is little need to measure pH and salt concentration of the eluant because the gel is swollen and washed in binding buffer and the chance of error is very small. The worst-case scenario is that the target protein will not bind (see Troubleshooting). The repeated washings are more for the removal of fines than for adjustment of pH or salt concentration.
6. Add enough binding buffer (∼100 ml) to produce an ∼50% (v/v) slurry and allow gel to stand in funnel until sample has been prepared. Adsorb sample to gel 7. Adjust pH and salt concentration of protein sample to initial optimal values. For a pilot experiment to determine optimal initial values for pH and salt concentration, see Support Protocol 1. Samples prepared by salt precipitation may require desalting prior to ion exchange (see UNIT 8.3). The sample can be dialyzed against the binding buffer. Also, the salt concentration of a sample can be reduced by adding distilled water until a desired salt concentration is achieved, as measured by a conductivity meter.
8. Combine gel and sample in 2000-ml beaker or wide-mouth flask. The ∼50% gel slurry is swirled and poured from the funnel; transfer may be aided with a rubber policeman, spoon, stirring rod, or wash bottle containing binding buffer.
Conventional Chromatographic Separations
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Supplement 15
9. Gently mix sample and gel by swirling every 15 min or shake on a platform shaker at sufficient rate to maintain gel in suspension. Allow 1 to 2 hr for binding at room temperature or 3 to 4 hr at 4°C. IMPORTANT NOTE: Excessive shaking may result in foaming and possible denaturation of the target protein.
10. Collect gel by filtration as in step 3. Save filtrate for assay. This filtrate is saved in case the target protein did not bind. All filtrates should be kept until the fraction containing the target protein has been identified. Storage conditions depend on the stability of the target protein; as a general rule they should be refrigerated to minimize microbial growth. Bacteriostatic agents may be added for prolonged storage.
11. Add 100 ml binding buffer to funnel and resuspend gel using a stirring rod. Allow resuspended gel to stand for 5 min, then apply suction to remove buffer. Repeat three or four times, pooling all washings with the filtrate from step 10. The gel containing the bound sample may be packed in a column for subsequent elution steps (see Basic Protocol 2).
Elute with step gradient of increasing salt concentration 12. Transfer the sintered-glass funnel to the top of a clean 2000-ml side-arm flask. 13. Add an equal volume of wash buffer to gel in funnel and resuspend using a stirring rod. Allow resuspended gel to stand for 5 min, then apply suction to remove buffer. Save filtrate. The wash buffer is the first step of the gradient. The filtrate will contain sample components that were weakly adsorbed to the gel (see annotation to step 10).
14. Measure absorbance of filtrate at 280 nm using wash buffer as blank. 15. Repeat steps 13 and 14 until the last filtrate shows no significant absorbance at 280 nm. Pool and save the filtrates. 16. Transfer the sintered-glass funnel to the top of a clean 2000-ml side-arm flask. 17. Add an equal volume of elution buffer to gel in funnel and resuspend using a stirring rod. Allow resuspended gel to stand 5 min, then apply suction to remove buffer. Save filtrate. The elution buffer is the second step of the gradient. The filtrate will contain the sample components that were more strongly adsorbed to the gel and should include the target protein.
18. Measure the absorbance of the filtrate at 280 nm using elution buffer as a blank. 19. Repeat steps 17 and 18 until the last filtrate shows no significant absorbance at 280 nm. Pool and save the filtrates for assay and subsequent purification of the target protein. If gel is to be reused, continue with remaining steps. Otherwise, discard the gel.
Regenerate ion-exchange medium for reuse 20. Transfer the sintered-glass funnel to the top of a clean 2000-ml side-arm flask. 21. Add an equal volume of regeneration buffer to gel in funnel and resuspend using a stirring rod. Allow resuspended gel to stand for 5 min, then apply suction to remove buffer. Repeat five times and discard filtrates. Ion-Exchange Chromatography
The regeneration buffer removes strongly bound materials from the gel. See Support Protocol 5 for additional considerations in regenerating ion-exchange media.
8.2.12 Supplement 15
Current Protocols in Protein Science
22. Repeat step 11 for a total of five washes to reequilibrate the gel with binding buffer. 23. Check the pH and conductivity of the last filtrate to ensure that the gel is properly equilibrated. See Support Protocol 5 for information on storing ion-exchange media.
pH-BASED STEP-GRADIENT ELUTION The net charge of a protein is pH dependent. Therefore, altering the pH of the mobile phase to make it closer to the pI of a protein can change its net charge, causing it to desorb and elute from an ion exchanger (see Figure 8.2.1). Continuous pH gradients are very difficult to produce at constant salt concentration because charge characteristics of sample components, buffering ions, and ion-exchange media depend on the pH of the system. pH-based step gradients are much easier to produce and more reproducible. For anionexchange media, elution occurs when the pH is decreased. For cation exchange, the pH is raised for elution.
ALTERNATE PROTOCOL
The first basic protocol can be modified for pH-gradient elution by selecting a buffer at a pH suitable for elution, based on results obtained in a pilot experiment (see Support Protocol 1). This elution buffer is then used in steps 17 to 19 of the first basic protocol. Incubation times and buffer volumes should be increased by 20% when using pH elution. When scouting methods for ion exchange, it is best to start with changes in salt concentration (see Basic Protocol 1) as this technique is simpler and more reproducible. Elution with pH change is only recommended when the ion-exchange behavior of the sample is well known and the resolution required cannot be attained with a change in salt concentration. pH elution may also be appropriate in cases where it provides a superior ionic/pH environment for loading in the next chromatographic step. A combination of increasing salt concentration and pH change may also be used for elution. COLUMN CHROMATOGRAPHY WITH LINEAR GRADIENT ELUTION Column chromatography is generally preferred to batch chromatography (see Basic Protocol 1), especially where high resolution is required, although it accommodates smaller sample volumes. Column chromatography requires specialized equipment to achieve high resolution, including a column containing the medium and a fluid-delivery system appropriate for the pressure and flow rate at which the column is to be operated. It is the technique preferred for the intermediate purification and polishing stages of protein purification (UNIT 8.1). Also in contrast to the first basic protocol, this technique uses a linear gradient for elution. Although more difficult to produce than step gradients (see Basic Protocol 1), linear gradients lead to better resolution (see Support Protocol 3).
BASIC PROTOCOL 2
The example presented here uses a RESOURCE Q anion-exchange column from Amersham Pharmacia Biotech, which may be used with either FPLC (fast protein liquid chromatography) or HPLC (high-performance liquid chromatography) systems. Optimal conditions for other columns should be obtained from the manufacturer. Procedures mentioned in the protocol steps should be carried out according to the manufacturer’s instructions for the particular chromatography system being used. NOTE: All buffers, media, and other system components should be filtered, degassed, and equilibrated to the same temperature before use. Conventional Chromatographic Separations
8.2.13 Current Protocols in Protein Science
Supplement 15
Materials Liquid chromatography system (FPLC or HPLC) Elution buffer: binding buffer (see Basic Protocol 1 and Support Protocol 1) containing 1 M NaCl Binding buffer (see Basic Protocol 1 and Support Protocol 1) RESOURCE Q chromatography column (1-ml packed bed volume; Amersham Pharmacia Biotech) Protein sample to be purified Conductivity meter 0.22-µm filter Prepare chromatography system 1. Set up the liquid chromatography system according to manufacturer’s instructions, without the column in-line. 2. Test system performance by running a blank gradient ranging from 0% to 100% elution buffer in 20 ml at a constant flow rate of 5 ml/min. See Figure 8.2.2 for a diagram of a typical column chromatography system. The gradient is formed by placing binding and elution buffer in the appropriate buffer reservoirs (see Support Protocol 3 for detailed discussion of continuous gradient formation).
pump
injector
mixer
buffer reservoirs column gradient maker
detector
recorder
fraction collector Ion-Exchange Chromatography
Figure 8.2.2 Liquid column chromatography system with gradient maker.
8.2.14 Supplement 15
Current Protocols in Protein Science
Stability of the column at extremes of pH must be considered when choosing binding and elution buffers. The working stability of the RESOURCE Q column is pH 2 to 12, and the stability for cleaning is pH 1 to 14. The flow rate for the test should be appropriate for the column to be connected. For the RESOURCE Q column, the recommended flow rate is 1 to 10 ml/min and the maximum operating pressure is 4 bar.
3. Test for system leaks and ascertain monitor stability according to system documentation. Test accuracy of gradient composition using conductivity meter. Adding 0.1% (v/v) acetone to the elution buffer used in step 2 will allow observation of gradient composition at 280 nm using the UV monitor. Do not include acetone in the elution buffer used for the chromatographic run.
Prepare column 4. Purge system with binding buffer to remove any air, then install column in system. New columns may require precycling prior to use; consult product documentation. Used columns may require cleaning or removal of storage solution prior to use (see Support Protocol 4).
5. Wash column with 5 Vc (5 ml for RESOURCE Q column) of elution buffer at a flow rate of 5 ml/min and check for leakage. Vc refers to the packed bed volume of a column and is calculated according to the equation Vc = πr2 × L, where r is the radius of the column and L is the bed height (i.e., the height of the packed medium in the column). As the RESOURCE Q column used here has a packed bed volume of 1 ml, the column is washed with 5 ml of elution buffer. Consult product literature to find Vc for other columns.
6. Equilibrate column with 5 to 10 Vc of binding buffer at 5 ml/min. Collect one fraction at the end of the equilibration stage and measure pH and conductivity to ensure the pH and salt concentration of the fraction are the same as that of binding buffer. If the pH and salt concentration of the fraction are not the same as that of the binding buffer, continue passing binding buffer through the column until equilibration is complete.
Purify protein sample and regenerate column 7. Adjust pH and salt concentration of sample to binding conditions determined in Support Protocol 1. Filter sample prior to injection using a 0.22-µm filter to remove particulates that may clog system or column. For initial experiments, a sample containing 25 mg total protein is appropriate. For columns packed with beads of average diameter ≥90 ìm, a 1-ìm filter may be used; for columns packed with beads of average diameter 34 to 90 ìm, a 0.45-ìm filter may be used. The 0.22-ìm filter is appropriate for columns packed with beads of average diameter <34 ìm (e.g., the RESOURCE Q). Alternatively, the sample may be centrifuged 5 min at 20,000 × g to remove particulates.
8. Inject sample into column by opening sample injection valve. Begin fraction collection. Fractions of 1-ml volume are generally sufficient. The flow rate during sample injection should be reduced for concentrated samples. It may be increased for dilute samples.
9. After all sample is injected, wash column with 3 to 5 Vc of binding buffer at 5 ml/min to remove unbound or weakly bound materials. The monitor signal should be allowed to approach the baseline before continuing.
Conventional Chromatographic Separations
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10. Elute with a linear gradient of 0% to 100% elution buffer in 20 Vc (20 ml). Sample-injection valves should be closed to reduce the system dead volume before beginning gradient.
11. Regenerate column by washing with 5 Vc of elution buffer. 12. Reequilibrate column by washing with 5 to 10 Vc of binding buffer. 13. Assay fractions for target protein. Pool fractions containing target protein for further processing. SUPPORT PROTOCOL 1
TEST TUBE PILOT EXPERIMENT TO DETERMINE STARTING CONDITIONS FOR ION-EXCHANGE CHROMATOGRAPHY Before beginning an ion-exchange separation (Basic Protocol 1 or 2), the pH and salt concentration required for binding the target protein and the change in pH or salt concentration required for elution must be determined empirically. Once the initial binding conditions have been determined, the available protein capacity (for batch chromatography) is determined to predict the amount of medium that must be used for a given amount of sample. Dynamic capacity (for column chromatography) may be determined as in Support Protocol 2. This protocol is most appropriate when the sample is not strictly limited—e.g., when milligram quantities of a recombinant protein are available. If sample is scarce, a column method should be substituted. As illustrated in Figure 8.2.3, aliquots of the protein-containing sample to be purified are mixed in test tubes with small quantities of ion-exchange gel that have been equilibrated with a series of buffers of different pH. The optimal binding pH is chosen on the basis of disappearance of target protein in the supernatant at a certain point in the series, then the process is repeated with another series of test tubes containing gel equilibrated with a series of salt solutions of different concentrations. The optimal salt concentrations for binding and elution are chosen, respectively, on the basis of disappearance, then reappearance at another salt concentration, of protein in the supernatant. Finally, aliquots containing increasing protein concentrations are mixed with the gel in a buffer under the optimal binding pH and salt concentrations previously determined. The available capacity of the gel is then determined on the basis of the point where unbound protein appears in the supernatant. The stability of the target protein at extremes of pH must be considered—i.e., the pH range chosen for the purification should be compatible with retention of structure and activity. In general, this can be evaluated with a crude preparation by incubating aliquots at a series of pH values, then performing the structural or functional assay at the optimal pH value. NOTE: The method presented here is for anion exchange with increasing salt concentration for elution. Differences for cation exchange are noted where appropriate. Select cation-exchange buffers from Table 8.2.1.
Ion-Exchange Chromatography
Materials 20 mM piperazine, pH 5.0, 5.5, and 6.0 (prepare from 100 mM stock) 20 mM 1,3-bis[tris{hydroxymethyl}methylamino]propane (bis-Tris propane), pH 6.5 and 7.0 (prepare from 100 mM stock) 20 mM Tris⋅Cl, pH 7.5, 8.0, and 8.5 (APPENDIX 2E; prepare from 100 mM stock) Q Sepharose Fast Flow (50% slurry in 20% ethanol; Amersham Pharmacia Biotech) or equivalent anion exchange resin in appropriate buffer Protein sample to be purified, containing known quantity of target and total protein
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4 M NaCl 15-ml test tubes Centrifuge with rotor accommodating 15-ml test tubes (optional) Determine optimal binding pH 1. Set up a series of test tubes, numbered 1 to 8, containing different buffers as follows: tube 1: 5 ml 20 mM piperazine, pH 5.0 tube 2: 5 ml 20 mM piperazine, pH 5.5 tube 3: 5 ml 20 mM piperazine, pH 6.0 tube 4: 5 ml 20 mM bis-Tris propane, pH, 6.5 tube 5: 5 ml 20 mM bis-Tris propane, pH 7.0 tube 6: 5 ml 20 mM Tris⋅Cl, pH 7.5 tube 7: 5 ml 20 mM Tris⋅Cl, pH 8.0 tube 8: 5 ml 20 mM Tris⋅Cl, pH 8.5. 2. Shake bottle of Q Sepharose Fast Flow to resuspend gel. Pour 25 ml of gel slurry into a graduated cylinder, then allow gel to settle. Adjust volume of liquid above gel to equal the volume of settled gel (i.e., prepare 50% slurry), then resuspend with a glass stirring rod.
A determine pH for binding and elution pH 5.0 pH 5.5 pH 6.0 pH 6.5 pH 7.0 pH 7.5 pH 8.0 pH 8.5 pH 9.0 pH 9.5
B determine salt concentration for binding and elution at binding pH (7.5)
0.05 M 0.1 M 0.15 M 0.2 M 0.25 M 0.3 M 0.35 M 0.4 M 0.45 M 0.5 M
C determine available capacity at binding pH (7.5) and NaCl concentration (0.1 M)
10
20
30
40
50
60
70
80
90
100
mg/ml protein
sample adsorbed
sample partially adsorbed
sample not adsorbed
Figure 8.2.3 Results of a typical test tube pilot experiment for selecting initial conditions for anion-exchange chromatography. Series of tubes with protein and ion-exchange medium at (A) varying pH (to select binding and elution pH); (B) constant pH, varying NaCl concentration (to select binding and elution salt concentrations); (C) constant pH and NaCl concentration, varying protein concentration (to determine available capacity). The binding pH here will be ≥7.5; the elution pH will be ≤5.0; the binding salt concentration will be ≥0.15 M; the elution salt concentration will be >0.3 M; and the available capacity will be >50 mg/ml.
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3. Add 2 ml of 50% slurry of Q Sepharose Fast Flow to each tube and mix. SP Sepharose Fast Flow (Amersham Pharmacia Biotech) or equivalent may be used for cation exchange.
4. Allow gel to settle to bottom of tube or centrifuge 1 min at ∼5000 × g, room temperature. 5. Decant supernatant from each tube and replace with 5 ml of the same buffer added in step 1. Gently shake or vortex to resuspend gel, then let tubes stand 2 min. 6. Repeat steps 4 and 5 three times to equilibrate gels in each tube. 7. Decant supernatants and resuspend each equilibrated gel in 1 ml of the same buffer added in step 1. 8. Set up eight identical aliquots (≥1 ml) of the protein to be purified in tubes numbered 1 to 8, each containing 0.1 to 1 mg total protein. Adjust the pH of each to obtain eight different pH values corresponding to those of tubes 1 to 8 in step 1. Quantity of protein to be used depends on availability of sample and extinction coefficient of target protein. pH may be adjusted by titrating with acid or base using pH paper or by diluting 1:1 with buffer of desired pH.
9. Add the corresponding pH-adjusted protein aliquot to each of the 8 tubes from step 7. Mix gently for 10 min by periodic swirling, then allow gel to settle. If the salt concentration of the sample is too high, no binding will occur. It is therefore recommended that the sample be desalted into the appropriate buffer as in UNIT 8.3.
10. Assay supernatant from each tube for the target protein. Measuring the absorbance of the supernatant at 280 nm is usually sufficient. Figure 8.2.3 shows a set of possible results. A decrease or absence of target protein activity (or absorbance) in the supernatant indicates binding. Specific assays for the target protein may also be used.
11. Select the lowest pH at which the highest quantity of target protein is bound to use as the binding pH for further work. Select the highest pH where no binding occurs as the elution pH. For anion exchange, it is best to select a binding pH no more than 0.5 to 1 pH unit (pH 7.5 to 8.0 in Fig. 8.2.3) above the highest pH at which no binding occurs. If too high a pH is chosen, elution becomes difficult and high salt concentrations may be required. For cation exchange, select a binding pH 0.5 to 1 unit below the lowest pH where no binding occurs. Select the lowest pH where no binding occurs as the elution pH.
Determine salt concentration for binding and elution 12. Set up a series of test tubes, numbered 1 to 10. Using the binding buffer selected in step 10 in place of the different buffers in step 1, perform steps 1 to 7 to equilibrate the Q Sepharose Fast Flow with buffer in all ten tubes. 13. To each tube, add the same amount of the protein to be purified that was added to the tubes in step 8, equilibrated in the binding buffer selected in step 10. 14. Mix contents of tubes gently for 10 min, then allow gel to settle. 15. Decant supernatants from each tube and discard. Wash gel two times with 5 ml of the binding buffer, allowing the gel to settle and decanting the supernatants after each addition of buffer. Ion-Exchange Chromatography
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16. Add 2 ml of the binding buffer to each tube, then sequentially add water and 4 M NaCl to each tube as follows: tube 1: 1.90 ml H2O and 0.10 ml 4 M NaCl (0.10 M NaCl final) tube 2: 1.80 ml H2O and 0.20 ml 4 M NaCl (0.20 M NaCl final) tube 3: 1.70 ml H2O and 0.30 ml 4 M NaCl (0.30 M NaCl final) tube 4: 1.60 ml H2O and 0.40 ml 4 M NaCl (0.40 M NaCl final) tube 5: 1.50 ml H2O and 0.50 ml 4 M NaCl (0.50 M NaCl final) tube 6: 1.40 ml H2O and 0.60 ml 4 M NaCl (0.60 M NaCl final) tube 7: 1.30 ml H2O and 0.70 ml 4 M NaCl (0.70 M NaCl final) tube 8: 1.20 ml H2O and 0.80 ml 4 M NaCl (0.80 M NaCl final) tube 9: 1.10 ml H2O and 0.90 ml 4 M NaCl (0.90 M NaCl final) tube 10: 1.00 ml H2O and 1.00 ml 4 M NaCl (1.00 M NaCl final). 17. Mix contents of tubes gently for 10 min, then allow gel to settle. 18. Assay supernatant from each tube for the target protein. Figure 8.2.3 shows a set of possible results. Presence of target protein in the supernatant indicates elution of the target protein.
19. Select the highest salt concentration at which no elution occurs as the maximum salt concentration allowable for binding target protein and for washing away unbound sample components. Select a salt concentration at least 0.05 M above the concentration where no protein binds to the gel as the salt concentration for elution. Determine available capacity 20. Prepare a buffer/salt solution with the binding pH selected in step 10 and the binding salt concentration selected in step 19. 21. Set up a series of test tubes numbered 1 to 10. Using the buffer/salt solution prepared in step 20 in place of the different buffers in step 1, perform steps 1 to 7 to equilibrate the Q Sepharose Fast Flow with buffer/salt solution in all ten tubes. 22. Add aliquots of sample to each tube to obtain the following amount of target protein: tube 1: 10 mg tube 2: 20 mg tube 3: 30 mg tube 4: 40 mg tube 5: 50 mg tube 6: 60 mg tube 7: 70 mg tube 8: 80 mg tube 9: 90 mg tube 10: 100 mg. 23. Mix gently for 10 min, then allow gel to settle. 24. Assay supernatant from each tube for the target protein. Figure 8.2.3 shows a set of possible results. Presence of target protein in supernatant indicates that the available capacity has been exceeded.
25. Select the highest protein concentration at which the target protein does not appear in the supernatant as the available capacity. Calculate the amount of ion-exchange medium needed for batch purification based on 50% of the available capacity (defined in UNIT 8.1). If a column system is used in pilot experiments, however, it is best to determine the dynamic capacity of the medium (see Support Protocol 2). However, 20% of the available capacity is a safe and reasonable starting point for pilot experiments using column chromatography.
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SUPPORT PROTOCOL 2
MEASUREMENT OF DYNAMIC (COLUMN) CAPACITY AND BREAKTHROUGH CAPACITY OF ION-EXCHANGE COLUMNS For chromatographic separations that will be performed on a routine basis as well as separations that will be scaled up, it is necessary first to optimize conditions with respect to resolution and to determine the capacity of the medium with respect to the target protein (see Support Protocol 1). Knowledge of the capacity allows optimal use of the gel medium in terms of cost and yield, and also allows working limits to be set on the sample load to ensure robustness of a particular purification step. After capacity has been determined, the separation can be optimized with respect to time. There are two capacity terms that commonly appear in the literature (in addition to available capacity; also see Support Protocol 1 and UNIT 8.1): dynamic capacity and breakthrough capacity. The dynamic (or column) capacity is the capacity of an ion-exchange column under defined conditions of flow rate, pH, salt concentration and sample concentration. The breakthrough capacity (QB) for a system is obtained by calculating the amount of protein that has been absorbed by the column when the sample is first detected in the effluent, or when the recorder signal reaches some arbitrarily defined percent of full-scale deflection. Breakthrough capacity values at 50% full-scale deflection (QB50) are commonly reported in the literature. However, it is generally recommended to use only 10% to 20% of the published QB as the practical capacity of the column to ensure high recovery of the target protein. Additional Materials (also see Basic Protocol 2) Ion-exchange gel of unknown capacity, and column Elution buffer capable of eluting target protein in single step of salt concentration or pH (e.g., 2 M NaCl; see Support Protocol 1) 1. Pack a defined amount of the ion-exchange gel into the column. See UNIT 8.3 for guidelines on column packing. Normally, a quantity of gel sufficient to give a packed bed volume of ∼1 ml is sufficient. In the case of a prepacked column, the amount of gel is predetermined. The packed bed volume (Vc) is calculated according to the equation Vc = π r2 × L, where r is the radius of the column and L is the bed height (i.e., the height of the packed medium in the column).
2. Prepare chromatography system and column (see Basic Protocol 2, steps 1 to 6) using the optimal binding and elution conditions determined in Support Protocol 1. Do not exceed 75% of the flow rate used for packing. The effect of flow rate on capacity should be determined in a true optimization.
3. Prepare protein solution and inject into column (see Basic Protocol 2, steps 7 and 8). Apply solution continually until the recorder shows >50% full-scale deflection, then stop. The absorbance of the binding buffer corresponds to 0% and the absorbance of the sample corresponds to 100% full-scale deflection.
4. Wash column with binding buffer until 0% full-scale deflection is approached. 5. Elute proteins with a buffer providing a single-step increase in salt concentration or pH. Continue collecting eluant until recorder shows ≤2% full-scale deflection, then pool fractions. A typical chromatogram for this procedure is presented in Figure 8.2.4.
6. Assay pooled fractions from step 5 for target protein. Calculate maximum amount of protein (A) that can be bound to the column according to the equation A = Cp × V, Ion-Exchange Chromatography
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Full-scale deflection (%)
sample application
washing
elution
100
50
pool x Elution volume (ml)
Figure 8.2.4 Typical chromatogram for determining the capacity of an ion-exchange column. The volume of eluant (x) at 50% full-scale deflection is used to calculate the breakthrough capacity (QB50).
where Cp is the protein concentration of the pooled fractions (mg/ml) and V is the volume of the pooled fractions (ml). 7. Calculate the column (dynamic) capacity (mg/ml) using the equation dynamic capacity = A/Vc. 8. Calculate breakthrough capacity (QB50) using the equation QB50 = (Cs × x)/Vc, where Cs is the concentration of protein in the original sample and x is the volume of sample that has been applied to the column at the point where 50% full-scale deflection has been attained. If QB50 greatly exceeds the dynamic capacity, then the target material may still be adsorbed on the column, and the elution conditions should be reexamined. See Figure 8.2.4 for graphical depiction of these variables.
GRADIENT-FORMATION TECHNIQUES
SUPPORT PROTOCOL 3
Simple Gradient Mixers Gradient mixers of the type shown in Figure 8.2.5 can generate linear pH or salt gradients in gravity-based or one-pump chromatography systems. Similarly designed gradient mixers used for casting electrophoresis gels may also be used for chromatography. The reservoir closest to the column is filled with the binding buffer and the second reservoir is filled with an equal volume of the elution buffer. The gradient begins when the valve between the two reservoirs is opened. Continuous mixing is required in the reservoir closest to the column to ensure gradient linearity and reproducibility.
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stirrer
gradient maker
mixing chamber
elution buffer
valve
binding buffer
Figure 8.2.5 Gradient mixer for forming gradients of pH and salt concentration used during ion-exchange chromatography. The apparatus shown is an Amersham Pharmacia Biotech Gradient Mixer GM-1, which can be used for preparing gradients up to 500 ml. A linear gradient of salt concentration is produced by filling the reservoirs with buffers at the same pH and different salt concentration. The mixing chamber contains the lower-salt-concentration (binding) buffer, and the other chamber contains the higher-salt-concentration (elution) buffer. A continuous linear or nonlinear pH gradient may be produced by filling the reservoirs with buffers at the same salt concentration and different pH. A gradient of increasing pH is used for cation-exchange separations, and a gradient of decreasing pH is used for anion-exchange separations. The buffer chambers are joined by a channel controlled by a valve, and outflow from the mixing chamber is controlled by another valve.
Gradient Mixing with Multichannel Peristaltic Pumps Peristaltic pumps that can accommodate three or more pumping channels simultaneously (Fig. 8.2.6) can be used to form linear or complex gradients (see Critical Parameters). Linear gradients are easily formed by using the same size tubing for all channels, pumping with one channel from the elution buffer reservoir to the binding buffer reservoir, and pumping with the other two channels from the binding buffer reservoir to the column. Continuous mixing is required in the binding buffer reservoir closest to the column to ensure gradient linearity and reproducibility. Switch-Valve-Based Gradient Mixing
Ion-Exchange Chromatography
Switch-valve-based gradient formation is useful for forming step, linear, and complex gradients (see Critical Parameters). Gradients are formed by proportioning variable amounts of binding buffer and elution buffer through a single three-way valve using a system controller. When programmed to deliver 10% elution buffer, the controller will open the valve port for binding buffer 90% of the time and the port for elution buffer 10% of the time. As higher switching rates are required for lower flow rates to ensure gradient accuracy, high-quality system controllers will vary the rate at which the valve switches (i.e., the number of switching events per unit time) as a function of flow rate. A dynamic mixer should be included in the flow path just after the proportioning valve to ensure gradient accuracy. Switch-valve-based gradients are generally not very accurate from 1% to 10% elution buffer (90% to 99% binding buffer) or from 1% to 10% binding buffer (90% to 99% elution buffer). Therefore, the ionic strength of the binding and elution buffers may be adjusted to allow a gradient of the desired composition to be formed by mixing the two buffers in the range of 20% to 80% elution buffer (80% to 20% binding buffer).
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channel 1 channel 2 channel 3
resembles a label
multichannel peristaltic pump
this
is
elution buffer
text
binding buffer
mixer
channel 3
to column
Figure 8.2.6 Peristaltic pump accommodating three pumping channels for continuous gradient formation.
Gradient Formation by Autoblending Some chromatography systems allow gradient formation over a range of both pH and salt concentration. Four solvents—a high-pH buffer, a low-pH buffer, a concentrated salt solution, and water—are blended via four two-way valves. A system controller proportions the high- and low-pH buffers to achieve the desired pH gradient while simultaneously proportioning the concentrated salt solution and water to achieve the desired salt-concentration gradient. Gradients formed in this way are usually difficult to reproduce but may be useful for scouting initial binding and elution conditions. Gradient Formation with Twin Pumps Use of two positive-displacement pumps, as in HPLC and FPLC systems, provides the highest degree of accuracy, precision, and reproducibility in gradient formation. One pump delivers binding buffer while the other delivers elution buffer. A system controller allows gradient formation simply by controlling the flow rate of each pump. A dynamic mixer should be included in the flow path, after the pumps and before the column, to ensure gradient accuracy. CLEANING AND REGENERATION OF ION-EXCHANGE MEDIA Proper maintenance of chromatography media is as much an art as protein purification itself. Knowing what kind of contaminating material from the sample is bound to the column helps in selecting a regime for maintaining good column hygiene. It is best to practice preventative maintenance by regenerating the medium after every run, using a high-salt buffer or large change in pH, and periodically cleaning the column and checking the system flow characteristics. If any increase in column back-pressure is noted, the column should be cleaned as soon as possible. Once enough contaminant has bound to a column to reduce or block the flow, it is probably too late to save the medium without investing considerable time and effort.
SUPPORT PROTOCOL 4
Conventional Chromatographic Separations
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Routine Washing of Columns and Media Columns or bulk medium should be washed after every run with salt solution until an ionic strength of ≥2 M is reached. This should remove any substances bound by ionic forces. This is recommended after every run. Removal of Contaminants Alkali-soluble contaminants. Contaminants such as lipids, proteins, and nucleic acids can usually be removed by washing with 2 to 3 column volumes of 0.1 M NaOH, followed by 2 to 3 column volumes of Milli-Q-purified (or equivalent) distilled water and then 2 to 3 column volumes of binding buffer. Many media can tolerate brief exposure to 1 M NaOH, if harsher conditions are required to remove contaminants; however, manufacturer’s guidelines should be consulted regarding pH stability of the medium. Hydrophobic contaminants. Lipids and other hydrophobic materials may be removed by washing with alcohol solutions (e.g., 70% ethanol) or nonionic detergents. When using alcohols or organic solvents to remove hydrophobic materials, it is often effective to wash first with a gradient from 0% to 100% and then with a gradient from 100% to 0%. Repeat washing with the alternating gradients until no contaminants are detected in the eluant. Metallic contaminants. Contamination with metals may cause blue or gray discoloration at the top of an ion-exchange column. These metals can originate from impurities in buffer salts or water, or they can leach from metallic system components in contact with buffer solutions. Metal contaminants may usually be removed by treating the column with several column volumes of 10 mM HCl (i.e., pH 2) saturated with EDTA. Manufacturer’s guidelines should be checked for pH stability of medium and column before exposure to mild or strong acid solutions. Precipitated materials. Precipitated materials that have accumulated in a column are very difficult to remove. Unless the precipitated material can be physically removed by removing the gel at the top of the column, the material will have to be resolubilized. Detergents, urea, and guanidine⋅Cl can be introduced to help dissolve contaminants. The viscosity of concentrated solutions used for cleaning may require very low flow rates. The column may be equilibrated and incubated in 6 M urea, then washed with distilled water and buffer. Degradative enzymes (e.g., proteases and lipases) may also be introduced in an attempt to degrade precipitates on the column; however, enzymatic cleaning can be quite expensive. It may be useful to reverse the direction of flow through the column (i.e., from bottom to top) when attempting to remove particulate material from the top of the bed. SUPPORT PROTOCOL 5
Ion-Exchange Chromatography
STORAGE OF ION-EXCHANGE MEDIA Bacterial and microbial growth can seriously interfere with the chromatographic properties of any chromatography medium and endanger the sample as well. Prior to storage, all media should be cleaned and sanitized. During prolonged experiments or storage (i.e., >24 hr at room temperature or 48 hr at 4°C), a bacteriostatic or antimicrobial agent should be added to the ion exchanger. Antimicrobials chosen to be added for storage should not bind to the ion exchanger and should be easily removed when the gel is to be reused. Azide is anionic and will bind to anion-exchange media. Treatment with 1 M NaOH for 1 hr will sanitize most media by lowering the bacterial count 100-fold, and will help solubilize dead cells (Pharmacia Biotech, 1997). Storage of sanitized media in 0.1 M NaOH will not leave any toxic materials on the column—a significant consideration in pharmaceutical production. Manufacturer’s guidelines should
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be checked for stability of the medium and column in NaOH. Silica-based media will not tolerate high pH. Exposure to 70% ethanol for 3 to 4 hr can be used to sanitize most media. Most ion-exchange media and columns can be stored for prolonged periods of time in 20% ethanol. Manufacturer’s guidelines should be checked for stability of the medium and column in ethanol. Effective antimicrobial agents appropriate for use with anion-exchange media include either 0.001% phenyl mercuric salts or 0.002% chlorhexidine, in weakly alkaline solution. An effective antimicrobial agent for use with cation-exchange media is 0.005% merthiolate in weakly acidic solution. An agent that is effective for use with either anion or cation exchangers is 0.05% trichlorobutanol in weakly acidic solution. COMMENTARY Background Information
Critical Parameters
The first ion exchangers were synthetic resins designed for applications such as demineralization, water treatment, and recovery of ions from waste. Early ion-exchange resins were tightly cross-linked hydrophobic polymers, highly substituted with ionic groups, and had very high capacity for small ions. The high degree of cross-linking provided mechanical strength, but the limited porosity restricted use of these media with large molecules. In addition, the high charge density resulted in very strong binding, and the hydrophobic matrix tended to adsorb and denature labile biological materials. The first ion exchangers designed for use with biomolecules were developed by Peterson and Sober (1956). These cellulose-matrix ion exchangers were very hydrophilic and had little tendency to denature proteins, but had low capacities and poor flow characteristics. Modern ion-exchange media have very high capacity, often >100 mg/ml. Modern particle technology has significantly increased the mechanical strength and flow characteristics of ion-exchange media so that separations that once required hours or days may now be completed in a few minutes. Ion-exchange can easily be coupled to other methods of protein separation in a multistep purification. Samples purified by ion exchange can typically be applied to a hydrophobic-interaction chromatography (HIC) medium simply by adding NaCl (or other salts) to a concentration sufficient for binding (UNIT 8.4). Samples to be further purified by chromatofocusing or an additional round of ion exchange will require desalting (UNIT 8.3). Samples may usually be applied directly to gel-filtration columns (UNIT 8.3) with no intervening treatment.
The most critical aspects of any ion-exchange experiment are selection of the appropriate ion-exchange medium and of the initial conditions for binding and elution of the target protein or contaminants. Once these parameters are selected, conditions can be optimized with respect to the three primary parameters of any chromatographic separation—i.e., resolution, capacity, and speed. These three parameters are mutually exclusive. Resolution will typically decrease when the sample load is increased and when the flow rate is increased (to increase speed). It is generally recommended that resolution be optimized first, then capacity, and finally speed. Optimization of resolution This parameter is most easily optimized by controlling the selectivity (UNIT 8.3) of the medium through manipulation of binding and elution conditions. Construction of chromatographic titration curves followed by optimization of the shape of the elution gradient is a common approach to achieving maximum resolution. Chromatographic titration curves can be constructed by performing a series of anionand cation-exchange experiments at different pH values as in Figure 8.2.7. For each of these experiments, the elution salt concentrations (in mM NaCl) for sample components A, B, and C are plotted on the y axis and the operating pH is plotted on the x axis. It is then simple to observe the pH ranges within which optimal resolution between the various sample components is achieved. The different sample components are usually detected by UV absorption, but may also be viewed by other means (e.g., fluorescence or bioactivity). Data for chroma-
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A
a. cation exchange b. cation exchange at pH 3.0 at pH 5.0
C B A
A280
C
B
B C
c. cation exchange at pH 7.0
A
A
Elution NaCl concentration d. anion exchange at pH 8.0
e. anion exchange at pH 10.0
C
BC
f. anion exchange at pH 11.0
A
B
B A
A
A280
C
Elution NaCl concentration
a cation exchange
Increasing mM NaCl
B
A
b
B c C 9
0 3
4
5
10
11
pH
6
Increasing mM NaCl
B C d
e A f anion exchange
Figure 8.2.7 Construction of a chromatographic titration curve. (A) Six chromatographic separations (a to f) of a sample containing proteins A, B, and C are carried out at six different operating pHs using a linear gradient of increasing salt concentration. (B) The elution ionic strengths for components A, B, and C (y axis) are plotted against the operating pH (x axis) for experiments a to f. Note that the salt concentration in mM NaCl (y axis) is ascending in both directions. The optimal resolution with respect to component A occurs below pH 5 with cation exchange and above pH 9 with anion exchange.
Ion-Exchange Chromatography
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tographic titration curves can often be generated by fast screening with low-volume gradients (e.g., 5 to 10 Vc) and low sample mass (e.g., <1 mg). After optimization of resolution with respect to pH, the shape of the elution gradient can be optimized. A gradient can be conceptually divided into segments, and each segment may have a different gradient shape. Gradient shapes include linear, concave, convex, and step. Complex gradients are constructed by using different gradient shapes in consecutive segments. The slope (e.g., mM NaCl per minute) of a linear segment is constant, that of a concave segment continually increases, that of a convex gradient continually decreases, and that of a step gradient approaches infinity. Resolution between sample components in any segment can usually be increased by decreasing the gradient slope in that segment. However, decreasing the gradient slope will increase the peak volume during elution, the gradient volume, and the time of the separation. Step gradients can provide the most concentrated fractions, and are desirable when minimizing sample volume is of primary importance. Use of concave gradients in the initial part of a segment and convex gradients in the later part of a segment can improve resolution. The capabilities of available equipment must be considered in the construction of complex gradients. Use of simple gradients that do not stress the performance limits of the system will result in robust methods that are insensitive to small changes in sample or buffer composition and gradient shape, thereby increasing probability of success. To construct a complete gradient profile, it is first necessary to identify the gradient range where desorption of the target protein occurs, as well as the gradient slope that provides optimal resolution. A single-step elution is then done at a salt concentration just below the range of where the target protein elutes, to remove weakly bound material. The target protein is then desorbed and eluted. A single-step elution at a high salt concentration will then remove tightly bound sample components and prepare the ion-exchange gel for regeneration. This approach, using step gradients before and after the range of interest, can provide simple, robust methods that are easy to scale up, and will save time and reagents. Different ion-exchange media offer different selectivities (UNIT 8.1). If adequate resolution for a specific stage in a purification cannot be
obtained with a specific ion-exchange medium, it may be desirable to select an alternate ion exchanger with a different functional group, or to use a different chromatographic technique at this stage in the separation. Salts other than NaCl may be used in elution, but there are no specific effects from different salts on the separation in ion-exchange chromatography, other than differences in concentration. Karlsson et al. (1998) provides a good review of the effects of nonbuffering ionic species in ion exchange. Optimization of capacity When optimum resolution has been achieved, the effect of increasing sample load on resolution should be evaluated to determine the optimal balance of resolution and capacity utilization. The procedure for determining the dynamic (column) capacity is described in Support Protocol 3 and can be performed using the conditions for optimal resolution (as determined above) to elute the target protein. Once the capacity is determined under the optimal elution conditions, the effect of changes in sample load on resolution can be investigated to identify optimal conditions. It is generally recommended that only 10% to 20% of the dynamic capacity be used to ensure high recoveries of the target protein. Optimization of throughput Once optimal conditions for resolution and capacity have been determined, the flow rate in each gradient segment may be increased and the effect on resolution and capacity monitored to complete the optimization process.
Troubleshooting Common problems encountered during ionexchange experiments, their potential causes, and suggested solutions are outlined in Table 8.2.5.
Anticipated Results A chromatogram illustrating typical results of an ion-exchange column chromatography experiment (i.e., Basic Protocol 1) is shown in Figure 8.2.8. The goal of any purification step is the pure target protein obtained at the end of the day. The chromatogram is only a record for comparison and is deceptive at best. Great care should be exercised in interpreting a chromatogram—i.e., the results from assays performed on fractions collected must be used in interpret-
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Table 8.2.5
Troubleshooting Guide for Ion-Exchange Chromatography
Problem
Possible cause
Protein does not elute in the Buffer pH is incorrect salt gradient Salt concentration is too low Protein does not elute in pH Buffer pH is incorrect gradient Protein elutes in wash phase Salt concentration of binding buffer is too high Salt concentration of sample is too high or pH is wrong Column is not properly equilibrated Ionic detergents or other additives are absorbed to column Resolution is less than Gradient slope is too steep expected Flow rate is too high Proteins or lipids have precipitated on columns Sample has not been filtered properly before application to the column Proteins in sample have formed aggregates that bind strongly to gel Column is poorly packed
Leading or very rounded peaks observed in chromatogram
Too much sample mass has been loaded onto column Detector flow cell volume or mixing spaces in or after column are too large Column is overloaded
Column is poorly packed
Column needs regeneration Tailing of peak is observed in the chromatogram
Sample is too viscous Precipitation of protein has occurred in column filter and/or at top of the gel bed Column is poorly packed
Previous elution profile cannot be reproduced
Ion-Exchange Chromatography
Solution Use buffer pH closer to pI of the protein Use more concentrated elution buffer Calibrate pH meter and prepare new solutions Decrease salt concentration of binding buffer Perform buffer exchange on sample Repeat or prolong equilibration until conductivity is constant Clean column Use shallower gradient or include plateau in gradient Run separation at lower flow rate Clean and regenerate column Regenerate column, filter sample, and repeat chromatography Use urea or detergents to clean column Check packing by running a colored compound and observing the band. Repack column if necessary. Clean and regenerate column. Decrease sample load and repeat run. Change flow cell or reduce all post-column volumes Decrease sample load and repeat run
Check packing by running a colored compound and observing the band. Repack column if necessary. Clean and regenerate column. If this does not help, replace with new one. Reduce amount of protein, or remove nucleic acids from sample. Clean column; exchange or clean filter. Check packing by running a colored compound and observing band. Repack column if necessary. Prepare new solutions
Buffer pH and/or ionic strength are incorrect Sample has deteriorated during Prepare fresh sample storage Proteins or lipids have precipitated Clean and regenerate column on column
continued
8.2.28 Supplement 15
Current Protocols in Protein Science
Table 8.2.5
Troubleshooting Guide for Ion-Exchange Chromatography, continued
Problem
Possible cause
Solution
Sample has not been filtered properly Equilibration was incomplete
Regenerate column, filter sample carefully, and repeat step Repeat or prolong equilibration until conductivity is constant Change elution conditions
Recovery of activity is low but recovery of protein is normal
Target protein is not stable in the elution buffer and is therefore inactivated Enzyme separated from cofactor or other substance needed for activity Microbial growth has occurred in the column Amount of protein in the Protein was degraded by eluted fractions is much less proteinases than expected Protein adsorbed to filter during sample preparation Microbial growth occurred in the column
Test by pooling all fractions and repeating assay Clean column and store gel in 20% ethanol or other antimicrobial agenta Add proteinase inhibitors to buffers
Use another type of filter or add detergents to buffer Clean column and store gel in 20% ethanol or other antimicrobial agenta
aMicrobial growth rarely occurs in columns during use, but steps should always be taken to prevent infection of packed
columns, buffers, and gel suspensions; see Support Protocol 5.
A280
100% elution buffer (high-salt)
100% binding buffers (low-salt)
equilibrate column (5 -10 8c)
inject wash away sample unbound sample components (3 - 5 8c)
elute with linear gradient (20 8c)
final wash with elution buffer (2 - 5 8c)
Time
Figure 8.2.8 Chromatogram representing typical results of stages of an ion-exchange column chromatography experiment. A linear gradient of 0% to 100% elution buffer is used as in Basic Protocol 2. The broken line superimposed on the chromatogram represents the composition of the elution gradient (100% binding buffer to equilibrate column and remove unbound sample components, followed by a linear gradient from 100% binding buffer to 100% elution buffer to elute bound proteins and a final wash with 100% elution buffer). Vc represents the packed bed volume of the column.
Conventional Chromatographic Separations
8.2.29 Current Protocols in Protein Science
Supplement 15
ing the chromatogram and inferring the identity of any given chromatographic peak.
Pharmacia Biotech. 1995. Ion Exchange Chromatography: Principles and Methods, ed. AA. Pharmacia Biotech AB, Uppsala, Sweden.
Time Considerations
Pharmacia Biotech. 1997. Application Note 181124-57: Use of Sodium Hydroxide for Cleaning and Sanitizing Chromatography Media and Systems. Pharmacia Biotech, Uppsala, Sweden.
If all necessary equipment is available and functioning properly, it should be possible to develop a chromatographic method in a day. With automated systems (e.g., FPLC and HPLC) and modern ion-exchange media, method development may require just a few hours. However, the most time-consuming aspect in developing a purification step is typically the determination of protein purity in the fractions collected, which will always depend on the specific protein of interest. The key to minimizing the time required for such determinations is to strive for high resolution and a minimum number of fractions.
Literature Cited Karlsson, E., Rydén, L., and Brewer, J. 1998. Ion exchange chromatography. In Protein Purification: Principles, High Resolution Methods and Applications, 2nd ed. (J.C. Janson and L. Rydén, eds.) pp. 145-205. John Wiley & Sons, New York. Gianazza, E. and Righetti, P.G. 1980. Size and charge distribution of macromolecules in living systems: J. Chromatog. 193:1-8. Peterson, E.A. and Sober, H.A., 1956. Chromatography of proteins. I. cellulose ion exchange adsorbents. J. Amer. Chem. Soc. 78:751-755.
Key References Cooper, E.H., Turner, R., Webb, J.R., Lindblom, H., and Fägerstam, L. 1985. Fast liquid protein chromatography scale-up procedures for the preparation of low molecular weight proteins from urine. J. Chromatogr. 327:269-277. Good example of methods development with respect to optimization of resolution. Pharmacia Biotech. 1985. FPLC Ion Exchange and Chromatofocusing: Principles and Methods. Pharmacia Biotech AB, Uppsala, Sweden. Contains detailed discussions of experimental approach, methodology, and applications for protein purification. Pharmacia Biotech, 1995. See above. Concise descriptions of theory and practice in planning and implementing ion-exchange purification.
Contributed by Alan Williams and Verna Frasca Amersham Pharmacia Biotech Piscataway, New Jersey
Ion-Exchange Chromatography
8.2.30 Supplement 15
Current Protocols in Protein Science
Gel-Filtration Chromatography
UNIT 8.3
Gel filtration (GF) separates proteins solely on the basis of differences in molecular size. Separation is achieved using a porous matrix to which the molecules, for steric reasons, have different degrees of access (smaller molecules have greater access and larger molecules less). The matrix is packed into a chromatographic column, the sample mixture applied, and the separation accomplished by passing an aqueous buffer (the mobile phase) through the column. Protein molecules that are confined in the volume outside the matrix beads will be swept through the column by the mobile phase and the sample is thus eluted in decreasing order of size. The protein zones eluted are detected by an in-line UV monitor and fractions are collected for subsequent specific analysis or further preparation steps. GF is often used to condition samples prior to further purification by adsorptive techniques. It is also frequently used in the final polishing stage (UNIT 8.1) to remove a few remaining contaminants with surface properties similar to the target protein (e.g., for polishing monomers from oligomers). Three basic protocols are provided in this unit, corresponding to the major applications of gel filtration. GF is used for desalting or group separation (see Basic Protocol 1), in which the target protein and contaminating solutes differ substantially in molecular size. It is also used for protein fractionation (see Basic Protocol 2), where proteins with only a small size difference must be separated. A third application is in characterizing the molecular dimensions of proteins (see Basic Protocol 3). A Support Protocol is also provided for calibrating GF columns to be used in estimating molecular size. NOTE: Among the items referred to in this unit, Sephadex, Sephacryl, Superdex, Superose, and HiLoad are trademarks of Amersham Pharmacia Biotech; Bio-Gel and Bio-Sil are trade marks of Bio-Rad; and Toyopearl and TSK SW are trademarks of Toyo Soda. STRATEGIC PLANNING Selecting Matrix and Column for Desalting The exclusion limit—i.e., the smallest-sized protein molecule that will be excluded from the pores of the matrix—is the most important factor in selecting a matrix for desalting. If the protein is able to permeate the matrix, flow-dependent zone broadening (UNIT 8.1) will occur, reducing the resolution. On the other hand, gels of very small pore size generally contain more matrix, and thus have smaller separating volumes. A gel of sufficiently small exclusion limit, but not smaller, should therefore be selected. Another factor to take into consideration is the particle size of the matrix. This should not be too small because the pressure drop over the column is inversely proportional to the square of the particle size; this factor is especially important for gravity flow columns. The selection of column dimensions (i.e., the bed volume of the column) is influenced by the amount of sample that is to be processed. Columns for desalting are typically short and wide to accommodate large sample volumes and keep the processing time short. Because the zone broadening of the column will have a limited effect on the peak width in desalting as compared to other applications of gel filtration—e.g., protein fractionation—short columns run at high flow rates are used for large-scale desalting. The typical length of columns for laboratory use is ∼10 cm. Columns for desalting vary greatly in size and sophistication, ranging from Pasteur pipets plugged with glass wool to prepacked high-performance GF columns. The scale of Contributed by Lars Hagel Current Protocols in Protein Science (1998) 8.3.1-8.3.30 Copyright © 1998 by John Wiley & Sons, Inc.
Conventional Chromatographic Separations
8.3.1 Supplement 14
Table 8.3.1
Gel Filtration Matrices Suitable for Desaltinga
Gel type Sephadex G-10 Bio-Gel P-2 Sephadex G-25 Bio-Gel P6DG Sephadex G-50 Bio-Gel P-30
Exclusion limit Mrb
Radius (Å)c
700 1,800 5,000 6,000 30,000 40,000
7 10 14 15 25 28
Supplierd PB BR PB BR PB BR
aData as reported by manufacturers. bMolecular mass of smallest protein excluded from matrix. cApparent radius of smallest spherical protein excluded from matrix, 1 calculated as R = 0.808 Mr ⁄3, where R is the radius and Mr the molecular
mass (Hagel, 1989). dAbbreviations: BR, Bio-Rad; PB, Amersham Pharmacia Biotech. Addresses and phone numbers of suppliers are provided in the SUPPLIERS APPENDIX.
operation for which they are used also varies widely, with applications ranging from micropreparative cleanup of a 50-µl reduction/alkylation mixture using a 0.9-ml column to industrial de-ethanolization of 12 liters of human serum albumin using a 75-liter column. Incidentally, columns for both these procedures may be packed with Sephadex G-25—of course, of different particle size. Some gel-filtration media suitable for laboratory-scale desalting are listed in Table 8.3.1, and some empty columns that can be used with these media are listed in Table 8.3.2. A partial compilation of disposable prepacked columns for desalting is found in Table 8.3.3. This table also contains information about spin columns for use with a centrifuge. Spin columns require special techniques, and they are presently not widely used for desalting proteins; therefore no protocols for their use have been included in this unit. More information about spin columns can be obtained from the suppliers. Selecting Matrix and Column for Protein Fractionation Suitability of a gel for protein fractionation may be judged from “typical runs” made with synthetic protein mixtures, the results of which are published by suppliers. The solute of interest should ideally be eluted in the first part of the chromatogram, but not too close to the void volume. A matrix with a distribution coefficient (see Basic Protocol 3) of ∼0.2 to 0.4 has been reported to yield maximum resolution (Hagel, 1989). This corresponds to an elution volume of ∼1.5 to 2 times the void volume. A long bed and a matrix with a large pore volume will increase the peak-to-peak distance between the proteins and thus increase the resolution of the system (also see Critical Parameters and UNIT 8.1). Another way to increase resolution is to employ a GF matrix of small particle size, which will reduce the zone width of the protein bands. The flow rate will also affect resolution. As in the case of desalting, the maximum sample volume that can be processed depends upon the bed volume and the pore volume of the packed bed. Thus, there are many parameters to consider when optimizing protein fractionation by GF. In the worst case, a suitable matrix may have to be found by trial and error.
Gel-Filtration Chromatography
Commercial prepacked columns for protein fractionation often sacrifice column length and pore volume in favor of small particle size. This results in columns that are optimized for fast separation at the expense of some reduction in the maximum sample volume that can be loaded. Therefore, traditional columns packed in the laboratory are often still a
8.3.2 Supplement 14
Current Protocols in Protein Science
good alternative for purification of large volumes of sample. If time is critical in the separation (e.g., in proteinase removal), fast column operation is of course advantageous. Matrices of large particle size will permit protein fractionation at low operating pressure. Some matrices suitable for protein fractionation are listed in Table 8.3.4, and some empty columns suitable for use with these media are listed in Table 8.3.5. Table 8.3.6 lists some prepacked columns for protein fractionation. Columns and media are selected on the basis of the difficulty of the separation (i.e., how small the size difference is between the proteins to be separated) and whether scaleup will be required (i.e., the availability of a variety of
Table 8.3.2
Empty Columns Suitable for Desaltinga
Column
Minimum bed volume (ml)
Maximum bed volume (ml)
Maximum operating pressure (MPa)
Supplierb
1 1 0 1 2
15 20 520 1374 1884
— — 0.1 0.1 0.5
PB BR PB BR PB
PD-10c Econo-Pacc C seriesd Econo-Columnd XK seriesd
aData as reported by the manufacturers. bAbbreviations: BR, Bio-Rad; PB, Amersham Pharmacia Biotech. Addresses and phone numbers of suppliers are
provided in the SUPPLIERS APPENDIX. cPlastic, disposable. dBorosilicate glass.
Table 8.3.3
Prepacked Columns Suitable for Desaltinga
Exclusion limit Prepacked column
Gel type
Mrb
Radius (Å)c
Maximum sample volume (ml)
Supplierd
Columns for use with a pump Hi Trap Desalting Sephadex G-25 SF Fast Desalting HR Sephadex G-25 SF 10/10 Econo-Pac P6 Bio-Gel P-6
5,000 5,000
14 14
1.5 3.0
PB PB
6,000
15
3.0
BR
Gravity-flow columns NAP-5 Sephadex G-25 M NAP-10 Sephadex G-25 M PD-10 Sephadex G-25 M Econo-Pac 10DG Bio-Gel P-6DG
5,000 5,000 5,000 6,000
14 14 14 15
0.5 1.0 2.5 3.0
PB PB PB BR
250,000 6,000 5,000 40,000
51 15 14 28
0.01-0.05 0.05-0.1 0.075-0.15 0.05-0.1
PB BR PB BR
Spin columns MicroSpin S-200 Bio-Spin 6 Nick Spin Column Bio-Spin 30
Sephacryl S-200 HR Bio-Gel P-6 Sephadex G-25 SF Bio-Gel P-30
aData reported by the manufacturer. bMolecular mass of smallest protein excluded from matrix.
cApparent radius of smallest spherical protein excluded from matrix, calculated as R = 0.808 M 1⁄3, where R is the radius r and Mr is the molecular mass (Hagel, 1989). dAbbreviations: BR, Bio-Rad; PB, Amersham Pharmacia Biotech. Addresses and phone numbers of suppliers are provided
in the SUPPLIERS APPENDIX.
Conventional Chromatographic Separations
8.3.3 Current Protocols in Protein Science
Supplement 14
columns with different diameters that can be packed with bulk media facilitates linear scaleup). A matrix of small particle size will permit preparation of columns of high efficiency (i.e., low zone broadening) at the expense of high pressure drops and greater cost. Also, smaller particles generally have smaller pore volumes. The net effect of the relationships between these parameters is that the prepacked columns listed in Table 8.3.6 have resolving power of the same order of magnitude, although protein fractionation may be carried out faster on the columns that have particles of small size. A final factor to consider in choosing a matrix for protein fractionation is the likelihood of matrix-solute interactions (e.g., as in the case of lectins on gels containing glucose-type structures). Selecting Matrix and Column for Molecular Size Determination A matrix with high resolving power is recommended for determination of molecular size as well as for protein fractionation (see Table 8.3.4, Table 8.3.5, and Table 8.3.6 for
Table 8.3.4
Gel-Filtration Matrices for Protein Fractionation or Size Determinationa
Gel type
Fractionation range (Mr)
Particle size (µm)
Supplierb
Toyopearl HW 40S Superdex 30 prep grade Toyopearl HW 50S Sephacryl S-100 HR Toyopearl HW 55S Superdex 75 prep grade Sephacryl S-200 HR Superose 6 prep grade Superdex 200 prep grade Sephacryl S-300 HR Sephacryl S-400 HR Sephacryl S-500 HR Toyopearl HW 65S Toyopearl HW 75S
100-10,000 200-10,000 500-80,000 1,000-100,000 1,000-700,000 3000-70,000 5,000-250,000 5,000-5,000,000 10,000-600,000 10,000-1,500,000 20,000-8,000,000 20,000-30,000,000 50,000-5,000,000 500,000-50,000,000
25-40 34 25-40 47 25-40 34 47 34 34 47 47 47 25-40 25-40
TH PB TH PB TH PB PB PB PB PB PB PB TH TH
aData as reported by manufacturers. bAbbreviations: PB, Amersham Pharmacia Biotech; TH, Toso Haas. Addresses and phone numbers of suppliers
are provided in the SUPPLIERS APPENDIX.
Table 8.3.5 Empty Columns Suitable for Protein Fractionation or Size Determinationa
Columns
Dimensionsb (length × inner diameter, cm)
HR series G series XK series
20 × 0.5–50 × 1.6 25 × 1.0–100 × 4.4 20 × 1.6–100 × 5
Maximum operating pressure (MPa) 3–5 3-7 0.5
Supplierc PB AM PB
aData as provided by manufacturers. All columns are made of borosilicate glass. bColumn dimensions given here are those suitable for protein fractionation and size
Gel-Filtration Chromatography
determination. Other column dimensions may also be available. cAbbreviations: AM, Amicon; PB, Amersham Pharmacia Biotech. Addresses and phone numbers of suppliers are provided in the SUPPLIERS APPENDIX.
8.3.4 Supplement 14
Current Protocols in Protein Science
Table 8.3.6
Prepacked Columns for Protein Fractionation or Size Determinationa
Dimensions available Supplierd (length × inner diameter, cm)
Fractionation range (Mr)c
Particle size (µm)
Superdex Peptide HiLoad Superdex 30 prep grade
100-7,000 200-10,000
13 34
30 × 1 60 × 1.6 60 × 2.6
PB PB
TSK SW 2000
500-60,000
10
30 × 0.75 60 × 0.75
TH
HiLoad Sephacryl S-100 HR TSK SW 3000
1,000-100,000
47
PB
1,000-300,000
10
Protein-Pak 60
2,000-30,000
10
60 × 1.6 60 × 2.6 30 × 0.75 60 × 0.75 30 × 0.78
Superdex 75 HR 10/30 HiLoad Superdex75 prep grade
3,000-70,000 3,000-70,000
13 34
30 × 1 60 × 1.6 60 × 2.6
PB PB
Bio-Sil SEC 125
5,000-100,000
10
BR
G2000SWXL
5,000-150,000
5
60 × 0.75 60 × 2.15 30 × 0.75
HiLoad Sephacryl S-200 HR TSK SW 4000
5,000-250,000
47
5,000-1,000,000
13
Superose 6 HR 10/30
5,000-40,000,000
13
30 × 1
PB
30 × 0.78
WA
30 × 1
DU
60 × 0.75 60 × 2.15 30 × 0.75
BR TH
Column typeb
60 × 1.6 60 × 2.6 30 × 0.75 60 × 0.75
TH WA
TH PB TH
Protein-Pak 125
10,000-80,000
10
Glass GF-250
10,000-250,000
4
Bio-Sil SEC 250
10,000-300,000
10
G3000SWXL
10,000-500,000
5
Superdex 200 HR 10/30 HiLoad Superdex 200 prep grade
10,000-600,000
13
30 × 1
PB
10,000-600,000
34
60 × 1.6 60 × 2.6
PB
HiLoad Sephacryl S-300 HR G4000SWXL
10,000-1,500,000
47
60 × 1.6
PB
20,000-10,000,000
7
30 × 0.75
TH
Glass GF-450
25,000-900,000
6
30 × 1
DU
aData as reported by the manufacturers. bIn general, matrices of natural polymers (e.g., Superdex, Sephacryl, or Superose) have larger pore volumes than
silica-based materials (e.g., TSK SW, Protein-Pak, Bio-Sil SEC, or GF-250/450). cSelectivity of materials decreases with increased width of the fractionation range and also with decreased pore volume. dAbbreviations: BR, Bio-Rad; DU, DuPont; PB, Amersham Pharmacia Biotech; TH, Toso Haas; WA, Waters. Addresses and phone numbers of suppliers are provided in the SUPPLIERS APPENDIX.
Conventional Chromatographic Separations
8.3.5 Current Protocols in Protein Science
Supplement 14
matrices and columns suitable for both these applications). However, certain other criteria that are important for selecting a matrix and column for size determination differ from those for protein fractionation. First, when analyzing samples containing several proteins of varying molecular size (e.g., protein digests) a matrix of sufficient separation range is needed. As selectivity inherently limits separation range, optimal separation range may not be achieved using a matrix of high selectivity. Second, because peak broadening is not critical as long as the peaks are resolved, short (e.g., 30-cm) columns operated at high flow rates may be used. However, in situations where the effluent is continuously monitored with a mass-sensitive detector (see Basic Protocol 3), complete resolution between peaks is essential. In this case the criteria used for protein fractionation may be applied. BASIC PROTOCOL 1
DESALTING (GROUP SEPARATION) Desalting, or group separation, is used to separate the target protein from low-molecularmass contaminants. The purpose might be to remove salt prior to lyophilization, or to remove low-molecular-mass reagents—e.g., a reductive cleavage mixture from a preparation step (UNIT 11.1) or polybuffer from chromatofocusing (UNIT 8.5). Desalting is also used to change from one buffer to another—e.g., to decrease ionic strength prior to ion-exchange chromatography (UNIT 8.2). The pore size of the matrix is selected to exclude the protein of interest while allowing maximum permeation of the contaminants. This is beneficial in three ways. First, because the protein does not penetrate into the pores, little dilution of the sample will take place except where the sample volume is very small. Second, because the protein is unable to diffuse into the pores, no flow-sensitive zone broadening (UNIT 8.1) that results from slow mass-transfer will take place. Hence, the flow rate may be kept high during the desalting. Third, because the protein and the low-molecular-mass solutes are maximally separated, very large sample volumes may be applied before contamination of the protein band becomes evident. Thus, very large sample volumes may be applied and desalted in a short time without dilution of the sample. The procedure described here uses a closed column system and a pump; annotations explain variations needed in some of the steps if an open-bed column and gravity flow are used. The GF matrix is swelled, the column is packed, and the void volume and total volume of the column are determined in preliminary runs using high- and low-molecular-weight solutes. This information is used to determine the maximum sample volume to apply for the unknown protein to be desalted. Guidelines for estimating approximate sample volumes to use for desalting are also given. Alternative steps are provided for using this procedure either to determine elution volume or to desalt a protein for preparative purposes (or further analysis). Elution volumes must be determined to ascertain the exact void volume and total volume of a column in preparation for desalting. It is also necessary to measure the elution volume for certain protein analyses—e.g., for determining molecular size (also see Basic Protocol 3). Materials GF matrix (Table 8.3.1) or prepacked GF column (Table 8.3.3) with appropriate exclusion limit for protein of interest (also see Strategic Planning and see Critical Parameters) GF desalting buffer (see recipe) Colored marker: 0.2 mg/ml Blue Dextran 2000 or 0.2 mg/ml vitamin B12 Void marker (see recipe) Total liquid volume marker (see recipe) Protein sample to be desalted
Gel-Filtration Chromatography
90°C water bath (optional) Buchner or sintered-glass funnel
8.3.6 Supplement 14
Current Protocols in Protein Science
A
B
C
pump
column extension/ packing reservoir
injector buffer reservoir
column column (with jacket to allow for thermostatation)
adaptor
detector
recorder
end piece
fraction collector
Figure 8.3.1 Equipment used for gel filtration. (A) Simple setup for desalting using an open-bed column made with a Pasteur pipet. (B) Column and attachments. (C) Complete automated chromatography system. Courtesy of Amersham Pharmacia Biotech.
GF chromatography column (see Critical Parameters and Table 8.3.2) with column extension (optional), adaptors, and buffer reservoir (Fig. 8.3.1) Carpenter’s level Peristaltic pump (Fig. 8.3.1) 0.22-µm filter: any 0.22-µm filter for buffer; protein-compatible for sample Detector (optional; Fig. 8.3.1) Chart recorder (optional; Fig. 8.3.1) Sample applicator: sample application loop (e.g., Superloop, Amersham Pharmacia Biotech) or syringe Fraction collector (optional; Fig. 8.3.1) NOTE: Small-scale desalting of multiple samples may be carried out using spin columns and a centrifuge. At present, spin columns are primarily used for preparing microlitersized samples of oligonucleotides; however, they are equally useful for small-volume protein samples. As the procedure is different from that described here, manufacturer’s instructions should be consulted (see Table 8.3.3 for suppliers). Conventional Chromatographic Separations
8.3.7 Current Protocols in Protein Science
Supplement 14
Prepare gel If GF matrix is supplied dry: 1a. Calculate the amount of dry matrix needed from the swelling factor given by the manufacturer and the approximate bed volume of the column to be packed. Add the calculated amount of dry gel to a volume of GF desalting buffer equal to twice the calculated final gel volume (i.e., twice the approximate bed volume of the column). 2a. Carefully stir suspension with a glass rod. Let gel swell overnight at room temperature or for 3 hr in a 90°C water bath, stirring as needed to keep gel in suspension. Do not use a magnetic stirrer, as this may break fragile gel particles.
3a. Allow gel bed to settle, then remove fines and broken beads by decanting the hazy solution. Repeat decantation (4 or 5 times if necessary) until bed settles as a sharp zone, then dilute so that final slurry is 50% (v/v) settled gel and 50% (v/v) GF buffer. Careful removal of fine particles will prevent the nets that support the column end pieces from becoming blocked, and will also permit a higher flow rate through the column.
If GF matrix is supplied preswollen: 1b. Wash gel with an excess of GF buffer on a Buchner or sintered-glass funnel to remove storage buffer. With some products, the gel slurry is obtained by simply shaking the gel bottle, and this can be used to pack the column without further preparation. If this type of gel is used, skip steps 1b to 3b and proceed directly to step 4. Gel particles may adhere to the glass wall of the funnel; therefore, to avoid contamination from other types of gels (e.g., ion-exchange media), it is important to ensure that glassware used is clean.
2b. If gel contains fines, remove by decantation as in step 3a. 3b. Dilute so that final slurry is 50% (v/v) settled gel and 50% (v/v) GF buffer. Pack column 4. Examine column to ensure that it is clean and that the support nets are undamaged. If necessary, supply column with new support nets. 5. Mount column on a stable laboratory stand, using a carpenter’s level to ensure that it is vertical. Equip column with an extension that together with the column will hold the complete volume of the gel slurry (i.e., twice the volume of the settled gel). If <50% of the column volume is to be packed, there is no need for a column extension.
6. Remove air from outlet tubing and end piece of column by injecting GF buffer into outlet tubing (using syringe or peristaltic pump) until support net is covered with 0.5 cm of buffer. Close outlet of column. 7. Inject GF buffer into inlet tubing of adaptor until net is wetted, while holding outlet upwards to ensure that air easily escapes from net. Place adaptor in a beaker containing GF buffer until it is time to use it. 8. Swirl gel slurry from step 3a or b and pour entire slurry into column along a glass rod tilted toward the inner wall of the column or column extension. Fill the remaining space with GF buffer by carefully pouring buffer along glass rod so as to cause minimum disturbance of the gel layer. Put lid on column extension (or put top adaptor on column). Gel-Filtration Chromatography
Small columns (e.g., 1-cm i.d., 5-cm length) to be used for preliminary experiments where optimum resolution is not critical may be quickly prepared by adding the gel slurry to the
8.3.8 Supplement 14
Current Protocols in Protein Science
chromatographic tube, letting the gel settle by gravity, and washing the bed with one column volume of GF buffer before applying the sample.
9. Fill buffer reservoir with GF buffer and place at a level higher than the pump. Connect reservoir to pump using a large-i.d. tube. This step is to ensure a free flow of buffer to the pump head.
10. Purge pump with GF buffer and attach outlet from pump to inlet of column or column extension. Open outlet of column and start pump at flow rate recommended by manufacturer for the gel/column combination used. Continue flow until height of gel bed becomes constant (typically 1 to 4 hr). If no manufacturer’s recommendation is available, use a flow rate 50% higher than that to be used in the separation. If column is to be packed by gravity flow—i.e., by attaching the buffer reservoir directly to the inlet of the column—the actual flow rate must be monitored (e.g., by continuously weighing the effluent, or in the simplest case, by using a graduated cylinder) or controlled by a pump placed after the column. In the latter case it is necessary to check that the free gravity flow exceeds the pump flow—i.e., that the pump is serving to reduce the flow to a constant value. The flow rate caused by gravity depends upon the hydrostatic pressure, determined by the difference in height between the air inlet in the buffer reservoir and the outlet tubing from the column. However, because the pressure drop over the column increases with the height of the packed bed, the free gravity flow rate will decrease during packing if the hydrostatic pressure is kept constant. Gravity flow rate may be increased by lowering the outlet tube or raising the buffer reservoir.
11. Turn pump off, close column outlet, remove column extension, adjust bed height if necessary, and adjust inlet adaptor so that it is flush with the bed surface. Avoid trapping air under the support net. Adjustment of the bed height is recommended only to remove excess gel. This is done by carefully swirling the upper part of the gel surface with a spatula and sucking the excess slurry out with a Pasteur pipet or tubing attached to a pump. If a crater or uneven surface is produced, the gel should be swirled so that a sharp bed surface is obtained before adjusting the inlet adaptor atop the bed. If the bed is too short the column should be repacked, although in most cases the old gel can be reused and added to new slurry prepared as in steps 1a to 3a or 1b to 3b before repacking.
12. Reattach column to pump, open column outlet, and resume flow conditions used in step 10 for 1 hr to stabilize column bed height. Readjust the adaptor so that it is flush with the new bed surface. In some cases it is desirable to lower the inlet adaptor a few millimeters into the gel surface to make sure that there is contact between adaptor and gel. Stabilizing the bed at a higher flow rate than that to be used during the separation will prevent the bed from shrinking as a result of “after-packing” during the run. If a gap appears between the adaptor and the bed surface, step 12 must be repeated at an increased flow rate before experiments are carried out. A space between the adaptor and the bed surface will act as a mixing chamber; the effect on separation will depend on the separation problem at hand.
13. Inspect packed bed visually for cracks, trapped air, and particle aggregates by observing the light scattered from a light source held behind the column. Chromatograph a colored marker (2 mg/ml Blue Dextran 2000 or 0.2 mg/ml vitamin B12) as in steps 20 to 23, and ascertain that the zone produced is horizontal and sharp. Quality of the packed bed need not be as high for desalting as for protein fractionation (see Basic Protocol 2); therefore evaluation of the packed bed may be performed by visual
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inspection. However, it is important for it to be free of cracks and for a colored sample to produce a sharp horizontal zone. Air bubbles would be a major problem and should be avoided at all costs (see Troubleshooting). If column is to be stored, GF buffer containing 0.02% (w/w) sodium azide or 20% (v/v) ethanol as antibacterial agents should be prepared, then 2 column volumes of this mixture should be pumped through the column. The outlet is then closed and the column stored in the buffer/antibacterial agent at room temperature for several months. See recipe for GF buffer for precautions concerning azide.
Prepare and test the GF system 14. Calculate amount of GF buffer necessary for the run and filter this amount plus a 50% excess through a 0.22-µm filter. Adjust pH if necessary and pour filtered buffer into buffer reservoir. If the column is going to be used for a prolonged time, it is good practice to continuously deaerate the GF buffer by heating the buffer slightly with heat from a light bulb, or to use a degasser.
15. Assemble GF system according to manufacturer’s instructions or Figure 8.3.1, placing detector and chart recorder (if used) in line but leaving column off line. Attach buffer reservoir to pump and purge pump with GF buffer with column off line. If the system is run by gravity flow alone, the risk of the column running dry may be eliminated by placing the end of the outlet tubing higher than the inlet adaptor or placing a loop of the tubing from the reservoir with its bend at a lower level than the outlet. Detection of salt may be accomplished in line by conductivity or off line by flame photometry or conductimetry. Colorimetric detection of proteins is readily done at 280 or 254 nm; the latter wavelength provides higher sensitivity. Refer to Chapter 3 for specific methods of protein detection.
16. Connect outlet of pump to column via the injection valve, run system with GF buffer at flow-rate settings to be used for separation, collect fractions with fraction collector, and note actual flow rate of the pump (e.g., by weighing collected fractions or measuring them with a graduated cylinder). If the concentration of the solute is detected by in-line monitoring, use of a two-channel recorder set at different sensitivities (e.g., 100% and 10% full-scale) is recommended to ensure that the peaks of unknown solutes are not off-scale. The actual flow rate of the system must be determined with the column installed, because the flow rate from a peristaltic pump or similar device depends upon the back-pressure of the system.
Determine separating volume of column 17. Determine the void volume of the system (Vo) by running a void marker and obtaining the elution volume according to steps 20 to 23. The sample volume for determination of void volume and total liquid volume (steps 17 and 18) is not critical and may typically be 1% to 2% of the bed volume. However, correction of the calculated elution volume is needed if large sample volumes are applied (see annotation to step 23a). See Figure 8.3.2 for graphical depiction of void and total liquid volume. The markers for void volume and total liquid volume may be mixed and run together. Void volume (Vo) is typically 30% to 33% of the bed volume of a nonrigid (e.g., agarose) gel and 36% to 40% of the bed volume of a rigid (e.g., silica) gel. Gel-Filtration Chromatography
18. Determine the total liquid volume of the system (Vt) by running a total liquid volume marker and obtaining the elution volume according to steps 20 to 23.
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Total liquid volume (Vt) is typically 85% to 95% of the geometric bed volume (πr2L, where r is the inner radius of the column) for a nonrigid gel and 70% to 80% of the bed volume for a rigid gel. Because solute-matrix interactions may occur, it is advisable to check that the total liquid volume is smaller than the bed volume and perhaps run several different total liquid volume markers to confirm the result.
19. Calculate the separating volume of the column (Vi) as Vt − Vo. The separating volume (Vi) is the amount of liquid that will pass through the column from the point where large proteins begin to emerge to the point where small solutes are eluted. It is equal to the pore volume of the bed, which is typically 50% to 65% of the bed volume of a nonrigid gel and 30% to 50% of the bed volume of a rigid gel.
Prepare, apply, and chromatograph the sample 20. Dissolve sample to be desalted in GF buffer. Filter through a 0.22-µm protein-compatible filter. If possible, let sample solution reach room temperature before being applied to column to reduce viscosity effects. This is more important for large sample volumes. The influence of the sample volume on the resolution (see Critical Parameters) is related to the need for column efficiency (see Basic Protocol 2 and UNIT 8.1). Too large a sample volume may ruin the separation of closely eluting proteins in a separation where high efficiency and narrow peaks are required. This is of primary interest in protein fractionation (see Basic Protocol 2), whereas in desalting, conditions are chosen to yield maximum separation of the protein from contaminants. To this end, column efficiency is less important, and very large sample volumes may be applied. See Critical Parameters for guidelines on optimal sample concentration. When GF is performed for analytical purposes, the sample concentration should be as small as possible within the sensitivity of the detection method employed. For preparative purposes, on the other hand, the concentration should be as high as possible. The limiting factor in the latter case is the viscosity of the sample divided by the viscosity of the GF buffer (relative viscosity). At relative viscosities >1.5, the hydrodynamic instability of the trailing edge of the zone may cause distortion of the protein zone (resulting from invasion of less viscous eluant into the more viscous zone; called “viscous fingering”), which will ruin the separation. It is generally safe to use protein concentrations up to 50 mg/ml and in some cases up to 70 mg/ml, whereas concentrations of 100 mg/ml often result in viscous
A280
Vi
Wh
a b
start
Vo
Vr
Vt
Vc
Elution volume
Figure 8.3.2 Chromatogram illustrating the separating volume (Vi) of gel filtration as determined by the void volume (Vo) and the total liquid volume (Vt). The total bed volume (Vc) is equal to the total liquid volume plus the matrix volume. Column efficiency is calculated as N = 5.54(Vr/Wh)2, where N is the efficiency of the column in terms of number of theoretical plates, Vr is the elution volume of the peak, and Wh is the peak width at half peak height. The symmetry of the peak is calculated at 10% peak height as b/a, where b is the width of the tailing part of the peak and a is the width of the leading part of the peak.
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fingering. However, when large sample volumes are applied—e.g., in desalting—the low dilution factor will result in a high concentration of protein in the peak, hence lower concentrations than these may be required to avoid viscosity effects.
21. Open outlet from column, turn on pump, and let two bed volumes of GF buffer pass through the column. Turn on detector and recorder and allow baseline to stabilize. 22. Load sample applicator with the appropriate volume of sample for desalting, not exceeding the separating volume determined in step 19. Switch the sample application valve to the load position and put a start mark on the chart recorder paper (or start a stopwatch if no chart recorder is used). Smaller sample volumes are used for protein fractionation than for desalting, and much smaller sample volumes are used for analysis. Before injecting, flush injector loop with excess sample to avoid trapping air bubbles, which otherwise may end up in the top of the column. If the separating volume of the column has not been determined, the maximum sample volume may be estimated as ∼50% of the bed volume for nonrigid media and ∼30% of the bed volume for rigid media. For a first attempt at desalting, 50% of these volumes should be applied (i.e., 25% and 15% of the bed volume for nonrigid and rigid media, respectively). If an open-bed column is used, gently add GF buffer to the top of the column with a Pasteur pipet or similar device, then open the outlet from the column and continue adding GF buffer until two bed volumes of buffer have passed through. Let the last portion of buffer penetrate the gel without the gel surface becoming dry; this is facilitated by using a top filter on the bed. Gently apply the sample to the clear gel surface or the top filter using a micropipet (or a Pasteur pipet if large volumes of sample are to be desalted).
23a. To determine elution volume: Pass GF buffer through the system at the appropriate flow rate and follow detector response with recorder while collecting fractions. Construct a chromatogram and calculate elution volume as the time from the start point to the apex of the peak for the protein (or other solute) multiplied by the flow rate of the pump (as determined in step 16). If large sample volumes are being applied, the correct elution volume of the peak is obtained by subtracting half the applied sample volume from the apparent elution volume. For determination of void volume as in step 17, the fractions that emerge between the point where a volume of buffer equal to 25% of the bed volume has passed through the column and the point where buffer volume equal to 50% of the bed volume has passed through the column are analyzed (either in line by monitoring detector response or off line by another method; see annotation below). For determination of the total liquid volume as in step 18, fractions emerging between the point where a buffer volume equal to 75% of the bed volume has passed through and that where 125% has passed through are analyzed. To determine the elution volume of an unknown protein sample (as in determining molecular size; see Basic Protocol 3), fractions emerging between the point where a buffer volume equal to 25% of the bed volume has passed and that where 125% has passed through are analyzed. Allowing 125% to pass through will permit the entire peak to be traced.
Gel-Filtration Chromatography
If fractions are to be collected for subsequent off-line analysis to determine where elution occurs (e.g., by photometry at 280 nm to detect Blue Dextran 2000 or proteins, or conductivity to trace the salt peak), start and stop the fraction collector to sample fractions at the volumes stated above. The fraction size should be large enough to permit accurate assay of the solute of interest; often 1 ml or 2% of the bed volume (whichever is smallest) is a suitable fraction size. The prefraction (i.e., the amount of liquid eluted from the point of sample application to the start of fraction collection for analysis) should be sampled in a vessel of known weight. The elution volume is calculated from the volume of the fractions by adding half the volume of the peak fraction and the volume of the prefraction, then subtracting half the sample volume. For dilute aqueous buffers (e.g., 0.5 M) a density of
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1 may be assumed for converting weight to volume. If the experiment will take a long period of time, it is advisable to minimize evaporation by sealing the fraction cups with Parafilm. If an open-bed (gravity flow) column is used, carefully add a volume of GF buffer that corresponds to the void volume of the column reduced by half the sample volume—unless the sample volume is <4% of the bed volume, in which case add 1 void volume of buffer. If the column is operating correctly, the effluent collected should contain no protein. Add GF buffer in ≤1-ml increments if the column has a bed volume <50 ml, collect fractions, assay for proteins, and calculate elution volumes as described above.
23b. To desalt a protein: Pass a volume of GF buffer equal to the void volume reduced by half the applied sample volume through the column, and collect a waste fraction. Place a vessel that can hold 1.5 times the applied sample volume under the column, then add a volume of GF buffer equal to the applied sample volume. Collect the desalted protein. Depending upon the efficiency of the desalting system, these quantities may have to be adjusted (i.e., the volume collected in the second fraction may need to be decreased in order to preclude contamination from substances that elute later). If system effects (e.g., mixing after the column) are substantial or the column is badly packed (e.g., contains cracks in the bed), it will be necessary to pass more GF buffer through the column than the void volume minus half the sample volume before sampling the protein (see Critical Parameters). Figure 8.3.3 shows a sample chromatogram of optimized desalting of 400 ml hemoglobin. The desalting step may be continuously monitored by simultaneous in-line detection of the protein zone by absorbance at 280 nm and in-line detection of the salt peak by conductivity.
24. Wash column with ≥1 column volume of GF buffer containing an antibacterial agent. Close column outlet and store column. See recipe for GF buffer for acceptable antibacterial agents.
protein peak 1.4
A280
salt peak 1.0
10
0.8
8
0.6
6
0.4
4
0.2
2
100
300
500
700
900
mg NaCl/ml
1.2
1100
Elution volume (ml)
Figure 8.3.3 Chromatogram illustrating desalting of proteins by gel filtration. A 4 × 85–cm column packed with Sephadex G-25 is used. The sample consists of 400 ml hemoglobin (protein peak; solid line) in NaCl (salt peak; broken line). Note that the sample volume is close to the pore volume of the packed matrix, i.e., 490 ml. No correction for sample volume has been made. Calculation from the figure yields Vo = 560 ml − 200 ml = 360 ml, which is 31% of the geometric column volume (Vc). Reproduced from Flodin (1961) with permission of the Journal of Chromatography.
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BASIC PROTOCOL 2
PROTEIN FRACTIONATION Protein fractionation refers to the separation of proteins of similar molecular size for purification purposes. The objective may be to separate dimer and higher aggregates from a pharmaceutically active protein or peptide. Requirements for the column and system are therefore substantially higher for protein fractionation than for desalting. The same materials and methodology described for desalting are used for protein fractionation. However, selection of matrix and column (see Strategic Planning) is more critical in this procedure, as are the quality of the column packing and control over experimental conditions (e.g., temperature and flow rate). The protocol includes steps for determining the column efficiency (also see UNIT 8.1) to evaluate the fitness of a packed column for use in protein fractionation. Materials GF matrix (see Table 8.3.4) or prepacked GF column (see Table 8.3.6) with appropriate selectivity for protein of interest (also see Strategic Planning and see Critical Parameters) GF fractionation buffer (see recipe) Colored marker: 0.2 mg/ml Blue Dextran 2000 or 0.2 mg/ml vitamin B12 Low-molecular-weight marker (e.g., 5 mg/ml acetone or 2 M sodium chloride) Protein sample to be fractionated GF chromatography column (see Critical Parameters and Table 8.3.5) Prepare column 1. Prepare GF matrix as described in the desalting procedure (see Basic Protocol 1, steps 1a to 3a or 1b to 3b) using GF fractionation buffer wherever GF buffer is indicated. It is generally difficult to prepare columns of maximum efficiency using a GF medium with a small (e.g., 10-ìm) particle size. The need to scale up the procedure should be considered when selecting medium and column (see Strategic Planning and Critical Parameters). The GF fractionation buffer must be selected in light of the method planned for testing purity of the final protein fraction (step 11), as this may put special restrictions on the buffer (e.g., sodium azide absorbs light at 254 nm and will also interfere with the anthrone reaction used for assaying carbohydrates).
2. Pack the column (see Basic Protocol 1, steps 4 to 12). Because protein fractionation puts high demands on the quality of the packed column, care must be taken in selecting columns of proper design. Columns for protein fractionation are typically 30 to 100 cm long, depending upon the difficulty of the separation and the particle size of the medium used. Columns used must be constructed so that parts coming into contact with buffer and sample are made of appropriate materials. Long tubings and other sources of dead space or mixing chambers should be avoided. The pressure resistance of the column should be compatible with the expected pressure drop of the bed. See Strategic Planning and Critical Parameters for details concerning column selection.
3. Check quality of column packing visually (see Basic Protocol 1, step 13). 4. Assemble and test the system (see Basic Protocol 1, steps 14 to 16). Place system and column in an environment not subject to temperature variations (e.g., the column should not be exposed to direct sunlight). If the system is to be used in a cold room, the influence of temperature on viscosity must be considered. Zone broadening of proteins and the pressure drop of the system are proportional to the viscosity of the buffer, and the flow rate will have to be adjusted accordingly.
Gel-Filtration Chromatography
Successful, reproducible protein fractionation requires a constant flow rate. This may be achieved using a peristaltic or high-precision pump whose flow rate is regularly checked. Continuous monitoring of the effluent is necessary to ensure proper sampling of the fraction
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of interest. Detection may be carried out at 280 or 254 nm to trace proteins containing aromatic amino acids, or at lower wavelength (e.g., 214 or 206 nm) to measure absorption by peptide bonds. Other parts of a protein fractionation system that might be of use include a fraction collector (although in the simplest case it is possible to collect the fraction of interest by using a switch valve) and a chromatography controller, which permits automatic fractionation of the protein of interest.
Evaluate column 5. Chromatograph a colored marker (2 mg/ml Blue Dextran 2000 or 0.2 mg/ml vitamin B12; see Basic Protocol 1, steps 20 to 23) and ascertain that the zone produced is horizontal and sharp. A volume of marker solution 1% to 4% of the bed volume should be applied. The zone produced by a colored sample must be horizontal and sharp for packing to be considered acceptable. If the supplier recommends a different method than this for evaluating column packing, that method should ordinarily be used.
6. Chromatograph a low-molecular-weight marker (e.g, 5 mg/ml acetone or 2 M sodium chloride; see Basic Protocol 1, steps 20 to 23) and construct a chromatogram to determine the column efficiency. The sample volume applied in this step may contribute considerably to the peak width and result in a falsely low value for column efficiency. It is therefore recommended that the sample volume not exceed 0.9%, 0.5%, and 0.4% of the bed volume for columns packed with GF matrices having average particle diameter of 100 ìm, 30 ìm, and 10 ìm respectively (Hagel, 1985).
7. Calculate the number of theoretical plates per column according to the equation N = 5.54(Vr/Wh)2, where N = number of theoretical plates per column, Vr = elution (retention) volume of peak, and Wh = width of the peak at half peak height. See Figure 8.3.2 for a graphical depiction of these variables. The efficiency of the column corresponds to the number of theoretical plates under given experimental conditions. See UNIT 8.1 for discussion of column efficacy.
8. Calculate the asymmetry factor of the peak according to the equation As = (b/a), where a is the width of the leading part of the peak and b is the width of the tailing part of the peak at 10% peak height. 9. Compare the plate number and asymmetry factor obtained for the column with the acceptance limits for these parameters in the manufacturer’s documentation. If no data for expected column performance is given, calculate the plate height (H) according to the equation H = L/N, where L is the length of the packed bed and N is the number of theoretical plates calculated in step 7. As a rule of thumb, H should be about three times the average particle diameter of the matrix for a good column and twice the average particle diameter for an excellent column. Plate heights of five times the average particle diameter or more are generally not accepted for high-resolution chromatography. The asymmetry factor should be 1.0 ± 0.2. The flow rate used will influence the broadening of the solute peak; suggested flow rates for GF matrices of different particle size are given in Table 8.3.7.
Perform chromatography and evaluate results 10. Dissolve, apply, and chromatograph protein sample to be fractionated (see Basic Protocol 1, steps 20 to 23). Start by applying a small fraction of the sample to get a rough idea of the separation and to calculate optimum conditions for the actual run. A sample volume of ∼1% to 4% of the bed volume has been found to be optimal for protein fractionation using a medium of 30-ìm
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particle size, depending upon the processing rate (i.e., amount of substance processed per unit time) of the system (Hagel et al., 1989). As a general rule, a sample volume of 2% of the bed volume can be applied. A chromatogram illustrating protein fractionation where dimers and higher aggregates are separated from the target monomeric form of the protein is given in Figure 8.3.4. Optimum sample volume depends upon many experimental factors; discussion of optimization strategies have been published (Hagel et al., 1989; Hagel and Janson, 1992). In general, the larger the bead size used, the larger the sample volume that may be applied before the initial resolution is lost. This is due to the fact that smaller bead sizes normally result in more efficient columns that produce narrower peaks; hence the influence of sample volume on peak width is more detrimental.
Table 8.3.7
Practical Velocities for Protein Fractionation or Size Determinationa
Nominal eluant velocities (cm/hr) at a support particle size of (µm):
Solute molecular mass
1,000 5,000 10,000 15,000 50,000 75,000 100,000 200,000 300,000
2
5
10
15
30
50
100
507 293 234 195 137 121 109 86 76
203 117 94 78 55 48 44 34 30
101 59 47 39 27 24 22 17 15
68 39 31 26 18 16 15 11 10
34 20 16 13 9 8 7 6 5
20 12 9 8 6 5 4 3 3
10 6 5 4 3 2 2 2 2
aAdapted from Hagel (1989) with permission from VCH Publishers.
monomer
A280
0.011
0.010
oligomer dimer
0.009 6
8
10
12
14
time (min)
Gel-Filtration Chromatography
Figure 8.3.4 Chromatogram illustrating protein fractionation by gel filtration. A Superdex 75 HR 10/30 column is used. The sample consists of recombinant human growth hormone. This figure is an enlarged section of the original chromatogram displayed to illustrate complete resolution between monomer and dimer. Reproduced from Hagel (1993) with permission of the publisher. Courtesy of B. Pavlu, Kabi-Pharmacia Peptide Hormones, and H. Lundström, Pharmacia Biotech.
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11. Check the purity of the collected fraction(s). Suitable methods of determining purity include HPLC and electrophoresis (see UNITS 10.1-10.4).
DETERMINATION OF MOLECULAR SIZE The third important application of gel filtration is in determining molecular size. To accomplish this, a GF column is calibrated with reference samples of known molecular size (see Support Protocol). Gel filtration is then carried out on an unknown protein solute as in Basic Protocols 1 and 2, and the molecular size of that solute is then inferred from the calibration curve, which relates elution volumes or distribution coefficients on the calibrated column to the molecular sizes of the different reference samples. If the relationship between the molecular size and mass is known, the mass of the solute may be calculated. However, because the shapes of biological molecules vary considerably (e.g., proteins may be spherical or ellipsoidal, or in some cases more or less rod-shaped), the calculation of molecular mass (Mr) from elution volume must be done with great care. If the shape of the molecule under study is not known, the experiment will yield only its estimated solvated size. The influence of solute shape on size may be exemplified by the fact 1 that the radius of gyration (one estimate of size) is proportional to Mr ⁄3 for spherical molecules 1 ⁄2 (e.g., some proteins), to Mr for molecules shaped like flexible coils (e.g., dextrans and denatured proteins), and to Mr for rod-shaped molecules (e.g., fibrous proteins and DNA fragments of intermediate length). Thus, gel filtration is very suitable for estimating the molecular size but generally not the molecular mass of unknown solutes.
BASIC PROTOCOL 3
Materials GF matrix or prepacked GF column (see Table 8.3.4 and Table 8.3.6; see Strategic Planning; see Critical Parameters) GF fractionation buffer (see recipe; note variations for size determination) GF chromatography column (Table 8.3.5) High-precision pump Protein sample for size determination 1. Prepare gel, pack column, then assemble and test GF system (see Basic Protocol 1, steps 1 to 16) using GF fractionation buffer wherever GF buffer is indicated. For high-precision analytical gel filtration, the temperature of the chromatographic bed is often kept constant by using a jacketed column and circulating water from a temperaturecontrolled water bath. This makes the instrument setup more complicated, and the need for this precaution must be assessed on a case-by-case basis (e.g., if the column is calibrated at every other run, as is possible with high-speed gel filtration, there is little need for temperature control). A high-precision pump must be used to ensure accurate and constant flow rate, which is of utmost importance for achieving reliable results in molecular size determinations. In addition to a UV detector for determining protein concentration, a detector for viscosity and/or light scattering—e.g., one employing multiple-angle laser light scattering (MALLS)—may be used for assay of molecular size or (indirectly) molecular mass. A mass spectrometry detector—e.g., one employing electrospray ionization mass spectrometry (ESI-MS)—may be used for assay of molecular mass. If denatured standards are to be used in the calibration to eliminate the influence of molecular shape (see Support Protocol), 6 M guanidine hydrochloride (see recipe for GF buffers in Reagents and Solutions) is used as the GF fractionation buffer in this and subsequent steps. Once a column has been calibrated (see Support Protocol), it is important to check column performance by determining the elution volume of one of the reference samples used for calibration as part of the routine testing of the system.
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2. Determine the void volume (Vo) and the total liquid volume (Vt) for the system (see Basic Protocol 1, steps 17 and 18). 3. Calibrate column (see Support Protocol). Check flow rate during calibration by sampling effluent and weighing fractions. Calibration of the column (and chromatography of the sample) may be carried out under denaturing conditions to eliminate the effects of molecular shape. Variations in procedure are described in the Support Protocol. There are no general restrictions on the flow rate as long as the protein of interest separates from contaminating solutes (ideally, baseline separation should be achieved). A high flow rate will not influence the qualitative information sought—i.e., the position of the peak maximum. Thus, qualitative information may be obtained by very fast gel filtration; however, at high flow rates the danger of shear effects on elongated solutes must be considered. If a high-precision pump is used, there is less need to check the flow rate for every run (though in the author’s experience this is a good practice because any pump may malfunction). If constant flow rate can be taken for granted, time may be substituted for volume in the calculations. For both the calibration (see Support Protocol) and the run to determine molecular size of the unknown sample, the elution volume should generally be corrected for the contribution from the sample volume (see Basic Protocol 1, step 23a). However, if the sample volume is kept constant throughout the entire investigation it may arbitrarily be regarded as a part of the dead volume of the system, and no correction is necessary.
4. Apply, elute, and chromatograph unknown sample (see Basic Protocol 1, steps 20 to 23). Prepare a chromatogram and calculate elution volume. Check flow rate by sampling the effluent and weighing fractions. The same applied sample volume should be used for the unknown sample as for the size standards (see Support Protocol). If denatured standards were used in the calibration to eliminate the effects of molecular shape (see Support Protocol), the sample must be denatured. Elution is then carried out using 6 M guanidine hydrochloride as the GF buffer. Often the calculations are performed on a time basis using an integrator or chromatography software (Amersham Pharmacia Biotech).
5. Calculate molecular size of unknown sample components using the calibration graph constructed in performing the support protocol. If two different detectors were used for the reference standards and the sample (e.g., a refractive index detector for calibration with dextran standards and a UV detector for the protein sample), a correction factor for the difference in volume between the two detectors must be used in the calculations. The estimate of molecular size will be correct only if there are no sample-matrix or sample-sample interactions (in fact, gel filtration has been used to study these self-association processes). The size estimate will be an apparent gel-filtration size which, together with data about the molecular mass (e.g., from mass spectrometry), will yield information about molecular shape.
6. Continue to run GF buffer through column until the next run is to be made.
Gel-Filtration Chromatography
It is important to continue flushing the column to maintain column characteristics and avoid the need for recalibration (see Critical Parameters). If the next run is not to be made for a long time, the flow rate may be reduced and buffer recirculated through a filter to the buffer reservoir (see Critical Parameters). Before reusing the column after a long period of rest, the validity of the calibration curve should be checked by running two or three reference samples covering the range of interest and calculating their molecular size from the old calibration curve. The apparent size should agree with the assigned size within ±5% error.
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COLUMN CALIBRATION Molecular size standards for column calibration should ideally be of the same shape and type as the molecules under study (e.g., globular proteins, fibrous proteins, or peptides). In cases where this requirement cannot be met, the column can be calibrated with other suitable reference substances of known size (e.g., dextrans). An apparent size may thus be assigned to the unknown and the actual size then inferred from the relationship between the shape of the different molecules. It is thus possible to use dextran as a reference for the estimation of Stokes radius for globular proteins (Hagel, 1993). However, a general shape parameter governing separation of all types of molecules by gel filtration has not yet been found (Dubin et al., 1990).
SUPPORT PROTOCOL
When protein standards are used, the calibration curve will reflect small natural variations in the molecular shapes of proteins, which will decrease the precision of the curve. However, the procedure described here is very simple and will readily yield an apparent molecular mass or molecular weight, which in many cases is quite satisfactory. The influence of protein shape on the calibration curve may be eliminated by disruption of noncovalent bonds with denaturing solvents (e.g., guanidine hydrochloride). Disulfide bonds are broken by reductive cleavage with a reagent such as DTT, and reoxidation is prevented by carboxymethylation. This treatment converts the shape of sample and reference proteins to random coils. The physical size of a random coil is larger than that of a compact molecule of equal mass; therefore a more porous gel is needed to separate the former species. Detergents may also be used to denature proteins. However, it must be noted that a protein-detergent complex is larger than the protein and that ionic detergents will introduce charged groups. Also, information about size or molecular mass obtained using detergent-denatured proteins may not be relevant to the native protein. As these types of treatment are frequently used to prepare proteins for analytical electrophoresis, the reader is referred to UNITS 10.1-10.4 for further details. The influence of variation in shape on the calibration curve may also be eliminated by using as molecular size standards reference substances from a homologous class of nonprotein polymers—e.g., dextran or polyethylene glycol. Dextran has been used extensively as reference substance for calibration of gel-filtration columns. Calibration can be done by running a number of narrow fractions or one sample of broad size distribution. Additional Materials (also see Basic Protocol 3) Calibration standards (see recipe and Table 8.3.8) 6 M guanidine hydrochloride (APPENDIX 3A) In-line refractive-index detector For calibration with native proteins 1a. Calibrate the column using protein standards (see Table 8.3.8) as in steps 20 to 23 of Basic Protocol 1 and construct a chromatogram. It is practical to mix a number of the calibration standards and chromatograph them simultaneously so that the calibration may be completed in a few runs (see example in Figure 8.3.5).
2a. Plot the elution volume (Vr; i.e., the volume on the chromatogram from the injection point to the apex of the peak for a particular size standard) versus the logarithm of the molecular size, log10R, where R is the molecular radius of the calibration standard. Table 8.3.8 gives the molecular radii of commonly used calibration standards. A sigmoid curve should be obtained that is approximated by a straight line in the middle section (see Fig. 8.3.5).
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If the sample volumes are not held constant, the elution volume must be normalized by setting the injection point at half injected volume. The calibration curve is often made system-independent by plotting distribution coefficient (KD) versus log10R. The distribution coefficient is calculated as KD = (Vr −Vo)/(Vt−Vo), where Vr is the elution volume for a particular peak, Vo is the void volume for the column, and Vt is the total liquid volume for the column. For substances belonging to a homologous series (e.g., dextrans), it is meaningful to plot elution volume versus the logarithm of the molecular mass (log10Mr). Where substances of different chemical nature are being used to calibrate the column, the preferable way to
A 0.4 5
4 0.3
2
6
3
A280
1 0.2 0.1 0 100
150 200 250 Elution volume (ml)
B
Elution volume (ml)
250 220 190 160 130 100 1.1
1.2
1.3
1.4
1.5
1.6
1.7
1.8
Log10 R
Gel-Filtration Chromatography
Figure 8.3.5 Calibration of GF column for determination of protein size. A K 26/70 column packed with Sephadex G-200 SF is used. (A) Chromatogram obtained for six size standards: 1, catalase; 2, aldolase; 3, bovine serum albumin; 4, ovalbumin; 5, chymotrypsinogen A; 6, ribonuclease A. (B) Calibration curve obtained by plotting the logarithms of the molecular radii (log10R) of the standards (x axis) versus the elution volumes for peaks 1, 2, 4, 5, and 6 from panel A (filled circles). The curve was used for evaluating the size of bovine serum albumin (filled square; elution volume corresponds to peak 3 in panel A), giving an apparent size of 34.8 Å. The excellent agreement with the nominal molecular size value of 35.5 Å may be incidental. Repeating the procedure for peak 4 in panel A showed that a deviation of ∼3 Å from the nominal value may be expected for proteins of globular shape. Panel A reproduced from Pharmacia Biotech (1991) with permission from the publisher.
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calibrate the column is by using the hydrodynamic volume, Vh. This is calculated according to the equation Vh = (η)(Mr)/Nν, where η is the intrinsic viscosity, Mr is the relative molecular mass, N is Avogadro’s number (6.02 × 1023 molecules per mole), and ν is a shape factor, which for a spherical solute is 2.5. It is wise to check the accuracy of the calibration by deleting one point at a time from the calibration data set and calculating the size or mass of the protein represented by that point from the calibration curve obtained with the remaining data. The average absolute difference from the nominal value indicates the accuracy of the calibration (see Fig. 8.3.5).
For calibration with denatured standards 1b. Denature calibration standards according to a standard denaturing protocol. 2b. Carry out steps 1a and 2a above, using a column equilibrated in 6 M guanidine hydrochloride and using 6 M guanidine hydrochloride as the GF buffer in the cross-referenced steps of Basic Protocol 1. Where denatured standards are used, denaturing conditions must be maintained in chromatography of the sample (see Basic Protocol 3).
For calibration with nonprotein polymer standards 1c. Using an in-line refractive index detector, run system until a stable baseline is obtained. Chromatograph a number of dextran calibration standards (see Basic Protocol 1, steps 20 to 23) that will give calibration points surrounding the protein under study, to obtain their elution volumes or distribution coefficients. The proper number and size of calibration standards to be run may be quickly found by running first the unknown sample and then the necessary calibration samples that will be eluted close to the unknown protein. Apply each standard individually or as a mixture of two dextrans of high and low molecular mass. Using a two-channel recorder set at two different sensitivities may prevent the peak from running off scale.
2c. Plot the elution volume (Vr) or distribution coefficient (KD) versus the logarithm of the viscosity radius of dextran (log10Rvis). The viscosity radius of dextran, Rvis, is related to the molecular mass (M) according to the equation Rvis = 0.271 × M0.498. Because dextran is a polydisperse polymer, the value of M used here is the molecular mass of dextran that corresponds to the peak apex, sometimes denoted Mp. When two different detectors are used for calibration and chromatography of the sample— e.g., a refractive index detector for dextran standards and a UV detector for the protein sample—the difference in volume between the two detectors must be taken into account in calculation of elution volume. Another procedure where the entire separation range of interest was calibrated in one run, using an integral calibration method, has been described (Hagel and Andersson, 1991; Hagel, 1993). Unfortunately, this simple and powerful method cannot yet be performed on routine basis because it requires data for the molecular mass distribution of the sample, and this information is, at the moment, not readily available in most cases.
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Calibration standards Table 8.3.8 provides a compilation of commercially available molecular size calibration standards and their suppliers. Some of these are available as kits (e.g., from Amersham Pharmacia Biotech). Protein standards should be prepared in the GF buffer that will be used in the molecular size determination. To prolong the life of the column, samples should be filtered through a protein-compatible filter prior to injection. The exact concentration to be prepared depends on the additives present (e.g., sucrose as stabilizer), detector sensitivity, and the zone broadening of the column. Concentrations of proteins in the range from 1 to 5 mg/ml are frequently used for calibration. It is possible to mix several standards to calibrate the separation range of interest in a few runs (it is seldom necessary to calibrate the entire separation range of the column). Table 8.3.8 also lists some dextran fractions that are useful for calibrating the column in hydrodynamic volume, which makes comparing solutes of different shape easier.
GF buffers For desalting: Use a volatile buffer—e.g., 0.05 M ammonium hydroxide, 0.05 M acetic acid, 0.05 M ammonium acetate, or 0.5 M ammonium bicarbonate—unless the step is for buffer exchange, in which case use the desired final buffer. Distilled (Milli-Q-purified or equivalent) water may be preferred to buffer as an eluant in some cases. This will yield adequate results with noncharged solutes; however, ionic interactions with the small amount of charged groups on the gel surface may disturb the purification of slightly charged molecules when water alone is used. Addition of a volatile bacteriostatic agent—e.g., chlorhexidine—may sometimes provide enough ionic strength to suppress such interactions.
For protein fractionation: Use 0.05 M sodium or potassium phosphate (pKa = 2.1, 7.2, 12.4), 0.05 M Tris⋅Cl (pKa = 8.1), or 0.05 M sodium or potassium acetate (pKa = 4.8). The buffering capacity of these buffers is adequate at a pH within ±0.5 units of the pKa value. If the protein is to be freeze-dried, use of a volatile buffer or water is advantageous. It is sometimes necessary to add 0.1 M sodium chloride to completely eliminate ionic interactions. If parts of the system that come into contact with the buffer are composed of steel, the sodium chloride should be replaced with 0.05 M sodium sulfate to prevent halide-induced corrosion. In some cases the ionic strength of the buffer will affect the sample, e.g., by disrupting aggregates. This effect can be eliminated only by changing the ionic strength.
For determining molecular size: Use the same buffers as for protein fractionation but select one that will preclude solute-matrix interactions (see Support Protocol). If the column is to be used for a long period, add preservatives to prevent bacterial growth. Make sure the buffer is compatible with the detection step (e.g., use a buffer with low salt content if mass spectrometry will be used for detection). If calibration is to be conducted under denaturing conditions, 6 M guanidine hydrochloride is used as the GF buffer. Guanidine hydrochloride (APPENDIX 3A) is a better denaturing agent than urea.
For all buffers: Add antimicrobial agent if buffer will be used for a long time, adjust the pH of the buffer, filter through a 0.22-µm filter, then recheck pH and readjust if necessary. Gel-Filtration Chromatography
It is also recommended to degas the buffer prior to prolonged use. It is common practice to prepare fresh buffer once a week, but if an antimicrobial agent has been added it may be continued
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stored for 1 month or more. Antimicrobial agents used for GF buffers include ethanol added to a concentration of 20% (v/v), sodium azide added to a concentration of 0.02% to 0.05% (w/v), or chlorhexidine added to a concentration of 0.002% (w/v). Ethanol is not recommended for use with Sephadex gels as organic solvents cause the bed to shrink when used with this matrix. IMPORTANT NOTE: Sodium azide is a very potent antimicrobial agent and therefore widely used. However, handling of azide, especially the dry powder, requires special attention; please refer to information from the supplier for safety precautions. Frequent exposure to the dry powder may be avoided by preparing a stock solution (e.g., 20% w/v) from which aliquots (e.g., 1 ml azide stock solution to 1 liter GF buffer) may be drawn conveniently with a micropipet. Azide may form explosive salts in lead-pipe waste disposal systems; collect the effluent in waste bottles.
Total liquid volume marker 5 mg/ml acetone, 10 mg/ml sodium azide, 0.1 mg/ml cytidine, 10 mg/ml sodium nitrate, and 2 M sodium chloride have been used for determination of total volume. A negative pulse from injection of water can also be used. In cases where interactions between matrix and solute must be eliminated, deuterium oxide can be used. Acetone and cytidine are detected by UV absorbance at 280 nm, azide at 254 nm, and nitrate at 254 nm. Sodium chloride can be detected by flame photometry, conductivity, a sodium or chloride electrode, or precipitation of chloride with silver nitrate solution. A negative pulse from injection of water is detected by measuring the absorbance or refractive index of the buffer or by monitoring the conductivity of the effluent. Deuterium oxide is detected with a refractive-index detector.
Void marker A suitable substance for determining void volume is a protein of the correct size (not too large and not too small), which may be selected by running a series of large proteins and observing which of them start to permeate the matrix. Other substances that have been used include 2 mg/ml Blue Dextran 2000 and DNA. Blue Dextran 2000 has the advantage of being visible to the eye. Disadvantages are that small amounts of free dye may stick to the gel and a broad permeating peak may be visible in addition to the sharp excluded peak if the pore size of the gel is large enough (e.g., as with Sepharose 4B). This occurs because of the polydispersity of dextran. If the molecule is too large, so-called secondary exclusion effects, in which the sample is excluded from the narrow space where beads intersect, will yield a falsely low void volume. However, this phenomenon is only important in high-precision gel filtration and rarely has to be considered in desalting.
COMMENTARY Background Information Although separation of molecules according to size had been noticed in the mid-1950s (Lindqvist and Storgårds, 1955; Lathe and Ruthven, 1956), the discovery of gel filtration (GF) may be attributed to Porath and Flodin (1959), who realized the potential of the technique and were the first to report using it to desalt proteins. Their discovery provided the basis for a new separation technology that rapidly gained widespread acceptance owing to the commercial availability of a suitable crosslinked gel-filtration matrix—Sephadex—introduced in 1959. Remarkably enough, after 35 years this matrix is still one of the premier choices for desalting of proteins (see Table 8.3.1). The use of gel filtration for determina-
tion of molecular size of proteins was described later (Andrews, 1970). Gel filtration was also rapidly established as an important tool for determining the molecular mass distribution of polymers in aqueous (Granath and Flodin, 1961) and organic solutions (Moore, 1964), for which suitable methods were lacking at that time. Moore used cross-linked polystyrenebased media for his application, naming the technique gel-permeation chromatography. Thus, within a few years after the discovery of gel filtration, applications of the technique basically covered the present range. During the following years, the focus was directed to investigating the mechanism behind gel filtration. Theories suggesting that the separation was regulated exclusively by diffusion
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of molecules were soon replaced by the explanation, accepted today, that the process is driven by loss of conformational entropy when free permeation of the matrix by a solute is hindered by steric factors (Casassa, 1967). Of course, diffusion affects peak width and thus also the degree of separation, but diffusion does not influence the elution volume of a solute. The definite impact of steric exclusion on the separation has led to the proposal to rename the technique “steric exclusion chromatography” (ASTM, 1978), but this term has not gained wide acceptance. Another generic designation Table 8.3.8
sometimes used in the literature is size-exclusion chromatography (SEC). It is important to realize that all of these designations refer to the same basic separation principle, which the majority of protein chemists refer to as gel filtration. Development of gel-filtration media was focused for a long period on natural polymers or hydrophilic polymers. The late 1970s saw the introduction of silica-based materials, which combined rigidity and small particle size to enable fast gel filtration. A few years later, natural polymers of enhanced rigidity were
Molecular Size Standards for Calibration of Gel Filtration Columns
Standard Proteinsb Thyroglobulin, bovine thyroid Ferritin, horse spleen Catalase, bovine liver Gamma globulin, bovine Aldolase, rabbit muscle Transferrin Albumin, bovine serum Ovalbumin, chicken Ovalbumin, hen egg Chymotrypsinogen A, bovine pancreas Myoglobin, equine Ribonuclease A, bovine pancreas Non-protein standard Vitamin B12 Dextransc Dextran T-10 Dextran T-40 Dextran T-70 Dextran T-500 Dextran 1 Dextran 5 Dextran 12 Dextran 25 Dextran 50 Dextran 80 Dextran 150
Molecular mass (Mr)
log10(Mr)
669,000 440,000 232,000 158,000 158,000 81,000 67,000 44,000 43,000 25,000
5.825 5.643 5.365 5.199 5.199 4.908 4.826 4.643 4.633 4.398
17,500 13,700 1,350 8,100 23,600 33,000 370,000 1,080 4,440 9,980 21,400 43,500 66,700 123,600
Molecular size (radius, Å)a
Suppliera
85.0 61.0 52.2 — 48.1 35.5 — 30.5 20.9
PB PB PB BR PB SI PB BR PB PB
4.243 4.137
— 16.4
BR PB
3.130
—
BR
23 40 50 150 9 18 26 39 55.3 68.4 93.1
PB PB PB PB PC PC PC PC PC PC PC
aAbbreviations and symbols: —, not stated by manufacturer; BR, Bio-Rad; PB, Amersham Pharmacia Biotech; PC, Pharmacosmos; SI, Sigma. Addresses and phone numbers of suppliers are provided in the SUPPLIERS APPENDIX. bThe size of proteins may vary according to the source of information and the methodology used for determination—e.g.,
a value of 67.1 Å has also been assigned to ferritin (de Haen, 1987). cDextran is a polydisperse polymer and values given here—e.g., peak molecular mass—may vary from lot to lot. In cases
where peak molecular mass is not stated by the supplier, the column is calibrated as elution volume versus M = (Mw⋅Mn) ⁄2, where Mw is the weight-average molecular mass and Mn is the average molecular mass. The molecular size of dextran is calculated according to the equation Rvis = 0.271 × M 0.498, where Rvis is the viscosity radius and M is the molecular mass (Hagel, 1989). 1
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introduced, which made it possible to pack columns with matrices of 10-µm bead size. To distinguish these media from the traditional type, the designation of high-performance sizeexclusion chromatography (HPSEC) was introduced. However, it has been recently pointed out that the advantage of HPSEC media lies not in performance (i.e., traditional media packed in long columns may have equal or even better resolving capacity) but in speed of operation (Hagel, 1992). In addition, none of these materials have the outstanding selectivity shown by some of the traditional matrices (e.g., Sephadex and Bio-Gel P). Therefore, development of matrices for gel filtration has now been directed toward increasing their selectivity, which is done by decreasing the apparent width of the pore-size distribution of the matrix. Development is also geared toward tailoring the apparent pore size of matrices for the separation of molecules in specific interesting areas of application (e.g., for polishing of recombinant peptides). Recently, some of these modern media have become available for separation of peptides and proteins of moderate size (Hagel, 1993). Even though the theoretical framework for the separation mechanism in gel filtration is well understood, there are still some uncertainties as to which molecular size estimate the elution volume reflects (Dubin et al., 1990). It is generally agreed that Stokes radius is a poor measure of molecular size, especially if different types of molecules are being studied. The hydrodynamic volume seems to be a general size parameter for polymers, globular proteins, and DNA of a certain length (Potschka, 1987). It appears that more work needs to be done to resolve this fundamental issue. Until recently, development of instrumentation for gel filtration has been modest. Highprecision pumps and automated systems for injection, fraction collection, and calculation were introduced more than a decade ago. New equipment and technology being introduced includes multiple-angle laser light scattering (MALLS) detectors and in-line viscosity detectors, which provide information about the shape and size of molecules, and electrospray ionization mass spectrometry (ESI-MS), which has the power to provide in-line detection of molecular mass. ESI-MS combined with highresolution gel-filtration columns provides a very powerful system for characterizing protein dimensions. Molecular size of solutes may also be determined with an in-line light-scattering or viscosity detector combined with a concentration-sensitive detector. It is now possible to
obtain the exact molecular mass of small-tomedium-sized proteins by connecting an inline mass spectrometer to the GF column. The advantages of using a mass-sensitive detector are that the system does not need to be calibrated, fluctuations in flow rate are not detrimental, and solute-matrix interactions or solute-solute interactions will not affect the molecular mass obtained. In this case, the column acts as a high-resolution separating device to resolve the sample components prior to determination of molecular mass by mass spectrometry.
Critical Parameters Optimization and scaleup of desalting Optimization of desalting starts with selection of a matrix of suitable exclusion limit with maximum pore volume and proper particle size. The bed volume is dictated by the sample volume to be purified. The actual elution protocol to be used depends upon the dispersion of the chromatography system, which in turn is related to the particle size and the column length. The particle size will influence the pressure drop over the column; this may be a limiting factor determining the maximum possible flow rate, the maximum column length, and the possibility of scaling up to columns of large diameters (i.e., the pressure specification is often reduced for columns of large diameter). However, in this case the column length can be traded off for an increase in column diameter to keep the bed volume high (also see Strategic Planning, Selecting Matrix and Column for Desalting). To maximize the sample volume that can be applied to a column A gel matrix should be selected from which the protein is excluded (see Strategic Planning) and in which the contaminants are eluted close to the total liquid volume (see Basic Protocol 1). It should also have a small matrix volume. The maximum sample volume that can be applied to a column is directly related to the pore volume of the packed bed (Vi). The pore volume is proportional to the geometric column volume. As a rule of thumb, the maximum sample volume that may be applied for desalting is 25% to 40% of the bed volume for the materials listed in Table 8.3.1. The lower figure applies to small columns (e.g., <10-ml bed volume) and the higher figure to large columns (e.g., 100-ml bed volume). After the bed length
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is determined on basis of the acceptable pressure drop (which is proportional to the bed length), the sample capacity is maximized by increasing the diameter of the column. This is done because the applicable volume is directly proportional to the cross-sectional area of the column. Limitations in applicable column length may be eliminated by using columns in series (“stacked columns”). The maximum volume that theoretically can be applied for desalting is equal to the separating volume of the system, which is calculated by subtracting the void volume from the total volume (see Basic Protocol 1 for steps to determine these quantities). To account for extra zone-broadening effects, it is generally advisable to apply not more than 80% of the separating volume or even less—e.g., 50%—for small columns in which eddy dispersion (see discussion below) will contribute proportionally more to the zone broadening. To optimize the yield of desalting Sampling of the protein fraction must be adjusted according to the zone broadening of the sample (i.e., the width of the protein zone). For a well-packed column with a plate height (see Basic Protocol 2) roughly equal to two particle diameters, this parameter may be estimated as the sample volume plus the dispersion of the zone that occurs through different pathways in the packed bed—i.e., eddy dispersion. The peak width (wb) is described by Equation 8.3.1: 1
2dp 2 w b = 4(Vt ) L
would in this case theoretically decrease the protein zone from 500 to 475 ml, at the expense of a pressure drop over the column three times higher than with the coarser medium. To minimize the time of the separation Use the highest possible flow rate for the desalting step. High flow rates are suitable for desalting because eddy dispersion effects are rather insensitive to flow rate. The limiting factors on the flow rate are the rigidity of the matrix and the pressure limitations of the system used. Because media with small pore volume are generally more rigid than those with large pore volume, the disadvantage of a smaller separating volume may be offset by increasing the applicable flow rate to keep productivity high when working with the former type of medium. Incomplete removal of fine particles from the medium prior to packing may reduce the flow rate of the column after use as a result of partial blockage of the outlet filter. Operating the column with the flow direction reversed may help to increase the applicable flow rate in such situations. The total process time for repetitive runs to desalt large volumes may be further minimized by utilizing the dead time of the void fraction. To do this, samples are added, not after total liquid volume + sample volume + eddy dispersion volume, but after subtracting the void volume from the total process volume (i.e., the next sample is added before the salt peak of the previous sample has emerged). The saving in time may be in the range of 20% to 30%, but this practice can only be recommended for well-understood processes.
Equation 8.3.1
Gel-Filtration Chromatography
where Vt is the total liquid volume of the column, dp is the particle size of the matrix, and L is the length of the packed bed. Thus, for a 5-ml NAP-5 column (0.9 × 2.8–cm bed dimensions) packed with Sephadex G-25 medium-particle matrix (particle size 175 µm), eddy dispersion adds ∼0.6 ml to the sample volume. This is roughly equal to the maximum sample volume (i.e., the separation volume; see Basic Protocol 1) and thus the eluted fraction is twice the sample volume. However, for a much larger 4 × 85–cm column packed with Sephadex G-25 coarse-particle matrix, which has an estimated particle size of 300 µm, the eddy dispersion adds ∼100 ml to the originally applied sample volume of 400 ml, giving a protein zone of 500 ml (see Fig. 8.3.3). If G-25 medium-particle matrix were used, the smaller particle size
Optimization and scaleup of protein fractionation There are two basic ways to optimize the resolution of a separation. One is to increase the peak-to-peak distance and the other is to decrease the peak widths. The resolution for gel filtration is described by Equation 8.3.2:
1 M r1 1 L2 1 Rs = 4 log10 b M r 2 Vo H + KD Vi Equation 8.3.2
where Mr1 and Mr2 are the molecular masses of the two sample components being separated, b is the slope of the selectivity curve
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(−∂KD/∂logMr), Vo is the void volume of the column, Vi is the pore volume of the bed, L the bed height, and H the plate height of the column (i.e., the term L/H = N, the plate number). The impact of these parameters will be described below. To increase the peak-to-peak distance 1. A matrix of high selectivity should be chosen. The most important factor for optimizing protein fractionation is selectivity, which is the inherent ability of the matrix to resolve proteins of similar size. This factor is a function of the pore-size distribution of the matrix. Selectivity is often illustrated by a plot of the elution volumes of a series of reference substances versus the logarithm of the molecular size or molecular mass of those substances. A function of the elution volume (e.g., the distribution coefficient; see Basic Protocol 3) may be substituted for the elution volume in constructing a selectivity curve. The steeper the selectivity curve (i.e., the greater the absolute value of the slope b, defined above), the better the resolvability of proteins differing in size. The selectivity of different gels can be judged from the fractionation range stated by the manufacturer. Gels with a small fractionation range (in terms of logMr) will generally have higher selectivity than gels with a large fractionation range. 2. A gel of high pore volume should be selected. The pore volume of a matrix is often related to the particle size. Smaller particles need to resist a higher friction force from the high flow rates that are used with matrices of smaller bead size; therefore the matrix volume is generally larger and pore volume smaller when using these matrices. A traditional softgel matrix may have up to 98% pore volume, whereas rigid matrices such as silica may have as low a pore volume as 52% (of the bead volume). 3. A column of sufficient length (L) should be selected. The resolution between peaks is proportional to the square root of the column length. The column length chosen for protein fractionation is usually optimized with respect to the particle size of the matrix—i.e., a 100-cm column is often chosen where the particle size is 100 µm, a 60-cm column where the bead size is 30 µm, and a 30-cm column where the bead size is 10 µm. To decrease peak width 1. A material that yields a small void volume (Vo) should be selected. Reducing the void
volume will decrease the dispersion of the protein band (as will minimization of all extra column dead volumes). Irregularly shaped or rigid particles generally yield beds with larger void volumes than spherical, nonrigid beads. 2. A material of small particle size should be used, as this will affect the plate height (H; also see Basic Protocol 3). Zone broadening of proteins within a column is directly proportional to the square of the particle size. Therefore, smaller particles will yield smaller peak widths unless very large sample volumes are applied or large system effects are causing dispersion of zones. 3. The separation should be run at optimum flow rate, as this will affect the plate height (see Basic Protocol 3). Zone broadening is caused by dispersion within the void volume of the chromatographic bed, or by nonequilibrium effects that occur in the separation process itself when the mobile-phase velocity in the extraparticle space is too high as compared with the diffusivities of the proteins and the diffusion distances in the matrix. Flow rates that will allow minimum zone broadening and complete equilibrium will in most cases be impractically low. Table 8.3.7 lists practical flow rates for different solute molecular weights and matrix particle sizes, which represent compromises with regard to these parameters. Dissolving the sample Care must be taken to assure that the sample is completely dissolved in the GF buffer. Conditions must be chosen so that spontaneously formed aggregates are dissolved and aggregates that are to be removed are not dissolved merely to reaggregate in the fraction tube. Because gel filtration permits rather free choice of pH, ionic strength, and additives, it is generally not difficult to find suitable conditions. Solute-matrix interactions Solute-matrix interactions are generally to be avoided. Even though mixed-mode separations (i.e., separations on the basis of both molecular size and the solute-matrix interaction) have sometimes given good resolution, one must be concerned about the reproducibility of such mechanisms. Solute-matrix interactions may be specific or nonspecific, and the chemical basis may be an ionic or hydrophobic interaction. In the case of specific solute-matrix interactions, the only thing to do is to change the matrix to one based on a different chemistry. Ionic interaction can be suppressed by addition of an electrolyte; hydrophobic interaction can
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be reduced by adding an organic modifier (e.g., methanol, acetonitrile, or isopropanol) or by reducing the ionic strength of the buffer. Of course, avoidance of solute-matrix interactions is a prerequisite for estimating molecular size using a calibrated column (see Basic Protocol 3 and Support Protocol). Solute-matrix interactions can be uncovered by varying the composition of the mobile phase or temperature and noting the effect. Because ideal gel filtration is driven by entropy, temperature should not affect the elution volume of proteins (unless the temperature change affects protein size or the pore size). Column maintenance Column life is drastically reduced by introduction of air into the column, deposition of material on the bed surface, and use of flow rates that exceed the maximum recommended flow rate. To protect valuable columns from air it is advisable to use an air trap, to degas the buffer, or to use a deaerator. Deposition of material is minimized by filtering both sample and buffer through a 0.22-µm filter prior to application. Exceeding the maximum flow rate or pressure (e.g., by failing to compensate for viscosity effects at low temperatures) will result in the bed becoming compressed. This may be avoided by using a flow or pressure sensor to shut down the pump if the limit is exceeded. To maintain the calibration of a column for molecular size determinations (see Basic Protocol 3 and Support Protocol), the flow through the column should never be turned off completely, but may be reduced (e.g., to one-tenth) and the effluent recycled to the buffer reservoir through a filter. The buffer should be treated with preservatives and must be discarded when the column is put back in service. According to the author’s experience, the lifetime of a column may be expected to exceed several years and calibration may remain valid for up to 1 year if the appropriate precautions are taken. A parameter that reflects the possibility of bed compression is the void fraction—i.e., the void volume divided by the bed volume. This parameter is 0.26 for beds made up of tetrahedral, closely packed spheres and 0.40 for beds made up of randomly packed spheres. Experimentally determined void fractions vary from 0.30 to 0.47 because of the surface and packing properties of the matrix and the particle shape. Void fractions <0.30 may indicate compression of beads. Gel-Filtration Chromatography
Flow rate Flow rate must be carefully controlled, especially if the column is used for characterization of molecular size as in Basic Protocol 3. The actual flow rate of a pump seldom agrees with the nominal flow rate, so it is essential that the flow rate be checked regularly.
Troubleshooting Gel filtration is a comparatively uncomplicated separation technique and troubleshooting is straightforward. Provided that the critical aspects of the technique (see Critical Parameters) are kept under control, gel filtration can be expected to be relatively trouble-free. Problems that may arise most frequently are compression of the bed and consequent decrease in flow rate, air bubbles in the column, tailing or leading peaks, and peaks that elute in the “wrong place.” Bed compression and decreased flow rate Compression of the bed most likely results from the maximum flow rate being exceeded during packing (see Critical Parameters). It may also arise from excessive packing of the column, which creates an inhomogeneously packed bed with a denser layer in the lower part of the column that will eventually collapse even at a moderate flow rate. A column found to have a compressed bed must be repacked. Compression may also result from partial clogging of the inlet filter and the upper part of the bed. To save the column in such a case, the adaptor should be removed, the filter cleaned, contaminated upper matrix removed, the upper part of the bed swirled, and the bed restabilized (see Basic Protocol 1, step 11). If the column still does not perform as expected, it must be repacked. Decrease in the flow rate often accompanies compression of the bed. If the gel contains fines, these may block parts of the outlet net, causing a decrease in flow rate. Reversing the direction of flow may restore normal conditions. If this is not successful, the column should be repacked after exhaustive decantation of fines (see Basic Protocol 1, steps 3a and 2b). Air bubbles in column Small air bubbles in the column may be due to failure to deaerate the buffer or to the introduction of bubbles during sample injection. Moving the column from a cold room directly to a warmer laboratory will also release air bubbles in the column and should be avoided.
8.3.28 Supplement 14
Current Protocols in Protein Science
An attempt may be made to remove the bubbles by pumping deaerated buffer through the column in the reverse direction. Heating the buffer slightly is sometimes successful, as the temperatures of the buffer and gel will equilibrate, allowing the warmer buffer to dissolve some air. This may have to be done at least overnight and perhaps for several days to be fully successful. A large air pocket situated right under the adaptor that has not been taken up by the buffer may be removed using the procedures for readjusting the adaptor to the bed surface (see Basic Protocol 1, step 11). Tailing and leading peaks Tailing peaks indicate a large dead volume somewhere in the system, solute-matrix interactions, or a bed that is packed too loosely. To check the system for extra dead volumes, another column or a dummy column of small volume should be packed and a sample run to see if the problem reproduces itself. If it does not, the problem can be attributed to the column, and a sample of a different substance should then be run to ensure that the effect is not caused by solute-matrix interactions. If the tailing problem is still noted, the defect probably lies in the column packing, and an attempt should be made to stabilize the bed at a higher flow rate (see Basic Protocol 1, step 11). If this does not help, the column should be repacked at a higher flow rate. Before doing this, however, it should first be ascertained that the tailing is really seen with the sample to be run under the actual conditions under which it is to be run. Slight tailing of a small solute may be negligible when compared to a large diffusionregulated zone broadening of a protein. Tailing may also be caused by partial separation of a small amount of a solute of slightly smaller size or slightly different surface properties. Leading (fronting) peaks indicate solutematrix interactions or cracks in the bed. Solutematrix interactions (e.g., ionic-exclusion effects) are detected by running a different substance and/or using a different buffer. If the problem is related to the column packing (i.e., cracks in the bed), the column must be repacked at a lower flow rate. As with tailing peaks, however, the actual sample should first be run under the actual conditions to be used to determine whether the effect of fronting on the separation is serious. Fronting may also be caused by partial separation of a small amount of slightly larger contaminant.
Peaks that elute in the wrong place Peaks that elute prior to the void volume indicate ionic-exclusion effects. To suppress these effects, the ionic strength of the GF buffer should be increased. Solutes that elute after the total liquid volume indicate adsorptive effects. These may be masked by adding an organic modifier or by increasing or decreasing the ionic strength of the buffer (see Critical Parameters for discussion of solute-matrix effects).
Anticipated Results Results from gel filtration are in general very predictable. Guidelines for optimization and scaleup are presented in Hagel et al. (1989) and Hagel and Janson (1992).
Time Considerations Gel filtration using media of small particle size may generally be carried out in half an hour. Gel filtration using traditional larger-particle matrices are typically run for 5 hr. Desalting is very fast: e.g., 1.4 ml of sample may be desalted in 30 sec. Preparation (i.e., swelling) of the gel will take 3 hr to overnight, depending on the procedure chosen. Packing of the column takes 1 to 4 hr, and the subsequent stabilization of the packed bed takes 1 hr. Thus, a column is typically prepared in a day, and testing of the packing quality is carried out overnight.
Literature Cited ASTM (American Society for Testing and Materials). 1978. Annual Book of ASTM Standards. ASTM D 3016-78. American Society for Testing and Materials, Philadelphia. Andrews, P. 1970. Estimation of molecular size and molecular weights of biological compounds by gel filtration. In Methods of Biochemical Analysis, Vol. 18 (D. Glick, ed.) pp. 1-53. Wiley-Liss, New York. Casassa, E.F. 1967. Equilibrium distribution of flexible polymer chains between a macroscopic solution phase and small voids. J. Polym. Sci. Polymer Letters Part B 5:773-778. de Haen, C. 1987. Molecular weight standards for calibration of gel filtration and sodium dodecyl sulfate–polyacrylamide gel electrophoresis: Ferritin and apoferritin. Anal. Biochem. 166:235245. Dubin, P.L., Kaplan, J.I., Tian, B.-S., and Metha, M. 1990. Size-exclusion chromatography dimensions for rod-like macromolecules. J. Chromatogr. 515:37-42. Flodin, P. 1961. Methodological aspects of gel filtration, with special reference to desalting operations. J. Chromatogr. 5:103-115.
Conventional Chromatographic Separations
8.3.29 Current Protocols in Protein Science
Supplement 14
Granath, K.A. and Flodin, P. 1961. Fractionation of dextran by the gel filtration method. Makromol. Chem. 48:160-171. Hagel, L. 1985. Effect of sample volume on peak width in high-performance gel filtration chromatography. J. Chromatogr. 324:422-427. Hagel, L. 1989. Gel filtration chromatography. In Protein Purification: Principles, High Resolution Methods, and Applications (J.-C. Janson and L. Rydén, eds.) pp. 63-106. VCH Publishers, New York. Hagel, L. 1992. Peak capacity of columns for sizeexclusion chromatography. J. Chromatogr. 591:47-54. Hagel, L. 1993. Size-exclusion chromatography in an analytical perspective. J. Chromatogr. 648:19-25. Hagel, L. and Andersson, T. 1991. A simple procedure for calibrating a column in solute radius. Presented at the 11th International Symposium on HPLC of Proteins, Peptides and Polynucleotides, October 20-23, Washington D.C. Hagel, L. and Janson, J.-C. 1992. Size-exclusion chromatography. In Chromatography, 5th ed., part A: fundamentals and techniques (E. Heftmann, ed.) pp. A267-A307. Elsevier/North-Holland, Amsterdam. Hagel, L., Lundström, H., Andersson, T., and Lindblom, H. 1989. Properties, in theory and practice, of novel gel filtration media for standard liquid chromatography. J. Chromatogr. 476:329-344. Lathe, G.H. and Ruthven, C.R.J. 1956. The separation of substances and estimation of their relative molecular size by the use of columns of starch in water. Biochem. J. 62:665-674. Lindqvist, B. and Storgårds, T. 1955. Molecularsieving properties of starch. Nature 175:511512.
Moore, J.C. 1964. Gel permeation chromatography. I. A new method for molecular weight distribution of high polymers. J. Polym. Sci. Part A 2:835-843. Pharmacia Biotech. 1991. Gel Filtration Principles and Methods (5th ed.), Lund, Sweden. Potschka, M. 1987. Universal calibration of gel permeation chromatography and determination of molecular shape in solution. Anal. Biochem. 162:47-64. Porath, J. and Flodin, P. 1959. Gel filtration: A method for desalting and group separation. Nature 183:1657-1659.
Key References Yau, W.W., Kirkland, J.J., and Bly, D.D. 1979. Modern Size-Exclusion Liquid Chromatography, Practice of Gel Permeation and Gel Filtration Chromatography. John Wiley & Sons, New York. Comprehensive overview of theory and use of gel filtration, covering the entire spectrum of applications (of which protein separation is only a small part). Describes in detail analytical determination of molecular size and size distribution of aqueous and nonaqueous polymers. Hagel, L. 1998. Gel filtration, In Protein Purification: Principles, High Resolution Methods, and Applications, 2nd ed. (J.-C. Janson and L. Ryden, eds.) pp. 19-144. John Wiley & Sons, New York. Describes practical implications of gel filtration theory, with special reference to proteins; a good complement to Yau et al. (1979), and particularly for protein chemists.
Contributed by Lars Hagel Amersham Pharmacia Biotech AB Uppsala, Sweden
Gel-Filtration Chromatography
8.3.30 Supplement 14
Current Protocols in Protein Science
Hydrophobic-Interaction Chromatography
UNIT 8.4
There are many separation techniques available to the modern chromatographer, all of which exploit either the physicochemical or biological properties of the target molecule. Chromatographic techniques are generally divided into two broad categories—nonadsorptive and adsorptive—based on whether or not the target molecule interacts with the chromatographic matrix. The major nonadsorptive method is gel-filtration chromatography (UNIT 8.3), which separates molecules on the basis of size. Gel filtration is essentially a sieving technique; the molecules being fractionated do not adsorb to or interact with the matrix. All other chromatographic methods—ion-exchange chromatography (UNIT 8.2); affinity methods (Chapter 9), including metal-chelate affinity chromatography (MCAC); reversed- and normal-phase chromatography; and hydrophobic-interaction chromatography (HIC; the subject of this unit)—are adsorptive techniques. The basis for separation in adsorptive techniques is the existence of varying degrees of interaction between the chromatographic matrix and the molecules to be resolved. Hydrophobic-interaction chromatography is used to separate proteins on the basis of the differing strengths of hydrophobic interaction between the proteins and a matrix containing immobilized hydrophobic groups. All proteins are to an extent hydrophobic. Most proteins contain some quantity of the hydrophobic amino acids (i.e., those with nonpolar R groups—alanine, phenylalanine, valine, tryptophan, leucine, isoleucine, and methionine). These amino acids are generally considered to be buried within the protein molecule as a result of the aqueous environment. However, because of folding constraints, some nonpolar amino acids are found on the surface of the molecule as distinct hydrophobic patches. These patches of hydrophobicity are most likely important in the maintenance of protein structure, and thus in determining the biological function of the molecule. The number of such hydrophobic patches and their degree of hydrophobicity is characteristic of a particular protein. The variation in relative hydrophobicity from one protein species to another makes it possible to selectively adsorb and elute proteins to and from a suitable hydrophobic matrix. Application of HIC involves loading a protein mixture onto a matrix containing hydrophobic ligands in a high-salt environment, then eluting the proteins by decreasing the salt in the environment (e.g., using a descending salt gradient) or by some other method of adjusting the polarity of the water phase (e.g., by adding nonionic detergents or organic solvents). HIC is often used in the early stages of a separation scheme, especially after a protein precipitation step carried out in the presence of salt, because the binding conditions used in HIC involve high salt concentrations. Elution conditions for HIC involve low salt concentrations; therefore the eluate from an HIC step is ready, in most cases, for application to an ion-exchange column, where binding occurs at low salt concentrations. For this reason, HIC and IEC are often used one after the other in purification strategies. Because binding of the target molecule to an HIC matrix is facilitated by high salt concentration, rather than by organic solvents as in reversed-phase chromatography, HIC is a mild technique that does not destroy the biological activity of the target molecule. HIC, IEC, and gel filtration are useful techniques to consider when developing a purification strategy for labile molecules whose biological activity would be affected by other separation methods. The Strategic Planning section discusses the important parameters in designing and optimizing a separation by HIC, including preparing the sample and choosing a matrix, column, and buffer. Protocols are provided for packing and testing a column (see Basic Contributed by Robert M. Kennedy Current Protocols in Protein Science (1995) 8.4.1-8.4.21 Copyright © 2000 by John Wiley & Sons, Inc.
Conventional Chromatographic Separations
8.4.1 CPPS
Protocol 1 and Basic Protocol 2 and Alternate Protocol); determining binding and elution conditions (see Support Protocol 1); eluting the sample (see Basic Protocol 3); and cleaning, regenerating, and storing HIC columns (see Support Protocol 2). STRATEGIC PLANNING The parameters to consider when selecting a matrix for hydrophobic-interaction chromatography (HIC) and optimizing a separation based upon hydrophobic interactions are: (1) ligand type and degree of substitution of the ligand in the matrix, (2) composition of the matrix support, (3) type and concentration of salt used in the buffer, and (4) pH, temperature, and additives used for the buffer. Table 8.4.1 lists some commercially available media for HIC. The most appropriate matrix will be one that allows the greatest amount of contaminant material to pass through the column during sample loading and initial rinsing. If scale-up is planned for the separation, the cost of items such as buffer additives and waste disposal should be considered for optimum process economy; for economic reasons the most appropriate matrix in such cases will also be one that allows use of the lowest concentration of salt in the binding buffer. Table 8.4.1 Representative Matrices for Hydrophobic-Interaction Chromatography
Matrix
Matrix support composition
Suppliera
45 to 165–ìm bead diameter (for capture and intermediate purification) Butyl Sepharose 4 Fast Flow Agarose PB Phenyl Sepharose 6 Fast Flow Agarose PB Octyl Sepharose 4 Fast Flow Agarose PB Methyl Macro-Prep Methacrylate BR Methacrylate BR t-Butyl Macro-Prep Bio-Beads SM2, 4, 16, 7 Polystyrene BR divinylbenzene Toyopearl butyl Methacrylate TH Toyopearl phenyl Methacrylate TH 30 to 50–ìm bead diameter (for intermediate and polishing) Phenyl Sepharose High Performance Agarose Bio-Beads SM PS/DVB Toyopearl ether Methacrylate Toyopearl butyl Methacrylate Toyopearl phenyl Methacrylate TSK ether 5-PW Silica TSK phenyl 5-PW Silica
PB BR TH TH TH TH TH
10 to 30–ìm bead diameter (for polishing and analytical applications) Phenyl Superose Agarose PB Alkyl (neopentyl) Superose Agarose PB Toyopearl ether Methacrylate TH Toyopearl butyl Methacrylate TH Toyopearl phenyl Methacrylate TH TSK ether 5-PW Silica TH TSK phenyl 5-PW Silica TH HydrophobicInteraction Chromatography
aBR, Bio-Rad; PB, Pharmacia Biotech; TH, TosoHaas. For addresses and phone numbers of suppliers, see SUPPLIERS APPENDIX. bSee UNIT 8.1 for definitions of the terms capture, intermediate purification, and polishing.
8.4.2 Current Protocols in Protein Science
If the matrix does not seem to work at all at first, it should be tested again at a later stage in the purification scheme. It is valuable to perform a number of empirical screenings and pilot experiments (see Support Protocol 1) at each step in the purification before establishing a complete multistep separation scheme. Each of these experiments should test several different matrices. See UNIT 8.1 for discussion of strategies in developing an overall separation scheme. Ligand Type and Degree of Substitution The type of immobilized ligand determines the selectivity of the HIC matrix. Generally, two types of ligand are used in HIC—alkyl-chain ligands of various lengths and the aryl (usually phenyl) ligand. Alkyl-chain ligands show true hydrophobic character. At constant ligand-substitution levels, the protein-binding capacities of HIC matrices increase with increasing alkyl-chain length—i.e., with increasing hydrophobicity. Matrices containing the aryl ligand demonstrate a type of mixed aromatic and hydrophobic interaction, because the phenyl ring has the potential for π-π interactions. The protein-binding capacity of the matrix will increase with increasing degrees of substitution of the immobilized ligand. However, at very high levels of ligand substitution the apparent binding capacity of a matrix will reach a plateau, although the strength of binding will continue to increase in proportion to the degree of substitution. Thus, proteins bound to highly substituted matrices will be difficult to elute. It is worth noting that binding of the surface of a protein to an immobilized HIC ligand is a multipoint attachment, although the attachment between a peptide and an immobilized ligand is a single-point interaction. Composition of Base Matrix The support (base matrix) upon which the ligand is immobilized contributes to the selectivity of the HIC medium. The two most common base matrices in use for HIC are 4% and 6% cross-linked agarose, which are strongly hydrophilic carbohydrates, and the synthetic copolymer materials. The selectivity of an agarose-based matrix and a polymerbased matrix substituted with the same ligand will be different. It is possible to achieve the same results using either type of support, but only if the the adsorption and elution conditions are modified appropriately. It is important to realize when evaluating matrices for HIC that a number of coupling chemistries can be used to attach the alkyl or aryl ligand to the base matrix. For HIC, a matrix should be employed that is prepared with a coupling chemistry in which no charge is imparted to the point of attachment between the base matrix and the hydrophobic ligand. Charge-neutral matrices are often prepared using ether coupling between the base matrix and the hydrophobic ligand. Ligands attached via cyanogen bromide chemistry will be charged, and should be avoided for true HIC. A third consideration relative to the base matrix is the particle size. The smaller the particle, the higher the resolution achieved in the separation. Matrices with particle sizes ≤34 µm are considered high-resolution media. UNIT 8.3 contains good discussion and references on the relationship between particle size and resolution. The practical drawback to using high-resolution media for all applications is that the pressure drop across a column bed is inversely proportional to the square of the matrix particle size. Thus, if larger columns (e.g., with bed heights >30 cm) are packed using a 34-µm matrix and the same pump, the back-pressure will increase, the flow rate will decrease, and the time required to achieve the separation will increase. Another complication is the contribution to pressure drop from the viscosity of the sample feed stream and wash buffers used in HIC, which often have a high salt content and are therefore more viscous than water at
Conventional Chromatographic Separations
8.4.3 Current Protocols in Protein Science
room temperature. Use of a gel matrix with particle size ≥90 µm allows use of reasonably high flow rates at lower back-pressures (e.g., 400 cm/hr at 1 bar). If two HIC steps are used in a separation scheme, it is practical to use a 90-µm matrix early in the purification, and a 34-µm matrix towards the end. Choosing a Column, Pump, Detector, and Other System Components Like most adsorptive techniques, HIC is carried out in short columns. A typical HIC column is packed to a bed height between 5 and 15 cm, but the bed height may be as long as 30 cm. Scale-up of an optimized separation is done by increasing column diameter while keeping the bed height constant. Wide columns (with an internal diameter between 1.6 and 5.0 cm) are appropriate for HIC. To achieve the appropriate bed height, it is helpful if the column is equipped with one or two flow adaptors. A pump should be chosen that is capable of delivering the appropriate flow rate required for HIC, typically between 100 and 600 cm/hr. Because of the high-salt buffers used, peristaltic or syringe pumps are recommended, rather than any type of pump with a ball-check valve. If a pump with a check valve must be used, care must be taken to ensure that no salt crystallizes in the valve seat. A detector capable of absorbance measurements at 280 nm is used to detect the protein peak in HIC. A chart recorder and fraction collector are recommended. Type and Concentration of Salt in Buffer The interaction between the immobilized ligand and the protein is facilitated by the addition of one of the various salting-out salts (i.e., “structure-forming” salts that increase precipitation of hydrophobic compounds; see Fig. 8.4.1) to the equilibration buffer and sample. As the concentration of salt is increased for a particular sample, the amount of protein bound to the immobilized ligand also increases, almost linearly, up to a specific salt concentration. The proteins are then eluted from the column by decreasing the salt concentration, in either a step or continuous descending gradient (see Basic Protocol 3). The effect of salts in hydrophobic interaction can be accounted for by reference to the well-known Hofmeister (lyophilic-lyophobic) series of anions and cations. Figure 8.4.1 reproduces the Hofmeister series for some anions and cations in terms of the degree to which they promote or inhibit protein precipitation. The most commonly used salts for HIC buffers are Na2SO4, NaCl, and (NH4)2SO4, all of which have a high salting-out (structure-forming) effect as seen in Figure 8.4.1. Salts can also be listed in order of their effect on the surface tension of water. For more detailed discussion of the Hofmeister
increasing structure-forming (salting-out) effect –,
anions
PO43
cations
NH4+, Rb+, K+, Na+, Cs+, Li +, Mg 2 + , Ca2+, Ba2 +
–
–
–
–
–
–
–
SO42 , CH3 COO , Cl , Br , NO3 , ClO4 , I , SCN
–
increasing chaotropic (salting-in) effect
HydrophobicInteraction Chromatography
Figure 8.4.1 The Hofmeister series illustrating the effects of some anions and cations in promoting protein precipitation. Ions with a higher salting-out effect promote binding to hydrophobic interaction chromatography matrices, whereas ions with a higher salting-in effect promote elution from those matrices.
8.4.4 Current Protocols in Protein Science
series, see Melander and Horvath (1977); for more detail on surface-tension salts, see Srinivasan and Ruckenstein (1980), Franks (1988), and Hodder et al. (1989). Effect of Buffer pH In published accounts of hydrophobic-interaction chromatography, pH is shown to be an important separation parameter. Generally, as the pH is increased, hydrophobic interaction is decreased. It is possible that increased titration of the charged groups that occurs with increased pH results in a protein that is more hydrophilic. A decrease in pH tends to increase hydrophobic interaction. Thus, proteins that do not bind to a hydrophobic-interaction matrix at neutral pH do bind at acidic pH. It is advisable to check the binding of the protein of interest at several pH values in the course of optimizing a HIC separation. Effect of Temperature On the basis of the hypothesis that interaction between hydrophobic moieties on the protein surface and the immobilized hydrophobic ligand is entropy-driven (see Background Information), one would expect that the interaction should increase with increasing temperature. Experimental evidence to this effect has been presented (Hjerten, 1977), but the role of temperature in HIC is evidently complex, because the opposite results have also been observed (Visser and Strating, 1975). The practical importance of this is that one must be aware that a process developed at room temperature may not be reproducible in the cold room (and vice versa). Elution of Proteins Elution of compounds bound to an immobilized hydrophobic ligand can be achieved in one of three ways: (1) by a linear or stepwise decrease in the concentration of salt, (2) by addition of some proportion of organic solvent to the elution buffer, or (3) by addition of neutral detergents to the elution buffer. The most common way to desorb the proteins from the HIC support is to reduce the salt concentration of the buffer (see discussion above under Type and Concentration of Salt in Buffer), or to use water alone as an eluant. An alternative elution scheme for desorption of the bound protein may exploit the fact that low concentrations of water-miscible alcohols, detergents, and salting-in salts (i.e., chaotropic salts that decrease precipitation of hydrophobic compounds; see Fig. 8.4.1) result in the weakening of the protein-ligand interaction. The nonpolar regions of the alcohols or detergents compete with the bound protein for the hydrophobic ligand, resulting in displacement of the protein. Salting-in salts disrupt the structure of the water, decrease the surface tension, and thereby weaken the hydrophobic interaction. These compounds can be used in the elution scheme to affect selectivity during desorption, but there is a risk that the protein will be denatured or inactivated after exposure to them. Detergents can be effective for cleaning HIC media that have very strongly hydrophobic proteins bound to them (see Support Protocol 2). Preparation of High-Salt Buffers As discussed above under Type and Concentration of Salt in Buffer, the binding of a protein to an HIC medium is facilitated by high salt concentrations. The sample should thus be in high salt prior to application to the HIC column. This can be done by direct dilution in high-salt solution or by buffer exchange through gel filtration (see UNIT 8.3). Before deciding to use gel filtration, it is necessary to ensure that none of the sample components precipitate in the high-salt buffer. This can be done in a small test tube by adding an aliquot of the high-salt buffer to the sample and checking for flocculation.
Conventional Chromatographic Separations
8.4.5 Current Protocols in Protein Science
As the use of ammonium sulfate in buffers for HIC is very common, preparation of such buffers requires some comment. Because ammonium sulfate can be contaminated with heavy metals, iron in particular, the best grade available (usually called enzyme grade) should be used. For the same reason it may be necessary to include micromolar amounts of EDTA in HIC buffers containing ammonium sulfate. Preparation of a saturated solution takes several hours; therefore it is best to prepare the solution a day before chromatography is to be attempted. Concentrations of ammonium sulfate are expressed in terms of molarity or percent saturation. For use in a saturation table, the sample can be considered to be 0% saturated or 0 M salt. Scopes (1994) presents two equations for calculating ammonium sulfate concentration. Also useful are the salting-out tables found in general biochemistry references including Segel (1976), which is aimed at students, and Englard (1990). Before buffer preparation, it is important to take into account that addition of ammonium sulfate to water causes an increase in volume; e.g., 761 g of ammonium sulfate is added to one liter of water to make a saturated solution, resulting in a final volume of ∼1.5 liters. If the water is heated to between 70°C and 80°C, the ammonium sulfate will go into solution faster (in 30 min to 1 hr). If the solution is heated, it should be cooled before use—e.g., overnight in a refrigerator. In a saturated solution there will be crystals of ammonium sulfate in the bottom of the flask. The pH of a saturated ammonium sulfate solution is ∼5; it can be adjusted with concentrated ammonium hydroxide. The aliquot of solution to be used should first be filtered through filter paper. Saturated ammonium sulfate solution may be stored several weeks at 4°C. NOTE: All glassware used in hydrophobic-interaction chromatography should be clean. Spills of highly concentrated salt solutions should be cleaned up immediately, especially if the salt splashes onto instruments. BASIC PROTOCOL 1
PACKING A COLUMN WITH AN HIC MATRIX OF ≥90-ìm BEADS This protocol describes preparation and degassing of the HIC gel medium and packing of the column. After these procedures are completed, a packed column will be ready for testing (Basic Protocol 2) and sample application. This protocol and that for high-resolution media (see Alternate Protocol) are generally applicable to all bulk carbohydratebased media, and can be skipped if a prepacked column is to be used. If non-carbohydrate-based media are to be used, consult the manufacturer’s literature for packing instructions. Materials Appropriate HIC matrix with bead size ≥90 µm (see Strategic Planning and Support Protocol 1) Packing solution: distilled H2O or low-ionic-strength buffer (e.g., 20 mM sodium phosphate, pH 7.0) 20% (v/v) ethanol Chromatography column with flow adaptors (see Strategic Planning) Sintered-glass funnel (medium or coarse grade) Carpenter’s level Pump (see Strategic Planning) 1. Equilibrate all materials to the temperature at which chromatography will be performed.
HydrophobicInteraction Chromatography
2a. To wash the HIC matrix by decantation: Allow gel to settle, decant off the preservative solution, and add an excess of packing solution. Repeat wash several times.
8.4.6 Current Protocols in Protein Science
2b. To wash the HIC matrix by filtration: Pour the gel into a sintered-glass funnel attached to a suction flask and vacuum line and allow the preservative solution to filter off. Release the suction, resuspend the gel in an excess of packing solution, and restore the suction to filter off the packing solution. Repeat wash several times. Gels are shipped in a preservative solution—usually 20% ethanol or a dilute thimerosal or sodium azide solution. Washing by filtration is a faster method than washing by settling and decantation. A sintered-glass filter must be used, not filter paper in a Buchner funnel.
3. Prepare a slurry consisting of 70% washed, settled gel in packing solution. The packing solution (water or dilute buffer) provides a low-viscosity liquid phase for packing the gel in the column.
4. Degas the gel slurry 3 to 5 min under vacuum with swirling. A water aspirator pulls enough vacuum for this degassing.
5. Mount column vertically. Flush the end pieces of the column with 20% ethanol, then flush with packing solution to replace the ethanol. Close outlet, leaving a few centimeters of packing solution in the bottom of the column. 6. Pour the gel slurry into the column with one motion, avoiding introduction of air bubbles. Pouring the slurry down a glass rod held up against the side of the column wall will minimize the introduction of air into the slurry. In the absence of a glass rod, pouring the gel down the column wall will have the same effect.
7. Immediately fill the remainder of the column with packing solution, mount the top adaptor, and connect the column to the pump. Use a carpenter’s level to check that the column is vertical. It is not necessary to push the adaptor down into the gel at this point, but it is convenient to move it into the column ∼1 cm above the settled gel to minimize the need for adjustment after the matrix is packed.
8. Fill the buffer reservoir with packing solution. Open the bottom of the column and set the pump to run at the desired flow rate. Continue passing the packing solution through the column until a constant bed height is reached, then pass an additional 3 bed volumes of solution through at the same flow rate. Different flow rates are recommended by manufacturers for different gel matrices—e.g., Sepharose 6 Fast Flow matrices are packed at a flow rate of 500 cm/hr, and Sepharose 4 Fast Flow gels at a flow rate of 400 cm/hr. Ideally, these two media should also be packed at constant pressure: 1.5 bar for Sepharose 6 Fast Flow and 1.0 bar for Sepharose 4 Fast Flow. However, constant-pressure packing requires a pressure gauge in the chromatography system and this is not always available. If the recommended flow rate cannot be achieved, the gel should be packed at the maximum flow rate that the pump can deliver. IMPORTANT NOTE: Do not exceed 70% to 75% of the packing flow rate in subsequent chromatographic procedures, because if the separation is run at a flow rate approaching that used for packing, there is a risk that the column will continue to pack during sample application and elution, leading to distorted peaks and nonreproducible elution patterns.
9. Mark the level of the packed bed on the column tube. Close the outlet of the column, then stop the pump. Disconnect the inlet tubing from the pump. Adjust the adaptor to the top of the gel medium and lower it 3 mm below the mark on the column tube. Tighten the adaptor and reconnect the inlet tubing to the pump. The column is now ready to use in a separation. For examples of protein purifications done by HIC, see Background Information. Conventional Chromatographic Separations
8.4.7 Current Protocols in Protein Science
ALTERNATE PROTOCOL
PACKING A COLUMN WITH AN HIC MATRIX OF ≤34-ìm BEADS This protocol describes the variations in packing procedure for high-resolution HIC matrices. Carry out Basic Protocol 1 with the following additional materials and alternate procedures at the indicated steps. Additional Materials (also see Basic Protocol 1) Appropriate HIC matrix with bead size ≤34 µm (see Strategic Planning and Support Protocol 1) Tween 20 Chromatography column rated at 5 bar, with flow adaptors (see Strategic Planning) Pressure gauge 3a. Add 250 µl (one drop) Tween 20 per 500 ml of gel slurry. Tween 20 will decrease the surface tension within the gel solution, making the slurry more homogeneous and easier to swirl and degas.
7a. Pack the column at a flow rate of 30 cm/hr until the bed reaches a constant height (or at least 20 min). Shut off the pump, then lower the adaptor to ∼1 cm above the settled gel bed. Turn on the pump and increase the flow rate until a pressure of 5.0 bar is reached. Continue packing at this pressure for 30 to 60 min. After the high-resolution gel is packed, some resistance may be encountered from the packed bed when attempting to push the adaptor down 3 mm into the bed (see Basic Protocol 1, step 9). It is nevertheless important to do this before using the column in a separation. BASIC PROTOCOL 2
HydrophobicInteraction Chromatography
TESTING THE PACKED BED Testing the packed column is easy, and is recommended for two reasons. First, it is better to find out whether or not the column is properly packed before the sample is applied. It is not unusual for someone who has never packed a column before to try two or three times before succeeding. Second, testing column performance with a tracer before the sample is applied and the column used and regenerated provides a baseline for comparison after a number of runs. Often, the theoretical plate number and/or asymmetry factor, determined as described below, can be used to predict column failure. Thus, it is recommended that the packed column be tested before it is used and then again before each additional use (or before every other use, or on some other regular schedule). To test a packed column, a low-molecular-weight test substance that has no interaction with the matrix is injected into the column. The number of theoretical plates (N) or the height equivalent to a theoretical plate (H, sometimes called HETP) is then calculated from the elution volume of the test substance. The asymmetry factor (As) is also calculated. The small molecular weight assures that the test substance has access to the entire chromatographic bed. 1% (v/v) acetone can be used as a test substance. The acetone sample is applied just as though it were any other chromatographic sample. The sample volume used for this test should be small: 2% of the column volume for 90-µm packings or 0.5% of the column volume for 34-µm packings. However, for evaluating column packing it is appropriate to minimize the volume between the injection point and the detector; thus, short lengths of tubing should be used in the chromatography system for the evaluation step even if longer lengths will be used for the chromatography of an unknown sample later on. The flow rates used for evaluation of the column should be kept low to minimize the zone broadening (UNIT 8.1) at the front and rear of the zones. For columns packed with 90-µm matrices, the flow rate should be 15 to 30 cm/hr; for columns packed with 34-µm matrices, the flow rate should be 30 to 60 cm/hr.
8.4.8 Current Protocols in Protein Science
Because acetone absorbs at 280 nm, it is not necessary to change the wavelength setting on the detector to test the packed bed. However, it will be necessary to change the sensitivity of the detector to keep the peak on-scale so that the peak apex can be seen. The variables N, H, and As should not be calculated from a peak that has gone off-scale. The following standard chromatographic equations are used to calculate the number of theoretical plates (N; also see discussion of column efficiency in UNIT 8.1) and then to calculate plate height (H): N = 5.54(Ve/Wh)2 and finally H =L/N where Ve is the volume eluted from the start of sample application to the peak maximum, Wh is the peak width measured at half peak height (as traced by the recorder), and L is the length of the packed bed. Because the plate count (N) is dimensionless, the measurement of Ve and Wh can be taken in distance on the chart paper (mm) or volume of eluant (ml). The units used should be the most convenient for the investigator, but both Ve and Wh should be expressed in the same units for a given calculation. Figure 8.4.2 shows a chromatogram with Ve and Wh marked. As a general rule, a good value for H is about two to three times the mean bead diameter of the gel being packed. Thus, a good H value for a 90-µm matrix is between 0.018 and 0.027 cm and a good H value for the 34-µm matrix is between 0.0070 and 0.010 cm.
A280
Ve
Wh
h
1/2 h a
b 10% h
Elution volume
Figure 8.4.2 Chromatogram illustrating parameters used in calculating plate number and asymmetry factor for a column. Ve is the elution volume for the protein whose peak is depicted, h is the height of that peak, Wh is the width of the protein peak at half the peak height, a is the width of the leading portion of the peak at 10% of the peak height, and b is the width of the tailing portion of the peak at 10% of the peak height.
Conventional Chromatographic Separations
8.4.9 Current Protocols in Protein Science
Another standard chromatographic data analysis procedure for testing a packed bed is calculation of the asymmetry factor As. On the same peak used to measure H on the chart recording, a line is drawn from the peak maximum to the baseline, dividing the peak in two parts lengthwise. Asymmetry is calculated simply by the equation: As = b/a where a is the width of the front (leading) portion of the peak measured at 10% peak height and b is the width of the back (tailing) portion of the peak measured at 10% peak height. The chromatogram in Figure 8.4.2 depicts the quantities a and b graphically. As should be as close as possible to 1, but a reasonable value for a short-bed HIC column is between 0.80 and 1.80. An extensive leading edge on a peak (As < 0.80) is usually an indication the column is packed too tightly, and an extensive trailing edge (As > 1.80) is an indication the column is packed too loosely. These problems can be solved by repacking the column. SUPPORT PROTOCOL 1
PILOT EXPERIMENT TO CHOOSE A MATRIX AND DETERMINE BINDING AND ELUTION CONDITIONS FOR HIC This protocol describes an experiment to identify the appropriate HIC matrix and initial salt concention to bind and elute a target molecule. This experiment is done with very small columns and consumes very little sample. A small quantity of the sample solution dissolved in a high-salt start buffer is applied to several different columns, each packed with a different HIC matrix. Elution is then carried out using a either a descending continuous gradient or a descending step gradient of salt concentration (see Basic Protocol 3 for more discussion of the two gradient types). The results will include a wash-through fraction (i.e, material that did not bind to the column at all) and one or more fractions that will elute during the chromatography. Each fraction is then analyzed by some sort of assay to identify the protein of interest (see Chapter 3), and the most appropriate matrix—i.e., the one that allows the greatest amount of contaminant material to pass through the column during sample loading and initial washing—is thus identified. For additional considerations in choosing a matrix, see Strategic Planning. Materials HIC matrices to be tested (e.g., a low-substitution phenyl, a higher-substitution phenyl, a butyl, and an octyl matrix; Table 8.4.1) General start buffer: 50 mM Na2HPO4/1.0 M (NH4)2SO4, pH 7.0 (prepare ∼1 liter) 5 to 10 mg/ml solution of protein mixture to be purified, in general start buffer 50 mM Na2HPO4containing 0.75 M, 0.50 M, 0.25 M and 0 M (NH4)2SO4 (for step gradient only; all solutions at pH 7.0; prepare 1 liter each) 1-ml chromatography columns Chromatography system with syringes or pump, detector, and test tubes or fraction collector (see Strategic Planning) Gradient mixer (UNIT 8.2) set to deliver descending gradient from 1.0 M to 0 M (NH4)2SO4 in 10 column volumes of Na2HPO4, pH 7.0 (for continuous gradient only; if gradient mixer is not built into chromatography system) 1. Pack each HIC matrix to be tested into a 1-ml column as in Basic Protocol 1 or the Alternate Protocol. 2. Equilibrate each column with at least 10 column volumes of general start buffer.
HydrophobicInteraction Chromatography
The flow rate for equilibration can be fast—e.g., 1 column volume/min. See Critical Parameters for guidelines on column equilibration.
8.4.10 Current Protocols in Protein Science
3. Apply 1 ml of protein solution to be purified in start buffer to each column. Perform initial washes in start buffer, and monitor the eluant using a detection scheme appropriate to the protein of interest. A slower flow rate is recommended for sample application and elution than for equilibration—e.g., 0.25 column volumes/min.
4. Elute the sample using either a gradient mixer set to deliver a descending gradient from 1.0 M to 0 M (NH4)2SO4 in 10 column volumes of Na2HPO4, pH 7.0 or a step gradient of 10 column volumes each of 0.75 M, 0.50 M, 0.25 M, and 0 M (NH4)2SO4 in 50 mM Na2HPO4, pH 7.0. Continue to monitor the eluant and construct a chromatogram. See Critical Parameters and Troubleshooting for guidelines on interpreting chromatograms obtained from this pilot experiment and for optimizing chromatography conditions. The most appropriate matrix is the one that allows the greatest amount of contaminant material to pass through the column during sample application and initial washing. If the protein of interest elutes during sample application or initial washing with start buffer, repeat the procedure with the same gel matrix but increase the salt concentration of the start buffer. Alternatively, more hydrophobic matrices may be investigated. The purification of the protein should be evaluated even where elution occurs in the initial wash, because a considerable amount of contaminating material may be bound to the HIC media and hence removed. If the protein of interest does not elute until a 0 M salt concentration has been reached, the salt concentration of the start buffer may be decreased from 1.0 M (NH4)2SO4 (e.g., to 0.25 M). Elution may then be done using 50 mM Na2HPO4, pH 7.0, or water alone. If the activity of the protein of interest is adversely affected by low salt concentrations, a less hydrophobic matrix that allows elution at a higher salt concentration may be investigated.
ELUTION OF PROTEINS FROM HIC COLUMNS
BASIC PROTOCOL 3
Continuous-Gradient Elution In continous gradients, salt concentration is increased or decreased gradually and continously using some sort of gradient-mixing device (see UNIT 8.2). Simple linear gradients (see UNIT 8.2 for definitions of gradient types) are the first choice in pilot experiments for determining initial conditions (see Support Protocol 1). After more experience is obtained with a particular separation, the gradient can be made more complex to optimize resolution between specific sample components. This is illustrated in Figure 8.4.3, where the linear gradient in panel A shows very poor resolution of the first two components in the mixture (i.e., peaks 1 and 2). By making the first portion of the gradient slope less steep, peaks 1 and 2 can be resolved, as shown in panel B. In areas of the chromatogram where resolution must be improved, the gradient can be made less steep, whereas in areas where resolution is adequate or not critical, the gradient can remain more steep. The general effect of gradient slope is illustrated in Figure 8.4.4. Increasing the total gradient volume—i.e., decreasing the gradient slope—will improve resolution in all parts of the chromatogram. The drawback to this is that it will increase the cycle time and dilute the separated fractions.
Conventional Chromatographic Separations
8.4.11 Current Protocols in Protein Science
A
B
1 2 A280
1
2
Elution volume
Elution volume
Figure 8.4.3 Chromatograms illustrating use of complex gradient elution to improve resolution. (A) A simple linear gradient gives poor resolution of protein peaks 1 and 2. (B) A complex gradient where the gradient slope is less steep in the region of peaks 1 and 2 gives better resolution of these two components.
A
B
1
A280
2
1 3
4
2 3
Elution volume
4
Elution volume
Figure 8.4.4 Chromatograms illustrating the general effect of a change in gradient slope on peak resolution. (A) A steep slope gives poor resolution among the four sample components. (B) A less steep slope gives better resolution (though the total elution volume is higher, meaning that the eluted sample components will be more dilute and that more buffer is needed for the elution).
Step-Gradient Elution
HydrophobicInteraction Chromatography
In step-gradient elution, one or more solutions of a particular discrete concentration are passed through a column. The principle of stepwise elution is to provide maximum resolution in the area of the gradient where the protein of interest elutes. Stepwise elution is preferred for large-scale preparative applications because it is technically simpler and more reproducible than continous-gradient elution. It is also advantageous in small-scale applications because the compound of interest can be eluted in a more concentrated form, provided there is no coelution of a more strongly bound compound. When stepwise elution is used, one must watch out for artifact peaks that can appear if the next step is started too
8.4.12 Current Protocols in Protein Science
A
B step 1
concentration for step 1
A280
concentration for step 2
step 2 step 3
concentration for step 3
Elution volume
Elution volume
Figure 8.4.5 Chromatograms illustrating use of the results of linear-gradient elution to devise a step-gradient elution scheme. The shaded area represents the target protein peak. (A) Elution by linear gradient yields poor resolution among sample components. However, it is possible to note the salt concentration at the points on the gradient where the peaks for the loosely bound contaminants, target protein, and tightly bound contaminants emerge. (B) Step-gradient elution with three discrete solutions prepared on the basis of the elution salt concentrations noted in panel A. The target protein peak and the two contaminant peaks are sharply resolved.
early—i.e., during a trailing peak. It is recommended that a stepwise elution scheme be developed after several continuous gradient schemes have been run to characterize sample behavior on the matrix. This development should include three stages. In the first stage a continuous gradient is optimized to elute all the compounds that bind to the gel less tightly than the protein of interest. The elution strength and gradient volume should be high enough to elute all weakly binding contaminants but not high enough to elute the protein of interest. In the second stage the elution strength is increased so that the protein of interest elutes with minimal dilution, but more tightly bound contaminants are not eluting. In the third step the elution strength is increased to elute the most tightly bound contaminants. Once these three regions of the chromatogram are identified, these gradient positions are converted to steps (i.e., solutions representing the concentration of eluant at these points in the chromatogram are prepared and used for step elution). Figure 8.4.5 illustrates this process of switching from a continuous to a step gradient. REGENERATION, CLEANING, AND STORAGE OF HIC COLUMNS
SUPPORT PROTOCOL 2
Regeneration of HIC Columns After each use, the matrix must be regenerated to restore its original functionality. There are two general regeneration procedures for HIC columns. The first is to simply wash the column with several column volumes of distilled water (Milli-Q purified or equivalent). The second is to wash the columns with at least two column volumes of 6 M urea or 6 M guanidine hydrochloride and then at least 10 column volumes of start buffer (see Support Protocol 1) or storage solution (see Storage of HIC columns, below). The first procedure is sufficient if the buildup of tightly bound protein contaminants is minimal; the second procedure is recommended for more tightly bound contaminants. Deciding which to use
Conventional Chromatographic Separations
8.4.13 Current Protocols in Protein Science
is situation-dependent, but a quick screen using each, noting if a peak of material emerges, can indicate the appropriate choice. If detergents were used in the elution scheme, the regeneration protocol is more complicated. In this case the column should be washed sequentially with one column volume of distilled water; one column volume each of 25%, 50%, and 95% ethanol; two column volumes of n-butanol; one column volume each of 95%, 50%, and 25% ethanol, and finally one column volume of distilled water. This should be followed by reequilibration with the start buffer (see Support Protocol 1) or the storage buffer (see Storage of HIC columns, below). Cleaning of HIC Columns Cleaning of HIC columns involves removal of very tightly bound precipitated or denatured proteins, as well as lipid substances, from the gel matrix. If not removed, these contaminants may remain on the matrix after the regeneration procedures, causing a decline in chromatographic performance. Because of buildup of these fouling materials on the gel, matrix selectivity will change, and in some cases the functioning of the column will also change—e.g., flow rates can decrease and back-pressure can increase. It is preferable to clean the matrix while it is still in the column, a practice referred to as “cleaning in place,” so that the column does not have to be unpacked, repacked, and tested again. There is no single recommended protocol for cleaning in place; the procedure must be designed with some knowledge of the contaminants present in the feed stream. In general, however, NaOH is a very effective cleaning agent that can be used for solubilizing both denatured and precipitated protein and lipids. Removal of precipitated proteins To remove precipitated proteins, a general recommendation is to wash the column with four column volumes of 0.5 to 1.0 N NaOH at a slow flow rate (e.g., 40 cm/hr), then with two to three column volumes of water. After the water wash, the column can be equilibrated with the start buffer (see Support Protocol 1) or put into the storage buffer (see Storage of HIC Columns, below). Removal of tightly bound proteins and lipids There are two general recommendations for the removal of strongly bound hydrophobic proteins, lipids, and lipoproteins. The first is to wash the column with four to ten column volumes of ethanol, starting with 30% ethanol and working up to 70%, then with six to ten column volumes of distilled water (Milli-Q purified or equivalent; this should reduce the alcohol content of the eluate to between 10 and 1 ppm). The 30% to 70% ethanol may be replaced with 30% isopropanol. The second procedure is to wash the gel matrix with one to two column volumes of a 0.5% solution of nonionic detergent (e.g., Triton X-100) in 1 M acetic acid, followed by five column volumes of 70% ethanol to remove the detergent. The ethanol is then washed out with six to ten column volumes of distilled water. After the water wash, the column can be equilibrated with the start buffer or put into the storage buffer. Sanitization of HIC Columns
HydrophobicInteraction Chromatography
Sanitization is the inactivation of microbial populations in the gel matrix. NaOH is very effective for sanitizing the matrix in addition to functioning as a general cleaning reagent. A general sanitizing procedure for HIC columns is to expose the matrix to 0.5 to 1.0 N NaOH for 30 to 60 min. The column should then be washed with at least ten column volumes of start buffer (see Support Protocol 1) if it is to be used right away. The large volume of wash solution is required because a large pH adjustment is being made. If the
8.4.14 Current Protocols in Protein Science
column is going to be stored it should be washed with storage buffer (see Storage of HIC Columns, below) until the pH of the effluent equals the original pH of the storage buffer. Sodium hydroxide is widely accepted as an agent for sanitizing chromatographic media, columns, and systems. For this purpose it is used at higher concentrations (e.g., 0.5 N to 1.0 N). At lower concentrations (e.g., 0.01 N) it is an effective bacteriostatic agent, and for the most common gram-negative contamination a good bactericidal effect is also seen at these concentrations. NaOH also destroys endotoxin on the column and is nontoxic as a contaminant in the end product. Sanitization is not synonymous with sterilization, which refers to the destructive elimination of a microbial population. The manufacturer’s recommendations for autoclaving the matrix should be followed if sterilization is required. Storage of HIC Columns Unused matrix should be stored in closed containers at a temperature between 4° and 25°C. The matrix should not be allowed to freeze as this will disrupt the bead structure and generate fines. Used matrix should be stored in the presence of a suitable bacteriostatic agent—e.g., 0.01 N NaOH or 20% ethanol—at a temperature between 4° and 8°C. Used matrix should also be protected against freezing. Packed columns should be stored under the same conditions. For long-term storage, the matrix should be cleaned (see Cleaning of HIC Columns) before equilibration with the storage buffer. Recycling the storage solution or flushing the column with fresh storage solution at least once a week during long-term storage is an effective method for preventing bacterial growth. Chromatography matrices from Pharmacia Biotech are supplied in 20% ethanol, and this solution can be used as an alternative to NaOH for storing columns. Other manufacturers ship media in other bacteriostatic solutions, which can also be used as alternative storage buffers. Other bacteriostatic compounds can be added to the storage buffer, including 0.002% chlorhexidine (which can, however, cause precipitation of sulfate ions if mixed with a high-molarity salt such as that used in start buffers for HIC); 0.5% trichlorobutanol (effective in weakly acidic solutions only), 0.005% thimerosal (effective in weakly acidic solutions); 0.001% to 0.01% phenyl mercuric acetate, borate, or nitrate (effective in weakly alkaline solutions); and 0.02% to 0.05% sodium azide. COMMENTARY Background Information Tiselius (1948) laid down the foundations of what today is called hydrophobic-interaction chromatography (HIC). Since then, a number of theories have been proposed to explain this type of chromatography, but no single one has gained universal acceptance. All of the explanations refer to the interactions of the structureforming salts (see Strategic Planning, Type and Concentration of Salt in Buffer) and the effects that they exert on other solutes and solvents in a system. Porath (1986) posed a general explanation for HIC based upon salt-promoted adsorption. In an extension of earlier work in
which the observations of Tiselius were extended and the behavior of hydrophobic molecules in solution were linked to thermodynamic theory (Porath, 1973), they concluded that the driving force of HIC is the gain in entropy arising from the structural changes in the water surrounding the interacting hydrophobic groups. Hjerten (1977) expressed this idea in terms of the thermodynamic relationship ∆G = ∆H − T∆S. The displacement of water molecules surrounding the hydrophobic ligands on the matrix and the hydrophobic amino acids on the surface of the protein leads to an increase in entropy (∆S), because more water is returned
Conventional Chromatographic Separations
8.4.15 Current Protocols in Protein Science
to the bulk solution after a hydrophobic interaction occurs. This leads to a negative value for the ∆G of the system, which implies that hydrophobic interaction is thermodynamically favorable. This also implies that the entropy of water is the driving mechanism for hydrophobic interaction; thus, HIC is unique among the adsorptive separations in that it occurs as a result of its environment. A second explanation for hydrophobic interaction was proposed by von der Haar (1976) and by Melander and Horvath (1977). According to these authors, hydrophobic interaction results from the increase in surface tension of water arising from dissolved structure-forming salts. A third explanation for hydrophobic interactions was proposed by Srinivasan and Ruckenstein (1980). This hypothesis proposes that hydrophobic interaction is a result of the increase in van der Waals attraction between the protein and the immobilized ligand that results from the increase in the ordered structure of water in the presence of structure-forming salts.
HydrophobicInteraction Chromatography
Elution by salt gradient An example of elution by salt gradient was published by Lau et al. (1987) as part of the purification of acid phosphatase from bovine bones by HIC. The active fractions from previous steps were pooled in 100 mM sodium acetate buffer, pH 6.5, and adjusted to 30% saturation by adding solid ammonium sulfate at 4°C with gentle stirring for ≥2 hr. This enzyme solution was applied to a phenyl HIC column equilibrated with 30% ammonium sulfate in 100 mM sodium acetate buffer, pH 6.5. The chromatography was done at room temperature. The column was eluted with a descending salt gradient starting at 30% ammonium sulfate/100 mM sodium acetate, pH 6.5, and ending with 0% ammonium sulfate/100 mM sodium acetate, pH 6.5. The yield was 92% and the purification was 13-fold. This was the last step in a four-step purification protocol and the purity of the isolated enzyme was determined by SDS-PAGE. HIC with gradient elution offers an alternative in monoclonal antibody purification, provided that the monoclonal antibody is stable in high salt. As most monoclonal antibody purifications start with an ammonium sulfate precipitation step, the antibody will normally be present either in a high salt–soluble fraction or as a precipitate that can be redissolved and applied to a HIC column. Monoclonal antibody samples containing albumin and transferrin are par-
ticularly suitable for HIC because the albumin and transferrin can be eluted as sharp peaks, minimizing contamination of the antibody. This was demonstrated by Sparrman et al. (1986), who used an alkyl Superose matrix. A clarified sample was equilibrated with 0.7 M ammonium sulfate by buffer exchange using gel-filtration chromatography. The column was then equilibrated with several bed volumes of the start buffer, 1.6 M ammonium sulfate. A linear gradient was produced by mixing buffer A (0.1 M phosphate, pH 7.0/2 M ammonium sulfate) and buffer B (0.1 M phosphate alone) starting at 20% buffer B and ending with 80% buffer B. The transferrin eluted at ∼40% buffer B, and the albumin eluted at ∼45% buffer B. The monoclonal IgG peak eluted at ∼55% buffer B and was therefore sharply separated from the contaminating proteins. Elution by detergents Addition of detergents to the HIC column is one method for increasing the resolution of elution. However, some detergents can bind very strongly to HIC matrices and are difficult to wash out with common solvents such as ethanol. Detergents should be used with care. The mechanism of detergent action in HIC is that the nonpolar end of the detergent molecule competes with the hydrophobic patches on the protein for the hydrophobic group on the matrix. An example of detergent elution was published by Hereld et al. (1986), describing the purification of phospholipase C from the protozoan Trypanosoma brucei. The enzyme was present in an ammonium sulfate cut prepared from the solubilized membranes of the protozoan. A phenyl column was equilibrated with 25 mM sodium succinate buffer (pH 6.0) containing ammonium sulfate at 50% saturation. The sample (already in ammonium sulfate) was applied to the column and eluted in a descending salt gradient starting with the 50% saturated ammonium sulfate in buffer A and ending with 0% salt (i.e., buffer only). The phospholipase was then eluted by application of buffer containing 1% CHAPS. Most of the protein eluted in a sharp peak immediately after application of the detergent, but there was some trailing in this elution. The yield was 62%, with a 22-fold purification. Elution by polarity-reducing compounds The addition of various polarity-reducing compounds to the buffer is another way to increase the resolving power of the elution
8.4.16 Current Protocols in Protein Science
buffer. This can be done only if the protein of interest is stable upon exposure to the compound used. The reagents most commonly used for this purpose are 40% ethylene glycol or 30% isopropanol, dissolved in salt-free buffer. These are used to decrease the polarity of the eluent, resulting in a decrease in binding strength and thus elution of the bound protein from the column. An example of this technique, using ethylene glycol to elute human pituitary prolactin, was published by Roos et al. (1979). The work was done in the cold, and started with whole organs that were homogenized and extracted at pH 9.8. Between 200 and 400 mg protein was applied to a phenyl column equilibrated with a buffer consisting of 0.2 M glycine adjusted to pH 9.8 with NaOH. Elution was carried out by stepwise reduction of the buffer concentration, first to 0.02 M glycine, pH 9.8, then to 0.002 M glycine, pH 9.8, with 50% (v/v) ethylene glycol added. The last buffer eluted the prolactin. Flow rate for this elution was ∼5 cm/hr and the recovery was 95%. HIC and protein polarity It has been estimated by Klotz (1970) and Lee and Richards (1971) that 40% to 50% of the surface of a protein is nonpolar. These nonpolar areas are the areas on the protein responsible for binding the protein to hydrophobic matrices in the presence of moderate to high concentrations of salting-out salts. Various hydrophobicity scales for the amino acids have been published; Ericksson (1989) presents several of these in tabular form. There is also a relationship between this salt-promoted interaction with a HIC ligand and the precipitation data for proteins (Melander and Horvath, 1977). HIC and other chromatographic techniques HIC is complementary to the other adsorptive techniques, as well as gel filtration, in that it separates on the basis of a very different physicochemical property of the protein than these other methods and can therefore be used in conjunction with them to achieve more complete separation (see UNIT 8.1). The physicochemical property exploited by HIC is the presence or absence of the various hydrophobic amino acids in the protein, as distinguished from the property exploited by ion-exchange chromatography, which is the presence or absence of the various charged amino acids. HIC also distinguishes proteins according to the hydrophobic amino acids on the surface of the protein, not those buried inside the folded pro-
tein. This distinguishes HIC from reversedphase chromatography, which also separates on the basis of hydrophobicity, but on the basis of the hydrophobicity of the denatured or unwound sequence. If there is reason to believe it should be possible to exploit the hydrophobicity of the surface of the protein of interest, or to exploit the hydrophobicity of the surface of the contaminants in the sample, then HIC should be included in a separation plan.
Critical Parameters and Troubleshooting Because the fundamental operating principle of HIC is that hydrophobic interactions are facilitated by high-salt environments, an incomplete equilibration of either the sample or the matrix with the high-salt buffer will have a negative effect on results and reproducibility. Sample equilibration The sample must be completely dissolved in the start buffer. This can be done either by buffer exchange through gel-filtration chromatography (UNIT 8.3), dialysis (APPENDIX 3B), or membrane filtration. Sample solubility should be checked in each salt environment to which the sample will be exposed to during chromatography, to make sure that there will be no precipitation when changing from a high-salt to a no-salt environment. Column equilibration The column must be equilibrated in the high-salt environment at the correct pH for sample application. Many chromatographers try short-cuts during column equilibration only to find that they must repeat the run. It is common that at least 10 column volumes of buffer must pass through a column to equilibrate it. The column should be equilibrated at the temperature at which the chromatography will be run. Thus, a column run overnight in the cold room is not equilibrated for a run the next morning at room temperature. Optimizing resolution Panels A to F in Figure 8.4.6 show some typical results that may be obtained on different HIC matrices using the methodology of Support Protocol 1. The shaded section of each chromatogram indicates the protein of interest. Target protein eluted early in the gradient; resolution not satisfactory (Fig. 8.4.6A). Not much can be gained in this situation by changing the salt concentration of the equili-
Conventional Chromatographic Separations
8.4.17 Current Protocols in Protein Science
B
C
D
E
F
A280
A280
A280
A
Elution volume
Elution volume
Figure 8.4.6 Chromatograms illustrating various optimal and suboptimal outcomes in HIC. (A) Target protein eluted early, resolution not satisfactory. (B) Target protein eluted late, resolution not satisfactory. (C) Target protein eluted in middle of gradient, resolution not satisfactory. (D) Target protein eluted early, resolution satisfactory. (E) Target protein eluted late, resolution satisfactory. (F) Target protein eluted in middle of gradient, resolution satisfactory (optimal results). These results and techniques for improving upon them are discussed in Critical Parameters and Troubleshooting.
HydrophobicInteraction Chromatography
bration buffer. Decreasing the salt concentration will decrease the binding capacity of the protein of interest and might even lead to its elution together with the unbound contaminant fraction. Increasing the salt concentration might lead to adsorption of unwanted impurities and a conseqent decrease in the selectivity of the matrix for the protein of interest. Changing the pH of the equilibration buffer might result in a stronger binding and higher
selectivity for the protein of interest. The effect of pH is variable for different proteins; usually lowering the pH leads to increased binding. Increasing the pH usually leads to a decreased binding of proteins, which in this particular case might result in the elution of the protein of interest together with the unbound fraction. The appropriate next step in this case would be to try the experiment at a lower and higher
8.4.18 Current Protocols in Protein Science
pH. If no improvement in selectivity is obtained, a matrix with a different ligand or (if available) the same ligand at a higher degree of ligand substitution should be tried. Target protein eluted near the end of the gradient; resolution not satisfactory (Fig. 8.4.6B). A decrease in buffer salt concentration will weaken the strength of binding, resulting in earlier elution of the protein of interest. A decrease in salt concentration may also have a positive effect on selectivity because a greater quantity of the less hydrophobic substances will be eluted with the unbound fraction. However, the effect of this approach on the resolution will be marginal, because the contaminants elute very close to the protein of interest—both before and after the target protein peak. Changing the pH of the equilibration buffer may have a positive effect on resolution and should be tried. The appropriate next step in this separation would be to try the experiment at a lower and higher pH. If no improvement in resolution is obtained, a gel matrix with a different ligand or (if available) the same ligand at a lower degree of ligand substitution should be tried. Target protein eluted in the middle of the gradient; resolution not satisfactory (Fig. 8.4.6C). Changing the concentration of the salt in the equilibration buffer will have a limited effect on resolution. However, either an increase or decrease in the pH of the equilibration buffer might have a favorable effect. The appropriate next step in this experiment is to repeat the separation at a higher and lower pH. If no improvement is seen, a gel matrix with a different ligand or (if available) the same ligand at a higher degree of ligand substitution should be tried. Target protein is eluted early in the gradient; resolution is satisfactory (Fig. 8.4.6D). The separation shown in panel D is an example of a good choice of matrix for the protein of interest. However, the fact that the protein of interest is eluted very early in the gradient indicates that the binding capacity of the matrix may be low. This may be compensated for, if necessary, by a moderate increase of salt in the equilibration buffer. This may in turn result in a decrease in the selectivity of the adsorbent because some of the unbound proteins might be adsorbed together with the protein of inter-
est. Another negative effect of increased salt concentration may be a decrease in resolution caused by an increase in gradient slope if the total gradient volume or the cycle time is kept constant. Increased salt concentration will also result in increased costs, which may be important in a manufacturing process. Not much can be gained here by changing the pH of the equilibration buffer, as the resolution obtained is satisfactory. There is no need to try to improve on this separation unless the low binding capacity is a problem. If that is the case and the problems with increased salt concentration mentioned above are encountered, a matrix with a different ligand or (if available) the same ligand at a higher degree of ligand substitution should be tried. Target protein of interest is eluted near the end of the gradient; resolution is satisfactory (Fig. 8.4.6E). This also illustrates a good choice of matrix for the protein of interest. Decreasing the concentration of salt in the equilibration buffer will give earlier elution of the protein of interest, reduced cycle time, and decreased cost for salt. A problem in this situation might be that some of the most hydrophobic-contaminating substances are bound so strongly that an organic solvent or chaotropic agent has to be used to clean the column (see Support Protocol 2). Not much can be gained here by changing the pH, because the selectivity is good. If the very strong binding of hydrophobic contaminants poses a problem, a medium with a different ligand or (if available) the same ligand at a lower degree of ligand substitution should be tried. Target protein elutes in the middle of the gradient; resolution is satisfactory (Fig. 8.4.6F). The choice of ligand here is good and there is little risk of strong binding of the most hydrophobic contaminants. This panel illustrates an example of good results in hydrophobic-interaction chromatography. The examples presented in Figure 8.4.6 do not consider two extreme cases that may arise—i.e., that in which the protein of interest is not bound to the column at all, and that in which the protein of interest binds so strongly that it is difficult to elute without the use of denaturing solvents. In both instances a different HIC medium or another chromatographic separation technique should be tried.
Conventional Chromatographic Separations
8.4.19 Current Protocols in Protein Science
Although it is assumed for the sake of discussion that the resolution is not satisfactory in some of the chromatograms shown in Figure 8.4.6, the requirements for resolution in any chromatographic step must be stipulated on a case-by-case basis. What seems to be fairly poor resolution at an advanced stage of a separation can often be good enough at an initial capture step where the main objective of the separation is reduction of volume, removal of some critical contaminants, and preparation for further chromatography using a high-resolution matrix. For more guidelines on troubleshooting chromatographic separations, the Troubleshooting section in UNIT 8.2 may be consulted.
Anticipated Results Hydrophobic-interaction chromatography (HIC) is intended for use as one of several chromatographic steps in a protein purification scheme. The product that elutes from a HIC column may not be chemically pure, as HIC will copurify proteins of similar hydrophobicity. Yields will be high (step yields of 60% to 90% are common), and because the conditions of HIC are nondenaturing, activity recovery will be high as well.
Time Considerations
Hodder, A.N., Aquilar, M.I., and Hearn, M.T.W. 1989. High performance liquid chromatography of amino acids, peptides and proteins LXXXIX. The influence of different displacer salts on the retention properties of proteins separated by gradient anion exchange chromatography. J. Chromatogr. 476:391-411. Klotz, I.M. 1970. Comparison of molecular structures of proteins: Helix content, distribution of apolar residues. Arch. Biochem. Biophys. 138:704-706. Lau, K.H., Freeman, T.K., and Baylink, D.J. 1987. Purification and characterization of an acid phosphatase that displays phosphotyrosyl-protein phosphatase activity from bovine cortical bone matrix. J. Biol. Chem. 262:1389-1397. Lee, B. and Richards, F.M. 1971. The interpretation of protein structures: Estimation of static accessibility. J. Mol. Biol. 55:397-400. Melander, W. and Horvath, C. 1977. Salt effects on hydrophobic interactions in precipitation and chromatography of proteins: An interpretation of the lyotropic series. Arch. Biochem. Biophys. 183:200-215. Porath, J., Sundberg, L., Fornstedt, N., and Olsson, I. 1973. Salting-out in amphiphilic gels as a new approach to hydrophobic adsorption. Nature 245:465-466.
Chromatography using prepacked HIC columns can be run in ∼30 min to 4 hr. It will take ∼8 hr to pack and equilibrate a column, then load and elute a sample. Cleaning, regeneration, and reequilibration can be carried out overnight. Samples can often be stored overnight in the high-salt start buffers, but the possibility of precipitation should be ruled out first.
Porath, J. 1986. Salt promoted adsorption: Recent developments. J. Chromatogr. 376:331-341.
Literature Cited
Sparrman, M. 1986. Purification of monoclonal antibodies by hydrophobic interaction chromatography. Presented at the 6th International Symposium on HPLC of Proteins, Peptides, and Polynucleotides, Baden-Baden, Germany.
Englard, S. and Seifter, S. 1990. Precipitation techniques. Methods Enzymol. 182:285-300. Ericksson, K.-O. 1989. Hydrophobic interaction chromatography. In Protein Purification: Principles, High Resolution Methods and Applications. (J.-C. Janson and L. Rydén, eds.) pp. 207226. VCH Publishers, New York. Franks, F. 1988. Characterization of Proteins. In Characterization of Proteins (F. Franks, ed.) p. 53. Humana Press, Clifton, N.J. Hereld, D., Krakow, J.L., Bangs, J.D., Hart, G.W., and Englund, P.T. 1986. A phospholipase C from Trypanosoma brucei which selectively cleaves the glycolipid on the variant surface glycoprotein. J. Biol. Chem. 261:13813-13819. HydrophobicInteraction Chromatography
erence to serum proteins. In Proceedings of the International Workshop on Technology for Protein Separation and Improvement of Blood Plasma Fractionation. pp. 410-421. Reston, Virginia.
Hjerten, S. 1977. Fractionation of proteins by hydrophobic interaction chromatography, with ref-
Roos, P., Nyberg, F., and Wide, L. 1979. Isolation of human pituitary prolactin. Biochim. Biophys. Acta. 588:368-379. Scopes, R.K. 1994. Protein Purification: Principles and Practice, 3rd ed. Springer-Verlag, New York. Segel, I.H. 1976. Biochemical Calculations. John Wiley & Sons, New York.
Srinivasan, R. and Ruckenstein, E. 1980. Role of physical forces in hydrophobic interactions chromatography. Sep. Purif. Methods 9:267370. Tiselius, A. 1948. Adsorption separation by salting out. Arkiv för Kemi, Mineralogi, Geologi 26B:1-5. Visser, J. and Strating, M. 1975. Separation of lipoamide dehydrogenase isoenzymes by affinity chromatography. Biochim. Biophys. Acta. 384:69-80. von der Haar, F. 1976. Fractionation of proteins by fractional interfacial salting out on unsubstituted agarose gels. Biochem. Biophys. Res. Commun. 70:1009-1013.
8.4.20 Current Protocols in Protein Science
Key Reference Ericksson, 1989. See above. Describes both the theory and practice of hydrophobic-interaction chromatography and includes a number of references to the theoretical as well as the applications literature.
Contributed by Robert M. Kennedy Pharmacia Biotech Piscataway, New Jersey
Conventional Chromatographic Separations
8.4.21 Current Protocols in Protein Science
Chromatofocusing
UNIT 8.5
Chromatofocusing (CF) is a versatile, high-resolution chromatographic technique suitable for separating proteins on the basis of their isoelectric points. It may be used to purify proteins with isoelectric points in the range of 3 to 11. Proteins elute from a CF column in descending order of isoelectric point. The unique aspect of this technique is the formation of a descending linear pH gradient in situ by elution with a single buffer containing a large number of buffering species (i.e., the CF elution buffer described here, which contains polybuffer)—the same buffering system used in isoelectric focusing (UNITS 10.2 & 10.4). This methodology is in contrast to the mixing of a high- and low-pH buffer prior to the column—the technique used to form pH gradients for elution in ion-exchange chromatography (see UNIT 8.2). The Strategic Planning section discusses choice of chromatofocusing medium and buffers, determination of the quantity of medium needed, and preparation of CF buffers. The Basic Protocol provides the procedure for a typical chromatofocusing experiment, Support Protocol 1 describes regeneration of the chromatofocusing column, and Support Protocol 2 presents several methods for separation of the polybuffer from the purified protein. A schematic representation of the steps in a CF experiment is provided in Figure 8.5.1. STRATEGIC PLANNING Selecting CF Medium and CF Buffers The first step in selecting a medium, start buffer, and elution buffer for chromatofocusing (CF) is to determine the appropriate pH gradient range on the basis of the isoelectric point of the target protein. After the pH gradient range has been selected, Tables 8.5.1 and 8.5.2 may be used to select the appropriate CF gel medium and buffers. Finally, after the CF buffers and CF gel medium have been selected, Table 8.5.1 may be used to determine the amount of buffer required and Table 8.5.2 may be used to determine the amount of gel required for the application. The CF media and buffers listed in Table 8.5.1 are designed to produce a gradient over a maximum interval of 3 pH units or a minimum of 1 pH unit. For initial experiments a pH gradient spanning 3 pH units should be used. Once the chromatographic behavior of the target protein is characterized, narrower gradients can be used to optimize resolution. If the isoelectric point of the protein is known, the pH range of the CF experiment is chosen so that the target protein will elute after one-third to one-half of the pH gradient. For example, if the isoelectric point of the target is 6.2, a pH range of 4 to 7 or 5 to 8 can be chosen for initial experiments. If the isoelectric point of the target protein is unknown, it may be determined by isoelectric focusing (UNIT 10.2) or by the test tube method for determining starting conditions for ion-exchange chromatography (UNIT 8.2). For an example of the latter method, see Figure 8.2.3, Panel A, where the isoelectric point of the protein is between 5.5 and 7.0. Alternatively, because most proteins have an isoelectric point in the range of 4 to 7 (Gianazza and Righetti, 1980), it is usually appropriate to simply select a pH gradient in that range for initial experiments. Polybuffer 96 and Polybuffer 74 (Pharmacia Biotech) are amphoteric buffers designed for chromatofocusing in the ranges of pH 6 to pH 9 and pH 4 to pH 7, respectively. They contain a large number of buffering species to provide an even buffering capacity, and
Conventional Chromatographic Separations
Contributed by Alan Williams
8.5.1
Current Protocols in Protein Science (1995) 8.5.1-8.5.10 Copyright © 2000 by John Wiley & Sons, Inc.
Supplement 2
choose appropriate CF medium and buffers according to pl of target protein (Strategic Planning and Table 8.5.1)
equilibrate CF medium with CF start buffer and pack into column (Basic Protocol, steps 1-7)
set up chromatography system (Basic Protocol, step 8)
adjust pH of CF start buffer to upper limit of pH gradient (Strategic Planning, Table 8.5.1, and Table 8.5.2)
purge chromatography system with CF start buffer (Basic Protocol, step 8 )
attach CF column to system and test for equilibration (Basic Protocol, steps 9 -10)
adjust pH of CF elution buffer to lower limit of pH gradient (Strategic Planning)
allow 5-10 ml CF elution buffer to run through CF column (Basic Protocol, step 12)
equilibrate sample with elution buffer if necessary (Basic Protocol, step 11)
apply sample to CF column and begin collecting fractions (Basic Protocol, step 12)
reequilibrate CF column with CF start buffer (Support Protocol 1)
develop pH gradient by elution with CF elution buffer (Basic Protocol, step 13)
regenerate gel (Support Protocol 1)
assay fractions and pool those containing the target protein (Basic Protocol, step 14)
remove polybuffer from protein, if necessary (Support Protocol 2)
Figure 8.5.1 Flow chart for a chromatofocusing experiment. Chromatofocusing
8.5.2 Supplement 2
Current Protocols in Protein Science
are used together with the polybuffer exchange gel PBE 94 or the prepacked Mono P column (both available from Pharmacia Biotech). For chromatofocusing in the range pH 8 to 11, Pharmalyte pH 8 to 10.5 is used in conjunction with the polybuffer exchange gel PBE 118 (both items also available from Pharmacia Biotech). Suitable CF start buffers for different selected pH gradient ranges are indicated in Table 8.5.1. The normal counter-ion for these anion-exchange media is chloride. Monovalent anions other than chloride may be used as counter-ions, but it is critical that these should have a pKa at least 2 pH units below the lowest point of the gradient chosen (Hutchens, 1989). As bicarbonate ions and atmospheric carbon dioxide can cause fluctuations in the pH gradient, all buffers must be degassed before use. Atmospheric carbon dioxide may cause a plateau in the region of pH 5.5 to 6.5, depending on the conditions. These effects are most marked with Polybuffer 96 in pH gradients ending at pH 6, and can be avoided by using acetate as the counter-ion (Pharmacia Biotech, 1985). Alternatively, the lower limit for gradients with Polybuffer 96 may be set at pH 6.5. Acetate is not usually recommended as a counter-ion with Polybuffer 74 because of its higher pKa. Multivalent anions are not recommended as counter-ions. Determination of Appropriate Quantity of CF Medium The amount of gel required for a chromatofocusing run depends on the amount of sample, the nature of the sample and contaminants, and the degree of resolution required. For most separations, a bed volume of 20 to 30 ml is sufficient for separation of samples containing from 1 to 200 mg of protein per pH unit covered by the gradient (e.g., 600 mg of protein for a 3-pH-unit gradient). However, the precise amount of gel used will depend on the resolution required from the experiment, a factor that should be considered during optimization. Preparation of CF Buffers CF start buffer is prepared as in Table 8.5.1. CF elution buffers are prepared by diluting Polybuffer 96 or 74 or Pharmalyte 8 to 10.5 with distilled water according to the dilution factors listed in Table 8.5.1. The pH of the CF elution buffer should then be adjusted to the lower limit of the selected pH gradient. Approximately 15 Vc of CF start buffer and 15 Vc of CF elution buffer per run are typically required. This should be enough for equilibration (system, column, and sample), separation, and regeneration. All buffers should be degassed under vacuum or by sonication prior to use. Buffers to be used with the prepacked column Mono P HR 5/20 should be filtered through a 0.45-µm filter membrane prior to use. SEPARATION OF PROTEINS BY CHROMATOFOCUSING In this procedure, the CF column is equilibrated in CF start buffer at a pH higher than that of the CF elution buffer. As the lower-pH CF elution buffer moves through the column, the ion-exchange groups of the medium in the column are titrated by the CF elution buffer, forming a descending linear pH gradient in the column. In a case where the isoelectric point (pI) of a protein is below the pH at which the column is equilibrated, the protein will have a net negative charge at the beginning of the experiment and therefore will bind to the anion-exchange medium in the CF column. As the descending pH gradient slowly forms in the column, the pH at the position where the protein is bound will eventually drop below the pI of the protein, and the net charge on the protein will become positive. The positively charged protein will then desorb from the anion-exchange medium and be carried through the column in the mobile phase. As the desorbed and now positively charged protein advances through the column in the mobile phase, it will pass through
BASIC PROTOCOL
Conventional Chromatographic Separations
8.5.3 Current Protocols in Protein Science
Supplement 2
Table 8.5.1
Selected pH range
Buffers and Gels for Chromatofocusing in Different pH Rangesa,b
CF gelc
Start buffer
Elution buffer
Dilution factord
Approx. volume of buffer needed (column volumes) Dead vol
10.5-9f
PBE 118
10.5-8
PBE 118
0.025 M Pharmalyte pH 8-10.5 triethylamine⋅HCl, pH 11 (pH 8.0)g
10.5-7
PBE 118
9-8h
—
—
—
Gradient Total vol vole
—
—
—
1:45
1.5
11.5
13
0.025 M Pharmalyte pH 8-10.5 triethylamine⋅HCl, pH 11 (pH 7.0)g
1:45
2
11.5
13.5
PBE 94 or Mono P
0.025 M Pharmalyte pH 8-10.5 ethanolamine⋅HCl, pH 9.4 (pH 8.0)g
1:45
1.5
10.5
12
9-7
PBE 94 or Mono P
0.025 M Polybuffer 96 (pH 7.0)g ethanolamine⋅HCl, pH 9.4
1:10
2
12
14
9-6
PBE 94 or Mono P
0.025 M ethanolamine acetate, pH 9.4
Polybuffer 96 (pH 6.0)i
1:10
1.5
10.5
12
8-7
PBE 94 or Mono P
0.025 M Tris.Cl, pH 8.3
Polybuffer 96 (pH 7.0)g
1:13
1.5
9
10.5
8-6
PBE 94 or Mono P
0.025 M Tris acetate, pH 8.3
Polybuffer 96 (pH 6.0)i
1:13
3
9
12
8-5j
PBE 94 or Mono P
0.025 M Tris acetate, pH 8.3
30%/70% (v/v) Polybuffer 96/ Polybuffer 74 (pH 5.0)i
1:10
2
8.5
10.5
7-6
PBE 94 or Mono P
0.025 M imidazole acetate, pH 7.4
Polybuffer 96 (pH 6.0)i
1:13
3
7
10
7-5
PBE 94 or Mono P
0.025 M imidazole⋅HCl, pH 7.4
Polybuffer 74 (pH 5.0)g
1:8
2.5
11.5
14
7-4
PBE 94 or Mono P
0.025 M imidazole⋅HCl, pH 7.4
Polybuffer 74 (pH 4.0)g
1:8
2.5
11.5
14
6-5
PBE 94 or Mono P
0.025 M histidine⋅HCl, pH 6.2
Polybuffer 74 (pH 5.0)g
1:10
2
8
10
6-4
PBE 94 or Mono P
0.025 M histidine⋅HCl, pH 6.2
Polybuffer 74 (pH 4.0)g
1:8
2
7
9
5-4
PBE 94 or Mono P
0.025 M piperazine⋅HCl, Polybuffer 74 (pH 4.0)g pH 5.5
1:10
3
9
12
aTable reproduced with permission of Pharmacia Biotech. bDegas all buffers before use. cPBE 94, Mono P, Pharmalyte 8-10.5, Polybuffer 96, and Polybuffer 74 are products of Pharmacia Biotech (see SUPPLIERS APPENDIX). dThe dilution factor given is not critical; optimal conditions will be found by experience. eGradient volume will vary with the exact conditions chosen. fGradients ending at pH 9 are not recommended because pH 9 is above the pH of Pharmalyte pH 8-10.5. gpH adjusted with HCl. hPBE 118 and Pharmalyte 8-10.5 as well as PBE 94 and Polybuffer 96 (all from Pharmacia Biotech) also cover this range. ipH adjusted with acetic acid. jMixing of Polybuffers 96 and 74 gives the best results; Polybuffer 74 gives better results than Polybuffer 96 when used alone.
Chromatofocusing
8.5.4 Supplement 2
Current Protocols in Protein Science
Table 8.5.2 pH Ranges, Loading Capacities, and Flow Rates for Chromatofocusing Gel Mediaa
CF gel medium PBE 118 PBE 94 Mono P HR 5/20
pH range 8-11 4-9 4-9
Recommended loading capacity
Recommended flow rate
10 mg protein/ml gel 10 mg protein/ml gel 25 mg protein/column 5 mg protein/peak
50 cm/hr 50 cm/hr 0.5-1 ml/min
aRecommended flow rates and loading capacities are for initial experiments; see UNIT 8.2 for optimiza-
tion procedures. A peristaltic pump is generally adequate for packing and operation of PBE 94 and PBE 118 columns; the Mono P HR 5/20 column is designed for use at higher pressures and should be used with HPLC or FPLC systems.
the gradient, in which the pH in the column is increasing. When the pH in the column finally exceeds the pI of the protein, the net charge of the protein again becomes negative; hence the protein is again adsorbed to the anion-exchange medium. The net effect, as the protein adsorbs and desorbs many times, is a focusing effect which results in zone sharpening, sample concentration, and very high resolution. Materials CF gel medium PBE 94 or PBE 118 or prepacked CF column Mono P (Pharmacia Biotech; see Strategic Planning, Table 8.5.1, and Table 8.5.2) CF start buffer (see Strategic Planning and Table 8.5.1) CF elution buffer (see Strategic Planning and Table 8.5.1) Protein sample to be purified Sintered-glass filter funnel, medium porosity Side-arm flask Chromatographic system including: Appropriate chromatographic column Fluid delivery system with appropriate pump Sample injection valve UV monitor suitable for use at 280 nm pH monitor with flowthrough electrode (optional) Recorder Fraction collector Additional reagents and equipment for checking packed chromatographic bed (UNIT 8.4), desalting via gel-filtration chromatography (UNIT 8.3), and protein assay (Chapter 3) Pack the column 1. Pour the appropriate amount of CF gel medium into a sintered-glass funnel attached to a side-arm flask. Using aspiration, pass start buffer through the gel in the funnel until the pH of the effluent is the same as that of the start buffer. Equilibration usually requires 10 to 15 bed volumes of start buffer. The gel should be stirred gently from time to time to ensure complete equilibration. Steps 1 to 7 are omitted where a prepacked CF column is used. CF columns may also be equilibrated after packing. If a column is used in which the gel medium was not equilibrated before packing, or if a prepacked column is used, the column should be washed with 2 to 3 Vc of 1 M NaCl to remove any bound substances, then with 10 to 15 Vc of CF start buffer until equilibrated. The column has reached equilibrium when the pH and conductivity of
Conventional Chromatographic Separations
8.5.5 Current Protocols in Protein Science
Supplement 2
the eluant match the corresponding values for the CF start buffer. When the column is equilibrated, proceed to step 8.
2. Disperse gel in a small amount of start buffer (∼50% of settled bed volume to be packed) to make a slurry. Degas under vacuum. 3. Mount chromatographic column vertically and flush any air bubbles from underneath the bed support nets. The column outlet should be closed at this point. See Figure 8.3.1 for an illustration of the chromatographic column. Best results are obtained on long, narrow columns. For high resolution in preparative applications, a bed height ≥30 cm is desirable. A column diameter of 2.5 cm at a 40-cm bed height (Vc = 200 ml) is usually sufficient for separating ≤2000 mg of protein.
4. Place 2 to 3 ml start buffer in the column and pour gel slurry down a glass rod into the column, taking care to avoid introducing air bubbles. If the volume of the slurry is greater than that of the column, a packing extension should be used (see Fig. 8.3.1).
5. Open column outlet and allow gel to settle into column. When all of the suspension has run into the column, remove packing extension (if used) and connect top piece (i.e., flow adaptor; see Fig. 8.3.1) to the column, taking care to exclude air bubbles. Connect column to pump. 6. Run start buffer through column at a flow rate of 100 cm/hr until the gel bed has completely settled. Equilibration of a column that has already been packed without previous equilibration can be carried out at a lower flow rate. The column has been equilibrated when the pH and conductivity of the eluant match the corresponding values for the start buffer (usually after 10 to 15 bed volumes of start buffer have passed through the column; see annotation to step 1).
7. Check the quality of the column packing (see UNIT 8.4). A well-packed column is essential to get the full benefits of chromatofocusing—i.e., the achievement of extremely narrow zones (0.02 pH units). Care must be taken in the choice of dye substance for checking beds, as they may be strongly charged. Bovine cytochrome c (pI = 10.5) is recommended as the dye marker for visual inspection of the bed in CF columns. Dilute acetone solutions are appropriate for efficiency tests (see UNIT 8.4).
Prepare the system 8. Set up the chromatography system according to the manufacturer’s instructions without the column in-line, but with the UV monitor in-line. Purge the system of air bubbles by washing with CF start buffer. Check for leakage. 9. Install the packed column in the system and continue to equilibrate with start buffer at the flow rate appropriate for the column (see Table 8.5.2). Do not exceed 75% of the maximum flow rate used in packing the column (i.e., 75 cm/hr for 100 cm/hr packing flow rate; see step 6).
10. Check that the UV monitor signal is stable and that the recorder is operational. Collect one fraction and measure the pH, which should be the same as that of the CF start buffer. If the pH is not the same, continue to equilibrate the system by running start buffer through the column. Prepare sample and apply to column 11. Equilibrate protein sample in CF elution buffer or start buffer by desalting via gel-filtration chromatography (UNIT 8.3). Chromatofocusing
8.5.6 Supplement 2
Current Protocols in Protein Science
Although the best way to ensure that the sample will bind to the column is by buffer exchange into start buffer, this step is only necessary when the ionic strength is high (typically >50 mM). Diluting the sample with distilled water can also be used to decrease the ionic strength. Sample pH is not critical if the buffering capacity of the sample is less than that of the start buffer. The amount of sample to be applied to the CF column depends on the amount of protein present in each zone. As a guideline for initial experiments, 10 mg protein can be applied for every 1 ml CF medium. This quantity will vary according to the number of proteins present in the sample. The volume of sample is unimportant so long as all of the sample is applied before the substance of interest is eluted from the column. The composition of the sample is important: it should not contain large amounts (>0.05 M) of salt.
12. Using a linear flow rate of 50 to 60 cm/hr, run 5 ml CF elution buffer through the equilibrated CF column, then run the sample in CF elution buffer or CF start buffer. Running elution buffer through the column before running the sample insures that sample proteins are never exposed to extremes of pH. NOTE: Do not monitor protein at 254 nm, because CF elution buffers absorb at that wavelength.
13. Still using a linear flow rate of 50 to 60 cm/hr, begin development of the pH gradient by eluting the sample with CF elution buffer. Begin fraction collection, and continue elution with CF elution buffer until the pH of the fractions collected reaches the desired pH (i.e., the pH of the CF elution buffer). When exploring conditions for a new separation, collect 20 fractions over the linear pH gradient. Once the separation has been refined, smaller fractions can be collected to optimize resolution; alternatively the peak collection feature of an automated fraction collector may be employed. There is a 1.5 to 3.0 Vc delay before the pH gradient begins to elute from the column. The exact delay volume depends on the buffering capacities of the CF column and CF elution buffer.
14. Assay the fractions collected (see Chapter 3) and pool those containing the target protein. The Polybuffer 74 or 96 contained in the CF elution buffer does not interfere with most enzyme assays or amino acid analyses (UNIT 3.3). Although CF elution buffer does not interfere with Coomassie blue–based protein assays, it forms a complex with copper ions and interferes with the Lowry protein assay (UNIT 3.4). Where necessary, CF elution buffer may be removed by any of several methods (see Support Protocol 2).
REGENERATION OF THE CF COLUMN Additional Materials (also see Basic Protocol) 1 M NaCl
SUPPORT PROTOCOL 1
1. Wash column with 2 to 3 Vc of 1 M NaCl to remove bound material. Strongly bound material can often be removed with 0.1 M HCl or 0.1 M NaOH, but the gel should be equilibrated to a pH of 4 to 9 as soon as possible after treatment with these acid or alkaline reagents.
2. Equilibrate column with 10 to 15 bed volumes of CF start buffer in preparation for the next chromatographic run. If column is being prepared for storage, equilibrate with 2 or 3 column volumes of 20% ethanol instead of start buffer.
Conventional Chromatographic Separations
8.5.7 Current Protocols in Protein Science
Supplement 2
SUPPORT PROTOCOL 2
REMOVAL OF POLYBUFFER FROM PROTEIN There are several methods for separating polybuffer from proteins where it is necessary to do this in preparation for analysis or further purification, including precipitation, hydrophobic interaction chromatography (HIC), gel filtration, and affinity chromatography. Precipitation The simplest method for separating protein from polybuffer is precipitation with ammonium sulfate (Scopes, 1994). Solid ammonium sulfate is added to the relevant fractions until a suitable concentration (80% to 100% saturated) is obtained. The sample is then allowed to stand 1 to 2 hr until the protein precipitates. This procedure is usually relatively simple, as the pH of the fraction is at or near the isoelectric point of the protein. Alternatively, the fractions of interest can be placed in dialysis tubing and dialyzed against saturated ammonium sulfate. Precipitation has the advantage of concentrating and stabilizing the sample, and is also inexpensive. Hydrophobic Interaction Chromatography (HIC) HIC (UNIT 8.4) can be used in conjunction with ammonium sulfate treatment. Ammonium sulfate is added to the fraction of interest obtained from chromatofocusing to a level of 80% saturation, which will favor hydrophobic interactions. This treatment may result in precipitation of the sample protein. If this happens, the purified sample can be harvested by centrifugation. Gel Filtration CF elution buffer can be removed from most proteins by gel filtration (UNIT 8.3) on a column packed with Superdex 75 prep grade medium (Pharmacia Biotech). If the molecular weight of the protein is <15,000, some difficulties may be encountered in achieving baseline separation in short columns. Affinity Chromatography The protein of interest can be easily be separated from CF elution buffer by taking advantage of the biological activity of the protein. The fractions of interest are passed through a column containing a bioselective adsorbent that binds the desired protein, allowing components of the CF elution buffer to elute unretarded. See Chapter 9 for a variety of affinity chromatography techniques. COMMENTARY Background Information Chromatofocusing was first described by Sluyterman and coworkers (Sluyterman and Elgersma, 1978; Sluyterman and Wijdenes, 1978). They proposed that a pH gradient could be produced on an ion exchanger by taking advantage of the buffering action of the charged group of the ion exchange medium. If a buffer that has been initially adjusted to one pH is run through an ion-exchange column that has been initially adjusted to a second pH, a pH gradient is formed, just as if two buffers at different pH were gradually mixed in the mixing chamber of a gradient maker. If such a pH gradient is used to elute proteins bound to the ion exchanger, the proteins elute in order of their
isoelectric points. Furthermore, focusing effects occur, resulting in band sharpening, sample concentration, and very high resolution. The high resolution attainable with CF makes this technique ideal for the intermediate purification and polishing stages of a purification process (UNIT 8.1). The combination of CF with techniques that separate proteins based on physiochemical attributes other than charge characteristics—i.e., HIC (UNIT 8.4), gel filtration (UNIT 8.3), and affinity chromatography (Chapter 9)—is recommended. If the presence of polybuffer in the sample interferes with the intended use of the target protein, an additional polishing step will be required to remove the polybuffer (see Support Protocol 2).
Chromatofocusing
8.5.8 Supplement 2
Current Protocols in Protein Science
Critical Parameters and Troubleshooting The most critical steps in chromatofocusing are: first, selection of the correct pH gradient range for separation, and then proper matching of the buffering capacity of the CF medium to the buffering capacities of the CF start and elution buffers. Monitoring pH and ionic strength of the column effluent during the run is advised for both the interpretation of results and use in troubleshooting. Conductivity and pH monitors equipped with flow cells may be included downstream from the column; these should be calibrated prior to the experiment. If the conductivity and pH are to be measured in the collected fractions, the measurements should be made as soon as possible after collection to avoid variations from uptake of atmospheric gases. Linear gradients are required for optimal
Table 8.5.3
resolution because nonlinear pH gradients give poor resolution in certain areas of the chromatogram. To produce a linear pH gradient, it is necessary that the CF eluant buffer and CF medium have an even buffering capacity over a wide pH range. A list of potential problems, causes, and solutions for use in troubleshooting are given in Table 8.5.3.
Anticipated Results Proteins elute from a CF column in descending order of isoelectric point. The pH at which a protein elutes from the CF column is typically less than its isoelectric point. Recovery from CF in terms of total protein is usually >80%. It is not unusual to notice fluctuations in the pH gradient during the delay in gradient formation (see Basic Protocol, step 14 annotation) or during the elution of concentrated proteins. A
Troubleshooting Guide for Chromatofocusing
Problem
Possible cause
Solution
Protein does not bind to column
Column is incorrectly equilibrated
Check that pH at bottom of column matches that of start buffer Choose a start buffer with a higher pH Use buffer composition recommended in Table 8.5.1 Remove salt in sample by gel filtration in elution or start buffer (Basic Protocol, step 11) Regenerate correctly and wash thoroughly before reuse (Support Protocol 1) Check pH of eluant
Start pH is too low Ionic strength of eluant is too high Ionic strength of sample is too high Column is contaminated with protein Protein binds to but does not elute from column Gradient does not reach desired pH Gradient fluctuates
Eluant composition is incorrect pH of eluant is too high Eluant composition is incorrect Eluant contains CO2 Incorrect counter-ion was used CO2 is present
Bands are skewed
Bed net is blocked Column was poorly packed
Try a lower pH range Check eluant pH; prepare fresh eluant if necessary Degas all buffers prior to use Use recommended counter-ions (Strategic Planning) Degas all buffers prior to use; in alkaline regions, degas water before making up eluant Remove bed net; clean or replace with a new net Check packing with cytochrome c (UNIT 8.4); repack bed if necessary.
Conventional Chromatographic Separations
8.5.9 Current Protocols in Protein Science
Supplement 2
2.0
9 1.5
pH
A 280
8 1.0
7 0.5
6
25
50
75
100
125
Volume (ml)
Figure 8.5.2 Chromatogram depicting separation by chromatofocusing with PBE 94 of a model mixture of proteins in the range of pH 9 to 7. An SR 10/50 column (Pharmacia Biotech) with a bed height of 18 cm was used. The sample consisted of 5 ml elution buffer containing 2 mg sperm whale myoglobin, 2 mg horse myoglobin, and 2 mg carboxyhemoglobin. The start buffer consisted of 0.025 M ethanolamine, pH 9.5, and the elution buffer consisted of 0.0075 mmol/pH unit/ml Polybuffer 96, pH 7. A flow rate of 15 cm/hr was used. (Courtesy of Pharmacia Biotech.)
typical chromatogram depicting the separation of a mixture of protein standards by CF is presented in Figure 8.5.2.
Time Considerations The time required for CF experiments depends primarily on the length of the column, the flow rate, and the volume of sample to be applied; 3 to 4 hr per run is common, excluding the time required for column packing. Separation times using the Mono P HR 15/20 are <1 hr. As with any chromatographic separation, the most time-consuming aspect is normally the analysis of the fractions collected. Increasing resolution with narrow gradients, achieved by optimizing the CF separation, will help reduce the number of fractions to be collected.
Literature Cited Gianazza, E. and Righetti, P.G. 1980. Size and charge distribution of macromolecules in living systems. J. Chromatogr. 193:1-8.
Chromatofocusing
Hutchens, W.T. 1989. Chromatofocusing. In Protein Purification: Principles, High Resolution Methods, and Applications (J.C. Janson and L. Rydén, eds.) pp. 149-174. VCH Publishers, New York.
Pharmacia Biotech. 1985. FPLC Ion Exchange and Chromatofocusing: Principles and Methods. Pharmacia Biotech AB, Uppsala, Sweden. Scopes, R.K. 1994. Protein Purification, Principles and Practice, 3rd ed., pp. 81-85. Springer-Verlag, New York. Sluyterman, L.A. Æ. and Elgersma, O. 1978. Chromatofocusing: Isoelectric focusing on ion exchange columns. I. General principles. J. Chromatogr. 150:17-30. Sluyterman, L.A. Æ. and Wijdenes, J. 1978. Chromatofocusing: Isoelectric focusing on ion exchange columns. II. Experimental verification. J. Chromatogr. 150:31-44.
Key Reference Pharmacia Biotech. 1984. Chromatofocusing with Polybuffer and PBE. Pharmacia Biotech AB, Uppsala, Sweden. An excellent discussion of CF theory including practical techniques and model applications for the separation of proteins by CF.
Contributed by Alan Williams Pharmacia Biotech Piscataway, New Jersey
8.5.10 Supplement 2
Current Protocols in Protein Science
Hydroxylapatite Chromatography
UNIT 8.6
Hydroxylapatite (also called hydroxyapatite), a form of calcium phosphate, can be used as a matrix for the chromatography of both proteins and nucleic acids. Molecular separation is not completely dependent on any single property of the molecule such as molecular weight, size, or isoelectric point; the complex interactions between the matrix and the sample confer unique separative properties on hydroxylapatite. It may be used early in purification schedules to fractionate crude mixtures or later on to separate components that have not been well resolved by other steps, such as affinity chromatography or gel filtration. Two basic protocols are set out in this unit. The first (see Basic Protocol 1) describes standard low-pressure chromatography of a protein mixture using a hydroxylapatite column prepared in the laboratory; the second (see Basic Protocol 2) describes an HPLC method, applicable to proteins and nucleic acids, that uses a commercially available column. Alternate protocols describe column chromatography using a step gradient (see Alternate Protocol 1) or batch binding and step-gradient elution (see Alternate Protocol 2). STRATEGIC PLANNING Selecting a Matrix Choosing the most suitable form of hydroxylapatite matrix for the application is crucial for success. Hydroxylapatite is available in powder form, as a hydrated gel, in a ceramic form, and prepacked into a variety of columns (Table 8.6.1). Most suppliers provide excellent instruction leaflets and/or technical bulletins, and are pleased to offer advice concerning their products. (Also see Critical Parameters.) Selecting a Column The optimal size of a hydroxylapatite column is determined by the amount of material to be processed. The binding capacity of the matrix is usually supplied by the manufacturer, and is used to calculate the approximate quantity required. The resin bed should not be too long in relation to its width to allow use of a reasonable flow rate and to minimize broadening of the eluted peaks (see UNIT 8.1). A bed height-to-width ratio between 5:1 and 12:1 is suggested. The best type of column to use is one with two adjustable end pieces, so the volume of the bed can be varied as required (see Figure 8.3.1 for illustrations of typical chromatography equipment; also see Critical Parameters). LOW-PRESSURE CHROMATOGRAPHY FOR PROTEIN PURIFICATION Low-pressure chromatography is frequently used for laboratory-scale purifications (i.e., a few milligrams to a few grams). By choosing a suitably sized column, small or large fractionations can be accommodated.
BASIC PROTOCOL 1
The procedure described here uses a closed-column pumped system. The matrix is prepared, the column is packed, the sample is chromatographed, and the eluted proteins are visualized by in-line UV detection. Variations on this procedure might include the use of prepacked columns and the detection of eluted proteins by dye binding (UNIT 3.4), functional assay, or PAGE (Chapter 10). Hydroxylapatite can be obtained commercially or, if desired, can be prepared in the laboratory (Atkinson et al., 1973). If the technical literature supplied with the product suggests steps additional to or different from those described in this protocol, always follow the manufacturer’s recommendations. Contributed by Anne V. Broadhurst Current Protocols in Protein Science (1997) 8.6.1-8.6.12 Copyright © 1997 by John Wiley & Sons, Inc.
Conventional Chromatographic Separations
8.6.1 Supplement 8
Hydroxylapatite Matrices for Standard Chromatography and HPLC
Table 8.6.1
Matrix
Form
Binding capacity (mg/g dry matrix) BSA
DNA
Lysozyme
Suppliera Remarks/applications
Bio-Gel HT
Hydrated
10
>0.5
—
BR
Suitable for general use in purification of proteins and nucleic acids Dehydrated form of Bio-Gel HT
Bio-Gel HTP
Dry
10
>0.5
—
BR
Bio-Gel HTP (DNA-grade)
Dry
10
>0.8
—
BR
Small particle size to increase capacity and selectivity for dsDNA; suitable for batch use or short columns only
Hydroxylapatite Fast Flow
Dry
>12
>0.7
—
CB
Hydroxylapatite High Resolution Hydroxyapatite Type I
Dry
>15
>1.2
—
CB
Recommended for applications requiring high flow rate and binding capacity Recommended for analytical applications
Hydrated
—
—
—
S
Hydroxyapatite Type III
Hydrated
—
—
—
S
HA-Ultrogel
Hydrated
—
—
—
S
Macro-Prep Ceramic Type I
Dry
—
—
>25
BR
Macro-Prep Ceramic Type II
Dry
—
—
>12.5
BR
Hydroxylapatite HPLC grade TSK gel HA-1000
Dry
>15
>1.2
—
CB
Suitable for HPLC
Prepacked column
>20
—
—
TH
Suitable for HPLC; available as prepacked glass and stainless steel columns; use with guard column kit HA-1000 recommended
Contains silica gel as crystal initiator Microcrystalline; requires no precycling; good flow rates even in large columns Ceramic, macroporous; high protein-binding capacity; best for acidic proteins; excellent flow rates and mechanical stability Macro-Prep Physical properties as Type I; recommended for purification of nucleic acids; low affinity for albamin, therefore recommended for immunoglobulin work
aAbbreviations: BR, Bio-Rad; CB, Calbiochem; S, Sigma; TH, TosoHaas. For addresses and phone numbers of suppliers, see SUPPLIERS APPENDIX.
Materials Hydroxylapatite matrix (see Table 8.6.1) Loading buffer: 10 mM sodium phosphate, pH 6.8 (APPENDIX 2E) Protein sample to be purified Gradient solutions: 0.01 and 0.4 M sodium phosphate, pH 6.8 (APPENDIX 2E) Loading buffer (see above) with 0.02% sodium azide
Hydroxylapatite Chromatography
Side-arm flask and vacuum source Glass chromatography column (see Strategic Planning) with two adaptors or removable end piece and one adaptor (Fig. 8.3.1), tubing, and optional packing reservoir (e.g., Pharmacia Biotech C or XK series)
8.6.2 Supplement 8
Current Protocols in Protein Science
Tubing clamps Peristaltic pump Buffer reservoirs Fraction collector UV monitor with 280-nm filter, connected to a chart recorder Gradient mixer Prepare gel 1. Calculate the amount of hydroxylapatite matrix required based on protein-binding capacity. Capacity of the matrix is usually supplied by the manufacturer
If matrix is supplied dry 2a. Degas a sufficient volume of loading buffer in a side-arm flask under vacuum. Add one part matrix to six parts loading buffer and resuspend by swirling gently. Do not use a magnetic stirrer as this will damage the hydroxylapatite crystals.
3a. Allow the slurry to settle for a minimum of 20 min, then decant the fines, which are in the cloudy solution above the settled matrix. 4a. Add a volume of loading buffer approximately equal to the volume of the settled bed and mix gently by swirling. Allow matrix to settle. 5a. If the matrix settles to give a sharp dividing line between the bed and clear buffer above, swirl gently to resuspend and proceed to packing of the column (step 6). If the supernatant buffer is still cloudy, repeat steps 3a and 4a, then proceed to step 6. If matrix is supplied hydrated 2b. Resuspend the matrix (in its own storage buffer as supplied by the manufacturer) by swirling gently. 3b. Pour a sufficient volume of the suspension into a beaker and allow to settle for a minimum of 20 min. Decant the fines. 4b. If the buffer in which the matrix was shipped is different from the loading buffer (10 mM sodium phosphate) add a volume of loading buffer approximately equal to that of the matrix, mix by swirling gently, allow to settle for a minimum of 20 min, then decant. 5b. Add a volume of loading buffer approximately equal to that of the matrix and swirl gently to resuspend. Proceed to packing of the column. Pack column 6. Examine the column to check that it is clean and undamaged and that sinters, support nets, and O rings are present and intact. 7. Insert bottom adaptor if used, or screw in the end piece. Mount column in a laboratory stand with clamps to ensure that it is vertical. If the volume of the resin slurry is greater than that of the column, fit a packing reservoir (Fig. 8.3.1). 8. Pour a small volume of loading buffer into the column and check that it flows freely through the bottom sinter. Clamp off the outlet tubing with a tubing clamp, leaving ∼2 cm of buffer above the sinter. 9. Swirl the hydroxylapatite slurry (step 5a or 5b) gently to resuspend, then pour it all into the column at once and allow 2 to 3 cm to settle under gravity. Release the tubing clamp and allow the rest of the bed to pack under gravity flow. 10. Attach a peristaltic pump to the bottom outlet and pump 5 bed volumes of loading buffer through the column to equilibrate.
Conventional Chromatographic Separations
8.6.3 Current Protocols in Protein Science
Supplement 8
As a rough guide, a standard general-purpose matrix such as Bio-Gel HT (see Table 8.6.1) may be used at a linear flow rate of 75 cm/hr. Fine-particle DNA columns should be run at ∼1⁄10 this rate. Macroporous material—e.g., Macro-Prep Ceramic (Table 8.6.1) can tolerate flow rates up to 5000 cm/hr. The linear flow rate (cm/hr) is equal to the volumetric flow rate (cm3/hr) divided by the column cross-sectional area (cm2). The column should be equilibrated at a flow rate 25% to 50% higher than that anticipated for the separation to ensure that there is no further compaction of the bed during the run. If the manufacturer recommends a maximum flow rate, this should never be exceeded.
11. Insert the top adaptor, lower it onto the column bed, and tighten. Buffer will emerge from the tubing, which should remain filled to prevent air from being drawn into the column.
12. Place the tubing from the top adaptor into a reservoir of loading buffer clamped to the support stand at the same level as the top of the column. 13. Pump a further 5 bed volumes of loading buffer through the column. If the bed shrinks so that there is a gap between the top adaptor and the surface of the bed, carefully loosen the top adaptor, lower it onto the bed and retighten. The column is now ready and may be used at once or stored for several weeks at 4°C.
Prepare sample 14. If necessary, free sample of particulate matter by centrifuging or passing it through a 0.45-µm filter. 15. Adjust sample to the same pH and a similar salt concentration as the loading buffer. If only the pH of the sample is different from that of the loading buffer, this can be carefully adjusted by adding 10 mM sodium dihydrogen phosphate (to decrease pH) or 10 mM disodium hydrogen phosphate (to increase pH). If the salt concentration is too high, the sample should be dialyzed against an appropriate buffer. Buffer exchange may also be effected by use of a desalting column (UNIT 8.3) or a membrane concentration device (UNIT 4.4). The sample may also be diluted with low-ionic-strength buffer.
Chromatograph sample and regenerate column 16. If the column has been stored for some time, pump a little loading buffer through it and check the bed for air bubbles or cracks and the tubing connections for leaks. 17. Transfer the top adaptor tubing from the buffer reservoir to the sample and pump the sample onto the column, collecting fractions with a fraction collector and monitoring the A280 using a UV monitor with a 280-nm filter, connected to a chart recorder. Take care not to introduce air into the column.
18. Wash the column with loading buffer until the A280 returns to baseline, collecting the flowthrough and wash fraction, as these contain the material which does not bind to the matrix. 19. Degas appropriate volumes of 0.01 and 0.4 M sodium phosphate, pH 6.8, as in step 2a. Connect gradient mixer to column and place the two gradient solutions in the respective reservoirs. Elute the bound material using a linear gradient from 0.01 to 0.4 M sodium phosphate with a gradient volume 6 to 10 times the bed volume. Elute any material still bound by washing with 0.4 M sodium phosphate, pH 6.8. Collect 20 to 40 fractions of a size appropriate to the volume of the gradient until all bound material has been eluted (as judged by the A280 of the eluant). Hydroxylapatite Chromatography
20. Regenerate column by washing with 5 bed volumes of loading buffer, then wash with 2 bed volumes of loading buffer containing 0.02% sodium azide. Store column at 4°C.
8.6.4 Supplement 8
Current Protocols in Protein Science
CAUTION: Sodium azide is toxic and potentially explosive. Avoid contact with metals and acids and do not lyophilize solutions containing azide. See APPENDIX 2A for additional guidelines on the handling and disposal of sodium azide. If the manufacturer recommends a different procedure for column regeneration, this should be followed. Figure 8.6.1 and Table 8.6.2 show results for low-pressure chromatography.
1.2
protein concentration
1.0
1.0
0.8
0.8
proteinase activity
0.6
0.6
0.4
0.4
0.2
0.2
A595 (HIV proteinase assay)
A599 (Bradford protein assay)
1.2
21 25 29 33 37 41 45 49 53 57 61 65 69 73 Fraction number
Figure 8.6.1 Chromatogram illustrating purification of HIV-2 proteinase on a hydroxylapatite column (A.V. Broadhurst and A.J. Ritchie, unpub. observ.). 139 g recombinant E. coli expressing HIV-2 proteinase were lysed by lysozyme digestion and sonication. The soluble fraction was treated with 1% streptomycin sulfate to precipitate nucleic acids. The supernatant was brought to 25% saturation with solid ammonium sulfate and the pellet was discarded. The ammonium sulfate was increased to 40% saturation and the pellet containing the proteinase was dialyzed against 10 mM sodium phosphate, pH 6.8, containing 0.0125% Tween 20. This fraction was applied to a 4.5 × 19–cm Bio-Rad HT hydroxylapatite column, bed volume 300 ml, at a linear flow rate of 10 cm/hr. The column was washed with 250 ml 10 mM sodium phosphate (pH 6.8)/0.0125% Tween followed by 250 ml 25 mM sodium phosphate (pH 6.8)/0.0125% Tween 20. Bound proteins were eluted with a 600 ml gradient of 25 to 250 mM sodium phosphate, (pH 6.8)/0.0125% Tween. The fractions were assayed for protein by the Bradford method (the dotted plot and the y axis on the left; Bradford, 1976; also see UNIT 3.4) and for proteinase activity using a colorimetric assay (the solid plot and the y axis on the right; Broadhurst et al., 1991). Table 8.6.2 Typical Purification Scheme for HIV-2 Proteinase Incorporating Hydroxylapatite Chromatographya
Purification stage Crude E.coli lysate 40% ammonium sulfate pellet Pooled peak fractions from hydroxylapatite column Pooled concentrated fractions from G-75 gel-filtration column
Total proteinase Specific activity activity (U) (U/mg) 79,120 54,590 31,995
13.9 48.2 194
18,333
561
U is arbitrarily defined as the amount of enzyme required to release 1 µmol product/min at 37° in a colorimetric assay (Broadhurst et al., 1991). a1
Conventional Chromatographic Separations
8.6.5 Current Protocols in Protein Science
Supplement 8
ALTERNATE PROTOCOL 1
SIMPLE COLUMN CHROMATOGRAPHY FOR PROTEIN PURIFICATION USING HYDROXYLAPATITE If only basic equipment is available, simplified methodology in which the pump is replaced by a gravity column may be applied successfully. The method described below is also useful for quick trial runs to assess the suitability of hydroxylapatite for particular applications. Additional Materials (also see Basic Protocol 1) Step gradient solutions: e.g., 0.025, 0.05, 0.10, 0.25, and 0.4 M sodium phosphate, pH 6.8 Column with integral bottom sinter and airtight lid: e.g., Kontes Flex-Column (Kontes Glass) or Bio-Rad Econo-Column Frit to place on top of resin bed (Kontes Glass) 1. Prepare matrix. (see Basic Protocol 1, steps 1 to 5a or 5b). 2. Pour matrix slurry into column and wash with 5 bed volumes of degassed loading buffer, allowing the column to drain by gravity. Do not allow column to run dry. 3. Place frit on top of bed to prevent disturbance of the surface. Avoid trapping air bubbles beneath it. 4. Prepare sample (see Basic Protocol 1, steps 14 and 15). Load sample on top of the frit and allow it run through the bed. When all the sample has been loaded, wash with 1.5 bed volumes of loading buffer. Combine the flowthrough and wash fractions, which will contain unbound material. 5. Wash with 8 bed volumes of loading buffer and discard the wash fractions. 6. Wash sequentially with 2.5 bed volumes each of step gradient solutions of increasing concentration—e.g., 0.025, 0.05, 0.10, 0.25, and 0.4 M sodium phosphate, pH 6.8 (degassed before use; see Basic Protocol 1, step 2a). Collect each wash separately.
ALTERNATE PROTOCOL 2
BATCH PROCESSING FOR PROTEIN PURIFICATION USING HYDROXYLAPATITE If hydroxylapatite is to be used with very crude material early in the purification schedule, or if the volume to be processed is very large, this batch separation procedure may be appropriate. Batch processing does not require sophisticated equipment; it can be done quickly and is useful for trial runs. Materials (also see Basic Protocol 1) Step gradient solutions: e.g., 0.025, 0.05, 0.10, 0.25, and 0.4 M sodium phosphate, pH 6.8 Sintered-glass funnel Side-arm flask and vacuum source NOTE: Degassing of buffers is not essential for batch processing. 1. Prepare matrix (see Basic Protocol 1, steps 1 to 5a or 5b). 2. Carefully remove all supernatant buffer from the settled matrix by decantation. 3. Prepare sample (see Basic Protocol 1, steps 14 and 15).
Hydroxylapatite Chromatography
If the sample volume is small, add sufficient 10 mM sodium phosphate, pH 6.8, to make it approximately twice the bed volume.
8.6.6 Supplement 8
Current Protocols in Protein Science
4. Mix the sample with the hydroxylapatite matrix by gentle swirling. Leave for 15 min. 5. Pour slurry into sintered-glass funnel and apply a very gentle vacuum until the buffer just reaches the top of the matrix. IMPORTANT NOTE: Do not allow the matrix to dry out.
6. Wash matrix in funnel with a small amount of loading buffer. Collect and combine the flowthrough and wash fractions. 7. Wash well with loading buffer and discard wash. 8. Elute bound proteins by washing with a step gradient of increasing phosphate concentration (see Alternate Protocol 1, step 6). HPLC OF PROTEINS AND NUCLEIC ACIDS USING HYDROXYLAPATITE Hydroxylapatite columns can be used for the chromatography of both proteins and nucleic acids. HPLC is especially useful for analytical applications, as the technique offers high resolution, excellent reproducibility, and fast analysis times. For occasional users, readypacked columns are recommended. These are usually of good quality and are tested and certified before sale.
BASIC PROTOCOL 2
The protocol described here is a general one applicable to the separation of proteins or nucleic acids. Using a method of this type, it is possible to quantitate the amount of a particular protein or nucleic acid in a sample by comparison with a range of standard samples of known concentration. Materials 0.01 and 0.4 M sodium phosphate buffer, pH 6.8 (APPENDIX 2E; prepare using boiled H2O) Protein or nucleic acid standard solutions Protein or nucleic acid test samples 0.2 M sodium hydroxide 0.01 M sodium phosphate, pH 6.8 (APPENDIX 2E) with 0.02% (w/v) sodium azide 0.45-µm filters (for buffers) 0.22-µm filters (for samples) ∼7.5-mm × 7.5-cm HPLC column packed with suitable grade of hydroxylapatite (Table 8.6.1) Guard column recommended for HPLC column used HPLC apparatus equipped with pumps, gradient mixer, autosampler or injection loop, integrator, and UV monitor Prepare buffers and sample 1. Filter 0.01 and 0.4 M sodium phosphate buffers through a 0.45-µm filter. Degas in side-arm flasks under vacuum before use. 2. Filter sample through a 0.22-µm filter. If sample is very turbid it may be necessary to centrifuge it first to avoid blocking the filter.
Prepare column and chromatograph samples 3. Follow any instructions supplied by the manufacturer relating to preparation of column and assembly of HPLC system. 4. Equilibrate column with 0.01 M sodium phosphate buffer, pH 6.8, prepared as in step 1.
Conventional Chromatographic Separations
8.6.7 Current Protocols in Protein Science
Supplement 8
4 2 3
A280
1
0
5
10
15
20
25
Elution time (min) Figure 8.6.2 Chromatogram illustrating purification of monoclonal antibody from mouse ascites fluid. A Bio-Rad Bio-Gel HPHT column with guard column was used and 0.5 ml mouse ascites fluid was applied. The column was eluted with a linear gradient of 0.01 to 0.30 M sodium phosphate, pH 6.8 at a flow rate of 1.0 ml/min. Peak 1 contains low-molecular-weight material, peak 2 contains albumin and serum protease, peak 3 contains other serum (ascites) proteins, and peak 4 contains essentially pure IgG. The linear gradient is traced via the conductance (dotted line). Adapted from Bio-Rad Technical Bulletin US/EG 1115.
5. Program the HPLC to inject 20 µl of sample and set the UV monitor to 260 nm for nucleic acids or 280 nm for proteins. Place 0.01 and 0.4 M sodium phosphate buffers, pH 6.8, in the appropriate reservoirs of the gradient mixer and run a 30-min linear gradient from 0.01 to 0.4 M sodium phosphate at a flow rate of 1 ml/min. 6. Run a standard through the column to to check its performance using the parameters in step 5. Collect 1-ml fractions. 7. Run the standard at a range of concentrations to generate a standard curve. 8. Run the test samples. 9. Use the integrated peak areas of the standards and test samples to calculate the concentration of the protein/nucleic acid of interest in the samples. See Figure 8.6.2 for representative results.
Regenerate and store column 10. At the end of each sample batch, inject 1 column volume of 0.2 M sodium hydroxide to remove any strongly bound material. Wash extensively with 0.4 M sodium phosphate. 11. Store column at room temperature in 0.01 M sodium phosphate, pH 6.8 containing 0.02% sodium azide. Hydroxylapatite Chromatography
CAUTION: Sodium azide is toxic and potentially explosive. Avoid contact with metals and acids and do not lyophilize solutions containing azide. See APPENDIX 2A for additional guidelines on the handling and disposal of sodium azide.
8.6.8 Supplement 8
Current Protocols in Protein Science
COMMENTARY Background Information Calcium phosphate gels have been used for chromatography for many years. The material originally used was unsatisfactory with respect to flow rate, stability, and elution characteristics, and fell out of favor as newer matrices such as HA Ultrogel and Macro-Prep Ceramic became available. In the 1950s, Tiselius and co-workers described a process for preparing hydroxylapatite, a crystalline form of calcium phosphate, and demonstrated its usefulness for protein separation (Tiselius et al., 1956). Material prepared by this method demonstrated superior flow rates and has been commercially available for many years. Recently, some new forms of hydroxylapatite with spherical particles have been developed, and these have demonstrated even better flow rates as well as enough mechanical strength to be used in highpressure systems. These technological advances, combined with hydroxylapatite’s unique mode of separation, make this medium a useful addition to the chromatographer’s repertoire. Protein binding to hydroxylapatite is mediated through interactions between the amino and carboxyl groups of the protein and the Ca2+ and PO43– groups of the hydroxylapatite crystal lattice. Basic proteins are thought to bind via electrostatic interactions between their amino groups and the surface PO43– groups, the positive charge rather than the chemical nature of the group being the important factor (Gorbunoff, 1984b). Acidic and neutral proteins bind to the Ca2+ sites via their carboxyl groups; this interaction encompasses both electrostatic interactions and specific effects (Gorbunoff, 1984b). As a general principle, basic proteins with an isoelectric point (pI) of 8 can be eluted by increasing the concentration of the phosphate buffer, and they are also displaced by very low concentrations of Mg2+ or Ca2+ ions and moderate concentrations of NaCl (Gorbunoff, 1984a). Acidic or neutral proteins can be eluted by increasing the phosphate concentration, but not by adding Mg2+ or Ca2+ ions or NaCl. Increasing the pH of the elution buffer promotes desorption of proteins irrespective of pI (Gorbunoff, 1984a). There are many proteins whose interactions with hydroxylapatite are not as easily predicted on the basis of pI alone—e.g., β-lactoglobulin, ovomucoid, trypsinogen (Gorbunoff and Timasheff, 1984), phosphoproteins (Bernardi et al., 1972), and metalloproteins (Atkinson et al.,
1973). Tertiary structure is also important for binding, and proteins with floppy or unstructured regions such as the glycoprotein ovomucoid, or those that have been denatured, show reduced affinity for hydroxylapatite (Gorbunoff, 1984a). The distribution of surface charge may also play a part (Gorbunoff, 1984b). Nucleic acids bind to hydroxylapatite by interaction of the charged phosphate backbone with the Ca2+ ions at the crystal face. Doublestranded DNAs bind more tightly than singlestranded DNAs, possibly because their planar nature maximizes the interaction. (Martinson, 1973). Small double- and single-stranded DNAs appear to separate by size. tRNAs can also be separated, and secondary and tertiary structures may be important in this context (Kawasaki et al., 1986). Hydroxylapatite chromatography is performed using mild conditions at moderate pH and salt concentration, which favors the preservation of biological activity. The matrix has low nonspecific binding, and small molecules such as amino acids do not bind well. It is chemically stable at pHs between 6 and 11 and can be used with denaturing agents, detergents, and reducing agents such as dithiothreitol. EDTA and chelating buffers such as PIPES should be avoided as these bind the calcium. The unique mode of separation of hydroxylapatite, combined with the improved handling characteristics of the newer forms available, make it a useful adjunct to the more commonly used techniques such as ion-exchange and hydrophobic-interaction chromatography. Hydroxylapatite has proved useful in the separation of IgG idiotypes that cannot easily be separated by other techniques (Juarez-Salinas et al., 1986), as well as for the isolation of immunoglobulins from ascites fluid and tissue culture media (Smith et al., 1984). Another application may be the secondary purification step for recombinant proteins with N-terminal affinity tags such as maltose-binding protein (MBP), glutathione-S-transferase (GST), or 6His. When these proteins have been isolated by affinity chromatography, any breakdown products produced by C-terminal degradation and any premature termination fragments will also bind to the resin, as they are tagged. These contaminants can be difficult to remove by gel filtration or ion-exchange chromatography as they may differ very little in size or charge from the protein of interest; hence hydroxylapatite chromatography may assist in the separation.
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As an example, during the preparation of human cytomegalovirus proteinase for crystallographic studies, protein that had been purified using Mimetic Green and Q Sepharose and that appeared to be homogeneous on PAGE, separated into two peaks on a hydroxylapatite column (A.V. Broadhurst, unpub. observ.). The ability of hydroxylapatite to bind and separate nucleic acids can be advantageous during chromatography of proteins derived from recombinant E. coli. Crude extracts contain large quantities of DNA, which can bind to proteins or ion-exchange resins and contaminate the product. Hydroxylapatite can be used to remove nucleic acids during the purification.
Critical Parameters Selecting format Careful consideration must be given to choosing the most appropriate chromatographic format for the application. If hydroxylapatite is to be used with very crude material early in the purification schedule, or if the volume to be processed is very large, a batch separation (see Alternate Protocol 2) may be suitable. Batch processing does not require sophisticated equipment; it can be done quickly and is useful for trial runs. Low-pressure chromatography (see Basic Protocol 1 and Alternate Protocol 1) can be used to prepare milligram to gram quantities of protein. Semiautomated or fully automated systems are available, but satisfactory results can be obtained with quite basic equipment. This type of chromatography is used primarily for preparative rather than analytical purposes. HPLC (see Basic Protocol 2), although it can be scaled up for preparative use, is particularly suitable for analytical applications. Its main advantages are high resolution, reproducibility, and rapid throughput.
Hydroxylapatite Chromatography
Selecting matrix Various grades and types of hydroxylapatite are commercially available (see Table 8.6.1). In general, hydroxylapatite with a small particle size will be suitable for analytical applications, giving excellent resolution but operating at low flow rates. Material with a larger particle size can be used at higher flow rates but with some loss of resolution. Hydroxylapatite can be difficult to work with as its crystalline structure is fragile and easily damaged, but some of the newer types (TSKgel HA-1000 from TosoHaas and MacroPrep ceramic hydroxylapatite from Bio-Rad) are purported to be more robust and
are suitable for use with-high pressure systems. There are several types of ready-prepared columns available commercially. These are designed for particular applications such as HPLC or low-pressure chromatography; some are of particular utility for the separation of proteins and others for separation of nucleic acids. Selecting buffers Sodium or potassium phosphate buffers at near-neutral pH are used for the majority of hydroxylapatite applications. Low phosphate concentration and low pH increase protein binding; high phosphate concentration and high pH promote desorption. In this context, low and high pH refer to the range 6 to 8, over which phosphate is an effective buffer. Potassium phosphate is more soluble than sodium phosphate at lower temperatures and is recommended for cold-room use. The protein-binding capacity of hydroxylapatite is maximal at pH 6.6 to 7 and decreases sharply above pH 7 (Atkinson et al., 1973). The matrix itself is chemically stable over a pH range of 6 to 11, but dissolves below pH 5.5. Buffers other than phosphate have been employed—e.g., MOPS with a phosphate gradient (de Gunzberg et al., 1984)—but should be used with caution as they may damage the matrix. 10 mM Tris⋅Cl can be used, but may cause some deterioration over time. Chelating buffers such as PIPES must be avoided. Buffer pH should always be within the range over which the sample protein is known to be stable. Various buffer additives can be employed. Glycerol, added to stabilize proteins, can be used at a concentration of 10% to 15%; it increases buffer viscosity so that a reduction in flow rate may be necessary. Reducing agents such as dithiothreitol and 2-mercaptoethanol, as well as nonionic detergents such as Triton X-100 and Tween 20, are compatible with hydroxylapatite. SDS has been used, but can reduce nonspecific binding and alter elution characteristics; it is also rather insoluble at 4°C and therefore is probably best avoided. Chelating agents such as EDTA are contraindicated as they bind to the calcium ions of hydroxylapatite and will rapidly destroy the column.
Troubleshooting Many of the problems encountered during hydroxylapatite chromatography are common to all types of column chromatography, but a few are unique owing to the crystalline mineral nature of the matrix.
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Column bed drying out or becoming filled with bubbles Drying out of the bed is usually caused by the system not being airtight. Ensure that all O rings and seals are present and in good condition. It may help to have the buffer reservoir slightly higher than the top of the column. Drying out can also occur if the top sinter or the top of the bed becomes clogged, causing the pump to create a vacuum in the bed. This might be alleviated by reducing the flow rate; if this does not work, remove the top sinter, wash it, and replace. If the problem persists it may be necessary to modify the sample preparation. Bubbles that appear in the gel are usually caused by inadequate degassing of the buffers. Dissolved gases will be released under reduced pressure or if the buffers/column are moved from a colder to a warmer environment. Degas all buffers (unless they contain volatile solvents) under vacuum before use. Bubbles may also appear if high concentrations of detergent are included in the buffers; this can be minimized by avoiding shaking or vigorous stirring of the buffers and using a moderate flow rate. Inadequate or decreasing flow rate This problem may be caused by packing the bed at too high a flow rate, causing it to become compressed. Manufacturers’ recommended flow rates should never be exceeded; this is particularly important with hydroxylapatite because of its inherently fragile nature. The presence of fines (resulting from inadequate removal before packing the bed) or degraded material can also cause poor flow rates. Ensure that the matrix is prepared as recommended; hydroxylapatite is easily damaged by rough treatment such as stirring with magnetic stirring bars and should be resuspended only by gentle swirling. If hydroxylapatite columns are stored for long periods the top of the bed may become hard and crusted, causing reduced flow rate. This results from the reaction of the calcium ions with carbonate from water used to prepare the buffers. Boiled, degassed water should be used. The hard, crusted material can be removed and replaced with a little fresh hydroxylapatite slurry. If the flow rate achievable is not high enough to allow the experiment to be performed in a reasonable time, consider whether a matrix with a larger particle size would be more suitable, or whether a batch process might be preferable.
If the flow rate is constant during the preliminary washing and equilibration of the column but decreases during sample loading, the problem may lie with the sample itself. It is important that particulate matter be removed from the sample before loading or it will block the sinter or the top of the gel bed. The sinter can be removed, washed, and replaced as an interim measure, but improved sample preparation is the cure. Crude extracts of recombinant E. coli, which are increasingly used as the starting point for protein purification, can be problematic. Even if the extract is clarified by centrifugation, it may be extremely viscous due to the high DNA content. It is often possible to reduce the viscosity enough to achieve a reasonable flow rate by diluting one part sample with three parts buffer before loading. If this fails, try precipitating the nucleic acids with 0.01% aqueous polyethyleneimine or 1% streptomycin sulfate, or treat the sample with DNase. UNIT 4.5 contains additional guidelines on clarification of samples and nucleic acid removal. Protein not bound to column If the protein of interest is only partially bound, with significant amounts appearing in the flowthrough, the binding capacity of the column may have been exceeded. Try increasing the ratio of matrix to total protein. If the protein is not bound using 10 mM sodium phosphate, pH 6.8, try reducing the molarity of the sodium phosphate. Concentrations as low as 1 mM can be used, although the buffering capacity will be small at this concentration. If the protein is stable at lower pH, the pH of the loading buffer can be decreased to 6 to facilitate binding. Do not reduce the pH below 5.8, as the hydroxylapatite matrix will dissolve at lower pH. Adding 0.005 mM CaCl2 to the buffer will increase binding of acidic proteins. CaCl2 should only be used with low molarities of phosphate (<200 mM) or calcium phosphate will precipitate out of solution. Even if none of the above modifications are successful, hydroxylapatite chromatography might still be a useful initial step if a significant amount of contaminating material is removed by binding to the matrix. Poor separation of proteins on gradient and problems with eluting bound protein If there are protein peaks that do not separate completely, try increasing the volume of the gradient. Do not exceed more than ∼12 bed
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volumes as there will be no benefit from this. If all the bound material elutes very early on in the gradient, try using a narrower gradient— e.g.,10 mM to 150 mM sodium phosphate, pH 6.8. If most of the proteins elute near the end of the gradient, try using a higher loading concentration of sodium phosphate—e.g., 50 to 100 mM. The molarity of the elution buffer can be increased. If phosphate concentrations >0.4 M are required for complete elution and the chromatography is to be done at 4C, use potassium phosphate buffer instead of sodium phosphate, as it is more soluble. Desorption of proteins can also be achieved by increasing the pH of the buffer. Basic proteins can be eluted with Ca2+ at concentrations of ∼0.003 M. This cannot be done in phosphate buffer as calcium phosphate will precipitate out. Wash the column with 1 mM NaCl, then with 0.003 M CaCl2 in 1 mM NaCl.
Anticipated Results High recoveries can be expected with hydroxylapatite; near-quantitative yields have been reported with 90% to 95% being common. Retention of activity is usually excellent because of the mild conditions employed (Kato et al., 1987). Depending on the nature of the sample to be chromatographed, >20-fold purification is achievable.
Time Considerations Small columns can be prepared and a chromatographic run carried out in a working day. Larger columns take longer to run and therefore will need to be prepared in advance.
Literature Cited
protein utilising the principle of protein-dye binding. Anal. Biochem. 72:248-254. Broadhurst, A.V., Roberts, N.A., Ritchie, A.J., Handa, B.K., and Kay, C. 1991. Assay of HIV-1 proteinase: A colorimetric method using small peptide substrates. Anal. Biochem. 193:280-286. de Gunzberg, J., Part, D., Guiso, N., and Veron, G. 1984. An unusual adenosine 3′, 5′-phosphate dependent protein kinase from Dictyostelium discoideum. Biochemistry 23:3805-3812. Gorbunoff, M.J. 1984a. The interaction of proteins with hydroxyapatite. 1. Role of protein charge and structure. Anal. Biochem. 136:425-432. Gorbunoff, M.J. 1984b. The interaction of proteins with hydroxyapatite. 2. Role of acidic and basic groups. Anal. Biochem. 136:433-439. Gorbunoff, M.J. and Timasheff, S.N. 1984. The interaction of proteins with hydroxyapatite. 3. Mechanism. Anal. Biochem. 136:440-445. Juarez-Salinas, H., Ott, G.S., Chen, J.-C., Brooks, T.L., and Stanker, L.H. 1986. Separation of IgG idiotypes by high performance hydroxylapatite chromatography. Methods Enzymol. 131:615622. Kato, Y., Nakamura, K., and Hashimoto, T. 1987. High performance hydroxyapatite chromatography of proteins. J. Chromatogr. 398:340-346. Kawasaki, T., Ikeda, K., Takahashi, S., and Kuboki, Y. 1986. Further study of hydroxylapatite highperformance liquid chromatography using both proteins and nucleic acids, and a new technique to improve chromatographic efficiency. Eur. J. Biochem. 155:249-257. Martinson, H.G. 1973. The basis of fractionation of single-stranded nucleic acids on hydroxylapatite. Biochemistry 12:2731-2736. Smith, G.J., McFarland, R.D., Reisner, H.M., and Hudson, G.S. 1984. Lymphoblastoid cell–produced immunoglobulins: Preparative purification from culture medium by hydroxylapatite chromatography. Anal. Biochem. 141:432-436. Tiselius, A., Hjerten, S., and Levin, O. 1956. Protein chromatography on calcium phosphate columns. Arch. Biochem. Biophys. 65:132-155.
Atkinson, A., Bradford, P.A., and Selmes, I.P. 1973. The large scale preparation of chromatographic grade hydroxylapatite and its application in protein separation procedures. J. Appl. Chem. Biotechnol. 23:517-529.
Key References
Bernardi, G., Giro, M.-G., and Gaillard, C. 1972. Chromatography of polypeptides and proteins on hydroxyapatite columns: Some new developments. Biochim. Biophys. Acta 278:409-420.
A series of three consecutive papers describing the basis of interaction of proteins with hydroxylapatite. Practical aspects are described, and there is extensive discussion of theoretical considerations.
Bio-Rad Technical Bulletin 1115 US/EG. Bio-Gel HPHT for protein and nucleic acid HPLC: New high performance hydroxyapatite column. BioRad Laboratories, Hercules, Calif. Bradford, M.M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of
Gorbunoff, 1984a. See above. Gorbunoff, 1984b. See above. Gorbunoff and Timasheff, 1984. See above.
Anne V. Broadhurst Roche Products Ltd. Welwyn Garden City, Hertfordshire United Kingdom
Hydroxylapatite Chromatography
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HPLC of Peptides and Proteins High-performance liquid chromatography (HPLC) is an essential tool for the purification and characterization of biomacromolecules. The choice of the chromatographic method and the type of high-performance equipment are determined by the molecular nature of the investigated molecules and the aim of the research. There are eight basic modes of HPLC currently in use for peptide and protein analysis and purification, namely size-exclusion chromatography (HP-SEC), ion exchange chromatography (HP-IEX), normal phase chromatography (HP-NPC), hydrophobic interaction chromatography (HP-HIC), reversed-phase chromatography (RP-HPLC), hydrophilic interaction chromatography (HP-HILIC), immobilized metal ion affinity chromatography (HPIMAC), and biospecific/biomimetic affinity chromatography (HP-BAC), and a number of subsets of these chromatographic modes, e.g., mixed mode chromatography (HP-MMC), charge transfer chromatography (HP-CTC), or ligand-exchange chromatography (HP-LEC). In terms of usage, versatility, and flexibility, RP-HPLC techniques dominate the application world with peptides and proteins at the analytical- and laboratory-scale preparative levels. All of these various chromatographic modes can be operated under isocratic or gradient elution conditions, and in analytical or preparative situations. They all have common start-up procedures, which are outlined in the initial sections of this unit, and specific standard conditions that are detailed for the major modes and which represent starting points for further method development. As an example for appropriate method development, the RP-HPLC mode is described in greater detail and discussed according to four possible intended purposes, i.e.: (1) the purification of one component out of a natural or synthesized sample; (2) the simultaneous purification of several components; (3) the desalting of proteins or polypeptides obtained from other purification procedures; and (4) the characterization of the physicochemical properties of peptides or proteins. They all require a good understanding of the underlying common principles of polypeptide-ligand interaction. The basics of these principles are touched upon with references to further reading. Finally, a short section of this unit is dedicated to troubleshooting; however, many of the “check-back” confirmatory procedures im-
plicit to sound operational practices and the identification of suitable alternatives for the separation strategy are included in the section on method development.
THE PROPERTIES OF PEPTIDES AND PROTEINS AND THEIR IMPLICATIONS FOR HPLC METHOD DEVELOPMENT Peptides and proteins are a class of molecules containing amino acids as the fundamental units. The chemical organization (i.e., the primary structure or amino acid sequence) and the folded structure (i.e., the secondary, tertiary and quaternary structure) are the essential features of a polypeptide or protein, around which a chromatographic separation can be designed. Two sets of factors must be considered. The first relates to the structural properties of the amino acid entities themselves; the second relates to the chemical and physical attributes of the separation system per se.
Biophysical Properties of Peptides and Proteins The 20 naturally occurring L-α-amino acids found in peptides and proteins vary dramatically in terms of the properties of the side chain or R-groups. Table 8.7.1 lists some of the fundamental properties of the common L-α-amino acids found in peptides and proteins. This chemical diversity becomes even greater in circumstances where some of these side chains have been post-translationally modified with carbohydrates or lipid moieties. The sidechains are generally classified according to their polarity (e.g., non-polar or hydrophobic versus polar or hydrophilic). The polar side chains are divided into three groups: uncharged polar, positively charged or basic, and negatively charged or acidic side chains. Peptides and proteins generally contain several ionizable basic and acidic functionalities. They therefore typically exhibit characteristic isoelectric points with the overall net charge and polarity in aqueous solution varying with pH, solvent composition and temperature. Cyclic peptides without ionizable side chains will have zero net charge, and they represent an exceptional subgroup. The number and distribution of charged groups will influence the polarizability and ionization status of a peptide or protein, as well
Contributed by Reinhard I. Boysen and Milton T.W. Hearn Current Protocols in Protein Science (2001) 8.7.1-8.7.40 Copyright © 2001 by John Wiley & Sons, Inc.
UNIT 8.7
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Table 8.7.1
Properties of the Common L-α-Amino Acid Residues
3-Letter code
1-Letter code
Ala Arg Asn Asp Cys Gln Glu Gly His Ile Leu Lys Met Phe Pro Ser Thr Trp Tyr Val α-aminoe α-carboxylf
A R N D C Q E G H I L K M F P S T W Y V — —
Mass (Da) 71.08 156.19 114.10 115.09 103.14 128.13 129.12 57.05 137.14 113.16 113.16 128.17 131.20 147.18 97.12 87.08 101.11 186.21 163.18 99.13 — —
Accessible pKa of Partial spec. Rel. surface area a c hydrophobicityd 3 vol. (Å ) side-chain (Å2)b 88.6 173.4 117.7 111.1 108.5 143.9 138.4 60.1 153.2 166.7 166.7 168.6 162.9 189.9 122.7 89 116.1 227.8 193.6 140 — —
115 225 160 150 135 180 190 75 195 175 170 200 185 210 145 115 140 255 230 155 — —
— 12.48 — 3.9 8.37 — 4.07 — 6.04 — — 10.54 — — — — — — 10.46 — 7.7-9.2g 2.75-3.2g
0.06 −0.85 0.25 −0.20 0.49 0.31 −0.10 0.21 −2.24 3.48 3.50 −1.62 0.21 4.8 0.71 −0.62 0.65 2.29 1.89 1.59 — —
aFrom Zamyatnin, 1972. bFrom Chothia, 1974. cFrom Dawson et al., 1986. dFrom Wilce et al., 1995. eα-amino group present on all primary amino acids. fα-carboxyl group present on all primary amino acids. gFrom Rickard et al., 1991.
as the microscopic and global hydrophobicity. These important factors ultimately determine the selection of the optimal separation conditions for the resolution of peptide and protein mixtures. Table 8.7.1 can be used to evaluate the impact of amino acid composition on retention behavior. For example, this information can be used to direct the choice of eluent composition or the gradient range in RP-HPLC; to assess the impact on retention of amino acid substitution or deletion with small peptides; or alternatively to guide the identification of peptide fragments derived from tryptic digestion of proteins for further sequence analysis.
HPLC of Peptides and Proteins
Parameters of the Mobile Phase/Stationary Phase These parameters directly impact on the molecular properties of the polypeptide or protein during liquid chromatographic separations, and are listed in Table 8.7.2. In solution, a polypeptide or protein can, in principle, explore a relatively large array of conformational space. For small peptides (up to ∼15 amino acid residues) a defined secondary structure (α-helical, β-sheet or β-turn motif) is generally absent. With increasing polypeptide chain length, depending on the nature of the amino acid sequence, specific regions/domains of a polypeptide or protein can adopt preferred secondary, tertiary or quaternary structures. In aqueous solutions this folding, which internalizes the hydrophobic residues and thus stabilizes the
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Table 8.7.2 Chemical and Physical Factors of the Chromatographic System that Contribute to the Variation in the Resolution and Recovery of Polypeptides, Proteins and Other Biomacromolecules in HPLC Systems (Hearn, 2000a)
Mobile phase contributions
Stationary phase contributions
Organic solvents pH Metal ions Chaotropic reagents Oxidizing or reducing reagents Temperature Buffer composition Ionic strength Loading concentration and volume
Ligand composition Ligand density Surface heterogeneity Surface area Pore diameter Pore diameter distribution Particle size Particle size distribution Particle compressibility
Table 8.7.3 Relevant Absorption Bands and Extinction Coefficients in Proteins (Campbell and Dwek, 1984)
Group Peptide bond His Cys Trp Tyr Phe
Wavelength (nm)
Log εmax
190-210 211 250 280 274 257
2.0-3.8 3.8 2.5 3.7 3.1 2.3
polypeptide structure, becomes a significant feature of peptides and proteins for chromatographic separations. A critical factor in the selection of an HPLC procedure is that the choice of experimental conditions will inevitably cause perturbations of the conformational status of these biomacromolecules. Although polypeptide and protein conformational stability can be manipulated in a number of ways (e.g., mobile and stationary phase composition, temperature) in HPLC, in most cases an integrated biophysical experimental strategy—including 1H 2-dimensional NMR (UNIT 17.5), Fourier transformed infrared (FTIR), ESI-MS (UNIT 16.1), or circular dichroism–optical rotatory dispersion (CD-ORD) spectroscopy—is required in order to determine the secondary and higher-order structure of a polypeptide or protein in solution or in the presence of specific ligands. Availability of such instrumentation is not mandatory, but the quality of the interpretation of the experimental results will become more substantial when additional results are
independently obtained with such spectrometric procedures to confirm the participation of conformational or self-self aggregation effects with peptides or proteins under HPLC conditions.
DETECTION OF PEPTIDES AND PROTEINS IN HPLC The peptide bond absorbs strongly in the far-ultraviolet (UV) region of the spectrum (∼λ = 205 to 215 nm). Hence UV detection is the most widely used method for detection of peptides and proteins in HPLC (Table 8.7.3). Besides absorbing in the far-UV range, the aromatic amino acid residues (and to some extent cysteine) also absorb light above 250 nm. Knowledge of the UV spectra, in particular the extinction coefficients of the non-overlapping absorption minima of these amino acids, allows, in conjunction with UV-diode array detection (DAD) and second derivative or difference UV-spectroscopy, verification of peak purity and determination of the aromatic amino
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Table 8.7.4 UV Cutoff Values of Different Organic Solvents in RP-HPLC
Eluent Methanol Acetonitrile Isopropanol
UV cutoff (nm)a 205 188 205
aWavelength at which the absorbance of a 1-cm-
long cell filled with the solvent was 1.0, measured against water as reference.
acid content of peptides and proteins. Moreover, the knowledge of the relative UV/VIS absorbancy of a peptide or protein is therefore crucial, since the choice of detection wavelength of peptides and proteins in RP-HPLC (and in the other HPLC modes) depends on the different UV cutoffs of the eluents used (Table 8.7.4). The common use of λ = 215 nm as the preferred detection wavelength for most analytical reversed-phase applications (and for those of other HPLC modes) with peptides and proteins is a good compromise between detection sensitivity and potential detection interference due to buffer absorption. However, wavelengths between 230 and 280 nm are frequently employed in preparative applications, where the use of more sensitive detection wavelengths could result in overloading of the detector response (usually above an absorbance value of 2.0 to 2.5 AU).
START-UP PROCEDURES Correct selection of these first important steps may take more time than the ultimate experimental procedure if a high-performance separation of high resolution, robustness, and reproducibility is to be achieved. They require good planning and thorough work. The following details are representative of the types of equipment, materials, chemicals, and experimental protocols that can be routinely required for isocratic or gradient elution HPLC. Sample Peptide or protein sample (kept at 4°C if not used)
HPLC of Peptides and Proteins
Apparatus Pump module Mixing chamber Spectrophotometer with analytical or preparative flow cell Injection valve
Analytical (10 to 100 µl) or preparative (500 to 1000 µl) sample loop Column oven or thermostated column coolant– jacket coupled to recirculating cooler Autosampler (optional) Computer, printer, and software, e.g., Beckman, System Gold; Hewlett-Packard HP1090A liquid chromatograph, or Waters 600/486 HPLC system with attendant data management systems and system automation controllers Chemicals Acetonitrile (HPLC grade) Methanol (HPLC grade) Acetone (HPLC grade) Thiourea or sodium nitrate Milli-Q water Glassware Two 1-liter eluent bottles Two 1-liter measuring cylinders 10-ml measuring cylinder Waste bottle All glassware coming into contact with sample before and during analysis should be rinsed three times with Milli-Q water Mobile phase filtration facility Vacuum pump 1-liter reservoir Support base with glass frit and integral vacuum connection Funnel Clamp 47-mm membrane filter (0.2 µm PTFE) Gases Helium Nitrogen (for autosampler) Columns See respective sections below
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HPLC peptide standards See respective sections below Tools Screwdrivers: 1⁄ -in. and 1⁄ -in. flat-head screwdrivers 8 4 Phillips screwdriver no. 2 Wrenches: 12-in. adjustable wrench (for compressed gas tank regulator) Three open-end wrenches (two 1⁄4-in. × 1⁄16-in.; one 1⁄2-in. × 1⁄16-in.) for fittings columns and valves Two long-jaw needle-nose pliers Two bastard files Two hex key (Allen) wrench sets (metric and nonmetric) Tweezers Pump seal insertion tool (if required) Inner reamer Teflon tape Flashlight Magnetic pick-up tool Logbook All steps must be documented to facilitate troubleshooting and reproduction Spare parts Zero dead volume union Ferrules (steel, rheodyne) 1⁄16-in. short and long (for rheodyne valves) Bushing nuts One-piece PEEK fitting Tubing (steel, PEEK) Column frits In-line filter Inlet filter Fuses Miscellaneous Graduated 25-µl Hamilton glass syringe 1-ml syringe with truncated needle Conical vials for autosampler Laboratory coat Gloves Safety glasses Stopwatch
PREPARATION OF THE SAMPLE The following considerations are relevant to the preparation of samples containing peptides and proteins. 1. Dissolve the sample with half the target volume of eluent A (weak mobile phase). If the sample is not soluble, a small amount of eluent B (strong mobile phase) may be added (typi-
cally <25% of total volume). Small amounts of strong mobile phase may cause pre-elution of the sample depending on the sample loop size, column dimensions, and starting mobile phase composition of the gradient. 2. Inspect the sample for clearness and filter through a 0.2 µm PTFE filter if insoluble, opalescent, or solid particles are present. Alternatively centrifuge the sample using the supernatant for injection. Samples that are not fully dissolved should not be injected, since they can block injector and column. 3. Store sample if not in use at 4°C or −20°C depending on the planned storage time and usage. Peptides and protein samples can degrade at room temperature. Avoid repeated freeze-thawing of the sample. Rather, prepare small aliquots of the peptide or protein sample, which are kept at −20°C and are used for one task.
PREPARATION OF THE MOBILE PHASE The following considerations are relevant to the preparation of mobile phases to be used with samples containing peptides and proteins. 1. Prepare 1 liter each of eluent A (weak mobile phase) and B (strong mobile phase). 2. Mix each eluent either by stirring with a magnetic “flea” or by shaking a stoppered cylinder (time depends on volume). 3. Filter the eluents, first A, then B, through a 0.2 µm PTFE filter. Filtering of eluents increases the column lifetime and contributes to degassing. 4. Close unused eluent bottles with a stopper to avoid evaporation of the organic solvent.
SETTING UP THE HPLC INSTRUMENT The following steps can be done either with or without a column connected to the HPLC instrumentation. 1. Switch on gas supply, nitrogen for the operation of the autosampler (if available), and helium for the degassing of the eluents. 2. Degas the eluents for 10 min at a rate of 100 ml/min, then reduce to a rate of 20 ml/min, which may be maintained throughout the experiments. Avoid excessive sparging since this will change the eluent composition. 3. Switch the detector lamp on in order to warm it up. Conventional Chromatographic Separations
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PRIMING OF THE PUMPS AND LOW PRESSURE LINES WITH ELUENTS The following steps can be done either with or without a column connected to the HPLC instrumentation. 1. Open purge valve. 2. Set the pump at 100% buffer A and a flow rate of 1 ml/min. Under these settings, the eluent will bypass the column and travel directly to the waste fluid container (all wastes are collected for appropriate disposal). 3. Prime all tubing lines by switching the selector to the “prime line” position or by opening an additional valve which has an outlet to attach a syringe (see manufacturers’ handbook for specific details). 4. Draw (via the opened valve) 20 ml out of the eluent bottle with the syringe. Close the valve and discharge eluent from syringe in waste. 5. Prime all tubing lines again by drawing 10 ml in syringe, but keep the eluent in the syringe. 6. Switch selector to the “prime pump” position and push eluent gently through the pump. 7. Repeat the last four steps with 100% buffer B. 8. Put the selector in the “operate” position, close the valve, and remove the syringe from instrument. 9. Close the purge valve (now the eluent will travel through the column). This procedure should expel air bubbles from the pump heads and replace the previous eluent (∼20 ml) in the solvent lines.
PREPARATION OF THE HPLC SYSTEM
HPLC of Peptides and Proteins
The considerations below are relevant to the preparation of the HPLC system for use with samples containing peptides and proteins. The following steps can be done either with or without a column connected to the HPLC instrumentation. 1. Test the pump delivery system with eluents comprising 100% A and 100% B each for 5 min at a flow rate of 1 ml/min, collecting the eluent into a 10 ml cylinder. Some HPLC systems allow on-line pump diagnostics (e.g., Beckman System Gold). This procedure tests the reliability of the inlet and outlet valve, which may be blocked or not closing properly, e.g., due to salt from previous eluents. Some instrument systems allow the removal of the valve, which then can be sonicated in methanol for 15 min (care must be taken not to mix up
the inlet and outlet valves, which often cannot be visually distinguished). 2. Flush the HPLC system with the eluent (e.g., 50% A and 50% B) and monitor the detector baseline. If spikes occur after 15 min of flushing in the absence of the column, the pressure in the detector cell is too low, leading to an outgassing of air and cycling bubble formation. Use the back-pressure restrictor (alternatively a restriction capillary) on the detector outlet to slightly, and very carefully, enhance the back-pressure. Every piece of capillary must be checked for blockage beforehand by connecting to a pump, thus bypassing the column and detector, since the detector cell is very sensitive to high pressure (consult manufacturer’s handbook for details). 3. Flush the needle port with the eluent B using a truncated 1-ml syringe in systems with manual injector. This procedure removes sample residues from previous injections. 4. Flush the Hamilton glass syringe (for manual injection) with methanol, then water to rinse it. 5. Flush the sample loop (in the load position) with three times the sample loop volume with eluent B using a Hamilton glass syringe.
THE GRADIENT DELAY (DWELL VOLUME) OF THE HPLC SYSTEM The following steps must be done without a column connected to the HPLC instrumentation. 1. Connect the injector directly to the detector with union piece (zero length column). 2. Prepare a special eluent A and B, 200 ml each: Eluent A: acetonitrile Eluent B: acetonitrile/0.2% acetone. 3. Run a gradient of 10% to 90% B in 10 min at a flow rate of 2.0 ml/min. Detection is carried out at 254 nm. The measured value of the dwell volume can be influenced by the injection technique. If after the injection the valve remains in the inject position, the dwell volume will include the volume of the sample loop; if the valve is put back in load position, the dwell volume will not. This effect can produce errors with sample loops >100 µl. The same consideration is valid if the sample loop is exchanged from an analytical separation (e.g., a sample loop of 50 µl) to a semipreparative separation (e.g., a sample loop of ≥500 µl) on the same column. 4. Determine the gradient delay and present the results graphically in a format similar to that shown as Figure 8.7.1.
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D V = [6.1 min – (10/2)] × 2 ml/min = 2.2 ml
0.3
Absorbance (254 nm)
90%
0.2
50% 0.1 6.1 min
10%
0.0 0
5
10
15
Time (min) Figure 8.7.1 Graphical illustration of the approach employed to determine the gradient delay volume, Dv. In this figure the gradient profile is illustrated for a defined flow rate (2 ml/min), with the gradient profile recorded from 10% to 90% buffer B at a specified wavelength such as 254 nm, as described in the text with acetonitrile-water-acetone mixtures.
The dwell volume, VD, is the volume of eluent from the pump heads to the column inlet (including the mixing chamber volume). The dwell volume values range from 2 to 7 ml; autosamplers in particular make a large contribution to the delay volume. It should be determined with an accuracy of ±0.5 ml. The profile can be used for diagnostic purposes, since the volume accuracy of the pump delivery is also monitored. Knowledge of the gradient delay is essential for method development, since it allows the accurate calculation of the S and k0 values (Ghrist and Snyder, 1988a; Hearn, 1991a). Its determination is particularly important when establishing segmented gradients (since various errors can accumulate here), and when an established HPLC method is transferred from one instrument to another instrument.
CONNECTING THE COLUMN The following considerations are relevant to the preparation of the HPLC column for use with samples containing peptides and proteins. 1. Flush the column with eluent B with the inlet connected to the injector and the outlet facing the waste collector (5 min at 1 ml/min for analytical columns). This procedure re-
moves air that may have been trapped and replaces the storage buffer. 2. Connect the column outlet to the detector. Start the flow with 0.5 ml/min and slowly increase the flow rate to 1 ml/min. 3. Equilibrate the column first with eluent B until a stable baseline is reached or alternatively with 10 column volumes (∼15 min for analytical column at 1 ml/min) and then with eluent A again with 10 column volumes. The pressure should be monitored and documented for each eluent since it can be used for diagnostic purposes.
PROGRAMMING THE HPLC INSTRUMENT The following considerations are relevant to the programming of the HPLC instrument for use with samples containing peptides and proteins. 1. Program, according to the manufacturer’s handbook, the pump, the detector, the integration module, and the autosampler. Test the method with a test run before leaving the instrument alone. 2. Program a shutdown method for overnight runs, which will switch off the lamp and pump. This approach prolongs lamp life and saves eluents.
Conventional Chromatographic Separations
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INJECTING THE SAMPLE The following considerations are relevant to the injection of the samples containing peptides and proteins onto the HPLC column. 1. Switch the sample loop to the load position and rinse with eluent A. This procedure removes eluent B (which may be present from rinsing the loop or previous runs). Failure to do so can cause pre-elution of the peptide or protein sample, particularly in conjunction with partially filled sample loops. 2. Load the sample slowly into the loop avoiding air bubbles. Do not squirt the sample into the loop too fast as it will end up in the waste. 3. Inject the sample by switching the valve swiftly into the inject position. If the switching is done too slowly, the pumps might shut down because the pressure limit is exceeded, as the valve is blocked in the intermediate switching position.
TESTING THE FUNCTIONAL HPLC SYSTEM The following considerations are relevant to the testing of the HPLC system for use with samples containing peptides and proteins. 1. Produce a blank run (inject eluent A) and run a gradient from 100% eluent A to 100% eluent B under the same conditions as intended for the peptide or protein sample. Repeat if peaks occur. This procedure cleans the column of peptides and proteins from previous separations, which have not been removed by the flushing process. 2. Measure the dead volume of the column with thiourea or sodium nitrate (or any other noninteractive solute). 3. Test the column performance with a gradient run and an appropriate test mixture (see below for details). First, this test allows the evaluation of the column bed integrity (low integrity will be associated with split, fronting or tailing peaks) and column performance (in terms of plate numbers). Second, this test allows, if repeated at regular intervals, the monitoring of the performance during the lifetime of a column, and the assessment of batch-tobatch differences of column fillings.
LOGBOOKS
HPLC of Peptides and Proteins
Record keeping is essential for liquid chromatography system maintenance and confirmation/substantiation of the experimental results. Three types of logbooks are recommended, the system logbook, the column
logbook, and the experiment/assay logbook (Dolan and Snyder, 1989).
System Logbook This logbook should contain information on: 1. The module identification (brand, model, serial number, purchase data, warranty information) for the entire liquid chromatography (LC) system: injector, autosampler, pump(s), detector, software, column. 2. Module replacements, maintenance records and upgrades. 3. Reference chromatograms and operating parameters. 4. Maintenance: activity, time, operator, and name/reliability of service engineer. 5. Column replacement (cross-reference to column logbook).
Column Logbook This logbook should contain information including a summary of use, column life in months and number and type of samples, cause of failure, and suggestions for extending life. For each column this would be tabulated as follows. 1. Date column first used. 2. Specification: mobile phase flow rate, plate number, peak shape, dead volume over the lifetime of the column. 3. Performance of new column (validated with a test mixture). 4. Record of use (instrument, operator, number and type of samples). 5. Storage information. 6. Maintenance performed (type of backflush protocols, frit replacement, etc.). 7. Revaluation of column performance. 8. Cause of failure.
Experiment/Assay Logbook This logbook should contain information on: 1. Equipment configuration. 2. Operating condition(s). 3. Mobile phase recipe(s) (literature references if available). 4. Sample pre-treatment method(s) (literature references if available). 5. Assay procedure(s) (literature references if available). 6. Data analysis procedure(s). In the following sections, illustrative examples of standard operating protocols for the major modes of HPLC are described based on
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Sample loop size: 20 to 200 µl Isocratic elution Eluent A: 50 mM KH2PO4, pH 6.5, 0.1 M KCl Flow rate: 0.5 ml/min Detection: 214 nm Temperature: room temperature Peptide standards for column testing as described, e.g., in Mant and Hodges (1991a) Method development in the HP-SEC of peptides and proteins can be performed via the following steps: 1. Select sorbent of the most appropriate average pore size, packed into a column of suitable length. 2. Check for “ideal” and “non-ideal” retention effects. 3. Optimize plate number (adjust the flow rate or change to a column of different length). An example of HP-SEC for the separation of peptides and proteins is illustrated in Figure 8.7.2. In this example, the resolution of the 50S ribosomal proteins from Thermus aquaticus was achieved on a tandem TSK-250 column (75-mm length × 7.5-mm i.d. and 300-mm length × 7.5-mm i.d.) at a flow rate of 0.5 ml/min with a 50 mM (NH4)2SO4/20 mM NaH2PO4 buffer system, pH 5.0.
the instrumental system validation procedures described above.
STANDARD OPERATING CONDITIONS FOR HP-SEC The separation of peptides and proteins by high-performance size-exclusion chromatography (HP-SEC) is based on the concept that molecules of different sizes (hydrodynamic volume, Stoke’s radius) permeate to different extents into porous SEC separation media and thus exhibit different permeation coefficients according to differences in their molecular weights (Regnier, 1983). However, many SEC materials are slightly hydrophobic or can weakly act as ion exchangers. These properties lead to nonideal behavior (specifically electrostatic or hydrophobic interactions between the peptide or protein and the matrix). This feature is not necessarily a disadvantage, since mixedmode selectivities can be achieved, but can be suppressed by the addition of a salt at a reasonably high ionic strength, i.e., ≥100 mM, to the mobile phase (Mant et al., 1987).
Chromatographic Conditions
Column: e.g., TSK-250 (10 µm, 300 Å, 300mm length × 7.5-mm i.d.) Sample size: <2 mg peptide/protein
Absorbance (205 nm)
0.064
0.032
0 0
10
20
30
40
50
Time (min)
Figure 8.7.2 Illustrative example of the use of HP-SEC in peptide and protein separation and analysis. In this example the resolution of 50S ribosomal proteins from Thermus aquaticus in 2% acetic acid (Molnar et al., 1989b) was achieved with a tandem TSK-250 column (75-mm length × 7.5-mm i.d. and 300-mm length × 7.5-mm i.d.) using a flow rate of 0.5 ml/min and a 50 mM (NH4)2SO4, 20 mM NaH2PO4, pH 5.0, with detection at 205 nm.
Conventional Chromatographic Separations
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STANDARD OPERATING CONDITIONS FOR HP-NPC
HPLC of Peptides and Proteins
Chromatographic systems in which the stationary phase is more polar than the mobile phase were developed at the beginning of the modern era of liquid chromatography and were known under the acronym NPC (“normal phase” liquid chromatography). In contrast to RPC with immobilized n-alkyl ligands, where the interaction of solute and stationary phase is based on solvophobic phenomena, the interaction in NP-LC is based on adsorption. The retention behavior of peptides and proteins in NP-LC is often described in terms of the classical concepts of multisite displacement and site occupancy theory (Snyder, 1970). Today, NP-HPLC is mainly used for the separation of polyaromatic hydrocarbons (PAHs), heteroaromatic compounds, nucleotides, nucleosides, etc., and much less frequently for protected synthetic peptides, deprotected small peptides in the “flash chromatographic mode,” and protected amino acid derivatives used in peptide synthesis (Ballschmiter and Wossner, 1998). Originally, normal phase chromatography was limited to unmodified silica columns. Recent work has, however, utilized polar bonded phases such as amino (-NH2), cyano (-CN), or diol (-COHCOH-) coated sorbents. Chromatography on such modified normal phase packing materials is also known as polar bonded phase chromatography (PBPC), which is used for the separation of peptides (Yoshida, 1998) and proteins (Buchholz et al., 1982). Today, one of the main applications of modified normal phase silica materials is in HPLC-integrated solid phase extraction procedures (SPE; Papadoyannis et al., 1995). These types of sorbents, particularly when used as precolumn packing materials in LC-LC column switching settings in conjunction with restricted access sorbents materials (RAM), allow multiple injections of untreated complex biological samples such as hemolyzed blood, plasma serum, fermentation broth, cell tissue homogenates, etc., for the isolation of bioactive peptides. Typically, with RAM materials, hydrophilic, electroneutral diol groups are immobilized onto the outer surface of spherical particles. This layer prevents nonspecific interactions between the support matrix and protein(s) or other high-molecular-weight biomolecules, which are thus excluded from the interior regions of the particle and elute as nonretained components. The inner surfaces of the porous RAM particles are, however, chemically modified with n-alkyl ligands, which are only freely
accessible for low molecular analytes, such as peptides. As a consequence, significant enrichment or partial resolution of peptide analytes can be achieved.
Chromatographic Conditions Column: e.g., diol-phase, aminopropyl-phase, cyano-phase column, 250-mm length × 4.6mm i.d. Sample size: <2 mg peptide/protein Sample loop size: 20 to 200 µl Linear A→B gradient Eluent A: 0.1% TFA in water Eluent B: 0.1% TFA in 20% acetonitrile/80% water (v/v) Gradient range and time: 0% to 100% eluent B in 60 min Flow rate: 1 ml/min Detection: 214 nm Temperature: room temperature An example of the use of HP-NPC for the separation of peptides and proteins is illustrated in Figure 8.7.3. In this example, the resolution of ferritin, bovine serum albumin, chymotrypsinogen, cytochrome c, and alanine was achieved on a LiChrosorb Diol column (250mm length × 16-mm i.d.) at flow rate of 2.5 ml/min and a 0.1 M phosphate buffer system, pH 5.0 (Buchholz et al., 1982).
STANDARD OPERATING CONDITIONS FOR HP-HIC As in reversed-phase chromatography, the hydrophobic interaction between the peptide or protein and the nonpolar ligands immobilized onto the surface of the sorbent represents the dominant effect in hydrophobic interaction chromatography (Fausnaugh et al., 1984; Fausnaugh and Regnier, 1986; Gooding et al., 1986; Wu et al., 1986a,b; Melander et al., 1989; Antia et al., 1995; Hearn, 2000b). In both techniques, peptides and proteins are eluted by lowering the surface tension of the mobile phase. However, in HP-HIC, this is achieved with a decreasing salt concentration, i.e., by increasing the water content of the eluent, in contrast to RP-HPLC where the decrease in surface tension of the eluent is achieved through an increase in the organic solvent content of the mobile phase. As such, the minimum surface tension reached in the HP-HIC of polypeptides or proteins with binary water-salt systems corresponds to the surface tension of pure water, i.e., 78 dynes/cm. These differences between RP-HPLC and HPHIC have fundamental effects on the recovery of proteins in bioactive state, as well as on the selectivity of the system. Typically, kosmot-
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ferritin 364,000
0.8
Absorbance (220 nm)
bovine serum albumin 67,000
0.6
chymotrypsinogen 25,000 cytochrome c 12,500
0.4
0.2
alanine 121
0.0 0
10
20
30
40
50
Time (min)
Figure 8.7.3 Illustrative example of the use of HP-NPC in peptide and protein separation and analysis. In this example the resolution of ferritin (Mr, 364,600), bovine serum albumin (Mr, 67,000), chymotrypsinogen (Mr, 25,000), cytochrome c (Mr, 12,500), and alanine (Mr, 121; Buchholz et al., 1982) was achieved on a LiChrospher Diol column (250-mm length × 16-mm i.d.) at a flow rate of 2.5 ml/min and a 0.1 M phosphate buffer system, pH 5.0. In this specific case, size exclusion effects dominated the global resolution, with polar phase effects contributing to the individual selectivities of these proteins, as assessed from the “nonideality” of the log Mr versus Ve plots.
ropic (anti-chaotropic) salts, i.e., ammonium sulfate, sodium sulfate, or magnesium chloride of high molal surface tension increment, are to be preferred for HP-HIC applications with polypeptides and proteins. In HP-HIC, non-polar ligands with lower hydrophobicity and lower ligand density (∼1⁄10th that of RP-HPLC sorbents) are employed. HP-HIC sorbents should be selected on the basis of the critical hydrophobicity concept (Hearn, 2000a,b). In combination with nondenaturing mobile phases, proteins, in particular, can potentially be eluted in their native conformation from HP-HIC sorbents.
Chromatographic Conditions Column: e.g., TSK Phenyl column, 75-mm length × 7.5-mm i.d., 10 µm Sample size: <2 mg peptide/protein Sample loop size: 20 to 200 µl Linear A→B gradient is the preferred method with “hold” options
Eluent A: 0.1 M NaH2PO4, 2.0 M (NH4)2SO4, pH 7.0 Eluent B: 0.1 M NaH2PO4, pH 7.0 Gradient rate: 5% eluent B/min Gradient range and time: 0 to 100% B in 20 min Flow rate: 1 ml/min Detection: 214 nm Temperature: room temperature The method development in HP-HIC can be performed in the following steps: 1. Select type of salt (antichaotropic) and concentration range, taking into account the concentration of the salt that is required to reach saturation, and then keep the maximum concentration below this value by ∼20%. 2. Optimize the gradient conditions (gradient run time, starting and final mobile phase composition). 3. Optimize the band spacing (pH, organic solvent). 4. Optimize the column conditions (flow, column length).
Conventional Chromatographic Separations
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3
1
6 4
Absorbance (280 nm)
8
2 7 5
0
15
30
Time (min)
Figure 8.7.4 Illustrative example of the use of HP-HIC in peptide and protein separation and analysis. In this example the resolution of: 1, cytochrome c; 2, ribonuclease; 3, lysozyme; 4, bovine serum albumin; 5, ovalbumin; 6, α-chymotrypsin; 7, α-chymotrypsinogen; and 8, myoglobin was achieved (Kato et al., 1983) on a Butyl-G3000 SW column (150-mm length × 6-mm i.d., 10 µm) with a 60-min linear gradient from 0% to 100% eluent B (eluent A: 1.5 M ammonium sulfate, 0.1 M phosphate buffer, pH 6.0; eluent B: 0.1 M phosphate buffer, pH 6.0).
Examples of the use of HP-HIC with proteins are described in Fausnaugh et al. (1984); Fausnaugh and Regnier (1986); Gooding et al. (1986); Melander et al. (1989); Wu et al. (1986a,b); Antia et al. (1995); and Hearn (2000a,b). Figure 8.7.4 illustrates the resolution of cytochrome c (1), ribonuclease (2), lysozyme (3), bovine serum albumin (4), ovalbumin (5), α-chymotrypsin (7), and myoglobin (8) on a Butyl-G3000 SW column (150-mm length × 6-mm i.d., 10 µm) with a 60-min linear gradient from 0% to 100% B using eluent A (1.5 M ammonium sulfate, 0.1 M phosphate buffer, pH 6.0) and eluent B (0.1 M phosphate buffer, pH 6.0).
STANDARD OPERATING CONDITIONS FOR HP-IEX
HPLC of Peptides and Proteins
Peptides and proteins can be eluted in ion exchange chromatography by either isocratic or gradient elution (Chang et al., 1976; Kopaciewicz and Regnier, 1983a, 1986; Reg-
nier, 1984; Kopaciewicz et al., 1985; Hearn et al., 1988; Heinitz et al., 1988; Hodder et al., 1990). Gradient elution is usually performed with a linear A→B gradient of a salt such as sodium or potassium chloride in phosphate buffer. The retention of peptides and proteins on ion-exchange sorbents arises from electrostatic interactions between the ionized surface of the solute and the charged surface of the sorbent. For peptide and protein separations, the use of a strong cation exchanger has a considerable advantage over other varieties of ion-exchanger (Mant and Hodges, 1985), since the column can retain its negatively charged character over the whole range from acidic to neutral pH. Both weak and strong cation exchangers, e.g., based on carboxymethyl or sulfonopropyl ligands, as well as weak and strong anion exchangers, e.g., dimethylamino or quaternary ammonium ligands, are available and can be applied to the HP-IEX of peptides and proteins.
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With peptides and proteins, at neutral pH, the side-chain carboxyl groups of the acidic amino acid residues—glutamic acid and aspartic acid—are completely ionized. Below pH 3.0 they are almost completely protonated. A change of pH therefore allows the retention of peptides and proteins to be varied according to the net charge of these biosolute(s). Even when the most dominant effect in IEX is electrostatic, in many cases the participation of a mixed mode separation, whereby hydrophobic interaction contributes, cannot be excluded (Burke et al., 1989). This is not necessarily a disadvantage, since it permits selectivity modulation with complex mixtures of peptides and proteins. Hydrophobic interactions can be suppressed by adding a nonpolar solvent, such as acetonitrile, or a nonionic detergent, such as Brij-25, to the mobile phase.
Chromatographic Conditions Column: e.g., strong cation exchanger with sulfonate functionality (5 µm, 300 Å, 75-mm length × 7.5-mm i.d.) Sample size: <2 mg peptide/protein Sample loop size: 20 to 200 µl Linear A→B gradient Eluent A: 5 mM KH2PO4, pH between 3.0 and 7.0 Eluent B: 5 mM KH2PO4/0.5 M KCl, pH between 3.0 and 7.0 Gradient rate: 3.3% eluent B/min
1.0
Gradient range and time: 0% to 100% eluent B in 30 min Flow rate: 1 ml/min Detection: 214 nm Temperature: room temperature Peptide standards for column testing as described, e.g., in Mant and Hodges (1991a) The method development in HP-IEX can be performed in the following steps: 1. Select type of anion or cation exchanger. 2. Optimize gradient conditions (gradient run time, starting and final mobile phase composition). 3. Optimize band spacing (pH, salt type). 4. Optimize column conditions (flow, column length). Examples of the use of HP-IEX with proteins are described in Chang et al. (1976); Regnier (1984); Kopaciewicz et al. (1985); Kopaciewicz and Regnier (1986); Hearn et al. (1988); Heinitz et al. (1988); Burke et al. (1989); Hodder et al. (1990); Mant and Hodges (1985, 1991b); and Hearn (2000a). Figure 8.7.5 illustrates the use of HP-IEC for the resolution under gradient elution conditions from 0% to 100% B (eluent A: 0.1 M Tris⋅Cl, pH 7.5: Eluent B: 0.1 M Tris⋅Cl/0.2 M NaCl, pH 7.5) of ovalbumin isoforms on a TSK-GEL IEX-545 DEAE SIL column (150-mm length × 6-mm i.d.) using a 90-min linear gradient at a flow rate of 1.0 ml/min.
fraction 1
Absorbance (280 nm)
a
fraction 2
b
0.0 0
20
40 Time (min)
60
Figure 8.7.5 Illustrative example of the use of HP-IEX in peptide and protein separation and analysis. In this example the resolution of ovalbumin isoforms (Kato et al., 1982), FrIa and FrIIb, was achieved on a TSK-GEL IEX-545 DEAE SIL column (150-mm length x 6-mm i.d.) using a 90 min linear gradient at a flow rate of 1.0 ml/min from 0-100% B (eluent A: 0.1 M Tris⋅Cl, pH 7.5: eluent B: to 0.1 M Tris⋅Cl, 0.2 M NaCl, pH 7.5).
Conventional Chromatographic Separations
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STANDARD OPERATING CONDITIONS FOR HP-HILIC Peptides can be separated on strong hydrophilic materials, i.e., poly(2-hydroxyethylaspartamide) silica (Alpert, 1990), whereby their selectivity changes with the concentration of the organic modifier. At high organic modifier concentrations (i.e., >55% n-propanol), the solute is retained on the column. The solute is subsequently eluted with a decreasing gradient of organic modifier, whereby with low organic modifier concentrations the separation of the solutes is governed by molecular sieving effects. HP-HILIC allows specifically the evaluation of highly polar compounds, which cannot be retained on traditional reversed-phase stationary phases. To date a variety of HP-HILIC sorbent packings, including amide-, poly(2-hydroxyethyl)-aspartamide-, cyclodextrin-, or teicoplanin-derivatized stationary phases are available (Strege, 1998; Risley and Strege, 2000). HP-HILIC can be applied in a variety of challenging separation tasks, e.g., the separation of glycopeptides (Zhang and Wang, 1998) or the desalting of electroeluted proteins from SDS-PAGE systems (Jeno et al., 1993). In this mixed-mode hydrophilic interaction/cation exchange chromatographic mode (Mant et al., 1998a,b; Litowski et al., 1999), peptides are usually subjected to linear, decreasing salt gradients in the presence of high levels of organic modifier. The following example is representative of the conditions that can be employed in the HP-HILIC of peptides and small proteins.
Chromatographic Conditions
HPLC of Peptides and Proteins
Column: e.g., HILIC PolySulfoethyl A sorbent with a poly(2-sulfonethylaspartimide) functionality (5 µm, 300 Å, 200-mm length × 4.6mm i.d.) Sample size: <2 mg peptide/protein Sample loop size: 20 to 200 µl Linear A→B gradient Eluent A: 20 mM triethylammonium phosphate/80% acetonitrile, pH 3.0 Eluent B: eluent A with 400 mM NaClO4, pH 3.0. Gradient rate: 2.5% B/min Gradient range and time: 0% to 100% B in 90 min Flow rate: 1 ml/min Detection: 214 nm Temperature: 30°C Method development in the HP-HILIC of peptides and proteins can be performed via the following steps:
1. Select sorbent of the most appropriate average pore size, packed into a column of suitable length. 2. Check for “ideal” and “non-ideal” retention effects. 3. Optimize selectivity through the changes in the composition or concentration of the organic solvent and kosmotropic/chaotropic salt additives. 4. Optimize plate number (by adjusting the flow rate, or changing to a column of different length). Examples of the use of HP-HILIC with peptide standards for column testing are described in Mant and Hodges (1991b). In Figure 8.7.6 is shown the separation of the tryptic glycopeptides of Asn97 for recombinant human interferon γ (Zhang and Wang, 1998) on a polyhydroxyethylaspartamide column (150mm length × 1-mm i.d.) using a flow rate of 50 µl/min. In this example, a 60-min gradient from 100% A (85% acetonitrile, 15% water, 10 mM triethylamine) to 100% B (20% acetonitrile, 15% water, 10 mM triethylamine, 25 mM sodium perchlorate), pH 6.0 was employed.
STANDARD OPERATING CONDITIONS FOR HP-IMAC Immobilized metal-chelate affinity chromatography (IMAC) exploits the affinities of the side chain moieties of specific surface accessible amino acids in peptides and proteins for the coordination sites of immobilized transition metal ions (Porath et al., 1975; Zachariou et al., 1993). The majority of investigations employed tri- or tetradentate ligands, such as iminodiacetic acid (IDA), nitrilotriacetic acid (NTA), tris-(carboxymethyl)ethylenediamine (TED), O-phosphoserine (OPS) or carboxy-methylaspartic acid (CMA) (Zachariou et al., 1993). Novel immobilized chelate systems, such as 2,6-diaminomethylpyridine or cis and trans carboxymethylproline, have shown binding properties different from the IMAC behavior of other chelating ligands (Hearn et al., 1997; Chaouk and Hearn, 1999). These techniques, in conjunction with soft gel matrices, have been predominantly applied for the preparative purification of globular proteins. Procedures to immobilize the IMAC ligand at the surface of silica supports, based on classical ligand exchange principles, have permitted useful guidelines to be developed for the design of HPIMAC applications for peptides and proteins at the analytical level (Wirth and Hearn, 1993). However, for the majority of the possible HPIMAC applications with peptides or proteins
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0.06 6 4
Absorbance (220 nm)
0.04
0.02
0 1
2
5 8
7
3
–0.02
25
30
35
40
Time (min)
Figure 8.7.6 Illustrative example of the use of HP-HILIC in peptide and protein separation and analysis. In this example the resolution of the tryptic Asn97 glycopeptides (1-8) of recombinant human interferon γ (Zhang and Wang, 1998) was achieved on a polyhydroxyethyl aspartamide column (150-mm length × 1-mm i.d.) using a flow rate of 50 µl/min and a 60-min gradient from 100% A (85% acetonitrile, 15% water, 10 mM triethylamine) to 100% B (20% acetonitrile, 15% water, 10 mM triethylamine, 25 mM sodium perchlorate), pH 6.
no standard chromatographic conditions exist. Rather, different elution regimes have tended to be employed with HP-IMAC systems, based on the selection of either empirical step- or gradient-elution protocols involving changes in pH, buffer composition, or the concentration of salts or another competitive binding reagent such as imidazole or malonic acid. Various applications of different IMAC systems for the separation of peptides and proteins have recently been reviewed elsewhere (Porath et al., 1975; Jeno et al., 1993; Mant et al., 1998a,b; Litowski et al., 1999; Hearn, 2000a). The following example is representative of the conditions that can be employed in the HP-IMAC of peptides and small globular proteins.
Chromatographic Conditions Column: e.g., HP-IMAC sorbent with a Cu2+ ion chelated to immobilized iminodiacetic acid functionality, i.e., IDA-Cu(II) TSK gel chelate5PW column (5 µm, 300 Å, 100-mm length × 4.6-mm i.d. or 75-mm length × 8-mm i.d.) Sample size: <1 mg peptide/protein Sample loop size: 20 to 200 µl
Linear A→B gradient Eluent A: 50 mM acetic acid/50 mM MES/50 mM HEPES/80 mM Na2SO4/2 × 10−6 M Cu2+, pH 8.0; or 1 mM imidazole/20 mM sodium phosphate buffer containing 0.5 M NaCl, pH 7.0 Eluent B: 50 mM acetic acid/50 mM MES/50 mM HEPES/50 mM ammonium acetate/2 × 10−6 M Cu2+, pH 5.5; or 20 mM sodium phosphate buffer containing 0.5 M NaCl, pH 7.0 with 20 mM imidazole gradient Gradient rate: 5% B/min Gradient range and time: 0% to 100% eluent B in 20 min Flow rate: 1 ml/min Detection: 280 nm Temperature: 25°C Method development in the HP-IMAC of peptides and proteins can be performed via the following steps: 1. Select sorbent of the most appropriate average pore-size, packed into a column of suitable length. 2. Check for “ideal” and “non-ideal” retention effects.
Conventional Chromatographic Separations
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Absorbance (280 nm)
0.06 38+39 20 0.04
30 35 29
37
0.02
Concentration of imidazole (mM)
24
0 0
20
40 Time (min)
60
Figure 8.7.7 Illustrative example of the use of HP-IMAC in peptide and protein separation and analysis. In this example the resolution of human gastrin-I (24), human GIP(21-42) (29), Trp(for)human GIP(21-42) (30), human big gastrin (35), human GIP (37), porcine GIP (38), and bovine GIP(39) (Yip et al., 1989) was achieved on an IDA-Cu(II) TSK gel chelate-5PW column (75-mm length × 8-mm i.d.) with a flow rate of 1 ml/min equilibrated with a 1 mM imidazole, 20 mM sodium phosphate buffer containing 0.5 m NaCl, pH 7.0, and eluted with an 1-20 mM imidiazole gradient (dotted line).
HPLC of Peptides and Proteins
3. Optimize selectivity through the changes in the type of chelating ligand and metal ion, i.e., whether borderline, hard or soft metal ion, the pH, the type and concentration of the buffer used at the loading and washing stages, and finally at the elution stage. 4. Optimize plate number (by adjusting the flow rate or changing to a column of different length). Examples of the use of HP-IMAC with proteins are described in Porath et al. (1975); Wirth and Hearn (1993); Zachariou et al. (1993); Jiang et al. (1998); Porath (1988); and Hearn (2000a). Figure 8.7.7 illustrates the use of HPIMAC for the resolution of human gastrin (24), human GIP (29), Trp(for)- human GIP (30), human big gastrin (35), porcine GIP (38), and bovine GIP on an IDA-Cu(II) TSK gel chelate5PW column (75-mm length × 8-mm i.d.) at a flow rate of 1 ml/min equilibrated with a 1 mM imidazole/20 mM sodium phosphate buffer
containing 0.5 M NaCl, pH 7.0 and eluted with a 1 to 20 mM imidiazole gradient (dotted line).
STANDARD OPERATING CONDITIONS FOR HP-BAC During the last 25 years, biospecific affinity chromatography (BAC) had evolved as a well established method for analytical separation (Porath, 1988), as well as the preparative scale purification of biopolymers of complex biological origin. Additionally, HP-BAC is a useful technique to investigate peptide-protein and protein-protein interactions. In HP-BAC, the separation is based on the propensity for biorecognition between the protein or peptide and its naturally occurring ligand (or suitable mimetic), in particular, on its relative binding affinity for a specific ligand. The ligand should have a high specificity for the protein or peptide, should bind to the protein or peptide reversibly with rapid on-and-off kinetics, and
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should be chemically stable under elution conditions. The introduction of HP-BAC dramatically increased the speed of affinity chromatographic separation of proteins, including sample loading and column regeneration (Ohlson et al., 1978). However, since the specificity and the selectivity of the HP-BAC both are very high, the range of applications is, paradoxically, much more limited than the other modes of HPLC. Every separation ideally demands its own ligand type, ligand density, column configuration, and particle type in order to enable the optimal purification of a particular component. Consequently, there are no “standard” HP-BAC conditions due to the broad range of ligand types. Some ligands are specific for a particular class of protein; however, other ligands can be multifunctional and bind to a number of related molecules. Currently, commercially available HP-BAC columns are supplied as activated supports (epoxy-, N-hydroxysuccinimido-, tresyl-, hydroxylpropyl-) ready for coupling appropriate ligands (via accessible SH, OH, or NH2 groups) or alternatively as preformed matrices with commonly
used ligands already attached (e.g., protein A). The following example is representative of the conditions that can be employed in the HPBAC with peptides and small globular proteins.
Chromatographic Conditions Column: custom designed, i.e., glycidoxypropyl-activated silica such as epoxy-activated LiChrosorb containing a suitable immobilized ligand functionality (10 µm, 10000 Å, 100-mm length × 4.6-mm i.d.) Sample size: <2 mg peptide/protein Sample loop size: 20 to 200 µl Linear gradient or step elution Eluent for equilibration: nondenaturing conditions with adequate buffer capacity Eluent for desorption: non-denaturing conditions of different pH to the equilibration conditions, with adequate buffer capacity and appropriate competitive species to affect efficient dissociation of the affinant-affinate cognate interaction Gradient rate: according to affinity of interaction, HP-BAC sorbent characteristics, and flow rate
Enzyme activity (dotted lines)
0.5 M NaCl
Absorbance at 276 nm (solid lines)
0.5
0.0
0
2
4 Time (min)
6
Figure 8.7.8 Illustrative example of the use of HP-BAC in peptide and protein separation and analysis. In this example, the enzyme activity is shown for rabbit muscle lactate dehydrogenase (LDH) (Anspach et al., 1988) separated on a Procion Red MX5B immobilized non-porous silica column (40-mm length × 6-mm i.d.) using a flow rate of 1 ml/min and step elution (eluent A: 10 mM phosphate buffer, pH 8.0: eluent B: 0.5 M sodium chloride in 10 mM phosphate buffer, pH 8.0).
Conventional Chromatographic Separations
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Flow rate: 0.5 to 1 ml/min Detection: 214 to 280 nm Temperature: 4°C In Figure 8.7.8 an example is shown of the use of HP-BAC for the resolution of rabbit muscle lactate dehydrogenase (LDH) on a Procion Red MX5B dye immobilized onto nonporous silica column (40-mm length × 6-mm i.d.) using a flow rate of 1 ml/min and step elution with eluent A: 10 mM phosphate buffer, pH 8.0, and eluent B: 0.5 M sodium chloride in 10 mM phosphate buffer, pH 8.0.
STANDARD OPERATING CONDITIONS FOR RP-HPLC As noted above, RP-HPLC procedures currently represent the majority of applications for peptide analysis and purification, and over 80% of all analytical studies with proteins. The dominant effect in reversed-phase chromatography is a hydrophobic interaction between the nonpolar amino acid residues of peptides or proteins and the nonpolar ligands, typically immobilized onto the surface of a spherical, porous silica particle (Hearn, 2001), although nonpolar polymeric sorbents derived, e.g., from cross-linked polystyrene-divinylbenzene, can also be employed. In this technique, isocratic elution, step elution, or gradient elution modes can be utilized to purify peptides and proteins. Besides the requirement for an organic solvent to be used as a surface tension modifier, ion-pair reagents (Mant and Hodges, 1991c) are utilized at low pH (e.g., pH 2.1) to suppress silanophilic interactions between free silanol groups on the silica surface and basic amino acid residues. Silica-based packing materials of 3 to 10 µm average particle diameter and ≥300 Å pore size, with n-butyl, n-octyl, or n-octadecyl ligands, are widely used.
Chromatographic Conditions Chemicals HPLC-grade acetonitrile, 2-propanol, methanol, or other suitable organic solvents with UV transparency down to 210 nm Trifluoroacetic acid (TFA) NaH2PO4 or other suitable salts H3PO4 Milli-Q water or equivalent TFA employed for protein sequencing may not be suitable because it can contain antioxidants.
Preparing the mobile phase 1. Prepare eluent A and B, e.g.: eluent A: 0.1% TFA in water and eluent B: 0.09% TFA in 60% acetonitrile/40% water (v/v). The volumes of organic solvent and water are measured in two different cylinders and then combined, because of the volume contraction upon mixing, which may be up to 30 ml per liter of prepared solvent (depending on the nature of the solvent). Failure to do so can lead to substantial errors in mobile phase composition. To compensate for the baseline shift in gradient elution (because organic components absorb more light at low wavelengths) when working with water/organic eluents, the amount of ion pair reagent (TFA, H3PO4) in eluent B is usually decreased by 10% to 15% in comparison with eluent A, yielding a flat baseline (Dolan, 1991). It is imperative that eluents be prepared in the fume hood. TFA is extremely corrosive; laboratory coat, gloves, and protective glasses must be worn. 2. Mix the solvent either by stirring with a Teflon-coated magnetic flea or by shaking a stoppered cylinder. Parafilm should not be used, under any circumstances, to cover the eluents, since the organic solvents in combination with acidic ionpair reagent components in the eluents will dissolve components in the Parafilm, yielding extra peaks in the chromatogram. Testing the RP-HPLC stationary phase The following considerations are relevant to the evaluation of HPLC stationary phases for use with samples containing peptides and proteins. 1. Test the column performance with a gradient run and a test mixture: RP-HPLC test mixture example: 0.15% (w/v) dimethylphthalate, 0.15% (w/v) diethylphthalate, 0.01% (w/v) diphenyl, 0.03% (w/v) O-terphenyl, 0.32% (w/v) dioctylphthalate in methanol. 2. Peptide standards for column testing are described, for example, in Mant and Hodges (1991b). Peptide purification by RP-HPLC In addition to the reagents and procedures described in the preceding section the following additional materials are required: Chemicals: acetonitrile, trifluoroacetic acid (TFA)
HPLC of Peptides and Proteins
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A
0.30
t R = 25.7 min
Absorbance (214 nm)
0.15
0.00
B
0.60
t R = 25.9 min
0.30
0.00 0
10
20
30 40 Time (min)
50
60
Figure 8.7.9 Illustrative example of the use of RP-HPLC in peptide separation and analysis. R e so lu ti on of N-acetyl lipocor tin-1 [2-26] peptide product [Ac-AMVSEFLKQAWFIENEEQEYVQTVK-CONH2] obtained by solid phase peptide synthesis on a TSK-ODS-120 T column (150-mm length x 4.6-mm i.d., 120 Å, 5 µm, end-capped) using a flow rate of 1 ml/min and a 60 min gradient from 100% A (0.1% TFA in water) to 100% B (90% acetonitrile 0.09% TFA), pH 2.1. (A) Crude peptide; (B) purified peptide.
Columns: analytical C-18 or other appropriate n-alkylsilica sorbent material (5 µm, 300 Å, 150-mm length × 4.6-mm i.d.) Preparative C-18 or other appropriate n-alkylsilica sorbent material (10 µm, 300 Å, 300-mm length × 21.5-mm i.d.) Apparatus: in addition to an analytical HPLC instrument with autosampler: preparative HPLC pump(s), manual injector, detector with preparative flow cell, and fraction collector are required Purification of peptides derived from SolidPhase Peptide Synthesis (SPPS) can be performed in the following steps. 1. Separation of the crude peptide (∼100 µg) with an analytical RP-HPLC procedure allows the assessment of the sample in terms of purity, peak profile and elution conditions. Appropriate equipment/conditions are as follows: Column: e.g., C-4, C-8, C-18, etc. (5 µm, 300 Å, 150-mm length × 4.6-mm i.d.) Sample size: <2 mg peptide/protein Sample loop size: 20 to 200 µl Linear A→B gradient Eluent A: 0.9% aqueous TFA Eluent B: 0.1% TFA in acetonitrile/water
Gradient rate: 1% B/min For example, for 60% ACN/water: gradient range and time: 0% to 100% eluent B in 60 min Flow rate: 1.0 ml/min Detection: 214 nm Temperature: room temperature An example of the analytical use of RPHPLC in peptide analysis is shown in Figure 8.7.9. In this case, resolution of the crude Nacetyl lipocortin-1[2-26] product from solid phase peptide synthesis (panel A), and the purified product (panel B) was achieved with a TSK-ODS-120 T column (150-mm length × 4.6-mm i.d., 120 Å, 5 µm, end-capped) at a flow rate of 1 ml/min and a 60 min gradient from 100% A (0.1% TFA in water) to 100% B (90% acetonitrile 0.09% TFA), pH 2.1. 2. Separation of the crude peptide (∼25 to 100 mg) with preparative RP-HPLC procedure. Appropriate equipment/conditions are as follows: Column: e.g., C-4, C-8, C-18, etc. (10 µm, 300 Å, 300-mm length × 21.5-mm i.d.) Sample size: <150 mg peptide/protein Sample loop size: 1 ml, multiple injection Linear A→B gradient
Conventional Chromatographic Separations
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0.15
Absorbance (254 nm)
t R = 103.9 min
0.10
0.05
0.00
0
20
40
60
80
100
120
Time (min)
Figure 8.7.10 Illustrative example of the use of RP-HPLC in the preparative isolation of synthetic polypeptides. Here, the isolation of the peptide, N-acetyl lipocortin-1[2-26]—Ac-AMVSEFLKQAWFIENEEQEYVQTVK-CONH2—(70 mg) from the crude mixture obtained from solid phase peptide synthesis using a TSK-ODS-120 T column (300-mm length × 21.5-mm i.d., 120 Å, 10 µm, end-capped) using a flow rate of 7.5 ml/min and a 135-min gradient from 25% B to 100% B (A: 0.1% TFA in water, B: 60% acetonitrile 0.09% TFA), pH 2.1.
Eluent A: 0.9% aqueous TFA Eluent B: 0.1% TFA in acetonitrile/water Gradient rate: 0.66% eluent B/min For example for 60% ACN/water: gradient range and time: 0% to 100% eluent B in 90 min Flow rate: 7.5 ml/min Detection: 254 nm Temperature: room temperature The preparative use of RP-HPLC in peptide purification is shown in Figure 8.7.10. In this example, the isolation of the synthetic N-acetyl lipocortin-1[2-26] peptide product obtained from solid-phase peptide synthesis was achieved on a TSK-ODS-120 T column (300mm length × 21.5-mm i.d., 120 Å, 10 µm, end-capped) using a flow rate of 7.5 ml/min and a 135 min gradient from 25% B to 100% B (A: 0.1% TFA in water, B: 60% acetonitrile 0.09% TFA), pH 2.1. In order to avoid detector response overloading effects, usually wavelengths from 230 to
280 nm are chosen. However, small amounts of chemical scavengers (used during the SPPS procedures) present in the crude peptide solution can absorb very strongly in this wavelength range. It may be worthwhile to sacrifice up to 1 mg of sample and to perform a preparative separation with detection at 214 nm in order to unambiguously determine the retention time of the main peptide product. 3. Collection of HPLC-fractions (3 to 7.5 ml). 4. Analysis of aliquots (30 to 50 µl) of the collected fraction with an analytical RP-HPLC (see step 4). A blank, the crude peptide solution, the fraction of interest and the 2 fractions before and 2 fractions after are typically analyzed. 5. Freeze drying of selected fraction. 6. Carry out off-line or on-line the ES-MS of purest fraction.
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S Absorbance (230 nm)
S
P
P
A 0
A 4
8
12
16
20
Time (min)
Figure 8.7.11 Illustrative example of the use of RP-HPLC in the desalting of peptide and protein samples isolated by other techniques. In this example, the 50S ribosomal protein from Thermus aquaticus on a C-4 column (40-mm length × 4-mm i.d., 300 Å, 5 µm) were desalted at a flow rate of 2.5 ml/min using a step elution protocol with 3 min elution with eluent A (0.1% TFA in water) and 3 min elution with eluent B (0.1% TFA in 2-propanol), pH 2.1. Abbreviations: S, salt; P, protein; A, time of injection.
DESALTING OF PEPTIDE AND PROTEIN MIXTURES BY RP-HPLC TECHNIQUES RP-HPLC can be utilized to desalt peptide or protein samples derived from extraction procedures or from previous HP-HIC, HP-IEC, HP-IMAC, HP-HILIC, or HP-BAC separations. Peptide or protein solutions are injected onto a small RP-HPLC column. An aqueous buffer is used to elute the salts, while the peptides or proteins are concentrated on the top of the column. After elution of the salts, monitored by UV detection, the peptides or proteins are eluted with water-acetonitrile or water 2propanol mobile phases. The loading capacity of an analytical column (100- to 300-mm length × 4-mm i.d.) is typically ∼8 mg, while the loading capacity for a semi-preparative column (30-mm length × 16-mm i.d.) is ∼34 mg (Pohl and Kamp, 1987). In addition to the reagents and procedures described in the preceding sections the following materials, reagents and conditions are required:
Chromatographic Conditions Chemicals: acetonitrile, 2-propanol, trifluoroacetic acid (TFA) Column: e.g., C-4, C-8, C-18, etc. (10 µm, 300 Å, 300-mm length × 21.5-mm i.d.) Sample size: 8 mg peptide or protein sample Sample loop size: 1 ml Step elution Eluent A: 0.1% aqueous TFA Eluent B: 0.1% TFA in acetonitrile or 2propanol Elution conditions: 100% eluent A for 3 min, then 100% eluent B for 3 min Flow rate: 2.5 ml/min Detection: 230 nm Temperature: room temperature Illustrative of the use of RP-HPLC in the desalting of peptide and protein samples isolated by other techniques is the example shown in Figure 8.7.11 for the step elution of a 50S ribosomal protein sample derived from Thermus aquaticus preparation on a C-4 column (40-mm length × 4-mm i.d., 300 Å, 5 µm) using a flow rate of 2.5 ml/min. Conventional Chromatographic Separations
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D
V3
RPC
V2
V1
SEC
P1,2
P3
Figure 8.7.12 Diagram of column switching unit employed in multi-dimensional HPLC. Abbreviations: D, detector; P1,2, two-pump module for gradient elution mode; P3, pump equilibrates offline SEC column, while RPC gradient is performed; V1, injection valve; V2 and V3, switching valves. The black field in the switching valve position symbolizes the joined valve positions. In the depicted position, the SEC and the RPC column are connected.
Table 8.7.5 Valve Positions and Pump Activities During the Four Steps in LC-LC Coupling
HPLC of Peptides and Proteins
Step
1
2
3
4
Event
HP-SEC
Start LC-LC
End LC-LC
RP-HPLC
Valve 2: HP-SEC Valve 3: RP-HPLC Mode Pump 1 Pump 2 Mode Pump 3
On Off Isocratic On Off — Off
On On Isocratic On Off — Off
On Off Isocratic On Off — Off
Off On Gradient On On Isocratic On
Figure 8.7.13 (At right) Illustrative example of the use of multimodal HPLC techniques for the separation and analysis of peptide and protein samples. (A) RP-HPLC resolution of the 50S ribosomal proteins from Thermus aquaticus in 2% acetic acid (Molnar et al., 1989b) on a C8 column (250-mm length × 4.6-mm i.d.) using a flow rate of 1.5 ml/min and a 30 min gradient from 10% to 100% eluent B [eluent A: 125 mM NaH2PO4, 125 mM H3PO4; eluent B: acetonitrile: 250 mM NaH2PO4, 250 mM H3PO4 (50:50), buffer system, pH 2.1]. (B) HP-SEC separation of the 50S ribosomal proteins from Thermus aquaticus in 2% acetic acid on a tandem TSK-250 column (75-mm length × 7.5-mm i.d. and 300-mm length × 7.5-mm i.d.) using a flow rate of 1.5 ml/min with a isocratic separation with 10% B (eluents as described above) with a fraction cut out and directed on to the reversed-phase column. (C) RP-HPLC separation of the fraction of the 50S ribosomal proteins from Thermus aquaticus in 2% acetic acid obtained on-line from the size-exclusion chromatography on a C8 column (250-mm length × 4.6-mm i.d.) using a flow rate of 1.5 ml/min and a 30 min gradient from 10% to 100% B (eluents as described above). Proteins 6, 9, and 10, which were coeluted with other proteins in chromatogram A, could now be obtained in relatively pure form for further investigations.
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A Absorbance (205 nm)
15,16 18 9,10 13,14 17 19 6,7
20 11 8 4
2
1
3
0
12
5
10
20
30
Time (min)
Absorbance (205 nm)
B
0
5
10
15
Time (min)
Absorbance (205 nm)
C
17 6 9
0
10
20
30
Time (min) Figure 8.7.13 (See legend on facing page.)
Conventional Chromatographic Separations
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MULTIMODAL HPLC: COLUMN SWITCHING Electronically controlled multimodal HPLC is a valuable technique (Kopaciewicz and Regnier, 1983b; Hulpke and Werthmann, 1986), which allows automated fractionation and sample cleanup. In addition to the reagents and procedures described in the preceding sections, the following materials, reagents, and conditions are required for such experiments: (1) an autosampler with two rheodyne 7010-080 valves, third pump (optional), and (2) columns (e.g., for HP-SEC and RP-HPLC). Automated fractionation, for example, with a HP-SEC-RP-HPLC coupling, can be performed with the setup shown in Figure 8.7.12. The valves V2 and V3 allow the on-line or off-line position of the HP-SEC and RP-HPLC column to be independently controlled. To establish an electronically controlled automated fractionation method, the following steps are needed: 1. Establish a suitable method for HP-SECRP-HPLC, whereby the HP-SEC is performed
in the isocratic mode and the RP-HPLC is performed in the gradient elution mode, and the HP-SEC eluent and the eluent A of the RPHPLC are identical. 2. Perform a HP-SEC run (the RP-HPLC column is off-line). 3. Perform a RP-HPLC run (the HP-SEC column is off-line). 4. Determine the valve switching times. 5. Equilibrate both columns with eluent A (with pump P1), with both columns connected. 6. Switch the RP-HPLC column off-line. 7. Perform a HP-SEC run according to step 1 (see Table 8.7.5). 8. Switch to LC-LC coupling according to step 2 (see Table 8.7.5). The fraction is collected at the inlet of the RP-HPLC column. 9. Switch back to the HP-SEC only mode according to step 3 (see Table 8.7.5). The fraction collection on the RP-HPLC column is stopped. The sample stays on the column (no flow) until the HP-SEC column run is completed.
Table 8.7.6 Methodological Requirements of the Different Tasks in RP-HPLC
HPLC of Peptides and Proteins
Task
Purification of one component
Purification of several components simultaneously
Desalting of peptide or protein sample
Characterization of physico-chemical properties
Example
Peptides from solid phase peptide synthesis, enzymatic digestion of a protein
Peptide from tryptic digests, ribosomal proteins, proteins from biological extracts
Fractions from IEX or HIC
Thermodynamic parameters of solute/ligand interaction
Gradient elution Yes Yes Yes Yes Yes Yes — — —
Gradient elution Yes Yes Yes Yes Yes Yes Yes Yes Yes
Step elution Yes Yes Yes Yes Yes — — — —
Isocratic elution Yes Yes Yes — Yes Yes — — Yes
Yes Yes
Yes Yes
— Yes
Yes Yes
—
Yes
Yes
Yes
Mode Analytical scale Preparative scale Measuring of t0 Measuring of tD Column testing A value X value Y value Extra-column band broadening (σ) Regression analysis Optimization procedures Computer simulation
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10. After completion of the HP-SEC run, the HP-SEC column is switched off-line, but flushed with buffer by pump 3, while a RPHPLC gradient elution run is performed according to step 4 (see Table 8.7.5). Illustrative of the use of multimodal HPLC techniques for the separation and analysis of peptide and protein samples is the example shown in Figure 8.7.13, panels A to C, for the separation of closely related 50S ribosomal proteins from Thermus aquaticus under RPHPLC, HP-SEC, and again RP-HPLC protocols under different elution conditions to achieve optimal resolution of specific proteins in high purity for further investigations.
METHOD DEVELOPMENT IN RP-HPLC For the separation of the diverse components of a sample containing peptides or proteins of unknown composition, usually a model is first developed which aims to create separation conditions that result in different retention times of the various components. Currently no algorithm can predict with absolute fidelity, on the basis of the amino acid sequence, the separation behavior of peptides or proteins. In many cases, the nature of the components is not known anyway. Moreover, changes in retention and peak shape will always occur when sample overload or volume overload conditions prevail. There are, however, empirical concepts that describe the retention behavior of peptides and proteins with a ligand in the presence of different solvent combinations. The most commonly adapted concepts are based on the solvophobic theory (Horvath et al., 1976, 1977) and the linear solvent strength theory (Snyder, 1980). These concepts allow the development of fast, robust, and cost-effective separation methods (Boysen et al., 1998). Various aspects of these theoretical approaches can be used to reach different aims with a specific separation (Table 8.7.6). These aims may be: a. the purification of one component, or alternatively several components simultaneously; b. the desalting of a peptide or protein sample; c. the characterization of physico-chemical properties of peptides and proteins in hydrophobic environments; d. examination of the unfolding behavior of proteins under different sorbent or mobile phase conditions;
e. determination of linear free energy dependencies between different members of a peptide analog family; selection and optimization of different elution protocols; f. examination of the effects of different hydrogen bonding solvents on the retention behavior of peptides or proteins; g. the generation of a large variety of empirical data that permits different sorbent types to be validated or batch-to-batch variations characterized. The requirements, in terms of preliminary measurements, for these optimization procedures and data analyses are outlined in Table 8.7.6.
SYSTEMATIC APPROACH TO METHOD DEVELOPMENT The quality of a separation is determined by the resolution of individual peak zones (Hearn, 1991a). Hence, method development is always an optimization of the resolution, with the trade-off either being the speed or the capacity (sample size) of the separation. The resolution of adjacent peak zones can be defined as: Rs =
( t1 − t2 ) (1 2 )( ω1 − ω2 )
Equation 8.7.1 Here, t1 and t2 are the retention times, while ω2 and ω2 are the peak widths of two adjacent peaks. The baseline separation (an overlap of the peaks less than 1%) is given by definition as RS = 1.5. The resolution of two peak zones depends on a large number of factors, including effects that influence the peak symmetry but also on the ratio of the peak areas. RS values are easily estimated using tables and graphs (Snyder et al., 1988). In a chromatogram, every peak pair has a different RS value. In developing a very high-resolution analytical separation of a complex mixture of peptide or protein components, method development always focuses on the least well resolved peak pair. If conditions can be developed to ensure that this “critical peak pair” is well resolved, then all other peaks are also well resolved. However, the peaks that constitute this critical pair can change as a consequence of changes in the experimental conditions. For preparative separations, method development always focuses on the peak of interest and the two adjacent contaminant peaks. In this case, all other peaks can be viewed as superfluous, and directed to the waste. Optimization of the resolution of the peak of interest from the adjacent peaks has to
Conventional Chromatographic Separations
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take into account the sample size and the relative abundances of the three components that form the basis of the separation task. In gradient elution, the resolution also depends on the plate number N, the selectivity α, and the capacity factor k, all of which can be experimentally influenced through systematic changes in individual chromatographic parameters. In this case, resolution is determined from:
k 1 + k Equation 8.7.2
Rs = (1 4 ) N
HPLC of Peptides and Proteins
1
2
( α − 1)
The plate number, N, is the band broadening of the peak zone caused by the column and is a measure of the column performance. The selectivity α describes the selectivity of a chromatographic system for a defined peak pair and is the ratio between the k value of the second peak zone and the k value of the first peak zone. The capacity factor k is a dimensionless parameter of the retention in gradient elution. Its calculation from gradient elution data will be described further below. In contrast to the isocratic elution, in a gradient elution system N, α, and k are the median values for these variables (Hearn, 1991b), since they change during the separation as the shape or duration of the gradient changes. The plate number N has no influence on the selectivity or the retention (except for conditions of temperature change). The selectivity α and the capacity factor k have only a minor influence on N. While N and α change only slightly during the solute migration through the column, the k value can change by a factor of 10 or more. The best chromatographic separation is achieved within a k value range between 1 and 20. Although the resolution is mainly influenced by the mobile phase variables α and k, and nearly independent of N for a given column, an optimization strategy should start, from a logistical point of view, with the optimization of the stationary phase, because many ab initio choices (like column dimensions, choice of ligand, etc.) are determined by the overall strategy (e.g., analytical versus preparative separations). It should be noted, from a theoretical point of view, that optimization of the stationary phase is often performed last (as in some computer simulation programs) when an established analytical method is up-scaled to a preparative method. Typically, a particular RP-HPLC sorbent will be selected empirically as the starting point for the separation. Rarely do most
investigators go to the trouble of optimizing the surface chemistries of their own sorbents, although a number of notable exceptions exist. Based on these considerations and in view of the implication of Equation 8.7.2, the optimization can be performed in three modules in the following order: (1) the optimization of the plate number N, then (2) optimization of the selectivity α, and finally, (3) optimization of the k values.
Optimization Module 1: The Stationary Phase The optimization of the peak efficiency, expressed as the theoretical plate number, N, requires an independent optimization of each of the contributing factors that influence the band broadening of the peak zones due to column and the extra-column effects. With a particular sorbent (ligand type, particle size, and pore size) and column configuration, this can be achieved through optimization of the linear velocity (flow rate), the temperature, the detector time constant, and the column packing characteristics, as well as by minimizing extra-column effects, e.g., by using zero–dead volume tubing and connectors. The temperature of the column and the eluents should be constant (±0.1°C) using a thermostatically controlled system in order to facilitate the reproducible determination of the various column parameters and to ensure reproducibility in the resolution. The theoretical plate numbers measured on the same column are smaller for proteins than for small peptides. This difference in peak width behavior between peptides and proteins can be explained directly from the Knox equation (Knox and Scott, 1983; Freebairn and Knox, 1984), if the N values are expressed as reduced plate heights, i.e., h = L/Ndp, where L is the column length, and dp is the average particle diameter of the sorbent. According to the Knox equation, h is related to the reduced mobile phase velocity, ν, through the relationship h = Aν0.33 + B ν−1 + Cν, where ν = udp/Dm, u is the linear velocity, and A, B, and C are constants that describe the packing and solute transport characteristics of a particular sorbent in a column. Observations of reduced plate heights (h values) in the range 3 < h < 15 for small peptides with reversed-phase sorbents of average particle size of 3 to 10 µm, and for proteins and other high molecular weight macromolecules where h values are >50, have frequently been made, confirming that the Knox equation can be approximated (Snyder et al.,
8.7.26 Supplement 23
Current Protocols in Protein Science
1983) for small peptides as h = Aν0.33 and for proteins as h = Cν, because, in the latter case, only the intraparticle pore transport and surface interaction (term C), contributes significantly to the band broadening. The following parameters can be used to assess column and extra column effects: The A value of the column (a term that reflects the quality of the packing of the sorbent in the column) can be determined according to the procedures described in Stadalius et al. (1987). The A values for well packed columns fall within the limits of 0.5 ≤ A ≤ 1. An A value equal to one corresponds to a column containing a theoretically ideal bed of particles in a dodecahedral close-packed arrangement (Stout et al., 1983). The X value (the column volume of mobile phase outside the pores of the particles) can de calculated according to the procedures described in Stadalius et al. (1987). The average Y value (an estimate of the diffusional restriction of proteins within the pores of the sorbent) can be calculated according to the procedures described in Stadalius et al. (1987). When no restricted diffusion occurs, the value of Y, which corresponds to the ratio (Dp/Dm) is equal to one. Here Dp is the diffusion constant of the protein within the pores of the particle and Dm is the diffusion coefficient of the peptide or protein in the bulk mobile phase. For proteins with molecular weights within the range 10,000 to 80,000, separated using RPHPLC sorbents of large pore diameters (e.g., ≥30 nm or larger), the Y values typically fall within the narrow range of 0.8 ± 0.1, indicating that limited restricted diffusion can occur but that the Dp/Dm ratio is largely independent of the molecular size and weight of the examined proteins. However, with particles of smaller pore size, such as ≤10 nm, a larger range of Y values (e.g., 0.05 ≤ Y ≤ 0.80) will occur with a significant dependency of the Y value on the molecular characteristics of the test proteins. The measurement of the extra-column band broadening σ (which describes the decrease in column performance due to instrumental design) can be determined with a “zero-length column” that connects the injector with a sample loop of 1 µl directly to the detector (Freebairn and Knox, 1984; Hupe et al., 1984). Its value can be calculated based on the calculated peak asymmetry values according to the procedures described in Stadalius et al. (1987). The extra-column band broadening or volume dispersion effects caused by the injector, tubing, and detector influence the performance
of the column and should be less than 10% of the plate number of the column. The extra-column band broadening expressed as peak standard deviation should therefore be less than a third of the band broadening due to the characteristics of the packed column, e.g., as given by: σ COL =
V0
with σ DI ≤ (1 3 ) σ COL N Equation 8.7.3
Hence, the LC system should be designed so as to have minimal extra-column band broadening. Care should be taken to attach fittings in such a way as to avoid unnecessary void volumes and to choose tubing with the smallest possible inner diameter, as well as to keep the volume of the detector cell low. For a standard analytical column, the maximum extra-column band broadening can be therefore be calculated as: σ COL =
1900 µl 10000
= 19 µl with σ DI ≤ 6.3 µl
Equation 8.7.4 In order to obtain this value, the detector cell volume must be 5 µl or less. The flow rate to achieve the minimum plate height, H, for a column can be taken from the literature or experimentally determined according to procedures described in Stadalius et al. (1987).
Optimization Module 2: The Mobile Phase Composition Change in selectivity of the separation with peptides and proteins is the most effective way to influence resolution. This is mainly achieved by changing the chemical nature or concentration of the organic solvent modifier (acetonitrile, methanol, isopropanol, etc.) or the choice of an ion pair reagent (Hancock et al., 1978). Choice of the organic solvent modifier A good starting point is the solvent selectivity triangle approach. Here, solvents are classified according to their relative dipole moment, basicity, or acidity in a triangle. Blends of three different solvents, plus water to provide an appropriate k range, are selected to differ as much as possible in their polar interactions. This selection permits the solvent combinations to mimic the selectivity that is possible for any given solvent, and confines the boundaries of the triangle (Snyder, 1974, 1978). At the
Conventional Chromatographic Separations
8.7.27 Current Protocols in Protein Science
Supplement 23
A
plot of log k ′ versus ϕ t G = 270 min
0.8
log k ′
13 and 14 overlap 11, 12 and 13 overlap
0.4 t = 90 min G
0.0
KEY 7 8 9 10 11 12 13 14 15
– 0.4 0.10
B
3.0
0.15 0.20 ϕ resolution vs. gradient run time t G = 90 min t G = 270 min
RS (in ∆tg time units; min)
2.5 KEY 7/8 8/9 9/10 10/11 11/12 12/13 13/14 14/15
2.0 1.5 1.0 13 and 14 11, 12 and 13 overlap overlap
0.5 0.0 60
C
90 120 150 180 210 240 270 300 t G (min) relative resolution map
3.0
R S (in ∆t g time units; min)
2.5 2.0 1.5
optimal gradient run time t G = 150 min
1.0 12/13
0.5 13/12
13/14 14/13
0.0 60
90 120 150 180 210 240 270 300 t G (min)
8.7.28
Figure 8.7.14 (A) Plots of logarithmic capacity factor ln k ′ versus the volume fraction of organic solvent, ϕ, for 9 different proteins, denoted 7 to 15, obtained from Thermus aquaticus on a C18 RP-HPLC column. As evident from this plot, the intersection of two plots represents complete overlap of the peak zones at the specific gradient run times tG 90 and 270 min. (Legend continues on next page.)
Supplement 23
Current Protocols in Protein Science
HPLC of Peptides and Proteins
same time, these solvents must be totally miscible with each other and with water. Three solvents that best meet these requirements are methanol, acetonitrile, and tetrahydrofuran. Four-solvent mobile phase optimization using three organic solvents and water provides a significant control over α values in reversedphase HPLC. If different organic solvents are used, the different eluotropic strengths (Schoenmakers et al., 1979; Patel and Jefferies, 1987) must be considered in order to elute the sample in the appropriate k range. Choice of ion pair reagent The retention of peptides can be influenced by the presence of ion pair reagents (Horvath et al., 1977; Hearn et al., 1979; Goldberg et al., 1984; Mant and Hodges, 1991c). The ion pair reagents interact with the ionized groups of the peptides: anionic counterions (e.g., hexanesulfonic acid, orthophosphoric acid, TFA, HFBA) interact with basic residues (arginine, lysine, histidine) of a peptide as well as with the protonated N terminus; cationic counterions (e.g., triethylammonium, tetrabutyl-ammonium) interact with ionized acidic residues (glutamic
and aspartic acid, tyrosine and cysteic acid) as well as ionized free C-terminal carboxylic groups. The actual effect on retention depends strongly on the hydrophobicity of the ion-pair reagent and the number of oppositely charged groups on the peptide. Typically, the retention of peptides and proteins in the presence of ion pairing reagents follow asymptotic dependencies on the concentration of the different ion pairing reagent. Once the selectivity parameter is fixed due to the initial choices of the mobile and stationary phase, the further optimization should concentrate on resolution optimization via achieving the most appropriate k value for the different peptides or proteins in the mixture.
Optimization Module 3: The Gradient Conditions For resolution optimization, the strategy takes advantage of the relationship between the gradient retention time of a protein (expressed as the median capacity factor, k) and the median volume fraction of the organic solvent modifier ϕ, in regular RP-HPLC systems based on the concepts of the linear-solvent-strength theory
Table 8.7.7 Parameters Required for the Calculation of ln k0 and S Values and for the Manual Optimization and Optimization with DryLabG/plus
Parameter Gradient delay volume VD (ml) Dead volume t0 (ml) Gradient range (%B) Gradient run time tG1, tG2 (min) Column length L (mm) Column diameter dC (mm) Flow rate F (ml/min) Column plate number N Retention times tg1 for all peaks Retention times tg2 for all peaks
S and logk
RRM
Manual
DryLab
Yes Yes Yes Yes — — — — Yes Yes
Yes — Yes Yes Yes Yes Yes Yes Yes Yes
Figure 8.7.14 (Continued) (B) Plots of the resolution RS (resolution taken as the difference in retention times, ∆tg, for two adjacent peaks) versus the gradient run time tG of these 9 different proteins from Thermus aquaticus. These data represent the blueprint for generation of the relative resolution map (RRM). Since the plotted values of the resolution are absolute values, negative resolution values of peaks that change their elution order are depicted as broken lines. (C) RRM is shown for the nine different proteins from Thermus aquaticus, depicting only the critical peak pairs including their change in retention order. The resolution optimum for this specific set of chromatographic conditions occurs at tG = 150 min.
Conventional Chromatographic Separations
8.7.29 Current Protocols in Protein Science
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(Horvath et al., 1976; Snyder, 1980; Hearn and Grego, 1983), such that: ln k = ln kO − Sϕ
Equation 8.7.5 where k0 is the capacity factor of the solute in the absence of the organic solvent modifier, and S is the slope of the plot of lnk versus ϕ. The values of lnk0 and S can be calculated by linear regression analysis. The underlying principles of an intuitively performed optimization and manually achieved optimization (using Excel spreadsheets, for example, to calculate the ln k0 and S values), or, alternatively, optimization via computer simulation software (e.g., Simplex methods, multivariant factor analysis programs, DryLab G/plus, etc.) are essentially the same. However, the outcomes result in different levels of precision. Two representative approaches are collectively outlined below. Resolution RS of peak zones is optimized through adjustment of k by successive change of the parameters tG and ∆ϕ in the gradient elution mode according to the following steps: Initial experiments; Peak tracking and assignment of the peaks; Calculation of lnk and S values from initial chromatograms; Optimization of gradient run time tG over the whole gradient range; Determination of new gradient range; Calculation of new gradient retention times tg; Change of gradient shape (optional); Verification of results.
HPLC of Peptides and Proteins
Initial experiments In initial experiments, the peptide or protein sample is separated under two linear gradient conditions differing by a factor of 3 in their gradient run times tG (all other chromatographic parameters being held unchanged; Dolan et al., 1989) to obtain the RP-HPLC retention times of each peptide or protein. Irrespective of what optimization strategy will be then used, it is advisable to separate any sample with at least two different gradient run times, in order to identify overlapping peaks. For optimization of the gradient shape and to achieve maximum resolution between adjacent peak zones, the ability to determine the retention times of the peptides or proteins and to classify the parameters that reflect contributions from the mobile phase composition and column dimensions (Table 8.7.7) is essential (Ghrist and Snyder, 1988a,b; Ghrist et al., 1988). The determination of the volume, Vmix,
is useful (Stadalius et al., 1984); determination of dead volume and gradient delay are essential (Snyder, 1980). With these compiled input parameters, various algorithms including DryLab G/plus can generate the relative resolution map (RRM), based on calculation of the corresponding S values and k0 values for each component. If no DryLab G/plus is available, the resolution information can be extracted directly from the vertical distances of the individual RS versus tG plots, whereby a cross-over of lines reflects a peak overlap as shown in Figure 8.7.14A,B,C. Over a small range of ϕ values, the S values and k0 values for an individual peptide or protein will remain essentially constant. It is assumed that no significant conformational change of the individual peptides or proteins occurs as a consequence of the differences in column residency time and gradient time (Purcell et al., 1993). The typical retention behavior of the peptides and proteins is depicted in Figure 8.7.14 as plots of lnk versus ϕ. These plots become nonparallel, and are frequently intersecting, as the mobile phase composition is varied. Choice of starting conditions The S values of peptides and proteins can be empirically correlated with their molecular weights (Snyder et al., 1983), based on a correlation of S = 0.48(MW)0.44. Thus S values, estimated with this equation or, alternatively, values of similar peptides or proteins taken from the literature, can be used to calculate a reasonable starting point for the initial experiments according to the following equation:
tG =
Vm × ∆ϕ × S × k 0.87 × F
Equation 8.7.6 The required gradient run time, tG, for a separation of peptides and proteins with expected S values of around 20 can be calculated for a gradient of 0% to 100% (60% ACN) at a flow rate of 1 ml/min and the ideal k value of 5 as follows:
tG =
1.9 ml × 0.6 × 20 × 5 0.87 × 1ml / min
= 131min
Equation 8.7.7 Based on this calculation, the initial gradients from 0% to 100% eluent B for gradient times of 1 and 3 hr duration can be selected. On the other hand, availability of the S values, derived from two gradient elution RP-
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Current Protocols in Protein Science
HPLC experiments, can be used as an analytical criterion early in the separation of the target peptide or protein from other components in soluble extracts of biological sources, to distinguish low- from high-molecular-weight molecules and, e.g., to exclude peptide fragments participating in the optimization process, without having to resort to SDS-PAGE experiments. The RP-HPLC behavior of small peptides with molecular weights from 300 to 1000 are approximately correlated (Hearn and Grego, 1981; Snyder, 1990) to S values between 3 and 10. For medium-molecular-weight globular proteins, S values above 20 are expected (Aguilar et al., 1985). As an example, cytochrome c, with a molecular weight of ∼12,000, has an S value of 28.8 on a Nucleosil C-18 column (Stadalius et al., 1984). Calculation of ln ko and S values The retention times tg1 and tg2 for a peptide or protein solute separated under conditions of two different gradient run times (tG1 and tG2, whereby tG1 < tG2) can be given by the following equations (Quarry et al., 1986; Ghrist et al., 1988):
t tg1 = 0 log ( 2.3k0 b1 ) + t0 + tD b1 Equation 8.7.8
t0 log ( 2.3k0 b2 ) + t 0 + tD b2
tg2 =
Equation 8.7.9 where:
b1 b2
=
tG2 tG1
=β
Equation 8.7.10 Here, tG1, tG2 are the gradient run time values of tG for two different gradient runs, resulting in different values of b (b1, b2) and tg (tg1, tg1) for a single solute; tg1, tg1 are the gradient retention times for a single solute in two different gradient runs; b1, b2 are the gradient steepness parameters for a single solute and two gradient runs differing only in their gradient times. Steep gradients correspond to large b values and small k values; k0 is the solute capacity factor at the initial mobile phase composition; β is the ratio of tG2 and tG1 which is equivalent to the ratio of b1 and b2; t0 is the column dead time; and tD is the gradient delay time.
For peptides and proteins there is an explicit solution (Stadalius et al., 1984; Quarry et al., 1986) for b and k0, namely: b1 =
t 0 log β
tg2 tg1 − β
(1 − β ) + ( t 0 + tD ) β
Equation 8.7.11
b1 ( tg1 − t0 − tD ) − log ( 2.3b1 ) t0
log k0 =
Equation 8.7.12 From the knowledge of b and k0 the values of k and ϕ can be calculated (Snyder, 1980; Hearn, 1991b):
k =
1 1.15b1
Equation 8.7.13
t0 tg1 − t0 − tD − b log2 1 ϕ= 0 tG1
Equation 8.7.14 Here, k is the value of k′ (capacity factor) for a solute when it reaches the column midpoint during elution; ϕ is the volume fraction of solvent in the mobile phase; ∆ϕ is the change in ϕ for the mobile phase during the gradient elution (∆ϕ = 1 for a 0% to 100% gradient); ϕ is the effective value of ϕ during gradient elution and the value of ϕ at band center when the band is at the midpoint of column, and t0G is the normalized gradient time with t0G1 = tG1/∆ϕ. By linear regression analysis, using k and ϕ, the S value (empirically related to the hydrophobic contact area between solute and ligand) can be derived from the slope of the logk versus ϕ plots, and ln k0 (empirically related to the affinity of the solute towards the ligand) as the y-intercept (Horvath et al., 1976): S=
( ln k0 − ln k ) ϕ
Equation 8.7.15 Peak tracking and assignment of the peaks Complex chromatograms that result from reversed-phase gradient elution can often exhibit changes in peak order when the gradient steepness is changed. Before ln k0 and S values
Conventional Chromatographic Separations
8.7.31 Current Protocols in Protein Science
Supplement 23
are calculated, or computer simulation is used, the peaks from the two initial runs need to be correctly assigned. Several approaches to peak tracking have been described, using algorithms based on relative retention and peak areas (Glajch et al., 1986; Lankmayr et al., 1989; Molnar et al., 1989a), or alternatively, based on diode-array detection (Berridge, 1986; Strasters et al., 1989). The assignment of peaks can be done in the following steps: 1. Integrate the chromatograms (using the integration software of the HPLC system or alternatively an integrator) of the initial runs and correct integration due to baseline drift or other instrumental causes where necessary. 2. Print out both chromatograms including the peak area percent reports. 3. Number all relevant peaks in the chromatogram with the better resolution. Relevant peaks have, e.g., an area >0.5% of the overall peak area. 4. Assign the peaks according to their peak areas allowing a reasonable elution window. The total peak areas, AT1 and AT2 for the initial runs 1 and 2 (whereby tG1 < tG2) are determined, as:
hood that a peak assignment hypothesis is correct:
%RT =
Optimization of the gradient time, tG, over the whole gradient range The capacity factor k is a linear function of the gradient run time tG if ∆ϕ is kept constant. Hence:
k tG
Equation 8.7.16 and their ratio is calculated:
0.87F Vm × ∆ϕ × S
= constant = C
Equation 8.7.20 The optimized gradient run time tGRRM can be obtained from the RRM or alternatively, from the plot of RS versus tG (see Fig. 8.7.14, panels A to C), and yields for each peptide or protein the new values of knew by tGRRM being multiplied by C: C tGRRM = knew
AT2
Determination of the new gradient range If the gradient run time tGRRM is changed in relation to ∆ϕ with t0G1 = constant, the k values do not change, as can be seen from the following equation:
AT1
0 = tG1
Equation 8.7.17 Consequently the ratio of the peak areas, Ai1 and Ai2 of a baseline separated single component in the initial runs 1 and 2, respectively, is:
Ai 2 Ai1
Equation 8.7.18
HPLC of Peptides and Proteins
=
i
i =1
Ri =
− 100
Equation 8.7.21
∑A
RT =
RT
Equation 8.7.19
n
AT =
100 Ri
If the peaks are correctly assigned, RT = Ri. Difficulties arise when peaks partially or completely overlap. If RT << Ri and the peak in run 2 is composed of a single component, the peaks are wrongly matched. If RT >> Ri then the peaks might be correctly matched, but the peak area in run 1 could be enlarged by a hidden additional peak. In order to resolve these difficulties, the deviation (%RT) of Ri from RT can be calculated and taken as a measure of the likeli-
tGRRM ∆ϕ
=
Vm × S × k 0.87 × F
Equation 8.7.22 where: ∆ϕopt =
tGopt 0 tG1
Equation 8.7.23 and the retention time tg of the first peak is >(t0 + tD) and the retention time tg of the last peak is <∆tGopt. Calculation of the new gradient retention times tg Based on the knowledge of the S and the ln k0 values, new gradient retention times can then be calculated. Change of gradient shape (optional). A multisegmented gradient should only be performed when the gradient delay has been meas-
8.7.32 Supplement 23
Current Protocols in Protein Science
ured. With multisegmented gradients, an error in the gradient delay will reoccur at the beginning and at the end of each gradient step. In addition, the effect of Vmix (which can be determined according to the procedures described in Ghrist et al., 1988), which modifies the composition of the gradient at the start and end (rounding of the gradient shape), can lead to deviation of the experimentally determined from the predicted “ideal” retention times in DryLab G/plus simulations. Verification of the results. After completion of the optimization process, the simulated chromatographic separation can now be verified experimentally using the predicted chromatographic conditions.
DETERMINATION OF THERMODYNAMIC PARAMETERS ASSOCIATED WITH PEPTIDE OR PROTEIN INTERACTIONS WITH IMMOBILIZED LIGANDS Isocratic elution can be utilized to determine thermodynamic parameters associated with the interaction of a peptide or protein with an immobilized ligand in all of the various HPLC modes. Thus, the enthalpy, ∆H0assoc, the entropy, ∆S0assoc, and the heat capacity, ∆C0p, of the association of a peptide or protein interacting with immobilized nonpolar ligand in RPHPLC can be evaluated from the dependency of ln k′ on T, i.e., from the van’t Hoff plots (Melander et al., 1984; Haidacher et al., 1996; Vailaya and Horvath, 1996; Boysen et al., 1999; Hearn and Zhao, 1999; Hearn, 2001), and from the Boltzmann-Helmholtz expression:
Table 8.7.8
ln k ′ =
o −∆Hassoc
RT
+
o ∆Sassoc
R
+ ln Φ
Equation 8.7.24 In order to derive this fundamental information, in addition to the procedures and reagents described in the sections above, the following materials and conditions are required: Two 5-liter flasks Peptide or protein (purity >95%) sample at a concentration of 1 mg/ml in a suitable buffer 1. Measure the dead volume t0 of the system as described above. 2. Determine the isocratic conditions at which the peptides or proteins elute in a k′ range of 2 to 10. Typically, these mobile phase conditions will encompass only a narrow range of values, e.g., 22% to 27% acetonitrile/water (v/v)/0.1% TFA for a RP-HPLC separation, and must be determined by running the sample under different eluent conditions. 3. Prepare stock solutions: e.g., 5 liters each of eluents of appropriate compositions that are ∼2% above and 2% below the determined elution range (per the example above 20% and 30% v/v acetonitrile/water/0.1% TFA). 4. Manually mix final buffer in 1% steps from the two stock solutions, according to mixing tables (e.g., Tables for the Laboratory, Merck). Anticipate different eluent consumption for different mobile phase compositions. 5. Inject the sample in order to determine the retention times of the peptide or protein in the temperature range of 5° to 65°C (or higher depending on column specifications) at different isocratic eluent compositions in 5°C increments, allowing no more than 0.5°C column temperature fluctuation, measuring each data point at least twice.
Troubleshooting Strategy and Resulting Spare Part Policy
Troubleshooting strategy
Spare part strategy
Parts replaced
Preventive maintenance
Have always available and change frequently
Anticipation of problems
Have available as backup parts Purchase as needed
Precolumn Guard column Column frits In-line filter Inlet filter Column Fitting and tubing valves Lamps Circuit boards Pump heads Fuses
Wait until complete breakdown
Conventional Chromatographic Separations
8.7.33 Current Protocols in Protein Science
Supplement 23
6. Establish the van’t Hoff plots and fit the data to a polynomial equation using linear regression analysis as described in Boysen et al. (1999). 7. Use the derived coefficients to calculate the thermodynamic parameters, the enthalpy ∆H0assoc of the association, the entropy ∆S0assoc of the association, and the heat capacity ∆C0p exemplified for a second-order approximation of the solute-ligand interaction by the following equations (Boysen et al., 1999):
ln k ′ = b( 0 ) +
b(1) T
+
b(2 ) T2
+ ln Φ
Equation 8.7.25
2b(2 )
T
o ∆Hassoc = − R b(1) +
Equation 8.7.26
b(2 )
T2
o ∆Sassoc = R b( 0 ) −
Equation 8.7.27
2b(2 ) 2 T
∆Cpo = R
Equation 8.7.28 The physical basis and applications of this approach to characterize peptide- or protein-ligand interactions have been extensively described in a variety of papers and reviews (Hearn, 2000a,b,c).
TROUBLESHOOTING There are three approaches to troubleshooting, which are usually practiced in combination: (1) preventive maintenance; (2) anticipation of problems during use by small signs of malfunction; and (3) complacency until complete breakdown. The choice of the troubleshooting approach ultimately affects the spare part strategy and general instrument management and maintenance approach (see Table 8.7.8). As a consequence, the concept of preventative maintenance addresses both the hardware and software, as well as the selection of sorbent characteristics, mobile phase composition, and sample preparation. HPLC of Peptides and Proteins
Preventive Maintenance By practicing preventive maintenance, the downtime of HPLC instrumentation used in peptide and protein separation can be substantially minimized. As a general principle, use clean buffer and reagents, use proper sample preparation techniques, keep air out of the LC system, check the system for leaks, remove all unwanted substances (sample, buffer components) from the system at the end of each day’s work, and be aware of how the system works under normal conditions. Mobile phase Filter all mobile phases except pure HPLCgrade solvents through 0.2-µm filters. During filtration, avoid contamination that might result from the use, e.g., of dirty glassware or rubber stoppers. Degas all solvents thoroughly. Ensure that all fittings on the low-pressure side of the HPLC pump are tight, so that air is not drawn in. Add sodium azide (10,000 ppm) to the mobile phase in order to stop microbial growth, particularly in phosphate and acetate buffers, if they are stored at room temperature for longer than one day. Use a solvent inlet line filter (usually 10 µm porosity) in order to prevent particulate matter entering the pump and to act as a weight to keep the inlet line at the bottom of the solvent reservoir. Cap reservoir loosely to prevent dust from entering the mobile phase and to allow air influx. Avoid cross-contamination of buffer, discard stale mobile phases, and clean the eluent reservoir regularly. Stationary phase Ensure compatibility of the sample solvent with the mobile phase. Incompatibility of the sample buffer with the mobile phase can cause a precipitation of sample in the pores of the column packing. Ensure compatibility of the mobile phase components with each other. Mobile-phase strength effects can cause precipitation of eluent components, such as salts, inside the column, resulting in column blockage or a packing void. Ensure compatibility of the column with the mobile phase. The set of guidelines supplied with the column reveal the pressure and temperature limit at which pH range the column can be used (the wrong pH can cause loss of bonded phase with many silica-based sorbents) and which solvents are compatible with the stationary phase (an incompatible solvent can
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shrink or swell column particles with the polymeric-type of sorbents). Use in-line filter or guard columns. For SEC and other chromatographic modes with impure samples, the use of an 0.5-µm in-line filter is recommended in order to prevent blockage of the column inlet frit. A guard column, preferably packed with the same material as the separation sorbent, prevents a blockage of the column inlet frit and deterioration of the column performance. The integrity of the guard column must be checked regularly, and it must be exchanged when necessary, since a poor guard column can adversely affect column performance. Avoid pressure shocks. Pressure shocks can be minimized by using a pulse damper and by avoiding drastic flow rate changes. Avoid exceeding the pressure limit by using the upper limit switches of the pump in the method program. Do not use default values. Select the upper limit of pressure according to the back-pressure of a new column with an associated method (depending on the viscosity of the mobile phase, it could be the mid-gradient pressure when using water-organic solvents, e.g., 1500 psi) and add a margin of 1000 psi in order to avoid stopping of overnight runs, etc. (yielding a pressure limit of, e.g., 2500 psi). Wash the column after use. All columns should be flushed with 10 column volumes of either an organic solvent/water mixture or water with 10,000 ppm sodium azide to suppress microbial growth. Store the column, when not in use, in the appropriate solvent (see manufacturers’ instructions)—either in >10% organic solvent or with sodium azide when organic solvents have to be avoided. The mobile phase in which the column is stored should not contain any salts, acids, or bases. Fittings and tubing Assemble the fittings so that the tubing contacts the bottom of the fitting body. Tighten the fitting sufficiently to prevent leaks, but not so much as to distort the ferrule. In order to minimize the extra column band broadening, all tubing on the high-pressure side from the injector to the detector (excluding detector outlet) should be 0.01-in. and as short as convenient. Larger i.d. tubing (e.g., 0.020-in.) is suitable for connecting the pump with the injector. The extra column band broadening can be measured with a zero length column.
Manual injector Filter all samples. Avoid pressure values above 5000 psi. Do not use syringes with sharp needles (e.g., from gas chromatography). Maintain adequate gas pressure for air- or nitrogen-operated valves. Flush injector at the end of each day’s work. If a leaking valve requires the replacement of a damaged seal, rebuild the valve using the kit and instructions supplied by valve vendor. Finally, store the injector in a nonbuffered mobile phase when not in use. Pump(s) Avoid air bubbles entering pump. If an air bubble arises in the pump see discussion of Priming of the Pumps and Low-Pressure Lines with Eluents, above. Always flush the HPLC after use to prevent salt deposits at the pump valves. If blocked see Preparing the HPLC System, above. Flush behind the pump seal at the end of each day’s LC operation, first with 5 to 10 ml water, then with 5 to 10 ml methanol or isopropanol in order to remove the water. Many LC systems have a flushing port. By design, the pump seal does not seal completely around the piston. Crystalline buffer residues may act as an abrasive on the pump seal when restarted, damaging the seal with possible scratching of the piston. Store pumps in nonbuffered mobile phase or pure organic solvent. Detector(s) Keep the detector cell clean by washing it at the end of each day’s work with the strong mobile phase. Do not perform acid washing on a routine basis. Switch the detector lamp off when not in use for the next 2 hr or longer. Avoid frequent “on and off” switching. Recorder, printer and data systems Replace ink when recorder ink begins to fade. Lift the pen from the paper and cap it, when recorder is not in use. Check paper supply before HPLC instrument is started and before leaving the HPLC instrument unattended. Ensure that adequate disk storage space exists within the computer for unattended runs.
Anticipation of Problems With this strategy parts are replaced when minor signs of malfunction occur. Conventional Chromatographic Separations
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Troubleshooting after HPLC Breakdown The first aim of troubleshooting is the reduction of HPLC instrument down time. Equally important is the prevention of further damage to the HPLC system, which includes the consequences of troubleshooting. It is therefore crucial to identify the point at which help is needed or whether an event is a case for the manufacturer’s service (as most electronic problems are). Before calling the service engineer, the following sources of help can be utilized if a problem occurs: Operation manual Internal laboratory records, including the logbooks Troubleshooting guides Other people in the laboratory Manufacturer’s technical support line. If technical support from the manufacturer is needed, agree on precise appointment times, be there in order to learn from the support personnel, or alternatively, to challenge them. Brief them thoroughly, and have all of the activities recorded for further reference. Check on warranty times of parts and on proper invoicing. With analytical thinking, and the following good practices, most of the problems can be solved: Describe the problem in as unprejudiced a way as possible. Observation of a problem should be confirmed twice before taking further action. Operate the instrument under reference conditions, if possible. Take notes of every change you make while trying to solve the problem. Label all removed parts. Make only one change at a time. Only use replacement parts that have been proven not to be faulty or worn out. Change a part back, if it did not resolve the problem. This does not apply if a risk exists to cause further damage to parts when they are replaced, if parts are inexpensive, if reinstallation creates the risk of damaging a module, or if parts are scheduled for replacement anyway. Document problem solution.
TERMINOLOGY In this chapter, the nomenclature follows that proposed for the chromatographic sciences by the IUPAC recommendations (Ettre, 1994). HPLC of Peptides and Proteins
SUMMARY In this overview, a variety of protocols and instrumental considerations have been introduced, and various factors involved in the correct selection of experimental procedures have been described. This information represents a starting point for good laboratory practices for the resolution of complex mixtures of peptides and proteins by HPLC methods—not the ultimate solution. As a consequence, the various sections have been prepared with the novice practitioners in mind, rather than the expert chromatographic scientist or peptide/protein chemist with advanced chromatographic skills. These introductory sections should thus facilitate the successful development by biologists and other life scientists with little formal training in the modern separation sciences, but who nevertheless face numerous experimental challenges requiring the high-resolution separation of complex mixtures of peptides and proteins by HPLC techniques.
LITERATURE CITED Aguilar, M.I., Hodder, A.N., and Hearn, M.T.W. 1985. Studies on the optimization of the reversed-phase gradient elution of polypeptides: Evaluation of retention relationships with β-endorphin related polypeptides. J. Chromatogr. 327:115-138. Alpert, A.J. 1990. Hydrophilic-interaction chromatography for the separation of peptides, nucleic acids and other polar compounds. J. Chromatogr. 499:177-196. Anspach, B., Unger, K.K., Davies, J., and Hearn, M.T.W. 1988. Affinity chromatography with triazine dyes immobilized onto activated nonporous monodisperse silicas. J. Chromatogr. 457:195-204. Antia, F.D., Fellegvari, I., and Horvath, C. 1995. Displacement of proteins in hydrophobic interaction chromatography. Ind. Engin. Chem. Res. 34:2796-2804. Ballschmiter, K. and Wossner, M. 1998. Recent developments in adsorption liquid chromatography (NP-HPLC): A review. Fresenius J. Anal. Chem. 361:743-755. Berridge, J.C. 1986. Techniques for the automated optimization of HPLC separations. Wiley Interscience, Chichester. Boysen, R.I., Erdmann, V.A., and Hearn, M.T.W. 1998. Systematic, computer-assisted optimization of the isolation of Thermus thermophilus 50S ribosomal proteins by reversed-phase highperformance liquid chromatography. J. Biochem. Biophys. Methods 37:69-89. Boysen, R.I., Wang, Y., Keah, H.H., and Hearn, M.T.W. 1999. Observations on the origin of the non-linear van’t Hoff behaviour of polypeptides in hydrophobic environments. J. Biophys. Chem. 77:79-97.
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Buchholz, K., Goedelmann, B., and Molnar, I. 1982. High-performance liquid chromatography of proteins: Analytical applications. J. Chromatogr. 238:193-202.
tographic gradients for the separation of either small or large molecules. III. An overall strategy and its application to several examples. J. Chromatogr. 459:43-63.
Burke, T.W., Mant, C.T., Black, J.A., and Hodges, R.S. 1989. Strong cation-exchange high-performance liquid chromatography of peptides. Effect of non-specific hydrophobic interactions and linearization of peptide retention behaviour. J. Chromatogr. 476:377-389.
Ghrist, B.F.D., Coopermann, B.S., and Snyder, L.R. 1988. Design of optimized high-performance liquid chromatographic gradients for the separation of either small or large molecules. Minimising errors in computer simulations. J. Chromatogr. 459:1-23.
Campbell, I.D. and Dwek, R.A. 1984. Biological Spectroscopy pp. 255-277. Benjamin/Cummings, Menlo Park, N.J.
Glajch, J.L., Quarry, M.A., Vasta, J.F., and Snyder, L.R. 1986. Separation of peptide mixtures by reversed-phase gradient elution. Use of flow rate changes for controlling band spacing and improving resolution. Anal. Chem. 58:280-285.
Chang, S., Noel, R., and Regnier, F.E. 1976. High speed ion exchange chromatography of proteins. Anal. Chem. 48:1839-1845. Chaouk, H. and Hearn, M.T.W. 1999. New ligand, N-(2-pyridylmethyl)aminoacetate, for use in the immobilised metal ion affinity chromatographic separation of proteins. J. Chromatogr. A 852:105-115. Chothia, C. 1974. Hydrophobic bonding and accessible surface area in proteins. Nature 248:338339. Dawson, R.M.C., Elliot, D.C., Elliot, W.H., and Jones, K.M. 1986. Data for Biomedical Research. Clarendon Press, Oxford. Dolan, J.W. 1991. Preventive maintenance and troubleshooting LC instrumentation. In High Performance Liquid Chromatography of Peptides and Proteins: Separation, Analysis and Conformation (C.T. Mant and R.S. Hodges, eds.) pp. 23-29. CRC Press, Boca Raton, Fla. Dolan, J.W. and Snyder, L.R. 1989. Troubleshooting LC Systems pp. 1-515. Humana Press, Clifton, N.J. Dolan, J.W., Lommen, D.C., and Snyder, L.R. 1989. DryLab computer simulation for high-performance liquid chromatographic method development. II. Gradient elution. J. Chromatogr. 485:91-112. Ettre, L.S. 1994. New, unified nomenclature for chromatography. Chromatographia 38:521526. Fausnaugh, J.L. and Regnier, F.E. 1986. Solute and mobile phase contributions to retention in hydrophobic interaction chromatography of proteins. J. Chromatogr. 359:131-146. Fausnaugh, J.L., Pfannkoch, E., Gupta, S., and Regnier, F.E. 1984. High-performance hydrophobic interaction chromatography of proteins. Anal. Biochem. 137:464-472. Freebairn, K.W. and Knox, J.H. 1984. Dispersion measurements on conventional and miniaturized HPLC systems. J. Chromatogr. 19:37-47. Ghrist, B.F.D. and Snyder, L.R. 1988a. Design of optimized high-performance liquid chromatographic gradients for the separation of either small or large molecules. II. Background and theory. J. Chromatogr. 459:25-41. Ghrist, B.F.D. and Snyder, L.R. 1988b. Design of optimized high-performance liquid chroma-
Goldberg, A.P., Nowakowska, A.E., Antle, P.E., and Snyder, L.R. 1984. Retention-optimization strategy for the high-performance liquid chromatographic ion-pair separation of samples containing basic compounds. J. Chromatogr. 316:241-260. Gooding, D.L., Schmuck, M.N., Nowlan, M.P., and Gooding, K.M. 1986. Optimization of preparative hydrophobic interaction chromatographic purification methods. J. Chromatogr. 359:331337. Haidacher, D., Vailaya, A., and Horvath, C. 1996. Temperature effects in hydrophobic interaction chromatography. Proc. Nat. Acad. Sci. U.S.A. 93:2290-2295. Hancock, W.S., Bishop, C.A., Prestidge, R.L., Harding, D.R., and Hearn, M.T.W. 1978. Reversedphase, high-pressure liquid chromatography of peptides and proteins with ion-pairing reagents. Science 200:1168-1170. Hearn, M.T.W. 1991a. High-performance liquid chromatography of peptides and proteins: General principles and basic theory. In High-Performance Liquid Chromatography of Peptides and Proteins: Separation, Analysis and Conformation (C.T. Mant and R.S. Hodges, eds.) pp. 95-104. CRC Press, Boca Raton, Fla. Hearn, M.T.W. 1991b. High-performance liquid chromatography of peptides and proteins: Quantitative relationships for isocratic and gradient elution of peptides and proteins. In High-Performance Liquid Chromatography of Peptides and Proteins: Separation, Analysis and Conformation (C.T. Mant and R.S. Hodges, eds.) pp. 105-122. CRC Press, Boca Raton, Fla. Hearn, M.T.W. 2000a. Physico-chemical factors in polypeptide and protein purification and analysis by high-performance liquid chromatographic techniques: Current status and future challenges. In Handbook of Bioseparation (S. Ahuja, ed.) pp. 72-235. Academic Press, New York. Hearn, M.T.W. 2000b. Reversed-phase and hydrophobic interaction chromatography of peptides and proteins. In HPLC of Biopolymers (K. Gooding and F.E. Regnier, eds.). Marcel Dekker, New York. Hearn, M.T.W. 2001. Conformational behaviour of polypeptides and proteins in reversed-phase environments. In Theory and Practice of Biochro-
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matography (M.A. Vijayalakshmi, ed.) pp. 72235. Harwood Academic Publishers, Chur, Switzerland. Hearn, M.T.W. and Grego, B. 1981. Organic solvent modifier effects in the separation of unprotected peptides by reversed-phase liquid chromatography. J. Chromatogr. 218:497-507. Hearn, M.T.W. and Grego, B. 1983. Further studies on the role of the organic modifier in reversedphase high-performance liquid chromatography of polypeptides. Implications for gradient optimization. J. Chromatogr. 255:125-136. Hearn, M.T.W. and Zhao, G.L. 1999. Investigations into the thermodynamics of polypeptide interaction with nonpolar ligands. Anal. Chem. 71:4874-4885. Hearn, M.T., Grego, B., and Hancock, W.S. 1979. Investigation of the effects of pH and ion-pair formation on the retention of peptides on chemically-bonded hydrocarbonaceous stationary phases. J. Chromatogr. 185:429-444. Hearn, M.T.W., Hodder, A.N., and Aguilar, M.I. 1988. Comparison of retention and bandwidth properties of proteins eluted by gradient and isocratic anion-exchange chromatography. J. Chromatogr. 458:27-44. Hearn, M.T.W., Middleton, S., Jackson, R.W., and Chaouk, W. 1997. Constrained immobilised metal ion affinity absorbents for protein purification. Int. J. Biochem. 2:153-190. Heinitz, M.L., Kennedy, L., Kopaciewicz, W., and Regnier, F.E. 1988. Chromatography of proteins on hydrophobic interaction and ion-exchange chromatographic matrices: Mobile phase contributions to selectivity. J. Chromatogr. 443:173182.
Kato, Y., Komiya, K., and Hishimoto, T. 1982. Study of experimental conditions in high-performance ion-exchange chromatography. J. Chromatogr. 46:13-22. Kato, Y., Kitamura, T., and Hashimoto, T. 1983. High-performance hydrophobic interaction chromatography of proteins. J. Chromatogr. 266:49-54. Knox, J.H. and Scott, H.P. 1983. B and C terms in the van Deemter equation for liquid chromatography. J. Chromatogr. 282:297-313. Kopaciewicz, W. and Regnier, F.E. 1983a. Mobile phase selection for the high-performance ion-exchange chromatography of proteins. Anal. Biochem. 133:251-259. Kopaciewicz, W. and Regnier, F.E. 1983b. A system for coupled multiple-column separation of proteins. Anal. Biochem. 129:472-482. Kopaciewicz, W. and Regnier, F.E. 1986. Synthesis of cation-exchange stationary phases using an adsorbed polymeric coating. J. Chromatogr. 358:107-117. Kopaciewicz, W., Rounds, M.A., and Regnier, F.E. 1985. Stationary phase contributions to retention in high-performance anion-exchange protein chromatography: Ligand density and mixed mode effects. J. Chromatogr. 318:157-172. Lankmayr, E.P., Wegscheider, W., and Budna, K.W. 1989. Global optimization of HPLC separations. J. Liq. Chromatogr. 12:35-58.
Hodder, A.N., Aguilar, M.I., and Hearn, M.T.W. 1990. The influence of the gradient elution mode and displacer salt type on the retention properties of closely related protein variants separated by high-performance anion-exchange chromatography. J. Chromatogr. 506:17-34.
Litowski, J.R., Semchuk, P.D., Mant, C.T., and Hodges, R.S. 1999. Hydrophilic interaction/cation-exchange chromatography for the purification of synthetic peptides from closely related impurities: Serine side-chain acetylated peptides. J. Peptide Res. 54:1-11.
Horvath, C., Melander, W., and Molnar, I. 1976. Solvophobic interactions in liquid chromatography with nonpolar stationary phases. J. Chromatogr. 125:129-156.
Mant, C.T. and Hodges, R.S. 1985. Separation of peptides by strong cation-exchange high-performance liquid chromatography. J. Chromatogr. 327:147-155.
Horvath, C., Melander, W., and Molnar, I. 1977. Liquid chromatography of ionogenic substances with nonpolar stationary phases. Anal. Chem. 49:142-154.
Mant, C.T. and Hodges, R.S. 1991a. Requirement for peptide standards to monitor ideal and nonideal size exclusion chromatography. In High Performance Liquid Chromatography of Peptides and Proteins: Separation, Analysis and Conformation (C.T. Mant and R.S. Hodges, eds.) pp. 125-134. CRC Press, Boca Raton, Fla.
Hulpke, H. and Werthmann, U. 1986. Column switching. In Practice of High Performance Liquid Chromatography (H. Engelhardt, ed.) pp. 147. Springer Verlag, New York. Hupe, K.P., Jonker, R.J., and Roing, G. 1984. Determination of band-spreading effects in high-performance liquid chromatographic instruments. J. Chromatogr. 285:253-265.
HPLC of Peptides and Proteins
Jiang, W., Graham, B., Spiccia, L., and Hearn, M.T.W. 1998. Protein selectivity with immobilized metal ion tacn sorbents: Chromatographic studies with human serum proteins and several other globular proteins. Anal. Biochem. 255:4758.
Jeno, P., Scherer, P.E., Manningkrieg, U., and Horst, M. 1993. Desalting electroeluted proteins with hydrophilic interaction chromatography. Anal. Biochem. 215:292-298.
Mant, C.T. and Hodges, R.S. 1991b. The use of peptide standards for monitoring ideal and nonideal behaviour in cation-exchange chromatography. In High Performance Liquid Chromatography of Peptides and Proteins: Separation, Analysis and Conformation (C.T. Mant and R.S. Hodges, eds.) pp. 171-185. CRC Press, Boca Raton, Fla. Mant, C.T. and Hodges, R.S. 1991c. The effects of anionic ion-pairing reagents on the peptide re-
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tention in reversed-phase chromatography. In High-Performance Liquid Chromatography of Peptides and Proteins: Separation, Analysis and Conformation, Vol. 327-341 (C.T. Mant and R.S. Hodges, eds.). CRC Press, Boca Raton, Fla. Mant, C.T., Parker, J.M., and Hodges, R.S. 1987. Size-exclusion high-performance liquid chromatography of peptides. Requirement for peptide standards to monitor column performance and non-ideal behaviour. J. Chromatogr. 397:99112. Mant, C.T., Litowski, J.R., and Hodges, R.S. 1998a. Hydrophilic interaction/cation-exchange chromatography for separation of amphipathic alphahelical peptides. J. Chromatogr. 816:65-78. Mant, C.T., Kondejewski, L.H., and Hodges, R.S. 1998b. Hydrophilic interaction/cation-exchange chromatography for separation of cyclic peptides. J. Chromatogr. 816:79-88.
Purcell, A.W., Aguilar, M.I., and Hearn, M.T.W. 1993. Dynamics of peptides in reversed-phase high-performance liquid chromatography. Anal. Chem. 65:3038-3047. Quarry, M.A., Grob, R.L., and Snyder, L.R. 1986. Prediction of precise isocratic retention data from two or more gradient elution runs. Analysis of some associated errors. Anal. Chem. 58:907917. Regnier, F.E. 1983. High-performance liquid chromatography of biopolymers. Science 222:245252. Regnier, F.E. 1984. High-performance ion-exchange chromatography. Methods Enzymol. 104:170-189. Rickard, E.C., Strohl, M.M., and Nielsen, R.G. 1991. Correlation of electrophoretic mobilities from capillary electrophoresis with physicochemical properties of proteins and peptides. Anal. Biochem. 197:197-207.
Melander, W.R., Corradini, D., and Horvath, C. 1984. Salt-mediated retention of proteins in hydrophobic-interaction chromatography. Application of solvophobic theory. J. Chromatogr. 317:67-85.
Risley, D.S. and Strege, M.A. 2000. Chiral separations of polar compounds by hydrophilic interaction chromatography with evaporative light scattering detection. Anal. Chem. 72:1736-1739.
Melander, W.R., el Rassi, Z., and Horvath, C. 1989. Interplay of hydrophobic and electrostatic interactions in biopolymer chromatography. Effect of salts on the retention of proteins. J. Chromatogr. 469:3-27.
Schoenmakers, P.J., Biliet, H.A.H., and Galan, L.D. 1979. Influence of organic modifiers on the retention behaviour in reversed-phase liquid chromatography and its consequences for gradient elution. J. Chromatogr. 185:179-195.
Molnar, I., Boysen, R.I., and Jekow, P. 1989a. Peak tracking in high-performance liquid chromatography based on normalized band areas. A ribosomal protein sample as an example J. Chromatogr. 485:569-579.
Snyder, L.R. 1970. Adsorption chromatography: Scope, technique and equipment. Methods Med. Res. 12:11-36.
Molnar, I., Boysen, R.I., and Erdmann, V.A. 1989b. Reversed-phase high-performance liquid chromatography of Thermus aquaticus 50S and 30S ribosomal proteins. Chromatographia 28:39-44. Ohlson, S., Hansson, L., Larsson, P.O., and Mosbach, K. 1978. High-performance liquid affinity chromatography (HPLAC) and its application to the separation of enzymes and antigens. FEBS Letts. 93:5-9. Papadoyannis, I.N., Zotou, A.C., and Samanidou, V.F. 1995. Solid-phase extraction study and RPHPLC analysis of lamotrigine in human biological fluids and in antiepileptic tablet formulations. J. Liq. Chromatogr. 18:2593-2609. Patel, H.B. and Jefferies, T.M. 1987. Eluotropic strength of solvents. Prediction and use in reversed-phase high-performance liquid chromatography. J. Chromatogr. 389:21-32. Pohl, T. and Kamp, R.M. 1987. Desalting and concentration of proteins in dilute solution using reversed-phase high-performance-liquid-chromatography. Anal. Biochem. 160:388-391. Porath, J. 1988. High-performance immobilizedmetal-ion affinity chromatography of peptides and proteins. J. Chromatogr. 443:3-11. Porath, J., Carlsson, J., Olsson, I., and Belfrage, G. 1975. Metal chelate affinity chromatography, a new approach to protein fractionation. Nature 258:598-599.
Snyder, L.R. 1974. Classification of the solvent properties of common liquids J. Chromatogr., Chromatogr. 92:223-230. Snyder, L.R. 1978. Classification of the solvent properties of common liquid solvents. J. Chromatogr. Sci. 16:223-234. Snyder, L.R. 1980. Gradient elution. In HPLC: Advances and Perspectives, Vol. 1 (C. Horvath, ed.) pp. 208-316. Academic Press, New York. Snyder, L.R. 1990. Gradient elution separation of large biomolecules. In HPLC of Biological Macromolecules. Methods and Application (K.M. Gooding and F.E. Regnier, eds.) pp. 231-259. Marcel Dekker, New York. Snyder, L.R., Stadalius, M.A., and Quarry, M.A. 1983. Gradient elution in reversed-phase HPLC separation of macromolecules. Anal. Chem. 55:1413-1430. Snyder, L.R., Glajch, J.L., and Kirkland, J.J. 1988. Practical HPLC Method Development pp. 1260. Wiley Interscience, New York. Stadalius, M.A., Gold, H.S., and Snyder, L.R. 1984. Optimization model for the gradient elution separation of peptide mixtures by reversedphase high-performance liquid chromatography. Verification of retention relationships. J. Chromatogr. 296:31-59. Stadalius, M.A., Ghrist, B.F., and Snyder, L.R. 1987. Predicting bandwidth in the high-performance liquid chromatographic separation of large
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biomolecules. II. A general model for the four common high-performance liquid chromatography methods. J. Chromatogr. 387:21-40. Stout, R.W., DeStefano, J.J., and Snyder, L.R. 1983. High-performance liquid chromatographic column efficiency as a function of particle composition and geometry and capacity factor. J. Chromatogr. 282:263-286. Strasters, J.K., Billiet, H.A.H., de Galan, L., Vandeginste, B.G.M., and Kateman, G. 1989. Automated peak recognition from photodiode array spectra in liquid chromatography. J. Liq. Chromatogr. 12:3-22. Strege, M.A. 1998. Hydrophilic interaction chromatography electrospray mass spectrometry analysis of polar compounds for natural product drug discovery. Anal. Chem. 70:2439-2445. Vailaya, A. and Horvath, C. 1996. Retention thermodynamics in hydrophobic interaction chromatography. Ind. Engin. Chem. Res. 35:2964-2981. Wilce, M.C.J., Aguilar, M.I., and Hearn, M.T.W. 1995. Physicochemical basis of amino acid hydrophobicity scales: Evaluation of four new scales of amino acid hydrophobicity coefficients derived from RP-HPLC of peptides. Anal. Chem. 67:1210-1219. Wirth, H.J. and Hearn, M.T.W. 1993. High-performance liquid chromatography of amino acids, peptides and proteins. CXXX. Modified porous zirconia as sorbents in affinity chromatography. J. Chromatogr. 646:143-151. Wu, S.L., Benedek, K., and Karger, B.L. 1986a. Thermal behavior of proteins in high-performance hydrophobic-interaction chromatography. On-line spectroscopic and chromatographic characterization. J. Chromatogr. 359:3-17.
Zhang, J.F. and Wang, D.I.C. 1998. Quantitative analysis and process monitoring of site-specific glycosylation microheterogeneity in recombinant human interferon-gamma from Chinese hamster ovary cell culture by hydrophilic interaction chromatography. J. Chromatogr. B 712:73-82.
KEY REFERENCES Dolan and Snyder, 1989. See above. An excellent practical manual. Glajch, J.L. and Snyder, L.R. 1990. Computer Assisted Method Development for High-Performance Liquid Chromatography. pp. 1-682. Elsevier Science Publishers, Amsterdam. An advanced text for experienced scientists. Hearn, M.T.W. 1991. HPLC of Peptides, Proteins and Polynucleotides, Fundamental Principles and Contemporary Applications. pp. 1-776. VCH Publ, Deerfield Beach, Fla. A comprehensive text encompassing most applications. Hearn, M.T.W. 2000. Physicochemical factors in polypeptide and protein purification by highperformance liquid chromatography: Current status and future challenges. In Handbook of Bioseparation (S. Ahuja ed.) pp. 1-652. Academic Press, New York. Hearn, M.T.W. 2001. RPC and HIC of peptides and proteins. In HPLC of Biological Macromolecules. Methods and Applications (K.M. Gooding and F.E. Regnier, eds.). Marcel Dekker, New York. A detailed survey of application and theory.
Wu, S.L., Figueroa, A., and Karger, B.L. 1986b. Protein conformational effects in hydrophobic interaction chromatography. Retention characterization and the role of mobile phase additives and stationary phase hydrophobicity. J. Chromatogr. 371:3-27.
Mant, C.T. and Hodges, R.S. 1991. High-Performance Liquid Chromatography of Peptides and Proteins: Separation, Analysis and Conformation. pp. 1-938. CRC Press, Boca Raton, Fla.
Yip, T.T., Nakagawa, Y., and Porath, J. 1989. Evaluation of the interaction of peptides with Cu(II), Ni(II), and Zn(II) by high-performance immobilized metal ion affinity chromatography. Anal. Biochem. 183:159-171.
McMaster, M.C. 1994. HPLC: A Practical User’s Guide. pp. 1-211. VCH Publishers, Inc., New York.
Yoshida, T. 1998. Calculation of peptide retention coefficients in normal-phase liquid chromatography. J. Chromatogr. 808:105-112.
Snyder et al., 1988. See above.
Zachariou, M., Traverso, I., and Hearn, M.T.W. 1993. O-phosphoserine as a new chelating ligand for use with hard Lewis metal ions in the immobilized-metal affinity chromatography of proteins. J. Chromatogr. 646:107-120.
Vijayalakshmi, M.A. 2000. Theory and Practice of Biochromatography. pp. 1-721. Harwood Academic Publishers.
Zamyatnin, A.A. 1972. Protein volume in solution. Progress Biophys. Mol. Biol. 24:107-123.
A comprehensive survey of applications.
A general guide to HPLC.
An excellent advanced text.
A useful guide to modern chromatography.
Contributed by Reinhard I. Boysen and Milton T.W. Hearn Monash University Victoria, Australia
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Expanded-Bed Adsorption Chromatography
UNIT 8.8
All protein purification processes can be divided into three main steps (UNIT 8.1). The first is the so-called capture step, since it involves immobilization of the target protein on an adsorptive surface. This step can also be viewed as a combination of clarification, concentration, stabilization, and initial purification. Most downstream processes require that the feed stream be free of particulate material, and so the initial capture phase of most processes includes some sort of particle reduction operation. If clarification can be achieved on a chemically active adsorptive surface, such as a chromatographic medium or a membrane, then particulate removal can be accomplished simultaneously with concentration, stabilization, and initial purification. Since the target molecule is immobilized along with only a few closely related molecules, the target is concentrated and somewhat purified after the capture step. It is also known that when a protein is immobilized, it becomes stabilized, because it is somewhat resistant to protease attack. Therefore, the most successful capture steps will result in some degree of concentration, purification, and stability enhancement, along with particulate removal. The second step in a purification process is called intermediate purification. In this step, most of the remaining contaminants are removed from the product. Chromatography is the most useful tool in this phase, providing high selectivity and a high rate of product recovery. The most useful and commonly used techniques are hydrophobic-interaction chromatography (HIC; UNIT 8.4), affinity chromatography (Chapter 9), and ion-exchange chromatography (UNIT 8.2). The last phase of a purification scheme, the polishing step, also typically involves chromatography. Every contaminant that has been purified along with the product to this point is very closely related to the product, and very high selectivity and efficiency are needed to separate the desired product from these closely related structural variants. The most useful and commonly employed techniques are gel-filtration chromatography (UNIT 8.3), hydrophobic-interaction chromatography (UNIT 8.4), ion-exchange chromatography (UNIT 8.2), and reversed-phase high-performance liquid chromatography (UNIT 8.7). More thorough discussions of the design of a separation protocol can be found in Sofer and Hagel (1997) and in other units in this chapter. The capture phase of the separation can be made more efficient if the separation power of chromatography can be combined with an operation that permits the use of particulateladen (i.e., nonclarified) feed. In recent years, expanded beds have been successfully used to combine clarification and concentration. The expanded bed also accomplishes an initial purification of the product. The references cited in the following three reviews provide a useful summary of these applications: Chase (1994); Hjorth (1997); and Shiloach and Kennedy (2000). In addition, two special issues of the journal Bioseparation (Volume 8, Iss. 1-5, 1999; Volume 10, Iss. 1-3, 2001) are dedicated to expanded-bed adsorption chromatography. Although expanded-bed adsorption (EBA) is a scalable technique, this unit focuses on EBA at the bench scale. EBA is useful in three protein purification/capture scenarios. The first is the recovery of molecules from cell cultures in which the recombinant cell line is very shear-sensitive. In these cases, clarification of the cell harvest by membrane filtration and/or centrifugation causes product degradation, because the cell lysis that results from these techniques leads to the release of proteases into the harvest broth. This scenario is common in the use of engineered CHO cell lines. The second scenario applies to situations in which the recombinant protein is attached to or strongly associated with
Contributed by Robert M. Kennedy Current Protocols in Protein Science (2005) 8.8.1-8.8.25 C 2005 by John Wiley & Sons, Inc. Copyright
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the cell membrane. Due to the high charge density of the resin, the resin will out-compete the cell for the protein, yielding very high recovery rates. (The chromatography resin has a charge density on the order of mmol/ml, which is very high compared with the charge density associated with the cell surface or cell surface debris; charged molecules are drawn to this higher charge density, similar to the way in which a stronger battery will make a bulb glow brighter than a weaker battery will.) This is the case in certain Pichia expression systems. The third scenario in which EBA is particularly useful is when exposure of the target protein to proteases must be avoided. In this scenario, the usual strategy is to work in the cold, as is the case in the recovery of some enzymes from natural sources.
STRATEGIC PLANNING The experimental strategy for the development of an expanded-bed process involves four major steps: (1) method scouting, (2) method optimization, (3) process verification, and (4) production. Most investigators can accomplish the first two steps and be satisfied that they have a reproducible method. Industrial applications typically require completion of all four steps if the process is to be scaled up for use in a production environment. The purpose of method scouting is to define the most suitable adsorbent and the optimal conditions for binding and elution. Method scouting is performed in both packed-bed mode (with a clarified feedstock sample) and expanded-bed mode. The alternate protocols in this unit describe initial method scouting procedures—namely, determining conditions for ion-exchange chromatography, HIC, and affinity chromatography. Employing these protocols using a clarified feed will only provide an approximate starting condition, however. Development of a method using only clarified feed does not take into account the charge carried by the biomass in the crude feedstock. This can lead to erroneous results, poor fluidization, and lowered capacity.
Method scouting with clarified feed The adsorbent is selected using the same principles that apply to packed-bed chromatography. The medium of choice will exhibit the strongest binding to the target protein while retaining only a minimum amount of contaminating proteins. To establish the chemical parameters of the separation, initial method scouting runs should be performed using feed that has been clarified. Later steps will be performed with unclarified feed to establish the physical parameters of the separation. It is critical to keep the characteristics of the feedstock in mind when designing a process. For method scouting, the bed height is typically 15 cm in a 2.5- or 2.6-cm-diameter column. This bed height allows the target protein sufficient time to bind at normal flow velocities of ∼300 cm/hr. The flow velocity used should be similar to what will subsequently be used in expanded-bed mode. It should be noted that the use of flow velocity (in cm/hr) instead of flow rate (in ml/min) throughout this unit is intentional, as this allows the flow to be normalized for any column diameter. The first step in the conversion from cm/hr to ml/min is to multiply the cross-sectional area of the column being used (π r2 , where r is the column radius in cm) by the specified velocity in cm/hr. This gives a result in units of cm3 /hr, which is equivalent to ml/hr. From there, the result can be divided by 60 to give the desired flow rate in units of ml/min. For a given flow velocity, actual flow rates will increase as a function of r2 . Knowledge of flow velocities is of practical utility when comparing runs between different columns, or when scaling a successful run to a column of larger diameter. Expanded Bed Adsorption Chromatography
Elution can be performed in stepwise fashion or by applying a gradient. Information from elution studies is used to optimize selectivity for the target molecule and to avoid the binding of contaminants that bind less avidly to the resin.
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Figure 8.8.1 Breakthrough curve for BSA on the STREAMLINE DEAE resin as determined by frontal analysis, in which the sample (BSA) is continuously applied to the column and C/Co , the ratio of BSA concentration in the flowthrough (C) to BSA concentration in the feedstock (Co ), is measured as a function of the amount of BSA applied. The resin becomes saturated (i.e., C/C0 approaches 1) as more and more BSA is applied to the column. As can be seen from the graph, the chromatographic behavior of the resin in expanded-bed mode is the same as in packed-bed format. Reproduced with permission from Amersham Biosciences (Expanded-Bed Adsorption: Principles and Methods; see Internet Resources) and Academic Press (Shiloach and Kennedy, 2000).
Once selectivity has been optimized, maximum dynamic binding capacity is determined from breakthrough tests conducted using the chosen binding conditions. Figure 8.8.1 shows breakthrough capacity curves in packed and expanded modes and in two STREAMLINE columns (Amersham Biosciences), representing laboratory scale and pilot scale. Note the similar shape of the breakthrough curves for packed-bed mode and expanded-bed mode, as well as for the different-sized STREAMLINE columns (STREAMLINE 50 versus STREAMLINE 200; numeric specification refers to the inner diameter of the column in mm). Thus, adsorption performance is the same in expandedbed and packed-bed modes, and STREAMLINE columns can be scaled up while maintaining the same performance. When defining a suitable maximum loading capacity for EBA chromatography, it is necessary to apply a safety margin to compensate for sources of variability in the sample that could affect binding capacity. The maximum loading amount should be ∼75% of breakthrough capacity.
Method scouting with feedstock containing biomass A second method scouting run should be performed starting with unclarified biomass. Two simple experiments that require no specialized equipment can be conducted to measure the interaction of the biomass with the resin (Kula et al., 2001; A. Liten and K. Kinealy, unpub. observ.). The first involves simply comparing the amount of biomass present in the sample before and after passage through the column by measuring light scattering at 600 nm (Fig. 8.8.2). This experiment can be done with flowthrough monitors and peak integration, but it can also be accomplished off-line using cuvettes and a spectrophotometer. (The reader is directed to UNIT 5.3, for instructions on measuring biomass with a spectrophotometer; briefly, for E. coli, an optical density reading of 0.5 at
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Figure 8.8.2 Schematic diagram of an experiment measuring gross interaction between the expanded-bed resin and the biomass in a feedstock mixture. A pulse of biomass is injected, and its light-scattering profile is measured at 600 nm before and after passage through the column. The resulting peaks are then integrated, with a change in peak area after passage of the biomass through the column indicating that the sample has interacted with the resin. Reproduced with permission from A. Liten.
600 nm will correspond to a concentration of ∼5 × 108 cells/ml.) The relevant calculation is a simple mass balance of biomass before and after passage through the resin, and the experiment should be followed by inspection of the resin for aggregation. Resin aggregation can be seen with the naked eye and is mostly due to interaction of the resin with the biomass. If aggregation has occurred, clumps of resin, rather than a uniform bed, will be evident. If aggregation of the resin is seen and appears to affect bed expansion or loading capacity for the target molecule, then dilution of the biomass, changes in salt concentration, or the use of a different salt may be necessary. If aggregation persists, and loading capacity and target recovery are unacceptably low, it may be the case that EBA is not an appropriate technique for harvesting the target molecule from the feedstock being used. In general, choose conditions that result in the lowest degree of sample interaction with the resin and in little or no aggregation. The second experiment involves estimating the equilibrium ratio of target protein to resin (see Support Protocol 1). The purpose of this procedure is to determine the maximum amount of sample that should be loaded onto a column (usually 30% of the equilibrium target protein binding capacity). The most important considerations in initial EBA method scouting are 1) to never choose conditions under which the resin is observed to aggregate; 2) to select conditions under which there is minimal interaction between the biomass and the resin, but under which the target protein still binds; and 3) to set the maximum amount of sample that can be loaded onto the column at 30% of the equilibrium capacity for the chosen conditions. After method scouting, the remaining three steps in the development of an expanded-bed process are performed in expanded-bed mode.
EBA method optimization Expanded Bed Adsorption Chromatography
The next step in the development of an expanded-bed process is optimization. The expanded-bed method is first optimized on a small scale (e.g., a 2.5-cm column) using crude, unclarified feed. The purpose of this optimization is to examine the effect of
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the crude feed on the stability of the expanded bed and on chromatographic performance and to make adjustments to the flow velocity if necessary. During method optimization, the process should be challenged by applying a sample load close to the maximum sample load as defined in the breakthrough study performed on the packed bed. In addition, a breakthrough capacity study should be performed to determine the effect of the crude feed on the binding capacity of the resin. Elution, wash, and re-equilibration volumes should also be determined as a part of method optimization, and finally, a suitable cleaning-in-place (CIP) procedure should be chosen. (Development of the CIP method should be initiated in the method scouting step.) CIP procedures for expanded-bed media are complicated by the fact that the debris load can be challenging to clean up. That is, unlike chromatographic media, expanded-bed media are typically exposed to a significant amount of contamination and particulates. CIP protocols for expanded-bed media are usually straightforward, but their identification requires an investment of time and effort. An expanded bed can be characterized by a variety of tracer techniques that can be used to monitor bed performance after CIP. See Batt et al. (1995) for a description of bed characterization.
Process verification Process verification, or pilot-scale work, is done using a pilot-scale column (e.g., a 20-cm column). For some applications, this column may also be suitable for the final production scale. The usual bed height of 15 cm provides a sedimented bed volume of 4.7 liters in a 20-cm column. Scale-up is performed by increasing the column diameter while maintaining the sedimented bed height, flow velocity, and expanded bed height.
STREAMLINE EBA columns Two main features characterize a STREAMLINE column. First, the design of the liquid distribution system at the base of the column ensures the formation of a stable expanded bed. Second, a movable adaptor allows the column height to be altered during the different stages of an expanded-bed adsorption cycle. There are STREAMLINE columns for each of the last three stages of process development—i.e., laboratory-scale, pilot-scale, and production-scale columns. These three stages of process development are also phases in the development of an optimized STREAMLINE expanded-bed unit operation. For the majority of applications, method optimization is done on the laboratory scale using STREAMLINE 25 or STREAMLINE 50 columns, process verification is done on the pilot scale using STREAMLINE 100 or STREAMLINE 200 columns, and finally, manufacture is carried out using productionscale columns that range in size from 400 to 1200 mm (inner diameter). This unit focuses on laboratory-scale EBA. The appropriate column for laboratory-scale EBA is the 25-mm-i.d. column, STREAMLINE 25. The STREAMLINE 50 column (i.d., 50 mm) can be used for laboratory-scale work as well, but it is top-heavy and not as userfriendly as the STREAMLINE 25 column. The STREAMLINE 25 column is available in two formats: manual and hydraulic. The equipment setup diagrams in this unit are for the hydraulic version. The manual version is set up in exactly the same manner, except that there is no need for the hydraulic pump or the hydraulic valve after the hydraulic pump. If only one peristaltic pump is available, the manual setup is more appropriate.
Resins Gel media for EBA are commercially available (Table 8.1.1). The fluidization properties of several types of media used in protein recovery are reported by Thommes (1997). Of these,
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STREAMLINE media are the most widely used. Commercially available resins include STREAMLINE SP, STREAMLINE DEAE, STREAMLINE SP XL, STREAMLINE Q XL, STREAMLINE rProtein A, STREAMLINE chelating resin, and STREAMLINE heparin resin (Amersham Biosciences); CM Hyper Z resin (Ciphergen); and Prosep A and Prosep G resins (Millipore). STREAMLINE adsorbents exhibit Gaussian-like distributions with respect to particle size and particle density. This particle polydispersity is an important factor that contributes to the stability of the expanded bed. The size and density gradients present cause the beads to be positioned at specific heights in the expanded bed depending on the sedimentation velocity of the individual adsorbent particles. The mean particle size of STREAMLINE beads is 200 µm, compared with ∼90 µm for packed-bed chromatography media. This larger diameter results from modifications made to increase particle density, which is ∼1.2 g/ml. In the bead population, smaller, lighter particles move to positions near the top of the expanded bed, while larger, heavier particles move to the bottom, resulting in a stable, uniform expansion. In other words, the beads find their ideal position in the column, which explains the low degree of axial dispersion in EBA. The sizes and densities of STREAMLINE adsorbents have been defined to allow optimal expansion at flow velocities up to 500 cm/hr. STREAMLINE Q XL is an anion-exchange resin based on highly cross-linked 6% agarose modified by the inclusion of an inert quartz core to produce the desired density. Long molecules of dextran are coupled to the agarose matrix, and ion-exchange groups are then attached to these dextran chains. This construction increases the effective interaction volume as well as the availability of ligands for adsorption of the target. The overall result is a large increase in binding capacity. The binding capacity of the STREAMLINE Q XL resin is >110 mg BSA per ml adsorbent, compared with 40 mg/ml for the normal DEAE Fast Flow ion exchanger (Amersham Biosciences). STREAMLINE SP XL is a cation exchanger that is also based on highly cross-linked 6% agarose modified by the inclusion of an inert quartz core to produce the desired density. As in the STREAMLINE Q XL resin, long molecules of dextran are coupled to the agarose matrix, and ion-exchange groups are then attached to these dextran chains. The binding capacity of the STREAMLINE SP XL resin is >140 mg lysozyme per ml adsorbent, compared with 60 mg/ml for the normal SP Fast Flow ion exchanger (Amersham Biosciences). The degree of expansion (calculated as the ratio of expanded bed height to sedimented bed height) for both of the above media is 2 to 3 at a flow rate of 400 cm/hr. The degree of expansion is a quick and useful measure of bed stability, although it is a less accurate measure compared with the number of theoretical plates (UNIT 8.1; also see Basic Protocol, steps 20 to 23).
Expanded Bed Adsorption Chromatography
STREAMLINE rProtein A is an affinity adsorbent for purifying monoclonal and polyclonal antibodies. It is based on highly cross-linked 4% agarose modified by the inclusion of an inert metal alloy core to provide the desired density. The binding capacity of the STREAMLINE rProtein A resin is ∼50 mg human IgG per ml of gel, and the degree of expansion is between 2.2 and 3 at 300 cm/hr. The ligand used in this medium is recombinant protein A, which has been engineered to include a C-terminal cysteine residue. The epoxy chemistry is controlled to favor a thioether coupling, providing single-point attachment of the protein A. This oriented coupling enhances the binding of IgG. The recombinant protein A has no mitogenic activity in human lymphocytes in vitro. (Mitogenic activity has sometimes been a concern when using native protein A from Staphylococcus aureus.) Leakage of protein A from the STREAMLINE rProtein A resin is generally low. In pharmaceutical production processes, protein A must be removed from the final
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product; leached recombinant protein A can be removed efficiently from an IgGcontaining fraction by gel filtration or ion-exchange chromatography. STREAMLINE chelating resin is used for immobilized metal affinity chromatography (IMAC; UNIT 9.4). This technique exploits the different affinities of proteins for metal ions bound to chelating groups on the adsorbent. The chelating support can be primed with metal ions by passing a salt solution through the column. The chelating group consists of iminodiacetic moieties on a spacer arm coupled to the matrix via stable covalent linkages. STREAMLINE heparin resin consists of heparin immobilized to the matrix. This adsorbent can be used in two modes: (1) as an affinity matrix for coagulation factors, or (2) as a high-capacity cation exchanger, due to the large number of anionic sulfate groups on the ligand. For example, when interacting with nucleic acid binding proteins, the ligand mimics a nucleic acid in terms of its polyanionic structure. STREAMLINE phenyl resin consists of phenyl groups immobilized to the matrix. This adsorbent is used for hydrophobic-interaction chromatography, or HIC (UNIT 8.4), a technique that separates proteins and peptides based on the strength of their hydrophobic interactions with hydrophobic groups attached to an uncharged matrix. It is estimated that as much as 30% to 40% of the accessible surface area of water-soluble proteins is nonpolar, which explains why HIC is such a widely applicable technique. STREAMLINE chelating resin, STREAMLINE heparin resin, and STREAMLINE phenyl resin, along with the DEAE and SP ion exchangers mentioned above, have average densities of 1.2 g/ml and expand to 2.5 times their original bed height in water at a flow rate of 300 cm/hr. CM Hyper Z is a weak cation exchanger in which ion exchange is mediated by carboxymethyl groups; the base material is zirconium oxide. The resin has a dynamic binding capacity of 30 mg IgG per ml at a flow rate of 300 cm/hr and is designed to function in a high-salt environment (15 to 20 millisiemens per cm), which is typical of fermentation conditions. Prosep A and Prosep G resins contain immobilized recombinant protein A and immobilized recombinant protein G, respectively. Both have very high densities, because their base material is 75- to 125-µm controlled-pore glass. These resins have a dynamic capacity of 25 mg IgG per ml at a flow rate of 750 cm/hr, and they support flow velocities of up to 1500 cm/hr for regeneration and equilibration.
EXPANDED-BED ADSORPTION CHROMATOGRAPHY This protocol covers the preparation of the EBA resin and the setup of the apparatus, as well as the typical steps in a chromatography experiment (equilibration of the resin, evaluation of the column, application of the sample, and elution of the target molecule). Cleaning and storage are described at the end of the protocol. The selection of buffers for equilibration and elution will be guided by the mode of chromatography being performed. Typically, the standard buffer systems for bimolecular purification are used. As mentioned before, the expanded-bed column can be operated manually or by a hydraulic pump. The figures related to this protocol show the hydraulic system, because it is the more complex of the two. In order to use the hydraulic system, two peristaltic pumps that can be dedicated to expanded-bed chromatography are needed. If only one peristaltic pump is available, then it will be necessary to use the manual system. Aside from making it possible to move the column flow adaptor hydraulically, the hydraulic
BASIC PROTOCOL
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system is no different from the manual system. Wherever necessary, separate instructions are given for manual and hydraulic columns. For methods development, ∼1 liter of feedstock is needed. Once the method is established, the volume of feedstock used will be determined by the amount of time required for feed application and washout. The protocol below is applicable for all of the resin types that can be used in EBA (see Alternate Protocol 1, Alternate Protocol 2, and Alternate Protocol 3).
Materials Chromatography resin (see Strategic Planning) Start buffer Equilibration buffer 0.25% (v/v) acetone in equilibration buffer Feedstock under constant stirring (e.g., by a magnetic stir plate or overhead mixer) Wash buffer Elution buffer Cleaning-in-place (CIP) solution 20% (v/v) ethanol (optional) Sintered glass funnel (optional) Water aspirator (optional) Chromatography column (see Strategic Planning) with or without hydraulic pump Chromatography apparatus (see Fig. 8.8.3, Fig. 8.8.4, Fig. 8.8.6, Fig. 8.8.7, and Fig. 8.8.8 for various schematics) consisting of the following: Peristaltic pump(s) (e.g., Cole-Parmer Masterflex tubing pump; do not use peristaltic pumps designed for laboratory chromatography, such as P1, Microperpex, or Rabbit pumps, as these pumps do not produce sufficiently high flow rates) attached to Masterflex size 16 silicon pump tubing Two-channel, four-port valve for flow reversal Single-channel, three- or four-port valves Pressure gauge Flowthrough UV monitor with S2-type or 6-mm-i.d. flow cell (do not use flow cells from analytical instruments, since cell debris may clog them) Chart recorder Flowthrough conductivity meter (optional) Flowthrough pH meter (optional) Wash bottle Spirit level Label tape Additional reagents and equipment for CIP (see Support Protocol 2) NOTE: See Expanded Bed Adsorption: Principles and Methods (Amersham Biosciences; see Internet Resources) for additional details regarding setup of the chromatography apparatus.
Expanded Bed Adsorption Chromatography
Prepare the resin for use NOTE: Since EBA resins are typically a mixture of particles of various sizes and densities, they, unlike other chromatographic resins, cannot simply be poured into a column. Instead, most EBA resins must be specially prepared to ensure an appropriate mixture of particle sizes and densities in the column. All particles in Hyper Z and Prosep media, however, have the same density, and so there is no need to perform the sampling procedure described in steps 2 to 6 if either of these media is being used; instead, simply pour the slurry supplied by the manufacturer to the desired bed height in step 8.
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1. Shake the resin container until the resin is completely slurried. 2. Immediately pour the entire contents of the container into a sintered glass funnel or (if the volume of the resin container is larger than is convenient for a glass funnel) into a beaker or bucket, and allow the resin to settle completely. 3a. If using a funnel: Allow the liquid component of the resin slurry to drain through the funnel. 3b. If using a beaker or bucket: Remove the liquid from the top of the resin cake by aspiration through a hose connected to a water aspirator, taking care not to aspirate the resin along with the liquid. 4. Using the following formula, calculate the volume of resin needed to obtain the desired sedimented bed height: V = π r2 h, where V is the volume of resin needed, r is the column radius, and h is the desired sedimented bed height. If column radius and bed height are entered into the formula in units of cm, then the calculated volume will be in units of cm3 , or ml. The column for initial method scouting should have a sedimented bed height of ∼15 cm.
5. Calculate the amount (in grams) of resin needed to obtain the desired bed height by multiplying the volume calculated in step 4 by the average density of the resin (in g/ml). The average density of the rProtein A STREAMLINE resin is 1.3 g/ml. The average density of all other STREAMLINE media is 1.2 g/ml.
6. Cut a wedge-shaped slice of the settled resin from the funnel, beaker, or bucket, and weigh it. Trim the slice or add more resin (also cut in wedges from the resin cake in the funnel, beaker, or bucket) as needed to obtain the desired amount of resin (as determined in step 5). Transfer the desired amount of resin to a beaker, and suspend in start buffer to obtain an ∼50% (w/v) slurry. Cutting wedge-shaped pieces ensures that a representative sample (in terms of particle sizes and densities) is obtained from the resin cake. The exact volume of start buffer used to make the slurry is not critical. The purpose of making a slurry in starting buffer is simply to make it easier to pour the resin into the column.
Set up the column for use 7. Connect the column to the chromatography apparatus. Set each valve as indicated in Figure 8.8.3. Remove the lid and adaptor from the column, and then use pump 1 to fill the column to approximately two-thirds of its capacity with start buffer via the bottom valve (valve 4). For columns with i.d. >25 mm, it may be necessary to remove air bubbles trapped under the column’s bottom net; this is done by drawing air out of the net using tubing connected to a water aspirator.
8. Make sure that the resin/start buffer slurry is uniform and, without letting the resin settle, quickly pour the slurry into the column. Use additional start buffer to rinse the sides of the container from which the slurry was poured, and pour this rinse liquid into the column as well. Use a wash bottle filled with start buffer to dislodge any resin stuck to the sides of the column. 9. After the resin has been poured into the column, add start buffer until the column is full to capacity. At this point, a tray or a tote can be placed under the column to catch any buffer that might spill out in subsequent steps.
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Figure 8.8.3 Schematic diagram showing the tubing connections, valve placement, pump configuration, and monitoring options for expanded-bed adsorption chromatography. Valves that control column operation are labeled V1, V2, V3, and V4. This diagram shows a hydraulic column; adaptor movement is controlled by pump 2. In a manual column, pump 2 and V3 are not present, and the column lid does not have a connector to the hydraulic line, but all other connections are the same. Buffer 1 and buffer 2 are typically equilibration buffer and elution buffer, respectively. P1 and P2 are fraction collection lines. CIP, cleaning-in-place solution (see Support Protocol 2); UV, flowthrough UV detector; cond., flowthrough conductivity meter; pH, flow-through pH meter. Reproduced with permission from Amersham Biosciences (Expanded-Bed Adsorption: Principles and Methods; see Internet Resources).
10. Tip the adaptor such that its bottom makes at least a 45◦ angle with respect to the surface of the buffer in the column, and then insert it into the column under the buffer, taking care not to introduce air bubbles. As soon as the entire adaptor O-ring is just inside the column, straighten the adaptor so that its bottom is parallel to the surface of the buffer, and then put the lid on the column and secure it with the screws provided by the manufacturer. IMPORTANT NOTE: No resistance should be felt when inserting the adaptor into the column. If resistance is felt, remove the adaptor from the column, make sure that the O-ring is wet and is seated properly in its groove, and then try again. Several attempts may be required to achieve proper insertion of the adaptor, as it is critical that no air bubbles be trapped in the adaptor net in the assembled column.
11a. If using a manual column: Position the adaptor so that its distance from the bottom of the column is ∼4× the height of the sedimented bed.
Expanded Bed Adsorption Chromatography
11b. If using a hydraulic column: Prime the hydraulic line with water. Open the flow path from the hydraulic pump to the column (through valve 3; Fig. 8.8.3), and set valve 2 open to waste. Use the hydraulic pump to pump water into the hydraulic chamber, and then stop the pump once the adaptor has been displaced downward by a few centimeters. Remove the column lid and aspirate the buffer/water mixture
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from the hydraulic chamber, and then refill the chamber to the top with water. Once the chamber has been refilled, put the lid back on the column and secure it as in step 10. The position of the adaptor is set to allow for expansion of the resin.
Fill the system 12. Set the bottom valve (valve 4) to the bypass (waste) position and run pump 1 to fill all tubing with the appropriate start buffer. 13. Put the column on-line by setting the bottom valve (valve 4) to allow direct flow into the column.
Equilibrate the column 14. Check the orientation of the column using a spirit level, and adjust the positioning of the column as necessary to ensure that it is mounted completely vertically. For optimal results, it is important that the column be completely vertical. If a tray was placed under the column to catch liquid spillage during setup, it can now be removed.
15. Mark the height of the sedimented bed on the column using label tape. This sedimented bed height should be measured and recorded so that the degree of bed expansion upon equilibration can be calculated. The author recommends marking the column with narrow label tape, although any laboratory tape is suitable for this purpose.
16. Set each valve in the appropriate position for column equilibration (Fig. 8.8.4). Using pump 1, begin pumping equilibration buffer upward through the column at a flow velocity of 200 to 500 cm/hr. Record the actual flow velocity. 17. Allow the bed to stabilize at the flow velocity being used. Assess bed stability either visually (step 19) or using a residence time distribution test (steps 20 to 23), and once the bed has stabilized, use a second piece of label tape to mark the height of the expanded bed on the column. Continue pumping buffer upward through the column. The bed is stable when no further expansion is evident. Bed stabilization should take ∼45 min.
18. As the equilibration buffer flows through the column, monitor the pH of the outlet stream. As soon as the pH of this stream is the same as that of the equilibration buffer, begin recirculating the equilibration buffer by removing the column outlet tube from the waste container and putting it into the flask containing the equilibration buffer.
Evaluate bed stability by visual inspection 19. If using a column made of glass (e.g., a STREAMLINE 25 column), visually inspect the bed to assess bed stability. If the bed is stable at a certain bed height, it is safe to assume that the entire chromatographic bed is stable. If constant liquid densities, viscosities, and temperatures are maintained, bed expansion should be predictable from run to run. A significant decrease in bed expansion after several runs could be indicative of air trapped under the bottom distributor of the column, blockage in the bottom of the column, fouled resin, or the column’s not being absolutely vertical. See the troubleshooting guide in Expanded Bed Adsorption: Principles and Methods (Amersham Biosciences; see Internet Resources) for more information on these topics. Bed stability can be evaluated more rigorously by performing a residence time distribution test (RTD; steps 20 to 23), which assesses the degree of mixing in a column by determining the number of theoretical plates.
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Figure 8.8.4 Schematic diagram of the appropriate valve positions for equilibrating and expanding the resin bed in an expanded-bed adsorption chromatography column. Valves that control column operation are labeled V1, V2, V3, and V4. Note that the adaptor is at an elevated position within the column. This is necessary only as long as the degree of expansion of the resin is not well characterized. Once bed expansion is well characterized, the adaptor should be kept as close as possible to the top of the expanded bed to minimize mixing. CIP, cleaning-in-place solution (see Support Protocol 2); UV, flowthrough UV detector; cond., flowthrough conductivity meter; pH, flow-through pH meter; P1 and P2, fraction collection lines. Reproduced with permission from Amersham Biosciences (Expanded-Bed Adsorption: Principles and Methods; see Internet Resources).
Evaluate bed stability using a residence time distribution (RTD) test (optional) NOTE: The RTD test is required for evaluating bed stability in steel columns, as the contents of a closed steel column cannot be assessed visually. 20. Lower the adaptor to within 1 cm of the top of the equilibrated expanded bed. 21. Set the UV detector to 280 nm, and start the recorder used to plot UV detector readings. When the baseline is stable, change the feed from equilibration buffer to 0.25% (v/v) acetone in equilibration buffer. Acetone absorbs strongly at 280 nm.
22. Look for a positive step signal to arise on the recorder plot (Fig. 8.8.5). Once this UV step signal is stable (at its maximum level, referred to as the 100% signal level), switch the feed back to equilibration buffer alone. On the recorder plot, label the point at which this switch was made as time 0.
Expanded Bed Adsorption Chromatography
23. Look for a negative step signal to appear on the recorder plot, and then wait for the UV signal to stabilize at its baseline level (0%). Calculate the number of theoretical plates (N) using the following formula: N = t2 /σ 2 , where t is the mean residence time, calculated as the time (from time 0) needed for the UV signal to fall to the 50% level, and σ is the standard deviation, taken to be half of the time needed for the UV signal to drop from the 84.15% level to the 15.85% level (Fig. 8.8.5).
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Figure 8.8.5 Chart paper output for a residence time distribution test using acetone as the tracer. A mark is placed on the paper to indicate time 0, the point at which a stable maximum signal is present and the feed is switched from tracer-containing equilibration buffer to tracer-free equilibration buffer. The quantity 2σ is calculated as the time (from time 0) needed for the UV signal (measured at 280 nm) to drop from the 84.15% level to the 15.85% level on the washout (negative-step) side of the tracer pulse. t, mean residence time; σ , standard deviation about the mean residence time. Reproduced with permission from Amersham Biosciences (Expanded-Bed Adsorption: Principles and Methods; see Internet Resources).
Note that 2σ , and not σ itself, is the quantity that the investigator measures directly from the recorder plot. Thus, it is necessary to divide this measurement by 2 to obtain the quantity that is entered into the formula for calculating N. Given a sedimented bed height of 15 cm and a flow velocity of 300 cm/hr, a value of N between 25 and 30 is desirable. A difference of ≥20% in N values from two runs performed under identical test conditions is indicative of air trapped under the bottom distributor of the column, blockage in the bottom of the column, fouled resin, or the column’s not being entirely vertical.
Apply sample to column 24. Raise the adaptor to a position very near the top of the column. Due to the higher viscosity of the sample mixture, bed expansion is greater when the sample, rather than buffer alone, is applied to the column. Thus, the adaptor is raised in this step (before application of the sample) to accommodate any additional bed expansion that might occur.
25. Set each valve in the appropriate position for applying the feedstock to the column (Fig. 8.8.6). Using pump 1, begin pumping the feedstock upward through the column at the same flow velocity used in step 16, continuously stirring the feedstock mixture that is fed into the pump to ensure uniformity throughout the course of sample application. Once any additional bed expansion resulting from feedstock application is complete, set the adapter ∼5 cm above the top of the expanded bed. 26. Continue pumping feedstock through the column, monitoring its application using the UV detector. Look for a large flowthrough peak, and direct this peak to waste. If a hydraulic column is being used, valve 3 should also be set open to waste during this step to allow for slight adaptor movement if pressure builds up in the bed.
Wash column 27. Once all of the feedstock has been applied to the column, change the flow to wash buffer. Using pump 1, pump the wash buffer upward through the column at the same flow velocity used for sample application. Look for some amount of bed contraction as the sample mixture is replaced with wash buffer. When this contraction is
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Figure 8.8.6 Schematic diagram of the appropriate valve positions for applying feedstock to an expanded-bed adsorption chromatography column. Valves that control column operation are labeled V1, V2, V3, and V4. Expect the bed to expand to a higher level during feed application (as compared with equilibration). Although it is not shown in this diagram, it is critical that the feedstock being fed to pump 1 be stirred throughout the course of application. CIP, cleaning-in-place solution (see Support Protocol 2); UV, flowthrough UV detector; cond., flowthrough conductivity meter; pH, flow-through pH meter; P1 and P2, fraction collection lines. Reproduced with permission from Amersham Biosciences (Expanded-Bed Adsorption: Principles and Methods; see Internet Resources).
complete, adjust the position of the adaptor so that it sits just above the top of the bed (Fig. 8.8.7). Record the position of the adaptor. Adjusting the position of the adaptor in this way will speed up the wash cycle and reduce buffer consumption by eliminating the large mixing chamber between the top of the bed and the bottom of the adaptor.
28. Continue pumping wash buffer through the column until the UV signal returns to its baseline level.
Elute product 29. Turn off pump 1 and allow the resin bed to sediment. It should be noted that the elution method presented here is known as elution in “packedbed” mode. The alternative to eluting in packed-bed mode is to immediately switch from wash buffer to elution buffer and elute the product while the bed is expanded; this is easier than eluting in packed-bed mode, but it dilutes the collected product by ∼40%.
30a. If using a manual column: When sedimentation is complete, lower the adaptor so that it sits just above the bed. Expanded Bed Adsorption Chromatography
30b. If using a hydraulic column: When sedimentation is complete, use the hydraulic pump to pump water into the hydraulic chamber (above the adaptor), and stop the pump once the adaptor sits just above the bed.
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Figure 8.8.7 Schematic diagram of the appropriate valve positions for washing an expandedbed adsorption chromatography column. Valves that control column operation are labeled V1, V2, V3, and V4. The bed height decreases as the switch is made from feedstock to wash buffer, and so the adaptor is lowered accordingly. CIP, cleaning-in-place solution (see Support Protocol 2); UV, flowthrough UV detector; cond., flow-through conductivity meter; pH, flowthrough pH meter; P1 and P2, fraction collection lines. Reproduced with permission from Amersham Biosciences (Expanded-Bed Adsorption: Principles and Methods; see Internet Resources).
31. Set each valve in the appropriate position for eluting product from the column (Fig. 8.8.8). Using pump 1, pump 1 to 2 bed volumes of elution buffer downward through the column at a flow velocity of 100 cm/hr, and collect the eluted fraction as it comes off the column (as indicated by a peak on the UV monitor).
Perform CIP and prepare column for storage 32a. If using a manual column: Immediately after collection of the product, set each valve in the appropriate position for column equilibration (Fig. 8.8.4). Raise the adaptor to the same height at which it sat at the end of step 27. 32b. If using a hydraulic column: Immediately after collection of the product, set valve 1 and valve 4 in the appropriate positions for column equilibration (Fig. 8.8.4); set valve 2 so that it forms a dead end in the outlet line coming from the top of the column (so that flow cannot exit the top of the column), and set valve 3 open to waste. Using pump 1, pump elution buffer upward through the column to lift the adaptor to the same height at which it sat at the end of step 27. Once the adaptor is at the appropriate height, set valves 2 and 3 as shown in Figure 8.8.4. With valve 2 creating a dead end in the outlet line and valve 3 open to waste, the only way for pressure to be released from the column as the elution buffer passes through is for the fluid in the hydraulic chamber to go to waste. This will cause the adaptor to move upward.
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Figure 8.8.8 Schematic diagram of the appropriate valve positions for product elution (in the downward direction) from an expanded-bed adsorption chromatography column. Valves that control column operation are labeled V1, V2, V3, and V4. Note that the product is collected as a fraction through the P1 line. CIP, cleaning-in-place solution (see Support Protocol 2); UV, flowthrough UV detector; cond., flowthrough conductivity meter; pH, flow-through pH meter; P1 and P2, fraction collection lines. Reproduced with permission from Amersham Biosciences (Expanded-Bed Adsorption: Principles and Methods; see Internet Resources).
33. Switch the feed from elution buffer to CIP solution, and carry out the CIP process developed in Support Protocol 2. CIP solution should always be pumped upward through the column. The resin bed may initially act as a plug during this process, but the plug will fall apart as CIP proceeds.
34a. If the column is to be reused immediately after CIP: Switch the feed from CIP solution to equilibration buffer. Using pump 1, pump equilibration buffer upward through the column as in step 16, and then proceed from step 17 onwards. If cleaning was adequate, the height of the expanded bed will be approximately the same as it was in the initial run. The sedimented bed height will not change unless more resin is added to the column.
Expanded Bed Adsorption Chromatography
34b. If the column is to be stored after CIP: Switch the feed from CIP solution to storage solution. Using pump 1, pump the storage solution upward through the column. When storage solution appears in the outlet line, stop all pumps and allow the bed to settle. Once the bed has settled, remove the lid from the column, and then remove all liquid from the hydraulic chamber using a hose attached to a water aspirator. Refill the chamber with 20% ethanol, put the lid back on the chamber, and secure the lid using the screws provided by the manufacturer. Store the column in accordance with the manufacturer’s instructions.
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ESTIMATION OF THE EQUILIBRIUM RATIO OF TARGET PROTEIN TO RESIN
SUPPORT PROTOCOL 1
While a good approximation of the resin’s capacity for binding the desired product can be obtained by working with clarified feed (i.e., the target protein isolated from the crude feedstock), a more practical option is to estimate the resin capacity using unclarified feedstock. The following is a method for estimating how much resin is needed to adsorb a given amount of product in unclarified feedstock.
Materials Unclarified feedstock containing 0.5 mg/ml target protein Chromatography resin (see Strategic Planning) End-over-end test tube mixer Additional reagents and equipment for protein assays (UNIT 3.4) 1. In the following way, prepare a series of three test tubes containing unclarified feedstock combined in different ratios with the chromatography resin to be used, making sure to maintain a constant volume of feedstock across all three test tubes.
Tube 1 10 ml unclarified feedstock containing 0.5 mg/ml target protein 0.125 g resin Ratio: 40 mg target protein per gram of resin Tube 2 10 ml unclarified feedstock containing 0.5 mg/ml target protein 0.250 g resin Ratio: 20 mg target protein per gram of resin Tube 3 10 ml unclarified feedstock containing 0.5 mg/ml target protein 0.5 g resin Ratio: 10 mg target protein per gram of resin 2. Cap tubes and place in an end-over-end mixer for 4 hr at room temperature (i.e., the temperature at which the EBA column will be run). 3. Clarify samples by performing low-speed centrifugation (1000 × g is more than adequate) at room temperature for 15 min. Assay the supernatant from each of the three tubes for target protein concentration (UNIT 3.4). Centrifugation time and speed may have to be adjusted empirically. All feedstock and resin debris must be in the pellet (leaving a clarified sample as the supernatant), because particulates will interfere with protein assays. Any of the colorimetric methods described in Unit 3 will be appropriate for assaying target protein concentration.
4. Identify the equilibrium concentration, Ceq , defined as the condition (i.e., the ratio of target protein to resin) under which the concentration of target protein in the supernatant is equal to half of the initial concentration of target protein in the feedstock. It may be necessary to extrapolate (by solving a proportionality equation based on the data points obtained in step 3) to determine Ceq . This extrapolation method assumes that Ceq falls somewhere between the highest and lowest tested ratios of target protein to resin.
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5. Calculate the target equilibrium capacity (Qeq ) using the following equation: Qeq = [(C0 − Ceq ) × Vl ]/Mad , where C0 is the initial concentration of target protein in the feedstock, Ceq is the equilibrium concentration identified in step 4, Vl is the volume of liquid in the test tube, and Mad is the amount (i.e., mass) of resin in the test tube. For a given condition, an EBA column should be loaded to 30% of its equilibrium capacity. SUPPORT PROTOCOL 2
DEVELOPMENT OF A CLEANING-IN-PLACE PROCESS FOR EXPANDED-BED ADSORPTION CHROMATOGRAPHY Expanded-bed adsorption (EBA) chromatography is unique among chromatographic methods in that the feed is not clarified before application to the column. As a result, EBA resins are exposed to cells, cell debris, and the products of cell lysis, as well as to culture media and their additives. In order to prolong the life of the EBA resin, a cleaning-in-place (CIP) protocol must be developed. The development of a CIP protocol for EBA resins is challenging. The appropriate CIP protocol depends on the feedstock used, but some general advice can be given. This protocol provides the basic steps for generating a CIP protocol and gives some suggestions for CIP solutions. In addition, a reference is provided in which the reader can find details on CIP methods developed for several different types of feedstock.
Additional Materials (also see Basic Protocol) CIP solution: 1.0 M NaOH, 6 M urea, or 6 M guanidine·HCl Additional components for inclusion in CIP solution (if necessary; e.g., isopropyl alcohol, acetic acid, benzonase) 1. Perform repeated expanded-bed chromatography runs (see Basic Protocol) on the same column (without performing CIP between consecutive runs) until it is evident that the degree of expansion or the chromatographic properties of the resin are changing. Changes in the chromatographic properties of the resin are signaled by, for example, the inability or decreased ability of the resin to bind the target protein, bleed-through of the target protein along with the feedstock, and the binding of contaminants that flowed through the resin in earlier runs.
2. Set all valves in the chromatography apparatus and adjust the height of the column adaptor as in Basic Protocol, step 32a (if using a manual column), or Basic Protocol, step 32b (if using a hydraulic column). Using pump 1, pump CIP solution (6 M urea or 6 M guanidine·HCl for protein A–containing media, and 1.0 M NaOH for all other types of media) upward through the column at a flow velocity of 100 to 300 cm/hr for a contact time of 8 hr or overnight. IMPORTANT NOTE: Do not start CIP with the CIP solution flowing downward through the column.
3. Switch the flow from CIP solution to equilibration buffer, adjust valve 1 so that pump 1 is connected directly to the top of the column, and then begin pumping equilibration buffer downward through the column.
Expanded Bed Adsorption Chromatography
4. Once two column volumes of equilibration buffer have passed through the column, adjust valve 1 so that pump 1 is connected directly to the bottom of the column, and then begin pumping equilibration buffer upward through the column. When the bed stabilizes, measure its height and use this measurement to calculate the degree of expansion of the resin. The resin is equilibrated when the pH of the outlet stream is the same as the pH of the equilibration buffer.
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5a. If the chromatographic bed is successfully regenerated (i.e., if the degree of expansion and the chromatographic properties of the resin return to normal): Repeat the CIP testing process (steps 1 to 4) multiple times, each time using a shorter contact time in step 2. Identify the minimum contact time needed for effective CIP. 5b. If the chromatographic bed is not successfully regenerated: Repeat the CIP testing process (steps 1 to 4) multiple times, each time adding new components (e.g., 30% v/v isopropyl alcohol, 25% v/v acetic acid, 5 µl/l benzonase) to the CIP solution used in step 2. Determine which of these components need to be added to the CIP solution for effective CIP. Once a suitable cleaning protocol has been identified, it can be used in all EBA runs involving the feedstock in question.
ION-EXCHANGE EXPANDED-BED ADSORPTION CHROMATOGRAPHY Resins for ion-exchange EBA chromatography are chosen in the same way that resins are chosen for other types of ion-exchange column chromatography—that is, based on what is known about the pI values of the target protein and all contaminants (UNIT 8.2).
ALTERNATE PROTOCOL 1
To determine the appropriate binding conditions for the target molecule in an ionexchange application, perform the basic test tube experiments described in UNIT 8.2, Support Protocol 1. Capacity determinations can be made as in UNIT 8.2, Support Protocol 2, but the results of these determinations should be treated only as approximations, since the capacity of the resin is impacted by the presence of debris. Support Protocol 1 in the current unit provides a method for estimating resin capacity in the presence of debris from unclarified feedstock.
HYDROPHOBIC-INTERACTION EXPANDED-BED ADSORPTION CHROMATOGRAPHY
ALTERNATE PROTOCOL 2
Resins for hydrophobic-interaction EBA chromatography are chosen in the same way that resins are chosen for other forms of hydrophobic-interaction column chromatography— that is, based on what is known about the polarities of the target protein and all contaminants (UNIT 8.4). To determine the appropriate binding conditions for the target molecule in a hydrophobicinteraction application, perform the basic experiments described in UNIT 8.4, Support Protocol 1. Capacity determinations made using clarified feed should be treated only as approximations, since the capacity of the resin is impacted by the presence of debris. Support Protocol 1 in the current unit provides a method for estimating resin capacity in the presence of debris from unclarified feedstock.
AFFINITY EXPANDED-BED ADSORPTION CHROMATOGRAPHY Resins for affinity EBA chromatography are chosen in the same way that resins are chosen for other forms of affinity column chromatography—that is, based on what is known about the potential for specific interaction between the target protein and the ligand immobilized on the resin. There are only a few commercially available affinity ligands for affinity EBA chromatography. The protocols described in Chapter 9 also apply to initial methods development for affinity EBA processes. Support Protocol 1 in the current unit provides a method for estimating resin capacity in the presence of debris from unclarified feedstock.
ALTERNATE PROTOCOL 3
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COMMENTARY Background Information Principles of expanded-bed operation The expanded-bed adsorption cycle is a five-step operation (Fig. 8.8.9). The first step is the creation of a sedimented bed, which must have a height of ∼15 cm to allow sufficient contact time for efficient binding of the target molecule to the resin. The column is then equilibrated, with this equilibration serving two purposes. First, it allows formation of a stable, fluidized bed when the suspended adsorbent beads attain an equilibrium between particle sedimentation velocity and upward liquid flow velocity. (At equilibrium, the bed is expanded to 2.5 to 3 times its sedimented height.) Second, it allows equilibration of the adsorbent with the sample buffer. After equilibration, crude, unclarified feedstock is applied to the expanded bed in the same upward direction; target proteins in the feedstock bind to the resin, while cells, cell debris, particulates, and contaminants pass through unhindered. Next, weakly bound material, such as residual cells, cell debris, and other types of particulate material, is washed out of the expanded bed, and once all weakly retained material has been removed, the liquid flow is stopped, allowing the adsorbent beads to quickly settle in the column due to their high density. The column adapter is then lowered to the surface
Expanded Bed Adsorption Chromatography
of the sedimented bed, the flow is reversed, the buffer is changed, and the captured proteins are eluted. These eluted proteins are concentrated, clarified, partially purified, and ready for further purification by packed-bed chromatography. After elution, the sedimented resin bed is regenerated by washing with downward flow using buffers specific for the type of chromatographic principle in use. This process removes the more strongly bound proteins not removed during the elution phase. Finally, a cleaning-inplace procedure is performed to remove nonspecifically bound, precipitated, or denatured substances from the bed and to restore its original performance. During this phase, a moderate upward flow is used, with the column adaptor positioned at approximately twice the sedimented bed height. The purpose of cleaning in place is to clean the adsorbent between purification runs. A review of five cleaning-in-place protocols has been published (Amersham Biosciences; see Internet Resources). Four of the five case studies in this review deal with E. coli expression systems—namely, E. coli homogenate on a DEAE resin, E. coli homogenate on an SP resin, E. coli lysate on an SP resin, and E. coli inclusion bodies on an SP resin. The fifth case study deals with S. cerevisiae homogenate on an SP resin. These case
Figure 8.8.9 Resin bed status throughout the steps of an expanded-bed adsorption chromatography experiment. Arrows indicate direction of feed flow. Cleaning in place (not shown) is the last step and is done in expanded-bed mode, similar to expansion and equilibration of the adsorbent. Reproduced with permission from Amersham Biosciences (Expanded-Bed Adsorption: Principles and Methods; see Internet Resources) and Academic Press (Shiloach and Kennedy, 2000).
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studies describe media reuse for 10 to 50 cycles, which is typical. In principle, an expanded bed is similar to a fluidized bed; the major difference is that during operation, there is minimal mixing in an expanded bed, while a fluidized bed is turbulent. This means that an expanded bed permits a more efficient adsorption process than does a fluidized bed. The most efficient expanded beds are stable, classified fluidized beds. (A classified fluidized bed is one in which particles of similar sizes and densities are found together, as a class, at the same position in the column.) A review comparing the properties of expanded beds and fluidized beds can be found in Thommes (1997). The pressure drop over an expanded bed is small, and so the distribution system must build up a pressure drop to produce a plug flow (i.e., the condition in which axial mixing is minimal compared with convective flow).
In addition to generating this pressure drop, the distributor also directs flow in a strictly vertical path. This eliminates any radial flow that might propagate turbulence through the column. A specially designed liquid distribution system is needed because of the small pressure drop over the expanded bed as compared with a packed bed. During operation, the pressure drop in an expanded bed is typically <1 bar. The stability of the bed arises from two properties of the adsorbent: its particle size distribution and its particle density distribution. Of these two properties, the particle size distribution is the major factor influencing bed stability. Although the density distribution of the beads can be altered to achieve higher flow velocities, this distribution does not play as large a role in bed stability. Figure 8.8.10 illustrates the distribution of particle size and density in the bed. The
Figure 8.8.10 Results of sampling of an expanded-bed column in operation after bed height stabilization. Mean particle size (d50V ) and density increase going from top to bottom, as smaller, less dense particles sit near the top of the column. The sedimented resin height is ∼15 cm in a 5.0cm-diameter column. The ‘ml’ column represents sampling the top 30 ml, then the next 49 ml, then the next 72 ml, then the final 104 ml for each d50V and density measurement. Sampling volumes were chosen for convenience and do not account for spilling or loss of sample. Reproduced with permission from Amersham Biosciences (Expanded-Bed Adsorption: Principles and Methods; see Internet Resources).
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particle size polydispersity causes beads to be positioned at specific heights in the expanded bed. The smaller, lighter particles move to positions at the top of the expanded bed, and the larger, heavier particles move to the bottom, resulting in a stable expansion. In other words, the beads find their ideal positions in the column, leading to low axial dispersion in expanded-bed mode. If all beads were the same size, this spatial distribution could not be achieved. Instead, many beads would compete for the same position in the column, causing movement and destabilization of the bed. Increased bead density is necessary so that high flow velocities can be used in expandedbed mode.
Critical Parameters There are both physical and chemical parameters that affect EBA chromatography. The physical parameters that influence expansion are related to feedstock viscosity and include temperature, DNA content, salt content, and the amount of biomass in the feedstock. The more viscous the feedstock, the more the bed will expand at a given flow velocity. Another important physical parameter is bed stability, which is influenced by the amount of biomass present and the binding conditions for the target protein. Since biomass is always charged and always has some degree of hydrophobicity, there is always some amount of interaction between the resin and the feedstock. Chemical parameters, such as pH and conductivity, influence binding capacity in EBA chromatography in the same manner as they do in standard protein chromatography. However, these parameters also influence the charge density and hydrophobicity of the biomass. Method scouting in EBA chromatography must be very thorough, since all of these interactions need to be identified and controlled. When defining a suitable maximum loading capacity for EBA chromatography, it is necessary to apply a safety margin to compensate for sources of variability in the sample that could affect binding capacity. The maximum load should be ∼75% of breakthrough capacity (see Support Protocol 1).
Troubleshooting Expanded Bed Adsorption Chromatography
Expanded Bed Adsorption: Principles and Methods (Amersham Biosciences; see Internet Resources) is highly recommended reading and contains an exhaustive troubleshooting table for EBA chromatography procedures.
EBA chromatography with mammalian cell culture feedstocks Mammalian cell culture feedstock will contain DNA from lysed cells. The presence of DNA will increase the feed viscosity. Viscosity will also be increased if the feedstock contains serum. Cell densities of up to 2 × 107 cells/ml are acceptable for EBA chromatography, but cell aggregates can present a problem. Since cell density can be high at the harvest of a cell culture, mixing is required during application of the feedstock to the column. In this way, the column will encounter the same cell density throughout the application of feedstock. The typical expression level for a target protein is ∼200 µg/ml. This translates to high load volumes (typically 50 to 100 times the settled bed volume) and long loading times. For example, given an expression level of 200 µg/ml, 5 liters of culture, or ∼66.6 times the volume of a 2.5-cm-diameter column with a bed height of 15 cm, will be required to isolate 1 gram of product. At a flow rate of 300 cm/hr, it will take 3.3 hr to load this volume of culture. Lowering the pH of a cell culture will force the cells into apoptosis and will lead to cell aggregation and lysis. If the cell culture feedstock is diluted, cells may undergo lysis due to osmotic shock, and so it is recommended that 200 mM glucose be added to the diluent to maintain feed osmolality. A strategy for diluting cell culture feedstock is illustrated in Figure 8.8.11. Adding salt to a cell culture feedstock will cause cells to shrink, which can result in aggregation and lysis as well, since cell surfaces are charged. When working with mammalian cell culture feedstocks, the same degree of expansion observed using the equilibration buffer can be expected. Stored feedstock should never be used, as cells that have been stored are likely to have lysed or formed aggregates. Feedstock should always be mixed during application to the column to prevent aggregation. The most successful EBA chromatography applications using mammalian cell culture feedstocks have involved protein A–based methods, while the least successful applications have involved anion-exchange methods. EBA chromatography with Pichia pastoris feedstocks The use of the Pichia expression system for commercial purposes requires license agreements and royalty payments. Pichia fermentations yield very high cell densities, up to 500 g cell wet weight per liter. Expression levels are
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Figure 8.8.11 Schematic diagram showing on-line dilution of mammalian cell culture feedstock for application to an expanded-bed adsorption column. Dilution is necessary when the conductivity of the culture medium is so high that it prevents adsorption of the product to the resin (e.g., in cation-exchange experiments using the SP STREAMLINE resin). The dilution mixture (feedstock plus diluent) must be continuously mixed during this procedure. Reproduced with permission from Amersham Biosciences (Expanded-Bed Adsorption: Principles and Methods; see Internet Resources).
also very high; levels of ∼1 mg/ml are typically observed, but levels as high as 10 mg/ml can be achieved. The use of Pichia feedstock will always require dilution to reduce the biomass concentration to an acceptable level for EBA. Biomass from Pichia is known to accumulate under the bottom nets of the EBA columns. Since the feed must be diluted 2- to 5-fold, there is the potential for long loading times due to the resulting large volume. Salts from the fermentation can precipitate and clog the nets in the EBA column. If this occurs, coarse filtration (through a 50-µm filter) is necessary before EBA. Pichia expression systems usually produce secreted recombinant protein. Pichia host proteins are very acidic, and so capture on an SP resin at pH >5 is very selective. The most successful applications have employed an SP resin at such pH levels. EBA chromatography with E. coli feedstocks E. coli feedstock is used in three scenarios: one in which the recombinant protein is soluble and is secreted into the fermentation broth, another in which the recombinant protein is secreted into the cell periplasm, and a third in which the recombinant protein is insoluble inside the cell and exists as an inclusion body. Recombinant proteins secreted directly into the broth can be harvested directly by EBA. In contrast, cells must be partially
broken to release product from the periplasm and completely homogenized to release inclusion bodies. When working with E. coli homogenates, at least two passes through the homogenizer (at 8000 psi) must be performed to shear all DNA present. Conditioning of the homogenate at pH <6 will cause precipitation of cellular components, and this will lead to clogging of the EBA columns. E. coli homogenates must be fresh, because their viscosity increases with time. Load volumes will be on the order of 10 to 20 settled bed volumes. The most successful applications using E. coli feedstock have involved solubilized inclusion bodies. Unsuccessful applications have involved the use of anion-exchange EBA with E. coli homogenates. However, anionexchange EBA chromatography can be performed successfully using whole or leaky cells. Successful cation-exchange EBA using E. coli homogenates at low pH is not possible.
Anticipated Results The typical chromatogram from an EBA chromatography run is unremarkable. There will be a large flowthrough fraction and a smaller fraction containing the target protein (Fig. 8.8.12). The second fraction will be free of debris and suitable for application to the next column in the purification process, but it may also be filtered through a 0.45-µm filter before application to the next column.
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Figure 8.8.12 Example of an expanded-bed adsorption chromatogram (bold line) showing feed flowthrough, product elution, and cleaning in place (CIP). Also shown are the results of an enzyme-linked immunosorbent assay for product breakthrough (C/Co ; thin line; horizontal axis label applies to this plot only) as a function of load volume. Separation was performed using the STREAMLINE SP resin (0.30 liters; sedimented bed height, 15 cm) in a STREAMLINE 50 column (i.d., 50 mm). E. coli homogenate containing a recombinant Fab fragment was used as the feedstock; 50 mM sodium acetate, pH 5.0, was used as buffer A (i.e., the wash buffer); 1 M NaCl/50 mM sodium acetate, pH 5.0, was used as buffer B (i.e., the elution buffer); and 1 M NaCl/0.5 M NaOH was used as the CIP solution. AU, arbitrary units (for absorbance measured at 280 nm); C/C0 , ratio of target concentration in flowthrough (C) to target concentration in feedstock (Co ). Reproduced with permission from Amersham Biosciences (Expanded-Bed Adsorption: Principles and Methods; see Internet Resources).
Time Considerations
Literature Cited
A thorough set of method scouting experiments can take a week. It is reasonable to expect that as the volume of feed increases, the time needed to load the column will increase as well. Thus, long loading times are to be expected, and this aspect must be compared with the time it takes to clarify the feedstock by filtration or centrifugation prior to the start of packed-bed chromatography. Beginners are cautioned not to automate the EBA process until it is well understood and the specific methodology established. By using a manual system for a number of experiments, the user will be forced to observe the process and react to problems as they occur. Once the process is well characterized, the EBA process can be automated.
Batt, B.V., Yabannavar, V.M., and Singh, S. 1995. Expanded bed adsorption process for protein recovery from whole mammalian cell culture broth. Bioseparation 5:42-53.
Acknowledgement Expanded Bed Adsorption Chromatography
The authors thank Alisa Liten and Kyle Kinealy for allowing their unpublished work to be cited in this unit.
Chase, H.A. 1994. Purification of proteins by adsorption chromatography in expanded beds. Trends Biotechnol. 12:296-303. Hjorth, R. 1997. Expanded bed adsorption in industrial bioprocessing: Recent developments. Trends Biotechnol. 15:230-235. Kula, M.R., Lin, D.Q., Knieps, E., Reichert, U., and Thommes, J. 2001. Process design in expanded bed adsorption: Integration of biomass influence into optimizing operation conditions. Presented at Recovery of Biological Products X, Cancun, Mexico. Shiloach, J. and Kennedy, R.M. 2000. Expanded bed adsorption process for protein capture. In Handbook of Bioseparations (S. Ahuja, ed.) pp. 431-451. Academic Press, New York. Sofer, G. and Hagel, L. 1997. Handbook of Process Chromatography: A Guide to Optimization, Scale-Up and Validation. Academic Press, New York.
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Thommes, J. 1997. Fluidized bed adsorption as a primary recovery step in protein purification. Adv. Biochem. Eng. Biotechnol. 58:185-230.
Internet Resources http://www.bo-conf.com Contains program information from the biennial international congress on EBA. Manuscripts from the second congress (1998) have been collected and published in Volume 8 of the journal Bioseparation (pp. 1-267). Manuscripts from the third congress (2000) have been collected and published in Volume 10 of Bioseparation (pp. 1-132). http://www.chromatography.amershambiosciences. com Collection of various chromatography resources published by Amersham Biosciences.
“STREAMLINE SP STREAMLINE DEAE” (publication no. 18-1115-27) provides a review of cleaning-in-place protocols for the STREAMLINE SP and STREAMLINE DEAE resins. “STREAMLINE Reference List” (publication no. 18-1162-07) contains references to 114 papers on EBA chromatography (includes papers published through 2002). “Expanded Bed Adsorption: Principles and Methods” (publication no. 18-1124-26) details the underlying principles of EBA chromatography and the practical considerations pertaining to the use of Amersham Biosciences STREAMLINE adsorbents and columns.
Contributed by Robert M. Kennedy GE Healthcare Piscataway, New Jersey
Conventional Chromatographic Separations
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CHAPTER 9 Affinity Purification INTRODUCTION ffinity chromatography in the context of this chapter refers to a variety of techniques that exploit the fact that proteins can act as ligands for a variety of ligand-binding proteins (e.g., antibodies, enzymes, and lectins) or are themselves ligand-binding proteins (i.e., act as receptors). Because receptor-ligand interactions are very specific and receptor and/or ligand sites are often unique to the protein of interest, exploitation of this methodology often yields pure or nearly pure proteins with a minimal number of purification steps.
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Some receptor or ligand sites, such as antigenic epitopes and the active sites of enzymes, are limited to the protein of interest and, once discerned, provide a powerful approach for purification of the protein. Other receptor or ligand sites, such as those with the ability to interact with certain dyes or lectins, respectively, are less specific, but nevertheless provide approaches for isolating particular proteins from complex mixtures of proteins. Receptor and/or ligand-binding sites present in a protein, such as reactivity with a particular substrate or monoclonal antibody, are often known prior to the initial biochemical characterization of the protein. This knowledge saves time and facilitates development of the purification procedure. In other cases, such as for dye binding or lectin reactivity, it is often necessary to perform pilot studies to determine the feasibility of using a particular affinity chromatography method. Because each protein contains multiple receptor and/or ligand-binding sites, the combination of several different affinity chromatography steps in a purification scheme can often be utilized to obtain pure proteins.
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Receptor and ligand-binding sites are usually natural properties of proteins; however, it has become popular in the case of recombinant proteins to engineer particular binding sites, such as metal-binding sites, as extensions from either the N- or C-terminus into the recombinant protein of interest. Such alterations of the natural protein greatly facilitate their purification. If structural alteration of the recombinant protein by the ligand-binding site is undesirable, protease cleavage sites can be positioned adjacent to the ligand-binding site so that this protein extension can be removed after it has been used for affinity purification (e.g., as when a polyhistidine Ni2+ binding site is removed by thrombin cleavage).
Essentially all forms of affinity chromatography involve the following steps: 1. Choice of appropriate ligand. 2. Immobilization of the ligand onto a support matrix. 3. Loading of the protein mixture of interest onto the matrix. 4. Removal of nonspecifically bound proteins. 5. Elution of the protein of interest in a purified form. This chapter presents an eclectic collection of affinity purification protocols that have proven extremely useful in protein purification.
Lectin affinity chromatography, described in UNIT 9.1, is frequently utilized in glycoprotein purification schemes. In this approach a protein is bound to an immobilized lectin Contributed by John E. Coligan Current Protocols in Protein Science (2004) 9.0.1-9.0.3 C 2004 by John Wiley & Sons, Inc. Copyright
Affinity Purification
9.0.1 Supplement 37
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through its sugar chain, unbound protein is washed away, and the bound protein is eluted. Sugar side chains present in glycoproteins are usually of sufficient complexity that they present the possibility of reacting with a number of different lectins in a fashion useful for purification. Suitable lectins are tabulated in this unit. However, the potential resolving power of this large battery of lectins is limited by the facts that (1) most or many glycoproteins in a crude mixture of proteins share identical or near-identical sugar side chains and (2) lectin specificities are often not absolute. Nonetheless, when appropriately applied, lectin affinity purification usually yields a significant enrichment of the glycoprotein of interest. The basic and alternate protocols describe the use of concanavalin A (Con A) or wheat germ agglutinin (WGA) for preparative glycoprotein purification. These lectins are commonly used because of their general applicability and availability. Others can be readily substituted using the procedures described.
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Dye affinity chromatography, described in UNIT 9.2, is a protein purification procedure based on the high affinity of immobilized dyes for binding sites on many proteins. It is a rapid and versatile method that is especially applicable for obtaining 10- to 100-fold purification of the desired protein from crude extracts. The dye ligand can be selected either to remove unwanted proteins (negative chromatography) or to bind to the protein of interest (positive chromatography); these two types of chromatography can also be coupled in series (tandem chromatography). Support protocols are provided to guide selection of appropriate dyes and support matrices.
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Affinity purification of proteins can also be accomplished by their interaction with natural ligands, as described in UNIT 9.3. These ligands may be macromolecules such as proteins and nucleic acids, or low-molecular-weight compounds such as substrates, inhibitors, cofactors, etc. Because such interactions tend to be specific for particular proteins or groups of proteins, high degrees of purification can be obtained in a single step. Because of this specificity, the protocols in UNIT 9.3 do not detail a specific purification scheme, but deal instead with choices of column matrices and methods of coupling potential ligands to these matrices.
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UNIT 9.4 describes a method for the facile purification of recombinant proteins by metalchelate affinity chromatography (MCAC). Proteins engineered to have six consecutive histidine residues on either the amino or carboxyl terminus can be purified in native or denatured forms using a resin containing nickel ions (Ni2+ ). Techniques for creating a fusion protein consisting of the protein of interest with a histidine tail are outlined in a Strategic Planning section. The protocols describe expression of the histidine-tailed fusion proteins and their purification in native form by MCAC, as well as the purification of the fusion protein under denaturing conditions and the subsequent renaturation procedure (by either dialysis or solid-phase renaturation). In addition, analysis of the purified product and regeneration of the resin is presented.
Introduction
Proteins can be used as immunogens to raise antibodies against them that have exquisite specificity. The resulting antisera can be used as analytical probes to detect the presence of proteins of interest in various biological preparations. Antisera and its component immunoglobulin fraction can have undesirable antibody reactivities that show up as “background” in analytical assays. This “background” is due to impurities in the protein immunogen or preexisting antibodies in the sera of the immunized animals. The latter can be controlled for by comparing the reactivity of preimmune sera to that of the immune sera. The complications of undesirable reactivities often present in antisera can be obviated by preparing monoclonal antibodies reactive with the protein of interest. UNIT 9.8 describes the use of antibodies as analytical probes to detect the presence of proteins in biological preparations. Basic Protocol 1 and Alternate Protocols 1 through 4 describe techniques for solubilizing cell associated proteins for antibody detection. These are followed by protocols that describe how to immobilize the antibody on a solid-phase
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matrix for use in analytical immunoprecipitation. Imunoprecipitated proteins can be dissociated from antibodies and reprecipitated by a protocol known as immunoprecipitation recapture (see Basic Protocol 2). This procedure can be used with the initial precipitating antibody for further purification of the reactive protein, or with a second antibody to identify components of multisubunit complexes or to study protein-protein interactions.
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UNIT 9.5 describes the isolation of soluble or membrane-bound protein antigens by immunoaffinity chromatography. By this technique, purifications of 10,000-fold or more can often be achieved in one step. Immunoaffinity columns most often employ monoclonal antibodies covalently attached to a solid-phase matrix; polyclonal antibodies have been used successfully, but they usually lack the specificity required for a single-step protein purification. The amount of protein that can be purified is solely dependent on the amount and affinity of the antibody employed. In addition to describing the overall procedure, this unit provides alternate conditions of elution and a method for attaching the antibody to the support matrix. A variation of this procedure is to purify antigens of interest by immunoprecipitation as described in UNIT 9.8. As protein antigens of interest often exist within the complex milieu of cells, several protocols are given for solubilizing such molecules. Precipitation of the antigen of interest is achieved by precipitating immunocomplexes with Protein A or G containing beads as described in Alternate Protocol 5 or with anti-Ig as described in Alternate Protocol 6. Basic Protocol 2 describes immunoprecipitation-recapture; a method to determine if the initially precipitated antigen can be reprecipitated by a second antibody. The primary antibody can be used a second time to confirm identity or achieve further purity, or a different antibody can be used to, for example, investigate subunit composition of the protein antigen.
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Many biological processes, such as recombination, replication, and transcription, involve the action of sequence-specific DNA-binding proteins. Affinity chromatography (as described in UNIT 9.6) is a very effective means of purifying a protein based on its sequencespecific DNA-binding properties, and is relatively straightforward if proper care is taken. The affinity chromatography procedure that is described in this unit uses DNA encoding specific recognition sites for the desired protein that has been covalently linked to a solid support. Researchers in the past have purified proteins based on the proteins’ ability to bind DNA, only to be disappointed to find that the DNA-binding activity of their highly purified products was not sequence-specific. This situation can be avoided by carefully performing all of the preliminary steps and experimental controls described in this unit. Figure 9.7.1 provides a summary diagram of the entire chromatography procedure. With this procedure, a high degree of purification can be achieved using a DNA affinity column.
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A similar degree of purification can be accomplished using biotinylated DNA fragments that contain the desired protein binding site and streptavidin-coated column matrices (UNIT 9.7). This purification scheme, outlined in Figure 9.7.1, utilizes the tight and nearly irreversible binding that occurs between biotin and streptavidin to fish out specific DNA binding proteins from crude cellular or nuclear lysates. After washing, the protein of interest is eluted with high-salt buffer. This protocol is relatively easy to optimize by following the success of the isolation using gel mobility-shift assays; it is also economical, as a single type of column matrix is compatible with any biotinylated DNA probe.
In summary, affinity chromatography at its best is the most powerful technique for protein purification available, because its high selectivity can, in principle, permit purification of a desired protein from a crude mixture of proteins in a single step. Moreover, the technique usually offers simultaneous concentration of the desired protein. John E. Coligan
Affinity Purification
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Lectin Affinity Chromatography
UNIT 9.1
Lectin affinity chromatography is the most frequently used specific purification procedure for glycoproteins. A protein is bound to an immobilized lectin through its sugar chain, the unbound protein is washed away, and the bound protein is eluted. Numerous plant lectins have been identified and in many cases their sugar specificity is known. This sugar specificity is the basis for the separation and structural analysis of many individual oligosaccharides and glycopeptides (Chapter 12) and for the identification of lectin-binding glycoproteins. Glycoproteins often contain multiple chains of a given sugar type; the resulting multiple interactions with the immobilized lectin may make it difficult to elute some of the proteins from the lectin column. The presence of multiple low-affinity sugar chains can still produce substantial interaction with a lectin-containing column even though individual sugar chains may have a low affinity. Thus, only limited information on the structure of a sugar chain can be obtained from the binding of a protein to a lectin column. These qualifications by no means belittle the usefulness of lectin affinity chromatography for glycoprotein purification; lectin affinity chromatography may be used to partially purify glycoproteins and to provide limited qualitative information about the nature of their carbohydrate components. Other preparative purification steps, such as ammonium salt precipitation and gel-filtration and ion-exchange chromatographies (UNIT 8.2 & UNIT 8.3), are usually done prior to using lectins, but this is not necessary. Many different lectins are commercially available in an immobilized form suitable for glycoprotein purification, but some can be rather expensive. Con A–Sepharose is the most commonly used lectin for glycoprotein purification. It is relatively inexpensive, stable, and able to bind to many different glycans. Bound proteins can be eluted with α-methylD-mannoside (αMM). Wheat germ agglutinin (WGA) is the next most popular lectin for the same reasons although it is somewhat more expensive. Many other lectins can be used to purify glycoproteins. A broad range of immobilized lectins is available from E-Y Labs, Vector, Sigma, and Pharmacia Biotech (SUPPLIERS APPENDIX). Most are also available in free form, if desired. However, this array of lectins is never used for routine protein purification. First, their higher price, shorter track records, and, in some cases, requirement for exotic sugars to elute bound proteins make them less popular for routine use. Second, the time and cost of screening and optimizing conditions for so many lectins would be prohibitive. Third, many have very similar specificities. Fourth, even using a battery of lectins to purify a protein would not yield a single species, since other proteins with similar sugar chains would copurify. Although these other lectins are less frequently used, the principles of Con A–Sepharose and WGA–agarose lectin affinity chromatography apply to them as well. The names, sugar specificity, elution conditions, and general types of sugar chains that bind to 25 different lectins are given in Table 9.1.1. This table is not comprehensive (there are at least twice as many lectins commercially available), but those listed are the most commonly used. The list is intended as a guide to possible alternatives, and an attempt has been made to group lectins of similar specificities together. However, specificities are not always hard and fast, and the comments in the table are not always sacrosanct. Some lectin–ligand binding requirements are well characterized, but many of the “rules” for lectin sugar specificity are based on screening simple sugars to inhibit lectin binding. These competitors can mimic only limited features of the binding of a lectin to its natural ligand. For instance, both pea and lentil lectins bind only sugar chains with α-fucose residues in the core region of N-linked chains, and yet mannose, not fucose, is used to elute bound proteins. Cummings (1994) provides an excellent perspective on the use of lectins. Contributed by Hudson H. Freeze Current Protocols in Protein Science (1995) 9.1.1-9.1.9 Copyright © 2000 by John Wiley & Sons, Inc.
Affinity Purification
9.1.1 CPPS
Table 9.1.1
Lectins Useful for Glycoprotein Purification
Acronym (organism and source)
Metal ions required
Con A (Canavalia ensiformis; jack bean seeds)
Ca2+, Mn2+ α-Man > α-Glc
0.1–0.5 M α-MeMan
High-Man, hybrid, and biantennary N-linked chains
LCA or LCH (Lens culinarus; lentil seeds)
Ca2+, Mn2+ α-Man > α-Glc
0.1–0.5 M α-MeMan
Bi- and triantennary N-linked chains with Fucα1-6 in core region
0.1–0.5 M α-MeMan
Similar to LCA/LCH
0.1–0.5 M GlcNAc
GlcNAc- and Sia-terminated chains, or clusters of O-GlcNAc; succinylated form selectively binds GlcNAc>Sia
0.1–0.5 M Gal or lactose
—
β-Gal β-GalNAc β-Gal
Galβ1-4 on N-linked chains; can also bind β-GalNAc β-GalNAc-terminated chains.
—
GalNAc
0.1–0.5 M GalNAc
—
GalNAc
0.1–0.5 M GalNAc
Sugar specificityb Elution conditions
PSA (Pisum sativum; Ca2+, Mn2+ α-Man peas) WGA (Triticum Ca2+, Mn2+ β-GlcNAc vulgaris; wheat germ)
RCA (Ricinus communis; castor bean) RCA-Ia RCA-IIa PHA (Phaseolus vulgaris; red kidney bean) E4PHA L4PHA GSL-1, BSL-1 [Griffonia (Bandeiraea); Simplicifolia seeds]
—
Mg2+, Ca2+ α-Gal α-GalNac
0.1–0.5 M Gal
Useful for binding
Gal-terminated N-linked chains with “bisecting” GlcNAcβ1-4 Tri- and tetrabranched N-linked chains
0.1–0.5 M melibiose
Chains terminated by α-Gal residues
Proteins with poly-N-acetyllactosaminecontaining chains; highly heterodispersed glycoproteins often contain variable lengths of this repeat Similar to DSA; prefers longer poly-N-acetyllactosamines than DSA Proteins with poly-N-acetyllactosamines
DSA (Datura stramonium; Jimsonweed seeds)
—
Galβ1-4GlcNAc
0.1–0.5 M GlcNAc or 20 mM N,N′,N′′chitotriose
LEL (Lycopersicon esculentum; tomato meat) STL (Solanum tuberosum; potato tubers) MAL (Maackia amurensis; seeds)
—
β-GlcNAc
—
β-GlcNAc
—
Siaα2-3Galβ1-4 GlcNAc
0.1–0.5 M β-GlcNAc or 20 mM chitobiose or chitotriose 0.1–0.5 M GlcNAc or 20 mM chitobiose or chitotriose 0.5 M lactose
α2-3-linked Sia to Gal in N-linked sugar chains; underlying sugars and peptide contribute to binding continued
Lectin Affinity Chromatography
9.1.2 Current Protocols in Protein Science
Table 9.1.1
Lectins Useful for Glycoprotein Purification, continued
Acronym (organism and source)
Metal ions required
Sugar specificityb Elution conditions
Useful for binding
EBL or SNA (Sambucus nigra; elderberry bark) LFA (Limax flavus; garden slug) GNL (Galanthus nivalis; snowdrop bulbs)
—
Siaα2-6Gal or GalNAc
0.1–0.5 lactose
α2-6-linked Sia in N and O sugar chains
—
Neu5Ac
10 mM Sia
—
α1-3Man
0.5 M α-MeMan
Sia-terminated chains irrespective of linkage High-Man-type chains, but not to Glc
UEA-I (Ulex europaeus; furze gorse seeds)
—
α-L-Fuc
0.1–0.5 M L-Fuc or methyl-α-L-Fuc
Sugar chains with terminal α-Fuc, especially in α1-2 linkage, but much less with α1-3 or α1-6 linkages
AAA (Anguilla anguilla; freshwater eel)
—
α-L-Fuc
0.1–0.5 M L-Fuc
Similar to UEA-I but more broadly binds any fucosylated oligosaccharide
Lotus (Lotus tetragonolobus; asparagus pea)
—
α-L-Fuc
0.1–0.5 M L-Fuc
Outer-branch α-Fuc residues; does not bind when Fuc is linked to chitobiose core of N-linked chains
HPA (Helix promatia; albumin gland of edible snail)
—
α-GalNAc
0.1–0.5 M GalNAc
Proteins with terminal α-GalNAc or GalNAcα-O-Ser/Thr (Tn antigen)
Jackalin (Artocarpus integrifolia; jackfruit seeds)
—
α-Gal
0.1–0.2 M melibiose
VVL or VVA (Vicia villosa; hairy vetch seeds) SBA (Glycine max; soybean)
—
GalNAc
0.1–0.5 M GalNAc
Proteins with only a single O-linked chain with T antigen Galβ1-3GalNAcα-O-Ser/Thr can bind; not inhibited by lactose; Sia in chain does not interfere with binding Proteins with Tn antigen, GalNAcα-O-Ser/Thr
—
GalNAc or Gal
0.1–0.5 M GalNAc
α- or β-linked GalNAc
PNA (Arachis hypogaea; peanut)
—
Galβ1-3GalNAc
0.1–0.5 M lactose
Proteins with T antigen, but does not bind if sugar chain is sialylated
DBA (Dolicholos biflorus; horse gram seeds)
—
Terminal α-GalNAc
0.1–0.5 M GalNAc
Terminal α-GalNAc (specific for blood group A), not those in the core of O-linked chains
0.1–0.5 M GalNAc
Proteins with blood group A structure GalNAcα1-3(Fucα12)Gal–
LBA (Phaseolus lunatus; lima bean)
Mn2+, Ca2+ Terminal αGalNAc
aCAUTION: Both RCA-I and RCA-II are extremely toxic, with lethal doses of 1 molecule/cell. bAbbreviations: Fuc, L-fucos; Gal, D-galactose; GalNAc, N-acetyl-D-galactosamine; Glc, D-glucose; GlcNAc, N-acetyl-D-glucosamine; Man, Dmannose; Neu5Ac, N-acetyl-D-neuraminic acid; Sia, sialic acid.
Affinity Purification
9.1.3 Current Protocols in Protein Science
In general, these immobilized lectins can be used in two ways. First is as an affinity step for routine purification. If a novel protein may be glycosylated, using several lectins that bind very different types of sugar structures (N-linked vs. O-linked) will probably give reasonable purification. It will also give some indication of the type of sugar chains on the protein. Both Vector and E-Y Labs sell kits with six or seven different lectins for this purpose. The second and more systematic approach is useful for better-characterized glycoproteins. Here, the lectins are chosen to resolve potentially different glycoforms that may have different biological activities. An excellent example of this second type of analysis is given by Li and Jourdian (1991). The Basic and Alternate Protocols describe the use of lectins for preparative glycoprotein purification. Con A–Sepharose and WGA–agarose were chosen for convenience and availability. The Support Protocol describes a small-scale pilot procedure to test for lectin binding and to determine elution conditions. There are many variations on the basic procedure in the literature, but all use the same principles: bind the protein to immobilized lectin through its sugar chain, wash away unbound protein, and elute bound protein with a simple sugar that resembles the sugar ligand of the bound protein. Because many proteins have sugar chains that can bind to a specific lectin, this procedure seldom yields a pure protein. BASIC PROTOCOL
CON A–SEPHAROSE AFFINITY CHROMATOGRAPHY Con A–Sepharose chromatography is used to partially purify glycoproteins that contain terminal mannose or glucose residues. The steps presented below give typical conditions for this type of chromatography, although a variety of approaches have been utilized. In this protocol, bound glycoproteins are eluted with α-methyl-D-mannoside (αMM) after the column is first washed to remove unbound and weakly bound proteins. Before proceeding, it is advisable to conduct a pilot study to test the protein of interest for lectin binding and elution conditions (see Support Protocol). In order to purify the protein effectively, a specific assay for detecting the protein of interest (i.e., activity or antibodybinding assay) must be available. Materials 10 mg/ml Con A–Sepharose (Pharmacia Biotech or Sigma) Column buffer (see recipe) 0.5 M αMM in column buffer Protein sample in column buffer 1.5 × 30–cm glass or disposable chromatographic column Glass wool NOTE: Carry out this procedure at room temperature if the protein to be isolated will tolerate this condition. If not, carry it out in a cold room, and prechill all solutions to maintain temperature. 1. Gently resuspend 50 ml settled Con A–Sepharose (10 mg lectin/ml packed resin) in 50 ml column buffer to make a slurry. Degas the slurry.
Lectin Affinity Chromatography
This volume of lectin beads should be sufficient to bind ∼100 mg glycoprotein. If a 1.5 × 30–cm column is not available, either the dimensions of the column or the total amount of resin can be modified. A 1.0 × 30–cm column will bind ∼50 mg total glycoprotein. If the amount of glycoprotein in the sample is substantially less than 50 to 100 mg, decrease the volume of the column accordingly. The ratio of input protein to lectin does not seem to matter as long as the column is not overloaded.
9.1.4 Current Protocols in Protein Science
2. Pack a glass wool plug over a sintered glass or polypropylene frit at the bottom of the column and pour the degassed slurry into the column. The glass wool prevents the beads from clogging the frit and slowing the flow rate. Alternatively, BioGel P2 or Sephadex G10 can be used in place of the glass wool.
3. Continue packing the column until the desired level is reached (for a 50-ml volume this is ∼28 cm). Wash the gel with 2 to 3 column volumes of column buffer to remove any loosely bound or degraded Con A. 4. Wash with 2 to 3 column volumes of 0.5 M αMM in column buffer or the highest concentration of αMM that will be used. 5. Wash the column with >5 column volumes of column buffer without αMM to reequilibrate. It is important to prewash the column with the eluting sugar and then to reequilibrate the column to remove any materials that may have previously bound to the column. Check completeness of washing by phenol–sulfuric acid assay (Manzi, 1993). Estimate the amount of residual sugar using αMM as a standard. An acceptable level is <0.1 mM or ∼20 µg/ml.
6. Slowly load the protein sample on the column to permit binding without disturbing the surface. A flow rate of ∼1 ml/min (∼0.5 ml/min/cm2 of area) is desirable.
7. Wash the column with column buffer and monitor the flowthrough and subsequent wash fractions by measuring the A280 until it approaches baseline value. If the A280 does not rapidly return to baseline, it may indicate weak interaction of some proteins with the lectin, or overloading of the column.
8. Using an appropriate specific assay, check flowthrough and wash fractions for the presence of the protein of interest (e.g., enzyme activity or antibody binding). To check the lectin-binding properties, see Support Protocol. To determine whether the column was overloaded, add a small amount of the flowthrough (1%) to a small volume of beads as described in the Support Protocol. If the sample is bound to the beads as determined by the appropriate assay, overloading of the original column is indicated. In this case, reapply the flowthrough with excess glycoprotein to the larger column after the bound proteins of the first run have been eluted with αMM and the column reequilibrated.
9. Elute the column with 0.5 M αMM in column buffer. Monitor fractions for A280 and for activity (using specific assay) and pool peak-activity fractions. Very broad peaks may result during elution with the sugar if the protein dissociates very slowly. When this happens, sample recovery can be improved by filling the column with 0.5 M αMM in column buffer and allowing the column to stand for a few hours. This allows dissociation of the bound material; when the flow is started again, it should give a very sharp peak. Other possible remedies include warming a cold column to room temperature in the presence of αMM, increasing the concentration of αMM to 1 M, or increasing the NaCl concentration to 1 M. A combination of these approaches may be needed to elute a tightly bound protein.
10. Regenerate the column by washing it with 10 vol column buffer or until the αMM concentration is <20 µg/ml. The column may now be used for another run or stored indefinitely at 4°C in column buffer containing 0.02% (w/v) NaN3. Reequilibrate in column buffer before using again. Affinity Purification
9.1.5 Current Protocols in Protein Science
SUPPORT PROTOCOL
PILOT STUDY TO DETERMINE LECTIN BINDING AND ELUTION CONDITIONS If the target protein can be detected easily, test a small sample to establish binding and elution conditions before applying the entire sample to a large column. The easiest approach is to mix a small amount of sample with a measured amount of Con A–Sepharose beads in a series of microcentrifuge tubes. If the material in the sample binds, elute it from the washed beads with various amounts of competing αMM to determine when the activated material is eluted into the supernatant. This shows whether binding occurs and what concentration of competing sugar (or other conditions) will be required to elute it from the column. An abbreviated version of this procedure can be done with a single sample to determine if the protein binds and if it can be eluted with only one concentration (0.75 M) of αMM. It is important to keep the temperature constant, because changes can affect binding and dissociation of the ligands. Additional Materials (also see Basic Protocol) Sepharose 4B (Pharmacia Biotech) or other beaded gel to fill space in the tubes αMM: 0, 0.1, 0.2, 0.4, 0.8, 1.0, and 1.5 M concentrations, in column buffer 1. Prepare a 50% slurry of Con A–Sepharose in column buffer. Cut ∼2 to 3 mm off the small end of a 1000-µl pipet tip. Resuspend the slurry immediately before each pipetting, and use the truncated tip to dispense 200 µl of slurry into a series of seven microcentrifuge tubes. Add an equal volume of 50% slurry of Sepharose 4B to one tube (to serve as no-lectin control). Allow gel to settle to see if the dispensing was reasonably accurate. Gel material is needed in the controls to account for the volume changes caused by adding the gel. Other nonionic beaded gels, such as Sephadex (Pharmacia Biotech) or Bio-Gel (Bio-Rad) may be substituted.
2. Add the protein sample in 50 µl column buffer and allow 15 min for binding (shake the tube occasionally). Microcentrifuge the gel beads 1 min at 1000 × g to sediment. Remove the supernatant and save for analysis (step 6). 3. Use an appropriate assay specific for the protein of interest to determine if it binds to Con A–Sepharose and not to control beads. 4. Wash gel beads by adding 1.4 ml column buffer, microcentrifuging 1 min at 1000 × g, and removing supernatant (save for analysis). Repeat wash twice more. 5. Resuspend gel beads in 200 µl of buffer containing 0, 0.1, 0.2, 0.4, 0.8, 1.0, or 1.5 M αMM for each of the seven tubes containing Con A–Sepharose. Add the same volume of column buffer or 1.5 M αMM in column buffer to the control. 6. Incubate 15 min and centrifuge the gel beads 1 min at 1000 × g. Remove supernatant. Analyze the supernatant (along with those from steps 2 and 4) using the appropriate detection assay to determine if the protein has been eluted and at what concentration of competing sugar this occurred. Increasing the concentration of αMM should elute the target protein from the Con A–Sepharose. If it is not eluted, longer incubation (10 hr) in the presence of αMM or higher temperature may be needed. Although the entire procedure can be done in the cold, binding is tighter at low temperature, and this may make elution more difficult. Once the conditions are established for a particular sample or a lectin, scaling up the preparation for the basic protocol is normally routine. Lectin Affinity Chromatography
9.1.6 Current Protocols in Protein Science
WHEAT GERM AGGLUTININ (WGA)–AGAROSE AFFINITY CHROMATOGRAPHY
ALTERNATE PROTOCOL
WGA-agarose chromatography is used to purify proteins that contain terminal N-acetylD-glucosamine (GlcNAc) or sialic acid residues. A protein sample is applied to the column, the column is washed to remove unbound and weakly bound proteins, and bound glycoproteins are eluted with GlcNAc. The advice for purification of glycoproteins on Con A–Sepharose columns applies to columns containing immobilized WGA. Test the binding of the target protein to the lectin using a pilot study like that for Con A–Sepharose (see Support Protocol) before running a large column, substituting GlcNAc for the αMM. Additional Materials (also see Basic Protocol) 5 mg/ml wheat germ agglutinin (WGA)–agarose (E-Y Labs, Pharmacia Biotech, or Sigma) PBS (APPENDIX 2E) 0.1 M N-acetyl-D-glucosamine (GlcNAc) in PBS Protein sample in PBS 1.0 × 10–cm glass or disposable column (or other column whose length is 10 times its diameter) 1. Resuspend 1 vol WGA–agarose in 2 to 3 vol PBS to make a slurry. One milliliter of gel should be enough to bind 1 to 2 mg glycoprotein. To be safe, assume ∼5% to 10% of total cell lysate can bind. If target protein requires detergent to solubilize it, WGA is still active in 25 mM Tris⋅Cl (pH 7.5)/1% Lubrol PX/0.1% sodium deoxycholate. Triton X-100 or Nonidet P-40 (∼2% to 3% v/v final) may also be used.
2. Place a glass wool plug over the frit at the bottom of a column whose length is 10 times its diameter. Pour the slurry into the column. Wash the column with ∼2 column volumes of PBS to remove any unbound or degraded WGA, then wash with 1 column volume PBS containing 0.1 M GlcNAc. Finally, wash with 5 column volumes PBS. The length of the column should be ∼10 times its diameter so that weakly bound proteins can be eluted by prolonged PBS wash in the absence of any competing sugar. This will separate them from proteins that do not interact, and differs from using Con A–Sepharose, where continued buffer wash in the absence of αMM does not usually elute weakly bound protein. The choice of these column dimensions for WGA and for most other lectins is based on the successful fractionation of individual glycopeptides by serial lectin affinity chromatography. A plug of BioGel P2 or Sephadex G10 can be used in place of the glass wool.
3. Add protein sample in ∼0.1 column volumes at a flow rate of ∼2 ml/cm2 per hour, and wash the column with PBS until the A280 returns to baseline. A small sample volume makes it easier to detect proteins that do not bind to the column and elute promptly as a sharp peak in the flowthrough. Proteins that bind weakly to the column are gradually washed off by continued elution with PBS alone; these proteins would “smear” as a trailing peak. If a preliminary test run shows that the target protein binds strongly to the lectin, and if the column is not overloaded, the sample can be added to the column in a larger volume. The flow rate can be varied. Sometimes the protein of interest can be absorbed to the lectin by gently mixing the two components for several hours before pouring the loaded gel into the column. The protein can then be eluted after washing as determined by the pilot study.
4. Elute the column with 2 to 3 column volumes of 0.1 M GlcNAc in PBS, and assay the fractions for the protein of interest. Generally, 0.1 M GlcNAc is sufficient to elute the protein. If this does not work, try a higher concentration (e.g., up to 0.25 M GlcNAc), temporarily turn off the column, or raise the
Affinity Purification
9.1.7 Current Protocols in Protein Science
temperature to elute the sample, as with Con A–Sepharose. Adding 0.5 M NaCl to the GlcNAc-containing elution buffer can sometimes improve recovery. A pilot study to determine elution conditions, similar to that described for Con A–Sepharose (see Support Protocol), is highly recommended.
5. Regenerate the column by washing with ≥10 column volumes of PBS or until reducing sugar content is below 20 µg/ml. The column can be reused immediately or stored at 4°C in column buffer containing 0.02% (w/v) NaN3. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Column buffer 0.01 M Tris⋅Cl, pH 7.5 0.15 M NaCl 1 mM CaCl2 1 mM MnCl2 Store indefinitely at room temperature Prepare MnCl2 within a day or two of use and add it to the buffer only after the pH has been adjusted. The final concentration of metal ions can be decreased 10-fold or more if needed. If detergents are required to solubilize the protein, either Triton X-100 or Nonidet P-40 at ≤2% has a negligible effect on lectins. Do not use glucoside-based detergents because they may interfere with binding.
COMMENTARY Background Information See the introductions to UNIT 12.1 and this unit for discussions concerning properties of glycoproteins that relate to their purification.
Critical Parameters
Lectin Affinity Chromatography
As with most protein purification procedures, successful lectin affinity chromotography is empirical, so few parameters are written in stone. However, in using lectins, it is important to first determine the binding and elution conditions in a pilot study. Blindly loading a sample onto a column and then trying different elution conditions may give very low yields, either because the protein becomes inactivated or because it cannot be eluted under those particular conditions. It is usually easy to bind the protein to the column, but elution may be more difficult. There are numerous ways to elute tightly bound protein: increase the concentration of sugar in the buffer used for elution, increase temperature, increase NaCl concentration, or stop the column flow for an extended period during the elution with sugar. A combination of these approaches may be needed. These modifications may cause a protein to elute differently from the same column. Thus, it may be useful to perform two consecutive
runs with different elution conditions to separate the protein of interest from different contaminants. Ketcham and Kornfeld (1992) describe the purification of a rare glycosyltransferase using WGA, whereby the lectin affinity chromatography step yielded a 160-fold purification. In addition, environmental carbohydrate contamination may contribute significantly to the total quantity of carbohydrate in a preparation. Follow the suggestions given within the protocols to minimize this effect. A few general points to remember. Most lectin–sugar binding relies on hydrophobic interactions, so moderate (0.2 M) to high (0.6 M) salt concentrations do not prevent binding. A high salt concentration also reduces nonspecific protein binding. Similarly, tightly bound proteins may be eluted by including 50% ethylene glycol with the sugar. The strength of lectin binding can be significantly affected by the density of lectins on the beads. Usually 3 to 5 mg/ml will bind most proteins, but bind- ing individual free sugar chains may require a higher concentration. Most useful lectins are supplied already bound to beads. If you must couple your own, CNBr-activated Sepharose 4B or Affi-Gel 10 (Bio-Rad) are good supports. Be sure to include any required metal ion and 0.2 M of the eluting sugar during the coupling.
9.1.8 Current Protocols in Protein Science
This will help generate the active conformation and prevent coupling the lectin through the sugar-binding site. Also, keep the soluble lectin concentration below 10 mg/ml during coupling or some may precipitate. Finally, do not couple free Ricinus communis II (RCA-II) from castor beans in your own lab because it is deadly: the LD50 in mice is 0.8 µg/100 g body weight. For all lectin column runs avoid high concentrations (>1%) of nonionic detergents or any SDS.
Anticipated Results
Lectins can give 2- to >100-fold purifications of glycoproteins and nearly quantitative yields, depending upon the source of the sample and the elution conditions. If the protein of interest is reasonably stable at high salt concentrations and room temperature, specific elution conditions can likely be found that will produce good yields.
Time Considerations Initial determination of the binding and elution conditions can easily be done in 1 day, assuming the detection assay for the target protein is simple and fast. The amount of time needed to conduct a single run once the elution conditions are known can vary depending upon the size of the sample and the column, and whether elution requires long-term incubation in the presence of the sugar. Even assuming the extremes of sample size and elution conditions, a single run should be completed in 2 to 3 days.
Literature Cited Cummings, R.D. 1994. Use of lectins in analysis of glycoconjugates. Methods Enzymol. 230:66-86. Ketcham, C.M. and Kornfeld, S. 1992. Purification of UDP-N-acetylglucosamine: Glycoprotein Nacetylglucosamine-1-phosphotransferase from Acanthamoeba castellanii and identification of a subunit of the enzyme. J. Biol. Chem. 267:11645-11653.
Li, M. and Jourdian, G.W. 1991. Isolation and characterization of the two glycosylation isoforms of low molecular weight mannose-6-phosphate receptor from bovine testis. J. Biol. Chem. 266:17621-17630. Manzi, A.E. 1993. Phenol–sulfuric acid assay for hexoses and pentoses. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 17.9.1-17.9.2. John Wiley & Sons, New York.
Key References Beeley, J.G. 1984. Glycoprotein and proteoglycan techniques. In Laboratory Techniques in Biochemistry and Molecular Biology, pp. 29-99. Elsevier Science Publishing, New York. Good discussions about general properties of glycoconjugates. Cummings, R.D. 1994. See above. Most current description of using lectins to characterize sugar chain structures. Lots of details on procedures to analyze free sugar chains rather than glycoproteins. Dulaney, J.T. 1979. Binding interactions of glycoproteins with lectins. Mol. Cell. Biochem. 21:4362. Lists many references and conditions used for lectins in protein purification. Li, M. and Jourdian, G.W. 1991. See above. Excellent example of separation of various glycoforms using lectin–affinity chromatography. Montreuil, J., Bouquelet, S., Debary, H., Fournat, B., Spik, G., and Strecker, G. 1986. Glycoproteins. In Carbohydrate Analysis: A Practical Approach (M.F. Chaplin and J.F. Kennedy, eds.) pp. 166-173. IRL Press, Washington, D.C. Lists several conditions for selected lectin-affinity purifications of proteins.
Contributed by Hudson H. Freeze La Jolla Cancer Research Foundation La Jolla, California
Affinity Purification
9.1.9 Current Protocols in Protein Science
Dye Affinity Chromatography
UNIT 9.2
Dye affinity chromatography is a protein purification procedure based on the high affinity of immobilized dyes for the binding sites on many proteins. It is a rapid, inexpensive, and versatile method that is applicable to the purification of crude cellular extracts. There are three types of dye affinity chromatography—negative chromatography, positive chromatography, and tandem chromatography. Negative chromatography (Basic Protocol 2) is the simplest of the three procedures. Some of the undesired proteins are retained by the immobilized dye while the remaining proteins, including the desired protein, flow through the column. This results in a modest-fold purification. Negative chromatography is particularly convenient for the rapid removal of degradative enzymes such as proteases and nucleases, and for the removal of very abundant proteins, such as serum albumin. A different immobilized dye is used for positive chromatography (Basic Protocol 3 and Alternate Protocol 2). This immobilized dye retains the desired protein as well as some undesired proteins. The desired protein is then selectively eluted using either a nonspecific reagent, such as a neutral salt, or a specific reagent, such as a substrate, cofactor, or competitive inhibitor. Although positive chromatography requires more investigator involvement, it results in a greater-fold purification than obtained using negative chromatography, and is more commonly used. In some cases, a desired protein can be purified to homogeneity from a crude cellular extract using positive chromatography. In tandem chromatography (Basic Protocol 4), the most elaborate of the three procedures, negative chromatography and positive chromatography are coupled sequentially. The protein mixture first passes through a column containing the immobilized dye chosen for negative chromatography. The desired protein flows through this column and immediately enters another column containing the immobilized dye chosen for positive chromatography. The desired protein, which is retained by the latter column, is then eluted using either batch or gradient elution. Tandem chromatography is particularly useful for situations in which the immobilized dye used for positive chromatography binds too many undesired proteins, making selective elution of the desired protein difficult. Use of dye affinity chromatography for protein purification requires selection of one or more immobilized dyes. In addition, positive chromatography requires selection of an elution reagent and its effective concentration. These selections can be made using chromatography, as outlined in Basic Protocol 1, or using centrifugation, as outlined in Alternate Protocol 1, depending on the inclination of the investigator. Immobilized dyes can be purchased as dry powders or suspensions or already poured into miniature columns in kit form. Free dyes also can be purchased and easily immobilized by the investigator (Support Protocol). Alternate Protocol 1 facilitates selection because many samples can be processed simultaneously, but is more expensive because more immobilized dye is required. SELECTION OF COMPONENTS USING CHROMATOGRAPHY The immobilized dye for negative or positive chromatography and the eluant for positive chromatography can be selected using a chromatographic procedure. A small volume of the protein mixture to be purified is applied to a series of miniature columns, each containing a different immobilized dye. The flowthrough and washes from each column are pooled and analyzed for total protein and for desired protein. The immobilized dye column whose pooled flowthrough and washes contain the least amount of undesired protein and the greatest amount of desired protein is appropriate for negative chromatography. Conversely, the immobilized dye column whose pooled liquid contains the greatest Contributed by Earle Stellwagen Current Protocols in Protein Science (1995) 9.2.1-9.2.16 Copyright © 2000 by John Wiley & Sons, Inc.
BASIC PROTOCOL 1
Affinity Purification
9.2.1 CPPS
amount of undesired protein and the least amount of desired protein is appropriate for positive chromatography. The immobilized dye column chosen for positive chromatography is used to select the elution reagent and its effective concentration. The column is washed with small volumes of a potential elution reagent, with the concentration of the reagent increased in small increments in each successive wash. The flowthrough for each wash is then analyzed for both total protein and desired protein. Additional potential elution reagents are examined following application of a fresh aliquot of the protein mixture to be purified. The concentration of elution reagent that provides the greatest-fold purification, recovery of desired protein, and economy of use is selected to be the elution solvent. A specific elution reagent will generally provide a greater increase in fold purification than a nonspecific elution reagent, but the specific elution reagent may cost more. Materials Protein mixture to be purified Appropriate application solvent (see step 1) Series of immobilized dyes (Table 9.2.1 and Support Protocol) Concentrated solution of an appropriate elution reagent (see step 11) Plastic disposable filtration unit with pore size ≤0.45 µm Small plastic disposable chromatography tubes with bed volume at least 1 ml
Table 9.2.1
Some Commercially Available Immobilized Dyesa
Dye
Immobilized dye
Supplierb
Reactive blue 2
Affi-Gel blue gel Blue Sepharose CL-6B Cibacron blue 3G-A-agarose Cibacron blue F3G-A DyeMatrex blue A gel Reactive blue 4-agarose Reactive blue 72-agarose Reactive brown 10-agarose Reactive green 5-agarose DyeMatrex green A gel Reactive green 19-agarose DyeMatrex red A gel Reactive red 120-agarose Red Sepharose CL-6B Reactive yellow 2-agarose Reactive yellow 3-agarose DyeMatrex orange A gel Reactive yellow 13-agarose Reactive yellow 86-agarose
Bio-Rad Pharmacia Biotech ICN Biomedicals, Sigma Pierce Amicon Sigma Sigma Sigma ICN Biomedicals, Sigma Amicon Sigma Amicon ICN Biomedicals, Sigma Pharmacia Biotech ICN Biomedicals, Sigma Sigma Amicon Sigma Sigma
Reactive blue 4 Reactive blue 72 Reactive brown 10 Reactive green 5 Reactive green 19 Reactive red 120
Reactive yellow 2 Reactive yellow 3 Reactive yellow 13 Reactive yellow 86
aScreening kits containing between 5 and 40 immobilized dyes in miniature chromatography columns can be purchased
from Affinity Chromatography, Amicon, Wako Pure Chemical, and Sigma. Alternatively, free reactive dyes such as those listed in Table 9.3.2 can be purchased, immobilized, and prepared for chromatography as described in the Support Protocol. bSupplier names and addresses are provided in SUPPLIERS APPENDIX.
Dye Affinity Chromatography
9.2.2 Current Protocols in Protein Science
Prepare protein sample for application to column 1. Prepare the crude protein mixture. If it is a solid, dissolve it in the application solvent. If the protein mixture to be purified is a suspension or a solution, dialyze it for several hours against the application solvent. A generic application solvent is 50 mM Tris⋅Cl buffer, pH 7.5, at room temperature. The identity of the buffer, its pH, and the temperature of the application solvent may be varied to maintain the integrity of the desired protein without compromising the chromatography. However, addition of reagents that increase the ionic strength of the application solvent above 50 mM may diminish retention of all proteins by the immobilized dye columns. Addition of millimolar concentrations of one or more specific reagents that bind to the desired protein to maintain its integrity may diminish the retention of the desired protein by the immobilized dye columns.
2. Clarify the protein mixture, if necessary, either by centrifugation or filtration. Filtration should be performed using a plastic disposable filtration unit having a pore size of ≤0.45 ìm.
3. Determine the amount of total protein and the amount of desired protein in 1 ml of the protein mixture. Prepare assortment of immobilized dye columns 4. Pour a sufficient volume of each immobilized dye suspension into a different chromatography tube to form a bed volume of 1 ml in each tube. Most commercial screening kits of immobilized dyes are supplied as chromatographic columns having bed volumes ranging from 0.1 to 2.5 ml; these may be used without alteration. The optimal bed volume for a particular situation depends on the total volume of the crude protein mixture available to the investigator; the greater this total volume, the more that can be used. The volume of available crude protein mixture is the principal reason the bed volumes of individual columns in the commercial kits range from 0.1 to 2.5 ml. If one or more of the immobilized dyes are available in powdered form, suspend about 0.5 g of each powdered immobilized dye in application solvent for at least 30 min with gentle stirring prior to pouring the columns.
5. Wash each immobilized dye column with 2 bed volumes of chromatographic application solvent. Let the column run dry and discard the washes. Perform chromatography and assess column performance 6. Apply the same volume of protein mixture to each immobilized dye column and collect the flowthrough in a container. The volume of protein mixture added should contain between 5 and 40 mg of total protein per milliliter of bed volume and sufficient desired protein (1% of total protein) for convenient analysis. The flowthrough can be collected in a disposable tube of sufficient volume to contain the application volume and 2 bed volumes of wash.
7. Wash each immobilized dye column with 2 bed volumes of application solvent and collect the flowthrough in the same container used in step 6. 8. Measure the concentrations of the total protein and the desired protein in the combined flowthrough from each column. 9. Compare the amount of total and desired protein in the combined flowthrough from each column. The immobilized dye whose combined flowthrough contains the least amount of total protein and the greatest amount of desired protein is most appropriate for negative chromatography (see Basic Protocol 2). The immobilized dye whose combined flowthrough Affinity Purification
9.2.3 Current Protocols in Protein Science
contains the greatest amount of total protein and the least amount of desired protein is most appropriate for positive chromatography (see Basic Protocol 3 or Alternate Protocol 2). If positive chromatography is to be performed, continue to use only the immobilized dye column selected for positive chromatography. This column should still retain the desired protein.
Elute desired protein from positive chromatography column 10. Wash the immobilized dye column whose flowthrough contained the least amount of desired protein (i.e., the column selected for positive chromatography) with 2 bed volumes of application solvent containing the lowest concentration of the elution reagent selected for testing. Collect the flowthrough in a separate container. A typical nonspecific elution reagent is NaCl, beginning with a lowest concentration of 0.1 M. A typical specific elution reagent is a substrate, cofactor, inhibitor, effector, or ligand, beginning with a lowest concentration of 1 ìM. The specific elution reagents chosen for testing should represent a good compromise between availability, economy, and high affinity for the desired protein.
11. Measure the total protein and the total desired protein in the flowthrough. 12. Increase the concentration of the elution reagent in the application solvent by increments and repeat steps 10 and 11. A representative increment for a nonspecific elution reagent is 0.2 M and that for a specific elution reagent is an order of magnitude. The application solvent containing the lowest concentration of a reagent that elutes at least 80% of the desired protein retained by the immobilized dye column is defined as the elution solvent. If alternative elution reagents are to be investigated for reasons of economy or availability, proceed with steps 13 to 15.
Repeat elution using alternative elution solvent (optional) 13. Wash the immobilized dye column selected for positive chromatography with 2 bed volumes of 2 M NaCl followed by 2 bed volumes of application solvent and discard the flowthrough. 14. Reapply a volume of the protein mixture as described in step 6 and wash with two column volumes of application solvent. Discard the flowthrough. 15. Repeat steps 10 to 12 using alternative elution reagent. ALTERNATE PROTOCOL 1
Dye Affinity Chromatography
SELECTION OF COMPONENTS USING CENTRIFUGATION The immobilized dye for negative or for positive chromatography and the eluant for positive chromatography can also be selected using a centrifugation procedure instead of the chromatography procedure described in Basic Protocol 1. A different immobilized dye is placed in a series of microcentrifuge tubes. The supernatant solution is discarded from each tube and replaced with a small volume of the protein mixture. The contents of each tube are mixed and centrifuged and the supernatants analyzed for the amount of total protein and desired protein. The immobilized dye whose supernatant contains the least amount of total protein and the greatest amount of desired protein is most appropriate for negative chromatography. Conversely, the immobilized dye whose supernatant contains the greatest amount of total protein and the least amount of desired protein is appropriate for positive chromatography. To select an elution reagent for positive chromatography, the same amount of the immobilized dye selected for positive chromatography is added to a series of microcentrifuge tubes. The protein mixture is added to each tube and the
9.2.4 Current Protocols in Protein Science
supernatant discarded. A different concentration of a potential elution reagent is added to each tube, mixed, and centrifuged, and each supernatant analyzed for the amount of total protein and desired protein present. Any number of different potential specific and nonspecific elution reagents can be evaluated in this manner. The concentration of elution reagent that provides the greatest-fold purification, recovery of desired protein, and economy of use is selected. The centrifugation procedure allows an investigator to select an elution solvent more expeditiously than does the chromatography procedure because many samples can be processed simultaneously. However, a larger amount of each immobilized dye is required. For a list of materials, also see Basic Protocol 1. Prepare the protein sample and set up immobilized dye series 1. Prepare protein sample for application to column (Basic Protocol 1, steps 1 to 3). 2. Add a sufficient volume of an immobilized dye suspension to a capped graduated 1.5-ml disposable plastic microcentrifuge tube to bring the solid/liquid boundary to the 0.4-ml mark. Repeat for each available suspension of immobilized dye. Cap each tube and centrifuge at full speed for 1 to 2 min and discard the supernatant. If one or more of the immobilized dyes are available in powdered form, suspend about 0.5 g of each powdered dye in application solvent for at least 30 min with gentle stirring prior to adding to the microcentrifuge tubes.
3. Add 1 ml application solvent to each microcentrifuge tube. Cap each microcentrifuge tube and mix its contents by inversion, shaking, or vortexing. Let stand for 30 min to swell the immobilized dye matrix. 4. Centrifuge each tube 1 to 2 min at full speed and discard the supernatant. 5. Add 1 ml application solvent to each tube. Cap each microcentrifuge tube and mix its contents by inversion, shaking, or vortexing. Centrifuge each tube 1 to 2 min at full speed and discard the supernatant. Apply protein sample to immobilized dye tubes 6. Add the same volume of the protein mixture to each tube. The volume of protein mixture should contain between 2.5 and 20 mg of total protein and sufficient desired protein for convenient measurement.
7. Cap each microcentrifuge tube and mix its contents by inversion, shaking, or vortexing. Centrifuge each tube 1 to 2 min at full speed. 8. Measure the amount of total protein and desired protein in each supernatant. The immobilized dye whose supernatant contains the least amount of total protein and the greatest amount of desired protein is most appropriate for negative chromatography (see Basic Protocol 2). The immobilized dye whose supernatant contains the greatest amount of total protein and the least amount of desired protein is most appropriate for positive chromatography (see Basic Protocol 3 or Alternate Protocol 2).
Test application solvents for positive chromatography 9. If positive chromatography is to be performed, add a sufficient volume of the immobilized dye suspension selected for positive chromatography to a series of graduated microcentrifuge tubes to bring the solid/liquid boundary in each tube to the 0.4-ml mark. 10. Repeat steps 3 to 6. 11. Construct a series of application solvents each containing concentration of the elution reagent to be tested.
Affinity Purification
9.2.5 Current Protocols in Protein Science
A typical nonspecific elution reagent is NaCl, beginning with a lowest concentration of 0.1 M and increasing in 0.2 M increments. A typical specific elution reagent is a substrate, cofactor, inhibitor, effector, or ligand having a lowest concentration of 1 ìM and increasing in order of magnitude increments.
12. Add 1 ml of each application solvent containing the elution reagent to a different centrifuge tube containing immobilized dye. Cap the microcentrifuge tube and mix the contents by inversion, shaking, or vortexing. Centrifuge each tube 1 to 2 min at full speed. 13. Measure the concentration of total protein and of desired protein in each supernatant. The application solvent containing the lowest concentration of a reagent that elutes at least 80% of the desired protein retained by the immobilized dye column and that represents a good choice with regard to cost and availability is defined as the elution solvent.
14. Repeat steps 11 to 13 to test additional elution reagents or to refine the concentration of the selected elution reagent, if desired. BASIC PROTOCOL 2
NEGATIVE CHROMATOGRAPHY A protein mixture is applied to an immobilized dye column selected for negative chromatography and the column is washed with application solvent. Many of the undesired proteins are retained by the column while the desired protein as well as some of the undesired proteins flow through the column. The concentrations of the total protein and total desired protein in the combined flowthrough are measured. These values should reflect a yield of >80% of desired protein and a modest-fold purification. Negative chromatography is the simplest to execute but, because the desired protein as well as some undesired proteins are not retained and appear in the column flowthrough, it is the least likely to dramatically increase the fold purification of a desired protein. However, it is particularly convenient for the rapid removal of troublesome contaminating proteins such as proteases, nucleases, or serum albumin. Materials Protein mixture to be purified Application solvent, with and without 2 M NaCl Immobilized dye selected for negative chromatography (see Basic Protocol 1 or Alternate Protocol 1) 0.02% (w/v) sodium azide Empty chromatographic column with height/diameter ratio ≤5 (sintered glass funnel can suffice) 1. Prepare the protein mixture for chromatography (see Basic Protocol 1, steps 1 to 3). 2. Divide the total protein concentration (in milligrams) to be purified by 5 to estimate the bed volume (in milliliters) of the immobilized dye needed. Add suspension of the immobilized dye to the chromatographic column until that bed volume is acquired. Division is by 5 because it is conservatively estimated that each milliliter of column bed volume can retain 5 mg of protein. Immobilized dyes may be purchased either as suspensions or as dry powders from commercial sources (see Table 9.2.1). The suspensions are easier to use because they are already swollen; therefore they are the form of choice. If the powdered form of the immobilized dye is used, it should be suspended in an application solvent for about an hour with gentle stirring prior to pouring the column.
Dye Affinity Chromatography
9.2.6 Current Protocols in Protein Science
3. Wash the column with application solvent until no further dye is eluted. The volume of application solution required for washing depends upon the storage of the column. The column matrix slowly but steadily hydrolyzes, generating soluble matrix fragments still containing covalently attached dye molecules. The longer the storage time and the higher the temperature, the greater the amount of solubilized dye. The volume of wash is determined by the investigator; when colored dye stops appearing in the wash, the column is ready for chromatography. This may require 1 or 20 column volumes. The presence of the colored dye can be detected in the washing either by visual inspection or by spectrophotometric measurement at an appropriate wavelength.
4. Apply the entire protein mixture to be purified to the immobilized dye column using a flow rate of at least 1 ml/min. Collect the flowthrough liquid in a single container. The flow rate is dictated by the size of the column and the density of its packing. A fast flow rate reduces experimental time. Equilibration of protein with immobilized dye is rapid; there is little advantage in using flow rates less than 1 ml/min.
5. Wash the column with 1 column volume of application solvent. Collect the flowthrough liquid in the same container as in step 4. 6. Measure the concentration of total protein and total desired protein in the combined flowthrough liquids. These values should reflect a good yield, >80% of desired protein, and a modest-fold purification. If the fold purification is less than expected from the results observed using Basic Protocol 1 or Alternate Protocol 1, add more immobilized dye to the column and reapply the combined flowthrough liquids.
7. Wash the column with 1 column volume of application solvent containing 2 M NaCl to displace the retained protein. Protein displacement can be ascertained by monitoring the amount of protein in the wash effluent.
8. Prepare the column for storage by washing with 2 column volumes of 0.02% sodium azide and storing the poured column at 4°C. POSITIVE CHROMATOGRAPHY USING BATCH ELUTION A protein mixture is passed through an immobilized dye column selected for positive chromatography. This immobilized dye retains the desired protein and some of the undesired proteins. The desired protein is then selectively eluted using either a nonspecific or a specific elution reagent (a higher-fold purification will likely be obtained using the specific eluant), and the concentrations of total protein and desired protein in each fraction are measured. This protocol for positive chromatography utilizes batch elution, which is simpler to execute than Alternate Protocol 2 utilizing gradient elution (the latter may yield a higher-fold purification than batch elution but requires more effort to set up).
BASIC PROTOCOL 3
Materials Protein mixture to be purified Appropriate application solvent Immobilized dye and elution solvent selected for positive chromatography (see Basic Protocol 1 or Alternate Protocol 1) Empty chromatographic column with height/diameter ratio >5 Fraction collector 1. Prepare protein sample and immobilized dye column (see Basic Protocol 2, steps 1 to 5) using the immobilized dye selected for positive chromatography.
Affinity Purification
9.2.7 Current Protocols in Protein Science
2. Measure the concentration of the total protein and the desired protein in the flowthrough liquids. If the majority of the desired protein is retained by the column, discard the flowthrough liquid. If a significant amount of desired protein is not retained, add more immobilized dye to the column and reapply the flowthrough liquid. Typically, at least 70% and, if possible, 90% or more of the desired protein is retained.
3. Wash the column with the elution solvent selected for positive chromatography using a flow rate of at least 1 ml/min. Collect the flowthrough in a fraction collector in constant-volume fractions, typically 1 ml. 4. Measure the concentration of total protein and desired protein in each fraction. 5. Pool fractions containing the desired protein. The specific activity of the desired protein in the fractions can be used to determine which fractions to include in the pool. Typically pool all tubes containing at least 10% of the total amount of desired protein applied to the column.
6. Wash and store the column (see Basic Protocol 2, steps 7 and 8). ALTERNATE PROTOCOL 2
POSITIVE CHROMATOGRAPHY USING GRADIENT ELUTION A protein mixture is passed through an immobilized dye column selected for positive chromatography. This immobilized dye retains the desired protein and some of the undesired proteins. The desired protein is selectively eluted using a linear gradient constructed from the elution solvent selected for positive chromatography, and the concentrations of total protein and desired protein in each fraction are measured. This protocol is more elaborate to set up and operate, but sometimes generates a greater-fold purification than obtained using the batch elution system of Basic Protocol 3. Additional Materials (also see Basic Protocol 3) Empty chromatographic column having a height/diameter ratio of >5 A coated magnet and magnetic stirrer Linear gradient maker (commercial or constructed from two beakers of the same size connected by a siphon) 1. Prepare protein sample and immobilized dye column (see Basic Protocol 3, steps 1 and 2). All solutions should be carefully introduced so that the column packing is minimally disturbed.
2. Place a volume of the elution solvent selected for positive chromatography in the nonmixing reservoir of the linear gradient maker. This volume is typically equal to the column volume. The nonmixing reservoir is the reservoir more distant from the column. Its contents flow into the mixing reservoir prior to introduction onto the column.
3. Place 10% of the volume of elution solvent used in step 2 in the mixing reservoir of the gradient maker. Add application solvent to make up the remaining 90% of the volume. This procedure will generate an order-of-magnitude gradient terminating in the eluant concentration employed in batch elution.
Dye Affinity Chromatography
9.2.8 Current Protocols in Protein Science
4. Place a magnetic stir bar in the mixing reservoir. Place the mixing reservoir on a magnetic stirrer. Position the nonmixing reservoir so that the bottom of this vessel is adjacent to and coplanar with the bottom of the mixing reservoir. 5. Provide for liquid flow between the two reservoirs and between the mixing reservoir and the column. 6. Gently mix the liquid in the mixing reservoir. Initiate liquid flow between the two reservoirs and between the mixing reservoir and the column. Use a flow rate of 1 ml/min and collect 1-ml fractions. Resolution of desired and undesired proteins may be improved by narrowing the concentration range of the specific or nonspecific elution reagent, by increasing the total volume of the elution solutions, or by decreasing the flow rate of the elution solutions.
7. Measure the protein concentrations, pool the appropriate fractions, and wash the column (see Basic Protocol 3, steps 4 to 6). TANDEM CHROMATOGRAPHY In tandem chromatography, columns for negative and positive immobilized dye chromatography are sequentially linked. The protein mixture is first passed through the column containing the immobilized dye chosen for negative chromatography, which preferentially removes undesired protein. The effluent from this column is immediately passed over a second column containing the immobilized dye chosen for positive chromatography, which retains the desired protein. The desired protein is then selectively eluted from the second column using either batch or gradient elution and the elution solvent selected for positive chromatography.
BASIC PROTOCOL 4
Materials Protein mixture to be purified Appropriate application solvent Immobilized dyes selected for negative and for positive chromatography (Basic Protocol 1 or Alternate Protocol 1) Selected elution solvent 2 M NaCl 0.02% (w/v) sodium azide Empty chromatographic column with height/diameter ratio >5 (two columns are required if positive chromatography is to be performed using batch elution) Empty chromatographic column with height/diameter ratio ≥5 (if positive chromatography is to be performed using gradient elution) Linear gradient maker (for gradient elution) Fraction collector Pipettors and disposable tips Prepare the protein sample and columns 1. Prepare the protein mixture for chromatography (see Basic Protocol 1, steps 1 to 3). 2. Prepare the negative and positive columns (see Basic Protocol 2, steps 2 and 3). The immobilized dye selected for negative chromatography is placed in a column with a height/diameter ratio ≤5. The immobilized dye selected for positive chromatography is placed in another identical column (if batch elution is to be employed) or in a column with a height/diameter ratio ≥5 (if gradient elution is to be employed). Affinity Purification
9.2.9 Current Protocols in Protein Science
3. Position the negative column immediately above the positive column. Connect the two columns so that the effluent of the upper column flows directly on top of the lower column. Pass sample through columns 4. Apply the protein mixture to the upper negative column. Collect the flowthrough liquid from the lower positive column in a single container. 5. Wash the tandem columns with 1 collective column volume of application solvent. Add the flowthrough from the washing to the container in step 4. 6. Measure the concentration of desired protein in the collective flowthrough to ascertain that the desired protein has been retained. 7. Uncouple the upper negative column and set it aside. Recover the protein 8. Wash the positive column with 2 column volumes of application solvent. 9. Subject the positive column to either batch or gradient elution using the elution solvent selected for positive chromatography. Perform and process batch elution (see Basic Protocol 3, steps 3 to 5) or gradient elution (see Alternate Protocol 2, steps 2 to 6). 10. Wash and store each column (see Basic Protocol 2, steps 7 and 8). SUPPORT PROTOCOL
IMMOBILIZATION OF REACTIVE DYES Reactive dyes must be covalently attached to an insoluble porous matrix (“immobilized”) to be used in a chromatographic protocol for protein purification. A reactive dye is a visible chromophore that has a good chemical leaving group, as illustrated in Figure 9.2.1. Immobilization of a reactive dye is a simple procedure that can be performed in the laboratory to generate immobilized dyes that cannot be purchased or to save the cost of commercial immobilization. Table 9.2.2 lists a selection of dyes that can be purchased at modest cost. Each of these dyes can be immobilized by reacting it with a chromatographic matrix in aqueous solvents using moderate conditions. The nature of the covalent attachment is illustrated in Figure 9.2.1B. This protocol is scaled for preparation of an immobilized dye column having a bed volume of 100 ml. Materials Reactive dye (Table 9.2.2) Concentrated solution of KCl Reactive chromatographic matrix: e.g., Sepharose CL-4B or CL-6B (Pharmacia Biotech) or other cross-linked agarose 4 M and 1 M NaCl 10 M NaOH 2 M NH4Cl Large sintered glass funnel Plastic disposable filter with pore size of 0.45 µm
Dye Affinity Chromatography
1. Remove any salt, buffer, or surfactant from the reactive dye sample by precipitation of the dye as the potassium salt from aqueous solution and filtration of the precipitate using a sintered glass funnel mounted on a filter flask. Place several grams of the dye in a beaker and add water to make a concentrated solution of the dye. Add concentrated solution of KCl dropwise to precipitate the dye from the solution. Pass the dye
9.2.10 Current Protocols in Protein Science
SO3–
A NH
N O
NH
N
NH SO3–
N Cl
SO3–
O
NH2
SO3–
B
NH
N O
NH
N
NH SO3–
N O
SO3–
O
MATRIX
NH2
N
C
CH2O
N
NH
SO3–
N Cl
N N
D
N (CH3)2N+H
Cl
N N
Cl N
NH N
CH3
Cl
Figure 9.2.1 Structures of some reactive dyes. The dyes are arranged so that their triazine rings, the rightmost rings in structures C and D, have a common position. (A) Reactive blue 2, color index 61211, a reactive monochlorotriazine dye, also designated as Cibacron blue 3G-A and Procion blue H-B. (B) Immobilized reactive blue 2, also designated as Affi-Gel blue gel, Blue Sepharose CL-6B, Cibacron blue 3G-A-agarose, and DyeMatrex blue A gel. (C) Reactive red 8, color index 17908, a reactive dichlorotriazine dye, also designated as Procion scarlet MX-G. (D) A positively charged reactive dye.
suspension through a plastic disposable filter with a pore size of 0.45 µm, wash the filtered precipitate with ∼100 ml of water, and air dry the washed precipitate. The most common reactive dyes—e.g., Procion (ICI) and Cibacron (Ciba-Geigy) dyes— have a sulfonated chromophore linked to a chlorotriazine group by an aminoether bridge.
2. Suspend 80 g chromatographic matrix in 280 ml water. 3. Dissolve 1.2 g reactive dye in 80 ml water and add it to the matrix suspension.
Affinity Purification
9.2.11 Current Protocols in Protein Science
Table 9.2.2
Some Commercially Available Reactive Dyesa
Generic name
Color index numberb
Reactive black 5 Reactive blue 2
61211
Reactive blue 4 Reactive blue 5
61205 61210
Reactive blue 15
74459
Reactive blue 19 Reactive blue 114 Reactive blue 160 Reactive brown 10 Reactive green 5 Reactive green 19 Reactive orange 14 Reactive orange 16 Reactive red 4
17757 18105
Reactive red 120 Reactive violet 5 Reactive yellow 2
18097 18972
Reactive yellow 3 Reactive yellow 81 Reactive yellow 86
13245
Some commercial names
Suppliers
Remazol black B Cibacron blue 3G-A Procion blue H-B Procion blue MX-R Cibacron brilliant blue BR-P Procion blue H-GR Cibacron turquoise blue GF-P Procion turquoise H-GF Remazol brilliant blue R Drimarene brilliant blue K-BL Procion blue HE-RD Procion brown MX-5BR Cibacron brilliant green 4G-A Procion green H-4G Procion green HE-4BD Procion yellow MX-4R Remazol brilliant orange 3R Cibacron brilliant red 3B-A Procion red H-7B Cibacron brilliant red 4G-E Procion red HE-3B Remazol brilliant violet 5R Cibacron brilliant yellow 3G-P Procion yellow H-5G Procion yellow H-A Procion yellow HE-3G Procion yellow M-8G
Aldrich, ICN Biomedicals, Sigma Aldrich, ICN Biomedicals Sigma, Spectrum Aldrich, ICN Biomedicals, Sigma ICN Biomedicals, Sigma Aldrich, ICN Biomedicals, Sigma ICN Biomedicals Sigma Sigma ICN Biomedicals, Sigma ICN Biomedicals, Sigma ICN Biomedicals, Sigma ICN Biomedicals, Sigma Aldrich Aldrich, ICN Biomedicals Sigma, Spectrum ICN Biomedicals, Sigma ICN Biomedicals, Sigma Aldrich, ICN Biomedicals Sigma, Spectrum Aldrich ICN Biomedicals, Sigma
aDichlorotriazine reactive dyes are denoted as Procion MX dyes; all the remaining dyes are monochlorotriazine dyes. bNumbers are taken from the Colour Index, The Society of Dyers and Colourists and the American Association of Textile Chemists and Colorists (1971).
4. Add 40 ml of 4 M NaCl to the matrix suspension. 5. If a monochlorotriazine reactive dye is used, add 4 ml of 10 M NaOH to the matrix suspension and gently stir it with a magnetic stirrer and stir bar for 72 hr at ambient temperature or 16 hr at 55° to 60°C. If a dichlorotriazine reactive dye is used, add 0.5 ml of 10 M NaOH and gently stir the matrix suspension with a magnetic stirrer and stir bar for 4 hr at ambient temperature. 6. Filter the suspension using the sintered glass funnel and wash the solid material on the filter with copious quantities of water, then 1 M NaCl, and then water again until the filtrate is clear. 7. Suspend the material on the filter in 2 M NH4Cl, pH 8.5, and gently stir with a magnetic stirrer and stir bar for 4 hr at ambient temperature. Dye Affinity Chromatography
This step eliminates any remaining chloro groups.
9.2.12 Current Protocols in Protein Science
8. Filter and wash the suspension as described in step 6. 9. Either store the immobilized dye as a suspension in water at 4°C or dry the immobilized dye on a sintered glass filter and store as a dry powder. COMMENTARY Background Information The surfaces of all proteins bind at least one biochemical with high affinity. Accordingly, immobilization of a biochemical on a chromatographic support should result in the retention of all proteins that have a binding site for that biochemical. The biochemical can exist in two forms: the mobile biochemical, which moves about freely in solution, and the immobilized biochemical, which is covalently attached to a chromatographic support and cannot move about freely. In chromatography, a competition exists between mobile and immobilized forms of the biochemical for the binding sites on the proteins. If the immobilized form is the dominant concentration of the biochemical, as in application to a column, the protein exists principally as a protein/immobilized dye complex and is not free to move in solution. If the mobile form is the dominant concentration of the biochemical, as in elution, the protein exists principally as a protein/mobile dye complex and moves freely in solution. Elution solutions containing a concentration gradient of the mobile form of the biochemical can often be used to distinguish among those proteins that bind the immobilized biochemical. Affinity chromatography, however, can present problems. The biochemical must be immobilized in such a manner that the protein can bind it with high affinity, often a substantial challenge. Each immobilized biochemical can bind only a limited number of different proteins, requiring construction of a large number of different affinity columns for purification of a series of proteins. Further, the degradative enzymes present in crude cellular extracts often cleave immobilized biochemicals into useless remnants, necessitating either the frequent replacement of affinity columns or the postponement of their use until the end of a purification scheme. Such postponement compromises the selective power of affinity chromatography, which should, in principle, purify a protein to homogeneity in a single step. Fortunately, immobilized dye affinity chromatography circumvents these problems. A single reactive dye can effectively bind a large number of different biochemicals. A reactive
dye is sufficiently complex that immobilization does not reduce its affinity. An immobilized reactive dye is not degraded by enzymes commonly found in crude protein mixtures, as the chemistry of the dye is quite distinct from that of typical biochemicals. An immobilized dye functions as a stable general affinity molecule that can be made selective by the nature and concentration of the mobile biochemical chosen as the elution reagent. A single immobilized dye, variously known as Cibacron blue (F)3G-A, Procion blue H-B, reactive blue 2, blue dextran, Affi-Gel blue, or DyeMatrex blue A, has contributed significantly to the purification of over 60 different proteins. Some of these proteins have been purified to homogeneity from a crude bacterial extract in a single step providing a several-thousand-fold purification with >80% yield. Immobilized dyes as a group have contributed significantly to the purification of several hundred different proteins.
Critical Parameters and Troubleshooting Several aspects of dye affinity chromatography are critical. Foremost is the ionic strength of the application solvent. Because application solvents of ionic strength >50 mM weaken the electrostatic interactions that contribute to the binding of desired proteins by immobilized dyes, the concentrations of ionic components in the application solvent should be kept to a minimum. Second is the presence of nonionic detergents in the application solvent. Nonionic detergents are frequently used to solubilize intrinsic protein located in subcellular organelles such as membranes. These nonionic detergents form micelles that encapsulate immobilized dyes, making them inaccessible to proteins. Addition of a modest concentration of a negatively charged detergent, such as SDS or deoxycholate, to an application solvent containing a nonionic detergent results in the formation of mixed micelles. The negatively charged immobilized dyes repel the negatively charged mixed micelles in solvents of modest ionic strength, preserving the accessibility of immo-
Affinity Purification
9.2.13 Current Protocols in Protein Science
Supplement 4
Dye Affinity Chromatography
bilized dyes to desired proteins (Robinson et al., 1980). The nonionic and ionic detergents should have a concentration ratio of ∼5:1. Third is the sign of the electrostatic charges on immobilized dyes. Most reactive dyes are negatively charged; this is normally an advantage, as most protein binding sites contain positively charged amino acid residues designed to bind negatively charged biochemicals, such as cAMP, glucose 6-phosphate, or oligo(dAT). However, some biochemicals have a positive charge; the proteins to which they bind must then contain negatively charged residues. These binding proteins are electrostatically repelled by negatively charged immobilized dyes in solvents of modest ionic strength. The immobilized form of a positively charged dye (Fig. 9.2.1D) retains proteins having binding sites for positively charged biochemicals, such as trypsin, thrombin, and carboxypeptidase (Clonis et al., 1987). This immobilized dye is particularly useful in removing contaminating proteases from cellular extracts by negative chromatography. The two primary problems encountered in immobilized dye affinity chromatography are the inability to retain a desired protein and the inability to elute a desired protein that has been retained. The inability to retain a desired protein may result from a number of situations. If a fresh immobilized dye column is being used, the following may pertain: The ionic strength of the protein mixture may be too high. Either dilute the protein mixture with distilled water or dialyze it against a low ionic strength application solvent, such as 50 mM Tris⋅Cl devoid of neutral salts. The concentration in the protein mixture of the biochemical that binds to the desired protein may be too high. Either dilute the mixture with distilled water or, if the biochemical is sufficiently small, dialyze the mixture against a low ionic strength application solvent. Retention of the desired protein may necessitate the presence of a metallic cation, such as Mg2+ or Zn2+. Add a minimal concentration of one or more likely metallic cations to the application solvent. A nonionic detergent may be present in the protein mixture. Add a sufficient amount of a negatively charged detergent, such as SDS or deoxycholate, to the crude protein mixture to maintain the accessibility of the immobilized dye. However, note that SDS, even in low concentrations, denatures many proteins. The immobilized dye may have a relatively weak affinity for the desired protein. Add more
immobilized dye to the column or screen a series of alternative immobilized dyes for one having a stronger affinity for the desired protein. The desired protein is designed to bind only positively charged biochemicals. Purchase or synthesize an immobilized dye having a net positive charge. The desired protein may be among a small group of proteins that is not retained by any immobilized dye. Negative chromatography can then be used to great advantage, particularly a series of negative columns in tandem. If an old immobilized dye column is being used, the following may pertain: Insufficient immobilized dye may be accessible. After repeated use of an immobilized dye column the dye may become bound with proteins, detergents, and other components in cellular extracts that are not removed by washing with 2 M NaCl. An immobilized dye column should be periodically washed with several column volumes of one or more of the following solutions to displace these dye-bound components: a strong denaturant such as 8 M urea, 6 M guanidine⋅HCl, or 1% SDS; a potent lyotropic reagent such as 3 M KSCN; a strong base such as 0.5 M NaOH; or a mixed solvent such as chloroform/methanol. Insufficient immobilized dye may be present. If an immobilized dye column has been used extensively or stored in solution, the glycosidic bonds linking the sugars in the agarose matrix will be hydrolyzed. Such hydrolysis results in loss of the immobilized dye. A new immobilized dye column is strongly colored. A depleted immobilized dye column is lightly colored and should be replaced. The inability to elute a desired protein that has been retained on an immobilized dye column presents a different problem, in that the affinity is too strong rather than too weak. Suggestions for weakening the affinity include the following: Performing the chromatography at a different pH. Most proteins exhibit a pH optimum for binding a biochemical. This pH optimum results in part from changes in the ionic character of some of the amino acid side chains in the protein that are directly involved in binding. Changes in pH that weaken the affinity of biochemicals are likely to weaken the affinity for the immobilized dye as well. Use Basic Protocol 1 to determine the effect of pH on the elution of the desired protein. Note, however, that pH values outside the range 4 to 11 may result in irreversible inactivation.
9.2.14 Supplement 4
Current Protocols in Protein Science
Performing the chromatography at a different temperature. The binding of a biochemical by a protein involves a combination of weak noncovalent bonds including hydrogen bonding, hydrophobic interactions, and electrostatic interactions. Changes in temperature weaken some of these bonds and strengthen others. Accordingly, changing the temperature may weaken the affinity of the immobilized dye for the desired protein. Note, however, that the temperature should not be increased above 50°C to prevent thermal inactivation of the desired protein. Increasing the potency of the elution solvent. Nonspecific elution solvents promote elution by weakening the electrostatic and hydrophobic contributions to affinity. The electrostatic contributions will usually be minimized in a 0.2 M solution of any neutral salt. How-ever, minimization of the hydrophobic contributions requires higher concentrations and judicious choice of the neutral salt. The lyotropic or Hofmeister series reflects the relative potency of neutral salts. A 3 M solution of the neutral salt KSCN is among the most potent for weakening hydrophobic interactions (Robinson et al., 1981). Specific elution solvents, by contrast, weaken affinity by competing with the immobilized dye for the desired protein. The higher the concentration of the biochemical and the stronger its affinity for the desired protein, the greater its effectiveness as an eluant. Accordingly, the concentration of a specific elution solvent should be increased in an effort to elute the desired protein. Clearly, a limiting concentration will be set by the solubility, availability, and cost of the specific eluant. Alternatively, other biochemicals known to bind to the desired protein as substrates, coenzymes, cofactors, competitive inhibitors, and allosteric effectors may be surveyed for their efficacy as elution reagents. Adding a chelation agent. Metallic cations often function as obligatory cofactors in the binding of biochemicals to proteins. Accordingly, the affinity of the desired protein for the immobilized dye may be weakened by the presence of a chelation agent such as 0.1 M EDTA or EGTA. Combining specific elution reagents. Immobilized dyes are sufficiently large that a single dye molecule may occupy adjacent biochemical binding sites on a protein: e.g., a dye may occupy the substrate site and the coenzyme site on an enzyme surface. Alternatively, two different immobilized dye molecules may simul-
taneously bind to two remote sites on the same protein molecule: e.g., different immobilized dye molecules may simultaneously bind to the substrate site and to the allosteric site on an enzyme. In each of these situations, the elution of a desired protein can be accomplished using a combination of specific elution reagents. Adding a protein denaturant such as 6 M guanidine⋅HCl. Most proteins can be eluted from an immobilized dye column using a denaturant that unfolds protein binding sites. Because most retained proteins have already been eluted, addition of a denaturant should result in a substantial-fold purification, provided the desired protein is denatured reversibly. Such reversibility is commonly achieved by dilution or dialysis of the column eluate with application solvent. Lowering the density of the immobilized dye. Many desired proteins are oligomers containing two or more identical binding sites. If the chromatographic matrix contains a high density of accessible immobilized dye molecules, it is likely that two or more immobilized dye molecules will simultaneously bind to a single desired oligomeric protein (Hogg and Winzor, 1985). Such multiple binding makes elution more difficult. Diluting an immobilized dye matrix with an unmodified matrix will not decrease the density of the immobilized dyes on each chromatographic bead. The only remedy is to obtain the same immobilized dye from an alternative supplier, hoping to obtain a less dense sample, or to immobilize the dye in the laboratory using the Support Protocol. Selecting an alternative immobilized dye that has a weaker affinity for the desired protein. Such weakening should facilitate improved results in positive chromatography.
Anticipated Results The yield of the desired protein following immobilized dye chromatography is commonly >80%. Random differences in yield occur depending upon how completely the desired protein is retained (positive chromatography) or not retained (negative chromatography) and how completely the retained protein is eluted. Lesser yields are usually the result of proteolysis either before or during chromatography. Proteolysis may generate remnants of the desired protein that have weaker affinities for an immobilized dye. Such remnants elute prior to the elution of the dominant unproteolyzed desired protein and would not be included in the pooled fractions. Good yields of desired proteins result in part from the speed of affinity
Affinity Purification
9.2.15 Current Protocols in Protein Science
chromatography, the temperature, the stabilization provided to the desired protein from binding an immobilized dye, and the respectful treatment of a crude protein mixture by an experienced investigator. The fold purification achieved by immobilized dye chromatography is largely controlled by the relative population of the desired protein in a protein mixture. If 10% of the mixture is the desired protein, then a 10-fold purification step is maximal. If <0.1% of the mixture is the desired protein, then a >1000-fold purification step is feasible and has been observed. However, in most instances 10- to 100-fold purifications are observed.
Time Considerations
The Society of Dyers and Colourists and the American Association of Textile Chemists and Colorists. 1971. Colour Index, 3rd edition. The Society of Dyers and Colourists, Bradford, England, and the American Association of Textile Chemists and Colorists, Research Triangle Park, NC.
Key References Clonis, Y.D., Atkinson, T., Bruton, C.J., and Lowe, C.R. 1987. Reactive Dyes in Protein and Enzyme Technology. Macmillan, New York. A comprehensive review of the preparation, immobilization, and utilization of reactive dyes for protein purification. Lowe, C.R., Burton, S.J., Burton, N., Stewart, D.J., Purvis, D.R., Pitfield, I., and Eapen, S. 1990. New developments in affinity chromatography. J. Mol. Recognit. 3:117-122.
The preliminary survey can be completed in a few hours or a day, depending on its breadth. Once a protocol is selected, negative chromatography should be completed in 1 hr and positive chromatography in a few hours. These time estimates greatly depend on the volume of the solutions to be processed and the flow rates employed.
A lively discussion of emerging developments.
Literature Cited
An alternative discussion of the topic with more emphasis on strategy and less on detail.
Clonis, Y.D., Stead, C.V., and Lowe, C.R. 1987. Novel cationic triazine dyes in protein purification. Biotechnol. Bioeng. 30:621-627. Hogg, P.J. and Winzor, D.J. 1985. Effects of solute multivalency in quantitative affinity chromatography: Evidence for cooperative binding of horse liver alcohol dehydrogenase to blue Sepharose. Arch. Biochem. Biophys. 240:70-76. Robinson, J.B., Jr., Strottmann, J.M., Wick, D.G., and Stellwagen, E. 1980. Affinity chromatography in nonionic detergent solutions. Proc. Natl. Acad. Sci. U.S.A. 77:5847-5851. Robinson, J.B., Jr., Strottmann, J.M., and Stellwagen, E. 1981. Prediction of neutral salt elution profiles for affinity chromatography, Proc. Natl. Acad. Sci. U.S.A. 78:2287-2291.
Scopes, R.K. 1994. Protein Purification, 3rd ed. Springer-Verlag, New York. A useful discussion of immobilized dye affinity chromatography by the advocate of tandem chromatography. Stellwagen, E. 1990. Chromatography on immobilized reactive dyes. Methods Enzymol. 182:343357.
Stellwagen, E. 1993. Affinity chromatography with immobilized dyes. In Molecular Interactions in Bioseparations (T.T. Ngo, ed.) pp. 247-255. Plenum Press, New York. A detailed review of the interaction of reactive blue 2 with a typical oligomeric protein, alcohol dehydrogenase.
Contributed by Earle Stellwagen University of Iowa Iowa City, Iowa
Dye Affinity Chromatography
9.2.16 Current Protocols in Protein Science
Affinity Purification of Natural Ligands
UNIT 9.3
Immobilization of proteins, nucleic acids, and other “bioligands” is not simple. The proper bioligand needs to be selected for the specific application of the product. This decision influences all others: the matrix is chosen and activated by the method that is most appropriate for this specific application and the ligand is coupled under conditions dictated by the activation method and the nature of the ligand (e.g., is it a labile protein or a sturdy enzyme cofactor?). There are many matrices and activation and coupling methods, and new ones are constantly being developed. The most frequently used are hydrophilic matrices, which exhibit the necessary low nonspecific binding, and coupling systems that react primarily with nucleophilic residues, such as lysyl amine groups in proteins. Orientation of the resulting biomolecule is extremely important, although these effects have been only superficially investigated. Immobilized IgG is only twofold more efficient when immobilized solely by the Fc portion (Murayama et al., 1978; Doman et al., 1990; Hermanson et al., 1992), whereas the interactions of large proteins that bind macromolecular ligands (lysozyme is a good example) are changed by several orders of magnitude depending on the orientation of the molecule (Fig. 9.3.1). For this reason, it is fortunate that both cationic (e.g., fluoromethyl pyridinium) and hydrophobic (e.g., Emphaze, tosyl agarose) activation methods are available. The choice of matrix and the stability of the bound ligand determine the choice of activation and coupling methods. Preactivated matrices are available but are very costly. Therefore they are best used when small amounts of ligand are to be immobilized for “test” columns and methods development. Larger projects and large-scale applications require ligand activation in the laboratory. Because of their frequent use in the past, agarose gel is often the matrix of choice and cyanogen bromide is often the activation and coupling agent of choice. However, depending on the procedure, neither may be optimal and before beginning a project it is best to explore the range of matrices and methods carefully, particularly if later industrial level scale-up with regulatory implications is anticipated. An overview of the major activation chemistries is given in Tables 9.3.1 and 9.3.2. Of the many activation systems available, three of the most frequently used are
A
B
S
L
L
S
Figure 9.3.1 Binding of small (S) versus large (L) substrates to an oriented (A) and randomly (B) immobilized enzyme. Arrows signify the movement of the enzyme to the ligand and its subsequent binding to the ligand. Reproduced from Voivodov et al. (1993) with permission of Hüthig & Wepf Publishers, Zug, Switzerland.
Affinity Purification
Contributed by William H. Scouten
9.3.1
Current Protocols in Protein Science (1995) 9.3.1-9.3.15 Copyright © 2000 by John Wiley & Sons, Inc.
CPPS
Table 9.3.1
Selected Reagents to Activate Matrix Hydroxyl Functionsa
Group that reacts
Reagent toxicity
Activation Type of bond time (hr)
Tresyl chloride, sulfonyl Excellent chlorides
Thiols, amines
Low
0.1-1.0
Secondary amine
Cyanogen bromide
Poor
Amines
High
0.1-0.4
Bisoxiranes (epoxides)
Excellent
Thiols
Moderate
5-18
Epichlorohydrin
Excellent
Moderate
2-24
High
0.5-2
Isourea or imidocarbonate Secondary amine Secondary amine Triazine ether
Activation method
Stability of bond
References Lawson et al. (1983); Gribnau (1977) Axen et al. (1967) Axen et al. (1967)
Low High
1-2 0.5-2
Anilinyl Secondary amine
Axen et al. (1967) Axen et al. (1967)
Glutaraldehyde
Thiols, amines Good Amines, thiols, hydroxyls Good Amines Poor in base, Amines fair at pH <7.0 Excellent Amine
Moderate
5-18
Axen et al. (1967)
Phosphoryl chloride
Excellent
Amine
High
18
Secondary amine (?) Amide
p-Nitrophenyl chloroformate
Good
Amine
Moderate
1.0
Urethane
N-Hydroxysuccinimidyl Good chloroformate
Amine
Moderate
1.0
Urethane
Carbonyldiimidazole Good Poor 1-Cyano-4-dimethylaminopyridinium tetrafluoroborate 2-Fluoro-1-methylpyrid- Excellent inium tosylate
Amine Amines
Moderate Moderate
Amine/thiol
Moderate
Triazine
Benzoquinone Divinyl sulfone
20 min
0.5-1.0
Urethane Isourea
Secondary amine
Axen et al. (1967) Gribnau (1977); Lily (1976)
Ngo and Lenhoff (1980) Wilchek and Miron (1982); Drobnik et al. (1982) Wilchek and Miron (1982); Drobnik et al. (1982) Bethell et al. (1979) Kohn and Wilchek (1984) BioProbe International (1986)
aReproduced from Scouten (1987) with permission of Academic Press.
described: cyanogen bromide (Basic Protocol), p-nitrophenyl chloroformate (Alternate Protocol 1), and tresyl chloride (Alternate Protocol 2). CAUTION: Many of the reagents used in these protocols are hazardous; manipulations should be performed in a chemical fume hood. BASIC PROTOCOL
Affinity Purification of Natural Ligands
CYANOGEN BROMIDE ACTIVATION This basic protocol describes the use of cyanogen bromide (CNBr) to activate and couple proteins and other nucleophilic ligands to beaded agarose gel (also see UNIT 9.5). The ligand is readily coupled by this method, which is the best studied and most widely used technique for protein immobilization in spite of the potential lack of stability of the resulting product and the safety hazards inherent in the use of CNBr. CAUTION: CNBr is very toxic, releases toxic vapor, and is reported to create explosive side products upon storage; therefore, it should always be used in a hood and handled with appropriate CNBr- and solvent-resistant gloves. Only a fresh bottle of CNBr should be
9.3.2 Current Protocols in Protein Science
Current Protocols in Protein Science
Nitrous acid yields azide None
Amide to hydrazide O-Alkylation
Acid pH
Concentrated nitric acid
Concentrated nitric acid
Hydrazine
Triethyloxonium tetrafluoroborate
Polyethylene
Polystyrene
Nylon
aReproduced from Scouten (1987) with permission of Academic Press.
Nitrate aromatic ring
Reduce with Zn/HCl, then diazotize
Oxidizes CH2 to COOH Water soluble carbodiimide, Woodward’s reagent K
Ester to carboxylic acid Water soluble + alcohol carbodiimide, Woodward’s reagent K
For stability, the imine is reduced by borohydride
Polyester
Anhydrogalactose yields free aldehyde
Acid pH
For stability, the imine is reduced by borohydride
Water-soluble carbodiimide
Agarose
Diol to aldehyde
Acid pH
Nitrous acid yields azide
Periodate
Amide to carboxyl
Hydrazine
Polyacrylamide
Subsequent activation needed
Cellulose, agarose
Amide to hydrazide
Reagent
Chemical change produced
Selected Reagents to Cleave/Modify and Activate Polymeric Matrix Backbonea
Polymer
Table 9.3.2
Affinity Purification
9.3.3
High?
High
Moderate
Moderate
Low
Low
Low
Low
High
Toxicity of reagents
0.2-0.5
Variable
Variable
Variable
1-5
3
1-20
1-20
1-5
Cleavage time (hr)
Amine
Amine
Histidine tyrosine
Amine
Amine
Amine
Amine
Amine
Amine
Protein group bonded
Fair
Good
Fair
Good
Good
Good if reduced
Good if reduced
Good
Good
Stability of bond
Hornby and Goldstein (1976)
Hornby and Goldstein (1976)
Grubhofer and Schleith (1953)
Ngo et al. (1979)
Rozprimova et al. (1978)
Stults et al. (1983)
Parikh et al. (1974)
Inman and Dintzis (1969)
Inman and Dintzis (1969)
References
employed and it should be opened in the hood. Because of the problems involved in handling CNBr, it is best to dissolve a complete bottle in a defined amount of anhydrous acetonitrile. Short-term storage of this solution (several weeks) at 0°C is possible, but prolonged storage is not recommended. Materials Agarose gel (e.g., Sepharose CL-4B) 2 M and 5 M potassium phosphate, pH 12.1 2 M sodium carbonate 2 g/ml CNBr in anhydrous acetonitrile (see recipe) 1 M NaOH 0.2 M sodium bicarbonate, pH 9.0 Sample to be coupled: typically 1 to 10 mg/ml protein, 10 to 200 mg/ml small ligand, or 0.1 to 10 mg/ml nucleic acid 0.1 M potassium phosphate, pH 7.0 Solvent- and CNBr-resistant gloves 1. Wash the agarose gel well with deionized water to remove any preservatives present, then exchange it into 2 M potassium phosphate, pH 12.1, or 2 M sodium carbonate by washing it with 5 vol of the buffer. Both buffers work equally well (March et al., 1974). Normally, after washing, the agarose gel is suction filtered with suction maintained until the surface of the agarose gel just barely “cracks.” This agarose gel is weighed to obtain the “wet weight” measurement. Although this is a technique-driven procedure and the results vary from person to person, it is the most convenient and widely used method of measuring agarose gel. Other methods, e.g., “dry weight,” may be more accurate, but are highly impractical. In most instances, the agarose gel weight is not a critical factor and the person-to-person variation in “wet weight” is not significant.
2. Working in the hood, add 10 g wet weight washed agarose gel at 0°C to 10 ml of ice-cold 5 M potassium phosphate, pH 12.1, or 2 M sodium carbonate, depending on which was used in step 1. Slowly add 1 ml of a 2 g/ml solution of CNBr in anhydrous acetonitrile, with constant stirring and monitoring of the temperature. The temperature of the reaction should not exceed 10°C. This can be controlled by the rate of addition of the CNBr solution, by the use of an ice-ethanol bath around the reactor vessel, and/or by adding pieces of ice to the reaction mixture as needed. The amount of activation can be varied by increasing or decreasing the volume of CNBr solution. It is preferable to do several test experiments with the total system than to rely on formulas to predict activation levels, even though several such formulas exist (Porath, 1974). The author usually uses the weight given by the manufacturer to make up the solution, since any resulting inaccuracy is relatively unimportant. Because the reaction is much more dependent on small changes in temperature, pH, addition rate, etc., than on the precise CNBr concentration, weighing the CNBr is not worth the danger and time involved. All subsequent reactions using CNBr must be done in the hood and wearing appropriate gloves. Excess reagent should be destroyed carefully according to the Material Safety Data Sheet instructions.
3. After the required amount of CNBr is added, keep the mixture under 5°C with stirring for 5 min. Pour the mixture into a suction filtrate apparatus in the hood and filter the agarose gel from the excess reaction mixture. 4. Immediately wash the agarose gel with 5 to 10 vol ice-cold water. Affinity Purification of Natural Ligands
Destroy the contents of the filter flask, which contains excess CNBr, by slowly adding—in the hood—a 5-fold molar excess relative to the total initial CNBr used, of
9.3.4 Current Protocols in Protein Science
1 M NaOH. Keep the reaction mixture cold by adding pieces of ice. The mixture should be kept in the hood until all CNBr is destroyed.
5. Add an ice-cold solution of protein (1 to 10 mg/ml), small ligand (10 to 200 mg/ml), or nucleic acid (0.1 to 10 mg/ml) in 10 ml of 0.2 M sodium bicarbonate buffer, pH 9.0, to the activated agarose gel. Although it is possible to store CNBr-activated agarose gel and activated agarose gel in dehydrated form (i.e., the commercially available CNBr-activated agarose gel) the only practical plan in the laboratory is to couple it to proteins or other biomolecules immediately after preparation (Kohn and Wilchek, 1983b).
6. Place the agarose gel/ligand/buffer solution in a sealed bottle and rotate slowly, end over end, overnight. Collect the agarose gel by suction filtration and wash with 100 ml of 0.1 M potassium phosphate buffer, pH 7.0. The temperature depends on ligand lability; for small molecules it is usually 25°C; for proteins it is 4°C. The agarose gel/ligand can be stored, but conditions and length of storage depend on the ligand.
ACTIVATION WITH p-NITROPHENYL CHLOROFORMATE Because some materials coupled to CNBr-activated agarose gel lack stability, other activation methods are frequently employed. This alternate protocol describes the use of p-nitrophenyl chloroformate in activating agarose gel to produce a stable activated agarose gel that, when coupled to protein or nucleophilic ligands, produces reasonably strong, nonionic coupling bonds to the bound ligand.
ALTERNATE PROTOCOL 1
Additional Materials (also see Basic Protocol) 30:70 and 70:30 (v/v) acetone/water Acetone, reagent grade, room temperature and ice cold Anhydrous acetone (dried over 4 Å molecular sieves) Anhydrous acetonitrile, room temperature and 4°C (see recipe) p-Nitrophenyl chloroformate, 800 mg in 12 ml anhydrous acetonitrile Dimethylaminopyridine 5% acetic acid in dioxane, ice cold Methanol, ice cold Anhydrous isopropanol, ice cold 0.5 and 0.1 M potassium phosphate, pH 7.5 0.1 M ethanolamine, pH 7.5 Prepare the agarose gel 1. Wash the agarose gel (Sepharose CL-4b, 10 g wet weight) on a sintered glass funnel successively with 100 ml of each of the following ice-cold solutions: 30:70 (v/v) acetone/water 70:30 (v/v) acetone/water 100% reagent-grade acetone anhydrous acetone anhydrous acetonitrile. 2. Suspend the agarose gel in 12 ml of 4°C anhydrous acetonitrile containing 800 mg p-nitrophenyl chloroformate. Slowly add 3 equivalents of dimethylaminopyridine in anhydrous acetonitrile and stir the mixture for 1 hr at 0° to 4°C. Affinity Purification
9.3.5 Current Protocols in Protein Science
Supplement 4
3. Wash the agarose gel beads successively with 100 ml of each of the following ice-cold solutions: reagent-grade acetone 5% acetic acid in dioxane methanol anhydrous isopropanol. 4. Store the agarose gel beads at 0° to 4°C in anhydrous isopropanol until used. If kept dry, the agarose gel beads may be stored in anhydrous isopropanol for 6 to 12 months.
Perform the coupling reaction 5. On a suction funnel, wash the stored agarose gel (10 g) with ice-cold distilled water to remove all isopropanol. 6. Transfer the agarose gel to 10 ml of 0.1 M potassium phosphate buffer, pH 7.5, containing the ligand at 1 to 10 mg/ml protein, 0.1 to 10 mg/ml nucleic acid, or 10 to 50 mg/ml low-molecular-weight nucleophilic ligand. 7. Place the agarose gel in a bottle, cap the bottle, and tumble it end over end at 4°C for 24 to 48 hr. With stable ligands, 25°C for 16 hr will probably be adequate.
8. Wash the agarose gel by suction filtration with 100 ml of 0.1 M phosphate buffer, pH 7.5. 9. Transfer the agarose gel to 100 ml of 0.1 M ethanolamine, pH 7.5. For stable low-molecular-weight ligands, solvents with higher pH values, e.g., pure ethanolamine, may be used.
10. Tumble overnight as in step 7. 11. Wash the agarose gel with 0.1 M potassium phosphate, pH 7.5. 12. Add a small sample of the agarose gel to 0.1 M NaOH. If an intense bright yellow color develops, repeat steps 9 to 11. 13. Assay the immobilized ligand as appropriate for its activity. ALTERNATE PROTOCOL 2
ACTIVATION WITH TRESYL CHLORIDE This alternate protocol describes the use of tresyl chloride to activate and couple proteins and other nucleophilic ligands to beaded agarose gel. This method is often employed to secure the ligand to the matrix by an exceptionally stable bond. It also produces an activated intermediate that can be stored for at least 6 months in 1 mM HCl at 0°C. Additional Materials (also see Basic Protocol) Pyridine (stored over NaOH pellets; see recipe) Tresyl chloride 5 mM and 1 mM HCl Protein ligand sample: typically 1 to 10 mg/ml of protein or 10 to 50 mg/ml low-molecular-weight nucleophilic ligand 0.2 M potassium phosphate, pH 8.0 1 M glycine, pH 8.0
Affinity Purification of Natural Ligands
9.3.6 Supplement 4
Current Protocols in Protein Science
1. Wash 10 g (wet weight) agarose gel successively with 100 ml of each of the following ice-cold solutions: 30:70 (v/v) acetone/water 70:30 (v/v) acetone/water 100% reagent-grade acetone anhydrous acetone anhydrous acetonitrile. Although totally anhydrous conditions are not essential, the use of anhydrous solvents is necessary for maximum activation.
2. Add the agarose gel to a bottle containing 5 ml anhydrous acetone and 0.2 ml pyridine (stored over NaOH pellets) and cool the mixture to 0°C in an ice bath. 3. Add 0.25 ml tresyl chloride to the mixture slowly with stirring. 4. After 10 min, wash the agarose gel with 100 ml of each of the following ice-cold solutions given in step 1, but adding 5 mM HCl to the aqueous component of the acetone/water mixtures: anhydrous acetonitrile anhydrous acetone 100% reagent-grade acetone 70:30 (v/v) acetone/water 30:70 (v/v) acetone water. 5. Wash the agarose gel with 100 ml ice-cold 1 mM HCl and store the agarose gel beads at 4°C in the same dilute HCl solution. 6. Dissolve the ligand to be bound in 10 ml of 0.2 M potassium phosphate buffer, pH 8.0. Usually from 1 to 10 mg/ml protein or 10 to 50 mg/ml low-molecular-weight nucleophilic ligand is employed.
7. Rapidly rinse the activated agarose gel with 100 ml ice-cold 0.2 M phosphate, pH 8.0. 8. Mix the washed agarose gel beads with the ligand solution. If a protein or temperature-sensitive ligand is employed, rotate the mixture at 4°C overnight. For more robust ligands and/or for higher yields, higher temperatures (25°C) can be employed.
9. Wash the coupled agarose gel beads with 0.2 M potassium phosphate buffer, pH 8.0, at the coupling temperature, and deactivate any residual tresyl groups by incubating the agarose gel overnight with 1 M glycine, pH 8.0. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Cyanogen bromide (Kohn and Wilchek, 1983b) CNBr is commercially available as a 5.0 M (∼530 mg/ml) solution in acetonitrile (Aldrich). Because CNBr is volatile and very hazardous, the premade solution is preferred. The Basic Protocol can be adjusted by using 3.8 ml of the 5 M solution in place of each milliliter of 2 g/ml CNBr needed. If, however, solid CNBr is purchased, it is most convenient and usually safest to make an appropriate (2 g/ml) solution in anhydrous acetonitrile. continued
Affinity Purification
9.3.7 Current Protocols in Protein Science
For example, a 25-g bottle of fresh CNBr may be cooled in a refrigerator overnight (stored upside down, since CNBr lyophilizes and crystallizes on the inside of the lid if stored refrigerated right side up), and then transferred to a hood where it is carefully opened and 50 ml of anhydrous acetonitrile at 0° to 4°C are added. The mixture is stirred with a magnetic stir bar until all of the CNBr is dissolved. The solution should be used within 2 to 4 weeks of preparation and kept at 0° to 4°C until used. As hydrogen cyanide can develop from the reaction of CNBr with moisture, it should never be stored in a cold room. NOTE: Because CNBr is very hazardous, all precautions noted in the Material Safety Data Sheet (MSDS) should be followed precisely and disposal should be done in accordance with the recommendations! The MSDS should be carefully read before handling CNBr. Old bottles should not be used, nor should CNBr be stored for a prolonged time, as it is reported that age will cause brown discoloration and the formation of explosive degradation products (see Kohn and Wilchek, 1983b).
Anhydrous solvents Anhydrous solvents can be prepared by a variety of methods, the most thorough of which are described by Vogel (1978) and Fieser and Fieser (1967). For our purposes, however, the following procedures are usually adequate. Acetonitrile. Using the commercial anhydrous solvent, to which 4 Å molecular sieves are added for storage, is probably the best procedure. A new bottle of anhydrous acetonitrile is opened and enough 4 Å molecular sieves are added to cover the bottom of the bottle to a depth of 1⁄4 to 1⁄2 in. Pyridine. The same procedure as for acetonitrile is applied, using a new bottle of anhydrous-grade pyridine and adding NaOH pellets instead of molecular sieves. If reagent-grade pyridine is used, NaOH pellets are added and the bottle is allowed to stand at 25°C for 2 to 3 days with occasional shaking (once or twice a day) before using. Acetone. Acetone is the most difficult solvent to dry and, no matter how dried, cannot be expected to be totally anhydrous. One procedure is to add 1⁄2 to 1 in. of calcium sulfate (not calcium chloride) to the bottom of 500 ml of acetone in a reagent bottle. The acetone is mixed occasionally by shaking and kept for 4 to 8 hr. It is then filtered and stored in a reagent bottle with 1⁄2 to 1 in. of 4 Å molecular sieves. This is not totally anhydrous, but works for these procedures. Consult Vogel (1978) if more stringent conditions are desired. COMMENTARY Background Information
Affinity Purification of Natural Ligands
Affinity chromatography has benefited both basic research and biotechnology. Without this tool, purification of restriction enzymes, antibodies, and polymerases, for example, would have been much more difficult. Affinity chromatography is a very diverse method of protein purification, encompassing many combinations of matrices, ligands, methods of activation, and coupling (Turkova, 1993). The basic considerations in designing an affinity chromatography system include the final application of the product, various cost and safety considerations, and potential scale-up concerns. These must all be evaluated when affinity systems are initially planned. The ap-
plication of the product dictates the amount of nonspecific binding that can be tolerated. Cost and safety considerations determine whether the matrix can have “acceptable” leakage levels and, therefore, whether inexpensive and less durable or expensive and therapeutically safe materials should be used. Scale-up considerations determine the porosity and mechanical stability required of the matrix. Matrix considerations The ideal matrix has sufficient porosity to maintain flow, mechanical and chemical stability, high capacity, and very low nonspecific absorption. Such an ideal matrix does not yet exist. The most commonly used matrices are
9.3.8 Current Protocols in Protein Science
naturally occurring hydroxylic polymers, such as agarose, cellulose, and cross-linked dextrans. Agarose gel, the most widely used matrix, is well suited to laboratory benchtop experiments in which small columns are employed, and in which many matrices containing different activation levels, ligands, and functions are often used. For scale-up, however, synthetic organic polymers may be preferred. A new material on the market, Emphaze (3M Company; Coleman et al., 1990), is a preactivated polymer of a vinyl azolactone. Its preactivation makes it very convenient to use and it exhibits relatively low nonspecific binding and high mechanical stability. Its usable flow rate, however, may be less than that of two-component systems, such as HyperD (BioSepra), Poros (PerSeptive Biosystems; Ugelstad et al., 1983; Afeyen et al., 1990, 1991; Regnier, 1991), and
Mono-Q/Mono-S (Pharmacia Biotech; Ugelstad et al., 1983). These matrices have relatively fast throughput rates due to the fact that they are multicomponent materials with large pores but reasonably high capacity. With the “gel-in-ashell” HyperD system, shown in Figure 9.3.2, the synthetic base provides the necessary mechanical stability and the coating of dextran provides an open hydroxylic framework, which minimizes nonspecific binding and maximizes capacity. Inorganic materials such as silica have often been successfully used in affinity systems when HPLC affinity chromatography is employed. Unfortunately, for large-scale purifications of proteins with a high degree of purity, the nonspecific binding exhibited by virgin silica may be a problem.
protein binding sites biomolecule diffusion solution flow
gel shell
Figure 9.3.2 Rigid HyperD media “gel-in-a-shell.” Illustration courtesy of BioSepra, Inc. HyperD is a trademark of BioSepra, Inc.
Affinity Purification
9.3.9 Current Protocols in Protein Science
Affinity Purification of Natural Ligands
Choice of ligand Biological ligands used in affinity chromatography can be macromolecular, e.g., proteins and nucleic acids, or low-molecular-weight biomolecular, e.g., substrates, inhibitors, cofactors, antigenic peptides, “phage display” peptides, and biomimetic dyes. Each of these has a biologically relevant interaction with the molecules being purified. Purification based on such immobilized ligands is termed “bioselective adsorption,” whereas purification based on resins that bind to a desired molecule via chemical rather than biological interaction is termed “chemiselective adsorption” (Scouten, 1981). Examples of the latter include ion exchange, hydrophobic, boronate, and active thiol (“covalent chromatography”) functions bound to a solid matrix. The choice of ligand depends upon which interaction is most nearly unique to the molecule to be purified relative to the other biomolecules in the sample. For example, if it is one of the few glycoproteins in a mixture, a boronate column—which binds selectively to vicinol diols, e.g., sugar groups of glycoproteins—can be used to effect significant purification by a chemiselective process. Boronates bind glycoproteins via weak covalent bonds (Bergold and Scouten, 1983) such that an equilibrium constant has little meaning. In contrast, biological ligands go from weak to strong noncovalent bonds; they may interact so weakly that a column of immobilized ligands retards the biomolecule being purified only slightly, or, conversely, it may bind almost covalently, as is the case with the biotin-avidin pair. Ligands at either end of the spectrum are unlikely to be useful for normal purification procedures. To be useful, the ligand should bind reasonably selectively, with equilibrium constants Ka from 104 to 108 M−1 (Turkova, 1993). Although affinity chromatography is considered to be specific for a particular ligandprotein pair, and thus produces remarkable purification in a single step (and sometimes it does!), many nonspecific factors decrease its effectiveness. First, most immobilized ligands have biologically relevant interactions with several proteins—e.g., glucose binds several lectins, many enzymes, and several receptors. Because a given cell-free extract may contain several proteins that bind a single ligand, affinity systems are unlikely to produce a single protein product. In addition, other proteins in the mixture may have an affinity for the immobilized ligand due to nonspecific attraction to the matrix, to the spacer arm, or to charged or hydrophobic components of the ligand. Such
nonspecific interactions need to be carefully considered, and minimized, in the design of the affinity media, either by trial and error or by using a matrix known for having little nonspecific binding. Finally, the ligands must be immobilized such that the protein-ligand interaction can still occur after immobilization, e.g., by immobilizing the ligand by means of a nucleophilic residue distant from the point of ligand-protein interaction. Often this means designing special ligands with a nucleophilic attachment site at an appropriate location on the ligand. Spacers In many cases it is essential to have a “spacer” between the matrix and the ligand. The spacer is added to eliminate steric hindrance by the matrix, which prevents the ligand from binding the enzyme (see Fig. 9.3.3). To determine if a spacer is needed, immobilize the ligand without it. If the resulting matrix will not capture the desired protein, new matrices with spacer arms must be created. If a spacer is needed, a 2- to 6-carbon diamine is usually coupled either to the ligand or the matrix. The use of bisepoxides creates a built-in spacer and obviates the need for the added step of preparing a spacer. More extensive discussion of spacers is provided by Hermanson et al. (1992). Ligand classes There are two distinct classes of ligands (Turkova, 1993): specific ligands that have a reasonably unique interaction (remembering the caveat cited above) for one or a few proteins, and general ligands that, as a result of biological functions, bind to a large class of protein. The former group includes antibodies, most receptors, and many enzymes; the latter group includes lectins, lectin-sugar interactions, and similar broad group–binding proteins. Often a sequence of two affinity steps—a broad group– based one, e.g., the purification of essentially all dehydrogenases on Cibacron Blue (UNIT 9.2; Clovis et al., 1987), followed by chromatography on a specific immobilized inhibitor— forms the core of a very simple and efficient purification procedure. Activation and coupling methods Hydroxylic matrices—the most commonly used matrices—are polysaccharides such as beaded agarose gel (Axen et al., 1967) and cellulose (Peška et al., 1976). These are generally chemically stable and exhibit minimal non-
9.3.10 Current Protocols in Protein Science
A
B
Figure 9.3.3 Effect of spacer arms on enzyme binding to immobilized ligands. (A) Direct attachment—the enzyme active site is prevented from reaching the substrate. (B) Attachment through a spacer arm—the enzyme active site can freely interact with the substrate. Reproduced from Scouten (1983).
specific adsorption. Most importantly, however, a wide variety of ligands, usually nucleophiles, can be readily coupled to them. Attaching ligands to hydroxylic matrices, e.g., agarose gel, occurs in two steps: first, activation of the matrix, and second, the coupling process per se. CNBr activation (Axen et al., 1967; Cuatrecasas, 1970; Porath et al., 1973; Porath, 1974; March et al., 1974; Kohn and Wilchek, 1982, 1983), as described in the Basic Protocol, is the most widely used method, primarily because it is the most studied system. The ligand is readily coupled by this method, but, unfortunately, it is undesirable from the point of view of the safety hazards involved in the activation procedure and the lack of stability of the resulting bond (Parikh et al., 1974; National Research Council, 1981). Furthermore, the procedure generates a positive isourea in the coupling process, which may impart some ion-exchange properties to the resulting product (Fig. 9.3.4A). p-Nitrophenyl chloroformate (Wilchek and Miron, 1982; Miron and Wilchek, 1987), as described in Alternate Protocol 1, can be used to activate agarose to produce a stable activated agarose gel that when coupled to protein or nucleophilic ligands produces reasonably strong, nonionic bonds to the ligand. Tresyl chloride (Nilsson and Mosbach, 1980, 1981, 1987; Scouten et al., 1987), as described in Alternate Protocol 2, can be used
to activate agarose, resulting in a reasonably stable, but reactive, ester. This can subsequently be coupled to a ligand by an exceptionally stable bond (Fig. 9.3.4B). Other coupling procedures for hydroxylic matrices worth noting include carbonyldiimidazole (Hearn, 1987), bisepoxide (Sundberg and Porath, 1974), and imine (Stults et al., 1983) systems. An additional method of creating an activated matrix with synthetic polymers is to form the polymer by polymerizing preactivated monomers. A recent example of this type of synthetic matrix is the previously discussed vinyl azolactone copolymer marketed as Emphaze (3 M Company and Pierce); other new synthetic matrices include two-component systems such as HyperD (gel-in-a-shell) and the macroporous-microporous material Poros, both of which have excellent mechanical stability and fast flow characteristics (see Matrix Considerations). These, along with Pharmacia Biotech’s hardened agarose, Superose and Sepharose Fast Flow, are being used in industrial affinity chromatography systems.
Critical Parameters As noted in Reagents and Solutions, CNBr is very toxic and hazardous. The Material Safety Data Sheet (MSDS) should be read carefully and all precautions should be carefully followed. Note the following in particular (also see Reagents and Solutions):
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9.3.11 Current Protocols in Protein Science
Supplement 2
A ⊕ NH2 H
Br OH + C
N
H2N O
C
L
N
O
C
N
L
activated cyanogen (usually used immediately)
B O CH2OH + CF3
CH2
S
CH2 O
Cl
O
H2N CH2
N
L
S
CH2 CF3
activated tresyl ester (can be stored for 1 year under proper conditions)
L
H very stable nonionic secondary amine bond between agarose and ligand
Figure 9.3.4 Activation reactions. (A) CNBr activation; (B) tresyl chloride coupling.
Affinity Purification of Natural Ligands
1. CNBr is very toxic and volatile and releases toxic cyanide fumes. 2. Because CNBr sublimes, bottles should be stored (briefly) in the refrigerator upside down. 3. CNBr should be used only in the hood, with proper equipment, per MSDS instructions, and never in a cold room. 4. Excess CNBr should be destroyed per the MSDS. 5. CNBr should not be stored for prolonged periods. 6. Use of solutions of CNBr in anhydrous acetonitrile is the most convenient method; use of prepared solutions (Aldrich) avoids some of the problems inherent in handling CNBr. Because of the many variables that influence coupling/activation yields, neither activation nor coupling of ligands to solid matrices is precisely reproducible. For maximum reproducibility, the temperature, rate of addition and mixing, pH (where appropriate), moisture (in anhydrous solvents), and reagent and/or ligand concentration must all be carefully controlled, which is not easy to do. Normally a range of
activation and coupling efficiencies is observed, largely because hydrolysis of the activated matrix is a significant process that competes with activation and coupling and that can, sometimes, even be the major reaction. Moreover, in coupling, the reactivity of nucleophilic residues of apparently similar ligands may be quite different, leading to dissimiliar coupling rates and efficiencies. In spite of this, sufficient control of the conditions is possible to achieve reasonably reproducible results for a given system, although results and the degree of reproducibility will vary from system to system.
Troubleshooting It is wise to assay the final degree of activation when using the system described in the Alternate Protocols. This can be done qualitatively for p-nitrophenyl chloroformate (Alternate Protocol 1) by adding a small sample of the agarose gel to 0.1 M NaOH. If activation is successful, assuming the washes in step 3 of Alternate Protocol 1 were carried out as described, a bright yellow color will develop. Activation in either Alternate Protocol can be
9.3.12 Supplement 2
Current Protocols in Protein Science
quantified by elemental analyses for nitrogen (Alternate Protocol 1) or fluoride (Alternate Protocol 2). For CNBr activation, methods of analysis are available (Kohn and Wilchek, 1983b), but usually it is best to base activation success on ligand coupling efficiency. A common problem in activation and coupling lies in the competition between activation or coupling and hydrolysis. This is usually the result of failing to control temperature, pH, or the dryness of anhydrous solvents, where applicable. Temperature during activation can be kept sufficiently low if mixing is thorough, reactants are added slowly (monitoring temperature), and the reaction vessels are held in an ice bath. CNBr activation mixtures can also be cooled by adding ice whenever the temperature exceeds 5°C.
Anticipated Results Perhaps the best way to gain an appreciation of the range of potential activation and coupling levels is to check the catalogs of major suppliers (e.g., Pierce, Sigma, and Pharmacia Biotech), where, for example, tresyl chloride is reported in terms of its ability to couple chymotrypsin (4 to 8 mg/ml of 4% cross-linked beaded agarose gel), p-nitrophenyl chloroformate is reported in micromoles of activation (10 to 30 µmol/ml of 4% cross-linked beaded agarose gel), and amino acids are immobilized on CNBr-activated 4% beaded agarose gel at from 1 µmol/ml (cysteine) to 20 µmol/ml (histidine). Such broad ranges represent, primarily, the extremes of batch-to-batch and ligand-to-ligand variations. Unfortunately, these values are based sometimes on wet weight and sometimes on dry weight agarose gel; thus, care needs to be taken in interpreting reported activation and/or coupling yields. Moreover, the matrix has a significant effect on yield. For example, Sepharose 4B is a 4% agarose gel with large pore diameters and small surface area, whereas Sepharose 6B is a 6% agarose gel with small pore diameters and large surface area. At a given activation level, Sepharose 6B would couple more of a small ligand, and Sepharose 4B, with its larger pores, might couple more of a very large ligand, e.g., a large protein or nucleic acid. In addition, cross-linked agarose gels (e.g., Sepharose CL-4B; CL = crosslinked) have smaller pores than non-crosslinked agarose gels. Given the many variables—pore size, surface area, ligand size, ligand reactivity, and lability—it is difficult to predict potential yields for established systems, let alone for
systems involving the immobilization of ligands that have never been immobilized. The best suggestion is to peruse catalogs and the literature for systems (e.g., agarose gel type and ligand type) that most closely approximate the one to be investigated and to use such a system as the best “predictor” of potential results.
Time Considerations For the Basic Protocol, activation takes approximately 10 min, washing 10 to 20 min, and coupling is overnight, followed by washing 10 to 15 min. For Alternate Protocol 1, washing the agarose gel takes 20 to 30 min, activation takes 1 hr, and washing requires 30 to 40 min. For coupling, a 5-min wash is followed by a 24- to 48-hr incubation, a 10-min wash, and an overnight deactivation. This is followed by a 10- to 20-min wash and a 5-min test for completeness of deactivation. For Alternate Protocol 2, washing the agarose gel takes 20 to 30 min, and activation requires 10 to 15 min, followed by 30 to 40 min of washing. For coupling, 2 to 3 min is needed for the rapid wash, after which the ligand solution is added to the agarose beads, mixed (2 to 3 min) and the mixture is incubated overnight. This is followed by washing, which takes 20 to 30 min, with a subsequent overnight deactivation step.
Literature Cited Afeyen, N.B., Gorden, N.F., Maszaroff, I., Varady, L. Fulton, S.P., Yang, Y.B., and Regnier, F.E. 1990. Flow-through particles for the high-performance liquid chromatographic separation of biomolecules: Perfusion chromatography. J. Chromatogr. 519:1-29. Afeyen, N.B., Fulton, S.P., and Regnier, F.E. 1991. Perfusion chromatography packing materials for protein and peptides. J. Chromatogr. 544:257279. Axen, R., Porath, J., and Ernback, S. 1967. Chemical coupling of peptides and proteins to polysaccharides by means of cyanogen halides. Nature 214:1302-1304. Bergold, A. and Scouten, W.H. 1983. Boronate chromatography. In Solid Phase Biochemistry: Analytical and Synthetic Aspects (W.H. Scouten, ed.) pp. 149-188. John Wiley & Sons, New York. Bethell, G.S., Ayers, J.S., Hancock, W.S., and Hearn, M.T.W. 1979. A novel method of activation of cross-linked agaroses with 1,1′-carbonyldiimidazole which gives a matrix for affinity chromatography devoid of additional charged groups. J. Biol. Chem. 254:2572-2574. BioProbe International. 1986. U.S. Patent 4,582,875. Clovis, Y.D., Atkinson, A., Bruton, C.J., and Lowe, C.R. 1987. Reactive Dyes in Protein and Enzyme Technology. Stockton Press, New York.
Affinity Purification
9.3.13 Current Protocols in Protein Science
Coleman, P.L., Walker, M.M., Milbroth, D.S., Stauffer, D.M., Rasmussen, J.K., Krepski, L.R., and Heilmann, S.M. 1990. Immobilization of protein-A at high-density on azolactone—functional polymeric beads and their use in affinity chromatography. J. Chromatogr. 512:345-363. Cuatrecasas, P. 1970. Protein-purification by affinity chromatography. Derivatizations of agarose and polyacrylamide beads. J. Biol. Chem. 245:3059-3065. Domen, P.L., Nevens, J.R., Mallia, A.F., Hermanson, G.T., and Klenk, D.C. 1990. Site-directed immobilization of proteins. J. Chromatogr. 510:293-302. Drobnik, J., Labsky, J., Kudwasarova, H., Saudek, V., and Svec, F. 1982. The activation of hydroxy groups of carriers with 4-nitrophenyl and N-hydroxysuccinimide chloroformates. Biotechnol. Bioeng. 24:487-493. Fieser, L.F. and Fieser, M. 1967. Reagents for Organic Synthesis. John Wiley & Sons, New York. Gribnau, T.C.J. 1977. Coupling of effector-molecules to solid supports. Ph.D. Thesis, University of Nijmegen, Nijmegen, The Netherlands. Grubhofer, N. and Schleith, L. 1953. Modifizierte Ionenaustauscher als spezifische Adsorbentien. Naturwissenschaften 40:508. Hearn, M.T.W. 1987. Carbonyldiimidazole-mediated immobilization of enzymes and affinity ligands. Methods Enzymol. 135:102-117. Hermanson, G.T., Mallia, A.K., and Smith, P.K. 1992. Immobilized Affinity Ligand Techniques. Academic Press, New York. Hornby, W.E. and Goldstein, L. 1976. Immobilization of enzymes on nylon. Methods Enzymol. 44:118-134. Inman, J.K. and Dintzis, H.M. 1969. The derivatization of cross-linked polyacrylamide beads. Controlled introduction of functional groups for the preparation of special-purpose, biochemical adsorbents. Biochemistry 8:4074-4082. Kohn, J. and Wilchek, M. 1982. Mechanism of activation of Sepharose and Sephadex by cyanogen-bromide. Enzyme Microb. Technol. 4:161163. Kohn, J. and Wilchek, M. 1983a. Para-nitrophenylcyanate—an efficient, convenient and nonhazardous substitute for cyanogen-bromide as an activating agent for Sepharose. Appl. Biochem. 8:277-285. Kohn, J. and Wilchek, M. 1983b. Activation of polysaccharide resins by CNBr. In Solid Phase Biochemistry: Analytical and Synthetic Aspects (W.H. Scouten, ed.) pp. 599-630. John Wiley & Sons, New York. Kohn, J. and Wilchek, M. 1984. The use of cyanogen bromide and other novel cyanylating agents for the activation of polysaccharide resins. Appl. Biochem. Biotechnol. 9:285-305. Affinity Purification of Natural Ligands
Lawson, T.G., Regnier, F.E., and Weith, H.L. 1983. Separation of synthetic oligonucleotides on columns of macro particulate silica coated with crosslinked polyethylene imine. Anal. Biochem. 133:85-93. Lily, M.D. 1976. Enzymes immobilized to cellulose. Methods Enzymol. 44:46-53. March, S.C., Parikh, I., and Cuatrecasas, P. 1974. Cyanogen-bromide activation of agarose for affinity chromatography. Anal. Biochem. 60:149152. Miron, T. and Wilchek, M. 1987. Immobilization of protein and ligands using chloroformates. Methods Enzymol. 135:84-90. Murayama, K., Shimada, K., and Yamamoto, T. 1978. Modification of immunoglobulin-G using specific reactivity of sugar moiety. J. Immunochem. 15:523-528. National Research Council, Committee on Hazardous Substances in the Laboratory. 1981. Prudent Practices for Handling Hazardous Chemicals in Laboratories. National Academy Press, Washington, D.C. Ngo, T.T. and Lenhoff, H.M. 1980. Immobilization of enzymes through activated peptide bonds of protein supports. J. Appl. Biochem. 2:373-379. Ngo, T.T., Laidler, K.J., and Yam, C.F. 1979. Kinetics of acetylcholinesterase immobilized on polyethylene tubing. Can. J. Biochem. 57:1200-1203. Nilsson, K. and Mosbach, K. 1980. p-Toluenesulfonyl chloride as an activating agent of agarose for the preparation of immobilized affinity ligands and proteins. Eur. J. Biochem. 112:397402. Nilsson, K. and Mosbach, K. 1981. Immobilization of enzymes and affinity ligands to various hydroxyl group carrying supports using high reactive sulfonyl chlorides. Biochemistry 102:449457. Nilsson, K. and Mosbach, K. 1987. Tresyl chloride– activated supports for enzyme immobilization. Methods Enzymol. 135:65-78. Parikh, I., March, S., and Cuatrecasas, P. 1974. Topics in the methodology of substitution reactions with agarose. Methods Enzymol. 34:70-102. Peška, J., Štamberg, J., Hradil, J., and Ilavsky, M. 1976. Cellulose in bead form: Properties related to chromatographic uses. J. Chromatogr. 125:455-469. Porath, J. 1974. Preparation of cyanogen bromide-activated agarose gels. Methods Enzymol. 34:13-30. Porath, J., Aspberg, K., Drevin, H., and Axen, R. 1973. Preparation of cyanogen bromide activated agarose gels. J. Chromatogr. 86:53-56. Regnier, F.E. 1991. Perfusion chromatography. Nature 350:634-635. Rozprimova, L., Franek, F., and Kubanek, V. 1978. Utilization of powder polyester in making insoluble antigens and pure antibodies. Czech Epidemiol. Mikrobiol. Immunol. 27:335-341.
9.3.14 Current Protocols in Protein Science
Scouten, W.H. 1981. Affinity Chromatography: Bioselective Absorption on Inert Matrices. Wiley-Interscience, New York. Scouten, W.H. 1983. Solid Phase Biochemistry: Analytical and Synthetic Aspects. John Wiley & Sons, New York. Scouten, W.H. 1987. Immobilization techniques for enzymes. Methods Enzymol. 135:30-65. Scouten, W.H., Van den Tweel, W., Kromenburg, H., and Dekker, M. 1987. Colored sulfonyl chloride as an activating agent for hydroxylic matrices. Methods Enzymol. 135:79-84.
Vogel, A. 1978. Textbook of Practical Organic Chemistry, 4th ed. Longman, New York. Voivodov, K., Chan, W.-H., and Scouten, W.H. 1993. Chemical approaches to oriented immobilization. Makromol. Chem. 70/71:275-283. Wilchek, M. and Miron, T. 1982. Immobilizations of enzymes and affinity ligands onto agarose via stable and uncharged carbamate linkages. Biochem. Int. 4:629-635.
Key References Hermanson, et al., 1992. See above.
Stults, N.L., Lin, P., Hardy, M., Lee, Y.G., Uchida, Y., Tsukada, Y., and Sugimori, T. 1983. Immobilization of proteins on partially hydrolyzed agarose beads. Anal. Biochem. 135:392-400.
A very good practical laboratory manual with good description of methods.
Sundberg, L. and Porath, J. 1974. Preparation of adsorbents for biospecific affinity chromatography: Attachment of group-containing ligands to insoluble polymers by means of bifunctional oxiranes. J. Chromatogr. 90:87-98.
An older, simple approach with a good mix of theory and practical laboratory methods.
Turkova, J. 1993. Bioaffinity Chromatography, 2nd ed. Elsevier, Amsterdam. Ugelstad, J., Söderberg, L., Berge, A., and Bergstrom, J. 1983. Monodisperse polymer particles: A step forward for chromatography. Nature 303:95-96.
Scouten, W.H. 1981. See above.
Turkova, J. 1993. See above. The most recent and comprehensive volume available in the field.
Contributed by William H. Scouten Utah State University Logan, Utah
Affinity Purification
9.3.15 Current Protocols in Protein Science
Metal-Chelate Affinity Chromatography
UNIT 9.4
Recombinant proteins engineered to have six consecutive histidine residues on either the amino or carboxyl terminus can be purified using a resin containing nickel ions (Ni2+) that have been immobilized by covalently attached nitrilotriacetic acid (NTA). This technique, known as metal-chelate affinity chromatography (MCAC), can readily be performed with either native or denatured protein. The Strategic Planning section discusses techniques for creating a fusion protein consisting of the protein of interest with a histidine tail attached (for purification by MCAC; see UNIT 6.5 for an example purification of HIV-1 integrase). The Basic Protocol describes expression of histidine-tail fusion proteins and their purification in native form by MCAC. Two alternate protocols describe purification of histidine-tail fusion proteins by MCAC under denaturing conditions and their renaturation by either dialysis or solid-phase renaturation. Support protocols are provided for analysis of the purified product and regeneration of the NTA resin. All of these protocols are easily adaptable to any protein expression system. Prior to large-scale preparation, the cells should be tested for expression of the protein in soluble form (see UNITS 6.1 & 6.2). Even if protein is expressed mostly in insoluble form (i.e., bacterial inclusion bodies; UNIT 6.3), there may be a small fraction that remains soluble and can therefore be purified using the Basic Protocol. If little or no soluble protein is observed, one of the denaturing protocols (Alternate Protocols 1 and 2) should be used. NOTE: All solutions and equipment coming into contact with live cells must be sterile. STRATEGIC PLANNING MCAC is designed to purify a specific protein whose complementary DNA (cDNA) sequence is available. A protein-expression system is selected and the cDNA is inserted into the appropriate expression vector (see UNIT 5.1). The cDNA sequence must encode a minimum of six histidines at either the amino or carboxyl terminus and must include an initiator methionine at the amino terminus and a termination codon at the carboxyl terminus. This can be accomplished by polymerase chain reaction (PCR) using primers
A
5 ′-GGGNNNNNNATGCATCATCATCATCATCAT . . . N15-30-3′ RE s i t e – Me t H i sH i sH i sH i sH i sH i s . . .
B
5 ′-GGGNNNNNNTTAATGATGATGATGATGATG . . . N15-30-3′ RE s i t e – ENDH i sH i s H i s H i sH i s H i s . . .
Figure 9.4.1 Sequences of primers required to create histidine tails at protein termini. Functions (e.g., protein sequence) are marked below. Three guanines are included at the 5′ ends of each primer to facilitate restriction enzyme digestion of the PCR product prior to subcloning. NNNNNN represents a unique restriction enzyme site compatible with the selected vector. N15-30 represents 15 to 30 additional nucleotides specific to the cDNA beginning with the second codon. Met represents an initiator methionine and END represents a termination codon. (A) Sequence of 5′ primer used to create a histidine tail at the amino terminus. The 3′ primer should include a second unique restriction enzyme site and the final 5 to 10 codons (including a stop codon) of the cDNA sequence. (B) Sequence of 3′ (antisense) primer used to create a histidine tail at the carboxyl terminus. The 5′ primer should contain a second unique restriction site and the first 5 to 10 codons of the cDNA sequence. Contributed by Kevin J. Petty Current Protocols in Protein Science (1996) 9.4.1-9.4.16 Copyright © 2000 by John Wiley & Sons, Inc.
Affinity Purification
9.4.1 Supplement 4
containing unique restriction-site sequences at the 5′ end (Fig. 9.4.1). Properly designed primers will permit insertion and expression of the cDNA in the correct reading frame. An alternative to PCR is to subclone the cDNA into an existing vector, synthesize complementary oligonucleotides containing the hexahistidine sequence with compatible restriction enzyme site overhangs, and insert them into the subcloned cDNA (Struhl, 1987). It is important to insert the oligonucleotide into the cDNA near one of the termini in the correct reading frame. Several companies (e.g., Qiagen, Novagen, and Invitrogen; see SUPPLIERS APPENDIX) sell expression systems that permit direct subcloning of cDNAs into vectors already containing oligohistidine tail sequences along with adjacent protease cleavage sites to allow removal of the tail after purification. Vectors have been developed for expression of histidine fusion proteins by bacteria, yeast, baculovirus, vaccinia, and various eukaryotic promoters. The protocols that follow are designed for use with a Novagen pET vector (Fig. 9.4.2), but may easily be modified for other vectors. BASIC PROTOCOL
NATIVE MCAC FOR PURIFICATION OF SOLUBLE HISTIDINE-TAIL FUSION PROTEINS This protocol describes expression and purification of histidine fusion proteins in E. coli. A desired protein is induced in bacteria, which are then harvested and lysed. The lysate is loaded directly onto a column containing Ni2+-NTA resin and the protein is eluted in nearly pure form with MCAC buffer containing imidazole. Materials M9ZB medium (see recipe) containing 50 µg/ml ampicillin and 25 µg/ml chloramphenicol E. coli BL21(DE3)pLysS or other suitable strain (Novagen) containing a pET vector (Fig. 9.4.2) expressing a histidine-tail fusion protein 0.1 M IPTG, filter sterilized NTA resin slurry: 50% (w/v) suspension in 20% (v/v) ethanol (Qiagen) 100 mM NiSO4⋅6H2O MCAC-0, MCAC-20, MCAC-40, MCAC-60, MCAC-80, MCAC-100, MCAC-200, and MCAC-1000 buffers (see recipe) 150× protease inhibitor cocktail (see recipe) 10% (v/v) Triton X-100 1 M MgCl2 MCAC-EDTA buffer (see recipe) DNase I solution (see recipe) Centrifuge with Beckman JA-20 rotor or equivalent 1 × 10–cm glass or polypropylene column Additional reagents and equipment for growth of bacteria in liquid medium (UNIT 5.3) and analysis and processing of purified proteins by SDS-PAGE (see Support Protocol 1) Express the protein 1. Inoculate 10 ml M9ZB/ampicillin/chloramphenicol with E. coli BL21(DE3)pLysS containing a pET vector expressing a desired histidine-tail fusion protein. Grow overnight with shaking at 37°C (see UNIT 5.3).
Metal-Chelate Affinity Chromatography
2. Inoculate 100 ml M9ZB/ampicillin/chloramphenicol with 1 ml of the overnight culture and grow with shaking at 37°C to OD600 = 0.7 to 1.0.
9.4.2 Supplement 4
Current Protocols in Protein Science
A
Eco RI (5706) Cla I (24) Aat II (5635) Hin dIII (29) Ssp I (5517) Sca I (5193) Pvu I (5083) Pst I (4958)
Apr
Bsa I (4774)
T7 transcription/ expression region
Esp I (267) Bam HI (319) Xho I (324) Nde I (331) Nco I (389) Xba I (428) Bgl II (494) Sgr AI (535) Sph I (691) Eco NI (751) Drd II (939) Mlu I (1216) Bcl I (1230)
pET-15b (5708 bp)
lacl
Bst EII (1397) Apa I (1427)
Alw NI (4236) Bss HII (1627)
ori
Hpa I (1722)
Sap I (3704) Psh AI (2061)
Sna I (3591) Acc I (3590) Bsa AI (3572) Tth lllI (3565) Bpu lOI (2926)
Xma III (2284) Nru I (2319) Bsp MI (2399) Bsm I (2704) Bal I (2791)
B T7 promoter lac operator Bgl II Xba l AGATCTCGATCCCGCGAAATTAATACGACTCACTATAGGGGAATTGTGAGCGGATAACAATTCCCCTCTAGAAATAATTTTGTTTA
Nco I
Nde I
ACTTTAAGAAGGAGATATACCATGGGCAGCAGCCATCATCATCATCATCACAGCAGCGGCCTGGTGCCGCGCGGCAGCCAT Me t G l y S e r S e r H i sH i sH i sH i sH i sH i s S e r S e r G l y L e u Va l P r o A r g G l y S e r H i s thrombin Xho I Bam HI Esp I ATGCTCGAGGATCCGGCTGCTAACAAAGCCCGAAAGGAAGCTGAGTTGGCTGCTGCCACCGCTGAGCAATAACTAGCATAACCC Me t L e uG l u A s pP r o A l a A l aAs n L y s A l a A r gL y s G l u A l aG l u Le u A l a A l a A l a Th r A l a G l u G l nEnd
Eco RV CTTGGGGCCTCTAAACGGGTCTTGAGGGGTTTTTTGCTGAAAGGAGGAACTATACCGGATATC
Figure 9.4.2 pET-15b, one of a series of pET vectors designed for cloning, expression, and purification of recombinant proteins. Reprinted with the permission of Novagen. (A) pET-15b vector; (B) sequence of pET-15b cloning/expression region. The target gene is cloned into the pET plasmid such that its expression is under the control of bacteriophage T7 transcription and translation signals (see UNIT 5.2). pET-15b encodes an amino-terminal His⋅Tag leader that allows purification of the resulting recombinant protein over Ni2+-NTA resin. Following purification, the His⋅Tag can be removed by thrombin cleavage (see UNIT 6.5). The pET series are derivatives of vectors originally described by Studier et al. (1990) and are available from Novagen.
Affinity Purification
9.4.3 Current Protocols in Protein Science
Supplement 4
3. Add 1 ml of 0.1 M IPTG (to 1 mM final) and continue shaking incubation 1 to 3 hr at 37°C. Incubation time will depend on the solubility of the expressed protein. A 3-hr incubation will allow maximal expression but may result in mostly insoluble protein. A 1-hr incubation will produce less protein but more of it may be soluble.
4. Centrifuge 10 min at 4400 × g (5000 rpm in a Beckman JA-20 rotor), 4°C. Discard supernatant. Freeze pellet at −70°C. It is not necessary to wash the cell pellet. The wet weight of the cell pellet will be ∼0.5 g. The pellet can be stored indefinitely at −70°C before proceeding. Alternatively, extract preparation (steps 9 to 13) can be carried out immediately and the column prepared during the centrifugation at step 13.
Prepare the affinity column 5. Add 0.2 ml NTA resin slurry to a 1 × 10–cm column and allow liquid to drain. During this and subsequent column washes, liquid should be allowed to drain to the top of the packed resin bed that forms as resin settles (packed bed volume should be 0.1 ml) and resin should not be allowed to dry.
6. Wash column with 1 ml (5 bed volumes) deionized water. 7. Charge column by washing with 1 ml (5 bed volumes) of 100 mM NiSO4⋅6H2O. 8. Wash column with 2 ml MCAC-0 buffer. NTA resin is nitrilotriacetic acid covalently coupled to Sepharose CL-6B. When charged with nickel ions, it has a light blue-green color, and when stripped of nickel, it is white. See supplier’s instructions for resin preparation. Charged resin can be stored at 4°C. If column is to be stored >1 day, wash with 10 bed volumes of 20% ethanol and add 1 bed volume of 20% ethanol prior to storage. Keep column sealed to prevent evaporation.
Prepare the extract IMPORTANT NOTE: Beginning with this step, all procedures should be performed on ice or in a cold room unless otherwise indicated. 9. Thaw cell pellet (from step 4) on ice. Add 5 ml MCAC-0 buffer and 33 µl of 150× protease inhibitor cocktail and resuspend by pipetting, sonication, or homogenization. All MCAC buffers contain phenylmethylsulfonyl fluoride (PMSF) as a protease inhibitor; to reduce expense, protease inhibitor cocktail is added only to the crude extract.
10. Add 0.05 ml of 10% (v/v) Triton X-100 (0.1% final). Mix thoroughly and subject the sample to 3 cycles of freezing at −70°C and thawing on ice. Ionic detergents may interfere with binding of the protein to the resin and should not be used. Cell lysis is evidenced by a visible increase in viscosity. The pLysS plasmid in these cells encodes an endogenous lysozyme that eliminates the need for exogenous lysozyme treatment to disrupt the bacterial cell wall.
11. Add 0.05 ml of 1 M MgCl2 (final concentration 10 mM) and 0.05 ml of DNase I solution (final concentration 10 µg/ml DNase I). Mix gently and incubate 10 min at room temperature. Metal-Chelate Affinity Chromatography
The DNase I treatment reduces the viscosity of the lysate.
9.4.4 Supplement 4
Current Protocols in Protein Science
12. Centrifuge 15 min at 27,000 × g (15,000 rpm in JA-20 rotor), 4°C. Decant the supernatant into a clean container on ice and discard the pellet. Set aside and freeze a 10-µl aliquot at −70°C for later analysis by SDS-PAGE (see Support Protocol 1). The supernatant can be frozen at −70°C indefinitely before continuing with the procedure.
Purify protein 13. If extract is frozen, thaw on ice. Load onto Ni2+-NTA column and allow to flow through at a rate of 10 to 15 ml/hr. Collect column flowthrough and save for SDS-PAGE (see Support Protocol 1). Charged NTA resin has a capacity of 5 to 10 mg histidine-tagged protein per milliliter of packed resin. The amount of extract that can be loaded on the column will depend on the amount of soluble histidine-tagged protein in the extract.
14. Wash column with 5 ml MCAC-0 buffer at a flow rate of 20 to 30 ml/hr. Discard flowthrough. 15. Wash column in stepwise fashion with 5 ml each of MCAC-20, MCAC-40, MCAC60, MCAC-80, MCAC-100, MCAC-200, and MCAC-1000 buffers at a flow rate of 10 to 15 ml/hr. Collect 0.5-ml fractions and save on ice for SDS-PAGE (see Support Protocol 1). Alternatively, the column can be eluted with a 5-ml linear gradient of 0 to 400 mM imidazole in MCAC buffer. The second and third fractions of each wash will contain most of the eluted proteins. Most proteins with hexahistidine tails will remain bound in 60 mM imidazole (MCAC-60) and elute with 100 to 200 mM imidazole (MCAC-100 or -200); therefore, the purified protein will elute in MCAC-100 or -200. Proteins with longer histidine tails (e.g., 10 residues) bind to Ni2+-NTA with greater affinity and require higher imidazole concentrations for elution. However, optimum washing and elution conditions must be determined for each protein. Once optimum washing and elution conditions are established, it is possible to prepare the crude extract in a buffer that contains the highest imidazole concentration in which the histidine tail remains bound to the Ni2+-NTA (e.g., MCAC-40 or -60 buffer). This decreases nonspecific binding of proteins to resin and permits use of a single buffer for extract preparation, column loading, and column washing.
16. Elute column with 1 ml MCAC-EDTA buffer at a flow rate of 10 to 15 ml/hr, collecting 0.5-ml fractions. The blue-green color of the column will disappear as nickel is removed by EDTA. More tightly bound proteins may be found in these fractions. The resin can now be recharged (repeat steps 6 to 8) and the column reused. If protein is eluted adequately with imidazole (as determined by overall yield for subsequent preparations), EDTA washing can be omitted. The column can be reequilibrated with MCAC-0 buffer and the purification repeated. The same column can be used three to five times before EDTA stripping and nickel recharging are necessary. Only one protein should be purified on any given column.
17. Analyze fractions for the presence of eluted protein. An ultraviolet (280-nm) absorbance flow monitor is helpful for following column elution but is not necessary. An alternative is to measure the OD280 of individual fractions to identify protein-containing fractions. However, imidazole will also absorb at 280 nm. A quick and easy method to determine which fractions contain eluted protein is to place 2 ìl undiluted Protein Assay Dye Reagent Concentrate (Bio-Rad) on a piece of Parafilm, add 8 ìl from fraction to be tested, and mix by pipetting up and down. Immediate appearance of blue color indicates that the fraction contains protein. This does not work in the presence
Affinity Purification
9.4.5 Current Protocols in Protein Science
Supplement 4
of Triton X-100 because the detergent itself produces an intense blue color; for this reason, Triton X-100 is excluded from the washing and elution buffers.
18. Combine the fractions containing eluted protein and remove a 10-µl aliquot for SDS-PAGE (see Support Protocol 1). Freeze the remainder in smaller aliquots at −70°C or in liquid nitrogen. If a different buffer for the protein is desired (e.g., for proteolytic removal of the histidine tail), the protein should be dialyzed against the buffer of choice to remove the MCAC buffer prior to storage at −70°C or in liquid nitrogen.
19. If time permits, proceed immediately to analysis of fractions by SDS-PAGE and processing of protein (see Support Protocol 1). Otherwise, freeze all samples at −70°C until ready for analysis and processing. ALTERNATE PROTOCOL 1
DENATURING MCAC FOR PURIFICATION OF INSOLUBLE HISTIDINE-TAIL FUSION PROTEINS High-level expression of foreign proteins in bacteria and other cells frequently results in poor solubility of the expressed protein (see UNITS 5.1 & 5.2). These insoluble proteins form inclusion bodies in bacteria, and strong chaotropic agents such as guanidine, urea, or SDS are usually required to solubilize them. These agents denature the protein and destroy the secondary structure that is essential to other affinity purification methods (e.g., maltosebinding protein or glutathione-S-transferase fusion proteins). A significant advantage of metal-chelate affinity chromatography is that the oligohistidine tail will bind to the Ni2+-NTA resin even when the protein is denatured. In denaturing MCAC, the protein extract is solubilized with 6 M guanidine and the entire affinity purification procedure is carried out in guanidine. The purified, denatured protein is renatured during dialysis. Additional Materials (also see Basic Protocol) GuMCAC-0, GuMCAC-20, GuMCAC-40, GuMCAC-60, GuMCAC-100, and GuMCAC-500 buffers (see recipe) GuMCAC-EDTA buffer (see recipe) Appropriate final buffer for protein (e.g., for proteolytic cleavage or long-term storage) Guanidine⋅HCl Additional reagents and equipment for analysis and processing of purified proteins (see Support Protocol 1) and dialysis (APPENDIX 3B & UNIT 4.4) Express the protein 1. Prepare the pellet of E. coli expressing a histidine-tail fusion protein (see Basic Protocol, steps 1 to 4). The pellet can be stored indefinitely at −70°C before proceeding. Alternatively, extract preparation (steps 5 and 6) can be carried out immediately and the column prepared during the centrifugation at step 6.
Prepare the affinity column 2. Prepare column (see Basic Protocol, steps 5 to 7). 3. Wash column with 2 ml GuMCAC-0 buffer. During this and subsequent column washes, liquid should be allowed to drain to top of packed resin bed and resin should not be allowed to dry. Metal-Chelate Affinity Chromatography
9.4.6 Supplement 4
Current Protocols in Protein Science
Prepare cell extract 4. Thaw cell pellet (from step 1) on ice. Resuspend in 5 ml GuMCAC-0 buffer by pipetting, sonication, or homogenization. 5. Freeze 10 min at −70°C and thaw at room temperature. Protease inhibitors are omitted because proteases are inactivated by guanidine. Triton X-100 is not needed at this step. Freezing is not necessary but is included because it ensures complete lysis of cells. Subsequent steps can be performed at room temperature. However, if solid-phase renaturation is used (see Alternate Protocol 2), it is better to maintain lower temperatures throughout the process.
6. Gently mix samples for 30 min using a rocker, rotating mixer, or magnetic stirrer. Centrifuge 15 min at 27,000 × g (15,000 rpm in Beckman JA-20 rotor), 4°C. Decant supernatant into a clean container and discard pellet. Set aside a 10-µl aliquot for analysis by SDS-PAGE (see Support Protocol 1). The supernatant can be frozen at −70°C indefinitely before continuing with the procedure.
Purify protein 7. If extract from step 5 is frozen, thaw at room temperature. Load onto Ni2+-NTA column and allow to flow through at a rate of 10 to 15 ml/hr. Collect flowthrough and save a 10-µl aliquot for SDS-PAGE (see Support Protocol 1). 8. Wash column with 5 ml GuMCAC-0 buffer at a rate of 20 to 30 ml/hr. Discard the flowthrough. 9. Wash column in stepwise fashion with 5 ml GuMCAC-20, -40, -60, -100, and -500 buffers at a rate of 10 to 15 ml/hr. Collect 0.5-ml fractions and save for SDS-PAGE (see Support Protocol 1). The second and third fractions from each wash will contain most of the unbound protein. The histidine tail binds slightly less avidly under denaturing conditions. Lower imidazole concentrations are therefore required for washing and elution than in the Basic Protocol.
10. Elute with 1 ml GuMCAC-EDTA buffer at a rate of 10 to 15 ml/hr, collecting 0.5-ml fractions. 11. Identify fractions containing the protein, pool together, transfer to dialysis tubing, and seal. Alternatively, fractions can be frozen at −70°C indefinitely before continuing with the procedure. Guanidine precipitates in the presence of SDS and must be removed by dialysis before SDS-PAGE. An alternative technique employs buffers that switch from 6 M guanidine to 8 M urea during affinity column washing (Stüber et al., 1990). This permits samples to be taken directly from urea fractions without dialysis and analyzed by SDS-PAGE or injected into animals for antibody production.
Renature purified protein by dialysis 12. Prepare appropriate final buffer for protein (e.g., for proteolytic cleavage or long-term storage) and add sufficient guanidine to bring final concentration to 4 M. 13. Dialyze purified protein from step 11 for ≥2 hr at 4°C against 500 ml buffer/4 M guanidine (see APPENDIX 3B). The MWCO of the dialysis membrane should be chosen to be smaller than the MW of the purified protein. In most cases an MWCO of 12 to 14 kDa is sufficient. Affinity Purification
9.4.7 Current Protocols in Protein Science
Supplement 4
14. Remove 250 ml buffer/guanidine and add 250 ml buffer without guanidine. Continue dialysis ≥2 hr. Repeat. With some proteins, renaturation by dialysis may require longer dialysis periods and more gradual decrements in the guanidine concentration of the buffer. Conditions for each protein must be determined empirically.
15. Remove dialysis bag to a container containing 500 ml of fresh buffer without guanidine at 4°C. Continue dialysis 2 hr to overnight. 16. Remove sample from dialysis bag, divide into aliquots, and freeze at −70°C or in liquid nitrogen. If protein precipitates during dialysis, solid-phase renaturation (see Alternate Protocol 2), in which protein bound to the column is renatured before elution, should be employed.
17. Analyze fractions and process protein (see Support Protocol 1). ALTERNATE PROTOCOL 2
SOLID-PHASE RENATURATION OF MCAC-PURIFIED PROTEINS In Alternate Protocol 1, removal of denaturants by dialysis will occasionally lead to precipitation of protein, possibly due in part to entanglement or aggregation of separate protein molecules as they refold. To avoid this problem, solid-phase renaturation may be attempted. In this procedure, protein extract is prepared and bound to the column under denaturing conditions. A series of washes removes the denaturing agent before the target protein is eluted and the resulting renatured protein is eluted from the column under native conditions. Additional Materials (also see Basic Protocol) 1:1 (v/v) MCAC-20/GuMCAC-20 buffer (see recipes) 3:1 (v/v) MCAC-20/GuMCAC-20 buffer (see recipes) 7:1 (v/v) MCAC-20/GuMCAC-20 buffer (see recipes) 1. Prepare protein extract, bind to column, and wash with GuMCAC buffers (see Alternate Protocol 1, steps 1 to 9). 2. Wash column with 5 ml of 1:1 (v/v) MCAC-20/GuMCAC-20 buffer. During this and subsequent washes, liquid should be allowed to drain just to top of packed resin bed and resin should not be allowed to dry.
3. Wash column with 5 ml of 3:1 (v/v) MCAC-20/GuMCAC-20 buffer. 4. Wash column with 5 ml of 7:1 (v/v) MCAC-20/GuMCAC-20 buffer. 5. Wash column with MCAC buffers, elute proteins, and analyze (see Basic Protocol, steps 15 to 19). Slow elution (between 1 and 2 hr) with a 5-ml linear gradient from 100% GuMCAC-20 buffer to 100% MCAC-20 buffer may also yield efficient renaturation.
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Current Protocols in Protein Science
ANALYSIS AND PROCESSING OF PURIFIED PROTEINS The success of the purification scheme (particularly during a small pilot study) should be monitored at each stage by SDS-PAGE. If the fusion protein contains a specific protease cleavage site, the histidine tail can be removed using an appropriate proteolytic procedure, if desired.
SUPPORT PROTOCOL 1
Materials Fractions from MCAC column purification (crude extract, flowthroughs, and purified protein; see Basic Protocol or Alternate Protocols 1 or 2) 2× SDS sample buffer (UNIT 10.1) MCAC-0 buffer (see recipe) Additional reagents and equipment for one-dimensional SDS-PAGE (UNIT 10.1), cleavage of proteins with factor Xa or thrombin (UNIT 6.5), and dialysis (UNIT 4.4 & APPENDIX 3B) 1. Thaw aliquots of fractions to be analyzed on ice. Mix 5 µl from crude extract and crude flowthrough fractions and 10 µl from the second and third fractions from each washing step with an equal volume of 2× SDS sample buffer. 2. Load samples onto a standard SDS-PAGE gel. Run gel and visualize to identify the fractions containing purified protein (UNIT 10.1). Guanidine must be removed by dialysis prior to addition of SDS sample buffer.
3. Thaw the remaining aliquots of fractions containing purified protein, dialyze against the appropriate proteolysis buffer, and carry out cleavage procedure if desired. If necessary, after cleavage dialyze the protein against an appropriate storage buffer and freeze in aliquots. The size of the cleaved histidine tail will generally be <3 kDa, depending on the design of the fusion protein. This fragment will usually be removed by the dialysis. It may also be removed by ultrafiltration (which will also concentrate the protein), size-exclusion chromatography, or a second MCAC (see Basic Protocol, steps 13 to 19). The advantage of a second MCAC is that the histidine tail and any uncleaved protein will bind to the column and only the cleaved protein will be found in the flowthrough from step 12. Aliquots of each fraction should be analyzed by SDS-PAGE to verify recovery of pure protein. If an expression vector other than pET has been used, and the fusion protein does not contain an appropriate cleavage site for factor Xa or thrombin, or if the histidine tail will not interfere with subsequent studies, omit the cleavage procedure. Presence of the histidine tail will not generally affect the protein’s biologic functions (see Background Information).
Affinity Purification
9.4.9 Current Protocols in Protein Science
Supplement 4
SUPPORT PROTOCOL 2
NTA RESIN REGENERATION NTA resin can be repeatedly charged with Ni2+ and stripped. Over time, however, some protein residue will accumulate and decrease the efficiency of the resin, resulting in slow flow rates or lack of blue-green color after charging with Ni2+. Periodic regeneration (e.g., after 5 to 10 cycles of charging and stripping) permits recycling of the resin for long-term use. Materials 2.5 ml used NTA resin (packed volume) Stripping solution: 0.2 M acetic acid/6 M guanidine⋅HCl 2% (w/v) SDS 20%, 25%, 50%, 75%, and 100% (v/v) ethanol 0.1 M EDTA, pH 8.0 1. Wash resin with 5 ml stripping solution. In this and all subsequent washes, liquid added to the column should be allowed to drain to top of packed resin bed and resin should not be allowed to dry.
2. Wash with 5 ml water. 3. Wash with 7.5 ml of 2% (w/v) SDS. 4. Wash with 2.5 ml of 25% (v/v) ethanol. 5. Wash with 2.5 ml of 50% (v/v) ethanol. 6. Wash with 2.5 ml of 75% (v/v) ethanol. 7. Wash with 12.5 ml of 100% (v/v) ethanol. 8. Wash with 2.5 ml of 75% (v/v) ethanol. 9. Wash with 2.5 ml of 50% (v/v) ethanol. 10. Wash with 2.5 ml of 25% (v/v) ethanol. 11. Wash with 2.5 ml water. 12. Wash with 12.5 ml of 0.1 M EDTA, pH 8.0. 13. Wash with 7.5 ml water. Proceed either to nickel charging (see Basic Protocol, step 7) or to step 14 below for long-term storage of the resin. 14. Add 2.5 ml of 20% (v/v) ethanol to resin and store at 4°C.
Metal-Chelate Affinity Chromatography
9.4.10 Supplement 4
Current Protocols in Protein Science
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
DNase I solution Dissolve lyophilized DNase I (U.S. Biochemical) at a final concentration of 1 mg/ml in 50% glycerol/1 mM CaCl2. Store ≤1 year at −20°C. GuMCAC buffers (store ≤6 months at 4°C) GuMCAC-0 buffer: 20 mM Tris⋅Cl, pH 7.9 0.5 M NaCl 10% glycerol 6 M guanidine⋅HCl GuMCAC-500 buffer: 20 mM Tris⋅Cl, pH 7.9 0.5 M NaCl 10% glycerol 6 M guanidine⋅HCl 0.5 M imidazole GuMCAC-20, GuMCAC-40, GuMCAC-60, and GuMCAC-100 buffers: These buffers (containing different concentrations of imidazole) are made by mixing GuMCAC-0 buffer and GuMCAC-500 buffer in the appropriate ratios: e.g., for GuMCAC-20 buffer, use 96:4 (v/v) GuMCAC-0/GuMCAC-500. GuMCAC-EDTA buffer 20 mM Tris⋅Cl, pH 7.9 0.5 M NaCl 10% glycerol 6 M guanidine⋅HCl 0.1 M EDTA, pH 8.0 Store ≤6 months at 4°C M9 medium, 10× 30 g Na2PO4 15 g KH2PO4 5 g NH4Cl 2.5 g NaCl 15 mg CaCl2 (optional) H2O to 1 liter Heat with stirring until dissolved. Pour aliquots into separate bottles, cap loosely, and autoclave 15 min at 15 lb/in2. After the bottles cool to <50°C, add nutritional supplements and/or antibiotics as desired. After the bottles cool to <40°C, tighten bottle caps and store indefinitely at room temperature (discard if medium becomes contaminated). M9ZB medium Dissolve 10 g N-Z-Amine A (Sigma) and 5 g NaCl in 889 ml water. Autoclave, cool, and add 100 ml of 10× M9 medium (see recipe), 1 ml of 1 M sterile MgSO4, and 10 ml of 40% (w/v) glucose (filter sterilized). Store ≤1 year at room temperature.
Affinity Purification
9.4.11 Current Protocols in Protein Science
Supplement 4
MCAC buffers (store ≤6 months at 4°C) MCAC-0 buffer: 20 mM Tris⋅Cl, pH 7.9 0.5 M NaCl 10% glycerol 1 mM PMSF (phenylmethylsulfonyl fluoride) MCAC-1000 buffer: 20 mM Tris⋅Cl, pH 7.9 0.5 M NaCl 10% glycerol 1 M imidazole 1 mM PMSF Add PMSF immediately before use from a 0.2 M stock in 100% ethanol stored at room temperature. All buffers used with Ni2+-NTA resin contain high salt concentrations to reduce nonspecific electrostatic interactions between proteins and resin. Lower salt concentrations can be used but may lead to nonspecific binding of unwanted proteins to the resin.
MCAC-20, MCAC-40, MCAC-60, MCAC-80, MCAC-100, and MCAC-200 buffers: These buffers (containing different concentrations of imidazole) are made by mixing MCAC-0 buffer and MCAC-1000 buffer in the appropriate ratios: e.g., for MCAC60 buffer, 94:6 (v/v) MCAC-0/MCAC-1000. MCAC-EDTA buffer 20 mM Tris⋅Cl, pH 7.9 0.5 M NaCl 10% glycerol 0.1 M EDTA, pH 8.0 1 mM PMSF Store ≤6 months at 4°C Protease inhibitor cocktail, 150× Stock solutions: 2 mg/ml aprotinin in H2O 1 mg/ml leupeptin in H2O 1 mg/ml pepstatin A in methanol Cocktail: Combine 1.5 vol of each stock solution with 5.5 vol sterile water. Protease inhibitors can be obtained from Sigma or U.S. Biochemical. The stock solutions and cocktail are stable ≥6 months at −20°C.
COMMENTARY Background Information
Metal-Chelate Affinity Chromatography
Metal-chelate affinity chromatography (also called immobilized metal affinity chromatography or IMAC) for protein purification was first described in 1975 (Porath et al., 1975). It is based on the ability of certain amino acids acting as electron donors on the surface of proteins (histidine, tryptophan, tyrosine, or phenylalanine) to bind reversibly to transitionmetal ions that have been immobilized by a chelating group covalently bound to a solid
support. Of these amino acids, histidine is quantitatively the most important in mediating the binding of most proteins to immobilized metal ions. Histidine binds selectively to immobilized metal ions even in the presence of excess free metal ions in solution (Hutchens and Yip, 1990b). Copper and nickel ions have the greatest affinity for histidine (Yip et al., 1989). Proteins and peptides containing phosphoamino acids (phophoserine, phosphotyrosine,
9.4.12 Supplement 4
Current Protocols in Protein Science
or phosphothreonine) have also been purified by MCAC via selective interaction of the terminal phosphate group with Fe3+ (Muszynska et al., 1992). Phosphoproteins also bind to less common metals such as lutetium, scandium, and thorium (Andersson and Porath, 1986). The chelating group that has been used most extensively for MCAC is iminodiacetic acid (IDA). The tridentate IDA group binds to three sites within the coordination sphere of divalent metal ions such as copper, nickel, zinc, and cobalt. When copper ions (coordination number of 4) are bound to IDA, only one site remains available for interaction with proteins (Hochuli et al., 1987). For nickel ions (coordination number of 6) bound to IDA, three sites are available for binding to proteins. Thus Cu2+IDA complexes are stable on the column but have lower capacity for protein binding. Conversely, Ni2+-IDA complexes bind proteins more avidly, but the Ni2+-protein complexes are more likely to dissociate from the solid support. A pentadentate chelating group—N,N,N′tris(carboxymethyl)ethylenediamine (TED)— has also been used for MCAC. This chelator binds five coordination sites of the metal ions, providing highly stable metal ion–TED groups. However, for metal ions with a coordination number of 6, this leaves only one site available for protein interaction. The development of a new metal-chelating adsorbent, nitrilotriacetic acid (NTA), has provided a convenient and inexpensive tool for purification of proteins containing histidine residues (Hochuli et al., 1987). The quadridentate NTA moiety covalently coupled via a spacer arm to Sepharose CL-6B forms an extremely stable complex with metal ions possessing a coordination number of 6 (e.g., Ni2+). After binding to the NTA, the Ni2+ ion has two sites within its octahedral coordination sphere available for binding to electron donor groups (i.e., histidine) on the surface of proteins. Thus the advantage of NTA over IDA is that the Ni2+ ion is bound by four rather than three of its coordination sites. This minimizes leaching of the metal from the solid support and allows for more stringent purification conditions. The NTA also binds Cu2+ ions with high affinity, but this occupies all of the coordination sites, rendering the resulting complex ineffective for MCAC. The affinity of histidine residues for immobilized Ni2+ ions allows selective purification of proteins containing a high proportion of histidine residues on the surface. Immobilized copper or nickel ions bind native proteins with
a Kd of 1–17 × 10−5 M (Hutchens and Yip, 1990a). This affinity is enhanced significantly by designing proteins to contain short stretches (6 to 10 residues) of histidines in regions likely to be surface exposed (amino or carboxyl termini) using recombinant DNA technology. Addition of a histidine tail results in a protein that binds to the Ni2+-NTA complex with a Kd of 10−13 M at pH 8.0 even in the presence of detergent, ethanol, 2 M KCl (Hoffmann and Roeder, 1991), 6 M guanidine (Hochuli et al., 1988), or 8 M urea (Stüber et al., 1990). The histidine tail binds to the Ni2+-NTA resin via the imidazole side chains of the histidine residues. At pH ≥7.0, the imidazole side chain is deprotonated, with a net negative charge; at pH 5.97 (the pK of the imidazole side chain of histidine), 50% of the histidines are protonated; and at pH ≤4.5, almost all of the histidines are protonated and do not interact with Ni2+-NTA. Thus, there are two methods of dissociating histidine tails from the Ni2+-NTA resin. The first method, presented in this unit, uses increasing concentrations of imidazole at constant pH to displace the histidine tail from the Ni2+-NTA. This technique provides great versatility because it can be used for both native and denaturing MCAC and buffer preparation is simplified. The second method uses buffers of decreasing pH to elute the histidine tail and is quite efficient for producing pure protein (Hochuli et al., 1988). Disadvantages are that the pH must be maintained accurately at all temperatures and that some proteins may not be able to withstand the extreme pH change. Creation of a small histidine tail has advantages over other fusion protein systems. Addition of six histidines to the protein adds only 0.84 kDa to the mass of the protein, whereas other fusion protein systems utilize much larger affinity groups that often must be removed to allow normal protein function (e.g., glutathione-S-transferase, protein A, maltosebinding protein, and lacZ have masses of 26, 30, 40, and 116 kDa, respectively). The small histidine tail is not immunogenic and therefore need not be removed before the purified protein is injected into animals for antibody production. Histidine tail fusion proteins often retain their normal biologic functions—e.g., dihydrofolate reductase and adenylylcyclase retain their enzymatic activities (Hochuli, 1990; Taussig et al., 1993) and the TATA box binding protein retains its transcriptional regulatory function (Parvin et al., 1992). The aforementioned studies also demonstrate the technique’s suitability for purification of membrane, cyto-
Affinity Purification
9.4.13 Current Protocols in Protein Science
Supplement 4
plasmic, and nuclear proteins. However, although it often is not necessary to remove the histidine tail, it is possible to include this step and still generate sufficient quantities of protein pure enough for detailed X-ray crystallographic studies (Nikolov et al., 1992). The histidine tail can also be used to detect the recombinant protein. Ni2+-NTA covalently coupled to alkaline phosphate (available from Qiagen; see SUPPLIERS APPENDIX) can bind directly to histidine tailed proteins, and the bound alkaline phosphatase can be quantitated by standard techniques (UNIT 10.10). This allows for direct quantitation of histidine-tailed proteins blotted to membranes without the need for primary and secondary antibodies (Botting and Randall, 1995). The protocols presented in this unit utilize protein produced in bacteria; an example protocol for the purification of HIV-1 integrase expressed in E. coli is presented in UNIT 6.5. Histidine-tail fusion proteins have also been expressed and purified from HeLa cells using transient transfection techniques (Janknecht and Nordheim, 1992; Papavassiliou et al., 1992) or a vaccinia expression system (Janknecht et al., 1991) and from baculovirus-infected insect cells (Gearing et al., 1993; Taussig et al., 1993). Therefore, MCAC can be used with any protein expression system currently available. This technique can also be used to prepare immunoaffinity columns (UNIT 9.5). Histidine tails have been covalently linked to oligosaccharide moieties of monoclonal antibodies and then subsequently bound to Ni2+-NTA columns (Loetscher et al., 1992). The advantage of this method over more conventional techniques of antibody immobilization is that the immobilized antibody will retain a much greater ability to bind antigen and is recoverable from the column.
Critical Parameters
Metal-Chelate Affinity Chromatography
NTA can be synthesized according to the protocol provided by Hochuli (1990); more commonly, it is purchased from Qiagen, which provides detailed protocols for its use. Binding and elution of proteins from the Ni2+-NTA resin depend upon the histidine content of the protein. Each protein will have a slightly different histidine content and optimal elution profile on the Ni2+-NTA resin. Therefore, it is important to establish the buffer imidazole concentrations that will give the best purification of a particular protein. As outlined
in the Basic Protocol, gradient or stepwise elution of small-scale preparations should be done before large-scale preparations are attempted. It is also possible to perform batch purification by adding charged resin to a tube containing the extract, mixing for 1 hr, loading into a column, washing, and eluting.
Troubleshooting The most common problem is low yield of purified protein in the expected fraction. This can be caused by insufficient protein loaded on the column, protein not binding to the column, or protein not eluting from the column. Insufficient protein It is important to verify that the desired protein is adequately expressed in the chosen system. Check the crude extract for the presence of the expressed protein by SDS-PAGE or other methods before proceeding to the affinity purification steps. Protein in the crude lysate may undergo significant proteolytic degradation in some cell types. Always use PMSF, minimize freezing and thawing, maintain low temperatures during purification, and use additional protease inhibitors whenever practical. Lack of binding If the protein does not bind to the column, either the protein lacks a histidine tail, the column is inadequately charged with nickel, or the histidine tail is buried in the protein and is inaccessible to the resin. Before expression and purification are attempted, verify that the DNA sequence of the histidine tail and adjacent protein-coding sequence are in frame with each other and that initiation and termination codons are present. If the histidine tail is at the amino terminus, it may be helpful to delete the endogenous codon for initiator methionine; if this is present, translation may initiate downstream of it, producing a full-length protein without a tail. Placing the histidine tail at the carboxyl terminus ensures that only the full-length protein will bind to the Ni2+-NTA resin (if protease activity is controlled). However, a histidine tail at the carboxyl terminus is more likely to be inaccessible to the resin. Chelating agents (e.g., EDTA and EGTA) must be excluded from all solutions because they will strip the Ni2+ ions from the affinity column. Strong reducing agents such as dithiothreitol should also be avoided because they reduce the Ni2+ ions to metallic nickel,
9.4.14 Supplement 4
Current Protocols in Protein Science
forming a brown precipitate. Some proteins will require the presence of reducing agents to maintain structure. It is possible to use up to 10 mM 2-mercaptoethanol with the Ni2+-NTA resin, although the lowest possible concentration should be used. The Ni2+-NTA should retain a distinct bluegreen color throughout the purification process until EDTA is applied. If the resin is white or if the blue-green color is faint after charging the column with Ni2+, check all buffers and make sure no chelating and reducing agents are present. Recharge the column with Ni2+ and if the color is still faint or absent, regenerate the NTA resin (see Support Protocol 2) and repeat Ni2+ charging (see Basic Protocol, steps 6 to 8). If the resin is still white, discard it and start over with a new batch. If the protein has or should have a histidine tail but is not binding to the resin, perform the purification under denaturing conditions (see Alternate Protocol 1 or 2) to unmask the inaccessible histidine tail. Placing the histidine tail at the opposite terminus of the protein may also be helpful. Failure to elute If the protein appears to be binding to the column but not eluting, make a final elution with EDTA-containing buffer. Any protein bound to the Ni2+-NTA will be removed with the EDTA. Occasionally the protein bound to the resin will precipitate and not elute; if this problem is suspected, purify the protein under denaturing conditions and analyze all eluted fractions by SDS-PAGE for the presence of the expressed protein.
Anticipated Results The maximum binding capacity of Ni2+NTA resin is 5 to 10 mg of protein per milliliter of packed resin. Under ideal conditions, as much as 80% of the bound protein can be recovered. The limiting factor in most instances will probably be the amount of protein loaded on the resin, which depends on the protein expression system.
Time Considerations Expression of protein and preparation of crude extract require 1 day for the system described here. Column preparation requires 30 to 60 min. Loading, washing, and eluting the column require 4 to 6 hr.
Literature Cited Andersson, L. and Porath, J. 1986. Isolation of phosphoproteins by immobilized metal (Fe3+) affinity chromatography. Anal. Biochem. 154:250-254. Botting, C.H. and Randall, R.E. 1995. Reporter enzyme–nitrilotriacetic acid–nickel conjugates: Reagents for detecting histidine-tagged proteins. Biotechniques19:362-363. Gearing, K., Göttlicher, M., Teboul, M., Widmark, E., and Gustafsson, J. 1993. Interaction of the peroxisome-proliferator-activated receptor and retinoid X receptor. Proc. Natl. Acad. Sci. U.S.A. 90:1440-1444. Hochuli, E., Bannwarth, W., Dobeli, H., Gentz, R., and Stüber, D. 1988. Genetic approach to facilitate purification of recombinant proteins with a novel metal chelate adsorbent. Bio/Technology 6:1321-1325. Hochuli, E., Dobeli, H., and Schacher, A. 1987. New metal chelate adsorbent selective for proteins and peptides containing neighbouring histidine residues. J. Chromatogr. 411:177-184. Hoffmann, A. and Roeder, R. 1991. Purification of His-tagged proteins in nondenaturing conditions suggests a convenient method for protein interaction studies. Nucl. Acids Res. 19:6337-6338. Hutchens, T.W. and Yip, T.-T. 1990a. Protein interactions with immobilized transition metal ions: Quantitative evaluations of variations in affinity and binding capacity. Anal. Biochem. 191:160168. Hutchens, T.W. and Yip, T.-T. 1990b. Differential interaction of peptides and protein surface structures with free metal ions and surface-immobilized metal ions. J. Chromatogr. 500:531-542. Janknecht, R., De Martynoff, G., Lou, J., Hipskind, R., Nordheim, A., and Stunnenberg, H. 1991. Rapid and efficient purification of native histidine-tagged protein expressed by recombinant vaccinia virus. Proc. Natl. Acad. Sci. U.S.A. 88:8972-8976. Janknecht, R. and Nordheim, A. 1992. Affinity purification of histidine-tagged proteins transiently produced in HeLa cells. Gene 121:321-324. Loetscher, P., Mottlau, L., and Hochuli, E. 1992. Immobilization of monoclonal antibodies for affinity chromatography using a chelating peptide. J. Chromatogr. 595:113-119. Muszynska, G., Dobrowolska, G., Medin, A., Ekman, P., and Porath, J.O. 1992. Model studies on iron(III) affinity chromatography. II. Interaction of immobilized iron(III) ions with phosphorylated amino acids, peptides and proteins. J. Chromatogr. 604:19-28. Nikolov, D., Hu, S., Lin, J., Gasch, A., Hoffman, A., Horikoshi, M., Chua, N., Roeder, R., and Burley, S. 1992. Crystal structure of TFIID TATA-box binding protein. Nature 360:40-46.
Affinity Purification
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Papavassiliou, A., Bohmann, K., and Bohmann, D. 1992. Determining the effect of inducible protein phosphorylation on the DNA-binding activity of transcription factors. Anal. Biochem. 203:302309. Parvin, J., Timmers, H., and Sharp, P. 1992. Promoter specificity of basal transcription factors. Cell 68:1135-1144. Porath, J., Carlsson, J., Olsson, I., and Belfrage, G. 1975. Metal chelate affinity chromatography, a new approach to protein fractionation. Nature 258:598-599. Struhl, K. 1987. Subcloning of DNA fragments. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 3.16.1-3.16.11. John Wiley & Sons, New York. Stüber, D., Matile, H., and Garotta, G. 1990. System for high-level production in Escherichia coli and rapid purification of recombinant proteins: Application to epitope mapping, preparation of antibodies, and structure-function analysis. Immunol. Methods 4:121-152.
Taussig, R., Quarmby, L., and Gilman, A. 1993. Regulation of purified type I and type II adenylylcyclases by G protein β-γ subunits. J. Biol. Chem. 268:9-12. Yip, T.-T., Nakagawa, Y., and Porath, J. 1989. Evaluation of the interaction of peptides with Cu(II), Ni(II), and Zn(II) by high-performance immobilized metal ion affinity chromatography. Anal. Biochem. 183:159-171.
Key Reference Hochuli, E. 1990. Purification of recombinant proteins with metal chelate adsorbent. In Genetic Engineering, Principles and Practice, Vol. 12 (J. Setlow, ed.) pp. 87-98. Plenum, New York. Describes basic principles of MCAC with detailed protocols.
Contributed by Kevin J. Petty University of Texas Southwestern Medical Center Dallas, Texas
Studier, F.W., Rosenberg, A.H., Dunn, J.J., and Dubendorff, J.W. 1990. Use of T7 polymerase to direct expression of cloned genes. Methods Enzymol. 185:60-89.
Metal-Chelate Affinity Chromatography
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Current Protocols in Protein Science
Immunoaffinity Chromatography
UNIT 9.5
This unit describes the isolation of soluble or membrane-bound protein antigens from cells or homogenized tissue by immunoaffinity chromatography. This technique involves the elution of a single protein from an immunoaffinity column after prior elution of nonspecifically adsorbed proteins. Specifically, antibodies are coupled to Sepharose (an insoluble, large-pore-size chromatographic matrix). High-molecular-weight antigens pass freely into and out of the pores and bind to antibodies covalently bound to the matrix. To elute the bound antigen from the immunoaffinity matrix, the antibody-antigen interaction is destabilized by brief exposure to high-pH (Basic Protocol) or low-pH (Alternate Protocol 1) buffer. The use of batch purification of antigens shortens the column loading time (Alternate Protocol 2). The detergent octyl β-D-glucoside can be used instead of Triton X-100 for elution. Because octyl β-D-glucoside has a high critical micelle concentration (CMC) it can be readily removed by dialysis (Alternate Protocol 3). The procedure for covalently linking an antibody to Sepharose using the cyanogen bromide activation method is given in the Support Protocol. ISOLATION OF SOLUBLE OR MEMBRANE-BOUND ANTIGENS Two different Sepharose columns in series—a precolumn to remove nonspecifically binding material and a specific column—are used to isolate antigens from a cell or tissue lysate. Column fractions are analyzed by SDS-PAGE and silver staining to detect the antigens.
BASIC PROTOCOL
Materials Antibody (Ab)-Sepharose (see Support Protocol) Activated, quenched (control) Sepharose, prepared as for Ab-Sepharose (see Support Protocol) but eliminating Ab or substituting irrelevant Ab during coupling Cells or homogenized tissue Tris/saline/azide (TSA) solution (see recipe), ice cold Lysis buffer (see recipe), ice cold 5% (w/v) sodium deoxycholate (Na-DOC; filter sterilize and store at room temperature) Wash buffer (see recipe) Tris/Triton/NaCl buffers, pH 8.0 and 9.0 (see recipe), ice-cold Triethanolamine solution (see recipe), ice cold 1 M Tris⋅Cl, pH 6.7 (APPENDIX 2E), ice cold Column storage solution (see recipe), ice cold Chromatography columns Ultracentrifuge Quick-seal centrifuge tubes (Beckman) Additional reagents and equipment for column chromatography (UNITS 8.1), SDS-PAGE (UNIT 10.1), and silver staining (UNIT 10.5) NOTE: Carry out all procedures involving antigen in a 4°C cold room or on ice. Prepare the columns 1. Prepare an Ab-Sepharose immunoaffinity column (5 ml; 5 mg/ml antibody per milliliter packed Sepharose) and an activated, quenched (control) Sepharose precolumn (5 ml packed bed volume) linked in series (Fig. 9.5.1). Irrelevant antibody can be coupled to the Sepharose in the precolumn. Column size can vary; adjust amounts of Sepharose and cells proportionally. Contributed by Timothy A. Springer Current Protocols in Protein Science (1996) 9.5.1-9.5.11 Copyright © 2000 by John Wiley & Sons, Inc.
Affinity Purification
9.5.1 Supplement 3
buffer reservoir
b
hydrostatic pressure head
a b pre column
c
immunoaffinity column
e d
b
f
safety loop
c b
Figure 9.5.1 Immunoaffinity chromatography. During the application of the sample, two Sepharose columns, a Sepharose precolumn (without covalently bound specific antibody or with a covalently bound irrelevant antibody) and an immunoaffinity column (with covalently bound antibody), are attached in series to a buffer reservoir containing the sample. After the sample has been washed through, the precolumn is removed and the tubing of the safety loop is connected to the immunoaffinity column. The hydrostatic pressure head is the distance between the top of the solution in the buffer reservoir and the tip of the tubing at the bottom of the immunoaffinity column. When the elution reservoir is emptied, the hydrostatic head becomes zero when the fluid level reaches the safety loop, preventing columns from running dry. Fluid remaining above the column beds can be removed by raising the safety loop. After rinsing the tubing, the next elution is begun by placing the end of the safety loop in another reservoir containing the next elution buffer.
Immunoaffinity Chromatography
Inset: Schematic diagram of an immunoaffinity column. (a) 50-µl disposable capillary micropipet. (b) Tubing: Tygon S-54-HL Microbore, 0.05-in. i.d., or Tygon R-3603, 1⁄16-in. i.d. (softer tubing). (c) Female Luer fitting, white nylon (Value Plastics), 1⁄16 in. (d) Kontes Flex-column (Kontes Glass). (e) Barbed nipple connector, polypropylene, 3⁄32-in. top, 1⁄16-in. bottom (Value Plastics, Series AD). (f) Luer-Lok two-way stopcock (Kontes Glass).
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Current Protocols in Protein Science
Prepare the lysate 2. Suspend 50 g of cells at 1–5 × 108 cells/ml in ice-cold TSA solution, or add 1 to 5 vol ice-cold TSA per vol packed cells or homogenized tissue. Add an equal volume of ice-cold lysis buffer and stir 1 hr at 4°C. For glycosylphosphatidylinositol (GPI)-anchored proteins (UNIT 12.5), incubate 10 min at 20°C for efficient solubilization.
3. Centrifuge 10 min at 4000 × g, 4°C, to remove nuclei. Decant supernatant and save. For purification of cytoplasmic (soluble) antigens, it is not necessary to add detergents to the solutions and buffer used in subsequent steps. Detergent is needed only for cell lysis and solubilization of integral membrane proteins.
4. For purification of membrane antigens, add 0.2 vol of 5% Na-DOC to the postnuclear supernatant, and leave 10 min at 4°C or on ice. Transfer to quick-seal centrifuge tubes and centrifuge 1 hr at 100,000 × g, 4°C. Carefully remove supernatant and save. Set up and wash the columns 5. Attach Sepharose precolumn to immunoaffinity column (Fig. 9.5.1). 6. Wash both columns using the following regimen: 10 column volumes of wash buffer 5 column volumes of Tris/Triton/NaCl buffer, pH 8.0 5 column volumes of Tris/Triton/NaCl buffer, pH 9.0 5 column volumes of triethanolamine solution 5 column volumes of wash buffer. Isolate the antigen 7. Apply the supernatant from steps 3 or 4 (reserving some for analysis as described in step 15 below) to the precolumn and allow it to flow through the precolumn and specific column linked in series at a flow rate of 5 column volumes/hr. Collect the flow-through fractions, each 1⁄10 to 1⁄100 the volume of the applied supernatant. “Fat” chromatography columns, filled with Sepharose to a height of ∼2× column diameter, are used to maximize flow rates. A 10- to 20-ml syringe is used for 5 ml of Sepharose. The flow rate is adjusted with a hydrostatic head of up to 250 cm (Fig. 9.5.1). Sample loading can routinely take up to 2 days with no deleterious effect, but longer periods would suggest the column is clogged or the lysate is too viscous. The latter is usually due to the presence of DNA.
8. Wash with 5 column volumes of wash buffer, then close the stopcocks on both columns and disconnect the precolumn from the immunoaffinity column. Open the stopcock of the immunoaffinity column and allow fluid above the top of the column to drain out to bed level. The Sepharose has some elasticity and draining can continue until there is no buffer above the Sepharose bed. Draining until cracks appear in the Sepharose should be avoided. Fractions of this wash and washes obtained below should be saved.
9. Wash the immunoaffinity column between each change of buffers (steps 10 to 14) as follows. Close the stopcock and remove the end cap of the column. With a syringe connected to the outlet of the tubing from the buffer reservoir, aspirate all buffer from the tubing. Place tubing into the next buffer contained in another reservoir. Aspirating with a syringe, fill the tubing from the reservoir and remove the syringe. Crimp the tubing to regulate flow and rinse the inside wall of the column with the buffer. Open the column stopcock and drain the buffer to bed level. Put end cap loosely on the
Affinity Purification
9.5.3 Current Protocols in Protein Science
Supplement 3
column and allow buffer to drain into the column to a level several centimeters above the bed. Secure end cap and commence washes or elution. 10. Wash with 5 column volumes of wash buffer. 11. Wash with 5 column volumes of Tris/Triton/NaCl buffer, pH 8.0. 12. Wash with 5 column volumes of Tris/Triton/NaCl buffer, pH 9.0. Some nonspecifically bound proteins may be eluted at this step. In addition, some monoclonal antibodies may release some ligand at this pH. This should be checked.
13. Elute the antigen with 5 column volumes of triethanolamine solution. Collect fractions of 1 column volume into tubes containing 0.2 vol of 1 M Tris⋅Cl, pH 6.7, to neutralize the fractions collected. In some cases it may be desirable to lower the pH of the triethanolamine solution to preserve the functional activity of the ligand. The ideal pH gives complete release of the ligand, as verified by SDS-PAGE evaluation of a sample (∼20 ìl) of the eluted column bed (AbSepharose) and eluate (50 ìl).
14. Wash the column with 5 column volumes of TSA solution. A column may be reused many times and remain active for several years after storage at 4°C in TSA solution. It is important to prevent drying out of a column during storage. The use of column storage solutions inhibits the growth of microorganisms.
15. Analyze fractions for the presence of antigen—50-µl aliquots of each eluate fraction should be analyzed by SDS-PAGE and silver staining. Analyze 0.5- to 1-ml aliquots of the sample applied to the column and representative flowthrough and wash fractions by immunoprecipitation with Ab-Sepharose, and detect by silver staining to determine whether the column was saturated. If antibody leaches off the column during elution, it may be removed from the eluate by passage through protein A–Sepharose (Ey et al., 1978). Even the weakly binding mouse IgG1 subclass can be quantitatively removed at pH 8 (M. Dustin, pers. comm.). Preliminary analysis of fractions can be done using A280 readings when octyl β-D-glucoside or sodium deoxycholate are used as detergents. ALTERNATE PROTOCOL 1
LOW-pH ELUTION OF ANTIGENS Some protein antigens may be eluted more completely with greater retention of native conformation and with fewer contaminants when low-pH buffers are employed. Additional Materials (also see Basic Protocol) Sodium phosphate buffer, pH 6.3 (see recipe) Glycine buffer (see recipe) 1 M Tris⋅Cl, pH 9.0 (APPENDIX 2E) 1. Prepare the columns and lysate, wash the columns, and isolate the antigen (see Basic Protocol, steps 1 to 11). It is essential to remove sodium deoxycholate from the column before acid elution, because it precipitates or forms a gel at acid to neutral pH. TSA and triethanolamine solutions used here may also be modified by substituting 1% octyl β-D-glucoside for 0.1% Triton X-100 (see Alternate Protocol 3).
2. Wash with 5 column volumes of sodium phosphate buffer, pH 6.3. Immunoaffinity Chromatography
9.5.4 Supplement 3
Current Protocols in Protein Science
3. Elute with 5 column volumes of glycine buffer. Collect fractions into tubes containing 0.2 vol of 1 M Tris⋅Cl, pH 9.0. Mix each fraction immediately after collection.
4. Analyze fractions for antigen (see Basic Protocol, step 15). BATCH PURIFICATION OF ANTIGENS Batch purification of antigens shortens the column loading time. This technique is valuable for viscous lysates that take too long to load on a column and for antigens especially susceptible to proteolysis, because less time is required to complete the steps. A precolumn is not utilized because the supernatant is mixed with Ab-Sepharose and poured into a column. The antigen is then eluted (see Basic Protocol). The drawbacks of this protocol are that more “hands-on” time is required by the investigator and that nonspecifically binding material is not removed by a precolumn.
ALTERNATE PROTOCOL 2
1. Suspend and centrifuge the cells and purify the membrane antigens (see Basic Protocol, steps 2 to 4) to obtain the postnuclear supernatant. 2. Suspend Ab-Sepharose in the supernatant in a flask. Shake gently on a rotary shaker for 3 hr. Stop shaking and allow the Sepharose to settle. 3. Decant most of the supernatant. Pour the Ab-Sepharose and the remainder of the supernatant into a column and open the stopcock. Continue draining the column until all the Sepharose has been added. Allow the fluid to drain to bed level and close the stopcock. 4. Wash the immunoaffinity column, elute the antigen, and analyze the fractions (see Basic Protocol, steps 9 to 15). ELUTION IN OCTYL β-D-GLUCOSIDE The detergent octyl β-D-glucoside has a high critical micelle concentration (CMC) of 0.73% and can therefore be readily removed by dialysis. Also, adsorption of membrane proteins to surfaces for ELISA or adhesion assays is much more efficient when solutions containing membrane proteins are diluted so that the concentration of octyl β-D-glucoside is below the CMC. Because octyl β-D-glucoside is expensive, initial steps requiring large volumes may be done as described in the Basic Protocol; the initial detergent is then replaced by octyl β-D-glucoside in the wash step prior to elution.
ALTERNATE PROTOCOL 3
Additional Materials (also see Basic Protocol) TSA solution (see recipe) containing 1% octyl β-D-glucoside 1. Prepare the columns and lysate, wash the columns, and isolate the antigen (see Basic Protocol, steps 1 to 11). 2. Wash with 5 column volumes of TSA solution containing 1% octyl β-D-glucoside. 3. Elute with 5 column volumes of triethanolamine solution with 1% octyl β-D-glucoside substituted for 0.1% Triton X-100. 4. Collect fractions of 1 column volume into tubes containing 0.2 vol of 1 M Tris⋅Cl, pH 6.7, to neutralize the fractions collected. 5. Wash the column and analyze the fractions (see Basic Protocol, steps 14 and 15). Affinity Purification
9.5.5 Current Protocols in Protein Science
Supplement 3
SUPPORT PROTOCOL
PREPARATION OF ANTIBODY-SEPHAROSE This protocol details the procedure for covalently linking an antibody to Sepharose (an insoluble, large-pore-size chromatographic matrix) using the cyanogen bromide activation method (also see UNIT 9.3). First, the antibody and Sepharose are prepared separately. The Sepharose is then activated with cyanogen bromide (alternatively, CNBr-activated Sepharose can be purchased from Pharmacia Biotech and used according to the manufacturer’s instructions). Finally, the CNBr-activated Sepharose is coupled to the antibody. CAUTION: CNBr is very toxic, releases toxic vapor, and is reported to create explosive side products upon storage; therefore, it should always be used in a hood and handled with appropriate CNBr- and solvent-resistant gloves. Only a fresh bottle of CNBr should be employed and it should be opened in the hood. Because of the problems involved in handling CNBr, it is best to dissolve a complete bottle in a defined amount of anhydrous acetonitrile. Short-term storage of this solution (several weeks) at 0°C is possible, but prolonged storage is not recommended. Materials 1 to 30 mg/ml antigen-specific monoclonal or polyclonal antibody 0.1 M NaHCO3/0.5 M NaCl Sepharose CL-4B (or Sepharose CL-2B for high-molecular-weight antigens; Pharmacia Biotech) 2 M Na2CO3 Cyanogen bromide (CNBr)/acetonitrile (see recipe) 1 mM and 0.1 mM HCl, ice-cold 0.05 M glycine (or ethanolamine), pH 8.0 Tris/saline/azide (TSA) solution (see recipe) Dialysis tubing (MWCO >10,000) Ultracentrifuge Whatman no. 1 filter paper Buchner funnel Erlenmeyer filtration flask Water aspirator Prepare the antibody 1. Dialyze 1 to 30 mg/ml antibody against 0.1 M NaHCO3/0.5 M NaCl at 4°C with three buffer changes during 24 hr. Use a volume of dialysis solution that is 500 times the volume of antibody solution. Dialysis is performed to remove all small molecules containing free amino or sulfhydryl groups.
2. Centrifuge 1 hr at 100,000 × g, 4°C, to remove aggregates. Save the supernatant. Removal of aggregates is important. Because only some of the antibody molecules in an aggregate will directly couple to the Sepharose, the noncoupled antibody molecules may leach out during elution.
3. Measure the A280 of an aliquot of the solution and determine the concentration of the antibody (mg/ml IgG = A280/1.44). Dilute with 0.1 M NaHCO3/0.5 M NaCl to 5 mg/ml (or to the same concentration as desired for Ab-Sepharose) and keep at 4°C. Measure the A280 of this solution for comparison of absorbance in step 10.
Immunoaffinity Chromatography
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Current Protocols in Protein Science
Prepare the Sepharose 4. Allow the Sepharose slurry to settle in a beaker or the container in which it is sold; decant and discard the supernatant. Weigh out the desired quantity of Sepharose (assume density = 1.0). 5. Set up a filter apparatus using Whatman no. 1 filter paper in a Buchner funnel and an Erlenmeyer filtration flask attached to a water aspirator. Wash the Sepharose on the filter apparatus with 10 vol water. Sintered-glass funnels are traditionally recommended but rapidly become clogged unless coarse-porosity funnels are used.
Activate Sepharose with CNBr 6. Transfer Sepharose to 50-ml beaker and add an equal volume of ice-cold 2 M Na2CO3. 7. Activate Sepharose in an ice bath using 3.2 ml CNBr/acetonitrile per 100 ml Sepharose. Add CNBr/acetonitrile dropwise with a Pasteur pipet over 1 min, while slowly stirring the slurry with a magnetic stir bar. Continue stirring slowly for 5 min. Excessive and vigorous stirring may fracture the Sepharose beads; this may cause flow problems during column chromatography. The protocol uses 2 g CNBr/100 ml Sepharose. Two to four grams of CNBr/100 ml Sepharose can be used to couple 1 to 20 mg of antibody/ml Sepharose. CAUTION: Activation should be carried out in a fume hood.
8. Rapidly filter the CNBr-activated Sepharose as in step 5. Aspirate to semidryness (i.e., until the Sepharose cake cracks and loses its sheen). Wash with 10 vol ice-cold 1 mM HCl, then with 2 vol ice-cold 0.1 mM HCl. Hydrate with enough ice-cold 0.1 mM HCl so that the cake regains its sheen, but there is no excess liquid above the cake. Washing is most efficient if the wash solution is added evenly over the surface of the cake at about the same rate as the solution is removed by filtration. CNBr-activated Sepharose is very unstable at the alkaline pH necessary for activation; it is much more stable in dilute HCl. CNBr-activated Sepharose can be purchased from Pharmacia Biotech, but the coupling capacity will be lower. Destroy the contents of the filter flask, which contains excess CNBr, by slowly adding a 5-fold molar excess (relative to the total initial amount of CNBr used) of 1 M NaOH. Keep the reaction mixture cold by adding pieces of ice. Perform this decomposition in the hood, and keep the mixture there until all CNBr is destroyed.
Couple antibody to CNBr-activated Sepharose 9. Immediately transfer a weighed amount of Sepharose (assume density = 1.0) to a beaker. Add an equal volume of a solution of antibody dissolved in 0.1 M NaHCO3/0.5 M NaCl (from step 3). Stir gently with a magnetic stir bar or rotate end-over-end 2 hr at room temperature or overnight at 4°C. 10. Add 0.05 M glycine (or ethanolamine), pH 8.0, to saturate the remaining reactive groups on the Sepharose and allow the slurry to settle. Remove an aliquot of the supernatant and centrifuge to remove any residual Sepharose. Measure the A280 and compare to the A280 of the antibody solution from step 3 to determine the percentage coupling. 11. Store the Ab-Sepharose in TSA solution. Affinity Purification
9.5.7 Current Protocols in Protein Science
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
CNBr/acetonitrile To 25 g of cyanogen bromide in the original bottle (CNBr should be white, not yellow, crystals) add 50 ml acetonitrile to make a 62.5% (w/v) solution. Store up to several weeks at −20°C in a desiccator over silica. Allow to warm before opening. CAUTION: CNBr is a highly toxic lachrymator; handle in a fume hood.
Column storage solution Prepare in TSA solution (see recipe) either 1 mM EDTA/20 µg/ml gentamicin or 0.01% thimerosal (Aldrich). Detergent stock solutions Prepare 10% Triton X-100 or 5% sodium deoxycholate in water. Sterilize either solution by Millipore filtration. Both solutions remain stable for 5 years at room temperature; Triton X-100 solution should be stored in the dark to prevent photooxidation. NP-40 can be used in place of Triton X-100.
Glycine buffer 50 mM glycine⋅HCl, pH 2.5 0.1% Triton X-100 (see recipe for detergent stock solutions) 0.15 M NaCl Lysis buffer TSA solution (see recipe) containing: 2% Triton X-100 (see recipe for detergent stock solutions) 5 mM iodoacetamide Aprotinin (0.2 trypsin inhibitor U/ml) 1 mM phenylmethylsulfonyl fluoride (add fresh from 100 mM stock solution prepared in absolute ethanol) IMPORTANT NOTE: Iodoacetamide is a protease inhibitor and prevents oxidation of free cysteines to disulfide-bonded cysteines. It should be omitted for enzymes that require cysteines for activity. NP-40 can be used in place of Triton X-100.
Sodium phosphate buffer, pH 6.3 50 mM sodium phosphate, pH 6.3 0.1% Triton X-100 (see recipe for detergent stock solutions) 0.5 M NaCl Triethanolamine solution 50 mM triethanolamine, pH ∼11.5 0.1% Triton X-100 (see recipe for detergent stock solutions) 0.15 M NaCl Other organic solutions, such as diethylamine, may be used in place of triethanolamine. The pH of this solution should be determined for each antibody as elution conditions may vary (see Critical Parameters).
Immunoaffinity Chromatography
9.5.8 Supplement 3
Current Protocols in Protein Science
Tris/saline/azide (TSA) solution 0.01 M Tris⋅Cl, pH 8.0 (at 4°C) 0.14 M NaCl 0.025% NaN3 CAUTION: Sodium azide (NaN3) is poisonous; wear gloves and handle cautiously.
Tris/Triton/NaCl buffer, pH 8.0 and 9.0 50 mM Tris⋅Cl, pH 8.0 or pH 9.0 0.1% Triton X-100 (see recipe for detergent stock solutions) 0.5 M NaCl Wash buffer 0.01 M Tris⋅Cl, pH 8.0 (at 4°C) 0.14 M NaCl 0.025% NaN3 0.5% Triton X-100 (see recipe for detergent stock solutions) 0.5% sodium deoxycholate (see recipe for detergent stock solutions) CAUTION: Sodium azide (NaN3) is poisonous; wear gloves and handle cautiously.
COMMENTARY Background Information The review of affinity chromatography by Wilchek et al. (1984) discusses available methods for activation of solid supports, coupling of ligands, adsorption of proteins, and elution of protein from affinity columns. Table III of that review lists numerous examples of proteins that have been purified by immunoaffinity chromatography and the elution conditions for each purification. Traditionally, purification of membrane proteins started with a membrane purification step, and in some cases this is still desirable. However, it is difficult to achieve more than a 5-fold purification of plasma membranes and yields are usually only 10% to 40%. Omission of membrane purification in this protocol (Williams and Barclay, 1986; Johnson et al., 1985) results in increased yield and decreased experiment time. Purification to homogeneity or near-homogeneity can usually be achieved for protein antigens present in ≥10,000 molecules per eukaryotic cell. This protocol can be used for both membrane and intracellular antigens. However, for soluble antigens, immunoaffinity chromatography is completed without detergent.
Critical Parameters Binding capacities of Ab-Sepharose columns (coupled at 5 mg monoclonal antibody/ml Sepharose) have been found to be 2% to 20% of the theoretical binding capacity. The lowest and highest binding capacity values were found for antigens of 150,000 and 18,000
Mr, respectively, suggesting that increased antigen size may constrain access to antibody in the pores of the affinity matrix. Because of its larger pore size, Sepharose CL-4B (a 4% crosslinked agarose) is far preferable to Sepharose 6B (a 6% agarose), which is the usual commercially available preactivated agarose. The cross-links in the CL series yield mechanically more robust beads and do not appreciably decrease activation with cyanogen bromide (CNBr). The Support Protocol for coupling protein antigens to CNBr-activated Sepharose is a modification of the methods of Cuatrecasas (1970) and March et al. (1974). As originally described, the washing was done at alkaline pH. Because activated Sepharose is very unstable at this pH, it was originally recommended that washing, adding the protein ligand, and mixing be done in <90 sec (Cuatrecasas, 1970). However, with the use of an acid wash buffer (Gelb, 1973) as described in the Support Protocol, activated groups remain stable for ≥30 min. Wilchek et al. (1984) described an alternative activation procedure with CNBr triethanolamine in acetone/water that is highly efficient and minimizes side-reaction derivatization of the Sepharose. The 0.1 M NaHCO3 buffer in which antibody is dissolved provides an optimal pH of ∼8.4 after mixing with activated Sepharose. The small amount of CNBr recommended is sufficient to obtain a coupling yield of 80% to 95%. Higher amounts of CNBr may result in multipoint attachment of IgG molecules to the matrix, thereby inactivating antigen
Affinity Purification
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Supplement 3
Immunoaffinity Chromatography
binding sites and reducing accessibility to antigen. Coupling efficiencies on the order of 99% are usually associated with lower antigen binding capacity and are to be avoided; coupling efficiencies of 80% to 90% are ideal. Smaller amounts of CNBr per milliliter of Sepharose also result in lower derivatization of the Sepharose with charged groups. These charged groups can cause nonspecific binding. The amount of CNBr used in commercially available activated Sepharose is not specified, but is probably high. Many investigators purchase CNBr-activated Sepharose; others activate their own. Commercially activated Sepharose is stabilized by drying. Not all Sepharose can withstand drying; e.g., Sepharose 2B (2% agarose beads) is less robust than Sepharose 4B (4% agarose) and is not available in a preactivated form. Activating your own Sepharose allows more control over coupling efficiency; enables use of CL (cross-linked) Sepharose, which is more robust and has faster flow rates; enables use of Sepharose CL-2B; and is more cost effective. A binding capacity of 40% was reported for coupling at 2 to 3 mg antibody/ml Sepharose. Successful purification has been achieved using monoclonal antibodies with affinity constants ranging from 2 × 107 to 4 × 108 M−1. The column should be saturated with antigen by allowing some of the antigen to flow through the column during loading. This will result in the highest antigen purity and a mass yield, and will diminish the relative level of antibody eluted along with the antigen when antibody is leaching off the column. Sodium deoxycholate is used in the solubilization protocol (Johnson et al., 1985) because it dissociates proteins from the membrane more effectively than Triton X-100. However, because sodium deoxycholate releases DNA from nuclei, it must be added to the lysate after the nuclei are removed. Sodium deoxycholate forms a mixed micelle with Triton X-100. Although sodium deoxycholate can be substituted for Triton X-100 during high-pH elution, it gels at low pH and in high salt. Both sodium deoxycholate and nonionic detergents may dissociate subunits of protein complexes, which interact within the membrane. Addition of phospholipid, low concentrations of Triton X-100, and mild detergents (e.g., digitonin, octylglucoside, and CHAPS) preserves membrane protein complexes (Helenius et al., 1979; Rivnay et al., 1982; Tsuchiya and Saito, 1984). Glycosylphosphatidylinositol (GPI)-anchored proteins (UNIT 12.5) are resistant to deter-
gent solubilization at 4°C (Schroeder et al., 1994). Preparation of the lysate (Basic Protocol, step 2) should be conducted for 10 min at 20°C for these antigens. Octyl β-D-glucoside is an expensive detergent that is readily removable by dialysis. Furthermore, adsorption of proteins to substrates is enhanced by dilution below a CMC of 0.73%. In Alternate Protocol 3, the initial detergent is replaced with octyl β-D-glucoside prior to elution of antigen from the affinity column. Protein antigens eluted by acid or base can frequently be renatured by neutralization. However, some protein antigens are irreversibly denatured. The structure of certain antigens is preserved after acid, but not base, elution (Plunkett and Springer, 1986), whereas the structure of other antigens is preserved after base, but not acid, elution (Johnson et al., 1985). Some antigens are eluted at low pH, but not at high pH; for others the reverse is true. For each antibody-antigen combination, the optimal pH for elution of specific antigens as well as contaminants must be empirically defined. Antibody binding capacity is usually retained after repeated exposure to low and high pH elution buffers (see Alternate Protocol 1 and see Basic Protocol, respectively). The use of chaotropic agents (e.g., potassium thiocyanate and urea) for elution is a seldom-employed alternative (Johnson et al., 1985).
Troubleshooting Immunoaffinity chromatography relies on the elution of a single protein from an immunoaffinity column after prior elution of all other nonspecifically adsorbed proteins. Thus, depending on the elution conditions used, the desired protein antigen may be contaminated with other proteins. It is very important to determine whether a contaminant is present if the protein is to be analyzed for amino acid composition or protein sequence. One-dimensional gel electrophoresis (UNIT 10.1) should be used to verify elution of contaminating proteins during washing of the immunoaffinity column, as well as the purity of the protein in the final eluate. If the protein is not pure, the wash steps must be optimized to ensure that other contaminating proteins are removed.
Anticipated Results Antigen yield is typically 40% to 70% of starting material (Kürzinger and Springer, 1982; Johnson et al., 1985) and purification factors of 1,000- to 10,000-fold may be achieved (Kürzinger and Springer, 1982; Williams and Bar-
9.5.10 Supplement 3
Current Protocols in Protein Science
clay, 1986; Plunkett and Springer, 1986). Further purification can usually be achieved by a second cycle of immunoaffinity chromatography. Monoclonal antibodies are most convenient, but affinity-purified polyclonal IgG can also be used.
Time Considerations
Plunkett, M.L. and Springer, T.A. 1986. Purification and characterization of the lymphocyte functionassociated-2 (LFA-2) molecule. J. Immunol. 136:4181-4187. Rivnay, B., Wank, S.A., Poy, G., and Metzger, H. 1982. Phospholipids stabilize the interaction between the alpha and beta subunits of the solubilized receptor for immunoglobulin E. Biochemistry 21:6922-6927.
Pouring the column takes a few minutes and lysate preparation requires ∼6 hr. Purification proceeds over 1 to 2 days depending on the flow rate of the immunoaffinity column. The majority of this time involves loading the sample on the column, which may be done from a reservoir and requires little hands-on time. The use of batch purification (Alternate Protocol 2) reduces the sample application time to 3 hr. Elution of an immunoaffinity column requires 5 to 6 hr.
Schroeder, R., London, E., and Brown, D. 1994. Interactions between saturated acyl chains confer detergent resistance on lipids and glycosylphosphatidylinositol (GPI)-anchored proteins: GPI-anchored proteins in liposomes and cells show similar behavior. Proc. Natl. Acad. Sci. U.S.A. 91:12130-12134.
Literature Cited
Williams, A.F. and Barclay, A.N. 1986. Glycoprotein antigens of the lymphocyte surface and their purification by antibody affinity chromatography. In Immunological Methods in Biomedical Sciences (D.M. Weir, L.A. Herzenberg, C.C. Blackwell, and L.A. Herzenberg, eds.) pp. 22.122.24. Blackwell Scientific, Oxford.
Cuatrecasas, P. 1970. Protein purification by affinity chromatography. J. Biol. Chem. 245:3059-3065. Ey, P.L., Prowse, S.J., and Jenkin, C.R. 1978. Isolation of pure IgG1, IgG2a and IgG2b immunoglobulins from mouse serum using protein A– Sepharose. Immunochemistry 15:429-436. Gelb, W.G. 1973. Affinity chromatography: For separation of biological materials. Am. Lab. 81:61-67. Helenius, A., McCaslin, D.R., Fries, E., and Tanford, C. 1979. Properties of detergents. Methods Enzymol. 56:734-749. Johnson, P., Williams, A.F., and Woollett, G.R. 1985. Purification of membrane glycoproteins with monoclonal antibody affinity columns. In Hybridoma Technology in the Biosciences and Medicine (T.A. Springer, ed.) pp. 163-175. Plenum, New York. Kürzinger, K. and Springer, T.A. 1982. Purification and structural characterization of LFA-1, a lymphocyte function-associated antigen, and Mac-1, a related macrophage differentiation antigen. J. Biol. Chem. 257:12412-12418. March, S.C., Parikh, I., and Cuatrecasas, P. 1974. A simplified method for cyanogen bromide activation of agarose for affinity chromatography. Anal. Biochem. 60:149-152.
Tsuchiya, T. and Saito, S. 1984. Use of n-octyl-β-Dthioglucoside, a new nonionic detergent, for solubilization and reconstitution of membrane proteins. J. Biochem. 96:1593-1597. Wilchek, M., Miron, T., and Kohn, J. 1984. Affinity chromatography. Methods Enzymol. 104:3-55.
Key References Harlow, E. and Lane, D. 1988. Antibodies: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Hjelmeland, J.M. and Chrambach, A. 1984. Solubilization of functional membrane proteins. Methods Enzymol. 104:305-318. Johnson et al., 1985. See above. Describes the critical parameters involved in immunoaffinity chromatography. Wilchek et al., 1984. See above. Describes the mechanism of activation of Sepharose by CNBr and alternative activation procedures, and lists numerous examples of proteins purified by affinity chromatography.
Contributed by Timothy A. Springer Center for Blood Research Harvard Medical School Boston, Massachusetts
Affinity Purification
9.5.11 Current Protocols in Protein Science
Supplement 3
Purification of Sequence-Specific DNA-Binding Proteins by Affinity Chromatography
UNIT 9.6
Many biological processes, such as recombination, replication, and transcription, involve the action of sequence-specific DNA-binding proteins. Analysis and purification of these proteins by conventional chromatographic methods is often difficult because DNA-binding proteins typically make up <0.01% of the total cellular protein. Various methods have been developed for purifying such proteins. Affinity chromatography is a very effective means of purifying a protein based on its sequence-specific DNA-binding properties and is relatively straightforward if proper care is taken. The affinity chromatography procedure described in this unit uses DNA containing specific recognition sites for the desired protein that has been covalently linked to a solid support. Researchers in the past have purified proteins based on the proteins’ ability to bind DNA, only to be disappointed to find that the DNA-binding activity of their highly purified products was not sequencespecific. This situation can be avoided by carefully performing all of the preliminary steps and experimental controls. Basic Protocol 1 describes preparation of a DNA affinity resin, including cyanogen bromide (CNBr) activation of the agarose support. The Alternate Protocol provides a method to couple DNA to commercially available CNBr-activated Sepharose. Support Protocol 1 describes how to purify crude synthetic oligonucleotides by gel electrophoresis prior to preparation of the affinity resin. Basic Protocol 2 outlines the affinity chromatography procedure. Support Protocol 2 describes determination of the appropriate type and quantity of nonspecific competitor DNA that should be used in the procedure and its preparation. Parameters essential to the success of an affinity chromatography experiment are discussed in detail in the Commentary. Figure 9.6.1 provides a summary diagram of the entire chromatography procedure. PREPARATION OF DNA AFFINITY RESIN Correct choice of oligonucleotide sequence (discussed in detail in the Commentary) and preparation of the affinity resin are probably the most important parts of the affinity chromatography procedure. Preparation of affinity resin can be broken down into four steps: (1) preparing oligonucleotides; (2) activating Sepharose; (3) coupling DNA to resin; and (4) blocking unreacted CNBr. The first step requires highly purified oligonucleotides that contain the recognition sequence for the desired protein. Once purified, the complementary oligonucleotides are annealed, phosphorylated with T4 polynucleotide kinase, and ligated into long, multimeric chains (averaging ≥10-mers) with T4 DNA ligase. Sepharose CL-2B is activated with CNBr, the ligated DNA added, and the coupling reaction carried out overnight. Because CNBr is very toxic, researchers may prefer to employ the Alternate Protocol, which uses commercially available CNBr-activated Sepharose and therefore avoids direct handling of CNBr. After the coupling reaction, the remaining reactive groups are blocked with ethanolamine. This protocol is designed to prepare 10 ml of affinity resin.
BASIC PROTOCOL 1
NOTE: Glass-distilled or other high-quality water should be used throughout these procedures.
Affinity Purification Contributed by Leslie A. Kerrigan and James T. Kadonaga Current Protocols in Protein Science (1998) 9.6.1-9.6.18 Copyright © 1998 by John Wiley & Sons, Inc.
9.6.1 Supplement 11
crude synthetic oligonucleotides
and
purified oligonucleotides
oligo A
oligo B
oligo A
oligo B
purify by acrylamide gel electrophoresis or HPLC
anneal
agarose resin
crude protein extract
phosphorylate
P treat with CNBr
P ligate
separate by conventional chromatography
multimerized oligonucleotides
CNBractivated resin
partially purified extract
couple to activated resin
add competitor DNA
affinity column starting material
block reactive groups
apply sample to affinity resin
increase salt concentration collect fractions assay for DNAbinding activity
resin
check purity with SDSPAGE
Figure 9.6.1 Purification of sequence-specific DNA-binding proteins with DNA affinity chromatography. Shown are the steps required to perform an affinity chromatography experiment using the methods described in this unit. Affinity Chromatography of DNA-Binding Proteins
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Materials 440 µg each of two synthetic oligonucleotides with desired binding site (see Support Protocol 1; or commercial HPLC-purified) TE buffer, pH 7.8 (APPENDIX 2E) 10× T4 polynucleotide kinase buffer (see recipe) 20 mM ATP (Na+ salt), pH 7.0 150 mCi/ml [γ-32P]ATP (6000 Ci/mmol) 10 U/µl T4 polynucleotide kinase (New England Biolabs) 10 M ammonium acetate (APPENDIX 2E) 25:24:1 (v/v/v) phenol/chloroform/isoamyl alcohol 24:1 (v/v) chloroform/isoamyl alcohol 3 M sodium acetate (APPENDIX 2E) 100% and 75% ethanol 10× linker/kinase buffer (see recipe) 6000 U/ml T4 DNA ligase (measured in Weiss units; New England Biolabs) Buffered phenol Isopropanol (2-propanol) Sepharose CL-2B (Pharmacia Biotech) Cyanogen bromide (CNBr; Aldrich) N,N-dimethylformamide 5 N NaOH (APPENDIX 2E) 10 mM and 1 M potassium phosphate buffer, pH 8.0 (APPENDIX 2E) 1 M ethanolamine hydrochloride, pH 8.0 (see recipe) NaOH, solid Glycine 1 M KCl (APPENDIX 2E) Column storage buffer (see recipe) 15-ml screw-cap polypropylene tubes Heating blocks or water baths, 15°C, 37°C, 65°C, and 88°C 60-ml coarse-sintered glass funnel Rotating wheel 5′-phosphorylate the oligonucleotides 1. In a 1.5-ml microcentrifuge tube, prepare a mixture containing 440 µg of each oligonucleotide in TE buffer in a total volume of 130 µl. Add 20 µl of 10× T4 polynucleotide kinase buffer. Incubate 2 min at 88°C, 10 min at 65°C, 10 min at 37°C, and 5 min at room temperature. A 1-ìmol synthesis of oligonucleotide should yield enough purified DNA for ∼20 ml of affinity resin.
2. Divide mixture in half in separate microcentrifuge tubes. To each 75-µl aliquot, add 15 µl of 20 mM ATP (pH 7.0), ∼5 µCi [γ-32P]ATP, and 10 µl of 10 U/µl T4 polynucleotide kinase (100 U total). Incubate 2 hr at 37°C. Dividing the reaction in half at this point facilitates the steps that follow. To add the labeled ATP, do not thaw, but simply touch (do not jab) the top of the frozen [γ-32P]ATP with a yellow pipet tip and transfer the resulting tiny amount to the reaction tube.
3. Inactivate the kinase by adding 50 µl of 10 M ammonium acetate and 100 µl water to each tube and heating 15 min at 65°C. Allow to cool to room temperature. Affinity Purification
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Purify phosphorylated oligonucleotides 4. Add 750 µl of 100% ethanol and mix by inversion. Microcentrifuge at high speed 15 min at room temperature to pellet DNA. Discard supernatant. 5. Resuspend each pellet in 225 µl TE buffer. 6. Add 250 µl of 25:24:1 phenol/chloroform/isoamyl alcohol to each tube. Vortex 1 min. Microcentrifuge 5 min at high speed to separate phases. Transfer the aqueous phase (upper layer) to a new tube. 7. Add 250 µl of 24:1 chloroform/isoamyl alcohol to aqueous phase. Vortex 1 min. Microcentrifuge 5 min at high speed to separate phases. Transfer the aqueous phase (upper layer) to a new tube. 8. Add 25 µl of 3 M sodium acetate to aqueous phase and mix by vortexing. Then add 750 µl of 100% ethanol and mix by inversion. Microcentrifuge 15 min at high speed to pellet DNA. Discard supernatant. 9. Wash pellet with 800 µl of 75% ethanol. Mix by vortexing. Microcentrifuge 5 min at high speed. Discard supernatant. 10. Dry pellet in vacuum evaporator (e.g., Speedvac). Ligate oligonucleotides 11. Add 65 µl water and 10 µl of 10× linker/kinase buffer to each pellet. Dissolve DNA by vortexing. Add 20 µl of 20 mM ATP (pH 7.0) and 5 µl of 6000 U/ml T4 DNA ligase (30 Weiss units). Incubate ≥2 hr at room temperature or overnight at 15°C. Depending upon the oligonucleotides used, the optimal temperature for ligation will vary from 4° to 30°C. Short oligonucleotides (≤15-mers) tend to ligate better at lower temperatures (4° to 15°C), whereas oligonucleotides that have a moderate degree of palindromic symmetry tend to self-anneal and therefore ligate better at higher temperatures (15° to 30°C).
12. Monitor the ligation reaction by agarose gel electrophoresis, using 0.5 µl of ligation reaction per gel lane and including lanes containing size markers. Visualize DNA by ethidium bromide staining and UV photography. 5′-phosphorylated oligonucleotides often do not ligate on the first attempt. If ligation has not occurred, extract the DNA once with 25:24:1 phenol/chloroform/isoamyl alcohol and once with 24:1 chloroform/isoamyl alcohol, then ethanol precipitate using sodium acetate as the precipitating salt. Dissolve the DNA in 225 ìl TE, add 25 ìl of 3 M sodium acetate, and reprecipitate with 750 ìl of 100% ethanol. Wash with 75% ethanol, dry in Speedvac, and repeat ligation. The average length of the ligated oligonucleotides should be ≥10-mers. Ligating the oligonucleotides increases the binding capacity of the affinity resin and helps avoid potential problems due to steric hindrance between the factor and the Sepharose support. However, oligonucleotide multimers <10-mers will probably also work.
Purify oligonucleotide multimers 13. Add 100 µl buffered phenol to the 100-µl ligation reactions. Vortex 1 min. Microcentrifuge 5 min at high speed. Transfer aqueous phase (upper layer) to a new tube.
Affinity Chromatography of DNA-Binding Proteins
14. Add 100 µl of 24:1 chloroform/isoamyl alcohol to aqueous phase. Vortex 1 min. Microcentrifuge 5 min at high speed. Transfer aqueous phase (upper layer) to a new tube. 15. Add 33 µl of 10 M ammonium acetate to the aqueous phase. Mix by vortexing.
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16. Add 133 µl isopropanol. Mix by inversion. Incubate 20 min at −20°C. Microcentrifuge 15 min at high speed to pellet DNA. Discard supernatant. The ammonium acetate/isopropanol precipitation removes residual ATP, which would otherwise interfere with coupling of the ligated DNA to the CNBr-activated Sepharose.
17. Add 225 µl TE buffer. Vortex to dissolve pellet. Add 25 µl of 3 M sodium acetate. Mix by vortexing. Add 750 µl of 100% ethanol. Mix by inversion. Microcentrifuge 15 min at high speed to pellet DNA. Discard supernatant. 18. Wash DNA twice with 75% ethanol. Dry pellet in vacuum evaporator. 19. Dissolve DNA in 50 µl water. Store at −20°C. Do not dissolve the DNA in TE buffer, as the Tris buffer in TE will interfere with the coupling reaction.
Prepare CNBr-activated Sepharose It is best to assemble all equipment and reagents required for the activation and coupling reactions before proceeding with the following steps. 20. Place 10 to 15 ml (settled bed volume) of Sepharose CL-2B in a 60-ml coarse-sintered glass funnel and wash extensively with 500 ml water. To wash, add water to resin in funnel, stir gently with a glass rod, and remove water by vacuum suction, making sure to release vacuum before the resin is suctioned into a dry cake. Repeat until all 500 ml water is used.
21. Transfer moist Sepharose resin to a 25-ml graduated cylinder, estimating 10 ml of resin. Add water to 20 ml final volume. Transfer the resulting slurry to a 150-ml glass beaker containing a magnetic stir bar. Place beaker in a water bath equilibrated to 15°C and set up over a magnetic stirrer in a fume hood. Turn on stirrer to slow medium speed. Keep the water bath at 15°C by periodically adding small chunks of ice as needed. Occasional variation of a degree or two is not critical.
22. In the fume hood, measure 1.1 g CNBr into a 25-ml Erlenmeyer flask, keeping the mouth of the flask covered with Parafilm or a ground glass stopper as much as possible (it is better to have slightly more than 1.1 g than slightly less). Add 2 ml N,N-dimethylformamide (the CNBr will dissolve instantly). Over the course of 1 min, add the resulting CNBr solution dropwise to the stirring Sepharose slurry. CAUTION: CNBr is highly toxic and volatile. Use only in a fume hood with extreme caution. Observe appropriate decontamination and disposal procedures.
23. Immediately add 5 N NaOH as follows: add 30 µl to the stirring mixture every 10 sec for 10 min until 1.8 ml NaOH has been added. It is convenient to measure 1.8 ml NaOH into a small tube before addition to the reaction to avoid having to change pipet tips during the 10-sec additions.
24. Immediately add 100 ml ice-cold water to the beaker and pour the mixture into a 60-ml coarse-sintered glass funnel. IMPORTANT NOTE: At this point, it is very important to avoid suction-filtering the resin into a dry cake. If the resin is accidentally dried into a cake, do not use; instead, repeat steps 20 to 24 with fresh resin.
25. Still working in the fume hood, wash the resin in the funnel with four 100-ml washes of ice-cold (≤4°C) water followed by two 100-ml washes of ice-cold 10 mM potassium phosphate, pH 8.0.
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26. Immediately transfer the resin to a 15-ml polypropylene screw-cap tube and add ∼4 ml of 10 mM potassium phosphate (pH 8.0) until the resin has the consistency of a thick slurry. Couple oligonucleotide multimers to CNBr-Sepharose 27. Immediately add the two 50-µl aliquots of DNA from step 19. Incubate on a rotating wheel overnight (≥8 hr) at room temperature. 28. In the fume hood, transfer the resin to a 60-ml coarse-sintered glass funnel and wash with two 100-ml washes of water and one 100-ml wash of 1 M ethanolamine hydrochloride, pH 8.0. Using a Geiger counter, compare the level of radioactivity in the first few milliliters of filtrate with the level of radioactivity in the washed resin to estimate the efficiency of incorporation of DNA to the resin. Usually, all detectable radioactivity is present in the resin.
29. In the fume hood, transfer the resin to a 15-ml polypropylene screw-cap tube and add 1 M ethanolamine hydrochloride (pH 8.0) until the mixture is a smooth slurry. Incubate the tube on a rotating wheel 2 to 4 hr at room temperature. This step inactivates unreacted CNBr-activated Sepharose. It is important to clean up all CNBr waste carefully. In the fume hood, add solid NaOH and glycine (∼10 to 20 mg/ml) to inactivate the CNBr. Soak contaminated instruments in a similar solution. Let sit overnight in the fume hood, then discard.
30. Wash the resin in a 60-ml coarse-sintered glass funnel with 100 ml of 10 mM potassium phosphate (pH 8.0), 100 ml of 1 M potassium phosphate (pH 8.0), 100 ml of 1 M KCl, 100 ml water, and 100 ml column storage buffer. 31. Store the resin at 4°C (stable at least 1 year; do not freeze). ALTERNATE PROTOCOL
COUPLING THE DNA TO COMMERCIALLY AVAILABLE CNBr-ACTIVATED SEPHAROSE The major advantage of this alternate procedure is that it begins with commercially available CNBr-activated chromatography resin, avoiding the need for preparation of CNBr-activated resin (see Basic Protocol 1). However, commercial CNBr-Sepharose is considerably more expensive than the homemade variety; moreover, the resulting column tends to run more slowly than one prepared as described above. Both resins are effective, leaving it up to the researcher’s discretion which to use. Additional Materials (also see Basic Protocol 1) 1 mM HCl (APPENDIX 2E), prepared fresh before use CNBr-activated Sepharose 4B (Pharmacia Biotech) 1. Prepare oligonucleotide multimers by phosphorylation and ligation (see Basic Protocol 1, steps 1 to 19). 2. Weigh out 3 g CNBr-activated Sepharose 4B (1 g freeze-dried resin gives ∼3.5 ml final gel volume).
Affinity Chromatography of DNA-Binding Proteins
3. Place the dry resin in a 15-ml conical polypropylene tube. Hydrate resin with 10 ml of 1 mM HCl and mix gently by flicking and inverting the tube. After 1 min, transfer slurry to a 60-ml coarse-sintered glass funnel. Wash and swell the beads by gradually pouring 500 ml of 1 mM HCl through the funnel (this will take ∼15 min).
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4. Wash the resin with 100 ml water and then with 100 ml of 10 mM potassium phosphate, pH 8.0. 5. Proceed with coupling of oligonucleotide multimers to the resin (see Basic Protocol 1, steps 26 to 31). PURIFICATION OF OLIGONUCLEOTIDES BY PREPARATIVE GEL ELECTROPHORESIS
SUPPORT PROTOCOL 1
Preparation of affinity resin requires a large amount (1-µmol synthesis) of purified synthetic oligonucleotides. The oligonucleotides used to prepare the affinity resin must be of high purity because contaminating, incompletely synthesized oligonucleotides can interfere with the ligation reaction. HPLC-purified oligonucleotides are of sufficient purity for preparation of DNA affinity resins; however, it is normally expensive to obtain such oligonucleotides. An alternative is to purify crude oligonucleotides by the following method. Additional Materials (also see Basic Protocol 1) 16% polyacrylamide-urea gel (see recipe) Oligonucleotides to be purified Formamide loading buffer (see recipe) sec-butanol (2-butanol) Diethyl ether 1 M MgCl2 (APPENDIX 2E) Saran wrap or other UV-transparent plastic wrap Intensifying screen (e.g., Lightning Plus, NEN Life Sciences) Hand-held short-wavelength UV light source Silanized glass wool Dry ice/ethanol bath (−78°C) Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (UNIT 10.3) Purify oligonucleotides by gel electrophoresis 1. Prepare a 20 cm × 40 cm × 1.5 mm denaturing polyacrylamide gel with four wells 3 cm in width, using 16% polyacrylamide/urea for separating oligonucleotides ∼10 to ∼45 bases long (or 8% or 6% polyacrylamide/urea for longer oligonucleotides). Let gel polymerize for ≥30 min, then prerun at 30 W for ≥1 hr. 2. Dissolve each oligonucleotide in formamide loading buffer to 200 µl final. Heat 15 min at 65°C to remove any secondary structure in the DNA. Load 50 µl of samples into separate wells, and run the gel at 30 W for ∼4 hr. The amount of oligonucleotide that is prepared in a 1-ìmol synthesis (∼1 to 2 mg) can be loaded on one gel (0.25 ìmol per 3-cm well). This is the maximum amount that can be applied to the gel without overloading it. It takes ∼4 hr for the bromphenol blue in the loading buffet to migrate three-quarters of the way down the gel, which is far enough to purify oligonucleotides of 15 to 30 bases. In a 16% gel, bromphenol blue comigrates with ∼10-base oligonucleotides and xylene cyanol comigrates with ∼30-base oligonucleotides. If the oligonucleotide is 25 to 35 bases long, it is recommended that the formamide loading buffer be made without xylene cyanol.
Visualize gel by UV shadowing 3. Remove one of the glass gel plates and cover the gel with Saran wrap. Flip gel over and remove the other plate so that the gel is lying on the Saran wrap. Cover the other side of the gel with Saran wrap.
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4. In a darkroom, lay the gel on an intensifying screen and hold a hand-held short-wavelength UV light source directly over it to visualize the DNA. Identify the major oligonucleotide band and mark its position directly on the Saran wrap using a marker, making sure that the light is directly over the band. The major band (usually the largest oligonucleotide but sometimes the next-to-largest) should be visible as a thick, dark band in the gel.
Purify oligonucleotides from gel 5. Carefully cut out the band with a razor blade, trying to avoid shredding the gel material or pulverizing the gel slice into small pieces. 6. Soak the gel piece in 5 ml TE buffer in a 15-ml polypropylene tube overnight at 37°C with shaking. 7. Place a silanized glass wool plug in a Pasteur pipet and prerinse with ∼5 ml water. Filter the supernatant containing the DNA through the glass wool. 8. Concentrate the DNA to ≤180 µl by repeated extractions with sec-butanol. 9. Extract the DNA once with diethyl ether and place in a vacuum evaporator (e.g., Speedvac) until all traces of ether are removed. 10. Adjust the volume of the liquid to 180 µl with TE buffer. Add 20 µl of 3 M sodium acetate and 2 µl of 1 M MgCl2. Mix by vortexing. 11. Add 600 µl of 100% ethanol. Mix by inversion. Chill 10 min in dry ice/ethanol. Let stand 5 min at room temperature. Microcentrifuge 15 min at high speed, room temperature, to pellet DNA. Discard supernatant. 12. Add 180 µl TE buffer. Dissolve pellet by vortexing. Add 20 µl of 3 M sodium acetate and 2 µl of 1 M MgCl2. Mix by vortexing. 13. Add 600 µl of 100% ethanol. Mix by inversion. Chill 10 min in dry ice/ethanol. Let stand 5 min at room temperature. Microcentrifuge 15 min at high speed, room temperature. Discard supernatant. 14. Add 800 µl of 75% ethanol. Mix by vortexing. Microcentrifuge 5 min at high speed, room temperature. Discard supernatant. 15. Carefully dry pellet in vacuum evaporator (sometimes static electricity will cause the pellet to jump out of the tube). 16. Dissolve DNA in 200 µl TE buffer and measure A260 and A280. Store at −20°C indefinitely. For sequence-specific DNA affinity resins, assume that 1 A260 unit = 40 ìg/ml DNA. BASIC PROTOCOL 2
Affinity Chromatography of DNA-Binding Proteins
DNA AFFINITY CHROMATOGRAPHY Affinity chromatography is performed by combining a partially purified protein sample with appropriate competitor DNA, pelleting the insoluble protein-DNA complexes by centrifugation, and loading the resulting soluble material by gravity flow onto the affinity resin. Nonspecific DNA-binding proteins flow through the column while the specific protein is retained by the column. The protein is then eluted by gradually increasing the salt concentration of the buffer and individual fractions are tested for DNA-binding activity and purity. Fractions containing appropriate DNA-binding activity can be reapplied to the affinity resin if further purification is desired. By using two sequential affinity chromatography steps, a typical protein can be purified 500- to 1000-fold with ∼30% yield. The method described below is performed with buffer Z, but many other buffers will work as well. Buffer choice is addressed in the Commentary.
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Materials Prepared DNA affinity resin (see Basic Protocol 1 or Alternate Protocol) Buffer Z or other column buffer (e.g., buffers Ze or TM; see recipes) made with varying KCl concentrations (buffers Z/0.1 M KCl through Z/1 M KCl) Partially purified protein fraction dialyzed against buffer Z/0.1 M KCl Nonspecific competitor DNA (see Support Protocol 2) Column regeneration buffer (see recipe) Column storage buffer (see recipe) Disposable chromatography column (Poly-Prep, Bio-Rad) Sorvall SS-34 rotor or equivalent Liquid nitrogen Narrow glass rod, silanized Additional reagents and equipment for DNA-binding assays (Brenowitz, et al., 1989; Buratowski and Chodosh, 1996; Baldwin et al., 1996), SDS-PAGE (UNIT 10.1), and silver staining (UNIT 10.5) 1. Equilibrate 1 ml settled bed volume of the DNA affinity resin in a disposable chromatography column with two 10-ml washes of buffer Z/0.1 M KCl. 2. Combine the partially purified protein fraction in buffer Z/0.1 M KCl with nonspecific competitor DNA as determined by DNA-binding studies (see Commentary and Support Protocol 2). 3. Incubate mixture 10 min on ice. 4. Centrifuge mixture 10 min at 12,000 × g (10,000 rpm in Sorvall SS-34 rotor), 4°C, to pellet insoluble protein-DNA complexes. 5. Load the supernatant onto the column at gravity flow (e.g., 15 ml/hr per column for Sepharose CL-2B). A single 1-ml column is sufficient for a standard nuclear extract (e.g., 12 liters of HeLa cells or 150 g of Drosophila embryos). When purifying a larger quantity of material, it is preferable to use multiple 1-ml columns. It is common practice to apply as much as 50 ml of protein sample onto a single 1-ml column. Typically, 1 ml of affinity resin contains ∼80 to 90 ìg DNA, which corresponds to a protein-binding capacity of 7 nmol/ml of resin, assuming one recognition site per 20 bp.
6. After loading the starting material, wash the column four times with 2-ml aliquots of buffer Z/0.1 M KCl, rinsing the sides of the column each time. It is very important to wash the affinity column thoroughly at this step by rinsing down the sides of the column with the 2-ml wash buffer aliquots; a single 8-ml wash is not effective. DNA affinity columns will often yield >100-fold purification of the desired factor. When a protein is purified 100-fold, however, a 1% contamination of the starting material due to inefficient washing of the column will lead to major contamination of the affinity-purified factor.
7. Elute the protein from the column by successive addition of 1-ml portions of buffer Z/0.2 M KCl, buffer Z/0.3 M KCl, buffer Z/0.4 M KCl, buffer Z/0.5 M KCl, buffer Z/0.6 M KCl, buffer Z/0.7 M KCl, buffer Z/0.8 M KCl, and buffer Z/0.9 M KCl, followed by three 1-ml aliquots of buffer Z/1 M KCl. Collect 1-ml fractions that correspond to the addition of the 1-ml portions of buffer. Quick-freeze the protein samples in liquid nitrogen and store at −80°C. Samples can generally be stored for at least 2 years.
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It is convenient to save separate aliquots of each fraction for DNA-binding assays (20 ìl each) and SDS-PAGE analysis (50 ìl each).
8. Assay the protein fractions for the sequence-specific DNA-binding activity using a DNA-binding assay (Brenowitz, et al., 1989; Buratowski and Chodosh, 1996; Baldwin et al., 1996). Estimate the purity of the protein fractions by SDS-PAGE (UNIT 10.1) followed by silver staining to visualize the protein. If further purification is desired, combine the fractions that contain the activity and, depending on the KCl concentration, either dilute (using buffer Z without KCl) or dialyze (against buffer Z/0.1 M KCl) to 0.1 M KCl. Then combine the protein fraction with nonspecific competitor DNA (the amount and type of which must be determined experimentally as described in Support Protocol 2) and reapply to either fresh or regenerated DNA affinity resin. In some instances, proteins might still be bound to the affinity resin after the 1 M salt elution step (step 7). Thus, if the desired protein is not detected in either the column fractions or flowthrough, it is possible that the protein is still on the column, and a wash at a higher salt concentration (e.g., 2 M KCl) may be needed to elute the protein from the resin.
9. Regenerate the affinity resin as follows: At room temperature, stop the column flow and add 5 ml column regeneration buffer to the column. Stir the resin with a silanized narrow glass rod to mix the resin with the regeneration buffer. Let the buffer flow out of the column. Repeat step. 10. To store column, add 10 ml column storage buffer and allow to flow through. Repeat this wash, then close the bottom of the column and add another 5 ml buffer. Cover top of column and store at 4°C. The column may be stored for ≤1 year. Alternatively, the affinity resin may be removed from the column after the two washes and stored in a clean polypropylene tube with the 5 ml buffer. SUPPORT PROTOCOL 2
Affinity Chromatography of DNA-Binding Proteins
SELECTION AND PREPARATION OF NONSPECIFIC COMPETITOR DNA Proper use of competitor DNA is essential for successful purification of DNA-binding proteins by affinity chromatography. Common competitor DNAs include poly(dI-dC), poly(dA-dT), poly(dG-dC), and calf thymus DNA. The amount and type of competitor to use in a given experiment must be determined experimentally. A typical method is to mix a protein sample (the exact fraction that will be applied to the affinity resin) with varying amounts of different competitor DNAs and evaluate DNA binding using assays such as DNase I footprinting (Brenowitz et al., 1989) or gel mobility shifts (Buratowski and Chodosh, 1996). The competitor DNA that inhibits DNA binding the least should be used for DNA affinity chromatography. It is perhaps easiest to think of these binding experiments as scaled-down affinity columns. First, determine the highest amount of competitor DNA that can be added to a DNA-binding reaction that does not interfere with the binding of the sequence-specific factor. To move up to a full-scale experiment, use one-fifth of the amount that would be required if the binding reaction were directly scaled up. The following example with a hypothetical factor M illustrates how to determine the amount of competitor to use in an affinity chromatography experiment. Using DNase I footprinting, a strong footprint was observed with 5 µl of a 0.4 M heparin fraction. Testing various competitors reveals that calf thymus DNA, poly(dI-dC), and poly(dG-dC) all strongly inhibit binding of factor M. However, no detectable inhibition of factor M binding is observed with 2 µg poly(dA-dT), weak but detectable inhibition is observed with 3 µg poly(dA-dT), and strong inhibition is observed with 4 µg poly(dA-dT). In this example,
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with 5 µl of the 0.4 M heparin fraction, the highest amount of poly(dA-dT) that does not inhibit factor M binding is 2 µg. If 5 ml of the 0.4 M heparin fraction is available, the direct scale-up would be 1000-fold, so 1000 × 2 µg = 2000 µg poly(dA-dT) would be needed. Because in an affinity chromatography experiment one-fifth of the direct scale-up amount is used, the appropriate amount of poly(dA-dT) to add to 5 ml of the 0.4 M heparin fraction is 2000 µg × 1/5 = 400 µg. The optimal amount of competitor DNA will vary with the purity of the partially purified protein sample. Fractions that contain more nonspecific DNA-binding proteins will normally require more competitor DNA and thus it is necessary to determine experimentally the optimal amount of DNA to use with each protein fraction. In addition, the amount of competitor DNA is estimated as the mass of DNA (in micrograms) to add per volume (in milliliters) of protein fraction; this is because it is likely that competitor DNA acts by forming a complex with high-affinity, nonspecific DNA-binding proteins in the crude extract. Different competitor DNAs can be used in a single experiment. To prepare poly(dI-dC), poly(dG-dC), and poly(dA-dT), dissolve the desired amount of competitor DNA to a final concentration of 10 A260 units in TE buffer per 100 mM NaCl. Heat the sample to 90°C and slowly cool to room temperature over 30 to 60 min. If the average length of the DNA is >1 kb, degrade it by sonication. Estimate the length of the DNA by agarose gel electrophoresis. REAGENTS AND SOLUTIONS Buffer TM 50 mM Tris⋅Cl, pH 7.9 (APPENDIX 2E) 0 M or 1 M KCl 12.5 mM MgCl2 1 mM dithiothreitol (DTT; add fresh just before use) 20% (v/v) glycerol 0.1% (v/v) Nonidet P-40 (NP-40) Do not make a 10× buffer. To generate aliquots of buffer with the range of KCl concentrations described in the protocol, make two 500-ml batches of 1× buffer containing no KCl and 1 M KCl respectively, and mix together appropriate quantities. Store at 4°C. Just prior to use, place 1.5-ml aliquots of each concentration in separate 1.5-ml microcentrifuge tubes, and add DTT. Buffer Z 25 mM HEPES (K+ salt), pH 7.6 0 M or 1 M KCl 12.5 mM MgCl2 1 mM DTT (add fresh just before use) 20% (v/v) glycerol 0.1% (v/v) NP-40 Adjust pH to 7.6 with KOH Do not make a 10× buffer. To generate aliquots of buffer with the range of KCl concentrations described in the protocol, make two 500-ml batches of 1× buffer containing no KCl and 1 M KCl respectively, and mix together appropriate quantities. Store at 4°C. Just prior to use, place 1.5-ml aliquots of each concentration in separate 1.5-ml microcentrifuge tubes, and add DTT.
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Buffer Ze 25 mM HEPES (K+ salt), pH 7.6 0 M or 1 M KCl 1 mM DTT (add fresh just before use) 20% (v/v) glycerol 0.1% (v/v) NP-40 Adjust the pH to 7.6 with KOH Do not make a 10× buffer. To generate aliquots of buffer with the range of KCl concentrations described in the protocol, make two 500-ml batches of 1× buffer containing no KCl and 1 M KCl respectively, and mix together appropriate quantities. Store at 4°C. Just prior to use, place 1.5-ml aliquots of each concentration in separate 1.5-ml microcentrifuge tubes, and add DTT. Column regeneration buffer 10 mM Tris⋅Cl, pH 7.8 (APPENDIX 2E) 1 mM EDTA, pH 8.0 2.5 M NaCl 1% (v/v) NP-40 Store at room temperature The solution will be cloudy and separate into two phases (NP-40 and aqueous) upon storage. Mix by swirling and shaking just before use. Column storage buffer 10 mM Tris⋅Cl, pH 7.8 (APPENDIX 2E) 1 mM EDTA, pH 8.0 0.3 M NaCl 0.04% (w/v) sodium azide Store at room temperature without sodium azide. Make a 4% (w/v) sodium azide stock solution and add just before use. Ethanolamine hydrochloride, pH 8.0, 1 M 1 M ethanolamine Adjust pH to 8.0 with HCl Filter sterilize Store at room temperature Formamide loading buffer 90 ml deionized formamide 10 ml 10× TBE (see recipe below) 40 mg xylene cyanol 40 mg bromphenol blue Store at −20°C Linker-kinase buffer, 10× 660 mM Tris⋅Cl, pH 7.6 (APPENDIX 2E) 100 mM MgCl2 100 mM DTT 10 mM spermidine Store at −20°C Add an extra 10 mM DTT just before use Affinity Chromatography of DNA-Binding Proteins
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16% polyacrylamide-urea gel 50 ml 40% (w/v) 19:1 acrylamide/bisacrylamide 12.5 ml 10× TBE (see recipe below) 62.5 g urea 17 ml H2O Mix, filter, briefly degas, and then add: 750 µl 10% (w/v) ammonium persulfate 20 µl TEMED Pour immediately into prepared gel plates.
T4 polynucleotide kinase buffer, 10× 500 mM Tris⋅Cl, pH 7.6 (APPENDIX 2E) 100 mM MgCl2 50 mM DTT 1 mM spermidine 1 mM EDTA, pH 8.0 Store at −20°C Add an extra 50 mM DTT just before use TBE (Tris/borate/EDTA) electrophoresis buffer, 10× 108 g Tris base (890 mM) 55 g boric acid (890 mM) 900 ml H2O 40 ml 0.5 M EDTA, pH 8.0 (20 mM) H20 to 1 liter COMMENTARY Background Information Purification of sequence-specific DNAbinding proteins has historically been a difficult task, mainly because such proteins are a small fraction of the total cellular protein. However, it is now possible to purify these factors quickly, simply, and effectively by using multimerized synthetic oligonucleotides that contain the recognition sequence for a particular DNA-binding protein. Early efforts with DNA-affinity chromatography involved adsorption or coupling of nonspecific DNA (such as calf thymus DNA) to either cellulose (Alberts and Herrick, 1971) or agarose (Arndt-Jovin et al., 1975) supports. These methods paved the way for the development of a variety of sequence-specific DNA affinity chromatography techniques, including the procedures described in this unit. Other methods have been described to purify sequence-specific DNA-binding proteins, including chromatography using biotinylated DNA fragments attached to various supports by biotin-avidin or biotin-streptavidin coupling (UNIT 10.6; Chodosh et al., 1986; Kasher et al., 1986; Leblond-Francillard et al., 1987), oligonucleotides synthesized onto Teflon-based beads (Duncan and Cavalier, 1988), or syn-
thetic oligonucleotide monomers attached to agarose supports (Wu et al., 1987; Blanks and McLaughlin, 1988; Hoey et al., 1993); and preparative gel mobility shifts (Gander et al., 1988). Although more than 50 sequence-specific DNA-binding proteins have been purified by the method described in this unit (Kadonaga, 1991, and references therein), it is likely that many of the techniques listed above are also effective for purifying sequence-specific DNAbinding proteins. There are a variety of reasons that CNBr activation is commonly used in the preparation of affinity resins: it is simple, works well with agarose matrices, and is mild enough to bind ligands such as DNA. Briefly, the chemistry of the CNBr activation reaction is as follows. At high pH, the hydroxyl groups on the agarose resin react with CNBr. The majority of the CNBr added to the reaction reacts with water to yield inert cyanate ions, which is part of the reason such a large amount of CNBr is required. Additionally, the majority of the cyanate esters that are formed on the agarose either are hydrolyzed to form inert carbamate or react with the matrix hydroxyls to form imidocarbonates. The imidocarbonates that form can act effectively
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as chemical cross-links, thus stabilizing the matrix (which is in most cases beneficial, particularly if the agarose resin chosen is not covalently cross-linked). The remaining active cyanate esters are coupled to the amino-containing ligands (in this case, oligonucleotides) at physiological pH. Finally, the unreacted cyanate esters are blocked with an excess of a suitable reagent, such as ethanolamine, to prevent coupling of the protein sample to the matrix (Janson and Rydén, 1989).
Critical Parameters and Troubleshooting Basic strategy The general approach to affinity purification of DNA-binding proteins is as follows. First, estimate the binding site of the desired protein by a technique such as DNase I footprinting (Brenowitz et al., 1989). If possible, it is best to survey a variety of promoters and enhancers. Next, determine the optimal conditions for protein binding to the DNA, considering factors such as temperature, ionic strength, pH, and Mg2+ concentration. Using conventional chromatography, partially purify the protein to remove contaminants such as nucleases that may degrade the affinity resin. Test a variety of nonspecific competitor DNAs, including poly(dI-dC), poly(dG-dC), poly(dA-dT), and calf thymus DNA to determine their effect on protein-DNA interactions. For best results, prepare two or more different DNA affinity resins with naturally occurring, high-affinity binding sites, preferably containing different flanking DNA sequences. The desired protein should bind with high affinity to both resins; if it does not, then it is likely that the protein that has been purified is not specific to the desired binding site. Finally, it is important to prepare a control resin that does not contain the recognition sequence for the desired protein. A control resin will make it possible to identify proteins that bind nonspecifically to DNA-Sepharose. Using this approach, it is possible to obtain a preparation containing a highly purified, sequencespecific DNA-binding protein.
Affinity Chromatography of DNA-Binding Proteins
Starting material and conventional chromatography Extensive purification of the DNA-binding protein is not required for effective affinity chromatography. However, partial purification of the protein by conventional chromatography is recommended prior to affinity chromatogra-
phy to remove proteases and nucleases that might degrade either the protein or the affinity resin. Keep in mind that sequence-specific and nonspecific DNA-binding proteins will often copurify in conventional chromatography, possibly because they may both interact with negatively charged polymers that resemble DNA. It is important to note that the most persistent contaminants in sequence-specific DNA-binding protein preparations are nonspecific, highaffinity DNA-binding proteins. Various conventional chromatographic methods may be employed prior to affinity purification of a DNA-binding protein; methods that have been used successfully in the past are discussed below. Ion-exchange chromatography. One obvious first step might be cation-exchange chromatography with resins such as S-Sepharose Fast Flow/Mono S (Pharmacia Biotech), CM52 (Whatman), CM Sepharose Fast Flow (Pharmacia Biotech), P11 phosphocellulose (Whatman), or Bio-Rex 70 (Bio-Rad). DNA-binding proteins (both specific and nonspecific) will normally bind to cation-exchange resins in buffers containing from 50 mM to 100 mM NaCl, and can be eluted with higher salt concentrations. Affinity chromatography. Nonspecific DNA-cellulose (Alberts and Herrick, 1971) or DNA-agarose (Arndt-Jovin et al., 1975) resins can be used for a preliminary purification step. These resins are typically prepared with salmon sperm or calf thymus DNA. Also, sequencespecific DNA affinity resins can be prepared with oligonucleotides that do not contain binding sites for the desired factor. A strategy for chromatography with either nonspecific DNA affinity resin (e.g., see Rosenfeld and Kelly, 1986) or sequence-specific DNA affinity resins that lack the binding site for the desired factor (e.g., see Kaufman et al., 1989) is as follows. Once the protein sample is applied to a nonspecific DNA affinity resin, the desired factor will either (1) flow through the resin or (2) elute from the resin at a low salt concentration. Highaffinity, nonspecific DNA-binding proteins should remain bound to the resin and will thus be separated from the desired protein. If this strategy is successful, it will not be necessary to add the competitor DNA prior to the subsequent sequence-specific affinity chromatography step because the high-affinity nonspecific DNA-binding proteins will already have been separated from the sample.
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A resin that works well if the desired protein contains O-linked N-acetylglucosamine monosaccharide residues is wheat germ agglutininagarose. This method has been used successfully in the past with transcription factors such as Sp1 (Jackson and Tjian, 1989) and HNF1 (Lichtsteiner and Schibler, 1989). Heparin-agarose chromatography. Heparin-agarose resins are also excellent for the purification of DNA-binding proteins because they possess properties similar to those of both ion-exchange and affinity resins. It is important to be aware that variability exists between different heparin-agarose preparations. This variability may be due to differences in the methods of coupling heparin to agarose, because alternate methods for coupling and blocking the agarose may produce different functional groups on the resin. If a particular batch of heparin-agarose is used successfully, it is wise to continue to use that batch to ensure reproducibility. Gel-filtration chromatography. Finally, gelfiltration chromatography may be used effectively for partial purification of a DNA-binding protein because separation is based on size and shape, not DNA-binding properties. For example, gel filtration may be useful for separating a desired protein from contaminating nucleases and other nonspecific DNA-binding proteins, in contrast to chromatography using ion-exchange, heparin-agarose, and nonspecific DNA affinity resins that may actually enrich for DNA-binding proteins. In some cases gel filtration may be undesirable because significant sample dilution occurs; however, further concentration can be obtained by subsequent affinity chromatography.
presence. Thus, in some cases it may be useful to omit Mg2+ from the chromatography buffer. In addition, although the procedures in this unit use buffers containing KCl, NaCl could probably be substituted. Although it is usually preferable to handle proteins at 4°C, proteins have been described that will not bind DNA at 4°C, yet bind perfectly well at room temperature; in such cases the temperature at which the affinity chromatography is performed should be adjusted accordingly. Finally, to minimize nonspecific adsorption of affinity-purified proteins to plastic and glass, chromatography and storage buffers should contain a nonionic detergent such as NP-40 at a concentration of ∼0.01% to 0.1% (v/v). Binding assays. There are many methods for identifying proteins that are bound to DNA, including DNAse I footprinting (Galas and Schmitz, 1978; Brenowitz et al., 1989), Fe-EDTA footprinting (Tullius et al., 1987), methidiumpropyl-EDTA⋅Fe2+ (MPE; Hertzberg and Dervan, 1982), methylation interference (Baldwin et al., 1996), and gel mobility shift assays (Buratowski and Chodosh, 1996). Although binding information may be obtained from all these techniques, it is most straightforward to use DNase I footprinting to identify a protein binding site. Additionally, DNase I is active over a broad range of buffer conditions, which allows for considerable variation in binding conditions. Visualization of the protected region by footprinting makes designing oligonucleotides for affinity chromatography relatively simple (as discussed later in this section). Both DNase I footprinting and gel mobility shift assays are useful for assaying affinity-column fractions.
DNA binding studies Binding conditions. In order to use affinity chromatography to purify a DNA-binding protein, it is necessary to optimize binding conditions. It is important to determine these conditions experimentally, as ionic strength, temperature, pH, and presence or absence of Mg2+ can affect protein-DNA interactions. In addition, although HEPES and PIPES are often considered to be superior to Tris buffers, some commercially available preparations of HEPES and PIPES buffers contain contaminants that inhibit binding of proteins to DNA. Therefore, in some respects, it may be safer to use Tris buffer. It is also important to note that some factors, such as Sp1, bind with higher affinity to DNA in the absence of Mg2+ than in its
Design of oligonucleotides Many different parameters are important in designing oligonucleotides for an affinity resin. The first step, as described above, is to determine the protein-binding site on the DNA. The DNA affinity resin should be prepared from naturally occurring, high-affinity binding sites. Using a consensus sequence may work in some instances, but in most situations a “real” binding site is best. Given that the length of the oligonucleotides can be reasonably flexible (14-mers to 61-mers have been used successfully), use of oligonucleotides that include the DNase I footprint as well as some flanking DNA (at least a few bases beyond the borders of the footprint) is recommended. In the early stages, it is better to copurify factors that bind
Affinity Purification
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Affinity Chromatography of DNA-Binding Proteins
to the flanking region than to fail to purify the desired factor because the binding site is too short. It may be wise to avoid using oligonucleotides that are 21 or 42 bases in length because the DNA, if bent, might have a greater tendency to circularize during the ligation step. The oligonucleotides should also be designed with a single-stranded overhang, for two reasons: first, the ligation reaction will be more efficient (as compared to a blunt-end ligation), and second, it is likely that the DNA couples to the resin via the primary amine groups of the single-stranded overhang of the ligated multimers. The sequence GATC-XXX-BINDING SITE-XXX (where XXX represents the DNA flanking the binding site) works well, though it may also be convenient to use the sequences flanking the binding site as the overhang.
centration of affinity-purified proteins is typically 5 to 50 µg/ml. About 0.2 to 1 µl of protein (2 to 10 ng) can be expected to be sufficient for a “blanked-out” footprint (with ∼10 fmol of DNA probe). Affinity resin is stable for >1 year if stored at 4°C. Affinity-purified proteins are typically stable for >2 years if stored at −80°C in an appropriate buffer and handled properly. Proteins should always be quick-frozen in liquid nitrogen and quick-thawed in either cold or room-temperature water (although if roomtemperature water is used to accelerate thawing, the sample must not be allowed to warm up above 4°C). Although many DNA-binding proteins remain active after several freeze-thaw cycles, it is preferable to divide protein preparations into small aliquots rather than to freeze and thaw the entire sample many times.
Inadvertent purification of nonspecific DNA-binding proteins Throughout this unit, an attempt has been made to emphasize how to avoid inadvertent purification of nonspecific DNA-binding proteins. Unfortunately, this problem is quite common, particularly when the gel mobility shift assay is used to monitor DNA binding. Proper use of the control resin (to identify proteins that bind nonspecifically to DNA-Sepharose), two different affinity resins (to identify proteins that might be interacting with the linker DNA), and competitor DNA (to deplete nonspecific DNAbinding proteins from the extract) should virtually eliminate purification of nonspecific DNA-binding proteins. In addition, DNase I footprinting is recommended for assaying DNA binding because it allows visualization of the actual protein binding site and to confirm sequence specificity. To correlate DNA binding with a particular polypeptide, it would be informative to purify the polypeptide from a polyacrylamide gel by using the denaturation/renaturation procedure described by Hager and Burgess (1980). Two common nonspecific DNA-binding proteins that have been purified from HeLa cells by affinity chromatography are poly(ADP-ribose) polymerase, which has an Mr of 116,100 (Ueda and Hayaishi, 1985; Slattery et al., 1983), and the Ku antigen, which consists of two polypeptides of Mr 70,000 and 80,000 (Mimori et al., 1986).
Time Considerations
Anticipated Results
Arndt-Jovin, D.J., Jovin, T.M., Bähr, W., Frischauf, A.-M., and Marquardt, M. 1975. Covalent attachment of DNA to agarose: Improved synthesis and use in affinity chromatography. Eur. J. Biochem. 54:411-418.
After two rounds of affinity chromatography, it is possible to achieve a 500- to 1000-fold purification with a 30% overall yield. The con-
The time required to optimize DNA-binding conditions of a particular protein is variable and depends on assay type, probe preparation time, and competitor choices. The time to prepare extracts and to perform conventional chromatography is also variable depending on factors such as starting material and type of chromatography. Beginning with purified oligonucleotides, preparation of the DNA affinity resin takes 2 days (with either commercially available or homemade CNBr-activated resin). If the oligonucleotides are to be gel-purified, an additional day and a half should be added. Affinity chromatography takes ∼4 to 8 hr, dependent mainly on the flow rate of the resin: Sepharose CL-2B runs at ∼15 ml/hr, while Sepharose-4B runs somewhat slower. Therefore, it is possible to estimate chromatography time by calculating the volume of sample that is to be applied to the column plus 8 ml for the wash steps and 11 ml for the elution steps. If the volume of starting material is not too large, it is possible to assay the affinity column on the day it is run (either by DNA-binding assays, SDS-PAGE, or both). It is important to freeze the affinity fractions in liquid nitrogen and store at −80°C as soon as the column is complete.
Literature Cited Alberts, B. and Herrick, G. 1971. DNA-cellulose chromatography. Methods Enzymol. 21:198217.
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Baldwin, A.S., Oettinger, M., and Struhl, K. 1996. Methylation and uracil interference assays for analysis of protein-DNA interactions. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 12.3.112.3.7. John Wiley & Sons, New York. Blanks, R. and McLaughlin, L.W. 1988. An oligodeoxynucleotide affinity column for the isolation of sequence specific DNA binding proteins. Nucl. Acids Res. 16:10283-10299. Brenowitz, M., Senear, D.F., and Kingston, R.E. 1989. DNase I footprint analysis of protein-DNA binding. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 12.4.1-12.4.16. John Wiley & Sons, New York. Buratowski, S. and Chodosh, L. 1996. Mobilityshift DNA-binding assay using gel electrophoresis. In Current Protocols in Molecular Biology. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 12.2.1-12.2.11. John Wiley & Sons, New York. Chodosh, L.A., Carthew, R.W., and Sharp, P.A. 1986. A single polypeptide possesses the binding and transcription activities of the Adenovirus major late transcription factor. Mol. Cell. Biol. 6:4723-4733. Duncan, C.H. and Cavalier, S.L. 1988. Affinity chromatography of a sequence-specific DNA binding protein using Teflon-linked oligonucleotides. Anal. Biochem. 169:104-108. Galas, D. and Schmitz, A. 1978. DNase footprinting: A simple method for the detection of protein-DNA binding specificity. Nucl. Acids Res. 5:3157-3170. Gander, I., Foeckler, R., Rogge, L., Meisterernst, M., Schneider, R., Mertz, R., Lottspeich, F., and Winnacker, E.L. 1988. Purification methods for the sequence-specific DNA-binding protein nuclear factor I (NFI)—generation of protein sequence information. Biochim. Biophys. Acta 951:411-418. Hager, D.A. and Burgess, R.R. 1980. Elution of proteins from sodium dodecyl sulfate polyacrylamide gels, removal of sodium dodecyl sulfate, and renaturation of enzymatic activity: Results with sigma subunit of Escherichia coli RNA polymerase, wheat germ DNA topoisomerase, and other enzymes. Anal. Biochem. 109:76-86. Hertzberg, R.P. and Dervan, P.B. 1982. Cleavage of double helical DNA by (methidiumpropylEDTA)iron(II). J. Am. Chem. Soc. 104:313-315. Hoey, T., Welnzierl, R.O., Gill, G., Chen, J.L., Dynlacht, B.D., and Tjian, R. 1993. Molecular cloning and functional analysis of Drosophila TAF110 reveal properties expected of coactivators. Cell 72:247-260.
Jackson, S.P. and Tjian, R. 1989. Purification and analysis of RNA polymerase II transcription factors using wheat germ agglutinin affinity chromatography. Proc. Natl. Acad. Sci. U.S.A. 86:1781-1785. Janson, J.-C. and Rydén, L. 1989. Protein Purification. Principles, High Resolution Methods, and Applications. VCH Publishers, New York. Kadonaga, J.T. 1991. Purification of sequence-specific DNA binding proteins by DNA affinity chromatography. Methods Enzymol. 208:10-23. Kasher, M.S., Pintel, D., and Ward, D.C. 1986. Rapid enrichment of HeLa transcription factors IIIB and IIIC by using affinity chromatography based on avidin-biotin interactions. Mol. Cell. Biol. 6:3117-3127. Kaufman, P.D., Doll, R.F., and Rio, D.S. 1989. Drosophila P element transposase recognizes internal P element DNA sequences. Cell 59:359371. Leblond-Francillard, M., Dreyfus, M., and Rougeon, F. 1987. Isolation of DNA-protein complexes based on streptavidin and biotin interaction. Eur. J. Biochem. 166:351-355. Lichtsteiner, S. and Schibler, U. 1989. A glycosylated liver-specific transcription factor stimulates transcription of the albumin gene. Cell 57:1179-1187. Mimori, T., Hardin, J.A., and Steitz, J.A. 1986. Characterization of the DNA-binding protein antigen Ku recognized by autoantibodies from patients with rheumatic disorders. J. Biol. Chem. 261:2274-2278. Rosenfeld, P.J. and Kelly, T.J. 1986. Purification of nuclear factor I by DNA recognition site affinity chromatography. J. Biol. Chem. 261:1398-1408. Slattery, E., Dignam, J.D., Matsui, T., and Roeder, R.G. 1983. Purification and analysis of a factor which suppresses nick-induced transcription by RNA polymerase II and its identity with poly(ADP-ribose) polymerase. J. Biol. Chem. 258:5955-5959. Tullius, T.D., Dombroski, B.A., Churchill, M.E.A., and Kam, L. 1987. Hydroxyl radical footprinting: A high resolution method for mapping protein-DNA contacts. Methods Enzymol. 155:537558. Ueda, K. and Hayaishi, O. 1985. ADP-ribosylation. Annu. Rev. Biochem. 54:73-100. Wu, C., Wilson, S., Walker, B., Dawid, I., Paisley, T., Zimarino, V., and Ueda, H. 1987. Purification and properties of Drosophila heat shock activator protein. Science 238:1247-1253.
Key References Kadonaga, J.T. 1991. See above. Techniques paper, though less descriptive than this unit, containing a table that lists (with references) >50 sequence-specific proteins that have been purified using the affinity chromatography method described herein.
Affinity Purification
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Kadonaga, J.T. and Tjian, R. 1986. Affinity purification of sequence-specific DNA binding proteins. Proc. Natl. Acad. Sci. U.S.A. 83:58895893. First paper to describe affinity chromatography with multimerized oligonucleotides; details purification of transcription factor Sp1.
Contributed by Leslie A. Kerrigan Osiris Therapeutics Baltimore, Maryland James T. Kadonaga University of California San Diego La Jolla, California
Affinity Chromatography of DNA-Binding Proteins
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Purification of DNA-Binding Proteins Using Biotin/Streptavidin Affinity Systems
UNIT 9.7
Short fragments of DNA—either natural or formed from oligonucleotides—containing a high-affinity site for a DNA-binding protein provide a powerful tool for purification. The biotin/streptavidin purification system is based on the tight and essentially irreversible complex that biotin forms with streptavidin. The experimental design of this system is illustrated in Figure 9.7.1. First, a DNA fragment is prepared that contains a high-affinity binding site for the protein of interest. A molecule of biotinylated nucleotide is incorporated into one of the ends of the DNA fragment. The protein of interest is allowed to bind to the high-affinity recognition site present in the biotinylated fragment. The tetrameric protein streptavidin is then bound to the biotinylated end of the DNA fragment. Next, the protein/biotinylated fragment/streptavidin ternary complex is efficiently removed by adsorption onto a biotin-containing resin. Since streptavidin is multivalent, it is able to serve as a bridge between the biotinylated DNA fragment and the biotin-containing resin. Proteins remaining in the supernatant are washed away under conditions that maximize the stability of the DNA-protein complex. Finally, the protein of interest is eluted from the resin with a high-salt buffer.
binding site for protein X DNA end-labeled with biotin-11-dUTP
+
protein X
+ streptavidin tetramer
+ biotin cellulose biotin-cellulose protein X
high-salt wash
Figure 9.7.1 Purification of DNA-binding proteins using the biotin/streptavidin affinity technique. The protocol involves the following steps: (1) a biotinylated, labeled DNA fragment is prepared containing a binding site for the protein to be purified; (2) the biotin-cellulose resin is prepared; (3) a binding reaction containing a crude protein fraction and the biotinylated probe is set up; (4) free streptavidin is added to the binding reaction; (5) the protein/biotinylated DNA fragment/streptavidin complex is bound to the biotin-cellulose resin; (6) unbound protein is removed by extensive resin washing; and (7) the protein is eluted from the resin with high-ionic-strength buffer. Each of these steps can be monitored and optimized in solution, using a mobility shift DNA-binding assay (see Support Protocol and Buratowski and Chodosh, 1996). Contributed by Lewis A. Chodosh and Stephen Buratowski Current Protocols in Protein Science (1998) 9.7.1-9.7.13 Copyright © 1998 by John Wiley & Sons, Inc.
Affinity Purification
9.7.1 Supplement 12
BASIC PROTOCOL
PURIFICATION USING BATCH METHOD Materials Plasmid DNA with binding site for the protein of interest Appropriate restriction endonucleases Biotin-11-dUTP (Life Technologies) Labeled and unlabeled dNTPs Klenow fragment of E. coli DNA polymerase I TE buffer (APPENDIX 2E) Biotin-cellulose (Pierce) Biotin-cellulose binding buffer (see recipe) BSA Bulk carrier DNA [e.g., poly(dI-dC)⋅poly(dI-dC), salmon sperm DNA, or E. coli DNA] Biotin-cellulose elution buffer (see recipe) Protein solution Streptavidin (Celltech; may be stored as 5 mg/ml stock for at least 2 months) DEAE membrane (Schleicher & Schuell NA45) 0.025-µm filter discs (Millipore VS) Additional reagents and equipment for mobility-shift DNA binding assay (see Support Protocol) Prepare biotinylated, labeled DNA fragment 1. Digest 50 µg of plasmid DNA in 100 µl with one or more appropriate restriction endonucleases to obtain DNA probe. 2. Add the following to the probe mixture: Biotin-11-dUTP to a final concentration of 20 µM Radioactive dNTP for incorporation into the 5′ overhang 100-fold molar excess of corresponding unlabeled dNTP Remaining two unlabeled dNTPs to a final concentration of 200 µM 5 U Klenow fragment. This reaction is identical to that used in standard DNA-binding assays (e.g., see Support Protocol), except biotinylated dUTP is incorporated into one end of the DNA fragment in place of TTP and the fragment is radiolabeled to low instead of high specific activity.
3. Precipitate the biotinylated probe. Isolate the probe by agarose gel electrophoresis using DEAE membrane. Gel purification of the biotinylated fragment removes unreacted biotin-11-dUTP. This is essential because any free biotin-dUTP binds to streptavidin and then to the biotin-cellulose column, reducing the apparent capacity of both materials to react with protein.
4. Resuspend the probe in TE buffer and measure an aliquot for Cerenkov counts. Estimate the DNA concentration by ethidium bromide dot quantitation. 5. Test the biotinylated probe to be certain it will efficiently bind to the protein of interest. Use a standard binding assay (e.g., see Support Protocol) with the biotinylated fragment as probe.
Purification of DNA-Binding Proteins Using Biotin/Streptavidin Affinity Systems
Prepare biotin-cellulose resin 6. Place 200 µl biotin-cellulose in a 1.5-ml microcentrifuge tube. Spin the resin for 30 sec in a microcentrifuge and remove the supernatant. Add to the pellet: 1.0 ml biotin-cellulose binding buffer 500 µg/ml BSA 200 µg carrier DNA
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Gently mix the tube 5 min on a rotating wheel. This step blocks nonspecific protein and nucleic acid binding sites present on the biotincellulose resin.
7. Spin the resin, remove the supernatant, and resuspend pellet in 1.0 ml of biotin-cellulose elution buffer. Gently mix 5 min on a rotating wheel. Repeat this wash. Washing removes molecules on the biotin-cellulose resin that might later be eluted from the resin by the biotin-cellulose elution buffer.
8. Spin the resin, remove the supernatant, and resuspend pellet in 1.0 ml of biotin-cellulose binding buffer. Repeat this wash. This 1:6 dilution of pretreated biotin-cellulose is ready for use and can be stored for several months.
Set up binding reaction 9. Determine the molar concentration of the protein to be purified via a mobility-shift assay (e.g., see Support Protocol). 10. Set up a standard binding reaction containing the protein to be purified, carrier DNA, and a 10-fold molar excess of biotinylated fragment relative to the protein to be purified. Allow the reaction to go to completion for ∼15 min. Use reaction conditions that optimize protein binding to its recognition site. The composition of the biotin-cellulose binding buffer should be the same as that optimized for protein-DNA binding.
11. Add a 5-fold molar excess of streptavidin relative to the biotinylated fragment. Continue the binding reaction for an additional 5 min at 30°C. Bind protein/DNA/streptavidin complex to biotin-cellulose resin 12. In a separate tube, place 2 µl pretreated biotin-cellulose (12 µl of the 1:6 dilution) for each picomole of biotinylated DNA fragment in the binding reaction. Spin the resin and remove the supernatant. One or two microliters of biotin-cellulose can easily be seen at the bottom of the microcentrifuge tube.
13. Transfer the binding reaction mix into the tube with the biotin-cellulose resin using a pipettor. Gently resuspend the resin and incubate on a rotating wheel for 30 min. This incubation can be done either at 4°C or at room temperature, depending on the stability of the protein.
14. Spin the resin and remove the supernatant. The supernatant should be measured using a mobility shift assay (e.g., see Support Protocol) to determine what percentage of the biotinylated fragment and the protein has been removed from the supernatant. It is also useful to assay the supernatant from the binding reaction for a control DNA-binding protein to determine whether the protein of interest has been specifically depleted or whether multiple DNA-binding proteins have been depleted nonspecifically. The latter observation would suggest that the matrix is acting as a nonspecific DNA-affinity column. For some applications, a protein fraction specifically depleted for a particular DNA-binding protein is a valuable reagent.
Wash the resin 15. Resuspend the biotin-cellulose pellet in 500 µl biotin-cellulose binding buffer. Mix by gently inverting the tube 1 to 2 min. Spin the resin and remove the supernatant.
Affinity Purification
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Repeat this procedure twice. The second time, transfer to a clean microcentrifuge tube. Transferring the reaction avoids elution of proteins that were bound nonspecifically to the walls of the tube in the first binding incubation.
Elute the protein 16. Resuspend the biotin-cellulose pellet in at least an equal volume of biotin-cellulose elution buffer. Mix gently on a rotating wheel for 20 min. The salt concentration in the biotin-cellulose elution buffer must be determined empirically. Successively higher salt concentrations may be tested until the concentration of eluted protein is maximal.
17. Spin the resin. Save the supernatant and assay for binding activity. Small volumes of protein solutions can be dialyzed effectively on 0.025-ìm filter discs. ALTERNATE PROTOCOL 1
PURIFICATION USING A MICROCOLUMN Although the batch method in the basic protocol is rapid and well-suited for analyticalscale purification, larger volumes of biotin-cellulose resin can be better handled in a microcolumn. This method is also used to elute the protein in as small a volume (i.e., as high a concentration) as possible. Additional Materials (also see Basic Protocol) Silanized glass wool 1.0-ml pipet tip Ring stand 1. Prepare the biotinylated DNA fragment and biotin-cellulose resin and set up binding reaction (see Basic Protocol, steps 1 to 11). 2. Place a small plug of silanized glass wool in the bottom of a 1.0-ml pipet tip. Firmly attach the pipet-tip microcolumn to a ring stand. Prewet the glass wool in biotin-cellulose binding buffer before insertion into the pipet tip to avoid trapping air bubbles which might denature proteins.
3. Add 500 µl binding buffer to the microcolumn. Maintain a steady flow through the glass wool plug. If the column does not flow smoothly, a pipettor can be gently inserted into the top of the microcolumn. Slightly depressing the plunger will start the column or increase the flow.
4. Add at least 40 µl of 1:1 biotin-cellulose slurry to the microcolumn. Allow the buffer to run down to the surface of the resin. 5. Equilibrate the resin with 3 column volumes of biotin-cellulose binding buffer if the resin has already been pretreated (Basic Protocol, steps 6 to 8). If the resin has not been pretreated, wash sequentially with 3 column volumes each of biotin-cellulose binding buffer, biotin-cellulose binding buffer with 500 µg/ml BSA and 200 µg/ml poly(dI-dC)⋅poly(dI-dC), and biotin-cellulose elution buffer. Finally, equilibrate with biotin-cellulose binding buffer. Purification of DNA-Binding Proteins Using Biotin/Streptavidin Affinity Systems
The biotin-cellulose microcolumn can be washed very rapidly. Each wash takes 2 or 3 min. The drop size from the pipet tip is ∼25 ìl but this will change with alterations in the ionic strength and protein concentration of the eluate.
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6. Load the binding reaction mix (Basic Protocol, step 11) onto the microcolumn and collect the flowthrough. The column can be run as fast as 6 to 10 column volumes/hr without affecting the amount of biotinylated fragment bound by the resin. If the flow rate is too slow, use a pipettor to apply pressure to the column. If the flow rate is too fast, plug the tip of the microcolumn with Parafilm in between drops.
7. Wash with 4 column volumes of biotin-cellulose binding buffer. Discard the flowthrough. 8. Wash with 3 column volumes of biotin-cellulose elution buffer. Collect 2-drop fractions and assay. Fractions as small as 5 ìl may be rapidly and effectively dialyzed on 0.025-ìm filter discs with minimal loss of volume and activity. Float a filter (shiny side up) in a petri dish on top of 20 ml dialysis buffer. Allow the filter 10 min to wet. Place the sample (5 to 100 ìl) to be dialyzed onto the surface of the filter. Surface tension will keep the sample confined in a drop unless there is a detergent in the sample. After 1 hr remove the sample. Once the protein has been eluted from the resin, the resin is effectively the same as a sequence-specific DNA-affinity microcolumn.
PURIFICATION USING STREPTAVIDIN-AGAROSE When high-quality free streptavidin is not available or cellulose is an inappropriate resin, a simple variation on the basic protocol may be employed. In this protocol, the same biotinylated DNA fragment is used but is removed from solution directly by streptavidinagarose (see Fig. 9.7.2).
ALTERNATE PROTOCOL 2
Additional Materials (also see Basic Protocol) Streptavidin-agarose 1. Prepare biotinylated DNA fragment and resin, substituting streptavidin-agarose for biotin-cellulose, and set up binding reaction (see Basic Protocol, steps 1 to 11).
binding site for protein X
+
DNA end-labeled with biotin-11-dUTP
protein X
+ streptavidin agarose beads
streptavidin protein X
high-salt wash
Figure 9.7.2 Purification of DNA-binding proteins using streptavidin-agarose.
Affinity Purification
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2. In a separate tube, add 50 µl pretreated streptavidin-agarose (300 µl of the 1:6 dilution) for each picomole of biotinylated DNA fragment in the binding reaction. Spin the resin and remove the supernatant. 3. Transfer the binding-reaction mix into the tube with the streptavidin-agarose using a pipettor. Gently resuspend the resin and incubate on a rotating wheel for 30 min to 2 hr. 4. Wash and elute the protein (see Basic Protocol, steps 14 to 17). Like biotin-cellulose, streptavidin-agarose may also be used in a microcolumn. Follow the microcolumn alternate protocol. SUPPORT PROTOCOL
MOBILITY-SHIFT ASSAY The DNA-binding assay using nondenaturing polyacrylamide gel electrophoresis (PAGE) provides a simple, rapid, and extremely sensitive method for detecting sequence-specific DNA-binding proteins. Proteins that bind specifically to an end-labeled DNA fragment retard the mobility of the fragment during electrophoresis, resulting in discrete bands corresponding to the individual protein-DNA complexes. The assay can be used to test binding of purified proteins (see Basic Protocol) or of uncharacterized factors found in crude extracts. This assay also permits quantitative determination of the affinity, abundance, association rate constants, dissociation rate constants, and binding specificity of DNA-binding proteins. The utility of this technique is underscored by the many proteins that have been characterized using this assay. It has become clear that there is no single protocol that works best for all proteins. Rather, several variables can be changed to optimize binding. There are several options available for the design of the DNA probe, the binding reaction conditions, and the gel running conditions. The reader is referred to Buratowski and Chodosh (1996) for additional details and protocols. This protocol can be divided into four stages: (1) preparation of a radioactively labeled DNA probe containing a particular protein binding site; (2) preparation of a nondenaturing gel; (3) a binding reaction in which a protein mixture is bound to the DNA probe; and (4) electrophoresis of protein-DNA complexes through the gel, which is then dried and autoradiographed. Consult Current Protocols in Molecular Biology (Ausubel et al., 1998) or Sambrook et al. (1989) for details and protocols for basic molecular biological techniques referred to below. DNA fragments from 20 to 300 bp long may be used as probes. However, longer fragments are likely to contain binding sites for multiple proteins, which may make interpretation of the gel difficult. The DNA probe can be prepared in one of several ways, outlined below.
Purification of DNA-Binding Proteins Using Biotin/Streptavidin Affinity Systems
Materials 10× electrophoresis buffer: e.g., TAE or TBE electrophoresis buffer (APPENDIX 2E) or Tris-glycine electrophoresis buffer (UNIT 10.3) 30% (w/v) ammonium persulfate, prepared fresh N,N,N′,N′-tetramethylethylenediamine (TEMED) Nondenaturing gel mix (see recipe) Bulk carrier DNA, e.g., poly(dI-dC)⋅poly(dI-dC) BSA Protein preparation containing DNA-binding protein (crude extract or purified fraction) 10× loading buffer (see recipe)
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Constant-temperature water bath Two-head peristaltic pump 10-µl glass capillary pipet (optional) Clay-Adams screw-top loader (optional) Whatman 3MM filter paper (or equivalent) Additional reagents and equipment for digesting DNA with restriction endonucleases, DNA labeling, agarose and nondenaturing polyacrylamide gel electrophoresis, recovery of DNA from gels, oligonucleotide synthesis, PCR, ethanol precipitation, ethidium bromide dot quantitation, and autoradiography (see Ausubel et al., 1998, or Sambrook et al., 1989) Prepare the DNA probe 1a. For restriction endonuclease fragments: Isolate a small DNA fragment containing the binding site of interest from a plasmid using a standard restriction endonuclease digestion. Label the fragment by filling in a 5′ overhang with the Klenow fragment of Escherichia coli DNA polymerase and 32P-labeled nucleotide or by end labeling using polynucleotide kinase. Separate the fragment from the plasmid by gel electrophoresis. Kinased probes should be avoided in experiments using crude protein preparations that might contain phosphatase activity. Agarose gels are useful for resolving fragments as small as 50 bp. Fragments can be recovered using low-melting-temperature agarose or DEAE paper. For smaller DNA fragments, nondenaturing polyacrylamide gels can be used.
1b. For synthetic oligonucleotides: Synthesize and anneal complementary oligonucleotides to generate a double-stranded DNA fragment containing the binding site of interest. Label the probe using polynucleotide kinase. The kinase reaction can be performed before annealing to label only one strand, or after annealing to label both strands. Alternatively, the oligonucleotides can be designed to leave an overhang that can be filled in with labeled nucleotide and Klenow fragment.
1c. For PCR fragments: Generate a DNA fragment containing the binding site of interest by polymerase chain reaction (PCR). End-label one of the primers with polynucleotide kinase before the PCR reaction, or label the double-stranded PCR product after purification. 2. Following isolation of the probe, determine its concentration by ethidium bromide dot quantitation. Concentrations in the range of 2 to 50 ng/ìl are convenient.
3. Count 1 µl for Cerenkov counts in a scintillation counter to determine specific activity (cpm/µl). A typical binding reaction will contain about ∼5,000 to 20,000 cpm and ∼10 to 100 fmol probe (10 fmol DNA in a final reaction volume of 10 ìl gives a total DNA concentration of 1 nM). If desired, probes can be stored up to 4 to 6 weeks at 4°C before proceeding with the experiment.
Prepare the nondenaturing gel 4. Dilute 10× electrophoresis buffer to prepare enough 1× electrophoresis buffer to fill the tank. 5. Assemble washed glass plates and 1.5-mm spacers for casting the gel. All traces of detergent must be removed because detergent will disrupt protein-DNA interactions.
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6. Add 150 µl of 30% ammonium persulfate and 70 µl TEMED to 60 ml nondenaturing gel mix prepared using the same buffer as that used for electrophoresis. Swirl gently to mix. The amounts of ammonium persulfate, TEMED, and nondenaturing gel mix can be scaled up or down as necessary depending on the size and number of gels.
7. Pour the gel mix between the plates and insert a comb. Allow the gel to completely polymerize for 20 min. For optimal results, use a comb with teeth that are ≥7 mm wide.
8. Remove the comb and bottom spacer and attach the plates to the electrophoresis tank after filling the lower reservoir with 1× electrophoresis buffer. Fill the upper reservoir of the tank with 1× electrophoresis buffer. With a bent-needle syringe, remove any air bubbles trapped beneath the gel and flush out the wells. Prerun the gel 30 to 60 min at 100 V. For low-ionic-strength buffers (≤0.5×), use a pump with two heads and a flow rate of 5 to 30 ml per min to exchange buffer between the upper and lower reservoirs. Recirculation of the buffer is essential to prevent polarization due to the low buffering capacity of the buffer.
Prepare the binding reactions 9. While the gel is prerunning, assemble the binding reaction by combining the following in a 0.5-ml or 1.5-ml microcentrifuge tube: 5,000 to 20,000 cpm radiolabeled probe DNA (0.1 to 0.5 ng, ≥10 fmol) 0.1 to 2 µg nonspecific carrier DNA 300 µg/ml BSA ≥10% (v/v) glycerol Appropriate buffer and salt Water or buffer to obtain a final reaction volume of 10 to 15 µl DNA-binding protein (∼15 µg crude extract or ∼5 to 25 ng purified protein). The protein should be added last.
10. Mix gently by tapping the bottom of the tube with a finger. Avoid introducing bubbles in the mix.
11. Incubate the binding reaction mix 15 to 30 minutes in a constant-temperature water bath. Optimal incubation temperatures for different proteins can vary from room temperature to 37°C.
Run the gel 12. Load each binding reaction into the appropriate well of the prerun gel using either a 10-µl glass capillary pipet and Clay-Adams screw-top loader, or a pipettor. Load a small volume of 10× loading buffer with dyes into a separate well. The dyes are used to monitor the progress of electrophoresis. There is no stacking gel in this system, so precise loading with little mixing with the gel buffer is necessary to obtain sharp bands on the gel. Allow the sample to fall along one side of the well to prevent dilution and avoid bubbles in the well.
Purification of DNA-Binding Proteins Using Biotin/Streptavidin Affinity Systems
13. Electrophorese at ∼30 to 35 mA for the minimum time required to give good separation of free probe and the protein-DNA complexes. Stop the gel before the bromphenol blue approaches the bottom of the gel (∼1.5 to 2 hr for a 15- to 20-cm gel). Longer run times may cause a weaker signal due to partial dissociation of complexes during electrophoresis.
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Bromphenol blue migrates at approximately the same position as a 70-bp DNA probe. For probes <70 bp, do not run the bromphenol blue to the bottom of the gel. If electrophoresis is performed at room temperature, the glass plates should be allowed to become only slightly warm. Decrease the voltage if the plates become any hotter. To run the gel faster, put the apparatus in a cold room, so that higher voltages may be used without heating the glass plates.
Analyze the gel 14. Remove the glass plates from the gel box and carefully remove the side spacers. 15. Using a spatula, slowly pry the glass plates apart, allowing air to enter between the gel and the glass plate. The gel should remain attached to only one of the plates. Prying the plates apart too quickly may tear the gel or cause it to stick to both plates. If this occurs or if the gel has become distorted, squirt a stream of water underneath it. This will reduce the stickiness of the gel. Be careful not to let the gel slide off the plate.
16. Lay the glass plate with the gel attached on the bench with the gel facing up. Place three sheets of Whatman 3MM filter paper cut to size on top of the gel. 17. Support both sides of the gel sandwich and carefully flip it over so that the filter paper is on the bottom and the glass plate is on the top. 18. Carefully lift up one end of the glass plate. Peel the filter paper with the gel attached to it from the plate. 19. Cover the gel with plastic wrap and dry under vacuum with heat. 20. Autoradiograph the dried filter overnight without an intensifying screen or 2 to 3 hr with an intensifying screen (UNIT 10.11). REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Biotin-cellulose binding buffer 12% glycerol 12 mM HEPES-NaOH, pH 7.9 4 mM Tris⋅Cl, pH 7.9 (APPENDIX 2E) 60 mM KCl 1 mM EDTA 1 mM DTT Store at −20°C upon addition of DTT This is a typical binding buffer. The composition of binding buffer—especially with respect to pH, ionic strength, and the presence or absence of MgCl2—should be determined by those conditions which optimize the binding of the protein of interest to its recognition site.
Biotin-cellulose elution buffer 12% glycerol 20 mM Tris⋅Cl, pH 6.8 (APPENDIX 2E) 1 M KCl 5 mM MgCl2 1 mM EDTA 1 mM DTT 200 µg/ml BSA Store at −20°C upon addition of DTT
continued
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This is a typical elution buffer. The composition of elution buffer—especially with respect to pH, ionic strength, and the presence or absence of MgCl2—should be determined by those conditions that maximize the dissociation rate of the protein from its recognition site. Another carrier protein, such as insulin or hemoglobin, may be substituted for BSA. The small size of insulin can be useful if protein gels will be run to determine the size of the regulatory factor.
Loading buffer, 10× 20% Ficoll 400 0.1 M Na2EDTA, pH 8 (APPENDIX 2E) 1.0% sodium dodecyl sulfate (SDS; APPENDIX 2E) 0.25% bromphenol blue 0.25% xylene cyanol (optional) Xylene cyanol runs ∼50% as fast as bromphenol blue and can interfere with visualization of bands of moderate molecular weight, but can be helpful for monitoring very long runs)
Nondenaturing gel mix For 60 ml: 6.0 ml 10× TAE or TBE elecrophoresis buffer (APPENDIX 2E) or Tris-glycine electrophoresis buffer (UNIT 10.3) 8.1 ml 30% (w/v) acrylamide 3.0 ml 2% (w/v) bisacrylamide 42.9 ml H2O Prepare fresh for each use CAUTION: Acrylamide monomer is neurotoxic. Gloves should be worn when handling the solution, and the solution should not be pipetted by mouth.
COMMENTARY Background Information
Purification of DNA-Binding Proteins Using Biotin/Streptavidin Affinity Systems
The biotin/streptavidin purification method works well because the interaction between biotin and avidin (or avidin-like proteins) is one of the strongest known noncovalent interactions. The dissociation constant for the streptavidin-biotin complex is ∼10−15 M. Avidin, from egg white, and streptavidin, from Streptomyces avidinii, are tetrameric proteins containing four high-affinity binding sites for the vitamin biotin. Since streptavidin is multivalent, it is able to serve as a bridge between the biotinylated DNA fragment and the biotin-containing resin. The strong interaction is extremely useful for purification of DNA-binding proteins, because DNA-affinity columns with streptavidin/biotin bridges can be washed under a wide variety of conditions (i.e., 2 M KCl and 1% SDS) without removing either the streptavidin or the biotinylated DNA fragment from the matrix. Two properties of streptavidin make it more suitable than avidin for use in DNA-affinity purification. The first is that streptavidin, unlike avidin, is not a glycoprotein, and the second is that streptavidin is slightly acidic whereas avidin is basic. Therefore, streptavidin is less
likely to bind nonspecifically to cellular glycoproteins and to acidically charged cell components such as nucleic acids. Several factors make this method simple, rapid, and effective. The same binding conditions and DNA fragment used in the mobilityshift DNA-binding assay to identify a protein (see Support Protocol; Buratowski and Chodosh, 1996) can be used to effect its purification. In addition, the binding of the protein to its DNA recognition site in solution is more efficient than protein-DNA interactions that take place on a column matrix (discussed below). Most importantly, binding in solution allows each reaction parameter to be optimized on an analytical scale by using the gel binding assay. As an analytical technique, biotin/streptavidin DNA-affinity purification permits the direct identification of a wide variety of sequence-specific DNA-binding proteins. It has already been successfully used to identify hormone receptors, components in mRNA splicing complexes, and RNA polymerase II and III transcription factors (Haeuptle et al., 1983; Chodosh et al., 1986; Grabowski and Sharp, 1986; Kasher et al., 1986).
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Another method is commonly used for purifying DNA-binding proteins, whereby DNA-protein binding interactions occur on a column matrix. In this technique, catenated DNA-binding sites are covalently coupled with cyanogen bromide to Sepharose CL-2B (Kadonaga and Tjian, 1986; Rosenfeld and Kelly, 1986). A partially purified protein fraction is combined with competitor DNA and passed through a DNA-Sepharose resin, binding the protein to the surface of the matrix. Generally, multiple passes through the column are used to effect purification, with 25- to 50fold purification achieved with each column passage. The biotin/streptavidin technique, on the other hand, typically yields 50- to 250-fold purification.
Critical Parameters Only one end of the DNA fragment should be labeled with biotin. If both ends are labeled, the biotinylated fragment may bind to the resin in such a fashion as to sterically displace the protein bound to the DNA fragment. Thus, one of the restriction enzymes used must be chosen to permit the incorporation of biotin-11-dUTP in place of TTP. Furthermore, since only one end of the fragment is biotin-labeled, the restriction enzyme generating the nonbiotinylated end should leave no overhang, a 3′ overhang, or a 5′ overhang that will not incorporate biotin-dUTP. For binding sites cloned into the pUC polylinker, an EcoRI and HincII fragment fulfills these requirements. Biotin-11-dUTP may be incorporated into the EcoRI overhang along with a low-specific-activity dATP radioactive label. The blunt HincII end of the fragment remains unlabeled. Alternately, if both ends have 5′ overhangs which permit incorporation of biotin-dUTP, the plasmid may first be digested with one of the enzymes and end-labeled with biotin-dUTP and a radioactive dNTP using the Klenow fragment. After heat-inactivating the Klenow fragment, the plasmid may then be digested with the second restriction endonuclease. Use of annealed complimentary oligonucleotides with the desired ends obviates the need to find appropriate restriction sites. The low-specific-activity label incorporated into the biotinylated DNA fragment is used to help monitor both the binding of the protein to the biotinylated fragment and the binding of the biotinylated fragment to the biotin-cellulose resin via a streptavidin bridge. The amounts of the radioactive dNTP and the nonradioactive dNTP added to the Klenow reaction should be
in a ratio such that 1% to 2% of the biotinylated fragments are labeled. For example, 100 µCi of 6000 Ci/mmol dATP (0.2 µM) may be added to a reaction containing 20 µM unlabeled dATP. If the radioactive dNTP and the biotin-dUTP residue are incorporated on the same end of the DNA fragment, as in an EcoRI site, decay of the radioactive nuclide will result in formation of an unlabeled, nonbiotinylated probe that binds to the protein but is incapable of binding to streptavidin. This molecule is a competitive inhibitor during protein purification and can be kept to a minimum by labeling to a lower specific activity. Over time, streptavidin tetramers dissociate into monomers. The integrity of the tetramer may be monitored by performing native (nondenaturing) gel electrophoresis. Since streptavidin monomers will bind to the biotinylated DNA fragment, but will be unable to bind to the biotin-cellulose resin, the monomeric form of streptavidin will inhibit purification. Most proteins can be eluted from their binding site with 1 M KCl. Since the added elution buffer is diluted by the binding buffer in the biotin-cellulose pellet, the actual salt concentration in the resuspended pellet will be lower than that of the elution buffer. As a result, it is useful to remove a small aliquot, dilute it, and measure the actual salt concentration with a conductivity meter. Otherwise, the time, temperature, and composition of the buffer used in the elution step should be chosen to maximize the protein’s dissociation rate from its binding site. Thus, if a protein’s dissociation rate is 15 sec in 1 M KCl at room temperature, and 3 hr in 1 M KCl at 4°C, a 5-min incubation in 1 M KCl at room temperature should be sufficient to elute the protein. Dissociation rates can easily be measured using a mobility shift DNAbinding assay described in this chapter (Support Protocol; Buratowski and Chodosh, 1996).
Troubleshooting A major advantage of the biotin/streptavidin method is the ability to monitor the purification on an analytical scale using a mobility-shift assay. Because different parameters can be independently examined, it is possible to pinpoint the particular step that is not working. Several parameters can be examined routinely. Using the biotinylated radiolabeled fragment, the specific binding of the DNA fragment to its protein can be directly determined. In addition, the binding conditions can be optimized in a binding reaction containing the DNA probe and protein. The probe can be a biotinylated or a
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Purification of DNA-Binding Proteins Using Biotin/Streptavidin Affinity Systems
nonbiotinylated radiolabeled DNA fragment. The complex is then resolved by native (nondenaturing) gel electrophoresis. Streptavidin binding to the probe can be monitored and titrated by varying the amount of streptavidin in a binding reaction. The resulting streptavidin-DNA complex is resolved from free DNA by native gel electrophoresis. Binding of this complex to the biotin-cellulose matrix can be directly monitored. First, biotin-cellulose is incubated with the complex. Next the resin is spun and aliquots of the supernatant are assayed for the presence of the radiolabeled fragment on a polyacrylamide gel. This basic procedure allows several relevant parameters to be examined, including the amount of streptavidin required for complete binding of the fragment to the resin, the amount of biotin-cellulose required to bind a given amount of fragment, the binding time, and the optimal binding conditions. These three final steps can also be monitored for the specific adsorption of the protein to the column. By assaying the amount of binding activity in the supernatant before and after addition of biotin cellulose, the adsorption efficiency can be determined. Nonspecific resin binding can be examined by assaying the binding activity in a supernatant containing the protein, the DNA carrier poly(dI-dC)⋅poly(dI-dC), streptavidin, and biotin-cellulose, but lacking the biotinylated DNA fragment with the protein’s binding site. Alternatively, a nonspecific biotinylated DNA fragment can be added. Interestingly, nonspecific protein binding to the resin is minimized with a lower poly(dI-dC)⋅poly(dI-dC) concentration than is required to generate a discrete band in the mobility shift assay. The conditions for protein elution from the column can also be directly examined and optimized. After protein adsorption onto the matrix, elution parameters such as time, temperature, pH, carrier protein, and ionic strength can be varied. The amount of protein released into the supernatant is then assayed by the mobility shift assay. In optimizing this step, it is very helpful to know as much as possible about the variables affecting the dissociation rate of the protein from its DNA binding site. These can be studied quickly and rigorously with the mobility shift assay. Troubleshooting guidelines for this assay are presented in Buratowski and Chodosh (1996).
Anticipated Results The strong interaction between biotin and avidin can result in 25 nmol of biotinylated DNA fragment binding to 1 ml of biotin-cellulose resin. This corresponds to 1.5 mg of a 60 kDa polypeptide. After parameter optimization, ∼50- to 250-fold purification of the DNAprotein complex can be obtained. Once the protein has been adsorbed to the column >99% of the remaining protein can be removed with repeated column washing using the binding buffer. Moreover, ∼30% to 80% of the binding activity can be recovered from the column. The recovery efficiency depends on how effectively nonspecific binding sites have been blocked. This purification scheme may be used either with crude extracts or with partially purified protein fractions.
Time Considerations Starting with a biotinylated DNA fragment, the batch protocol (Basic Protocol) can be performed in ∼1.5 hr, the microcolumn (Alternate Protocol 1) in ∼2.5 hr, and the streptavidinagarose (Alternate Protocol 2) protocol in ∼1.5 hr. The DNA-binding assay (Support Protocol) may take 4 to 5 hr to complete from the time the gel is poured to the time it is dried onto filter paper. Only a small fraction of this time is labor intensive. Thus, several DNA-binding gels can be run simultaneously while other experiments are being performed. Furthermore, the results are often available within 2 to 3 hr after completion of the experiment.
Literature Cited Ausubel, F.A., Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G., Smith, J.A., and Struhl, K. (eds.) 1998. Current Protocols in Molecular Biology. John Wiley & Sons, New York. Buratowski, S. and Chodosh, L. 1996. Mobilityshift DNA-binding assay using gel electrophoresis. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 12.2.1-12.2.11. John Wiley & Sons, New York. Chodosh, L.A., Carthew, R.W., and Sharp, P.A. 1986. A single polypeptide possesses the binding and transcription activities of the adenovirus major late promoter. Mol. Cell. Biol. 6:4723-4733. Grabowski, P.J. and Sharp, P.A. 1986. Affinity chromatography of splicing complexes: U2, U5, and U4 + U6 small nuclear ribonucleoprotein particles in the spliceosome. Science 233:1294-1299.
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Haeuptle, M.-T., Aubert, M.L., Kjiane, J., and Kraehenbuhl, J.-P. 1983. Binding sites for lactogenic and somatogenic hormones from rabbit mammary gland and liver. J. Biol. Chem. 258:305314. Kadonaga, J.T. and Tjian, R. 1986. Affinity purification of sequence-specific DNA binding proteins. Proc. Natl. Acad. Sci. U.S.A. 83:58895893. Kasher, M.S., Pintel, D., and Ward, D.C. 1986. Rapid enrichment of HeLa transcription factors IIIB and IIIC by using affinity chromatography based on avidin-biotin interactions. Mol. Cell. Biol. 6:3117-3127. Rosenfeld, P.J. and Kelly, T.J. 1986. Purification of nuclear factor I by DNA recognition site affinity chromatography. J. Biol. Chem. 261:1398-1408. Sambrook, J., Fritsch, E.F., and Maniatis, T. 1989. Molecular Cloning: A Laboratory Manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
Key References Chodosh et al., 1986. See above. Describes the biotin/streptavidin-agarose microcolumn procedure from which the second alternate protocol of this unit was drawn. Kasher et al., 1986. See above. Describes a biotin-cellulose/streptavidin procedure similar to the basic protocol of this unit.
Contributed by Lewis A. Chodosh University of Pennsylvania Philadelphia, Pennsylvania Stephen Buratowski Harvard Medical School Boston, Massachusetts
Affinity Purification
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Immunoprecipitation
UNIT 9.8
Immunoprecipitation is a technique in which an antigen is isolated by binding to a specific antibody attached to a sedimentable matrix. The source of antigen for immunoprecipitation can be unlabeled cells or tissues, metabolically or extrinsically labeled cells (UNIT 3.7), subcellular fractions from either unlabeled or labeled cells (see Chapter 4), or in vitro–translated proteins. Immunoprecipitation is also used to analyze protein fractions separated by other biochemical techniques such as gel filtration (UNIT 8.3) or sedimentation on density gradients (UNIT 4.2). Either polyclonal or monoclonal antibodies from various animal species can be used in immunoprecipitation protocols. Antibodies can be bound noncovalently to immunoadsorbents such as protein A– or protein G–agarose, or can be coupled covalently to a solid-phase matrix. Immunoprecipitation protocols consist of several stages (Fig. 9.8.1; see Basic Protocol 1). In stage 1, the antigen is solubilized by one of several techniques for lysing cells. Soluble and membrane-associated antigens can be released from cells grown either in suspension culture (see Basic Protocol 1) or as a monolayer on tissue culture dishes (see Alternate Protocol 1) with nondenaturing detergents. Alternatively, cells can be lysed under denaturing conditions (see Alternate Protocol 2). Soluble antigens can also be extracted by mechanical disruption of cells in the absence of detergents (see Alternate Protocol 3). All of these procedures are suitable for extracting antigens from animal cells. By contrast, yeast cells require disruption of their cell wall in order to allow extraction of the antigens (see Alternate Protocol 4). In stage 2, a specific antibody is attached, either noncovalently or covalently, to a sedimentable, solid-phase matrix to allow separation by low-speed centrifugation. In this unit, two methods for achieving this are described: the noncovalent attachment of antibody to protein A– or protein G–agarose beads (see Basic Protocol 1) and covalent coupling to Sepharose (see Alternate Protocol 5 and Support Protocol). Stage 3 is the actual immunoprecipitation, which can be achieved by incubating the solubilized antigen from stage 1 with the immobilized antibody from stage 2, followed by extensive washing to remove unbound proteins (see Basic Protocol 1). Another method is to precipitate the immune complexes using antibodies contained in an anti-immunoglobulin (anti-Ig) serum (see Alternate Protocol 6). Immunoprecipitated antigens can be dissociated from antibodies and reprecipitated by a protocol referred to as “immunoprecipitation-recapture” (see Basic Protocol 2). This procedure can be used with the same antibody for further purification of the antigen, or with a second antibody to identify components of multisubunit complexes or to study protein-protein interactions (Fig. 9.8.3). Immunoprecipitated antigens can be analyzed by one-dimensional electrophoresis (UNIT 10.1), two-dimensional electrophoresis (UNIT 10.4), or immunoblotting (UNIT 10.10). In some cases, immunoprecipitates can be used for structural or functional analyses of the isolated antigens. Immunoprecipitates can also be used as sources of immunogens for production of monoclonal or polyclonal antibodies. IMMUNOPRECIPITATION USING CELLS IN SUSPENSION LYSED WITH A NONDENATURING DETERGENT SOLUTION
BASIC PROTOCOL 1
In this protocol, cells in suspension (labeled or unlabeled) are extracted by incubation in nondenaturing lysis buffer containing the nonionic detergent Triton X-100 (steps 1 to 7). This procedure results in the release of both soluble and membrane proteins; however, many cytoskeletal and nuclear proteins, as well as a fraction of membrane proteins, are Affinity Purification Contributed by Juan S. Bonifacino, Esteban C. Dell’Angelica, and Timothy A. Springer Current Protocols in Protein Science (1999) 9.8.1-9.8.28 Copyright © 1999 by John Wiley & Sons, Inc.
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not efficiently extracted under these conditions. The procedure allows immunoprecipitation with antibodies to epitopes that are exposed in native proteins. For immunoprecipitation, a specific antibody is immobilized on a sedimentable, solidphase matrix (steps 8 to 14). Although there are many ways to attach antibodies to matrices (see Commentary), the most commonly used methods rely on the property of immunoglobulins to bind Staphylococcus aureus protein A, or protein G from group G Streptococcus (Table 9.8.1). The best results are obtained by binding antibodies to protein A or protein G that is covalently coupled to agarose beads. In this protocol, Sepharose beads are used (Sepharose is a more stable, cross-linked form of agarose). Immunoprecipitation is most often carried out using rabbit polyclonal or mouse monoclonal antibodies, which,
protein A–agarose bead
animal cell specific antigen
antibody
1
cell lysis (see Basic Protocol 1, steps 1 to 7; Alternate Protocols 1 to 4)
2
antibody binding to protein A–agarose bead (see Basic Protocol 1, steps 8 to 14)
3 antigen isolation on antibody bead (see Basic Protocol 1, steps 18 to 21)
washing (see Basic Protocol 1, steps 22 to 26) and analysis (step 27)
Immunoprecipitation
Figure 9.8.1 Schematic representation of the stages of the immunoprecipitation protocol presented in Basic Protocol 1. (1) Cell lysis: antigens are solubilized by extraction of the cells in the presence or absence of detergents. To increase specificity, the cell lysate can be precleared with protein A–agarose beads (steps 15 to 17, not shown). (2) Antibody immobilization: a specific antibody is bound to protein A–agarose beads. (3) Antigen capture: the solubilized antigen is isolated on antibody-conjugated beads.
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with some exceptions (e.g., mouse IgG1), bind well to protein A (Table 9.8.1). Antibodies that do not bind to protein A–agarose can be adsorbed to protein G–agarose (Table 9.8.1) using exactly the same protocol. For optimal time management, incubation of antibodies with protein A–agarose can be carried out either before or during lysis of the cells. The final stage in immunoprecipitation is combining the cell lysate with the antibodyconjugated beads and isolating the antigen (steps 18 to 26). This can be preceded by an optional preclearing step in which the lysate is absorbed with either “empty” protein A–agarose beads or with an irrelevant antibody bound to protein A–agarose (steps 15 to 17). The need for preclearing depends on the specific experimental system being studied and the quality of the antibody reagents. The protocol described below incorporates a preclearing step using protein A–agarose. Protein fractions separated by techniques such as gel filtration (UNIT 8.3) or sedimentation on sucrose gradients (UNIT 4.2) can be used in Table 9.8.1 Ga,b,c
Binding of Antibodies to Protein A and Protein
Protein A binding
Protein G bindingd
Monoclonal antibodiese Human IgG1 Human IgG2 Human IgG3 Human IgG4 Mouse IgG1 Mouse IgG2a Mouse IgG2b Mouse IgG3 Rat IgG1 Rat IgG2a Rat IgG2b Rat IgG2c
++ ++ − ++ + ++ ++ ++ + − − ++
++ ++ ++ ++ ++ ++ ++ ++ + ++ ++ ++
Polyclonal antibodies Chicken Donkey Goat Guinea pig Hamster Human Monkey Mouse Rabbit Rat Sheep
− − + ++ + ++ ++ ++ ++ + +
− ++ ++ + ++ ++ ++ ++ ++ + ++
Antibody
a++, moderate to strong binding; +, weak binding; −, no binding. bA hybrid protein A/G molecule that combines the features of protein A and
protein G, coupled to a solid-phase matrix, is available from Pierce. cInformation from Harlow and Lane (1999), and from Amersham Pharmacia
Biotech, Pierce, and Jackson Immunoresearch. dNative protein G binds albumin from several animal species. Recombinant variants of protein G have been engineered for better binding to rat, mouse, and guinea pig IgG, as well as for avoiding binding to serum albumin. eProtein A binds some IgM, IgA, and IgE antibodies in addition to IgG, whereas protein G binds only IgG.
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place of the cell lysate at this stage. After binding the antigen to the antibody-conjugated beads, the unbound proteins are removed by successive washing and sedimentation steps. Materials Unlabeled or labeled cells in suspension PBS (APPENDIX 2E), ice cold Nondenaturing lysis buffer (see recipe), ice cold 50% (v/v) protein A–Sepharose bead (Sigma, Amersham Pharmacia Biotech) slurry in PBS containing 0.1% (w/v) BSA and 0.01% (w/v) sodium azide (NaN3) Specific polyclonal antibody (antiserum or affinity-purified immunoglobulin) or monoclonal antibody (ascites, culture supernatant, or purified immunoglobulin) Control antibody of same type as specific antibody (e.g., preimmune serum or purified irrelevant immunoglobulin for specific polyclonal antibody; irrelevant ascites, hybridoma culture supernatant, or purified immunoglobulin for specific monoclonal antibody; see Critical Parameters) 10% (w/v) BSA Wash buffer (see recipe), ice cold Microcentrifuge with fixed-angle rotor (Eppendorf 5415C or equivalent) Tube rotator (capable of end-over-end inversion) Pasteur pipet attached to a vacuum trap CAUTION: When working with radioactivity, take appropriate precautions to avoid contamination of the experimenter and the surroundings. Carry out the experiment and dispose of wastes in an appropriately designated area, following the guidelines provided by the local radiation safety officer (also see APPENDIX 2B). NOTE: All solutions should be ice cold and procedures should be carried out at 4°C or on ice. Prepare cell lysate 1. Collect cells in suspension by centrifuging 5 min at 400 × g, 4°C, in a 15- or 50-ml capped conical tube. Place tube on ice. Approximately 0.5–2 × 107 cells are required to yield 1 ml lysate, which is the amount generally used for each immunoprecipitation. Labeled cells are likely to have been pelleted earlier as part of the labeling procedure. If the cells are frozen, they should be thawed on ice before solubilization.
2. Aspirate supernatant with a Pasteur pipet attached to a vacuum trap. CAUTION: Dispose of radioactive materials following applicable safety regulations (APPENDIX 2B).
3. Resuspend cells gently by tapping the bottom of the tube. Rinse cells twice with ice-cold PBS as in steps 1 and 2, using the same volume of PBS as in the initial culture. 4. Add 1 ml ice-cold nondenaturing lysis buffer per ∼0.5–2 × 107 cells and resuspend pellet by gentle agitation for 3 sec with a vortex mixer set at medium speed. Do not shake vigorously, as this could result in loss of material or protein denaturation due to foaming.
Immunoprecipitation
5. Keep suspension on ice 15 to 30 min and transfer to a 1.5-ml conical microcentrifuge tube.
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Tubes can have flip-top or screw caps. Screw-capped tubes are preferred because they are less likely to open accidentally during subsequent procedures. They are also recommended for work with radioactivity.
6. Clear the lysate by microcentrifuging 15 min at 16,000 × g (maximum speed), 4°C. Centrifugation can be carried out in a microcentrifuge placed in a cold room or in a refrigerated microcentrifuge. Take precautions to ensure that the 4°C temperature is maintained during the spin (e.g., use a fixed-angle rotor with a lid, as the aerodynamics of this type of rotor reduces generation of heat by friction). If it is necessary to reduce background, the lysate can be spun for 1 hr at 100,000 × g in an ultracentrifuge.
7. Transfer the supernatant to a fresh microcentrifuge tube using an adjustable pipet fitted with a disposable tip. Do not disturb the pellet, and leave the last 20 to 40 µl of supernatant in the centrifuge tube. Keep the cleared lysate on ice until preclearing (step 15) or addition of antibody beads (step 18). NOTE: Resuspension of even a small amount of sedimented material will result in high nonspecific background due to carryover into the immunoprecipitation steps. A cloudy layer of lipids floating on top of the supernatant will not adversely affect the results of the immunoprecipitation. When the lysate is highly radioactive—as is the case for metabolically labeled cells—the use of tips with aerosol barriers is recommended to reduce the risk of contaminating internal components of the pipet. Cell extracts can be frozen at −70°C until used for immunoprecipitation. However, it is preferable to lyse the cells immediately before immunoprecipitation in order to avoid protein degradation or dissociation of protein complexes. If possible, freeze the cell pellet from step 3 rather than the supernatant from step 7.
Prepare antibody-conjugated beads 8. In a 1.5-ml conical microcentrifuge tube, combine 30 µl of 50% protein A–Sepharose bead slurry, 0.5 ml ice-cold PBS, and the following quantity of specific antibody (select one): 1 to 5 µl polyclonal antiserum 1 µg affinity-purified polyclonal antibody 0.2 to 1 µl ascitic fluid containing monoclonal antibody 1 µg purified monoclonal antibody 20 to 100 µl culture supernatant containing monoclonal antibody. The quantities of antibody suggested are rough estimates based on the expected amount of specific antibodies in each preparation. Quantities can be increased or decreased depending on the quality of the antibody preparation (see Commentary). Substitute protein G for protein A if antibodies are of a species or subclass that does not bind to protein A (see Table 9.8.1). If the same antibody will be used to immunoprecipitate multiple samples (e.g., samples from a pulse-chase experiment; UNIT 3.7), the quantities indicated above can be increased proportionally to the number of samples and incubated in a 15-ml capped conical tube. In this case, the beads should be divided into aliquots just prior to the addition of the cleared cell lysate (step 18). Antibody-conjugated beads can be prepared prior to preparation of the cell lysate (steps 1 to 7), in order to minimize the time that the cell extract is kept on ice.
9. Set up a nonspecific immunoprecipitation control in a 1.5-ml conical microcentrifuge tube by incubating 30 µl of 50% protein A–Sepharose bead slurry, 0.5 ml ice-cold PBS, and the appropriate control antibody (select one):
Affinity Purification
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1 to 5 µl preimmune serum as a control for a polyclonal antiserum 1 µg purified irrelevant polyclonal antibody (an antibody to an epitope that is not present in the cell lysate) as a control for a purified polyclonal antibody 0.2 to 1 µl ascitic fluid containing irrelevant monoclonal antibody (an antibody to an epitope that is not present in the cell lysate and of the same species and immunoglobulin subclass as the specific antibody) as a control for an ascitic fluid containing specific monoclonal antibody 1 µg purified irrelevant monoclonal antibody as a control for a purified monoclonal antibody 20 to 100 µl hybridoma culture supernatant containing irrelevant monoclonal antibody as a control for a hybridoma culture supernatant containing specific monoclonal antibody. The amount of irrelevant antibody should match that of the specific antibody and the antibody should be from the same species as the specific antibody.
10. Mix suspensions thoroughly. Tumble incubation mixtures end over end ≥1 hr at 4°C in a tube rotator. Addition of 0.01% (w/v) Triton X-100 may facilitate mixing of the suspension during tumbling. Incubations can be carried out for as long as 24 hr. This allows preparation of the antibody-conjugated beads prior to immunoprecipitation.
11. Microcentrifuge 2 sec at 16,000 × g (maximum speed), 4°C. 12. Aspirate the supernatant (containing unbound antibodies) using a fine-tipped Pasteur pipet connected to a vacuum aspirator. 13. Add 1 ml nondenaturing lysis buffer and resuspend the beads by inverting the tube three or four times. For lysates prepared with detergents (this protocol and see Alternate Protocols 1 and 2), use 1 ml nondenaturing lysis buffer; for lysates prepared by mechanical disruption (see Alternate Protocol 3), use detergent-free lysis buffer (see recipe). Use of a repeat pipettor is recommended when processing multiple samples.
14. Wash by repeating steps 11 to 13, and then steps 11 and 12 once more. At this point the beads have been washed twice with lysis buffer and are ready to be used for immunoprecipitation. Antibody-bound beads can be stored up to 6 hr at 4°C until used.
Preclear lysate (optional) 15. In a microcentrifuge tube, combine 1 ml cell lysate (from step 7) and 30 µl of 50% protein A–Sepharose bead slurry. The purpose of this step is to remove from the lysate proteins that bind to protein A–Sepharose, as well as pieces of insoluble material that may have been carried over from previous steps. If the lysate was prepared from cells expressing immunoglobulins—such as spleen cells or cultured B cells—the preclearing step should be repeated at least three times to ensure complete removal of endogenous immunoglobulins. If cell lysates were frozen and thawed, they should be microcentrifuged 15 min at 16,000 × g (maximum speed), 4°C, before the preclearing step.
16. Tumble end over end 30 min at 4°C in a tube rotator. Immunoprecipitation
17. Microcentrifuge 5 min at 16,000 × g (maximum speed), 4°C.
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Immunoprecipitate 18. Add 10 µl of 10% BSA to the tube containing specific antibody bound to protein A–Sepharose beads (step 14), and transfer to this tube the entire volume of cleared lysate (from step 7 or 17). If a nonspecific immunoprecipitation control is performed, divide lysate in two ∼0.4-ml aliquots, one for the specific antibody and the other for the nonspecific control. In order to avoid carryover of beads with precleared material, leave 20 to 40 µl of supernatant on top of the pellets in the preclearing tubes. Discard beads and remaining supernatant. The BSA quenches nonspecific binding to the antibody-conjugated beads during incubation with the cell lysate.
19. Incubate 1 to 2 hr at 4°C while mixing end over end in a tube rotator. Samples can be incubated overnight, although there is an increased risk of protein degradation, dissociation of multiprotein complexes, or formation of protein aggregates.
20. Microcentrifuge 5 sec at 16,000 × g (maximum speed), 4°C. 21. Aspirate the supernatant (containing unbound proteins) using a fine-tipped Pasteur pipet connected to a vacuum aspirator. The supernatant can be kept up to 8 hr at 4°C or up to 1 month at −70°C for sequential immunoprecipitation of other antigens or for analysis of total proteins. To reutilize lysate, remove the supernatant carefully with an adjustable pipet fitted with a disposable tip. Before reprecipitation, preabsorb the lysate with protein A–Sepharose (as in steps 15 to 17) to remove antibodies that may have dissociated during the first immunoprecipitation. CAUTION: Dispose of radioactive materials following applicable safety regulations.
22. Add 1 ml ice-cold wash buffer, cap the tubes, and resuspend the beads by inverting the tube 3 or 4 times. Use of a repeat pipettor is recommended when processing multiple samples.
23. Microcentrifuge 2 sec at 16,000 × g (maximum speed), 4°C. 24. Aspirate the supernatant, leaving ∼20 µl supernatant on top of the beads. 25. Wash beads three more times (steps 22 to 24). Total wash time (steps 22 to 26) should be ∼30 min, keeping the samples on ice for 3 to 5 min between washes if necessary (see Critical Parameters).
26. Wash beads once more using 1 ml ice-cold PBS and aspirate supernatant completely with a drawn-out Pasteur pipet or an adjustable pipet fitted with a disposable tip. The final product should be 15 µl of settled beads containing bound antigen. Immunoprecipitates can either be processed immediately or frozen at −20°C for later analysis. For subsequent analysis of the isolated proteins prior to electrophoresis (e.g., comparison of the electrophoretic mobility of the antigen with or without treatment with glycosidases), samples can be divided into two or more aliquots after addition of PBS. Transfer aliquots of the bead suspension to fresh tubes, centrifuge and aspirate as in the previous steps.
27. Analyze immunoprecipitates by one-dimensional electrophoresis (UNIT 10.1), two-dimensional electrophoresis (UNIT 10.4), or immunoblotting (UNIT 10.10).
Affinity Purification
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ALTERNATE PROTOCOL 1
IMMUNOPRECIPITATION USING ADHERENT CELLS LYSED WITH A NONDENATURING DETERGENT SOLUTION Immunoprecipitation using adherent cells can be performed in the same manner as with nonadherent cells (see Basic Protocol 1). This protocol is essentially similar to steps 1 to 5 of Basic Protocol 1, but describes modifications necessary for using the same nondenaturing detergent solution to lyse cells attached to tissue culture plates. It is preferable to use cells grown on plates rather than in flasks, because the cell monolayer is more easily accessible. Additional Materials (also see Basic Protocol 1) Unlabeled or labeled cells grown as a monolayer on a tissue culture plate (UNIT 3.7) Rubber policeman NOTE: All solutions should be ice cold and procedures should be carried out at 4°C or on ice. 1. Rinse cells attached to a tissue culture plate twice with ice-cold PBS. Remove the PBS by aspiration with a Pasteur pipet attached to a vacuum trap. CAUTION: Dispose of radioactive materials following applicable safety regulations.
2. Place the tissue culture plate on ice. 3. Add ice-cold nondenaturing lysis buffer to the tissue culture plate. Use 1 ml lysis buffer for an 80% to 90% confluent 100-mm-diameter tissue culture plate. Depending on the cell type, a confluent 100-mm dish will contain 0.5–2 × 107 cells. For other plate sizes, adjust volume of lysis buffer according to the surface area of the plate.
4. Scrape the cells off the plate with a rubber policeman, and transfer the suspension to a 1.5-ml conical microcentrifuge tube using an adjustable pipettor fitted with a disposable tip. Vortex gently for 3 sec and keep tubes on ice for 15 to 30 min. Tubes can have flip-top or screw caps. Screw-capped tubes are preferred because they are less likely to open accidentally during subsequent procedures. They are also recommended for work with radioactivity.
5. Clear the lysate and perform immunoprecipitation (see Basic Protocol 1, steps 6 to 27). ALTERNATE PROTOCOL 2
IMMUNOPRECIPITATION USING CELLS LYSED WITH DETERGENT UNDER DENATURING CONDITIONS If epitopes of native proteins are not accessible to antibodies, or if the antigen cannot be extracted from the cell with nonionic detergents, cells should be solubilized under denaturing conditions. This protocol is based on that for nondenaturing conditions (see Basic Protocol 1, steps 1 to 7), with the following modifications. Denaturation is achieved by heating the cells in a denaturing lysis buffer that contains an ionic detergent such as SDS or Sarkosyl (N-lauroylsarcosine). The denaturing lysis buffer also contains DNase I to digest DNA released from the nucleus. Prior to immunoprecipitation, the denatured protein extract is diluted 10-fold with nondenaturing lysis buffer, which contains Triton X-100; this step protects the antigen-antibody interaction from interference by the ionic detergent. Immunoprecipitation is performed as described (see Basic Protocol 1).
Immunoprecipitation
The following protocol is described for cells in suspension culture, although it can be adapted for adherent cells (see Alternate Protocol 1). Only antibodies that react with denatured proteins can be used to immunoprecipitate proteins solubilized by this protocol.
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Additional Materials (also see Basic Protocol 1) Denaturing lysis buffer (see recipe) Heating block set at 95°C (Eppendorf Thermomixer 5436 or equivalent) 25-G needle attached to 1-ml syringe 1. Collect cells in suspension culture (see Basic Protocol 1, steps 1 to 3). Place tubes on ice. 2. Add 100 µl denaturing lysis buffer per ∼0.5–2 × 107 cells in the pellet. 3. Resuspend the cells by vortexing vigorously 2 to 3 sec at maximum speed. Transfer suspension to a 1.5-ml conical microcentrifuge tube. The suspension may be very viscous due to release of nuclear DNA. Tubes can have flip-top or screw caps. Screw-capped tubes are preferred because they are less likely to open accidentally during subsequent procedures. They are also recommended for work with radioactivity.
4. Heat samples 5 min at 95°C in a heating block. 5. Dilute the suspension with 0.9 ml nondenaturing lysis buffer. Mix gently. The excess 1% Triton X-100 in the nondenaturing lysis buffer sequesters SDS into Triton X-100 micelles.
6. Shear DNA by passing the suspension five to ten times through a 25-G needle attached to a 1-ml syringe. If the DNA is not digested by DNase I in the denaturing lysis buffer or thoroughly sheared mechanically, it will interfere with the separation of pellet and supernatant after centrifugation. Repeat mechanical disruption until the viscosity is reduced to manageable levels.
7. Incubate 5 min on ice. 8. Clear the lysate and perform immunoprecipitation (see Basic Protocol 1, steps 6 to 27). IMMUNOPRECIPITATION USING CELLS LYSED WITHOUT DETERGENT Immunoprecipitation of proteins that are already soluble within cells (e.g., cytosolic or luminal organellar proteins) may not require the use of detergents. Instead, cells can be mechanically disrupted by repeated passage through a needle, and soluble proteins can be separated from insoluble material by centrifugation. The following protocol describes lysis of cells in a PBS-based detergent-free lysis buffer. Other buffer formulations may be used for specific proteins.
ALTERNATE PROTOCOL 3
Additional Materials (also see Basic Protocol 1) Detergent-free lysis buffer (see recipe) 25-G needle attached to 3-ml syringe NOTE: All solutions should be ice-cold and procedures should be carried out at 4°C or on ice. 1. Collect and wash cells in suspension (see Basic Protocol 1, steps 1 to 3). 2. Add 1 ml of ice-cold detergent-free lysis buffer per ∼0.5–2 × 107 cells in a pellet. 3. Resuspend the cells by gentle agitation for 3 sec with a vortex mixer set at medium speed.
Affinity Purification
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4. Break cells by passing the suspension 15 to 20 times through a 25-G needle attached to a 3-ml syringe. Extrusion of the cell suspension from the syringe should be rapid, although care should be exercised to prevent splashing and excessive foaming. Cell breakage can be checked under a bright-field or phase-contrast microscope. Repeat procedure until >90% cells are broken. It is helpful to check ahead of time whether the cells can be broken in this way. If the cells are particularly resistant to mechanical breakage, they can be swollen for 10 min at 4°C with a hypotonic solution containing 10 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) before mechanical disruption.
5. Clear the lysate and perform immunoprecipitation (Basic Protocol 1, steps 6 to 27). ALTERNATE PROTOCOL 4
IMMUNOPRECIPITATION USING YEAST CELLS DISRUPTED WITH GLASS BEADS Unlike animal cells, yeast cells have an extremely resistant, detergent-insoluble cell wall. To allow extraction of cellular antigens, the cell wall needs to be broken by mechanical, enzymatic, or chemical means. The most commonly used procedure consists of vigorous vortexing of the yeast suspension with glass beads. The breakage can be done in the presence or absence of detergent, as previously described for animal cells (see Basic Protocol 1, Alternate Protocol 2, and Alternate Protocol 3). The protocol described below is suitable for mechanical disruption of most yeast species, including Saccharomyces cerevisiae and Schizosaccharomyces pombe. A protocol for metabolic labeling for yeast has been described by Franzusoff et al. (1991). Additional Materials (also see Basic Protocol 1) Unlabeled or radiolabeled yeast cells Lysis buffer, ice cold: nondenaturing, denaturing, or detergent-free lysis buffer (see recipes) Glass beads (acid-washed, 425- to 600-µm diameter; Sigma) NOTE: All solutions should be ice-cold and procedures should be carried out at 4°C or on ice. 1. Collect 10 ml of yeast culture at 1 OD600 per immunoprecipitation sample, and centrifuge 5 min at 4000 × g, 4°C. Place tube on ice. 2. Remove supernatant by aspiration with a Pasteur pipet attached to a vacuum trap. CAUTION: Dispose of radioactive materials following applicable safety regulations.
3. Loosen pellet by vortexing vigorously for 10 sec. Rinse cells twice with ice-cold distilled water as in steps 1 and 2. Radiolabeled yeast cells are likely to have been pelleted earlier as part of the labeling procedure. If the pellets are frozen, they should be thawed on ice prior to cell disruption.
4. Add 3 vol ice-cold lysis buffer and 3 vol glass beads per volume of pelleted yeast cells. Use nondenaturing lysis buffer or detergent-free lysis buffer as required for the antigen under study. If the experiment requires denaturation of the antigen, the procedure can be adapted to include this (see Alternate Protocol 2 for higher eukaryotic cells); however, the yeast cells must be broken with glass beads before heating the sample at 95°C.
Immunoprecipitation
5. Shake cells by vortexing vigorously at maximum speed for four 30-sec periods, keeping the cells on ice for 30 sec between the periods.
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Check cell breakage under a bright-field or phase-contrast microscope. It is helpful to check ahead of time if the cells can be broken in this way.
6. Remove the yeast cell lysate from the beads using a pipettor with a disposable tip. Transfer to a fresh tube. 7. Add 4 vol (see step 4) lysis buffer to the glass beads, vortex for 2 sec, and combine this supernatant with the lysate from step 6. 8. Clear the lysate and perform immunoprecipitation (see Basic Protocol 1, steps 6 to 27). IMMUNOPRECIPITATION WITH ANTIBODY-SEPHAROSE This protocol, which follows the steps presented in Figure 9.8.2, relies on the formation of an insoluble immune complex between a protein antigen and an antigen-specific monoclonal (or polyclonal) antibody covalently bound to Sepharose.
ALTERNATE PROTOCOL 5
Materials Unlabeled cells, surface-labeled cells (e.g., with 125I or biotin; UNIT 3.6) or biosynthetically 35S-, 3H-, or 14C-labeled cells (UNIT 3.7) Triton X-100 lysis buffer (see recipe) Dilution buffer (see recipe) Antibody (Ab)-Sepharose (see Support Protocol) Activated, quenched (control) Sepharose, prepared as for Ab-Sepharose (see Support Protocol) but eliminating Ab or substituting irrelevant Ab during coupling Tris/saline/azide (TSA) solution (see recipe) 0.05 M Tris⋅Cl, pH 6.8 2× SDS sample buffer (UNIT 10.1) NOTE: Carry out all procedures in a 4°C cold room or on ice. Lyse cells and preclear the lysate 1. Incubate cells in Triton X-100 lysis buffer (5 × 107 cells/ml) for 1 hr at 4°C. 2. Centrifuge the lysate 10 min at 3000 × g to remove nuclei and save the supernatant. 3. Centrifuge the supernatant 1 hr at 100,000 × g and save the supernatant. Supernatants may also be prepared by microcentrifugation (10,000 × g) for 30 min. The supernatant must be used within several days or stored at −70°C. The length of storage is limited by autoradiolysis and the half-life of the isotope. 3H- and 14C-labeled samples can often be stored frozen for years. Storage of 125I-labeled samples is usually limited to 1 to 2 months because of autoradiolysis, while the usefulness of 35S-labeled samples is usually limited to 6 months because of half-life. Repeated freezing and thawing may disrupt antigenic determinants and dissociate some protein complexes, especially those that are noncovalently associated.
4. Preclear supernatant to be used in one batch by adding 10 µl activated, quenched (control) Sepharose per 200 µl supernatant. Shake on an orbital shaker 2 hr at room temperature or overnight at 4°C. Centrifuge 1 min at 200 × g and save supernatant. Preclearing removes nonspecifically absorbing material. Control Sepharose can be prepared without antibody or coupled with irrelevant (nonspecific) antibody. Irrelevant antibody is an antibody directed against an unrelated protein, and could also be whole IgG; it must not cross-react with the protein being immunoprecipitated. Affinity Purification
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Immunoprecipitate the antigen 5. Precoat 1.5-ml microcentrifuge tubes by filling with Triton X-100 lysis buffer 10 min at room temperature. Remove the solution by aspiration. Precoating minimizes antigen absorption to the tube.
6. Add 105 to 106 cpm of radiolabeled (125I or 35S) supernatant containing antigen (from step 4) to a precoated microcentrifuge tube and bring the volume to 200 µl with dilution buffer. The recommended amount of radioactivity is appropriate for eukaryotic cells with >1000 molecules of antigen/cell. It is assumed that detection on slab gels of 125I-labeled proteins will be carried out with enhancing screens and 35S-labeled proteins with fluorography. For nonradiolabeled samples, use 0.2 to 1 ml of precleared lysate.
cell containing unlabeled or radiolabeled protein antigens
1
1
2
2
1 lyse (detergent) 2 immunoprecipitate 3 wash antigen-specific 4 dissociate monoclonal (or polyclonal) antibody-Sepharose
antigen-specific monoclonal antibody
anti-lg antibody
3
3
4
4
(unlabeled) (radiolabeled) analyze by electrophoresis and silver stain
Immunoprecipitation
analyze by electrophoresis and autoradiography or colorimetric detection
Figure 9.8.2 Schematic representation of the stages of the immunoprecipitation protocols using either antibody-Sepharose (left, see Alternate Protocol 5) or anti-Ig serum (right, see Alternate Protocol 6). (1) Cell lysis. (2) Immunoprecipitation using specific antibodies coupled covalently to Sepharose beads (left) or specific antibodies combined with anti-Ig serum (right). (3) Washing. (4) Dissociation of the antigen-antibody complex in sample buffer for electrophoresis.
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7. Add ∼10 µl of a 1:1 slurry of Ab-Sepharose/dilution buffer and shake 1.5 hr at 4°C on an orbital shaker. The antibody coupled to Sepharose is antigen specific. As described in the following support protocol, 5 mg/ml antibody per milliliter Sepharose is coupled, and the amount actually coupled can be estimated as described in step 10 of the support protocol. Shaking must be vigorous enough to suspend the Sepharose. Shaking may be extended to 3 hr; longer periods may increase background.
Wash, dissociate, and analyze the immunoprecipitate 8. Wash the Ab-Sepharose with 1 ml of the buffers listed below. After each wash, centrifuge 1 min at 200 × g or microcentrifuge 5 sec. Then, carefully aspirate the supernatant with a fine-tipped Pasteur pipet and leave 10 µl of fluid above the pellet. After the fourth wash, centrifuge again to bring down any residual drops on the side of the tube, aspirate, and leave 10 µl over the pellet. First wash: dilution buffer Second wash: dilution buffer Third wash: TSA solution Fourth wash: 0.05 M Tris⋅Cl, pH 6.8. Prepare a fine-tipped Pasteur pipet by pulling the pipet in a flame, scoring with a diamond pen, and breaking at the score.
9. Add 20 to 50 µl of 2× SDS sample buffer. Because the sample buffer has a higher density than the wash solution, it will sink into the Sepharose; do not vortex, because Sepharose may stick to side of tube above buffer level. Cap the tube securely and incubate 5 min at 100°C. 10. Vortex and centrifuge 1 min at 200 × g or microcentrifuge 5 sec. Load the supernatant, carefully avoiding the Sepharose, into a gel lane and analyze by SDS-PAGE (UNIT 10.1). 11. Detect labeled proteins by autoradiography (UNIT 10.11) with an enhancing screen (125I), by fluorography (35S, 14C, or 3H), or by colorimetric or chemiluminescent detection (biotinylated proteins; UNIT 3.6). PREPARATION OF ANTIBODY-SEPHAROSE This protocol details the procedure for covalently linking an antibody to Sepharose (an insoluble, large-pore-size chromatographic matrix) using the cyanogen bromide activation method. It is necessary to first prepare the antibody and Sepharose separately. Next, the Sepharose is activated with cyanogen bromide (alternatively, CNBr-activated Sepharose can be purchased from Amersham Pharmacia Biotech and used according to the manufacturer’s instructions). Finally, the CNBr-activated Sepharose is coupled to the antibody. Materials 1 to 30 mg/ml antigen-specific monoclonal or polyclonal antibody 0.1 M NaHCO3/0.5 M NaCl Sepharose CL-4B (or Sepharose CL-2B for high-molecular-weight antigens; Amersham Pharmacia Biotech) 0.2 M Na2CO3 Cyanogen bromide (CNBr)/acetonitrile (see recipe) 1 mM and 0.1 mM HCl, ice-cold (APPENDIX 2E) 0.05 M glycine (or ethanolamine), pH 8.0 Tris/saline/azide (TSA) solution (see recipe)
SUPPORT PROTOCOL
Affinity Purification
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Dialysis tubing (molecular weight cutoff >10,000) Whatman no. 1 filter paper Buchner funnel Erlenmeyer filtration flask Water aspirator Prepare the antibody 1. Dialyze 1 to 30 mg/ml antibody against 0.1 M NaHCO3/0.5 M NaCl at 4°C with three buffer changes during 24 hr. Use a volume of dialysis solution that is 500 times the volume of antibody solution. Dialysis is performed to remove all small molecules containing free amino or sulfhydryl groups (see UNIT 4.4 and APPENDIX 3C).
2. Centrifuge 1 hr at 100,000 × g, 4°C, to remove aggregates. Save the supernatant. Removal of aggregates is important. Because only some of the antibody molecules in an aggregate will be directly coupled to the Sepharose, the noncoupled antibody molecules may leach out during elution.
3. Measure the A280 of an aliquot of the solution and determine the concentration of the antibody (mg/ml IgG = A280/1.44). Dilute with 0.1 M NaHCO3/0.5 M NaCl to 5 mg/ml (or to the same concentration as desired for Ab-Sepharose) and keep at 4°C. Measure the A280 of this solution for later use in step 11. Prepare the Sepharose 4. Allow the Sepharose slurry to settle in a beaker and decant and discard the supernatant. Weigh out the desired quantity of Sepharose (assume density = 1.0). 5. Set up a filter apparatus using Whatman no. 1 filter paper in a Buchner funnel and an Erlenmeyer filtration flask attached to a water aspirator. Wash the Sepharose on the filter apparatus with 10 vol water. Sintered-glass funnels are traditionally recommended but rapidly become clogged unless coarse-porosity funnels are used.
Activate Sepharose with cyanogen bromide 6. Transfer Sepharose to 50-ml beaker and add an equal volume of 0.2 M Na2CO3. 7. Activate Sepharose at room temperature using 3.2 ml CNBr/acetonitrile per 100 ml Sepharose. Add CNBr/acetonitrile dropwise with a Pasteur pipet over 1 min, while slowly stirring the slurry with a magnetic stirrer. Continue stirring slowly for 5 min. Excessive and vigorous stirring may fracture the Sepharose beads. The protocol uses 2 g CNBr/100 ml Sepharose. Two to four grams of CNBr/100 ml Sepharose can be used to couple 1 to 20 mg of antibody/ml Sepharose. CAUTION: Activation should be carried out in a fume hood.
8. Rapidly filter the CNBr-activated Sepharose as in step 5. Aspirate to semidryness (i.e., until the Sepharose cake cracks and loses its sheen). 9. Wash with 10 vol ice-cold 1 mM HCl, then with 2 vol of ice-cold 0.1 mM HCl. Hydrate the cake with enough ice-cold 0.1 mM HCl so the cake regains its sheen, but so there is no excess liquid above the cake. Washing is most efficient if the wash solution is added evenly over the surface of the cake at about the same rate as the solution is removed by filtration. CNBr-activated Sepharose is very unstable at the alkaline pH necessary for activation; it is much more stable in dilute HCl. CNBr-activated Sepharose can be purchased premade from Amersham Pharmacia Biotech, but the coupling capacity will be lower. Immunoprecipitation
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Couple antibody to CNBr-activated Sepharose 10. Immediately transfer a weighed amount of Sepharose (assume density = 1.0) to a beaker. Add an equal volume of a solution of antibody dissolved in 0.1 M NaHCO3/ 0.5 M NaCl (from step 2). Stir gently with a magnetic stirrer or rotate end over end 2 hr at room temperature or overnight at 4°C. 11. Add 0.05 M glycine (or ethanolamine), pH 8.0, to saturate the remaining reactive groups on the Sepharose and allow the slurry to settle. Remove an aliquot of the supernatant, centrifuge to remove any residual Sepharose, and measure A280. Compare absorbance to that of the A280 of the antibody solution from step 2 to determine the percentage coupling. 12. Store the Ab-Sepharose in TSA solution. IMMUNOPRECIPITATION OF RADIOLABELED ANTIGEN WITH ANTI-Ig SERUM
ALTERNATE PROTOCOL 6
This protocol relies on the formation of soluble immune complexes between a protein and an antigen-specific antibody, followed by immunoprecipitation of the immune complexes by antibodies contained in anti-immunoglobulin (Ig) serum. This procedure is usually only used with radiolabeled or biotinylated antigen, as the unlabeled antibody remains in the precipitate and greatly complicates the use of any other detection method. Additional Materials (also see Alternate Protocol 5) Normal serum Anti-Ig serum (Zymed Laboratories) Antigen-specific antiserum or antigen-specific purified monoclonal antibody or antigen-specific hybridoma culture supernatant Follow the procedures in Alternate Protocol 5, with the following modifications at the indicated steps: 4a. Preclear by adding normal serum at a concentration of 2 µl/ml radiolabeled antigen. Add the proper amount of anti-Ig serum and let stand 12 to 18 hr at 4°C. Centrifuge 10 min at 1000 × g and reserve supernatant. Normal serum is the source of carrier Ig. The proper amount of anti-Ig serum must be determined by titration with radiolabeled antigen or Ig. For high-titered anti-Ig serum, this amount would be 20× to 40× the volume of antigen-specific antiserum, 2 to 4 µl/µg purified MAb, or one-third the volume of hybridoma culture supernatant.
7a. Add 1 µl antigen-specific antiserum, 3 µg antigen-specific purified MAb, or antigenspecific hybridoma culture supernatant (30 µl cloned line or 100 µl uncloned line). Vortex and allow to stand 2 hr at 4°C. Then add the proper amount of anti-Ig serum, vortex, and allow to stand 12 to 18 hr at 4°C. 8a. Wash the immunoprecipitate (see Alternate Protocol 5, step 8), except centrifuge 7 min at 1000 × g. 9a. Add 20 to 50 µl of 2× SDS sample buffer. Do not vortex, as immunoprecipitates may stick to side of tube above buffer level. Cap the tube securely. For immunoprecipitates, incubate first 1 hr at 56°C and then 5 min at 100°C. The initial 56°C incubation enhances the dissolution of the immunoprecipitates by reducing irreversible aggregation which occurs when precipitated protein is rapidly heated to 100°C. Proteolytic degradation has never been noted, probably because of the high IgG protein concentration.
Affinity Purification
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BASIC PROTOCOL 2
IMMUNOPRECIPITATION-RECAPTURE Once an antigen has been isolated by immunoprecipitation, it can be dissociated from the beads and reimmunoprecipitated (“recaptured”) either with the same antibody used in the first immunoprecipitation or with a different antibody (Fig. 9.8.3). Immunoprecipitationrecapture with the same antibody allows identification of a specific antigen in cases where the first immunoprecipitation contains too many bands to allow unambiguous identification. By using a different antibody in the second immunoprecipitation, immunoprecipitation-recapture can be used to analyze the subunit composition of multi-protein complexes (Fig. 9.8.4). The feasibility of this approach depends on the ability of the second antibody to recognize denatured antigens. Dissociation of the antigen from the beads is achieved by denaturation of antigen-antibody-bead complexes at high temperature in the presence of SDS and DTT. Prior to recapture, the SDS is diluted in a solution containing Triton X-100, and the DTT is neutralized with excess iodoacetamide. Recapture is then performed as in the first immunoprecipitation (see Basic Protocol 1, steps 6 to 26). Materials Elution buffer (see recipe) Beads containing bound antigen (see Basic Protocol 1, step 26) 10% (w/v) BSA Nondenaturing lysis buffer (see recipe) Heating block set at 95°C (Eppendorf Thermomixer 5436 or equivalent) 1. Add 50 µl elution buffer to 15 µl beads containing bound antigen. Mix by vortexing. The DTT in the elution buffer reduces disulfide bonds in the antigen and the antibody, and the SDS contributes to the unfolding of polypeptide chains.
2. Incubate 5 min at room temperature and 5 min at 95°C in a heating block. Cool tubes to room temperature. 3. Add 10 µl of 10% BSA. Mix by gentle vortexing. BSA is added to prevent adsorption of antigen to the tube, and to quench nonspecific binding to antibody-conjugated beads.
4. Add 1 ml nondenaturing lysis buffer. The iodoacetamide in the nondenaturing lysis buffer reacts with the DTT and prevents it from reducing the antibody used in the recapture steps. The presence of PMSF and leupeptin in the buffer is not necessary at this step.
5. Incubate 10 min at room temperature. 6. Clear the lysate and perform second immunoprecipitation (see Basic Protocol 1, steps 6 to 26). REAGENTS AND SOLUTIONS Use deionized or distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
CNBr/acetonitrile To 25 g of cyanogen bromide (CNBr should be white, not yellow, crystals), add 50 ml acetonitrile to make a 62.5% (w/v) solution. This may be stored indefinitely at −20°C in a desiccator over silica. Allow to warm before opening. Immunoprecipitation
CAUTION: CNBr is a highly toxic lachrymator; handle in fume hood.
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antigen 2 antigen 1 antibody 1 antibody 2
protein A– agarose bead
1
protein A–agarose bead
denaturation (see Basic Protocol 2)
2
antibody 2 binding to protein A–agarose bead (see Basic Protocol 1, steps 8 to 14)
3 recapture
wash and analysis
Figure 9.8.3 Scheme showing the stages of immunoprecipitation-recapture. (1) Dissociation and denaturation of the antigen: an antigen immunoprecipitated with antibody 1 bound to protein A–agarose beads is dissociated and denatured by heating in the presence of SDS and DTT. (2) Immobilization of the second antibody: antibody 2 is bound to protein A–agarose beads. (3) Recapture: the denatured antigen 2 (striped oval) is recaptured on antibody 2 bound to protein A–agarose beads. Alternatively, antibody 1 can be used again for further purification of the original antigen (square).
Affinity Purification
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Ab to:
1st
IP
2nd
IP
BSA
σ3
σ3
µ3
BSA
1
2
3
4
5
200
98 66 46
30 22
15
Figure 9.8.4 Example of an immunoprecipitation-recapture experiment. Human M1 fibroblasts were labeled overnight with [35S]methionine (UNIT 3.7) and extracted with nondenaturing lysis buffer (see Basic Protocol 1). The cell extract was then subjected to immunoprecipitation with antibodies to BSA (irrelevant antibody control; lane 1) and to the AP-3 adaptor (σ3; lane 2), a protein complex involved in protein sorting. Notice the presence of several specific bands in lane 2. The AP-3 immunoprecipitate was denatured as described in Basic Protocol 2 and individual components of the AP-3 complex were recaptured with antibodies to two of its subunits: σ3 (Mr ∼22,000; lane 3) and µ3 (Mr ∼47,000; lane 4). An immunoprecipitation with an antibody to BSA was also performed as a nonspecific control (lane 5). The amount of immunoprecipitate loaded on lanes 1 and 2 is ∼1⁄10 the amount loaded on lanes 3 to 5. Notice the presence of single bands in lanes 3 and 4. The positions of Mr standards (expressed as 10−3 × Mr) are shown at left. IP, immunoprecipitation.
Denaturing lysis buffer 1% (w/v) SDS 50 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) 5 mM EDTA (APPENDIX 2E) Store up to 1 week at room temperature (SDS precipitates at 4°C) Add the following fresh before use: 10 mM dithiothreitol (DTT; from powder) 1 mM phenylmethylsulfonyl fluoride (PMSF; store 100 mM stock in 100% ethanol up to 6 months at −20°C) 2 µg/ml leupeptin (store 10 mg/ml stock in H2O up to 6 months at −20°C) 15 U/ml DNase I (store 15,000 U/ml stock solution up to 2 years at −20°C)
Immunoprecipitation
1 mM 4-(2-aminoethyl)benzenesulfonyl fluoride (AEBSF), added fresh from a 0.1 M stock solution in H2O, can be used in place of PMSF. AEBSF stock can be stored up to 1 year at −20°C.
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Detergent-free lysis buffer PBS (APPENDIX 2E) containing: 5 mM EDTA (APPENDIX 2E) 0.02% (w/v) sodium azide Store up to 6 months at 4°C Immediately before use add: 10 mM iodoacetamide (from powder) 1 mM PMSF (store 100 mM stock in 100% ethanol up to 6 months at −20°C) 2 µg/ml leupeptin (store 10 mg/ml stock in H2O up to 6 months at −20°C) 1 mM AEBSF, added fresh from a 0.1 M stock solution in H2O, can be used in place of PMSF. AEBSF stock can be stored up to 1 year at −20°C.
Dilution buffer TSA solution (see recipe) containing: 0.1% Triton X-100 (store at room temperature in dark) 0.1% bovine hemoglobin (store frozen) Elution buffer 1% (w/v) SDS 100 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) Store up to 1 week at room temperature 10 mM DTT (add fresh from powder before use) Nondenaturing lysis buffer 1% (w/v) Triton X-100 (store at room temperature in dark) 50 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) 300 mM NaCl 5 mM EDTA (APPENDIX 2E) 0.02% (w/v) sodium azide Store up to 6 months at 4°C Immediately before use add: 10 mM iodoacetamide (from powder) 1 mM PMSF (store 100 mM stock in 100% ethanol up to 6 months at −20°C) 2 µg/ml leupeptin (store 10 mg/ml stock in H2O up to 6 months at −20°C) 1 mM AEBSF, added fresh from a 0.1 M stock solution in H2O, can be used in place of PMSF. AEBSF stock can be stored up to 1 year at −20°C.
Tris/saline/azide (TSA) solution 10 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 140 mM NaCl 0.025% NaN3 CAUTION: Sodium azide (NaN3) is poisonous; wear gloves.
Triton X-100 lysis buffer TSA solution (see recipe) containing: 1% Triton X-100 (store at room temperature in dark) 1% bovine hemoglobin (store frozen) 1 mM iodoacetamide (from powder) Aprotinin (0.2 trypsin inhibitor U/ml) 1 mM PMSF (store 100 mM stock in 100% ethanol up to 6 months at −20°C) Prepare fresh 1 mM AEBSF, added fresh from a 0.1 M stock solution in H2O, can be used in place of PMSF. AEBSF stock can be stored up to 1 year at −20°C. Affinity Purification
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Wash buffer 0.1% (w/v) Triton X-100 (store at room temperature in dark) 50 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) 300 mM NaCl 5 mM EDTA (APPENDIX 2E) 0.02% (w/v) sodium azide Store up to 6 months at 4°C COMMENTARY Background Information
Immunoprecipitation
The use of antibodies for immunoprecipitation has its origin in the precipitin reaction (Nisonoff, 1984, and references therein). The term precipitin refers to the spontaneous precipitation of antigen-antibody complexes formed by interaction of certain polyclonal antibodies with their antigens. The precipitation arises from formation of large networks of antigen-antibody complexes, due to the bivalent or polyvalent nature of immunoglobulins and to the presence of two or more epitopes in some antigens. This phenomenon was quickly exploited to isolate antigens from protein mixtures; however, its use remained limited to antibodies and antigens that were capable of multivalent interaction. In addition, the efficiency of precipitate formation was highly dependent on the concentrations of antibody and antigen. Thus, the precipitin reaction was not generally applicable as a method for immunoprecipitation. A significant improvement was the use of secondary anti-immunoglobulin reagents (generally anti-immunoglobulin serum) to crosslink the primary antibodies, thus promoting the formation of a precipitating network. In the 1970s, immunoprecipitation became widely applicable to the study of cellular antigens as a result of several technological advances. A critical development was the introduction of methods for the production of monoclonal antibodies (Köhler and Milstein, 1975). The ability to produce unlimited amounts of antibodies with specificity against virtually any cellular antigen had a profound impact in many areas of biology and medicine. The fact that preparation of monoclonal antibodies did not require prior purification of the antigens accelerated the characterization of cellular proteins and organelles, a process in which immunoprecipitation protocols played a major role. To this day, monoclonal antibodies produced in mice or rats continue to be among the most useful tools in biology. Another important development was the discovery of bacterial Fc receptors, proteins found
on the surface of bacteria that have the property of binding a wide range of immunoglobulins. Two of the most widely used bacterial Fc receptors are protein A from Staphylococcus aureus (Kessler, 1975) and protein G from group G streptococci. Protein A and protein G bind both polyclonal and monoclonal antibodies belonging to different subclasses and deriving from different animal species (Table 9.8.1). Protein A was initially used to adsorb immunoglobulins as part of fixed, killed S. aureus particles. Both protein A and protein G are now produced in large quantities by recombinant DNA procedures and are available coupled to solid-phase matrices such as agarose. In most cases, the binding of polyclonal or monoclonal antibodies to immobilized protein A (or G) avoids the need to use a secondary antibody to precipitate antigen-antibody complexes. Because of their broad specificity and ease of use, protein A–agarose and protein G–agarose (and related products) are the state-of-the-art reagents for the isolation of soluble antigen-antibody complexes in immunoprecipitation protocols. Recent progress in the field of antibody engineering (reviewed by Rapley, 1995; Irving et al., 1996) promises to make antibody production a less time-consuming and haphazard process. Antibody fragments with high affinity for specific antigens can now be selected from phage display antibody libraries. Selected recombinant antibodies can then be produced in large quantities in Escherichia coli. Techniques have been developed for producing antibodies in soluble, secreted form. Affinity tags are added to the recombinant antibody molecules to facilitate purification, detection, and use in procedures such as immunoprecipitation. While attractive in principle, the production of recombinant antibodies has been plagued by technical difficulties that so far have limited their widespread use in biology. However, as technical problems are overcome, recombinant techniques will progressively replace immunization of animals as a way of producing anti-
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bodies for immunoprecipitation and for other applications.
Critical Parameters Extraction of antigens Isolation of cellular antigens by immunoprecipitation requires extraction of the cells so that the antigens are available for binding to specific antibodies, and are in a physical form that allows separation from other cellular components. Extraction with nondenaturing detergents such as Triton X-100 (see Basic Protocol 1, Alternate Protocol 1, and Alternate Protocol 5) or in the absence of detergent (see Alternate Protocol 3) allows immunoprecipitation with antibodies to epitopes that are exposed on native proteins. Other nondenaturing detergents such as Nonidet P-40, CHAPS, digitonin, or octyl glucoside are also appropriate for extraction of native proteins (APPENDIX 1B). Some of these detergents (e.g., digitonin) preserve weak protein-protein interactions better than Triton X-100. If the antigen is part of a complex that is insoluble in nondenaturing detergents (e.g., cytoskeletal structures, chromatin, membrane “rafts”) or if the epitope is hidden within the folded structure of the protein, extraction under denaturing conditions is indicated (see Alternate Protocol 2). Alternate Protocol 2 may also be indicated for in vitro–translated products, which often tend to form aggregates (Anderson and Blobel, 1983). The number of cells necessary to detect an immunoprecipitated antigen depends on the cellular abundance of the antigen and on the efficiency of radiolabeling. The protocols for radiolabeling (UNIT 3.7) and immunoprecipitation described in this book are appropriate for detection of antigens that are present at low to moderate levels (10,000 to 100,000 copies per cell), as is the case for most endogenous integral membrane proteins, signal transduction proteins, and transcription factors. For more abundant antigens, such as cytoskeletal and secretory proteins or proteins that are expressed by viral infection or transfection, the quantity of radiolabeled cells used in the immunoprecipitation can be reduced accordingly. Production of antibodies Immunoprecipitation can be carried out using either polyclonal or monoclonal antibodies (see discussion of selection below). Polyclonal antibodies are most often prepared by immunizing rabbits, although polyclonal antibodies produced in mice, guinea pigs, goats, sheep,
and other animals are also suitable for immunoprecipitation. Antigens used for polyclonal antibody production can be whole proteins purified from cells or tissues, or whole or partial proteins produced in bacteria or insect cells by recombinant DNA procedures. Another useful procedure is to immunize animals with peptides conjugated to a carrier protein. Production of polyclonal antibodies to recombinant proteins and peptides has become the most commonly used approach to obtain specific probes for immunoprecipitation and other immunochemical techniques, because it does not require purification of protein antigens from their native sources. The only requirement for making these antibodies is knowledge of the sequence of a protein, which is now relatively easy to obtain as a result of cDNA library production and genomic DNA sequencing projects. Polyclonal antibodies can be used for immunoprecipitation in the form of whole serum, ammonium sulfate–precipitated immunoglobulin fractions, or affinity-purified immunoglobulins. Although all of these are suitable for immunoprecipitation, affinity-purified antibodies often give lower backgrounds and are more specific. Most monoclonal antibodies are produced in mice or rats. The sources of antigen for monoclonal antibody production are the same as those for production of polyclonal antibodies, namely proteins isolated from cells or tissues, recombinant proteins or protein fragments, and peptides. A significant advantage of using monoclonal antibodies is that antigens do not need to be purified to serve as immunogens, as long as the screening method is specific for the antigen. Another advantage is the unlimited supply of monoclonal antibodies afforded by the ability to grow hybridomas in culture or in ascitic fluid. Many monoclonal antibodies can now be produced from hybridomas deposited in cell banks or are directly available commercially. Ascitic fluid, cell culture supernatant, and purified antibodies are all suitable sources of monoclonal antibodies for immunoprecipitation. Ascitic fluid and purified antibodies should be used when a high antibody titer is important. Cell culture supernatants have lower antibody titers, but tend to give cleaner immunoprecipitations than ascitic fluids due to the lack of contaminating antibodies. Selection of antibodies: Polyclonal versus monoclonal What type of antibody is best for immunoprecipitation? There is no simple answer to this
Affinity Purification
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Immunoprecipitation
question, as the outcome of both polyclonal and monoclonal antibody production protocols is still difficult to predict. Polyclonal antibodies to whole proteins (native or recombinant) have the advantage that they frequently recognize multiple epitopes on the target antigen, enabling them to generate large, multivalent immune complexes. Formation of these antigenantibody networks enhances the avidity of the interactions and increases the efficiency of immunoprecipitation. Because these antibodies recognize several epitopes, there is a better chance that at least one epitope will be exposed on the surface of a solubilized protein and thus be available for interaction with antibodies. Thus, the likelihood of success is higher. These properties can be a disadvantage, though, as some polyvalent antibodies can cross-react with epitopes on other proteins, resulting in higher backgrounds and possible misidentification of antigens. Because they are directed to a short peptide sequence, anti-peptide polyclonal antibodies are less likely to cross-react with other proteins. However, their usefulness is dependent on whether the chosen sequence turns out to be a good immunogen in practice, as well as on whether this particular epitope is available for interaction with the antibody under the conditions used for immunoprecipitation. Unfractionated antisera are often suitable for immunoprecipitation. However, there is a risk that serum proteins other than the antibody will bind nonspecifically to the immunoadsorbent, and in turn bind proteins in the lysate that are unrelated to the antigen. For instance, transferrin can bind nonspecifically to immunoadsorbents, potentially leading to the isolation of the transferrin receptor as a contaminant (Harford, 1984). Polyclonal antisera can also contain antibodies to other antigens (e.g., viruses, bacteria) to which the animal may have been exposed, and these antibodies can also crossreact with cellular proteins during immunoprecipitation. Affinity-purified antibodies are a better alternative when antisera do not yield clean immunoprecipitations. Affinity-purification can lead to loss of high-affinity or low-affinity antibodies; however, the higher specificity of affinity-purified antibodies generally makes them “cleaner” reagents for immunoprecipitation. The specificity, high titer, and limitless supply of the best immunoprecipitating monoclonal antibodies are unmatched by those of polyclonal antibodies. However, not all monoclonal antibodies are useful for immunopre-
cipitation. Low-affinity monoclonal antibodies can perform acceptably in immunofluorescence microscopy protocols but may not be capable of holding on to the antigen during the repeated washes required in immunoprecipitation protocols. The use of ascitic fluid has the same potential pitfalls as the use of polyclonal antisera, as ascites may also contain endogenous antibodies to other antigens and proteins such as transferrin that can bind to other proteins in the lysate. In conclusion, an informed empirical approach is recommended in order to select the best antibody for immunoprecipitation. In general, it is advisable to generate and/or test several antibodies to a particular antigen in order to find at least one that will perform well in immunoprecipitation protocols. Antibody titer The importance of using the right amount of antibody for immunoprecipitation cannot be overemphasized. This is especially the case for quantitative immunoprecipitation studies, in which the antibody should be in excess of the specific antigen. For instance, in pulse-chase analyses of protein degradation or secretion (UNIT 3.7), it is critical to use sufficient antibody to deplete the antigen from the cell lysate. This is particularly important for antigens that are expressed at high levels, a common occurrence with the growing use of high-yield protein expression systems such as vaccinia virus (UNITS 5.11-5.15) or replicating plasmids in COS cells. Consider, for example, a protein that is expressed at high levels inside the cell, and of which only a small fraction is secreted into the medium. If limiting amounts of antibody are used in a pulse-chase analysis of this protein, the proportion of protein secreted into the medium will be grossly overestimated, because the limiting antibody will bind only a small proportion of the cell-associated protein and a much higher proportion of the secreted protein. The same considerations apply to degradation studies. Thus, it is extremely important in quantitative studies to ensure that the antibody is in excess of the antigen in the cell samples. This can be ascertained by performing sequential immunoprecipitations of the samples (see Basic Protocol 1, annotation to step 21). If the second immunoprecipitation yields only a small amount of the antigen relative to that isolated in the first immunoprecipitation (<10%), then the antibody titer is appropriate. If, on the other hand, the amount of antigen isolated in the second immunoprecipitation is
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>10%, either more antibody or less antigen should be used. Too much antibody can also be a problem, as nonspecific immunoprecipitation tends to increase with increasing amounts of immunoglobulins bound to the beads. Thus, titration of the antibody used for immunoprecipitation is strongly advised. Immunoadsorbent If cost is not an overriding issue, the use of protein A– or protein G–agarose is recommended for routine immunoprecipitation (see Basic Protocol 1). Protein A– or protein G– agarose beads (or equivalent products) have a very high capacity for antibody binding (up to 10 to 20 mg of antibody per milliliter of gel). Both protein A and protein G bind a wide range of immunoglobulins (Table 9.8.1). Backgrounds from nonspecifically bound proteins are generally low. Protein A– and protein G– agarose beads are also stable and easy to sediment by low-speed centrifugation. A potential disadvantage, in addition to their cost, is that some polyclonal or monoclonal antibodies bind weakly or not at all to protein A or protein G (Table 9.8.1). This problem can be solved by using an intermediate rabbit antibody to the immunoglobulin of interest. For example, a goat polyclonal antibody can be indirectly bound to protein A–agarose by first incubating the protein A–agarose beads with a rabbit antigoat immunoglobulin, and then incubating the beads with the goat polyclonal antibody. Antiimmunoglobulin antibodies (e.g., rabbit anti– goat immunoglobulins) coupled covalently to agarose can also be used for indirect immunoprecipitation in place of protein A– or protein G–agarose. A less expensive alternative to protein A– or protein G–agarose is the use of anti-Ig serum to crosslink the primary antibody (see Alternate Protocol 6). This procedure can result in very low backgrounds, although it requires proper titration of the anti-Ig serum. Protein A– agarose can also be substituted by fixed Staphylococcus aureus particles (Pansorbin). They have a lower capacity, can give higher backgrounds, and take longer to sediment. However, they work quite well in many cases. In order to establish if they are appropriate for a particular experimental setup, conduct a preliminary comparison of the efficiency of protein A– agarose with Staphylococcus aureus particles as immunoadsorbent. Specific antibodies coupled covalently to various affinity matrices can also be used for
direct immunoprecipitation of antigens (see Alternate Protocol 5). After binding to protein A–agarose, antibodies can be cross-linked with dimethylpimelimidate (Gersten and Marchalonis, 1978). Purified antibodies can also be coupled directly to derivatized matrices such as CNBr-activated Sepharose (see Support Protocol). This latter approach avoids having to bind the antibody to protein A–agarose. Covalently bound antibodies should be used when elution of immunoglobulins from the beads complicates further analyses of the complexes. This is the case when proteins in immunoprecipitates are analyzed by one- or two-dimensional gel electrophoresis (UNITS 10.1-10.4) followed by Coomassie blue or silver staining, or are used for microsequencing. Also, the released immunoglobulins could interfere with detection of some antigens by immunoblotting (UNIT 10.10) following immunoprecipitation. The support protocol for coupling protein antigens to CNBr-activated Sepharose is a modification of the methods of Cuatrecasas (1970) and March et al. (1974). As originally described, the washing was done at alkaline pH. Because activated Sepharose is very unstable at this pH, it was originally recommended that washing, adding the protein ligand, and mixing be done in <90 sec (Cuatrecasas, 1970). However, using an acid wash buffer (Gelb, 1973) as described in the support protocol causes activated groups to remain stable for ≥30 min. The 0.1 M NaHCO3 buffer in which antibody is dissolved provides an optimal pH of ∼8.4, after mixing with activated-Sepharose. The low amount of CNBr recommended is sufficient to obtain a coupling yield of 80% to >99%. Higher amounts of CNBr may result in multipoint attachment of IgG molecules to the matrix, thereby reducing accessibility to antigen. Most investigators purchase CNBr-activated Sepharose, while others, to achieve a higher coupling efficacy or to avoid the expense of the commercial product, prefer to prepare it themselves. Quantities and ratios recommended in these protocols have been found to work well with several hundred monoclonal antibodies and more than 40 different antigens. However, titration of Ab-Sepharose or sandwich reagents versus the protein antigen may further optimize a given immunoprecipitation. Nonspecific controls For correct interpretation of immunoprecipitation results, it is critical to include appropriate nonspecific controls along with the spe-
Affinity Purification
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additional wash with 0.1% SDS/0.1% DOC
PI I
PI I
200
46
30
22
15 1
2
3
4
Figure 9.8.5 Lowering background by washing with SDS and sodium deoxycholate (DOC). In this experiment, BW5147 cells (mouse thymoma) labeled with [35S]methionine for 1 hr were extracted with nondenaturing lysis buffer (see Basic Protocol 1). The extracts were subjected to immunoprecipitation with protein A–agarose beads incubated with either preimmune (PI) or immune (I) serum from a rabbit immunized with the ribosomal protein L17 (doublet at Mr ∼22,000). Lanes 1 and 2 correspond to immunoprecipitates obtained using the protocols described in this unit. Notice the presence of nonspecific bands and/or associated proteins in lane 2. Lanes 3 and 4 correspond to beads that were washed an additional time with 0.1% (w/v) SDS and 0.1% (w/v) DOC. Notice the disappearance of most of the nonspecific bands and/or associated proteins. The positions of Mr standards (expressed as 10−3 × Mr) are shown at left.
Immunoprecipitation
cific samples. One type of control consists of setting up an incubation with an irrelevant antibody in the same biochemical form as the experimental antibody (e.g., serum, ascites, affinity-purified immunoglobulin, antibody bound to protein A–agarose or directly conjugated to agarose), and belonging to the same species and immunoglobulin subclass as the experimental antibody (e.g., rabbit antiserum, mouse IgG2a). For an antiserum, the best control is preimmune serum (serum from the same animal obtained before immunization). Nonimmune serum from the same species is an accept-
able substitute for preimmune serum in some cases. “No-antibody” controls are not appropriate because they do not account for nonspecific binding of proteins to immunoglobulins. In immunoprecipitation-recapture experiments, control immunoprecipitations with irrelevant antibodies should be performed for both the first and second immunoprecipitation steps (Fig. 9.8.4). Another type of control is to perform an immunoprecipitation from cells that do not express a specific antigen in parallel with immunoprecipitation of the antigen-expressing cells. For instance, untransfected cells are a
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perfect control for transfected cells. In yeast cells, null mutants that do not express a specific antigen are an ideal control for wild-type cells. Order of stages In the immunoprecipitation procedure described in Basic Protocol 1, the antibody is prebound to protein A–agarose before addition to the cell lysate containing the antigen. This differs from other methods in which the free antibody is first added to the lysate and the antigen-antibody complexes are then collected by addition of the immunoadsorbent. Although both procedures can give good results, the authors prefer the protocol described here because this method allows better control of the amount of antibody bound to the immunoadsorbent. Prebinding antibodies to the immunoadsorbent beads allows removal of unbound antibodies. The presence of unbound antibodies in the incubation mixture could otherwise result in decreased recovery of the antigen on the immunoadsorbent beads. Another advantage of the prebinding procedure is that most proteins other than the immunoglobulin in the antibody sample (e.g., serum proteins) are removed from the beads and do not come in contact with the cell lysate. This eliminates potential adverse effects of these proteins on isolation of the antigen. Washing The five washes described in Basic Protocol 1 (four with wash buffer and one with PBS) are sufficient for maximal removal of unbound proteins; additional washes are unlikely to decrease the background any further. The last wash with PBS removes the Triton X-100 that can lead to decreased resolution on SDS-PAGE. It also removes other components of the wash buffer that could interfere with enzymatic treatment of immunoprecipitates. It is not advisable to complete all the washes quickly (e.g., in 5 min), because this may not allow enough time for included proteins to diffuse out of the gel matrix. Instead, beads should be washed over ∼30 min, which may require keeping the samples on ice for periods of 3 to 5 min between washes. In order to reduce nonspecific bands, samples can be subjected to an additional wash with wash buffer containing 0.1% (w/v) SDS, or with a mixture of 0.1% (w/v) SDS and 0.1% (w/v) sodium deoxycholate (Fig. 9.8.5). This wash should be done before the last wash with PBS (in Basic Protocol 1) or before the wash with 0.05 M Tris⋅Cl, pH 6.8 (in Alternate Protocol 5).
Troubleshooting Two of the most common problems encountered in immunoprecipitation of metabolically labeled proteins are failure to detect specific antigens in the immunoprecipitates, and high background of nonspecifically bound proteins for antigens that were radiolabeled in vivo and analyzed by SDS-PAGE (UNIT 10.1) followed by autoradiography or fluorography (UNIT 10.11). When immunoprecipitates are analyzed by immunoblotting (UNIT 10.10), an additional problem may be the detection of immunoprecipitating antibody bands in the blots (Table 9.8.2).
Anticipated Results For antigens that are present at >10,000 copies per cell, the radiolabeling and immunoprecipitation protocols described in this book can be expected to result in the detection of one or more bands corresponding to the specific antigen and associated proteins in the electrophoretograms. Specific bands should not be present in control immunoprecipitations done with irrelevant antibodies. If antigens are labeled with [35S]methionine (UNIT 3.7), specific bands should be visible within 2 hr to 2 months of exposure. Due to the relatively low yield of the immunoprecipitation-recapture procedure (<10% of that of a single immunoprecipitation step), detection of specific bands is likely to require longer exposure times. This may turn out to be problematic due to the radioactive decay of 35S (half-life = 88 days). In immunoprecipitation-recapture experiments in which the antibodies used for the first and second immunoprecipitation steps recognize different antigens (e.g., for the study of protein-protein interactions; Fig. 9.8.3), it is advisable to include in the second immunoprecipitation step a positive control containing either the antibody used for the first immunoprecipitation (if the antibody recognizes both the native and denatured forms of the antigen) or a different antibody with specificity for the same antigen (Fig. 9.8.4, lane 3).
Time Considerations Preparation of cell extracts (Basic Protocol 1 and Alternate Protocols 1 to 4) takes 1 to 3 hr to complete, and isolation of the antigen on antibody-conjugated beads takes 2 to 3 hr. Binding antibodies to immunoadsorbent beads can be done prior to or simultaneously with preparation of the cell extracts and also takes 1 to 3 hr. Therefore, the whole immunoprecipitation procedure can be completed in 1 day. Similar time requirements apply for Alternate
Affinity Purification
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Table 9.8.2
Troubleshooting Guide for Immunoprecipitation
Problem
Cause
Solution
Gel is completely blank after prolonged autoradiographic exposure
Poorly labeled cells: too little radiolabeled precursor, too few cells labeled, lysis/loss of cells during labeling, too much cold amino acid in labeling mix, wrong labeling temperature
Check incorporation of label by TCA precipitation (UNIT 3.8); troubleshoot the labeling procedure
Only nonspecific bands present
Antigen does not contain the amino acid used for labeling
Label cells with another radiolabeled amino acid or, for glycoproteins, with tritiated sugar
Antigen expressed at very low levels
Substitute cells known to express higher levels of antigens as detected by other methods; transfect cells for higher expression
Protein has high turnover rate and is not well labeled by long-term labeling
Use pulse labeling
No specific radiolabeled antigen band
Protein has a low turnover rate and is Use long-term labeling not well labeled by short-term labeling Protein is not extracted by lysis buffer used to solubilize cells
Solublize with a different nondenaturing detergent or under denaturing conditions
Antibody is nonprecipitating
Identify and use antibody that precipitates antigen
Epitope is not exposed in native antigen
Extract cells under denaturing conditions
Antibody does not recognize denatured antigen
Extract cells under nondenaturing conditions
Antibody does not bind to immunoadsorbent
Use a different immunoadsorbent (Table 9.8.1); use intermediate antibody
Antigen is degraded during immunoprecipitation
Ensure that fresh protease inhibitors are present
Isolated lanes on gel with high background
Random carryover of detergent-insoluble proteins
Remove supernatant immediately after centrifugation, leaving a small amount with pellet; if resuspension occurs, recentrifuge
High background in all lanes
Incomplete washing
Cap tubes and invert several times during washes
Poorly radiolabeled protein
Optimize duration of labeling to maximize signal-to-noise ratio
Incomplete removal of detergent-insoluble proteins
Centrifuge lysate 1 hr at 100,000 × g
Insufficient unlabeled protein to quench nonspecific binding
Increase concentration of BSA
High background of nonspecific bands
continued
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Current Protocols in Protein Science
Table 9.8.2
Troubleshooting Guide for Immunoprecipitation, continued
Problem
Cause
Solution
Antibody contains aggregates
Microcentrifuge antibody 15 min at maximum speed before binding to beads
Antibody solution contains nonspecific antibodies
Use affinity-purified antibodies; absorb antibody with acetone extract of cultured cells that do not express antigen; for yeast cells, absorb antibody with null mutant cells
Too much antibody
Use less antibody
Incomplete preclearing
Preclear with irrelevant antibody of same species of origin and immunoglobulin subclass bound to immunoadsorbent
Nonspecifically immunoprecipitated proteins
Fractionate cell lysate (e.g, ammonium sulfate precipitation, lectin absorption, or gel filtration) prior to immunoprecipitation; after washes in wash buffer, wash beads once with 0.1% SDS in wash buffer or 0.1% SDS/0.1% sodium deoxycholate
Immunoprecipitating antibody detected in immunoblots Complete immunoglobulin or heavy and/or light chains visible in immunoblot
Protein A conjugate or secondary antibody recognizes immunoprecipitating antibody
Protocols 5 and 6. Reaction between antibody and antigen can be extended overnight, but this may increase the background when using AbSepharose. Antigen-antibody complexes should not be left overnight in the midst of washing, as significant dissociation may occur. Immunoprecipitation-recapture experiments (Basic Protocol 2) require an additional 1 to 2 hr to denature and prepare the antigen for immunoprecipitation, and 2 to 3 hr to isolate the antigen. Completion of an entire immunoprecipitation-recapture experiment requires a very long workday. Alternatively, samples can be frozen after the first immunoprecipitation, and the elution and recapture can be carried out another day. Immunoprecipitates can be analyzed immediately (e.g., resolved by SDSPAGE) or frozen and analyzed another day.
Use antibody coupled covalently to solid-phase matrix for immunoprecipitation; probe blots with primary antibody from a different species and the appropriate secondary antibody specific for immunoblotting primary antibody
Literature Cited Anderson, D.J. and Blobel, G. 1983. Immunoprecipitation of proteins from cell-free translations. Methods Enzymol. 96:111-120. Cuatrecasas, P. 1970. Protein purification by affinity chromatography. J. Biol. Chem. 245:3059. Franzusoff, A., Rothblatt, J., and Schekman, R. 1991. Analysis of polypeptide transit through yeast secretory pathway. Methods Enzymol. 194:662-674. Gelb, W.G. 1973. Affinity chromatography: For separation of biological materials. Am. Lab. 81:61-67. Gersten, D.M. and Marchalonis, J.J. 1978. A rapid, novel method for the solid-phase derivatization of IgG antibodies for immune-affinity chromatography. J. Immunol. Methods 24:305-309. Harford, J. 1984. An artefact explains the apparent association of the transferrin receptor with a ras gene product. Nature 311:673-675. Affinity Purification
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Harlow, E. and Lane, D. 1999. Antibodies: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Irving, R.A., Hudson, P.J., and Goding, J.W. 1996. Construction, screening and expression of recombinant antibodies. In Monoclonal Antibodies: Principles and Practice (J.W. Goding, ed.). Academic Press, London. Kessler, S.W. 1975. Rapid isolation of antigens from cells with a staphylococcal protein A–antibody adsorbent: Parameters of the interaction of antibody-antigen complexes with protein A. J. Immunol. 115:1617. Köhler, G. and Milstein, C. 1975. Continuous cultures of fused cells secreting antibody of predetermined specificity. Nature 256:495-497. March, S.C., Parikh, I., and Cuatrecasas, P. 1974. A simplified method for cyanogen bromide activation of agarose for affinity chromatography. Anal. Biochem. 60:149-152. Nisonoff, A. 1984. Introduction to Molecular Immunology. Sinauer Associates, Sunderland, Mass. Rapley, R. 1995. The biotechnology and applications of antibody engineering. Mol. Biotech. 3:139-154.
Key References Harlow and Lane, 1988. See above. Helenius, A., McCaslin, D.R., Fries, E., and Tanford, C. 1979. Properties of detergents. Methods Enzymol. 56:734-749. Hjelmeland, J.M. and Chrambach, A. 1984. Solubilization of functional membrane proteins. Methods Enzymol. 104:305-318. The above references describe various detergents and solubilization conditions.
Contributed by Juan S. Bonifacino and Esteban C. Dell’Angelica National Institute of Child Health and Human Development Bethesda, Maryland Timothy A. Springer (precipitation with antibody-Sepharose and with anti-Ig) Center for Blood Research Harvard Medical School Boston, Massachusetts
Immunoprecipitation
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Current Protocols in Protein Science
Overview of Affinity Tags for Protein Purification EARLY AFFINITY TAGS
Polyhistidine
The first affinity tags used were large proteins utilized almost exclusively for protein expression and purification in E. coli. Staphylococcal protein A is 280 amino acids in length and, because of its size and proteolytic stability, can increase the solubility and/or expression of heterologous proteins (Sambrook et al., 1989). Protein A fusions can be purified by affinity chromatography on IgG Sepharose and eluted with a low pH buffer (UNIT 9.5); however, the large size of the tag and/or the low pH elution can alter the functional activity of the fusion protein. In addition, the binding of fusion proteins to IgG complicates immunological analysis, and thus Protein A fusions are unsuitable for use as immunological reagents. LacZ, also known as β-galactosidase or βgal, is a proteolytically stable 1024–amino acid protein which can be used as an affinity tag to increase expression of fusion proteins in E. coli (Sambrook et al., 1989). LacZ fusions can be purified by substrate affinity chromatography on immobilized p-amino-phenyl-β-D-thiogalactosidase (APTG) and eluted with a high pH borate buffer. Detection is then achieved using a colorimetric enzymatic assay. Because LacZ is relatively immunologically neutral, it is a useful fusion partner for the generation of antibodies to a protein of interest. The LacZ affinity tag can alter the functional activity of the purified fusion protein because of its extremely large size and tendency to form homotetramers in solution. In addition, LacZ fusion proteins are often insoluble. This insolubility has advantages and disadvantages: protein targeting to inclusion bodies can allow the expression of gene products that are toxic to E. coli, but because LacZ must be correctly folded to bind APTG, LacZ fusion proteins must be refolded prior to purification and/or detection (UNIT 6.5).
The polyhistidine affinity tag, also known as the His-tag or His6, usually consists of six consecutive histidine residues, but can vary in length from two to ten histidine residues. Polyhistidine was first used to purify recombinant galactose dehydrogenase by immobilized metal affinity chromatography (IMAC; UNIT 9.4) in 1991 (Lilius et al., 1991). Previously, IMAC had been used to purify proteins with naturally occurring histidine- or tryptophan-containing sequences, but these sequences were not as generally applicable to protein expression and purification as would be desired for an affinitytagging system (Smith et al., 1988; Ljungquist et al., 1989). Polyhistidine is such a ubiquitous affinity tag that most companies providing expression vectors or protein expression and purification reagents offer products related to this tag (see Table 9.9.1 for examples). Histidine readily coordinates with immobilized transition metal ions. Immobilized Co2+, Cu2+, Ni2+, Zn2+, Ca2+, and Fe3+ can all be used to purify polyhistidine fusion proteins, but Ni2+ is the most commonly used. Empirical determination of the most effective transition metal ion for purification of a specific polyhistidine fusion protein can be performed if purification by Ni2+ is unsatisfactory. There are several companies that offer IMAC resin. Iminodiacetic acid agarose (chelating Sepharose, Amersham Biosciences), nitrilotriacetic acid agarose (Ni-NTA resin, Qiagen), and carboxymethylaspartate agarose (Talon resin, Clontech) are all used for the immobilization of transition metal ions. Commercially available IMAC resins are unaffected by protease or nuclease activity, and are appropriate for purification of fusion proteins from crude cell lysates. Most of these resins can be regenerated and reused indefinitely. Although polyhistidine does not usually cause a protein to be targeted to E. coli inclusion bodies, IMAC is amenable to denaturing agents (i.e., 8 M urea, 6 M guanidine⋅HCl, ionic and nonionic detergents, and low concentrations of reducing agents) for the purification of insoluble or membrane-bound proteins. High concentrations of reducing agents such as dithiothreitol (DTT) can reduce the immobilized metal ion and should be avoided. The relatively small size and charge of the polyhistidine tag rarely af-
COMMONLY USED AFFINITY TAGS Although protein A and lacZ are still used as affinity tags today (see Table 9.9.1 for commercially available systems), many other affinity tags have recently been developed that improve upon these original protein fusion partners.
Contributed by Michelle E. Kimple and John Sondek Current Protocols in Protein Science (2004) 9.9.1-9.9.19 Copyright ©2004 by John Wiley & Sons, Inc.
UNIT 9.9
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fects protein function, and elution by imidazole gradient is relatively mild, preserving the immunogenicity of polyhistidine fusion proteins. While purification of a highly-expressed polyhistidine fusion protein can lead to relatively (>80%) pure protein in one chromatographic step, purification from insect and mammalian cells, which contain a higher percentage of His residues in their proteins than E. coli, can lead to significant background binding to immobilized metal ions. This may be circumvented by using stringent wash conditions (e.g., 5 to 10 mM imidazole), although a stringent wash may cause premature elution of the protein of interest. The location of the tag (N terminal, C terminal, or internal) can also have an effect on IMAC. If a change in tag location does not increase the effectiveness of IMAC, a denaturing purification can be attempted. Primary antibodies have also been developed for the detection of polyhistidine fusion proteins in vitro. Again, because of the predominance of histidine residues in mammalian and insect systems, anti-polyhistidine antibodies are notoriously promiscuous. Ni2+ resin can also be used to precipitate a polyhistidinetagged protein for the detection of protein-protein interactions.
Glutathione S-Transferase
Overview of Affinity Tags for Protein Purification
The pGEX E. coli expression vectors, which encode for N-terminal glutathione S-transferase (GST) molecules followed by protease cleavage sites, were first designed and used to express and purify antigens of the parasite Taenia ovis in 1988 (Smith and Johnson, 1988; Smith, 2000). Currently, pGEX vectors are available from Amersham Biosciences in all three reading frames and with three different protease cleavage sites (e.g., thrombin, factor Xa, and PreScission). GST fusion proteins can be purified by affinity chromatography (UNIT 6.6) on commercially available glutathione (γ-glutamylcysteinylglycine) Sepharose, which is affected by γ-glutamyl transpeptidase activity in crude cell lysates. Therefore, glutathione resin has a finite lifetime and can only be regenerated and reused between four and twenty times. Glutathione affinity chromatography is amenable to low concentrations of denaturing agents (2 to 3 M urea or guanidine hydrochloride), reducing agents (<10 mM 2-mercaptoethanol or dithiothreitol), and nonionic detergents (2% v/v Tween 20), depending on the nature of the fusion protein. Elution of GST fusion proteins with 10 mM glutathione is relatively mild, often preserving protein function and antigenicity. A
70-kDa E. coli heat-shock-induced chaperonin often copurifies with eluted GST fusion proteins (Thain et al., 1996). This contaminant can be removed by treatment of cell lysates with 5 mM MgCl2 and 5 mM ATP prior to purification. Furthermore, GST can be cleaved from its fusion protein while still bound to glutathione agarose, providing a convenient method for separating the 26-kDa GST from the protein of interest. GST fusion proteins are often expressed at high levels in E. coli (typical yields ∼10 mg/ liter), which may result in accumulation of aggregated protein in inclusion bodies. Purification from inclusion bodies (UNIT 6.5) has both advantages and disadvantages. Some advantages are that protein targeting to E. coli inclusion bodies allows the high-level expression of toxic genes and the separation of inclusion bodies serves as a significant purification step from whole-cell lysate. Unfortunately, since glutathione affinity chromatography depends on the proper three-dimensional fold of GST, insoluble fusion proteins must be refolded and buffer exchanged before purification; however, some insoluble proteins may not refold correctly or into a soluble form. Another potential disadvantage of the GST tag is that the large size of the tag and its dimerization in solution may affect the properties of the fusion protein. GST fusion proteins can be detected by a colorimetric assay with the GST substrate 1chloro-2,4-dinitrobenzene (CDNB) or with anti-GST antibodies (e.g., Amersham Biosciences, Sigma, BD Biosciences). Precipitations of GST fusion proteins with glutathionecoupled beads are commonly used for the detection of protein-protein or pretein-DNA interactions.
Maltose Binding Protein pMAL E. coli expression vectors were designed in 1988 (di Guan et al., 1988; Maina et al., 1988). Maltose binding protein (MBP) is often used to increase the expression level and/or solubility of its fusion partner, with typical yields of 10 to 40 mg fusion protein per liter culture. pMAL vectors are available for cytoplasmic or periplasmic expression in all three reading frames, with factor Xa, enterokinase, or genenase I protease cleavage sequences (New England Biolabs). Other MBP fusion vectors include pIVEX (Roche), which can be used for coupled in vitro transcription/translation. MBP fusion proteins can be purified by affinity chromatography on cross-linked amy-
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lose resin. Amylose resins are commercially available, but are affected by amylase activity in crude cell lysates and can be regenerated and reused only three to five times. Adding glucose to bacterial growth media helps suppress amylase expression (see New England Biolabs pMAL fusion system FAQ; http://www.circuit. neb.com/neb/faqs/faq_pfp.html). Amylose affinity chromatography is not amenable to denaturing or reducing agents. Low concentrations of nonionic detergents (0.2% v/v Triton-X or Tween 20) may be used depending on the nature of fusion protein. Like GST fusion proteins, high-level expression of MBP fusion proteins in E. coli may result in accumulation of insoluble protein aggregates in inclusion bodies. The large size of the MBP tag (45 kDa) may also affect protein function. Unlike glutathione affinity chromatography (UNIT 6.6), proteolytic cleavage of the tag while bound to amylose resin is not effective and the fusion protein must be eluted by free maltose before cleavage. The 10 mM maltose elution is mild though, and often preserves protein function and antigenicity. If proteolytic removal of the MBP tag is performed, free maltose must be removed from the cleaved MBP by an additional chromatographic step if MBP is to rebind the amylose column (NEB recommends standard DEAE chromatography.) Anti-MBP antibodies are available for the detection of MBP fusion proteins (e.g., BD Biosciences, Sigma, Zymed ).
Intein-Chitin Binding Domain The intein-chitin binding domain (inteinCBD) tag is a combination of a protein selfsplicing element (intein) with a chitin-binding domain, and allows for the purification of a native recombinant protein without need for a protease. Intein-CBD E. coli expression vectors were designed in 1997 (Chong et al., 1997). Commercially available vectors provide for intein-CBD expression on the N-terminus, C-terminus, or both termini of a heterologous protein of interest (IMPACT system, New England Biolabs). CBD fusion proteins are purified by affinity chromatography on chitin resin. Commercially available resins can be regenerated at least five times. Nonspecific binding to chitin resin can be reduced by a stringent wash including high salt or detergent (2 M NaCl, 0.5% Triton X). Chitin affinity chromatography is not amenable to denaturing reagents such as urea or guanidine hydrochloride. Low concentrations of reducing agents (<1 mM dithiothreitol or <5
mM 2-mercaptoethanol) may be used during purification, but higher concentrations will prematurely activate the intein self-cleavage reaction. Low concentrations of nonionic detergents (i.e., 0.2% Triton-X or Tween 20) may also be used depending on the nature of the fusion protein. The intein self-cleavage reaction is induced by overnight incubation with 50 mM DTT at 4°C. 2-mercaptoethanol, cysteine, or hydroxylamine may also be used but are less effective (Chong et al., 1997), with the latter two forming permanent complexes with the cleaved protein. The production of proteins possessing an N-terminal cysteine and/or C-terminal thioester can be useful for protein labeling, ligation, or cyclization (Chong et al., 1997). Typical yields from intein-CBD expression and purification are ∼0.5 to 5 mg cleaved protein per liter bacterial culture.
Biotin Carboxyl Carrier Protein In vivo biotinylation of heterologous proteins containing a biotinylation signal peptide (BCCP) was first reported in 1990 (Cronan, 1990). The PinPoint Xa vectors provide all three reading frames and a factor Xa cleavage site (Promega). Typical yields from the PinPoint expression and purification system are in the range of 1 to 5 mg fusion protein per liter bacterial culture. Biotinylated proteins can be purified by affinity chromatography on avidin resin, although purification by traditional avidin resin requires a denaturing elution (e.g., heat, urea, or guanidine hydrochloride). Commercially available resins (i.e., SoftLink avidin resin, Promega) can be regenerated at least ten times and allow elution of biotinylated fusion proteins with a mild 10 mM biotin buffer. Denaturing reagents are not compatible with avidin affinity chromatography. Low concentrations of reducing agents or nonionic detergents may be used depending on the nature of the fusion protein. There is little nonspecific binding to avidin in E. coli systems. Native BCCP does not bind to avidin; however, mammalian systems have at least four biotinylated protein species that can co-elute with the biotinylated protein of interest (Hollingshead et al., 1997). Biotinylated proteins can be detected by alkaline phosphatase (AP)- or horseradish peroxidase (HRP)-coupled streptavidin, primary antibodies against biotin, or labeling with fluorescent or radioactive biotin. Biotin precipitation assays with streptavidin-coupled beads (UNIT 9.7)
Affinity Purification
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can be used for the detection of protein-protein or protein-DNA interactions. In addition, the high-affinity interaction of biotin and streptavidin makes the biotin tag useful for the immobilization of fusion proteins on streptavidincoated surfaces, such as surface plasmon resonance chips (Sensor Chip SA, Biacore). In addition to the in vivo biotinylated BCCP tag, other purification systems utilize the highaffinity interaction between biotin and streptavidin or avidin. Streptavidin-binding peptide (SBP) is 38 residues in length and can be used to immobilize fusion proteins on a streptavidin matrix. In addition, the Strep-Tag and StrepTag II (Sigma) are eight- to nine-residue peptides that bind a mutant form of streptavidin (Strep-Tactin). Because the affinity of biotin for streptavidin is higher than that of any of the streptavidin-binding peptides described above, streptavidin-bound SBP-tagged or Streptagged fusion proteins can be efficiently eluted by free biotin. Streptavidin itself can also be used as an affinity tag for purification or immobilization of fusion proteins on a biotinylated matrix.
Epitope Tags Relatively short epitope tags such as FLAG, hemagglutinin (HA), c-myc, T7, and Glu-Glu, among others (Table 9.9.1), are used for the detection of fusion proteins in vitro and in cell culture. Their short, linear recognition motifs rarely affect the properties of the heterologous protein of interest and are usually very specific for their respective primary antibodies. One exception is the anti-myc antibody, which is somewhat promiscuous; however, specificity can be increased by using an enzyme-linked secondary antibody to detect a conjugated antimyc primary antibody, instead of using an HRP- or AP-anti-myc conjugate alone. Epitope-tagged proteins can be purified using immobilized primary antibodies to the epitope tag although antibody affinity chromatography often involves a low or high pH elution which can irreversibly affect the properties of the fusion protein, and employs resin which has limited reusability. Epitope tags are therefore not the first choice when the main goals are high-level expression and fusion protein purification.
Reporter Tags Overview of Affinity Tags for Protein Purification
Enzymes such as LacZ, alkaline phosphatase (AP), and chloramphenicol acetyl transferase (CAT) serve as convenient reporters of protein expression and protein-protein inter-
action. These reporter tags can be used for protein purification, but are frequently used as reporter tags for detecting protein expression. β-galactosidase LacZ interacts with its substrate, 5-bromo4-chloro-3-indoyl-β-D-galactopyranoside (Xgal), to yield a blue precipitate. The advantages and disadvantages of lacZ as a protein purification tag have been discussed above (see Early Affinity Tags). Alkaline phosphatase AP interacts with 5-bromo-4-chloro-3-indolyl phosphate/nitro blue tetrazolium (BCIP/NBT) to yield a purple precipitate. AP is commonly fused to primary antibodies for immunoblotting but can also be fused to any number of nonimmune proteins for far-western blotting (i.e., detection of a membrane-transferred protein by interaction with another protein; UNIT 19.7). Since AP is normally targeted to the E. coli periplasmic space, crude AP fusion protein lysates can be purified by periplasmic extraction (UNIT 5.2). AP fusion proteins can be further purified by immunoaffinity chromatography. Chloramphenicol acetyl transferase The ease and sensitivity of the assay for CAT activity made CAT one of the first reporter genes used for studies of mammalian gene expression (Podbielski et al., 1992). The CAT gene is not found in eukaryotes, and therefore eukaryotic cells contain no background CAT activity, which can be detected using the radioisotope-labeled or fluorescently-labeled substrates acetyl-CoA or chloramphenicol. CAT fusion proteins can be purified by affinity chromatography on immobilized chloramphenicol (i.e., chloramphenicol caproate agarose, Sigma) and eluted by free chloramphenicol.
DEVELOPING AFFINITY TAG TECHNOLOGIES Some newer affinity tags have yet to be rigorously evaluated and consequently cannot be recommended above the tried-and-true affinity tags described above (see Commonly Used Affinity Tags). Still, many of these novel affinity tags contain unique properties that may make them quite useful in cases of troublesome purifications.
His-Patch ThioFusion Thioredoxin has the ability to accumulate to ∼40% of the total cellular protein in E. coli. In
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large quantity (>1 mg) of high purity, untagged protein
cleavable His-tag
expression tests
insoluble, His-tagged protein
cleavable GSTor MBP-tag
extract
soluble, His-tagged protein
optimize expression
refold
soluble GSTor MBP-tagged protein
insoluble GSTor MBP-tagged protein
new technology: His-patch thloredoxin?
Figure 9.9.1 Flow chart describing the general scheme for choosing an affinity tag for protein purification if a large amount (>1 mg) of highly pure untagged protein is required. Experimental methods requiring this high quantity and purity of native protein include, but are not limited to, protein crystallography (UNITS 17.3 & 17.4), NMR spectroscopy (UNIT 17.5), isothermal titration calorimetry (UNIT 20.4), and surface plasmon resonance. Solid lines indicate forward progression in purification optimization, dashed lines indicate steps backward, and thick boxes indicate the final desired product, a protein of high purity and quality.
addition, the thioredoxin moiety can in some cases confer solubility to formerly insoluble heterologous proteins expressed in E. coli. Unfortunately, the phenylarsine oxide matrix used to purify thioredoxin fusion proteins does not provide high-yield, high-purity product (Terpe, 2003). To address this problem, Invitrogen developed a mutant thioredoxin molecule that has a cluster of histidines, such that when the thioredoxin molecule is properly folded, the histidines form a patch amenable to IMAC (His-patch ThioFusion expression system; see Polyhistidine). In addition, the tag is cleavable by Factor Xa, allowing the fusion protein to be separated from the His-Patch ThioFusion molecule, which is >100 residues in length and readily dimerizes in solution. While the cost of this expression system (>$600.00) might discourage its use as a first-line affinity tag, the HisPatch ThioFusion tag might be useful when
other attempts at producing large amounts of soluble protein have failed.
NorpA The type II N-terminal PDZ domain (PDZ1) of InaD binds the C-terminus of no receptor potential A (NorpA), the relevant phospholipase C β isozyme in the Drosophila phototransduction pathway. The crystal structure of PDZ1 in complex with a peptide corresponding to the NorpA C-terminus shows that only the last five residues of NorpA contact PDZ1 and that a disulfide bond is the major intermolecular interaction (Kimple et al., 2001). The short PDZ1 binding motif of NorpA, coupled with its covalent yet dissociable interaction, led to the hypothesis that the NorpA C-terminal residues (Thr-Glu-Phe-Cys-Ala) could be used as an affinity tag for protein detection and purification by appropriately modified PDZ1
Affinity Purification
9.9.5 Current Protocols in Protein Science
Supplement 36
medium quantity 0.1–1.0 mg purity tagged or untagged protein
tagged only
both tagged and untagged
untagged only
Is detection of tag important?
BCCP tag
intein-CBD tag
yes
no
HIS-tag
GST-tag
Figure 9.9.2 Flow chart describing general scheme for selecting an affinity tag for protein purification if a mid-range amount (100 µg to 1 mg) of highly pure tagged and/or untagged protein is needed. An example of an experimental method requiring this quantity of tagged protein is surface plasmon resonance (SPR), where a biotin moiety could be used to attach a protein ligand to a streptavidin-coated SPR chip. Other possible applications are in GST and Ni2+ precipitation assays, and mass spectrometry. Certain variants of the intein-CBD tag are useful for methods requiring a completely native protein, as it is possible to cleave off every intein residue from the fusion protein, leaving a native N or C terminus. Finally, the His- and GST-tags are useful because both tagged and untagged proteins can easily be, prepared during the same protein preparation, and there are many secondary reagents, such as antibodies, resins, SPR chips, and ELISA assays that are based on these fusion systems. Thick lines indicate the desired result has been reached.
small quantity (< 100 µg ) of moderate-tohigh purity tagged protein Will tag be used as a reporter?
Overview of Affinity Tags for Protein Purification
yes
no
AP or CAT
Epitope tag
Figure 9.9.3 Flow chart describing general scheme for selecting an affinity tag for purification of a small amount (<100 µg) of moderate-to-highly pure tagged protein—e.g., for co-immunoprecipitation, western blotting (i.e., immunoblotting), far-western blotting, ELISA, gel overlay assays. Since only small amounts of protein are required, it is usually not necessary to optimize protein expression, leaving the choice of affinity tag to be based on the final use for the fusion tag—i.e., as a reporter or for general detection. Examples of epitope tags include the T7-tag, B-tag, Glu-Glu, HA, and c-Myc.
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(Kimple and Sondek, 2002). In fact, PDZ1 specifically recognizes not only its physiological ligand, NorpA, but also a heterologous protein expressing the NorpA C-terminal five residues. Because of its short linear recognition sequence and its specific interaction with PDZ1, which is both covalent and dissociable, the NorpA tag may improve upon most currently available affinity tags. Optimization of the purification protocols, such as addition of a flexible linker to PDZ1 and different coupling chemistries, could yield a moderately reusable, high affinity resin comparable to glutathione Sepharose. This would make the NorpA C-terminus one of the most useful affinity tags: comparable with the His6 tag in length but potentially more useful for protein detection because of the specific high affinity interaction of NorpA with PDZ1. Specifically, the covalent interaction between PDZ1 and NorpA is ideal for protein-protein interaction techniques such as surface plasmon resonance, where strong attachment of the protein of interest to a biosensor chip is required, but covalent attachment of proteins via primary amino groups can sometimes result in loss of biological activity. A PDZ1 surface would “catch” NorpA-tagged proteins, which would then be analyzed for their ability to bind various ligands. The entire surface could be regenerated by exposure to a reducing buffer, allowing the covalent attachment of a different NorpAtagged protein for a new set of binding experiments.
AFFINITY TAGS AND PROTEIN PURIFICATION Figures 9.9.1, 9.9.2, and 9.9.3 give general schemes for selecting an affinity tag based on three different desired protein purification outcomes: a large quantity of highly pure, untagged protein (Fig. 9.9.1), a medium quantity of highly pure tagged or untagged protein (Fig. 9.9.2), and a small amount of moderate-tohighly pure tagged protein (Fig. 9.9.3). These flow charts should be used as an initial guide but more difficult protein purification problems may require the use of a system different than those recommended.
LITERATURE CITED Berlot, C.H. 1999. Expression and functional analysis of G-protein alpha subunits in mammalian cells. In G-proteins: Techniques of Analysis (D.R. Manning, ed.) pp. 37-57. CRC Press, New York.
Bornhorst, J.A. and Falke, J.J. 2000. Purification of proteins using polyhistidine affinity tags. Methods Enzymol. 326:245-254. Chong, S., Mersha, F.B., Comb, D.G., Scott, M.E., Landry, D., Vence, L.M., Perler, F.B., Benner, J., Kucera, R.B., Hirvonen, C.A., Pelletier, J.J., Paulus, H., and Xu, M.Q. 1997. Single-column purification of free recombinant proteins using a self-cleavable affinity tag derived from a protein splicing element. Gene 192:271-281. Crespo, P., Schuebel, K.E., Ostrom, A.A., Gutkind, J.S., and Bustelo, X.R. 1997. Phosphotyrosinedependent activation of Rac-1 GDP/GTP exchange by the vav proto-oncogene product. Nature 385:169-172. Cronan, J.E. Jr. 1990. Biotination of proteins in vivo. A post-translational modification to label, purify, and study proteins. J. Biol. Chem. 265:1032710333. di Guan, C., Li, P., Riggs, P.D., and Inouye, H. 1988. Vectors that facilitate the expression and purification of foreign peptides in Escherichia coli by fusion to maltose-binding protein. Gene 67:2130. Fritze, C.E. and Anderson, T.R. 2000. Epitope tagging: General method for tracking recombinant proteins. Methods Enzymol. 327:3-16. Gerdes, H.H. and Kaether, C. 1996. Green fluorescent protein: Applications in cell biology. FEBS Lett. 389:44-47. Goldstein, D.J., Toyama, R., Dhar, R., and Schlegel, R. 1992. The BPV-1 E5 oncoprotein expressed in Schizosaccharomyces pombe exhibits normal biochemical properties and binds to the endogenous 16-kDa component of the vacuolar proton-ATPase. Virology 190:889-893. Hollingshead, M., Sandersen, J., and Vaux, D.J. 1997. Anti-biotin antibodies offer superior organelle-specific labeling of mitochondria over avidin or streptavadin. J. Histochem. Cytochem. 45:1053-1058. Jones, C., Patel, A., Griffin, S., Martin, J., Young, P., O’Donnell, K., Silverman, C., Porter, T., and Chaiken, I. 1995. Current trends in molecular recognition and bioseparation. J. Chromatogr. 707:3-22. Kaldalu, N., Lepik, D., Kristjuhan, A., and Ustav, M. 2000. Monitoring and purification of proteins using bovine papillomavirus E2 epitope tags. Biotechniques 28:456-462. Karp, M. and Oker-Blom, C. 1999. A streptavidinluciferase fusion protein: Comparisons and applications. Biomol. Eng. 16:101-104. Kimple, M.E., Siderovski, D.P., and Sondek J. 2001. Functional relevance of the disulfide-linked complex of the N-terminal PDZ domain of InaD with NorpA. EMBO J. 20: 4414-4422. Kimple, M.E. and Sondek, J. 2002. Affinity tag for protein purification and detection based on the disulfide-linked complex of InaD and NorpA. Biotechniques 33:578-588. Affinity Purification
9.9.7 Current Protocols in Protein Science
Supplement 36
Kolodziej, P.A. and Young, R.A. 1991. Epitope tagging and protein surveillance. Methods Enzymol. 194:508-519. Kuliopulos, A. and Walsh, C.T. 1994. Production, purification, and cleavage of tandem repeats of recombinant peptides. J. Am. Chem. Soc. 116:4599-4607. Kwatra, M.M., Schreurs, J., Schwinn, D.A., Innis, M.A., Caron, M.G., and Lefkowitz, R.J. 1995. Immunoaffinity purification of epitope-tagged human beta 2-adrenergic receptor to homogeneity. Protein Expr. Purif. 6:717-721. Lazzaroni, J.C., Atlan, D., and Portalier, R.C. 1985. Excretion of alkaline phosphatase by Escherichia coli K-12 pho constitutive mutants transformed with plasmids carrying the alkaline phosphatase structural gene. J. Bacteriol. 164:1376-1380. Lilius, G., Persson, M., Bulow, L., and Mosbach, K. 1991. Metal affinity precipitation of proteins carrying genetically attached polyhistidine affinity tails. Eur. J. Biochem. 198:499-504. Ljungquist, C., Breitholtz, A., Brink-Nilsson, H., Moks, T., Uhlen, M., and Nilsson, B. 1989. Immobilization and affinity purification of recombinant proteins using histidine peptide fusions. Eur. J. Biochem. 186:563-569. Maina, C.V., Riggs, P.D., Grandea, A.G. III, Slatko, B .E., Mora n, L.S., Ta gliam onte, J .A., McReynolds, L.A., and Guan, C.D. 1988. An Escherichia coli vector to express and purify foreign proteins by fusion to and separation from maltose-binding protein. Gene 74:365-373.
Rubinfeld, B., Munemitsu, S., Clark, R., Conroy, L., Watt, K., Crosier, W.J., McCormick, F., and Polakis, P. 1991. Molecular cloning of a GTPase activating protein specific for the Krev-1 protein p21rap1. Cell 65:1033-1042. Sambrook, J. and Fritsch, E.F. 1989. Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Sano, T., Vajda, S., and Cantor, C.R. 1998. Genetic engineering of streptavidin, a versatile affinity tag. J. Chromatogr. B. Biomed. Appl. 715:85-91. Skerra, A. and Schmidt, T.G. 2000. Use of the StrepTag and streptavidin for detection and purification of recombinant proteins. Methods Enzymol. 326:271-304. Smith, D.B. 2000. Generating fusions to glutathione S-transferase for protein studies. Methods Enzymol. 326:254-270. Smith, D.B. and Johnson, K.S. 1988. Single-step purification of polypeptides expressed in Escherichia coli as fusions with glutathione Stransferase. Gene 67:31-40. Smith, M.C., Furman, T.C., Ingolia, T.D., and Pidgeon, C. 1988. Chelating peptide-immobilized metal ion affinity chromatography. A new concept in affinity chromatography for recombinant proteins. J. Biol. Chem. 263:7211-7215.
Makrides, S.C. 1996. Strategies for achieving highlevel expression of genes in Escherichia coli. Microbiol. Rev. 60:512-538.
Stevens, R.C. 2000. Design of high-throughput methods of protein production for structural biology. Structure Fold Des. 8:R177-R185.
McLean, P.J., Kawamata, H., and Hyman, B.T. 2001. Alpha-synuclein-enhanced green fluorescent protein fusion proteins form proteasome sensitive inclusions in primary neurons. Neuroscience 104:901-912.
Tai, T.N., Havelka, W.A., and Kaplan, S. 1988. A broad-host-range vector system for cloning and translational lacZ fusion analysis. Plasmid 19:175-188.
Morandi, C., Perego, M., and Mazza, P.G. 1984. Expression of human dihydrofolate reductase cDNA and its induction by chloramphenicol in Bacillus subtilis. Gene 30:69-77. Nelson, R.W., Jarvik, J.W., Taillon, B.E., and Tubbs, K.A. 1999. BIA/MS of epitope-tagged peptides directly from E. coli lysate: Multiplex detection and protein identification at low-femtomole to subfemtomole levels. Anal. Chem. 71:28582865. Nilsson, J., Bosnes, M., Larsen, F., Nygren, P.A., Uhlen, M., and Lundeberg J. 1997a. Heat-mediated activation of affinity-immobilized Taq DNA polymerase. Biotechniques 22:744-751. Nilsson, J., Stahl, S., Lundeberg, J., Uhlen, M., and Nygren, P.A. 1997b. Affinity fusion strategies for detection, purification, and immobilization of recombinant proteins. Protein Expr. Purif. 11:116. Overview of Affinity Tags for Protein Purification
Podbielski, A., Peterson, J.A., and Cleary, P. 1992. Surface protein-CAT reporter fusions demonstrate differential gene expression in the vir regulon of Streptococcus pyogenes. Mol. Microbiol. 6:2253-2265.
Terpe, K. 2003. Overview of tag protein fusions: From molecular and biochemical fundamentals to commercial systems. Appl. Microbiol. Biotechnol. 60:523-533. Thain, A., Gaston, K., Jenkins, O., and Clarkie, A.R. 1996. A method for the separation of GST fusion proteins from co-purifying GroEL. Trends Genet. 12:209-210. Wang, L.F., Yu, M., White, J.R., and Eaton, B.T. 1996. BTag: A novel six-residue epitope tag for surveillance and purification of recombinant proteins. Gene 169:53-58.
Contributed by Michelle E. Kimple Duke University Medical Center Durham, North Carolina John Sondek University of North Carolina Chapel Hill Chapel Hill, North Carolina
9.9.8 Supplement 36
Current Protocols in Protein Science
9.9.9
Current Protocols in Protein Science
Supplement 36
6 (TDFYLK)
AU5
N-term or C-term
N-term or C-term
C-term
6 (DTYRYI)
AU1
C-term
Bacteriophage 14 V5 epitope (GKPIPNPL (V5-tag) LGLDST)
444
AP
N-term or C-term
N-term or internal
137
ABP
Position
Bacteriophage 11 T7 epitope (MASMTGGQ (T7-tag) QMG)
Lengthc (sequence)
NA
mAb/low pH
mAb/low pH
mAb/low pH
mAb/low pH
Albumin/low pH or denaturation (e.g., heat, urea)
Matrix/elution
Characteristics of Protein Affinity Tagsa,b
Tag
Table 9.9.1
Selected pET directional TOPO, pBAD, and Gateway systems (Invitrogen)
T7-tag affinity purification kit (Novagen), pGEMEX vectors (Promega)
NA
NA
PhoA* color system (Qbiogene); gWIZ secreted AP expression vectors (Gene Therapy Systems)
NA
Commercial systems (supplier)
Detection
Purification and increased expression
Detection and purification
Detection and purification
Detection
Purification, increased expression, increased solubility
Typical use(s)
Lazzaroni et al. (1985)
Nilsson et al. (1997a)
References
Short, linear recognition motif; antibody specific in bacterial lysates, although some cross-reactivity in mammalian lysates; often used in combination with His-tag for protein purification
N-term eleven amino acids of phage T7 gene 10; may increase expression of fusion proteins; antibody purification does not give high yields; low pH elution may irreversibly affect protein properties; matrix is of limited reusability
continued
McLean et al. (2001)
Makrides (1996)
Antibody purification does not give high yields; Crespo et al. low pH elution may irreversibly affect protein (1997) properties; matrix is of limited reusability
Antibody purification does not give high yields; Goldstein et low pH elution may irreversibly affect protein al. (1992) properties; matrix is of limited reusability
Colorimetric detection; useful for far-western blotting, southern blotting, protein quantification, sandwich ELISA, and subcellular localization in bacterial and mammalian expression systems; convenient purification of crude periplasmic extract from bacteria; antibody purification does not give high yields, low pH elution may irreversibly affect protein properties; matrix is of limited reusability; AP dimerization and very large size may affect properties of fusion
May increase proteolytic stability of fusion proteins to increase expression; may improve fusion solubility; high background binding to albumin in eukaryotic systems; matrix has limited reusability; large tag or low pH elution conditions may affect fusion properties
Comments
9.9.10
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Current Protocols in Protein Science
Cellulose 27–189 binding domain (CBD)
218
N-term, C-term, or internal (domaindependent)
N-term
N-term or C-term
CAT
6 (QYPALT)
B-tag
N-term or C-term
N-term or C-term
100
BCCP
Position
Calmodulin 26 binding peptide (CBP)
Lengthc (sequence)
gWIZ CAT mammalian expression vector (Gene Therapy Systems)
pCAL system (Stratagene)
NA
PinPoint system (Promega)
Commercial systems (supplier)
Cellulose/denatura- pET CBD Fusion tion (family I System (Novagen) CBDs) or ethylene glycol (family II or III CBDs)
Chloramphenicol/ chloramphenicol
Calmodulin/EGTA or EGTA, and high salt
mAb/low pH
Avidin or streptavidin/biotin or denaturation (e.g., heat, urea)
Matrix/elution
Characteristics of Protein Affinity Tagsa,b, continued
Tag
Table 9.9.1
Purification and immobilization
Detection, purification, and increased solubility
Purification
Detection and purification
Detection, purification, and immobilization
Typical use(s)
Irreversible binding of some CBDs to cellulose useful for immobilization; reversible binding of CBD I, II, and III families useful for purification; purification of family I CBD fusion necessitates protein refolding; purified family II CBD fusions must be buffer exchanged
Enzymatic assay available for protein quantification; chloramphenicol purification does not give high yields; large tag may affect properties of fusion protein
Relatively short recognition motif; no endogenous E. coli proteins that bind calmodulin; PKA target sequence allows 32P-labelling; high yield matrix is compatible with reducing agents and detergents, but of limited reusability; not useful for purification from eukaryotic cells; tag at N terminus may reduce translation efficiency
VP7 region of bluetongue virus; antibody purification does not give high yields; low pH elution may irreversibly affect protein properties; matrix is of limited reusability
Tag is biotinylated in vivo; one-step purification of fusion protein; protein may be secreted for convenient purification; biotin tag can be used to immobilize fusion to streptavidin-coated surfaces—e.g., surface plasmon resonance (SPR) chips; high background binding to avidin in eukaryotic systems; elution with biotin may be inefficient; denaturing elution requires refolding of fusion protein; Promega SoftLink avidin, part of the PinPoint system, allows elution under mild conditions
Comments
continued
Terpe (2003)
Podbielski et al. (1992)
Terpe (2003)
Wang et al. (1996)
Nilsson et al. (1997b)
References
9.9.11
Current Protocols in Protein Science
Supplement 36
N-term or C-term
227
10 (SSTSSDFR DR)
DHFR
E2-tag
N-term, C-term, or internal
N-term
Choline-bindin 145 g domain (CBD)
Position N-term or C-term
Lengthc (sequence) Commercial systems (supplier) Typical use(s)
NA
mAb/low pH
E2-Tagging and Detection Kit (Abcam), E2 tagging technology (Quattromed)
Methotrexate/folate pQE vector set (Qiagen)
Choline/choline or CBD peptide
Detection and purification
Increased expression
Purification and immobilization
Chitin (when fused IMPACT system (New Purification with England Biolabs) intein)/thiol-containi ng reducing agent (e.g., DTT or β-ME)
Matrix/elution
Characteristics of Protein Affinity Tagsa,b, continued
Chitin 51 binding domain (CBD)
Tag
Table 9.9.1
Derived from bovine papilloma-virus type-1 transactivator protein E2; primary antibodies available for colorimetric and chemiluminescent detection; antibody purification does not give high yields, low pH elution may irreversibly affect protein properties; matrix is of limited reusability
May increase proteolytic stability of fusion proteins to increase expression; little immunogenicity in mouse and rat, therefore, useful for generation of antibodies to fusion; methotrexate purification does not give high yields; usually coupled with a short affinity tag (e.g., His-tag, Strep-tag) for purification
N-term CBD can immobilize fusion protein on choline-coated gold chip for SPR or microscopy studies; large tag or C-term tag placement may affect fusion properties
Typically used in conjunction with self-splicing intein tag; one-step purification of nearly 100% pure protein with low milligram yields; matrix compatible with high salt and nonionic detergents; purification must be done in absence of thiol-containing reducing agents until elution step; fusion protein may have effect on intein cleavage efficiency
Comments
continued
Kaldalu et al. (2000)
Morandi et al. (1984)
Jones et al. (1995)
Terpe (2003)
References
9.9.12
Supplement 36
Current Protocols in Protein Science
8 (DYKDDDK)
509
220
6 (EYMPME or EFMPME)
GBP
GFP
Glu-Glu (EE-tag)
Lengthc (sequence)
N-term, C-term, or internal
N-term or C-term
N-term or C-term
N-term or C-term
Position
mAb/low pH or 30°C incubation
NA
Typical use(s)
NA
gWIZ GFP mammalian expression vector (Gene Therapy Systems); CT-GFP Fusion TOPO Cloning kit (Invitrogen)
Detection and purification
Detection
Purification and increased solubility
FLAG system (Sigma) Detection and purification
Commercial systems (supplier)
Galactose/galactose NA
mAb/Low pH, EDTA, or FLAG peptide
Matrix/elution
Characteristics of Protein Affinity Tagsa,b, continued
FLAG
Tag
Table 9.9.1
Short, linear recognition motif; available antibody recognizes only EYMPME motif; antibody purification does not give high yields, low pH or 30°C elution may irreversibly affect protein properties; matrix is of limited reusability
continued
Rubinfeld et al. (1991)
Gerdes and Kaether (1996)
Jones et al. (1995)
Fusion protein may be targeted to periplasm for convenient purification of crude periplasmic extracts; may increase solubility of fusion proteins; large tag may affect properties of fusion protein Intrinsic fluorescence can permit native detection without antibody; anti-GFP antibodies specific; expression in prokaryotic and eukaryotic expression systems, as well as whole organisms (e.g., worms, plants, mice); useful to monitor gene expression, protein folding, and targeting, as well as protein-protein interactions, by FRET; some GFP fusions nonspecifically targeted to nucleus; very large tag or GFP dimerization may affect properties of fusion
Terpe (2003)
References
Short, linear recognition motif; moderately pure protein in one step; enterokinase cleaves after C-term Lys to completely remove tag, depending on identity of first amino acid of fusion; M1 antibody can only bind tag at N-term; antibody purification does not give high yields; low pH elution may irreversibly affect protein properties; matrix is of limited reusability
Comments
9.9.13
Current Protocols in Protein Science
Supplement 36
Lengthc (sequence)
211
31
19 (KDHLIHNV HKEFHAH AHNK)
11 (QPELAPED PED)
125
GST
HA
HAT
HSV-tag
KSI
N-term
C-term
N-term or C-term
N-term, C-term, or internal
N-term or C-term
Position
Commercial systems (supplier)
NA
mAb/low pH
Divalent metal (Ni2+, Co2+, Cu2+, or Zn2+)/imidazole or low pH
mAb/low pH or HA peptide
pET-31b(+) vector (Novagen)
Increased expression
Purification
Purification
HAT protein expression and purification system (Clontech)
NA
Detection and purification
Detection, purification, increased expression, increased solubility, and immobilization
Typical use(s)
RTS pIVEX HA-tag vector set (Roche)
Glutathione/reduced pGEX system glutathione (Amersham Biosciences), GST-bind system (Novagen)
Matrix/elution
Characteristics of Protein Affinity Tagsa,b, continued
Tag
Table 9.9.1
References
Tai et al. (1988)
Insoluble protein is targeted to inclusion bodies; usually coupled with a short affinity tag (His-tag, Strep-tag) for purification; convenient denaturing purification of toxic proteins, but refolding necessary
C-term tag placement only; antibody purification does not give high yields; low pH elution may irreversibly affect protein properties; matrix is of limited reusability
continued
Kuliopulos and Walsh (1994)
Fritze and Anderson (2000)
Natural protein sequence that was found to Terpe (2003) chelate divalent metals; short recognition motif; contains six histidines interspersed among other amino acids; semipure protein in one step; matrix may be regenerated and reused almost infinitely; does not bind metal affinity resin as tightly as His-tag; tag or elution conditions may affect fusion properties
Anti-HA antibodies specific; useful in mammalian expression systems; antibody purification does not give high yields; low pH elution may irreversibly affect protein properties; matrix is of limited reusability
Very common purification tag; one-step Smith (2000) purification of relatively pure protein; detection antibodies specific; kits for GST fusion proteins common (e.g., SPR, enzymatic assays); matrix relatively reusable (four to twenty times); GST is highly antigenic; purification under native conditions only; some GST fusions insoluble; GST dimerization and/or glutathione elution may affect fusion protein properties
Comments
9.9.14
Supplement 36
Current Protocols in Protein Science
11 (KPPTPPPE PET)
1024
551
396
LacZ
Luciferase
MBP
Lengthc (sequence)
N-term or C-term
N-term
N-term or C-term
N-term or C-term
Position
Cross-linked amylose/maltose
NA
APTG/high pH borate buffer
mAb/low pH
Matrix/elution
Characteristics of Protein Affinity Tagsa,b, continued
KT3
Tag
Table 9.9.1
pMAL system (New England Biolabs), RTS pIVEX MBP fusion vector (Roche)
gWIZ luciferase mammalian expression vector (Gene Therapy Systems), luciferase assay system (Promega)
pMC1871 fusion vector (Amersham Biosciences), gWIZ β-galactosidase mammalian expression vector (Gene Therapy Systems)
NA
Commercial systems (supplier)
Detection, purification, increased expression, and increased solubility
One-step purification of relatively pure protein (>70%); matrix compatible with nonionizing detergents and high salt, but not reducing agents; can increase expression of eukaryotic proteins in bacteria; anti-MBP antibodies specific; tag at N-term can decrease translation efficiency; very large size of tag may affect fusion protein properties
continued
Nilsson et al. (1997b), Terpe (2003)
Karp and Luminescent; can serve as a reporter immediately upon translation; useful for studies Oker-Blom (1999) involving in situ hybridization, RNA processing, RNA transfection or coupled in vitro transcription/translation, protein folding, and imaging; can be labeled with 35S; no more than five codons can be removed from the N- or C-term to maintain enzymatic activity; very large tag may affect properties of fusion
Tai et al. (1988)
Also known as β-galactosidase or β-Gal; enzymatic assay available for protein quantification; may increase proteolytic stability of fusion proteins to increase expression; fusion proteins may be insoluble; extremely large tag which forms tetramers in solution, potentially affecting properties of fusion protein
Detection, purification, and increased expression
Detection
Kwatra et al. (1995)
References
Short, linear recognition motif; antibody purification does not give high yields; low pH elution may irreversibly affect protein properties; matrix is of limited reusability
Comments
Detection and purification
Typical use(s)
9.9.15
Current Protocols in Protein Science
Supplement 36
11 (CEQKLISE EDL)
5 (TEFCA)
495
5–6 (usually 5; RRRRR)
5–16 (DDDDD)
4 (CCCC)
NorpA
NusA
Polyarginine (Arg-tag)
Polyaspartate (Asp-tag)
Polycysteine (Cys-tag)
Lengthc (sequence)
N-term
C-term
C-term
N-term or C-term
C-term
N-term, C-term, or internal
Position pDual Expression System (Stratagene), PRO bacterial expression system (Clontech)
Commercial systems (supplier)
NA
NA
NA
Thiopropyl-Sepharo NA sethiol-containing reducing agent (e.g., DTT, β-ME)
Anion exchange resin/low-neutral pH salt gradient
Cation exchange resin/high pH salt gradient
NA
PDZ1/thiol-containi NA ng reducing agent (e.g., DTT, β-ME)
mAb/low pH
Matrix/elution
Characteristics of Protein Affinity Tagsa,b, continued
Myc
Tag
Table 9.9.1
Purification
Purification
Purification and immobilization
Increased expression and solubility
Detection, purification, and immobilization
Detection and purification
Typical use(s)
Terpe (2003)
Terpe (2003)
Kimple and Sondek (2002)
Kolodziej and Young (1991)
References
Short, linear recognition motif; moderately pure protein in one step; purification must be performed in absence of thiol-containing reducing agents until elution step; reducing elution may disrupt properties of fusion protein
continued
Stevens (2000)
Short, linear recognition motif; polar tag may Stevens (2000) affect tertiary structure of protein and/or protein properties
Can immobilize targets on mica for microscopy studies; short, linear recognition motif; very pure protein in one step; charged tag may affect tertiary structure of protein and/or protein properties; limited success of tag cleavage by carboxypeptidase B
Anti-transcription termination factor; increases solubility and expression of fusion proteins; must be used in conjunction with another affinity tag for protein purification; large tag may affect properties of fusion protein
Short, linear recognition motif; alkaline phosphatase-coupled PDZ1 allows antibody-independent detection; PDZ1-NorpA interaction highly specific; PDZ1 can couple fusion proteins to SPR resonance chip; purification must be performed in absence of thiol-containing reducing agents until elution step; reducing elution may disrupt properties of fusion protein; tag must be at C-term to bind PDZ1
Short, linear recognition motif; anti-myc antibody somewhat promiscuous; antibody purification does not give high yields; low pH elution may irreversibly affect protein properties; matrix is of limited reusability
Comments
9.9.16
Supplement 36
Current Protocols in Protein Science
9 (NANNPDWD F)
S1-tag
N-term or C-term
N-term or C-term
12
Protein C
N-term or C-term
Position
N-term
2–10 (usually 6; HHHHHH)
Lengthc (sequence)
NA
pXB, pBX, pXM, and pMX vectors (Roche)
mAb/Ca2+ buffer
mAb/low pH
NA
QIAexpress system (Qiagen), Selected pET directional TOPO, pBAD, and Gateway systems (Invitrogen)
Commercial systems (supplier)
Phenyl-Sepharose/ ethylene glycol
Divalent metal (i.e., Ni2+, Co2+, Cu2+, or Zn2+)/imidazole or low pH
Matrix/elution
Characteristics of Protein Affinity Tagsa,b, continued
Polyphenylalan 11 ine tag (FFFFFFFF (Phe-tag) FFF)
Polyhistidine (His-tag)
Tag
Table 9.9.1
Detection and purification
Detection and purification
Purification
Detection, purification and immobilization
Typical use(s)
Hepatitis B virus S1 region; short, linear recognition motif; AP1 antibody specific; has been tested in bacterial and mammalian expression systems; relatively pure protein in one step; antibody purification does not give high yields, low pH elution may irreversibly affect protein properties, and matrix is of limited reusability
Short, linear recognition motif; anti-PC antibody binds in Ca2+-dependent manner; elution by Ca2+ in physiological buffer conditions; antibody purification does not give high yields
Short, linear recognition motif; moderately pure protein in one step; nonpolar tag or ethylene glycol elution may disrupt properties of fusion protein
Most common purification tag; short, linear recognition motif; one-step purification of 20%–80% pure protein, depending on fusion protein expression levels; denaturing purification possible; matrix may be regenerated and reused indefinitely; can be used to immobilize fusion to Ni-NTA SPR chip, but significant dissociation complicates data analysis; Tag or elution may affect protein properties; detection antibodies highly promiscuous
Comments
continued
Berlot (1999)
Fritze and Anderson (2000)
Stevens (2000)
Bornhorst and Falke (2000)
References
9.9.17
Current Protocols in Protein Science
Supplement 36
N-term or C-term
8–9 (WSHPQFEK or AWAHPQPG G)
159
Strep-tag
Streptavidin
N-term or C-term
N-term or C-term
Staphylococcal 280 protein G (Protein G)
N-term, C-term or internal
Position
N-term
15 (KETAAAKF ERQHMDS)
Lengthc (sequence) S-Tag system (Novagen)
Commercial systems (supplier)
Biotin/biotin or denaturation (e.g., heat, urea)
Strep-Tactin (modified streptavidin)/biotin or desthiobiotin
Amylose/low pH or amylose
NA
Strep-tag II system (Sigma-Genosys), pASK75 vector (Biometra)
NA
IgG/Low pH or IgG pEZZ 18 and pRIT2T vectors (Amersham Biosciences)
S-fragment of RNase A/low pH
Matrix/elution
Characteristics of Protein Affinity Tagsa,b, continued
Staphylococcal 280 protein A (Protein A)
S-tag
Tag
Table 9.9.1
Detection, purification, increased expression, and immobilization
Detection, purification, and immobilization
Purification and increased solubility
Purification and increased solubility
Detection and purification
Typical use(s)
Skerra and Schmidt (2000)
Nilsson et al. (1997b)
Nilsson et al. (1997b)
Fritze and Anderson (2000)
References
continued
May increase proteolytic stability of fusion Sano et al. proteins to increase expression; extremely high (1998) affinity for biotin useful for immobilization of fusion on surfaces such as SPR chips; large size or tetramer formation may disrupt properties of fusion protein; fusion protein may not be released upon addition of free biotin, necessitating denaturing elution followed by refolding; newer streptavidin mutants that have lower affinities for biotin useful for purification
Short, linear recognition motif; matrix regenerable; useful for purification under anaerobic conditions, eukaryotic cell surface display, and immobilization to streptavidin-coated surfaces (e.g., SPR chips); specific binding conditions may be unsuitable for some fusions
Proteolytically stable; may increase solubility of fusion; fusion proteins secreted; purification does not give high yields; large tag size and/or low pH elution may irreversibly affect protein properties; matrix is of limited reusability
Proteolytically stable; may increase solubility of fusion; fusion proteins secreted; purification does not give high yields; large tag size and/or low pH elution may irreversibly affect protein properties; matrix is of limited reusability
Short, linear recognition motif; RNase S assay possible for quantitative assay of expression levels; colorimetric assays used for detection without antibody; tag or low pH elution may irreversibly affect protein properties; matrix is of limited reusability
Comments
9.9.18
Supplement 36
Current Protocols in Protein Science
Lengthc (sequence)
38
260
109
25–336
76
6 (HTTPHH)
SBP
T7
Trx
TrpE
Ubiquitin
Universal
N-term or C-term or internal
N-term
N-term or C-term
N-term or C-term
N-term
C-term
Position
NA
NA
Commercial systems (supplier)
mAb/low pH
NA
mAb/low pH
NA
NA
NA
ThioFusion System Phenylarsinine oxide/thiol-containin (Invitrogen) g reducing agent (e.g. DTT, β-ME)
mAb/low pH
Streptavidin/biotin
Matrix/elution
Characteristics of Protein Affinity Tagsa,b, continued
Tag
Table 9.9.1
Detection and purification
Increased solubility
Purification and increased expression
Increased solubility
Purification and increased expression
Purification and immobilization
Typical use(s)
References
Stevens (2000)
Terpe (2003)
Sequence HTTPHH is translated regardless of reading frame for ease in cloning; multiple tag copies increase antibody specificity; immobilized mAb can bind multiple tag copies in SPR studies; antibody purification does not give high yields; low pH elution may irreversibly affect protein properties; matrix is of limited reusability
continued
Nelson et al. (1999)
May increase solubility of proteins expressed in Stevens (2000) E. coli; not useful for expression in eukaryotic cells.
Larger constructs expressed may be targeted to inclusion bodies (allowing high-level expression of toxic genes; antibody purification does not give high yields; low pH elution may irreversibly affect protein properties; matrix is of limited reusability; large tag may affect properties of fusion protein
Heat stable; may increase solubility of fusion proteins; convenient purification of crude periplasmic extract from bacteria; purification must be done in absence of thiol-containing reducing agents until elution step; large tag or elution conditions may affect properties of fusion protein
May increase expression of fusion proteins; Stevens (2000) insoluble protein is targeted to inclusion bodies; denaturing purification of toxic proteins necessitates refolding
Relatively short recognition motif; Terpe (2003) immobilization of protein to various media (e.g., streptavidin-coated beads, SPR chips); tag at C-term only
Comments
Current Protocols in Protein Science
11 (YTDIEMNR LGK)
Lengthc (sequence) C-term
Position mAb/low pH
Matrix/elution
Characteristics of Protein Affinity Tagsa,b, continued
pVB6, pBV, pVM6, and pMV vector set (Roche)
Commercial systems (supplier) Detection and purification
Typical use(s)
C-term residues of VSV-G; relatively pure protein in one step; antibody purification does not give high yields; low pH elution may irreversibly affect protein properties; matrix is of limited reusability
Comments
Fritze and Anderson (2000)
References
cSequence lengths are reported in amino acids.
bAdapted, with permission, from BioMedicalPDA (http://www.biomedicalpda.com), Professional PDA Publishing, LLC, Larchmont, N.Y.
aAbbreviations: ABP, albumin-binding protein; AP, alkaline phosphatase; APTG, p-amino-phenyl-beta-D-thiogalactosidase; β-ME, 2-mercaptoethanol; BCCP, biotin carboxyl carrier protein; CA, chloramphenicol acetyl transferase; C-term, C terminal; DHFR, dihydrofolate reductase; DTT, dithiothreitol; FRET, fluorescence resonance energy transfer GBP; galactose-binding protein; GFP, green fluorescent protein; GST, glutathione S-transferase; HA, hemagglutinin; HAT, histidine-affinity tag; HSV, Herpes simplex virus peptide; KSI, ketosteroid isomerase; mAb, monoclonal antibody; MBP, maltose-binding protein; NA, not applicable; N-term, N terminal; SBP, streptavidin-binding peptide; T7-tag, T7 gene 10 tag; Trx, thioredoxin; VSV-G, vesicular stomatitis virus glycoprotein peptide.
VSV-G
Tag
Table 9.9.1
Affinity Purification
9.9.19
Supplement 36
CHAPTER 10 Electrophoresis INTRODUCTION
E
lectrophoresis of protein samples in polyacrylamide gels is an indispensable analytical and, in some cases, preparative tool for the protein scientist. Electrophoresis can be used to separate and compare complex protein mixtures, evaluate purity of a protein during the course of its isolation, and provide estimates of physical characteristics such as subunit composition, isoelectric point, size, and charge. Each type of electrophoretic separation can be conducted in a variety of gel sizes ranging from microgels (e.g., Phast gels from Hoefer Pharmacia) only slightly larger than a postage stamp to giant gels much larger than this page (Garrels, 1979; Young et al., 1983). In general, the time required to electrophorese, stain, and destain small gels is very short, and a minimal amount of sample is consumed. In contrast, larger gels consume more reagents, sample, and time, but provide increased resolution. Small gels are therefore recommended for rapid screening, and larger gels are indicated when maximum resolution is required, as in analysis of complex mixtures or samples containing very similar components.
The most common one-dimensional gel methods utilize the detergent sodium dodecyl sulfate (SDS) to solubilize, denature, and impart a strong negative charge to proteins (UNIT 10.1). Although many one-dimensional SDS gel methods have been published over the past 30 years, the single most widely used method is that initially described by Laemmli (1970). SDS-based gel separations are very robust as most, but not all, proteins are readily solubilized in SDS solutions. Most proteins bind a uniform amount of SDS per microgram of protein, which imparts a uniform charge density per unit mass to provide a separation based on the mass of the polypeptide chain. An alternative one-dimensional separation methodology is based on the protein’s isoelectric point and is usually performed in the presence of denaturants such as urea (UNIT 10.2). Like one-dimensional SDS gel separations, this denaturing isoelectric focusing method is capable of separating a large number of components in a single dimension. A third one-dimensional method uses native conditions to separate proteins based on intrinsic charge (UNIT 10.3). Although this method has less resolving power than the previous two methods, it can be particularly useful for associating a specific electrophoretically separated component with a biological activity, since no denaturants are used. Two-dimensional electrophoresis (UNIT 10.4) involves the orthogonal combination of two different electrophoretic methods. Any two electrophoresis techniques can be combined in this manner to produce useful separations. The most common two-dimensional approach combines two individually high-resolution methods, consisting of isoelectric focusing (UNIT 10.2) followed by SDS gel electrophoresis (UNIT 10.1). Each of these methods is individually capable of resolving up to about 100 protein bands; when the two are combined, more than 1,000 proteins can be separated on a single two-dimensional gel (O’Farrell, 1975; Garrels, 1979). Hence this method can be used to analyze and compare complex protein mixtures, including whole-cell or tissue extracts. One important application of two-dimensional gels is the systematic analysis and quantitative comparison of whole-cell or tissue extracts for the study of the proteome, or complete protein profile, of Electrophoresis Contributed by David W. Speicher Current Protocols in Protein Science (1999) 10.0.1-10.0.3 Copyright © 1999 by John Wiley & Sons, Inc.
10.0.1 Supplement 17
the cell or tissue. In recent years a growing emphasis has been focused on such studies, particularly for organisms whose complete genome sequence has been determined. Other applications of two-dimensional electrophoresis include assessment of the purity and homogeneity of a purified protein, including recombinant proteins (Chapter 7), and evaluation of some post-translational modifications (Chapters 12 and 13). After electrophoretic separation, proteins are usually detected using general protein staining methods, such as Coomassie blue staining or the more sensitive silver staining. These staining methods “fix” the proteins in the gel matrix by either chemical cross-linking or denaturation to prevent subsequent diffusion of the protein bands (UNIT 10.5). When proteins are to be eluted or extracted from the gels for subsequent analyses, however, fixation-based stains should usually be avoided. In these cases, methods for protein detection without fixation should be used (UNIT 10.6) when it is necessary to stain a gel to locate the protein of interest. A useful method of isolating proteins from gels for subsequent analysis involves electroblotting onto an inert support, such as polyvinylidene difluoride (PVDF) membranes (UNIT 10.7). As a result of the versatility of PVDF membranes, electroblotting has largely replaced electroelution as the method of choice for isolating proteins from most types of polyacrylamide gels (LeGendre and Matsudaira, 1988). PVDF membranes have very high protein binding capacities, good handling characteristics when either dry or wet, and are highly chemically inert. Hence they are compatible with many methods for subsequent analysis, including amino acid analysis, in situ chemical modification, in situ protease digestion, and N-terminal sequence analysis (Chapter 11). A wide range of general protein stains exist that are compatible with most blotting membranes (UNIT 10.8). Alternatively, radiolabeled proteins can be detected by autoradiography either in the gel or after transfer to a PVDF membrane (UNIT 10.11). The latter is normally preferable; as long as the proteins of interest are electrotransferred in high yield, autoradiography signals of weak β emitters such as 14C or 35S are usually higher on electroblotted membranes than in dried gels (even when fluorophores are impregnated in the gel), because the protein is primarily located on the surface of the membrane that was in contact with the gel during electroblotting. Historically, autoradiography was performed using medical X-ray films. More recently, a relatively wide range of commercially available autoradiography films and exposure methods have been developed that provide superior contrast and higher sensitivity for specific isotopes (UNIT 10.11). The dimensional stability and chemical resistance of PVDF membranes provides an optimal medium for combining multiple detection methods, either sequentially on the same membrane, or on parallel lanes that can be reassembled after different detection methods have been performed. For example, sets of lanes containing replicate samples can be run on a single gel separated by prestained standards, which are used as guides to cut the resulting membrane. Each lane set can be subjected to different detection methods such as protein staining and immunoblotting with different antibodies (UNIT 10.10). The analyzed membranes can then be precisely reassembled to determine the relationships between bands detected by different methods. As there is no shrinkage or swelling of the membrane, this approach is particularly valuable for detecting slight differences in migration between components detected by different methods.
Introduction
An approach that has recently emerged as a useful analytical method for separating peptides and, to a more limited extent, proteins is capillary electrophoresis (CE; UNIT 10.9). CE protocols have also been developed for preparative peptide isolation in adequate amounts for subsequent high sensitivity measurements such as micro-sequencing (Chapter 11) and mass spectrometry (Chapter 16). CE employs a fused-silica capillary column that may contain either free solution or a fluid matrix. One of CE’s limitations for protein
10.0.2 Supplement 17
Current Protocols in Protein Science
and peptide separations is the tendency of these molecules to interact with silanol groups on the capillary tubing surface. As described in UNIT 10.9, these interactions can be minimized through the use of coated capillaries, a separation pH several units above the protein pI, or buffer additives that minimize interactions with the silica walls. An interesting alternative approach is separation of proteins and peptides using capillary zone electrophoresis (CZE) in acidic amphoteric buffers (UNIT 10.13). The low conductivity of these buffers permits higher voltages to be used, resulting in faster separations, and the low pH minimizes ionization of silanol groups on underivatized capillaries. In this mode, nearly all proteins and peptides have strong positive charges and high mobilities. In contrast to the relatively simple equipment requirements of polyacrylamide-based electrophoretic methods, commercial CE instruments are similar in complexity and expense to high-performance liquid chromatography (HPLC) instruments, due to the necessity for a high-voltage supply, a high-performance cooling system, and a very sensitive detector. Because CE separates components by charge rather than by hydrophobicity as in reversedphase HPLC, it is a highly useful orthogonal separation method that is frequently used to complement HPLC peptide separation strategies. Compared with polyacrylamide methods, CE offers much higher resolution, flexibility, sensitivity, and speed for separating peptides (<6 kDa), whereas polyacrylamide methods are more generally applicable for protein separations. Documentation of gel electrophoresis results for either archival or publication purposes historically used primarily silver halide–based photography. In recent years, the availability of economical hardware and software for capturing, storing, and analyzing digital images has led to the replacement of photography by digital analysis as the preferred documentation method (UNIT 10.12). Advantages of digital-based methods include more convenient storage and retrieval, inexpensive and rapid production of backup or additional copies, facile manipulation such as resizing and inserting into reports, and rapid data distribution to colleagues over the Internet. Digital processing of electrophoretic results is particularly essential for comparing large numbers of gels, developing databases of gel patterns and related information, and performing quantitative comparisons such as proteome studies utilizing two-dimensional gels. LITERATURE CITED Garrels, J.I. 1979. Two-dimensional gel electrophoresis and computer analysis of proteins synthesized by cloned cell lines. J. Biol. Chem. 54:7961-7977. Laemmli, U.K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680-685. LeGendre, N. and Matsudaira, P. 1988. Direct protein microsequencing from Immobilon-P transfer membrane. BioTechniques 6:154-159. O’Farrell, P.H. 1975. High resolution two-dimensional electrophoresis of proteins. J. Biol. Chem. 250:40074021. Young, D.A., Voris, B.P., Maytin, E.V., and Colbert, R.A. 1983. Very high resolution two-dimensional electrophoretic separation of proteins on giant gels. Methods Enzymol. 91:190-214.
David W. Speicher
Electrophoresis
10.0.3 Current Protocols in Protein Science
Supplement 17
One-Dimensional SDS Gel Electrophoresis of Proteins
UNIT 10.1
Electrophoresis is used to separate complex mixtures of proteins, to investigate subunit compositions, and to verify homogeneity of protein samples. It can also serve to purify proteins for use in further applications. In polyacrylamide gel electrophoresis, proteins migrate in response to an electrical field through pores in the gel matrix; pore size decreases with higher acrylamide concentrations. The combination of gel pore size and protein charge, size, and shape determines the migration rate of the protein. Basic Protocol 1 gives the standard Laemmli method used for discontinuous gel electrophoresis under denaturing conditions, that is, in the presence of sodium dodecyl sulfate (SDS). In Basic Protocol 2, the standard method for full-size gels (e.g., 14 × 14 cm) is adapted for the minigel format (e.g., 7.3 × 8.3 cm). Minigels provide rapid separation but give lower resolution. Several alternate protocols are provided for specific applications. The first and second alternate protocols cover electrophoresis of peptides and small proteins, separations that require modification of standard buffers; Alternate Protocol 1 uses a Tris-tricine buffer system, and Alternate Protocol 2 employs modified Tris buffers in the absence of urea. Alternate Protocol 3 describes continuous SDS-PAGE, a simplified method in which the same buffer is used for both gel and electrode solutions and the stacking gel is omitted. Other protocols cover the preparation and electrophoresis of various types of gels: ultrathin gels (Alternate Protocol 4), multiple single-concentration gels (Support Protocol 1), gradient gels (Alternate Protocol 5), multiple gradient gels (Support Protocol 2), and multiple gradient minigels (Support Protocol 3). Two-dimensional electrophoresis is discussed in UNIT 10.4. CAUTION: Before any protocols are used, it is extremely important to read the following section about electricity and electrophoresis. ELECTRICITY AND ELECTROPHORESIS Many researchers are poorly informed concerning the electrical parameters of running a gel. It is important to note that the voltages and currents used during electrophoresis are dangerous and potentially lethal. Thus, safety should be an overriding concern. A working knowledge of electricity is an asset in determining what conditions to use and in troubleshooting the electrophoretic separation, if necessary. For example, an unusually high or low voltage for a given current (milliampere) might indicate an improperly made buffer or an electrical leak in the chamber. Safety Considerations 1. Never remove or insert high-voltage leads unless the power supply voltage is turned down to zero and the power supply is turned off. Always grasp high-voltage leads one at a time with one hand only. Never insert or remove high-voltage leads with both hands. This can shunt potentially lethal electricity through the chest and heart should electrical contact be made between a hand and a bare wire. On older or homemade instruments, the banana plugs may not be shielded and can still be connected to the power supply at the same time they make contact with a hand. Carefully inspect all cables and connections and replace frayed or exposed wires immediately. 2. Always start with the power supply turned off. Have the power supply controls turned all the way down to zero. Then hook up the gel apparatus: generally, connect the red Contributed by Sean R. Gallagher Current Protocols in Protein Science (1995) 10.1.1-10.1.34 Copyright © 2000 by John Wiley & Sons, Inc.
Electrophoresis
10.1.1 Supplement 3
high-voltage lead to the red outlet and the black high-voltage lead to the black outlet. Turn the power supply on with the controls set at zero and the high-voltage leads connected. Then, turn up the voltage, current, or power to the desired level. Reverse the process when the power supply is turned off: i.e., to disconnect the gel, turn the power supply down to zero, wait for the meters to read zero, turn off the power supply, and then disconnect the gel apparatus one lead at a time. CAUTION: If the gel is first disconnected and then the power supply turned off, a considerable amount of electrical charge is stored internally. The charge will stay in the power supply over a long time. This will discharge through the outlets even though the power supply is turned off and can deliver an electrical shock.
Ohm’s Law and Electrophoresis Understanding how a gel apparatus is connected to the power supply requires a basic understanding of Ohm’s law: voltage = current × resistance, or V = IR. A gel can be viewed as a resistor and the power supply as the voltage and current source. Most power supplies deliver constant current or constant voltage. Some will also deliver constant power: power = voltage × current, or VI = I2R. The discussion below focuses on constant current because this is the most common mode in vertical SDS-PAGE. Most modern commercial equipment is color-coded so that the red or positive terminal of the power supply can simply be connected to the red lead of the gel apparatus, which goes to the lower buffer chamber. The black lead is connected to the black or negative terminal and goes to the upper buffer chamber. This configuration is designed to work with vertical slab gel electrophoreses in which negatively charged proteins or nucleic acids move to the positive electrode in the lower buffer chamber (an anionic system). When a single gel is attached to a power supply, the negative charges flow from the negative cathode (black) terminal into the upper buffer chamber, through the gel, and into the lower buffer chamber. The lower buffer chamber is connected to the positive anode (red) terminal to complete the circuit. Thus, negatively charged molecules, such as SDS-coated proteins and nucleic acids, move from the negative cathode attached to the upper buffer chamber toward the positive anode attached to the lower chamber. SDSPAGE is an anionic system because of the negatively charged SDS. Occasionally, proteins are separated in cationic systems. In these gels, the proteins are positively charged because of the very low pH of the gel buffers (e.g., acetic acid/urea gels for histone separations) or the presence of a cationic detergent (e.g., cetyltrimethylammonium bromide, CTAB). Proteins move toward the negative electrode (cathode) in cationic gel systems, and the polarity is reversed compared to SDS-PAGE: the red lead from the lower buffer chamber is attached to the black outlet of the power supply, and the black lead from the upper buffer chamber is attached to the red outlet of the power supply. Most SDS-PAGE separations are performed under constant current (consult manufacturer’s instructions to set the power supply for constant current operation). The resistance of the gel will increase during SDS-PAGE in the standard Laemmli system. If the current is constant, then the voltage will increase during the run as the resistance goes up.
One-Dimensional SDS Gel Electrophoresis of Proteins
Power supplies usually have more than one pair of outlets. The pairs are connected in parallel with one another internally. If more than one gel is connected directly to the outlets of a power supply, then these gels are connected in parallel. In a parallel circuit, the voltage is the same across each gel. In other words, if the power supply reads 100 V, then each gel has 100 V across its electrodes. The total current, however, is the sum of the individual currents going through each gel. Therefore, under constant current it is necessary to
10.1.2 Current Protocols in Protein Science
power supply
power supply
+ –
gel tank
+ –
gel tank
gel tank
gel tank connected in series to one outlet (gel voltages are additive)
gel tank
power supply
+ –
gel tank
two gels connected in parallel to adjacent outlets (gel currents are additive)
gel tank
gel tank
two gels connected in parallel to one outlet (gel currents are additive)
Figure 10.1.1 Series and parallel connections of gel tanks to power supply.
increase the current for each additional gel that is connected to the power supply. Two identical gels require double the current to achieve the same starting voltages and electrophoresis separation times. Multiple gel apparatuses can also be connected to one pair of outlets on a power supply. This is useful with older power supplies that have a limited number of outlets. When connecting several gel units to one outlet, make certain the connections between the units are shielded and protected from moisture. The gels can be connected in parallel or in series (Fig. 10.1.1). In the case of two or more gels running off the same outlet in series, the current is the same for every gel. If 10 mA is displayed by the power supply meter, for example, each gel in series will experience 10 mA. The voltage, however, is additive for each gel. If one gel at a constant 10 mA produces 100 V, then two identical gels in series will produce 200 V (100 V each) and so on. Thus, the voltage can limit the number of units connected in series on low-voltage power supplies. Gel thickness affects the above relationships. A 1.5-mm gel can be thought of as consisting of two 0.75-mm-thick gels run in parallel. Because currents are additive in parallel circuits, a 0.75-mm gel will require half the current of the 1.5-mm gel to achieve the same starting voltage and separation time. If a gel thickness is doubled, then the current must also be doubled. There are limits to the amount of current that can be applied. Thicker gels require more current, generating more heat that must be dissipated. Unless temperature control is available in the gel unit, a thick gel should be run more slowly than a thin gel. NOTE: Milli-Q-purified water or equivalent should be used throughout the protocols. DENATURING (SDS) DISCONTINUOUS GEL ELECTROPHORESIS: LAEMMLI GEL METHOD
BASIC PROTOCOL 1
One-dimensional gel electrophoresis under denaturing conditions (i.e., in the presence of 0.1% SDS) separates proteins based on molecular size as they move through a polyacrylamide gel matrix toward the anode. The polyacrylamide gel is cast as a separating gel (sometimes called resolving or running gel) topped by a stacking gel and secured in an electrophoresis apparatus. After sample proteins are solubilized by boiling in the presence of SDS, an aliquot of the protein solution is applied to a gel lane, and the individual proteins are separated electrophoretically. 2-Mercaptoethanol (2-ME) or dithiothreitol (DTT) is added during solubilization to reduce disulfide bonds. This protocol is designed for a vertical slab gel with a maximum size of 0.75 mm × 14 cm × 14 cm. For thicker gels, or minigels (see Basic Protocol 2 and Support Protocol 3),
Electrophoresis
10.1.3 Current Protocols in Protein Science
Supplement 14
the volumes of stacking and separating gels and the operating current must be adjusted. Support protocols describe the preparation of ultrathin gels (see Alternate Protocol 4) and gradient gels (see Alternate Protocol 5), as well as the use of gel casters to make multiple gels, both single-concentration gels (see Support Protocol 1) and gradient gels (see Support Protocol 2). Materials Separating and stacking gel solutions (Table 10.1.1) H2O-saturated isobutyl alcohol 1× Tris⋅Cl/SDS, pH 8.8 (dilute 4× Tris⋅Cl/SDS, pH 8.8; Table 10.1.1) Protein sample to be analyzed 2× and 1× SDS sample buffer (see recipe) Protein molecular-weight-standards mixture (Table 10.1.2) 6× SDS sample buffer (see recipe; optional) 1× SDS electrophoresis buffer (see recipe) Electrophoresis apparatus: Protean II 16-cm cell (Bio-Rad) or SE 600/400 16-cm unit (Hoefer Pharmacia) with clamps, glass plates, casting stand, and buffer chambers 0.75-mm spacers 0.45-µm filters (used in stock solution preparation) 25-ml Erlenmeyer side-arm flask Vacuum pump with cold trap 0.75-mm Teflon comb with 1, 3, 5, 10, 15, or 20 teeth 25- or 100-µl syringe with flat-tipped needle Constant-current power supply (see introduction) Pour the separating gel 1. Assemble the glass-plate sandwich of the electrophoresis apparatus according to manufacturer’s instructions using two clean glass plates and two 0.75-mm spacers. If needed, clean the glass plates in liquid Alconox or RBS-35 (Pierce). These aqueous-based solutions are compatible with silver and Coomassie blue staining procedures.
2. Lock the sandwich to the casting stand. 3. Prepare the separating gel solution as directed in Table 10.1.1, degassing using a rubber-stoppered 25-ml Erlenmeyer side-arm flask connected with vacuum tubing to a vacuum pump with a cold trap. After adding the specified amount of 10% ammonium persulfate and TEMED to the degassed solution, stir gently to mix. Table 10.1.1 was prepared as a convenient summary to aid in the preparation of separating and stacking gels. The stacking gel is the same regardless of the separating gel used. The desired percentage of acrylamide in the separating gel depends on the molecular size of the protein being separated. Generally, use 5% gels for SDS-denatured proteins of 60 to 200 kDa, 10% gels for SDS-denatured proteins of 16 to 70 kDa, and 15% gels for SDS-denatured proteins of 12 to 45 kDa (Table 10.1.1).
4. Using a Pasteur pipet, apply separating gel solution to the sandwich along an edge of one of the spacers until the height of the solution between the glass plates is ∼11 cm. Use the solution immediately; otherwise it will polymerize in the flask.
One-Dimensional SDS Gel Electrophoresis of Proteins
Sample volumes <10 µl do not require a stacking gel. In this case, cast the resolving gel as you normally would, but extend the resolving gel into the comb to form the well. The proteins are then separated under the same conditions as used when a stacking gel is present. Although this protocol works well with single-concentration gels, a gradient gel is recommended for maximum resolution (see Alternate Protocol 5).
10.1.4 Supplement 14
Current Protocols in Protein Science
Table 10.1.1
Recipes for Polyacrylamide Separating and Stacking Gelsa
SEPARATING GEL Stock solutionb 30% acrylamide/ 0.8% bisacrylamide 4× Tris⋅Cl/SDS, pH 8.8 H2O 10% (w/v) ammonium persulfated TEMED
5
Final acrylamide concentration in separating gel (%)c 6 7 7.5 8 9 10 12 13 15
2.50
3.00
3.50
3.75
4.00
4.50
5.00
6.00
6.50
7.50
3.75
3.75
3.75
3.75
3.75
3.75
3.75
3.75
3.75
3.75
8.75 0.05
8.25 0.05
7.75 0.05
7.50 0.05
7.25 0.05
6.75 0.05
6.25 0.05
5.25 0.05
4.75 0.05
3.75 0.05
0.01
0.01
0.01
0.01
0.01
0.01
0.01
0.01
0.01
0.01
Preparation of separating gel In a 25-ml side-arm flask, mix 30% acrylamide/0.8% bisacrylamide solution, 4× Tris⋅Cl/SDS, pH 8.8 (see reagents, below), and H2O. Degas under vacuum about 5 min. Add 10% ammonium persulfate and TEMED. Swirl gently to mix. Use immediately. STACKING GEL (3.9% acrylamide) In a 25-ml side-arm flask, mix 0.65 ml of 30% acrylamide/0.8% bisacrylamide, 1.25 ml of 4× Tris⋅Cl/SDS, pH 6.8 (see reagents, below), and 3.05 ml H2O. Degas under vacuum 10 to 15 min. Add 25 µl of 10% ammonium persulfate and 5 µl TEMED. Swirl gently to mix. Use immediately. Failure to form a firm gel usually indicates a problem with the persulfate, TEMED, or both. REAGENTS USED IN GELS 30% acrylamide/0.8% bisacrylamide Mix 30.0 g acrylamide and 0.8 g N,N′-methylenebisacrylamide with H2O in a total volume of 100 ml. Filter the solution through a 0.45-µm filter and store at 4°C in the dark. The 2× crystallized grades of acrylamide and bisacrylamide are recommended. Discard after 30 days, as acrylamide gradually hydrolyzes to acrylic acid and ammonia. CAUTION: Acrylamide monomer is neurotoxic. Mask should be worn when weighing acrylamide powder. Gloves should be worn while handling the solution, and the solution should not be pipetted by mouth.
4× Tris⋅Cl/SDS, pH 6.8 (0.5 M Tris⋅Cl containing 0.4% SDS) Dissolve 6.05 g Tris base in 40 ml H2O. Adjust to pH 6.8 with 1 N HCl. Add H2O to 100 ml total volume. Filter the solution through a 0.45-µm filter, add 0.4 g SDS, and store at 4°C up to 1 month. 4× Tris⋅Cl/SDS, pH 8.8 (1.5 M Tris⋅Cl containing 0.4% SDS) Dissolve 91 g Tris base in 300 ml H2O. Adjust to pH 8.8 with 1 N HCl. Add H2O to 500 ml total volume. Filter the solution through a 0.45-µm filter, add 2 g SDS, and store at 4°C up to 1 month. aThe recipes produce 15 ml of separating gel and 5 ml of stacking gel, which are adequate for a gel of dimensions
0.75 mm × 14 cm × 14 cm. The recipes are based on the SDS (denaturing) discontinuous buffer system of Laemmli (1970). bAll reagents and solutions used in the protocol must be prepared with Milli-Q-purified water or equivalent. cUnits of numbers in table body are milliliters. The desired percentage of acrylamide in the separating gel depends on the molecular size of the protein being separated. See annotation to step 3, Basic Protocol 1. dBest to prepare fresh.
Electrophoresis
10.1.5 Current Protocols in Protein Science
5. Using another Pasteur pipet, slowly cover the top of the gel with a layer (∼1 cm thick) of H2O-saturated isobutyl alcohol, by gently layering the isobutyl alcohol against the edge of one and then the other of the spacers. Be careful not to disturb the gel surface. The overlay provides a barrier to oxygen, which inhibits polymerization, and allows a flat interface to form during gel formation. The H2O-saturated isobutyl alcohol is prepared by shaking isobutyl alcohol and H2O in a separatory funnel. The aqueous (lower) phase is removed. This procedure is repeated several times. The final upper phase is H2O-saturated isobutyl alcohol.
6. Allow the gel to polymerize 30 to 60 min at room temperature. A sharp optical discontinuity at the overlay/gel interface will be visible on polymerization. Failure to form a firm gel usually indicates a problem with the ammonium persulfate, TEMED (N,N,N′,N′-tetramethylethylenediamine), or both. Ammonium persulfate solution should be made fresh before use. Ammonium persulfate should “crackle” when added to the water. If not, fresh ammonium persulfate should be purchased. Purchase TEMED in small bottles so, if necessary, a new previously unopened source can be tried.
Pour the stacking gel 7. Pour off the layer of H2O-saturated isobutyl alcohol and rinse with 1× Tris⋅Cl/SDS, pH 8.8. Residual isobutyl alcohol can reduce resolution of the protein bands; therefore, it must be completely removed. The isobutyl alcohol overlay should not be left on the gel longer than 2 hr.
8. Prepare the stacking gel solution as directed in Table 10.1.1. Use the solution immediately to keep it from polymerizing in the flask.
9. Using a Pasteur pipet, slowly allow the stacking gel solution to trickle into the center of the sandwich along an edge of one of the spacers until the height of the solution in the sandwich is ∼1 cm from the top of the plates. Be careful not to introduce air bubbles into the stacking gel.
10. Insert a 0.75-mm Teflon comb into the layer of stacking gel solution. If necessary, add additional stacking gel to fill the spaces in the comb completely. Again, be careful not to trap air bubbles in the tooth edges of the comb; they will cause small circular depressions in the well after polymerization that will lead to distortion in the protein bands during separation.
11. Allow the stacking gel solution to polymerize 30 to 45 min at room temperature. A sharp optical discontinuity will be visible around wells on polymerization.
Prepare the sample and load the gel 12. Dilute a portion of the protein sample to be analyzed 1:1 (v/v) with 2× SDS sample buffer and heat 3 to 5 min at 100°C in a sealed screw-cap microcentrifuge tube. If the sample is a precipitated protein pellet, dissolve the protein in 50 to 100 µl of 1× SDS sample buffer and boil 3 to 5 min at 100°C. Dissolve protein-molecular-weight standards mixture in 1× SDS sample buffer according to supplier’s instructions as a control (Table 10.1.2).
One-Dimensional SDS Gel Electrophoresis of Proteins
For dilute protein solutions, consider adding 5:1 protein solution/6× SDS sample buffer to increase the amount of protein loaded. Proteins can also be concentrated by precipitation in acetone, ethanol, or trichloroacetic acid (TCA), but losses will occur.
10.1.6 Current Protocols in Protein Science
Table 10.1.2 Molecular Weights of Protein Standards for Polyacrylamide Gel Electrophoresisa
Protein Cytochrome c α-Lactalbumin Lysozyme (hen egg white) Myoglobin (sperm whale) β-Lactoglobulin Trypsin inhibitor (soybean) Trypsinogen, PMSF treated Carbonic anhydrase (bovine erythrocytes) Glyceraldehyde-3-phosphate dehydrogenase (rabbit muscle) Lactate dehydrogenase (porcine heart) Aldolase Ovalbumin Catalase Bovine serum albumin Phosphorylase b (rabbit muscle) β-Galactosidase RNA polymerase, E. coli Myosin, heavy chain (rabbit muscle)
Molecular weight 11,700 14,200 14,300 16,800 18,400 20,100 24,000 29,000 36,000 36,000 40,000 45,000 57,000 66,000 97,400 116,000 160,000 205,000
aProtein standards are commercially available in kits (e.g., Hoefer Pharma-
cia Biotech, GIBCO/BRL, Bio-Rad, or Sigma).
For a 0.8-cm-wide well, 25 to 50 µg total protein in <20 µl is recommended for a complex mixture when staining with Coomassie blue, and 1 to 10 µg total protein is needed for samples containing one or a few proteins. If silver staining is used, 10- to 100-fold less protein can be applied (0.01 to 5 µg in <20 µl depending on sample complexity). To achieve the highest resolution possible, the following precautions are recommended. Prior to adding the sample buffer, keep samples at 0°C. Add the SDS sample buffer (room temperature) directly to the 0°C sample (still on ice) in a screw-top microcentrifuge tube. Cap the tube to prevent evaporation, vortex, and transfer directly to a 100°C water bath for 3 to 5 min. Let immunoprecipitates dissolve for 1 hr at 56°C in 1× SDS sample buffer prior to boiling. DO NOT leave the sample in SDS sample buffer at room temperature without first heating to a 100°C to inactivate proteases (see Critical Parameters and Troubleshooting). Endogenous proteases are very active in SDS sample buffer and will cause severe degradation of the sample proteins after even a few minutes at room temperature. To test for possible proteases, mix the sample with SDS sample buffer without heating and leave at room temperature for 1 to 3 hr. A loss of high-molecular-weight bands and a general smearing of the banding pattern indicate a protease problem. Once heated, the samples can sit at room temperature for the time it takes to load samples.
13. Carefully remove the Teflon comb without tearing the edges of the polyacrylamide wells. After the comb is removed, rinse wells with 1× SDS electrophoresis buffer. The rinse removes unpolymerized monomer; otherwise, the monomer will continue to polymerize after the comb is removed, creating uneven wells that will interfere with sample loading and subsequent separation.
14. Using a Pasteur pipet, fill the wells with 1× SDS electrophoresis buffer. Electrophoresis
10.1.7 Current Protocols in Protein Science
If well walls are not upright, they can be manipulated with a flat-tipped needle attached to a syringe.
15. Attach gel sandwich to upper buffer chamber using manufacturer’s instructions. 16. Fill lower buffer chamber with the recommended amount of 1× SDS electrophoresis buffer. 17. Place sandwich attached to upper buffer chamber into lower buffer chamber. 18. Partially fill the upper buffer chamber with 1× SDS electrophoresis buffer so that the sample wells of the stacking gel are filled with buffer. Monitor the upper buffer chamber for leaks and if necessary, reassemble the unit. A slow leak in the upper buffer chamber may cause arcing around the upper electrode and damage the upper buffer chamber.
19. Using a 25- or 100-µl syringe with a flat-tipped needle, load the protein sample(s) into one or more wells by carefully applying the sample as a thin layer at the bottom of the wells. Load control wells with molecular weight standards. Add an equal volume of 1× SDS sample buffer to any empty wells to prevent spreading of adjoining lanes. Preparing the samples at approximately the same concentration and loading an equal volume to each well will ensure that all lanes are the same width and that the proteins run evenly. If unequal volumes of sample buffer are added to wells, the lane with the larger volume will spread during electrophoresis and constrict the adjacent lanes, causing distortions. The samples will layer on the bottom of the wells because the glycerol added to the sample buffer gives the solution a greater density than the electrophoresis buffer. The bromphenol blue in the sample buffer makes sample application easy to follow visually.
20. Fill the remainder of the upper buffer chamber with additional 1× SDS electrophoresis buffer so that the upper platinum electrode is completely covered. Do this slowly so that samples are not swept into adjacent wells. Run the gel 21. Connect the power supply to the cell and run at 10 mA of constant current for a slab gel 0.75 mm thick, until the bromphenol blue tracking dye enters the separating gel. Then increase the current to 15 mA. For a standard 16-cm gel sandwich, 4 mA per 0.75-mm-thick gel will run ∼15 hr (i.e., overnight); 15 mA per 0.75-mm gel will take 4 to 5 hr. To run two gels or a 1.5-mm-thick gel, simply double the current. When running a 1.5-mm gel at 30 mA, the temperature must be controlled (10° to 20°C) with a circulating constant-temperature water bath to prevent “smiling” (curvature in the migratory band). Temperatures <5°C should not be used because SDS in the running buffer will precipitate. If the level of buffer in the upper chamber decreases, a leak has occurred.
22. After the bromphenol blue tracking dye has reached the bottom of the separating gel, disconnect the power supply. See Safety Considerations in the introduction.
Disassemble the gel 23. Discard electrode buffer and remove the upper buffer chamber with the attached gel sandwich. One-Dimensional SDS Gel Electrophoresis of Proteins
24. Orient the gel so that the order of the sample wells is known, remove the sandwich from the upper buffer chamber, and lay the sandwich on a sheet of absorbent paper or paper towels.
10.1.8 Current Protocols in Protein Science
25. Carefully slide one of the spacers halfway from the edge of the sandwich along its entire length. Use the exposed spacer as a lever to pry open the glass plate, exposing the gel. 26. Carefully remove the gel from the lower plate. Cut a small triangle off one corner of the gel so the lane orientation is not lost during staining and drying. Proceed with protein detection. The gel can be stained with Coomassie blue or silver (UNIT 10.5), or proteins can be electroeluted, electroblotted onto a polyvinylidene difluoride (PVDF) membrane for subsequent staining or sequence analysis (UNIT 10.7), or transferred to a membrane for immunoblotting. If the proteins are radiolabeled (UNIT 3.3), they can be detected by autoradiography.
ELECTROPHORESIS IN TRIS-TRICINE BUFFER SYSTEMS Separation of peptides and proteins under 10 to 15 kDa is not possible in the traditional Laemmli discontinuous gel system (Basic Protocol 1). This is due to the comigration of SDS and smaller proteins, obscuring the resolution. Two approaches to obtain the separation of small proteins and peptides in the range of 5 to 20 kDa are presented: this Tris-tricine method that follows and a system using increased buffer concentrations (see Alternate Protocol 2). The Tris-tricine system uses a modified buffer to separate the SDS and peptides, thus improving resolution. Several precast gels are available for use with the tricine formulations (Table 10.1.3).
ALTERNATE PROTOCOL 1
Additional Materials (also see Basic Protocol 1) Separating and stacking gel solutions (Table 10.1.4) 2× tricine sample buffer (see recipe) Peptide molecular-weight-standards mixture (Table 10.1.5) Cathode buffer (see recipe) Anode buffer (see recipe) Coomassie blue G-250 staining solution (see recipe) 10% (v/v) acetic acid
Table 10.1.3
Vertical Format Precast Gel Compatibility
Gel supplier Gel type and compatibility SDS-PAGE gel type offered Peptide (tricine) Single concentration Gradient Minigel size Standard gel size
Bio-Rad ISS/Daiichi × × × ×
× × × × ×
Compatibility of gel with equipment manufactured by Hoefer Pharmacia Biotech × Bio-Rad × × GIBCO/BRL × × Novex × ISS/Daiichi ×
Jule
Millipore Novex
× × × × ×
× × × ×
× × × ×
× × ×
× × × × ×
×
×
× ×
Electrophoresis
10.1.9 Current Protocols in Protein Science
Table 10.1.4
Recipes for Tricine Peptide Separation Gelsa
SEPARATING AND STACKING GELS Stock solutionb 30% acrylamide/0.8% bisacrylamide Tris⋅Cl/SDS, pH 8.45 H2O Glycerol 10% (w/v) ammonium persulfatec TEMED
Separating gel
Stacking gel
9.80 ml 10.00 ml 7.03 ml 4.00 g (3.17 ml) 50 µl 10 µl
1.62 ml 3.10 ml 7.78 ml — 25 µl 5 µl
Prepare separating and stacking gel solutions separately.
In a 25-ml side-arm flask, mix 30% acrylamide/0.8% bisacrylamide solution (Table 10.1.1), Tris⋅Cl/SDS, pH 8.45 (see reagents, below), and H2O. Add glycerol to separating gel only. Degas under vacuum 10 to 15 min. Add 10% ammonium persulfate and TEMED. Swirl gently to mix, use immediately. Failure to form a firm gel usually indicates a problem with the persulfate, TEMED, or both.
ADDITIONAL REAGENTS USED IN GELS Tris⋅Cl/SDS, pH 8.45 (3.0 M Tris⋅Cl 0.3% SDS) Dissolve 182 g Tris base in 300 ml H2O. Adjust to pH 8.45 with 1 N HCl. Add H2O to 500 ml total volume. Filter the solution through a 0.45-µm filter, add 1.5 g SDS, and store at 4°C up to 1 month. aThe recipes produce 30 ml of separating gel and 12.5 ml of stacking gel, which are adequate for two gels of
dimensions 0.75 mm × 14 cm × 14 cm. The recipes are based on the Tris-tricine buffer system of Schagger and von Jagow (1987). bAll reagents and solutions used in the protocol must be prepared with Milli-Q-purified water or equivalent. cBest to prepare fresh.
1. Prepare and pour the separating and stacking gels (see Basic Protocol 1, steps 1 to 11), using Table 10.1.4 in place of Table 10.1.1. 2. Prepare the sample (see Basic Protocol 1, step 12), but make the following changes for tricine gels. Substitute 2× tricine sample buffer for the 2× SDS sample buffer. Dilute an aliquot of the protein or peptide sample to be analyzed 1:1 (v/v) with 2× tricine sample buffer. Treat the sample at 40°C for 30 to 60 min prior to loading. If proteolytic activity is a problem, heating samples to 100°C for 3 to 5 min before loading the wells may be required (see Basic Protocol 1, annotation to step 12). Use the peptide molecular-weight-standards mixture for peptide separations (Table 10.1.5).
3. Load the gel and set up the electrophoresis apparatus (see Basic Protocol 1, steps 13 to 20) with the following alterations. Remove comb and, using the tricine-containing cathode buffer or water, rinse once and fill wells. Fill the lower buffer chamber with anode buffer, assemble the unit, and attach the upper buffer chamber. Fill the upper buffer chamber with cathode buffer and load the samples. One-Dimensional SDS Gel Electrophoresis of Proteins
4. Connect the power supply to the cell and run 1 hr at 30 V (constant voltage) followed by 4 to 5 hr at 150 V (constant voltage). Use heat exchanger to keep the electrophoresis chamber at room temperature.
10.1.10 Current Protocols in Protein Science
Table 10.1.5 Molecular Weights of Peptide Standards for Polyacrylamide Gel Electrophoresisa
Peptide Myoglobin (polypeptide backbone) Myoglobin 1-131 Myoglobin 56-153 Myoglobin 56-131 Myoglobin 1-55 Glucagon Myoglobin 132-153
Molecular weight 16,950 14,440 10,600 8,160 6,210 3,480 2,510
aPeptide standards are commercially available from Sigma.
5. After the tracking dye has reached the bottom of the separating gel, disconnect the power supply. See Safety Considerations in the introduction. Coomassie blue G-250 is used as a tracking dye instead of bromphenol blue because it moves ahead of the smallest peptides.
6. Disassemble the gel (see Basic Protocol 1, steps 23 to 26). Stain proteins in the gel for 1 to 2 hr in Coomassie blue G-250 staining solution. Follow by destaining with 10% acetic acid, changing the solution every 30 min until background is clear (3 to 5 changes). For higher sensitivity, use silver staining as a recommended alternative (UNIT 10.5). Prolonged staining and destaining will result in the loss of resolution of the smaller proteins (<10 kDa). Proteins diffuse within the gel and out of the gel, resulting in a loss of staining intensity and resolution.
NONUREA PEPTIDE SEPARATIONS WITH TRIS BUFFERS A simple modification of the traditional Laemmli buffer system presented in Basic Protocol 1, in which the increased concentration of buffers provides better separation between the stacked peptides and the SDS micelles, permits reasonable separation of peptides as small as 5 kDa.
ALTERNATE PROTOCOL 2
Additional Materials (also see Basic Protocol 1) Separating and stacking gel solutions (Table 10.1.6) 2× SDS electrophoresis buffer (see recipe) 2× Tris⋅Cl/SDS, pH 8.8 (dilute 4× Tris⋅Cl/SDS, pH 8.8; Table 10.1.1) 1. Prepare and pour the separating gel (see Basic Protocol 1, steps 1 to 6) using Table 10.1.6 in place of Table 10.1.1. 2. Prepare and pour the stacking gel (see Basic Protocol 1, steps 7 to 11), using 2× Tris⋅Cl/SDS, pH 8.8, rather than 1× Tris⋅Cl/SDS buffer, for rinsing the gel after removing the isobutyl alcohol overlay. 3. Prepare the sample and load the gel (see Basic Protocol 1, steps 12 to 20) and substitute 2× SDS electrophoresis buffer for the 1× SDS electrophoresis buffer. Table 10.1.5 lists the standards for small protein separations. Electrophoresis
10.1.11 Current Protocols in Protein Science
Table 10.1.6
Recipes for Modified Laemmli Peptide Separation Gelsa
SEPARATING AND STACKING GELS Stock solutionb 30% acrylamide/0.8% bisacrylamide 8× Tris⋅Cl, pH 8.8 4× Tris⋅Cl, pH 6.8 10% (w/v) SDS H2O 10% (w/v) ammonium persulfatec TEMED
Separating gel
Stacking gel
10.00 ml 3.75 ml — 0.15 ml 1.00 ml 50 µl 10 µl
0.65 ml — 1.25 ml 50 µl 3.00 ml 25 µl 5 µl
Prepare separating and stacking gel solutions separately. In a 25-ml side-arm flask, mix 30% acrylamide/0.8% bisacrylamide solution (see Table 10.1.1), 8× Tris⋅Cl, pH 8.8 (separating gel) or 4× Tris⋅Cl, pH 6.8 (stacking gel), 10% SDS (see reagents, below), and H2O. Degas under vacuum 10 to 15 min. Add 10% ammonium persulfate and TEMED. Swirl gently to mix; use immediately. Failure to form a firm gel usually indicates a problem with the persulfate, TEMED, or both.
ADDITIONAL REAGENTS USED IN GELS 4× Tris⋅Cl, pH 6.8 (0.5 M Tris⋅Cl) Dissolve 6.05 g Tris base in 40 ml H2O. Adjust to pH 6.8 with 1 N HCl. Add H2O to 100 ml total volume. Filter the solution through a 0.45-µm filter and store at 4°C up to 1 month. 8× Tris⋅Cl, pH 8.8 (3.0 M Tris⋅Cl) Dissolve 182 g Tris base in 300 ml H2O. Adjust to pH 8.8 with 1 N HCl. Add H2O to 500 ml total volume. Filter the solution through a 0.45-µm filter and store at 4°C up to 1 month. 10% (w/v) SDS Mix 1 g SDS in 10 ml of H2O. Use immediately. aThe recipes produce 15 ml of separating gel and 5 ml of stacking gel, which are adequate for one gel of
dimensions 0.75 mm × 14 cm × 14 cm. The recipes are based on the modified Laemmli peptide separation system of Okajima et al. (1993). bAll reagents and solutions used in the protocol must be prepared with Milli-Q-purified water or equivalent. cBest to prepare fresh.
4. Run the gel (see Basic Protocol 1, steps 21 and 22). Note that the separations will take ∼25% longer than those using Basic Protocol 1. The increased buffer concentrations lead to faster transit through the stacking gel but lower mobility in the resolving gel.
5. Disassemble the gel (see Basic Protocol 1, steps 23 to 26). One-Dimensional SDS Gel Electrophoresis of Proteins
If Coomassie blue staining is used, stain using the solutions outlined in UNIT 10.5 for 1 hr followed by destaining with several changes of 25% methanol/7% acetic acid. Prolonged staining will result in the loss of the smaller proteins.
10.1.12 Current Protocols in Protein Science
CONTINUOUS SDS-PAGE With continuous SDS-PAGE, the same buffer is used for both the gel and electrode solutions. Although continuous gels lack the resolution of the discontinuous systems, they are extremely versatile, less prone to mobility artifacts, and much easier to prepare. The stacking gel is omitted.
ALTERNATE PROTOCOL 3
Additional Materials (also see Basic Protocol 1) Separating gel solution (Table 10.1.7) 2× and 1× phosphate/SDS sample buffer (see recipe) 1× phosphate/SDS electrophoresis buffer (see recipe) 1. Prepare and pour a single separating gel (see Basic Protocol 1, steps 1 to 4), except use solutions in Table 10.1.7 in place of those in Table 10.1.1 and fill the gel sandwich to the top. Omit the stacking gel. Insert the comb (see Basic Protocol 1, step 10) and allow the gel to polymerize 30 to 60 min at room temperature.
Table 10.1.7
Recipes for Separating Gels for Continuous SDS-PAGEa
SEPARATING GEL Final acrylamide concentration in separating gel (%)c
Stock solutionb 5 30% acrylamide/ 0.8% bisacrylamide 4× phosphate/SDS, pH 7.2 H2O 10% (w/v) ammonium persulfated TEMED
2.5
6
7
8
9
10
11
12
13
14
15
3.00 3.50 4.00 4.50 5.00 5.50 6.00 6.50 7.00 7.50
3.75 3.75 3.75 3.75 3.75 3.75 3.75 3.75 3.75 3.75 3.75 8.75 8.25 7.75 7.25 6.75 6.25 5.75 5.25 4.75 4.25 3.75 0.05 0.05 0.05 0.05 0.05 0.05 0.05 0.05 0.05 0.05 0.05 0.01 0.01 0.01 0.01 0.01 0.01 0.01 0.01 0.01 0.01 0.01
Preparation of separating gel In a 25-ml side-arm flask, mix 30% acrylamide/0.8% bisacrylamide solution (see Table 10.1.1), 4× phosphate/SDS, pH 7.2, and H2O. Degas under vacuum about 5 min. Add 10% ammonium persulfate and TEMED. Swirl gently to mix. Use immediately.
ADDITIONAL REAGENTS USED IN GELS 4× phosphate/SDS, pH 7.2 (0.4 M sodium phosphate/0.4% SDS) Mix 46.8 g NaH2PO4⋅H2O, 231.6 g Na2HPO4⋅7 H2O, and 12 g SDS in 3 liters H2O. Store at 4°C for up to 3 months. aThe recipes produce 15 ml of separating gel, which is adequate for one gel of dimensions 0.75 mm × 14 cm × 14 cm. The recipes are based on the original continuous phosphate buffer system of Weber et al. (1972). The stacking gel is omitted. bAll reagents and solutions used in the protocol must be prepared with Milli-Q-purified water or equivalent. cUnits of numbers in table body are milliliters. The desired percentage of acrylamide in the separating gel depends on the molecular size of the protein being separated. See Basic Protocol 1, annotation to step 3. dBest to prepare fresh.
Electrophoresis
10.1.13 Current Protocols in Protein Science
2. Mix the protein sample 1:1 with 2× phosphate/SDS sample buffer and heat to 100°C for 2 min. For large sample volumes or samples suspended in high ionic strength buffers (>50 mM), dialyze against 1× sample buffer prior to electrophoresis. Note that the precautions about proteases (see Basic Protocol 1, step 12) apply here.
3. Assemble the electrophoresis apparatus and load the sample (see Basic Protocol 1, steps 13 to 20), using the phosphate/SDS electrophoresis buffer and loading empty wells with 1× phosphate/SDS sample buffer. 4. Connect the power supply and start the run with 15 mA per 0.75-mm-thick gel until the tracking dye has entered the gel. Continue electrophoresis at 30 mA for 3 hr (5% gel), 5 hr (10% gel), 8 hr (15% gel), or until the dye reaches the bottom of the gel. Use temperature control if available to maintain the gel at 15° to 20°C. SDS will precipitate below 15°C in this system.
5. Disassemble the gel (see Basic Protocol 1, steps 23 to 26). Proteins in the gel may be stained using the protocols given in UNIT 10.5. ALTERNATE PROTOCOL 4
CASTING AND RUNNING ULTRATHIN GELS Ultrathin gels provide superb resolution but are difficult to handle. In this application, gels are cast on Gel Bond, a Mylar support material. Silver staining is recommended for the best resolution. Combs and spacers for gels <0.5 mm thick are not readily available for most protein electrophoresis units. However, by adapting combs and spacers used for DNA sequencing, casting gels from 0.2 to 0.5 mm thick is straightforward. Additional Materials (also see Basic Protocol 1) 95% (v/v) ethanol Gel Bond (FMC) cut to a size slightly smaller than the gel plate dimensions Glue stick Ink roller (available from art supply stores) Combs and spacers (0.19 to 0.5 mm; sequencing gel spacers and combs can be cut to fit) 1. Wash gel plates with water-based laboratory detergent followed by successive rinses with hot tap water, deionized water, and finally 95% ethanol. Allow to air dry. Gel plates must be extremely clean for casting thin gels.
2. Apply a streak of adhesive from a glue stick to the bottom edge of the glass plate. Quickly position the Gel Bond with the hydrophobic side down (a drop of water will bead up on the hydrophobic surface). Apply pressure with Kimwipe tissue to attach the Gel Bond firmly to the plate. Last, pull the top portion of the Gel Bond back, place a few drops of water underneath, and roll flat with an ink roller. Make sure the Gel Bond does not extend beyond the edges of the upper and lower sealing surface of the plate. This will cause it to buckle on sealing. Reposition the Gel Bond if needed to prevent it from extending beyond the glass plate. Material may also be trimmed to fit flush with the plate edge.
One-Dimensional SDS Gel Electrophoresis of Proteins
3. Assemble the gel cassette according to the manufacturer’s instructions (also see Basic Protocol 1, steps 1 and 2). Just prior to assembly, blow air over the surface of both the Gel Bond and the opposing glass surface to remove any particulate material (e.g., dust). Sequencing gel spacers can be easily adapted. First, cut the spacers slightly longer
10.1.14 Current Protocols in Protein Science
than the length of the gel plate. Position a spacer along each edge of the glass plate and assemble the gel sandwich, clamping in place. With a razor blade, trim the excess spacer at top and bottom to get a reusable spacer exactly the size of the plate.
4. Prepare and pour the separating and stacking gels (see Basic Protocol 1, steps 3 to 9). In place of the Teflon comb, insert a square well sequencing comb cut to fit within the gel sandwich. Allow the stacking gel to polymerize 30 to 45 min at room temperature. Less solution is needed for ultrathin gels. For example, a 0.5-mm-thick gel requires 33% less gel solution than a 0.75-mm gel.
5. Prepare the sample and load the gel (see Basic Protocol 1, steps 12 to 20). When preparing protein samples for ultrathin gels, 3 to 4 µl at 5 µg protein/µl is required for Coomassie blue R-250 staining, whereas 10-fold less is needed for silver staining.
6. Run the gel (see Basic Protocol 1, steps 21 and 22), except conduct the electrophoresis at 7 mA/gel (0.25-mm-thick gels) or 14 mA/gel (0.5-mm-thick gels) for 4 to 5 hr. 7. When the separation is complete, disassemble the unit and remove the gel (see Basic Protocol 1, steps 23 to 26). With a gloved hand, wash away the adhesive material under a stream of water before proceeding to protein detection. Either Coomassie blue or silver staining (UNIT 10.5) may be used, but silver staining produces particularly fine resolution with thin Gel Bond–backed gels. Compared to staining thicker (>0.75 mm) gels, thin (<0.75 mm) gels stain and destain more quickly. Although the optimum staining times must be empirically determined, all steps in Coomassie blue and silver staining procedures are generally reduced by half. Gloves are needed throughout these procedures to prevent contamination by proteins on the surface of skin.
CASTING MULTIPLE SINGLE-CONCENTRATION GELS Casting multiple gels at one time has several advantages. All the gels are identical, so sample separation is not affected by gel-to-gel variation. Furthermore, casting ten gels is only slightly more difficult than casting two gels. Once cast, gels can be stored for several days in a refrigerator.
SUPPORT PROTOCOL 1
Additional Materials (also see Basic Protocol 1) Separating and stacking gels for single-concentration gels (Table 10.1.8) H2O-saturated isobutyl alcohol Multiple gel caster (Bio-Rad, Hoefer Pharmacia) Extra plates and spacers 14 × 14–cm acrylic blocks or polycarbonate sheets 250- and 500-ml side-arm flasks (used in gel preparation) Long razor blade or plastic wedge (Wonder Wedge, Hoefer Pharmacia) Resealable plastic bags Pour the separating gel 1. Assemble the multiple gel caster according to the manufacturer’s instructions. With the Hoefer Pharmacia unit make sure to insert the large triangular space filler plug in the base of the caster. The plug is removed when casting gradient gels (see Support Protocol 2).
2. Assemble glass sandwiches and stack them in the casting chamber. Stack up to ten 1.5-mm gels and fill in extra space with acrylic blocks or polycarbonate sheets to
Electrophoresis
10.1.15 Current Protocols in Protein Science
Table 10.1.8
Recipes for Multiple Single-Concentration Polyacrylamide Gelsa
SEPARATING GEL Final acrylamide concentration in separating gel (%)c
Stock solutionb 30% acrylamide/ 0.8% bisacrylamide 4× Tris⋅Cl/SDS, pH 8.8 H2O 10% (w/v) ammonium persulfated TEMED
5
6
7
8
9
10
11
12
13
14
15
52
62
72
83
93
103
114
124
134
145
155
78
78
78
78
78
78
78
78
78
78
78
181 1.0
171 1.0
160 1.0
150 1.0
140 1.0
129 1.0
119 1.0
109 1.0
98 1.0
88 1.0
78 1.0
0.21
0.21
0.21
0.21
0.21
0.21
0.21
0.21
0.21
0.21
0.21
Preparation of separating gel In a 500-ml side-arm flask, mix 30% acrylamide/0.8% bisacrylamide solution (see Table 10.1.1), 4× Tris⋅Cl/SDS, pH 8.8 (Table 10.1.1), and H2O. Degas under vacuum about 5 min. Add 10% ammonium persulfate and TEMED. Swirl gently to mix; use immediately. STACKING GEL In a 250-ml side-arm flask, mix 13.0 ml 30% acrylamide/0.8% bisacrylamide solution, 25 ml 4× Tris⋅Cl/SDS, pH 6.8 (Table 10.1.1), and 61 ml H2O. Degas under vacuum about 5 min. Add 0.25 ml 10% ammonium persulfate and 50 µl TEMED. Swirl gently to mix. Use immediately. Failure to form a firm gel usually indicates a problem with the persulfate, TEMED, or both. aThe recipes produce about 300 ml of separating gel and 100 ml of stacking gel, which are adequate for ten gels of dimensions 1.5 mm × 14 cm × 14 cm. Volumes were measured using 1.5-mm spacers (or fewer gels). For thinner spacers or fewer gels, calculate volumes using the equation in the annotation to step 4. The recipes are based on the SDS (denaturing) discontinuous buffer system of Laemmli (1970). bAll reagents and solutions used in the protocol must be prepared with Milli-Q-purified water or equivalent. cUnits of numbers in table body are milliliters. The desired percentage of acrylamide in separating gel depends on the molecular size of the protein being separated. See Basic Protocol 1, annotation to step 3. dBest to prepare fresh.
hold the sandwiches tightly in place. Make sure the spacers are straight along the top, right, and left edges of the glass plates and that all edges of the stack are flush. The presence of loosely fitting sandwiches in the caster will lead to unevenly cast gels, creating distortions during electrophoresis. Polycarbonate inhibits gel polymerization. Therefore, if polycarbonate sheets are placed in the caster before and after the set of glass sandwiches, the entire set will slide out as one block after polymerization. Placing polycarbonate sheets between each gel sandwich makes them easier to separate from one another after polymerization.
3. Place the front sealing plate on the casting chamber, making sure the stack fits snugly. Secure the plate with four spring clamps and tighten the bottom thumb screws. 4. Prepare the separating (resolving) gel solution (Table 10.1.8). A 12-cm separating gel with a 4-cm stacking gel is recommended. One-Dimensional SDS Gel Electrophoresis of Proteins
If fewer than ten gels are prepared (Table 10.1.8), use the following formula to estimate the amount of separating gel volume needed: Volume = gel number × height (cm) × width (cm) × thickness (cm) + 4 × gel number + 10 ml.
10.1.16 Current Protocols in Protein Science
5. Using a 100-ml disposable syringe with flat-tipped needle, inject the resolving gel solution down the side of one spacer into the multiple caster. A channel in the silicone plug distributes the solution throughout the whole caster. Avoid introducing bubbles by giving the caster a quick tap on the benchtop once the caster is filled. 6. Overlay the center of each gel with 100 µl H2O-saturated isobutyl alcohol and let polymerize for 1 to 2 hr. 7. Drain off the overlay and rinse the surface with 1× Tris⋅Cl/SDS, pH 8.8. If the gels will not be used immediately, skip to step 12. Pour the stacking gel 8. Prepare the stacking gel solution either singly (see Basic Protocol 1, step 8) or for all the gels at once (Table 10.1.8). The stacking gel solution should be prepared just before pouring the gel.
9. Fill each sandwich in the caster with stacking gel solution. 10. Insert a comb into each sandwich and let the gel polymerize for 2 hr. Insert the combs at a 45° angle to avoid trapping air underneath the comb teeth. Air bubbles will inhibit polymerization and cause dents in the wells and a distorted pattern of protein bands.
11. Remove the combs and rinse wells with 1× SDS electrophoresis buffer. Remove the gels from the caster 12. Remove the gels from the caster and separate by carefully inserting a long razor blade or knife between each gel sandwich. A plastic wedge (Hoefer Pharmacia’s Wonder Wedge) also works well. 13. Clean the outside of each gel plate with running water to remove the residual polymerized and unpolymerized acrylamide. 14. Overlay gels to be stored with 1× Tris⋅Cl/SDS, pH 8.8, place in a resealable plastic bag, and store at 4°C until needed (up to 1 week). SEPARATION OF PROTEINS ON GRADIENT GELS Gels that consist of a gradient of increasing polyacrylamide concentration resolve a much wider size range of proteins than standard uniform concentration gels (see Critical Parameters and Troubleshooting). The protein bands, particularly in the low-molecularweight range, are also much sharper. Unlike single-concentration gels, gradient gels separate proteins in a way that can be plotted easily to give a linear curve from 10 to 200 kDa. This facilitates molecular weight estimations.
ALTERNATE PROTOCOL 5
The quantities given below provide separating gel solution sufficient for two 0.75-mm gels (∼7 ml of each concentration) or one 1.5-mm gel (∼14 ml of each concentration). Volumes can be adjusted to accommodate gel sandwiches of different dimensions. Additional Materials (also see Basic Protocol 1) Light and heavy acrylamide gel solutions (Table 10.1.9 and Table 10.1.10) Bromphenol blue (optional; for checking practice gradient) TEMED Gradient maker (30 to 50 ml, Hoefer Pharmacia SG30 or SG50; or 30 to 100 ml, Bio-Rad 385) Tygon tubing with micropipet tip
Electrophoresis
10.1.17 Current Protocols in Protein Science
light solution heavy solution
peristaltic pump Tygon tubing
pipet tip magnetic stirrer vertical gel unit
gradient maker reservoir chamber mixing chamber interconnecting valve stir-bar outlet valve
ring stand
Figure 10.1.2 Gradient gel setup. A peristaltic pump, though not required, will provide better control.
Peristaltic pump (optional; Markson A-13002, A-34040, or A-34105 minipump) Whatman 3MM filter paper Set up the gradient maker and prepare the gel solutions 1. Assemble the magnetic stirrer and gradient maker on a ring stand as shown in Figure 10.1.2. Connect the outlet valve of the gradient maker to Tygon tubing attached to a micropipet tip that is placed over the vertical gel sandwich. If desired, place a peristaltic pump in line between the gradient maker and the gel sandwich. A peristaltic pump will simplify casting by providing a smooth flow rate.
2. Place a small stir-bar into the mixing chamber of the gradient maker (i.e., the chamber connected to the outlet). 3. Using the recipes in Table 10.1.9 and Table 10.1.10, prepare light and heavy acrylamide gel solutions. Do not add ammonium persulfate until just before use (step 7). 4. With the outlet port and interconnecting valve between the two chambers closed, pipet 7 ml of light (low-concentration) acrylamide gel solution into the reservoir chamber for one 0.75-mm-thick gradient gel. Recommended gradient ranges are 5% to 20% for most applications (to separate proteins of 10 to several hundred kilodaltons). A practice run with heavy and light solutions is recommended. Bromphenol blue should be added to the heavy solution to demonstrate linearity of the practice gradient.
5. Open the interconnecting valve briefly to allow a small amount (∼200 µl) of light solution to flow through the valve and into the mixing chamber. The presence of air bubbles in the interconnecting valve may obstruct the flow between chambers during casting. One-Dimensional SDS Gel Electrophoresis of Proteins
Deaeration is not recommended for either the light or heavy solution. Omitting the deaeration will allow polymerization to proceed more slowly, letting the gradient establish itself in the gel sandwich before polymerization takes place.
10.1.18 Current Protocols in Protein Science
Table 10.1.9
Light Acrylamide Gel Solutions for Gradient Gelsa
Acrylamide concentration of light gel solution (%)b
Stock solution 30% acrylamide/ 0.8% bisacrylamidec 4× Tris⋅Cl/SDS, pH 8.8c H2O 10% (w/v) ammonium persulfate
5
6
7
8
9
10
11
12
13
14
2.5
3.0
3.5
4.0
4.5
5.0
5.5
6.0
6.5
7.0
3.75 8.75 0.05
3.75 8.25 0.05
3.75 7.75 0.05
3.75 7.25 0.05
3.75 6.75 0.05
3.75 6.25 0.05
3.75 5.75 0.05
3.75 5.25 0.05
3.75 3.75 4.75 4.25 0.05 0.05
aTo survey proteins ≥10 kDa, 5-20% gradient gels are recommended. To expand the range between 10 and 200 kD, a
10-20% gel is recommended. bNumbers in body of table are milliliters of stock solution. Deaeration is not required. Keep solution at room temperature
prior to adding TEMED no longer than 1 hr. cSee Table 10.1.1 for preparation.
Table 10.1.10
Heavy Acrylamide Gel Solutions for Gradient Gelsa
Stock solution 30% acrylamide/ 0.8% bisacrylamidec 4× Tris⋅Cl/SDS, pH 8.8c H2O Sucrose (g) 10% (w/v) ammonium persulfateb
Acrylamide concentration of heavy gel solution (%)b 10
11
12
13
14
15
16
17
18
19
20
5.0
5.5
6.0
6.5
7.0
7.5
8.0
8.5
9.0
9.5
10.0
3.75 3.75 3.75 3.75 3.75 3.75 3.75 3.75 3.75 3.75
3.75
5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 2.25 2.25 2.25 2.25 2.25 2.25 2.25 2.25 2.25 2.25 0.05 0.05 0.05 0.05 0.05 0.05 0.05 0.05 0.05 0.05
0 2.25 0.05
aDeaeration is not recommended for gradient gels. bNumbers in body of table are milliliters of stock solution (except sucrose). Do not add the ammonium persulfate until
just before use. The heavy acrylamide will polymerize, albeit more slowly, without the addition of TEMED. Keep the heavy solution on ice after adding ammonium persulfate. cSee Table 10.1.1 for preparation.
6. Add 7 ml of heavy (high-concentration) acrylamide gel solution to the mixing chamber. Keep the heavy solution on ice until use. Once the ammonium persulfate is added to the heavy solution, it will polymerize without TEMED, albeit more slowly; keeping the solution on ice prevents this. The gel solution will come to room temperature during casting. The higher the percentage of acrylamide, the more severe the problem of premature polymerization is.
7. Add specified amount of ammonium persulfate and ∼2.3 µl TEMED per 7 ml acrylamide solution to each chamber. Mix the solutions in each chamber with a disposable pipet. Form the gradient and cast the gel 8. Open the interconnecting valve completely. Some of the heavy solution will flow back into the reservoir chamber containing light solution as the two chambers equilibrate. This will not affect the formation of the gradient. Electrophoresis
10.1.19 Current Protocols in Protein Science
9. Turn on the magnetic stirrer and adjust the rate to produce a slight vortex in the mixing chamber. 10. Open the outlet of the gradient maker slowly. Adjust the outlet valve to a flow rate of 2 ml/min. If using a peristaltic pump, calibrate the flow rate with a graduated cylinder prior to casting the gel. Some adjustment of the flow rate may be necessary during casting. If the light solution is not flowing into the mixing chamber, a bubble may be caught in the interconnecting valve. Quickly close the outlet and cover the top of the reservoir chamber with a gloved thumb. Push down with the thumb to increase the pressure in the chamber and force the air bubble out of the center valve.
11. Fill the gel sandwich from the top. Place the pipet tip against one side of the sandwich so the solution flows down one plate only. The heavy solution will flow into the sandwich first, followed by progressively lighter solution. 12. Watch as the last of the light solution drains into the outlet tube and adjust the flow rate to ensure that the last few milliliters of solution do not flow quickly into the gel sandwich and disturb the gradient. 13. Overlay the gradient gel with H2O-saturated isobutyl alcohol. Allow the gel to polymerize ∼1 hr. In this gel system, the gel will polymerize from the bottom (i.e., heavy solution) up. Because polymerization is an exothermic reaction, heat can be felt evolving from the bottom of the gel sandwich during polymerization. A sharp optical discontinuity at the gel-overlay interface indicates that polymerization has occurred. In general, 1 hr is adequate for polymerization.
14. Remove the H2O-saturated isobutyl alcohol and rinse with 1× Tris⋅Cl/SDS, pH 8.8. Cast the stacking gel (see Basic Protocol 1, steps 8 to 11). The gel can be covered with 1× Tris⋅Cl/SDS, pH 8.8, sealed in a plastic bag, and stored for up to 1 week.
Load and run the gel 15. Prepare the protein sample and protein molecular-weight-standards mixture (see Basic Protocol 1, steps 13 to 26). Load and run the gel. The gel can be stained with Coomassie blue or silver (UNIT 10.5).
16. After staining, dry the gels onto Whatman 3MM or equivalent filter paper. Gradient gels >0.75 mm thick require special handling during drying to prevent cracking. The simplest approach to drying gradient gels is to use thin gels; ≤0.75-mm gradient gels with ≤20% acrylamide solutions will dry without cracking as long as the vacuum pump is working properly and the cold trap is dry at the onset of drying. For gradient gels >0.75 mm thick, add 3% (w/v) glycerol to the final destaining solution to help prevent cracking. Another method is to dehydrate and shrink the gel in 30% methanol for up to 3 hr prior to drying. Then place the gel in distilled (Milli-Q-purified or equivalent) water for 5 min before drying.
One-Dimensional SDS Gel Electrophoresis of Proteins
10.1.20 Current Protocols in Protein Science
CASTING MULTIPLE GRADIENT GELS Casting gradient gels in a multiple gel caster has several advantages. In addition to the time savings, batch casting gives gels that are essentially identical. This is particularly important for gradient gels, where slight variations in casting technique can cause variations in protein mobility. The gels may be stored for up to 1 week after casting to ensure internal consistency from run to run during the week. Furthermore, gels with several ranges of concentrations (e.g., 5% to 20% and 10% to 20% acrylamide) can be cast and stored, giving much more flexibility to optimize separations.
SUPPORT PROTOCOL 2
Additional Materials (also see Alternate Protocol 5) Plug solution (see recipe) Light and heavy acrylamide gel solutions for multiple gradient gels (Table 10.1.11 and Table 10.1.12) TEMED H2O-saturated isobutyl alcohol Multiple gel caster (Bio-Rad, Hoefer Pharmacia) Peristaltic pump (25 ml/min) 500- or 1000-ml gradient maker (Bio-Rad, Hoefer Pharmacia) Tygon tubing Set up system and pour separating gel 1. Assemble the multiple caster as in casting multiple single-concentration gels (see Support Protocol 1, steps 1 to 3), making sure to remove the triangular space filler plugs in the bottom of the caster. The plug is used only when casting single-concentration gels.
2. Set up the peristaltic pump (Fig. 10.1.3). Using a graduated cylinder and water, adjust the flow rate so that the volume of the gradient solution plus volume of plug solution is poured in ∼15 to 18 min (∼25 ml/min). 3. Set up the gradient maker. Close all valves and place a stir-bar in the mixing chamber, which is the one with the outlet port. Attach one end of a piece of Tygon tubing to the outlet of the gradient maker. Run the other end of the tubing through the peristaltic pump and attach it to the red inlet port at the bottom of the caster. Choose a gradient maker that holds no more than four times the total volume of the gradient solution to be poured (i.e., a 1000- or 500-ml gradient maker).
4. Prepare solutions for the gradient maker (Table 10.1.11 and Table 10.1.12). 5. Add the TEMED to both heavy and light solutions (54 µl/165 ml) and immediately pour the light (low concentration) solution into the mixing chamber (the one with the port). Open the mixing valve slightly to allow the tunnel to fill and to avoid air bubbles. Close the valve again and pour the heavy (high-concentration) acrylamide solution into the reservoir chamber. 6. Start the magnetic stirrer and open the outlet valve; then start the pump and open the mixing valve. In units for casting multiple gels, acrylamide solution flows in from the bottom. To use a multiple casting unit, the light solution is placed in the mixing chamber and the heavy in the reservoir. This is the reverse of casting a single gel (see Alternate Protocol 5). Thus, light solution enters the multiple caster first, followed by progressively heavier solution. Finally, the acrylamide solution is stabilized in the multiple caster with a heavy plug solution and allowed to polymerize (see step 8 and manufacturer’s instructions). Electrophoresis
10.1.21 Current Protocols in Protein Science
Table 10.1.11
Light Acrylamide Gel Solutions for Multiple Gradient Gelsa,,b
Acrylamide concentration of light separating gel solution (%)c
Stock solution 30% acrylamide/0.8% bisacrylamided 4× Tris⋅Cl/SDS, pH 8.8d H2O 10% (w/v) ammonium persulfate
5
6
7
8
9
10
11
12
13
14
28
33
39
44
50
55
61
66
72
77
41 96 0.55
41 91 0.55
41 85 0.55
41 80 0.55
41 74 0.55
41 69 0.55
41 63 0.55
41 58 0.55
41 52 0.55
41 47 0.55
aTo survey proteins ≥10 kDa, 5-20% gradient gels are recommended. To expand the range between 10 and 200 kDa, a 10-20% gel is recommended. bRecipes produce ten 1.5-mm-thick gradient gels with 10 ml extra solution to account for losses in tubing. cNumbers in body of table are milliliters of stock solution. Deaeration is not required. Keep solution at room temperature prior to adding TEMED no longer than 1 hr. dSee Table 10.1.1 for preparation.
7. When almost all the acrylamide solution is gone from the gradient maker, stop the pump and close the mixing valve. Tilt the gradient maker toward the outlet side and remove the last milliliters of the mix. Do not allow air bubbles to enter the tubing. 8. Add the plug solution to the mixing chamber and start the pump. Make sure that no bubbles are introduced. Continue pumping until the bottom of the caster is filled with plug solution to just below the glass plates; then turn off the pump. Clamp the tubing close to the red port of the casting chamber. 9. Quickly overlay each separate gel sandwich with 100 µl H2O-saturated isobutyl alcohol. Use the same amount on each sandwich. Failure to use the same amount of overlay solution will cause the gel sandwiches to polymerize at different heights.
10. Drain off the overlay and rinse the surface of the gels with 1× Tris⋅Cl/SDS, pH 8.8.
grid maker reservoir chamber (heavy solution)
mixing chamber (light solution) outlet valve
mixing valve
outlet port
inlet port
stir-bar
stir plate
One-Dimensional SDS Gel Electrophoresis of Proteins
peristaltic pump
multiple gel caster
Figure 10.1.3 Setup for casting multiple gradient gels. Casting multiple gradient gels requires a peristaltic pump and a multiple gel caster. Gel solution is introduced through the bottom of the multiple caster.
10.1.22 Current Protocols in Protein Science
Table 10.1.12
Heavy Acrylamide Gel Solutions for Multiple Gradient Gelsa,b
Stock solution 30% acrylamide/0.8% bisacrylamided 4× Tris⋅Cl/SDS pH 8.8d H2O Sucrose (g) 10% (w/v) ammonium persulfate
Acrylamide concentration of heavy gel solution (%)c 10
11
12
13
14
15
16
17
18
19
20
55
61
66
72
77
83
88
94
99
105
110
41 55 25 0.55
41 50 25 0.55
41 44 25 0.55
41 39 25 0.55
41 33 25 0.55
41 28 25 0.55
41 22 25 0.55
41 17 25 0.55
41 11 25 0.55
41 5.5 25 0.55
41 0 25 0.55
aDeaeration is not recommended for gradient gels. bRecipes produce 10 ml extra solution to account for losses in tubing. cNumbers in body of table are milliliters of stock solution (except sucrose). Do not add the ammonium persulfate until just before use. The heavy acrylamide will polymerize, albeit more slowly, without the addition of TEMED. Keep the heavy solution on ice after adding ammonium persulfate. dSee Table 10.1.1 for preparation.
Pour stacking gel and remove gels from caster 11. Prepare and cast the stacking gel as in casting multiple single-concentration gels (see Support Protocol 1, steps 8 to 11). 12. Remove gels from the caster and clean the gel sandwiches (see Support Protocol 1, steps 12 and 13). Store gels, if necessary, according to the instructions for multiple single-concentration gels (see Support Protocol 1, step 14). ELECTROPHORESIS IN SINGLE-CONCENTRATION MINIGELS Separation of proteins in a small-gel format is becoming increasingly popular for applications that range from isolating material for peptide sequencing to performing routine protein separations. The unique combination of speed and high resolution is the foremost advantage of small gels. Additionally, small gels are easily adapted to singleconcentration, gradient, and two-dimensional SDS-PAGE procedures. The minigel procedures described are adaptations of larger gel systems.
BASIC PROTOCOL 2
This protocol describes the use of a multiple gel caster. The caster is simple to use, and up to five gels can be prepared at one time with this procedure. Single gels can be prepared using adaptations in the manufacturer’s instructions. A multiple gel caster is the only practical way to produce small linear polyacrylamide gradient gels (see Support Protocol 3). Materials Minigel vertical gel unit (Hoefer Pharmacia Mighty Small SE 250/280 or Bio-Rad Mini-Protean II) with glass plates, clamps, and buffer chambers 0.75-mm spacers Multiple gel caster (Hoefer Pharmacia SE-275/295 or Bio-Rad Mini-Protean II multicasting chamber) Acrylic plate (Hoefer Pharmacia SE-217 or Bio-Rad 165-1957) or polycarbonate separation sheet (Hoefer Pharmacia SE-213 or Bio-Rad 165-1958) 10- and 50-ml syringes Combs (Teflon, Hoefer Pharmacia SE-211A series or Bio-Rad Mini-Protean II) Long razor blade Micropipet
Electrophoresis
10.1.23 Current Protocols in Protein Science
Additional reagents and equipment for standard denaturing SDS-PAGE (see Basic Protocol 1) Pour the separating gel 1. Assemble each gel sandwich by stacking, in order, the notched (Hoefer Pharmacia) or small rectangular (Bio-Rad) plate, 0.75-mm spacers, and the larger rectangular plate. Be sure to align the spacers properly with the ends flush with the top and bottom edge of the two plates when positioning the sandwiches in the multiple gel caster (Fig. 10.1.4). The protocol described is basically for the Hoefer Pharmacia system. For other systems, make adjustments according to the manufacturer’s instructions. Alternatively, precast minigels can be purchased from a number of suppliers (see Table 10.1.3). The multiple casters from Hoefer Pharmacia have a notch in the base designed for casting gradient gels. A silicone rubber insert fills up this space when casting single-concentration gels. The Hoefer Pharmacia spacers are T-shaped to prevent slipping. The flanged edge of the spacer must be positioned against the outside edge of the glass plate. Placing a sheet of wax paper between the gel sandwiches will help separate the sandwiches after polymerization.
2. Fit the gel sandwiches tightly in the multiple gel caster. Use an acrylic plate or polycarbonate separation sheet to eliminate any slack in the chamber. Loosely fitting sandwiches in the caster will lead to unevenly cast gels, creating distortions during electrophoresis.
faceplate inlet fitting
gasket bottom
gel sandwich
high acrylamide concentration
filler glass plate caster body
low acrylamide concentration T-shaped spacer
One-Dimensional SDS Gel Electrophoresis of Proteins
Figure 10.1.4 Minigel sandwiches positioned in the multiple gel caster. Extra glass or acrylic plates or polycarbonate sheets are used to fill any free space in the caster and to ensure that the gel sandwiches are held firmly in place.
10.1.24 Current Protocols in Protein Science
3. Place the front faceplate on the caster, clamp it in place against the silicone gasket, and verify alignment of the glass plates and spacers. 4. Prepare the separating gel solution as directed in Table 10.1.1. For five 0.75-mm-thick gels, prepare ∼30 ml solution (i.e., double the volumes listed). To compute the total gel volume needed, multiply the area of the gel (e.g., 7.3 × 8.3 cm) by the thickness of the gel (e.g., 0.75 mm) and then by the number of gels in the caster. If needed, add about 4 to 5 ml of extra gel solution to account for the space around the outside of the gel sandwiches. Do not add TEMED and ammonium persulfate until just before use.
5. Fill a 50-ml syringe with the separating gel solution and slowly inject it into the caster until the gels are 6 cm high, allowing 1.5 cm for the stacking gel. 6. Overlay each gel with 100 µl H2O-saturated isobutyl alcohol. Allow the gels to polymerize for ∼1 hr. Pour the stacking gel 7. Remove the isobutyl alcohol and rinse with 1× Tris⋅Cl/SDS, pH 8.8. Stacking gels can be cast one at a time with the gel mounted on the electrophoresis unit, or all at once in the multiple caster.
8. Practice placing a comb in the gel sandwiches before preparing the stacking gel solution. Press the comb against the rectangular or taller plate so that all teeth of the comb are aligned with the opening in the gel sandwich, then insert into the sandwich. Remove combs after practicing. 9. Prepare the stacking gel solution (2 ml per gel) as directed in Table 10.1.1. Fill a 10-ml syringe with stacking gel solution and inject the solution into each gel sandwich. 10. Insert combs, taking care not to trap bubbles. Allow gels to polymerize 1 hr. 11. Remove the front faceplate. Carefully pull the gels out of the caster, using a long razor blade to separate the sandwiches. If the gels are left to polymerize for prolonged periods, they will be difficult to remove from the caster. The gels can be stored tightly wrapped in plastic wrap with the combs left in place inside a sealable bag to prevent drying for ∼1 week. Without the stacking gel, the separating gel can be stored for 2 to 3 weeks. Keep gels moist with 1× Tris⋅Cl/SDS, pH 8.8, at 4°C. Do not store gels in the multiple caster.
Prepare the sample, load the gel, and conduct electrophoresis 12. Remove the combs and rinse the sample wells with 1× SDS electrophoresis buffer. Place a line indicating the bottom of each well on the front glass plate with a marker. 13. Fill the upper and lower buffer chambers with 1× SDS electrophoresis buffer. The upper chamber should be filled to 1 to 2 cm over the notched plate. 14. Prepare the protein sample and protein-standards mixture (see Basic Protocol 1, step 12). 15. Load the sample using a micropipet. Insert the pipet tip through the upper buffer and into the well. The mark on the glass plate will act as a guide. Dispense the sample into the well. For a complex mixture, 20 to 25 µg protein in 10 µl SDS sample buffer will give a strongly stained Coomassie blue pattern. Much smaller amounts (1 to 5 µg) are required for highly purified proteins, and 10- to 100-fold less protein amount in the same volume (e.g., 10 µl is required for silver staining).
Electrophoresis
10.1.25 Current Protocols in Protein Science
16. Electrophorese samples at 10 to 15 mA per 0.75-mm gel until the dye front reaches the bottom of the gel (∼1 to 1.5 hr). 17. Disassemble the gel (see Basic Protocol 1, steps 23 to 26). Proceed with detection of proteins. SUPPORT PROTOCOL 3
PREPARING MULTIPLE GRADIENT MINIGELS Polyacrylamide gradients not only enhance the resolution of larger format gels but also greatly improve protein separation in the small format. Casting gradient minigels one at a time is not generally feasible because of the small volumes used, but multiple gel casters make it easy to cast several small gradient gels at one time. The gels are cast from the bottom in multiple casters, with the light acrylamide solution entering first. This is the opposite of casting one gel at a time, in which the heavy solution enters from the top of the gel sandwich and flows down to the bottom. Additional Materials (also see Basic Protocol 2) Plug solution (see recipe) Additional reagents and equipment for preparing gradient gels (see Alternate Protocol 5) Set up the system and prepare the gel solutions 1. Assemble minigel sandwiches in the multiple gel caster as described for single-concentration minigels (see Basic Protocol 2, steps 1 to 3). 2. Set up the 30-ml gradient maker, magnetic stirrer, peristaltic pump (optional), and Tygon tubing as in Figure 10.1.3. Connect the outlet of the 30-ml gradient maker to the inlet at the base of the front faceplate of the caster. The monomer solution will be introduced through the inlet at the bottom of the front faceplate of the caster first, followed by progressively heavier solution.
3. Prepare light (Table 10.1.9) and heavy (Table 10.1.10) acrylamide gel solutions. Use ∼12 ml of each solution for five 0.75-mm-thick minigels. Adjust volumes if a different thickness or number of gels is needed. Do not add ammonium persulfate until just before use. Deaeration is not recommended for gradient gels.
4. With the outlet and interconnecting valve closed, add the heavy solution to the reservoir chamber. Briefly open the interconnecting valve to let a small amount of heavy solution through to the mixing chamber, clearing the valve of air. 5. Fill the mixing chamber with light solution. Add 4 µl TEMED per 12 ml acrylamide solution to each chamber and mix with a disposable pipet. Form the gradient and cast the gels 6. Turn on the magnetic stirrer. Open the interconnecting valve and allow the chambers to equilibrate. Then slowly open the outlet port to allow the solution to flow from the gradient maker to the multiple caster by gravity (a peristaltic pump may be used for better control). Adjust the flow rate to 3 to 4 ml/min. Faster flow rates are possible and will also produce good gradients. However, a fast flow increases the potential for introduction of bubbles into the caster.
One-Dimensional SDS Gel Electrophoresis of Proteins
7. Close the outlet port as the last of the gradient solution leaves the mixing chamber, just before air enters the outlet tube. Fill the two chambers with plug solution and slowly open the outlet once again.
10.1.26 Current Protocols in Protein Science
8. Allow the plug solution to push the acrylamide in the caster up into the plates. Close the outlet when the plug solution reaches the bottom of the plates. A discontinuity between the bottom of the gels and the plug solution will be obvious.
9. Quickly add 100 µl H2O-saturated isobutyl alcohol to each gel sandwich. Let the gels polymerize undisturbed for ∼1 hr. 10. Prepare and pour the stacking gel (see Basic Protocol 2, steps 9 and 10). Disassemble the system 11. Disconnect the gradient maker, place the caster in a sink, and remove the front faceplate. The plug solution will drain out from bottom of the caster. 12. Remove the gels (see Basic Protocol 2, step 11). Gradient minigels can be stored as described for single-concentration minigels (see Basic Protocol 2, step 11 annotation). For instructions on preparing, loading, and running the gels, see Basic Protocol 2, steps 12 to 17.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Anode buffer 121.1 g Tris base 500 ml H2O Adjust to pH 8.9 with concentrated HCl Dilute to 5 liters with H2O Store at 4°C up to 1 month Final concentration is 0.2 M Tris⋅Cl, pH 8.9.
Cathode buffer 12.11 g Tris base 17.92 g tricine 1 g SDS Dilute to 1 liter with H2O Do not adjust pH Store at 4°C up to 1 month Final concentrations are 0.1 M Tris, 0.1 M tricine, and 0.1% (w/v) SDS.
Coomassie blue G-250 staining solution 200 ml acetic acid 1800 ml H2O 0.5 g Coomassie blue G-250 Mix 1 hr and filter (Whatman no. 1 paper) Store at room temperature indefinitely The final solution is 0.025% (w/v) Coomassie blue G-250 in 10% (v/v) acetic acid.
Phosphate/SDS electrophoresis buffer Dilute 500 ml of 4× phosphate/SDS, pH 7.2 (Table 10.1.7) with H2O to 2 liters. Store at 4°C up to 1 month. Final concentrations are 0.1 sodium phosphate (pH 7.2)/0.1% (w/v) SDS.
Electrophoresis
10.1.27 Current Protocols in Protein Science
Phosphate/SDS sample buffer, 2× (for continuous systems) 0.5 ml 4× phosphate/SDS, pH 7.2 (Table 10.1.7) 0.2 g SDS 0.1 mg bromphenol blue 0.31 g DTT 2.0 ml glycerol Add H2O to 10 ml and mix Final concentrations are 20 mM sodium phosphate, 2% (w/v) SDS, 0.001% (w/v) bromphenol blue, 0.2 M DTT, and 2% (v/v) glycerol.
Plug solution 0.125 M Tris⋅Cl, pH 8.8 (APPENDIX 2E) 50% (w/v) sucrose 0.001% (w/v) bromphenol blue Store at 4°C up to 1 month Recrystallized SDS (optional) High-purity SDS is available from several suppliers, but for some sensitive applications (e.g., protein sequencing) recrystallization is useful. Commercially available electrophoresis-grade SDS is usually of sufficient purity for most applications. Add 100 g SDS to 450 ml ethanol and heat to 55°C. While stirring, gradually add 50 to 75 ml hot H2O until all SDS dissolves. Add 10 g activated charcoal (Norit 1, Sigma) to solution. After 10 min, filter solution through Whatman no. 5 paper on a Buchner funnel to remove charcoal. Chill filtrate 24 hr at 4°C and 24 hr at −20°C. Collect crystalline SDS on a coarse-frit (porosity A) sintered-glass funnel and wash with 800 ml −20°C ethanol (reagent grade). Repeat crystallization without adding activated charcoal. Dry recrystallized SDS under vacuum overnight at room temperature. Store in a desiccator over P2O5 in a dark bottle. If proteins will be electroeluted or electroblotted for protein sequence analysis, it may be desirable to crystallize the SDS twice from ethanol/H2O (Hunkapiller et al., 1983).
SDS electrophoresis buffer, 5× 15.1 g Tris base 72.0 g glycine 5.0 g SDS H2O to 1000 ml Dilute to 1× or 2× for working solution, as appropriate Do not adjust the pH of the stock solution, as the solution is pH 8.3 when diluted. Store at 0° to 4°C until use (up to 1 month).
SDS sample buffer, 2× (for discontinuous systems) 25 ml 4× Tris⋅Cl/SDS, pH 6.8 (Table 10.1.1) 20 ml glycerol 4 g SDS 2 ml 2-ME or 3.1 g DTT 1 mg bromphenol blue Add H2O to 100 ml and mix Store in 1-ml aliquots at −70°C To avoid reducing proteins to subunits (if desired), omit 2-ME or DTT (reducing agent) and add 10 mM iodoacetamide to prevent disulfide interchange. One-Dimensional SDS Gel Electrophoresis of Proteins
10.1.28 Current Protocols in Protein Science
SDS sample buffer, 6× (for discontinuous systems) 7 ml 4× Tris⋅Cl/SDS, pH 6.8 (Table 10.1.1) 3.0 ml glycerol 1 g SDS 0.93 g DTT 1.2 mg bromphenol blue Add H2O to 10 ml (if needed) Store in 0.5-ml aliquots at −70°C Tricine sample buffer, 2× 2 ml 4× Tris⋅Cl/SDS, pH 6.8 (Table 10.1.1) 2.4 ml (3.0 g) glycerol 0.8 g SDS 0.31 g DTT 2 mg Coomassie blue G-250 Add H2O to 10 ml and mix Final concentrations are 0.1 M Tris, 24% (w/v) glycerol, 8% (w/v) SDS, 0.2 M DTT, and 0.02% (w/v) Coomassie blue G-250.
COMMENTARY Background Information Polyacrylamide gels form after polymerization of monomeric acrylamide into polymeric polyacrylamide chains and cross-linking of the chains by N,N′-methylenebisacrylamide. The polymerization reaction is initiated by the addition of ammonium persulfate, and the reaction is accelerated by TEMED, which catalyzes the formation of free radicals from ammonium persulfate. Because oxygen inhibits the polymerization process, deaerating the gel solution before the polymerization catalysts are added will speed up polymerization; deaeration is not recommended for the gradient gel protocols because slower polymerization facilitates casting of gradient gels. Precast gels for commonly used vertical minigel and standard-sized SDS-PAGE apparatuses are available from several manufacturers (Table 10.1.3). Flatbed (horizontal) isoelectric focusing (IEF) and SDS-PAGE gels are not listed. Pharmacia supplies a range of horizontal gels for a variety of applications and should be consulted for further information. When using precast gels, pay strict attention to shelf life. In general, manufacturers overrate the shelf life, and the sooner the gels are used, the better. When reasonably fresh, precast gels provide excellent resolution that is as good or better than a typical gel cast in the laboratory. The most widely used method for discontinuous gel electrophoresis is the system described by Laemmli (1970). This is the denaturing (SDS) discontinuous method used in
Basic Protocol 1. A discontinuous buffer system uses buffers of different pH and composition to generate a discontinuous pH and voltage gradient in the gel. Because the discontinuous gel system concentrates the proteins in each sample into narrow bands, the applied sample may be more dilute than that used for continuous electrophoresis. In the discontinuous system the sample first passes through a stacking gel, which has large pores. The stacking gel buffer contains chloride ions (called the leading ions) whose electrophoretic mobility is greater than the mobility of the proteins in the sample. The electrophoresis buffer contains glycine ions (called the trailing ions) whose electrophoretic mobility is less than the mobility of the proteins in the sample. The net result is that the faster migrating ions leave a zone of lower conductivity between themselves and the migrating protein. The higher voltage gradient in this zone allows the proteins to move faster and to “stack” in the zone between the leading and trailing ions. After leaving the stacking gel, the protein enters the separating gel. The separating gel has a smaller pore size, a higher salt concentration, and higher pH compared to the stacking gel. In the separating gel, the glycine ions migrate past the proteins, and the proteins are separated according to either molecular size in a denaturing gel (containing SDS) or molecular shape, size, and charge in a nondenaturing gel. Proteins are denatured by heating in the presence of a low-molecular-weight thiol (2-
Electrophoresis
10.1.29 Current Protocols in Protein Science
200
(A)
myosin
β-galactosidase –3
Molecular weight (x 10 )
100 80 60
phosphorylase albumin, bovine (B)
albumin, bovine
albumin, egg
albumin, egg
40 glyceraldehydecarbonic 3-phosphate anhydrase dehydrogenase trypsinogen
20
carbonic anhydrase
trypsin inhibitor, soybean
α-lactalbumin 10 0.2
0.4
0.6
0.8
1.0
Relative mobility (R f )
Figure 10.1.5 Typical calibration curves obtained with standard proteins separated by nongradient denaturing (SDS) discontinuous gel electrophoresis based on the method of Laemmli (1970). (A) Gel with 7% polyacrylamide. (B) Gel with 11% polyacrylamide. (Redrawn with permission from Sigma.)
One-Dimensional SDS Gel Electrophoresis of Proteins
ME or DTT) and SDS. Most proteins bind SDS in a constant-weight ratio, leading to identical charge densities for the denatured proteins. Thus, the SDS-protein complexes migrate in the polyacrylamide gel according to size, not charge. Most proteins are resolved on polyacrylamide gels containing from 5% to 15% acrylamide and 0.2% to 0.5% bisacrylamide (see Table 10.1.1). The relationship between the relative mobility and log molecular weight is linear over these ranges (Fig. 10.1.5). With the use of plots like those shown in Figure 10.1.5, the molecular weight of an unknown protein (or its subunits) may be determined by comparison with known protein standards (Table 10.1.2). Basic Protocol 1 relies on denaturing proteins in the presence of SDS and 2-ME or DTT. Under these conditions, the subunits of proteins are dissociated and their biological activities are lost. A true estimate of a protein’s molecular size can be made by comparing the relative mobility of the unknown protein to proteins in
a calibration mixture (Fig. 10.1.5). Gradient gels (Alternate Protocol 5) simplify molecularweight determinations by producing a linear relationship between log molecular weight of the protein and log % T over a much wider size range than single-concentration gels. Although percent acrylamide monomer is a more common measure of gel concentration, % T, the percentage of total monomer (acrylamide plus bisacrylamide) in the solution or gel, is used for molecular weight calculations in gradient gels. The % T is estimated assuming the acrylamide gradient is linear. For example, proteins in the gel shown in Figure 10.1.6 were separated in a 5% to 20% T acrylamide gradient. The % T of the point halfway through the resolving gel is 12.5% T. Simply plotting log molecular mass versus distance moved into the gel (or Rf) also produces a relatively linear standard curve over a fairly wide size range. If two proteins have identical molecular sizes, they more than likely will not be resolved with one-dimensional SDS-PAGE, and two-di-
10.1.30 Current Protocols in Protein Science
Molecular mass (kDa)
Molecular mass (kDa)
205 116 97.4 66
66
45 36
45
29 24
29
20.1 14.2
Figure 10.1.6 Separation of membrane proteins by 5.1% to 20.5% polyacrylamide gradient SDS-PAGE. Approximately 30 µl of 1× SDS sample buffer containing 30 µg of Alaskan pea (Pisum sativum) membrane proteins was loaded in wells of a 14 × 14–cm, 0.75-mm-thick gel. Standard proteins were included in the outside lanes. The gel was run at 4 mA for ∼15 hr.
mensional SDS-PAGE should be considered (UNIT 10.4). Unusual protein compositions can cause anomalous mobilities during electrophoresis (see Critical Parameters and Troubleshooting), but similar-sized proteins of widely different amino acid composition or structure may still be resolved from one another using one-dimensional SDS-PAGE. Purified protein complexes or multimeric proteins consisting of subunits of different molecular size will be resolved into constituent polypeptides. Comparison of the protein bands obtained under nonreducing and reducing conditions provides information about the molecular size of disulfide cross-linked component subunits. The individual polypeptides can be isolated by electroelution (Smith, 1993) or electroblotting (UNIT 10.8), and the amino acid sequences can be determined. Both the tricine (Schagger and von Jagow, 1987) and the modified Laemmli (Okajima et al., 1993) peptide separation procedures presented here (Alternate Protocols 1 and 2) are simple to set up and provide resolution down to 5 kDa. To separate peptides below 5 kDa, the tricine procedure must be modified by preparing a 16.5% T, 2.7% C resolving gel that uses a 10% T spacer gel between the stacking and resolving gel (Schagger and von Jagow, 1987).
% C is the percentage of cross-linker (bisacrylamide) in the total monomer (acrylamide plus bisacrylamide). Continuous electrophoresis, where the same buffer is used throughout the tank and gel, is popular because of its versatility and simplicity. The phosphate system described in Alternate Protocol 3 is based on that of Weber et al. (1972). Although unable to produce the highresolution separations of the discontinuous SDS-PAGE procedures, continuous SDSPAGE uses fewer solutions with one basic buffer and no stacking gel. Artifacts are also less likely to occur in continuous systems. Pepsin, for example, migrates anomalously on Laemmli-based discontinuous SDS-PAGE but has the expected mobility after electrophoresis in the phosphate-based continuous system described here. This is also true of cross-linked proteins. Multiple gel casting (Support Protocols 1 to 3) is appropriate when gel-to-gel consistency is paramount or when the number of gels processed exceeds 5 a week. The variety of multiple gel casters, gradient makers, and inexpensive pumps available from major suppliers simplifies the process of casting gels in the laboratory. Alternatively, precast gradient gels are availElectrophoresis
10.1.31 Current Protocols in Protein Science
able for most major brands of gel apparatuses (Table 10.1.3). Minigels (Basic Protocol 2) are generally considered to be in the 8 × 10–cm size range, although there is considerable variation in exact size. Every technique that is used on larger systems can be translated with little difficulty into the minigel format. This includes standard and gradient SDS-PAGE and separations for immunoblotting and peptide sequencing. Twodimensional SDS-PAGE electrophoresis also adapts well, but here the limitation of separation area becomes apparent; for high-resolution separations, large-format gels are required. Gradient minigels (Support Protocol 3) are popular due to the combination of separation range and resolution (Matsudaira and Burgess, 1978). They are particularly useful for separation of proteins prior to peptide sequencing. Mylar support (Gel Bond) provides a practical way of casting, running, and, staining extremely thin gels. When gels <0.75 mm thick are used, reagents have much better access both into and out of the gel, reducing staining time in both Coomassie blue and silver staining (UNIT 10.5). Double and broadened images caused by differential migration of the protein across the thickness of the gel are minimized, improving resolution.
Critical Parameters and Troubleshooting
One-Dimensional SDS Gel Electrophoresis of Proteins
If an electrophoretically separated protein will be electroeluted or electroblotted (UNIT 10.8) for sequence analysis, the highest-purity reagents available should be used. If necessary, SDS can be purified by recrystallization following the procedure given in Reagents and Solutions. If the gels polymerize too fast, the amount of ammonium persulfate should be reduced by one-third to one-half. If the gels polymerize too slowly or fail to polymerize all the way to the top, fresh ammonium persulfate should be used or the amount of ammonium persulfate should be increased by one-third to one-half. The overlay should be added slowly down the spacer edge to prevent the overlay solution from crashing down and disturbing the gel interface. After a separating gel is poured, it may be stored with an overlay of the same buffer used in the gel. Immediately prior to use, the stacking gel should be poured; otherwise, there will be a gradual diffusion-driven mixing of buffers between the two gels, which will cause a loss of resolution. The protein of interest should be present in
0.2- to 1-µg amounts in a complex mixture of proteins if the gel will be stained by Coomassie blue. Typically, 30 to 50 µg of a complex protein mixture in a total volume of <20 µl is loaded on a 0.75-mm-thick slab gel (16 cm, 10 wells). When casting multiple gradient gels, all bubbles in the outlet tubing of the gradient maker should be eliminated. If air bubbles get into the outlet tube, they may flow into the caster and then up through the gradient being poured, causing an area of distortion in the polymerized gel. Air bubbles are not so great a problem when casting single gradient gels from the top. As the gels are cast, the stirrer must be slowed so that the vortex in the mixing chamber does not allow air to enter the outlet. Uneven heating of the gel causes differential migration of proteins, with the outer lanes moving more slowly than the center lanes (called smiling). Increased heat transfer eliminates smiling and can be achieved by filling the lower buffer chamber with buffer all the way to the level of the sample wells, by maintaining a constant temperature between 10° to 20°C, and by stirring the lower buffer with a magnetic stirrer. Alternatively, the heat load can be decreased by running at a lower current. If the tracking dye band is diffuse, prepare fresh buffer and acrylamide monomer stocks. If the protein bands are diffuse, increase the current by 25% to 50% to complete the run more quickly and minimize band diffusion, use a higher percentage of acrylamide, or try a gradient gel. Lengthy separations using gradient gels generally produce good results (Fig. 10.1.6). Check for possible proteolytic degradation that may cause loss of high-molecularweight bands and create a smeared banding pattern. If there is vertical streaking of protein bands, decrease the amount of sample loaded on the gel, further purify the protein of interest to reduce the amount of contaminating protein applied to the gel, or reduce the current by 25%. Another cause of vertical streaking of protein bands is precipitation, which can sometimes be eliminated by centrifuging the sample or by reducing the percentage of acrylamide in the gel. Proteins can migrate faster or slower than their actual molecular weight would indicate. Abnormal migration is usually associated with a high proportion of basic or charged amino acids (Takano et al., 1988). Other problems can occur during isolation and preparation of the protein sample for electrophoresis. Proteolysis of proteins during cell fractionation by endog-
10.1.32 Current Protocols in Protein Science
enous proteases can cause subtle band splitting and smearing in the resulting electrophoretogram (electrophoresis pattern). Many endogenous proteases are very active in SDS sample buffers and will rapidly degrade the sample; thus, first heating the samples to 70° to 100°C for 3 min is recommended. In some cases, heating to 100°C in sample buffer will cause selective aggregation of proteins, creating a smeared layer of Coomassie blue–stained material at the top of the gel (Gallagher and Leonard, 1987). To avoid heating artifacts and also prevent proteolysis, the use of specific protease inhibitors during protein isolation and/or lower heating temperatures (70° to 80°C) have been effective (Dhugga et al., 1988). Although continuous gels suffer from poor band sharpness, they are less prone to artifacts caused by aggregation and protein cross-linking. If streaking or aggregation appear to be a problem with the Laemmli system, then the same sample should be subjected to continuous SDS-PAGE to see if the problem is intrinsic to the Laemmli gel or the sample. If the protein bands spread laterally from gel lanes, the time between applying the sample and running the gel should be reduced in order to decrease the diffusion of sample out of the wells. Alternatively, the acrylamide percentage should be increased in the stacking gels from 4% to 4.5% or 5% acrylamide, or the operating current should be increased by 25% to decrease diffusion in the stacking gel. Use caution when adding 1× SDS electrophoresis buffer to the upper buffer chamber. Samples can get swept into adjacent wells and onto the top of the well arm. If the protein bands are uneven, the stacking gel may not have been adequately polymerized. This can be corrected by deaerating the stacking gel solution thoroughly or by increasing the ammonium persulfate and TEMED concentrations by one-third to one-half. Another cause of distorted bands is salt in the protein sample, which can be removed by dialysis (APPENDIX 3B), gel filtration (UNIT 8.3), or precipitation. Skewed protein bands can be caused by an uneven interface between the stacking and separating gels, which can be corrected by starting over and being careful not to disturb the separating gel while overlaying with isobutyl alcohol. If a run takes too long, the buffers may be too concentrated or the operating current too low. If the run is too short, the buffers may be too dilute or the operating current too high. If double bands are observed, the protein
may be partially oxidized or partially degraded. Oxidation can be minimized by increasing the 2-ME concentration in the sample buffer or by preparing a fresh protein sample. If fewer bands than expected are observed and there is a heavy protein band at the dye front, increase the acrylamide percentage in the gel.
Anticipated Results Polyacrylamide gel electrophoresis done under denaturing and reducing conditions should resolve any two proteins, except two of identical size. Resolution of proteins in the presence of SDS is a function of gel concentration and the size of the proteins being separated. Under nondenaturing conditions, the biological activity of a protein will be maintained. Comparison of reducing and nonreducing denaturing gels can also provide valuable information about the number of disulfide crosslinked subunits in a protein complex. If the subunits are held together by disulfide linkages, the protein will separate in denaturing gels as a complex or as smaller sized subunits under n on redu cin g or redu cin g con ditio ns, respectively. However, proteins separated on nonreducing denaturing gels appear more diffuse and exhibit less overall resolution than those separated on reducing gels. Gradient gels provide superior protein-band sharpness and resolve a larger size range of proteins, making them ideal for most types of experiments in spite of being more difficult to prepare. Molecular-weight calculations are simplified because of the extended linear relationship between size and protein position in the gel. Increased band sharpness of both highand low-molecular-weight proteins on the same gel greatly simplifies survey experiments, such as gene expression studies where the characteristics of the responsive protein are not known. Furthermore, the increased resolution dramatically improves autoradiographic analysis. Preparation of gradient gels is straightforward, although practice with gradient solutions containing dye is recommended. The gradient gels can be stored for several days at 0° to 4°C before casting the stacking gel.
Time Considerations Preparation of separating and stacking gels requires 2 to 3 hr. Gradient gels generally take 5 min to cast singly. Casting multiple singleconcentration gels requires an additional 10 min for assembly. Casting multiple gradient gels takes 15 to 20 min plus assembly time. It takes 4 to 5 hr to run a 14 × 14–cm, 0.75-mm
Electrophoresis
10.1.33 Current Protocols in Protein Science
gel at 15 mA (70 to 150 V), and 3 to 4 hr to run a 0.75-mm gel at 20 mA (80 to 200 V). Overnight separations of ∼12 hr require 4 mA per 0.75-mm gel. It takes 4 to 5 hr to run a 1.5-mm gel at 30 mA. Electrophoresis is normally performed at 15° to 20°C, with the temperature held constant using a circulating water bath. For air-cooled electrophoresis units, lower currents and thus longer run times are recommended. It takes ∼1 hr to run a 0.75-mm minigel at 20 mA (100 to 120 V). Separation times are not significantly different for gradient minigels.
Schagger, H. and von Jagow, G. 1987. Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa. Anal. Biochem. 166:368-379.
Literature Cited Dhugga, K.S., Waines, J.G., and Leonard, R.T. 1988. Correlated induction of nitrate uptake and membrane polypeptides in corn roots. Plant Physiol. 87:120-125.
Takano, E., Maki, M., Mori, H., Hatanaka, N., Marti, T., Titani, K., Kannagi, R., Ooi, T., and Murachi, T. 1988. Pig heart calpastatin: Identification of repetitive domain structures and anomalous behavior in polyacrylamide gel electrophoresis. Biochemistry 27:1964-1972.
Gallagher, S.R. and Leonard, R.T. 1987. Electrophoretic characterization of a detergent-treated plasma membrane fraction from corn roots. Plant Physiol. 83:265-271.
Weber, K., Pringle, J.R., and Osborn, M. 1972. Measurement of molecular weights by electrophoresis on SDS-acrylamide gel. Methods Enzymol. 26:3-27.
Hunkapiller, M.W., Lujan, E., Ostrander, F., and Hood, L.E. 1983. Isolation of microgram quantities of proteins from polyacrylamide gels for amino acid sequence analysis. Methods Enzymol. 91:227-236. Laemmli, U.K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680-685. Matsudaira, P.T. and Burgess, D.R. 1978. SDS microslab linear gradient polyacrylamide gel electrophoresis. Anal. Biochem. 87:386-396. Okajima, T., Tanabe, T., and Yasuda, T. 1993. Nonurea sodium dodecyl sulfate-polyacrylamide gel electrophoresis with high-molarity buffers for the separation of proteins and peptides. Anal. Biochem. 211:293-300.
Sigma. Molecular weight markers for proteins kit (Technical Bulletin MWS-877L). Sigma Chemical Company, St. Louis, Mo. Smith, J.A. 1993. Electroelution of proteins from stained gels. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 10.5.1-10.5.5. John Wiley & Sons, New York.
Key Reference Hames, B.D. and Rickwood, D. (eds.) 1990. Gel Electrophoresis of Proteins: A Practical Approach, 2nd ed. Oxford University Press, New York. An excellent book describing gel electrophoresis of proteins.
Contributed by Sean R. Gallagher Hoefer Pharmacia Biotech San Francisco, California
One-Dimensional SDS Gel Electrophoresis of Proteins
10.1.34 Current Protocols in Protein Science
One-Dimensional Isoelectric Focusing of Proteins in Slab Gels
UNIT 10.2
Separation of proteins in complex mixtures for further analysis or purification can be achieved by isoelectric focusing (IEF), a special type of electrophoresis in which movement of proteins through pores in a polyacrylamide gel matrix is modulated by the presence of a pH gradient created by soluble ampholytes. The types of reagents and equipment used are similar to those needed for standard Laemmli sodium dodecyl sulfate (SDS) gels (UNIT 10.1) and IEF tube gels (UNIT 10.4). Use of IEF in the slab-gel format provides the advantage of permitting the analysis of large numbers of samples in a single experiment. The Basic Protocol describes isoelectric focusing of polypeptides in urea-containing polyacrylamide slab gels. A pH gradient forms when ampholytes (small charged organic molecules) migrate through the gel in response to an electric field. Denaturing of proteins in special solubilizing buffer allows maximum exposure of amino acid side chains that contribute to a protein’s isoelectric point (pI), as well as dissociation of multisubunit proteins. Separation on the basis of pI values produces bands of polypeptides that can be electrotransferred to suitable membranes for further analysis, for example, by staining (UNIT 10.8) or immunodetection. If samples have been radiolabeled, they can be visualized by fluorography. This unit contains support protocols detailing special conditions for application of denaturing IEF slab gels for electrotransfer (Support Protocol 1) and fluorography (Support Protocol 2) in which a scintillator is incorporated into the gel after completion of the run to facilitate detection of radioactively labeled polypeptides. ISOELECTRIC FOCUSING IN SLAB GELS UNDER DENATURING CONDITIONS
BASIC PROTOCOL
The preparation and running of an IEF slab gel containing nonionic detergent and urea is presented. An acrylamide solution supplemented with urea, Triton X-100, and ampholytes is used to cast a slab gel; the stacking gel normally used in protein electrophoresis is omitted. Protein samples (preferably salt-free) are solubilized in buffer containing urea, 2-mercaptoethanol (2-ME), a nonionic detergent such as Triton X-100, and ampholytes before electrophoresis. A variety of electrophoresis units may be used with this protocol, but gel lengths of 15 to 20 cm and thicknesses of 0.75 to 1.0 mm are recommended. Thicker gels can be run, but consume correspondingly larger quantities of expensive ampholytes, and cannot be run at high power at the onset of the run. Soluble ampholytes (Hoefer Pharmacia, Oxford Glycosystems, Serva) in the desired pH range are commercially available. The choice of pH range is dictated by the isoelectric points of the proteins to be resolved. This protocol results in a rather broad pH gradient (∼pH 4.5 to 7.5) and can be modified as needed, provided the final ampholyte concentrations (∼2% w/v) are maintained as indicated. Materials Urea (any high-grade urea will usually prove satisfactory) 10% (v/v) Triton X-100 Acrylamide stock solution (see recipe) Ampholytes pH 5-7 Ampholytes pH 3.5-10 10% (w/v) ammonium persulfate (store 500-µl aliquots of freshly prepared stock at −20°C) Contributed by Hidde L. Ploegh Current Protocols in Protein Science (1995) 10.2.1-10.2.8 Copyright © 2000 by John Wiley & Sons, Inc.
Electrophoresis
10.2.1 CPPS
TEMED (N,N,N′-N′-tetramethylenediamine) 20 mM H3PO4 Protein samples to be analyzed IEF solubilization buffer (see recipe) Overlay solution: IEF solubilization buffer diluted 1:3 (v/v) with H2O 20 mM NaOH (freshly made, degassed immediately prior to use) Electrophoresis apparatus (e.g., Studier type or equivalent) with glass plates, spacers, clamps, casting stand, and buffer chambers Side-arm flask Comb Power supply (capable of delivering at least 1000 V and 10 W total power) Hamilton syringe or mechanical pipet CAUTION: Acrylamide is a neurotoxin; wear gloves when handling solutions. Cast the isoelectric focusing gel 1. Prepare the electrophoresis apparatus by thoroughly cleaning glass plates. Assemble gel mold using spacers and clamps and mount in casting stand. Agarose (1% w/v) is a useful sealing agent. It is dispensed from a freshly boiled solution using a Pasteur pipet. Be sure that the hot agarose flows well between glass plates and spacer. Some minor spillage of agarose into the inside of the gel mold may be tolerated without affecting polymerization of the gel or quality of the separation. Most types of equipment for vertical slab gel electrophoresis can be used. Dimensions of gels are not specified, as these will differ widely between laboratories.
2. Prepare acrylamide gel solution in a side-arm flask; for 10 ml gel mix use the following: 5.5 g urea (9.1 M final) 2 ml 10% (v/v) Triton X-100 (2% v/v final) 2 ml Milli-Q water 1.35 ml acrylamide stock solution 0.4 ml ampholytes pH 5-7 0.1 ml ampholytes pH 3.5-10. To promote rapid dissolution of urea, the gel solution may be swirled in a 37°C water bath. Avoid overheating to minimize decomposition of urea. The total volume of the gel mix to be prepared is easily calculated from the dimensions of the gel and the thickness of the spacer. For a 200 × 150 × 1–mm gel, a total of 30 ml gel mix would be required; multiply quantities given above by three to obtain the appropriate amounts. The side-arm flask size (volume) should be at least 3× the volume of the amount of gel mix to be accommodated.
3. Degas solution briefly (up to 5 min). Because oxygen inhibits the polymerization process, it is important that the gel solution be deaerated before the polymerization catalysts are added.
4. Add 20 µl of 10% (w/v) ammonium persulfate and 10 µl TEMED to initiate polymerization. Mix well by swirling. One-Dimensional Isoelectric Focusing of Proteins in Slab Gels
5. Pour gel all the way to the top of the glass plates. Wear gloves when handling unpolymerized gel solutions. Some of the gel mix will overflow from the mold. Remove spillage with tissue paper. By filling the mold to the
10.2.2 Current Protocols in Protein Science
top, the insertion of the comb without causing bubbles to get trapped underneath the wells (step 6) is facilitated.
6. Insert comb, being careful to avoid air bubbles. Allow gel to polymerize ∼45 min. The gel should be used shortly after polymerization is complete (the same day it is cast). In view of the decomposition of urea in aqueous solution, storage of poured gels is not recommended.
Perform the electrophoretic separation 7. Remove comb and place gel-plate sandwich in electrophoresis apparatus. Fill lower buffer chamber (anode) with 20 mM H3PO4. 8. Prepare protein samples to be analyzed in IEF solubilization buffer. Apply samples in gel wells. Fill any empty wells with a similar volume of IEF solubilization buffer. Samples may be applied with a Hamilton syringe or a mechanical pipet. The volume of the sample loaded is not particularly important, but it is recommended that no greater than 60% to 70% of the well’s capacity be loaded so that overlay solution may be subsequently applied. Generally, sample volumes are <100 ìl. Samples are loaded immediately adjacent to one another, without the need for empty lanes in between. Acrylamide teeth should be in good contact with both glass plates to eliminate the potential of spillover between wells. When in doubt, the integrity of the sample wells can be checked by first loading the overlay solution (which is colored); spillage from a freshly loaded well into an adjacent one indicates a faulty sample well, not to be used. If the wells are intact, samples can then be layered underneath the overlay using a Hamilton-type syringe. If unequal sample volumes are added to wells, the lane with the larger volume will spread during electrophoresis and constrict adjacent lanes, causing distortions. Preparing samples at approximately the same concentration and loading an equal volume in each well will ensure that all lanes are the same width and that the polypeptides run evenly. Fill any extra lanes (i.e., those not receiving a protein sample) with a similar volume of sample buffer, such that all wells are filled. The total amount of protein that may be loaded in a volume of 100 ìl should not exceed 200 ìg, and should be loaded over a total cross-section of 10 to 15 mm2. The sample capacity of IEF gels is considerably greater than that of SDS-PAGE gels (UNIT 10.1), as it is an equilibrium separation method. The guideline given is recommended, but larger amounts of protein can in all likelihood be analyzed without serious loss of resolution. Samples should be salt-free, if possible, or of low ionic strength, to obtain reproducible separations. Immune complexes can be dissociated by the addition of IEF solubilization buffer. Most proteins in solution can be analyzed successfully when diluted with IEF solubilization buffer (4 parts buffer to 1 part sample). Samples that have been denatured in solubilization buffer for SDS-PAGE can likewise be diluted with IEF solubilization buffer and analyzed in parallel.
9. Apply overlay solution with a syringe or mechanical pipet, taking care not to disturb the sample layer. Use ∼1⁄5 to 1⁄3 of the sample volume for the overlay. It is most convenient to add the tracking dye to the overlay solution, as this will generate a colored layer that separates sample from the upper reservoir electrolyte.
10. Using a Pasteur pipette, gently layer 20 mM NaOH over the samples, until the NaOH solution is flush with the top of the glass plate of the gel mold. Fill the upper reservoir with the remainder of the NaOH solution. 11. Connect power supply and apply voltage, starting at 20 V/cm. Limit voltage to 50 V/cm. CAUTION: The voltages and currents used during electrophoresis are dangerous and potentially lethal. Safety considerations are given in the introduction to UNIT 10.1.
Electrophoresis
10.2.3 Current Protocols in Protein Science
The cathode is at the top (NaOH-containing reservoir) and the anode is at the bottom (H3PO4-containing reservoir). The most convenient manner of running the gel is to use a power supply with a constant power setting and voltage limitation. The power is adjusted to a limit such that the desired starting voltage is obtained. Over the course of the run, the current will drop and the voltage will increase until the desired preset value is attained.
12. Run gel 13 to 16 hr (for a gel of a separating distance of ∼20 cm). An overnight run is convenient. Over the course of the run, bromphenol blue included with samples will migrate the entire length of the gel and accumulate at the acidic end of the gel as a greenish-yellow band. When gels are run for longer times (>16 hr), no bromphenol blue may be discernible at all by the end of the run.
13. On termination of the run disconnect the power supply and disassemble the electrophoresis apparatus. Remove one of the glass plates from the gel-plate sandwich. The IEF gel will present a rippled appearance. The ripples correspond to positions of high local ampholyte concentrations that osmotically attract water. The ripple pattern is usually a good indicator of whether the gel ran straight and may allow one to gauge edge effects, “smiling” (curvature of migrating band), or the presence of samples with salt content drastically different from that of adjacent samples.
14. Process the gel, if desired, for staining (UNIT 10.5), electroblotting (see Support Protocol 1), or fluorography (see Support Protocol 2). Invert the glass plate with the gel still attached and hold it directly over the dish containing the appropriate solution. Using a spatula, peel off one corner of the gel, and continue loosening the gel until it begins to detach spontaneously from the glass plate. While holding the glass plate horizontally over the liquid-filled dish, minimize the distance that the gel will have to “fall” into the receptacle to reduce the risk of breakage. SUPPORT PROTOCOL 1
ELECTROBLOTTING FROM DENATURING ISOELECTRICFOCUSING SLAB GELS The Basic Protocol produces gels with separated polypeptides that can be subjected to standard electrotransfer procedures (UNIT 10.7). It is necessary, however, to remove the nonionic detergent (included in the IEF gel recipe) from the gel as it interferes with a number of commonly used staining protocols; the nonionic detergent also very significantly reduces the amount of protein transferred to membranes in electroblotting experiments. It is also necessary to bind SDS to the proteins. Both tasks are accomplished by repeated washings of the gel as follows: The gel is submerged in a shallow dish containing 50% methanol/1% (w/v) SDS/5 mM Tris⋅Cl (pH 8.0); the volume of wash solution should be 10× the volume of the gel. The gel is washed five times, 10 min each time, changing the solutions between washes. A convenient way to restrain the gel while decanting the used wash solution is to place paper towels or filter paper of a size that approximates that of the gel directly on the surface of the gel. Alternatively, liquid may be removed by aspiration; again, paper towels or filter paper may be used to hold the gel in the container. After the washes, the gel is ready for transfer as described in UNIT 10.7.
One-Dimensional Isoelectric Focusing of Proteins in Slab Gels
10.2.4 Current Protocols in Protein Science
FLUOROGRAPHY OF DENATURING ISOELECTRIC-FOCUSING SLAB GELS
SUPPORT PROTOCOL 2
Slab gels containing separated polypeptides radiolabeled with 3H, 14C, or 35S can be analyzed by fluorography. The use of the scintillant PPO (2,5-diphenyloxazole) is recommended to increase detectability of these weak β emitters and make the gels relatively easy to handle. Materials DMSO DMSO/PPO solution (see recipe) Polyacrylamide gel containing polypeptides of interest Film cassette Film Intensifying screen CAUTION: Solutions of PPO in DMSO readily permeate the skin, causing the PPO to precipitate subcutaneously. It is therefore imperative that gloves be worn at all times while handling DMSO/PPO solutions. 1. Fill a shallow dish with DMSO, using at least 10× the volume of the gel. Agitate for 20 min. 2. Remove DMSO by decantation or aspiration, and repeat step 1, using the same volume of DMSO. At this stage, the gel often presents an opaque appearance, with lines running perpendicular to the direction of the electrical field. We have not determined the identity of the material that forms this banding pattern, but it almost certainly relates to the distribution of the ampholytes. Like the “ripple” pattern alluded to in the Basic Protocol (step 13), it is a very good indicator of the quality of the separation and would reveal distortions due to overloaded lanes or high salt content in some of the samples.
3. Remove DMSO as in step 2, then add DMSO/PPO. Use at least 10× the volume of the gel. Agitate for 45 to 60 min. The DMSO/PPO solution may be reused multiple times and can be replenished by addition of the appropriate amount of PPO (assume that PPO consumed is equivalent to the gel volume). Spent DMSO/PPO solution may be diluted with water to precipitate remaining PPO. The PPO is then recovered by filtration and dried. In this manner, the (expensive) PPO can be used to maximum advantage.
4. Pour off and save DMSO/PPO. Add tap water (use at least 10× the gel volume, or as much as the container will allow) to precipitate the PPO in the gel, imparting mechanical strength to the gel. Agitate for 10 min. 5. Replace water. Wash at least two more times, replacing water at 20 min intervals. 6. Dry gel and place in film cassette. Cover with film, close cassette, and expose at −70°C. A brief (≤1 msec) exposure of the film to a flash of light from a photographic flash unit or stroboscope hypersensitizes the film, allowing a linear relationship to be drawn between the film image and the amount of radioactivity in the sample.
Electrophoresis
10.2.5 Current Protocols in Protein Science
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Acrylamide stock solution: 30% (w/v) acrylamide, 1.6% (w/v) bisacrylamide Use the recipe in Table 10.1.1, increasing the amount of bisacrylamide to 1.6 g per 100 ml final volume. Any high-quality electrophoresis-grade acrylamide should be satisfactory. It may be advisable to treat the solution with activated charcoal. Use 5 g charcoal (Merck) per 500 ml acrylamide stock solution and remove the charcoal by filtration. Filter the final solution through a 0.45-µm filter. Store ≤30 days at 4°C in the dark. DMSO/PPO solution Dissolve 220 g PPO in 800 ml DMSO (technical-grade DMSO may be used). Store at room temperature. IEF solubilization buffer 5.7 g urea 2.0 ml 10% (v/v) Triton X-100 0.2 ml ampholytes pH 3.5-10 0.5 ml 2-mercaptoethanol H2O to 10 ml final volume Bromphenol blue may be added to samples at a final concentration of 0.01% (w/v) to improve visibility of proteins during loading and overlay. Alternatively, it may be added to the overlay solution. COMMENTARY Background Information
One-Dimensional Isoelectric Focusing of Proteins in Slab Gels
Electrophoretic methods for separating polypeptides are powerful analytical tools. Separation according to size is achieved by electrophoresis in polyacrylamide matrices in the presence of sodium dodecyl sulfate (SDS; see UNIT 10.1). Separation of polypeptides according to charge is accomplished by isoelectric focusing: a pH gradient stabilized by an electric field allows the polypeptides to migrate until they reach their isoelectric point. The combination of separation according to charge in one dimension and separation according to molecular weight in the second dimension has become a widespread analytical method capable of resolving complex protein mixtures (see UNIT 10.4). Separation in a single dimension is informative when the complexity of the polypeptide mixture to be resolved is small, or when methods for detecting the specific polypeptide of interest are available. For example, when a complex mixture of proteins is resolved by separation under nondenaturing conditions (UNIT 10.3), gel overlay methods that measure enzyme activity or ligand-binding activity can be used to detect the protein of interest. The most commonly used separation methods,
however, involve prior denaturation of the protein mixture, so that multisubunit proteins dissociate and each polypeptide migrates as a single species, according to its mass or charge. In this case, specific detection of polypeptides is usually accomplished by transferring the resolved polypeptides to a suitable membrane (UNIT 10.7) and using antibodies to detect the polypeptide of interest (UNIT 10.10), or lectins to reveal the position of glycoproteins. In many instances the complexity of the mixture of proteins to be resolved is reduced by prior purification. In some cases, for example, radiolabeled protein samples are subjected to immunoprecipitation, and the proteins thus recovered are resolved electrophoretically. Other methods of labeling (e.g., biotinylation) could be used, bearing in mind that the attachment of label may affect the charge of the protein or result in the introduction of charge heterogeneity in an otherwise homogeneous sample. When examining the post-translational modifications on a single polypeptide backbone (e.g., by glycosylation, Chapter 12, or phosphorylation, Chapter 13), it may be advantageous to analyze the samples by isoelectric focusing instead of SDS-PAGE, especially if
10.2.6 Current Protocols in Protein Science
the modifications in question result in a charge difference (e.g., addition of sialic acids to protein-bound glycans; phosphorylation; and sulfation). Such charge changes are often difficult to detect by SDS-PAGE, because whereas SDSPAGE is sensitive to charge differences in proteins to be resolved, such changes affect mobility in an often unpredictable manner, or result in very minor differences in migration. The comparison of multiple samples by two-dimensional methods, although feasible, is clearly a rather cumbersome procedure with the inherent pitfalls of variation between individual separations. The protocol described here for one-dimensional IEF is an adaptation of the method developed by O’Farrell (1975) for two-dimensional gel electrophoresis. In O’Farrell’s technique, the IEF gel is cast in a glass tube, and later serves as the starting sample for the second-dimension gel. We have adapted gels of a composition similar to those described by O’Farrell to the slab-format gels. Most equipment currently in use for SDS-PAGE should be satisfactory (UNIT 10.1). No cooling of the electrophoresis unit is required, as long as the power applied to the gel is held within reasonable limits. The technique as described here allows high resolution of membrane proteins and has been used to detect isoelectric variants of membrane proteins not separable by size, to detect N-linked glycan modifications such as the addition of sialic acids, and to detect phosphorylated species of membrane proteins (Eichholtz et al., 1992; Benaroch et al., 1995). The method also lends itself to immunoblotting procedures, provided the high Triton X-100 concentrations in the gel are reduced and SDS is added by extensive washing of the gel in a methanol/SDS-containing solution. The method described is particularly useful when comparing polypeptides that may show only minute differences in charge: such differences would be more difficult to detect on two-dimensional separations, unless sophisticated methods are used to standardize gel pouring, electrophoresis, and workup of the gels (see Garrels, 1989). In addition, this method may be used to resolve short peptides, provided they can be detected (radiolabeled peptides).
Critical Parameters and Troubleshooting It is important that the samples to be analyzed be salt-free, if possible. For immune complexes, this is easily accomplished by a final rinse of the complex adsorbed onto protein
A–Sepharose, or some similar matrix, with water. Dissociation of the samples from the Sepharose is then achieved by addition of IEF solubilization buffer. Staining of IEF gels as described in this protocol requires that the proteins are initially fixed in the gel using 20% (w/v) TCA (note that exposure of skin to TCA can cause blistering and other damage). This is followed by repeated washes in 10% (w/v) acetic acid/30% methanol. The gel can then be stained with most conventional SDS-PAGE stains (see UNIT 10.5). Given the extreme variability of available electrophoresis equipment, one consideration is the size of the gel to be used. We have found a length of 15 to 20 cm to be satisfactory. Shorter gels may induce loss of the gel: the gel may simply slide from between the glass plates, as the combination of Triton and urea in the gel makes for poor adhesion to the glass plates. To avoid this, one may increase the surface area of the gel (pour larger gels), or assemble the gel mold with spacers such that the gel produced will taper towards the bottom (i.e., 1 cm narrower at the bottom than at the top). Occasionally one may find that a gel has polymerized in an irregular fashion. Because the acrylamide gel serves as an anticonvection medium to stabilize the pH gradient, and does not contribute to resolution of the polypeptide mixture, such irregularities are usually inconsequential. If the gel does not polymerize all the way to the top and hence no depressions of adequate size are present to form the sample wells, one may prepare a small batch of fresh gel solution and repour the top part separately. In this case, care should be taken to remove any remaining liquid from the top of the first gel, using blotting paper. The concentrations of ammonium persulfate and TEMED may be increased twofold to obtain rapid polymerization of the second, top layer. Any discontinuity in the gel that results from this procedure is unlikely to affect the separation. In this manner, an initially defective gel can be rescued with minimal loss of costly ampholytes. Samples in SDS-PAGE solubilization buffer may be used for IEF, provided the same amount of buffer is present in each lane of the gel. Try to keep the amount of SDS sample buffer to be loaded as low as possible (30 µl or less). Dilute these samples with IEF solubilization buffer as much as the capacity of the sample well will allow (preferably more than a twofold dilution), and make sure that every sample well in the gel will receive the same amount of SDS
Electrophoresis
10.2.7 Current Protocols in Protein Science
sample buffer (including lanes that will not receive protein samples). Polypeptides that are rather basic will be poorly resolved on the gradients specified in this protocol. This may be compensated, but only in part by using ampholytes of a higher pH range. Although so-called nonequilibrium IEF gels, in which the samples are loaded at the acidic end of the gel and the run is terminated before equilibrium has been reached, have been successful when run in glass tubes, we have generally not been successful in adapting slab gels to nonequilibrium IEF conditions, except for runs of very short duration to resolve peptides (Heemels et al., 1993). When using equipment that can be cooled, care should be taken not to cool the gels to the point where urea comes out of solution in the gel. Nonetheless, it is recommended that the temperature of the gel not be allowed to rise over 40°C (i.e., warm to the touch). Even though the high concentration of ampholytes in the gel is likely to scavenge most of the decomposition products, decomposition of urea in aqueous solution accelerates at higher temperatures and is a cause for concern (i.e., results in carbamylation of proteins). For fluorography of gels, the use of the DMSO/PPO protocol is recommended, because the final product, the PPO-impregnated gel, is easier to handle and less fragile than gels processed using commercially available fluorography reagents such as EN3HANCE (DuPont NEN) and similar products.
Anticipated Results For representative examples of the types of separation that can be achieved using these protocols, the reader is referred to Eichholtz et al. (1992) and Benaroch et al. (1995).
Time Consideration Preparing the gel mix, pouring the gel, and completing the polymerization process can be accomplished in <2 hr. Gels ∼20 cm in length
are conveniently electrophoresed overnight. Because the separation is essentially taken to equilibrium, variations in running time have minor impact on the resolution actually achieved. Runs should not be extended beyond 18 hr, however, as cathodic drift may begin to seriously affect the pH gradient in the basic portion of the gel. The time required for electroblotting will depend somewhat on the conditions used, but preparation of the gel for transfer should take 1 hr. The entire fluorography procedure requires 2.5 to 3 hr, not including drying of the gel, which can take 0.5 to 1 hr depending on the equipment used.
Literature Cited Benaroch, P., Yilla, M., Raposo, G., Ito, K., Miwa, K., Geuze, H.J., and Ploegh, H.L. 1995. How MHC Class II molecules reach the endocytic pathway. EMBO J. 14:37-49. Eichholtz, T.E., Vossebeld, P., Van Overveld, M., and Ploegh, H.L. 1992. Activation of protein kinase C accelerates internalization of transferrin receptor but not major histocompatibility complex class I molecules, independent of their phosphorylation status. J. Biol. Chem. 267:22490-22495. Garrels, J.I. 1989. The QUEST system for quantitative analysis of two-dimensional gels. J. Biol. Chem. 264:5269-5282. Heemels, M.T., Schumacher, T.N.M., Wonigeit, K., and Ploegh, H.L. 1993. Different peptides are translocated by allelic variants of the transporter associated with antigen presentation (TAP). Science 262:2059-2063. O’Farrell, P.H. 1975. High resolution two-dimensional electrophoresis of proteins. J. Biol. Chem. 250:4007-4021.
Key References Benaroch, et al., 1995. See above. Eichholtz et al., 1992. See above. These contain representative examples of good quality separations of membrane proteins attained by the IEF methods described here.
Contributed by Hidde L. Ploegh Massachussetts Institute of Technology Cambridge, Massachusetts
One-Dimensional Isoelectric Focusing of Proteins in Slab Gels
10.2.8 Current Protocols in Protein Science
One-Dimensional Electrophoresis Using Nondenaturing Conditions
UNIT 10.3
Nondenaturing or “native” electrophoresis—i.e., electrophoresis in the absence of denaturants such as detergents and urea—is an often-overlooked technique for determining the native size, subunit structure, and optimal separation of a protein. Because mobility depends on the size, shape, and intrinsic charge of the protein, nondenaturing electrophoresis provides a set of separation parameters distinctly different from mainly size-dependent denaturing sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE; UNITS 10.1 & 10.4) and charge-dependent isoelectric focusing (IEF; UNITS 10.2 & 10.4). Two protocols are presented below. Continuous PAGE (Basic Protocol) is highly flexible, permitting cationic and anionic electrophoresis over a full range of pH. The discontinuous procedure (Alternate Protocol) is limited to proteins negatively charged at neutral pH but provides high resolution for accurate size calibration. CONTINUOUS ELECTROPHORESIS IN NONDENATURING POLYACRYLAMIDE GELS
BASIC PROTOCOL
Separation of proteins by nondenaturing electrophoresis requires the same type of equipment used for denaturing slab gels (UNIT 10.1) and is adaptable to a range of gel sizes (e.g., from 7.3 × 8.3–cm minigels to 14 × 16–cm full-size gels) and matrix types (e.g., single-concentration and gradient gels). This protocol outlines straightforward procedures for making acrylamide solutions, casting separating gels (stacking gels are omitted), loading samples, and conducting electrophoresis. Continuous systems, although flexible, do not give the high-resolution separation found in discontinuous systems (see Alternate Protocol). Separation in a continuous system (i.e., in which the same buffer is used for preparing acrylamide solutions and filling electrophoresis chambers) is governed by pH, and this protocol describes four types of buffers useful over discrete ranges from pH 3.7 to pH 10.6. Use of unadjusted acetic acid gel buffer can extend the range to pH 2.0. The choice of pH and thus the buffer system depends on the protein being studied (i.e., its isoelectric point) and often must be determined empirically. In general, the system should be between pH 5.0 and 8.0 for optimal results. Extremes of pH can lead to precipitation or denaturation of the protein. Acrylamide concentrations are empirically determined, but the higher the percent acrylamide, the sharper the protein bands. It is important to include native protein standards in the electrophoresis runs. Several manufacturers supply standards for isoelectric focusing that are also suitable for native electrophoresis. The standards have a range of isoelectric points and will carry a net positive, negative, or zero charge depending on the pH of the gel system. Alternatively, Sigma supplies a standard kit that is useful for calculating molecular weights under neutral pH, nondenaturing conditions. The samples and standard proteins, should be used at concentrations of ∼1 to 2 µg/µl. Materials 4× acetic acid gel buffer (200 mM acetic acid, pH 3.7 to 5.6; see recipe) 4× phosphate gel buffer (400 mM sodium phosphate, pH 5.8 to 8.0; see recipe) 4× Tris gel buffer (200 mM Tris⋅Cl, pH 7.1 to 8.9; see recipe) 4× glycine gel buffer (200 mM glycine, pH 8.6 to 10.6; see recipe) 300 mM sodium sulfite (0.38 g in 10 ml H2O; used in acetic acid gel preparation) Protein samples to be analyzed Native protein standards
Electrophoresis
Contributed by Sean R. Gallagher
10.3.1
Current Protocols in Protein Science (1995) 10.3.1-10.3.11 Copyright © 2000 by John Wiley & Sons, Inc.
CPPS
Electrophoresis buffer: appropriate 4× gel buffer diluted to 1× with H2O 75-ml side-arm flask (used in gel preparation) Additional reagents and equipment for gel electrophoresis (UNIT 10.1) and staining proteins in gels (UNIT 10.5) Prepare the gel 1. Assemble the glass-plate sandwich of the gel electrophoresis unit and secure it to the casting stand. Either single-concentration or gradient gels can be used in the minigel or standard-size format. Gradient gels will enhance the band sharpness of the separated proteins.
2. Prepare acrylamide solutions according to the recipes in Table 10.3.1, Table 10.3.2, Table 10.3.3, or Table 10.3.4, adding the ammonium persulfate and TEMED just before use. Deaeration of the solution before the polymerization catalysts are added will speed polymerization by removing inhibitory oxygen, but is not generally required. The pH used depends on many factors. The most important are the pI values of both the protein of interest and any contaminants, as well as protein mobility and protein solubility. Determining which pH and thus which buffer system to use is largely empirical. However, extremes of pH (<4.0 and >9.0) can lead to denaturation and should be avoided. Prior knowledge of the pI of a protein (UNITS 10.2 & 10.4) allows determination of the net charge under the separation conditions (i.e., if gel pH < protein pI, the protein
Table 10.3.1 to 5.6b
Recipes for Acetic Acid Nondenaturing Polyacrylamide Gelsa: pH range 3.7
Stock solutionc 30% acrylamide/0.8% bisacrylamide 300 mM sodium sulfitee 4× acetic acid gel buffer H2O 10% (w/v) ammonium persulfatee TEMEDf
Final acrylamide concentration in gel (%)d 5
7.5
10
12.5
15
17.5
20
6.7
10
13.3
16.8
20
23.32
26.6
0.4 10 22.58 0.3
0.4 10 19.28 0.3
0.4 10 15.98 0.3
0.4 10 12.48 0.3
0.4 10 9.28 0.3
0.4 10 5.96 0.3
0.4 10 2.68 0.3
0.02
0.02
0.02
0.02
0.02
0.02
0.02
Preparation of gel In a 75-ml side-arm flask, mix 30% acrylamide/0.8% bisacrylamide solution (see Table 10.1.1), 300 mM sodium sulfite, 4× acetic acid gel buffer (see Reagents and Solutions), and H2O. If desired to speed polymerization, degas under vacuum ∼5 min. Add 10% ammonium persulfate and TEMED. Swirl gently to mix. Use immediately.
One-Dimensional Electrophoresis Using Nondenaturing Conditions
aThe recipes produce 40 ml gel solution, which is adequate for one gel of dimensions 1.5 mm × 14 cm × 16 cm or two gels of dimensions 0.75 mm × 14 cm × 16 cm. bThe pH range can be extended to ∼2.0 (the pH of acetic acid) by using unadjusted acetic acid in place of 4× acetic acid gel buffer, although there is little buffering capacity at this pH. cAll reagents and solutions used in the protocol must be prepared with Milli-Q-purified water or equivalent. dUnits of numbers in table body are milliliters. The desired percentage of acrylamide in the gel solution depends on the molecular size of the protein being separated. eMust be freshly made. Sodium sulfite is needed for efficient polymerization at acid pH. fAdded just before polymerization.
10.3.2 Current Protocols in Protein Science
Table 10.3.2 to 8.0
Recipes for Phosphate Nondenaturing Polyacrylamide Gelsa: pH range 5.8
Stock solutionb 30% acrylamide/0.8% bisacrylamide 4× phosphate gel buffer H2O 10% (w/v) ammonium persulfated TEMEDe
Final acrylamide concentration in gel (%)c 5
7.5
10
12.5
15
17.5
20
6.7
10
13.3
16.8
20
23.32
26.6
10 23.08 0.2
10 19.78 0.2
10 16.48 0.2
10 12.98 0.2
10 9.78 0.2
10 6.46 0.2
10 3.18 0.2
0.02
0.02
0.02
0.02
0.02
0.02
0.02
Preparation of gel In a 75-ml side-arm flask, mix 30% acrylamide/0.8% bisacrylamide solution (see Table 10.1.1), 4× phosphate gel buffer (see Reagents and Solutions), and H2O. If desired, degas under vacuum ∼5 min to speed polymerization. Add 10% ammonium persulfate and TEMED. Swirl gently to mix. Use immediately. aThe recipes produce 40 ml gel solution, which is adequate for one gel of dimensions 1.5 mm × 14 cm × 16 cm or two gels of dimensions 0.75 mm × 14 cm × 16 cm. bAll reagents and solutions used in the protocol must be prepared with Milli-Q-purified water or equivalent. cUnits of numbers in table body are milliliters. The desired percentage of acrylamide in the gel solution depends on the molecular size of the protein being separated. dMust be freshly made. eAdded just before polymerization.
will have a net positive charge; if gel pH > protein pI, the protein will be negatively charged).
3. Pour gel to 2 cm from the top of the gel mold and insert the comb. Avoid trapping air bubbles under the comb teeth. Air bubbles will cause small semicircular depressions in the well and lead to distortions in the protein banding pattern.
4. Allow gel solution to polymerize 1 to 2 hr. Polymerization is indicated by a sharp optical discontinuity around the wells.
Prepare samples and load the wells 5. Solubilize the protein sample to be analyzed using 5% (w/v) sucrose in water or dilute (1 to 5 mM) gel buffer if possible. Also prepare native protein standards. The concentration of protein will vary depending on the sample complexity and detection method. For Coomassie blue staining of highly enriched samples such as the standards, use 1 to 2 mg/ml (1 to 2 ìg/ìl). For more complex mixtures, use 5 to 10 mg/ml (5 to 10 ìg/ìl). Load 10- to 100-fold less for silver staining. In general, samples should be loaded in a minimum volume, preferably 10 to 20 ìl for 0.75- and 1.5-mm-thick gels, respectively. With thin gels, this means using a more concentrated protein sample.
6. Remove comb carefully and rinse wells with electrophoresis buffer (appropriate 4× gel buffer diluted to 1×). Rinsing with electrophoresis buffer is needed to remove residual unpolymerized acrylamide monomer, which will continue to polymerize after comb removal, creating uneven wells that may interfere with sample loading. Electrophoresis
10.3.3 Current Protocols in Protein Science
Table 10.3.3
Recipes for Tris Nondenaturing Polyacrylamide Gelsa: pH range 7.1 to 8.9
Stock solutionb 30% acrylamide/0.8% bisacrylamide 4× Tris gel buffer H2O 10% (w/v) ammonium persulfated TEMEDe
Final acrylamide concentration in gel (%)c 5
7.5
10
12.5
15
17.5
20
6.7
10
13.3
16.8
20
23.32
26.6
10 23.08 0.2
10 19.78 0.2
10 16.48 0.2
10 12.98 0.2
10 9.78 0.2
10 6.46 0.2
10 3.18 0.2
0.02
0.02
0.02
0.02
0.02
0.02
0.02
Preparation of gel In a 75-ml side-arm flask, mix 30% acrylamide/0.8% bisacrylamide solution (see Table 10.1.1), 4× Tris gel buffer (see Reagents and Solutions), and H2O. If desired, degas under vacuum ∼5 min to speed polymerization. Add 10% ammonium persulfate and TEMED. Swirl gently to mix. Use immediately. aThe recipes produce 40 ml gel solution, which is adequate for one gel of dimensions 1.5 mm × 14 cm × 16 cm or two gels of dimensions 0.75 mm × 14 cm × 16 cm. bAll reagents and solutions used in the protocol must be prepared with Milli-Q-purified water or equivalent. cUnits of numbers in table body are milliliters. The desired percentage of acrylamide in the gel solution depends on the molecular size of the protein being separated. dMust be freshly made. eAdded just before polymerization.
7. Fill wells with electrophoresis buffer. If desired, prerun gel. The gel can be prerun at this point to remove any charged material such as ammonium persulfate from the gel prior to loading the sample. Assemble the electrophoresis unit and fill the buffer chambers with electrophoresis buffer. Run the gel at 300 V until the current no longer drops. This should take ∼30 min. Disassemble the unit, discard the buffer, and proceed to the next step.
8. Carefully load up to 10 µl (0.75-mm gels) or 20 µl (1.5-mm gels) sample per lane as a thin layer at the bottom of the wells. Load control wells with native protein standards. Add an equal volume of electrophoresis buffer to any empty wells to prevent spreading of adjoining lanes. Mobility (Rf) markers require special consideration in nondenaturing gel systems. For cationic systems, cytochrome c (pI ∼9 to 10, 5 to 10 ìg/lane) works well as an Rf marker. Bromphenol blue (10 ìg/ml) is a suitable marker for anionic systems. The marker should be included in the solubilization buffer with the sample.
Perform the separation 9. Assemble the gel unit, fill the upper and lower buffer chambers with electrophoresis buffer, and connect the unit to the power supply. Set current to 30 mA for a 1.5-mm-thick gel (15 mA for a 0.75-mm-thick gel).
One-Dimensional Electrophoresis Using Nondenaturing Conditions
If the protein is negatively charged under the separation conditions, then the standard SDS-PAGE electrode polarity should be used (proteins will migrate to the anode or positive electrode). If the protein is positively charged, then the electrodes should be reversed at the power supply (i.e., red high-voltage cable to the black output and black high-voltage lead to the red output) so the positively charged protein migrates to the negative cathode.
10.3.4 Current Protocols in Protein Science
Table 10.3.4 Recipes for Glycine Nondenaturing Polyacrylamide Gelsa: pH range 8.6 to 10.6
Stock solutionb 30% acrylamide/0.8% bisacrylamide 4× glycine gel buffer H2O 10% (w/v) ammonium persulfated TEMEDe
Final acrylamide concentration in gel (%)c 5
7.5
10
12.5
15
17.5
20
6.7
10
13.3
16.8
20
23.32
26.6
10 23.08 0.2
10 19.78 0.2
10 16.48 0.2
10 12.98 0.2
10 9.78 0.2
10 6.46 0.2
10 3.18 0.2
0.02
0.02
0.02
0.02
0.02
0.02
0.02
Preparation of gel In a 75-ml side-arm flask, mix 30% acrylamide/0.8% bisacrylamide solution (see Table 10.1.1), 4× glycine gel buffer (see Reagents and Solutions), and H2O. If desired, degas under vacuum ∼5 min to speed polymerization. Add 10% ammonium persulfate and TEMED. Swirl gently to mix. Use immediately. aThe recipes produce 40 ml gel solution, which is adequate for one gel of dimensions 1.5 mm × 14 cm × 16 cm or two gels of dimensions 0.75 mm × 14 cm × 16 cm. bAll reagents and solutions used in the protocol must be prepared with Milli-Q-purified water or equivalent. cUnits of numbers in table body are milliliters. The desired percentage of acrylamide in the gel solution depends on the molecular size of the protein being separated. dMust be freshly made. eAdded just before polymerization.
10. Continue electrophoresis until the Rf marker reaches the bottom of the gel. For minigels, electrophoresis will take 1 to 2 hr. Standard gels require 4 to 6 hr runs.
11. Turn off power supply, disassemble the unit, and remove gel from sandwich. 12. Stain the gel according to UNIT 10.5. NATIVE DISCONTINUOUS ELECTROPHORESIS AND GENERATION OF MOLECULAR WEIGHT STANDARD CURVES (FERGUSON PLOTS)
ALTERNATE PROTOCOL
One straightforward approach to discontinuous native electrophoresis is to leave out the SDS and reducing agent (DTT) from the standard Laemmli SDS-PAGE protocol (UNIT 10.1). The gels are prepared as described in UNIT 10.1 except that the sample buffer contains no SDS or DTT (samples are not heated), and the gel and electrophoresis solutions are prepared without SDS. This protocol illustrates the separation of standard proteins at four different concentrations of acrylamide and how the results are used to construct a molecular weight standard curve (Ferguson plot) without the need for SDS. By plotting relative mobility against %T (percentage weight per volume of acrylamide plus bisacrylamide in the gel), the presence of isoforms and multimeric proteins can also be detected. Materials 4× Tris⋅Cl, pH 8.8 (1.5 M Tris⋅Cl; APPENDIX 2E) 4× Tris⋅Cl, pH 6.8 (0.5 M Tris⋅Cl; APPENDIX 2E) Protein sample of interest 2× Tris/glycerol sample buffer (see recipe)
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10.3.5 Current Protocols in Protein Science
Native protein standards (e.g., Sigma nondenatured protein molecular weight kit) Tris/glycine electrophoresis buffer (see recipe) 1. Assemble the glass-plate sandwich of the gel electrophoresis unit and place it in the casting stand. 2. Prepare and cast the gels, using 4× Tris⋅Cl, pH 8.8, for the separating gel and 4× Tris⋅Cl, pH 6.8, for the stacking gel instead of the SDS-containing counterparts (Table 10.1.1). Prepare a minimum of four separate gels at different acrylamide concentrations. A typical range of concentrations is from 5% to 12.5% (e.g., 5%, 7.5%, 10%, 12.5% acrylamide). As with SDS-PAGE, typical gel thickness ranges from 0.75 to 1.5 mm. The 0.75-mm-thick gels are recommended because they offer a combination of fast staining and high resolution.
3. Mix protein sample of interest 1:1 with 2× Tris/glycerol sample buffer to attain a 1 to 2 µg/µl final concentration. Also prepare native protein standards. Remove comb, rinse wells, and load 10 to 20 µl per well for Coomassie brilliant blue staining, and 1 to 2 µl for silver staining. Some proteins must be dissolved in 50 mM NaCl or water to become fully solubilized prior to mixing with the sample buffer (Sigma, 1986).
4. Assemble gel electrophoresis unit, using Tris/glycine electrophoresis buffer to fill both lower and upper buffer chambers. Connect power supply and conduct electrophoresis. Conditions for separation are the same as for discontinuous SDS-PAGE (i.e., 30 mA for 1.5-mm-thick gels, 15 mA for 0.75-mm-thick gels). For standard-size gels the separation takes 4 to 5 hr; for minigels, 1 to 2 hr is required. Alternatively, standard gels can be run at 4 to 6 mA/gel overnight.
5. After the bromphenol blue Rf marker has reached the bottom of the gel, fix and stain the proteins in the gels according to UNIT 10.5. Estimate relative mobilities of the proteins. An example of a stained gel is shown in Figure 10.3.1. A minimum of four gel concentrations is recommended. In Figure 10.3.1, Sigma native molecular weight standards were separated on 5%, 7.5%, 10%, and 12.5% acrylamide gels (5.1%, 7.7%, 10.3%, and 12.8% T, respectively).
6. Plot log Rf against gel concentration (% T) (Fig. 10.3.2). Determine the slope of Kr using linear regression. 7. Plot −log Kr of the curves from step 6 against log molecular weight of the standards (Fig. 10.3.3). Determine the slope using linear regression. 8. Estimate the size of the standards and unknowns from the generated curve (Ferguson plot). Use the curve generated by linear regression to estimate the predicted size of the standard for comparison to the actual size stated by the supplier. This indicates the accuracy of the curve. The −log Kr value (y) of the unknown is then used to predict the molecular weight (x).
One-Dimensional Electrophoresis Using Nondenaturing Conditions
10.3.6 Current Protocols in Protein Science
Figure 10.3.1 Separation of native protein standards under nondenaturing conditions by discontinuous polyacrylamide gel electrophoresis at 12.8% T. Approximately 20 µg protein was loaded per lane on a 1.5-mm-thick, 16-cm-long gel. The gel and samples were prepared according to the Alternate Protocol and were electrophoresed 16 hr at 6 mA. Proteins were stained with Coomassie blue.
A
B
Log Rf
1.0
0.10 4
6
8
10 %T
12
14 4
6
8
10
12
14
%T
Figure 10.3.2 Effect of %T on the relative mobility of several native proteins. The relative mobility (Rf) of the standard proteins shown in Figure 10.3.1 was determined at four different gel concentrations and plotted as log Rf against %T. See text for details. (A) BSA monomer (squares) and dimer (circles); (B) carbonic anhydrase isoforms.
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0.3 r 2 = 0.9863
–Log K r
0.2
0.1 0.09 0.08 0.07 0.06 0.05 0.04 0.03 0.02
0.01 14
29
45 66
132
545
Log molecular weight
Figure 10.3.3 Native molecular weight standard curve. The −log slope of the line (Kr) from Figure 10.3.2 is plotted against log molecular weight of the standards.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Acetic acid gel buffer, 4× (200 mM acetic acid, pH 3.7 to 5.6) 11.49 ml glacial acetic acid Add to 500 ml H2O Adjust to pH 3.7 to 5.6 with 1 M NaOH Add H2O to 1000 ml Store up to 1 month at 4°C Glycine gel buffer, 4× (200 mM glycine, pH 8.6 to 10.6) 15.01 g glycine Add to 500 ml H2O Adjust to pH 8.6 to 10.6 with 1 M NaOH Add H2O to 1000 ml Store up to 1 month at 4°C Phosphate gel buffer, 4× (400 mM sodium phosphate, pH 5.8 to 8.0) 55.2 g NaH2PO4⋅H2O Add to 500 ml H2O Adjust to pH 5.8 to 8.0 with 1 M NaOH Add H2O to 1000 ml Store up to 1 month at 4°C One-Dimensional Electrophoresis Using Nondenaturing Conditions
10.3.8 Current Protocols in Protein Science
Tris gel buffer, 4× (200 mM Tris⋅Cl, pH 7.1 to 8.9) 24.23 g Tris base Add to 500 ml H2O Adjust to pH 7.1 to 8.9 with 1 M HCl Add H2O to 1000 ml Store up to 1 month at 4°C 2× Tris/glycerol sample buffer 25 ml 0.5 M Tris⋅Cl, pH 6.8 (APPENDIX 2E) 20 ml glycerol 1 mg bromphenol blue Add H2O to 100 ml and mix Store in 1-ml aliquots up to 6 months at −70°C Tris/glycine electrophoresis buffer 15.1 g Tris base 72.0 g glycine H2O to 5000 ml Store up to 1 month at 4°C COMMENTARY Background Information Under nondenaturing conditions, in which protein activity, native charge, and conformation are sustained, electrophoretic separation depends on many factors, including size, shape, and charge. Characteristics such as intrinsic molecular weight (i.e., in the absence of denaturation), the number of isoforms, and the presence of multimeric proteins can be determined with nondenaturing electrophoresis (often called native electrophoresis). The most important application of nondenaturing electrophoresis is in the determination of native protein size (Alternate Protocol). Ferguson plots (reviewed by Andrews, 1986) were first described for starch gels (Ferguson, 1964) and then for polyacrylamide gels (Hedrick and Smith, 1968). Ferguson plots are prepared by separating proteins under nondenaturing conditions at several different gel concentrations. As the acrylamide concentration (%T) is increased, the relative mobility (Rf) of the protein decreases. This is plotted as log relative mobility (on the y axis) versus %T (on the x axis) to produce a straight line. The slope of this line is referred to as the retardation coefficient (Kr) and measures how effectively a protein is slowed by the increase in %T. Large proteins will be retarded much more significantly than small proteins with increasing gel concentration, with the size of the protein being proportional to the slope of the curve. Once the Kr plots for several size standards are generated (Fig. 10.3.2), the Kr values are plotted against the molecular weight of the standard proteins
using a log-log graph (Fig. 10.3.3). The retardation coefficient also depends on a large number of other variables including temperature, pH, buffer type, ionic strength, and %C (percent bisacrylamide cross-linker). All these factors should be kept constant for a given experiment. In addition to estimated size, other types of information are available from the Ferguson plots (Rodbard and Chrambach, 1971; Andrews, 1986). For example, if two components differ in size but have the same charge per unit size (e.g., for a multimeric protein with identical subunits), curves similar to those illustrated by BSA monomer and dimer (Fig. 10.3.2A) will result. Note that when the curve is extrapolated back to 0% T, it is evident that the monomer and the dimer have similar free solution mobilities. Furthermore, as the acrylamide concentration is increased, the separation between the two also increases. However, if two proteins have similar sizes but different amounts of charge, the curves will be parallel on the log plot. This is illustrated by the carbonic anhydrase isoforms (Fig. 10.3.2B). In this example, optimal separation of the isoforms occurs at the lower concentrations of acrylamide as this is a log plot. Further applications of nondenaturing electrophoresis include preparative purification. The pH of the gel determines the net charge on the protein. Below its isoelectric point (pI) a protein will have a net positive charge, whereas above its pI it will have a net negative charge. In general, most proteins will be positively charged at pH 2.0 to 4.0; above pH 8.0, most
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proteins will be negatively charged. As these general guidelines imply, the majority of proteins have isoelectric points between pH 4.0 and 8.0. There are, however, many exceptions. A protein with a highly acidic isoelectric point (e.g., pepsin, with a pI of 2.2) will remain negatively charged at a pH down to its pI. Although a full range of pH options are given, extremes of pH (<4.0 and >9.0) should be avoided, if possible, to minimize denaturation or inactivation. By picking an appropriate electrophoresis pH, it is possible to ensure that the protein of interest will be either positively or negatively charged so that it can be selectively run into the gel, excluding a large proportion of contaminants that have the opposite or no charge. Furthermore, the pH conditions determine the mobility and can be adjusted to ensure a difference in mobility between the protein of interest and contaminants. Continuous gel systems (Basic Protocol) offer the most flexibility in terms of separation design. The pH can be tailored so that a given protein has a net positive, neutral, or negative charge. Depending on the polarity of the gel, the protein can then be excluded from or electrophoresed into the gel. Discontinuous gels have a fixed pH and gel polarity. For the nondenaturing Laemmli gel presented in the Alternate Protocol, the proteins of interest should have an isoelectric point of ≤7.0 in order to be negatively charged so that they move into the gel. Other more basic and more acidic discontinuous gel systems can be found in Hames (1990) and Schägger (1994).
Critical Parameters
One-Dimensional Electrophoresis Using Nondenaturing Conditions
The success of a gel separation under nondenaturing conditions depends on many factors, and two of the most important are protein solubility and isoelectric point. The protein must be soluble at the pH and the ionic strength of the gel, and it must be charged at that pH in order to move into the gel. If the protein experiences a pH below its isoelectric point, then it will have a net positive charge and will move to the negative electrode. Note that this is the reverse of typical SDS-PAGE. If the protein experiences a pH above its isoelectric point, it will have a net negative charge and will migrate to the positive electrode. Solubility is a complex issue. Membrane-associated and other hydrophobic proteins are difficult to separate by nondenaturing electrophoresis (Schägger, 1994). Nonionic detergents at concentrations up to 1% and solubilizing reagents such as urea (4 to 8 M) can be used,
but these reagents, especially urea, are likely to alter the protein’s conformation and most likely the isoelectric point by exposing previously hidden charged groups. If detergent or urea must be included for solubilization, the minimum required to solubilize the protein should be used. Schägger (1994) lists several nonionic detergents suitable for solubilization. Among the more popular are octylglucoside and CHAPS. In general, detergents should be used near the critical micelle concentration (CMC; 0.001% to 1%, depending on the detergent). The gel concentration has a dramatic effect on resolution and should be optimized in order to achieve the best separation and band sharpness. In general, increasing the %T will improve band sharpness.
Troubleshooting Gel polymerization at acid pH can be problematic, and sodium sulfite is needed for efficient polymerization (Andrews, 1986). Both the ammonium persulfate and the sodium sulfite must be freshly made, and the highest quality reagents available should be used. Furthermore, the gel solutions should be at room temperature for effective polymerization. If the protein does not enter the gel and no stained material is present at the well surface, try reversing the polarity of the electrode. If material concentrates at the top of the gel, try lowering the acrylamide concentration. Stained material at the top of the gel may also indicate poor solubilization, and increasing the ionic strength of the solubilization buffer or adding a small amount of urea and/or nonionic detergent may be required.
Anticipated Results Proteins will resolve depending on their solubility and native charge at the chosen pH. Ideally, a distinct band representing the protein of interest will be visible. If the band is diffuse, then increasing the gel concentration or using a gradient gel will improve resolution. If the band is not visible, then the protein may be at its isoelectric point or may have moved out of the gel because it had the wrong charge. Continuous gel systems, although more versatile, will give lower resolution than discontinuous gels. Detergents or other solubilizing agents such as urea may be needed to fully solubilize and resolve the protein. Once the conditions that resolve the protein are determined, Ferguson plots will give indications of multiple subunit structure, native size, and potential isoform relationships.
10.3.10 Current Protocols in Protein Science
Time Considerations Separations will be complete when the tracking dye or protein reaches the bottom of the gel. For minigels, this generally takes 1 to 2 hr, using 15 or 30 mA for 0.75- or 1.5-mmthick gels, respectively. Standard-format gels require 4 to 5 hr at 15 or 30 mA for 0.75- or 1.5-mm-thick gels, respectively. Standardformat gels can also run overnight at 4 to 6 or 8 to 12 mA for 0.75- or 1.5-mm-thick gels, respectively.
Literature Cited
Hedrick, J.L. and Smith, A.J. 1968. Size and charge isomer separation and estimation of molecular weights of proteins by disc gel electrophoresis. Arch. Biochem. Biophys. 126:155-164. Rodbard, D. and Chrambach, A. 1971. Estimation of molecular radius, free mobility, and valence using polyacrylamide gel electrophoresis. Anal. Biochem. 40:95-134. Schägger, H. 1994. Native gel electrophoresis. In A Practical Guide to Membrane Protein Purification (G. Von Jagow and H. Schägger, eds.) pp. 81-104. Academic Press, San Diego. Sigma. 1986. Nondenatured protein molecular weight marker kit (Technical Bulletin No. MKR137). Sigma Chemical Company, St. Louis, Mo.
Andrews, A.T. 1986. Electrophoresis: Theory, Techniques and Biochemical and Clinical Applications, 2nd ed. Oxford University Press, New York.
Key Reference
Ferguson, K.A. 1964. Starch-gel electrophoresis— application to the classification of pituitary proteins and polypeptides. Metabolism 13:985-1002.
Covers a variety of electrophoretic techniques, including nondenaturing electrophoresis and Ferguson plots.
Hames, D. 1990. One-dimensional polyacrylamide gel electrophoresis. In Gel Electrophoresis of Proteins: A Practical Approach, 2nd ed. (B.D. Hames and D. Rickwood, eds.) pp. 1-147. Oxford University Press, New York.
Andrews, 1986. See above.
Contributed by Sean R. Gallagher Hoefer Pharmacia Biotech San Francisco, California
Electrophoresis
10.3.11 Current Protocols in Protein Science
Two-Dimensional Gel Electrophoresis
UNIT 10.4
Two-dimensional gel electrophoresis combines two different electrophoretic separating techniques in perpendicular directions to provide a much greater separation of complex protein mixtures than either of the individual procedures. The most common two-dimensional technique uses isoelectrofocusing (IEF) in a tube gel (see Basic Protocol 1) followed by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) in a perpendicular direction (Basic Protocol 3). This combination of isoelectric point (pI) and size separation is the most powerful tool for protein separations currently available. After staining, proteins appear on the final two-dimensional gel as round or elliptical spots instead of the rectangular bands observed on one-dimensional gels. Although the total separating power of large-format two-dimensional gels is estimated to be as high as 5000 spots per gel, in practice a single two-dimensional separation of a complex mixture such as a whole-cell or tissue extract may produce 1000 to 2000 well-resolved spots when a sensitive detection method is used. The most common IEF procedures are based on the use of soluble ampholytes, which are relatively small organic molecules with various isoelectric points and buffering capacities. The pH gradient for IEF gels is produced when the soluble ampholytes migrate in the gel matrix until they reach their isoelectric point. Because stable pH gradients outside the pH 3.0 to 8.0 range are difficult to create, alternative protocols using nonequilibrium conditions are required to resolve proteins with pI values below 3.0 to 4.0 (see Alternate Protocol 1 for acidic proteins) or above 8.0 (see Alternate Protocol 2 for basic proteins). One of the more important limitations of soluble ampholytes is the difficulty in obtaining highly reproducible pH profiles, especially when very narrow pH ranges are needed. An increasingly attractive alternative to soluble ampholytes is the use of immobilized pH gradient (IPG) gels (see Basic Protocol 2). In this system, the buffering side chains are covalently incorporated into the acrylamide matrix, and any pH range and curve shape can be generated by pouring a gradient gel using two solutions that differ in ampholyte composition rather than acrylamide concentration. As with tube gels, the initial electrophoresis is followed by a second separation using SDS-PAGE in a perpendicular direction (see Basic Protocol 4). The use of IPG gels has recently increased, for at least three major reasons: many of the technical problems associated with their use have been solved or substantially minimized, reproducible premade IPG gels are now commercially available, and lately strong interest has arisen in using two-dimensional gels for proteome analysis studies (analyzing and comparing the complete protein profiles of cell lines, tissue samples, or single-celled organisms). Another common two-dimensional electrophoresis format is a nonreducing/reducing electrophoretic separation (see Alternate Protocol 3), which provides useful information about intersubunit disulfides or protein-protein complexes that have been cross-linked using a bifunctional chemical cross-linker containing a disulfide bond within the linker region. This unit also includes support protocols describing pI standards and pH profile measurements (see Support Protocol 1), casting Immobiline gels (see Support Protocol 3), preparation of tissue culture cells and solid tissues for isoelectricfocusing (see Support Protocols 4 and 5), preparation of molecular weight standards for two-dimensional gels (see Support Protocol 6), and two-dimensional protein databases (see Support Protocol 7). NOTE: High-purity water (e.g., Milli-Q water or equivalent) is essential for all solutions. For cautions relating to electricity and electrophoresis, see Safety Considerations in the introduction to UNIT 10.1. Electrophoresis Contributed by Sandra Harper, Jacek Mozdzanowski, and David Speicher Current Protocols in Protein Science (1998) 10.4.1-10.4.36 Copyright © 1998 by John Wiley & Sons, Inc.
10.4.1 Supplement 11
BASIC PROTOCOL 1
HIGH-RESOLUTION EQUILIBRIUM ISOELECTROFOCUSING IN TUBE GELS This protocol describes the preparation of broad-range first-dimension gels using soluble ampholytes that resolve proteins with pI values between approximately 3.0 and 8.0, and is based on the original procedure described by O’Farrell (1975). The procedure presented here refers specifically to 3-mm IEF tube gels (first-dimension) combined with 1.5-mmthick 16 × 16–cm (size of separating gel) second-dimension gels (see Basic Protocol 3) and may be easily adapted to a variety of different gel sizes (see Table 10.4.1). A 3-mm IEF gel has a total protein capacity of ∼500 µg for complex protein mixtures such as whole-cell extracts. The maximum capacity of any single protein spot is ∼0.5 to 5 µg, depending on the solubility of the protein near its isoelectric point and the separation distance from any near neighbors. In this protocol, gels are cast and prefocused before the sample is loaded. The proteins are then separated according to isoelectric point, and the gels are extruded from the tubes and stored. Measuring pH profiles in IEF gels is a convenient and accurate method for determining pI (see Support Protocol 1). To provide optimal reproducibility, multiple gels should be cast and run simultaneously. This is especially important for comparative studies involving complex mixtures of proteins. The IEF gels may be cast either by pouring the gel solution into the gel tubes (steps 3a to 7a) or by using hydrostatic pressure (steps 3b to 7b). Pouring the gel solution into the gel tubes is convenient for 3-mm-diameter IEF gels and requires only a minimal excess of reagents. Because the gels are cast using a long needle and syringe, for narrower gels, where the needle does not fit inside the gel tube, casting using hydrostatic pressure is more appropriate. This method requires a larger excess of reagents and special casting cylinders. Many types of ampholytes are readily available from different suppliers to form the desired pH profiles. As ampholytes may vary significantly in their performance, careful selection of the appropriate ampholytes is usually necessary (see Commentary). Materials Chromic acid, in acid-resistant container Urea (ultrapure) 30% acrylamide/0.8% bisacrylamide (see recipe) 20% (w/v) Triton X-100 (see recipe) Ampholytes (e.g., pH 3-10/2D; ESA) TEMED (N,N,N′,N′-tetramethylethylenediamine) 2.5% (w/v) ammonium persulfate (see recipe; prepare immediately before use) 8 M urea (see recipe; prepare immediately before use) 0.1 M orthophosphoric acid (H3PO4; see recipe) 0.1 M NaOH (APPENDIX 2E; make fresh daily) Lysis buffer (see recipe) Protein samples to be analyzed Equilibration buffer (see recipe) 2-Mercaptoethanol
Two-Dimensional Gel Electrophoresis
Isoelectrofocusing apparatus (e.g., Protean II xi 2D from Bio-Rad or equivalent) with glass tubes, casting stand, buffer chambers, rubber grommets, and plugs 37°C water bath 110°C oven 10-ml syringe equipped with filter capsule (0.22 or 0.45 µm, e.g., Costar µStar LB) 10-ml syringe equipped with blunt needle [e.g., 20-G × 6 in. (15 cm) or 18-G × 6 in. (15 cm)]
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Current Protocols in Protein Science
Large glass cylinder sealed at bottom with Parafilm (optional, for hydrostatic pressure casting method only) 2000-V power supply 60-ml syringe Metal or plastic scoop Dry ice pellets Wash tubes and prepare the gel mixture 1. Remove the glass tubes from a chromic acid–filled container. Extensively wash the tubes with water, using high-purity water for the last wash. Dry the tubes at least 1 hr in an oven at 110°C and store them at room temperature, covered with aluminum foil. To prevent gels from sticking to the glass tubes, gel tubes have to be very clean. Satisfactory results are obtained by storing the tubes in chromic acid between uses and washing them shortly before use. Because drying the tubes requires at least 1 hr, cleaning steps should be performed the day before gels will be cast. CAUTION: Chromic acid is highly corrosive; follow supplier’s precautions carefully.
2. Prepare the gel solution by mixing: 16.9 g urea 4.0 ml of 30% acrylamide/0.8% bisacrylamide 3.0 ml of 20% (w/v) Triton X-100 7.5 ml water 3.0 ml ampholytes. Briefly warm the mixture in a 37°C water bath to solubilize urea if needed. To minimize decomposition of urea, never warm any solutions containing urea above 37°C, use ultrapure urea, and prepare solutions immediately before use. Choice of ampholyte composition is one of the key factors determining the quality of isoelectrofocusing separations. Substantial differences in performance, resolution, and shape of the pH gradient formed may be observed with different combinations of ampholytes and with ampholytes from different suppliers. ESA’s ampholytes (pH 3-10/2D) are suited for most applications and give reproducible results. Although purity of all reagents is important, the purity of urea and choice of ampholytes are among the most critical factors for the quality and performance of isoelectrofocusing. Most commercially available reagents marketed specifically for two-dimensional gel electrophoresis should be suitable, although individual lots of reagents from any supplier may provide variability and/or unacceptable results.
Cast gels by pouring 3a. Wrap one end of each glass tube with Parafilm and mount the tube in a casting stand. Mark all the tubes to indicate the desired gel height. For reproducible results, all gels should be the same height.
4a. Filter the gel solution using a 10-ml syringe equipped with a syringe-tip filter capsule. Briefly degas the gel solution (∼5 min) either by sonication or under vacuum. Then add 42.5 µl TEMED and 187.5 µl of 2.5% (w/v) ammonium persulfate solution to the filtered gel mixture and swirl gently to mix. 5a. Using a 10-ml syringe with a blunt needle, fill each glass tube with gel solution to the desired height. Make sure there are no air bubbles trapped in the gel. A needle is the best choice for casting gels if tubes of 3-mm inner diameter are used. For narrower tubes, the use of hydrostatic pressure is more appropriate (see steps 3b to 7b,
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below). For long gels the needle can be extended by inserting a piece of capillary polyethylene tubing over the needle tip. The amount of gel solution described in step 2 is sufficient for sixteen 3-mm tube gels that are 16 cm long.
6a. Immediately overlay each gel with ∼50 µl of 8 M urea. A pipettor with a capillary pipet tip is a convenient tool for overlaying with urea. Avoid mixing the overlay and gel solutions. Polymerization starts to occur ∼15 min after the addition of TEMED and ammonium persulfate. It is essential that the gels be poured and overlaid before significant polymerization has occurred.
7a. Let the gels polymerize at least 3 hr prior to use. Urea decomposes at a substantial rate at room temperature; therefore, the gels should be used the same day they are cast.
Cast gels using hydrostatic pressure 3b. Place a rubber band around the gel tubes so they form a tight bundle. Place the bundle inside a larger glass cylinder that is sealed at the bottom with several layers of Parafilm. All tubes must be precisely vertical. The dimensions of the larger cylinder depend on the dimensions and number of gel tubes. Excessive space will require more gel solution to cast the gels.
4b. Filter the gel solution using a 10-ml syringe and filter capsule. Degas the gel solution briefly (∼5 min) either with sonication or under vacuum. Add 42.5 µl TEMED and 187.5 µl of 2.5% ammonium persulfate solution and swirl. 5b. Pipet the gel solution into the bottom of the glass cylinder. Gently run water down the outside of the tube bundle using a wash bottle. Keep adding water until the gel mix reaches the desired height. Hydrostatic pressure will force the gel solution into the tubes. Sufficient gel solution must be used to obtain the desired gel height while avoiding forcing any water into the tubes. The volume of gel solution required can be estimated as follows: number of gels × 3.14 × (tube internal radius in cm)2 × height in cm + ∼10 ml to keep a safe level of gel mix at the bottom of the casting cylinder. As water is less dense than the gel solution, the water level will be slightly higher than the level of gel solution inside the tubes.
6b. Overlay the gels with 8 M urea. Urea decomposes at a substantial rate at room temperature; therefore, the gels should be used the same day they are cast.
7b. Let the gels polymerize at least 3 hr prior to use. Mount the gels in the electrophoresis unit 8. Prepare the lower electrode solution by degassing the proper amount of 0.1 M H3PO4 under vacuum with stirring for at least 5 min. Fill the bottom electrophoresis chamber. The amount of phosphoric acid depends on the length of the gel tubes and the type of electrophoresis unit. The solution should cover the entire gel for good heat dissipation. Approximately 3 liters are required for Protean II xi 2D electrophoresis units.
9. Remove the gel tubes from the casting stand, remove the Parafilm from the tube bottoms, and inspect gels for irregularities or trapped air bubbles. Discard imperfect gels. If using gels cast with hydrostatic pressure, remove the bundle of tubes en bloc, cut off excess acrylamide with a razor blade, and then rinse away remaining acrylamide particles from the outside of each tube. Two-Dimensional Gel Electrophoresis
10. Place a rubber grommet on the top of the tube. Approximately 5 mm of the tube should be visible above the upper edge of the grommet.
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11. Mount the tube with the grommet in the upper reservoir and plug any unused holes. After the tube is seated, its lower end must be submerged in the lower electrode solution. Be sure to remove any air bubbles trapped at the bottom of the tube by shaking or tapping the tube gently. Alternatively, with some units bubbles can be dislodged by raising and lowering the tubes or by using a long curved needle and syringe.
Prefocus the gels 12. Prepare the 0.1 M NaOH upper electrode solution by degassing under vacuum with stirring for at least 5 min. The amount of upper electrode solution necessary depends on the type of electrophoresis chamber. If a Bio-Rad Protean II xi 2D apparatus is used, 1 liter of 0.1 M NaOH is sufficient for both prefocusing and the separation.
13. Remove the 8 M urea overlay from the top of the gels using a Pasteur pipet and place ∼50 µl lysis buffer on the top of each gel. 14. Overlay lysis buffer with the degassed 0.1 M NaOH to fill the gel tubes. Avoid mixing of NaOH with the lysis buffer. 15. Pour the degassed 0.1 M NaOH into the upper chamber, making sure that all the gel tubes are covered with the electrode solution. Check carefully for leaks and air bubbles, then place lid on apparatus. 16. Connect the electrodes to a power supply by the red (+) lead to the lower chamber and the black (−) lead to the upper chamber. The voltages and currents used during electrophoresis are dangerous and potentially lethal. Safety considerations are given in the Electricity and Electrophoresis section of UNIT 10.1.
17. Prefocus for 30 min using 500 V constant voltage. Load the samples 18. Turn off power supply (see Safety Considerations in UNIT 10.1), disconnect leads, and remove lid. Using a 60-ml syringe, remove the electrode solution (0.1 M NaOH) from the upper chamber. 19. Remove the electrode solution and the overlay solution from each tube. Be careful not to damage the gel surface. 20. Place ∼50 µl lysis buffer on the top of each gel. Wait at least 2 min. 21. Remove the lysis buffer from the tubes. Rinsing the gels with lysis buffer removes any residual NaOH and protects the samples against exposure to high pH.
22. Load protein samples to be analyzed and carefully overlay each sample with ∼50 µl lysis buffer diluted with water 8:2 (v/v). Avoid mixing the buffer with the sample. The overlay solution protects samples from direct contact with the strong base used as an upper electrode solution. Dilution of the lysis buffer with water is necessary to decrease the density so the overlay does not mix with the sample. A 3-mm-i.d. × 16-cm-long IEF gel has a total protein capacity of ∼500 ìg for whole-cell extracts and other complex protein mixtures. The maximum capacity for any single protein spot is ∼0.5 to 5 ìg, depending on its solubility near its isoelectric point and the separation distance from any near neighbors. Preparation of relatively pure protein samples for isoelectrofocusing is generally straightforward. The sample usually may be prepared in one of the following ways: dialyze into any compatible low-ionic-strength buffer, lyophilize in a volatile or compatible low-ionic-strength buffer and dissolve in lysis buffer, or
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precipitate the protein using trichloroacetic acid (TCA) and redissolve in lysis buffer. For preparing extracts from cultured cells and from tissue samples, see Support Protocol 4 and Support Protocol 5, respectively. The minimum sample concentration of protein or radioactivity has to be sufficient for the desired detection method. For complex protein mixtures such as tissue or cell extracts, a 500-ìg total load is recommended for Coomassie blue staining (UNIT 10.5) or electroblotting (UNIT 10.7) for subsequent structural analysis, a 50-ìg total protein load should be sufficient for silver staining (UNIT 10.5) or immunoblotting, and no less than 100,000 counts/gel is recommended for proteins labeled with 3H, 14C, or 35S for autoradiography purposes. Sample volumes should be <150 ìl for 3-mm gels and <40 ìl for 1.5-mm gels. This implies at least a 5 ìg/ìl protein concentration in the sample for gels to be stained with Coomassie blue.
23. Carefully fill all tubes with 0.1 M NaOH. Avoid mixing the NaOH solution with the overlay solution and the sample. 24. Fill the upper reservoir with 0.1 M NaOH. Be sure that all gel tubes are covered with the solution. Run the gels 25. Connect the electrodes to a power supply with red (+) to the lower chamber and black (−) to the upper chamber. 26. Focus for a total of 12,000 Vhr. Unlike other electrophoretic techniques, in IEF the volt-hour is the most common unit describing the “time” of isoelectrofocusing. The initial voltage is usually set according to the desired number of volt-hours in a way that is convenient for the operator (i.e., so that the separation will run overnight), but it should not be <400 V. The upper voltage limit is restricted by heat released in the gels during isoelectrofocusing. At constant voltage the current will be the highest during the first hour of separation. The initial current will be strongly influenced by the ionic strength of the samples loaded onto the gels. An initial voltage of <800 V is recommended for 3-mm gels loaded with samples containing less than 100 mM salts/buffers; the voltage could be increased to 1200 V after ∼1 hr, if cooling is used. The current is a derivative of voltage and is never preset for isoelectrofocusing purposes. Some power supplies allow preprogramming the desired number of volt-hours and continuously adjust voltage and current during the isoelectrofocusing procedure (constant power). The total number of volt-hours is a major factor that affects separation in the first dimension. Optimal focusing time will vary for different ampholyte combinations, but 12,000 Vhr is a reasonable value for most systems. To achieve a total of 12,000 Vhr set the power supply to 667 V for 18 hr. These conditions are convenient for an overnight separation and do not require use of a cooling unit. Higher voltages can be used but may cause overheating of gels unless a highly efficient cooling system is employed. The maximum practical voltage decreases with increased gel tube inner diameter. Focusing for too long may cause cathodic drift and result in a shifted pH profile in the gel, whereas focusing for a short time will decrease resolution.
Extrude and store gels 27. Turn off power supply and carefully disconnect leads. Detach the lid and remove the NaOH solution from the upper reservoir of the electrophoresis chamber using a 60-ml disposable plastic syringe. 28. Remove one gel tube at a time from the chamber. 29. Using a 10-ml syringe equipped with a blunt needle, slowly and carefully inject water between the gel and glass tube. Start from the bottom of the tube, then repeat the procedure from the top. The gel should slide out of the glass tube. Two-Dimensional Gel Electrophoresis
It is convenient to let the gel slide from the glass tube onto a metal or plastic scoop, which facilitates transfer of the gel into a storage vial. It is relatively easy to break the gel during
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extrusion, and practicing on several unused gels is recommended. To extrude smaller-diameter gels, use water pressure generated by a syringe connected to the gel tube with Tygon tubing. If clean, unscratched glass tubes are used, extrusion should be easy.
30. Using the scoop, slide the gel into a 4.5-ml cryovial containing 3 ml equilibration buffer and 50 µl 2-mercaptoethanol. Close the vial, incubate exactly 5 min at room temperature, then freeze by placing the tube horizontally on top of dry ice pellets. Do not move or agitate the tube while the sample is freezing. The IEF gels may be run on a second-dimension gel immediately (see Basic Protocol 3), or can be stored at −80°C for many weeks. Even when the second-dimension is to be run immediately, extruded gels should be frozen after a carefully controlled incubation time at room temperature, such as the 5 min cited above for 3-mm-i.d. gels, to minimize diffusion of proteins out of the IEF gel. This short incubation before freezing will allow glycerol to diffuse into the gel. Too short an incubation or agitation during freezing can result in gel breakage. The total incubation time in equilibration buffer (sum of the time prior to freezing and after thawing) is critical and should be carefully controlled. Insufficient incubation time in equilibration buffer will not allow sufficient time for SDS to diffuse into the gel and saturate sites on the proteins. Excessive incubation times can result in appreciable protein losses due to diffusion out of the highly porous IEF gel.
CONDUCTING pH PROFILE MEASUREMENTS Standards with different isoelectric points can help in evaluating the performance of a specific system and determining the effective pH range in the isoelectrofocusing gel. Many pI standards are commercially available from different suppliers. It is most useful to separate a mixture of standard proteins that is prepared from several individual proteins or purchased as a preformulated kit. This mixture should be run in parallel with experimental samples on a separate reference gel. It is generally not recommended to run pI standards together in the same gel with samples because of possible interference with migration and identification of proteins of interest. Instead of analyzing standard proteins, a more precise evaluation of the pH profile can be made by directly measuring the pH throughout the gel using either a surface pH electrode or the following procedure.
SUPPORT PROTOCOL 1
1. Prepare and focus one or two gels (see Basic Protocol 1, steps 1 to 26) without any sample in parallel with experimental samples. 2. Prepare 20 to 40 glass test tubes each containing 1 ml high-purity, degassed water for each gel that will be used to measure the pH gradient (measurements on duplicate gels are recommended). The number of tubes required per gel equals twice the gel length (in cm).
3. After electrofocusing is completed, extrude the blank gels (see Basic Protocol 1, steps 27 to 29). Briefly rinse the gels with water. After extrusion, gel surfaces may be contaminated with electrode solutions. Rinsing with water is essential for obtaining reliable pH profiles.
4. Place the gel on a glass plate with a plastic ruler below the plate. Cut the gel into 0.5-cm pieces using a sharp razor blade. 5. Place each gel piece in a test tube containing 1 ml water. Do not mix the order of samples because each gel piece represents a single pH profile data point.
6. Place all test tubes on a shaker and shake gently for 1 hr at room temperature. 7. Read the pH of each solution and plot the pH profile as a function of the distance from the top of the gel.
Electrophoresis
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ALTERNATE PROTOCOL 1
NONEQUILIBRIUM ISOELECTROFOCUSING OF VERY ACIDIC PROTEINS Basic Protocol 1 is sufficient for separating proteins with isoelectric points greater than ∼3.0 to 3.5. For very acidic proteins, however, a nonequilibrium system is needed. The major features of this method are utilization of a shorter focusing time (without reaching equilibrium), a modified ampholyte mixture, and different electrode solutions. Additional Materials (also see Basic Protocol 1) 10% (w/v) ammonium persulfate (prepare immediately before use) Concentrated sulfuric acid (used in lower chamber electrode solution) Ampholytes, pH 2-11 (used in upper chamber electrode solution) To analyze very acidic proteins, follow Basic Protocol 1 with these exceptions in the indicated steps: 2. When preparing the gel solution, use the following mixture of ampholytes: 2.4 ml ampholytes pH 2.5-4 and 0.6 ml ampholytes pH 2-11. 4. Following the procedure for casting gels by pouring, add 100 µl of 10% ammonium persulfate solution, swirl, add 42.5 µl TEMED, and swirl again. Gel mixtures containing entirely or predominantly very acidic or very basic ampholytes are generally difficult to polymerize. Use of an increased ammonium persulfate concentration and adherence to the proper order of adding the reagents should ensure polymerization.
8. Prepare the bottom chamber electrode solution by adding 4.5 ml concentrated sulfuric acid to 3 liters water. Degas at least 5 min. Omit steps 12 to 19 (do not prefocus the gels). 20. Remove the 8 M urea (polymerization overlay solution) and place ∼50 µl lysis buffer on top of each gel. Wait at least 2 min, then remove the lysis buffer. 23. Carefully fill all tubes with the upper chamber electrode (anode) solution prepared by mixing pH 2-11 ampholytes with water in a 1:40 ratio. 24. Fill the upper buffer chamber (anode) with the solution described in step 23. Iminodiacetic acid (10 mM) may be a more economical alternative anode solution.
26. Focus for a total of 4000 Vhr. ALTERNATE PROTOCOL 2
NONEQUILIBRIUM ISOELECTROFOCUSING OF BASIC PROTEINS In general, most equilibrium IEF gel systems using soluble ampholytes produce pH gradients that do not exceed pH 8.0 on the basic end, yet many proteins have higher pI values. For this reason samples containing very basic proteins are usually focused using a nonequilibrium system. In an equilibrium system, proteins are loaded on the basic end of the gel and migrate toward the acidic end until they reach a pH equal to their pI. In nonequilibrium systems, the sample is loaded on the acidic end of the gel, and focusing is terminated after a relatively short time (fewer volt-hours). To run nonequilibrium IEF gels, follow the procedure previously described (see Basic Protocol 1) with these alterations in the indicated steps: 8. Use 0.1 M NaOH as the lower electrode solution.
Two-Dimensional Gel Electrophoresis
Electrode solutions and electrodes are reversed in this procedure relative to equilibrium isoelectrofocusing.
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Omit steps 12 to 19 (do not prefocus the gels). 20. Remove the 8 M urea (polymerization overlay solution) and place 50 µl lysis buffer on top of each gel. Wait at least 2 min, then remove the lysis buffer. 23. After loading the samples and overlaying with lysis buffer diluted with water 8:2 (v/v) as in Basic Protocol 1, use 0.1 M H3PO4 instead of NaOH to fill all gel tubes. 24. Use 0.1 M H3PO4 as the upper electrode solution. 25. Reverse the connection of electrodes—i.e., connect the red (+) lead to the upper chamber and the black (−) lead to the lower chamber. 26. Focus for a total of 3000 to 5000 Vhr. The optimal number of volt-hours depends on the nature of the sample and the ampholytes used. The values recommended above may need to be adjusted empirically.
ISOELECTROFOCUSING USING IMMOBILIZED pH GRADIENT GEL STRIPS
BASIC PROTOCOL 2
In immobilized pH gradient (IPG) gels, the ampholytes are covalently linked to the acrylamide matrix, which facilitates production of highly reproducible gradients as well as very narrow pH gradients for optimal resolution of minor charge differences. A variety of precast gels and all the necessary equipment are commercially available from Hoefer Pharmacia. Equipment and chemicals are also available for the user to cast gels in the laboratory (see Support Protocol 3), although precast gels are likely to suffice for the majority of applications. Narrow strips of precast IEF gels (Immobiline DryStrips) may be used to achieve a first-dimension separation for two-dimensional gel electrophoresis, and broader precast slab gels (Immobiline DryPlates) can be used to compare multiple samples after IEF separation only (see Support Protocol 2 and Table 10.4.3). In this protocol, precast Immobiline DryStrips from Hoefer Pharmacia are rehydrated overnight using the reswelling cassette (one to twelve sample strips may be handled at a time); samples are applied using sample cup holders and gel strips are isoelectrofocused overnight. This procedure has been adapted from instruction booklets provided by Hoefer Pharmacia with Immobiline Dry Strip Kits and with the Immobiline DryStrip reswelling tray. Since there is a greater selection of pH ranges for premade Immobiline DryPlates than DryStrips, it is sometimes convenient to cut DryPlates into strips prior to rehydrating the gel to obtain narrower pH ranges where needed. See Basic Protocol 4 for details concerning preparing and running the second-dimension gel. Wear gloves throughout the procedure and handle the Immobiline DryStrips with forceps where feasible to prevent extraneous protein contamination of the gels and gel solutions. Thoroughly clean all equipment with a mild laboratory detergent solution, rinse well with Milli-Q water, and allow to dry before using. Solutions containing 10 M urea may be heated briefly to 30° to 40°C to aid in solubilization. Materials Urea (ultrapure) CHAPS or Triton X-100 Pharmalyte 3-10, 4-6.5, and/or 8-10.5 soluble ampholytes (see Table 10.4.1; Hoefer Pharmacia) Ampholine pH 6-8 (Hoefer Pharmacia) DTT (dithiothreitol) Bromphenol blue Precast Immobiline DryStrips (Hoefer Pharmacia)
Electrophoresis
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Table 10.4.1
Rehydration Solutions for Immobiline DryStripsa
Component Ultrapure urea CHAPSc Pharmalyte pH 3-10 Pharmalyte pH 4-6.5 Pharmalyte pH 8-10.5 Ampholine pH 6-8 DTT Bromphenol blue Milli-Q water
DryStrip type
Final conc. 9 Mb 2% 1:50
0.3% Trace
3-10L
3-10NL
4-7L
2.7 g 0.1 g 100 µl
2.7 g 0.1 g
2.7 g 0.1 g
50 µl 25 µl 25 µl 75 mg A few grains To 5 ml
100 µl
75 mg A few grains To 5 ml
75 mg A few grains To 5 ml
aRehydration
solutions should be prepared fresh immediately before use and should be filtered using a 0.2-µm filter. Minimize total time the solution is at room temperature prior to use to minimize decomposition of urea. If the reswelling tray is used, ~250 or 400 µl rehydration solution is required per 11- or 18-cm DryStrip, respectively. bUrea concentrations between 8 M and 10 M are typically used. Higher concentrations promote better protein solubility during isofocusing while increasing the risk of urea crystallization. If 10 M urea is used, extra care should be taken to minimize evaporation and the temperature should be maintained at 20°C or slightly higher at all times to avoid crystallization of the urea. cThe optimal detergent and detergent concentration should be empirically determined. Other common alternatives are Triton X-100 and octyl-glucoside. The detergent used must be nonionic or zwitterionic to avoid high current and consequent overheating during isoelectrofocusing.
DryStrip cover fluid (Hoefer Pharmacia) Immobiline DryStrip kit (Hoefer Pharmacia) including: Cathode electrode Anode electrode Sample cup bar Tray Sample cups Immobiline strip aligner IEF electrode strips Sample application pieces Instruction manual Protein sample to be analyzed Lysis buffer (see recipe) Immobiline DryStrip reswelling tray (Hoefer Pharmacia) Forceps Filter paper Glass plate Flatbed electrophoresis unit (Hoefer Pharmacia Multiphor II or equivalent) Recirculating cooling water bath Power supply (minimum capacity of 3000 to 3500 V) Petri dishes Additional reagents and equipment for protein detection by staining (UNIT 10.5) and/or for electroblotting (UNIT 10.7, optional) Two-Dimensional Gel Electrophoresis
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Rehydrate the Immobiline DryStrip(s) 1. Prepare an appropriate rehydration solution for the type of DryStrips to be used as described in Table 10.4.1 (∼400 µl rehydration solution per 18-cm DryStrip). The urea concentration in the rehydration solution should be 8 to 10 M, and 2% CHAPS or another appropriate detergent (zwitterionic or nonionic) such as Triton X-100, NP-40, or n-octylglucoside should be included in the rehydration solution to aid in sample solubility. The optimal detergent and detergent concentration may vary with type of sample and should be determined empirically. One possible method of loading large sample volumes onto IPG gels is to add the sample directly to the rehydration solution. However, this sample loading method is not typically recommended since it tends to result in poor and variable protein loading into the gel. During rehydration of the gel, proteins are only poorly adsorbed into the gel matrix, apparently as a result of charge repulsion effects caused by the highly charged gel matrix. Solutions containing urea should be filtered using a 0.2-ìm filter before use.
2. Slide the protective lid off the reswelling tray and level the tray by adjusting the leveling feet until the leveling bubble is centered. 3. For an 18-cm gel, pipet 350 to 400 µl of rehydration solution into a slot of the reswelling tray. Move the pipet along the length of the well while adding the solution to spread it evenly throughout the length of the slot. Avoid excessive air bubble formation while pipetting this solution. 4. Remove the protective cover from the Immobiline DryStrips and gently place them, gel side down, into the prepared slot. To facilitate their removal after rehydration, the strips should be oriented with their pointed ends at the sloped end of the slots in the rehydration tray. Be careful not to trap any air bubbles under the gel strips.
5. Overlay each strip with 2 to 3 ml of DryStrip cover fluid to prevent evaporation and urea crystallization. Slide the protective lid into place and allow gels to rehydrate overnight (∼16 hours) at room temperature. Shorter rehydration times may not completely and reproducibly rehydrate the gels. Do not substantially exceed 16 hr as extensive incubation, especially rehydrating gels over a weekend, increases potential problems due to evaporation and subsequent urea crystallization. In addition, long incubation times increase the extent of urea decomposition, which will increase the risk of amino group modification on proteins by the cyanate produced from urea decomposition.
6. After the overnight rehydration, slide the lid off the reswelling tray. Place a forceps tip into the slight depression under each strip and remove the strip. Gently blot any excess oil or moisture from the plastic backing of the rehydrated strips with filter paper. A damp piece of filter paper may also be used to blot the surface of the gel. Some of the paper may adhere to the gel and should be gently peeled away. The gel is now ready to be placed in the strip aligner on the cooling plate of the electrophoresis unit. Do not allow the gel to dehydrate prior to placing it on the cooling plate in step 11. (Steps 7 to 10 should be completed prior to removing the strips from the reswelling tray.)
Run the first dimension 7. Level the Multiphor II electrophoresis unit, then connect it to a circulating cooling water bath. Allow it to cool to 15°C for 1 to 2 hr to ensure even cooling. Do not cool below 15°C to prevent precipitation of urea in the gels. Electrophoresis
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8. Pipet ∼5 ml DryStrip cover fluid onto the surface of the Multiphor II cooling plate. Position the Immobiline DryStrip tray on the cooling plate oriented with the red (+, anodic) electrode at the top, near the cooling tubes. Avoid large air bubbles between the cooling plate and the tray (small bubbles should not cause a problem).
9. Connect the red and black electrode leads on the tray to their respective positions on the Multiphor II unit. Pour 10 ml of DryStrip cover fluid into the tray. Place the Immobiline DryStrip aligner on top of the oil, groove side up. Avoid getting oil on top of the strip aligner. The possible presence of small air bubbles under the strip aligner is not important.
10. Cut two electrode strips to a length of 11 cm (regardless of the number of DryStrips used). Place the electrode strips onto a clean glass plate and soak each one with 0.5 ml Milli-Q water. Blot with a Kimwipe or tissue paper to remove excess water. The electrode strips should be evenly soaked and just damp after blotting. Excessive water could cause sample streaking.
11. Transfer the strips from step 6 to adjacent grooves in the aligner tray. Position the rounded (acidic) end of each strip near the top of the tray at the red electrode (anode) near the cooling tubes, and the square end at the bottom of the tray near the black electrode (cathode). Be sure that the edges of all gel strips at the anode end are lined up evenly. 12. Place the blotted electrode strips from step 9 on top of the gel surface of the DryStrips near the anode and cathode ends of the gel. Position the red (anode) and black (cathode) electrodes on top of the electrode strips at their respective ends. After the electrodes have been pressed down on top of the electrode strips, check that the gel strips have not shifted position.
13. Push the sample cups onto the sample cup bar. Place the sample cup bar near the anode end of the gel so that the small spacer arm just touches the electrode and the sample cups are nearest to the electrode, but do not allow the cups to touch the gel. The sample cups should face the nearest electrode. The acidic end of the gel can usually be used for sample application; however, the optimal loading position may need to be determined empirically for different types of samples. At high protein concentrations and/or at non-optimal pH, samples may precipitate in the gel at the loading position.
14. Position one sample cup above each gel strip and push down to ensure good contact between the bottom of the sample cup and the gel strip. Make sure the gel strips have not shifted position. 15. Pour 70 to 80 ml of DryStrip cover fluid into the tray (it will cover the gels). If oil leaks into the sample cups, adjust the cups to stop leakage. When there is no leakage into the sample cups, add enough cover fluid to the tray to completely cover the sample cups (~150 ml). 16. Pipet protein samples (in lysis buffer) into the sample cups by underlaying. The sample should sink to the bottom of the cup. Check for leakage of the sample out of the sample cup.
Two-Dimensional Gel Electrophoresis
Samples should either be lyophilized and then solubilized in lysis buffer, or diluted 9 parts lysis buffer to 1 part sample. The maximum volume each sample cup holds is 100 ìl. The complexity of the sample, the sample solubility at the loading concentration and pH used, the thickness of the second-dimension gel, and the detection method to be employed should be considered when deciding how much protein to load. As a starting reference, typical
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loading ranges for 1.0- to 1.5-mm-thick 18-cm × 18-cm gels would be ∼5 to 20 ng per major spot for silver staining and ∼1 to 5 ìg per major spot for Coomassie blue staining. When very complex samples are used such as whole cell extracts, total protein loads are likely to be ∼20 to 100 ìg for silver staining and ∼200 to 1000 ìg for Coomassie blue staining. The salt concentration in samples should be kept <50 mM and, if the sample contains SDS, the final SDS concentration should be <0.25%.
17. Place the lid on the Multiphor II unit and connect the leads to a power supply. Focus the gels with constant voltage for 2 to 3 hr at 500 V followed by 12 to 16 hr at 3500 V for a total of 40 to 60 kVhr. Refer to the user manual for exact recommended voltage conditions for each type of Immobiline DryStrip. The optimal number of Vhr will depend upon the pH range of the Immobiline Strip used, the type of sample, and the sample load and volume; therefore, the optimal Vhr should be empirically determined for different applications.
18. When isoelectrofocusing is complete, disconnect the power supply and remove the cover from the Multiphor II unit. Remove the electrodes, electrode strips, and sample cup bar from the tray. If gels are to be run in the second dimension immediately after isofocusing, steps 1 to 3 of Basic Protocol 4 should be completed prior to terminating isofocusing.
19. Using forceps, remove the DryStrips from the tray. If the gels are to be run in the second dimension immediately, place in a petri dish with the support film along the wall of the dish and proceed directly to equilibration of the gel (see Basic Protocol 4, step 4). Alternatively, gels may be stored sealed in a plastic bag at −80°C until ready to run the second-dimension gel. Gels may be stored at least 2 to 3 months at −80°C. Do not place in the equilibration buffers required for the second dimension prior to storage.
ELECTROPHORESIS ON IMMOBILIZED pH GRADIENT GELS In this protocol, after precast IEF gels (Immobiline DryPlates) from Hoefer Pharmacia are rehydrated, samples are loaded and subjected to isoelectrofocusing. Gels are typically run at 2500 to 3500 V and require focusing times of 2 to 7 hr. Protein samples may be detected by conventional methods such as Coomassie blue or silver staining (UNIT 10.5). Isoelectric points can be determined with the use of pI calibration proteins; alternatively, because the gradient is linear, one can measure the migration distance across the gel and estimate the pI at each location. As noted in Basic Protocol 2, Immobiline DryPlates can be cut into 3-mm-wide strips to use as the first dimension of two-dimensional gels, since DryPlates are available in narrower pH ranges than DryStrips. DryPlates can also be used as described in this protocol to simultaneously separate multiple samples in a single dimension. Applications of this method include initial screening of samples to determine the optimal pH gradient prior to running more time-consuming two-dimensional gels, prescreening fractions from a chromatographic purification step prior to running two-dimensional gels, and evaluation of charge heterogeneity of purified proteins. Additional Materials (also see Basic Protocol 2) Precast DryPlate gel (Hoefer Pharmacia) Repel-Silane (Hoefer Pharmacia) Paraffin oil Protein samples to be analyzed Reswelling Cassette kit (Hoefer Pharmacia) including: 125 × 260 × 3–mm glass plate with 0.5-mm U frame 125 × 260 × 3–mm glass plate continued
SUPPORT PROTOCOL 2
Electrophoresis
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Silicone tubing Pinchcock Clamps 20-ml syringe Roller (Hoefer Pharmacia) Whatman no. 1 filter paper Flatbed electrophoresis unit (Hoefer Pharmacia Multiphor II) 10° or 15°C cooling water bath Electrode strips Sample applicator strip or sample application pieces Power supply (minimum capacity 3000 to 3500 V) Additional reagents and equipment for protein detection by staining (UNIT 10.5) and for electroblotting (UNIT 10.7; optional) Rehydrate the gel 1. Remove precast gel from packaging. If the entire gel is not needed, cut off the required number of lanes and reseal the unused gel. Mark the polarity of the gel section to be used by cutting a small triangle off the anode corner. Handle the gel by the support film only. It is critical that the lanes are cut from the gel in the proper orientation to preserve the pH gradient (see Fig. 10.4.1), and polarity must be indicated for proper orientation of electrodes later in the procedure.
2. Use the Reswelling Cassette to rehydrate the gel. Connect silicone tubing through hole in the bottom corner of the U-frame plate, seal with silicone glue, and connect the pinchcock to the other end of the tubing. Place a glass plate on a clean flat surface and wet with a few drops of water. Place the gel on the plate, gel side up. Gently roll with a clean rubber roller to remove any air bubbles. 3. Cover the plate and gel with the plate fitted with the U frame. The U-frame plate should be coated with a thin layer of Repel-Silane to prevent the gel from sticking to the plate.
4. Place clamps around the edges of the plates, making sure the seal is tight.
anode side (+) cut corner (anode side)
cut in this direction
Two-Dimensional Gel Electrophoresis
anode corner cut by manufacturer
cathode side (–)
Figure 10.4.1 Marking orientation of a precast IPG gel when only a portion of the gel is used.
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5. Slowly fill the cassette with the desired rehydration solution using a 20-ml syringe connected to the silicone tubing and let stand for the recommended amount of time. Precool electrophoresis unit 1 to 2 hr prior to electrophoresis (see step 9). Reswelling with water for 2 to 3 hr is normally sufficient. If using additives such as urea, Triton, glycerol, or reducing agents, allow the gel to rehydrate overnight. Additives can be used to improve solubility of proteins near their isoelectric point. Reducing reagents such as 2-mercaptoethanol or DTT are used to reduce disulfide bonds.
6. When gel has been allowed to rehydrate completely, remove the clamps and gently pry the plates apart. 7. Moisten a piece of filter paper with water and place on top of the gel, then layer with a piece of dry filter paper. 8. Gently blot the gel by rolling over the dry filter paper with the rubber roller to remove excess water. The gel is now ready to be placed on the cooling plate. Do not let the gel dehydrate prior to placing it on the cooling plate in step 11.
Run the gel 9. Connect the flatbed electrophoresis unit to a recirculating cooling water bath. Allow to cool to 10°C for 1 to 2 hr to ensure even cooling. If the gel has been rehydrated in the presence of urea, do not cool below 15°C so that urea does not precipitate. 10. Pipet 2 to 3 ml paraffin oil onto the surface of the cooling plate. 11. Position the gel on the cooling plate, being careful not to trap air bubbles between the gel and the plate. Orient the gel so that the polarity of the gel matches the polarity of the cooling plate. 12. Soak two electrode strips with ∼3 ml water, then blot to remove excess water. 13. Lay a blotted electrode strip along each long edge of the gel. Cut off the ends of the electrode strip so that it does not extend beyond the edge of the gel. 14. Load protein samples to be analyzed onto the gel. Use an applicator strip for sample volumes between 5 and 20 µl (make sure contact between the strip and the gel is uniform). Use sample application pieces for sample volumes >20 µl. Remove the application pieces halfway through focusing. For sample volumes of 2 to 10 µl, samples may be spotted directly on the gel without using applicator strips. See manufacturer’s instructions for further details. An important experimental consideration is the position in the pH gradient where the sample is applied. The acidic end of the gel can usually be used for sample application; however, the optimal loading position may need to be determined empirically for different types of samples. At high protein concentrations and/or at nonoptimal pH’s, samples may precipitate in the gel at the loading position. Samples should contain <50 mM salt or buffer components; greater concentrations will cause local overheating of the gel. If possible, salt-free samples should be solubilized or dialyzed in the rehydration buffer.
15. Align the electrodes with the electrode strips, put the safety lid in place, and connect the apparatus to the power supply. Conduct electrophoresis at 3000 V. Broad-range IPG gels, such as Immobiline DryPlate, pH 3-10, should be run at ∼3000 V for 2 to 4 hr. Narrower-range gels are also run at 3000 V but may require a focusing time of 4 to 7 hr. Electrophoresis
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16. After removing gels from the electrophoresis apparatus, detect proteins using any conventional staining technique such as Coomassie blue or silver staining. 17. Preserve the gels by sealing in a plastic bag or by drying for a permanent record. Alternatively, electrotransfer the proteins on the gel to a membrane. To dry a gel, presoak it first in a preservation solution. For silver-stained gels, use a solution of 5% to 10% (w/v) glycerol/30% (v/v) ethanol; for Coomassie blue–stained gels, use a solution of 5% to 10% (w/v) glycerol/16% (v/v) ethanol/8% (w/v) acetic acid. After soaking the gel, place it on a glass plate gel side up, cover with a cellophane sheet soaked in preservation solution, and allow to dry at room temperature. For electrotransfer (UNIT 10.7), use film remover to remove the plastic support film from the gel. Electrotransfer of proteins to a polyvinylidene difluoride (PVDF) membrane using a Multiphor II NovaBlot transfer kit (Hoefer Pharmacia) is recommended. Transferring IPG gels requires special procedures; see the transfer kit manual for instructions. SUPPORT PROTOCOL 3
CASTING AN IMMOBILINE GEL An alternative to precast IPG gels is the use of Hoefer Pharmacia Immobilines to cast immobilized pH gradient gels with customized pH gradients and ranges, including very narrow pH ranges, to improve separation of proteins with small charge differences. This protocol describes the general procedure of casting custom-made Immobiline gels. The Reswelling Cassette used in Basic Protocol 2 for rehydrating gels is employed for casting the gels, which are polyacrylamide gels poured with a gradient of Immobilines following instructions provided by Hoefer Pharmacia application note 324. Additional Materials (also see Support Protocol 2) GelBond PAG film (Hoefer Pharmacia) Immobiline solutions (Hoefer Pharmacia) 2.5% (v/v) glycerol Gradient maker Orbital shaker Additional materials and equipment for rehydrating immobilized pH gradient gels (see Basic Protocol 2) Cast the gel 1. Coat the plate with the U frame with Repel-Silane to prevent the gel from sticking to the glass plate. 2. Place a glass plate on a clean, flat surface and wet with several drops of water. Cover with a sheet of GelBond PAG film, hydrophilic side up. Use the roller to remove any air bubbles trapped between the film and the glass plate. 3. Place the plate with the U frame on top of the GelBond PAG film. Clamp the plates together on three sides. 4. Mix the Immobiline solutions following the instructions provided by Hoefer Pharmacia application note 324 to prepare the desired pH range. Cast the pH gradient gel using a gradient maker. Once the catalysts have been added to the gel solution, it is important to work quickly to ensure that the gradient is poured before polymerization occurs. See UNIT 10.1 for instructions for using gradient makers.
Two-Dimensional Gel Electrophoresis
5. Do not disturb the gel during the first 10 min to allow the gradient to stabilize. Allow the gel to polymerize 1 hr in a 50°C oven.
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Dry and store the gel 6. Allow the gel to cool to room temperature, then disassemble the cassette. Cut off a small corner to label the anode end of the gel. 7. Wash the gel 2 to 3 hr with 200 to 300 ml water. Use an orbital shaker and change the water two or three times. At this stage the gel may be used immediately (if no additives are needed) or dried as described in steps 8 to 10.
8. Wash the gel 30 min to 1 hr with 200 to 300 ml of 2.5% (v/v) glycerol. 9. Place the gel on a glass plate (gel side up) in a dust-free environment and allow to dry at room temperature overnight. 10. Store the dried gel in a sealed plastic bag at −20°C for up to 2 months. The gels may be rehydrated when needed (see Support Protocol 2).
PREPARING TISSUE CULTURE CELL EXTRACTS FOR ISOELECTROFOCUSING
SUPPORT PROTOCOL 4
Preparation of samples for isoelectrofocusing containing relatively pure proteins is generally straightforward (see Basic Protocol 1, step 22). In contrast, complex samples such as whole-cell extracts, tissue extracts, or subcellular fractions are more difficult to prepare for successful isoelectrofocusing. Solubility limitations both prior to isoelectrofocusing and during focusing restrict analysis of these complex samples to protocols that include nonionic detergents and urea (see Basic Protocol 1). In addition, the presence of DNA and RNA in crude cell extracts further complicates isoelectrofocusing. The protocol presented below is suitable for preparing samples from cell cultures and is based on quantities compatible with silver staining or Coomassie blue staining. If smaller cell numbers and high-sensitivity detection methods such as autoradiography are used, volumes and quantities should be adjusted as needed. In this protocol the cells are harvested and washed in phosphate-buffered saline (PBS) with proteolysis inhibitors, then lysed in Tris/SDS buffer using sonication, after which the total protein concentration is determined in the lysate. The lysate is further treated with a mixture of DNase and RNase, and additional SDS and reducing agent are added. At this stage, the samples can be stored at −80°C for an extended time or, after addition of urea and lysis buffer, may be loaded directly onto prefocused IEF gels. Filtration of the final sample prior to loading onto the IEF gel is essential for quality of isoelectrofocusing. Materials Cell culture flasks containing cells of interest PBS with proteolysis inhibitors (PBS/I buffer; see recipe) Dry ice/ethanol (optional, for freezing samples) Tris/SDS buffer (see recipe) BCA protein assay kit (Pierce) DNase and RNase solution (see recipe) 20% (w/v) SDS (APPENDIX 2E) 2-Mercaptoethanol Urea (ultrapure) Lysis buffer (see recipe) 50-ml centrifuge tube Centrifuge with rotor (e.g., Beckman JS-4.2), 4°C
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1- to 2-ml cryovials Microcentrifuge, 4°C Sonicator with microtip 0.2-µm microcentrifuge filter units (e.g., Millipore Ultrafree-MC filter units) Harvest and wash the cells 1. Place cell culture flasks containing cells of interest on ice. 2. Rapidly wash cells three times with 2 to 6 ml PBS/I buffer. Keep the flasks on ice. The required volume of PBS/I buffer depends on the flask size. For example, use 2 ml for a 25-cm2 tissue culture flask and 6 ml for a 75-cm2 tissue culture flask.
3. Add 2 to 6 ml PBS/I buffer to the flask, scrape the cells using a cell scraper, and transfer the suspension to a 50-ml centrifuge tube. Repeat this step with another 2 to 6 ml buffer to ensure complete transfer of the cells. Cells grown in suspension are washed in an analogous manner using repetitive centrifugation.
4. Collect the cells by centrifuging 15 min at 2600 × g (3000 rpm in Beckman JS-4.2 rotor), 4°C. 5. Discard the supernatant and resuspend the cells in a small volume of PBS/I buffer. 6. Transfer the cell suspension to a labeled cryovial. The weight of the empty cryovial can be determined prior to use if the wet weight of the cell pellet is desired as a reference value rather than cell number, radioactivity, or another criterion.
7. Microcentrifuge the cells 15 min at maximum speed, 4°C. 8. Remove and discard the supernatant using a pipettor or Pasteur pipet. Weigh the vial containing the cell pellet. Record the wet weight of the cell pellet (in mg). 9. Freeze the cell pellet in a dry ice/ethanol mixture (optional). Frozen cells can be stored at −80°C for at least several months.
Prepare the cell pellets for isoelectrofocusing 10. Retrieve cell pellets from −80°C storage if samples were frozen. 11. Add 400 µl Tris/SDS buffer per 50 to 100 mg cell pellet wet weight. Keep the cells on ice at all times. The total amount of protein in the pellet is roughly 5% of the wet pellet weight.
12. Sonicate the sample three times for 3 sec using a sonicator with a microtip at medium power. Keep the samples on ice during sonication. Use pulse sonication or wait at least 5 min between sonications. Minimizing heat generation is essential because substantial proteolysis can occur if the sample warms appreciably. If additional sonication is necessary (i.e., if the sample is not homogeneous), let the sample cool down on ice before the next series of sonications.
13. Run a BCA protein assay to determine the protein concentration if sample will be loaded on that basis. For maximum accuracy use the same amount of Tris/SDS buffer in standards as in experimental samples. 14. Prepare labeled cryovials to store aliquots of the sample, if desired. Precool vials on ice prior to making aliquots. Two-Dimensional Gel Electrophoresis
The amount of protein per aliquot depends on the anticipated future uses of the sample, as repeated freezing and thawing should be avoided. About 500 ìg/gel whole-cell extract is
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a maximum load for preparative purposes using 3-mm gels (i.e., isolation of proteins for sequencing or other structural work). Approximately 50 ìg/gel is an appropriate load for silver staining. The final protein concentration after completion of the protocol (steps 15 to 20) will equal the concentration found by protein assay divided by 1.1, owing to the addition of reagents after the protein assay step.
15. Add 20 µl DNase and RNase solution per 400 µl Tris/SDS buffer used for sonication (step 11). Incubate 10 min on ice. 16. Add 20 µl of 20% SDS solution and 5 µl of 2-mercaptoethanol per 400 µl Tris/SDS buffer used in step 11. Incubate 5 min at 37°C. 17. Quickly divide samples into previously prepared cryovials and immediately freeze aliquots using a dry ice/ethanol bath. Store at −80°C. Samples stored at −80°C are stable ≥1 year. Work quickly to minimize potential proteolysis. This step may be omitted if the samples are to be loaded on IEF gels immediately. Generally, the total amount of sample greatly exceeds the amount required for an IEF gel, and freezing aliquots is beneficial. To avoid potential reproducibility problems, all samples should be processed identically.
Prepare the samples for isoelectrofocusing 18. If cell extracts were frozen, thaw samples and immediately add dry urea to 9 M final concentration. The amount of urea (in mg) equals 0.83 times the sample volume (in ìl). For example, use 83 mg urea per 100 ìl sample. The final volume of the sample (with urea added) equals 1.6 times the initial volume (160 ìl in the same example).
19. Add an equal volume of lysis buffer (160 µl in the above example) and warm briefly if necessary to dissolve urea. 20. Filter samples using a 0.2-µm microcentrifuge filter unit by microcentrifuging at maximum speed, room temperature, until the entire sample has passed through the filter. Load the desired volume onto the IEF gel. If a 500-ìg total protein load per 3-mm gel is desired (a practical maximum load for most whole-cell extracts), the protein concentration determined during the protein assay has to be ≥5 ìg/ìl. If the sample is less concentrated, the sample volume required will be too large for a 3-mm IEF gel. Alternatively, sample loads can be based on cell numbers, radioactivity, or any other appropriate reference (see Basic Protocol 1, step 22).
PREPARING PROTEINS IN TISSUE SAMPLES Tissue samples are usually solubilized in lysis buffer using homogenization. After centrifugation, the protein sample can be loaded onto the first-dimension gel. In general, much higher sample-to-sample variability is expected when tissue samples are analyzed.
SUPPORT PROTOCOL 5
Materials Tissue samples Lysis buffer (see recipe) Dounce homogenizer or equivalent Ultracentrifuge and rotor (e.g., Beckman Ti70), 2°C 1. Place tissue sample in a Dounce homogenizer, add ∼2 ml lysis buffer per 100 mg tissue, and homogenize the sample on ice (e.g., 3 to 5 strokes). 2. Let the mixture stand a few minutes, then transfer to an appropriately sized ultracentrifuge tube depending on total sample volume.
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3. Centrifuge 2 hr at 100,000 × g (e.g., 33,000 rpm in a Beckman Ti70 rotor for 100,000 × g), or 1 hr at 200,000 × g 2°C. 4. Divide the supernatant into aliquots and freeze at −80°C or immediately load an appropriate volume onto the IEF gel. BASIC PROTOCOL 3
SECOND-DIMENSION ELECTROPHORESIS OF IEF TUBE GELS Second-dimension gels are identical to those described in UNIT 10.1 except for sample loading, which requires a broad, flat well. A broad well can be cast using an appropriate two-dimensional comb if the second-dimension gel thickness is slightly larger than that of the first-dimension gel. Alternatively, when the second-dimension gel is being cast, water can be layered over the entire surface of the gel to produce a flat surface that will accommodate the first-dimension gel. Narrow analytical isoelectrofocusing gels (≤1.5 mm) that fit between the glass plates of the second-dimension gel do not generally require a stacking gel, although a stacking gel may improve resolution under some circumstances. Stacking gels are essential when first-dimension gels >1.5 mm are loaded on reduced-thickness second-dimension gels, for example, when 3-mm first-dimension gels are loaded on 1.5-mm second-dimension gels. To ensure the best reproducibility, casting multiple second-dimension gels in a multigel casting stand is strongly recommended. This is especially important when gradient gels are used for the second-dimension and/or critical comparisons of multiple samples are planned. This protocol describes all the specific steps required for successfully casting and running the second-dimension gel. The use of beveled plates and an agarose overlay is especially important when 3-mm IEF gels are loaded onto 1.5-mm second-dimension gels. Materials 2% (w/v) agarose (see recipe) Equilibration buffer (see recipe) Isoelectrofocusing gels containing protein samples to be analyzed (see Basic Protocol 1) Piece of agarose containing molecular weight standards (see Support Protocol 6) Beveled glass plates Boiling water bath Metal or plastic scoop Additional reagents and equipment for linear and gradient Laemmli gels (UNIT 10.1) Cast the second-dimension gels 1. Assemble the glass-plate sandwich of an electrophoresis apparatus, using a beveled plate for the shorter side of the gel sandwich. A beveled plate provides more space for a thicker IEF gel and will accommodate a first-dimension gel that is at least 1 to 2 mm larger than the thickness of the second-dimension gel.
Two-Dimensional Gel Electrophoresis
2. If the thickness of the first-dimension gel exceeds that of the second-dimension gel, pour a separating gel of the desired acrylamide concentration and immediately overlay with water to produce a smooth surface. The separating gel height should be a minimum of 2 cm below the top of the beveled plate to accommodate the stacking gel.
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A
B
agarose overlay IEF gel
water overlay electrophoresis central cooling core
stacking gel, unpolymerized
polymerized stacking gel outer plate
inner plate
outer plate
inner plate
Figure 10.4.2 Casting the second-dimension gel and loading the IEF gel. (A) The stacking gel solution should reach to the upper edge of the beveled plate, and then the gel solution has to be overlaid with a minimum volume of water. The water will stay on the surface because of surface tension. (B) After polymerization, the gel is mounted on the central cooling core of the electrophoresis unit, and the equilibrated IEF gel is placed on top of the polymerized stacking gel. Excess buffer is removed, and the IEF gel is overlaid with hot agarose/equilibration buffer mixture. After the agarose solidifies, the upper electrophoresis chamber is filled with buffer.
3. After the separating gel has polymerized (a sharp interface between the polymerized gel and the water overlay will reappear), remove the overlay, rinse the gel surface with water, and pour the stacking gel. The stacking gel solution should reach to the top of the bevel. Immediately overlay the stacking gel solution with a minimum amount of water, which will adhere owing to the surface tension (see Fig. 10.4.2A). A water overlay of the stacking gel provides a smooth surface and better contact between the IEF gel and second-dimension gel. A small volume of water has to be used to avoid lowering the upper edge of the stacking gel below the edge of the beveled plate. The stacking gel height must be between 1.5 and 2 cm. The solution is filled to the top of the bevel so that after the slight shrinkage that occurs during polymerization the top of the polymerized gel will be near the bottom of the bevel (see Fig. 10.4.2B).
Load the isoelectrofocusing gels onto the second-dimension gels 4. Assemble second-dimension gels in an electrophoresis chamber. Do not pour electrophoresis buffer into the upper chamber. 5. Melt 2% (w/v) agarose in a boiling water bath and add an equal volume of equilibration buffer for use in step 11. Keep agarose/equilibration buffer in the boiling water bath until step 11 is completed. 6. Retrieve isoelectrofocusing gels containing protein samples to be analyzed from storage. Incubate cryotubes containing frozen IEF gels in a 37°C water bath for 15 min for a 3-mm tube gel. A 5- to 7-min incubation is sufficient for 1.5-mm or thinner IEF gels. Do not agitate during thawing, as vigorous agitation of a partially thawed gel can break the gel. During this thawing/equilibration step, SDS in the equilibration solution in which the gels were frozen diffuses into the gel matrix and binds to proteins in the IEF gel. The length of incubation in the equilibration buffer is critical because insufficient saturation of proteins with SDS will contribute to vertical streaks on staining. On the other hand, extended incubation in equilibration buffer will result in excessive loss of proteins owing to diffusion of protein out of the gel, which is especially critical for thin IEF gels. For this reason, it is recommended that after extrusion from the IEF tube IEF gels be initially incubated for
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5 min to allow adequate diffusion of glycerol into the gel to minimize gel breakage, followed by freezing on dry ice (see Basic Protocol 1, step 30). This is desirable even if the second-dimension gel will be run directly after isoelectrofocusing, as it is the most feasible way of precisely controlling the equilibration time while the remaining gels in the IEF run are extruded.
7. Pour the gel and equilibration solution out of the cryovial onto a metal or plastic scoop. Carefully remove excess equilibration buffer with a pipet. 8. Place a few milliliters of electrophoresis buffer on the top of the second-dimension gel. 9. Slowly slide the IEF gel off the scoop and onto the top of the second-dimension gel. Remove all air bubbles trapped between the gels. Remove excess electrophoresis buffer from the top of the second-dimension gel. The basic end of the gel may be placed on either the left or right side of the second-dimension gel. However, once a convention is established, all gels should be oriented the same way. The acidic end of the IEF gel can be recognized in two ways: the bromphenol blue will usually be yellow, and a bulge (increased gel diameter) will be present.
10. Place a piece of agarose containing molecular weight standards (see Support Protocol 6) beside the basic side of the IEF gel (optional). Note that when molecular weight standards are used, the isoelectrofocusing gel has to be shorter than the width of the second-dimension gel.
11. Carefully overlay the IEF gel (and the gel piece with standard proteins) with the hot agarose/equilibration buffer mixture (∼2 ml/gel) prepared in step 5. Let the agarose solidify. The agarose prevents the IEF gel from shifting position and ensures good contact between the IEF and second-dimension gels.
12. Carefully pour electrophoresis buffer into the upper reservoir, taking care to avoid disturbing the agarose-covered IEF gel. 13. Connect electrodes and run the gels. See UNIT 10.1 for electrophoresis conditions. BASIC PROTOCOL 4
SECOND-DIMENSION ELECTROPHORESIS OF IPG GELS In this protocol, vertical gel electrophoresis is used as the second dimension for IPG gels in an analogous manner to the protocol described for the second dimension of IEF tube gels (see Basic Protocol 3). One difference is the use of second-dimension gel spacers or gel apparatus that will accommodate an 18-cm-long Immobiline DryStrip; Bio-Rad offers a conversion kit to increase the gel width from 16 cm to 18 cm, and Hoefer Pharmacia offers the Iso-Dalt gel system. The use of beveled plates is not necessary as the 0.5-mm strips are narrower than the second-dimension gel (1.0 or 1.5 mm thick). Another change involves a two-step equilibration of the strips prior to electrophoresis. Additional Materials (also see Basic Protocol 3) DryStrip equilibration solutions 1 and 2 (see recipes; prepare fresh in step 4) Immobiline IPG DryStrip with focused protein (see Basic Protocol 2) Platform shaker Cast the second-dimension gel 1. Assemble the glass-plate sandwich of an electrophoresis apparatus, using gel plates wide enough to accommodate an 18-cm-long DryStrip gel.
Two-Dimensional Gel Electrophoresis
Beveled plates are not necessary. If the spacers are not wide enough to accommodate an 18 cm gel, the ends of the gel strip may be trimmed away from the IPG gel so that it will fit on top of the second dimension; however, some very basic or acidic proteins may be lost.
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2. Pour a separating gel of the desired acrylamide concentration and immediately overlay with water to produce a smooth surface. The separating gel should be a minimum of ∼2.5 cm below the top of the inner plate to accommodate a 2-cm stacking gel.
3. After the separating gel has polymerized, remove the water overlay, rinse the gel surface with water to remove any unpolymerized acrylamide, and pour the stacking gel to a height of 0.5 cm from the top of the plate. Overlay with water to produce a smooth surface. A water overlay provides a smooth surface for better contact between the Immobiline DryStrip and the second dimension gel. The stacking gel height should be ∼2 cm.
Load the Immobiline IPG DryStrip gel onto the second-dimension gel 4. Prepare Immobiline DryStrip equilibration solutions 1 and 2 (see recipe). 5. Assemble the second-dimension gels in a electrophoresis chamber. Do not pour electrophoresis buffer into the upper chamber. 6. Melt 2% (w/v) agarose in a boiling water bath. Mix a solution of 1 part 2% agarose to 2 parts equilibration solution 2. Keep agarose/equilibration buffer mixture in boiling water bath until step 11 is completed. The agarose prevents the IPG DryStrip from shifting position and ensures good contact between the IEF and second-dimension gels.
7. Using forceps, remove the IPG gels from the electrophoresis tray after isoelectrofocusing is complete or from the −80°C freezer (see Basic Protocol 2, step 19) and place each strip in a separate petri dish with the support film side of the strip facing the petri dish wall. Add 15 ml of DryStrip equilibration buffer 1. Cover and place on a platform shaker for 10 min. Strips may be run in the second dimension immediately after isofocusing or after storage at −80°C. If the strips have been stored at −80°C, remove them from the freezer, then place in petri dish as stated and continue with the equilibration procedure.
8. Discard equilibration buffer 1 and add 15 ml of equilibration buffer 2. Cover and place on a platform shaker for 10 min. 9. Dampen a piece of filter paper and place on a glass plate. Remove the DryStrips from equilibration buffer 2. Place each strip on its edge on the filter paper to remove any excess buffer. Strips should not be left in this position for >10 min, or spot sharpness may be affected.
10. Add a small amount of SDS electrode buffer along the glass plate above the second-dimension gel. Place the DryStrip gel in the well with the gel facing out and the basic side to the left. Push the DryStrip down so that it is firmly in contact with the stacking gel of the second-dimension gel. Remove excess running buffer. 11. Overlay the IPG gel strip with the agarose/equilibration buffer (from step 6) and allow agarose to solidify. 12. Carefully pour electrophoresis buffer into the upper reservoir, taking care to avoid disturbing the agarose-embedded IPG DryStrip. 13. Connect electrodes and run the gels. See UNIT 10.1 for electrophoresis conditions and UNIT 10.5 for gel staining conditions. Electrophoresis
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SUPPORT PROTOCOL 6
PREPARING MOLECULAR WEIGHT STANDARDS FOR TWO-DIMENSIONAL GELS Molecular weight markers are usually necessary for the identification of proteins or as references to describe experimental proteins on two-dimensional gels. In many cases, molecular weight markers are required only at the beginning of a project. Once the system is established, common proteins in the sample (e.g., actin or tubulin) provide sufficient references for molecular weight identification on subsequent gels. To minimize any differences in migration of the molecular weight standards and isoelectrofocused proteins, the standard proteins should be loaded on the second-dimension gel in the same manner as the IEF gel. This protocol describes the preparation of standards in solidified agarose. The agarose pieces may be stored at −80°C for at least a year and provide a convenient source of standards for the second-dimension gel. The procedure described is recommended for 3-mm IEF gels. Narrow standards in solidified agarose (made in tubes ≤1.5 mm in diameter) can be prepared by the same method, but extrusion of the thinner agarose gel without breaking is more difficult. The protocol supplies molecular weight markers containing ∼2.5 µg of each standard suitable for Coomassie blue staining or 0.25 µg of each standard for silver staining. Materials Molecular weight standards (Table 10.1.2) 1× SDS sample buffer (UNIT 10.1) 2% (w/v) agarose (see recipe) Boiling water bath Glass tubes (3-mm inner diameter) Plastic or metal tray 1. Prepare 3 ml molecular weight standards in 1× SDS sample buffer using 250 µg of each standard. The stated amount is appropriate for Coomassie blue staining of gels. If silver staining is planned, use 25 ìg of each standard.
2. Mix the standards with 2 ml of 2% (w/v) agarose melted in a boiling water bath. 3. Prepare clean glass tubes by wrapping one end with Parafilm. Pour the hot mixture into the tubes and let the agarose solidify. 4. Carefully extrude the agarose from the tubes. 5. Cut agarose rods into 5-mm pieces using a razor blade. 6. Freeze all pieces separately on a plastic or metal tray using dry ice. 7. Collect frozen pieces in a plastic bottle and store at −80°C. The standards may be stored ≥1 year.
Two-Dimensional Gel Electrophoresis
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DIAGONAL GEL ELECTROPHORESIS (NONREDUCING/ REDUCING GELS)
ALTERNATE PROTOCOL 3
Protein subunit compositions and cross-linked protein complexes can be analyzed by two-dimensional gel electrophoresis using separation under nonreducing conditions in the first dimension followed by reduction of disulfide bonds and separation under reducing conditions in the second dimension. Most proteins will migrate equal distances in both dimensions, forming a diagonal pattern. Proteins containing interchain disulfide bonds will be dissociated into individual subunits and can be resolved in the second-dimension gel. The approach is similar to that described for two-dimensional gel electrophoresis (see Basic Protocol 3) except, in this protocol, the first-dimension gels are nonreducing (i.e., 2-mercaptoethanol or dithiothreitol is omitted from sample buffer) SDS-denaturing gels instead of isoelectrofocusing gels. Use of 1.2-mm tube gels for the first-dimension separation and 1.5-mm slab gels for the second-dimension run is recommended. Additional Materials (also see Basic Protocol 3) Separating and stacking gel solutions (see Table 10.1.1) 1× SDS sample buffer without reducing agents (UNIT 10.1) Reducing buffer (see recipe) 1.5% (w/v) agarose in reducing buffer (see recipe; optional, for securing first-dimension gel on second-dimension gel) Two-dimensional comb (optional) Additional reagents and equipment for casting tube gels (see Basic Protocol 1), SDS-PAGE (UNIT 10.1), and protein staining (UNIT 10.5) Pour and run the first-dimension gel 1. Clean and dry 1.2-mm glass gel tubes for the first-dimension gel (see Basic Protocol 1, step 1). 2. Prepare a separating gel solution with the desired percentage acrylamide (Table 10.1.1); omit the stacking gel for the first dimension. Stacking gels can usually be avoided in the first dimension by keeping sample volumes small (i.e., ≤10 ìl). Less than 200 ìl of gel solution is required to cast a single 1.2-mm tube gel 12 cm in length. Adjust the amounts from Table 10.1.1 accordingly.
3. Cast the first-dimension polyacrylamide gels in 1.2-mm tubes using a syringe with a long needle (see Basic Protocol 1, step 5a). Overlay with water and allow the gels to polymerize. 4. Prepare samples in 1× SDS sample buffer without any reducing reagents (i.e., no 2-mercaptoethanol or DTT). Load the samples and electrophorese until the tracking dye is ∼1 cm from the bottom of the tube. Reduce sample and run the second-dimension gel 5. Extrude the gel from the tube (see Basic Protocol 1, steps 27 to 29). 6. Place the extruded gel in a test tube containing 5 ml reducing buffer. Equilibrate 15 min at 37°C with gentle agitation. 7. Cast the second-dimension separating and stacking gels (see Basic Protocol 3, steps 1 to 3), making sure that the top of the stacking gel is at least 5 mm below the top of Electrophoresis
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the short glass plate. Layer water across the entire stacking gel or use a two-dimensional comb. Most two-dimensional gel combs have a separate small well for a standard or reference sample. The use of beveled plates (see Basic Protocol 3, steps 1 to 3) is not essential but is still preferred because it will facilitate loading of the first-dimension gel. In this procedure, the first-dimension gel will fit between the glass plates if 1.2-mm tubes are used for the first dimension and 1.5-mm gels are used for the second dimension.
8. Load the first-dimension gel onto the second-dimension gel. Remove any air bubbles trapped between the gels. If the first-dimension gel does not remain securely in place, it can be embedded using 1.5% (w/v) agarose in reducing buffer.
9. Carefully pour electrophoresis buffer into the upper electrophoresis chamber and electrophorese using voltages and times appropriate for the gel type selected. Parameters for electrophoresis are given in UNIT 10.1. SUPPORT PROTOCOL 7
USING TWO-DIMENSIONAL PROTEIN DATABASES Computerized image acquisition and manipulation constitute the only practical method for systematic qualitative and quantitative evaluation of complex protein patterns from different samples that are to be compared by high-resolution two-dimensional gel analysis. Examples of experimental applications include comparisons of tumor cells or tissues with appropriate normal controls and comparisons of a single cell line under different experimental conditions. There are currently a number of commercially available image acquisition/computer systems specifically designed for comparing two-dimensional gels and storing associated information in a database (UNIT 10.12). The systems include both hardware and the necessary software for comparing different gels and producing databases containing the two-dimensional protein patterns, with options for annotating specific spots and producing quantitative comparisons among large numbers of different samples. With most systems, images can be acquired from either stained gels or autoradiographs. The equipment used to obtain two-dimensional gel images includes laser scanners, video cameras, and phosphoimagers. After image acquisition, software running on a microcomputer or workstation is used to refine the image, detect spots, and match spots between different gels. It is essential that very high-quality, reproducible gels be used for computerized comparisons. The greatest dynamic range in protein abundance for a single two-dimensional gel can be obtained using autoradiography (UNIT 10.12) or phosphoimaging. With these methods, up to several thousand spots can be compared and tracked. A representative reference gel or a composite image can be stored and used as a reference for future experiments.
Two-Dimensional Gel Electrophoresis
Information related to each spot on the two-dimensional pattern, including the quantity of protein in the indicated spot on different gels used in the comparison, can be archived and updated. Other known information related to a specific spot can also be added to the investigator-built database, including the pI, molecular weight, amino acid composition, sequence, and/or identity of the protein and any other important attributes correlated with the indicated spot. A number of research groups, including those of Garrels and Celis (Garrels, 1989; Garrels and Franza, 1989; Celis et al., 1991), have extensively characterized hundreds of spots from specific cell lines and have used multiple methods to characterize proteins of interest. The most definitive methods for establishing the identi-
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ties for proteins of interest detected by computer-assisted comparisons are protein sequence analysis and, more recently, mass spectrometry of tryptic fragments. Both methods are compatible with the quantities of protein that can be recovered from two-dimensional gels. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
30% acrylamide/0.8% bisacrylamide 30 g acrylamide 0.8 g bisacrylamide H2O to 100 ml Filter solution through 0.2- to 0.45-µm filter (e.g., Micro Filtration Systems, cellulose nitrate, 0.2 µm). Store at 4°C (stable at least 3 months). Acrylamide is a neurotoxin. Wear gloves and a dust mask when handling solid acrylamide. Wear gloves when working with acrylamide solution. Never pipet acrylamide solutions by mouth.
Agarose in reducing buffer, 1.5% (w/v) Mix 0.15 g agarose and 10 ml reducing buffer (see recipe). Heat in boiling water bath until dissolved. Prepare immediately before use. Agarose, 2% (w/v) Mix 2 g agarose and 100 ml water. Stir on a hot plate until dissolved. Keeping the solution near 100°C, divide by placing 5-ml aliquots in 25-ml glass screw-cap tubes. Let the aliquots solidify. Store at 4°C (stable at least 3 months). Ammonium persulfate, 2.5% (w/v) 0.25 g ammonium persulfate H2O to 10 ml Prepare immediately before use DNase and RNase solution Mix 2.50 ml of 1 M Tris⋅Cl, pH 7.0 (APPENDIX 2E), 250 µl of 1 M MgCl2, (APPENDIX 2E), and 2.2 ml water. Add to 5 mg DNase (Worthington) and dissolve DNase. Add 2.5 mg RNase (in solution; Worthington). Mix well, divide into 50- or 100-µl aliquots, and store at −80°C (stable at least 1 year). The volume of RNase solution needed is variable and depends on the protein concentration, which is reported on the vial label (in mg/ml). The volume of RNase added should be less than 200 ìl; if a larger volume is used, the amount of H2O should be decreased proportionally. Final concentrations are 0.5 M Tris⋅Cl, pH 7.0, 0.1 M MgCl2, 0.1% (w/v) DNase, and 0.05% RNase.
DryStrip equilibration solutions 20 ml 1 M Tris⋅Cl, pH 6.8 (APPENDIX 2E) 72 g ultrapure urea 60 ml glycerol 2 g sodium dodecyl sulfate (SDS) 67 ml Milli-Q water For solution 1: Add 50 mg DTT per 10 ml of equilibration buffer For solution 2: Add 0.45 g iodoacetamide and a few grains bromphenol blue per 10 ml of equilibration buffer Make fresh immediately before use Electrophoresis
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Final concentrations are 50 mM Tris⋅Cl, pH 6.8; 6 M urea; 30% glycerol; and 1% SDS, in a final volume of 200 ml.
EDTA, 2% (w/v) 2 g Na2EDTA H2O to 100 ml Adjust to pH 7.0 with NaOH Store at room temperature (stable several months) Titrate while dissolving. EDTA is difficult to dissolve without addition of NaOH even when the disodium salt is used.
Equilibration buffer 3 g SDS 7.4 ml 2% (w/v) EDTA, pH 7.0 (see recipe) 10 ml glycerol 2 ml 1.0 M Tris⋅Cl, pH 8.65 (APPENDIX 2E) 0.3 ml bromphenol blue (saturated solution in H2O) H2O to 100 ml Store at room temperature (stable for several weeks) Final concentrations are 3% (w/v) SDS, 0.4 mM EDTA, 10% (v/v) glycerol, and 20 mM Tris⋅Cl, pH 8.65.
Leupeptin, 2 mg/ml 20 mg leupeptin 10 ml water Divide into convenient volumes Store at −20°C (stable at least 1 year) Lysis buffer 2.59 g urea (ultrapure) 1.6 ml H2O 0.25 ml 2-mercaptoethanol 0.3 ml ampholytes 1.0 ml 20% (w/v) Triton X-100 solution (see recipe) Prepare immediately before use Use same ampholytes as for the IEF gel formulation. To dissolve urea, warm the mixture in a 30°C water bath if necessary.
Orthophosphoric acid (H3PO4), 0.1 M 13.7 ml 85% phosphoric acid Water to 2 liters Make fresh daily Must be degassed prior to use.
PBS, 10× 152 g NaCl 24 g monobasic sodium phosphate, anhydrous 1600 ml H2O Adjust pH to ∼6.7 with NaOH Add H2O to 2 liters Store at room temperature (stable at least 1 month) The 1× solution should be pH 7.3 to 7.5. Final concentrations are 130 mM NaCl and 10 mM sodium phosphate. Two-Dimensional Gel Electrophoresis
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PBS with proteolysis inhibitors (PBS/I buffer) 20 ml 10× PBS (see recipe) 20 ml 2% (w/v) EDTA, pH 7.0 (see recipe) 200 µl 0.15 M phenylmethylsulfonyl fluoride (PMSF) in 2-propanol 100 µl 2 mg/ml leupeptin (see recipe) 200 µl 1 mg/ml pepstatin (see recipe) Adjust to pH 7.2 with HCl Add H2O to 200 ml Prepare immediately before use Diisopropyl fluorophosphate (DFP) is a better serine protease inhibitor than PMSF at lower temperatures (0° to 4°C); however, although both compounds are toxic, exceptional caution must be exercised with DFP owing to its volatility. If DFP is used, work in a chemical fume hood and carefully follow the supplier’s precautions. DFP and PMSF have half-lives on the order of hours in aqueous neutral solutions, and the degradation rate increases rapidly as the pH is increased above neutral. Make aqueous solutions immediately before use. Use of 1 M NaOH is convenient to inactivate residual DFP or PMSF. Final concentrations are 10 mM sodium phosphate, 130 mM NaCl, 0.2% EDTA, 0.15 mM PMSF; 1ìg/ml leupeptin, and 1 ìg/ml pepstatin.
Pepstatin, 1 mg/ml 10 mg pepstatin 10 ml anhydrous ethanol Divide into convenient volumes Store at −20°C (stable at least 1 year) Thaw aliquots and mix well immediately before use.
Reducing buffer 0.5 g dithiothreitol (DTT) 0.1 g SDS 1.51 g Tris base Adjust to pH 6.8 with HCl Add H2O to 100 ml Prepare fresh every time Final concentrations are 0.5% (w/v) DTT, 0.1% (w/v) SDS, and 125 mM Tris⋅Cl, pH 6.8.
Tris/SDS buffer 0.3 g SDS 0.6 g Tris base Adjust to pH 8.0 with HCl Add H2O to 100 ml Divide into 5-ml aliquots Store at −80°C (stable at least 1 year) Final concentrations are 0.3% (w/v) SDS and 50 mM Tris⋅Cl, pH 8.0.
Triton X-100 solution, 20% (w/v) 3 g Triton X-100 12 ml H2O Warm in 37°C water bath to dissolve Triton X-100 Store at 4°C (stable ∼2 weeks) Urea, 8 M 0.75 g ultrapure urea 1.0 ml H2O Prepare immediately before use Avoid heating above room temperature.
Electrophoresis
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COMMENTARY Background Information
Two-Dimensional Gel Electrophoresis
Two-dimensional gel electrophoresis, using isoelectricfocusing followed by SDS-PAGE, is the single most powerful analytical method currently available for separating complex protein mixtures such as whole-cell or tissue extracts. It is therefore a valuable method for following disease-related changes or for detecting changes in protein expression under diverse experimental conditions. To achieve maximum reproducibility between samples to be compared, multiple gels should be cast and run simultaneously (Anderson and Anderson, 1978a,b). Despite the exceptionally high resolving power of the method, the total number of proteins that can be resolved in a single two-dimensional gel is only ∼1000 to 2000, whereas the total number of proteins in a single mammalian cell type is likely to be at least 10 to 20 times higher. Therefore, the old guideline that a single spot on a two-dimensional gel is a single protein needs to be revised as analytical detection methods improve. Conversely, a single protein (single gene product) can produce multiple, usually adjacent spots in the isoelectrofocusing dimension owing to variable degrees of chemical or post-translational modification. Common examples of variable posttranslational modifications that can usually be detected on two-dimensional gels include phosphorylation and glycosylation. Examples of chemical modifications that cause charge heterogeneity include deamidation of sidechain amines (usually asparagines), oxidation of sensitive side chains, and modification of lysines. Potential modification of lysines by urea is particularly important because urea rapidly decomposes to form cyanate, which readily reacts with amino groups, especially above pH 7. Despite potential side reactions, urea is the most useful IEF additive for maintaining solubility of proteins near their isoelectric points. Combined with electroblotting to PVDF membranes, two-dimensional PAGE has become a valuable preparative tool for protein isolation in addition to its historical role as an analytical method. The sensitivity of many protein analysis methods has improved to the point where one or several spots from two-dimensional gels are sufficient for N-terminal or internal sequence analysis. Commercially available equipment for running two-dimensional gels can be divided into
four groups based on size: microgels (Hoefer Pharmacia Phast system), minigels (e.g., BioRad or Hoefer Pharmacia), standard or fullsized gels (e.g., Bio-Rad or Hoefer Pharmacia), and large or “giant” gels (ESA Investigator 2D gel system or Hoefer Pharmacia Iso-Dalt gel system). In general, the larger the gel, the better the final resolution, but as gel size increases so do costs, difficulty of gel handling, and time requirements. Standard-size gels provide adequate resolution for most applications and are relatively easy to handle. A 3-mm soluble ampholyte tube IEF gel or a 0.5 mm × 3 mm × 18-cm Immobiline gel has a total protein capacity of ∼500 µg for complex protein mixtures such as wholecell extracts. The maximum capacity for any single protein spot is ∼0.5 to 5 µg, depending on the solubility of the protein near its isoelectric point and the separation distance from any near neighbors. A variety of alternative gel sizes, their limits, and their advantages are summarized in Table 10.4.2. The lower protein limit for any of the systems is determined strictly by the available detection methods (UNIT 10.5). Proteins can be detected in two-dimensional gels by the same wide range of techniques used for one-dimensional gels. Autoradiography (UNIT 10.11), silver staining (UNIT 10.5), and electroblotting to PVDF membranes (UNIT 10.7) followed by colloidal gold or colloidal silver staining (UNIT 10.8) or immunodetection (UNIT 10.10) are among the most sensitive techniques available. If a larger amount of protein is available, Coomassie blue staining of the gel (UNIT 10.5) or amido black staining of a PVDF membrane after electrotransfer (UNIT 10.8) would be the detection methods of choice. The major technical limitation in two-dimensional gel electrophoresis is gel-to-gel variation. Even when extreme care is exercised to produce highly reproducible first- and seconddimension gels, some gel-related variability among gels cast at the same time is likely to persist. Another source of variability includes differences in extraction or recovery of proteins during sample solubilization and handling. Maximizing resolution and reproducibility is especially important if computerized comparisons of two-dimensional gels of complex protein mixtures such as cell or tissue extracts are being attempted.
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Table 10.4.2
Size Options for Two-Dimensional Gel Electrophoresis
First-dimension gel Gel type
Microgels/minigelsc
Full-size gelse
Giant gelsf
Diameter D Length L (cm) (mm) <1.5 <1.5 >1.5 <1.5 <1.5 >1.5 <1.5 >1.5
<10 <10 <10 12-18 12-18 12-18 >20 >20
Second-dimension gela Thickness T (mm)
Height H (cm)
D D D
<10 <10 <10 12-18 12-18 12-18 >20 >20
Purpose
Analytical Analytical Analytical/preparative Analytical Analytical Analytical/preparative Analytical Analytical/preparative
Commentsb
1-4 3-6 1,2,7,8 1-4 3-6 1,2,7,8 4-6,9 1-3,7
aThe
second-dimension gel width has to be at least equal to the IEF tube gel height. to comments: (1) tube gel cannot be placed directly on top of second-dimension gel, and use of agarose is recommended; (2) use of stacking gel is recommended; (3) extrusion and handling are relatively difficult; (4) total protein load is limited to usually ≤50 µg for whole-cell or tissue extracts; (5) tube gel can be placed directly on top of second-dimension gel, and use of agarose is not necessary; (6) use of a stacking gel is not necessary; (7) total protein load capacity is relatively large; (8) extrusion and handling are relatively simple; (9) extrusion and handling are very difficult. cMinigel systems provide rapid separations with moderate resolution. Microgels (Phastgels) are precast gels that are slightly smaller than most minigels. dUse of second-dimension gels thicker than 1.5 mm is generally not recommended owing to difficulty with either efficient staining or efficient electroblotting. eFull-size gels provide resolution satisfactory for most applications. fGiant gels provide very good resolution. Specialized equipment is required, such as Investigator 2D (ESA), Iso-Dalt (Hoeffer Pharmacia), or homemade giant-size gel systems. bKey
Isoelectrofocusing using soluble ampholytes Soluble ampholytes are mixtures of lowmolecular-weight organic compounds with differing side-chain pKa values that provide buffering capacity. In an IEF gel, the ampholytes migrate to their isoelectric point, where they provide buffering capacity and hence produce stable pH gradients. In theory, any desired pH gradient could be produced by blending ampholytes with appropriate pKa values. In practice, it is relatively easy to produce pH gradients from ∼pH 3.0 or 4.0 to pH 8.0, but stable soluble gradients outside this range are usually not technically feasible. Within these pH limits, some manipulation of the gradient shape and pH range is possible by blending different amounts of specific pH range ampholytes. For example, 0.50 ml of pH 5-7 ampholytes plus 0.25 ml of pH 4-8 ampholytes can be used instead of 0.75 ml of pH 4-8 ampholytes alone to increase the separation distance of proteins in the pH 5.0 to 7.0 range. Basic Protocol 1 is based on use of 3-mm first-dimension isoelectrofocusing gels and 1.5-mm second-dimension gels using the BioRad two-dimensional gel apparatus (Protean II
xi 2D). The method can be easily adapted to equipment from other suppliers or to differentsized gels by adjusting the quantities of reagents used. The protocol uses 8 M urea and Triton X-100 as solubilizing agents. Solubilization of the protein sample applied to the gel as well as maintenance of solubility during electrofocusing are the most critical factors influencing the quality of separation in the first dimension. The most common modification to Triton X-100–based procedures is addition of 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS) to the gel and solubilizing buffer mixtures. Addition of SDS to complex samples such as tissue or cell extracts can also enhance reproducible solubilization of the largest possible subset of proteins. Although SDS is charged, in the presence of Triton X-100 it is separated from proteins during focusing and migrates to the acidic end of the gel. Regardless of the method used to maintain solubility, some proteins (especially those >100 kDa) tend to precipitate at pH values approaching their isoelectric point and thus produce horizontal smears on the final seconddimension gel.
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Table 10.4.3
Commercially Available Precast IPG Gels and Immobiline Chemicalsa
Name
Use
Immobiline DryPlate
Running one-dimensional immobilized pH pH 4.0-7.0, pH 4.2-4.9, pH 4.5-5.4, gradient gels pH 5.0-6.0, pH 5.6-6.6 Running first dimension in two-dimensional gels pH 4-7L, pH 3-10L pH 4-7L, pH 3-10Lb, pH 3-10NLc Creating custom gradient immobilized pH pK 3.6, pK 4.6, pK 6.2, pK 7.0, gradient gels pK 8.5, pK 9.3
Immobiline DryStrip 110 mm 180 mm Immobiline II
Available pH range
aFrom
Hoefer Pharmacia. linear gradient with maximum resolution above pH 7.0. cA nonlinear gradient with best resolution at pH 5.0-7.0. bA
Basic Protocol 1 requires several modifications for successful separation of very acidic (Alternate Protocol 1) or very basic (Alternate Protocol 2) proteins. In both cases, prefocusing of the gels must be avoided, and the isoelectrofocusing time has to be reduced. A short separation time does not allow the system to reach equilibrium and is used to establish the desired pH gradient. Separation of very acidic proteins requires modifications of gel and electrode solutions. Separation of very basic proteins has to be performed with the positions of electrodes and electrode solutions reversed in the electrophoresis chamber. The total protein load per gel depends on the complexity of the sample, the solubility of proteins in the sample, and the diameter of the first-dimension gel. Approximately 4 times as much total sample can be applied to a 3-mm gel compared with a 1.5-mm gel. Another advantage of a larger-diameter isoelectrofocusing gel is that extrusion and gel handling are easier owing to improved strength of the gel. If care is exercised in loading the first-dimension gel onto the 1.5-mm second-dimension gel, the final resolution will be similar to that obtained using smaller-diameter IEF gels. The separating or resolving power of a system is dependent on the quality of ampholytes used, the slope of the pH gradient, and the lengths of both firstand second-dimension gels.
Two-Dimensional Gel Electrophoresis
Immobilized pH gradient gels In immobilized pH gradient (IPG) gels (Basic Protocol 2), the pH gradient is an integral part of the polyacrylamide matrix (Strahler and Hanash, 1991). Because the pH gradient is covalently associated with the polyacrylamide gel matrix, precise, reproducible, and very high-resolution separations can be achieved. A
variety of precast gels and all the necessary equipment are commercially available from Hoefer Pharmacia. Reproducible two-dimensional gels can be obtained by running a sample on a narrow strip of immobilized pH gradient gel and then running it in a vertical SDS second-dimension gel. Isoelectrofocusing is performed in a horizontal electrophoresis unit in which multiple gel strips may be run simultaneously. Equipment and reagents are also available for the user to cast custom Immobiline gels in the laboratory. Some of the major advantages of using precast IPG gels are their ease of use and high reproducibility, the time savings realized, and the fact that precision narrow-range gradients can be used to resolve small charge differences. Because the pH gradient is covalently coupled to the polyacrylamide gel matrix, the pH gradient remains stable and linear during prolonged electrophoresis, thus ensuring reproducibility. This is in contrast to conventional IEF gels, where gradient drift occurs during prolonged electrophoresis. Additionally, the precast gels can be rehydrated in water or in solutions with one or more additives such as urea, CHAPS or Triton X-100, carrier ampholytes, glycerol, and reducing agents, which may help to increase protein solubility. Precast gel strips are available with pH ranges such as pH 3.0 to 10.0 or pH 4.0 to 7.0 as well as narrower ranges (as DryPlates). In addition, a variety of Immobilines permits the user to cast IPG gels with customized pH gradients of any gradient range and shape between pH 3.0 and pH 10.0. Table 10.4.3 lists types of commercially available gels and Immobilines. With the Immobiline system, the apparent pI of a given protein may be slightly different from that determined by other methods. Therefore, it is
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recommended that a broad-range gradient be tried initially, followed by a narrower-range gradient, if needed. Diagonal gel electrophoresis Diagonal gel electrophoresis is a form of two-dimensional analysis useful for investigating the subunit composition of multisubunit proteins containing interchain disulfide bonds (Goverman and Lewis, 1991). Proteins are electrophoresed in the first-dimension in a tube gel (or a slab gel) under nonreducing conditions. The proteins are then reduced in situ, and the first gel (or a strip thereof) is layered onto a second gel and electrophoresed. In the second gel, the proteins migrate at right angles to the original, first-dimension migration. Most cellular proteins are not disulfide-linked and will fall on the “diagonal” in this system; that is, they migrate approximately equal distances in both directions during electrophoresis and lie approximately on the diagonal line connecting opposite corners of the gel. On reduction, component subunits of proteins connected by interchain disulfide bonds will resolve below the diagonal because the individual subunits migrate faster than the disulfide-linked complex during the second electrophoresis. Some proteins with internal disulfide bonds, but no interchain disulfides, may migrate slightly above the diagonal because internal disulfides can produce a more compact molecular shape (causing faster migration in the first dimension). Investigator 2D gel system The Investigator 2D gel system was introduced in 1990 by Millipore as the first commercial “large” or “giant” format two-dimensional gel system designed for analytical purposes, although analogous homemade units had been reported earlier (Garrels, 1979; Young et al., 1983). This product line, including precast first- and second-dimension gels, can now be purchased from ESA. The Investigator 2D gel system uses larger gel sizes than the gels described in this unit as well as a number of novel approaches and reagents designed to enhance resolution and gel-to-gel reproducibility. The major features of this system include the following: an increased length of the first-dimension gel (20 cm), an increased length and width of the second-dimension gel (20 × 22 cm), a thread reinforcement of the isoelectrofocusing gel, temperature control during electrophoresis of the second-dimension gel, and use of a special high-tensile-strength acrylamide to facilitate handling of the large, thin second-dimen-
sion gel. Narrow gel tubes (1.2 mm) for the first-dimension are standard, but 3-mm-innerdiameter tubes are available to accommodate larger protein loads. The 3-mm IEF gels do not have the thread reinforcement. One modification related to the above protocol includes addition of 0.3% (w/v) CHAPS in the gel solution, which is intended to improve the solubility of proteins and their migration and separation in the gel. The system is supplied with a manual that adequately describes the method.
Critical Parameters and Troubleshooting Although no individual steps in two-dimensional gel analysis are exceptionally difficult, the large number of steps involved increase the likelihood and possible severity of errors or problems. Several steps are especially critical and may require optimization. The first is sample preparation. Proteins applied to IEF gels have to be completely solubilized. Residual precipitate or even soluble aggregates are likely to cause artifacts on the end of the tube gel or position on the IPG strip where the sample is loaded. Precipitated or aggregated proteins may also interact with soluble proteins, causing components with normally good solubility to coprecipitate or migrate anomalously. Two general rules of thumb apply to sample solubility: (1) the more complex the sample or the more crude the extract, the more likely problems will be encountered with sample solubility, and (2) the higher the protein load applied to the gel, the more likely solubility problems will arise. Care must also be taken to avoid proteolysis during sample preparation, especially when complex, impure samples are used. If samples are frozen either before or after solubilization, they should be stored at −80°C and should not be subjected to repeated freezethawing. Addition of SDS to the solubilization solution increases the solubilization of some proteins; however, SDS should only be used when the IEF gels contain urea and Triton X-100, as the solubilizing agents are required for effective separation of the strongly anionic SDS molecules from the proteins during isoelectrofocusing. Heating of samples in ureacontaining solutions must be avoided because urea readily decomposes to cyanate, which reacts with amino groups and causes charge heterogeneity. High concentrations of salts and buffers in the sample should be avoided. Ionic compounds increase the conductivity of the sample and can result in localized overheating, especially for Immobiline gels.
Electrophoresis
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Two-Dimensional Gel Electrophoresis
Samples analyzed on immobilized pH gradient gels should not contain precipitates. The concentration of salts and buffer ions in the sample should be kept to a minimum (<50 mM) to avoid local overheating of the gel during electrophoresis. If the sample forms aggregates or precipitates at the point of application, apply the sample to a different location (different pH) on the gel. Early decisions that must be made include the size of the gels required and whether a soluble ampholyte system or an Immobiline gel will be used. The major consideration affecting appropriate gel size is the degree of resolution needed. In general, the smallest gel format should be selected that will provide the needed degree of resolution, because smaller gels are easier, less expensive, and faster to run. Therefore, quick screening of samples or analysis of relatively simple samples can easily be accomplished with microgels or minigels. In contrast, if detailed qualitative or quantitative comparisons of cell or tissue extracts are planned, standard-size or large gels are indicated. Similarly, 3-mm or larger first-dimension tube gels followed by 1.5-mm second-dimension gels in the standard or large format are indicated if the two-dimensional gels will be used for preparative isolation of a protein for applications such as raising antibodies or conducting structural analysis. Immobilized pH gradient gels should very seriously be considered as an alternative to soluble ampholyte gels for most separations owing to their stable and reproducible pH profiles. The IPG gels are particularly appropriate when a narrow pH range is required. Immobilized pH gradient gels are typically run at 2500 to 3500 V and typically require a focusing time of 16 to 18 hr. Optimal focusing conditions may be experimentally determined by applying the sample to different positions on the gel and estimating the time for the migration patterns to coincide. Some proteins may require longer run times to reach their pI. Because there is no gradient drift, the potential problems with longer run times are limited to sample denaturation or drying out of the gel. These problems usually can be minimized by including a reducing agent in the rehydration solution and/or coating the top surface of the gel with paraffin oil. The quality of the first-dimensional separation is strongly dependent on the purity of the reagents used, especially the urea and ampholytes. One fairly commonly encountered frustration, when soluble ampholyte gels are used, is that different batches of ampholytes from the
same supplier will sometimes produce markedly different two-dimensional gel patterns. Therefore, it is advisable to purchase an adequate supply of a single lot of ampholytes to meet anticipated needs for an entire study where such an approach is feasible. However, whereas ampholytes usually have a reasonably long shelf life at 4°C (usually up to a year), shelf life as well as total ampholyte requirements often cannot be predicted with much certainty. When any doubt arises about the purity or quality of ampholytes or any other reagent, the reagent should be replaced immediately. Constant monitoring of the system performance, especially when changing lots of ampholytes, urea, or acrylamide, can help minimize potential reagent-associated problems. In most cases, the best standard for a given two-dimensional gel system is an experimental sample or control that is available in sufficient quantity so that many replicate aliquots can be frozen and stored for an extended time at −80°C; alternatively, a sample that can be reproducibly prepared over a long time frame would make an acceptable standard. Such an experimental standard or reference is more likely to detect subtle, but experimentally important, changes in the two-dimensional gel system than commercially available IEF or SDS gel standard mixes. Another critical factor is equilibration of the first-dimension gel in the second-dimension equilibration buffer. During this step, urea diffuses out of the IEF gel while SDS and reducing reagent diffuse into the gel. If the gel is inadequately saturated with SDS, vertical streaking will result. However, if the gel is incubated in the equilibration buffer for an extended time, a substantial amount of the protein can rapidly diffuse out of the large-pore IEF gel. Losses arising from diffusion can be critical for any experiment because different proteins will diffuse at different rates, but rigorous control of the incubation step is especially important if quantitative comparisons among gels are planned. The simplest method of controlling the incubation time is to freeze the extruded IEF gel after a carefully controlled 5-min incubation in the equilibration buffer; any additional equilibration incubation time required can then be incorporated and carefully controlled when the sample is thawed for loading onto the second-dimension gel. Another crucial step is loading of the equilibrated IEF gel onto the top of the second-dimension gel. Any irregularity or obstruction between the two gels, including particles of dirt or air bubbles, will affect the flow of current
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and disrupt the resolution of proteins in the final gel. Similarly, poor contact or any movement of the IEF gel during electrophoresis of the proteins out of the IEF gel into the seconddimension gel will lead to artifacts. Therefore, it is advisable to embed the IEF gel in a buffered agarose matrix to ensure good electrical contact between the gels and to prevent gel movement after electrophoresis is initiated. Finally, the choice of second-dimension gel composition and separation conditions can influence the quality of results. A proper percentage of acrylamide should be selected to optimize resolution within the desired molecular mass range. If gradient gels are needed, use of a multiple gel casting stand is the best way to ensure reproducibility among samples within a single experiment. Further details on optimization of two-dimensional gel systems are presented by Hochstrasser et al. (1988). When no technical, sample-related, or reagent-related problems are encountered, the final stained two-dimensional gels should contain numerous rounded or slightly elliptical spots. Typically, more than 1000 spots can be detected on a standard 16 × 14–cm gel when using a sensitive staining protocol such as silver staining or autoradiography and a whole-cell extract as a sample. Some horizontal streaks for most high-molecular-mass proteins (proteins exceeding ∼100 kDa) are common owing to the decreased solubility of larger proteins near their isoelectric points, even in the presence of urea and nonionic detergent. However, excessive horizontal smearing of proteins smaller than 100 kDa indicates poor isoelectrofocusing, which could be related to one or more of the following factors: sample improperly solubilized or contaminated with interfering substances such as large nucleic acid molecules, poor purity of reagents (check the urea first), poor-quality ampholytes, or insufficient isoelectrofocusing (total volt-hours too low). It is important to note that in general the solubility of any protein is the lowest near its isoelectric point, but there are vast differences among proteins in terms of both the minimum concentration where precipitation becomes a problem and the degree to which precipitation can be prevented by adding different solubilization agents. The best conditions for maintaining solubility during isoelectricfocusing for a given sample type must be determined empirically, although the most universal conditions are those described in the protocols in this unit. In contrast, IEF systems that do not use any detergents or denaturants are limited to that fairly
small percentage of proteins which maintain good solubility near their isoelectric point. If high-molecular-weight proteins are expected but are not present in the final two-dimensional gel, check the sample preparation protocol as well as the sample storage conditions. The most likely problem is proteolysis during sample preparation. Multiple freezethawing cycles could contribute to this problem. Vertical smears on the two-dimensional gel suggest (1) insufficient equilibration of the IEF gel (not enough SDS bound to the proteins), (2) poor contact between the IEF and second-dimension gels, or (3) problems related to the stacking gel (too short or wrong buffer). Use of a stacking gel is especially important when large-diameter IEF gels are loaded onto smaller second-dimension gels. Omission of Triton X-100 or other nonionic or zwitterionic detergent from the final sample loaded on the gel can yield poor results, especially for samples containing SDS, because the amount of detergent in the IEF gels alone may be insufficient to remove bound SDS from proteins. The presence of Triton X-100 in the sample is especially important when SDS sample buffer is used to solubilize protein samples (i.e., after immunoprecipitation). The amount of Triton X-100 in the lysis buffer is normally sufficient for effective dissociation of SDS from proteins. If poor results are encountered with SDS-containing samples, try decreasing the final SDS concentration and/or increasing the final Triton X-100 concentration in the sample. If no proteins are detected on the gel, check whether (1) the total protein load is appropriate for detection method used, (2) the orientation of electrical connections is wrong or electrical connection during isoelectrofocusing is poor (all gels from that run will be blank), (3) an air bubble obstructs current in a single IEF tube, or (4) the electrical connection is incorrect or is poor during the second-dimension gel separation. Careful monitoring of current and voltage at the beginning, during, and at the end of electrophoretic separations is strongly recommended. Recording the initial and final current and voltage will also facilitate troubleshooting. Additional guidelines for troubleshooting and evaluating artifacts in two-dimensional gel electrophoresis are described by Dunbar (1987).
Anticipated Results A two-dimensional electrophoretic separation of proteins should produce a pattern of round or elliptical spots separated from one another.
Electrophoresis
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The pI range of the separated proteins as well as the observed molecular weight range depend on the first-dimension isoelectrofocusing protocol and the percentage of acrylamide used for the second-dimension gel. A complex protein mixture such as a whole-cell extract should produce more than 1000 silver-stained spots distributed over most of the gel area. Fewer spots will be detected with less sensitive detection techniques such as Coomassie blue staining. On the other hand, separation of radiolabeled proteins and use of multiple exposures permit detection of many low-abundance proteins.
Time Considerations Time requirements are very dependent on gel size and whether an external cooling unit is used to permit faster separations. Isoelectricfocusing using the standard-size gel format described in Basic Protocol 1 with 3-mm tubes is most conveniently done in an overnight run of ∼16 to 18 hr. This separation time can be decreased to ∼5 to 6 hr using higher voltages and an external cooling device. Isoelectrofocusing of 18-cm-long IPG gels requires ∼16 to 18 hr. Extruding a set of sixteen IEF tube gels and freezing them in equilibration buffer takes 1 to 2 hr, including setup time. Preparing and running SDS gels is described in UNIT 10.1. It takes ∼30 min to thaw, equilibrate, and load two second-dimension gels. Overall, if standard-size gels are used without external cooling, it will take ∼3 working days before the results of two-dimensional electrophoresis are obtained. A single person can conveniently run about 16 two-dimensional gels in one week, depending on the amount of electrophoresis equipment available. The ratelimiting step in most laboratories is running the second-dimension gels because 16 soluble ampholyte IEF gels or 12 IPG gels can be focused in one run, but loading, running, and detecting results from 12 to 16 second-dimension gels requires substantial operator time and electrophoresis equipment.
Literature Cited Anderson, N.G. and Anderson, N.L. 1978a. Two-dimensional analysis of serum and tissue proteins: Multiple isoelectrofocusing. Anal. Biochem. 85:331-340.
Celis, J.E., Rasmussen, H.H., Leffers, H., Madsen, P., Honore, B., Gesser, B., Dejgaard, K., and Vandekerckhove, J. 1991. Human cellular protein patterns and their link to genome DNA sequence data: Usefulness of two-dimensional gel electrophoresis and microsequencing. FASEB J. 5:2200-2208. Dunbar, B. 1987. Troubleshooting and artifacts in two-dimensional polyacrylamide gel electrophoresis. In Two-Dimensional Electrophoresis and Immunological Techniques (B.S. Dunbar, ed.) pp. 173-195. Plenum, New York. Garrels, J.I. 1979. Two-dimensional gel electrophoresis and computer analysis of proteins synthesized by cloned cell lines. J. Biol. Chem. 54:7961-7977. Garrels, J.I. 1989. The QUEST system for quantitative analysis of two-dimensional gels. J. Biol. Chem. 264:5269-5282. Garrels, J.I. and Franza, Jr., B.R. 1989. The REF52 protein database: Methods of database construction and analysis using the QUEST system and characterizations of protein patterns from proliferating and quiescent REF52 cells. J. Biol. Chem. 264:5283-5298. Goverman, J. and Lewis, K. 1991. Separation of disulfide-bonded polypeptides using two-dimensional diagonal gel electrophoresis. Methods 3:125-127. Hochstrasser, D.F., Harrington, M.C., Hochstrasser, A.C., Miller, M.J., and Merril, C.R. 1988. Methods for increasing the resolution of two-dimensional protein electrophoresis. Anal. Biochem. 173:424-435. O’Farrell, P.H. 1975. High resolution two-dimensional electrophoresis of proteins. J. Biol. Chem. 250:4007-4021. Strahler, J.R. and Hanash, S.M. 1991. Immobilized pH gradients: Analytical and preparative use. Methods 3:109-114. Young, D.A., Voris, B.P., Maytin, E.V., and Colbert, R.A. 1983. Very high resolution two-dimensional electrophoretic separation of proteins on giant gels. Methods Enzymol. 91:190-214.
Key Reference Hochstrasser et al., 1988. See above. Discusses methods for improving and troubleshooting two-dimensional separation.
Contributed by Sandra Harper, Jacek Mozdzanowski, and David Speicher The Wistar Institute Philadelphia, Pennsylvania
Anderson, N.L. and Anderson, N.G. 1978b. Two-dimensional analysis of serum and tissue proteins: Multiple gradient-slab gel electrophoresis. Anal. Biochem. 85:341-354. Two-Dimensional Gel Electrophoresis
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Protein Detection in Gels Using Fixation
UNIT 10.5
Detection of proteins in polyacrylamide gels is essential for purification and analysis of proteins. In addition, 1-D and 2-D gels have become the preferred methods for microscale isolation of small amounts of proteins for identification using internal cleavage with proteases followed by mass spectrometry analysis methods (Chapter 16). This unit presents basic protocols for detection of proteins by Coomassie blue staining (see Basic Protocol 1), staining with SYPRO Ruby a preformulated, noncovalent fluorescent stain (see Basic Protocol 2), and silver staining (see Basic Protocol 3). All protocols involve fixing the protein within the gel. Alternate protocols for Basic Protocol 1 detail rapid colloidal Coomassie blue staining (Alternate Protocol 1), which is slightly less sensitive, and acid-based colloidal Coomassie blue staining (Alternate Protocol 2), which avoids the need for destaining. Basic Protocol 3 for silver staining is accompanied by three alternate protocols: nonammoniacal silver staining (Alternate Protocol 3), which uses more stable solutions; rapid silver staining (Alternate Protocol 4), which involves fixation with formaldehyde instead of glutaraldehyde; and the enhanced-background, two-stage method (Alternate Protocol 5), which employs fixation without cross-linking to enhance sensitivity and also includes an optional reduction step to reduce background staining. A Support Protocol describes methods to obtain images of gels for archiving and quantitative analyses. The protocols in this unit and UNIT 10.6 require proteins that have been previously separated by one- or two-dimensional electrophoresis (UNITS 10.1-10.4). Coomassie blue staining is relatively quick and inexpensive, whereas SYPRO Ruby and silver staining are considerably more expensive but between 10 and 100 times more sensitive. When subsequent removal of the protein from the gel is desired, detection methods without fixation should be used (UNIT 10.6). In recent years, the number of commercially available preformulated gel stains has steadily increased to the point where multiple variations of every common type of stain can be purchased. Commercial staining kits offer convenient alternatives to preparing gel-staining solutions in the laboratory. They generally save time, have defined shelf lives, and often provide greater quality control and reproducibility than stains formulated in research laboratories. In many cases, staining kits have been modified by the manufacturers to enhance specific features such as sensitivity, staining speed, and environmental compatibility. These modifications and differences in stain formulations are usually proprietary; therefore, the effects of specific commercial products on different proteins and gel systems may vary and cannot be readily predicted from product descriptions. For example, not all commercial colloidal Coomassie stains will have the same sensitivity, background, and ease of use. Hence, to identify the best staining kit for a specific purpose, it may be necessary to purchase and compare different products based on the application of interest. There are particularly wide ranges of choice for silver stain kits due to the relative complexity of the basic method and the alternative chemistries that can be used. One potential limitation of staining kits for some applications might be the “black box” nature of working with undefined proprietary stains. Another limitation may be their generally higher cost compared with staining solutions formulated on site, especially when only the costs of consumables are considered. However, preformulated stains can often be more economical when labor costs are considered as well. The most common types of preformulated gel stains are summarized in Table 10.5.1.
Electrophoresis Contributed by Lynn A. Echan and David W. Speicher Current Protocols in Protein Science (2002) 10.5.1-10.5.18 Copyright © 2002 by John Wiley & Sons, Inc.
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Table 10.5.1
Common Gel Staining Kits
Type of stain Coomassie stains Coomassie Brilliant blue (CBB) Colloidala
Environment-friendly Rapid stains
Silver stains Conventional
Mass-spectrometry compatible
Rapid stains
Fluorescent dyes SYPRO Rubya
SYPRO Orangea SYPRO Reda SYPRO Tangerinea
Features
Representative vendors
Rapid, simple one-step stain and destain; good protein-to-protein consistency; linear quantitation range of ∼2 orders of magnitude Rapid one-solution preparation; typically 5- to 10-fold more sensitive than conventional CBB; destain optional; minimal background
Amersham Pharmacia Biotech; Bio-Rad
∼100-fold more sensitive than CBB for most proteins; much greater protein-to-protein variation in staining; kits contain multiple components and also require some common reagents that are not supplied; linear quantitation range is ∼1 order of magnitude; band intensities and background can vary substantially with small differences in reaction times and temperatures Milder chemical conditions used to minimize protein modifications; often less sensitive than conventional silver stains; reduced interference with trypsin digestion and mass spectroscopy analysis Reduced number of steps and reaction times; total staining times ∼1 to 2 hr
Amersham Pharmacia Biotech, Bio-Rad, ICN Biomedicals, Invitrogen, Pierce, Sigma
Bio-Rad, Diversified Biotech, Geno Technologies, ICN Biomedicals, Invitrogen, Pierce, Roche, Sigma Nonhazardous; no alcohol/acid fixatives or destains; BioRad, Invitrogen, Pierce, easy disposal of reagents Roche Bands usually develop >1h, sensitivity is often Amresco, Diversified somewhat reduced Biotech, ESA Inc., Invitrogen, Life Technologies, Pierce
Rapid, single-solution stain; low protein-protein variability; mass-spectrometry compatible; linear range over 3 orders of magnitude; staining sensitivity is highly dependent upon method used to acquire fluorescent image (see Basic Protocol 2 and Support Protocol); can rival silver staining sensitivity (1 to 2 ng/band); compatible with 1- and 2-D SDS-PAGE and IEF gels; available only as preformulated stain Good sensitivity (4 to 8 ng/band); applicable to 1-D SDS-PAGE Good sensitivity (4 to 8 ng/band), applicable to 1-D SDS-PAGE Good sensitivity (4 to 8 ng/band), applicable to 1-D SDS-PAGE, blotting
Bio-Rad, Invitrogen
Amersham Pharmacia Biotech, Bio-Rad, Geno Technologies, ICN Biomedicals, Invitrogen, Pierce, Sigma Bio-Rad, Molecular Probes
Bio-Rad, Molecular Probes Bio-Rad, Molecular Probes Bio-Rad, Molecular Probes
aColloidal Coomassie and SYPRO stains typically consist of a single stock staining reagent provided by the manufacturer. Any additional fixative or destaining steps may require common laboratory reagents. Silver stain kits generally consist of multiple components which may also require supplemental reagents.
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COOMASSIE BLUE R-250 STAINING Coomassie brilliant blue R-250 binds nonspecifically to almost all proteins, which allows detection of protein bands in polyacrylamide gels. This stain is very popular because it is relatively rapid, simple, inexpensive, and stains most proteins with similar intensities. Its major limitation is that it is less sensitive that the other stains described in this unit. In this procedure, proteins separated on a polyacrylamide gel are simultaneously fixed and stained in the Coomassie blue staining solution, which turns the entire gel blue. Destaining eliminates the background while the protein bands retain the blue color. Gels can be photographed or dried to maintain a permanent record. Alternatively the “wet” gel can be sealed in a plastic bag with a small amount of destaining solution and stored for many months at 4°C.
BASIC PROTOCOL 1
Materials Coomassie blue R-250 staining solution (see recipe) Polyacrylamide gel containing protein of interest (UNITS 10.1-10.4) Destaining solution: 15% methanol/10% acetic acid (v/v; store up to 1 month at room temperature) 7% (v/v) acetic acid (optional) Plastic or glass container with tight-fitting lid of size appropriate for gel Platform shaker Protein-based destaining material (optional): white 100% wool yarn, cellulose sponge, or Whatman 3MM filter paper Resealable plastic bags Gel dryer (optional) NOTE: Only high-purity water such as Milli-Q-purified water or equivalent should be used throughout the protocol, unless otherwise specified. 1. Add ∼10 gel volumes Coomassie blue R-250 staining solution to a plastic or glass container with lid. 2. Remove polyacrylamide gel from the electrophoresis unit and place in staining solution, then place container on a platform shaker and agitate gel slowly to prevent it from adhering to the container. CAUTION: Rapid agitation can result in gel breakage. The appropriate length of time to stain a gel depends on the gel thickness as well as the percentage of acrylamide. In general, gels ≤1 mm thick require only 30 min to 1 hr of staining, whereas gels >1 mm thick require a minimum of 1 to 3 hr of staining. Also, the higher the percentage of acrylamide, the longer it will take to stain the gel. Gels can be left in staining solution overnight without adverse effects.
3. Remove staining solution and rinse the gel briefly with deionized water to remove excess stain. Do not reuse staining solution.
4. Add 5 to 10 gel volumes destaining solution and place the container with gel on shaker. For more efficient destaining and to reduce waste, a protein-based destaining material such as a piece of wool yarn (6 to 12 in. long), a cellulose sponge (1 in. × 1 in.), or Whatman 3MM filter paper (1 in. × 1 in.) can be added to a minimal amount of destaining solution to bind the dye as it is released from the gel. Alternatively, the destaining solution can be changed several times until the gel shows a clear background with blue protein bands. Electrophoresis
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CAUTION: Overdestaining can occur if the gel is left in destaining solution for a prolonged period (>24 hr) or if the dye-binding material is left with the gel in the destaining solution for more than 2 to 3 hr.
5. For storage, place gel in a thermally sealed or resealable plastic bag with 1 to 2 ml destaining solution or 7% acetic acid. Gels can be stored sealed in plastic bags for several weeks at room temperature or for many months at 4°C.
6. For a permanent record, scan or photograph the gel (see Support Protocol), or dry the gel as follows. Place a wet piece of Whatman 3MM filter paper on a gel dryer and position the gel (rinsed with water) on top of the filter paper. Cover gel with a piece of plastic wrap and then add a second piece of wet filter paper. Dry 1 to 2 hr at ∼80°C or according to gel dryer instructions. As an alternative gel drying method, use a porous membrane to sandwich the gel in a supporting frame. In this case, dry the gel on the countertop at room temperature overnight. ALTERNATE PROTOCOL 1
RAPID COOMASSIE BLUE G-250 STAINING This protocol utilizes Coomassie brilliant blue G-250 in place of the Coomassie brilliant blue R-250 used in Basic Protocol 1. Also called xylene brilliant cyanin G, Coomassie brilliant blue G-250 (greenish hue) usually stains with a slightly higher sensitivity than Coomassie brilliant blue R-250 (reddish hue). The G-250 stain produces a somewhat brighter blue coloration than the deep blue of the R-250 stain. The G-250 dye is less soluble than R-250 and is used in a colloidal state rather than as a solution. Due to its lower solubility, G-250 preferentially binds to the protein bands and not to the polyacrylamide gel itself; therefore, little or no destaining is necessary. The procedure requires fixation of gels prior to staining. Protein staining then develops rapidly and can be observed easily throughout the staining solution. This procedure provides faster detection of proteins in gels than Basic Protocol 1. Additional Materials (also see Basic Protocol 1) Isopropanol fixing solution: 25% isopropanol/10% acetic acid (v/v; store several months at room temperature) Rapid Coomassie blue G-250 staining solution: 10% (v/v) acetic acid/0.006% (w/v) Coomassie brilliant blue G-250 (Bio-Rad; store several months at room temperature) 10% (v/v) acetic acid 1. To a plastic container with lid add ∼10 gel volumes isopropanol fixing solution. Remove gel from the electrophoresis unit and place in fixing solution. Agitate container on a platform rocker or shaker for 10 to 15 min for 1-mm-thick gel or 30 to 60 min for 1.5-mm-thick gel. 2. Discard fixing solution and add 5 to 10 gel volumes rapid Coomassie blue G-250 staining solution. Agitate 30 min to 1 hr for 1-mm-thick gel or 2 to 3 hr for 1.5-mm-thick gel. Staining of protein bands becomes apparent within 5 to 15 min.
3. Remove staining solution after appropriate time. Add 5 to 10 gel volumes of 10% acetic acid and agitate slowly. Destain (1 to 2 hr) until a clear background with blue protein bands appears. Protein Detection in Gels Using Fixation
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4. For storage, place gel in a resealable plastic bag at 4°C with a small amount of 10% acetic acid. 5. To scan or photograph gel, see Support Protocol. To dry gel, see Basic Protocol 1 (step 6). ACID-BASED COOMASSIE BLUE G-250 STAINING The acid-based Coomassie brilliant blue G-250 staining procedure fixes and stains the gels simultaneously and, unlike the first basic and alternate protocols, does not require a destaining solution. Color intensifies when the gels are placed in water after the staining period, and as little as 50 to 100 ng protein per band can be detected. Acid-based Coomassie blue G-250 which is also used in a colloidal state rather than a solution, staining has a sensitivity similar to that of the basic Coomassie blue R-250 staining. More heavily loaded protein bands can be detected within 30 min, but ∼5 hr is needed to visualize light bands.
ALTERNATE PROTOCOL 2
Additional Materials (also see Basic Protocol 1) Acid-based Coomassie blue G-250 staining solution (see recipe) 1. Remove polyacrylamide gel from the electrophoresis unit. If gel contains SDS, incubate in 5 to 10 gel volumes deionized water for a minimum of 5 to 15 min depending on gel thickness to allow SDS to diffuse out. SDS will precipitate in the staining solution and produce an opaque background.
2. Place gel into a container with ∼100 ml acid-based Coomassie blue G-250 staining solution. Place gel on a platform shaker and agitate gently for 5 hr to overnight. Heavily loaded proteins will be visible within 30 min. However, greatest sensitivity is achieved by staining for at least 5 hr.
3. Decant staining solution and replace with deionized water. Even after the gel has been stained overnight, protein bands will not be completely resolved until the gel is placed into deionized water. Bands will appear bright blue against a clear background within 2 to 5 min.
4. For storage, place gel in 1 to 2 ml water in a resealable plastic bag at 4°C. 5. To scan or photograph gel, see Support Protocol. To dry gel, see Basic Protocol 1 (step 6). SYPRO RUBY STAINING There are several commercially available fluorophores which interact with proteins in polyacrylamide gels through noncovalent mechanisms similar to Coomassie stains and that are more sensitive than Coomassie brilliant blue or colloidal Coomassie stains. These fluorescent stains have wider dynamic ranges than colorimetric stains. The most sensitive of these dyes, SYPRO Ruby, can be visualized using a standard 302-nm UV transilluminator or imaging system containing the appropriate excitation and emission filters (see Support Protocol). The maximal excitation and emission wavelengths are 450 and 610 nm, respectively. SYPRO Ruby does not interfere with common downstream analysis methods such as mass spectrometry, Edman sequencing, or immunodetection procedures. Depending upon the imaging method used, the sensitivity of this stain can be comparable to most silver staining methods. The protocol for SYPRO Ruby staining described below
BASIC PROTOCOL 2
Electrophoresis
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has been adapted from the method described by the manufacturer of the fluorophore (Molecular Probes). Materials SDS 1- or 2-D polyacrylamide gels containing proteins of interest (UNITS 10.1-10.4) Fixing solution: 40% (v/v) methanol/10% (v/v) acetic acid in H2O SYPRO Ruby protein gel stain (Molecular Probes) Destain/wash solution: 2% (v/v) acetic acid 2% (w/v) glycerol Plastic container with tight-fitting lid, of size appropriate for gel Platform shaker Additional reagents and equipment for acquiring gel images (see Support Protocol) 1. Remove polyacrylamide gel from the electrophoresis unit and place in a plastic container containing at least 10 gel volumes of 40% methanol/10% acetic acid. Cover with a tight-fitting lid and fix the gel for 1 hr on a platform shaker with continuous, gentle agitation. Plastic containers are optimal because they absorb a minimal amount of the dye.
2. Remove staining solution and incubate gel in 10 gel volumes of undiluted SYPRO Ruby stain for 3 hr to overnight with gentle agitation, protected from light. All reagents should be carefully removed by vacuum aspiration. The gel should be protected from light from this step forward, until an image has been obtained for archiving. The manufacturer does not recommend using less than 10 gel volumes of stain or reusing the stain because reduced staining sensitivity can result. However, if at least 10 gel volumes of stain are used and the stain is protected from light, the reduction in sensitivity is minor when the stain, which is quite expensive, is reused once. However,with further use of the stain, staining intensity will be more dramatically reduced.
3. Wash the gel by immersing in 10 gel volumes of 2% acetic acid for 30 min to reduce background fluorescence and increase sensitivity. For convenience, the gel may be washed overnight without a loss in sensitivity when 2% acetic acid is used as the destain solution. If 10% methanol (or ethanol) is used, background fluorescence reduction is more rapid, but protein band intensities will begin to fade appreciably after 30 min.
4. Acquire an image of the gel for a permanent record (see Support Protocol). The stained gel can be stored for several days in 2% acetic acid in the dark at 4°C for at least a week with minimal reduction in signal intensity.
5. Optional: For longer-term storage, transfer the gel into a 2% glycerol solution, incubate for 30 min, then dry gel in a gel dryer. BASIC PROTOCOL 3
SILVER STAINING Silver staining is about 50 to 100 times more sensitive than Coomassie blue staining; however, staining responses can vary greatly from protein to protein. After alcohol/acid fixation and treatment with silver nitrate, the gel is developed until a desired staining intensity is reached. Because the absolute staining intensity is controlled by the length of reaction in the developer, and staining intensity is variable for different proteins, estimation of protein amounts using silver staining is not advisable unless known quantities of
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the protein of interest are included on the same gel to calibrate staining intensity. Reagent volumes, reaction times, and temperature should be carefully controlled, and the very narrow linear detection range must be considered when silver staining is used to compare protein patterns on 2-D gels. The following protocol is applicable to 1-D and 2-D SDS-PAGE, and is based on the ExPASy SWISS-2D PAGE method, which has been optimized to give uniformly brown spots on a light background to facilitate image acquisition and quantitative comparisons. However, this method is not suitable for protein detection using mass spectrometry because of the extensive chemical modification of proteins during the staining procedure, especially the chemical cross-linking of proteins by glutaraldehyde. Materials Polyacrylamide gel containing proteins of interest (see UNITS 10.1-10.4; also see recipe) Fixing solution A: 40% (v/v) ethanol/10% (v/v) acetic acid in H2O Fixing solution B: 5% (v/v) ethanol/5% (v/v) acetic acid in H2O 0.5 M sodium acetate containing 1% (v/v) glutaraldehyde 0.05% (w/v) 2,7-napthalene disulfonic acid Ammoniacal silver nitrate solution (see recipe) Citric acid developing solution (see recipe) Stop solution: 5% (w/v) Tris base containing 2% (v/v) acetic acid Glass or plastic container with tight-fitting lid, of size appropriate for gel Platform shaker Resealable plastic bags Additional reagents and equipment for acquiring gel images (see Support Protocol) CAUTION: Ammoniacal silver nitrate solution becomes explosive on drying. Discard immediately after use by adding 1 M HCl to precipitate the silver as silver chloride and dispose of as required by local regulations. Fix gels 1. After electrophoresis (UNITS 10.1-10.4), place gels in a glass or plastic container and wash in deionized water for 5 min on a platform shaker with continuous, gentle agitation. Since silver nitrate reacts with skin proteins, wear gloves that have been thoroughly rinsed with deionized water when handling all materials. All reagents should be removed from staining trays using vacuum aspiration to avoid any contact between skin or gloves and gels, as any physical contact with the gels can create staining artifacts. IMPORTANT NOTE: All subsequent incubations and washes are performed with continuous gentle agitation on a platform shaker as described above.
2. Remove water and add ∼10 to 20 gel volumes fixing solution A. Fix gel by incubating for 1 hr in this solution. 3. Remove fixing solution A and add ∼10 to 20 gel volumes fixing solution B. Fix gel by incubating for 2 hr to overnight in this solution. 4. Remove fixing solution B and wash with ∼10 to 20 gel volumes deionized water for 5 min. 5. Remove water and incubate gel in ∼5 to 10 gel volumes 0.5 M sodium acetate containing 1% glutaraldehyde for 30 min. Electrophoresis
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CAUTION: Glutaraldehyde is an irritant. Wear gloves and prepare solution in a fume hood.
6. Carefully remove glutaraldehyde solution and wash gel with three changes of ∼10 to 20 gel volumes deionized water, each time for 10 min. 7. Remove water and incubate gel in ∼5 to 10 gel volumes in two changes of 0.5% (w/v) 2,7-napthalene disulfonic acid, each time for 30 min. The solution in these steps provides homogeneous dark brown staining of proteins.
8. Remove the 2,7-napthalene disulfonic acid solution and rinse gel with four changes of ∼10 to 20 gel volumes deionized water, each time for 15 min. Stain gels 9. Remove water and stain gel with ∼5 to 10 gel volumes of freshly prepared ammoniacal silver nitrate solution for 30 min. 10. After staining, wash the gel with four changes of ∼10 to 20 gel volumes deionized water, each time for 4 min. 11. Remove water and develop gels in ∼5 to 10 gel volumes citric acid developing solution for 5 to 10 min. Develop gels until a slight background stain appears and bands are a homogeneous brown color; be careful not to overdevelop. As noted above, all gels that will be used for quantitative comparisons should be processed and developed using identical conditions.
12. When gels are fully developed, quickly remove citric acid solution and stop development by incubating for 30 min in ∼10 to 20 gel volumes stop solution. 13. Remove stop solution and rinse gel with ∼10 to 20 gel volumes deionized water for at least 30 min. Record and store gels 14. Scan gels using a flatbed scanner (see Support Protocol). Gels should be scanned promptly after development, i.e., within about 24 hr, because the stained spots may fade with time of storage.
15. Store gels in resealable plastic bags at 4°C after scanning. ALTERNATE PROTOCOL 3
NONAMMONIACAL SILVER STAINING The nonammoniacal silver staining procedure is similar to Basic Protocol 3 except that more stable solutions are used for staining. Using this procedure, some proteins may be detected that are not recognized by other silver staining methods. Materials Fixing solution A: 50% (v/v) methanol/10% (v/v) acetic acid (store up to 1 month at room temperature) Fixing solution B: 5% (v/v) methanol/7% (v/v) acetic acid (store up to 1 month at room temperature) 10% (w/v) glutaraldehyde 5 µg/ml dithiothreitol (DTT; make fresh daily) 0.1% (w/v) silver nitrate Carbonate developing solution (see recipe) 2.3 M citric acid
Protein Detection in Gels Using Fixation
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0.03% (w/v) sodium carbonate Plastic or glass container with tight-fitting lid, of size appropriate for gel Platform shaker Fix and stain proteins 1. Remove gel from electrophoresis apparatus and place in a plastic or glass container equipped with a lid. Add 5 to 10 gel volumes fixing solution A. Agitate for 30 min on a platform shaker. 2. Decant fixing solution A, and add 5 to 10 gel volumes fixing solution B. Agitate for 30 min. 3. Remove fixing solution B, add 50 ml of 10% glutaraldehyde, and agitate for 10 min in a fume hood. CAUTION: Glutaraldehyde is an irritant. Wear gloves and work only in a fume hood.
4. Decant glutaraldehyde solution. Add 5 to 10 gel volumes water and agitate for 15 min. Repeat eight times. Alternatively, wash gel overnight in 30 to 40 gel volumes water with slow agitation. The next day add fresh water and agitate for 30 min to ensure that all traces of glutaraldehyde are removed so a clear background is attained.
5. Decant water and add 5 gel volumes of 5 µg/ml DTT. Agitate 30 min. 6. Pour off DTT and, without rinsing, add 5 gel volumes of 0.1% silver nitrate. Agitate 30 min. Develop stain and process gel 7. Decant silver nitrate solution. Rinse gel once for 15 sec with water by filling the container with water and then pouring it out. Rinse gel quickly two times with a few milliliters of carbonate developing solution. Finally, add 100 ml carbonate developing solution and agitate until bands appear as desired. 8. When proper staining has been attained, add 5 ml of 2.3 M citric acid directly to the carbonate developing solution to stop the staining reaction. Agitate for 10 min, replace the solution with water, and agitate for 30 min. Repeat four more times. Always use a 1:20 (v/v) ratio of 2.3 M citric acid to carbonate developing solution to neutralize the solution, thus stopping the reaction. If the pH is too high, color development will continue; if the pH is too low, the color will bleach away.
9. To store gel, soak the gel in 5 to 10 gel volumes of 0.03% sodium carbonate for ∼1 hr with agitation to prevent bleaching, then place in a resealable plastic bag. Store gel at 4°C. 10. To scan or photograph gel, see Support Protocol. To dry gel, see Basic Protocol 1 (step 6). RAPID SILVER STAINING Unlike the previous two silver staining procedures (see Basic Protocol 3 and Alternate Protocol 3) which use glutaraldehyde as a fixative, rapid silver staining employs formaldehyde to fix the gel. The procedure has a sensitivity similar to that of the previous protocols. Also, the developing solution contains thiosulfate to inhibit background stain-
ALTERNATE PROTOCOL 4
Electrophoresis
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ing by dissolving unspecific surface staining. This method is rapid but may not be quite as sensitive as other methods, especially for small proteins. Materials 50% methanol/12% acetic acid (v/v) 95% ethanol (optional) Formaldehyde fixing solution (see recipe) 0.02% (w/v) sodium thiosulfate 0.1% (w/v) silver nitrate Thiosulfate developing solution (see recipe) 50% methanol Plastic or glass container with tight-fitting lid, of size appropriate for gel Platform shaker Fix and stain proteins 1. To prevent protein diffusion and improve background, fix gel after electrophoresis, if desired, for a minimum of 1 hr to overnight in 50% methanol/12% acetic acid. Wash gel with 95% ethanol three times, 20 min each time, before proceeding with step 2. 2. Add 3 gel volumes formaldehyde fixing solution. Place container on a platform shaker, then agitate 15 min for a 1-mm-thick gel or 30 min for a 1.5-mm-thick gel. 3. Decant fixing solution, then wash gel by adding 5 to 10 gel volumes water and agitating for 5 min. Repeat two times. 4. Remove water and add 3 gel volumes of 0.02% sodium thiosulfate. Agitate for 1 min. 5. Decant sodium thiosulfate and add 5 to 10 gel volumes water. Agitate for 1 min. Repeat two times. 6. Decant water and add 3 gel volumes of 0.1% silver nitrate. Agitate for 10 min. Develop stain and process gel 7. Remove silver nitrate, add 5 to 10 gel volumes water, and agitate for 1 min. Repeat two times. 8. Decant water. Add a few milliliters of thiosulfate developing solution until a brown precipitate forms (30 sec), decant, and add 3 gel volumes fresh thiosulfate developing solution. 9. Just before bands reach desired intensity, wash the gel quickly with water to remove excess thiosulfate (to avoid background staining). Stop the reaction by adding 50% methanol/12% acetic acid and agitate for 10 min. 10. Wash the gel with 5 to 10 gel volumes of 50% methanol with agitation for ∼30 min to remove acetic acid. Repeat one time. The methanol washes are important to prevent darkening of protein bands over a prolonged period.
11. Store gel in a resealable plastic bag at 4°C. 12. For a permanent record, scan or photograph gel (see Support Protocol) or dry gel (see Basic Protocol 1, step 6). Protein Detection in Gels Using Fixation
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ENHANCED-BACKGROUND (TWO-STAGE) RAPID SILVER STAINING The enhanced-background rapid silver staining method is a two-stage staining procedure that enhances the sensitivity of most proteins and decreases background staining. In this method, however, proteins are not chemically cross-linked by either glutaraldehyde or formaldehyde; hence some proteins, especially those of low molecular weight, may be selectively lost during staining. The first part of the procedure is a rapid silver staining method that is easier and quicker than both basic silver staining and nonammoniacal silver staining. The second part of the procedure, which is optional, improves sensitivity and lessens background staining of the gel.
ALTERNATE PROTOCOL 5
Materials Rapid fixative 1: 40% methanol/10% acetic acid (v/v; store 1 month at room temperature) Rapid fixative 2: 10% methanol/5% acetic acid (v/v; store 1 month at room temperature) 1× oxidizer (see recipe) 0.17% (w/v) silver nitrate Thiosulfate developing solution (see recipe) 5% (w/v) acetic acid Farmers reducer (see recipe) Plastic or glass container with tight-fitting lid, of size appropriate for gel Platform shaker Rapid Fixation Silver Staining Fix proteins 1. After electrophoresis, soak gel in 5 gel volumes rapid fixative 1 in a glass or plastic container for 60 min, agitating container on a platform shaker. 2. Pour off used rapid fixative 1 and add 5 gel volumes rapid fixative 2. Agitate for 30 min. 3. Pour off used rapid fixative 2 and add another 5 gel volumes fresh rapid fixative 2. Agitate for 30 min. Stain and develop gel 4. Pour off used rapid fixative 2 and add 100 ml of 1× oxidizer. Agitate for 10 min. 5. Pour off oxidizer, add 5 gel volumes water, and agitate for 15 min. Repeat four times. 6. Add 100 ml of 0.17% silver nitrate and agitate for 45 min. 7. Wash gel with running water for 2 min. 8. Add 1 gel volume thiosulfate developing solution. After a brown precipitate appears (∼30 sec), decant solution and add 5 gel volumes fresh thiosulfate developing solution. 9. After protein bands have reached the desired intensity, stop the reaction by pouring off developing solution and replacing with 5% acetic acid. 10. Pour off acetic acid, add 5 gel volumes water, and agitate 15 min. Repeat three times.
Electrophoresis
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11. To improve background and sensitivity, continue to step 12. Otherwise, store the gel at 4°C in a resealable plastic bag with 5% acetic acid. Alternatively, scan or photograph (see Support Protocol) or dry the gel according to step 6 in Basic Protocol 1. Improving Sensitivity (Optional) 12. Remove water, add 5 gel volumes Farmers reducer, and agitate for 10 min. After this step the gel should be transparent and yellow, and void of any silver-stained bands.
13. Remove reducer and wash gel by adding 5 gel volumes water and agitating for 15 min. Repeat wash five times. The gel must be colorless before continuing; therefore, it is preferable to do the last wash overnight.
14. Repeat steps 6 to 10. Store the gel in a sealed plastic bag at 4°C; scan or photograph (see Support Protocol), or dry the gel (see Basic Protocol 1, step 6) for a permanent record. SUPPORT PROTOCOL
GEL IMAGING Historically, gel images were obtained using Polaroid or 35-mm photography. These older techniques have been largely replaced by digital-imaging methods, which have become reasonably economical and facile. Digital images are essential for quantitative analyses of gel images; they also facilitate data archiving and transfer over the Web, and streamline illustration of data in presentations and manuscripts. The choice of imaging method depends on the types of imaging devices available, the quality of image desired (resolution and bit depth), and the type of stain used (colorimetric or fluorescent). The critical factors to consider in acquiring digital gel images are briefly summarized here. For a more detailed discussion of digital imaging, see UNIT 10.12. Resolution, Bit Depth, and File Size Most instruments used for digitizing gel images offer multiple options for spatial resolution and bit depth. Using the highest resolution and bit depth available is not always the best option, primarily because file sizes can become very large, which not only results in consumption of large amounts of disk space but can slow performance of subsequent analysis programs. Hence, when setting image acquisition parameters, the requirements of downstream analysis software should be considered.
Protein Detection in Gels Using Fixation
The required spatial resolution of the acquired image will be influenced by the purpose of the resulting images. Applications with low to moderate resolution requirements include preparation of laboratory notebook prints, small manuscript figures, and PowerPoint figures. Higher resolution is generally required when software programs will be used to identify and quantitate bands and spots on 1-D and 2-D gels. For these applications, sufficient detail must be captured so that the smallest and highest-resolution features are preserved and do not limit the capacity of the quantitation software. In some cases, spatial resolution may be influenced by software requirements or limitations of the imaging system used. However, as a general guide, 2-D gel images should be acquired using a spatial resolution that yields at least 15 to 20 data points (pixels) for the smallest spot, and the sharpest band on a 1-D gel should contain at least 6 to 10 pixels of data across the band. Depending upon the imaging device, resolution will be specified in either
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micrometers (µm) or dots per inch (dpi). For most purposes, images of minigels at 50 or 100 µm or 300 to 600 dpi (∼85 µm to 42 µm) are adequate. Signal intensity resolution is dependent upon the resolution capacity of the detection device and the bit depth, or the number of bits used to report the brightness of a particular spot on an image (Goforth, 2001). Bit depth defines the maximum quantitative resolution of signal intensity or number of grayscale units. The most common options are 8-, 12-, or 16-bit resolution. The major advantage of 8-bit resolution, which provides 256 grayscale units, is small file size. This format was compatible with older imaging systems and software but provides very limited quantitative dynamic range. It is adequate for photographic-type prints and computer-generated displays, but is not optimal for quantitative analyses. Some current imaging systems and software use 12-bit data with 4096 grayscale levels; however, due to the availability of faster computers and economical data storage devices, 16-bit data with 65,536 gray scale units is rapidly becoming the most common, preferred format. The most common digital imaging devices include flatbed scanners, cooled CCD cameras, CCD cameras, and laser scanners. Brief summaries of each imaging source are given below. It is important to note that, for all of these methods, all gels must be destained properly for best results; otherwise signal-to-noise will not be optimal. Flatbed Scanners A simple, inexpensive, effective method of acquiring digital images of Coomassie, colloidal Coomassie, or silver-stained gels is to use a flatbed scanner in transparency rather than reflective mode. Data quality will be substantially compromised if images are acquired in reflective mode. Scanning resolution is typically specified in dpi rather than micrometers. Scanners typically offer greater uniformity of illumination and provide more reliable density analyses (Miura, 2001). Charged-Coupled Device (CCD) Cameras CCD cameras usually offer illumination sources for both colorimetric and fluorescent images. These cameras provide high resolution and signal intensity by dissecting an image into many small picture elements, or pixels, which are stored on a CCD chip and later converted to a video output (Miller et al., 2001). Most imaging devices offer a range of 50-, 100-, or 250-µm resolution. Inexpensive units do not use cooling of the CCD chip, often have manual adjustments for focus and image size, and may have simple software that does not adjust for nonuniform light source intensity and camera lens distortion. Such cameras are useful for archiving visual images but will generally have limited utility for quantitative comparisons. Cooled CCD cameras use reduced temperatures (e.g., −30°C) to reduce background noise, thereby allowing longer exposures with associated increased sensitivity for fluorescent and chemiluminescent images. These cameras often have fixed focal distances and image sizes. Objects that are larger than the acquisition size of the camera are imaged sequentially and then combined into a composite image using software. Software is also used to correct for light-intensity differences. Another feature of these cameras is that they usually acquire images at the maximum possible resolution, and, when reduced image resolution is selected, data points are averaged. Hence, a cooled CCD camera with 50-µm resolution will average 4 data points on the 100-µm setting, which will improve signal-to-noise and reduce file size by four-fold. Therefore, a lower resolution setting can be advantageous for some imaging applications with these devices, provided that spatial resolution is still adequate for analysis software (see above). Cooled Electrophoresis
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CCD cameras with appropriate excitation and emission filters will typically produce the greatest-sensitivity images when using fluorescent dyes such as SYPRO Ruby (see Basic Protocol 2). Laser Scanners An alternative to using a CCD camera for fluorescence imaging is to use a laser scanner. Laser-scanner-based systems pass an illumination beam over each point in a sample in a 2-dimensional fashion to excite the specific fluorophore (Patton, 2000). When imaging these gels, sensitivity is determined by the signal-to-noise ratio, and sensitivity can only be increased by increasing the sensitivity of the photomultiplier tube, provided the background is low (Yan, 2000). Therefore, it is essential to adequately stain the sample to achieve maximum sensitivity, as well as to properly wash the gel to minimize background fluorescence. Excitation wavelengths are constrained by the availability of a small number of available laser wavelengths; therefore it may not be possible to excite some fluorophores at optimal wavelengths. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
NOTE: Only high-purity water such as Milli-Q-purified water or equivalent should be used to prepare all solutions, unless otherwise specified. Acid-based Coomassie blue G-250 staining solution Prepare 0.2% (w/v) Coomassie brilliant blue G-250 (Bio-Rad) in water. Add an equal volume of 2 N H2SO4. Mix well and let stand for 3 hr minimum. Filter through Whatman no. 1 filter paper. Measure recovered volume and add 1/9 vol of 10 N KOH. Finally, add 100% (w/v) trichloroacetic acid (TCA) to a final concentration of 12% (w/v). The final concentration of Coomassie blue is ∼0.08% (w/v). Store up to several months but maintain solution below pH 1.0. Ammoniacal silver nitrate solution Stock solution I: Dissolve 2 g of silver nitrate in 10 ml of water (prepare fresh). Stock solution II: Mix 53 ml of deionized water, 2.8 ml of 30% ammonium hydroxide, and 0.265 ml of 50% sodium hydroxide (v/v), and stir vigorously. Prepare fresh. CAUTION: Wear protective glasses and take other appropriate precautions for strongly alkaline solutions (also see APPENDIX 2A) when handling stock solution II.
Working solution: Slowly pipet stock solution I into solution II while the latter solution is being vigorously stirred. A transient brown precipitate will form briefly; after it clears, add deionized water to bring the total volume 250 ml. Carbonate developing solution 0.5 ml 37% (v/v) formaldehyde 30 g sodium carbonate (3%, w/v) H2O to 1 liter Make fresh each time
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Citric acid developing solution 0.05 g citric acid monohydrate (0.01% w/v final) 0.5 ml formaldehyde (0.1% w/v final) H2O to 500 ml Make fresh each time. Coomassie blue R-250 staining solution Concentrated stock: Mix 24 g Coomassie brilliant blue R-250 (Bio-Rad or Pierce) and 600 ml methanol. Add 120 ml acetic acid. Stir for 2 hr minimum, and preferably overnight. Store up to several months at room temperature. Staining solution: Mix 1 liter methanol and 60 ml Coomassie blue R-250 concentrated stock. Add 800 ml H2O, then 200 ml concentrated acetic acid. If precipitation occurs, filter the solution through Whatman no. 1 filter paper prior to use. Store up to several months at room temperature. The staining solution has approximate final concentrations of 50% (v/v) methanol, 0.1% (w/v) Coomassie brilliant blue R-250, and 10% (v/v) acetic acid.
Farmers reducer 5 g potassium ferricyanide 8 g sodium thiosulfate H2O to 500 ml Make fresh every time Formaldehyde fixing solution 400 ml methanol 0.5 ml 37% (v/v) formaldehyde H2O to 1 liter Store ≤1 month at room temperature Oxidizer, 10× and 1× 10× stock: 5 g potassium dichromate 1 ml 70% (w/v) nitric acid H2O to 500 ml Store up to several months at 4°C 1× working solution: Dilute 10× oxidizer 1/10 with H2O Make fresh each time Polyacrylamide gel (for use in Basic Protocol 3) The basic silver staining protocol (see Basic Protocol 3) involves casting the polyacrylamide gels using piperazine-diacrylyl (PDA) as the cross-linker to provide a gel that is more stable to the alkaline pH of the ammoniacal silver nitrate solution. These gels are also cast containing 5 mM sodium thiosulfate to reduce gel background. See UNITS 10.1-10.4 for casting of PAGE gels; see http://www.expasy.com for these modifications. Thiosulfate developing solution 15 g sodium carbonate 0.5 ml 37% (w/v) formaldehyde 0.2 ml 1% (w/v) sodium thiosulfate H2O to 500 ml Make fresh each time Electrophoresis
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COMMENTARY Background Information Coomassie blue and silver staining are the two most common approaches used for protein detection in gels with fixation. Silver staining relies on binding of silver to amino acid side chains, primarily sulfhydryl and carboxyl groups, and Coomassie blue binds nonspecifically to proteins. Silver staining is typically about 10 to 100 times more sensitive than Coomassie blue staining but it is more complex and time-consuming. There are many variations of silver stain methods both in the scientific literature and in the form of commercial kits (see Table 10.5.1). Recently, fluorescent stains for gels, such as the SYPRO series have been introduced (Patton, 2000). These stains are available only as preformulated solutions and the most sensitive stain, SYPRO Ruby (see Basic Protocol 2), can be as sensitive as some silver stain methods when high-performance laser scanners or cooled CCD cameras are used to acquire the images. This staining method is much simpler than silver staining and uses less harsh conditions, which can be an advantage when gel bands or spots will be used to identify proteins using mass spectrometry methods. Another advantage of the SYPRO fluorescence stains is that they typically have about a threeorder-of-magnitude dynamic range of detection, which is much wider than the dynamic ranges for the colorimetric stains. The Basic and Alternate Protocols described for Coomassie blue staining differ most significantly in time required. The rapid Coomassie blue G-250 staining procedure (Alternate Protocol 1) uses isopropanol for fixing instead of methanol, and the colloidal stain produces a low background, which allows for faster protein staining (bands appear within 15 min); however, it is not as sensitive as Basic Protocol 1 involving Coomassie blue R-250 (Weber et al., 1972). The acid-based Coomassie blue staining procedure (Blakesley and Boezi, 1977) also uses colloidal Coomassie blue G-250, but it does not involve a separate fixation step (Alternate Protocol 2). Acid-based staining achieves a sensitivity similar to that of Coomassie R-250, although for maximum sensitivity gels must be stained 5 hr to overnight. Basic Protocol 2 describes an optimized protocol for SYPRO Ruby staining of 1-D and 2-D gels (IEF gels can also be stained with minor modifications) that largely follows the
supplier’s recommendations. However, several versions of the staining protocol have been circulated. The impact of these variations has not been well defined and may be at least partially dependent upon the types of gels used. Basic Protocol 3 for silver staining (http://www.expasy.com) is a more sensitive protein detection method compared to the nonammoniacal and rapid two-stage alternate silver staining protocols; however, the procedure uses unstable ingredients and tends to require more expensive reagents. Nonammoniacal silver staining (Alternate Protocol 3) is based on a method described by Morrissey (1981). The two-stage silver staining procedure (Alternate Protocol 5) is a modification of the method described by Gorg (1985) and consists of two parts: the first stage, involving rapid silver staining, is followed by an optional series of steps to reduce background. The rapid silver staining procedure (Alternate Protocol 4; Blum et al., 1987) is a quick and easy way to detect proteins within 50 min and, like the alternate two-stage silver staining method, uses thiosulfate to reduce background impurities.
Critical Parameters and Troubleshooting All solutions used in preparing the polyacrylamide gels as well as the staining solutions must contain high-purity reagents. In addition, gel solutions should be filtered to avoid artifacts in the gel. This is especially important in silver staining, which is based on a photochemical process where impurities can increase background and cause artifactual staining. Powderfree gloves should be used at all times, and care should be taken to avoid any physical contact with the gels, using vacuum aspiration, rather than pouring, to remove staining reagents. One common contaminating protein is skin keratin, which shows up as multiple bands with molecular weights between 55,000 and 60,000, especially when high-sensitivity stains are used. If such bands appear in blank lanes where no protein was loaded, most likely the stacking gel, solubilizing buffer, and/or electrode buffer are contaminated. The problem can be eliminated by avoiding contact of skin and hair with gel and buffer solutions. There can be great variations in protein and background staining with different silver staining methods or even with seemingly minor
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modifications of a single protocol. In addition, these protocols are very sensitive to exact reaction conditions for critical steps, including reaction times and temperatures. In general, the most critical steps are the wash step(s) to remove excess silver reagent, the development time, effectiveness of the step that stops development, and whether or not sodium thiosulfate is added to the gel during casting (see Basic Protocol 3). Addition of sodium thiosulfate to the gel typically improves the level of background staining, but increases development times. Because SYPRO stains are relatively new, the impact of variations in gel fixation and destaining are not yet well defined. As noted above, the instructions from the manufacturer recommend a number of alternative fixatives and a relatively wide range of staining times; the manufacturer has also recommended several different destaining solutions that can yield quite different results. Hence it is advisable to explore the impact of staining/destaining conditions if maximum sensitivity and consistent staining results are desired, since different gel types, especially with regard to gel porosity and gel size, might not have the same optimal staining conditions. The method described here (see Basic Protocol 2) is based on preliminary comparisons of different methods of fixation, staining, and destaining of minigels. These results suggest that the fixative is not particularly critical but stronger fixatives such as 40% methanol/10% acetic acid might be advantageous, especially if the gel will be archived for potential future uses such as protein identifications. At least with some gel systems, staining for 3 hr is inadequate for maximal staining, and overnight staining yields much stronger signals. The destaining conditions also appear to be quite critical for achieving maximum sensitivity. If the manufacturer-recommended destain solution of 10% methanol or ethanol is used, the background fluorescence decreases rapidly and a maximum signal-to-noise ratio is obtained within the recommended 30 min. But incubation of the gel in these solutions for longer than 30 min can markedly decrease band intensities, with complete loss of some bands after overnight incubation. When water or 2% acetic acid is used as the destain solution, the background continues to decrease for a number of hours and optimal signal-to-noise ratio is obtained after overnight destaining. While this
method is slower than a dilute alcohol solution, overdestaining of bands does not occur.
Time Considerations In general, Coomassie blue and SYPRO Ruby staining tends to be easier and faster than silver staining. Basic Coomassie staining of standard-sized gels (11 × 16 cm) that are 1.5 mm thick require at least 2 to 3 hr of staining followed by destaining for at least several hours to overnight. Minigels require less time, ∼30 min for staining and about the same time for destaining. The rapid colloidal Coomassie blue staining procedures (Alternate Protocol 1) allow visualization of protein bands within 5 to 15 min. SYPRO Ruby staining requires an hour of fixation followed by overnight incubation with the stain, and minimal (30 min) destain. Silver staining, on the other hand, is very timeconsuming because the gels must be treated with many different solutions. Also, gels require successive washes with water to remove glutaraldehyde for good background in the silver staining and nonammoniacal silver staining methods (Basic Protocol 3 and Alternate Protocol 3), and to remove Farmers reducer in the second part of the two-stage silver staining procedure (Alternate Protocol 5). Rapid silver staining (Alternate Protocol 4), however, is the easiest and the most time-efficient of all the silver staining methods as it takes only ∼50 min. In general, using a pre-manufactured gel stain will significantly cut down on solution preparation time, and will provide highly consistent results.
Literature Cited Blakesley, R.W. and Boezi, J.A. 1977. A new staining technique for protein in polyacrylamide gels using Coomassie Brilliant Blue G-250. Anal. Biochem. 82:580-582. Blum, H., Beier, H., and Goss, H.J. 1987. Improved silver staining of plant proteins, RNA and DNA in polyacrylamide gels. Electrophoresis 8:93-99. Goforth, S. 2001. Exposing gel documentation systems. The Scientist. 15:28, Gorg, A. 1985. Horizontal SDS electrophoresis in ultrathin pore gradient gels for the analysis of urinary proteins. Sci. Tools 32:5-9. Miller, M.D., Acey, R.A., Lee, L.Y.T., and Edwards, A.J. 2001. Digital imaging considerations for gel electrophoresis analysis systems. Electrophoresis 22:791-800. Miura, K. 2001. Imaging and detection technologies for image analysis in electrophoresis. Electrophoresis 22:801-813. Electrophoresis
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Morrissey, J.H. 1981. Silver stain for proteins in polyacrylamide gels: A modified procedure with enhanced uniform sensitivity. Anal. Biochem. 117:307-310.
Key References
Patton, W.F. 2000. A thousand points of light: The application of fluorescence detection technologies to two-dimensional gel electrophoresis and proteins. Electrophoresis. 21:1123-1144.
Patton, 2000. See above.
Weber, K., Pringle, J.R., and Osborn, M. 1972. Measurement of molecular weights by electrophoresis on SDS-acrylamide gel. Methods Enzymol. 26:3-27. Yan, J.X., Harry, R.A., Spibey, C., and Dunn, M.J. 2000. Postelectrophoretic staining of proteins separated by two-dimensional gel electrophoresis using SYPRO dyes. Electrophoresis 21:3675-3665.
Miller et al., 2001. See above. Overview of gel-imaging devices.
Describes fluorescent stains. Rabilloud, T. 1992. A comparison between low background silver diammine and silver nitrate protein stains. Electrophoresis 13:429-439. Compares many different silver staining methods. Wilson, C. 1983. Staining of proteins on gels: Comparison of dyes and procedures. Methods Enzymol. 91:236-247. Reviews different Coomassie blue staining procedures.
Contributed by Lynn A. Echan and David W. Speicher The Wistar Institute Philadelphia, Pennsylvania
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Protein Detection in Gels Without Fixation
UNIT 10.6
describes protein detection in gels using methods that fix the protein within the gel. Such methods are not always suitable, particularly when protein recovery from the gel is the ultimate goal. In general, fixatives remove SDS from gels, causing the denatured protein to become entwined in the gel matrix; subsequent extraction from the gel is usually not practical without partial cleavage of the protein or dissolution of the gel matrix. In this unit, methods for the detection of proteins without fixation are described. In general, the sensitivity of these methods is lower than that of protein detection with fixation. For any given method, sensitivity is dependent on the volume of the protein band within the gel; hence, sensitivity is the highest for sharp, narrow bands. Techniques described in this unit include basic protocols for protein detection in gels by SDS precipitation (Basic Protocol 1), preparation of contact blots (Basic Protocol 2), staining with Imidazole-zinc (Basic Protocol 3), and use of the fluorescent labels IAEDANS (Basic Protocol 4) and fluorescamine (Basic Protocol 5) to detect proteins in gels. Several additional methods, including the use of tryptophan fluorescence, guide strips, and minimal protein staining, are discussed in the Commentary. UNIT 10.5
PROTEIN DETECTION IN GELS BY SDS PRECIPITATION 0◦
Although proteins can be detected in SDS-containing gels by simply chilling at to ◦ 4 C for 3 to 5 hr to precipitate free SDS, resolution is low and special precautions must be taken in preparing and running the gels. The method presented here involves the use of high-ionic-strength solutions to precipitate the SDS and produces the same effect more reliably. Using this method, the SDS in the gel precipitates and forms an opaque background, whereas the SDS bound to the proteins does not precipitate, creating a clear protein band.
BASIC PROTOCOL 1
Materials SDS-polyacrylamide gel containing protein of interest (UNITS 10.1 to 10.4) 4 M sodium acetate, pH unadjusted (APPENDIX 2E; store at 4◦ C; stable up to 6 months) Black, nonreflective surface Fluorescent desk lamp NOTE: All steps are performed at room temperature. 1. Immediately after electrophoresis, place the polyacrylamide gel containing the protein of interest into a covered container with 10 to 12 gel volumes of 4 M sodium acetate. 2. Place on a platform shaker at low speed for 40 to 50 min for 1-mm-thick gels or longer for thicker gels. 3. View the gel over a black, nonreflective surface using a fluorescent desk lamp to illuminate the gel from the side. If the gel bands are not yet visible, allow gels to continue to soak. Monitor gels about every 15 min, or more often if incubation must continue longer than 1 hr. Be careful not to allow the incubation in high-ionic-strength solution to proceed unattended as the resolution will begin to decrease after a few hours. Electrophoresis Contributed by Won-A Joo and David W. Speicher Current Protocols in Protein Science (2007) 10.6.1-10.6.9 C 2007 by John Wiley & Sons, Inc. Copyright
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4. For recovery of proteins by electroelution, soak gel slices in water to reduce the ionic strength. In general, soaking gel slices in 50 ml water for 1 hr, changing the water every 15 min, should be sufficient. Some elution of proteins by diffusion is likely to occur, especially small proteins, i.e., <30 to 50 kDa. BASIC PROTOCOL 2
PREPARATION OF CONTACT BLOTS Unlike electroblot procedures (UNIT 10.7) where it is desirable to transfer all of the protein in the gel onto the membrane surface, contact blots are used to create merely a surface replica of the gel by transferring a small amount of protein to the transfer membrane while leaving most of the protein in the gel. The method involves making a gel/membrane sandwich that is soaked in a Tris/glycine transfer buffer for a short time to effect transfer of proteins from the gel to the membrane. The membrane is then stained to visualize proteins. Because the membrane will contain very little protein, a high-retention polyvinylidene difluoride (PVDF) membrane should be used, and it must be treated with a sensitive stain such as AuroDye (GE Healthcare), MemCode (Pierce Biotechnology), or colloidal gold (UNIT 10.8). For the recognition of specific proteins on a membrane using antibodies, the initial stain method should be reversible, such as Ponceau S or MemCode (Pierce Biotechnology) staining. Protein bands can then be identified in the gel by direct comparison with the transfer membrane.
Materials Polyacrylamide gel containing protein of interest (UNITS 10.1 to 10.4) 100% methanol 1× transfer buffer (see recipe) PVDF transfer membrane: 0.2-µm polyvinylidene difluoride (PVDF) membrane (Bio-Rad) Whatman no. 1 filter paper Additional reagents and equipment for staining membranes (UNIT 10.8) NOTE: Use powder-free gloves rinsed in Milli-Q water when handling all materials for this procedure, and handle membranes with forceps at the edges to avoid potential artifactual staining.
Prepare transfer membrane and gel 1. Cut PVDF transfer membrane slightly larger than the polyacrylamide gel containing the protein of interest. Cut three slightly larger pieces of Whatman no. 1 filter paper. 2. Wet PVDF membrane in 100% methanol in a clean glass dish for 5 sec. Decant methanol and pour in 1× transfer buffer. Incubate membrane in transfer buffer for at least 5 min. The PVDF membrane will wet almost immediately with the 100% methanol. Because the membrane is strongly hydrophobic it will not wet efficiently in either water or transfer buffer alone. Do not allow membrane to dry between wetting with methanol and addition of the transfer buffer.
3. Remove polyacrylamide gel from the electrophoresis unit and mark the first lane by cutting off a small piece of the gel on the lower corner of that side. Rinse gel briefly in transfer buffer and place face down into a plastic container with lid. Protein Detection in Gels Without Fixation
The cut edge should now be on the lower right-hand edge of the gel if the gel was loaded from the left-hand side.
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Make gel/membrane sandwich 4. Pour ∼2 ml transfer buffer on top of the gel. 5. Starting at a place on the gel that has the most buffer on it, lower the prepared PVDF membrane onto the gel. Check carefully for bubbles. Trim the edges of the membrane to the exact size of the gel, including the cut corner, marking the first lane described in step 3. Do not cover the gel with too much buffer or the transferred protein bands will appear diffuse on the membrane. Use only enough buffer so that the membrane can be placed on top of the gel without creating bubbles. If a bubble is detected, lift that portion of the membrane to release the bubble and lower the membrane slowly back onto the gel.
6. Wet the three precut pieces of filter paper with transfer buffer and layer one at a time on top of the membrane. Smooth gently by rolling a test tube or pipet over the surface after adding each layer to eliminate bubbles.
Transfer and stain protein 7. Top with a glass plate to ensure tight contact between the gel and membrane. Cover the plastic container with lid and allow transfer to continue by diffusion for 15 to 30 min. The glass plate should be slightly larger than the gel/membrane sandwich.
8. After opening the container and removing the glass plate, remove the filter paper from the membrane. To ensure proper realignment after staining the membrane, mark three corners of the gel/membrane sandwich with a syringe needle dipped in permanent ink by piercing through both membrane and gel. The membrane and gel should be adequately marked for proper realignment before disassembling the gel/membrane sandwich.
9. Remove the membrane and stain it with a sensitive stain. Keep the gel covered in plastic wrap at 4◦ C to limit diffusion while the membrane is being stained. 10. Once the membrane is stained, identify the position of protein bands in the unstained gel by realigning the gel over the stained blot. Mark bands of interest with a needle dipped in permanent ink as in step 8 or excise the band with a sharp scalpel.
IMIDAZOLE-ZINC STAINING OF PROTEINS Imidazole-zinc stain is a negative stain that utilizes precipitation of free dodecyl sulfate by forming a salt with zinc cations. Because protein-bound ions (e.g., dodecyl sulfate or the heavy cation zinc) are chemically less active than free ions in the gel, precipitation of an insoluble salt is much slower in those regions of gel occupied by proteins than in the gel background where zinc dodecyl sulfate precipitates. The result is a “negative stain,” with translucent proteins and an opaque gel background.
BASIC PROTOCOL 3
Materials Polyacrylamide gel containing protein of interest (UNITS 10.1 to 10.4) 0.2 M imidazole solution containing 0.1% (w/v) SDS, pH is not adjusted 0.2 M zinc sulfate solution Transparent plastic tray 1. After electrophoresis, place the polyacrylamide gel containing the protein of interest in a transparent plastic tray and rinse for 30 sec in Milli-Q water. 2. Incubate in 0.2 M imidazole solution with constant gentle agitation. A recommended incubation time in the imidazole solution is 15 min for 10% to15% of the gel. Current Protocols in Protein Science
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3. Discard the imidazole solution and rinse gel for a few seconds in fresh Milli-Q water. Discard rinse. 4. Incubate gel in 0.2 M zinc sulfate while manually shaking the gel for ∼1 min. The gel background will “stain,” becoming white due to the precipitation of a zincimidazole-SDS complex on the gel surface. Prolonging the incubation in the zinc solution may result in indiscriminate staining of the gel surface and loss of protein band detection. Increasing the concentration of imidazole (i.e., 0.3 M or higher) generates a deeper white background that may render the detection of less-abundant proteins difficult. BASIC PROTOCOL 4
IAEDANS MODIFICATION OF PROTEINS N-Iodoacetyl-N -(5-sulfo-1-naphthyl)ethylenediamine (1,5-I-AEDANS or IAEDANS) is a modified iodoacetic acid (IAA) reagent formed by linkage of IAA to an aminonaphtholsulfonic acid fluorophor. This reagent specifically modifies cysteine residues. IAEDANS-modified cysteines have an excitation maximum of ∼340 nm and an emission maximum of 418 nm. In this procedure, modification of protein with the fluorescent label is performed prior to electrophoresis. Several dialysis steps may be needed to provide labeled proteins suitable for electrophoresis. A major advantage of this method is that proteins can be modified in crude mixtures and labeled proteins can be followed through all subsequent steps, including elution from the gel as well as visualization in the gel.
Materials Protein sample (<5 mg protein in 0.3- to 3-ml volume) 0.75 M Tris·Cl (pH 8.6)/15 mM EDTA (make with 4◦ C water, measure pH at that temperature, and use within 1 week; store at 4◦ C) Ultrapure Urea (Bio-Rad) Dithiothreitol (DTT or Cleland’s reagent; Calbiochem) Argon or nitrogen gas N-Iodoacetyl-N -(5-sulfo-1-naphthyl)ethylenediamine (IAEDANS; Sigma; keep light yellow crystals desiccated in the dark at −20◦ C) 2-mercapthoethanol Additional reagents and equipment for dialysis (APPENDIX 3B) and electrophoresis (UNITS 10.1 to 10.4) CAUTION: Use UV-rated eye protection when using UV lamps and minimize skin exposure to direct or reflected UV light. 1. Dialyze 0.3 to 3 ml protein sample (<5 mg total protein) twice against 1 liter Tris/EDTA buffer at 4◦ C, for a minimum of 3 hr each time. The Tris scavenges the cyanate (CN- ) produced by decomposing urea (see below). The indicated volumes are convenient ranges, but the method can be adapted for smaller or larger volumes if needed.
2. Remove protein from the dialysis tubing and measure the volume. Place sample in a tube and add dry urea to a final concentration of 9 M (0.833 g/ml original volume). Bring the solution to room temperature to dissolve urea. When the urea is added the sample volume will increase to ∼1.5 ml for each 1 ml original volume. It is important to take this into account in calculating amounts of reagents. Protein Detection in Gels Without Fixation
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3. Immediately add DTT to a final concentration of 20 mM (4.62 mg/1.5 ml reaction solution, based on volume after urea addition). DTT will reduce any oxidized components including artificial disulfides that may have formed during protein purification procedures or storage. 2-mercaptoethanol should not be substituted for DTT because it is a weaker reducing reagent.
4. Flush the tube with argon or nitrogen and seal to provide an inert atmosphere. Incubate the tube 1 hr at 37◦ C. 5. Add 30.1 mg IAEDANS/1.5 ml reaction mixture. Mix to dissolve the reagent, flush with argon or nitrogen, and reseal the tube. Wrap the entire tube carefully in aluminum foil. Because IAEDANS is light-sensitive, minimize the time and amount of exposure to light. The indicated quantity of IAEDANS is sufficient to neutralize the DTT (2 moles IAEDANS per mole DTT) plus a moderate excess over thiol groups in the protein. Because IAEDANS modifies any reactive thiol groups, this excess is necessary for complete labeling of cysteine residues in the protein.
6. Incubate the sample 1 hr at room temperature. Stop the reaction by adding 2mercaptoethanol to 100 mM final concentration. The sample will appear greenish-yellow in color.
7. Dialyze the protein for ≥4 hr against the desired buffer using approximately a 1000fold larger volume of dialysis buffer relative to sample volumes. Repeat with fresh dialysis buffer until the protein sample is colorless. Conduct the first dialysis in the dark. IAEDANS equilibrates across dialysis membranes very slowly, so each dialysis should be performed for a minimum of 4 hr. Longer times are better. The dialysis can be followed by using a hand-held UV lamp. The protein should be colorless and strongly fluorescent, whereas unreacted reagent is yellow and weakly fluorescent. Samples do not have to be completely dialyzed to colorless prior to electrophoresis. One to two dialyses following the labeling should be sufficient to reduce the level of unreacted IAEDANS such that it will not interfere with electrophoretic separation; the weakly fluorescent underivatized reagent will migrate near the dye front.
8. Perform gel electrophoresis, following movement of IAEDANS-modified protein during and after the electrophoresis by exposing gels to a hand-held UV lamp. If desired, excise fluorescent bands from the gel with a sharp scalpel and monitor the efficiency of subsequent removal from the gel using a UV light.
FLUORESCAMINE LABELING OF PROTEINS 4-phenylspiro[furan-2(3H),1 -phthalan]-3,3 -dione,
Fluorescamine, or reacts with αamino groups of proteins and with the ε-amino groups of lysine residues. The sensitivity of fluorescamine labeling is therefore dependent on the number of lysine residues in a particular protein, but in general as little as 0.1 µg of protein should be easily detected. The excitation maximum for fluorescamine is 390 nm and the emission maximum is 475 nm. This method involves covalent labeling of proteins prior to electrophoresis like the IAEDANS procedure, but is quicker because only a single round of dialysis is needed following labeling.
BASIC PROTOCOL 5
Materials Protein sample (0.5 to 1 mg/ml) 15 mM sodium phosphate buffer, pH 8.5 (APPENDIX 2E; prepare with 4◦ C water just before use)
Electrophoresis
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SDS (Bio-Rad) Sucrose Dithiothreitol (DTT; optional) 1 mg/ml fluorescamine (Sigma) in acetone 5 mg/ml bromphenol blue (optional) 37◦ C water bath Additional reagents and equipment for dialysis (APPENDIX 3B) and electrophoresis (UNIT 10.1) CAUTION: Use UV-rated eye protection when using UV lamps and minimize skin exposure to direct or reflected UV light. 1. Dialyze 100 µl protein sample against 1 liter of 15 mM sodium phosphate buffer, pH 8.5, at 4◦ C for a minimum of 3 hr. 2. Remove protein from the dialysis tubing and add dry SDS and sucrose to a final concentration of 5% (w/v) each. Alternatively, lyophilize the sample and dissolve the protein (0.5 to 1 mg) in 100 µl of 15 mM sodium phosphate, pH 8.5, containing 5% (w/v) SDS and 5% (w/v) sucrose. Use of Tris should be avoided because it will react with the fluorescamine reagent.
3. Heat the sample 15 min at 37◦ C to dissolve protein. Higher temperatures can be used if needed to effectively solubilize the protein. If reduction with DTT is necessary to reduce disulfides, add the reducing reagent (20 mM DTT final concentration) prior to heating and increase time at 37◦ C to 1 hr after flushing the sample reaction tube with argon or nitrogen.
4. Add 5 µl of 1 mg/ml fluorescamine in acetone to the protein mixture and mix well. Labeling will occur rapidly, and the degree of modification can be controlled to some extent by the time of incubation prior to electrophoresis. For example, 1 min is sufficient for moderate labeling and 30 min for extensive labeling.
5. Transfer the desired volume of sample to a clean microcentrifuge tube. Add 1 µl of 5 mg/ml bromphenol blue as a tracking dye, if desired, and apply to a polyacrylamide gel well. 6. Conduct electrophoresis. Visualize labeled proteins during or after separation with a hand-held UV lamp.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Transfer buffer
Protein Detection in Gels Without Fixation
20× stock solution: 200 mM Tris base 2.0 M glycine Do not adjust pH Stable up to several months at room temperature 1× working solution: 50 ml 20× stock 100 ml methanol H2 O to 1 liter Do not adjust pH Make 1× working solution fresh daily
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COMMENTARY Background Information Polyacrylamide gel electrophoresis is widely used for the separation of protein mixtures (UNITS 10.1 to 10.4), and a variety of methods to detect the separated proteins have been published. Methods involving fixation of the protein within the gel are presented in UNIT 10.5. Fixation is used to inhibit diffusion of the proteins within and out of the gel matrix. Unfortunately, the fixation process eliminates biological activity of the protein and inhibits recovery of intact proteins from gels. If the protein is needed in its native form and/or recovery of the intact protein from the gel is needed, methods for detection in polyacrylamide gels that do not involve fixation are necessary. In general, the latter methods are required when the protein must be eluted from a polyacrylamide gel. Historically, a standard procedure for locating a desired protein has involved making 1-mm slices of an entire gel or reagent of a gel using prestained proteins as a guide. Protein is then eluted from each slice, and the eluants assayed for the desired protein. That procedure is time-consuming, and it is likely that the gel will not be cut in the best place. This makes it preferable to visualize specifically the protein in the gel, without fixation. Several methods currently available for this purpose are discussed in this unit. The choice of label for detecting proteins in polyacrylamide gels without fixation is largely dependent on the ultimate use to which the protein will be put. If it is desirable to modify the protein as little as possible, SDS precipitation or contact blot preparation offer the least manipulation of the proteins (Basic Protocols 1 and 2). The most reliable method for precipitating free SDS uses high-ionic-strength solutions (Higgins and Dahmus, 1979), although simply chilling gels at 0◦ to 4◦ C to precipitate SDS can be used under certain running conditions (Wallace et al., 1974). Making contact blots involves preparing a gel/membrane sandwich similar to that described for electroblotting (UNIT 10.7); however, in a contact blot transfer is achieved by diffusion. The transfer buffer listed in Basic Protocol 2 is approximately half-strength Towbin buffer (Towbin et al., 1979), but other transfer buffers may be suitable as well. Typical protein detection sensitivity with the imidazole-zinc staining procedure approach is in the low-nanogram range (1 to 10 ng/band). This protein staining method (Basic Protocol 3) is based on using a procedure re-
ported by Hardy and Castellanos-Serra (1992). The stain is reversible when divalent cation chelators such as EDTA are used. This method allows very high sensitivity detection followed by efficient recovery of intact proteins for further structural or biological analyses. The reproducibility is better than that of copper or zinc negative staining. Fluorescent labels chemically modify the protein and may affect migration in gels prepared either with or without SDS, but they offer the advantage that the protein can be monitored at all subsequent steps including removal from the gel, which can be problematic when proteins cannot be directly visualized and monitored. The fluorescent label IAEDANS is especially attractive and can assist in localizing proteins containing cysteine residues. The IAEDANS modification method described in Basic Protocol 4 is based on that of Speicher et al. (1983). Another fluorescent label, fluorescamine, modifies N-terminal amino groups and ε-amino groups on lysine residues; it is preferred over the alternative fluorescent label dansyl chloride because neither its free form nor its hydrolysis products are fluorescent. Labeling conditions are usually selected that minimally modify any individual sites. The simple method of fluorescamine labeling presented in Basic Protocol 5 is based on a procedure described by Eng and Parkes (1974). The fluorescent labels can also be advantageous when the whole protein or peptide products after specific protein cleavage are subjected to HPLC because peaks containing the desired product can be identified by fluorescence. In addition to the protocols given in this unit, several alternative simple, nonperturbing methods may be used. Intrinsic tryptophan fluorescence. In this procedure, the gel is frozen in a suitable container which is floated in liquid nitrogen. Proteins are detected using a hand-held UV lamp. Sensitivity depends on the final temperature achieved, the intensity of the lamp, and the number of tryptophan residues in the protein. In most cases, sensitivity is similar to that obtained with Coomassie blue staining. Guide strip staining. Multiple identical lanes are run on a polyacrylamide gel. The end lanes of the gel (and the center lane, for improved accuracy) are cut out and stained by one of the methods listed in UNIT 10.5. Meanwhile, the unstained portion of the gel is kept in plastic wrap at 4◦ C. Once the stained portions
Electrophoresis
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Supplement 48
of the gel have been destained, the stained gel slices can be lined up with the unstained gel. From this alignment, the region of the desired protein band can be estimated and the corresponding portion of the gel excised. This method works reasonably well, provided the sizes of the stained and unstained gel sections are identical. Minimal protein staining. With this rapid detection method, the gel is treated briefly (1 to 5 min) with Coomassie blue R-250 (UNIT 10.5) so that the stain and fixative do not have time to penetrate into the gel matrix, and only the protein molecules near the surface are actually fixed and stained. This method, however, has two drawbacks. The brief staining procedure will emphasize gel surface artifacts, which could lead to incorrect localization of the protein band, and the portion of sample that is fixed in the gel is likely to vary appreciably from gel to gel and is difficult to control.
Critical Parameters
Protein Detection in Gels Without Fixation
In all procedures, it is essential to use highpurity reagents. In addition, if the protein of interest is to be eluted from the gel, it is important to consider all the factors that may affect protein recovery and activity after isolation. Although fluorescent probes provide the most sensitive and easiest detection, the impact of the modifications must be assessed through the use of appropriate controls to evaluate potential shifts in migration resulting from the chemical modification. When reaction conditions are optimized for complete modification, IAEDANS labeling can be used for twodimensional electrophoresis using isoelectric focusing (UNIT 10.4) with little difficulty (Speicher et al., 1983). The use of fluorescent probes offers a sensitive alternative for detecting proteins in unstained gels. It is crucial, however, to follow the protocols carefully to obtain efficient and specific labeling of proteins. The modification with IAEDANS is dependent on the pH (pH 8.4 to 8.6 is optimal), state of denaturation (8 to 10 M urea is preferred), level of reduction (50- to 200-fold excess of DTT over cysteines in the protein under argon at 37◦ C for 1 to 2 hr), and the concentration of IAEDANS (allow 2 moles IAEDANS for every mole of DTT plus at least a 10- to 20-fold molar excess of IAEDANS over SH groups). A larger excess of IAEDANS as well as changes in reaction time or pH can lead to modification of other amino acids. Incomplete reduction can lead to incomplete modification. It is therefore important
that DTT be used instead of the weaker reducing agent 2-mercaptoethanol. Because modification is performed prior to electrophoresis, incomplete modification of cysteines will result in heterogeneity in isoelectric focusing separations as each modified residue adds negative charge to the molecule. Appreciable molecular weight shifts can also occur, especially when a large number of cysteines are modified in a given protein. A useful alternative approach is to delete the reduction step and/or forgo the use of urea to modify only cysteines not involved in disulfide bonds or solvent-exposed cysteines, respectively. If the reduction step is deleted, the amount of IAEDANS should be reduced to 4.3 mg/1.5 ml of reaction solution (Basic Protocol 4). The fluorescamine-protein interaction changes the net charge of proteins so that mobilities are changed in polyacrylamide gels run in the absence of SDS. As the charge of the protein is less important in SDS-polyacrylamide gels, moderately labeled proteins should run with similar mobilities to the unlabeled proteins.
Troubleshooting There are several problems associated with detecting proteins in unstained gels. Perhaps the most common problem arises from the quantity of protein to be detected. With the exception of the fluorescent labeling and imidazole-zinc methods, these procedures are not very sensitive. Also, some practice may be required before proteins can be detected reliably. One problem that may occur with contact blots is blurring of the transferred bands. If this occurs, it is most likely due to the presence of too much buffer in the transfer sandwich or to insufficient contact between the gel and membrane. Only enough buffer to keep the gel and membrane moist is needed. Improved contact can be achieved by placing a 100-ml beaker filled with water on top of the glass plate. The cover to the gel box must be omitted in this case. Also, if the gel is not rinsed with transfer buffer prior to exposure to the transfer membrane, protein binding to the membrane may be inhibited. On the other hand, if the gel is left in the transfer buffer too long, too much SDS will be removed from the gel and the transfer will not be efficient; therefore, only a brief rinse of the gel is recommended (∼10 sec). If insufficient protein is transferred to the membrane by contact blotting, a brief electrotransfer (UNIT 10.7) can be used. Transfer time and transfer voltage should be kept as low as consistent with adequate detection sensitivity
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to ensure that a minimal amount of protein is transferred from the gel. With the fluorescent labels, if the modification is incomplete or appears to be nonspecific, the reaction conditions should be carefully reviewed. Check calculations for the amount of fluorescent label added as well as reaction pH, reduction conditions, and length of labeling as all these parameters are important for successful labeling of the protein.
complete; although the labeling procedure itself takes only 2 to 3 hr, the reaction mixture should be dialyzed at least 4 hr before the protein is run on a polyacrylamide gel to minimize interference of reaction by-products with the gel separation.
Anticipated Results
Hardy, E. and Castellanos-Serra, L.R. 1992. Reverse staining of sodium dodecyl sulfatepolyacrylamide gels by imidazole-zinc salts: Sensitive detection of unmodified proteins. BioTech. 12:564-573.
SDS precipitation and fluorescent labels yield similar sensitivities and should detect as little as 0.05 to 0.5 µg protein. This is only a general estimate, however, as the level of fluorescent labeling is dependent on the number of modified residues in the protein sample. The sensitivity of a contact blot depends on both the amount of protein transferred and the membrane staining technique (see UNIT 10.8). The most sensitive staining techniques can detect as little as 1 ng protein on the blot.
Time Considerations The detection of proteins in unstained gels can take as little as 5 min or as long as 1 day, depending on the method. The quickest method available is monitoring of fluorescent bands in the gel. SDS precipitation with high ionic strength (Basic Protocol 1) occurs within 10 min for protein bands containing 1 to 5 µg protein. Contact blots (Basic Protocol 2) are relatively quick to set up and require a short period for protein transfer to occur (30 min). However, the transfer membranes must be stained with a sensitive stain such as MemCode (Pierce Biotechnology) which takes a minimum of 30 min. The imidazole-zinc stain (Basic Protocol 3) takes about 30 min. With fluorescent labeling, dialysis of the proteins both before and after labeling consumes the majority of time required. Dialysis of unlabeled protein into the modification buffer can be done the night before labeling. With fluorescamine (Basic Protocol 5), labeling of the dialyzed protein can be completed within 30 min, and the sample is ready to load on a gel immediately. IAEDANS labeling (Basic Protocol 4) requires more time to
Literature Cited Eng, P.R. and Parkes, C.O. 1974. SDS electrophoresis of fluorescamine-labeled proteins. Anal. Biochem. 59:323-325.
Higgins, R.C. and Dahmus, M.E. 1979. Rapid visualization of protein bands in preparative SDSpolyacrylamide gels. Anal. Biochem. 93:257260. Speicher, D.W., Davis, G., and Marchesi, V.T. 1983. Structure of human erythrocyte spectrin. II. The sequence of the α-I domain. J. Biol. Chem. 258:14938-14947. Towbin, H., Staehlin, T., and Gordon, J. 1979. Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: Procedure and some applications. Proc. Natl. Acad. Sci. U.S.A. 76:4350-4354. Wallace, R.W., Yu, P.H., Dieckert, J.P., and Dieckert, J.W. 1974. Visualization of proteinSDS complexes in polyacrylamide gels by chilling. Anal. Biochem. 61:86-92.
Key Reference Hames, B.D. and Rickwood, D. (eds.) 1990. Gel Electrophoresis of Proteins: A Practical Approach. Oxford University Press, New York. A general reference for gel electrophoresis and protein detection methods. Simpson, R. 2003. Peptide mapping and sequence analysis of gel-resolved proteins. In Protein and Proteomics: A Laboratory Manual (R. Simpson, ed.) pp. 394-395. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. A general reference for gel electrophoresis and protein detection methods.
Contributed by Won-A Joo and David W. Speicher The Wistar Institute Philadelphia, Pennsylvania
Electrophoresis
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Electroblotting from Polyacrylamide Gels
UNIT 10.7
Electroblotting of proteins from polyacrylamide gels onto retentive membranes is usually performed to facilitate procedures leading to protein identification and characterization. It has been used for immunoblotting (or Western blotting), N-terminal protein sequencing, in situ protease or chemical cleavage of proteins for further structural analysis (UNIT 11.2 & UNIT 11.5), protein band detection using sensitive stains such as AuroDye colloidal gold (UNIT 10.8), and protein elution. The electroblotting methods in this unit require that the proteins have already been separated on a polyacrylamide gel (UNITS 10.1-10.4). However, if the blotted protein bands will be used for N-terminal protein sequencing or sequencing of cleavage/digestion products, refer to the specific requirements detailed in Alternate Protocol 1, which may affect electrophoresis conditions. Including protein markers or prestained standards in gels is useful because they will transfer to the membrane and can be used to indicate the size of proteins after immunostaining. This unit contains procedures for electrophoretically transferring proteins onto a variety of membranes including polyvinylidene difluoride (PVDF; Basic Protocol) and nitrocellulose (Alternate Protocol 2) and derivatized membranes. The choice of membrane type for electrotransfer is dependent on the ultimate application for the blot membrane. High-retention PVDF binds proteins tightly and is well suited for applications such as sequencing where high protein amount is critical (Alternate Protocol 1), whereas low-retention membranes may be advantageous for subsequent protein extraction or immunoblot analysis. Most protocols are designed for use with tank transfer setups; Alternate Protocol 3 presents procedures for electroblotting in semidry systems. In some cases, stained blots are used only to identify protein band patterns while leaving the gel unmodified for subsequent steps (UNIT 10.8). If such minimal protein transfer is desired, contact blotting is a suitable alternative (UNIT 10.6). This unit also describes procedures for eluting proteins from membranes using detergents (Basic Protocol 2) or acidic extraction with organic solvents (Alternate Protocol 4). Unless otherwise indicated, the following protocols are primarily designed for electrotransferring proteins from SDS gels (UNITS 10.1-10.4). As proteins in other types of gels have much lower charge densities and, sometimes, opposing net charges for different proteins, substantial modifications to the protocols must be made for these specialized applications (UNIT 10.2). ELECTROBLOTTING ONTO PVDF MEMBRANES The following procedure is based on electrotransfer from polyacrylamide gels containing 0.2% sodium dodecyl sulfate (SDS) in the gel solution and electrophoresis and sample buffers. It uses a Bio-Rad solid plate tank transfer apparatus with 7 cm spacing between the electrode plates, which can accommodate gels up to 14 × 18 cm in size and requires ∼2.5 liters transfer buffer. The choice of PVDF membrane depends on the planned subsequent use: high-retention Trans-Blot membranes (Bio-Rad) are suitable for protein sequencing, whereas low-retention Immobilon-P membranes (Millipore) are recommended for low-background immunodetection or staining and for recovering proteins from membranes.
BASIC PROTOCOL 1
Electrophoresis Contributed by Jeanine A. Ursitti, Jacek Mozdzanowski, and David W. Speicher Current Protocols in Protein Science (1995) 10.7.1-10.7.14 Copyright © 2000 by John Wiley & Sons, Inc.
10.7.1 CPPS
Materials 1× transfer buffer (see recipe) Polyacrylamide gel containing proteins of interest (UNITS 10.1-10.4) 100% methanol Powder-free gloves Electroblotting apparatus: “solid” plate electrode tank transfer system (e.g., Trans-Blot Cell, Bio-Rad) Glass dishes and trays Gel support sheet (e.g., porous polyethylene sheet, Curtin Matheson) PVDF transfer membrane: 0.2 µm PVDF membrane (Bio-Rad) or Immobilon-P (Millipore) Whatman no. 1 filter paper Power supply (500 V, 300 mA) Additional reagents and equipment for staining gels (UNIT 10.5) NOTE: Use powder-free gloves when handling all materials for this procedure and handle membranes with forceps at the edges to avoid potential artifactual staining. Equilibrate the gel and membrane in transfer buffer 1. Prior to blotting, prepare the blotting apparatus by thoroughly rinsing with high-purity water. The optional porous polyethylene sheet is hydrophobic, and it is difficult to get water or buffer into the pores. Be sure to hydrate or “wet” it properly by spraying water under pressure through the sheet or by submerging the entire polyethylene sheet in methanol and then submerging in high-purity water.
2. Fill the transfer tank with 1× transfer buffer. Submerge the gel cassette holder, fiber pads, and polyethylene sheet (after initial hydration as described in step 1) in transfer buffer. Place them inside the tank or submerge in transfer buffer using a separate tray. The transfer buffer can be left in a covered transfer tank for up to 2 hr prior to use. Longer times should be avoided because the methanol in the buffer will evaporate, significantly changing the buffer composition.
3. Prepare the polyacrylamide gel containing proteins of interest by disassembling the gel apparatus, removing the stacking gel with a razor blade, and cutting a small piece from the lower left-hand corner of the gel (near lane 1 for gels loaded left to right; see Fig. 10.7.1) to aid in identifying the lanes in subsequent steps. 4. Briefly equilibrate the gel in a glass dish containing ∼250 ml of 1× transfer buffer for an appropriate time based on gel thickness (0.5-mm-thick gels for 1 min; 0.75- to 1-mm gels for 5 min; 1.5-mm gels for 15 min). Longer equilibrations in transfer buffer prior to electrotransfer may extract too much SDS from the gel and hence reduce transfer efficiency. This equilibration step will remove excess SDS and prevent the gel from swelling during transfer. The negatively charged SDS allows proteins to move toward the transfer membrane in an electric field. On the other hand, too much SDS decreases binding of protein to PVDF membranes. Therefore, the amount of SDS is critical for transfer yield (see Critical Parameters and Troubleshooting).
5. Cut the PVDF transfer membrane about 0.5 to 1 cm larger than the gel. Cut a piece of Whatman no. 1 filter paper ∼0.5 cm larger than the PVDF membrane. Electroblotting from Polyacrylamide Gels
6. Wet the PVDF membrane in a clean glass dish containing 100% methanol for 5 sec. Transfer membrane to a tray containing transfer buffer.
10.7.2 Current Protocols in Protein Science
A
B
C transfer membrane (mirror image of gel)
lane 1
gel face up
lane 1
gel turned over to create mirror image
transfer membrane (with proteins)
lane 1
gel after transfer
Figure 10.7.1 Aligning the gel for transfer. (A) If the polyacrylamide gel has been loaded from left to right, cut a small piece from the corner of the lower left-hand edge of the gel near the first lane. (B)When preparing the transfer sandwich, turn the gel over so that the cut edge is on the lower right-hand corner of the gel. This will ensure that the transferred proteins will appear in the same order as in the original gel. (C)After transfer, trim the membrane above the cut corner of the gel to mark orientation. Dots indicate where the proteins were before transfer.
The PVDF membrane will wet almost immediately with the 100% methanol. The hydrophobic membrane will not wet in either water or transfer buffer alone. After initial wetting in methanol, do not let the membrane dry; if drying occurs, rewet the membrane in 100% methanol.
Prepare the gel/membrane transfer sandwich 7. Place the opened gel holder cassette on a clean, flat surface. Place one transfer buffer–wetted fiber pad (see step 2) on the top surface, followed by the gel support sheet. A sheet of porous polyethylene is preferred as a support because it is rigid and facilitates handling, but filter paper can be used as a simple alternative. If the polyethylene sheet is used, place it with the smooth side up (toward the gel).
8. Place the gel face down on the support, so that the cut edge is now on the right-hand side (see Fig. 10.7.1B). Pour 5 to 10 ml transfer buffer on top of the gel. 9. Remove wet PVDF membrane from the tray containing transfer buffer (step 6). Eliminate air bubbles on both surfaces of membrane by sliding the membrane across the edge of the glass tray. Resubmerge membrane in transfer buffer for a few seconds and then position membrane above gel, letting its center contact the gel, and slowly lower the membrane from the center outward to force any bubbles to the edge of the gel. Rinse gloved hands with high-purity water and smooth the membrane gently to Electrophoresis
10.7.3 Current Protocols in Protein Science
pads
cathode
anode +
+
plastic gel holder cassette polyethylene sheet filter paper
gel transfer membrane transfer buffer direction of protein transfer
Figure 10.7.2 Electroblotting with a tank transfer unit. The polyacrylamide gel containing the protein(s) to be transferred is placed on the smooth side of the polyethylene sheet (or filter paper sheets) and covered with the PVDF membrane and then a single sheet of filter paper. This stack is sandwiched between two fiber pads and secured in the plastic gel holder cassette. The assembled cassette is then placed in a tank containing transfer buffer. For transfer of negatively charged protein, the membrane is positioned on the anode side of the gel. Charged proteins are transferred electrophoretically from the gel onto the membrane.
ensure uniform contact with the gel. Inspect the membrane for any trapped air bubbles. This step is extremely important. Air bubbles will prevent the transfer of proteins.
10. Briefly wet the filter paper with transfer buffer and cover the PVDF membrane. Smooth gently to remove any air bubbles. Place the other fiber pad on top of the membrane and close the holder. Do not allow any area of the PVDF membrane to dry during assembly, or transfer will not occur in these areas. The transfer sandwich should fit together snugly to provide good contact between the membrane and gel. The complete gel sandwich should look like that in Figure 10.7.2.
Conduct protein electrotransfer 11. Slide the assembled transfer cassette into the tank with the gel on the cathode side and the membrane on the anode side. Proteins in an SDS-polyacrylamide gel have a net negative charge and will migrate toward the anode. For transfer from another type of gel, the orientation of the cassette may need to be reversed.
12. Fill the transfer tank with 1× transfer buffer so that the buffer completely covers the electrode panels but does not touch the electrical connectors. Electroblotting from Polyacrylamide Gels
13. Connect the power supply and perform the transfer at constant current compatible with complete transfer from the gel. For a Bio-Rad Trans-Blot tank transfer apparatus
10.7.4 Current Protocols in Protein Science
with solid plate electrodes, use 200 to 250 mA constant current. Transfer 0.5-mm gels for 1 hr, 0.75- to 1-mm gels for 2 hr, and 1.5-mm gels for 3 hr. Longer times do not appear to increase transfer yield but may lead to overheating of the gel. Transfers can be carried out at constant voltage as well, but the current may increase during the run. Use caution to prevent overheating and/or unacceptably high current. Because of the amount of heat generated at higher transfer rates, it may be necessary to run the transfer in a cold room or with a cooling core to prevent overheating.
14. At the end of the transfer period, turn off the power and disconnect the power supply. Open the transfer sandwich and remove the membrane and gel (Fig. 10.7.1C). Thoroughly rinse the membrane with high-purity water, three times, 5 min each. The membrane can be stained as described in UNIT 10.8 or used for other detection methods. Stained or unstained membranes can be air dried (for ∼30 min) and stored in resealable plastic bags at room temperature for several weeks or at −20°C for a permanent record. Dried membranes can be rehydrated in 50% or 100% methanol and used for subsequent detection procedures.
15. To assess the efficiency of transfer, stain the proteins remaining on the gel with Coomassie blue or with more sensitive silver solutions, depending on the amount of protein originally loaded on the gel. ELECTROBLOTTING OF PROTEINS FOR SEQUENCE ANALYSIS One-dimensional or two-dimensional gel electrophoresis combined with electroblotting to PVDF membranes provides a method to obtain proteins of high purity that can be used directly for N-terminal protein sequence analysis. The major concern is avoiding possible chemical modification of proteins such as N-terminal blocking. Simple precautions such as casting gels in advance and modifying the composition of reagents to protect proteins minimize the possibility of side reactions during electrophoresis and electroblotting and are described in this protocol.
ALTERNATE PROTOCOL 1
NOTE: Use high-quality water such as Milli-Q-purified water or equivalent and high-purity electrophoresis reagents (i.e., Bio-Rad) throughout for best results. Additional Materials (also see Basic Protocol 1) Thioglycolate (thioglycolic acid, sodium salt; Sigma) 2× or 6× SDS sample buffer (for discontinuous systems; UNIT 10.1) PVDF transfer membrane: 0.2-µm PVDF membrane (Bio-Rad), ProBlott (Perkin-Elmer), or Immobilon-PSQ (Millipore) Additional reagents and equipment for one- or two-dimensional gel electrophoresis (UNITS 10.1 & 10.4) and for staining membranes (UNIT 10.8) and gels (UNIT 10.6) 1. Cast polyacrylamide gel including stacking gel at least 24 hr but not more than 48 hr prior to use. Store the gel at room temperature until needed, being sure to protect it from dehydration (e.g., store submerged in high-purity water). For preparing gels choose an acrylamide concentration such that the protein will migrate as a sharp, tight band with an Rf between 0.2 and 0.8. Casting a gel in advance allows complete polymerization, reduces the amount of oxidants and free radicals, and minimizes the possibility of blocking the N terminus or other amino groups of the protein.
2. Add thioglycolate to the electrophoresis buffer in the cathode buffer chamber to a final concentration of 0.1 mM (11.4 mg per liter buffer). Thioglycolate scavenges remaining free radicals and oxidants in the gel. Electrophoresis
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3. Solubilize samples in either 2× or 6× SDS sample buffer (for discontinuous systems; UNIT 10.1), containing sucrose or glycerol but not urea. Heat to 37°C for 15 min. If possible, avoid boiling the sample, as the high temperature increases the risk of chemical modification of the protein. Boiling may be necessary for complete solubilization of some samples, however (e.g., whole viruses). DTT or 2-ME in the sample buffer is acceptable.
4. Conduct gel electrophoresis. Proceed with electroblotting (see Basic Protocol 1, steps 1 to 14). 5. After transfer is complete, rinse the membrane six times for 5 min each with a large volume of water (at least 200 ml each time). The membrane must be thoroughly rinsed with water immediately after transfer is complete. It is critical that the membrane is not allowed to dry prior to thorough rinsing because even partial drying can prevent effective removal of Tris and glycine.
6. Stain the membrane with amido black or Ponceau S to detect proteins. Stain the gel after transfer as appropriate to judge transfer efficiency. ALTERNATE PROTOCOL 2
ELECTROBLOTTING ONTO NITROCELLULOSE MEMBRANES The electroblotting procedure for nitrocellulose membranes differs little from that for PVDF membranes. Nitrocellulose is compatible for use with a moderate amount of SDS (up to 0.1%) in the transfer buffer. The delicate membranes must, however, be handled carefully and protected from high concentrations of organic solvents. Additional Materials (also see Basic Protocol 1) 0.45-µm nitrocellulose transfer membrane (Schleicher & Schuell) Additional reagents and equipment for membrane (UNIT 10.8) and gel staining (UNIT 10.6) 1. Prepare the transfer apparatus and gel as described for PVDF membranes (see Basic Protocol 1, steps 1 to 4). 2. Cut the nitrocellulose transfer membrane to a size slightly larger than the gel and cut a piece of filter paper slightly larger than the membrane. 3. Wet the membrane by slowly introducing the membrane from one corner into a glass dish of 1× transfer buffer. Equilibrate the membrane for 15 min. IMPORTANT NOTE: Never soak the membrane in 100% methanol. During electrotransfer and staining, avoid exposure to high concentrations of organic solvents, which will dissolve the membrane. Up to 20% methanol can be used in the transfer buffer without affecting the membrane.
4. Proceed with electrotransfer as above for PVDF membranes (see Basic Protocol 1, steps 7 to 13). 5. When transfer is complete, rinse membrane three times for 5 min each with high-purity water; detect proteins on the membrane using an aqueous stain such as Ponceau S if desired. 6. Assess efficiency of transfer by staining the gel after transfer as appropriate. Electroblotting from Polyacrylamide Gels
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Current Protocols in Protein Science
PROTEIN ELECTROBLOTTING IN SEMIDRY SYSTEMS An alternative to the tank transfer system is the semidry transfer system. In this procedure, the gel is stacked horizontally on top of the membrane in the transfer apparatus. Because only a small volume of transfer buffer is used, SDS from the gel is less effectively diluted, which may result in incomplete binding and lower yields, especially with PVDF membranes. For this reason, semidry transfer units are not recommended when reproducible high recoveries of electroblotted proteins are desired (e.g., for subsequent sequence analysis). Some procedures recommend stacking multiple transfer sandwiches to achieve several transfers simultaneously. To prevent unbound protein from migrating through the next gel and onto the membrane in the next transfer stack, sheets of porous cellophane sheets or dialysis membrane are placed between adjacent transfer stacks (see Fig. 10.7.3). Semidry electrotransfer requires shorter transfer times than tank transfer.
ALTERNATE PROTOCOL 3
Additional Materials (also see Basic Protocol 1) Semidry transfer apparatus Mylar mask (optional) Prepare the transfer sandwiches 1. Cut transfer membrane just slightly larger than the gel containing proteins to be transferred, then equilibrate in the appropriate transfer buffer (see Basic Protocol 1, step 6). 2. Cut six sheets of filter paper the same size as the transfer membrane and wet thoroughly in transfer buffer.
-
cathode
top of gel
buffer-soaked filter paper gel membrane buffer-soaked filter paper
transfer stack
cellophane or dialysis membrane gel membrane buffer-soaked filter paper
transfer stack
buffer-soaked filter paper
Mylar mask
+ anode Figure 10.7.3 Electroblotting with a semidry transfer unit. In most cases, the lower electrode is the anode, as shown. Position the Mylar mask (optional) directly over the anode. Layer on three sheets of filter paper that have been wetted in transfer buffer. For negatively charged proteins, place the preequilibrated transfer membrane on top of the filter paper followed by the gel and three additional sheets of wetted filter paper. If multiple gels are to be transferred, separate the transfer sandwiches by inserting a sheet of porous cellophane or dialysis membrane between each stack. Place the cathode on top of the assembled transfer stack(s). Transfer the proteins by applying a maximum current of 0.8 mA/cm2 gel area.
Electrophoresis
10.7.7 Current Protocols in Protein Science
3. Remove gel containing proteins of interest from the electrophoresis unit. Using a razor blade, cut the lower corner of the gel at the first lane. Equilibrate the gel in transfer buffer (see Basic Protocol 1, step 4). 4. Layer a Mylar mask (optional) followed by three sheets of filter paper on the anode of the transfer unit. Smooth each piece of filter paper to avoid trapping air bubbles between the layers. It is important to remove all air bubbles from the transfer stack as they will block the flow of current through that area of the gel, creating blank spots on the transfer membrane.
5. Layer the prepared transfer membrane on top of the filter paper. Gently roll a test tube or Pasteur pipet over the membrane to push out any air bubbles. 6. Place the gel on top of the transfer membrane so that the cut edge is on the left-hand side. With a pair of scissors, cut the lower corner of the membrane even with the gel (to aid in realigning gel and blot after final staining). Again, remove any air bubbles (see Basic Protocol 1, step 9). When assembling this transfer sandwich, the gel is placed on top of the membrane, in contrast to the opposite order for tank type sandwich assembly as in Basic Protocol 1. Therefore, it is unnecessary to turn the gel over to obtain the proper orientation (compare Figs. 10.7.1 and 10.7.3).
7. Layer the remaining three sheets of filter paper individually on top of the gel, rolling out any air bubbles after each addition. It is possible to transfer multiple gels simultaneously using semidry blotting. As shown in Figure 10.7.3, a sheet of porous cellophane (Hoefer Pharmacia) or dialysis membrane (Bio-Rad or Sartorius), preequilibrated for 5 min in transfer buffer, can be placed between the transfer stacks to prevent proteins from migrating onto an adjacent transfer stack membrane. However, proteins on the gel closest to the anode tend to be transferred more efficiently; thus, transferring multiple gels simultaneously is not recommended for critical applications such as protein sequencing.
Perform the protein electrotransfer 8. Attach the cathode unit on top of the transfer sandwich and connect leads to the power supply. Transfer proteins at constant current for no more than 1 hr. Do not exceed 0.8 mA/cm2 gel surface area. If the outside of the unit becomes warm during transfer, the current is too high and should be lowered.
9. After the transfer period, turn off the power supply and disconnect the leads. Remove the cathode and uppermost three layers of filter paper. Cut the corner of the transfer membrane above the cut corner of the gel. Alternatively, mark the transfer membrane by tracing the gel with a soft lead pencil. 10. Proceed with protein staining or immunodetection, as desired, or dry and store membranes at −20°C for later use (optional).
Electroblotting from Polyacrylamide Gels
10.7.8 Current Protocols in Protein Science
PROTEIN ELUTION FROM PVDF MEMBRANES USING DETERGENTS Binding affinities of most proteins to PVDF membranes are relatively strong. The most efficient protocol for protein recovery from PVDF membranes requires the use of detergents, which limits the possible use of extracted samples because detergents are often incompatible with subsequent procedures. This protocol is a simple procedure to elute proteins from the membrane into a Triton/SDS solution. A protocol that employs acidic extraction with organic solvents is also described (Alternate Protocol 4).
BASIC PROTOCOL 2
Materials Polyacrylamide gel containing proteins of interest Ponceau S stain (UNIT 10.8; optional) 50% methanol (optional) Triton/SDS elution buffer (see recipe) PVDF transfer membrane (Immobilon-P, Millipore) Transfer apparatus: solid plate electrode tank transfer system (e.g., Trans-Blot, Bio-Rad) Additional reagents and equipment for staining membranes (UNIT 10.8) 1. Prepare gel and membrane for transfer as for PVDF membranes (see Basic Protocol 1, steps 1 to 6). 2. Assemble the gel/membrane transfer sandwich and place in transfer tank (see Basic Protocol 1, steps 7 to 10). 3. Conduct electrotransfer (see Basic Protocol 1, steps 11 to 14), using lower field strengths for step 13 (e.g., transfer for 6 hr at 100 mA for 1.5-mm gels). Temperatures >20°C during the transfer can increase the interaction between protein and matrix and make extraction more difficult. If problems arise in eluting proteins from the membrane, try running the transfer in a cold room overnight at low currents (50 mA).
4a. Visualize transferred proteins with any membrane-compatible stain (UNIT 10.8). If bound stain will interfere with subsequent steps, use one of the following alternative detection methods. 4b. Visualize proteins with Ponceau S stain (UNIT 10.8). After staining with Ponceau S, wash the membrane with water, place under clear plastic wrap, and mark bands with a soft pencil by outlining the protein band through the plastic. Completely destain the membrane with water (a permanent indentation made by the pencil will be left on the membrane). 4c. For visualizing bands after partial drying of the membrane, fill a glass dish with 50% methanol and place the dry membrane on the surface of the solution. Do not submerge the membrane. Identify protein bands (portions of the membrane containing proteins will wet more quickly and dry slower than areas that do not contain protein). Remove the membrane from the surface of the methanol solution and place on a clean glass surface. Cover the membrane with clear plastic wrap and use a pencil to mark the protein bands as described in step 4b. 5. Excise the band(s) of interest by carefully cutting the membrane with a clean razor blade or scalpel. Do not allow the membrane to dry during this process. Keep the membrane wet (e.g., submerged in water) at all times (unless using the partial drying procedure).
6. Place the excised membrane band in a 1.5-ml microcentrifuge tube containing 0.2 to 0.5 ml of Triton/SDS elution buffer for every square centimeter of membrane.
Electrophoresis
10.7.9 Current Protocols in Protein Science
Microcentrifuge the sample 10 min at maximum speed, at room temperature, to release the protein from the membrane. 7. Remove supernatant, place it in a new microcentrifuge tube, and microcentrifuge again to remove any particulate material. Multiple extractions with the elution mixture significantly aid quantitative recovery. In most cases, three extractions remove 90% to 100% of the protein from the membrane. ALTERNATE PROTOCOL 4
PROTEIN ELUTION FROM PVDF MEMBRANES USING ACIDIC EXTRACTION WITH ORGANIC SOLVENTS Blotted proteins are usually of high purity and thus good candidates for additional characterization. If the protein cannot be further characterized on the blot and contamination of samples with large amounts of detergent (Support Protocol 1) is unacceptable, acidic elution with organic solvents may be an alternative. The use of organic solvents limits this protocol to PVDF membranes, which have high chemical resistance. This protocol may produce highly variable recoveries for different proteins and is best suited for relatively low-molecular-weight proteins and peptides (<50 kDa). Additional Materials (also see Basic Protocol 2) Extraction solution (see recipe) 50% methanol 1:2 (v/v) acetonitrile/formic acid NOTE: All solutions should be made with high-purity water and can be stored at room temperature for at least 1 month. 1. Rinse the desired number of 1.5-ml microcentrifuge tubes (two tubes per sample) three times with extraction solution, and air dry under an aluminum foil dust cover. Minimize possible contamination with dust at all times.
2. Visualize and mark the protein bands by partial drying of the membrane using 50% methanol (see Support Protocol 1, step 4c). Use a scalpel or razor blade to cut out the protein bands. 3. Place pieces of membrane containing the protein of interest in a clean microcentrifuge tube. Combining several bands from multiple gels is usually necessary. Use a pair of tweezers cleaned with 100% methanol to transfer the membrane pieces to the microcentrifuge tube. Avoid contaminating the membrane pieces and microcentrifuge tube with airborne dust or residues from fingers.
4. Add 200 µl extraction solution to each microcentrifuge tube. An alternative mixture of 1:2 acetonitrile/formic acid seems to be as efficient as the extraction solution described above for some proteins.
5. Agitate tubes for 48 hr at 4°C. 6. Repeat the extraction using 1:2 acetonitrile/formic acid. Combine the extracts. Multiple extractions (up to three) with the same or different solvent mixtures are often beneficial. The extracts should be combined and kept on ice at all times.
Electroblotting from Polyacrylamide Gels
7. Lyophilize protein extracts to provide highly purified samples suitable for most subsequent experiments.
10.7.10 Current Protocols in Protein Science
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Extraction solution (40% acetonitrile/1% trifluoroacetic acid) Add 1.0 ml trifluoroacetic acid to 59 ml high-purity water, then add 40 ml acetonitrile. Store at room temperature. Solution is stable for at least 1 month in a tightly sealed container. Transfer buffer, 20× 200 mM Tris base 2.0 M glycine Do not adjust pH of final solution (stable at room temperature for at least 1 month) (Prepare 1× transfer buffer by mixing 200 ml 20× stock, 400 ml methanol, and water to 4 liters.) Prepare 1× transfer buffer on the same day of use. Keep at room temperature. Triton/SDS elution buffer 50 mM Tris⋅Cl, pH 9.0 (APPENDIX 2E) 2% (w/v) SDS (APPENDIX 2E) 1% (v/v) Triton X-100 Prepare solution on the same day of use. Keep at room temperature. COMMENTARY Background Information The transfer of proteins from polyacrylamide gels onto blot membranes offers many benefits. Transferred proteins can be eluted from the membrane, probed with antibodies (immunoblotting), used for N-terminal protein sequencing as well as other structural analyses including amino acid analysis, protease cleavage (UNIT 11.2), and chemical cleavages (UNIT 11.5), or stained by a variety of highly sensitive techniques. In addition, several manipulations can be performed on the same blot by loading samples in multiple lanes on the gel before electroblotting. Prestained protein standards are useful guides for cutting the membrane prior to different analysis procedures, such as using one part for immunoblotting and the other for protein staining. Prestained radiolabeled standards are useful for aligning the blot membrane with autoradiograms. Prestained standards will migrate differently on the gel compared with the unmodified standard proteins, usually showing higher molecular weight. During electrotransfer, proteins migrate out of gels in an electric field according to the charge on the protein. Most electrotransfers employ a tank transfer apparatus (Fig. 10.7.2) in which the gel/membrane transfer sandwich is mounted in a cassette and placed in a tank of
Tris/glycine/methanol transfer buffer. The semidry transfer system (Fig. 10.7.3) is an alternative. The major advantages of the semidry system are its use of substantially less buffer and the shortening of transfer time due to the close positioning of the electrodes, which produces a high field strength with minimal heating. A variety of transfer membranes, which can effectively bind proteins based on different types of protein-membrane interactions, are commercially available. These include PVDF, nitrocellulose, and nylon membranes as well as a number of derivatized membranes. PVDF membranes bind proteins primarily through hydrophobic interactions and are commonly used for their chemical resistance as well as physical stability. High-affinity PVDF membranes such as Trans-Blot (Bio-Rad), ProBlott (Perkin-Elmer), and Immobilon-PSQ ( Millipore) are preferred for blots intended for use in N-terminal protein sequencing, whereas low-retention membranes such as ImmobilonP (Millipore) may produce lower background in both immunoblotting and common staining procedures. In addition, low-retention membranes are preferred when proteins will be extracted from the membrane. Nitrocellulose binds proteins primarily by hydrophilic and/or electrostatic interactions and for this reason is less sensitive than PVDF
Electrophoresis
10.7.11 Current Protocols in Protein Science
Electroblotting from Polyacrylamide Gels
membranes to the concentration of SDS in both the gel and transfer buffer. Its tolerance to SDS allows relatively good recovery of poorly migrating proteins because higher concentrations of SDS can be left in the gel or added to the transfer buffer without adversely affecting protein binding to the membrane. In addition, nitrocellulose is often used for immunoblots, although PVDF membranes have replaced nitrocellulose for this application in many laboratories. The major limitations of nitrocellulose are its poor binding of low-molecularweight proteins and peptides, mechanical weakness, and lack of resistance to organic solvents. PVDF can be derivatized to modify the nature of protein-membrane interactions and to potentially increase the strength of protein binding. Derivatization most commonly involves adding a charge to the membrane so that proteins will be held on the membrane by electrostatic charges as well the usual hydrophobic or hydrophilic interactions. The procedure for electrotransfer onto derivatized membranes is essentially identical to that for PVDF or nitrocellulose membranes described in this unit. Although derivatized membranes are better for binding RNA and DNA, most derivatized membranes have not offered clear advantages for blotting proteins. One commercially available derivatized PVDF membrane that has potentially useful binding properties for proteins is ImmobilonCD (Millipore), which has been derivatized to produce a cationic surface so that it behaves like an ion-exchange support (Patterson et al., 1992). In addition, the membrane is treated to prevent nonspecific protein adsorption. The well-defined nature of the protein/ImmobilonCD interaction allows good recoveries of electroblotted proteins under relatively mild conditions. Proteins may be recovered from CD membranes nearly quantitatively using denaturing detergent solutions such as 4 M guanidine hydrochloride/0.1% Triton X-100. Proteins can be fragmented using a variety of enzymes and proteases either directly on the Immobilon-CD membrane or after elution. If the proteins are to be digested after elution, the enzymes should be tested for compatibility with the elution buffer. Proteins can also be eluted from ImmobilonP membranes, and two procedures for doing so are described in this unit: protein elution using detergents (Support Protocol 1; Szewczyk and Summers, 1988) and acidic elution with organic solvents (Support Protocol 2; D.W.S.,
unpub. observ.). Although gel electrophoresis combined with subsequent electroblotting is often the most powerful procedure for microscale protein isolation, identification, and characterization, one limitation of this system is that subsequent elution of proteins from PVDF membranes in good yields is often difficult.
Critical Parameters and Troubleshooting The protocols listed in this unit should be sufficient for the transfer of most proteins. The efficiency of protein transfer is easily assessed by staining both the transfer membrane and the gel after transfer. When difficulty is encountered in the transfer recovery of a specific protein, several factors can be considered to improve transfer efficiency. A first consideration is the type of gel used to separate proteins for transfer. Higher percentages of acrylamide make the transfer of proteins out of the gel more difficult. Gel thickness also affects transfer: the thicker the gel, the farther the protein has to migrate to reach the membrane. In general, effective transfer of proteins out of gels up to 1.5 mm thick can be achieved; thicker gels, although having increased protein capacity, offer little advantage owing to reduced removal of proteins from the thicker gel matrix. In this unit, the transfer buffer described is essentially half-strength Towbin buffer (Towbin et al., 1979). This buffer contains a mixture of Tris and glycine and is the buffer of choice if the electroblotted proteins will be used for sequence analysis. Tris, glycine, and most other low-molecular-weight buffer components do not normally bind to PVDF membranes and can be completely removed by thorough rinsing with high-purity water. However, incomplete rinsing or partial drying of the membrane prior to rinsing can result in high residual levels of these compounds, which could be detrimental for sequence analysis. The amino groups in the buffer act as scavengers for contaminants that could potentially modify proteins. Several simple precautions noted in Alternate Protocol 1 can be taken to minimize side reactions of proteins during electrophoresis and electroblotting (Speicher, 1989). If possible chemical modifications are not critical, several other buffers may be used. CAPS buffer (cyclohexylaminopropanesulfonic acid; LeGendre and Matsudaira, 1988) has a high pH (pH 11) and low ionic strength and, therefore, because of its low conductivity,
10.7.12 Current Protocols in Protein Science
permits the use of high electric fields. For the transfer of very basic proteins that bind SDS poorly, high pH may prove advantageous in keeping the total net protein charge negative; however, the high pH of the CAPS buffer can also lead to deamination of sensitive side chains. Other types of electrotransfer buffers such as Tris/borate (Vandekerckhove et al., 1985) do not differ significantly in overall performance from the Tris/glycine buffer and may be used if N-terminal protein blockage is not a major concern. Selection of membrane type is dependent on the ultimate use of the transferred protein, although high-retention PVDF membranes are most often chosen because they bind proteins more tightly than nitrocellulose membranes and have greater chemical and mechanical stability. For immunoblots, either nitrocellulose or Immobilon-P (Millipore) are preferred by some investigators because they produce a lighter background than high-retention PVDF membranes. It is not uncommon in electroblot procedures to observe less than optimal transfer of proteins. If the membrane is entirely blank and little or no protein is left in the gel after transfer, it is likely that the electrodes were connected in a reversed order or that the membrane was placed on the wrong side of the gel. Blank spots on membranes are caused by air bubbles, which block the flow of current. Another problem often encountered with poor transfer of proteins relates to the amount of SDS in the gel, which coats the proteins and improves mobility during electrotransfer. Nitrocellulose membranes are not sensitive to the amount of SDS in the gel and can even tolerate addition of SDS to the transfer buffer to encourage transfer of recalcitrant proteins. PVDF membranes are more sensitive to excess SDS, which can inhibit protein binding to the hydrophobic membrane; nevertheless, in a limited number of cases, addition of SDS to the transfer buffer may be beneficial. Low-retention PVDF membranes are particularly sensitive to excess SDS concentrations, whereas high-retention PVDF membranes are less so. Methanol is a common component of many transfer buffers because it facilitates dissociation of bound SDS from proteins. Therefore, if proteins transfer from the gel efficiently but do not bind well to PVDF membranes, the methanol concentration can be increased to 20% and/or the gel can be preequilibrated in transfer buffer for 15 to 30
min prior to transfer to reduce the SDS concentration (Mozdzanowski et al., 1992). Conversely, if proteins remain in the gel after transfer, the methanol concentration can be reduced or eliminated, and if necessary a low concentration of SDS (0.005%) can be added to the transfer buffer to facilitate electrotransfer from the gel. If proteins have transferred efficiently but the stained membrane has blurred bands or a swirled pattern, it is likely that there was insufficient contact between the gel and membrane during transfer. The transfer sandwich should be held together firmly both to avoid having the gel shift position during transfer and to ensure close contact between the gel and membrane. Another common problem with transfers is the presence of bright white spots within protein bands on the membrane. These are due to air bubbles trapped between the gel and membrane or, in the case of PVDF membranes, possibly due to local drying of the membrane during transfer unit assembly. As indicated in the protocols, it is very important to check for air bubbles before electrotransfer. Air bubbles can be rolled out from the two surfaces by using a test tube, or the membrane can be carefully lifted to release the bubble and replaced without shifting the position of either gel or membrane.
Anticipated Results Proteins in the middle two-thirds of the gel usually transfer to high-retention PVDF membranes with an average yield of 50% to 80%. The protein pattern on PVDF membranes stained with amido black or Coomassie blue should closely resemble the pattern on duplicate lanes of the gel stained with Coomassie blue. The staining intensity on the blot should be slightly higher than on the gel because the proteins are concentrated on the surface of the blot rather than distributed throughout the thickness of the gel. Proteins below the dye front usually will not be recovered on the membrane. Proteins in the top 20% of the gel are often incompletely transferred out of the gel.
Time Considerations
Assembling the transfer unit takes ∼30 min. Electroblotting can be completed in 2 to 4 hr in most cases using a tank transfer unit and in ∼1 hr using a semidry blotting unit.
Electrophoresis
10.7.13 Current Protocols in Protein Science
Literature Cited LeGendre, N. and Matsudaira, P. 1988. Direct protein microsequencing from Immobilon-P transfer membrane. BioTechniques 6:154-159. Mozdzanowski, J., Hembach, P., and Speicher, D. 1992. High yield electroblotting onto polyv inylid ene d ifluo ride membranes from polyacrylamide gels. Electrophoresis 13:59-64. Patterson, S.D., Hess, D., Yungwirth, T., and Aebersold, R. 1992. High-yield recovery of electroblotted proteins and cleavage fragments from a cationic polyvinylidene fluoride–based membrane. Anal. Biochem. 202:193-203. Speicher, D.W. 1989. Microsequencing with PVDF membranes: Efficient electroblotting, direct protein adsorption and sequencer program modifications. In Techniques in Protein Chemistry (T. Hugli, ed.) pp. 24-35, Academic Press, San Diego. Szewczyk, B. and Summers, D.F. 1988. Preparative elution of proteins blotted to Immobilon membranes. Anal. Biochem. 168:48-53. Towbin, H., Staehelin, T., and Gordon, J. 1979. Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: Procedure and some applications. Proc. Natl. Acad. Sci. U.S.A. 76:4350-4354.
Vandekerckhove, J., Bauw, G., Puypi, M., Van Damme, J., and Van Montagu, M. 1985. Proteinblotting on Polybrene-coated glass-fiber sheets. Eur. J. Biochem. 152:9-19.
Key References LeGendre, N. and Matsudaira, P. 1988. See above. Thoroughly reviews electroblotting using PVDF membranes. Mozdzanowski, J. and Speicher, D.W. 1992. Microsequence analysis of electroblotted proteins. I. Comparison of electroblotting recoveries using different types of PVDF membranes. Anal. Biochem. 207:11-18. Compares electroblotting recoveries using different PVDF membranes.
Contributed by Jeanine A. Ursitti, Jacek Mozdzanowski, and David W. Speicher The Wistar Institute Philadelphia, Pennsylvania
Electroblotting from Polyacrylamide Gels
10.7.14 Current Protocols in Protein Science
Detection of Proteins on Blot Membranes
UNIT 10.8
Staining of blot transfer membranes permits visualization of proteins and allows the extent of transfer to be monitored. In the protocols described in this unit, proteins are stained after electroblotting from one-dimensional or two-dimensional polyacrylamide gels to blot membranes such as polyvinylidene difluoride (PVDF), nitrocellulose, or nylon membranes (UNIT 10.7). PVDF is the preferred, more universal membrane and is emphasized here; however, most stains work similarly on nitrocellulose, and many can be used on alternative blotting membranes. The first six Basic Protocols describe the use of six general protein stains—amido black, Coomassie blue, Ponceau S, colloidal gold, colloidal silver, and India ink. In addition, the fluorescent stains fluorescamine and IAEDANS, which covalently react with bound proteins, are described in Basic Protocol 7 and the Alternate Protocol. Table 10.8.1 lists approximate detection limits for each nonfluorescent stain as well as membrane compatibilities. The dimensional stability of blotted membranes facilitates direct, precise comparisons of staining patterns using different detection methods. For example, protein staining can be directly compared with immunoreactivity by staining one portion of a blot with a general protein stain while subjecting another portion containing duplicate samples to immunoblotting. When it is desirable to cut replicate lanes from the blot prior to any staining, prestained standards provide very useful visual reference points. NOTE: High-purity water (from a Milli-Q purification system or equivalent) should be used throughout the protocols, and all plastic and glass boxes must be thoroughly cleaned before use to avoid staining artifacts. Membranes should be handled only by the edges with forceps. All steps should be performed at room temperature (unless otherwise described) and with gentle agitation. Use of an orbital shaker is recommended for steps that take longer than 1 min. If a PVDF membrane is allowed to dry after transfer, wet for 5 sec in 100% methanol and rinse with water before staining. Volumes of stain, destain, and wash solutions should be sufficient to cover the membrane and allow it to float freely. Unless noted otherwise, solutions may be stored for several months at room temperature.
Table 10.8.1 Staining Sensitivities and Membrane Compatibilities for Nonfluorescent Stainsa
Stain Amido black Coomassie blue Ponceau S Colloidal gold Colloidal silver India ink
Membrane typec Minimum amount detectedb PVDF Nitrocellulose Nylon 50 ng 50 ng 200 ng 2 ng 5 ng 5 ng
+ + + + + +
+ − + + + +
+ + + − − −
aSensitivity of fluorescent stains is very dependent on protein amino acid composition. bMinimum amount detected based on amount of protein loaded onto gel. The actual amount
on the blot will be slightly lower because of losses during electrotransfer. Values are based on use of a full-sized gel (11 cm × 16 cm × 1.5 mm). Sensitivity will be ∼2 to 5 times higher when minigels (8 cm × 10 cm × 1.0 mm) are used because the protein bands are concentrated on a smaller area of membrane. c+ indicates stains well; − indicates membrane not compatible.
Electrophoresis Contributed by Sandra Harper and David W. Speicher Current Protocols in Protein Science (1995) 10.8.1-10.8.7 Copyright © 2000 by John Wiley & Sons, Inc.
10.8.1 CPPS
BASIC PROTOCOL 1
AMIDO BLACK STAINING Amido black is used to stain proteins on blot transfer membranes. Transferred proteins (>50 ng/band) appear as dark blue bands on a light blue background. Amido black has a sensitivity similar to that of Coomassie blue, but it stains faster. It is the preferred stain for protein sequencing and in situ cleavage of proteins for determining internal sequences because the mild staining and destaining conditions minimize the likelihood that any protein will be extracted during the procedures. Materials Protein sample electroblotted to PVDF, nitrocellulose, or nylon membrane (UNIT 10.7) Amido black 10B stain: 0.1% (w/v) amido black (naphthol blue black 10B, Sigma) in 10% (v/v) acetic acid 5% (v/v) acetic acid Plastic boxes 1. Place blot transfer membrane in a plastic box and wash with water three times for 5 min each. 2. Stain membrane with amido black 10B stain for 1 min. Longer staining times only increase background and hence decrease sensitivity.
3. Destain the membrane with 5% acetic acid twice for 1 min each. 4. Rinse the membrane with water twice for 10 min each, then air dry. BASIC PROTOCOL 2
COOMASSIE BLUE R-250 STAINING Coomassie blue R-250 can be used with most types of blot membranes except nitrocellulose (high concentrations of organic solvents can dissolve nitrocellulose membranes). Coomassie blue has a similar sensitivity to amido black. Coomassie blue–stained proteins (>50 ng/band) appear as dark blue bands against a light blue background. The sequence of washing with water, staining, and destaining is similar to that for amido black staining (Basic Protocol 1), but the staining step is lengthened to 5 min. Materials Blot transfer membrane (UNIT 10.7) Coomassie blue stain: 0.025% (w/v) Coomassie brilliant blue R-250 (Bio-Rad) in 40% methanol/7% acetic acid (v/v) 50% methanol/7% acetic acid (v/v) Plastic box 1. Place blot transfer membrane in a clear plastic box. Wash with water three times for 5 min each. 2. Stain membrane with Coomassie blue stain for 5 min. If proteins on PVDF membranes are to be subjected to N-terminal sequencing, acetic acid should be omitted from the staining and destaining solutions to minimize protein extraction.
3. Destain membrane with 50% methanol/7% acetic acid for 5 to 10 min. 4. Rinse with water several times, then air dry. Detection of Proteins on Blot Membranes
10.8.2 Current Protocols in Protein Science
PONCEAU S STAINING Ponceau S is the least sensitive general protein stain described here. Transferred proteins (>200 ng/band) appear as red bands on a pink background. Major advantages of Ponceau S staining are that it is simple, rapid, and reversible. If desired, essentially all of the stain can be removed by extended destaining as described in steps 4 to 6. This can be particularly advantageous if the blot is to be reused after initial protein staining for a second detection method such as immunoblotting.
BASIC PROTOCOL 3
Materials Blot transfer membrane (UNIT 10.7) Ponceau S stain: 0.5% (w/v) Ponceau S (Sigma) in 1% (v/v) acetic acid 200 µM NaOH/20% (v/v) acetonitrile Plastic box 1. Place blot transfer membrane in a plastic box and wash with water three times for 5 min each. 2. Stain membrane with Ponceau S stain for 30 sec to 1 min. 3. Destain membrane with several changes of water for 30 sec to 1 min each, then air dry. If the stain is to be extracted from protein bands (steps 4 to 6) prior to employing a second detection method, omit drying. Do not overdestain because the protein bands will be difficult to detect. Stop destaining when the background has a very slight pink tinge.
4. Make a permanent record of the staining pattern by photocopying or photographing the blot. Alternatively, mark the positions of the bands of interest directly on the blot by overlaying the wet blot with plastic wrap or a plastic bag and using a pencil or pen to outline the bands. Press with moderate pressure to make a permanent indentation on the membrane. 5. Extract Ponceau S stain from the protein bands with 200 µM NaOH/20% acetonitrile for 1 min. 6. Wash membrane with water three times for 5 min each, then air day. COLLOIDAL GOLD STAINING Colloidal gold is a highly sensitive stain. AuroDye (Amersham) is a ready-to-use commercial source of colloidal gold stain in a low-pH buffer. AuroDye should be used without dilution. Transferred proteins (>2 ng/band) will appear as red bands on a pink background. A higher signal may be obtained with alkali treatment of the membrane prior to staining by washing the membrane with 1% KOH followed by several rinses with phosphate-buffered saline (PBS). Glass rather than plastic boxes must be used to hold the membranes and solutions. Glass is easier to clean than plastic and is less likely to give artifacts. Ideally, a set of glass trays should be dedicated to the procedure.
BASIC PROTOCOL 4
Materials Blot transfer membrane (UNIT 10.7) Tween 20 solution: 0.3% (v/v) Tween 20 in PBS (APPENDIX 2E; prepare solution fresh weekly and store at 4°C) AuroDye colloidal gold reagent (Amersham; store at 4°C) Glass box Electrophoresis
10.8.3 Current Protocols in Protein Science
1. Place blot transfer membrane in a glass box. Wash with water three times for 5 min each. 2. Incubate membrane with Tween 20 solution for 30 min at 37°C with gentle agitation. 3. Wash membrane with Tween 20 solution three times for 5 min each at room temperature. 4. Rinse membrane several times with water. 5. Stain membrane with AuroDye for 2 to 6 hr (until desired color formation). Use only enough stain to cover the membrane completely.
6. Rinse membrane thoroughly with water, then air dry. BASIC PROTOCOL 5
COLLOIDAL SILVER STAINING Colloidal silver is a more economical stain than colloidal gold. The staining procedure is rapid, although the ferrous sulfate solution must be prepared immediately before use. Transferred proteins (>5 ng/band) appear as black bands on a light brown background for nitrocellulose and on a dark background for PVDF. Materials Blot transfer membrane (UNIT 10.7) 40% (w/v) sodium citrate (store at 4°C for several months) 20% (w/v) ferrous sulfate (FeSO4⋅7H2O), prepared fresh 20% (w/v) silver nitrate (store at 4°C for several months) Glass box 1. Place blot transfer membrane in a glass box and wash with water three times for 5 min each. 2. Add 5 ml of 40% sodium citrate and 4 ml of 20% ferrous sulfate to 90 ml water. Stir vigorously and slowly add 1 ml of 20% silver nitrate over ∼1 to 2 min to form a suspension. 3. Immediately use the suspension to stain the membrane for ∼5 min. The staining suspension should be used within 30 min.
4. Rinse membrane with water, then air dry. BASIC PROTOCOL 6
INDIA INK STAINING India ink is used to stain electroblotted proteins on blot transfer membranes. Transferred proteins (>5 ng/band) appear as black bands on a gray background. Sensitivity may be enhanced by brief alkali treatment of the membrane with 1% KOH followed by several rinses with PBS. Materials Blot transfer membrane (UNIT 10.7) Tween 20 solution: 0.3% (v/v) Tween 20 in PBS (APPENDIX 2E; prepare solution fresh weekly and store at 4°C) India ink solution: 0.1% (v/v) India ink (Pelikan 17 black) in Tween 20 solution (store 1 month at room temperature) Plastic box
Detection of Proteins on Blot Membranes
1. Place blot transfer membrane in a plastic box. Wash with water three times for 5 min each.
10.8.4 Current Protocols in Protein Science
2. Wash membrane with Tween 20 solution four times for 10 min each. 3. Stain membrane with India ink solution for 2 hr or overnight. 4. Rinse with water until an acceptable background is obtained, then air dry. FLUORESCAMINE LABELING Fluorescamine, or 4-phenylspiro[furan-2(3H),1′-phthalan]-3,3′-dione, is used to introduce a fluorescent label on electroblotted proteins via reaction with free amines. Transferred proteins are visualized on blot transfer membranes with UV light. This stain can be very sensitive and can be used in conjunction with a second detection method such as immunoblotting (also see Basic Protocol 3). However, the protein is irreversibly modified because fluorescamine reacts with available amino groups (i.e., lysines and the protein N terminus if it was not previously blocked).
BASIC PROTOCOL 7
Materials Blot transfer membrane (UNIT 10.7) Sodium bicarbonate solution: 100 mM sodium bicarbonate in 0.3% (v/v) Tween 20, pH 9.0 (prepare fresh weekly and store at 4°C) Fluorescamine stain: 0.25 mg/ml fluorescamine (Sigma) in sodium bicarbonate solution (prepare fresh daily) Plastic box 1. Place blot transfer membrane in a plastic box. Wash with water three times for 5 min each. 2. Wash membrane with sodium bicarbonate solution twice for 10 min each. 3. Label protein bands with fluorescamine stain for 15 min. Use enough staining solution to cover the membrane completely. 4. Wash membrane with bicarbonate solution three times for 5 min each. 5. Rinse membrane several times with water. 6. Visualize transferred proteins with UV light. IAEDANS LABELING N-iodoacetyl-N′-(5-sulfo-1-naphthyl)ethylenediamine (IAEDANS or 1,5-I-AEDANS) is used for fluorescent labeling of electroblotted proteins on blot transfer membranes. Because IAEDANS reacts with free cysteines, disulfides in the sample must first be reduced with dithiothreitol (DTT). Transferred protein bands are visualized under UV light.
ALTERNATE PROTOCOL
Additional Materials (also see Basic Protocol 7) DTT solution: 200 mM dithiothreitol (DTT) in 100 mM Tris⋅Cl, pH 8.6 (APPENDIX 2E; prepare immediately before use) 100 mM Tris⋅Cl, pH 8.6 (APPENDIX 2E) N-iodoacetyl-N′-(5-sulfo-1-naphthyl)ethylenediamine (IAEDANS; Sigma; store desiccated in the dark at −20°C) Glass box 1. Place blot transfer membrane in a glass box. Wash with water three times for 5 min each. 2. Incubate membrane with DTT solution for 30 min to reduce disulfides in the sample. 3. Wash membrane with 100 mM Tris⋅Cl (pH 8.6) three times for 5 min each. Electrophoresis
10.8.5 Current Protocols in Protein Science
This washing step probably could be deleted if proteins have been electroblotted from a reducing gel (i.e., containing DTT and/or 2-mercaptoethanol). However, it is advisable to include this simple step routinely as some oxidation may occur during electrotransfer or on the blotted membrane itself during drying or storage.
4. Dissolve 86 mg IAEDANS (2 mM final concentration) in 100 ml of 100 mM Tris⋅Cl (pH 8.6). This step should be carried out in the dark (solution should be made fresh daily).
5. Add IAEDANS solution to the membrane and allow the reaction to proceed in the dark for 30 min with shaking. 6. Wash membrane with 100 mM Tris⋅Cl (pH 8.6) twice for 5 min each. Thoroughly rinse membrane with water to remove excess reagent. 7. Visualize the transferred proteins under UV light. COMMENTARY Background Information
Detection of Proteins on Blot Membranes
The recent rapid expansion in the use of electrophoretic transfer of separated proteins to different types of membranes has necessitated the adaptation of existing protein staining techniques to transfer membranes. On-blot staining techniques serve multiple purposes, including detection of proteins for structural analysis and use in parallel with antibody reactivity to correlate precisely immunoreactivity with protein staining patterns. For the latter purpose an advantage of on-blot staining is that duplicate sections of membrane can be cut out and one section used for immunoblotting (UNIT 10.10) while a second section with duplicate lanes is stained with a general protein stain. The two pieces can then be precisely realigned. In contrast, a direct comparison of an immunoblot with a stained polyacrylamide gel is much less precise because of shrinking and swelling of the gel. The most common membranes used for electroblotting are polyvinylidene difluoride (PVDF) and nitrocellulose. PVDF membranes have become increasingly popular because they are easy to handle and store, whereas nitrocellulose membranes are brittle and tend to break easily when dry. A number of PVDF membranes are now commercially available that have subtle but important differences in protein binding properties resulting from different proprietary manufacturing processes. For example, BioRad Trans-Blot or Millipore Immobilon-PSQ PVDF membranes generally show higher protein binding capacities and higher binding affinities compared to Immobilon-P (Mozdzanowski and Speicher, 1992). Use of high-retention PVDF membranes usually results in higher and more consistent electroblotting recoveries
of most proteins than the use of either low-retention PVDF membranes or nitrocellulose. However, high-retention PVDF membranes tend to exhibit higher staining backgrounds, and it is more difficult to extract proteins or peptides from such membranes. The protocols for staining with amido black, Coomassie blue, Ponceau S, and AuroDye follow the suppliers’ recommendations. It should be noted that when staining PVDF membranes with Coomassie blue before N-terminal sequencing, omitting acetic acid from both the stain and destain solution is recommended to minimize potential extraction of protein from the membrane (Speicher, 1989). The protocol for India ink staining of electroblotted proteins is essentially that of Hancock and Tsang (1983). Different brands of India ink may be used, but staining sensitivity may vary as a result. The colloidal gold stain is the most sensitive membrane stain described here. As an alternative to commercially available colloidal gold stains, Moeremans et al. (1985) describe methods for preparing gold and iron solutions. Fluorescent labels are advantageous because they can be used not only for sequential detection methods on the same blot with minimal potential interference, but also for detection prior to protein extraction from the membrane. For example, after visualization of proteins with a fluorescent label, the blot can be photographed and specific bands marked with a pencil, either directly on the membrane or through a plastic bag. The latter method leaves a permanent indentation on the membrane. The blot can then be probed with antisera (i.e., immunoblotted; UNIT 10.10).
10.8.6 Current Protocols in Protein Science
The protocol for fluorescamine labeling is based on the procedure described by Vera and Rivas (1988). Fluorescamine itself is not strongly fluorescent; however, when combined with the primary amines of proteins (i.e., N termini and lysine residues) it yields a highly fluorescent product. IAEDANS is an iodoacetic acid analog containing a naphthalene ring and is fluorescent under UV light. When the sample protein is reduced either prior to gel electrophoresis or with DTT after blotting, all cysteines are potentially available for reaction with IAEDANS. When the protein is not reduced, some cysteines may be involved in disulfide bonds and therefore not available for reaction. Additionally, some cysteines may be sterically inaccessible because of adsorption to the membrane and therefore will not react. An additional visualization technique for PVDF membranes is transillumination, described by Reig and Klein (1988). In that technique, the membrane is dried at room temperature, then wet with 20% methanol and viewed on a white light box. Protein bands appear as clear areas. Sensitivity is usually comparable to that of Coomassie blue staining.
Critical Parameters and Troubleshooting High-quality water (from a Milli-Q purification system or equivalent) should be used throughout these protocols. All plastic and glass boxes must be thoroughly cleaned by rinsing with water before use to avoid staining artifacts. Blot membranes should be handled by the edges only with gloves or, preferably, with forceps. This precaution is most critical for the more sensitive stains (i.e., colloidal gold, colloidal silver, and India ink). When using PVDF membranes, it is especially critical that the membrane does not dry between steps. If drying occurs, wet the PVDF membrane for 5 sec with 100% methanol, then rinse several times with water. A brief alkali treatment can enhance staining with India ink or colloidal gold. In a procedure described by Sutherland and Skerritt (1986), the membrane is washed with 1% (w/v) KOH for 5 min followed by several rinses with PBS. The alkali treatment can easily be incorporated at the beginning of the procedures if desired.
lamide in the gel used in preparation of the blot transfer membrane. The sensitivity of fluorescent stains is related to the number of reactive amino groups (fluorescamine) or cysteine residues (IAEDANS) present in the protein of interest.
Time Considerations The total time required for staining with amido black, Coomassie blue, and Ponceau S is 30 min to 1 hr; colloidal gold requires 4 to 6 hr; colloidal silver requires 1 hr; whereas staining with India ink requires 2 hr to overnight. Fluorescent labeling requires ∼1 hr. The optional alkali enhancement (see Critical Parameters and Troubleshooting) requires an additional 30 min at the beginning of the India ink and colloidal gold staining procedures.
Literature Cited Hancock, K. and Tsang, V.C.M. 1983. India ink staining of proteins on nitrocellulose paper. Anal. Biochem. 133:157-162. Moeremans, M., Daneels, G., and De Mey, J. 1985. Sensitive colloidal metal (gold or silver) staining of protein blots on nitrocellulose membranes. Anal. Biochem. 145:315-321. Mozdzanowski, J. and Speicher, D.W. 1992. Microsequence analysis of electroblotted proteins. Anal. Biochem. 207:11-18. Reig, J. and Klein, D. 1988. Submicrogram quantities of unstained proteins are visualized on polyvinylidene difluoride membranes by transillumination. Appl. Theor. Electrophor. 1:59-60. Speicher, D.W. 1989. Microsequencing with PVDF membranes: Efficient electroblotting, direct protein adsorption and sequencer program modifications. In Techniques in Protein Chemistry (T.E. Hugli, ed.) pp. 24-35. Academic Press, San Diego. Sutherland, M.W. and Skerritt, J.H. 1986. Alkali enhancement of protein staining on nitrocellulose. Electrophoresis 7:401-406. Vera, J.C. and Rivas, C. 1988. Fluorescent labeling of nitrocellulose-bound proteins at the nanogram level without changes in immunoreactivity. Anal. Biochem. 173:399-404.
Key References Moeremans, M., Daneels, G., and De Mey, J. 1985. See above. Describes a method for preparation of colloidal metal stains. Vera, J.C. and Rivas, C. 1988. See above. Describes the use of multiple detection methods.
Anticipated Results Approximate detection limits and membrane compatibilities are listed in Table 10.8.1. It should be noted that detection limits may vary with gel size and the percentage of polyacry-
Contributed by Sandra Harper and David W. Speicher The Wistar Institute Philadephia, Pennsylvania
Electrophoresis
10.8.7 Current Protocols in Protein Science
Supplement 5
Capillary Electrophoresis of Proteins and Peptides
UNIT 10.9
Capillary electrophoresis (CE) is a high-resolution technique for the separation of a wide variety of molecules of biological interest such as metabolites, drugs, amino acids, nucleic acids, and carbohydrates. This unit focuses on the use of CE to separate proteins and peptides. As with polyacrylamide gel electrophoresis (PAGE; UNIT 10.1), CE separations of proteins and peptides are based on charge-to-mass ratios. While PAGE separations are restricted to polyacrylamide matrices and a relatively small number of buffer systems; CE separations can be achieved in a variety of different matrices using a wide range of electrophoresis buffers. Consequently, there is a much greater flexibility in the design of optimal separation protocols. CE employs a fused-silica capillary column, which may or may not be derivatized, either in free solution or in the presence of a fluid matrix. In contrast to PAGE, the resulting protein or peptide bands cannot be fixed or stained. The CE separation is, in essence, a dynamic one and is more analogous to high-performance liquid chromatography (HPLC) than PAGE. However, CE separation is based on electrophoretic parameters, so it can be viewed as orthogonal to HPLC, which is based primarily on solvent partitioning principles. The combination of these two techniques is very powerful for analyzing and characterizing proteins and peptides. The initial expectation that CE would play a major role in protein separations has not been realized. However, it is particularly useful for determining the purity of a sample and for rapid, efficient, and quantitative evaluation of protein purification techniques. For best separation results, it is necessary to optimize the procedure. If some properties of the protein are known—e.g., its isoelectric point (pI)—the selection of a suitable running buffer is relatively straightforward. In fact, CE itself can be used to determine the isoelectric point of a protein, either in purified form or in a mixture, by focusing the sample in a pH gradient that is generated within the capillary during electrophoresis (Basic Protocol 1). After refocusing, the sample is mobilized either by changing the anode buffer, or by applying a hydrodynamic force to the column to move the separated components past the detector. However, there is little information available that will predict the potential of a protein to interact with the capillary wall. Underivatized silica capillaries have a strong tendency to adsorb proteins, thereby immobilizing them and making protein separations impractical. If such interactions occur, then an additive or a coated capillary column is required. A certain amount of trial and error is involved, and it may be necessary to perform several CE runs to optimize the separation conditions for a given protein. One approach to optimization is described in Basic Protocol 2. CE is most useful for separations of peptides (Basic Protocol 3) because it offers great flexibility in separation parameters. It can be used to monitor proteolytic digestions and optimize digestion conditions for the production of a representative peptide fingerprint of a protein. This profile can subsequently be used to provide structural information about the protein, especially when used in conjunction with reversed-phase HPLC (RP-HPLC). It can be used to screen peptide fractions that are obtained from a preparative RP-HPLC separation of a protease digestion. Fractions that contain single or major components are suitable candidates for protein sequence analysis. Similarly, CE can be used to assess the purity of synthetic peptides. In the presence of an internal standard, it can provide quantitative information about the various components that are present in a peptide mixture. CE can also be used as a micropreparative technique—with either multiple separations that are pooled (Basic Protocol 4) or a single, larger-scale separation (AlterContributed by Dean Burgi and Alan J. Smith Current Protocols in Protein Science (1995) 10.9.1-10.9.13 Copyright © 2000 by John Wiley & Sons, Inc.
Electrophoresis
10.9.1 Supplement 2
nate Protocol 1)—for the isolation of peptides from a protease digestion (in much the same way that RP-HPLC is currently used). In most of these examples the same capillary column can be used for all the separations. Only changes in buffer composition, ionic strength, and the presence or absence of additives are required for each specific application. INSTRUMENTATION CE separation in its simplest form can be achieved by passing a high voltage between two buffer reservoirs that are joined by a liquid-filled fused silica capillary (Fig. 10.9.1). This results in the generation of electroosmotic flow (EOF; see Separation Theory), which allows the molecules of interest to be carried from one end of the capillary to the other. The capillaries are generally 30 to 50 cm long with 50 to 75 µm i.d. The net total volume of these capillaries is in the low microliter range. For comparison, the volume of a slab gel lane is ∼1000 µl. The capillaries are thin walled; this allows rapid and efficient exchange of the Joule heating resulting from the high voltages (20 to 25 kV) that are necessary for electrophoretic separations. This heat dissipation minimizes the negative convective effects that could result in band broadening during electrophoresis. The fused-silica capillary is coated on the outside with a polyimide layer that confers excellent tensile strength to the otherwise fragile capillary. The polyimide sheathing is carefully burned from a small portion of the capillary to expose a section of the silica. This clear section of the capillary is inserted into the light path of a UV detector and becomes the flow cell. As the protein and peptide molecules are swept through the capillary by EOF, they pass through the detector light path and are registered on the UV monitor. In effect the capillary becomes a very-low-volume flow cell. IMPORTANT NOTE: The removal of the polyimide coating makes the capillaries susceptible to breakage. Capillaries that are not provided in cartridges by the manufacturer should be handled with care to avoid breakage. The combination of high field strength and large surface-to-volume ratios results in rapid and very efficient separations of both proteins and peptides. Sample loading volumes are routinely in the nanoliter range, with starting sample concentrations of ∼0.1 µg/µl for UV
Capillary Electrophoresis of Proteins and Peptides
Figure 10.9.1 Schematic of a CE instrument.
10.9.2 Supplement 2
Current Protocols in Protein Science
Table 10.9.1
Commercially Available CE Instruments
Capillary Cooling of heating sample/buffer
Capillary format
Software available
UV: 190-700 nm Controlled by software
FAC
Sample
Freehanging
DA
Manual UV: filter; Diode-array: 190-600 nm; Fluorescence: argon laser; MS adapters UV: 190-800 nm Controlled by software
Liquid (Peltier)
Sample/buffer
Cartridge
C, DA
Liquid
Sample/buffer
Cartridge
C, DA
FAC (Peltier) FAC (Peltier)
Sample/buffer
Cartridge
C, DA
Sample/buffer
Freehanging
DA
Manufacturer
Model
Detection method
Applied Biosystems
270-HT
Beckman
P/ACE 5000 P/ACE 5510 P/ACE 5510
Bio-Rad
Biofocus 3000
Hewlett-Packard HP 3D CE Waters
Diode-array: 190-600 nm Quantra 4000E UV: filter
Reversal of polarity
Controlled by software Manual
aAll instruments have hydrostatic/electrokinetic sample loading and fraction collecting. bAbbreviations: C, CE system control software; DA, data acquisition; FAC, forced-air convection.
detection. On-capillary preconcentration protocols are available when starting concentrations are below these levels. Clearly, with respect to sensitivity, speed, and versatility, CE can offer some significant advantages over gel electrophoresis for the separation of proteins and peptides. As shown in Figure 10.9.1, CE can require minimal instrumentation. However, in reality, due to the high voltages that are utilized, safety issues require that the capillary column be incorporated into a dedicated CE instrument. These instruments can provide efficient capillary cooling and online detection of analytes. Table 10.9.1 lists a number of commercially available instruments and some of their characteristics. For the purpose of this discussion it is assumed that a generic CE system possessing the following capabilities is used: an active capillary cooling system, a UV detector capable of monitoring at 200 nm, a thermostatted autosampler, and a chromatographic data package. Given this basic unit a variety of separations can be performed, depending upon the nature of the proteins or peptides that are being separated. SEPARATION THEORY CE is part of the family of electrophoretic techniques that separate species based upon their size and ionic properties. An ion (i) placed into an electric field will move in the direction parallel to the field with a velocity (vi) as follows: vi = µi E
Here µi is the electrophoretic mobility of species i and E is the electric field, as defined by the following equation, where V is the voltage and L is the total length from electrode to electrode: E=
V L Electrophoresis
10.9.3 Current Protocols in Protein Science
Supplement 2
The electrophoretic mobility of an ion (µi) is: µi =
q 6 π η ai
where q is the charge on the ion, η is the viscosity of the solution, and ai is the radius of the ion. As seen from these three equations, the movement of the molecule in the column is dependent on the applied voltage, the length of the column, the charge on the molecule, and the size of the molecule. In CE, buffer flow is generated inside the column when the electric field is applied. This flow is from the cathode electrode to the anode electrode in a fused-silica capillary column (from left to right in Fig. 10.9.1); this movement of the buffer is called the electroosmotic flow (EOF). In addition, normal electrophoretic separation occurs whereby a positively charged molecule moves in the same direction as the EOF, a negatively charged molecule moves in a direction opposite to the flow, and a neutral molecule is carried along by the flow. Thus, the total velocity of the molecule (µt) is given by the following equation, where µeo is the EOF in the column. µt = ( µeo + µi ) E
The magnitude of the EOF will vary as a function of the pH of the carrier buffer. In uncoated fused-silica columns, a low pH generates a slow EOF, whereas a high pH generates a fast flow. The flow can be suppressed or completely reversed depending on the coating on the column or organic modifier added to the carrier buffer (Landers et al., 1992; Tsuiji and Little, 1992). Thus, EOF is another parameter that can be optimized to aid in difficult separations. Neutral molecules, because they are all carried along by the flow, can be difficult to separate. Addition of a micelle to the carrier buffer results in the partitioning of molecules between the micelle and carrier buffer and can effectively resolve neutral molecules (Khaledi, 1994). STRATEGIC PLANNING Proteins can be separated on coated or uncoated capillary columns, and the choice of separation protocol depends on the specific properties of the target protein. The most important property is the pI, which can be determined by either conventional gel electrophoresis or CE. The use of a buffer ∼2 pH units above the pI is optimal for CE separations. Table 10.9.2 lists a number of buffers and their pH ranges. The presence of high salt concentrations in the sample can interfere with the separation process, so dialysis or dilution may be required. Protein concentrations of microgram per microliter are optimal for CE separations; however, on-capillary concentration protocols exist that allow the separation of proteins with concentrations 1 to 2 orders of magnitude lower.
Capillary Electrophoresis of Proteins and Peptides
Peptides can be effectively separated in open-tube fused-silica capillary columns. The EOF generated within the capillary causes separation of both charged and neutral peptides. The respective migration times are dependent upon both the pH of the electrophoretic separation and the charge-to-mass ratios of the peptides. For example, at low pH, peptides with net positive charge migrate towards the anode faster than neutral or negatively charged peptides. Even negatively charged peptides are swept towards the anode by EOF. In practice this migration order is further modified by the mass of the peptides, with small positively charged peptides having faster migration rates. The choice of buffer for CE is of primary importance. The use of buffer systems that separate at high pH significantly alters the migration positions of peptides relative to those
10.9.4 Supplement 2
Current Protocols in Protein Science
Table 10.9.2
Useful Electrolytes for CE of Peptides
Electrolyte
Useful pH range
Minimum useful wavelength (nm)
Phosphate Citrate Acetate MES Phosphate Tris Borate
1.14-3.14 3.06-5.40 3.76-5.76 5.15-7.15 6.20-8.20 7.30-9.30 8.14-10.14
195 260 220 230 195 220 180
of low-pH separations. The selection of a specific buffer is dictated both by its buffering capacity at a selected pH and by its minimum usable UV absorbance wavelength. Characteristics of some useful buffer systems are shown in Table 10.9.2. Small differences in the pH or composition of the buffer can have a significant impact on the absolute mobilities of the peptides, so it is advisable to adjust the pH of buffers accurately and reproducibly. Alternatively, a suitable internal standard can be incorporated into the separation in order to obtain relative mobilities. For the separation of peptides from a tryptic digest of a protein, where almost every peptide carries at least two positive charges, a phosphate buffer of pH 2.0 to 3.0 is commonly used. Likewise, for V8 protease digestions (which cleaves C-terminal to glutamic acid residues), the preferred separation conditions use a borate buffer with a pH range of 8.0 to 9.0. SEPARATION OF PROTEINS BY ISOELECTRIC FOCUSING Isoelectric focusing (IEF) separations can be performed readily by CE; this can be a useful first step in selecting subsequent buffer systems for protein separations. Proteins and peptides are amphoteric in nature, so their charge is dictated by the surrounding carrier buffer. When a protein or peptide is placed in an electric field, it moves to a region where the surrounding pH equals its isoelectric point (pI); at this point the molecule stops moving. A pH gradient is generated in a coated capillary column by filling the column with a sample solution that contains ampholytes. A high-pH solution (sodium hydroxide) is placed in the cathode reservoir and a low-pH solution (phosphoric acid) is placed in the anode reservoir. An electric field is then applied across the column and the ampholytes and the sample move to a location in the column corresponding to their respective pIs. If the molecule drifts out of the isoelectric region, a charge is induced by the surrounding pH solution and the molecule moves back to the region of zero charge. Narrow regions are formed with pI resolutions on the order of 0.01 pH units. To determine the pI location of a particular protein in the column, markers of known pIs are added to the sample mixture, and extrapolation between markers yields the pI of the protein of interest.
BASIC PROTOCOL 1
After separation of the sample in the absence of EOF, mobilization of the proteins past the detector is necessary. A simple replacement of the anode reservoir with the buffer in the cathode reservoir establishes EOF and causes the sample zone to migrate pass the detector. The separation discussed here assumes there is no EOF in the column during focusing. However, separations may also be done in the presence of EOF (Mazzeo and Krull, 1992).
Electrophoresis
10.9.5 Current Protocols in Protein Science
Supplement 2
Materials Ampholyte mixture, pH 3 to 10 (Bio-Rad) Sample containing 0.5 to 1.0 mg/ml protein IEF markers (optional) 20 mM sodium hydroxide (NaOH; store at 4°C) 10 mM phosphoric acid (store at 4°C) 50-µm-i.d. coated silica capillary column CE instrument (see Table 10.9.1) 1. Add ampholyte mixture to 0.5 ml protein sample to give a final concentration of 2.5% (v/v) ampholytes. If desired, add IEF markers to a final concentration of 0.1 mg/ml to calibrate the column.
2. Add the solution to the sample reservoir. Fill the capillary by pressurizing the reservoir (0.5 psi). 3. Place 10 mM phosphoric acid in the anode reservoir (negative end). Place 20 mM NaOH in the cathode reservoir (positive end). 4. Focus the sample 4 to 6 min at 8 to 10 kV constant voltage. Monitor the current until it reaches steady state. The instrument should be operated in accordance with the manufacturer’s instructions.
5. Mobilize the sample by placing 20 mM NaOH in the anode reservoir and setting the voltage to 10 kV. Monitor the mobilization of the proteins past the detector 15 to 20 min. Another method for mobilizing the proteins past the detector is to apply a hydrodynamic force to one end of the column and push the zones past the detector. Commercially available CE instruments have this capacity as an option. To minimize distortion of the pI regions, a constant electric field (10 kV) must be applied at the same time. The pressure applied is small (0.5 psi) and is maintained for the entire time of mobilization.
6. Wash the column after each run with 10 mM phosphoric acid for 1 min at 0.5 psi. Store the column at room temperature in running buffer (short-term) or water (long-term). BASIC PROTOCOL 2
SEPARATION OF PROTEINS To use CE for protein analyses, the more that is known about the particular protein, the higher the success rate for developing a separation method. Knowing the pI of a protein permits selection of the appropriate run buffer to maximize differences in the charge-tosize ratio. IEF separation performed using CE (Basic Protocol 1) can be a useful first step in selecting subsequent buffer systems for protein separations on an underivatized capillary column. If no protein is detected, it has probably adsorbed to the silica surface of the column. Repeating the separation in the presence of an ionic detergent such as SDS will coat the protein with negative charge and prevent adsorption. However, this means that proteins will separate on the basis of size alone. Alternatively, if the pI of the protein is unknown, the separation can often be achieved by using a coated capillary column in the presence of high-pH, high-ionic-strength buffer.
Capillary Electrophoresis of Proteins and Peptides
10.9.6 Supplement 2
Current Protocols in Protein Science
Materials Sample containing 10 mg/ml protein 50 mM and 500 mM sodium borate, pH 8.0 to 9.5 (for separation of proteins with unknown pI) 5 mM and 50 mM buffer at pH >pI (for separation of proteins with known pI; see Table 10.9.2) 50-µm-i.d. coated (unknown pI) or uncoated (known pI) fused-silica capillary column CE instrument (see Table 10.9.1) For separations of proteins with unknown pI 1a. Dilute protein sample 1:10 (v/v) with 50 mM sodium borate buffer to give a final concentration of 1.0 mg/ml. Alternatively, dialyze sample (at a concentration of 1 mg/ml) in 50 mM sodium borate buffer.
2a. Fill a coated column with 500 mM sodium borate buffer. 3a. Inject the sample hydrodynamically, e.g., 3 sec at 1 psi. Consult the manufacturer’s instruction manual for the proper procedure.
4a. Separate using the following conditions: Detector wavelength: 200 nm Temperature: 25°C Run voltage: 10 kV Run time: 30 min. For separations of proteins with known pI 1b. Dilute protein sample 1:10 (v/v) with a 5 mM buffer that has a pH above the pI of the protein to induce a charge on the protein (final protein concentration = 1.0 mg/ml). Alternatively, dialyze sample (at a concentration of 1 mg/ml) in 5 mM buffer.
2b. Fill an uncoated capillary column with the same buffer at 50 mM. 3b. Inject the sample hydrodynamically, e.g., 3 sec at 1 psi. Consult the manufacturer’s instruction manual for the proper procedure.
4b. Separate using the following conditions: Detector wavelength: 200 nm Temperature: 25°C Run voltage: 25 kV Run time: 30 min. If the protein of interest is not detected, add 10 mM SDS to the run buffer to induce a charge on the molecule. If the peaks are too broad, add 0.1% (w/v) methylcellulose to coat the column and reduce adsorption of the molecule onto the wall of the column. The addition of SDS and/or methylcellulose will increase the run time of the separation.
Electrophoresis
10.9.7 Current Protocols in Protein Science
Supplement 2
BASIC PROTOCOL 3
ANALYTICAL PEPTIDE SEPARATIONS CE is particularly useful for separating specific peptides from complex mixtures such as the products of proteolytic digestion. In this procedure, the peptide mixture to be analyzed is placed in a sample vial at the cathode end of the capillary column. The anode reservoir contains running buffer. The sample is loaded into the capillary either electrokinetically or by pressure or vacuum, then the sample vial is replaced by running buffer and electrophoresis begins. The following protocol describes the use of a P/ACE 5000 CE instrument (Beckman) using low-pressure sample injection. However, other instruments (see Table 10.9.1) are capable of similar separations when operated according to the manufacturer’s instructions. Materials Peptide mixture: e.g., tryptic digest of β-lactoglobulin (Applied Biosystems) 0.05 M and 0.25 M sodium phosphate buffer, pH 2.30 (store at 4°C) 0.1 M sodium hydroxide (NaOH) 75-µm-i.d. fused-silica capillary column (Beckman) CE instrument (e.g., P/ACE 5000, Beckman, or equivalent; see Table 10.9.1) 1. Precondition the capillary by flushing with the following solutions: 10 column volumes of 100 mM NaOH at low pressure (0.5 psi) 10 column volumes water 4 column volumes of 0.25 M sodium phosphate buffer, pH 2.3. Store the column in 0.25 M sodium phosphate buffer, pH 2.3, at 25°C. When changing to a different separation buffer, equilibrate the column 4 hr with the new buffer (Strickland and Strickland, 1990). If a new capillary is being used, it is essential that this preconditioning step be performed prior to attempting separations.
2. Prepare the peptide mixture by dissolving 10 nmol (∼300 µg) in 10 ml of 0.05 M sodium phosphate buffer, pH 2.30. Freeze unused mixture in 100-µl aliquots. 3. Apply 10 to 20 nl sample using low-pressure injection for 10 sec at 0.5 psi. Consult the manufacturer’s manual for the proper procedure.
4. Separate the peptide mixture using the following conditions: Electrolyte: 0.05 M sodium phosphate buffer, pH 2.3 Detector wavelength: 200 nm Temperature: 25°C Voltage: 25 kV. Consult the manufacturer’s manual for proper operating conditions.
5. Rinse the column with the following solutions: 0.5 min with water 1.0 min with 0.1 N NaOH 1.5 min with water 1 min with 0.25 M sodium phosphate buffer, pH 2.30. Store the column in running buffer or water at room temperature. Capillary Electrophoresis of Proteins and Peptides
An example of a separation protocol for a Beckman P/ACE 5000 instrument with a 37-min run time is described in Table 10.9.3.
10.9.8 Supplement 2
Current Protocols in Protein Science
Table 10.9.3
Operating Program for Peptide Analysisa
Step
Process
Duration
Inletb
Outletb
Control summaryc
1 2 3 4 5 6 7 8 9 10 11
Set Temp Set Detector Rinse Rinse Rinse Inject Separate Rinse Rinse Rinse Rinse
— — 1.0 min 1.5 min 0.5 min 10.0 sec 30.0 min 0.5 min 1.0 min 1.5 min 1.0 min
— — 33 29 29 11 29 33 32 33 34
— — 7 7 9 9 9 7 7 7 9
Temp 25°C UV 200nm Forward: HP Forward: HP Forward: HP Low pressure Voltage 25.00 kV Forward: HP Forward: HP Forward: HP Forward: HP
aFor analysis of proteolytic digests, HPLC fractions, and synthetic peptides. bVial contents: 7, waste vial (water level just to contact capillary effluent); 9, electrolyte (50 mM sodium
phosphate buffer, pH 2.30); 11, sample; 29, electrolyte (50 mM sodium phosphate buffer, pH 2.30); 32, regeneration solution (0.1 N NaOH); 33, water; and 34, 0.25 H phosphate buffer, pH 2.30. cAbbreviation: HP, high pressure.
MICROPREPARATIVE CAPILLARY ELECTROPHORESIS: MULTIPLE SEPARATIONS
BASIC PROTOCOL 4
Although CE has been most commonly used for analytical separations, considerable interest has developed in using its high resolving power in micropreparative applications. Two basic approaches have evolved, one utilizing multiple separations and collections from an analytical capillary (Bergman and Jornvall, 1992) and the other utilizing a single separation and collection on a much larger-diameter (150-µm-i.d.) capillary (see Alternate Protocol). For the multiple collection approach to be effective, the elution times of the peptides must be reproducible. The following method uses the P/ACE 5000 capillary electrophoresis instrument to separate a mixture of peptides. Materials Peptides: ACTH 4-100, angiotensin I, and angiotensin II (Sigma) 0.05 mM and 0.25 mM sodium phosphate buffer, pH 2.30 (store at 4°C) 0.1 M sodium hydroxide (NaOH) 75-µm-i.d. fused-silica capillary column (Beckman) CE instrument (e.g., P/ACE 5000, Beckman, or equivalent; see Table 10.9.1) Conical microvials (Beckman) 1. Precondition the capillary column by flushing with the following solutions at low pressure (0.5 psi): 10 column volumes of 0.1 M NaOH 10 column volumes of water 4 column volumes of 0.25 M sodium phosphate buffer, pH 2.3. Store in 0.25 M sodium phosphate, pH 2.30. When changing to a different separation buffer, equilibrate the column 4 hr with the new buffer (Strickland and Strickland, 1990).
2. Prepare a peptide mixture by dissolving 0.2 mg each of ACTH 4-10, angiotensin I, and angiotensin II in 1 ml of 0.05 M sodium phosphate buffer, pH 2.3 (final concentration = 0.6 mg/ml peptide). Store 100-µl aliquots at −20°C.
Electrophoresis
10.9.9 Current Protocols in Protein Science
Supplement 2
3. Load 10 to 20 nl peptide mixture by low-pressure injection for 10 sec at 0.5 psi. Consult the manufacturer’s manual for the proper procedure.
4. Separate the mixture using the following conditions: Electrolyte: 0.05 M sodium phosphate, pH 2.30 Detector wavelength: 200 nm Temperature: 25°C Run voltage: 25 kV Fraction size: 1 min. Consult the manufacturer’s manual for proper operating conditions.
5. Replace the standard outlet reservoir (anode) with a series of conical microvials, each containing 10 µl of 0.05 M sodium phosphate buffer, pH 2.30. Collect 3-min fractions into each of the vials for the length of the separation. 6. Repeat the injection and separation (steps 3 to 5) four times and combine the contents from the corresponding fraction numbers. It will take 2 to 3 hr to complete the fractionation.
7. Screen fractions for peptide content using the analytical separation previously described (see Basic Protocol 3). ALTERNATE PROTOCOL
MICROPREPARATIVE CAPILLARY ELECTROPHORESIS: SINGLE SEPARATION In this micropreparative protocol, a larger quantity of the peptide mixture is loaded onto a 150-µm-i.d. capillary column and the fractions are collected from a single separation (Kenny et al., 1993). For the single-collection approach the CE instrument must be able to effectively control the capillary temperature at the elevated power levels that are required for electrophoresis. Forced air cooling is generally inadequate for this purpose. Additional Materials (also see Basic Protocol 4) 4:1 (v/v) 0.5 M sodium phosphate buffer (pH 2.50)/ethylene glycol 150-µm-i.d. fused-silica capillary column (Polymicro) 1. Prepare peptide mixture and precondition the column (see Basic Protocol 4, steps 1 and 2). 2. Load 0.1 µl peptide mixture by low-pressure injection for 10 sec at 0.5 psi. Consult the manufacturer’s manual for the proper procedure.
3. Separate the peptide mixture using the following conditions: Electrolyte: 0.05 M sodium phosphate buffer, pH 2.30 Detector wavelength: 200 nm Temperature: 25°C Run voltage: 7.5 kV Fraction size: 3-min per collection vial. Consult the manufacturer’s manual for proper operating conditions.
Capillary Electrophoresis of Proteins and Peptides
4. Replace the standard outlet reservoir (anode) by a series of conical microvials each containing 10 µl of 4:1 (v/v) 0.5 M sodium phosphate buffer/ethylene glycol. Collect 3-min fractions into each vial for the length of the separation. It will take ∼2 hr to fractionate the sample.
10.9.10 Supplement 2
Current Protocols in Protein Science
5. Screen fractions for peptide content using the analytical separation protocol (see Basic Protocol 3). COMMENTARY Background Information Protein separation is one of the more difficult types of separation to perform by capillary electrophoresis (CE; Novotny et al., 1990). Each protein has it own set of optimal separation conditions, and one optimized protocol will not necessarily transfer well to the separation of a different protein. The more that is known about the properties of the molecule, the easier it will be to optimize the separation conditions. The most important property is probably the pI, because it will dictate the selection of pH and ionic strength conditions for the initial separation attempt. However, a variety of other properties must also be taken into account when optimizing a separation protocol: shape (globular or fibrous), aggregation tendencies, solubility in dilute solutions, salt requirements, thermal stability, and hydrophobic nature. Several parameters can be varied within CE separations in order to utilize this knowledge—e.g., temperature, buffer/salt selection, ionic and nonionic detergents, and matrix additives (polyacrylamide, methylcellulose). Several protocols have been developed that can serve as useful starting points for most protein separations. Table 10.9.4 lists references for some groups of proteins. Furthermore, many of the manufacturers listed in Table 10.9.1 have developed additional protocols for protein separations. Ultimately, the selection of an appropriate separation protocol for a protein depends on the specific properties of that protein. However, there are separation approaches that utilize specific properties of proteins. IEF-CE separates on the basis of charge alone; CE in the presence of ionic detergents such as SDS separates on the basis of size alone; and CE using underivatized capillaries at low pH or derivatized capillaries at high salt and high pH separates on the basis of both size and charge. Successful CE Table 10.9.4
separations also depend on the behavior of other contaminants that are present in the mixture. It may also be possible to take advantage of the knowledge gained from previous purification steps, e.g., ion-exchange or size-exclusion chromatography, to select CE separation conditions. Analytical-scale separation of peptides by CE has found application in a variety of areas. Proteolytic digestions can be monitored by placing the digestion mixture in a sample vial on the sample table that is equilibrated to 37°C. The instrument can then be programmed to analyze an aliquot at desired intervals, such as every 4 hr. Digestion is complete when a stable CE profile is obtained. Also, fractions collected from an HPLC separation of a protease digest can readily be screened orthogonally by CE to determine the homogeneity of any given peak. Fractions with a single major component can then be subjected to protein sequence analysis. The sensitivity of UV detection at 200 nm is such that digest levels of ≥100 pmol can be effectively screened by this method. Below these load levels the fractions have to be reduced in volume or preconcentrated on the capillary to achieve a sufficiently high concentration for visualization (Dolnik et al., 1990; Burgi and Chien, 1992). Selection of an appropriate micropreparative protocol will depend on the type of instrumentation that is available. The single-separation, large-diameter capacity protocol will only work with a CE instrument that has adequate cooling capabilities. The multiple-separation, analytical capillary protocol will work on any CE instrument that has reproducible and stable electrophoretic mobilities.
Critical Parameters For protein analysis, the charge on the protein is the most important parameter for sepa-
References for Separations of Different Protein Types
Protein type
Reference
Basic proteins Glycoproteins Antibody-antigen complexes Milk proteins
Wiktorowicz (1990); Bullock (1991) Tran et al. (1991); Tsuiji (1992) Nielsen et al. (1991) Chen et al. (1992)
Electrophoresis
10.9.11 Current Protocols in Protein Science
Supplement 2
ration. However, the charge-to-shape ratio also plays a role in effective separation of proteins. The charge on the protein is controlled by the buffer and the additives used for the run. The shape is determined by the denaturing conditions of the solutions. The surface charge on a protein may ultimately dictate the extent of wall interactions in addition to electrophoretic mobility, whereas shape has its greatest effect upon mobility alone. The recovery of peptides from underivatized silica capillary columns appears to be very good—even at subpicomole levels. However, proteins are frequently irreversibly bound to these same capillaries. Clearly, at some point a “peptide” must become a “protein” in the context of CE. There is little published data available to clarify this issue, although good recoveries of peptides 30-40 amino acids in length have been obtained from underivatized capillary columns. If the recoveries of higher molecular weight peptides are poor, then the separation should be repeated using a coated capillary column. During peptide synthesis, single-amino-acid-deletion products are frequently produced. The properties of these deletion products are very similar to those of the expected species, so the deletion products may not be readily resolved by reversed-phase HPLC. In this case, CE has proven to be a very useful orthogonal technique because of its high resolution power and separation based on charge-to-mass ratios. However, it must be emphasized that the effective separation and detection of proteins and peptides requires relatively high solute starting concentrations. When using uncoated columns, a sodium hydroxide rinse is required to maintain the capillary surface. However, this rinse is never used with coated capillaries because it will remove the coating. Commercially available coated columns are satisfactory but linear polymers (e.g., methylcellulose) have been used with more success (Palmieri and Nolan, 1994). If a protocol can be worked out for the molecule of interest, rapid CE analysis with little loss of starting material can augment the tools used now for protein analysis (e.g., HPLC).
Troubleshooting
Capillary Electrophoresis of Proteins and Peptides
Mechanical and electrical problems that might be encountered during instrument use are addressed in the troubleshooting sections of the manufacturers’ manuals. One of the common difficulties is the loss of electrical contact during a run. This loss of current can be caused if a small bubble forms in the column during
injection or if the power generated by the run is so great that the solution boils or outgasses. Purging the column after a failed run removes any bubbles in the column. The sample vial must contain enough sample to cover the end of the column during injection to prevent injection of air into the column. The sample vial can dry out during a long run, so sample temperature control or the use of the proper cap on the vial is required to slow evaporation. Reducing the buffer concentration or the run voltage can eliminate bubble formation during a run. Degassing the run buffer is also useful if the outgassing problems continue. The instrument sometimes arcs during a run; this is caused by salt deposits or moisture on the high-voltage electrode stations. Particular attention should be given to keeping the electrode stations salt free. Moisture buildup can be minimized by reducing the separation voltage by 5 kV. Frequent visual inspection is also advised. The column can plug up because of salt deposition if the column dries out. Washing the column with water dissolves the plug; storing the column in water if it is not going to be used for a long period of time prevents plugging. Proteins can precipitate during a run and plug the column. Using lower protein concentrations prevents this type of plugging.
Anticipated Results Electrophoretic profiles for both protein and peptide separations should be highly reproducible. However, absolute migration times may vary from run to run, especially if the separation temperature is not adequately controlled. For this reason, the use of an appropriate internal standard is highly recommended (see the manufacturer’s instructions). At load levels <50 pmol of digest, the recovery of peptides from a narrow-bore (1-mm) HPLC separation is poor. A convenient upper separation range for micropreparative CE is ∼50 pmol of digest. The use of CE as a preparative separation technique therefore complements conventional HPLC capabilities. Recoveries from micropreparative CE separations are essentially quantitative down to the low picomole level, although postseparation handling losses can be significant. To use the multiple separation approach, the CE instrument must be capable of maintaining very stable migration times. Not all instruments have this capability, so it is advisable that the method be used for simple mixtures of peptides. The single-separation approach is recommended for more com-
10.9.12 Supplement 2
Current Protocols in Protein Science
plex mixtures. Because the fractions are collected “blind,” they should be screened by matrix-assisted laser desorption/ionization (MALDI; UNIT 11.6) mass spectrometry prior to sequencing.
Time Considerations
Complete analysis times for IEF are ∼30 min. Analysis times for protein separations are ∼30 min for each step in method development. Peptide separations are normally completed in 40 min and frequently in as little as 20 min. Micropreparative peptide separations using either the multiple or single-separation approach are normally achievable within 90 min, but may take 2 hr or longer. It should be noted that these times are approximate, and sufficient time should be allowed to electrophorese all the components completely from the capillary.
Literature Cited Bergman, T. and Jornvall, H. 1992. Capillary electrophoresis for preparation of peptides and direct determination of amino acids. In Techniques in Protein Chemistry III (R.A. Hogue Angeletti, ed.) pp. 129-134. Academic Press, San Diego. Bullock, J.A. and Yuan, L.-C. 1991. FSCE of basic proteins in uncoated fused-silica capillary tubing. J. Microcol. Sep. 3:241-248. Burgi, D.S. and Chien, R.-L. 1992. On-column sample concentration using field-amplification in CZE. Anal. Chem. 64:489A-495A. Chen, F.A., Kelly, L., Palmieri, R., Biehler, R., and Schwartz, H. 1992. Use of high ionic strength buffers for the separation of proteins and peptides with CE. J. Liq. Chromatogr. 15:11431161. Dolnik, V., Cobb, K.A., and Novontny, M. 1990. Capillary zone electrophoresis of dilute samples with isotachophoretic preconcentration. J. Microcol. Sep. 2:127-140. Kenny, J.W., Ohms, J.I., and Smith, A.J. 1993. Micropreparative capillary electrophoresis (MPCE) and micropreparative HPLC of protein digests. In Techniques in Protein Chemistry, IV (R.A. Hogue Angeletti, ed.) pp. 363-370. Academic Press, San Diego. Khaledi, M.G. 1994. Micellar electrokinetic capillary chromatography. In Handbook of Capillary Electrophoresis (J.P. Landers, ed.) pp. 43-94. CRC Press, Boca Raton, Fla.
Mazzeo, J.R. and Krull, I.S. 1992. Improvements in the methods developed for performing isoelectric focusing in uncoated capillaries. J. Chromatogr. 606:291-296. Nielsen, R.G., Richard, E.C., Santa, P.F., Sharknas, D.A., and Sittampalam, G.S. 1991. Separation of antibody-antigen complexes by CZE, IEF, and HPSEC. J. Chromatogr. 539:177-185. Novotny, M.V., Cobb, K.A., and Liu, J. 1990. Recent advances in CE of proteins, peptides, and amino acids. Electrophoresis 11:735-749. Palmieri, R. and Nolan, J.A. 1994. Protein capillary electrophoresis: Theoretical and experimental considerations for methods development. In Handbook of Capillary Electrophoresis (J.P. Landers, ed.) pp. 325-368. CRC Press, Boca Raton, Fla. Strickland, M. and Strickland, N. 1990. FSCE using phosphate buffer and acidic pH. Am. Lab. 22:6065. Tran, A.D., Parks, S., Lisi, P.J., Huynh, O.T., Ryall, R.R., and Lane, P.A. 1991. Separations of carbohydrate-mediated microheterogeneity of human erythropoietin by free solution CE. J. Chromatogr. 542:459-471. Tsuiji, K. and Little, R.J. 1992. Charge-reversed, polymer-coated capillary column for the analysis of a recombinant chimeric glycoprotein. J. Chromatogr. 594:317-324. Wiktorowicz, J.E. and Colburn, J.C. 1990. Separation of cationic proteins via charge reversal in CE. Electrophoresis 11:769-773.
Key References McCormick, R.M. 1994. Capillary zone electrophoresis of peptides. In Handbook of Capillary Electrophoresis (J.P. Landers, ed.) pp. 287-324. CRC Press, Boca Raton, Fla. Excellent review of both theory and practice for separation of peptides by CE. Palmeiri and Nolan, 1994. See above. Good review of methods development for protein separations.
Contributed by Dean Burgi Genomyx Foster City, California Alan J. Smith Stanford University Medical Center Stanford, California
Landers, J.P., Madden, B.J., Oda, R.P., and Spelsberg, T.C. 1992. The use of modifiers of endoosmotic flow for analysis of microheterogeneity. Anal. Biochem. 205:115-124.
Electrophoresis
10.9.13 Current Protocols in Protein Science
Supplement 2
Immunoblot Detection
UNIT 10.10
Immunoblotting (often referred to as western blotting) is used to identify specific antigens recognized by polyclonal or monoclonal antibodies. Protein samples are solubilized, usually with sodium dodecyl sulfate (SDS) and in selected cases with reducing agents such as dithiothreitol (DTT) or 2-mercaptoethanol (2-ME); some antibody epitopes are destroyed if reducing conditions are used. Following solubilization, the material is separated by SDS-PAGE (using either one- or two-dimensional gels; UNITS 10.1 & 10.4). The antigens are then electrophoretically transferred in a tank or a semidry electroblotting unit to a nitrocellulose, polyvinylidene difluoride (PVDF), or nylon membrane (UNIT 10.7). When nitrocellulose or PVDF membranes are used (UNIT 10.8), the process can be monitored by a reversible staining procedure with Ponceau S. After staining, protein bands on the membrane can be photographed and/or the positions of the detected proteins can be marked with indelible ink (e.g., Paper-Mate pen). The membrane is then completely destained by soaking in water for an additional 10 min. At this point the transferred proteins are bound to the surface of the membrane, providing access for reaction with immunodetection reagents. All remaining binding sites are blocked by immersing the membrane in a solution containing either a protein or detergent blocking agent. After being probed with primary antibody, the membrane is washed and the antibody-antigen complexes are identified using horseradish peroxidase (HRPO) or alkaline phosphatase (AP) enzymes coupled to the secondary anti-immunoglobulin-G (anti-IgG) antibody (e.g., goat anti-rabbit IgG). The enzymes are attached directly (Basic Protocol) or via an avidin-biotin bridge (Alternate Protocol) to the secondary antibody. Chromogenic or luminescent substrates (Support Protocols 1 and 2) are then used to visualize the activity. IMMUNOPROBING WITH DIRECTLY CONJUGATED SECONDARY ANTIBODY
BASIC PROTOCOL
After electrophoretic transfer to the membrane (UNIT 10.7), the immobilized proteins are probed with specific antibodies to identify and quantitate any antigens present. The membrane is first immersed in blocking buffer to fill all protein-binding sites with a nonreactive protein or detergent. Next, the membrane is placed in a solution containing an antibody directed against the antigen (primary antibody). The blot is washed and then exposed to an enzyme-antibody conjugate directed against the primary antibody (secondary antibody; e.g., goat anti-rabbit IgG). Antigens are identified by chromogenic or luminescent visualization (see Support Protocols 1 and 2) of the antigen/primary antibody/secondary antibody/enzyme complex bound to the membrane. Tween 20 is a common alternative to protein blocking agents for use with nitrocellulose or PVDF filters. Materials Membrane with transferred proteins (UNIT 10.8) Blocking buffer (see recipe) appropriate for membrane and detection protocol Primary antibody specific for protein of interest TTBS (nitrocellulose or PVDF) or TBS (neutral or positively charged nylon; see recipes for both solutions) Secondary antibody conjugate: horseradish peroxidase (HRPO)- or alkaline phosphatase (AP)-anti-Ig (Cappel, Vector, Kirkegaard & Perry, or Sigma; dilute as indicated by manufacturer and store frozen in 25-µl aliquots until use) Heat-sealable plastic bags Powder-free gloves Electrophoresis Contributed by Sean Gallagher Current Protocols in Protein Science (1996) 10.10.1-10.10.12 Copyright © 2000 by John Wiley & Sons, Inc.
10.10.1 Supplement 4
Plastic box Additional reagents and equipment for chromogenic or luminescent visualization (see Support Protocol 1 or see Support Protocol 2) 1. Place membrane in heat-sealable plastic bag with 5 ml blocking buffer and seal bag. Incubate 30 min to 1 hr at room temperature with agitation on an orbital shaker or rocking platform. Usually 5 ml buffer is sufficient for two to three membranes (14 × 14–cm size). Plastic incubation trays are often used in place of heat-sealable bags, and can be especially useful when processing large numbers of strips in different primary antibody solutions.
2. Dilute primary antibody in blocking buffer. Primary antibody dilution is determined empirically but is typically 1/100 to 1/1000 for a polyclonal antibody (Fig. 10.10.1; Cooper and Paterson, 1995; Andrew and Titus, 1991a,b,c, 1993), 1/10 to 1/100 for hybridoma supernatants (Yokoyama, 1991a), and ≥1/1000 for murine ascites fluid containing monoclonal antibodies (Yokoyama, 1991b). Ten- to one-hundred-fold higher dilutions can be used with alkaline phosphatase– or luminescence–based detection systems. Both primary and secondary antibody solutions can be used at least twice, but long-term storage (i.e., >2 days at 4°C) is not recommended.
3. Open bag and pour out blocking buffer. Replace with diluted primary antibody and incubate 30 min to 1 hr at room temperature with constant agitation. Usually 5 ml diluted primary antibody solution is sufficient for two to three membranes (14 × 14–cm size). Incubation time may vary, depending on conjugate used.
1/3200 1/6400
1/1600
1/50 1/100 1/200 1/400 1/800
Serum Dilution
Size (kDa) 200
Figure 10.10.1 Serial dilution of primary antibody directed against the 97-kDa catalytic subunit of the plant plasma membrane ATPase. The blot was developed with HRPO-coupled avidin-biotin reagents according to the Alternate Protocol and visualized with 4-chloro-1-naphthol (4CN). Note how background improves with dilution.
116 97
66
43
24
18
Immunoblot Detection
10.10.2 Supplement 4
Current Protocols in Protein Science
When using plastic trays, the primary and secondary antibody solution volume should be increased to 25 to 50 ml. For membrane strips, incubation trays with individual slots are recommended. Typically, 0.5 to 1 ml solution/slot is needed.
4. Remove membrane from plastic bag with gloved hand. Place in plastic box and wash 4 times by agitating with 200 ml TTBS (nitrocellulose or PVDF) or TBS (nylon), 10 to 15 min each time. 5. Dilute secondary antibody HRPO- or AP-anti-Ig conjugate in blocking buffer. Commercially available enzyme-conjugated secondary antibody is usually diluted 1/200 to 1/2000 (i.e., 20 ìl/ml to 2 ìl/ml) prior to use (Harlow and Lane, 1988).
6. Place membrane in fresh heat-sealable plastic bag, add diluted HRPO- or AP-anti-Ig conjugate, and incubate 30 min to 1 hr at room temperature with constant agitation. When using plastic incubation trays, see step 3 annotation for proper antibody solution volumes.
7. Remove membrane from bag and wash as in step 4. Develop according to appropriate visualization protocol (see Support Protocol 1 or see Support Protocol 2). IMMUNOPROBING WITH AVIDIN-BIOTIN COUPLING TO SECONDARY ANTIBODY
ALTERNATE PROTOCOL
The following procedure is based on the Vectastain ABC kit from Vector (SUPPLIERS APPENDIX). It uses an avidin-biotin complex to attach horseradish peroxidase (HRPO) or alkaline phosphatase (AP) to the biotinylated secondary antibody. Avidin-biotin systems are capable of extremely high sensitivity because multiple reporter enzymes are bound to each secondary antibody. In addition, the detergent Tween 20 is a popular alternative to protein blocking agents when using nitrocellulose or PVDF membranes. Additional Materials (also see Basic Protocol) Vectastain ABC (HRPO) or ABC-AP (AP) kit (Vector) containing the following: reagent A (avidin), reagent B (biotinylated HRPO or AP), and biotinylated secondary antibody (request membrane immunodetection protocols when ordering) 1. Equilibrate membrane in appropriate blocking buffer in heat-sealed plastic bag with constant agitation using an orbital shaker or rocking platform. For nitrocellulose and PVDF, incubate 30 to 60 min at room temperature. For nylon, incubate ≥2 hr at 37°C. TTBS is well suited for avidin-biotin systems. For nylon, protein binding agents are recommended. Because nonfat dry milk contains residual biotin that will interfere with the immunoassay, its use must be restricted to the blocking step only. Plastic incubation trays are often used in place of heat-sealable bags, and can be especially useful when processing large numbers of strips in different primary antibody solutions.
2. Prepare primary antibody solution in TTBS (nitrocellulose or PVDF) or TBS (nylon). Dilutions of sera containing primary antibody generally range from 1/100 to 1/10,000. This depends in large part on the sensitivity of the detection system. With high-sensitivity avidin-biotin systems, dilutions from 1/1000 to 1/100,000 are common. Higher dilutions can be used with AP- or luminescence-based detection systems. To determine the appropriate concentration of primary antibody, a dilution series is easily performed with membrane strips. Separate antigens on a preparative gel (i.e., with a single large sample well) and immunoblot the entire gel. Cut 2- to 4-mm strips by hand or with a membrane cutter (Schleicher & Schuell or Inotech) and incubate individual strips in a set of serial dilutions of primary antibody. The correct dilution should give low background and high specificity (Fig. 10.10.1).
Electrophoresis
10.10.3 Current Protocols in Protein Science
Supplement 4
3. Open bag, remove blocking buffer, and add enough primary antibody solution to cover membrane. Reseal bag and incubate 30 min at room temperature with gentle rocking. When using plastic trays, the primary and secondary antibody solution volume should be increased to 25 to 50 ml. For membrane strips, incubation trays with individual slots are recommended. Typically, 0.5 to 1 ml solution/slot is needed.
4. Remove membrane from bag and place in plastic box. Wash membrane 3 times over a 15-min span in TTBS (nitrocellulose or PVDF) or TBS (nylon). Add enough TTBS or TBS to fully cover the membrane (e.g., 5 to 10 ml/strip or 25 to 50 ml/whole membrane). 5. Prepare biotinylated secondary antibody solution by diluting 2 drops biotinylated antibody with 50 to 100 ml TTBS (nitrocellulose or PVDF) or TBS (nylon). This dilution gives both high sensitivity and enough volume to easily cover a large (14 × 14–cm) membrane.
6. Transfer membrane to fresh plastic bag containing secondary antibody solution. Incubate 30 min at room temperature with slow rocking, then wash as in step 4. When using plastic incubation trays, see step 3 annotation for proper antibody solution volumes.
7. While membrane is being incubated with secondary antibody, prepare avidin-biotinHRPO or -AP complex. Mix 2 drops Vectastain reagent A and 2 drops reagent B into 10 ml TTBS (nitrocellulose or PVDF) or TBS (nylon). Incubate 30 min at room temperature, then further dilute to 50 ml with TTBS or TBS. Diluting the A and B reagents to 50 ml expands the amount of membrane that can be probed without greatly affecting sensitivity. Azide is a peroxidase inhibitor and should not be used as a preservative for long-term storage of the antibody solution. Casein, nonfat dry milk, serum, and some grades of bovine serum albumin (BSA) may interfere with the formation of the avidin-biotin complex and should not be used in the presence of avidin or biotin reagents (Gillespie and Hudspeth, 1991; see also instructions from Vector).
8. Transfer membrane to fresh plastic bag containing avidin-biotin-enzyme solution. Incubate 30 min at room temperature with slow rocking, then wash over a 30-min span as in step 4. Hybridization in a plastic bag requires 5 to 10 ml avidin-biotin-enzyme solution. Membrane strips require 5 to 10 ml/strip, whereas blots from standard-sized gels (i.e., 14 × 16 cm) require 50 ml for convenient handling in a tray.
9. Develop membrane according to the appropriate visualization protocol (see Support Protocol 1 or see Support Protocol 2).
Immunoblot Detection
10.10.4 Supplement 4
Current Protocols in Protein Science
VISUALIZATION WITH CHROMOGENIC SUBSTRATES After incubation with primary and secondary antibody conjugates (see Basic Protocol or see Alternate Protocol), bound antigens are typically visualized with chromogenic substrates. The substrates 4CN, DAB/NiCl2, and TMB are commonly used with horseradish peroxidase (HRPO)–based immunodetection procedures, whereas BCIP/NBT is recommended for alkaline phosphatase (AP)–based procedures (see Table 10.10.1). After incubation with primary and secondary antibodies, the membrane is placed in the appropriate substrate solution. Protein bands usually appear within a few minutes.
Table 10.10.1
SUPPORT PROTOCOL 1
Chromogenic and Luminescent Visualization Systemsa
System
Reagentb
Reaction/detection
Commentsc
Chromogenic HRPO-based
4CN
Oxidized products form purple precipitate
DAB/NiCl2d
Forms dark brown precipitate
TMBe
Forms dark purple stain
BCIP/NBT
BCIP hydrolysis produces indigo precipitate after oxidation with NBT; reduced NBT precipitates; dark blue-gray stain results
Not very sensitive (Tween 20 inhibits reaction); fades rapidly on exposure to light More sensitive than 4CN but potentially carcinogenic; resulting membrane is easily scanned More stable and less toxic than DAB/NiCl2; may be somewhat more sensitivee; can be used with all membrane types More sensitive and reliable than other AP-precipitating substrates; note that phosphate inhibits AP activity
Luminol/H2O2/ p-iodophenol
Oxidized luminol substrate gives off blue light; p-iodophenol increases light output Dephosphorylated substrate gives off light
AP-based
Luminescent HRPO-based
AP-based
Substituted 1,2dioxetane phosphates (e.g., AMPPD, CSPD, Lumigen-PPD, Lumi-Phos 530f)
Very convenient, sensitive system; reaction is detected within a few seconds to 1 hr Protocol described gives reasonable sensitivity on all membrane types; consult instructions of reagent manufacturer for maximum sensitivity and minimum background (see Troubleshooting)
aAbbreviations: AMPPD or Lumigen-PPD, disodium 3-(4-methoxyspiro{1,2-dioxetane-3,2′-tricyclo[3.3.1.13,7]-decan}-4-yl)phenyl phosphate; AP, alkaline phosphatase; BCIP, 5-bromo-4-chloro-3-indolyl phosphate; 4CN, 4-chloro-1-napthol; CSPD, AMPPD with substituted chlorine moiety on adamantine ring; DAB, 3,3′-diaminobenzidine; HRPO, horseradish peroxidase; NBT, nitroblue tetrazolium; TMB, 3,3′,5,5′-tetramethylbenzidine. bRecipes and suppliers for all reagents except TMB are listed in Reagents and Solutions. Kits containing TMB are available from Kirkegaard & Perry, TSI Center for Diagnostic Products, Moss, and Vector. cSee Commentary for further details. dDAB/NiCl can be used without the nickel enhancement, but sensitivity is greatly reduced. 2 eFirst treating nitrocellulose filters with 1% dextran sulfate for 10 min in 10 mM citrate-EDTA (pH 5.0) causes TMB to precipitate onto the membrane
with a sensitivity much greater than that seen for 4CN or DAB and equal to or better than that for BCIP/NBT (McKimm-Breschkin, 1990). fLumi-Phos 530 contains dioxetane, MgCl , cetyltrimethylammonium bromide (CTAB), and fluorescent enhancer in a pH 9.6 buffer. 2
Electrophoresis
10.10.5 Current Protocols in Protein Science
Supplement 4
Materials Membrane with transferred proteins and probed with antibody-enzyme complex (see Basic Protocol or see Alternate Protocol) TBS (see recipe) Chromogenic visualization solution (Table 10.10.1; see recipes) Additional reagents and equipment for gel photography (UNIT 10.5) 1. If final membrane wash (see Basic Protocol, step 7, or see Alternate Protocol, step 9) was performed in TTBS, wash membrane 15 min at room temperature in 50 ml TBS. The Tween 20 in the TTBS interferes with 4CN development (Bjerrum et al., 1988).
2. Place membrane into chromogenic visualization solution. Bands should appear in 10 to 30 min. 3. Terminate reaction by washing membrane in distilled water. Air dry and photograph (UNIT 10.5) for a permanent record. SUPPORT PROTOCOL 2
VISUALIZATION WITH LUMINESCENT SUBSTRATES After incubation with primary and secondary antibody conjugates (see Basic Protocol and see Alternate Protocol), antigens can also be visualized with luminescent substrates. Detection with light offers both greater speed and enhanced sensitivity over chromogenic and radioisotopic procedures. After the final wash, the blot is immersed in a substrate solution containing luminol for horseradish peroxidase (HRPO) systems, or dioxetane phosphate for alkaline phosphatase (AP) systems, sealed in thin plastic wrap and placed firmly against film. Exposures range from a few seconds to several hours, although typically strong signals appear within a few seconds or minutes. Additional Materials (also see Support Protocol 1) Luminescent substrate buffer: 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E; HRPO), or dioxetane phosphate substrate buffer (see recipe; AP) Nitro-Block solution (AP reactions only): 5% (v/v) Nitro-Block (Tropix) in dioxetane phosphate substrate buffer (see recipe), prepared just before use Luminescent visualization solution (Table 10.10.1; see recipes) Clear plastic wrap NOTE: See Troubleshooting section for suggestions concerning optimization of the protocol, particularly when employing AP-based systems. 1. Equilibrate membrane in two 15-min washes with substrate buffer each. For blots of whole gels, use 50 ml substrate buffer; for strips, use 5 to 10 ml/strip.
2. For AP reactions using nitrocellulose or PVDF membranes: Incubate 5 min in Nitro-Block solution, followed by 5 min in substrate buffer. For blots of whole gels, use 50 ml Nitro-Block solution and substrate buffer; for strips, use 5 to 10 ml/strip. Nitro-Block enhances light output from the dioxetane substrate in reactions using AMPPD, CSPD, or Lumigen-PPD concentrate. It is required for nitrocellulose and recommended for PVDF membranes. It is not needed for Lumi-Phos 530, AP reactions on nylon membranes, or HRPO–based reactions on any type of membrane. Lumi-Phos 530 is not recommended for nitrocellulose membranes. Immunoblot Detection
3. Transfer membrane to luminescent visualization solution. Soak 30 sec (HRPO reactions) to 5 min (AP reactions).
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Current Protocols in Protein Science
Alternatively, lay out a square of plastic wrap and pipet 1 to 2 ml visualization solution into the middle. Place membrane on the plastic so that the visualization solution spreads out evenly from edge to edge. Fold wrap back onto membrane, seal, and proceed to step 5.
4. Remove membrane, drain, and place face down on a sheet of clear plastic wrap. Fold wrap back onto membrane and seal with tape to form a liquid-tight enclosure. To ensure an optimal image, only one layer of plastic should be present between the membrane and film. Sealable bags are an effective alternative. Moisture must not come in contact with the X-ray film.
5. In a darkroom, place membrane face down onto film. Do this quickly and do not reposition; a double image will be formed if the membrane is moved while in contact with the film. A blurred image is usually caused by poor contact between membrane and film; use a film cassette that ensures a tight fit.
6. Expose film for a few seconds to several hours. Typically, immunoblots produce very strong signals within a few seconds or minutes; however, weak signals may require several hours to an overnight exposure. If no image is detected, expose film 30 min to 1 hr and, if needed, overnight (see Troubleshooting).
7. If desired, wash membrane in two 15-min washes of 50 ml TBS and process for chromogenic development (see Support Protocol 1). It is possible to develop the same membrane with chromogenic substrates after luminescent visualization.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Alkaline phosphate substrate buffer 100 mM Tris⋅Cl, pH 9.5 100 mM NaCl 5 mM MgCl2 Blocking buffer Colorimetric detection For nitrocellulose and PVDF: 0.1% (v/v) Tween 20 in TBS (TTBS; see recipe). For neutral and positively charged nylon: Tris-buffered saline (TBS; see recipe) containing 10% (w/v) nonfat dry milk. TTBS can be stored up to 1 week at 4°C. Prepare blocking buffer containing nonfat dry milk immediately prior to use, as the milk blocking solution is not stable.
Luminescent detection For nitrocellulose, PVDF, and neutral nylon (e.g., Pall Biodyne A): 0.2% (w/v) casein (e.g., Hammarsten grade or I-Block; Tropix) in TTBS (see recipe). For positively charged nylon: 6% (w/v) casein/1% (v/v) polyvinylpyrrolidone (PVP) in TTBS (see recipe). For each solution: With constant mixing, add casein and PVP to warm (65°C) TTBS. Stir for 5 min, then cool. Prepare each solution just before use.
Electrophoresis
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Chromogenic visualization solutions BCIP/NBT visualization solution: Mix 33 µl NBT stock [100 mg NBT in 2 ml 70% (v/v) dimethylformamide (DMF), stored <1 year at 4°C] and 5 ml alkaline phosphate substrate buffer (see recipe). Add 17 µl BCIP stock (100 mg BCIP in 2 ml 100% DMF, stored <1 year at 4°C) and mix. Stable 1 hr at room temperature. Recipe is from Harlow and Lane (1988). Alternatively, BCIP/NBT substrates may be purchased from Sigma, Kirkegaard & Perry, Moss, and Vector (SUPPLIERS APPENDIX).
4CN visualization solution: Mix 20 ml ice-cold methanol with 60 mg 4-chloro-1naphthol (4CN). Separately mix 60 µl of 30% (w/v) H2O2 with 100 ml TBS (see recipe) at room temperature. Rapidly mix the two solutions and use immediately. DAB/NiCl2 visualization solution: 5 ml 100 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 100 µl DAB stock (40 mg/ml in H2O, stored in 100-µl aliquots at −20°C) 25 µl NiCl2 stock (80 mg/ml in H2O, stored in 100-µl aliquots at −20°C) 15 µl 3% (w/v) H2O2 Mix just before use CAUTION: Handle DAB carefully, wearing gloves and mask; it is a carcinogen. Suppliers of chromogenic HRPO substrates (4CN and DAB/NiCl2) are Sigma, Kirkegaard & Perry, Moss, and Vector (SUPPLIERS APPENDIX). For selection of appropriate chromogenic solutions, and for definition of abbreviations, see Table 10.10.1.
Dioxetane phosphate substrate buffer 1 mM MgCl2 0.1 M diethanolamine 0.02% (w/v) sodium azide (optional) Adjust to pH 10 with HCl and use fresh Traditionally, the AMPPD substrate buffer has been a solution containing 1 mM MgCl2 and 50 mM sodium carbonate/bicarbonate, pH 9.6 (Gillespie and Hudspeth, 1991). The use of diethanolamine results in better light output (Tropix Western Light instructions). Alternatively, 100 mM Tris⋅Cl (pH 9.5)/100 mM NaCl/5 mM MgCl2 can be used (Sandhu et al., 1991).
Luminescent visualization solutions Dioxetane phosphate visualization solution: Prepare 0.1 mg/ml AMPPD or CSPD (Tropix) or 0.1 mg/ml Lumigen-PPD (Lumigen) substrate in dioxetane phosphate substrate buffer (see recipe). Prepare just before use. Lumi-Phos 530 (Boehringer Mannheim or Lumigen) is a ready-to-use solution and can be applied directly to the membrane. This concentration (240 ìM) of AMPPD substrate is the minimum recommended by Tropix. Ten-fold lower concentrations can be used but require longer exposures.
Luminol visualization solution: 0.5 ml 10× luminol stock [40 mg luminol (Sigma) in 10 ml dimethyl sulfoxide (DMSO)] 0.5 ml 10× p-iodophenol stock [optional; 10 mg (Aldrich) in 10 ml DMSO] 2.5 ml 100 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 25 µl 3% (w/v) H2O2 H2O to 5 ml Prepare just before use Recipe is from Schneppenheim et al. (1991). Premixed luminol substrate mix (Mast Immunosystems, Amersham, Du Pont NEN Renaissance, or Kirkegaard & Perry LumiImmunoblot Detection
continued
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GLO) may also be used. For selection of appropriate luminescent solutions, and for definition of abbreviations, see Table 10.10.1. p-Iodophenol is an optional enhancing agent that increases light output. Luminol and p-iodophenol stocks can be stored ≤6 months at −20°C.
Tris-buffered saline (TBS) 100 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 0.9% (w/v) NaCl Store up to several months at 4°C Tween 20/TBS (TTBS) 0.1% (v/v) Tween 20 in Tris-buffered saline (TBS; see recipe) Store up to several months at 4°C COMMENTARY Background Information Immunoprecipitation has been widely used to visualize the antigens recognized by various antibodies, both polyclonal and monoclonal. However, there are several problems inherent to immunoprecipitation, including the need to radiolabel the antigen, coprecipitation of tightly associated macromolecules, occasional difficulty in obtaining precipitating antibodies, and insolubility of various antigens (Talbot et al., 1984). To circumvent these problems, electroblotting (Towbin et al., 1979)—subsequently popularized as western blotting or immunoblotting (Burnette, 1981)—was conceived. Immunoblotting is a rapid and sensitive assay for the detection and characterization of proteins that works by exploiting the specificity inherent in antigen-antibody recognition. It involves the solubilization and electrophoretic separation of proteins, glycoproteins, or lipopolysaccharides by SDS-PAGE (UNITS 10.1 & 10.4) or urea-PAGE (UNIT 10.2), followed by quantitative transfer and irreversible binding to nitrocellulose, PVDF, or nylon membranes (UNIT 10.8). This technique has been useful in identifying specific antigens recognized by polyclonal or monoclonal antibodies, and is highly sensitive (1 ng of antigen can be detected). Protein blotting has important clinical applications—notably, it is the confirmatory test for human immunodeficiency virus type 1 (HIV1). SDS-PAGE-separated virus proteins are blotted onto nitrocellulose and processed with patient sera. After washing and incubating (using a Tween 20/nonfat dry milk diluting and washing solution) with an anti-human IgG coupled to horseradish peroxidase (HRPO) or alkaline phosphatase (AP), the antigens are identified by chromogenic development. Typically,
the prepared blots and all reagents needed for the test are purchased commercially (Bio-Rad Novapath, Organon Teknika Cappel HIV-1 Western Blot Kit, or Du Pont NEN HIV Western Blot Kit). Immunoblotted proteins can be detected by chromogenic or luminescent assays (see Table 10.10.1 for a description of the reagents available for each system, their reactions, and a comparison of their advantages and disadvantages). Luminescent detection methods offer several advantages over traditional chromogenic procedures. In general, luminescent substrates increase the sensitivity of both HRPO and AP systems without the need for radioisotopes. Substrates for the latter have only recently been applied to protein blotting (see Gillespie and Hudspeth, 1991; Sandhu et al., 1991; Bronstein et al., 1992). Luminescent detection can be completed in as little as a few seconds; exposures rarely exceed 1 hr. Depending on the system, the luminescence can last up to 3 days, permitting multiple exposures of the same blot. Furthermore, the signal is detected by film, and varying the exposure can result in more or less sensitivity. Luminescent blots can be easily erased and reprobed because the reaction products are soluble and do not deposit on the membrane. Compared to chromogenic development, a luminescent image recorded on film is easier to photograph and to quantitate by densitometry. AP–based luminescent protocols that achieve maximum sensitivity with minimum background can be complex, and the manufacturer’s instructions should be consulted (see Reagents and Solutions). The procedure described in Support Protocol 2 gives reasonable sensitivity on nitrocellulose, PVDF, and nylon membranes with a minimum of steps. Electrophoresis
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Critical Parameters
Troubleshooting
First and foremost, the antibody being used should recognize denatured antigen. Nonspecific binding of antibodies can occur, so control antigens and antibodies should always be run in parallel. Time of transfer and dilutions of primary antibody and conjugate should always be optimized. A variety of agents are currently used to block binding sites on the membrane after blotting (Harlow and Lane, 1988). These include Tween 20, PVP, nonfat dry milk, casein, BSA, and serum. A 0.1% (v/v) solution of Tween 20 in TBS (TTBS), a convenient alternative to protein-based blocking agents, is recommended for chromogenic development of nitrocellulose and PVDF membranes (Blake et al., 1984). In contrast to dry milk/TBS blocking solution, TTBS is stable and has a long shelf life at 4°C. Furthermore, TTBS generally produces a clean background and permits subsequent staining with India ink. Two types of nylon membranes are used for transfer—neutral (e.g., Pall Biodyne A) and positively charged (e.g., Pall Biodyne B). Although the positively charged membranes have very good protein-binding characteristics, they tend to have a higher background. These membranes remain positively charged from pH 3 to pH 10. Neutral nylon membranes are also charged, having a mix of amino and carboxyl groups that give an isoelectric point of 6.5. Because of their high binding capacity, positively charged membranes are popular for protein applications using luminescence. Nylon membranes require more stringent blocking steps. Here 10% (w/v) nonfat dry milk in TBS is recommended for chromogenic development. During development of luminescence, however, background is a more significant problem. When compared to dry milk, purified casein has minimal endogenous AP activity (such activity leads to high background) and is therefore recommended as a blocking agent for nitrocellulose, PVDF, and nylon membranes. Positively charged nylon requires much more stringent blocking with 6% (w/v) casein and 1% (v/v) polyvinylpyrrolidone (PVP-40). Because nonfat dry milk and casein may contain biotin, which will interfere with avidin-biotin reactions, subsequent steps are done without protein blocking agents when using these systems. If background is a problem, highly purified casein (0.2% to 6%) added to the antibody incubation buffers may help.
There are several problems associated with immunoblotting. The antigen is solubilized and electrophoresed in the presence of denaturing agents (e.g., SDS or urea), and some antibodies may not recognize the denatured form of the antigen transferred to the membrane. The results observed may be entirely dependent on the denaturation and transfer system used. For example, zwitterionic detergents have been shown to restore the antigenicity of outer membrane proteins in immunoblotting (Mandrell and Zollinger, 1984). Gel electrophoresis under nondenaturing conditions can sometimes preserve antigenicity (UNIT 10.3). Other potential problems include high background, nonspecific or weak cross-reactivity of antibodies, poor protein transfer or membrane binding efficiency, and insufficient sensitivity. For an extensive survey and discussion of immunoblotting problems and artifacts, see Bjerrum et al. (1988). Insufficient blocking or nonspecific binding of the primary or secondary antibody will cause a high background stain. A control using preimmune sera or only the secondary antibody will determine if these problems are due to the primary antibody. Try switching to another blocking agent; protein blocking agents may weakly cross-react. Lowering the concentration of primary antibody should decrease background and improve specificity (Fig. 10.10.1). Because of the nature of light and the method of detection, certain precautions are warranted when using luminescent visualization (e.g., Harper and Murphy, 1991). Very strong signals can overshadow weaker signals nearby on the membrane. Because light will pipe through the membrane and the surrounding plastic wrap, overexposure will produce a broad, diffuse image on the film. The signal can also saturate the film, exposing it to a point whereby increased exposure will not cause a linear increase in the density of the image on the film. With the AP substrate AMPPD, nitrocellulose, PVDF, and nylon membranes require 2, 4, and 8 to 12 hr, respectively, to reach maximum light emission. In addition, PVDF is reported to give a stronger signal than nitrocellulose (Tropix Western Light instructions). Positively charged nylon requires special blocking procedures to minimize background (Gillespie and Hudspeth, 1991). These procedures include using a blocking and pri-
Immunoblot Detection
10.10.10 Supplement 4
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mary antibody solution containing 6% (w/v) casein, 1% (v/v) polyvinylpyrrolidone-40 (PVP-40), 3 mM NaN3, 10 mM EDTA, and phosphate-buffered saline (PBS), pH 6.8. Prior to use, the casein must be heated to 65°C to reduce AP activity in the casein itself. In addition, maximum sensitivity has been observed when free biotin or biotinylated proteins are removed by pretreating the casein with avidinagarose (Sigma).
Anticipated Results Immunoblotting should result in the detection of one or more bands. Although antibodies directed against a single protein should produce a single band, degradation of the sample (e.g., via endogenous proteolytic activity) may cause visualization of multiple bands of slightly different size. Multimers will also form spontaneously, causing higher-molecular-weight bands on the blot. If one is simultaneously testing multiple antibodies directed against a complex protein mixture (e.g., using a patient sera against SDS-PAGE-separated viral proteins in the AIDS western blot test), multiple bands will be visualized.
Time Considerations The entire immunoblotting procedure can be completed in 1 to 2 days, depending on the transfer time and type of gel. Gel electrophoresis requires 4 to 6 hr on a regular gel and 1 hr on a minigel. Transfer time can be 1 hr (for high-power transfer) to overnight. Blocking, conjugate incubation, and washing each take 30 min to 1 hr. Finally, substrate incubation requires 10 to 30 min (chromogen) and a few seconds to several hours (luminescence).
Literature Cited Andrew, S.A. and Titus, J.A. 1991a. Purification of immunoglobulin G. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. 2.7.1-2.7.12. John Wiley & Sons, New York. Andrew, S.A. and Titus, J.A. 1991b. Fragmentation of immunoglobulin G. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. 2.8.1-2.8.10. John Wiley & Sons, New York. Andrew, S.A. and Titus, J.A. 1991c. Fragmentation of immunoglobulin M. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. 2.10.1-2.10.4. John Wiley & Sons, New York. Andrew, S.A. and Titus, J.A. 1993. Purification of immunoglobulin M. In Current Protocols in Im-
munology (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. 2.9.1-2.9.3. John Wiley & Sons, New York. Bjerrum, O.J., Larsen, K.P., and Heegaard, N.H.H. 1988. Nonspecific binding and artifacts—specificity problems and troubleshooting with an atlas of immunoblotting artifacts. In CRC Handbook of Immunoblotting of Proteins, Vol. I: Technical Descriptions (O.J. Bjerrum and N.H.H. Heegaard, eds.) pp. 227-254. CRC Press, Boca Raton, Fla. Blake, M.S., Johnston, K.H., Russell-Jones, G.J., and Gotschlich, E.C. 1984. A rapid, sensitive method for detection of alkaline phosphatase– conjugated anti-antibody on Western blots. Anal. Biochem. 136:175-179. Bronstein, I., Voyta, J.C., Murphy, O.J., Bresnick, L., and Kricka, L.J. 1992. Improved chemiluminescent Western blotting procedure. BioTechniques 12:748-753. Burnette, W.N. 1981. Western blotting: Electrophoretic transfer of proteins from sodium dodecyl sulfate–polyacrylamide gels to unmodified nitrocellulose and radiographic detection with antibody and radioiodinated protein A. Anal. Biochem. 112:195-203. Cooper, H.M. and Paterson, Y. 1995. Production of polyclonal antisera. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. 2.4.1-2.4.9. John Wiley & Sons, New York. Gillespie, P.G. and Hudspeth, A.J. 1991. Chemiluminescence detection of proteins from single cells. Proc. Natl. Acad. Sci. U.S.A. 88:25632567. Harlow, E. and Lane, D. 1988. Immunoblotting. In Antibodies: A Laboratory Manual, pp. 471-510. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Harper, D.R. and Murphy, G. 1991. Nonuniform variation in band pattern with luminol/horseradish peroxidase Western blotting. Anal. Biochem. 192:59-63. Mandrell, R.E. and Zollinger, W.D. 1984. Use of zwitterionic detergent for the restoration of antibody-binding capacity of electroblotted meningococcal outer membrane proteins. J. Immunol. Methods 67:1-11. McKimm-Breschkin, J.L. 1990. The use of tetramethylbenzidine for solid phase immunoassays. J. Immunol. Methods 135:277-280. Sandhu, G.S., Eckloff, B.W., and Kline, B.C. 1991. Chemiluminescent substrates increase sensitivity of antigen detection in Western blots. BioTechniques 11:14-16. Schneppenheim, R., Budde, U., Dahlmann, N., and Rautenberg, P. 1991. Luminography—a new, highly sensitive visualization method for electrophoresis. Electrophoresis 12:367-372. Talbot, P.V., Knobler, R.L., and Buchmeier, M. 1984. Western and dot immunoblotting analysis of viral antigens and antibodies: Application to Electrophoresis
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murine hepatitis virus. J. Immunol. Methods 73:177-188. Towbin, H., Staehelin, T., and Gordon, J. 1979. Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: Procedure and some applications. Proc. Natl. Acad. Sci. U.S.A. 76:4350-4354. Yokoyama, W. 1991a. Production of monoclonal antibodies. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. 2.5.12.5.17. John Wiley & Sons, New York. Yokoyama, W. 1991b. Monoclonal antibody supernatant and ascites fluid. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. 2.6.1-2.6.7. John Wiley & Sons, New York.
Key References Gillespie and Hudspeth, 1991. See above. Describes alkaline phosphatase–based luminescent detection methods. Harlow and Lane, 1988. See above. Details alternative detection methods. Schneppenheim et al., 1991. See above. Details horseradish peroxidase–based luminescent detection methods.
Contributed by Sean Gallagher Hoefer Scientific Instruments San Francisco, California
Immunoblot Detection
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Autoradiography Autoradiography is a radioisotope-detection method that records the spatial distribution of radioisotope-labeled substances within a membrane or specimen material. The ionizing radiation (usually β particles) that is emitted from the radioisotope interacts with the silver halide grains within the emulsion of the photographic film, forming a latent image. The latent image is amplified by the action of the developer during processing, thus creating a visible image. The pattern and density of the image are used to locate and quantify the radioactive distribution. Commercially available autoradiography films are best suited for autoradiography because of their high sensitivity to both light and ionizing radiation. Autoradiography films generally yield higher-contrast images with a shorter exposure time than medical X-ray films and other commercially available photographic films. The most appropriate choice of autoradiography film depends upon the detection requirements of the experiment and the type and quantity of the radiolabeled isotope. Several autoradiography films are designed and used with very specific exposure methods and radioisotopes—e.g., BioMax MR and BioMax MS (Eastman Kodak). BioMax MR is designed for direct-exposure detection of 14C and 35S radioisotopes and BioMax MS is designed for exposure detection of 32P and 125I radioisotopes with an intensifying screen. The following descriptions of films and exposure procedures will assist in optimizing autoradiography techniques.
TYPES OF FILM The structure of autoradiography film includes either one emulsion layer or two on a 7-mil (178-µm) polyester support. Doublecoated film includes emulsion layers on both sides of the sheet of film. Emulsion layers consist of light-sensitive silver halide grains and gelatin. The polyester support can be either clear or blue-tinted.
Single-Coated Films Autoradiography films that are single-emulsion coated are optimized for use with mediumenergy radioisotopes—e.g., 14C, 35S, and 33P— and direct-exposure techniques (see discussion of Direct Exposure) for a wide range of biologi-
UNIT 10.11 cal applications such as DNA sequencing. At maximum energy levels of 150 to 170 KeV, the majority of the β particles do not penetrate through the film base, so an additional emulsion on the other side of the film support, such as that found in double-coated films, does not increase sensitivity to 14C and 35S radioisotopes. Single-coated films—e.g., BioMax MR—have greater clarity because they do not have the second emulsion layer. BioMax MR is highly sensitive to direct radioisotope exposure. It is twice as sensitive to 14C and 35S as X-Omat AR film (Eastman Kodak; see discussion of Double-Coated Films). BioMax MR film sensitivity to light has been reduced in order to increase allowable safelight handling time in the darkroom, thus reducing its sensitivity to fluorography or intensifying-screen exposures.
Double-Coated Films Double-coated films are generally used with intensifying screens (see discussion of Intensifying-Screen Exposure) at reduced temperatures to detect high-energy radioisotopes such as 32P and 125I. High-energy β particles, X rays, and γ rays penetrate and expose both emulsion layers. These high-energy particles are primarily absorbed by an intensifying screen if it is placed on the opposite side of the film from the specimen. Light is emitted by the phosphor grains in the intensifying screen upon absorbing the β particle, and this greatly amplifies detection. Double-coated X-Omat AR film is a widely used film for general autoradiography. This film is highly sensitive to blue light and is used with a blue-light-emitting calcium tungstate (CaWO4) intensifying screen. The green-light-sensitive BioMax MS film is a double-coated film that gives greatest sensitivity to 32P when used with a BioMax MS intensifying screen or high-energy TranScreen (Eastman Kodak) at −70°C. Both of these screens emit green light when exposed to ionizing radiation. These film/screen systems are approximately four times more sensitive than the X-Omat AR film/CaWO4 intensifying screen system. Most fluorographic exposures (see discussion of Fluorographic Exposure) emit light in the blue spectrum. X-Omat AR film is a bluesensitive film that is generally used with scintillants such as PPO (2, 5-diphenyloxazole) and Electrophoresis
Contributed by Daniel C. Bundy Current Protocols in Protein Science (1997) 10.11.1-10.11.6 Copyright © 1997 by John Wiley & Sons, Inc.
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sodium salicylate, as well as prepared fluorographic solutions and sprays (Amersham, DuPont NEN). X-Omat AR film is at least two times more sensitive than other blue-sensitive autoradiography films for most fluorographic exposure techniques.
HANDLING OF AUTORADIOGRAPHY FILM Safelighting Safe darkroom illumination is dependent upon the type of safelight filter, the wattage of the bulb, and the distance from the safelight bulb to the sheet of film. A red safelight filter (e.g., Kodak GBX-2) is recommended for the majority of autoradiography films. The bulb wattage should not exceed 15 W with direct-illuminating safelight lamps. The minimum safelight-to-film distance is 4 feet. If one is unsure of the safelight conditions in a darkroom or the safelight sensitivity of a film, it is wise to perform a safelight safety test. In the dark, put the film on the workbench and place a coin on top of the film. Next, turn on the safelight for 1 to 2 min or more, then turn off the safelight and process the film in the dark. If the coin’s shadow image is not visible on the film, then the safelight conditions are suitable for the tested exposure time. If the safelight safety test fails, measures to be taken may include replacing the safelight filter, decreasing the bulb wattage, increasing the safelight-to-film distance, or decreasing the time of exposure to safelight conditions.
EXPOSURE PROCEDURES There are many factors to consider when choosing an autoradiographic exposure procedure. The choices include the isotope, the radioactivity level, whether to use double- or single-coated film, film treatment, and exposure temperature. Each researcher needs to determine the best procedure for each particular experimental protocol (see Table 10.11.1).
Storage and Care of Film
Direct Exposure
BioMax MS, BioMax MR, and X-Omat AR films are highly efficient detectors of background radiation. With these films, the background fog can be expected to increase with film age. It is advisable to keep down the inventory of these films and order them more frequently to minimize background fog. Most laboratories routinely order just enough film for 3 to 5 months. Film sheets should be handled carefully to prevent black or white marks and lines that result from pinching and scratching the emulsion. Individual film sheets should be handled only by the edges.
For direct exposure, the radiolabeled specimen is placed in direct contact with autoradiography film and enclosed in a light-tight cassette for a predetermined, or suitable, amount of time. At the end of exposure, the film is removed from the cassette after noting the orientation of the film sheet with respect to the specimen by bending a corner of the film towards the specimen or by marking a corner of the film with a no. 2 pencil. The film is then photographically processed. Exposing autoradiography film at reduced temperatures (see discussion of Fluorographic Exposure) does not improve the sensitivity with direct β-particle exposures as it does with fluorographic and intensifying-screen exposures. The β particle forms a very stable latent image in the silver grain of the emulsion. This latent image does not significantly fade with time as in the case with light exposures from fluors
QUANTITATIVE AUTORADIOGRAPHY Autoradiography
standards (e.g., 3H and 14C tissue equivalent standards from American Radiolabeled Chemicals or Amersham), self-prepared radioactive scales, or radioisotope-labeled dilution series are used during an autoradiographic exposure to expose the film or storage phosphor screen to quantitate the image. The autoradiography image (e.g., from a tissue specimen, membrane, or gel) and the calibration strip require digitization or densitometry to convert the information to radioactivity from a plot of radioactivity versus density or pixel value. A common method of preparing a calibration strip is to serially dilute the radioisotope label by a factor of 2 or more and apply it to a substrate or membrane. It is important to extend the dilution series beyond the estimated range of the radioisotope label concentration. The individual radioisotope-label concentrations are spotted (3 to 5 µl), blotted, or hybridized onto the support material. The shape and configuration of each spot or band should be similar and uniform. The density or pixel value is plotted versus the radioactivity value to obtain the relative radioisotope label calibration curve.
A radioactive calibration strip is very effective in autoradiography quantification techniques. Commercially available radioactive
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Table 10.11.1
Autoradiography Exposure Procedures
Application(s)
Isotope
Protein gels, western blots Nucleic acid sequencing
Priority
3H, 14C, 35S 32P 35S
Exposure procedurea
Speed
Fluorography TranScreen LE
Resolution Speed Resolution Speed
Direct Direct Fluorography TranScreen LE Direct Intensifying screen
Oligonucleotide separation
32P
Resolution Speed
Northern, Southern, and dot blots; library screening
32P
Speed Speed
Intensifying screen Fluorography TranScreen LE
Resolution Speed
Direct Fluorography TranScreen LE
Speed
Fluorography TranScreen LE
In situ hybridization
35S 14C, 35S
3H
aTranScreen
LE is a product of Eastman Kodak (see SUPPLIERS APPENDIX).
(scintillants) and intensifying screens (see discussions of Intensifying-Screen Exposure and Fluorographic Exposure). Direct-exposure techniques are effective with all β-particle-emitting radioisotopes except tritium (3H). Tritium does not have enough energy to penetrate the overcoat layer on top of autoradiography film and expose the silver halide emulsion. 14C and 35S direct-exposure techniques provide the highest imaging resolution when used with a single-coated autoradiographic film such as BioMax MR film (Steinfeld et al., 1994), although longer exposure times are required than for fluorographic exposures with X-Omat AR film. Wet or moist specimens should be sealed in a plastic bag or wrapped in plastic wrap. Water and other solvents should not come in contact with film. The plastic wrap attenuates radiation from medium-energy β emitters (35S or 14C) by a factor of 2. The effect with 32P is less, but still noticeable. The investigator should be aware of the thickness of the plastic wrap used. Commercially available plastic wraps are acceptable; extremely thin plastic wraps are better but more expensive.
Fluorographic Exposure Fluorography enhances the detection of low levels of low- and medium-energy radioisotopes such as 3H, 14C, and 35S. The scintillant
(i.e., a reagent that emits visible light upon absorbing ionizing radiation) is placed either on or in the radiolabeled specimen. When the scintillant absorbs the β-particle energy it emits blue light, which is detected by autoradiographic film. In fluorographic exposure, the radiolabeled specimen (e.g., gel, nitrocellulose membrane, or TLC plate) is soaked or sprayed with the scintillant. The specimen is then dried according to the instructions provided by the scintillant manufacturer and published protocols. Next, under safelight conditions, the specimen is placed against autoradiography film in a light-tight cassette. The temperature for optimum exposure sensitivity is between −70° and −100°C. At the end of the exposure, the cassette is warmed to ambient temperature before opening. The film is carefully removed from the specimen after marking the orientation of the film sheet with respect to the specimen by bending a corner of the film towards the specimen or by marking a corner of the film with a no. 2 pencil. Then the film is photographically processed. If the film and specimen are separated too rapidly, static discharge may occur. This static may appear on the film as many small black dots or as large, branched black streaks. X-Omat AR film has a spectral sensitivity that is optimal for detecting fluorescence from
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film. 32P and 125I are the most common highenergy β and γ emitters that are exposed with intensifying screens. In this procedure, under safelight conditions, the intensifying screen is placed in the cassette and a sheet of double-coated autoradiography film is placed on top of it. Next, a radiolabeled specimen is placed on top of the film with its radioactive side against the film and exposed at optimum exposure temperature—i.e., −70° to −100°C. At the end of the exposure, the cassette is warmed to ambient temperature before opening it. The film is removed carefully from the specimen after noting the orientation of the film sheet with respect to the specimen by bending a corner of the film towards the specimen or by marking a corner of the film with a no. 2 pencil. Then the film is photographically processed. Inorganic phosphors in the intensifying screen convert ionizing radiation to blue light or blue and green light, depending on the type of intensifying screen. Generally, the spectral emission of the intensifying screen is matched to the spectral sensitivity of the film to produce optimal sensitivity. X-Omat AR film is spectrally matched with blue-light-emitting intensifying screens and BioMax MS film is spectrally matched to the blue- and green-lightemitting BioMax MS intensifying screen.
the scintillant 2,5-diphenyloxazole (PPO), which emits at 388 nm, sodium salicylate, which emits at 420 nm, and other commercially available fluorographic solutions and sprays. Commercially available scintillants and fluorographic products are available from several companies, including Amersham Corporation and DuPont NEN. The invisible latent image is the forerunner of the visible, metallic silver image that arises from photographic development after exposure to ionizing radiation. This darkening of the photographic film is the result of the latent image, on the silver halide grain, being converted to metallic silver by reducing agents in the developer. Two or more photons of light are required to make a latent image center on a silver halide grain, whereas only one mediumenergy β particle (from 14C or 35S) usually forms many latent image centers on numerous silver halide grains. Generally the latent image centers formed from the interaction with two or more light photons are not stable and fade via low-intensity reciprocity failure mechanisms. The latent image centers formed from direct β-particle interactions are very stable and do not exhibit fading from reciprocity failure (Hamilton, 1977). Detection of light emitted from scintillants is significantly improved by reducing the exposure temperature to −70° to −100°C. At these low temperatures, the latent image center forming in the silver grain becomes more stable, thus reducing latent-image fading. The detection sensitivity increases because the latent image accumulates with additional light exposure (see Table 10.11.2).
TranScreen Intensifying-Screen Exposure In this method, ionizing radiation is amplified by placing a thin and uniform phosphor coating between the radiolabeled specimen and autoradiography film. The TranScreen ~LE intensifying screen is used to enhance the detection sensitivity with low-energy radioisotopes such as 3H. The BioMax TranScreen LE intensifying screen folder is placed in the cassette and opened. Next, under safelight conditions, a sheet of BioMax MS film is placed inside the
Intensifying-Screen Exposure For use with an intensifying screen, the radioisotope must have sufficient energy to pass through autoradiography film and into the screen, which is located on the other side of Table 10.11.2
Exposure procedure Direct Fluorographic Fluorographic
Sensitivity of X-Omat AR Film
Temperature (°C)
Sensitivity (µCi/cm2)a
20 20 −70
8.00 0.30 0.07
aSensitivity is expressed as the amount of radioactivity required
Autoradiography
to produce a 0.10 optical density on X-Omat AR film (Eastman Kodak). [3H]Glutaraldehyde bis bisulfite was spotted on Kodak Chromagram sheet and exposed to film for 24 hours (Laboratory and Research Products, 1993).
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folder and the folder is closed on the film. The 3H-radiolabeled specimen is placed on top of the folder and the light-tight cassette is closed and exposed at the optimum exposure temperature—between −70° and −80°C. At the end of the exposure, the cassette is warmed to ambient temperature before opening it. The film is carefully removed from the TranScreen folder after noting the orientation of the film sheet with respect to the specimen by bending a corner of the film towards the specimen or by marking a corner of the film with a no. 2 pencil. The film is then photographically processed. BioMax MS film is spectrally matched to the BioMax TranScreen LE screen phosphor. The spectral output of the BioMax TranScreen LE is similar to BioMax MS intensifying screen. Using the TranScreen LE to detect 3H in a protein gel with western blot (10–4 cell fraction of a 10-atom label per protein of 30-kDa mass and a 10-µg gel load), the number of total discharges per mm2 required to produce a 0.10 optical density above background was reduced ∼16-fold, from 7.10 × 106 for direct exposure (Vizard et al., 1996) to 0.42 × 106 for TranScreen LE exposure.
PREFLASHING THE FILM For very-low-light autoradiographic applications, such as with fluorography, detection may be improved by hypersensitizing the film and exposing it at reduced temperature. Hypersensitizing the film is achieved by preflashing it prior to autoradiographic light exposure. For this treatment to be beneficial, light must be the primary source of the autoradiographic film exposure, as in fluorography or intensifying-screen procedures. Preflashing does not provide any advantages with direct β-particle exposure procedures (see discussion of Direct Exposure). Equipment needed for preflashing includes an electronic flash—e.g., Vivitar Auto 252 Electronic Flash or Sensitize Pre-Flash (Amersham), and light-filtering material. The Vivitar flash requires filtering the light through several sheets of Whatman 3-mm paper, a Kodak Wratten 22 gelatin filter, and a 3.0 neutral-density filter to control intensity. The electronic preflash unit from Amersham will most likely not require any additional filtering material. The preflash density is the film-density difference between the exposed and the unexposed film. The optimum increase in preflash density is 0.10 to 0.15 optical density units.
Preflashing produces the following various effects on photographic film. 1. It increases background density. 2. It lowers the contrast in the low-dose region of the film’s characteristic curve (the characteristic film curve is a graph of film response to light exposure as measured by optical density). 3. It increases in net density above background. 4. It produces a more linear density-versusexposure film response. Information will be lost if preflashing is not in the optimal range. Thus, it is very important to develop a repeatable and effective preflashing procedure. Amersham provides detailed comprehensive instructions and calibration protocols with their preflashing unit (also see Laskey and Mills, 1975).
FILM-PROCESSING RECOMMENDATIONS Autoradiography films are processed either in automated medical X-ray film processors or manually in 5-gallon tanks or small trays placed in a sink. With the exception of Direct Exposure Film (Eastman Kodak) and Hyperfilm-βmax (Amersham), most films are processable using the standard cycles on automated medical Xray film processors. Direct Exposure Film and Hyperfilm-βmax must be processed manually. It is best to standardize film processing by using one processor and one method of processing, because film development in a manual method is slightly different than automated film development in a medical X-ray film processor.
Automated Processing Automated medical X-ray film processing affords the most reliable and convenient processing of autoradiography films. Generally, the total processing time is <3 min. Besides the ease of use, most low-volume medical X-ray film processors produce high-quality autoradiograms.
Manual Processing There are two different methods of manual processing for autoradiography films. The difference is primarily in the size and configuration of the containers used to hold the processing reagents. Both methods will yield highquality autoradiograms. The deep-tank method involves several 5-gallon tanks surrounded by a temperature- and flow-controlled water bath. Film is attached to metal film hangers, and the Electrophoresis
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Table 10.11.3
Manual Processing Guidelines for Autoradiography Filmsa
Step
Solution
Agitation
Develop
Kodak GBX developer and replenisher
Do not agitate film sheet
Rinse
Kodak indicator stop bath or running water rinse
Fix
Kodak GBX fixer and replenisher
Wash
Clean running water with 2 changes every 15 min
Dry
aFor
Temperature (°C)
Time
20
5 min
Use continuous agitation
16 to 24
30 sec
Intermittently agitate film 5 sec for every 60 sec
16 to 24
5 to 10 min
16 to 24
10 min
Let stand in dustfree environment until dry
<50
X-Omat AR, BioMax MR, BioMax MS, and BioMax Light films (Eastman Kodak; see
SUPPLIERS
APPENDIX).
hangers are moved from solution to solution by hand. The tray method involves at least three photographic processing trays on a bench or in a sink. The trays must be 2- to 3-cm longer and wider than the film sheet. The processing chemistries, times, temperatures, and agitation specifications shown in Table 10.11.3 are a guide for manual processing of autoradiography film in either deep tanks or trays. The processing cycle indicated there is appropriate for most autoradiography, medical, and industrial films. Other types of developers and fixers may require different processing times and temperatures. Autoradiographic imaging that is consistent and quantifiable relies upon consistent and reproducible processing. The developer and fixer solutions must be replenished daily or made fresh every 1 to 2 weeks to maintain solution activity level.
Laskey, R.A. and Mills, A.D. 1975. Quantitative film detection of 3H and 14C in polyacrylamide gels by fluorography. Eur. J. Biochem. 56:335341.
LITERATURE CITED
Contributed by Daniel C. Bundy Eastman Kodak Company New Haven, Connecticut
Hamilton, J.F. 1977. The mechanism of formation of the latent image. In The Theory of the Photographic Process, 4th ed. (T.H. James, ed.) pp. 105-145. Eastman Kodak Company, Rochester, N.Y.
Steinfeld, R., McLaughlin, W., Vizard, D., and Bundy, D. 1994. Advances in autoradiography: A new film brings quality improvements. In The Biotechnology Report pp. 147-149. Campden Publishing Ltd., London. Vizard, D., Attwood, J., and McLaughlin, W. 1996. A universal intensifying screen system for enhanced detection of low- and high energy isotopes. Am. Biotechnol. Lab. 14(11):74-76.
INTERNET RESOURCES http://www.kodak.com:80/daiHome/SIS/ksis.shtml Site dedicated to scientific imaging systems and autoradiography products. Contains technical information, analysis software demos, and sales and service information for all Kodak Scientific Imaging Systems products.
Laboratory and Research Products. 1993. Autoradiographic Detection Principles. Eastman Kodak Company, Rochester, New York.
Autoradiography
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Digital Electrophoresis Analysis Gel electrophoresis has become a ubiquitous method in molecular biology for separating biomolecules. This prominence is the result of several factors, including the robustness, speed, and potentially high throughput of the technique. The results of this method are traditionally documented using silver halide–based photography followed by manual interpretation. While this remains an excellent method for qualitative documentation of single-gel results, digital capture offers a number of significant advantages when documentation requires quantitation and sophisticated analysis. Digital images of gel electropherograms can be obtained rapidly using an image-capture device, and the images can be easily manipulated using image analysis software.
REASONS FOR DIGITAL DOCUMENTATION AND ANALYSIS There are several reasons to consider digital documentation and analysis of electrophoresis results. These justifications can usually be categorized into issues of ease of handling, accuracy, reproducibility, and cost.
Ease of Handling A major advantage of the digital revolution has been in storage and retrieval of information. Storage in notebooks and filing cabinets previously meant that searching for specific data or experiments was a tedious manual process. With digital information, modern search engines can quickly find specific information in a fraction of the time usually required for a manual search. Making backup copies of nondigital data can be difficult, expensive, and time-consuming since it requires copying, retyping, or photographic reproduction. Copies of digital data can be generated more easily and at reduced costs. Manipulation of information is also easier when it is in a digital format. While the cut-andpaste analogy comes from physical documentation, it takes on a new perspective when applied digitally. Electrophoresis images can be resized, cropped, and inserted into reports. Data can be passed to spreadsheets and statistical packages for analysis and later insertion into notebooks and reports. These reports can be passed out via the Internet to colleagues throughout the world. A single individual can do all this in a few hours.
UNIT 10.12
Digital analysis also provides an easier method for handling the data when comparing large numbers of results or large numbers of separate experiments. Research that requires comparing the banding patterns on 1000 gels containing 50 lanes each can be an undertaking of heroic proportions if the analysis is performed manually. Database software can dramatically speed the analysis and handle the more mundane tasks, leaving the researcher free to interpret the data.
Accuracy The human eye is an extremely versatile measuring instrument. It can handle light intensities covering a range of nearly nine orders of magnitude and is sensitive to a fairly wide spectrum of light (Russ, 1995). Yet the eye cannot accurately and reproducibly quantitate density and patterns, nor can it deal with large numbers of bands or spots. Accuracy of measurement is a primary reason for using digital analysis on electrophoretically separated proteins and nucleic acids. Two categories of accuracy are key to digital analysis: positional accuracy, which is important for mobility determinations such as molecular weight, and quantitative accuracy. Positional accuracy is based on both resolution of the recording medium and measuring accuracy. Silver halide–based recording has a theoretical resolution based on ∼2000 imaging elements (silver grains) per inch. Measurement traditionally occurs using a ruler, with an accuracy of ∼20 to 40 elements (50 to 100 elements per inch). In comparison, typical digital systems have 200 to 600 picture elements (pixels) per inch. The advantage that digital systems have is in measuring accuracy, which can occur at the level of a single imaging element. Quantitative accuracy is also an issue. The amount of material represented by a band or spot is difficult to determine accurately from an image of a gel unless it is a digital image. On a digital image, the amount present is directly correlated with the derived volume of the band or spot—the volume is calculated using the intensity values of the pixels within the object.
Reproducibility Any technique or measurement is only as good as its ability to be faithfully replicated. With software-defined routines, measurements Electrophoresis
Contributed by Scott Medberry and Sean Gallagher Current Protocols in Protein Science (1998) 10.12.1-10.12.14 Copyright © 1998 by John Wiley & Sons, Inc.
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are performed in the same manner every time. Allowing the computer to do repetitive tasks and complicated calculations minimizes the chance for individual errors. This does not imply that such measurements are correct, just that they are reproducible. An incorrect routine or algorithm can also invalidate data.
Cost A consideration when evaluating any laboratory method is cost. Digital electrophoresis analysis equipment can be expensive. In many cases, however, it offers the only method for achieving acceptable analysis performance. In other cases, equal performance can be achieved using silver halide technology. However, traditional photography can also be expensive when the costs of consumable supplies such as film and developers, as well as other expensive requirements such as developing tanks and dark rooms, are included. Often, digital methods can be a good choice when all costs are considered.
KEY TERMS FOR IMAGING There are several specialized terms encountered during digital image analysis. The most commonly encountered are contrast, brightness, gamma, saturation, resolution, and dynamic range. They describe controls on how the light detectors report a range of light intensities. Below is a brief description of each.
Contrast
Digital Electrophoresis Analysis
Contrast describes the slope of the light intensity response curve. An increase in the contrast increases the slope of the curve. The result is a more detailed display over a narrowed range of intensities with less detail in the remaining portions of the intensity range. This is depicted in Figure 10.12.1A and 10.12.1B, where a normal, unadjusted image and a contrast-adjusted image are displayed, respectively. The contrast was increased on midrange intensity values in Figure 10.12.1B to highlight band intensity differences at the expense of background information. Images with a narrow range of informative intensities can benefit from increasing the contrast, since that effectively increases the scale and improves detection of minor differences in intensity. Contrast settings should be lowered if information is being lost outside of the contrast range. For example, in Figure 10.12.1B, loss of background information between peaks indicates that this image should not be used for quantitation.
Brightness While brightness can have many different definitions, only one will be considered here. Brightness shifts the light intensity response curve without changing its slope as is shown in Figure 10.12.1C. Another name for brightness is black level, since it is commonly used to control the number of black picture elements (pixels) in an image. Incorrect brightness levels can lead either to high background and potential image saturation or, as is illustrated in Figure 10.12.1C, to a total loss of background information and partial loss of band information.
Gamma Nonlinear corrections are often applied to images to compensate for how the eye perceives changes in intensity, how display devices reproduce images, or both. The most common correction is an exponential one, with the exponent in the equation termed the gamma. A typical gamma value for camera-based systems is 0.45 to 0.50, and is illustrated in Figure 10.12.1D. This is a compromise value that compensates for the 2.2 to 2.5 gamma present in most video monitors and the print dynamics of most printers. Since it is a nonlinear correction, special care must be taken if quantitation is desired. Unless directed to by the manufacturer, gamma values other than 1.0 should be avoided when quantitating. More information on gamma correction can be found on Poynton’s Gamma FAQ (www.inforamp.net/∼poynton/Poynton-color.html).
Dynamic Range Dynamic range describes the breadth of intensity values detectable by a system and is usually expressed in logarithmic terms such as orders of magnitude, decades, or optical density (OD) units. A large dynamic range is important when trying to quantitate over a wide range of concentrations. The most accurate quantitation occurs in the linear part of the dynamic range, which is usually not the complete dynamic range of the system. An additional consideration is the dynamic range of the visualization method. Many popular visualization methods have linear dynamic ranges of 1 to 2.5 orders of magnitude. An imaging system with greater dynamic range analyzing the results of such a visualization method will not improve the dynamic range.
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A
normal
Output
100
0 0
Input
B
100 200 300 400 500
contrast
Output
100
0 0 100 200 300 400 500
Input
C
brightness
Output
100
0 0 100 200 300 400 500
Input
D
gamma
Output
100
0 0
Input
E
100 200 300 400 500
saturation 100
0 0
100 200 300 400 500
Figure 10.12.1 Examples of how altering image capture settings affects the image and the analysis. The graph on the left displays the light intensity response curve used for image capture while the image and resulting lane profile on the right display how the setting affects the image. The lane profile displays pixel position versus normalized pixel intensity (A). In this case, the output has not been altered, giving a straight line with a slope of 1 on the response curve. (B) The image acquisition was adjusted to increase the contrast of the displayed image. Although useful for images with a narrow range of informative intensity values, increasing the contrast can lead to a loss of low and high values. (C) Decreasing the brightness reduces peak values but also leads to a loss of the weak bands and original background. (D) Gamma adjusts raw data to appear more visually accurate. Note that this leads to a loss of fidelity between the adjusted image and the original. (E) Saturation indicates that the detector is reporting its maximum value or that the dynamic range for the visualization method has been exceeded.
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high-resolution peaks
medium-resolution peaks
42 µm
Intensity
168 µm
840 µm
0
100 Pixel position
Figure 10.12.2 The effect of spatial resolution on the ability to detect closely spaced objects. Whole-cell protein lysates from E. coli were separated using SDS-PAGE and visualized with Coomassie blue staining. An image was captured at 42 µm (600 dpi), 168 µm (150 dpi), and 840 µm (30 dpi) from a segment of the lane, and a lane profile was generated for each image. The lane profiles have offset intensities to allow for comparison. Only major bands can be detected with the low-resolution image (840 µm); at higher resolutions more bands are detectable.
Saturation Saturation occurs when a detector or visualization method receives input levels beyond the maximum end of the dynamic range. This results in a loss of detail and quantitative information from those data points that are saturated. For fluorescent and luminescent samples, reduction in the sampling time can sometimes correct saturation problems. Optical densitybased visualization techniques can also generate saturated images, as is illustrated in Figure 10.12.1E; this can sometimes be avoided with longer sampling times or increased detection source intensities. More often, it will be necessary to perform another electrophoresis with more dilute samples or to alter the visualization process to generate a less optically dense material.
Resolution
Digital Electrophoresis Analysis
Resolution is the ability of a system to distinguish between two closely placed or similar objects. Three types of resolution are important for analysis—spatial resolution, intensity resolution, and technique-dependent resolution. Spatial resolution is the ability to detect two closely placed objects in one-, two-, or three-
dimensional space. It is most accurately described as the closest distance two objects can be placed and still be detected as separate objects. In practice, it is often defined nominally in terms of the number of detectors per unit area such as dots per inch (dpi) or the number of detectors present in total or in each dimension such as 512 × 512 (262,144 total detectors). Actual resolution is less than half the nominal resolution due to the need for two detectors for every resolvable object (one for the object and one for the separation space) and the effects of optical resolution. Figure 10.12.2 demonstrates how spatial resolution can affect detection of objects. The 42-µm resolution image allows detection of closely spaced bands, the 168-µm resolution image detects fewer bands, and the 840-µm resolution image detects only major bands. For instruments with on-line detection systems, a pseudo spatial resolution is often reported in units of time from the start of the separation or the time interval between two objects crossing the detection path. Intensity resolution is the ability to identify small changes in intensity. It is a function of both the dynamic range of a detector and the number of potential values that detector can
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report. Greater dynamic range decreases the intensity resolution of a given detector. The number of potential values a detector reports is described by its bit depth. An 8-bit detector can report 256 (28) different possible values, while a 12-bit detector can report 4,096 (212) values, and a 16-bit detector can report 65,536 (216) values. The higher the bit depth, the greater the intensity resolution. Technique-dependent resolution directly affects the spatial and intensity resolution. Electrophoretic separation techniques that generate overlapping objects or that have object separation distances shorter than the spatial resolution will fail to provide reliable data. Many factors, including the amount of sample loaded, gel pore size, buffer constituents, and electrophoresis field strengths, can dramatically affect separation and resolution of biomolecules. Likewise, detection methods that can only generate a small range of discrete intensity values will not benefit from systems with improved intensity resolution.
IMAGE CAPTURE Devices Capturing digital images involves a detection beam or source, a sensor for that beam or source, and some method of assembling a twodimensional image from the data generated. Most systems use a light source for detection. The light wavelengths used range from ultraviolet (UV) to infrared (IR) and can be broad spectrum or narrow wavelength. Broad-spectrum detection is more versatile since it can often be used for more than one detection wavelength. However, when compared to narrowwavelength sources such as lasers, broad-spectrum detection suffers from reduced sensitivity and reduced dynamic range. While many types of light sensors have been used, including charge-coupled devices (CCDs), charge-injection devices (CIDs), and photon multiplier tubes (PMTs), technology advances in CCDs have led to their dominance. CCDs are semiconductor imaging devices that convert photons into charge. This charge is then read and converted into a digital format via an analogto-digital converter (ADC). The method of image assembly depends on the light source and detector geometry. One method is to capture the image all at once using a two-dimensionally arrayed CCD detector similar to the detectors found in digital and video cameras. Typically a camera-type sensor is paired with a light source that evenly illumi-
nates the sample. This same sensor is often used with fluorescent and chemiluminescent detection methods, as its ability to detect light continuously over the entire sample reduces image capture times. Another method of image assembly is to capture the image a line at a time. This typically involves a linearly arrayed CCD scanning slowly across the sample in conjunction with the detection beam of light. The data from each line is then compiled into a composite image. Spatial resolution in this method can be significantly better on large-format samples than the resolution of a camera-based system. This method is also advantageous when ODbased detection is used, since the more focused light beam is usually of higher intensity and can penetrate denser material. A third method of image assembly is to use a point light source and single-element detector on each point on a sample. The image is then compiled from each point sampled. This method is slower than the others but can offer extremely high resolution and sensitivity. A fourth commonly encountered m eth od is that of generating a pseudoimage of electrophoresis results through the use of a finish line type of detection system. This is comprised of a light source positioned at the bottom of the gel (i.e., the end opposite of the site of sample loading) and light detectors positioned next to each lane to detect the transmitted light or emitted fluorescence. A lane trace is generated using time on the x axis and light intensity on the y axis. The pseudoimage is then generated from this data (Sutherland et al., 1987).
Capture Process Prior to image capture, electrophoretic separation and any visualization steps are performed. To calibrate the separation process, standards are usually run at the edges of the gel and often at internal positions. If quantitation of specific proteins or nucleic acids is to be performed, a dilution series of standards with similar properties to the experimental samples should also be included. After separation, the protein or nucleic acid is visualized if necessary. Visualization can include binding of a fluorochrome or chromophore such as Coomassie blue, precipitation of metal ions such as copper, silver, or gold, enzymatic reactions, and exposure of film or phosphor screens to radiant sources. These methods can be grouped based on the type of detection into optical density, fluorescence, chemiluminescence, and radioactivity. The suitability of popular detection devices with these methods
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Table 10.12.1 Compatability of Popular Image-Capture Devices with Common Visualization Methodsa
Image-capture device Visualization method Optical densityb Fluorescence Chemiluminescence Radioactivity
Silver halide photography
CCD camera
Desktop scanner
Storage phosphor
Fluorescent scanner
+
+
++
−
−
+ + +
+ ++ −
− − −
− ± ++
++ − −
aThe device with the highest sensitivity and greatest dynamic range for a visualization method is marked with a ++,
other devices that can detect this visualization method are indicated with a +, and devices that are not suitable for a visualization method are indicated with a −. A ± indicates that only some devices of this type can be used with this visualization method. bOptical density methods include Coomassie blue staining.
Digital Electrophoresis Analysis
is described in Table 10.12.1. Once visualization has occurred, image capture consists of the following steps: previewing the image while adjusting capture parameters, capturing the image, and saving the image for later analysis. During the preview process, capture parameters are optimized for data content and for ease and rapidity of later processing steps. Typically, the first step is to place the sample so that when the image is captured, the rectangular edges of the gel are horizontal and vertical on the monitor and any lanes are either horizontal or vertical. Since band and spot detection will be much easier if the image is properly oriented, this eliminates the need to later rotate the image digitally. Image rotation is time-consuming and can result in spatial linearity errors (a change in the size and shape of objects in image) caused by rectangular image-capture device geometries. The next step for camera-based systems is to adjust magnification and to focus the sample image. For thicker samples, it might be necessary to reduce the aperture on camerabased systems to get a sufficient depth of field to focus the entire sample. Often at this point image imperfections—e.g., dust, liquid, or other foreign objects that will detract from later analysis—are detected, and they need to be removed. Next, image intensity is set. Within the area of interest on the image, band or spot peaks should have values less than the maximum saturated value, and the background should have nonzero values. This is usually accomplished through adjustment of the lightsource intensity or the sensor signal integration. If the device allows precapture optimization of other parameters such as spatial resolution, contrast, brightness, gain, or gamma, they are adjusted next. Note that this only applies to
controls that affect the response of the sensor or processing of the image prior to a data reduction step and not to controls that affect the image at later stages. The latter process can enhance visualization of specific features but is best left to adjustments in look-up tables (LUTs) in later analysis steps rather than during image capture, since there is a risk of data loss during postacquisition image processing. LUTs are indexed palettes or tables where each index value corresponds to color or gray-scale intensity values present in an image. Many image analysis programs alter LUTs instead of image values directly, since it both is faster and does not change the original image data. Once all the capture parameters are optimized, the image capture process is initiated. This might take less than a second for images captured with camera-based detectors and up to hours for scanning single-point detectors. When the image has been captured, it should be carefully examined for content. It should fully capture the area of interest and the parameters should have been set so that all necessary information is detectable. Furthermore, it should be in a form that will allow for easy analysis. Extra time spent optimizing the capture parameters will often result in a reduction in total image analysis time and in an increase in data quality. When the best possible image has been captured, it often contains information outside the area of interest. While this is unlikely to cause problems with later analysis, it is often advantageous to crop the image so that the only portion that is saved contains the area of interest. This reduces the amount of disk space necessary to store the image, and the image usually will load and analyze faster with the analysis software.
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The last step in image capture is to save the image. Several options are available at this point, including choosing which location to save the image at, what file type or format to use, and whether to use some form of compression. The location where the image is saved is not as trivial a question as it might seem if the image will need to be transferred to another computer at some point. File sizes can easily exceed 15 megabytes on high-resolution images. This is a manageable size for hard drives but exceeds current floppy drive sizes by an order of magnitude. There are software utilities available that will subdivide files into disk-size chunks and then reassemble them at the next computer, but this is an inconvenient and slow method. If the computer used to help capture the image is connected to a network, the image files can easily be transferred this way or potentially saved on a central server. Alternatively, several types of high-capacity removable media are available (e.g., Zip or Jazz). This usually requires the installation of additional hardware onto two or more computers but does make backing up data easier. Since image files can be very large, compression techniques are sometimes used to reduce disk space requirements. Compression algorithms use several methods, typically by replacing frequent or repetitive values or patterns with smaller reference values and by replacing pixel values with the smaller difference values describing the change in adjacent pixels. When the file is later decompressed, the compressed values are then replaced with the original information. Not all images compress equally, with simple images containing mostly repetitive motifs compressible by ≥90%, while complex images will benefit much less from compression. Because compression is a much slower method of saving files and will not benefit every file, compression is not used to save all files. Several different forms of compression are available but are separable into two main classes, lossless and lossy. Lossless methods faithfully and completely restore the image when it is decompressed (no loss of data) but offer only moderate file compression; compression values range from ∼10% to 90%, depending on the image. Examples of lossless compression include Huffman coding (Huffman, 1952), RLE (Run Length Encoding), and LZW (Lempel, Ziv, and Welch; Welch, 1984). In comparison, lossy methods such as JPEG (Joint Photographic Experts Group), MPEG (Moving
Picture Experts Group), or fractal compression schemes can reach compression values of ≥98% (Russ, 1995). The trade-off is that not all information from the original file is recovered during decompression. Lossy compression is sometimes necessary for applications with extremely large image files such as real time video capture, but it usually represents an unacceptable loss of data if used with electrophoresis image capture. Many different file types have been developed to store digital images. Some of these file types are proprietary or hardware specific. For example, PICT is a Macintosh format and BMP is a PC-compatible format. Each file type has its own structure. Some types do not allow compression, for others it is optional, and for some it is mandatory. File types vary in the types of images they support, particularly in the number of colors or gray levels. Below is a brief description of a few of the more prevalent file types. TIFF (Tagged-Image File Format) is one of the most commonly used formats. It is particularly versatile since it is an open format that can be modified for specific applications. One reason for its versatility is the ability to attach or tag data to the image. The tags can include information such as optical density calibration, resolution, experimenter, date of capture, and any other data that the application software supports. TIFF images can be monochrome, 4-, 8-, or 16-bit gray-scale, or one of many colorimage formats. Compression is optional, with LZW, RLE, and JPEG often supported (Russ, 1995). Since TIFF is supported by both Macintosh and PC computers, it is a good choice for multiple-platform environments. The versatility of TIFF can also be a weakness. Since there are many different tagging schemes and since not all programs support all possible compression and color schemes, it is sometimes not possible for one program to access the information in a TIFF file generated by a different program. GIF (Graphics Interchange Format) is a file format that is widely encountered on the Internet due to its compactness and standardization. Its compactness is attributable to a mandatory modified LZW compression. Another feature of GIF is the use of a LUT to index the values in the image. One interesting ability of GIF is that it supports storing multiple images within a single file. This can offer some advantages for applications such as time-lapse image capture. A GIF image can contain no Electrophoresis
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more than 256 individual color or gray levels and does not support intensity resolutions higher than 8 bit. In addition, since the image is implemented as a LUT, it also is not a true gray-scale image. Due to these limitations and others, alternative formats such as PNG have been developed to replace GIF. PICT is a file format and graphics metafile language (it contains commands that can be played back to recreate an image) designed for the Apple Macintosh. It can contain both bitmap images and vector-based objects such as polygons and fonts. It supports a ≤256-graylevel LUT, and monochrome images can be RLE compressed. Because it only offers a 256gray-level LUT, it has the same weaknesses that GIF does with true gray-scale and high-intensity-resolution images. In addition, any vector objects in the image are difficult to translate on a PC since they are designed to be interpreted by Macintosh QuickDraw routines. BMP is the native bitmap file format present on Windows-based PCs. It supports 2-, 16-, 256-, or 16-million-level images. With images of ≤256 gray levels, it implements a LUT, while the highest-resolution image is implemented directly. RLE compression is optional for 16and 256-gray-level images. Since compression is prohibited on 16 million-gray-level images and there is no intermediate level supported beyond 256 levels, BMP is not a good choice for images with high-intensity resolution requirements.
ANALYSIS
Digital Electrophoresis Analysis
Once the image has been captured, the data needs to be analyzed and distilled into information about the results of the electrophoresis experiment. Through the use of standards and experimenter input, this software-driven process can estimate mass and quantity of objects in an image and detect relationships between objects within one image and between similar images. The type of software used depends on the analysis to be performed. Images from single electrophonetic separations are examined by one-dimensional analysis software optimized for lane-based band detection. Images from two-dimensional electrophoresis are best handled by specific programs designed to detect spots and to assign two mobility values and a quantity value to the spot. After the initial characterization of bands and spots, comparisons are often made between bands or spots from different experiments through the use of database programs and matching algorithms.
Software for One-Dimensional Analysis Lane positioning For one-dimensional analysis, the first activity is to detect the lanes on the image. One of three different methods is commonly employed for this. For images with straight, welldefined lanes with a large number of bands, automatic lane-detection algorithms can quickly and accurately place the lanes. On images with very well-defined lanes, such as pseudoimages from finish-line type electrophoresis equipment, automated lane calling based on image position is possible. For images with “smiling,” bent, or irregular lanes, manual positioning of the lanes is often the fastest and most accurate method of lane definition. Regardless of the method of identifying the lanes, the lane boundaries need to be carefully set for accurate quantitation and mass determinations. Lane widths should be wide enough so that the entire area of all bands in that lane are included, but they should not be so wide as to include bands from adjacent lanes. To accomplish this, curved or bent lanes might need to be used in order to follow the electrophoresis lane pattern. Lane length and position also must be adjusted as necessary so that all bands of interest are included. If mass determinations are necessary, the sample loading point should probably also be included in the lane or be the start of the lane. At this point, lines of equal mobility (often called Rf or iso-molecular-weight lines) are added to the image as necessary. These lines allow for correction of lane-to-lane deviations in the mobility of reference bands and generate more accurate measurements of mass. A similar form of correction is also possible for withinlane correction of mobilities. This correction is important for accurate detection and quantitation of closely spaced bands. Band detection Once the lanes have been defined, the bands present in each lane need to be detected. There are many methods for detecting bands. One method is to systematically scan the lane profile from one end to the other, identifying regions of local maxima as bands. Another common method is to use first- and second-order derivatives of the lane image or lane profile in order to find inflection points in the change of slope in pixel intensity values (Patton, 1995). Regardless of the method used, it is often necessary to alter the search parameters so that they perform reliably under a given experimental condition.
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Typical search parameters include ones for detection sensitivity, smoothing, minimum interband gaps, and minimum or maximum band peak size. Smoothing reduces the number of bands detected due to noise in the image. A minimum interband gap is often used to avoid detection of false secondary bands on the shoulders of primary bands. Limitations on peak sizes, especially for within-lane comparison to the largest band’s peak, can be a useful way to allow sensitive detection of bands in underloaded lanes without detecting false bands in overloaded lanes. Band edges are often detected in addition to band peaks in order to further define bands or to quantitate band amounts. This can be accomplished by using local minima, derivatives of the lane profile, or fixed parameters such as image distances or a percentage of band peak height. The band edges can be applied as edges perpendicular to the long axis of the lane or as a contour of equal intensity circling the band. The perpendicular method is advantageous for bands with uneven distribution of material across the face of the band, while the latter method is better for “smiling” or misshapen bands. Background subtraction With nearly all electrophoresis procedures, the most informative images have a low level of signal intensity at each pixel that does not result from protein or nucleic acid. Instead, this background intensity is attributable to the gel medium, the visualization method, electronic noise, and other factors. Since this background tends to be nonuniformly distributed throughout the image, failure to subtract it can make band detection and quantitation less accurate. Many methods of background subtraction are possible. Sometimes it is possible to generate a second image under conditions that do not detect the protein or nucleic acid. The second image is then digitally subtracted from the data-containing image to remove the background. More often, background information must be obtained from a single image. If the background varies uniformly across the image, a line that crosses the variation can be defined at a point where no bands are present. The intensity values at each point on the line can be used as the background value for the pixels perpendicular to the line at that point. Commonly, background is also present as variations in intensity along the long axis within each lane. One simple method is to take the lowest point in the lane profile as the background. Another
method is to use an average value of the edge of each band as the background for that band. More complicated methods such as valley-tovalley and rolling-disk use local minima points in the lane profile to define a variable background along the length of the lane. Because there can be many different causes and distributions of background, no single method of background determination can be recommended for all experiments. Characterization Once lanes and bands have been detected, it is possible to interpret the mobility of the nucleic acid or protein bands. Depending on the method of electrophoretic separation, information on mass (length or size), pI, or relative mobility (Rf) can be inferred from mobility information. The mobility is characterized using a standard curve with internal standards of known properties. The type of curve depends on several parameters. By definition, with Rfbased separation, a linear first-order curve is used since it represents the linear relationship between mobility and Rf. Similarly, pI and mobility are generally linear in isoelectric focusing separations. For separations based on size, a curve generated from mobility versus the log of the molecular weight provides a relatively good fit as measured by the correlation coefficient (R2). Several other curves have been suggested for size-based separations, including modified hyperbolic curves and curves of mobility versus (molecular weight)2/3 that have good correlation coefficients (Plikaytis et al., 1986). In some cases, no single curve equation can adequately represent the data, and methods of fitting smooth contiguous curves using only neighboring points, such as a Lagrange or spline fit (described in Hamming, 1973), are necessary. This is most common for size separations with a very large range of separation sizes and with nonlinear gradient gels. Care must be taken with multiple-curve techniques since they rely on only a few data points for any one part of the composite curve, and outlying data points can drastically affect the outcome. For size and Rf determination, a uniform position must be found in each lane as a point from which to measure the mobility of each band. Many software analysis packages use the end of the lane as the measuring start point, so for them it is important to position each lane start point at an iso-molecular-weight or iso-Rf point. A convenient point is the well or sampleloading position since it is usually easily de-
Electrophoresis
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tectable and at an equal mobility position in each lane. A consistent point on each band must also be chosen to measure mobility. A band’s peak is easily defined in digital image analysis and is commonly used. Since peak positions are harder to detect visually than edges on silver halide images, the leading band edge is sometimes used when comparing digital results with silver halide–based results. Once lanes and bands have been detected, it is also possible to quantify the amount or at least relative amount of nucleic acid or protein present in each band. The amount in a band is related to the sum total of the intensity values of each pixel subtracted by the background value for each pixel in a band. For absorptively detected bands, intensity values are converted to OD values. The total value that is calculated is equivalent to the volume of the band and can be directly compared to other bands that are within the linear range for the visualization method. If standards of known amounts are loaded onto the same gel, they can be used to generate a standard curve that converts band volume into standard units such as micrograms. For greatest accuracy, it is important to be able to generate multiple standard curves when using visualization methods, such as Coomassie blue staining, that are affected by band or spot composition. Quantitation becomes more complicated when bands are not fully resolved. In this case, material from one band is contributing to the volume measurement of an adjacent band and vice versa. The simplest method for handling this is to partition into each band only the volume within its edges. Alternatively, a Gaussian curve can be fitted to each band and the volume contained within the curve used to estimate the amount of the band. Since most electrophoresis bands have a pronounced skew towards the leading edge of the band, modified Gaussian curves have also been used (Smith and Thomas, 1990). In either case, the curvefitting process is calculation intensive and can significantly increase analysis times for images with many bands.
Software for Two-Dimensional Analysis
Digital Electrophoresis Analysis
In two-dimensional analysis, the first-dimension separation is performed in a single column or lane and then a second separation is performed perpendicular to the first. The result after visualization is a rectangular image of up to 10,000 spots. The most common two-dimensional gel type is one in which protein is sepa-
rated first by apparent pI and second by molecular weight, although two-dimensional separation of nucleic acids is also possible. While many of the concepts and analysis techniques used with one-dimensional gels are applicable to two-dimensional gels, the complex nature of most two-dimensional gels requires somewhat different methodology. For example, spots are more difficult to detect since they are not conveniently arranged in lanes and can vary more in shape and overlap than bands. In addition, two-dimensional experiments usually require some method of comparing between two images, whereas one-dimensional images usually contain all of the information from an experiment. Spot detection Probably the most difficult aspect of two-dimensional analysis is efficient and accurate spot detection. If it is incorrectly done, it can lead to hours of manual editing. Due to the complexity and computational intensity of some algorithms, the detection process itself can last hours on relatively fast desktop computers. One theoretically effective but computationally intense method is to treat the image as essentially a three-dimensional image with spots treated as hills and background as valleys. A large number of Gaussian curves are then combined to describe the topology of the image. Many other methods make use of a digital-imaging technique known as filtering. In essence, filtering is a way to weight the value of a pixel and its neighbors in order to generate a new value for a pixel. By passing a filter across an image pixel by pixel, a secondary image is generated. Filters can be designed for many tasks, including sharpening an image or removing high-frequency noise. Filters can also be generated to help detect spots by making images that are first and second derivatives of the original image. The derivative images indicate inflection points in the intensity pattern and can be used to detect spot centers and edges. In a different method, called thresholding, filters can be used to detect the edges of objects. Instead of looking for inflection points, threshold filters identify intensities above a set level or at ratios between central and edge pixels above a set value. Since the edges on two-dimensional spots tend to be diffuse, sharpening filters are sometimes used prior to the thresholding filter. In some cases, multiple techniques are used to detect spots (Glasbey and Horgan, 1994).
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Unlike one-dimensional detection, detection on images of two-dimensional experiments usually requires secondary processing to get acceptable performance. One example of a secondary process is to discard spots with sizes below a set minimum or above a set maximum. Another is to analyze spots that are oval for possible splitting into two spots. Even after secondary processing, it is likely that a small amount of manual editing will be necessary. When manually editing an image, care must be taken to use as objective criteria as possible, especially if two or more images are to be matched and spot volumes compared. Characterization In a two-dimensional system, determination of protein or nucleic acid mobility is complicated by the fact that there are two mobilities to account for and that the second-dimension separation tends to make estimation of the separation which occurred in the first dimension more difficult. One method for dealing with this is to have a series of markers in the sample that, after both separations are completed, are evenly distributed within the gel and image. It is also possible to estimate separation characteristics from calibration points located at the periphery of the gel. For example, distance measurements can be used to pass calibration data from the first dimension separation, and standards can be separated at the ends of the gel to calibrate for mobility in the second dimension. Regardless of the method used, in many instances, a series of related images will be examined and similar spots in each image will be matched. When this occurs, it is possible to calibrate one image and then pass the calibration information via the matches to the related images. Quantity determination is similar in many regards to that which occurs in one-dimensional analysis, but there are some differences. If spot edges are detected, a simple method of determining spot volume is to take the sum of the intensity value of each pixel in the spot reduced by a background intensity value. Multiple Gaussian curves can also be fitted to the spot to approximate the volume (Garrels, 1989). More difficult is attempting to compensate for a skewed distribution in a size-separating dimension while trying to use a regular Gaussian fit for a pI separation, such as is encountered with the most common form of two-dimensional protein separations. The distribution of background makes quantitation more difficult in two-dimensional gels. There is no lane-dependent component, so it is nec-
essary to use other methods such as image stripes, finding local minimum values or using values derived from the spot edges to determine background values.
Matching Matching is the process in which proteins or nucleic acids with similar separation properties are linked or clustered together. Matching can occur within one image or between multiple images as long as a frame of reference is established. Matching allows for comparisons between samples. It also makes annotation and data entry easier, since if one spot or band is matched to others and is characterized or annotated, this information is easily passed to all the other matches. An underlying assumption of matching is that objects with similar separation properties are actually similar. Care must be taken to confirm the identity of matched spots or bands by other methods on critical experiments. A simple form of matching is to link bands or spots at similar positions on the gel images. This works well when separation and imaging conditions are uniform. This is very seldom the case, since slight differences in the electrophoresis, visualization, and imaging conditions across a gel and between gels generates incorrect matching with this method. Since bands on one-dimensional gels are relatively easy to calibrate for mobility, matching can occur along contours of equal mobility. This dramatically decreases but does not eliminate the variability in detecting similar bands. Much of the remaining variability can be attributed to calibration errors. This error can often be compensated for by allowing a small tolerance in mobility values in determining whether a band is matched or not. Because of the difficulties in calibrating mobility in two-dimensional gels, it is often more practical to use matched spots for calibrating mobility than vice versa. Spot matching between two two-dimensional images starts with finding a small number of landmark spots that are used as seeds for subsequent matches (Appel et al., 1991; Monardo et al., 1994). There are many methods for finding the landmarks in both images, including finding the highest-intensity spots, finding spots in unique clusters, and manual positioning. The most common procedure from this point is to derive a vector that describes the direction and extent of the path from one matched spot to the other when the two images are superimposed. The vector is used as the basis for finding more matches near the landmark matches. To allow
Electrophoresis
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Figure 10.12.3 Example of a dendrogram generated from similarity data on band matching between lanes. DNA samples from 22 isolates of Listeria were subjected to Random Amplification of Polymorphic DNA (RAPD) analysis and the resulting electrophoresis image was analyzed with ImageMaster software (Amersham Pharmacia Biotech). Clustering was performed using the Dice coefficient with a tree structure based on the Unweighted Pair-Group Method using Arithmetic Averages (UPGMA). Similarity values between isolates can be determined by locating the node that connects the isolates and reading the value from the scale on the lower left edge of the dendrogram.
Digital Electrophoresis Analysis
for error, the area within a small radius is searched extending from the end of the vector. Once another match is found, its vector is computed and used as the starting point for finding neighboring matches. From this progression, the entire gel is matched. If all vectors are displayed graphically when matching is complete, questionable matches can often be identified as vectors that are significantly different from neighboring vectors. One specialized use of matching is as an estimator of the similarity and potential genetic relatedness of organisms. For example, on a one-dimensional gel image, a ratio of the matched to unmatched bands for each pairwise combination of lanes can be calculated. This
ratio can be used as an indicator of similarity, with values near 1 indicating a pair of highly similar lanes, and values near zero indicating very dissimilar lanes. Assuming that the contents of the lanes are valid samples of the originating organism’s genetic makeup, the information on lane similarity can be converted to estimates of genetic similarity. A convenient way to display this similarity data graphically is to generate a dendrogram with similar objects close to each other and less similar objects more distantly placed. An example of such a dendrogram is presented in Figure 10.12.3, where samples from Listeria isolates are arranged based on banding pattern.
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Databases In many cases, image analysis is not the last step in the process. The image and analysis data need to be archived in a searchable format. There may be a need to analyze the data from multiple experiments conducted at different sites or in laboratories around the world. Bioinformatic links to diverse data sources might be desired to help develop a unified understanding of the biology behind particular phenomena. When these situations arise, database programs can be utilized to store, link, and search image analysis results. As the number of images that are captured and analyzed grows, it becomes increasingly more difficult to find particular information from the large number of files that are stored. Relatively simple databases can be used if the major requirement is to find previously analyzed images and associated data. Such databases often display a miniaturized version of each image to aid in visual scanning for the file as well as simple searching for image-specific information such as date of analysis, file name, or other information that was entered at the time of image capture. More powerful database products are also available that can perform complex searches on data generated during the analysis. For an example, a search on a two-dimensional database might include finding proteins exhibiting a specific expression profile and having a molecular weight >20 kDa with a pI between 3 and 5 or 8 and 10 with an amount <50 ng in a series of experiments conducted <1 year ago. Such searches can quickly target potentially interesting molecules for further analysis. With the increasing ease of transferring data through the Internet as well as local- and widearea networks, it has become practical to quickly find and examine data from distant locations. Of course, great care must be taken to ensure that similar experimental conditions are employed, as otherwise the results will be difficult to compare. In this manner it is sometimes possible to dramatically increase the sample size and statistical accuracy as well as the probability of detecting rare events. In addition, if one data set is more completely characterized, this extra information can be extracted and applied to the other data set. For example if there is a band in common in two databases and there is sequence information for it in one database, that sequence information can be added to the other database. Currently most public electrophoresis database sites are twodimensional protein databases. A list with links
to many of these Internet database sites can be found at http://www-lmmb.ncifcrf.gov/EP/table2Ddatabases.html. With biological questions becoming more complicated and the answers to the questions often requiring information from a variety of sources, it is becoming increasingly important to be able to move easily between information sources. A relational type of database can help achieve this. Unlike a conventional database with a fixed arrangement of data, relational databases have links between related files that allow for easy movement from one file to another. Another approach to interconnecting electrophoresis data with data from other sources is to generate a series of hypertext links between data sets, similar to what occurs on the Internet. Selecting a specific link moves the search to the related network site and the related information. Regardless of the method, the end goal is similar. An example of what is possible: a researcher selects a protein spot on a two-dimensional gel image, which triggers accessing of related information on this protein. The protein sequence is accessed from mass spectroscopy analysis of the spot on a separate gel. The sequence of the gene and the cDNA that generated the protein is retrieved. The expression pattern of the gene in various tissues and conditions, as well as information on similar genes in other organisms, is incorporated. Citations and annotations to this are retrieved as well. All of this information is compiled automatically into an interactive report about the protein. From this report, the researcher can formulate a more refined hypothesis and plan the most appropriate experiments to test it.
LITERATURE CITED Appel, R.D., Hochstrasser, D.F., Funk, M., Vargas, J.R., Muller, A.F., and Scherrer, J.-R. 1991. The MELANIE project: From a biopsy to automatic protein map interpretation by computer. Electrophoresis 12:722-735. Garrels, J.I. 1989. The QUEST system for quantitative analysis of two-dimensional gels. J. Biol. Chem. 264:5269-5282. Glasbey, C.A. and Horgan, G.W. 1994. Image Analysis for the Biological Sciences. John Wiley & Sons, Chichester, England. Hamming, R.W. 1973. Numerical methods for scientists and engineers, 2nd ed. Dover Publications, New York. Huffman, D.A. 1952. A method for the construction of minimum-redundancy codes. Proc. Inst. Elect. Radio Eng. 40:9-12. Electrophoresis
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Monardo, P.J., Boutell, T., Garrels, J.I., and Latter, G.I. 1994. A distributed system for two-dimensional gel analysis. Comput. Appl. Biosci. 10:137-143. Patton, W.F. 1995. Biologist’s perspective on analytical imaging systems as applied to protein gel electrophoresis. J. Chromatogr. A. 698:55-87. Plikaytis, B.D., Carlone, G.M., Edmonds, P., and Mayer, L.W. 1986. Robust estimation of standard curves for protein molecular weight and linear-duplex DNA base-pair number after gel electrophoresis. Anal. Biochem. 152:346-364. Russ, J.C. 1995. The Image Processing Handbook. CRC Press, Boca Raton, Fla. Smith, J.M. and Thomas, D.J. 1990. Quantitative analysis of one-dimensional gel electrophoresis profiles. Comput. Appl. Biosci. 6:93-99. Sutherland, J.C., Lin, B., Monteleone, D.C., Mugavero, J., Sutherland, B.M., and Trunk, J. 1987. Electronic imaging system for direct and rapid quantitation of fluorescence from electrophoretic gels: Application to ethidium bromide– stained DNA. Anal. Biochem. 163:446-457. Welch, T.A. 1984. A technique for high performance data compression. IEEE Computer. 17:21-32.
KEY REFERENCES Glasbey and Horgan, 1994. See above. Describes general image-processing techniques as they are applied to biological images.
Sutherland, J.C. 1993. Electronic imaging of electrophoretic gels and blots. In Advances in Electrophoresis, Vol. 6. (A. Chrambach, M.J. Dunn, and B.J. Radola, eds.) pp. 1-41. VCH Verlagsgesellschaft mbH, Weinheim, Germany. Provides an overview of image capture with particular emphasis on types of capture equipment.
INTERNET RESOURCES rsb.info.nih.gov/nih-image NIH Image is free software that provides basic image analysis tools for the Macintosh. www.inforamp.net/∼poynton/Poynton-color.html Contains an excellent description of gamma correction in the Gamma FAQ. www-immb.ncifcrf.gov/EP/table2Ddatabases.html A list of links to many two-dimensional databases that are available via the Internet.
Contributed by Scott Medberry Amersham Pharmacia Biotech San Francisco, California Sean Gallagher Motorola Corporation Tempe, Arizona
Russ, 1995. See above. A general reference book on digital image capture and analysis.
Digital Electrophoresis Analysis
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Capillary Electrophoresis of Peptides and Proteins Using Isoelectric Buffers
UNIT 10.13
Capillary electrophoresis (CE) is a high-resolution separation technique that has become a valuable tool in the analysis of many molecules. UNIT 10.9 describes the basic concepts, instrumentation, and theory of this technology. Recent developments in the field have resulted in improvements to the procedure, and one such methodology, capillary zone electrophoresis (CZE) in amphoteric, acid buffers, is presented here. CZE has some marked advantages over standard CE technology. For example, by using isoelectric, amphoteric buffers that have very low conductivity, CZE can accommodate high-voltage gradients (up to 800 V/cm), allowing fast separation. In addition, the acidic buffers—typically with isoelectric point (pI) values in the pH range 2.5 to 3.5—allow the use of naked, fused-silica capillaries (i.e., capillaries in which the inner surface has not been derivatized in order to suppress silanol ionization). This is an important advantage, as it drastically lowers the costs of the separation column (the cost of a fused-silica capillary soars from a few dollars/meter for an untreated material to a few hundred dollars/meter for passivated surfaces). Finally, the acidic pH values of the background electrolyte permits maximization of the net positive charge of peptide/protein analytes via protonation of Glu and Asp residues. This further accelerates the migration process and thus further reduces analysis time. This unit presents methods for fast protein and peptide analysis in uncoated capillaries by CZE. Because the amphoteric buffers used are key to the success of the experiment, the authors have carefully studied the properties and uses of the buffers for the achievement of optimum results, and these parameters are outlined below (see Strategic Planning). Due to the acidic environment, it cannot be expected that proteins will maintain their native structure and integrity. Thus, these separations are routinely performed in 6 to 7 M urea, which accelerates the process of denaturation and helps keep the proteins in solution. As a consequence, if a protein is oligomeric, the present CZE technique will detect only the component subunits. Basic Protocol 1 details the procedure for analytical peptide separations, while Basic Protocol 2 outlines the separation of protein mixtures. An Alternate Protocol describes the detection of point mutations, a variation on Basic Protocol 2 that has applications in clinical studies. For additional information on the fundamentals of capillary electrophoresis, see UNIT 10.9. STRATEGIC PLANNING Properties of Amphoteric, Isoelectric Buffers To perform CZE peptide/protein separation in uncoated capillaries, the buffer must have a pH close to the neutralization point of the silica wall, determined to be pH 2.3. Given an average pK value of silanols of 6.3, essentially all silanols should be protonated at pH 2.3, and thus should be unable to adsorb proteinaceous samples by an ion-exchange mechanism. Four such acidic, amphoteric buffers have been tested for use in this system: cysteic acid (Cys-A; pI 1.85), imino diacetic acid (IDA; pI 2.23), aspartic acid (Asp; pI 2.78), and glutamic acid (Glu; pI 3.22). Their properties are listed in Figure 10.13.1. The following parameters should also be considered when choosing a buffer. Isoelectric point The pI is not an absolute value, but a limiting pH value. At vanishing concentrations of the amphotere in solution, the pH will tend to the pI of pure water (pH 7.0); at high enough Electrophoresis Contributed by Pier Giorgio Righetti, Alessandra Bossi, and Cecilia Gelfi Current Protocols in Protein Science (1999) 10.13.1-10.13.16 Copyright © 1999 by John Wiley & Sons, Inc.
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Chemical formulas
Electrolyte
Cysteic acid
Imino diacetic acid
pK1
pK2
pK3
pl
pl– pKprox
1.55
2.15
8.7
1.85
0.3
1.73
2.73
ND
2.23
0.5
1.88
3.66
9.9
2.77
0.89
2.19
4.25
9.5
3.22
1.03
H O
O
S OH O
OH NH2
O
O NH
OH
OH O
Aspartic acid
O
Glutamic acid
HO
OH OH NH2 O
O OH NH2
Figure 10.13.1 Properties of amphoteric, acidic, isoelectric buffers. ND, not determined.
concentrations, the pH will tend to the true pI of the amphotere. For Cys-A and IDA, the given pI value corresponds to a 100 mM solution; for Asp and Glu, it corresponds to a 50 mM solution. The (pI − pKprox) difference pKprox indicates a protolytic group active in the proximity of the pI value. In the case of Figure 10.13.1, it refers to either pK1 or pK2. The absolute value of the difference between pI and pKprox represents a hallmark of the amphotere. Very small values (≤1.0) indicate a very good species, possessing a high buffering capacity at pH = pI. Larger values (≥2) indicate a poor compound, unable to properly buffer at its pI value. Note that it is quite irrelevant which pK value will be displayed by the amino group in the molecule; any pK value above 6.0 will be compatible with the given buffer pIs. Solubility Cys-A and IDA display very high solubilities in water, whereas Asp and Glu are soluble only up to 50 mM (at 25°C). Compatibility with organic solvents Cys-A and IDA are compatible with a number of organic solvents, such as trifluoroethanol (TFE) and hexafluoropropanol (HFP), which impart unique electrophoretic properties to peptides. In contrast, Asp and Glu are compatible with only 10% TFE.
Capillary Electrophoresis Using Isoelectric Buffers
Conductivity The lower the pI, the higher the conductivity of the amphotere, due to dissociation of bulk water. Thus, Cys-A and IDA should preferably be used in the presence of organic solvents (such as TFE and HFP) or 7 M urea, which, in addition to modulating the mobility of peptides and proteins, also act as conductivity quenchers.
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2.1
0.90 2.0
0.85 0.80
1.9
0.75
1.8
Asp
0.072
D
3.2
3.0
0.064 2.9
0.060
2.8
0.056
2.7
0.052 0
5 10 15 20 25
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0.32
2.4
0.28
2.3 2.2
0.24
3.1
0.068
0.36
2.6
0.052
Glu
0.048
3.6 3.5
0.044 3.4 0.040
pH
0.95
IDA
pH
2.2
0.076 Conductivity (mS/cm)
Conductivity (mS/cm)
1.00
0.40
Conductivity (mS/cm)
C
B
2.3
Cys-A
pH
Conductivity (mS/cm)
1.05
pH
A
3.3
0.036 0.032 0
Time (days)
3.2 5 10 15 20 25 Time (days)
Figure 10.13.2 Conductivity and pH profiles of Cys-A (A), IDA (B), Asp (C), and Glu (D) upon aging for three weeks at room temperature. The four amphoteres were dissolved at 50 mM in plain water (unpublished data).
Addition of liquid polymers Although the acidic pH values adopted should ensure the absence of interaction between peptides or proteins and the silica wall, in reality this risk always exists. Thus, in all buffer formulations reported, small amounts of liquid polymers are added—typically 0.5% (w/v) hydroxyethyl cellulose (HEC; mol. wt. 27,000 Da). HEC (or other liquid polymer) is dissolved at concentrations well below the entanglement threshold and does not exert any sieving of the macromolecular analytes. However, it is very effective in forming a “dynamic coating” on the silica wall, thus preventing binding of polypeptides. Use of neutral and alkaline amphoteres Although histidine and lysine could be excellent amphoteric ions for such separations, the authors have chosen not to address them. Histidine has been excellent for oligonucleotide separations (Gelfi et al., 1996), and the use of lysine has been reported for analysis of native proteins (Hjertèn et al., 1995). However, coated capillaries should be used with these amphoteres, considerably adding to the burden and expenses of the technique. Long-Term Stability of Amphoteric, Isoelectric Buffers Shelf life An assessment of the shelf life of isoelectric buffers, dissolved as pure species in plain water as a solvent and stored at room temperature, determined that the final pH of such solutions approaches the pI value of each compound (P.G. Righetti, unpub. observ.). Because quite a number of solutions have strongly acidic pI values, they could potentially degrade at such local pH values. As indicators of potential degradation, pH and conductivity changes as a function of storage time were evaluated. Figure 10.13.2 gives these variations for four compounds, in order of increasing pI values: Cys-A, IDA, Asp, and
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2.8
0.48
2.7
0.44
2.6
0.40
0.24
2.5
Conductivity (mS/cm)
Cys-A
2.9
0.36
0.23
3.2 3.1 3.0
0.22
2.9
0.21
2.8 0.20
3.9
0.10
3.8 0.08
3.7 3.6
0.06
3.5 0.04
4.6
0.10
Conductivity (mS/cm)
4.0
Asp
Glu 4.4
0.08
4.2 0.06 4.0 0.04
3.8
0.02 0
5 10 15 20 25 Time (days)
pH
D
0.12
pH
Conductivity (mS/cm)
C
IDA
pH
B
0.52
pH
Conductivity (mS/cm)
A
3.6 0
5 10 15 20 25 Time (days)
Figure 10.13.3 Conductivity and pH profiles of Cys-A (A), IDA (B), Asp (C), and Glu (D) upon aging for three weeks at room temperature. The four amphoteres were dissolved at 50 mM in an aqueous 6 M urea solution (unpublished data).
Glu. Changes were monitored for a period of up to 3 weeks. There seems to be a general trend emerging in all four compounds: the pH and conductivity of all solutions remain stable for a period of ∼10 days, after which there appear to be fluctuations in both parameters. In some cases, the pH changes appear to be very minute (e.g., Fig. 10.13.2C, Asp) or to be nonmonotonic (e.g., Fig. 10.13.2A and B), so it would seem that they could be neglected. On the contrary, the conductivity increases steadily for all solutions analyzed, and this increase (in some cases quite marked) appears to be substantial after a period of 2 weeks. Figure 10.13.3 displays the monitoring of the same parameters for the set of four compounds dissolved in 6 M urea. Here too it is difficult to draw any conclusions from the shape of the pH curves: they either fluctuate or give minimal absolute changes with time, except for Asp and Glu, which show a steady increase in pH for a total of 0.2 and 0.6 pH units, respectively, over a period of 3 weeks. Conductivity changes, on the other hand, are quite marked and appear to increase constantly for all buffers studied. In addition, the onset of such conductivity increments occurs quite early compared to the same solutions in the absence of urea (typically after only a few days).
Capillary Electrophoresis Using Isoelectric Buffers
Making and using new stocks of buffers Another practical point of interest to users of CZE is how often one should replenish the anolyte and catholyte solutions, so as to avoid electrode polarization as well as marked changes of buffer pH and conductivity due to buffering ion and counter ion depletion. This point, in principle, should not be critical when using isoelectric buffers; their depletion at either electrode should be negligible, because they should not bear a net charge. Figure 10.13.4 explores potential pH and conductivity variations as a function of time of operation of the CZE instrument under a constant electric field of 20,000 V, using
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10.00
0.30
0.20 6.00
4.00 0.10
pH (cathodic – anodic)
Conductivity (anodic – cathodic)
8.00
2.00
0.00 0.00
10.00
20.00
30.00
40.00
0.00 50.00
Time (hours)
Figure 10.13.4 Conductivity and pH profiles of 40 mM Asp with 6M urea and 0.5% HEC, used for up to 50 hr as anolyte and catholyte. Electrophoretic conditions: 20 kV at room temperature without any change of electrolytes. The data are expressed as the difference in conductivity and pH measured in the two electrodic vessels (unpublished data).
Asp as the sole buffering species in all compartments (anode, cathode, capillary lumen). The data are expressed as the difference in pH and conductivity between the two electrode vessels. Note that in continuous operation the pH increases at the cathode, whereas the conductivity increases at the anode. This means that there must be a net transport of Asp from the cathodic to the anodic vessel. If that is the case, the pH should keep increasing at the cathode and decreasing at the anode. Decrements of pH in the anodic vessel should bring about increments of conductivity. This is to be expected. In fact, at progressively more acidic pH values, a fraction of the amphotere should be nonisoelectric in order to neutralize the excess positive charges brought about by bulk water ionization. For example, in a solution of Asp at pH 3.0, there must be 1 mM proton concentration, which must be neutralized by a 1 mM fraction of Asp being nonisoelectric and negatively charged. This excess, nonisoelectric, molar fraction will constantly migrate towards the anode, creating an imbalance in the concentration of Asp between the two electrodic vessels. This phenomenon can be neglected during the first 10 hr of operation, but after that it becomes quite appreciable. Practical approaches Considering the information given above, the following guidelines should apply to CZE in isoelectric, acidic buffers: 1. Buffers, especially if made in the presence of urea, should be prepared fresh every week, stored at 4°C, and then discarded. 2. Anolyte and catholyte solutions should be refreshed after each run when using the most acidic buffer (Cys-A). As the pI increases, the electrodic solutions can be progressively used for a few sequential runs before replenishing (e.g., in case of Asp, buffer can be used for a full day of operation before being discarded).
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3. As a corollary to the above, monitoring the electrodic vessels for pH and conductivity changes will give a sure indication of when to refresh these solutions. BASIC PROTOCOL 1
ANALYTICAL PEPTIDE SEPARATIONS CZE is particularly useful for generating peptide maps (fingerprinting) from complex mixtures, such as the products of proteolytic digestion (Bossi and Righetti, 1997; Capelli et al., 1997; Righetti and Nembri, 1997; McLaughlin et al., 1998). In this procedure, the peptide mixture to be analyzed is placed in a sample vial at the cathodic end of the capillary column. The anodic reservoir contains running buffer. The sample is loaded into the capillary either electrokinetically or by pressure or vacuum, and then the sample vial is replaced by running buffer and electrophoresis begins. The following protocol describes the use of a Bio-Rad Bio Focus 3000 instrument using low-pressure sample injection. Other instruments are capable of similar separations when operated according to the manufacturer’s instructions. Materials Peptide mixture (e.g., β-casein digestion; see recipe) 0.1 M NaOH Asp buffer: 50 mM aspartic acid (pH = pI = 2.77) containing 0.5% (w/v) hydroxyethyl cellulose (HEC; mol. wt. 27,000 Da, conductivity at 25°C 0.42 mS/cm) Capillary: 37-cm length (32.4 cm to the detection window); 75-µm i.d., 375-µm o.d. (Polymicro Technologies) Capillary zone electrophoresis (CZE) instrument (Bio Focus 3000, Bio-Rad, or equivalent) 1. Precondition a new capillary by flushing it with the following solutions, and then store the column in Asp buffer at 25°C. 1 M NaOH (1 hr) H2O (15 min) 1 M HCl (2 hr) H2O (15 min) 0.1 M NaOH at high pressure (30 min) H2O (5 min) Asp buffer (30 min). When changing to a different separation buffer, equilibrate the capillary for 4 hr with the new buffer by flushing at high pressure for 10 min and monitoring the baseline for 1 hr. The preconditioning step is essential if a new capillary is being used. If the capillary has already been properly equilibrated in separation buffer, it is best to avoid any conditioning with solutions of widely different pH, so as not to disturb the silica surface and alter its ionization properties. The capillary should be stored no more than 5 days
2. Load 10 to 20 nl peptide mixture using pressure injection for 0.2 to 1 sec at 5 psi according to manufacturer’s instructions. 3. Separate the peptide mixture using CZE instrument according to manufacturer’s instructions and using the following conditions:
Capillary Electrophoresis Using Isoelectric Buffers
Electrolyte: Asp buffer used as anolyte, catholyte, and for filling the capillary Detector wavelength: 214 nm Temperature: 25°C Voltage: 400 to 600 V/cm Run time: up to 20 min.
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4. Between runs, rinse the column with the same Asp buffer used for separations. When using the same capillary for a series of separations of the same type of sample, store the column in running buffer at room temperature. The capillary can be stored in this state for no more than 5 days. Figure 10.13.5 gives an example of the development of these peptide maps at 400 or 600 V/cm. Note that in this last case the entire map is developed in only 11 min. The same separation, when using a standard high conductivity buffer (80 mM phosphate, pH 2.0) takes ∼70 min, with severe loss of resolution due to diffusional peak spreading.
SEPARATION OF PROTEINS IN ACIDIC ISOELECTRIC BUFFERS Knowledge of the pIs of the components of the mixture would greatly help in selecting an appropriate run buffer. One capable of maximizing differences in the charge-to-mass ratio and ensuring that the proteins will all migrate towards the detector window. If the pI of a protein is unknown, separations can be achieved by using either a very alkaline or a very acidic buffer. However a coated column must be utilized with very alkaline buffers, so as to suppress any interaction of the analyte with the capillary wall. Therefore, very acidic buffers are preferred, as they should effectively quench adsorption of proteins onto the silanols. There is a catch to this strategy, however—in such an acidic environment proteins will most likely unfold and denature, due to disruption of the hydrogen bonds. Such denaturation could also induce protein aggregation and precipitation. The addition of 6 to 7 M urea in the background electrolyte ensures full denaturation while maintaining solubility of the analyte. Since the addition of 7 M urea will increase the apparent pH of the background electrolyte by ~1 pH unit, it is suggested that low pI buffers, such as IDA (pI 2.33) or Asp (pI 2.77), be used rather than Glu (pI 3.22). Please note that for oligomeric proteins, it is the subunits that will be separated (unless they are held together by disulfide bridges). The following protocol is based on a procedure routinely used for separation of wheat gliadins (Capelli et al., 1998).
BASIC PROTOCOL 2
Materials Protein sample (e.g., gliadin extraction; see recipe) 0.1 M NaOH 70% (v/v) ethanol Asp/urea buffers: 40 mM aspartic acid (pH = pI = 2.77), containing 0.5% (w/v) hydroxyethyl cellulose (HEC; mol. wt. 27,000 Da) and 7 M and 5 M urea (apparent pH 3.9; conductivity at 25°C: 0.7 mS/cm; both values refer to 7 M urea solutions) Capillary: 30-cm length (25.4 cm to the detection window), 50-µm i.d., 375-µm o.d. (Polymicro Technologies) Capillary zone electrophoresis (CZE) instrument (Bio Focus 3000, Bio-Rad, or equivalent) 1. Precondition a new capillary by flushing it with the following solutions. After equilibration, store in water at 25°C. 1 M NaOH (1 hr) H2O (15 min) 1 M HCl (2 hr) H2O (15 min) 0.1 M NaOH at high pressure (30 min) H2O (5 min) Asp buffer/7 M urea buffer (30 min) When changing to a different separation buffer, equilibrate the capillary for 4 hr with the new buffer by flushing at high pressure for 10 min and monitoring the baseline for 1 hr. Electrophoresis
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Absorbance (214 nm), mAU
A
8.00
(114-169) pl 6.1
(49-97) pl 6.93
(33-48) pl 3.95
4.00
0.00
5
B
10
15
20
20.00
Absorbance (214 nm), mAU
(114-169) pl 6.1 (49-97) pl 6.93
16.00
(33-48) pl 3.95
12.00
8.00
4.00
0.00
4
8
12
Time (min)
Figure 10.13.5 CZE of tryptic digests of β-casein in a 37-cm × 100-µm-i.d. capillary bathed in 50 mM isoelectric Asp (pH approximating the pI value of 2.77) with 0.5% HEC. (A) 400 V/cm (current: 33 µA); (B) 600 V/cm (current: 58 µA). The three major labeled peaks are (from left to right) the pI 6.1 fragment (β-CN 114-169); the pI 6.9 fragment (β-CN 49-97) and the pI 3.95 fragment (β-CN 33-48). CN, casein. Reproduced from Righetti and Nembri (1997), with permission from Elsevier. Capillary Electrophoresis Using Isoelectric Buffers
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The preconditioning step is essential if a new capillary is being used. If the capillary has already been properly equilibrated in separation buffer, it is best to avoid any conditioning with solutions of widely different pH, so as not to disturb the silica surface and alter its ionization properties.
2. Dilute the protein sample 1:5 (v/v) with 70% ethanol to give a final concentration of 2 mg/ml. 3. Fill the uncoated capillary with Asp/urea buffer containing 7 M urea. Fill the anode and cathode reservoirs with Asp/urea buffer containing 5 M urea. 4. Inject the sample hydrodynamically (e.g., 3 sec at 1 psi) according to manufacturer’s instructions. For a 2 mg/ml sample, 0.2 sec at 5 psi is sufficient.
5. Separate using the following conditions: Electrolyte: Asp/urea buffer containing 5 M urea used as anolyte and catholyte, and containing 7 M urea for filling the capillary Detector wavelength: 214 nm Temperature: 25°C Run voltage: 800 to 1000 V/cm Run time: 15 min. Figure 10.13.6 gives an example of a CZE run of four different cultivars of durum wheat, and Figure 10.13.7 gives an analogous separation for four cultivars of bread wheat. Note that these separations are typically over in 10 to 12 min. In contrast, the same separations performed in standard 100 mM phosphate buffer, pH 2.5, last for >30 min.
6. Between runs, rinse column with the same Asp/urea buffer used for separations. SEPARATION OF POINT MUTATIONS IN POLYPEPTIDE CHAINS Detection of point mutations is important in identifying several congenital defects of major clinical interest. For example, in the case of hemoglobinopathies, >600 defective variants of hemoglobin (Hb) have been reported. Among those, approximately one third involve a neutral-to-charged amino acid substitution, easily detectable by routine electrophoretic techniques, since they bring about a variation of the net charge of the expressed polypeptide. However, approximately two thirds represent the so-called electrophoretically silent mutations, in that they involve a neutral-to-neutral amino acid substitution. In this last case, not only is there no appreciable change in net charge, but the variation of polypeptide mass is also so minute as to escape detection by SDS-PAGE (UNIT 10.1). It has recently been demonstrated that addition of a neutral surfactant can in fact induce separation of the wild-type and mutant chains, probably by hydrophobic interaction of the more hydrophobic mutant with the detergent micelle, via a mechanism similar to micellar electrokinetic chromatography (Terabe et al., 1985). This is a protocol the authors routinely utilize for separation of point mutations in human globin chains (Righetti et al., 1998a; see also Regnier and Lin, 1998; Righetti et al., 1998b).
ALTERNATE PROTOCOL
Additional Materials (also see Basic Protocol 2) Protein sample (e.g., globin chain preparation; see recipe) IDA buffer: 50 mM imino diacetic acid (pH = pI = 2.23), containing 0.5% (w/v) hydroxyethyl cellulose (HEC; mol. wt. 27,000 Da) and 7 M and 5 M urea (apparent pH 3.2; conductivity at 25°C: 176 µS/cm) Uncoated capillary, 33-cm length (26 cm to the detection window), 50-µm i.d., 375-µm o.d. (Polymicro Technologies) Capillary zone electrophoresis (CZE) instrument (Bio Focus 3000, Bio-Rad, or equivalent)
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1. Condition the capillary (see Basic Protocol 2, step 1). 2. Fill the uncoated capillary with IDA buffer solution containing 7 M urea buffer. Fill the anode and cathode reservoirs with IDA buffer solution containing 5 M urea. If studying neutral point mutations, add 2% reduced Triton X-100 to the buffer.
3. Inject the sample hydrodynamically (e.g., 3 sec at 1 psi or 0.2 sec at 5 psi) according to manufacturer’s instructions. 4. Separate using the following conditions: Electrolyte: IDA buffer containing 5 M urea used as anolyte and catholyte, and containing 7 M urea for filling the capillary Detector wavelength: 214 nm Temperature: 15°C Run voltage: 600 V/cm Run time: 10 min.
Duilio
Absorbance (214 nm)
Creso
Capeiti
Appio
3
5
7
9
11
13
15
Time (min)
Capillary Electrophoresis Using Isoelectric Buffers
Figure 10.13.6 Representative CZE run of gliadins, obtained by sequential extraction of flour from four different cultivars of durum wheat. Run performed in 40 mM Asp with 7 M urea and 0.5% HEC (apparent pH 3.9). Conditions: 50-µm-i.d. × 30-cm-long capillary, run at 1000 V/cm, room temperature. From Capelli et al. (1998), with permission from Wiley-VCH.
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Figure 10.13.8 gives an example of a separation of a β-chain mutant in both control (in the absence of surfactant) and background electrolytes containing 2% (v/v) reduced Triton X-100. Notice, in the lower panel, the splitting of the two β-chains, which coelute in the absence of surfactant. Note that these separations are typically over in 6 to 7 min. The same separations performed in standard 100 mM phosphate buffer, pH 2.2, take ∼20 min.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
β-casein digestion Dissolve 10 mg/ml β-casein in 50 mM 3-(cyclohexylamino)-1-propanesulphonic acid (CAPS) buffer, pH 8.2. Add trypsin at a trypsin/β-casein ratio of 2% (w/w). Allow to hydrolyze for 4 hr at 50°C. Stop the reaction by adding 4 M acetic acid to pH 4.0. Store the tryptic peptide digest up to 6 months at −20°C.
Absorbance (214 nm)
Bussard
Australiano
Astron
Arabo
3
5
7
9
11
13
15
Time (min)
Figure 10.13.7 Representative CZE run of gliadins, obtained by sequential extraction of flour from four different cultivars of bread wheat. Run performed in 40 mM Asp with 7 M urea and 0.5% HEC (apparent pH 3.9). Conditions: 50-µm-i.d. × 30-cm-long capillary, run at 1000 V/cm, room temperature. From Capelli et al. (1998), with permission from Wiley-VCH.
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Gliadin extraction In the direct extraction, extract 30 mg bread or durum wheat flour while vortexing for 30 min at room temperature with 0.5 ml cold 70% (v/v) aqueous ethanol. Centrifuge for 15 min at 11,000 × g, 20°C. Use the supernatant, containing mostly gliadins (10 mg/ml), as is for CZE analysis. Store up to 6 months at −20°C. Globin chain preparation Wash red blood cells in physiological saline (0.9% NaCl), lyse in distilled water, and pellet the resulting “ghosts” by centrifuging 5 min at 5000 × g, 4°C. Add 20 µl concentrated hemoglobin (Hb) solution dropwise to 1.5 ml acetone containing 2% (v/v) HCl (at room temperature) in a microcentrifuge tube. Centrifuge again as before, discard the supernatant and resuspend the pellet in water. Repeat the centrifugation again, discarding the supernatant, in order to obtain a white globin powder. Dry the
A Absorbance (214 nm), mAU
α chains 6.00
β chains Hb Knossos beta 27 Ala control
Ser
4.00
2.00
0.00 5.00
6.00 Time (min)
7.00
Absorbance (214 nm), mAU
B α chains
8.00
+ 2% Triton X-100
6.00 β chains
4.00
5.20
Capillary Electrophoresis Using Isoelectric Buffers
5.60 Time (min)
6.00
Figure 10.13.8 CZE separation of hemoglobin Knossos (β27 Ala → Ser). (A) Control run; (B) buffer added with 2% reduced Triton X-100. CZE conditions: 50 mM IDA with 7 M urea and 0.5% HEC (apparent pH 3.2). Capillary: uncoated, 33-cm (length to detector 26 cm) × 50-µm-i.d. × 375-µm-o.d. Run at 600 V/cm (25 µA current) at 15°C. from Righetti et al. (1998a), with permission from Wiley-VCH.
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pellet in a Speedvac evaporator. Prior to analysis, dissolve the globin powder at a concentration of 2 mg/ml in background electrolyte containing 2% (v/v) 2-mercaptoethanol and only 5 (instead of 50) mM IDA. Store up to 6 months at −20°C. COMMENTARY Background Information CZE of proteins and peptides has usually been performed either at very acidic (pH 2 to 2.5) or rather alkaline (pH 9 to 10) pH values. Under alkaline conditions, a coated capillary has to be used, in order to quench the electroendoosmotic flow (EOF) and to minimize residual adsorption to fully ionized silanols. Under acidic conditions, a naked capillary can be used, but the very high conductivity of such acidic buffers (notably phosphate), allows only lowvoltage gradients (e.g., maximum 100 V/cm) with concomitant very long analysis times. The first report on the use of amphoteric buffers, used as the sole background electrolyte at pH = pI, came from Hjertèn’s group in 1995 (Hjertèn et al., 1995). This group could separate albumin and transferrin in only 40 sec using 15-cm-long capillaries at 2000 V/cm in lowconductivity polymeric buffers (pH 8.6). In 5 mM Lys (pH = pI = 9.7), a mixture of different protein markers (soybean trypsin inhibitor, ovalbumin, conalbumin, and myoglobin) could also be separated in 240 to 40 sec at voltage gradients ranging from 330 to 2000 V/cm. However, while Hjertèn’s procedure solved the problem of fast analysis times, it had to adopt capillaries with an inner coating of polymeric material (usually polyacrylamide) covalently anchored to the silanols. The first article published dealing with the use of amphoteric, isoelectric buffers was that of Mandecki and Hayden (1988), on the use of His as the sole buffering ion in slab gel electrophoresis of DNA. Although this methodology appeared in 1988, it went largely unnoticed until it was rediscovered by Hjertèn’s group. The authors have also adopted isoelectric His in CZE for the fast separation of antisense oligonucleotides in liquid-sieving polymers (Gelfi et al., 1996). The main difference between previous data and the present technique resides in the use of acidic, isoelectric buffers, with all the benefits inherent to this methodology, such as fast analysis times, high resolution, and the ability to use untreated fused-silica capillaries.
Critical Parameters For separation of peptides (see Basic Protocol 1), it is generally not necessary to add urea,
as these fragments are well soluble in plain water solvent because of their relatively small size and good hydrophilicity. One exception would be peptides obtained from hydrophobic (e.g., membrane) proteins—these samples run the severe risk of aggregating and/or precipitating. In this case, addition of urea, possibly also combined with surfactants, is recommended. If detergents are used, neutral or zwitterionic species (such as reduced Triton X-100 or CHAPS) work best. Do not use Nonidet P-40 or other detergents that contain aromatic rings, as their absorbance at 200 nm will lower the sensitivity of detection. In mixed hydro-organic solvents of the water/urea type, it should be remembered that, due to the heat developed in a capillary unit and to the strong air streams in equipment that uses forced-ventilation cooling, there is a strong risk of solvent evaporation in the two electrode vessels. Urea will concentrate and crystallize, blocking the inlet of the capillary. Thus, as a safety precaution, only 5 M urea should be added to the electrode vessels when using buffers containing urea. For the running buffer in the capillary, the urea concentration can be increased to 7 M urea, but not higher than that. With high levels of urea, be sure that the extremities of the capillary are always kept inserted in a vessel containing liquid. When left in the air, the solvent will quickly evaporate from the capillary tips and urea will crystallize. For protein separations (see Basic Protocol 2 and see Alternate Protocol), there is often the risk that small amounts of protein will be adsorbed onto the silica wall. After repeated injections, there might be a significant alteration of the silica surface, leading to irreproducible results and even to spurious peaks. A simple solution is to dispose of the capillary and start with a new one. However, if the same capillary has to be reused, there are several procedures for regenerating the silica wall. In one (drastic) treatment, the capillary can be rinsed with 0.1 N (or even 1 N) NaOH. Alkaline treatments are very effective in removing and solubilizing proteinaceous material and most adsorbed impurities. Unfortunately, such a drastic procedure will ionize most of the surface silanols and deeply alter the surface properties of the silica
Electrophoresis
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Capillary Electrophoresis Using Isoelectric Buffers
wall. In addition, because alkaline treatments are abrasive, part of the silica surface can be dissolved. As a result, it might be very difficult to recondition the capillary wall to the original properties and obtain reproducible results in electroendoosmotic flow and transit times. A better way to remove adsorbed material is by rinsing the capillary with running buffer containing micellar SDS (e.g., 30 mM SDS). However, if the running buffer has too low a pI, the SDS micelles might not be effective—in this case, dissolve the SDS in Good’s buffers (e.g., MES or MOPS, titrated at pH 6.8). The running buffer/SDS solution is placed at the cathode and the SDS micelle is transported electrophoretically into the capillary. The migrating SDS micelles act by sweeping any silica-bound material, which will be adsorbed into the micelle and migrate with the SDS front. If this operation is conducted under UV monitoring, a peak of desorbed material will be seen at the passage of the SDS micelles. This electrophoretic desorption can be continued for 30 min under constant electric field. This process is the electrophoretic analog of frontal chromatography. After this step, the SDS micelles can be removed by rinsing with water and then the capillary can be reconditioned by purging with the chosen isoelectric running buffers (10 column volumes; see Basic Protocol 1, step 1), in the presence of the proper additives, if any. It appears that the two best buffers, in general, are IDA and Asp. Cys-A is strongly acidic and does not offer particular advantages over such a strongly conducting buffer as phosphate, titrated at pH 2.0 to 2.5. Cys-A is best used in presence of conductivity quenchers, such as urea and/or organic solvents (TFE and HFP) that are compatible with this buffer and with bulk water at any concentration (although, typically, no advantage is obtained at concentrations greater than 40%, v/v). Cys-A seems to be particularly attractive when analyzing small (di-, tri-, and tetra-) peptides that bear the terminal carboxy and amino groups as their only charged groups. In fact, such peptides are isoelectric over a broad pH range (typically at least 4 to 8), so that either strongly acidic or strongly basic solutions are needed in order to provide sufficient charge for a fast separation. Glu is also an interesting buffer, because its higher pI (3.22, especially when in presence of urea, apparent pH 4.3) allows modulation of the net charge of large peptides, due to partial deprotonation of Asp and Glu residues. Thus, in general, the titration curves of polypeptides tend to fan out at and above pH 4, allowing
mobility modulation and increased selectivity. The risk with Glu, however, is that at this higher pH (especially in the presence of 7 M urea) polypeptides might bind to ionized silanols, resulting in loss of resolution. Thus, when adopting Glu as a background electrolyte, one should perform preliminary runs in order to identify potential interaction of the silica wall with the components of the protein mix under analysis.
Troubleshooting Potential mechanical and/or electrical problems that might be encountered during instrument use are usually addressed in the manufacturer’s manual. Therefore, this discussion will focus on problems encountered with the sample and background electrolyte. The most common problems encountered in CZE are discussed below. Loss of current during a run A typical problem is the sudden or gradual loss of electric contact during a run. Sudden loss of electric contact could be due to a breaking of the capillary or the trapping of a large air bubble spanning the entire lumen of the capillary. Gradual loss of current could be caused by small air bubbles that develop during the run (e.g., due to excess power forcing the solution to outgas), which adhere to the silica wall but are too minute to completely obstruct the lumen. During the run, more air bubbles could develop and further reduce the lumen available to the passage of electric current. On a long run, such tiny air bubbles could coalesce and suddenly break the current (remember that air bubbles can be negatively charged due to adsorption of ions in their diffuse double layer, and thus can migrate in the electric field). Bubbles can be avoided by thoroughly degassing all solutions used in the instrument (including the reservoirs). The authors often adopt a double degassing procedure: under vacuum and also by sonication. It is also a good procedure to check the total applied electric power and see that it does not exceed the power recommended by the manufacturer. One should also make sure that the sample vial contains enough sample to cover the extremity of the capillary at the injection site, so as to prevent injection of air into the column. Remember that, although the sample volume injected in the capillary is minute (typically 5 to 20 nl), for proper immersion of the capillary end in the analyte solution the vial should contain 10 µl. In addition, the sample vials can
10.13.14 Supplement 16
Current Protocols in Protein Science
dry out during a long run, especially when performing sequential runs and placing a large number of samples in the carousel. One would then notice that the later samples give exaggeratedly huge peaks due to excessive liquid evaporation. In order to avoid that, when programming a sequence of runs (e.g., for overnight analyses), the authors have adopted a neat safety procedure, consisting of overlaying the analyte solution with a layer of light paraffin oil. This oil will float on top of the liquid layer of analyte, effectively sealing it from the air and completely preventing evaporation. Arcing CZE instruments can arc during a run. This is often caused by salt deposits or moisture on the high-voltage electrode stations. In instruments using a free capillary, not encapsulated in a cartridge, moisture can be deposited all along the capillary’s outer surface, causing creeping current on its surface. This can be particularly true in a hot and humid summer environment in the absence of proper air conditioning and dehumidification. A proper cure would be to purge the capillary chamber with dry inert gas and to place Drierite bags in the capillary holding compartment. Column plugging The column can plug because of salt deposition as the liquid inside dries out. This is particularly true when using urea-laden buffers. As a safety precaution, if no further use of the capillary is expected at the end of a working day, the column should be emptied and stored overnight in plain water or in urea-free buffer. In general, washing the column with water dissolves the plug if a standard buffer is used; however, in the case of urea-containing buffers, it might be necessary to discard the capillary if extensive deposition of urea has occurred in a substantial portion of its length. Proteins too can precipitate during a run and plug the column. Using lower protein concentrations usually prevents this type of plugging. Sudden loss of sample resolution When using very acidic isoelectric buffers (e.g., Cys-A), poor resolution can be caused by ion depletion at one electrode due to electrophoretic migration of the nonisoelectric molar fraction. When this occurs, most of the voltage gradient falls on the electrode vessel depleted of ions, with consequently poor or no migration of the analyte components. Check the electrode pH and conductivity, and refresh the buffer.
Sample tailing Sample tailing could be due to adsorption of sample ions to ionized silanols. It might occur in two particular cases: when using proteins (peptides are generally too small to elicit substantial binding to the silica wall) and when using progressively higher pI amphoteres, such as Glu. Adsorption of polypeptides to the capillary can be demonstrated by frontal analysis, i.e., by making a blank run in which SDS micelles are added to the cathodic vessel. On their migration to the anode, the SDS micelles will sweep away bound proteins and a sharp sample peak will be detected. The remedy is to adopt lower pI amphoteric buffers and/or increase the amount of sieving liquid polymer (HEC, which should be used below 1%, as it will give too high a viscosity). Unreproducible transit times A possible cause of unreproducible transit times could be too short an equilibration period of the capillary in a given buffer. Let the capillary equilibrate overnight, instead of only 4 hr, with frequent periods of pumping the buffer through the column. This will also favor deposition of the HEC polymer onto the inner capillary surface, a process called dynamic coating. Another cause could be a drastic washing step with alkaline solutions (e.g., 0.1 N NaOH). This will ionize a large number of silanols and deeply alter the charge state of the surface. It might then take extensive washings in the original acidic buffer solution in order to reestablish the proper starting surface conditions. Often, the biggest offender is accumulation of adsorbed material from the sample plug. This material might not necessarily be the protein or peptide of interest, but some impurities in the sample matrix (e.g., lipids). One remedy to it is to try to pre-purify the sample solution, e.g., by a lipid-extraction step. The other remedy is to wash the capillary with a front of SDS micelles, which will quite efficiently desorb bound material by forming mixed micelles.
Anticipated Results Results of separations obtainable with the experimental conditions outlined here are discussed with each of the protocols (also see Figs. 10.13.5-10.13.8).
Time Considerations As a virtue of the high voltage gradients that can be applied in isoelectric buffers, the total analysis time is much reduced compared to analogous techniques in conventional, non-
Electrophoresis
10.13.15 Current Protocols in Protein Science
Supplement 16
isoelectric buffers. For example, in Asp buffer at the standard voltage of 600 V/cm in a 37-cmlong capillary, analysis is usually over in 10 to 12 min. The total analysis time, including sample loading, running, and capillary regeneration prior to the following run, is typically on the order of 15 min.
Literature Cited Bossi, A. and Righetti, P.G. 1997. Generation of peptide maps by capillary zone electrophoresis in isoelectric iminodiacetic acid. Electrophoresis 18:2012-2018. Capelli, L., Stoyanov, A.V., Wajcman, H., and Righetti, P.G. 1997. Generation of tryptic maps of α- and β-globin chains by capillary electrophoresis in isoelectric buffers. J. Chromatogr., A 791:313-322.
Righetti, P.G. and Nembri, F. 1997. Capillary electrophoresis of peptides in isoelectric buffers. J. Chromatogr., A 772:203-211. Righetti, P.G., Saccomani, A., Stoyanov, A.V., and Gelfi, C. 1998a. Human globin chain separation by capillary electrophoresis in acidic, isoelectric buffers. Electrophoresis 19:1733-1737. Righetti, P.G., Olivieri, E., and Viotti, A. 1998b. Identification of maize lines via capillary electrophoresis of zeins in isoelectric, acidic buffers. Electrophoresis 19:1738-1741. Terabe, S., Otsuka, K., and Ardo, T. 1985. Micellar electrokinetic chromatography. Anal. Chem. 57:834-841.
Key References Righetti, P.G., Gelfi, C., Perego, M., Stoyanov, A.V. and Bossi, A. 1997. Capillary zone electrophoresis of oligonucleotides and peptides in isoelectric buffers: Theory and methodology. Electrophoresis 18:2145-2153.
Capelli, L., Forlani, F., Perini, F., Guerrieri, N., Cerletti, P., and Righetti, P.G. 1998. Wheat cultivar discrimination by capillary electrophoresis of gliadins in isoelectric buffers. Electrophoresis 19:311-318.
Righetti, P.G., Bossi, A., and Gelfi, C. 1997. Capillary isoelectric focusing and isoelectric buffers: An evolving scenario. J. Cap. Elec. 4:47-59.
Gelfi, C., Perego, M., and Righetti, P.G. 1996. Capillary electrophoresis of oligonucleotides in sieving liquid polymers in isoelectric buffers. Electrophoresis 17:1470-1475.
Stoyanov, A.V. and Righetti, P.G. 1997. Fundamental properties of isoelectric buffers for capillary zone electrophoresis. J. Chromatogr., A 790: 169-176.
Hjertèn, S., Valtcheva, L., Elenbring, K., and Liao, J.L. 1995. Fast, high-resolution (capillary) electrophoresis in buffers designed for high field strengths. Electrophoresis 16:584-594.
Three good reviews of both theory and practice for separation of peptides and proteins by CZE in isoelectric buffers.
Mandecki, W. and Hayden, M. 1988. Electrophoresis of DNA fragments in isoelectric histidine buffer. DNA 7:57-62. McLaughlin, G.M., Anderson, K.W., and Hauffe, D.K. 1998. Peptide analysis by capillary electrophoresis: Method development and optimization, sensitivity enhancement strategies and applications. In High-Performance Capillary Electrophoresis (M.G. Khaledi, ed.) pp. 637-681. John Wiley & Sons, New York. Regnier, F. and Lin, S. 1998. Capillary electrophoresis of proteins. In High-Performance Capillary Electrophoresis (M.G. Khaledi, ed.) pp. 683728. John Wiley & Sons, New York.
Contributed by Pier Giorgio Righetti and Alessandra Bossi University of Verona Verona, Italy Cecilia Gelfi Instituto Tecnologie Biomediche Avanzate and Consiglio Nazionale delle Richerce Milan, Italy
The authors wish to acknowledge the support of grants from the Agenzia Spaziale Italiana (No. ARS-98-179) and from MURST (Coordinated Project 40%, Folding/Unfolding of Proteins).
Capillary Electrophoresis Using Isoelectric Buffers
10.13.16 Supplement 16
Current Protocols in Protein Science
CHAPTER 11 Chemical Analysis INTRODUCTION
C
hapter 11 is devoted to methodologies related to determining the primary structure of proteins. Although the techniques detailed can be applied to proteins available in large amounts, particular emphasis is placed on analyzing proteins available in limited (picomole) quantities. Amino acid analysis, described in UNIT 11.9, is one of the best methods to quantify peptides and proteins. It also serves as a valuable analytical method for evaluating synthetic peptide or recombinant protein purity, identifying proteins based on composition, and detecting odd amino acids. Primary sequence analyses (UNIT 11.10) are used to identify proteins and to obtain information that can be used to design oligonucleotide probes that can be used to clone the gene encoding the protein of interest. Also, with the proviso that contaminating proteins may not be sequenceable, and hence undetectable due to blocked N termini, some assessment of purity can be made from N-terminal sequence analyses. The usual approach for obtaining primary sequence analysis is to attempt to sequence the N terminus of the intact protein; however, for one reason or another, proteins may lack a free N-terminal amino group (i.e., they are blocked) and are consequently refractory to Edman degradation. One common N-terminal blocking group, pyrrolidone carboxylic acid, formed by cyclization of glutamine or glutamic acid, can often be removed by treatment with the enzyme pyroglutamate amino peptidase, as described in UNIT 11.7. This unit also describes methods for obtaining N-terminal sequence of Nα-acyl-proteins, but in this case, the protein must first be fragmented. If purified, blocked protein cannot be “unblocked” by these or other means, it is usually necessary to fragment the protein by enzymatic and/or chemical methods (UNITS 11.1 & 11.4, respectively) and to isolate individual protein fragments by reversed-phase high performance liquid chromatography (RPHPLC; UNIT 11.6) or other means in order to obtain primary sequence data. Even if the N terminus of a protein is not blocked, such an approach frequently constitutes the most economical use of protein, in that much more sequence can often be obtained from a limited amount of protein using this approach than if just the N terminus were sequenced. Carboxy-terminal sequence analysis detailed in UNIT 11.8 is used for direct confirmation of the C-terminal sequence of native and expressed proteins, for identifying post-translational proteolytic cleavages, and for obtaining partial sequence information in order to design oligonucleotide probes for gene cloning. UNIT 11.8 contains an automated chemical method and a manual method based on carboxypeptidase digestion for obtaining such information. A major advantage of the automated method is that no proteolytic cleavage is required and as little as 10 pmol of a protein can be directly analyzed. The procedure allows combination of N- and C-terminal sequence analyses of the same sample aliquot immobilized on a PVDF membrane. The manual methods employ carboxypeptidase digestion of the sample. Using Basic Protocol 2, the sequence is deduced by determining the mass differentials between the undigested and digested sample at different time points using MALDI-TOF instrumentation (see Chapter 16). The advantages of this manual method are the relatively low sample requirements, the rapidity of mass analysis and the ease of data interpretation. Basic Protocol 3 describes the classical manual method in which the released amino acids are determined by amino acid analysis (see UNIT 3.2 and Chemical Analysis Contributed by John E. Coligan Current Protocols in Protein Science (2003) 11.0.1-11.0.2 Copyright © 2003 by John Wiley & Sons, Inc.
11.0.1 Supplement 31
UNIT 11.9). This method requires more protein, but has the potential for more accuracy, especially in the case of larger proteins.
Because the final step in protein purification is usually SDS-PAGE, electroblotting to solid supports such as polyvinylidene difluoride (PVDF) membrane is the simplest and most common procedure for recovering proteins in high yield and free of contaminants such as SDS. However, because elution of proteins from such solid supports is often very inefficient, procedures for enzymatic and chemical digestion of proteins on solid supports have been developed (UNITS 11.2 & 11.5, respectively). In some cases protein losses are incurred during transfer from the SDS-polyacrylamide gel to the solid support, or digestion on the solid support may be particularly inefficient. For such cases, special enzymatic digestion protocols have been developed to digest proteins directly in the resolving gels (UNIT 11.3). These protocols are tailored to deal with the high concentrations of SDS present in such gels, which normally inhibits enzymatic hydrolysis. A support protocol provides a method for determining sample concentration by amino acid analysis. As described in UNIT 11.6, RP-HPLC is a fundamental tool for the isolation and analysis of peptides generated by various cleavage procedures. Peptides are separated on a hydrophobic stationary phase and eluted with a gradient of increasing organic solvent concentration. Depending on the amount of peptide, different-sized columns are recommended—e.g., narrow-bore, microbore, and capillary; protocols for each column type are provided. The support protocol in UNIT 11.6 details the construction of a capillary HPLC system that allows flow rates of 3 to 5 µl/min for analysis of peptides at the 500 fmol level. UNIT 11.6 also describes the use of MALDI-TOF mass spectrometry (Basic Protocol 3, also see Chapter 16) or capillary electrophoresis (Basic Protocol 4, also see UNIT 10.9) analyses to assess the purity of the peptides present in the HPLC fractions. These protocols can also be used to directly analyze protein digests. In the end, data obtained through these procedures can be used to confirm the identity of a known protein, correlate a protein with a known gene sequence, or form the basis for the synthesis of oligodeoxynucleotides (primers or probes) to clone a gene corresponding to the protein sequence. John E. Coligan
Introduction
11.0.2 Supplement 31
Current Protocols in Protein Science
Enzymatic Digestion of Proteins in Solution
UNIT 11.1
Analysis of protein covalent structure is less complex and more accurate when performed on peptides derived from the larger protein. In contrast to acid-promoted total hydrolysis (e.g., in 6 N HCl), peptides are typically generated by selective proteolysis—i.e., by specifically cleaving peptide bonds with endoproteases that have varying degrees of specificity. The basic protocol can be used to generate peptide fragments from intact, undenatured proteins. Fragments can be analyzed directly by mass spectrometry (MS) or, more often, are first separated by reversed-phase HPLC (RP-HPLC) and then analyzed by MS or automated sequencing. Expertise with these analytical and chromatographic techniques is required, or else the procedures must be performed in association with a skilled operator. Most proteins are resistant to enzymatic proteolysis under nondenaturing conditions (see Table 11.1.2) or are not soluble in aqueous solution. Digestion procedures performed in the presence of chaotropic agents and SDS are described in Alternate Protocols 1 and 2. Support Protocols 1 and 2 provide instructions for preparing enzyme stocks and reducing and alkylating peptides prior to sequencing or HPLC analysis. DIGESTION OF PROTEINS UNDER NONDENATURING CONDITIONS Protein is solubilized in an appropriate digestion buffer and subjected to enzymatic proteolysis. Fragmentation is monitored by SDS-PAGE or RP-HPLC, followed by preparative RP-HPLC of the completed digest. The procedure is recommended for checking protease sensitivity of unknown proteins or for preparing proteolytic fragments that can be sequenced when working with proteins already shown to be sensitive to a particular protease. The minimal amount of protein required is 100 pmol.
BASIC PROTOCOL
Materials 100 pmol to 5 nmol protein sample, as pellet or solution 1 × and 10× digestion buffer (see Table 11.1.1) 20% 3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate (CHAPS), 20% octyl glucoside, or 20% Nonidet P-40 (NP-40; Calbiochem) 100% acetonitrile 1 µg/µl enzyme stock (see Support Protocol 1; store at −20°C) Trifluoroacetic acid (TFA; Pierce) Solvent A: 0.1% (v/v) TFA in water Solvent B: 0.09% (v/v) TFA in 70% (v/v) acetonitrile (Burdick & Jackson) Sonicator bath (Branson) Phast Gel system (Pharmacia), optional HPLC system, with C18 or C4 reversed-phase column, 4.6-mm or 2.1-mm (e.g., Vydac);UV detector; chart recorder Additional reagents and materials for SDS-PAGE (UNIT 10.1), gel staining (UNIT 10.5), and HPLC analysis of peptides (UNIT 11.6) 1. Dissolve protein pellet in minimal volume of appropriate 1× digestion buffer. If protein sample is already in solution, add 0.1 vol of 10× digestion buffer. Final protein concentration must be >1 µg/20 µl. If sample is too dilute and volume is over 0.5 ml, use Centricon microconcentrator (see manufacturer’s instructions) to reduce the volume. Chemical Analysis Contributed by Lise R. Riviere and Paul Tempst Current Protocols in Protein Science (1995) 11.1.1-11.1.19 Copyright © 2000 by John Wiley & Sons, Inc.
11.1.1 CPPS
11.1.2
Current Protocols in Protein Science
d
—
Reducing agent 2-ME —
30% 20%
0.1%c 1% 1%
0.5%
40% 40%
1% 2% 2%
0.1%
40% 40%
<0.1% 2% 2%
0.1 M AB 0.2 M TC 18 hr
0.5%
20%c 20%c
0.1%c 2%c 2%c
18 hr
0.05 M AB
1
Mc
1
5 hr
Mc
2 Mc 0.05 M AB
— —
5 hr
S
2
ND
40% 40%c
1% 2%c 2%c
2 hr
0.1 M AB
Mc
1 hr
8 Mc 0.1 M TC
— 1M
1 hr
0.1 M AB
Enzyme
0.05 M AB
EC
5 hr
2M 0.1 M AB
— —
5 hr
0.1 M AB
DN
ND
40% 40%c
0.1%c 2% 2%
—
8 Mc 0.1 M AB/ +5 mM Ca 0.1 M TC/ +5 mM Ca 2 hr
— —
0.1 M AB/ 1 mM CaCl2 2 hr
H
ND
20%c 20%c
ND
ND
ND
2 hr
12 mM HCl
2M
ND
+ —
1 hr
0.1% TFA
P
2
ND
40% 20%
0.1% 1% 1%
18 hr
0.2 M TC
Mc
18 hr
2M 0.2 M TC
— —
18 hr
0.2 M TC
E
ND
40% 40%
— 1% 2%c
0.1 M AB/ 1 mM EDTA 18 hr
2M
5 hr
8 Mc 0.1 M AB/ 1 mM EDTA
— —
0.1 M AB/ 1 mM EDTA 5 hr
PA
bAbbreviations: Enzymes: C, chymotrypsin; DN, endoproteinase Asp-N; E, elastase; EC, endoproteinase Glu-C; H, thermolysin; KC, endoproteinase Lys-C; P, pepsin; PA, papain; S, subtilisin; T, trypsin. Other: —, does not work; +, works at pH 2.0; AB, ammonium bicarbonate; Gu⋅HCl, guanidine hydrochloride; IPA, isopropanol; 2-ME, 2-mercaptoethanol; MeCN, acetonitrile; OG, octyl glucoside; TC, Tris⋅Cl, pH 8.5; TFA, trifluoroacetic acid. cRestricted digest. Note that for the 8M urea conditions, the final concentration is actually 7.3 M after addition of the buffer. dConcentrations higher than 2% CHAPS, octyl glucoside, and NP-40, and 40% acetonitrile and isopropanol were not tested.
aBuffers listed here are 1× stocks; it may be necessary to prepare 10× stocks for certain parts of the protocols. Listed here are the highest concentrations of additives that still permit adequate proteolysis.
40% 40%
<0.1% 2% 2%
Organic solvents MeCN IPA
Detergents SDS CHAPS OG and NP-40
5 hr
24 hr
Time
d
2M 0.2 M TC
2M 0.2 M AB
—
5 hr
Gu⋅HCl Buffer
5 hr
8M 0.2 M TC
— 0.1 M
2 hr
0.1 M AB
KC
15 hr
2M 0.1 M AB
— —
2 hr
0.1 M AB
C
Time
4M 0.1 M AB/ +5 mM CaCl2
— —
Chaotropes Urea Buffer
2 hr
Low/high pH Acid NH4OH
0.1 M AB
T
Conditions for Endoprotease Activitya,b
Time
Nondenaturing Buffer
Condition
Table 11.1.1
2. Incubate sample 5 min at 37°C, then sonicate 1 min. Repeat until pellet is completely dissolved. If pellet does not dissolve, add CHAPS (2% final), octyl glucoside (2% final), NP-40 (2% final), or acetonitrile (20% to 40% final; Table 11.1.1) to digestion solution. These solubilization agents do not interfere with digestion (see Table 11.1.1). NOTE: Digestion solution should not contain protease inhibitors. All proteases are inhibited by leupeptin. In addition, trypsin, chymotrypsin, endoproteinase Lys-C, endoproteinase Glu-C, subtilisin, and elastase are inhibited by phenylmethylsulfonyl fluoride (PMSF), diisopropyl fluorophosphate (DFP), and aprotinin; thermolysin and endoproteinase Asp-N are inhibited by EDTA.
3. Transfer 1 µl sample to pH indicator strip. If pH is not correct, add sufficient 10× digestion buffer to increase buffer strength by 100 mM (see Table 11.1.1) and recheck the pH. For pepsin, the pH must be 2.0. Subtilisin and endoproteinase Lys-C work well in a pH range of 7 to 10.5. For all other proteases, pH should be 8.0 to 8.5.
4. Add 1 µg/µl enzyme stock solution to a final enzyme/substrate ratio of 1:50 to 1:10 (w/w). Final enzyme concentration should be >1 µg/100 µl.
5. Incubate at 37°C for the amount of time recommended for the selected protease (see Table 11.1.1). 6. Determine extent of digestion by analyzing 0.5 to 1 µg digest by SDS-PAGE (e.g., using a Phast Gel). Stain with Coomassie brilliant blue (UNIT 10.5). Alternatively, analyze 25 to 500 pmol by RP-HPLC (use a 2.1-mm column for 25 to 200 pmol protein or a 4.6-mm column for 200 to 500 pmol protein). Keep remaining sample on ice. The procedure should not consume >10% of the sample. Phast Gels are useful when protein amounts are limited.
7. If sample is digested, proceed to preparative RP-HPLC (step 8). If sample is not digested, check pH and adjust if necessary. Add a second aliquot of enzyme and incubate at 37°C for an additional 30 min to full incubation time. Check extent of digestion as in step 6. If sample is still not digested after incubation with a second aliquot of enzyme, try a different enzyme. If the second enzyme is ineffective, denature the substrate (see Alternate Protocol 1 and Alternate Protocol 2). If HPLC analysis cannot be performed immediately, TFA should be added to a final concentration of 2% and the sample should be stored at −70°C. Digestion samples may be kept indefinitely under these conditions.
8. Using a clean Hamilton syringe, inject remaining digested sample onto column. Program HPLC to run isocratically at 5% solvent B until baseline returns to its original position. Set flow rate to 100 µl/min for a 2.1-mm column or 1 ml/min for a 4.6-mm column. Run gradient as follows: 5% to 50% solvent B in 45 min; 50% to 100% solvent B in 25 min. A 4.6-mm column can easily accommodate 1 to 2 ml sample volume. If the sample contains organic solvent, it should be diluted five-fold with solvent A. See UNIT 11.6 for further details on HPLC separation of peptides.
Chemical Analysis
11.1.3 Current Protocols in Protein Science
ALTERNATE PROTOCOL 1
DIGESTION OF PROTEINS IN THE PRESENCE OF UREA OR GUANIDINE⋅HCl Because many proteins are resistant to protease digestion in their native state, the majority of digests are carried out in the presence of urea and guanidine⋅HCl. The highest concentrations of these chaotropes compatible with the activity of the most commonly used proteases are listed in Table 11.1.1. Including chaotropes in the digest mixture avoids having to denature (or reduce and alkylate) substrate proteins prior to digestion. This, in turn, eliminates the need to remove the denaturants by chromatography or dialysis before adding the protease, a step which can cause heavy losses of substrate proteins present in subnanomole quantities. Proteins are solubilized and denatured in high concentrations of urea or guanidine⋅HCl, and the denaturant is then diluted to a concentration high enough to prevent refolding of the substrate but low enough to allow protease digestion. Fragmentation is monitored by RP-HPLC. The procedure has a high success rate and is most often used as the method of first choice. The protocol requires ≥200 pmol of protein. Additional Materials (also see Basic Protocol) 8 M urea (sequenal grade, Pierce; store up to several weeks at room temperature over Bio-Rad AG 501-X8 mixed-bed resin) 6 M guanidine⋅HCl (sequenal grade, Pierce), prepared fresh immediately before use Milli-Q water or equivalent 50°C water bath 1a. If sample is solid: Dissolve protein pellet in a minimal vol of 8 M urea or 6 M guanidine⋅HCl. Heat sample 5 min at 37°C, then sonicate 1 min. Repeat heating and sonication until pellet is dissolved. 1b. If sample is already in solution and precipitation is not desirable: Add solid guanidine⋅HCl to give a final concentration of 6 M: i.e., add 104 mg guanidine⋅HCl per 100 µl sample volume (final volume will then be 182 µl). 2. Heat sample 30 min at 37°C (for 8 M urea) or 30 min at 50°C (for 6 M guanidine⋅HCl). 3. Allow sample to cool to room temperature. Add water and appropriate 10× digestion buffer to achieve desired final concentration of chaotropic agent as follows: For 8 M urea: undiluted, add 0.1 vol 10× buffer For 4 M urea: 0.8 vol water and 0.2 vol 10× buffer For 2 M urea: 2.6 vol water and 0.4 vol 10× buffer For 2 M guanidine⋅HCl: 1.7 vol water and 0.3 vol 10× buffer. Final substrate concentration should be >1 µg/10 µl; e.g., for a fourfold dilution (e.g., 8 M to 2 M urea), the initial protein concentration (before denaturation) should be >1 µg/2.5 µl. If the sample is too dilute, use a Centricon microconcentrator (see manufacturer’s instructions) to reduce the volume. For the 8 M urea conditions, after addition of the buffer the final urea concentration is actually 7.3 M.
4. Transfer 1 µl sample to pH indicator strip. If pH is not correct, add sufficient 10× digestion buffer to increase buffer strength by 100 mM and recheck the pH. Enzymatic Digestion of Proteins in Solution
For pepsin, the pH must be 2.0. Subtilisin and endo Lys-C work well in a pH range of 7.0 to 10.5. For all other proteases, pH should be 8.0 to 8.5. Guanidine⋅HCl requires double buffer strength to adjust to mild alkaline pH.
11.1.4 Current Protocols in Protein Science
5. Add 1 µg/µl enzyme stock to a final enzyme/substrate ratio of 1:25 to 1:10 (w/w). Vortex sample lightly to mix. Microcentrifuge 10 sec at high speed. Enzyme concentration should be >1 µg/50 µl. Autolysis of proteases occurs in 8 M urea and in 2 M guanidine⋅HCl. The resulting peaks (run an auto-digestion control in parallel) must be accounted for when sample digest is analyzed by RP-HPLC.
6. Incubate sample at 37°C for the amount of time recommended for the selected protease (see Table 11.1.1). 7. Determine extent of digestion by analytical RP-HPLC (see Basic Protocol step 6). Store remainder of sample on ice. 8. If digestion is complete, proceed to step 9. If digestion is incomplete, check and adjust pH, then redigest (see Basic Protocol, step 7). If more enzyme is added to continue digestion, enzyme/substrate ratio must be <1:10. Do not use SDS-PAGE to check digests performed in the presence of guanidine⋅HCl, as guanidine⋅DS salts are insoluble. The analytical procedure should not consume >10% of the sample. If HPLC analysis cannot be performed immediately, samples may be stored indefinitely at −70°C.
9. If desired, reduce and alkylate peptides (see Support Protocol 2). NOTE: Samples alkylated with 4-vinylpyridine (Support Protocol 2) must be injected onto an HPLC column immediately. They cannot be stored even for 1 hr! On a 2.1–mm microbore column, operated at a 100 µl/min flow, it may take up to an hour for the 2-ME and 4-vinylpyridine to wash off the column.
10. Separate peptides by RP-HPLC (see Basic Protocol, step 8). DIGESTION OF PROTEINS IN THE PRESENCE OF SDS Proteins must be in solution before they can be adequately digested. Adding SDS to a protein sample is an excellent way to keep hydrophobic proteins (e.g., membrane proteins) in aqueous solution. At concentrations of ≥1%, and at elevated temperatures, SDS will also efficiently denature the substrates. The highest SDS concentrations compatible with the most commonly used proteases are listed in Table 11.1.1.
ALTERNATE PROTOCOL 2
Protein is solubilized (and denatured, if necessary) in SDS and digested with the desired protease. The extent of digestion can easily be monitored by SDS-PAGE. SDS is precipitated as the guanidine⋅DS salt and removed by centrifugation prior to separating the digested peptides by RP-HPLC. SDS can also be used to generate larger peptide fragments by providing conditions favorable for limited protease digestion; the fragments can then be separated by SDS-PAGE, followed by electroblotting onto a membrane, if desired. Additional Materials (also see Basic Protocol) 1× digestion buffer with 1% or 0.1% (w/v) SDS (see Table 11.1.1) 10% and 1% (w/v) SDS (Bio-Rad) 10× digestion buffer (no SDS) 1 M guanidine⋅HCl (sequenal grade, Pierce), prepared immediately before use 60°C and 95°C water baths 1a. If sample is solid: Dissolve protein sample in a minimal volume of digestion buffer containing 1% (or 0.1%) SDS. Heat sample 5 min at 37°C and sonicate 1 min. If Chemical Analysis
11.1.5 Current Protocols in Protein Science
pellet does not dissolve, heat sample 5 min at 60°C and sonicate 1 min. Repeat heating and sonication until protein dissolves. 1b. If sample is already in solution: Add 0.1 vol of 10% or 1% SDS (1% or 0.1% final) and 0.1 vol of 10× digestion buffer. Buffer composition and optimal SDS concentration differ for each protease. See Table 11.1.1 for recommended buffer conditions. IMPORTANT NOTE: Solution should not contain guanidine⋅HCl, as guanidine⋅DS salt will precipitate. Final substrate concentration should be >1 µg/10 µl. If protein sample is too dilute, use a Centricon concentrator (see manufacturer’s instructions) to reduce the sample volume.
2. Transfer 1 µl sample to pH indicator strip. If pH is not correct, add sufficient 10× digestion buffer to increase buffer strength by 100 mM and recheck pH. For pepsin, pH must be 2.0. Subtilisin and endoproteinase Lys-C work well in a pH range of 7.0 to 10.5. For all other proteases, pH should be 8.0 to 8.5.
3. If nondenatured protein is protease resistant, heat sample 5 min at 95°C to denature protein. Allow sample to cool to room temperature. For a partial list of protease-resistant proteins, see Table 11.1.2.
4. Add 1 µg/µl enzyme stock to a final enzyme/substrate ratio of 1:25 to 1:10 (or 1:100 to 1:50 for a limited digestion). Vortex sample lightly to mix. Microcentrifuge sample 10 sec at high speed. Enzyme concentration should be >1 µg/50 µl (or less—e.g., 1 µg/200 µl—for a limited digestion).
Table 11.1.2
Results of Proteolysis of Undenatured Substratesa,b,c
Substrate
#AA
Intrachain disulfide bonds
Cytochrome c Triosephosphate isomerase Carbonic anhydrase Trypsin inhibitor RNase Lysozyme Superoxide dismutase β-Lactoglobulin Ovalbumin Bovine serum albumin
104 248
0 0
259 56 124 129 151 162 385 577
0 3 3 4 1 2 1 17
T
C
KC
DN
EC
S
H
P
+++ +++ − −
+++ −
+++ −
− −
+++ −
+++ −
+++ −
− − − − − − − − − − +++ +++ − − +++ −
+++ − − − − +++ − +++
+ − − − − +++ − +
+ − − − − +++ − −
+++ − +++ +++ − +++ +++ +++
− − − − − +++ − −
+++ − ++ − − − − +++
aDigests were carried out at 37°C with enzyme/substrate ratios of 1:25 (w/w); buffer compositions and incubation times
were as listed in Table 11.1.1 (top rows) except for subtilisin digest of lysozyme, which was done for 5 hr. Analysis was done by RP-HPLC.
Enzymatic Digestion of Proteins in Solution
bAbbreviations: #AA, length of protein in amino acids; C, chymotrypsin; DN, Pseudomonas fragi endoprotease Asp-N; EC, Staphylococcus aureus V8 endoprotease Glu-C; H, thermolysin; KC, Achromobacter protease I, or endoprotease Lys-C; P, pepsin; S, subtilisin; and T, trypsin. cResults: +++, complete digest; ++, incomplete digest; +, very incomplete digest (>50% substrate left); −, no digest (no peptides, only substrate).
11.1.6 Current Protocols in Protein Science
5. Incubate at 37°C for the amount of time recommended for the selected protease (see Table 11.1.1). For a limited digest, incubate 15 to 30 min at 37°C. 6. Determine extent of digestion by analyzing 0.5 to 1 µg sample by SDS-PAGE (e.g., using a Phast Gel). Stain sample with Coomassie brilliant blue (UNIT 10.5). Phast Gels are useful when protein amounts are limited.
7. If sample is fully digested, proceed to step 8. If sample is not fully digested, check and adjust pH, then redigest (see Basic Protocol, step 7). If a limited digest is desired, proceed to step 8. If more enzyme is added to achieve a more extensive digest, the enzyme/substrate ratio must be <1:10. The analytical procedure should not consume more than 10% of the sample. Do not use RP-HPLC to check digest, because SDS and RP-HPLC are not compatible.
8. To remove SDS, add 1 vol of 1 M guanidine⋅HCl to sample and tap tube lightly to mix. A white precipitate will form immediately.
9. Microcentrifuge sample 10 min at high speed. Transfer supernatant to clean microcentrifuge tube or inject directly onto RP-HPLC. If HPLC analysis cannot be performed immediately, sample may be stored indefinitely at −70°C.
10. Reduce and alkylate sample if desired (see Support Protocol 2) 11. Separate peptides by RP-HPLC (see Basic Protocol, step 8). Do not inject solids onto HPLC column.
PREPARING AND USING ENZYME STOCKS This protocol details the handling of proteases—the solubilization of the enzyme from manufacturer stocks, the adjusting of concentrations and testing of activities under the most stringent conditions (listed in Table 11.1.1), and the thawing of enzyme stocks prior to experimental use. Materials Enzymes: Trypsin (sequencing grade, Boehringer Mannheim 1047-841 or Promega V5111) Chymotrypsin (sequencing grade, Boehringer Mannheim 1334-131) Endoproteinase Glu-C (endo Glu-C; sequencing grade, Boehringer Mannheim 1047-817) Endoproteinase Asp-N (endo Asp-N; sequencing grade, Boehringer Mannheim 1054-859) Endoproteinase Lys-C (endo Lys-C; Wako Chemicals 129-02541) Subtilisin (Sigma P5380) Thermolysin (Sigma P1512) Pepsin (Sigma P6887) Elastase (Sigma PE0258) Papain (Sigma P3125) Milli-Q water or equivalent Trifluoroacetic acid (TFA; Pierce) 5 mg/ml cytochrome c (Sigma) 10× digestion buffer (see Table 11.1.1) 5 mg/ml β-lactoglobulin (Sigma)
SUPPORT PROTOCOL 1
Chemical Analysis
11.1.7 Current Protocols in Protein Science
5 mg/ml triosephosphate isomerase (Sigma) 8 M urea (sequenal grade, Pierce; store up to several weeks at room temperature over Bio-Rad AG 501-X8 mixed-bed resin) 6 M guanidine⋅HCl (sequenal grade, Pierce) HPLC system and 4.6-mm reversed-phase column (see Basic Protocol) Solubilizing the Protease from Manufacturer Stocks For sequencing-grade proteases 1a. Dissolve protease (powder) in the shipping vial by adding distilled water to yield a final enzyme concentration of 1 µg/µl. 2a. Aliquot 5- or 10-µl volumes of 1 µg/µl stock solution into 0.5-ml microcentrifuge tubes. Keep one aliquot on ice for testing activity under nonstringent or stringent conditions. 3a. Place remaining aliquots inside a labeled 50-ml tube and store up to 1 year at −20°C. For proteases purchased in amounts ≥1 mg 1b. Transfer protease to a 1.5-ml microcentrifuge tube. Dissolve powder in water to obtain a final concentration of 5 mg/ml. 2b. Aliquot 200-µl volumes of 5 mg/ml stock solution to 0.5-ml microcentrifuge tubes. Keep one aliquot on ice for further dilution. Place remaining tubes inside a labeled 50-ml tube and store up to 1 year at −20°C. 3b. Dilute remaining 5 mg/ml stock solution with water to a final concentration of 1 mg/ml. Aliquot 5- and 10-µl volumes. Keep one aliquot on ice for testing activity under nonstringent or stringent conditions. Store remaining aliquots as described in step 2b. IMPORTANT NOTE: If proteases do not readily dissolve, tap microcentrifuge tube gently and warm <5 min at 37°C. If enzyme still does not dissolve, add TFA to final concentration of 0.1% (v/v). Tap tube gently and warm <5 min at 37°C. Frozen stocks are stable for < 1 year and trypsin may be stable for several years. Do not use 5 mg/ml stocks until 1 mg/ml stocks are depleted. Another tube of concentrated stock can then be thawed, diluted, and aliquoted as above.
4. Test protease activity under nonstringent conditions (steps 5 through 8) or stringent conditions (step 9 through 15). It is best to test under stringent conditions. If the enzyme passes that test, it is fine for everything. If it fails, it might still be useful for experiments under nonstringent conditions.
Testing Protease Activity Test under nonstringent conditions 5. In a 0.5-ml microcentrifuge tube, mix (25 µl total): 5 µl 5 µg/µl cytochrome c 16.5 µl water 2.5 µl 10× digestion buffer 1 µl 1 µg/µl enzyme stock. Vortex tube lightly to mix. Microcentrifuge 10 sec at high speed. Enzymatic Digestion of Proteins in Solution
11.1.8 Current Protocols in Protein Science
6. Incubate sample at 37°C for the amount of time recommended for the selected protease (see Table 11.1.1). 7. Analyze 1⁄4 of the sample (500 pmol) by RP-HPLC using a 4.6-mm column (see Basic Protocol step 8). β-lactoglobulin should be used as substrate for endoproteinase Glu-C, as this protease cannot degrade undenatured cytochrome c. Stocks should be tested periodically (every 6 months) or before a crucial experiment is to take place.
Test under stringent conditions 8. Prepare digests as follows: For trypsin, prepare digestion buffer containing 4 M urea (final); for endoproteinase Lys-C, subtilisin, or thermolysin, prepare a digest mixture containing 7.3 M urea (final) and cytochrome c as substrate or 2 M guanidine⋅HCl and triosephosphate isomerase as substrate. 9. Pipet 5 µl of 5 mg/ml cytochrome c (2 nmol) or 5 µl of 5 mg/ml triosephosphate isomerase (900 pmol) into a 0.5-ml microcentrifuge tube. Dry in a Speedvac evaporator. 10. Dissolve pellet in 20 µl of 8 M urea or 10 µl of 6 M guanidine⋅HCl, as follows: For 7.3 M urea (final): Add 2 µl 10× buffer For 4 M urea (final): Add 15 µl water and 4 µl 10× buffer For 2 M guanidine⋅HCl (final): Add 16 µl water and 3 µl 10× buffer. 11. Allow substrate to denature (see Alternate Protocol 2, steps 1 to 3). 12. Add 1 µl of 1 µg/µl enzyme stock. Vortex sample lightly to mix. Microcentrifuge 10 sec at high speed. 13. Incubate at 37°C for the amount of time recommended for the selected protease (see Table 11.1.1). 14. Analyze 1⁄4 (500 pmol) of the cytochrome c digests or 1⁄2 (450 pmol) of the triosephosphate isomerase digests by RP-HPLC using a 4.6-mm column (see Basic Protocol, step 8). Thawing Protease Stock Aliquots for Experimental Use 15. Remove tube containing 1 µg/µl stock solution of desired enzyme from −20°C freezer and place on ice. Allow entire sample to thaw. Tap tube lightly to mix solution and microcentrifuge 10 sec at high speed. 16. Pipet desired volume of stock solution into microcentrifuge tube containing substrate in buffer. 17. If ≤5 µl enzyme remains in stock tube, discard. If >5 µl enzyme remains, recap stock tube, mark cap with black dot, and return to −20°C. This tube can be used one more time; always discard any stock protease remaining after second time thawed.
Chemical Analysis
11.1.9 Current Protocols in Protein Science
SUPPORT PROTOCOL 2
REDUCTION AND S-ALKYLATION OF PEPTIDES IN DIGEST MIXTURES Unless they are reduced, disulfide bonds remain intact during the digestion procedures outlined in the Basic and Alternate Protocols. Several peptides will therefore be disulfidelinked and elute as single peaks during HPLC. In addition, underivatized cysteine residues cannot be identified easily by standard chemical sequencing. Derivatization will also prevent random disulfide bond formation when the reducing agent is removed and peptides are exposed to oxidative conditions. In this protocol, the peptide mixture is reduced, then S-alkylated with 4-vinylpyridine. The resulting cysteine derivative, pyridylethyl cysteine, is easily detectable during chemical sequencing. Materials Peptide digest mixture (see Basic Protocol or Alternate Protocols 1 or 2) 2-mercaptoethanol (2-ME; Bio-Rad): undiluted and 10% (v/v) solution (prepare in Milli-Q water or equivalent immediately before use) 4-vinylpyridine (Aldrich; store at −20°C), undiluted and 10% (v/v) solution (prepare in high-purity grade ethanol immediately before use) Argon (prepurified) HPLC system and reversed-phase column (see Basic Protocol) Reduce sample 1. Add 2 µl of 2-ME per 1 nmol original substrate protein. For smaller quantities of protein (e.g., 100 pmol), add 2 µl of 10% 2-ME. 2. Attach a Pasteur pipet to a length of Tygon tubing connected to the regulator on the argon tank. Adjust argon flow until it is gentle enough to be felt only barely when directed to the back of the hand. Direct argon stream over sample for 20 sec. 3. Cap tube, vortex lightly, and microcentrifuge briefly. 4. Incubate 30 min in 37°C water bath. If S-alkylation is unnecessary (i.e, if the protein will not be sequenced), proceed to step 8. Alkylate sample 5. Add 6 µl of 4-vinyl pyridine per 1 nmol reduced substrate protein. For smaller quantities of protein (e.g., 100 pmol), add 6 µl of 10% 4-vinylpyridine. IMPORTANT NOTE: Not all lots of 4-vinylpyridine are good, and there is also an aging effect. Quality control should be done on all new batches, and periodically thereafter, by derivatizing a model substrate (e.g., insulin) and checking for pyridylethyl incorporation (by HPLC analysis, monitoring absorbance at 253 nm). Aliquots (20 µl) are then stored under argon at −20°C for one-time use.
6. Direct stream of argon over sample for 20 sec. Cap tube, vortex lightly, and microcentrifuge briefly. 7. Incubate 30 min at room temperature. Do not start alkylation until the HPLC apparatus is completely ready for use (performance-tested and equilibrated at initial conditions); if necessary, the digest should be kept on ice or stored at −70°C prior to alkylation, until HPLC is ready.
Separate peptides by RP-HPLC 8. Inject sample immediately onto RP-HPLC column (see Basic Protocol, step 8). Enzymatic Digestion of Proteins in Solution
HPLC analysis must be performed immediately on S-alkylated samples. Do not let the reaction mixture sit for more than 15 to 20 min after the 30-min incubation, and do not try to store sample at −70°C; either action will severely affect the HPLC profile.
11.1.10 Current Protocols in Protein Science
COMMENTARY Background Information Detailed analysis of protein covalent structure usually requires analysis of peptides derived from the larger protein. In contrast to acid-promoted total hydrolysis (e.g., 6 N HCl), peptides are typically generated by selective proteolysis. Endoproteases fragment proteins by cleaving certain peptide bonds, with varying degrees of specificity (Beynon and Bond, 1989; Keil, 1991). In nature, most proteases digest nutrients, aid in cellular defense, or influence other physiological processes. When purified, the proteases with the greatest degree of specificity are useful tools for protein chemical analysis. Specificities of the most frequently used endoproteases are well established (Keil, 1981, 1991; see Table 11.1.3). When digesting substrates of known sequence, proteases can therefore be rationally selected. The number of fragments resulting from protease digestion will be proportional to the abundance of the targeted peptide bonds in the substrate protein. Average fragment size, on the other hand, is inversely related to the frequency of the targeted peptide bond. In cases where the protease specificity is determined solely by the P1 or P′1 residue (Table 11.1.3), the number of cleavages depends on the distribution of these particular amino acids. It is estimated that lysine (Lys), arginine (Arg), glutamic acid (Glu), and aspartic acid (Asp) each account for about 5% of all amino acids in the protein databases (Henzel et al., 1994). In contrast, tryptophan (Trp) is less abundant and glycine (Gly) more abundant. Thus, a tryptic or an endoproteinase Lys-C digest should yield about 11 or 6 fragments per 100 residues, respectively. Enzymatic digestion of a protein results in a pool of variously sized peptides. The complexity of the mixture is correlated to the molecular weight of the substrate. Although some information may be obtained by analyzing the mixture directly (e.g., by mass spectrometry), it is standard procedure to first perform some type of fractionation (e.g., reversed-phase (RP)-HPLC). The combined technique of digestion followed by separation is commonly called “peptide mapping,” particularly when used for comparative purposes. Because RP-HPLC can detect differences as small as a few carbon or oxygen atoms, peptides that differ by a single amino acid or by a post-translational modifica-
tion (either naturally occurring or experimentally induced) will elute with different retention times (i.e., at a different position in the peptide map). In principle, single amino acid substitutions or modifications can be detected and the distinct peptides isolated. More detailed information about molecular composition is then obtained from peptide sequencing and mass analysis. If the sequence of the substrate protein is known, any residue of interest can be precisely located. Similarly, sites of phosphorylation (Chapter 13), glycosylation (Chapter 12), or other posttranslational modifications can be located as long as the unmodified protein is digested as a control. Modification can be induced in vivo or performed in vitro. If a protein is known to be constitutively modified in vivo, the modification can be selectively removed from the “control” protein (e.g., by phosphatases or endoglycosidases). Peptide mapping is facilitated when the modifying group is also detectable by radioisotopic or spectrophotometric monitoring. Peptide mapping can also be used to detect and localize a protein disulfide array. In this case, reduced and unreduced peptides will be differently positioned in the comparative map, enabling identification of paired cysteines. Finally, mapping is also a preferred tool to verify the authenticity of overexpressed proteins (Christianson and Paech, 1994). When digesting proteins of unknown sequence, fragmentation was traditionally performed, not for mapping purposes, but to generate sets of overlapping peptide sequences (e.g., tryptic and chymotryptic peptide digests) essential for deducing the full protein structure (Tempst and Van Beeumen, 1983). With the advent of DNA cloning technology, internal peptide sequences facilitate conventional and PCR-based cloning of the corresponding genes, which can then be easily sequenced (Tempst et al., 1990). Moreover, “internal” sequencing is most often required to overcome the problem of blocked N-termini, a major obstacle for protein chemical analysis. Efficient proteolytic cleavage is often hampered or prevented due to the insolubility or protease resistance of the substrates. A rather large percentage of membrane proteins and secreted proteins present these problems. Denaturing or chemically modifying the substrate protein can improve digestion efficiency by enhancing solubility or breaking up any tightly packed structures. However, such manipula-
Chemical Analysis
11.1.11 Current Protocols in Protein Science
Table 11.1.3
Specificity of Selected Proteolytic Enzymesa
Enzyme
Specificity
Comments
Trypsin
P1 = Arg > Lys
Inhibited with P2 - - - - - P1 - - - - - - ↓ (1) X - - - - - Arg/Lys - - ↓ (2) Asp - - - - Lys - - - - - ↓ (3) Arg - - - - Arg - - - - - ↓ Reduced cleavage seen with (1) X - - - - - Lys - - - - - ↓ (2) X - - - - - Arg - - - - - ↓ (3) Asp - - - - Lys - - - - - ↓ (4) Asp - - - - Arg - - - - - ↓ (5) X - - - - - Arg/Lys - - ↓ (6) Arg - - - - Arg/Lys - - ↓ (7) Lys - - - - Lys - - - - - ↓ (8) Lys - - - - Arg - - - - ↓
-------------
P1′ Pro Asp Arg/His
-------------------------
Asp/Glu Asp Asn/Gln/His/Phe/Leu Glu/His Arg/Lys X X His
Endoproteinase Lys-C (KC) Endoproteinase Glu-C
P1 = Lys
Not inhibited when P1′ = Pro
P1 = Glu >>> Asp
Endoproteinase Asp-N Chymotrypsin
P1′ = Asp
Reduced cleavage with P1′ = Leu/Phe or with P1 surrounded by Glu/Asp Ammonium bicarbonate strongly inhibits, making it selective for the Glu-X bond under standard digest conditions. Adding more enzyme or extending incubation times might help cleave at Asp. In phosphate buffer, Asp-X bonds cleaved (at a 15% rate of Glu-X bonds). —
Subtilisin
Thermolysin
Pepsin
Elastase
Papain
P1 = Trp > Tyr > Phe >> Leu > Met >> His >>> Asn/Gly
Cleavage after Arg/Lys possible, but may be unspecific or the result of contaminating trypsin Inhibited with P1′ = Pro Enhanced cleavage seen with (1) P3, P1′, P2′, or P3′ = Arg (Lys) (2) P2 = Pro Reduced cleavage seen with (1) P2, P1′, or P2′ = Asp (Glu) (2) P3 or P2′ = Pro P1 = neutral or acidic amino Enhanced cleavage seen when acid (broad specificity) (1) P1′ = Gly (2) P2 = bulky/hydrophobic amino acid Inhibited with P2′ = Pro; not inhibited with P1 = Pro P1′ = Leu/Ile > Phe > Val >> Tyr > Ala Enhanced fragmentation with P1 = Phe/Tyr/Trp Reduced fragmentation with P1 = Glu/Asp — P1 or P1′ = Phe >> Leu >> Trp > Ala > other hydrophobic amino acids — P1 or P1′ = Ile > Val > Ala Gly/Ser (and other neutral, nonaromatic amino acids) P2 = hydrophobic amino Very broad specificity; extensive degradation acid Enhanced fragmentation with P1 = Lys/Arg
aCleavage site nomenclature: P - - - P - - - P - - - ↓ - - - P ′ - - - P ′ - - - P ′, where ↓ marks the site of cleavage. 3 2 1 1 2 3
11.1.12 Current Protocols in Protein Science
tions (e.g., reduction and alkylation in the presence of guanidine⋅HCl, followed by dialysis or chromatography) are tedious and frequently result in heavy losses, particularly when working with protein quantities <1 nmol. Moreover, some unstable chemical groups (e.g., carbetoxy histidines) or bonds (e.g., disulfides) get destroyed in the process and can therefore no longer be mapped. Instead, it is easier to supplement the digestion buffer with agents that promote substrate solubility and denature the protein without interfering with protease activity. For this reason, protocols for protease digestion in the presence of organic solvents, detergents, urea, and guanidine⋅HCl have been developed (see Alternate Protocol 1 and Alternate Protocol 2). Further advantages of such approaches include improved ability to dissolve protein pellets, avoidance of the need to remove additives present after chromatography or electrophoresis, increased reproducibility (e.g., in chaotrope solutions), and limited proteolysis of solubilized proteins in their native state (e.g., in nondenaturing detergents such as CHAPS).
preferentially using protease-resistant substrates (e.g., superoxide dismutase or triosephosphate isomerase). All endoproteases should digest cytochrome c quite easily under nondenaturing conditions, except endoproteinase Glu-C which works best on β-lactoglobulin (see Table 11.1.2). If any of these low-stringency quality control tests fail, the investigator should complain to the enzyme’s manufacturer and demand a new batch at no extra charge. All buffer components must be “ultra-pure” or “reagent-grade” quality, and the water of Milli-Q purity. Guanidine⋅HCl solutions must always be made fresh; many problems have been associated with solutions that have aged for only a few days. Urea solutions can be stored up to several weeks at room temperature over Bio-Rad AG 501-X8 mixed-bed resin to capture any cyanates that might form with time. Do not use the blue indicator dye (“D”) form of the resin, as the dye has been found to leach out into the urea and interfere with RP-HPLC. Newly prepared stock solutions should be tested with enzyme batches previously found to work well.
Critical Parameters and Troubleshooting
Protease compatibilities Endoproteases have been systematically tested in the presence of increasing concentrations of urea, guanidine⋅HCl, SDS, nonionic and zwitterionic detergents, and acetonitrile and isopropanol, using cytochrome c and β-lactoglobulin as model substrates (Riviere et al., 1991; L.R.R. and P.T., unpub. observ.). The highest concentrations of additives that still permit a fairly complete digest are listed in Table 11.1.1. Cytochrome c peptide maps resulting from these digests are shown in Fig. 11.1.1. It should be noted that the conditions listed for digests containing urea and guanidine⋅HCl have been carefully optimized. Small changes may result in a drastically different outcome of an experiment, and, if so desired, modified conditions should be tested prior to the real experiment. It is advisable to not change conditions within any set of comparative peptide mapping experiments. In the presence of 8 M urea, Tris⋅Cl buffers work best for endoproteinase Lys-C and subtilisin digestion, tending to give higher yields and fewer partial cleavages than bicarbonate buffers. Both types of buffer are equally compatible with thermolysin digestion in 8 M urea, although Tris⋅Cl digests consistently generate larger fragments. Trypsin activity in 4 M urea is borderline and requires the presence of Ca2+ and a lengthy (24-hr) incubation to ensure even
In practice, with many substrates, proteolytic digests under nondenaturing conditions are essentially a waste of time; they usually don’t work (see Table 11.1.2). Unless special reasons exist to subject a protein to proteolytic attack in its native state, it is advisable to include urea or guanidine⋅HCl in the digest mixture by default. Provided that the conditions summarized in Table 11.1.1 are followed, and enzymes and buffer components are of good quality (and have been stored correctly), the failure rate for protease digestion will be minimal. Access to a well-maintained HPLC system is also prerequisite. Enzymes and buffer components Enzyme activity must be assessed when a new lot (or brand) is received, and then periodically thereafter, either after prolonged storage or whenever concentrated stock solutions are diluted and aliquoted (see Support Protocol 2). The suppliers and grades that have worked consistently well in our hands are listed in the Materials section of Support Protocol 1. Repeated cycles of freezing and thawing should be avoided by preparing small (single-use) aliquots (see Support Protocol 1). Activities should be checked under conditions at least as stringent as those of the planned experiment,
Chemical Analysis
11.1.13 Current Protocols in Protein Science
no additives
0.1 M NH4OH
2 M urea
1% SDS
4 M urea CaCl2
8 M urea
1 M Gu•HCI
2 M Gu•HCI
2% OG
2% OG
40% MeCN
40% MeCN
2% NP-40
Figure 11.1.1 Effects of additives on proteolytic digestion of 1 nmol of cytochrome c with 1 µg trypsin (left) or 1 µg endoproteinase Lys-C (right). Digest conditions are listed on the panels. Digest mixtures were separated by reversed-phase HPLC on a 4.6-mm Vydac C18 column, using gradient conditions and flow rate given in the Basic Protocol. Abbreviations: MeCN, acetonitrile; OG, octyl glucoside.
Enzymatic Digestion of Proteins in Solution
partial digestion (Fig. 11.1.1). In general, artificial blocking (i.e., carbamylation) of the newly generated N-termini during digestion in the presence of urea is not evident, as determined by sequencing. Whereas endoproteinase Lys-C and subtilisin work well in the presence of 2 M guanidine ⋅HCl, chymotrypsin does so only at higher
bicarbonate concentrations and after prolonged incubation (24 hr). Trypsin does not digest proteins in guanidine⋅HCl solutions because guanidine is a competitive inhibitor of the enzyme (Fig. 11.1.1). Some batches of trypsin, less pure than the Boehringer-Mannheim or Promega sequencing grades, have been found to work under these conditions. However, lim-
11.1.14 Current Protocols in Protein Science
trypsin inhibitor
superoxide dismutase
triosephosphate isomerase
A
B
C
D
E
Figure 11.1.2 Enzymatic digestion of protease-resistant substrates. 2 nmol trypsin inhibitor (left panels), 1 nmol superoxide dismutase (middle panels), and 1 nmol triosephosphate isomerase (right panels) were digested by 1 µg endoproteinase Lys-C. Digestion conditions were as follows: (A) no additives; (B) 8 M urea; (C) 1 M guanidine⋅HCl (D) 2 M guanidine⋅HCl; (E) 1% SDS (with guanidine⋅DS precipitation). Digest mixtures were separated by reversed-phase HPLC on a 4.6-mm Vydac C18 column, using gradient conditions and flow rate as given in the Basic Protocol.
ited sequencing of the resulting peptides indicated chymotryptic-like cleavages. Although enzymatic digestion in the presence of SDS has been described (Cleveland et al., 1977), it has not found widespread use in combination with HPLC, primarily because dodecyl sulfate interferes with peptide separation on reversed-phase packings. Excess SDS can be removed by guanidine⋅HCl precipitation (as guanidine⋅DS salt) prior to RP-HPLC. Chymotrypsin and endoproteinase Glu-C retain activity in 0.1% SDS. Although endo Glu-C digests are usually incomplete, chymotryptic di-
gestion is quantitative but restricted (i.e., only larger fragments are reproducibly generated). If the substrate is first heated at 95°C, endoproteinase Lys-C and subtilisin can degrade almost any protein in 1% SDS (Fig. 11.1.2). Recovery after guanidine⋅DS precipitation is highly variable, with the worst yields for larger or hydrophobic peptides (Figs. 11.1.1 and 11.1.2). This is not a problem with SDS concentrations ≤0.1%. With few exceptions, enzymatic proteolysis works quite well in 2% nonionic or zwitterionic detergents, and in acetonitrile or isopropanol
Chemical Analysis
11.1.15 Current Protocols in Protein Science
at concentrations up to 40%. Endoproteinase Lys-C even works in 8 M urea/2% NP-40, a very strong solubility-promoting combination. When heated, several proteins precipitate in organic solvent concentrations over 40%. Very detailed studies on digestion with organic solvents have been described by Welinder (1988). Under certain conditions, digests are restricted; i.e., there are fewer peaks than usual but all in good yields. Combinations of enzymes and additives that yield restricted digests are indicated in Table 11.1.1. With no exceptions, even with enzymes of low specificity (e.g., subtilisin, thermolysin, and pepsin), peptide patterns are very reproducible under a strictly controlled set of conditions. All enzymes and additives can therefore be used in peptide mapping experiments.
Enzymatic Digestion of Proteins in Solution
Additional experimental parameters Enzymatic proteolysis is generally done at 37°C. However, most enzymes work fairly well in the 20° to 50°C range. Optimal temperature for thermolysin activity is actually 60°C, but this offers no apparent advantage over its use at 37°C. Whereas pH optima of most proteases are quite narrow, considerable flexibility exists for subtilisin and endoproteinase Lys-C digests, which can proceed at pH 10.5 (Table 11.1.1). Reaction times listed in Table 11.1.1 are usually adequate. Prolonged incubation provides absolutely no advantage. If no favorable result is obtained at the end of the indicated time periods, something is wrong with the experiment or the chosen combination of enzyme and denaturant is not optimal for the particular substrate. The former situation can be monitored by performing the proper control digests. The latter can be alleviated by trying a different combination of enzyme and denaturant. To ensure complete digestion, the final enzyme concentrations should be >1 µg/100 µl of reaction mixture under native conditions and >1 µg/50 µl of reaction mixture in the presence of guanidine⋅HCl, urea, or SDS. This restriction essentially determines the enzyme/substrate ratios, with the clear rule that those ratios should be kept as low as possible to avoid excessive autolytic digestion of the protease. Finally, it is imperative that the substrate be in solution before the protease is added. Never try to improve solubility (e.g., by heating, sonicating, or vortexing) in the presence of the enzyme, as this will result in loss of activity.
Protease-resistant substrates Proteases active in 2 M guanidine⋅HCl and 8 M urea provide efficient tools to digest substrates whose physical properties prevent normal enzymatic degradation. A brief survey of the literature along with additional tests indicate that ribonuclease, ADP-ribosyl cyclase, lysozyme, amylase, superoxide dismutase, triosephosphate isomerase, xylose isomerase, pancreatic trypsin inhibitor, and many other proteins are quite resistant to protease digestion (see Table 11.1.2). After heating in 6 M guanidine⋅HCl and subsequent dilution to 2 M urea, they apparently do not refold to a compact structure, rendering them amenable to digestion. Alternatively, 8 M urea can be used to denature the protein. All the protease-resistant substrates listed above have been digested successfully using one or the other technique, but not always by both (Vangrysperre et al., 1989; Fig. 11.1.2). When the guanidine⋅HCl or urea concentrations are lowered to 1 M or 4 M, respectively, the digests no longer proceed. This excludes the use of endoproteinase Glu-C and trypsin for these purposes. In general, guanidine⋅HCl in combination with endoproteinase Lys-C is the method of choice. Endoproteinase Lys-C and subtilisin undergo a substantial amount of autolysis in 2 M guanidine⋅HCl, 8 M urea, or 1% SDS. Care must be taken to not confuse these peaks with the real peptide map. Autolysis profiles are fairly reproducible. Thus, with enzyme/substrate ratios of ≤1:10 and appropriate enzyme blank experiments, mistakes can usually be avoided. Should an autolytic fragment be accidentally analyzed by sequencing or mass spectrometry, the error can be quickly traced by comparing the sequence with the known sequence of the protease (Table 11.1.4). Limited and partial digestion Although the usual goal of a proteolytic digest is to fully cleave all susceptible bonds and generate a complete peptide map, sometimes restricted digestion is advantageous. Cleavage of an artificially low number of bonds, each one to completion, will yield fewer (and bigger) fragments and result in less complicated chromatograms. Addition of chaotropes will sometimes lead to exactly such an effect. Mild digestion of a protein in its native state may provide useful information on the domain structure and surface topography (Marks et al., 1990). The protein is thereby kept soluble in a nondenaturing detergent (e.g., CHAPS) solu-
11.1.16 Current Protocols in Protein Science
Table 11.1.4
Protease Sequences
Enzyme
Sequence
α-Trypsin (bovine) Chain 1
IVGGYTCGAN
TVPYQVSLNS
GYHFCGGSLI
NSQWVVSAAH
CYKSGIQVRL
GEDNINVVEG
NEQFISASKS
IVHPSYNSNT
LNNDIMLIKL
KSAASLNSRV
ASISLPTSCA
SAGTQCLISG
WGNTK (125)
SSGTSYPDVL
KCLKAPILSD
SSCKSAYPGQ
ITSNMFCAGY
LEGGKDSCQG
DSGGPVVCSG
KLQGIVSWGS
GCAQKNKPGV
YTKVCNYVSW
IKQTIASN (98)
IVGGYTCAAN
SIPYQVSLNS
GSHFCGGSLI
NSQWVVSAAH
CYKSRIQVRL
GEHNIDVLEG
NEQFINAAKI
ITHPNFNGNT
LDNDIMLIKL
SSPATLNSRV
ATVSLPRSCA
AAGTECLISG
WGNTK (125)
SSGSSYPSLL
QCLKAPVLSD
SSCKSSYPGQ
ITGNMICVGF
LEGGKDSCQG
DSGGPVVCNG
QLQGIVSWGY
GCAQKNKPGV
YTKVCNYVNW
IQQTIAAN (98)
GVSGSCNIDV
VCPEGDGRRD
IIRAVGAYSK
SGTLACTGSL
VNNTANDRKM
YFLTAHHCGM
GTASTAASIV
VYWNYQNSTC
RAPNTPASGA
NGDGSMSQTQ
SGSTVKATYA
TSDFTLLELN
NAANPAFNLF
WAGWDRRDQN
YPGAIAIHHP
NVAEKRISNS
TSPTSFVAWG
GGAGTTHLNV
QWQPSGGVTE
PGSSGSPIYS
PEKRVLGQLH
GGPSSCSATG
TNRSDQYGRV
FTSWTGGGAA
ASRLSDWLDP
ASTGAQFIDG
LDSGGGTP (268)
VILPNNDRHQ
ITDTTNGHYA
PVTYIQVEAP
TGTFIASGVV
VGKDTLLTNK
HVVDATHGDP
HALKAFPSAI
NQDNYPNGGF
TAEQITKYSG
EGDLAIVKFS
PNEQNKHIGE
VVKPATMSNN
AETQVNQNIT
VTGYPGDKPV
ATMWESKGKI
TYLKGEAMQY
DLSTTGGNSG
SPVFNEKNEV
IGIHWGGVPN
EFNGAVFINE
NVRNFLKQNI
EDIHFANDDQ
PNNPDNPDNP
NNPDNPNNPD
EPNNPDNPNN
PDNPDNGDNN
NSDNPDAA (268)
Chain 2 α-Trypsin (pig) Chain 1
Chain 2
Endoproteinase Lys-C (Achromobacter lyticus strain M497-1)
Endoproteinase Glu-C (Staphylococcus aureus strain V8)
Chymotrypsin A (bovine) CGVPAIQPVL A chain B chain
C chain
SGL (13)
IVNGEEAVPG
SWPWQVSLQD
KTGFHFCGGS
LINENWVVTA
AHCGVTTSDV
VVAGEFDQGS
SSEKIQKLKI
AKVFKNSKYN
SLTINNDITL
LKLSTAASFS
QTVSAVCLPS
ASDDFAAGTT
CVTTGWGLTR (130)
YTNANTPDRL
QQASLPLLSN
TNCKKYWGTK
IKDAMICAGA
SGVSSCMGDS
GGPLVCKKNG
AWTLVGIVSW
GSSTCSTSTP
GVYARVTALV
NWVQQTLAAN (100)
tion and the enzyme activity attenuated through reduced enzyme/substrate ratios (e.g., <1:100), lower digestion temperatures (≤20°C), and shorter incubation times (≤30 min). Fragments can be separated by HPLC or SDS-PAGE. In contrast to experimentally designed limited proteolysis, partial digestions also occur quite often unintentionally. The problem derives from incomplete (10% to 90%) cleavage of certain peptide bonds which, through combinatorial overlaps, results in many more peptides than expected and lower yields. Partial
digestions are typically caused by incompletely denatured substrates or proteases with low activity. The necessary corrective measures are self-evident. Preventing failures Provided that the fractionation techniques were carried out properly, the absence of a peptide profile during HPLC or after SDSPAGE indicates that the digest didn’t work or that there was a lack of sufficient substrate to begin with. The protein could have been lost
Chemical Analysis
11.1.17 Current Protocols in Protein Science
during final preparation (e.g., dialysis or precipitation) or transfer, or not been properly solubilized. The quantity of the starting substrate should be estimated by SDS-PAGE. Other factors that may contribute to failure include inactive proteases, the presence of protease inhibitors (carried over from the protein purification procedure), an incorrect pH, or a concentration of denaturants that was either too low (so that the substrate refolded) or too high (so that the protease was inactive). Preliminary testing of enzyme and digest conditions on model substrates, and pilot optimization experiments using dispensable amounts of the “real” protein, will largely prevent such mishaps from occurring. In addition, when a preparative experiment is carried out, analyzing 5% to 10% of the sample, while keeping the rest on ice or frozen, prior to preparative chromatography or electrophoresis will prevent a preparative “total loss” in case something goes wrong unexpectedly. It should be noted that undigested guanidine⋅HCl–denatured protein cannot usually be recovered from reversed-phase columns, because the exposed hydrophobic core of the protein becomes irreversibly adsorbed. This could lead the investigator to conclude that there was no protein present at the onset of the experiment.
HPLC separation takes 1 to 1.5 hr. The entire procedure can be completed in 1 day (for the shorter digest times), but enzymes, reagents, and the HPLC apparatus should be checked the day before. The HPLC should be checked again using a standard peptide mixture a few hours before running the actual digest mixture. A set of samples may be digested simultaneously, with the restriction that only three or four HPLC experiments can be completed in one (long) day on one instrument using manual collection. This is a critical consideration when peptide mixtures are S-alkylated with 4vinylpyridine (because they cannot be stored). Underivatized analytical digests can be processed using an autosampler.
Anticipated Results
Henzel, W.J., Billeci, T.M., Stults, J.T., Wong, S.C., Grimley, C., and Watanabe, C. 1994. Identification of 2-D gel proteins at the femtomole level by molecular mass searching of peptide fragments in the protein sequence database. In Techniques in Protein Chemistry V (J.W. Crabb, ed.) pp. 3-9. Academic Press, San Diego.
In the basic protocol, 100 pmol of cytochrome c digested with any of the enzymes listed in Table 11.1.1 will yield a nice peak profile after separation on a 4.6-mm C4 or C18 reversed-phase column, with an absorbance detector setting of 0.05 AUFS at 214 nm. Fifty pmol may be digested and separated on a 2.1mm column. Digestion in the presence of chaotropes and SDS requires about twice the amount of starting material. When substrate protein is available in quantities <100 pmol, in situ digests are more efficient than those in solution, both in terms of sample preparation and experimental losses. The drawback is that not all peptides are recovered, which makes in situ procedures less than ideal for peptide mapping experiments.
Time Considerations
Enzymatic Digestion of Proteins in Solution
Time required for enzymatic digestion of protein sample in solution can be broken down as follows: dissolving and denaturing the sample should take 15 to 45 min; preparing solutions for the digest will take 20 min; protease digestion requires from 2 to 24 hr; reduction and alkylation of sample will take 1 hr; and
Literature Cited Beynon, R.J. and Bond, J.S. 1989. Proteolytic Enzymes: A Practical Approach. Oxford University Press, Oxford. Christianson, T. and Paech C. 1994. Peptide mapping of subtilisins as a practical tool for locating protein sequencing errors during extensive protein engineering projects. Anal. Biochem. 223:119-129. Cleveland, D.W., Fischer, S.G., Kirschner, M.W., and Laemmli, U.K. 1977. Peptide mapping by limited proteolysis in sodium dodecyl sulfate by gel electrophoresis. J. Biol. Chem. 252:11021106.
Keil, B. 1981. Enzymic cleavage of proteins. In Methods in Protein Sequence Analysis (M. Elzinga, ed.) pp. 291-304. Humana Press, Clifton, N.J. Keil, B. 1991. Specificity of Proteolysis. SpringerVerlag, Heidelberg. Marks, A.R., Fleischer, S., and Tempst, P. 1990. Surface topography analysis of the ryanodine receptor/junctional channel complex based on proteolysis sensitivity mapping. J. Biol. Chem. 265:13143-13149. Riviere, L.R., Fleming, M., Elicone, C., and Tempst, P. 1991. Study and applications of the effects of detergents and chaotropes on enzymatic proteolysis. In Techniques in Protein Chemistry II (J.J. Villafranca, ed.) pp. 171-179. Academic Press, San Diego. Tempst, P. and Van Beeumen, J. 1983. The amino acid sequence of cytochrome C-556 from Agrobacterium tumefaciens strain Apple 185. Eur. J. Biochem. 135:321-330.
11.1.18 Current Protocols in Protein Science
Tempst, P., Link, A.J., Riviere, L.R., Fleming, M., and Elicone, C. 1990. Internal sequence analysis of proteins separated on polyacrylamide gels at the sub-microgram level: Improved methods, applications and gene cloning strategies. Electrophoresis 11:537-553.
Welinder, K.G. 1988. Generation of peptides suitable for sequence analysis by proteolytic cleavage in reversed-phase high-performance liquid chromatography solvents. Anal. Biochem. 174:54-64.
Vangrysperre, W., Ampe, C., Kersters-Hilderson, H., and Tempst, P. 1989. Single active-site histidine in D-xylose isomerase from Streptomyces violaceoruber: Identification by chemical derivatization and peptide mapping. Biochem. J. 263:195-199.
Contributed by Lise R. Riviere OsteoArthritis Sciences, Inc. Cambridge, Massachusetts Paul Tempst Memorial Sloan-Kettering Cancer Center New York, New York
Chemical Analysis
11.1.19 Current Protocols in Protein Science
Enzymatic Digestion of Proteins on PVDF Membranes
UNIT 11.2
Enzymatic digestion of membrane-bound proteins is one of the most widely used procedures for determining the internal amino acid sequence of proteins that either have a blocked amino terminus or require two or more stretches of sequence data for DNA cloning or confirmation of protein identification. Because the final step of protein purification is usually SDS-PAGE, electroblotting to either polyvinylidene difluoride (PVDF) or nitrocellulose is the simplest and most common procedure for recovering protein free of contaminants (e.g., SDS or acrylamide) with a high yield. As described in this unit, PVDF is preferred over nitrocellulose because it can be used for a variety of other structural analysis procedures, such as amino-terminal sequence analysis and amino acid analysis. In addition, peptide recovery from PVDF membranes is higher than from nitrocellulose, particularly from higher-retention PVDF (e.g., ProBlott, Transblot, Westran, or Immobilon Psq). Finally, PVDF-bound protein can be stored dry, as opposed to nitrocellulose-bound protein, which must remain wet during handling and storage to prevent loss of peptides during digestion. DIGESTION OF PVDF-BOUND PROTEINS IN A HYDROGENATED TRITON X-100 BUFFER
BASIC PROTOCOL
This procedure describes enzymatic digestion of proteins bound to PVDF following SDS-PAGE (UNITS 10.1-10.4) and electroblotting (UNIT 10.7). PVDF-bound proteins are visualized by staining and protein bands are excised from the membrane and placed directly in hydrogenated Triton X-100 (RTX-100) digestion buffer. RTX-100 acts both as an elution reagent for removing peptides from the membrane and as a blocking reagent that prevents any proteinase from adsorbing to the PVDF membrane. After a 24-hr incubation with a proteinase, the peptide fragments are recovered from the digestion buffer. Further washes of the membrane remove any remaining peptides, and the pooled digestion buffer and washes can be analyzed by reversed-phase HPLC (UNIT 11.6). The amino acid sequence can then be determined by either gas-phase or pulsed liquid-phase sequence analysis. NOTE: The key to success with this procedure is cleanliness. Use of clean buffers, tubes, and staining/destaining solutions, as well as only hydrogenated Triton X-100, greatly reduces contaminant peaks during reversed-phase HPLC. A corresponding blank piece of membrane must always be analyzed at the same time a sample is digested, as a negative control. Materials Protein samples electrophoresed on SDS-polyacrylamide gel (UNITS 10.1-10.4) PVDF membrane (e.g., ProBlott, Immobilon Psq, or Westran) Milli-Q water or equivalent Digestion buffer (see recipe and Table 11.2.1) Appropriate 0.1 µg/µl enzyme solution (depending on the cleavage site required; see recipe) 0.1% (v/v) trifluoroacetic acid (TFA) in Milli-Q water or equivalent 1% (v/v) diisopropylfluorophosphate (DFP) in ethanol (see recipe) Powder-free gloves Glass plate Forceps Sonicating water bath Contributed by Joseph Fernandez and Sheenah M. Mische Current Protocols in Protein Science (1995) 11.2.1-11.2.10 Copyright © 2000 by John Wiley & Sons, Inc.
Chemical Analysis
11.2.1 CPPS
Table 11.2.1
Enzyme
Digestion Buffers for Various Enzymesa
Digestion buffer
Recipe
Comments
Trypsin or 1% RTX-100/10% 100 µl 10% RTX-100 stock endoproteinase acetonitrile/100 100 µl acetonitrile Lys-C mM Tris⋅Cl, pH 8.0 300 µl HPLC-grade water 500 µl 200 mM Tris⋅Cl, pH 8.0
RTX-100 prevents enzyme adsorption to membrane and increases recovery of peptides Endoproteinase 1% RTX-100/100 100 µl 10% RTX-100 stock Acetonitrile is omitted Glu-C mM Tris⋅Cl, pH 8.0 400 µl HPLC-grade water because it decreases 500 µl 200 mM Tris⋅Cl, pH 8.0 digestion efficiency of endoproteinase Glu-C Clostripain 1% RTX-100/10% 100 µl 10% RTX-100 stock DTT and CaCl2 are acetonitrile/2 mM 100 µl acetonitrile necessary for DTT/1 mM 45 µl 45 mM DTT clostripain activity CaCl2/100 mM 10 µl 100 mM CaCl2 Tris⋅Cl, pH 8.0 245 µl HPLC-grade water 500 µl 200 mM Tris⋅Cl, pH 8.0 aSee APPENDIX 2E for Tris⋅Cl and DTT stock solution recipes. Abbreviations: DTT, dithiothreitol; RTX-100, hydrogenated
Triton X-100 (Tiller et al., 1984).
Injection vials for HPLC Microbore HPLC Clean capless 1.5-ml microcentrifuge tubes and separate caps (e.g., Sarstedt) Additional reagents and equipment for electroblotting (UNIT 10.7), staining transferred proteins (UNIT 10.8), and reversed-phase HPLC (UNIT 11.6) Transfer sample to PVDF membrane and stain protein bands 1. Electroblot proteins from polyacrylamide gel to PVDF membrane. For most efficient transfer, follow protocol for tank-transfer system rather than semi-dry system. When separating the protein sample by SDS-PAGE, concentrate as much protein into each lane as possible; however, if protein resolution is a concern, up to 20 individual bands of protein (∼ 5cm2) can be excised and combined in a single proteinase digestion reaction.
2. Stain PVDF membrane with Ponceau S, amido black, India ink, or Coomassie brilliant blue-G. Destain membrane until background is clean enough to visualize stained protein bands. Rinse destained membrane three times with distilled water to remove excess acetic acid. Ponceau S and amido black are preferred because they generally contain the fewest contaminants. Most commercial sources of Coomassie brilliant blue contain a large amount of contaminating material and should be avoided. If using Coomassie brilliant blue, we have found that preparations with greater than 90% dye content (e.g., from Aldrich) is suitable.
3. Excise desired protein bands from membrane and place in 1.5-ml microcentrifuge tube. Also excise a blank region of membrane, approximately the same size as protein band, to serve as a negative control. The negative control should be cut from the same blot as the protein sample and should have undergone the same manipulations (e.g., electroblotting, staining, destaining).
4. Air dry PVDF-bound protein at room temperature and store at −20° or 4°C. Enzymatic Digestion of Proteins on PVDF Membranes
IMPORTANT NOTE: If nitrocellulose must be used, the membrane-bound protein must be stored wet at −20°C and never allowed to completely dry out. If the membrane dries, peptide recovery can be greatly diminished.
11.2.2 Current Protocols in Protein Science
Digest membrane-bound protein with appropriate proteinase IMPORTANT NOTE: Gloves should be worn during all steps to avoid contamination of membrane with skin keratins. 5. Place ∼100 µl Milli-Q water onto a clean glass plate. Submerge membrane samples into water droplet and transfer wet membrane to a dry area of the plate. Cut membrane lengthwise into 1-mm-wide strips with a clean razor blade. Cut membrane strips into 1 × 1–mm squares. Treat negative control using identical conditions as the samples. Keeping the membrane wet will simplify manipulation of the sample as well as minimize any static charge, which could cause PVDF membrane to “jump” off the plate. The 1 × 1–mm pieces of membrane will settle to the bottom of the tube and require less digestion buffer to completely immerse the membrane.
6. Slide membrane squares onto forceps and place in a clean 1.5-ml microcentrifuge tube. Use the cleanest tubes possible to minimize contamination during peptide mapping. Surprisingly, many UV-absorbing contaminants can be found in microcentrifuge tubes, and this appears to vary with supplier and lot number. Tubes can be cleaned by rinsing with 1 ml HPLC-grade methanol followed by 2 rinses with 1 ml Milli-Q water prior to adding the protein band.
7. Add 50 µl of appropriate digestion buffer to membrane squares. Vortex sample 10 to 20 sec and incubate 5 to 30 min at room temperature. Add 50 µl digestion buffer to an empty microcentrifuge tube to serve as an RTX-100 blank for HPLC analysis (optional). Digestion buffers for typical proteases are listed in Table 11.2.1. The amount of digestion buffer can be increased or decreased depending on the amount of membrane; however, the best results will be obtained with a minimum amount of digestion buffer. PVDF membrane will float in the solution at first, but will submerge after a short time, depending on the type and amount of PVDF.
8. Add a volume of 0.1 µg/µl enzyme solution that yields an enzyme-to-substrate ratio of ∼1:10 (w/w). Incubate samples (including RTX-100 blank) 22 to 24 hr at 37°C. The amount of substrate protein on membrane can be estimated by comparing staining intensity of band to known quantities of stained standard proteins. The 1:10 ratio of enzyme to substrate is a general guideline. Ratios of 1:2 through 1:50 can be used without loss of enzyme efficiency or peptide recovery. An enzyme that will produce peptides greater than 10 amino acids in length should be selected. Prior amino acid analysis (UNIT 3.2) is helpful for estimating the number of cleavage sites in the protein. A second aliquot of enzyme may be added after 4 to 6 hr.
Extract the digested peptides 9. Vortex membrane sample 5 to 10 sec and sonicate 5 min at full speed in a sonication bath. Microcentrifuge sample 2 min at ∼4000 rpm (1800 × g), room temperature. Transfer supernatant to an HPLC injection vial. 10. Add another 50 µl of digestion buffer to the membrane in the 1.5-ml centrifuge tube and repeat step 9. Pool supernatant with original buffer supernatant in HPLC vial. 11. Add 100 µl of 0.1% TFA to membrane, vortex, and transfer supernatant to HPLC vial (total volume of samples should be 200 µl). Add 150 µl digestion buffer to RTX-100 blank to bring final volume to 200 µl. Most of the peptides (∼80%) are recovered in the original digestion buffer; however, these additional washes ensure maximum recovery of peptides from the membrane. Chemical Analysis
11.2.3 Current Protocols in Protein Science
12. Terminate enzymatic digestion reaction by immediately analyzing sample on HPLC or by adding 2 µl of 1% DFP solution to sample. CAUTION: DFP is a dangerous neurotoxin and must be handled with double gloves under a chemical fume hood. Please follow all precautions listed for this chemical.
Analyze samples by reversed-phase HPLC 13. Inspect pooled supernatants for small pieces of membrane or particles that might clog HPLC tubing. Remove any membrane fragments with a clean probe such as pointed tweezers, wire, or long pipet tip. Alternatively, microcentrifuge sample 2 min at 4000 rpm (1800 × g), room temperature. Transfer supernatant to a clean vial. A precolumn filter will help increase the life of HPLC columns, which frequently have problems with clogged frits.
14. Analyze samples in a microbore HPLC. Collect fractions in capless 1.5-ml microcentrifuge tubes. Cap samples and store at −20°C until ready for sequencing. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Diisopropylfluorophosphate (DFP) solution, 1% Add 10 µl DFP to 990 µl absolute ethanol in a capped microcentrifuge tube. Store at −20°C (stable at least 1 month). CAUTION: DFP is a dangerous neurotoxin. Handle with double gloves in a chemical fume hood. Please follow all precautions listed with this chemical. Digestion buffer Prepare digestion buffer in 1-ml aliquots according to recipes listed in Table 11.2.1. Store at −20°C up to 1 week. NOTE: Use only hydrogenated Triton X-100 (10% RTX-100 solution, protein grade; Calbiochem) because UV-absorbing contaminants present in ordinary Triton X-100 make identification of peptides on subsequent HPLC analysis impossible.
Enzyme solutions Trypsin: Solubilize 25 µg trypsin (sequencing grade, Boehringer Mannheim) in 25 µl of 0.01% trifluoroacetic acid (TFA) and let stand ∼10 min. Transfer 5-µl (5 µg) aliquots into clean microcentrifuge tubes, dry in a Speedvac evaporator, and store at −20°C (stable at least 1 month). Immediately before use, reconstitute enzyme in 50 µl of 0.01% TFA to obtain a 0.1 µg/µl working solution. Trypsin cleaves at arginine and lysine residues. One aliquot provides enough enzyme to digest 50 ìg of protein.
Endoproteinase Lys-C: Solubilize 3 U endoproteinase Lys-C (Boehringer Mannheim) in 200 µl Milli-Q water to obtain a 0.015 U/µl working solution. Let stand ∼10 min. Transfer 5-µl aliquots into clean tubes and store at −20°C (stable at least 1 month). Endoproteinase Lys-C cleaves only at lysine residues. One aliquot provides enough enzyme to digest ∼5 ìg protein.
Enzymatic Digestion of Proteins on PVDF Membranes
Endoproteinase Glu-C: Solubilize 50 µg endoproteinase Glu-C (sequencing grade, Boehringer Mannheim) in 50 µl of Milli-Q water and let stand ∼10 min. Transfer 5-µl aliquots (5 µg) into clean tubes, dry in a Speedvac evaporator, and store at continued
11.2.4 Current Protocols in Protein Science
−20°C (stable at least 1 month). Immediately before use, reconstitute enzyme in 50 µl Milli-Q water to obtain a 0.1 µg/µl working solution. Endoproteinase Glu-C cleaves predominately at glutamic acid, but can sometimes cleave at aspartic acid. One aliquot provides enough enzyme to digest 50 ìg of protein.
Clostripain: Solubilize 20 µg clostripain (sequencing grade, Promega) in 200 µl of manufacturer’s supplied buffer to obtain a concentration of 0.1 µg/µl. Store enzyme solution at −20°C (stable up to 1 month). Clostripain cleaves at arginine residues only. Enzyme solution provides enough enzyme to digest 50 ìg of protein.
COMMENTARY Background Information The basic protocol described in this unit offers a simple method for obtaining amino-terminal and internal amino acid sequence data for PVDF membrane–bound proteins. The procedure is highly reproducible and is compatible with subsequent peptide mapping by reversedphase HPLC (UNIT 11.6). Although proteins as large as 300 kDa have been successfully digested with this procedure, the average size of PVDF-bound proteins is ∼80 kDa. The limits of the procedure appear to depend on the HPLC used for peptide isolation and the limitations of the protein sequencer. Enzymatic digestion of a nitrocellulosebound protein for internal sequence analysis was first reported by Aebersold et al. (1987), with a more detailed procedure later reported by Tempst et al. (1990). These procedures included treatment of the nitrocellulose-bound protein with PVP-40 (polyvinylpyrrolidone, Mr of 40,000) to prevent adsorption of the selected proteinase to any remaining nonspecific protein binding sites on the membrane. After extensive washing to remove excess PVP-40, the nitrocellulose-bound sample was cut into 1 × 1–mm pieces and then digested. Aebersold et al. (1987) digested the protein overnight at 37°C with either trypsin (in a buffer of 5% acetonitrile/100 mM Tris⋅Cl, pH 8.2) or V8 protease (in 5% acetonitrile/100 mM sodium phosphate), with no additional washing of the membrane after digestion. Tempst et al. (1990) digested the protein for 15 hr at 37°C with trypsin, endoproteinase Asp-N, endoproteinase Lys-C, chymotrypsin, and subtilisin (in 5% acetonitrile/100 mM NH2HCO3), followed by one wash with fresh digestion buffer. They also reported no success with endoproteinase Glu-C. Using the above procedures to digest PVDF-bound protein yielded poor results and generally required >25 µg of protein (Bauw et al., 1989; Aebersold, 1993).
Enzymatic digestion of both PVDF- and nitrocellulose-bound protein in the presence of 1% hydrogenated Triton X-100 (RTX-100) buffers (listed in Table 11.2.1) was first performed after treating the protein band with PVP-40, as described above (Fernandez et al., 1992). The RTX-100 buffer is a strong elution reagent that allows peptide recovery from the hydrophobic, high-binding-capacity PVDF membrane as well as from nitrocellulose. Unfortunately, the RTX-100 buffer also removes PVP-40 from the membrane, which can interfere with subsequent reversed-phase HPLC. Recent reports (Fernandez et al., 1994a,b) demonstrate that treatment with PVP-40 is unnecessary when RTX-100 is used in the digestion buffer. In addition to being a strong elution reagent, RTX-100 also acts as a blocking reagent, preventing proteinase adsorption to PVDF. There are several clear advantages to using the RTX-100 digestion buffer procedure. First, it is applicable to PVDF (especially high-retention PVDF membranes such as ProBlott or Immobilon Psq). PVDF membranes are preferred over nitrocellulose because they allow higher recovery of peptides after digestion, result in fewer contaminants on subsequent HPLC analysis, and are compatible with other procedures used to analyze protein structure or amino acid analysis. The older procedures (Aebersold et al., 1987; Tempst et al., 1990) have not been successful with PVDF. In addition, recovery of peptides from nitrocellulose is also higher when RTX-100 is used in the digestion buffer (Fernandez et al., 1992). The RTX-100 procedure does not require pretreatment of the protein band with PVP-40 and can be performed in a single step. Because this protocol does not require all the washes that the PVP-40 procedures do, there is less chance of protein loss during washing. Finally, this protocol requires considerably less time than the Chemical Analysis
11.2.5 Current Protocols in Protein Science
other procedures. Overall, it is the simplest and quickest method for obtaining quantitative recovery of peptides.
Critical Parameters
Enzymatic Digestion of Proteins on PVDF Membranes
The largest source of sample loss is generally not proteinase digestion but the electroblotting of the sample. Electroblotting onto PVDF (preferably a high-binding type, such as ProBlott or Immobilon Psq) rather than nitrocellulose is recommended because peptide recovery after digestion is usually higher from PVDF (Fernandez et al., 1992; Best et al., 1994). If nitrocellulose must be used because the protein is obtained already bound to nitrocellulose prior to digestion, never allow the membrane to dry out, as this will decrease peptide yields. Always electroblot the protein using a transfer tank system because yields from semi-dry systems are lower (Mozdzanowski and Speicher, 1990). Using cleaner stains such as Ponceau S, amido black, or Coomassie brilliant blue-G with >90% dye content will increase detection of peptide fragments during reversed-phase HPLC. The key to the success of the procedure and quantitative recovery of peptides from both PVDF and nitrocellulose membranes is the use of RTX-100 in the buffer. This is desirable because non-hydrogenated Triton X-100 has several strong UV-absorbing contaminants (Fig. 11.2.1; Tiller et al., 1984). In addition, RTX-100 does not inhibit enzyme activity or interfere with peak resolution during HPLC as do ionic detergents such as SDS (Fernandez et al., 1992). Finally, the concentration of RTX100 can be decreased to 0.1% with no loss in peptide yield (Fernandez et al., 1994b). The addition of a second aliquot of enzyme after 4 to 6 hr initial digestion can improve peptide recovery (Best et al., 1994). A membrane should be cut into 1 × 1–mm pieces while keeping it wet to avoid buildup of static charge. These small pieces allow using the minimum amount of buffer to cover the membrane. The volume of digestion buffer used should be enough to cover the membrane (∼50 µl) but can be increased or decreased depending on the amount of membrane present. The enzyme solution should be selected based on any additional knowledge of the protein available, such as its amino acid composition and whether it is basic or acidic. If the protein is a complete unknown, endoproteinase Lys-C or Glu-C would be a good choice. The enzyme-to-substrate ratio should be ∼1:10; however, if the exact amount of protein is unknown, ratios of
1:2 through 1:50 are suitable for digestion and will not affect the quantitative recovery of peptides. After digestion, most of the peptides (∼80%) are recovered in the original buffer and the additional washes are performed to ensure maximum recovery. Microbore reversed-phase HPLC is the best isolation procedure for peptides.
Troubleshooting The greatest source of failure in obtaining internal sequence data is insufficient transfer of protein to the PVDF membrane, which leads to an inability to detect peptides during HPLC analysis. After staining the PVDF-bound protein, if the protein band cannot be detected by amido black staining but is observable with India ink (which is ∼10-fold more sensitive), the protein quantity may be insufficient for this procedure. Similarly, if the protein band is detectable by radioactivity or immunostaining but not by protein stain, the quantity may be insufficient for subsequent HPLC analysis. Amino acid analysis (UNIT 3.2), amino-terminal sequence analysis, or at the very least, comparison with stained standard proteins on the blot, should be performed to help determine if enough material is present. When a sufficient but small (less than 10 µg) amount of protein is available, problems may arise from misidentification of peptides on reversed-phase HPLC due to artifact peaks and contaminants. Although elimination of every contaminant is usually impossible, there are several strategic points and steps that can be taken to help reduce contamination. Simultaneous processing of a negative control (a protein-free segment excised from the PVDF membrane) will help to identify contaminants associated with the membrane and digestion buffer. The negative control must be processed through the same purification steps as the sample, including electroblotting and staining, and should be analyzed by HPLC immediately before or after the sample. A positive control (membrane-bound standard protein) is generally unnecessary but should be performed if the activity of the enzyme is in question or if a new lot number of enzyme is to be used. Major sources of contaminants include the stains used to visualize the protein on the PVDF membrane, the microcentrifuge tubes used for digestion, reagents used during digestion and extraction of peptides, and the HPLC itself. Stains are the greatest source of contaminants, and Coomassie brilliant blue in particular frequently gives problems. Amido black and Pon-
11.2.6 Current Protocols in Protein Science
A 120
Absorbance at 220 nm (mAU)
80
40
B 120
80
40
20
40
60
80
Time (min)
Figure 11.2.1 HPLC profiles of digestion buffer blanks. (A) Blank for 50 µl of 1% hydrogenated Triton X-100 (RTX-100)/10% acetonitrile/100 mM Tris⋅Cl, pH 8.0. (B) Blank for 1% Triton X-100/10% acetonitrile/100 mM Tris⋅Cl, pH 8.0. Both samples were incubated 20 hr at 37°C. Sample volumes were brought to 200 µl with 150 µl of 0.1% TFA and samples were analyzed on a Vydac C18 column (2.1 × 250–mm) using chromatographic conditions previously described (Fernandez et al., 1992). Peaks eluting at 50 to 100 min in panel B are UV-absorbing contaminants present only in Triton X-100.
ceau S are generally the cleanest, and Coomassie brilliant blue-G which has been chromatographically purified with a dye content >90% (e.g., Aldrich) appears to generate fewer contaminants than other less pure Coomassie brilliant blue stains. Surprisingly, microcentrifuge tubes can produce significant artifact peaks, which seem to vary with supplier and lot number. A blank containing only digestion buffer from a microcentrifuge tube should be included
because some contaminants only appear after incubation in the RTX-100 buffer. The major concern with the digestion buffer is the hydrogenated Triton X-100 (see Figure 11.2.1), which is purchased as a 10% stock solution. Additional late-eluting peaks may be observed with certain lots of RTX-100, whereas other lots are completely free of UV-absorbing contaminants. Milli-Q water or water prepared as described by Atherton (1989) should be used
Chemical Analysis
11.2.7 Current Protocols in Protein Science
A
Immobilon P
B
Immobilon Psq
C
ProBlott
20
Absorbance at 220 nm (mAU)
10
20
10
20
10
Time
Figure 11.2.2 Peptide maps of trypsin digestion of human transferrin bound to different types of membrane. (A) Immobilon P; (B) Immobilon Psq; and (C) ProBlott. Samples were prepared as described in the basic protocol. Four micrograms (∼53 pmol) of transferrin was analyzed by SDS-PAGE, electroblotted to PVDF, and stained with Ponceau S prior to digestion. Chromatographic conditions were as previously described (Fernandez et al., 1992).
Enzymatic Digestion of Proteins on PVDF Membranes
for all solution preparation. An HPLC blank (i.e., a gradient run with no injection) should always be performed to determine which peaks are related to the HPLC. As discussed in Critical Parameters, the concentration of RTX can be decreased without loss of peptide recovery. However, with a large amount of membrane this may not be the case. Previous procedures (Aebersold et al, 1987; Tempst et al., 1990; Bauw et al., 1989; Fernandez et al., 1992) required pretreatment of the membrane with PVP-40 to prevent any proteinase adsorption to the membrane. RTX-100 is essential for quantitative recovery of peptides from the membrane; however, RTX-100 also strips PVP-40 from the membrane, resulting in
a broad, large, UV-absorbing contaminant that can interfere with peptide identification. The PVP-40 contaminant does not depend on the age or lot number of PVP-40; making fresh solutions does not prevent the problem (Aebersold, 1993). This appears to be more prevalent with nitrocellulose and higher-binding PVDF (ProBlott and Immobilon Psq) than with lowerbinding PVDF (Immobilon P), and depends on the amount of membrane used. The PVP-40 contaminant also appears to elute earlier in the chromatogram as the HPLC column ages, becoming more of a nuisance in visualizing peptides. Therefore, using PVP-40 to prevent enzyme adsorption to the membrane should be avoided.
11.2.8 Current Protocols in Protein Science
Peptide mapping by reversed-phase HPLC is described in detail in UNIT 11.6; however, there are a few considerations that are worth discussing here. A precolumn filter (Upchurch Scientific) must be used to prevent small membrane particles from reaching the HPLC column. Inspection of the pooled supernatants for visible pieces of PVDF can prevent clogs in the microbore tubing. Membrane fragments can be removed either with a clean probe (e.g., pointed tweezers, wire or thin pipet tip) or by spinning in a centrifuge and transferring the sample to clean vial.
Anticipated Results Peptide mapping by reversed-phase HPLC after digestion of the membrane-bound protein should result in several peaks on the HPLC. Representative peptide maps from trypsin digestion of human transferrin bound to different PVDF membrane types—Immobilon P, Immobilon Psq, and ProBlott—are shown in Figure 11.2.2. Peptide maps should be reproducible when performed under the same digestion and HPLC conditions as described in this unit, as demonstrated by Figure 11.2.2. In addition, the peptide maps from proteins digested on PVDF membranes are comparable if not identical to maps derived from proteins digested in solution, indicating that the same number of peptides are recovered from the membrane as from solution. The average peptide recovery is generally 40% to 70% based on the amount of protein analyzed by SDS-PAGE, and 70% to 100% based on the amount of protein bound to PVDF (as determined by amino acid analysis). Recovery of peptides from the membrane tends to be quantitative, and the greatest loss of sample seems to occur during electroblotting.
Time Considerations
The entire procedure can be done in ∼24 hr plus the time required for peptide mapping by reversed-phase HPLC (see UNIT 11.6). Cutting the membrane takes ∼10 min, incubation after the digestion buffer is added takes 5 to 30 min, digestion at 37°C takes 22 to 24 hr, and extraction of the peptides requires ∼20 min.
Literature Cited Aebersold, R. 1993. Internal amino acid sequence analysis of proteins after in situ protease digestion on nitrocellulose. In A Practical Guide to Protein and Peptide Purification for Microsequencing, 2nd Ed. (P. Matsudaira, ed.) pp. 105154. Academic Press, New York.
Aebersold, R.H., Leavitt, J., Saavedra, R.A., Hood, L.E., and Kent, S.B. 1987. Internal amino acid sequence analysis of proteins separated by oneor two-dimensional gel electrophoresis after in situ protease digestion on nitrocellulose. Proc. Natl. Acad. Sci. U.S.A. 84:6970-6974. Atherton, D. 1989. Successful PTC amino acid analysis at the picomole level. In Techniques in Protein Chemistry (T. Hugli, ed.) pp. 273-283. Academic Press, New York. Bauw, G., Van Damme, J., Puype, M., Vandekerckhove, J., Gesser, B., Ratz, G.P., Lauridsen, J.B., and Celis, J.E. 1989. Protein-electroblotting and -microsequencing strategies in generating protein data bases from two-dimensional gels. Proc. Natl. Acad. Sci. U.S.A. 86:7701-7705. Best, S., Reim, D.F., Mozdzanowski, J., and Speicher, D.W. 1994. High sensitivity sequence analysis using in situ proteolysis on high retention PVDF membranes and a biphasic reaction column sequencer. In Techniques in Protein Chemistry V (J. Crabb, ed.) pp. 205-213. Academic Press, New York. Fernandez, J., DeMott, M., Atherton, D., and Mische, S.M. 1992. Internal protein sequence analysis: Enzymatic digestion for less than 10 µg of protein bound to polyvinylidene difluoride or nitrocellulose membranes. Anal. Biochem. 201:255-264. Fernandez, J., Andrews, L., and Mische, S.M. 1994a. An improved procedure for enzymatic digestion of polyvinylidene difluoride-bound proteins for internal sequence analysis. Anal. Biochem. 218:112-118. Fernandez, J., Andrews, L., and Mische, S.M. 1994b. A one-step enzymatic digestion procedure for PVDF-bound proteins that does not require PVP-40. In Techniques in Protein Chemistry V (J. Crabb, ed.) pp. 215-222. Academic Press, New York. Mozdzanowski, J. and Speicher, D.W. 1990. Quantitative electrotransfer of proteins from polyacrylamide gels onto PVDF membranes. In Current Research in Protein Chemistry: Techniques, Structure, and Function. (J. Villafranca, ed.) pp. 87-94. Academic Press, New York. Tempst, P., Link, A.J., Riviere, L.R., Fleming, M., and Elicone, C. 1990. Internal sequence analysis of proteins separated on polyacrylamide gels at the submicrogram level: Improved methods, applications and gene cloning strategies. Electrophoresis 11:537-553. Tiller, G.E., Mueller, T.J., Dockter, M.E., and Struve, W.G. 1984. Hydrogenation of Triton X100 eliminates its fluorescence and ultraviolet light absorbance while preserving its detergent properties. Anal. Biochem. 141:262-266.
Chemical Analysis
11.2.9 Current Protocols in Protein Science
Key References Fernandez et al., 1994a. See above. Describes digestion with and without PVP-40 and applies it to unknown proteins.
Contributed by Joseph Fernandez and Sheenah M. Mische The Rockefeller University New York, New York
Fernandez et al., 1994b. See above. Describes digestion procedure and emphasizes applicability to different types of PVDF membranes and the concentration of RTX-100 buffer.
Enzymatic Digestion of Proteins on PVDF Membranes
11.2.10 Current Protocols in Protein Science
Digestion of Proteins in Gels for Sequence Analysis
UNIT 11.3
A high percentage of eukaroytic proteins have blocked amino termini, so it is usually necessary to cleave an “unknown” protein chemically or enzymatically to obtain the partial sequences needed for cDNA cloning. Because SDS-polyacrylamide gel electrophoresis (SDS-PAGE; UNIT 10.1) is the current method of choice for the final purification of the >25 pmol amounts of protein that are usually required for internal sequencing, procedures that can be used to digest proteins in situ in SDS-polyacrylamide gels are often the most useful and have the added benefit of eliminating losses that may occur during blotting. Two alternative strategies have been developed to respond to this need and to deal with the unique problems posed by SDS, which are that SDS inhibits trypsin, one of the enzymes that is most commonly used for internal sequencing studies, and also interferes with reversed-phase HPLC. In the Basic and Alternate Protocol 2, SDS is removed from the gel prior to enzymatic cleavage by the staining and subsequent washing steps. The Basic Protocol calls for an acetonitrile wash to remove residual SDS from the protein sample. A different detergent, Tween 20, is then added back to the sample to maintain the solubility of the denatured protein. In Alternate Protocol 2, the gel slices are washed with ammonium bicarbonate and the protein samples digested in the absence of detergent. Alternate Protocol 1 utilizes lysyl endopeptidase, an enzyme resistant to SDS; the digestion can therefore be carried out without prior removal of the SDS. Ultimately, SDS is removed via an anion-exchange precolumn that immediately precedes the reversedphase HPLC column. In all cases, the peptides resulting from in-situ digestion are extracted from the gel matrix, then separated via reversed-phase HPLC prior to amino acid sequencing. Before beginning the enzymatic digests described in these protocols, it is helpful to determine the amount of protein present in the sample via amino acid analysis, as described in Support Protocol 1. Reducing and alkylating proteins separated by SDSPAGE, as described in Support Protocol 2, facilitates the identification of cysteine residues during subsequent peptide sequencing reactions. DIGESTION OF PROTEINS IN GELS IN THE PRESENCE OF TWEEN 20 In this protocol, which is a slight modification of the Rosenfeld et al. (1992) procedure, an acetonitrile wash is used to remove residual SDS and Coomassie brilliant blue from the excised gel slice containing the protein of interest. The washed gel is then partially dried prior to rehydrating in the presence of the enzyme of choice—usually trypsin or lysyl endopeptidase—in a buffer containing Tween 20. The Tween 20 presumably helps both to remove residual SDS from the protein and to maintain the solubility of the denatured protein and its resulting cleavage fragments. After digestion, the peptides are alkylated with iodoacetic acid to facilitate identification of cysteine residues during amino acid sequencing (see Support Protocol 1).
BASIC PROTOCOL
After digestion, the peptides are extracted from the gel and separated on a C18 reversedphase HPLC column. Refer to UNIT 11.6 and to Stone et al. (1990, 1991, 1993) for further discussion of HPLC mapping and peptide isolation.
Chemical Analysis Contributed by Kathryn L. Stone and Kenneth R. Williams Current Protocols in Protein Science (1995) 11.3.1-11.3.13 Copyright © 2000 by John Wiley & Sons, Inc.
11.3.1 CPPS
Materials Protein sample separated on SDS-polyacrylamide gel (UNIT 10.1; include appropriate standard protein on gel) and stained with Coomassie brilliant blue (UNIT 10.5) 50% (v/v) acetonitrile in 0.2 M ammonium carbonate, pH 8.9 0.02% (v/v) Tween 20 (Sigma) in 0.2 M ammonium carbonate, pH 8.9 0.1 mg/ml modified trypsin in manufacturer’s dilution buffer (Promega; stable at least 2 years when stored at −20°C) 0.1 mg/ml lysyl endopeptidase (Achromobacter Protease I, Wako Chemicals USA) in 2 mM Tris⋅Cl, pH 8.0 (store <2 years at −20°C) 0.2 M ammonium carbonate, pH 8.9 0.1% (v/v) trifluoroacetic acid (TFA)/60% (v/v) acetonitrile 2 M urea/0.1 M NH4HCO3 Buffer A: 0.06% (v/v) TFA Buffer B: 0.052% (v/v) TFA in 80% (v/v) acetonitrile Water-bath sonicator 0.22 µm Ultrafree filter unit (Millipore) HPLC system (Hewlett Packard 1090M or equivalent), equipped with 250-µl injection loop Vydac C18 2.1 × 250–mm, 300-Å pore size, 5-µm particle size reversed-phase HPLC column (Separations Group) or equivalent Fraction collector with peak separator (e.g., Model 2150; Isco) Prepare the sample for digestion 1. Excise protein band of interest from stained gel. Also excise two control sections of gel: one containing protein standard (positive control), and one approximately equal in size to band of interest that does not contain any protein (“blank” negative control). Estimate the total volume of gel slices: total volume (mm3) = length × width × gel thickness. Although more than one gel lane can be pooled for digestion, every effort should be made to keep the protein in as few lanes as possible and to attain the highest possible ratio of protein to gel volume. The blank control section of gel is useful for readily identifying artifact peaks in the final HPLC chromatogram that may arise from reagents, stains, or autolysis of the protease. For discussion of appropriate standard proteins for positive controls, see Critical Parameters and Troubleshooting.
2. Remove a portion of gel slice containing protein (i.e., 10% to 15%) and subject samples to amino acid analysis (see Support Protocol 1). Amino acid analysis is used to determine the amount of protein present in the sample that is to be digested. The gel slice should contain ≥25 pmol of protein and the minimum recommended protein density is 0.05 ìg protein/mm3 gel. If this analysis indicates that a sufficient amount of protein is present at a sufficiently high density, continue with step 3. Otherwise, additional protein should (if possible) be prepared and pooled with the sample.
3. Cut each gel slice (sample and control) into ∼1 × 2–mm sections. Place each set of fragments in a 1.5-ml microcentrifuge tube and add ∼150 µl of 50% acetonitrile in 0.2 M ammonium carbonate, pH 8.9.
Digestion of Proteins in Gels for Sequence Analysis
A sufficient volume should be added so that the sample can be shaken efficiently. If more than one lane of protein is being digested, an additional ∼150 ìl acetonitrile/ammonium carbonate may be added per lane and a total of about 8 lanes can fit reasonably well in a 1.5-ml microcentrifuge tube. From this step onward, process both controls in parallel with the sample containing the protein of interest.
11.3.2 Current Protocols in Protein Science
4. Place sample on a rocking table and incubate 20 min at room temperature. 5. Remove wash buffer and save in a clean 1.5-ml microcentrifuge tube. 6. Repeat wash with another 150 µl acetonitrile/ammonium carbonate. 7. Remove wash buffer and add to previous sample. Lyophilize gel pieces in a Speedvac evaporator until their volume is reduced by ∼50%. Digest the protein 8. Dilute enzyme of choice (modified trypsin or lysyl endopeptidase) to a concentration of 33 µg/ml using 0.02% Tween 20 in 0.2 M ammonium carbonate, pH 8.9. Add a volume of diluted enzyme approximately equal to original gel volume determined in step 1. Assume that 1 mm3 = 1 ìl.
9. If gel pieces are not completely immersed, add 0.02% Tween 20 in 0.2 M ammonium carbonate until pieces are covered with buffer. 10. Incubate samples 24 hr at 37°C. Alkylate the digested sample 11. Add 50 µl of 0.2 M ammonium carbonate (pH 8.9) to each sample. If necessary, add additional 0.2 M ammonium carbonate until pieces are completely covered with buffer. This may be necessary because the gel swells during the 37°C incubation in step 10.
12. Reduce and alkylate sample (see Support Protocol 2). Extract the peptides from the gel 13. Add 100 µl of 0.1% TFA/60% acetonitrile. If necessary, add additional TFA/acetonitrile until pieces are completely covered with buffer. Incubate 30 to 40 min on rocking table at room temperature. If a rocking table is not available, vortex the sample intermittently.
14. Sonicate 5 min in a water-bath sonicator. 15. Remove wash buffer (containing the extracted peptides) to a clean 1.5-ml microcentrifuge tube. Repeat extraction steps 13 and 14 with fresh TFA/acetonitrile. 16. Remove wash buffer and combine with buffer from first TFA/acetonitrile wash. Discard gel pieces. Add 100 µl of 2 M urea/0.1 M NH4HCO3 to combined wash buffer samples. 17. Lyophilize sample in a Speedvac evaporator until final volume is <100 µl. This will ensure sufficient removal of the CH3CN so that the resulting peptides can be directly loaded on a reversed-phase HPLC column.
18. Dilute sample to 110 µl with HPLC-grade water. 19. Transfer sample to a 0.22-µm filter unit and microcentrifuge for 5 min at maximum speed. 20. Equilibrate Vydac C18 reversed-phase HPLC column at 0.15 ml/min in 98% buffer A/2% buffer B. Inject the filtrate from step 19 (containing eluted peptides) onto column. A Vydac C18 (2.1 × 250–mm, 300-Å pore size, 5-ìm particle size) or comparable column
Chemical Analysis
11.3.3 Current Protocols in Protein Science
is recommended for isolation of 25 to 250 pmol of digested peptides. Larger amounts may be separated on 4.6-mm columns (Stone et al., 1991).
21. Elute with buffer A/buffer B mixture according to the following gradient: 0 to 60 min 2% to 37% buffer B 60 to 90 min 37% to 75% buffer B 90 to 105 min 75% to 98% buffer B. 22. Collect eluate in 1.5-ml capless microcentrifuge tubes placed in the top of 13 × 100–mm test tubes. Cap all sample tubes immediately following the completion of the collection run to prevent evaporation of the acetonitrile. Store eluted peptides at 5° to 10°C. ALTERNATE PROTOCOL 1
DIGESTION OF PROTEINS IN GELS IN THE PRESENCE OF SODIUM DODECYL SULFATE In this protocol, the gel slice containing the protein of interest is presoaked in a sodium dodecyl sulfate (SDS)/Tris buffer at pH 9.0. The slice is then digested in situ with lysyl endopeptidase (Achromobacter protease I) in the presence of SDS. After digestion, the gel pieces are crushed with a nylon mesh and the peptides extracted using an SDS/Tris buffer. The SDS is removed from the sample prior to reversed-phase separation of the peptides by a DEAE (or equivalent anion exchange) HPLC precolumn. The following protocol is a slight modification of the original procedure described by Kawasaki et al. (1990). In this modified protocol, the amount of protein present in the samples is determined by amino acid analysis and there is an additional carboxymethylation step at the end of the procedure. Additional Materials (also see Basic Protocol) 0.1% (w/v) SDS in 100 mM Tris⋅Cl, pH 9.0 Nylon mesh (109 mesh) DEAE HPLC precolumn or equivalent Prepare the sample for digestion 1. Soak the stained and destained gel 1 hr in H2O. This will reduce the concentrations of acetic acid and methanol in the gel.
2. Excise the protein band of interest from the gel, along with sections for positive and negative controls, and estimate protein concentration in protein band sample as in steps 1 and 2 of the Basic Protocol. 3. Cut each gel slice into ∼1 × 2–mm pieces. Add ∼100 µl of 0.1% SDS in 100 mM Tris⋅Cl (pH 9.0) to each sample. Preincubate sample 1 hr at 37°C. Try to keep volume of SDS to a minimum. Add only enough to cover gel pieces. From this step onward, process controls in parallel with the sample containing the protein of interest.
Digest the protein 4. Add a volume of lysyl endopeptidase that yields a 1:10 (w/w) ratio of enzyme to sample protein. Incubate 24 hr at 37°C.
Digestion of Proteins in Gels for Sequence Analysis
Dilutions of the 1 mg/ml lysyl endopeptidase stock should be made in 2 mM Tris⋅Cl, pH 8.0. According to the original protocol (Kawasaki et al., 1990), the final enzyme concentration must be >2 ìg/ml. Therefore, if a 1:10 (w/w) ratio yields an enzyme
11.3.4 Current Protocols in Protein Science
concentration below this level, increase the ratio until this minimum concentration is reached.
Extract the peptides from the gel 5. Transfer supernatant to a clean 1.5-ml microcentrifuge tube. 6. Crush the gel pieces through a nylon mesh as follows: Cut a hole in the bottom of the filter unit section of a 0.22-µm filter. Replace filter with a 2-in. (5-cm) square of nylon mesh and place filter in a 1.5-ml microcentrifuge tube. Place gel pieces on top of nylon mesh. Microcentrifuge tube 10 to 15 min at maximum speed. 7. Remove filter from microcentrifuge tube. Add 500 µl of 0.1% SDS in 100 mM Tris⋅Cl (pH 9.0) to crushed gel pieces at bottom of tube and incubate with shaking 1 hr at room temperature. Shake sample on rocking table or vortex intermittently.
8. Transfer supernatant to a 0.22-µm filter unit and microcentrifuge 10 to 15 min at maximum speed. 9. Combine supernatant with sample from step 5. 10. Lyophilize sample in a Speedvac evaporator until volume is <200 µl. 11. Dilute sample to 210 µl with water. Alkylate the digested protein 12. Reduce and alkylate the protein (see Support Protocol 2). 13. Separate alkylated sample on 2.1 × 250–mm Vydac C18 reversed-phase HPLC column that is equipped with a DEAE precolumn as in steps 20 to 22 of the Basic Protocol. DIGESTION OF PROTEINS IN GELS IN THE ABSENCE OF DETERGENT
ALTERNATE PROTOCOL 2
In this protocol, enzyme digestion is carried out in the absence of detergent. The protein is separated by SDS-PAGE and washed extensively with 0.2 M ammonium bicarbonate. After washing, the protein is reduced, alkylated, and digested with either modified trypsin or lysyl endopeptidase. The resulting peptides are then eluted from the gel using passive elution in 2 M urea/0.1 M ammonium bicarbonate prior to separation by reversed-phase HPLC. Additional Materials (also see Basic Protocol) 0.2 M ammonium bicarbonate 2 M urea/0.1 M ammonium bicarbonate Prepare the sample for digestion 1. Excise the protein of interest from the gel, along with sections for positive and negative controls, and determine the protein concentration as in steps 1 and 2 of the Basic Protocol. 2. Cut each gel slice into 1 × 2–mm pieces and add 500 µl of 0.2 M ammonium bicarbonate to each sample. Gel pieces should be totally covered with 0.2 M ammonium bicarbonate to enable efficient shaking. If gel pieces are not totally immersed, add extra ammonium bicarbonate to completely cover sample.
Chemical Analysis
11.3.5 Current Protocols in Protein Science
From this step onward, process controls in parallel with the sample containing the protein of interest.
3. Incubate sample on rocking table 30 min to 1 hr at room temperature. If rocking table is not available, vortex samples intermittently.
4. Transfer wash buffer to clean 1.5-ml microcentrifuge tubes and measure pH. 5. If pH is acidic, add an additional 500 µl of 0.2 M ammonium bicarbonate to gel pieces and incubate 2 hr at room temperature. Check the pH again after 2 hr. If pH of wash buffer is still acidic, repeat wash again; otherwise proceed to step 6. 6. Add sufficient volume of 0.2 M ammonium bicarbonate to gel pieces to completely immerse sample. Alkylate the protein 7. Reduce and alkylate protein sample (see Support Protocol 2). Digest the protein 8. Add an appropriate volume of enzyme to the alkylated sample to obtain a final enzyme concentration of 6 µg/ml for modified trypsin or 12 µg/ml for lysyl endopeptidase. Incubate 24 hr at 37°C. 9. Add 500 µl of 2 M urea/0.1 M ammonium bicarbonate to each sample. Incubate on a rocking table 6 hr at room temperature to extract peptides. If a rocking table is unavailable, vortex samples intermittently.
10. Transfer supernatant to a clean 1.5-ml microcentrifuge tube and freeze. 11. Add 500 µl of 0.2 M ammonium bicarbonate to gel pieces. Incubate on rocking table overnight at room temperature. 12. Combine this extract with supernatant from step 10. Lyophilize sample in a Speedvac evaporator until the volume is <200 µl. 13. Adjust volume to 210 µl with water. 14. Transfer supernatant to a 0.22-µm filter unit and microcentrifuge 10 min at maximum speed. 15. Subject the filtrate containing eluted peptides to HPLC as in steps 20 to 22 of the Basic Protocol. SUPPORT PROTOCOL 1
AMINO ACID ANALYSIS OF PROTEINS SEPARATED BY SDS-PAGE Prior to beginning the enzymatic digests described in the basic and alternate protocols, an amino acid analysis should be carried out on 10% to 15% of the gel slice to accurately determine the amount of protein that is present. Materials Gel slice (Basic Protocol, Alternate Protocol 1, or Alternate Protocol 2) 0.2% (v/v) phenol in 6 N HCl, containing 2 nmol/100 µl norleucine internal standard 10 × 75-mm Pyrex test tube Amino acid analyzer (Beckman 7300 or equivalent)
Digestion of Proteins in Gels for Sequence Analysis
1. Add 100 µl of 0.2% phenol in 6 N HCl (with 2 nmol/100 µl norleucine) to gel slice. Hydrolyze sample 16 to 24 hr at 115°C.
11.3.6 Current Protocols in Protein Science
2. After hydrolysis, transfer supernatant to a 10 × 75-mm Pyrex test tube and lyophilize to dryness using a Speedvac evaporator. 3. Dissolve hydrolysate in buffer recommended for the amino acid analyzer. Load sample onto analyzer. 4. Determine protein concentration as described in UNIT 3.2. Continue with digestion as described in the Basic Protocol, Alternate Protocol 1, or Alternate Protocol 2. REDUCTION AND ALKYLATION OF PROTEINS The major benefit of reducing and alkylating proteins separated by SDS-PAGE is that it facilitates the subsequent identification of cysteine residues via Edman degradation. Reduction and alkylation convert cysteine to a more stable phenylthiohydantoin (PTH) derivative.
SUPPORT PROTOCOL 2
Materials Gel pieces in buffer (basic or alternate protocols) 45 mM dithiothreitol (DTT) 100 mM iodoacetic acid (IAA) 50°C incubator 1. Estimate the total volume of sample (gel plus buffer). Sample volumes can be most easily estimated by comparing to a “standard” set of microcentrifuge tubes containing known volumes of water.
2. Calculate volume of DTT (“x”) needed to yield a final DTT concentration of 1 mM: (x) × (0.045 M stock) = (total volume) × (0.001 M). 3. Add DTT and incubate sample 20 min at 50°C. 4. Remove sample from incubator and let cool to room temperature. Add a volume of 100 mM IAA equal to the volume (x) of DTT added in step 3. 5. Incubate sample 20 min at room temperature in the dark, to prevent formation of iodine which could react with tyrosine residues. 6. Continue with protein digestion (see Basic Protocol, step 13; Alternate Protocol 1, step 13; or Alternate Protocol 2, step 8). COMMENTARY Background Information Because a high percentage of eukaryotic proteins have blocked amino termini (Brown and Roberts, 1976; Dreissen et al., 1985), direct amino terminal sequencing often ends in failure. In fact, when the amount of a eukaryotic protein is extremely limiting (i.e., <50 pmol), it is usually far better to assume the amino terminus is blocked and proceed directly with enzymatic or chemical cleavage. Because SDSPAGE is often the last step in protein purification, cleavage protocols that can be directly carried out on proteins in SDS polyacrylamide gels are often the most useful, as they avoid the losses that may occur during elution of the
proteins from the gels or during electroblotting of proteins onto membranes. Trypsin and lysyl endopeptidase are often the preferred enzymes for cleaving proteins in SDS-polyacrylamide gels. These enzymes have high specificity and produce peptides of a size that separate optimally via reversedphase HPLC (typically 5 to 25 residues). Based on the average occurrence of lysine (5.7%) and arginine (5.4%) in proteins listed in the Protein Identification Resource Database, digestion with either of these enzymes would be expected to produce peptides with an average length of 17 residues (for lysyl endopeptidase) or ∼9 residues (for trypsin).
Chemical Analysis
11.3.7 Current Protocols in Protein Science
Enzymatic digestions of proteins in gels have been carried out both in the absence (Stone and Williams, 1993; Ward et al., 1990) and the presence of detergents such as SDS (Kawasaki et al., 1990) or Tween 20 (Rosenfeld et al., 1992). In all instances, the protease is allowed to diffuse into the gel and the resulting peptides are subsequently obtained by diffusion out of the gel. The Basic Protocol is a slightly modified version of the Rosenfeld et al. (1992) procedure in which the SDS is removed from the gel slice (prior to carrying out the digest) by two quick washes with 50% acetonitrile/0.2 M ammonium carbonate (pH 8.9). After removal of the SDS, the gel slices are dried to half their original volume and then partially rehydrated in a 200 mM ammonium carbonate (pH 8.9) buffer containing Tween 20 and the enzyme. It is possible to digest proteins in the presence of SDS by using lysyl endopeptidase, an enzyme that is active in the presence of SDS, and by using an anion-exchange precolumn to remove the SDS immediately prior to reversedphase HPLC (Kawasaki and Suzuki, 1990). This modified version of a procedure described by Kawasaki et al. (1990) is presented in Alternate Protocol 1. In this procedure, the SDS binds to the precolumn and the peptides pass through to the reversed-phase HPLC column. Although the precolumn is regenerated by the high concentration of acetonitrile present at the end of the gradient, in practice, the precolumn should be replaced at least every five runs or after a total of 1 mg SDS has been applied to the column (Williams et al., 1993). There is currently a lack of definitive data regarding which of the three procedures described in this unit provides the highest yield of peptides from a given amount of protein separated on an SDS-polyacrylamide gel (Williams et al., 1993). The procedure described in the Basic Protocol is preferred because it is the simplest and can be used for a wide range of proteases, including trypsin, lysyl endopeptidase, chymotrypsin, and endoproteinase GluC. It was observed that one sample that was resistant to digestion in the absence of a detergent, as described in Alternate Protocol 2, was successfully digested using the procedure described in the Basic Protocol, perhaps due to the presence of Tween 20 (K.L.S. and K.R.W., unpub. observ.). Digestion of Proteins in Gels for Sequence Analysis
Critical Parameters and Troubleshooting Two of the most critical questions that need
to be addressed prior to digesting a protein in a polyacrylamide gel are whether or not the protein is pure and whether or not the Coomassie blue–stained band that has been selected is actually responsible for the activity of interest. The first question can often be addressed by subjecting a small aliquot of the protein to two-dimensional gel electrophoresis (UNIT 10.4), where the first direction is an isoelectric focusing step. Keep in mind, however, that although the appearance of a single Coomassie blue– stained spot following two-dimensional electrophoresis provides good evidence for homogeneity, microheterogeneity with respect to the isoelectric point may result from post-translational modifications, such as phosphorylation, which would have little impact on internal amino acid sequence. To ensure that the protein band selected is responsible for the activity of interest, aliquots of fractions eluted from the last chromatography step can be subjected to another round of SDS-PAGE. It should be possible to directly correlate the protein’s activity with the intensity of the candidate band on SDS-PAGE and to thus confirm the identity and purity of this band. There are several other approaches that may be taken to ensure that the appropriate band has been selected for internal amino acid sequencing. Gel filtration can be used to independently verify the approximate molecular weight of the protein of interest. A protein separated by SDSPAGE often can be renatured to confirm its activity. In the case of nucleic acid binding proteins, the protein preparation can be photocrosslinked to a 32P-labeled nucleic acid target site. When the nucleic acid is degraded with a nuclease, if the candidate band is the correct DNA-binding protein, it should be labeled with 32P (Safer et al., 1988). Once the identity of the Coomassie blue– stained protein band has been confirmed, it is important to determine whether a sufficient amount of protein is present for the experiment to proceed. Investigators who rely on relative Coomassie blue staining and colorimetric assays often overestimate the amount of protein present (sometimes by as much as 5- to 10fold). To obtain an accurate estimate of protein concentration, an aliquot of the Coomassie blue–stained gel band should be subjected to hydrolysis and ion-exchange amino acid analysis as described in Support Protocol 1. In the case of 50 pmol of a 50 kDa protein, 15% of the band (∼0.4 µg of protein) would be a rea-
11.3.8 Current Protocols in Protein Science
sonable fraction of the sample to commit to this step. Although glycine, cysteine, tryptophan, and sometimes histidine and arginine cannot be quantified by this analysis, it nonetheless provides a more accurate estimate of the amount of protein than does Coomassie blue staining. The latter approach can be affected by how well different proteins stain with Coomassie brilliant blue. Because the “background” obtained from hydrolyzing and analyzing blank sections of SDS-polyacrylamide gels averages ∼0.09 µg (Williams et al., 1993), an aliquot of a control section of gel that appears not to contain any protein should also be hydrolyzed along with the slices containing the protein of interest. The blank values should be subtracted from those obtained for the unknown protein sample. Although the Basic Protocol has been used successfully on as little as 16 pmol of transferrin and 26 pmol of an unknown protein, a minimum of 75 pmol (as determined by hydrolysis/amino acid analysis) should be used both to ensure a high probability of success and to improve the quality and length of the resulting sequence data. A final parameter to consider is the “density” (i.e., in µg/mm3) of the protein in the gel band. All five of the unsuccessful digests listed in Table 11.3.1 contained proteins at densities ≤0.05 µg/mm3 (with a mean density for these five samples of 0.03 µg/mm3). Because the group of unsuccessful samples included two samples containing 101 to 200 pmol of protein and two samples with >200 pmol of protein, it appears that, within reasonable limits, the density of the protein in the gel band is a more important determinant of success than is the absolute amount of protein. Although samples with densities as low as 0.02 have been digested successfully, to ensure success, samples should have a minimum density of ≥0.05 µg/mm3. Obviously, everything possible should be done to ensure that the sample is loaded in one or two lanes as opposed to three or more. Typically, gels that are ≤0.75 mm in thickness should be used and a minimum of 1 µg of the protein of interest should be included in each lane. If a minimum of 25 pmol protein (at a density > 0.05 µg/mm3) is digested, the success rate should be well above 90% (see Table 11.3.1). The laboratory carrying out the work should be accustomed to routinely isolating <10 pmol amounts of peptides and to sequencing in the several hundred femtomole range.
Because human transferrin (Mr = 75,000) digests as well or better using the Basic Protocol than does any other “standard” protein, it serves as an excellent positive control for proteins in the >50 kDa range. For proteins <50 kDa, carbonic anhydrase (Mr = 29,000) provides a reliable positive control. The positive control is just as important as the negative control (i.e., the blank section of gel that does not contain any protein). Hence, positive and negative controls should be performed along with all samples subjected to enzymatic digestion in a gel. In other words, a similar amount of transferrin should be run on the same gel as the sample and should be digested, carboxymethylated, and subjected to analytical HPLC along with the sample. Obviously, if neither the transferrin nor the sample digests, the fault does not lie with the sample. If the above criteria have been met and there is no trivial reason (e.g., failure to add the enzyme solution to the sample) for the failure of a sample to digest, then a lack of digestion may reflect the intrinsic resistance of the sample to enzymatic cleavage. A common reason for this is extensive glycosylation. Although <10% (by weight) carbohydrate does not seem to adversely affect cleavage (e.g., of transferrin), high levels of glycosylation may well prevent or severely limit cleavage. If the extent of glycosylation is significantly above 10%, the carbohydrate should be enzymatically removed immediately prior to SDS-PAGE. Finally, even after complete deglycosylation, a few proteins, such as the major subunit of the Duffy blood group system (Chaudhuri et al., 1993), may be totally resistant to digestion in gels by trypsin, lysyl endopeptidase, or chymotrypsin (with the latter enzyme so far uniformly failing to cleave any protein that was resistant to trypsin). In this instance, cyanogen bromide (CNBr) can be used to chemically digest the protein sample (see UNIT 11.4). Assuming that SDS-PAGE is a necessary final purification step, CNBr cleavage has to be carried out in the gel (Jahnen et al., 1990). Because CNBr-generated peptides do not usually separate well by reversed-phase HPLC, the resulting peptides should be eluted from the gel and then separated via subsequent SDSPAGE. The separated peptides can then be electroblotted onto a high-capacity PVDF membrane (UNIT 10.7), stained (UNIT 10.8), and sequenced. Alternatively, CNBr cleavage could be carried out prior to the Basic Protocol. Of course, prior CNBr cleavage will make at least Chemical Analysis
11.3.9 Current Protocols in Protein Science
B
A210
A
40.00
60.00
80.00
Time (min)
Figure 11.3.1 (A)HPLC profile of tryptic digestion of 42 pmol of transferrin. Protein was separated on a 12.5% SDS-polyacrylamide gel and digested as described in the Basic Protocol. Following trypsin digestion, the resulting peptides were fractionated on a Vydac C18 column via reversedphase HPLC as described in the Basic Protocol. The Y-axis is absorbance at 210 nm (full scale = 0.02) and the X-axis is in terms of minutes. (B) Analytical HPLC profile for a control “digest” that was carried out on a section of gel that appeared not to contain any protein.
some regions of the protein susceptible to further cleavage via proteolysis. It is important to point out that, based on several hundred proteins subjected to analysis in the Keck Facility over the last few years, <2% of proteins fall into this intractable category (K.L.S. and K.R.W., unpub. observ.). Hence, assuming that the protein is not extensively glycosylated, resistance of the protein to digestion should be one of the last rather than one of the first reasons advanced to account for a failed digest.
Anticipated Results
Digestion of Proteins in Gels for Sequence Analysis
Figure 11.3.1 shows the HPLC profile from a tryptic digestion (following the Basic Protocol) of 42 pmol transferrin that was separated in a 12.5% SDS-polyacrylamide gel at a protein/gel-volume density of 0.19 µg/mm3. By comparing the transferrin sample (A) to the “blank” chromatogram (B), it is possible to quickly identify most of the artifact absorbance peaks that arise from trypsin autolysis products, reagents, and Coomassie blue (which typically
gives rise to one or more peaks eluting after ∼80 min). Table 11.3.1 summarizes the results for 53 proteins digested according to the Basic Protocol or Alternate Protocol 2 (in the absence of detergent). One of the first conclusions that can be drawn from this data is that the overall success rate of these two procedures (even when some of the samples do not meet the criteria given above) appears to be at least 90%. Another conclusion is that the major benefit of going from the <50 pmol to the >200 pmol level is a progressive increase in the percent yield of an average peptide (i.e., from <6% to >14%), which directly translates into a larger number of positively identified residues for each peptide sequenced (i.e., from <9 to >14). It should be noted that because the percent initial yields (i.e., overall recovery of peptide from a given amount of protein that is digested) are based on the initial sequencing yield, which is typically equal to only ∼50% to 75% of the actual amount of peptide loaded, the actual recovery of tryptic peptides is probably from
11.3.10 Current Protocols in Protein Science
Table 11.3.1
Effect of Sample Amount on Digestion of Proteins in Gelsa
Amount of protein analyzed Parameter
<50 pmol Range
Number of proteins Mass of protein (kDa) Amount of protein digested (pmol) Density of protein bank (µg/mm3) % Initial yieldb Results of Sequencing # of Peaks sequenced per protein ≥4 Residues called (%) Mixture (%) No sequence obtained (%) # of Residues called per peptide sequenced Overall success ratec (%)
51-100 pmol
Mean or number
Range
Mean or number
>200 pmol
101-200 pmol Range
Mean or number
Range
Mean or number
15-106 26-49
5 69 39
40-120 54-100
8 72 80
18-285 115-181
20 71 139
17-285 204-850
20 58 364
0.03-0.12
0.07
0.02-0.09
0.05
0.02-0.31
0.09
0.05-0.66
0.21
2-11.1
5.7
1-37.6
10.6
0.6-68.9
11.7
0.005-50.5
14.4
1.4
2.0
2.1
2.5
88 12 0
76 0 24
76 12 12
80 2 18
8.71
10.6
12.0
14.2
100
88
90
90
aRepresents the analysis of 53 recent in-gel digests carried out in the W.M. Keck Facility at Yale University. bCalculated as the yield of Pth-amino acid sequencing at the first or second cycle compared to the amount of protein that was digested as determined by hydrolysis and amino acid analysis of a 10% to 15% aliquot of the gel slice. Because initial yields of purified peptides directly applied onto the sequencer are usually about 50%, the actual % recovery of peptides from the above digests is probably twice the % initial yields given above. cAs judged by the ability to positively identify the sequence of at least 15 residues in one or two peptides or to obtain sufficient sequence to positively
identify the protein via database searches. 34 of the above 53 proteins (64%) were identified based on these database searches.
1.3- to 2-fold above that given in Table 11.3.1 (i.e., it probably ranges from ∼12% greater recovery at the <50 pmol level to >28% at the >200 pmol level). These average recoveries are particularly useful for judging whether the peptide sequenced was derived from the major component in the gel band that was digested. For instance, if the initial yield of a peptide from a digest of 350 pmol of protein in a gel is only 1%, it raises the possibility this peptide may have derived from a 10% contaminant. Another conclusion from Table 11.3.1 is that, on average, >80% of peptides selected for sequencing provide usable sequences, as assessed by their symmetrical absorbance profile and laser desorption mass spectra (LDMS). The remaining 20% of samples either fail to sequence, contain mixtures of peptides, or turn out to be derived from autolysis of the protease. Occasionally, an autolysis product observed in the sample will not be present in the blank.
Without the benefit of LDMS, the fraction of symmetrical HPLC peptide peaks that provide usable sequences decreases to ∼67% (Stone et al., 1992). When the criteria for selecting candidate peaks for sequencing are increased to include both the appearance of a symmetrical HPLC peak and a major/minor mass spectrometric peak ratio of at least 10 as judged by LDMS, the probability of obtaining useful data from sequencing any given HPLC absorbance peak increases to 90% (based on the last 200 peptides sequenced in the Keck Facility at Yale University; K.L.S. and K.R.W., unpub. observ.). Obviously, because the sensitivity of LDMS is such that spectra can be routinely obtained on 25 to 500 fmol amounts of most tryptic peptides, this technique provides an extremely valuable adjunct to internal sequencing. In addition to providing an independent measure of peak purity, LDMS readily identifies Coomas-
Chemical Analysis
11.3.11 Current Protocols in Protein Science
sie blue, protease autolysis products, and other reagent peaks. In addition, the resulting peptide mass can be used to confirm the results of Edman degradation as well as to detect and often identify post-translational modifications such as phosphorylation. Finally, the high percentage of proteins that are identified based on database searching of internal peptide sequences (64% based on the 53 proteins listed in Table 11.3.1) points out the absolute necessity of these searches. Unfortunately, in many instances these searches demonstrate that the major component in the Coomassie blue–stained band sequenced is not the protein of interest. Some of the proteins that have recently been identified via these searches (and which were obviously not responsible for the activity being purified) include: nucleolin, lactotransferrin, keratin, human serum albumin, vinculin, ubiquitin, collagen, alcohol dehydrogenase, tubulin, and—particularly when the purification involves immunoaffinity chromatography—immunoglobulins (K.L.S. and K.R.W., unpub. observ.). Because as few as 5 to 6 residues of sequence are often sufficient to identify a protein via a database search, the search can sometimes be performed while the peptide is actually being sequenced. In this way, sequencing can be terminated if the peptide is found to derive from a known protein.
Time Considerations Enzymatic digestions of proteins in gels carried out in the presence of Tween 20 or SDS can be completed in 24 hr. Alternate Protocol 2 (carried out in the absence of detergent) requires 3 to 4 days to complete. The amino acid analysis will delay the start of the digestion for 2 to 3 days, but it is well worth the extra time, because proceeding with 50 pmol instead of 10 pmol may well mean the difference between success and failure. Isolating the peptide fragments via reversed-phase HPLC requires only a few hours and subsequent peptide sequencing requires 1 to 2 days.
Literature Cited Brown, J.L. and Roberts, W.K. 1976. Evidence that approximately eighty percent of the soluble proteins from Ehrlich Ascites Cells are amino terminally acetylated. J. Biol. Chem. 251:1009-1014.
Digestion of Proteins in Gels for Sequence Analysis
Chaudhuri, A., Polyakova, J., Zbrzezna, V., Williams, K.R., Gulati, S., and Pogo, O. 1993. Cloning of glycoprotein D cDNA, which encodes the major subunit of the Duffy blood group system and the receptor for the Plasmodium vivax malaria parasite. Proc. Natl. Acad. Sci. U.S.A. 90:10,793-10,797.
Driessen, H.P., De Jong, W.W., Tesser, G.I., and Bloemendal, H. 1985. The mechanism of N-terminal acetylation of proteins. In Critical Reviews in Biochemistry (G.D. Fasman, ed.) Vol. 18, pp. 281-325. CRC Press, Boca Raton, Fla. Jahnen, W., Ward, L.D., Reid, G.E., Moritz, R.L., and Simpson, R.J. 1990. Internal amino acid sequencing of proteins by in situ cyanogen bromide cleavage in polyacrylamide gels. Biochem. Biophys. Res. Commun. 166:139-145. Kawasaki, H. and Suzuki, K. 1990. Separation of peptides dissolved in a sodium dodecyl sulfate solution by reversed-phase liquid chromatography: Removal of SDS from peptides using an ion-exchange precolumn. Anal. Biochem. 186:264-268. Kawasaki, H., Emori, Y., and Suzuki, K. 1990. Production and separation of peptides from proteins stained with Coomassie brilliant blue R-250 after separation by SDS polyacrylamide gel electrophoresis. Anal. Biochem. 191:332-336. Laemmli, U. 1970. Cleavage of structural proteins during the assembly of bacteriophage T4. Nature 227:680-685. Rosenfeld, J., Capdeville, J., Guillemot, J., and Ferrara, P. 1992. In-gel digestion of proteins for internal sequence analysis after 1 or 2 dimensional gel electrophoresis. Anal. Biochem. 203:173-179. Safer, B., Cohen, R.B., Garfinkel, S., and Thompson, J.A. 1988. DNA affinity labeling of adenovirus type 2 upstream promoter sequencebinding factors identifies two distinct proteins. Mol. Cell. Biol. 8:105-113. Stone, K.L., LoPresti, M.B., Williams, N.D., Crawford, J.M., DeAngelis, R., and Williams, K.R. 1989. Enzymatic digestion of proteins and HPLC peptide isolation in the sub-nanomole range. In Techniques in Protein Chemistry (T.E. Hugli, ed.) pp. 377-391. Academic Press, San Diego. Stone, K.L., Elliott, J.I., Peterson, G., McMurray, W., and Williams, K.R. 1990. Reversed-phase high performance liquid chromatography for fractionation of enzymatic digests and chemical cleavage products of proteins. Methods Enzymol. 193:389-412. Stone, K.L., LoPresti, M.B., Crawford, J.M., DeAngelis, R., and Williams, K.R. 1991. Reversedphase HPLC separation of sub-nanomole amounts of peptides obtained from enzymatic digests. In High-Performance Liquid Chromatography of Peptides and Proteins: Separation, Analysis, and Conformation. (C. Mant and R. Hodges, eds.) pp. 669-677. CRC Press, Boca Raton, Fla. Stone, K.L., McNulty, D.E., LoPresti, M.L., Crawford, J.M., DeAngelis, R., and Williams, K.R. 1992. Elution and internal sequencing of PVDFblotted proteins. In Techniques in Protein Chemistry III (R. Angeletti, ed.) pp. 22-34. Academic Press, San Diego.
11.3.12 Current Protocols in Protein Science
Stone, K.L. and Williams, K.R. 1993. Enzymatic digestion of proteins and HPLC peptide isolation. In A Practical Guide to Protein and Peptide Purification for Microsequencing, 2nd ed. (P. Matsudaira, ed.) pp. 43-69. Academic Press, San Diego. Ward, L.D., Reid, G.E., Moritz, R., and Simpson, R.J. 1990. Peptide mapping and internal sequencing of proteins from acrylamide gels. In Current Research in Protein Chemistry: Techniques, Structure, and Function (J.J. Villafranca, ed.) pp. 179-190. Academic Press, San Diego.
Williams, K.R., Kobayashi, R., Lane, W., and Tempst, P. 1993. Internal amino acid sequencing: Observations from four different laboratories. ABRF News 4:7-12.
Contributed by Kathryn L. Stone and Kenneth R. Williams Yale University New Haven, Connecticut
Chemical Analysis
11.3.13 Current Protocols in Protein Science
Chemical Cleavage of Proteins in Solution
UNIT 11.4
Detailed structural analysis of proteins frequently involves digestion of samples (enzymatically or chemically) into smaller fragments. Enzymatic digestion (UNIT 11.1) generally produces a mixture of a large number of peptides, which require additional purification (e.g., by HPLC) before they can be analyzed further. Purification becomes more challenging when certain proteases, such as trypsin and Staphylococcus aureus V8, are used, owing to the high proportion of these recognition site residues in proteins. Chemical cleavage, in contrast to enzymatic cleavage, usually targets residues and specific dipeptide linkages that occur at low frequencies in proteins. Thus, on average, fewer fragments will be generated and these fragments will be larger in size than those produced by standard enzymatic treatment. Consequently, in addition to reversed-phase, ion-exchange (UNIT 8.2), and size-exclusion (UNIT 8.3) chromatographies, analytical highresolution electrophoresis (UNIT 10.1) can be used to purify resulting protein fragments. In this unit, the five basic protocols present widely used reactions that are specific and efficient for the chemical cleavage of proteins in solution. Cyanogen bromide (CNBr; Basic Protocol 1) cleaves at methionine (Met) residues; 2-(2′-nitrophenylsulfonyl)-3methyl-3-bromoindolenine (BNPS-skatole; Basic Protocol 2) cleaves at tryptophan (Trp) residues; formic acid (Basic Protocol 3) cleaves at aspartic acid–proline (Asp-Pro) peptide bonds; hydroxylamine (Basic Protocol 4) cleaves at asparagine-glycine (Asn-Gly) peptide bonds, and 2-nitro-5-thiocyanobenzoic acid (NTCB; see Basic Protocol 5) cleaves at cysteine (Cys) residues. Because the above loci are at relatively low abundance in most proteins, digestion with these agents will yield relatively long peptides. In general, reactions are chosen based on a priori knowledge of a protein’s target residues. If no information is available, perform the reactions in the order listed. Solid-phase digestions can be performed using similar chemical reactions to cleave proteins attached to membranes (UNIT 11.5). CAUTION: Most chemicals used in the following protocols are toxic. Therefore, it is imperative that good laboratory practice be followed. All procedures and incubations should be performed in a well-ventilated chemical fume hood and laboratory personnel should wear appropriate protective laboratory clothing, including safety glasses and gloves. All chemical waste should be properly disposed of according to local regulations. CLEAVAGE AT THE C-TERMINUS OF MET RESIDUES WITH CNBr Cyanogen bromide (CNBr) cleaves proteins on the C-terminal side of unoxidized methionine (Met) residues. For proteins with multiple Met residues, fragments generated from this reaction can be incomplete (i.e., contain an internal unmodified Met or homoserine). After digestion with CNBr, resulting peptide fragments are fractionated by HPLC or electrophoresis prior to further characterization. This protocol also provides instructions for neutralizing CNBr, which is extremely toxic, after use.
BASIC PROTOCOL 1
Materials Lyophilized protein sample in 1.5-ml capped polypropylene microcentrifuge tube 88% (v/v) formic acid 5 M CNBr in acetonitrile (Aldrich) 10 M NaOH (APPENDIX 2E) Milli-Q-purified water or equivalent continued Contributed by Dan L. Crimmins, Sheenah M. Mische, and Nancy D. Denslow Current Protocols in Protein Science (2000) 11.4.1-11.4.11 Copyright © 2000 by John Wiley & Sons, Inc.
Chemical Analyis
11.4.1 Supplement 19
Aluminum foil or opaque container 5-ml glass or polypropylene tube Additional reagents and equipment for reversed-phase (UNIT 11.6) or size-exclusion chromatography (UNIT 8.3), electrophoresis (UNIT 10.1), or electroblotting to PVDF membranes (UNIT 10.7) 1. Add 80 µl of 88% formic acid, 15 µl water, and 5 µl of 5 M CNBr in acetonitrile to lyophilized sample containing 1 to 50 µg total protein. The quantity of CNBr used for chemical cleavage of proteins at Met residues is based on a molar ratio of 200:1 (CNBr/Met). This ratio can be altered if the protein contains >2.3% Met residues.
2. Cap sample and vortex until the protein is completely solubilized in the CNBr solution. Cover sample tube with aluminum foil or place in an opaque container. Incubate 24 hr at room temperature in a chemical fume hood. IMPORTANT NOTE: This reaction must be performed in complete darkness to minimize unwanted side reactions with other amino acid side-chains.
3. Neutralize any unused CNBr by transferring remaining CNBr slowly to a 5-ml tube containing 5 to 10 vol of 10 M NaOH. Rinse the pipet with 10 M NaOH before discarding. CAUTION: Reaction of CNBr with NaOH is exothermic, generating heat and bumping of the solution. Therefore, add the CNBr solution slowly to minimize violent reactions. Once the CNBr is neutralized, it can be discarded according to halogen chemical waste disposal procedures.
4. Add 10 vol Milli-Q water to sample tube. Lyophilize overnight or dry sample completely (1 to 2 hr) in a Speedvac evaporator. CAUTION: After lyophilization or drying, defrost the cold trap and neutralize its contents by adding 10 M NaOH until the pH reaches ≥7.0. Discard contents according to halogen chemical waste disposal procedures.
5. Resolubilize sample in appropriate buffer for separation by reversed-phase (UNIT 10.6) or size-exclusion chromatography (UNIT 8.3) or for analysis by electrophoresis (UNIT 10.1) followed by electroblotting to PVDF membranes (UNIT 10.7). Some cleavage products can be difficult to resolubilize unless chaotropic agents (e.g., 6 M guanidine hydrochloride or 8 M urea) or detergents (e.g., 1% SDS or 1% cetyltrimethylammonium bromide) are used (see UNIT 1.3 & UNIT 6.1). Sonication is helpful for resolubilization, as is mild heat (i.e., in a 37°C water bath). Use of such agents may necessarily dictate the method chosen for fractionation. For example, detergent-treated samples are best analyzed by electrophoresis, whereas column chromatography should be used for guanidine- or urea-treated samples. Tris-tricine gels are recommended for electrophoretic separation of peptide fragments <10 kDa (UNIT 10.1). BASIC PROTOCOL 2
Chemical Cleavage of Proteins in Solution
CLEAVAGE AT THE C-TERMINUS OF TRP RESIDUES WITH BNPS-SKATOLE BNPS-skatole, or 2-(2′-nitrophenylsulfonyl)-3-methyl-3-bromoindolenine, cleaves proteins on the C-terminal side of unoxidized tryptophan (Trp) residues. For proteins with multiple Trp residues, fragments generated from this reaction can be incomplete (i.e., contain an internal unmodified or oxidized Trp). After digestion with BNPS-skatole, the resulting peptide fragments are fractionated by HPLC or electrophoresis prior to further characterization.
11.4.2 Supplement 19
Current Protocols in Protein Science
Materials Lyophilized protein sample Milli-Q-purified water or equivalent Dilute acid: e.g., 0.1% trifluoroacetic acid (TFA) or 1% acetic acid or formic acid 1 mg/ml BNPS-skatole (Pierce; store desiccated at −20°C) in glacial acetic acid (aldehyde-free; Mallinckrodt Specialty Chemicals), prepared immediately before use Aluminum foil or opaque container Beckman Microfuge 11 equipped with type 13.2 fixed-angle rotor, or equivalent microcentrifuge Additional reagents and equipment for reversed-phase (UNIT 11.6) or size-exclusion chromatography (UNIT 8.3), electrophoresis (UNIT 10.1), or electroblotting to PVDF membranes (UNIT 10.7) 1. In a 0.5-ml microcentrifuge tube, solubilize sample containing 1 to 50 µg total protein in solvent such as Milli-Q-purified water or dilute acid (0.1% TFA or 1% acetic or formic acid) to a final protein concentration of 1 mg/ml. Which solvent should be used for a particular protein may need to be determined empirically. Additional chaotropic agents—e.g., 8 M urea or 6 M guanidine hydrochloride—may be added to aid solubilization.
2. Add 60 µl BNPS-skatole to 20 µl protein. Cover tube with aluminum foil or place an opaque container over sample tube. Incubate 30 min at room temperature. It may be beneficial to increase the incubation time and temperature as well as adding chaotropic agents to the original protein solution to increase the yield from cleavage. If sufficient protein is available, an experiment covering an incubation time of 30 min to 24 hr, at room temperature and 47°C, can be used to optimize the reaction for a specific protein. IMPORTANT NOTE: This reaction must be performed in complete darkness to minimize unwanted side reactions with other reactive amino acid side-chains.
3. Add 80 µl Milli-Q-purified water to sample to precipitate BNPS-skatole. Centrifuge sample 5 min at 10,000 × g (12,000 rpm in Beckman Microfuge 11 with type 13.2 fixed-angle rotor), room temperature. Some researchers use an organic extraction with ethyl acetate (Gardner et al., 1994) or ethyl ether (Beach et al., 1992) to remove excess reagent and byproducts. A comparison with water extraction is presented in Rahali and Gueguen (1999).
4. Transfer supernatant to clean 0.5-ml microcentrifuge tube and discard pellet. Lyophilize or dry sample completely in a Speedvac evaporator. The resultant pellet may be slightly yellow due to residual reagent and byproducts.
5. Resolubilize sample in an appropriate buffer for separation by reversed-phase (UNIT 11.6) or size-exclusion chromatography (UNIT 8.3) or for analysis by electrophoresis (UNIT 10.1) followed by electroblotting to PVDF membranes (UNIT 10.7). Some cleavage products can be difficult to resolubilize unless chaotropic agents (e.g., 6 M guanidine hydrochloride or 8 M urea) or detergents (e.g., 1% SDS or 1% cetyltrimethylammonium bromide) are used (see UNIT 1.3 & UNIT 6.1). Sonication is helpful for resolubilization, as is mild heat (i.e., in a 37°C water bath). Use of such agents may necessarily dictate the method chosen for fractionation. For example, detergent-treated samples are best analyzed by electrophoresis, whereas column chromatography should be used for guanidine- or urea-treated samples. Tris-tricine gels are recommended for electrophoretic separation of peptide fragments <10 kDa (UNIT 10.1). Chemical Analyis
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BASIC PROTOCOL 3
CLEAVAGE AT ASP-PRO PEPTIDE BONDS WITH FORMIC ACID Formic acid cleaves proteins on the C-terminal side of the Asp residue in aspartic acid–proline (Asp-Pro) peptide bonds. For proteins with multiple Asp-Pro bonds, fragments generated from this reaction can be incomplete (i.e., contain an internal Asp-Pro bond). After digestion with formic acid, resulting peptide fragments are fractionated by HPLC or electrophoresis prior to further characterization. Materials 1 to 50 µg lyophilized protein sample in capped 0.5-ml microcentrifuge tube 70% (v/v) formic acid prepared immediately before use from 88% formic acid (Fluka) Milli-Q-purified water or equivalent Additional reagents and equipment for reversed-phase (UNIT 11.6) or size-exclusion (UNIT 8.3) chromatography, electrophoresis (UNIT 10.1), or electroblotting to PVDF membranes (UNIT 10.7) 1. Add 50 µl of 70% formic acid to lyophilized protein sample. Vortex thoroughly. Incubate 48 hr at 37°C in a chemical fume hood. 2. Add 150 µl Milli-Q water to sample. Lyophilize or dry sample completely. 3. Resolubilize sample in appropriate buffer for separation by reversed-phase (UNIT 11.6) or size-exclusion chromatography (UNIT 8.3) or for analysis by electrophoresis (UNIT 10.1) followed by electroblotting to PVDF membranes (UNIT 10.7). Some cleavage products can be difficult to resolubilize unless chaotropic agents (e.g., 6 M guanidine hydrochloride or 8 M urea) or detergents (e.g., 1% SDS or 1% cetyltrimethylammonium bromide) are used. Sonication is helpful for resolubilization, as is mild heat (i.e., in a 37°C water bath; see UNIT 1.3 & UNIT 6.1). Use of such agents may necessarily dictate the method chosen for fractionation. For example, detergent-treated samples are best analyzed by electrophoresis, whereas column chromatography should be used for guanidine- or urea-treated samples. Tris-tricine gels are recommended for electrophoretic separation of peptide fragments <10 kDa (UNIT 10.1).
BASIC PROTOCOL 4
CLEAVAGE AT ASN-GLY PEPTIDE BONDS WITH HYDROXYLAMINE Hydroxylamine cleaves proteins on the C-terminal side of Asn residues in asparagineglycine (Asn-Gly) peptide bonds. For proteins with multiple Asn-Gly peptide bonds, fragments generated from this reaction can be incomplete (i.e., contain an internal Asn-Gly). After digestion with hydroxylamine, resulting peptide fragments are fractionated by HPLC or electrophoresis prior to further characterization. Materials 1 to 50 µg lyophilized protein sample in a 0.5-ml capped microcentrifuge tube 1.8 M hydroxylamine solution (see recipe) 10% TFA 45°C oven or water bath Additional reagents and equipment for reversed-phase (UNIT 11.6) or size-exclusion chromatography (UNIT 8.3) 1. Add 200 µl of 1.8 M hydroxylamine solution to lyophilized protein sample. Vortex thoroughly. Incubate sample 3 hr at 45°C in a chemical fume hood.
Chemical Cleavage of Proteins in Solution
2. Allow sample to cool to room temperature and slowly add 10 µl of 10% TFA to quench the reaction.
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3. Fractionate or desalt the sample by reversed-phase (UNIT chromatography (UNIT 8.3).
11.6)
or size-exclusion
The high concentration of salt (1.8 M hydroxylamine) in the reaction mixture makes direct sample analysis by SDS-PAGE extremely problematic. The reaction products can be analyzed by electrophoresis, however, if the sample is desalted prior to analysis (UNIT 8.3). Tris-tricine gels are recommended for electrophoretically separating peptide fragments <10 kDa (UNIT 10.1).
CLEAVAGE AT THE N-TERMINUS OF CYS RESIDUES WITH NTCB Cleavage of proteins at cysteine residues following modification with 2-nitro-5-thiocyanobenzoate (NTCB) is a well-established procedure (Catsimpoolas and Wood, 1966; Jacobsen et al., 1973; Degani and Patchornick, 1974; Price, 1976; Lu and Gracy, 1981; Denslow and Nguyen, 1996) that has been paid little attention in recent years. NTCB cyanylates reduced cysteine thiols at pH 8. The chain is then cleaved by cyclization under alkaline conditions. The N-terminus of the released peptide fragment becomes modified by the iminothiazolidine-carboxyl group during the cleavage reaction (Catsimpoolas and Wood, 1966; Jacobsen et al., 1973), thus rendering the fragments blocked for N-terminal sequence analysis by Edman chemistry. For this reason, this cleavage method has not been much used. The advent of mass spectrometry, especially matrix-assisted laser-desorption ionization time-of-flight (MALDI TOF) mass spectrometry (UNIT 16.2) with delayed extraction makes this problem less significant, since amino acid sequences of fragments can now be identified (Denslow and Nguyen, 1996; Daniel et al., 1997).
BASIC PROTOCOL 5
Cysteine plays an important role in protein secondary structure. Only reduced cysteines are able to react with NTCB, making this reagent an important one to distinguish free sulfhydryls from those involved in disulfide linkages. In addition, cysteines are relatively scarce in proteins; thus cleavage with NTCB can generate rather large fragments that can then be identified and used for further structural studies. Materials Protein samples: 25 to 50 pmol in 20 µl of 200 mM Tris-acetate, pH 8 (see recipe)/1 mM EDTA/0.1% SDS Ekathiol resin (Ekagen) Nitrogen source Acidified acetone, pH 3, ice-cold (see recipe) 20 mM NTCB (see recipe) 100 mM sodium borate, pH 9 2 mM NaOH Ultramicrospin columns, gel filtration G10 or G25 media (AmiKa) or acidified acetone, pH 3, ice-cold (see recipe) 100% acetonitrile 50% and 60% acetonitrile in 0.1% (v/v) trifluoroacetic acid (TFA) 0.5% and 0.1% (v/v) TFA 40° and 50°C water baths Centrifuge and rotor (e.g., Eppendorf 5415C, F-45-18-11 rotor) Zip tip (C18-filled tip; Millipore) Additional reagents and equipment for MALDI-TOF mass spectrometry (UNIT 16.2) Reduce disulfides and purify the protein 1. Treat protein samples with 10× to 20× molar excess of Ekathiol to fully reduce cysteine thiols. Seal tubes under N2 and shake 2 hr at room temperature. Chemical Analyis
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Ekathiol resin contains 0.3 to 0.7 mmol of SH-substitution per gram resin. This treatment will reduce all Cys in the protein and give maximum cleavage with NTCB. If the resin is not available, 1 mM dithiothreitol (DTT) can be used to reduce the protein. However, in this case, make sure that in the subsequent step there is an excess of NTCB over all thiols, including those coming from DTT. The advantage of the resin is that it can be easily separated from the protein after reduction. Omit this reduction step if the position of disulfides is to be mapped.
2. Microcentrifuge 5 min at top speed to pellet the resin and transfer the supernatant containing the reduced protein to a new tube containing 5 µl of 20 mM NTCB, resulting in a final concentration of 4 mM NTCB. Seal tube under N2 and incubate 20 min at 40°C. During this step, NTCB forms a covalent bond with reduced Cys-thiols in the protein. NTCB concentration should be ∼10-fold molar excess over total thiol groups.
To purify the protein through a gel-filtration spin column 3a. Add 100 µl distilled or Milli-Q-purified water to the sample tube and place on top of a gel-filtration ultramicrospin column prepared according to manufacturer’s specifications. 4a. Centrifuge the column 5 min at 1000 × g (3,500 rpm in an Eppendorf microcentrifuge 5415C, F-45-18-11 rotor) and collect supernatant in a 0.5-ml microcentrifuge collecting tube. At this point, the sample is essentially free of NTCB, which is necessary to prevent side reactions.
5a. Decrease the sample volume in a Speedvac evaporator to ∼20 µl and then add an equal volume of 100 mM sodium borate, pH 9. Incubate 1 hr at 50°C. To purify the protein by acetone precipitation 3b. Add 9 vol ice-cold acidified acetone, pH 3. Incubate overnight at −20°C, or 30 min in a dry-ice/ethanol bath to fully precipitate the protein. 4b. Microcentrifuge 5 min at top speed, then carefully remove the supernatant to avoid disturbing the pellet. Rinse the pellet once with ice-cold acidified acetone and centrifuge again. Remove the acetone and let the pellet air dry. Acetone precipitation works well for quantities of at least 1 ìg/ml protein.
5b. Redissolve the sample in 20 µl 100 mM sodium borate, pH 9 and then add 20 µl of water. Incubate 1 hr at 50°C. Cleave and analyze the protein 6. Remove a 1-µl aliquot and spot onto pH paper to make sure pH is 9. Adjust pH, if necessary, with 2 mM NaOH. Vortex to make sure samples are well dissolved. Incubate 1 hr at 50°C. A basic pH is required to cleave the protein. This happens when the NTCB-modified thiol of the Cys cyclizes with the amino nitrogen cleaving the protein to produce an amino-terminal iminothiazolidine-4-carboxyl residue.
Chemical Cleavage of Proteins in Solution
7. While incubation is proceeding, prepare a Zip tip by following the manufacturer’s instructions. Attach the Zip tip to the tip of a standard micropipettor. Wet the Zip tip with 10 µl of 100% acetonitrile, rinse once with 10 µl of 50% acetonitrile in 0.1% TFA, and finally rinse twice with 10 µl of 0.1% TFA. Keep Zip tip wet with 0.1% TFA until needed.
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8. Quench the sample by adding 10 µl of 0.5% TFA solution, so that the final concentration of TFA is 0.1%. 9. Immediately adsorb the sample onto the Zip tip by pipetting 10-µl aliquots through the C18 reverse-phase material of the Zip tip several times. 10. Rinse the Zip tip twice with 10 µl of 0.1% TFA. This step is important to remove traces of NTCB and buffers in order to improve the signal-to-noise ratio in the mass spectrometer.
11. Elute with 10 µl of 60% acetonitrile/0.1% TFA or directly with the matrix of choice. α-cyano-4-hydroxycinnamic acid matrix works well for most fragments. To prepare this matrix, weigh 10 mg of re-crystallized α-cyano-4-hydroxycinnamic acid matrix into a tube. Dissolve in 1 ml of 50% acetonitrile/0.1% TFA. Vortex 1 min to dissolve as much of the resin as possible. This should be a saturated solution; therefore not all of the matrix will go into solution. Pellet by centrifuging and use the supernatant. Other matrices, e.g., 3,5-dimethoxy-4-hydroxycinnamic acid (sinapinic acid), can be tried as well (see UNIT 16.2).
12. Place 1 µl sample (equal parts eluted sample and matrix) on a sample plate and analyze by MALDI-TOF mass spectrometry (see UNIT 16.2). REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Acidified acetone, pH 3.0 Add 1 drop of 0.1 M HCl to 100 ml acetone, mix, and place in a −20°C freezer ≥2 hr to chill. Hydroxylamine solution, 1.8 M Place 5 g hydroxylamine (Fluka) in a 50-ml graduated cylinder. Add 20 ml of 5 M NaOH (pH should be ∼7.2). If salt does not dissolve, continue adding NaOH until pH is 7 to 8. Add 1 g Na2CO3 and adjust pH to 9 with 12 M HCl. Add Milli-Q water to 40 ml and readjust pH to 9 if needed. Prepare immediately before use. 2-Nitro-5-thiocyanobenzoate (NTCB), 20 mM Weigh 9 mg NTCB (mol. wt. 224.2; Sigma) into a clean tube. Add 2 ml of 200 mM Tris-acetate, pH 8 (see recipe). Vortex to dissolve. Make fresh for each use. Tris-acetate, pH 8.0, 200 mM Add 969 mg Tris base (mol. wt. 121.1) to 35 ml distilled water. Dissolve completely and adjust to pH 8 with acetic acid. Bring volume to 40 ml. COMMENTARY Background Information The most specific and effective way to fragment proteins into relatively large peptides is by chemical cleavage. CNBr and BNPS-skatole cleave specifically at methionine (Met; Gross, 1967) and tryptophan (Trp; Fontana, 1972) residues, respectively. These two amino acids are in relatively low abundance (Crawford, 1990) and are each encoded by a single codon in the genetic code. As a consequence,
protein fragmentation at these residues is desirable because relatively long peptides are obtained (Jay, 1984; Jue and Doolittle, 1985), and it is ideally suited for the design of low-degeneracy nucleotide probes. In addition to the protocols presented in this chapter, chemical cleavage procedures specific for other low-frequency residues (Crawford, 1990) have been successful (Smith, 1994). Proteins can be cleaved at tyrosine (Tyr) residues
Chemical Analyis
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with N-bromosuccinimide (NBS), but this reagent will also cleave at Trp residues. The large excess of NBS required to obtain a reasonable yield typically results in modification of sulfurcontaining amino acids (Met and Cys) and histidine (His; Fontana and Gross, 1986). Trp residues can also be cleaved with o-iodosobenzoic acid but with a reported specificity and yield less than that of the preferred BNPS-skatole method. Reaction of NTCB with cysteine thiols has been well characterized in the literature (Catsimpoolas and Wood, 1966; Jacobsen et al., 1973; Degani and Patchornick, 1974; Price, 1976; Lu and Gracy, 1981). The predominant reaction under these conditions is the cyanylation of the protein in the first step of the reaction followed by cleavage (N-terminal to the Cys) of the protein under alkaline conditions. Several side reactions, however, are also possible, including β-elimination of the thiocyanoalanine residue instead of cyclization and cleavage (Degani and Patchornick, 1974) and the formation of mixed disulfides (Price, 1976). Using the Ekathiol resin to reduce the sample and removing excess NTCB promptly after cyanylation minimizes the number of side reactions. In order to favor the cyanylation reaction, a 10-fold molar excess of NTCB over total thiol groups should be used at pH 8 for a short incubation period—15 to 60 min at 37° to 50°C. Immediate acidification will prevent most side reactions. Prompt removal of the NTCB reagent will also help reduce side reactions. Cyclization and cleavage of the peptide backbone is accomplished by raising the pH to 9.0 for a longer incubation period. This method is ideally suited to determining the positions of disulfides in proteins. In this case, the protein is not reduced prior to cyanylation, but instead is treated directly with NTCB. If sufficient protein is available, the reaction can be performed on both the native and reduced protein. This would control for those Cys that may be oxidized but not involved in disulfide interactions.
Critical Parameters
Chemical Cleavage of Proteins in Solution
A general disadvantage of any chemical cleavage procedure is that chemical fragmentation tends to be incomplete (Smith, 1994), leaving some peptide fragments with undigested internal target residues. One can attempt to increase fragment yield by a combination of increasing chemical reagent concentration, reaction time, and temperature, and adding denaturants during the cleavage reaction. Further,
the high concentration of reagents used may cause unwanted modification of non-target amino acid residues in the protein (a problem that is exacerbated if the reagent concentrations are increased as just suggested). Side reactions (e.g., whereby BNPS-skatole may oxidize Cys or Met) can be minimized by performing the cleavage in complete darkness, as described in the protocols. In addition, all reagents must be pure and fresh, and solutions should be stored as recommended and made fresh immediately prior to their use. Taking these few precautions will result in optimal cleavage with minimal side reactions. If possible, mass spectrometry (UNIT 16.1) should be used to characterize the resulting cleavage products and will greatly aid in assessing the extent of side reactions. This approach was successfully employed to determine side-reaction modifications formed during solution-phase BNPS-skatole cleavage of proteins and peptides (Vestling et al., 1995). A detailed study comparing the removal of excess BNPS-skatole, its byproducts, and cleavage fragment recovery via water or ethyl ether extraction with dialysis or gel filtration has recently appeared (Rahali and Gueguen, 1999). Using alkylated bovine β-lactoglobulin as a model protein, these authors found that the highest fragment recovery and greatest reagent removal was achieved by following the water extraction step with a gel filtration step. Successful cleavage and recovery of fragments for sequence analysis can be performed with as little as 5 µg of pure protein. However, obtaining unambiguous protein sequence data requires that the protein sample be pure. Correct estimation of protein quantity is the single most important factor in determining whether sequence data can be obtained. All too often, an estimated 10 pmol turns out to be 1 pmol. Therefore, care should be taken to prepare adequate amounts of protein for cleavage and to estimate protein concentration accurately. Obtaining an accurate estimate of the amount of protein available can be difficult. The best method involves amino acid compositional analysis (UNIT 3.2). This also provides information about the amino acid composition of the protein, which is also important because it should ultimately correspond to the predicted sequence. If the protein concentration cannot be determined by amino acid compositional analysis, estimate the concentration by comparing the sample with known quantities of a standard protein by SDS-PAGE (UNIT 10.1). It is best to use a standard protein that approximates the molecular weight and possible secondary
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Table 11.4.1 Commercially Available Proteins Useful as Positive Controls for Chemical Cleavage Reactionsa,b
Cleavage reaction/target residue(s)
Locationc
Human immunoglobulin heavy and light chains
CNBr/Met
Bovine serum albumin
BNPS-skatole/Trp
Light chain: Met at 4 and 89 (214 residues total) Heavy chain: Met at 82, 265, and 459 (478 residues total) Trp at 134 and 212 (582 residues total)
Protein
Human serum transferrin Formic acid/Asp-Pro
Asp-Pro at 90-91 and 548-549 (679 residues total) Bovine brain calmodulin Hydroxylamine/Asn-Gly Asn-Gly at 61-62 and 97-98 (148 residues total) Rabbit cardiac α-tropomyosin
NTCB/Cys
Cys at 190 (284 residues total)
aSDS-PAGE is by far the preferred analytical technique to monitor the reaction. In addition, the reaction products can be
electroblotted to a PVDF membrane (UNIT 10.7) for N-terminal sequence analysis to assess the efficiency of the chemical cleavage reaction. bAbbreviations: Asn, asparagine; Asp, aspartic acid; BNPS-skatole, 2-(2′-nitrophenylsulfonyl)-3-methyl-3-bromo indo-
lenine; CNBr, cyanogen bromide; Cys, cysteine; Gly, glycine; Met, methionine; NTCB, 2-nitro-5-thiocyanobenzoic acid; Pro, proline; Trp, tryptophan. cThe extent of cleavage at each site is not the same, resulting in a skewed distribution of fragments. For a protein with n
cleavage sites (n amino acids not at the C-terminus), there are a total of [(n + 1)(n + 2)]/2 possible fragments, (n + 1) of which are complete digest fragments, and [n(n + 1)]/2 of which are incomplete digestion fragments, including the intact, undigested protein substrate (Crimmins et al., 1990).
structure of the unknown protein. After staining with either Coomassie blue or amido black (UNIT 10.5), the gel can be scanned using a densitometer and a linear standard curve can be generated based on the amount of protein versus staining intensity. Analytical SDS-PAGE can also be used to estimate the quantity of an unknown protein prior to committing the remainder of the protein preparation to a particular cleavage method. Molar concentrations can be calculated from quantitation and molecular weight.
Troubleshooting Commercial proteins of known sequence are useful for monitoring and optimizing the efficiency of chemical cleavage reactions. Individual heavy and light chains of human immunoglobulin G (IgG) proteins can be used to monitor CNBr cleavage; BSA can be used to monitor BNPS-skatole cleavage; bovine brain calmodulin (CAL) can be used to monitor hydroxylamine cleavage at Asn-Gly peptide bonds; human serum transferrin (HST) can be used to monitor formic acid cleavage at AspPro peptide bonds; and rabbit cardiac α-tropomyosin (Tm) can be used to monitor NTCB cleavage. Appropriate protein standards must be included as positive controls for any chemi-
cal cleavage experiment on any protein; reaction products thus generated are easily analyzed by SDS-PAGE (UNIT 10.1). Table 11.4.1 provides the locations of the chemical cleavage sites for the five standard proteins discussed above. The efficiency of chemical cleavage can be compromised if the protein to be analyzed contains any disulfide bonds. For Basic Protocols 1 through 4, if the sample has Cys residues that are involved in disulfide bond formation, it should be reduced and alkylated prior to cleavage to improve cleavage efficiency and yield. This step will also permit unalkylated Cys residues that would be missed by Edman degradation to be identified by sequence analysis. The protein should be denatured by dissolution in 6 M guanidine⋅HCl or 8 M urea (see UNIT 6.3). Reduction is performed under an inert atmosphere (nitrogen or argon) with dithiothreitol (DTT), 2-mercaptoethanol, or tributyl phosphine (the latter is commonly used because of its volatility). It is critical to take extreme care in following all precautions recommended by the manufacturer when using this reagent. Alkylating reagents include iodoacetamide, iodoacetate, and vinyl pyridine, which modify Cys to carboxamidomethyl-Cys, carboxymethyl-Cys, and pyridylethyl-Cys, respectively.
Chemical Analyis
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Reduction and alkylation are best performed in solution prior to cleavage: the sample should then be desalted by conventional chromatographic methods (UNIT 8.3). An alternative is to use PVDF sample cleanup filters manufactured by Applied Biosystems (ProSpin or ProBlot cartridge) or Millipore. These are easy to use and require no more complex equipment than a centrifuge that can hold a 1.5-ml microcentrifuge tube. The sample mixture is applied, then centrifuged for several hours (depending on the volume and viscosity), and the end result is the protein bound to a PVDF membrane. At this point, the protein sample can be washed and subjected to chemical cleavage for membrane-bound protein (UNIT 11.5). For Basic Protocol 5, the Cys residues must be reduced and unmodified for successful cyanylation prior to cleavage. This is readily accomplished with the preferred method of Ekathiol reduction. If there is poor or no cleavage with CNBr, BNPS-skatole, or NTCB, the Met, Trp, or Cys residues respectively may be oxidized. Methionine sulfoxide oxidation can be reversed by the use of N-methylmercaptoacetamide (Houghton and Li, 1979), but this method will not reverse methionine sulfone oxidation. Unfortunately, there is no method available for reversing Trp oxidation. Most of these oxidations occur during the isolation and characterization of the protein of interest, and steps should be taken to minimize these occurrences. In addition, Met-Thr (methionine-threonine) and Met-Ser (methionine-serine) cleavage yields with CNBr tend to be poor using the standard protocol. A mechanistic study (Kaiser and Metzka, 1999) has shown that increasing the water content of the cleavage reaction mixture improves the cleavage yield at these bonds. Deamidation of Asn residues occurs spontaneously during protein degradation, causing the formation of a succinimide intermediate (Aswad and Guzzetta, 1995). If this occurs, hydroxylamine will not cleave at the Asn-Gly peptide bond under the conditions detailed in Basic Protocol 4. However, cleavage will occur if denaturants are included in the cleavage solution (Kwong and Harris, 1994). This ability to differentiate modified Asn residues may be helpful in the elucidation of the primary structure of a protein containing these residues.
Anticipated Results Chemical Cleavage of Proteins in Solution
The protocols detailed in this unit have been successfully applied to a variety of proteins. As previously discussed, fewer fragments will be generated by chemical cleavage and these frag-
ments will be larger in size than those produced by standard enzymatic treatment. Consequently, in addition to reversed-phase (UNIT 11.6), ion-exchange (UNIT 8.2), and size-exclusion (UNIT 8.3) chromatographies, analytical high-resolution electrophoresis (UNIT 10.1) can be used to purify the peptide fragments. Electrophoresis is often the method of choice for separating small quantities of peptides because it is a technique most laboratories are equipped to perform and the recovery of the cleaved peptides is relatively high. Tris-tricine gels are recommended for peptides of lower molecular weight (<10 kDa) because this buffer system offers good resolution (UNIT 10.1). Electroblotting to PVDF membranes is the only additional step required for preparing the peptides for sequence analysis (UNIT 10.8). As a rule of thumb, 100 pmol of pure protein should yield 25 pmol of cleaved peptide on PVDF after electrophoresis and electroblotting (assuming a 50% efficiency for both cleavage and electrotransfer). These numbers may vary, but it is best to start with at least 100 pmol of pure protein.
Time Considerations In general, allow a full two days for completion of both the cleavage and isolation of peptides. It is best to proceed directly from the cleavage reaction to peptide isolation without stopping in order to minimize protein damage.
Literature Cited Aswad, D.W. and Guzzetta, A.W. 1995. Methods for analysis of deamidation and isoasparate formation in peptides and proteins. In Deamidation and Isoaspartate Formation in Peptides and Proteins (D. Aswad, ed.) pp. 7-29. CRC Press, Boca Raton, Fla. Beach, C.M., DeBeer, M.C., Sipe, J.D., Loose, L.D., and DeBeer, F.C. 1992. Human serum amyloid A protein. Complete amino acid sequence of a new variant. Biochem. J. 282:615-620. Catsimpoolas, N. and Wood, J.L. 1966. Specific cleavage of cystine peptides by cyanide. J. Biol. Chem. 241:1790-1796. Crawford, M. 1990. Protein compositions: What is average? ABRF News 1:7. Crimmins, D.L., McCourt, D.W., Thoma, R.S., Scott, M.G., Macke, K., and Schwartz, S.D. 1990. In situ chemical cleavage of proteins immobilized to glass fiber and polyvinylidenedifluoride membranes: Cleavage at tryptophan residues with (2-(2′-nitrophenylsulfonyl)-3methyl-3-bromoindolenine) to obtain internal amino acid sequence. Anal. Biochem. 187:27-38. Daniel, R., Camide, E., Martel, A., LeGoffic, F., Canosa, D., Carrascal, M., and Abian, J. 1997. Mass spectrometric determination of the cleavage sites in Escherichia coli dihydroorotase in-
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duced by a cysteine-specific reagent. Biochemistry 272:26934-26939. Degani, Y. and Patchornick, A. 1974. Cyanylation of sulfhydryl groups by 2-nitro-5-thiocyanobenzoic acid. High yield modification and cleavage of peptides at cysteine residues. Biochemistry 13:1-11. Denslow, N.D. and Nguyen, H.P. 1996. Specific cleavage of blotted proteins at cysteine residues after cyanylation: analysis of products by MALDI-TOF. In Techniques in Protein Chemistry VII (D.R. Marshak, ed.) pp. 241-248. Academic Press, San Diego. Fontana, A. 1972. Modification of tryptophan with BNPS-skatole (2-(2′-nitrophenylsulfonyl)-3methyl-3′- bromoindolenine). Methods Enzymol. 25:419-423. Fontana, A. and Gross, E. 1986. Fragmentation of polypeptides by chemical methods. In Methods in Protein Chemistry: A Handbook (A. Darbre, ed.), pp. 68-120. John Wiley & Sons, Chichester, U.K., and New York. Gardner, A.M., Vaillancourt, R.R., Lange-Carter, C.A., and Johnson, G.J. 1994. MEK-1 phosphorylation by MEK kinase, Raf, and mitogen-activated protein kinase: Analysis of phosphopeptides and regulation of activity. Mol. Biol. Cell 5:193-201.
Kwong, M.Y. and Harris, R.J. 1994. Identification of succinimide sites in proteins by N-terminal sequence analysis after alkaline hydroxylamine cleavage. Protein Sci. 3:147-149. Lu, H.S. and Gracy, R.W. 1981. Specific cleavage of glucosephosphate isomerases at cysteinyl residues using 2-nitro-5-thiocyanobenzoic acid: Analyses of peptides eluted from polyacrylamide gels and localization of active site histidyl and lysyl residues. Arch. Biochem. Biophys. 212:347-359. Price, N.C. 1976. Alternative products in the reaction of 2-nitro-5-thiocyanatobenzoic acid with thiol groups. Biochem. J. 159:177-180. Rahali, V. and Gueguen, J. 1999. Chemical cleavage of bovine beta-lactoglobulin by BNPS-skatole for preparative purposes: Comparative study of hydrolytic procedures and peptide characterization. J. Prot. Chem. 18:1-12. Smith, B. 1994. Chemical cleavage of proteins. In Methods in Molecular Biology, Vol. 32: Basic Protein and Peptide Protocols (J. Walker, ed.) pp. 297-309. Humana Press, Totowa, N.J. Vestling, M.M., Kelly, M.A., and Fenselau, C. 1995. Optimization by mass spectrometry of a tryptophan-specific protein cleavage reaction. Rapid Commun. Mass Spectrom. 8:786-790.
Gross, E. 1967. The cyanogen bromide reaction. Methods Enzymol. 11:238-255.
Key References
Houghton, R.A. and Li, C.-H. 1979. Reduction of sulfoxides in peptides and proteins. Anal. Biochem. 98:36-46.
Excellent source of chemical cleavage procedures.
Jacobsen, G.R., Schaffer, M.H., Stark, G.R., and Vanaman, T.C. 1973. Specific chemical cleavage in high yield at the amino peptide bonds of cysteine and cystine residues. J. Biol. Chem. 248:6583-6591.
Valuable, up-to-date summary of several chemical cleavage procedures.
Jay, D.G. 1984. A general procedure for the end labeling of proteins and positioning of amino acids in the sequence. J. Biol. Chem. 269:1557215578. Jue, R.A. and Doolittle, R.F. 1985. Determination of the relative positions of amino acids by partial specific cleavages of end-labeled proteins. Biochemistry 24:162-170. Kaiser, R. and Metzka, L. 1999. Enhancement of cy an o gen bro mid e cleavag e yield s for methionyl-serine and methionyl-threonine peptide bonds. Anal. Biochem. 266:1-8.
Fontana, A. and Gross, E. 1986. See above.
Smith, B. 1994. See above.
Contributed by Dan L. Crimmins Washington University School of Medicine St. Louis, Missouri Sheenah M. Mische Boehringer Ingelheim Pharmaceuticals Ridgefield, Connecticut Nancy D. Denslow University of Florida Gainsville, Florida
Chemical Analyis
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Chemical Cleavage of Proteins on Membranes
UNIT 11.5
Sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) followed by electrotransfer to polyvinylidene difluoride (PVDF) membranes (UNIT 10.8) has become the most widely-used method of preparing protein samples for primary structural analysis. Very often SDS-PAGE (UNIT 10.1) or two-dimensional isoelectrophoresis/SDS-PAGE (UNIT 10.4) is used as a final preparative step for proteins that can only be recovered in very small quantities. The protocols in this unit describe in situ chemical cleavage methods that have been optimized for proteins that are noncovalently bound to PVDF membranes. These procedures can also be used to analyze PVDF-bound protein samples that have already been subjected to Edman degradation. This allows a sample with a blocked N-terminus to be chemically cleaved and resubmitted for sequence analysis to obtain internal sequence data. Alternatively, the reaction fragments can be eluted from the membrane and fractionated by HPLC or analytical high-resolution electrophoresis (UNIT 10.1) prior to further analysis. The fragments generated from chemical digestion tend, on average, to be fewer in number and larger in size than those produced via proteolysis. These chemical cleavage fragments are thus useful as substrate species for secondary cleavage reactions or for directly obtaining internal protein sequence data for use in the subsequent design of DNA probes for genomic or cDNA-library screening. In this unit, five basic protocols present widely used reactions that are specific and efficient for the in situ chemical cleavage of proteins bound to membranes (related protocols for cleaving proteins in solution, utilizing the same chemical reactions, are presented in UNIT 11.4). Cyanogen bromide (CNBr; Basic Protocol 1) cleaves proteins at methionine (Met) residues; 2-(2′-nitrophenylsulfonyl)-3-methyl-3-bromoindolenine (BNPS-skatole; Basic Protocol 2) cleaves proteins at tryptophan (Trp) residues; formic acid (Basic Protocol 3) cleaves proteins at aspartic acid–proline (Asp-Pro) peptide bonds; hydroxylamine (Basic Protocol 4) cleaves proteins at asparagine-glycine (Asn-Gly) peptide bonds, and 2-nitro5-thiocyanobenzoic acid (NTCB; Basic Protocol 5) cleaves proteins at cysteine (Cys) residues. In general, reactions are chosen based on a priori knowledge of a protein’s target residues. If no information is available, perform the reactions in the order listed. In addition, an Alternate Protocol describes CNBr cleavage of PVDF-bound protein previously analyzed by Edman degradation. Finally, a Support Protocol discusses preferred methods of separating and analyzing peptide fragments generated by the chemical cleavage reactions described in the basic protocols. CAUTION: Most chemicals used in the following protocols are toxic. Therefore, it is imperative that good laboratory practice be followed. All procedures and incubations should be performed in a well-ventilated chemical fume hood and laboratory personnel should wear appropriate protective clothing, including safety glasses and gloves. All chemical waste should be properly disposed of according to local regulations. CLEAVAGE AT THE C-TERMINUS OF MET RESIDUES WITH CNBr CNBr cleaves proteins on the C-terminal side of unoxidized methionine (Met) residues. For proteins with multiple Met residues, fragments generated from this reaction can be incomplete (i.e., contain an internal unmodified Met or homoserine). After the reaction, the PVDF-bound peptides may be sequenced directly or eluted from the PVDF membrane (UNIT 10.7) and fractionated prior to further characterization (if multiple N-termini are
BASIC PROTOCOL 1
Chemical Analysis Contributed by Dan L. Crimmins, Sheenah M. Mische, and Nancy D. Denslow Current Protocols in Protein Science (2000) 11.5.1-11.5.13 Copyright © 2000 by John Wiley & Sons, Inc.
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expected from a protein that contains more than one Met residue). Additionally, if the N-terminus of a protein is thought to be blocked (because no sequence could be obtained from the first sequencing attempt although a sufficient amount of the protein was analyzed), in situ CNBr cleavage can be used to confirm N-terminal blockage by directly sequencing the PVDF membrane after the cleavage reaction. Materials Dried PVDF membrane containing electroblotted protein sample (UNIT 10.7) 0.25 M CNBr in 70% formic acid (see recipe) 10 M NaOH Milli-Q water or equivalent Clean razor blade Aluminum foil or opaque container 47°C water bath or oven (optional) Additional reagents and equipment for analysis and separation of cleavage fragments (see Support Protocol) 1. Feather PVDF membrane containing electroblotted protein by making parallel cuts running most of its length with a clean razor blade to give a pattern similar to that of a rake head. Feathering the PVDF membrane increases its surface area.
2. Place feathered membrane in a clean 0.5-ml microcentrifuge tube and add 150 µl of 0.25M CNBr in 70% formic acid. Cap tube, then wrap completely with aluminum foil or place in an opaque container. Incubate with agitation in a chemical fume hood either 24 to 48 hr at room temperature or 1 hr at 47°C. Traditionally, CNBr incubations are performed for 24 to 48 hr at room temperature; however, we find that the shorter incubation at 47°C produces comparable results for most proteins. IMPORTANT NOTE: This reaction must be performed in complete darkness to minimize unwanted side reactions with other reactive amino acid side-chains.
3. Neutralize unused CNBr by slowly transferring remaining CNBr to a 5-ml tube containing 5 to 10 vol of 10 M NaOH. Rinse the pipet with 10 M NaOH before discarding. CAUTION: Reaction of CNBr with NaOH is exothermic, generating heat and bumping of the solution. Therefore, add the CNBr solution slowly to minimize violent reactions. Once the CNBr has been neutralized, discard it according to halogen chemical waste disposal procedures.
4. Dry entire reaction mixture (membrane plus liquid) 1 to 2 hr in a Speedvac evaporator. 5. Add 50 µl Milli-Q water to dried sample. Vortex and dry sample again. The water wash removes residual acid; this is important for subsequent electrophoresis or chromatography.
6. Analyze peptide sequence directly from membrane or elute and separate peptide fragments (see Support Protocol).
Chemical Cleavage of Proteins on Membranes
CAUTION: After lyophilization or drying, defrost the cold trap and neutralize its contents by adding 10 M NaOH until the pH reaches ≥7.0. Discard contents according to halogen chemical waste disposal procedures.
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CNBr CLEAVAGE OF PVDF-BOUND PROTEIN PREVIOUSLY ANALYZED BY EDMAN DEGRADATION
ALTERNATE PROTOCOL
This protocol is used to obtain internal protein sequence from a PVDF-bound protein that has been previously sequenced via Edman degradation. It combines in situ CNBr cleavage and o-phthalaldehyde (OPA) blockage to obtain protein sequence data from samples that contain a small amount of protein. It should be noted that this cleavage is relatively inefficient, usually generating a small number of N-termini (1 to 4) that can be sequenced. In favorable circumstances, one gets only a single or major sequence, thus allowing unambiguous assignment. Additional Materials (also see Basic Protocol 1) 1 M CNBr in 70% formic acid (see recipe) 0.5 mg/ml o-phthalaldehyde (OPA), in butyl chloride (prepared immediately before use) Preconditioned polybrene-treated glass-fiber filters 1. Feather PVDF membrane containing bound protein sample using a clean razor blade (see Basic Protocol 1, step 1). Place membrane on top of preconditioned polybrenetreated glass-fiber filter. Preconditioning of polybrene-treated glass-fiber filters is done as a prerequisite to protein sequencing. Refer to Atherton et al. (1993a) for details regarding sequence analysis of proteins.
2. Apply 30 to 42 µl of 1 M CNBr in 70% formic acid to sample until filter and PVDF membrane are saturated. 3. Wrap wet filters in Parafilm to prevent evaporation, then wrap with aluminum foil or cover with opaque container. Incubate 24 hr at room temperature in chemical fume hood. The glass reaction-cartridge block from an ABI sequencer can be used as the support for this reaction. Alternately, the filter and PVDF membrane can be placed on top of a piece of Parafilm large enough to create an envelope around the membrane and filter when folded. Take care to avoid contact between the Parafilm and the membrane. IMPORTANT NOTE: This reaction must be performed in complete darkness to minimize unwanted side reactions with other reactive amino acid side-chains.
4. Using a clean razor blade, slice a hole in Parafilm and place the packet in a Speedvac evaporator. Dry sample for 10 min. 5. Prepare PVDF membrane for sequence analysis using a fresh preconditioned polybrene-treated glass-fiber filter. 6. Subject ∼25% of membrane to Edman degradation (UNIT positions of Pro residues.
11.10)
to ascertain the
7. Block other residues on remaining membrane with OPA. Subject blocked membrane to further sequence analysis via Edman degradation. OPA blockage will result in a sequence beginning with one or more Pro residues. Consult Brauer et al. (1984) for specific details regarding modification of sequencer cycles to optimize OPA blockage, and Atherton et al. (1993a) for details regarding sequence analysis.
Chemical Analysis
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BASIC PROTOCOL 2
CLEAVAGE AT THE C-TERMINUS OF TRP RESIDUES WITH BNPS-SKATOLE BNPS-skatole cleaves proteins on the C-terminal side of unoxidized tryptophan (Trp) residues. For proteins with multiple Trp residues, fragments generated from this reaction can be incomplete (i.e., contain an internal unmodified or oxidized Trp). After the reaction, the PVDF membrane may be sequenced directly or the fragments eluted from the PVDF membrane (UNIT 10.7) and fractionated prior to further characterization (if multiple N-termini are expected because the protein contains more than one Trp). Additionally, if the N-terminus of a protein is thought to be blocked (because no sequence is obtained from the first sequencing attempt although a sufficient amount of protein was analyzed), in situ BNPS-skatole cleavage can be used to confirm N-terminal blockage by directly sequencing the PVDF-bound fragments after the cleavage reaction. Materials Dried PVDF membrane containing electroblotted protein sample (UNIT 10.7) Milli-Q water or equivalent 1 mg/ml BNPS-skatole (Pierce; store desiccated at −20°C) in 75% glacial acetic acid (aldehyde-free, Mallinckrodt Specialty Chemicals), prepared immediately before use Clean razor blade Aluminum foil or opaque container 47°C water bath or oven Beckman Microcentrifuge equipped with type 13.2 fixed-angle rotor, or equivalent microcentrifuge Additional reagents and equipment for analysis and separation of cleavage fragments (see Support Protocol) 1. Feather PVDF membrane containing protein of interest using a clean razor blade (see Basic Protocol 1, step 1). 2. Place feathered membrane in clean 1.5-ml microcentrifuge tube and add 100 µl BNPS-skatole. Cap tube, then wrap with aluminum foil or place in an opaque container. Incubate 1 hr at 47°C with occasional agitation. IMPORTANT NOTE: This reaction must be performed in complete darkness to minimize unwanted side reactions with other reactive amino acid side-chains.
3. Remove liquid portion of sample to a clean 0.5-ml microcentrifuge tube and save until use in step 6. 4. Add 500 µl Milli-Q water to tube containing PVDF membrane and vortex vigorously. Aspirate water from membrane and discard. 5. Repeat wash four more times. Dry membrane in a Speedvac evaporator. 6. Add 100 µl Milli-Q water to liquid sample from step 3. Centrifuge 5 min at 10,000 × g (12,000 rpm in Beckman Microfuge with type 13.2 fixed-angle rotor), room temperature. This step precipitates BNPS-skatole from the reaction mixture.
7. Transfer supernatant to clean microcentrifuge tube and dry in a Speedvac evaporator. Chemical Cleavage of Proteins on Membranes
8. Add 100 µl Milli-Q water to dried sample and vortex ∼5 min. Dry resulting solution in a Speedvac evaporator. Initial pellet may appear yellow due to residual reagent and byproducts.
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9. Analyze sample (using both dried membrane and liquid sample) by direct sequence analysis or separate the peptide fragments (see Support Protocol). CLEAVAGE AT ASP-PRO PEPTIDE BONDS WITH FORMIC ACID Formic acid cleaves proteins primarily at the C-terminal side of the Asp residue in aspartic acid–proline (Asp-Pro) peptide bonds. However, secondary cleavages may occur at acid-labile bonds such as aspartic acid–threonine (Asp-Thr) and aspartic acid–serine (Asp-Ser). In addition, for proteins with multiple Asp-Pro peptide bonds, fragments generated from this reaction can be incomplete (i.e., contain internal Asp-Pro peptide bonds). Therefore, sequence analysis following dilute acid cleavage is usually accompanied by treatment with o-phthalaldehyde (OPA) to block all N-termini that may have been generated except for the N-terminal Pro. Alternately, if sample quantities allow, the fragments can be eluted from the PVDF membrane (UNIT 10.7) and fractionated prior to further characterization.
BASIC PROTOCOL 3
Materials Dried PVDF membrane containing electroblotted protein sample (UNIT 10.7) 88% formic acid (Fluka) Preconditioned polybrene-treated glass-fiber filter 45°C oven or water bath Additional reagents and equipment for analysis and separation of cleavage fragments (see Support Protocol) 1. Feather PVDF membrane containing protein sample using a clean razor blade (see Basic Protocol 1, step 1). 2. Place feathered membrane and corresponding preconditioned polybrene-treated glass-fiber filter on a sheet of Parafilm large enough to form an envelope around membrane and filter. 3. Apply 88% formic acid to sample until membrane and filter are saturated. 4. Cover sample completely with Parafilm and seal to prevent evaporation. 5. Incubate sealed packet 24 hr at 45°C in a chemical fume hood. 6. Remove sample from oven, place in chemical fume hood, and carefully cut Parafilm to expose membrane and filter to air. 7. Dry sample 5 min in a Speedvac evaporator. 8. Analyze sample using direct sequence analysis or separate the peptide fragments (see Support Protocol). CLEAVAGE AT ASN-GLY PEPTIDE BONDS WITH HYDROXYLAMINE Hydroxylamine cleaves proteins on the C-terminal side of Asn residues in asparagineglycine (Asn-Gly) peptide bonds. For proteins with multiple Asn-Gly bonds, fragments generated from this reaction can be incomplete (i.e., contain an internal Asn-Gly peptide bond). After the reaction, the PVDF membrane may be sequenced directly or the fragments eluted from the PVDF membrane (UNIT 10.7) and fractionated prior to further characterization (if multiple N-termini are expected because the protein contains more than one Asn-Gly peptide bond). Additionally, if the N-terminus of a protein is thought to be blocked (because no sequence was obtained from the first sequencing attempt although a sufficient amount of protein was analyzed), in situ hydroxylamine cleavage
BASIC PROTOCOL 4
Chemical Analysis
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can be used to confirm N-terminal blockage by directly sequencing the PVDF membrane after the cleavage reaction. Materials Dried PVDF membrane containing electroblotted protein sample (UNIT 10.7) 2 M hydroxylamine solution (see recipe) Milli-Q water or equivalent 45°C oven or water bath Additional reagents and equipment for analysis and separation of cleavage fragments (see Support Protocol) 1. Feather PVDF membrane containing protein sample using a clean razor blade (see Basic Protocol 1, step 1). 2. Place feathered PVDF membrane in clean 1.5-ml microcentrifuge tube. Working in a chemical fume hood, add 20 to 30 µl hydroxylamine solution until membrane is wet. 3. Cap tube and incubate 9 hr at 45°C in a chemical fume hood with agitation ad libitum. 4. Add 1 ml Milli-Q water to sample. Vortex vigorously. Aspirate water from membrane and discard. 5. Repeat wash two more times. Dry membrane in Speedvac evaporator or air dry. 6. Analyze sample using direct sequence analysis or separate the peptide fragments (see Support Protocol). BASIC PROTOCOL 5
CLEAVAGE AT THE N-TERMINUS OF CYS RESIDUES WITH NTCB 2-Nitro-5-thiocyanobenzoic acid (NTCB) can be used to cyanylate proteins at Cys residues; this is then followed by cleavage at a basic pH (Catsimpoolas and Wood, 1966; Jacobson et al., 1973; Degani and Patchornick, 1974; Price, 1976; Lu and Gracy, 1981; Denslow and Nguyen, 1996). The protein cleaves along the backbone, at a position that is N-terminal with respect to the modified Cys residue. The mechanism of cleavage involves the amino group of the cysteine, thereby blocking it and making it refractory to sequence analysis by N-terminal Edman chemistry. However, with the advent of MALDITOF mass spectrometry over the past few years, blockage of the amino terminus is no longer a problem. The new models of MALDI-TOF mass spectrometers, equipped with delayed extraction and reflectors, are ideal for this type of analysis (see UNIT 16.2). It is possible to treat proteins that are blotted to PVDF membranes with NTCB. Cleavage of the protein occurs readily on the membrane with an efficiency almost equal to that of cleavage in solution (UNIT 11.4; Denslow and Nguyen, 1996). A great advantage of this method is that there is minimal protein handling, since the protein is adsorbed to the membrane during all steps of the procedure. Thus, purification steps that are included in the protocol for cleavage in solution are not required for cleavage of the protein on a membrane. At the end of the procedure, the cleaved fragments are released from the membrane by acetonitrile/TFA washes, and are ready for analysis by mass spectrometry.
Chemical Cleavage of Proteins on Membranes
Materials Dried PVDF membrane containing electroblotted protein sample (UNIT 10.7). Reducing buffer (see recipe) 10 mM 2-nitro-5-thiocyanobenzoic acid (NTCB; Sigma) in 200 mM Tris-acetate, pH 8 (see recipe for Tris-acetate/EDTA but omit EDTA) continued
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Nitrogen source 10 mM MES buffer, pH 5 (see recipe) 200 mM Tris acetate/1 mM EDTA, pH 8 (see recipe) 50 mM sodium borate, pH 9 (see recipe) 60% (v/v) acetonitrile/2.5% (v/v) TFA α-Cyano-4-hydroxycinnamic acid matrix for small fragments or other appropriate matrix (also see UNIT 16.2) 40° and 50°C water baths Sonicator Additional reagents and equipment for MALDI-TOF mass spectrometry (UNIT 16.2) 1. Feather PVDF membrane by making parallel cuts with a clean razor blade (see Basic Protocol 1, step 1). Wash PVDF membrane two times over 10 min with 1 ml distilled water each time. If PVDF membrane is dry, it must be prewetted by dipping into 50% acetonitrile and rinsing twice with water.
2. Place feathered membrane in a clean 0.5-ml microcentrifuge tube and reduce the protein by adding 50 to 100 µl reducing buffer. Incubate 1 hr at 50°C. Enough reducing buffer should be added to cover the PVDF membrane.
3. Remove buffer and wash membrane twice with 0.5 ml distilled water to remove all traces of reducing agent. 4. Add 50 to 100 µl of 10 mM NTCB in 200 mM Tris-acetate, pH 8. Flush tube with N2 and seal. Incubate 20 min at 40°C. Enough NTCB solution should be added to cover the PVDF membrane.
5. Remove feathered membrane from tube and rinse one time with 0.5 ml 10 mM MES buffer, pH 5 to stop the reaction and then three times with 0.5 ml distilled water to remove NTCB and byproducts of the cyanylation reaction. 6. Place feathered membrane in a clean tube. Add 50 to 100 µl of 50 mM sodium borate, pH 9, to cleave the protein at the cyanylation sites. Incubate 1 hr at 50°C under N2. Enough sodium borate should be added to cover the PVDF membrane.
7. Rinse membrane twice with 0.5 ml distilled water to remove excess reagents. 8. Sonicate membrane 30 min with 20 µl of 60% acetonitrile/2.5% TFA to extract fragments. Remove and save the acetonitrile/TFA solution. 9. Rinse with an additional 10 µl of 60% acetonitrile/2.5% TFA. Combine with solution from step 8. 10. Mix sample with an excess of matrix of choice. For best results, sample should be at 1 to 10 pmol per µl. α-cyano-4-hydroxycinnamic acid matrix works well for most fragments. However, other matrices such as 3,5-dimethoxy-4-hydroxycinnamic acid (sinapinic acid) can be tried as well. See UNIT 16.2 for a complete procedure for this step.
11. Place sample on a sample plate and analyze by MALDI-TOF mass spectrometry (UNIT 16.2). Chemical Analysis
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SUPPORT PROTOCOL
ANALYSIS AND SEPARATION OF CLEAVAGE FRAGMENTS After performing any of the in situ cleavage reactions described in the five basic protocols, the resultant fragments can be analyzed by several different methods. The fragments can be separated by SDS-PAGE (UNIT 10.1) or two-dimensional electrophoresis (UNIT 10.4). They can then be eluted from the gel for direct sequence analysis, or electroblotted to PVDF or another suitable membrane (UNIT 10.7) for sequence analysis on the membrane. Cleaved fragments from any of the chemical cleavage protocols can also be purified by reversedphase (UNIT 11.6) or size-exclusion chromatography (UNIT 8.3) following elution from the PVDF membrane (UNIT 10.7; Szewczyk and Summers, 1988). Direct Sequence Analysis If no additional purification of the cleavage fragments is necessary, subject the membrane to N-terminal sequence analysis, with the proviso, of course, that multiple N-termini may be observed. This can be useful, however, in confirming the presence of a blocked N-terminus and may yield interpretable sequence information due to variable cleavage or sequence yields. NOTE: Regardless of the in situ cleavage method used, an ethyl acetate wash cycle should be used prior to the first sequencing cycle (Atherton et al., 1993a). For BNPS-skatole-cleaved protein fragments: Combine the dried membrane and the dried original liquid portion of the in situ digest, the latter resuspended in a minimal volume of 0.1% (v/v) TFA/50% (v/v) acetonitrile. Load the mixture onto a sequencer for N-terminal sequence analysis. The results are subject to the same caveats stated above. For proteins cleaved with formic acid at Asp-Pro bonds: Use OPA to block the N-termini of other sites that may have been generated with the formic acid (Brauer et al., 1984). Load onto a sequencer for N-terminal sequence analysis. Elution of Cleaved Fragments by a Second Round of SDS-PAGE Add 50 µl of 2% SDS/1% Triton X-100/50 mM Tris⋅Cl (pH 9.1) elution buffer to the PVDF membrane containing cleaved fragments (Szewczyk and Summers, 1988). Incubate 90 min at room temperature with occasional vortexing. Remove membrane from elution buffer and add a minimal volume of a concentrated stock of an appropriate SDS-PAGE loading buffer (Crimmins et al., 1990). Load on an SDS-polyacrylamide gel and perform electrophoresis (UNIT 10.1); if peptides <10 kDa are expected, use of Tristricine gels is recommended. Elute the cleaved fragments directly from the gel for further digestion or sequence analysis or electroblot fragments to PVDF membrane for subsequent N-terminal sequence analysis (UNIT 10.7). In addition, for larger cleavage fragments, enzymatic digestion can be performed to recover smaller fragments for N-terminal sequence analysis or mass-spectrometric analysis. An alternative elution procedure uses an alcohol/acid mixture (Aroor et al., 1994). Extract the dried membrane (see Basic Protocol 1, step 5) for 1 hr with 200 µl of 70% isopropanol/5% TFA at room temperature in a 1.5-ml microcentrifuge tube. Remove the membrane, dry the extracted supernatant, and resuspend it in 100 µl water. Precipitate fragments with 900 µl ice-cold acetone and collect the sample by centrifugation. Finally, process the pellet for gel electrophoresis.
Chemical Cleavage of Proteins on Membranes
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2; for suppliers, see SUPPLIERS APPENDIX.
Hydroxylamine solution, 2 M Weigh 5.5 g hydroxylamine⋅HCl (Fluka) into a 100-ml beaker. Slowly add 4.5 M LiOH, stirring vigorously, until hydroxylamine salt is completely dissolved. Adjust pH to 11.5 with 4.5 M LiOH. Add Milli-Q water (or equivalent) to 40 ml. Prepare immediately before use. LiOH is used instead of NaOH because it is volatile and hence more easily removed.
CNBr in 70% formic acid, 0.25 M and 1 M For 0.25 M CNBr (Basic Protocol 1): 160 µl 88% formic acid 30 µl H2O 10 µl 5 M CNBr in acetonitrile For 1 M CNBr (Alternate Protocol): 80 µl 88% formic acid 30 µl H2O 20 µl 5 M CNBr in acetonitrile Prepare solutions immediately before use. CAUTION: CNBr is extremely toxic. Therefore, all manipulations should be performed as described with extreme caution. Do not transfer dried or solubilized CNBr outside of a chemical fume hood unless it is in a sealed container.
MES buffer, pH 5, 10 mM Take 976 mg 2-(N-morpholino)ethanesulfonic acid (MES; mol. wt. 195.2). Bring the volume up to ∼470 ml with distilled water and adjust the pH to 5 with HCl. Finally, bring the volume up to 500 ml. Store up to 1 month at 4°C. Reducing buffer 969 mg Tris base (mol. wt. 121.1) 15 mg EDTA (mol. wt. 292.2) 22.92 g guanidinium⋅HCl (mol. wt. 95.5) 31 mg DTT (mol. wt. 154.3) Bring volume up to ∼35 ml with distilled H2O Adjust pH to 8 with acetic acid Bring volume up to 40 ml with H2O Prepare immediately before use. Sodium borate, pH 9, 50 mM Add 35 ml distilled water to 0.765 g sodium borate (mol. wt. 381.4), dissolve completely, then adjust pH to 9 with 2 M NaOH. Bring volume up to 40 ml. Prepare immediately before use. Tris-acetate/ EDTA, pH 8 969 mg Tris base (mol. wt. 121.1) 15 mg EDTA (mol. wt. 292.2) Bring volume to ∼35 ml with distilled H2O Adjust pH to 8 with acetic acid Bring volume up to 40 ml with H2O Store up to 1 month at 4°C Chemical Analysis
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COMMENTARY Background Information
Chemical Cleavage of Proteins on Membranes
Protein structural analysis requires the integration of peptide isolation techniques with amino acid sequence determination. A problem that frequently arises when working with small quantities of protein is that the sample can be lost between isolation and sequence determination. Transferring electrophoretically separated proteins to membrane supports through electroblotting has become a standard technique for efficient recovery of proteins from the gel matrix (UNIT 10.7; Matsudaira, 1987; LeGendre, 1990). The protocols detailed in this unit have been optimized for chemical cleavage of proteins bound to PVDF membranes. Alternatively, these protocols can be used for proteins bound to cationic-derived polyvinylidene fluoride membranes (CD-PVDF) or spotted onto glass-fiber filters. Refer to the original literature cited for each protocol for specific details regarding in situ cleavage of proteins on these membranes (Simpson and Nice, 1984; Scott et al., 1988; Miczka and Kula, 1989; Patterson et al., 1992; Landon, 1977; Bauw et al., 1988; Crimmins et al., 1990; Wadsworth et al., 1992; Kwong and Harris, 1994; Aroor et al., 1994; Denslow and Nyugen, 1996). Nitrocellulose, often used for protein binding studies, is not stable when exposed to the chemicals used in these protocols and should not be used. There are several advantages to working directly from PVDF-bound protein. Preparing the protein involves no more than electrophoresis followed by electroblotting to the PVDF membrane. PVDF-bound protein can be subjected to sequence analysis prior to chemical cleavage, which permits N-terminal sequencing. If the N-terminal appears blocked, chemical cleavage can be performed and the protein sequenced directly from the same PVDF membrane. In all likelihood, a peptide sample will contain multiple N-termini, but chemical cleavage will provide confirmation that the protein is present and may provide valuable internal sequence data. As detailed in the alternate CNBr protocol, OPA treatment (Brauer et al., 1984) can provide amino acid sequence data starting from very small sample quantities (Wadsworth et al., 1992). In situ membrane cleavage with BNPS-skatole has proven to be a valuable bioanalytical procedure. For instance, complementarity determining region II (CDR II) light chain sequence was obtained for several human oligo-
clonal antibodies (Scott et al., 1989, 1991), MEK-1 (mitogen-activated protein kinase/extracellular signal-related kinase) phosphorylation sites were determined (Gardner et al., 1994), the oligomeric contact domains of heterotrimeric G-protein were partially mapped (Vaillancourt et al., 1995; Liu et al., 1996), and the sequence of serum amyloid A protein was reported (Beach et al., 1992). Two variations of the BNPS-skatole cleavage protocol have been reported. The first (described in Bauw et al., 1988) uses minimally different cleavage conditions (80% acetic acid and a 36-hr incubation at room temperature). However, the original reaction supernatant is not saved and the PVDF is sequenced directly after digestion (see Support Protocol). This may result in lower recovery of the cleaved peptides. In the second procedure (Patterson et al., 1992), digested fragments are eluted from a CD-PVDF membrane by incubation for 1 hr at room temperature with 70% formic acid; the eluate is then combined with the initial reaction mixture supernatant prior to further analysis.
Critical Parameters The protocols described here have been performed on protein samples ranging from 1 to 100 µg, with an average of 10 µg. This quantity of protein is usually obtained from one to ten stained or unstained bands of electroblots from an SDS-PAGE minigel (UNIT 10.1) or ten to twenty spots from an electroblot of a two-dimensional gel (UNIT 10.4). Although there is no clear consensus as to how important density is to the success of cleavage, keeping the protein as concentrated as reasonable without compromising resolution is recommended. In general, either “high-retention” or “lowretention” PVDF membranes (Mozdzanowski and Speicher, 1992; UNIT 10.7) can be used for these in situ chemical cleavage protocols, although the “low-retention” type of PVDF membranes yield better peptide recoveries. CD-PVDF has also been used and may have some advantages for the successful elution of larger peptides (Patterson et al., 1992). Proteins bound to PVDF can be stained with amido black, Coomassie blue, or Ponceau S (UNIT 10.8) or radiolabeled with the isotope of choice. Proteins bound to CD-PVDF must be negatively stained, as the cationic nature of this membrane prohibits the use of standard protein stains. All cleavage solutions should be stored as recommended and made fresh immediately prior
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to use with pure, fresh reagents. A general disadvantage of any chemical cleavage procedure is that chemical fragmentation tends to be incomplete and the high concentration of reagents used may cause unwanted modification of nontarget amino acid residues in the protein. One can attempt to increase fragment yield by a combination of increasing chemical reagent concentration (with the caveat discussed above), reaction time, and temperature, and adding denaturants during the cleavage reaction. These side reactions can be minimized by performing the cleavage in complete darkness, as detailed for both the CNBr and BNPS-skatole protocols. If possible, mass spectrometry (UNIT 16.1) should be used to characterize the resulting cleavage products and will greatly aid in assessing the extent of side reactions.
Troubleshooting Consult the Troubleshooting section of UNIT for guidelines on sample preparation and optimization of the chemical cleavage reactions. It is important to include an appropriate protein standard as a positive control in any chemical cleavage experiment. See Table 11.4.1 for a list of commercially available protein standards. For Basic Protocols 1 to 4, if the protein to be cleaved has cystine (Cys) residues that are involved in disulfide bonds, it is strongly suggested that the sample be reduced and alkylated prior to cleavage (UNIT 11.4, see Troubleshooting). Reduction and alkylation should be performed in solution and the sample desalted using either an Applied Biosystems Prospin or Problot PVDF cartridge or a Millipore PVDF sample clean-up cartridge. This allows transferral of protein to PVDF using centrifugation while removing salts and other contaminants that accumulate during sample preparation or modification. Alternately, reduction and alkylation can be performed directly on PVDF (Atherton et al., 1993b) or glass-fiber filters (Andrews and Dixon, 1987). For Basic Protocol 5, the predominant reaction is the cyanylation of reduced Cys residues in the first step of the reaction followed by cleavage (N-terminal to the Cys) of the protein under alkaline conditions. Several side reactions, however, are also possible, including βelimination of the thiocyanoalanine residue instead of cyclization and cleavage (Degani and Patchornick, 1974) and the formation of mixed disulfides (Price, 1976). The side reactions are 11.4
an annoyance but do not interfere with the analysis, especially if the protein sequence is known and expected fragment sizes can be predicted. For a protein whose sequence is not known, the side reactions become more significant. For the most part, these side reactions can be minimized by keeping the ratio of NTCB to protein in the 10-fold molar range, and by acidifying the reaction mixture promptly after cyanylation to inhibit disulfide exchange. Finally, the excess NTCB should be removed prior to the cyclization step. If the object of the experiment is to map disulfides in the protein backbone, then the protein sample should be divided into two aliquots, one that is treated with disulfide reagent to reduce all the Cys residues, and another aliquot that is not treated, but rather is kept in the native state. In this case, nonreducing SDSPAGE (i.e., with gel buffers lacking reducing agents) should be performed. Remember that oxidized Cys residues will not react with NTCB. When working with PVDF membranes, it is always advisable to wear talc-free gloves at all times. Please consult UNIT 10.7 for additional troubleshooting information.
Anticipated Results Successful cleavage and recovery of fragments for primary structural analysis can be performed with a little as 5 µg of pure protein. However, it is impossible to overemphasize the importance of having a pure protein for obtaining unambiguous protein sequence data. It should be emphasized that chemical cleavages are often incomplete reactions, and typically generate larger peptide fragments than those obtained with enzymatic digestion. Therefore, they may not be the optimal methods for obtaining a definitive peptide map, because neither cleavage nor recovery is 100%. As a rule of thumb, an initial protein sample of 100 pmol should yield 25 pmol of a resultant peptide on PVDF membranes after electrophoresis and electroblotting (assuming a 50% efficiency for both cleavage and electrotransfer). Of course, recovery can be higher or lower, but it’s best to start with at least this much protein. We have found that for the larger fragments generated using chemical in situ cleavage protocols, the efficiency of elution from PVDF is generally better for the less sensitive and more weakly bound Ponceau-S-stained bands than with other methods. Also, if the cleaved fragments will be subjected to an additional purification step, Ponceau-S-stained protein samples are preferred. If the additional round of purifi-
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cation is gel electrophoresis, the type of stain apparently does not matter, and all are equally eluted from the PVDF (Szewczyk and Summers, 1988) into the gel.
Time Considerations In general, two days is sufficient for the completion of both the cleavage and elution of fragments for further characterization. Once the PVDF-bound protein is cleaved and dried, it can be stored at −20°C for a short period (several days) until elution or sequence analysis is performed. However, excess reagent remaining on the PVDF membrane may cause unwanted modification of the protein. Therefore, the fragments should be eluted or the PVDF mixture analyzed as soon as possible following cleavage.
Literature Cited Andrews, P.C. and Dixon, J.E. 1987. A procedure for in situ alkylation of cystine residues on glass fiber prior to protein microsequence analysis. Anal. Biochem. 161:524-528. Aroor, A.R., Denslow, N.D., Singh, L.P., O’Brien, T.W., and Wahba, A.J. 1994. Phosphorylation of rabbit reticulocyte guanine nucleotide exchange factor in vivo. Identification of putative casein kinase II phosphorylation sites. Biochemistry 33:3350-3357. Atherton, D., Fernandez, J., DeMott, M., Andrews, L., and Mische, S.M. 1993a. Routine protein sequence analysis below ten picomoles: One sequencing facility’s approach. In Techniques in Protein Chemistry (R.H. Angeletti, ed.) pp. 409418. Academic Press, San Diego. Atherton, D., Fernandez, J., and Mische, S.M. 1993b. Identification of cysteine residues at the 10 pmol level by carboxamidomethylation of proteins bound to sequencer membrane supports. Anal. Biochem 212:98-105. Bauw, G., Van den Bulcke, M., Van Damme, J., Puype, M., VanMontagu, M., and Vandekerckhove, J. 1988. NH2-terminal and internal microsequencing of proteins electroblotted on inert membranes. In Methods in Protein Sequence Analysis (B. Wittmann-Liebold, ed.), pp. 220233, Springer-Verlag, Berlin. Beach, C.M., DeBeer, M.C., Sipe, J.D., Loose, L.D., and DeBeer, F.C. 1992. Human serum amyloid A protein. Complete amino acid sequence of a new variant. Biochem. J. 282:615-620. Brauer, A.W., Oman, C.L., and Margolies, M.N. 1984. Use of o-phthalaldehyde to reduce background during automated Edman degradation. Anal. Biochem. 137:134-142.
Chemical Cleavage of Proteins on Membranes
Catsimpoolas, N. and Wood, J.L. 1966. Specific cleavage of cystine peptides by cyanide. J. Biol. Chem. 241:1790-1796. Crimmins, D.L., McCourt, D.W., Thoma, R.S., Scott, M.G., Macke, K., and Schwartz, S.D.
1990. In situ chemical cleavage of proteins immobilized to glass fiber and polyvinylidenedifluoride membranes: Cleavage at tryptophan residues with (2-(2′-nitrophenylsulfonyl)-3methyl-3-bromoindolenine) to obtain internal amino acid sequence. Anal. Biochem. 187:27-38. Degani, Y. and Patchornick, A. 1974. Cyanylation of sylfhydryl groups by 2-nitro-5-thiocyanobenzoic acid. High-yield modification and cleavage of peptides at cysteine residues. Biochemistry 13:1-11. Denslow, N.D. and Nguyen, H.P. 1996. Specific cleavage of blotted proteins at cysteine residues after cyanylation: Analysis of products by MALDI-TOF. In Techniques in Protein Chemistry VII (D.R. Marshak, ed.) pp. 241-248. Academic Press, San Diego. Gardner, A.M., Vaillancourt, R.R., Lange-Carter, C.A., and Johnson, G.J. 1994. MEK-1 phosphorylation by MEK kinase, Raf, and mitogen-activated protein kinase: Analysis of phosphopeptides and regulation of activity. Mol. Biol. Cell 5:193-201. Jacobson, G.R., Schaffer, M.H., Stark, G.R., and Vanaman, T.C. 1973. Specific chemical cleavage in high yield at the amino peptide bonds of cysteine and cystine residues. J. Biol. Chem. 248:6583-6591. Kwong, M.Y. and Harris, R.J. 1994. Identification of succinimide sites in proteins by N-terminal sequence analysis after alkaline hydroxylamine cleavage. Protein Sci. 3:147-149. Landon, M. 1977. Cleavage at aspartyl-prolyl bonds. Methods Enzymol. 47:145-149. LeGendre, N. 1990. Immobilon P transfer membrane: Applications, utility and protein biochemical analyses. BioTechniques 9:788-805. Liu, Y., Arshavsky, V.Y., and Ruoho, A.E. 1996. Interaction sites of the COOH-terminal region of the gamma subunit of cGMP phosphodiesterase with the GTP-bound alpha subunit of transducin. J. Biol. Chem. 271:26900-26907. Lu, H.S. and Gracy, R.W. 1981. Specific cleavage of glucosephosphate isomerases at cysteinyl residues using 2-nitro-5-thiocyanobenzoic acid: Analyses of peptides eluted from polyacrylamide gels and localization of active site histidyl and lysyl residues. Arch. Biochem. Biophys. 212:347-359. Matsudaira, P. 1987. Sequence from picomole quantities of proteins electroblotted onto polyvinylidene difluoride membranes. J. Biol. Chem. 262:10035-10038. Miczka, G. and Kula, M.R. 1989. The use of polyvinylidene difluoride membranes as blotting matrix in combination with sequence: Application to pyruvate decarboxylase from Zymomonas mobilis. Anal. Lett. 22:2771-2782. Mozdzanowski, J. and Speicher, D.W. 1992. Microsequence analysis of electroblotted proteins. I. Comparison of electroblotting recoveries using different types of PVDF membrane. Anal. Biochem. 207:11-18.
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Patterson, S.D., Hess, D., Yungwirth, T., and Aebersold, R. 1992. High-yield recovery of electroblotted proteins and cleavage fragments from a cationic polyvinylidene fluoride-based membrane. Anal. Biochem. 202:193-203. Price, N.C. 1976. Alternative products in the reaction of 2-nitro-5-thiocyanatobenzoic acid with thiol groups. Biochem. J. 159:177-180. Scott, M.S., Crimmins, D.L., McCourt, D.W., Tarrand, J.J., Eyerman, M.C., and Nahm, M.H. 1988. A simple in situ cyanogen bromide cleavage method to obtain internal amino acid sequence of proteins electroblotted to polyvinylidenedifluoride membranes. Biochem. Biophys. Res. Commun. 155:1353-1359. Scott, M.G., Crimmins, D.L., McCourt, D.W., Zocher, I., Thiebe, R., Zachau, H.G., and Nahm, M.H. 1989. Clonal characterization of the human IgG antibody repertoire to Haemophilus influenzae type b polysaccharide. III. A single VkII gene and one of several JK genes are joined by an invariant arginine to form the most common L chain V region. J. Immunol. 143:4110-4116. Scott, M.G., Crimmins, D.L., McCourt, D.W., Chung, G., Schable, K.F., Thiebe, R., Quenzel, E.-M., Zachau, H.G., and Nahm, M.H. 1991. Clonal characterization of the human IgG antibody repertoire to Haemophilus influenzae type b polysaccharide. IV. The less frequently expressed VL are heterogenous. J. Immunol. 147:4007-4013. Simpson, R.J. and Nice, E.C. 1984. In-situ cyanogen bromide cleavage of N-terminally blocked proteins in a gas-phase sequencer. Biochem. Int. 8:787-791. Szewczyk, B. and Summers, D.F. 1988. Preparative elution of proteins blotted to immobilon membranes. Anal. Biochem. 168:48-53. Vaillancourt, R.R., Dhanasekaran, N., and Ruoho, A.E. 1995. The photoactivatable NAD+ analogue [32P]2-azido-NAD+ defines intra- and inter-molecular interactions of the C-terminal domain of G-protein G alpha t. Biochem. J. 311:987-993. Wadsworth, C.L., Knuth, M.W., Burrus, L.W., Olwin, B.B., and Niece, R.L. 1992. Reusing PVDF electroblotted protein samples after N-terminal sequencing to obtain unique internal amino acid sequence. In Techniques in Protein Chemistry III (R.H. Angeletti, ed.) pp. 61-68. Academic Press, San Diego.
Key References Fontana, A. and Gross, E. 1986. Fragmentation of polypeptides by chemical methods. In Practical Protein Chemistry: A Handbook (A. Darbre, ed.) pp. 68-120. John Wiley & Sons, Chichester, U.K., and New York. Excellent source of chemical cleavage procedures. Matsudaira, P. (ed.) 1989, 1993. A Practical Guide to Protein Purification, Editions 1 and 2. Academic Press, San Diego. Various protocols for PVDF-bound proteins are described. Angeletti, R.H. (ed.) 1992, 1993. Techniques in Protein Chemistry, Vols. III and IV. Academic Press, San Diego. Crabb, J. (ed.) 1994, 1995. Techniques in Protein Chemistry, Vols. V and VI. Academic Press, San Diego. Hugli, T. (ed.) 1989, 1991. Techniques in Protein Chemistry, Vols. I and II. Academic Press, San Diego. Marshak, D. 1996. 1997. Techniques in Protein Chemistry, Vol. VII and VIII. Academic Press, San Diego. Villafranca, J. (ed.) 1990. Current Protocols in Protein Chemistry: Technique, Structure and Function. Academic Press, San Diego. The above five references are superb compendia of selected, top-notch papers (presented at the 2nd through 9th Symposia of the Protein Society) related to all areas of protein chemistry, including various solid-phase PVDF membrane applications. Smith, B. 1994. Chemical cleavage of proteins. In Methods in Molecular Biology, Vol. 32: Basic Protein and Peptide Protocols (J. Walker, ed.) pp. 297-309. Humana Press, Towata, NJ. Valuable, up-to-date summary of several chemical cleavage procedures.
Contributed by Dan L. Crimmins Washington University School of Medicine St. Louis, Missouri Sheenah M. Mische Boehringer Ingelheim Pharmaceuticals Ridgefield, Connecticut Nancy D. Denslow University of Florida Gainesville, Florida
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Reversed-Phase Isolation of Peptides
UNIT 11.6
Reversed-phase high-performance liquid chromatography (HPLC) is a fundamental tool for the isolation and analysis of peptides. Peptides are separated on a hydrophobic stationary phase and eluted with a gradient of increasing organic solvent concentration. Peptides in 5- to 500-pmol quantities are separated on a narrow-bore (2-mm-i.d.) or microbore (1-mm-i.d.) column (see Basic Protocol 1). Separation of peptides in quantities <5 pmol can be accomplished using capillary HPLC columns (see Basic Protocol 2). Capillary HPLC columns, however, require a gradient flow rate of 3 to 5 µl/min, which most current HPLC pumps cannot attain without modifications. A procedure is therefore provided for constructing a capillary HPLC system using readily available components (see Support Protocol). HPLC peaks that appear to be symmetrical may actually contain coeluting peptides. Matrix-assisted laser desorption/ionization (MALDI) mass spectrometry (see Basic Protocol 3 and Alternate Protocol; also see UNITS 16.1 & 16.2) and capillary electrophoresis (see Basic Protocol 4; also see UNIT 10.9) can be used to analyze a small portion of an HPLC fraction and to determine the number of components present in a small sample. These methods can be utilized to screen fractions prior to automated sequencing. REVERSED-PHASE PEPTIDE SEPARATION AT THE 5- TO 500-PMOL LEVEL
BASIC PROTOCOL 1
A 2-mm narrow-bore column requires a flow rate of 100 to 200 µl/min. Many commercial HPLC instruments are capable of providing flow rates in this range, although some have tubing with a large internal volume that may add excessive delay time to the separation. Some HPLC instruments that are suitable without modification for 1- and 2-mm columns are listed in the materials section below. Many older HPLC instruments can be utilized for 2-mm columns, but most cannot reproducibly deliver the 50- to 100-µl/min flow rates required by 1-mm columns. Materials Mobile-phase solvent A: 0.1% (v/v) trifluoroacetic acid (TFA; Pierce) in Milli-Q water (TFA sold for protein sequencing may not be suitable because some manufacturers add antioxidants that can generate artifacts) Mobile-phase solvent B: 0.07% to 0.1% (v/v) TFA (Pierce) in acetonitrile or 1- or 2-propanol (Burdick & Jackson or Baker; HPLC-grade) Solvent modifier: TFA (Pierce) Milli-Q grade water or equivalent (distilled water is not suitable) Peptide sample HPLC peptide standards, commercial (e.g., PE Biosystems) or tryptic digest (see Troubleshooting), for testing column HPLC system (e.g., Hewlett-Packard HP-1090 liquid chromatograph; PE Biosystems model 170A; Michrom BioResources Ultrafast Microprotein Analyzer; Beckman System Gold; Waters Alliance System) Detector flow cell (see recipe) 2-channel strip-chart recorder (Kipp & Zonen or equivalent) C18, C8, or C4 reversed-phase columns, 300 Å, 1- or 2-mm i.d. (e.g., SynChrom or Vydac; many other columns from numerous manufacturers can be used)
Chemical Analysis Contributed by William J. Henzel and John T. Stults Current Protocols in Protein Science (2001) 11.6.1-11.6.16 Copyright © 2001 by John Wiley & Sons, Inc.
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1. Prepare and degas the mobile phase and set up an appropriate gradient of mobilephase solvents A and B. Select a gradient based on number of peaks expected and resolution desired. Normal gradient: 0% to 70% solvent B in 70 min Fast gradient: 0% to 70% solvent B in 35 min Slow gradient: 0% to 70% solvent B in 90 to 140 min. A normal, fast, or slow gradient should be used when the expected number of peaks is ∼30 to 60, <30, or >60, respectively.
2. Program flow rate to 200 µl/min or 100 µl/min for 2-mm- or 1-mm-i.d. columns, respectively. Set detector to 0.1 and 0.02 AUFS and the wavelengths on chart recorder to 214 and 280 nm for channels 1 and 2, respectively. Equilibrate column with initial conditions and establish a flat baseline. The 280-nm pen on the chart recorder should be set to a scale five times more sensitive than the 214-nm pen. If only one wavelength is available, use 214 nm.
3. Fill injector loop with starting solvent and run a blank gradient consisting of the same gradient as will be utilized for separation of sample. This will wash the column, removing any peptides that may be present from previous injections.
4. Prepare sample for injection as follows (or prepare peptide standards, when using a new column or when troubleshooting): a. If the sample contains organic solvent, lower the organic solvent concentration to <10% (v/v) by evaporating it in a Speedvac evaporator or by diluting it with water or aqueous buffer. b. Check that the sample volume is appropriate for the volume of the injector sample loop; if not, either reduce the sample volume by evaporation or change to a larger sample loop. c. Spin the sample in a microcentrifuge to remove particulates prior to injection. The presence of organic solvent in the sample may prevent peptides from adsorbing to the stationary phase. The size of the sample loop will not have a significant effect on the separation. However, if a larger loop is used, an appropriate delay should be added to the gradient program to allow the entire sample volume to load on the column and to allow salts that may be in the sample to wash off the column before the gradient begins.
5. Change injector to load position and rinse injector with starting solvent (mobile-phase solvent A). If the sample loop was switched from inject to load position during the gradient run, the sample loop will contain organic solvent. If the sample only partially fills the sample loop, the presence of organic solvent in the loop might prevent the sample from binding to the column; rinsing with starting solvent averts this.
6. Inject sample, being careful to avoid injecting air into sample loop. Air that is injected onto the column can produce air bubbles that may lodge in the detector.
7. Prepare a rack with 1.5-ml microcentrifuge tubes for collecting peak fractions. Number tubes with felt-tipped pen. Reversed-Phase Isolation of Peptides
8. Calculate the delay for peak collection by dividing the volume of tubing exiting the flow cell by the flow rate.
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Table 11.6.1
Volumes of Tubing Exiting Flow Cell
Tubing inner diameter
Volume (µl/cm)
0.075-mm (75-µm) FSC 0.005-in. (127-µm) PEEK
0.045 0.127
0.007-in. (178-µm) PEEK
0.249
This is particularly important to prevent miscollection of peaks with low flow rates (<200 ìl/min). Table 11.6.1 provides a list of volumes for different diameters of PEEK and FSC tubing. Powder-free gloves should be worn during peak collection to avoid sample contamination by free amino acids.
9. Collect peaks by monitoring absorbance change on a strip-chart recorder and allowing appropriate delay time (calculated in step 8) before changing tubes. A stopwatch can be used to accurately measure the delay time when collecting poorly resolved peaks. Data systems on most HPLC systems have signal processing delays that can make manual peak collection difficult. Monitoring a strip-chart recorder provides a real-time measurement of absorbance and eliminates this problem.
REVERSED-PHASE PEPTIDE SEPARATION AT ≤5 PMOL Separation of peptides that are present in quantities ≤5 pmol requires the use of a capillary column. These columns have internal diameters <1 mm and provide high-resolution peptide mapping of peptides collected from in-gel or in situ digests of proteins electroblotted from one- and two-dimensional gels. A 0.32-mm-i.d. capillary column is capable of separating peptides at the 500-fmol level, but requires a flow rate of 3 to 5 µl/min. Several commercial HPLC systems are currently available that are capable of delivering gradient flow rates in this range; other HPLC pumps can be adapted to deliver capillary gradient flow rates using a flow splitter (see Support Protocol).
BASIC PROTOCOL 2
Materials Solvent A: 0.1% (v/v) trifluoroacetic acid (TFA) in Milli-Q water Solvent B: 0.05% to 0.1% (v/v) TFA in acetonitrile Peptide sample Detector flow cell (LC Packings) Strip-chart recorder (Kipp & Zonen or equivalent) Capillary HPLC system (see Support Protocol) Graduated 10-µl Hamilton glass syringe connected to outlet of flow cell with 0.25-mm-i.d. Teflon tubing (LC Packings) Two stopwatches Additional reagents and equipment for reversed-phase peptide separation at 5 to 500 pmol (see Basic Protocol 1) and capillary HPLC system assembly (see Support Protocol) 1. Prepare and degas the mobile phase (see Basic Protocol 1, step 1). 2. Set detector to 0.1 AUFS and the wavelength on the chart recorder to 195 nm. 3. Equilibrate capillary column in solvent A until a flat baseline is established. Connect a graduated 10-µl Hamilton glass syringe to the outlet tubing of capillary flow cell with 0.25-mm-i.d. Teflon tubing. Measure the time it takes for liquid to reach a given volume and calculate flow rate as follows: flow rate (in µl/min) = volume/time.
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4. Accurately measure the length of tubing from the flow cell to its outlet end and calculate the delay according to the following formula: delay (min) = volume of outlet tubing (µl)/flow rate (µl/min). See Table 11.6.1 for a list of tubing internal volumes. A typical delay time is 35 sec.
5. If necessary, adjust the flow rate to between 3 and 5 µl/min (see Support Protocol, step 1). 6. Run blank gradient, prepare sample, and inject (see Basic Protocol 1, steps 3 to 6). 7. Collect peaks in 0.5-ml microcentrifuge tubes using two stopwatches. Mark the start of the peak with one stopwatch and time the end of the peak with the second watch. Peak collection is the most difficult aspect of preparative capillary HPLC and requires a little practice. However, the degree of peak purity will be surprisingly high for closely eluting peaks if the delay time is calculated accurately. With Z-shaped flow cells, the flow cell is a length of fused silica capillary tubing. Very minor resolution loss occurs as the peaks travel from the end of the column to the outlet of the detector tubing. Only one stopwatch will be needed if the capillary flow cell is modified as described for capillary HPLC assembly (see Support Protocol, step 3b). SUPPORT PROTOCOL
CAPILLARY HPLC SYSTEM ASSEMBLY Peptide separations at the low picomole level require columns with an i.d. of <1 mm, which are called capillary columns. A 0.32-mm-i.d. capillary column is capable of separating peptides at the 500-fmol level. However, these columns require a flow rate of 3 to 5 µl/min, and at present only a few commercial HPLC systems can deliver gradient flow rates in this range without modifications. Using the material listed below, it is quite easy to construct or modify an existing HPLC using readily available components. PE Biosystems, Waters Associates, Micro-Tech Scientific, and Michrom BioResources have capillary HPLC pumps. Materials Capillary flow cell (LC Packings) Piston- or syringe-pump splitter (LC Packings) or laboratory-constructed splitter: 1/16 in. tee (Valco); fingertight fittings; 0.012-in.-i.d. sleeve for capillary tubing; fingertight fitting for 0.012-in. sleeve (Upchurch Scientific); and 150 cm of 0.05-mm-i.d., 300-mm-o.d. FSC tubing (Polymicro Technologies) or 0.250-mm-i.d. Teflon tube inserted into a PEEK fingertight nut and ferrule Capillary pump (PE Biosystems 140-D) or HPLC piston pump (see Basic Protocol 1) for use with an LC Packings splitter, or HPLC syringe pump (PE Biosystems models 120A, 140A, or 170A) for use with a laboratory-constructed splitter Injector equipped with a 20-µl loop (Rheodyne) In-line precolumn filter (Upchurch Scientific) 0.32-mm × 15-cm C18 column (LC Packings, Keystone Scientific, Metachem Technologies, Micro-Tech Scientific, or Michrom BioResources) 1a. If using a capillary pump that can provide a flow rate of 3 to 5 ìl/min: Proceed to step 3a or 3b. 1b. If assembling a splitter: Connect a 150-cm length of 0.05-mm-i.d. FSC tubing to Valco tee with a 0.012-in.-i.d. sleeve or with a 0.250-mm-i.d. Teflon tube inserted into a PEEK fingertight nut and ferrule. Connect the inlet side of splitter to HPLC pump with 0.005-in.-i.d. PEEK tubing.
Reversed-Phase Isolation of Peptides
A schematic diagram of the capillary fittings and the splitter assembly is shown in Figure 11.6.1.
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A splitter must be connected to the HPLC pump to allow the flow rate to be reduced from 50 to 200 ìl/min to the 3 to 5 ìl/min required for a 0.32-mm-i.d. column. A splitter can be purchased commercially or assembled as described. The split ratio is controlled by varying the flow rate from the pump or by adjusting the length of the 0.05-mm-i.d. capillary tubing connected to the Valco tee. The pump flow rate should be set to the lowest flow that the pump can reliably deliver. A homemade splitter requires a pulse-free pump that can reliably deliver a flow rate of <200 ìl/min. PE Biosystems syringe pumps can accurately deliver a pulse-free flow rate of 70 ìl/min. These pumps have a dynamic mixer and a static mixer,
A to waste
fused silica tubing (50-µm i.d., 300-µm o.d.)
Teflon tube (0.25-mm i.d., 1/16-in o.d.)
or
to waste
capillary sleeve
PEEK fitting and ferrule
to capillary column
from pump tee
B from capillary column
UV
detector fused silica 0.075-mm i.d.
Teflon sleeve 0.025-mm i.d.
Figure 11.6.1 Modifications for capillary HPLC system. (A) Assembly of capillary fittings and splitter. (B) Modification of U or Z flow cell.
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splitter
solvent A solvent B
sample injector UV detector
HLPC pump precolumn filter waste
capillary column
microcentrifuge tube
Figure 11.6.2 Capillary HPLC system.
which are removed to reduce the gradient delay caused by their large internal volumes. The high-pressure outlet of the mixing tee is connected directly to the splitter. Figure 11.6.2 shows a schematic diagram of the capillary HPLC system.
2. Connect outlet of splitter to injector and connect the injector to the in-line precolumn filter with 0.005-in.-i.d. PEEK tubing. The precolumn filter protects the capillary tubing and the column from clogging. The injector is fitted with a 20-ìl sample loop. Any size sample loop can be used; however, a large loop will require a long wait for the sample to load on a 0.32-mm-i.d. column at a flow rate of 3 to 5 ìl/min.
3a. Install capillary flow cell. The small peak volumes of capillary HPLC prevents the use of conventional flow cells. Instead, a length of capillary tubing with a short section of its polyamide coating removed is used as a flow cell. LC Packings has developed a capillary tube that is bent in the shape of a U or Z. The UV beam is aligned with the short axis of the U, increasing the path length of the flow cell. Although these flow cells have shorter path lengths than many conventional ones, the short path length and thin walls of the capillary tubing allow peptide detection at 195 nm.
3b. Optional: Modify the capillary flow cell (Fig. 11.6.1B) to reduce the delay volume by carefully cutting the 0.075-mm-i.d. outlet tubing of the flow cell, leaving ~1 to 2 cm adjacent to the Z cell (the fused silica tubing can be easily cut with a ceramic tubing cutter). Connect a 30-cm length of 0.025-mm-i.d., 0.280-mm-o.d. fused silica tubing with a 0.25-mm-i.d. Teflon sleeve to the remaining short outlet of the Z cell. The connection should be made inside the detector housing and the glass capillary tubing must be touching each other to avoid introducing any dead volume. This modification eliminates the need for two stopwatches for peak collection. The flow cell modification reduces the delay volume to 0.45 ìl, corresponding to a delay of 6 sec for a flow rate of 3.5 ìl/min. The short delay greatly facilitates hand collection of HPLC fractions and the small-i.d. outlet tubing provides backpressure that prevents bubble formation in the flow cell.
4. Connect the 0.32-mm-i.d. C18 capillary column to the sample prefilter with the 1⁄16-in. PEEK fitting supplied with the column. Connect outlet tubing (75-µm-i.d.) of capillary column to capillary flow cell with a short length of 0.25-mm-i.d. Teflon tubing, also supplied with the column. The 0.32-mm × 15-cm C18 column from LC Packings is the best general-purpose column for tryptic peptides. Keystone Scientific, Metachem Technologies, and Micro-Tech Scientific also sell capillary columns.
Reversed-Phase Isolation of Peptides
The Teflon tubing slips over the ends of the capillary tubing, creating a zero-dead-volume union between the column outlet and the detector union. Resolution will be significantly diminished if a gap is present between the adjacent pieces of capillary tubing.
5. Prepare the capillary HPLC system for sample run (see Basic Protocol 2).
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PEPTIDE MAPPING BY MATRIX-ASSISTED LASER DESORPTION/IONIZATION (MALDI) MASS SPECTROMETRY
BASIC PROTOCOL 3
Protein digests are mass analyzed either directly as an unfractionated mixture or as individual fractions from reversed-phase HPLC. The peptide-containing solution is mixed with a UV-absorbing matrix and applied to the sample stage of the mass spectrometer. Crystals of the peptide/matrix mixture form on the stage. This stage is inserted into the time-of-flight mass spectrometer vacuum system and irradiated with a UV laser to obtain a spectrum (see UNITS 16.1 & UNIT 16.2). Greater mass accuracy is achieved by addition of known peptide(s) as internal mass standards. Alternate protocols are found in UNIT 16.3. Materials Peptide sample 30% (v/v) acetonitrile/0.1% (v/v) trifluoroacetic acid (TFA) MALDI matrix solution (see recipe) Mass standard solution (see recipe) Time-of-flight mass spectrometer (e.g., PE Biosystems, Amersham Pharmacia Biotech, Micromass, Brucker, or Kratos) 1. Dilute samples in 30% acetonitrile/0.1% TFA to a final concentration of 0.05 to 5 pmol/µl. Salt and buffer concentrations should be <50 mM. Most detergents, especially SDS, are not tolerated. If necessary, reduce the ionic strength by dilution, HPLC (UNIT 8.7), or size-exclusion chromatography (UNIT 8.3). If volatile buffers have been used, they may be removed by lyophilization and the dried sample redissolved in 2% TFA/50% acetonitrile.
2. Mix 2 µl of sample with 2 µl MALDI matrix solution in a 0.5-ml microcentrifuge tube. 3. Apply 1 to 2 µl of the mixture to the sample target, depending on the target size. Allow sample to dry completely and crystals to form without heating or applying vacuum. Alternatively, 0.5 ìl of sample solution and 0.5 ìl of matrix solution may be mixed directly on the target. The solutions should be mixed well by repeatedly drawing the liquid into a pipettor. The α-cyano-4-hydroxycinnamic acid (4HCCA) matrix solution works well for most peptides. An alternative is 2,5-dihydroxybenzoic acid (DHB). Another alternative is 3,5-dimethoxy-4-hydroxy-trans-cinnamic acid (sinapinic acid), but this produces a large background below 2000 Da and is not recommended for smaller peptides. Sinapinic acid or 4HCCA work best for larger peptides (>3000 Da). Due to preferential ionization of particular peptides in mixtures, it is advisable to obtain spectra for peptide mixtures with more than one matrix.
4. Insert sample into vacuum chamber of the mass spectrometer and follow the mass measurement procedure recommended by the instrument’s manufacturer. Laser intensity (fluence) should be set just above the threshold for appearance of ion intensity. The observed signal (m/z) corresponds to M+H+. Larger peptides may yield some (M+2H)2+, which will appear at half the peptide mass (M/2). At high laser fluence, dimers may be formed as artifacts of the ionization process. Peptide mixtures may display preferential ionization for some peptides, and the relative peptide levels may vary from crystal to crystal. For this reason, spectra should be acquired from several different crystals on the target to obtain a representative display of peaks. The relative peak abundances in some cases (but not always!) give a semiquantitative image of the relative concentrations of peptides in the mixture, but they should not be relied upon for quantitation. Additional spectra acquired from the peptide mixture in a different matrix sometimes provide signals for peptides with suppressed ionization.
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5. Calibrate the mass spectrometer with a mixture of peptides of known mass. Obtain calibrant peptide mass spectra as described in steps 2 to 4. Calibrate the instrument according to the manufacturer’s recommendation. An equimolar solution (∼1 pmol/ìl of each component) of des-Arg-bradykinin (e.g., Sigma) and ACTH-CLIP (18-39) provides two peaks that cover the masses of most peptides. The centroid of the peak corresponds to the isotopically averaged mass and should be used for calibration and mass assignment. The average masses (M+H+) for des-Arg-bradykinin (905.1 Da) and ACTH-CLIP (18-39) (2466.7 Da) are generally reliable as calibrants. When sufficient resolution is available, the monoisotopic peaks should be used for calibration: des-Arg-bradykinin 904.468, ACTH-CLIP (18-39) 2465.199. Additional calibrant masses are found in Table 16.2.2. For higher mass peptides bovine insulin (M+H+ = 5734.5) may be used for calibration. Mass accuracy is best when the calibration peaks are close in mass and abundance to the unknown sample being analyzed. The laser fluence should be constant for the calibrant and unknown sample spectra.
6a. If >500 fmol of unknown peptide sample is available: Mix an equimolar concentration of des-Arg-bradykinin/ACTH-CLIP (18-39) standard solution with the unknown sample. Repeat mass analysis of sample with internal mass standards as described in steps 2 to 4. 6b. If no additional unknown peptide sample is available: Redissolve sample and matrix remaining on target after analysis in step 4 with 1 µl of 2% TFA/50% acetonitrile. Add an equimolar concentration of mass standard solution. Allow mixed sample to dry on sample target and repeat mass analysis as described in step 4. Internal mass calibration provides greater mass accuracy than external calibration. The amount of internal standard may need to be varied in several samples in order to obtain peak intensities that are approximately the same for the standard and the unknown sample. Avoid adding an excess of internal standard, which can suppress ionization of the unknown sample. ALTERNATE PROTOCOL
PEPTIDE MAPPING BY MALDI MASS SPECTROMETRY USING THE MATRIX FAST-EVAPORATION METHOD Preparation of a thin layer of matrix onto which the sample is applied offers advantages for small quantities of peptides and for contaminated samples. Matrix is applied to the sample stage in a highly volatile solvent that leaves a thin layer of matrix after evaporation. The uniform layer yields more uniform, intense signal across the sample. The sample is easily washed for efficient removal of contaminating salts and other buffer components that may interfere with ionization. Nitrocellulose is added to the matrix to provide greater peptide binding during the rinse step and to give greater structural support to the matrix, especially in the presence of higher concentrations of organic solvent that readily dissolve the matrix. This method is applicable to any type of MALDI instrument and gives superior results for a broad range of peptide and protein samples. Additional Materials (also see Basic Protocol 3) Fast-evaporation MALDI matrix solution (see recipe) 10% and 0.1% formic acid 1. Apply 0.5 to 2.0 µl fast-evaporation MALDI matrix solution to a clean sample target. The solution must be applied quickly due to the high volatility of the acetone. A uniform, white surface 3 to 5 mm in diameter will form in 3 to 5 sec.
2. Apply 0.3 to 2.0 µl peptide sample to target surface. Allow sample to dry completely. Reversed-Phase Isolation of Peptides
The matrix dissolves easily in basic solution or in the presence of a high concentration of organic solvent (e.g., >30% acetonitrile). If the peptide sample is not already acidic, apply
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1.0 ìl of 10% formic acid to the target before applying sample. For samples that contain high concentrations of organic solvent, apply 1.0 ìl of water to the target before applying the sample.
3. Rinse sample with 4.0 µl cold 0.1% formic acid. After 10 sec, carefully remove the solution from the target surface with a pipet, or blow solution off target with a stream of nitrogen. Repeat rinse. Rinse step may be eliminated if the peptide sample is free of salts. If the salt content of the sample is unknown, try to obtain signal without rinsing. If the results are unsatisfactory, remove the sample from the mass spectrometer and perform the rinse steps.
4. Insert sample into vacuum chamber of the mass spectrometer and follow the mass measurement procedure recommended by the instrument’s manufacturer (see Basic Protocol 3). Best results are obtained when samples are analyzed immediately. The authors have observed a decrease in signal intensity when samples are stored >24 hr. They have not observed any decrease in signal when samples are prepared according to Basic Protocol 3 using α-cyano-4–hydroxycinnamic acid (4HCCA).
5. Calibrate the mass spectrometer with a mixture of peptides of known mass (see Basic Protocol 3). Because the peptide sample often covers only a part of the matrix spot, a small volume (0.1 to 0.3 ìl) of the mass standard solution may be applied to the side of the matrix spot (after the peptide sample is dried on the other side of the matrix spot and rinsed) to provide a pseudo-internal standard. The mass spectrum of this small calibration sample may be taken immediately before or after the peptide spectrum to provide an external calibration with high accuracy. Alternatively, the signal from this sample may be included in the sample spectrum to provide internal calibration without the risk of ion suppression by the calibration compounds.
CAPILLARY ELECTROPHORESIS ANALYSIS Capillary electrophoresis (also see UNIT 10.9) separates peptides by electrophoretic migration and electroosmotic flow using a high-voltage electric field in a fused silica capillary. The high voltage (20 to 25 kV) and the small internal volume of the fused silica capillary provide resolution that often exceeds 100,000 plates/meter. This high resolution combined with a different separation mechanism will often provide baseline resolution of peaks that coeluted during a reversed-phase HPLC separation. Very small volumes are required (10 to 200 nl), allowing the majority of the sample to be used for other methods of analysis or, if necessary, for repurification.
BASIC PROTOCOL 4
Materials 0.1 N NaOH Peptide sample 20 mM sodium phosphate, pH 2.5 Capillary electrophoresis (CE) instrument 75-µm-i.d. uncoated capillary column with separating length of 50 cm (from manufacturer of CE instrument or Polymicro Technologies) 1. Set detector wavelength to 200 nm. If excessive baseline noise occurs at 200 nm, increase wavelength to 214 nm.
2. Wash capillary with 0.1 N NaOH for 1 min, followed by Milli-Q water for 1 to 2 min. 3. Equilibrate capillary with 20 mM sodium phosphate buffer, pH 2.5, for 5 min or until a stable baseline is established.
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4. Inject sample using a pressure or vacuum injection of 5 to 30 sec. Some instruments inject the sample by creating a vacuum, whereas others apply a gas pressure to the sample reservoir. Both methods are capable of producing reproducible injections. Other instruments employ electrokinetic injection, in which an electric field is used to move the sample into the separating capillary. A drawback to this procedure is that the velocity of sample migration is dependent on the charge of the sample. This can result in a disproportionate injection of the sample components that have a higher charge. Also see UNIT 10.9. The peptide sample does not generally require any special preparation before injection.
5. Separate the components of the sample at 20 to 25 kV. 6. Repeat steps 2 to 5 for each additional sample. Replace capillary inlet and outlet buffer reservoir with fresh 20 mM sodium phosphate after every five runs. Buffer ion depletion can affect the reproducibility of the separation.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Detector flow cell A flow cell with an internal volume <12 µl is required for 2-mm columns and with internal volume <5 µl for 1-mm columns. Many of the newer HPLC systems designed for 2-mm columns are supplied with flow cells that have appropriately small (0.005-in.-i.d.) outlet tubing that allows peaks to be collected without loss of resolution caused by mixing. Older instruments designed only for 4.6-mm-i.d. columns, such as the Hewlett-Packard HP-1090, may need to be modified. This can easily be accomplished for most instruments by connecting 75-µm-i.d. FSC tubing (Polymicro Technologies) or 0.005-in.-i.d. PEEK tubing (Upchurch Scientific) to the outlet of the flow cell. This tubing should be kept as short as possible to prevent loss of resolution from mixing and to minimize backpressure on the flow cell (excessive backpressure can result in a broken flow cell). Fast-evaporation MALDI matrix solution 5 mg nitrocellulose 20 mg α-cyano-4-hydroxycinnamic acid (4HCCA) Dissolve nitrocellulose and 4HCCA in 500 µl acetone in a 1.5-ml microcentrifuge tube. Add 500 µl isopropanol and mix completely. Store at −20°C 1 to 2 months. Keep the cap tightly sealed at all times—evaporation of the solvent is the main problem encountered in repeated use. Before adding isopropanol, it may be necessary to vortex the material for 1 to 2 min to dissolve the nitrocellulose completely. Use only pure nitrocellulose (e.g., from Schleicher & Schuell). Some nitrocellulose membranes contain other material supports.
MALDI matrix solution Prepare in 30% (v/v) acetonitrile/0.1% (v/v) trifluoroacetic acid either: 1. 20 mg/ml α-cyano-4-hydroxycinnamic acid (use only the supernatant as matrix solution); 2. 50 mg/ml 2,5-dihydroxybenzoic acid (DHB); or 3. Saturated solution of 3,5-dimethoxy-4-hydroxy-trans-cinnamic acid (sinapinic acid; use only the supernatant as matrix solution). Reversed-Phase Isolation of Peptides
Trifluoroacetic acid from Pierce is recommended because some manufacturers add antioxidants that can generate artifacts. The 4HCCA may yield better signal if it is further purified by recrystallization (UNIT 16.3).
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Mass standard solution Dissolve 1 pmol/ml des-Arg-bradykinin (e.g., Sigma) and 2 pmol/ml ACTH-CLIP (18-39) in 30% (v/v) acetonitrile/0.1% (v/v) trifluoroacetic acid (Pierce). COMMENTARY Background Information Reversed-phase HPLC has been widely utilized for analysis and preparative fractionation of peptides. Although many different buffers have been described in the literature, trifluoroacetic acid (TFA) has become the most widely used ion-pairing agent for reversed-phase chromatography of peptides. The volatility of TFA enables it to be compatible with many analytical methods including protein sequencing, online electrospray mass spectrometry, amino acid analysis, and capillary electrophoresis. The development of columns with small internal diameters (2 and 1 mm) has greatly facilitated the ease of handling of small amounts (<200 pmol) of proteolytic digests of proteins (see Basic Protocol 1; Schlabach and Wilson, 1987). The use of 2-mm-i.d. columns generally results in resolution equivalent or superior to that obtained on a 4.6-mm-i.d. column, with significant cost savings in solvent purchase and disposal. Another advantage of smaller-i.d. columns is the increase in sensitivity they provide. The increase in detector signal is a function of the inverse square of the column radius. Reducing the column diameter from 4.6 to 1 mm results in a signal that is 20 times larger. Microbore 1-mm-i.d. columns have been more problematic than 2-mm-i.d. columns. The inability of many HPLC pumps to deliver samples at a rate of 50 to 100 µl/min reliably has prevented widespread use of these smaller columns. Many commercially packed 1-mm-i.d. columns have had low theoretical plate counts compared to 2-mm-i.d. columns. This is primarily due to difficulty in packing these columns in small stainless steel tubes. The problems associated with packing 1mm stainless steel columns are generally not observed with columns of diameters <1 mm. These columns are packed into fused silica capillaries that have extremely smooth walls compared to stainless steel tubing. Capillary columns have a high theoretical plate count that often exceeds 100,000 plates per meter, resulting in higher resolution than can usually be achieved in a 1- or 2-mm-i.d. column. The high resolution and small internal diameters of capillary columns are ideal for high-sensitivity peptide mapping at the 5-pmol level and below (see Basic Protocol 2; Cobb and Novotny,
1989; Davis and Lee, 1992). Capillary columns can be fabricated in the laboratory using a slurry packing technique with an HPLC pump (Moritz and Simpson, 1992). Many different types and sizes of capillary columns are commercially available. Capillary columns of 0.8mm i.d. can be used on some HPLC systems without using a splitter. These HPLC systems must be capable of delivering a flow rate of 25 to 50 µl/min and be equipped with a micro flow cell (<1 µl) to prevent a loss of resolution caused by mixing in the flow cell. The short path-length of these flow cells can often outweigh any gain in sensitivity. Replacing a micro flow cell with a 200-µmi.d. U-shaped capillary cell can increase detector sensitivity. This type of capillary cell is available for a variety of UV detectors from LC Packings. An 0.32-mm-i.d. capillary column can provide high sensitivity and good recovery for analytical and micropreparative collection of peptides from in situ digests of proteins electroblotted on one- and two-dimensional gels (UNIT 10.7; Wong et al., 1993). An 0.32-mmi.d. capillary column requires a capillary flow cell to prevent loss of resolution from mixing. Although capillary flow cells typically have short path lengths, the thin walls of the capillary and the short path length of the detector cell allow transmission of UV light at 195 nm. The shorter wavelength allows higher-sensitivity peptide detection than can be obtained at 214 nm. Peak collection of capillary HPLC fractions can be tedious. However, a significant advantage to this method is the high concentration present in the collection tube as a result of the small peak volumes (0.5 to 3 µl). The high concentration in the sample tube minimizes sample loss by adsorption to the tube walls. When a 5-pmol sample is diluted to 100 to 200 µl—the typical peak volume obtained from a separation on a 1- to 2-mm column—adsorption to the tube walls may occur. Despite the small internal volume of capillary columns, the capacity can be relatively large (>1 nmol). Large-volume injections (>1 ml) may require a wait of several hours for the sample to load on the column. The increased volume will not have a significant effect on resolution for peptide separations, however, because of the absorption/desorption mecha-
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Reversed-Phase Isolation of Peptides
nism of peptide retention. Another advantage of capillary HPLC is its compatibility with on-line electrospray ionization mass spectrometry (UNIT 16.1), a result of the low flow rates (3 to 5 µl/min) employed with these columns. On-line electrospray ionization mass spectrometry allows mass measurement of each component that elutes from the column and in some cases provides sequence information (Griffin et al., 1991; Huang and Henion, 1991; Hess et al., 1993). Small-i.d. columns can provide high-resolution separations, but some peaks may still contain multiple components. MALDI mass spectrometry (see Basic Protocol 3) and capillary electrophoresis (see Basic Protocol 4) are rapid methods for analyzing peak purity. MALDI mass spectrometry (Hillenkamp et al., 1991; Beavis and Chait, 1996; UNIT 16.2) is capable of analyzing low-femtomole quantities of peptide mixtures or isolated peptide fractions (Billeci and Stults, 1993). For example, an in situ tryptic digest of 2 to 5 pmol of a PVDFelectroblotted protein may yield 1- to 2-pmol peptide fractions after enzymatic digestion and separation on a capillary HPLC column. Only a small aliquot (1% to 10%) of a 1-pmol peptide fraction is usually required to determine both purity and molecular weight by MALDI mass spectrometry (Henzel et al., 1993). This information can be useful in deciding which peptide fraction to sequence as well as deciding on the number of cycles with which to program the sequencer. The peptide mass is indispensable for corroborating sequence data and identifying post-translational modifications (Geromanos et al., 1994). Peptide masses may also be used for protein identification by peptide mass mapping (Henzel et al., 1993; Patterson and Aebersold, 1995; Gevaert and Vandekerckhove, 2000). Capillary electrophoresis (CE; UNIT 10.9) can also provide an indication of peak purity. The electropherogram (chromatogram) produced by the CE instrument can be used to approximate the degree of purity of peptide fractions. This is in contrast to MALDI mass spectrometry, which is not quantitative and in which two components with the same concentration may result in significantly different signals. CE generally requires a peptide concentration of 10 pmol/µl, which is 2 to 3 orders of magnitude larger than that required by MALDI mass spectrometry. Although some progress has been made in injecting larger volumes by concentrating the sample within the CE column (Aebersold and Morrison, 1990; Chien and Burgi,
1992; Figeys et al., 1996; Tomlinson et al., 1996), CE injection volumes are generally limited to <50 nl.
Critical Parameters Selection of stationary phase A large variety of reversed-phase stationary phases have been developed. The most commonly employed packings for peptide separation are based on porous silica derivatized with C4, C8, or C18. These columns are available with pore sizes ranging from 100 to 4000 Å. Columns of small particles containing small pores generally provide higher resolution than columns made of materials with larger pores, but peptide recovery is often lower. Peptides obtained from trypsin, chymotrypsin, or pepsin digests, which are often short (<15 residues), are preferably separated on a C18 column. Peptides from Lys-C and cyanogen bromide cleavages, which are often >15 residues long, are usually separated on a C4 column with 300or 1000-Å pores. Selection of mobile-phase solvent Another important variable is the choice of the mobile phase. Mixtures of trifluoroacetic acid (TFA) in water with either acetonitrile or propanol as the organic solvent are the most widely utilized types of mobile phase for peptide separation. The choice of acetonitrile or propanol can affect peptide recovery and resolution. Acetonitrile will generally result in greater resolution than propanol; however, propanol usually provides higher peptide recovery. Complex peptide separations resulting from digestion of large proteins (>100 kDa) demand high-resolution separations and hence are best done using acetonitrile. Increasing the length of the gradient can significantly increase resolution. A tryptic digest of a 100-kDa protein may require a 2- to 3-hr gradient. Peptides that fail to resolve in the course of an extended gradient can often be separated by repeating the chromatography using a different column (C4 or C8 in place of C18). The selectivity of the separation also can be affected by substituting a different organic solvent (i.e., propanol in place of acetonitrile) or by changing the pH of the separation. Rechromatography on the same column using a different organic modifier will often resolve coeluting peaks. Rechromatography can be repeated as many times as necessary but at the expense of some losses with each step. The sample must be diluted with water or concen-
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trated in a Speedvac to lower the organic solvent concentration prior to rechromatography in order to allow the peptides to adsorb to the reversed-phase column. Figure 11.6.3 shows the relationship of pore size and alkyl chain length to resolution and peptide recovery. Sample preparation and storage Peptide fractions should be stored at 5°C in polypropylene tubes, not in polystyrene or glass. Prolonged storage can result in some evaporation of organic solvent from the sample, which may lead to sample adsorption to the microcentrifuge tube. Adding neat TFA to the sample tube (33% final concentration) can resolubilize peptides that may have precipitated or adsorbed to the tube walls (ErdjumentBromage et al., 1993). Solubility of peptides for MALDI can also be enhanced by addition of up to 50% acetonitrile and 2% TFA. Alternatively, up to 1% octyl-β-glucoside may assist in solubilization and is compatible with MALDI (Gharahdaghi et al., 1996). As fractions isolated by RP-HPLC with a volatile mobile phase are normally quite clean, no further cleanup of the sample is usually needed before analysis by MALDI. Preparation of samples for MALDI is easily done by taking a 0.3- to 0.6-µl aliquot of an HPLC fraction straight from the fraction collection tube and mixing it with matrix directly on the sample target. Mass measurement of unfractionated proteolytic digests is useful for rapidly confirming the sequence of known proteins (Billeci and Stults, 1993), for identifying proteins by searching a sequence database with peptide masses (Henzel et al., 1993), and for assessing completeness of digestion. The digestion is preferably performed in a volatile buffer, such as ammonium bicarbonate, ammonium acetate, or N-ethylmorpholine. The buffer should be removed by vacuum centrifugation and the
peptides solubilized in 50% acetonitrile/2% TFA. If nonvolatile buffer components are used, they must be diluted below 50 mM. Samples can be washed after application when the fast-evaporation matrix is used (Shevchenko et al., 1996; see Alternate Protocol). Analysis of subpicomole amounts of peptide may only be successful at substantially lower buffer concentrations. Buffer components may also be removed by pipet-tip purification methods (Erdjument-Bromage et al., 1998; Gobom et al., 1999). Similar pipet tips are available as ZipTips from Millipore. Additional details on sample preparation are found in UNIT 16.3. Special consideration for MALDI and CE Mass accuracy for peptide measurement (1to 3-kDa peptides) with a linear time-of-flight instrument is typically ±1 to 2 Da with external calibration of the mass axis. Higher mass accuracy is achieved by measuring the sample with an internal mass standard, usually a pair of peptides of known mass with signal intensity approximately the same as the unknown sample. Repeated experiments may be necessary to achieve the proper ratio of standard to sample. With proper internal mass standards and an adequate signal-to-noise ratio, the mass accuracy can be ±0.01% (±0.2 Da at m/z 2000). Higher mass accuracy and resolution may be obtained with reflector instruments (UNIT 16.1) or delayed extraction (Vestal et al., 1995). The main variable in capillary electrophoresis of peptides is the choice of the separation buffer. Peptides that comigrate can often be separated by changing the pH of the buffer (Grossman et al., 1988). Changing the pH from 2.5 to 7.0 will often have a major effect on the separation. The ionic strength of the buffer can be increased for peptides that have slow migration rates. A different buffer can be tried when pH and ionic strength fail to improve the separation. Buffer additives can also be utilized to
resolution high
low
small pore size (30 Å)
large pore size (4000 Å)
longer alkyl chain (C18)
shorter alkyl chain (C4)
recovery low
high
Figure 11.6.3 Effects of pore size and alkyl chain length on the resolution and recovery of peptides.
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further enhance selectivity (Cobb and Novotny, 1992).
Troubleshooting
Reversed-Phase Isolation of Peptides
One of the main problems associated with reversed-phase HPLC is the occurrence of a drifting baseline. This is usually the result of contaminants in the mobile phase or inadequate mobile phase degassing. It is essential to use high-purity HPLC reagents and Milli-Q-quality water and to wear powder-free gloves for all manipulations. A baseline rise that increases with increasing organic modifier (solvent B) can be corrected by adjusting the amount of TFA added to the organic solvent. Titrating the amount of TFA in the solvents is especially important for capillary HPLC. In general a concentration of 0.05% to 0.08% TFA in solvent B will usually result in an acceptable baseline. Baseline artifacts, which may appear as negative fluctuations in the baseline, can be caused by a number of HPLC pump malfunctions, including leaks (Dolan, 1991). It is essential to keep HPLC pumps in optimum condition for high-sensitivity HPLC. It is sometimes difficult to detect minor leaks, which can cause a major baseline disturbance during use of capillary HPLC columns. These leaks can usually be detected by placing a dry Kimwipe on the suspected fitting. Occasionally the reversed-phase column can become contaminated by the sample, causing ghost peaks to appear. These are peaks that appear in blank runs when a gradient is run without injecting a sample. Sometimes several blank runs may be necessary before the ghost peaks completely disappear. Extended washing with 80% to 100% organic modifier can also be tried. Another method of cleaning columns, tubing, and flow cells is to use methanol as a solvent. The HPLC is run in isocratic mode with 100% methanol in solvents A and B for 30 to 60 min. If all of the above methods have been tried without success, the column should be replaced. Increased backpressure (>500 p.s.i.) above the normal backpressure of the column may indicate a clogged frit or a contaminated column. Use of a precolumn filter can prevent the column frit from clogging. The precolumn filter is easily replaced and relatively inexpensive. Column inlet frits can be replaced, but care must be exercised to prevent loss of column packing material. Capillary columns are particularly prone to clogging and should always be used in conjunction with a precolumn filter.
Peptide standards have been developed to monitor HPLC performance. Commercial standards (e.g., PE Biosystems) should be run when a new column is first utilized and when resolution appears to deteriorate (Mant and Hodges, 1990). Alternatively, a tryptic digest of a protein that results in well-resolved peptides on reversed-phase chromatography can be utilized to monitor column resolution. The digest should be solubilized in 0.1% TFA and stored frozen at −70°C between uses. Reversed-phase columns should be stored in organic solvent that does not contain TFA, as TFA can slowly dissolve silica. Capillary columns will last significantly longer if this practice is followed. Unsuccessful mass measurement by MALDI may be due to insufficient sample (<10 to 50 fmol). Sample loss is often a result of poor solubility of the peptide or the presence of contaminants that suppress ionization or crystal formation. Peptide solubility is aided by addition of acid, organic solvent, or octyl-β-glucoside. Alternative solvents such as formic acid or methanol may also assist in solubility and ionization (see Table 16.3.1; Cohen and Chait, 1996). In the case of an unfractionated digest or a nonvolatile buffer system, incompatible buffer components may be accommodated simply by dilution of the sample, if the peptide concentration permits, or by pipet-tip purification. Alternatively, salts and buffers are often excluded from the less soluble peptide/matrix crystal, so a brief (∼5-sec) wash of the sample target with cold water may remove many of the interfering salts. CE instruments are relatively simple and are less problematic than HPLC equipment. The two major problems in CE are clogging of the capillary tubing and artifacts resulting from reagent or sample contaminants. Complete absence of current flow usually indicates a blocked capillary. The clog is often found on the sample inlet side and can be usually corrected by removing a short section (2 to 5 mm) of capillary from the column inlet end. The high sensitivity of CE requires high-purity reagents. A number of chemical distributors now sell CE-purity regents.
Anticipated Results Reversed-phase HPLC should result in resolution and good recovery of the majority of the peptides (<30 residues) present in a typical proteolytic digest of a protein (<30 kDa). The use of capillary HPLC for tryptic peptides should allow recovery of peptides at the 1- to
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2-pmol level. CE should be capable of separating peptides that coelute as a single peak on HPLC; however, the concentration must be at least 10 pmol/µl. Peptides in the concentration range of 10 fmol/µl to 50 pmol/µl can be analyzed by MALDI mass spectrometry.
Time Considerations Reversed-phase separations usually require 30 min to 1 hr per analysis. When additional blank gradients are run, several hours may be required. CE analysis of a purified peptide fraction may require from 15 to 60 min, depending on the charge and mass of the sample and the buffer utilized. Mass analysis by MALDI requires 1 to 2 min for preparation of each sample target, 5 to 10 min drying time, and 2 to 5 min per sample for data acquisition. Data processing may add an additional 5 to 10 min.
Literature Cited Aebersold, R. and Morrison, H.D. 1990. Analysis of dilute peptide samples by capillary zone electrophoresis J. Chromatogr. 516:79-88. Beavis, R.C. and Chait, B.T. 1996. Matrix-assisted laser desorption ionization mass spectronomy of proteins. Methods Enzymol. 270:519-551. Billeci, T.M. and Stults, J.T. 1993. Tryptic mapping of recombinant proteins by matrix-assisted laser desorption/ionization mass spectrometry. Anal. Chem. 65:1709-1716. Chien, R.L. and Burgi, D.S. 1992. On-column sample concentration using field amplification in CZE. Anal. Chem. 64:489A-496A. Cobb, K.A. and Novotny, M. 1989. High-sensitivity peptide mapping by capillary zone electrophoresis and microcolumn liquid chromatography using immobilized trypsin for protein digestion. Anal. Chem. 61:2226-2231. Cobb, K.A. and Novotny, M. 1992. Peptide mapping of complex proteins at the low-picomole level with capillary electrophoretic separations. Anal. Chem. 64:879-886. Cohen, S.L. and Chait, B.T. 1996. Influence of matrix solution conditions on the MALDI-MS analysis of peptides and proteins. Anal.Chem. 68:31-37. Davis, M.T. and Lee, T.D. 1992. Analysis of peptide mixtures by capillary high performance liquid chromatography: A practical guide to smallscale separations. Protein Sci. 1:935-944. Dolan, J.W. 1991. Preventive maintenance and troubleshooting LC instruments. In HPLC of Peptides and Proteins: Separation, Analysis, and Conformation (C.T. Mant and R.S. Hodges, eds.) pp. 23-30. CRC Press, Boca Raton, Fla.
Erdjument-Bromage, H., Geromanos, S., Chodera, A., and Tempst, P. 1993. Successful peptide sequencing with femtomole level PTH-analysis: A commentary. In Techniques in Protein Chemistry IV (R.H. Angeletti, ed.) pp. 419-426. Academic Press, San Diego. Erdjument-Bromage, H., Lui, M., Lacomis, L., Grewal, A., Annan, R.S., McNulty, D.E., Carr, S.A., and Tempst, P. 1998. Examination of micro-tip reversed-phase liquid chromatographic extraction of peptide pools for mass spectrometric analysis. J. Chromatogr. A. 826:167-181. Figeys, D., Ducret, A., Yates, J.R., III, and Aebersold, R. 1996. Protein identification by solid phase microextraction-capillary zone electrophoresis-microelectrospray-tandem mass spectrometry. BioTechnology 14:1579-1583. Geromanos, S., Casteels, P., Elicone, C., Powell, M., and Tempst, P. 1994. Combined Edman-chemical and laser-desorption mass spectrometric approaches to micro peptide sequencing: Optimization and applications. In Techniques in Protein Chemistry V (J.W. Crabb, ed.) pp. 143-150. Academic Press, San Diego. Gevaert, K. and Vandekerckhove, J. 2000. Protein identification methods in proteomics. Electrophoresis 21:1145-1154. Gharahdaghi, F., Kirchner, M., Fernandez, J., and Mische, S.M. 1996. Peptide-mass profiles of polyvinylidene difluoride-bound proteins by matrix-assisted laser desorption ionization timeof-flight mass spectrometry in the presence of nonionic detergents. Anal.Biochem. 233:94-99. Gobom, J., Nordhoff, E., Mirgorodskaya, E., Ekman, R., and Roepstorff, P. 1999. Sample purification and preparation technique based on nanoscale reversed-phase columns for the sensitive analysis of complex peptide mixtures by matrixassisted laser desorption/ionization mass spectrometry. J. Mass Spectrom. 34:105-116. Griffin, P.R., Coffman, J.A., Hood, L.E., and Yates, J.R., III. 1991. Structural analysis of proteins by capillary HPLC electrospray tandem mass spectrometry. Int. J. Mass Spectrum Ion Process. 111:131-149. Grossman, P.D., Wilson, K.J., Petrie, G., and Laura, H.H. 1988. Effect of buffer pH and peptide composition on the selectivity of peptide separations by capillary zone electrophoresis. Anal. Biochem. 173:265-270. Henzel, W.J., Billeci, T.M., Stults, J.T., Wong, S.C., Grimley, C., and Watanabe, C. 1993. Identifying proteins from two-dimensional gels by molecular mass searching of peptide fragments in protein sequence databases. Proc. Natl. Acad. Sci. U.S.A. 90:5011-5015. Hess, D., Covey, T.C., Winz, R., Brownsey, R.W., and Aebersold, R. 1993. Analytical and micropreparative peptide mapping by high performance liquid chromatography/electrospray mass spectrometry of proteins purified by gel electrophoresis. Protein Sci. 2:1342-1351. Chemical Analysis
11.6.15 Current Protocols in Protein Science
Supplement 24
Hillenkamp, F., Karas, M., Beavis, R.C., and Chait, R.C. 1991. Matrix-assisted laser desorption/ionization mass spectrometry of biopolymers. Anal. Chem. 63:1193A-1203A. Huang, E.C. and Henion, J.D. 1991. Packed-capillary liquid chromatography/ion-spray tandem mass spectrometry determination of biomolecules. Anal. Chem. 63:732-739. Mant, C.T. and Hodges, R.S. 1990. HPLC of peptides. In HPLC of Biological Macromolecules (K.M. Gooding and F.E. Regnier, eds.) pp. 301332. Marcel Dekker, New York. Moritz, R.L. and Simpson, R.J. 1992. Application of capillary reversed-phase high-performance liquid chromatography to high-sensitivity protein sequence analysis J. Chromatogr. 599:119130. Patterson, S.D. and Aebersold, R. 1995. Mass spectrometric approaches for the identification of gel-separated pr oteins. Electrophoresis 16:1791-1814. Schlabach, T.D. and Wilson, K.J. 1987. Microbore flow-rates and protein chromatography. J. Chromatogr. 385:65-74. Shevchenko, A., Wilm, M., Vorm, O., and Mann, M. 1996. Mass spectrometric sequencing of proteins from silver-stained polyacrylamide gels. Anal.Chem. 68:850-858.
Key References Davis and Lee, 1992. See above. Provides a guide to setting up and using a capillary HPLC. Grossman, P.D. and Colburn, J.C. (eds.) 1992. Capillary Electrophoresis—Theory and Practice. Academic Press, San Diego. Provides an introduction to capillary electrophoresis. Mant, C.T. and Hodges, R.S. (eds.) 1991. HPLC of Peptides and Proteins: Separation, Analysis, and Conformation. CRC Press, Boca Raton, Fla. A comprehensive guide to HPLC of peptides and proteins Wilm, M. 2000. Mass spectrometric analysis of proteins. Adv. Protein Chem. 54:1-30. A general description of mass spectrometers and their utility for solving problems in protein chemistry.
Contributed by William J. Henzel and John T. Stults Genentech, Inc. South San Francisco, California
Tomlinson, A.J., Benson, L.M., Guzman, N.A., and Naylor, S. 1996. Preconcentration and microreaction technology on-line with capillary electrophoresis. J. Chromatogr. 744:3-15. Vestal, M.L., Juhasz, P., and Martin, S.A. 1995. Delayed extraction matrix-assisted laser desorption time-of-flight mass spectrometry. Rapid Commun. Mass. Spectrom. 9:1044-1050. Wong, S.C., Grimley, C., Padua, A., Bourell, J., and Henzel, W.J. 1993. Peptide mapping of 2-D gel proteins by capillary HPLC. In Techniques in Protein Chemistry IV (R. H. Angeletti, ed.) pp. 371-378. Academic Press, San Diego.
Reversed-Phase Isolation of Peptides
11.6.16 Supplement 24
Current Protocols in Protein Science
Removal of N-Terminal Blocking Groups from Proteins
UNIT 11.7
Two enzymatic methods are commonly used in N-terminal sequence analysis of blocked proteins, one for Nα-pyrrolidone carboxyl-proteins in solution (Basic Protocol 1) or blotted onto a membrane (Alternate Protocol 1) and the other for Nα-acyl-proteins blocked with other acyl groups (Basic Protocol 2, Alternate Protocol 2). The enzyme involved in the first of these reactions, pyroglutamate aminopeptidase (EC 3.4.19.3), works on intact proteins as well as on peptides, and represents an excellent reagent in the unblocking/sequencing arsenal. The Support Protocol describes a colorimetric assay for pyroglutamate aminopeptidase activity. The enzyme used for unblocking N-terminal acetyl/formyl amino acids, acylaminoacyl-peptide hydrolase (EC 3.4.19.1; also referred to as acyl-peptide hydrolase, N-acetylalanine aminopeptidase, and acylamino acid–releasing enzyme in the literature) is more restricted in that it cannot use intact proteins as substrate, but can be used to obtain N-terminal sequence from peptides containing up to 20 residues. Consequently, any sequencing protocol with this latter enzyme must include fragmentation of the protein before unblocking can be carried out. To avoid extensive purification after fragmentation, newly generated peptides are chemically blocked with either succinic anhydride (Basic Protocol 2) or phenylisothiocyanate/performic acid (Alternate Protocol 2), and hydrolase is applied to the total mixture of peptides, only one of which, the acylated N-terminal peptide, should be a substrate for hydrolase. After incubation, the mixture of peptides is subjected to sequence analysis. A great variety of chemical methods have been applied to the problem of unblocking N-terminally blocked proteins. The methods include acid-catalyzed hydrolysis (Basic Protocol 3) or methanolysis, hydrazinolysis, and β-elimination after acid-catalyzed N-O shift. For each of these methods, a variety of conditions have been adapted to satisfy the unique properties of different blocking groups and different proteins/peptides. For chemical removal of acetyl and longer-chain alkanoyl groups (Alternate Protocol 3), conditions may have to be harsh enough to cause extensive cleavage of, for example, aspartyl peptide bonds. For chemical removal of formyl groups and opening of the cyclic imide of pyrrolidone carboxylate (Alternate Protocol 4), reaction conditions can generally be sufficiently mild that few or no peptide bonds are cleaved. Blocked proteins that are devoid of aspartate and other labile peptide bonds are not often encountered, so the best substrates for chemical unblocking methods are aspartate-free N-terminal peptides. Because the yield of unblocked peptide varies extensively, the amounts of protein recommended for these protocols are considerably larger than those needed for the direct sequencing of an unblocked protein. It is important to keep in mind that the side chain amides of asparagine and glutamine are among the sensitive amide bonds that are likely to be modified by the various treatments. In general, it should be safe to conclude that even low yields of amide together with high yields of the acid originated as the amide in the acid-treated sequence under study. However, if hydrolysis is extensive enough that the amide cannot be observed at all, the wrong amino acid will obviously be identified. The choice of method for removing N-terminal blocking groups depends on the nature of the blocking group and the N-terminal amino acid. In some cases information about the groups will be available from the prior history of the sample, from mass analysis, or from studies of homologous proteins. In the absence of such information, the process is trial and error. Testing for hydrazine lability, acid lability, and enzyme susceptibility provides clues as to the nature of the blocking group. Successful removal of the group should make it possible to obtain sequence information beyond the blocked residue.
Chemical Analysis
Contributed by Elizabeth Fowler, Mary Moyer, Radha G. Krishna, Christopher C.Q. Chin, and Finn Wold
11.7.1
Current Protocols in Protein Science (1996) 11.7.1-11.7.17 Copyright © 2000 by John Wiley & Sons, Inc.
Supplement 3
BASIC PROTOCOL 1
REMOVAL OF PYRROLIDONE CARBOXYLIC ACID WITH PYROGLUTAMATE AMINOPEPTIDASE TREATMENT OF PROTEINS IN SOLUTION Pyrrolidone carboxylic acid (PCA), also called pyroglutamic acid (see Fig. 11.7.1), is formed by cyclization of glutamine or glutamic acid to form an amide bond between the α-amino group and the δ-carboxyl group. PCA is found on many naturally occurring proteins and peptides. In addition, PCA may be formed by cyclization of amino-terminal glutamine residues during protein isolation, proteolysis and separation of peptides, and/or protein sequencing (Crimmins et al., 1988). The presence of PCA precludes direct amino-terminal sequence analysis because the α-amino group is not available for reaction with the Edman reagent. The enzyme pyroglutamate aminopeptidase (EC 3.4.19.3) can be used to remove pyroglutamate and leave a free α-amino group on the adjacent residue accessible for Edman degradation and other chemical reactions. The enzyme may be used to digest proteins and peptides either in solution or after electroblotting to membranes (see Alternate Protocol 1). The following protocol describes enzymatic removal of pyrrolidone carboxylic acid from proteins or peptides in solution. The reaction is generally more efficient in solution than on electroblotted proteins (see Alternate Protocol 1). Materials Protein or peptide, reduced and alkylated (UNIT 11.1) PGA digestion buffer (see recipe) Calf liver pyroglutamate aminopeptidase (EC 3.4.19.3; sequencing grade, Boehringer Mannhein) 0.05 M acetic acid Nitrogen (research grade) Screw-cap vial Magnetic stirring plate Additional reagents and equipment for dialysis (UNIT 4.4 & APPENDIX 3B) 1. Prepare an 0.5 to 1 mg/ml solution of protein or peptide in PGA digestion buffer. Before PGA digestion, the protein should be reduced and alkylated (UNIT 11.1). If the protein preparation contains components other than those specified for PGA digestion buffer, the protein solution should be dialyzed against PGA digestion buffer at 4°C (UNIT 4.4 & APPENDIX 3B) or otherwise desalted.
2. Transfer the protein solution to a screw-cap vial containing a stir bar. The vial should have a volume no greater than three times the volume of the sample.
2HC
CH2
C
C H
O
Figure 11.7.1 Pyrrolidone carboxylic acid in peptide linkage.
O C
peptide
N H Removal of N-Terminal Blocking Groups from Proteins
11.7.2 Supplement 3
Current Protocols in Protein Science
3. Dissolve an aliquot of calf liver pyroglutamate aminopeptidase in water to give a concentration of 0.25 mg/ml (enzyme stock solution). Sequencing-grade enzyme is supplied by the manufacturer (Boehringer Mannheim) in vials containing 25 ìg enzyme protein. The lyophilized material supplied by the manufacturer contains ∼70% (w/w) sucrose, 17% (w/w) potassium phosphate, 8% (w/w) EDTA, and 5% (w/w) enzyme. A single vial should be reconstituted in 0.1 ml water, then flushed with nitrogen and stored ≤1 week at 4°C. If bulk enzyme (Boehringer Mannheim) is used, reconstitute at 0.25 mg/ml. Some authors report successful storage of this preparation up to 1 month at −20°C in screw-cap vials that have been flushed with nitrogen.
4. Add sufficient enzyme stock solution to the protein solution in the screw-cap vial to give an enzyme/substrate ratio of ∼1/300 (w/w). Flush the vial with nitrogen and stir the solution 7 to 9 hr at 4°C. Enzyme/substrate ratio and digestion time are dependent on the particular protein or peptide being digested. The rate of removal of PCA varies with the adjacent residue (see Background Information). Small peptides may be digested at higher temperatures (40°C) for shorter times, generally 2 to 3 hr (Dimaline and Reeve, 1983).
5. Add a second aliquot of enzyme stock solution (equal in volume to that added in step 4). Flush the vial with nitrogen and stir the solution an additional 14 to 16 hr at room temperature. 6. Dialyze the reaction mixture against 0.05 M acetic acid (UNIT 4.4 & APPENDIX 3B). The protein is now suitable for sequence analysis. Alternatively, the protein or peptide may be separated from the reagents by any suitable chromatographic or electrophoretic technique (see Chapters 8 and 10). The reaction mix may also be applied directly to the sequencer support.
REMOVAL OF PYRROLIDONE CARBOXYLIC ACID WITH PYROGLUTAMATE AMINOPEPTIDASE TREATMENT OF ELECTROBLOTTED PROTEINS
ALTERNATE PROTOCOL 1
Identification of proteins separated by gel electrophoresis is conveniently achieved by performing amino-terminal sequence analysis after electroblotting the proteins onto polyvinylidene difluoride (PVDF) membranes (UNITS 10.7 & 11.3). If no sequence is obtained when the blotted protein is sequenced, the amino terminus of the protein may be blocked by pyrrolidone carboxylic acid (PCA) or another modification of the α-amino group. Digestion with pyroglutamate aminopeptidase (EC 3.4.19.3) is useful for removing PCA prior to sequencing. Additional Materials (also see Basic Protocol 1) Single protein band on PVDF membrane (UNIT 10.7) containing ≥50 pmol protein 0.5% (w/v) polyvinylpyrrolidone (mol. wt. 40,000, PVP-40; Sigma) in 0.1 M acetic acid PGA digestion buffer (see recipe) Screw-cap polypropylene microcentrifuge tube 1. Dissolve an aliquot of calf liver pyroglutamate aminopeptidase in water to give a concentration of 0.25 mg/ml (enzyme stock solution). Sequencing-grade enzyme is supplied by the manufacturer (Boehringer Mannheim) in vials containing 25 ìg of enzyme protein. The lyophilized material supplied by the manufacturer contains ∼70% (w/w) sucrose, 17% (w/w) potassium phosphate, 8% (w/w) EDTA, and 5%
Chemical Analysis
11.7.3 Current Protocols in Protein Science
Supplement 3
(w/w) enzyme. A single vial should be reconstituted in 0.1 ml water, then flushed with nitrogen and stored ≤ 1 week at 4°C. If bulk enzyme (Boehringer Mannheim) is used, reconstitute at 0.25 mg/ml. Some authors report successful storage of this preparation up to 1 month at −20°C in screw-cap vials that have been flushed with nitrogen.
2. Place single protein band on PVDF membrane in a screw-cap polypropylene microcentrifuge tube and rinse well with water. Add 0.5% PVP-40 in 0.1 M acetic acid to cover. Incubate 30 min at 37°C. After rinsing, as much as possible should be drained off without allowing the membrane to dry.
3. Rinse ten times with water. 4. Cover the PVDF membrane with PGA digestion buffer. 5. Estimate the amount (micrograms) of blotted protein present. The amount of protein present in the band may be estimated from the amount applied to the gel, or, preferably, by comparing the intensity of the stained band with a dilution series of a known protein. An estimate +/−100% is generally adequate.
6. Add enzyme stock solution to achieve an enzyme/substrate ratio of 1/20 (w/w). Flush the vial with nitrogen and close. Incubate 5 hr at 4°C, then 18 hr at room temperature. 7. Rinse the PVDF membrane with water several times. Air dry. The pieces are now suitable for sequence analysis. SUPPORT PROTOCOL
COLORIMETRIC ASSAY FOR PYROGLUTAMATE AMINOPEPTIDASE ACTIVITY Pyroglutamate aminopeptidase activity is not particularly stable. One cause of failure to remove an amino-terminal blocking group may be lack of activity in the enzyme stock solution. The following protocol describes the method used by Boehringer Mannheim and several authors for monitoring enzyme activity based on generation of β-naphthylamine from L-pyroglutamyl-β-naphthylamide. An alternative method for monitoring enzyme activity—assaying for the hydrolysis of a synthetic peptide with ninhydrin—is described in Doolittle (1972). Materials PGA digestion buffer (see recipe) PGA-N substrate solution (see recipe) 25% (v/v) trichloroacetic acid 0.1% (w/v) sodium nitrite 0.5% (w/v) ammonium sulfamate NED solution (see recipe) 13 × 100–mm test tubes 37°C incubator or water bath 1. Prepare a 13 × 100–mm test tube for each enzyme sample and a “no enzyme” blank. Add 1.0 ml PGA digestion buffer and 0.1 ml PGA-N substrate. Incubate 5 min at 37°C.
Removal of N-Terminal Blocking Groups from Proteins
The no enzyme blank is prepared identically to the enzyme sample, except that the enzyme solution is omitted from the incubation; it is added after the reaction has been stopped.
11.7.4 Supplement 3
Current Protocols in Protein Science
2. Add 0.1 ml enzyme stock solution to enzyme sample tube; add nothing to blank tube. Mix well. Incubate exactly 15 min at 37°C. 3. Add 1.0 ml of 25% trichloroacetic acid to all tubes. Add 0.1 ml enzyme stock solution to blank tube. Mix well. If precipitate forms, filter through medium-grade filter paper into clean tubes.
4. Transfer 1.0 ml from each tube into a clean tube. 5. Add 1.0 ml of 0.1% sodium nitrite and mix well. Incubate 3 min at 37°C. 6. Add 1.0 ml of 0.5% ammonium sulfamate and mix well. Let stand ∼5 min at room temperature. 7. Add 2.0 ml NED solution and mix well. Incubate 20 min at 37°C. 8. Mix samples and read absorption at 578 nm. 9. Use A578 to calculate activity (U/ml enzyme solution) using the following formulas: ∆A578 = A578,sample − A578,blank Activity = 0.163 × ∆A578
REMOVAL OF ACYLAMINO ACIDS BY HYDROLASE TREATMENT USING SUCCINIC ANHYDRIDE AS BLOCKING REAGENT
BASIC PROTOCOL 2
Instead of purifying the N-terminal peptide after fragmentation of a blocked protein, the generated peptides can be rendered inert by chemical blocking using one of two blocking reactions: succinylation with succinic anhydride (as in this protocol) and carbamoylation by reaction with isothiocyanate followed by oxidation with performic acid to convert thiocarbamate to carbamate (Alternate Protocol 2). The general strategy for establishing N-terminal sequences in Nα-acylated proteins is outlined in Figure 11.7.2. All steps— fragmentation of the blocked protein, blocking the newly generated N-termini, treatment with aminoacyl-peptide hydrolase and finally with acylase—are performed in a single test tube to ensure optimum yield. Because the enzyme releases an N-terminal acylamino acid, special considerations must be given to identification of this first residue. Direct identification by HPLC is possible; in the protocol below the acetylamino acid is hydrolyzed with acylase, and the liberated amino acid is identified by direct amino acid analysis. Other protocols involve derivatization with phenacyl bromide and analysis of the resulting phenacylate acetylamino acids by reversed-phase HPLC (Hirano et al., 1993). Intact protein is digested with specific endoproteases (UNIT 11.1) or cleaved with cyanogen bromide (UNIT 11.4), the newly generated amino groups are blocked, and the N-terminal Ac-peptide is unblocked with acylaminoacyl-peptide hydrolase for sequencing. The blocked N-terminal amino acid is treated with acylase and identified. In this protocol succinic anhydride is used to block newly generated peptides; in Alternate Protocol 2 phenylisothiocyanate and performic acid are used for blocking to yield phenyl carbamates. NOTE: This protocol was developed using an LKB Alpha Plus (ninhydrin-based) Analyzer (Pharmacia Biotech) for amino acid analysis. This analyzer requires ≥0.5 to 1 nmol amino acid for a significant analysis. With more sensitive amino acid analysis methods, the amount of sample can obviously be reduced. Chemical Analysis
11.7.5 Current Protocols in Protein Science
Supplement 3
(NH2)
H
NH2 protein
AcN–XYZ fragment protein (NH2)
H AcN–XYZ
NH2 H2N
H2N
peptides
block newly created N-terminals
H
(NHR)
AcN–XYZ
H
NHR
RN
H Ac-peptide, R-peptides
RN treat with hydrolase
(NHR) H
H AcN–X + H2 N–YZ
RN
sequence peptide sequencing (YZ – – )
NHR
H RN
identify original blocked N-terminal amino acid amino acid analysis (X)
Figure 11.7.2 Strategy for the sequencing of Nα-acetylated proteins, unblocking with acylaminoacyl-peptide hydrolase. Symbols: Ac, acetyl group; R, amino acid side chain; X, Y, and Z, amino acid residues.
Materials 1 nmol/10 µl protein/peptide dissolved in a small volume of suitable volatile solvent 20% (v/v) acetic acid Succinic anhydride 12% (v/v) triethylamine (TEA) 20% (v/v) trifluoroacetic acid (TFA) Ethyl ether Hydrolase buffer I: 50 µM sodium phosphate buffer (pH 7.2; APPENDIX 2E)/ 1 mM EDTA/2 mM MgCl2 0.5 M NaOH Acylaminoacyl-peptide hydrolase (EC 3.4.19.1; Pierce, Takara Biochemical, Boehringer Mannheim, or Sigma) 70% (v/v) formic acid Acylase buffer: 100 mM sodium phosphate buffer, pH 7.0 (APPENDIX 2E) Acylase I (3.5.1.14; e.g., Sigma) 0.2 M sodium citrate with pH adjusted to pH 2.2 using concentrated HCl 0.9 × 7.5–cm glass test tubes Removal of N-Terminal Blocking Groups from Proteins
Additional reagents and equipment for endoprotease digestion (UNIT 11.1) or cyanogen bromide cleavage (UNIT 11.4), and reversed-phase HPLC (UNIT 11.6)
11.7.6 Supplement 3
Current Protocols in Protein Science
Generate peptides 1. Transfer 1 to 2 nmol protein in a suitable volatile solvent to a 0.9 × 7.5–cm glass test tube and lyophilize. 2. Generate peptides by digesting the protein with an endoprotease (UNIT 11.1) or by cleaving it with cyanogen bromide (UNIT 11.4). For endoprotease digestion, select an endoprotease with defined cleavage. Resuspend the protein in 50 ìl of an appropriate buffer and add enzyme to give an enzyme/substrate ratio of 1/20 (w/w). Incubate briefly.
3. Heat the reaction mixture 5 min at 100°C or acidify it with 20% acetic acid (when carbonate buffers are used) to stop proteolysis. 4. Lyophilize the reaction products. Succinylate α- and ε-NH2 groups 5. Dissolve lyophilized sample in 50 µl water or carbonate buffer. Add a 200-fold molar excess of solid succinic anhydride in small portions over a span of 1 hr with vigorous mixing on a vortex stirrer. Maintain pH at 10.0 by adding 12% triethylamine. 6. Let the reaction mixture (∼pH 10) stand overnight at room temperature. 7. Acidify the reaction mixture to pH 2 to 2.5 with 20% TFA. Lyophilize thoroughly. 8. Extract the residue six to ten times with 2 ml ethyl ether to remove all succinic acid and salts of triethylamine. The combined ether extracts can be air dried and screened for any possible presence of peptides by reversed-phase HPLC on a C18-100/300Å column (UNITS 11.6) and/or by amino acid analysis after hydrolysis.
Unblock N-terminal peptide and sequence 9. Dry residual blocked peptides. The residue is now invisible.
10. Add 50 µl hydrolase buffer I and adjust the pH to 7.2 with 1 to 3 µl of 0.5 M NaOH. 11. Add 2 to 5 µg acylaminoacyl-peptide hydrolase to the total peptide mixture and incubate 6 hr at 37°C. More enzyme can be used because the enzyme itself is an acetylated protein and hence does not give any background in sequencing. When pure enzyme is used, incubation times ≤12 hr do not give any side reactions. Although the hydrolase is quite active on short peptides, larger quantities of enzyme and longer incubation times may well be needed for longer peptides. Acetylated peptides with three to eight amino acids are essentially completely hydrolyzed under standard conditions unless they contain positively charged amino acids or proline; longer peptides such as the acetylated 16-residue fibrinopeptide A and the natural 13-residue Ac-α-MSH were hydrolyzed ∼50% in 12 hr, but the 14-residue Ac-renin substrate was only hydrolyzed ∼2% in 20 hr with 4 ìg of hydrolase (Kobayashi and Smith, 1987).
12. Heat-inactivate the reaction 5 min at 100°C. 13. Lyophilize the sample. 14. Dissolve sample in 50 µl of 70% formic acid. Apply an aliquot equivalent to 0.5 nmol protein to the sequencer to identify the N-terminal sequence from the second residue onwards. Chemical Analysis
11.7.7 Current Protocols in Protein Science
Supplement 3
Lysine is detected as the phenylthiohydantoin (PTH) derivative of Nε-succinyl-lysine, which elutes ∼1 min in front of PTH-alanine (about at the same time as Nε-acetyl-lysine) in standard analysis.
Identify the blocked N-terminal amino acid 15. After removing the required aliquot for sequencing, lyophilize the remaining 0.5 to 1.5 nmol protein. 16. Dissolve lyophilized sample in 50 µl acylase buffer. Add 5 U acylase I at pH 7.0 and digest 6 hr at 37°C. 17. Lyophilize the reaction mixture. 18. Dissolve the residue in 50 µl citrate buffer, pH 2.2. Apply entire sample to an amino acid analyzer. ALTERNATE PROTOCOL 2
REMOVAL OF ACYLAMINO ACIDS BY HYDROLASE TREATMENT USING PHENYLISOTHIOCYANATE/PERFORMIC ACID AS BLOCKING REAGENTS This protocol is the basis for a commercially available kit for unblocking proteins (Tsunasawa et al., 1990; Hirano et al., 1993). The catalog/Technical Guide 1 from Takara Biochemical describes kits and methods for chemical and enzymatic removal of blocking groups and sequencing of N-terminally blocked protein. As indicated in Figure 11.7.2, the general strategy in this protocol is essentially the same as that in Basic Protocol 2. The two protocols differ in the chemistry for blocking newly generated peptides. Additional Materials (also see Basic Protocol 2) 12.5% (v/v) trimethylamine Phenylisothiocyanate Benzene Ethyl acetate Performic acid, freshly prepared by mixing 95 µl of 99% formic acid and 5 µl of 30% hydrogen peroxide Hydrolase buffer II: 10 mM sodium phosphate buffer (pH 7.2; APPENDIX 2E)/ 0.1 mM dithiothreitol (DTT) 0.5% (v/v) trifluoroacetic acid (TFA) 0.9 × 7.5–cm glass test tubes IEC CL clinical centrifuge or equivalent Additional reagents and equipment for endoprotease digestion (UNIT 11.1) or cyanogen bromide cleavage (UNIT 11.4), and reversed-phase HPLC (UNIT 11.6) Generate peptides 1. Prepare sample and generate peptides by enzymatic or chemical cleavage, and lyophilize the reactions products as described in Basic Protocol 2, steps 1 to 4. Block α- and ε-NH2 groups 2. Dissolve lyophilized peptides in 100 µl of 12.5% trimethylamine. Add 10 µl phenylisothiocyanate and mix well. Incubate 1 hr at 50°C.
Removal of N-Terminal Blocking Groups from Proteins
3. Add 250 µl benzene and mix by vortexing. Centrifuge 5 min at maximum speed in an IEC CL centrifuge (5000 rpm). Remove the upper layer. Extract the lower phase two more times with benzene. 4. Similarly, extract the remaining lower phase three times with 250 µl ethyl acetate.
11.7.8 Supplement 3
Current Protocols in Protein Science
5. Lyophilize the final lower phase. 6. Repeat steps 2 to 5 once. 7. Dissolve lyophilized sample in 50 µl performic acid and mix well. Incubate 1 hr at 0°C, to convert the thiocarbamate to stable phenyl carbamate. 8. Remove the reagent under reduced pressure in a vacuum desiccator. Dissolve the residue in 100 µl cold water and lyophilize repeatedly to ensure complete removal of organic solvents. Unblock N-terminal peptide 9. Dissolve lyophilized blocked peptides in 50 µl hydrolase buffer II. Add 2 to 5 µg acylaminoacyl-peptide hydrolase to the total peptide mixture. Incubate 6 hr at 37°C. The lyophilized peptides are not visible in the tube. If there is any difficulty in dissolving the substrate, sonicate the sample. The authors recommend 0.05 to 1.0 U of commercial acylaminoacyl-peptide hydrolase per nmol of substrate in their kit.
10. Heat inactivate the mixture 5 min at 100°C. 11. Lyophilize the reaction mixture. Sequence the peptide 12. Dissolve reaction products in 50 µl of 0.5% TFA (or other suitable solvent). An appropriate aliquot may now be applied to the glass-fiber disc of a sequencer. REMOVAL OF ACETYL GROUPS BY ACID HYDROLYSIS The protein is digested with endoprotease (see Table 11.1.3) to release peptides with a minimum of acid-labile peptide bonds. The preferred method for preparing peptides is to digest the protein with the protease from Staphylococcus aureus strain V8 (endoproteinase Glu-C), which cleaves peptide bonds at the carboxyl side of aspartate and glutamate in phosphate buffer, pH 7.8 (UNIT 11.1). With this cleavage, the aspartyl bond (generally the most labile peptide bond) is at the C-terminus, and, at worst, acid treatment will give a high background of aspartate in the first cycle only, with relatively little interference in subsequent cycles. Peptide purification using either ion-exchange chromatography (UNIT 8.2) or high-performance liquid chromatography (HPLC) can be quite tedious because of the complex assay for the blocked N-terminal peptide—the blocked peptide is identified after complete enzymatic digestion with pronase and treatment of the resulting acylamino acid with acylase to release free amino acid (Chin and Wold, 1985, 1986). Because of the anticipated low yield of pure N-terminal peptide, the amount of starting protein required is about five times greater than the amount of peptide analyzed. Blocked peptides are hydrolyzed with HCl, lyophilized, and sequenced.
BASIC PROTOCOL 3
Materials 2 to 5 nmol blocked peptide 1 N HCl 70% (v/v) formic acid or other suitable solvent 1-ml glass vial 14-oz. propane torch 110°C oven 1. Transfer 2 to 5 nmol blocked peptide to a 1-ml vial and lyophilize. Chemical Analysis
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2. Add 200 to 500 µl of 1 N HCl and seal the vial using a 14-oz. propane torch. Incubate 10 min in 110°C oven. The incubation time may have to be adjusted for each individual peptide. The yield of unblocked N-terminus increases with time, but background from peptide bond cleavage also increases to complicate sequence determination. Based on the results with a limited number of proteins, 10 to 20 min appears to be optimal.
3. Immediately cool the vial on ice. Break the seal and lyophilize the hydrolysate. 4. Dissolve sample in 40 to 100 µl of 70% formic acid. An appropriate aliquot (e.g., 10 to 20 µl, or 0.5 to 1.0 nmol peptide) can now be applied to the sequencer. As a solvent, 70% formic acid has been found to be uniformly useful, especially for small amounts of material of low solubility in water. Additional exposure to acid does not appear to affect results. Also, 0.5% (v/v) trifluoroacetic acid (TFA) is a good solvent. Because unblocking is likely to be incomplete, the amount of material subjected to sequencing is increased. ALTERNATE PROTOCOL 3
REMOVAL OF ACETYL GROUPS FROM N-TERMINAL SER AND THR WITH ANHYDROUS TRIFLUOROACETIC ACID Treatment of proteins containing N-terminal Ac-Ser or Ac-Thr with anhydrous trifluoroacetic acid causes an N→O acyl shift and β-elimination of the acetyl esters. This procedure is adapted from Wellner et al., 1990. Materials Polybrene solution: 100 mg/ml Polybrene in 6.7 mg/ml NaCl (0.115 M final) 0.1 nmol/µl protein or peptide in a suitable volatile solvent Anhydrous trifluoroacetic acid (TFA) 12-mm-diameter glass-fiber filter disc, treated with TFA (standard sequencer disc, Applied Biosystems) 1.5-ml polypropylene microcentrifuge tube with cap 45° and 65°C oven 1. Place a 12-mm glass-fiber filter disc in a 1.5-ml polypropylene microcentrifuge tube. Wet disc with 30 µl polybrene solution and dry. 2. Apply 10 to 20 µl protein or peptide solution to the filter disc and dry again. 3. Add 30 µl anhydrous TFA to the thoroughly dried filter. Close tube and incubate 4 min at 45°C. 4. Open the tube in a fume hood and let stand ∼5 min to allow most of the TFA to evaporate. 5. Incubate open tube 10 min at 45°C. Close tube and incubate 16 hr at 65°C or 3 days at 45°C. The filter can now be applied to the pad of a sequencer.
Removal of N-Terminal Blocking Groups from Proteins
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Current Protocols in Protein Science
REMOVAL OF N-TERMINAL FORMYL GROUPS AND UNBLOCKING OF PYRROLIDONE CARBOXYL GROUPS WITH ANHYDROUS HYDRAZINE
ALTERNATE PROTOCOL 4
Protein blocked at the N-terminus is incubated in the presence of anhydrous hydrazine to remove formyl groups and open the cyclic imide of pyrrolidone carboxylate; it is then used for sequencing. In treatment with hydrazine, asparagine and glutamine are converted to the corresponding hydrazides and arginine is converted to ornithine, so three additional PTH standards are required to identify these amino acids in the sequence. This procedure is taken from Miyatake et al., 1993. Materials ∼100 pmol protein, free or bound to polyvinylidine difluoride (PVDF) membrane 3-phenyl-2-thiohydantoin (PTH) derivatives of β-hydrazidyl-aspartate, γ-hydrazidyl-glutamate, and ornithine (standards) Anhydrous hydrazine 4 × 44–mm (small) test tubes 13 × 100–mm (large) test tubes 1. Place ∼100 pmol protein in a 4 × 44–mm test tube and dry. The sample may be protein on a membrane removed from a sequencer after obtaining evidence that the sample is blocked.
2. Place small test tube in a 13 × 100–mm test tube that contains 100 µl anhydrous hydrazine. Seal the large test tube under a vacuum. 3. Incubate the sample 8 hr at −5°C or 4 hr at 20°C. Treatment at −5°C is sufficient to remove formyl blocking groups, and treatment at 20°C is sufficient to open the cyclic imide of pyrrolidone carboxylate. Under the latter conditions, removal of acetyl blocking groups appears to be very low (<10%). At the lowest temperature, side reactions are minimal; at 20°C, only the most labile peptide bonds may be cleaved, but the side-chain amides of asparagine and glutamine are converted to the corresponding hydrazides, and arginine is converted to ornithine. The two conditions can be used in succession if the first one fails. It should be noted that the PTH derivatives of the two hydrazides elute well after leucine in a standard HPLC program; they may not be observed unless the elution time is extended.
4. Open the tube and dry 16 hr in a vacuum desiccator. The treated sample may now be sequenced. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
NED solution Dissolve 25 mg N-(naphthyl)-ethylenediamine⋅2 HCl (Sigma; 2 mM final) in 50 ml absolute ethanol. Prepare fresh daily. PGA digestion buffer 0.1 M NaHPO4, pH 8.0 5 mM dithiothreitol (DTT) 10 mM EDTA 5% (v/v) glycerol Cool to 4°C Prepare fresh before use.
Chemical Analysis
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PGA-N substrate Dissolve 5.5 mg L-pyroglutamyl-β-naphthylamide (Sigma; 0.02 M final) in 1 ml methanol. Prepare fresh daily. COMMENTARY Background Information Of the approximately 200 covalent modifications known to exist in proteins, the sixteen shown in Table 11.7.1 involve the N-terminus (Krishna and Wold, 1993), and all but one of these result in “blocked” N-termini that cannot be sequenced by regular Edman sequencing. The exception is acylation of the N-terminus with another amino acid, leaving a free N-terminus in which the original N-terminal amino acid has become the second residue in the sequence. Blocked N-termini have represented a major hurdle to investigators who wish to establish the N-terminal sequence of a protein. A good deal of effort has been expended in establishing methods that can be used to undo the blocking reaction and make the protein or peptide susceptible to Edman sequencing. Methods for unblocking include both chemical and enzymatic reactions. The choice of method depends on the nature of both the blocking group and the blocked N-terminal amino acid. In these protocols only the two most common blocking structures, pyrrolidone carboxyl and acetylaminoacyl terminals, are considered in detail; others (listed in Table 11.7.1) are mentioned briefly.
Table 11.7.1
N-terminal derivatives in proteinsa
Derivative Nα-formyl Nα-acetyl Nα-acyl
Removal of N-Terminal Blocking Groups from Proteins
Enzymatic methods for unblocking N-termini are more specific than chemical methods. When it was first established that all prokaryotic proteins started with an initial Nα-formylMet, the search for the enzyme(s) responsible for the removal of the formyl (or the formylMet) group was immediately started, and a formyl-hydrolase was found. The occurrence of proteins with an N-terminal formyl group still attached is rare, and the reaction catalyzed by this enzyme has not become a part of the standard sequencing procedures. When it was subsequently found that a large number of eukaryotic proteins contained acylated N-termini, a similar search for acyl-hydrolases was initiated. In spite of much work in this area, no such enzyme has yet been found. Considering that initiation of protein synthesis in eukaryotes involves free Met and that any acylated derivative is thus made co- or post-translationally, presumably for a specific purpose, the absence of the sought-after unblocking hydrolases is perhaps not surprising. Similarly, there is no enzyme known to catalyze the hydrolysis (deacylation) of the cyclic amide of pyrrolidone carboxylate to convert this N-terminal blocking group to glutamate. However, there are
Nα-myristoyl Nα-lauroyl Nα-tetradeca(mono and di)enoyl α N -pyrolidone carboxyl α N -aminoacyl Nα-αketoacyl Nα-glucuronyl Nα-glycosyl Nα-methyl
No. carbon atoms
Reference
C1 C2 C2, C4, C6, C8, C10 C14 C12 C14:1, C14:2
Capecchi (1966) Arfin and Bradsaw (1988) Neubert et al. (1992)
—
Orlowski and Meister (1971)
— — — — —
Kaji (1976) Recsei and Snell (1984) Kolattukudy (1984) Kennedy and Baynes (1984) Siegel (1988)
Carr et al. (1982) Moscarello et al. (1992) Moscarello et al. (1992)
aTaken from Krishna and Wold (1993).
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enzymes that will remove the entire blocked amino acid. One of these, pyroglutamate aminopeptidase (5-oxoprolyl peptidase; Orlowski and Meister, 1971), catalyzes removal of pyrrolidone carboxylate (pyroglutamate) from the N-terminus of proteins and peptides, and, as illustrated in the protocols in this unit, is being used very effectively to unblock proteins for sequencing. Another type of enzyme, acylaminoacyl-peptide hydrolase, catalyzes the removal of other acylamino acids from the N-terminus. In both cases, the acylated N-termini are unblocked by removing the entire N-terminal acylamino acid, leaving residue 2 as the new free N-terminus. Pyroglutamate is found at the amino terminus of a number of naturally occurring proteins. In some cases the cyclized residue has been found to be essential for biological activity, but in other cases, such as in immunoglobulin chains, formation of pyrocarboxylic acid (PCA) does not appear to affect function (Abraham and Podell, 1981). Pyroglutamate can also form by cyclization of glutamine or glutamic acid during protein purification and sequencing. This cyclization is more likely to occur under acid conditions (Crimmins et al., 1988). The presence of PCA precludes amino-terminal sequence analysis by the widely used Edman degradation method. The enzyme pyroglutamate aminopeptidase is present in a wide variety of plant and animal tissues and in numerous species of bacteria (Szewczuk and Kwiatkowska, 1970; McDonald and Barrett, 1986). Although the first preparations used for deblocking peptides prior to sequencing were isolated from bacteria (Doolittle and Armentrout, 1968), the present commercially available enzymes are of mammalian origin. The enzyme acylaminoacyl-peptide hydrolase (see Basic Protocol 2 and Alternate Protocol 2) is most active with short peptides containing acetylated alanine, methionine, glycine, serine, or threonine as the N-terminus. It will also remove formyl-, propionyl-, and butyryl derivatives of these amino acids, albeit at a slower rate than for the acetyl derivatives (Gade and Brown, 1987). It is not known whether longerchain alkanoyl derivatives are also acceptable as substrates; as more proteins are found with N-terminal saturated and unsaturated fatty acyl groups, the suitability of these derivatives as substrates for the hydrolase becomes increasingly relevant. In addition to peptide length, the nature of the amino acids in the vicinity of the N-terminus also affects enzyme activity
(Krishna and Wold, 1992; Sokolik et al., 1994). An immobilized derivative of the enzyme has been prepared, making it more stable and reuseable (Farries et al., 1991). As in the case of the chemical methods, definition of a specific set of unblocking and sequencing conditions may have to be modified empirically to give optimal information yield for a particular protein. Many of the methods for unblocking proteins use simple chemical reagents; these methods have evolved to take advantage of the unique chemical properties of each derivative. Thus, specific unblocking of acetyl-serine or acetyl-threonine can be effected through a putative β-elimination after N-O-acyl shift in anhydrous acid (Wellner et al., 1990), pyruvoyl can be converted to alanine by reductive amination (Huynh et al., 1984), and formyl and pyrrolidone carboxylate groups can be removed by mild treatment with acid or hydrazine under conditions where acetyl and longerchain acyl groups are not cleaved (Miyatake et al., 1993). It is possible to achieve preferential cleavage of acetyl blocking groups as well (see Basic Protocol 3 and Alternate Protocol 3). The main difficulty with chemical unblocking methods is that, in the typical protein, there will always be some particularly labile peptide bonds that may be cleaved by the treatment; in fact, the various acid treatments that have been used to remove N-terminal acetate are typically equal to or harsher than those used to specifically cleave aspartate peptide bonds (Schultz, 1967; Landon, 1977). Because of this, most of the chemical methods are more reliable and specific if they can be applied to short blocked peptides, selected to be void of aspartate if possible, rather than to the intact parent proteins. This in turn means that potentially complex protein fragmentation and peptide isolation steps (UNITS 11.1-11.6) must be added to the general unblocking procedure. A number of strategies have been explored to produce pure blocked N-terminal peptides. The most obvious one is to digest the protein with trypsin followed by carboxypeptidase B to remove all C-terminal lysines and arginines in the resulting peptides. In this reaction mixture, all the peptides except the N-terminal one will have the positive charge of the N-terminus; thus, these charged peptides can be retarded on a cation-exchange column, and the N-terminal blocked peptide can be found in the unretarded fraction (Chin and Wold, 1986). Because of all the variable and unique properties of individual proteins, it is necessary to
Chemical Analysis
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empirically develop an approach suitable to each protein. The protocols given in this unit are based on a limited number of cases, but they should provide useful starting points for development of appropriate reactions for a particular protein.
Critical Parameters Removal of pyrrolidone carboxylic acid Pyroglutamate aminopeptidase activities isolated from all sources are thiol exopeptidases. The enzyme requires the presence of thiols for activation, so dithiothreitol (DTT) is included in the digestion buffer. The enzyme is inhibited by thiol-binding reagents such as iodoacetate, chloromercuribenzoate, Hg2+, Cu2+, Co2+, and Zn2+ (Szewczuk and Kwiatkowska, 1970). The pH optimum of the enzyme is ∼8.0. Pyroglutamate aminopeptidase is sensitive to denaturing agents. Urea may be used at concentrations ≤1 M, but 0.5 M guanidine hydrochloride decreases enzyme activity to 79% of control values, and the enzyme is essentially inactivated at higher concentrations (Boehringer-Mannheim product information). Enzyme specificity Pyroglutamate aminopeptidase is specific for pyrrolidone carboxylic acid (PCA) residues at the amino terminus of peptides and proteins. The rate of removal of PCA is dependent on the adjacent residue and the enzyme does not cleave PCA-proline (McDonald and Barrett, 1986). Highly purified pyroglutamate aminopeptidase should remove only the amino-terminal PCA residue; however, cleavage after an internal proline has been reported (Nagasawa et al., 1988). Early preparations often contained contaminating endoproteases which produced unwanted internal cleavages. When using such preparations, it was essential to repurify the desired intact protein after digestion. The protein-sequencing grade of pyroglutamate aminopeptidase presently available from Boehringer Mannheim is subjected to a quality control test designed to detect contaminating proteases. Although this should eliminate the problem, the prudent investigator will continue to be alert for evidence of additional cleavages. Removal of N-Terminal Blocking Groups from Proteins
Removal of Nα-acetylamino acids It is well established that persistent buffering capacity in the sample to be sequenced seriously jeopardizes the effectiveness of sequenc-
ing cycles. For this reason, one of the main concerns in the various unblocking procedures is ensuring that they do not leave an excessive amount of buffers to interfere with sequencing. The two major trouble-makers in these protocols are the succinate used in blocking peptides (Basic Protocol 2), and the phosphate used in enzymatic unblocking (Basic Protocol 2 and Alternate Protocol 2). Once aware of these potential problems, it is relatively simple to develop methods to avoid them. Ether extraction and repeated lyophilization are recommended after succinylation and appear to be sufficient to eliminate interfering succinate from the sample. Using small volumes and low concentrations of phosphate buffer in enzymatic unblocking renders interference from phosphate insignificant. It is important to be aware of these potential hazards from introduced impurities to ensure that a successfully unblocked protein or peptide yields meaningful sequence information.
Troubleshooting To the extent that unknown proteins really are unknowns, most of the protocols are strictly empirical and essentially constitute a series of troubleshooting steps. The proposed sequential exploration of hydrazine-labile, acid-labile, and enzyme-susceptible N-termini (Hirano et al., 1993), for example, is designed specifically as a progressive hunting expedition to identify the N-terminus and get sequence information beyond it. Similarly, for unknown blocked proteins, selection of appropriate unblocking procedures, choice of proper enzyme to cause fragmentation, and identification of conditions for the unblocking reaction must be made on a trial-and-error basis. When the amount of protein is limited, this is obviously not an attractive approach. For proteins whose mRNA sequence has been determined, and for which the experiments are designed to explore the extent of N-terminal processing of a given predicted structure, selection of the proper methods can be much more direct. Pyroglutamate digestion is generally used after a sequencing attempt on a protein or peptide fails to yield sequence information, suggesting that the amino terminus is blocked. The protein is then digested with pyroglutamate aminopeptidase and a second sequencing attempt is made. A second failure to obtain amino-terminal sequence data may indicate either that the blocking group is not pyroglutamate or that enzymatic digestion has not been successful. Strategies for removing other
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Current Protocols in Protein Science
blocking groups are discussed below. Several controls to demonstrate enzyme activity of the pyroglutamate aminopeptidase may be employed. Pyroglutamate aminopeptidase activity is fairly labile and sensitive to various inhibitors. Activity of the enzyme may be tested by the colorimetric method described in the Support Protocol. An internal control using a commercially available PCA-containing peptide such as bombesin may be added to the reaction. After digestion, the mixture is separated chromatographically and the peptide subjected to sequence analysis. Crimmins et al. (1988) and Dimaline and Reeve (1983) describe digestion of bombesin and other peptides and separation of unblocked from blocked forms. If the enzyme is active in the protein mixture, variations in enzyme/substrate ratio and digestion times should be investigated. The rate of cleavage of PCA is dependent on the nature of the adjacent residue. If the rate is low, increasing digestion time or temperature may be necessary. Additional aliquots of enzyme may be added at intervals during digestion to compensate for enzyme lability. If the substrate protein is large with substantial tertiary structure, the amino terminus may be inaccessible to the enzyme. The protein should be reduced and alkylated before digestion with pyroglutamate aminopeptidase. Denaturing the protein by making a concentrated solution in 8 M urea or 6 M guanidine⋅HCl and then diluting to ≤1 M urea or ≤0.5 M guanidine⋅HCl may be helpful in exposing the aminoterminus. It may be necessary to cleave the protein with an endoprotease and isolate the amino-terminal peptide before digestion with pyroglutamate aminopeptidase. Digestion of small peptides tends to occur more easily than digestion of large proteins. If more than one residue is exposed at the amino terminus, it may indicate that the enzyme preparation contains additional endoprotease activities. A purer preparation of enzyme may alleviate this problem. Alternatively, the digestion products may be separated chromatographically before sequence analysis. Two amino-terminal residues have also been detected when the residue adjacent to the original PCA is a glutamine (Crimmins et al., 1988). Presumably some of this second glutamine cyclized to PCA, which was then removed by the enzyme.
Anticipated Results With Basic Protocol 1, 200 pmol protein should yield 40 to 100 pmol unblocked protein ready for sequence analysis. Digestion of electroblotted proteins (Alternate Protocol 1) is somewhat less efficient but fewer sample-handling steps are required. Depending on the sensitivity of the protein sequencer, 50 pmol starting protein should produce a detectable signal of 10 to 20 pmol. When combining treatment with hydrolase and proteolysis (Basic Protocol 2 and Alternate Protocol 2), the results can be quite variable because the activity of the hydrolase depends on peptide structure. In a test of eight known acetylated proteins, 500 pmol starting material produced sequences as short as two residues and as long as twelve residues. Under optimized conditions for acid hydrolysis (Basic Protocol 3), 40% to 60% of total peptide yields unblocked N-terminal residues. Treatment with trifluoroacetic acid (Alternate Protocol 3) has given yields of 3% to 40% unblocked N-terminals. Treatment with hydrazine (Alternate Protocol 4) has yielded 50% to 80% unblocking of formylated proteins and 69% to 90% of pyrrolidone-carboxylated proteins.
Time Considerations For Basic Protocol 1 with a reduced and alkylated protein sample that can be placed directly into PGA digestion buffer, setting up the digestion takes ∼30 min and should be started early in the morning so the first digestion step, which requires 7 to 9 hr, can be carried out during the remainder of the working day; the second digestion is carried out overnight. Dialysis requires 3 to 4 hr; alternatively, reversed-phase desalting combined with isolation of the blocked protein can generally be accomplished in 60 to 90 min by reversedphase HPLC. In Alternate Protocol 1, which starts with a blotted protein, preparation and preliminary incubation requires ∼90 min and digestion requires 23 hr. Support Protocol 1 takes ∼3.5 hr to complete. Treatment with hydrolase combined with proteolysis (Basic Protocol 2 and Alternate Protocol 2) requires 1 to 2 days depending on the method for blocking newly generated amino groups. Acid hydrolysis (Basic Protocol 3) can be completed in 0.5 day. Treatment with trifluoroacetic acid (Alternate Protocol 3) requires 2 to 4 days, and treatment with hydrazine (Alternate Protocol 4) takes 1.5 days. Chemical Analysis
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Abraham, G.N. and Podell, D.N. 1981. Pyroglutamic acid. Mol. Cell. Biochem. 38:181-190.
Kaji, H. 1976. Amino-terminal arginylation of chromosomal proteins of arginyl-tRNA. Biochemistry 15:5121-5125.
Arfin, S.M. and Bradshaw, R.A. 1988. Cotranslational processing and protein turnover in eukaryotic cells. Biochemistry 27:7979-7984.
Kennedy, L. and Baynes, J.W. 1984. Nonenzymatic glycosylation and the chronic complications of diabetes: An overview. Diabetologia 26:93-98.
Capecchi, M.R. 1966. Initiation of E. coli proteins. Proc. Natl. Acad. Sci. U.S.A. 55:1517-1524.
Kobayashi, K. and Smith, J. 1987. Ac-peptide hydrolase from rat liver: Characterization of enzyme reaction. J. Biol. Chem. 262:11435-11445.
Literature Cited
Carr, S.A., Biemann, K., Shoji, S., Parmelee, D.C., and Titani, K. 1982. N-tetradecanoyl is the NH2 terminal blocking group of the catalytic subunit of cyclic AMP-dependent protein kinase from bovine cardiac muscle. Proc. Natl. Acad. Sci. U.S.A. 79:6128-6131. Chin, C.C.Q. and Wold, F. 1985. Studies on Nα-acetylated proteins: The N-terminal sequences of two muscle enolases. Biosci. Rep. 5:847-854. Chin, C.C.Q. and Wold, F. 1986. Reinventing the wheel: General approaches to the elucidation of blocked N-terminal sequences. In Methods in Protein Sequence Analysis (K.A. Walsh, ed.) pp. 505-512. Humana Press, Clifton, N.J. Crimmins, D.L., McCourt, D.W., and Schwartz, B.D. 1988. Facile analysis and purification of deblocked N-terminal pyroglutamyl peptides with a strong cation-exchange sulfoethyl aspartamide column. Biochem. Biophys. Res. Comm. 156:910-916. Dimaline, R. and Reeve, J.R., Jr. 1983. Reversedphase high-performance liquid chromatography used to monitor enzymatic cleavage of pyrrolidone carboxylic acid from regulatory peptides. J. Chromatogr. 257:355-360. Doolittle, R.F., 1972. Terminal pyrrolidonecarboxylic acid: Cleavage with enzymes. Methods Enzymol. 25:231-244. Doolittle, R.F. and Armentrout, R.W. 1968. Pyrrolidonyl peptidase. An enzyme for selective removal of pyrrolidonecarboxylic acid residues from polypeptides. Biochemistry 7:516-521. Farries, T.C., Harris, A., Auffret, A.D., and Aitken, A. 1991. Removal of N-acetyl groups from blocked peptides with acylpeptide hydrolase: Stabilization of the enzyme and its application to protein sequencing. Eur. J. Biochem. 196:679685.
Removal of N-Terminal Blocking Groups from Proteins
Kolattukudy, P.E. 1984. Detection of an N-terminal glucuronamide linkage in proteins. Methods Enzymol. 106:210-217. Krishna, R.G. and Wold, F. 1992. Specificity determinants of acetylaminoacyl-peptide hydrolase. Protein Sci. 1:582-589. Krishna, R.G. and Wold, F. 1993. Post-translational modification of proteins. Adv. Enzymol. Relat. Areas Mol. Biol. 67:265-298. Landon, M. 1977. Cleavage at aspartyl-prolyl bonds. Methods Enzymol. 47:145-149. McDonald, J.K. and Barrett, A.J. 1986. Pyroglutamyl peptidase I. In Mammalian Proteases: A Glossary and Bibliography, Vol.2. Exopeptidases (J.K. McDonald and A.J. Barrett, eds.) pp. 305-307. Academic Press, New York. Miyatake, N., Kamo, M., Satake, K., Uchiyama, Y., and Tsugita, A. 1993. Removal of N-terminal formyl groups and deblocking of pyrrolidone carboxylic acid of proteins with anhydrous hydrazine vapor. Eur. J. Biochem. 212:785-789. Moscarello, M.A., Pang, H., Pace-Asciak, C.R., and Wood, D.D. 1992. The N-terminus of human myelin basic protein consists of C2, C4, C6, and C8 alkyl carboxylic acids. J. Biol. Chem. 267:9779-9782. Nagasawa, H., Maruyama, K., Sato B., Hietter, H., Isogai, A., Tamura, S., Ishizaki, H., Semba, R., and Suzuki, A. 1988. Structure and synthesis of bombesin from the silkworm Bombyx mori. In Peptide Chemistry (T. Shiba and S. Sakakibara, eds.) pp. 123-126. Protein Research Foundation, Osaka, Japan. Neubert, T.A., Johnson, R.S., Hurley, J.B., and Walsh, K.A. 1992. The rod transducin subunit amino terminus is heterogeneously fatty acylated. J. Biol. Chem. 267:18274-18277.
Gade, W. and Brown, J.L. 1987. Purification and partial characterization of N-acyl peptide hydrolase from liver. J. Biol. Chem. 253:5012-5018.
Orlowski, M. and Meister, A. 1971. Enzymology of pyrrolidone carboxylic acid. Enzymes 4:123151.
Hirano, H., Komatsu, S., Kajiwara, H., Tsunasawa, S. 1993. Microsequence analysis of the N-terminally blocked proteins immobilized on polyvinylidene difluoride membrane by Western blotting. Electrophoresis 14:839-846.
Recsei, P.A. and Snell, E.E. 1984. Pyruvoyl enzymes. Annu. Rev. Biochem. 53:357-387.
Huynh, Q.K., Vaaler, G.L., Recsei, P.A., and Snell, E. E. 1984. Histidine carboxylase of Lactobacillus 30a. Sequences of the cyanogen bromide peptides from the α chain. J. Biol. Chem. 259:2826-2832.
Schultz, J. 1967. Cleavage at aspartic acid. Methods Enzymol. 11:255-263.
Sakiyama, F. 1990. New tools for protein sequence analysis. Trends Biotech. 8:282-288.
Siegel, F.L. 1988. Enzymatic N-methylation of calmodulin, In Advances in Post-translational Modification of Proteins and Aging (V. Zappia, P. Galletti, R. Porta, and F. Wold, eds.) pp. 341352. Plenum Press, New York.
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Sokolik, C.W., Liang, T.C., and Wold. F. 1994. Studies on the specificity of acetylaminoacyl-peptide hydrolase. Protein Sci. 2:126-131. Szewczuk, A. and Kwiatkowska, J. 1970. Pyrrolidonyl peptidase in animal, plant and human tissues: Occurrence and some properties of the enzyme. Eur. J. Biochem. 15:92-96. Tsunasawa, S., Takakura, H., and Sakiyama, F. 1990. Microsequence analysis of N-acetylated proteins. J. Prot. Chem. 9:265-266. Wellner, D., Panneerselvam, C., and Horecker, B.L. 1990. Sequencing of peptides and proteins with blocked N-terminal amino acids: N-acetylserine or N-acetylthreonine. Proc. Natl. Acad. Sci. U.S.A. 87:1947-1949.
Moyer, M., Harper, A., Payne, G., Ryals, J., and Fowler, E. 1990. In situ digestion with pyroglutamate aminopeptidase for N-terminal sequencing of electroblotted proteins. J. Prot. Chem. 9: 282-283. Describes enzymatic unblocking of electroblotted proteins. Podell, D.N. and Abraham, G.N. 1978. A technique for the removal of pyroglutamic acid from the amino terminus of proteins using calf liver pyroglutamate aminopeptidase. Biochem. Biophys. Res. Comm. 81:176-185. Describes the basic method for digestion of milligram quantities of protein. Tsunasawa et al., 1990. See above.
Key References Chin and Wold, 1986. See above. Describes chemical unblocking using aqueous acid. Crimmins et al., 1988. See above. Describes digestion of peptides and a cation-exchange method for isolating digestion products. Dimaline and Reeve, 1983. See above. Describes digestion of small amounts of peptides and a reversed-phase HPLC method for monitoring digestion and isolating digestion products. Hirano et al., 1993. See above.
Describes hydrolysis with phenylcarbamate blocking. Van Beeumen, J., Van Driessche, G., Huitema, F., Duine, J.A., and Canters, G.W. 1993. N-terminal heterogeneity of methylamine dehydrogenase f ro m Thiobacillus versutus. FEBS Letters 333:188-192. Describes digestion and subsequent sequencing of microgram quantities of protein. Wellner et al., 1990. See above. Describes chemical unblocking with anhydrous acid.
A general overview of unblocking. Krishna, R.G. 1992. N-Acylaminoacyl-peptide hydrolase: Specificity and use to unblock N-acetylated proteins. In Techniques in Protein Chemistry III (R.H. Angeletti, ed.) pp. 77-84. Academic Press, San Diego.
Contributed by Elizabeth Fowler (pyrrolidone carboyxlic acid) AutoImmune, Inc. Lexington, Massachusetts
Krishna, R.G., Chin, C.C.Q., and Wold, F. 1991. N-terminal sequencing of N-acetylated proteins after unblocking with N-acetylaminoacyl-peptide hydrolase. Anal. Biochem. 199:45-50.
Mary Moyer (pyrrolidone carboxylic acid) Glaxo Research Institute Research Triangle Park, North Carolina
Describes the use of hydrolase with succinic acid blocking.
Radha G. Krishna, Christopher C.Q. Chin, and Finn Wold (removal of acylamino acids, chemical unblocking) University of Texas Medical School Houston, Texas
Miyatake et al., 1993. See above. Describes chemical unblocking using hydrazine.
Chemical Analysis
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C-Terminal Sequence Analysis
UNIT 11.8
Sequence analysis from the C-terminus of a protein provides important complementary information to data acquired by mass spectrometry and Edman degradation. A major advantage is that a portion of the C-terminal sequence, from even very large proteins, can be directly analyzed without the need of proteolytic cleavage to generate smaller peptides for analysis. Basic Protocol 1 includes several important modifications in the standard procedure for automated C-terminal protein sequencing (Bergman et al., 2001) based on the principles of Boyd et al. (1992, 1995). This methodology has been tested in the analysis of a broad range of polypeptides with improved results in relation to standard protocols, particularly regarding sensitivity, length of degradation, Pro-passage, and a combination of N- and C-terminal sequence analyses (Bergman et al., 2002). In this procedure, the sample proteins or peptides are electroblotted or immobilized onto polyvinylidene difluoride (PVDF) membranes before either C- or a combination of N- and C-terminal degradation. Over 600 polypeptides ranging from 10 to 600 residues have been analyzed using this protocol in amounts as low as 10 pmol. This technique allows for the determination of up to 15 residues using 100 to 500 pmol, the quantitative characterization of C-terminal sequences and truncation patterns, and the efficient analysis of N-terminally blocked polypeptides. One of the more common techniques for manually determining C-terminal sequence information involves digestion of the purified, intact protein or peptide with one or more carboxypeptidases, which are exopeptidases that remove amino acids one at a time from the C-terminus of a protein. Two methods are commonly used for analysis of the digestion products. (1) The set of peptides generated by sequential removal is analyzed by mass spectrometry and the sequence is then deduced from the mass differences between adjacent peaks in the mass spectrum profile (see Basic Protocol 2). (2) In Basic Protocol 3, after digestion with carboxypeptidase, the released amino acids are separated from the protein and analyzed by amino acid analyses (see UNITS 3.2 & 11.9). Amino acid analysis requires more protein, but provides better quantitation. Carboxypeptidases have a wide range of specificity for individual amino acids, e.g., they cleave C-terminal Lys and Arg residues at very different rates. Carboxypeptidase A, P, and Y cleave most amino acids, although carboxypeptidase A will not cleave Pro or Arg residues. In addition, cleavage by carboxypeptidase A is greatly slowed by Gly residues; carboxypeptidases P and Y are also greatly retarded by Ser and Asp, respectively. These enzymes can be used individually or combined to provide further sequence information and to help with amino acid assignments (see Anticipated Results and Fig. 11.8.1). AUTOMATED C-TERMINAL SEQUENCE ANALYSIS This protocol begins with application of the sample onto a PVDF membrane via adsorption from solution or electroblotting, followed by treatment with phenylisocyanate (PIC) to block amino groups that would otherwise interfere with the degradation chemistry. The membrane-bound sample is then applied to the C-terminal sequencer and the analysis initiated. If the sample is first analyzed by Edman degradation in an N-terminal sequencer, the PVDF membrane must be treated with PIC after N-terminal sequencing and then applied to the C-terminal instrument and subjected to several ethyl acetate washes to remove byproducts from the Edman chemistry before initiating C-terminal analysis. When N-terminal sequence analysis is performed first, the subsequent C-terminal sequence analysis is not compromised in terms of performance, but the amount of polypeptide available for sequencing is reduced due to the unavoidable loss of material in the
BASIC PROTOCOL 1
Chemical Analysis Contributed by Tomas Bergman, Ella Cederlund, Hans Jörnvall, and Elizabeth Fowler Current Protocols in Protein Science (2003) 11.8.1-11.8.17 Copyright © 2003 by John Wiley & Sons, Inc.
11.8.1 Supplement 31
preceding N-terminal degradation. Modifications of reagents and solutions, as well as modifications of the sequencer program and hardware originally described by the manufacturer (Applied Biosystems), are detailed below. Chemical degradation of proteins from both termini using a single-sample application is attractive since it maximizes the sequence information from limited amounts of protein. The sample can be analyzed in the N-terminal sequencer for 5 to 25 cycles, the membrane removed, treated with PIC, and transferred to the reaction cartridge (blot cartridge) of the C-terminal instrument. Lysine residues will not be derivatized with PIC since their ε-amino group has already reacted with phenylisothiocyanate during Edman degradation, which results in formation of phenylthiocarbamyl (PTC)-Lys derivatives. Subsequent C-terminal sequencing can regularly recover 4 to 8 residues. Proteins are commonly blocked at their N-terminus, often by acetylation. Sequence analysis of blocked proteins can be carried out by tandem mass spectrometry of the corresponding N-terminal proteolytic peptide (Jonsson et al., 2000) and by sequencer degradation after application of deblocking protocols to the intact protein (Gheorghe et al., 1997). In this respect, C-terminal sequence analysis is an attractive route, particularly since C-terminal blocking of proteins is not common. Materials Gel-separated protein sample (one- or two-dimensional separations; UNITS 10.1-10.4) Coomassie blue (UNIT 10.5) Analytical-grade chemicals for N- and C-terminal sequence analysis (Table 11.8.1 provides a list of reagents and solutions employed in the C-terminal sequencer; Applied Biosystems) Methanol 0.5% and 0.1% (v/v) trifluoroacetic acid (TFA) 1.25% (v/v) diisopropylethylamine (DIEA) in heptane 1% (v/v) phenylisocyanate (PIC; Sigma) in acetonitrile PVDF membranes (e.g., Immobilon-P, Millipore; ProBlott, Applied Biosystems) 1.5-ml microcentrifuge tubes ProSorb cartridges for sample immobilization and desalting (Applied Biosystems) Petri dishes 60°C heating block For C-terminal sequencer degradation: Procise C instrument (Applied Biosystems) equipped with a blot cartridge (Applied Biosystems) and a 2 × 220–mm reversed-phase (C18) column (Brownlee Spheri-5 PTC, 5 µm) operated at 0.3 ml/min and with detection at 257 nm For sequential N-terminal and then C-terminal degradation: Procise HT instrument (Applied Biosystems) equipped with the same C18 column described for the Procise C instrument, but with detection at 269 nm Additional reagents and equipment for electroblotting (UNIT 10.7), N-terminal degradation (UNIT 11.10) NOTE: Only the modifications of the standard program are given here. The standard program comes with the instrument and the operator has only to include the extra points into the existing program.
C-Terminal Sequence Analysis
Apply sample to the PVDF membrane via electroblotting 1a. Transfer gel-separated proteins (one- or two-dimensional separations) to PVDF membranes using standard electroblotting techniques (UNIT 10.7). Stain the proteins with Coomassie blue (typically R-250; UNIT 10.5) and then dry the membrane. Excise
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Table 11.8.1
Reagents, Solutions, and Conditions for C-Terminal Sequencer Analysisa
Parameter changed
Manufacturer’s protocol
Modified protocol
100 pmol/injection Use as delivered Use as delivered
10 pmol/injection Dilute 1:1 with acetonitrile Dilute 1:1 with acetonitrile
Use as delivered
Dilute 1:1 with acetonitrile
Use as delivered
Dilute 1:1 with acetonitrile
Use as delivered
Dilute 1:1 with acetonitrile
Use as delivered According to Applied Biosystems recommendation
Diluted 1:1 with heptane Dilute 1:1 with acetonitrile
Solvent B
3.5% THF in 82.5 mM sodium acetate pH 3.8, DIEA/acetone 18% THF in acetonitrile
3.8% ethanol in 30 mM sodium acetate, pH 3.8, acetone 7.1% ethanol in acetonitrile
Temperature Column Transfer flask
45°C 45°C
40°C 40°C
Dilution (C1)b ATH aa standard (C3)b N-methylimidazole/ acetonitrile (C4) b piperidine thiocyanate/acetonitrile (C6)b acetic anhydride/ lutidine/acetonitrile (C8)b bromomethyl-naphthalene/acetonitrile (C10)b tetrabutylammonium thiocyanate/acetonitrile (C11)b DIEA/heptane 2% phenylisocyanate/acetonitrile Chromatography Solvent A
aParameters were modified in relation to the manufacturer’s protocol to improve sensitivity and length of degradation. bLabel (bottle position) according to Applied Biosystems nomenclature.
the bands or spots of interest and begin analysis or store in 1.5-ml microcentrifuge tubes at –20°C until use. 2a. For combinations of N- and C-terminal sequence analysis, subject the membranebound sample to N-terminal degradation (UNIT 11.10) first without further pretreatment. After a sufficient number of residues are determined, remove the sample membrane from the N-terminal sequencer and place it in a 1.5-ml microcentrifuge tube for PIC treatment as described for solution samples (step 7) and then apply the sample to the C-terminal instrument. Apply sample to the PVDF membrane in solution 1b. Apply the sample to a PVDF membrane using the ProSorb sample preparation cartridge. Wet the membrane with 5 µl methanol. Add 100 µl of 0.5% TFA followed by the sample solution (5 to 800 µl volumes have been tested with success). Wash three times with 100 µl of 0.1% TFA to remove salt and low molecular weight contaminants. 2b. Thoroughly dry the membrane by turning the insert (according to the ProSorb manufacturer’s instructions) upside-down and placing it on a petri dish (the PVDF membrane with the immobilized sample should face up). Cover with a beaker and place the assembly on a 60°C heating block for ∼20 min. The sample is now ready for application to N-terminal sequence analysis in a combined N-/C-terminal approach. Chemical Analysis
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Perform combined N- and C-terminal sequence analysis (optional) 3. Perform N-terminal degradation of the membrane-bound sample typically for 5 to 25 residues in the N-terminal sequencer (UNIT 11.10). 4. Remove the membrane, place it in a 1.5-ml microcentrifuge tube and treat with PIC as described in step 7. 5. Apply the membrane to the C-terminal sequencer (blot cartridge). Begin the first cycle with ethyl acetate washes in the sequencer cartridge to remove byproducts from the Edman chemistry as follows: 30-sec wash, 30-sec argon dry, 60-sec wash, and 30-sec argon dry. Perform C-terminal sequence analysis The C-terminal degradation method employed in this protocol basically follows the thiohydantoin method (Stark, 1968). The main differences are (1) the C-terminus is activated only once with acetic anhydride and (2) bromomethylnaphthalene is added to S-alkylate the thiohydantoin formed in each cycle which generates a better leaving group in the cleavage. This practice allows combination of peptide cleavage and activation for the next cycle using tetrabutylammonium thiocyanate in TFA (Boyd et al., 1992, 1995). A blot cartridge (Applied Biosystems) intended for N-terminal degradation of electroblotted samples (Procise HT) is used (see Critical Parameters) for improved sensitivity in the C-terminal degradation. 6. Add 5 µl of 1.25% DIEA in heptane (reagent C11 diluted 1:1 with heptane) to the PVDF membrane and allow the heptane to evaporate at 60°C. The heptane is usually gone within ∼1 min.
7. Add 5 µl of 1% PIC in acetonitrile and incubate for 5 min at 60°C. 8. Apply the PIC-derivatized membrane-bound sample to C-terminal sequence analysis using a blot cartridge (see Commentary). 9. Use the following modifications of the standard C-terminal sequencer program. a. Perform extra washing steps with ethyl acetate in each sequencer cycle: 90-sec wash, 30-sec argon dry, 90-sec wash, 30-sec argon dry, 45-sec wash, 30-sec argon dry, 45-sec wash, 30-sec argon dry. The washing steps are accommodated in the sequencer program immediately before the C10/C12 combined cleavage/cyclization step. Additional modifications concerning chromatography and temperatures are given in Table 11.8.1 and below.
b. The elution program is dependent on the particular reversed-phase column used. A typical program consists of a gradient with linear segments: 14% to 27% B from 0 to 4 min 27% to 44% B from 4 to 28 min 44% to 47% B from 28 to 34 min 47% to 90% B from 34 to 35 min followed by isocratic washing of the column at 90% B for 3 min (see Table 11.8.1).
C-Terminal Sequence Analysis
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MANUAL C-TERMINAL SEQUENCING USING CARBOXYPEPTIDASES FOLLOWED BY MASS SPECTROMETRY
BASIC PROTOCOL 2
This method involves digestion of peptide or protein with carboxypeptidases followed by mass analysis of the digestion products. Mass analysis is typically performed using a matrix-assisted laser desorption/ionization time of flight [MALDI-TOF] instrument for its speed of analysis and ease of use, but mass analysis can also be performed using an electrospray instrument. Since the sequence is determined from the mass differences between adjacent peaks in the mass spectrum, the maximum size of the starting sequence is limited by the ability of the mass spectrometer to resolve species differing in the loss of the smallest amino acid, glycine. The advantages of this method include the low sample requirements, the rapidity of mass analysis, and the ease of data interpretation. The following procedure describes the use of the Sequazyme C-peptide sequencing kit available from Applied Biosystems for use with MALDI analysis. The procedures can be adapted for other enzymes provided the appropriate digestion conditions are used (see manufacturer’s recommendations). Materials Peptide or protein of interest (1 to 3 pmol, ≥90% purity) ∼2 × 10–3 U/µl carboxypeptidase Y (CPY) in 30 mM ammonium citrate, pH 6.1 ∼2 × 10–3 U/µl carboxypeptidase P (CPP) in 30 mM ammonium citrate, pH 4.5 Sequazyme kit (Applied Biosystems) containing (or individual reagents): MALDI matrix: α-cyano-4-hydroxy cinnamic acid MALDI matrix diluent: 50% acetonitrile in 0.3% trifluoroacetic acid (TFA) 1 pmol/µl ACTH 18-39 peptide sequencing standard prepared in HPLC-grade water 1 pmol/µl ACTH 18-39 calibration standard prepared in HPLC-grade water 1 pmol/µl angiotensin I calibration standard prepared in HPLC-grade water C-terminal sequencer Sequence analysis software NOTE: If the peptide of interest is considerably larger than the recommended standards, additional calibration standards in the appropriate mass range should be used. Prepare solutions and matrix 1. Dissolve the peptide of interest at 0.1 to 0.5 pmol/µl in HPLC-grade water. Alternatively, prepare two solutions: one in 30 mM ammonium citrate, pH 6.1 to be used for CPY digestion and one in 30 mM ammonium citrate, pH 4.5 to be used for CPP digestion. It is recommended to verify enzyme activity and calibrate the instrument (steps 1 to 14) with the standards before analyzing experimental samples, especially if only limited quantities of sample are available.
2. Prepare four serial 1:3 dilutions of the 2 × 10–3 U/ µl CPY stock, diluting in 30 mM ammonium citrate, pH 6.1. 3. Prepare four serial 1:3 dilutions of the 2 × 10–3 U/µl CPP stock, diluting in 30 mM ammonium citrate, pH 4.5. 4. Prepare MALDI matrix by reconstituting in MALDI matrix diluent per manufacturer’s instructions. If kit is not used, a suitable solution is 5 mg/ml matrix in 50% acetonitrile, 0.05% TFA. Check enzyme activity by digesting and mass analyzing the peptide sequencing standard.
Chemical Analysis
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Spot wells 5. Spot 6 wells of the MALDI plate with 0.5 µl of 1 pmol/µl ACTH 18-39 peptide sequencing standard solution. Add nothing to well 1; this will serve as the no-enzyme control. 6. Add the following volumes of CPY solutions to the corresponding wells: add to well 2: 0.5 µl of 2 × 10–3 U/ µl CPY stock solution add to well 3: 0.5 µl of 1:3 CPY dilution add to well 4: 0.5 µl of 1:9 CPY dilution add to well 5: 0.5 µl of 1:27 CPY dilution add to well 6: 0.5 µl of 1:81 CPY dilution. 7. Repeat steps 5 and 6 using the CPP solutions. 8. Spot 0.5 µl of 1 pmol/µl ACTH 18-39 and 0.5 µl of 1 pmol/µl angiotensin I calibration standards in separate clean wells. 9. Allow the reaction mixtures to evaporate to dryness. Drying should take 5 to 10 min; if the mixtures evaporate in less time, the enzyme digestion may be too limited. Refer to Troubleshooting.
10. Add 0.5 µl matrix solution to each sample and standard well and allow to dry. Visually inspect plate to ensure wells are dry. Acquire spectra, analyze and assign residues 11. Acquire one spectrum from the calibration standards. Calibrate the instrument using the appropriate average or monoisotopic masses (Table 11.8.2). The spectra for the CPY peptide sequencing standard should show a continuous sequence of at least 8 residues with no gap in the sequence. CPY is fully active if 8 residues are digested from the C-terminus. The spectra for the CPP peptide sequencing standard (ACTH 18-39) should show production of a peak with a mass of about 2188 by the more dilute enzyme solutions and both a 2188 mass peak and a 2075 mass peak by the more concentrated solutions. A sequence gap for the first two C-terminal residues is normal for CPP. CPP is fully active if three residues are digested from the C-terminus.
12. Acquire at least 3 spectra per well for the digestion series, including a no-enzyme control. 13. Calculate the peak masses. Set the threshold high enough to detect only the peaks of interest or manually mark the peaks. 14. Either use sequence analysis software to automatically assign residues or perform a manual analysis (see UNIT 16.7 for ladder sequencing by mass spectrometry). Most mass spectrometer manufacturers provide software for performing sequence analysis by ladder methods. Data Explorer and DEcode Sequence Analysis are two packages available from Applied Biosystems for use with the VoyagerBiospectrometry MALDI instrument.
Table 11.8.2
Calibrant
C-Terminal Sequence Analysis
Angiotensin I ACTH 18-39
Calibration Standards for C-Terminal Sequencer
Monoisotopic mass (Da) Average mass (Da) 1296.68 2465.2
1297.51 2466.72
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Digest the peptide of interest 15. Digest the peptide of interest by spotting 0.5 µl of the sample followed by addition of varying dilutions of enzyme (step 6). 16. Allow the reaction mixtures to evaporate to dryness. Drying should take 5 to 10 min; if the mixtures evaporate in less time, the enzyme digestion may be too limited. Refer to Troubleshooting.
17. Add 0.5 µl matrix solution to each filled well and allow to dry. Visually inspect plate to ensure wells are dry. 18. Acquire spectra and analyze as described in steps 11 to 14. C-TERMINAL SEQUENCING USING CARBOXYPEPTIDASES FOLLOWED BY AMINO ACID ANALYSIS
BASIC PROTOCOL 3
This protocol assumes that an unlimited quantity of protein is available. It can also be performed for some proteins with as little as 300 pmol of protein (Michel et al., 1993). This protocol describes a reaction using carboxypeptidases P and Y. Other carboxypeptidases may be used in the same way, provided the appropriate buffer is used (see manufacturer’s instructions). Materials Purified protein Digestion buffer: 100 mM N-ethylmorpholine (NEM, Aldrich), adjusted to pH 6.0 with acetic acid; store at 4°C Aminobutyric acid (Sigma) Carboxypeptidases A, B, P, and/or Y (20 µg/vial sequencing grade, Boehringer Mannheim) Trifluoroacetic acid (TFA, Fluka) Titertubes with plugs (Bio-Rad) 5-kDa-MWCO and (optionally) 30- and 10-kDa-MWCO ultrafree filtration devices (Millipore) Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and amino acid analysis (UNIT 3.2) Prepare sample, controls, and enzymes 1. Dissolve 4 mg (∼40 nmol) purified protein to a concentration of 1 mg/ml in digestion buffer. Heat 3 min at 100°C to denature the protein. The protein concentration should be as high as possible. Using 100 mM N-ethylmorpholine, pH 6.0, as the digestion buffer provides good reaction conditions for carboxypeptidases P and Y.
2. Add 400 ng (∼4 nmol) aminobutyric acid to the 4-ml protein solution to monitor recovery. Also add 100 ng (∼1 nmol) aminobutyric acid to 1 ml digestion buffer. Aminobutyric acid is used as an internal standard, as it is not usually found in proteins. Norleucine may also be used (see Critical Parameters).
3. Add 1 ml protein solution to each of three reaction Titertubes labeled 1, 2, and 3. Add 200 µl protein solution to each of three control Titertubes labeled 4, 5, and 6. Titertubes are preferred because they tend to give high recoveries with low amounts of protein. Chemical Analysis
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4. Dissolve one vial (20 µg) each of carboxypeptidases P and Y in separate 50-µl aliquots of water. 5. Heat 10 µl of each enzyme 3 min at 100°C to inactivate the enzyme. Cool to room temperature. Digest the protein 6. Add 20 µl carboxypeptidase P to tubes 1 and 2. Add 20 µl carboxypeptidase Y to tubes 2 and 3. Add 5 µl inactivated carboxypeptidase to tubes 5 and 6 as controls. Incubate at room temperature. Tube 4 serves as the protein-only (no-peptidase) control. 7. Remove 200-µl aliquots from reaction tubes 1, 2, and 3 at 1, 10, 60, and 240 min and place the aliquots into 22 µl TFA to terminate the reaction. Similarly, remove 200-µl aliquots from control tubes 4, 5, and 6 at 240 min only and quench into 22 µl TFA. Analyze released amino acids 8. Pass each aliquot through a 5-kDa-MWCO ultrafree filtration device to separate released amino acid residues from the shortened peptide and the carboxypeptidases. A combination of 30-kDa-, and 50-kDa-MWCO filters may be used to separate large proteins from the enzyme(s) and amino acids (see Critical Parameters).
9. Wash the retentate on the filter with 1 vol digestion buffer. 10. Collect the combined filtrates (containing the amino acid residues) and use either rotary evaporation or lyophilization to dry them completely. 11. Optional: Wash the retentate with digestion buffer or other suitable buffer and retain this sample, which contains protein and carboxypeptidases, for future analysis by SDS-PAGE (UNIT 10.1) or comparative peptide mapping (see Troubleshooting). Store at –20°C. 12. Reconstite the dried filtrate in appropriate sample buffer, and then perform amino acid analysis on all samples and controls. For each amino acid, plot amount recovered versus time of digestion (see Fig. 11.8.1). From the preliminary experimental data, choose a time for quantitative digestion that produces >50% of expected yield for at
Absorbance
1.20 1.00
Ile
0.80
Tyr
0.60
Glu
0.40
Pro
0.20
His
0.00 0.00
1.00
2.00
3.00
4.00
Time (hr)
C-Terminal Sequence Analysis
Figure 11.8.1 Manual C-terminal sequence analysis of a protein ending in the amino acids His-Pro-Glu-Tyr-Ile.
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least one amino acid and intermediate times that enable discrimination of rates of release. If rates of release are extremely slow, the ratio of enzyme to substrate or the incubation temperature can be increased. Alternatively, the reaction can be slowed by decreasing the temperature or the amount of enzyme.
COMMENTARY Background Information Automated methods A novel method for C-terminal sequence analysis using automated chemical degradation was introduced by Boyd and coworkers in 1992. Since then, major developments presented by Applied Biosystems include the ability to sequence samples electroblotted onto PVDF membranes, to sequence most amino acid residues, and to determine longer stretches of C-terminal sequence (up to ∼10 residues, with an average of 3 to 5 residues, depending on the sequence). The chemistry employed in the Procise C sequencer instrument (Applied Biosystems) involves an initial one-time activation cycle to render the C-terminus susceptible to derivatization into an amino acid thiohydantoin using thiocyanate anion. Selective alkylation of the thiohydantoin at the sulfur atom follows, and thereafter, cleavage of the alkylated thiohydantoin with the thiocyanate anion under acidic conditions. The instrument performs stepwise sequence analysis with an average initial yield of 15% and repetitive yields of ∼70%. The improvements made by Boyd et al. (1992, 1995) using alternative protocols are (1) more efficient cleavage of the thiohydantoin derivative when the thiohydantoin is S-alkylated and (2) simultaneous cleavage and derivatization/cyclization of the newly formed Cterminus in the same step, which eliminates the need to return to a free carboxy terminus. Manual methods Manual methods of determining C-terminal sequences have relied on enzymatic digestion with carboxypeptidases followed by analysis of the released amino acids by conventional amino acid analysis. Carboxypeptidases are exopeptidases that remove amino acids one at a time from the C-terminus of the protein. Ideally, amino acids are released in order of their position from the C-terminus; thus the C-terminal sequence is deduced from the rate of appearance of released amino acids. In practice, the rates of release are strongly influenced by the nature of the side-chains of both terminal
and penultimate amino acids, as well as by the general protein structure. Thus, the kinetics of release are highly dependent on the nature of the particular protein and in the most unfavorable situations will not yield interpretable sequence data. Recent methods using mass spectrometry to examine the “ladder” of partially digested peptides in the reaction mixture (Rosnack and Stroth, 1992; Patterson et al., 1995) offer a means of identifying the C-terminal sequence that avoids the problems inherent in monitoring digestion kinetics. The mass spectrometry method (Basic Protocol 2) is more rapid than the amino acid analysis method (Basic Protocol 3) and requires significantly less material, ∼1 to 23 pmol. It is limited to proteins small enough to be able to accurately distinguish 1 to 2 mass unit differences. For larger proteins, isolation of the Cterminal peptide is necessary prior to mass analysis. Analysis of the digestion products by amino acid analysis requires on the order of 40 nmol of protein. The procedure requires more time to perform, but offers a more quantitative assessment of the C-terminus and an opportunity of studying larger proteins without degrading them to peptides. Carboxypeptidases have a wide range of specificity for individual amino acids and therefore cleave different C-terminal amino acid residues at very different rates. Carboxypeptidase B predominantly cleaves Lys and Arg residues. Carboxypeptidases A, P, and Y cleave most amino acids, but the rates of cleavage are reduced significantly by the presence of Gly residues. Carboxypeptidase A will not cleave Pro or Arg residues. Cleavage rates for carboxypeptidases P and Y are also greatly retarded by Ser and Asp, respectively. Enzymes can be used individually or combined to provide further sequence information and to help with amino acid assignments. Table 11.8.3 summarizes information on the rates of release of particular amino acids by various carboxypeptidases. The four carboxypeptidases (A, B, P, and Y) described are commercially available; the protocols for their use are straightforward and
Chemical Analysis
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Table 11.8.3
Carboxypeptidase A
B P
Relative Rate of Release of Amino Acids by Carboxypeptidases
Release ratea
References
Noneb
Slow
Y,F,W,L,I,M,T Q,H,A,V,Hsr, N,S,K,MetSO2 K,R
G,D,E,CysSO3H, P,R,Hypro CMCys
Ambler, 1972
-
-
D,T
S,G
All except K,R, Penultimate P Penultimate P,G
P,A,E,S,T,Q, N,CMCys
D,H,R,K,G
Penultimate G
R,A,N,E,Q,H,I,L, K,M,F,P,W,Y,V Y,F,W,L,I,M,V
Y
Very slowb
Rapid
Ambler, 1972 Hofmann, 1976; Lu et al., 1988 Jones, 1986
aAmino acids are indicated using single-letter abbreviations (see Table A.1.A.1). Modified amino acid abbreviations: CysSO H, cysteic acid; 3 CMCys, carboxymethyl cysteine; Hsr, homoserine; Hypro, hydroxyproline; MetSO2, methionine sulfone. bThe presence of one of these amino acids in the penultimate position generally decreases the rate of release of the terminal amino acid.
quick. The digestion products may be analyzed by amino acid analysis or mass spectrometry of the truncated peptides (see Chapter 16). Thus, manual sequencing using enzymatic digestion offers a relatively low-cost method of obtaining C-terminal sequence information without the need for a dedicated instrument. Analysis of the digestion products by mass spectrometry produces a ladder of masses in which the product of each step differs from the next by the mass of the removed amino acid. This method is generally applied to peptides rather than intact proteins in order to more easily resolve the various mass species. The C-terminal peptide may be isolated from an intact protein by several methods. A typical method uses an anhydrotrypsin column to enrich for the C-terminal peptide by removing internal peptides generated by trypsin or endoproteinase lysine-C (Sechi and Chait, 2000). Anhydrotrypsin selectively binds peptides with C-terminal lysine or arginine. Only the peptide originating from the C-terminus of the parent protein will flow through the column.
Critical Parameters Automated method (see Basic Protocol 1)
C-Terminal Sequence Analysis
Chemicals and separation of amino acid derivatives. The chromatographic background in the reversed-phase analysis of alkylated thiohydantoin (ATH) amino acids from C-terminal degradation is lowered and the baseline improved if chemicals and reagents are diluted (Table 11.8.1). The sensitivity is increased, allowing 10 pmol of ATH amino acid standard to be used
(Fig. 11.8.2). To block the amino groups, pretreatment with 1% PIC is employed and one treatment is sufficient. For separation of ATH amino acids, the manufacturer’s program has been modified. Ethanol is used instead of tetrahydrofuran (THF) in both solvents A and B (Table 11.8.1). Ethanol affects the elution time for most ATH amino acids and the ethanol concentration can be varied to optimize the separation, e.g., when a new column is used. The separation of ATHAsn and ATH-Gln is improved and baseline resolution is achieved for the pairs ATHAsp/ATH-Glu and ATH-Trp/ATH-Thr (Fig. 11.8.2) if ethanol is used instead of THF. Ethanol is also less toxic and less hazardous than THF, which can create peroxides and the risk of explosion. Acetone is used in solvent A to balance the baseline (DIEA is excluded because it tends to generate a U-shaped baseline at high sensitivity), improving the chromatographic behavior. The column temperature is decreased from 45°C to 40°C, which results in better resolution and stability for the ATH amino acids, in particular, that of ATH-Ser. The recovery of ATHSer and ATH-Thr is further improved by lowering the temperature of the transfer flask, from 45°C to 40°C. To reduce the level of byproducts from degradation and alkylation, the sequencer program is modified to include three extra washing steps in each cycle with ethyl acetate. By employing four instead of a single ethyl acetate wash, the chromatography is clean with a low background.
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A257 (mAU)
Time (min)
Figure 11.8.2 Chromatography of ATH standard amino acids (10 pmol). The abbreviation, nmtc, denotes naphtylmethylthiocyanate, the major byproduct formed during sequence analysis.
Polypeptides Over 600 proteins and fragments from 10 to 600 residues have been analyzed using the modified protocol. The average determination is 5 residues, but extended degradations have also been achieved of up to 15 residues and longer. Samples are applied to PVDF membranes either via electroblotting from gel separations followed by Coomassie staining and excision, or via direct application of sample in solution using a ProSorb sample preparation cartridge. The amount of sample that can be analyzed with these formats varies from a few hundred pmol to 10 pmol. The sequencer recoveries and lengths of degradation are very much sequence-dependent. In particular, Pro and acidic or hydroxylated residues can lower the yields or stop degradation altogether. However, using the modified protocol, passage of Pro residues and identification of the next residue in the C-terminal sequence has been demonstrated.
Blot cartridge Use of a blot cartridge (Applied Biosystems) with a vertical slit for the PVDF membrane, instead of the standard horizontal slit, results in increased sensitivity. In the blot cartridge, the liquid flow is along the entire length of the membrane instead of perpendicular to it, which results in more efficient solvent extraction and penetration of reagents. The increased liquid hold-up time improves the recovery of ATH amino acids, resulting in better sensitivity. This type of cartridge was originally designed for N-terminal sequence analysis of electroblotted samples and now provides an overall increased C-terminal sequencer yield, which is important in combined N- and C-terminal sequence analysis (Fig. 11.8.3). It is particularly important not to apply more PVDF membrane material to the sequencer cartridge than corresponding to a normal protein band excised from two to three electrophoretic lanes. Otherwise, there is a substantial risk that the liquid flow through the blot cartridge will be affected or even blocked. It is also
Chemical Analysis
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B
A269 (mAU)
A257 (mAU)
A
Time (min)
Time (min)
Figure 11.8.3 Combined N- and C-terminal sequence analysis of 100 pmol of a 28-kDa protein. The sample was first subjected to N-terminal degradation for 5 cycles (A), followed by C-terminal analysis for an additional 5 cycles (B). The letter K in B denotes the ATH-PTC-Lys derivative or a breakdown product thereof. The combined sequence result is H-Thr-Ile-Val-Pro-Ala- - - - -Leu-PheIle-Gln-Lys-OH.
crucial to completely dry the PVDF membrane with the immobilized sample before PIC treatment so that no moisture is left. When analyzing small peptides, it is necessary to apply more sample than what is required for large proteins that bind well to the PVDF (up to two to three fold more) in order to compensate for wash-out losses. With regard to reagents, C4 (piperidine thiocyanate/acetonitrile) and C6 (acetic anhydride/lutidine/acetonitrile) bottles should be replaced after no more than 4 weeks in the instrument so as to minimize the risk of degradation and suboptimal sequencer performance.
C-Terminal Sequence Analysis
Manual sequencing (see Basic Protocols 2 and 3) Enzymes and specificities. Several sequencing-grade carboxypeptidases are available from commercial vendors. The substance specificities are diverse enough that at least one enzyme or a combination of enzymes should be able to remove amino acids from the C-terminus of almost any protein. Mixtures of two or more carboxypeptidases can initiate further digestion that can provide insight into the sequence of a protein. An enzyme/substance ratio of 1:100 is usually sufficient; however, altering the ratio is one easy way of manipulating the reaction rate (Ambler, 1972). Most carboxypeptidases have significant catalytic activity at 25°C as well as 37°C (Lu et al., 1988), and the reaction tem-
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perature can also be modified to change the reaction rate as desired. Stopping the reaction can be easily accomplished by adding enough trifluoroacetic acid (TFA) to the reaction mixture to bring the pH to 2.5 to 3.0 (adding TFA to 10% of final volume is usually sufficient to inactivate the enzyme; Lu et al., 1988). Buffer system. When mass spectrometry is used for analysis of the digestion products, the digestion buffer must be suitable for mass analysis. Ammonium salts are frequently used for this application. Buffer salts also interfere with amino acid analysis; when amino acid analysis is used as the read-out, volatile buffers that do not contain ammonia are most suitable. N-ethylmorpholine (NEM) is commonly used with acetic acid to provide a volatile buffering system compatible with most carboxypeptidases. A 200 mM NEM/acetic acid buffer is usually sufficient, but it is best to follow the enzyme supplier’s recommendations for ionic strength and pH conditions. Inappropriate digestion buffer conditions are a typical cause for lack of digestion. Denaturing protein. Unless the C-terminus is known to be accessible in the protein’s native state, denaturation is usually recommended. Product literature from enzyme suppliers lists the compatibility of denaturants with enzymatic activity. Disulfide bridges are usually not problematic during exopeptidase digestion. If the C-terminus is not accessible after addition of denaturants, reduction and alkylation of the disulfides should be considered. Time course. The classic limitation of carboxypeptidases is that the rate of release is highly dependent on the amino acid side chains of both the terminal and penultimate amino acids. For this reason, it is highly recommended that a preliminary experiment be performed to provide an initial understanding of the kinetics of the reaction for a particular protein. When mass analysis is the read-out, the kinetic parameters are best adjusted by altering the amount of enzyme added, since the carboxypeptidase is unlikely to have the same mass as the sample. When amino acid analysis is the read-out, the kinetic experiment is best performed by altering the digestion time. Suggested time points for a preliminary experiment are 0, 1, and 5 hr. These suggested time points should provide enough information to choose time points for the quantitative experiment; the selected time points should enable discrimination of the rates of release of several amino acids and provide a final yield of amino acid that is a substantial portion of the input substance (i.e.,
>50%). In general, the reaction is sampled more frequently during the initial stages. Digestion for up to 16 to 24 hr may be required. Preparing samples for amino acid analysis. Previous investigators have used precipitation with trichloroacetic acid (TCA) or sulphosalicylic acid to separate the protein from the released amino acids (Mondino et al., 1972). Alternatively, disposable filtration units are available with different membranes that provide different specific molecular weight cutoffs. Ultrafree units (Millipore) are designed to filter ∼350 µl in a standard microcentrifuge. Depending on the size of the protein under analysis, one round of filtration through a 5kDa-MWCO filter, followed by a single washing step, can very effectively separate amino acids from protein. If the protein under scrutiny is particularly large, it may be necessary to pass the sample first through a wide-pore (e.g., 30kDa) filter and then through a 5-kDa filter. If protein of interest is significantly different in molecular weight from the carboxypeptidase used, then sequential filtration can also potentially yield relatively pure digested protein sample free of carboxypeptidase contamination. Carboxypeptidase Y, for example, has a molecular weight of 61 kDa. By passing the mixture of carboxypeptidase Y (61 kDa), lowmolecular-weight protein (e.g., 25 kDa), and amino acids (<0.2 kDa) first through a 30-kDa filter and then through a 5-kDa filter, each component can be purified. Controls. A no-enzyme or inactivated enzyme control should always be included. When using amino acid analysis as the read-out, it is particularly important to run a control of inactivated enzyme alone because sequencinggrade enzymes contain salts and possibly impurities that create artifacts, which are most easily identified by comparing the controls to the samples. Internal standards are essential for interpreting the amino acid analysis data. An internal standard is added to the initial reaction mixture at an amount usually equal to 10% of the theoretical amount of each amino acid to be released; this places the standard in the range of the released amino acids (∼10% of the maximum theoretical yield is usually recovered). Aminobutyric acid and norleucine are suitable amino acid standards that are not usually found in proteins.
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Troubleshooting Manual sequencing using carboxypeptidases followed by MALDI analysis If digestion does not occur or there are gaps in the sequence, confirm that the reaction mixtures did not evaporate too quickly. Drying may be slowed by adding 1 to 2 µl of HPLC-grade water to the well. Also confirm that the enzyme is active. Lack of digestion may also reflect the fact that the digestion rate of some amino acid sequences is pH dependent, while other sequences are resistant to particular enzymes. Therefore, examine the effects of pH, time, temperature, and the amount of enzyme on the digestion. In performing these experiments, it is important to ensure that the samples do not dry prematurely. If these adjustments fail to result in a digested sample, it may be necessary to use a different carboxypeptidase or a combination of enzymes. If the peptide is digested so completely that no parent ion is present, slow the digestion by diluting the enzyme. If matrix crystallization is poor, ensure that the pH of the matrix is below 3; the trifluoroacetic acid level in the matrix may be increased to lower the pH. Poor crystallization may result from degraded matrix or frozen matrix; in this case, prepare fresh matrix using 10 mg/m l á-cyano-4-hydroxy cinnamic acid in 50% acetonitrile/0.3% trifluoroacetic acid. Unidentified or unexpected peaks in the mass spectrum with a ∆mass of 22 or 38 Da from the parent are generally Na+ or K+ adducts. Desalting the sample or increasing the trifluoroacetic acid may decrease formation of these adducts. Alternatively, they may be ignored. Similarly, peaks with a ∆mass of 16 typically arise by oxidation of methionine and/or tryptophan. Unidentified peaks of other masses may result from degraded enzyme or degraded matrix. Enzyme degradation may be assessed by performing mass analysis on the enzyme alone.
C-Terminal Sequence Analysis
Manual sequencing using carboxypeptidases followed by amino acid analysis If no amino acids are detected by the preliminary experiment, the first suggestion is to perform a new digestion with a different enzyme or a combination of enzymes. If a combination of enzymes is used, the active pH ranges for the enzymes must overlap. Most
often, the use of a different enzyme or a combination of enzymes will expose the reason(s) for failure of the preliminary experiment. If the digested protein from the experiment was not discarded, it can be analyzed by comparative peptide mapping (Isobe et al., 1986). Peptide maps on both the intact and digested proteins can usually be used to infer which peptide or peptides in the chromatogram is the C-terminal peptide, based on the loss or decrease in one or more peaks and the appearance of one or several other peaks. This peptide can, of course, be collected and analyzed further. An additional method for C-terminus identification is the use of a phenylenediisothiocyanate (DITC) membrane or resin in combination with Endo-Lys C digestion. The DITC moiety binds to peptides with a C-terminal Lys. Shimadzu (see SUPPLIERS APPENDIX) has recently introduced a method that uses a resin to which the DITC moiety has been bound (Nokihara, 1992). Eluting the unbound peptide is easily accomplished, and subsequent analysis (e.g., mass spectrometry and Edman degradation) can be performed immediately. Aside from the automated technique, the most promising method for obtaining C-terminal sequence data is the use of mass spectrometry (MS; see Chapter 16) coupled with “laddergenerating” techniques (Patterson et al., 1995), usually either carboxypeptidase digestion (see Basic Protocol 3; Rosnack and Stroh, 1992) or C-terminal chemical truncation (Takamoto et al., 1995). Recent improvements in the accuracy and feasibility of MS procedures have allowed for accurate mass determinations of mixtures of protein molecules differing by only one amino acid. The C-terminal sequence of β-lactoglobulin, an 18,800-Da protein, has been determined using carboxypeptidase Y digestion followed by MALDI analysis (Patterson et al., 1996). There are several limitations with this type of method; one is that the accuracy of the information obtained decreases quickly with increasing protein size, so that it is currently most promising for peptides. This technique can be performed without the use of a truncation agent to assess the level of carboxyterminal heterogeneity (Horn et al., 1995).
Anticipated Results As little as 10 pmol of polypeptides ranging in size from 10 to 600 residues (or above) can be analyzed using the automated system described in Basic Protocol 1. The technique allows for determination of up to 15 residues using 100 to 500 pmol. Initial C-terminal se-
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A 2466.72
No CPP (well 6) 2465.86
2189.51 Dilution 5 (well 5) 2189.43 Dilution 4 (well 4)
2076.44 2188.94
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1632.55 1562.43 1761.87 1832.90
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1561.53 1633.01
Dilution 1 1400
1600 Ala
1800 Glu
Ala
m/z (Da) Phe
2000 Pro
2200 Leu
2400 Glu
2600
Phe
Figure 11.8.4 Expected mass spectral results for C-terminal sequence analysis of peptide sequencing standard. The ACTH 18-39 peptide sequencing standard was digested with varying dilutions of carboxypeptidase P (A) or Y (B). The resulting digests were analyzed on a Voyager Biospectrometry Workstation. Figure reproduced with permission from Applied Biosystems.
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quencer yields range from 10% to 50% for proteins and 5% to 30% for peptides. The repetitive yield is typically 50% to 80% for proteins and 30% to 50% for peptides. Passage of Pro-residues is possible with identification of the subsequent residue in the C-terminal sequence. Manual sequencing using carboxypeptidases followed by MALDI mass analysis Mass analysis of digestion mixtures should produce a ladder of masses decreasing from the mass of the parent species. The difference in mass between two adjacent peaks corresponds to the mass of the released amino acid. Results from sequencing the peptide adrenocortocotropic hormone 18-39 are shown in Figure 11.8.4. Manual sequencing using carboxypeptidases followed by amino acid analysis The data from manual C-terminal sequencing are usually expressed according to the following function (Hayashi, 1977): [AA released]theoretical =
[standard ] theoretical [AA released]found × [standard ]found and are plotted with nanomoles released on the y axis and time on the x axis (see Fig. 11.8.1). This plot is a best-case scenario for a protein that ends in His-Pro-Glu-Tyr-Ile. In practice, the data can be more difficult to infer because of the different reaction rates. Also, loss of individual amino acids, presence of repetitive amino acids in a sequence, and certain other factors increase the difficulty in making specific sequence assignments using only this method. However, the method is particularly useful to confirm an automated analysis or other data suggesting the C-terminal sequence. It is also particularly useful in comparing the C-terminal sequence of preparations of the same material.
Time Considerations
C-Terminal Sequence Analysis
Sample application and washing using a ProSorb sample-preparation cartridge takes ∼10 min. If the sample is applied via electroblotting to the PVDF membrane, the time required varies depending on the particular protocol, but on average, this process including staining/destaining takes 2 to 4 hr. Drying of the PVDF membrane with an immobilized
sample takes ∼20 min and the PIC treatment 5 min. The first C-terminal sequencer cycle including blank and standard (Fig. 11.8.2) takes 4.5 hr. Subsequent cycles each take 75 min. Manual C-terminal sequencing using mass analysis typically requires only 30 to 60 min. Manual C-terminal sequencing using quantitative amino acid analysis can be easily performed in the course of 1 day including an overnight automated amino acid analysis of the samples.
Literature Cited Ambler, R.P. 1972. Enzymatic hydrolysis with carboxypeptidases. Methods Enzymol. 25:143-154. Bergman, T., Cederlund, E., and Jörnvall, H. 2001. Chemical C-terminal protein sequence analysis: Improved sensitivity, length of degradation, proline passage, and combination with Edman degradation. Anal. Biochem. 290:74-82. Bergman, T., Cederlund, E., and Jörnvall, H. 2003. Carboxy-terminal sequence analysis. In Proteins and Proteomics: A Laboratory Manual. (R. Simpson, ed.) Cold Spring Harbor Laboratory Press. Cold Spring Harbor. N.Y. pp. 310-317 and 332-335. Boyd, V.L., Bozzini, M., Guga, P.J., DeFranco, R.J., and Yuan, P.-M. 1995. Activation of the carboxy terminus of a peptide for carboxy-terminal sequencing. J. Organic Chem. 60:2581-2587. Boyd, V.L., Bozzini, M., Zon, G., Noble, R.L., and Mattaliano, R.J. 1992. Sequencing of peptides and proteins from the carboxy terminus. Anal. Biochem. 206:344-352. Geng, M., Zhang, X., Bina, M., and Regnier, F. 2001. Proteomics of glycoproteins based on affinity selection of glycopeptides from tryptic digests. J. Chromatography B 752:293-306. Gheorghe, M.T., Jörnvall, H., and Bergman, T. 1997. Optimized alcoholytic deacetylation of N-acetyl-blocked polypeptides for subsequent Edman degradation. Anal. Biochem. 254:119-125. Hayashi, R. 1977. Carboxypeptidase Y in sequence determination of peptides. Methods Enzymol. 47:84-93. Hofmann, T. 1976. Penicillocarboxy peptidases S-1 and S-2. Methods Enzymol. 45:587-599. Horn, M.J., Mayhew, J.W., and O’Dea, K.C. 1995. A method for the characterization of the C-terminus of peptides and proteins. Protein Sci. 2:155. Isobe, T., Ichimura, T., and Okuyama, T. 1986. Identification of the C-terminal portion of a protein by comparative peptide mapping. Anal. Biochem. 155:135-140. Jones, B.N. 1986. Microsequence analysis by enzymatic methods. In Methods of Protein Microcharacterization. (J.E. Shively, ed.) pp. 337-361. Humana Press, Totowa, N.J. Jonsson, A.P., Griffiths, W.J., Bratt, P., Johansson, I., Strömberg, N., Jörnvall, H., and Bergman, T.
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2000. A novel Ser O-glucuronidation in acidic proline-rich proteins identified by tandem mass spectrometry. FEBS Lett. 475:131-134. Lu, H.S., Klein, M.L., and Lai, P.H. 1988. Narrowbore high-performance liquid chromatography of phenylthiocarbamyl amino acids and carboxypeptidase P digestion for protein C-terminal sequence analysis. J. Chromatogr. 447:335-364. Michel, G., Chauvet, M.T., Clarke, C., Bern, H., and Acher, R. 1993. Chemical identification of the mammalian oxytocin in a holocephalian fish, the ratfish (Hydrolagus colliei). Gen. Comp. Endocrinol. 92:260-268. Mondino, A., Bongiovanni, G., Fumero, S., and Rossi, L. 1972. An improved method of plasma deproteination with sulphosalicylic acid for determining amino acids and related compounds. J. Chromatogr. 74:255-263.
Key References Boyd et al., 1992. See above. Describes the application of the thiohydantoin chemistry to a sequencer instrument and presents the one-time-activation and S-alkylation for improved yield and recovery. Bergman et al., 2001. See above. Describes the current modifications of the original sequencer system (Boyd et al. 1992, 1995) and provides details about performance and applications. Jones, B.N. 1986. See above. A detailed discussion of enzymatic methods for Cand N-terminal sequence determination; includes examples and information about enzyme properties, kinetic studies, and analytical methods.
Patterson, D.H., Tarr, G., Regnier, F.E., and Martin, S.A. 1995. C-terminal ladder sequencing via matrix-assisted laser-desorption mass spectrometry coupled with carboxypeptidase Y timedependent and concentration-dependent digestions. Anal. Chem. 67:3971-3978.
Patterson, et al. 1995. See above.
Patterson, D.H., Tarr, G.E., Hine, W.M., Vestal, M.L. and Martin, S.A. 1996. Proceedings from the 1996 ASMS conference.
Ward, C.W. 1986. Carboxy terminal sequence analysis. In Practical Protein Chemistry: A Handbook (A. Darbee, ed.) pp. 517-522. John Wiley & Sons, New York.
Rosnack, K.J. and Stroh, J.G. 1992. C-terminal sequencing of peptides using electrospray ionization mass spectrometry. Rapid Commun. Mass Spectrom. 6:637-640. Sechi, S. and Chait, B.T. 2000. A method to define the carboxyl terminal of proteins. Anal. Chem. 72:3374-3378. Stark, G.R. 1968. Sequential degradation of peptides from their carboxyl termini with ammonium thiocyanate and acetic anhydride. Biochemistry 7:1796-1807.
Comparison of results generated by liquid phase and on-plate digestions; includes discussion of a statistical analysis strategy for ladder sequencing data.
General description of the techniques and carboxypeptidases.
Contributed by Tomas Bergman, Ella Cederlund, and Hans Jörnvall Karolinska Institutet Stockholm, Sweden Elizabeth Fowler Millennium Pharmaceuticals Cambridge, Massachusetts
Acknowledgements: This work was supported in part by grants from the Swedish Research Council (projects 03X-3532, 03X-10832, and K5104-20005891), the Swedish Cancer Society (project 4159), the European Commission (Bio4CT97-2123), and the Swedish Society of Medicine.
Chemical Analysis
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Amino Acid Analysis
UNIT 11.9
Amino acid analysis (AAA) is one of the best methods to quantify peptides and proteins. AAA also provides valuable qualitative measurement, such as quality control evaluation of synthetic peptide and recombinant protein structures, identification of proteins based on composition, and detection of odd amino acids. AAA has been used for years in primary structural studies, for example, in designing fragmentation strategies, identifying exopeptidase reaction products, and quantifying sequence samples in order to discriminate between useful data, background contamination, and N-terminally blocked samples. In addition, AAA has applications in the characterization of feed and food samples and physiological fluids. Two general approaches to quantitative AAA exist, namely, classical postcolumn derivatization following ion-exchange chromatography and precolumn derivatization followed by reversed-phase HPLC (RP-HPLC). Excellent instrumentation and several specific methodologies are available for both approaches, and both have advantages and disadvantages as outlined in Background Information. This unit focuses on picomole-level AAA of peptides and proteins using the most popular precolumn-derivatization method, namely, phenylthiocarbamyl amino acid analysis (PTC-AAA). It is directed primarily toward those interested in establishing the technology with a modest budget. The Basic Protocol describes PTC derivatization and analysis conditions, and Support and Alternate Protocols describe additional techniques necessary or useful for most any AAA method— e.g., sample preparation, hydrolysis, instrument calibration, data interpretation, and analysis of difficult or unusual residues such as cysteine, tryptophan, phosphoamino acids, and hydroxyproline. STRATEGIC PLANNING Although straightforward in principle, it is far from simple in practice to achieve picomole-level quantitative accuracy in analysis of amino acids in peptides and proteins. Before embarking on amino acid analysis (AAA) bench work, strategic consideration is encouraged regarding sample amount, work load, costs, time commitment, and resource facility options (see Background Information). With a low or intermittent need for the technology, utilizing the services of an appropriate resource facility will probably be more cost-effective and expedient than performing the analysis oneself. For those who decide to establish laboratory AAA capability, the following strategic issues should be considered prior to bench work. Most AAA methods involve the following steps: (1) peptide or protein sample preparation; (2) hydrolysis of the peptide or protein into amino acids; (3) derivatization of the amino acids for detection; (4) calibration of the HPLC system with standard amino acids and chromatographic analysis of the experimental hydrolysate; and (5) calculations and data interpretation. For postcolumn derivatization AAA, steps 3 and 4 are reversed. Various chemicals can be used for derivatizing the amino acids, but all five steps must succeed for high-quality results. AAA is complicated by the variable susceptibility of amino acids and peptide bonds to acid hydrolysis, variable levels of background contamination, and potentially variable instrument and human performance. Overall success and useful results require meticulous work habits, reproducible instrument performance, and care at each step of the procedure. Satisfactory instrument performance will be easier to maintain if an HPLC system is dedicated to the analysis of amino acids rather than used part-time for some other purpose. Chemical Analysis Contributed by John W. Crabb, Karen A. West, W. Scott Dodson, and Jeffrey D. Hulmes Current Protocols in Protein Science (1997) 11.9.1-11.9.42 Copyright © 1997 by John Wiley & Sons, Inc.
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Furthermore, if a significant, long-term need for the technology exists, a dedicated PTC amino acid analyzer supported by an instrument vendor is strongly recommended. The most expensive piece of equipment is the HPLC system, and the available budget will dictate HPLC options; a simple gradient HPLC system with a recording integrator will suffice; however, an automatic sample injector is also recommended. This technology is labor-intensive. Successful AAA requires technical, mechanical, analytical, and quantitative skills, as well as common sense. Laboratory experience with general protein chemistry techniques and with HPLC instrumentation is very useful. Sample Preparation To obtain the most accurate PTC-AAA data, peptide and protein samples should be homogeneous (i.e., exhibit a single SDS-PAGE band and/or a single N-terminal amino acid). In addition, the sample should be dissolved in a volatile solvent, free of salts and detergents. Common volatile solvents useful for transferring readily soluble samples to hydrolysis tubes include water (HPLC grade), aqueous 0.1% trifluoroacetic acid/acetonitrile, 30% to 50% methanol, ethanol, or acetonitrile, 0.1% to 1% N-ethylmorpholine acetate, pH 8.5, and dilute (0.1%) HCl. Less soluble samples can be dissolved in 75% to neat organic acids such as formic, acetic, and trifluoroacetic acids. Whenever possible, enough sample should be prepared for duplicate analyses per hydrolysis (plan on ∼0.5 to 1.0 µg per analysis). Useful data can be obtained with <0.5 µg of sample, but ubiquitous contamination makes routine analyses below this range more labor-intensive and difficult for any AAA procedure. Useful PTC-AAA data can be obtained from some dilute salt solutions (Dupont et al., 1989). For example, samples dried from 1 mM MOPS, pH 7 (3-[N-morpholino]propanesulfonic acid), or from 25 mM Tris (pH 7.2)/1 mM DTT/1 mM EDTA can yield high-quality data provided the total salt in the hydrolysis/PTC derivatization reaction is kept low (e.g., <50 µg per hydrolysis). Several specific methods for sample preparation are described: reversed-phase HPLC (see Support Protocol 1), micropreparation concentration (see Support Protocol 2), microdialysis (see Support Protocol 3), acetone preciptation (see Support Protocol 4), and ether precipitation (see Support Protocol 5). As an aid for quantitation, a defined amount of an internal standard may be added to the sample prior to hydrolysis and/or derivatization. Subsequent data analysis is used to normalize recovery to the initial conditions. Use of internal standards in amino acid analysis is essential for some experimental applications, such as C-terminal sequencing with carboxypeptidases (UNIT 11.8), but in general it is optional and a matter of personal preference. Good internal standards are stable to the hydrolysis, derivatization, and analysis conditions to which they are subjected and must not coelute with any other peak in the chromatography system. Odd amino acids such as α-aminobutyric acid, norvaline, norleucine, and ornithine can be useful as internal standards in PTC-AAA. Peptide/Protein Hydrolysis
Amino Acid Analysis
Key to the success of most AAA is the hydrolysis of peptides and proteins into free amino acids. Complete hydrolysis is generally performed by heating the sample in constant-boiling HCl; however, variable lability of the residues complicates the process, and only sixteen of the common twenty amino acids are usually measured from HCl hydrolysates— Asp, Glu, Ser, Gly, His, Arg, Thr, Ala, Pro, Tyr, Val, Met, Ile, Leu, Phe, and Lys. For example, Trp and Cys are destroyed by standard HCl hydrolysis, Ser and Thr are partially destroyed, Ile and Val are slow to cleave, Met is subject to oxidation, and Asn and Gln are deamidated. Shorter hydrolysis times can be used to optimize recovery of Ser and Thr, and longer hydrolysis times may be used to optimally quantify Ile and Val. Cys must be
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modified for quantification by AAA (see Support Protocols 12 and 13). Special hydrolysis conditions must be used to measure Trp (see Support Protocols 14 and 15) and phosphoamino acids (see Support Protocols 16 and 17). Manual vapor-phase (see Support Protocol 7) and liquid-phase HCl hydrolysis (see Support Protocol 8) protocols work well and are widely used for analysis of the common residues in hydrolysates. The liquid-phase HCl hydrolysis method presented is a simple approach for hydrolyzing multiple samples of small amounts. Vapor-phase hydrolysis methods are now preferred by many analysts because they are more rapid than the liquid-phase methods, and they reduce the possibility of introducing background contaminants with HCl. Directions for cleaning hydrolysis tubes (see Support Protocol 6) are also provided. Microwave hydrolysis (see Support Protocol 10) and HCl hydrolysis on polyvinylidene difluoride (PVDF) membranes (see Support Protocol 9) are optional approaches. Phenylisothiocyanate Derivatization PTC-AAA typically identifies the sixteen common amino acids in peptide/protein hydrolysates, including proline. The derivatization reaction is relatively tolerant to variations in pH and molar ratios of reagents to sample, but a slow, variable degradation of the derivatives can occur in solution at room temperature. The derivatization protocol may also be successfully performed with ethanol in place of methanol, and triethylamine (TEA) substituted for N,N-diisopropylethylamine (DIEA), but resulting reagent by-product peaks will differ slightly from those shown in Figure 11.9.1 (Bidlingmeyer et al., 1984; Tarr, 1986). The Basic Protocol describes a procedure for manual PTC derivatization. Alternatively, automatic PTC derivatization with on-line HPLC analysis can be performed with instrumentation marketed by PE Applied Biosystems. This instrumentation provides savings in time and labor as well as reduction in variability due to time-dependent degradation of the PTC derivatives. Automatic on-line vapor-phase HCl hydrolysis can also be performed with the PE Applied Biosystems model 420H hydrolyzer/derivatizer. Detailed comparisons of the performance of automatic and manual hydrolysis/PTC derivatization methods and applications of the automatic PTC-AAA technology are available elsewhere (West and Crabb, 1990, 1992). Quantitative Chromatographic Analysis Reversed-phase conditions for separating PTC amino acids are well established for a number of HPLC instruments with a variety of columns; HPLC vendors are usually willing to recommend and sometimes even demonstrate specific columns, solvents, and gradient conditions useful for their HPLC system. Request HPLC system information regarding optimization of the HPLC analysis of PTC amino acids from the vendor. The PTC derivatization process normally produces UV-absorbing by-products that appear on the chromatogram (Fig. 11.9.1). These by-products, which come from reactions between PITC and other derivatization reagents and sample components, commonly include phenylthiourea (PTU, reaction with ammonia), aniline (hydrolysis product), ethylphenylthiourea (EPTU, reaction with ethylamine from DIEA), isopropylphenylthiourea (IPTU, reaction with isopropylamine from DIEA), and diisopropylphenylthiourea (DIPTU, reaction with diisopropylamine from DIEA). These chromatography peaks usually are not problematic provided they are well resolved from the amino acids. Increased amounts of by-products usually indicate that fresh derivatization reagents are needed. To minimize PTU, avoid ammonia-containing solvents, and keep ammonia vapors away from the derivatization reaction and PTC workstation. Chemical Analysis
11.9.3 Current Protocols in Protein Science
Supplement 7
In addition to the column and HPLC system, important chromatography parameters for reversed-phase resolution of PTC derivatives are acetonitrile concentration, ionic strength and pH of the solvents, and temperature of the chromatography. Constant temperature is needed for reproducibility. Increasing the acetonitrile concentration and temperature causes all the PTC derivatives to elute with shorter retention times. Lowering the solvent pH increases the retention of Asp and Glu, whereas raising the solvent pH generally causes Asp and Glu to elute sooner. High-purity solvents and reagents (HPLC grade) are essential. The elution position of all the PTC amino acids in the chromatography system should be determined before attempting to calculate response factors (i.e., identify all the peaks from analysis of Pierce Standard H). Calculations and Data Interpretation Amino acid compositional data are commonly expressed as mole % or residues per molecule. The use of a desktop computer with spreadsheet software (e.g., Microsoft Excel) greatly facilitates amino acid analysis data reduction. Mole % (i.e., residues per 100 residues) may be used for data interpretation without knowing the identity or molecular weight of the sample. Peptide or Protein Identification Amino acid compositional data can be used to search computer databases using PROPSEARCH and/or ExPASy to identify unknown samples (see Support Protocol 19). BASIC PROTOCOL
Amino Acid Analysis
PHENYLTHIOCARBAMYL AMINO ACID ANALYSIS Phenylthiocarbamyl amino acid analysis (PTC-AAA) is an established method for picomole-level quantitative analysis based on the reaction of phenylisothiocyanate (PITC) with amino groups to form phenylthiocarbamyl amino acid derivatives, which are then separated by reversed-phase HPLC and identified and quantified by ultraviolet absorbance. Since its introduction in the 1980s (Koop et al., 1982; Heinrikson and Meredith, 1984; Tarr, 1986), PTC-AAA has become the most popular precolumn-derivatization approach to AAA and may be used today with greater frequency than the classic ninhydrin postcolumn-derivatization method developed in the 1950s (Moore and Stein, 1949; Moore et al., 1958). During the 1990s, the majority of PTC-AAA has been performed with instrumentation from two vendors. Waters Associates first commercialized the PTC method under the trade name PicoTAG amino acid analysis (Bidlingmeyer et al., 1984, 1986), and PE Applied Biosystems Division introduced instrumentation in 1986 that performs automatic PTC derivatization with on-line HPLC analysis. The PE Applied Biosystems instrument was subsequently modified to also perform automatic on-line HCl hydrolysis. Both companies have provided reliable instrumentation and valuable technical and service support for PTC-AAA. However, most other companies marketing HPLC systems also provide instructions for how to separate PTC amino acids using their respective instrumentation. Protocols in this unit can be used with essentially any HPLC system. Materials Peptide or protein samples Volatile solvents, e.g., HPLC-grade water (Burdick and Jackson); aqueous 0.1% trifluoracetic acid/acetonitrite; 30% to 50% methanol, ethanol, or acetonitrite; 0.1% to 1% N-ethylmorpholine acetone, pH 8.5; or 0.1% HCl. Internal standard, e.g., α-aminobutyric acid (optional) 2:1 (v/v) methanol (Burdick and Jackson)/diisopropylethylamine (DIEA; PE Applied Biosystems)
11.9.4 Supplement 7
Current Protocols in Protein Science
7:2:2 (v/v/v) methanol/DIEA/5% (w/v) phenylisothiocyanate (PITC; Pierce or PE Applied Biosystems) in heptane (Burdick and Jackson) EDTA-containing transfer buffer compatible with HPLC system, e.g., 29 mM sodium acetate buffer pH 5.2 (Mallinkrodt), containing 0.25 mg/ml K3EDTA (Sigma) Amino acid Standard H (Pierce) Peptide or protein standard, e.g., insulin subunits, myoglobin, and lysozyme (Sigma) or 7% (w/v) bovine serum albumin (National Institute for Standards and Technology, formerly National Bureau of Standards) Vacuum system consisting of: Vacuum pump (e.g., Leybold-Heraeus Trivac D2A) Vacuum gauge (e.g., Fredericks Televac) Vacuum hose and connectors Cold trap and Dewar flask Glass three-way stopcock Polytetrafluoroethylene (PTFE) 6-mm bore Rotofol valve (Corning) or two-way stopcock Argon (or nitrogen) gas supply consisting of: Tank of prepurified gas Regulator for gas tank Silicone tubing Gradient HPLC system consisting of: Two HPLC pumps Mixing chamber Controller UV detector Manual injection valve Automatic sample injector (recommended but optional) Recording integrator (e.g., Shimadzu C-R6A Chromatopac) Mininert slide valves (Pierce) 40-ml screw-cap vials (Pierce) with Teflon/silicone discs 6 × 50–mm glass Pyrex tubes (Corning) Spreadsheet software (e.g., Microsoft Excel) Additional reagents and equipment for preparing samples (see Support Protocols for Sample Preparation), hydrolyzing samples (see Support Protocols for Hydrolysis), performing quantitative RP-HPLC (UNITS 8.2 & 11.6), plotting results to determine linear response ranges, and calculating standard deviations to determine reproducibility Set up the PTC-AAA workstation 1. Assemble key equipment and accessories close at hand in a dedicated PTC-AAA work area near a fume hood. A PTC-AAA workstation consists of a vacuum system, a fume hood, an argon (or nitrogen) flush system, an oven, freezer, HPLC system, automatic pipettor, 6 × 50–mm glass hydrolysis/derivatization tubes, 40-ml screwcap vials, and Mininert slide valves. Waters markets a PicoTAG Work Station (Bidlingmeyer et al., 1986) designed after the manual sequencing and PTC-AAA station developed by Tarr (1986). Original workstation descriptions with schematic diagrams appear in the above articles, and photographs of workstations are also available (Kuhn and Crabb, 1986). The work area can be arranged according to individual preference. The vacuum/argon flush system, 6 × 50–mm tubes, and 40-ml reaction vials with Mininert slide valves particularly facilitate manual acid hydrolysis and PTC derivatization protocols. Chemical Analysis
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2. Modify a few Mininert slide valves. First, remove the Teflon slider and silicone plug, enlarge the bore to ∼1/8 in. (∼3 mm) with an electric drill, and connect the modified valve to the vacuum system via vacuum hose and a PTFE Rotofol valve or two-way stopcock. This allows vacuum drying of PTC-derivatized samples in a 40-ml screw-cap vial.
3. On other Mininert valves, replace the silicone plug with the unused Teflon slider from the first modified valves, cut off the excess Teflon plug with a razor blade, push the valve open (green position in), and enlarge the bore slightly to ∼3/64 in. (∼1.2 mm) with an electric drill (Bidlingmeyer et al., 1986; Kuhn and Crabb, 1986; Tarr, 1986). This allows effective argon flushing and evacuation of samples in a 40-ml screw-cap vial in preparation for acid hydrolysis. CAUTION: With extended use, the Teflon liner of the valve compresses and may no longer seal properly. Discard and replace worn valves as necessary—e.g., when the indentation in the valve becomes significant or the vacuum no longer holds.
4. Modify several 40-ml screw-cap vials. Have a glass blower form a bulge around the circumference of the vial ∼50 mm from the bottom. This prevents HCl condensate from running down the inside wall of the vial into the 6 × 50–mm sample tubes during acid hydrolysis.
5. Connect the vacuum and argon flush systems via a three-way stopcock (Tarr, 1986) to provide alternating gas flush and vacuum to samples in preparation for hydrolysis. Attach a short piece of vacuum hose to the third port on the three-way stopcock and notch the other end to fit snugly around the slider knobs on the Mininert valve. Prepare the sample 6. Prepare the peptide or protein sample (see Support Protocols for Sample Preparation) and dissolve in a volatile solvent. Whenever possible, prepare enough sample for duplicate analyses per hydrolysis (∼0.5 to 1.0 µg sample is needed per analysis; additional sample material is necessary to quantify unusual amino acids). Best results are obtained with homogeneous samples, free of salts and detergents. Common volatile solvents for readily soluble samples are HPLC-grade water; aqueous 0.1% trifluoroacetic acid/acetonitrile; 30% to 50% methanol, ethanol, or acetonitrile; 0.1% to 1% N-ethylmorpholine acetate, pH 8.5; and 0.1% HCl. Less soluble samples may require 75% to 100% formic, acetic, or trifluoroacetic acids. Samples dried from 1 mM MOPS, pH 7, or from 25 mM Tris⋅HCl, pH 7.5/1 mM DTT/1 mM EDTA may yield satisfactory data, if the total salt is <50 µg. A defined amount of an internal standard (e.g., α-aminobutyric acid) may be added to the sample prior to hydrolysis and/or derivatization; indeed, an internal standard is essential for C-terminal sequencing with carboxypeptidases (UNIT 11.8). The internal standard should be stable to hydrolysis, derivatization, and analysis conditions and should be clearly resolved on the HPLC system.
Hydrolyze the sample 7. Hydrolyze the sample (see Support Protocols for Hydrolysis). Hydrolysis, a key step in determining success in PTC-AAA, is usually performed by heating samples in scrupulously clean hydrolysis tubes (see Support Protocol 6), in constant-boiling HCl (see Support Protocols 7 to 9). An alternate method is microwave hydrolysis (see Support Protocol 10). Automatic on-line vapor-phase HCl hydrolysis can also be done with the PE Applied Biosystems model 420H hydrolyzer/derivatizer. Amino Acid Analysis
Shorter hydrolysis times may be used to optimize recovery of Ser and Thr, which are partially destroyed by HCl hydrolysis, whereas longer times may help with Ile and Val,
11.9.6 Supplement 7
Current Protocols in Protein Science
which are slow to cleave. Other modifications of this basic procedure may be necessary to resolve amino acids that are adversely affected by hydrolysis (see Support Protocols for Amino Acid Analysis of Difficult or Unusual Residues).
Derivatize the sample The following manual protocol is adapted from the Applied Biosystems PTC amino acid analysis instrument manual (version 3.22, May 1989). 8. Dry <20 µg hydrolyzed sample in a clean 6 × 50–mm tube. 9. Add 20 µl of 2:1 (v/v) methanol/DIEA and redry. It is possible to substitute ethanol for the methanol and triethylamine (TEA) for the DIEA; reagent by-product peaks in the chromatogram, however, will differ slightly from those shown in Figure 11.9.1.
10. Add 15 µl of 7:2:2 (v/v/v) methanol/DIEA/5% PITC in heptane. 11. Mix samples and incubate 20 min at room temperature. 12. Dry sample in a Speedvac evaporator or 40-ml screw-cap vial. Store at −20°C under argon until ready to analyze. Use of instrumentation for automatic PTC derivatization with on-line HPLC analysis (PE Applied Biosystems) reduces variability caused by time-dependent degradation of PTC derivatives.
13. Dissolve the derivatized amino acids in an EDTA-containing transfer buffer compatible with the HPLC system. A typical transfer buffer is 29 mM sodium acetate, pH 5.2, containing K3 EDTA (0.25 mg/ml) which works well for the HPLC separation system shown in Figure 11.9.1. Sample amounts of 0.5 to 1.0 µg protein analysis provide useful results for most PTC-AAA HPLC systems. Dissolved samples should be analyzed within 12 to 15 hr. The derivatization reaction is relatively tolerant to variations in pH and molar ratios of reagents to sample, but PTC derivatives may slowly degrade in solution at room temperature.
Analyze the sample 14. Perform quantitative reversed-phase HPLC analysis of the PTC amino acids. An example of a typical PTC amino acid separation with UV detection at 254 nm is shown in Figure 11.9.1 along with a description of the chromatography conditions. UV-absorbing by-products from reactions between PITC and other derivatization reagents and sample components (e.g., aniline, phenylthiourea compounds) usually are not problematic provided they are well resolved from amino acid peaks. If increased amounts of by-products are observed, fresh derivatization reagents may be needed. Highly pure solvents and reagents are essential. Conditions for separating PTC amino acids by RP-HPLC are well established, and HPLC vendors can provide additional information. Aside from the actual column and HPLC system, important chromatography parameters are temperature, ionic strength and pH of the solvent, and the acetonitrile gradient used for elution. Increasing the temperature and acetonitrile concentration causes all PTC amino acids to elute faster. Asp and Glu are retained longer at low solvent pH and are eluted sooner at higher pH.
Calibrate the system 15. Dilute amino acid Standard H to an appropriate working concentration in water (e.g., 50 to 100 pmol/µl). Store 1 to 2 ml at −20°C. The amino acid Standard H from Pierce is convenient for calibration and contains the sixteen amino acids commonly found in protein hydrolysates (Asp, Glu, Ser, Gly, His, Arg,
Chemical Analysis
11.9.7 Current Protocols in Protein Science
Supplement 7
1 2 345 67 D
E
Aniline
8
RA
Y V 9 M
11 12 13
I L
F
K
DIPTU
P
ETPU
G S H T
10
IPTU
C PTU
0
5
10
15
20
Minutes
Figure 11.9.1 RP-HPLC of PTC amino acid standards. PTC-derivatized amino acid Standard H (150 pmol; Pierce) was chromatographed using the PE Applied Biosystems Model 130 HPLC and PE Applied Biosystems PTC C18 column (2.1 × 220–mm) with a flow rate of 300 µl/min, column temperature of 36°C, and the following gradient: time 0, 5% B; time 10 min, 32% B; time 20 min, 60% B; and time 25 min, 100% B. Solvent A was 50 mM sodium acetate, pH 5.4; solvent B was 31.5 mM sodium acetate containing 70% acetonitrile. The elution positions of sixteen amino acids are identified using the single-letter code (see Table A.1A.1) The elution positions of several other amino acids and PTC derivatization by-products like aniline, phenylthiourea (PTU), ethylphenylthiourea (EPTU), isopropylphenylthiourea (IPTU), and diisopropylphenylthiourea (DIPTU) are also indicated by arrows and numbers: 1, cysteic acid; 2, carboxymethyl cysteine; 3, hydroxyproline; 4, asparagine; 5, glutamine; 6, homoserine; 7, methionine sulfoxide; 8, α-aminobutyric acid; 9, norvaline; 10, pyridylethylcysteine; 11, norleucine; 12, ornithine; 13, tryptophan.
Thr, Ala, Pro, Tyr, Val, Met, Ile, Leu, Phe, and Lys) at the defined concentration of 2.5 µmol/ml plus cystine at 1.25 µmol/ml. Alternatively, quantitative standard solutions may be prepared by weighing individual dry amino acid standards (Pierce) and suspending in 0.1 N HCl to a defined concentration. (See Critical Parameters for further discussion of performance overview using amino acid standards.)
16. Derivatize 1 to 2 nmol (e.g., dry 10 to 20 µl of Standard H at 100 pmol/µl in a 6 × 50–mm tube and follow the PTC derivatization protocol, steps 9 to 13). 17. Perform six to nine PTC amino acid analyses (step 14) with a defined amount of Standard H (e.g., 100 to 300 pmol). It is essential that the amount used for calibration fall within the linear response range of the analysis system yet be above background contamination levels. Linear responses are typically obtainable in the range 1 to 5000 pmol; however, background contamination varies from laboratory to laboratory. Background can be problematic at the 50-pmol level and usually will be problematic below 20 pmol.
18. Determine the average peak area (or peak height) for each amino acid from the multiple analyses using an integrator. Amino Acid Analysis
This calculation is greatly facilitated by a computer or inexpensive integrator (such as the Shimadzu C-R6A Chromatopac, which has the ability to store and reprocess a number of analyses).
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Current Protocols in Protein Science
19. Calculate a response factor for each amino acid by dividing the average peak area (or peak height) by the picomoles of amino acid analyzed. response factor =
average peak height average peak area or pmol amino acid pmol amino acid
20. Create a calibration file in the integrator that lists each amino acid by name, retention time, and determined response factor. Set the integrator to divide amino acid peak areas (or peak heights) from unknown samples by the response factors to determine the picomoles of each amino acid in experimental samples. 21. Evaluate the quantitative accuracy of the system every day of use by running at least three standards; update the calibration file as needed. Recalibrate at least once a week. Determine the linear response range 22. Analyze in replicate five different amounts of Pierce Standard H covering the presumed linear response range (e.g., 1 to 5 nmol). To obtain accurate quantification, the detected amount of amino acids must fall within the linear response range of the analyzer. It is important to recognize data that exceed the useful range of measurement (see Anticipated Results).
23. Plot average peak area versus amount analyzed for each amino acid (e.g., see Fig. 11.9.10). 24. Determine the linear response range from the graph and strive to keep all residues in the experimental analyses below the upper limit. Background contamination and hydrolysis losses, not sensitivity, are generally the limiting factors at the lower end of the response range.
25. Hydrolyze and analyze in replicate different amounts of a peptide or protein standard of known concentration, calculate the amount of protein analyzed, and plot the results as amount hydrolyzed versus amount recovered. Determine reproducibility 26. Monitor the reproducibility of residue retention times and standard amino acid peak areas (or peak heights) in the analysis system (see Anticipated Results) by performing several analyses of a defined amount of Pierce Standard H (e.g., 100 to 300 pmol) as described for instrument calibration. 27. Compile retention time and peak area/height data for about six analyses. 28. Calculate the reproducibility (i.e., precision) of retention times and peak areas (or peak heights) as the relative standard deviation (RSD) or % standard deviation about the mean for each amino acid. Calculate average reproducibility by averaging the RSD values for all amino acids measured. Reproducibility (i.e., precision) not only is dependent on instrument performance but also is related to amount of sample analyzed, efficiency of PTC derivatization, background contamination levels, pipetting accuracy, and efficiency of hydrolysis for peptides and proteins. In general, variability increases with decreased sample amount and increased background. Reproducible performance must be demonstrated to justify use of the analysis system!
29. Perform several analyses of a defined amount of a standard peptide or protein hydrolysate and calculate the RSD for average peak area (or peak height) for each amino acid. 30. Repeat the analyses and statistical calculations every week over several weeks and months to demonstrate the long-term reproducibility of the system.
Chemical Analysis
11.9.9 Current Protocols in Protein Science
Supplement 7
Acceptable PTC-AAA reproducibility is reflected by average RSD values of ∼0.3% for retention times and 6 to 7% for peak areas with either Standard H (150-pmol level) or standard proteins (300-ng level). Lower RSD values are attainable; higher RSD values should be avoided.
Verify system performance with a standard peptide or protein 31. Every day when in use, verify the accuracy of the AAA system: hydrolyze, derivatize, and analyze in duplicate a peptide or protein standard. Any peptide or protein of known composition can be used following quantitation—e.g., insulin, myoglobin, or lysozyme (Sigma). A convenient quantitative standard for this purpose is a 7% solution of bovine serum albumin available from NIST (formerly the National Bureau of Standards).
Calculate amino acid composition in mole % 32. Using data from chromatographic peaks that are accurately identified and integrated and a spreadsheet (e.g., Fig. 11.9.2), enter pmol analyzed for each amino acid (Column B). 33. Sum the total pmol of amino acids in the analysis (B27). 34. Divide pmol of each residue by the sum total (Column B) and multiply by 100. Tabulate results as mole % (Column C). A mole % calculation example using PTC-AAA data from the analysis of bovine serum albumin is shown in Figure 11.9.3. This information can be used with PROPSEARCH and
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 Amino Acid Analysis
A DATE RUN TYPE IN ID# NAME AND METHOD TOTAL VOL IN UL VOL HYD ul RATIO APPLIED
B
C
3/31/96
AMINO ACID ASP (D) GLU (E) SER (S) GLY (G) (H) HIS ARG (R) THR (T) ALA (A) PRO (P) TYR (Y) VAL (V) MET (M) (I) ILE LEU (L) PHE (F) LYS (K)
z
TOTAL PMOL ANALYZED =
=SUM(B10:B25)
COMPOSITION MOLE % =(B10/B27) =(B11/B27) =(B12/B27) =(B13/B27) =(B14/B27) =(B15/B27) =(B16/B27) =(B17/B27) =(B18/B27) =(B19/B27) =(B20/B27) =(B21/B27) =(B22/B27) =(B23/B27) =(B24/B27) =(B25/B27)
=SUM(C10:C25)
Figure 11.9.2 Spreadsheet for calculating mole %. Microsoft Excel data entry commands are shown.
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ExPASy to search databases to provide information about the identity of the peptide or protein (see Support Protocol 19).
Calculate amino acid composition in residues per molecule for unknown sample 35a. For a sample for which the approximate molecular weight is known (e.g., from SDS-PAGE; UNIT 10.1), enter the approximate molecular weight (B28) into a spreadsheet, e.g., Figure 11.9.4. 36a. Enter the µl volume hydrolyzed (B5) and the fraction (ratio) of the amount applied to the analyzer that was actually analyzed (B6). The amount analyzed is a defined fraction of the amount applied and depends upon how the instrument is set up.
37a. Enter pmol analyzed for each amino acid (B9 to B24). 38a. Divide the molecular weight by 112 (assumed average residue weight) to estimate the total residues per molecule (B30). 39a. Sum total pmol analyzed (column B). Divide total pmol analyzed (B26) by total residues per molecule (B30) to estimate pmol protein analyzed (D29). 40a. Divide pmol each amino acid (column B) by pmol protein (D29) to estimate residues per molecule (column C).
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27
A DATE RUN TYPE IN ID# NAME AND METHOD TOTAL VOL IN UL VOL HYD ul RATIO APPLIED
B
C
3/31/96 BSA 10 1.0
AMINO ACID ASP (D) GLU (E) SER (S) GLY (G) HIS (H) ARG (R) THR (T) ALA (A) PRO (P) TYR (Y) VAL (V) MET (M) ILE (I) LEU (L) PHE (F) LYS (K)
pmol analyzed 1606.1 2251.26 760.99 545.79 469.49 637.16 887.22 1316.07 832.03 621.56 967.31 112.87 298.79 1844.82 799.26 1102.34
COMPOSITION MOLE % 10.67% 14.96% 5.06% 3.63% 3.12% 4.23% 5.89% 8.74% 5.53% 4.13% 6.43% 0.75% 1.98% 12.26% 5.31% 7.32%
TOTAL PMOL ANALYZED =
15053.06
100.00%
Figure 11.9.3 Calculation example—mole %. PTC-AAA data from the analysis of bovine serum albumin is shown. Chemical Analysis
11.9.11 Current Protocols in Protein Science
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Amino Acid Analysis
11.9.12
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Current Protocols in Protein Science
B
=SUM(B9:B24)
TOTAL PMOL ANALYZED =
=SUM(C9:C24)
UG PROTEIN ANALYZED
PMOL PROTEIN ANALYZED
CALCULATED MW PROTEIN
EXPERIMENTAL RESIDUES =B9/D29 =B10/D29 =B11/D29 =B12/D29 =B13/D29 =B14/D29 =B15/D29 =B16/D29 =B17/D29 =B18/D29 =B19/D29 =B20/D29 =B21/D29 =B22/D29 =B23/D29 =B24/D29
C
=(D29*D26)/1000000
=B26/B30
=SUM(F9:F24)+18
COMPOSITION MOLE % =(B9/B26) =(B10/B26) =(B11/B26) =(B12/B26) =(B13/B26) =(B14/B26) =(B15/B26) =(B16/B26) =(B17/B26) =(B18/B26) =(B19/B26) =(B20/B26) =(B21/B26) =(B22/B26) =(B23/B26) =(B24/B26)
D
ug/ul
=D32/B6/B5
=D29/B6/B5
=D26/B30
AV. RES. WT.
PMOL/ul
SUM RESIDUE WTS. =C9*E9 =C10*E10 =C11*E11 =C12*E12 =C13*E13 =C14*E14 =C15*E15 =C16*E16 =C17*E17 =C18*E18 =C19*E19 =C20*E20 =C21*E21 =C22*E22 =C23*E23 =C24*E24
F
RESIDUE WT. 115.09 129.12 87.08 57.05 137.14 156.19 101.10 71.08 97.12 163.18 99.07 131.19 113.16 113.16 147.18 128.17
E
Spreadsheet for calculating residues per molecule for unknown samples. Microsoft Excel data entry commands are shown.
EXP TOTAL RESIDUES
=B28/112 ESTIMATED TOTAL RESIDUES (ASSUMING AVE RESIDUE WT=112)
estimated molecular weight=
pmol analyzed
Unknown sample
3/31/96
AMINO ACID ASP (D) GLU (E) SER (S) GLY (G) HIS (H) ARG (R) THR (T) ALA (A) PRO (P) TYR (Y) VAL (V) MET (M) ILE (I) LEU (L) PHE (F) LYS (K)
AMOUNT HYD RATIO APPLIED
A DATE RUN TYPE IN ID# NAME AND METHOD
Figure 11.9.4
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33
41a. To calculate the original sample concentration (F29) in pmol/µl, divide the pmol protein analyzed (D29) by the µl volume hydrolyzed (B5) and the ratio of the applied amount that was actually analyzed (B6). To simplify the ratio applied, the integrator can be set to normalize the pmol values to 100% analyzed.
42a. Calculate mole % in column D by dividing by total pmol analyzed (B26). 43a. Enter the average residue weight for each amino acid in column E. Calculate the sum of the residue weights in column F by multiplying residue values (column C) by residue weights (column E). 44a. Calculate the molecular weight of the sample (D26) by summing F9 through F24 and adding 18 for water. 45a. Calculate µg protein analyzed (D32) by multiplying pmol analyzed (D29) by the molecular weight (D26) and by 10−6. 46a. Calculate the original sample concentration (F32) in µg/µl: divide the µg amount analyzed (D32) by the µl volume hydrolyzed (B5) and the ratio of the applied amount that was actually analyzed (B6). A calculation of the residues per molecule for PTC-AAA of bovine serum albumin is shown in Figure 11.9.5.
Calculate amino acid composition residues per molecule for known sample 35b. For samples of known mass and composition, use the spreadsheet in Figure 11.9.6 to calculate residues per molecule. Enter the known molecular weight (B6), the µl volume hydrolyzed (B4), and the fraction (ratio) of the amount applied to the analyzer that was actually analyzed (B5). This method allows sample quantitation from partial compositional data and provides a rapid evaluation of the accuracy of the analysis. The calculation methods outlined in Figures 11.9.2 to 11.9.5 can also be applied to samples of known mass and composition. The amount analyzed is a defined fraction of the amount applied and depends upon how the instrument is set up.
36b. Enter pmol analyzed for each amino acid (column C) and the known residues per molecule (column B). 37b. Divide pmol of each residue (column C) by the known residue value (column B) to determine pmol protein based on each amino acid (column D). 38b. Estimate the amount analyzed (B27) by averaging all the values in column D. 39b. Discard pmol protein values with unacceptable deviation from the mean (e.g., greater than ±15%). This is an arbitrary window that can be adjusted (for example, to ±30%) to optimize the number of relevant residues in the recalculated mean (F26).
40b. Calculate the average pmol protein analyzed by averaging the remaining relevant values (F26). 41b. Calculate the experimental composition in residues per molecule (column G) by dividing the amount of each residue by the calculated pmol protein analyzed (F26). 42b. Round off experimental residue values (column G) to integer values (column H). This is the determined amino acid composition.
Chemical Analysis
11.9.13 Current Protocols in Protein Science
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Amino Acid Analysis
11.9.14
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Current Protocols in Protein Science
UG PROTEIN ANALYZED 589.29
EXP TOTAL RESIDUES
1.71
25.54
67138
COMPOSITION MOLE % 10.67% 14.96% 5.06% 3.63% 3.12% 4.23% 5.89% 8.74% 5.53% 4.13% 6.43% 0.75% 1.98% 12.26% 5.31% 7.32%
D
ug/ul
0.171
2.554
113.92 AV. RES. WT.
PMOL/ul
SUM RESIDUE WTS. 7236 11379 2594 1219 2521 3896 3512 3662 3163 3971 3754 580 1324 8172 4605 5532
F
RESIDUE WT. 115.09 129.12 87.08 57.05 137.14 156.19 101.11 71.08 97.12 163.18 99.13 131.19 113.16 113.16 147.18 128.17
E
Calculation example—residues per molecule for unknown samples. PTC-AAA data from the analysis of bovine serum albumin is shown.
PMOL PROTEIN ANALYZED 589
66000
CALCULATED MW PROTEIN
EXPERIMENTAL RESIDUES 62.9 88.1 29.8 21.4 18.4 24.9 34.7 51.5 32.6 24.3 37.9 4.4 11.7 72.2 31.3 43.2
C
ESTIMATED TOTAL RESIDUES (ASSUMING AVE RESIDUE WT=112)
estimated molecular weight =
15053.06
pmol analyzed 1606.1 2251.26 760.99 545.79 469.49 637.16 887.22 1316.07 832.03 621.56 967.31 112.87 298.79 1844.82 799.26 1102.34
AMINO ACID (D) ASP (E) GLU (S) SER (G) GLY (H) HIS (R) ARG (T) THR (A) ALA (P) PRO (Y) TYR (V) VAL (M) MET (I) ILE (L) LEU (F) PHE (K) LYS TOTAL PMOL ANALYZED =
10 1
AMOUNT HYD RATIO APPLIED
B
Unknown sample
3/31/96
A DATE RUN TYPE IN ID # NAME AND METHOD
Figure 11.9.5
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33
Current Protocols in Protein Science B
=SUM(B8:B23) ANALYZED =AVERAGE(D8:D23) =B27*0.15 =B27-B28 =B27+B28
known Composition
KNOWN
3/31/96
pmol analyzed
C
TOTAL PMOL TOTAL
PMOL PROTEIN =IF(B8=0,FALSE,C8/B8) =IF(B9=0,FALSE,C9/B9) =IF(B10=0,FALSE,C10/B10) =IF(B11=0,FALSE,C11/B11) =IF(B12=0,FALSE,C12/B12) =IF(B13=0,FALSE,C13/B13) =IF(B14=0,FALSE,C14/B14) =IF(B15=0,FALSE,C15/B15) =IF(B16=0,FALSE,C16/B16) =IF(B17=0,FALSE,C17/B17) =IF(B18=0,FALSE,C18/B18) =IF(B19=0,FALSE,C19/B19) =IF(B20=0,FALSE,C20/B20) =IF(B21=0,FALSE,C21/B21) =IF(B22=0,FALSE,C22/B22) =IF(B23=0,FALSE,C23/B23)
D
PMOL PROTEIN =AVERAGE(F8:F23) =SUM(C8:C23) =F26/B5 =F29/B4
AMINO ACID PMOL HYDROL CONC pmol/µl
UNDER 15% =IF(E8>B29,E8,FALSE) =IF(E9>B29,E9,FALSE) =IF(E10>B29,E10,FALSE) =IF(E11>B29,E11,FALSE) =IF(E12>B29,E12,FALSE) =IF(E13>B29,E13,FALSE) =IF(E14>B29,E14,FALSE) =IF(E15>B29,E15,FALSE) =IF(E16>B29,E16,FALSE) =IF(E17>B29,E17,FALSE) =IF(E18>B29,E18,FALSE) =IF(E19>B29,E19,FALSE) =IF(E20>B29,E20,FALSE) =IF(E21>B29,E21,FALSE) =IF(E22>B29,E22,FALSE) =IF(E23>B29,E23,FALSE)
F
NEW ESTIMATE WITHIN 15%
OVER 15% =IF(D8
E
Total ugrams CONC µg/µl
EXP COMP =C8/(F26) =C9/(F26) =C10/(F26) =C11/(F26) =C12/(F26) =C13/(F26) =C14/(F26) =C15/(F26) =C16/(F26) =C17/(F26) =C18/(F26) =C19/(F26) =C20/(F26) =C21/(F26) =C22/(F26) =C23/(F26)
G
=(F29*B6)/1000000 =H29/B4
AVERAGE % ERROR
INT COMP =ROUND(G8,0) =ROUND(G9,0) =ROUND(G10,0) =ROUND(G11,0) =ROUND(G12,0) =ROUND(G13,0) =ROUND(G14,0) =ROUND(G15,0) =ROUND(G16,0) =ROUND(G17,0) =ROUND(G18,0) =ROUND(G19,0) =ROUND(G20,0) =ROUND(G21,0) =ROUND(G22,0) =ROUND(G23,0)
H
Spreadsheet for calculating residues per molecule for known samples. Microsoft Excel data entry commands are shown.
TOTAL #AA ESTIMATED AMOUNT PMOL PROTEIN 15% OF PROTEIN -15% OF PROTEIN +15% OF PROTEIN
A DATE RUN TYPE IN NAME/ID # VOL HYDROLYZED RATIO APPLIED Molecular weight AMINO ACID (D) ASP (E) GLU (S) SER (G) GLY (H) HIS ARG (R) (T) THR (A) ALA PRO (P) (Y) TYR (V) VAL (M) MET (I) ILE (L) LEU (F) PHE (K) LYS
Figure 11.9.6
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30
Chemical Analysis
11.9.15
Supplement 7
=AVERAGE(I8:I23)
% INT ERR =IF(B8=0,FALSE,(ABS(B8-H8)/B8)*100) =IF(B9=0,FALSE,(ABS(B9-H9)/B9)*100) =IF(B10=0,FALSE,(ABS(B10-H10)/B10)*100) =IF(B11=0,FALSE,(ABS(B11-H11)/B11)*100) =IF(B12=0,FALSE,(ABS(B12-H12)/B12)*100) =IF(B13=0,FALSE,(ABS(B13-H13)/B13)*100) =IF(B14=0,FALSE,(ABS(B14-H14)/B14)*100) =IF(B15=0,FALSE,(ABS(B15-H15)/B15)*100) =IF(B16=0,FALSE,(ABS(B16-H16)/B16)*100) =IF(B17=0,FALSE,(ABS(B17-H17)/B17)*100) =IF(B18=0,FALSE,(ABS(B18-H18)/B18)*100) =IF(B19=0,FALSE,(ABS(B19-H19)/B19)*100) =IF(B20=0,FALSE,(ABS(B20-H20)/B20)*100) =IF(B21=0,FALSE,(ABS(B21-H21)/B21)*100) =IF(B22=0,FALSE,(ABS(B22-H22)/B22)*100) =IF(B23=0,FALSE,(ABS(B23-H23)/B23)*100)
I
Supplement 7
B
KNOWN (NBS-BSA) 10 1.00 66328 known Composition 54 79 28 16 17 24 34 46 28 19 36 4 14 61 27 59
3/31/96
pmol analyzed 1606.1 2251.26 760.99 545.79 469.49 637.16 887.22 1316.07 832.03 621.56 967.31 112.87 298.79 1844.82 799.26 1102.34
C
TOTAL PMOL TOTAL
PMOL PROTEIN 29.74 28.50 27.18 34.11 27.62 26.55 26.09 28.61 29.72 32.71 26.87 28.22 21.34 30.24 29.60 18.68
D
PMOL PROTEIN 28.25 15053.06 28.25 2.825 AMINO ACID PMOL HYDROL µl CONC pmol/µ
UNDER 15% 29.74 28.50 27.18 FALSE 27.62 26.55 26.09 28.61 29.72 FALSE 26.87 28.22 FALSE 30.24 29.60 FALSE
F
NEW ESTIMATE WITHIN 15%
OVER 15% 29.74 28.50 27.18 FALSE 27.62 26.55 26.09 28.61 29.72 FALSE 26.87 28.22 21.34 30.24 29.60 18.68
E
Total ugrams CONC µg/µl
EXP COMP 56.9 79.7 26.9 19.3 16.6 22.6 31.4 46.6 29.5 22.0 34.2 4.0 10.6 65.3 28.3 39.0
G
H
1.87 0.187
AVERAGE % ERROR
INT COMP 57 80 27 19 17 23 31 47 30 22 34 4 11 65 28 39
Calculation example—residues per molecule for known samples. PTC-AAA data from the analysis of bovine serum albumin is shown.
TOTAL #AA 546 ESTIMATED AMOUNT ANALYZED PMOL PROTEIN 27.86 15% OF PROTEIN 4.18 -15% OF PROTEIN 23.68 +15% OF PROTEIN 32.04
A DATE RUN TYPE IN NAME/ID # VOL HYDROLYZED RATIO APPLIED Molecular weight AMINO ACID ASP (D) GLU (E) SER (S) GLY (G) HIS (H) ARG (R) THR (T) ALA (A) PRO (P) TYR (Y) VAL (V) MET (M) ILE (I) LEU (L) PHE (F) LYS (K)
Figure 11.9.7
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30
Amino Acid Analysis
11.9.16
Current Protocols in Protein Science
8.65
% INT ERR 5.56 1.27 3.57 18.75 0.00 4.17 8.82 2.17 7.14 15.79 5.56 0.00 21.43 6.56 3.70 33.90
I
43b. Calculate % error for each residue (column I) based on integer values (column H). 44b. Calculate the original sample concentration in pmol/µl (F30) by dividing the pmol protein analyzed (F26) by the ratio applied (B5) and the µl volume hydrolyzed (B4). 45b. Calculate total µg hydrolyzed (H29) by multiplying [pmol protein analyzed (F26)] × [ratio applied (B5)] × [molecular weight (D26)] × [10−6]. 46b. Calculate the original sample concentration in µg/µl (H30): divide total µg protein hydrolyzed (H29) by the µl volume hydrolyzed (B4). A calculation of residues per molecule for PTC-AAA of bovine serum albumin is shown in Figure 11.9.7.
Determine compositional accuracy 47. Evaluate compositional accuracy by calculating average compositional % error relative to the known composition of the sample (i.e., known from the amino acid sequence). This value usually excludes Trp and Cys but reflects all other residue values unadjusted for partial destruction, incomplete hydrolysis, or contamination.
48. Calculate % error per amino acid after rounding the experimentally observed number of residues to the nearest integer (see Figs. 11.9.6 and 11.9.7). % error = 100 ×
known number of residues − experimental number of residues known number of residues
average % compositional error =
Σ % error for n amino acids n
Usually, n = 16.
49. Check compositional and quantitative accuracy daily. Strive to maintain compositional and quantitative accuracy within 10% of the known values. AAA requires system performance verification with a standard peptide or protein of defined concentration and composition.
SUPPORT PROTOCOLS FOR SAMPLE PREPARATION The purpose of sample preparation is to produce homogeneous, salt- and detergent-free samples. Samples for amino acid analysis (AAA) may be desalted by one of several methods such as microdialysis (see Support Protocol 3), organic solvent precipitation (see Support Protocol 4 or 5), RP-HPLC (see Support Protocol 1), and centrifugation using microseparation concentrators (Amicon; see Support Protocol 2). Recovery from these procedures can be variable and sample dependent and can complicate the process of determining the concentration of stock preparations. Remember to check for and record sample volume changes that may occur during desalting and concentration steps. Sample Preparation by RP-HPLC Additional Materials (also see Basic Protocol) Solvent, e.g., trifluoroacetic acid (Sequenal-grade TFA, Pierce)/acetonitrile (UV HPLC-grade, Burdick and Jackson) RP-HPLC column, minicolumn, or disposable cartridge support (e.g., C4 or C18 support with 300-Å pore size, Vydac)
SUPPORT PROTOCOL 1
Chemical Analysis
11.9.17 Current Protocols in Protein Science
Supplement 7
Desalt samples by RP-HPLC (see UNIT 11.6) using a column and solvent appropriate for peptides and proteins. Classic aqueous trifluoroacetic acid (TFA)/acetonitrile solvents, are excellent for RP-HPLC desalting purposes. A variety of standard HPLC columns, minicolumns, and disposable cartridge style supports may be useful. SUPPORT PROTOCOL 2
Sample Preparation by Microseparation Concentrators
SUPPORT PROTOCOL 3
Sample Preparation by Microdialysis
CentriPrep, Centricon, and Microcon concentrators (Amicon) are particularly useful for simultaneously concentrating samples and exchanging solvents, and they are available in different volumes and exclusion limits (specifications and detailed instructions for use are supplied by the vendor). Apply the sample to the tube, centrifuge until the volume is appropriately reduced, add volatile solvent (see Basic Protocol, annotation to step 6 for examples), and repeat centrifugation. Repeat the exchange two to three times, depending on the original salt concentration.
Microdialysis can be used to prepare 100- to 200-µl samples in a method adapted from Orr et al. (1995). Additional Materials (also see Basic Protocol) Solvent (e.g., see Basic Protocol, step 6 annotation) 1.5-ml clear microcentrifuge tube Spectra/Por dialysis membrane (Spectrum Medical), rehydrated 1. Cut the cap off a microcentrifuge tube (∼1.5 ml) with scissors and then cut the tube in two. 2. Place the cap upside down on the bench and pipet the peptide or protein into the “cup” of the cap, which typically holds ∼200 µl. If the sample contains a high concentration of salt (e.g., 2 to 8 M urea or 6 M guanidine), apply ≤100 µl in order to leave room for volume expansion during dialysis.
3. Cut open a small piece of rehydrated Spectra/Por dialysis membrane of appropriate exclusion limit. Cover the cap and sample with a single layer of membrane and secure with the top piece of the cut tube. 4. Float the cap in a beaker of solvent (200 to 500 ml) and cover the beaker; dialyze 30 min with vigorous mixing. Change solvent and repeat three times. 5. Remove cap and collect the sample by puncturing the membrane with a micropipet. SUPPORT PROTOCOL 4
Amino Acid Analysis
Sample Preparation by Acetone Precipitation for Proteins This procedure is suitable for recovering proteins from most aqueous solvents and from SDS-containing buffers. It is not recommended for proteins dissolved in urea or guanidine or for peptides. Practice the precipitation procedure with a readily available protein before attempting use with a precious sample. This procedure is useful for removing SDS from protein preparations after electrophoretic purification and electroelution from polyacrylamide gels; it can also be used for concentrating samples for Edman sequence analysis and/or proteolytic digestion. Additional Materials (also see Basic Protocol) Acetone (Burdick and Jackson) Ethyl acetate (Burdick and Jackson) N-ethylmorpholine (Sequenal grade, Pierce)
11.9.18 Supplement 7
Current Protocols in Protein Science
HCl (ULTREX II ultrapure reagent, J.T. Baker) Trifluoroacetic acid (Sequenal-grade TFA, Pierce) Centrifuge 1. Ensure that the sample volume is in the range of 20 µl to 2 ml. For <10 µg protein, the volume should be reduced to <500 µl. This can be accomplished by partial drying in a Speedvac evaporator or by several extractions with excess ethyl acetate.
2. Add 2 vol acetone and vortex. The precipitated protein will be visible, usually as a flocculent, white particulate material. A magnifying glass can help in visualizing the precipitate.
3. If no precipitate is apparent, add ∼0.5 vol ethyl acetate; vortex again. 4. If still no precipitate is apparent, change pH slightly by adding a few microliters of N-ethylmorpholine to raise the pH or HCl to lower the pH. Check pH before and after additions by pipetting ∼1 µl to an appropriate range of pH paper. The protein will precipitate at its isoelectric pH.
5. Centrifuge and carefully remove the supernatant. Save the supernatant if the location of the sample is questionable. 6. For amino acid or sequence analysis, desalt the sample (e.g., wash the pellet carefully with a small volume of 67% acetone, blow dry with a gentle stream of nitrogen, and suspend in 100% TFA for necessary transfers). For proteolytic digestion, the pellet usually need not to be washed.
Peptide Sample Preparation by Ether Precipitation This procedure is suitable for precipitating peptides from organic acids and is used routinely for concentrating synthetic peptides from the trifluoroacetic acid cleavage cocktail following 9-fluorenylmethyloxycarbonyl (Fmoc) solid-phase peptide synthesis. It can be used to desalt peptides in nonvolatile buffers but is not recommended for removing detergents like SDS. Take appropriate safety precautions when using this technique; ether is highly flammable, and a refrigerated centrifuge and fume hood are recommended. Practice the precipitation procedure with 5- to 25-µg amounts of a standard peptide before attempting use with a precious sample. Maintain cold solution temperatures for best results.
SUPPORT PROTOCOL 5
Additional Materials (also see Basic Protocol) 75% trifluoroacetic acid (Sequenal grade TFA, Pierce) Ether, ice-cold 6 × 50–mm tube Centrifuge, 4°C CAUTION: Ether is highly flammable; work with a fume hood. 1. Vacuum dry an aliquot of the peptide preparation in a 6 × 50–mm tube. Redissolve the peptide in 50 to 200 µl of 75% TFA and cool to 4°C on ice. Alternatively, the peptide preparation can be diluted with 100% TFA to a final concentration of 75% TFA and cooled to 4°C. Keep final sample volumes small (e.g., ~200 µl).
2. Add 3 vol ice-cold ether, mix, and incubate 15 to 30 min on ice or in a −20°C freezer. Flocculent peptide is usually visible above 20 µg but increasingly difficult to see below 20 µg.
Chemical Analysis
11.9.19 Current Protocols in Protein Science
Supplement 7
3. Centrifuge 5 min at 4°C. 4. Decant supernant immediately while cold; vacuum dry the pellet. For amounts below ∼5 µg, a pellet may not be visible. Hydrolyze and analyze anyway (see Basic Protocol, steps 7 to 14).
5. Repeat steps 1 to 4 as needed for very salty samples. Typically very little precipitate is visible in the tube following vacuum drying. Substantial precipitate suggests excess salt remains.
SUPPORT PROTOCOLS FOR HYDROLYSIS The hydrolysis process has a major impact on the quality of results from amino acid analysis (AAA). Liquid-phase (see Support Protocol 8) and vapor-phase (see Support Protocol 7) HCl hydrolysis procedures are both popular as standard, complete hydrolysis methods. Examples of each method are presented. However, vapor-phase methods receive more wide-spread use. Samples can also be hydrolyzed on PVDF membranes (see Support Protocol 9) or using a microwave hydrolysis system (see Support Protocol 10). Vapor-phase and liquid-phase hydrolysis require specially prepared tubes and vials (see Support Protocol 6). For more information regarding the influence of various hydrolysis conditions, see the AAA studies sponsored by the Association of Biomolecular Resource Facilities, particularly Tarr et al. (1991), Strydom et al. (1992), and Yücksel et al. (1995). SUPPORT PROTOCOL 6
Preparation of Hydrolysis Tubes and Vials 1. Use 6 × 50–mm hydrolysis tubes for either vapor- or liquid-phase hydrolyses (see Support Protocols 7 and 8). 2. Rinse the tubes at least five times with water followed by five times with methanol (Burdick and Jackson) and then pyrolyze by heating 5 hr at 500°C. Clean tubes batchwise. 3. Store clean tubes in a dust-free, closed container. Discard used tubes—do not reuse. 4. Use clean bulge-modified 40-ml screw-cap vials (see Basic Protocol, step 4) for either vapor- or liquid-phase hydrolyses (see Support Protocols 7 and 8). Clean the modified vials with soap, rinse exhaustively with water and then methanol, and dry. Reuse bulge vials but always clean first.
SUPPORT PROTOCOL 7
Vapor-Phase HCl Hydrolysis of Samples Additional Materials (also see Basic Protocol) 6 N HCl (Sequenal grade, Pierce) Phenol (Aldrich) Diamond pencil 1. Vacuum dry purified, salt-free peptide/protein samples in clean 6 × 50–mm tubes (∼0.5 to 5 µg/tube) using a Speedvac evaporator. Use a diamond pencil to etch identifying marks on each tube. 2. Place 6 × 50–mm sample tube in a bulge modified 40-ml screw-cap vial containing ∼300 µl of 6 N HCl plus a crystal of phenol (∼1 to 2 mg). Seal the vial with a modified Mininert slide valve (see Basic Protocol, step 3).
Amino Acid Analysis
Phenol serves as a scavenger to prevent halogenation of Tyr. The bulge modification of the screw-cap vial prevents HCl condensate from running down the inside wall of the vial into the 6 × 50–mm sample tube during hydrolysis.
11.9.20 Supplement 7
Current Protocols in Protein Science
3. Evacuate briefly (15 to 20 sec) then flush with argon or nitrogen 1 to 2 sec. Repeat this alternating process three times, closing the slide valve (red position in) on the fourth vacuum step. 4. Heat 1 hr at 150°C. Following hydrolysis, release the pressure within a fume hood by pointing the cap away from your face and pushing the slide valve open. Perform time course hydrolyses for 1, 2, and 4 hr if desired. 5. Transfer the sample tubes to a clean, unmodified 40-ml vial and vacuum dry. Store at −20°C under argon until ready to perform AAA. Liquid-Phase HCl Hydrolysis of Samples
SUPPORT PROTOCOL 8
Additional Materials (also see Basic Protocol) 6 N HCl (Sequenal grade, Pierce) containing 1% (v/v) phenol (Aldrich) Diamond pencil 1. Follow steps 1 and 2 for manual vapor-phase hydrolysis (see Support Protocol 7) but add 20 µl of 6 N HCl containing 1% phenol directly to each sample. Seal the vial with a modified Mininert slide valve (see Basic Protocol, step 3). 2. Evacuate 15 to 20 sec and flush 1 to 2 sec with argon or nitrogen. Repeat three times. Watch for bubbles during the vacuum phase; avoid bumping when switching to argon flush. Close the slide valve (red position in) on the last vacuum step. 3. Heat 20 to 24 hr at ∼110°C. Following hydrolysis, release the pressure within a fume hood by pointing the cap away from your face and pushing the slide valve open. Perform time course hydrolyses for 24, 48, and 72 hr if desired. 4. Transfer the sample tubes to a clean, unmodified 40-ml vial and vacuum dry. Store at −20°C under argon until ready to perform AAA. Hydrolysis of Samples on PVDF Membranes
SUPPORT PROTOCOL 9
Additional Materials (also see Basic Protocol) 5% acetic acid/50% methanol (Burdick and Jackson) 50% acetonitrile (UV HPLC grade; Burdick and Jackson) or 50% methanol (Burdick and Jackson) Polyvinylidene difluoride (PVDF) membrane (PE Applied Biosystems, Millipore) ProSpin cartridge membrane (PE Applied Biosystems; optional) PVC gloves, powder-free Additional reagents and equipment for vapor-phase hydrolysis (see Support Protocol 7) or liquid-phase hydrolysis (see Support Protocol 8) 1. Apply samples to PVDF membranes by electroblotting, by centrifuging onto a ProSpin cartridge membrane, by absorbing, or by spotting with a micropipettor. Be aware that variable amino acid analysis results are frequently obtained from samples spotted or absorbed on PVDF membrane, presumably due to difficult-to-extract background contamination within the membrane (Tarr et al., 1991; Mahrenholz et al., 1996).
2. Wear powder-free PVC gloves, maintain clean technique, and use clean glassware and reagents. Cut PVDF membrane to fit into a 6 × 50–mm hydrolysis tube. Prior to hydrolysis wash briefly in 5% acetic acid/50% methanol to remove any excess stain. Prepare blank control pieces of PVDF from membrane treated as for the experimental samples. Forceps are useful for handling the small pieces of membrane.
Chemical Analysis
11.9.21 Current Protocols in Protein Science
Supplement 7
3. Hydrolyze control and experimental PVDF samples using standard vapor- or liquidphase HCl hydrolysis methods (see Support Protocol 7 or 8). 4. Following hydrolysis, add 100 µl of 50% acetonitrile or 50% methanol to each tube, incubate 10 min, then transfer the solution to a second 6 × 50–mm tube. Repeat the extraction once and vacuum dry the combined extracts. If necessary, samples extracted from PVDF membranes can be stored at −20°C under argon until ready to perform AAA.
5. PTC derivatize and analyze as usual (see Basic Protocol). Results from control samples can be subtracted from experimental samples. However, because background contamination is extremely variable, subtracting background does not always improve the results. Higher quality results are more readily obtained with sample amounts >1 µg (Gharahdaghi et al., 1992). SUPPORT PROTOCOL 10
Microwave Hydrolysis of Samples Microwave hydrolysis can reduce the time required to perform HCl vapor-phase hydrolysis. The following procedure was adapted from Hsi and Yamane (1994) and is based on the use of a CEM MDS 2000 Microwave Sample Preparation System equipped with a Teflon vapor-phase hydrolysis vessel and special microvials. The instrumentation is expensive compared with kitchen microwaves but is specially designed for vapor-phase hydrolysis for amino acid analysis. For a description of this microwave instrument, see Engelhart (1990). Additional Materials (also see Basic Protocol) 6 N HCl Microwave Sample Preparation System (CEM) consisting of: Microwave oven Teflon vapor-phase hydrolysis vessel Microvials 1. Dry samples in microvials; place up to ten vials in the Teflon vapor-phase hydrolysis vessel. 2. Add 5 ml of 6 N HCl to the hydrolysis vessel, flush with nitrogen, and seal. 3. Microwave hydrolyze 20 min at 150°C. 4. Vacuum dry, PTC derivatize, and analyze (see Basic Protocol, steps 8 to 14). SUPPORT PROTOCOLS FOR AMINO ACID ANALYSIS OF DIFFICULT OR UNUSUAL RESIDUES The Basic Protocol provides for analysis of only sixteen of the common amino acids. Difficult and unusual residues—e.g., cysteine, tryptophan, and phosphoamino acids—are often adversely affected by standard hydrolysis, so special hydrolysis conditions are required. In addition, chromatographic conditions may need to be modified to optimize resolution, as in the case of hydoroxyproline which may coelute with other residues. The response factors for these residues must be carefully determined using the appropriate standards (see Support Protocol 11).
Amino Acid Analysis
Quantification of cysteine is difficult because hydrolysis in 6 N HCl destroys the residue and partially destroys the disulfide-linked dimeric form of the residue, cystine. This sulfur-containing residue must be modified for accurate measurement; two useful modification methods that receive widespread use are oxidative hydrolysis (see Support
11.9.22 Supplement 7
Current Protocols in Protein Science
Protocol 12) and pyridylethylation (see Support Protocol 13). A comparison of the performance of these and other methods for cysteine analysis can be found in Strydom et al. (1993). Tryprophan is destroyed by standard HCl hydrolysis and is even more challenging to quantify than cysteine. Two successful alternate hydrolysis strategies for quantifying tryptophan are vapor-phase HCl hydrolysis in the presence of dodecanethiol (see Support Protocol 13) and hydrolysis with 4 N methanesulfonic acid (see Support Protocol 15). A comparison of the performance of these and other methods for tryptophan analysis can be found in Strydom et al. (1993). Phosphoamino acid analysis is not yet routine for most laboratories that perform amino acid analysis (AAA), and no current AAA methodology allows quantification of phosphoamino acids in a single analysis in conjunction with common amino acids (Yücksel et al., 1994). Hydrolysis is the most critical part of the analytical process because of the acid lability of the phosphate moiety. Optimal hydrolysis conditions (see Support Protocols 16 and 17) must be determined for each sample peptide or protein. Hydrolysis conditions do not generally provided for quantitative cleavage of peptide bonds and often produce short peptides that may interfere with the measurement of other amino acids. The peptide or protein concentration must be verified in a separate analysis using standard conditions for complete hydrolysis. Hydroxyproline can be hydrolyzed and derivatized using standard conditions, but it may coelute with other residues such as glutamic acid, so chromatography conditions may need to be modified to optimize resolution of hydroxyproline in HPLC (see Support Protocol 18). Determining Response Factors for Difficult Residues Determine response factors for difficult residues such as cysteic acid (Cya) and tryptophan (Trp) from protein hydrolysates.
SUPPORT PROTOCOL 11
1. Perform the appropriate hydrolysis (see Basic Protocol, step 7) with more than one standard peptide or protein containing the residue of interest, e.g., oxidative hydrolysis for Cya (see Support Protocols 12 and 13) and hydrolysis with methanesulfonic acid (see Support Protocol 15) or dodecanethiol (see Support Protocol 14) for Trp . 2. Determine the amount of standard protein analyzed (see Figs. 11.9.6 and 11.9.7). 3. Divide the peak area (or peak height) of the amino acid of interest (e.g., Cya or Trp) by the amount of protein analyzed. 4. Finally, divide by the known number of residues of interest in the standard protein (e.g., for lysozyme, determine the response factor for Cya by dividing by 8). 5. Use the hydrolysate-derived response factor for converting Trp and Cya peak area (or peak height) to picomoles of amino acid analyzed. 6. Interpret data for the other residues using response factors derived from the routine quantitative amino acid standard (e.g., Standard H).
Chemical Analysis
11.9.23 Current Protocols in Protein Science
Supplement 14
SUPPORT PROTOCOL 12
Oxidative Hydrolysis for Quantification of Cysteine Oxidation of cysteine/cystine to cysteic acid (Cya) by hydrolysis in the presence of dimethylsulfoxide (DMSO) is a simple and convenient method of Cys quantification. The method was originally introduced by Spencer and Wold (1969) and more recently demonstrated to be effective in PTC-AAA (West and Crabb, 1992). Additional Materials (also see Basic Protocol) 2% (v/v) dimethylsulfoxide (DMSO; HPLC grade, Aldrich) 2% (w/v) diisopropylethylamine (DIEA; PE Applied Biosystems) Cysteic acid (Sigma) Standard protein containing cysteine, e.g., cytochrome c, or β-lactoglobulin, lysozyme, PTC-derivatized Additional reagents and equipment for vapor-phase hydrolysis (see Support Protocol 7) and determining response factor for cysteic acid (see Support Protocol 11) 1. Subject the peptide or protein to manual vapor-phase HCl hydrolysis (see Support Protocol 7) except add 2% (v/v) DMSO to the 6 N HCl and omit the phenol. 2. Follow the manual PTC derivatization protocol (see Basic Protocol, steps 8 to 11) and vacuum dry. 3. Dissolve the derivatized sample in EDTA-containing transfer buffer containing 2% (w/v) DIEA and incubate 10 min at room temperature prior to HPLC analysis. This step was developed as a vapor-phase “bubbling” step during automatic PTC derivatization and helps neutralize the negative charge on Cya through ion pairing, resulting in increased Cya retention time (Hsi and Yamane, 1994). Optimization of HPLC analysis of PTC-Cya (see Fig. 11.9.1) may also require lowering the solvent pH a few tenths of a unit to increase its retention.
4. Determine response factors for cysteic acid (see Support Protocol 11) by oxidative hydrolysis and PTC-AAA of standard cysteine-containing proteins such as cytochrome c (2 Cys/mol), β-lactoglobulin (5 Cys/mol), and lysozyme (8 Cys/mol). A cysteic acid standard is useful for optimizing chromatography, but response factors derived from oxidative hydrolysates of standard proteins are more reliable in the range of 1 µg protein hydrolyzed (West and Crabb, 1992).
5. Always perform concurrent oxidative hydrolysis and analysis of control and unknown samples to verify the efficacy and accuracy of the analytical process. Incomplete compositional analysis results from oxidative hydrolysis—His, Met, Tyr, and Trp are also modified. Quantitation of these residues should be obtained in separate analyses using other hydrolysis conditions. SUPPORT PROTOCOL 13
Amino Acid Analysis
Pyridylethylation for Quantification of Cysteine Alkylation of cysteine with 4-vinylpyridine produces pyridylethylcysteine, which is stable to acid hydrolysis (Friedman et al., 1970) and readily measured by PTC-AAA. Pyridylethylcysteine (Pec) is commercially available (Pierce) and can be used to determine an appropriate Pec response factor as described for other standard amino acids. The following protocol is adapted from Hawke and Yuan (1987). Additional Materials (also see Basic Protocol) 1:3 (v/v) 1 M Tris⋅HCl (pH 8.5) containing 4 mM EDTA (Sigma)/8 M guanidine⋅HCl or 1 M Tris⋅HCl/1 mM EDTA, pH 8.5, containing 8 M urea (ultrapure) 1 M Tris⋅HCl, pH 8.5, or 1 N HCl, for adjusting pH if necessary
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10% (w/v) 2-mercaptoethanol 4-vinylpyridine (Aldrich) Additional reagents and equipment for desalting sample (see Support Protocols for Sample Preparation) 1. Dry the sample and dissolve in <50 µl of either 1:3 (v/v) 1 M Tris⋅HCl, pH 8.5, containing 4 mM EDTA/8 M guanidine⋅HCl or 1 M Tris⋅HCl/1 mM EDTA, pH 8.5, containing 8 M urea. Check pH by spotting ∼1 µl on pH paper; adjust pH to ∼8.5 with 1 M Tris⋅HCl or 1 N HCl. Avoid guanidine⋅HCl if the sample contains SDS.
2. Add 2.5 µl freshly prepared 10% (w/v) 2-mercaptoethanol, flush with argon, seal, and incubate 2 hr at room temperature. 3. Add 2 µl 4-vinylpyridine neat, mix, and incubate 2 hr under argon. Always use fresh 4-vinylpyridine for pyridylethylation.
4. Desalt immediately by RP-HPLC, dialysis, solvent exchange (with a Microcon concentrator), or precipitation (see Support Protocols for Sample Preparation). Avoid SDS if possible; remove SDS by acetone precipitation.
5. Hydrolyze, PTC derivatize, and analyze for pyridylethylcysteine (see Fig. 11.9.1). Hydrolysis with HCl/Dodecanethiol for Quantification of Tryptophan The dodecanethiol/HCl hydrolysis method was the most successful method for Trp quantification in the comparative study of Strydom et al. (1993). The method involves automatic vapor-phase hydrolysis in the PE Applied Biosystems model 420H Amino Acid Analysis system.
SUPPORT PROTOCOL 14
Additional Materials (also see Basic Protocol) 40% (w/v) dodecanethiol (Fluka) in heptane (Burdick and Jackson) Standard protein containing tryptophan, e.g., cytochrome c, α-lactalbumin, or lysozyme Amino Acid Analysis system (PE Applied Biosystems model 420H) Additional reagents and equipment for determining response factors for difficult residues (see Support Protocol 11) 1. Apply 40 µl of 40% dodecanethiol (in heptane) to the sample frit of the PE Applied Biosystems model 420H Amino Acid Analysis system 10 to 15 min prior to sample application. Apply the sample and proceed with the normal autohydrolysis process. For further details, see Bozzini et al. (1991) and West and Crabb (1992).
2. Determine response factors for Trp by hydrolysis and PTC-AAA of standard Trpcontaining proteins such as cytochrome c (1 Trp/mol), α-lactalbumin (4 Trp/mol), and lysozyme (6 Trp/mol; see Support Protocol 11). A Trp standard is useful for optimizing chromatography, but response factors derived from hydrolysates of standard proteins are more reliable in the range of 1 µg protein hydrolyzed (West and Crabb, 1992).
3. Perform concurrent dodecanethiol/HCl hydrolysis and analysis of control and unknown samples to verify the efficacy and accuracy of the analytical process.
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SUPPORT PROTOCOL 15
Hydrolysis with 4 N Methanesulfonic Acid for Quantification of Tryptophan This alternative procedure, hydrolysis with 4 N methanesulfonic acid, is adapted from Hsi and Yamane (1994). Response factors for Trp are determined from 4 N methanesulfonic acid hydrolysates of standard proteins and concurrent hydrolysis and analysis of control and unknown samples. Additional Materials (also see Basic Protocol) 4 N methanesulfonic acid 4 N NaOH 1. Add 30 µl of 4 N methanesulfonic acid to the dried sample in a clean 6 × 50–mm tube. Seal sample tubes in bulge-modified 40-ml screw-cap vials and alternately flush with argon and evacuate as described for routine vapor- and liquid-phase HCl hydrolysis (see Support Protocol 7 or 8). Seal under argon rather than vacuum. Batchwise hydrolyses are recommended.
2. Heat 1 hr at 165°C, then cool to room temperature. Add 30 µl of 4 N NaOH to neutralize, then vacuum dry. 3. PTC derivatize and analyze (see Basic Protocol, steps 8 to 14). SUPPORT PROTOCOL 16
Quantification of pSer, pThr, and pTyr Chromatographic resolution and measurement of PTC-phosphoamino acids are straight forward and relatively simple (see Fig, 11.9.8). However, the acid lability of the phosphate moiety makes hydrolysis the most critical part of the analytical process. Phosphoserine (pSer), phosphothreonine (pThr), and phosphotyrosine (pTyr) are analyzed using a time course of liquid-phase HCl hydrolysis (see Support Protocol 8). Commercially prepared standards of the three phosphoamino acids are used to optimize chromatographic conditions and to determine response factors (see Support Protocol 11). Synthetic phosphopeptides can also be used to determine the response factors following limited HCl hydrolysis, but any such peptide must contain a well-defined amount of the phosphoamino acid of interest. Additional Materials (also see Basic Protocol and Support Protocol 8) Standard phosphoamino acids Additional reagents and equipment for liquid-phase hydrolysis (see Support Protocol 8) and determining response factors for difficult residues (see Support Protocol 11) 1. Perform a time course liquid-phase HCl hydrolysis at 110°C for 1, 2, 3, and 4 hr (see Support Protocol 8). Vacuum dry the hydrolysate. 2. PTC derivatize and analyze (see Basic Protocol, steps 8 to 14). Establish the chromatography conditions needed for optimum resolution of PTC-phosphoamino acids on the HPLC system using standard phosphoamino acids. An example of PTC-AAA resolution of pSer, pThr, pTyr, and hydroxyproline is presented in Figure 11.9.8. CAUTION: Extended use of silica-based RP-HPLC columns near neutral pH (as in Fig. 11.9.8) can shorten the useful lifetime of the column.
3. Estimate response factors from analysis of standard phosphoamino acids and by hydrolysis and PTC-AAA of standard synthetic phosphopeptides of defined phosphate content (see Support Protocol 11). Amino Acid Analysis
4. Perform concurrent HCl hydrolysis and analysis of control and unknown samples to verify the efficacy and accuracy of the analytical process.
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K
Hyp
IPUT pThr pTyr
PTU
Y
ETPU I L F
V P
D pSer E
SG
R H
M
A T
Aniline C
0
5
10 Minutes
DIPTU
15
20
Figure 11.9.8 RP-HPLC resolution of PTC-phosphoamino acids and hydroxyproline. PTC-derivatized amino acid Standard H plus pSer, pThr, pTyr, and Hyp (120 pmol) were chromatographed using the PE Applied Biosystems 130 HPLC and PE Applied Biosystems PTC C18 column (2.1 × 220–mm) with a flow rate of 300 µl/min, column temperature of 36°C, and the following gradient: time 0, 7% B; time 4 min, 14% B; time 10 min, 34% B; time 20 min, 66% B; and time 25 min, 100% B. Solvent A was 120 mM sodium acetate, pH 6.45, and solvent B was 31.5 mM sodium acetate containing 70% acetonitrile. Amino acids are identified using the single-letter amino acid code (see Table A.1A.1). Abbreviations (also see Fig. 11.9.1): pSer, phosphoserine; pThr, phosphothreonine; pTyr, phosphotyrosine; Hyp, hydroxyproline.
Quantification of pSer as PTC-S-Ethylcysteine Phosphoserine (pSer) can be quantified by first converting it to S-ethylcysteine (see Meyer et al., 1991), then hydrolyzing and derivatizing it for PTC-AAA. A synthetic phosphopeptides containing a well-defined amount of pSer is used as a standard.
SUPPORT PROTOCOL 17
Additional Materials (also see Basic Protocol and Support Protocols 7 and 8) Performic acid solution: 95:5 (v/v) performic acid (Sequenal grade)/hydrogen peroxide (30% by mass) Modification mixture (see recipe) Acetic acid Standard synthetic phosphopeptides with defined phosphate concentration Additional reagents and equipment for vapor-phase hydrolysis (see Support Protocol 7) or liquid-phase hydrolysis (see Support Protocol 8) 1. Dry sample in a 6 × 50–mm tube. Add 5 µl performic acid solution and incubate 1 hr on ice to oxidize cysteine to avoid false positives. Redry. 2. Add 20 µl modification mixture. Incubate 1 hr at 50°C under nitrogen. Acidify with 5 µl acetic acid and vacuum dry. 3. Perform standard vapor- or liquid-phase HCl hydrolysis (see Support Protocol 7 or 8). 4. PTC derivatize and analyze (see Basic Protocol, steps 8 to 14) in comparison with synthetic standard phosphopeptides. pThr and pTyr do not form equivalent derivatives. O-Glycosylated Ser will also yield S-ethylcysteine in this procedure.
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SUPPORT PROTOCOL 18
SUPPORT PROTOCOL 19
Analysis for Hydroxyproline Perform standard HCl hydrolysis (see Support Protocol 7 or 8) and PTC derivatization (see Basic Protocol). No special hydrolysis or modification methods are required to quantify hydroxyproline; however, PTC-hydroxyproline may coelute with other residues such as glutamic acid (Glu). Prepare a hydroxyproline amino acid standard (Pierce) and optimize its elution position in the HPLC system by changing the pH of the chromatography solvents, generally to a slightly more alkaline pH. For example, changing the solvent pH from 5.4 to ∼5.6 with 5 N NaOH in the PE Applied Biosystems PTC analysis system provides adequate resolution of hydroxyproline from Glu without changes in the gradient conditions (Fig. 11.9.1). To resolve hydroxyproline from pThr and pTyr in the PE Applied Biosystems system, increase the ionic strength of chromatography solvent A from 50 to ~120 mM sodium acetate (Mallinkrodt) and the pH from 5.4 to ~6.5 (Fig. 11.9.8). IDENTIFICATION OF PROTEINS FROM AAA DATA Amino acid analysis in combination with computer search programs (see UNITS 2.4 & 2.5) provides a valuable and rapid method for the identification of known and homologous proteins. Two useful search programs readily accessible via the World Wide Web are 1. PROPSEARCH (http://www.EMBL-Heidelberg.de/aaa.html) and 2. ExPASy (http://expasy.hcuge.ch/ch2d/aacompi.html). These programs are relatively tolerant to error and provide a very good chance of identifying proteins in the database. More information about PROPSEARCH can be found in Hobohm et al. (1994), about ExPASy in Wilkins et al. (1996), and about applications of the methods in resource facilities in Schegg et al. (1997). 1. Contact the Website for PROPSEARCH at http://www.EMBL-Heidelberg.de/aaa.htm or ExPASy at http://expasy.hcuge.ch/ch2d/aacompi.html. Enter compositional mole % data (see Basic Protocol, step 34) on the Internet forms; also enter optional information about molecular weight and isoelectric point (pI) if available. 2. Execute both search programs and compare results. 3. Evaluate the computer hit lists. Smaller scores are better for both programs. For PROPSEARCH, scores <1.5 are worthy of attention, scores between 1.5 and 2.5 generally indicate less reliable identification, and scores >2.5 indicate unreliable identification. For ExPASy, scores of 0 to 25 are worthy of attention, scores of 25 to 50 provide less reliable identification, and scores >50 indicate unreliable identifications. (For examples, see Anticipated Results and Figs. 11.9.9 and 11.9.13.)
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Modification mixture 80 µl ethanol 65 µl 5 N NaOH 60 µl ethanethiol 400 µl H2O, added last Amino Acid Analysis
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COMMENTARY Background Information This unit is meant primarily for readers with little or no experience in amino acid analysis (AAA) and focuses on the PTC method because (1) it is an established and reliable method of high-sensitivity AAA, (2) it is the most widely used method for picomole-level AAA, and (3) it is easier to find advice and knowledgeable assistance regarding the PTC method than any other method of high-sensitivity AAA. Distinguishing characteristics of amino acid analysis AAA is deceptively simple in concept, but it is one of the most difficult analytical methods in all of protein chemistry. The greatest value and benefit of the analysis is its quantitative capability, but measuring many different amino acids with quantitative accuracy is precisely what makes the analysis so difficult. AAA is a statistical type of measurement; peptide bonds and amino acids vary widely in their stability/lability to acid hydrolysis, oxidation, and modification. As noted previously, Trp and Cys are destroyed by HCl hydrolysis, Ser and Thr are partially destroyed, Ile and Val are slow to cleave, Met is subject to oxidation, and Asn and Gln are completely deamidated. Variable but ubiquitous background contamination plus variable instrument and analyst performance contribute to the statistical nature of the measurement and can confound the analysis, particularly at and below the low picomole level. Nevertheless, AAA remains a powerful tool. The technology offers the best way to quantify peptides and proteins in many biochemical studies, and it provides invaluable support to primary structure studies. It also provides quantitative compositional characterizations useful in verification of synthetic and recombinant polypeptide structures, identification of odd or modified residues, and identification of proteins by way of computer database searches. Overview of AAA methods Several methods of quantitative AAA exist, but all essentially involve either postcolumn derivatization following ion-exchange chromatography or precolumn derivatization followed by reversed-phase HPLC. The classical AAA methodology involves ion-exchange chromatography and postcolumn continuous reaction with ninhydrin (Moore and Stein, 1949; Moore et al., 1958). Moore and Stein were awarded the 1972 Nobel Prize in Chemistry for devel-
opment of quantitative AAA technology. Advantages of the classical method include excellent instrumentation, valuable technical support from instrument vendors, and the ability to analyze samples in the presence of salt and other contaminants. In fact, protein in SDSPAGE bands may be quantified using ion-exchange amino acid analyzers following HCl hydrolysis of the polyacrylamide gel band (Stein and Brink, 1981a; Williams and Stone, 1995). The primary disadvantage of the ion-exchange postcolumn ninhydrin method is that its sensitivity is limited to the high picomole/low nanomole range, which means that >2 µg of sample per analysis is usually required to obtain useful information (typically, 5 to 10 µg per analysis is required). However, fluorescence detection in place of ninhydrin enhances the sensitivity of the classical methodology to the low picomole range by using reagents such as fluorescamine or o-phthalaldehyde (OPA). A disadvantage of fluorescamine and OPA fluorescence–detection, postcolumn ion-exchange amino acid analyzers is that proline, a secondary amine, is not detected without an additional and relatively intricate oxidation step to convert the residue to a detectable primary amine (Stein and Brink, 1981b). A number of precolumn-derivatization reversed-phase HPLC methods for AAA have evolved that provide useful and more sensitive alternatives to classical AAA. The precolumn methods are distinguished from one another by different derivatization chemistry and are more sensitive and generally less expensive to set up than the ninhydrin postcolumn method. Common precolumn derivatizing reagents for AAA include phenylisothiocyanate (PITC; Koop et al., 1982; Heinrikson and Meredith, 1984; Tarr, 1986), dimethylaminoazobenzenesulfonyl chloride (DABSYL; Knecht and Chang, 1986), OPA (Jones and Gilligan, 1983), 9-fluorenylmethyl chloroformate (Fmoc; Bank et al., 1996), OPA plus Fmoc (Blankenship et al., 1989), and 6-aminoquinnolyl-N-hydroxysuccinyl carbamate (AQC; Strydom and Cohen, 1994). Excellent instrumentation and valuable technical support are also available from instrument vendors for many of the precolumn approaches. Disadvantages of precolumn methods include sensitivity to derivatization interference from salt and other contaminants, multiple reaction by-products that may obscure chromatographic resolution of the amino acids, and variable stability of the derivatives. How-
Chemical Analysis
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ever, effective ways of avoiding, minimizing, or eliminating these complications have been established for most of the precolumn methods. Performance comparisons of the various precolumn and postcolumn AAA methods and documentation of the instrumentation used in studies spanning almost a decade and involving up to 77 different resource laboratories can be found in the annual AAA studies of the Association of Biomolecular Resource Facilities (ABRF; Niece et al., 1989; Crabb et al., 1990; Tarr et al., 1991; Strydom et al., 1992, 1993; Yücksel et al., 1994, 1995; Mahrenholz et al., 1996; Schegg et al., 1997). From 1989 through 1996, ~38% of the participants in these studies used the postcolumn ninhydrin methodology, ~48% used precolumn PTC methods, and 14% used other AAA methods. Overall these studies demonstrate that more than one method of AAA works well. It is important to recognize that the skill and experience of the analyst are probably the most important factors for successful AAA. Costs of PTC-AAA Excluding the investment in instrumentation, labor is the largest expense in PTC-AAA. Labor costs may be estimated on a per analysis basis by multiplying the hourly wage of the analyst by 1 to 2. Supply costs are modest and typically account for ~25% of the per analysis cost. For heavy-use AAA systems, a service contract from the instrument vendor may be worthwhile; the service contract can cost up to ~$8000 per year. A gradient HPLC system with at least a simple integrator is essential ($25,000 to $35,000 in 1996); add another ~$10,000 for an automatic sample injector. In 1996, the list price of a dedicated PTC-AAA system ranged from $50,000 to $80,000 depending on options and vendor.
Amino Acid Analysis
Where to find help Any biomolecular resource facility that routinely performs AAA is an excellent source of general information about AAA and also may be of assistance in troubleshooting. The ABRF sponsors an Amino Acid Analysis Research Committee that can help answer many questions concerning how to perform amino acid analysis. In addition, the ABRF publishes a Yellow Pages listing of Resource Facilities that will perform AAA and/or other analytical and synthetic services for out-of-house clients, generally for a fee. The ABRF Yellow Pages and Amino Acid Analysis Research Committee membership are accessible via the World Wide
Web at the ABRF homepage (http://www.medstv. unimelb.edu.au/abrf.html) or by contacting the ABRF Business Office, FASEB, 9650 Rockville Pike, Bethesda, MD 20814-3998. As previously noted, the service and technical support offered by instrument companies selling amino acid analyzers can also be useful.
Critical Parameters Performance overview Essentially all low-picomole level AAA methods are labor-intensive, high-maintenance processes subject to a variety of possible complications. High-quality PTC-AAA performance should be expected but not without costs in time and energy. Vigilant effort is required to verify that the PTC-AAA system is functioning in a quantitatively reliable and trustworthy fashion. At least three analyses of an amino acid standard (e.g., Pierce Standard H) should be performed every day the system is used to ensure accurate calibration. In addition, daily analysis of a peptide or protein standard of known concentration is recommended to demonstrate that the performance provides accurate quantitation and low compositional error. Analysis of blank samples (e.g., solvent and reagent aliquots, empty PVDF pieces, and empty hydrolysis tubes) should be performed regularly and as needed to quantify background levels of amino acids and “junk” peaks in the chromatography. If the analysis system sits idle for a few weeks, expect to spend several days to a week reestablishing reliable performance levels. To maintain a reliably accurate high-sensitivity analysis system, expect that half of the overall analyses performed on the system may involve standards, blanks, and various tests (e.g., to establish Trp or Cys response factors), especially if the demand for experimental sample analysis is modest. Maintenance Instrument and system maintenance is a key factor in achieving successful performance from PTC-AAA. A systematic schedule for instrument maintenance should be established and all maintenance activities documented. Good record keeping practices will facilitate troubleshooting efforts when performance problems occur. Maintenance activity will vary from system to system, but daily maintenance activities should include preparing new derivatization reagents (for manual derivatization), cleaning the vacuum system cold trap, and checking for vacuum system leaks, HPLC de-
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tector lamp stability, HPLC leaks, and the quality of HPLC resolution of the amino acids. Weekly maintenance activities should include changing chromatography buffers and derivatization reagents, performing available selftests on automatic instrumentation, cleaning away dust on and around the instrumentation, changing the vacuum pump oil, and cleaning the Speedvac evaporator. Monthly maintenance activities should include changing inline filters and fan filters on the HPLC and/or automatic derivatizer. Periodically, HPLC pump seals (about twice a year), the UV detector lamp (1 to 2 per year), and HPLC columns (2 to 4 per year) must be replaced. Automatic micropipettors must also be calibrated and maintained in clean condition to achieve accurate quantitation in amino acid analysis. Contamination Background contamination plagues all high-sensitivity AAA procedures, not just PTC-AAA, and it seems to come from everywhere when attempting to perform measurements at the low-picomole level. Dust in the air can be one of the most difficult sources to deal with; dust can contribute high levels of glycine, serine, and alanine to the results. Keep the laboratory room as clean and tidy as possible. Locate the AAA workstation and instrumentation in a low-traffic area, and avoid placing the work area directly under heater and air conditioner vents. All reagents and solvents can be sources of contaminants (e.g., water, HCl, derivatizing chemicals, and sample preparation buffers). Use the highest purity solvents and reagents available. To minimize UV-absorbing phenylthiourea by-products, avoid ammonia-based solvents. Be aware of the age of the solvents and reagents purchased and use the newest lots available. Date the chemicals as they are received in the laboratory to assist with troubleshooting when problems arise. During the analysis, keep solvents and reagents in covered amber bottles away from sunlight. Store phenylisothiocyanate (PITC), diisopropylethylamine (DIEA), triethylamine (TEA) amino acid Standard H, and peptide/protein standards at −20°C under argon or nitrogen. Monitor the chromatography solvents for “floaters”; bacteria grow well in sodium acetate buffers, and sunlight enhances their growth. Meticulous sample handling technique is also essential for high-quality analyses. Glassware and other lab ware that comes in contact
with the sample or is used in preparation of reagents and chemicals for hydrolysis, derivatization, and analysis are other sources of contaminants and should always be cleaned before use. Keep micropipet tips and 6 × 50–mm tubes in a dust-free, covered box. Avoid touching pipet tips, avoid unnecessary opening and closing of sample vials, and wear powder-free gloves when handling samples. A more detailed description of sources of contamination and ways to reduce background in PTC-AAA can be found in Atherton (1989).
Troubleshooting Table 11.9.1 is a brief, generic troubleshooting guide for common problems encountered in PTC-AAA; it is not meant to be all-inclusive. The analyst should query instrument companies providing support for PTC-AAA about the availability of more detailed troubleshooting guides for PTC-AAA on their particular instrumentation.
Anticipated Results Average PTC-AAA data The quality of AAA results is significantly influenced by the amount of sample hydrolyzed, as shown in the PTC-AAA results from pyridylethyl ribonuclease (see Table 11.9.2). These data represent average results from two to three hydrolyses and illustrate that compositional error and variability increase as the amount of protein hydrolyzed decreases. Lower recovery is also evident from the smaller amounts of sample hydrolyzed. With clean sample preparations and careful technique, an average compositional error of ≤10% can be obtained with 0.5 to 1 µg of sample hydrolyzed, with the best analyses showing 0% to 5% error from a single hydrolysis/analysis. Data exhibiting relatively high compositional error can still provide useful estimates of the quantity of protein adequate for many applications (e.g., estimating whether there is enough sample to justify proteolysis and subsequent peptide purification and sequence analysis). Furthermore, the results from 0.2 µg of hydrolysate in Table 11.9.2 allow correct identification of the sample via the PROPSEARCH and ExPASy computer database queries (Fig. 11.9.9), despite 21% average compositional error. Realistic expectations for the quality of AAA results can be obtained from reviewing the published AAA studies from the ABRF (Niece et al., 1989; Crabb et al., 1990; Tarr et al., 1991; Strydom et al., 1992, 1993; Yücksel
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Table 11.9.1
Troubleshooting Guide for PTC Amino Acid Analysisa
Problem
Possible cause
Possible solution
No peaks in HPLC chromatogram
Sample is not injected
Unclog or replace tubing; purge autosampler lines
UV lamp is not working
Check that the lamp is on; replace the lamp
No peaks on chromatogram but injection front present
Sample not being derivatized
Check/replace derivatizing reagents; replace lines or bottle seals; replace old reagents
No amino acid peaks detected but background “junk peaks” are present
No hydrolysis
Make certain hydrolysis vessel has a proper seal; make certain proper vacuum or inert gas is present Try with ten times more sample
Not enough protein/peptide in sample Poor peak size reproducibility and irregular recovery
Pipetting errors Incomplete hydrolysis Incomplete derivatization
Recalibrate pipettors Improper seal on hydrolysis vessel; replace slide valve Check deliveries of derivatizing reagents; replace lines, bottle seals, or reagents
Too much salt or detergent in sample
Desalt sample by RP-HPLC, microdialysis, Centricon, or precipitation
Improper sample injection UV lamp getting old
Unclog or replace tubing Replace UV lamp
Erroneous calibration file Incorrect concentration used for amino acids and/or protein standards
Recalibrate Replace standards
Incomplete hydrolysis and/or derivatization
Repeat hydrolysis and derivatization with new sample
Low Asp and Glu
Metal contamination from glass hydrolysis tubes
Prior to hydrolysis, dry samples in 6 × 50–mm tubes from 0.25 µg/µl EDTA
High amino acid background in control blanks
Contamination in solvents, reagents, glassware, micropipets, etc.
Change reagents/utensils; use more care during hydrolysis/derivatization
Dust (= Gly, Ser, and Ala) Hands and fingers
Clean workstation and instrument Wear powder-free gloves
Dirty column
Clean or replace column
High by-product (PTU, IPTU, aniline, etc.) and “junk” peaks in control blanks
Old derivatization reagents Inadequate drying step
Replace PITC, DIEA, etc. Vacuum dry longer
Unusually large and unidentifiable chromatography peaks
Air injected with sample
Completely fill (overfill) the sample loop with liquid; clean/replace clogged delivery lines; degas solvents; maintain constant solvent temperature Install narrower tubing and/or back pressure device on flow cell outlet
Inaccurate quantitation of standard peptides and proteins
Air bubble in flow cell
Amino Acid Analysis
continued
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Table 11.9.1
Troubleshooting Guide for PTC Amino Acid Analysisa, continued
Problem
Possible cause
Possible solution
Poor retention time reproducibility
Improper equilibration and/or gradient cycle
Extend equilibration time and/or redesign gradient
Leaky HPLC system Inconsistent column temperature
Tighten or replace leaky fittings, pump seals, tubing, column, and flow cell connections Control column temperature electronically
Faulty gradient or flow rate control
Repair or replace microprocessor controller and mixing chamber
Improper pH and/or composition of solvents and/or transfer buffer
Reevaluate buffer recipe, change formula as needed, and remake buffers
HPLC column too old Injection volume too large
Replace column Decrease injection volume
Old UV lamp
Replace lamp
Improper flow or mixing Poor electrical connections
Repair/replace leaks, microprocessor, and mixer Tighten/replace electrical connections between detector and integrator
Dirty/old column
Replace column
Poor peak shape
Noisy HPLC baseline
aAbbreviations:
DIEA, diisopropylethylamine; PITC, phenylisothiocyanate
et al., 1994, 1995; Mahrenholz et al., 1996; Schegg et al., 1997). These reports provide an overview of the accuracy obtained by all participants of the study itemized by method (e.g., postcolumn ninhydrin versus precolumn PTC, OPA/Fmoc, AQC) and usually highlight the best results from both precolumn and postcolumn methods. Reproducibility and calibration data The average peak areas (or peak heights) and retention times for standard amino acids vary with derivatization conditions (e.g., reagent concentration and age), chromatography conditions (e.g., injection reproducibility, solvent, column age, and temperature), pipetting accuracy, and background contamination. The AAA system reproducibility must be routinely evaluated. An example of relevant data for this purpose using two of the sixteen standard amino acids is shown in Table 11.9.3. This type of statistical evaluation should be performed for all sixteen amino acids used for calibration and summarized with average relative standard deviation (RSD) values as shown in Table 11.9.4. The reproducibility of the system must be maintained such that average RSD values for all sixteen amino acids do not exceed an upper limit, for example, 8% RSD for peak area and 0.4% RSD for retention time. RSD values typi-
cally increase with smaller sample amounts (see Table 11.9.2), indicating less reproducibility and more variability. Once the average peak area (or average peak height) has been satisfactorily determined for all the amino acids, response factors may be calculated by dividing the average areas by the picomoles analyzed. A calibration file should be compiled as illustrated in Table 11.9.5. The calibration file should include a retention time window which sets retention time limits for the integrator to identify and quantify the various peaks (e.g., ±0.15 min in Table 11.9.5). Sensitivity range Amino acids must be measured in the linear response range of the analysis system to obtain meaningful results. The appropriate amount of sample should allow quantification of both the low-abundancy and high-abundancy residues in a given peptide or protein. However, in practice, the results sometimes fall outside the meaningful response range because too much or too little sample was analyzed. The analyst must learn to recognize unreliable data beyond the upper limit of the range as well as be cognizant of the influences of background level contamination and hydrolysis losses in the lower end. An example of PTC-AAA data demonstrating a linear response between the
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amount analyzed and peak area is shown for the sixteen standard amino acids in Figure 11.9.10. For this particular analysis system, the response of most residues plateaus above 5 nmol; lysine, which has two PITC-reactive amino groups, plateaus at a lower level (~4 nmol). Another useful quantitative evaluation of every PTC-AAA system is a comparison of the amount of sample recovered following HCl hydrolysis. Average recovery data from 123 manual vapor-phase HCl hydrolyses of nine different proteins performed between ~65 and 2600 ng are shown in Figure 11.9.11 (West and Crabb, 1990). These data show that essentially quantitative recovery was obtained from amounts hydrolyzed in 6 × 50–mm tubes in the 400 to 2600–ng range but that losses occurred
Table 11.9.2
for amounts hydrolyzed at or below ~230 ng. Average data from 336 automatic vapor-phase HCl hydrolyses of the same nine proteins showed markedly better recovery from automatic than manual hydrolyses below the 400ng level and a concomitant improvement in compositional accuracy (West and Crabb, 1990). For the majority of analysts without automatic instrumentation, the sensitivity of PTC-AAA is not limited by the detection chemistry or current chromatography capabilities but rather by complications introduced by hydrolysis and background contamination. Sample amounts of 0.5 to 2 µg usually provide reasonable results for routine PTC-AAA following standard HCl hydrolysis conditions.
Average PTC AAA Results from Pyridylethyl Ribonucleasea,b
Amino acid
Known compositionc
Amount hydrolyzed (µg) 0.06
0.2
0.4
1.0
2.6
Residues per molecule Asx Glx Ser Gly His Arg Thr Ala Pro Tyr Val Met Ile Leu Phe Lys Pec
15 12 15 3 4 4 10 12 4 6 9 4 3 2 3 10 8
18.3 11.7 16.2 8.6 3.3 5.9 8.8 11.1 2.3 10.9 5.9 0.8 0.6 2.0 1.7 12.0 6.2
16.8 12.3 14.9 5.3 3.5 5.2 9.6 12.6 3.6 7.3 8.5 1.3 1.5 2.2 2.6 10.4 8.3
17.2 13.2 15.3 4.3 4.3 5.2 10.0 13.5 3.9 6.5 8.5 2.0 1.5 2.1 2.9 11.5 9.7
16.0 12.7 12.5 3.3 3.2 5.2 9.6 12.6 3.7 6.6 9.1 2.1 2.1 2.2 3.3 11.7 8.3
16.6 12.7 13.4 3.4 3.1 5.1 10.2 12.6 3.7 6.5 8.8 3.7 1.8 2.2 3.3 7.6 9.1
10 12
19 24
66 71
189 192
pmol analyzed pmol hydrolyzed
2.7 4.0
Number of analyses
2
2
3
3
3
15.7 34 67
13.2 21 82
3.7 17 78
2.8 11 108
2.2 9 96
Average peak area RSD Average % error Average % recovery aManual
Amino Acid Analysis
vapor-phase HCl hydrolysis was performed at 150°C for 1 hr. Automatic PTC derivatization and analysis were performed with PE Applied Biosystems models 420H, 130, and 920 instrumentation as described in West and Crabb (1990). bAbbreviations: Pec, pyridylethyl cysteine; RSD, % standard deviation or relative standard deviation. Amino acids are identified using the three-letter abbreviations (see Table A.1A.1). cRelative standard deviation (standard deviation of the mean/mean) × 100.
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Results from PROPSEARCH
Rank 1 2 3 4 5 6 7 8 9 10
ID
DIST
rnp_antam rnp_girca rnp_conta rnp_sheep rnp_bubar rnp_gazth rnp_aepme rnp_hipam rnp_macru >p1;s44712
1.89 1.95 2.12 2.14 2.29 2.30 2.32 2.62 2.65 2.65
LEN2 POS1 POS2
pl
DE
1 1 1 1 1 1 1 1 1 1
8.43 7.22 8.24 7.83 7.83 8.43 8.43 7.59 8.06 10.22
Ribonuclease Pancreatic Ribonuclease Pancreatic Ribonuclease Pancreatic Ribonuclease Pancreatic Ribonuclease Pancreatic Ribonuclease Pancreatic Ribonuclease Pancreatic Ribonuclease Pancreatic Ribonuclease Pancreatic Opacity Protein-Neisseria
124 124 124 124 124 121 124 124 122 192
Results from ExPASy
Rank 1 2 3 4 5 6 7 8 9 10
(http://www.embl-heidelberg.de/aaa.html)
124 124 124 124 124 121 124 124 122 192
(http://expasy.hcuge.ch/ch2d/aacompi.html))
Score
Protein
pl
Mw
Description
28 36 37 38 38 39 39 40 40 40
RNP_BOVIN RNP_CONTA RNP_DAMKO POLG_POL1M POLH_POL1M POLG_POL32 POLG_POL3L POLG_POL2L POLG_POL2W RNP_TAUOR
8.64 8.64 9.60 8.25 8.25 8.20 8.20 8.25 8.25 9.30
13690 13686 13709 7385 7385 7313 7313 7339 7339 13742
Ribonuclease Pancreatic Ribonuclease Pancreatic Ribonuclease Pancreatic Coat Protein VP4 Coat Protein VP4 Coat Protein VP4 Coat Protein VP4 Coat Protein VP4 (P1A) Coat Protein VP4 (P1A) Ribonuclease Pancreatic
Figure 11.9.9 Identification of RNase from PTC-AAA data. The PTC-AAA data in Table 11.9.2 from hydrolysis of 12 pmol (∼0.2 µg) ribonuclease (RNase) was converted to mole % and entered into the PROPSEARCH and ExPASy database query programs for identification of the protein. The following mole % data were entered: Arg 4.42, Met 1.11, Glx 10.46, Ile 1.28, Ser 12.67, Leu 1.87, His 2.98, Phe 2.21, Gly 4.51, Lys 8.84, Thr 8.16, Asx 14.29, Ala 10.71, Pro 3.06, Tyr 6.21, Val 7.23, sum 99.81. The top ten scores from both programs are displayed. Despite the 21% compositional error in the AAA data, both programs correctly identified the protein. Abbreviations for the PROPSEARCH results: DIST, score; LEN2, length of sequence; POS1 and POS2, begin and end of the sequence; pI, isoelectric pH; DE, description. PROPSEARCH identification scores <1.5 are best, 1.5 to 2.5 are less reliable, and scores >2.5 are unreliable; for ExPASy, scores 0 to 25 are best, 25 to 50 less reliable, and >50 unreliable.
Using raw AAA data to identify a protein Once the AAA system is calibrated and the precision and linear response range of the system are established, experimental results can be generated as shown in Figure 11.9.12. Raw data from PTC-AAA of a 36-kDa protein are presented; the picomoles of amino acids have been calculated from peak area using a calibration file stored in the integrator (Fig. 11.9.12). Before proceeding to any other calculation, the analyst should verify that the amino acid peaks have been correctly identified. The amino acid composition of the protein may then be calculated from the picomole values as described and exemplified in Figures 11.9.2 through 11.9.7. The easiest amino acid compositional calculation is mole %, and this is the format that is
normally required to identify the protein by computer query of composition databases. Entering the mole % data calculated from Figure 11.9.12 into the PROPSEARCH and ExPASy database query programs provides correct identification of the protein as the cellular retinaldehyde-binding protein (CRALBP; Fig. 11.9.13). Ambiguity exists regarding the species in this identification (bovine and human CRALBP are 91% identical); however, the source of most AAA samples is known, and this does not constitute a major problem. The raw data in Figure 11.9.12 were generated from hydrolyzing ~1.9 µg of protein and analyzing 50% of the hydrolysate. The data exhibit ~6.9% average compositional error using the calculation method described in Figure
Chemical Analysis
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Table 11.9.3
Amino Acid Analysis Reproducibility and Calibration Dataa
Sample ID
Retention time (min)
Peak height
pmol by height
Peak area
pmol by area
Peak ID: Alanine 31102001 31102002 31102003 31202001 31202002
9.60 9.65 9.67 9.62 9.67
45424 45252 43070 43397 46658
210.1 209.3 199.2 200.7 215.8
197668 197040 190578 192789 203824
260.1 259.3 250.8 286.3 302.7
Minimum Maximum Average Std dev RSD
9.60 9.67 9.64 0.03 0.31
43070 46658 44760 1500 3
199.2 215.8 207.0 6.9 3.4
190578 203824 196380 5101 3
250.8 302.7 271.8 21.8 8.0
31102001 31102002 31102003 31202001 31202002
8.82 8.87 8.88 8.83 8.88
46723 47794 48632 44862 49562
210.5 215.3 219.1 202.1 223.3
199489 207920 210325 195017 211775
222.5 231.9 234.5 243.8 264.8
Minimum Maximum Average Std dev RSD
8.82 8.88 8.86 0.03 0.34
44862 49562 47515 1815 4
202.1 223.3 214.0 8.2 3.8
195017 211775 204905 7293 4
222.5 264.8 239.5 16.8 6.7
Peak ID: Arginine
aAmino
acid Standard H (300 pmol, Pierce) was analyzed five times; results from two of the sixteen amino acids are shown before instrument calibration. Std dev, standard deviation of the mean; RSD, % standard deviation of the mean or relative standard deviation. Automatic PTC derivatization, analysis, and statistical calculations were performed with PE Applied Biosystems models 420H, 130, and 920 instrumentation.
11.9.6. Note that the PROPSEARCH and ExPASy scores for CRALBP in Figure 11.9.13 are much lower and in a more reliable range than those for RNase in Figure 11.9.9, where the PTC-AAA data were derived from a smaller amount of sample (0.2 µg hydrolyzed) and exhibited greater error (21%). Despite ambiguities and a margin for error, computer searches of amino acid composition databases offer a valuable tool for the identification of proteins from minute amounts of sample. This relatively new application of amino acid analysis will likely become more important and widespread as more compositional data become available from gene sequencing.
Time Considerations
Amino Acid Analysis
Amino acid analysis is a time-consuming process, and new practitioners should anticipate several weeks to initially establish a system in addition to ongoing effort to maintain the system performance in a trustworthy fash-
ion. The following time allocations for the various steps in PTC-AAA are reasonable after developing some experience with the methodology. Sample preparation is the most variable and potentially the longest step. Assuming purification has been accomplished, desalting and/or sample concentration can usually be achieved within a few hours. The standard HCl hydrolysis step requires an hour for the vapor-phase method and 20 to 24 hr for the liquid-phase method; microwave vapor-phase HCl hydrolysis requires 20 min or less. Many samples can be hydrolyzed at once by any of these methods; allow ~1 hr total for handling 20 samples before and after heating (e.g., setting up the hydrolysis and drying the hydrolysates). The manual PTC derivatization reaction requires a 20-min incubation; allow ~30 min for handling 20 samples (e.g., setting up the reaction and drying the final product).
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Table 11.9.4
Peak ID
Summary of Reproducibility and Calibration Dataa
Number Average retention of peaks time (min)
Asx Glx Ser Gly His Arg Thr Ala Pro Tyr Val Met Ile Leu Phe Lys
5 5 5 5 5 5 5 5 5 5 5 5 5 5 5 5
5.81 6.16 7.59 7.99 8.26 8.86 9.31 9.64 9.88 12.40 13.35 13.78 15.49 15.72 16.56 17.69
Average RSD
RSD
Average peak height
RSD
Average peak area
RSD
0.51 0.41 0.40 0.38 0.37 0.34 0.33 0.31 0.27 0.19 0.21 0.20 0.16 0.19 0.18 0.17
61112 46318 39202 35917 40133 47515 44320 44760 29321 60110 50073 49079 51292 48384 53671 80504
2.95 2.67 3.17 2.88 2.62 3.82 2.87 3.35 2.52 3.91 3.29 7.28 3.72 4.06 3.59 4.98
253139 201011 177504 163656 177585 204905 192943 196380 127821 263636 220594 216244 229503 211669 234958 357280
9.20 7.34 6.23 7.45 6.94 6.70 5.57 8.03 7.05 7.74 6.85 7.83 7.42 7.10 7.04 6.69
0.29
3.61
7.20
aSummary
of average retention times, peak heights, peak areas, and relative standard deviation (RSD) values (reproducibility) from five PTC analyses of 300 pmol of amino acid Standard H. Response factors are calculated by dividing average peak heights and average peak areas by picomoles of amino acid analyzed (300 pmol in this case).
Table 11.9.5
Peak ID Asx Glx Ser Gly His Arg Thr Ala Pro Tyr Val Met Ile Leu Phe Lys
Amino Acid Standard Calibration Filea
Response factors
Retention time (min)
Retention time window (min)
Height per pmol
Area per pmol
5.8 6.2 7.6 8.0 8.3 8.9 9.3 9.7 9.9 12.4 13.4 13.8 15.5 15.7 16.6 17.7
0.15 0.15 0.15 0.15 0.15 0.15 0.15 0.15 0.15 0.15 0.15 0.15 0.15 0.15 0.15 0.15
203.7 154.4 130.7 119.7 133.8 158.4 147.7 149.2 97.7 200.4 166.9 163.6 171.0 161.3 178.9 268.4
843.8 670.0 591.7 545.5 592.0 683.0 643.1 654.6 426.1 878.8 735.3 720.8 765.0 705.6 783.2 1190.9
aResponse
factors were calculated from the average peak heights and peak areas listed in Table 11.9.4. The retention time window restricts amino acid peak identification to a defined time span, in this case ±0.15 min about each retention time.
Chemical Analysis
11.9.37 Current Protocols in Protein Science
Supplement 7
7.0 6.3 5.0 3.7
Asp
1.8
Area (x106)
7.8 6.8 5.8 4.0
Glu
1.9
8.0 7.9 5.4 3.7 1.8
His
7.8 7.2 5.7 3.9
Arg
9.7 8.5 6.7 4.4
Ser
8.9 7.9 5.8 4.1
Gly
8.5 7.4 5.8 4.0
9.2 8.5 6.8 4.7
Thr
9.3 8.6 6.6 4.6
1
2
3
4
5
Tyr
9.8 8.8 6.5 4.5
Leu
2.0
Val
9.7 8.6 6.5 4.4
2.1
Ala
10.4 9.5 6.7 4.6 2.1
1.9
1.9
lle
2.2
1.9
9.1 8.2 6.1 4.2
9.4 8.6 6.6 4.6 2.1
2.2
1.9
8.5 7.4 5.7 4.0 1.9
Pro
1
2
3
4
5
Phe
2.0
Met
14.1 12.9 10.4
Lys
3.1 1.0 1
2
3
4
5
1
2
3
4
5
Amount analyzed (pmol x 103)
Figure 11.9.10 Linear response range for sixteen PTC amino acids. PTC amino acid Standard H (Pierce) was analyzed in replicate using PE Applied Biosystems models 420H, 130, and 920 instrumentation and PTC-AAA column (2.1 × 220–mm). A linear response of peak area versus amount analyzed was obtained between 10 pmol and 5 nmol for most residues (data <1 nmol not shown).
Percent recovery
100 2000
75 50 1000
25 0
0 0
Amino Acid Analysis
Amount recovered (ng)
3000
1000 2000 Amount hydrolyzed (ng)
3000
Figure 11.9.11 PTC-AAA linear response range following HCl hydrolysis. Nine different proteins were subjected to manual vapor-phase HCl hydrolysis in replicate and 123 analyses performed using PE Applied Biosystems PTC-AAA models 420H, 130, and 920 instrumentation. Average amounts hydrolyzed (in nanograms) are plotted versus both the average amounts recovered (in nanograms) and the percent recovery. These results demonstrate a linear response from ∼400 to 2600 ng hydrolyzed but also show that losses occur for hydrolyses ≤230 ng (West and Crabb, 1990).
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Time requirements vary for the chromatography step depending on the HPLC system but typically involve 30 to 60 min from injection to injection (e.g., 15 to 30 min for separation followed by another 15 to 30 min to clean and reequilibrate the column). Allow ∼30 min to dissolve 20 samples and to load or program an autosampler. If no autosampler is available,
0
5
allow all day for manual injections. Automatic derivatization and analysis with or without automatic hydrolysis generates about one analysis per hour with significantly reduced time required for sample handling. Once the calibration process has become routine, plan on ~30 to 45 min to check and adjust retention time windows, reprocess data,
10 Minutes
15
20
Peak ID
Retention time (min)
Peak area
pmol by area
Asx Glx Ser Gly His Arg Thr Ala Pro PTU Aniline Tyr Val Met Ile Leu Phe Lys IPTU DIPTU
5.05 5.32 7.15 7.59 7.99 8.68 9.08 9.40 9.67 9.98 11.40 11.98 12.70 13.14 14.43 14.60 15.43 16.41 17.17 19.54
3250442 6767374 1235546 1952006 451547 2097680 1576640 2837580 1668884 564676 228556 1340244 2790862 753814 1360616 3705736 3253882 3545286 340468 896444
1162.44 2729.84 728.13 1175.61 271.38 1114.78 763.55 1294.95 680.52 –––– –––– 450.25 960.13 323.62 521.45 1624.57 117.95 789.21 –––– ––––
Figure 11.9.12 Raw data from PTC-AAA. A typical PTC-AAA chromatogram and raw data reduction of peak area to picomoles of amino acid is shown for one analysis of a 36-kDa protein, human recombinant cellular retinaldehyde-binding protein (CRALBP; 1.9 µg hydrolyzed, 50% or 950 ng analyzed). Amino acids are identified using the three-letter code (see Table A.1A.1).
Chemical Analysis
11.9.39 Current Protocols in Protein Science
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Results from PROPSEARCH (http://www.embl-heidelberg.de/aaa.html) Rank DIST LEN2 POS1 POS2 pl ID DE 1 2 3 4 5 6 7 8 9 10
cral_bovin cral_human cral_human y701_haein >p1;c64012 spsd_bacsu >p1;s39721 >p1;s43135 sys_coxbu gba1_soybean
0.68 0.71 0.71 1.79 1.79 1.80 1.82 1.89 1.89 1.93
Results from ExPASy Protein Rank Score 1 2 3 4 5 6 7 8 9 10
6 31 32 32 36 37 38 38 38 38
1 1 1 1 1 1 1 1 1 1
316 316 318 339 339 289 289 423 423 385
CRAL_HUMAN RO52_HUMAN GRK6_HUMAN RYNR_HUMAN CP27_HUMAN SYD_HUMAN VAV_HUMAN STA5_HUMAN RFP_HUMAN RTC1_HUMAN
316 316 316 339 339 289 289 423 423 385
4.84 4.82 4.82 5.87 5.87 6.42 6.42 6.07 6.07 5.41
Cellular Retinaldehyde-Binding Cellular Retinaldehyde-Binding Cellular Retinaldehyde-Binding Hypothetical protein Hl0701 Hypothetical protein Hl0701 Spore Coat Polysaccharide Hypothetical Protein-Bacillus Serine-tRNA Ligase-Coxiella Seryl-tRNA Synthetase Gaunine Nucleotide-Binding
(http://expasy.hcuge.ch/ch2d/aacompi.html) Description pl Mw
4.98 5.98 8.33 5.19 8.43 6.20 6.20 5.98 5.83 6.34
36343 54169 65968 64429 56908 57192 98354 90646 58489 23294
Cellular Retinaldehyde-Binding 52 kd RO Protein (Sjogren synd) G Protein-Coupled Receptor Kinase Ryanodine Receptor, Skeletal Sterol 26-Hydroxylase Aspartyl-tRNA Synthetase Vav Oncogene Signal Transducer Transforming Protein RFP Ras-Like Protein TC21
Figure 11.9.13 Identification of CRALBP from PTC-AAA. The raw PTC-AAA data in Figure 11.9.12 from analysis of ∼27 pmol recombinant human CRALBP was converted to mole % and entered into the PROPSEARCH and ExPASy database query programs for identification of the protein. The following mole % data were entered: Arg 7.07, Met 2.05, Glx 17.32, Ile 3.31, Ser 4.62, Leu 10.31, His 1.72, Phe 7.44, Gly 7.46, Lys 5.01, Thr 4.84, Asx 7.37, Ala 8.22, Pro 4.32, Tyr 2.86, Val 6.09, sum 100.01. The top ten scores from both programs are displayed. No species designation was used in the PROPSEARCH query; however, human was the designated species in the ExPASy search. The AAA data exhibit 6.9% compositional error. Abbreviations and scoring parameters are as in Figure 11.9.9.
and adjust response factors in the integrator. Data reduction of experimental samples using the computer-based spreadsheets and methods of calculation presented in this unit requires up to 1 hr for 5 to 10 analyses. Protein identification using the Internet and PROPSEARCH and ExPASy search programs typically requires 15 to 30 min per search. Allow another 15 to 30 min per day for general instrument maintenance.
Literature Cited Applied Biosystems. 1989. Model 420A Derivatizer/Analyzer Amino Acid System User’s Manual, Version 3.2.2 pp. 4-5. Applied Biosystems, Foster City, Calif. Amino Acid Analysis
Atherton, D. 1989. Successful PTC amino acid analysis at the picomole level. In Techniques in
Protein Chemistry (T. Hugli, ed.) pp. 273-283. Academic Press, San Diego. Bank, R.A., Jansen, E.J., Beekman, B., and te Koppele, J.M. (1996). Amino acid analysis by reverse-phase high performance liquid chromatography: Improved derivatization and detection conditions with 9-fluorenylmethyl chloroformate. Anal. Biochem. 240:167-176. Bidlingmeyer, B.A., Cohen, S.A., and Tarvin, T.L. 1984. Rapid analysis of amino acids using precolumn derivatization. J. Chromatogr. 336:93104. Bidlingmeyer, B.A., Tarvin, T.L., and Cohen, S.A. 1986. Amino acid analysis of submicrogram hydrolyzate samples. In Methods in Protein Sequence Analysis (K.A. Walsh, ed.) pp. 229-245. Humana Press, Clifton, N.J. Blankenship, D.T., Krivanek, M.A., Ackermann, B.L., and Cardin, A.D. 1989. High-sensitivity amino acid analysis by derivatization with o-
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phthalaldehyde and 9-fluorenylmethyl chloroformate using fluorescence detection: Applications in protein structure determination. Anal. Biochem. 178:227-232. Bozzini, M., Bello, R., Cagle, N., Yamane, D., and Dupont, D. 1991. Amino acid analysis, tryptophan recovery from autohydrolyzed samples using dodecanethiol. Appl. Biosystems Res. News, February, pp. 1-4.
unique isozyme of cytochrome P450 from liver microsomes of ethanol-treated rabbits. J. Biol. Chem. 257:8472-8480. Kuhn, C.C. and Crabb, J.W. 1986. Modern manual microsequencing methods. In Advanced Methods in Protein Microsequence Analysis (B. Wittmann-Liebold, J. Salnikow, and V.A. Erdmann, eds.) pp. 64-76. Springer-Verlag, Berlin.
Crabb, J.W., Ericsson, L., Atherton, D., Smith, A.J., and Kutny, R. 1990. A collaborative amino acid analysis study from the Association of Biomolecular Resource Facilities. In Current Research in Protein Chemistry (J.J. Villafranca, ed.) pp. 49-61. Academic Press, San Diego.
Mahrenholz, A.M., Denslow, N.D., Andersen, T.T., Schegg, K.M., Mann, K., Cohen, S.A., Fox, J.W., and Yücksel, K.Ü. 1996. Amino acid analysis– recovery from PVDF membranes: ABRF95AAA. In Techniques in Protein Chemistry VII (D.R. Marshak, ed.) pp. 323-330. Academic Press, San Diego.
Dupont, D.R., Keim, P.S., Chui, A.H., Bello, R., Bozzini, M., and Wilson, K.J. 1989. A comprehensive approach to amino acid analysis. In Techniques in Protein Chemistry (T. Hugli, ed.) pp. 284-294. Academic Press, San Diego.
Meyer, H.E., Hoffman-Posorske, E., and Heilmeyer, L.M.G. 1991. Determination and location of phospogerine in proteins and peptides by convers io n to S-ethylcysteine. Methods Enzymol. 201:169-185.
Engelhart, W.G. 1990. Microwave hydrolysis of peptides and proteins for amino acid analysis. Am. Biotechnol. Lab. 15:31-34.
Moore, S. and Stein, W.H. 1949. Photometric ninhydrin method for use in the chromatography of amino acids. J. Biol. Chem. 176:367-388.
Friedman, M., Krull, L.H., and Cavins, J.F. 1970. The chromatographic determination of cystine and cysteine residues in proteins as S-β-(4pyridylethyl)cysteine). J. Biol. Chem. 245:38683871.
Moore, S., Spackman, D.H., and Stein, W.H. 1958. Chromatography of amino acids on sulfonated polystyrene resins: An improved system. Anal. Chem. 30:1185-1190.
Gharahdaghi, F., Atherton, D., DeMott, M., and Mische, S.M. 1992. Amino acid analysis of PVDF bound proteins. In Techniques in Protein Chemistry III (R.H. Angeletti, ed.) pp. 249-260. Academic Press, San Diego. Hawke, D. and Yaun, P. 1987. S-Pyridylethylation of cystine residues. Appl. Biosystems User Bull. 28:1-8. Heinrikson, R.L. and Meredith, S.C. 1984. Amino acid analysis by reverse-phase high performance liquid chromatography: Precolumn derivatization with phenylisothiocyanate. Anal. Biochem. 136:65-75. Hobohm, U., Houthaeve, T., and Sanders, C. 1994. Amino acid analysis and protein database compositional search as a rapid and inexpensive method to identify proteins. Anal. Biochem. 222:202-209. Hsi, K.-L. and Yamane, D. 1994. PTC-Amino acid analysis using an automated amino acid analyzer. In Protein Analysis Renaissance, pp. 3740. PE Applied Biosystems, Foster City, Calif. Jones, B.N. and Gilligan, J.P. 1983. o-Phthaldialdehyde precolumn derivatization and reversedphase high-performance liquid chromatography of polypeptide hydrolysates and physiological fluids. J. Chromatogr. 266:471-482. Knecht, R. and Chang, J.-Y. 1986. High sensitivity amino acid analysis using DAB-Cl precolumn derivatization method. In Advanced Methods in Protein Microsequence Analysis (B. WittmannLiebold, J. Salnikow, and V.A. Erdmann, eds.) pp. 56-61. Springer-Verlag, Berlin. Koop, D.R., Morgan, E.T., Tarr, G.E., and Coon, M.J. 1982. Purification and characterization of a
Niece, R.L., Williams, K.R., Wadsworth, C.L., Elliott, J., Stone, K.L., McMurray, W.J., Fowler, A., Atherton, D., Kutny, R., and Smith, A. 1989. A synthetic peptide for evaluating protein sequences and amino acid analyzer performance in core facilities: Design and results. In Techniques in Protein Chemistry (T.E. Hugli, ed.) pp. 89101. Academic Press, San Diego. Orr, A., Ivanova, V.S., and Bonner, W.M. 1995. “Waterbug” dialysis. Biotechniques 19:204-206. Schegg, K.M., Denslow, N.D., Anderson, T.T., Bao, Y.A., Cohen, S.A., Mahrenholz, A.M., and Mann, K. 1997. Quantitation and identification of proteins by amino acid analysis. In Techniques in Protein Chemistry VIII (D.R. Marshak, ed.) in press. Academic Press, San Diego. Spencer, R.L. and Wold, F. 1969. A new convenient method for estimation of total cystine-cysteine in proteins. Anal. Biochem. 32:185-190. Stein, S. and Brink, L. 1981a. Amino acid analysis of protein bands on stained polyacrylamide gels. Methods Enzymol. 79:25-27. Stein, S. and Brink, L. 1981b. The fluorescamine amino acid analyzer. Methods Enzymol. 79:2025. Strydom, D. and Cohen, S. 1994. Comparison of amino acid analyses by phenylisothiocyanate and 6-aminoquinolyl-N-hydroxysuccinimidyl carbamate precolumn derivatization. Anal. Biochem. 222:19-28. Strydom, D.J., Tarr, G.E., Pan, Y.-C.E., and Paxton, R.J. 1992. Collaborative trial analysis of ABRF91AAA. In Techniques in Protein Chemistry III (R.H. Angeletti, ed.) pp. 261-274. Academic Press, San Diego. Chemical Analysis
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Strydom, D.J., Andersen, T.T., Apostol, I., Fox, J.W., Paxton, R.J., and Crabb, J.W. 1993. Cysteine and tryptophan amino acid analysis of ABRF92AAA. In Techniques in Protein Chemistry IV (R.H. Angeletti, ed.) pp. 279-288. Academic Press, San Diego. Tarr, G.E. 1986. Manual edman sequencing system. In Methods of Protein Microcharacterization, A Practical Handbook (J.E. Shively, ed.) pp. 155194. Humana Press, Clifton, N.J. Tarr, G.E., Paxton, R.J., Pan, Y.-C.E., Ericsson, L.H., and Crabb, J.W. 1991. Amino acid analysis 1990: The third collaborative study from the Association of Biomolecular Resource Facilities (ABRF). In Techniques in Protein Chemistry II (J.J. Villafranca, ed.) pp. 139-150. Academic Press, San Diego. West, K.A. and Crabb, J.W. 1990. Performance evaluation, automatic hydrolysis and PTC amino acid analysis. In Current Research in Protein Chemistry (J.J. Villafranca, ed.) pp. 37-48. Academic Press, San Diego. West, K.A. and Crabb, J.W. 1992. Applications of automatic PTC amino acid analysis. In Techniques in Protein Chemistry III (R.H. Angeletti, ed.) pp. 233-242. Academic Press, San Diego. Wilkins, M., Pasquali, C., Appel, A., Ou, K., Golaz, O., Sanchez, J.-C., Yan, J.X., Gooley, A.A., Hughes, G., Humphrey-Smith, I., Williams, K.L., and Hochstrasser, D.F. 1996. From proteins to proteomes: Large-scale protein identification by two-dimensional electrophoresis and amino acid analysis. BioTechnology 14:61-65.
Williams, K.R. and Stone, K.L. 1995. In-gel digestion of SDS PAGE–separated proteins: Observations from internal sequencing of 25 proteins. In Techniques in Protein Chemistry VI (J.W. Crabb, ed.) pp. 143-152. Academic Press, San Diego. Yücksel, K.Ü., Andersen, T.T., Apostol, I., Fox, J.W., Crabb, J.W., Paxton, J.L., and Strydom, D.J. 1994. Amino acid analysis of phosphopeptides: ABRF-93AAA. In Techniques in Protein Chemistry V (J.W. Crabb, ed.) pp. 231-242. Academic Press, San Diego. Yücksel, K.Ü., Andersen, T.T., Apostol, I., Fox, J.W., Paxton, J.L., and Strydom, D.J. 1995. The hydrolysis process and the quality of amino acid analysis: ABRF-94AAA collaborative trial. In Techniques in Protein Chemistry VI (J.W. Crabb, ed.) pp. 185-192. Academic Press, San Diego.
Key Reference Cohen, S.A. and Strydom, D.J. 1988. Amino acid analysis utilizing phenylisothiocyanate derivatives. Anal. Biochem. 174:1-16. Reviews PTC-AAA methods and applications, including information about other precolumn derivatization methods and analysis of physiological fluid samples.
Contributed by John W. Crabb, Karen A. West, W. Scott Dodson, and Jeffrey D. Hulmes W. Alton Jones Cell Science Center Lake Placid, New York
Amino Acid Analysis
11.9.42 Supplement 7
Current Protocols in Protein Science
N-Terminal Sequence Analysis of Proteins and Peptides
UNIT 11.10
Amino-terminal (N-terminal) sequence analysis is used to identify the order of amino acids of proteins or peptides, starting at their N-terminal end. The complexity of the chemical reactions involved (see Fig. 11.10.1) has resulted in development of sophisticated instruments that process the protein or peptide in an automated manner. In order to undergo this cyclic process, an unmodified α-amino group is required at the N-terminal end of the molecule. After modification with phenylisothiocyanate (PITC), the derivatized terminal amino acid is removed by acid cleavage as its phenylthiohydantoin (PTH) derivative and a new α-amino group on the next amino acid is now available to react with
N
C
S + H2N
H
O
C
C
O Asp Phe Arg Asp Trp
CH3
PITC
N C S
NH
H
O
C
C
Ser
C O−
basic pH
O Asp Phe Arg Asp Trp
Ser
C O−
CH3
sequencer reactions
+ TFA O
H2N
Asp Phe Arg Asp Trp
H
O
C N
C O−
+
CH3
Ser
C S
C
HN thiazolinone derivative
H+ CH3
conversion (flask) reactions
O
H
C
H
N
C C
N
S PTH amino acid
PTH amino acid analysis
HPLC
Figure 11.10.1 Edman chemistry and automated N-terminal sequence analysis. Modification of the free N-terminus of a protein or peptide sample with phenylisothiocyanate (PITC) at high pH, followed by acid cleavage of the modified terminal residue, results in release of a peptide one residue shorter in length and with a free N-terminus. This process represents a single sequencer cycle; subsequent residues can be removed by repeating this series of reactions (sequencer reactions). The cleaved 2-anilino-5-thiazolinone derivative (ATZ amino acid) is extracted from the sample support and transferred to a “flask” for conversion to the phenylthiohydantoin (PTH) amino acid (conversion reactions). The sample is then injected onto an HPLC column connected in line to the sequencer for separation analysis (PTH amino acid analysis). Contributed by David F. Reim and David W. Speicher Current Protocols in Protein Science (1997) 11.10.1-11.10.38 Copyright © 1997 by John Wiley & Sons, Inc.
Chemical Analysis
11.10.1 Supplement 8
PITC. The series of sequencer reactions shown in Figure 11.10.1 represents a sequencing cycle that results in identification of the N-terminal amino acid present on the peptide or protein at the beginning of that cycle. If each step were 100% efficient, it would be possible to sequence an entire protein in a single sequencer run. In practice, multiple factors limit the amount of sequence information that can be obtained, and only rarely is it feasible to obtain >50 residues from a single sequencer run even when the amount of available protein is not limiting. With current technology, it is fairly routine to obtain at least 20 to 40 residues of sequence from the N-terminus of proteins and large peptides in the low picomole range (<50 pmol). However, a complication when working with an intact protein is that between 50% and 80% of all proteins are estimated to have a modified (blocked) N-terminal amino group and hence cannot be sequenced directly (Brown and Roberts, 1976). Several methods are available to deblock modified N-termini (see UNIT 11.7), although these methods usually require relatively large amounts of protein and may not always be successful, especially if the nature of the blocking group is not known. Because most proteins have blocked N-termini, the preferred first step in analyzing an unknown protein is to proteolytically fragment the protein and obtain internal peptide sequences (see Strategic Planning). This multistep process involves separation of the protein of interest on either a one-dimensional (UNIT 10.1) or two-dimensional (UNIT 10.4) SDS polyacrylamide gel, possibly followed by electrotransfer to a polyvinylidene difluoride (PVDF) membrane (UNIT 10.7). This is followed by cleavage either in the gel (UNIT 11.3) or on the PVDF membrane with a specific protease such as trypsin or endoproteinase Lys-C (UNIT 11.2). Separation of the peptides is then accomplished by reversed-phase HPLC and the peptides are analyzed by MALDI mass spectrometry (UNIT 11.6). Finally, sequence analysis of one or more of the resulting peptides is carried out according to the methods in this unit (see Basic Protocols 1 and 3). Most peptides produced by these methods are <35 residues in length and their complete sequence can usually be determined in a single sequence run. Currently, the most common applications for protein sequence analysis include the following. 1. Peptide sequences may be obtained and used to identify a protein of interest by searching them against protein and translated DNA databases (UNITS 2.1 & 2.5). 2. In the increasingly rare cases where the target protein is not in available databases, the peptide sequence may be used either to design oligonucleotide probes or to confirm putative cDNA clones. 3. The sequences may be used to characterize recombinant proteins and fragments of natural and recombinant proteins used in structure-function studies. 4. The sequences may be used to characterize recombinant proteins used as pharmaceutical reagents. 5. The sequences may be used to determine the sites and nature of post-translational modifications implicated in biological function.
N-Terminal Sequence Analysis of Proteins and Peptides
This unit describes the sequence analysis of protein or peptide samples in solution (see Basic Protocol 1) or bound to PVDF membranes (see Basic Protocol 2) using a PerkinElmer Procise Sequencer. Sequence analysis of protein or peptide samples in solution (see Basic Protocol 3) or bound to PVDF membranes (see Basic Protocol 4) using a Hewlett-Packard Model G1005A sequencer is also described. (See SUPPLIERS APPENDIX for contact information regarding the manufacturers of these instruments.) Methods are provided for optimizing separation of PTH amino acid derivatives on Perkin-Elmer instruments (see Support Protocol 1) and for increasing the proportion of sample injected onto the PTH analyzer on older Perkin-Elmer instruments by installing a modified sample
11.10.2 Supplement 8
Current Protocols in Protein Science
loop (see Support Protocol 2). The amount of data obtained from a single sequencer run is substantial, and careful interpretation of this data by an experienced scientist familiar with the current operation performance of the instrument used for this analysis is critically important. A discussion of data interpretation is therefore provided (see Support Protocol 3). Finally, discussion of optimization of sequencer performance as well as possible solutions to frequently encountered problems is included (see Support Protocol 4). STRATEGIC PLANNING Instrumentation N-terminal protein/peptide sequencing instruments have improved dramatically over the past 30 years with the net effect that sequencer sensitivity has increased from sample amounts in the micromole level to the low picomole level. Modern sequencing instruments can be categorized into two groups based on the types of sequencer support that are used (Fig. 11.10.2). The most common sequencer support is a glass fiber filter (GFF). Most instruments in this group have been manufactured by Applied Biosystems (now owned by Perkin-Elmer), including the prototype Model 470A gas-phase sequencer initially produced in the early 1980s (Hewick et al., 1981). The latest models in this group are the Procise series Models 491, 492, and 494, which have one, two, or four sample cartridges, respectively. The availability of multiple sample cartridges allows automated
A
B
C
D
Figure 11.10.2 Sequencer sample cartridges. (A) Glass fiber filter (GFF) sample cartridge for a Perkin-Elmer sequencer, used to analyze samples in solution. The sample is loaded onto a Polybrene-treated GFF, inserted into the top reaction cartridge block, and the two halves of the cartridge are sealed with a Zitex (Teflon) cartridge seal. (B) Blott cartridge for a Perkin-Elmer sequencer, used to analyze proteins bound to PVDF membranes, which are inserted into the semicircular slot in the top of the Blott cartridge. (C) Biphasic sequencer reaction cartridge for a Hewlett-Packard G1005A sequencer, used to analyze samples in solution. The sample is loaded on the top half of the column (cross-hatched area) which contains a hydrophobic C18-type resin. The bottom half of the unit contains a hydrophilic resin (solid area). (D) A Hewlett-Packard sample cartridge for analyzing samples bound to PVDF membranes. The PVDF strips are inserted into an empty upper column (note absence of cross-hatched area). The bottom half of this unit is identical to the bottom half of the unit in panel C (solid area represents hydrophilic resin).
Chemical Analysis
11.10.3 Current Protocols in Protein Science
Supplement 8
tandem analysis of multiple sequence samples, which substantially increases instrument throughput and flexibility. The second type of sequencer is the Model G1005A (HewlettPackard) sequencer, which utilizes a biphasic sequencer sample support consisting of an upper column unit packed with a hydrophobic bonded phase on silica beads, and a lower hydrophilic column unit (Miller et al., 1990). This unique sample support is especially well suited to loading large-volume samples and/or samples contaminated with buffers or reagents that are incompatible with loading directly to a glass fiber filter support. This was the first commercial instrument to introduce a highly reproducible PTH analysis method using an ion-pairing reagent. Both the Perkin-Elmer and Hewlett-Packard have modified the design of the liquid sample cartridges (Fig. 11.10.2) to accommodate proteins bound to PVDF membranes (Sheer et al., 1991; Reim and Speicher, 1994). Although there are substantial differences between the two instruments in hardware design and the controlling software, differences in the overall sequencing sensitivity and sample throughput are relatively minor. Both sequencer systems include integration and coordination of multiple components required for sequence analysis: i.e., a computer, the sequencer module, and a dedicated in-line HPLC for analysis of the PTH amino acid derivatives obtained from each sequencer cycle (PTH analyzer). The computer controls the overall operation of all components and handles data storage and analysis from the PTH amino acid analyzer. Figure 11.10.3, which is a schematic of the reagents, solvents, and flow paths for a Perkin-Elmer Procise Model 494 sequencer (with four sample cartridges) highlights the complexity of the instrument and the three major stages of sequence analysis: the Edman chemistry conducted on the sample support (sequencer reactions); sample conversion to a stable PTH derivative (flask reactions); and HPLC analysis (PTH analyzer). Each of these three stages takes a similar amount of time (∼30 to 40 min), and to minimize total sample-analysis times, each of these three stages is performed in parallel. Therefore, while the first residue is being separated on the PTH analyzer, the second residue is being converted in the flask, and derivatization and cleavage of the third residue is occurring on the sample column. Sample Preparation To ensure the success of a sequencing experiment, the following considerations must be taken into account. Should internal peptides or intact protein be sequenced? When amino acid sequence information is needed to identify a protein of interest, an important decision is whether one should attempt an N-terminal sequence of the intact
N-Terminal Sequence Analysis of Proteins and Peptides
Figure 11.10.3 (at right) Perkin-Elmer Procise 494 reagent schematic, illustrating the current complexity of instrumentation used for automated sequence analysis. Bottles for chemistry involved in sequencer reactions include: base (R2) and PITC (R1); trifluoroacetic acid (R3) for cleavage, which can be delivered in either the gas phase or as a small pulse of liquid reagent; solvents for removing reaction byproducts (S1 and S2) and extraction of the cleaved amino acid derivative to the conversion flask (S3); and additional reagent positions for addition of optional chemistries or solvents (X1 and X3). The conversion reactions section includes: aqueous acid (R4) for conversion of the amino acid derivative to PTH amino acids; acetonitrile/water (S4) to redissolve the PTH amino acids for HPLC analysis; PTH standard (R5) used for calibration; and additional positions for user-designated chemistries or solvents (X2 and X3). Delivery of appropriate chemicals or solvents to the sample cartridges or conversion flask are regulated by the related reagent or solvent valve blocks. The HPLC injector transfers the PTH amino acids from the conversion flask to an HPLC column. The HPLC pumps and detector are controlled through the sequencer computer during the sequencing process in the automated mode. Adapted with permission from Perkin-Elmer Procise User’s Manual.
11.10.4 Supplement 8
Current Protocols in Protein Science
cartridge reagent block
1
2
3
4
X1 R2 gas gas
X3 liq
*
*
5 *
X1 liq
6
cartridge solvent block
7
R1 liq
8
9
*
*
10 11 12 13 14 15
R3 R3 liq gas
S2 liq
S3 liq
S4 liq
W
46 com
cartridge input block
NC
NO
16 17 18 19 20 21 22 23 argon argon high low
#6 #5 W A
B
C
sequencer reactions
W W D SAMPLE CARTRIDGES
#1 #2 #3 #4
F/C W #7
34 35 36 37 38 39 40 flask reagent block
cartridge output block
24 25 26 27 28 29 30 *
S4 X3 liq liq
*
X2 liq
R4 liq
47 com NC
R5 liq
# 11 W
NO
flask input block argon argon high low LEGEND
= output to waste = normally open port = normal ly closed port = common port = output to fraction collector = flow direction (reagents/solvents) = flow direction (argon) = fluid sensor = check valve = valve (two-way) com= valve (three-way)
45 W
NO
#
# 10
conversion (flask) reactions
X2 gas
W flask output block
W NO NC com F/C
NC
31 32 33
41 42 43 44 W 48 com flask
NC
NO
argon argon high low
injector to column 4 3 2 #9 6 1 5 #8 from pump
HPLC
Chemical Analysis
11.10.5 Current Protocols in Protein Science
Supplement 8
protein or cleave the protein into peptides and determine internal sequences. Until several years ago, the large difference in the amount of sample needed for internal sequencing (>100 pmol) as compared to N-terminal sequencing (<10 pmol) tended to sway the decision for most sequencing laboratories toward initially attempting an N-terminal sequence, despite the high probability that the N-terminus would be naturally blocked and no sequence would be obtained. As noted above, between 50% and 80% of all proteins are naturally blocked and cannot be sequenced directly. However, improvements in producing and isolating proteolytic fragments in the low picomole range (see UNITS 11.2, 11.3 & 11.6) have reduced the amount of protein needed for obtaining internal sequences to the 10- to 20-pmol range for many sequencing laboratories. As a result, if the amount of protein that can be isolated is very limited, it is now usually advisable to initially attempt internal sequencing. Because similar amounts of protein are now required for either sequencing strategy, attempting internal sequencing provides a much higher probability of successfully obtaining the needed sequence information with the minimum investment of sample, effort, and expense. Matching biological function with the right protein (the right band on a gel) The most challenging protein isolation problems are those that arise where only a very limited amount of a protein can be isolated from a natural source. In the most typical case, a biological event or process is being studied and the goal is to identify a specific protein or proteins associated with a particular biological observation. In addition to isolating sufficient quantities of what is often a relatively low-abundance protein within a cell line or tissue, it is critical to ensure that the correct protein is isolated and sequenced. Although this latter point may seem so obvious that it does not deserve mention, a substantial portion of the samples submitted to sequencing laboratories actually are contaminants rather than the desired protein. Three helpful steps to ensure that the right protein is being sequenced are: (1) make an order of magnitude estimate of the amount of protein expected, (2) consider likely contaminants, and (3) run appropriate controls to test for likely contaminants. Order of magnitude estimate: An order of magnitude estimate should be made to determine how much protein can feasibly be isolated from a given amount of source material. Even if little is known about the target protein, making a rough estimate about the protein’s likely abundance may avoid effort wasted in attempting to isolate the protein from a source that can not possibly contain enough of the desired protein. Similarly, if the protein yields during purification dramatically exceed expectations, the observed protein band is probably a contaminant rather than the target protein.
N-Terminal Sequence Analysis of Proteins and Peptides
As noted above, for those laboratories that have optimized the sensitivity of their procedures, the minimum amount of protein required for obtaining some sequence information from internal peptide sequencing is ∼10 to 20 pmol. Although not every sequencing laboratory can analyze samples at this level, a 20-pmol minimum value will be assumed in this unit; the minimum value should be adjusted accordingly depending upon the anticipated sequencing sensitivity of the laboratory that will be used for the sequence analysis. There are 6 × 1023 molecules/mol or 6 × 1011 molecules/pmol × 20 pmol desired = 1.2 × 1013 molecules of purified protein required. To obtain the number of molecules that must be present in the initial sample this number is divided by an estimated overall purification yield—e.g., 0.25 (25% yield) for a high-yield purification involving only a few efficient steps such as an antibody affinity column followed by SDS-PAGE and electroblotting (lower overall yields would be likely for more complex purifications). With a 25% overall purification yield, 4.8 × 1013 molecules of the desired protein must be in the starting tissue. If the protein is to be purified from tissue culture cells, it is simple to estimate the number of cells required if some idea of the copy number
11.10.6 Supplement 8
Current Protocols in Protein Science
Table 11.10.1
Estimating Cell Amounts Needed To Obtain Sequence Informationa
Copies/ cell
Cells needed
106
5 × 107
Only the most abundant proteins including major cytoskeletal and structural proteins
105
5 × 108
Abundant proteins usually easy to detect on 2-dimensional gels of whole-cell homogenates
104
5 × 109
Generally below the detection limit on 2-dimensional gels of whole-cell homogenates
103-102
Not practical
Examples include many enzymes and regulatory proteins
Comments
aBased on a 25% overall purification yield and analysis by a sequencing laboratory capable of obtaining internal sequence information on 20 pmol of a target protein.
Table 11.10.2
Probable Protein Contaminants When Purifying Low-Abundance Proteins
Mol. wt. (kDa)a
Probable contaminants
200
Myosin
68 50-60
Serum albumin from source tissue or serum in tissue culture media Intermediate filament proteins from source cells or human skin tissue keratins from airborne or direct-contact contamination
55 43
Antibody heavy chainb Actin (usually the most abundant cell protein)
25-28 Varied
Antibody light chainb Any added protein including enzymes such as DNase, RNase, soybean trypsin inhibitor, pancreatic trypsin inhibitor, or other protein-type protease inhibitors
aApproximate
apparent size on an SDS gel run using reducing conditions.
bDefinite major contaminant in immunoprecipitates and a likely contaminant when using an immunoaffinity
column (column bleed).
per cell can be made (Table 11.10.1). These estimates illustrate that in most cases it is only feasible to use tissue culture cells as source material for major cellular proteins (>100,000 copies/cell). Purification of low-abundance proteins (100 to 1000 copies/cell) usually requires a substantial amount of a solid tissue; more purification steps are also often needed. Contaminants and controls: The four most likely groups of protein contaminants are: (1) major cell proteins; (2) proteins used as reagents in the purification, such as antibodies; (3) proteins from culture media or serum proteins; and (4) keratins from skin. Residual contamination with a very small quantity of a major cell protein such as myosin or actin in a purified fraction can often exceed the amount of a lower-abundance target protein that has been recovered in high yield. A number of commonly observed protein contaminants are listed in Table 11.10.2. The best guard against focusing on a contaminant protein during isolation of a low-abundance protein is to analyze a parallel control sample that has been exposed to the purification conditions used, but which lacks the target protein. For example, if a cell extract is purified on a monoclonal antibody column, two useful controls can be run in parallel with the protein eluted from the specific antibody column. First, preelute the specific antibody column prior to the preparative purification with the same elution conditions as used to elute the target protein. Second, one can pass the cell
Chemical Analysis
11.10.7 Current Protocols in Protein Science
Supplement 8
extract through an unrelated monoclonal antibody column prior to the specific column and elute the nonspecific column using the same elution conditions. The controls and purified protein can then be analyzed in parallel on an SDS gel. Avoiding chemical modification of the protein It is especially critical to avoid the use of any reagent during the purification that might react with amino groups if the N-terminal amino group is not naturally blocked and sequencing of the intact protein is planned (e.g, to map a protease cleavage site or if the N-terminus is known to be unblocked). However, even if the planned strategy is to immediately pursue internal peptide sequences, it is very advisable to avoid any chemical modification of reactive groups on the protein. Modification of side-chain amino groups on lysines will interfere with trypsin or endoproteinase Lys-C digestion, and modification of any reactive groups on the protein will introduce heterogeneity that may reduce yields during protein purification steps. This microheterogeneity will reduce the yields of specific peptides and increase the complexity of HPLC peptide maps, as the different chemical forms of each peptide will probably be separated from each other at the reversed-phase HPLC step. The reagent most frequently used in protein purifications that leads to protein modification is urea (also see APPENDIX 3A). Although urea itself is uncharged and does not directly react with proteins, it decomposes rapidly to form cyanate, which reacts with amino groups. If possible, urea should be eliminated from the purification procedure. If its use cannot be eliminated, the following precautions should be taken: (1) purchase an “ultrapure” grade of urea (the dry powder is stable at room temperature; solutions are not stable); (2) prepare urea-containing solutions using ultrapure urea immediately before use and keep the temperature as low as possible when using the solution; (3) include a scavenger in the urea solution that will react with cyanate as it forms, e.g., 200 mM Tris⋅Cl or 50 mM glycine; (4) keep the pH at 7.0 or lower (if feasible), as amino groups are ∼10-fold more reactive at pH 8.0 than at pH 7.0 and cyanate is less stable at acidic pHs. For example, it should be safe to expose a protein to a 7 M urea solution containing 50 mM glycine at pH 7.0 for at least 24 hr at 4°C or for several hours at room temperature. Note that an appropriate buffer such as sodium phosphate should be included in this solution, as neither glycine nor Tris⋅Cl have much buffering capacity at pH 7. Avoiding adsorptive losses It is well known that small amounts of proteins and peptides adsorb to glass and plastic surfaces and various strategies are commonly employed to minimize such losses. Adsorptive losses are especially problematic in late stages of purifications where low pmol amounts (low µg amounts) of proteins are being isolated for sequence analysis. Addition of a blocking protein such as 0.1% BSA to solutions is not feasible, as minor contaminants and cross-linked forms of the blocking protein are almost certain to interfere with sequence analysis of the target protein even if the target protein has a very different molecular weight.
N-Terminal Sequence Analysis of Proteins and Peptides
The severity of protein adsorptive losses is often underestimated. As shown in Table 11.10.3, appreciable protein adsorption to both glass and plastic tubes occurs within seconds. The adsorbed protein is tightly bound to the container surface and can only be removed with very harsh conditions, e.g., 1% SDS. Because the values in Table 11.10.3 represent brief exposure of a dilute protein to a single tube, the actual adsorptive losses that can occur during late stages of a purification would be expected to be far higher than those shown here. The best method of minimizing such losses is to add a detergent, preferably SDS, to the protein sample in late purification steps; this is likely to be especially important for any step where the total protein concentration of the sample is
11.10.8 Supplement 8
Current Protocols in Protein Science
Table 11.10.3
Adsorptive Losses of Proteins from Dilute Solutionsa
Protein
Phosphorylase B BSA Ovalbumin Myoglobin
Glass test tube
Siliconized glass test tube
Polypropylene microcentrifuge tube
µg
pmol
µg
pmol
µg
pmol
0.4 1.2 0.9 1.9
4.3 18 21 112
0.7 2.0 1.3 2.5
7.5 30 30 147
0.6 0.2 0.3 1.0
6.4 3.0 6.9 58
aAmount of protein lost by adsorption to the indicated container surface within 15 sec of addition of a 300-µl aliquot of dilute protein solutions in phosphate buffer, pH 7. Samples were pipetted directly into the bottoms of the tubes to minimize exposure to the container surfaces. Losses were determined by quantitative amino acid analysis of the protein left in solution. Protein adsorbed to the container surface could be extracted using 1% SDS. Adsorptive losses were higher than the values shown here when the solution was vortexed or when protein solutions were applied to the top of the tube and allowed to run down the side.
<0.1 µg/µl. As discussed below, the final purification step should be an SDS gel, therefore the addition of SDS to the sample during late steps of most purifications should be feasible. Of course, biological activity will be irreversibly lost in most cases upon addition of SDS, and if maintaining biological activity is critical for monitoring purification steps, an alternative blocking reagent such as a nonionic detergent (e.g., Triton X-100 or Tween 20) should be considered. Final purification step SDS-PAGE gels are the preferred final purification step for most protein sequencing applications. SDS gels are ideally suited for handling low microgram to submicrogram amounts (low picomole levels) of protein. They have high resolving power, and most buffer contaminants that would interfere with direct sequencing or with a protease digestion are separated from the target protein in the gel. Protein losses resulting from adsorption are negligible in comparison to those incurred with virtually any other method, and SDS gels are compatible with addition of SDS or another detergent to samples in preceding purification steps to minimize adsorptive losses (see above). After SDS-PAGE, the protein can be electroblotted to a high-retention PVDF membrane (UNIT 10.7) for direct N-terminal sequencing (see Basic Protocols 2 and 4) or else the protein can be digested on the membrane with trypsin, endoproteinase Lys-C, or another protease to obtain internal peptides (UNIT 11.2) for sequence analysis. Alternatively, the protein can be digested with a protease directly in the gel (UNIT 11.3). Another advantage of isolating proteins in this manner is that the protein in the gel or on the PVDF membrane is in a denatured form that facilitates proteolytic digestion. Because SDS gels are usually used as the final purification step for sequence analysis, the protein of interest does not need to be purified to homogeneity by other methods. In some cases, quite crude extracts have been applied to a one-dimensional gel and the protein of interest identified. The limitations to this approach are that: (1) the migration position of the protein of interest on the gel must be known, (2) the target protein must be resolved from other proteins in the sample by the gel system used, and (3) the target protein must be present in the sample in a sufficiently high proportion so that contaminating proteins do not cause a gel-overload condition when sufficient target protein for sequence analysis is applied to the gel. Complex samples that may contain multiple proteins at the target-protein migration position on a one-dimensional gel can usually be adequately separated by two-dimensional gel electrophoresis (UNIT 10.4), and the target protein from one or several replicate two-dimensional gels (or electroblots from the
Chemical Analysis
11.10.9 Current Protocols in Protein Science
Supplement 8
two-dimensional gels) can be combined and digested with trypsin. Frequently, high-abundance proteins can be isolated for internal sequencing by loading a whole-cell homogenate directly to a series of replicate two-dimensional gels. However, despite the very high resolving power of two-dimensional gels, a single spot on a two-dimensional gel may contain multiple proteins, especially when very complex samples such as whole-cell extracts are used. A useful guideline is to purify proteins larger than ∼6 kDa using SDS-PAGE as the last purification step; peptides smaller than 6 kDa from either in situ protease digests or from other sources should be purified using reversed-phase HPLC (UNIT 11.6). If <100 pmol of peptide is being isolated, use of a 1.0- or 2.1-mm-diameter C18 column with acetonitrile and 0.1% TFA as the reversed-phase solvents is recommended. Amount of protein used for sequence analysis and observed sequencer yields Occasionally, enthusiastic sequencer manufacturers as well as research scientists claim that 1 pmol or less of a protein can be sequenced. A critical point that is often not emphasized is the cited sensitivity usually refers to the observed sequence yield in early sequencer cycles, not the total amount of protein or peptide actually used in the experiment. The average initial sequence yield for proteins or peptides (i.e., the signal observed in early sequencer cycles divided by the amount loaded into the sequencer multiplied by 100) averages ∼50% (with the typical range 20% to 80%). Because adsorptive losses and artifactual modification of the N-terminus are likely to increase as the quantity of sample decreases, where 1 pmol of a protein or peptide is loaded into a sequencer, a realistic initial yield would be 0.1 to 0.5 pmol. Only a few laboratories that have carefully optimized all aspects of the sequence analysis procedure can obtain significant N-terminal sequence information at this level. Obtaining internal sequences involves multiple steps that are individually reasonably efficient but that have the effect of further reducing the observed sequence yield relative to the starting protein. A reasonable guideline is that initial sequencing yields of internal peptides relative to the amount of a protein loaded onto a gel is ∼15% (with a typical range of 5% to 30%). This means that 20 pmol of a target protein applied to a gel will yield internal sequences with initial yields between 1 and 6 pmol. Although these overall yields may appear to be quite poor, the yields at each step of the procedure must be quite good to achieve such overall yields. An example of yields at each step that might be obtained in a laboratory that has carefully optimized the procedure might be as follows: • recovered in the major gel band relative to the amount applied to the top of the
gel—90% • electroblotting efficiency—70% • trypsin digestion efficiency and recovery of cleaved peptides from either gel or
PVDF membrane—70% • recovery from the HPLC—90% • recovery from the collection tube—80% • initial yield in the sequencer—50%.
The cumulative yield in this example would be: 0.9 × 0.7 × 0.7 × 0.9 × 0.8 × 0.5 = 0.16, or 16%.
N-Terminal Sequence Analysis of Proteins and Peptides
Estimating amount of protein used for sequence analysis Estimating the amount of target protein that has been isolated and that will be used for either obtaining a N-terminal sequence or internal sequences contributes to proper interpretation of results. For example, if a protein band that has been electroblotted to a PVDF membrane does not produce a detectable sequence, knowing the amount loaded
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into the sequencer should indicate whether the protein was blocked or insufficient protein was present. Alternatively, a low-level sequence may arise from a minor contaminant rather than the target protein, which might be blocked. For example, assuming that a PVDF-bound band yielded a 2-pmol sequence, if ∼4 pmol were present on the membrane, this sequence would clearly correspond to the major component in that band, but if 40 pmol were present it would be considered likely that the major component was blocked and that a minor contaminant in the gel band produced the minor sequence observed. Ideally, accurate quantitation of a portion of the protein or peptide sample should be made prior to either N-terminal sequencing or protease digestion. Unfortunately, when only low picomole amounts of protein are available, a substantial portion of the total sample is required simply to obtain a reasonable estimate of protein concentration by amino acid analysis (AAA), which is the most reliable method of quantitating protein concentrations in solution, in gels, or on PVDF membranes (see UNITS 3.2, 11.3 & 11.9 for AAA and related techniques). Therefore, if an unacceptable proportion of the sample would be consumed by quantitative AAA, the amount of protein on the gel or PVDF membrane can be estimated by comparing its staining intensity to the staining intensity of a series of standard protein lanes prepared using 2-fold dilutions—e.g., adjacent lanes containing 2 µg/band, 1 µg/band, 0.5 µg/band, 0.25 µg/band, and 0.125 µg/band. If the protein standards are run on the same gel as the experimental sample, one can readily estimate the amount of target protein available for sequencing to within a factor of ∼2 to 4, with the important limitation that there is some variability in staining response for different proteins. However, Coomassie blue, the preferred stain for in-gel digestions (UNIT 11.3), and Amido black, the preferred stain for on-PVDF digestions (UNIT 11.2) show much less protein-specific variability in staining than silver staining. MALDI mass spectrometry If access to a mass spectrometer is available, it is highly advantageous to prescreen all peptides prior to sequence analysis using MALDI mass spectrometry (UNITS 11.6 & 16.2) to complement peptide sequence analysis. Mass analysis of ∼10 to 15 peaks from HPLC separation of an in situ trypsin digest is recommended. The longest peptides (i.e., with the largest masses) should be selected for sequencing, as longer sequences provide more informative database-search results, especially when related proteins, but not the identical protein, are in the sequence database searched. Similarly, if the protein is not known, longer sequences provide a wider range of options for oligonucleotide probe design. After the sequence run is complete, comparison of the assigned sequence with the previously determined mass provides confirmation of the residue assignments and can extend interpretation of the sequencer data if most, but not all, of the residues in the peptide are clearly assigned. For example, reconciliation of the mass with the sequence can confirm one or two tentative assignments, suggest possibilities for an unassigned residue, or indicate the presence of a post-translational modification. In theory, prescreening HPLC peptides by MALDI mass analysis can also potentially identify peptide mixtures. In practice, only some mixtures are detected by this method, because mass signals obtained by this method are not quantitative and some peptides can suppress the signals of other peptides.
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BASIC PROTOCOL 1
SEQUENCING LIQUID SAMPLES ON GLASS FIBER FILTERS This protocol describes application of a highly purified protein or peptide in solution to a Perkin-Elmer Procise model 494 sequencer. Sample preparation and loading procedures would be similar for other sequencers that use glass fiber filter (GFF) supports. The sample should ideally be in a small volume of high-purity water or a volatile buffer/solvent. Materials Liquid peptide or protein sample(s) to be sequenced Volatile solvents for reversed-phase chromatography: e.g., 0.1% trifluoroacetic acid (TFA) and acetonitrile (UNIT 11.6) Methanol Argon or nitrogen gas source Polybrene solution (Biobrene Plus from ABI Biotechnology; store in original container up to 3 months at 4°C; discard if contamination is suspected; alternatively store 30- to 100-µl aliquots in clean microcentrifuge tubes up to 1 year at −20°C) Sequencer solvent/reagent kit (ABI Biotechnology) including: R1 (phenylisothiocyanate) R2B (n-methylpiperidine) R3 (trifluoroacetic acid) R4A (25% trifluoroacetic acid) R5 (PTH sequencing standard) S2B (ethyl acetate) S3 (n-butyl chloride) S4B (20% acetonitrile) Premix PTH analyzer kit (ABI Biotechnology) including: HPLC Solvent A3 (3.5% tetrahydrofuran) HPLC Solvent B2 (isopropanol/acetonitrile) Premix Buffer Concentrate DMPTU 1.0- or 2.1-mm-i.d. reversed-phase column (UNIT 11.6) TFA-treated glass fiber filters (GFF; ABI Biotechnology) Cartridge seals (ABI Biotechnology) Perkin-Elmer Procise sequencing system Model 494 (ABI Biotechnology) including: Sequencer module with glass sample cartridge blocks On-line PTH analyzer (dedicated HPLC and detector) Computer controller Powder-free gloves Stainless steel or Teflon-coated forceps Additional reagents and equipment for reversed-phase purification of peptides (UNIT 11.6), concentration of proteins and microdialysis (UNIT 4.4), quantitation of proteins/peptides by amino acid analysis (UNITS 3.2 & 11.9), and spectrophotometric quantitation of protein (UNIT 3.4) NOTE: Prepare solutions with Milli-Q water or equivalent taken directly from the purification unit. Water stored for any length of time in glass or plastic containers supports microbial and algal growth, which results in high amino acid background in early sequencer cycles.
N-Terminal Sequence Analysis of Proteins and Peptides
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1. Prepare peptide or protein samples in a small volume (preferably <50 µl) of high-purity water or volatile buffer/solvent. Small amounts (<10 ìmol) of nonvolatile, non-amine reactive salts such as NaCl can usually be tolerated, but at higher levels, salts may interfere with binding of the protein or peptide to the GFF and can interfere with retention times of His and Arg in early cycles. Nonvolatile buffers can interfere with sequencer performance by altering the actual pHs achieved during the coupling (basic pH) and cleavage steps (acid pH). Any nonvolatile components or contaminants such as Tris buffer that react with amine-specific reagents are especially problematic. Larger volumes can be reduced in a Speedvac evaporator, but this will concentrate nonvolatile impurities and it is likely to increase adsorptive losses on the tube holding the sample.
Purify and/or concentrate sample 2a. If analyzing a peptide sample: Purify the peptide on a 1- or 2.1-mm-i.d. reversedphase column using volatile solvents such as 0.1% TFA and acetonitrile (UNIT 11.6). 2b. If analyzing a protein sample: Concentrate the sample to ∼100 µl, then desalt the sample by exhaustive microdialysis (UNIT 3.4) against water taken directly from a Milli-Q purification unit or equivalent. Estimate amount of protein or peptide sample 3a. If >100 pmol of sample is available: Use 20% of the protein or peptide sample for quantitative amino acid analysis using a high-sensitivity method such as precolumn phenylthiocarbamyl (PTC) derivatization (UNIT 11.9). This will accurately define the amount to be loaded and detect potential contaminants in the sample that can react with amine reactive reagents.
3b. If <100 pmol of sample is available: Estimate the quantity of sample to be loaded by an indirect method such as absorbance at 280 nm (for proteins) or 215 nm (for peptides) as described in UNIT 3.4. Precycle glass fiber filter (GFF) 4. Thoroughly rinse the glass sample cartridge blocks with methanol and dry with a stream of argon or nitrogen while wearing powder-free gloves that have been prerinsed with Milli-Q water. Refer to manufacturer’s instructions for the Procise system for this and all subsequent steps. Do not touch with bare hands any surfaces that are part of the sequencer chemistry flow path or that will contact the samples. These include, e.g., the glass sample cartridge, the GFF, solvent pickup tubes, and bottle seals. Also avoid exposing these surfaces to airborne contamination.
5. Place a TFA-treated GFF on the top half of the cartridge block with a pair of stainless steel or Teflon-coated forceps. Saturate filter by applying 15 µl (for 9-mm filters) or 30 µl (for 12-mm filters) of Polybrene solution. 6. Dry Polybrene solution completely using a gentle argon or nitrogen stream, or air dry GFF under a piece of aluminum foil to prevent dust from settling on it. 7. Assemble the two cartridge halves using a new cartridge seal and insert the assembled unit into the sequencer. Pressure test the cartridge to detect leaks. 8. Run at least two cycles using the appropriate filter-conditioning program. Samples can be loaded immediately after completing the conditioning cycles or the GFF can be stored in a gas-tight bottle up to 3 days. However, the increased handling involved
Chemical Analysis
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in removing the filter from the sample cartridge and reinserting it after storage can result in an increased background in early sequencer cycles, especially for Ser and Gly. The filter-conditioning program can also include a flask program that injects the PTH standard so that the quality of the PTH amino acids can be evaluated. Alternatively, a flask blank can be injected on the HPLC during filter-conditioning cycles. After the conditioning cycles have been completed and prior to loading the sample, one complete sequencing cycle can be run to evaluate any background that is contributed by the Edman sequencing chemistry.
Apply sample and perform sequence analysis 9. Remove the sample cartridge from the sequencer (wearing powder-free gloves) and apply 15 to 30 µl (depending upon filter diameter) of the sample to the GFF in the upper half of the sample cartridge using a pipettor. Dry the filter using a gentle stream of argon or nitrogen, then repeat application and drying until the entire sample has been applied. Alternatively, air dry the sample between repetitive applications using an aluminum foil “tent” to prevent airborne contaminants from settling on the GFF. Do not overload the filter with more liquid than it can hold. Most sample volumes are larger than the filter capacity, hence the need for repetitive applications. The number of applications will depend on the total volume of the sample being loaded (volumes >100 ìl are quite time-consuming to load and increase the risk of contamination). If HPLC-purified peptide samples containing <20 pmol are used for sequence analysis, addition of trifluoracetic acid (25% final concentration) to the sample may increase the amount of sample actually transferred into the sequencer (Erdjument-Bromage et al., 1993). Estimate the sample volume by comparing the liquid height to an identical tube calibrated by marking volumes bracketing the expected volume on the tube. Add a volume of neat TFA (reagent R3 from ABI Biotechnology) equal to one-third the sample volume (25% TFA final concentration). It is very important not to vortex the sample! Use a pipettor to mix the TFA/peptide solution completely. After loading the entire sample, wash the sides of the empty sample tube with 20 ìl neat TFA and apply to the sample filter.
10. After the sample has been completely loaded and dried, place a new cartridge seal on the bottom glass block, reassemble the sample cartridge, and insert the assembly into the sequencer. Pressure test the cartridge assembly to detect leaks. 11. Check the levels of all needed items—e.g., sequencing reagents, PTH analyzer solvents, argon, and printer paper—to ensure that sufficient quantities are available to complete the projected number of sequence cycles. Program the computer appropriately using manufacturer’s recommendations and carry out the sequence run.
N-Terminal Sequence Analysis of Proteins and Peptides
At least one PTH standard or a PTH analyzer blank run followed by a PTH standard should be injected immediately prior to each sequence run to verify retention times and to determine calibration factors for each amino acid. The number of Edman cycles that should be run depends upon the characteristics of the sample and the purpose of the analysis. For most applications, it is recommended that peptides be sequenced through their C-terminus plus two cycles (to ensure that the entire sequence has been obtained and to assess background levels of amino acids; see Support Protocol 3). If the peptide’s mass has been determined prior to the sequence run, the number of cycles to run can be estimated by: cycles = (peptide mass/100) + 3 (rounded up to next integer). For example, if the observed mass is 1644.5 Da, the sequencer can be set to complete 20 sample cycles. When intact proteins or large peptides are being analyzed and a maximum amount of sequence data is desired, set a number of cycles greater than the number that will be completed in an overnight run; the next morning review the data and continue the sequence until the decreasing signal-to-noise level limits sequence assignments (see Support Protocol 3). If the protein sequence is known and the purpose is to verify an intact N-terminal sequence or to define a cleavage site, 5 to 6 cycles should be sufficient. If the protein is to be searched
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against sequence databases to identify the protein, analysis of at least 20 cycles is recommended. Procise sequencers can be programmed to automatically analyze up to four samples in tandem. Therefore, care must be taken to ensure that sufficient reagents/solvents are available for the total cycles programmed and that the appropriate sequencing programs have been selected for the sample(s) to be analyzed.
SEQUENCING PVDF-BOUND SAMPLES USING A BLOTT CARTRIDGE For most applications where N-terminal sequence analysis of intact proteins or large peptides (>6 kDa) is desired, the preferred final purification method is to use one-dimensional (UNIT 10.1) or two-dimensional (UNIT 10.4) gels followed by electroblotting to a high-retention PVDF membrane (UNIT 10.7) and staining with Amido black (UNIT 10.8). For more detailed discussion of these preliminary steps, see Strategic Planning. PVDF membranes have a high protein-binding capacity, bind large peptides and proteins with high affinity, and are inert to the harsh chemical environment in a protein sequencer. However, the wetability characteristics of these hydrophobic membranes are quite different from those of the hydrophilic glass filters (see Basic Protocol 1). These differences have necessitated development of specific sequencer programs and sample holders for PVDF membranes (see Fig. 11.10.2). The preferred method for loading proteins onto a PVDF membrane is by electroblotting from a gel. Dilute samples can be concentrated and/or separated from an incompatible buffer by direct adsorption to PVDF membranes. If detergents are not present in the protein solution, the protein can be adsorbed by immersing a small piece of PVDF membrane (which must be prewetted in 100% methanol and rinsed with water) into the sample solution and incubating it with agitation in a cold room for several hours to overnight. Alternatively, the sample can be loaded onto a PVDF disc in the bottom of a ProSorb device (ABI Biotechnology). Directly spotting a protein sample onto a prewetted PVDF membrane is not recommended as this is an unreliable loading method. The total liquid volume that can be applied to the PVDF membrane is quite small, and adsorption to the PVDF membrane competes with sample drying. After the liquid has dried, further adsorption does not occur. When the sample is rewet, either prior to loading in the sequencer or during sequence analysis, the dried protein that did not bind is usually washed from the membrane, which results in reduced sequencing yields.
BASIC PROTOCOL 2
Materials PVDF-bound sample(s) to be sequenced Methanol in wash bottle Argon or nitrogen gas source Glass gel plate Stainless steel scalpel and stainless steel or Teflon-coated forceps Bath sonicator Blott cartridge for Perkin-Elmer Procise sequencer (ABI Biotechnology) Additional reagents and equipment for sequencing on glass fiber filters (see Basic Protocol 1) NOTE: Prepare solutions with Milli-Q water or equivalent taken directly from the purification unit. Water stored for any length of time in glass or plastic containers supports microbial and algal growth, which results in high amino acid background in early sequencer cycles. Chemical Analysis
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Prepare sample 1. Using a scalpel, excise the desired stained protein band from PVDF membrane, keeping the membrane on a clean dry surface such as a glass gel plate and handling the membrane with a forceps. The glass plate, scalpel, and forceps should be thoroughly cleaned before use with Milli-Q water and methanol. Wear powder-free gloves rinsed in Milli-Q water. Do not touch the membrane or surfaces that contact the membrane with bare hands. Dry PVDF membranes are electrostatic and can be difficult to handle. The electrostatic nature of the membranes also attracts airborne contamination that invariably contains amino acid and protein contaminants. Special caution must be exercised to minimize contamination of the sample when working with small amounts of protein (<20 pmol). Two to three replicate 3-mm-wide one-dimensional gel bands on PVDF (50 to 75 mm2 in area) or an equivalent area of membrane containing replicate two-dimensional gel spots can typically be loaded into a Procise cartridge. The larger Blott cartridge on older Perkin-Elmer sequencers can accommodate about twice as much membrane surface area.
2. While holding the membrane(s) with stainless steel or Teflon-coated forceps, thoroughly rinse with a stream of methanol from a wash bottle. Immediately rinse the membranes with a stream of Milli-Q water delivered directly from the Milli-Q purification unit. PVDF membranes cannot be solvated directly with aqueous solutions and must initially be wetted in a strong organic solvent such as methanol. After the membranes are wetted with methanol in the above step, do not allow them to dry out until drying is indicated in step 3.
3. Place the membrane(s) in a clean beaker containing 5 ml Milli-Q water and sonicate 5 min in a bath sonicator. Remove the membrane(s) with forceps, rinse with methanol, and dry under a gentle stream of argon or nitrogen. Load sample and perform sequence analysis 4. Insert the membrane(s) in the slot in the top half of the Blott cartridge. Refer to the manufacturer’s instructions for this and subsequent steps. If needed, an additional piece of membrane can be placed protein-side-up within the recess on the top half of the cartridge.
5. Reassemble the Blott cartridge using a new cartridge seal and insert the assembly into the sequencer. Pressure test the cartridge assembly to detect leaks. 6. Check the levels of all needed items—e.g., sequencing reagents, HPLC solvents, argon, and printer paper to ensure that sufficient quantities are available to complete the projected number of sequence cycles. Program the computer appropriately using manufacturer’s recommendations and carry out the sequence run.
N-Terminal Sequence Analysis of Proteins and Peptides
At least one PTH standard or a PTH analyzer blank run followed by a PTH standard should be injected immediately prior to each sequence run to verify retention times and to determine calibration factors for each amino acid. The number of Edman cycles that should be run depends upon the purpose of the analysis. When intact proteins are sequenced and a maximum amount of sequence data is desired, set a number of cycles greater than the number that will be completed in an overnight run; the next morning review the data and continue the sequence until the decreasing signal-to-noise level limits sequence assignments (see Support Protocol 3). If the protein sequence is known and the purpose is to verify an intact N-terminal sequence or to define a cleavage site, 5 to 6 cycles should be sufficient. If the protein is to be searched against sequence databases to identify the protein, analysis of at least 20 cycles is recommended.
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SEQUENCING LIQUID SAMPLES ON A BIPHASIC CARTRIDGE SEQUENCER
BASIC PROTOCOL 3
The unique sample cartridge in the Hewlett-Packard sequencer (Fig. 11.10.2) results in quite different sample loading constraints than those encountered with sequencers utilizing a glass fiber filter (see Basic Protocol 1). Since samples are loaded onto the hydrophobic half of the cartridge unit, sample loading constraints are similar to those involved in applying a sample to a C18 reversed-phase column (also see UNIT 11.6). Large volumes of aqueous samples can be readily loaded. Most buffers, as well as high levels of nonvolatile salts that would severely interfere with sequence analysis on a glass filter, are easily accommodated because any hydrophilic buffers including those containing Tris and glycine can be readily washed out of the column. Because the binding of proteins or peptides to the sample cartridge is via hydrophobic interactions, only strong organic solvents or detergents are likely to interfere with sample binding. Samples in high concentrations of organic solvents can usually be successfully loaded by simply diluting the sample with 2% TFA or Milli-Q water to reduce the solvent strength, unless the peptide does not remain soluble in the more hydrophilic solvent. Samples in detergents may bind to the column either directly or after the sample is diluted; however, efficient sample loading should not be assumed in this case and the unbound fraction should be analyzed by an appropriate method such as quantitative amino acid analysis (UNIT 11.9), if feasible, or using SDS gels to detect unbound proteins and HPLC to detect unbound peptides. Materials Methanol Source of argon or nitrogen Liquid peptide or protein sample(s) to be sequenced Sequencer and PTH analyzer solvent/reagent kit (Hewlett-Packard) including: R1 (phenylisothiocyanate) R2 (diisopropylethylamine) R2A (octylamine) R3 (trifluoroacetic acid) R4 (25% trifluoroacetic acid) S2A (ethyl acetate) S2/3 (23% acetonitrile/77% toluene) S3 (15% acetonitrile/85% toluene) S3A (aqueous trifluoroacetic acid solution) S4 (10% acetonitrile) Std (PTH sequencing standard) L1 (acetonitrile) L2 (methanol) HPLC solvent A (aqueous triethylamine/acetonitrile) HPLC solvent B (aqueous triethylamine/propanol) HPLC solvent C (acetonitrile) Sample cartridges for liquid samples (Hewlett-Packard) Sample loading funnels (Hewlett-Packard) Hewlett-Packard Model G1005A sequencing system including: Sequencer module On-line PTH analyzer (dedicated HPLC and detector) Computer controller Sample loading station Powder-free gloves Chemical Analysis
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NOTE: Refer to the manufacturer’s instructions for the Hewlett-Packard Model G1005A sequencing system at all steps. NOTE: Prepare solutions with Milli-Q water or equivalent taken directly from the purification unit. Water stored for any length of time in glass or plastic containers supports microbial and algal growth, which results in high amino acid background in early sequencer cycles. 1. Precondition the sample cartridge in the sequencer using the version 2.0 column preparation program for one cycle. Sample cartridges can be preconditioned and stored in a dry, closed container ≤1 week. Airborne contamination and sample handling contamination are less of a concern with this sample unit than when working with a glass fiber filter (see Basic Protocol 1) because of its design (i.e., it does not expose a large sample surface area). Mark the cartridges with the date immediately after preconditioning using a permanent laboratory marker. Newer versions of the Hewlett-Packard column preparation program are ∼40 min in length but do not exhibit apparent advantages over the shorter (∼20-min) version 2.0 program. The shorter column preparation program is therefore recommended.
2. Prepare the sample loading funnel prior to inserting it in the sample loading station by rinsing extensively with ∼30 ml Milli-Q water and then rinsing a large number of times with methanol (∼30 ml total). Fill the funnel with one volume of sample loading solution and allow to completely drain through the funnel. Dry the funnel with a gentle stream of argon or nitrogen. Use powder-free gloves that have been rinsed with Milli-Q water when cleaning the loading funnel and applying samples to the sample cartridges. In all instances, avoid contacting the inside of the sample loading funnel and the ends of the sample cartridges with bare fingers.
3. Attach the top half of a conditioned sample cartridge to a washed sample loading funnel and insert into the sample loading station. Add 1 ml methanol to the loading funnel and apply nitrogen pressure to force the methanol through the sample column. Release the pressure when ∼5 to 10 µl remains in the funnel. Add 1 ml of sample loading solution to the funnel, then force the liquid through the sample column until ∼5 to 10 µl remains in the funnel. 4. Estimate the volume of the sample to be analyzed. Add an appropriate volume of sample loading solution to the funnel, calculated as follows: 1.0 ml − sample volume = sample loading solution volume. If HPLC-purified peptide samples containing <20 pmol are used for sequence analysis, addition of TFA (25% final concentration) to the sample may increase the amount of sample actually transferred into the sequencer (Erdjument-Bromage et al., 1993). Estimate the sample volume and add neat TFA (reagent R3 from ABI Biotechnology) equal to 25% TFA final concentration. It is very important not to vortex the sample! Use a pipettor to mix the TFA/peptide solution completely.
5. Load the sample by pipetting it below the surface of the sample loading solution in the loading funnel. Wash the sides of the empty sample tube with 20 µl neat TFA and add to the solution in the sample funnel. Rinse the pipet tip by pipetting the solution in the funnel into and out of the tip several times.
N-Terminal Sequence Analysis of Proteins and Peptides
6. Load the sample onto the sample column using nitrogen pressure until all liquid has passed through the column. Allow nitrogen to pass through the column for several minutes to ensure complete drying of the column.
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7. Remove the top half of the sample cartridge from the loading station and reattach it to the bottom half of the sample cartridge. Insert the cartridge assembly into the sequencer. Check the levels of all needed items—e.g., sequencing reagents, HPLC solvents, argon, and printer paper—to ensure that sufficient quantities are available to complete the desired number of Edman cycles. Carefully select the appropriate sequencing programs for the samples to be analyzed and carry out the sequence run. When multiple samples are to be analyzed, verify that the sequencer recognizes the presence of each cartridge before starting the first analysis. This is accomplished after all sample cartridges have been loaded and inserted onto the sequencer by manually selecting each sample cartridge position in turn from the protein sequencer menu. At least one PTH standard or a PTH analyzer blank run followed by a PTH standard should be injected immediately prior to each sequence run to verify retention times and to determine calibration factors for each amino acid. The number of Edman cycles that should be run depends upon the characteristics of the sample and the purpose of the analysis. For most applications, it is recommended that peptides be sequenced through their C-terminus plus two cycles (to ensure that the entire sequence has been obtained and to assess background levels of amino acids (see Support Protocol 3). If the peptide’s mass has been determined prior to the sequence run, the number of cycles to run can be estimated by: cycles = (peptide mass/100) + 3 (rounded up to next integer). For example, if the observed mass is 1644.5 Da, the sequencer can be set to complete 20 sample cycles. When intact proteins or large peptides are being analyzed and a maximum amount of sequence data is desired, set a number of cycles greater than the number that will be completed in an overnight run; the next morning review the data and continue the sequence until the decreasing signal-to-noise level limits sequence assignments (see Support Protocol 3). If the protein sequence is known and the purpose is to verify an intact N-terminal sequence or to define a cleavage site, 5 to 6 cycles should be sufficient. If the protein is to be searched against sequence databases to identify the protein, analysis of at least 20 cycles is recommended. The Hewlett-Packard G1005A sequencer can be programmed to automatically analyze up to four samples in tandem. Therefore, care must be taken to ensure that sufficient reagents/solvents are available for the total cycles programmed and that the appropriate sequencing programs have been selected for the sample(s) to be analyzed.
SEQUENCING PVDF-BOUND SAMPLES ON A BIPHASIC CARTRIDGE SEQUENCER
BASIC PROTOCOL 4
PVDF-bound samples are prepared in a similar fashion regardless of the sequencer to be used (see Basic Protocol 2). PVDF samples sequenced in the biphasic cartridge sequencer are loaded to a reaction cartridge that has an empty top half rather than a top half packed with C18 resin like that used with liquid samples (see Fig. 11.10.2). Materials PVDF-bound sample(s) to be sequenced Methanol in wash bottle Source of argon or nitrogen Reaction cartridge(s) for PVDF samples (Hewlett-Packard) Glass gel plate Stainless steel scalpel and stainless steel or Teflon-coated forceps Bath sonicator Additional reagents and equipment for sequencing on a biphasic cartridge sequencer (see Basic Protocol 3) NOTE: Refer to the manufacturer’s instructions for the Hewlett-Packard Model G1005A sequencing system at all steps.
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NOTE: Prepare solutions with Milli-Q water or equivalent taken directly from the purification unit. Water stored for any length of time in glass or plastic containers supports microbial and algal growth, which results in high amino acid background in early sequencer cycles. 1. Precondition the reaction cartridge (see Basic Protocol 3, step 1). 2. Using a scalpel, excise the desired stained protein band from a PVDF membrane, keeping the membrane on a clean, dry surface such as a glass gel plate and handling the membrane with a forceps. The glass plate, scalpel, and forceps should be thoroughly cleaned before use with Milli-Q water and methanol. Wear powder-free gloves rinsed in Milli-Q water. Do not touch the membrane or surfaces that contact the membrane with bare hands. Dry PVDF membranes are electrostatic and can be difficult to handle. The electrostatic nature of the membranes also attracts airborne contamination that invariably contains amino acid and protein contaminants. Special caution most be exercised to minimize contamination of the sample when working with small amounts of protein (<20 pmol). A total membrane area of up to 60 mm2 can be loaded in the top half of the reaction cartridge.
3. While holding the membrane(s) with a stainless steel or Teflon-coated forceps, thoroughly rinse with a stream of methanol from a wash bottle. Immediately rinse the membranes with a stream of Milli-Q water delivered directly from the Milli-Q purification system. PVDF membranes can not be solvated directly with aqueous solutions and must initially be wetted in a strong organic solvent such as methanol. After the membranes are wetted with methanol in the above step, do not allow the membranes to dry out until drying is indicated in step 4.
4. Place the membrane(s) in a clean beaker containing 5 ml Milli-Q water and sonicate 5 min in a bath sonicator. Remove the membrane(s) with forceps, rinse with methanol, and dry under a gentle stream of argon or nitrogen. 5. Insert the membrane(s) in the top half of the sample cartridge. Because the opening is small (1-mm i.d.), membrane(s) can be cut into strips along their long axis and loaded as a stack or singly. An alternative to cutting strips is to fold the membrane along its longest axis with the protein side facing out. Load carefully so as not to damage the inside of the reaction cartridge chamber or the column-column joint within the sample cartridge assembly.
6. Reassemble the sample cartridge and insert the cartridge assembly into the sequencer. When multiple samples are to be analyzed, verify that the sequencer recognizes the presence of each cartridge before starting the first analysis (see Basic Protocol 3, step 7).
7. Check the levels of all needed items—e.g., sequencing reagents, HPLC solvents, argon, and printer paper—to ensure that sufficient quantities are available to complete the projected number of sequence cycles. Carefully select the appropriate sequencing programs for the samples to be analyzed and carry out the sequence run.
N-Terminal Sequence Analysis of Proteins and Peptides
At least one PTH standard or a PTH analyzer blank run followed by a PTH standard should be injected immediately prior to each sequence run to verify retention times and to determine calibration factors for each amino acid. The number of Edman cycles that should be run depends upon the purpose of the analysis. When intact proteins are sequenced and a maximum amount of sequence data is desired, set a number of cycles greater than the number that will be completed in an overnight run; the next morning review the data and continue the sequence until the decreasing signal-to-noise level limits sequence assign-
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ments (see Support Protocol 3). If the protein sequence is known and the purpose is to verify an intact N-terminal sequence or to define a cleavage site, 5 to 6 cycles should be sufficient. If the protein is to be searched against sequence databases to identify the protein, analysis of at least 20 cycles is recommended.
OPTIMIZING SEPARATION OF PTH AMINO ACIDS Successful N-terminal sequencing is dependent on both the N-terminal sequencer and the in-line HPLC system that separates the PTH amino acids from each sequencer cycle. Both systems must be operating optimally to obtain a maximum amount of sequencing information, especially when analyzing samples in the low picomole range. The HPLC system should provide complete separation of all the commonly occurring PTH-derivative amino acids as well as sequencer reagent–related peaks. Also, retention times of all peaks should be highly consistent over the course of a sequence run. For high-sensitivity work (<10 pmol), baseline shifts must be minimal and reagent purity must be adequate so that labile amino acid derivatives are not destroyed.
SUPPORT PROTOCOL 1
The HPLC connected to the HP G1005A biphasic reaction column sequencer (see Basic Protocols 3 and 4) is a three-solvent system using preformulated solvents provided by the manufacturer. This HPLC system typically needs little or no adjustment in the parameters discussed below—e.g., gradient and solvents. The following guidelines are for an HPLC system connected to the Perkin-Elmer Procise Model 491, 492, or 494 sequencer; however, the same separation system can also be used on older sequencers. A typical gradient program is shown in Table 11.10.4 and a summary of typical solvent and gradient adjustments is illustrated in Table 11.10.5.
Table 11.10.4 Gradient for Separation of PTH amino acidsa
Step no.
Time (min)
%B
1 2
0.0 0.2
8 11
3 4
0.4 18.0
14 44
5 6
18.5 21.0
90 90
aFlow
Table 11.10.5
rate of 325 µl/min used throughout.
Optimizing the Separation of PTH Amino Acids
Problem
Corrective action
Rise in baseline late in chromatogram
Add small aliquots of 1% acetone to buffer A
Negative baseline between DTT and Glu His too close to Ala or eluting after Ala
Add small aliquots of 1.0 M KH2PO4 to buffer A Increase Premix in buffer A
Arg too close to Tyr or eluting after Tyr Ile/Lys not well separated (Lys early)
Increase Premix in buffer A Decrease %B at 18-min point in gradient
Lys/Leu not well separated (Lys late)
Increase %B at 18-min point in gradient Chemical Analysis
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Additional Materials (also see Basic Protocol 1) 1% (v/v) acetone in Milli-Q water 1.0 M KH2PO4 in Milli-Q water PTH Analyzer Standard mixture (standard mixture of PTH amino acids; ABI Biotechnology) 1-liter bottles and three-valve caps (Rainin) NOTE: Prepare solutions with Milli-Q water or equivalent taken directly from the purification unit. Water stored for any length of time in glass or plastic containers supports microbial and algal growth, which results in high amino acid background in early sequencer cycles. 1. Prepare the “A solvent” by adding 15 ml Premix Buffer Concentrate to 1 liter of solvent A3. Unopened bottles of solvent A3 and the Premix Buffer Concentrate are stored at 4°C; partially used bottles of solvent A3 with additives can be stored at room temperature after displacing the air in the bottle with argon. Solvent A3 should be replaced after being in use for 7 to 10 days at room temperature. Peroxides will form with time in this solvent at room temperature and will adversely affect the yield of labile amino acids, especially lysine. The Premix Buffer Concentrate contains an ion-pairing additive that affects the peak shape and elution position of PTH-Arg, PTH-His, and the pyridylethyl derivative of cysteine. Addition of 15 ml Premix Buffer Concentrate to 1 liter A3 should position the peak for His in front of Ala and the peak for Arg in front of Tyr but after the dehydroalanine derivative of Ser.
2. Prepare the “B solvent” as follows. Dissolve the contents of one tube (500 nmol) of DMPTU in 1 ml solvent B2, vortex the tube, and allow to sit at least 15 min with additional vortexing every 5 min. Add the entire contents to 1 liter of solvent B2. Solvent B2 should resolve PTH-Trp from the typical Edman degradation product diphenylurea (DPU). Addition of 500 nmol DMPTU improves the recovery of PTH-Ile, PTH-Lys, and PTH-Leu, especially when analyzing <10 pmol. The rise in baseline late in the chromatogram that is caused by the addition of DMPTU may need to be adjusted with the addition of acetone to the A3 solvent (see step 6).
3. Transfer the freshly prepared solvents from steps 1 and 2 to two separate clean, dry 1-liter bottles, each sealed with a three-valve cap. The caps from Rainin are more robust than those supplied by the sequencer manufacturer and are easily replaced if leaking of the argon used to pressurize the bottles occurs.
4. Attach the bottle containing the A solvent to the A pump of the sequencer and the bottle containing the B solvent to the B pump. Purge the A and B pumps with the fresh solvents to eliminate any air bubbles in the pumps or solvent supply lines. The purge also replaces any old solvents in the pumps so that the first gradient is run entirely with the fresh solvents.
Optimize baseline 5. Repeatedly run the (blank) gradient illustrated in Table 11.10.4 using the “Run Gradient” program, each time adjusting the composition of the A solvent to render the baseline as flat as possible (steps 6, 7, and 8). Each time adjustments are made to the solvent composition, purge the pump with the adjusted solvent and run another blank gradient to observe the effects of the additions on the baseline. 6. Flatten late baseline rise by adding acetone in Milli-Q water to the A solvent. N-Terminal Sequence Analysis of Proteins and Peptides
Small amounts of 1% acetone in 100-ìl increments can be added to the A solvent to produce an absorbance match between the A and B solvents. This results in a flat baseline when a
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linear gradient is used. Typically, ∼2 ml of 1% acetone is required per liter of A solvent to achieve this effect in a case where DMPTU is added to B solvent as described in step 2.
7. Correct early negative baseline slope by adding 1.0 M KH2PO4 to the A solvent. Frequently, a negative slope is observed in the beginning of the chromatogram between DTT peak and Glu (see Fig. 11.10.4). Addition of up to 100 ìl of 1.0 M KH2PO4 in 10-ìl increments will flatten the baseline in this area.
8. For high-sensitivity sequence analysis (<10 pmol), continue to adjust the A solvent as necessary with 1% acetone and 1.0 M KH2PO4 until the rise in baseline is no more than 0.1 mAU between 0 to 21 min. Optimize positions of PTH amino acid peaks 9. After the baseline has been optimized, inject the PTH Analyzer Standard mixture by running the “PTH-Standards” program and evaluate the separation. If needed, repeat the run adjusting the composition of the A solvent to optimize the separation (steps 10 and 11). Each time adjustments are made to the solvent composition, purge the pump with the adjusted solvent and carry out another standard run to observe the effects of the additions on the chromatogram. The chromatogram illustrated in Figure 11.10.4 shows good separation of the common PTH amino acids and the Edman degradation byproducts DMPTU, DPTU, and DPU.
10. Add additional Premix Buffer Concentrate to the A solvent to move the His and Arg peaks earlier in the chromatogram if needed. Aging of the PTH analyzer column may necessitate increasing the amount of Premix Buffer Concentrate in order to maintain the position of His and Arg. Add Premix Buffer Concen-
D 0.50
Q TG
0.00 DTT
m AU
−0.50
N
S
E DMPTU
DPU
A
Y
H
K
W M V DPTU
F
L
P R
−1.00
I
−1.50 S'
−2.00 −2.50 6.0
9.0
12.0 min
15.0
18.0
Figure 11.10.4 Chromatogram of PTH standard, illustrating an example of a typical separation of PTH amino acids and Edman degradation byproducts for the Perkin-Elmer Procise Models 491, 492, and 494 sequencers. The common amino acids recovered during the sequencing process are well separated, as are the normal byproducts of the Edman process: DMPTU (dimethylphenylthiourea), DPTU (diphenylthiourea), and DPU (diphenylurea). Unmodified cysteine is completely destroyed. Dithiothreitol (DTT) is typically present in the R4 reagent. S′ is an adduct between DTT and PTH-dehydroalanine, the latter of which is a degradation product of PTH-serine. Unmodified cysteine also forms S′, and can be distinguished from a serine residue because only the S′ peak increases in cycles containing unmodified cysteine. However, assigning cysteine based solely on this minor, often somewhat variable peak is risky. Abbreviations: mAU, milli–absorbance units; see Table A.1A.1 for one-letter amino acid abbreviations.
Chemical Analysis
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trate to solvent A in 2-ml increments to position His before Ala and Arg before Tyr. The ability of Premix Buffer Concentrate to shift His and Arg retention times is lost if >25 ml are added to the A solvent. In this case, the A solvent can be prepared again as in step 1, but with the amount of Premix Buffer Concentrate added to a liter of A3 reduced to 12 ml, which should position His after Ala and Arg after Tyr. Alternatively, the PTH analyzer column can be replaced with a new one.
11. If further optimization of the separation is necessary for Ile, Lys, and Leu, reposition these peaks by making changes to the gradient as noted in Table 11.10.5. Ideally, Lys should be centered between Ile and Leu. Consult the manufacturer’s instructions if any other separation problems are observed. SUPPORT PROTOCOL 2
INSTALLING AN INTERNAL HPLC RESTRICTER LOOP AND OPTIMIZING MAXIMUM INJECTION VOLUME ON OLDER SEQUENCERS The Perkin-Elmer Procise Model 491, 492, and 494 sequencers monitor the volume of sample injected onto the in-line HPLC by using fluid sensors on the inlet and outlet of the sample loop. However, most older Perkin-Elmer/ABI Biotechnology sequencers have a 50-µl injection loop that results in injection and analysis of only ∼42% of the total sample from the converter flask (∼120 µl). Clogging of the external restricter also frequently occurs, which results in lost sequence data. The proportion of sample injected can be increased to ∼85% and the risk of clogging the injector can be avoided by modifying the sample loop using a 100-µl loop with an internal restricter. Injecting a larger portion of the sample onto the HPLC increases sensitivity and is particularly helpful for high-sensitivity sequencing. Simply adding a larger loop would result in a very narrow time window for proper loop loading, which would result in variable injections due to inconsistent filling of the larger loop. The addition of an internal restricter avoids variable injections, as sample flow slows markedly before the loop is completely filled. The time window for loading the loop prior to injection is hence much longer and results in more consistent injections. Using the modified loop, a maximum amount of sample, ∼102 µl out of 120 µl total or 85%, of the total volume is injected. In addition, the probability of missed injections resulting from a clogged restricter line is dramatically reduced because the internal restricter is back-flushed upon each injection. Materials Stainless steel tubing cutter 100-µl stainless steel rheodyne injection loop (Rainin) Two restricter lines: 30-cm (length) × 0.007-in. (i.d.) stainless steel tubing with appropriate stainless steel nuts and ferrules (Upchurch) Zero-dead-volume coupler (Upchurch) Rheodyne injector on older Perkin-Elmer sequencer (Fig. 11.10.5) Add modified injector loop 1. Using a stainless steel tubing cutter, cut the end fitting off one end of any commercially available rheodyne 100-µl stainless steel loop, immediately behind the ferrule (if in place on loop).
N-Terminal Sequence Analysis of Proteins and Peptides
2. Connect the cut end of the 100-µl loop to one piece of restricter tubing using a zero-dead-volume coupler by inserting one piece of tubing halfway into the zerodead-volume fitting and holding it in place while tightening the nut onto the ferrule. Insert the second piece of tubing as far as possible into the coupler and hold it firmly in place while tightening the connection. Care should be taken to ensure that there is no gap between the two pieces of tubing in the zero-dead-volume connector.
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0.007-in. x 30-cm restrictors
A
B pump
1
1
2 zero-deadvolume fitting
2 column
6
6 3
3 5
5
4 from sequencer
LOAD
4
INJECT
100-µl loop
Figure 11.10.5 Illustration of an injector loop from an older model sequencing system, modified with an internal restricter. One end of the internal restricter is connected by a zero-dead-volume fitting to a 100-µl loop and the other end is connected to port 1 of the injector. The external restricter is connected to port 6 (overflow tube) and the two restricters aid in controlling the proper filling of the 100-µl loop/internal restricter assembly. As the sample is loaded (A), the restricters are encountered last and slow the speed of flow as the loop is nearly filled. When the sample is injected (B), the flow though the internal restricter is reversed. This reversal of flow on each injection prevents clogging of the restricter lines, which is normally the major cause of lost sequence data.
3. Connect the restricter end of the modified loop with port 1 of the injector valve and the full-bore end of the modified loop to port 4 (see Fig. 11.10.5). 4. Attach the second restricter line to port 6 of the rheodyne injector. 5. Flush injector loop with 100% acetonitrile in the Load position, switch injector to Inject position, run HPLC at typical flow rate, and check for leaks. Correct as required. SEQUENCE DATA INTERPRETATION After a sequencer run is completed, the next step is to assign a sequence by analyzing the data from all cycles. Although assigning some sequences can be straightforward—e.g., from a short, pure peptide at the 100-pmol level—accurate sequence assignment is often complex and usually requires an experienced scientist who is familiar with the recent sequencing performance history of the instrument used. Although Perkin-Elmer instruments have a software feature that automatically assigns sequences, an independent study that analyzed sequence assignment accuracy on an unknown sample showed that software assignments were much less accurate than those obtained by the average experienced sequencer operator using manual assignments (Yuksel et al., 1991). This same study showed that operators using the software-assigned sequence to assist them in their manual assignments achieved less accurate results than those operators who did not use software assignments at all. Care must be particularly exercised when interpreting the sequence of larger proteins, peptides containing contaminating (secondary) sequences, long peptide or protein sequence runs, and analyses where the signal is below the 10-pmol level. Factors that must be considered when assigning sequence include the expected recovery of amino acids, the repetitive yield, signal carryover (lag) in subsequent cycles, and increases in background from nonspecific acid cleavage of peptide bonds. In addition,
SUPPORT PROTOCOL 3
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Table 11.10.6
Expected Recoveries of PTH Amino Acidsa
Recovery (%)
Amino acid Arg Asn Gln His Ile Lys Ser Thr Trp
Procise
HP G1005A
50-75 95b 90c 50 100 75-90 30-50 50 10-40
75-100 95b 90c 50-75 50d 50-100 10-20 25-40 20-70
aThe common amino acids not listed are essentially quantitatively recovered on both sequencers (except cysteine which is completely destroyed during sequencing unless chemically modified to form a stable derivative). bAsparagine is partially deamidated and detected as aspartic acid. cGlutamine is partially deamidated (∼10%) and detected as glutamic acid. dIsoleucine on this system is resolved as its two isomers; one isomer migrates with phenylalanine.
accurate assignment of early cycles in a sequence can be problematic because of high amino acid background resulting from contamination of the sample (from many sources) prior to loading into the sequencer. In general, careful manual examination of the tabulated data from all cycles of the analysis, together with a comparison of the corresponding chromatograms, will produce the most accurate sequence assignment. Expected Recoveries of PTH Amino Acids Many PTH amino acids are essentially quantitatively recovered during the sequencing process, whereas others are partially destroyed or incompletely extracted (see Table 11.10.6). However, the recoveries of these problematic amino acids are usually fairly consistent for a given instrument, sequencer method, and set of reagents; they should normally be within the ranges indicated in Table 11.10.6. Lower-than-normal or variable recoveries are usually indicative either of chemical modification of the sample during isolation or problematic sequencer performance (resulting from hardware or reagent quality). When assigning sequences, the amino acid recoveries recently observed on the instrument used must be considered, especially when analyzing low-level or difficult sequence data sets. Effects of Repetitive Yield on Number of Cycles Assigned
N-Terminal Sequence Analysis of Proteins and Peptides
The background-corrected signal observed at each cycle of a sequence relative to the background-corrected signal observed in the previous cycle is the repetitive yield. Repetitive yield can be most easily calculated from the slope of the best-fit line when the logarithm of the net sequence signal observed (in pmol) is plotted versus cycle number. The size of the sequence signal always steadily decreases as a sequence run progresses, as a result of two factors. First, the efficiency of the Edman chemistry for each sequencer cycle is high but slightly less than 100%. A reasonable estimate for the overall efficiency of the reactions in each cycle when using an optimized sequencer and high-purity reagents
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20
Yield (pmol)
10
95% 92% 90% 1 85%
0.5
2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 34 36 38 40 Cycle number
Figure 11.10.6 Effects of repetitive yield on number of cycles assigned. Theoretical amino acid yields for several different repetitive yields that are frequently encountered in automated sequence analysis illustrate the importance of this parameter on the amount of sequence that can be assigned. An initial yield of 10 pmol and a sequence detection limit of 0.5 pmol were used for this example.
would be ∼96% to 98%. The second factor that decreases the yield is loss of sample from the sample support (sample washout). This second factor is very dependent upon the specific characteristics of individual samples, the type of sequencer support used, optimization of solvent deliveries in the sequencer, and related factors. The relationship between repetitive yield and the amount of sequence that can be obtained is illustrated in Figure 11.10.6. In this example, an initial yield of 10 pmol and a sequencing detection threshold of 0.5 pmol is used. As shown, a repetitive yield of 95% would allow sequence assignment of >40 residues in this case, whereas lower repetitive yields decrease the maximum amount of sequence information that can be obtained. The observed repetitive yield for any sequencer run is related to both sample characteristics and sequencer operation. Optimization of all sequencer parameters to yield the highest possible repetitive yields, using an appropriate standard protein or peptide, will insure that the maximum amount of sequence can be assigned when sequencing experimental samples. Carryover (Lag) Can Complicate Sequence Assignments in Later Cycles Incomplete coupling and incomplete cleavage of the N-terminal residue during each sequencer cycle results in partial transfer of an amino acid signal into subsequent cycles. This effect is cumulative, so that carryover is usually but not always low in early cycles and progressively increases throughout the sequence. Cleavage at prolines is especially difficult, and an obvious jump in carryover at prolines is sometimes observed. The increase in carryover in later cycles and its influence on assigning sequences in later cycles is shown in Figure 11.10.7. The alanine signal in cycle 3 is very clear, with a small amount of carryover into cycle 4. The alanine signal at cycle 13 is much smaller, and the alanine value in cycle 14 is nearly equal to the value in cycle 13. The alanine in cycle 14 results from carryover and not an Ala-Ala sequence. Cycle 14 is a threonine, as indicated by the increase in signal of this residue in this cycle. Another alanine occurs in cycle 23 and the further increase in alanine signal in cycle 24 is due to a progressive further increase in
Chemical Analysis
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12
Yield (pmol)
10 8 6 4 2 0 0
2
4
6
8
10 12 14 16 18 20 22 24 26 28 Cycle number
Figure 11.10.7 Yields of alanine and threonine from a sequence exhibiting somewhat higher-thannormal carryover. Uncorrected values of Ala (crosses) and Thr (triangles) in each cycle of a sequence are shown. The amount of carryover in this sequence was larger than the average for this instrument, but within the range of carryover seen with some samples. The high carryover in later cycles complicates sequence interpretation. This sequence contained Ala at cycles 3, 13, and 23, and Thr at cycle 14.
carryover. In this worse-than-average sequence, by cycles 13 and 14 carryover of the alanine and threonine signals, respectively, extend into the n + 1, n + 2, n + 3, and n + 4 cycles before the value of the amino acid returns to background levels. Background in Early Sequencer Cycles Assigning the first several residues in a sequence can often be difficult because of contaminants introduced during sample loading or from the sample itself, especially when the initial sequence signal is <10 pmol. These contaminants are primarily free amino acids and small peptides from such sources as airborne contamination, ungloved fingers, and buffers used in the purification. They result in a high level of multiple amino acids (background), especially glycine and serine, in the first one or more sequencer cycles.
N-Terminal Sequence Analysis of Proteins and Peptides
The first four cycles from a sequence run using 10 pmol of β-lactoglobulin loaded to a GFF are shown in Figure 11.10.8A as an example of a very clean sample with minimal initial background. The first residue, leucine, as well as subsequent cycles can be easily assigned. The sequence assignment for the four cycles shown is LIVT (i.e., Leu-Ile-ValThr; see Table A.1A.1 for one-letter amino acid abbreviations). The sequence in Figure 11.10.8B is shown at the same sensitivity as the β-lactoglobulin sequence; however this experimental sample has substantial free amino acid and small-peptide contamination that results in a high background of several amino acids in early cycles. The multiple signals present in the first three cycles make assignment of sequence in these cycles difficult, but a clear glutamate (E) signal in cycle 4 and additional signals in subsequent cycles show that this protein is not blocked; hence a definitive sequence assignment from cycle 4 on was made. Very tentative assignments for cycles 2 and 3 were made based on knowledge of how rapidly background normally declined in analogous samples in this sequencer; however, the more conservative assignment for the cycles shown would be XXXE (where X represents an unassigned residue).
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A
B L
1.0 I
m AU
0 .0
V
− 1.0 − 2.0 T − 3.0
E
− 4.0 4
6
8
10 12 14 16 18 20 4 min
6
8
10 12 14 16 18 20
Figure 11.10.8 Chromatograms of the first four cycles from two different low-pmol-level sequences. (A) 10 pmol β-lactoglobulin were loaded onto a Procise 494. The first four residues can be clearly identified above a moderate initial background. (B) A tryptic peptide loaded to the same sequencer. In this case, the high background starting in cycle 1 interferes with sequence assignment of early cycles and the first unambiguous assignment that can be made is E, in cycle 4. Abbreviations: mAU, milli–absorbance units; see Table A.1A.1 for one-letter amino acid abbreviations.
Calculation of Background-Corrected Signals In addition to early-cycle background from contaminants, most sequences exhibit a detectable level of most amino acids in most cycles of the sequence. The major source of this background is “nonspecific” acid cleavage of peptide bonds that free new internal sequences during the cleavage step of each sequencer cycle. Because there are fewer peptide bonds in peptides as compared with proteins, background is usually minimal for peptides, but it becomes increasingly higher for larger proteins as they have more peptide bonds to be randomly cleaved. The new internal N-termini start to produce many very low-level sequences in subsequent cycles. As the degree of cleavage is extremely low and somewhat random, a fairly uniform background results, which steadily increases throughout the sequence as more peptide bonds are cleaved at each cycle. Peptide bonds involving serine, threonine, and aspartic acid are the most labile; proteins or peptides that have a high content of these residues usually show higher background levels than other proteins of the same size. Table 11.10.7 illustrates data from 10 pmol of β-lactoglobulin loaded to a GFF and sequenced for 15 cycles with the Procise 494. The picomole values listed for each amino acid/cycle are the uncorrected or raw values automatically calculated versus the PTH standard (see Fig. 11.10.4) injected at the beginning of the run. Because β-lactoglobulin is a fairly small protein, there is detectable background for most amino acids; this increases steadily throughout the sequence run but it does not interfere with sequence assignment. The accuracy of the reported values in this table should be verified by examining each chromatogram and correcting any problems such as incorrect peak identification or
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Supplement 8
1.81 0.92 0.70 0.66 0.76 0.78 0.74 0.77 0.74 0.84 0.87 0.88 0.85 0.87 0.90
0.45 0.47 1.20b 0.78 0.98 0.56 0.88 0.63 0.66 0.62 [3.66] 0.99 0.72 0.67 0.64
D 0.79 0.60 0.57 0.59 1.15c 0.71 0.76 0.80 0.76 0.75 0.75 0.76 1.16c 0.83 0.76
E 0.45 0.34 0.28 0.24 0.25 0.17 0.30 0.26 0.34 0.36 0.37 0.37 0.35 0.42 0.41
F 0.36 0.15 0.16 0.20 0.16 0.11 0.19 0.16 0.14 0.09 0.09 0.13 0.14 0.14 0.08
H
dSer contamination is common in cycle 1.
cGlu value has increased due to partial deamidation of Gln in cycle.
bValue incorrect due to incorrect baseline.
2.41 1.91 1.54 1.52 1.49 1.48 1.53 1.68 [5.05] 2.05 1.83 1.83 1.87 1.89 1.92
G
K
L
M
0.80 1.95 [6.86] 0.32 [7.76] 0.86 0.75 0.15 0.76 0.58 0.66 0.13 0.53 0.53 0.67 0.13 0.51 0.52 0.83 0.10 0.41 0.42 1.03 0.13 0.44 0.43 1.02 [4.36] 0.30 [3.59] 0.98 0.51 0.60 0.90 1.02 0.23 0.53 0.55 [4.99] 0.20 1.61 0.26 0.00b 0.51 [4.85] 0.56 1.29 0.26 1.10 0.49 1.26 0.24 0.74 [2.97] 1.21 0.23 0.67 0.94 1.27 0.23
I 0.48 0.45 0.42 0.28 0.38 0.38 0.38 0.38 0.37 0.35 0.35 0.26 0.37 0.39 0.42
N 0.41 0.48 0.60 0.54 0.56 0.60 0.56 0.49 0.47 0.46 0.45 0.42 0.45 0.22 0.51
P
Q 1.03 0.59 0.51 0.61 [5.24] 0.91 0.64 0.66 0.69 0.69 0.68 0.67 [4.07] 1.15 0.80
PTH Amino Acid Yields (Uncorrected) from 15-Cycle Sequence Using 10 pmol β-lactoglobulina
aThe assigned sequence is indicated by brackets.
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15
A
Table 11.10.7
N-Terminal Sequence Analysis of Proteins and Peptides
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0.48 0.62 0.47 0.66 0.60 0.72 0.68 0.66 0.70 0.80 0.69 0.73 0.68 0.79 0.79
R
T
V
3.21d 0.93 0.83 1.73 0.45 0.46 0.94 0.37 [5.95] 0.79 [4.00] 0.60 0.67 0.48 0.47 0.65 [3.47] 0.58 0.64 0.53 0.55 0.72 0.40 0.52 0.67 0.35 0.52 0.72 0.32 0.60 0.78 0.27 0.61 0.77 0.27 0.62 0.73 0.26 0.58 0.81 0.30 0.57 0.79 0.32 [3.71]
S
0.10 0.00 0.00 0.13 0.00 0.00 0.00 0.09 0.03 0.04 0.00 0.00 0.05 0.03 0.04
W
0.65 0.44 0.42 0.43 0.42 0.50 0.50 0.52 0.55 0.56 0.59 0.54 0.62 0.58 0.57
Y
inconsistent peak integration. In some cases, it may be necessary to reintegrate and reprint the entire sequence run prior to manually computing the net or background-corrected amino acid yields of assigned residues in the sequence. After the accuracy of reported values is verified, an appropriate background level can be estimated for each observed sequence signal and subtracted from the sequence signal observed in cycle n. In general, the most appropriate background value is the amount observed in the cycle before the sequence signal (n – 1). For example, in Table 11.10.7 the background-corrected value for Val in cycle 3 would be 5.5 pmol (5.95 − 0.46 = 5.49). An alternative to evaluating a table of raw data values as shown in Table 11.10.7 is to plot the values for each residue versus cycle number—i.e., the type of plots shown in Figure 11.10.7 for an unrelated sequence. Careful evaluation of net sequence yields for each cycle is especially important for complex sequence data sets and can be useful in identifying potential errors in initial sequence assignments. Especially when one or more lower-level (secondary) sequences are present in addition to a major (primary) sequence, evaluation of sequence yields minimizes the risk of confusing signals from the secondary sequence with positions in the primary sequence, where labile or post-translationally modified residues might occur. Sequence Assignment The simplest data set from a sequenced sample is one signal per cycle, as illustrated by the data set in Table 11.10.7. In this case the sequence is easily assigned—e.g., LIVTQ TMKGL DIQKV. Assignments are typically reported using the following conventions: uppercase letters for positive calls (confidence level should be >99%); lowercase letter or letter enclosed by parentheses for a tentative call (confidence level between 50% and 99%); and X for no assignment possible. When two or more signals are present in each cycle from the sample sequenced and the sequence is unknown, assignment becomes more challenging. A single major sequence in the presence of one or more sequences at much lower levels (less than a third the major sequence level) can usually be assigned with substantial confidence. Sorting of multiple sequences must rely heavily on quantitative yields and must consider corrections for residues not recovered quantitatively (Table 11.10.6). In some cases, a minor sequence can also be assigned, but assignment of some residues in this sequence may be complicated by interference that is due to carryover of the major sequence. Mass Analysis Can Aid Sequence Assignments As briefly discussed in Strategic Planning, obtaining a mass for HPLC-purified peptides by MALDI mass spectrometry (UNITS 11.6 & 16.2) can aid in assigning sequences. When the entire sequence is assigned by Edman sequencing, comparing the calculated and observed masses provides a useful check that the correct sequence assignment has been made. Similarly, when one or two residues have been assigned tentatively and the remainder of the peptide sequence has been positively assigned, the confidence level of the tentative calls can often be increased or decreased by considering the observed mass. Finally, when a single residue cannot be assigned and the remainder of the sequence has been positively assigned, the difference between the observed and calculated masses will yield the mass of the unassigned residue, which frequently suggests a likely assignment for this position. Examples where this approach is useful include: an X (unassignable residue) in the first cycle resulting from initial background, a missing last residue in a tryptic peptide resulting from washout of the last residue, an unmodified cysteine, or a post-translationally modified residue not detected by Edman sequencing. Mass analysis of peptides is therefore a very valuable complementary method when used in conjunction with Edman
Chemical Analysis
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sequencing. However, caution must be exercised when using mass analysis to assign residues where no or multiple Edman signals are present. In some cases, more than one possible interpretation of tentative or unassigned residues may fit within the error limits of the mass analysis, which can be up to ±0.1% or more, depending upon the calibration method and instrument used. SUPPORT PROTOCOL 4
OPTIMIZING SEQUENCER PERFORMANCE Optimal performance of a protein sequencer depends on careful and frequent evaluation of multiple parameters including proper instrument delivery of solvents and reagents, quality of sequencer reagents and solvents, sequencer performance, and yields of nonquantitatively recovered amino acids when analyzing appropriate peptide and protein standards (Tempst and Riviere, 1989; Atherton et al., 1993; Tempst et al., 1994). For additional information on problems in sequencer performance and their solutions, see Troubleshooting. Sequencer Standards Commonly used sequencing standards include: human serum albumin (HSA; typically used for the Hewlett-Packard G1005A sequencer); β-lactoglobulin (typically used for the Procise 491, 492, 494, and other Perkin-Elmer sequencers). Horse apomyoglobin is also sometimes used as a protein standard. Peptide standards suitable for either sequencer include melittin or the sequencing test peptide from Sigma. A sequence of a standard sample should be evaluated at least once for every 10 to 20 experimental samples sequenced, to detect noncatastrophic sequence-performance problems. The standard sample should be analyzed at a level similar to the average levels of experimental samples that are being analyzed—e.g., 5, 10, or 50 pmol. Standard protein or peptide solutions should be prepared in adequate quantity for multiple analyses, then quantified by amino acid analysis, divided into multiple aliquots, and stored at −20°C. A fresh aliquot can be used for every third standard run to minimize possible variation resulting from degradation or contamination of the aliquot during handling. Results from all sequences for each type of standard can be listed in a spreadsheet-format database that includes such items as background corrected yield for each cycle, repetitive yield, and carryover at a specific reference cycle. Sequences that exhibit optimal initial yields, maximum repetitive yields, low carryover, and good recoveries of all amino acids can then be used as reference point for evaluating sequencer performance in future sequence runs. Proper Delivery of Solvents and Reagents During Sequencer Cycles
N-Terminal Sequence Analysis of Proteins and Peptides
Each manufacturer publishes guidelines for evaluating and adjusting deliveries of all solvents and reagents used in the sequencing process for their respective sequencer. Direct observation should be made of all solvent and reagent deliveries for a complete Edman cycle (both reaction and conversion portions of the cycle) at least once every three to six sequencer runs; these should then be compared to the manufacturers recommendations. These observations should be kept in a log book containing a record of pertinent information corresponding to each sequence run. A sample log sheet for this purpose is shown in Figure 11.10.9. These records are useful for identifying trends, such as a gradual reduction in wash solvent volume delivery, before they become problematic to the point where sequencer performance is significantly affected. Significant adjustments to such parameters as delivery volumes and pressures should be evaluated by sequencing an appropriate standard to verify good sequencer performance prior to sequencing experimental samples.
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Quality of Solvents and Reagents All solvents and reagents should be marked with the date of receipt prior to being stored according to the manufacturer’s recommendations. Solvents and reagents should be free of any kind of particulate matter, haze, or cloudiness. The lot numbers should be changed in the sequencer-bottle menus whenever a bottle is refilled or replaced on the sequencer. The actual date that the bottle is placed on the sequencer should also be written on the label of the bottle and the appropriate information should be recorded in the sequencer log book (see Fig. 11.10.9). When analyzing samples at the <10-pmol level, many changes in sequencer performance that are not due to hardware problems can quickly be correlated
Figure 11.10.9 A sample log sheet for recording critical observations for each sequence. This is for documenting reagent lot numbers, gas pressures, and observations of reagent/solvent deliveries, which facilitates troubleshooting when sequencer performance problems are encountered. If a log sheet is completed for each sequence, it can also serve as a checklist to ensure that adequate reagents, solvents, gases, and other necessary items are available for the entire sequencer run.
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with a lot change for a specific sequencer reagent by careful tracking of reagent lot numbers and installation dates. In general, dramatic changes in sequencer performance are related to hardware failures or a marked decrease in the quality of the most recently changed solvent or reagent. COMMENTARY Background Information
N-Terminal Sequence Analysis of Proteins and Peptides
Most protein sequencing projects fall into one of two groups: (1) partial sequence analysis of one or more proteins isolated from natural sources that correlate with a biological process of interest, or (2) analysis of recombinant proteins or fragments of these proteins as part of structure-function studies or verification of structural integrity for pharmaceutical applications. The chemical reactions used in modern sequencers involve the cyclic removal of the N-terminal residue from a protein or peptide after modification with phenylisothiocyanate (PITC). This basic chemistry remains largely unchanged since it was first described in 1949 by Pehr Edman (Edman, 1949). However, the amount of protein required for routine sequence analysis has steadily declined from micromole quantities using early automated sequencers (Edman and Begg, 1967) to the current 10- to 100-pmol levels (Yuksel et al., 1991; Crimmins et al., 1992). Although most sequencing laboratories can effectively sequence samples at the >100-pmol level, substantial optimization of instrumentation and associated methods must be invested to ensure that a sequencing laboratory can routinely obtain sequence data at the 10- to 100-pmol level. Only a small proportion of sequencing laboratories have optimized their methods sufficiently to permit fairly routine high-sensitivity sequence analysis at the 1- to 10-pmol level. Hence, the selection of an appropriate sequencing facility can be an important consideration for projects where the amount of sample is limited. The capacity to sequence samples at maximum sensitivity appears to be largely a function of the rigor that is devoted to instrument optimization, monitoring, and maintenance—rather than of differences in specific instrumentation or methods used (Tempst and Riviere, 1989; Yuksel et al., 1991; Crimmins et al., 1992). In general, N-terminal sequence analyses of intact proteins and large peptide fragments are most effectively performed by electroblotting the sample from an appropriate one- or two-dimensional gel onto a high-retention PVDF membrane (UNIT 10.7). Electrotransfer efficiencies of 50% to 80% can usually be obtained for
most proteins, and total losses including artifactual blocking of free N-terminals and transfer losses are usually <35% even at the <10pmol level when appropriate precautions are taken (Mozdzanowski and Speicher, 1992; Speicher, 1994). Substantial N-terminal sequence information can be obtained from an electroblotted protein when as little as 2 to 5 pmol of protein is present on the blot and extensive blocking of the N-terminus has been avoided during sample preparation. When 2 pmol of a desired unblocked protein is electroblotted onto a PVDF membrane, the initial signal observed in the sequencer is expected to be between 0.4 and 1.4 pmol. As noted above, sequence analysis at this level is currently feasible, but certainly not routine. In some cases, it may be preferable to sequence a soluble, highly purified protein that is already in a solution compatible with direct sequence analysis (see Basic Protocol 1 or 3) instead of using SDS gels and electroblotting. Because 50% to 80% of all proteins isolated from natural sources are physiologically blocked (Brown and Roberts, 1976), the current preferred strategy for obtaining partial sequence data from an unknown protein is to immediately attempt internal sequencing rather than N-terminal sequencing of the intact protein. This approach involves four sequential steps: in situ digestion with a specific protease such as trypsin either in the gel or on a PVDF membrane, microbore or narrow-bore reversed-phase HPLC separation of the tryptic peptides, MALDI mass analysis of selected peak fractions, and finally N-terminal sequencing of one or more of the largest peptides as indicated by the mass analysis. Over the past several years, protein identification by mass fingerprinting has emerged as an increasingly useful alternative approach to Edman sequencing of unknown proteins. In this method, peptides from in-gel digestion of a protein band using a specific protease such as trypsin are analyzed as a mixture using MALDI mass spectrometry (UNITS 11.6 & 16.2). These multiple peptide masses are then searched against a protein database that has been transformed from a sequence database to a database
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Table 11.10.8
Troubleshooting and Optimizing Sequencer Performancea,b
Problem
Possible cause(s)
No sequence and no increase in back- Insufficient sample ground in later cycles No sequence; background does Sample is N-terminally blocked increase in later cycles
Suggested solution(s) Load larger amount of sample and try to quantify amount loaded Cleave protein using trypsin and sequence internal peptides and/or evaluate isolation procedure for conditions that might block N-terminus Replace with full bottle of R4 and/or check delivery Replace extraction solvent with full bottle; check delivery of extraction solvent; fix injector
No sequence; DMPTU, DPTU, and DPU peaks present No sequence and no injection solvent front
R4 bottle empty or reagent not delivered Extraction solvent bottle empty or solvent not delivered; injector clogged or not working
High DMPTU, DPTU, DPU
Poor washing of sample after coupling; poor R4 drydowns
Observe washes and optimize per manufacturer’s recommendations; observe dry times and adjust as needed
PTC analogs of PTH amino acids present
Poor R4 drydowns
Observe dry times and adjust as needed
Low PTH amino acid yields in all cycles High background in initial cycles
Insufficient transfer of ATZs from sample support Sample excessively contaminated
Optimize extractions
Low initial background increases too Acid cleavage time is excessive rapidly High lag, low RY, low DMPTU, DPTU, DPU High lag after Pro
Low initial yield
Large peak early in chromatogram (air injected onto HPLC column)
Clean up sample, buffers, buffer water source, etc. Reduce delivery of TFA (R3)
R2 depleted or too old; R2 delivery too short Typical after Pro residue
Replace R2; check R2 delivery and adjust if needed Can increase R3 reaction time for Pro-rich samples (but will increase background) R1 bottle empty or poor delivery; R1 Replace R1 and/or check delivery; too old; argon gas contaminated replace if >30 days old; replace argon tank with higher-purity gas Check S4 delivery and adjust if needed; Low delivery of S4; injection loop adjust loop fill time; check values and over/under filled; clogged flask sensors; clean pickup tube or replace; pickup tube; S4 clinging to flask clean or replace flask walls
aIf dramatic changes in delivery times are observed or if delivery is variable, possible hardware problems should be considered instead of simply adjusting delivery times for solvents and reagents. bAbbreviations:
ATZ, 2-anilino-5-thiazolinone; DMPTU, dimethylphenylthiourea; DPTU, diphenylthiourea; DPU, diphenylurea; PTC, phenylthiocarbamyl; PTH, phenylthiohydantoin; RY, repetitive yield; TFA, trifluoroacetic acid.
of tryptic peptide masses (Henzel et al., 1993; Mann et al., 1993; Pappin et al., 1993; Yates et al., 1993). The validity of protein identification by this method is dependent upon multiple factors including the accuracy of the masses used for the search. The stringency of the comparison can be increased dramatically if partial sequence information from tandem mass spectrometry methods (MS/MS; UNIT 16.2) are used in addition to the tryptic peptide masses, either using postsource decay on a reflectron MALDI mass spec-
trometer or using an electrospray quadruple mass spectrometer (triple quadruple or ion trap). Of course, if the specific protein sequence from the species being studied is not in the database, a reliable match is unlikely. However, as the sequence databases are rapidly expanding to include complete genomes of multiple species, the utility of this approach will continue to rapidly expand. Most importantly, this approach has the potential to reliably identify proteins by mass matching with limited se-
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Table 11.10.9
Troubleshooting and Optimizing HPLC Performancea,b
Problem
Possible cause(s)
No peaks and no changes in baseline
Suggested solution(s)
HPLC pumps not running; detector lamp not on Retention times for all PTH amino HPLC pump seals worn; A or B acids are variable pump delivery erratic Retention times for early PTH amino Insufficient reequilibration prior to acids are variable injection
Evaluate and correct HPLC problem(s); turn on lamp or replace if needed Replace pump seals; identify problem pump and repair or replace Increase reequilibration time
PTH amino acid peaks become broader with time Noisy baseline; erratic small sharp spikes in baseline
Replace guard column at least every 30 days; replace PTH column Replace lamp
aAlso
Contaminants on guard column; PTH column aging HPLC lamp aging
see Table 11.10.5 for baseline adjustments and optimization of PTH amino acid separation.
bAbbreviations:
HPLC, high-performance liquid chromatography, PTH, phenylthiohydantoin.
quence analysis by MS/MS methods at one to two orders of magnitude higher sensitivity (100 to 1000 femtomoles of protein) than the current maximum sensitivity for internal sequencing (10 to 20 pmol) using the Edman sequencing approach described in this unit (Shevchenko et al., 1996).
Critical Parameters When isolating proteins from natural sources, the most difficult part of the project is usually the isolation of a sufficient amount of the protein of interest in a form that is free of contaminating proteins, amino acids, and incompatible inorganic compounds. Purifying relatively low-abundance proteins from natural sources is particularly challenging. In such situations it is advisable to keep the purification scheme to the fewest possible steps and to use either one-dimensional SDS gels or two-dimensional gels for the final purification step. Regardless of the abundance of the target protein or purification method, the vast majority of proteins currently analyzed in sequencing facilities are separated by SDS-PAGE prior to either N-terminal sequence analysis or internal sequencing. Detailed guidelines for sample isolation are described above (see Strategic Planning) and should be carefully followed.
Troubleshooting
N-Terminal Sequence Analysis of Proteins and Peptides
The first step in troubleshooting sequencer problems is to distinguish between sequence analysis problems and problems arising from the sample—i.e., sample heterogeneity, blocked N-terminal sequence, or insufficient sample loaded into the sequencer. Suspected
sequence analysis problems can be readily evaluated by immediately analyzing an appropriate standard peptide. Table 11.10.8 lists some of the more common problems encountered in sequencer operation and optimization; Table 11.10.9 lists common problems for the HPLC analyzer. In general, dramatic changes in sequencer performance are related to hardware failures or a marked decrease in the quality of the most recently changed solvent or reagent.
Anticipated Results When the N-terminus of the protein or peptide loaded to the sequencer is not blocked, an initial sequence yield should be observed equal to ∼20% to 80% of the amount of sample loaded into the sequencer. The first several cycles of any sequence may have signals from multiple amino acids, which can complicate making positive identifications for these cycles; however, tentative assignments can often be confirmed if the mass of the peptide has been determined prior to sequence analysis. The high initial background, if observed, should disappear by cycle 2 or 3 unless contamination is extremely severe. Some peptide and protein samples will contain minor contaminating sequences, and these secondary sequences can often be distinguished from the major sequence if a large quantitative difference exists. However, caution must be exercised in this case, as the repetitive yields of the major and minor sequences may be different because of differing degrees of sample washout. For example, a short major peptide may exhibit a lower repetitive yield than a minor longer peptide, and
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similar signals might be observed near the end of the major sequence, resulting in potential misassignment to the major sequence of subsequent residues in the minor sequence. When homogeneous peptides are being sequenced, the entire sequence can usually be assigned unless the peptide exhibits unusually high washout. When proteins or large peptides are being sequenced, several factors can limit the amount of sequence that can be obtained, including repetitive yield, progressive increase in carryover as the sequence proceeds, and increased background from nonspecific acid cleavage. Typically, most proteins exhibit high repetitive yields (>92%); hence repetitive yield is rarely the most limiting factor when sequencing proteins except when the sequencer is not optimized or the initial sequence yield is near the detection limit (usually 0.1 to 1 pmol). As a general rule, when sequencing proteins >100 kDa, increased background is likely to be the most important limiting factor. For proteins <100 kDa, the limiting factor is usually the rising carryover.
Time Considerations The time required for preparation of the sequencer and HPLC prior to loading the samples can range from ∼5 min (if sufficient sequencer reagents/solvents and HPLC solvents are on the instrument for the projected number of cycles) to up to 2 hr when multiple reagent replacements are needed. Loading PVDFbound samples to the Procise sequencer requires ∼15 min, which includes excising the sample from the membrane(s), washing it, drying it, and loading it into a clean Blott cartridge. Loading PVDF-bound samples into the Hewlett-Packard biphasic cartridge sequencer requires an additional 20 min, as the sample cartridge must be precycled on the sequencer. Loading liquid samples onto the HewlettPackard biphasic cartridge sequencer takes ∼20 to 40 min for precycling the sample cartridge while simultaneously cleaning the sample loading funnel, followed by 20 min to load the sample onto the sample cartridge. Loading liquid samples onto the Procise or other sequencers using a glass fiber filter (GFF) sample support requires the most time. It takes ∼15 min to load Polybrene on a GFF and set up two filter precycles that take ∼35 min each to precondition the GFF on the sequencer prior to sample application. The time required for sample application will vary widely depending upon the volume to be loaded. It takes ∼5 min to apply and dry 15 µl of sample on a preconditioned
GFF. Loading large volumes using repetitive application can be quite time-consuming; for example, it would take about 1 hr to load 150 µl in 15-µl aliquots. After the run is complete, the time required for analyzing a sequence is related to the length of the sequence and the complexity of the data. A straightforward 20-cycle peptide sequence may require as little as 30 min to analyze. In contrast a very complex 20-cycle multiple-sequence data set with integration problems may require at least 2 hr for careful data analysis. It is also advisable for two experienced sequencer operators to independently assign very complex sequences, to improve the accuracy of the assignments.
Literature Cited Atherton, D., Fernandez, J., DeMott, M., Andrews, L., and Mische, S.M. 1993. Routine protein sequence analysis below ten picomoles: One sequencing facility’s approach. In Techniques in Protein Chemistry IV (R. Angelletti, ed.) pp. 409-418. Academic Press, San Diego. Brown, J.L. and Roberts, W.K. 1976. Evidence that ∼80% of the soluble proteins from Ehrlich ascites cells are N-alpha acetylated. J. Biol. Chem. 251:1009-1014. Crimmins, D.L., Grant, G.A., Mende-Muller, L.M., Niece, R.L., Slaughter, C., Speicher, D.W., and Yuksel, K.U. 1992. Evaluation of protein sequencing core facilities: Design, characterization, and results from a test sample (ABRF91SEQ). In Techniques in Protein Chemistry III. (R. Angelletti, ed.) pp. 35-43. Academic Press, San Diego. Edman, P. 1949. A method for the determination of the amino acid sequence in peptides. Arch. Biochem. Biophys. 22:475-480. Edman, P. and Begg, G. 1967. A protein sequenator. Eur. J. Biochem. 1:80-91. Erdjument-Bromage, H., Geromanos, S., Chodera, A., and Tempst, P. 1993. Successful peptide sequencing with femtomole level PTH-analysis: A commentary. In Techniques in Protein Chemistry IV. (R. Angelletti, ed.) pp. 419-426. Academic Press, San Diego. Henzel, W.J., Billeci, T.M., Stults, J.T.,Wong, S.C., Grimley, C., and Watanabe, C. 1993. Identifying proteins from two-dimensional gels by molecular mass searching of peptide fragments in protein sequence databases. Proc. Natl. Acad. Sci. U.S.A. 90:5011-5015. Hewick, R.M., Hunkapiller, M.W., Hood, L.E., and Dryer, W.J. 1981. A gas-liquid solid phase peptide and protein sequencer. J. Biol. Chem. 256:7990-7997. Mann, M., Hojrup, P., and Roepstorff, P. 1993. Use of mass spectrometric molecular weight information to identify proteins in sequence databases. Biol. Mass Spectrom. 22:338-345.
Chemical Analysis
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Miller, C.G., Bente, H.B., Fisher, S., Myerson, J., Wagner, G., Widmayer, R., and Horn, M.J. 1990. Sequence analysis of diverse protein systems using a novel column-based protein sequencer. Abstract T143, Fourth Symposium of the Protein Society, San Diego. Mozdzanowski, J. and Speicher, D.W. 1992. Microsequence analysis of electroblotted proteins I. Comparison of electroblotting recoveries using different types of PVDF membranes. Anal. Biochem. 207:11-18. Pappin, D.J.C., Hojrup, P., and Bleasby, A.J. 1993. Rapid identification of proteins by peptide-mass fingerprinting. Curr. Biol. 3:327-332. Reim, D.F. and Speicher, D.W. 1994. A method for high-performance sequence analysis using polyvinylidene difluoride membranes with a biphasic reaction column sequencer. Anal. Biochem. 216:213-222. Sheer, D.G., Yuen, S., Wong J., Wasson, J., and Yuan, P.M. 1991. A modified reaction cartridge for direct sequencing on polymeric membranes. Biotechniques 11:526-534. Shevchenko, A., Wilm, M., Vorm, O., and Mann, M. 1996. Mass spectrometric sequencing of proteins from silver-stained polyacrylamide gels. Anal. Chem. 68:850-858. Speicher, D. W. 1994. Methods and strategies for the sequence analysis of proteins on PVDF membranes. Methods 6:262-273. Tempst, P. and Riviere, L. 1989. Examination of automated polypeptide sequencing using standard phenylisothiocyanate reagent and subpicomole high-performance liquid chromatographic analysis. Anal. Biochem. 183:290-300.
Tempst, P., Geromanos, S., Elicone, C., and Erdjument-Bromage, H. 1994. Improvements in microsequencer performance for low picomole sequence analysis. Methods 6:248-261. Yates, J.R., Speicher, S., Griffin, P.R., and Hunkpiller, T. 1993. Peptide mass maps: A highly informative approach to protein identification. Anal. Biochem. 214:397-408. Yuksel, K.U., Grant, G.A., Mende-Muller, L., Niece, R.L., Williams, K.R., and Speicher, D.W. 1991. Protein sequencing from polyvinylidenefluoride membranes: Design and characteristics of a test sample (ABRF-90SEQ) and evaluation of results. In Techniques in Protein Chemistry II. (J.J. Villafranca, ed.) pp. 151-162. Academic Press, San Diego.
Key References Edman and Begg, 1967. See above. First description of the basic Edman chemistry. Hewick et al., 1981. See above. Describes the first sequencer capable of picomolelevel sequencing. Tempst et al., 1994. See above. Recent review containing additional suggestions for improving sequencing sensitivity.
Contributed by David F. Reim and David W. Speicher The Wistar Institute Philadelphia, Pennsylvania
N-Terminal Sequence Analysis of Proteins and Peptides
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Determination of Disulfide-Bond Linkages in Proteins
UNIT 11.11
The formation of disulfide bonds in proteins is an important post-translational modification that is essential for stabilizing and maintaining the three-dimensional structure of proteins, a property which is critical for their biological activity. The disulfide linkages in a protein cannot be predicted from its amino acid sequence; therefore, determination of disulfide linkages is an important step toward a detailed understanding of the structural properties of a protein. The primary method given in this unit (see Basic Protocol) describes disulfide-bond mapping using immobilized trypsin as the cleavage agent for proteins. Disulfide-containing peptides are then identified from chromatographic separations of the tryptic digest under reducing and nonreducing conditions. In Alternate Protocol 1, chemical cleavage of proteins with cyanogen bromide (CNBr) is presented. CNBr cleavage generates larger fragments and is especially suited for use as the first cleavage step for disulfide bond mapping of a large protein (>50 kDa). CNBr is also able to cleave proteins with extensive secondary structure that are resistant to most proteolytic cleavage. Alternate Protocol 2 describes a partial reduction and alkylation strategy to determine disulfide linkages of proteins/peptides with multiple disulfide-linked cysteine residues that have no appropriate cleavage sites between half-cystinyl residues or are resistant to cleavage. The protein of interest is converted into a mixture of isoforms that ideally have only one of its disulfide bonds reduced. Subsequent alkylation and analysis of the isoforms allow assignment of the disulfide linkages. In the Support Protocol, a method is presented to determine the number of disulfide bonds present in the protein. This protocol should be performed first and the information obtained used to confirm the completeness of the disulfide bond assignments. NOTE: The major problem in disulfide bond determination is disulfide bond scrambling: the exchange of partners between thiols and disulfides (see Strategic Planning and Critical Parameters and Troubleshooting). Disulfide scrambling occurs at neutral or alkaline pH, especially after the native structure is disrupted with denaturants or by partial hydrolysis of the protein with proteases or chemical cleavage reagents. Disulfide scrambling can usually be suppressed at low pH. Hence, all procedures should be performed in buffers with pH <7 whenever possible.
STRATEGIC PLANNING Overall Strategy The basic strategy for disulfide bond mapping involves four major steps (Fig. 11.11.1). First, the protein is cleaved by proteases and/or chemical cleavage reagents under nonreducing and acidic conditions to minimize disulfide bond rearrangement. Second, protein fragments are separated from one another by chromatographic methods. Third, disulfidecontaining complexes are identified and confirmed by their ability to be reduced into constituent peptides. Finally, the identified disulfide-linked peptides are characterized by mass spectrometry (MS; see UNIT 16.1 for overview) and N-terminal Edman sequencing (UNIT 11.10). Multiple disulfide-containing peptides are subjected to further chemical/protease cleavages or analyzed by partial reduction and alkylation to produce simpler disulfide-linked peptides suitable for unambiguous determination of cysteine residues involved in disulfide bond formation. Figure 11.11.2 illustrates the determination of Chemical Analysis Contributed by Hsin-Yao Tang and David W. Speicher Current Protocols in Protein Science (2004) 11.11.1-11.11.20 C 2004 by John Wiley & Sons, Inc. Copyright
11.11.1 Supplement 37
Figure 11.11.1
Overall strategy for location of disulfide bond in proteins.
disulfide linkages of a protein (GA733-2) with six disulfide bonds, and also shows the structure of some commonly encountered disulfide-linked peptides.
Determination of Disulfide-Bond Linkages in Proteins
Special Instruments The procedures listed here require the use of a reversed-phase high performance liquid chromatography (RP-HPLC) system (UNIT 11.6), a matrix-assisted laser desorption/ionization (MALDI)-time of flight (TOF) mass spectrometer (UNIT 16.2), and/or a tandem mass spectrometer (UNIT 16.10). Frequently, an automated Edman sequencer (UNIT 11.10) is also helpful. Availability and expertise with these instruments are essential; otherwise the procedures can be performed by utilizing a protein chemistry/proteomics core facility or by collaborating with a biochemical research laboratory with the needed
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Figure 11.11.2 Disulfide bond determination of the GA733-2 antigen. The protein contains six disulfide bonds (gray lines) located within the first 115 residues of the protein. The locations of the cysteine residues involved in disulfide linkages and some of the tryptic cleavage sites are shown. Digestion of the protein with trypsin generated three major disulfide-containing complexes: T1, T2, and T3. The T1 complex was deglycosylated and further cleaved chemically with BNPS-skatole to generate single disulfide-linked peptides. The partial reduction and alkylation strategy was used to determine the three disulfide linkages of the T2 complex. The alkylated cysteine residues are indicated by the italicized lowercase “c.” MALDIMS (UNIT 16.2) and Edman sequencing (UNIT 11.10) of the T2-R3 and T2-R5 complexes revealed disulfide-bond formation between Cys6-Cys36, and between Cys15-Cys25, respectively. Based on the two disulfide bond assignments, the third disulfide bond is deduced to form between Cys4-Cys23.
resources. To assist in the identification of the numerous peptides generated by chemical/proteolytic cleavage, bioinformatics software capable of correlating masses with peptide sequences is indispensable. The authors’ typically use the GPMAW program http://welcome.to/gpmaw, but free web-based analysis software, such as Protein Prospector (Clauser et al., 1999) and SearchXLinks (Schnaible et al., 2002a), are also suitable.
Sample Preparation The assignment of correct disulfide linkages in proteins containing multiple disulfide bonds is a challenging task and becomes increasingly difficult as the number of cysteine residues increases. In addition, as noted above, disulfide bond exchange can contribute to the difficulty of disulfide bond assignment. Very often, a protein requires multiple consecutive or simultaneous cleavages to produce suitable disulfide-linked peptides for unambiguous identification. To ensure the success of the project, a relatively large amount of protein (i.e., 10 to 50 nmol) is desirable, especially for proteins with high disulfide bond content. This is usually not a problem because disulfide linkages are often determined from purified recombinant proteins with known amino acid sequences. More
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importantly, protein purification strategies must be modified to avoid potential disulfide bond rearrangements. Whenever possible, buffers used in protein purification should be slightly acidic (pH ∼6.5) to minimize disulfide scrambling, particularly during steps that may perturb tertiary structure, and reducing agents should not be used. If proteases will be used for fragmentation of the protein, protease inhibitors should be omitted from the purification (see UNIT 11.1) or removed after purification.
Production of Disulfide-Containing Peptides Cleavage of the protein at sites that will produce simple disulfide-linked peptides is a critical step toward successful assignment of disulfide linkages. Cleavage is often performed in the presence of denaturing agents (urea or guanidine·HCl) to allow optimal access to the cleavage sites. The choice of protease (UNIT 11.1) or chemical cleavage reagent (UNIT 11.4) to use for protein cleavage is not straightforward and in part depends on the susceptibility of the protein to the cleaving reagents and the position of cysteine residues in the protein sequence. Ideally, the cleavage agent selected should produce peptides with only one disulfide bond and should be active at a pH <7. In practice, a combination of cleavage agents is often required to produce a series of peptides suitable for disulfide bond assignment (Chong and Speicher, 2001; Roszmusz et al., 2001; Chong et al., 2002). Specific cleavage agents such as trypsin, endoproteinase Glu-C, and CNBr have the advantage of producing easily identified fragments because the N- or C-terminal residues are known; however, the specificity of these reagents may not permit sufficient cleavages for production of peptides linked by a single disulfide bond. To circumvent this problem, broad specificity cleavage methods (e.g., partial acid hydrolysis, pepsin, subtilisin) can be employed. The major disadvantage of using broad specificity cleavage agents is that the complexity of the peptides generated is greatly increased, making the isolation and identification of disulfide-linked peptides much more difficult. In addition, a larger amount of the protein is usually required when nonspecific cleavage agents are used because the yields of specific peptides are reduced due to nested cleavages. Cleavage agents such as CNBr (see Alternate Protocol 1), partial acid hydrolysis, endoproteinase Glu-C, and pepsin are favored because of their capacity to work effectively at low pH. The repertoire of proteases with low pH optima is limited, but often sufficient cleavage can be achieved at suboptimum pH conditions. To compensate, proteases with alkaline pH optima, such as trypsin, are typically used at higher concentrations with longer incubation times. In this respect, immobilized proteases are preferred to avoid contamination of the sample with large amounts of protease autolytic products (see Basic Protocol). A number of potentially useful immobilized proteases (e.g., endoproteinase Glu-C, trypsin, pepsin, α-chymotrypsin) are available from MoBiTec and other suppliers. A list of cleavage agents with typical reaction conditions for disulfide bond mapping is shown in Table 11.11.1. In situations where proteolytic and chemical cleavages are not able to produce peptides with single disulfide-linkages, partial reduction and alkylation methods can be employed (see Alternate Protocol 2). This strategy depends on Edman sequencing (UNIT 11.10) or tandem mass spectrometry (UNIT 16.10) to identify the cysteine residues and will work well with cleavage agents, such as trypsin, that produce peptide fragments with less than ∼30 residues.
Determination of Disulfide-Bond Linkages in Proteins
Separation and Identification of Disulfide-Containing Peptides For separation of cleavage products, the method chosen depends mainly on the fragment sizes. For large fragments such as those generated by CNBr cleavage, gel-filtration chromatography is often useful (see Alternate Protocol 1). For separation of smaller fragments, such as those produced by protease digestion, RP-HPLC with 0.1% TFA in a water/acetonitrile binary system is the separation method of choice. This system is
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Table 11.11.1 Proteases and Chemical Cleavage Agents Suitable for Use in Disulfide Bond Mapping
Cleavage residue
Buffera
Temp
Time
Reference
BNPS-skatole
Trp
0.1% TFA
37◦ C
1 hr
Chong and Speicher (2001)
Cyanogen bromide
Met
70% formic acid
23◦ C
16 hr
Chong et al. (2002)
Cleavage agent Chemical
Partial acid hydrolysis
◦
Broad
0.25 M oxalic acid
100 C
8 hr
Zhou and Smith (1990)
Chymotrypsinb
Trp, Tyr, Phe
50 mM sodium phosphate, pH 5.5
37◦ C
16 hr
Schnaible et al. (2002a)
Endoproteinase Glu-C (V8 proteinase)
Glu, Asp
100 mM sodium acetate, pH 4.0
24◦ C
16 hr
Lippincott and Apostol (2002)
Endoproteinase Lys-C
Lys
25 mM Tris·Cl, pH 6.8
37◦ C
24 hr
Leal et al. (1999)
Pepsin
Broad
0.02 N HCl, pH 2.0
37◦ C
16 hr
Bures et al. (1998)
◦
Protease
Subtilisin
Broad
50 mM sodium phosphate, pH 6.5
37 C
16 hr
Chong et al. (2002)
Thermolysin
Broad
0.1M triethylamine· HCl, pH 6.0
37◦ C
16 hr
Bures et al. (1998)
Trypsin
Lys, Arg
50 mM sodium phosphate, pH 6.0
37◦ C
16 hr
Schnaible et al. (2002a)
Trypsin (immobilized)
Lys, Arg
50 mM sodium phosphate, pH 6.5
23◦ C
16 hr
Chong and Speicher (2001)
a Urea or guanidine·HCl is commonly included in the buffer to denature the protein. See Appendix 2A for sodium phosphate and Tris·Cl buffer recipes. b Chymotrypsin also cleaves a number of other peptide bond with widely varying efficiency.
capable of detecting minor changes in the hydrophobic character of the peptides, such as those caused by reduction of intrachain disulfide bonds, making it a highly sensitive method for detecting disulfide-containing peptides (see below). In addition, the RP-HPLC solvents have a pH of ∼2, which is optimal for preventing disulfide scrambling. The general strategy to identify disulfide-containing peptides is to detect alterations to peptide mobility as a consequence of disulfide bond reduction. An early yet elegant approach was the use of diagonal paper electrophoresis of peptides with performic acid cleavage of disulfide bonds between the first and second dimension separation (Brown and Hartley, 1966). Similar approaches are still used, except that paper electrophoresis is replaced by higher sensitivity methods, such as comparison of peptide mobilities by RP-HPLC in the nonreduced and reduced states (see Basic Protocol). Changes in the hydrophobicity of dissociated peptides after reduction can easily be detected by this method (UNIT 11.6). Mass spectrometry can also be used to directly analyze the unfractionated nonreduced and reduced states to rapidly provide information on disulfide linkages (Morris and Pucci, 1985). Interchain disulfides are identified by the appearance of two signals in the reduced digest that correspond to the mass of a single signal in the nonreduced digest, while intrachain disulfides are identified by the mass increase of 2 Da after reduction. The mass spectrometric approach works best with simpler proteins, and high performance mass spectrometers with high mass accuracy and resolution are required for analyzing relatively high molecular masses, especially peptides with intrachain disulfide bonds. While it is feasible to perform direct mass spectrometric analysis of cleavage
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fragments, it is often desirable to isolate the peptide fragments by RP-HPLC prior to mass analysis as this allows repeated and additional analysis, such as Edman sequencing, to be performed on the isolated peptides.
Sequence Determination of Disulfide-Containing Peptides The sequence of the disulfide-containing peptides and location of the disulfide bonds can be interpreted using amino acid analysis (UNIT 11.9), tandem mass spectrometry (UNIT 16.10), or automated Edman sequencing (UNIT 11.10). Amino acid analysis requires relatively pure samples, which in turn requires well resolved peaks on the RP-HPLC separation. Tandem mass spectrometry, in the form of MALDI postsource decay or electrospray ionization collision-induced dissociation (ESI-CID), requires the least amount of sample and does not require well-separated peptides (Jones et al., 1998; Pitt et al., 2000; Schnaible et al., 2002b). By coupling liquid chromatography with tandem mass spectrometry (LCMS/MS), disulfide linkages can be determined from the unfractionated tryptic digest using <100 pmol protein (Yen et al., 2000). The mass spectrometric approach is comparatively faster and requires relatively small quantities of samples, making it the method of choice when sample is limiting. Another major advantage of tandem mass spectrometry is the ability to determine the disulfide linkages of simple multiple disulfide-containing complexes, such as three peptide chains linked by two disulfide bonds or two peptide chains with one intra- and one interchain disulfide bond. However, the fragmentation data from a disulfide-containing complex is complicated and can often be hard to interpret. For unambiguous sequence determination, investigators frequently use MALDI-MS analysis followed by Edman sequencing where needed. The availability of peptide masses and predicted sequences helps in the interpretation of Edman sequencing data when disulfidecontaining peptides are not totally resolved by RP-HPLC separation.
BASIC PROTOCOL
TRYPSIN CLEAVAGE AND DISULFIDE-BOND MAPPING OF PROTEINS In this method, the protein is denatured and digested with immobilized trypsin under nonreducing conditions in the presence of intermediate concentrations of a denaturant such as urea. The proteolytic fragments generated are separated by RP-HPLC and disulfide-containing complexes are identified by comparing the reduced and nonreduced chromatograms. The peaks of interest are further characterized by MALDI-TOF mass spectrometry (UNIT 16.2) and/or N-terminal Edman sequencing (UNIT 11.10).
Materials
Determination of Disulfide-Bond Linkages in Proteins
Urea buffer (see recipe) Protein sample, lyophilized or aqueous (i.e., in mildly acidic buffer without serine protease inhibitors and with known concentration) Argon Urea, solid (for aqueous proteins only) 1M HCl 1M NaOH 50 mM sodium phosphate, pH 6.5 (APPENDIX 2E) Reaction buffer: 3 M urea/50 mM sodium phosphate, pH 6.5 Trifluoroacetic acid (TFA) Reducing solution, pH 8.0 (see recipe) Solvent A: 0.1% TFA in H2 O Solvent B: 0.1% TFA in 95% acetonitrile 50% acetonitrile/0.1% TFA
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Immobilized tosylphenylalanine chloromethylketone (TPCK)-treated trypsin F7m column (MoBiTec) 10-ml syringe HPLC system, with 2.1-mm C18 reversed-phase column (e.g., ZORBAX 300SB-C18), UV detector, fraction collector, peak detector, and chart recorder C18 ZipTip (Millipore) Additional reagents and materials for RP-HPLC (UNIT 11.6) and MALDI mass spectrometry analysis of peptides (UNIT 16.2), and tandem mass spectrometry (UNIT 16.10) Denature protein 1a. For lyophilized proteins: Add 100 µl of urea buffer to 10 nmol lyophilized protein in a microcentrifuge tube. Blanket with argon, seal, and vortex lightly to dissolve. Use freshly prepared buffer and high purity urea to minimize accumulation of cyanate from urea.
1b. For aqueous proteins: Dissolve solid urea in 10-nmol protein solution to a final concentration of 9 M and readjust pH to 6.5 with 1 M HCl or NaOH as appropriate. The protein should be prepared in a mild acidic buffer without phenylmethylsulfonyl fluoride, diisopropyl fluorophosphate, aprotinin, or other serine protease inhibitors to prevent inactivation of trypsin (UNIT 11.1). As in step 1a, use freshly prepared buffer and high purity urea to minimize accumulation of cyanate from urea. The final volume should be 100 µl.
2. Incubate sample 30 min at 37◦ C to ensure protein denaturation. Avoid temperature higher than 37◦ C and prolonged incubation (>1 hr) to minimize carbamylation of cysteine and lysine residues. The rate of carbamylation is greatly reduced at pH 6.5 compared with higher pH. As denatured protein is more susceptible to disulfide scrambling (see Strategic Planning and Critical Parameters and Troubleshooting), proceed to cleavage immediately. Do not store denatured protein for disulfide mapping.
3. Dilute denatured protein solution with 200 µl of 50 mM sodium phosphate, pH 6.5, to lower the urea concentration to 3 M. Other proteases have different tolerance for urea (UNIT 11.1).
Equilibrate immobilized-trypsin F7m column 4. Equilibrate a TPCK-treated trypsin F7m column with 10 ml reaction buffer at room temperature. Connect the top of the column to a 10-ml syringe filled with reaction buffer and apply pressure to assist the flow. The column contains 200 µl F7m matrix, which has large pores and is suitable for proteins up to 107 Da. Immobilized trypsin is preferred because a larger amount of immobilized protease can be used without appreciable contamination of reaction solutions with protease autolytic products.
5. Microcentrifuge the column 5 sec at 2000 rpm to remove residual buffer.
Digest protein 6. Load the protein solution onto the column as follows. a. Apply protein solution to the column and allow it to flow through by gravity over a period of ∼1 hr. b. During this time, continue to reapply the sample onto the column as it elutes from the column. c. After passing the entire sample through the column approximately six times, cap the column outlet and incubate overnight at room temperature.
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Repeated application of the sample of the column and overnight incubation ensure more efficient protease cleavage.
7. Microcentrifuge the column 5 sec at 2000 rpm to collect the digested sample. 8. Remove residual peptides with two successive washes of 200 µl reaction buffer, followed by microcentrifuging 5 sec at 2000 rpm. Column can be reused after re-equilibration and should be stored in buffer without urea.
9. Combine digest and washes. Remove a 1.5-nmol aliquot for reduction with Tris(2carboxyethyl)phosphine (TCEP; see steps 10 to 12). Add TFA to the remainder of the sample to a final concentration of 2% (pH ∼2) and proceed to step 13 with this aliquot. Samples should be analyzed by RP-HPLC within ∼24 hr (store at 0◦ to 4◦ C until analyzed). Otherwise, store samples at −80◦ C. At pH 2, disulfide scrambling is largely inhibited.
Reduce disulfide bonds of digest with TCEP 10. Add an equal volume of reducing solution, pH 8.0, to the 1.5-nmol aliquot. Mix and blanket reaction mixture with argon. Although TCEP can function at acidic pH, reduction of disulfide bonds is most effective at pH 7 to 8. An inert atmosphere prevents reoxidation of cysteines to disulfides.
11. Incubate the mixture 1 hr at 37◦ C. 12. Adjust the pH of the reduced tryptic digest by adding TFA to a final concentration of 2%. This reduces the pH to ∼2 for RP-HPLC analysis.
Separate reduced and nonreduced peptides by RP-HPLC 13. Equilibrate an RP-HPLC column at 0.2 ml/min in 98% solvent A/2% solvent B (UNIT 11.6). A ZORBAX 300SB-C18 (2.1-mm inner diameter × 150 mm, 300-Å pore size, 5-µm particle size) or comparable column is suitable for this purpose. See UNIT 11.6 for details on RPHPLC separation of peptides.
14. If necessary, determine the optimal separation gradient for the protein or set of digestion conditions. Once this has been determined, inject the reduced (step 12) and nonreduced (step 9) samples separately onto the column. The separation gradient will need to be optimized for different proteins/digests to obtain the best separation for all the components. Refer to UNIT 11.6 for examples. For optimization, inject 20 pmol aliquots of each sample. Once a satisfactory gradient has been established, inject 80% of the remaining samples in 500-pmol aliquots to avoid overloading the column.
15. Elute peptides using the optimized acetonitrile gradient and collect all fractions with the help of a peak detector and fraction collector. Store eluted peptides at 4◦ C (up to a month) or −80◦ C (indefinitely).
Identify disulfide-linked peptides by MALDI-MS 16. Analyze all peaks in the nonreduced and reduced RP-HPLC chromatogram by MALDI-MS (UNIT 16.2).
Determination of Disulfide-Bond Linkages in Proteins
Large peptides are mixed 1:1 with a saturated solution of 3,5-dimethoxy-4hydroxycinnamic acid (sinapinic acid) in 33% CH3 CN and 0.1% TFA. Peptides <5 kDa are applied to a MALDI sample plate precoated with a saturated solution of nitrocellulose and α-cyano-4-hydroxycinnamic acid (1:4 w/w) in 2-propanol and acetone (1:1 v/v). Since each chromatographic peak could have a mixture of peptides with differing ionization efficiency, it is advisable to perform the MALDI-MS with more than one matrix (see UNIT 16.2). MALDI-MS is usually sufficient to identify peptides generated by high
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Figure 11.11.3 RP-HPLC separation and identification of the disulfide-linked tryptic peptides. Tryptic digest of the recombinant GA733-2 protein was separated on a ZORBAX 300SB-C18 column before (bottom panel) and after (top panel) reduction with TCEP. Major peaks, which disappeared following reduction are indicated by T1 to T3. The peptide constituents of the complexes are shown in Figure 11.11.2. T2* and T3* indicate incomplete tryptic cleavages of T2 and T3, respectively. Single peptides that appear as a result of reduction are indicated in the top panel by their residue numbers. Peptides were identified by MALDI-MS and, in some cases, by automated Edman sequencing (UNIT 11.10). Adapted with permission of ASBMB from Chong and Speicher (2001).
specificity proteolytic or chemical cleavages. Confirm ambiguous peptides by N-terminal Edman sequencing (UNIT 11.10) or tandem mass spectrometry (UNIT 16.10).
17. Compare RP-HPLC chromatograms of reduced and nonreduced tryptic digests. Confirm the presence of disulfide complexes by reducing the appropriate fractions. Peaks in the nonreduced chromatogram that disappear or have a decreased peak area after reduction probably contain disulfide-linked peptides. New peaks corresponding to constituent peptides of these disulfide-linked complexes are usually present in earlier eluting fractions of the reduced chromatogram. Figure 11.11.3 shows an example of the reduced and nonreduced chromatograms of a tryptic digest.
Reduce RP-HPLC fractions to confirm presence of disulfide complexes 18. Add 8 µl reducing solution to 2 µl RP-HPLC fractions from the nonreduced digest that contains the suspected disulfide-linked peptides. Mix and blanket reaction mixture with argon. 19. Incubate 1 hr at 37◦ C to reduce disulfide bonds. 20. Desalt the reaction solution using a Millipore C18 ZipTip or equivalent following manufacturer’s instructions.
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TCEP can interfere with efficient ionization of peptides and should be removed prior to MS analysis. Alternatively, if the sample is concentrated enough, dilute the reduced sample at least 2- to 4-fold with 0.1% TFA for MALDI-MS analysis.
21. Elute peptides with 2 µl of 50% acetonitrile/0.1% TFA and analyze by MALDI-MS (UNIT 16.2). Appearance of two new peptides having a combined mass (MH+) that is 3 Da higher than the nonreduced peptide complex indicates a single interpeptide disulfide bond. An internal peptide disulfide is identified by a mass increase of 2 Da compared with the nonreduced peptide. The disulfide assignment is complete if the complex contains a total of two cysteine residues (e.g., T3 complex in Figure 11.11.2). Cysteine residues not involved in disulfide linkages are identified from RP-HPLC fractions containing cysteinyl peptides that do not display retention time shifts or mass changes after reduction.
Confirm peptide sequence and (if necessary) choose an alternate cleavage strategy 22. Confirm sequence of disulfide-containing peptides by Edman sequencing (UNIT 11.10) or tandem mass spectrometry (UNIT 16.10) as needed. The disulfide-containing peptides from nonreducing fractions can be sequenced directly and the multiple residues at each cycle can be readily sorted based on the known protein sequence.
23. If reduction of the disulfide complex resulted in more than two peptides or at least one of two peptides contains multiple cysteines perform one of the following steps: a. Dry the RP-HPLC fractions completely (∼1 hr) in a Speedvac. Choose a new cleavage agent (Table 11.11.1), resuspend in appropriate denaturing buffer, and digest. Repeat steps 9 to 22. b. Alternatively, analyze the fractions by partial reduction and alkylation (see Alternate Protocol 2). ALTERNATE PROTOCOL 1
CYANOGEN BROMIDE CLEAVAGE OF PROTEIN Proteins or large peptide complexes are cleaved at the C-terminal side of methionine residues in 70% formic acid using CNBr. The low pH of the cleavage reaction prevents disulfide scrambling from occurring. The fragmentation is usually monitored by SDSPAGE and the cleavage products are subsequently separated by preparative HPLC gelfiltration for disulfide mapping. Due to the low occurrence of methionine residues in most proteins, CNBr cleavage usually generates relatively large fragments. Therefore, this procedure is often used as the first cleavage step for disulfide mapping of large proteins (>50 kDa) or proteins with large domains containing multiple disulfide bonds, which are usually resistant to denaturation and proteolysis.
Additional Materials (also see Basic Protocol) Glacial and 5% acetic acid Solution of purified protein with known sequence 88% and 70% formic acid CNBr Acetonitrile
Determination of Disulfide-Bond Linkages in Proteins
Desalting column (i.e., Bio-Rad Econo-Pac 10DG or equivalent) Fraction collector Glass vial with Teflon-lined cap Aluminum foil Additional reagents and materials for desalting protein samples (UNIT 8.3) and SDS-PAGE (UNIT 10.1)
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Prepare sample for CNBr cleavage 1. Add glacial acetic acid to ∼50 nmol purified protein solution to a final concentration of 5%. Vortex to mix. CNBr cleavage, like most chemical cleavages, tend to be incomplete; therefore, a higher amount of starting material is required. The protein quantity given (50 nmol) is a good starting amount, especially when CNBr is used as the first cleavage step in multistep disulfide mapping experiments.
2. Desalt the acidified protein sample on a desalting column (UNIT 8.3). Use 5% acetic acid to elute the protein. The acidic pH inhibits disulfide bond scrambling.
3. Collect 0.5-ml fractions for a total of 20 fractions. Determine the absorbance at 280 nm for each fraction and analyze by 1-D SDS-PAGE (UNIT 10.1) to verify elution of the protein. Pool all protein-containing fractions. 4. Lyophilize samples and reconstitute with 400 µl of 88% formic acid. Vortex to dissolve sample. Keep a 20-µl aliquot for SDS-PAGE.
Cleave protein with CNBr 5. Prepare a CNBr stock solution (0.5 mg/µl) immediately before use in a chemical fume hood using the following procedure: a. Weigh an empty glass vial and Teflon-lined cap, then move to the hood and transfer a small amount of CNBr into the vial. b. Tightly cap the vial, and weigh it again. c. Determine the amount of CNBr in the vial (in grams) and dissolve in the hood using 1 ml acetonitrile per 0.5 g CNBr. d. Wrap the vial in aluminum foil and keep it in the hood. CAUTION: CNBr is extremely toxic and must be neutralized with NaOH. See UNIT 11.4 for details. This procedure is necessary as balances generally do not function properly in fume hoods.
6. Determine the Met content of the protein and calculate the amount of CNBr required for 100-fold molar excess. For example, 50 nmols of a protein with 10 Met residues will require 50 nmol × 10 × 100 = 50 µmol CNBr. This is equivalent to 50 µmol × 0.106 mg/µmol = 5.3 mg CNBr.
7. Add appropriate amount of CNBr stock solution (0.5 mg/µl) to the sample from step 4 and add water to a final volume of 500 µl. Vortex to mix. Blanket reaction mixture with argon. The final concentration of formic acid is ∼70%.
8. Wrap the tube with aluminum foil. Incubate 24 hr at room temperature in the hood. The reaction must be carried out in complete darkness to avoid unwanted side-reactions. CNBr cleavage will result in peptide bond cleavage on the C-terminal side of methionines with conversion of the methionine to a mixture of homoserine and homoserine lactone. The predominant form is homoserine lactone which has a residue mass of 83.04 Da. The change in mass should be taken into consideration when identifying CNBr peptides using mass spectrometry.
Determine cleavage success or failure 9. Dry the reaction mixture with a Speedvac (∼1 hr) and resolubilize the sample in 400 µl of 70% formic acid. Repeat the lyophilization and reconstitute in 500 µl urea buffer.
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CAUTION: The CNBr will be in the trap of the Speedvac. Transfer to the chemical hood to discard CNBr and drain trap.
10. Analyze the cleavage reactions with reducing and nonreducing SDS-PAGE (UNIT 10.1). By loading proportional amounts of the total sample at each step, the success of the cleavage reaction can be easily determined from the intensities of protein staining and cleavage failure or excessive losses can be easily traced to a particular step.
Separate and analyze cleavage products 11. Separate the cleavage products by HPLC gel-filtration chromatography (UNIT 8.3). The authors typically use two TSK columns, G3000 SWXL and G2000 SWXL , connected in series with a running buffer of 10 mM sodium phosphate (pH 6.5)/150 mM NaCl/7 M urea, pH 6.5, at a flow rate of 0.6 ml/min at 4◦ C. (The running buffer is prepared fresh daily.) If the sample is separated at 4◦ C, the urea in the sample must be decreased from 9 to 7 M using buffer without urea.
12. Analyze separated fragments with reducing and nonreducing SDS-PAGE (UNIT 10.1) and MALDI-MS (UNIT 16.2). Fragments containing disulfide linkages can be identified easily after reduction by changes in migration.
13. Analyze disulfide-containing fragments as described (see Basic Protocol, steps 3 to 23). ALTERNATE PROTOCOL 2
PARTIAL REDUCTION AND ALKYLATION OF DISULFIDE BONDS Protein fragments with multiple disulfide linkages are partially reduced with Tris(2carboxyethyl)phosphine (TCEP) to generate a series of isoforms with only one disulfide bond reduced. The nascent free thiols are immediately alkylated with excess iodoacetamide. The partially reduced and alkylated fragments are separated by RPHPLC. Each isoform is characterized by MALDI-MS (UNIT 16.2) and Edman sequencing (UNIT 11.10) to reveal the position of the alkylated cysteine residues. This procedure is useful for dissecting protein fragments with multiple disulfide bonds when cleavage with proteases/chemicals fails to generate appropriate peptides with a single disulfide bond.
Additional Materials (also see Basic Protocol) Reducing solution, pH 3.5 (see recipe) RP-HPLC-purified peptide complex (see Basic Protocol) 1 M alkylation solution (see recipe) 0.3% and 10% TFA 1. Add an equal volume of reducing solution, pH 3.5 to ∼100 pmol (∼80 µl) of a RPHPLC purified peptide complex containing multiple disulfides. Incubate reaction mixture 3 min at room temperature. The reduction rate can be controlled by altering the amount of TCEP in the reaction mixture and the incubation time. It is necessary to optimize the reaction to generate yield of different disulfide forms. The yield of the different partially reduced isoforms can be conveniently determined by RP-HPLC. Determination of Disulfide-Bond Linkages in Proteins
2. Remove an aliquot (15% of reaction volume) and add to two volumes of 0.3% TFA. Store the reaction mixture on dry ice until ready for RP-HPLC analysis. Dilution with 0.3% TFA reduces the pH to ∼2 and decreases the acetonitrile content of the reaction mixture so that peptides can bind to the RP-HPLC column.
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Figure 11.11.4 RP-HPLC analysis of partial reduced and alkylated T2 disulfide-containing complex. Peptides were separated on a ZORBAX 300SB-C18 column. (A) T2 control before partial reduction and alkylation. (B) T2 after partial reduction with TCEP and alkylation with iodoacetamide. The structure of the labeled peptides is shown in Figure 11.11.2. Peptides were identified by MALDI-MS (UNIT 16.2) and the position of the alkylated cysteines was determined by Edman sequencing (UNIT 11.10). The small peak marked with an asterisk showed a molecular mass indicative of T2 with two alkylated cysteines, although the amount of this species was negligible compared with the major species (T2-R3). Adapted with permission of ASBMB from Chong and Speicher (2001).
3. Vigorously pipet the remaining partially reduced peptide solution into a microcentrifuge tube containing an equal volume of 1 M alkylation solution. Mix immediately by pipetting up and down a few times. To minimize disulfide scrambling, it is important to add the partially reduced peptides into the alkylation buffer and not vice versa. This will prevent a transient pH increase before a sufficient concentration of alkylation reagent is present.
4. Wrap reaction tube with aluminum foil and incubate 30 min at 37◦ C. Protect the alkylation solution and reaction from light to minimize photolytic production of iodine, a strong oxidant for thiols.
5. Terminate the reaction by adding 10% TFA to a final concentration of 1.3% (pH ∼2.0). Acidify the reaction quickly to prevent disulfide scrambling. Analyze the reaction immediately by RP-HPLC to prevent artifactual alkylation of other amino acid residues by the high concentration of iodoacetamide.
6. Separate both reaction mixtures (with and without alkylation) immediately by RPHPLC (UNIT 11.6). The alkylated peptides have different retention times than the nonalkylated peptides. Ideally, the number of peptide peaks from the partially reduced sample with alkylation should be the same as the partially reduced sample without alkylation. In practice, more peaks
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are normally observed in the alkylated sample due to some disulfide scrambling during alkylation; however, these peaks are relatively minor and should not pose a problem in the disulfide bond assignments.
7. Analyze all the RP-HPLC peaks by MALDI-MS (UNIT 16.2). Disulfide-linked peptides with a single reduced disulfide have a mass increase of 116 Da due to alkylation. When subjected to MALDI-MS analysis, disulfide-linked peptides often undergo in-source fragmentation of the disulfide bond to yield the constituent peptides. Alkylated constituent peptides have a mass increase of 57 Da over free cysteine. If the MALDI in-source fragmentation is not sufficient, the masses of the constituent peptides can be determined after reduction as described (see Basic Protocol, steps 18 to 21).
8. Verify the sequence and locate the alkylated cysteines by Edman sequencing (UNIT 11.10). Figure 11.11.4 shows an example of the partial reduction and alkylation of a multiple disulfide-containing complex. Disulfide-linked cysteine residues do not give any signal in an Edman sequencer whereas Scarboxamidomethylcysteine derivatives produce identifiable signals. Hence, the disulfide linkages should be easy to interpret. Theoretically, one isoform is sufficient to define a two disulfide-linked complex, and two isoforms for a three disulfide-linked complex, etc. In practice, additional isoforms are often obtained that can provide confirmation to the disulfide assignments. SUPPORT PROTOCOL
DETERMINATION OF TOTAL DISULFIDE BONDS The total number of disulfide bonds can be determined by comparing the molecular masses of the protein of interest obtained after (1) reduction and alkylation of all SH groups and (2) alkylation of the original free SH groups without reduction of disulfide bonds. Reduction is carried out with TCEP, and alkylation is performed with iodoacetamide. An alternative method to determine the number of disulfide bonds is to determine the free sulfhydryl content of the protein with Ellman’s reagent (UNIT 15.1). The colorimetric method is, however, less sensitive and requires nanomoles of protein.
Additional Materials (also see Basic Protocol) Protein solution Guanidine buffer (see recipe) Reducing solution, pH 6.5 (see recipe) 80 mM alkylation solution (see recipe) 0.3% TFA 50% acetonitrile/0.1% TFA 1. Mix 2 µl (∼10 pmol) protein solution with 15 µl guanidine buffer. Blanket with argon. 8 M urea can be used in place of guanidine-HCl.
2. Incubate sample 30 min at 37◦ C to denature protein. 3. Remove 5 µl denatured protein (step 2) and reduce by adding an equal volume of reducing solution, pH 6.5. Blanket with argon and incubate reaction 1 hr at 37◦ C.
Determination of Disulfide-Bond Linkages in Proteins
4. Remove 5 µl reduced protein (step 3) and alkylate with an equal volume of 80 mM alkylation solution. Blanket with argon and incubate reaction 1 hr at 37◦ C. To minimize disulfide scrambling and avoid incomplete modification, perform a series of alkylations with different incubation times to determine the minimal incubation period required for successful alkylation. The success of the reduction and alkylation procedure
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can be gauged from the experimental total number of cysteine residues (NCys , steps 9 to 10) compared to the expected number determined from the protein sequence.
5. In parallel, alkylate 5 µl denatured but nonreduced protein (step 2) as described in step 4. 6. Add an equal volume of 0.3% TFA to the remaining denatured protein (step 2), reduced protein (step 3), reduced and alkylated protein (step 4) and nonreduced alkylated protein (step 5). 7. Remove reaction buffer components using C18 ZipTips according to the instructions provided by the manufacturer. 8. Elute peptides from ZipTips with 2 µl of 50% acetonitrile/0.1% TFA. 9. Determine the molecular masses of denatured protein (Mstep2 ), reduced protein (Mstep3 ), reduced and alkylated protein (Mstep4 ), and nonreduced alkylated protein (Mstep5 ) by MALDI-MS (UNIT 16.2). Mix sample with 1:1 with a saturated solution of 3,5-dimethoxy-4-hydroxycinnamic acid (sinapinic acid) in 33% CH3 CN and 0.1% TFA, and apply to target plate.
10. Calculate the number of cysteine residues (NCys ): (NCys ) = (Mstep4 −Mstep3 )/57. where Mstep4 is the mass of the denatured, reduced, alkylated protein and Mstep3 is the mass of the denatured reduced protein. S-Carboxamidomethylation of a cysteine residue with iodoacetamide results in a mass increase of 57 Da.
11. Calculate the number of free sulfhydryl groups (NSH ): (NSH ) = (Mstep5 −Mstep2 )/57. where Mstep5 is the mass of the denatured, nonreduced, alkylated protein and Mstep2 is the mass of the denatured nonreduced protein. 12. Use this information to calculate the number of disulfide bonds: No. disulfide bonds = (NCys −NSH )/2.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Alkylation solution, 80 mM Dissolve the following in 8 ml H2 O: 0.15 g iodoacetamide (SigmaUltra; 80 mM final) 5.73 g guanidine hydrochloride (Pierce; 6 M final) 0.06 g sodium phosphate monobasic, anhydrous (50 mM final) Adjust pH to 7.5 with 10 M NaOH H2 O to 10 ml Protect solution from light Prepare fresh daily
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Alkylation solution, 1 M 1.85 g iodoacetamide (SigmaUltra; 1 M final) 2 ml 1 M HEPES, pH 7.5 (200 mM final) 40 µl 0.5 M EDTA, pH 7.5 (2 mM final) H2 O to 10 ml Protect solution from light Prepare fresh daily Guanidine buffer Dissolve the following in 8 ml H2 O: 5.73 g guanidine hydrochloride (Pierce; 6 M final) 0.06 g sodium phosphate monobasic, anhydrous (50 mM final) Adjust pH to 6.5 with 10 M NaOH H2 O to 10 ml Store up to one week at 23◦ C Reducing solution, pH 3.5 Dissolve the following in 9 ml H2 O: 0.143 g with Tris(2-carboxyethyl)phosphine (TCEP; Pierce; 50 mM final) 0.105 g citric acid monohydrate (50 mM final) Adjust pH to 3.5 with 10 M NaOH H2 O to 10 ml Prepare fresh daily Reducing solution, pH 6.5 Dissolve the following in 8 ml H2 O: 0.057 g TCEP (Pierce; 20 mM final) 5.73 g guanidine hydrochloride (Pierce; 6 M final) 0.06 g sodium phosphate monobasic, anhydrous (50 mM final) Adjust pH to 6.5 with 10 M NaOH Adjust volume to 10 ml with H2 O Prepare fresh daily Reducing solution, pH 8.0 Dissolve the following in 9 ml H2 O: 0.057 g TCEP (Pierce; 20 mM final) 0.158 g ammonium bicarbonate (200 mM final) Adjust pH to 8.0 with 10 M NaOH Adjust volume to 10 ml with H2 O Prepare fresh daily
Determination of Disulfide-Bond Linkages in Proteins
Urea buffer Dissolve the following in about 10 ml H2 O: 0.12 g sodium phosphate monobasic, anhydrous (50 mM final) 10.8 g urea (9 M final) Adjust pH to 6.5 with 10 M NaOH Adjust volume to 20 ml with H2 O Prepare immediately before use
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COMMENTARY Background Information The thiol (SH) group of cysteine is one of the most reactive amino acid side chains. In proteins, cysteines are usually present in the reduced form (with a free SH group) or in the oxidized form as cystines (two cysteines linked by a disulfide bond). Free SH groups are often found in the active site of proteins where they are typically involved in substrate binding and catalysis. Disulfide bonds are common posttranslational modifications that are critical for stabilizing the native structures of extracellular domains of proteins. Disulfide-bond linkages can occur between two cysteines in the same polypeptide chain (intrachain disulfide) or between two different polypeptides (interchain disulfide). Disulfide bond linkages cannot be predicted with any certainty and must be determined experimentally. Determination of disulfide bond linkages in proteins will provide insights into their three-dimensional structures, and because disulfide linkages are strongly conserved evolutionarily, conserved disulfide pairing patterns can be used to confirm common origins of domains with very low sequence homology. Cleavage-based strategies Due to the importance of disulfide bonds in the structure and function of proteins, a number of strategies have been devised to determine disulfide linkages. In a common strategy (see Basic Protocol) that has been applied successfully to numerous proteins, the protein of interest is subjected to extensive chemical and/or proteolytic cleavage in its nonreduced state to generate disulfide-linked peptides which are identified by comparing the RP-HPLC separation of peptides produced in the nonreduced versus the reduced state. The disulfide-linked peptides are subsequently isolated and analyzed by mass spectrometry and Edman sequencing (Chong and Speicher, 2001; Chong et al., 2002). In some situations, the need to perform HPLC separations can be circumvented by the use of the mass spectrometer to simultaneously separate and analyze the unfractionated digests (see Strategic Planning and below). Partial reduction and alkylation strategies In cases where protein domains are extensively cross-linked by multiple disulfide bonds, the partial reduction and alkylation strategy (see Alternate Protocol 2) introduced by Gary (1993) is often used to selectively reduce one or two disulfide bonds at low pH with
TCEP while leaving the remaining disulfides unmodified. The partially reduced disulfides are rapidly alkylated with iodoacetamide and separated for characterization by mass spectrometry (UNIT 16.2) and Edman sequencing (UNIT 11.10). Two major advantages of this approach are that accessible cleavage sites are not required and the low pH reduction reaction with TCEP minimizes disulfide scrambling. In a variation of the partial reduction and alkylation method, nascent thiols of partially reduced proteins are cyanylated with the reagent 2-nitro-5-thiocyanobenzoic acid (NTCB) at pH 8, or with 1-cyano-4dimethylamino-pyridinium tetrafluoroborate (CDAP) at pH 3 (Wu et al., 1996; Wu and Watson, 1997). Both cyanylating reagents are able to react with TCEP quantitatively, which is useful to terminate the partial reduction reaction (Schnaible et al., 2002a). Cyanylation with CDAP is preferred as the reaction can be carried out at low pH where disulfide scrambling is minimized. The cyanylated cysteines are susceptible to cleavage at the N-terminal side of the residues at alkaline pH (Jacobson et al., 1973). Following cleavage of the cyanylated cysteines and complete reduction of the remaining disulfide linkages, mass spectrometric analysis allows the identification of the initially reduced disulfide bonds (Wu and Watson, 1997). The advantages of this method are that the reactions are performed at low pH and the ability to cleave at the cyanylated cysteines avoids the need for separate proteolytic/chemical cleavage steps; however, after cleavage, the cyanylated cysteine is converted to a cyclic derivative (Jacobson et al., 1973), resulting in peptides with blocked N termini that are refractory to Edman sequencing. Therefore, peptide identity can only be determined by use of mass spectrometric approaches. MALDI mass spectrometry In addition to reducing agents, disulfide bonds can undergo spontaneous dissociation when analyzed by MALDI mass spectrometry (Patterson and Katta, 1994). This observation has led to mass spectrometry being used to directly analyze HPLC fractions of digested proteins for disulfide-containing peptides without the need for chemical reduction (Qin and Chait, 1997; Schnaible et al., 2002a,b). In addition to producing the constituent peptides, fragmentation of the disulfide bonds by MALDI in-source decay also produces various combinations of
Chemical Analysis
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disulfide-linked peptides which can be used for successful determination of disulfide linkages. The identity of the peptides and location of the disulfide bonds can subsequently be determined by tandem MS approaches. The mass spectrometric approach is comparatively faster and requires relatively small quantities of samples, making it the method of choice when sample is limited.
Critical Parameters and Troubleshooting
Determination of Disulfide-Bond Linkages in Proteins
Disulfide scrambling pH. In disulfide mapping, the most critical concern is disulfide scrambling. Disulfide scrambling occurs readily at pH ≥8, when the protein conformation is perturbed by chaotropic reagents or other nonphysiological conditions. This pH dependency is due to conversion of thiol groups (pKa 8.3) to thiolate anions (S− ). The nucleophilic thiolate anion can attack the sulfur of another disulfide bond, forming a new “scrambled” disulfide bond. The other sulfur is released as a new thiol/thiolate which can continue the exchange with other disulfide bonds. The exchange process occurs more readily when the protein contains free cysteines which can provide the catalytic source of thiolate anions, but it also occurs when proteins whose cysteines are all participating in disulfide bonds are exposed to denaturants and mild reducing conditions at alkaline pH. Since the exchange mechanism is caused by the thiolate anion, the process can be reduced by a factor of ten for each pH unit below the thiol pKa. For example, with trypsin digestion, a >30-fold reduction in disulfide exchange can be achieved by performing the digest at pH 6.5 instead of the optimum, pH 8. Therefore, acidic conditions are always preferred in disulfide mapping. See Strategic Planning for more information on the pH issue with regard to sample preparation and choice of cleavage agents. Alkylation. As mentioned above, proteins with both disulfide bonds and free cysteines are more susceptible to disulfide scrambling, especially under denaturing conditions. To overcome this problem, free cysteines can be alkylated prior to protein denaturation (Yen et al., 2000; Zhang et al., 2002). For this purpose, Nethylmaleimide (NEM) or maleimide derivatives are useful alkylating agents that have been used successfully at acidic pH, from 3.0 to 6.5 (Bures et al., 1998; Yen et al., 2000; van den Hooven et al., 2001; Schnaible et al., 2002a; Zhang et al., 2002). Alternatively, the free cys-
teines can be cyanylated with CDAP, which works well at pH 3.0 where disulfide scrambling is minimized (Wu and Watson, 1998); however, cyanylated cysteines can undergo undesired cleavage at basic pH, such as those used during Edman degradation. Peptides with cyanylated cysteines are best analyzed by mass spectrometric approaches (Wu and Watson, 1998; Schnaible et al., 2002a). Following alkylation, the alkylated protein can be purified by gel filtration with mobile phase containing denaturing agent. In addition to minimizing disulfide scrambling, alkylation will also prevent the highly reactive free SH group from undergoing oxidation to form disulfide or carbamylation when urea is used as a protein denaturant (Lippincott and Apostol, 1999). Partial reduction and alkylation. Among the protocols presented in this unit, disulfide scrambling is most likely to occur in the partial reduction and alkylation protocol during the alkylation step with iodoacetamide (Gary, 1993). In the authors experience, disulfide scrambling is usually minor when the protocol conditions are followed, and the disulfideexchanged peptides can be easily identified by comparing the RP-HPLC chromatograms of the alkylated and nonalkylated partial reduction products (see Alternate Protocol 2). In addition, multiple isoforms are often obtained that can be related to the same disulfide linkage and this redundancy increases the confidence of disulfide assignments; however, if disulfide scrambling is significant, the alkylation step can be performed with NEM instead of iodoacetamide (Bures et al., 1998; van den Hooven et al., 2001; Schnaible et al., 2002a). As mentioned above, NEM, although a bulkier reagent, has been reported to work at the low pH of 3, where disulfide scrambling is minimal. Cleavage agents Very often, due to the multitude of disulfide bonds, the protein of interest cannot be cleaved readily with specific proteases to yield satisfactory amounts of proteolytic fragments, even though denaturing agents are used. The problem can be aggravated in some cases with the use of proteolytic cleavages at suboptimum pH conditions. If this occurs, the authors suggest using chemical cleavages such as CNBr (see Alternate Protocol 1) to provide initial fragmentation (Chong et al., 2002). Alternatively, limited cleavage with broad specificity agents such as pepsin can be used (Roszmusz et al., 2001); however, as mentioned above (see Strategic Planning), the use of broad specificity
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Current Protocols in Protein Science
cleavage agents can complicate the separation and identification of the fragments. An alternative to chemical or broad specificity cleavages is the partial reduction and alkylation procedure (Bures et al., 1998; Schnaible et al., 2002b). If partial reduction and alkylation is performed prior to cleavage, the protein of interest is first denatured to ensure that the reducing agent will ideally have similar access to all disulfide bonds. This will result in the generation of a maximum number of isoforms that can be used to locate and confirm disulfide bonds. Because some chemical reagents used for cleavage and modification of proteins are highly reactive, it is important to keep in mind that prolonged incubations can often result in undesirable artifactual modifications or cleavages. Therefore, do not incubate reactions longer than required and remove or inactivate the reagent promptly after the reaction has been completed. Commonly observed modifications are oxidation of methionine and tryptophan residues, carbamylation of lysines and cysteine residues in the presence of urea, and conversion of N-terminal glutamine to pyroglutamate. With iodoacetamide, extended reaction can lead to undesirable alkylation of lysine, histidine, and methionine residues. These modifications increase the heterogeneity of the cleavage products which will complicate peptide separation and data analysis as well as reduce the yields of desired products. In addition, formation of N-terminal pyroglutamate will make peptides refractory to Edman sequencing. If this happens, the N-terminal blocked residue can be removed using pyroglutamate aminopeptidase (Chong and Speicher, 2001).
linked peptides that are <20 residues. With a protein of ∼25 kDa, most of the peptides produced can easily be resolved by RP-HPLC with good recovery. A starting amount of 10 nmols should yield time point analyses of disulfide-linked peptides that are at least 200 pmol, which is usually sufficient for downstream analysis strategies including additional cleavages or partial reduction and alkylation. Peptides in the range of 10 fmol/µl to 1 pmol/µl are usually sufficient for MALDI-MS analysis (UNIT 16.2), whereas at least 10 pmol peptides is preferred for automated Edman sequencing (UNIT 11.10).
Time Considerations The time needed to complete the disulfide mapping depends largely on the complexity and number of disulfide bonds. In general, chemical and proteolytic cleavage usually requires an overnight incubation. An RPHPLC gradient of ∼1 hr is normally sufficient. With control and blanks included, several hours may be required. MALDI-MS analysis coupled with data analysis will require one or two days. Edman sequencing will also require a day. In general, the entire Basic Protocol will require 1 to 2 weeks, while the Alternate Protocols 1 and 2 require ∼3 days. The Support Protocol can be completed within a day. Overall, assignment of all disulfide linkages in a 30-kDa protein containing six disulfides is likely to be a 3- to 12-month project due to the multiple step strategies required and the fact that some methods need to be optimized and a few methods will not work satisfactory even after optimization.
Literature Cited Anticipated Results The protocols detailed in this unit have been successfully used to determine the disulfide linkages of a number of proteins. Cleavage of proteins to generate fragments suitable for disulfide assignment is the most critical step. With a starting amount of 5 to 10 nmol, the whole procedure can usually be successfully performed without much problem. More protein may be required if it contains a large number of disulfide bonds, or it is very large and requires CNBr cleavage. Since determination of disulfide linkages is performed using recombinant proteins, protein availability is not a concern. Digestion with specific proteases, such as trypsin, will normally generate disulfidelinked peptides that are <50 residues and un-
Brown, J.R. and Hartley, B.S. 1966. Location of disulfide bridges by diagonal paper electrophoresis. Biochem. J. 101:214-228. Chong, J.M. and Speicher, D.W. 2001. Determination of disulfide bond assignments and Nglycosylation sites of the human gastrointestinal carcinoma antigen GA733-2 (CO17-1A, EGP, KS1-4, KSA, and Ep-CAM). J. Biol. Chem. 276:5804-5813. Chong, J M., Uren, A., Rubin, J.S., and Speicher, D.W. 2002. Disulfide bond assignments of secreted Frizzled-related protein-1 provide insights about Frizzled homology and netrin modules. J. Biol. Chem. 277:5134-5144. Clauser, K.R., Baker, P., and Burlingame, A.L. 1999. Role of accurate mass measurement (±10 ppm) in protein identification strategies employing MS or MS/MS and database searching. Anal. Chem. 71:2871-2882. Bures, E.J., Hui, J.O., Young, Y., Chow, D.T., Katta, V., Rohde, M.F., Zeni, L., Rosenfeld, R.D.,
Chemical Analysis
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Stark, K.L., and Haniu, M. 1998. Determination of disulfide structure in agouti-related protein (AGRP) by stepwise reduction and alkylation. Biochemistry 37:12172-12177. Gary, W.R. 1993. Disulfide structures of highly bridged peptides: A new strategy for analysis. Protein Sci. 2:1732-1748. Jacobson, G.R., Schaffer, M.H., Stark, G.R., and Vanaman, T.C. 1973. Specific chemical cleavage in high yield at the amino peptide bonds of cysteine and cystine residues. J. Biol. Chem. 248:6583-6591. Jones, M.D., Patterson, S.D., and Lu, H.S. 1998. Determination of disulfide bonds in highly bridged disulfide-linked peptides by matrixassisted laser desorption/ionization mass spectrometry with post0source decay. Anal. Chem. 70:136-143. Leal, W.S., Nikonova, L., and Peng, G. 1999. Disulfide structure of the pheromone binding protein from the silkworm moth, Bombyx mori. F.E.B.S. Lett. 464:85-90. Lippincott, J. and Apostol, I. 1999. Carbamylation of cysteine: A potential artifact in peptide mapping of hemoglobins in the presence of urea. Anal. Biochem. 267:57-64. Morris, H.R. and Pucci, P. 1985. A new method for rapid assignment of S-S bridges in proteins. Biochem. Biophys. Res. Commun. 126:11221128. Patterson, S.D. and Katta, V. 1994. Prompt fragmentation of disulfide-linked peptides during matrixassisted laser desorption ionization mass spectrometry. Anal. Chem. 66:3727-3732. Pitt, J.J., Da Silva, E., and Gorman, J.J. 2000. Determination of the disulfide bond arrangement of Newcastle disease hemagglutinin neuraminidase. J. Biol. Chem. 275:6469-6478. Qin, J., and Chait, B.T. 1997. Identification and characterization of posttranslational modifications of proteins by MALDI ion trap mass spectrometry. Anal. Chem. 69:4002-4009. Roszmusz, E., Patthy, A., Trexler, M., and Patthy, L. 2001. Localization of disulfide bonds in the frizzled module of Ror1 receptor tyrosine kinase. J. Biol. Chem. 276:18485-18490.
Schnaible, V., Wefing, S., Resemann, A., Suckau, D., Bucker, A., Wolf-Kummeth, S., and Hoffmann, D. 2002b. Screening for disulfide bonds in proteins by MALDI in-source decay and LIFTTOF/TOF-MS. Anal. Chem. 74:4980-4988. van den Hooven, H.W., van den Burg, H.A., Vossen, P., Boeren, S., de Wit, P.J.G.M., and Vervoort, J. 2001. Disulfide bond structure of the AVR9 elicitor of the fungal tomato pathogen Cladosporium fulvum: Evidence for a cystine knot. Biochemistry 40:3458-3466. Wu, J. and Watson, J.T. 1997. A novel methodology for assignment of disulfide bond pairings in proteins. Protein Sci. 6:391-398. Wu, J., and Watson, J.T. 1998. Optimization of the cleavage reaction for cyanylated cysteinyl proteins for efficient and simplified mass mapping. Anal. Biochem. 258:268-276. Wu, J., Gage, D.A., and Watson, J.T. 1996. A strategy to locate cysteine residues in proteins by specific chemical cleavage followed by matrix-assisted laser desorption/ionization timeof-flight mass spectrometry. Anal. Biochem. 235:161-174. Yen, T.-Y., Joshi, R.K., Yan, H., Seto, N.O.L., Palcic, M.M., and Macher, B.A. 2000. Characterization of cysteine residues and disulfide bonds in proteins by liquid chromatography/electrospray ionization tandem mass spectrometry. J. Mass Spectrom. 35:990-1002. Zhang, W., Marzilli, L.A., Rouse, J.C., and Czupryn, M.J. 2002. Complete disulfide bond assignment of a recombinant immunoglobulin G4 monoclonal antibody. Anal. Biochem. 311:19. Zhou, Z. and Smith, D.L. 1990. Assignment of disulfide bonds in proteins by partial acid hydrolysis and mass spectrometry. J. Protein Chem. 9:523-532.
Contributed by Hsin-Yao Tang and David W. Speicher The Wistar Institute Philadelphia, Pennsylvania
Schnaible, V., Wefing, S., Bucker, A., WolfKummeth, S., and Hoffmann, D. 2002a. Partial reduction and two-step modification of proteins for identification of disulfide bonds. Anal. Chem. 74:2386-2393.
Determination of Disulfide-Bond Linkages in Proteins
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CHAPTER 12 Post-Translational Modification: Glycosylation INTRODUCTION
T
his chapter is devoted to detecting and characterizing glycosyl moieties that occur as post-translation additions to proteins. These additions are of two major types: (1) N-glycosylation, which potentially occurs as an attachment to the Asn side chain of proteins containing the sequon AsnXSer/Thr (where X is any amino acid except Pro) or (2) O-glycosylation, which begins with the addition of a N-acetylgalactosamine to the oxygen of specific Ser or Thr side chains of folded proteins; however, increasing data indicate that N-acetylglucosamine addition is also a common modification of this type. There is some indication that the latter modification may play an important regulatory function, as its addition to proteins is often reciprocal with phosphorylation.
UNIT 12.1 provides the nomenclature employed and a general description of the known types
of animal cell glycoconjugates, including not only glycoproteins, but also glycolipids and glycosaminoglycans. The fact that many membrane-bound proteins are anchored via a terminal glycosyl phosphatidyl-inositol (GPI) moiety is also noted. Detailed methods for determining if a membrane-bound protein is GPI-anchored are provided in UNIT 12.5. Such proteins are usually detected by their susceptibility to phospholipase cleavage (Basic Protocol 2 and Alternate Protocol 2), but can also be detected by susceptibility to chemical cleavage with nitrous acid (Basic Protocol 3). The remainder of Chapter 12 provides methods for detecting the presence of N- and O-linked glycosylation and for determining the structure of these glycosyl units at various levels of sophistication. UNIT 12.2 describes procedures for detecting glycoconjugates by tagging them through radiolabeling with glycoconjugate precursors, including [3H] or [14C]monosaccharides, [35S]sulfate, [3H]acetate or [32P]orthophosphate. An alternate protocol describes pulse or pulse-chase labeling with deficient medium. UNIT 12.3 provides methods for inhibiting N-bound glycosylation in cultured cells. Such procedures can be used to classify glycoproteins and to examine the role of glycosylation in protein conformation and function. UNIT 12.4 describes the use of endoglycosidases and glycosamidases to remove N-linked oligosaccharides from proteins. Susceptibility to these enzymes is used to detect the presence of N-linked oligosaccharides and their stage of maturation in bioprocessing experiments. UNIT 12.8 concentrates on techniques for the detection and analyses of proteins modified by O-N-acetylglucosamine, including the analysis of enzymes responsible for the addition and removal of this moiety. UNITS 12.6 and 12.7 provide state-of-the-art methods for sequencing the oligosaccharide side
chain components of glycoproteins. The strategy described in these units was developed at the Glycobiology Institute in Oxford. The technology is sensitive and precise, but does not require sophisticated instrumentation and expertise. The protocols provided enable the glycosylation of 5 to 30 µg of glycoprotein to be analyzed either directly from SDS-PAGE gels (N-linked sugars) or after subsequent hydrazinolysis of the glycoproteins (O-linked sugars). The strategy is based on HPLC technology (UNIT 12.6), which can be used alone or combined with mass spectrometric techniques (UNIT 12.7). John E. Coligan
Post-Translational Modification: Glycosylation
Contributed by John E. Coligan Current Protocols in Protein Science (2001) 12.0.1
12.0.1
Copyright © 2001 by John Wiley & Sons, Inc.
Supplement 26
Overview of Glycoconjugate Analysis The modern revolution in molecular biology has been driven to a large extent by advances in methods for analysis and manipulation of DNA, RNA, and proteins. Although oligosaccharides (sugar chains) are also major macromolecules of the typical cell, they did not initially share in this molecular revolution. The reasons for this are to a large extent technical. Whereas DNA, RNA, and proteins are linear polymers that can usually be directly sequenced, oligosaccharides show substantially more complexity, having branching and anomeric configurations (α and β linkages). Thus, whereas three amino acids or nucleotides can be combined into six possible sequences, three hexose monosaccharides can theoretically generate 1056 possible oligosaccharides. Also, the syntheses of DNA, RNA, and proteins are template-driven, and the sequence of one can generally be predicted from that of another. In contrast, the biosynthesis of oligosaccharides, termed glycosylation, is extremely complex, is not template-driven, varies among different cell types, and cannot be easily predicted from simple rules. It is clear that oligosaccharides have important, albeit varied, effects upon the biosynthesis, folding, solubility, stability, subcellular trafficking, turnover, and half-life of the molecules to which they are attached. These are matters of great importance to the cell biologist, protein chemist, biotechnologist, and pharmacologist. On the other hand, the successful growth of several glycosylation mutants as permanent tissue culture cell lines indicates that the precise structure of many oligosaccharides is not critical for the growth and viability of a single cell in the protected environment of the culture dish. Thus, until recently, it was possible for many researchers working with in vitro single-cell systems to ignore the existence of oligosaccharides. However, with the increasing emphasis on studying cell-cell interactions in normal development, tissue morphogenesis, immune reactions, and pathological conditions such as cancer and inflammation, the study of oligosaccharide structure and biosynthesis has become very important. The term “glycobiology” has found acceptance for denoting studies of the biology of glycoconjugates in both simple and complex systems. Many technical advances have occurred in the analysis of oligosaccharides, making it now feasible to study them in detail. These
advances include the development of sensitive and specific assays and the availability of numerous purified enzymes (glycosidases) with high degrees of specificity. In spite of all these recent advances, many analytical techniques in glycobiology have remained in the domain of the few laboratories that specialize in the study of oligosaccharides. Likewise, published compendia of carbohydrate methods are designed mainly for use by experts. This chapter attempts to place some of this technology within easy reach of any laboratory with basic capabilities in biochemistry and molecular biology. The techniques described here include modern versions of timehonored methods and recently developed methods, both of which have widespread applications. However, it is important to emphasize that the protocols presented here are by no means comprehensive. Rather, they serve as a starting point for the uninitiated scientist who wishes to explore the structure, biosynthesis, and biology of oligosaccharide chains. In most cases, further analysis using more sophisticated techniques will be required to obtain final and definitive results. Nonetheless, armed with results obtained using the techniques described here, the average researcher can make intelligent decisions about the need for such further analyses. The appearance of many commercial kits for analysis of glycoconjugates is another sign that the technology has arrived and that many laboratories have developed an interest in glycobiology. It is worth noting that although some of these kits are designed to simplify the use of well-established techniques, others employ methodologies that have been newly developed by the companies themselves. Experience with the latter methodologies in academic scientific laboratories may be limited, and the techniques in question may therefore not be represented in this chapter. However, although the admonition caveat emptor is appropriate, some kits may well become useful adjuncts to the methods presented here. Different types of glycosylation are perhaps best defined by the nature of the linkage region of the oligosaccharide to a lipid or protein (Fig. 12.1.1). Although the linkage regions of these molecules are unique, the sugar chains frequently tend to share common types of outer sequences. It is important to note that this chapter deals only with the major forms of glycosyla-
Contributed by Ajit Varki, Hudson H. Freeze, and Adriana E. Manzi Current Protocols in Protein Science (1995) 12.1.1-12.1.8 Copyright © 2000 by John Wiley & Sons, Inc.
UNIT 12.1
Post-Translational Modification: Glycosylation
12.1.1 CPPS
glycosaminoglycan chains
S
Ser - O
S
S
S S
NS
P
S
S
S
O - Ser
NS
N-linked chains O-linked chain
glycophospholipid anchor
Ac
glycosphingolipid
Etn P
P S
Ac
O Ser/Thr
N Asn
NH2 inositol
N Asn
P
OUTSIDE
INSIDE
O-linked GIcNAc
= = = = = =
glucose mannose galactose N-acetylglucosamine N-acetylgalactosamine polypeptide
= = = = = =
fucose xylose sialic acid glucuronic acid iduronic acid lipid
O Ser Ac P S NS NH2 Acy
= = = = = =
O-acetyl pyrophosphate O-sulfate N-sulfate N-free amino group O-acyl group
Figure 12.1.1 Common oligosaccharide linkage regions on animal cell glycoconjugates. The most common types of oligosaccharides found in animal glycoconjugates are shown, with an emphasis upon the linkage region between the oligosaccharide and the protein or lipid. Other rarer types of linkage regions and free oligosaccharides that can exist naturally (e.g., hyaluronan) are not shown.
Overview of Glycoconjugate Analysis
tion found in higher-animal glycoconjugates— N-acetylglucosamine (GlcNAc)-N-Asn-linked, N-acetylgalactosamine (GalNAc)-O-Ser/Thrlinked oligosaccharides on glycoproteins, xylose-O-Ser-linked glycosaminoglycans on proteoglycans, ceramide-linked glycosphingolipids, phosphatidylinositol-linked glycophospholipid anchors, and O-linked N-acetylglucosamine (GlcNAc-O-Ser). These structures are depicted in Figure 12.1.1. Other less common forms of glycosylation may need to be considered, especially if prior literature suggests their existence in a given situation. Examples of rarer sugar chains include (1) O-linked
glucose, mannose, and fucose; (2) N-linked glucose; (3) glucosyl-hydroxylysine; and (4) GlcNAc-P-Ser (phosphoglycosylation). Likewise, this chapter deals only with the most common forms of shared outer sequences (e.g., sialylated and fucosylated lactosamines, p oly lactosamines, O-glycosaminoglycan chains, and blood group sequences), and not with rarer sequences (e.g., bisecting xylose residues, N-linked glycosaminoglycans, and βlinked GalNAc residues on N-linked oligosaccharides). For further information, the reader is directed to the Key References section at the end of this unit.
12.1.2 Current Protocols in Protein Science
STEREOCHEMISTRY AND DIAGRAMMATIC REPRESENTATION OF MONOSACCHARIDES AND OLIGOSACCHARIDES Basic Stereochemistry of Monosaccharides In addition to this discussion, the reader is referred to Allen and Kisalius (1992) for a more detailed discussion of the principles of carbohydrate structure. The italicized terms in the discussion below are defined in the glossary at the end of this unit. Glyceraldehyde, the simplest monosaccharide, has a single chiral (asymmetric) carbon (C-2). Therefore, it is a chiral molecule that shows optical isomerism; it can exist in the form of two nonsuperimposable mirror images called enantiomers (see Fig. 12.1.2). These enantiomers have identical physical properties, except for the direction of rotation of the plane of polarized light (−, left hand; +, right hand). Historically, the (+)-glyceraldehyde was arbitrarily assigned the prefix D (for dextrorotatory), and the (−)-glyceraldehyde, the prefix L (for levorotatory). The pair of enantiomers also have identical chemical properties, except toward optically active reagents. This fact is particularly important in biological systems, because most enzymes and the compounds they work on are optically active. The configuration of the highest-numbered asymmetric carbon atom in the carbon chain of a higher monosaccharide is determined by comparison with the configuration of the chiral (or asymmetric) carbon of glyceraldehyde. Thus, a prefix D- is added to the name of each monosaccharide having the configuration of D-glyceraldehyde at the highest-numbered asymmetric carbon, and the prefix L- to those having the configuration of L-glyceraldehyde
CHO
Diagrammatic Representations of Monosaccharides Monosaccharides are conventionally presented using the Fisher projection or the Haworth representation. Some examples are shown in Figure 12.1.4.
Figure 12.1.2 Enantiomers of glyceraldehyde.
CHO OH
H
at the highest-numbered asymmetric carbon. The direction in which the compound rotates the plane of polarized light needs to be determined experimentally, and is indicated by a − or + sign, in parentheses, immediately after the prefix D or L (e.g., D-(+)-glucose). In biological systems, where stereochemical specificity is the rule, D molecules may be completely active and L molecules completely inert, or vice versa. All known naturally occurring monosaccharides in animal cells are in the D configuration, except for fucose and iduronic acid, which are in the L configuration. Each aldose sugar is usually present in a cyclic structure, because the hemiacetal produced by reaction of the aldehyde group at C-1 with the hydroxyl group at C-5 gives a sixmembered ring, called a pyranose (see Fig. 12.1.3). When five-membered rings are formed by reaction of the C-1 aldehyde with the C-4 hydroxyl, they are called furanoses. Hexoses commonly form pyranosidic rings, and pentoses form furanosidic rings, although the reverse is possible. Similarly, ketoses (e.g., sialic acids) can form hemiketals by reaction of the keto group at C-2 with the hydroxyl group at C-6. These ring forms are the rule in intact oligosaccharides. The formation of the ring produces an additional chiral center at C-1 (or C-2 for keto sugars). Thus, two additional isomers are possible, which are called anomers. These anomers are designated α and β. Because glycosidic linkages (see above) occur via anomeric centers, they are called α linkages and β linkages.
CH2OH
D -(+)- glyceraldehyde
HO
H
CH2OH
L -(–)- glyceraldehyde
Post-Translational Modification: Glycosylation
12.1.3 Current Protocols in Protein Science
Fisher projections H
1
α
OH
H
1
β HO
O
C
C OH HO
C
OH
O
HO
OH
HO
HO
HO
HO OH
H
H
1
H
D CH2OH
CH2OH 6
D CH2OH
6
α - D -galactopyranose
O
6
β -D -galactopyranose
6
6
CH2OH D
O
HO
CH2OH D 1
α
OH
O
HO OH
1
OH
β
OH OH
OH Haworth representations
Figure 12.1.3 Formation of cyclic structures: open and ring forms of galactose. The first and last carbon in each chain have been numbered as 1 and 6, respectively. The spatial arrangement of groups that determine the series (D or L) and the anomericity (α and β) of the monosaccharides is indicated by dotted rectangles.
The Fisher projection The carbon chain of a monosaccharide is written vertically with carbon atom number 1 at the top. The horizontal lines represent bonds projecting out from the plane of the paper, whereas the vertical lines represent bonds projecting behind the plane of the paper. When the hydroxyl group at the highest-numbered asymmetric carbon is on the right, the monosaccharide belongs to the D series. When the hydroxyl group at the highest-numbered asymmetric carbon is on the left, the monosaccharide belongs to the L series.
Overview of Glycoconjugate Analysis
The Haworth representation The six-membered ring is represented with the oxygen at the upper right corner, approximately perpendicular to the plane of the paper, and with the groups attached to the carbons above or below the ring. All groups that appear to the right in a Fisher projection appear below the plane of the ring in a Haworth repre-
sentation. The D-series hexoses have the carboxymethyl group (C-6) above the ring. The reverse is true for the L series. Therefore, the only difference betwen the α and β anomers of a given hexose would be the relative position of the hydroxyl and the hydrogen at C-1. In the α anomer, the hydrogen is below the ring for the D series (above the ring for the L series); in the β anomer the hydrogen is above the ring for the D series (below it for the L series).
Conformation of Monosaccharides in Solution The Haworth representation does not depict the real conformation of these molecules. The preferred conformation in solution of the sixmembered ring monosaccharides is what is known as a chair conformation. In this conformation, each different group can adopt either an equatorial or axial position. There are two possible chair conformations called conformers, that exist in equilibrium.
12.1.4 Current Protocols in Protein Science
Fisher projections
Haworth representations
2-acetamido-2-deoxy-D -glucose N-acetyl-D -glucosamine 1
6
CHO
CH2OH D
NHAc
O
HO
H, OH
OH
OH
1
HO
OH D
NHAc CH2OH 6
pyranosidic hemiacetal (mixture of α and β anomers)
Open-chain form
L O CH3
1
CHO L HO
OH
HO
HO
α
HO
HO
O
6
CH3
OH
OH
OH
H, OH 1
L O
HO
CH3
HO
L CH3 6
β
HO OH
HO OH
6-deoxy-L -galactose L-fucose
Figure 12.1.4 Fisher projections and Haworth representations of monosaccharides. The first and last carbon in each chain have been numbered as 1 and 6, respectively. The spatial arrangement of groups that determine the series (D or L) of the monosaccharides, and the anomericity (α or β), is indicated with dotted rectangles.
The position of this equilibrium differs from one monosaccharide to another depending on the relative positions of hydroxyl groups or other substituents. The preferred conformation is that with the lowest number of bulky groups in an axial position (Fig. 12.1.5).
of the component monosaccharides. Thus, the simple tetrasaccharide commonly called sialylLewisx (attached to an underlying oligosaccharide, R) would be written as follows:
Formulas for Representation of Oligosaccharide Chains
In more common practice, the D and L configurations and the nature of ring structures are assumed, and the formula is written in one of the ways shown in Figure 12.1.6.
Full and correct representation of the formula of an oligosaccharide chain requires notation describing the complete stereochemistry
α-D-Neup5Ac-2,3β-D-Galp1,4(α-L-Fucp1,3)β-D-GlcpNAc-R
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12.1.5 Current Protocols in Protein Science
CH2OH CH2OH OH
HO
HO
OH
O
HO
OH OH
HO
α-D-mannopyranose
Figure 12.1.5 D-Mannose chair conformation.
Neu5Acα2,3Galβ1,4(Fucα1,3)GlcNAcβ1–R Neu5Acα2–3Galpβ1–4GlcNAcβ1–R 3 1 Fucα
Figure 12.1.6 Two common short forms for denoting oligosaccharide structures. The tetrasaccharide known as sialyl-Lewisx, is shown attached to an underlying oligosaccharide, R.
Higher-Order Structures in Oligosaccharides
epimers two monosaccharides differing only in the configuration of a single chiral carbon.
The presence of monosaccharides with different types of glycosidic linkages in oligo- and polysaccharides creates further complexity, causing the development of secondary and tertiary structures in these molecules. The approximate shape adopted in solution by a given carbohydrate chain can be predicted according to the type of glycosidic linkages involved. These higher-order structures may be critical for the biological roles of glycoconjugates.
Fuc (L-fucose) type of deoxyhexose (see also types of monosaccharides below).
GLOSSARY The following definitions are presented for the terms most commonly used throughout this chapter. aldose monosaccharide with a carbonyl group at the end of the carbon chain (aldehyde group); the carbonyl group is assigned the lowest possible number (i.e., carbon 1).
Overview of Glycoconjugate Analysis
Gal (D-galactose) type of hexose (see also types of monosaccharides below). GalNAc (N-acetyl-D-galactosamine) type of hexosamine (see also types of monosaccharides below). ganglioside anionic glycolipid containing one or more units of sialic acid. Glc (D-glucose) type of hexose (see also types of monosaccharides below). GlcNAc (N-acetyl-D-glucosamine) type of hexosamine (see also types of monosaccharides below). glyceraldehyde simplest monosaccharide (an aldotriose; i.e., containing three carbon atoms).
carbohydrates polyhydroxyaldehydes or polyhydroxyketones, or compounds that can be hydrolyzed to them.
glycoconjugate natural compound in which one or more monosaccharide or oligosaccharide units are covalently linked to a noncarbohydrate moiety.
diastereoisomers compounds with identical formulas that have a different spatial distribution of atoms (e.g., galactose and mannose).
GlUA or GlA (D-glucuronic acid) type of uronic acid (see also types of monosaccharides below).
enantiomers nonsuperimposable mirror images of any compound (e.g., D- and L-glucose).
12.1.6 Current Protocols in Protein Science
glycolipid or glycosphingolipid oligosaccharide attached via glucose or galactose to the terminal primary hydroxyl group of ceramide, which is composed of a long-chain base (i.e., sphingosine) and a fatty acid. Glycolipids can be neutral or anionic (negatively charged).
which is in turn β-linked to a chitobiosyl group and (2) a chitobiosyl group that is β-linked to the asparagine amide nitrogen. The N-linked oligosaccharides can be divided into three main classes: high-mannose type, complex type, and hybrid type.
glycophospholipid anchor glycan bridge between phosphatidylinositol and a phosphoethanolamine in amide linkage to the C terminus of a protein; constitutes the sole membrane anchor for such proteins.
Neu5Ac (N-acetyl-D-neuraminic acid)
glycoprotein glycoconjugate in which a protein carries one or more oligosaccharide chains covalently attached to a polypeptide backbone via N-GlcNAc- or O-GalNAc-linkages. glycosaminoglycan linear copolymers of disaccharide repeating units, each composed of a hexosamine and a hexose or hexuronic acid; these are the oligosaccharide chains that define the proteoglycans. The type of disaccharide unit can define the glycosaminoglycans as chondroitin or dermatan sulfate, heparan or heparin sulfate, hyaluronic acid, and keratan sulfate. The glycosaminoglycans (except hyaluronic acid) also contain sulfate esters substituting either hydroxyl or amino groups (N- or O-sulfate groups). glycosidic linkage linkage of a monosaccharide to another residue via the anomeric hydroxyl group. IdUA or IdA (L-iduronic acid) type of uronic acid (see also types of monosaccharides below). ketose monosaccharide with a carbonyl group in an inner carbon; the carbonyl group is assigned the lowest possible number (i.e., carbon 2). Man (D-mannose) type of hexose (see also types of monosaccharides below). monosaccharide carbohydrate that cannot be hydrolyzed into simpler units (see also types of monosaccharides below). mucin large glycoproteins that contain many (up to several hundred) O-GalNAc-linked oligosaccharide chains that are often closely spaced. N-linked oligosaccharide oligosaccharide covalently linked to an asparagine residue of a polypeptide chain in the consensus sequence -Asn-X-Ser/Thr. Although there are many different kinds of N-linked oligosaccharides, they have certain common features: (1) a common core pentasaccharide containing two mannosyl residues α-linked to a third mannosyl unit,
type of sialic acid (Sia; see also types of monosaccharides below). O-linked oligosaccharide oligosaccharide linked to the polypeptide via N-acetylgalactosamine (GalNAc) to serine or threonine. Note that other types of O-linked oligosaccharides also exist (e.g., O-GlcNAc) but that the O-GalNAc linkage, being the most well-known, is often described by this generic term. oligosaccharide branched or linear chain of monosaccharides attached to one another via glycosidic linkages (see also polysaccharides below). polylactosaminoglycan long chain of repeating units of the disaccharide β-Gal(1-4)GlcNAc. These chains may be modified by sialylation, fucosylation, or branching. When sulfated, they are called keratan sulfate (see glycosaminoglycans above). polysaccharides branched or linear chain of monosaccharides attached to each other via glycosidic linkages that usually contain repetitive sequences. The number of monosaccharide residues that represents the limit between an oligosaccharide and a polysaccharide is not defined. A tetrafucosylated, sialylated tetraantennary carbohydrate moiety containing eighteen monosaccharide units, present in a glycoprotein, is considered an oligosaccharide. On the other hand, a carbohydrate composed of eighteen glucose residues linked β-(1-4) present in a plant extract is considered a polysaccharide. polysialic acid homopolymer of sialic acid selectively expressed on a few vertebrate proteins and on the capsular polysaccharides of certain pathogenic bacteria. proteoglycan glycoconjugate having one or m ore O-xylose-linked glycosaminoglycan chains (rather than N-GlcNAc- or O-GalNAclinked oligosaccharides) linked to protein. The distinction from a glycoprotein is otherwise arbitrary, because some proteoglycans can have both glycosaminoglycan chains and N- or Olinked oligosaccharides attached to them. reducing sugar sugar that undergoes typical reactions of aldehydes (e.g., is able to reduce
Post-Translational Modification: Glycosylation
12.1.7 Current Protocols in Protein Science
Supplement 4
Ag+ or Cu++). Mono-, oligo-, or polysaccharides can be reducing sugars when the aldehyde group in the terminal monosaccharide residue is not involved in a glycosidic linkage. saccharide modifications hydroxyl groups of different monosaccharides can be subject to phosphorylation, acetylation, sulfation, methylation, or fatty acylation. Amino groups can be free, N-acetylated, or N-sulfated. Carboxyl groups are occasionally subject to lactonization to nearby hydroxyl groups. Sia (sialic acid) generic name for a family of acidic nine-carbon monosaccharides (e.g., Neu5Ac). types of monosaccharides monosaccharides may have a carbonyl group at the end of the carbon chain (aldehyde group) or in an inner carbon (ketone group). The carbonyl group is assigned the lowest possible number, e.g., carbon 1 (C-1) for the aldehyde group; carbon 2 (C-2) for the most common ketone groups. These two types are named aldoses and ketoses accordingly. The simplest monosaccharide is glyceraldehyde (see Fig. 12.1.2), an aldotriose (i.e., containing three carbon atoms). Natural aldoses with different number of carbon atoms in their chain are named accordingly (e.g., aldohexoses, containing six carbon atoms). Two monosaccharides differing only in the configuration of a single chiral carbon are called epimers. For example, glucose and galactose are epimers of each other at C-4. The common monosaccharides present in animal glycoconjugates are (1) deoxyhexoses (e.g., L-Fuc); (2) hexosamines, usually N-acetylated (e.g., DGalNAc and D-GlcNAc); (3) hexoses (e.g., DGlc, D-Gal, and D-Man); (4) pentoses (e.g., D-Xyl); (5) sialic acids (Sia; e.g., Neu5Ac); and (6) uronic acids (e.g., D-GlA and L-IdA). Xyl (D-xylose) type of pentose (see also types of monosaccharides above). For further detailed information on the analysis of oligosaccharides, the reader is referred to the specific literature cited in the individual protocol units. In addition, the following sources can be used for general information, for details on specific methods, and for many additional methods that are not included in this chapter.
LITERATURE CITED Overview of Glycoconjugate Analysis
Allen, H.J. and Kisalius, E.C. (eds.). 1992. Glycoconjugates: Composition, Structure, and Function. Marcel Dekker, New York.
KEY REFERENCES Beeley, J.G. 1985. Glycoprotein and Proteoglycan Techniques. Elsevier/North-Holland, Amsterdam and New York. Chaplin, M.F. and Kennedy, J.F. (eds.). 1987. Carbohydrate Analysis: A Practical Approach. IRL Press, Oxford. Clamp, J.R., Bhatti, T., and Chambers, R.E. 1971. The determination of carbohydrates in biological materials by gas-liquid chromatography. Methods Biochem. Anal. 19:229-344. Cummings, R.D., Merkle, R.K., and Stults, N.L. 1989. Separation and analysis of glycoprotein oligosaccharides. Methods Cell Biol. 32:141183. Dell, A. 1987. F.A.B.-mass spectrometry of carbohydrates. Adv. Carbohydr. Chem. Biochem. 45:19-72. Esko, J.D. 1991. Genetic analysis of proteoglycan structure, function and metabolism. Curr. Opin. Cell Biol. 3:805-816. Fleischer, S. and Fleischer, B. (eds.). 1983. Biomembranes, Part L. Methods Enzymol. Vol. 98. Fukuda, M. (ed.). 1993. Glycobiology: A Practical Approach. IRL Press, Oxford. Ginsburg, V. (ed.). 1972-1989. Complex Carbohydrates, Parts B-F. Methods Enzymol. Vols. 28, 50, 83, 138 and 179. Hart, G. and Lennarz, W. 1993. Guide to Techniques in Glycobiology. Methods Enzymol. Vol. 230. Lane, D.G. and Lindahl, U. (eds.). 1990. Heparin: Chemical and Biological Properties. CRC Press, Boca Raton, Fla. McCloskey, J.A. (ed.). 1990. Mass Spectrometry. Methods Enzymol. Vol. 193. Stanley, P. 1992. Glycosylation engineering. Glycobiology 2:99-107. Varki, A. 1991. Radioactive tracer techniques in the sequencing of glycoprotein oligosaccharides. FASEB J. 5:226-235. Vliegenthart, J.F.G., Dorland, L., and van Halbeek, H. 1983. NMR spectroscopy of carbohydrates. Adv. Carbohydr. Chem. Biochem. 41:209-378. Whistler, R. and Wolfram, F. (eds.). 1968-1980. Methods in Carbohydrate Chemistry, Vols I– VIII. Academic Press, San Diego.
Contributed by Ajit Varki University of California San Diego La Jolla, California Hudson H. Freeze La Jolla Cancer Research Foundation La Jolla, California Adriana E. Manzi University of California San Diego La Jolla, California
12.1.8 Supplement 4
Current Protocols in Protein Science
Metabolic Radiolabeling of Animal Cell Glycoconjugates
UNIT 12.2
Useful information about glycoconjugates can be obtained by labeling their aglycone (noncarbohydrate) portions—e.g., labeling proteins with radioactive amino acids—then using techniques described elsewhere in this chapter to infer the presence, type, and nature of oligosaccharide chains. This unit describes metabolic labeling techniques that provide more specific information about the structure, sequence, and distribution of the sugar chains of glycoconjugates. Following metabolic labeling, the radioactive glycoconjugate of interest is isolated, individual glycosylation sites are identified and separated if necessary, and the labeled oligosaccharides are subjected to structural analysis. Although these techniques provide less information than would complete sequencing of the sugar chains, the partial structural information derived is sufficient for many purposes. Metabolic labeling is simple and easy to perform, and requires no sophisticated instrumentation other than a scintillation counter. In addition, purification of the glycoconjugate to radiometric homogeneity is sufficient for further analysis (i.e., other contaminating molecules that are not labeled do not have to be removed). Important practical considerations for metabolic labeling experiments include selecting the type of experiment and the labeled precursor, understanding the specificity of labeling, and maximizing uptake and incorporation (see Critical Parameters). Before proceeding with experiments to label glycoconjugates metabolically, labeling conditions should be evaluated to ensure that cell viability and metabolism are not altered, and optimized for uptake and incorporation of the precursor (this should be done for each combination of precursor and cell line; see Basic Protocol and Alternate Protocol 1). In the Basic Protocol, cells in culture are grown through several population doublings in complete medium supplemented with radiolabeled glycoconjugate precursors to reach a steady-state level of incorporation. In the alternate protocols, cells are cultured for a short period of time in a deficient medium that contains a high concentration of radiolabeled precursor. A pulse or pulse-chase labeling procedure (Alternate Protocol 1) can be used to analyze precursor-product relationships. With sequential pulse-labeling (Alternate Protocol 2), it is possible to obtain quantities of labeled glycoconjugates while using a minimum amount of labeled precursor by using the same batch of medium to pulse-label a series of cultures. A support protocol describes the preparation of multiply deficient medium (MDM) for use in making appropriate deficient media. NOTE: Conventional protocols for handling, monitoring, shielding, and disposing of radioactivity and radioactive waste (APPENDIX 2B) and for tissue culture of cells and sterile handling of media should be followed throughout these protocols. STEADY-STATE LABELING WITH RADIOACTIVE PRECURSORS When studying glycoconjugates in an established tissue culture cell line, molecules of interest can be labeled for structural characterization. In the following protocol, cells in culture are grown to a steady-state level of incorporation in complete medium supplemented with radiolabeled glycoconjugate precursor.
BASIC PROTOCOL
Optimizing Conditions An attempt should be made to label the molecules as close as possible to the metabolic steady state—i.e., a constant level of radioactivity per mass unit of a given monosaccharide in all glycoconjugates in the cell—under conditions of normal growth. In practice, Contributed by Sandra Diaz and Ajit Varki Current Protocols in Protein Science (1995) 12.2.1-12.2.15 Copyright © 2000 by John Wiley & Sons, Inc.
Post-Translational Modification: Glycosylation
12.2.1 CPPS
this is somewhat difficult to achieve with most cell types. The maximum possible number of cells should be grown for the longest possible period of time in the minimum possible volume of complete medium supplemented with the maximum possible amount of radioactive label. Before labeling is attempted, the following growth characteristics should be determined for the cell type of interest: (1) population doubling time, (2) maximum degree of dilution upon splitting that is compatible with proper regrowth, (3) maximum cell density compatible with healthy growth and metabolism (confluence), and (4) minimum volume of medium that will sustain regrowth from a full split to confluence. If these parameters are properly defined, growth from full split to confluence will allow at least three population doublings during the labeling period for most cell lines. In some cases, however (e.g., with very slowly growing cell lines, lines requiring frequent medium changes, or cells that cannot be diluted to low density when splitting), this may not be feasible. Materials Radioactive precursor: 3H- or 14C-labeled monosaccharide, [35S]sulfate, [3H]acetate, or [32P]orthophosphate; at highest available specific activity Complete tissue culture medium appropriate for long-term growth of tissue culture cell line, supplemented as necessary Established tissue culture cell line, either suspension or monolayer PBS, pH 7.2, (APPENDIX 2E), ice cold Disposable 50-ml vacuum-suction filter device: filter flask fitted with 0.22-µm filter Disposable 0.22-µm filter attached to sterile plastic syringe, both with Luer-Lok fittings Tissue culture plates or flasks Screw-cap centrifuge tubes Tabletop centrifuges, at room temperature and 4°C 1. If necessary, dry radioactive precursor (in a ventilated hood) to completely remove any organic solvents. Dissolve in a small volume of complete tissue culture medium. 2. Filter sterilize radioactive medium, using a standard disposable 50-ml vacuum suction filter device (filter flask fitted with a 0.22-µm filter) for volumes ≥10 ml or a 0.22-µm filter attached directly to the tip of a disposable sterile syringe for smaller volumes. Wash out the original container with additional medium and use this fluid to wash through the filter (this maximizes recovery of label). CAUTION: Dispose of the radioactive filter and/or syringe appropriately. The radioactive precursor may be available in a sterile aqueous medium, ready for use. However, such preparations are more expensive, and after repeated opening of such packages, filtering is recommended to ensure continued sterility.
3. Add additional medium if necessary to bring the volume to the amount needed for the experiment. Warm the medium in an incubator or water bath to the temperature at which labeling will be done. 4. Using a sterile pipet tip, aliquot a small amount of radioactive medium and save for scintillation counting in step 8.
Metabolic Radiolabeling of Animal Cell Glycoconjugates
5. Split monolayer cells according to standard method (e.g., detach by trypsinization), resuspend in radioactive medium, and plate onto tissue culture plates. For suspension cultures, dilute directly in radioactive medium or concentrate as follows: place culture in an appropriately sized screw-cap centrifuge tube and centrifuge in a tabletop centrifuge 5 min at ∼500 × g, room temperature. Discard supernatant and resuspend
12.2.2 Current Protocols in Protein Science
cell pellet in radioactive medium. Incubate under standard conditions for the cell line being studied. The number of cells needed, and therefore the culture vessels and volume of medium used, will depend on the growth characteristics of the particular cell line being studied and should be optimized as described in the protocol introduction.
6. When the cells reach maximum growth (confluence or late-log phase), chill culture on ice and harvest cells, scraping monolayer cells from plate using a rubber policeman or cell scraper. Pellet cells 5 min at ∼500 × g, 4°C. Decant the labeled medium and save an aliquot for scintillation counting in step 8. If the glycoconjugates in the conditioned medium are to be studied, the medium should be filtered through a 0.22-µm filter to remove any cell debris remaining after the centrifugation.
7. Wash cell pellet twice in a >50-fold excess of ice-cold PBS, pH 7.2. At this stage, the labeled cell pellet can be frozen at −20°C to −80°C for later analysis if desired.
8. Using a scintillation counter, determine the efficiency of incorporation by measuring the radioactivity incorporated into an aliquot of the cells and the sample of conditioned medium from step 6. As a control, measure the radioactivity of the sample of sterile radioactive medium from step 4. It may be necessary to dilute the medium to get an accurate count.
PULSE OR PULSE-CHASE LABELING WITH RADIOACTIVE PRECURSORS
ALTERNATE PROTOCOL 1
If steady-state labeling (see Basic Protocol) does not yield sufficient incorporation of label into the glycoconjugate of interest, it may be desirable to provide labeled precursor in the absence of, or with decreased amounts of, the unlabeled form. The pulse procedure described in this protocol—in which cells are cultured briefly in deficient medium supplemented with a radioactive precursor—may be used in these cases. Alternatively, the procedure can be expanded into a pulse-chase by adding a chase of nonradioactive medium; this permits brief labeling of the glycoconjugate of interest to establish precursor-product relationships. These procedures are most useful for labeling monosaccharides that compete with glucose for uptake—i.e., Gal, Glc, Man, and GlcNH2 (see Commentary). They are of limited value for monosaccharides that are not usually taken up efficiently by cells—i.e., GlcNAc, GalNAc, ManNac, Neu5Ac, Fuc, Xyl, and GlcA—unless very large quantities of such labeled molecules can be used for pulse labeling. Because pulse labelings are typically done for a short time (e.g., minutes or hours), cells are usually used in a nearly confluent state, to maximize uptake and incorporation. Optimizing Conditions Before the experiment is performed, several pilot experiments should be carried out to determine the optimal conditions for labeling. Determining base incorporation level and the effect of glucose. MDM (multiply deficient medium) appropriate for growing cultures of the cell line of interest should be reconstituted (see Support Protocol) to 100% levels of all components except the one being presented as a radiolabeled precursor or competing molecule (e.g., glucose). If required by the cells, dialyzed serum should be added to the appropriate final concentration. Finally, radiolabeled precursor should be added. A small-scale pilot labeling should be carried out by culturing cells in the labeled medium and monitoring incorporation of the label into the macromolecule of interest or into whole-cell glycoproteins. If the
Post-Translational Modification: Glycosylation
12.2.3 Current Protocols in Protein Science
radiolabeled precursor is a monosaccharide, the effect of glucose on incorporation of radiolabeled precursor should be assessed by carrying out this pilot experiment in duplicate, using one lot of medium prepared as described and one prepared the same way but without glucose. Determining optimal concentration of radiolabeled precursor. Enough MDM for several pilot experiments should be reconstituted to 100% levels of all components except the one being presented as a radiolabeled precursor. The radiolabeled precursor should then be added and the medium divided into aliquots. The aliquots should be supplemented with an unlabeled 100× stock solution of the same precursor to yield a series of media containing different concentrations of the unlabeled precursor (e.g., 0%, 5%, 10%, 20%, 50%, and 100% of the concentration in normal medium). These should be used for small-scale pilot labelings of the cell line of interest for defined periods of time and incorporation of label should be monitored as described above. Incorporation should be plotted against the total precursor concentration. The point at which the curve breaks— i.e., where the percentage of label incorporated is markedly decreased by further addition of unlabeled compound—is the point at which the precursor concentration is no longer limiting for biosynthetic reactions (Fig. 12.2.1). The optimal concentration of unlabeled precursor will be a little higher than the break point for the curve, where unlabeled precursor is not limiting but incorporation is still good.
35
SO4 incorporated (cpm)
Checking cell growth at optimal precursor concentration. Reconstituted MDM containing radioactive precursor plus the optimal concentration of unlabeled precursor should be prepared and cells should be grown in this medium for the desired period of time. Incorporation of radiolabeled precursor should be monitored and linearity of uptake and incorporation of label over time should be assessed. Measures of cell growth, stability, and general biosynthetic capability—e.g., cell counts and trypan blue exclusion (UNIT 5.5) and radioactive amino acid incorporation—should also be assayed to determine if reducing the concentration of the precursor into this range has any detrimental effect upon the
0
20
40
60
Sulfate concentration (µM)
Metabolic Radiolabeling of Animal Cell Glycoconjugates
Figure 12.2.1 Incorporation of radiolabeled sulfates into macromolecules in cells labeled in sulfate-free medium and the effect of adding increasing amounts of unlabeled sulfate. The arrow indicates the point at which the sulfate is no longer limiting.
12.2.4 Current Protocols in Protein Science
cells. It is sometimes necessary to reach a compromise between the opposing factors of lowered concentration, labeling time, and cell viability. An attempt should be made to determine the lowest concentration of the unlabeled precursor that can be used for the desired period of time without affecting cell growth, viability, and biosynthesis of macromolecules in general. It should be kept in mind that the use of partially deficient medium may selectively alter glycoconjugate makeup (e.g., lowering sulfate concentration too much can result in undersulfation of glycosaminoglycans in some cell types). Additional Materials (also see Basic Protocol) Multiply deficient medium (MDM; see Support Protocol and Table 12.2.1), supplemented as appropriate Fetal bovine serum (FBS; GIBCO/BRL), dialyzed (APPENDIX 3B) against sterile 0.15 M NaCl (glucose concentration ∼250 µM)
Table 12.2.1
Composition of Multiply Deficient Medium (MDM)
Component
Final concentration in complete medium
CaCl2 KCl MgCl2b NaCl NaH2PO4⋅H2O Phenol red Sodium pyruvate L-Alanine L-Arginine L-Asparagine L-Aspartic acid L-Cysteine L-Glutamic acid L-Glutamine L-Glycine L-Histidine L-Isoleucine L-Leucine L-Lysine L-Methionine L-Phenylalanine L-Proline L-Serine L-Threonine L-Tryptophan L-Tyrosine L-Valine MEM vitamins (mixture)
200 mg/l 400 mg/l 75 mg/l 6800 mg/l 140 mg/l 10 mg/l 110 mg/l 25 mg/l 126 mg/l 50 mg/l 30 mg/l NONEc 75 mg/l NONEc 50 mg/l 42 mg/l 52 mg/l 52 mg/l 72 mg/l NONEc 32 mg/l 40 mg/l NONEc 48 mg/l 10 mg/l 36 mg/l 46 mg/l 1×
Stocka Salt, reagent grade Salt, reagent grade Salt, reagent grade Salt, reagent grade Salt, reagent grade 10× 100× 100× 100× 100× 100× — 100× — 100× 100× 100× 100× 100× — 100× 100× — 100× 100× 100× 100× 100×
a10× and 100× stock solutions are available commercially (e.g., from GIBCO/BRL). bDo not use MgSO in place of MgCl . 4 2 cNot present in MDM; add as needed for specific experiments.
Post-Translational Modification: Glycosylation
12.2.5 Current Protocols in Protein Science
1. Determine optimal conditions for labeling as described in the protocol introduction. Grow monolayer cells in complete medium to near confluence or suspension cells to late-log phase. 2. Based on the pilot experiments for optimizing conditions described in the protocol introduction, reconstitute MDM to prepare a stock of medium partially deficient in only the precursor of interest. 3. Add an appropriate quantity of radioactive precursor to the reconstituted MDM. Add dialyzed FBS if required for growth of the cell line being studied. Filter sterilize, warm the medium to 37°C, and reserve an aliquot of radioactive medium (see Basic Protocol, steps 2 to 4). 4. For monolayer cells, aspirate culture medium from plate; for suspension cells, microcentrifuge or centrifuge briefly and decant the supernatant medium. Add an appropriate quantity of radioactive labeling medium from step 3. Incubate cells for the desired period of time. Labeling time should be based on the length of time the cells can survive in deficient medium as determined in the optimizing pilot experiment. Save radioactive medium for analysis, if necessary.
5. For pulse labeling, proceed to step 6. For pulse-chase labeling, remove medium as described in step 4, add a chase of nonradioactive complete medium, and incubate for the desired period of time. For a pulse-chase experiment, several vessels of each cell culture should be set up and chased for varying periods of time.
6. Harvest and wash the cells. Determine efficiency of incorporation (see Basic Protocol, steps 6 to 8). ALTERNATE PROTOCOL 2
Metabolic Radiolabeling of Animal Cell Glycoconjugates
SEQUENTIAL PULSE OR PULSE-CHASE LABELING WITH REUSE OF RADIOACTIVELY LABELED MEDIUM In some cases, incorporation of label into the glycoconjugate of interest may be inadequate, even under defined conditions with selectively deficient medium. In addition, prolonged exposure to deficient medium may result in alterations in synthesis of proteins or other macromolecules. In these situations, it may be desirable to expose a series of plates of cells sequentially to a small volume of medium containing a high concentration of label. This also conserves expensive radiolabeled precursor. In this protocol, multiple sets of cell cultures are sequentially pulse-labeled. One set is incubated in labeling medium for the pulse period, then the labeling medium is removed and added to the next set. For a pulse experiment (designed to obtain a sizable quantity of material with a high level of incorporation), the cells are harvested at this point; for a pulse-chase, a chase of nonradioactive complete medium is added to the first set of cultures. This process is repeated until all cells have been labeled (Fig. 12.2.2). Because the incubation period is relatively short (e.g., a few hours), it is assumed that only a very small fraction of the radiolabeled precursor is consumed from the medium during each labeling cycle and that the concentrations of other essential components are not substantially changed. To further conserve labeled medium, some investigators freeze it after limited usage and thaw it to reuse (the pH of thawed medium should be adjusted before use). When reusing medium, experimental results should be interpreted carefully, especially with regard to the specific activity of labeled products and the possibility of metabolic effects of secreted molecules.
12.2.6 Current Protocols in Protein Science
Additional Materials (also see Basic Protocol) Multiply deficient medium (MDM; see Support Protocol and Table 12.2.1), supplemented as appropriate Fetal bovine serum (FBS; GIBCO/BRL), dialyzed (APPENDIX 3B) against sterile 0.15 M NaCl (glucose concentration ∼250 µM) 1. Determine optimal conditions for labeling (see Alternate Protocol 1 introduction). Set up several identical sets of monolayer or suspension cells to be labeled sequentially and grow to confluence or late-log phase. For suspension cultures in which it is possible to use <1 ml labeling medium, cells may be labeled in tightly capped 1.5-ml screw-cap microcentrifuge tubes and incubated at 37°C on a rotating end-over-end mixer.
2. Prepare radioactive labeling medium and label the first set of cell cultures for the appropriate length of time (see Alternate Protocol 1, steps 2 to 4). Labeling time will vary from minutes to hours, depending upon the cell type and the objective of the labeling.
3. For monolayer cells, aspirate the radioactive medium from the first set of cultures; for suspension cells, microcentrifuge briefly (∼10 sec at top speed) and decant the supernatant radioactive medium. Transfer the radioactive medium to the second set of cultures. 4. For pulse labeling, harvest cells from first set of cultures. For pulse-chase labeling, add a chase consisting of an adequate quantity of nonradioactive complete culture medium to the first set of cultures and continue incubating. 5. Repeat transfers until all sets of cells have been labeled. 6. Harvest all plates. For pulse labeling, pool all the pellets together. For pulse-chase labeling, keep the individual cell pellets separate (see Fig. 12.2.2).
TIME: HARVEST PULSE
CHASE CHASE
PULSE
PULSE
*
CHASE PULSE
* *
* Transfer labeling medium to next plate. Add complete medium to labeled cells.
Figure 12.2.2 Scheme for sequential pulse labeling of cells reutilizing radioactive medium.
Post-Translational Modification: Glycosylation
12.2.7 Current Protocols in Protein Science
SUPPORT PROTOCOL
PREPARATION AND SUPPLEMENTATION OF MULTIPLY DEFICIENT MEDIUM (MDM) Some types of selectively deficient media are available commercially at reasonable prices, and some companies will custom-prepare selectively deficient media on request. For a laboratory where labeling experiments with a variety of deficient media are frequently carried out, it is convenient to prepare a basic multiply deficient medium (MDM) that is completely lacking in several commonly studied components. This medium can be used to make up different selectively deficient media as needed. The following MDM, based on α-MEM medium (GIBCO/BRL), supports growth of most tissue culture cells. For labeling experiments, MDM can be reconstituted with 3H- or 14C-labeled monosaccharides, [35S]sulfate, 35S-labeled methionine or cysteine, or [3H]serine. If desired, other specific components can also be omitted and replaced with their radiolabeled forms. NOTE: Tissue culture grade reagents (including distilled, deionized water) should be used in making MDM. To ensure that reagents do not become contaminated by compounds such as endotoxin as portions are being removed from stock bottles, it is important to pour liquids carefully and use disposable spatulas for removing solids. For the same reason, clean glassware must be used; preferably, a set of glassware should be set aside for this purpose. 100× stock solutions of many components can be purchased commercially in sterile form, or may be made up from solids and filter sterilized individually. Additional Materials (also see Basic Protocol) Stock solutions for multiply deficient medium (MDM; Table 12.2.1) 100× stock solutions for reconstituting MDM (Table 12.2.2) 1. Based on the experiment to be performed, determine appropriate components for MDM. 2. Make up MDM according to the ingredient list in Table 12.2.1 by dissolving salts and phenol red one by one in ∼800 ml water, adding sufficient pyruvate, amino acid, and vitamin stocks for a 1× final concentration in 1 liter, and adding water to 1 liter. 3. Filter sterilize. Do not adjust the pH. Freeze 25-ml aliquots in 50-ml tubes at −20°C. The solution will be yellow.
Table 12.2.2 Stock Solutions for Reconstitution of MDMa
Component
Final concentration in complete medium
Stockb
NaHCO3c HEPES⋅HClc Na2SO4 D-Glucose L-Cysteine L-Glutamine L-Methionine L-Serine
1× (2200 mg/l) 20 mM (4.76 g/l) 0.81 mM (115 mg/l) 1× (1000 mg/ml) 1× (100 mg/l) 1× (292 mg/l) 1× (15 mg/l) 1× (25 mg/l)
100× 2 M, pH 7.3 100 mM, sterile 100× 100× 100× 100× 100×
aIndividual components are added to MDM at full strength, lower concentration, or
left out altogether, depending upon the experiment planned.
Metabolic Radiolabeling of Animal Cell Glycoconjugates
b100× stock solutions are available commercially (e.g., from GIBCO/BRL). cUse either NaHCO /CO or HEPES⋅HCl (not both); these control the pH of the 3 2
final medium.
12.2.8 Current Protocols in Protein Science
4. Add appropriate components (from those listed in Table 12.2.2) to reconstitute MDM for the labeling experiment. The following example describes the selective reconstitution of MDM with missing components. It is a recipe for 50 ml of a supplemented MDM for optimal labeling of endothelial cells with [35S]sulfate and [6-3H]glucosamine, which requires a medium with no cysteine, low glucose, and low concentrations of methionine and sulfate (see Roux et al., 1988, for detailed rationale). Stock solutions are as listed in Table 12.2.2. To 50 ml MDM, add:
500 µl 100× NaHCO3 stock solution 500 µl 100× glutamine stock solution 500 µl 100× serine stock solution 50 µl 100× glucose stock solution (200 mg/L final) 10 µl 100 mM inorganic sulfate stock solution (20 µM final) 50 µl 100× methionine stock solution (0.1× final). Just before use, add 0.5 mCi each of [35S]sulfate and [6-3H]glucosamine and mix. Filter sterilize directly into culture vessel. The total volume of reconstituted medium will be slightly more than 50 ml; for practical purposes, this minor discrepancy can be ignored. The medium should be orange-red, indicating the correct pH range.
COMMENTARY Background Information If sufficient quantities of pure molecules (i.e., in the nanomole range) are available, complete sequencing of oligosaccharides is best performed by conventional techniques not requiring radioactivity. However, isolation of sufficient quantities of material may not be practical (e.g., in analysis of biosynthetic intermediates or rare molecules). Alternatively, the biological questions underlying the experiment may be adequately answered by less definitive means of structural analysis. In these cases, metabolic labeling with radioactive sugars or donors that can transfer label to sugars may provide sufficient structural information about the oligosaccharide chains (see Cummings et al., 1989). Sugar chains of glycoconjugates can be successfully labeled by short-term labeling experiments using radioactive precursors (Tabas and Kornfeld, 1980; Goldberg and Kornfeld, 1981; Roux et al., 1988; Muchmore et al., 1989). Although sugar nucleotides are the immediate donors for glycosylation reactions, they cannot be taken up by cultured cells. Hence, metabolic labeling of sugar chains is accomplished by providing the cells with radiolabeled monosaccharides (Yurchenco et al., 1978; Yanagishita et al., 1989; Varki, 1991). These are taken up by the cells, activated to sugar nucleotides, and transported into the Golgi apparatus, where lumenally oriented transferases add the monosaccharides to lumenally oriented ac-
ceptors (Hirschberg and Snider, 1987). The few known exceptions to this topology are cytosolic and nuclear forms of glycosylation (Hart et al., 1989). For oligosaccharides that contain modifications—such as sulfate, acetate, and phosphate ester groups—an alternative metabolic labeling technique is to use a [35S]sulfate, [3H]acetate, or [32P]orthophosphate label, respectively. Although metabolic labeling can provide useful information regarding glycoprotein oligosaccharides, there are significant limitations to its use. First, it is difficult to determine when true steady-state labeling of a cell is reached (as a rule of thumb, 3 to 4 doublings are usually assumed to be sufficient); thus the numerical ratio between labeled glycoconjugates may not reflect steady-state ratios. Second, individual precursors show greatly differing uptake and incorporation in different cell types. Third, almost all labeled precursors are only partially specific for certain monosaccharides, and the degree of this specificity can vary depending on cell type. Fourth, although lowering glucose concentration improves the incorporation of monosaccharides that compete with glucose for uptake, the low glucose supply may also directly affect oligosaccharide precursors in some cell types.
Critical Parameters As diagrammed in Figure 12.2.3, the labeling protocol most suitable for a particular study
Post-Translational Modification: Glycosylation
12.2.9 Current Protocols in Protein Science
determine type of study (structural or biosynthetic)
determine type of glycoconjugate to be labeled
choose labeling protocol
select radiolabeled precursor(s)
check incorporation of radioactivity in complete medium
if insufficient for analysis
if sufficient for analysis
try using more labeled precursor
decrease level of unlabeled precursor in medium
check effects on incorporation of label
check effects on cells and molecule(s) of interest
select optimal concentration of unlabeled precursor
proceed with steady-state, pulse, pulse-chase, or sequential transfer protocols for labeling
Figure 12.2.3 Strategy for planning metabolic labeling of animal cell glycoconjugates.
depends on several different considerations. Some of these arise from the objectives of the study and must be investigated by performing pilot experiments to determine optimal conditions (see Optimizing Conditions sections of the Basic Protocol and Alternate Protocol 1).
Metabolic Radiolabeling of Animal Cell Glycoconjugates
General considerations If the goal of a study is to obtain labeled material for structural identification and partial characterization, the yield of radioactivity should be maximized by either steady-state or pulse labeling. For quantitating the relative mass of a molecule in two different samples or comparing the masses of two molecules in the
same sample, the oligosaccharide should be labeled to constant specific activity—i.e., steady-state distribution (although this may not always be achievable). For establishing precursor-product relationships between molecules, a pulse-chase protocol should be used. If the radioactive precursor is expensive, a sequential procedure may be employed in pulse and pulsechase experiments to minimize the quantity needed. Selecting a labeled precursor Selection of the labeled precursor to be used is based upon several factors, including efficiency of uptake (see section on Factors affect-
12.2.10 Current Protocols in Protein Science
Table 12.2.3
Distribution of Monosaccharides and Modifications in Glycoconjugatesa
Type of glycoconjugate GlycoXyloseGlycophospholipid linked sphingolipid anchor proteoglycan
N-GlcNAclinked glycoprotein
O-GalNAclinked glycoprotein
Man Fuc Gal
+++ ++ +++
− ++ +++
− +/− ++
− + +++
Glc
−
−
++
GlcNAc
+ (precursor) +++
+++
++
++
GalNAc
+/−
+++
++
++
Sia
+++
+++
−
+++
GlcA SO4 Pi esters
+? + Man-6-P
− + −
+++ ++++ Xyl-P
+ + −
O-Acetyl Acyl
Sia-OAc −
Sia-OAc −
− −
Sia-OAc ++++
Precursor
O-linked GlcNAc
+++ − +/− (side chain) −
− − −
+ (free amine) +/− (side chain) +/− (side chain) − − M6P (in core) − +/− (on inositol)
++
−
− − − − − − −
aThe relative distribution of the different monosaccharides and modifications in the commonly occurring glycoconjugates, indicated by a relative scale from + to ++++, is generally valid over many cell types. However, an uncommon monosaccharide may be commonly found in some cell types or glycoconjugate (e.g., most pituitary glycoprotein hormones have GalNAc as a major component of their N-linked oligosaccharides). Other symbols: +?, possible distribution; +/−, variably present; −, not identified to date.
ing uptake and incorporation) and type of glycoconjugate to be labeled. In most cases, the precursor will be a monosaccharide; for an oligosaccharide chain containing one or more ester modifications of sugars, it is also possible to use a precursor such as acetate, sulfate, or orthophosphate. Factors affecting specificity and final distribution. In selecting a monosaccharide precursor, it should be noted that the distribution of monosaccharides among different types of vertebrate oligosaccharides is nonrandom (Table 12.2.3). It is also important to consider the metabolic pathways for uptake, activation, utilization, and interconversion of the various monosaccharides and their nucleotide sugars, which have been studied extensively (Fig. 12.2.4 and Table 12.2.4). The final distribution and specific activity of label from a given radiolabeled monosaccharide can be significantly affected by dilution from an unlabeled compound generated by endogenous pathways, the characteristics of the particular cell type being labeled, and the labeling conditions (Kim and Conrad, 1976; Yurchenco et al., 1978; Yanagishita et al., 1989). The exact position of the tritium label within a monosaccharide can
affect the label’s ultimate fate (Diaz and Varki, 1985). Interconversion between monosaccharides (e.g., conversion to glucose; Fig. 12.2.4), which can be expected to occur with prolonged labeling, will eventually result in labeling of non-oligosaccharide components. An exception is labeling with [2-3H]mannose: this is extremely specific, because conversion of the labeled mannose can yield only [2-3H]fucose or unlabeled fructose-6-phosphate. In the latter case, the label is lost as tritiated water, which is diluted into the pool of cellular water. Thus, in most instances, label from [2-3H]mannose remains confined to mannose and fucose residues, regardless of how long the labeling continues. Factors affecting uptake and incorporation. With radioactive amino acids, high-specific-activity labeling can be obtained by omitting the unlabeled molecule from the medium. Labeling with radioactive sugars or other oligosaccharide precursors, however, is usually less efficient. Thus, incorporation into the glycoconjugate of interest must be optimized empirically, taking into consideration the following factors:
Post-Translational Modification: Glycosylation
12.2.11 Current Protocols in Protein Science
monosaccharides competing with glucose for active transport cannot be taken up very well. Thus, lowering glucose concentration can improve uptake. On the other hand, monosaccharides that do not compete with glucose seem to be taken up only by inefficient noncompetitive, passive mechanisms; thus, glucose concentration has little effect upon their already poor uptake. Experience indicates that glucosamine, galactosamine, galactose, and mannose can compete with glucose for uptake into most cells, whereas N-acetylglucosamine, N-acetylmannosamine, mannosamine, fucose, and xylose do not (Yurchenco et al., 1978). However, members of the glucose transporter family of gene products are tissue-specific in expression; a given cell type may express more than one,
Experimental parameters. Amount of label, concentration of label in the medium, cell number, duration of labeling, and number of cell doublings during labeling are obviously important. The goal is to expose the maximum number of cells to the maximum amount of label in the minimum volume of medium for the longest period of time. Some of these factors are at odds with one another, and the correct balance must be tailored to the particular cell type and the question being investigated. Competition with glucose. When labeling using a radioactive monosaccharide precursor, it is important to determine whether the monosaccharide competes with glucose for uptake. The high concentration of glucose in normal tissue culture medium (∼5 mM) means that
Gal
Gal-1-P
Glc
Glc-6-P
UDP-Gal
Glc-1-P
UDP-Glc GlcA
Man-6-P
Man
Man-1-P
Fruc-6-P
UDP-GlcA
GDP-Man UDP-Xyl
GlcNH2
Xyl
GlcNH2 -6-P
GDP-Fuc GlcNAc-6-P
Fuc CMP-Neu5Gc
CMP-Neu5Ac
Neu5Ac
GlcNAc-1-P ManNAc-6-P
GlcNAc
GalNAc
UDP-GalNAc
UDP-GlcNAc
ManNAc
Neu5Ac
Neu5Ac-9-P
active uptake, competing with glucose passive uptake, not competing with glucose
Metabolic Radiolabeling of Animal Cell Glycoconjugates
Figure 12.2.4 Cytosolic pathways for the interconversion of monosaccharides and their nucleotide sugar forms.
12.2.12 Current Protocols in Protein Science
Table 12.2.4
Cellular Monosaccharides Labeled with Radioactive Precursorsa
Product identified as: [2-3H]Man [3H]Man [3H]Fuc [3H]Gal [3H]GlcA [3H]Glc [3H]GlcNAc [3H]GalNAc [3H]Sia
+++ +++ − − − − − −
Radiolabeled precursor [6-3H]Gal
[1-3H]Gal
[1-3H]Glc
[6-3H]GlcNH2
[6-3H]ManNAc
[6-3H]GalNAc
+/− +/− ++++ − ++ +/− +/− −
+/− +/− ++++ ++ ++ +/− +/− −
++ ++ +++ ++ ++++ ++ ++ ++
+/− +/− +/− − +/− ++++ +++ ++
+/− +/− +/− − +/− ++ + ++++
+/− +/− +/− − +/− +++ ++++ +/−
aThe extent of conversion from the original monosaccharide into minor pathways will vary considerably, depending upon the cell type studied and
the length of the labeling (longer time periods tend to result in more conversion into other monosaccharides). The extent of incorporation is indicated by a relative scale from + to ++++. Other symbols: +/−, variable low-level incorporation; −, not identified to date.
each with distinctive kinetic and stereospecific uptake properties (Gould and Bell, 1990). Thus, each monosaccharide must be tested in the specific cell type under study to see if it competes with glucose. A final point that should be considered is that in some cell types, lipid-linked oligosaccharide precursors may become altered when glucose-free medium is used (Rearick et al., 1981). Other intracellular metabolic factors. Dilution of label by endogenously synthesized monosaccharides, pool size of individual monosaccharides and nucleotides, and flux rates between interconverting pathways can all affect the final specific activity and distribution of the label. If cells are cultured in labeling medium for prolonged periods of time, the medium may become depleted of glucose (Kim and Conrad, 1976; Varki and Kornfeld, 1982). The specific activity of the labeled sugar nucleotides in the cells may then actually rise as the medium glucose concentration falls below the Km of the cellular glucose transporter(s). These factors must be considered in designing the labeling protocol, although they can be controlled only to a limited extent. The goal is to obtain sufficient label in the glycoconjugate of interest without significantly altering the metabolic state of the cell. Metabolic labeling using nonmonosaccharide precursors. Acetylation, sulfation, and phosphorylation can significantly affect the behavior of oligosaccharides both in biological systems and during analysis. Labeled precursors such as [3H]acetate, [35S]sulfate, and [32P]orthophosphate can be used to label these modified oligosaccharides. These precursors are expected to enter a variety of other macro-
molecules, and release or isolation of the labeled oligosaccharides with specific endoglycosidases is often required before further analysis. Double-labeling of both the modified oligosaccharide and the underlying sugar chain (e.g., with a 14C- or 3H-labeled monosaccharide precursor) can be quite useful in monitoring purification and in subsequent structural analysis. With [35S]sulfate, labeling efficiency varies widely with cell type. In many cells, if the endogenous pool of the precursor 3′-phosphoadenosine-5′-phosphosulfate (PAPS) is very small, the specific activity of the exogenously added [35S]sulfate can be practically unaltered inside the cell (Yanagishita et al., 1989). In some cell types, however, the endogenous sulfate pool is constantly diluted by breakdown of the sulfur-containing amino acids methionine and cysteine (Esko et al., 1986; Roux et al., 1988). In this situation, reducing the cysteine and methionine concentrations in the medium can improve incorporation of label into macromolecules (Roux et al., 1988). Sulfate levels can also be affected by certain antibiotics (e.g., gentamycin) that are sulfate salts. Determining the specific activity of incorporated label In most instances, the precise specific activity of each monosaccharide pool need not be determined. If steady-state labeling is attempted, a plateau in the rate of incorporation of label per milligram of cell protein can be taken as a rough indication that a steady state has been reached. In some cases, however, the precise specific activity of a given monosaccharide may be of interest. Discussions of how this can be determined can be found in Kim and Conrad
Post-Translational Modification: Glycosylation
12.2.13 Current Protocols in Protein Science
(1976), Yurchenco et al. (1978), Yanagishita et al. (1989), and Varki (1991). Endogenous glucose is the normal precursor for hexoses and hexosamines (see Fig. 12.2.4). Experimental manipulations that alter the concentration of glucose can thus alter the concentration of the internal pool of other hexoses and hexosamines. Exogenously added labeled hexosamines become diluted within the cell, making specific radioactivity in the cell lower than that of the starting material. With the advent of high-performance anionexchange chromatography with pulsed amperometric detection (HPAE-PAD), measurement of monosaccharide levels in the low picomole range is now possible. Thus, it is now feasible to measure the specific activity of a labeled monosaccharide component of a purified oligosaccharide by acid hydrolysis of a portion of the oligosaccharide (Hardy et al., 1988) followed by HPAE-PAD. High-performance liquid chromatography (HPLC)–based methods for accurate measurement of low quantities of sulfate are also now available.
Troubleshooting Ideally, the radiochemical purity of the precursor should be checked using an appropriate chromatographic procedure. As with all tissue culture work, cells should be checked for mycoplasma infection (Strober, 1991) before the experiments are carried out. If bacterial contamination poses a problem for long-term labeling experiments, appropriate antibiotics should be added to the medium.
Anticipated Results Levels of incorporation will be highly variable, depending upon the cell type, the culture conditions, and the precursor used. If adequate incorporation into the glycoconjugate of interest is obtained, analyses can be carried out.
Time Considerations
Metabolic Radiolabeling of Animal Cell Glycoconjugates
Optimizing conditions for uptake and incorporation of label for each precursor and cell line combination (introductions to the first two protocols) may take one to two weeks, but should not be omitted: the time will be well-spent. The time required for the actual labeling protocols will vary widely depending on the goal of the experiment. Preparation of MDM can take an entire day, especially if some components must be prepared from scratch. If the amount required for a series of experiments is planned ahead of time, however, this need only be done occasionally. Reconstituting MDM takes up to
1 hr (assuming all the stock solutions are already available).
Literature Cited Cummings, R.D., Merkle, R.K., and Stults, N.L. 1989. Separation and analysis of glycoprotein oligosaccharides. Methods Cell Biol. 32:141183. Diaz, S. and Varki, A. 1985. Metabolic labeling of sialic acids in tissue culture cell lines: Methods to identify substituted and modified radioactive neuraminic acids. Anal. Biochem. 150:32-46. Esko, J.D., Elgavish, A., Prasthofer, T., Taylor, W.H., and Weinke, J.L. 1986. Sulfate transport-deficient mutants of Chinese hamster ovary cells. Sulfation of glycosaminoglycans dependent on cysteine. J. Biol. Chem. 261:15725-15733. Goldberg, D. and Kornfeld, S. 1981. The phosphorylation of beta-glucuronidase oligosaccharides in mouse P388D1 cells. J. Biol. Chem. 256:13060-13067. Gould, G.W. and Bell, G.I. 1990. Facilitative glucose transporters: An expanding family. Trends Biochem. Sci. 15:18-23. Hardy, M.R., Townsend, R.R., and Lee, Y.C. 1988. Monosaccharide analysis of glycoconjugates by anion exchange chromatography with pulsed amperometric detection. Anal. Biochem. 170:5462. Hart, G.W., Haltiwanger, R.S., Holt, G.D., and Kelly, W.G. 1989. Glycosylation in the nucleus and cytoplasm. Annu. Rev. Biochem. 58:841874. Hirschberg, C.B. and Snider, M.D. 1987. Topography of glycosylation in the rough endoplasmic reticulum and Golgi apparatus. Annu. Rev. Biochem. 56:63-87. Kim, J.J. and Conrad, E.H. 1976. Kinetics of mucopolysaccharide and glycoprotein biosynthesis by chick embryo chondrocytes. Effect of D-glucose concentration in the culture medium. J. Biol. Chem. 251:6210-6217. Muchmore, E.A., Milewski, M., Varki, A., and Diaz, S. 1989. Biosynthesis of N-glycolylneuraminic acid: The primary site of hydroxylation of N-acetylneuraminic acid is the cytosolic sugar nucleotide pool. J. Biol. Chem. 264:20216-20223. Rearick, J.I., Chapman, A., and Kornfeld, S. 1981. Glucose starvation alters lipid-linked oligosaccharide biosynthesis in Chinese hamster ovary cells. J. Biol. Chem. 256:6255-6261. Roux, L., Holoyda, S., Sunblad, G., Freeze, H.H., and Varki, A. 1988. Sulfated N-linked oligosaccharides in mammalian cells I: Complex-type chains with sialic acids and O-sulfate esters. J. Biol. Chem. 236:8879-8889. Strober, W. Diagnosis and treatment of mycoplasma-contaminated cell cultures. 1991. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. A.3.9-A.3.12.
12.2.14 Current Protocols in Protein Science
Tabas, I. and Kornfeld, S. 1980. Biosynthetic intermediates of beta-glucuronidase contain high mannose oligosaccharides with blocked phosphate residues. J. Biol. Chem. 255:6633-6639.
Yanagishita, M., Salustri, A., and Hascall, V.C. 1989. Specific activity of radiolabeled hexosamines in metabolic labeling experiments. Methods Enzymol. 179:435-445.
Varki, A. 1991. Radioactive tracer techniques in the sequencing of glycoprotein oligosaccharides. FASEB J. 5:226-235.
Yurchenco, P.D., Ceccarini, C., and Atkinson, P.H. 1978. Labeling complex carbohydrates of animal cells with monosaccharides. Methods Enzymol. 50:175-204.
Varki, A. and Kornfeld, S. 1982. The spectrum of anionic oligosaccharides released by endo-β-Nacetylglucosaminidase H from glycoproteins. J. Biol. Chem. 258:2808-2818.
Contributed by Sandra Diaz and Ajit Varki University of California San Diego La Jolla, California
Post-Translational Modification: Glycosylation
12.2.15 Current Protocols in Protein Science
Inhibition of N-Linked Glycosylation Treatment of cells with inhibitors of the enzymes that synthesize N-linked oligosaccharide chains results in production of glycoproteins containing missing or altered chains. This approach is useful for examining potential functional role(s) of this class of oligosaccharides on specific proteins or intact cells. Table 12.3.1 lists the enzymes blocked by each inhibitor and the consequences; Figure 12.3.1 diagrams the early processing steps in the assembly of N-linked oligosaccharides, showing the enzymes blocked by each inhibitor except tunicamycin (which affects an earlier stage). This unit describes the use of these inhibitors to prevent N-linked glycosylation of proteins in cultured cells. First, the optimal concentration of inhibitor for the experiment (i.e., highest nontoxic concentration) is determined by monitoring [35S]methionine incorporation as a measure of protein biosynthesis. The inhibitor’s ability to inhibit oligosaccharide processing is then determined by analyzing cells labeled with [3H]mannose using TCA precipitation or endo H digestion. A Support Protocol details a method for concentrating proteins by acetone precipitation. Although not specifically covered in this unit, these inhibitors can be used to examine specific glycoproteins that have been identified and/or purified (e.g., by immunoprecipitation, immunoblotting) in a scheme established by the investigator. As an example, several of the inhibitors render the oligosaccharide chains on a glycoprotein susceptible to removal by endo H, and the change in size of the glycoprotein is frequently large enough to be detected by SDS-PAGE (UNIT 10.1). Thus, radiolabeled samples (e.g., labeled with [35S]Met, 125I) from inhibitor-treated cells can be purified, and the effects of endo H digestion utilized to determine the effect of the inhibitor on oligasaccharide processing.
UNIT 12.3 BASIC PROTOCOL
NOTE: All media and solutions should be made with Milli-Q-purified (or equivalent) water. All media and equipment coming into contact with cells should be sterile. Culture incubations should be performed in a humidified 37°C, 5% CO2 incubator. Materials Cultured cell line, either adherent or suspension Complete culture medium Inhibitor of N-linked glycosylation (one or more of the following: tunicamycin, deoxynojirimycin, castanospermine, deoxymannojirimycin, or swainsonine; see recipe for inhibitor stock solutions and Table 12.3.2) Solvent used for making inhibitor solution (see recipe) Multiply deficient medium (MDM; UNIT 12.2) supplemented with 200 µM glucose [3H]mannose (5 to 20 Ci/mmol) Phosphate-buffered saline (PBS; APPENDIX 2E), ice cold Lysis buffer (see recipe) Sepharose, activated 0.5 U/ml endoglycosidase H (endo H) and endo H digestion buffer (see recipe) 20% (w/v) SDS (APPENDIX 2E) Sephacryl S-200 column (UNIT 8.3), calibrated and equilibrated in 25 mM ammonium formate/0.1% SDS 25 mM ammonium formate/0.1% (w/v) SDS 24-well tissue culture plate 100-mm tissue culture plates Disposable plastic scraper or rubber policeman Centrifuges and appropriate rotors: low-speed and ultracentrifuge 1.5-ml, 15-ml, or 50-ml conical polypropylene centrifuge tube
Contributed by Leland D. Powell Current Protocols in Protein Science (1995) 12.3.1-12.3.9 Copyright © 2000 by John Wiley & Sons, Inc.
Post-Translational Modification: Glycosylation
12.3.1 CPPS
Table 12.3.1
Inhibitors of Enzymes that Synthesize N-linked Oligosaccharides
Type of inhibitor
Target enzyme
Effective concentration range
Effect on oligosaccharide structure
Glycosylation inhibitor Tunicamycin GlcNAc transferase Prevents assembly of GlcNAc-PP-dolichol and thus assembly of G3M9GlcNAc2-PP-dolichola; glycosylation of Asn residues does not occur; proteins migrate faster on SDS-PAGE and generally show less size heterogeneity; proteins will not shift to a faster mobility on SDS-PAGE after PNGase F treatment Processing inhibitors Deoxynojirimycin Glucosidase I Prevents removal of first glucose residue, Castanospermine and/or II thereby inhibiting any further processing of the oligosaccharide chain; proteins may migrate with a larger or smaller size on SDS-PAGE, depending on extent of processing that normally occurs; sensitive to endo Hb DeoxymannoPrevents removal of mannose residues on the α-mannosidase I jirimycin α1-3 arm of the high mannose structure, thereby blocking the activity of GlcNAc T I and thus α-mannosidase II; proteins generally run as a smaller size on SDS-PAGE, depending on size of the normal oligosaccharide; structures remain sensitive to endo Hc Swainsonine Prevents removal of mannose residues on the α1α-mannosidase II 6 arm of the high mannose structure, preventing the activities of GlcNAc T II and V (which add GlcNAc to the α1-6 mannose residue); addition of GlcNAc, galactose, and sialic acid to the α1-3 mannose can occur normally, although the structures remain sensitive to endo H digestion
0.5-10 µg/ml
0.5-20 mM 1-50 µg/ml
1-5 mM
1-10 µg/ml
aAbbreviations: G M GlcNAc -PP-dolichol, Glc Man GlcNAc-PP-dolichol; GlcNAc T I, II, V, N-acetylglucosaminyltransferase I, II, V; PNGase F, 3 9 2 3 9 peptide N:glycosidase F. bMature processed oligosaccharide structures cannot be cleaved from the protein by endoglycosidase H; structures retaining mannose residues on the α1-6 arm can be removed, generally resulting in a difference in size easily detected by SDS-PAGE. cEndomannosidases can bypass this block.
Additional reagents and equipment for metabolic radiolabeling of glycoconjugates (UNIT 12.2), gel-filtration chromatography (UNIT 8.3), and one-dimensional SDS-PAGE (UNIT 10.1) Determine optimal inhibitor concentration 1. For each inhibitor tested, set up 15 wells of cells in a 24-well tissue culture plate by splitting the cells (using trypsinization for adherent cells or dilution for suspension cells) into 1.8 ml final volume per well of complete medium. Incubate 24 hr. The ratio of splitting should be designed to yield confluent (but not overgrown) cultures by the third day (i.e., 48 hr after splitting).
Inhibition of N-Linked Glycosylation
2. While cells are incubating, make a series of 1:1 (v:v) dilutions of inhibitor(s) as follows. Place the volume of inhibitor stock solution indicated in Table 12.3.2 in a sterile microcentrifuge tube and dilute to 800 µl final volume with complete culture medium. Transfer 400 µl to a second tube and dilute with 400 µl complete culture medium. Repeat for a total of seven tubes. As a control, prepare an identical series of dilutions of the solvent used for making the inhibitor solution.
12.3.2 Current Protocols in Protein Science
Figure 12.3.1 Structures of early processing intermediates in the assembly of N-linked oligosaccharides. Structure A is initially assembled on the lipid carrier dolichol phosphate, a process blocked by tunicamycin, and then transferred to the Asn residue. The initial trimming glucosidases I and II convert A into B; these glucosidases are inhibited by castanospermine and deoxynojirimycin. Deglucosidation is followed by removal of the outer four mannose residues by mannosidase I (inhibited by deoxymannojirimycin) to form C. Removal of these mannose residues is the signal for the first N-acetylglucosaminyltransferase (GlcNAc transferase I) to form D. Structure D, but not C, is a substrate for mannosidase II, resulting in the formation of E, which can be further extended into bi-, tri-, and tetraantennary structures (sialylated and neutral) by a series of N-acetylglucosaminyltransferases, galactosyltransferases, and sialyltransferases plus the appropriate sugar-nucleotide donors. Post-Translational Modification: Glycosylation
12.3.3 Current Protocols in Protein Science
Table 12.3.2
Range of Inhibitor Concentrations for Dilution Series
Inhibitor
Stock solution
Tunicamycin Swainsonine Deoxynojirimycin Deoxymannojirimycin Castanospermine
1 mg/ml 1 mg/ml 400 mM 400 mM 400 mM
Stock added to Concentration first dilution (µl) range 80 80 200 200 200
0.15-10 µg/ml 0.15-10 µg/ml 0.15-10 mM 0.15-10 mM 0.15-10 mM
3. Add 200 µl from each tube (from step 1) to a well of the tissue culture plate containing cells (2 ml final). Add 200 µl complete culture medium alone to the remaining well (as a zero-inhibitor control point). Incubate plate 24 hr. Store unused inhibitor-containing media at 4°C. If the inhibitor is tunicamycin, dilutions should be prepared fresh each day, as it may precipitate out of solution during storage.
4. Perform short-term labeling of cells with [35S]methionine using 0.2 mCi/ml label and 1.8 ml final volume per well. 5. Add to wells the remaining 200 µl of each dilution of inhibitor (from step 2) and incubate a further 4 hr. 6. Harvest cells and determine the incorporation of [35S]methionine into macromolecules by TCA precipitation. Plot label incorporation versus inhibitor concentration. CAUTION: Safety precautions for disposing of radioactive waste should be followed. Comparing incorporation by cells incubated with and without inhibitor will indicate the maximum concentration of inhibitor that does not inhibit protein synthesis. This is the optimal concentration for the following steps.
Label cells in the presence of N-linked glycosylation inhibitor 7. Split a new cell sample into complete medium in a 100-mm tissue culture plate and incubate 24 hr. 8. Wash the cells using the same procedure as employed during labeling (step 4). To each plate, add sufficient MDM to cover the cells (e.g., 5 ml for a 100-mm plate). Add inhibitor to the optimal concentration determined above. Set up a control plate containing the same quantity of stock solution solvent but no inhibitor. Add [3H]mannose to 0.02 to 0.1 mCi/ml. Incubate 4 to 12 hr. The duration of incubation will depend on how well the cells label with [3H]mannose. The amount of [3H]mannose used will depend on the cell line examined. However, for most cells, a single 100-mm plate with 5 ml media incubated with 0.02 mCi/ml [3H]mannose for 6 hr should yield sufficient radioactive material for the analyses described below. “Glucose-free” MDM is described in UNIT 12.2; in fact, cells should not be labeled in the total absence of glucose, as this causes alterations in glycan processing. For this reason, the glucose concentration should be kept above 200 µM. For metabolic labeling, preincubation of the cells with inhibitor is not essential. However, a 24-hr preincubation with inhibitor is recommended for the cell-surface expression of a sufficient population of glycoproteins with altered glycan structures.
Inhibition of N-Linked Glycosylation
9. Harvest the cells by scraping with a disposable scraper or rubber policeman (for adherent cells) or centrifugation (for suspension cells) into ice-cold PBS. CAUTION: Safety precautions for disposing of radioactive waste should be followed.
12.3.4 Current Protocols in Protein Science
Measure incorporated macromolecular radioactivity 10. If tunicamycin was used as the inhibitor, determine the amount of incorporated macromolecular radioactivity by TCA precipitation and proceed to step 18. If another inhibitor was used, carry out steps 11 to 18. The amount of radioactivity incorporated into macromolecules reflects the effect of the inhibitor(s) on oligosaccharide biosynthesis: the presence of inhibitor should block incorporation of [3H]mannose into macromolecules.
11. Wash the cells twice by resuspending the pellet in ice-cold PBS, centrifuging, and removing the supernatant. Resuspend the pellet in lysis buffer (5 × 107 cells/ml) and incubate 1 hr at 4°C. 12. Centrifuge the lysate 10 min at 3000 × g, 4°C, to remove nuclei and save the supernatant. 13. Centrifuge the supernatant 1 hr at 100,000 × g, 4°C, and save the supernatant. Supernatants may also be prepared by microcentrifugation (10,000 × g) for 30 min. The supernatant must be used within several days or stored at −70°C. The length of storage is limited by autoradiolysis and the half-life of the isotope. 3H- and 14C-labeled samples can often be stored frozen for years. Storage of 125I-labeled samples is usually limited to 1 to 2 months because of autoradiolysis. The usefulness of 35S-labeled samples is usually limited to 6 months because of half-life. Repeated freezing and thawing may disrupt antigenic determinants and dissociate some protein complexes, especially those that are noncovalently associated.
14. Divide each sample into two equal aliquots in conical polypropylene centrifuge tubes and precipitate protein with acetone (see Support Protocol). Use 1.5-ml, 15-ml, or 50-ml centrifuge tubes, depending on the quantity of sample (8 vol acetone must be added).
15. Resuspend pellets in 20 to 40 µl endo H digestion buffer per 107 cells and boil 10 min to inactivate endogenous hydrolases. 16. Add 5 µl endo H per 100 µl sample to one tube of each sample pair and incubate both overnight at 37°C. 17. Add 1⁄10 vol of 20% (w/v) SDS to each sample and boil 3 to 5 min to terminate the digestion. 18. Apply one sample at a time to a calibrated Sephacryl S-200 column equilibrated in 25 mM ammonium formate/0.1% SDS, in a sample volume that is <5% of the bed volume. Collect the eluate and use a scintillation counter to measure the radioactivity associated with glycoproteins (eluting in the void volume of the column, Vo) and with free oligosaccharides (eluting after Vo) for each of the four samples. Wash column with 2 to 4 vol 25 mM ammonium formate/0.1% SDS between sample applications. In the absence of endo H digestion, all the radioactivity from both the inhibitor-treated and untreated samples should elute in the Vo. Elution of any radioactive material after the Vo may indicate that the wash steps following acetone precipitation did not completely remove low-molecular-weight contaminants present in the cell lysate. For samples digested with endo H, elution profiles should demonstrate a marked increase in the amount of endo H-releasable radioactivity in samples from inhibitor-treated cells relative to treated cells. For samples not treated with inhibitor, the oligosaccharides cleaved from the peptide backbone by endo H represent immature oligosaccharides still being processed in the Golgi apparatus and a small population of endo H-sensitive structures found on some mature glycoproteins. For samples treated with inhibitor, a much higher percentage of the total radioactivity should be released, as the inhibitors prevent the
Post-Translational Modification: Glycosylation
12.3.5 Current Protocols in Protein Science
processing of the oligosaccharides into mature oligosaccharide structures (Fig. 12.3.1) and the underprocessed oligosaccharides contain more mannose residues than the processed ones. Theoretically, 100% of the radioactivity should be releasable by endo H digestion of the inhibitor-treated samples (see Critical Parameters), although in reality no inhibition is ever complete. To save time, analyze only the first and last tubes of the control cells in this fashion, i.e., those not treated at all and those treated with the highest concentration of the inhibitor solvent (from step 2 above). If there is no difference in the results from these two groups of cells, it is reasonable to assume that those cells treated with lower concentrations of the inhibitor solvent will yield the same results. However, if the solvent is found to be toxic, then the cells exposed to lower concentrations will need to be examined to find the maximally tolerated concentration of inhibitor solvent.
19. If desired, and if methods are available for purifying and identifying a specific glycoprotein from cells radiolabeled with [35S]methionine, examine the effect of a given inhibitor by SDS-PAGE and autoradiography. Each N-linked carbohydrate chain contributes ∼2000 to 4000 Da to a protein’s mass, so synthesis of a protein in the presence of tunicamycin will result in faster mobility on SDS-PAGE. The effect of other inhibitors can be demonstrated by an increased sensitivity to digestion by endo H, which is also detectable as an increase in migration rate in SDS-PAGE. SUPPORT PROTOCOL
ACETONE PRECIPITATION Acetone precipitation is a useful step for concentrating proteins and for exchanging them from one buffer to another. As little as 10 ng of protein can be precipitated successfully. Additional Materials (also see Basic Protocol) 100% acetone (HPLC or ACS grade), −20°C 1. Place the protein solution on ice, add 8 vol acetone (−20°C), mix gently, and precipitate at −20°C overnight. If the sample volume is too large to be conveniently diluted with 8 vol, concentrate it by lyophilizing and resuspending in a smaller volume. If the sample contains NP-40 or Triton X-100 detergent, however, there is a limit to how much it can be concentrated, as high concentrations of detergent (e.g., >1%) will partition out of acetone solutions, appearing as an oily pellet after centrifugation (step 2). If a large amount of protein is present (i.e., a precipitate is readily visible after the acetone is added), it can be precipitated after standing at −20°C for only a few hours instead of overnight. Leaving the material in acetone for longer than 1 day should be avoided, as it may become increasingly difficult to redissolve the protein.
2. Collect the precipitate by centrifuging 15 min at 3000 × g, 4°C (a small pellet should be visible). Carefully invert the tube and pour out the acetone into a clean tube. If the pellet is oily due to the presence of detergent, it should be extracted by adding 85% (v/v) acetone (−20°C), mixing, incubating at −20°C for >1 hr, and recentrifuging.
3. Centrifuge the tube containing the pellet briefly to concentrate the remaining acetone and remove it with a micropipettor. Allow pellet to dry until moist but not powder-dry, and resuspend in an appropriate quantity of the desired solvent.
Inhibition of N-Linked Glycosylation
Care must be taken in drying the pellet, as great difficulty may be encountered in resuspending some proteins if the pellet dries completely. This insolubility depends on both the individual protein and the total amount present.
12.3.6 Current Protocols in Protein Science
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Endo H digestion buffer 15 µl 0.5 M sodium citrate, pH 5.5 80 µl water 1 µl 100 mM PMSF in absolute ethanol Prepare just before use (PMSF is unstable in water) This is sufficient for 5 µl of 0.5 U/ml endo H (100 µl digestion volume). The PMSF prevents proteolysis. Nonionic detergent is not required.
Inhibitor stock solutions Dissolve castanospermine (mol. wt. 189.2), 1-deoxymannojirimycin (mol. wt. 199.6), or 1-deoxynojirimycin (mol. wt. 163.2) in water to 400 mM; dissolve swainsonine (mol. wt. 173.2) in water to 1 mg/ml. Sterilize by filtration through a 0.2-µm filter. Store frozen at −20°C (stable indefinitely). Dissolve tunicamycin (mol. wt. 840) in dimethylsulfoxide (DMSO), dimethylformamide (DMF), 95% ethanol, or 25 mM NaOH to 1 mg/ml. Store frozen at −20°C (stable ∼1 yr). If NaOH is used, store stock solution in a small tube that is tightly capped to prevent adsorption of carbon dioxide, which will lower the pH. Tunicamycin’s solubility in neutral aqueous solution is <1 mg/ml (attempting to make such a solution will produce a visible precipitate).
Lysis buffer 1% Triton X-100 1 mM iodoacetamide (freshly prepared) Aprotinin (0.2 trypsin inhibitor U/ml) 1 mM PMSF (add fresh from 100 mM stock in absolute ethanol) Prepare in TSA solution (see recipe) Tris/saline/azide (TSA) solution 0.01 M Tris⋅Cl, pH 8.0 0.14 M NaCl 0.025% NaN3 CAUTION: Sodium azide (NaN3) is poisonous; wear gloves.
COMMENTARY Background Information Assembly of N-linked oligosaccharide chains involves the sequential action of several glycosidases and glycosyltransferases (reviewed by Hubbard and Ivatt, 1981, and Kobata and Takasaki, 1992). Growth of cells in the presence of inhibitors of these enzymes produces glycoproteins with underprocessed or missing oligosaccharide chains. Depending on the protein, these changes may alter biological functions and/or rates of transport, secretion, or turnover (Elbein, 1987; McDowell and Schwarz, 1988). N-linked glycosylation begins with the assembly of the lipid-linked precursor glucose3-
m an no se 9 N- acety lglu co sam ine 2 - pyr ophosphate-dolichol (Glc3Man9GlcNAc2-PPdolichol; similar to structure A, Fig. 12.3.1). Assembly of this precursor requires N-acetylg l u c o s a m i n e - p y r o p h o s p h a t e - d o l ich o l (GlcNAc-PP-dolichol), whose formation is blocked by tunicamycin (Tkacz and Lampen, 1975). Thus, the presence of tunicamycin results in proteins missing some or all of their N-linked side chains; however, those that are added due to incomplete inhibition are processed normally. Tunicamycin may also inhibit protein synthesis; not all proteins will be affected to the same degree. Tunicamycin also blocks the assembly of type II keratan sulfate
Post-Translational Modification: Glycosylation
12.3.7 Current Protocols in Protein Science
Supplement 4
chains, because these glycans also utilize GlcNAc-PP-dolichol (Hart and Lennarz, 1978). Glycolipid biosynthesis may also be inhibited, although the mechanism underlying this has not been established (Yusuf et al., 1983; Guarnaccia et al., 1987). The glycosidase inhibitors castanospermine, deoxynojirimycin, deoxymannojirimycin, and swainsonine are considerably less toxic than tunicamycin. Castanospermine inhibits glucosidase I, and deoxynojirimycin inhibits both glucosidases I and II (Fig. 12.3.1; Pan et al., 1983; Saunier et al., 1982). In some studies, the metabolic block produced by these two drugs is not complete, yielding a mixture of normally processed and underprocessed structures. More recently, an endo-α-D-mannosidase has been described that cleaves the tetrasaccharide Glc3-Man1 from Glc3Man9GlcNAc2, effectively bypassing the block produced by the glucosidase inhibitors (Lubas and Spiro, 1987). Although the question has not been examined fully, the level of this endo-αD-mannosidase may vary between different cell lines. Deoxymannojirimycin and swainsonine are inhibitors of mannosidases I and II, respectively (Fig. 12.3.1; Fuhrmann et al., 1984; Tulsiani et al., 1982). By inhibiting mannosidase I, deoxymannojirimycin prevents addition of the α1-2 N-acetylglucosamine (GlcNAc) residue required to produce structure D, Figure 12.3.1, which is a necessary step for the action of mannosidase II. Thus, deoxymannojirimycin prevents addition of any GlcNAc residues, blocking galactosylation and sialylation as well. Swainsonine blocks mannosidase II. However, it does not prevent the single GlcNAc residue (structure D, Fig. 12.3.1) from being galactosylated and sialylated, resulting in hybrid structures (Tulsiani and Touster, 1983). Some of the glycosidase inhibitors are also active against lysosomal enzymes, although the significance of this effect in tissue culture cells has been little explored. Deoxynojirimycin can inhibit synthesis of the lipid-linked precursor in cell-free extracts (Romero et al., 1985); this also has not been thoroughly investigated with intact cells. No information is available on the metabolism or half-lives of these inhibitors in cultured cells.
Critical Parameters and Troubleshooting Inhibition of N-Linked Glycosylation
The two most critical variables are duration of exposure and inhibitor concentration. The treatment times suggested in this unit (24 hr)
will permit a cell to replace most of its endogenous glycoproteins with glycoproteins synthesized in the presence of inhibitor. For proteins that turn over slower or faster than average, longer or shorter incubation times may be appropriate. If pulse-chase studies with radiolabeled sugar or amino acid precursors (UNIT 12.2) are being carried out, it is necessary to incubate cells in inhibitor for only 1 to 2 hr before adding label. N-linked glycosylation inhibitors may be toxic to the cells, as indicated by decreased cell viability and/or reduced protein synthesis. The concentration of tunicamycin that can be used is generally limited by its toxicity, which may be considerable in some cell lines (more so with tumor cells than nontransformed cells). Other glycosidase inhibitors are usually not toxic in the concentration ranges suggested in Table 12.3.1 (the highest concentrations listed represent those used in most published studies). It is important to realize that in a given cell line, not all glycoproteins will be affected to the same extent by a given inhibitor. Moreover, it is possible that not all sites on the same glycoprotein will be affected equally. These caveats should be kept in mind when interpreting the results from studies employing these inhibitors. Failure to observe an effect from an inhibitor may indicate that concentration of inhibitor was too low and/or length of exposure was too short. Cell lines differ in their susceptibility to these compounds.
Anticipated Results These experiments will demonstrate the chosen inhibitor’s toxicity, as assessed by protein synthesis, and its ability to alter oligosaccharide biosynthesis, as assessed by total [3H]mannose incorporation (for tunicamycin) or endo H release of the oligosaccharides (for the other inhibitors listed in Table 12.3.1).
Time Considerations Setting up the initial round of cell cultures for determining the optimal inhibitor concentration takes 2 hr, after which they are incubated 24 hr. On the second day, the dilution series of medium containing inhibitor is set up, the cells are transferred to these media (<1 hr), and the plates incubated 24 hr. On the third day, the incubation of the cells with the radiolabeled precursor is set up (∼4 hr); the actual incubation takes 4 to 16 hours. Following incubation, the cells are assayed for radiolabel incorporation by TCA precipitation (2 to 4 hr).
12.3.8 Supplement 4
Current Protocols in Protein Science
The main part of the experiment follows a similar time course if the inhibitor used is tunicamycin. If a different inhibitor is used, analysis of endo H susceptibility by Sephacryl S-200 chromatography will take ∼1 day, including the overnight incubation.
Literature Cited Elbein, A.D. 1987. Inhibitors of the biosynthesis and processing of N-linked oligosaccharide chains. Annu. Rev. Biochem. 56:497-534. Fuhrmann, U., Bause, E., Legler, G., and Ploegh, H. 1984. Novel mannosidase inhibitor blocking conversion of high mannose to complex oligosaccharides. Nature 307:755-758. Guarnaccia, S.P., Shaper, J.H., and Schnaar, R.L. 1987. Tunicamycin inhibits ganglioside biosynthesis in neuronal cells. Proc. Natl. Acad. Sci. U.S.A. 80:1551-1555. Hart, G.W. and Lennarz, W.J. 1978. Effects of tunicamycin on the biosynthesis of glycosaminoglycans by embryonic chick cornea. J. Biol. Chem. 253:5795-5801. Hubbard, S.C. and Ivatt, R.J. 1981. Synthesis and processing of asparagine-linked oligosaccharides. Annu. Rev. Biochem. 50:555-583. Kobata, A. and Takasaki, S. 1992. Structure and biosynthesis of cell surface carbohydrates. In Cell Surface Carbohydrates and Cell Development (M. Fukuda, ed.) pp. 1-24. CRC Press, Boca Raton, Fla.
Romero, P., Friedlander, P., and Herscovics, A. 1985. Deoxynojirimycin inhibits the formation of Glc3Man9GlcNAc2-PP-dolichol in intestinal epithelial cells in cultures. FEBS Lett. 183:2932. Saunier, B., Kilker, R.D., Tkacz, J.S., Quaroni, A., and Herscovics, A. 1982. Inhibition of N-linked complex oligosaccharide formation by 1-deoxynojirimycin, an inhibitor of processing glucosidases. J. Biol. Chem. 257:14155-14161. Tkacz, J.S. and Lampen, J.O. 1975. Tunicamycin inhibition of polyisoprenyl N-acetylglucosaminyl pyrophosphate formation in calf liver microsomes. Biochem. Biophys. Res. Commun. 65:248-55. Tulsiani, D.R.P., Harris, T.M., and Touster, O. 1982. Swainsonine inhibits the biosynthesis of complex glycoproteins by inhibition of Golgi mannosidase II. J. Biol. Chem. 257:7936-7939. Tulsiani, D.R.P. and Touster, O. 1983. Swainsonine causes the production of hybrid glycoproteins by human skin fibroblasts and rat liver Golgi preparations. J. Biol. Chem. 258:7578-7585. Yusuf, H.K.M., Pohlentz, G., Schwarzmann, G., and Sandhoff, K. 1983. Ganglioside biosynthesis in Golgi apparatus of rat liver. Eur. J. Biochem. 134:47-54.
Key References Elbein, A.D. 1987. See above. Useful general review.
Lubas, W.A. and Spiro, R.G. 1987. Golgi endo-αD-mannosidase from rat liver, a novel N-linked carbohydrate unit processing enzyme. J. Biol. Chem. 262:3775-3781.
McDowell, W. and Schwarz, R.T. 1988. See above.
McDowell, W. and Schwarz, R.T. 1988. Dissecting glycoprotein biosynthesis by the use of specific inhibitors. Biochimie 70:1535-1549.
Good review of early work on processing pathway.
Pan, Y.T., Hori, H., Saul, R., Sanford, B.A., Molyneux, R.J., and Elbein, A.D., 1983. Castanospermine inhibits the processing of the oligosaccharide portion of the influenza viral hemagglutinin. Biochemistry 22:3975-3984.
Good review of recent work on some of the complexities in oligosaccharide processing.
Useful general review. Hubbard, S.C. and Ivatt, R.J. 1981. See above.
Kobata, A. and Takasaki, S. 1992. See above.
Contributed by Leland D. Powell University of California San Diego La Jolla, California
Post-Translational Modification: Glycosylation
12.3.9 Current Protocols in Protein Science
Endoglycosidase and Glycoamidase Release of N-Linked Oligosaccharides
UNIT 12.4
Carbohydrate chain modifications are often used to monitor glycoprotein movement through the secretory pathway. This is because stepwise sugar-chain processing is unidirectional and generally corresponds to the forward or anterograde movement of proteins. This unit offers a group of techniques that will help analyze the general structure of carbohydrate chains on a protein and, therefore, oligosaccharide processing mileposts. The minimum requirements are that the protein can be labeled metabolically (UNIT 3.7) and immunoprecipitated (UNIT 9.8) and clearly seen on a gel or blot (UNITS 10.5-10.8). The sugar chains themselves are not analyzed, but their presence and structure are inferred from gel mobility differences after one or more enzymatic digestions. This approach is most often used in combination with [35S]Met pulse-chase metabolic labeling protocols, but they can be applied to any suitably labeled protein (e.g., biotinylated or 125I-labeled). As the oligosaccharide chains mature, they become either sensitive or resistant to highly specific glycosidases. Some of these enzymes cleave intact oligosaccharide chains from the protein—e.g., endo H, endo F2, endo F3, peptide:N-glycosidase F (PNGase F), endo D, and O-glycosidase. Others strip only terminal sugars (e.g., sialidase) or degrade a selected portion of the chain (e.g., endo-β-galactosidase). The techniques can be adapted to count the number of N-linked oligosaccharide chains on a protein. One unusual protease, O-sialoglycoprotease, degrades only proteins containing tight clusters of Olinked sialylated sugar chains. These techniques work best on average size proteins (<100 kDa) that contain a few percent carbohydrate by weight, where a gel shift of 1 kDa can be seen. A summary of the enzymes and their applications is shown in Table 12.4.1. This unit provides information on how to measure changes in carbohydrate structure and how these changes relate to protein trafficking. Fortunately, the techniques are independent of mechanistic views, although it should be borne in mind that the organization and distribution of many of these indicator enzymes are cell type dependent. Table 12.4.1 Enzymes Described in This Unit
Enzyme
Indications and uses
Endo D
Transient appearance of highly processed, sensitive forms prior to addition of GlcNAc by GlcNAc transferase I Presence of biantennary chains ± core fucose Endo F2 Endo F3 Presence of core fucosylated biantennary α chains and/or triantennary chains ± core fucosylation Endo-β-galactosidase Presence of polylactosamines Endo H Conversion of high mannose to complex type N-linked chains O-Glycosidase Presence of Galβ1,3GalNAc-α-Thr/Ser O-linked chains PNGase F Presence of N-linked chains cleaves; nearly all N-linked chains; only enzyme that cleaves tetrantennary chains Sialidase Acquisition of sialic acids O-Sialoglycoprotease Presence of mucin-like proteins with cluster of sialylated oligosaccharides aAbbreviation: TGN, trans-Golgi network.
Contributed by Hudson H. Freeze Current Protocols in Protein Science (2000) 12.4.1-12.4.26 Copyright © 2000 by John Wiley & Sons, Inc.
Monitorsa Cis to medial Golgi
Medial Golgi Medial Golgi
Trans-Golgi and TGN Cis to medial Golgi Cis to medial Golgi Medial Golgi
Trans-Golgi and TGN Trans-Golgi and TGN Post-Translational Modification: Glycosylation
12.4.1 Supplement 22
The starting material for these protocols is assumed to be [35S]Met-labeled, immunoprecipitated protein bound to ∼20 µl of protein A–Sepharose beads (as described in UNIT 9.8). The trace amount of protein is eluted by heating in a small volume of 0.1% SDS, diluted in the appropriate buffer and then digested with one or more enzymes in a small volume. The digest is analyzed on an appropriate SDS-PAGE system that can detect a 1- to 2-kDa size change. A change in the mobility of the protein after digestion is evidence that the carbohydrate chain was sensitive to the enzyme and therefore, that the protein had encountered a certain enzyme in the processing pathway. Alternatively, the analysis can be done by two-dimensional isoelectric focusing (IEF)/SDS-PAGE or two-dimensional nonequilibrium pH gradient electrophoresis (NEPHGE)/SDS-PAGE (Adams and Gallagher, 1992) to see the loss of charged sugar residues or of anionic oligosaccharide chains. The same digestions and SDS-PAGE analysis also apply to proteins that are radioiodinated or biotinylated, or to immunoprecipitates derived from subcellular fractions separated on sucrose or Percoll gradients. It is important to present the glycosylation pathways, as a detailed description of the pathways is needed to appreciate how they will be used in this unit. A single protein can have more than one kind of oligosaccharide (N-linked and O-linked), and each individual N-linked chain can mature into a different final form. The same is true for O-linked chains. Each is described below. THE N-LINKED PATHWAY The N-linked oligosaccharide maturation pathway is most frequently used for tracking protein movement through the Golgi complex. A common feature of all N-linked chains is the core region pentasaccharide shown in Figure 12.4.1, which consists of three mannose units and two N-acetylglucosamine units. The mannose units comprise the trimannosyl core, and two of these residues are α-linked to the only β-linked mannose in the molecule. The β-linked mannose is bound to one of the two N-acetylglucosamines. Because they are β1-4 linked to each other, resembling the polysaccharide chitin, this is called a chitobiose disaccharide. Initially, all N-glycosylated proteins begin life when a preformed, lipid-associated oligosaccharide is transferred within the lumen of the endoplasmic reticulum (ER) to Asn of proteins having an Asn-X-Thr/Ser sequence. This precursor oligosaccharide contains three glucose (Glc), nine mannose (Man), and two N-acetylglucosamine (GlcNAc) sugar residues, and has the structure shown in Figure 12.4.1. There are several ways to depict this structure. The short-hand symbol method used in Figure 12.4.1 is the most convenient, but be sure to note the linkages of the individual sugars, as they are important. The α and β symbols denote the anomeric configuration of the sugar, and the number indicates which hydoxyl group of the next sugar is involved in the glycosidic linkage. In all cases, the anomeric position is 1, except in sialic acid where it is 2.
Release of N-Linked Oligosaccharides
The details of the pathway are presented in Figures 12.4.2 and 12.4.3, along with the sensitivity to each endoglycosidase or glycoamidase. The figures show the steps between high-mannose and hybrid types (Fig. 12.4.2) and complex types (Fig. 12.4.3). The three Glc residues (filled triangles) are removed from properly folded proteins within the ER by two different oligosaccharide-processing α-glucosidases. The first α1-2Glc is cleaved by α-glucosidase I, and the next two α1-3Glc residues by α-glucosidase II (Fig. 12.4.2, step 1). An ER-associated α-mannosidase removes one Man residue (open circle; Fig. 12.4.2, step 2). The protein then moves on to the first step in Golgi-localized processing— the removal of the three remaining α1-2 Man units by Golgi α-mannosidase I to produce Man5GlcNAc2 (Fig. 12.4.2, steps 3 and 4). Many proteins have only high-mannose-type oligosaccharides with five to nine Man residues, and no further processing occurs.
12.4.2 Supplement 22
Current Protocols in Protein Science
A
B
α 2 α 3 α 3 α 2 α 2
α 2
6 α3 α 36 α α β 4
α 2 α
3 6
α β 4 β 4
=
Asn
=
β 4
Asn
Asn
Asn
core structure
precursor oligosaccharide
glucose (Glc)
N-acetylgalactosamine (GalNAc)
mannose (Man)
fucose (Fuc)
galactose (Gal)
xylose (Xyl)
N-acetylglucosamine (GlcNAc)
sialic acids (Sia)
Figure 12.4.1 Symbol structures for the core region and precursor of N-linked sugar chains. Each sugar is given a symbol and abbreviation at the bottom of the figure. Each one except sialic acid uses its anomeric carbon (C-1) for linking to other sugars. Sialic acid uses C-2 for glycosidic linkage to other sugars. Glycosidases and glycosyltransferases are anomeric specific and distinguish α or β configurations of each sugar. The core structure (A) is common to all N-linked chains and is composed of three Man and two GlcNAc residues. The α or β configuration of each sugar is indicated, and the OH group to which that sugar is linked is shown on the bar linking the two symbols. Thus, GlcNAcβ1-4GlcNAcβ is represented by two filled squares with β and 4 between them. When a structure is first presented, it will have full display such as that on the left side; if it is repeated, only the symbols will be used as shown immediately to the right. The precursor oligosaccharide (B) for all N-linked chains is synthesized in the ER and transferred cotranslationally to the peptide containing an available Asn-X-Thr/Ser sequon.
Alternatively, one to five GlcNAc residues (filled squares) can be added to the trimannosyl core, and these are usually extended with galactose (Gal; filled circles) and sialic acid (Sia; filled diamonds) residues. These extensions, called antennae, are the hallmarks of complex-type oligosaccharides. The transformation of the precursor sugar chain into various high-mannose or complex types is called oligosaccharide processing (Kornfeld and Kornfeld, 1985). Man5GlcNAc2 is an important intermediate because it can have several fates. The first is the well-established addition of one GlcNAc residue by GlcNAc transferase I (Fig. 12.4.2, step 6). This is the first step toward the formation of complex chains. However, simply adding Gal and Sia to the terminal GlcNAc of this oligosaccharide forms a hybrid structure (Fig. 12.4.2, steps 12 and 13), where the left side of the molecule looks like a complex chain having one antenna, and the right side still resembles a high-mannose chain. The GlcNAc1Man5GlcNAc2 structure is the required substrate for α-mannosidase II, which removes the two terminal Man units from the upper branch of the chain (i.e., the α1-3Man and α1-6 Man units; Fig. 12.4.2, step 7). This enzyme only works after the addition of the first GlcNAc.
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high mannose type
hybrid type Golgi complex
ER
α 3/6
1
β 4
13
5 2
6
12 β 2
α 2
3
6
4
7
endo H
+
+
+
–
+
–
D
–
–
+
+
–
–
F2 –
–
–
–
–
–
F3 –
–
–
–
–
–
+
+
+
+
+
PNGase F
+
12.4.4
Figure 12.4.2 N-linked oligosaccharide maturation pathway for high-mannose and hybrid types, and sensitivities to various enzymes (see Fig. 12.4.1 for key). Brackets (top) show the structures designated as high-mannose and hybrid chains. The boxes indicate ER or Golgi localization.The pathway begins with the precursor oligosaccharide (see Figure 12.4.1). Each successive numbered step in circles represents a glycosidase or glycosyl transferase that generates a new sugar chain with different sensitivities to the various endoglycosidases or PNGase F. (1) precursor oligosaccharide is trimmed by α-glucosidases I and II, removing three Glc. (2) ER mannosidase removes one Man. (3) α-Mannosidase I in Golgi complex removes two Man to make Man6GlcNAc2, with a single remaining α1-2Man. (4) The final α1-2Man is removed by a Golgi complex α-mannosidase I. (5) α-Mannosidase III removes the α1-3 and α1-6Man units to make Man3GlcNAc2. (6) GlcNAc transferase I adds GlcNAc to either Man5GlcNAc2 or Man3GlcNAc2. (7) α-Mannosidase II removes the α1-3 and α1-6Man units to make GlcNAc1Man3GlcNAc2. Sensitivity to various enzymes (bottom) changes when moving from left to right, but remains the same within vertical columns. NOTE: This continued maturation to form complex chains is shown in Figure 12.4.3. Additionally, these figures are not comprehensive; many glycosylation steps have not been included, but they do not affect the sensitivities to the enzymes listed.
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complex type Golgi complex
α 3/6
13
13
13
13
12
12
12
β 4
12
β
β
β 2
4
6
9 8
10
11 α 6 endo H –
–
–
–
–
D –
–
–
–
–
F2 –
+
+
–
–
F3 –
–
+
+
–
PNGase F +
+
+
+
+
Figure 12.4.3 N-linked oligosaccharide maturation pathway for complex types, and sensitivities to various enzymes (see Fig. 12.4.1 for key; see Fig. 12.4.2 for additional details). (8) GlcNAc transferase II adds a second GlcNAc to initiate a biantennary chain. (9) GlcNAc transferase IV adds a third GlcNAc to initiate a triantennary chain. (10) GlcNAc transferase V adds a fourth GlcNAc to initiate a tetraantennary chain. (11) Fucosyltransferase adds α1-6Fuc to the core region of complex chains. (12) β1-4Gal is added to available GlcNAc residues of hybrid and complex chains. (13) α2-3 or α2-6Sia is added to Gal residues of hybrid and complex chains.
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The other fate for the Man5GlcNAc2 chain is to act as a substrate for the newly identified α-mannosidase III (Chui et al., 1997), which removes the same two Man units as α-mannosidase II, but does not require the prior addition of the first GlcNAc (Fig. 12.4.2, step 5). α-Mannosidases II and III show partial overlap in the Golgi complex, but α-mannosidase III may occur preferentially in an earlier compartment. The Man3GlcNAc2 product of α-mannosidase III is also a substrate for GlcNAc transferase I. Thus, the same product, GlcNAc1Man3GlcNAc2 can be formed in two ways: first, by the sequential action of α-mannosidase III and GlcNAc transferase I (Fig. 12.4.2, steps 5 and 6) or, second, by GlcNAc transferase I and α-mannosidase II (Fig. 12.4.2, steps 6 and 7). Sensitivity to specific enzyme digestions (endo H and endo D) can distinguish which route was taken (Fig. 12.4.2). GlcNAc transferase II now adds a second GlcNAc to the α1-6-linked Man (Fig. 12.4.3, step 8). This molecule can also have several fates. First, fucose (Fuc) can be added to GlcNAc residue linked to the Asn of the protein (Fig. 12.4.3, step 11). Second, one to three more GlcNAc residues can be added to the core mannose residues to initiate tri- and tetraantennary chains (Fig. 12.4.3, steps 9 and 10), and even pentaantennary chains (not shown). GlcNAc additions are considered to occur in the medial Golgi regions. Each GlcNAc-based branch can be individually modified, but they are usually extended by one Gal (Fig. 12.4.3, step 12) and terminated by a Sia (Fig. 12.4.3, step 13). Both of these sugars are usually thought to be added in trans-Golgi cisternae or in the trans-Golgi network (TGN). Sometimes selected antennae are also fucosylated in the TGN. One or more terminal Gal residues can be extended by variable-length polylactosamines (Galβ4GlcNAc repeats) capped by a Sia. GlcNAc and Gal can be sulfated as a late, perhaps even final, step of processing. These extensions/modifications are thought to occur in the late Golgi complex and TGN, but their order and compartmental segregation are not well understood. Other modifications of N-linked sugar chains are known, but there are fewer tools available to analyze their biosynthetic localization. THE O-LINKED PATHWAY For practical purposes, only a portion of the O-linked pathway—i.e., the addition of the first few sugars—will be presented. However, it is very important to remember that some of the same outer chain structures such as Sia, polylactosamines, and Fuc residues are common to both N- and O-linked oligosaccharides. α-N-Acetylgalactosamine (α-GalNAc; open square) is the lead-off sugar for the O-linked pathway (Fig. 12.4.4; also see Fig. 12.4.1 for symbols). It is added to Ser/Thr residues that occur in the proper configuration, generating a broad variety of acceptor sequences. These sequences often cluster as repeats within mucin-like domains. GalNAc is added in the earliest parts of the Golgi complex, not cotranslationally. GalNAc can be further extended by at least six different sugars. The most common is the addition of a β1-3Gal (Fig. 12.4.4, step 1), forming a disaccharide that is one of the few O-linked chains that can be diagnosed by enzymatic digestions. This disaccharide is often capped by a Sia (Fig. 12.4.4, step 2). Additional sugars such as Sia (Fig. 12.4.4, step 3) or GlcNAc followed by Gal (Fig. 12.4.4, steps 4 and 5) can be added. Structural analysis can be done by sequential exoglycosidase digestion, but given the complexity and heterogeneity of the sugar chains, such analysis is not a very useful indicator for tracking protein movement through the Golgi complex. Many O-linked chains have terminal Sia residues and, when tightly clustered on Ser/Thr residues, these chains promote proteolysis by O-sialoglycoprotease regardless of the structure of the underlying sugar chain. Release of N-Linked Oligosaccharides
Another type of O-linked glycosylation is the addition of glycosaminoglycan (GAG) chains to form proteoglycans. This occurs by a different pathway than the α-GalNAc
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α 6
α 3
3 Ser/Thr
β3 α
1
-Ser/Thr-
2
4
α 3
-Ser/Thr- Ser/Thr β 4
β 6 -Ser/Thr-
6
5 Ser/Thr
Ser/Thr
Figure 12.4.4 A small portion of the O-GalNAc pathway (see Fig. 12.4.1 for key). The first step of the O-linked pathway occurs in the early Golgi complex with the addition of α-GalNAc. There are at least six other sugars that can be added at this point in this complex pathway. Often β1-3Gal is added (1), quickly followed by α2-3Sia (2). The presence of these structures can be detected with a combination of O-glycosidase and sialidase. Additional sugars can be added as shown. α2-6Sia (3) or β1-6GlcNAc (4) followed by β1-4Gal (5) and α2-3Sia (6) on Gal. Each of these sugars must be removed before O-glycosidase can cleave the disaccharide.
linkage. Instead, the chains begin by addition of a β-Xylose (Xyl; open inverted triangle) residue to Ser and are then elongated by two Gal residues and a glucuronic acid (GlcA; half-filled diamond) residue. This core structure can be further elongated by the addition of GlcAβ1-3GalNAcβ disaccharides to form the backbone of chondroitin/dermatan sulfate chains, or by GlcAβ1-3GlcNAcα to form the backbone of heparan sulfate chains. Biosynthesis and movement of these proteins have also been followed through the Golgi complex. Initiation begins in late ER/early Golgi complex, and the core tetrasaccharide is probably finished within the medial Golgi, but the addition of chondroitin chains appears to be confined to the TGN. In addition to the well-known O-linked GAG chains, there is clear evidence for the existence of a class of N-linked GAG chains. ENDOGLYCOSIDASE H DIGESTION Endoglycosidase H (endo H) cleaves N-linked oligosaccharides between the two N-acetylglucosamine (GlcNAc) residues (Fig. 12.4.5) in the core region of the oligosaccharide chain (Fig. 12.4.1) on high-mannose and hybrid, but not complex, oligosaccharides. In this protocol, a fully denatured protein is digested with endo H to obtain complete release of sensitive oligosaccharides. Materials Immunoprecipitated protein of interest (UNIT 9.8) 0.1 M 2-mercaptoethanol (2-ME)/0.1% (w/v) SDS (ultrapure electrophoresis grade; prepare fresh) 0.5 M sodium citrate, pH 5.5 1% (w/v) phenylmethylsulfonyl fluoride (PMSF) in isopropanol 0.5 U/ml endoglycosidase H (endo H; natural or recombinant; Sigma, Glyko, or Boehringer Mannheim) 10× SDS sample buffer (UNIT 10.1) Water baths, 30° to 37°C and 90°C
BASIC PROTOCOL 1
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X
α
Y
3
6
β 4 β 4 PNGase F
β
α endo H D F1 F2 F3
Figure 12.4.5 PNGase F and endoglycosidase-sensitive bonds in the core of N-linked oligosaccharides (see Fig. 12.4.1 for key). PNGase F is a glycoamidase that severs the bond between GlcNAc and Asn, liberating the entire sugar chain and converting Asn into Asp. The endoglycosidases (H, D, and Fs) cleave the bond between the two GlcNAc residues in the core region, leaving one GlcNAc still bound to the protein. The differential specificity of the endoglyosidases is based on the structure of the sugar chain in a fully denatured protein. Incomplete denaturation may not expose all sensitive linkages. X and Y are unspecified sugar residues.
Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11) 1. Add 20 to 30 µl of 0.1 M 2-ME/0.1% SDS to immunoprecipitate in a microcentrifuge tube, mix well, and heat denature 3 to 5 min at 90°C. Use the larger amount of reagent (≥30 ìl) for more complete recovery. This treatment may also release some (unlabeled) antibody molecules. Protein solubilization in nonionic detergents such as Triton X-100 or Nonidet P-40 is not always sufficient to completely expose all susceptible cleavage sites. Only strong denaturation with SDS exposes all sites for maximum cleavage.
2. Cool and microcentrifuge for 1 sec at 1000 × g to collect condensed droplets in the bottom of the tube. 3. Place 10-µl aliquots of solubilized, denatured protein (supernatant) in each of two clean microcentrifuge tubes, one for a control (no enzyme) and the other to digest (plus enzyme). 4. Add in the following order, mixing after each addition: 6 µl 0.5 M sodium citrate, pH 5.5 20 µl H2O 2 µl 1% PMSF (in isopropanol) 1 µl 0.5 U/ml endo H (enzyme digest only; substitute with water in control). The PMSF prevents proteolysis. Nonionic detergent is not required to prevent inactivation of endo H as long as high-purity SDS is used. The amount of water can be varied. Other solutions (or an increased amount of sample) may be substituted for water if needed, but potassium buffers should be avoided because they precipitate SDS as a potassium salt. The reaction volume can be scaled up proportionally if required, but in nearly all cases the enzyme will be in vast excess.
5. Incubate overnight at 30°C to 37°C. 6. Immediately prior to electrophoresis, inactivate endo H by adding 4 µl of 10× SDS sample buffer and heating 5 min at 90°C. Release of N-Linked Oligosaccharides
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7. Analyze protein by one-dimensional SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11). The presence of high mannose and/or hybrid N-linked oligsaccharide chains will be evidenced by increased mobility of the digested proteins on SDS-PAGE.
ENDOGLYCOSIDASE D DIGESTION Like endo H, endo D also cleaves between the two GlcNAc residues in the core of the N-linked sugar chains (Fig. 12.4.5). However, its narrow substrate specificity makes it useful for detecting the transient appearance of just a few early processing intermediates. It requires that the 2 position of the α1-3-linked core Man be unsubstituted. This intermediate arises after processing by either α-mannosidase I or III, but prior to addition of the first GlcNAc or action of α-mannosidase II (see Fig. 12.4.2, steps 3 to 5). Cells with a defect in GlcNAc I transferase (e.g., Lec 1 CHO cells) do not add the first GlcNAc residue (Fig. 12.4.2, step 6), and N-linked oligosaccharides will remain sensitive to endo D because they cannot modify the α1-3Man residue.
BASIC PROTOCOL 2
Materials Immunoprecipitated protein of interest (UNIT 9.8) 0.1 M 2-mercaptoethanol (2-ME)/0.1% (w/v) SDS (ultrapure electrophoresis grade; prepare fresh) 0.5 M NaH2PO4, pH 6.5 10% (w/v) Triton X-100 or Nonidet P-40 (NP-40) 0.5 U/ml endoglycosidase D (endo D; Boehringer Mannheim) 10× SDS sample buffer (UNIT 10.1) Water baths, 37° and 90°C Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11) 1. Denature immunoprecipitated protein in 20 to 30 µl of 0.1 M 2-ME/0.1% SDS by heating for 3 to 5 min at 90°C. Use the larger amount of reagent (≥30 ìl) for more complete recovery. This treatment may also release some (unlabeled) antibody molecules.
2. Cool and microcentrifuge at 1000 × g for 1 sec to collect condensed droplets in the bottom of the tube. 3. Transfer 10-µl aliquots of supernatant into two microcentrifuge tubes, one for a control (no enzyme) and the other to digest (plus enzyme). 4. Add in the following order, mixing after each addition: 2 µl 10% Triton X-100 or NP-40 (20-fold excess over SDS) 2 µl 0.5 M NaH2PO4, pH 6.5 5 µl H2O 1 µl 1 IU/ml endo D (enzyme digest only; substitute with water in control). The 20-fold excess of nonionic detergent is essential to prevent inactivation of endo D by SDS. The amount of water can be varied. Other solutions (or an increased amount of sample) may be substituted for water if needed, but potassium buffers should be avoided because they precipitate SDS as a potassium salt. The reaction volume can be scaled up proportionally if required, but in nearly all cases the enzyme will be in vast excess.
5. Incubate at 37°C overnight.
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6. Immediately prior to electrophoresis, inactivate by adding 2 µl of 10× SDS sample buffer and heating for 5 min at 90°C to 95°C. 7. Analyze protein by one-dimensional SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11). Endo D sensitivity is detected by increased electrophoretic mobility of the digested proteins on SDS-PAGE. BASIC PROTOCOL 3
ENDOGLYCOSIDASE F2 DIGESTION Endo F2, like endo H and endo D, cleaves between the two GlcNAc residues in the chitobiose core (Fig. 12.4.5). It preferentially releases biantennary complex-type oligosaccharide chains from glycoproteins, but does not cleave tri- or tetraantennary chains. Materials Immunoprecipitated protein of interest (UNIT 9.8) 0.1 M 2-mercaptoethanol (2-ME)/0.1% (w/v) SDS (ultrapure electrophoresis grade; prepare fresh) 0.5 M sodium acetate, pH 4.5 (APPENDIX 2E) 10% (w/v) Triton X-100 or Nonidet P-40 (NP-40) 0.1 M 1,10-phenanthroline in methanol 200 mU/ml endoglycosidase F2 (endo F2; Glyko) 4× SDS sample buffer (UNIT 10.1) Water baths, 30° to 37°C and 90°C Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11) 1. Denature immunoprecipitated protein in 20 to 30 µl of 0.1 M 2-ME/0.1% SDS by heating for 3 to 5 min at 90°C. Use the larger amount of reagent (≥30 ìl) for more complete recovery. This treatment may also release some (unlabeled) antibody molecules.
2. Cool and microcentrifuge at 1000 × g for 1 sec to collect condensed droplets in the bottom of the tube. 3. Transfer 10-µl aliquots of supernatant into two microcentrifuge tubes, one for a control (no enzyme) and the other to digest (plus enzyme). 4. Add in the following order, mixing after each addition: 15 µl 0.5 M sodium acetate, pH 4.5 3 µl 0.1 M 1,10-phenanthroline in methanol 2 µl 10% Triton X-100 or NP-40 (20-fold excess over SDS) 1 µl 200 mU/ml endo F2 (enzyme digest only; substitute 0.5 M sodium acetate in control). A 10- to 20-fold excess of nonionic detergent is required to stabilize the enzyme. The amount of water can be varied. Other solutions (or an increased amount of sample) may be substituted for water if needed, but potassium buffers should be avoided because they precipitate SDS as a potassium salt. The reaction volume can be scaled up proportionally if required, but in nearly all cases the enzyme will be in vast excess.
5. Incubate the mixture at 30° to 37°C overnight. Release of N-Linked Oligosaccharides
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Some inactivation of the enzyme occurs at 37°C, even with nonionic detergent present; however, if the enzyme is present in sufficient excess, incubation can generally be carried out successfully at 37°C.
6. Immediately before electrophoresis, inactivate by adding 8 µl of 4× SDS sample buffer and heating for 5 min at 90°C to 95°C. 7. Analyze the protein by one-dimensional SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11). Sensitivity to endo F2 is detected by increased electrophoretic mobility on SDS-PAGE.
ENDOGLYCOSIDASE F3 DIGESTION Endoglycosidase F3 (endo F3) is another endoglycosidase with a narrow substrate range and, therefore, high specificity: it cleaves triantennary chains, but not high-mannose, hybrid, nonfucosylated biantennary or tetraantennary chains. A core-fucosylated biantennary chain is the only other demonstrated substrate. When both endo F3 and endo F2 digestions are done in parallel on a sample, it can provide evidence for chain branching and core fucosylation. The approach is essentially the same as for the other endoglycosidases.
BASIC PROTOCOL 4
Materials Immunoprecipitated protein of interest (UNIT 9.8) 0.1 M 2-mercaptoethanol (2-ME)/0.1% (w/v) SDS (ultrapure electrophoresis grade; prepare fresh) 0.5 M sodium acetate, pH 4.5 (APPENDIX 2E) 10% (w/v) Triton X-100 or NP-40 0.1 U/ml endoglycosidase F3 (endo F3; Glyko) 10× SDS sample buffer (UNIT 10.1) Water baths, 37° and 90°C Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11) 1. Denature immunoprecipitated protein in 20 to 30 µl of 0.1 M 2-ME/0.1% SDS and heat denature 3 to 5 min at 90°C. Use the larger amount of reagent (≥30 ìl) for more complete recovery. This treatment may also release some (unlabeled) antibody molecules.
2. Cool and microcentrifuge for 1 sec at 1000 × g to collect condensed droplets at the bottom of the tube. 3. Transfer 10-µl aliquots of supernatant into two microcentrifuge tubes, one for a control (no enzyme) and the other to digest (plus enzyme). 4. Add in the following order, mixing after each addition: 2 µl 10% Triton X-100 or NP-40 (20-fold excess over SDS) 4 µl 0.5 M sodium acetate, pH 4.5 5 µl H2O 1 µl 0.1 U/ml endo F3 (enzyme digest only; substitute with water in control). The amount of water can be varied. Other solutions (or an increased amount of sample) may be substituted for water if needed, but potassium buffers should be avoided because they precipitate SDS as a potassium salt. The reaction volume can be scaled up proportionally if required, but in nearly all cases the enzyme will be in vast excess.
5. Incubate at 37°C overnight.
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6. Immediately prior to electrophoresis, inactivate by adding 2 µl of 10× SDS sample buffer and heating for 5 min at 90°C to 95°C. 7. Analyze by one-dimensional SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11). Sensitivity to endo F3 is detected by increased mobility on SDS-PAGE. BASIC PROTOCOL 5
PEPTIDE: N-GLYCOSIDASE F DIGESTION PNGase F is a glycoamidase that cleaves the bond between the Asp residue of the protein and the GlcNAc residue that joins the carbohydrate to the protein (Fig. 12.4.5). Because it liberates nearly all known N-linked oligosaccharides from glycoproteins, it is the preferred enzyme for complete removal of N-linked chains. It is the only enzyme that releases tetra- and pentaantennary chains. The glycoprotein sample must be denatured and digested with PNGase F to remove N-linked oligosaccharides completely. Materials Immunoprecipitated protein of interest (UNIT 9.8) 0.1 M 2-mercaptoethanol (2-ME)/0.1% (w/v) SDS (ultrapure electrophoresis grade; prepare fresh) 0.5 M Tris⋅Cl, pH 8.6 as determined at 37°C (APPENDIX 2E) 10% (w/v) Triton X-100 or Nonidet P-40 (NP-40) 200 to 250 mU/ml peptide:N-glycosidase F (PNGase F; Sigma or Glyko) 10× SDS sample buffer (UNIT 10.1) Water baths, 30° to 37°C and 90°C Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11) 1. Denature immunoprecipitated protein in 20 to 30 µl of 0.1 M 2-ME/0.1% SDS by heating 3 to 5 min at 90°C. Use the larger amount of reagent ≥30 ìl) for more complete recovery. This treatment may also release some (unlabeled) antibody molecules.
2. Cool and microcentrifuge for 1 sec at 1000 × g to collect condensed droplets in the bottom of the tube. 3. Transfer 10-µl aliquots of supernatant into two microcentrifuge tubes, one for a control (no enzyme) and the other to digest (plus enzyme). 4. Add in the following order, mixing after each addition: 3 µl 0.5 M Tris⋅Cl, pH 8.6 5 µl H2O 2 µl 10% NP-40 or Triton X-100 5 µl 200 to 250 mU/ml PNGase F (enzyme digest only; substitute with 0.5 M Tris⋅Cl in control). Sodium phosphate or HEPES buffer, pH 7.0, can be used instead of Tris⋅Cl. Avoid potassium buffers because these may cause precipitation of a potassium SDS salt. Use of a nonionic detergent is essential, because SDS inactivates PNGase F. A 10-fold weight excess of any of the above nonionic detergents over the amount of SDS will stabilize the enzyme. The amount of water can be varied. Other solutions (or an increased amount of sample) may be substituted for water if needed, but potassium buffers should be avoided because they precipitate SDS as a potassium salt. The reaction volume can be scaled up proportionally if required, but in nearly all cases the enzyme will be in vast excess. Release of N-Linked Oligosaccharides
5. Incubate overnight at 30° to 37°C.
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6. Immediately prior to electrophoresis, inactivate the enzyme by adding 2.5 µl of 10× SDS sample buffer and heating 3 to 5 min at 90°C. 7. Analyze the protein by one-dimensional SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11). The presence of N-linked oligosaccharide chains will be evidenced by increased electrophoretic mobility on SDS-PAGE.
ESTIMATING THE NUMBER OF N-LINKED OLIGOSACCHARIDE CHAINS ON A GLYCOPROTEIN
SUPPORT PROTOCOL
One widely used application of endo H or PNGase F digestion is estimation of the number of N-linked oligosaccharide chains on a given glycoprotein. This is done by creating a ladder of partially digested molecules that each differ by only one N-linked sugar chain. The number of separate bands in a one-dimensional polyacrylamide gel (less one for the totally deglycosylated protein) provides an estimate of the number of N-linked chains. The conditions used to generate partially deglycosylated protein must be determined for each protein studied, because the sensitivity of each chain may be different, even when all of them are completely exposed by denaturation. For this protocol, either the incubation time or the amount of enzyme can be varied to determine the best conditions to produce a ladder of partial digests. Usually five or six points are enough to provide a reasonable estimate (Fig. 12.4.6). Of course, it is important to use enough enzyme to obtain complete deglycosylation. This is best done by monitoring the effects of endo H or PNGase F on newly synthesized [35S]Met pulse-labeled protein just after synthesis, but before any N-linked oligosaccharide processing has occurred. Pulse labeling of protein for 10 min with [35S]Met followed by digestion is the best way to be sure that all chains are removed.
1 kDa
92.5–
2
3
4
5 kDa
85
66–
45– 39.5 30–
Figure 12.4.6 Data from the estimation of the number of glycosylation sites on lysosome-associated membrane protein 1 (LAMP-1; Viitala et al., 1988). LAMP-1 contains eighteen potential N-linked sites. Graded digestion with increasing amounts of PNGase F was used to generate this ladder of glycoforms. Each band contains at least one less N-linked chain than the band above it. An average N-linked carbohydrate chain has an apparent mass of ∼1.5 to 3 kDa. Lysosomal membrane glycoprotein was immunoprecipitated from [35S]Met-labeled cells and the sample was digested with PNGase F for 0 min (lane 1), 5 min (lane 2), 20 min (lane 3), 45 min (lane 4), and 24 hr (lane 5). Figure courtesy of Dr. Minoru Fukuda.
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1. Add 0.1 M 2-ME/0.1% SDS solution to the total volume of immunoprecipitated protein required and heat denature by incubating 3 to 5 min at 90°C. Each digestion reaction requires 20 ìl of immunoprecipitate. Thus, 120 to 140 ìl is sufficient for one control plus five or six digests.
2. Cool and centrifuge for 1 sec at 1000 × g to collect condensed droplets at the bottom of the tube. 3. Aliquot 10 µl supernatant to the number of microcentrifuge tubes required to cover the concentration range (e.g., 0.01 to 1 mU/ml PNGase F) or incubation times (e.g., 5 to 60 min) plus one for an undigested control. 4. Add remaining reagents as specified for endo H (see Basic Protocol 1, step 4) or PNGase F (see Basic Protocol 5, step 4), adjusting the enzyme concentration as desired. 5. Incubate at 30°C for the desired length of time. High enzyme concentration (10 mU/ml) and prolonged incubation (16 hr) must be among the conditions included, in order to ensure that there is a data point for maximum deglycosylation. For varying enzyme concentrations, incubate for the same amount of time, but the duration of incubation should be shorter than what would give complete digestion because the goal is to obtain increasing extent of incomplete cleavage.
6. After the desired incubation time, inactivate enzyme by adding 0.1 volume of µl of 10× SDS sample buffer and heating 5 min at 90° to 95°C. 7. Analyze the sample from each concentration/time point, including undigested sample, by one-dimensional SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11). Most newly formed N-linked chains will have a molecular weight in the range of 1500 to 2200, and loss of one chain is sufficient to change the migration of a protein. This procedure has been used to count up to eighteen N-linked sites on one molecule. A sample result is shown in Figure 12.4.6. BASIC PROTOCOL 6
SIALIDASE (NEURAMINIDASE) DIGESTION Sialic acids are the terminal sugars on many N- and O-linked oligosaccharides. The great majority are released with broad-specificity sialidases (neuraminidases) such as that from Arthrobacter ureafaciens. Because sialic acids are charged, their loss usually changes the mobility on one-dimensional SDS polyacrylamide gels, but it will always change the mobility on a two-dimensional gel. Since one-dimensional analysis is easier, it can be tried first. Materials Immunoprecipitated protein of interest (UNIT 9.8) 0.1 M 2-mercaptoethanol (2-ME)/0.1% (w/v) SDS (ultrapure electrophoresis grade; prepare fresh) 10% (w/v) Triton X-100 or Nonidet P-40 (NP-40) 0.5 M sodium acetate, pH 5.0 (APPENDIX 2E) 1 IU/ml neuraminidase from Arthrobacter ureafaciens (Sigma or Glyko) 10× SDS sample buffer (UNIT 10.1) Water baths, 37° and 90°C Additional reagents and equipment for SDS-PAGE (UNIT 10.1), for IEF/SDS-PAGE or NEPHGE/SDS-PAGE (UNIT 10.4), and for autoradiography (UNIT 10.11)
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1. Denature immunoprecipitated protein in 20 to 30 µl of 0.1 M 2-ME/0.1% SDS by heating 3 to 5 min at 90°C. Denaturation is less important here, because the sialic acids are exposed at the ends of the sugar chains. In most instances, the denaturation step can probably be omitted and the digestion done while the protein is still bound to the beads.
2. Cool and microcentrifuge for 1 sec at 1000 × g to collect condensed droplets in the bottom of the tube. 3. Transfer 10-µl aliquots of supernatant to two clean microcentrifuge tubes, one for a control (no enzyme) and the other to digest (plus enzyme). 4. Add in the following order, mixing after each addition: 2 µl 10% Triton X-100 or NP-40 (20-fold excess over SDS) 4 µl 0.5 M sodium acetate, pH 5.0 5 µl H2O 1 µl 1 IU/ml neuraminidase (enzyme digest only; substitute with water for control). This amount of neuraminidase should be in great excess. Addition of nonionic detergent is not needed if the digestion is done while the protein was still bound to the beads. The amount of water can be varied. Other solutions (or an increased amount of sample) may be substituted for water if needed, but potassium buffers should be avoided because they precipitate SDS as a potassium salt. The reaction volume can be scaled up proportionally if required, but in nearly all cases the enzyme will be in vast excess.
5. Incubate at 37°C overnight. This time can be shortened to 2 hr, if necessary, but longer incubations are better.
6. Immediately prior to electrophoresis, inactivate the enzyme by adding 2 µl of 10× SDS sample buffer and heating 3 to 5 min at 90°C. If the protein will be analyzed by IEF or NEPHGE, addition of sample buffer is replaced by lysis buffer used for these techniques.
7. Analyze the protein using either the appropriate one-dimentional SDS-PAGE system (UNIT 10.1) or a two-dimentional IEF/SDS-PAGE or NEPHGE/SDS-PAGE system (UNIT 10.4), and detect by autoradiography (UNIT 10.11). Removal of sialic acids usually results in a decrease in apparent molecular weight on one-dimentional gel analysis, or an increase in the isoelectric point of the protein analyzed by two-dimentional gel analysis.
ENDO-â-GALACTOSIDASE DIGESTION The endo-β-galactosidase from Bacillus fragilis degrades polylactosamine chains (Galβ1-4GlcNAcβ1-3)n found on both N- and O-linked oligosaccharides. The variable length of these repeating units usually causes the protein to run as a broad band or a smear on the gel. Although not all linkages are equally cleaved by this enzyme (see Fig. 12.4.7), sensitive proteins that often run as broad bands or smears on gels—e.g., lysosome-associated membrane protein 1 (LAMP-1)—produce both sharper bands and lower molecular weight species after digestion.
BASIC PROTOCOL 7
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α
β
β
β
β
β
3
4
3
4
3
4
α 3
β
β
β 6S β
3
4
3
4
Figure 12.4.7 Endo-β-galactosidase-sensitive linkages in polylactosamines (see Fig. 12.4.1 for key). Linear, unsubstituted polylactosamine units (GlcNAcβ1-3Galβ1-4) are sensitive to digestion with endo-β-galactosidase, while substitutions—such as sulfate esters (S) or branches starting with GlcNAc (not shown)—completely block digestion. Substitution of neighboring GlcNAc with Fuc or sulfate esters slows the rate, but does not block cleavage. Sensitive sites are shown with bold arrows, slowly hydrolyzed sites with a lighter arrow, and resistant bonds are struck out. Various substitutions are possible, leading to broad bands on gels. This will create variable sensitivities, but even partial sensitivity should give a sharper, more defined band.
Materials Immunoprecipitated protein of interest (UNIT 9.8) 0.1 M 2-mercaptoethanol (2-ME)/0.1% (w/v) SDS (ultrapure electrophoresis grade; prepare fresh) 0.5 M sodium acetate buffer, pH 5.8 (APPENDIX 2E) 10% (w/v) Triton X-100 or Nonidet P-40 (NP-40) 100 mU/ml endo-β-galactosidase (Boehringer Mannheim, Oxford GlycoSystems, Sigma) 10× SDS sample buffer (UNIT 10.1) Water baths, 37° and 95°C Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11) 1. Denature immunoprecipitated protein in 20 to 30 µl of 0.1 M 2-ME/0.1% SDS by heating 3 to 5 min at 95°C. Use the larger amount of reagent (≥30 ìl) for more complete recovery. This treatment may also release some (unlabeled) antibody molecules.
2. Cool and microcentrifuge 1 sec at 1000 × g to collect condensed droplets in the bottom of the tube. 3. Transfer 10-µl aliquots of supernatant into two microcentrifuge tubes, one for a control (no enzyme) and the other to digest (plus enzyme). 4. Add in the following order, mixing after each addition: 2 µl 10% Triton X-100 or NP-40 (20-fold excess over SDS) 4 µl 0.5 M sodium acetate, pH 5.8 5 µl H2O 1 µl 100 mU/ml endo-β-galactosidase (enzyme digest only; substitute with water in control).
Release of N-Linked Oligosaccharides
The amount of water can be varied. Other solutions (or an increased amount of sample) may be substituted for water if needed, but potassium buffers should be avoided because they precipitate SDS as a potassium salt. The reaction volume can be scaled up proportionally if required, but in nearly all cases the enzyme will be in vast excess.
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5. Incubate at 37°C overnight. 6. Immediately prior to electrophoresis, inactivate by adding 3 µl of 10× SDS sample buffer and heating for 5 min at 90°C. 7. Analyze protein by one-dimensional SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11). If the protein has polylactosamine chains, its mobility should increase after digestion.
ENDO-á-N-ACETYLGALACTOSAMINIDASE DIGESTION This enzyme (also known as O-glycosidase or O-glycanase) has limited utility because it is highly specific for cleaving only one O-linked disaccharide, Galβ1-3GalNAcαSer/Thr. Adding any more sugars, including sialic acid, renders the molecule resistant to cleavage and requires removal of each residue before the enzyme will work. Prior sialidase digestion is sometimes used (see Basic Protocol 6), and this can be done while the protein is still bound to the immunoprecipitation beads.
BASIC PROTOCOL 8
Materials Immunoprecipitated protein of interest (UNIT 9.8) 0.1 M 2-mercaptoethanol (2-ME)/0.1% (w/v) SDS (ultrapure electrophoresis grade; prepare fresh) 0.5 M sodium citrate phosphate buffer, pH 6.0, containing 500 µg/ml BSA (complete buffer supplied with enzyme) 10% (w/v) Triton X-100 or Nonidet P-40 (NP-40) 300 mU/ml endo-α-N-acetylgalactosaminidase (5× concentrate from Glyko; use according to directions) 10× SDS sample buffer (UNIT 10.1) Water bath, 37° and 95°C Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11) 1. Denature immunoprecipitated protein in 20 to 30 µl of 0.1 M 2-ME/0.1% SDS by heating 3 to 5 min at 95°C. Use the larger amount of reagent (≥30 ìl) for more complete recovery. This treatment may also release some (unlabeled) antibody molecules.
2. Cool and microcentrifuge 1 sec at 1000 × g to collect condensed droplets in the bottom of the tube. 3. Transfer 10-µl aliquots of supernatant into two microcentrifuge tubes, one for a control (no enzyme) and the other to digest (plus enzyme). 4. Add in the following order, mixing after each addition: 2 µl 10% Triton X-100 or NP-40 (20-fold excess over SDS) 4 µl 0.5 M sodium citrate phosphate buffer, pH 6.0, with 500 µg/ml BSA 3 µl H2O 1 µl 300 mU/ml endo-α-N-acetylgalactosidase (enzyme digest only; substitute with water in control). The amount of water can be varied. Other solutions (or an increased amount of sample) may be substituted for water if needed, but potassium buffers should be avoided because they precipitate SDS as a potassium salt. The reaction volume can be scaled up proportionally if required, but in nearly all cases the enzyme will be in vast excess.
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5. Incubate at 37°C overnight. 6. Immediately prior to electrophoresis, inactivate by adding 2 µl of 10× SDS sample buffer and heating for 5 min at 90°C. 7. Analyze protein by one-dimensional SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11). If the protein contains the disaccharide unit, the mobility of the protein should increase. Presence of only a single unit (mol. wt. ∼400 Da) may be difficult to detect unless a high-resolution gel is used. BASIC PROTOCOL 9
O-SIALOGLYCOPROTEASE DIGESTION Digestion with O-sialoglycoprotease requires that the substrate have a tight cluster of sialylated O-linked oligosaccharides. Proteins with a single O-linked chain or a few widely spaced chains will not be cleaved. This property makes the enzyme a valuable diagnostic tool. Materials Immunoprecipitated protein of interest (UNIT 9.8) 0.1 M 2-mercaptoethanol (2-ME)/0.1% (w/v) SDS (ultrapure electrophoresis grade; prepare fresh) 0.4 M HEPES buffer, pH 7.4 10% (w/v) Triton X-100 or Nonidet P-40 (NP-40) 2.4 mg/ml O-sialoglycoprotease (O-sialoglycoprotein endoglycoprotease; Accurate Chemical & Scientific; reconstituted according to directions) 10× SDS sample buffer (UNIT 10.1) Water baths, 37° and 95°C Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11) 1. Denature immunoprecipitated protein in 20 to 30 µl of 0.1 M 2-ME/0.1% SDS by heating 3 to 5 min at 95°C. Use the larger amount of reagent (≥30 ìl) for more complete recovery. This treatment may also release some (unlabeled) antibody molecules.
2. Cool and microcentrifuge 1 sec at 1000 × g to collect condensed droplets in the bottom of the tube. 3. Transfer 10-µl aliquots of supernatant into two microcentrifuge tubes, one for a control (no enzyme) and the other to digest (plus enzyme). 4. Add in the following order, mixing after each addition: 2 µl 10% Triton X-100 or NP-40 (20-fold excess over SDS) 4 µl 0.4 M HEPES buffer, pH 7.4 5 µl H2O 2 µl 2.4 mg/ml O-sialoglycoprotease (enzyme digest only; substitute with water in control). O-Sialoglycoprotein endopeptidase is a partially purified enzyme, and the specific activity is relatively low. A quantity of 1.0 ìg of this enzyme preparation will cleave 5 ìg of sensitive substrate per hour at 37°C. Human glycophorin A can serve as a positive control.
Release of N-Linked Oligosaccharides
The amount of water can be varied. Other solutions (or an increased amount of sample) may be substituted for water if needed, but potassium buffers should be avoided because they precipitate SDS as a potassium salt. The reaction volume can be scaled up proportionally if required, but in nearly all cases the enzyme will be in vast excess.
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5. Incubate at 37°C overnight. 6. Immediately prior to electrophoresis, inactivate by adding 2.5 µl of 10× SDS sample buffer and heating for 5 min at 90°C. 7. Analyze protein by one-dimensional SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11). If the digestion was successful, the target protein will be undetectable or may be cleaved into small fragments.
COMMENTARY Background Information The results of digestion of a hypothetical protein with two N-linked carbohydrate chains as it moves through the ER and Golgi complex with various enzymes are shown in Figure 12.4.8. At 0 min, both N-linked chains are high-mannose type. They have lost their Glc residues and one Man residue in the ER. Both are sensitive to endo H and PNGase F digestion, yielding a protein with only two remaining GlcNAc residues in the case of endo H diges-
time after pulse labeling enzyme C H digestion
0 min D F2 PNG Sia
tion, and no carbohydrate at all in the case of PNGase F digestion. These sugar chains are resistant to the other enzyme digestions. At 45 min, the protein is in the medial Golgi complex and both sugar chains have been processed by Golgi α-mannosidase I. However, one of the chains (left) has been partially processed by Golgi α-mannosidase III to an endo H–resistant/endo D–sensitive chain. The other chain (right) has received a GlcNAc residue from GlcNAc transferase I, but has not encountered
45 min
120 min
C H D F2 PNG Sia C H D F2 PNG Sia
SDS-PAGE gel pattern
+
+
+
+
+
sugar chains
protein
Figure 12.4.8 Schematic diagram showing results that could be obtained for a hypothetical protein with two N-linked glycosylation sites as it moves through the Golgi complex. Assume that the protein has been biosynthetically labeled with an amino acid precursor (such as [35S]Met) for 10 min and chased in the absence of label for 45 min and 120 min. The protein is then precipitated with a specific antibody. At each time point, equal amounts of the sample are analyzed by flourography after one-dimensional SDS-PAGE, either without any digestion (control; C) or following digestion with endo H (H), endo D (D), endo F2 (F2), PNGase F (PNG), or sialidase (Sia). Oligosaccharide structures consistent with the banding patterns are shown below the schematic gel pattern.
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Golgi α-mannosidase II. This chain is still endo H sensitive and endo D resistant. Both chains are sensitive to PNGase F, but neither is sensitive to endo F2 or sialidase digestion. At 120 min, both chains have fully matured to complex-type chains. One is a sialylated biantennary chain and the other is a sialylated triantennary chain. Note that each sialic acid is marked ±, indicating that not all molecules are fully sialylated, accounting for the broader bands. Neither sugar chain is sensitive to endo H or endo D digestion. The biantennary chain (left) is sensitive to endo F2 cleavage, leaving one GlcNAc residue on the protein, but the triantennary chain (right) is not sensitive to this enzyme. Both chains are cleaved by PNGase F. All sialic acids are removed by sialidase to produce a sharp band, but the underlying sugar chains remain.
Release of N-Linked Oligosaccharides
Endoglycosidase H Endo H from Streptomyces plicatus cleaves the bond between the two GlcNAc residues in the core of N-linked oligosaccharides. One GlcNAc residue remains attached to the protein or peptide and the remainder of the chain is released as an intact unit (Tarentino et al., 1989). The oligosaccharide structures cleaved by endo H are shown schematically in Figure 12.4.5. Substrates for endo H include all high-mannose oligosaccharides and certain hybrid types, but not bi-, tri-, tetra- or pentaantennary (complex) chains. Endo H also cleaves oligosaccharides that have α1-6 fucose residues bound to the reducing (protein to carbohydrate linkage) GlcNAc residue (Tarentino et al., 1989). Endo H sensitivity is the most common way to trace the movement of newly synthesized glycoproteins from the endoplasmic reticulum (ER) into the Golgi complex. Proteins remain sensitive to endo H while they are in the ER and in early regions of the Golgi complex; they become endo H–resistant after they are processed by enzymes located in the medial Golgi complex. Endo H cleaves the N-linked oligosaccharides from proteins as long as they have not lost the α1-3Man residue cleaved by Golgi mannosidase II or mannosidase III (Fig. 12.4.2). After removal of that mannose, the oligosaccharide becomes endo H resistant. Endo H is the best enzyme for identifying high-mannose and hybrid chains and their general transition toward complex chains. Using endo H alone gives an incomplete map of subcellular trafficking. The resolution of this picture can be enhanced by combining it with endo D digestions. In general, N-linked
chains become endo H resistant at about the same time they become transiently sensitive to endo D. However, the discovery of the alternative pathway for complex N-glycan processing using α-mannosidase III now gives a different perspective. Both α-mannosidase II and III are expressed in most mouse tissues except for hematopoietic cells, which seem to have only α-mannosidase II. So it is conceivable that some proteins, and even different N-linked chains on a single protein, can be processed by α-mannosidase II and others by α-mannosidase III, yielding complex digestion patterns and kinetics of endo H and endo D sensitivity. Endoglycosidase D The narrow substrate specificity of endo D makes it useful for detecting a very restricted set of processing intermediates, particularly for distinguishing α-mannosidase II from αmannosidase III processing. When the results of endo D digestion are combined with the results of an endo H digestion of the same sample, it can help determine which of the two alternate processing pathways is being used. The key requirement for endo D cleavage is an unsubstituted 2 position on the α1-3Man that forms the trimannosyl core (Fig. 12.4.1). This window of endo D sensitivity occurs only after removal of the α1-2Man residue by α-mannosidase I and before the addition of a GlcNAc residue to this same position by GlcNAc transferase I (Beckers et al., 1987; Davidson and Balch, 1993). The immediate precursor of both pathways is Man6GlcNAc2, which is endo D resistant/endo H sensitive. The better-known α-mannosidase II pathway involves the removal of the last α1-2Man residue by α-mannosidase I (making the oligosaccharide endo D sensitive/endo H sensitive), and then the addition of GlcNAc via GlcNAc transferase I (making it endo D resistant/endo H sensitive). This product, and only this one, is the substrate for α-mannosidase II, which removes two of the remaining five Man residues, specifically the α1-3Man and α1-6Man that form the upper branch. Once this occurs, this endo D–resistant molecule also becomes endo H resistant. This structure is also a substrate for GlcNAc transferase II on the way to forming a biantennary chain. The second pathway involves α-mannosidase III, which was identified as a functionally important enzyme in nearly all tissues when the α-mannosidase II pathway was ablated in mice (Chui et al., 1997). After α-mannosidase I removes the final α1-2Man from
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Man6GlcNAc2 to generate Man5GlcNAc2, αmannosidase III cleaves the same two Man residues as α-mannosidase II; however, αmannosidase III cleavage does not require addition of the first GlcNAc that α-mannosidase II requires. α-Mannosidase III creates the endo D–sensitive/endo H–resistant trimannosyl core. This sugar chain serves as an acceptor for GlcNAc transferase I to form an endo D–resistant/endo H–resistant molecule that can be modified by GlcNAc transferase II, and then on to biantennary chains. As summarized in Figure 12.4.2, using the two endoglycosidases together will distinguish whether the oligosaccharide was processed by α-mannosidase II or α-mannosidase III. This may be important since there is evidence that distribution of the two enzymes in the Golgi complex is different with α-mannosidase III in an earlier compartment. It is important to point out that all chains of even a single protein are not necessarily processed using all the same enzymes. Mutant cell line CHO Lec1 lacks GlcNAc transferase I activity and cannot synthesize either complex or hybrid chains. These cells can be used to measure the kinetics of acquiring endo D sensitivity (Beckers et al., 1987). This cell line can be obtained through ATCC (CRL-1735) . The chains become permanently endo D sensitive, because they lack GlcNAc transferase I and cannot be converted back to an endo D–resistant form. These chains will also remain endo H sensitive, because α-mannosidase II requires GlcNAc transferase I. However, if any chains are processed by αmannosidase III, they would be become endo H resistant. Again, the combination of endo D and endo H digestions can reveal which pathway was used. CHO Lec1 cells are also useful for tracking the movement of a protein from the ER into the earliest Golgi compartment where α-mannosidase I is located. Acquisition of endo D sensitivity requires the action of this enzyme. The advantage of using Lec1 cells is that the proteins remain permanently endo D sensitive and there is no risk of kinetically missing that small window of sensitivity before the sugar chain might become endo D resistant once again. Even if α-mannosidase III acts on the protein, it would still remain endo D sensitive, and no further processing would occur.
Endoglycosidase F and peptide N-glycosidase F Elder and Alexander (1982; Alexander and Elder, 1989) made a landmark discovery when they identified an enzyme in culture filtrates of the bacterium Flavobacterium meningosepticum that cleaved N-linked oligosaccharides from glycoproteins. This preparation had a broad substrate specificity. The endoglycosidase activity in this preparation (endo F) was actually due to a set of enzymes, each with a more restricted substrate range. Like endo H, the endo F enzymes cleave the sugar chains between the two core GlcNAc residues (Fig. 12.4.5). Endo F was originally thought to be a single enzyme, but it is now known that each of the three enzymes has a distinct specificity (Plummer and Tarentino, 1991). The specificity of endo F1 is very similar to that of endo H, while endo F2 prefers biantennary chains, and endo F3 will cleave core fucosylated biantennary and triantennary, but not tetraantennary, chains (Fig. 12.4.2 and Fig. 12.4.3). All are commercially available from Glyko. Plummer et al. (1984) carefully analyzed Flavobacterium filtrates and found that the very broad substrate range was actually due to a glycoamidase activity rather than an endoglycosidase activity. The glycoamidase releases the entire carbohydrate chain from the protein by cleaving the Asp-GlcNAc bond (Fig. 12.4.5). The enzyme is called by various names, including peptide:N-glycosidase F (PNGase F), glycopeptidase F, and N-glycanase (previously available from Genzyme), but the proper name is peptide N-4(N-acetyl-β-glucosaminyl)asparagine amidase F. PNGase F has the broadest specificity, and it releases most of the N-linked oligosaccharide chains from proteins. Endo F2 and Endo F3 Endo F2 prefers biantennary chains over high-mannose chains by ∼20 fold. Thus, endo H and PNGase F are better choices for broadly distinguishing high-mannose from complex chains as described above in the endo H protocol (Tarentino and Plummer, 1994). Many proteins have core-fucosylated Nlinked glycans, and the addition of fucose can be used as an additional trafficking marker. Endo F3 will hydrolyze triantennary chains, but endo F2 will not. Endo F2 hydrolyzes biantennary chains; however, endo F3 will also hydrolyze core-fucosylated biantennary chains only
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a bit more slowly than it does triantennary chains. Thus, if all the chains on a protein are sensitive to endo F2 (biantennary) and to endo F3, this is evidence for the presence of a core fucose on those chains. The enhancement of endo F3 activity on biantennary chains with a core α1-6Fuc points out that some specificities are really a matter of relative rates of cleavage. If both endo F2 (biantennary) and endo F3 (triantennary and biantennary with core fucose) cleave the protein, they may be acting on different chains on the same molecule. If there is only a single chain, repeat the experiment under the same conditions using 10- to 20-fold dilutions of each enzyme. If both still cleave the chain about equally, it is evidence for core fucosylation of a biantennary chain. PNGase F PNGase F has the broadest specificity of all the enzymes that cleave N-linked oligosaccharides. It is indifferent to all extended structures on the chains, such as sulfate, phosphate, polylactosamines, polysialic acids, and even the occasional glycosaminoglycan chain. Most of the modifications in the Man3GlcNAc2 core region also make no difference in chain cleavage. The only oligosaccharide structural feature that confers PNGase F resistance is the presence of an α1-3Fuc on the GlcNAc bound to Asn (Tretter et al., 1991). This modification is commonly found in plants and in some insect glycoproteins, but it is rare in most mammalian cell lines. However, caution is warranted, as there is evidence that some mammalian cells do have the critical α1-3Fuc transferase, and some studies show that a majority of N-linked chains of bovine lung are actually PNGase F resistant! It is not known how common this resistance may be. It is thus important to document N-glycosylation with proteins still in the ER (see Support Protocol) before they might be processed to a PNGase F–resistant form.
Release of N-Linked Oligosaccharides
Sialidase Sialidases are also called neuraminidases because the most common form of sialic acid is N-acetyl neuraminic acid. The sialic acids are a family with over forty different members, but fortunately the very great majority of them can be removed from the oligosaccharides by the broad-spectrum sialidase from Arthrobacter ureafaciens (AUS). It can even digest polysialic acids, a rare modification found on only a few proteins such as neuronal cell adhesion molecule (NCAM). Sialidases with selected speci-
ficities from other sources are available but would not usually be needed. AUS has an optimum pH of 5.0, with ∼30% of maximum activity at pH 7. Because sialic acids are charged, they affect gel mobility of proteins more than would be expected from their nominal molecular weight. The magnitude of the gel shift depends on the number of residues. It is difficult to estimate their number by sialidase digestion, but the mobility change is usually sufficient if there are several sialic acid residues. On the other hand, if a protein has only one sialic acid, its presence could be missed using standard one-dimensional SDS-PAGE. To be certain of the effects of sialidase, the sample can be analyzed by a two-dimensional system, using IEF or NEPHGE in the first dimension (UNIT 10.4). The loss of even a single sialic acid will be evident because it changes the isoelectric point. AUS will remove sialic acids from both Nand O-linked chains, so the type of chain carrying them must be determined independently using PNGase F or possibly O-glycosidase in combination with sialidase. A protein will generally be partially or completely resistant to O-glycosidase because the required disaccharide, Galβ1-3GalNAc, is usually extended and often sialylated. Until the sialic acid is removed, it will be resistant. The presence of sialic acid (sialidase sensitivity) is often used as an indication of the transport of a protein into the trans-Golgi network (TGN). This may be true in general, but it is important to remember that the distribution of Golgi enzymes is cell type dependent. For instance, α-mannosidases I and II, which are typically considered cis/medial Golgi enzymes, are strongly expressed on the brush border of enterocytes—hardly a Golgi compartment. There are other similar examples of various distributions of sialyl transferase. Moreover, there are different sialyl transferases and each may have its own unique distribution. Although one should be cautious, it is probably safe to place sialyl transferase in the late Golgi compartment rather than an early one. Endo-α-N-acetylgalactosaminidase This enzyme from Diplococcus pneumoniae also goes by various names, including O-glycosidase and O-Glycanase. The last name is a trade name from Genzyme, which no longer sells enzymes; the enzyme is now available from Glyko. This enzyme has a narrow substrate range and cleaves only Galβ1-3GalNAcα-Ser/Thr. These are only the first two
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sugars added in the diverse O-linked pathway that can produce glycans with a dozen or more sugar units. A portion of the pathway is shown in Figure 12.4.4. Fortunately, many, but far from all, O-linked chains have the simple trisaccharide structure and would be sensitive to cleavage after removing the Sia. Thus, sequential individual digestions or mixed digestions can be used. As both Gal and GalNAc (and probably Sia) are added in the early Golgi complex, sensitivity to the enzyme shows that the protein carries O-linked chains, but matching enzyme sensitivity and a Golgi compartment to further chain extension is difficult. Combining a battery of exoglycosidases (sialidase, α-fucosidase, α-N-acetylgalactosaminidase, and β-hexosaminidase) with endo-α-Nacetylgalactosaminidase will probably remove most O-linked sugar chains, except sulfated ones. The bottom line is that it is easy to use the enzyme in combination with sialidase to show that a protein has simple O-linked chains, but it is difficult to conclude much more concerning either the structure of the sugar chain or intracellular trafficking. Endo-β-galactosidase Bacterioides fragilis endo-β-galactosidase is one of several enzymes that specifically degrade polylactosamines by cleaving linear chains of GlcNAcβ1-3Galβ1-4 repeats at the Galβ1-4 linkage. Any substitution on the galactose itself blocks cleavage; however, modifications of the neighboring sugars can slow hydrolysis (Fig. 12.4.7). For instance, fucosylation and/or sulfation of nearby GlcNAc slows cleavage, but chain branching or sulfation at Gal block it. Even with these potential complexities, digestion with endo-β-galactosidase will sharpen a broad band even if it does not cleave every linkage. The repeating GlcNAcβ1-3Galβ1-4 units can be found on both N- and O-linked chains, so sensitivity to PNGase F digestion can potentially distinguish the location. Lysosome-associated membrane protein 1 (LAMP-1) has polylactosamine repeats on N-linked chains. Remember that glycosylation is not template driven, so oligosaccharides often exist as a continuum of different structures on individual proteins. For example, heavily sulfated polylactosamine repeats are also known as keratan sulfate and are degraded by keratanases. O-Sialoglycoprotease O-Sialoglycoprotease (also called O-glycoprotease or O-sialoglycoprotein endopepti-
dase; Mellors and Lo, 1995) is a neutral metalloprotease produced by Pasteurella haemolytica. This enzyme requires clusters of sialylated oligosaccharides on Ser or Thr residues (Norgard et al., 1993). Having a single sialylated O-linked sugar chain on a protein will not lead to degradation, nor will having a nonsialylated sugar chain. Therefore, this enzyme can be used in a typical pulse-chase experiment to indicate whether there are tightly grouped Olinked chains and when they are sialylated. Adding the initial α-GalNAc in the early Golgi complex is insufficient to cause proteolysis; sialylation is specifically required, and this may occur in a later Golgi compartment. Proteolysis can generate smaller fragments of a target protein that are still visible on gels, or the fragments may be so small that they are not even seen on the gels. It depends on the protein. Many leukocyte antigens such as the P-selectin ligand, or others such as CD43, CD44, CD45, and CD34 found on hematopoietic stem cells, are all substrates. As sialic acids are found on both N- and O-linked chains (in clusters and not), sequential digestions using PNGase F, sialidase, and O-sialoglycoprotease endopeptidase in different orders can reveal different kinds of sialylated glycans. If used as part of a pulse-chase protocol, they can reveal different kinetic subcompartments. Not all sialyl transferases are necessarily within the same Golgi compartment.
Critical Parameters For nearly all of the digestions, complete denaturation of the protein can be important, as maximum deglycosylation occurs only when the sugar chains of glycoproteins are completely exposed. This is not important for sialidase digestions since sialic acids are exposed. Usually endo H digestions will work without full denaturation, but the unprocessed highmannose chains remain unprocessed because they are often less exposed to the processing enzymes. This may or may not be true of the target protein, but it is better to be safe and denature it completely before digestion. Heating the immunoprecipitate with 0.1 M 2-mercaptoethanol/0.1% SDS is the best way to release the protein in a denatured state. Assuming that the immunoprecipitate contains <10 µg of protein, 30 µl of 0.1% SDS still provides a three-fold excess over the protein. Adding SDS presents another problem: free SDS may denature the digesting enzyme before it has a chance to finish its job. For most enzymes, be sure that the free SDS concentration is <0.01%. The best
Post-Translational Modification: Glycosylation
12.4.23 Current Protocols in Protein Science
Supplement 22
way to do this is to add a 10- to 20-fold weight excess of nonionic detergent with a low critical micellar concentration (e.g., Triton X-100 or NP-40). These detergents will form mixed micelles with the free SDS and keep it from denaturing the added enzymes. The amount of enzyme and the incubation time recommended in the protocols are in excess and should be sufficient to cleave any of the sensitive linkages. The incubation times can be shortened, if necessary, but it is better to keep the enzyme concentration as indicated. Many of the digestions (e.g., sialidase, Osialoglycoprotease) can be adapted for use on membrane preparations or on live cell surfaces by simply omitting the ionic and nonionic detergents and decreasing the incubation time. The problem is that some linkages may not be exposed and/or sensitive to the digestion. Thus, the usefulness of this approach needs to be determined on a case-by-case basis. Endo H Endo H has a broad pH optimum between 5.5 and 6.5, and phosphate or citrate/phosphate buffers can be used in place of citrate. Endo H is very stable to proteases, freezing and thawing, and prolonged incubations. No additives are required for storage of the enzyme. At concentrations below 5 to 10 µg/ml (200 to 400 mU/ml), endo H will bind to glass, so it should be stored in plastic vials (e.g., screw-cap microcentrifuge tubes). Endo D Endo D has a broad pH optimum of 4 to 6.5. One unit of enzyme activity will cleave 1 µmole of a Man5GlcNAc3 glycopeptide per min at 37°C . I t is sup plied from Boehringer Mannheim as a powder containing 0.1 U of activity. Adding 0.1 ml of water gives a 20 mM phosphate buffer, pH 7, 0.05% sodium azide, and 5 mg/ml BSA. It is stable for 3 months at 4°C or at −20°C, but freezing and thawing should be avoided. There may be a slight contamination (<0.2%) with β-N-acetylglucosaminidase activity.
Release of N-Linked Oligosaccharides
Endo F Commercial endo F preparations are mixtures of endo F1, F2, and F3. Endo F preparations should not be used for routine deglycosylation or to draw conclusions about the structure of the released oligosaccharides unless the specificity is clearly defined.
Endo F2 Endo F2 has a broad pH optimum of 4 to 6 and retains >50% of its activity at pH 7. The enzyme is sensitive to SDS, but adding nonionic detergents prevents denaturation of the enzyme by SDS. Although the enzyme is stable at 4°C for months, it can be frozen in aliquots at −70°C as long as repeated freeze/thaw cycles are avoided. The 1,10-phenanthroline can be used to inhibit a trace of a zinc metalloprotease that may be present. PNGase F The pH optimum for PNGase F is 8.6, but 80% of full activity occurs between 7.5 to 9.5 with a range of buffers including phosphate, ammonium bicarbonate, Tris⋅Cl, and HEPES. Borate buffers inhibit the enzyme. Commercial PNGase F is endo F free and is stable for 6 months at 4°C, or indefinitely at −70°C. However, it should be stored in small aliquots and repeated freeze/thaw cycles should be avoided. PNGase F will bind to glass and plastic surfaces and should not be stored in dilute solutions (<0.1 mU/ml). All of the unit activities of commercial preparations are based on cleavage of dansylated glycopeptides; they are expressed in nmoles/min, which are actually mU, not true International Units (1 International Unit = 1 µmole/min). SDS inactivates PNGase F, but adding a ten-fold weight excess of nonionic detergents protects the enzyme (Tarentino et al., 1989; Tarentino and Plummer, 1994). Sialidase AUS is available as a lyophilyzed powder from typical commercial sources such as Sigma, Boehringer Mannheim, and Glyko. It should be reconstituted in water at 1 to 10 mU/µl according to manufacturer’s directions. It is stable for 6 months at 4°C. Treatment with sialidase is also used in assays of protein transport to the cell surface. Endo-α-N-acetylgalactosaminidase (O-glycosidase) Endo-α-N-acetylgalactosaminidase has a pH optimum of 6.0 and has 50% activity at 5.5 to 7.0. The thiol inhibitor parachloromercuric benzoate (PCMB; 1 mM) inactivates the enzyme, and 1 mM EDTA inhibits it (63%), as do Mn2+ and Zn2+ (50%). Chloride also inhibits the enzyme, so HCl-containing buffers should be avoided. The enzyme will have full access to the sugar chains only after denaturating the protein with SDS, but the excess SDS needs to be removed by forming mixed micelles with
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Current Protocols in Protein Science
nonionic detergents. The enzyme is stable at 4°C and at −20°C, but freeze/thaw cycles should be avoided. Endo-â-galactosidase The enzyme is supplied by several commercial sources, including Glyko and Sigma. The enzyme is free of contaminating endo- and exoglycosidases. It has a pH optimum of 5.8 and should be stored at −20°C, but is stable for ≥2 months at 4°C. Glyko supplies the enzyme as a lyophilyzed powder with BSA for stability. EDTA, Ca2+, Mn2+, and Mg2+ do not affect stability or activity, but PCMB inactivates it. O-Sialoglycoprotease The partially purified enzyme is supplied by Accurate Chemical and Scientific as a lyophilyzed powder containing nonsubstrate bovine serum proteins and HEPES buffer. The enzyme should be reconstituted according to the manufacturer’s instructions, divided into aliquots appropriate for a single use, and stored at −20°C. Freeze/thaw cycles inactivate the enzyme, and it is inhibited by EDTA or 1,10phenanthroline. It is possible to check the activity with a positive control of glycophorin A, which is available through Sigma.
1 . The smallest N-linked chain (Man3GlcNAc2) will have a mass of ∼0.9 kDa. 2. Two sialic acids on a single N-linked biantennary chain will have a mass of ∼0.6 kDa, but their loss may appear larger. If they occur on clustered O-linked chains (sialoglycoprotease sensitive), the apparent size difference will be even larger. 3. Most polylactosamines are three or more repeats, and therefore their mass would be ∼1 kDa. The protein will probably run as a heterogeneous smear or broad band before digestion. 4. A single O-glycosidase-sensitive disaccharide (0.4 kDa) may be below detection limits.
Time Considerations All digestions can be done overnight for convenience, but the amount of enzymes should be sufficient for complete digestion in less time. The gels are run the next day, but the time needed for the development of autoradiograms will depend on the strength of the signal. A low-abundance protein labeled for 10 to 30 min with [35S]Met may give a weak signal and require long exposures (e.g., 2 weeks). Trafficking of abundant glycoproteins such as viral coat proteins require only short exposure times (e.g., a few hours).
Troubleshooting Most of the procedures should work as described, but there is a chance that the enzyme is inactive because of a variety of factors such as age, poor storage, or excess SDS. To check activity, it is worthwhile to run a positive control digestion using the same solutions including SDS and nonionic detergents as for the samples. Since the positive controls are simply glycoproteins that are visualized by Coomassie or silver staining, this requires running a separate gel for staining. This should not be required on a routine basis if the enzymes are used and stored as directed. The most likely culprit in failed digestions is using SDS solutions that are too old or too impure.
Anticipated Results If the digestions are effective, the labeled band will usually show increased mobility on the gel. In rare instances, digestions can actually decrease mobility. The amount of change will depend on the contribution of that component to the overall mass of the protein. As mentioned before, a gel system that allows visualization of a 1-kDa change should be used. Proteins that are >100 kDa may cause problems for fine resolution. Here are a few numbers to keep in mind.
Literature Cited Adams, L.D. and Gallagher, S.R. 1992. Two-dimensional gel electrophoresis using the O’Farrell system. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 10.4.1-10.4.13. John Wiley & Sons, New York. Alexander, S. and Elder, J.H. 1989. Endoglycosidases from Flavobacterium meningosepticum: Application to biological problems. Methods Enzymol. 179:505-518. Beckers, C.J.M., Keller, D.S., and Balch, W.E. 1987. Semi-intact cells permeable to macromolecules: Use in reconstitution of protein transport from the endoplasmic reticulum to the Golgi complex. Cell 50:523-534. Chui, D., Oh-Eda, M., Liao, Y.-F., Panneerselvam, K., Lal, A., Marek, K.W., Freeze, H.H., Moremen, K.W., Fukuda, M.N., and Marth, J.D. 1997. Alpha-mannosidase-II deficiency results in dyserythropoiesis and unveils an alternate pathway in oligosaccharide biosynthesis. Cell 90:157-167. Davidson, H.W. and Balch, W.E. 1993. Differential inhibition of multiple vesicular transport steps between the endoplasmic reticulum and trans Golgi network. J. Biol. Chem. 268:4216-4226.
Post-Translational Modification: Glycosylation
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Elder, J.H. and Alexander, S. 1982. Endo-β-N-acetylglucosaminidase F: Endoglycosidase from Flavobacterium meningosepticum that cleaves both high mannose and complex glycoproteins. Proc. Natl. Acad. Sci. U.S.A. 79:4540-4544. Kornfeld, R. and Kornfeld, S. 1985. Assembly of asparagine-linked oligosaccharides. Annu. Rev. Biochem. 54:631-664. Mellors, A. and Lo, R.Y. 1995. O-Sialoglycoprotease from Pasteurella haemolytica. Methods Enzymol. 248:728-740. Norgard, K.E., Moore, K.L., Diaz, S., Stults, N.L., Ushiyama, S., McEver, R.P., Cummings, R.D., and Varki, A. 1993. Characterization of a specific ligand for P-selectin on myeloid cells. A minor glycoprotein with sialylated O-linked oligosaccharides. J. Biol. Chem. 268:12764-12774. Plummer, T.H. Jr. and Tarentino, A.L. 1991. Purification of the oligosaccharide-cleaving enzymes of Flavobacterium meningosepticum. Glycobiology 1:257-263. Plummer, T.H. Jr., Elder, J.H., Alexander, S., Phelan, A.W., and Tarentino, A.L. 1984. Demonstration of peptide:N-glycosidase F activity in endo-β-Nacetylglucosaminidase F preparations. J. Biol. Chem. 259:10700-10704. Tarentino, A.L. and Plummer, T.H. Jr. 1994. Enzymatic deglycosylation of asparagine-linked glycans: Purification, properties, and specificity of oligosaccharide-cleaving enzymes from Flavobacterium meningosepticum. Methods Enzymol. 230:44-57.
Tretter, V., Altmann, F., and März, L. 1991. PeptideN4-(N-acetyl-β-glucosaminyl) asparagine amidase F cannot release glycans with fucose attached α1→3 to the asparagine-linked N-acetylglucosamine residue. Eur. J. Biochem. 199:647-652. Viitala, J., Carlsson, S.R., Siebert, P.D., and Fukuda, M. 1988. Molecular cloning of cDNAs encoding lamp A, a human lysosomal membrane glycoprotein with apparent Mr approximately equal to 120,000. Proc. Natl. Acad. Sci. U.S.A. 85(11):3743-3747.
Key References Beckers et al., 1987. See above. Describes the use of Lec 1 CHO cells and endo D to study processing. Chui et al., 1997. See above. Demonstrates the importance of α-mannosidase III. Kornfeld and Kornfeld, 1985. See above. Landmark review of processing. Tarentino and Plummer, 1994. See above. Best and most recent review of the use of these enzymes.
Contributed by Hudson H. Freeze The Burnham Institute La Jolla, California
Tarentino, A.L., Trimble, R.B., and Plummer, T.H. Jr. 1989. Enzymatic approaches for studying the structure, synthesis, and processing of glycoproteins. Methods Cell Biol. 32:111-139.
Release of N-Linked Oligosaccharides
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Detection of Glycophospholipid Anchors on Proteins
UNIT 12.5
Many eukaryotic proteins are tethered to the plasma membrane by glycosyl phosphatidylinositol (GPI) membrane anchors. Figure 12.5.1 schematically depicts the minimal (core) structure common to all GPI protein anchors characterized to date. Detailed structural information is described in reviews by Englund (1993) and McConville and Ferguson (1993). This unit provides a general approach for detecting GPI-anchored proteins. First, the detergent-partitioning behavior of a protein of interest is examined for characteristics of GPI-linked species. The partitioning of total cellular and isolated proteins with Triton X-114 is described in Basic Protocol 1 and Alternate Protocol 1, respectively. Precondensation of Triton X-114, which is necessary to remove hydrophilic contaminants before partitioning, is outlined in Support Protocol 1. The protein may also be subjected to specific enzymatic or chemical cleavages to release it from its GPI anchor. Phospholipase cleavage (starting with intact cells or membranes, or with isolated protein, respectively) is detailed in Basic Protocol 2 and Alternate Protocol 2, and chemical cleavage with nitrous acid is described in Basic Protocol 3. If GPI-anchored proteins are radiolabeled with fatty acids, it facilitates the detection of the GPI protein products following the cleavage reactions. Separation of lipid moieties released from proteins is described in Support Protocol 3 and base hydrolysis of proteins is presented in Basic Protocol 4. Figure 12.5.2 outlines the relationships between the various protocols.
N
Protein
C
EthN
P
Man
Man
Man
GlcN HONO Inositol GPI-PLD P
PI
PI-PLC, GPI-PLC
PA DAG
Figure 12.5.1 Schematic representation of the glycan core structure common to all GPI anchors. The sites of cleavage of phospholipase C enzymes (PI-PLC, GPI-PLC), phospholipase D (GPIPLD), and nitrous acid (HONO) are indicated, as are lipid products resulting from these cleavages: PI (phosphatidylinositol), PA (phosphatidic acid), and DAG (diacylglycerol). Other abbreviations: Man, mannose; GlcN, glucosamine; EthN, ethanolamine; P, phosphate group. Phospholipase C treatment generates inositol cyclic phosphate, the major epitope contributing to the cross-reacting determinant (CRD). Lipid products may vary from those depicted, depending on features of the anchor (see Background Information).
Glycosylation
Contributed by Tamara L. Doering, Paul T. Englund, and Gerald W. Hart
12.5.1
Current Protocols in Protein Science (1995) 12.5.1-12.5.14 Copyright © 2000 by John Wiley & Sons, Inc.
Supplement 2
cells, membranes radiolabeling (UNIT 3.3)
proteins
Triton X-114 partitioning (Basic Protocol 1 and Alternate Protocol 1) phospholipase digestion (basic Protocol 2 and Alternate Protocol 2)
anti-CRD Ab detection (Support Protocol 2)
one-dimensional gel electrophoresis (UNIT 10.1) immunoblotting immunoprecipitation
nitrous acid cleavage (Basic Protocol 3)
separation of lipid moiety (Support Protocol 3)
Triton X-114 partitioning
immunoblotting immunoprecipitation
Triton X-114 partitioning immunofluorescence
base hydrolysis (Basic Protocol 4)
autoradiography
TLC
Figure 12.5.2 Flow chart for the detection of GPI-anchored proteins.
STRATEGIC PLANNING The techniques described below require that a method for identifying the protein of interest be available. Proteins may be visualized after one-dimensional gel electrophoresis (UNIT 10.1) and Coomassie blue or silver staining (UNIT 10.5), or they may be identified by immunoprecipitation or immunoblotting using available specific antibodies. Enzymes may also be detected by their known activities (e.g., see Kodukula et al., 1992). It is convenient, particularly for low-abundance species, to use proteins that are labeled with radioactive amino acids (UNIT 3.3) or GPI components (see Fig. 12.5.1). The ability to radiolabel proteins with GPI precursors such as fatty acids, ethanolamine, myo-inositol, and certain monosaccharide precursors (UNIT 12.2) is in itself suggestive of GPI anchorage. Because GPI-anchored proteins commonly represent a small fraction of eukaryotic glycoconjugates, metabolic radiolabeling may be inefficient. Tunicamycin may be used to increase relative incorporation of monosaccharide precursors into GPIs. Detection of Glycophospholipid Anchors
Rigorous identification of specific GPI anchor structures requires detailed analysis. These methods, which are beyond the scope of this unit, are described in Ferguson (1992).
12.5.2 Supplement 2
Current Protocols in Protein Science
EXTRACTION AND PARTITIONING OF TOTAL PROTEINS FROM CELLS OR MEMBRANES WITH TRITON X-114
BASIC PROTOCOL 1
Extraction of cells or tissues with the detergent Triton X-114 provides a profile (and potentially an initial purification) of amphiphilic proteins, including integral membrane proteins and species bearing GPI anchors. Once a GPI-containing protein is released from the lipid component of its anchor, it will no longer partition into the detergent-enriched phase. This alteration of partitioning behavior provides a rapid assay for the presence or absence of anchors. It may be used to monitor the changes in proteins produced by the other procedures. In this protocol, cells or membrane fractions are first resuspended in TBS, then incubated on ice with Triton X-114. The mixture is then warmed and separates into two proteincontaining phases due to aggregation of detergent micelles (see Background Information). The upper (detergent-depleted) phase contains hydrophilic proteins; amphiphilic proteins (including those with GPI anchors) partition to the lower (detergent-enriched) phase. Both phases are then analyzed to determine the group to which each protein belongs. Materials Cells, membrane fraction, or other source of protein Tris-buffered saline (TBS; see recipe), ice-cold Precondensed Triton X-114 stock solution in TBS (see Support Protocol 1), ice-cold 15-ml polypropylene centrifuge tubes Centrifuges: low-speed (tabletop) and equipped with appropriate rotor (e.g., SS-34), at 4°C and room temperature 1. Resuspend cells (or membranes) in ice-cold TBS to a final protein concentration of ≤4 mg/ml in a 15-ml tube. The amount of protein required is that which will give detectable levels at the end of the procedure, which will depend on the system the individual researcher is using. As a guideline, use several times the amount of protein that is believed to be needed, to account for partitioning and losses (see Critical Parameters).
2. Add 1⁄5 vol precondensed Triton X-114 stock solution (∼2% final concentration). 3. Extract cells by incubating 15 min on ice with occasional mixing. At this point in the procedure there will only be one phase.
4. Centrifuge cell mixture 10 min at 10,000 × g (9000 rpm in an SS-34 rotor), 4°C, and transfer supernatant to a fresh tube. Resuspend pellet in ice-cold TBS and save on ice for analysis in step 7. The cold supernatant fraction contains both soluble and detergent-extracted proteins; the latter group includes proteins anchored by GPI structures or by transmembrane polypeptide domains. The pellet contains additional GPI species, as well as cell elements insoluble in nonionic detergents.
5. Warm supernatant fraction to 37°C in a water bath; the solution will become cloudy as it reaches 37°C. 6. Centrifuge the solution in a tabletop centrifuge 10 min at 1000 × g, room temperature. Collect upper and lower phases in separate tubes. The upper (detergent-depleted) phase contains soluble proteins; the lower (detergent-enriched) phase contains proteins anchored by GPI structures or by transmembrane domains. Glycosylation
12.5.3 Current Protocols in Protein Science
Supplement 2
7. Analyze the resuspended pellet from step 4 and each phase from step 6 for the proteins of interest. Analysis may be by an activity assay, one-dimensional gel electrophoresis and staining of proteins (UNITS 10.1 & 10.5), or immunoprecipitation or immunoblotting using specific antibodies (see Strategic Planning for discussion). ALTERNATE PROTOCOL 1
PARTITIONING OF ISOLATED PROTEINS WITH TRITON X-114 A modification to the basic protocol for Triton X-114 extraction of proteins from whole cells or membranes is used to partition isolated proteins (see Basic Protocol 1). 1. Dilute or dissolve the protein in 1 ml ice-cold TBS in a 1.5-ml microcentrifuge tube. Add 0.2 ml Triton X-114 and mix. See Critical Parameters for guidelines on the total amount of protein to use.
2. Warm protein mixture several minutes (or until it becomes cloudy) in a 37°C water bath. Microcentrifuge 5 min at maximum speed. 3. Collect the upper and lower phases and analyze each phase for the protein(s) of interest. SUPPORT PROTOCOL 1
PRECONDENSATION OF TRITON X-114 DETERGENT Triton X-114 often contains hydrophilic contaminants, and must be precondensed by several rounds of phase separation before use. Materials Triton X-114 detergent Tris-buffered saline (TBS; see recipe) 50-ml centrifuge tubes Tabletop centrifuge 1. Dissolve 1.5 g Triton X-114 in 50 ml TBS, place in a 50-ml centrifuge tube, then chill on ice. At this point, the solution will be clear.
2. Warm the solution to 37°C in a water bath. Once the solution has warmed to the bath temperature, it will become turbid.
3. Centrifuge the solution in a tabletop centrifuge 10 min at 1000 × g, room temperature. 4. Remove and discard the upper (detergent-depleted) phase and redissolve the lower (detergent-enriched) phase in an equal volume of ice-cold TBS. Hydrophilic contaminants partition into the upper phase.
5. Repeat partitioning (steps 2 to 4) three times. The final detergent-enriched phase contains ∼12% detergent. It may be stored at 4°C for routine use or frozen for long-term storage.
Detection of Glycophospholipid Anchors
12.5.4 Supplement 2
Current Protocols in Protein Science
IDENTIFICATION OF GPI-ANCHORED PROTEINS BY PI-PLC DIGESTION OF INTACT CELLS
BASIC PROTOCOL 2
Phospholipase C (PLC) cleaves within the GPI membrane anchors of glycoproteins (see Fig. 12.5.1), causing the release of protein from cell membranes. In this protocol, intact cells or cell membranes are digested with phosphatidylinositol-specific phospholipase C (PI-PLC). Following incubation with the enzyme, the cell mixture is centrifuged and the resulting pellet and supernatant are analyzed for the presence of the protein of interest. Proteins released from GPI anchors are found in the supernatant; lack of release may indicate absence of a GPI anchor, modification of the GPI protein, or other factors (see Background Information). Materials Cells (or membrane preparation) Bacterial phosphatidylinositol-specific phospholipase C (PI-PLC; e.g., from Bacillus thuringiensis, Oxford GlycoSystems) Hanks balanced salt solution (HBSS; e.g., Life Technologies), buffered saline, or culture medium Tabletop centrifuge and appropriate centrifuge tubes 1. Obtain cells (or membranes) of interest in dispersed suspension. Wash duplicate aliquots twice in HBSS, buffered saline, or culture medium, then resuspend in any of these solutions. The final protein concentration of the samples (typically 0.1 to 0.5 mg/ml) should be based on supplier specifications for the enzyme preparation used. See Critical Parameters for guidelines on the total amount of protein to use.
2. Add bacterial PI-PLC to one aliquot. Add no enzyme to the second aliquot (control). Incubate tubes 1 hr at 37°C. The amount of enzyme should be determined by supplier recommendations.
3. Centrifuge cells in a tabletop centrifuge 5 min at 1000 × g. Remove each supernatant to a fresh tube and save for analysis in step 4. Resuspend pellet in a volume of buffered saline equal to the volume of supernatant removed. 4. Assay supernatant and pellet fractions for protein(s) of interest. The proteins may be assayed by Triton X-114 detergent partitioning (see Basic Protocol 1), one-dimensional SDS-PAGE (UNIT 10.1) followed by Coomassie blue or silver staining (UNIT 10.5), immunoprecipitation or immunoblotting.
IDENTIFICATION OF GPI ANCHORAGE BY PHOSPHOLIPASE TREATMENT OF ISOLATED PROTEINS
ALTERNATE PROTOCOL 2
Several specific phospholipases that effectively degrade only GPI species are useful for analyzing GPI anchorage of isolated proteins (see Background Information). In this protocol, aliquots of the protein are digested in parallel with the following phospholipases: bacterial phosphatidylinositol-specific phospholipase C (PI-PLC), a GPI-specific phospholipase C (GPI-PLC) from Trypanosoma brucei, and a GPI-specific phospholipase D (GPI-PLD) from mammalian serum (rat, rabbit, or human). Proteins are partitioned by Triton X-114 extraction (see Alternate Protocol 1), then analyzed. Specific cleavage of proteins by phospholipase treatment strongly indicates GPI anchorage.
Glycosylation
12.5.5 Current Protocols in Protein Science
Supplement 2
Materials Isolated protein (see Basic Protocol 1) Acetone, −20°C Appropriate enzyme buffer: GPI-PLD buffer, GPI-PLC buffer, and PI-PLC buffer (see recipes) Phospholipase enzymes: GPI-PLD from rat, rabbit, or human whole serum, GPI-PLC from Trypanosoma brucei (Oxford GlycoSystems), and PI-PLC from Bacillus thuringiensis (Oxford GlycoSystems) or B. cereus (Boehringer Mannheim or Sigma) Tris-buffered saline (TBS; see recipe) Precondensed Triton X-114 solution (see Support Protocol 1) Centrifuge and rotor (e.g., SS-34) 1. If the protein sample is lyophilized or concentrated in a buffer similar to the phospholipase buffers, begin at step 3. If it is not, acetone-precipitate the protein by adding 8 vol of −20°C acetone to the protein solution and incubating it ≥3 hr at −20°C. See Critical Parameters for guidelines on the total amount of protein to use. For low-abundance material, efficiency of precipitation may be increased by including 15 µg of a carrier protein (e.g., cytochrome c or bovine serum albumin).
2. Centrifuge the sample 20 min at 15,000 × g (∼11,000 rpm in an SS-34 rotor), 4°C. Aspirate supernatant and allow pellet to air dry. 3. For each enzyme to be tested, dissolve two aliquots of the protein in 100 µl of the appropriate enzyme buffer in 1.5-ml microcentrifuge tubes. 4. To one aliquot from each pair add the appropriate enzyme. Add an equal volume of the appropriate enzyme buffer to the second aliquot (control). Mix tubes and incubate 1 hr at 37°C. The amount of enzyme added should be based on the activity of the lipase preparation (see supplier specifications) and the estimated amount of protein present. It may be desirable to titrate the enzyme, using various amounts in parallel reactions, to be sure that all sensitive proteins are completely digested. GPI-PLD is not generally available, but whole serum (rat, rabbit, or human) may serve as a source of GPI-PLD activity. Aliquots of serum may be stored frozen and thawed before use. For the assay, use 2 µl whole serum in a 100-µl reaction.
5. Dilute reaction with 200 µl ice-cold TBS, then add 60 µl Triton X-114. Chill 15 min on ice. 6. Partition the proteins (see Alternate Protocol 1, steps 2 and 3). GPI-anchored proteins partition into the detergent-enriched phase of Triton X-114 in the control reaction (uncleaved) and into the detergent-depleted phase after cleavage of the GPI anchor by phospholipase.
7. Analyze each phase for the presence of protein(s) of interest (see Basic Protocol 1, step 7). Alternatively, PI-PLC and GPI-PLC cleavage products can be detected by reactivity with anti-CRD antibody (see Support Protocol 2). In addition, if the anchor is radiolabeled with fatty acids (UNIT 12.2), cleavage may be assessed by analysis of radiolabeled lipid products.
Detection of Glycophospholipid Anchors
Migration of phospholipase-digested proteins on electrophoretic gels is not predictable and should not be used alone to assess cleavage. Either no shift or a shift in either direction (relative to uncleaved protein migration) may be observed upon GPI anchor removal, depending on the properties of the protein.
12.5.6 Supplement 2
Current Protocols in Protein Science
DETECTION OF PRODUCTS AFTER PHOSPHOLIPASE TREATMENT BY REACTIVITY WITH ANTI-CRD ANTIBODY
SUPPORT PROTOCOL 2
Cleavage of GPI structures with phospholipase C reveals a cryptic antigen, termed the cross-reacting determinant (CRD). The major epitope contributing to the CRD is an inositol 1,2-(cyclic) monophosphate formed during phospholipase C cleavage; other structural features may also contribute to the antigenicity. Reactivity of a protein with anti-CRD antibody after solubilization by phospholipase C strongly indicates the presence of a GPI anchor. Anti-CRD antibody (Oxford GlycoSystems) may be used after phospholipase C treatment to immunoprecipitate or immunoblot proteins of interest. It should be noted that phospholipase D cleavage does not create the epitope required for anti-CRD activity (see Fig. 12.5.1). NITROUS ACID CLEAVAGE OF GPI-ANCHORED PROTEINS An unusual property common to all GPI structures is the presence of a nonacetylated glucosamine that is glycosidically linked to inositol. The glycosidic bond is specifically cleaved when the glucosamine undergoes nitrous acid deamination, thereby cleaving the GPI anchor (see Fig. 12.5.1). Because nonacetylated glucosamine rarely occurs outside of GPIs, nitrous acid deamination may be used to identify GPI linkage to protein. Due to side reactions, deamination is not always quantitative or precisely reproducible, but it does serve a useful diagnostic purpose.
BASIC PROTOCOL 3
In this protocol, the protein is incubated in nitrous acid to cleave the GPI anchors. Cleaved and uncleaved proteins are then separated by Triton X-114 partitioning. Materials Protein of interest 0.1 M acetate buffer, pH 3.5 (see recipe) 0.5 M NaNO2, made fresh 0.5 M NaCl 1. Dry two aliquots of the protein of interest in 1.5-ml microcentrifuge tubes and dissolve each in 0.1 ml of 0.1 M acetate buffer, pH 3.5. See Critical Parameters for guidelines on the total amount of protein to use. Nonidet P-40 (NP-40) detergent (0.1% v/v final) may be added to aid solubilization.
2. Add 0.1 ml of 0.5 M NaNO2 to one tube. To the second tube (control), add 0.1 ml of 0.5 M NaCl. Incubate 3 hr at room temperature. 3. Detect the release of protein from GPI anchor by Triton X-114 partitioning and analyzing the reaction products (see Alternate Protocol 1). If the anchor is radiolabeled with fatty acids (UNIT 12.2), separation of lipid moiety may be used (see Support Protocol 3).
Glycosylation
12.5.7 Current Protocols in Protein Science
Supplement 2
SUPPORT PROTOCOL 3
SEPARATION OF LIPID MOIETY TO DETECT CLEAVAGE OF GPIANCHORED PROTEINS If GPI-anchored proteins are radiolabeled with fatty acids, the efficacy of GPI anchor cleavage may be conveniently assessed by examining the released radiolabeled lipid products (see Fig. 12.5.1). In this protocol, radiolabeled proteins that have been cleaved by phospholipase or nitrous acid treatment are extracted with butanol. Released lipid moieties are extracted into the butanol phase, then analyzed and quantitated. Materials Radiolabeled protein, cleaved by phospholipase or nitrous acid treatment (see Alternate Protocol 2 and Basic Protocol 3) Water-saturated n-butanol (see recipe) 1. Add 100 µl water to each of two 1.5-ml microcentrifuge tubes. To one tube, add 100 µl cleaved radiolabeled protein. To the second, add 100 µl control (uncleaved) protein. 2. Add 200 µl water-saturated n-butanol to each tube and vortex vigorously. 3. Microcentrifuge 30 sec at maximum speed, room temperature, to separate the two phases. 4. Remove the upper (butanol) phase to a fresh tube, and set aside. Add another 200 µl of water-saturated butanol to the original tubes. 5. Vortex and centrifuge as in steps 2 and 3 above. 6. Remove the second butanol phase and pool it with the first, retaining the lower (aqueous) phase for analysis. 7. Quantitate the percentage of lipid moieties released by the cleavage reactions by liquid scintillation counting of both butanol and aqueous phases. Compare counts recovered in butanol phases of the cleaved material to those recovered from control reactions to assess specific release. To characterize released products fully, examine the butanol-extracted products by thinlayer chromatography (TLC) and compare to standards (Doering et al., 1990b). Products of phospholipase C digestion include diacylglycerol, alkyl-acyl glycerol, or ceramide, depending upon the original structure. Phosphatidylinositol (from nitrous acid treatment) or phosphatidic acid (released by phospholipase D) may also be analyzed and quantitated. This assay has been used to purify phospholipase activity (Hereld et al., 1986).
BASIC PROTOCOL 4
BASE HYDROLYSIS OF RADIOLABELED PROTEINS For proteins radiolabeled with fatty acids, it is useful to ascertain whether incorporated fatty acids are actually part of a GPI structure or are present in amide linkage (N-myristoylation) or thioester linkage (as in palmitoylation of cysteine residues). This can be characterized by assessing the lability of the fatty acid linkage to various chemical treatments. In this protocol, aliquots of radiolabeled proteins are resolved by electrophoresis and the various lanes are treated with KOH in methanol or hydroxylamine. Methanol and Tris⋅Cl buffer are used to treat the control lanes. The lanes are then processed for fluorography. If the chemical bonds in the protein are susceptible to these treatments, a radiolabeled signal will no longer be produced corresponding to the protein band of interest.
Detection of Glycophospholipid Anchors
12.5.8 Supplement 2
Current Protocols in Protein Science
Materials Protein radiolabeled with fatty acid (UNIT 12.2) 0.2 M KOH in methanol Methanol 1 M hydroxylamine⋅HCl, pH 7.5, made fresh (see recipe) 1 M Tris⋅Cl, pH 7.5 (APPENDIX 2E) Additional reagents and equipment for one-dimensional SDS (UNIT 10.1) and staining of gels (UNIT 10.5) 1. Resolve five identical samples of protein radiolabeled with fatty acid by SDS-PAGE (UNIT 10.1). Include appropriate markers. 2. Cut apart the five gel lanes, and stain one to visualize protein and markers (UNIT 10.5). 3. Soak the other four lanes in one of the following solutions for 1 hr at room temperature: Lane 1: 0.2 M KOH in methanol Lane 2: methanol alone Lane 3: 1 M hydroxylamine⋅HCl, pH 7.5 Lane 4: 1 M Tris⋅Cl, pH 7.5. 4. Soak the treated gel slices in water for three washes (10 min each). 5. Process gel pieces for autoradiography. If bonds are susceptible to the treatments above, radiolabel from fatty acids will no longer produce a signal at the position of the protein band of interest. KOH in methanol cleaves thio- and oxyesters but not amide linkages. Hydroxylamine treatment cleaves thioesters only. Fatty acids that are part of a GPI moiety will be released by KOH in methanol, but not by hydroxylamine treatment. Methanol and Tris⋅Cl treatments serve as controls. Ceramide-containing anchors are not sensitive to mild base treatment (see Anticipated Results and Conzelmann et al., 1992).
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Acetate buffer (pH 3.5), 0.1 M Dissolve 0.82 g sodium acetate in 95 ml distilled water. Adjust pH to 3.5 with acetic acid, then bring volume to 100 ml with distilled water. Store indefinitely at room temperature. GPI-PLC buffer 50 mM Tris⋅Cl, pH 8.0 5 mM EDTA 1% NP-40 Store indefinitely at 4°C GPI-PLD buffer 50 mM Tris⋅Cl, pH 7.4 10 mM NaCl 2.5 mM CaCl2 0.1% NP-40 Store indefinitely at 4°C Glycosylation
12.5.9 Current Protocols in Protein Science
Supplement 2
Hydroxylamine⋅HCl (pH 7.5), 1 M Dissolve 6.95 g hydroxylamine⋅HCl in 95 ml distilled water. Adjust pH to 7.5 with NaOH, then increase volume to 100 ml with water. Make fresh for each use. PI-PLC buffer (for B. thuringiensis enzyme) 25 mM Tris acetate, pH 7.4 0.1% (w/v) sodium deoxycholate (Na-DOC) Store indefinitely at 4°C Tris-buffered saline (TBS) 10 mM Tris⋅Cl, pH 7.5 150 mM NaCl Store indefinitely at room temperature Water-saturated n-butanol Mix equal volumes of distilled water and n-butanol. Cap tightly and shake vigorously. Let stand to allow phases to separate; the upper phase is the butanol. Store indefinitely at room temperature. COMMENTARY Background Information
Detection of Glycophospholipid Anchors
Glycosyl phosphatidylinositol (GPI) membrane anchors are one mode whereby proteins are anchored to cell surfaces. These structures consist of a glycan bridge between phosphatidylinositol and phosphoethanolamine; the phosphoethanolamine is in amide linkage to the C-terminus of the protein (see Fig. 12.5.1). The glycan core of GPIs (shown schematically in Fig. 12.5.1) is remarkably conserved throughout evolution, although it is modified in many cell types by the addition of side chains. The lipid moieties of GPIs, however, are quite heterogeneous. This portion may consist of diacyl glycerol, alkyl acyl glycerol, lyso-acyl compounds, or ceramide. The hydrocarbon chains also vary in length and degree of saturation, and may be present as mixtures of species. GPI structures are reviewed in Englund (1993) and McConville and Ferguson (1993). Much of our understanding of GPI anchors is derived from studies of the variant surface glycoprotein (VSG) of trypanosomes. Early work showed that VSG is associated with the membrane via a lipid moiety that contains dimyristoyl phosphatidylinositol (Cardoso de Almeida and Turner, 1983; Ferguson and Cross, 1984; Ferguson et al., 1985, 1986). Other investigations had shown that bacterial phospholipase C releases certain proteins from cell membranes (reviewed by Low, 1989). Together, these studies led to the definition of a class of proteins bearing a C-terminal glycolipid anchor. Further investigations in this field have led to the elucidation of the pathway of GPI bio-
synthesis (reviewed by Doering et al., 1990a, and Field and Menon, 1991), identification of mutations in this pathway and several of the genes involved (Leidich et al., 1994, and reviews cited above), as well as the purification of a GPI-specific phospholipase C (GPI-PLC), and the development of anti-cross-reacting determinant (anti-CRD) antibodies as probes for GPIs. Phosphatidylinositol-specific phospholipase C (PI-PLC) and GPI-specific phospholipase D (GPI-PLD) are also useful in identification of GPI-anchored proteins. A number of compounds found to inhibit GPI biosynthesis are reviewed in McConville and Ferguson (1993). One is mannosamine, which is taken up by cells and incorporated into the growing glycan, preventing completion of GPI precursor biosynthesis and subsequent anchorage (Pan et al., 1992; Ralton et al., 1993). This inhibitor prevents the cell-surface localization of proteins, as shown by Lisanti et al. (1991). PMSF and thiol alkylating reagents inhibit GPI anchor biosynthesis in vitro, as do several compounds affecting the synthesis of the mannose donor Dol-P-mannose (2-fluoro2-deoxy-D-glucose and amphomycin; references in Lisanti et al., 1991). Data obtained from use of these compounds must be interpreted with care, as many also affect other forms of glycosylation, and their effects on GPIs are not always quantitative (see McConville and Ferguson, 1993). Although altered localization of a cell-surface protein of interest upon mannosamine treatment may indicate GPI anchorage, other methods must be used to confirm this, as such changes may be secondary to other
12.5.10 Supplement 2
Current Protocols in Protein Science
adverse effects on the cell. The protocols included here are designed to allow identification of GPI-anchored proteins without detailed structural analysis. Because these techniques are rather diverse, further explanatory information is outlined below. Triton X-114 partitioning. Triton X-114 forms a clear micellar solution at low temperatures; the solution separates into two phases when warmed above 20°C, due to aggregation of detergent micelles (Bordier, 1981). When cellular material is extracted in Triton X-114 at low temperatures, the solution contains soluble proteins, integral membrane proteins, and GPIanchored proteins. Centrifugation results in a pellet containing some of the GPI-anchored species as well as cellular components that are insoluble in nonionic detergents (Hooper and Bashir, 1991). When the detergent solution is warmed, amphiphilic proteins (including those with GPI anchors) associate with the detergentenriched phase, while hydrophilic ones partition to the detergent-depleted phase. Some GPI-anchored proteins exhibit detergent insolubility that develops shortly after synthesis (Brown and Rose, 1992); this is thought to be due to aggregation of the anchored proteins with glycosphingolipids, possibly as a step in directed protein transport. Once a GPI-containing protein is released from the lipid component of its anchor, it will no longer partition into the detergent-enriched phase. This alteration of partitioning behavior provides a rapid assay for the presence or absence of anchors. It may be used to monitor the changes in proteins produced by the other procedures. Phospholipase digestion. Bacterial phosphatidylinositol-specific phospholipase C (PIPLC) efficiently cleaves GPI anchors, releasing proteins in aqueous soluble forms. This enzyme also cleaves phosphatidylinositol (PI), lyso-PI, and PI-linked glycans. Proteins specifically released from membranes by PI-PLC have all been found to be GPI-anchored, making this an extremely reliable mode of analysis (for review see Ikezawa, 1991). Several other specific phospholipases that effectively degrade only GPI species are also useful for analysis. These enzymes are generally more expensive than PI-PLC. One of these, initially purified from Trypanosoma brucei, is a GPI-specific phospholipase C (GPI-PLC). Another, present in mammalian serum (rat, rabbit, or human), is a GPI-specific phospholipase D (GPI-PLD). These enzymes are convenient tools for identifying the presence of a GPI
anchor. Release of a protein from its GPI anchor is followed by use of appropriate protein detection methods—e.g., enzyme activity assays, electrophoretic separation (UNIT 10.1) followed by protein staining (UNIT 10.5), and, if specific antibodies are available, immunoprecipitation or immunoblotting. Alternatively, treated cells may be studied for loss of a specific protein using immunofluorescence detection methods (Watkins, 1989), including fluorescence-activated cell sorting. Also, because release by PLC reveals a specific epitope (termed the cross-reacting determinant, or CRD), antibodies directed against this epitope may be employed to identify cleavage products resulting from PLC treatment (Support Protocol 2). Lack of release may indicate absence of a GPI anchor, although some caution is required in interpreting negative results. Certain GPI molecules, bearing an additional fatty acid on inositol, are not susceptible to cleavage by either bacterial PI-PLC or trypanosomal GPIPLC. Some proteins are partly sensitive, suggesting that only a fraction of these polypeptides bear anchors with acyl-inositol. Serum phospholipase D (GPI-PLD) cleaves GPI structures that contain acylated inositols, and is useful in their identification. However, it is not effective on membrane-bound proteins in the absence of detergent. When intact cells are treated with phospholipase, partial or total resistance to cleavage can also occur due to inaccessibility of the protein to the enzyme, expression of a protein on the cell surface in both GPI-anchored and transmembrane-anchored forms, or tight association of a protein with a nonsusceptible protein on the cell surface (reviewed by Rosenberry, 1991).
Critical Parameters Some details of these procedures will be affected by the method used to detect each protein under study. For example, specific quantities of starting materials are not provided because detection methods vary widely in terms of sensitivity. Also, for activity assays, the conditions of each procedure and whether the activity would survive such treatment must be considered. In general, plan to use two to ten times the amount of protein that would be reasonable for the mode of detection employed, depending upon the availability of material. This should allow for potential losses sustained during the procedure, as well as for the fact that proteins may be recovered in several fractions.
Glycosylation
12.5.11 Current Protocols in Protein Science
Supplement 2
Detection of Glycophospholipid Anchors
Several GPI-anchored proteins, which may be useful control reagents for these protocols, are now available from Oxford GlycoSystems. Triton X-114 partitioning provides a first hint of GPI anchorage, although transmembrane domains are also concentrated in the detergent phase of this partition. For example, if a protein is known from sequence data to have no membrane-spanning domain, its detection in the detergent phase of a Triton X-114 partition would be suggestive of lipid modification. In general, however, this method is most useful as a means of assessing release of a protein from its anchor, as shown by an alteration of partitioning behavior. Many of the other methods described specifically release proteins from their anchors to demonstrate GPI linkage (e.g., nitrous acid deamination or phospholipase digestion); Triton X-114 is a convenient tool for assessing such release. Triton X-114 often contains hydrophilic contaminants, and should be precondensed by several rounds of phase separation before use (Bordier, 1981). Phospholipase digestion is useful for determining GPI linkage, and bacterial PI-PLC is perhaps the best enzyme to try initially for reasons of cost and efficacy. Although GPIPLC is more specific, it is generally more expensive. Some GPI anchors are not susceptible to cleavage by phospholipase C. These structures have an additional fatty acid covalently attached to the inositol of the anchor (Rosenberry, 1991). For this reason, failure of cleavage by PLC does not necessarily imply that a protein is not GPI-anchored. In such cases GPI-PLD may be useful, although this enzyme does not expose the epitope for anti-CRD detection. Extraction and analysis of radiolabeled lipid products from phospholipase digestions provides rapid assessment of cleavage. As mentioned earlier, radiolabeling the protein of interest is often helpful and permits less material and reagents to be used. In particular, if a protein may be radiolabeled with fatty acids, base hydrolysis is a helpful analytical tool, especially if lipid modification is suspected but other methods have yielded negative results.
detergent-enriched phase. These proteins can be chemically cleaved by nitrous acid deamination and enzymatically hydrolyzed by treatment with GPI-PLC, PI-PLC, or GPI-PLD. The protein cleaved with PLC will react with antiCRD antibody, and appropriate lipid products will be generated from all cleavage reactions (Fig. 12.5.1). Radiolabel incorporated into the anchor as fatty acid is released by treatment with KOH in methanol, but not by hydroxylamine treatment, if the fatty acid is hydroxyesterified to glycerol (as in diacyl or alkyl-acyl anchors). Some PI-PLC and nitrous acid-sensitive anchors (e.g., in yeast) have been found to contain ceramide instead of diacyl or alkylacyl glycerol. These anchors are resistant to mild base treatment but sensitive to conditions that hydrolyze amides (Conzelmann et al., 1992). If the GPI anchor includes an acylated inositol, the above characteristics will hold true except that PLC treatment will not cleave the anchor. In these cases, nitrous acid deamination, GPI-PLD digestion, and radiolabel studies are used to characterize the linkage. As mentioned above, removal of a GPI anchor may not alter the electrophoretic mobility of a protein at all, or may even decrease its apparent mobility. Size shifts may be used reliably to assess anchorage only after other methods (such as partitioning, anti-CRD reactivity, and specific radiolabel removal) have been used to ascertain that an observed change does correspond to anchor cleavage.
Anticipated Results
Literature Cited
Proteins anchored by GPI structures without acylated inositols should require detergent for solubilization. They may be extracted by Triton X-114 or remain in the cell pellet after such extraction (depending on the solubility properties of the protein). In a warmed Triton X-114 solution, anchored protein(s) partition into the
Bordier, C. 1981. Phase separation of integral membrane proteins in Triton X-114 solution. J. Biol. Chem. 256:1604-1607.
Time Considerations In general, the methods described above are fairly rapid, with enzyme incubations or chemical reactions and subsequent partitioning requiring 1 to 2 hr. The time required for completion of these analyses depends primarily on the method used to detect the protein(s) of interest. For example, if a protein is radiolabeled in the fatty acid portion of the anchor, cleavage may be assessed by lipid extraction and scintillation counting in ≤1 hr. If immunoprecipitation or immunoblotting is used for product detection, analysis will be more lengthy.
Brown, D.A. and Rose, J.K. 1992. Sorting of GPIanchored proteins to glycolipid-enriched membrane subdomains during transport to the apical surface. Cell 68:533-544.
12.5.12 Supplement 2
Current Protocols in Protein Science
Cardoso de Almeida, M.L. and Turner, M.J. 1983. The membrane form of variant surface glycoproteins of Trypanosoma brucei. Nature 302:349352. Conzelmann, A., Puoti, A., Lester, R.L., and Desponds, C. 1992. Two different types of lipid moieties are present in glycophosphoinositolanchored membrane proteins of Saccharomyces cerevisiae. EMBO J. 11:457-466. Doering, T.L., Masterson, W.J., Hart, G.W., and Englund, P.T. 1990a. Biosynthesis of glycosyl phosphatidylinositol membrane anchors. J. Biol. Chem. 265:611-614. Doering, T.L., Raper, J., Buxbaum, L.U., Hart, G.W., and Englund, P.T. 1990b. Biosynthesis of glycosyl phosphatidylinositol protein anchors. Methods 1:288-296.
cleavage and GPI attachment. J. Cell Biol. 120:657-664. Leidich, S.D., Drapp, D.A., and Orlean, P.A. 1994. A conditionally lethal yeast mutant blocked at the first step in glycosyl phosphatidylinositol anchor synthesis. J. Biol. Chem. 269:1019310196. Lisanti, M.P., Field, M.C., Caras, I.W., Menon, A.K., and Rodriguez-Boulan, E. 1991. Mannosamine, a novel inhibitor of glycosyl-phosphatidylinositol incorporation into proteins. EMBO J. 10:1969-1977. Low, M.G. 1989. The glycosyl-phosphatidylinositol anchor of membrane proteins. Biochim. Biophys. Acta 988:427-454.
Englund, P.T. 1993. The structure and biosynthesis of glycosyl phosphatidylinositol protein anchors. Ann. Rev. Biochem. 62:121-138.
McConville, M.J. and Ferguson, M.A.J. 1993. The structure, biosynthesis, and function of glycosylated phosphatidylinositols in parasitic protozoa and higher eukaryotes. Biochem. J. 294:305324.
Ferguson, M.A.J. 1992. The chemical and enzymatic analysis of GPI fine structure. In Lipid Modifications of Proteins: A Practical Approach (A.J. Turner and N. Hooper, eds.) pp. 191-230. IRL Press, Oxford.
Pan, Y.-T., Kamitani, T., Bhuvaneswaran, C., Hallaq, Y., Warren, C.D., Yeh, E.T.H., and Elbein, A.D. 1992. Inhibition of glycosylphosphatidylinositol anchor formation by mannosamine. J. Biol. Chem. 267:21250-21255.
Ferguson, M.A.J. and Cross, G.A.M. 1984. Myristylation of the membrane form of Trypanosoma brucei variant surface glycoprotein. J. Biol. Chem. 259:3011-3015.
Ralton, J.E., Milne, K.G., Guther, M.L.S., Field, R.A., and Ferguson, M.A.J. 1993. The mechanism of inhibition of glycosylphosphatidylinositol anchor biosynthesis in Trypanosoma brucei by mannosamine. J. Biol. Chem. 268:2418324189.
Ferguson, M.A.J., Duszenko, M., Lamont, G.S., Overath, P., and Cross, G.A.M. 1986. Biosynthesis of Trypanosoma brucei variant surface glycoprotein: N-glycosylation and addition of a phosphatidylinositol membrane anchor. J. Biol. Chem. 261:356-362. Ferguson, M.A.J., Haldar, K., and Cross, G.A.M. 1985. Trypanosoma brucei variant surface glycoprotein has a sn-1,2-dimyristoyl glycerol membrane anchor at its COOH-terminus. J. Biol. Chem. 260:4963-4968. Field, M.C. and Menon, A.K. 1991. Biosynthesis of glycolipid anchors in Trypanosoma brucei. Trends Genet. 3:107-115.
Rosenberry, T.L. 1991. A chemical modification that makes glycoinositol phospholipids resistant to phospholipase C cleavage: Fatty acid acylation of inositol. Cell Biol. Int. Rep. 15:1133-1149. Watkins, S. 1989. Immunohistochemistry. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 14.6.1-14.6.13. John Wiley & Sons, New York.
Key References
Hereld, D., Krakow, J.L., Bangs, J.D., Hart, G.W., and Englund, P.T. 1986. A phospholipase C from Trypanosoma brucei which selectively cleaves the glycolipid on the variant surface glycoprotein. J. Biol. Chem. 261:13813-13819.
Ferguson, M.A.J., Homans, S.W., Dwek, R.A., and Rademacher, T.W. 1988. Glycosyl-phosphatidylinositol moiety that anchors Trypanosoma brucei variant surface glycoprotein to the membrane. Science 239:753-759.
Hooper, N.M. and Bashir, A. 1991. Glycosyl-phosphatidylinositol-anchored membrane proteins can be distinguished from transmembrane polypeptide-anchored proteins by differential solubilization and temperature-induced phase separation in Triton X-114. Biochem. J. 280:745751.
This landmark paper first described the core glycan of GPI anchors, providing a scheme for their structural analysis.
Ikezawa, H. 1991. Bacterial PIPLcs—unique properties and usefulness in studies on GPI anchors. Cell Biol. Int. Rep. 15:1115-1131. Kodukula, K., Gerber, L.D., Amthauer, R., Brink, L., and Udenfriend, S. 1992. Biosynthesis of glycosylphosphatidylinositol (GPI)-anchored membrane proteins in intact cells: Specific amino acid requirements adjacent to the site of
McConville and Ferguson, 1993. See above. Provides an excellent overview of the GPI field, with an extensive bibliography and useful compilations of structural information. Post-translational Modifications of Proteins by Lipids: A Laboratory Manual. 1988. (U. Brodbeck and C. Bordier, eds.) Springer-Verlag, N.Y. A useful laboratory manual of methods dealing with various post-translational modifications of proteins by lipids; many relate to GPI-linked proteins. Glycosylation
12.5.13 Current Protocols in Protein Science
Supplement 2
Methods: A Companion to Methods in Enzymology, Vol. 1, Number 3. (P. Casey, ed.) Academic Press, San Diego, Calif. This issue of Methods, titled “Covalent modification of proteins by lipids,” contains two chapters specifically about GPIs—Doering et al., 1990b (see above) and Mayor and Menon (pp. 297-305)—as well as material on related topics.
Contributed by Tamara L. Doering University of California Berkeley, California Paul T. Englund Johns Hopkins Medical School Baltimore, Maryland Gerald W. Hart University of Alabama Birmingham, Alabama
Detection of Glycophospholipid Anchors
12.5.14 Supplement 2
Current Protocols in Protein Science
Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
UNIT 12.6
Many proteins involved in biological events are glycosylated. A glycoprotein consists of a mixture of glycosylation variants of a single polypeptide chain, known as glycoforms. It has become clear that a detailed understanding of the roles that glycosylation plays in the biosynthesis, transport, biological function, and degradation of a glycoprotein can only be achieved when the protein and sugar(s) are viewed as an entity. Many glycoproteins can now be modeled by combining glycan sequencing data and oligosaccharide structural information with protein structural data. Pivotal to this approach is sensitive, state-of-the-art oligosaccharide sequencing technology, which can give a rapid insight into the glycosylation of a glycoprotein without the need for sophisticated equipment and expertise. To this end the strategy described in this unit has been developed in the Glycobiology Institute, Oxford, where it is in daily use. These protocols enable the glycosylation of 5 to 30 µg of glycoprotein to be analyzed either directly from SDS-PAGE gels (N-linked sugars) or following hydrazinolysis of the glycoprotein (O-linked sugars). The strategy is based on high-performance liquid chromatographic (HPLC) technology, which can be used alone or combined with mass spectrometric techniques (UNIT 12.7). Using HPLC or mass spectrometry (MS), the main characteristics of the oligosaccharides in a glycan pool can be determined within a few days. The preparative nature of HPLC allows individual sugars to be isolated for more detailed analysis if this is required. These advances open up the possibility of ranking high-throughput oligosaccharide sequencing alongside that of proteins and DNA. The increasing efficiency of the methods used to release the glycans and the improved sensitivity of the detection systems suggest that it will soon be possible to characterize oligosaccharides directly from two-dimensional gels used in proteomics. GLYCAN ANALYSIS COMPLEMENTS PROTEIN STRUCTURAL ANALYSIS TO GIVE A MORE COMPLETE VIEW OF A GLYCOPROTEIN Many proteins involved in major biological events such as reproduction, immune defense, the fibrinolytic pathway, and remodeling of the extracellular matrix are glycosylated, and the importance of the attached sugars for the function of these molecules is now well established (Varki, 1993; Rudd and Dwek, 1997a). Whereas the synthesis of the polypeptide chain of a glycoprotein is under genetic control, the oligosaccharides are attached to the protein and processed by a series of enzyme reactions without the rigid direction of nucleic acid templates. N-glycosylation is a cotranslational modification available to, but not necessarily used by, all proteins that contain the sequon AsnXSer/Thr (where X is any amino acid except Pro) in their primary sequence. As the nascent protein enters the endoplasmic reticulum (ER) a block of sugars known as the oligosaccharide precursor (Glc3Man9GlcNAc2) is transferred to nitrogen in the side chain of some Asn residues that are part of the sequon. This process involves a ternary interaction of the glycosylation sequon with an oligosaccharyl transferase and the dolichol phosphate–linked oligosaccharide precursor, both of which are located in the membrane of the ER. Each sugar chain is subsequently processed in the ER and Golgi by a series of glycosidases and glycosyltransferases to either an Glycosylation Contributed by Pauline M. Rudd and Raymond A. Dwek Current Protocols in Protein Science (2000) 12.6.1-12.6.47 Copyright © 2000 by John Wiley & Sons, Inc.
12.6.1 Supplement 22
oligomannose or a hybrid- or complex-type glycan. In mammals oligomannose sugars contain five to nine mannose residues attached to the GlcNAcGlcNAc (chitobiose) core. Hybrid glycans are those in which only the Manα1,3Manβ1,4GlcNAcβ- arm of the core contains GlcNAc (Fig. 12.6.5). The addition of this residue by GlcNAc transferase I is the starting point for the synthesis of both hybrid and complex glycans. In the case of complex glycans, GlcNAc transferase II adds a GlcNAc residue to the Manα1,6Manβ1,4GlcNAcβ- arm of the core, allowing future processing to bi-, tri-, and tetraantennary complex glycans. The process of assigning monosaccharide, linkage type, and position to glycans in oligosaccharide pools, which is the subject of this unit, is made considerably easier if a symbolic, rather than the classical IUPAC nomenclature, is adopted to represent glycan structures. The scheme here uses shapes to represent monosaccharides, shading to represent modifications to the monosaccharide, bond angle to represent linkage position, and line type to represent anomericity. This symbolic representation is designed for ease of use, and does not represent the true conformation of the glycans. The nomenclature is explained in Figures 12.6.1 and 12.6.2, and the structures of N-glycan core structures, hybrid, complex, and oligomannose-type N-linked oligosaccharides, and the O-glycan core structures are shown in Figures 12.6.1 to 12.6.8. Glycans are frequently as large as the protein domains to which they are attached. Moreover, in addition to stabilizing the protein (Joao et al., 1992), both the sugars themselves and the N-glycosidic linkage to the Asn side chain are flexible. This feature enables the glycans to shield large areas of the protein surface (Woods, 1995). As mentioned above, it is clear that a detailed understanding of the roles of glycosylation can only be achieved when the protein and sugar(s) are viewed as an entity. By combining glycan sequencing data and oligosaccharide structural information (Petrescu et al., 1999) with protein structural data, many glycoproteins can now be modeled in their entirety. This means that, in many cases, it is possible to visualize the sugars in the context of the glycoproteins to which they are attached. An example of this is shown in Figure 12.6.9, in which the N-linked, O-linked, and GPI anchors of CD59 and CD55 have been modeled by combining protein and glycan structural data with the oligosaccharide database. Rapid, sensitive, state-of-the-art oligosaccharide sequencing technology is needed to deal with the complex heterogeneity of the glycoform populations of many natural glycoproteins. Figure 12.6.10 highlights the strategies that have been developed at the Glycobiology Institute that enable the overall N-glycosylation of a few micrograms of glycoprotein to be analyzed in a few days using HPLC (see Basic Protocol 7 and Guile et al., 1994, 1996) and/or MS (see Chapter 16 and UNIT 12.7; Rudd et al., 1997a). N-linked glycans can be released directly from SDS-PAGE gels (UNIT 10.2; Küster et al., 1997), or the N- and O-glycans can be released from purified glycoproteins by hydrazine (see Basic Protocols 4 and 5). These advances open up the possibility of ranking high-throughput sequencing of oligosaccharides alongside that of proteins and DNA. This unit deals with the practical aspects of a state-of-the-art strategy for analyzing the structure of glycosyl side-chains on glycoproteins (for an overview of glycosylation analysis, see UNIT 12.1). This involves four stages: (1) glycan release, (2) labeling the glycan pool, (3) separating (profiling) the glycans in the pool, and (4) characterization. Each of these steps is discussed in the sections that follow. Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
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GLc
xylose
P
unknown pentose
GlcNAc
ribose
H
unknown hexose
Gal
arabinose
GalNH2
glucuronic acid
GalNAc
iduronic acid
Man
NeuNAc HO
Fuc
NeuNGc
unknown HexNAc
s
unknown sialic acid P NS OS NMe
KDO
6
8
4 3
OMe
phosphate N-sulfate O-sulfate N-methyl O-methyl
β linkage α linkage unknown linkage
2 linkage position
Figure 12.6.1 Various principles of diagrammatic nomenclature. (1) Each monosaccharide is represented by a particular symbol (e.g., square for Glc) whose shape does not relate to structure. (2) The shading of the shape represents substituents on the monosaccharide (e.g., open, not substituted; black, N-acetyl group; dot, deoxy sugar). Hence, Gal is represented by an open diamond and deoxy-Gal (Fuc) by an open diamond with a dot inside. When the exact monosaccaride is unknown (e.g., from MS data), an appropriately shaded symbol is used (e.g., open hexagon, hexose; black hexagon, HexNAc). (3) Linkage positions are represented by the angle of the line linking adjacent monosaccharides. As most sugars are linked via the C1 reducing end (except sialic acids, which are linked via C2), this information is not represented in these diagrams. It is the position to which the reducing end of the sugar to the left attaches to the sugar on the right that is represented by bond angle. (4) The type of line represents anomericity. A straight line indicates a β linkage and a dotted line represents an α linkage. Unknown linkages are represented by a wavy line. (5) Substituents can be represented as P, S, and so on. This simple visual representation of both sugars and linkages clearly conveys the structural information needed when working with complex glycan structures. Abbreviations: Fuc, fucose; Gal, galactose; Glc, glucose; Man, mannose; NAc, N-acetyl; Neu, neuraminic acid; NGc, N-glycolyl.
RELEASE OF GLYCANS FROM GLYCOPROTEINS Intact N-linked glycans can be released from glycoproteins or glycopeptides enzymatically with peptide N-glycosidase F (PNGase F; Roche Diagnostics), as described in Basic Protocol 1. This enzyme is an amidase that cleaves the linkage between the core GlcNAc and the A residue of all classes of N-glycans (Fig. 12.6.3; Kuhn et al., 1994), with the exception of some plant and insect N-glycans that contain fucose α1,3-linked to the core GlcNAc residue attached to the protein. The glycoprotein should be fully denatured, since unfolding promotes enzyme accessibility to the cleavage site, which may, in some cases (as with RNase B), be protected by the native protein (Fig. 12.6.11). In some cases it may also be necessary to cleave the peptide chain with trypsin before PNGase F treatment to ensure that all sugars are released. Glycosylation
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2
2 6 1 41
1
1
6 1 4 14 6 3
3
reducing terminus
2 1 1 41
2
structure illustrated using the scheme in Figure 12.6.1 annotated to demonstrate linkage position
structure as it would conventionally appear
structure as it would appear in figures
Figure 12.6.2 Examples showing how glycan structures are constructed.
Asn chitobiose (NN)
Asn
Asn
4′-β-mannosyl chitobiose (MNN) conserved trimannosyl core (M3N2)
Figure 12.6.3 Native N-linked oligosaccharide core structures. For nomenclature, see Table 12.6.8.
Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
PNGase F can be used to release sugars from glycoproteins that are either in solution (see Basic Protocol 1) or in an SDS-PAGE gel band (see Basic Protocols 2 and 3). One advantage of working from SDS-PAGE gels is that the electrophoresis step is a useful means of purifying many glycoproteins to homogeneity without prohibitive loss of material. This means that it is possible to analyze the glycosylation of low levels of biologically relevant glycoproteins where either limited quantities of material or protein purification problems would previously have precluded a detailed glycan analysis. In practice, releasing the sugars directly from a glycoprotein in a gel band results in a higher recovery of glycans than that obtained when the glycoprotein is incubated with PNGase F in solution. A third application for the in-gel method is to release the sugars from peptide fragments resulting from protease digests when these can be resolved by SDS-PAGE. In some cases, this may enable a rapid analysis of glycosylation sites in glycoproteins. Reducing gels also allow the straightforward separation of proteins into their component subunits. For example, IgG can be resolved into heavy and light chains (Küster et al., 1997), and the surface coat proteins of the hepatitis B virus or particle can be separated into three major glycoprotein components (Mehta et al., 1997). Also, when gelatinase B is purified on gelatin-Sepharose, it is contaminated with its natural inhibitor (tissue inhibitor of metalloproteinase I or TIMP-1). This enzyme-inhibitor complex is stable, but
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Asn
oligomannose 6 (Man-6)
Asn
oligomannose 7 D1 (Man-7 D1)
oligomannose 7 D2 (Man-7 D2)
Asn
Asn
oligomannose 8 D1,D2 (Man-8 D1,D2)
oligomannose 7 D3 (Man-7 D3)
Asn
oligomannose 8 D2,D3 (Man-8 D2,D3)
Asn
Asn
oligomannose 8 D1,D3 (Man-8 D1,D3)
Asn
oligomannose 9 (Man-9)
Figure 12.6.4 Oligomannose-type N-linked oligosaccharides. D1, D2, and D3 refer to the terminal mannose residues attached Manα1,2 to the Manα1,6Manα1,6 (D1), the Manα1,3Manα1,6 (D2), and the Manα1,2Manα1,3 (D3) arms of the glycan core.
SDS-PAGE run under reducing conditions can be used to resolve the two proteins. N-linked glycans can then be released directly from the gels with PNGase F (Rudd et al., 1999a). A final advantage of the in-gel release method is that the protein remains in the gel after the sugars have been removed. In-gel proteolysis of the protein (for example, by trypsin) and analysis of the peptide fragments by nanospray MS enables the protein to be identified. This is achieved by using a protein data base to compare the molecular weights of the tryptic fragments with those of the predicted sequences of fragments from tryptic digests of known proteins (Küster et al., 1997). While PNGase F is the enzyme of choice to release N-linked sugars for glycan analysis, in certain circumstances other enzymes may be required (see UNIT 12.4 for more details). For example, PNGase A cleaves sugars containing α1,3-linked core fucose, which are not susceptible to PNGase F. Two enzymes that cleave between the two N-acetylglucosamine (GlcNAc) residues within the chitobiose core are endoglycosidase D (endo D), which
Glycosylation
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Asn
Asn
A2G0
A2G0B
Asn
A2G2B
Asn
A2G2F(α1,6)
Asn
Asn
A3G0
A2G2F(α1,6)B
Asn
A3G3
Asn
Asn
A3G3
A4G0
Asn
A4G4
Asn
A2G1
A2G2
A2G0F(α1,6)
A2G0FB
Asn
Asn
Asn
Asn
hybrid with bisecting GIcNAcA2G2
Figure 12.6.5 Hybrid and complex native N-linked oligosaccharides. For nomenclature, see Table 12.6.8.
releases all classes of N-linked sugars, and endo H, which selectively cleaves oligomannose- and hybrid-type structures. Another useful group of enzymes is the endo-β-N-acetylglucosaminidases F1, F2, and F3. All three enzymes cleave within the chitobiose core, but while endo F1 digests only oligomannose and hybrid structures, endo F2 cleaves oligomannose and biantennary glycans, and endo F3 is specific for bi- and triantennary sugars. For deglycosylation, NaHCO3 not NH4HCO3 should be used as a buffer since the latter leaves an amine group at the reducing terminus of the glycans, which will result in severe losses during 2-AB labeling (Küster et al., 1997).
Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
Both N- and O-linked sugars can be released chemically with hydrazine (Fig. 12.6.12), either manually (see Basic Protocol 4) or with the automated GlycoPrep system (Oxford GlycoSciences; see Basic Protocol 5). O-linked glycans may only be released chemically, given that a generic O-glycanase is not as yet available.
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LABELING THE RELEASED GLYCANS The classical method of attaching radioactive tritium to the released oligosaccharides has largely been replaced by labeling with fluorescent compounds. As a result, it is now possible to detect sugars in the femtomole compared with the picomole range. A suitable label must have a high molar labeling efficiency and be capable of labeling the oligosaccharide components of a glycan pool nonselectively so that they are detected in their correct molar proportions. As described in Basic Protocol 6, 2-aminobenzamide (2-AB)
Asn
Asn
A2G2S
A2G2S(α2,3)
Asn
Asn
A2G2S(α2,6)
A2G2FS(α2,6)
Asn
Asn
Asn
Asn
A3G3FS(α2,6)2
A3G3FS3(α2,6)2(α2,3)1
(thus, six oligomers of this structure may exist)
(thus, six oligomers of this structure may exist)
Figure 12.6.6 Sialylated complex native N-linked oligosaccharides. For nomenclature, see Table 12.6.8.
....
core I
.... Ser/Thr core V
.... Ser/Thr core II
.... Ser/Thr core III
.... Ser/Thr
.... Ser/Thr
core VI
core VII
Figure 12.6.7 O-glycan core structures.
.... Ser/Thr core IV
....
.... Ser/Thr
.... Ser/Thr core VIII
Glycosylation
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core 1
core 2
core 3
core 4
core 5
core 6
core 7
core 8
Figure 12.6.8 Molecular models of O-glycan core structures.
fulfills these conditions and, in addition, it is compatible with a range of separation techniques including normal-phase (NP), weak anion-exchange (WAX), and reversedphase (RP) HPLC, and matrix-assisted laser desorption/ionization MS (MALDI-MS) and electrospray ionization MS (ESI-MS). It was for these reasons that 2-AB was selected as the label for the HPLC-based glycan analysis strategy described below. PROFILING BY HPLC Three different HPLC systems (see Basic Protocol 7) are used to assign structures to the released sugars: NP-HPLC, RP-HPLC, and WAX-HPLC. If material is limited, NP-HPLC should be used, as it gives the maximum information from the minimum sample. Although it is possible to recover the sample from a MALDI-MS target, in practice, roughly twenty NP- or RP-HPLC or five WAX-HPLC runs can be carried out with the amount of sample needed for one MALDI-MS analysis. Ten to fifty femtomoles of a single glycan give a reasonable signal-to-noise ratio on NP- or RP-HPLC. The amount of sample needed from a pool depends on the number of different glycans in the pool and on its concentration. The best way to work out how much to load is to run a small amount of the sample (e.g., 5% for WAX and 2% for NP- or RP-HPLC) and adjust the volume for subsequent runs depending on the ratio of signal to noise in the trial run.
Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
A major advantage of all HPLC systems is that they can be used preparatively. This means, for example, that pools of sugars separated according to charge using WAXHPLC can be collected and individually analyzed by NP-HPLC. This procedure may be used to limit the number of structures in the normal-phase profiles, thus simplifying the interpretation. Preparative HPLC also allows individual peaks to be collected and analyzed by mass spectrometry (UNIT 12.7) or other techniques to validate assignments (Rudd et al., 1997a).
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WAX-HPLC WAX chromatography using a GlycoSep C column (Glyko) is used to separate glycan mixtures according to the number of charged residues they contain (Guile et al., 1994). Within one charge band several peaks may be resolved, since larger structures elute before smaller ones. Charge due to sialic acid can be assigned by reference to a standard run of fetuin oligosaccharides, which contain mono-, di-, tri,- and tetrasialylated glycans (Fig.
N-glycan
O-glycans
A
B O-glycan
N-glycan
GPI anchor
CD55
CD59
Figure 12.6.9 (A) CD55 (a.k.a., decay accelerating factor or DAF) and (B) CD59 both contain N- and O-linked glycans and are attached to the membrane by a glycosylphosphatidylinositol (GPI) anchor. CD59 is a glycoprotein consisting of one domain (Ly6) and it is located close to the membrane surface of many cells, including erythrocytes. Both the classical and alternative complement pathways terminate with the formation of the membrane attack complex (MAC) in the cell surfaces of bacteria and other pathogens, and this leads to cell lysis. Host cells are normally protected from destruction through inhibitors of the complement pathway, such as CD55, which destabilizes the C3 convertase components C3b/Bb and C4b2a, and CD59, which binds C8 and/or C9 preventing the formation of the fully assembled MAC complex. CD59 contains all three types of post-translational modifications that involve glycosylation. It has one N-linked sugar at Asn-18 and one or two O-linked glycans (Rudd et al., 1997b). CD55 is also linked to the erythrocyte cell surface by means of a GPI anchor. The protein consists of four complement control protein domains. Three of these domains, together with an O-linked region that acts as a rigid spacer, act to position the N-linked glycan some 20 nm above the membrane surface (Kuttner-Kondo et al., 1996).
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purified glycoprotein manual hydrazinolysis dialysis: 16-48 hr lyophilization: 12-48 hr cryogenic drying: 48-72 hr
O-links: 6 hours at 60°C N-links: 12 hours at 85°C
automated hydrazinolysis dialysis: 16-48 hr lyophilization: 48 hr GlycoPrep procedure: 14 hr for one sample, 22 hr for two samples
post hydrazinolysis: re-N-acetylation, desalting and chromatography: 3 days
SDS-PAGE 4 hr excision of bands of interest and preparation 4 hr in-gel PNGase F digest to release N-linked glycans 20 hr
protein in-gel trypsin digest
glycan pool extract peptides fluorescent labeling 4 hr HPLC 3 hr per run
MALDI-MS
glycan profiling, exoglycosidase sequencing
MALDI-MS
protein sequence confirmation from database searching
Figure 12.6.10 Flowchart for analysis of N-glycans.
12.6.13A). Neutral sugars (and noncarbohydrate material) elute in the void. Incubation of the pool with Arthrobacter ureafaciens sialidase will digest sialylated glycans to neutral (Fig. 12.6.13A, lower profile). Sugars containing charge due to groups other than sialic acid, such as the sulfated structures attached to the glycoforms of sialoadhesin that bind the cysteine-rich domain of the mannose receptor (Martinez-Pomares et al., 1999), will remain charged after the sialidase digestion, and can, therefore, be isolated for further analysis (Fig. 12.6.13B, peak S1). In Figure 12.6.13C, which shows the analysis of glycoforms of sialoadhesin that do not bind the cysteine-rich domain of the mannose receptor (upper profile), peak S1 does not contain sulfated glycans and is digested to neutral with A. ureafaciens sialidase (lower profile). NP-HPLC
Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
This is performed using an amide column in which the polar functional groups interact with the hydroxyl groups on the sugars (Guile et al., 1996). Samples applied in high concentrations of organic solvent adsorb to the column surface and are eluted with an aqueous gradient so that glycans are resolved on the basis of hydrophilicity. Typically, larger oligosaccharides are more hydrophilic than smaller ones, and require higher concentrations of aqueous solvent to elute. The system described here is sensitive and highly reproducible, and has been developed using the GlycoSep N column (Glyko). It is capable of resolving subpicomolar quantities of mixtures of fluorescently labeled neutral and acidic N- and O-glycans simultaneously and in their correct molar proportions. A typical profile is shown in Figure 12.6.14. Elution positions are expressed as glucose units (GU) by comparison with the elution times of a dextran ladder. The contribution of individual monosaccharides to the overall GU value of a given glycan can be calculated,
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N-glycan
active site RNase B
N-glycan
PNGase F
soluble CD59
Figure 12.6.11 Structures of CD59, RNase B, and PNGase F. The structure of PNGase F is based on the X-ray crystal structure (Norris et al., 1994). The protein is folded into two domains, each with an eight-stranded antiparallel β–jelly roll configuration similar to the types of structures found in lectins. CD59 was modeled according to the NMR solution structure (Kieffer et al., 1994). RNase B was modeled according to the X-ray crystal structure (Williams et al., 1987). All three molecules are modeled to the same scale. The active site for PNGase F is in a cleft flanked by five Trp residues and one Phe residue (Kuhn et al., 1994). In contrast to the CD59 Asp-sugar amide linkage, which is susceptible to the enzyme, the same linkage in RNase B is protected by the protein and is not accessible. The arrow in CD59 indicates the attachment site for the GPI anchor in the membranebound form.
and these incremental values can be used to predict the structure of an unknown sugar from its GU value (Tables 12.6.1 and 12.6.2). The system is able to resolve arm-specific substitutions of a multiantennary sugar, and the linkage positions of individual monosaccharides, such as fucose, also affect retention times, giving a further level of specificity. RP-HPLC Reversed-phase HPLC, using a GlycoSep R C18 column (Glyko), separates sugars on the basis of hydrophobicity. Samples are applied in an aqueous solvent and eluted with increasing concentrations of organic solvent. The column is calibrated with an arabinose ladder rather than the dextran ladder used for the NP-HPLC, and retention measurements are made in arabinose units (AU; Guile et al., 1998). Sugars that co-elute by NP-HPLC can often be separated using RP-HPLC (Fig. 12.6.15). The effect of linkage position is much more marked on reversed-phase separations than on normal phase. For example, the two trisaccharides Lewis-X and Lewis-A, which have the same monosaccharide
Glycosylation
12.6.11 Current Protocols in Protein Science
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typical O-linked glycan R2 O peptide O N O O O O N H peptide H R1 base (B)
(R2) Ser/Thr
(R1)
base catalyzed β-elimination
R2 R2
R2 OH O
O
N
O degradative ‘peeling’
R1
reaction with excess hydrazine
labeling
R2 OH O
O O
O
O
OH H
B O
O OH
N
R1
O
O R1 ‘peeled’ glycan
re-N-acetylation and O acetohydrazone cleavage
NH2
O
R2
OH O N
O
O
R1
R1
reaction with 2-aminobenzamide
stable 2-AB-labeled O-glycan reduction with sodium R2 OH H2N O cyanoborohydride N O O O N
O R1
OH O N
R2
O
OH H2N O N N O
R1
unstable Schiff’s base
Figure 12.6.12 The chemistry of the hydrazine reaction to release intact O-glycans from proteins, the 2-AB fluorescent labeling step, and the “peeling” reaction. The exact mechanism of release of intact O-glycans has not been determined but is believed to take place via an initial base-catalyzed β-elimination reaction cleaving the bond between GalNAc and Ser/Thr, followed by immediate reaction of the released aldehyde with excess hydrazine to form a hydrazone derivative. In the presence of excess water, further β-elimination may take place when the GalNAc is substituted at the 3 position (as shown by the Gal R1 group in this figure), which leads to loss of the GalNAc and may proceed even further to total destruction of the glycan. The hydrazone is then re-N-acetylated, the acetohydrazone at C1 is cleaved, and a glycan with a free reducing terminus is produced. This may then be reacted with 2-aminobenzamide (2-AB) to give an unstable Schiff’s base intermediate, which is stabilized by addition of sodium to the reaction mixture to form a reduced, fluorescently labeled product of the glycan.
Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
composition—Galβ1,4Fucα1,3GlcNAc and Galβ1,3Fucα1,4GlcNAc, respectively— have similar elution times on normal phase (Lex GU value: 2.59; Lea GU value: 2.85), but are well separated by reversed phase (Lex AU value: 0.00; Lea AU value: 2.39). RP-HPLC is particularly useful for identifying oligosaccharides containing bisecting GlcNAc. Structures that differ only by the presence or absence of bisecting GlcNAc are well separated on reversed-phase columns, but elute at similar positions on normal-phase columns.
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A fetuin pool
Fluorescence
S3 S2 S4 S1
S5
ABS
B Sn-b
b1 b2 Fluorescence
b3 b4 b5
ABS
C
Fluorescence
nb1
Sn-nb
nb2
nb3 nb4
nb5
ABS
0
10
20 Time (min)
Figure 12.6.13 WAX separation. (A) A standard mixture of neutral, mono-, di-, tri-, and tetrasialylated fetuin sugars used for calibrating the column (upper profile), and the profile of the same sugars following desialylation with Arthrobacter ureafaciens sialidase (ABS) showing that all structures have become neutral. (B) Glycans from a set of glycoforms of sialoadhesin that binds the cysteine-rich domain of the mannose receptor (Sn-b; upper profile; Martinez-Pomares et al., 1999).The lower profile shows the same sample following desialylation. Some of the glycans remain acidic, consistent with the presence of sulfate on these oligosaccharides. (C) Glycans from sialoadhesin that do not bind the cysteine-rich domain of the mannose receptor (Sn-nb; upper profile). The lower profile shows that, after digestion with sialidase, all these sugars become neutral and, therefore, do not contain sulfate substitutions.
Glycosylation
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Fluorescence
6
7
Sn-b
MW kd 205
8
9
10
11
12
13 gu Sn-b glycan pool
116 97 84 66 55 45
80
90
100
110
120
130
Time (min)
Figure 12.6.14 NP-HPLC separation of sialoadhesin. The NP-HPLC profile of sialoadhesin glycoforms that bind the cysteine-rich domain of the mannose receptor (Sn-b). Insert: Gel from which the Coomassie blue–stained band of sialoadhesin was excised.
CHARACTERIZATION OF N- AND O-LINKED OLIGOSACCHARIDES Preliminary Assignment of Peaks From NP- and RP-HPLC Profiles Tables 12.6.1 and 12.6.2 and Figure 12.6.16 show the GU and AU values for common Nand O-linked glycans. They are used to list the possible structures that correspond to the GU (NP-HPLC analysis) and AU (RP-HPLC analysis) values of the sample peaks. These values are highly reproducible from column to column and system to system (Guile et al., 1996), giving typical standard deviations of <0.05 GU or AU units. Oligosaccharide Structural Analysis Using Exoglycosidase Sequencing Enzymatic analysis of oligosaccharides using highly specific exoglycosidases is a powerful means of determining the sequence and structure of glycan chains (see Basic Protocols 8 and 9). The preliminary assignments of structures made from GU and AU values are confirmed by digesting the entire glycan pool with arrays of exoglycosidases, and the digestion products are analyzed by NP-HPLC using the same conditions as for the original glycan pool. However, until recently, this has only been possible after isolation of single sugars from the total glycan pool and digesting each—with exoglycosidase enzymes, either sequentially or using enzyme arrays. It is often impractical and always time consuming to purify each individual sugar to 80% purity. Therefore, the simultaneous analysis of the total glycan pool as presented in Basic Protocol 8 (for N-glycans) and Basic Protocol 9 (for O-glycans) is a major development. Detection of Structures by Mass Spectrometry Mass spectrometry is used mainly to determine the molecular weights and hence the composition of the glycans in the pool, but does not give linkage information directly. UNIT 12.7 presents methods to analyze glycan structure by this method. Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
CAUTION: Acrylamide is a neurotoxin, although it is not as dangerous once it has been polymerized. N,N,N′,N′-Tetramethylethylenediamine (TEMED) is also toxic. Wear gloves at all times and use these compounds in a fume hood. Refer to materials safety data sheets (MSDSs) for further information.
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ENZYMATIC RELEASE OF N-LINKED OLIGOSACCHARIDES BY PNGase F DIGESTION IN SOLUTION
BASIC PROTOCOL 1
This enzymatic procedure (Goodarz and Turner, 1998) may be used to release N-linked glycans from glycoproteins when the glycoprotein sample is pure and when use of hydrazinolysis (Basic Protocol 4) is not feasible. As there may be variable steric hindrance depending on the nature of the peptide and/or the glycans, the glycoprotein is denatured by reduction and alkylation in the presence of SDS. The SDS must then be exchanged for Table 12.6.1
N-Linked Oligosaccharides
Structurea
Massb
GUc
AUc
Core structures: M1N2
586.32
2.65
2.72
M1N2F M3N2
732.38 910.44
3.17 4.41
4.95 2.56
M3N2F Oligomannose structures:
1056.5
4.90
4.65
Man-9 Man-8
1882.80 1720.74
9.56 8.83
0.83 0.83
Man-7 Man-6
1558.68 1396.62
7.96 7.10
1.3 1.52
Man-5 Bi-, tri-, and tetraantennary complex structures:
1234.56
6.21
2.17
A2G0 A2G0B
1316.59 1519.67
5.50 5.63
3.14 3.10
A2G0F(α1,6) A2G2
1462.64 1640.50
5.93 7.15
5.18 3.22
A2G2B A2G2F(α1,6)
1843.60 1786.60
7.30 7.57
4.96 5.18
A2G2F(α1,6)B A3G0
1989.7 1519.6
7.67 5.91
7.59 3.65
A3G3 A4G0
2005.6 1722.70
8.32 6.53
3.81 2.70
A4G4 Sialylated structures:
2371.00
9.67
2.70
A2G2S(α2,3) A2G2S(α2,6)
1931.69 1931.69
7.49 7.89
3.84 3.48
A2G2FS(α2,3) A3G3FS2(α2,6)
2077.75 3845.37
8.14 9.61
5.86 4.46
A3G3FS3(α2,6)2(α2,3)1
4136.46
9.54
9.78
aN-linked structures are selected as examples only; this is not an exhaustive list.
All of these structures are illustrated schematically in Figures 12.6.3 to 12.6.6. For nomenclature, see Table 12.6.9. bMass given is for closed-ring unlabeled structures; add 120.069 for addition of
2-AB label and 22.9 for sodium adduct in MALDI-MS. cGlucose unit (GU) and arabinose unit (AU) values are given for 2-AB-labeled
structures.
Glycosylation
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Table 12.6.2 Incremental Glucose Units for Addition of Monosaccharides to N-Linked Oligosaccharides
Monosaccharide
Linkage
To
Increment
Core fucose
α1,6
Chitobiose and M3N2
0.5
Core fucose Core fucose
α1,6 α1,3
A2G0, A2G2, A2G2B, A3G3 Any structure
0.42 0.8
Antennary fucose Mannose
α1,3; α1,6 α1,2; α1,3; α1,6
Any structure M3N2 up to Man-7
0.40-0.80a 0.9
Mannose Bisecting GlcNAc
α1,2; α1,3; α1,6 β1,4
Man-8 and above First mannose
0.72 0.10-0.15
Galactose NeuNAc
β1,4 α2,3
Any structure Galactose of any structure
0.90-1.00 0.7
NeuNAc
α2,6
Galactose of any structure
1.15
aDepending on location.
another detergent, since SDS inactivates PNGase F. Finally, since it is possible that glycan removal may be incomplete, it is advisable to check for the completion of release by SDS-PAGE. Incomplete removal is indicated by the presence of more than one band. Materials Glycoprotein sample Buffer A (see recipe) Buffer B (see recipe) 10% (w/v) dithiothreitol (DTT) 1000 U/ml peptide N-glycosidase F (PNGase F), recombinant glycerol-free lyophilisate (Roche Life Sciences) Toluene Sub-boiling-point-distilled (SBPD) water Vacuum centrifuge 100°C temperature-controlled heating block Protein-binding membrane (e.g., 1.9-ml Microspin filter, 0.45-µm CN, PP NON ST; Radley and Co.) PCR tubes Additional reagents and equipment for SDS-PAGE (UNIT 10.1) 1. Dry a glycoprotein sample in a 1.5-ml microcentrifuge tube in a vacuum centrifuge. Generally 50 to 200 µg of glycoprotein is required. The glycoprotein should be isolated according to usual procedures. If the sample is in a low-molarity or volatile salt solution, no dialysis is required. The incubation of a control glycoprotein with known glycosylation alongside experimental samples is recommended. Suitable control glycoproteins include ribonuclease B, bovine serum fetuin, and haptoglobin. All are available from Sigma, although there may be some variation in glycosylation profiles between different batches.
2. Dissolve sample in 10 µl buffer A and incubate 3 min at 100°C. Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
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3. Cool to room temperature and add the following in the order indicated: 8 µl buffer B 2 µl 10% DTT 20 µl 1000 U/ml PNGase F (20 U) 5 µl toluene. Toluene is added to prevent bacterial growth.
RP-HPLC AU values AU scale
NP-HPLC GU values GU scale
1
2
3
4
2AB 3.58
5
6
7
2
3
undig
undig 2AB 1.63
A
2AB 2.73
B
C
1
SPG + SPH
2AB 1.79
D 2AB
SPG
2.73
BTG
2AB 1.63
SPG
SPG + SPH
2AB 1.35
1.63
BTG 2AB
0.9 1.79
E
2.15
2AB 0.9
2.80
2AB BTG + SPH
10 20 30 40 50 60 70 80 90 Time (min)
2.14
BTG + SPH
20 25 30 35 40 45 50 55 60 65 70 Time (min)
Figure 12.6.15 Two-dimensional separation of O-glycans using NP- and RP-HPLC. GU and AU values for a set of digests carried out on an O-linked tetrasaccharide and monitored by NP- and RP-HPLC. On the NP column, the peak that results from digesting the glycan with SPG (B; GU 2.73) elutes at a smaller GU than the starting glycan (A; GU 3.58); while on the RP column, the starting glycan and the product of the digest elute in the same position. However, comparing the BTG digest to the starting glycan (D and A), the GU of the product (1.79) is again less than the GU of the starting material (3.58) on the NP column, whereas the AU of the product (2.80) is greater than the AU of the starting glycan (1.63) on the RP column. Thus, the two separation techniques are complementary, allowing two structures that co-elute on one column to be resolved on the other. Abbreviations: BTG, bovine testes β-galactosidase; SPG, Streptococcus pneumoniae β-galactosidase; SPH, Streptococcus pneumoniae β-hexosaminidase; undig, undigested.
Glycosylation
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Structure
Mass
GU
AU
monosaccharides
341
0.91
2.14
300
1.00
0.39
284
0.53
2.07
341
1.07
1.62
300
1.00
0.54
503
1.80
1.35
794
2.97
1.97
794
3.29
2.49
1085
4.40
2.62
544
1.79
2.80
706
2.73
1.63
868
3.58
1.63
1014
4.31
1.38
1159
4.41
3.23
1305
5.09
3.04
1450
5.40
3.42
591
2.26
2.11
446
1.58
5.18
503
1.82
0.92
core 1
core II
galactose
Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
Figure 12.6.16 Structure and mass of O-linked oligosaccharides. The structures are selected as examples only and do not represent an exhaustive list. Mass given is for closed-ring unlabeled structures; add 120.069 for addition of 2-AB label, and 22.9 for sodium adduct in MALDI-MS. GU and AU values are given for 2-AB-labeled structures.
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4. Incubate 24 hr at 37°C. 5. Remove 5 µl and analyze by SDS-PAGE (UNIT 10.1). When the sample is completely deglycosylated, proceed with step 6. Otherwise continue incubation. If the reaction mixture is analyzed alongside a control sample that has not been incubated with PNGase F, a shift to a lower molecular weight, corresponding to the deglycosylated protein, will be observed. A series of bands may be observed that can indicate deglycosylation at different sites of the glycoprotein.
6. Filter samples through a protein-binding membrane into PCR tubes by centrifuging 5 min at 1200 × g. 7. Wash glycan sample off the filter with two washes of 10 µl SBPD water, centrifuging 5 min at 1200 × g. 8. Dry sample in a vacuum centrifuge. The sample is ready for 2-AB labeling or may be stored frozen (up to 1 year at −20°C) after redissolving in 20 µl SBPD water.
IN-GEL ENZYMATIC RELEASE OF N-LINKED GLYCANS BY PNGase F FROM PROTEINS SEPARATED IN SDS-PAGE GEL BANDS
BASIC PROTOCOL 2
The following in-gel release method has been developed to release N-glycans from Coomassie blue–stained protein bands on SDS-PAGE gels by PNGase F (Küster et al., 1997). The released glycan pool can be analyzed by matrix-assisted laser desorption/ionization time-of-flight MS (MALDI-TOF-MS) and, after fluorescent labeling, by HPLC. The individual sugars in the pool can be analyzed simultaneously using enzyme arrays (Guile et al., 1996) monitored by either MALDI-TOF-MS or HPLC. Materials 30% acrylamide/0.8% bis-acrylamide solution (e.g., Acrylagel; National Diagnostics) 0.5 M Tris base, pH 8.8 1.5 M Tris base, pH 6.6 Sub-boiling-point-distilled (SBPD) water 10% (w/v) SDS 10% (w/v) ammonium persulfate (APS) N,N,N′,N′-Tetramethylethylenediamine (TEMED) Glycoprotein sample 0.5 M dithiothreitol (DTT) 5× SDS sample buffer (see recipe) 100 mM iodoacetamide 25 mM Tris/190 mM glycine/0.5% (w/v) SDS 50% (v/v) methanol/7% (v/v) acetic acid 5% (v/v) methanol/7% (v/v) acetic acid 20 mM NaHCO3, pH 7.0 1:1 (v/v) acetonitrile/20 mM NaHCO3 1000 U/ml peptide N-glycosidase F (PNGase F) in 20 mM NaHCO3 Acetonitrile Dowex AG50X12 (H+ form; 100 to 200 mesh; Bio-Rad), activated (see UNIT 12.7 for conditioned resin) 1 M HCl Glycosylation
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Supplement 22
Table 12.6.3 Preparation of Separating and Stacking Gels
Separating gels
Stacking gel
6%
10%
30% acrylamide/0.8% bis-acrylamide
2.4 ml
0.5 M Tris base, pH 8.8 1.5 M Tris base, pH 6.2
3.0 ml
Distilled water 10% SDS
6.6 ml 120 µl
5.0 ml 120 µl
3.0 ml 120 µl
6.1 ml 100 µl
10% APS TEMED
120 µl 12 µl
120 µl 12 µl
120 µl 12 µl
50 µl 10 µl
Total
15%
4%
4.0 ml
6.0 ml
1.33 ml
3.0 ml
3.0 ml 2.5 ml
12.252 ml 12.252 ml 12.252 ml
10.09 ml
70°C water bath or heating block Vacuum centrifuge Sonicator 0.45-µm Millex-LH/hydrophilic PTFE filter (Millipore) 2.5-ml syringe Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and Coomassie blue staining (UNIT 10.5) NOTE: Iodoacetamide is light sensitive. It should be prepared fresh before each use. Run gel 1. Prepare an 80 × 80 × 0.75–mm polyacrylamide gel as described in UNIT 10.1 (Laemmli gel method), but use the gel recipes given in Table 12.6.3. Always add APS and TEMED immediately before pouring gels. The 10% APS solution must be made fresh every time. If it fizzes, it is still active. The greater the percentage of bis-acrylamide the greater the cross-linking of the gel and hence the smaller the pore size. Higher bis-acrylamide concentrations lead to reduced glycan yield. Gels should be 15% for 10 to 70 kDa proteins, 10% for 70 to 200 kDa proteins, or 6% for ∼200 kDa proteins.
Reduce and alkylate protein 2. To 5 to 30 µg glycoprotein, add 0.5 M DTT to a final concentration of 50 mM and 5× SDS sample buffer to a final concentration of 1×. Incubate 10 min at 70°C. Reduction and alkylation are necessary to ensure optimal efficiency in digests using PNGase F and/or trypsin. Clearly the amount of protein required to release picomoles of sugar for analysis will depend on the molecular weight of the protein as well as how many glycosylation sites are occupied. For example, in the case of ovalbumin, which has a mol. wt. of 45 kDa and one glycosylation site, 4.5 µg protein equals 100 pmol. Note that overloading gels does not always yield more glycans. The protein bands should be as tight as possible. The method works better on a small scale; therefore, the gel pieces to be processed should be small as well. Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
3. Add 100 mM iodoacetamide to a final concentration of 10 mM and incubate 30 min at room temperature in the dark.
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4. Apply sample to a gel slot in a vertical mini-gel system and run at 25 mA of constant current per gel. Use 25 mM Tris/190 mM glycine/0.5% SDS as the running buffer. Stain/destain protein 5. Visualize protein in the gel by staining with Coomassie blue (UNIT 10.5) for 2 hr. 6. Partially destain by rinsing with 50% methanol/7% acetic acid for a few minutes and then overnight with 5% methanol/7% acetic acid at 4°C. Use <7.5% acetic acid, since this concentration is sufficient to fix the protein in the gel and minimizes the risk of losing sialic acid residues from acidic glycans. Incomplete destaining does not appear to have any adverse effect on the subsequent in situ deglycosylation.
7. Cut out the bands of interest, cut them into pieces of ∼1 mm2, and place in microcentrifuge tubes that have been rinsed with SBPD water. 8. Wash gel pieces with 300 µl of 20 mM NaHCO3, vortex, centrifuge briefly, and soak 30 min at room temperature. Discard wash. Repeat. This procedure replaces the previous buffers.
9. Wash 60 min in 300 µl of 1:1 acetonitrile/20 mM NaHCO3 to remove residual SDS. 10. Dry gel pieces in a vacuum centrifuge. Perform in situ digestion of N-glycans with PNGase F 11. To the gel pieces add 3 µl of 1000 U/ml PNGase F and 27 µl of 20 mM NaHCO3. Leave for ∼5 min for the gel to reswell. The volumes given are for two to three gel bands.
12. Cover the gel with additional buffer (for a total of 70 to 100 µl), adding 10 µl at a time until the gel is covered. Incubate 12 to 16 hr at 37°C. 13. Vortex the sample and centrifuge 5 min at 1100 × g, room temperature. Extract glycans from gel pieces 14. Remove and retain the supernatant, placing it in a 1.5-ml microcentrifuge tube. 15. Add 200 µl SBPD water to the gel and sonicate 30 min. Remove buffer and add to supernatant. 16. Repeat two additional times with 200 µl water and once with 200 µl acetonitrile. Add each wash to the supernatant. The acetonitrile shrinks the gel.
17. Add 30 µl activated Dowex AG50X12 (H+) slurry to the sample, vortex, and incubate 5 min at room temperature to desalt. 18. Centrifuge 5 min at 550 × g, room temperature. 19. Filter the supernatant with a 0.45-µm Millex-LH/hydrophilic PTFE filter attached to a 2.5-ml syringe, collecting the filtrate in a 1.5-ml microcentrifuge tube. The sample can now be stored frozen at −20°C for up to a year. It can be cleaned up as described later for direct analysis by MALDI-MS or can be labeled with 2-AB (see Basic Protocol 6).
Glycosylation
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BASIC PROTOCOL 3
IDENTIFICATION/CONFIRMATION OF GLYCOPROTEIN BY IN-GEL TRYPSIN DIGESTION This procedure is based on the method of Küster et al. (1997). After the sugars have been released, the identity of the protein can be confirmed by in situ trypsin digestion, MS analysis of the recovered fragments, and comparison of the obtained masses or peptide sequences with a protein database. Trypsin digests can be performed either directly on the reduced and alkylated glycoprotein or after in situ deglycosylation with PNGase F. Materials Gel pieces containing glycoprotein of interest Acetonitrile 20 and 50 mM ammonium hydrogen carbonate (NH4HCO3) in sub-boiling-point-distilled (SBPD) water 0.1 mg/ml sequencing-grade trypsin (Roche Diagnostics) solution in SBPD water 10% (v/v) formic acid in SBPD water Vacuum centrifuge Wash gel pieces 1. Place gel pieces in 1.5-ml microcentrifuge tubes. 2. Wash 15 min in 500 µl acetonitrile. Discard acetonitrile. 3. Wash 15 min in 500 µl of 50 mM NH4HCO3. Discard NH4HCO3. 4. Repeat steps 2 and 3. 5. Dry gel pieces in a vacuum centrifuge. Digest with trypsin 6. Dilute 0.1 mg/ml sequencing-grade trypsin solution 1/8 (v/v) in 20 mM NH4HCO3 (final 12.5 µg/ml). 7. Add 15 µl of 12.5 µg/ml sequencing-grade trypsin solution and 5 µl of 20 mM NH4HCO3 to the gel pieces. 8. Incubate 15 min at room temperature. Retrieve peptide 9. Remove solution from gel pieces and collect into 0.5-ml microcentrifuge tubes. 10. Add 10 µl of 10% formic acid and 10 µl acetonitrile to the gel pieces. 11. Incubate 15 min at room temperature. 12. Remove solution from gel pieces and pool with previously collected solution in the microcentrifuge tube. 13. Add 15 µl acetonitrile to the gel pieces. 14. Incubate 15 min at room temperature. 15. Remove solution from gel pieces and pool with previously collected solution in the microcentrifuge tube. 16. Dry sample in a vacuum centrifuge.
Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
The sample is now ready for sequence analysis by mass spectrometry (UNIT 12.7).
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MANUAL HYDRAZINOLYSIS TO RELEASE N- AND O-GLYCANS This procedure involves cleavage of the N- or O-glycosidic linkage with anhydrous hydrazine under conditions designed to optimize the release and minimize the degradation of sugars. In the case of O-linked sugars, “peeling” (Fig. 12.6.12) is difficult to eliminate, but the conditions in this protocol generally give <15% of peeled structures. At present there is no other method for the release of O-linked glycans, which leaves the reducing end of the glycan clear for labeling with a fluorescent tag. Specialized equipment is needed for the handling of hydrazine and for cryo-drying of the sample (Ashford et al., 1987). The methods set out below cover the preparation of the sample and work-up following hydrazinolysis.
BASIC PROTOCOL 4
IMPORTANT NOTE: All glassware should be soaked in 4 M nitric acid overnight, then rinsed with at least ten washes of purified water. Powder-free gloves should be worn so as not to contaminate the sample. A sheet of clean aluminium foil makes a good clean surface to work on. Purified water (TOC < 10 ppb) should be used throughout. Materials 0.01 to 2 mg/ml glycoprotein sample Sub-boiling-point-distilled (SBPD) or purified water 0.1% (v/v) trifluoroacetic acid (TFA) Liquid nitrogen Lyophilizer Dry hydrazine (spec: ∼0.1% v/v H2O by GC – measured on a 10 ft × 1⁄4 column of Fluoropak/20% UCOW 550; thermal conductivity detector; Ashford et al., 1987) Argon Anhydrous toluene Acetic anhydride Saturated sodium bicarbonate Dowex AG50X12 resin (H+ form; 200 to 400 mesh; Bio-Rad) 1 M HCl 8:2:1 and 4:1:1 (v/v/v) butanol/ethanol/water Flow dialyzer (use membrane with appropriate pore size for sample) Hydrazinolysis tubes (covered with perforated Teflon tape) New acid-washed long Pasteur pipets Incubation equipment at 60° and 85°C Vacuum pump and vapor trap Poly-Prep disposable chromatography columns (Bio-Rad) Rotary evaporator with clean glass tubes Whatman 3MM chromatography paper Pinking shears Chromatography tank: 60 cm tall, equipped with solvent troughs and lid Waxed paper Teflon 2.5-ml Fortuna Norm-ject syringe 0.45-µm 4-mm hydrophilic PTFE Millex-LH filter (Millipore) Dialyze and freeze dry sample 1. Dialyze 10 µg to 2 mg glycoprotein sample (in a volume of ≤1 ml) in a flow dialyzer against water (purified or SBPD) or 0.1% trifluoroacetic acid ≥16 hr at a flow rate of ∼0.5 ml/min to remove all traces of salts or detergents.
Glycosylation
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Supplement 22
It is preferable to use a flow dialyzer, as dialysis bags can introduce carbohydrate impurities. If using dialysis bags, use a flask with nitrogen sparge at 4°C. If using TFA to solubilize the protein, dialyze at 4°C to avoid hydrolyzing sialic acid residues.
2. Transfer dialyzed glycoprotein into a hydrazinolysis tube. Use new acid-washed long Pasteur pipets for the transfer and avoid spilling the sample on the neck of the tube. Rinse neck with water if spillage does occur. Do not fill bulb more than half full.
3. Cover the tube with perforated Teflon tape. Place tube in a clean 50-ml plastic tube (to catch sample in case glass breaks during freezing). 4. Freeze carefully in liquid nitrogen while rotating the tube. Rotating the tube helps avoid expansion of the sample as it freezes, and thus breakage of the glass tube.
5. Lyophilize overnight. 6. Remove final traces of water by subjecting the sample to cryogenic drying for 2 days. Perform hydrazinolysis 7. Add 100 µl dry hydrazine by syringe to the 0.5 mg glycoprotein in each hydrazinolysis tube and flame seal under argon at atmospheric pressure. Carry out this and all subsequent manipulations under argon.
8. Incubate 6 hr at 60°C for O-link glycan release. For N-link release, ramp up the temperature to 85°C at 10°C/min, then incubate 12 hr at 85°C. O-glycan release conditions may also release low levels of N-glycans. N-glycan release conditions will release O-glycans, but they will be partially degraded.
9. Remove hydrazine under vacuum and condense hydrazine vapor in a trap. Complete hydrazine removal by repeated addition of anhydrous toluene (five additions of 10 drops each), followed by evaporation. Immediately place samples on ice. Hydrazinolysis causes the partial removal of acetyl groups from GlcNAc, GalNAc, and N-acetylneuraminic acid (NeuNAc), as well as cleaving peptide bonds. The exposed amino groups are re-N-acetylated to redress this change.
Perform re-N-acetylation 10. Add a five-fold molar excess of acetic anhydride and enough saturated sodium bicarbonate to bring the acetic anhydride to 0.5 M. Incubate 10 min at room temperature. First calculate the number of millimoles of amino acids in the sample, which is the mass of sample in mg divided by 110 (e.g., a 2-mg sample has 0.018 mmol amino acids). Then calculate the volume of acetic anhydride required to give a five-fold molar excess. Acetic anhydride is 10.6 M (0.018 mmol amino acids will require 0.09 mmol acetic anhydride, i.e., 8.6 µl). Finally, calculate the volume of saturated NaHCO3 needed to adjust the acetic anhydride to 0.5 M. The total volume should be 8.6 µl × 10.6/0.5 = 182 µl. Therefore, 182 − 8.6 = 173.4 µl saturated NaHCO3 is required. In practice, for samples of 2 mg or less, 200 µl NaHCO3 and 10 µl acetic anhydride are used. Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
11. If the acetic anhydride has all been consumed, add a second aliquot. A phase separation indicates that the acetic anhydride is in excess.
12. Incubate 50 min at room temperature.
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Desalt sample 13. Prepare an AG50X12 column in a Poly-Prep disposable plastic column. Calculate the volume of resin needed to give a five-fold excess of H+ resin over Na+ from the NaHCO3 (assumed 1 M). For example, for a 2-mg sample, 173.4 µl NaHCO3 was added. AG50X12 exchanges 2.3 meq Na+/ml. Therefore, 173.4 × 5/2.3 = 377 µl of resin is needed.
14. Wash resin with 10 vol of 1 M HCl followed by ≥10 vol of water until the pH of the eluant is the same as water. 15. Apply the re-N-acetylated sample to the column and elute with 5 column vol of water into a clean glass rotary evaporator tube. Evaporate to dryness. To avoid losing sialic acid do not allow the temperature to exceed 30°C.
16. Reconstitute the sample in 1 ml water and repeat the evaporation procedure three times to remove all traces of acetic anhydride. 17. Add 0.5 ml water and concentrate the sample to the bottom of the tube. 18. Centrifuge 5 min at 500 × g, room temperature, and repeat step 17 with 0.25 ml water. Separate released glycans from peptides 19. Cut 50-cm lengths of Whatman 3MM chromatography paper with pinking shears, as this helps the solvent to flow evenly. Wash with purified water by descending chromatography for 24 hr. Hang up to dry. 20. Reconstitute the sample in 25 µl water and apply to prewashed chromatography paper, allowing the spot to dry in between applications. Wash tube with 25 µl water and apply to paper. Allow spot to dry thoroughly. 21. Separate glycans from peptides by descending chromatography using a tank with fresh solvent (∼2 in. deep) and filter paper–lined walls (allow time for the tank to equilibrate). For O-linked glycans, run for 48 hr with 8:2:1 butanol/ethanol/water. For N-linked glycans, run for 60 hr with 4:1:1 butanol/ethanol/water. N-linked and large O-linked glycans will remain at the origin. Small O-links (di- and trisaccharides) will move down the paper behind the peptide. Monosaccharides elute with the peptide and cannot easily be recovered.
22. Dry paper in a fume hood. Elute glycans 23. Cut out the section of the paper containing the sugars (approximately −1 cm to +5 cm from origin). Holding the section in waxed paper (so as not to handle sample), roll it up and place it in a Teflon 2.5-ml Fortuna Norm-ject syringe with a 0.45-µm 4-mm hydrophilic PTFE Millex-LH filter. 24. To elute the sugars apply 1.5 ml water to the top of the paper, leave for 5 min, and push through the filter into a clean tube. Repeat three more times and evaporate all the filtrate to dryness. 25. Reconstitute the sample in 25 µl water and transfer to a microcentrifuge tube. Wash the collection tube three times with 25 µl water. Store samples in solution at −20°C until ready for labeling (stable for several months). The sample can now be stored frozen at −20°C for up to one year. It can be purified as described for direct analysis by MALDI-MS (UNIT 12.7) or can be labeled with 2-AB (see Basic Protocol 6).
Glycosylation
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BASIC PROTOCOL 5
AUTOMATED CHEMICAL RELEASE OF N- AND O-GLYCANS Currently there is no alternative to manual hydrazinolysis for optimal O-glycan release (<10% degradation). In practice, however, useful information can usually be derived from the automated O-mode. Please note that detailed operating instructions for the GlycoPrep are in the manufacturer’s instructions. Materials Glycoprotein sample 0.1% (v/v) trifluoroacetic acid (TFA) Sub-boiling-point-distilled (SBPD) water Microflow dialyzer with membrane with cutoff appropriate for sample GlycoPrep 1000 and reaction vials Lyophilizer Rotary evaporator or similar apparatus Acid-washed Pasteur pipets PCR tubes Vacuum centrifuge 1. Dialyze glycoprotein sample against 0.1% TFA at 4°C in a microflow dialyzer at a flow rate of 0.5 ml/min for ≥16 hr. Complete analysis may be performed on <250 µg material, and frequently 50 µg of glycoprotein may be used. The total volume should be <1 ml after dialysis. If necessary, lyophilize and redissolve in a smaller volume before dialysis. The hydrazinolsysis reaction is inhibited by metal salts, detergents, and many other ions. The samples should be essentially salt free. Some protein may precipitate if dialyzed against water, but they may be kept in solution in very dilute TFA. Since desialylation can occur at low pH, it is essential that the sample be maintained at 4°C during dialysis.
2. Transfer sample to a GlycoPrep reaction vial, making sure that the total volume is <1 ml. 3. Lyophilize 24 hr or until the sample is ready to run. If the sample is stored frozen after lyophilization, it will contain some condensed moisture. It should be lyophilized for an additional 2 hr before hydrazinolysis.
4. Set up a GlycoPrep 1000 with new column sets (either N+O or O only) and collection vials. 5. Select desired program (N+O, N*, or O). The programs differ in the reaction times and conditions: 5 hr at 100°C for N*, 5 hr at 95°C for N+O, and 5 hr at 65°C for O. Additional peptide removal material is included in the O only columns.
6. Load one or two samples directly from the lyophilizer. 7. Start the instrument run. The instrument performs a system check and then commences the run. The complete run takes ∼18 hr. Samples in collection vials may be removed when the protocol “System Wash” commences. This is ∼2 hr before the end of the run. The sample is delivered in an acid solution, and desialylation may occur if it is left standing. Thus, the sample should be removed immediately after collection and dried at once.
Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
8. Dry sample down on a rotary evaporator in two stages using 2.5-ml aliquots. Transfer sample using an acid-washed Pasteur pipet. Wash the tube twice with 0.5 ml SBPD water to maximize recovery. If a vacuum centrifuge is used, it is necessary to divide the sample into smaller aliquots (∼1 ml) for drying.
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9. Redissolve sample in 200 µl SBPD water and transfer to a PCR tube for subsequent labeling. Dry in a vacuum centrifuge. The sample can now be stored frozen at −20°C for up to one year. It can be purified as described for direct analysis by MALDI-MS (UNIT 12.7) or can be labeled with 2-AB (see Basic Protocol 6).
FLUORESCENT LABELING OF THE GLYCAN POOL WITH 2-AB The Glyko Signal 2-AB labeling kit (K-404) is used for efficient (∼100%) nonselective labeling of any salt-free glycan or glycan pool. Sugars are labeled at their reducing terminus (Fig. 12.6.12) for high-sensitivity fluorescent detection (10 to 50 fmol sugar gives an acceptable signal-to-noise ratio) following WAX-, NP-, and RP-HPLC.
BASIC PROTOCOL 6
Materials Glyko Signal 2-aminobenzamide (2-AB) labeling kit (K-404) Glycoprotein sample Sub-boiling-point-distilled (SBPD) water 30% (v/v) acetic acid in SBPD water Acetonitrile PCR tubes 65°C incubator or heating block GlycoClean S cartridges (Glyko) Lubricant-free 2.5-ml Fortuna syringe 0.45-µm hydrophilic PTFE Millex-LG filters (Millipore) Rotary evaporator and tubes, acid-washed Vacuum centrifuge 1. Mix the kit components as directed by the manufacturers. Excess labeling mixture can be stored for use later; if frozen immediately and stored at −20°C until just before use, it may be kept for ∼3 months.
2. Completely dry down a glycoprotein sample into the bottom of a PCR tube. Add 5 µl labeling solution per 50 nmol glycan dried in a 200-µl PCR tube. Incubate 2 hr at 65°C. 3. Immediately before use, wash a GlycoClean S cartridge eight times with 1 ml SBPD water. The filter may not allow flow when first wetted. Flow may be started by the application of slight pressure to the top of the filter.
4. Wash four times with 1 ml of 30% acetic acid. 5. Wash twice with 1 ml acetonitrile. 6. Remove 2-AB reaction tubes from oven and put in a freezer for 5 min to cool down. 7. Briefly microcentrifuge the tube and spot all sample over the surface of the cartridge. The cartridge should be still wet when applying sample. If it has dried out, an additional 1 ml acetonitrile should be applied.
8. Leave 15 min, rinse tube with 100 µl acetonitrile, and add to cartridge. Leave another 5 min. 9. Wash four times with 1 ml acetonitrile. 10. Place cartridge over a 2.5-ml Fortuna syringe fitted with a 0.45-µm hydrophilic PTFE Millex-LG filter. Glycosylation
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Supplement 22
11. Add 0.5 ml SBPD water and allow to pass through the cartridge into the syringe. 12. Wash cartridge three times with 0.5 ml SBPD water. 13. Push sample in the syringe through the filter into a rotary evaporator tube. Dry down completely in a rotary evaporator. 14. Redissolve sample in 200 µl SBPD water and transfer to a PCR tube for subsequent labeling. Dry in a vacuum centrifuge. 15. Redissolve in a suitable volume of SBPD water for HPLC analysis. For initial HPLC analysis, generally no more than 10% of the sample is required, and frequently less than that. Thus, reconstitution in volumes of 200 µl to 2 ml are appropriate. The sample is stable in solution for several weeks at 4°C, but should be frozen for extended storage. BASIC PROTOCOL 7
MULTIDIMENSIONAL SEPARATION AND PROFILING OF 2-AB-LABELED GLYCANS USING NP-, RP-, AND WAX-HPLC Complementary information can be obtained by resolving a glycan pool on several systems. Figure 12.6.17 shows such an analysis of the glycan pool released from prion protein isolated from Syrian hamster brains (Rudd et al., 1999b). An HPLC system capable of delivering a highly reproducible gradient and a sensitive fluorescence detector are required. The Waters system is detailed below, but others are also appropriate. Materials Glycoprotein sample 80% (v/v) acetonitrile HPLC standards (see recipe): dextrose for NP-HPLC, arabinose for RP-HPLC, and fetuin N-glycans for WAX-HPLC HPLC solvents appropriate for method: Solvent A (see recipe; NP): 50 mM formate, adjusted to pH 4.4 with ammonia Solvent B (NP): HPLC-grade acetonitrile Solvent C (see recipe; RP): 50 mM formate, adjusted to pH 5 with triethylamine Solvent D (RP): 50:50 (v/v) solvent C/acetonitrile Solvent E (see recipe; WAX): 500 mM formate, adjusted to pH 9 with ammonia Solvent F (WAX): 10:90 (v/v) methanol/water Waters Alliance 2690XE Separations Module: combined autosampler, injector, and column heater Waters 474 or Jasco Fluorescence Detector (for 2-AB, excitation: λmax 330 nm, band width 16 nm; emission λmax 420 nm, band width 16 nm) Pentium PC Waters Bus SAT/IN interface for data collection for 474 Waters Bus LAC/E interface, internal board for PC In-line degasser Column: Normal phase: GlycoSep N (Glyko), 4.6 × 250–mm (or equivalent, e.g., TSK amide silica columns), column temperature 30°C Reversed phase: GlycoSep R (Glyko), 4.6 × 150–mm, column temperature 30°C WAX: Vydac protein WAX, 7.5 × 50–mm (P/N 301 VHP 575) or GlycoSep R (Glyko), 4.6 × 100–mm, column temperature ambient
Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
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NP-HPLC
80
85
90
95
100
105
110
115
120
125
130
Fluorescence
RP-HPLC
40
50
60
70
80
90
100
110
120
130
140
150
WAX-HPLC
0
5
10
15
20 Time (min)
25
30
35
40
Figure 12.6.17 Resolution of the N-glycans attached to prion protein from scrapie-infected hamster brain (PrPSc27-30) using NP-, RP-, and WAX-HPLC.
Prepare sample 1. Prepare samples in 80% acetonitrile for NP-HPLC or in 100% water for WAX-and RP-HPLC. Store samples at 4°C. It is advisable to add water to the vial first, followed by the sample in aqueous solution, and then the acetonitrile, as this reduces the risk of precipitation.
Perform HPLC 2a. For NP-HPLC:Run the start-up method (to preserve the life of the column) followed by the separation method with no injected sample (this helps reproducibility). Then run the dextran standard followed by the samples.
Glycosylation
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Start-up method (SUM-NP), run time 30 min: 0 min 0 ml/min 20:80 (v/v) solvent A/solvent B 4 min 1 ml/min 20:80 8 min 1 ml/min 95:5 13 min 1 ml/min 95:5 16 min 1 ml/min 20:80 25 min 1 ml/min 20:80 26 min 0.4 ml/min 20:80 60 min 0.4 ml/min 20:80 61 min 0 ml/min 20:80 Separating method (SM-NP), run time 180 min: 0 min 0.4 ml/min 20:80 (v/v) solvent A/solvent B 152 min 0.4 ml/min 58:42 155 min 0.4 ml/min 100:0 157 min 1 ml/min 100:0 162 min 1 ml/min 100:0 163 min 1 ml/min 20:80 177 min 1 ml/min 20:80 178 min 0.4 ml/min 20:80 260 min 0 ml/min 20:80 261 min 0 ml/min 20:80 2b. For RP-HPLC: Run the start-up method (to preserve the life of the column) followed by the separation method with no injected sample (this helps reproducibility). Then run the arabinose standard followed by the samples. Start-up method (SUM-RP), run time 30 min: 0 min 0.05 ml/min 5:95 (v/v) solvent C/solvent D 5 min 1 ml/min 5:95 10 min 1 ml/min 5:95 20 min 1 ml/min 5:95 21 min 1 ml/min 95:5 28 min 1 ml/min 95:5 29 min 0.5 ml/min 95:5 90 min 0.5 ml/min 95:5 91 min 0 ml/min 95:5 Separating method (SM-RP), run time 180 min: 0 min 0.5 ml/min 95:5 (v/v) solvent C/solvent D 30 min 0.5 ml/min 95:5 160 min 0.5 ml/min 85:15 165 min 0.5 ml/min 76:24 166 min 1.5 ml/min 5:95 172 min 1.5 ml/min 5:95 173 min 1.5 ml/min 95:5 178 min 1.5 ml/min 95:5 179 min 0.5 ml/min 95:5 220 min 0.5 ml/min 95:5 221 min 0 ml/min 95:5 Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
If the RP system has been unused for a while, the system should be purged with buffers because the concentration of acetonitrile in the system decreases with time. Dextran hydrolysate is not suitable as a standard for the RP column, as it elutes in too short a time scale.
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Solvent D is designed to assist the reproducibility of the shallow gradient. The last two lines of SM-RP enable the system to be shut down automatically.
2c. For WAX-HPLC (charge separation): Run the start-up method (to preserve the life of the column) followed by the separation method with no injected sample (this helps reproducibility). Then check the system by running a standard set of charged glycans released from fetuin before loading samples. Start-up method (SUM-WAX), run time 30 min: 0 min 0 ml/min 0:0 (v/v) solvent E/solvent F 5 min 1 ml/min 0:100 90 min 1 ml/min 0:100 91 min 0 ml/min 0:100 Separating method (SM-WAX), run time 80 min: 0 min 1 ml/min 0:100 (v/v) solvent E/solvent F 12 min 1 ml/min 5:95 25 min 1 ml/min 21:79 50 min 1 ml/min 80:20 55 min 1 ml/min 100:0 65 min 1 ml/min 100:0 66 min 2 ml/min 0:100 77 min 2 ml/min 0:100 78 min 1 ml/min 0:100 120 min 1 ml/min 0:100 121 min 0 ml/min 0:100 CALIBRATION OF THE HPLC SYSTEM It is essential to calibrate the normal- and reversed-phase HPLC systems against temporal or other variations in factors such as column condition, pressure gradient, eluent concentration, or environmental conditions. A relatively complex calibration method is necessary for this purpose, employing a range of standard peaks and the use of curve fitting. Two methods are presented below by which the required calculations may be performed either manually or by the use of PeakTime, a special-purpose computer program (E.R. Hart, P.M. Rudd, and R.A. Dwek, unpub. data).
SUPPORT PROTOCOL
An external standard (dextran ladder for NP or arabinose ladder for RP) is run independently of the sample but under the same conditions. The elapsed time between the sample run and the standard run must be kept low. The maximum time allowed will depend on local conditions and should be kept under review; however, under stable conditions, one standard run per day should suffice, allowing a single standard run to be applied to several different samples. The standards used contain a mixture of oligomers of glucose or arabinose with elution times covering the whole range in which sample peaks are expected. Standard mixtures of glycans are sold for the purpose (Glyko). These have been defined above and contain linear oligomers of all lengths up to and beyond ∼20, in consistent proportions. Chromatograms produced from such mixtures take the form of a ladder, as shown in Figure 12.6.18. A curve-fitting technique is used to extract information from the standard HPLC log in a suitable form to calibrate the sample. PeakTime is designed both to fit the curve and to Glycosylation
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A 2
3
4
5
6
0.2
0.1 ×
0.0
×
×
×
×
×
15
7 8 9 10 12 11 × × × 13 × × × ×
10
GU
Fluoresence
1
5
0
0
100
50
150
B 1
2
3
0.2 ×
0.0 0
×
×
4
×
5
×
6
×
7 ×
10
8 ×
AU
Fluoresence
0.4
5
0 50
100
150
Time (min)
Figure 12.6.18 Standard ladder chromatograms and calibration curves for (A) dextran and (B) arabinose.
apply the results to the sample automatically; otherwise a computer spreadsheet or a specialized graphics package is the best option. There are several possible curve-fitting techniques and algorithms known; however, it is important to use the same one consistently for the results to be comparable with each other. Two standard scales are defined—the GU scale for NP-HPLC and the AU scale for RP-HPLC—each based on the glycan chain length of standard linear oligosaccharides. These scales both employ a least-squares fifth-order polynomial curve-fitting algorithm. The standard mixtures on which these scales are defined are derived by partial acid hydrolysis of dextran for the GU scale and of arabinose for the AU scale. The GU scale has the convenient property of increasing monotonically with glycan size. Manual Calibration of the HPLC System 1. Select a region of the ladder to use, such that each constituent ladder peak is clearly defined and distinguished from impurity peaks. The region should be as large as possible; however, peaks at the ends of the ladder should not be used if there is any difficulty in assigning them. It is very important to include the whole of the region in which sample peaks are expected. The highest reproducibility will be obtained if the same region is used on every occasion.
Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
2. Label the ladder peaks within the chosen region with integer values representing the associated glycan chain length. Check once again that the correct peaks have been identified, and that the numbers are consecutive and correctly represent the glycan chain lengths.
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Current Protocols in Protein Science
3. Record the integer value and the elution time for each peak within the chosen region. 4. Create a calibration curve by applying a curve-fitting algorithm to the pairs of data, treating the elution time as the independent variable (x axis) and the integer values as the dependent variable (y axis). Use the least-squares fifth-order polynomial algorithm. Note that this choice of axes is the reverse of that usually employed in the least-squares theory. This is not a problem as long as the same is done every time.
5. Plot the curve and inspect for irregularities. Within the selected region, the curve should be monotonic and without pronounced gradients, and should not be overly dissimilar in shape to others plotted previously. It is acceptable and common, however, for it to veer away sharply outside. Neither is it a problem if the curve does not pass exactly through every point, as this is the nature of a least-squares curve fit. However, the discrepancy should not be too large. Examples of GU and AU calibration curves are shown in Figure 12.6.18.
6. Use the calibration curve to determine a GU/AU value for the sample peaks. Be careful not to use any part of the curve outside of the selected region, as this would lead to nonreproducible results. Calibration of the HPLC System Using PeakTime PeakTime is a computer program developed at the Oxford Glycobiology Institute that automates the calibration process. PeakTime is an interactive rather than a fully automatic program; the user directs it through a series of steps that correspond exactly to those of the manual process described above. PeakTime has the advantage over the manual process that the numerical manipulation is done internally, including the automatic importation of results from the chromatography software. A further advantage is that it can quickly and easily calibrate an entire chromatogram and replot it to the nonlinear GU/AU scale axis, which would be difficult to do manually. This ability to plot a chromatogram to a calibrated scale is useful for the method of successive enzyme digests (see Basic Protocol 8), which depends on the detection of shifts in the peaks of a series of chromatograms. When plotted to the same scale, the shift due to a particular enzyme is likely to be of the same length at any point on the axis. Future updates planned for PeakTime include the addition of a database of known glycan GU/AU values, and the ability to automate the method of successive enzyme digests itself. SEQUENCING AN ENTIRE N-GLYCAN POOL SIMULTANEOUSLY USING EXOGLYCOSIDASE ARRAYS
BASIC PROTOCOL 8
In this protocol, the N-glycan pool is digested with specific exoglycosidases to establish an array of digestion patterns. Enzyme specificities and digest conditions are noted in Tables 12.6.4 and 12.6.5, and a schematic representation is given in Figure 12.6.19. The glycan pool is profiled by NP-, RP-, and WAX-HPLC (see Basic Protocol 7) and MALDI-MS (UNIT 12.7). If sample is limited, use only NP-HPLC, as this will give the most information for the minimum amount of material. Figure 12.6.20 and Table 12.6.6 show data for sequencing of the N-linked glycans of human neutrophil gelatinase B–associated lipocalin (Rudd et al., 1999a). Glycosylation
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Supplement 22
20 µU + 2 µl water 0.1 U + 10 µl water
Fuc α1-3, 4 Fuc α1-6 (>2,3,4)
GlcNAc, GalNAc β1-2, 3, 4, 6 As is GlcNAcβ1-4, 6 GlcNAcβ1-3, 6 Gal 30 mU + 100 µl water
AMF BKFd-i
JBHd,j,k SPHd,l,m,n
Man β1-4 GlcNAc
0.2 U in 10 µl water
10%
100% 8%
20% 40%
10% 10%
20% 20%
10% 10%
% of final volc
2 mU/µl
67 mU/µl 5.4 mU/µl
10 mU/µl 120 µU/µl
1 µU/µl 1 mU/µl
1 mU/µl 80 µU/µl
1 mU/µl 200 µU/µl
Final enzyme concentration
100 mM Na citrate-phosphate
100 mM Na acetate, 2 mM Zn 100 mM Na acetate, 2 mM Zn
100 mM Na citrate-phosphate 100 mM Na citrate-phosphate
50 mM Na acetate 100 mM Na citrate
100 mM Na citrate-phosphate 100 mM Na acetate
100 mM Na acetate 50 mM Na acetate
Final buffer concentration
4
5 5
5 6
5 6
5 6
5 5.5
Optimum pH
See manufacturers’ inserts for further details.
The following notes refer to inhibitory metal ions: dHg2+; Ca2+.
Add second aliquot at 12 to 18 hr.
o
n
d-n
Also includes 20% 5× incubation buffer.
c
b
e
L-fuctose;
Ni2+; gCu2+; hAg2+; i2 µM 4-chloromercuriphenylsulfonic acid; jAg+; kFe3+; lCd2+; mMg2+;
f
This table shows most commonly used enzymes. For further details refer to manufacturers’ inserts. Abbreviations: ABS, Arthrobacter ureafaciens sialidase; AMF, almond meal fucosidase; BKF, bovine kidney α-fucosidase; BTG, bovine testes β-galactosidase; HPM, Helix pomatia α-mannosidase; JBH, jack bean β-N-acetylhexosaminidase; JBM, jack bean α-mannosidase; NDVS, Newcastle disease virus sialidase; SPG, Streptococcus pneumoniae β-galactosidase; SPH, Streptococcus pneumoniae βhexosaminidase.
a
HPM
2 U in 30 µl 1× buffer 2 U in 30 µl 1× buffer
As is 40 mU + 100 µl water
Gal β1-3, 4 > 6 Gal β1-4
BTG SPG
JBM (high)d,j,o Man α1-2, 3, 6 Man α1-2, 6 JBM (low)d,j
0.2 U + 20 µl water 20 mU + 10 µl water
Make up as
Sialic acid α2-6 > 3,8 Sialic acid α2-3, 8
Specificityb
Conditions for Digestion of Glycan Pools by Exoglycosidase Enzymes
ABS NDVS
Enzymea
Table 12.6.4
Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
12.6.34
Current Protocols in Protein Science
Exoglycosidase Arrays Commonly Used in N-Glycan
Table 12.6.5 Sequencing
Enzyme
1
ABS
X
2
NDVS AMF
3
4
5
6
X
X
X
X
X
X
X
X
X
X X
X X
7a
X
BTG BKF SPH JBM (high)
X X
aJBM is used only when the presence of oligomannose structures is indicated, i.e., structures
resistant to other enzyme arrays.
JBM
BKF SPG,BTG
BKF
HPM ABS
BKF, AMF
JBM
BKF, AMF
JBH,SPH
ABS, NDVS
Figure 12.6.19 Diagram showing specificities of exoglycosidases. For enzyme abbreviations, see Table 12.6.4.
Materials N-glycan pool (2-AB labeled; see Basic Protocol 6) Exoglycosidase enzymes and buffers (Table 12.6.4) 5% and 100% (v/v) acetonitrile 50-µl microcentrifuge tubes Speedvac evaporator 0.45-µm high-protein-binding nitrocellulose microcentrifugal filter unit (e.g., Pro-Spin filters, P85100 Pro-Mem, Radley and Co.) Additional reagents and equipment for NP-HPLC (see Basic Protocol 7) Perform enzymatic digests 1. Pipet aliquots of the entire N-glycan pool into 200-µl microcentrifuge tubes and evaporate to dryness using a Speedvac evaporator. Glycosylation
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total N-glycan pool
10 11 7 89
7 9
4
6 ABS
Fluorescence
3 5
2
1
ABS BTG 3
ABS BTG BKF
1 1
ABS BTG BKF AMF
5
Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
6
7
8 Glucose units
9
10
11
Figure 12.6.20 Sequencing N-linked glycans from neutrophil gelatinase B–associated lipocalin. For enzyme abbreviations, see Table 12.6.4.
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Table 12.6.6 N-Glycans
Peak
HPLC Analysis of Exoglycosidase Digestions of the PNG-released, 2-AB-labeled Pool of NGAL
% Assigned undigested peaks
Assignment
Undigested peaks
Digests ABS
ABS/BTG
ABS/BTG/ ABS/BTG/ BKF BKF/AMF
1
A2G0
5.50
5.50
2 3
A2G0F A2G1F (to GN)
5.93 7.07
7.06
4 5
A2G2 A2G1F (core) F (to GN)
7.15
6 7
14.7
A2G2F A2G2F (to GN)
7.96
7.57 7.96
8 9
6.2 8.9
A2G2S2 A2G2F (core) F (to GN)
8.28 8.39
8.38
10 11
38.2 32.0
A2G2SF (to GN) A2G2SF (core) F (to GN)
8.64 9.04
5.53
7.50
2. Perform enzymatic digest as described in Table 12.6.4, following the arrays set up in Table 12.6.5, in a volume of 10 to 20 µl buffer. Digest 18 hr in a 37°C water bath. For example, for a triple digest with ABS, AMF, and BTG, use: 1 µl ABS 1 µl AMF 2 µl BTG 2 µl 5× incubation buffer (250 mM sodium acetate, pH 5.5) 4 µl water. Gently spin the contents to the bottom of the tube from time to time, so that water condensing on the lid does not cause the concentrations in the digest to change. In the case of combined digests, use the average optimum pH conditions of the constituent enzymes. All enzyme digests should be carried out with reference to the suppliers’ literature. The enzymes noted in Table 12.6.4 are supplied by Glyko (or, alternatively, Seikagaku Kogyo, Roche, and Sigma). Full technical information and storage conditions can be found in the manufacturers’ information and catalogs.
3. Remove reaction vial from the water bath and briefly spin down the contents to the bottom of the tube. Remove enzymes 4. Prewash a 0.45-µm high-protein-binding nitrocellulose microcentrifugal filter unit with 100 µl of 5% acetonitrile in water and spin. Discard wash. The sample is filtered to remove the majority of enzymes and the carrier protein, BSA, in order to prevent contamination of the HPLC column. The NP-HPLC column (Glyko GlycoSep N) is not sensitive to buffer salts. However, sialidase activity has been noted on the columns following many analyses of enzyme digestions. Filtering the samples prior to HPLC is good laboratory practice. IMPORTANT NOTE: Do not denature the enzyme by heating, as this may remove some of the 2-AB label. Glycosylation
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5. Wash with 100 µl of double-distilled water and spin. Discard wash. 6. Add the sample to the filter and allow to disperse into the membrane for 30 min. 7. Spin the sample through the membrane and then add 20 µl of 5% acetonitrile in water. 8. Leave 5 min and then spin through the filter. Repeat once to maximize the recovery of glycans from the membrane. IMPORTANT NOTE: It is very important that the 5% acetonitrile solution is made fresh every time. Over time, the percentage of acetonitrile will reduce as a result of evaporation, eventually leading to reduced recovery from the filter membrane.
9. Dry the sample and dissolve in 20 µl water. Add 80 µl acetonitrile and inject onto an NP-HPLC column (see Basic Protocol 7). 10. Assign GU values to the NP-HPLC profiles of the digests and stack them in order (1 to 6) so that the changes in the elution position of each peak can be followed from the initial profile through the different enzyme digests. Peaks are assigned from a knowledge of the specificity of the enzymes in the arrays (Table 12.6.6) and the incremental values of individual monosaccharide residues (Table 12.6.1; Guile et al., 1996). The final enzyme array for N-glycan pools is designed to digest all the oligosaccharides to their basic cores. This means that sugars containing less-common oligosaccharides such as xylose, or sugars containing substitutions such as sulfate or phosphate, will be only partially digested and can be recovered and examined independently. BASIC PROTOCOL 9
SEQUENCING O-GLYCANS This protocol represents the authors’ current progress in developing HPLC-based methods for analyzing O-glycosylation. O-linked glycans are more complicated to sequence than N-linked sugars. This is partly because the heterogeneity can be very complex, particularly in glycan pools released from mucins (Fig. 12.6.21). In addition, O-linked sugars are constructed from eight different core structures and have more complicated branching patterns than N-glycans. Of the O-linked core structures listed in Figure 12.6.7, four are diHexNAc and two are HexHexNAc, so mass alone cannot be used to identify them. The elution positions of these disaccharides on NP-HPLC are very similar, so RP-HPLC is required as a complementary separation method. A greater range of exoglycosidases than is currently available is needed to determine the branching of the O-glycans. The enzymes routinely used for N-link sequencing can all be used, but it is important to test out their specificity on standard O-linked glycans first. Mass data from MALDI or quadrupole time-of-flight (Q-TOF) analysis and fragmentation data from Q-TOF—by on-line liquid chromatography MS (LC-MS) or single injections—may also be needed to complement HPLC data to enable structures to be assigned. Figure 12.6.16 shows the GU and AU values for a range of O-glycans. Some incremental values for individual monosaccharide linkages are emerging, suggesting that as information accumulates the system will be able to be used predictively. The strategy for O-glycan analysis is shown in Figure 12.6.22. Sequencing the released glycans involves essentially the same enzymes as those used for N-linked glycans, but the arrays are different.
Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
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0.30
Fluorescence (V)
0.25 0.20 0.15 0.10 0.05 0.00 0.00
10.00
20.00
30.00
40.00 50.00
60.00
70.00
80.00
90.00 100.00 110.00 120.00
Time (min) 1
2
3
4
5
6
7
8
9
10
GU value
Figure 12.6.21 Profile of 2-AB-labeled glycan pool released from porcine stomach mucin.
Materials Exoglycosidase enzymes (Table 12.6.4) Chicken liver β-N-acetylhexosaminidase (CLH) Additional reagents and equipment for HPLC (see Basic Protocol 7) and arrays of enymatic digests (see Basic Protocol 8) 1. Resolve samples of the glycan pool by NP-, RP-, and WAX-HPLC (see Basic Protocol 7, Fig. 12.6.23). Assign preliminary structures to peaks on the basis of GU and AU values. 2. Take aliquots of the glycan pool and digest with the appropriate enzymes (see Basic Protocol 8 for procedure; see Table 12.6.7 for enzyme array). Analyze the digestion products by NP- and RP-HPLC. 3. If ABS digests, digest with NDVS. If ABS digests but NDVS does not, sialic acid is linked α2,6.
4. If BTG digests, digest with SPG. If this also digests, galactose is linked β1,4.
5. If BKF digests, digest with AMF. If BKF digests but AMF does not, fucose is α1,2 linked.
6. If JBH digests, digest with SPH. SPH digests only GlcNAc, whereas JBH digests both GalNAc and GlcNAc. Figures 12.6.24 and 12.6.25 show a selection of these digests, which enable the O-glycans attached to fetuin to be analyzed. Glycosylation
12.6.39 Current Protocols in Protein Science
Supplement 22
pool 2AB O-glycans MALDI-MS RP-HPLC or ESI-MS NP-HPLC MS/MS GU
AU
mass
fragmentation
compare to standards
complex profile collect peaks from NP-or RP-HPLC run
simple profile individual peaks to confirm peak allocations
to confirm peak allocations enzyme digests MALDI-MS or ESI-MS MS/MS
NP-HPLC RP-HPLC GU
AU
mass
fragmentation
compare to standards and initial allocations O-glycan sequence and linkages
Figure 12.6.22 A strategy for the analysis of O-glycans. Table 12.6.7 Sequencing
Exoglycosidase Arrays Commonly Used in O-Glycan
Enzyme
1
ABS
X
BTG BKF
2
3
4
X
5
6
7
X
X
X
X X
JBH
X X
X
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
Buffer A 5 ml 2 M ammonium formate, pH 4.4 50 ml Milli-Q-purified water Adjust pH to 8.6 Add 0.4 g SDS Bring to 100 ml Store up to 6 months at 4°C
12.6.40 Supplement 22
Current Protocols in Protein Science
Figure 12.6.23 Resolution of the O-glycans attached to fetuin using NP-, RP-, and WAX-HPLC.
Glycosylation
12.6.41 Current Protocols in Protein Science
Supplement 22
NP-HPLC
RP-HPLC
GU scale
AU scale
1
2
3
4
5
1
2
d
i
d j
b
Fluorescence
h
c
b
b
j
g
c h
ABS
h a
a b
b
e
e
ABS + BTG
c
b
c b h
b
h
ABS +JBH
c
b e
0
4
c
NDVS
g h
i
undig
c
b
3
f
f
20
40 Time (min)
e
ABS + SPG 60
80
g
0
20
40 Time (min)
60
80
Figure 12.6.24 Sequencing fetuin O-linked glycans using NP- and RP-HPLC. For enzyme abbreviations, see Table 12.6.4.
Buffer B 5 ml 2 M ammonium formate, pH 4.4 50 ml Milli-Q-purified water Adjust pH to 8.6 Add 1.2 g CHAPS (1.2% w/v) Add 3.72 g EDTA (0.1 M) Bring to 100 ml Store up to 6 months at 4°C HPLC standards Dextran ladder (NP-HPLC): 2-AB labeled homopolymer stranded (Glyko) Arabinose ladder (RP-HPLC): Label individual arabinose homopolymers with 2-AB and co-inject onto the RP column. Arabinose is available from Fluka, and arabinobiose to arabinooctaose forms are available from Dextra Laboratories.
Fetuin N-linked glycans (WAX-HPLC): Label fetuin N-linked glycans (Glyko) with 2-aminobenzamide. The mono-, di-, tri-, and tetrasialylated glycans are separated by WAX-HPLC. Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
12.6.42 Supplement 22
Current Protocols in Protein Science
GU (AU) values % in Peak undigested
Structure assigned
Undigested
NDVS
ABS
ABS + BTG
ABS + JBH
ABS + SPG
a
(0.82)
2AB
0.90(2.19)
0.90(2.19) 0.90(2.17)
0.90(2.17)
0.91(2.14)
0.90(2.17)
b
(0.37)
2AB
1.00(0.42)
1.00(0.42) 1.00(0.42)
1.00(0.42)
1.00(0.42)
1.00(0.42)
c
2.66
2AB
1.81(1.35)
1.81(1.41) 1.80(1.40)
1.81(1.38)
1.81(1.43)
1.80(1.40)
d
18.15
2AB
2.24(2.19)
e
2.73(1.68)
2AB
f
33.26
g
0.36
2AB
2AB
2.19(2.06) 3.22(2.54)
2AB
h i
7.81
j
1.73
2AB
2.72(1.70)
3.23(2.42) 3.58(1.68) 3.58(1.66)
3.59(1.68)
4.40(2.71)
2AB 5.38(3.50)
Figure 12.6.25 Structures, GU values, and AU values corresponding to peaks in Figure 12.6.24. For enzyme abbreviations, see Table 12.6.4.
SDS sample buffer, 5× 625 µl 0.5 M Tris⋅Cl, pH 6.6 (APPENDIX 2E) 1.0 ml 10% (w/v) SDS (APPENDIX 2E) 2.875 ml 10% glycerol 0.4 g bromphenol blue Store up to 1 year at −20°C Solvent A (for NP-HPLC) Stock solution (2 M formate): Prepare a 2-liter 2 M formate stock solution as for solvent E (see recipe), but adjust pH to ∼4.0 with ammonia (∼260 ml), allow to cool and settle outside the ice bath, and then adjust to pH 4.4. Working solution (50 mM): Dilute 50 ml stock to 2 liters using purified water. Prepare fresh. Solvent C (for RP-HPLC) Stock solution (1 M formate): Prepare a 1 M formate stock solution as described for solvent E (see recipe), but start with 92.06 g formic acid and adjust pH to 5 with triethylamine (BDH HiPerSolv for HPLC), which acts as an ion-pairing agent. Working solution (50 mM): Dilute 100 ml stock to 2 liters using purified water. Prepare fresh. Glycosylation
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Solvent E (for WAX-HPLC) Stock solution (2 M formate): Place a 2-liter beaker in an ice bath containing salt to lower the temperature of the ice to −10°C. Weigh 184.12 g formic acid (BDH Aristar grade) into the beaker using a glass acid-washed pipet. Turn off the fume hood while weighing, since air flow will affect the balance. Add 1 liter water while stirring. Adjust pH to ∼8.5 by dropwise addition of 35% ammonia solution (BDH Aristar grade). Remove beaker from ice and allow the warm solution to cool to room temperature (∼1 hr). Adjust pH to 9.0 with ammonia solution, added dropwise. Transfer solution to a 2-liter volumetric flask, bring to volume using purified water, and note the final pH. Transfer to a brown bottle and store up to several months at room temperature. Working solution (500 mM): Dilute 500 ml stock to 2 liters using purified water. Check pH before using. Prepare fresh. CAUTION: DO NOT add ammonia solution in large quantities as this will produce a rapid and potentially dangerous increase in temperature. When using formic acid or ammonia, place the pH meter and balance in an empty fume hood. Wear a laboratory coat, gloves, and goggles.
COMMENTARY Background Information
Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
Within the past three years major advances in technology available for separating and sequencing oligosaccharides from glycoproteins have brought glycan analysis of micromolar amounts of biological samples within the reach of any well-founded laboratory. It is now possible to release N-linked oligosaccharides directly from 5 to 30 µg of protein in an SDSPAGE gel band and analyze them by mass spectrometry (UNIT 12.7) or HPLC (Basic Protocol 7). HPLC strategies allow structures of subpicomolar levels of neutral and sialylated oligosaccharides to be predicted from a single run, while further information can be obtained rapidly using multiple exoglycosidase arrays to analyze all of the components of a glycan pool simultaneously. When the three-dimensional structure of the protein and oligosaccharide analysis are both available, the new database of preminimized oligosaccharide structural units can be assembled appropriately and modeled onto the protein. The justification for this stems from the fact that, in contrast to proteins, where tertiary and quaternary interactions play a significant role, the main structural features of oligosaccharides are determined by their secondary structure, which involves interactions across individual glycosidic linkages. Molecular modeling of glycoproteins requires an oligosaccharide analysis of the sugars associated with the protein. However, the classical strategies and techniques used to locate glycans and separate and analyze the oligosac-
charides released from the heterogeneous mixture of glycoforms (for review, see Dwek et al., 1993), although effective, were time consuming and difficult. Moreover, the poor resolution achieved by the separation technologies and the insensitivity of the detection systems precluded the analysis of the glycosylation of many biologically important glycoproteins that are only available in microgram quantities or less. It is important to retain a flexible and interactive approach to sequencing strategies to obtain specific information required to answer particular biological questions. However, there is also a need for a robust technology that is generally applicable to all glycoproteins, so that oligosaccharide sequencing at the subpicomolar level can become as rapid and automated as peptide sequencing, and routinely available to protein chemists as well as glycobiologists. It is such a technology that is described in this unit and in UNIT 12.7, which is based solely on methodology used in the Glycobiology Institute.
Critical Parameters and Troubleshooting When carrying out procedures on samples that might be analyzed by mass spectrometry (UNIT 12.7), it is essential to use the highest quality solvents for every purpose. In particular, purified water sometimes contains polymeric material (principally polyethylene glycol) that will co-concentrate with the sugars and impair MALDI-MS detection. To avoid this problem, purified water should be further
12.6.44 Supplement 22
Current Protocols in Protein Science
Table 12.6.8 Nomenclature (Terms and Examples) for Describing N-linked Oligosaccharide Structures
Nomenclature
Definition
Terms A(1-4) B
Number of antennae linked to trimannosyl core Bisecting N-acetylglucosamine (GlcNAc)
F Fo
Core fucose α1,6 linked Outer arm fucose α1,3 linked
G(0-4) Gal
Number of (sub)terminal galactose residues in the structure Galactose
GalNAc
Reducing terminal N-acetylgalactosamine of mucin-like O-linked glycans
M N
Mannose N-Acetylglucosamine in the chitobiose core of N-glycans
S Examples
Sialic acid; the conformation of the linkage is noted in brackets
MNNF M3N2
Fucosylated N-linked trisaccharide core Conserved N-linked pentasaccharide core
M3N2F A2G0, A3G0, and A4G0
Fucosylated N-linked pentasaccharide core Agalactosyl bi-, tri-, and tetraantennary complex glycans
A2G0F A2G0FB
Core fucosylated agalactosyl biantennary complex glycan Core fucosylated and bisected agalactosyl biantennary complex glycan
A2G2, A3G3, and A4G4 A2G2S(3)
Bi-, tri-, and tetraantennary complex glycans α2,3-linked monosialylated biantennary complex glycan
A2G2S2(3)2 A2G2S(6)
α2,3-linked disialylated biantennary complex glycan α2,6-linked monosialylated biantennary complex glycan
A2G2S2(6)(3)
Disialylated biantennary complex glycan with one α2,3- and one α2,6-linked sialic acid α2,6-linked disialylated biantennary complex glycan
A2G2S2(6)2 A3G3Fo
Triantennary complex glycan with one outer arm fucose α1,3-linked to GlcNAc
treated, e.g., by distilling at sub-boiling-point temperatures (SBPD) to prevent organic contaminants from being carried across with the steam. For purposes where purified water is adequate the quality should be at least TOC <10 ppb (e.g., Milli-Q-purified water). Powder-free gloves should be worn at all times, and work should ideally be carried out on ceramic or glass bench tops to avoid contaminating the sample with carbohydrate from the environment. All glassware should be washed with 4 M nitric acid for 24 hr and rinsed with ten changes of purified water and finally with SBPD water.
Acetonitrile is a source of artefacts both on the HPLC and on MS. A suitable source of acetonitrile for this purpose can be obtained from Allied Signal Riedel-de Haen (available via Sigma). Glycans should not be left dried in tubes for long periods, as they may subsequently be hard to remove from vial/tube walls.
Anticipated Results Representative chromatograms and profiles are presented in figures within each of the specific protocols. The structures should be consistent with known biosynthetic pathways. If not, care must be taken to confirm the assign-
Glycosylation
12.6.45 Current Protocols in Protein Science
Supplement 22
ments, as novel structures may imply novel pathways.
Time Considerations The time required for each protocol is below. Figure 12.6.10 offers additional information on interrelation of the protocols. PNGase F release of N-linked glycans (Basic Protocol 1) requires 2 days. In-gel enzymatic release (Basic Protocol 2) will also take 2 days, as will the identification of glycoproteins through in-gel trypsin digestion (Basic Protocol 3). Manual hydrazinolysis (Basic Protocol 4) can take anywhere from 5 to 8 days to complete. Automated hydrazinolysis (Basic Protocol 5) will require 4 to 5 days to complete. Fluorescent labeling of the glycan pool with 2-AB (Basic Protocol 6) takes 3 to 5 hr. HPLC separation (Basic Protocol 7; Support Protocol) of the pool can take up to 6 hr, depending on the system used. Sequencing the N-glycan (Basic Protocol 8) or O-glycan (Basic Protocol 9) pools requires 2 days each.
Literature Cited Ashford, D., Dwek, R.A., Welply, J.K., Amatayukul, S., Homans, S.W., Lis, H., Taylor, G.N., Sharon, N., and Rademacher, T.W. 1987. The β1-2-D-xylose and α1-3-L-fucose substituted N-linked oligosaccharides from Erythrina crystagalli lectin. Isolation, characterisation and comparison with other legume lectins. Eur. J. Biochem. 166:311320. Clausen, H. and Bennett, E.P. 1996. A family of UDP-GalNAc:polypeptide N-acetylgalactosaminyl-transferases control the initiation of mucin-type O-linked glycosylation. Glycobiology 6:635-646. Dwek, R.A., Edge, C.J., Harvey, D.A., Wormald, M.W., and Parekh, R.B. 1993. Analysis of glycoprotein associated oligosaccharides. Annu. Rev. Biochem. 62:65-100. Goodarzi, M.T., and Turner, G.A. 1998. Reproducible and sensitive determination of charged oligosaccharides from haptoglobin by PNGase F digestion and HPAEC/PAD analysis: glycan composition varies with disease. Glycoconj. J. 15:469-475. Guile, G.R., Wong, S.Y., and Dwek, R.A. 1994. Analytical and preparative separation of anionic oligosaccharides by weak anion-exchange highperformance liquid chromatography on an inert polymer column. Anal. Biochem. 222:231-235.
Determining the Structure of Oligosaccharides N- and O-Linked to Glycoproteins
Guile, G.R., Rudd, P.M., Wing, D.R., Prime, S.B., and Dwek, R.A. 1996. A rapid high-resolution high-performance liquid chromatographic method for separating glycan mixtures and analyzing oligosaccharide profiles. Anal. Biochem. 240:210-226.
Guile, R.R., Harvey, D.J., O’Donnell, N., Powell, A.K., Hunter, A.P., Zamze, S., Fernandes, D.L., Dwek, R.A., and Wing, D.R. 1998. Identification of highly fucosylated N-linked oligosaccharides from the human parotid gland. Eur. J. Biochem. 258:623-656. Joao, H.C., Scragg, I.G., and Dwek, R.A. 1992. Effect of glycosylation on protein conformation and amide proton exchange rates in RNase B. FEBS Lett. 307:343-346. Kieffer, B., Driscoll, P.C., Campbell, I.D., Willis, A.C., van der Merwe, P.A., and Davis, S.J. 1994. Three-dimensional solution structure of the extracellular region of the complement regulatory protein CD59, a new cell-surface protein domain related to snake venom neurotoxins. Biochem. 33:4471-4482. Kuhn, P., Tarentino, A.L., Plummer, T.H. Jr., and Van Roey, P. 1994. Crystal structure of peptide-N4(N-acetyl-β-D-glucosaminyl)asparagine amidase F at 2.2-A resolution. Biochemistry 33:11699-11706. Küster, B., Wheeler, S.F., Hunter, A.P., Dwek, R.A., and Harvey, D.J. 1997. Sequencing of N-linked oligosaccharides directly from protein gels: Ingel deglycosylation followed by matrix-assisted laser desorption/ionization mass spectrometry and normal-phase high-performance liquid chromatography. Anal. Biochem. 250:82-101. Kuttner-Kondo, L., Medof, M.E., Brodbeck, W., and Shoham, M. 1996. Molecular modeling and mechanism of action of human decay-accelerating factor. Prot. Eng. 9:1143-1149. Martinez-Pomares, L., Crocker, P., Da Silva, R., Holmes, N., Colominas, C., Rudd, P.M., Dwek, R.A., and Gordon, S. 1999. Cell specific glycoforms of sialoadhesin and CD45 are counter receptors for the cystein-rich domain of the mannose receptor. J. Biol. Chem. 274:35211-35218. Mehta, A., Lu, X., Block, T.M., Blumberg, B.S., and Dwek, R.A. 1997. Hepatitis B virus (HBV) envelope glycoproteins vary drastically in their sensitivity to glycan processing: Evidence that a single N-linked glycosylation site on an HBV envelope protein is crucial for virus secretion—a potential therapeutic target. Proc. Natl. Acad. Sci. U.S.A. 94:1822-1827. Norris, G.E., Stillman, T.J., Anderson, B.F., and Baker, E.N. 1994. The three-dimensional structure of PNGase F, a glycosylasparaginase from Flavobacterium meningosepticum. Structure 2:1049-1059. Petrescu, A.J., Petrescu, S.M., Dwek, R.A., and Wormald, M.R. 1999. A statistical analysis of Nand O-glycan linkage conformations from crystallographic data. Glycobiology 9:343-352. Rudd, P.M. and Dwek, R.A. 1997a. Glycosylation: Heterogeneity and the 3D structure of proteins. Crit. Rev. Biochem. Mol. Biol. 32:1-100. Rudd, P.M. and Dwek, R.A. 1997b. Rapid, sensitive sequencing of oligosaccharides from glycoproteins. Curr. Opin. Biotechnol. 8:488-497.
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Rudd, P.M., Guile, G.R., Küster, B., Harvey, D.J., Opdenakker, G., and Dwek, R.A. 1997a. Oligosaccharide sequencing technology. Nature 388:205-207. Rudd, P.M., Morgan, B.P., Wormald, M.R., Harvey, D.J., van den Berg, C.W., Davis, S.J., Ferguson, M.A.J., and Dwek, R.A. 1997b. The glycosylation of the complement regulatory protein, CD59, derived from human erythrocytes and human platelets J. Biol. Chem. 272:7229-7244. Rudd, P.M., Mattu, T.S., Masure, S., Bratt, T., Van den Steen, P., Wormald, M.R., Küster, B., Harvey, D.J., Borregaard, N., Van Damme, J., Dwek, R.A., and Opdenakker, G. 1999a. Glycosylation of natural human neutrophil gelatinase B and neutrophil gelatinase B-associated lipocalin. Biochemistry 38:13937-13950. Rudd, P.M., Endo, T., Colominas, C., Groth, D., Wheeler, S.F., Harvey, D.J., Wormald, M.R., Serban, H., Prusiner, S.B., Kobata, A., and Dwek, R.A. 1999b. Glycosylation differences between the normal and pathogenic prion protein isoforms. Proc. Natl. Acad. Sci. U.S.A. 96:1304413049. Varki, A. 1993. Biological roles of oligosaccharides: All of the theories are correct. Glycobiology 3:97-130. Williams, R.L., Greene, S.M., and McPherson, A. 1987. The crystal structure of ribonuclease B at 2.5-A resolution. J. Biol. Chem. 262:1602016031. Woods, R.J. 1995. Three dimensional structures of oligosaccharides. Curr. Opin. Struct. Biol. 5:591-598.
Key References Dwek et al., 1993. See above. Although some of the technology in this review has been superseded, the chapter remains essential background reading for an understanding of the new technologies. It describes specific analytical techniques required for oligosaccharide analysis, with emphasis on those required for the elucidation of oligosaccharide primary structure. Guile et al., 1994. See above. This paper describes the methodology for weak anion-exchange chromatography. Guile et al., 1996. See above. This study describes a sensitive technology capable of resolving subpicomolar quantities of mixtures containing both neutral and charged fluorescently labeled oligosaccharides in a single run using normal-phase HPLC. It describes how incremental values for the addition of monosaccharides to oligosaccharide cores have been calculated, enabling the prediction of structures from the elution positions of oligosaccharides.
Rudd et al., 1999b. See above. A paper showing the N-glycan analysis of a glycan pool containing 52 glycans with different compositions. It also shows how to use enzyme digestion profiles to make comparisons of glycosylation.
Internet Resources http://www.cbs.dtu.dk/databases/OGLYCBASE Site for O-GlycBase, a database of O-glycosylated proteins, and for O-Unique, a database of O-glycosylated mucins. ftp://ncbi.nlm.nih.gov/repository/carbbank/ Site for the complex carbohydrate structure database (CCSD16), which can be used to perform searches for carbohydrate structures. Additionally, SUGABASE is a carbohydrate-NMR database that combines CCSD with proton and carbon chemical shift values. http://www.ccrc.uga.edu The site for the Complex Carbohydrate Research Centre Neural Networks (CCRC-Net) contains mass spectra of partially methylated alditol acetate derivatives generated during carbohydrate methylation analysis. http://expasy.cbr.nrc.ca/tools/glycomod/ A tool for predicting oligosaccharide structures on proteins from experimentally determined masses. Can be used for free or derivatized oligosaccharides or glycopeptides. http://www.vei.co.uk/TGN Site for The Glycoscience Network, a site of general interest to researchers in this field. http://www.dkfz-heidelberg.de/spec/sweet2/doc/ index.html A program for constructing three-dimensional models of saccharides from their sequences. http://www.bioch.ox.ac.uk/glycob/ Web site for the Glycobiology Institute. Petrescu et al., 1999. See above. A recently compiled database that allows a more full understanding of the implications of N- and O-glycosylation for the structure and function of a protein, which can only be reached when a glycoprotein is viewed as a single entity.
Contributed by Pauline M. Rudd and Raymond A. Dwek University of Oxford Oxford, United Kingdom
The authors wish to thank Neil Murphy, Cristina Colominas, Edmund Hart, Catherine M. Radcliffe, Louise Royle, Tony Merry, and David J. Harvey for their assistance.
Glycosylation
12.6.47 Current Protocols in Protein Science
Supplement 22
Determining the Structure of Glycan Moieties by Mass Spectrometry
UNIT 12.7
Mass spectrometry provides a complementary method to HPLC (UNIT 12.6) for the analysis of glycans. In general, its resolution, defined as its ability to detect carbohydrates in a complex mixture, is higher than in HPLC, particularly for larger structures. In addition, it provides a mass from which the composition of the glycan, in terms of its isobaric monosaccharide composition, may usually be determined directly. Analysis by mass spectrometry is also faster than by HPLC. On the other hand, mass spectrometry does not usually allow isomeric compounds to be distinguished unless they give diagnostic fragmentation spectra, and, because mass spectrometry is more susceptible to the presence of salts and residual buffers, efficient sample preparation is much more critical and sensitivity generally lower. The high sensitivity and specificity of HPLC is a result of the fluorescent derivatives that are necessary for detection. Although the same technique of derivative preparation can be applied to mass spectrometry in order to improve detection, there does not yet appear to be a single derivative that improves detection in both techniques. Because of the need to analyze minute amounts of sample, there is not usually enough for the preparation of two sets of derivatives, and the first choice for sample analysis is usually the more sensitive technique of HPLC coupled with exoglycosidase digestion. Intact glycans may be detected directly as singly charged ions by matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS; see Basic Protocol 1 and UNIT 16.1; Harvey, 1999) or, following permethylation or peracetylation, by fast atom bombardment mass spectrometry (FAB-MS; see Basic Protocol 2). Electrospray ionization mass spectrometry (ESI-MS; see Basic Protocol 3) also gives good signals for the smaller glycans, but produces singly, doubly, or even triply charged ions, depending on the conditions. This technique is not as good as the other two for glycan profiling; additionally, the sensitivity tends to fall as the glycans become larger. All three techniques give ions in the form of adducts, usually with sodium adducts in the case of MALDI-MS, or as mixtures of sodium and hydrogen in the case of ESI-MS. Adducts can be controlled by addition of organic acids (to promote protonation) or suitable salts to the MALDI or FAB matrices or to the electrospray solvent. For rapid profiling, MALDI-MS is the most appropriate technique. For obtaining structure-revealing fragment ions, FAB-MS or ESI-MS followed by collisional activation gives superior results. CALCULATION OF RESIDUE MASS Mass spectrometry is used mainly to determine the molecular weights and, hence, the composition of the glycans, both neutral or acidic, in a pool, but it will not give linkage information directly. Compositional information, in terms of the isobaric monosaccharide content, is possible because the glycans typically contain only a few mass-different moieties and any combination of these will give a unique mass. It is not, however, possible to differentiate the various isobaric monosaccharides, such as mannose, glucose, and galactose. Masses may be calculated by adding the residue masses of the monosaccharides (Table 12.7.1; either native or derivatized), the mass of the two end groups (Table 12.7.2), the mass of any reducing-terminal derivative (Table 12.7.3), the mass of any additional groups such as sulfate (Table 12.7.4), and the mass of the adduct (Table 12.7.5). Masses calculated in this way can then be matched to the experimentally derived mass, either by computer or by matching with a suitable table of calculated masses. It is important to be aware of the type of mass measurement being used: monoisotopic if the mass spectrometer has resolved the isotopes, or average if no isotopic resolution has been achieved. Both relevant masses are given in the tables. Contributed by David J. Harvey, Raymond A. Dwek, and Pauline M. Rudd Current Protocols in Protein Science (2000) 12.7.1-12.7.15 Copyright © 2000 by John Wiley & Sons, Inc.
Glycosylation
12.7.1 Supplement 22
FRAGMENTATION Fragment ions are often present in FAB spectra but are usually absent from MALDI spectra unless the molecules contain sialic or other relatively unstable residues when a series of focused and unfocused fragment ions can be observed. The unfocused peaks are formed from ions arising from fragmentation while the ions are traveling through the mass spectrometer. These ions can be focused in reflectron-time-of-flight (TOF) instruments to yield postsource decay (PSD) spectra. Such spectra are often of relatively poor quality Table 12.7.1 Residue Masses of Common Monosaccharides and Their Methyl and Acetyl Derivativesa,b
Oligosaccharide
Residue formula
Residue mass
Deoxypentose
C5H8O3
Pentose
Methyl
Acetyl
Residue mass
No. Me
Residue mass
No. Ac
116.047 116.117
130.063 130.144
11
158.058 158.154
1
C5H8O4
132.042 132.116
160.074 160.170
2
216.063 216.191
2
Deoxyhexose
C6H10O4
230.079 230.218 288.084 288.254
2
C6H10O5
174.089 174.197 24.100 204.223
2
Hexose
146.078 146.143 162.053 162.142
Hexosamine
C6H11NO4 161.069 161.157
217.131 217.265
4
287.100 287.269
3
HexNAcc
C8H13NO5 203.079 203.179 C6H8O6 176.032 176.126
245.126 245.276 218.079 218.207
3
287.100 287.269 260.053 260.200
3
KDOc
C8H12O7
220.058 220.179
276.121 276.287
4
346.090 346.291
3
N-Acetylneuraminic acid
C11H17NO8 291.095 291.258 C11H17NO9 307.090 307.257
361.174 361.392 391.184 391.419
5
417.127 417.370 475.133 475.406
Hexuronic acid
N-Glycoylneuraminic acid
3
3
6
3
2
aTop: monoisotopic mass (based on C = 12.000000, H = 1.007825, N = 14.003074, O = 15.994915); bottom: average
mass (based on C = 12.011, H = 1.00794, N = 14.0067, O = 15.9994). bThe masses of the intact glycans can be obtained by addition of the individual residue masses plus the masses for the
relevant terminal groups listed in Table 12.7.2, the mass of the reducing-terminal derivative (if any) in Table 12.7.3, and the mass of any additional groups from Table 12.7.4. For the mass of the molecular ion, also add the mass of the adduct from Table 12.7.5. cAbbreviations: HexNAc, N-acetylhexosamine; KDO, 2-keto-3-deoxyoctulosonic acid.
Table 12.7.2 Masses of Terminal Groups for Calculating Molecular Weights of Carbohydratesa
Reducing end
Determining the Structure of Glycan Moieties by Mass Spectrometry
Underivatized
Methyl
Acetyl
Free
18.011a
46.042
102.090
Reduced
18.153 20.026
46.069 62.073
102.032 146.058
20.031
62.112
146.143
aTop: monoisotopic mass; bottom: average mass.
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from glycans and frequently require sample amounts of 100 pmol or more. In addition, most ions appear in the upper mass range of the spectrum. Much better fragmentation spectra can be obtained using collisional activation. Instruments with a TOF analyzer for separating the fragments, such as the Micromass tandem quadrupole time-of-flight (Q-TOF) instrument, are greatly superior to those with a scanning analyzer, as sensitivity is considerably higher. Newer instruments of the Q-TOF type, incorporating a MALDI ion source rather than an electrospray source, offer improved performance for the analysis of glycans because of the superior ionization characteristics of MALDI sources for the analysis of glycans. Fragmentation of glycans occurs by three processes: elimination of the metal adduct, cleavage between the sugar rings (glycosidic cleavage), and cleavage across the rings. The first process provides no useful structural information. The second Table 12.7.3 Masses of Reducing Terminal Groups for Calculating Molecular Weights of Carbohydrates
Reducing-terminal Formula derivativea
Monoisotopic mass
Average mass
2-AB 2-AA
C7H8N2 C7H7NO
120.069 121.053
120.154 121.139
2-AMAC
C13H10N2
194.084
194.236
aAbbreviations: 2-AA, 2-aminobenzoic acid/anthranilic acid; 2-AB, 2-aminobenzamide;
2-AMAC, 2-aminoacridone.
Table 12.7.4 Masses of Additional Groups Found on Carbohydratesa
Group
Underivatized
Methyl
Acetyl
Methyl
14.015 14.027
0
0
Acetyl
42.011 42.038
0
0
Sulfate
79.957 80.064
65.941b 66.037
37.946b 38.027
Phosphate
79.966 79.980
93.982 94.007
37.956b 37.943
Inositol
162.053 162.142
218.115 218.250
330.095 330.292
aTop: monoisotopic mass; bottom: average mass. bAssuming that no derivatization occurs.
Table 12.7.5 Masses of Common Adducts for Carbohydrates Ionized by MALDI, FAB, or ESI
Adduct
Monoisotopic massa
Average mass
Hydrogen
1.007825
1.00794
Sodium Lithium
22.9898 7.0160
22.9898 6.941
Potassium Cesium
38.9637 132.9054
39.0983 132.9054
aMost abundant isotope.
Glycosylation
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Y2
Z2
Y1
Z1
Y0
CH2OH
CH2OH
CH2OH
O
O
O
OH
O
OH
O
O
Z0
R
OH
HO OH
2,5A 1
B1
OH
C1
2,4A 2
B2
OH
C2
O,2A 3
B3
C3
Figure 12.7.1 Fragmentation nomenclature, after Domon and Costello (1988).
gives information on composition, sequence, and sometimes branching. Cross-ring cleavages provide linkage information. Glycosidic cleavages are the most common, but if several occur during the formation of a given ion, the results can be ambiguous as the resulting “internal cleavage” ion may be formed by several competing processes. Crossring cleavage ions are usually absent from the spectra of protonated ions, but appear, usually as minor peaks, in the spectra of sugars ionized as sodium or lithium adducts. Abundant cross-ring cleavage ions can be produced in instruments capable of producing high-energy collisions. The accepted nomenclature for fragment ions is taken from that proposed by Domon and Costello (1988) and is summarized in Figure 12.7.1. Ions retaining the charge at the reducing terminus are labeled A (cross-ring), B (cleavage of the C1-to-O bond), or C (cleavage of the other glycosidic bond), whereas ions charged at the nonreducing terminus are labeled X (cross-ring), Y, and Z. Subscript numbers denote the bond broken, starting at the reducing terminus for the X, Y, and Z ions, and at the nonreducing terminus for the others. Cross-ring ions are given an additional superscript number to denote the bonds broken (e.g., 2,4A2 representing cleavage of the 2,3 and 4,5 bonds of the second monosaccharide from the nonreducing terminus). SAMPLE PURITY Purity requirements for mass spectrometry are very different from those for HPLC (UNIT 12.6). Many contaminants, such as buffers and detergents, can cause problems by, for example, preventing efficient crystallization. If the sample is concentrated, then it may be possible to dilute it to the point where the contaminants have little effect. However, as a rule, the purer the sample, the cleaner, sharper, and more intense the spectrum.
Determining the Structure of Glycan Moieties by Mass Spectrometry
The Support Protocols describe different methods of sample purification, depending on the starting material. Support Protocol 1 presents the removal of contaminants by microdialysis or membrane absorption techniques. Support Protocol 2 describes a technique for removing contaminants with a graphitized carbon column. Clean-up of 2-AB derivitives of neutral glycans is described in Support Protocol 3. SDS gel purification of neutral or sialylated glycans is presented in Support Protocols 4 and 5, respectively.
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Table 12.7.6 Upper Tolerated Limits of Buffers and Detergents for MALDI-MS Analysis
Reagent
Upper limit
Phosphate
20 mM
Tris Ammonium bicarbonate Alkali metal salts
50 mM 30 mM (readily removed by lyophilization) 1M
Guanidine Glycerol
1M 1% (v/v)
SDS Other detergents
0.01% (w/v) 0.1% (w/v)
When considering which purification to perform, it should be noted that levels of buffer and detergent that exceed the limits in Table 12.7.6 will probably cause noticeable degradation of MALDI spectra (Mock et al., 1992). ANALYSIS OF GLYCANS BY MATRIX-ASSISTED LASER DESORPTION/IONIZATION MASS SPECTROMETRY
BASIC PROTOCOL 1
Most laboratories have a favorite method for preparation of MALDI targets; this protocol represents the technique that this laboratory has found to be reliable. Best results are obtained by loading a minimum of 1 to 10 pmol sugar onto the target for recording molecular ions only, and upwards of 100 pmol of each glycan to obtain PSD spectra. In practice, this requires the sample concentration to be in the range of 2 × 10−6 M to 1 × 10−4 M. In cases where the sample concentration is unknown, it may be necessary to use trial and error. If the sample concentration is not known, there is a good chance that unknown contaminants may also be present. If success simply cannot be achieved, try spiking the solution with something that is known to run well, such as insulin. If the spectrum is still a blank, this is a strong indication that the sample solution contains a problem contaminant. Materials Glycan sample, purified (see Support Protocols 1 to 5 and UNIT 12.6) Matrix solution (see recipe) Mass spectrometer (e.g., Micromass TofSpec2, PerSeptive Biosystems Voyager series TOF mass spectrometer, Bruker Reflex MALDI mass spectrometer, or Kratos Kompact) 1. Deposit 0.5 µl glycan sample to the target. 2. Using a separate tip, add 0.5 µl matrix solution and allow the mixture to dry at room temperature. Inspect the dried sample deposit. The dried sample should look fairly uniform across the area of the spot. If the deposit is sticky, then either the sample is much too concentrated or it contains a high concentration of detergent or glycerol. Such deposits are unlikely to yield good spectra and the sample should be purified further. Some investigators prefer to premix the matrix and sample solutions prior to application to the MALDI target. In the authors’ hands, this method does not appear to offer any advantages and may result in loss of sample. Glycosylation
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3. If good crystals are present, redissolve the spot in the minimum amount of ethanol and allow it to recrystallize. Only ∼0.1 ìl ethanol is necessary, and care must be taken to prevent larger amounts of ethanol from spreading the sample over a larger area.
4. Introduce the target into the mass spectrometer and operate in positive ion mode. The mass spectrometer should be calibrated. Suitable calibration compounds are a mixture of dextran oligomers (e.g., Fluka dextran from Leuconostoc ssp.) for positive ion mode, and the same compounds derivatized with 2-aminobenzoic acid for calibrating negative ion spectra. Masses of the calibration peaks are given in Table 12.7.7 (dextran for positive ion) and Table 12.7.8 (2-AA derivative of dextran for negative ion).
5. Acquire the spectrum and analyze results. With linear TOF instruments without delayed extraction (time-lag focusing) ion sources (e.g., ThermoQuest LaserMat), it is essential to keep the laser power as low as practical in order to avoid peak broadening and consequent mass errors. This effect is less pronounced with delayed-extraction instruments, but increased laser power can still lead to reduced resolution. The laser spot should be moved to different areas of the target during acquisition, as not all areas of the target will yield a signal. It is frequently found with 2,5-hydroxybenzoic Table 12.7.7 Masses of MNa+ Ions from Dextran For Calibrating the MALDI-MS in Positive Ion Mode
Monoisotopic
Average
Monoisotopic
Average
Monoisotopic
Average
203.05
203.15
1823.58
1824.57
3444.11
3446.00
365.11 527.16
365.29 527.43
1985.63 2147.69
1986.71 2148.86
3606.16 3768.22
3608.14 3770.28
689.21 851.26
689.57 851.72
2309.74 2471.79
2311.00 2473.14
3930.27 4092.32
3932.42 4094.57
1013.32 1175.37
1013.86 1176.00
2633.85 2795.90
2635.28 2797.43
4254.37 4416.43
4256.71 4418.85
1337.42 1499.48
1338.14 1500.29
2957.95 3120.00
2959.57 3121.71
4578.48 4740.53
4580.99 4743.13
1661.53
1662.43
3282.06
3283.85
4902.59
4905.28
Table 12.7.8 Masses of [M–H]− Ions from the 2-AA Derivative of Dextran Sugars For Calibrating the MALDI-MS in Negative Ion Mode
Determining the Structure of Glycan Moieties by Mass Spectrometry
Monoisotopic
Average
Monoisotopic
Average
Monoisotopic
Average
300.11 462.16
300.29 462.43
1758.58 1920.64
1759.57 1921.71
3541.17 3703.22
3543.14 3705.28
624.21 786.27
624.57 786.72
2082.69 2244.74
2083.86 2246.00
3865.27 4027.32
3867.42 4029.56
948.32 1110.37
948.86 1111.00
2406.80 2568.85
2408.14 2570.28
4189.38 4351.43
4191.71 4353.85
1272.43 1434.48
273.14 1435.29
2730.90 2892.95
2732.42 2894.57
4513.48 4675.53
4515.99 4678.13
1596.53 300.11
1597.43 300.29
3055.01 3217.06
3056.71 3218.85
4837.59 3541.17
4840.28 3543.14
12.7.6 Supplement 22
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2151.8
100
1745.6
1339.5
1257.4
1501.5
Relative abundance (%)
1419.5
1663.6 1542.6 1704.6
1136.4
2313.9
1907.7
1866.7
1948.7
2110.8 1298.5
50
2028.7
1460.5 1622.6
2476.5 2272.8
2069.7
0 1100
1300
1500
1700
m/z
1900
2100
2300
2500
Figure 12.7.2 An example of a MALDI spectrum.
acid that an active spot will only give a signal for a few laser shots due either to depletion of the sample/matrix mixture or vertical inhomogeneities within the crystal. The laser spot should, therefore, be moved continuously and spectra averaged until a satisfactory signalto-noise ratio is obtained (Figure 12.7.2). For PSD spectra, the laser spot should be moved slowly over the surface throughout the acquisition, as it is sometimes difficult to see weak ions until the final spectrum is assembled (Figure 12.7.3).
Relative abundance (%)
Because sialylated glycans frequently give abundant peaks due to loss of sialic acid when their spectra are recorded with reflectron-TOF instruments, their spectra are often recorded
Y5 Y4 Y 3
Y2 Y1 MNa+ 1783.5
2-AB
100 B1 B2 B3
B4 B5
B2 Y1 388.1 364.1
50
200
400
B5 Y4 1077.3
B4 Y3 712.2
B4 Y4 874.2 Y4 Y4 1053.3 915.2 948.3 753.2
B3 550.1 509.1
226.1 346.1
0
Y2 567.2
600
800
1000 m/z
Figure 12.7.3 An example of a MALDI-PSD spectrum.
Y4 1418.4 B5 1442.3 B4 1239.3 Y3 1256.4 1400.4
1200
1400
Y5 1621.4
1600
1800
Glycosylation
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in linear mode in order to minimize sialic acid loss. However, it must be borne in mind that the resulting mass measurement may well be an average rather than a monoisotopic mass because of the lower resolution.
6. If desired, recover the remaining sample from the MALDI target after MS analysis by dissolving the spot in several repetitive additions of 2 µl of 50% acetonitrile in water and removal by pipet. The matrix can then removed from the resultant solution by drop dialysis or use of a C18 ZipTip (see Support Protocol 3). BASIC PROTOCOL 2
ANALYSIS OF GLYCANS BY FAST ATOM BOMBARDMENT MASS SPECTROMETRY FAB is less sensitive than MALDI, and samples must be permethylated or peracetylated prior to analysis. Permethylation requires either sodium hydride or, more conveniently, sodium hydroxide catalysis (Ciucanu and Kerek, 1984). Materials Sodium hydroxide (NaOH) pellets Dry dimethyl sulfoxide (DMSO) Purified glycan sample (see Support Protocols 1 to 5 and UNIT 12.6) Methyl iodide Chloroform Source of dry nitrogen 50% (v/v) methanol/water 75% (v/v) acetonitrile/water 1:1 (v/v) glycerol/thioglycerol Mass spectrometer (e.g., Micromass AutoSpec or Finnigan ThermoQuest 900) Additional reagents and equipment for cleaning sample (see Support Protocols 1, 3, and 4) Permethylate glycan samples 1. Crush one pellet of NaOH in a dry atmosphere and make into a slurry with dry DMSO. 2. Dissolve dried glycan sample in 25 µl DMSO/NaOH slurry, mix well, and allow to stand 1 hr at room temperature. 3. Add 25 µl methyl iodide and leave at room temperature for another hour. CAUTION: This procedure must be performed in a fume hood, as methyl iodide is toxic.
4. Cool in ice and slowly add 50 µl cold water. It is important to minimize contact between the permethylated glycans and aqueous base to avoid decomposition and hydrolysis of methyl ester groups (from sialic acids).
5. Extract the permethylated glycans with 500 µl chloroform and wash with 1-ml portions of water until the chloroform (bottom layer) is no longer cloudy (typically three washes). Blow to dryness with a nitrogen stream to ensure that all of the methyl iodide has been removed. 6. Dissolve products in 20 to 30 µl of 50% methanol/water and clean (see Support Protocol 1 or 4). Determining the Structure of Glycan Moieties by Mass Spectrometry
If the sample is labeled at the reducing terminus, it can also be cleaned with a C18 ZipTip or Sep-Pak cartridge (see Support Protocol 3), but using 75% acetonitrile/water to elute the glycans. Some investigators prefer to elute with various concentrations (25%, 50%, 75%, and 100%) of acetonitrile in order to ensure complete recovery of glycans.
7. Remove acetonitrile with a stream of nitrogen and resuspend in methanol.
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Record FAB spectrum 8. Deposit a small drop of 1:1 glycerol/thioglycerol to the FAB probe. 9. Add ≥100 pmol permethylated glycans in methanol and mix well. 10. Insert the probe into the mass spectrometer and analyze. The beam should last for several minutes without the need for any movement of the probe.
ANALYSIS OF GLYCANS BY ELECTROSPRAY IONIZATION MASS SPECTROMETRY
BASIC PROTOCOL 3
This technique is currently not as satisfactory for glycan profiling as MALDI, but provides a steady signal for the acquisition of fragmentation spectra. Satisfactory spectra from most glycans can be obtained from mixtures of water and either methanol or acetonitrile. Formation of hydrogen, sodium, or other adducts can be induced by the addition of formic or acetic acid (0.1% to 1.0%) for protonation, or an alkali metal salt for formation of metal adducts. Furthermore, ESI-MS can be conveniently interfaced with HPLC. The glycan solution should be as free from other compounds as possible, as these compounds may compete for the ionization and reduce the signal strength. Electrospray is not as sensitive for detection of carbohydrates as it is for peptides, and concentrations of 1 to 10 pmol/µl solvent are typically needed. However, certain reducing-terminal derivatives, such as that formed from the N,N-diethylaminoethyl ester of 4-aminobenzoic acid (ABDEAE derivatives; Mo et al., 1999) provide increases in sensitivity on the order of 100- to 1000-fold over that of the underivatized glycan. Such derivatives may be prepared in the same way as the 2-aminobenzamide derivatives with the appropriate amine substituted for 2-AB. Glycans derivatized with 2-AB or 2-AMAC (Charlwood et al., 1999) may be cleaned (see Support Protocol 3) and then dissolved in 1:1 (v/v) methanol/water or 1:1 (v/v) acetonitrile/water containing 1 to 50 mM sodium iodide. Sodium iodide is used to produce [M + metal]+ ions; 0.1% formic acid is used to produce [M + H]+ ions. This solution is then infused into the mass spectrometer at a rate determined by the inlet system (0.05 to 10 µl/min). A Micromass Q-TOF or LCT, or a ThermoQuest TSQ7000 or LCQ, is acceptable. The cone voltage should be set to a high value (∼180 V on the Micromass Q-TOF instrument) to maximize the intensity of the MNa+ ion. Unfortunately, under these conditions, some cone-voltage fragmentation will occur and may distort the glycan profile. However, these conditions will be optimal for subsequently recording the collision-induced fragmentation spectrum. If samples do not contain added salt, and spectra are recorded at lower cone voltages (50 to 80 V), abundant doubly charged ions will be present. These ions are a mixture of species such as MH22+, MHNa2+, MHK2+, or MNa22+. Their relative percentages depend on the composition of the solution and the derivative used. To maximize the percentage of the MHNa2+ ion in, for example, the spectra of the ABDEAE derivatives where they form the most abundant species, 0.1% formic acid and 10 nmol/µl sodium iodide should be added to the solution. Fragment ions from MH+ and MHNa2+ molecular ions generally consist of glycosidic fragments only, whereas those from MNa+ and MNa22+ ions are more complex and additionally contain cross-ring fragments (Fig. 12.6.3). Spectra from the MNa+ ions are generally superior and do not contain mixtures of singly and doubly charged ions. The extent of fragmentation will depend on the collision cell voltage, with higher voltages being required for the larger molecules and for singly charged metal-containing ions.
Glycosylation
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SUPPORT PROTOCOL 1
REMOVAL OF CONTAMINANTS BY MICRODIALYSIS OR ADSORPTION ONTO MEMBRANES Salts and small molecules can be removed by microdialysis using membranes with a 500 Da MWCO for N-linked glycans or 100 Da for smaller sugars. Peptides can be removed with a Nafion membrane (Börnsen et al., 1995). Materials Sample prepared by PNGase F digestion or hydrazinolysis (UNIT 12.6) Membrane (select one): Dialysis membrane (e.g., Spectra/Pore CE membrane, MWCO 500) Nafion 117 membrane (Aldrich) 1. Cut a small square of a relevant membrane ∼1 cm on each side and float this on the surface of water contained in a small container (e.g., a petri dish). If dialysis tubing has been used as the source of the membrane, place the outside surface down.
2. Add one or more 1-µl drops of an aqueous solution of the glycans to the upper surface of the membrane, cover the apparatus to prevent evaporation, and leave 10 to 15 min at room temperature. 3. Remove the glycan solution and apply directly to a MALDI target (see Basic Protocol 1). SUPPORT PROTOCOL 2
PURIFICATION OF N-LINKED OLIGOSACCHARIDES ON A POROUS GRAPHITIZED CARBON COLUMN Removal of detergents and anionic buffers is problematical for the isolation and cleanup of N-linked sialylated oligosaccharides for mass spectrometry. Neutral and acidic glycans can be separated from buffer salts, protein, and detergents on porous graphitized carbon columns (Packer et al., 1998). The oligosaccharides are fractionated depending on their anionic charge by increasing concentrations of acetonitrile and trifluoroacetic acid. Materials PNGase F digestion sample (UNIT 12.6) 1 M NaOH 25% (v/v) aqueous acetonitrile 25% (v/v) aqueous acetonitrile/0.05% (v/v) trifluoroacetic acid 80% (v/v) aqueous acetonitrile/ 0.10% (v/v) trifluoroacetic acid 100-mg prepacked Hypersep porous graphitized carbon (PGC) columns (ThermoQuest) 1. Prewash PGC column three times with 2 ml of each of the following solutions: 1 M NaOH Distilled water 80% aqueous acetonitrile/0.05% trifluroacetic acid 25% aqueous acetonitrile/0.05% trifluroacetic acid 25% aqueous acetonitrile Distilled water.
Determining the Structure of Glycan Moieties by Mass Spectrometry
NOTE: PGC columns require extensive washing before use and high water purity is essential.
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2. Load a PNGase F digest onto the column and wash 5 times with 300 µl water to remove all buffer salts, neutral monosaccharides, and detergents such as Triton X-100. If SDS is used for PNGase F digestion, the addition of an excess of Nonidet P-40 or a similar nonionic detergent is required prior to loading onto PGC columns.
3. Elute neutral oligosaccharides from the column with three 300-µl additions of 25% aqueous acetonitrile. 4. Recover anionic glycans with three 300-µl additions of 25% acetonitrile/0.05% trifluroacetic acid. All fetuin oligosaccharides are eluted at this concentration; however, for highly negatively charged glycans, concentrations up to 40% acetonitrile/0.05% TFA can be used. This method is not as effective for the cleanup of polysialylated glycans such as colominic acid.
5. Freeze-dry all fractions and reconstitute in water prior to mass spectrometry analysis. Although samples are best analyzed immediately, they can be kept for up to several weeks at −20°C. All fractions must be frozen and freeze-dried as quickly as possible after collection to avoid acid hydrolysis of sialic acid groups.
PURIFICATION OF 2-AB DERIVATIVES OF NEUTRAL N-LINKED GLYCANS USING A C18 ZIPTIP In this procedure, the hydrophobic nature of 2-AB or other reducing-terminal derivative allows the compound to be adsorbed onto C18 and the contaminants to be removed by washing.
SUPPORT PROTOCOL 3
Materials Sample prepared by PNGase F digestion or hydrazinolysis and derivatized by reductive amination (UNIT 12.6) C18 ZipTips (Millipore) 1:1 (v/v) acetonitrile/water 1. Attach a C18 ZipTip to a micropipettor and condition twice by aspirating 20 µl of 1:1 acetonitrile/water. 2. Wash tip twice with 20 µl water. 3. Draw up ∼20 µl of aqueous glycan solution and return to the main solution. Repeat approximately ten times. This procedure should saturate the C18 with the derivatized glycan if concentrated solutions are used.
4. Wash tip twice with 20 µl water. 5. Elute glycans with ∼10 µl of 1:1 acetonitrile/water. The solution can be used directly for MALDI- or ESI-MS.
PURIFICATION OF NEUTRAL GLYCANS RELEASED FROM SDS GELS Glycan solutions recovered from SDS-PAGE gels (Küster et al., 1997), as described in UNIT 12.6, are inevitably contaminated with residual salts and organic material such as SDS. These compounds should be removed prior to MALDI mass spectrometry in order to obtain a strong signal. If the samples are to be examined by both HPLC, after derivatization, and by mass spectrometry, it is better to perform the MALDI analysis on underivatized sample. Although MALDI produces satisfactory spectra from most derivatives, the derivatization process invariably introduces additional contaminants whose removal
SUPPORT PROTOCOL 4
Glycosylation
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Supplement 22
incurs additional sample loss. Also, prompt analysis of the glycans usually produces stronger MALDI signals than those obtained from stored samples. Materials Sample prepared by in-gel PNGase F release (UNIT 12.6) Conditioned AG50X12 resin (see recipe) Conditioned AG3X4 resin (see recipe) C18 resin: obtain from a SepPak cartridge and make a slurry in ethanol Pressurized nitrogen GelLoader pipet tip (Eppendorf) SpeedVac evaporator 1. Reconstitute sample in ∼40 µl distilled water. 2. Fill a GelLoader pipet tip with ethanol, then add: ∼10 µl conditioned AG3X4 resin ∼25 µl conditioned AG50X12 resin ∼10 µl C18 resin. 3. Using a nitrogen line to pressurize liquid through the column, wash the column three times with 50-µl aliquots of distilled water. Ensure that the resin stays wet throughout. The nitrogen is used to apply a slight pressure to the column in order to speed up the passage of liquid. The pressure should be adjusted so that the flow is approximately one drop every three to four seconds.
4. Apply sample to the top of the column and use nitrogen to push all of the solution onto the column. Immediately begin to collect eluent. As the glycans are not adsorbed onto the column, collection of the eluent should be started as soon as the glycans are applied in order to avoid losses.
5. Rinse the column four times with 50-µl aliquots of distilled water and collect the eluant. At the end of the final rinse, blow the column dry to ensure that all of the sample has been eluted. 6. Lyophilize the eluant in a SpeedVac evaporator and reconstitute in 1 to 5 µl distilled water (depending on sample quantity) for MALDI analysis. SUPPORT PROTOCOL 5
Determining the Structure of Glycan Moieties by Mass Spectrometry
PURIFICATION OF SIALYLATED GLYCANS RELEASED FROM SDS GELS This procedure is similar to that described in Support Protocol 4. It has been found that the use of the AG3X4 resin is essential in order to obtain MALDI signals from the in-gel released glycans. However, its presence in the above microcolumn removes acidic glycans. This loss can be avoided if the sialic acids are converted into their methyl esters prior to use of the above procedure (Powell and Harvey, 1996). Methyl ester formation is also advantageous for MALDI-MS because it increases the signal strength of the positive ions from sialylated glycans by preventing negative ion formation. Furthermore, it prevents metastable loss of sialic acid and inhibits salt formation. Consequently, single positive ion peaks are produced, comparable to those from the neutral glycans. Additional Materials (also see Support Protocol 4) Sample of sialyted glycans prepared by in-gel PNGase F release (UNIT 12.6) AG50X12 resin (wash 100 µl resin with 1 ml of 1 M NaOH solution followed by an equivalent amount of water) Anhydrous dimethyl sulfoxide (DMSO), stored over a molecular sieve Methyl iodide 20% (v/v) acetonitrile in water GelLoader pipet tips (Eppendorf)
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For Standard Glycan Samples 1a. Pass 5 to 20 µl of aqueous glycans through a short column of AG50X12 (Na+) packed into a GelLoader pipet tip to a bed height of ∼5 mm. A small plug of C18 will prevent the resin from being washed out of the tip.
2a. Lyophilize or freeze dry in a SpeedVac evaporator. Do not leave the sample in the SpeedVac for too long in order to avoid loss of sialic acids.
3a. Add 1 µl anhydrous DMSO and 1 µl methyl iodide. Vortex and microcentrifuge briefly. 4a. Allow to stand 2 hr at room temperature in a fume hood. 5a. Add 5 µl anhydrous DMSO, vortex, microcentrifuge, and blow off excess methyl iodide using a nitrogen line. 6a. Lyophilize sample in a SpeedVac evaporator and reconstitute in ∼40 µl distilled water. 7a. Repeat step 1 but remove the C18 plug from the column since its presence will retain the methylated sugars. For Glycans Obtained as Standard Compounds or as HPLC Fractions 1b. Prepare a C18 microcolumn containing ∼20 to 30 µl resin and rinse thoroughly with distilled water. 2b. Load the sample onto the column and rinse through five times with 50 µl distilled water. 3b. Elute glycans with four 50-µl additions of 20% acetonitrile. 4b. Lyophilize the eluent and reconstitute in 1 µl distilled water. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Conditioned AG50X12 resin Wash ∼1 ml AG50X12 resin (H+ form; Bio-Rad) with 10 ml of 10% HCl. Rinse with ≥10 ml distilled water until neutralized. Store up to several weeks at 4°C. Take resin from the settled layer at the bottom of the container. Conditioned AG3X4 resin Wash ∼1 ml AG3X4 resin (OH− form; Bio-Rad) with 10 ml of 1 M NaOH. Rinse with ≥10 ml distilled water until neutralized. Store up to several weeks at 4°C. Matrix solution Prepare a saturated solution of 2,5-dihydroxybenzoic acid (2,5-DHB) in acetonitrile. Store up to several weeks at room temperature. Commercial DHB (Aldrich) appears sufficiently pure for direct use. Sufficient sodium is usually present that it is not necessary to add extra. Solutions containing additives appear less stable and should be prepared more frequently. For PSD work, it is recommended that only fresh solutions be used.
COMMENTARY Background Information MALDI mass spectrometry usually produces one positive ([M + adduct]+) singly
charged ion from neutral glycans, but a mixture of positively and negatively charged ([M–H]−) ions from acidic sugars. In addition, the latter Glycosylation
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Determining the Structure of Glycan Moieties by Mass Spectrometry
compounds can also give fragment ions and form additional peaks as the result of salt formation. Signals are thus usually weak and complex. The matrix most commonly used for MALDI analysis of glycans is 2,5-dihydroxybenzoic acid (2,5-DHB; molecular mass 154). Unfortunately, DHB forms large needleshaped crystals that usually start growing at the periphery of the target and project towards the center, often leaving a considerable area covered with an amorphous material containing some of the glycans. Several procedures can be used to overcome this problem. One is to add ∼10% of 5-methoxy2-hydroxybenzoic acid (Karas et al., 1993), 1hydroxyisoquinoline (Mohr et al., 1995), or 2,4,6-trihydroxyacetophenone to the matrix solution in order to catalyze the formation of small even crystals. The second is to recrystallize the dry target spot from ethanol (Harvey, 1993). Addition of 5-methoxy-2-hydroxybenzoic acid produces a matrix commonly referred to as “super DHB.” Spermine has been added to DHB for examination of sialylated glycans in order to reduce salt formation (Mechref and Novotny, 1998). Other useful matrices are 3-aminoquinoline (Metzger et al., 1994), 2-(4-hydroxyphenylazo)benzoic acid (HABA), and arabinosazone (Chen et al., 1997), the latter being particularly useful for recording spectra of acidic glycans in the negative mode. Another matrix that has been reported to be suitable for these glycans is a mixture of 2,4,6-trihydroxyacetophenone (THAP) and ammonium citrate (1 mM THAP in a 1:1 mixture of acetonitrile and 20 mM ammonium citrate; Pitt and Gormon, 1996).
Time Considerations
Critical Parameters and Troubleshooting
Charlwood, J., Langridge, J., Tolson, D., Birrell, H., and Camilleri, P. 1999. Profiling of 2-aminoacridone derivatised glycans by electrospray ionization mass spectrometry. Rapid Commun. Mass Spectrom. 13:107-112.
It is essential to use the highest quality solvents for every purpose. In particular, purified water sometimes contains polymeric materials (principally polyethylene glycol) that will coconcentrate with the sugars and impair MALDIMS detection. To avoid this problem, purified water should be further treated—e.g., by distilling at sub–boiling point temperatures (SBPD) to prevent organic contaminants from being carried across with the steam. All glassware should be washed with 4 M nitric acid for 24 hr and rinsed with ten changes of purified water and finally with SBPD water. Acetonitrile is a source of artefacts in mass spectrometers. A suitable source of acetonitrile for this purpose can be obtained from Allied Signal Riedel-de Haen (available via Sigma).
The procedures for the preparation and purification of the glycan samples do not require much time. Microdialysis or membrane adsorption (Support Protocol 1) requires <1 hr. Graphitized carbon column cleanup (Support Protocol 2) requires 2 to 3 hr. Neutral glycan purification procedures (Support Protocols 3 and 4 ) take 1 hr. Purification of sialylated glycans (Support Protocol 5) can take up to 1 day. The preparation of samples for MALDI analysis (Basic Protocol 1) takes 20 min; the actual running of the sample and analysis of results requires less than a minute if the sample gives a strong signal. For problem samples that give very weak signals, spectra may be accumulated for as long as 5 min in order to achieve a reasonable signal:noise ratio. Calibration of the instrument takes ∼5 min, but this need only be done once or twice for a batch of samples. FAB spectrometry (Basic Protocol 2) can take 3 to 4 hr to complete the sample preparation and running and analysis of the sample. Electrospray samples run by direct infusion (Basic Protocol 3) take only ∼1 min, but several minutes are needed to wash the injector system between samples and allow the fresh sample to infuse into the ion source. This time will depend on the infusion rate and the volume of the injection system.
Literature Cited Börnsen, K.O., Mohr, M.D., and Widmer, H.M. 1995. Ion exchange and purification of carbohydrates on a Nafion membrane as a new sample pretreatment for matrix-assisted laser desorptionionization mass spectrometry. Rapid Commun. Mass Spectrom. 9:1031-1034.
Chen, P., Baker, A.G., and Novotny, M.V. 1997. The use of osazones as matrices for the matrix-assisted laser desorption/ionization mass spectrometry of carbohydrates. Anal. Biochem. 244:144-151. Ciucanu, I. and Kerek, F. 1984. A simple and rapid method for the permethylation of carbohydrates. Carbohydrate Res. 131:209-217. Domon, B. and Costello, C.E. 1988. A systematic nomenclature for carbohydrate fragmentations in FAB-MS/MS spectra of glycoconjugates. Glycoconjugate J. 5:397-409. Harvey, D.J. 1993. Quantitative aspects of the matrixassisted laser desorption mass spectrometry of complex oligosaccharides. Rapid Commun. Mass Spectrom. 7:614-619.
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Harvey, D.J. 1999. Matrix-assisted laser desorption/ionization mass spectrometry of carbohydrates. Mass Spectrom. Rev. 18:349-451. Karas, M., Ehring, H., Nordhoff, E., Stahl, B., Strupat, K., Hillenkamp, F., Grehl, M., and Krebs, B. 1993. Matrix-assisted laser desorption/ionization mass spectrometry with additives to 2,5-dihydrobenzoic acid. Org. Mass Spectrom. 28:14761481. Küster, B., Wheeler, S.F., Hunter, A.P., Dwek, R.A., and Harvey, D.J. 1997. Sequencing of N-linked oligosaccharides directly from protein gels: Ingel deglycosylation followed by matrix-assisted laser desorption/ionization mass spectrometry and normal-phase high-performance liquid chromatography. Anal. Biochem. 250:82-101. Mechref, Y. and Novotny, M.V. 1998. Matrix-assisted laser desorption/ionization mass spectrometry of acidic glycoconjugates facilitated by the use of spermine as a co-matrix. J. Am. Soc. Mass Spectrom. 9:1292-1302. Metzger, J.O., Woisch, R., Tuszynski W., and Angermann, R. 1994. New type of matrix for matrix-assisted laser desorption mass spectrometry of polysaccharides and proteins. Fresenius J. Anal. Chem. 349:473-474. Mo, W., Sakamoto, H., Nishikawa, A., Kagi, N., Langridge, J.I., Shimonishi, Y., and Takao, T. 1999. Structural characterisation of chemically derivatised oligosaccharides by nanoflow electrospray ionization mass spectrometry. Anal. Chem. 71:4100-4106. Mock, K.K., Sutton, C.W., and Cottrell, J.S. 1992. Sample immobilization protocols for matrix-assisted laser-desorption mass spectrometry. Rapid Commun. Mass Spectrom. 6:2333-238. Mohr, M.D., Börnsen, K.O., and Widmer, H.M. 1995. Matrix-assisted laser desorption/ionization mass spectrometry: Improved matrix for oligosaccharides. Rapid Commun. Mass Spectrom. 9:809-814. Packer, N.H., Lawson, M.A., Jardine, D.R., and Redmond, J.W. 1998. A general approach to desalting oligosaccharides released from glycoproteins. Glycoconjugate J. 15:737-747. Pitt, J.J. and Gormon, J.J. 1996. Matrix-assisted laser desorption/ionization time-of-flight mass spectrometry of sialylated glycopeptides and proteins using 2,6-dihydroxyacetophenone as a matrix. Rapid Commun. Mass Spectrom. 10:1786-1788. Powell, A.K. and Harvey, D.J. 1996. Stabilisation of sialic acids in N-linked oligosaccharides and gangliosides for analysis by positive ion matrix-assisted laser desorption-ionization mass spectrometry. Rapid Commun. Mass Spectrom. 10:1027-1032.
Key References Dwek, R.A., Edge, C.J., Harvey, D.J., Wormald, M.W., and Parekh, R.B. 1993. Analysis of glycoprotein associated oligosaccharides. Annu. Rev. Biochem. 62:65-100. Although some of the technology in this review has been superseded, the chapter remains essential background reading for an understanding of the new technologies. It describes specific analytical techniques required for oligosaccharide analysis, with emphasis on those required for the elucidation of oligosaccharide primary structure. Küster et al., 1997. See above. This paper describes a sensitive technique for releasing N-linked glycans from within SDS-PAGE gels and for their analysis by MALDI mass spectrometry. Harvey, D.J. 1999. Matrix-assisted laser desorption/ionization mass spectrometry of carbohydrates. Mass Spectrom. Rev. 18:349-451. This is a comprehensive review of MALDI mass spectrometry as applied to carbohydrate analysis covering instrumentation, techniques, sample preparation, and applications from the literature to the end of 1998.
Internet Resources http://www.cbs.dtu.dk/services/NetOGlyc/ Site for O-GlycBase, a database of O-glycosylated proteins. CCSD16; ftp://ncbi.nlm.nih.gov/repository/ carbbank/ Site for the complex carbohydrate structure database (CCSD16), which can be used to perform searches for carbohydrate structures. Additionally, SUGABASE is a carbohydrate-NMR database that combines CCSD with proton and carbon chemical shift values. CCRC-Net; http://www.ccrc.uga.edu The site for the Complex Carbohydrate Research Centre Neural Networks (CCRC-Net) contains mass spectra of partially methylated alditol acetate derivatives generated during carbohydrate methylation analysis. http://www.bioch.ox.ac.uk/glycob/ Web site for the Glycobiology Institute.
Contributed by David J. Harvey, Raymond A. Dwek, and Pauline M. Rudd University of Oxford Oxford, United Kingdom
The authors wish to thank Neil Murphy, Louise Royle, and Susan F. Wheeler for their assistance. Glycosylation
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Detection and Analysis of Proteins Modified by O-Linked N-Acetylglucosamine
UNIT 12.8
The modification of Ser and Thr residues with O-linked β-N-acetylglucosamine (OGlcNAc) is a common, dynamic, and essential modification of nuclear and cytoplasmic proteins. O-GlcNAc is ubiquitous, having been identified in all complex eukaryotes studied, including filamentous fungi and plants. O-GlcNAc has been found on a diverse range of proteins including cytoskeletal proteins, nuclear pore proteins, RNA polymerase II (RNA Pol II), transcription factors, proto-oncogene products, tumor suppressors, hormone receptors, phosphatases, and kinases (Comer and Hart, 2000). The functional consequences of modifying proteins with O-GlcNAc is unclear, but it is required for survival at the single-cell level (Shafi et al., 2000). Three features suggest that O-GlcNAc performs a regulatory role: (1) O-GlcNAc occurs at sites on the protein backbone that are similar to those modified by protein kinases; (2) O-GlcNAc is reciprocal with phosphorylation on some well-studied proteins, such as RNA Pol II, estrogen receptor-β, SV40 large T-antigen, and the c-Myc proto-oncogene product; and (3) like phosphorylation, O-GlcNAc is highly dynamic, with rapid cycling in response to cellular signals or cellular stages. Perturbations in the metabolism of GlcNAc, which alter the regulation of many O-GlcNAc proteins, have been implicated in Alzheimer’s disease, diabetes, and cancer (Wells et al., 2001). This unit concentrates on techniques for the detection and analysis of proteins modified by O-GlcNAc, as well as methods for the analysis of enzymes responsible for the addition and removal of O-GlcNAc. We have focused on methods that require standard laboratory equipment. However, in some cases we also discuss more specialized technology. The unit is set out in a stepwise manner. First, a protocol for increasing the stoichiometry of O-GlcNAc on proteins is given (see Basic Protocol 1). This is followed by simple techniques for the detection/screening of O-GlcNAc modified proteins either by western blotting or lectin affinity chromatography (see Basic Protocols 2 to 4). Separate protocols verify that the glycan is O-linked GlcNAc (see Support Protocols 1 and 2). These methods are followed by protocols for more comprehensive analysis of O-GlcNAc modified proteins, including labeling of O-GlcNAc residues with [3H]Gal, and subsequent product analysis (see Alternate Protocol 1, see Basic Protocols 5 to 8, and see Support Protocols 3 and 4). The final two protocols assay for O-GlcNAc transferase and O-GlcNAcase activity, respectively (see Basic Protocols 9 and 10 and Support Protocol 5). INCREASING THE STOICHIOMETRY OF O-GlcNAc ON PROTEINS BEFORE ANALYSIS
BASIC PROTOCOL 1
In many cultured mammalian cells, the number of O-GlcNAc moieties per protein molecule can be increased by treating cells with O-(2-acetamido-2-deoxy-D-glucopyranosylidene)-amino-N-phenyl-carbamate (PUGNAc), a potent and cell-permeable inhibitor of O-GlcNAcase (Haltiwanger et al., 1998). Alternatively, streptozotocin (STZ; Roos et al., 1998) and glucosamine (Han et al., 2000) have been used to increase the stoichiometry of O-GlcNAc on proteins. However, STZ has been shown to induce poly-(ADP-ribose) polymerase–mediated apoptosis in Min6 cells (Gao et al., 2000) and should be used with caution. In addition, STZ is only effective in cells that express the glucose transporter GLUT-2 (Schnedl et al., 1994). Post-Translational Modification: Glycosylation Contributed by Natasha E. Zachara, Robert N. Cole, Gerald W. Hart, and Yuan Gao Current Protocols in Protein Science (2001) 12.8.1-12.8.25 Copyright © 2001 by John Wiley & Sons, Inc.
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Materials Cells of interest growing in monolayer culture, and appropriate culture medium 20 mM PUGNAc (CarboGen Laboratories) stock in Milli-Q water (filter sterilize and store in aliquots up to 6 to 12 months at −80°C) 500 mM glucosamine stock in 500 mM HEPES, pH 7.5 (make just prior to use; filter sterilize) 500 mM streptozotocin (STZ; Sigma) in 100 mM citrate buffer, pH 4.5 (make just prior to use; filter sterilize) 100-mm tissue culture dishes 1. Grow cells in monolayer culture in a sufficient number of 100-mm dishes. 2. Add (or replace growth medium with fresh medium containing) 40 to 100 µM PUGNAc (added from 20 mM stock), 5 mM glucosamine (added from 500 mM stock), or 2 to 5 mM STZ (added from 500 mM stock). Incubate cells in an incubator for 6 to 18 hr. PUGNAc is taken up by both dividing and stationary cells. If necessary, cells can be treated by PUGNAc for several days without any apparent cell toxicity. However, prolonged treatment does not appear to result in additional increase in O-GlcNAc compared to the 6 to 18 hr treatment. When using glucosamine, mannitol is often added at the same concentration. This controls for changes in osmolarity due to the additional sugar in the medium (Heart et al., 2000).
3. At the end of treatment, take the dishes out of the incubator and place on ice. Extract as desired. Separate proteins by SDS-PAGE (UNIT 10.1) and electroblot onto appropriate membrane (UNIT 10.7). Alternatively, extract proteins and proceed with protein purification or immunoprecipitation. BASIC PROTOCOL 2
DETECTION OF PROTEINS MODIFIED BY O-GlcNAc USING ANTIBODIES Recently Comer et al. (2001) showed that an antibody (CTD 110.6) raised against the glycosylated C-terminal domain of the RNA polymerase II large subunit was a general O-GlcNAc antibody. Unlike many lectins, CTD 110.6 shows little cross-reactivity toward terminal GlcNAc on complex glycans. Several other antibodies, HGAC 85 (Turner et al., 1990) and RL2 (Snow et al., 1987), have been reported as O-GlcNAc specific antibodies. However, these antibodies only recognize a subset of O-GlcNAc modified proteins, and RL2 in particular is dependent on the structure of the peptide backbone around the glycosylation site (Holt et al., 1987). It is important to include an appropriate negative control (100 ng ovalbumin) for cross-reactivity toward N-linked glycans. Using the standard Amersham Pharmacia Biotech enhanced chemiluminescent (ECL) system, the authors have found that 10 µg of cytoplasmic, nuclear, or total cell extract is sufficient. For purified proteins, Comer and co-workers found that 25 to 50 ng of a neoglycoconjugate was sufficient (Comer et al., 2001).
Detection and Analysis of Proteins Modified by O-Linked NAcetylglucosamine
Materials Purified or crude protein (e.g., Basic Protocol 1) separated by SDS-PAGE (UNIT 10.1) and electroblotted to polyvinylidene difluoride (PVDF; UNIT 10.7) or nitrocellulose (duplicate blots are needed) TBS-HT (see recipe)
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Antibody: CTD 110.6 ascites (Covance) diluted 1/2500 in TBS-HT (see recipe for TBS-HT) N-acetylglucosamine (GlcNAc; Sigma) HRPO-conjugated anti–mouse IgM diluted 1/5000 in TBS-HT TBS-HD (see recipe) ECL kit (Amersham Pharmacia Biotech) Additional reagents and equipment for visualization with chromogenic and luminescent substrates (UNIT 10.10) 1. Block blots by incubating with TBS-HT for 60 min at room temperature. Batteiger et al. (1982) have shown that high concentrations of Tween 20 substitute for blocking with milk or bovine serum albumin (BSA).
2. Incubate blots with CTD 110.6 (1/2500 dilution in TBS-HT), in duplicate, with and without 10 mM GlcNAc, overnight at 4°C. To control for specificity it is important to perform a control blot. Here, the antibody is preincubated with 10 mM GlcNAc for ∼5 min on ice before being applied to the control blot. Note that the concentration of antibody should be optimized with each new preparation. We find that 1/2500 is a good place to start with ascites (containing antibody in the mg/ml range).
3. Wash blots in TBS-HD twice, each time for 10 min at room temperature. As the blots are washed under these conditions, prestained markers will fade. This does not appear to be the result of proteins being solubilized off the membranes, but of the dye used to stain the markers. While the markers fade, they do not completely disappear. The authors usually double the amount of marker used, to compensate for fading.
4. Wash blots in TBS-HT three times, each time for 10 min, room temperature. 5. Incubate blots with HRPO-conjugated anti–mouse IgM (1/5000 dilution in TBS-HT) for 50 min, room temperature. The concentration of secondary antibody varies from lot to lot and should be optimized each time with each new preparation.
6. Wash blots in TBS-HD twice, each time for 10 min, room temperature. 7. Wash blots in TBS-HT three times, each time for 10 min, room temperature. 8. Develop the HRPO reaction using, e.g., the ECL system (UNIT 10.10). Note that the antibody often cross-reacts with pre-stained markers.
DETECTION OF PROTEINS MODIFIED BY O-GlcNAc USING THE LECTIN sWGA
BASIC PROTOCOL 3
Many lectins are reportedly specific for β-GlcNAc residues. The authors have typically used succinylated wheat germ agglutinin (sWGA), which is widely available and is derivatized with a number of useful functional groups including horseradish peroxidase (HRPO). Before succinylation, WGA will recognize both silaic acid and GlcNAc (Monsigny et al., 1980). For additional information concerning lectin chromatography, see UNIT 9.1. The amount of “test” protein used is dependent on the technique(s) used to develop the HRPO reaction. Using the standard Amersham Pharmacia Biotech ECL system the authors find that 7.5 µg of cytoplasmic or nuclear extract is sufficient.
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It is important to include an appropriate positive (100 ng ovalbumin) and negative (100 ng of BSA) control. As a control, a portion of the sample should also be treated with hexosaminidase (see Support Protocol 2), to show that reactivity is toward GlcNAc. In addition, the sample should be subjected to reductive β-elimination to verify that lectin/antibody reactivity is towards O-linked glycans (see Support Protocol 1). Materials Purified or crude protein separated by SDS-PAGE (UNIT 10.1) and electroblotted to polyvinylidene difluoride (PVDF; UNIT 10.7) or nitrocellulose (duplicate blots are needed) 5% (w/v) BSA in TBST (see recipe for TBST) TBST (see recipe) 0.1 µg/ml HRPO-conjugated sWGA (EY Labs) in TBST (see recipe for TBST): the lectin can be stored at 1 mg/ml in 0.01 M PBS pH 7.4 (APPENDIX 2E), at −20°C for at least 1 year N-acetylglucosamine (GlcNAc; Sigma) High-salt TBST (HS-TBST): TBST (see recipe) containing 1 M NaCl Tris-buffered saline (TBS; see recipe) ECL kit (Amersham Pharmacia Biotech) Additional reagents and equipment for visualization with chromogenic and luminescent substrates (UNIT 10.10) 1. Wash duplicate blots for 10 min in 5% BSA/TBST, room temperature. 2. Block by incubating blots in 5% (w/v) BSA/TBST for at least 60 min at room temperature. IMPORTANT NOTE: Milk cannot be used as the blocking agent, since many of the proteins in milk are modified by glycans that react with sWGA.
3. Wash blots three times, each time for 10 min in TBST, room temperature. 4. Incubate blots in 0.1 µg/ml sWGA-HRPO in TBST, in duplicate, with and without 1 M GlcNAc, overnight at 4°C. To control for lectin specificity it is important to perform a control blot. Here, the lectin is preincubated with 1 M GlcNAc for ∼5 min on ice before being applied to the control blot.
5. Wash blots six times, each time for 10 min, in HS-TBST. 6. Wash blots once in TBS for 10 min. 7. Develop the HRPO-reaction using, e.g., the the ECL system (UNIT 10.10). Using the ECL system described, 100 ng of ovalbumin should be visualized in 5 to 15 sec. SUPPORT PROTOCOL 1
Detection and Analysis of Proteins Modified by O-Linked NAcetylglucosamine
CONTROL FOR O-LINKED GLYCOSYLATION Traditionally, mild alkaline reduction (reductive β-elimination) has been used to release O-linked carbohydrates from proteins (Amano and Kobata, 1990). This method has been adapted for blots to show that lectin/antibody reactivity is toward O-linked rather than N-linked glycans (Duk et al., 1997). Proteins blotted to PVDF are treated with 55 mM NaOH overnight (releasing O-linked sugars) and then probed using lectins or antibodies. There are a number of reasons why lectin/antibody reactivity could be lost after NaOH treatment, e.g., the sugars were destroyed instead of being released, or the protein was degraded. To control for these, it is important to have control proteins with N- and O-linked sugars, and to stain one blot for protein after treatment preferably with an antibody. The
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authors suggest a control blot of bovine asialofetuin (Sigma) which contains both N- and O-linked sugars terminating in GlcNAc, treated and not treated with PNGase F (UNITS 12.4 & 12.6). Materials Protein samples and controls blotted to PVDF (triplicate blots are needed; nitrocellulose is not suitable as it dissolves in 55 mM NaOH) Tris-buffered saline (TBS; see recipe) 55 mM NaOH 3% (w/v) BSA in TBST (see recipe for TBST) 40°C water bath Additional reagents and equipment for probing protein blots with protein-specific antibodies (see Basic Protocol 2) or lectins (see Basic Protocol 3) 1. Wash blots once in TBS for 10 min. 2. Incubate two blots in 55 mM NaOH at 40°C overnight; incubate the control blot in Milli-Q water at 40°C overnight. The blots treated with NaOH will yellow slightly.
3. Wash blots three times, each time for 10 min at room temperature, in TBST. 4. Block by incubating blots in 3% w/v BSA/TBST for 60 min at room temperature. 5. Probe blots (one treated and one untreated) with carbohydrate-specific lectins (see Basic Protocol 3) or antibodies (see Basic Protocol 2). Probe the second NaOHtreated blot with a protein-specific antibody (see Basic Protocol 2). On the untreated blot, asialofetuin ± PNGase F should react with sWGA, as both the Nand O-linked sugars contain terminal GlcNAc residues. On the treated blot only the asialofetuin − PNGase F should react with sWGA.
DETECTION AND ENRICHMENT OF PROTEINS USING sWGA-AGAROSE sWGA lectin affinity chromatography provides a convenient method for enriching and detecting O-GlcNAc modified proteins. This procedure has been adapted for detecting proteins that are difficult to purify or are present in low copy number, such as transcription factors. In this protocol, the protein of interest is synthesized in a rabbit reticulocyte lysate (RRL) in vitro transcription translation (ITT) system (Promega) and labeled with either [35S]Met, [35S]Cys, or [14C]Leu. After desalting, the proteins are tested for their ability to bind sWGA agarose in a GlcNAc-specific manner (Roquemore et al., 1994).
BASIC PROTOCOL 4
Alternatively, the lectin Ricinus communis agglutinin 1 (RCA1) has been used to select for O-GlcNAc proteins that have previously been labeled by galactosyltransferase (see Alternate Protocol 1). Proteins modified by terminal Gal are specifically retained on a RCA1 affinity column. Labeled O-GlcNAc proteins are released under mild conditions, while those containing N-linked structures require lactose addition to the buffer before elution results (Hayes et al., 1995; Greis and Hart, 1998). The method described in this protocol can be adapted for RCA1 affinity chromatography by substituting RCA1-agarose (EY Labs) for sWGA-agarose and changing the order of the Gal and GlcNAc elution buffer. Materials cDNA subcloned into an expression vector with an SP6 or T7 promoter (∼0.5 to 1 µg/µl) Kit for RRL ITT system (Promega)
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Supplement 25
Label: [35S]Met, or [35S]Cys, or [14C]Leu sWGA-agarose (Vector Laboratories) sWGA wash buffer: PBS (APPENDIX 2E) containing 0.2% (v/v) NP-40 sWGA Gal elution buffer (see recipe) sWGA GlcNAc elution buffer (see recipe) 1-ml tuberculin syringe with glass wool plug at bottom to support chromatography matrix or Bio-Rad Bio-Spin disposable chromatography column Additional reagents and equipment for digesting proteins with hexosaminidase (see Support Protocol 2), desalting (see Support Protocol 5), SDS-PAGE (UNIT 10.1), and autoradiography (UNIT 10.11) Prepare proteins 1. Synthesize proteins to incorporate the desired label ([35S]Met, [35S]Cys, or [14C]Leu) using the RRL ITT system according to the manufacturer’s instructions. Include the protein of interest, a positive control for sWGA binding (for example, the nuclear pore protein p62), a negative control (luciferase, supplied with kit), and a no-DNA control. 2. Treat half of each sample with hexosaminidase (see Support Protocol 2). 3. Desalt samples using spin filtration (e.g., Amersham Pharmacia Biotech Microspin G-50 columns) or a 1-ml G-50 desalting column (as for desalting O-GlcNAc transferase; see Support Protocol 5). The following procedure is carried out at 4°C.
Apply protein samples to chromatography columns 4. Equilibrate sWGA-agarose and pack column as follows: a. If resin is supplied as 50% slurry (i.e., 50% resin/50% storage solution) remove 300 µl (double the volume required) and pipet into a 1-ml tuberculin syringe or disposable chromatography column. b. Let storage solution drain from resin. c. Equilibrate resin by washing column four times, each time with 1 ml of sWGA wash buffer. Cap column. The volumes given are appropriate for a sample derived from an ITT. For enrichment of other protein samples, the volume of sWGA should be optimized for the protein sample applied. The authors find that 50 mg of cell extract requires 1 ml of sWGA-agarose, assuming that 1% to 2% of the total cell extract is modified by O-GlcNAc.
5. Apply sample (∼30 µl of an ITT reaction) to the column and let stand at 4°C for 30 min, or cap and incubate at 4°C for 30 min with rotating or rocking. Wash column and elute GlcNAc 6. At the end of the 30-min incubation, uncap the column and allow the sample to “run through” the resin. Collect this as the “run through” fraction. Wash column with 15 ml of sWGA wash buffer at 10 ml/hr, collecting 0.5-ml fractions. 7. Load the column with 300 µl of sWGA Gal elution buffer and let stand at 4°C for 20 min. 8. Wash column with 5 ml of sWGA Gal elution buffer, collecting 0.5 ml fractions. Detection and Analysis of Proteins Modified by O-Linked NAcetylglucosamine
9. Repeat steps 6 to 8 using GlcNAc elution buffer. 10. Count 25 µl of each fraction using a liquid scintillation counter.
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11. Pool positive fractions that elute in the presence of GlcNAc and precipitate using TCA or methanol. To precipitate proteins with methanol, mix 1 vol of sample with 10 vol of ice-cold methanol. Incubate overnight at −20°C. Recover protein by microcentrifuging 10 min at 16,000 × g, 4°C, in a microcentrifuge tube (which is the most efficient procedure) or in 15-ml conical centrifuge tubes for 10 min at 3000 × g, 4°C. Resuspend samples in SDS-PAGE sample buffer (UNIT 10.1). As many proteins in rabbit reticulocyte lysate contain O-GlcNAc and bind WGA, the authors do not recommend the addition of carrier proteins at this point. Typically, a fraction of the GlcNAc elution containing a total of 1000 to 2000 dpm [35S]Met is precipitated and analyzed by SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11). The use of acetone to precipitate proteins is not recommended, as free GlcNAc will also precipitate.
12. Analyze pellet by SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11). DIGESTION OF PROTEINS WITH HEXOSAMINIDASE Terminal GlcNAc and O-GlcNAc can be removed from proteins using commercially available hexosaminidases; these enzymes will also cleave terminal GalNAc residues. Unlike O-GlcNAcase, commercial hexosaminidases have low pH optima, typically pH 4.0 to 5.0.
SUPPORT PROTOCOL 2
Materials Protein sample for digestion (include a positive control, e.g., ovalbumin) 2% (w/v) SDS (see APPENDIX 2E for 20× stock) 2× hexosaminidase reaction mixture (see recipe) Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and electroblotting (UNIT 10.7) 1. Mix sample 1:1 with 2% SDS and boil for 5 min. 2. Mix sample 1:1 with 2× hexosaminidase reaction mixture and incubate at 37°C for 4 to 24 hr. 3. To assess completeness of the digestion, separate an aliquot of the reaction by SDS-PAGE (UNIT 10.1) and electroblot (UNIT 10.7) onto an appropriate membrane. Probe blots with carbohydrate-specific lectins or antibodies. DETECTION OF PROTEINS MODIFIED BY O-GlcNAc USING GALACTOSYLTRANSFERASE The enzyme β-1,4-galactosyltransferase (from bovine milk) will label any terminal GlcNAc residue with Gal, using uridine diphospho-D-Gal (UDP-Gal) as a donor substrate (Brew et al., 1968). Hart and colleagues have exploited this property, using the enzyme to label terminal GlcNAc residues on proteins with [6-3H]Gal, forming a [3H]-βGal14βGlcNAc (Torres and Hart, 1984; Roquemore et al., 1994; Greis and Hart, 1998). The labeled sugar can be chemically released (via β-elimination) and analyzed by size-exclusion chromatography on a BioGel-P4 column, using the 3H radiolabel to detect the fraction of interest (Roquemore et al., 1994). Labeling the O-GlcNAc allows for the subsequent detection of the proteins and peptides of interest during SDS-PAGE, HPLC, protease digestion, and Edman degradation steps. Researchers have been able to identify glycosylation sites on as little as 10 pmol using these methods (Greis et al., 1996).
ALTERNATE PROTOCOL 1
Post-Translational Modification: Glycosylation
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To achieve efficient labeling of some proteins, it is necessary to denature samples, for example by boiling in the presence of 10 mM DTT and 0.5% (w/v) SDS. Galactosyltransferase has been shown to be active in solutions containing 5 mM DTT, 0.5 M NaCl, up to 2% (v/v) Triton X-100, up to 2% (v/v) NP-40, and 1 M urea. Up to 0.5% (w/v) SDS can be used, if it is titrated with a 10-fold molar excess of either Triton X-100 or NP-40 in the final reaction mixture. Digitonin, which is commonly used to solubilize cells, should be used with caution, as it is a substrate for galactosyltransferase. The total ionic strength should be less than 0.2 M. Galactosyltransferase requires 1 to 5 mM Mn2+ for activity, but is inhibited by Mg2+ and concentrations of Mn2+ >20 mM. EDTA (or analogs) should be avoided unless titrated with appropriate levels of Mn2+. Note that 1 mol of EDTA binds 2 mol of Mn2+. Free UDP is also an inhibitor of galactosyltransferase. For studies where complete labeling of the GlcNAc is preferable, such as site mapping, calf intestinal alkaline phosphatase is included in the reaction, as it degrades UDP (Unverzagt et al., 1990). While this increases the efficiency of the reaction, it is important to add this to the control as some preparations of alkaline phosphatase contain proteins what will label with galactosyltransferase (R.N. Cole, pers. commun.). NOTE: Protease inhibitors, such as PIC 1, PIC 2, and PMSF (see recipe for 1000× protease inhibitors in Reagents and Solutions), can be included (final concentrations, 1×), but GlcNAc and 1-amino GlcNAc should be removed prior to labeling by spin filtration or another method of desalting. Materials Protein sample(s) Dithiothreitol (DTT) Sodium dodecyl sulfate (SDS; see APPENDIX 2E for 20% stock solution) Label: 1.0 mCi/ml UDP-[3H]Gal, (17.6 Ci/mM; Amersham Pharmacia Biotech) in 70% v/v ethanol Nitrogen source 25 mM 5′-adenosine monophosphate (5′-AMP), in Milli-Q water, pH 7.0 Buffer H (see recipe) 10× galactosyltransferase labeling buffer (see recipe) Galactosyltransferase, autogalactosylated (see Support Protocol 3) Calf intestinal alkaline phosphatase Unlabeled UDP-Gal Stop solution: 10% (w/v) SDS/0.1 M EDTA 100°C water bath 30 × 1–cm Sephadex G-50 column equilibrated in 50 mM ammonium formate/0.1% (w/v) SDS Additional reagents and equipment for acetone precipitation of protein (UNIT 4.5), PNGase F digestion of proteins (UNITS 12.4 & 12.6), SDS-PAGE (UNIT 10.1), and product analysis (see Basic Protocol 6) Prepare the reaction 1. Denature protein sample by adding DTT to 10 mM and SDS to 0.5% (w/v), then boiling the sample for 10 min. Detection and Analysis of Proteins Modified by O-Linked NAcetylglucosamine
2. Decide how many reactions are going to be carried out and thus how much label will be needed (∼1 to 2 µCi/reaction).
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A positive control (ovalbumin, 2 ìg), a negative control (because galactosyltransferase can label itself), and a sample-minus-enzyme control will be needed.
3. Remove solvent from label in a Speed-Vac evaporator or under a stream of nitrogen. Ethanol can inhibit the galactosyltransferase reaction, but if <4 ìl is required the label can be added directly to the reaction (final reaction volume, 500 ìl).
4. Resuspend appropriate amount of label for each reaction, respectively, in 50 µl of 25 mM 5′-AMP. The AMP is included to inhibit possible phosphodiesterase reactions, which might compete for label during the labeling experiment.
5. Set up reactions as follows: Up to 50 µl protein sample (final concentration 0.5 to 5 mg/ml) 350 µl buffer H 50 µl 10× galactosyltransferase labeling buffer 50 µl UDP-[3H]Gal/5′-AMP mixture from step 4 30 to 50 µl autogalactosylated galactosyltransferase 1 to 4 U calf intestinal alkaline phosphatase Milli-Q water to final volume of 500 µl Reaction volumes can be scaled down to 50 ìl.
6. Labeling at 37°C for 2 hr or at 4°C overnight. These are the typical conditions. Galactosyltransferase is active over a range of temperatures.
7. Add unlabeled UDP-Gal to a final concentration of 0.5 to 1.0 mM and another 2 to 5 µl of galactosyltransferase. For studies where complete labeling of the GlcNAc is required, such as site mapping, the reactions are chased with unlabeled UDP-Gal and fresh galactosyltransferase.
8. Add 50 µl of stop solution to each sample and heat to 100°C for 5 min in a water bath. Isolate the product 9. Resolve the protein from unincorporated label using a Sephadex G-50 column equilibrated in 50 mM ammonium formate/0.1% w/v SDS. Collect 1-ml fractions. Size-exclusion chromatography using Sephadex G-50 is traditionally used to desalt samples. However, TCA precipitation, spin filtration/buffer exchange, or other forms of size-exclusion chromatography (e.g., Pharmacia PD-10 desalting column) can be used. The addition of carrier proteins such as BSA (∼67 kDa) and cytochrome c (∼12.5 kDa) to samples and buffers will reduce the amount of protein lost due to nonspecific protein adsorption.
10. Count a 50-µl aliquot of each fraction using a liquid scintillation counter. Approximately 2 × 106 dpm of [3H]Gal should be incorporated into 2 ìg of ovalbumin.
11. Combine the void volume and lyophilize to dryness. 12. Resuspend samples in Milli-Q water and precipitate with acetone (UNIT 4.5). 13. Treat samples with PNGase F (UNITS 12.4 & 12.6), separate by SDS-PAGE (UNIT 10.1) and detect by autoradiography (UNIT 10.11). Alternatively, subject samples to “product analysis” to confirm that the label was incorporated onto O-GlcNAc (see Basic Protocol 6).
Post-Translational Modification: Glycosylation
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SUPPORT PROTOCOL 3
AUTOGALACTOSYLATION OF GALACTOSYLTRANSFERASE As galactosyltransferase contains N-linked glycosylation sites, it is necessary to block these before using this enzyme to probe other proteins for terminal GlcNAc. Materials 10× galactosyltransferase labeling buffer (see recipe) 10,000 U/ml aprotinin 2-mercaptoethanol UDP-Gal Saturated ammonium sulfate: >17.4 g (NH4)2SO4 in 25 ml Milli-Q water 85% ammonium sulfate: 14 g (NH4)2SO4 in 25 ml Milli-Q water Galactosyltransferase storage buffer (see recipe) 30- to 50-ml centrifuge tubes Refrigerated centrifuge 1. Resuspend 25 U of galactosyltransferase in 1 ml of 1× galactosyltransferase labeling buffer. 2. Transfer sample to 30- to 50-ml centrifuge tube. The centrifuge tubes should be able to withstand a centrifugal force of 15,000 × g.
3. Remove a 5-µl aliquot for an activity assay. This is the “Pre-Gal” sample to be used in Support Protocol 4.
4. Add 10 µl of 10,000 U/ml aprotinin, 3.5 µl of 2-mercaptoethanol, and 1.5 to 3.0 mg of UDP-Gal. 5. Incubate the sample on ice for 30 to 60 min. 6. Add 5.66 ml of prechilled saturated ammonium sulfate in a dropwise manner. Incubate on ice for 30 min. 7. Centrifuge 15 min at >10,000 × g, 4°C. Pour off supernatant. 8. Resuspend pellet in 5 ml cold 85% ammonium sulfate and incubate on ice for 30 min. 9. Centrifuge 15 min at >10,000 × g, 4°C, and pour off supernatant. 10. Resuspend pellet in 1 ml of galactosyltransferase storage buffer and divide into 50-µl aliquots, saving 5 µl for an activity assay as the “Auto-Gal” sample. Assay that aliquot for activity (see Support Protocol 4). 11. Store remaining aliquots up to 1 year at −20°C pending use in Alternate Protocol 1. SUPPORT PROTOCOL 4
Detection and Analysis of Proteins Modified by O-Linked NAcetylglucosamine
ASSAY OF GALACTOSYLTRANSFERASE ACTIVITY As sample and activity may be lost during the autogalactosylation procedure, it is important to assess the activity of the enzyme. Materials 1.0 mCi/ml UDP-[3H]Gal, (17.6 Ci/mM; Amersham Pharmacia) in 70% v/v ethanol Nitrogen source 1× galactosyltransferase dilution buffer: galactosyltransferase storage buffer (see recipe) supplemented with 5 mg/ml BSA 10× galactosyltransferase labeling buffer (see recipe)
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25 mM 5′-adenosine monophosphate (5′-AMP) in Milli-Q water, pH 7.0 “Pre-Gal” sample aliquot (see Support Protocol 3, step 3) and “Auto-Gal” sample aliquot (see Support Protocol 3, step 10) 200 mM GlcNAc Dowex AG1-X8 resin (PO4 form) slurry in 20% (v/v) ethanol Glass wool 1. Dry 40 µl of 0.1 µCi/µl of UDP-[3H]Gal in a Speed-Vac evaporator or under a stream of nitrogen. 2. Resuspend in 90 µl of 25 mM 5′-AMP. 3. Make 1/1000, 1/10,000, and 1/100,000 serial dilutions of the “Pre-Gal” and “AutoGal” sample aliquots, in 1× galactosyltransferase dilution buffer. Using these dilutions, 200 mM GlcNAc, and 10× galactosyltransferase labeling buffer, prepare reaction mixtures as described in Table 12.8.1. 4. Start the reaction by adding 10 µl of 0.05 µCi/µl UDP-[3H]Gal (see step 2) to each tube. 5. Incubate samples at 37°C for 30 min. While the samples are incubating, prepare the columns.
6. Pour 1 ml of Dowex AG1-X8 slurry (PO4 form) into 13 Pasteur pipets, each plugged with a small amount of glass wool. The glass wool prevents the resin from flowing out of the column. If too much glass wool is used, it will reduce the flow rate of the column. The glass wool plug should be 0.3 to 0.5 mm long and should not be over-compressed.
7. Wash with at least 3 ml of Milli-Q water. Do not let the columns run dry. 8. When almost all the Milli-Q water has eluted, place each column over a separate 15-ml scintillation vial. 9. Stop the reaction (still incubating from step 5) by adding 500 µl of Milli-Q water. 10. Load each sample onto the corresponding column and add a 500 µl water wash of the tube. Collect eluate as fraction A. 11. Elute with two 1-ml additions of Milli-Q water. Collect eluates as fractions B and C, respectively.
Table 12.8.1
Sample
Blank Pre-Gal
Auto-Gal
Reaction Mixtures for Assay of Galactosyltransferase Activity
Dilution
1/1000 1/10,000 1/100,000 1/1000 1/10,000 1/100,000
Vol. (µl) Vol. (µl) of Vol. (µl) of 10× Gal 200 mM diluted labeling GlcNAc sample buffer
Vol. (µl) Milli-Q water
0 10 10 10 10 10 10
70 60 60 60 60 60 60
10 10 10 10 10 10 10
10 10 10 10 10 10 10
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12. Count 100 µl of the sample and 10 µl of the UDP-[3H]Gal using a liquid scintillation counter. 13. Calculate activity. a. Calculate the total moles of UDP-Gal transferred to GlcNAc as follows: {cpm(with GlcNAc) − cpm(no GlcNAc)/total cpm} × 1.76 nmol Note, cpm(with GlcNAc) represents counts in the sample, cpm(no GlcNAc) represents counts in the no-enzyme control, and total cpm represents the total counts available to transfer. {cpm(with GlcNAc) − cpm(-GlcNAc)/total cpm} represents the portion of [3H]Gal transferred in the reaction. This proportion is multiplied by the total number of moles of UDP-Gal in the reaction. If the specific activity of the UDP-[3H]Gal is 17.6 Ci/mM and 10 ìl of label was used, then there are 1.76 nmol of UDP-Gal in 10 ìl.
b. Calculate the activity; one unit of activity (U) is defined as 1 µM of Gal transferred to GlcNAc per minute at 37°C. BASIC PROTOCOL 5
DETECTION OF PROTEINS MODIFIED BY O-GlcNAc USING METABOLIC LABELING Metabolic labeling of O-GlcNAc-bearing proteins provides a useful means to test if a protein of interest is modified by O-GlcNAc, to observe the gross dynamic changes in O-GlcNAc levels and to study the subcellular localization in response to stimulation or during cell cycle. In the protocol described below, cells are labeled with [3H]glucosamine, which is metabolized to UDP-[3H]GlcNAc in the hexosamine synthetic pathway. For labeling, it is critical that the labeled sugar compete with glucose import; this ensures efficient uptake of the label. While glucosamine is a good competitor, N-acetylglucosamine is not. For further discussions on metabolic labeling of glycoconjugates, readers are encouraged to consult Varki (1994) for details. Materials Cells of interest, growing in culture Biosynthetic labeling medium (see recipe) Phosphate-buffered saline (PBS; APPENDIX 2E) Additional reagents and equipment for PNGase F digestion of proteins (UNITS 12.4 & 12.6), SDS-PAGE (UNIT 10.1), and autoradiography (UNIT 10.11) 1. Replace growth medium with biosynthetic labeling medium. Label cells for 5 to 24 hr in an incubator. The labeling is done in a glucose-free medium to maximize labeling efficiency. The addition of nonessential amino acids to the medium reduces the influx of glucosamine into the amino acid biosynthetic pathways (Medina et al., 1998).
2. Wash labeled cells twice, each time by resuspending in 5 ml of PBS and centrifuging 10 min at 500 × g, 4°C. 3. Extract as desired. See Critical Parameters for a discussion on reducing O-GlcNAcase activity.
Detection and Analysis of Proteins Modified by O-Linked NAcetylglucosamine
There are many types of cell fractionation and extraction techniques available. Those that minimize plasma membrane and organelle contamination are preferable. If a particular study concerns one or a few proteins, and specific antibodies are available, the protein(s) can be isolated by immunoprecipitation after metabolic labeling.
4. Treat sample with PNGase F (UNITS 12.4 & 12.6).
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5. Separate proteins by SDS-PAGE (UNIT 10.1) and visualize labeled proteins by autoradiography (UNIT 10.11). CHARACTERIZATION OF LABELED GLYCANS BY â-ELIMINATION AND CHROMATOGRAPHY
BASIC PROTOCOL 6
This protocol has three steps: (1) the release of carbohydrates as sugar alditols by reductive β-elimination; (2) desalting the sample, while confirming the size of the labeled sugar alditol(s); and (3) confirmation that the product is [3H] βGal1-4βGlcNAcol (from galactosyltransferase labeling) or [3H]GlcNAcol (from metabolic labeling). Materials Labeled proteins β-elimination reagent: 1 M NaBH4/0.1 M NaOH (prepare fresh) 4 M acetic acid Screw-cap microcentrifuge tubes Additional reagents and equipment for acetone or methanol precipitation of proteins (UNIT 4.5), size-exclusion (gel-filtration) chromatography (UNIT 8.3), and Dionex chromatography (Townsend et al., 1990; Hardy and Townsend, 1994) 1. Acetone or methanol precipitate labeled proteins (UNIT 4.5) in screw-cap microcentrifuge tubes. The β-elimination reaction is performed in screw-cap microcentrifuge tubes to prevent the lids from popping open during the lengthy incubation, which would result in evaporation of the sample. The tubes should be tightly sealed. Take care in opening the tubes at the end of the reaction, as gas generated during the reaction will escape.
2. Resuspend sample in 500 µl of β-elimination reagent and incubate at 37°C for 18 hr. After several hours check that the pH is >13. Add more β-elimination reagent if needed.
3. Cool the sample on ice. 4. Neutralize the reaction by adding 5 µl cold 4 M acetic acid in a stepwise manner. Check that the pH is between pH 6 and 7. Samples can be desalted either by chromatography on a Sephadex G-50 column (1 × 30 cm, equilibrated in 50 mM ammonium formate, 0.1% SDS) or by anion-exchange chromatography on a 1-ml Bio-Rad Dowex AG 50W-X2 200-400 mesh (H+ form) column equilibrated in water. Fractions containing [3H]GlcNAc or [3H]Gal are pooled and lyophilized. Residual NaBH4 is removed by washing the sample with methanol; NaBH4 is volatile in the presence of methanol and is removed in a Speed-Vac evaporator or under a stream on nitrogen (Fukuda, 1990).
5. Resuspend the sugar alditols in Milli-Q water. Analyze by size-exclusion chromatography or by Dionex chromatography. To determine the size of the oligosaccharide, labeled glycans released by β-elimination are subjected to size exclusion chromatography. Readers are referred to several standard methods using BioGel P4 (Kobata, 1994) or TSK Fractogel (Fukuda, 1990) chromatography. Alternatively, the Amersham Pharmacia Biotech Superdex Peptide column, equilibrated in 30% v/v CH3CN, 0.1% v/v TFA, has been used to size oligosaccharides (R.N. Cole, pers. commun.). To determine the nature of the monosaccharide alditol or disaccharide alditol generated from either metabolic labeling or galactosyltransferase labeling, samples released by β-elimination can be analyzed by high-voltage paper electrophoresis or high-pH anion exchange chromatography (HAPEC) with pulsed amperometric detection on a Dionex CarboPac PA100 column (Townsend et al., 1990; Hardy and Townsend, 1994).
Post-Translational Modification: Glycosylation
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BASIC PROTOCOL 7
SITE MAPPING BY MANUAL EDMAN DEGRADATION This protocol describes methods for mapping sites on peptides that have been labeled with [3H]Gal (see Alternate Protocol 1) or [3H]glucosamine (see Basic Protocol 5). Proteins of interest can be digested in solution or in gel (UNITS 11.1 & 11.3) and the peptides separated by conventional reversed-phase HPLC. Glycopeptides are covalently attached to arylamine-derivatized PVDF disks via the activation of peptide carboxyl groups using water-soluble N-ethyl-N′-dimethylamine-nopropylcarbodiimide (EDC; Kelly et al., 1993). Note that both the C-terminus and any acidic residue will couple to the membrane and as such, cycles with acidic amino acids will yield blanks. Some researchers have shown that glycosylated amino acids can be visualized during automated Edman degradation. However, this technique requires at least 20 pmol of starting material, where at least 20% of the sample is glycosylated. For more details, readers are referred to Zachara and Gooley (2000). Materials Acetonitrile (CH3CN; HPLC-grade) Trifluoroacetic acid (TFA; sequencing grade) Sequelon-AA Reagent Kit (Millipore) containing: Mylar sheets Carbodiimide Coupling buffer N-ethyl-N′-dimethylamine-nopropylcarbodiimide (EDC) Labeled sample and control peptides (must contain ≥1000 dpm and not be in amine-containing buffers such as Tris) Methanol (HPLC-grade) Sequencing reagent (see recipe) 100 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) Heating block Screw-capped polypropylene microcentrifuge tubes Couple the sample to a PVDF disk 1. Resuspend peptide in 10% to 30% (v/v) CH3CN and up to 0.1% (v/v) TFA. 2. Place disk on a Mylar sheet (from Sequelon kit) on a heating block at 55°C. 3. Wet disk with 10 µl of methanol and allow excess methanol to evaporate. 4. Apply sample in 10-µl aliquots, allowing the membrane to come to near-dryness between aliquots. 5. Dry the membrane after all of the sample has been applied. Place the disc in the lid of a screw-cap microcentrifuge tube for subsequent reactions. 6. Prepare coupling reagent by combining 1 mg carbodiimide per 100 µl of coupling buffer (both provided with Sequelon kit). Add ∼1 mg of EDC per 100 µl of coupling reagent and carefully pipet ∼50 µl onto each disk. 7. Incubate samples at 4°C for 30 min. At the end of the reaction, remove the coupling reagent to a microcentrifuge tube.
Detection and Analysis of Proteins Modified by O-Linked NAcetylglucosamine
Incubation of the sample at 4°C increases the yield. The membrane should not be incubated for >30 min.
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8. Place the membrane in a microcentrifuge tube. Wash the membrane alternately three times with 1 ml methanol and 1 ml Milli-Q water, vortexing briefly after each addition. Combine each wash with the coupling reagent from step 7. 9. Dry the membrane. Samples are stable at −20°C on disks for at least 6 months. 10. Count an aliquot of the pooled coupling reagent and washes (step 8) and determine the coupling efficiency. Sequencing peptides The times given in this procedure are critical. Steps 12 to 17 represent a “cycle.” This method is only efficient for 10 to 20 cycles. 11. Place disk in a screw-capped polypropylene microcentrifuge tube. 12. Add 0.5 ml of the sequencing reagent to the disk and incubate 10 min at 50°C in a heating block. This step derivatizes the N-terminus of the peptide, forming the phenylthiocarbamyl derivative.
13. Wash disk five times in microcentrifuge tube, each time with 1 ml of methanol, vortexing after each addition. Dry disk in a Speed-Vac evaporator for 5 min. 14. Add 0.5 ml TFA and incubate at 50°C for 6 min. Remove and save the supernatant. In this step, the derivatized amino acid is released from the peptide.
15. Wash the disk with 1 ml of methanol. Save the supernatant and combine the supernatant from step 14. 16. Wash disk five times, each time with 1 ml methanol. 17. Return to step 12 and repeat the cycle. Steps 12 to 17 represent a “cycle,” with one amino acid being released per cycle from the N-terminus of the protein or peptide. This method is only efficient for 10 to 20 cycles. As each cycle represents the release of an amino acid, it is not necessary to perform 20 cycles if the peptide in question is 10 amino acids long. The authors usually perform 15 cycles, as the data become hard to interpret after this.
18. Dry the combined supernatants from steps 14 and 15 on a 50°C heating block, or on the bench overnight. 19. Add 0.5 ml of 100 mM Tris⋅Cl, pH 7.4 and 15 ml liquid scintillation fluid. Count on a liquid scintillation counter. Edman degradation is a sequential process, with one amino acid being released from the N-terminus of the protein or peptide per cycle. In the method described here, glycosylated amino acids are being detected by following the [3H]Gal label. To interpret the data, plot the dpm per cycle as a bar graph. A cycle with a glycosylated amino acid should have much higher counts than either the preceding or following cycle, unless there are two sites in a row. Manual Edman degradation is often performed in concert with other techniques that will identify the peptide being sequenced, e.g., conventional Edman degradation (UNIT 11.5) or MALDI-TOF MS (Chapter 16). When looking at data sets from several techniques, the cycle with the counts should line up with a Ser or Thr residue. Post-Translational Modification: Glycosylation
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BASIC PROTOCOL 8
SITE MAPPING BY MASS SPECTROMETRY AFTER β-ELIMINATION As mass spectrometry (MS; UNITS 16.1 & 16.2) has become more readily available, several MS techniques have been applied to the characterization of O-GlcNAc proteins and peptides (Reason et al., 1992; Greis et al., 1996; Haynes and Aebersold, 2000). When analyzing proteins or peptides modified by O-GlcNAc via mass spectrometry, several challenges need to be overcome. Some researchers have shown that when using matrixassisted laser desorption ionization-time of flight mass spectrometry (MALDI-TOF; UNIT 16.2) with a conventional matrix (such as α-cyano-4-hydroxycinnaminic acid) the addition of a single GlcNAc to a peptide may reduce the signal by approximately five-fold. In addition, the presence of the unglycosylated peptide further suppresses the signal (Hart et al., 2000). Secondly, the β-O-GlcNAc bond is more labile in the mass spectrometer than other bonds formed by protein glycosylation. Thus, in conventional electrospray ionization-MS, O-GlcNAc is released at lower orifice potentials and collision energies than is required to sequence peptides. Greis et al. (1996) developed a technique where peptides are analyzed before and after β-elimination. During β-elimination Ser (89 atomic mass units) and Thr (101 atomic mass units) residues are converted to 2-aminopropenoic acid (69 atomic mass units) and α-aminobutyric acid (83 atomic mass units), respectively. These amino acids have different masses than their parent amino acids and can be used to map the site of attachment. It should be noted that this method will also release phosphate linked to Ser and Thr residues. In the protocol described here, glycans are released from peptides by β-elimination, and then analyzed using conventional MS-MS strategies. Materials Samples from β-elimination (see Basic Protocol 7) 200 mM NaOH 300 mM acetic acid Screw-cap polypropylene microcentrifuge tubes 45°C heating block Additional reagents and equipment for ESI-MS (UNITS 16.8 & 16.9), reversed-phase HPLC of peptides (UNITS 8.7 & 11.6), and MALDI-TOF MS (UNIT 16.2) 1. Combine the following in a screw-cap polypropylene microcentrifuge tube: ≤25 µl sample from β-elimination 25 µl 200 mM NaOH Milli-Q water to a final volume of 50 µl 2. Incubate at 45°C for 4 hr in a heating block. Time is critical, as increased incubation times will result in the peptide backbone hydrolyzing.
3. Place sample on ice and add 25 µl of 300 mM acetic acid in 5-µl increments. The reaction mixture will boil over unless the acetic acid is added in a stepwise manner.
4. Dry sample in a Speed-Vac evaporator and store at −80°C until analyzed by ESI-MS (UNITS 16.8 & 16.9). Alternatively, analyze peptides by MALDI-TOF MS (UNIT 16.2) after desalting using reversed-phase chromatography (UNITS 8.7 & 11.6). Detection and Analysis of Proteins Modified by O-Linked NAcetylglucosamine
Zip-Tips (Millipore) are ideal for desalting small quantities of peptide before MALDI-TOF MS.
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ASSAY FOR OGT ACTIVITY The detection and analysis of O-GlcNAc on proteins is only the first step in the analysis of O-GlcNAc and the protein(s) of interest. More important is determining the function of the modification. Protocols for the analysis of the enzymes that add and remove O-GlcNAc have been included, as they may aid in understanding the role of O-GlcNAc. Recent examples where studies such as this have been critical include those which have shown the reciprocity between O-GlcNAc and O-phosphate on the C-terminal domain of RNA Pol II; studies showing elevated activity of enzymes in certain tissue/cell lines and tissue fractions; and, finally, studies which have indicated that the enzymes responsible for the addition and removal of O-GlcNAc copurify with kinases and phosphatases.
BASIC PROTOCOL 9
O-GlcNAc transferase (OGT), or uridine diphospho-N-acetylglucosamine:polypeptide β-N-acetylglucosaminyltransferase, transfers GlcNAc to the hydroxyl groups of Ser and Thr residues of proteins and peptides using UDP-GlcNAc as a donor substrate (Haltiwanger et al., 1992; Kreppel et al., 1997). OGT activity is assayed by determining the rate at which [3H]GlcNAc is transferred to an acceptor peptide. A number of peptides have been i dent ifi ed as substr at es for OGT i n vi tr o, but a pept ide (340PGGSTPVSSANMM352) from the α-subunit of casein kinase II (CKII) is the most efficient in vitro substrate known to date (Kreppel and Hart, 1999). OGT activity can be assayed in crude preparations (Haltiwanger et al., 1992) or using recombinant protein (Kreppel and Hart, 1999). OGT activity is sensitive to salt inhibition, so it is important to desalt the preparation before assaying if high salt concentrations are present (see Support Protocol 5). For pure preparations 0.2 to 1 µg is typically used per assay, in crude preparations 20 to 50 µg, though the latter is precipitated using ammonium sulfate (40% to 60%). Materials 0.1 mCi/ml UDP-[3H]GlcNAc (20 to 45 Ci/mmol; NEN Life Science Products) in 70% ethanol 25 mM 5′-adenosine monophosphate (5′-AMP), in Milli-Q water, pH 7.0 Nitrogen source Crude or purified OGT sample, desalted (see Support Protocol 5) 10× OGT assay buffer (see recipe) CKII peptide substrate (+H2N-PGGSTPVSSANMM-COO−): dissolve in H2O to 10 mM and adjust to pH 7 if necessary 50 mM formic acid Methanol (HPLC-grade) Waters Sep-Pak C18 cartridges 1. Dry down an aliquot of UDP-[3H]GlcNAc in a Speed-Vac evaporator or under a stream of nitrogen just prior to use. Resuspend in an appropriate volume of 25 mM 5′-AMP, so that the concentration is 0.02 µCi to 0.1 µCi/µl. AMP is included in the assay to competitively inhibit any pyrophosphatase in the sample that will hydrolyze the UDP-GlcNAc.
2. Set up assay reactions as follows: 5 µl of 10× OGT assay buffer 10 µl of 10 mM CKII peptide substrate 5 µl of 0.02 to 0.1 µCi/µl UDP-[3H]GlcNAc (from 0.1 to 0.5 µCi total) ≤25 µl of desalted OGT to be analyzed H2O to 50 µl
Post-Translational Modification: Glycosylation
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It is critical to include a negative control. A mimic of the CKII peptide where the Ser and Thr residues are replaced with Ala is appropriate, or, simply, a “no-enzyme” control can be included. The results generated are variable and the reactions should be set up in duplicate.
3. Incubate at room temperature for 30 min. An incubation time of 15 to 30 min at room temperature is usually sufficient.
4. Stop reaction by adding 450 µl of 50 mM formic acid. 5. Wet a Waters Sep-Pak C18 cartridge with methanol, then wash the cartridge with 5 ml H2O. 6. Load the reaction (500 µl total) onto the cartridge with a syringe. Wash with 5 ml H2O. The CKII peptide binds to the matrix of the C18 cartridge. Unincorporated UDP[3H]GlcNAc is eliminated by the wash.
7. Elute the peptide with 2 to 4 ml methanol, directly into a 15-ml liquid scintillation counter tube. 8. Add 10 ml scintillation fluid and count 3H. Calculate OGT activity according to the following equations. µCi of GlcNAc incorporated = (dpm in sample − dpm in blank)/(2.22 × 106 dpm ⋅ µCi−1) mmol of GlcNAc incorporated = (µCi of GlcNAc incorporated)/(specific activity in µCi/mmol) This number should be expressed in terms of mg of OGT or cell extract. If the assay is done at several time points, it can be expressed as mmol/min. The activity can be expressed either as dpm 3H incorporation into the peptide, or as ìmol 3 H incorporation (1 ìCi = 2.22 × 106 dpm). SUPPORT PROTOCOL 5
DESALTING THE O-GlcNAc TRANSFERASE OGT activity is sensitive to salt inhibition (IC50 = 40 to 50 mM NaCl). It is important to desalt the enzyme preparation before assay if high concentration of salt is present. Materials Sephadex G-50 slurry (Pharmacia Biotech) OGT desalting buffer (see recipe) Protein sample for OGT assay in volume ≤200 µl 1-ml tuberculin syringe 1.5-ml tubes, prechilled 1. Pack a column containing exactly 1 ml of Sephadex G-50 slurry in a 1-ml tuberculin syringe. Wash the column with 5 ml of OGT desalting buffer. Sephadex G-50 is usually supplied in 20% (v/v) ethanol.
2. Load protein sample onto column. Detection and Analysis of Proteins Modified by O-Linked NAcetylglucosamine
The volume of sample can be up to 200 ìl.
3. Wash column with desalting buffer so that the total volume of this wash and the protein sample is 350 µl. For example, if sample volume is 150 µl, add 200 µl desalting buffer to the column at this step.
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4. Transfer the syringe column to a clean, prechilled 1.5-ml tube. Elute protein with 200 µl desalting buffer. Keep on ice. This is the desalted sample. Alternatively, PD-10 desalting columns (Amersham Pharmacia Biotech) can be used if O-GlcNAc transferase is in larger volume (1 to 2.5 ml).
ASSAY FOR O-GlcNAcase ACTIVITY O-GlcNAcase, also known as N-acetylglucosaminidase or hexosaminidase C (EC 3.2.1.52), is a cytosolic glycosidase specific for O-linked β-GlcNAc. The activity of O-GlcNAcase can be conveniently assayed in vitro with a synthetic substrate, p-nitrophenol N-acetylglucosaminide (pNP-β-GlcNAc). The cleavage product, pNP, has an absorbance peak at 400 nm.
BASIC PROTOCOL 10
Materials Partially purified O-GlcNAcase (0.2 to 1 µg) or cell extract sample (20 to 50 µg, precipitated with 30% to 50% ammonium chloride) 10× O-GlcNAcase assay buffer (see recipe) 100 mM (50×) p-nitrophenol N-acetylglucosaminide (pNP-GlcNAc) in DMSO 500 mM Na2CO3 96-well flat bottom plates or 1.5 ml microcentrifuge tubes Plate reader or spectrophotometer 1. Prepare O-GlcNAcase. Native or recombinant O-GlcNAcase can be partially purified from animal tissues or cultured cells by several chromatographic steps (Dong and Hart, 1994; Gao et al., 2001). Alternatively, a crude enzyme preparation can be generated by passing cell extract over a 1-ml Con-A column. Most of the interfering acidic hexosaminidases are modified by N-linked sugars and bind to Con-A, while neutral O-GlcNAcase is in the flow-through (Izumi and Suzuki, 1983).
2. Precool 96-well plate or microcentrifuge tubes on ice. 3. Set up reactions in the precooled plate wells or tubes as follows: 1 to 50 µl partially purified O-GlcNAcase enzyme or cell extract 10 µl 10× O-GlcNAcase assay buffer 2 µl 100 mM pNP-GlcNAc H2O to 100 µl The total reaction volume can be scaled up to 500 ìl in microcentrifuge tubes. pNP-GlcNAc breaks down chemically. A blank reaction without enzyme should be included to determine the background. 50 mM GalNAc is included in the reaction to inhibit lysosomal hexosaminidases A and B which may be present in the enzyme preparation. O-GlcNAcase is not inhibited by 50 mM GalNAc.
4. Mix well and cover. 5. Incubate at 37°C for 30 min to 4 hr. Yellow color will develop as pNP-GlcNAc is hydrolyzed by O-GlcNAcase. Reactions should be optimized to keep the absorbance within the linear range of the spectrophotometer. The authors find that 20 to 50 ìg of cell extract used in a reaction of 100 ìl, with a 1 to 2 hr incubation time is appropriate.
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6. At the end of incubation, add an equal volume of 500 mM Na2CO3 to each well (100 µl or 500 µl). Na2CO3 raises the pH to >pH 9.0, intensifying the yellow color and stopping the reaction, as O-GlcNAcase has little activity at pH 9 to 10.
7. Read the absorbance at 400 nm on a plate reader or spectrophotometer. 8. Calculate O-GlcNAcase activity according to the following equation: mM of GlcNAc released = A400/(17.4 × 10 mM−1 ⋅ cm−1 × pathlength) One unit is the amount of enzyme catalyzing the release of 1 ìmol/min of pNP from pNP-GlcNAc. The molar extinction coefficient for pNP is 17.4 × 10 mM−1⋅cm−1 at pH 10. The path length for 200 ìl on a 96-well plate is 0.71 cm.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Biosynthetic labeling medium Glucose-free culture medium containing: 50 µCi/ml D-[6-3H]glucosamine (22 Ci/mmol; Amersham Pharmacia Biotech) 10% (v/v) FBS Prepare fresh Buffer H 50 mM HEPES, pH 6.8 50 mM NaCl 2% (v/v) Triton X-100 Store up to 1 month at room temperature Citrate-phosphate buffer, pH 4.0, 2× Dissolve 12.9 g citric acid monohydrate (mol. wt. 210) and 20.6 g disodium hydrogen phosphate heptahydrate (Na2HPO4⋅6H2O) in 300 ml Milli-Q water. Bring volume to 500 ml. Galactosyltransferase labeling buffer, 10× 100 mM HEPES, pH 7.5 100 mM galactose 50 mM MnCl2 Store up to 1 month at 4°C Galactosyltransferase storage buffer 2.5 mM HEPES, pH 7.4 2.5 mM MnCl2 50% (v/v) glycerol Store up to 1 month at room temperature
Detection and Analysis of Proteins Modified by O-Linked NAcetylglucosamine
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Hexosaminidase reaction mixture, 2× Per reaction: 25 µl 2× citrate-phosphate buffer (see recipe) 1 U N-acetyl-β-D-glucosaminidase (V-Labs) 0.01 U aprotinin 1 µg leupeptin 1 µg α2-macroglobulin O-GlcNAcase assay buffer, 10× 500 mM sodium cacodylate, pH 6.4 500 mM N-acetylgalactosamine (GlcNAc) 3% (w/v) bovine serum albumin (BSA) Prepare fresh OGT assay buffer, 10× 500 mM sodium cacodylate, pH 6.0 10 mg/ml bovine serum albumin (BSA) 10 mM 1-amino-GlcNAc (2-acetamido-1-amino-1,2-dideoxy-β-D-glucopyranose; Sigma) Prepare fresh OGT desalting buffer 20 mM Tris⋅Cl, pH 7.8 (APPENDIX 2E) 1 mg/ml bovine serum albumin (BSA) 20% (v/v) glycerol 0.02% (w/v) NaN3 Store up to 1 week at 4°C Protease inhibitors, 1000× PIC 1, 1000×: Dissolve the following in 10,000 U/ml aprotinin solution (Sigma) 1 mg/ml leupeptin 2 mg/ml antipain 10 mg/ml benzamide PIC 2, 1000×: Prepare in DMSO 1 mg/ml chemostatin 2 mg/ml pepstatin PMSF, 1000×: 0.1 M phenylmethylsulfonyl fluoride in 95% ethanol Sequencing reagent 7 ml methanol (HPLC-grade) 1 ml triethylamine (TEA; sequencing grade) 1 ml phenylisothiocyanate (PITC; sequencing grade) 1 ml Milli-Q water Stable 24 hr at 4°C sWGA Gal elution buffer Phosphate-buffered saline (PBS; APPENDIX 2E) containing: 0.2% (v/v) NP-40 1 M D-(+)-galactose (Gal) Store up to 1 week at 4°C Post-Translational Modification: Glycosylation
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sWGA GlcNAc elution buffer Phosphate-buffered saline (PBS; APPENDIX 2E) containing: 0.2% (v/v) NP-40 1 M N-acetylglucosamine (GlcNAc) Store up to 1 week at 4°C TBS-HD 10 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 150 mM NaCl 1% (v/v) Triton X-100 0.1% (w/v) SDS 0.25% (w/v) deoxycholic acid Store up to 1 month at 4°C TBS-HT 10 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 150 mM NaCl 0.3% (v/v) Tween 20 Store up to 1 month at room temperature TBST 10 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 150 mM NaCl 0.05% (v/v) Tween 20 Store up to 1 month at room temperature Tris-buffered saline (TBS) 10 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 150 mM NaCl Store up to 1 month at room temperature COMMENTARY Background Information
Detection and Analysis of Proteins Modified by O-Linked NAcetylglucosamine
(β)-D-1-4-galactosylaminyltransferase from bovine milk recognizes terminal N-acetylglucosamine (GlcNAc) residues and modifies them by the addition of a single Gal residue. Torres and Hart (1984) first used this enzyme in combination with UDP-[3H]Gal to demonstrate that bovine lymphocytes contain proteins modified by O-linked GlcNAc. Further refinements of this experiment led them to propose that the product, βGal1-4βGlcNAc, was the result of the galactosyltransferase recognizing and modifying a single GlcNAc residue Olinked to Ser/Thr residues of nuclear and cytoplasmic proteins (Holt and Hart, 1986). Since this report, many cytosolic and nuclear proteins from mammalian cells were shown to be modified by O-GlcNAc. This method has remained the “gold standard” technique for the detection of O-GlcNAc-modified proteins, as the label provides a “tag” for subsequent analyses, such as those described under Product Characterization, below (see Critical Parameters and Troubleshooting).
In subsequent years, methods such as WGA affinity and western blotting with GlcNAc-specific lectins and antibodies have become popular as simple techniques for the initial characterization of target proteins.
Critical Parameters and Troubleshooting Extraction of proteins from cells The O-GlcNAc modification can be removed from proteins either by cytosolic OGlcNAcase or lysosomal hexosaminidases. The inclusion of inhibitors during the extraction and purification process will preserve the levels of O-GlcNAc on proteins. Commonly used inhibitors (Dong and Hart, 1994) include 1-amino-GlcNAc (1 mM), GlcNAc (100 mM), and PUGNAc (5 µM). Note that these may have to be removed, as they will act as inhibitors in other methods.
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Product characterization Product characterization is a critical step showing that a protein is modified by OGlcNAc, and not other glycans. While many proteins modified by O-GlcNAc have been identified, there is evidence based on metabolic labeling (Medina et al., 1998) and lectin labeling studies (Hart et al., 1989) that indicate that O-GlcNAc is not the only intracellular carbohydrate post-translational modification. In addition, at least one peptide mimic of O-GlcNAc has been identified in cytokeratins (Shikhman et al., 1994). Moreover, many techniques used for breaking open cells also release proteins that are modified by complex N- and O-linked sugars, which may contain terminal GlcNAc. Many of the techniques described in this unit will recognize any GlcNAc residue, and it is important to perform the described controls such as PNGase F digestion to show specificity. Product analysis is critical for metabolic labeling with glucosamine. While UDPGlcNAc is the major product, glucosamine can enter other biosynthetic pathways, such as those used for amino acid synthesis. This issue was highlighted by studies of the SV40 large T-antigen. Some researchers have found that the SV40 large T-antigen labels with a number of different tritiated carbohydrates. However, O-GlcNAc is the only carbohydrate post-translational modification of the SV40 large T-antigen. The incorporation of glucosamine into amino acid biosynthetic pathways could be reduced by growing cells in the presence of excess nonessential amino acids (Medina et al., 1998). Lastly, while galactosyltransferase is specific for terminal GlcNAc residues, researchers (Elling et al., 1999) have shown that galactosyltransferase will modify GlcNAc linked in either the α- or β-anomeric conformation. The authors of this unit have shown that proteins modified by α-O-GlcNAc will be labeled using the procedure described (N. Zachara, unpub. observ.). While α-O-GlcNAc has not been identified in complex eukaryotes, it is a common modification of cell surface proteins of simple eukaryotes such as trypanosomes and Dictyostelium. Product analysis, such as HPAEC of the sugar alditols, will resolve many of the issues discussed.
Time Considerations Detection of O-GlcNAc proteins using antibodies (see Basic Protocol 2) and lectins (see Basic Protocol 3) will take approximately 2 to
3 days after the extraction of the proteins from cells. Samples and controls must be treated with PNGase F and/or hexosaminidase (1 hr to overnight), before SDS-PAGE and blotting. In either case, overnight incubation at 4°C provides the best signal-to-noise ratio. WGA affinity chromatography of lowcopy-number proteins will take 2 to 3 days. The ITT and WGA affinity chromatography can be completed in 1 day; subsequent analysis of the product by SDS-PAGE will take 1 to 2 days depending on the label used and the amount of label incorporated into proteins eluting from the WGA-agarose. Autogalactosylation of the galactosyltransferase and subsequent analysis of the activity will take 1 to 2 days. As the enzyme is stable for 6 to 12 months at −20°C, autogalactosylation does not need to be repeated for each analysis. Labeling of the proteins can take several hours to overnight, though optimization of the conditions may take a few days. The subsequent analysis, as well as desalting (dependent on the technique used), PNGase F digestion (1 hr to overnight), precipitation of protein (3 hr to overnight), SDS-PAGE, and autoradiography (1 to 10 days), can take up to 2 weeks. The length of time allotted to product analysis is dependent on the methods chosen, but will almost certainly require 7 to 10 days. Further analysis, including digestion of labeled proteins and subsequent purification of peptides, will take at least 3 days.
Literature Cited Amano, J. and Kobata, A. 1990. Quantitative conversion of mucin-type sugar chains to radioactive oligosaccharides. Methods Enzymol. 179:261-270. Batteiger, B., Newhall, W.J., and Jones, R.B. 1982. The use of Tween 20 as a blocking agent in the immunological detection of proteins transferred to nitrocellulose membranes. J. Immunol. Methods 55:297-307. Brew, K., Vanaman, T.C., and Hill, R.L. 1968. The role of alpha-lactalbumin and the A protein in lactose synthetase: A unique mechanism for the control of a biological reaction. Proc. Natl. Acad. Sci. U.S.A. 59:491-497. Comer, F.I. and Hart, G.W. 2000. O-Glycosylation of nuclear and cytosolic proteins: Dynamic interplay between O-GlcNAc and O-phosphate. J. Biol. Chem. 275:29179-29182. Comer, F.I., Vosseller, K., Wells, L., Accavitti, M.A., and Hart, G.W. 2001. Characterization of a mouse monoclonal antibody specific for Olinked N-acetylglucosamine. Anal. Biochem. 293:169-177.
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Dong, D.L. and Hart, G.W. 1994. Purification and characterization of an O-GlcNAc selective Nacetyl-β-D-glucosaminidase from rat spleen cytosol. J. Biol. Chem. 269:19321-19330. Duk, M., Ugorski, M., and Lisowska, E. 1997. Betaelimination of O-glycans from glycoproteins transferred to immobilon P membranes: Method and some applications. Anal. Biochem. 253:98102. Elling, L., Zervosen, A., Gallego, R.G., Nieder, V., Malissard, M., Berger, E.G., Vliegenthart, J.F., and Kamerling, J.P. 1999. UDP-N-Acetyl-alphaD-glucosamine as acceptor substrate of beta-1,4galactosyltransferase: Enzymatic synthesis of UDP-N-acetyllactosamine. G lycoco n. J. 16:327-336. Fukuda, M. 1990. Characterization of O-linked saccharide structures from cell surface glycoproteins. Methods Enzymol. 179:17-29. Gao, Y., Parker, G.J., and Hart, G.W. 2000. Streptozotocin-induced β-cell death is independent of its inhibition of O-GlcNAcase in pancreatic Min6 cells. Arch. Biochem. Biophys. 383:296302. Gao, Y., Wells, L., Comer, F.I., Parker, G.J., and Hart, G.W. 2001. Dynamic O-glycosylation of nuclear and cytosolic proteins: Cloning and characterization of a neutral, cytosolic β-N-acetylglucosaminidase from human brain. J. Biol. Chem. 276:9838-9845. Greis, K.D. and Hart, G.W. 1998. Analytical methods for the study of O-GlcNAc glycoproteins and glycopeptides. Methods Mol. Biol. 76:19-33. Greis, K.D., Hayes, B.K., Comer, F.I., Kirk, M., Barnes, S., Lowary, T.L., and Hart, G.W. 1996. Selective detection and site-analysis of OGlcNAc-modified glycopeptides by beta-elimination and tandem electrospray mass spectrometry. Anal. Biochem. 234:38-49. Haltiwanger, R.S., Blomberg, M.A., and Hart, G.W. 1992. Glycosylation of nuclear and cytoplasmic proteins: Purification and characterization of a uridine diphospho-N-acetylglucosamine:polypeptide β-N-acetylglucosaminyltransferase. J. Biol. Chem. 267:9005-9013. Haltiwanger, R.S., Grove, K., and Philipsberg, G.A. 1998. Modulation of O-linked N-acetylglucosamine levels on nuclear and cytoplasmic proteins in vivo using the peptide O-GlcNAc-β-Nacetylglucosaminidas e inhibitor O-(2acetamido-2-deoxy-D-glucopyranosylidene) amino-N-phenylcarbamate. J. Biol. Chem. 273:3611-3617. Han, I., Oh, E.-S., and Kudlow, J.E. 2000. Responsiveness of the state of O-linked N-acetylglucosamine modification of nuclear pore protein p62 to the extracellular glucose concentration. Biochem. J. 350:109-114. Detection and Analysis of Proteins Modified by O-Linked NAcetylglucosamine
Hardy, M.R. and Townsend, R.R. 1994. High-pH anion exchange chromatography of glycoprotein-derived carbohydrates. Methods Enzymol. 230:208-225.
Hart, G.W., Holt, G.D., Haltiwanger, R.S., and Kelly, W.G. 1989. Cytoplasmic and nuclear glycosylation. Annu. Rev. Biochem. 58:841-874. Hart, G.W., Cole, R.N., Kreppel, L.K., Arnold, C.S., Comer, F.I., Iyer, S., Cheng, X., Carroll, J., and Parker, G.J. 2000. Glycosylation of proteins: A major challenge in mass spectrometry and proteomics. In Proceedings of the 4th International Symposium on Mass Spectrometry in the Health and Life Sciences (A. Burlingame, S. Carr, and M. Baldwin, eds.) pp. 365-382. Humana Press, Totowa, N.J. Hayes, B.K., Greis, K.D., and Hart, G.W. 1995. Specific isolation of O-linked N-acetylglucosamine glycopeptides from complex mixtures. Anal. Biochem. 228:115-122. Haynes, P.A. and Aebersold, R. 2000. Simultaneous detection and identification of O-GlcNAc-modified glycoproteins using liquid chromatographymass spectrometry. Anal. Chem. 72:5402-5410. Heart, E., Choi, W.S., and Sung, C.K. 2000. Glucosamine-induced insulin resistance in 3T3-L1 adipocytes. Am. J. Physiol. Endocrinol. Metab. 278:E103-E112. Holt, G.D. and Hart, G.W. 1986. The subcellular distribution of terminal N-acetylglucosamine moieties: Localization of a novel protein-saccharide linkage, O-linked GlcNAc. J. Biol. Chem. 261:8049-8057. Holt, G.D., Snow, C.M., Senior, A., Haltiwanger, R.S., Gerace, L., and Hart, G.W. 1987. Nuclear pore complex glycoproteins contain cytoplasmically disposed O-linked N-acetylglucosamine. J. Cell Biol. 104:1157-1164. Izumi, T. and Suzuki, K. 1983. Neutral β-N-Acetylhexosaminidases of rat brain: Purification and enzymatic and immunological characterization. J. Biol. Chem. 258:6991-6999. Kelly, W.G., Dahmus, M.E., and Hart, G.W. 1993. RNA polymerase II is a glycoprotein: Modification of the COOH-terminal domain by OGlcNAc. J. Biol. Chem. 268:10416-10424. Kobata, A. 1994. Size fractionation of oligosaccharides. Methods Enzymol. 230:200-208. Kreppel, L.K. and Hart, G.W. 1999. Regulation of a cytosolic and nuclear O-GlcNAc transferase: Role of the tetratricopeptide repeats. J. Biol. Chem. 274:32015-32022. Kreppel, L.K., Blomberg, M.A., and Hart, G.W. 1997. Dynamic glycosylation of nuclear and cytosolic proteins: Cloning and characterization of a unique O-GlcNAc transferase with multiple tetratricopeptide repeats. J. Biol. C hem. 272:9308-9315. Medina, L., Grove, K., and Haltiwanger, R.S. 1998. SV40 large T antigen is modified with O-linked N-acetylglucosamine but not with other forms of glycosylation. Glycobiology 8:383-391. Reason, A.J., Morris, H.R., Panico, M., Marais, R., Treisman, R.H., Haltiwanger, R.S., Hart, G.W., Kelly, W.G., and Dell, A. 1992. Localization of O-GlcNAc modification on the serum response
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transcription factor. J. Biol. Chem. 267:1691116921. Roos, M.D., Xie, W., Su, K., Clark, J.A., Yang, X., Chin, E., Paterson, A.J., and Kudlow, J.E. 1998. Streptozotocin, an analog of N-acetylglucosamine, blocks the removal of O-GlcNAc from intracellular proteins. Proc. Assoc. Am. Physicians 110:422-432. Roquemore, E.P., Chou, T.Y., and Hart, G.W. 1994. Detection of O-linked N-acetylglucosamine (OGlcNAc) on cytoplasmic and nuclear proteins. Methods Enzymol. 230:443-460. Schnedl, W.J., Ferber, S., Johnson, J.H., and Newgard, C.B. 1994. STZ transport and cytotoxicity: Specific enhancement in GLUT-2 expressing cells. Diabetes 43:1326-1333. Shafi, R., Iyer, S.P., Ellies, L.G., O’Donnell, N., Marek, K.W., Chui, D., Hart, G.W., and Marth, J.D. 2000. The O-GlcNAc transferase gene resides on the X chromosome and is essential for embryonic stem cell viability and mouse ontogeny. Proc. Natl. Acad. Sci. U.S.A. 97:57355739. Shikhman, A.R., Greenspan, N.S., and Cunningham, M.W. 1994. Cytoker atin peptide SFGSGFGGGY mimics N-acetyl-beta-D-glucosamine in reaction with antibodies and lectins, and induces in vivo anti-carbohydrate antibody response. J. Immunol. 153:5593-5606. Snow, C.M., Senior, A., and Gerace, L. 1987. Monoclonal antibodies identify a group of nuclear pore complex glycoproteins. J. Cell Biol. 104:11431156. Torres, C.R. and Hart, G.W. 1984. Topography and polypeptide distribution of terminal N-acetylglucosamine residues on the surfaces of intact lymphocytes. Evidence for O-linked GlcNAc. J. Biol. Chem. 259:3308-3317.
Townsend, R.R., Hardy, M.R., and Lee, Y.C. 1990. Separation of oligosaccharides using high-performance anion-exchange chromatography with pulsed amperometric detection. Methods Enzymol. 179:65-76. Turner, J.R., Tartakoff, A.M., and Greenspan, N.S. 1990. Cytologic assessment of nuclear and cytoplasmic O-linked N-acetylglucosamine distribution by using anti-streptococcal monoclonal antibodies. Proc. Natl. Acad. Sci. U.S.A. 87:56085612. Unverzagt, C., Kunz, H., and Paulson, J.C. 1990. High-efficiency synthesis of sialyloligosaccharides and sialoglycopeptides. J. Am. Chem. Soc. 112:9308-9309. Varki, A. 1994. Metabolic radiolabeling of glycoconjugates. Methods Enzymol. 230:16-32. Wells, L., Vosseller, K., and Hart, G.W. 2001. Glycosylation of nucleocytoplasmic proteins: Signal transduction and O-GlcNAc. Science 291:23762378. Zachara, N.E. and Gooley, A.A. 2000. Identification of glycosylation sites in mucin peptides by Edman degradation. Methods Mol. Biol. 125:121128.
Contributed by Natasha E. Zachara, Robert N. Cole, and Gerald W. Hart The Johns Hopkins University Medical School Baltimore, Maryland Yuan Gao Deakin University Victoria, Australia
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CHAPTER 13 Post-Translational Modification: Phosphorylation and Phosphatases INTRODUCTION
A
polypeptide sequence deduced from a nucleic acid sequence tells only part of a protein’s story: covalent modifications often play a key role in determining the final properties of a protein. Some modifications may be relatively constant over time; immediately obvious in this category is the important role played by disulfide bonds, which stabilize the folded structure of a protein and whose presence cannot be deduced from the sequence. Other modifications (e.g., N-linked glycosylation, covered in Chapter 12) may also affect protein properties in a relatively static sense by controlling (at least in part) solubility, antigenic structure, and, on occasion, the ability to interact with appropriate partner molecules. In other cases post-translational modifications are more dynamic: these are often of critical importance to protein function, directly controlling enzymatic activity or the ability to interact with partner molecules. Such modifications often take place on a time scale of minutes or even seconds. Foremost in this category is phosphorylation: changes in the phosphorylation state control the activity of a wide range of polypeptides, from transcription factors to soluble enzymes and cell-surface receptors (see UNIT 13.1 for an overview of this topic).
The ability to detect the presence of phosphorylated amino acids and changes in phosphorylation status is essential in the study of phosphoprotein function. A method for labeling cells with inorganic phosphate, usually the first step in establishing whether a given polypeptide is phosphorylated in a particular cell type, is provided in UNIT 13.2. It is also important to distinguish among the different phosphoaminoacids. Methods for distinguishing between P-Ser, P-Thr, and P-Tyr are presented in UNIT 13.3. Phosphoproteins containing phosphotyrosine (but not other phosphoamino acids) can be detected by immunological means; this powerful and convenient technique is described in UNIT 13.4. Kinase reactions permit the addition of phosphate residues to polypeptides; it should also be kept in mind that the reverse reaction may be equally important as a means of controlling the activity of phosphoproteins. Similarly, phosphatases may be used analytically to demonstrate the presence of phosphorylated amino acid residues in phosphoproteins; this approach is presented in UNIT 13.5. Production of antiphosphopeptide antibodies (i.e., antibodies that recognize phosphorylated peptides) is described in UNIT 13.6. This technology is an important tool for analysis of a protein’s functional state, as these antibodies have unique specificity toward the cognate phosphorylated proteins and will not cross-react with unphosphorylated proteins or other phosphoproteins. The assays for protein kinases are manifold and can include a variety of substrates. UNIT provides the strategies and protocols for assaying protein kinases using exogenous substrates. While the purpose of the unit is not to catalog all combinations of kinases and substrates, it provides specific examples of protein kinase assays that will assist in optimizing conditions for individual protein kinases. 13.7
Contributed by Hidde L. Ploegh, Ben M. Dunn, and John E. Coligan Current Protocols in Protein Science (2003) 13.0.1-13.0.2 Copyright © 2003 by John Wiley & Sons, Inc.
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In many cases, the activities of kinases are contingent on the cellular context in which such activities are measured and on interactions with other proteins in the cell. In some cases prior activation of cell surface receptors is required to achieve full activation of the kinase(s) of interest and may determine the outcome of measuring the activity or the biochemical consequences of such kinase activity. Experiments with purified kinases may not accurately reflect these conditions, or the kinase(s) responsible may not be available in purified form. Under these circumstances, it is highly advantageous to conduct activity measurements in semi-intact or permeabilized cells or in subcellular fractions derived from them. UNIT 13.8 describes different strategies for studying protein phosphorylation in permeabilized cells. Precise control of biological function often requires phosphorylation of specific residues in a protein sequence. The identification of the site of phosphorylation within a protein through enzymatic digestion, gel electrophoresis, manual or automated sequencing, or mass spectroscopy is described in UNIT 13.9. UNIT 13.10 describes the use of protein phosphatase inhibitors to prevent the turnover of protein-bound phosphate intracellularly or during protein isolation procedures, thereby enhancing the detection of phosphoproteins by standard techniques such as SDS-PAGE (UNIT 10.1) followed by imaging through immunoblotting (UNIT 13.4). Consequently, these procedures have the potential to enhance detection of sites of phosphorylation as described in UNIT 13.9. Hidde L. Ploegh, Ben M. Dunn, and John E. Coligan
Introduction
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Overview of Protein Phosphorylation HISTORY Phosphorylation is the most common and important mechanism of acute and reversible regulation of protein function. Studies of mammalian cells metabolically labeled with [32P]orthophosphate suggest that as many as one-third of all cellular proteins are covalently modified by protein phosphorylation. Protein phosphorylation and dephosphorylation function together in signal transduction pathways to induce rapid changes in response to hormones, growth factors, and neurotransmitters. Most polypeptide growth factors (platelet-derived growth factor and epidermal growth factor are among the best studied; Heldin, 1995) and cytokines (e.g., interleukin 2, colony stimulating factor 1, and γ-interferon; Ihle et al., 1994) stimulate phosphorylation upon binding to their receptors. Induced phosphorylation in turn activates cytoplasmic protein kinases, such as raf, MEK, and MAP kinases (Marshall, 1995). Additionally, in all nucleated organisms, cell cycle progression is regulated at both the G1/S and the G2/M transitions by cyclin-dependent protein kinases (Doree and Galas, 1994). Differentiation and development are also controlled by phosphorylation. Development of the R7 cell in the Drosophila retina (Simon, 1994) and of the vulva in Caenorhabditis elegans (Eisenmann and Kim, 1994) are both dependent on the function of receptor and cytoplasmic protein kinases. Finally, metabolism—in particular, the interconversion of glucose and glycogen and the transport of glucose—is regulated by phosphorylation (Cohen, 1985). Biologists of all stripes therefore find, often unexpectedly and occasionally reluctantly, that they must study protein phosphorylation in order to understand the regulation and function of their favorite gene and its product. In the early 1940s, soon after their discovery that glycogen phosphorylase existed in two distinct forms in skeletal muscle, Carl and Gerty Cori identified a cellular activity that converted active phosphorylase a into a less active phosphorylase b. Initial studies of what they called the prosthetic-group removing (PR) enzyme suggested that the inactivation of phosphorylase a was mediated via proteolysis (Cori and Cori, 1945). More than ten years later, this converting enzyme was shown to be a protein phosphatase. Thus, a protein phosphatase was studied nearly 20 years prior to the identifica-
tion of the first protein kinase. The finding that phosphorylase phosphatases were more widely expressed in tissues than phosphorylase a led to the current realization that these enzymes have a broad substrate specificity and regulate many physiological processes. The use of substrates other than phosphorylase a also identified several other protein serine/threonine phosphatases.
LABELING STUDIES Protein phosphorylation is usually studied by biosynthetic labeling with 32P-labeled inorganic phosphate (32Pi; UNIT 13.2). This is intrinsically quite simple—the label is just added to growth medium. It is this step of an experiment, however, that makes many investigators the most nervous, given the perceived danger of radioactive exposure and the real danger of contamination of laboratory equipment with radioactivity. Neither problem is insurmountable. With proper shielding and technique, exposure of the investigator can be limited to the hands and contamination of the laboratory can be avoided. A general protocol for biosynthetic labeling with 32Pi that maximizes incorporation and minimizes radioactive exposure of workers in the lab and contamination of lab equipment is described in UNIT 13.2. (For a general discussion of radiation safety consult Safe Use of Radioisotopes, APPENDIX 2B.) Many protein kinases can be assayed by the transfer of radiolabel from [32P]ATP to surrogate protein and peptide substrates. Synthetic peptide substrates have been particularly useful in establishing consensus motifs that define the substrate for protein kinases. By comparison, protein phosphatases show very low activity against synthetic phosphopeptides, greatly preferring the phosphoprotein substrates (Shenolikar and Ingebritsen, 1984). Therefore, attempts to define structural requirements for protein dephosphorylation using synthetic phosphopeptides have failed to establish the molecular basis for substrate specificity of the major cellular protein phosphatases. The rate of 32P incorporation into proteins in vivo, at either basal or hormone-stimulated levels, depends heavily on the rate of turnover of phosphate in the protein. For example, low phosphatase activity against specific sites or proteins will reduce the turnover of proteinbound phosphates and hinder their metabolic labeling in the intact cell. Conversely, physi-
Contributed by Bartholomew M. Sefton and Shirish Shenolikar Current Protocols in Protein Science (1995) 13.1.1-13.1.5 Copyright © 2000 by John Wiley & Sons, Inc.
UNIT 13.1
Post-Translational Modification: Phosphorylation and Phosphatases
13.1.1 CPPS
ological stimuli that increase phosphatase activity may facilitate metabolic labeling of proteins, allowing rapid replacement of endogenous “cold” phosphate with the radiolabel. Thus, 32P incorporation by itself gives no indication about the extent of phosphorylation of a particular protein. As most phosphoproteins are phosphorylated on multiple sites, this also makes it difficult to correlate changes in protein function with 32P labeling of a given protein in the intact cell.
SITES OF PHOSPHORYLATION Most proteins are found to be phosphorylated at serine or threonine residues, and many proteins involved in signal transduction are also phosphorylated at tyrosine residues. These three hydroxyphosphoamino acids exhibit sufficient chemical stability at acidic pH that they can be recovered after acid hydrolysis and identified in a straightforward manner. Proteins that contain covalently bound phosphate at histidine, cysteine, and aspartic acid residues, either as phosphoenzyme intermediates or as stable modifications, have also been described. Each of these phosphoamino acids is chemically labile and impossible to study with the standard techniques used for the acid-stable phosphoamino acids. Indeed, they are often identified by inference or elimination. A technique for identifying phosphoserine, phosphothreonine, and phosphotyrosine by acid hydrolysis and two-dimensional thin-layer electrophoresis is described in UNIT 13.3. Techniques for analyzing acid-labile forms of protein phosphorylation are described in Ringer, 1991; Kamps, 1991; and Duclos et al., 1991. Phosphotyrosine is not an abundant phosphoamino acid. Therefore, its detection in samples labeled with 32Pi is often difficult, especially if the samples contain large quantities of proteins phosphorylated at serine residues or are contaminated with RNA. Detection of phosphotyrosine, as well as of phosphothreonine, can be enhanced considerably by incubation of gel-fractionated samples in alkali. This hydrolyzes RNA and dephosphorylates phosphoserine, allowing visualization of minor tyrosineand threonine-phosphorylated proteins. A simple procedure for alkaline treatment is described in the alternate protocol of UNIT 13.3.
DETECTION OF UNLABELED PHOSPHOAMINO ACIDS Overview of Protein Phosphorylation
If a protein is modified by phosphorylation, identification of the phosphoamino acid can
often be accomplished without resorting to biosynthetic labeling. For example, tyrosine phosphorylation can be studied because proteins containing this rare phosphoamino acid can be detected with great specificity and sensitivity by antibodies to phosphotyrosine (UNIT 13.4). By comparison, the quantitation of phosphoserine and phosphothreonine is more difficult and requires partial acid hydrolysis of the phosphoprotein and subsequent separation of phosphoamino acids by high-voltage electrophoresis on thin-layer cellulose acetate plates. Attempts to generate antibodies that recognize phosphoserine or phosphothreonine have failed to produce reagents with the required specificity and/or sufficient sensitivity to be useful. However, once the primary sequence around a phosphorylation site containing phosphoserine or phosphothreonine has been determined, it is possible to make antibodies against synthetic phosphopeptides modeled on these phosphorylation sites (Czernik et al., 1991). Such antiphosphopeptide antibodies have been very useful tools for monitoring the phosphorylation of the parent protein at specific sites. The phospho-specific antibodies have also been used to inhibit the dephosphorylation of individual sites and demonstrate their role in protein function. Several recent studies have produced phospho-specific antibodies against phosphorylation sites predicted by the primary sequence of a protein. These reagents provide new insights into phosphorylations that control protein function. More generally, because phosphorylation often alters the mobility of a protein during SDS–polyacrylamide gel electrophoresis and almost always alters its isoelectric point, the presence of phosphorylated residues in an unlabeled protein can be deduced from altered gel mobility after incubation of the protein with a phosphatase. Most phosphorylation sites are thought to reside near the surface of phosphoproteins. Therefore, they are equally accessible to phosphatases and proteases. Indeed, limited proteolysis often produces functional changes in phosphoproteins that closely mimic their in vitro dephosphorylation by phosphatase. As monitoring the release of radioactivity from a 32P-labeled phosphoprotein does not distinguish between a phosphatase and a protease reaction, additional steps must to be taken to differentiate orthophosphate from small phosphopeptides. This is particularly important when dephosphorylation reactions are carried out in crude extracts (UNIT 13.5).
13.1.2 Current Protocols in Protein Science
PROTEIN KINASES Protein kinases exhibit a strict specificity for phosphorylation of either serine/threonine or tyrosine residues. Yet these enzymes share structural homology in several domains such as the ATP-binding and catalytic sites, emphasizing their common function as phosphotransferases. A third group of protein kinases more closely resembles the serine/threonine kinases in their primary sequence but phosphorylate both serine/threonine and tyrosine residues. MEK, a MAP kinase activator (Crews and Erickson, 1992) and wee1, an inhibitor of cdc2 kinase (Parker et al., 1992) are examples of such dual-specificity protein kinases that phosphorylate threonine and tyrosine residues which are closely located in substrate proteins. Kinases have been used in a number of ways to analyze protein phosphorylation. Mutations that abrogate ATP-binding inactive protein kinases. Such inactive kinases retain the ability to recognize substrates and therefore act as dominant-negative reagents to analyze the physiological function of kinases in the intact cell (Thorburn et al., 1994; Alberola-Ila et al., 1995). Cell-permeable compounds modeled on ATP also inhibit selected kinases in the intact cell. Growth factors promote dimerization and activation of transmembrane receptor kinases. Mutant receptors that can dimerize in the absence of the natural ligand offer a novel approach to activate receptor kinases and initiate the cellular responses to growth factors (Spencer et al., 1993). Several cytoplasmic kinases are also activated by binding of intracellular second messengers. The allosteric activation of these kinases results from conformation changes that eliminate internal autoinhibitory interactions which silence the kinases in the absence of ligand. Deletion of the autoinhibitory sequences generates a constitutively active kinase that provides important insights into its physiological role (Planas-Silva and Means, 1992). Synthetic peptides modeled on the autoinhibitory sequences can also be used to selectively inhibit these kinases and reveal their role in controlling protein phosphorylation. Still other kinases, which are not directly regulated by ligand binding, are activated by phosphorylation. Some of these kinases participate in cascades of sequential phosphorylations that amplify physiological signals. Substitution of the amino acids phosphorylated during the activation of these kinases with nonphosphorylated residues generates dominant-negative enzymes that can block the functions of such
kinases in intact cells. Alternately, acidic amino acids may be substituted for phosphoamino acids to produce a constitutively active kinase (Brunet et al., 1994). Such reagents have been used to validate the role of protein kinases in specific phosphorylation events.
PROTEIN PHOSPHATASES Many protein phosphatases also show a strict specificity for either phosphoserine/phosphothreonine or phosphotyrosine residues. However, unlike kinases, the serine/threonine and tyrosine phosphatases are not evolutionarily related and exhibit no primary sequence homology (Shenolikar and Nairn, 1991; Charbonneau and Tonks, 1992). Acid and alkali phosphatases can also dephosphorylate phosphoproteins in vitro but share no structural homology with protein phosphatases. A new family of phosphatases, such as cdc25 and CL100, are distantly related to the tyrosine phosphatases but dephosphorylate both phosphotyrosine and phosphothreonine in their target substrates. Mutation of an essential cysteine in tyrosine phosphatases has produced dominant-negative enzymes that have been shown to shield their substrates from dephosphorylation by endogenous phosphatases, thereby prolonging the biological effects of protein phosphorylation (Sun et al, 1993). Mutating several conserved amino acids can inactivate a serine/threonine phosphatase (Zhou et al, 1994), but the ability of such mutant phosphatases to modulate serine/threonine phosphorylation in cells has not yet been demonstrated. A number of potent phosphatase inhibitors have been identified in recent years. Okadaic acid and several other toxins inhibit protein (serine/threonine) phosphatase 1 and 2A, and vanadate and phenylarsenoxide inhibit tyrosine phosphatase. These compounds have implicated reversible phosphorylation as a regulatory mechanism in many physiological processes (Cohen, 1989; Hardie et al., 1991; Shenolikar and Nairn, 1991; Shenolikar, 1994); under certain conditions, they may be the dominant regulators of these cellular processes. These phosphatase inhibitors can also be used to distinguish protein dephosphorylation from proteolysis in crude tissue extracts (Cohen, 1991). However, by far the most important contribution of these reagents has been that they have allowed assessment of the role played by phosphorylation in cellular processes where neither the identity of the phosphoprotein involved nor that of the kinase(s) that regulates its function are known.
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13.1.3 Current Protocols in Protein Science
The units in this chapter describe techniques that detect protein phosphorylation and identify amino acids that have been covalently modified (UNITS 13.2, 13.3, & 13.4). The next step is to use phosphatases to dephosphorylate the substrate protein in vitro and address the functional relevance of the covalent modification (UNIT 13.5). Once the regulatory role of phosphorylation has been clearly established, the more sophisticated approaches discussed above can be used to to identify the specific kinases and phosphatases involved and thereby begin to elucidate the physiological mechanism that regulates the substrate in the intact cell.
Eisenmann, D.M. and Kim, S.K. 1994. Signal transduction and cell fate specification during Caenorhabditis elegans vulval development. Curr. Opin. Genet. Dev. 4:508-516.
LITERATURE CITED
Kamps, M.P. 1991. Determination of phosphoamino acid composition by acid hydrolysis of protein blotted to Immobilon. Methods Enzymol. 201:21-27.
Alberola-Ila, J., Forbush, K.A., Seger, R., Krebs, E.G., and Perlmutter, R.M. 1995. Selective requirement for MAP kinase activation in thymocyte differentiation. Nature 373:620-623. Brunet, A., Pages, G., and Poussegur, J. 1994. Constitutively active mutants of MAP (MEK1) induce growth factor-relaxation and oncogenicity when expressed in fibroblasts. Oncogene 9:3379- 3387. Charbonneau, H. and Tonks, N.K. 1992. 1002 protein phosphatases? Annu. Rev. Cell Biol. 8:463493. Cohen, P. 1985. The role of protein phosphorylation in the hormonal control of enzyme activity. Eur. J. Biochem. 15:439-448. Cohen, P. 1989. The structure and regulation of protein phosphatases. Annu. Rev. Biochem. 58:453-508. Cohen, P. 1991. Classification of protein serine/threonine phosphatases: Identification and quantitation in cell extracts. Methods Enzymol. 201:389-398.
Heldin, C.-H. 1995. Dimerization of cell surface receptors in signal transduction. Cell 80:213223. Ihle, J.N., Witthuhn, B.A., Quelle, F.W., Yamamoto, K., Thierfelder, W.E., Kreider, B., and Silvennoinen, O. 1994. Signalling by the cytokine receptor superfamily: JAKS and STATS. Trends Biochem. Sci. 19:222-227.
Marshall, C.J. 1995. Specificity of receptor tyrosine kinase signalling: Transient versus sustained extracellular signal-regulated kinase activation. Cell 80:179-185. Parker, L.L., Atherton-Fessler, S., and PiwinicaWorms, H. 1992. p107 wee1 is a dual-specificity kinase that phosphorylates p34cdc2 on tyrosine 15. Proc. Natl. Acad. Sci. U.S.A. 89:2917-2921. Planas-Silva, M.D. and Means, A.R. 1992. Expression of a constitutive form of calcium/calmodulin-dependent protein kinase II leads to arrest of the cell cycle in G2. EMBO J. 11:507517. Ringer, D.P. 1991. Separation of phosphotyrosine, phosphoserine, and phosphothreonine by highperformance liquid chromatography. Methods Enzymol. 201:3-10. Shenolikar, S. 1994. Protein serine/threonine phosphatases: New avenues for cell regulation. Annu. Rev. Cell Biol. 10:55-86.
Cori, G.T. and Cori, C.F. 1945. Enzymatic conversion of phosphorylase a to b. J. Biol. Chem. 158:321-345.
Shenolikar, S. and Ingebritsen, T.S. 1984. Protein (serine, threonine) phosphate phosphatases. Methods Enzymol. 107:102-130.
Crews, C.M. and Erickson, R.L. 1992. Purification of a murine protein-tyrosine/threonine kinase that phosphorylates and activates the erk-1 gene product: Relationship to the fission yeast byr1 gene product. Proc. Natl. Acad. Sci. U.S.A. 89:8205-8209.
Shenolikar, S. and Nairn, A.C. 1991. Protein phosphatases: Recent progress. Adv. Second Messenger Phosphoprotein Res. 23:1-121.
Czernik, A.J., Girault, J.A., Nairn, A.C., Chen, J., Snyder, G., Kebabian, J., and Greengard, P. 1991. Production of phosphorylation state–specific antibodies. Methods Enzymol. 201:264-283. Doree, M. and Galas, S. 1994. The cyclin-dependent protein kinases and the control of cell division. FASEB J. 85:1114-1121.
Overview of Protein Phosphorylation
Hardie, D.G., Haystead, T.A.J., and Sim, A.T.R. 1991. Use of okadaic acid to inhibit protein phosphatases in intact cells. Methods Enzymol. 201:469-477.
Duclos, B., Marcandier, S., and Cozzone, A.J. 1991. Chemical properties and separation of phosphoamino acids by thin-layer chromatography and/or electrophoresis. Methods Enzymol. 201:10-21.
Simon, M. 1994. Signal transduction during the development of the Drosophila R7 photoreceptor. Dev. Biol. 166:431-442. Spencer, D.M., Wandless, T.J., Schreiber, S.L., and Crabtree, G.R. 1993. Controlling signal transduction with synthetic ligands. Science 262:1019-1024. Sun, H., Charles, C.H., Lau, L.F., and Tonks, N.K. 1993. MKP-1 (3CH134), an immediate early gene product, is a dual-specificity phosphatase that dephosphorylates MAP kinase in vivo. Cell 75:487- 493.
13.1.4 Current Protocols in Protein Science
Thorburn, J., McMahon, M. and Thorburn, A. 1994. Raf-1 kinase activity is necessary and sufficient for gene expression changes but not sufficient for cellular morphology changes associated with cardiac myocyte hypertrophy. J. Biol. Chem. 269: 30580-30586. Zhou, S., Clemens, J.C., Stone, R.L., and Dixon, J.E. 1994. Mutational analysis of a ser/thr phosphatase. Identification of residues important in phosphatase substrate binding and catalysis. J. Biol. Chem. 269:26234-26238.
Contributed by Bartholomew M. Sefton The Salk Institute San Diego, California Shirish Shenolikar Duke University Medical Center Durham, North Carolina
Post-Translational Modification: Phosphorylation and Phosphatases
13.1.5 Current Protocols in Protein Science
32
Labeling Cultured Cells with Pi and Preparing Cell Lysates for Immunoprecipitation
UNIT 13.2
This unit describes 32Pi labeling and lysis of cultured cells to be used for subsequent immunoprecipitation of proteins. The procedure is suitable for insect, avian, and mammalian cells and can be used with both adherent and nonadherent cultures. The same general approach—biosynthetic labeling with 32Pi in medium containing a reduced concentration of phosphate—can also be applied to bacteria and yeast; however, specific techniques to accomplish this are not presented here. This approach can also be used to label any cellular constituents with 32P; in these cases the Basic Protocol must be modified beginning with step 8. The first procedure described (see Basic Protocol) is 32Pi labeling of adherent or nonadherent (e.g., hematopoietic) cells with subsequent lysis in a detergent buffer containing either Nonidet P-40 (NP-40), 3[(3-cholamidopropyl)-dimethylammonio]-1-propane-sulfonate (CHAPS), or RIPA (RadioImmunoPrecipitation Assay buffer), a combination of NP-40, sodium deoxycholate, and SDS. More rigorous lysis conditions to be used for working with proteins that are difficult to solubilize are also described (see Alternate Protocol). CAUTION: Unshielded 32P will penetrate ∼1 cm into flesh. Exposure to the skin and eyes is, therefore, of concern. Gloves and protective eyeware should always be worn when handling significant amounts of 32P. A 1-in.-thick (2.5-cm) Plexiglas shield, tall enough to look through when seated or standing comfortably, should be used when handling samples containing 32P. LABELING CULTURED CELLS WITH 32Pi AND LYSIS USING MILD DETERGENT
BASIC PROTOCOL
The first six steps of this protocol describe culturing and labeling procedures for cultures of adherent cells; modifications appropriate for nonadherent cells are described in alternate steps. The remaining steps describe lysis of cells in detergent buffer to prepare the sample for immunoprecipitation. For a more detailed discussion of mammalian cell culture conditions and reagents, see APPENDIX 3C. Materials Cell culture to be labeled Labeling medium: phosphate-free tissue culture medium (e.g., DMEM; APPENDIX 3C) supplemented with the usual concentration of serum or serum dialyzed against phosphate-free saline, 37°C 500 mCi/ml H332PO4 in HCl (carrier-free; ICN) Tris-buffered saline (TBS, see recipe) or PBS (see recipe), cold Mild lysis buffer or RIPA lysis buffer (see recipes) 1-in.-thick Plexiglas shield (APPENDIX 2B) Plugged, aerosol-resistant pipet tips Plexiglas box (APPENDIX 2B), warmed to 37°C Screw-cap microcentrifuge tubes Plugged disposable pipet or disposable one-piece transfer pipet Rubber policeman
Contributed by Bartholomew M. Sefton Current Protocols in Protein Science (1997) 13.2.1-13.2.8 Copyright © 1997 by John Wiley & Sons, Inc.
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13.2.1 Supplement 10
Sorvall refrigerated centrifuge with SM-24 rotor and rubber adaptors, refrigerated microcentrifuge, or equivalent, 4°C Plexiglas sheet (10 × 10 × 1⁄4–in.) or Plexiglas tube holder, 4°C (APPENDIX 2B) Additional reagents and equipment for cell culture (APPENDIX 3C) and gel electrophoresis (UNIT 10.1) NOTE: All culture incubations are performed in a humidified 37°C, 10% CO2 incubator unless otherwise specified. Label the cell culture 1. Culture the cells to be labeled to an appropriate stage of growth. Phosphate transport is maximal in rapidly growing cells. Therefore, except in those cases where phosphorylation of a protein in quiescent cells is to be examined, cells to be labeled should be subconfluent (adherent cells) or at less than maximal density (nonadherent cells). It is useful to change the medium to fresh growth medium 3 to 18 hr prior to labeling. Brief cultivation of cells in low-phosphate medium (to reduce the phosphate pool by starvation prior to labeling) is of minimal value.
For adherent cells 2a. Remove growth medium by aspiration. Wash away any residual phosphate-containing medium by adding 37°C labeling medium supplemented with serum, but lacking the label, and removing the wash medium by aspiration. Phosphate-free DMEM is routinely used for labeling medium because RPMI has a very high concentration of phosphates.
3a. Add prewarmed labeling medium to cultures, using 0.5 to 1 ml per 35-mm dish, 1 to 2 ml per 50-mm dish, or 2 to 4 ml per 100-mm dish of adherent cells. For nonadherent cells 2b. Gently centrifuge the culture 1 min at 1800 × g and aspirate the medium away from the cell pellet. Resuspend the cells in labeling medium supplemented with serum but lacking the label. Centrifuge again for 1 min at 1800 × g and remove the medium. 3b. Add 2 ml prewarmed labeling medium per 107 cells and transfer to an appropriate size petri dish. 4. Working behind a Plexiglas shield, use a micropipettor with plugged, aerosol-resistant pipet tips to add 32Pi to a final concentration of 0.1 to 2 mCi/ml. Use of plugged, aerosol-resistant pipet tips will minimize contamination of the micropipettor. 32Pi is usually shipped in HCl and is generally sterile, so it is not necessary to sterilize the labeling medium after adding the radioisotope. Additionally, except in the rare cases when steady-state labeling with 32P is desired, the labeling interval is usually so short (<6 hr) that microbial growth resulting from added isotope is undetectable. Therefore, addition of 32P to cells can be performed on the lab bench rather than in a tissue culture hood.
5. Place dishes in a warmed Plexiglas box, and put the box in the incubator. Labeling for 1 to 2 hr is usually sufficient, but cells will tolerate as much as 2 mCi/ml for 6 hr and lower concentrations (0.1 to 0.5 mCi/ml) for 18 hr.
Labeling Cells and Preparing Lysates for Immunoprecipitation
Wash and lyse the cells 6. At the end of the labeling period, carry the labeled cells, still in the Plexiglas box, into the cold room. Place the cells behind a Plexiglas shield.
13.2.2 Supplement 10
Current Protocols in Protein Science
For adherent cells 7a. Take the dish out of the box and remove the labeling medium manually, using either a plugged disposable pipet or a disposable one-piece plastic Pasteur transfer pipet. Discard the medium and pipet as radioactive waste. Labeling medium should be removed without using a vacuum aspirator. Vacuum aspiration generates radioactive aerosols and leaves a radioactive film on the equipment.
8a. Wash cells once with 2 to 10 ml cold TBS. Remove the wash buffer manually, as in step 7, and discard as radioactive waste. The uptake of phosphate is efficient and often a majority of the added 32Pi is found within the labeled cells. This necessitates continued shielding of the labeled cells following removal of the labeling medium.
9a. Add lysis buffer to cells, using 0.3 ml per 35-mm dish, 0.6 ml per 50-mm dish, or 1.0 ml per 100-mm dish of adherent cells. Dislodge adherent cells by scraping with a rubber policeman, but leave the lysate in the dish. Incubate 20 min at 4°C. With a rubber policeman, scrape the lysate of adherent cells to the side of the dish and transfer lysate to a screw-cap microcentrifuge tube using a disposable one-piece plastic transfer pipet. If a low background of nonspecific contaminants is critical, use RIPA lysis buffer. If maintenance of enzymatic activity or the structure of protein complexes is critical, use a milder lysis buffer containing either CHAPS or NP-40 as the only detergent. If complete solubilization of the cells and denaturation of the protein is desired, use SDS for lysis (see Alternate Protocol).
For nonadherent cells 7b. Take the dish out of the box, transfer the cells to a screw-cap centrifuge tube, pellet them by centrifugation (1 min at 1800 × g), and remove the medium. Do not use a vacuum aspirator to remove medium.
8b. Resuspend pelleted cells gently in a small volume of cold TBS, transfer to a screw-cap microcentrifuge tube, and pellet the cells by microcentrifuging 1 min at 1800 × g. Continue to use shielding with cells and lysate.
9b. Add 0.5 to 1 ml lysis buffer per 107 cells and resuspend the pellet by gentle agitation with a disposable plastic Pasteur pipet. Incubate 20 min at 4°C. Choose the lysis buffer as in annotation to step 9a.
10. Cap the tube, and clarify the lysate by centrifuging 30 min at 26,000 × g (17,000 rpm in Sorvall SM-24 rotor), 4°C. Use an ice bucket with a sheet of Plexiglas over it, a chilled Plexiglas tube holder, or a prechilled centrifuge rotor to transport the tube of lysate. It is best to half-fill the tube to prevent spilling. A Sorvall SM-24 rotor with rubber adaptors for microcentrifuge tubes is ideal for clarifying lysates. Alternatively, a refrigerated microcentrifuge can be used for 30 min at maximum speed. Do not use a nonrefrigerated microcentrifuge in the cold room because the centrifuge will warm to 20°C during a 30-min spin. Lysates prepared with RIPA buffer often become viscous due to lysis of nuclei. If this occurs, increase the time of centrifugation to 90 min, or add 50 ìl of fixed Staphylococcus aureus bacteria (Pansorbin, Calbiochem) in RIPA buffer to the lysate prior to centrifugation. Either modification will cause the solubilized DNA to pellet.
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13.2.3 Current Protocols in Protein Science
Supplement 10
11. After centrifugation, transfer the supernatant (lysate) to a new tube and discard the tube and pellet in radioactive waste. If the pellet is very viscous, so that it is impossible to remove the supernatant cleanly, suck part of the pellet into a micropipet tip, lift the viscous material out of the tube, and discard it in the radioactive waste. The residual liquid in the tube is the supernatant and can be used for immunoprecipitation.
12. Analyze the labeled lysate using gel electrophoresis (UNIT 10.1), immunoprecipitation, or protein purification. Carry out all analytical procedures at 4°C using adequate shielding. ALTERNATE PROTOCOL
LYSIS OF CELLS BY BOILING IN SDS Some proteins, such as eukaryotic RNA polymerase II, are difficult to solubilize with mild lysis buffer or RIPA lysis buffer, and some analytical procedures use antibodies that recognize epitopes exposed only in denatured proteins. In these cases, it is useful to solubilize labeled cells completely in SDS and then adjust the composition of the lysate solution to match that of RIPA buffer for immunoprecipitation. To avoid the formation of spurious disulfide bonds, lysis and washing during immunoprecipitation are carried out in the presence of fresh 1 mM dithiothreitol (DTT). The protocol describes the procedure for adherent cells; modifications for working with nonadherent cells are described as alternate steps. Additional Materials (also see Basic Protocol) SDS lysis buffer (see recipe) RIPA correction buffer (see recipe) Immunoprecipitate wash buffer (see recipe) Fixed Staphylococcus aureus bacteria (Pansorbin, Calbiochem; optional) Boiling water bath 1. Label and wash cells (see Basic Protocol steps 1 to 7). 2a. For adherent cells: Add SDS lysis buffer using 0.1 ml for a 35-mm dish, 0.25 ml for a 50-mm dish, or 0.5 ml for an 100-mm dish. Immediately scrape off the dish with a rubber policeman and transfer the cell lysate to a screw-cap microcentrifuge tube. 2b. For nonadherent cells: Vortex briefly to loosen the cell pellet, add 1 ml SDS lysis buffer per 5 × 107 cells and vortex again. 3. Boil the samples 2 to 5 min, then add 4 vol RIPA correction buffer and mix well. 4. Clarify the cell lysate by centrifuging 90 min at 26,000 × g (17,000 rpm in Sorvall SM-24), 4°C, or at maximum speed in a refrigerated (4°C) microcentrifuge. Lysate may also be clarified by adding 50 ìl fixed Staphylococcus aureus and centrifuging 30 min at 26,000 × g, 4°C.
5. Carry out immunoprecipitation as usual using immunoprecipitate wash buffer for washes.
Labeling Cells and Preparing Lysates for Immunoprecipitation
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Current Protocols in Protein Science
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Immunoprecipitate wash buffer for boiled sample RIPA lysis buffer (see recipe) 1 mM DTT, added fresh DTT is added from a 1 M stock solution stored at −20°C.
Mild lysis buffer 10 mM 3[(3-cholamidopropyl)-dimethylammonio]-1-propane-sulfonate (CHAPS) or 1% (w/w) Nonidet P-40 (NP-40) 0.15 M NaCl 0.01 M sodium phosphate, pH 7.2 2 mM EDTA 50 mM sodium fluoride 0.2 mM sodium vanadate added fresh from 0.2 M stock solution 100 U/ml aprotinin (Trasylol, Pentex/Miles) Store buffer without vanadate at 4°C up to 1 year CHAPS is a milder detergent than NP-40, but yields precipitates with a higher background and may solubilize some proteins less efficiently. Sodium vanadate stock solution can be stored in plastic at room temperature.
Phosphate-buffered saline (PBS) Dissolve the following in 800 to 900 ml H2O: 8 g sodium chloride (136.8 mM final) 0.2 g potassium chloride (2.5 mM final) 0.115 g dibasic sodium phosphate (anhydrous; 0.8 mM final) 0.2 g monobasic potassium phosphate (anhydrous; 1.47 mM final) 0.1 g calcium chloride (anhydrous; 0.9 mM final) 0.1 g magnesium chloride hexahydrate (0.5 mM final) Add H2O to 1 liter Dispense 50- or 100-ml aliquots into glass bottles Autoclave and store indefinitely at room temperature RIPA (RadioImmunoProtection Assay) correction buffer for boiled sample 1.25% (w/w) Nonidet P-40 (NP-40) 1.25% (w/v) sodium deoxycholate 0.0125 M sodium phosphate, pH 7.2 2 mM EDTA 0.2 mM sodium vanadate added fresh from 0.2 M stock solution 50 mM sodium fluoride 100 U/ml aprotinin (Trasylol, Pentex/Miles) Store buffer without vanadate at 4°C up to 1 year Sodium vanadate stock solution can be stored in plastic at room temperature.
RIPA (RadioImmunoPrecipitation Assay) lysis buffer 1% (w/w) Nonidet P-40 (NP-40) 1% (w/v) sodium deoxycholate 0.1% (w/v) SDS 0.15 M NaCl 0.01 M sodium phosphate, pH 7.2 2 mM EDTA continued
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Supplement 10
50 mM sodium fluoride 0.2 mM sodium vanadate added fresh from 0.2 M stock solution 100 U/ml aprotinin (Trasylol, Pentex/Miles) Store buffer without vanadate at 4°C up to 1 year Sodium vanadate stock solution can be stored in plastic at room temperature.
SDS lysis buffer 0.5% (w/v) SDS 0.05 M Tris⋅Cl, pH 8.0 (APPENDIX 2E) 1 mM DTT, added fresh SDS and Tris⋅Cl solutions can be made in advance and stored at room temperature. DTT is added from a 1 M stock solution stored at −20°C.
Tris-buffered saline (TBS) 8 g sodium chloride (136.8 mM final) 0.38 g potassium chloride (5.0 mM final) 0.1 g calcium chloride (anhydrous; 0.9 mM final) 0.1 g magnesium chloride hexahydrate (0.5 mM final) 0.1 g dibasic sodium phosphate (anhydrous; 0.7 mM final) 800 to 900 ml H2O Add 25 ml 1 M Tris⋅Cl, pH 7.4 (APPENDIX 2E) Add H2O to 1 liter Dispense 50- or 100-ml aliquots into glass bottles Autoclave and store indefinitely at room temperature COMMENTARY Background Information
Labeling Cells and Preparing Lysates for Immunoprecipitation
The extent to which a protein becomes radiolabeled via biosynthetic labeling with 32Pi depends on the rate of transport of phosphate into the cells being labeled, abundance of the protein, stoichiometry of phosphorylation of the protein, and rate of phosphate turnover in the protein. The rate of turnover of the protein itself is less important because phosphate in protein usually turns over at a much faster rate than does the protein. Most of the phosphate taken up by cells is not incorporated into phosphoproteins—the vast majority is incorporated into phospholipid, RNA, and DNA. The efficiency of incorporation of 32Pi into proteins is therefore low—only ∼1% of incorporated radioactivity is found in phosphoproteins. An obvious and immediate question is how many cells and how much isotope should be used. The answer is that it depends on the protein being studied. If the protein is abundant (i.e., it constitutes 0.5% to 1% of total cellular protein) and highly phosphorylated (i.e., it contains 1 mole of phosphate per mole of protein), it can be labeled by incubating 106 cells with 0.2 to 0.5 mCi 32Pi in 2 ml labeling medium for 1 hr. In contrast, if the protein is rare or nonabundant, it may be necessary to label 107 cells with 5 to 10 mCi of 32Pi in 3 to 5 ml labeling
medium for 4 to 6 hr. A pilot experiment can be useful: if 50 cpm are recovered in the protein of interest, use more cells and more isotope; if 200,000 cpm are recovered in the protein of interest, use fewer cells with less isotope.
Critical Parameters Incorporation of 32Pi during biosynthetic labeling is greatest if labeling is done in medium lacking any phosphate except the added radioisotope. Labeling can also be accomplished at a somewhat reduced efficiency in complete growth medium. Phosphate-free medium is commercially available but can also be easily formulated in-house by omitting sodium and potassium phosphate from a recipe and replacing them with either sodium chloride or potassium chloride or both. Serum dialyzed against phosphate-free saline can be used instead of complete serum to further reduce the level of phosphate in labeling medium. Dialysis of serum against phosphate-free saline reduces but never entirely eliminates the phosphate in serum. Washing or rinsing cells in reducedphosphate medium just prior to labeling significantly increases labeling efficiency, but starvation of cells by incubation in reduced-phosphate medium prior to labeling is of very limited value.
13.2.6 Supplement 10
Current Protocols in Protein Science
Labeling cells with 32P almost certainly induces radiation damage in the labeled cells and will affect or arrest cell-cycle progression. If phosphorylation of the protein under study varies during the cell cycle, labeling with 32P may alter its phosphorylation. At the end of the labeling interval, the radioactive medium should be discarded, except in experiments where labeled virions or labeled secreted proteins are being studied. Labeling medium should be removed without using a vacuum aspirator. Vacuum aspiration generates radioactive aerosols and leaves a radioactive film inside the aspirator hose. As a result the hose can become an intense source of radiation exposure. It is best to use a plugged disposable pipet or disposable plastic Pasteur pipet for removing labeling medium. The medium and pipet should be discarded immediately as radioactive wastes. In general, labeled cells are lysed and subjected to gel electrophoresis, immunoprecipitation, or protein purification. If the labeled cells are to be centrifuged, they should be transferred to capped, disposable centrifuge tubes. Tubes should be no more than half full if they are to be spun in a fixed-angle rotor; tubes that are too full may leak radioactive lysate and contaminate the centrifuge. Screw-cap microcentrifuge tubes are ideal for centrifuging lysates. If large volumes are being handled, multiple partially filled tubes are preferable to tubes filled to the top. An ice bucket can be used for storage and transport of samples to keep the samples cold and provide considerable shielding. Radiation exposure from the tops of the tubes can be minimized by covering the ice bucket with a 1⁄ -in. Plexiglas sheet. 4 RadioImmunoPrecipitation Assay (RIPA) buffer (Brugge and Erikson, 1977) is the lysis buffer of choice because it solubilizes proteins well, gives a low background of nonspecific proteins, and is tolerated by most antigens and antibodies. It does however denature some antigens and disrupt some protein:protein complexes. If 1% (w/w) Nonidet P-40 (NP-40) is used as the only detergent, most of these problems are solved without increasing background unacceptably (Sefton et al., 1980). Some workers like to use digitonin as an extremely mild detergent for cell lysis. Digitonin sometimes is tricky and idiosyncratic, giving erratic results and high backgrounds during immunoprecipitation. 3[(3-cholamidopropyl)-dimethylammonio]-1-propane-sulfonate (CHAPS; Chen et
al., 1990) and Brij 96 (Osman et al., 1992) are mild detergents that appear to give more reproducible results. Besides the usual concern about proteolysis following cell lysis, the cells must be handled carefully to prevent protein phosphorylation and dephosphorylation following lysis of the cells. Both problems can be minimized by proper formulation of the lysis buffer and by keeping the sample at 4°C. Inclusion of 2 mM EDTA in the lysis buffer will minimize phosphorylation in the lysate by chelating both Mg2+ and Mn2+, which are essential for protein kinase activity. Addition of phosphatase inhibitors to the lysis buffer will reduce dephosphorylation significantly. For example, 50 mM sodium fluoride is used to inhibit serine and threonine phosphatases, and 0.2 mM sodium vanadate is used to inhibit all known tyrosine phosphatases. Sodium vanadate appears to lose its effectiveness if it is stored diluted in lysis buffer, so it should be added fresh each time. The concentrated stock solution should be stored in plastic at room temperature.
Anticipated Results This protocol can be used to label phosphoproteins or other phosphorylated cellular constituents. Phosphoproteins will contain ∼1% of incorporated label.
Time Considerations Proteins can and should be labeled biosynthetically with 32Pi and isolated by immunoprecipitation in a single day. As is the case with all 32P-labeled samples, it is important to work quickly because the specific activity of the sample declines 5% per day. If it is absolutely necessary to stop during the immunoprecipitation, it is best to leave the samples as precipitated pellets after aspiration of the wash buffer. Such samples can be stored at 4°C or −20°C overnight. Freezing the cell lysate tends to increase the background. Frozen lysates should be reclarified by centrifugation prior to immunoprecipitation.
Literature Cited Brugge, J.S. and Erikson, R.L. 1977. Identification of a transformation-specific antigen induced by an avian sarcoma virus. Nature 269:346-348. Chen, J., Stall, A.M., Herzenberg, L.A., and Herzenberg, L.A. 1990. Differences in glycoprotein complexes associated with IgM and IgD on normal murine B cells potentially enable transduction of different signals. EMBO J. 9:2117-2124.
Post-Translational Modification: Phosphorylation and Phosphatases
13.2.7 Current Protocols in Protein Science
Supplement 10
Osman, N., Ley, S.C., and Crumpton, M.J. 1992. Evidence for an association between the T cell receptor/CD3 antigen complex and the CD5 antigen complex in human T lymphocytes. Eur. J. Immunol. 22:2995-3000.
Contributed by Bartholomew M. Sefton The Salk Institute San Diego, California
Sefton, B.M., Hunter, T., and Beemon, K. 1980. Temperature-sensitive transformation by Rous sarcoma virus and temperature-sensitive protein kinase activity. J. Virol. 33:220-229.
Labeling Cells and Preparing Lysates for Immunoprecipitation
13.2.8 Supplement 10
Current Protocols in Protein Science
Phosphoamino Acid Analysis
UNIT 13.3
It is often valuable to identify the phosphorylated residue in a protein. In the case of proteins phosphorylated at serine, threonine, or tyrosine, this is readily accomplished by partial acid hydrolysis in HCl followed by two-dimensional thin-layer electrophoresis of the labeled phosphoamino acid (Basic Protocol). Phosphothreonine and phosphotyrosine are more stable to hydrolysis in alkali than are RNA and phosphoserine. Therefore, mild alkaline hydrolysis of protein samples can be used to enhance the detection of phosphothreonine and phosphotyrosine (Alternate Protocol). Although this procedure can be carried out with a protein eluted from a preparative gel and concentrated by trichloroacetic acid (TCA) or acetone precipitation, it is most easily accomplished by transfer of the protein of interest to a PVDF membrane. This technique is obviously not ideal if the protein being studied does not transfer efficiently. NOTE: Wear gloves and use blunt-end forceps to handle membranes. ACID HYDROLYSIS AND TWO-DIMENSIONAL ELECTROPHORETIC ANALYSIS OF PHOSPHOAMINO ACIDS The protein to be acid hydrolyzed is transferred to a PVDF membrane using the same technique used for immunoblotting or for microsequencing. It is useful, but not absolutely essential, to keep the filter wet following transfer. Following acid hydrolysis, phosphoamino acids are separated by two-dimensional thin-layer electrophoresis. Because electrophoresis equipment differs considerably in design, the details of the assembly and placement of the plate are not discussed here. It is assumed that a suitable apparatus is available for use by an experienced operator. Electrophoresis conditions are described for using the HTLE 7000 (CBS Scientific). They are almost certainly not correct for other equipment and will need to be altered according to the equipment manufacturer’s directions.
BASIC PROTOCOL
Materials 32 P-labeled phosphoprotein (UNIT 13.2) India ink solution: 1 µl/ml India ink in TBS (see recipe)/0.02% (v/v) Tween 20, pH 6.5 (prepare fresh or store indefinitely at room temperature); or radioactive or phosphorescent alignment markers 6 M HCl (APPENDIX 2E) Phosphoamino acid standards mixture (see recipe) pH 1.9 electrophoresis buffer (see recipe) pH 3.5 electrophoresis buffer (see recipe) 0.25% (w/v) ninhydrin in acetone in a freon (aerosol, gas-driven) atomizer/sprayer PVDF membrane (Immobilon-P, Millipore) 110°C oven Screw-cap microcentrifuge tubes 20 cm × 20 cm × 100 µm glass-backed cellulose thin-layer chromatography plate (EM Sciences) Large blotter: two 25 × 25–cm layers of Whatman 3MM paper sewn together at the edges, with four 2-cm holes that align with the origins on the TLC plate Glass tray or plastic box Whatman 3MM paper Thin-layer electrophoresis apparatus (e.g., HTLE 7000, CBS Scientific) Fan
Contributed by Bartholomew M. Sefton Current Protocols in Protein Science (1995) 13.3.1-13.3.8 Copyright © 2000 by John Wiley & Sons, Inc.
Post-Translational Modification: Phosphorylation and Phosphatases
13.3.1 Supplement 4
Small blotters: 4 × 25–cm, 5 × 25–cm, and 10 × 25–cm pieces of Whatman 3MM paper 50° to 80°C drying oven Sheets of transparency film for overhead projector Additional reagents and equipment for SDS-PAGE (UNIT 10.1), electroblotting onto PVDF membranes (UNIT 10.7), and staining membranes (UNIT 10.8) Prepare sample 1. Run radiolabeled phosphoprotein on a preparative SDS-polyacrylamide gel (UNIT 10.1). 2. Transfer proteins electrophoretically to a PVDF membrane (UNIT membrane several times with water. Do not let the membrane dry.
10.7).
Wash the
These washes remove buffer and detergent.
3. Locate the band of interest by staining the membrane 5 to 10 min in 30 to 50 ml India ink solution with shaking until bands are detectable, or by wrapping the membrane in plastic wrap, applying radioactive or phosphorescent alignment markers, and performing autoradiography. 4. Excise the piece of membrane containing the band of interest with a clean razor blade. Rewet the piece of membrane with methanol for 1 min and then rewet it in >0.5 ml water. Place the piece of membrane paper in a screw-cap microcentrifuge tube. Keep the excised piece as small as possible.
Hydrolyze sample 5. Add enough 6 M HCl to submerge the piece of membrane. Screw the cap on the tube tightly and incubate 60 min in 110°C oven. 6. Let cool. Microcentrifuge 2 min at maximum speed. Transfer the liquid hydrolysate to a fresh microcentrifuge tube and dry with a Speedvac evaporator. Drying takes ∼2 hr. Simultaneous drying of the hydrolysate and deblocked oligonucleotides in NH4OH must be avoided, as this will generate a cloud of ammonium chloride that will collect in the centrifuge tube and render the hydrolysate unsuitable for thin-layer electrophoresis.
7. Dissolve the sample in 6 to 10 µl water by vortexing vigorously. Microcentrifuge 5 min at maximum speed. Prepare plate for first-dimension electrophoresis 8. Spot 25% to 50% of the sample, in 0.25- to 0.50-µl aliquots, on one origin of a 20 cm × 20 cm × 100 µm glass-backed cellulose thin-layer chromatography plate (see Fig. 13.3.1 for arrangement of samples). Between each application, dry the sample spot with compressed air delivered through a Pasteur pipet plugged with cotton. Use long, thin plastic micropipet loading tips for loading, and do not let the tip touch the plate. Four samples can be analyzed simultaneously. The complete hydrolysate can be spotted on a single origin, but some streaking in the first dimension may be observed due to overloading. This problem can be avoided by using a fraction of the sample.
Phosphoamino Acid Analysis
9. Spot 1 µl nonradioactive phosphoamino acid standards mixture (containing phosphoserine, phosphothreonine, and phosphotyrosine) on top of each sample in 0.25to 0.50-µl aliquots as above.
13.3.2 Supplement 4
Current Protocols in Protein Science
A
B
25 cm 8 cm
3 cm 25 cm
8 cm 3 cm plate with samples applied at 4 origins (+)
C
first-dimension blotter for electrophoresis at pH 1.9
D
phosphamino acids blotter applied to plate in order to wet it
results of first-dimension electrophoresis at pH 1.9
Figure 13.3.1 First-dimension electrophoretic separation of phosphoamino acids at pH 1.9. (A) Positions of the four origins on a single 20 × 20–cm plate; (B) blotter used for wetting the plate with pH 1.9 electrophoresis buffer; (C) placement of the blotter on the plate (underneath; indicated by dashed outline); and (D) orientation of the plate between the + and − electrodes with the positions of the phosphoamino acids after electrophoresis.
10. Wet the large blotter (with four holes) by submerging it in pH 1.9 electrophoresis buffer in a large glass tray or plastic box. Briefly allow the excess buffer to drain off. Lower the wet blotter onto the prespotted plate with the origins on the plate in the centers of the four holes in the blotter (Fig. 13.3.1). Press on the blotter gently to achieve even wetting of the cellulose and concentration of the samples. When the plate is uniformly wet, remove the blotter. The blotter should be quite damp but not sopping wet. Excess buffer can be wicked off onto filter paper. Areas of the plate that are too dry can be seen through the blotter and will appear to be whiter than the rest of the plate. If this happens, dab the blotter with a Kimwipe wetted with pH 1.9 electrophoresis buffer. If there are puddles of buffer on the plate, let them dry before carrying out electrophoresis.
11. Place the thin-layer plate in the electrophoresis apparatus and overlap 0.5 cm of the right and left sides of the plate with wicks made of Whatman 3MM paper. If the
Post-Translational Modification: Phosphorylation and Phosphatases
13.3.3 Current Protocols in Protein Science
A
B 10 cm
5 cm
4 cm 25 cm second-dimension blotters for electrophoresis at pH 3.5
C
blotters applied to plate in order to wet it
D
phosphamino acids
90o counterclockwise rotation of plate
results of second-dimension electrophoresis at pH 3.5 showing ninhydrin-stained standards
Figure 13.3.2 Second-dimension electrophoretic separation of phosphoamino acids at pH 3.5. (A) The three pieces of Whatman 3MM paper used for wetting the plate with pH 3.5 electrophoresis buffer; (B) proper placement of the blotters on the plate (underneath; indicated by dashed outline); (C) reorientation of the plate for electrophoresis in the second dimension; and (D) orientation of the plate between the + and − electrodes with the position of the phosphoamino acids after electrophoresis.
apparatus has an air bag, be sure to inflate it. Close the cover and start electrophoresis. With an HTLE 7000, double-thickness Whatman 3MM wicks, and a plate with four samples, electrophorese 20 min at 1.5 kV. For the HTLE 7000 apparatus, use folded-over Whatman 3MM wicks that are 20 cm wide (the same as the plate) and not overly wet. Overly wet wicks will flood the plate and cause sample diffusion. For other electrophoresis, apparatuses the appropriate duration of electrophoresis can be determined empirically by examining the rate of migration of the phosphoamino acid standards.
Phosphoamino Acid Analysis
12. Following electrophoresis, remove the plate and quickly air dry with a fan without heating. It takes ∼20 min to dry the plate.
13.3.4 Current Protocols in Protein Science
Pi
phosphoserine phosphothreonine phosphotyrosine phosphopeptides generated by partial hydrolysis
origin pH 3.5
pH 1.9
Figure 13.3.3 Hypothetical autoradiogram of a two-dimensional separation. Four samples of acid-hydrolyzed, 32P-labeled proteins are applied at the origins, one in each of the four quandrants. This diagram shows the origins, the directions of electrophoresis, the positions of phosphoserine, phosphothreonine, and phosphotyrosine, the position of Pi, and the position of partially hydrolyzed fragments of the proteins for the upper right-hand sample. Every protein generates different partial hydrolysis peptide fragments.
Perform second-dimension electrophoresis 13. Wet the small blotters in pH 3.5 electrophoresis buffer and use them to wet the plate using the method described in step 10 to achieve even wetting without puddling (Fig. 13.3.2). After electrophoresis at pH 1.9, phosphoamino acids are present as a streak extending from the origin towards the + electrode. Blotters are not applied directly over the phosphoamino acids to prevent sample blurring or smearing.
14. Remove the blotters, rotate the plate 90° counterclockwise, and electrophorese 16 min at 1.3 kV in pH 3.5 electrophoresis buffer if using the HTLE 7000 apparatus. 15. At the end of the electrophoresis run, remove the plate and dry 20 to 30 min in an oven at 50° to 80°C. When dry, spray with 0.25% ninhydrin in acetone, then reheat in the oven 5 to 10 min to visualize the phosphoamino acid standards. 16. Place radioactive or phosphorescent alignment marks on the plate and autoradiograph with an intensifying screen overnight to 10 days at −70°C. 17. Following autoradiography, trace the alignment markers and the stained phosphoamino acid markers onto a transparent sheet used for overhead projectors. Save this template. Align the film with the plate and identify radioactive phosphoamino acids (Fig. 13.3.3).
Post-Translational Modification: Phosphorylation and Phosphatases
13.3.5 Current Protocols in Protein Science
ALTERNATE PROTOCOL
ALKALI TREATMENT TO ENHANCE DETECTION OF TYR- AND THR-PHOSPHORYLATED PROTEINS BLOTTED ONTO FILTERS Because phosphothreonine and phosphotyrosine are much more stable to hydrolysis in alkali than RNA or phosphoserine, detection of proteins containing phosphothreonine and phosphotyrosine can often be enhanced by mild alkaline hydrolysis of gel-fractionated samples. Although this technique was first developed for the treatment of fixed polyacrylamide gels, it is much more easily performed with proteins that have been first transferred to a PVDF membrane. Alkaline hydrolysis does not preclude subsequent phosphoamino acid analysis. A band from a blot that has been treated with alkali can be excised and subjected to acid hydrolysis as described in the Basic Protocol. Additional Materials (also see Basic Protocol) 1 M KOH TN buffer: 10 mM Tris⋅Cl (pH 7.4 at room temperature)/0.15 M NaCl 1 M Tris⋅Cl, pH 7.0 at room temperature (APPENDIX 2E) Covered plastic container (e.g., Tupperware box) 55°C oven or water bath 1. Run radiolabeled phosphoprotein on a preparative SDS–polyacrylamide gel and transfer proteins electrophoretically to a PVDF membrane (see Basic Protocol, steps 1 and 2). A nylon membrane may be used in place of a PVDF membrane, but in that case, the bands cannot subsequently be analyzed by acid hydrolysis, as nylon membrane will dissolve in 6 M HCl.
2. Wash membrane thoroughly with water: three 2-min incubations in 1 liter water are sufficient. These washes remove buffer and detergent.
3. Incubate membrane 120 min at 55°C in an oven or water bath in sufficient 1 M KOH to cover the filter in a covered Tupperware container. 4. Discard KOH. Wash membrane and neutralize remaining KOH by rinsing once for 5 min in 500 ml TN buffer, once for 5 min in 500 ml of 1 M Tris⋅Cl (pH 7.0), and twice for 5 min in 500 ml water. Wrap the membrane in plastic wrap and autoradiograph overnight with flashed film and an intensifying screen at −70°C. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
pH 1.9 electrophoresis buffer 50 ml 88% formic acid (0.58 M final concentration) 156 ml glacial acetic acid (1.36 M final concentration) 1794 ml H2O Store indefinitely in a sealed bottle at room temperature
Phosphoamino Acid Analysis
pH 3.5 electrophoresis buffer 100 ml glacial acetic acid (0.87 M final concentration) 10 ml pyridine [0.5% (v/v) final concentration] 10 ml 100 mM EDTA (0.5 mM final concentration) 1880 ml H2O Store indefinitely in a sealed bottle at room temperature
13.3.6 Current Protocols in Protein Science
Phosphoamino acid standards mixture Prepare a solution of phosphoserine, phosphothreonine, and phosphotyrosine (Sigma) in water at a final concentration of 0.3 µg/ml each. Store in 1-ml aliquots indefinitely at −20°C. Tris-buffered saline (TBS) For 100 ml, add 10 ml solution A to 89 ml H2O. While stirring rapidly, add 1 ml solution B slowly, drop by drop. Filter sterilize and store at 4°C. Solution A: Solution B: 80 g/liter NaCl 15 g/liter CaCl2 3.8 g/liter KCl 10 g/liter MgCl2 2 g/liter Na2HPO4 Filter sterilize 30 g/liter Tris base Store at −20°C Adjust pH to 7.5 Filter sterilize Store at −20°C Make certain that the stock TBS solution is mixed well just before use by bringing the solution into and out of a sterile 10-ml pipet several times.
COMMENTARY Background Information Phosphoamino acid analysis by the two-dimensional electrophoretic technique described in the Basic Protocol was first carried out with proteins isolated by elution from unfixed SDSpolyacrylamide gels (Hunter and Sefton, 1980). However, this technique is laborious, especially if it involves grinding up pieces of high-percentage acrylamide gels, and the yields can be disappointing. Additionally, because the eluted protein must be precipitated in the presence of a carrier protein, spotting the whole sample on a single origin usually yields a badly smeared pattern. The grind-and-elute technique is, however, advantageous with proteins that are very refractory to electrophoretic transfer to PVDF membranes. The alkaline treatment of protein described in the Alternate Protocol was first developed by Jon Cooper and Tony Hunter, who treated fixed gels with KOH (Cooper and Hunter, 1981). The original technique is tricky because the gel becomes extremely sticky during incubation with KOH and swells. Additionally, the manipulations needed to recover proteins from the gel following treatment are very involved because the proteins are contaminated with products of the hydrolysis of polyacrylamide.
Critical Parameters and Troubleshooting It is essential to use PVDF membranes to immobilize proteins for acid hydrolysis rather than nylon or nitrocellulose membranes, both
of which dissolve in 6 M HCl. Proteins immobilized on either PVDF or nylon membranes may be subjected to alkaline hydrolysis with KOH (Contor et al., 1987), but nitrocellulose membranes are not suitable. Proteins immobilized on nylon cannot subsequently be analyzed by acid hydrolysis because nylon is dissolved by 6 M HCl. Two-dimensional thin-layer electrophoresis is required for unambiguous identification of phosphorylated residues, as some spots after one-dimensional electrophoresis do not represent pure species. For example, uridine monophosphate, which is generated during acid hydrolysis of RNA (a frequent contaminant of phosphoproteins), comigrates with phosphotyrosine during one-dimensional electrophoresis at pH 3.5. Streaking of the sample in the first dimension is a symptom of overloading, either with the phosphoprotein itself or with contaminants in the sample. This problem can be corrected by loading less sample. Streaking in the second dimension is usually the result of problems with wetting or running the plate and cannot be corrected by loading less sample. Some batches of blotting paper contain calcium, which interferes with electrophoresis of phosphoamino acids (probably by precipitating them). Inclusion of EDTA in the pH 3.5 buffer alleviates this problem. In the author’s experience Whatman 3MM paper is quite reliable; other blotting papers are probably suitable as well.
Post-Translational Modification: Phosphorylation and Phosphatases
13.3.7 Current Protocols in Protein Science
This unit calls for glass-backed cellulose thin-layer plates rather than the plastic-backed variety, which are lighter and less expensive. This is because plastic-backed plates can under some circumstances cause sample streaking. They are, however, probably satisfactory for most experiments. If use of plastic-backed plates results in streaking, try glass-backed plates to see if that corrects the problem.
Anticipated Results To detect a phosphoamino acid by autoradiography, a minimum of 10 cpm must be spotted and the plate exposed for a week with flashed film and an intensifying screen. Only 15% to 20% of the radioactivity in a phosphoprotein is recovered as phosphoamino acids. The majority is present as 32Pi, which is released by dephosphorylation of phosphoamino acids, with the remainder being peptide products resulting from partial acid hydrolysis. As a result, the thin-layer plates will contain a number of radioactive spots that are not phosphoamino acids (see Fig. 13.3.3). Partial hydrolysis products remain near the origin during electrophoresis at pH 1.9, but exhibit some mobility at pH 3.5 (see Fig. 13.3.3). After two-dimensional electrophoresis, they are found above the origin and below the phosphoamino acids. 32Pi has a high mobility at both pH 1.9 and pH 3.5 and is found in the upper left-hand corner of each quadrant of the plate (see Fig. 13.3.3). Because of these additional radioactive spots, it is essential to localize internal phosphoamino acid standards by staining with ninhydrin.
Time Considerations After the preparative gel has been run and the protein transferred to the membrane, isolation of the membrane fragment containing the protein, followed by acid hydrolysis, takes <2 hr. Two to three hours are required for drying the sample with a Speedvac evaporator. First-
and second-dimension electrophoresis and staining the internal phosphoamino acid standards takes no more than ∼2 hr. Overnight autoradiographic exposure with flashed film and an intensifying screen is usually sufficient; however, exposures can be carried out for up to 10 days before the background from the screen becomes objectionable. Detection of phosphorylated protein after alkali treatment, as described in the Alternate Protocol, is a very quick procedure requiring little more time than it takes to carry out the alkaline hydrolysis. In general, samples of this sort can be detected after overnight exposure with flashed film and an intensifying screen at −70°C.
Literature Cited Contor, L., Lamy, F., and Lecocq, R.E. 1987. Use of electroblotting to detect and analyze phosphotyrosine containing peptides separated by two-dimensional gel electrophoresis. Anal. Biochem. 160:414-420. Cooper, J.A. and Hunter, T. 1981. Four different classes of retroviruses induce phosphorylation of tyrosines present in similar cellular proteins. Mol. Cell. Biol. 1:394-407. Hunter, T. and Sefton, B.M. 1980. The transforming gene product of Rous sarcoma virus phosphorylates tyrosine. Proc. Natl. Acad. Sci. U.S.A. 77:1311-1315.
Key Reference Kamps, M.P. and Sefton, B.M. 1989. Acid and base hydrolysis of phosphoproteins bound to Immobilon facilitates the analysis of phosphoamino acids in gel-fractionated proteins. Anal. Biochem. 176:22-27. Discusses all of the variables involved in subjecting filter-bound proteins to acid and base hydrolysis.
Contributed by Bartholomew M. Sefton The Salk Institute San Diego, California
Phosphoamino Acid Analysis
13.3.8 Current Protocols in Protein Science
Detection of Phosphorylation by Immunological Techniques
UNIT 13.4
Phosphorylation of unlabeled proteins can be detected using immunological or enzymatic techniques (UNIT 13.5). Anti-phosphotyrosine antibodies are used with immunoblots to detect tyrosine phosphorylation. These antibodies can be detected by 125I-labeled protein A (Basic Protocol) or enhanced chemiluminescence (ECL; Alternate Protocol). IMMUNOBLOTTING WITH ANTI-PHOSPHOTYROSINE ANTIBODIES AND DETECTION USING [125I]PROTEIN A
BASIC PROTOCOL
Initial determination of whether a protein is phosphorylated on tyrosine is carried out by immunoblotting with antibodies to phosphotyrosine. This technique is often superior to the use of biosynthetic labeling with 32Pi because of its sensitivity and low background, and it can be used to detect tyrosine phosphorylation of proteins in tissues that do not incorporate 32Pi efficiently. Materials Protein sample: Cultured cells, tissue, lysate, or immunoprecipitate Transfer buffer containing 100 µM sodium vanadate Blocking buffer (see recipe) Anti-phosphotyrosine antibody: 2 µg/ml rabbit polyclonal anti-phosphotyrosine (UBI) or mouse monoclonal anti-phosphotyrosine—e.g., py20 (Leinco, ICN Biomedicals, Zymed, Transduction Laboratories) or 4G10 (UBI)—in blocking buffer TN buffer: 10 mM Tris⋅Cl (pH 7.4 at room temperature)/0.15 M NaCl (APPENDIX 2E) TNA solution: TN buffer (recipe above)/0.01% (v/v) sodium azide (store at room temperature) NP-40 wash solution: 0.05% (v/v) Nonidet P-40 (NP-40)/TNA solution (store at room temperature) 0.5 µCi/ml 125I-labeled protein A (30 mCi/mg, ICN) in blocking buffer India ink solution: 1 µl India ink in TBS (UNIT 13.3)/0.02% (w/v) Tween 20, pH 6.5 (prepare fresh or store indefinitely at room temperature; optional) Plastic container with lid (e.g., Tupperware box) Blotting paper Additional reagents and solutions for SDS-PAGE (UNIT 10.1) and fluorography (UNIT 10.2) NOTE: Wear gloves and use blunt-end forceps to handle membrane. 1. Dissolve cultured cells, tissue, or lysate immunoprecipitate in an equal volume of 2× SDS sample buffer and boil 5 min. Lysates or immunoprecipitates in sample buffer can be stored at −70°C indefinitely. Storage >1 week at −20°C will lead to deterioration. Cells or tissues can be lysed by other means, but care must be taken to prevent phosphorylation or dephosphorylation from occurring following cell lysis. This can be accomplished by using 50 mM sodium fluoride to inhibit serine and threonine phosphatases, 0.2 mM sodium vanadate to inhibit tyrosine phosphatases, and 2 mM EDTA to inhibit kinases. Viscosity resulting from DNA can be reduced somewhat by boiling the sample 5 min and/or by shearing the sample by passing it five to ten times through a 22-G needle and five to ten times through a 27-G needle.
Contributed by Bartholomew M. Sefton Current Protocols in Protein Science (1995) 13.4.1-13.4.5 Copyright © 2000 by John Wiley & Sons, Inc.
Post-Translational Modification: Phosphorylation and Phosphatases
13.4.1 CPPS
2. Electrophorese sample on an SDS-polyacrylamide gel (UNIT 10.1). Transfer proteins to a membrane suitable for immunoblotting using transfer buffer that contains 100 µM sodium vanadate. The lysate from ∼105 cells can be analyzed in a single 0.5-cm wide lane of a 1-mm-thick SDS-polyacrylamide slab gel. Sodium vanadate should be included in the transfer buffer to prevent tyrosine dephosphorylation.
3. Incubate membrane in 30 to 50 ml blocking buffer ≥30 min at room temperature. Use of Blotto or other blocking mixtures containing dry milk is unsuitable because anti-phosphotyrosine antibodies bind to a number of the protein constituents of milk. Blocking buffer can be reused several times.
4. Incubate the membrane in anti-phosphotyrosine antibody 60 to 120 min at room temperature with occasional agitation. This step is most conveniently done in a covered plastic box using 30 to 50 ml antibody solution for a typical 15 × 15–cm blot. Although the volume of antibody solution seems large, it reduces the background. The antibody solution can be saved in the plastic box, stored at 4°C for at least a month, and reused up to five to ten times.
5. Remove the membrane from the staining box and wick off excess liquid with blotting paper. In a new plastic box, wash the membrane twice for 10 min with 50 ml TBSA, twice for 10 min with 50 ml NP-40 wash solution, and twice for 5 min with 50 ml TBSA. Discard washes. 6. Incubate membrane in 30 to 50 ml of 0.5 µCi/ml 125I-labeled protein A in blocking buffer 60 min at room temperature with gentle agitation in a covered plastic box. 125
I-labeled protein A is used to detect bound antibody.
7. Remove the membrane and wick off excess liquid with blotting paper. Wash the membrane as described in step 5 in a clean plastic box. The blotting paper and first wash should be discarded as radioactive wastes. The amount of radioactivity in the subsequent washes should not be great, and it is generally acceptable to pour them down the drain. Protein A solution can be stored at least 1 month at 4°C and reused ∼3 to 5 times.
8. If desired, stain the membrane 5 to 10 min with 30 to 50 ml India ink solution until bands are detectable. Wash the membrane with water to remove excess ink. 9. Remove excess moisture with blotting paper, wrap in plastic wrap, attach radioactive or fluorescent alignment markers, and expose by fluorography using an intensifying screen and preflashed film for 18 hr to 10 days.
Detection of Phosphorylation by Immunological Techniques
13.4.2 Current Protocols in Protein Science
DETECTION OF BOUND ANTIBODIES BY ENHANCED CHEMILUMINESCENCE (ECL)
ALTERNATE PROTOCOL
Bound anti-phosphotyrosine antibodies can also be detected using enhanced chemiluminescence (ECL). This technique uses a second antibody coupled to horseradish peroxidase to detect anti-phosphotyrosine antibodies. The bound second antibody is visualized by ECL and the image is captured on film. The technique is faster than that employing [125I]-labeled protein A (Basic Protocol) because it requires autoradiographic exposure of only minutes rather than days. It also avoids the use of radioactivity. However, results obtained with the two techniques are not always identical and exhibit quantitative differences. Additionally, if immunoprecipitated proteins are being analyzed, the secondary antibody may stain the precipitating antibodies intensely, making detection of proteins of a size similar to those of the heavy and light chains of immunoglobulin difficult. The technique is fundamentally the same as that using [125I]-labeled protein A, except that azide must be omitted from all solutions because it interferes with chemiluminescence chemistry. This makes storage and reuse of antibodies more difficult. Additionally, the technique appears to work somewhat better with nitrocellulose membranes than with Immobilon-P. Therefore, handling of the membranes requires somewhat more delicacy. Additional Materials (also see Basic Protocol) Blocking buffer (see recipe) without azide 0.05% (v/v) Nonidet P-40 (NP-40)/TN buffer (see Basic Protocol) Horseradish peroxidase–conjugated secondary antibody: anti-rabbit or anti-mouse antibody diluted 1/1000 to 1/2000 in blocking buffer without azide ECL detection reagents A and B (Amersham) Luminol reagent for ECL detection (Amersham) Oxidizing reagent for ECL detection (Amersham) Nitrocellulose membrane Plastic container slightly larger than the membrane Plastic sheet protector NOTE: The reagents for ECL may be purchased as a kit, the Enhanced Chemiluminescence Western Blotting Detection System, from Amersham. NOTE: Wear gloves and use blunt-end forceps to handle membrane. Prepare membrane 1. Prepare sample in 2× SDS-sample buffer (see Basic Protocol, step 1) and electrophorese on an SDS-polyacrylamide gel. Transfer proteins to a nitrocellulose membrane using a transfer buffer that contains 100 µM sodium vanadate. 2. In a plastic container, incubate membrane in blocking buffer without azide ≥30 min at room temperature. Expose membrane to antibodies 3. Incubate membrane in 30 to 50 ml anti-phosphotyrosine antibody 60 to 120 min at room temperature with occasional agitation. The anti-phosphotyrosine antibody can be either polyclonal rabbit serum or mouse monoclonal antibodies.
4. In a clean plastic container, wash the membrane twice for 10 min with TN buffer, twice for 10 min with 0.05% NP-40/TN buffer, and twice for 5 min with TN buffer. Discard the washes.
Post-Translational Modification: Phosphorylation and Phosphatases
13.4.3 Current Protocols in Protein Science
5. In a plastic container, incubate the membrane in 30 to 50 ml of horseradish peroxidase–conjugated secondary antibody 60 min at room temperature with gentle agitation. Use a secondary antibody appropriate for the primary antibody.
6. Remove membrane and wash it in a clean plastic container as described in step 4. Detect bound antibodies with ECL 7. In a plastic container only slightly larger than the membrane, mix equal volumes of the Luminol reagent and the oxidizing reagent for ECL detection. Amersham recommends that 0.125 ml of each reagent be used per square centimeter of membrane. This is probably excessive with large blots.
8. Put the membrane, with the side containing protein facing up, into the mixed reagents. Agitate gently 60 sec. 9. Remove the blot, wick off excess moisture, and place it, protein-side up, in a sheet protector. Alternatively, place the blot on a sheet of transparency film used for overhead projectors and cover with a second sheet of transparency film.
10. In the darkroom, expose the blot to X-ray film. Exposure times can range from 15 sec to 30 min.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Blocking buffer 5% (w/v) BSA 1% (w/v) hen ovalbumin 10 mM Tris⋅Cl, pH 7.4 at room temperature 0.15 M NaCl 0.01% (w/v) sodium azide Store ≤6 months at 4°C For enhanced chemiluminescence, omit the sodium azide.
COMMENTARY Background Information
Detection of Phosphorylation by Immunological Techniques
Immunoblotting with antibodies to phosphotyrosine is an extremely useful technique for the study of tyrosine phosphorylation. It appears to be very specific. Although there is a report that antibodies to phosphotyrosine recognize phosphohistidine (Frackelton et al., 1983), nonspecific cross-reactivity is rarely seen. It is possible to be misled, however, if a large amount of a single protein is subjected to immunoblotting. For example, nonspecific staining of molecular weight standards is often observed. This almost certainly does not indicate that they contain phosphotyrosine. Rather, it illustrates the fact that the primary antibodies and the secondary detection reagents exhibit a low level of nonspecific binding that can be
detectable if a large amount of a single protein is present in a sample.
Critical Parameters Not all anti-phosphotyrosine antibodies give the same staining patterns for immunoblots of the same sample. Each antibody appears to react with some proteins more strongly than with others. This presumably reflects the amino acid sequence surrounding the phosphotyrosine residue, but the cause has not been examined carefully. Correspondingly, a given anti-phosphotyrosine antibody will not react equally strongly with all sites of tyrosine phosphorylation. For example, rabbit polyclonal anti-phosphotyrosine antibodies raised against phosphoTyr-Gly-Ala react more strongly with
13.4.4 Current Protocols in Protein Science
phosphorylated Tyr-394 in the Lck tyrosine protein kinase than they do with phosphorylated Tyr-505. The differential reactivity of phosphotyrosine should be kept in mind when drawing quantitative conclusions. Diluted polyclonal anti-phosphotyrosine antibodies can be reused several times. Their properties, however, change subtly with reuse. The staining of some bands becomes less intense and the staining of others becomes apparently more prominent. This may be a consequence of preferential removal of high-affinity antibodies during early uses. It should not be a problem with monoclonal antibodies. Analysis of immunoprecipitated proteins by immunoblotting can be problematic. The reagent used to detect the staining antibody on the membrane will often bind to the heavy and light chains of the precipitating antibodies. This occurs to a varying degree with [125I]-labeled protein A, which under some circumstances can bind detectably to the heavy chain of rabbit antibodies even after they have been denatured on a membrane. It is a greater problem when second antibodies reactive with the species of antibody used for immunoprecipitation are used as detection reagents. For example, horseradish peroxidase–conjugated anti-mouse antibodies will, in general, bind strongly to both the heavy and light chains of mouse monoclonal antibodies used for immunoprecipitation. This can complicate the use of enhanced chemiluminescence (ECL) as a means for analyzing immunoblots of immunoprecipitated proteins. Immunoprecipitates can be run on nonreducing gels to cause interfering antibody bands to migrate near the top of the gel. Unreduced IgG runs as a 150-kDa band. Nonreducing sample buffer is the same as SDS sample buffer except that 50 mM iodoacetamide replaces the reducing agents 2-mercaptoethanol (2-ME) and DTT. The mobility of many, but not all, proteins is altered profoundly under nonreducing conditions and the suitability of nonreducing gel electrophoresis must be evaluated carefully for each case. It is often difficult to analyze reduced and nonreduced samples on the same gel because 2-ME, which is present in most sample buffers at very high concentrations, diffuses rapidly and can reduce nonreduced samples, even in the presence of 50 mM iodoacetamide. The ECL technique (Alternate Protocol) is very rapid and sensitive. It is unclear, however, whether it exhibits the same linearity of re-
sponse as does autoradiography with [125I]-labeled protein A. Immunoblotting should be used with caution for quantitative comparisons. For accurate quantitation, it is essential to determine the linearity of the detection system by measuring the signal obtained from multiple known dilutions of a standard sample. Not all proteins exhibit altered gel mobility when they undergo phosphorylation or dephosphorylation. Lack of effect of phosphatase digestion on the mobility of a protein is therefore not evidence of lack of phosphorylation.
Anticipated Results Immunoblotting with anti-phosphotyrosine antibodies is very sensitive. Even the extremely low levels of tyrosine phosphorylated proteins present in unactivated lymphoid cells can be detected when total lysates of cells are analyzed by gel electrophoresis and immunoblotting.
Time Considerations Immunoblotting with enhanced chemiluminescence is the most rapid technique described in this unit, because staining can be accomplished in less than a day and bound antibodies visualized within 30 min. Staining with [125I]labeled protein A can also be accomplished in a single day, but autoradiography usually requires 18 to 100 hr.
Literature Cited Frackelton, A.R., Ross, A.H., and Eisen, H.N. 1983. Characterization and use of monoclonal antibodies for isolation of phosphotyrosyl proteins from retrovirus-transformed and growth factor-stimulated cells. Mol. Cell. Biol. 3:1343-1352.
Key References Kamps, M.P. and Sefton, B.M. 1988. Identification of multiple novel polypeptide substrates of the v-src, v-yes, v-fps, v-ros, and v-erb-B oncogenic tyrosine protein kinases utilizing antisera against phosphotyrosine. Oncogene 2:305-315. Examines many of the variables that affect immunoblotting with antibodies to phosphotyrosine. Morla, A. and Wang, J.Y.J. 1986. Protein tyrosine phosphorylation in the cell cycle of BALB/c 3T3 fibroblasts. Proc. Natl. Acad. Sci. U.S.A. 83:8191-8195. Establishes the ability of immunoblotting with antibodies to phosphotyrosine to detect the low levels of tyrosine-phosphorylated proteins in normal cells.
Contributed by Bartholomew M. Sefton The Salk Institute San Diego, California
Post-Translational Modification: Phosphorylation and Phosphatases
13.4.5 Current Protocols in Protein Science
Detection of Phosphorylation by Enzymatic Techniques
UNIT 13.5
Reversible protein phosphorylation has been increasingly recognized as an important mechanism for regulating physiological processes in both plant and animal cells. There are a number of techniques to demonstrate the presence of covalently bound phosphate in proteins. Metabolic labeling of cells with 32P and subsequent isolation of the 32P-labeled protein is the most direct method for demonstrating protein phosphorylation. This approach is difficult when the specific polypeptides involved in the physiological phosphorylation event have not been well characterized or when these proteins are present in such low abundance in the cell that significant incorporation of [32P]phosphate cannot be achieved. Metabolic labeling of proteins (UNIT 13.2) may also be compromised by a slow turnover of the protein-bound phosphate in the intact cell. As the extent of phosphorylation of given protein reflects the relative activities of the protein kinases and phosphatases that act on it, knowledge of physiological stimuli that modulate the protein function might be expected to establish the experimental conditions which increase the metabolic labeling of the substrate protein. In the absence of this information it may be difficult to optimize protein phosphorylation in the intact cell. This situation can be further exacerbated by the lack of experimental tools, such as monospecific antibodies or affinity probes, that permit the rapid isolation of the substrate protein and preserve its phosphorylation. If significant 32P-labeling of the desired protein cannot be easily demonstrated, a different approach must be taken. Recent studies have utilized one or more phosphatases to reverse existing covalent modifications in proteins, resulting in changes in the function or activity of the substrate. In vitro dephosphorylation by one or more commercially available phosphatases has implicated phosphorylation as a regulatory mechanism in many physiological processes. These studies have been undertaken using cell extracts and subcellular fractions as well as partially purified proteins. The methodologies discussed in this unit can be used to detect protein phosphorylation where actual incorporation of [32P]phosphate into a specific polypeptide cannot be demonstrated. The overall strategy is to first examine the functional effects elicited by nonspecific acid or alkaline phosphatases that dephosphorylate many phosphoproteins in vitro (Basic Protocol 1 and Alternate Protocol 1). Nonspecific phosphatases isolated from plant and animal tissues show significant activity against a wide variety of phosphoester-containing compounds, including many phosphoproteins. These enzymes reverse all phosphorylations whether they occur as phosphotyrosine, phosphothreonine, phosphoserine, or phosphohistidine so they can define the regulatory consequence of protein dephosphorylation without necessarily identifying the specific phosphoamino acid. Protein phosphatases that selectively hydrolyze phosphoserine and phosphothreonine or phosphotyrosine residues can then be used to identify the functionally important covalent modification. Additional protocols describe digestion of phosphoproteins with a protein serine/threonine phosphatase (Basic Protocol 2) and protein tyrosine phosphatase (Alternate Protocol 2). A support protocol has been included to identify the radiolabel as 32Pi based on its ability to form a complex with ammonium molybdate. Protein dephosphorylation can be analyzed as the loss of [32P]phosphate from metabolically labeled proteins following their separation by SDS-PAGE. Some phosphoproteins also show changes in their electrophoretic migration following the loss of covalently bound phosphate. This can provide an additional monitor of protein dephosphorylation that can be used even in the absence of metabolic labeling. Finally, measurements of protein function (e.g., opening or closing of ion channels, binding of nuclear factors to specific DNA elements, or altered enzyme activity) can define the consequences of
Post-Translational Modification: Phosphorylation and Phosphatases
Contributed by Shirish Shenolikar
13.5.1
Current Protocols in Protein Science (1995) 13.5.1-13.5.8 Copyright © 1997 by John Wiley & Sons, Inc.
Supplement 5
dephosphorylation even when little information is available about the specific phosphopeptides involved. BASIC PROTOCOL 1
DIGESTION OF PHOSPHOPROTEINS WITH NONSPECIFIC ACID PHOSPHATASES Potato acid phosphatase shows a broad substrate specificity and hydrolyzes many metabolites containing phosphoester bonds. This enzyme shows comparatively low activity against phosphoproteins. Moreover, metabolites present in cell extracts or some subcellular fractions may inhibit its ability to dephosphorylate protein substrates. In this protocol, a sample containing a phosphoprotein is digested with potato acid phosphatase and analyzed for evidence of structural and functional changes. Materials Sample containing 100 to 200 µg total protein 50 mM piperazine-N,N′-bis(2-hydroxypropanesulfonic acid) (PIPES), pH 6.0 Sephadex G-25 column (optional) PIPES/2-ME or PIPES/DTT buffer, pH 6.0: 50 mM PIPES containing 15 mM 2-mercaptoethanol or 1 mM dithiothreitol (prepare fresh) Potato acid phosphatase 2× SDS-PAGE sample buffer: 50 mM Tris⋅Cl/0.4 M glycine (pH 8.3)/0.2% (w/v) SDS 100 mM sodium pyrophosphate or other general phosphatase inhibitor 90°C water bath or heating block Additional reagents and equipment for electrophoresis (UNIT 10.1) 1. Prepare a protein sample containing 100 to 200 µg total protein. Remove inhibitory metabolites or particulate material. If the substrate is particulate, soluble contaminants can be removed by washing 2 to 3 times with 50 mM PIPES, pH 6.0. Inhibitory metabolites can be removed by dialyzing against the same buffer, or if the substrate is a soluble protein, it can be desalted on a Sephadex G-25 column.
2. Incubate the sample in 100 µl PIPES/2-ME or PIPES/DTT, pH 6.0, for 10 min at 30°C. The pH optimum for potato acid phosphatase hydrolysis is between pH 4.0 and 5.0. However, many phosphoprotein substrates may be irreversibly inactivated at low pH, so incubation at pH 6.0 is recommended.
3. Add 5 to 10 U potato acid phosphatase and incubate 15 min at 30°C. If it is not possible to incorporate [32P]phosphate into the substrate protein by metabolic labeling, control reactions should include a general phosphatase inhibitor—e.g., sodium vanadate, sodium pyrophosphate, sodium glycerophosphate, sodium orthophosphate, or p-nitrophenyl phosphate (PNPP) at a final concentration ≥10 mM—to distinguish the effects produced by protein dephosphorylation by potato acid phosphatase from those produced by contaminating proteases. One unit of acid phosphatase hydrolyzes 1 nmol PNPP in 1 min at 30°C and pH 4.0.
4. Remove 10 µl of the sample and add an equal volume of 2× SDS-PAGE sample buffer. Heat 5 min at 90°C. Detection of Phosphorylation by Enzymatics Technique
5. Electrophorese control and dephosphorylated material on an SDS polyacrylamide gel (UNIT 10.1). For a radiolabeled sample, analyze the loss of radiolabel in the substrate protein by autoradiography or phosphoimage analysis; for unlabeled sample, analyze the change in substrate mobility by immunoblot analysis.
13.5.2 Current Protocols in Protein Science
6. Add 10 µl of 100 mM sodium pyrophosphate (10 mM final concentration) to the remaining dephosphorylated material to terminate the dephosphorylation reaction. Use an appropriate assay of protein function to characterize the changes that accompany substrate dephosphorylation. DIGESTION OF PHOSPHOPROTEINS WITH NONSPECIFIC ALKALINE PHOSPHATASE
ALTERNATE PROTOCOL 1
Calf intestine alkaline phosphatase has been widely utilized to dephosphorylate both proteins and nucleic acids. With the widespread use of this enzyme in molecular biology most commercial preparations are essentially free of contaminating proteases. Alkaline phosphatase has a broad pH profile with an optimum between pH 8.0 and 8.5. In contrast to acid phosphatases, this enzyme can be utilized for in vitro dephosphorylation of proteins under conditions that do not denature the substrate proteins. Calf intestine alkaline phosphatase effectively dephosphorylates proteins containing phosphoserine and phosphothreonine residues, which together account for >97% of the protein-bound phosphate in eukaryotic cells. Under defined conditions, preferential dephosphorylation of phosphotyrosines by alkaline phosphatases has also been reported (Swarup et al., 1981). Alkaline phosphatase activity is stimulated by Mg2+ ions, although significant activity is seen even in the absence of divalent cations. Additional Materials (also see Basic Protocol 1) Tris/MgCl2, pH 7.5 or HEPES/MgCl2, pH 7.5 buffer: 50 mM Tris⋅Cl (pH 7.5; APPENDIX 2E)/1 mM MgCl2 or 50 mM N,-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid (HEPES; pH 7.5)/1 mM MgCl2 Calf intestine alkaline phosphatase (molecular biology grade) 1. Incubate the sample (containing 100 to 200 µg total protein in 100 µl Tris/MgCl2 or HEPES/MgCl2) for 10 min at 30°C. 2. Add 20 to 30 U calf intestine alkaline phosphatase and incubate 15 min at 30°C. One unit of calf intestine alkaline phosphatase hydrolyzes 1 nmol p-nitrophenyl phosphate (PNPP) in 1 min at 30°C and pH 8.5.
3. Terminate the dephosphorylation reaction by adding an equal volume of 2× SDSPAGE sample buffer or a general phosphatase inhibitor. Analyze changes in substrate mobility and function (see Basic Protocol 1). DIGESTION OF PHOSPHOPROTEINS WITH PROTEIN SERINE/THREONINE PHOSPHATASES
BASIC PROTOCOL 2
Protein phosphatase 1, 2A, and 2B (PP1, PP2A, and PP2B, respectively) represent three major classes of protein serine/threonine phosphatases found in eukaryotic cells. These enzymes are currently available from several suppliers; they will retain full enzymatic activity for many months when stored below −20°C (see supplier’s recommendations). This protocol describes the use of PP2A, a broad-specificity phosphatase, to dephosphorylate the phosphoproteins. Materials Sample containing 100 µg total protein Tris/DTT/MnCl2 buffer, pH 7.5: 50 mM Tris⋅Cl (pH 7.5; APPENDIX 2E)/1 mM dithiothreitol (DTT)/1 mM MnCl2 (prepare fresh) Microcystin-LR Protein phosphatase 2A (PP2A), catalytic subunit
Post-Translational Modification: Phosphorylation and Phosphatases
13.5.3 Current Protocols in Protein Science
1. Prepare the sample (containing 100 µg total protein) in 100 µl Tris/DTT/MnCl2 buffer, pH 7.5. Prepare a control reaction in the same buffer with 1 µM microcystinLR. Incubate 10 min at 37°C. Microcystin-LR is a potent inhibitor of PP1 and PP2A.
2. Add 0.2 to 0.5 U PP2A and incubate 10 to 30 min at 37°C. One unit of PP2A or PP1 activity hydrolyzes 1 nmol phosphorylase A in 1 min at 30°C and pH 7.5.
3. Add microcystin-LR to 1 µM final concentration to terminate the dephosphorylation reaction. Unlike the general phosphatases discussed above, protein phosphatases show some selectivity for the dephosphorylation of different sites in substrate phosphoproteins and can be used to gain insight into the functional role of individual protein-bound phosphates.
4. Analyze the dephosphorylated material by SDS-PAGE or functional assays (see Basic Protocol 1). ALTERNATE PROTOCOL 2
DIGESTION OF PHOSPHOPROTEINS WITH PROTEIN TYROSINE PHOSPHATASES Transmembrane tyrosine phosphatases such as CD45 possess low in vitro activity against many substrates, leading to the suggestion that association with extracellular ligands activates these receptor phosphatases. In this regard, protein tyrosine phosphatase 1B (PTP-1B) and src homology-protein tyrosine phosphatases (SH-PTP1 or SH-PTP2) are highly active phosphotyrosine phosphatases that demonstrate no measurable activity against phosphoserine and phosphothreonine and may be the preferred reagents to investigate the regulatory role of tyrosine phosphorylation. Materials Sample containing 10 to 100 µg total protein 50 mM imidazole, pH 7.5 Protein tyrosine phosphatase (e.g., PTP-1B or SH-PTP) 2× SDS-PAGE sample buffer (see Basic Protocol 1) or 100 mM sodium vanadate 1. Incubate the sample (containing 10 to 100 µg total protein) in 50 mM imidazole, pH 7.5, for 10 min at 37°C. 2. Add 1 to 5 U protein tyrosine phosphatase and incubate 15 to 30 min at 37°C. One unit of phosphatase activity hydrolyzes 1 nmol p-nitrophenyl phosphate (PNPP) in 1 min at 37°C and pH 7.0.
3. Terminate the dephosphorylation reaction by adding an equal volume of 2× SDSPAGE sample buffer or 100 mM sodium vanadate to a final concentration of 0.1 mM. Recombinant PTP-1B and SH-PTP1 are also available as GST fusion proteins that can be bound to glutathione-agarose and conveniently removed by centrifugation or filtration following dephosphorylation of the substrate.
4. Analyze digested material by gel electrophoresis, immunoblotting, or a functional assay (see Basic Protocol 1, steps 5 and 6). Detection of Phosphorylation by Enzymatics Technique
13.5.4 Current Protocols in Protein Science
MEASUREMENT AND IDENTIFICATION OF RELEASED 32P Time-dependent dephosphorylation of radiolabeled proteins is usually monitored by precipitating the protein substrate with 10% to 15% (w/v) trichloroacetic acid (TCA) at 4°C. Following removal of the TCA-denatured phosphoprotein by centrifugation, radioactivity that accumulates in the TCA-soluble fraction provides a monitor of [32P]phosphate release. However, this procedure cannot distinguish between the release of orthophosphate and TCA-soluble phosphopeptides. To define a dephosphorylation event, the TCA-soluble radioactivity must be characterized as 32Pi. This procedure takes advantage of the unique ability of 32Pi orthophosphate to form a complex with ammonium molybdate. The [32P]phosphomolybdate complex is soluble in organic solvent and can be readily separated from 32P-labeled phosphopeptides. Therefore, this assay specifically monitors protein dephosphorylation (Shenolikar and Ingebritsen, 1984).
SUPPORT PROTOCOL
Materials Trichloroacetic acid Radiolabeled protein/phosphatase reaction mixture (see Basic Protocol 1) 1.25 mM potassium phosphate (KH2PO4)/1 N H2SO4 1:1 (v/v) isobutanol/toluene 5% (w/v) ammonium molybdate Scintillation fluid Liquid scintillation counter 1. Add trichloroacetic acid (TCA) to dephosphorylated radiolabeled protein to give a final concentration of 10% to 15% (w/v). Incubate 10 min at 4°C. 2. Microcentrifuge 5 min at maximum speed. Remove 0.1 ml of the TCA supernatant and place in a microcentrifuge tube. Add 0.2 ml of 1.25 mM KH2PO4/1 N H2SO4 and mix. 3. Add 0.5 ml of 1:1 isobutanol/toluene to the microcentrifuge tube and mix thoroughly. 4. Add 0.1 ml of 5% ammonium molybdate and mix. 5. Microcentrifuge 5 min at maximum speed to separate aqueous and organic phases. 6. Remove 0.3 ml of the upper (organic phase), mix with 1 ml scintillation fluid, and count in a liquid scintillation counter. COMMENTARY Background Information A key feature of protein phosphorylation (as compared to other protein modifications, such as proteolysis) is its reversibility. The ability to phosphorylate a substrate in vitro requires that the kinase responsible for phosphorylation be known. However, many cellular protein kinases are themselves activated via protein phosphorylation, making it difficult to use them in vitro to phosphorylate the substrate. Establishing the physiological relevance of many in vitro phosphorylations is also made difficult because these studies often do not take account of the physiological concentrations of the kinase and its substrate. In vitro experiments also tend to ignore the importance of the subcellular local-
ization of the substrate and the kinase, sometimes leading to modifications that do not occur in vivo. For these reasons, knowledge of the physiological stimuli that regulate the substrate’s function can be used to promote substrate phosphorylation as well as to provide important clues in the identification of protein kinases that phosphorylate the substrate. Most cellular phosphoproteins are subject to covalent modification at multiple sites by more than one protein kinase. Therefore, in vitro studies with purified protein kinases often fail to recreate the spectrum of phosphorylations seen in the intact cell, and this also makes it difficult to define the role of individual covalent modifications. Thus, a new strategy of
Post-Translational Modification: Phosphorylation and Phosphatases
13.5.5 Current Protocols in Protein Science
Detection of Phosphorylation by Enzymatics Technique
analyzing the role of multisite phosphorylation has been developed using proteins that have been modified in the intact cell. However, it is particularly important to isolate such substrates in the presence of phosphatase inhibitors and to preserve the protein-bound phosphates during lengthy purification procedures. Biochemical studies of protein phosphatases have established the in vitro substrate specificity of these enzymes, which distinguish between different phosphoamino acids (phosphoserine/phosphothreonine versus phosphotyrosine) and different phosphoprotein substrates. These enzymes can also show preferential dephosphorylation of specific sites (Shenolikar and Nairn, 1991; Charbonneau and Tonks, 1992; Shenolikar, 1994). In vitro dephosphorylation by selected phosphatases may not only define the functional role of selected covalent modifications but also identify “silent” or “structural” phosphorylations that do not directly regulate the protein function. These phosphorylations may play an indirect role in modulating protein phosphorylation or dephosphorylation of other sites. This unit has focused on commercially available phosphatases that have been used to characterize the functional importance of protein phosphorylation. For example, several growth-associated kinases, including the mitogen-activated protein kinases (MAP kinases) and the cyclin-dependent kinases such as cdc2, are phosphorylated on adjacent threonine and tyrosine residues. Phosphorylation of these enzymes is controlled by kinases and phosphatases that are themselves highly regulated in the intact cell. In vitro dephosphorylation by serine/threonine-specific or tyrosine-specific phosphatases both established the functional importance for phosphorylations even prior to the identification of the kinases and phosphatases that regulate these proteins. The most direct method for detecting protein phosphorylation is clearly metabolic labeling of cells with 32Pi (see UNIT 13.2) and subsequent isolation of the 32P-labeled substrate protein. However, it is often difficult to develop a rapid purification procedure for the specific protein of interest without selective reagents such as monospecific antibodies or other affinity ligands. Significant labeling of proteins is also a problem given the low abundance of many regulatory proteins and the slow turnover of their protein-bound phosphate under “basal” conditions. In cases where [32P]phosphate can be readily incorporated into the protein, in vitro
dephosphorylation by selected phosphatases can establish the functional importance of the modification. Ability to correlate loss of [32P]phosphate from a specific polypeptide with a functional change provides the best evidence for the physiological importance of protein phosphorylation. Initial experiments should use a general phosphatase such as potato acid phosphatase or calf intestine alkaline phosphatase which can successfully dephosphorylate a wide variety of phosphoproteins modified on several different amino acids. It should be noted that acid phosphatases from animal tissues (e.g., bone, prostate, liver, and heart) represent a class of lowmolecular-weight (18,000- to 25,000-kDa) protein tyrosine phosphatases that selectively dephosphorylate phosphotyrosine residues. These enzymes were defined as acid phosphatases because they hydrolyzed p-nitrophenyl phosphate (PNPP) at a pH optimum between 4.0 and 5.0. However, at physiological pH, these enzymes appear to specifically dephosphorylate proteins and peptides containing phosphotyrosine residues. The mammalian acid phosphatases show no significant activity against phosphoserine/phosphothreonine-containing proteins. With phosphoprotein substrates, potato acid phosphatase shows a preference for dephosphorylating phosphotyrosine residues, followed by phosphothreonine residues. By comparison, this enzyme shows very low activity against phosphoserine-containing substrates. Wheat germ acid phosphatase can dephosphorylate phosphoserines but still shows significantly higher activity against phosphotyrosine and phosphothreonine (Van Etten and Waymack, 1991). In this regard, alkaline phosphatase is often preferred over potato or wheat germ acid phosphatase for dephosphorylation of phosphoserine, the predominant protein modification in eukaryotic cells. When a general phosphatase, such as alkaline phosphatase, is used, effects independent of protein dephosphorylation must be considered. For instance, alkaline phosphatase hydrolyses ATP and inhibits ATP-dependent processes, such as protein kinases and ATP-dependent ion channels that are also inhibited by protein dephosphorylation (Berger et al., 1993). To distinguish between these differing effects, use of immobilized alkaline phosphatase is recommended. Agarose or dextran beads covalently linked to alkaline phosphatase should be washed extensively with the assay buffer to remove storage solutions and any
13.5.6 Current Protocols in Protein Science
unbound alkaline phosphatase. Following incubation with the substrate protein, immobilized alkaline phosphatase can be removed by centrifugation or filtration. Functional analysis of the substrate can then be undertaken without concern for the potential effects of the phosphatase on such analysis. However, if the substrate is present in a particulate fraction (e.g., in subcellular organelles), it cannot be easily separated from the beads by centrifugation or filtration. In that case, the phosphatase should be inhibited with a general inhibitor such as sodium pyrophosphate. In contrast to the general phosphatases, protein phosphatases are inefficient at hydrolyzing ATP and do not hinder analysis of ATP-dependent processes. Once phosphorylation of a substrate is established using a general phosphatase, the next step is to use protein phosphatases that have much higher activities against phosphoprotein substrates than either acid or alkaline phosphatase and are highly selective for specific phosphoamino acids. Protein phosphatases 1, 2A, and 2B (PP1, PP2A, and PP2B) represent three major classes of protein serine/threonine phosphatases in eukaryotic cells. These enzymes differ in their subunit structure, substrate specificity, and regulation by divalent cations. PP1 and PP2A account for >90% of the total protein serine/threonine phosphatase activity in many cell extracts (Shenolikar and Nairn, 1991) and demonstrate a broad in vitro substrate specificity. Therefore, PP1 and PP2A are the most commonly used reagents for in vitro dephosphorylation of phosphoserine- and phosphothreonine-containing proteins. PP1 and PP2A are inhibited by microcystin-LR as well as several other recently discovered inhibitors, including okadaic acid, calyculin A, and tautomycin, albeit with differing IC50 values. Inhibitors, such as okadaic acid (with IC50 for PP2A of 0.1 to 1.0 nM and for PP1 of 10 to 100 nM), calyculin A (with an IC50 for PP1 of 1 nM and for PP2A of 10 nM), and microcystin-LR (with an IC50 for both PP1 and PP2A of 0.1 nM) are used to exclude nonspecific effects of PP1 and PP2A. These inhibitors represent important experimental tools for establishing the physiological importance of reversible protein phosphorylation. PP2B, also called calcineurin, is a Ca2+/calmodulin-dependent phosphatase and demonstrates a narrow in vitro substrate specificity compared to PP1 or PP2A. The enzyme purified from mammalian tissues consists of a catalytic subunit that is tightly associated with a calcium-binding regulatory subunit. PP2B has
no activity in the absence of 1 µM calmodulin and 1 mM CaCl2 and can be activated by either Ca2+/calmodulin or 1 mM MnCl2. Many highly purified PP2B preparations show no activity in the presence of Ca2+/calmodulin but are fully activated by Mn2+. The immunosuppressive drugs cyclosporin A and FK506 selectively inhibit PP2B in vivo and in vitro. PP2B in the presence of Ni2+ ions and PP2A in association with an endogenous regulator protein or viral antigens can dephosphorylate proteins and peptides containing phosphotyrosines. However, under the conditions described in Basic Protocol 2, PP1, PP2A, and PP2B are highly selective for dephosphorylation of phosphoserine and phosphothreonine residues. PP1 and PP2A represent the major phosphohistidine phosphatase activity measured in tissue extracts, but the regulatory importance of this modification has not yet been established.
Critical Parameters Some commercial preparations of potato acid phosphatase contain contaminating protease activity. Because phosphorylation sites reside near the surface of the substrate protein, they are often accessible to proteases, so functional changes that result from limited proteolysis can be incorrectly attributed to dephosphorylation. To limit the effects of proteases, high enzyme concentrations or prolonged incubations with the general phosphatases should be avoided. Aside from including protease inhibitors in the reaction, two strategies should be considered to distinguish proteolysis from dephosphorylation. Formation of a complex b etween tr ich lor oacetic acid–soluble [32P]phosphate and ammonium molybdate (Support Protocol) is particularly useful in distinguishing [32P]phosphate from 32P-labeled phosphopeptides. The use of phosphatase inhibitors in control reactions also identifies those effects that can be attributed to reactions in which there are contaminating proteolytic enzymes. Commercial preparations of PP2A and PP1 contain only the free catalytic subunits. In vivo these subunits associate with regulatory subunits, which modulate their substrate specificity. However, the catalytic subunits are the most readily purified to homogeneity and can be stored for prolonged periods without significant changes in their enzymatic properties. Both PP1 and PP2A catalytic subunits retain high activity in the absence of divalent cations; PP2A activity may be further stimulated by 1
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mM Mn2+ in the enzyme reaction. Long-term storage of PP1 and PP2A results in an apparent loss of enzyme activity. However, these inactive enzymes can be fully reactivated by including 1 mM MnCl2 in the assay. The Mn2+-dependent PP1 enzyme shows slightly altered substrate specificity when compared with PP1 purified from tissues. Moreover, PP1 is potentially inhibited by several endogenous inhibitors. Thus, PP2A may be the preferred reagent for in vitro dephosphorylation of substrates in many cell extracts.
Troubleshooting The absence of a functional change following incubation of the substrate with one or more phosphatases may indicate either that the protein was not phosphorylated or that the dephosphorylation of existing phosphates has no effect on protein function. It is also possible that components in the reaction mixture inhibited the phosphatases. The latter possibility can be readily established if the substrate protein has been metabolically labeled with [32P]phosphate. SDS-PAGE followed by autoradiography or phosphoimage analysis can directly monitor the time-dependent loss of radiolabel from substrate protein. Coincidence between the removal of radiolabeled phosphate and change of function of the substrate protein can be used to distinguish between important regulatory modifications and “silent” phosphorylations. In analyses of unlabeled substrates, the reaction should include an internal standard or a known 32P-labeled protein or peptide. Dephosphorylation of the radiolabeled marker in the presence or absence of the sample should establish the extent to which activity of the added phosphatase has been compromised by components in the sample. If several different phosphatases fail to elicit a change in protein function under conditions that dephosphorylate the marker, the substrate protein may contain no protein-bound phosphate or phosphorylation may not be functionally important for the protein.
Anticipated Results
Detection of Phosphorylation by Enzymatics Technique
Most phosphoproteins should be dephosphorylated by the general phosphatases, leading to complete removal of protein-bound phosphate and functional changes indicating that reversible phosphorylation controls the protein function. In a number of cases, meta-
bolic labeling and phosphoamino acid analysis or the use of specific protein (phosphoserine/phosphothreonine or phosphotyrosine) phosphatases has successfully identified the amino acids modified. Such studies have then located the phosphorylated residues, especially in proteins whose primary structures have been determined by amino acid sequencing or cDNA cloning. A combination of biochemical studies and site-directed mutagenesis has then been used to establish the functional role of phosphorylation at individual sites. Increasing evidence points to the presence of both positive and negative regulatory phosphorylations in proteins. Thus, it is difficult to predict the functional outcome of complete in vitro dephosphorylation of many substrates.
Time Considerations Approximately 1 hr is required to complete the dephosphorylation reaction. SDS-PAGE and analysis of the gel require 1.5 hr. The time required for functional analysis depends on the particular assay used.
Literature Cited Berger, H.A., Travis, S.M., and Welsh, M.J. 1993. Regulation of the cystic fibrosis transmembrane conductance regulator Cl− channel by specific protein kinases and protein phosphatases. J. Biol. Chem. 268:2037-2047. Charbonneau, H. and Tonks, N.K. 1992. 1002 protein phosphatases? Annu. Rev. Cell Biol. 8:463493. Shenolikar, S. 1994. Protein serine/threonine phosphatases: New avenues for cell regulation. Annu. Rev. Cell Biol. 10:55-86. Shenolikar, S. and Ingebritsen, T.S. 1984. Protein (serine and threonine) phosphate phosphatases. Methods Enzymol. 107:102-129. Shenolikar, S. and Nairn, A.C. 1991. Protein phosphatases: Recent progress. Adv. Second Messenger Phosphoprotein Res. 23:1-123. Swarup, G., Cohen, S., and Garbers, D.L. 1981. Selective dephosphorylation of proteins containing phosphotyrosine by alkaline phosphatases. J. Biol. Chem. 256:8197-8201. Van Etten, R.L. and Waymack, P.P. 1991. Substrate specificity and pH dependence of homogenous wheat germ acid phosphatase. Arch. Biochem. Biophys. 288:634-645.
Contributed by Shirish Shenolikar Duke University Medical Center Durham, North Carolina
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Preparation and Application of Polyclonal and Monoclonal Sequence-Specific Anti-Phosphoamino Acid Antibodies
UNIT 13.6
Sequence-specific anti-phosphoamino acid antibodies provide powerful new tools to study phosphorylation events within cells and can provide new insights into protein function, including mechanisms of signal transduction, cell differentiation, cell growth, cell death, and a multitude of other physiologically important molecular events. These complex processes involve interaction of proteins and enzymatic changes in which phosphorylation of Ser, Thr, and Tyr residues play critical roles. Thus, unlike conventional antibodies, sequence-specific anti-phosphopeptide antibodies provide information regarding not only the abundance of a protein, but also its functional state. Unlike general anti-phosphoamino acid antibodies (e.g., commercially available anti-phosphotyrosine antibodies), which have broad reactivity against a protein phosphorylated at any number of phosphoamino acids, sequence-specific anti-phosphoamino acid antibodies have unique specificity toward a modified protein. Such antibodies not only facilitate structural analyses of phosphoproteins, but also allow heretofore impossible applications; e.g., differential isolation of a particular protein that has been phosphorylated at a particular site, as well as analysis of the functional state of a protein in situ by immunohistochemical protocols. Traditional methods of analysis of protein kinases and their substrates involve in vitro protein kinase assays (UNIT 13.7), 2-D tryptic phosphopeptide mapping (UNIT 13.9), phospho-amino acid analysis (UNIT 13.3), and phosphopeptide sequencing. Protein kinase assays involve phosphorylation of amino acid sequences that may not always reflect the phosphorylation events that occur inside the cell. Investigation of the phosphorylation of specific Tyr or Ser residues within cells requires phospho-amino acid analysis and 2-D tryptic peptide mapping of proteins labeled either by in vitro kinase assays or in vivo [32P]orthophosphate labeling. In addition, phosphopeptide sequencing is needed to identify the exact phosphorylation site. None of these techniques is easy to master and there are technical problems that frequently cannot be solved, e.g., large peptides may be insoluble and thus cannot be separated (see below). The use of a general anti-pTyr antibody (e.g., commercial monoclonal antibody 4G10 from Upstate Biotech) provides an easy and fast way to analyze Tyr phosphorylation in vivo. However, it cannot be used for the analysis of phosphorylation of a particular Tyr of a specific protein. Anti-pTyr peptide antibodies have been developed against phosphopeptides derived from proteins to overcome this problem (Bangalore et al., 1992; Epstein et al., 1992). Anti-pSer peptide antibodies have also been developed (Coghlan et al., 1994; Ishida et al., 1996; Alberta and Stiles, 1997; Banin et al., 1998; Canman et al., 1998; Bodnar et al., 1999). These antibodies are very valuable reagents for the study of protein kinases and their target proteins, either alone or in combination with traditional methods. For example, specific anti-pSer peptide antibodies have been used to study the in vivo phosphorylation of Ser-15 of p53 (Alberta et al., 1997; Banin et al., 1998 ; Canman et al., 1998). There are commercial suppliers of sequence-specific anti-phosphoamino acid antibodies, e.g., Sigma and Bethyl Laboratories. The primary advantage of sequence-specific, anti-phosphoamino acid antibodies is that they permit rapid analysis of intracellular phosphoproteins without the need to employ complex structural protein analyses (e.g., protease digestion of orthophosphate-labeled proteins). Such biochemical analyses are fraught with difficulties including the use of high levels of [32P]orthophosphate to label cellular proteins. Besides the health risk of Contributed by Tong Sun and Ralph B. Arlinghaus Current Protocols in Protein Science (2003) 13.6.1-13.6.27 Copyright © 2003 by John Wiley & Sons, Inc.
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exposure to beta particles, labeling of cells with orthophosphate can produce artifacts that affect cellular events. Moreover, the analyses are not always productive due to a number of inherent problems. These include: 1. incomplete digestion of the protein under study (yielding insoluble peptide fragments that contain the phosphorylation site of interest); 2. removal of the phosphate moiety by phosphatases during cell lysis and the immunoprecipitation steps required to isolate the protein under study; 3. difficulties in identifying the protein fragment that contains the phosphoamino acid residue under study; 4. once the correct fragment is identified, obtaining an appropriate size fragment that can be sequenced is often problematic; and 5. automated Edman degradation methods (UNIT 13.9) do not identify phosphoamino acids residues released in the sequencing steps. This unit includes a section of Strategic Planning, which describes the overall steps of producing sequence-specific anti-phosphoamino acid antibodies, including polyclonal and monoclonal antibodies. It covers details of producing polyclonal sequence-specific anti-phosphoamino acid antibodies and discusses general issues such as immunogen selection and peptide crosslinking. Basic Protocol 1 provides a detailed protocol for affinity purification of polyclonal sequence-specific anti-phosphoamino acid antibodies. Basic Protocol 2 provides procedures for production and purification of monoclonal sequence-specific anti-phosphoamino acid antibodies that are not covered in Basic Protocol 1. For production of monoclonal antibodies, ELISAs are performed to screen candidate hybridoma culture fluid against the cognate phosphopeptide, as well as against related phosphorylated and nonphosphorylated peptides, until one is found with the desired reactivity and specificity. Although monoclonal antibodies have a number of advantages, production of polyclonal antibodies is likely to be more predictable, as a monoclonal hybridoma clone of stringent specificity may occur with very low frequency. Production of monoclonal antibodies is also generally more time-consuming and expensive. The relative merits of monoclonal and polyclonal antibodies are discussed in other general references (also see Key References). Support Protocols 1 and 2 cover procedures for cross-linking peptides or phosphotyrosine, repectively, to an affinity matrix. The Commentary addresses and discusses some important issues in the production of these antibodies; and finally, an example of production and application of sequence-specific anti-phosphoamino acid antibodies is provided. STRATEGIC PLANNING Generation of site-specific, sequence-specific phosphoamino acid antibodies is relatively simple and straightforward. The procedure involves the following steps: selecting the phosphopeptide sequence to be used as an immunogen, cross-linking it to a carrier protein, injecting the cross-linked peptide/carrier complex emulsified in oil/water adjuvant into rabbits or mice, and characterizing the resulting antiserum for reactivity to the peptide immunogen (see Fig. 13.6.1 and 13.6.2 for flow charts that outline the steps required to generate and characterize polyclonal and monoclonal sequence-specific anti-phosphoamino acid antibodies). An example of the use of crude serum to obtain useful data is described in Anticipated Results.
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design and synthesize pTyr or pSer peptide and its cognate non-phospeptide; coupling peptide to carrier protein (core facility/Support Protocol 1/Strategic Planning)
immunize rabbits to produce crude antiserum
test serum by ELISA with phospho- and nonphosphopetides
concentrate antibody by ammonium sulfate precipitation or protein A/protein G affinity binding and elution (optional)
peptide blocking strategy
affinity purification strategy
remove the non-specific antibodies by passing through negative selection affinity matrices, such as, carrier protein affinity matrix, phosphotyrosine affinity matrix, cognate nonphospho-peptide afffinity matrix, and test the specificity of the Ab homologous phosphopeptide affinity matrix
test serum by immunoprecipitation or immunoblotting with wild type and target Tyr or Ser mutant proteins, either alone, with purify the antibody by phospho- or nonphosphobinding the antibody to peptide blocking; as an positive selection affinity alternative testing method test the specificity of the Ab matrices containing target if mutant protein is not available, phospho-peptide, elute test with wild type protein the antibody and dialzye and dephosphorylated protein against PBS/azide
application use the antiserum in immunoblotting, immunopreciptiation, immunohistochemistry, immunofluorescence, without blocking with phospho- or cognate nonphosphopeptide
application use the purified antibody in immunoblotting, immunopreciptiation, immunohistochemistry, immunofluorescence, without blocking with phospho- or cognate nonphosphopeptide
Figure 13.6.1 Flow chart of the steps involved in production of polyclonal phosphoamino acid sequence-specific antibodies (see Basic Protocol 1). Post-Translational Modifications: Phosphorylation and Phosphates
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design and synthesize pTyr or pSer peptide and its cognate non-phosphopeptide; couple peptide to carrier protein (core facility/Support Protocol 1/Strategic Planning)
immunize mice to produce crude antiserum
test serum by ELISA with phospho- and nonphosphopeptides
test serum by immunoprecipitation or immunoblotting with wild type and target Tyr or Ser mutant proteins, either alone, with phospho- or nonphosphopeptide blocking; as an alternative testing method if mutant protein is not available, test with wild type protein and dephosphorylated protein select positive and relatively specific serum-producing mice produce hybridoma cell line using spleen cells from the selected mice Basic Protocol 2 screen cell lines by ELISA with phospho- and nonphosphopeptides
screen cell lines by immunoprecipitation or immunoblotting with wild type and target Tyr or Ser mutant proteins, either alone, or with phosphopeptide blocking; as an alternative testing method if mutant protein is not available, test with wild type protein and dephosphorylated protein
screen cell lines by immunohistochemistry, immunofluorescence, alone or blocking with phosphopeptide (optional)
expand and freeze positive and specific cell lines
optional processing: subclones may be screened using above tests to select optimal clones; perform isotype analysis; perform large-scale production and purification if necessary
application use crude hybridoma cell culture fluid, ascites, or purified antibody in immunoblotting, immunoprecipitation, immunohistochemistry, immunofluorescence, without blocking peptides
Sequence-Specific Anti-Phosphoamino Acid Antibodies
Figure 13.6.2 Flow chart of the steps involved in production of monoclonal sequence-specific anti-phosphoamino acid antibodies (see Basic Protocol 2).
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Protein Versus Peptide Immunization For conventional antibody generation, there are typically two major strategies. The first is direct immunization of animals with purified native protein or recombinant protein. The other strategy involves immunizing animals with a synthetic short peptide. Both strategies have advantages and disadvantages. The first strategy normally generates antibodies with higher titers because multiple B cell clones can be activated targeting multiple epitopes in their natural conformation in the protein. In contrast, anti-peptide antibodies usually exhibit higher specificity and are very useful in detecting specific domains of the protein of interest, albeit with lower titers. This is due to the fact that the short peptide sequence can constitute one or very few epitopes, and the conformation of the synthetic short peptide may not resemble that of the same sequence in the native protein. However, the anti-peptide antibody strategy is better suited for production of sequence-specific phosphoamino acid antibodies because (1) immunizing animals with synthetic short peptides containing target phosphoamino acids will generate more specific antibodies targeting the defined short sequence; and (2) the analogous phosphorylated peptide sequences in the native protein are likely to be exposed on the surface of the protein because of their accessibility to kinases and because they bear a charged phosphate group. The phosphopeptide in the native protein is therefore likely to be in a conformation recognizable by the anti-phosphopeptide antibodies. Select a Peptide that Contains the Phosphoamino Acid of Interest A peptide containing 12 to 15 amino acids with the phosphorylation site Tyr or Ser in the central part of the sequence is typically employed. This sequence includes amino acids from the actual gene sequence of the protein plus an N-terminal or C-terminal cysteine to enhance coupling of the peptide to a carrier protein, e.g., bovine serum albumin (BSA) or keyhole limpet hemocyanin (KLH; see UNIT 18.3 for peptide-carrier protein coupling methods). This linker amino acid is usually added to a selected peptide sequence, but may be naturally occurring in the sequence and does not have to be at a terminus. Because the method of cross-linking the peptide to a carrier protein is critical in generating a high titer, specific antibody, an appropriate cross-linking amino acid residue must be included when designing a peptide immunogen. More detailed discussion on cross-linking methods is given below (see Cross-Link the Phosphopeptide to a Carrier Protein). Synthesize Selected Peptide with Phosphate Coupled to the Appropriate Ser/Thr or Tyr Peptides are synthesized with phosphate coupled to the appropriate Ser/Thr or Tyr. Many institutions have core facilities for synthesis of peptides using automated solid-phase peptide synthesizers. Once synthesized, peptides are typically purified by HPLC and analyzed by mass spectrometry prior to use. Currently, most phosphotyrosyl-containing peptides are synthesized with 9-fluorenylmethyloxycarbonyl (Fmoc) phosphotyrosine. The lack of a side chain protecting group on the phosphotyrosine (Kitas et al., 1994) allows standard Fmoc synthesis and cleavage procedures to be employed (Chang and Meienhofer, 1978). However, a higher molar excess of activated Fmoc amino acids than that used in standard procedures may be required for efficient coupling of amino acids after unprotected phosphotyrosine has been added. In some circumstances, a double coupling procedure may be employed to add the few amino acids following phosphotyrosine to the growing peptide chain. This alternative requires the sequential, repeated addition of the next particular amino acid to be added to the peptide. A protocol for the coupling of peptides to an affinity matrix is described in Support Protocol 1, and general methods for coupling peptides to carrier proteins are described in UNIT 18.3.
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Unphosphorylated peptides should also be synthesized. They are used in immunoblots to block the nonspecific antibodies raised against the non-phosphorylated sequences. Phosphorylated peptides can also be used as blocking peptides to confirm the specific affinity for the desired phosphorylated sequence. In addition, the Ser→Ala, Thr→Ala, or Tyr→Phe mutant protein should be expressed and purified for use in assays for estimating the titer of the immune response in the sera of the immunized rabbits. Convenient screening assays for this purpose include tests for the reactivity of crude sera by immunoblotting of lysates expressing the wild-type protein or the Tyr→Phe or Ser→Ala mutant protein. Cross-Link the Phosphopeptide to a Carrier Protein The phosphopeptide must be cross-linked to a suitable carrier protein, e.g., KLH or BSA, to elicit a strong T cell response and a high titer, anti-phosphopeptide antibody. The method of cross-linking appears to be critical. An appropriate method should be selected with consideration of the sequence of the peptide. Typically, there are three ways to cross-link a peptide to a carrier protein (UNIT 18.3): (1) glutaraldehyde cross-links the peptide N-terminal α-amino group or the ε-amino group of either a native or added Lys to the amino groups on the carrier protein; (2) cross-link the sulfhydryl group of an added N- or C- terminal or native Cys in the peptide to amino groups on the carrier protein; or (3) carbodiimides cross-link free carboxyl and amino groups on the C- or N-terminal and side chains of Lys, Asp, and Glu to form amide bonds. Amide bonds are extremely rigid and the carbodiimide cross-linking is not specific, therefore, the third method is less frequently used. It is important not to cross-link with agents that will alter the sequence immediately surrounding the phosphoamino acid. Thus, if Lys residues are near the phosphoamino acid site, the use of glutaraldehyde to cross-link the peptide to the carrier protein might compromise the antigenic site. A similar concern should be applied to the design of the peptide if Cys cross-linking is to be used. The average occurrence of Lys in proteins is much higher than Cys, 5.8% to 1.8%, repectively. Cys cross-linking is preferred if the peptide sequence does not contain a native Cys. Typically, an N- or C-terminal Cys is added for cross-linking purposes. For example, to raise the anti-pSer-354 Bcr antibody, the peptide GQSRVpSPSPTTY(C) with an added C-terminal Cys was used as the immunogen (see Anticipated Results for further discussion of anti-Bcr antibody generation and characterization). If a peptide with more than one Cys and Lys in the natural sequence has to be used as the immunogen, the authors suggest to use a conjugation method to crosslink the peptide to the carrier protein in a way that least affects the phosphoamino acid region. For generation and characterization of monoclonal antibodies, a different carrier protein must be employed for immunization than will be used as substrate for the ELISA screen. This is done to avoid detection of anti-carrier antibodies in the ELISA. BSA is used for screening because many of the clones are likely to recognize the KLH carrier used in the immunization of the mice and because unconjugated peptides have not been found to function well as ELISA substrates (Doolittle, 1986). In addition, because antibodies may be generated against the cross-linking reagent, the method used for coupling the phosphopeptide to the BSA to produce the substrate for ELISAs should be different from that used to link peptides to the KLH carrier for immunization (Doolittle, 1986; Czernik et al., 1991; UNIT 18.3).
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For cross-linking to KLH. The peptide is coupled to KLH using the bifunctional reagent, N-hydroxy-succinimide ester of N-(carboxy cyclohexyl-methyl)-maleimide (smcc; Sigma) in a two-step procedure. First, KLH is reacted with smcc, coupling amines of KLH to the succinimide moiety of smcc. The smcc-modified KLH is then reacted (1 mg:1 mg) with the phosphopeptide containing a Cys, linking the maleimide of the smcc-KLH to the sulfhydryl of the phosphopeptide Cys. Excess unlinked peptide is removed by gel filtration. This method offers several advantages over alternative coupling methods such as glutaraldehyde or a carbodiimide. (1) KLH is modified, removing excess free smcc, to produce a relatively stable activated carrier that will not crosslink itself, thereby avoiding aggregation. Aggregation can produce an insoluble, particulate preparation; such preparations exhibit poor immunogen qualities. (2) The peptide reacts with the activated KLH at a single cysteine sulfhydryl group, thereby avoiding peptide-to-peptide “scaffolding.” Such scaffolding may affect the proper loading of peptide onto the carrier, which is necessary for optimal immunogenicity. (3) Linking the peptide through a single coupling site allows relatively unrestricted chain folding that mimics folding of the native protein, thereby enhancing cross-reactivity of the anti-peptide antibodies with the native protein. (4) Maximum conjugation to the carrier occurs with a molar excess of peptide, at neutral pH, in a brief (2-hr) incubation period. These conditions are compatible with modified amino acids that may be unstable at acidic or basic pH. The reaction is compatible with DMSO or SDS and with low or high pH if required for solubility. The concentration of DMSO and SDS will not affect the reaction, therefore, whatever concentration of DMSO and SDS that can solubilize the peptide is acceptable. The authors prefer to use a pH range of 6.5 to 8.5. (5) Maximum carrier loading can be effected with peptides of essentially any amino acid sequence; the solubility of the peptide is essentially the only limitation of the reaction. The smcc bifunctional conjugation provides a very reproducible method applicable to any peptide except those with multiple (≥2) cysteine residues and those with an internal cysteine that is close to the phosphoamino acid epitope. Immunogenicity of peptide/KLH conjugates has been superior to glutaraldehyde and carbodiimide (EDC) conjugates in comparative studies (Arlinghaus, unpub. observ.). For cross-linking to BSA. To prepare a phosphopeptide-BSA carrier conjugate for immunization of rabbits, couple a 30-fold molar excess of phosphopeptide (e.g., 15.3 mg of phosphoserine peptide J27, mol. wt. 1705, see Table 13.6.1) dissolved in water and neutralized with 1 N NaOH) to BSA (20 mg dissolved in 0.4 M sodium phosphate buffer, pH 7.5), using glutaraldehyde as the cross-linking reagent as described by Doolittle (1986). The reaction is allowed to proceed for 30 min at room temperature; the development of a yellow color in the coupling solution is an indication that the glutaraldehyde has reacted. General methods for coupling of peptides to carrier proteins for use in immunization are described in UNIT 18.3. Conjugation can be confirmed by performing an anti-phosphotyrosine immunoblot (UNITS 10.7 & 10.10) on a small aliquot of the conjugate (although a smear should be expected on the blot, resulting from variation in the amount of peptide coupled per BSA molecule, as well as from possible multimers of BSA).
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Immunize Animals with Cross-Linked Phosphopeptide to Generate a Useful Antibody The next step is to raise antisera in rabbits or a suitable animal (e.g., goat). Guidelines for immunization of rabbits are provided by Grant (2003), and many institutions have core facilities for the immunization of rabbits and production of antisera. The immunogen (the peptide-carrier conjugate dissolved in PBS) is combined 1:1 with complete Freund’s adjuvant (CFA) to deliver a total of 0.1 to 1 mg of conjugate protein per 1-ml injection. This is administered subcutaneously at multiple sites. For subsequent booster injections, mix the antigen 1:1 with incomplete Freund’s adjuvant (IFA) and administer at 2-week intervals. Collect blood 1 week after injection, beginning with the third injection. An adequate immune response can be seen as early as week 6. Screen Serum by ELISA for the Presence of Useful Antibodies Titers of antisera are tested by ELISA with phosphopeptides (see Basic Protocol 1 for special concerns related to sequence-specific anti-phosphoamino acid antibodies). Purify the Anti-Phosphopeptide Antibodies (optional) The purification of polyclonal antibodies consists of multiple affinity-chromatography steps for negative and positive selection of antibodies of the appropriate reactivity (see Basic Protocol 1 and Fig. 13.6.1). All of the chromatographic steps may be carried out at room temperature. Determine the Specificity of Antiserum by Immunoblotting The specificity of the final antibody can be demonstrated most simply by immunoblot analysis (UNIT 10.10) using a panel of relevant phosphorylated and nonphosphorylated proteins and peptides. Electrophoresis is carried out with purified cognate protein in both its phosphorylated and unphosphorylated states, as well as with any proteins toward which there could conceivably be cross-reactivity (e.g., the Ser→Ala, Thr→Ala, or Tyr→Phe mutant proteins). Probing immunoblots of wild type and mutant proteins with the purified anti-phosphopeptide antibody will show whether other Tyr or Ser/Thr sites within the selected protein are recognized by the antibody. This immunoblot assay may be used after each chromatographic step to monitor the progress of the purification. An example of this type of cross-reactivity with anti-Bcr pSer-354 antibody is illustrated in Anticipated Results.
Sequence-Specific Anti-Phosphoamino Acid Antibodies
Determine and Deplete Cross-Reactivity In the production and characterization of polyclonal anti-phosphopeptide antibodies, cross-reactivity is a major issue. Cross-reactivity is most likely to occur when the target protein is a member of a family of closely related homologous proteins—e.g., of the subfamilies of tyrosine protein kinases. For example, a monoclonal anti-phosphopeptide antibody with specificity for the Tyr-1248 autophosphorylation site of Neu has been produced (DiGiovanna and Stern, 1995). A homologous site exists in the epidermal growth factor receptor (EGFR). The anti-Neu antibody mentioned above does not cross-react with the homologous EGFR site, despite having identical residues in 7 of the 14 amino acids of the peptide sequence (5 of which are N-terminal to the phosphotyrosine, 1 is C-terminal, and the phosphotyrosine itself). Thus, specificity is achievable even with at least up to 50% identity. Nevertheless, such a close relationship among the phosphorylation sites of different proteins highlights the importance of considering all known related proteins. Cross-reactive proteins are usually most easily removed by immunoprecipitation (UNIT 9.8) using conventional antibodies. Blocking (pre-incubation of) the antiserum with either the non-phosphorylated peptide or phosphorylated peptide can be performed to determine the utility of the antiserum.
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Another consideration for depleting cross-reactivity is the quantity of cross-reactive antibodies in a given aliquot of anti-serum relative to the capacity of the peptide columns used for the negative selection. Because of the expense of the peptides, it is recommended to prepare small columns (with 2- to 3-ml bed volumes) and pass the sera over the columns multiple times, empirically determining the number of passes necessary to quantitatively deplete cross-reacting material. In these columns, 3 µmol of ligand is coupled per milliliter of Affi-Gel 10. The manufacturer states that the resin contains ∼15 µmol of active ester per milliliter of gel, and that the gel has a capacity for 35 mg of protein (or 15 to 20 µmol of a low-molecular-weight ligand) per milliliter of gel. The BSA and phosphotyrosine columns are relatively inexpensive to produce; thus, a single large column of each is practical and sufficient. AFFINITY PURIFICATION OF POLYCLONAL SEQUENCE-SPECIFIC ANTI-PHOSPHOPEPTIDE ANTIBODIES For the purification of polyclonal antibodies, a multi-step chromatographic purification procedure with several negative- and positive-selection steps is performed to produce a final antibody preparation having the desired reactivity and specificity (Fig. 13.6.1). Note that the positive-selection procedures usually denature the antibody, requiring an inefficient renaturation procedure, often resulting in reduction of the antibody titer and potentially altered specificity.
BASIC PROTOCOL 1
A series of negative-selection affinity-chromatography columns are used first to adsorb antibodies from the serum that cross-react with epitopes other than the cognate phosphopeptide. These negative-selection columns are composed of matrix coupled to various synthetic peptides (see Support Protocol 1)—principally the cognate non-phosphorylated peptide (to remove phosphorylation-independent antibodies) and phosphotyrosine itself (to remove indiscriminant anti-phosphotyrosine reactivity; see Support Protocol 2 for preparation of phosphotyrosine affinity columns). Additional negative-selection steps may be used if needed, depending on the target chosen, to remove antibodies that cross-react with phosphorylation sites of other proteins having known homology to the target site. This cross-reactivity occurs particularly when the target protein is a member of a family of homologous proteins, e.g., a subgroup of related receptor tyrosine protein kinases. If such cross-reactivity is anticipated, it is possible to synthesize not only the cognate phosphorylated and nonphosphorylated peptides, but also phosphopeptides based upon the related homologous sequences, to be used for negative selection. The serum is applied to a column consisting of immobilized carrier protein (commercially available BSA-agarose) to remove the majority of the antibodies generated against the much larger carrier protein. The cognate phosphopeptide is also immobilized on an affinity-chromatography column support (see Support Protocol 1), and this affinity matrix is ultimately used to purify the antibody from the rabbit serum. Thus, a typical purification scheme might consist of the following: Negative-selection affinity-purification columns • BSA (carrier) • phosphotyrosine • cognate nonphosphopeptide • homologous phosphopeptides (if desired) Positive-selection affinity-purification column • cognate phosphopeptide.
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Materials BSA-agarose affinity matrix (Sigma) packed (see Support Protocol 1) in a 10-ml bed volume column Phosphotyrosine affinity matrix column (10-ml bed volume; see Support Protocol 2), if an anti-phosphotyrosine antibody is being made Clarified, crude antiserum from rabbit immunized with phosphopeptide-carrier conjugate PBS/azide: PBS (APPENDIX 2E) containing 0.02% (w/v) sodium azide (store indefinitely at 4°C or room temperature) 3 M NaSCN Cognate nonphosphopeptide affinity matrix column (3-ml bed volume; see Support Protocol 1 and Strategic Planning) Homologous phosphopeptide affinity matrix columns (optional; 3-ml bed volume; see Support Protocol 1 and Strategic Planning) Positive-selection phosphopeptide affinity matrix column (3-ml bed volume; see Support Protocol 1 and Strategic Planning) 3.5 M and 4.5 M MgCl2 (optional) Spectrophotometer (optional) Dialysis tubing (MWCO 12,000 to 14,000; 10-mm width, 6.4-mm diameter; e.g., Spectra/Por 4 from Spectrum) Additional reagents and equipment for dialysis (UNIT 4.4), protein quantitation (UNITS 3.1 & 3.4), and analysis of antibodies by ELISA or immunoblotting (UNIT 10.10) Deplete reactivity of serum toward phosphotyrosine and carrier protein 1. Connect the washed BSA-agarose and phosphotyrosine affinity (if an anti-phosphotyrosine antibody is being made) matrix columns in series. 2. Pass ∼15 ml clarified crude antiserum through the two columns, allowing it to flow by gravity. Wash with PBS/azide until all of the yellow color of the serum has passed through the columns (or monitor the A280 of the column effluent spectrophotometrically until baseline absorbance is reached), then wash with an additional 5 to 10 ml PBS/azide, collecting all washings in the flowthrough fraction which will contain the antibody of interest. After the serum passes through, regenerate the columns by washing with 10 bed volumes of 3 M NaSCN followed by 10 bed volumes of PBS/azide, and store at 4°C in PBS/azide for future use. Aliquots of the crude serum as well as this first flow-through may be saved (at −20°C or 4°C with 0.05% sodium azide) for subsequent analysis and comparison with individual purification fractions. The serum may be analyzed at this time for elimination of anti-phosphotyrosine reactivity by immunoblotting (UNIT 10.10) samples containing a variety of phosphotyrosyl proteins; alternatively, it is possible to save an aliquot for later analysis and proceed to the next step (see Troubleshooting for a discussion of stepwise analyses). Expect that the volume of the sample will increase with each passage through a column. This will prolong the amount of time needed for subsequent columns. Note that if flowthrough is monitored by the passage of the yellow color of the serum, this will become more difficult as the serum becomes more dilute.
3. To deplete reactivity of serum toward unphosphorylated cognate peptide, pass the flow-through from the columns in step 2 through the cognate nonphosphopeptide affinity matrix column by gravity as many times as necessary to deplete cross-reactivity, regenerating the column between passes by washing with 10 bed volumes of 3 M NaSCN followed by 10 bed volumes PBS/azide. Sequence-Specific Anti-Phosphoamino Acid Antibodies
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A 15-ml starting serum sample may have to be passed over this column several times to quantitatively deplete cross-reactivity. The serum may be analyzed after each or several passes, or an aliquot can be saved for analysis at a later time.
4. If cross-reactivity with homologous phosphoprotein(s) is anticipated, pass the flowthrough through the homologous phosphopeptide affinity matrix column(s) using the methodology described in step 3. Refer to Strategic Planning for discussion of cross-reactivity with homologous phosphoproteins.
Purify antibodies 5. To purify antibodies by positive-selection affinity chromatography and dialysis, hydrate and wash ∼25-cm strips of dialysis tubing in advance for collection of fractions from the positive-selection affinity purification. Secure one end of each length of tubing with a dialysis clamp and check for leaks (UNIT 4.4). Also prepare 6 liters PBS/azide and cool to 4°C in advance for use as the dialysis solution. To minimize the time that the antibodies are exposed to the elution solution, fractions (∼3-ml each) from the positive-selection affinity column should be collected directly into prepared dialysis tubing.
6. Pass the flow-through from the previous chromatography step through the positiveselection phosphopeptide affinity column three times, without washing between passes. Three passes are performed to maximize the interaction of the antibody with the affinity matrix.
7. Collect the flow-through from the final pass, then wash the column with 5 to 20 ml PBS/azide (depending upon the pre-column volume and column bed volume) and combine the washings with the flow-through. Wash the column with an additional 20 ml PBS/azide, and collect the washings separately as the “wash” fraction. 8. Elute with chaotropic agent of choice: either 20 ml of 3 M NaSCN, or 10 ml of 3.5 M MgCl2, followed by 10 ml of 4.5 M MgCl2. Upon starting the elution, immediately begin collecting ∼3-ml fractions directly into the dialysis bags prepared in step 5, securing the proximal end of the tubing with dialysis clamps and dropping the bag immediately into the stirring 4°C PBS/azide dialysis solution. Collect at least six fractions. This elution step can be repeated one to three times for better recovery. Because antibodies are eluted from the positive-selection affinity column in strongly chaotropic solutions that are potentially deleterious to the stability of the antibody, it is highly recommended to collect the fractions in prepared dialysis tubing so that they may be immediately placed into the PBS dialysate, thus, minimizing the time that the antibodies are exposed to the eluting solutions. Although MgCl2 is thought to be “gentler,” the authors have found that gravity-driven flow rates from phosphopeptide columns eluted with this salt quickly become extremely slow, possibly because of precipitation of the salt in the column or an interaction of the Mg2+ ion with the phosphate groups. Thus, use of NaSCN is preferred, and this salt appears to permit recovery of comparable activity.
9. Dialyze all fractions collected in step 8 exhaustively against PBS/azide at 4°C (UNIT 4.4). Analyze and store purified antibody 10. Determine protein concentration of dialyzed fractions by measuring absorbance at 280 nm or by a colorimetric protein assay (UNIT 3.4) and calculate yield. Generally, ≥1 mg of purified antibody is recovered from a 15-ml serum sample.
11. Store the antibody in aliquots at −20°C or colder for long term storage; store working aliquots at 4°C with 0.05% sodium azide to prevent bacterial growth.
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12. Assay the reactivity and cross-reactivity of the final samples, as well as aliquots saved from previous steps, using ELISA or immunoblotting (UNIT 10.10). BASIC PROTOCOL 2
PRODUCTION OF MOUSE MONOCLONAL SEQUENCE-SPECIFIC ANTI-PHOSPHOPEPTIDE ANTIBODIES Similar to the method used for production of polyclonal anti-phosphopeptide antibodies (see Basic Protocol 1), the first step of the planning phase for making anti-phosphopeptide monoclonal antibodies is the design and synthesis of the required peptides (see Strategic Planning, Fig. 13.6.2, and Background Information). The same recommendations regarding design of the immunogen peptide apply, but a different carrier protein must be employed for immunization than will be used as substrate for the ELISA screen. This is done to avoid detection of anti-carrier antibodies in the ELISA. For example, KLH is used as the carrier for immunization and BSA as the carrier for the ELISA substrate. General methods for coupling peptides to carrier proteins for use in immunization are described in UNIT 18.3. The cognate phosphopeptide can be immobilized on an affinity-chromatographic column support (see Support Protocol 1) and used to affinity-purify the antibody produced by the desired hybridoma clone. A number of additional peptides must be produced for use in ELISA screening of candidate hybridoma supernatants, to screen both for the presence of the desired reactivity and the absence of undesired cross-reactivity.
Sequence-Specific Anti-Phosphoamino Acid Antibodies
The next step is to raise antibodies in mice. General guidelines for the immunization of mice and the production of monoclonal antibodies are described by Cooper and Paterson (2003), and many institutions have core facilities that will perform that task. The authors have immunized BALB/c mice by intraperitoneal injection with 1 mg/ml of immunogen (dissolved in PBS) in a 50% emulsion with complete Freunds adjuvant on day 1 and boosted intraperitoneally on days 15 and 37 with immunogen in incomplete Freunds adjuvant. Test bleeds from day 47 were analyzed by ELISA. On day 57, the mouse with the best titer was boosted intravenously, and its spleen was harvested on day 60. Freshly harvested spleen cells were prepared for cell fusion to generate hybridoma lines, which were subsequently screened by ELISA to identify clones that react with the cognate phosphopeptide and to eliminate clones that cross-react with other epitopes in addition to the cognate phosphopeptide. Epitopes screened by ELISA generally consist of the cognate nonphosphorylated peptide (to eliminate clones with phosphorylation-independent reactivities) and at least one unrelated phosphopeptide (to eliminate clones with indiscriminate anti-phosphotyrosine reactivity). In addition, depending upon the particular target chosen, it may be necessary to screen for reactivity with phosphopeptides from proteins of known homology. This occurs particularly when the target protein is a member of a family of homologous proteins—e.g., the subgroups of related receptor tyrosine protein kinases. If such cross-reactivity is anticipated, it is essential to synthesize not only the cognate phosphorylated and nonphosphorylated peptides and at least one unrelated phosphopeptide, but also phosphopeptides based upon the related homologous sequences. Thus, a typical ELISA screening scheme might consist of the following steps. (1) Screen for clones that exhibit reactivity against the cognate phosphopeptide. These clones, which will likely be <10% of all the clones, are potential candidates for the desired clones. (2) Screen clones from step 1 for reactivity against the cognate nonphosphopeptide. These clones, which are phosphorylation state-independent, are then eliminated. (3) Screen the resulting pool of clones for reactivity against at least one unrelated phosphopeptide. These clones, which are likely to represent indiscriminate anti-phosphotyrosine reactivity, are also eliminated. (4) If reactivity against potential cross-reacting homologous peptides is anticipated, screen the remaining clones as appropriate and eliminate those showing
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cross-reactivity. Monoclonal antibodies can also be screened by immunoblotting against COS1 cells expressing the wild-type and Y→F mutant or the S→A mutant. For ELISA screening, 96-well polyvinyl chloride assay plates are generally used and bound monoclonal antibody is detected using biotinylated horse anti-mouse secondary antibody followed by horseradish peroxidase (HRPO)-conjugated avidin D (Vector Labs) and 4-aminoantipyrine (Sigma)/H2O2 as the substrate. This system yields a deep red color in positive wells and no color in negative wells, allowing easy visual scoring without the need for quantitative measurements. The clones that pass all the screening steps are candidates for desirable hybridoma cell lines. These are expanded, frozen as a pool, subcloned by limiting dilution, then re-screened against cognate phosphopeptide to ensure continued production of antibody through the early passages. Those continuing to produce antibody are expanded further and a larger quantity of either culture supernatant or ascites fluid is screened by immunoassay. Reactivity is demonstrated most simply by immunoblot analysis (UNIT 10.10) using a panel of relevant phosphorylated and nonphosphorylated proteins. Electrophoresis is carried out with purified (generally by immunoprecipitation, UNIT 9.8, using conventional antibodies) cognate protein in both its phosphorylated and unphosphorylated states, as well as with any proteins toward which there could conceivably be cross-reactivity. The blot is probed with the monoclonal antibody preparation to demonstrate appropriate reactivity. The antibody preparation can ultimately be purified on a cognate phosphopeptide affinity column (see Support Protocol 1). This protocol presents procedures for isolating hybridoma clones of the desired reactivity, starting from the point at which candidate hybridoma clones have been seeded in 96-well culture plates. Candidate hybridoma supernatants are screened in ELISA assays against a series of peptides or lysates of the desired proteins (as described in Strategic Planning). Identified clones are expanded, subcloned by limiting dilution, and further purified. Materials Candidate hybridoma cell lines from fusion HT medium (see recipe) Screening diluent (see recipe) BSA-conjugated cognate phosphopeptide (for use as ELISA antigen; see Support Protocol 1 and UNIT 18.3) Pre-immune serum from mouse used to produce hybridoma line (negative control) Immune serum from mouse used to produce hybridoma line (positive control) BSA-conjugated cognate nonphosphopeptide (for use as ELISA antigen; UNIT 18.3) BSA-conjugated non-cognate phosphotyrosyl peptide (for use as ELISA antigen; UNIT 18.3) BSA-conjugated homologous phosphopeptides (optional, for use as ELISA antigens; UNIT 18.3) 96-well polystyrene tissue culture plates Grid note sheets Additional reagents and equipment for cloning by limiting dilution, ELISA, and freezing and recovery of hybridoma cell lines NOTE: All solutions and equipment coming into contact with cells must be sterile, and proper sterile technique should be used accordingly.
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Identify and sample conditioned media from candidate hybridomas 1. Seed hybridoma cells from fusion at a low density (aiming for one cell per three wells) in multiple 96-well polystyrene tissue culture plates using HT medium. Examine at ∼2 weeks and identify all single-colony hybridoma culture wells as potential candidates by recording the number of colonies per well on a 96-grid note sheet. Mark the underside of each well containing one colony for easy identification. The culture medium from wells possessing a single colony is screened for reactivity.
2. Using sterile technique, remove an aliquot of supernatant (e.g., 100 µl from a 200-µl culture) from each potential candidate well and transfer to a separate 96-well polystyrene plate (the “screen plate”). Record the original plate number and well position and the corresponding plate number and well position for the screen plate (e.g., on a grid sheet representing the screen plate, record the original plate number and well position from which the supernatant in each well was transferred). Replace the aliquot removed from each original well with 100 µl of 37°C fresh HT medium. A brightly colored dot sticker the size of one well, attached to the tissue culture hood surface, will help to keep the plate in register, as the plate may be moved one well at a time along the sticker. If the supernatants are not to be tested immediately, store the screen plates at 4°C, wrapped in plastic wrap.
3. To each supernatant in the screen plates, add 150 µl screening diluent. The components of the screening diluent prevent microbial contamination; this reagent also serves to expand the volume should multiple rounds of screening be required.
4. To screen candidate hybridomas by ELISA, analyze an aliquot (e.g., 50 µl) of each candidate hybridoma supernatant from the screen plate in an ELISA using the cognate phosphopeptide-BSA conjugate as antigen. Record the screen-plate number and well position and corresponding original culture plate number and well position for each positive sample. Also assay pre-immune serum from the mouse used for fusion (negative control) and immune serum from the same mouse (positive control), each at a 1:100 dilution in a 1:1 mixture of HT medium and screening diluent. Include an ELISA with screening diluent only (additional negative control). It is important to use a different carrier protein in the ELISAs (BSA) than that used for the immunizations (KLH), to avoid detecting reactivity against the carrier (see Strategic Planning).
5. Screen an aliquot of each positive sample from step 4 in an ELISA against the following: a. cognate nonphosphopeptide-BSA conjugate (to eliminate clones with phosphorylation-independent reactivity); b. unrelated phosphotyrosylpeptide-BSA conjugate (to eliminate clones with indiscriminate phosphotyrosine reactivity); c. phosphopeptide-BSA conjugates corresponding to any potentially cross-reacting homologous phosphoproteins (if cross-reactivity with homologous phosphoprotein(s) is anticipated). These screens may be done sequentially or, if the number of candidate hybridomas has diminished sufficiently, simultaneously. Isolation of clones with phosphorylation-independent cognate peptide specificity may also be useful.
6. Expand clones that satisfy all screening requirements and freeze aliquots as backup; use remainder for subcloning by limiting dilution. Sequence-Specific Anti-Phosphoamino Acid Antibodies
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7. Screen culture supernatants from subclones as in steps 4 and 5 to check for continued production and specificity of the antibody. Keep representative independent subclones from each original parental colony, as it cannot be predicted whether each parental monoclonal antibody will recognize the native phosphoprotein or function in all types of immunoassays desired in the future. Candidate subclones should then also be expanded and frozen. It is a good idea to continue to check for production of the appropriate antibody after several rounds of serial passaging and after freezing, thawing, and re-expansion.
8. To characterize the subclones further and identify the most useful clones, assay the culture supernatants for reactivity with cognate holophosphoprotein in individual immunoassays (e.g., immunoprecipitations as described in UNIT 8.3, or immunoblot analyses as described in UNIT 10.10). Antibody from the desired clones can be produced on a large scale by harvesting large volumes of culture supernatant and performing affinity purification (see Support Protocol 1), or by producing ascites fluid, which can be purified by ammonium sulfate precipitation followed by affinity purification. Isotype analysis can be performed as described by Hornbeck et al. (1991) or by using commercially available kits (Pierce or Sigma).
COUPLING PEPTIDES TO AFFI-GEL 10 MATRIX This protocol describes coupling of phosphopeptides or non-phosphopeptides to Affi-Gel 10 affinity matrix for use in affinity column chromatographic purification of antibodies. Anhydrous coupling, detailed here, is the most efficient coupling method for peptides. The steps here result in production of 3 ml final bed volume of affinity resin; 3 µmol of peptide is coupled per milliliter Affi-Gel 10. According to the manufacturer’s specifications, Affi-Gel 10 resin contains ∼15 µmol of active ester per milliliter of gel, and the gel has a capacity for 35 mg of protein or 15 to 20 µmol of a low-molecular-weight ligand per milliliter of gel. Note that these capacities are dependent upon the molecular weight and nature of the protein. After coupling, the matrix may be poured into columns of the appropriate size.
SUPPORT PROTOCOL 1
Materials Synthetic oligopeptide for coupling (see Strategic Planning) Dimethyl sulfoxide (DMSO) N-methylmorpholine (99% purity; Acros Organics) Affi-Gel 10 (Bio-Rad) or equivalent activated support matrix Ethanolamine 0.1 M ethanolamine⋅Cl, pH 8.0 High-salt/high-pH solution: 0.5 M NaCl/0.4% (w/v) sodium bicarbonate High-salt/low-pH solution: 0.5 M NaCl/100 mM sodium acetate, pH 4.2 PBS/azide: PBS (APPENDIX 2E) containing 0.02% (w/v) sodium azide 0.5 M NaCl 3 M NaSCN Polypropylene screw-cap centrifuge tubes (do not use polystyrene tubes with DMSO) End-over-end rotator IEC Clinical centrifuge (or equivalent) Vacuum aspirator Glass chromatography columns, ≥5-ml capacity (Bio-Rad) Post-Translational Modifications: Phosphorylation and Phosphates
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Prepare peptide solution 1. Dissolve synthetic oligopeptide in DMSO at a volume ∼0.5 to 4 times the desired bed volume. For a 3-ml bed volume, dissolve a total of 9 ìmol peptide, which for an average 15-mer peptide of mol. wt. ∼1700 would be 15.3 mg.
2. Neutralize the peptide solution by titration as follows: add peptide synthesis-grade N-methylmorpholine (full-strength or diluted 1:1 to 1:9 in DMSO) in 1- to 2-µl increments, remove a 2- to 3-µl aliquot after each addition, dilute this with 50 µl water, then spot it onto a strip of pH paper. Repeat this process until a pH in the range of 7 to 8 is reached. IMPORTANT NOTE: Titrate carefully, as it is very easy to overshoot the desired pH. Typically, 7 to 8 ìl of N-methylmorpholine is required per 3 ìmol peptide, although the precise amount will depend greatly upon the particular amino acid sequence. The aliquots must be mixed with water because pH is a measure of hydrogen ion concentration, and hence the pH of a nonaqueous solution cannot be measured.
Add peptide to Affi-Gel 10 3. Mix contents of the bottle of Affi-Gel 10 matrix well to resuspend the resin, then transfer the required amount of resin to a screw-cap polypropylene centrifuge tube and wash three times in DMSO, each time by centrifuging 5 min at 700 × g (2000 rpm in an IEC clinical centrifuge, setting 5), room temperature, aspirating the supernatant, resuspending the resin in 5 vol DMSO, then centrifuging again at 700 × g. IMPORTANT NOTE: Be sure to start with at least twice as much Affi-Gel 10 as ultimately will be needed, because the volume will shrink. The resin is supplied as a 50% slurry; therefore, it is necessary to use at least four times the required bed volume. Avoid centrifugation at higher-than-recommended speed as this may result in collapse of the resin.
4. Aspirate the excess DMSO from the resin and add the peptide solution prepared in step 2. Incubate overnight at room temperature with end-over-end rotation. Quench and wash the resin 5. Add 2 µl pure ethanolamine per milliliter resin and incubate 2 hr at room temperature with end-over-end rotation. Ethanolamine is added directly to the coupling reaction to quench unreacted ester groups.
6. Wash two times in DMSO as in step 3. 7. Wash two times in 0.1 M ethanolamine⋅Cl, pH 8.0, as in step 3 (performing the first wash on ice as a great deal of heat will be generated). Remove the 0.1 M ethanolamine⋅Cl from the second wash and replace with fresh, incubate tube overnight at 4°C with end-over-end rotation, then centrifuge at low speed (1000 × g) at 4°C and aspirate the supernatant. 8. Wash three times with high-salt/high-pH solution using the technique described in step 3. 9. Wash three times with high-salt/low-pH solution using the technique described in step 3. 10. Wash three times with PBS/azide using the technique described in step 3 and store washed resin at 4°C until ready to pack in column. Sequence-Specific Anti-Phosphoamino Acid Antibodies
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11. Mix matrix into a slurry with 0.5 M NaCl and pour into glass chromatography columns. Commercially available glass chromatography columns can be used; alternatively, 5-ml plastic syringes with 1-cc plugs of silanized glass wool in the bottom may function as columns.
12. Wash each column with 10 column volumes 3 M NaSCN followed by 10 column volumes PBS/azide. COUPLING PHOSPHOTYROSINE TO AN AFFI-GEL MATRIX This protocol describes coupling of phosphotyrosine to Affi-Gel 10 affinity matrix for use in affinity column chromatographic purification of antibodies. Phosphotyrosine is insoluble under the anhydrous conditions used for coupling peptides (see Support Protocol 1); thus coupling is done under aqueous conditions. The steps here result in production of 10 ml final bed volume of affinity resin; as for peptides, 3 µmol phosphotyrosine is coupled per milliliter of Affi-Gel 10. According to the manufacturer’s specifications, Affi-Gel 10 resin contains ∼15 µmol of active ester per ml of gel, and the gel has a capacity for 35 mg of protein per ml gel or 15 to 20 µmol of a low-molecular-weight ligand per ml gel. Note that these capacities may vary depending upon the molecular weight and nature of the protein. Phosphotyrosine-agarose affinity matrix is also available commercially from Sigma.
SUPPORT PROTOCOL 2
Materials Phosphotyrosine 0.4% (w/v) sodium bicarbonate 1 M NaOH (optional) Fritted-glass funnel and vacuum aspirator Glass chromatography column, ≥14-ml capacity (Bio-Rad) Additional reagents and equipment for coupling peptides to Affi-Gel 10 affinity matrix (see Support Protocol 1) Prepare phosphotyrosine solution 1. Dissolve phosphotyrosine in 0.4% sodium bicarbonate at a volume of about one-half to four times the desired bed volume. For a 10-ml bed volume, dissolve a total of 30 ìmol phosphotyrosine. Based on its molecular weight of 261, this amounts to 7.8 mg, although it is acceptable (and may be easier) to use an excess, as phosphotyrosine is relatively inexpensive.
2. Check that the pH is in the range of 7 to 8 using pH paper. If adjustment is necessary, neutralize the solution by titration—i.e., adding small amounts of 1 M NaOH, removing a 2- to 3-µl aliquot after each addition, then spotting the aliquots onto a strip of pH paper. Repeat this process until a pH in the range of 7 to 8 is reached. Add phosphotyrosine to Affi-Gel 10 3. Mix contents of the bottle of Affi-Gel 10 matrix well to resuspend resin, then transfer the required amount of resin to a fritted-glass funnel attached to a vacuum aspirator. Wash three times, each time by pouring cold distilled water over the resin in the funnel and aspirating, then perform one final wash in the same manner using ice-cold 0.4% sodium bicarbonate. Post-Translational Modifications: Phosphorylation and Phosphates
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IMPORTANT NOTE: Be sure to take at least twice as much Affi-Gel 10 as ultimately will be needed, as the volume will shrink. The resin is supplied as a 50% slurry; therefore, it is necessary to take at least four times the required bed volume. Do not allow >20 min to elapse from the time the resin is removed from the bottle to the time it is mixed with the phosphotyrosine solution.
4. Remove most of the excess liquid from the gel by vacuum aspiration without letting it dry completely, transfer it to a screw-cap centrifuge tube, and add the phosphotyrosine solution from step 2. 5. Incubate overnight at 4°C with end-over-end rotation. Quench and wash resin 6. Add 2 µl pure ethanolamine per milliliter resin and incubate 2 hr at room temperature with end-over-end rotation. Ethanolamine is added directly to the coupling reaction to quench unreacted ester groups.
7. Wash resin two times, each time by centrifuging at low speed (see Support Protocol 1, step 3), aspirating the supernatant, resuspending the resin in 5 vol of 0.4% sodium bicarbonate, then centrifuging again at low speed. 8. Wash resin two times with 0.1 M ethanolamine⋅Cl, pH 8.0, as in step 7. Remove the 0.1 M ethanolamine⋅Cl from the second wash and replace with fresh 0.1 M ethanolamine⋅Cl, incubate tube overnight at 4°C with end-over-end rotation, then centrifuge at low speed and aspirate the supernatant. 9. Wash, store, and pack resin in columns (see Support Protocol 1, steps 8 to 12). REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
HT medium To complete DMEM-20 medium (APPENDIX 3C) containing 10 mM HEPES and 1 mM pyruvate, add 100× HAT (hypoxanthine/aminopterin/thymidine; Sigma) or 100× HT (Sigma) supplement to 1× final. Store up to 1 month at 4°C. Screening diluent 6.18 g boric acid (100 mM final) 9.54 g sodium borate (47 mM final) 4.38 g NaCl (75 mM final) 10 g bovine serum albumin (BSA; 1% w/v final) 1 g sodium azide (1% w/v final) H2O to 1 liter The pH of the solution will be 8.4 to 8.5. The screening diluent contains sodium azide to allow storage up to 1 year at 4°C.
COMMENTARY Background Information
Sequence-Specific Anti-Phosphoamino Acid Antibodies
Conventional antibodies have proven to be invaluable tools in numerous techniques for the biochemical analysis of proteins. In the past, antibodies with specificity toward the phosphorylated forms of proteins were produced serendipitously. Using techniques such as those
described in this unit, such antibodies may now be produced by design. Whereas phosphorylated holoproteins might in general be poor candidates for immunogens in the production of such antibodies as a result of their susceptibility to dephosphorylation, it has been suggested that short phosphopeptides are relatively
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resistant to phosphatases (Czernik et al., 1991), thereby providing a more stable antigen. A dramatic advance in the analysis of protein tyrosine phosphorylation and in understanding the regulation of signal transduction pathways by such phosphorylation occurred with the production of polyclonal and monoclonal anti-phosphotyrosine antibodies (Ross et al., 1981; Frackelton et al., 1983; reviewed in Stern, 1991). These antibodies proved capable of recognizing phosphorylated tyrosine residues in the context of virtually any flanking peptide sequence, and were shown to be exquisitely specific in their requirement for a phosphorylated tyrosine, yet at the same time remarkably promiscuous in their acceptance of any peptide sequence. Techniques using polyclonal and monoclonal anti-phosphotyrosine antibodies have supplanted the standard methodology of metabolically labeling proteins with 32Pi, which was cumbersome, time-consuming, hazardous, and of lower sensitivity. Antibodies that recognize a specific tyrosylphosphorylated peptide, as described in this unit, represent a marriage of anti-phosphotyrosine technology with the stringent sequencedependent specificity of conventional antibodies (Roussel, 1990; Bangalore et al., 1992; Epstein et al., 1992; DiGiovanna and Stern, 1995). Conceptually, they may be thought of either as anti-phosphotyrosine antibodies that have strict sequence specificity, or as conventional antibodies possessing the additional specificity of phosphorylation dependence. Anti-phosphotyrosine antibodies have been most useful in the analysis of tyrosine phosphorylation of proteins using a technique that combines immunoprecipitation and immunoblotting (known as the “IP-western” method). In this procedure, a protein typically is immunoprecipitated either with conventional anti-protein antibody or with anti-phosphotyrosine antibody, then immunoblotted with whichever of these two antibodies was not used for the immunoprecipitation. Because anti-phosphopeptide antibodies recognize the cognate protein, but only when it is phosphorylated, they are capable of performing both functions in a single step—i.e., either an immunoprecipitation alone or an immunoblot alone is sufficient to supply the desired information. Using anti-phosphopeptide antibodies, it is possible to identify and isolate distinct phosphorylated species of a phosphoprotein containing multiple phosphorylation sites. Thus, phosphorylation at each site can be examined independently, and a preparative separation of
individual phosphospecies of a holoprotein can be achieved. Another unique application is analysis of the abundance and phosphorylation state of individual proteins in situ in preparations of cells or tissues. Because phosphorylation is a major mode of regulation of protein function, the phosphorylation state is often an indicator of the functional status of a protein. The mere identification of a protein in a cell or tissue specimen gives no indication of its functional status. The ability of anti-phosphopeptide antibodies to demonstrate the phosphorylation state, and by extrapolation the functional state, of a single protein with high specificity places these reagents in a unique class. Hence, identity and functional status may be probed simultaneously using one simple assay. In tissue sections, one may probe to determine whether a particular protein is present, and, if present, in what functional state it is found. Immunohistochemical staining of formalinfixed, paraffin-embedded human breast tumors using antibody to the phosphorylated form of the receptor tyrosine kinase Neu (DiGiovanna and Stern, 1995) has demonstrated that Neu is phosphorylated, and therefore functionally active, in only a subset of the tumors that overexpress this protein. Recently, a novel approach of using sequence-specific anti-phosphoamino acid antibodies has emerged (Zhang et al., 2002). In this approach, phosphopeptide libraries with fixed residues corresponding to a consensus motif (e.g., Akt phosphorylation motif, XXXRXRXXpTXXX) were used as antigens to generate polyclonal antisera in rabbits. These antibodies can recognize many different phosphoproteins containing the consensus phosphorylated motif for a given protein kinase (e.g., Akt). These antibodies provide a new approach to rapidly indentify protein kinase substrates. From a technical standpoint, production of antibodies with specificity for the nonphosphorylated state is achievable with similar methods to those described in this unit. Numerous examples have been reported (Nairn et al., 1982; Roussel, 1990; Czernik et al., 1991; Epstein, 1995; Tzartos et al., 1995; Kawakatsu et al., 1996).
Critical Parameters and Troubleshooting In the production of polyclonal anti-phosphopeptide antibodies, the most common unfavorable outcomes are persistence of cross-reactivity, loss of specific reactivity, and poor yield. Although it may be tempting, in the
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Sequence-Specific Anti-Phosphoamino Acid Antibodies
interest of saving time, to perform the entire purification and postpone all analyses until the end (a reasonable course of action once a scheme is known to work), it is recommended, especially with a new protocol, to analyze the serum at each step. If loss of specific reactivity is also a problem, stepwise analyses can also demonstrate where the loss is occurring. Loss of specific reactivity can occur for three major reasons. The first possible reason is that the antibody is not stable over the course of the purification, which may be the case if the procedure is very prolonged. If this is suspected, the pace of the purification may be hastened or the procedure may be carried out at 4°C instead of room temperature. With prolonged purifications, it is especially important to include sodium azide in the PBS used to dilute the serum, to prevent microbial growth. Activity may also be lost nonspecifically as a result of the multiple manipulations in a protocol requiring multiple columns and/or multiple passes through each column. In this case, fraction analysis may indicate which columns are absolutely essential, and also indicate the minimum necessary number of passes through each column. Also, creating larger columns with greater capacity should reduce the number of times the serum will need to be passed over each to deplete cross-reactivity. Finally, specific activity may be lost as a result of the presence in one of the negative-selection columns of a cross-reacting peptide that is so similar to the peptide of interest that essentially any antibody that recognizes the cognate phosphopeptide will recognize this phosphopeptide as well. In this last theoretical scenario, ultimate unique specificity may not be achievable. The columns used in the purifications can be regenerated and used repeatedly if stored at 4°C in sodium azide–containing buffer to prevent microbial contamination and consequent damage to the matrix. A theoretical consideration is that with repeated use, phosphatases in serum may eventually cleave enough phosphate groups from the peptide moieties on the column to render it ineffective as a phosphopeptide column. Thus, it may be prudent to periodically prepare fresh columns. An alternative would be to add phosphatase inhibitors to the serum prior to purification. Similarly, proteases in serum may eventually cleave peptides from the resin, and this may be prevented by adding protease inhibitors to the serum. The main technical hurdle in the production of monoclonal antibodies appears to be the large number of clones that it may be necessary
to screen to find one with the desired reactivity. Here, time and persistence are the primary defenses. Another obstacle may be the failure of hybridoma cells to continue to produce antibody upon subcloning, which is likely due to the continuing genetic instability of hybridomas at early passage. If the parental hybridoma colony has been expanded and frozen prior to subcloning, this will serve as a potential source for additional subclones, although it is possible that these additional daughter clones may also be nonproducing. The use of the final monoclonal antibody in a variety of immunoassays is more likely to require potentially extensive optimization of protocols and higher concentrations (because of lower affinities) than the use of polyclonal antibodies.
Anticipated Results In the production of polyclonal antibodies, yields of ∼1 mg of purified anti-phosphopeptide antibody can be obtained from a 15-ml serum sample. These antibodies have been used in the immunoprecipitation of phosphoproteins from both denaturing and nondenaturing solutions. They have also been used in immunoblotting, in immunofluorescence on fixed cultured cells, and in immunohistochemistry on formalin-fixed, paraffin-embedded tissue sections. Specificity can be demonstrated when such assays are inhibited by cognate phosphopeptide but not by cognate nonphosphopeptide or unrelated phosphopeptides. Through the production of monoclonal antibodies a permanent supply of a uniform reagent is ensured. Many milligrams can be obtained from the ascites produced by a single mouse. Such an antibody can be used in all of the same assays as described above for polyclonal antibodies. More extensive optimization of protocols is likely to be required with monoclonal antibodies, and these may be expected to have a lower affinity than typical polyclonal antibodies. Isotype analysis can easily be performed as described by Hornbeck et al. (1991) or by use of commercially available kits (Pierce or Sigma). In the production of monoclonal anti-phosphotyrosyl peptide antibodies, a major technical hurdle is the low frequency with which clones of the desired specificity may be produced. In one case (DiGiovanna and Stern, 1995), antibody was produced against the phosphorylated form of the receptor tyrosine kinase Neu, >1200 hybridomas (obtained from a single fusion) were screened and a single clone was identified that satisfied all requirements.
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Table 13.6.1 Antisera
Peptide Antigen Sequences and Anti-Phosphopeptide Titers of Anti-Bcr
Peptide antigen sequences Phosphoserine 354 peptide J27 Serine 354 peptide I81 Phosphotyrosine 360 peptide J138 Phosphotyrosine 360 peptide J119 Tyrosine 360 peptide J118
GQSSRVS*PSPTTYR(C) GQSSRVSPSPTTYR SPSPTTY*RMFRDKC (antigen) SPSPTTY*RMFRDK (blocking peptide) SPSPTTYRMFRDKC
Titers of the antisera anti-phosphoserine 354 (J27)a Rabbit no. 5402 5403
Titer 1/10,400 1/70,900 Titers of the antisera anti-phosphotyrosine 360 (J138)a
Rabbit no. 5792 5793
Titer 1/18,800 1/78,300
a Two rabbits were injected with each antigen. Titers are shown as dilutions of the antiserum that produced an O.D.
of 1.0 in an ELISA assay in a peptide-coated microplate.
A
3
1
2
Anti-PSer354 Ab 5402
Anti-Bcr Ab G156 (Bcr 298-310)
A
B
A
C
B
Anti-PSer354 Ab 5403
A
C
B
C
Bcr 1-413
B
1
2
3
4
Anti-PSer354 Ab 5402 +Phospho Peptide J27
Anti-PSer354 Ab 5402 +Unphospho Peptide I81
Anti-PSer354 Ab 5403 +Phospho Peptide J27
Anti-PSer354 Ab 5403 +Unphospho Peptide I81
A
B
C
A
B
C
A
B
C
A
B
C
Bcr 1-413
Figure 13.6.3 Characteristics of anti-pSer-354 antisera. Immunoblot analyses were performed on extracts from COS1 cells overexpressing Bcr(1-413) wild-type or S354A mutant proteins. Lane A is COS1 cells without transfection. Lane B is COS1 cells transfected with wild-type BCR(1-413). Lane C is COS1 cells transfected with S354A mutant BCR(1-413). Lane M represents molecular weight markers. (A) Immunoblotting using anti-pSer-354 antisera 5402 (panel 1), anti-Bcr(298-310) Ab G156 (panel 2) and anti-pSer-354 antisera 5403 (panel 3). (B) Immunoblotting using antisera 5402 pre-incubated with pSer-354 peptide J27 (panel 1) or Ser-354 peptide I81 (panel 2), or using antisera 5403 pre-incubated with pSer-354 peptide J27 (panel 3) or Ser-354 peptide I81 (panel 4).
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forms of the Bcr-Abl protein are major factors in the induction and maintenance of these types of leukemia, as determined by mouse studies (Daley et al., 1990; Heisterkamp et al., 1990), and both forms exhibit elevated tyrosine kinase activity as compared to normal c-Abl Tyr kinase (Ponticelli et al., 1982; Konopka et al., 1984; Kloetzer et al., 1985). The elevated Tyr kinase activity is crucial for the oncogenic function of Bcr-Abl oncoprotein. Previous studies identified several phosphorylation sites on Bcr, including Tyr-177, 283, 328, and 360 (Liu et al., 1993; Puil et al., 1994; Liu et al., 1996a; Wu et al., 1998). The following section describes the development and characterization of sequence-specific anti-phosphopeptide antibodies, specific for phosphorylated residues Tyr-360 and Ser-354, to examine the in vivo phosphorylation status these residues in Bcr and Bcr-Abl.
Example of generation and characterization of sequence-specific anti-phosphoamino acid antibodies (Sun et al., 2001) The breakpoint cluster region (BCR) gene is expressed in all human tissues (Campell and Arlinghaus, 1991). It encodes a novel protein kinase, p160 BCR, within its first exon that phosphorylates itself and target proteins on Ser/Thr residues. The Philadelphia (Ph) chromosome is found in >95% of patients with chronic myelogenous leukemia (CML) and ∼15% to 20% of patients with adult acute lymphoblastic leukemia (ALL). This abnormal chromosome contains 5′ BCR gene sequences from chromosome 22 fused to the second exon of the ABL gene from chromosome 9 (Heisterkamp et al., 1985; Shtivelman et al., 1985; Stam et al., 1985). Two major forms of BCRABL gene products have been described. Depending on the location of the breakpoint within BCR, either a 210- (P210 BCR-ABL) or 185-kDa (P185 BCR-ABL) protein (also some times identified as P190) is produced. P210 Bcr-Abl is found predominantly in CML and some Ph-positive ALL patients. P185 BcrAbl is found in Ph-positive ALL patients. Both
Anti-PSer354 Ab5402 +S354 Phospho Peptide J49
A
B
C
Expression of wild-type and mutant BCR species BCR, P185 Bcr-Abl wild type and mutant constructs, and P210 Bcr-Abl wild type and mutant constructs were subcloned into the pSG5 vector according to established methods.
Anti-PSer354 Ab5402 +S356 Phospho Peptide J69
A
B
C
Anti-PSer354 Ab5402 +S354 S356 double Phospho Peptide J70
A
B
C
Anti-PSer354 Ab5402 +Unphospho Peptide G95
A
B
C
Bcr 1-413
Sequence-Specific Anti-Phosphoamino Acid Antibodies
Phosphoserine 354
J49
SSRVS*PSPTTRAIRS
Phosphoserine 356
J69
SSRVSPS*PTTRAIRS
Phosphoserine 354/356
J70
SSRVS*PS*PTTRAIRS
Unphosphorylated
G95
SSGQSSRVSPSP
Figure 13.6.4 pSer-356 peptide inhibits the cross-reactivity with S354A mutant, but not the reactivity with wild-type Bcr(1-413). Membranes from Figure 13.6.3 were stripped, and immunoblotted using antiserum 5402 mixed with different peptides. Panel 1, pSer-354 peptide J49; panel 2, pSer-356 peptide J69; panel 3, double pSer-354/356 peptide J70; and panel 4, non-phosphopeptide G95 containing Ser-354 and 356. The sequences of the peptides are shown below the figure. The phosphoresidues are marked by *. Lane A is COS1 cell without transfection. Lane B is COS1 cells transfected with wild-type BCR(1-413). Lane C is COS1 cells transfected with S354A mutant BCR(1-413). Lane M represents molecular weight markers.
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1
2
3
4
A
5
6
7
8 Bcr-Abl p210
anti-phosphotyr 360 Ab 5792 + nonphosphopeptide I118
B anti-phosphotyr 360 Ab5792 + phosphopeptide I119
Bcr-Abl p210
C
Bcr-p160
anti-Bcr (1-16) Ab G143
Figure 13.6.5 Characterization of the sequence-specific Bcr pTyr-360 antiserum. COS1 extracts expressing various Bcr-Abl and Bcr proteins were analyzed by immunoblotting with anti-pTyr-360 antiserum (5792, see Table 13.6.1) mixed with excess non-phosphopeptide I118 (see Table 13.6.1) (panel A); anti-pTyr-360 blocked with excess pTyr-360 peptide (I119) (panel B); anti-Bcr(1-16) (G143) as a loading control (panel C). The lanes are: vector transfected COS1 cells (lane 1); P210 Bcr-Abl transfected COS1 cells (lane 2); Y328F mutant P210 Bcr-Abl (lane 3); Y360F mutant P210 Bcr-Abl (lane 4); Y328F/Y360F double mutant P210 Bcr-Abl (lane 5); S354A mutant P210 Bcr-Abl (lane 6); P160 BCR (lane 7); and Y328F/Y360F double mutant P160 BCR (lane 8).
BCR, BCR-ABL cDNAs, and P185 Bcr-Abl Y360F constructs were from Dr. John Groffen (Children’s Hospital Los Angeles Research Institute). Other mutant BCR constructs and BCR-ABL constructs [BCR Y328F, BCR Y360F, BCR(1-413), BCR(1-413) S354A, BCR-ABL Y328F, BCR-ABL Y360F, BCRABL Y328F/Y360F, BCR-ABL S354A] were made by Dr. Yun Wu from the Arlinghaus Laboratory (Liu et al., 1996a; Wu et al., 1998; Wu et al., 1999). COS-1 cells were transiently transfected with BCR-ABL or BCR constructs in the pSG5 vector with LipofectAmine (Life Technologies) according to the manufacturer’s protocol. Immunoblotting and immunoprecipitation Hematopoietic cells or COS-1 cells expressing Bcr or Bcr-Abl proteins were lysed in (SDS)/2-mercaptoethanol sample buffer and boiled for 5 min. The lysates were clarified by centrifugation and the proteins in the super-
natant were resolved by SDS-PAGE. The gelembedded proteins were electroblotted onto Immobilon P membranes and probed. Immunoblotting was performed using the ECL method with commercial reagents (e.g., Amersham Biosciences). Mouse monoclonal antibodies, including 8E9 (anti-Abl SH2 domain; Guo et al., 1991) and 4G10 (anti-pTyr, from Upstate Biotech) were used at 1:5000 dilution. Rabbit antibodies, including G143 [anti-Bcr(116)], G156 [anti-Bcr(298-310)], G157 [antiBcr(181-194)], anti-pTyr-360, and anti-pSer354 were used at either 1:1000 or 1:2000 dilution. For peptide blocking experiments, excess non-phosphorylated peptides or phosphopeptides was added to the antiserum, incubated 10 min at room temperature and then used for immunoblotting. For immunoprecipitation, cells were lysed, homogenized, and centrifuged. The supernatant was incubated with anti-Abl(51-64) monoclonal antibody (P6D; Guo et al., 1991)
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followed by addition of protein A–Sepharose beads. After several washings, the beads were collected, boiled in SDS/2-mercaptoethanol sample buffer, and analyzed by SDS-PAGE followed by immunoblotting as described above. Production of anti-Bcr antibodies Anti-phosphopeptide antibodies, initially focusing on Tyr 360 and Ser-354 of Bcr, were raised to examine the in vivo phosphorylation status of these residues in Bcr and Bcr-Abl. Two rabbits were injected with each antigen. The peptide antigen sequences and the anti-phosphopeptide titers of the antisera are shown in Table 13.6.1.
Sequence-Specific Anti-Phosphoamino Acid Antibodies
Characterization of anti-Bcr pSer-354 antisera Anti-pSer-354 antisera were immunoblotted with both wild-type and an S354A mutant of a truncated form of Bcr, Bcr(1-413), which represents the amino terminal 413 residues of the Bcr protein. Bcr(1-413) functions as a Ser/Thr kinase with activity that is similar to, if not higher than, that of the full length Bcr protein (Liu and Arlinghaus, unpub. observ.) It has been shown that this truncated protein is phosphorylated on Ser-354 in kinase assays (Hawk et al., 2002). Immunoblotting with rabbit anti-Bcr peptide 298-310 antibody (G156), which does not detect the pSer epitope, was used as a loading control (Fig. 13.6.3A, panel 2). With similar amounts of wild-type and mutant Bcr(1-413) proteins, the high titer rabbit 5403 antiserum (Table 13.6.1) detected the wild-type protein (Fig. 13.6.3A, panel 3, lane B), but not the S354A mutant protein (Fig. 13.6.3A, panel 3, lane C). However, the low titer rabbit serum 5402 (Table 13.6.1) recognized both wild-type (Fig. 13.6.3A, panel 1, lane B) and mutant proteins (Fig. 13.6.3A, panel 1, lane C), although it had very low reactivity towards the S354A mutant. To determine whether the cross-reactivity was due to the antibodies raised against the non-phosphorylated peptide sequence and to confirm the specificity of the antisera, immunoblotting was performed with antisera preincubated with various forms of the antigenic peptides (Fig. 13.6.3B). As expected, the preincubation with pSer-354 peptide J27 (Table 13.6.1) greatly reduced the reactivity towards the Bcr(1-413) (Fig. 13.6.3B, panels 1 and 3), which indicated that the reactive antibodies in the serum recognized the pSer-354 form of the Bcr protein. Phosphopeptides blocked the an-
tisera obtained from both rabbits. Surprisingly, the non-phosphorylated peptide I81 did not block the cross-reactivity of antiserum 5402 with S354A mutant (Fig. 13.6.3B, panel 2, lane C) compared to reactivity to the serum in the absence of blocking peptides (Fig. 13.6.3A, panel 1). These data suggest that this reactivity in the 5402 rabbit is directed to an epitope quite similar to the pSer-containing Bcr sequence. In support of this interpretation, the pSer-354 peptide J27 blocked this cross-reactivity (Fig. 13.6.3B, panel 1, lane C), confirming that the low-titer serum (5402) detects another pSer epitope in Bcr(1-413) resembling pSer-354. Nevertheless, the results with high-titer antiserum 5403 confirmed that it contains an antibody that is specific for the pSer-354 Bcr sequence (compare Fig. 13.6.3A, panel 3 to Fig. 13.6.3B, panel 4). Bcr Ser-356 is a potential phosphorylation site in vivo The results of Figure 13.6.3B, panels 3 and 4, show that the high-titer antiserum 5403 was reactive with wild-type Bcr(1-413), but not with the S354A mutant, and this reactivity was greatly reduced by incubating the antiserum with pSer-354 peptide J27. These results indicated that Ser-354 is phosphorylated in vivo in the Bcr(1-413) protein. The fact that the crossreactivity of the lower titer anti-pSer-354 antiserum 5402 with S354A mutant Bcr(1-413) could not be reduced or partially reduced by non-phosphorylated peptide and could be blocked totally by phosphopeptide suggested the cross-reactive epitope detected by this rabbit serum is very likely to be similar to the pSer-354 sequence. Ser-356, which is in the same peptide context as Ser-354 and shares the same repeat sequence of 354SP356SP (the leftsuperscripted number denotes the position of the amino acid in Bcr protein) is a candidate for the alternative epitope. Immunoblotting analyses using various peptides as blocking reagents were performed with anti-pSer-354 antiserum 5402 (Fig. 13.6.4). The sequences of the blocking peptides are shown below the figure. The non-phosphorylated peptide G95 did not reduce the cross-reactivity with S354A mutant Bcr(1-413) (Fig. 13.6.4, panel 4). The Ser354/356 double phosphorylated peptide J70 also did not reduce the cross-reactivity (Fig. 13.6.4, panel 3). A second and different pSer354 peptide J49 blocked reactivities with both wild type and mutant proteins (Fig. 13.6.4, panel 1) similar to that blocked by phosphopeptide J27 (Fig. 13.6.3B, panel 1). This further
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confirmed that the antiserum contains the specific affinity towards pSer-354. Interestingly, pSer-356 peptide J69 inhibited the cross-reactivity with S354A mutant, but did not block the reactivity with wild-type protein (Fig. 13.6.4, panel 2). This result suggests that Ser-356 is phosphorylated in Bcr(1-413) in vivo and that lower-titer antiserum 5402 recognizes two epitopes, the high-affinity pSer-354 site and the low-affinity pSer-356 site. Experiments with an S356A Bcr mutant and an S354/356A double Bcr mutant will be required to confirm this conclusion. Characterization of anti-pTyr-360 antiserum Previous studies established that Tyr-360 of Bcr is a site of phosphorylation in P210 Bcr-Abl and P160 BCR (Liu et al., 1996a). Therefore, it was of interest to confirm these studies by generating a pTyr sequence–specific antibody involving Tyr-360. Figure 13.6.5 shows the results of immunoblots with antiserum obtained with rabbit 5792 injected with a pTyr360 peptide cross-linked to KLH. As expected the antiserum pre-incubated with the non-phosphorylated peptide detected wild type Bcr-Abl and Bcr-Abl Y328F (Fig. 13.6.5A, lanes 2 and 3), but not the Y360F Bcr-Abl mutant and the Y328/360F double mutant (Fig. 13.6.5A, lanes 4 and 5), establishing that the antiserum is specific for the pTyr-360 form of Bcr-Abl. Blocking the antiserum with excess pTyr-360 peptide removed reactivity with the wild type Bcr-Abl protein, which is known to be phosphorylated on Tyr-360 (Fig. 13.6.5B; Liu et al., 1996a). Of interest, the antiserum to pTyr-360 preincubated with the non-phosphopeptide did not detect the wild-type Bcr protein (Fig. 13.6.5A, lane 7), which is not phosphorylated on Tyr unless expressed in cells that express Bcr-Abl proteins (Liu et al., 1996a), further confirming that the reactivity of the anti-pTyr360 antibody is specific for Bcr sequences containing pTyr-360. Analyses of the blot with anti-Bcr(1-16) (G143), shown in Fig. 13.6.5C, served as a loading control for Fig. 13.6.5A. The antiserum from a second rabbit (5793) turned out to have cross-reactivity with another protein (data not shown).
Time Considerations For Basic Protocols 1 and 2, the principal time-consuming step is the production of an immune response in the animals. For the production of polyclonal antibodies, high titers of reactive sera can be obtained as early as 6 to 8 weeks in animals that have been boosted every
2 weeks. Later bleeds with continued boosts can yield even higher titers. Once adequate bleeds have been obtained and the necessary affinity columns prepared, purification can generally be carried out within 1 week, with the exact length of time depending upon the number of columns and the necessity of analyzing the product after each step. In the production of monoclonal antibodies, splenocytes can typically be harvested at ∼2 months from the initial immunization, with several boosts in between. The screening of hybridoma supernatants can take 1 to 2 weeks, and their expansion and subcloning several weeks more.
Literature Cited Alberta, J.A. and Stiles, C.D. 1997. Phosphorylation-directed antibodies in high flex screens for compounds that modulate signal transduction. BioTechniques 23:490-493. Bangalore, L., Tanner, A.J., Laudano, A.P., and Stern, D.F. 1992. Antiserum raised against a synthetic phosphotyrosine-containing peptide selectively recognizes p185neu/erbB-2 and the epidermal growth factor receptor. Proc. Natl. Acad. Sci. U.S.A. 89:11637-11641. Banin, S., Moyal, L., Shieh, S.-Y., Taya, C., Anderson, C.W., Chessa, L., Smorodinsky, N.I., Prives, C., Reiss, Y., Shiloh, Y., and Ziv, Y. 1998. Enhanced phosphorylation of p53 by ATM in response to DNA damage. Science 281:16741677. Bodnar, R.J., Gu, M., Li, Z., Englund, G.D., and Du, X. 1999. The cytoplasmic domain of the platelet glycoprotein Ib is phosphorylated at serine 609. J. Biol. Chem. 274:33474-33479. Campbell, M.L. and Arlinghaus, R.B. 1991. Current status of the BCR gene and its involvement with human leukemia. Adv. Cancer Res. 57:227-256. Canman, C.E., Lim, D.-S., Cimprich, K.A., Taya, Y., Tamai, K., Sakaguchi, K., Appella, E.A., Kastan, M.B., and Siliciano, J.D. 1998. Activation of the ATM kinase by ionizing radiation and phosphorylation of p53. Science 281:1677-1679. Chang, C.D. and Meienhofer, J. 1978. Solid-phase peptide synthesis using mild base cleavage of N alpha-fluorenylmethoxycarbonylamino acids, exemplified by a synthesis of dihydrosomatostatin. Int. J. Dept. Protein Res. 11:246-249. Coghlan, M.P., Pillay, T.S., Tavare, J.M., and Siddle, K. 1994. Site-specific anti-phosphopeptide antibodies: Use in assessing insulin receptor serine/threonine phosphorylation state and identification of serine-1327 as a novel site of phorbol ester-induced phosphorylation. Biochem. J. 303:893-899. Czernik, A.J., Girault, J.-A., Nairn, A.C., Chen, J., Snyder, G., Kebabian, J., and Greengard, P. 1991. Production of phosphorylation state-specific antibodies. Methods Enzymol. 201:264283.
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Daley, G.Q., van Etten, R.A., and Baltimore, D. 1990. Inductin of chronic myelogenous leukemia in mice by the P210bcr-abl gene of the Philadelphia chromosome. Science 247:824-830. DiGiovanna, M.P. and Stern, D.F. 1995. Activation state-specific monoclonal antibody detects tyrosine phosphorylated p185neu/erbB-2 in a subset of human breast tumors overexpressing this receptor. Cancer Res. 55:1946-1955. Doolittle, R.F. 1986. Of URFS and ORFS: A Primer on How to Analyze Derived Amino Acid Sequences. University Science Books, Mill Valley, Calif. Epstein, R.J. 1995. Preferential detection of catalytically inactive c-erbB-2 by antibodies to unphosphorylated peptides mimicking receptor tyrosine autophosphorylation sites. Oncogene 11:315323. Epstein, R.J., Druker, B.J., Roberts, T.M., and Stiles, C.D. 1992. Synthetic phosphopeptide immunogens yield activation-specific antibodies to the c-erbB-2 receptor. Proc. Natl. Acad. Sci. U.S.A. 89:10435-10439. Frackelton, A.R. Jr., Ross, A.H., and Eisen, H.N. 1983. Characterization and use of monoclonal antibodies for isolation of phosphotyrosyl proteins from retrovirus-transformed cells and growth factor-stimulated cells. Mol. Cell. Biol. 3:1343-1352. Cooper, H.M. and Paterson, Y. 1995. Production of polyclonal antisera. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. 2.4.1-2.4.9. John Wiley & Sons, New York. Guo, J.Q., Wang, J.Y.J., and Arlinghaus, R.B. 1991. Detection of BCR-ABL proteins in blood cells of benign phase chronic myelogenous leukemia patients. Cancer Res. 51:3048-3051.
Kitas, E., Kung, E., and Bannwarth, W. 1994. Chemical synthesis of O-thiophosphotyrosyl peptides. Int. J. Pept. Protein Res. 43:146-153. Kloetzer, W., Kurzrock, R., Smith, L., Talpaz, M., Spiller, M., Gutterman, J., and Arlinghaus, R. 1985. The human cellular abl gene product in the chronic myelogenous leukemia cell line K562 has an associated tyrosine protein kinase activity. Virology 140:230-238. Konopka, J.B., Watanabe, S.M., and Witte, O.N. 1984. An alteration of the human c-abl protein in K562 leukemia cells unmasks associated tyrosine kinase activity. Cell 37:1035-1042. Liu, J., Campbell, M., Guo, J.Q., Xian, Y.M., Anderssen, B.S., and Arlinghaus, R.B. 1993. BcrAbl tyrosine kinase is autophosphorylated or transphosphorylates P160 Bcr on tyrosine predominantly within the first BCR exon. Oncogene 8:101-109. Liu, J., Wu, Y., Ma, G.Z., Lu, D., Haataja, L., Heisterkamp, N., Groffen, J., and Arlinghaus, R.B. 1996. Inhibition of Bcr serine kinase by tyrosine phosphorylation. Mol. Cell. Biol. 16:998-1005. Nairn, A.C., Detre, J.A., Casnellie, J.E., and Greengard, P. 1982. Serum antibodies that distinguish between the phospho- and dephospho-forms of a phosphoprotein. Nature 299:734-736. Ponticelli, A.S., Whitlock, C.A., Rosenberg, N., and Witte, O.N. 1982. In vivo tyrosine phosphorylations of the Abelson virus transforming protein are absent in its normal cellular homolog. Cell 29:953-960.
Hawk, N., Sun, T., Xie, S., Wang, Y., Wu, Y., Liu, J., and Arlinghaus, R. 2002. Inhibition of the BcrAbl oncoprotein by Bcr requires phosphoserine 354. Cancer Res. 62:386-390.
Puil, L., Liu, J., Gish, G., Mbamalu, G., Bowtell, D., Pelicci, P.G., Arlinghaus, R., and Pawson, T. 1994. Bcr-Abl oncoproteins bind directly to activators of the Ras signalling pathway. EMBO J. 13:763-773.
Heisterkamp, N., Stam, K., Groffen, J., de Klein, A., and Grosveld, G. 1985. Structural organization of the bcr gene and its role in the Ph′ translocation. Nature. 315:758-761.
Ross, A.H., Baltimore, D., and Eisen, H.N. 1981. Phosphotyrosine-containing proteins isolated by affinity chromatography with antibodies to a synthetic hapten. Nature 294:654-656.
Heisterkamp, N., Jenster, G., ten Hoeve, J., Zovich, D., Pattengale, P.K., and Groffen, J. 1990. Acute leukemia in Bcr-Abl transgenic mice. Nature. 344:251-253.
Roussel, R. 1990. Synthetic Phosphopeptides Modeled on the Carboxyl Terminus of pp60C-SRC: Specific Antibodies, Binding and Effects on Kinase Activity. M.S. thesis, University of New Hampshire, Durham.
Hornbeck, P., Fleisher, T.A., and Papadopoulos, N.M. 1991. Isotype determination of antibodies. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. 2.2.1-2.2.6. John Wiley & Sons, New York.
Sequence-Specific Anti-Phosphoamino Acid Antibodies
Kawakatsu, H., Sakai, T., Takagaki, Y., Shinoda, Y., Saito, M., Owada, M.K., and Yano, J. 1996. A new monoclonal antibody which selectively recognizes the active form of src tyrosine kinase. J. Biol. Chem. 271:5680-5685.
Ishida, R., Iwai, M., Marsh, K.L., Austin, C.A., Yano, T., Shibata, M., Nozaki, M., and Hara, A. 1996. Threonine-1342 in human topoisomerase II alpha is phosphorylated throughout the cell cycle. J. Biol. Chem. 271: 30077-30082.
Shtivelman, E., Lifshitz, B., Gale, R.P., and Canaani, E. 1985. Fused transcript of abl and bcr genes in chronic myelogenous leukaemia. Nature 315:550-554. Smith, R.G., Dev, V.G., and Shannon, W.A. Jr. 1981. Characterization of a novel human pre-B leukemia cell line. J. Immunol. 126:596-602. Stam, K., Heisterkamp, N., Reynolds, F.H. Jr, and Groffen, J. 1987. Evidence that the phl gene encodes a 160,000-dalton phosphoprotein with associated kinase activity. Mol. Cell. Biol. 7:1955-1960.
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Stern, D.F. 1991. Antiphosphotyrosine antibodies in oncogene and receptor research. Methods Enzymol. 198:494-501. Sun, T., Campbell, M., Gordon, W., and Arlinghaus, R.B. 2001. Preparation and application of antibodies to phosphoamino acid sequences. Biopolymers 60:61-75. Tzartos, S.J., Kouvatsou, R., and Tzartos, E. 1995. Monoclonal antibodies as site-specific probes for the acetylcholine-receptor δ-subunit tyrosine and serine phosphorylation sites. Eur. J. Biochem. 228:463-472. Wu, Y., Liu, J., and Arlinghaus, R.B. 1998. Requirement of two specific tyrosine residues for the catalytic activity of Bcr serine/theronine kinase. Oncogene 16:141-146. Wu, Y., Ma, G., Lu, D., Lin, F., Xu, H-J., Liu, J., and Arlinghaus, R.B. 1999. Bcr: A negative regulator of the Bcr-Abl oncoprotein. Oncogene 18:44164424. Zhang, H., Zha, X., Tan, Y., Hornbeck, P.V., Mastrangelo, A.J., Alessi, D.R., Polakiewicz, R.D., and Comb, M.J. 2002. Phosphoprotein analysis using antibodies broadly reactive against phosphorylated motifs. J. Biol. Chem. 277:3937939387.
Key References Sun et al., 2001 (see above) A description of the production of polyclonal antiphosphopeptide antibody by the method described in this unit. DiGiovanna, M.P., Stern, D.F., and Roussel, R.R. 2003. Production of antibodies that recognize specific tyrosine-phosphorylated peptides. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. 11.6.1-11.6.19. John Wiley & Sons, New York. Source of Basic Protocol 2 and Support Protocols 1 and 2. Harlow, E. and Lane, D. 1988. Antibodies: A Laboratory Manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. A comprehensive text describing the production and utilization of antibodies.
Contributed by Tong Sun and Ralph B. Arlinghaus University of Texas M.D. Anderson Cancer Center Houston, Texas
Post-Translational Modifications: Phosphorylation and Phosphates
13.6.27 Current Protocols in Protein Science
Supplement 34
Assays of Protein Kinases Using Exogenous Substrates
UNIT 13.7
In studies of the regulation of specific biochemical events by reversible phosphorylation, assaying the protein kinases themselves can often lead to significant progress in understanding the mechanistic details of a system under study. This unit describes assays for a variety of protein kinases that require different conditions to detect and measure their activities—cyclic nucleotide–dependent kinases (see Basic Protocol 1), protein kinase C and isoforms (see Basic Protocol 2), casein kinases (see Basic Protocol 3 and Alternate Protocol), Ca2+/calmodulin–dependent kinases (see Basic Protocol 4), and tyrosine kinase (see Basic Protocol 5). The unit is not meant to be a catalog of individual protein kinase assays; however, the general principles of these assays should apply to most if not all known protein kinases. It is also possible to perform in-gel assays for specific kinase activity (see Basic Protocol 6). To assay the phosphotransfer reactions catalyzed by protein kinases, it is necessary first to identify a target substrate for the transfer reaction. Essentially this means a substrate that is quite specific—i.e., is not phosphorylated very well or at all by other protein kinases under the appropriate assay conditions (see Strategic Planning). The source of the enzyme activity can be crude cell lysate (see Support Protocol 1), a cellular fraction (see UNIT 4.3), an immunoprecipitate, a partially purified protein, or a purified enzyme. The results of the assay can be evaluated using trichloroacetic acid (TCA) precipitation to measure incorporation of radioactivity (see Support Protocol 2), adsorption of the labeled material onto P81 phosphocellulose paper (see Support Protocol 3), or electrophoretic separation and autoradiography (see Support Protocol 4). STRATEGIC PLANNING The basic strategy for protein kinase assays involves the use of a labeled donor substrate so that when phosphotransferase activity is present in the enzyme sample, accumulation of the label in the protein or peptide acceptor substrate can be easily detected. The most often used protocol requires [γ-32P]ATP as the donor substrate and a specific protein or peptide as the acceptor substrate. Phosphotransfer is detected as the accumulation of [32P]-labeled protein or peptide. The main problem with this assay is that it requires 32P, a high-energy β emitter. There are a few examples of kinase assays that do not require [γ-32P]ATP, but they are far more specialized assays and generally not as widely applicable. Hopefully, further development of nonradioactive kinase assays will provide good alternatives and reduce the use of radioisotopes. Enzyme Sources Of all the choices to be made in designing protein kinase assays, of paramount importance is the source of enzyme activity: very crude sources such as whole-cell lysates (see Support Protocol 1 and UNIT 13.2), immunoprecipitates, partially purified preparations, and homogeneous purified protein can be the source of the enzyme activity. The first requirement is that the kinase activity be relatively stable under the conditions used to prepare the enzyme and under the conditions used for the assay. Often enzymes are unstable in whole-cell lysates or homogenates and become very stable only after an initial purification step. Protein kinases commonly exist in intact cells as phosphoproteins themselves, and the phosphorylation state is often important for activity. Thus, it is essential to attempt to preserve the phosphorylation state, e.g., by inhibiting protein phosphatases. Including specific kinase inhibitors in the lysis or homogenation buffer is Contributed by Nigel Carter Current Protocols in Protein Science (1997) 13.7.1-13.7.22 Copyright © 1997 by John Wiley & Sons, Inc.
Phosphorylation and Phosphatases
13.7.1 Supplement 9
a common procedure to inhibit serine/threonine kinases (sodium fluoride, β-glycerophosphate, okadaic acid) or tyrosine kinases (sodium vanadate, phenylarsinine oxide). Other common inclusions in lysis buffer are chelating agents and protease inhibitors. Many enzymes, including kinases, are sensitive to limited proteolysis. Chelators, such as EDTA and EGTA, chelate calcium and reduce the activity of calcium-activated proteases. The most commonly used protease inhibitors include phenylmethylsulfonyl fluoride (PMSF), leupeptin, pepstatin A, antipain, and benzamidine. It is always a good idea to add these inhibitors to the lysis buffer just before it is used to lyse cells, especially in the early stages of a new project; the need for specific inhibitors should be investigated at a later time. The oxidation state of cysteines and the state of disulfide linkages may influence enzyme activity, so inclusion of a reducing agent—e.g., 2-mercaptoethanol (2-ME) or dithiothreitol (DTT)—is sometimes essential to preserve enzyme activity. Low concentrations of a detergent such as Brij35, at or just above the critical micelle concentration, may also stabilize enzyme activity. Choice of Substrate Selecting a substrate normally depends on the type of experiment and the enzyme preparation, i.e., are there any contaminating kinases in the enzyme sample that phosphorylate the same or similar substrates. The type of experiment may range from following a particular activity through purification to determining substrate specificity or kinetics for the kinase-catalyzed reaction. Most often specific substrate information is available from previously published work or from initial experiments to identify a suitable substrate. It is always helpful to evaluate the suitability of several substrates in parallel experiments in order to identify the best substrate. For crude extracts, the best substrate will be one that is phosphorylated by the kinase of interest and by no other kinase. In practice this is rarely the case, so a compromise must be reached. If an antibody that recognizes the kinase is available, it is easier to use that antibody to immunoprecipitate the protein and use the immunoprecipitate as the source of enzyme activity. Immunoprecipitation is useful because it provides a quick one-step partial purification of the kinase that also concentrates the activity. With a partially purified enzyme, it may be easier to select an appropriate specific substrate. Histones are very basic proteins found complexed with DNA in the nucleus of the cell. They have been used for many years as protein kinase substrates because of their relative abundance, ease of purification, and ability to function as very good substrates for a large number of protein kinases. Distinct fractions of mixed histones and purified histones are available commercially from a wide variety of sources. Different kinases have different specificities with respect to phosphorylation of histones, hence the type of histone preparation that should be used depends on the type of kinase to be assayed. Histones have been used to assay many protein kinases and may be used for cAMP- and cGMPdependent protein kinases. However, phophorylation of histone substrates is generally not enzyme specific—e.g., histone H1 is phosphorylated by cAMP-dependent protein kinases, most cyclin/cdk complexes, and some protein kinase Cs albeit on different sites, and histone 2B is phosphorylated by both cAMP- and cGMP-dependent protein kinases and protein kinase B—so more specific substrates are generally preferred.
Assays of Protein Kinases Using Exogenous Substrates
Initially, it is much easier to use intact proteins as substrates for kinase assays. However, once a substrate is identified, specific determinant(s) within the protein that allow phosphorylation and specific primary sequence requirements can be identified, and the appropriate peptide sequences modeled on the phosphorylation site can be synthesized and used as substrate.
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Table 13.7.1 Substrate Specificities of Some Protein Serine/Threonine Kinases
Enzyme
Consensus sequencea,b
cAMP-dependent protein kinase cGMP-dependent protein kinase Protein kinase Cα Cyclin B/cdc2 Casein kinase I
XArgArgXSerΦ XArg/LysXSerΦ XLys/ArgXXSerX XSerProXLys/ArgX Glu/Ser(P) XXSer XArgXXSerX
Ca2+/calmodulin–dependent protein kinase type II MAP kinase Casein kinase II
XProSerXX XSerXGluGlu/AspX
aSee Table A.1A.1 for amino acid codes. X indicates any amino acid, Φ indicates
a hydrophobic residue, underlining indicates the sites of phosphorylation (which, although marked Ser in each case, may be either Ser or Thr residues), and (P) indicates a phosphorylated residue. bSequence information based on Pearson and Kemp (1991).
Table 13.7.2 Substrate Specificities of Some Protein Tyrosine Kinases
Enzyme
Optimal sequencea,b
MT/c-src c-abl
GluGluIleTyrGlyGluPheGlu AlaGluValIleTyrAlaAlaProPhe
c-fps/fes lck
GluGluGluIleTyrGluGluIleGlu XGluXIleTyrGlyValLeuPhe
EGF receptor PDGF receptor
GluGluGluGluTyrPheGluLeuVal GluGluGluGluTyrValPheIleGlu
Insulin receptor
XGluGluGluTyrMetMetMetMet
aThese sequences were determined through the use of peptide libraries
(Songyang et al., 1995), and their relevance to in vivo phosphorylation specificities is unclear. bSee Table A.1A.1 for amino acid codes. X indicates any amino acid;
underlining indicates the site of phosphorylation.
When synthetic peptide substrates are used to assay protein kinase activity, it is possible to study the specificity of different kinases. Synthetic peptide substrates are available from a number of suppliers, e.g., Peninsula Labs, Bachem Biosciences, and CalbiochemNovabiochem. Peptides are easy to use: they are potentially available in large quantities as chemically defined molecules, their purification is usually straightforward, and they provide a single phosphorylation site. In addition, it is relatively simple to synthesize a series of peptides with single amino acid changes to further define the specificity of an enzyme. However, peptides do have their disadvantages. They are expensive, $10 to $20 per residue. The kinase may not recognize the disordered structure usually adopted by small peptides in solution, or it may promiscuously phosphorylate the peptide. Finally, it is very difficult to use peptides to model reactions in which prior derivitization (i.e., phosphorylation) is required for specific kinase activity. Table 13.7.1 summarizes the sequence specificity of a number of protein serine/threonine kinases, and Table 13.7.2 provides similar information for protein tyrosine kinases.
Phosphorylation and Phosphatases
13.7.3 Current Protocols in Protein Science
Supplement 13
Assay Conditions The assay conditions can have profound effects on any enzyme activity: if pepsin is assayed at pH 7, it is essentially inactive, but at pH 3, it has good protease activity. Similarly T4 DNA ligase is inhibited by salt concentrations >150 mM. For kinases, it is a good idea to use preliminary experiments to identify the effects of pH, salt concentration, concentrations of cations such as Mg2+ or Mn2+, and temperature to optimize the assay conditions for the kinase of interest. It is often advantageous to perform kinase assays at 30°C rather than 37°C because the lower temperature makes it easier to stay within the linear range of the kinase, thus providing more control of the assay. Another factor to be considered when optimizing an assay for a newly identified kinase is the concentration of ATP in the reaction mixture. In these protocols and in general, detection of kinase activity is based on the transfer of radiolabeled phosphate from ATP to the substrate, so the concentration of ATP can be varied only a little. Also, the specific activity of the [γ-32P]ATP must be known in order to actually measure phosphotransfer. Most kinases have a KM for ATP of 1 to 100 µM. If there is too much ATP in the reaction mixture, it will be difficult to measure phosphotransfer. An ATP concentration of 50 to 100 µM tends to work well. At that concentration, the enzyme should be working at ≥50% of maximum, depending on its apparent KM for ATP, and addition of sufficient [γ-32P]ATP to measure phosphotransfer is not prohibitively expensive. Usually the substrate concentration is high so that the enzyme is working at or close to Vmax. Controls Including the correct controls for a kinase assay is critical to the success of the assay, especially when the enzyme source is a cell or tissue extract. The appropriate controls should always include a no substrate control, a no enzyme source control, a heat-denatured enzyme control, and, for enzymes that require an activator or cofactor, controls that use irrelevant activator or cofactor and controls without activator or cofactor. CAUTION: These assays use [γ-32P]ATP which should be handled and disposed of according to safety regulations. See APPENDIX 2B and the institutional Radiation Safety Office for guidelines for proper handling and disposal of 32P. BASIC PROTOCOL 1
ASSAY FOR CYCLIC NUCLEOTIDE–DEPENDENT PROTEIN KINASES Cyclic adenosine monophosphate (cAMP)–and cyclic guanosine monophosphate (cGMP)–dependent protein kinases (PrKs) are quite similar in structure and require similar assay conditions. cAMP-PrK is a heterotetramer consisting of two regulatory (R) subunits and two catalytic (C) subunits, R2C2. In each case the regulatory subunits have binding sites for two molecules of cAMP. When the cAMP concentration is elevated in cells after activation of receptor-linked adenylyl cyclases, cAMP binds to the R subunits, causing the affinity of the C subunits for R subunits to drop by about four orders of magnitude. The formation of R2cAMP4 dimers and free C-subunits is favored, and the C-subunits are now active. The case of cGMP-PrK is slightly different in that there are no free catalytic subunits released. cGMP-PrK is composed of two identical subunits. Each subunit has a regulatory domain and a catalytic domain, which are homologous to the R and C subunits of cAMP-PrK. On binding four molecules of cGMP the enzyme is activated, presumably by a conformational change.
Assays of Protein Kinases Using Exogenous Substrates
The assay procedures for these two kinases are very similar and the same protein substrate can be used for each one as detailed below. More recently, both of these kinases have been assayed using peptide substrates.
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Materials 5× cyclic nucleotide–dependent kinase reaction buffer (see recipe) 10 mg/ml histone 2B in H2O [γ-32P]ATP solution (see recipe) 20× cyclic nucleotide solution: 20 µM cyclic AMP in H2O/20 µM cyclic GMP in H2O Enzyme sample containing cyclic nucleotide–dependent kinase activity (see Support Protocol 1), kept on ice until use 30° water bath Additional reagents and equipment for TCA precipitation (see Support Protocol 2), adsorption onto P81 phosphocellulose paper (see Support Protocol 3), or electrophoretic analysis (see Support Protocol 4) 1. For each assay reaction, add the following to a 1.5-ml microcentrifuge tube kept on ice: 4 µl 5× cyclic nucleotide–dependent kinase reaction buffer 1 µl of 10 mg/ml histone 2B 1 µl [γ-32P]ATP solution (to give 5 µCi/µl and 5 µM ATP final) 1 µl 20× cyclic nucleotide solution 0 to 13 µl H2O. Cap tube and warm the mixture in a 30°C water bath. Perform each assay in triplicate and include no substrate, no enzyme, and no cyclic nucleotide controls. The total reaction mix volume is 20 ìl per reaction. The amount of water required depends on how much enzyme sample is used. Sufficient reaction mix can be prepared in a single tube for all the reactions by multiplying the quantities for a single reaction by the total number of reactions + 1. Then the total amount of reaction mixture (minus enzyme) for one reaction can be added to each tube. This is often very convenient and reduces the potential for pipetting errors.
2. Start the reaction by adding 1 to 14 µl ice-cold enzyme sample containing cyclic nucleotide–dependent kinase activity. The volume of enzyme used depends on the amount of activity in the enzyme sample. In a preliminary experiment, the maximum indicated amount can be used to gauge the extent of the phosphotransfer reaction. The volume can be reduced as appropriate in order to allow for linear incorporation of phosphate into the substrate during the assay. For enzyme samples that are immunoprecipitates adsorbed onto Sepharose beads, prepare the reaction mix minus enzyme source first, warm it, then dispense it into the immunoprecipitate. For immunoprecipitates, it is advisable to scale up the reaction for a total volume of 100 ìl: add 75 ìl reaction mix to 25 ìl immunoprecipitate bound to beads.
3. Incubate 10 min in a 30°C water bath. 4. Stop the reaction using the reagent appropriate for the analytic method to be used—20 µl ice-cold 10% TCA for TCA precipitation (see Support Protocol 2), or 10 µl or 20 µl ice-cold 2× SDS-PAGE sample buffer for electrophoretic analysis (see Support Protocol 4). Use 10 µl of the reaction mix for adsorption to P81 phosphocellulose paper (see Support Protocol 3). Proceed with analysis by one of those methods.
Phosphorylation and Phosphatases
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Supplement 9
BASIC PROTOCOL 2
ASSAY FOR PROTEIN KINASE C ISOFORMS There are presently at least ten members of the protein kinase C (PKC) family of kinases. Protein kinase C was originally identified as a kinase activity that was reversibly activated by calcium (Ca2+), phospholipid (usually phosphatidylserine), and the neutral lipid diacylglycerol (DAG; Takai et al., 1979). Subsequently, these kinases were shown to be the sites of action for the tumor-promoting phorbol esters, which bind to some PKC isoforms and activate the kinase activity by substituting for DAG. The initial isoforms of PKC that were purified were called α, β, and γ, and it is these forms that are the most clearly regulated by Ca2+, phospholipid, and DAG or phorbol esters. They are termed the “conventional” subfamily of PKCs (cPKCs). Another subfamily contains δ, ε, ν, and θ, the “novel” PKCs (nPKCs); these kinases are not regulated by Ca2+ but are activated by phorbol esters and phospholipids. A third subfamily, the “atypical” PKCs (aPKCs), consist of ζ, λ, and ι; these are not regulated by Ca2+ nor do they bind phorbol esters or DAG, but there is still a requirement for phospholipids. When the isoforms are analyzed, the reaction buffer must include the appropriate activating cofactors. Materials 5× PKC reaction buffer (see recipe) 10 mg/ml histone H1 in H2O [γ-32P]ATP solution (see recipe) Enzyme sample containing PKC activity (see Support Protocol 1) 30°C water bath Additional reagents and equipment for TCA precipitation (see Support Protocol 2), adsorption onto P81 phosphocellulose paper (see Support Protocol 3), or electrophoretic analysis (see Support Protocol 4) 1. For each assay reaction, add the following to a 1.5-ml microcentrifuge tube: 4 µl 5× PKC reaction buffer 1 µl 10 mg/ml histone H1 1 µl [γ-32P]ATP solution (to give 5 µCi/µl and 5 µM ATP final) 0 to 14 µl H2O. Cap tube and warm the mixture 10 min in a 30°C water bath. Perform each assay in triplicate and include no substrate and no enzyme controls. The total reaction mix volume is 20 ìl per reaction. The amount of water required depends on how much enzyme sample is used. Sufficient reaction mix can be prepared in a single tube for all the reactions by multiplying the quantities for a single reaction by the total number of reactions + 1. Then the total amount of reaction mixture (minus enzyme) for one reaction can be added to each tube. This is often very convenient and reduces the potential for pipetting errors.
2. Start reaction by adding 1 to 14 µl enzyme sample containing PKC activity to the warmed reaction mixture. The volume of enzyme used depends on the amount of activity in the enzyme sample. In a preliminary experiment, the maximum indicated amount can be used to gauge the extent of the phosphotransfer reaction. The volume can be reduced as appropriate in order to allow for linear incorporation of phosphate into the substrate during the assay. Assays of Protein Kinases Using Exogenous Substrates
For enzyme samples that are immunoprecipitates adsorbed onto Sepharose beads, prepare the reaction mix minus enzyme source first, warm it, then dispense it into the immunoprecipitate. For immunoprecipitates, it is advisable to scale up the reaction for a total volume of 100 ìl: add 75 ìl reaction mix to 25 ìl immunoprecipitate bound to beads.
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Current Protocols in Protein Science
3. Incubate 10 min in a 30°C water bath. 4. Stop the reaction using the reagent appropriate for the analytic method to be used—20 µl ice-cold 10% TCA for TCA precipitation (see Support Protocol 2), or 10 µl, or 20 µl ice-cold 2× SDS-PAGE sample buffer for electrophoretic analysis (see Support Protocol 4). Use 10 µl of the reaction mix for adsorption to P81 phosphocellulose paper (see Support Protocol 3). Proceed with analysis by one of those methods. ASSAY FOR CASEIN KINASES USING β-CASEIN Casein is a very acidic milk protein available in reasonable purity and quantity. Casein kinases were named for their ability to phosphorylate casein rather than other substrates available at the time. Because of its acidic nature, the use of casein as a substrate presents a number of problems, one of which is the fact that it does not carry a net positive charge at low pH and thus does not bind to P81 phosphocellulose paper. In recent years this problem has been circumvented by using short acidic peptides engineered to bind to P81 paper as substrates for the casein kinases (see Alternate Protocol).
BASIC PROTOCOL 3
There are currently two known mammalian casein kinases, casein kinase I and casein kinase II, named for their order of elution from anion-exchange chromatography columns. Casein kinases show a marked preference for substrates containing acidic residues close to the serine or threonine to be phosphorylated. Casein kinase I is a 37-kDa monomer that phosphorylates serine and threonione residues (see Table 13.7.1) and utilizes ATP only as phosphoryl group donor. The basic phosphorylation consensus sequence is Glu/Ser(P)XXSer (where the underlined residue is the phosphate acceptor, and may be either Ser or Thr). Mammalian casein kinase II is a tetramer of α2β2 structure: the α-subunit is 24 to 28 kDa and the β-subunit is 37 to 44 kDa. Its preferred substrate has aspartate or glutamate residues C-terminal to the phosphorylation site serine or threonine and a consensus of SerGluGlu/Asp. Casein kinase II is unusual in that it can utilize both ATP and GTP as phosphoryl group donor with similar KM values, ∼10 µM for ATP and ∼30 µM for GTP. Mammalian casein kinase II is also inhibited by heparin, whereas casein kinase I is not. The IC50 for the inhibition of casein kinase II by heparin is in the 10 to 20 nM range and at these concentrations casein kinase I is not inhibited. Assays of either casein kinase I or II can be performed using casein as substrate. When isolated from milk, casein is a phosphoprotein. However, it is preferable to use dephosphorylated casein as substrate; this is commercially available in a relatively pure state as either the α or β isoform. Materials 5× casein kinase reaction buffer (see recipe) 10 mg/ml β-casein in H2O [γ-32P]ATP solution (see recipe) Enzyme sample containing casein kinase activity (see Support Protocol 1), kept on ice 30°C water bath Additional reagents and equipment for TCA precipitation (see Support Protocol 2) or electrophoretic analysis (see Support Protocol 4) Phosphorylation and Phosphatases
13.7.7 Current Protocols in Protein Science
Supplement 9
1. For each assay reaction, add the following to a 1.5-ml microcentrifuge tube kept on ice: 4 µl 5× casein kinase reaction buffer 1 µl 10 mg/ml β-casein 1 µl [γ-32P]ATP solution (to give 5 µCi/µl and 5 µM ATP final) 1 to 14 µl H2O. Cap tube and warm the mixture 10 min in a 30°C water bath. This reaction mix can also be used for α-casein. The total reaction mix volume is 20 ìl per reaction. The amount of water required depends on how much enzyme sample is used. Sufficient reaction mix can be prepared in a single tube for all the reactions by multiplying the quantities for a single reaction by the total number of reactions + 1. Then the total amount of reaction mixture (minus enzyme) for one reaction can be added to each tube. This is often very convenient and reduces the potential for pipetting errors.
2. Start the reaction by adding 1 to 14 µl enzyme sample containing casein kinase activity. The volume of enzyme used depends on the amount of activity in the enzyme sample. In a preliminary experiment, the maximum indicated amount can be used to gauge the extent of the phosphotransfer reaction. The volume can be reduced as appropriate in order to allow for linear incorporation of phosphate into the substrate during the assay. For enzyme samples that are immunoprecipitates adsorbed onto Sepharose beads, prepare the reaction mix minus enzyme source first, warm it, then dispense it into the immunoprecipitate. For immunoprecipitates, it is advisable to scale up the reaction for a total volume of 100 ìl: add 75 ìl reaction mix to 25 ìl immunoprecipitate bound to beads.
3. Incubate 10 min in a 30°C water bath. 4. Stop the reaction using the reagent appropriate for the analytic method to be used—20 µl ice-cold 10% TCA for TCA precipitation (see Support Protocol 2), or 10 µl or 20 µl ice-cold 2× SDS-PAGE sample buffer for electrophoretic analysis (see Support Protocol 4). Proceed with analysis by one of those methods. ALTERNATE PROTOCOL
ASSAY FOR CASEIN KINASES USING A PEPTIDE SUBSTRATE Casein kinase I can be assayed with the peptide AspAspAspGluGluSerIleThrArgArg. Casein kinase II has most recently been assayed using specific peptide substrates, e.g., ArgArgArgGluGluGluThrGluGluGlu (where the underlined residue is the phosphate acceptor). In both cases the arginine residues allow binding to P81 phosphocellulose paper. The first peptide is relatively specific for casein kinase I, but the kinetics of its phosphorylation are not ideal; the KM is in the millimolar range. Thus, it is preferable to use this assay for casein kinase I only if the kinase is highly purified. Additional Materials (also see Basic Protocol 3) 10 mM synthetic peptide substrate solution (see recipe) for casein kinase: e.g., AspAspAspGluGluSerIleThrArgArg (for casein kinase I) or ArgArgArgGluGluGluThrGluGluGlu (for casein kinase II)
Assays of Protein Kinases Using Exogenous Substrates
Additional reagents and equipment for TCA precipitation (see Support Protocol 2), adsorption onto P81 phosphocellulose paper (see Support Protocol 3), or electrophoretic analysis (see Support Protocol 4)
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1. For each assay, add the following to a microcentrifuge tube: 10 µl 5× casein kinase reaction buffer 5 µl 10 mM synthetic peptide substrate solution for casein kinase 5 µl [γ-32P]ATP solution (to give 5 µCi/µl and 5 µM ATP final) 0 to 30 µl H2O. Cap the tube and warm 10 min in a 30°C water bath. The total reaction mix volume is 50 ìl per reaction. The amount of water required depends on how much enzyme sample is based. Sufficient reaction mix can be prepared in a single tube for all the reactions by multiplying the quantities for a single reaction by the total number of reactions + 1. Then the total amount of reaction mix minus enzyme for one reaction can be added to each tube. This is often very convenient and reduces the potential for pipetting errors.
2. Start the reaction by adding 1 to 30 µl enzyme sample containing casein kinase activity. The volume of enzyme used depends on the amount of activity in the enzyme sample. In a preliminary experiment, the maximum indicated amount can be used to gauge the extent of the phosphotransfer reaction. The volume can be reduced as appropriate in order to allow for linear incorporation of phosphate into the substrate during the assay. For enzyme samples that are immunoprecipitates adsorbed onto Sepharose beads, prepare the reaction mix minus enzyme source first, warm it, then dispense it into the immunoprecipitate. For immunoprecipitates, it is advisable to scale up the reaction for a total volume of 100 ìl: add 75 ìl reaction mix to 25 ìl immunoprecipitate bound to beads.
3. Incubate 10 min in a 30°C water bath. 4. Stop the reaction using the reagent appropriate for the analytic method to be used—20 µl ice-cold 10% TCA for TCA precipitation (see Support Protocol 2), or 10 µl or 20 µl ice-cold 2× SDS-PAGE sample buffer for electrophoretic analysis (see Support Protocol 4). Use 10 µl of the reaction mix for adsorption to P81 phosphocellulose paper (see Support Protocol 3). Proceed with analysis by one of those methods. ASSAY FOR Ca2+/CALMODULIN–DEPENDENT KINASES 2+
2+
As the name implies, Ca /calmodulin–dependent kinases are dependent on Ca and calmodulin for their activity. Ca2+/calmodulin–dependent kinase I (CaM kinase I) is a monomeric protein of ∼40 kDa. Ca2+/calmodulin–dependent kinase II (CaM kinase II) is composed of the products of four distinct genes for α, β, γ, and δ subunits. In its native form, CaM kinase II is an oligomer of Mr ∼500,000 to 600,000. As its name suggests, both Ca2+ and calmodulin are required for activity.
BASIC PROTOCOL 4
Little is known of the physiological function of CaM kinase I. Activity can be assayed using the protein synapsin 1 as substrate, but both CaM kinase 1 and CaM kinase II phosphorylate full-length synapsin 1. This problem can be overcome by using synthetic peptide substrates that include the specific phosphorylation site for the enzyme being assayed. CaM kinase I can be assayed using a peptide containing site 1 of synapsin I, TyrLeuArgArgArgLeuSerAspSerAsnPhe (where the underlined residue is the phosphate acceptor). CaM kinase II can be assayed using autocamtide, LysLysAlaLeuArgGlnGluThrValAspAlaLeu (Hanson et al., 1989), which is modeled on the autophosphorylation site of the kinase itself. Phosphorylation and Phosphatases
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Materials 5× CaM kinase reaction buffer (see recipe) 10 mM synthetic peptide substrate solution (see recipe): e.g., TyrLeuArgArgArgLeuSerAspSerAsnPhe (for CaM kinase I) or LysLysAlaLeuArgGlnGluThrValAspAlaLeu (autocamtide for CaM kinase II) [γ-32P]ATP solution (see recipe) 1 mg/ml calmodulin in Milli-Q-purified water (store small aliquots at −70°C) Enzyme sample containing CaM kinase activity (see Support Protocol 1), kept on ice 30°C water bath Additional reagents and equipment for adsorption onto P81 phosphocellulose paper (see Support Protocol 3) 1. Set up and label duplicate 1.5-ml microcentrifuge tubes. For each reaction, prepare a mixture containing: 5 µl 5× CaM kinase reaction buffer 5 µl 10 mM synthetic peptide substrate solution 5 µl [γ-32P]ATP solution (to give 5 µCi/µl and 5 µM ATP final) 5 µl 1 mg/ml calmodulin or 5 µl H2O 0 to 30 µl H2O. Cap tube and warm the mixture 10 min in a 30°C water bath. The reaction mix is described for 50 ìl per reaction, but it can be carried out in volumes of 25 to 100 ìl by adjusting the amounts of the ingredients proportionally. Control reactions should include one reaction without calmodulin so that any background phosphotransfer reactions can be accounted for and only calcium2+/calmodulin–dependent phosphotransfer is measured.
2. Start each set of reactions sequentially every 30 sec by adding 1 to 25 µl enzyme sample containing CaM kinase activity to one of the tubes. The volume of enzyme used depends on the amount of activity in the enzyme sample. In a preliminary experiment, the maximum indicated amount can be used to gauge the extent of the phosphotransfer reaction. The volume can be reduced as appropriate in order to allow for linear incorporation of phosphate into the substrate during the assay. For enzyme samples that are immunoprecipitates adsorbed onto Sepharose beads, prepare the reaction mix minus enzyme source first, warm it, then dispense it into the immunoprecipitate. For immunoprecipitates, it is advisable to scale up the reaction for a total volume of 100 ìl: add 75 ìl reaction mixture to 25 ìl immunoprecipitate bound to beads.
3. Incubate for 10 min in a 30°C water bath. 4. Once the desired incubation time has elapsed, pipet 30 µl of the contents of the tube quickly onto the surface of prepared P81 paper squares (see Support Protocol 3). Only adsorption onto P81 phosphocellulose (see Support Protocol 3) is used to measure CaM kinase–mediated transfer because peptides are not efficiently precipitated by TCA, and they require specialized techniques for analysis by PAGE.
Assays of Protein Kinases Using Exogenous Substrates
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ASSAY FOR TYROSINE KINASES Acid-denatured rabbit muscle enolase was first used as an exogenous tyrosine kinase substrate in the early 1980s, and it is still a useful substrate in a number of experimental situations. Usually enolase is used as a substrate for cytosolic tyrosine kinases such as pp60c-src. Once the reaction is complete, phosphorylation of enolase is detected by separating proteins in the reaction mixture by SDS-PAGE (UNIT 10.1).
BASIC PROTOCOL 5
The following method is optimized for pp60c-src, but may be applied to other tyrosine kinases. It is also useful as a framework for the assay of any tyrosine kinase, but optimization of the immunoprecipitation and assay buffers is required for the tyrosine kinase of interest. Materials Rabbit muscle enolase (enolase EC 4.2.1.11; ammonium sulfate precipitate or suspension purified from rabbit muscle) 1 mM DTT/50 mM HEPES, pH 7.0 Glycerol 100 mM acetic acid 5× tyrosine kinase reaction buffer (see recipe) [γ-32P]ATP solution (see recipe) Enzyme sample containing tyrosine kinase activity (see Support Protocol 1), kept on ice 2× SDS-PAGE sample buffer (UNIT 10.1), ice-cold 30°C and boiling water bath Additional reagents and equipment for SDS-PAGE (see Support Protocol 4 and UNIT 10.1) 1. Microcentrifuge the equivalent of 100 µg rabbit muscle enolase 5 min at maximum speed, 4°C. Discard the supernatant. 2. Add 10 µl of 1 mM DTT/50 mM HEPES, pH 7.0, to the pellet, mix thoroughly, and incubate 30 to 60 min on ice. 3. Add 10 µl glycerol, mix, and then keep on ice if assays are to be performed the same day. This preparation can also be stored at −70°C until required.
4. Immediately prior to assay, add 20 µl of 100 mM acetic acid to the enolase solution and mix thoroughly. Incubate this mixture 5 min in a 30°C water bath, and then keep on ice until the reaction mixture is prepared. Do not store acid-denatured enolase on ice in this state for >1 hr.
5. For each reaction, add the following to a 1.5-ml microcentrifuge tube: 4 µl 5× tyrosine kinase reaction buffer 1 µl acid-denatured enolase (step 4; 2.5 µg enolase total) 1 µl [γ-32P]ATP (to give 5 µCi/µl and 5 µM ATP final) 0 to 14 µl H2O. Cap the tube and warm the mixture 10 min in a 30°C water bath. The amount of water required depends on the amount of enzyme sample to be added; the total reaction volume should be 20 ìl. Sufficient reaction mix can be prepared in a single tube for all the reactions by multiplying the quantities for a single reaction by the total number of reactions + 1. Then the total
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amount of reaction mix minus enzyme for one reaction can be added to each tube. This is often very convenient and reduces the potential for pipetting errors.
6. Start the reactions by adding 1 to 14 µl enzyme sample containing tyrosine kinase activity. The volume of enzyme used depends on the amount of activity in the enzyme sample. In a preliminary experiment, the maximum indicated amount can be used to gauge the extent of the phosphotransfer reaction. The volume can be reduced as appropriate in order to allow for linear incorporation of phosphate into the substrate during the assay. For enzyme samples that are immunoprecipitates adsorbed onto Sepharose beads, prepare the reaction mix minus enzyme source first, warm it, then dispense it into the immunoprecipitate. For immunoprecipitates, it is advisable to scale up the reaction for a total volume of 100 ìl: add 75 ìl reaction mixture to 25 ìl immunoprecipitate bound to beads.
7. Incubate the tube 10 min in a 30°C water bath. 8. Stop the reaction by adding 20 µl ice-cold 2× SDS-PAGE sample buffer. Mix thoroughly and heat the tube 3 min in a boiling water bath. Analyze 20 µl of the reaction by SDS-PAGE (see Support Protocol 4 and UNIT 10.1). Most tyrosine kinases autophosphorylate on a number of tyrosine residues and in in vitro assays are promiscuous with respect to substrate. Backgrounds are often very high so TCA precipitation and adsorption to P81 phosphocellular paper are not very useful for analysis of results. SDS-PAGE (see Support Protocol 4) gives more specific and cleaner results. BASIC PROTOCOL 6
IN-GEL PROTEIN KINASE ASSAYS In-gel protein kinase assays (Kameshita and Fujisawa, 1989) involve the copolymerization of a kinase substrate in the separating gel layer of an SDS-PAGE gel. Protein samples are run on the modified gel as usual (UNIT 10.1). After electrophoresis is complete, the separated proteins are allowed to refold by various gel treatments. The gel is then incubated with [γ-32P]ATP at 30°C for enough time to allow the ATP to penetrate the entire gel. After further washes to remove unincorporated [γ-32P]ATP, the gel is stained and fixed as normal. 32P-containing proteins are revealed by autoradiography. Although in general these assays are not particularly sensitive, they are sometimes very useful for identifying kinases in complex mixtures. Materials 10 mg/ml kinase substrate: e.g., myelin basic protein Enzyme sample containing kinase activity (see Support Protocol 1), kept on ice 20% (v/v) 2-propanol/50 mM Tris⋅Cl (pH 8.0 at room temperature; APPENDIX 2E) 1 mM DTT/50 mM Tris⋅Cl (pH 8.0 at room temperature) 6 M guanidine⋅HCl/1 mM DTT/50 mM Tris⋅Cl (pH 8.0 at room temperature) or 8 M urea/1 mM DTT/50 mM Tris⋅Cl (pH 8.0 at room temperature) 1 mM DTT/0.05% (v/v) Tween 20/50 mM Tris⋅Cl (pH 8.0 at 4°C) Appropriate kinase reaction buffer (see recipes) 10 mM Mg/ATP solution (see recipe) 10 mCi/ml [γ-32P]ATP (3000 Ci/mmol; Amersham, DuPont NEN, or ICN Biomedicals) 5% (w/v) trichloroacetic acid (TCA) 1% (w/v) sodium pyrophosphate/5% (w/v) TCA
Assays of Protein Kinases Using Exogenous Substrates
Container for radioactive incubation: e.g., small tray with tight cover, or heat-sealable polyethylene bag (Seal-a-Meal) Seal-a-Meal apparatus (optional) Additional reagents and equipment for SDS-PAGE (UNIT 10.1)
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CAUTION: Because the samples are radioactive, great care should be exercised in sample preparation and loading. Perform the reactions and subsequent manipulations in screwcap microcentrifuge tubes to minimize the risk of 32P contamination. Run gels until the dye front passes out of the gel, as all residual [γ-32P]ATP will migrate with the dye front. This means that the SDS-PAGE running buffer will be contaminated with 32P, and it should be disposed of as radioactive waste according to safety regulations. See APPENDIX 2B for proper handling and disposal. Separate proteins on modified gel 1. Prepare the mixture for a polyacrylamide gel of the appropriate %T (see UNIT 10.1). Add 10 mg/ml kinase substrate to give a final concentration of 1 mg/ml in the gel mixture. Polymerize as usual in a cassette prepared for casting the gel. Myelin basic protein has been used with some success for assays of the MAP kinase family of enzymes. However, any protein is suitable if it is a substrate for the kinase of interest. This assay can also be used to identify potential substrates.
2. Prepare the enzyme samples containing kinase activity for SDS-PAGE in the usual way and carry out electrophoresis (see UNIT 10.1). 3. Once separation is complete, place the gel in 200 ml of 20% 2-propanol/50 mM Tris⋅Cl, pH 8.0. Wash the gel for 20 min, change to fresh buffer, and repeat twice for a total of three 20-min washes. Washing with propanol removes SDS from the gel.
4. Wash the gel with 200 ml of 1 mM DTT/50 mM Tris⋅Cl (pH 8.0) three times, 20 min each. 5. Wash the gel with 250 ml of either 6 M guanidine⋅HCl/1 mM DTT/50 mM Tris⋅Cl (pH 8.0), or 8 M urea/1 mM DTT/50 mM Tris⋅Cl (pH 8.0) twice, 30 min each. Either guanidine⋅HCl or urea can be used to denature the proteins. Depending on the kinase, one or the other may be more successful; guanidine⋅HCl should be tried first, because it seems to work best for most kinases.
6. Wash the gel eight to ten times at 4°C over a period of 18 hr with 250 ml of 1 mM DTT/0.05% Tween 20/50 mM Tris⋅Cl (pH 8.0), to renature proteins. Assay for kinase activity 7. Incubate the gel with 250 ml of an appropriate kinase reaction buffer containing 10 mM MgCl2, for 20 min at 30°C. 8. Remove all traces of buffer and place the gel in a small tray or a Seal-a-Meal bag (the container is to be used for radioactive incubation). Add as small an amount of kinase reaction buffer as possible to just cover the gel. Add 1/4 of the reaction buffer volume of 10 mM Mg/ATP solution to give a final concentration of 50 µM ATP. Add 20 µCi/ml [γ-32P]ATP. 9. Incubate 1 hr at 30°C. The duration of this incubation can be adjusted after viewing the results of the initial experiment.
10. Incubate the gel in 250 ml 5% TCA twice, 15 min each. Wash 10 min in 500 ml of 1% sodium pyrophosphate/5% TCA. Repeat this wash until the solution contains little or no radioactivity. 11. Dry the gel onto filter paper and autoradiograph.
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SUPPORT PROTOCOL 1
PREPARING A CELL LYSATE FOR KINASE ASSAYS This protocol describes a method for detergent-induced cell lysis to prepare a crude extract containing kinase enzyme activity. Other methods of cell lysis may be appropriate. Hypotonic lysis and isolation of P100 and S100 fractions can also provide useful data on the recovery of a protein kinase as a soluble or membrane-bound activity. Materials Cultured cells: adherent cells at ∼70% confluence in 100-mm tissue culture dishes or suspension cells at 106 cells/ml PBS (APPENDIX 2D), ice-cold Lysis buffer (see recipe) Protease inhibitor stock solutions (see recipe) Microcentrifuge, 4°C 1. Wash the cultured cells twice in ice-cold PBS. Completely aspirate the final wash solution. 2. Add 0.75 ml lysis buffer to ∼2 × 107 cells; for adherent cells check that the monolayer is completely covered. Add protease inhibitors from stock solutions to the appropriate final concentration: 1 mM PMSF, 100 µM benzamidine, 5 µg/ml leupeptin, 5 µg/ml pepstatin A, and 5 µg/ml antipain. A 0.75-ìl lysate should provide sufficient material for immunoprecipitation (use 25% to 50% of the lysate for an initial experiment and scale up or down depending on the results) or simple purification. If the purification procedure involves more than one chromatography step, prepare a larger lysate because only 10% to 20% of the activity is recovered from each chromatography step.
3. Incubate the cells 10 min at 4°C. Scrape the cells from the dish and transfer the lysate to a labeled microcentrifuge tube. 4. Microcentrifuge the lysate 10 min at maximum speed, 4°C. Carefully remove the supernatant and transfer it to a clean microcentrifuge tube. Lysates should be used immediately until proper storage conditions (e.g., −70°C, liquid nitrogen) can determined experimentally. For some kinases, it may not be possible to store the lysate at all.
SUPPORT PROTOCOL 2
TCA PRECIPITATION TO DETERMINE INCORPORATION OF RADIOACTIVITY One of the classical methods for separating a reaction product from the reactants is differential precipitation. In the case of protein kinase assays using a protein substrate and [γ-32P]ATP, it is very easy to precipitate the protein and leave the [γ-32P]ATP in the soluble fraction. Most proteins are quantitatively precipitated by trichloroacetic acid (TCA), so TCA can be used to precipitate proteins phosphorylated during a kinase assay. Protein precipitate can be captured by filtration and the filter can be washed with TCA. TCA precipitation is a quick and reproducible way to determine the extent of [32P]phosphate transfer to protein substrates. However, peptides are not precipitated by TCA, so adsorption of labeled peptides to P81 phosphocellulose paper (see Support Protocol 3) is a more suitable method for analyzing the results of kinase assays that use a peptide as substrate.
Assays of Protein Kinases Using Exogenous Substrates
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Materials Assay samples (see Basic Protocols 1 to 5 and Alternate Protocol) 5% and 10% (w/v) trichloroacetic acid (TCA), ice-cold 95% ethanol Diethyl ether Whatman GF-C glass-fiber filters Vacuum manifold (e.g., Fisher) 20-ml scintillation vials Scintillation counter 1. Stop the reaction by adding 20 µl ice-cold 10% TCA. Mix thoroughly and incubate 10 min on ice to precipitate protein. 2. Pipet sample onto Whatman GF-C glass-fiber filter held in a vacuum manifold. Allow the sample to pass through the filter. Rinse the tube with 500 µl ice-cold 5% TCA. Add this wash to the appropriate filter. 3. Wash each filter with 5 ml ice-cold 5% TCA solution four times. Wash once with 10 ml of 95% ethanol, then three times with 10 ml diethyl ether. Allow each filter to dry for a few minutes. CAUTION: Diethyl ether is extremely flammable and washes should be performed in a fume hood away from any flame or heat source. Ether washes should be disposed of in an appropriate manner.
4. Place the dry filters in a 20-ml scintillation vial and count in a scintillation counter by detection of Cerenkov radiation. Alternatively, if dpm quantitation is desired, place dry filter in 10 ml scintillation fluid and count. The filter can also be counted by Cerenkov radiation after the washes with 95% ethanol. If the filters are to be counted in scintillant, however, the diethyl ether washes are essential, because TCA (which is removed by the ether) causes significant chemiluminescence when placed in scintillation fluid, making the counts produced by the scintillation counter meaningless.
ADSORPTION ONTO P81 PHOSPHOCELLULOSE PAPER One of the main methods to separate [32P]-labeled proteins or peptides from [γ-32P]ATP after a protein kinase assay reaction is by adsorbing the protein or peptide to P81 phosphocellulose paper. P81 paper is an ion-exchange matrix with net negative charge at most pHs. At low pH (such as in 75 mM orthophosphoric acid, which is used to wash the paper in this protocol), the excess [γ-32P]ATP left after a kinase assay will not bind. Under the same conditions, however, the phosphorylated peptide will bind to the P81 paper if it carries a net positive charge at low pH. In practice, this is usually achieved by tagging a peptide with two or three arginine or lysine residues at the N- or C-terminus. At the pH of 75 mM orthophosphoric acid the arginyl or lysyl side chains are positively charged and will bind to P81 paper. It is necessary, of course, that the activity of the kinase in question is not affected by the addition of basic residues to the substrate. Usually five or more residues are placed between the hydroxy-amino acid phosphorylation site and the basic residues required for binding to P81 paper. When proteins are used as substrates for kinases, the net charge of the protein is also a consideration for adsorption to P81 paper. As for peptides, adsorption to P81 paper is more suitable for basic proteins than for acidic proteins. Under the conditions described in this protocol, histones bind very well to P81 paper because they are highly basic, but casein binds very poorly or not at all because it is a very acidic protein.
SUPPORT PROTOCOL 3
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fold here label here in pencil P81 paper
board of nonabsorbent material
Figure 13.7.1 Addition of sample to P81 phosphocellulose paper. The 2 × 2–cm squares of P81 paper are labeled using a pencil (ink is dissolved by the acetone wash), and 50% to 75% of the reaction product is applied to the paper.
Materials Assay samples (see Basic Protocols 1 to 5 and Alternate Protocol) 75 mM orthophosphoric acid (stored up to 6 months at room temperature) Acetone 2 × 2–cm squares of P81 phosphocellulose paper (Whatman) 500-ml plastic beaker with the bottom replaced with solvent-resistant plastic or wire mesh (see Fig. 13.7.2) 20-ml scintillation vial Scintillation counter 1. Cut 2 × 2–cm squares of P81 paper, fold the end over, and label the folded end with pencil as shown in Figure 13.7.1. Place each square on a large piece of nonabsorbent material, such as plastic or acrylic sheeting or an aluminum foil–covered fiberboard. Use a soft pencil to label the squares of P81 paper, as the final acetone wash will remove or destroy ink labels.
2. Pipet 15 or 30 µl of the contents of each assay tube quickly onto the surface of the prepared P81 paper squares. Immediately place the paper squares in a 500-ml plastic beaker with the bottom replaced with solvent-resistant plastic or wire mesh (Fig. 13.7.2). Assays of Protein Kinases Using Exogenous Substrates
Usually 50% to 75% of the final reaction mixture is spotted onto PB1 paper to ensure that there is sufficient signal to be detected. If the plastic mesh used in the beaker is not solvent-resistant, the final acetone washes will dissolve it!
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P81
solvent- resistant mesh
stir bar
magnetic stirrer
Figure 13.7.2 Setup for washing P81 phosphocellulose paper with reaction samples. The paper squares are washed five times for 5 min each in 75 mM orthophosphoric acid and once in acetone. They are then air dried and counted in a scintillation counter.
3. Place this beaker into a larger beaker containing 75 mM orthophosphoric acid and incubate 5 min with stirring. 4. Pour the used orthophosphoric acid wash into a container suitable for 32P-containing liquid waste. Refill the large beaker with 75 mM orthophosphoric acid and incubate 5 min more. Repeat for a total of five 5-min washes. 5. Fill the large beaker with acetone and wash the P81 papers for 5 min. Discard the acetone and allow the papers to dry. 6. Place each square of P81 paper in a 20-ml scintillation vial and count by detection of Cerenkov radiation. Alternatively, if the absolute dpm value is desired and a 32P quenching curve is available for the scintillation counter, add scintillation fluid to the vials and count.
ELECTROPHORETIC ANALYSIS OF PHOSPHORYLATION Electrophoretic analysis of phosphorylation allows a more specific determination of protein substrate phosphorylation, because other phosphoproteins are separated by electrophoresis. It is advantageous to use the resolving power of SDS-PAGE to assay relatively crude enzyme samples; this method of analysis may also give useful information on kinase autophosphorylation.
SUPPORT PROTOCOL 4
Stop the reaction (see Basic Protocols 1 to 5 or Alternate Protocol) by adding 20 µl ice-cold 2× SDS-PAGE sample buffer (UNIT 10.1). Mix thoroughly, heat the sample 3 min in a boiling water bath, then analyze by SDS-PAGE (UNIT 10.1). Stain, fix, dry, and autoradiograph the gel.
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REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
[γ-32P]ATP solution Dilute 10 mM Mg/ATP solution (see recipe) 1/10 in water to give a 1 mM solution. For the assay, add 10 µl of 1 mM Mg/ATP to 80 µl water and then add 10 µl of 10 mCi/ml [γ-32P]ATP (3.3 pmol ATP). The final ATP concentration is ∼100 ìM (plus 33 nM from the [γ-32P]ATP, which is negligible), and the 100 ìCi (∼2.2 × 108 dpm) in 100 ìl gives a specific activity of 1 ìCi/ìl or ∼2.2 × 107 dpm/nmol. Suppliers for [γ-32P]ATP include Amersham, DuPont NEN, and ICN; it is usually available at a variety of specific activities. A good specific activity to use is 3000 Ci/mmol, normally provided by the manufacturer at a concentration of 10 mCi/ml. Amersham’s Redivue is a convenient [γ-32P]ATP solution to use as it is stored at 4°C instead of −20°C. As 1 Ci is ∼ 2.2 × 1012 dpm, then for a specific activity of 3000 Ci/mmol = 3 Ci/ìmol = 6.6 × 1012 dpm/ìmol = 6.6 × 106 dpm/pmol = 2.2 × 106 dpm ∼ 1 ìCi ∼ 0.33 pmol. The stock Mg/ATP solution prepared earlier can be diluted to a working ATP solution of the desired concentration (in this example, 0.1 mM).
CaM kinase reaction buffer, 5× 100 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 25 mM MgCl2 1 mM CaCl2 Store up to 6 months at −20°C Casein kinase reaction buffer, 5× 100 mM Tris⋅Cl, pH 7.5 at 30°C (APPENDIX 2E) 25 mM MgCl2 2.5 mM DTT 750 mM KCl Store up to 6 months at −20°C Cyclic nucleotide–dependent protein kinase reaction buffer, 5× 250 mM Tris⋅Cl, pH 7.5 at 30°C (APPENDIX 2E) 25 mM MgCl2 Store up to 6 months at −20°C Lysis buffer 50 mM HEPES (N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid), pH 7.4 100 mM NaCl 50 mM sodium fluoride 5 mM β-glycerophosphate 2 mM EDTA 2 mM EGTA 1 mM sodium vanadate (Na3VO4) 1% (v/v) Nonidet P-40 (NP-40) or Triton X-100 Store up to 6 months at −20°C Assays of Protein Kinases Using Exogenous Substrates
To stabilize kinase activity, specific kinase inhibitors, protease inhibitors, and reducing agents may be added to the lysis buffer (see Strategic Planning, Enzyme Sources).
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Mg/ATP solution, 10 mM Stock solutions: 20 mM MgCl2 or MgSO4 20 mM Na2ATP Working solution: Mix equal volumes of the two stock solutions and measure the pH. Adjust the pH slowly to pH 7.4 with 100 mM HCl or 100 mM NaOH. Store up to 1 year in small aliquots at −20°C. If a precipitate forms, the solution should be stirred continually and the pH maintained at 7.4. Adenosine trisphosphate is normally purchased as a sodium salt. Most kinases require ATP to be complexed with magnesium for its efficient utilization as a substrate. In some cases manganese can substitute for Mg2+.
PKC reaction buffer, 5× Solvent: 100 mM Tris⋅Cl, pH 7.5 at 30°C (APPENDIX 2E) 25 mM MgCl2 1 mM CaCl2 Store up to 6 months at −20°C Lipids: Just before the buffer is to be used, transfer enough 5 µg/ml phosphatidylserine stock solution in chloroform to provide 100 µg phosphatidylserine/ml buffer (final concentration) to a clean tube. Add sufficient 0.5 µg/ml Diolein stock solution (Sigma) in chloroform to provide 10 µg Diolein/ml buffer (final concentration) to the same tube and mix. Dry the lipids under a stream of nitrogen. Working solution: Add the required volume solvent (see above) and place on ice for 10 min. Sonicate the mixture at medium power for 1 min while the sample is on ice. Let the sample sit on ice 2 min. Repeat sonication 4 more times for a total of 5 min. Discard any unused buffer. Diolein is the old name for various preparations that contain mixed isomers of diacylglycerol, the cofactor for PKC. Diolein is a suitable source for the cofactor in these assays, and it is less expensive than pure preparations of 1,2-diacylglycerol.
Protease inhibitor stock solutions 100 mM phenylmethylsulfonyl fluoride (PMSF) in 100% ethanol 100 mM benzamidine 1 mg/ml leupeptin 1 mg/ml pepstatin A 1 mg/ml antipain Store up to 6 months at −20°C Synthetic peptide substrate solution Prepare 10 mM solutions of each peptide (highest quality available) in Milli-Q water and adjust pH to 7.4 before storage at −70°C or use. If the peptide is insoluble, check the sequence—a peptide with a high proportion of acidic residues may be more soluble at alkaline pH, and one with a high proportion of basic residues may be more soluble at acidic pH. Hydrophobic peptides may be solubilized by adding up to 20% (v/v) acetonitrile. Whatever is added must not compromise the assay conditions: e.g., if a peptide must be dissolved at pH 10, adding the solution to the reaction mix must not change the pH of the reaction. Most synthetic peptides are quite stable when stored at −70°C; the stability of individual peptide substrates should be determined for the storage conditions used, however. If peptides are custom synthesized they should ideally be purified by HPLC before use.
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Tyrosine kinase reaction buffer, 5× 100 mM HEPES (N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid), pH 7.4 5 mM MnCl2 5 mM DTT 500 µM sodium vanadate (Na3VO4) Store up to 6 months at −20°C COMMENTARY Background Information
Assays of Protein Kinases Using Exogenous Substrates
In recent years there has been an explosion of interest in protein kinases. It is now obvious that this class of enzyme plays many essential roles in a large array of biological processes, including the cellular responses to hormones, control of most stages of the cell cycle, and development of the nervous system in systems as diverse as fruit fly and man. Because of this interest, a very large number of protein kinase and protein phosphatase genes have been identified; current estimates are that there are well over 2000 genes encoding these enzymes in mammalian genomes. Obviously the study of protein kinases and phosphatases can provide fundamental information regarding the regulation of many diverse systems; the protocols provided in this unit include enough information to enable researchers new to this type of experiment to embark on the study of protein kinases. Protein kinases can be assayed using two types of substrate, intact proteins or synthetic peptides, and there are advantages and disadvantages to the use of each. A number of intact proteins may be used as general protein kinase substrates, and those in common use are available in relatively pure form from a number of manufacturers at reasonable prices. The main disadvantage to the use of proteins as substrates is the lack of specificity, sometimes making interpretation of experimental data difficult. Intact proteins such as myelin basic protein (MBP) contain many different phosphorylation sites. MBP can be phosphorylated by MAP kinase, some protein kinase C isoforms, and Ca2+/calmodulin kinase II, each at one or more sites. Hence, it is often impossible to differentiate which kinase is responsible for phosphorylating MBP in a complex mixture like a whole cell lysate. Some proteins cause headaches because they cannot be used with simple experimental protocols, e.g., casein. The use of synthetic peptides improves the specificity of the reaction by providing a single, well-defined phosphorylation site. They also allow experiments to define the concensus se-
quence for phosphorylation by a particular protein kinase and make it possible to design a substrate that has excellent kinetic properties for the kinase of interest. Peptides are often easier to use than their protein counterparts as in the case of casein kinases, where peptides can be designed that allow the use of phosphocellulose paper rather than other more involved methods for assessment of experimental results. Use of peptides is not without disadvantages though, the most significant being the cost of producing reasonable quantities of large numbers of peptides. When this is coupled with the observation that sometimes peptides are not recognized by the kinase because some structural element is missing, using peptide substrates can be a frustrating and expensive business. However, this is the exception rather than the rule, and the use of specific peptides to assay protein kinases is ultimately the best choice. One unit of kinase activity is usually defined as the amount of enzyme required to transfer 1 µmol of phosphate to a substrate from ATP per minute. From the following example it is possible to calculate a number of the reaction parameters: enzyme + [γ-32P]ATP + substrate → enzyme + ADP + [32P]product For this example, 1 mg protein (based on protein assay; see UNIT 3.4) was used for the assay. The [γ-32P]ATP in the assay mixture had a specific activity of 106 dpm/nmol. After the 10-min reaction the 32P-labeled product had a specific activity of 105 dpm; therefore, there were (105/106) nmol of 32P-labeled product, and the concentration was 10−1 nmol or 100 pmol. Hence, the enzyme transferred 10 pmol of phosphate per minute, and represents 0.01 U of enzyme activity when measured with this substrate. Put another way, this represents an enzyme specific activity of 10 pmol/min/mg of protein. This latter expression of enzyme activity is a useful measure when comparing different preparations of a kinase or, during purification of a kinase, as a measure of the extent of purification through different steps.
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Critical Parameters Although it may seem obvious, the most critical element in performing kinase assays is using the correct controls. This is especially true when performing assays on tissue samples or cell extracts. Controls should always include addition of no substrate, addition of no enzyme source, addition of heat-denatured enzyme source, and, for reactions that require activator or cofactor, addition of any activator or cofactor and addition of no activator or cofactor. Another important aspect is the concentration of ATP in the assay. The ATP concentration can vary depending on the type of experiment to be performed. For derivation of kinetic data, the concentrations of both ATP and peptide or protein substrate are critical. If kinase detection during purification is required, assay linearity is not as important as long as the right controls are present. For mapping in vitro phosphorylation sites on a kinase substrate, very often the ATP concentration is kept extremely low but the specific activity very high. When designing potential peptide substrates for individual protein kinases, there are a number of considerations to remember. For example, phosphorylated peptide will bind to the P81 phosphocellulose paper. To be able to analyze the assay with this paper, it is necessary that the peptide carry a net positive charge at low pH. In practice this is usually achieved by tagging a peptide with two or three arginine or lysine residues at the N- or C-terminus. At the pH of 75 mM orthophosphoric acid, the arginyl or lysyl side chains are positively charged and will bind to the P81 paper. It is important to determine that the activity of the kinase of interest is relatively unaffected by the addition of basic residues to its peptide substrates in this way. To help ensure this is the case, it is a good idea to place five or more residues between the hydroxy-amino acid to be phosphorylated and the basic residues required for binding to P81 paper. Net charge considerations also pertain to some extent to proteins used as substrates for protein kinases. Under the conditions described in Support Protocol 3, histones will bind very well to P81 paper, because they are highly basic, but casein, which is very acidic, binds very poorly or not at all. In these cases, the P81 paper assay is very good when histones are used to assay kinase activity, but worthless when casein is used. When crude enzyme sources, such as cell lysates, are analyzed, many other proteins are present and phosphorylation of multiple proteins in the crude mixture is possible, as is
autophosphorylation of the protein kinase. Because many proteins will be precipitated by trichloroacetic acid (TCA) or will bind to P81 paper, there can be elevated backgrounds and severe overestimates of peptide phosphorylation when Support Protocols 2 and 3 are used. When performing kinetic studies, this overestimation of phosphorylation can be very misleading, as substrates other than the peptide under study will undoubtedly be phosphorylated and with different kinetics.
Troubleshooting Any troubleshooting of kinase assay procedures should be very straightforward. If the assay does not work, verify that both [γ-32P]ATP and the peptide or protein substrate have been added at the correct concentrations. It is also very important to keep the enzyme source on ice before use and to store it appropriately between experiments. Very often when this type of experiment does not work it is because of a labile enzyme, poor storage of the enzyme, or a combination. It is important to know how long a particular enzyme is stable under the conditions used and the rate of inactivation or denaturation, if relevant. It is also essential to know if the enzyme activity is comparable between different preparations, especially for purified proteins.
Anticipated Results Depending on the requirements, information can be obtained about the specific activity of a particular enzyme preparation, kinetic parameters for the reaction catalyzed, or whether a particular kinase has been identified or substantially purified.
Time Considerations Kinase assays are quite simple to set up and perform. If the samples for assay are available, it is a matter of only ∼3 to 4 hr to go from assay set up to data interpretation when either TCA precipitation (Support Protocol 2) or adsorption to phosphocellulose paper (Support Protocol 3) is used to separate phosphorylated protein from contaminating [γ-32P]ATP. With SDS-PAGE analysis (Support Protocol 4), the assays can be set up, performed, and the samples run on a SDS-PAGE gel in ∼4 to 5 hr. Gel drying and autoradiography can take from 2 hr to a number of days depending on the level of protein phosphorylation. In general, if individual protein bands have ∼20,000 cpm of 32P associated with them, they will be visible with 1 hr of autoradiography.
Phosphorylation and Phosphatases
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Literature Cited Hanson, P.I., Kapiloff, M.S., Lou, L.L., Rosenfeld, M.G., and Shulman, H. 1989. Expression of a multifunctional Ca2+/calmodulin–dependent protein kinase and mutational analysis of its autoregulation. Neuron 3:59-70. Kameshita, I. and Fujisawa, H. 1989. A sensitive method for detection of calmodulin-dependent protein kinase II activity in sodium dodecyl sulfate polyacrylamide gel. Anal. Biochem. 183:139-143. Pearson, R.B., and Kemp, B.E. 1991. Protein kinase phosphorylation site sequences and consensus specificity motifs. Methods Enzymol. 200:62-81. Songyang, Z., Carraway, K.L. III, Eck, M.J., Harrison, S.C., Feldman, R.A., Mohammadi, M.,
Schlessinger, J., Hubbard, S.R., Smith, D.P., Eng, C., Lorenzo, M.J., Ponder, B.A.J., Mayer, B.J., and Cantley, L.C. 1995. Catalytic specificity of protein tyrosine kinases is critical for selective signalling. Nature 373:536-539. Takai, Y., Kishimoto, A., Kikkawa, U., Mori, T., and Nishizuka, Y. 1979. Unsaturated diacylglycerol as a possible messenger for the activation of calcium-activated, phospholipid-dependent protein kinase system. Biochem. Biophys. Res. Commun. 91:1218-1224.
Contributed by Nigel Carter The Salk Institute for Biological Studies La Jolla, California
Assays of Protein Kinases Using Exogenous Substrates
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Permeabilization Strategies to Study Protein Phosphorylation
UNIT 13.8
This unit deals with the use of nucleotide triphosphates to label proteins in vitro in permeabilized cells and isolated cellular fractions. These experiments generally utilize [γ-32P]ATP as an exogenously added phosphate donor, although [γ-32P]GTP can be used in specific cases. The method is very straightforward, although numerous considerations must be made before applying it to each new system. describes labeling of phosphoproteins in intact cells with 32P-labeled inorganic phosphate (32Pi). After ascertaining that the protein of interest is a phosphoprotein using the protocols detailed in that unit, the phosphorylation event itself may be studied either in cell-free systems or in a more intact cellular system. In vitro, most protein kinases are capable of phosphorylating many substrates in addition to those phosphorylated in vivo, thus making it desirable to identify and study a protein kinase–catalyzed phosphorylation reaction in a more intact system than a cell lysate or homogenate. Permeabilization of intact cells provides a powerful experimental approach for accomplishing this. Brief permeabilization of cells is possible using a variety of agents. This procedure gives access to the intracellular milieu but still allows signal transduction events to be initiated by hormone stimulation of the cells via cell-surface receptors. UNIT 13.2
Brief chemical permeabilization of cells allows the introduction of cell-impermeable reagents into the intracellular compartment in order to study particular signal-transduction processes. In this type of experiment, relatively large molecules such as the Fab′ fragments of specific antibodies may be introduced into the permeabilized cell, although the kinetics of entry may be too slow for the study of some processes. This technique is often more useful when introducing relatively small molecules into the permeabilized cell, for example non-cell-permeable activators (e.g., hydrophilic or lipophobic small molecules that cannot partition into the membrane) or inhibitors of protein kinases or phosphatases (see, e.g., Calbiochem catalog). The kinetics of entry into permeabilized cells for small molecules such as [γ-32P]ATP are fast, and the system equilibrates the cellular ATP and [γ-32P]ATP with intracellular processes within 2 to 3 min. Figure 13.8.1 shows the
40,000
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32
P (cpm)
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10,000 A 0 0
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15 20 Time (min)
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Figure 13.8.1 Equilibration of the monoester phosphates of the lipid phosphatidylinositol-4,5bisphosphate with the ATP pool after permeabilization of human platelets. The letter A marks the point of addition of [γ-32P]ATP.
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equilibration of the monoester phosphates of the lipid phosphatidylinositol-4,5-bisphosphate with the ATP pool after permeabilization of human platelets. As these monoester phosphates are known to equilibrate with the ATP pool extremely quickly, this experiment provides a good indication of the speed of equilibration of cellular ATP pools with intracellular processes after addition of [γ-32P]ATP to permeabilized cells. Procedures are outlined for performing a protein phosphorylation experiment using permeabilized cells (see Basic Protocol 1) and isolated intracellular organelles (see Basic Protocol 2). Both of these procedures result in lysates from which the protein of interest may be easily immunoprecipitated; however alternative techniques are described for preparing the final lysate from either Basic Protocol 1 or Basic Protocol 2 for electrophoretic analysis (see Alternate Protocols 1, 2, 3, and 4). A related procedure that does not involve permeabilization is outlined for direct analysis of cytosolic or membranebound kinases (see Alternate Protocol 5). Two different methods for determining the specific radioactivity of 32P-containing compounds are also included (see Support Protocols 1 and 2). CAUTION: It cannot be emphasized enough that each investigator should be fully familiarized with their local regulations regarding the safe handling and disposal of 32P and also have the appropriate equipment for dealing with the containment of 32P. See APPENDIX 2B for more information on the safe use of radioisotopes. NOTE: Milli-Q purified water or its equivalent should be used to prepare all solutions used throughout this unit. BASIC PROTOCOL 1
Permeabilization Strategies to Study Protein Phosphorylation
ANALYSIS OF PROTEIN PHOSPHORYLATION IN PERMEABILIZED CELLS When setting out to study a protein phosphorylation event via a permeabilization protocol, the general procedure is as follows. First, cells are incubated in a buffer which has been designed to mimic the composition of the intracellular compartment and which contains a permeabilizing agent. The preferred permeabilization agent of choice is streptolysin O, as this agent allows many signal-transduction systems to remain intact for a number of minutes after its addition (Stephens et al., 1994). This is certainly not true of all permeabilization agents or protocols; if another agent (e.g., saponin) is to be tried, the methods detailed below should be used as a general guideline only. Depending on the experiment to be performed, the permeabilization buffer may also contain [γ-32P]ATP and the particular reagents under study. The [γ-32P]ATP provides the “hot” phosphate, and will equilibrate with the intracellular processes utilizing ATP within 2 to 3 min (A.N. Carter, unpub. observ.). After a brief time the experimental reagents, such as inhibitors, can be introduced into the permeabilization medium and hence into the inside of the cell. At this point the cells may be stimulated with the appropriate agent in the same way as for intact cell preparations. The phosphorylation of the protein under study is initially investigated by immunoprecipitation as described in UNIT 13.2, followed by phosphopeptide mapping as described in UNIT 13.3. Materials Cell culture to be labeled Cell culture medium appropriate for cells being studied, buffered with 20 mM HEPES, pH 7.2, prewarmed to 37°C Cells under study (normally cultured cell lines or isolated primary cells in culture) Extracellular buffer (see recipe), 37°C Permeabilization buffer (see recipe), 37°C 60 U/ml streptolysin O working solution (see recipe), freshly prepared 10 mM cold ATP stock solution (see recipe) 10 mCi/ml [γ-32P]ATP (3000 Ci/mmol; e.g., DuPont NEN)
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Pharmacological agents, peptides, or Fab′ fragments to be studied Appropriate receptor agonists 2× lysis buffer for immunoprecipitation (see recipe), ice-cold Protease inhibitor stock solutions in absolute ethanol (store up to 6 months at −20°C): 100 mM phenylmethylsulfonyl fluoride (PMSF) 100 mM benzamidine 1 mg/ml pepstatin A 1 mg/ml leupeptin 1 mg/ml antipain 100 mM DTT Dry ice/ethanol bath Noncirculating 37°C water bath containing a flat shelf (preferably made of wire mesh) with space for the number of cell plates in the experiment 50-ml centrifuge tubes Tabletop centrifuge Cell scrapers Screw-cap microcentrifuge tubes Permeabilize cells For monolayer cultures 1a. Approximately 1 hr before starting the experiment, change the medium in the cell culture plates to be labeled to equivalent medium containing 20 mM HEPES. Return cells to incubator and allow the cells to equilibrate with the new medium before use. The number of cells used for any experiment of this type is totally dependent on how easy it is to detect the end product being studied. In some cases, a 60-mm dish of cells that has just reached confluency, with ∼1 × 106 cells, is adequate; in others a 100-mm dish of cells with ∼1 × 107 cells is required. It is advisable to start with ∼1 × 106 cells and see if the signal is detectable. Use of a 60-mm dish allows a smaller volume of permeabilization medium (normally of the order of 0.5 to 2 ml) to be used. Standard cell culture medium is buffered by bicarbonate ions in the medium and the CO2 in the atmosphere of the incubator. Outside the incubator this buffering system is ineffective and the pH will quickly rise if the culture is left in the normal atmosphere. The pH of HEPES buffers are minimally affected by temperature (unlike Tris buffers), hence they provide good pH control for cell culture medium. One drawback is that HEPES may be toxic to some cells above a concentration of ∼20 mM, so it is important to check possible cellular toxicity before using HEPES-buffered culture media.
2a. Place the required number of plates containing cells in the 37°C water bath and allow 10 min for the medium to reequilibrate. The cells are stable in this state for an hour or so, depending on the rate of evaporation of the culture medium.
3a. Aspirate the culture medium from each plate and quickly rinse the cells twice, each time with 10 ml of 37°C extracellular buffer. Aspirate the extracellular buffer, then add 0.5 to 2 ml permeabilization buffer to each plate. Be careful when rinsing, as some cell lines are less adherent than others. Usually, most epithelial or mesenchymal lines adhere well and will not slough off the dish unless extreme force is used. Transformed cells usually adhere poorly and can easily be rinsed off the dish if care is not taken.
4a. Add streptolysin O (from the 60 U/ml working solution) to each plate to a final concentration of 0.6 U/ml. Add cold ATP (from the 10 mM stock solution) to a final concentration of 100 µM.
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For suspension cultures 1b. Approximately 1 hr before starting the experiment, transfer cells to sterile 50-ml centrifuge tubes and centrifuge 5 min at 1000 × g, room temperature. Remove the supernatant, then rinse cells twice, each time by adding ∼40 ml of 37°C extracellular buffer, centrifuging 5 min at 1000 × g, room temperature, and removing the supernatant. The cell suspension may be split into aliquots, if desired, after the addition of the final rinse of extracellular medium. To do this, add the final (40-ml) rinse of extracellular medium, resuspend cells gently, and dispense aliquots into fresh tubes. Centrifuge the aliquots and remove the supernatants to complete the final rinse before proceeding to step 2b.
2b. Resuspend the cells in 37°C extracellular buffer at a density of 1 × 107 cells/ml. Allow the cells to equilibrate 15 to 30 min at 37°C with gentle swirling. This step may have to be modified if a cell is used that could be activated by this procedure, (e.g., platelets). If a sensitive cell is used, equilibration time may be reduced to 5 to 10 min and the agitation may be omitted.
3b. Centrifuge cells again for 5 min at 1000 × g, room temperature, and aspirate the buffer. 4b. Add streptolysin O (from the 60 U/ml working solution) to fresh 37°C permeabilization buffer to a final concentration of 0.6 U/ml. Add cold ATP (from the 10 mM stock solution) to a final concentration of 100 µM. Add this permeabilization solution containing streptolysin O and ATP to the cells and mix gently but thoroughly. Addition of cold ATP at the same time as streptolysin O allows cellular ATP levels to be maintained and will therefore preserve cell viability. One important consideration is whether the cells under study possess cell-surface receptors for ATP. The ATP acts as a full agonist for some types of purinergic receptors, and this can obviously cause confusion when analyzing signal-transduction mechanisms. Prior to performing permeabilization studies, first determine, using intact cells, whether extracellular ATP activates signaltransduction mechanisms that interfere with the system under investigation. Stephens et al. (1994) have developed a way of inactivating extracellular receptors for ATP in neutrophils prior to permeabilization. The utility of this method for other cell types is not known to date, although it may be used as a guideline if required. Readers are referred to the original paper for full discussions of this method.
Permeabilize and label cells 5. Permeabilize cells by incubating 1 to 5 min at 37°C. The time of incubation depends very much on the particular system under study and the cell type, and must be determined empirically. Permeabilization for >5 min usually reduces hormone responsiveness of cells dramatically. This is probably due to a number of processes, including the loss of cytosolic components into the extracellular pearmeabilization buffer and physical uncoupling of receptors from signal-transduction processes.
6. Add 10 mCi/ml [γ-32P]ATP to a final concentration of 50 to 500 µCi/ml. [γ-32P]ATP is usually purchased at a specific activity of ∼3000 Ci/mmol, and a concentration of 10 mCi/ml. Hence 5 µl of the stock is 50 µCi [γ-32P]ATP. The label may be used at this concentration. If the stock [γ-32P]ATP is to be diluted into another solution it is better to do this directly before use. The criterion for the amount of radiolabeled ATP to add (between 50 and 500 µCi/ml) is the detection limit. Initial experiments must be performed to see how much label must be added. Permeabilization Strategies to Study Protein Phosphorylation
The alternative to this separate labeling step is to add the [γ-32P]ATP at the same time as the streptolysin O and cold ATP in step 4a or b. The type of experiment to be performed will determine at which point the [γ-32P]ATP is added to the cells.
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7. Add the pharmacological agents, peptides, or Fab′ fragments under study and allow time for their action as required. Many pharmacological agents amenable to testing by this protocol are listed in the Calbiochem catalog. For example, a peptide containing residues 280 to 305 of the CaM kinase II α subunit can be a potent inhibitor of CaM kinase II and is non-cell-permeable. There are other peptides modeled on pseudosubstrate inhibitors of a variety of protein kinases that can be used as specific inhibitors, all of which are non-cell-permeable. Any Fab′ fragment may be tested by this protocol as long as it has an effect on enzyme activity, either positive or negative. Similarly, if an antibody stops an enzyme from reaching its site of action, it may also be tested via this procedure. The time for a particular agent to act as required should be determined experimentally as part of the initial experimental protocols. As with the [γ-32P]ATP addition, these agents may also be added at the same time as streptolysin O and cold ATP.
8. Add receptor agonists as required. The same considerations are true for receptor agonist additions as for inhibitor and [γ-32P]ATP additions to the cells. One must perform preliminary experiments to empirically determine the timings for these additions for each cell type under study. The remaining steps of this protocol are to prepare the cells for analysis by immunoprecipitation. If electrophoretic analysis is to be performed, see Alternate Protocols 1 and 2.
Prepare cell lysates for immunoprecipitation 9a. For monolayer cultures: Stop the reactions by rapidly aspirating the permeabilization buffer and adding 0.75 ml ice-cold 2× lysis buffer for immunoprecipitation to each plate. Add protease inhibitors and DTT (as stock solutions) to the following final concentrations: PMSF to 1 mM final Benzamidine to 1 mM Pepstatin A to 5 µg/ml final Leupeptin to 5 µg/ml final Antipain to 5 µg/ml final DTT to 1 mM final. Place plates on ice for 10 min, then scrape the cell debris from the bottoms of the plates into the buffer using a separate cell scraper for each plate. 9b. For suspension cultures: Stop the reactions by adding an equal volume of ice-cold 2× lysis buffer for immunoprecipitation. Add protease inhibitors and DTT to the final concentrations in step 9a, then place the tubes of cells on ice for 10 min. 10. Transfer lysates to screw-cap microcentrifuge tubes (one plate or cell aliquot/tube). Microcentrifuge lysates at maximum speed, 4°C, then transfer the supernatants to fresh screw-cap microcentrifuge tubes and freeze immediately in a dry ice/ethanol bath. Store at −70°C until needed. The lysates are now ready for immunoprecipitation. CAUTION: The permeabilization buffer in the latter steps of this protocol contains [γ-32P]ATP, and great care should be taken to remove the medium safely and dispose of it correctly as liquid radioactive waste. The lysates and cell scrapers are also very radioactive at this point. Screw-cap tubes should be used at all subsequent stages of processing the cell lysates as these provide the best containment of the 32P-containing samples. The same microcentrifuge should be used for all centrifugation steps; after use it will undoubtedly be contaminated with 32P. It can then be set aside for future use with 32P-containing samples with the appropriate safety precautions.
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INTACT CELL SAMPLE PREPARATION FOR ELECTROPHORETIC ANALYSIS OF PROTEIN PHOSPHORYLATION The preceding protocol (see Basic Protocol 1) is set up to allow immunoprecipitation of specific proteins after the permeabilization experiment. Two alternatives to this approach, described below, involve using the separating power of polyacrylamide gel electrophoresis (PAGE) to fractionate proteins, instead of immunoprecipitation. Fractionation can be either by one-dimensional SDS-PAGE (UNIT 10.1), or by two-dimensional PAGE in which isoelectric focusing is performed as the first dimension, followed by SDS-PAGE (UNIT 10.4). In each case, sample loading must be optimized so that best resolution of the protein of interest is achieved. These protocols are written for both monolayer cells and suspension cells. To perform the procedure with suspension cells, the cells must be centrifuged down as quickly as possible after reacting with the appropriate pharmacological agents/receptor agonists, then treated as described for monolayer cultures. Obviously, the timing of the reactions is less precise this way, but experiments can be done successfully and reproducibly if reaction and centrifugation times are kept exactly the same. CAUTION: Samples for SDS-PAGE or isoelectric focusing will still contain a great deal 32 P at the latter stages of the protocol. Great care should be exercised in sample preparation. In particular do not use a probe sonicator to shear DNA, as 32P-containing aerosols will be produced. In the case of SDS-PAGE, the gels should be run until the dye front passes out of the gel. This means that the SDS-PAGE running buffer will be also be contaminated with 32P. In the case of isoelectric focusing the 32P will focus out of the gel during electrophoresis; hence the running buffer will be contaminated with 32P. These solutions should be disposed of as radioactive waste as local regulations dictate. ALTERNATE PROTOCOL 1
Intact Cell Sample Preparation for SDS-PAGE Additional Materials (also see Basic Protocol 1) 2× SDS-PAGE sample buffer (see recipe), 4°C Bath sonicator Boiling water bath 1. Prepare cells and perform permeabilization and labeling (see Basic Protocol 1, steps 1 through 8). 2a. For monolayer cultures: Stop the reactions by rapidly aspirating the permeabilization buffer and adding 100 to 250 µl of 4°C 2× SDS-PAGE sample buffer to each plate. Quickly scrape the cells into the buffer using a separate cell scraper for each plate and quantitatively transfer the sample to a screw-cap microcentrifuge tube. 2b. For suspension cultures: Microcentrifuge the reaction tubes 1 min at maximum speed, room temperature. Quickly aspirate the supernatants, taking care not to disturb the cell pellets. Add 100 to 250 µl of 2× SDS-PAGE sample buffer to each tube and vortex thoroughly. When performing this step for suspension cultures it is very important to use exactly the same times for centrifugations if reproducible data are to be obtained.
3. If samples are very viscous as a result of release of DNA from cell nuclei, shear the DNA by placing the capped sample tubes in a bath sonicator filled with ice water and sonicating 5 min. Permeabilization Strategies to Study Protein Phosphorylation
4. Boil samples 3 min in a boiling water bath. To perform the analysis, carefully apply up to 50 µl of the samples to the sample wells of a precast SDS-PAGE gel. Proceed as described in UNIT 10.1.
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Intact Cell Sample Preparation for Isoelectric Focusing
ALTERNATE PROTOCOL 2
Additional Materials (also see Basic Protocol 1) Two-dimensional-PAGE lysis buffer (see recipe), 4°C Two-dimensional-PAGE sample buffer (see recipe) Bath sonicator Dry ice/ethanol bath Lyophilizer 1. Prepare cells and perform permeabilization and labeling (see Basic Protocol 1, steps 1 through 8). 2a. For monolayer cultures: Stop the reactions by rapidly aspirating the permeabilization buffer and adding 100 to 250 µl of 4°C two-dimensional-PAGE lysis buffer. Scrape the cells into the buffer using a separate cell scraper for each plate and transfer the sample to a screw-cap microcentrifuge tube. 2b. For suspension cultures: Microcentrifuge the reaction tubes 1 min at maximum speed, room temperature. Quickly aspirate the supernatant, taking care not to disturb the cell pellet. Add 100 to 250 µl of two-dimensional-PAGE lysis buffer to each tube and vortex thoroughly. When performing this step for suspension cultures it is very important to use exactly the same times for centrifugations if reproducible data are to be obtained.
3. If samples are very viscous as a result of release of DNA from cell nuclei, shear the DNA by placing the samples in a bath sonicator filled with ice water and sonicating 5 min. Freeze samples in a dry ice/ethanol bath, then lyophilize. 4. Redissolve the lyophilized samples in 50 to 100 ml of two-dimensional-PAGE sample buffer. Warm the samples to 37°C for 3 to 5 min. To perform the analysis, apply 10 to 20 µl of each sample to a prepared isoelectric focusing gel of the required pH gradient (see UNITS 10.2 & 10.4). Samples for isoelectric focusing must have a low salt concentration because a high salt concentration often causes band broadening during isolectric focusing.
ANALYSIS OF PROTEIN PHOSPHORYLATION USING ISOLATED SUBCELLULAR FRACTIONS
BASIC PROTOCOL 2
When the protein under study exists in a particular organelle and is phosphorylated in intact cells, one consideration when studying the phosphorylation event in vitro in isolated organelles is whether the protein is accessible to exogenously added kinases or phosphatases, or whether it is accessible to endogenous kinases or phosphatases. For instance, if it is a protein of the endoplasmic reticulum, is its location transmembrane or intraluminal? If transmembrane, is it normally phosphorylated on the cytosolic side or luminal side of the membrane? Permeabilization protocols can also be used with intact intracellular organelles as well as intact cells. This is obviously an important consideration when dealing with proteins found inside the organelle in question. Another consideration here is whether the organelle can be isolated in an intact condition, or whether its contents will be partially or completely lost during purification. In the case of mitochondria this does not pose a problem, but for more fragile organelles such as the Golgi apparatus this may be more problematic. The study of protein phosphorylation in isolated organelles (UNIT 4.2) can be a straightforward extension of the permeabilization protocols described above (see Basic Protocol 1).
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The isolated organelle is treated in the same way as one would treat cultured cells in suspension. However, as there is little or no requirement for the maintenance of signaltransduction mechanisms in isolated organelles, and the permeabilization procedure can be more prolonged than for intact cells. This allows the introduction of larger molecules such as purified protein kinases into the organelles under study. Materials Suspension of isolated intracellular organelles (UNIT 4.2) Intracellular buffer (see recipe), 4°C Permeabilization buffer (see recipe), 37°C 60 U/ml streptolysin O working solution (see recipe) 10 mM cold ATP stock solution (see recipe) 10 mCi/ml [γ-32P]ATP (3000 Ci/mmol; e.g., DuPont NEN) Reagents under study: e.g., kinase/phosphatase inhibitors, protein kinases, or phosphatases 2× lysis buffer for immunoprecipitation (see recipe), 4°C Screw-cap microcentrifuge tubes Benchtop ultracentrifuge (e.g., Beckman TL-100 or Airfuge) 37°C circulating water bath 1. Wash the isolated intracellular organelle preparation three times with 4°C intracellular buffer, each time by ultracentrifuging 10 min at ≥100,000 × g. Transfer the equivalent of 1 mg protein to individual screw-cap microcentrifuge tubes and keep on ice. The best way to wash the organelle preparations is to use a Beckman TL-100 benchtop ultracentrifuge or the older Beckman Airfuge. Both of these instruments will produce g-forces >100,000 × g. Because of the difficulty in centrifuging some organelles (e.g., plasma membrane preparations), it is essential to use ultracentrifugation for all organelles except nuclei, which can be recovered by microcentrifuging 30 sec at maximum speed. See UNIT 4.2 for details of organelle isolation.
2. Ultracentrifuge the organelle preparation 10 min at ≥100,000 × g, 4°C, and aspirate the supernatant. Add 200 ml 37°C permeabilization buffer, mix gently, and transfer to the 37°C circulating water bath. Work quickly at this point to minimize any proteolytic and other unwanted side reactions.
3. Add streptolysin O (from the 60 U/ml working solution) to each tube to a final concentration of 0.6 U/ml. Add cold ATP (from the 10 mM stock solution) to a final concentration of 100 µM. Exactly the same considerations apply when performing permeabilization of organelles as when performing permeabilization of intact cells (see Basic Protocol 1). However, because there is no intact signal-transduction machinery in isolated organelle preparations, the range of possible experiments is fewer, so the system is usually more robust than for intact-cell experiments.
4. Add 10 mCi/ml [γ-32P]ATP to a final concentration of 50 to 500 µCi/ml. The criterion for the amount of radiolabeled ATP to add (between 50 and 500 µCi/ml) is the detection limit. Initial experiments must be performed to see how much label must be added.
5. Add the reagents under study—e.g., kinase/phosphatase inhibitors, protein kinases, or phosphatases. Permeabilization Strategies to Study Protein Phosphorylation
6. Add 200 ml of 4°C 2× lysis buffer to each tube to lyse the organelles. Mix well and place on ice for 5 min.
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7. Microcentrifuge the samples 10 min at maximum speed, 4°C. Store lysates at −70°C. The lysates are now ready for immunoprecipitation.
ORGANELLE SAMPLE PREPARATION FOR ELECTROPHORETIC ANALYSIS OF PROTEIN PHOSPHORYLATION Exactly the same considerations apply to the electrophoretic analysis of protein phosphorylation in isolated organelles as apply to permeabilized cells (see Alternate Protocols 1 and 2). Organelle preparations must first be ultracentrifuged and the supernatant removed before adding the appropriate buffer for electrophoresis. For this, the use of a Beckman TL-100 benchtop ultracentrifuge, which can provide g-forces >300,000 × g, is recommended. Complete removal of the supernatant is especially important if isoelectric focusing for two-dimensional-PAGE (Alternate Protocol 4) is used, as the salt concentration must be kept low. CAUTION: Samples for SDS-PAGE or isoelectric focusing will contain a large amount of 32P at the latter stages of the protocol. Great care should be exercised in sample preparation. In the case of SDS-PAGE, the gels should be run until the dye front passes out of the gel. This means that the SDS-PAGE running buffer will be also be contaminated with 32P. In the case of isoelectric focusing, the 32P will focus out of the gel during electrophoresis; hence the running buffer will be contaminated with 32P. These solutions should be disposed of as radioactive waste as local regulations dictate. Organelle Sample Preparation for SDS-PAGE
ALTERNATE PROTOCOL 3
Additional Materials (also see Basic Protocol 2) 2× SDS-PAGE sample buffer (see recipe) Boiling water bath 1. Prepare organelles and perform permeabilization and labeling (see Basic Protocol 2, steps 1 through 5). 2. Ultracentrifuge samples 10 min at 100,000 × g, 4°C, and aspirate the supernatant. Add 50 µl of 2× SDS-PAGE sample buffer and mix well. 3. Boil the samples 3 min in a boiling water bath. The samples are now ready for analysis by SDS-PAGE (UNIT 10.1).
Organelle Sample Preparation for Isoelectric Focusing
ALTERNATE PROTOCOL 4
Additional Materials (also see Basic Protocol 2) Two-dimensional-PAGE lysis buffer (see recipe) Lyophilizer 1. Prepare organelles and perform permeabilization and labeling (see Basic Protocol 2, steps 1 through 5). 2. Ultracentrifuge samples 10 min at >100,000 × g, 4°C, and aspirate the supernatant. Add 200 µl of two-dimensional-PAGE lysis buffer and mix well. 3. Lyophilize the samples overnight. The samples are now ready for analysis by isoelectric focusing (UNITS 10.2 & 10.4). Samples for isoelectric focusing must have a low salt concentration because a high salt concentration often causes band broadening during the isoelectric focusing dimension.
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ALTERNATE PROTOCOL 5
DIRECT ANALYSIS OF CYTOSOLIC OR MEMBRANE-BOUND KINASES This protocol utilizes isolated subcellular fractions in straightforward kinase assays by adding exogenous, purified protein kinases to the reaction mixture containing the organelle in question. This works only if the protein under study is phosphorylated by a cytosolic or membrane-bound kinase and its substrate is on the cytosolic face of the organelle. One can then determine the phosphorylation of proteins by analyzing the reaction mixture by SDS-PAGE. It is a good idea to perform the reactions and subsequent manipulations in screw-cap microcentrifuge tubes to minimize the risk of 32P contamination. This protocol may be used to extend findings based on the use of the permeabilization strategies and assays with isolated organelles (see Basic Protocol 2). Hence, if a protein of interest is phosphorylated, it can be useful to determine the subcellular localizations of the kinase(s) responsible, as this may make it possible to infer clues as to the identity of the kinase(s). Materials Appropriate kinase assay buffer (UNIT 13.7), 4°C [γ-32P]ATP solution (see recipe in UNIT 13.7) Suspension of isolated intracellular organelles (UNIT 4.2), 4°C Purified protein kinase of interest (keep on ice) 2× SDS-PAGE sample buffer (see recipe), ice-cold Screw-cap microcentrifuge tubes 30°C and boiling water baths 1. Prepare the following reaction mixture in a screw-cap microcentrifuge tube at 4°C: 10 µl 5× kinase assay buffer 5 µl [γ-32P]ATP solution (5 µCi/5 µM ATP final) 1 to 35 µl isolated intracellular organelle suspension (i.e., volume equivalent to 1 mg protein) H2O to 50 to 100 µl. After reaction mix has been assembled, warm to 30°C for 5 min. 2. Start reaction by adding the purified protein kinase of interest. Incubate 15 min at 30°C. In the case of immunoprecipitated enzymes adsorbed onto beads, the assay mix minus the enzyme source can be made up complete first. This can then be dispensed, prewarmed, onto the immunoprecipitates. Incubation time can be adjusted as necessary depending on the extent of protein phosphorylation observed and the linearity of the reaction.
3. Stop the reactions by the addition of 100 µl of ice-cold 2× SDS-PAGE sample buffer. After thorough mixing, heat the samples 3 min in a boiling water bath. The samples are then ready for resolution by SDS-PAGE (UNIT 10.1). Samples for SDS-PAGE contain all the 32P originally in the assays. Great care should be exercised in sample preparation and loading. The gels should be run until the dye front passes out of the gel. This means that the SDS-PAGE running buffer will be contaminated with 32P, and this should be disposed of as radioactive waste as local regulations dictate.
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DETERMINATION OF SPECIFIC RADIOACTIVITY The specific radioactivity (specific activity) of any radioactive compound is defined as the number of atoms of the radioactive isotope in a molecule as a proportion of the total number of atoms of the same type. The structure of ATP is shown in Figure 13.8.2, and the form used in the study of protein kinases is [γ-32P]ATP, indicating that only the γ phosphate group has the radioactive 32P atom. There are, however, two other atoms of 31P. Hence, to attain the maximum possible specific activity, one should replace 100% of all phosphorus atoms with the 32P isotope. This is obviously not necessary in the case of kinase phosphotransfer reactions, as only the γ phosphate is transferred to a substrate; therefore in practice it is irrelevant whether the α and β phosphate groups have the 32P label. The units used by vendors of [γ-32P]ATP are curies per mole (Ci/mol), even though the curie is no longer the international unit of radioactivity.
SUPPORT PROTOCOL 1
NH2 N
N O H
O
P HO
γ
O O
P
O O
P
OH
OH
β
α
O
CH2
HO
N
N O
OH
Figure 13.8.2 Structure of ATP highlighting the α, β, and γ phosphates/phosphorus atoms.
It is important to know the specific radioactivity of the ATP in a kinase assay, as important data can be derived from this information. Consequently this can be determined as required, although one can make assumptions based on the volume of [γ-32P]ATP added to cold ATP prior to the assay. Obviously, the specific activity of [γ-32P]ATP cannot be assumed when adding [γ-32P]ATP to permeabilized cells or after labeling intact cells with 32 Pi. In these cases, one can determine the specific activity experimentally as described below. The specific radioactivity of [γ-32P]ATP can be determined by a number of methods, but the following are reliable and relatively simple. Methodologies for the extraction of ATP from either 32Pi-labeled cells or from cell lysates are detailed below (Stephens and Downes, 1990). The basis for the extraction is the same in each case, but slightly different alternative steps are required. Protein or cell debris is first precipitated with perchloric acid and removed by centrifugation. The supernatants are then neutralized with a mixture of tri-n-octylamine and 1,1,2-trichlorotrifluoroethane (Freon). The aqueous phase from this step can then be directly analyzed for ATP concentration and 32P content. ATP may be quantified using anion-exchange HPLC to separate nucleotides and a UV detector to detect nucleotides via UV absorption at 254 nm. The peak areas associated with injection of different amounts of ATP onto the column are then calculated and a standard curve constructed. The radioactivity associated with the ATP peak is easily determined by scintillation counting. Post-Translational Modification: Phosphorylation and Phosphatases
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Materials 32 P-labeled cells or cell lysates Phosphate-buffered saline (PBS; APPENDIX 2E) Standard ATP samples of known concentration in 4% perchloric acid 4% or 8% (v/v) perchloric acid, ice-cold Tri-n-octylamine (Sigma) 1,1,2-trichlorotrifluoroethane (Freon; e.g., Aldrich, Sigma) Linear gradient consisting of: Solution A: Milli-Q water filtered through 0.2-mm filter Solution B: 1.25 M NH4H2PO4, pH 3.8 (adjust pH with H3PO4; filter through 0.2-mm filter) Nucleotide standard mix: e.g., 1 mM each of AMP, ADP, ATP, GMP, GDP, and GTP Cell scrapers Polypropylene screw-cap microcentrifuge tubes or 5-ml polypropylene tubes with very tight-fitting caps Tabletop centrifuge HPLC system consisting of: Whatman Partisphere SAX HPLC column and guard column Apparatus for producing linear gradient UV detector set at 254 nm Fraction collector Whatman low-dead-volume 0.2-mm syringe filters Extract ATP from cells or cell lysates For labeled cells 1a. Aspirate the medium from the labeled cells and rinse twice with PBS. Aspirate the final wash thoroughly and place the cells on ice. Quickly add 0.5 ml of ice-cold 4% perchloric acid and mix gently to lyse the cells. Incubate 5 min on ice. 2a. Scrape the cells into the acid solution using a cell scraper and quantitatively transfer the lysate to a screw-cap microcentrifuge tube. Microcentrifuge 5 min at maximum speed, 4°C, to bring down the cell debris. 3a. Transfer the supernatant to a fresh tube and keep on ice. Freshly prepare a mixture of 1.2 parts tri-n-octylamine to 1 part Freon (on a v/v basis) and mix well. Add 0.6 ml of this mixture to the perchloric acid supernatants, cap the tubes tightly, and vortex for at least 1 min. Extract four to six standard ATP samples of known concentration in parallel with the test supernatants. It is very important that the samples be mixed extremely thoroughly at this point to ensure complete neutralization.
For labeled cell lysates 1b. In a 5-ml polypropylene tube add an equal volume of ice-cold 8% perchloric to the amount of labeled cell lysate required and place on ice. Mix thoroughly and incubate 5 min on ice. 2b. Centrifuge 5 min (or until protein is completely pelleted) at 3800 × g, 4°C.
Permeabilization Strategies to Study Protein Phosphorylation
3b. Transfer the supernatant to a fresh tube and keep on ice. Freshly prepare a mixture of 1.2 parts tri-n-octylamine to 1 part Freon (on a v/v basis) and mix well. Add 1.2 vol of this mixture to the volume of sample remaining after protein precipitation and
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vortex extremely well. Extract four to six standard ATP samples of known concentration in parallel with the test supernatants. It is very important that the samples be mixed extremely thoroughly at this point to ensure complete neutralization.
Recover neutralized aqueous sample 4. After complete mixing, centrifuge 5 min at 3800 × g, room temperature. Three layers should be evident following centrifugation. The bottom layer is excess tri-n-octylamine/Freon, the central layer is octylamine perchlorate (which is often yellow in color), and the upper layer is the neutralized aqueous sample.
5. Transfer the upper layer to a fresh tube and measure the pH using pH paper. The pH should be pH 6 to 7. If the pH of the sample is <6, repeat the treatment with tri-n-octylamine/Freon, although this is rarely necessary.
Quantitate ATP concentration 6. Equilibrate the SAX column in water for 30 min at 1 ml/min flow rate. As an alternative to this step and subsequent steps it is possible to quantitate ATP using a luminescent assay. A very easy, sensitive and convenient assay for ATP is available from Sigma and is based on using the enzyme luciferase, which produces light from luciferin and ATP. Hence, at constant luciferin/luciferase concentrations, standard curves can be constructed for the quantum light yield at different ATP concentrations. Unknown samples derived from cell lysates can thus be easily quantitated. If, however, one needs to determine the specific radioactivity of the γ phosphate of ATP (as for 32Pi-labeled cells), proceed as described in this protocol .
7. Subject the column to a blank gradient run—i.e., run the linear gradient to be used (from 100% solution A to 100% solution B) through the column but do not inject the sample. A typical simple gradient is shown in Table 13.8.1. The initial 5 min allows the contents of the injection loop to be cleared and passed onto the column, assuming use of up to a 5-ml-volume loop. It is very important that the ionic strength of the sample be <200 mM for the inorganic phosphate to bind to the column. In practice, sample loops of up to 5-ml volume can be used successfully, allowing 0.5-ml samples to be diluted 10-fold with water before analysis.
Table 13.8.1 Gradient Profile for the Separation of Nucleotides by HPLC and Approximate Retention Times for Selected Nucleotidesa,b
Time (min)
%A
0 5 70 85 90 120 125
100 100 40 40 0 0 0
%B 0 0 60 60 100 100 0
Approximate retention time (min) AMP/GMP
ADP/GDP
ATP/GTP
30/35
45/52
75/85
aA = Milli-Q water; B = 1.25 M NH H PO buffer, pH 3.8. 4 2 4 bColumn = Whatman Partisphere SAX, 25 cm × 4.6–mm i.d. The retention times for the nucleotides
are approximate, as different HPLC systems produce slight differences in gradient delivery and different HPLC columns give different separations.
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8. Filter standard mixture of nucleotides through a low-dead-volume syringe filter, then run on the SAX column. A good mix to use is AMP, ADP, ATP, GMP, GDP, and GTP, all at a concentration of 1mg/ml. A 10-ml injection will result in easily detected peaks at 254 nm using the UV detector. Once consistent separation of the required standard nucleotides is made, the samples can be analyzed.
9. Filter the four to six ATP samples of known concentration (from step 5) through a low-dead-volume syringe filter and inject them into the SAX column. Construct a standard curve. The unknown samples should fall around the center of the curve. In practice, this can be estimated according to the fact that the intracellular concentration of ATP is normally ∼1 mM, so the starting cell volume can be used to roughly estimate the ATP concentration in the cell lysates.
10. Filter the unknown samples (from step 5) through a low-dead-volume syringe filter, then run them on the SAX column and collect the ATP peak (which absorbs at 254 nm) using a fraction collector connected in series after the UV monitor. The concentration of ATP is then calculated.
11. Transfer an aliquot of the ATP peak to a scintillation vial and determine cpm. The amount of [γ-32P]ATP is then calculated from the amount of 32P associated with the peak absorbing at 254 nm. SUPPORT PROTOCOL 2
DETERMINATION OF THE SPECIFIC ACTIVITY OF THE γ-PHOSPHATE OF [32P]ATP The above technique (see Support Protocol 1) details methods for the determination of the total ATP concentration and the amount of 32P associated with that concentration. For the case of permeabilized cells where [γ-32P]ATP has been introduced specifically as the phosphate donor, the amount of radioactivity associated with ATP will give the specific activity of the γ phosphate. However, when cells are labeled with [32P]inorganic phosphate (32Pi) as in UNIT 13.2, the 32P will be distributed between all of the phosphorus atoms in ATP and hence the total radioactivity will not give a meaningful result regarding the specific activity of the γ phosphate of ATP. Also, in permeabilization studies such as the ones described in this unit, it is good to know the final specific activity of the ATP because the [γ-32P]ATP is diluted with the ATP contained within the cells. A good method for obtaining this specific activity information is the one developed by Hawkins et al. (1983), which uses cyclic AMP–dependent protein kinase (cA-PrK) to phosphorylate the substrate histone 2A under conditions where the purified [32P]ATP is the sole ATP in the assay. The 32 P-labeled ATP is completely used up during the phosphorylation of histone. Phosphorylated histone is easily separated from other 32P-containing metabolites; hence the amount of 32P associated with the γ phosphate of the labeled ATP is easily determined. The method detailed below is a slight modification of the Hawkins method and uses a peptide substrate in place of histone to allow greater specificity. This procedure is the basis for many protein kinase assays and is described in greater detail in UNIT 13.7. The details of assays for cA-PrK are dealt with in detail in that unit, along with specifics about assay buffers and substrate preparation.
Permeabilization Strategies to Study Protein Phosphorylation
Materials Cell lysates containing unknown [32P]ATP concentrations cA-PrK (purified subunit from bovine heart; Calbiochem) and 10× cA-PrK assay buffer (see recipe) 0.5 mM cyclic AMP (cAMP; Calbiochem)
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10 mg/ml kemptide (Sigma) 1 M DTT (store at −20°C) 0.5 M MgCl2 75 mM orthophosphoric acid Acetone Screw-cap microcentrifuge tubes P 81 phosphocellulose paper, precut and labeled (UNIT 13.7, Support Protocol 3) 30°C circulating water bath Additional reagents and equipment for precipitating 32P-labeled peptide on P 81 paper (UNIT 13.7, Support Protocol 3) 1. For each lysate to be assayed, prepare the following reaction mixture in a screw-cap microcentrifuge tube on ice: 25 µl [γ-32P]ATP-containing lysate 5 µl 10× cA-Prk assay buffer 1 µl 0.5 mM cAMP 5 µl 10 mg/ml kemptide 1 µl 1 M DTT 1 µl 0.5 M MgCl2 2 µl H2O. Once all assay tubes have been set up and labeled, cap tubes and keep on ice. 2. Start one reaction every 30 sec by directly adding 15 U cA-PrK to the reaction mixture and transferring the tube to a 30°C water bath. Incubate for the optimal period of time. Because the objective is to transfer all of the γ-phosphate of ATP if possible, a relatively large amount of cA-PrK is required. Initially, 15 U per tube can be used; this can be adjusted for later experiments. The optimal assay time must be determined empirically by performing a time course. Times of 60, 120, 180, and 240 min should be tried; the time chosen should be one after the incorporation of 32P into kemptide has reached a plateau.
3. Pipet 30 µl of the contents of each tube onto the surface of prepared P 81 paper squares (see UNIT 13.7, Support Protocol 3). Immediately place the squares in a beaker of 75 mM orthophosphoric acid. 4. Once all the assay tubes have been treated as in step 3, wash the P 81 paper squares five times with 75 mM orthophosphoric acid (see UNIT 13.7, Support Protocol 3). 5. Fill the beaker with acetone and wash the papers an additional 5 min (see UNIT 13.7, Support Protocol 3). Discard the acetone and allow the papers to dry at room temperature. 6. Place each square of P 81 paper in a separate 20-ml scintillation vial and determine the associated radioactivity using a scintillation counter. Once the number of dpm of 32P transferred is known, it is a simple task to calculate the specific activity of the ATP input into the experiment. For example—if the amount of total ATP has been determined to be 10 µmol, the radioactivity associated with ATP is 220,000 dpm, the amount of 32P transferred to kemptide is 110,000 dpm, and the amount of 32P remaining is 110,000 dpm—then there are 125,000 dpm of 32P in the γ-phosphate of ATP as there are ∼2.2 × 106 dpm per µCi. Hence, 110,000 dpm is equivalent to 0.05 µCi, so the specific activity of the γ-phosphate of ATP is 0.5 mCi/mmol.
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REAGENTS AND SOLUTIONS Use Milli-Q purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
cA-PrK assay buffer, 10× 500 mM Tris⋅Cl, pH 7.5 at 30°C (APPENDIX 2E) 50 mM MgCl2 Store up to 6 months at −20°C Cold ATP stock solution, 10 mM Mix equal volumes of 20 mM MgCl2 and 20 mM disodium ATP. Slowly raise the pH of the solution to 7.4 with 1 M NaOH using constant stirring. Filter sterilize and store in small aliquots up to 1 year at −20°C. Extracellular buffer 110 mM NaCl 10 mM KCl 1 mM MgCl2 1.5 mM CaCl2 30 mM HEPES 10 mM glucose Filter sterilize and store up to 1 month at 4°C Intracellular buffer 25 mM NaHCO3 120 mM KCl 1 mM KH2PO4 10 mM MgCl2 20 mM HEPES 10 mM glucose Filter sterilize and store up to 1 month at 4°C Lysis buffer for immunoprecipitation, 2× 100 mM HEPES 200 mM NaCl 40 mM EDTA 4 mM EGTA 100 mM NaF 20 mM β-glycerophosphate 2 mM Na3VO4 2% (v/v) NP-40 or Triton X-100 Filter sterilize and store up to 6 months at 4°C This buffer has a 10× concentration of EDTA to chelate the excess magnesium used in kinase reactions.
PBS/BSA Dissolve 10 mg bovine serum albumin (BSA) in 100 ml PBS (APPENDIX 2E). Filter sterilize using a low-protein-binding membrane (e.g., 0.2-µm Millipore GV filters) and store up to several months at 4°C.
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Permeabilization buffer 120 mM KCl 25 mM NaHCO3 5 mM HEPES 10 mM MgCl2 1 mM KH2PO4 1 mM EGTA 300 mM CaCl2 Filter sterilize and store up to several months at 4°C This buffer has a free ionized calcium concentration of ∼100 nM.
SDS-PAGE sample buffer, 2× 4 ml 0.125 M Tris⋅Cl, pH 6.8 (APPENDIX 2E) 4 ml 10% (w/v) SDS 1 ml glycerol 0.4 ml 2-mercaptoethanol 0.6 ml H2O 20 mg bromphenol blue Filter sterilize and store up to 6 months at −20°C The pH of the Tris buffer is measured at room temperature.
Streptolysin O solution 600 U/ml stock solution: Make up a stock solution of 600 U/ml streptolysin O in PBS (APPENDIX 2E) and snap-freeze in liquid nitrogen. Store at −70°C in small aliquots up to several months. 60 U/ml working solution: Immediately before the experiment, dilute the 600 U/ml stock solution to 60 U/ml with PBS/BSA (see recipe). Keep on ice until ready to use. Two-dimensional-PAGE lysis buffer 20 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 0.3% (w/v) SDS 1% (v/v) 2-mercaptoethanol Filter sterilize and store up to 6 months at −20°C Two-dimensional-PAGE sample buffer 6 g urea 4 ml NP-40 0.5 ml 40% ampholytes of the required pH range (UNIT 10.2) 0.154 g DTT 1.12 ml H20 Filter sterilize and store in small aliquots up to 1 month at –70°C NOTE: Use the highest-purity urea available so that the concentrations of the breakdown products of urea—carbamic acid and ammonia—are minimized. This is important because proteins can be carbamylated in solution by these breakdown products, thereby altering the native protein’s net charge and hence its isoelectric point.
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COMMENTARY Background Information
Permeabilization Strategies to Study Protein Phosphorylation
The permeabilization of cells has become a popular method for the study of the regulation of signal-transduction mechanisms. In practice, similar methodologies can be used to study the different types of receptor-regulated intracellular communication mechanisms such as those regulated by heterotrimeric G-protein–coupled receptors, receptor tyrosine kinases, and cytokine receptors (Hawkins et al., 1983; Stephens and Downes, 1990; Alexander et al., 1992; Stutchfield and Cockcroft, 1993; Slowiejko et al., 1994; Stephens et al., 1994; Cunningham et al., 1995; Martys et al., 1995; Taylor et al., 1995). Although permeabilization protocols are very simple in theory, they are very much more problematic in practice. This is because different cells have different permeabilization properties; hence each time permeabilization protocols are applied to a new cell type, it is neccesary to reexamine the experimental protocol in detail before reproducible data may be obtained. Similarly, different permeabilization-inducing agents are available, but they are not easily interchangeable, nor do they show predictable properties in the maintainence of signal-transduction processes in different cell types. The current agent of choice is streptolysin O, a bacterial toxin from the organism Streptococcus pyogenes. Streptolysin O has been used to study G-protein–coupled receptors and receptor tyrosine kinases; for these systems it appears to work quite reproducibly (Hawkins et al., 1983; Stephens and Downes, 1990; Alexander et al., 1992; Stutchfield and Cockcroft, 1993; Slowiejko et al., 1994; Stephens et al., 1994; Cunningham et al., 1995; Martys et al., 1995; Taylor et al., 1995). A series of experiments utilizing cell permeabilization will not produce quick results. It will, however, produce data not normally obtainable using other experimental approaches. A number of preliminary experiments must be performed to determine the optimal conditions for the permeabilization of intact cells, as well as for the time of addition for various reagents. Aspects to be considered include the following. 1. The kinetics of phosphorylation of the protein under study in intact cells must be determined. 2. The time required for cold ATP/[γ32P]ATP equilibration with intracellular processes—e.g., phosphatidic acid formation and phosphatidylinositol-4-phosphate or phospha-
tidylinositol-4,5-bisphosphate synthesis— must be determined. The timing of the addition of [γ-32P]ATP to the permeabilization medium must be optimized—e.g., should the [γ-32P]ATP be added at the same time as the permeabilization agent, or some time later? 3. If possible, the integrity and time course of receptor activation of a known signaltransduction mechanism should be determined—e.g., determine whether MAP kinase is activated under the conditions of permeabilization indicated in the experiments performed under point 2, above, as well as the optimal time for activation and how long the response is detectable. 4. The timing for addition of receptor agonists should be tested—e.g., can a receptor agonist be added at the same time as the permeabilization agent, at the same time as the [γ-32P]ATP, or some time after these reagents? 5. It should be determined whether the protein under study is phosphorylated under the same conditions as the reporter activity observed under point 3, above, in the permeabilized cells. 6. The timing of addition of inhibitors/activators or other reagents should be determined, as well as their concentration for optimal function in the permeabilized cells. Once these issues have been resolved, experiments can easily be optimized and the phosphorylation event under study examined in detail.
Critical Parameters and Troubleshooting A number of problems can be experienced when using permeabilization protocols. Highquality streptolysin O must be used, and each preparation of streptolysin O should be tested according to the supplier’s instructions before use. The timing for addition of all reagents to the cells must be strictly reproduced within experiments and between experiments. If large variability is noted between experiments and the various timings have been strictly observed, then the problem is almost always due to the streptolysin O preparation used. The solution should then be checked per the supplier’s instructions. Another problem that may be encountered is hydrolysis of [γ-32P]ATP by endogenous ATPases. This should always be assayed in any experiment by determining the production of
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[32P]inorganic phosphate during the time course of the experiment. If the ATP concentration is depleted by >10% during the course of the experiment, general ATPase inhibitors can be included in the permeabilization buffer. One such inhibitor is ouabain; however the potential exists for the ATPase inhibitor to inhibit whatever process is being studied; hence more experiments are required to determine the result empirically for each inhibitor to be used.
Anticipated Results Once the optimal conditions for permeabilization, ATP addition, agonist addition, and inhibitor addition have been determined, it is relatively straightforward to study a novel phosphorylation event. Initially, one can detect the phosphorylation of the protein under study as a result of receptor activation. If pharmacological tools are available—e.g., inhibitors/activators of specific protein kinases—it is possible to make a provisional identification of the kinase responsible for the phosphorylation event being examined. It is also possible to determine the proximity of the phosphorylation event to receptor activation.
Time Considerations The entire permeabilization procedure can be completed in ∼2 hr. After this time, the samples generated can be prepared for analysis by immunoprecipitation or directly analyzed by SDS-PAGE. Immunoprecipitation times are dependent on the antibodies being used, but high-affinity interactions should only require 1 to 2 hr of incubation at 4°C. Harvesting the immunoprecipitates with protein A–Sepharose will take 1 hr at 4°C, and the washing procedure then takes 30 min before the samples can be prepared for SDS-PAGE. One-dimensional SDS-PAGE gels can generally be run in ∼3 hr for a regular-sized gel or 1 hr for a minigel. The gels can then be stained to the required level in 1 to 2 hr before placing on a gel dryer for 1 to 2 hr. Once the gel has been stained and dried, it can be analyzed by either standard autoradiography or using a Phosphorimager. The exposure times required are dependent on the radioactivity contained in the protein bands. For regular autoradiography using an intensifying screen, it is possible to visualize a band containing a few hundred cpm with an overnight exposure at −70°C. A Phosphor screen will give similar data with ∼4 hr exposure.
For two-dimensional-PAGE analysis, the samples can be placed in the lyophilizer overnight prior to electrophoresis. The two-dimensional-PAGE procedure will take one day to complete before the gel can be dried and analyzed by autoradiography or with a Phosphorimager.
Literature Cited Alexander, D.R., Brown, M.H., Tutt, A.L., Crumpton, M.J., and Shivnan, E. 1992. CD3 and CD2 antigen–mediated CD3 γ-chain phosphorylation in permeabilized human T cells: Regulation by cytosolic phosphatases. Biochem. J. 288:69-77. Cunningham, E., Thomas, G.M., Ball, A., Hiles, I., and Cockcroft, S. 1995. Phosphatidylinositol transfer protein dictates the rate of inositol triphosphate production by promoting the synthesis of PIP2. Curr. Biol. 5:775-783. Hawkins, P.T., Michell, R.H., and Kirk, C.J. 1983. A simple assay method for determination of the specific radioactivity of the γ-phosphate group of 32P-labeled ATP. Biochem. J. 210:717-720. Martys, J.L., Shevell, T., and McGraw, T.E. 1995. Studies of transferrin recycling reconstituted in streptolysin O–permeabilized Chinese hamster ovary cells. J. Biol. Chem. 270(43):2597625984. Slowiejko, D.M., Levey, A.I., and Fisher, S.K. 1994. Sequestration of muscarinic cholinergic receptors in permeabilized neuroblastoma cells. J. Neurochem. 62:1795-1803. Stephens, L.R. and Downes, C.P. 1990. Product-precursor relationships amongst inositol polyphosphates: Incorporation of [32P]Pi into myo-inositol 1,3,4,6-tetrakisphosphate, myo-inositol 1,3,4,5-tetrakisphosphate, myo-inositol 3,4,5,6tetrakisphosphate and myo-inositol 1,3,4,5,6pentakisphosphate in intact avian erythrocytes. Biochem. J. 265:435-452. Stephens, L.R., Jackson, T., and Hawkins, P.T. 1994. Synthesis of phosphatidylinositol-3,4,5trisphosphate in permeabilized neutrophils regulated by receptors and G-proteins. J. Biol. Chem. 268:17162-17172. Stutchfield, J. and Cockcroft, S. 1993. Correlation between secretion and phospholipase D activation in differentiated HL60 cells. Biochem. J. 293:649-655. Taylor, J.A., Karas, J.L., Ram, M.K., Green, O.M., and Seidel-Dugan, C. 1995. Activation of the high-affinity immunoglobulin E receptor Fc ε RI in RBL-2H3 cells is inhibited by Syk SH2 domains. Mol. Cell. Biol. 15:4149-4157.
Contributed by A. Nigel Carter The Salk Institute for Biological Studies La Jolla, California
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Phosphopeptide Mapping and Identification of Phosphorylation Sites
UNIT 13.9
Many proteins in the cell are modified by phosphorylation. Protein phosphorylation can affect catalytic activity, localization of a protein in the cell, protein stability, and the ability of a protein to dimerize or form a stable complex with other molecules. There are several techniques available to find out whether or not a protein is modified by phosphorylation. To understand exactly why a particular protein becomes phosphorylated, it may be necessary to identify precisely which amino acid residues are phosphorylated. These residues can then be changed by site-directed mutagenesis, and the mutant protein can be examined for changes in activity, intracellular localization, and association with other proteins in the cell. Studies geared towards understanding the phosphorylation of a particular protein usually start with labeling of the protein in intact cells followed by phosphoamino acid analysis (UNITS 13.2 & 13.3). Proteolytic digestion of a 32P-labeled protein, followed by separation of the digestion products in two dimensions on a TLC plate, will give rise to a phosphopeptide map (see Basic Protocol 1). Phosphopeptides present on the TLC plate are visualized by autoradiography. These maps give information about the number of phosphate-containing peptides in the digest, and this is related to the number of phosphorylation sites present in the protein. Phosphopeptide maps can also be used to find out whether the sites of phosphorylation on a protein change upon treatment of cells with certain agents. Treatment of cells could lead to a reduction in the labeling of certain peptides and an increase in the labeling of other peptides present on the peptide map. This suggests that treatment results in a loss of phosphorylation at certain sites and an increase in phosphorylation at other sites. The identification of the sites of phosphorylation requires further analysis of the phosphopeptides present on these maps (see Support Protocol 1 and Basic Protocols 2 and 3). If 10 pmol of phosphorylated material can be generated, phosphopeptides can be purified by HPLC and identified directly by mass spectrometry or peptide microsequencing (see Support Protocol 2). TRYPTIC PHOSPHOPEPTIDE MAPPING OF PROTEINS ISOLATED FROM SDS-POLYACRYLAMIDE GELS
BASIC PROTOCOL 1
32
P-labeled proteins are resolved by SDS-PAGE and visualized following autoradiography. Protein bands are cut out of the dried gel and the protein of interest is isolated by extraction from the gel and TCA precipitation in the presence of carrier protein. The precipitated protein is oxidized in performic acid and digested with trypsin. The bicarbonate buffer is evaporated by several rounds of lyophilization, the tryptic peptide mix is spotted on a TLC plate, and peptides are resolved by electrophoresis and chromatography in two dimensions and visualized by autoradiography. Materials Samples containing 32P-labeled proteins of interest (UNIT 13.2) Fluorescent ink or paint (can be obtained from most arts and crafts supply stores) 50 mM ammonium bicarbonate, pH 7.3 to 7.6 (when freshly prepared the buffer has a pH of ∼7.5) , and pH 8.0 (the pH drifts overnight to ∼8.0, ideal for digestion with trypsin or chymotrypsin as in step 18) 2-Mercaptoethanol (2-ME) 20% (w/v) SDS (APPENDIX 2E) 50 mM ammonium bicarbonate, pH 7.3 to 7.6, containing 0.1% (w/v) SDS and 1.0% (v/v) 2-ME
Contributed by Jill Meisenhelder, Tony Hunter, and Peter van der Geer Current Protocols in Protein Science (1999) 13.9.1-13.9.27 Copyright © 1999 by John Wiley & Sons, Inc.
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2 mg/ml carrier protein (RNase A, BSA, or immunoglobulins) in deionized water (store in aliquots at −20°C or −70°C) 100% (w/v) trichloroacetic acid (TCA) 96% ethanol, ice cold 30% (w/v) hydrogen peroxide 98% (w/v) formic acid 1 mg/ml TPCK-treated trypsin (e.g., Worthington) in deionized water or 0.1 mM HCl (store in aliquots at −70°C or under liquid nitrogen) Electrophoresis buffers (see recipe): pH 1.9, 3.5, 4.72, 6.5, and 8.9 Green marker dye (see recipe) Chromatography buffer (see recipe; also see Critical Parameters for discussion of buffer selection) Single-edge razor blades or surgical blades Scintillation counter appropriate for Cerenkov counting 1.7-ml screw-cap microcentrifuge tubes (Sarstedt) Disposable tissue-grinder pestles (Kontes) Platform rocker Tabletop centrifuge with swinging-bucket rotor 1.5 ml microcentrifuge tube, pretested for suitability (see Critical Parameters) Glass-backed TLC plates (20 × 20 cm, 100 µm cellulose; EM Science) Low-volume adjustable pipet with long disposable tips made of flexible plastic, e.g., gel-loading tips Air line fitted with filter to trap aerosols and particulate matter HTLE 7000 electrophoresis apparatus (CBS Scientific) Polyethylene sheeting (35 × 25 cm; CBS Scientific) Electrophoresis wicks (20 × 28 cm sheet of Whatman 3MM paper folded lengthwise to give double thickness sheets of 20 × 14 cm) Chromatography tank (CBS Scientific) Fan for drying TLC plates 65°C drying oven Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11) Isolate 32P-labeled protein by SDS-PAGE and recover from the gel 1. Resolve the samples containing the 32P-labeled protein of interest by SDS-polyacrylamide gel electrophoresis (SDS-PAGE; UNIT 10.1). 2. Following electrophoresis, dry the gel, mark it around the edges with fluorescent ink, and expose to X-ray film (autoradiography; UNIT 10.11). 3. Localize the protein of interest in the gel by aligning fluorescent markers around the gel precisely with their images on the film. Staple the film and gel together and place this sandwich film-side-down on a light box. Mark the position of 32P-labeled protein bands on the back of the gel using a soft pencil or ballpoint pen (do not use a felt-tip marker).
Phosphopeptide Mapping and Identification of Phosphorylation Sites
4. Separate the gel from the film and cut out the protein bands from the individual lanes of the gel using a single-edge razor blade. Strip the paper backing from the gel slices and remove residual bits of paper by scraping gently with a razor blade. Try not to shave pieces from the gel because this will reduce recovery of the protein of interest. Place each gel slice in a 1.7-ml screw-cap tube and determine the amount of radioactivity by Cerenkov counting in a scintillation counter.
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5. Rehydrate each dry gel slice in 500 µl of 50 mM ammonium bicarbonate, pH 7.3 to 7.6, for 5 min at room temperature. Mash the gel slice using a Kontes tissue-grinder pestle until no bits are seen when the tube is held up to the light. Add 500 µl of 50 mM ammonium bicarbonate, pH 7.3 to 7.6, 10 µl 2-ME, and 10 µl of 20% SDS. Boil 2 to 3 min. 6. Extract the protein from the gel by incubation on a rocker for at least 4 hr at room temperature or for at least 90 min at 37°C. For convenience, extractions can be done overnight.
7. Collect the gel slurry at the bottom of the tube by centrifuging 5 min at 500 × g, room temperature, in a tabletop centrifuge with a swinging-bucket rotor. Transfer the supernatant to a new 1.5-ml microcentrifuge tube. IMPORTANT NOTE: The brand of tube is important. From this point on use a brand of tubes that does not produce unwanted side reactions or retain too many cpm at the final transfer step (see Critical Parameters). The authors use microcentrifuge tubes from Myriad Industries.
8. Before starting the second elution, measure the volume of the first eluate and calculate the volume to be used for the second elution so that the volume of the combined eluates will measure ∼1300 µl. For the second elution, resuspend the gel pellet in the calculated appropriate volume of 50 mM ammonium bicarbonate containing 0.1% SDS and 1.0% 2-ME. Vortex, then boil 2 to 3 min and extract again by incubation on a rocker for at least 90 min. 9. Separate the gel from the eluate again by centrifugation as in step 7 and transfer the supernatant to the tube containing the first eluate. 10. To clear the combined eluate of gel slurry that has been inadvertently transferred, microcentrifuge 5 to 10 min at full speed, then transfer the supernatant to a new microcentrifuge tube. Before discarding the gel bits, monitor by Cerenkov counting to ensure that 60% to 90% of the 32P-labeled protein has been extracted. It is important to remove all gel fragments, and it may be worthwhile to repeat the last centrifugation step one more time.
11. Cool the eluates by placing them on ice. Add 20 µg carrier protein (10 µl of a 2 mg/ml stock), mix well, add 250 µl ice-cold 100% TCA, mix well, and incubate for 1 hr on ice. 12. Collect the protein precipitate by microcentrifuging 5 to 10 min at full speed, 4°C. Decant the supernatant, then microcentrifuge again for 3 min at 4°C and aspirate the last traces of TCA. 13. Wash the TCA precipitate by adding 500 µl ice-cold 96% ethanol, invert the tube a few times, and microcentrifuge 5 min at full speed, 4°C. Decant the bulk of the supernatant, microcentrifuge again for 3 min at 4°C, and aspirate the residual liquid. Air dry the protein pellet (do not lyophilize). 14. Monitor the precipitate by Cerenkov counting to make sure that the majority of the 32 P-labeled protein has been recovered. There should be as many or slightly more cpm in the sample at this point as compared to the eluate (see step 10), since the liquid of the eluate will have quenched the counting somewhat. Post-Translational Modification: Phosphorylation and Phosphatases
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Incubate with performic acid to achieve oxidation of the 32P-labeled protein 15. Generate performic acid by incubating 9 parts formic acid with 1 part 30% hydrogen peroxide for 60 min at room temperature. Cool the performic acid by placing it on ice. 16. Resuspend the TCA-precipitated protein pellet in 50 µl of the ice-cold performic acid and incubate 60 min on ice. 17. Add 400 µl deionized water, mix, and freeze on dry ice. Evaporate the performic acid under vacuum in a SpeedVac evaporator. It is extremely important to dilute and then freeze the sample before evaporating it, as otherwise the elevated temperature of the SpeedVac evaporator may cause acid hydrolysis of the sample. A sample (5% to 10% of the digest, at least 200 cpm) can be taken at this point for phosphoamino acid analysis. Lyophilize and proceed at described in UNIT 13.3.
Perform proteolytic digestion with trypsin 18. Resuspend the protein pellet in 50 µl of 50 mM ammonium bicarbonate, pH 8.0, and add 10 µg trypsin (10 µl of a 1 mg/ml stock). Digest for 3 to 4 hr or overnight at 37°C. 19. Add a second 10 µg aliquot of trypsin and digest again for 3 to 4 hr or overnight at 37°C. 20. Add 400 µl deionized water, and lyophilize in a SpeedVac evaporator. Resuspend the pellet in 400 µl deionized water and lyophilize again. Repeat these steps until at least four lyophilizations have been achieved. At this stage there should be no visible pellet.
21. Resuspend the tryptic digest in 400 µl electrophoresis buffer or deionized water. The authors use pH 1.9 buffer or pH 4.72 buffer for samples that will be analyzed by electrophoresis at pH 1.9 and pH 4.72, respectively, and deionized water for samples that will be analyzed by electrophoresis at pH 8.9.
22. Clear the peptide mix of all particulate matter by microcentrifuging 5 to 10 min at full speed, transfer the supernatant to a new microcentrifuge tube, and lyophilize. Measure the amount of 32P radioactivity in the final sample by Cerenkov counting. It is very important that there be no particulate matter in this final supernatant, and it may be worthwhile to repeat this centrifugation step one more time.
23. Resuspend the digest in at least 5 µl of pH 1.9 electrophoresis buffer, pH 4.72 electrophoresis buffer, or deionized water, and collect the sample at the bottom of the tube by microcentrifuging 2 to 5 min at full speed. Perform first-dimension electrophoresis on a TLC plate 24. Mark the sample and dye origins on the cellulose side of a glass-backed TLC plate with a small cross using a blunt extra-soft pencil, making sure not to perturb the cellulose layer (or, alternatively, mark on the reverse side with a permanent marker). This is most easily done by placing the plate on top of a “life-sized” marking template on a light box (Fig. 13.9.1).
25. Spot each sample onto the corresponding origin using an adjustable low-volume pipet fitted with a long flexible tip (round, gel-loading tips work well). Apply 0.2- to 0.5-µl drops, and dry between applications using an air line fitted with a filter to trap aerosols and particulate matter, and a 1-ml syringe or a Pasteur pipet to focus the air flow. Phosphopeptide Mapping and Identification of Phosphorylation Sites
Avoid touching the plate with the air nozzle or the pipet tip, since gouges on the cellulose may affect electrophoresis or chromatography. Ideally, at least 1000 cpm should be loaded onto the TLC plate. A brown ring around the circumference of the spot is normal.
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26. Spot 0.5 µl of green marker on the dye origin at the top of the plate (Figs. 13.9.1 and 13.9.2). This marker dye is green, but separates into its blue (xylene cyanol FF) and yellow (ε-dinitrophenyllysine) components during electrophoresis (Fig. 13.9.2D).
27. Prepare the HTLE 7000 apparatus as described below, referring to Figure 13.9.3. a. Fill the buffer tanks so that the level of the electrophoresis buffer is ∼5 cm deep. Place a sheet of polyethylene on the Teflon cover that protects and insulates the base and tuck the ends down between the base and the buffer tanks to hold the sheet in place. b. Wet the electrophoresis wicks in electrophoresis buffer and slide them into the slots of the buffer tank with the folded edge up. Fold the ends of the wicks over the polyethylene sheet on the base. Place the second polyethylene sheet over the base, the electrophoresis wicks, and part of the buffer tank.
A
B dye origin
2 cm
dye origin
15 cm
15 cm
sample origin
sample origin 3 cm
6 cm
2 cm
3 cm 10 cm
14 cm
C
10 cm
D
7.5 cm
17.5 cm
4.5 cm
4.5 cm
15.5 cm
15.5 cm
5.0 cm
5.0 cm 12.5 cm
12.5 cm
Figure 13.9.1 Sample and dye origins and blotter dimensions for separation of peptides at different pH values. (A) Location of the sample and dye origins for electrophoresis at pH 1.9 and pH 4.72 and (B) at pH 8.9. To mark a TLC plate, the plate is placed on top of a life-sized template. This is then placed on top of a light box and the origins are marked on the cellulose side using a very blunt extra-soft pencil. Dimensions of the blotter and the location of two holes that fit over the sample and dye origins are shown (C) for electrophoresis at pH 1.9 and 4.72 and (D) for pH 8.9.
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c. Place the second Teflon sheet and the neoprene pad on top, close the apparatus, and secure the lid with the two pins. Inflate the airbag by turning up the air pressure to 10 psi to squeeze out excess buffer from the electrophoresis wicks. Keep the air pressure on until ready to start the first run. d. After the samples have been loaded on the TLC plates (steps 25 and 26) and one is ready to start the electrophoresis, shut off the air pressure and open the apparatus. Remove the neoprene pad and the top Teflon and polyethylene sheets, and fold back the electrophoresis wicks. Wipe both polyethylene sheets dry with tissue paper.
A
B
C
D
+
Phosphopeptide Mapping and Identification of Phosphorylation Sites
Figure 13.9.2 Separation of peptides by electrophoresis. (A) The sample and dye are spotted on their respective origin at the bottom and the top of the TLC plate as described in the text. (B) The blotter is soaked briefly in the electrophoresis buffer, and excess liquid is removed by blotting briefly on a piece of 3MM filter paper. (C) The TLC plate is wetted by placing the wetted blotter on top, with the sample and marker origins in the centers of the two holes. The blotter is pressed onto the TLC plate around the sample and marker origins to ensure uniform flow of electrophoresis buffer from the blotter towards the sample and marker origins. This will result in concentration of the sample and marker dye on their respective origins, and will improve resolution. The rest of the blotter is pressed with a flat hand onto the TLC plate, the blotter is removed, and the plate is examined; it should be dull gray with no shiny puddles of buffer. Excess buffer should be allowed to evaporate or be blotted very carefully with tissue paper. The plate is placed on the apparatus and the electrophoresis run for 20 to 30 min at 1 kV. This results in separation of the peptides in the first dimension (D, peptides shown in black). The position of the anode and cathode are indicated in panel D.
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e. Wet the TLC plate containing a sample as described in Figure 13.9.2 and place the plate on the polyethelene sheet covering the base. Fold the wicks over the plate so they cover ∼1 cm of the plate at each end and carefully reassemble the apparatus as described above. Avoid lateral movement of the polyethylene sheet when it is in contact with the TLC plate. Secure the lid with the pins, inflate the airbag to 10 psi, turn on the cooling water flow, and start the electrophoresis. At this point the authors concentrate the sample by wetting the TLC plates with electrophoresis buffer using a blotter with holes around the origin (Figs. 13.9.1 and 13.9.2). The blotter is made from two layers of Whatman 3MM paper, stitched together around the edges. The 1.50-cm holes that surround the origins (Fig. 13.9.1) are cut with a sharp cork borer. These blotters can be reused many times; it is best to keep a separate blotter for each buffer. The buffer has to move with similar speed from the entire circumference towards the origin. The sample will inevitably streak if the buffer takes a long time to wet the spot, or moves unevenly through the spot.
f. Perform electrophoresis for 20 to 30 min at 1.0 kV. Electrophoresis results in separation of the peptides in one dimension (Fig. 13.9.2).
28. After completing the run, disassemble the apparatus and air dry the plate with the help of a fan for at least 30 min after electrophoresis is completed. Do not dry in an oven as this will bake the peptides onto the cellulose, thereby interfering with the separation in the chromatography dimension.
Perform second-dimension separation by chromatography 29. Apply a drop (∼0.5 µl) of green marker dye in the left- or right-hand margin of the plate at the same level as the sample origin, avoiding the area that has been compressed by contact with the electrophoresis wick (Fig. 13.9.4). Place the dried plates in an almost upright position in the chromatography tanks with the appropriate chromatography buffer and replace the lid. Allow chromatography to proceed until the buffer runs to within 1 to 2 cm of the top of the plate.
top plate and
attached airbag
neoprene pad 6
5
4 3
base 1
.
2 1 2
3
Teflon cover for bottom plate4 bottom polyethylene sheet 5 6 TLC plate
. electrophoresis wick top polyethylene sheet Teflon sheet
Figure 13.9.3 Preparation of the HTLE 7000 electrophoresis system.
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A
B
C
D
Figure 13.9.4 Separation of peptides in the second dimension by chromatography. After electrophoresis, air-dry the plate. A fan may be used to facilitate this. Add a small amount of green marker dye in the left hand (A) or right hand margin at the same level as the sample origin. Place the plate(s) almost upright in a chromatography tank, replace the lid, and run the chromatography until the buffer front reaches to within 1 to 2 cm from the top of the TLC plate (B and C). This results in separation of the peptides in the second dimension (D; peptides shown in black). Open the tank, take out all plates, let the plates air dry, apply fluorescent ink at the margins of the plate, and expose to X-ray film.
IMPORTANT NOTE: Do not disturb or open a tank while chromatography is in progress. See Figure 13.9.4 for illustrations of these procedures.For information on selecting an appropriate chromatography buffer, see Critical Parameters.
30. Remove all TLC plates from the chromatography tank at the time the tank is opened. Allow the plates to dry for 1 hr in a fume hood or for 15 min in a 65°C oven. Do not use the oven if peptides are to be extracted from these plates for further analysis.
Phosphopeptide Mapping and Identification of Phosphorylation Sites
31. Mark the dried plates with fluorescent ink around the edge; these reference marks can be used later to align the autoradiogram with the TLC plate. Expose the plates to X-ray film in the presence of an intensifier screen for autoradiography (UNIT 10.11), or to a phosphorimager screen. If needed, recover peptides for further analysis (see Support Protocol) X-ray film may be presensitized for increased sensitivity (see UNIT 10.11).
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PROTEOLYTIC DIGESTION OF IMMOBILIZED PROTEINS Isolation of proteins from polyacrylamide gels is a lengthy and laborious procedure (see Basic Protocol 1). In addition, recoveries can be poor. Saving time can be important if one is working with the limited amounts of 32P radioactivity present in proteins labeled in intact cells. As an alternative, proteins can be transferred to nitrocellulose or PVDF membranes, followed by digestion of the immobilized protein. The peptides are oxidized after digestion and elution from the membrane. Obviously, this approach is not a good choice for proteins that transfer with poor efficiency.
ALTERNATE PROTOCOL
Additional Materials (also see Basic Protocol 1) Methanol 0.5% (w/v) PVP-360 in 100 mM acetic acid (see recipe) 50 mM ammonium bicarbonate, pH 8.0 PVDF membrane (Immobilon P, Millipore) or nitrocellulose membrane (UNIT 10.10) Saran Wrap or Mylar Additional reagents and equipment for wet or semidry protein transfer (UNIT 10.10) 1. Resolve the 32P-labeled samples by SDS-polyacrylamide gel electrophoresis (UNIT 10.1) and transfer the proteins to a PVDF or nitrocellulose membrane using a standard wet or semidry protein-transfer protocol (UNIT 10.10). 2. Air dry the membrane and wrap it in Saran Wrap or Mylar to prevent the membrane from sticking to the film, mark with fluorescent ink (see Basic Protocol 1, step 2), and expose the blot to X-ray film (autoradiography; UNIT 10.11). 3. Align the film with the membrane using the fluorescent markers and their images on the film to identify the exact position of the protein of interest on the membrane (see Basic Protocol 1, step 3). 4. Cut out the strips of membrane containing the protein of interest with a single-edge razor blade, then cut this strip into several smaller pieces. Place all pieces of membrane containing a particular phosphate-labeled protein in a single microcentrifuge tube. Quantify the amount of radioactivity present on these strips of membrane by Cerenkov counting. 5. Rewet the membrane by adding 500 µl methanol, wash the membrane strips several times with deionized water, and incubate 30 min at 37°C with 0.5% PVP-360 in 100 mM acetic acid. 6. Wash the membrane at least five times, each time with 1 ml deionized water, then two times, each time with 1 ml of 50 mM ammonium bicarbonate, pH 8.0. 7. Add enough 50 mM ammonium bicarbonate to cover the pieces of membrane (usually 200 to 400 µl), then add 10 µg TPCK-trypsin (10 µl of a 1 mg/ml stock). Incubate for at least 2 hr at 37°C. 8. Add another 10 µl aliquot of 1 mg/ml TPCK-trypsin and incubate again for 2 hr at 37°C. 9. Vortex briefly, then microcentrifuge briefly at full speed to collect all liquid at the bottom of the tube and transfer the supernatant to a fresh microcentrifuge tube. Rinse the membrane pieces with 500 µl of deionized water, microcentrifuge briefly, and add the rinse to the supernatant. 10. Lyophilyze in a SpeedVac evaporator and quantitate the elution of peptides by Cerenkov counting. 80% to 90% of the radioactivity should be in the eluate.
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P-labeled Post-Translational Modification: Phosphorylation and Phosphatases
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11. Oxidize the peptides by incubation in performic acid (see Basic Protocol, steps 15 to 17). 12. Add 500 µl of deionized water, lyophilize, and proceed with electrophoresis on TLC plate (see Basic Protocol, steps 21 to 31). SUPPORT PROTOCOL 1
ISOLATION OF PHOSPHOPEPTIDES FROM THE CELLULOSE PLATE Individual phosphopeptides can be isolated from the TLC plate for further analysis. The location of the phosphopeptides on the TLC plate is determined by aligning the autoradiogram with the TLC plate. The cellulose containing the phosphopeptide of interest is scraped off the plate and sucked into a pipet tip fitted with a 25-µm filter. Peptides are eluted from the cellulose and lyophilized in a SpeedVac evaporator. Materials TLC plate with resolved phosphopeptides and corresponding autoradiogram (see Basic Protocol 1 or Alternate Protocol) Electrophoresis buffer, pH 1.9 (see recipe) Single-edge razor blades 1000-µl (blue) pipet tips Small sintered polyethylene disk to fit inside blue tips (Kontes) Glass rod or similar instrument to push filters into tips Prepare the elution tips 1. Using a sharp razor blade, carefully remove the collar portion of the wide end of the blue tip. Trim ∼3 mm off the small end of the tip as well. 2. Using a glass rod, push a sintered polyethylene disk in through the wide end of the blue tip until it fits snugly in the tip. Use of a glass rod with the same diameter as the disk helps keep the disk straight, i.e., perpendicular to the length of the tip. Do not push the disk down too far or it will cause the tip to bulge out and crack. The sintered disk will serve as a barrier across the pipet tip to catch the cellulose as it is scraped from the plate. It therefore must fit securely in the tip, able to withstand the pull of the vacuum line. This is the most difficult part of this protocol—but be consoled by the fact that once made, a good tip will last forever!
3. Test the placement of the disk in the tip by attaching a piece of tubing to the wide end of the tip, with the other end of the tubing connected to a vacuum line. Now apply a strong vacuum and use one finger to block off the small end of the elution tip. Examine to make sure that the sintered disk stays in place. If the sintered disk stays in place, the elution tip is ready to use.
Mark the location of peptides to be eluted 4. Hold the TLC plate, cellulose side up, over a light box. Place the autoradiogram directly onto the cellulose layer of the plate, precisely aligning the reference marks on the plate with their images on the film. 5. Using a dark laboratory marker, trace the outline of the spot(s) of interest on the (glass) underside of the TLC plate. Phosphopeptide Mapping and Identification of Phosphorylation Sites
Be conservative—when vacuuming two adjacent spots there should always be cellulose left on the plate between the two.
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6. Remove the film, put the plate down on the lightbox, and use a soft lead pencil to trace the marker outline, this time on the cellulose side so it will be possible to see the outline without the benefit of the lightbox. Vacuum the cellulose and elute the peptide(s) 7. Connect the elution tip to a vacuum via a piece of tubing and turn the vacuum on. 8. Use the small end of the elution tip to scrape the cellulose off a spot of interest; the cellulose will be sucked up against the filter barrier in the tip as it is scraped from the plate. When the spot is completely removed from the plate, ease the tubing off the wide end of the elution tip, keeping the small end upright. The same spot can be vacuumed from multiple plates into one elution tip. However, after repeated use, the small end of the elution tip becomes “dull,” and it becomes increasingly difficult to scrape the cellulose from the plate. When this happens, use a razor blade to trim a thin sliver of plastic from the small end of the tip to recreate the sharp edge.
9. Place the elution tip into a 1.5-ml microcentrifuge tube with the wide end down and the side of the sintered disk containing the vacuumed cellulose up. The elution tip now becomes a little column.
10. Immediately pipet 100 µl of electrophoresis buffer, pH 1.9 (elution buffer), onto the cellulose; let this soak in while other spots are vacuumed from the plate. The elution buffer used here should be pH 1.9. If pH 1.9 buffer fails to elute all the phosphopeptide, try deionized water.
11. When all spots have been vacuumed and the last one has been left to soak in buffer for ∼5 min, place the microcentrifuge tubes, tips and all, into a microcentrifuge. Run the microcentrifuge at full speed for ∼3 sec, then shut it off. Pipet another 100 µl elution buffer onto the cellulose in each tube and let it sit and soak for another 5 min before centrifuging it through the column. Repeat the elutions five times to give 600 µl of eluate in each tube. This is usually enough to elute >90% of the radioactivity from the cellulose.
12. Remove the elution tip from each of the microcentrifuge tubes, being careful to leave all the eluate in the tube (some may cling to the sides of the tip as drops, which should be removed and added back to the contents of the tube). Save the elution tip. If eluting more than one spot, keep track of which tip was used for which peptide. 13. Clarify the eluate(s) by microcentrifuging 5 min at full speed (a small cellulose pellet will be visible after centrifugation; its size will depend on how snugly the sintered disk fits into the elution tip). Transfer the supernatant to a fresh microcentrifuge tube. It is very important to remove all traces of cellulose at this point, as contamination of the phosphopeptide with cellulose can ruin further analyses.
14. Count both the eluate and the “empty” elution tips by Cerenkov counting. Approximately ~90% of the radioactivity should be in the eluates, with little remaining in the cellulose left in the tips. Given the pain and frustration involved in their manufacture, a good elution tip should be saved and reused. To clean these tips, apply a vacuum to the small end of the tip and suck the cellulose out (into a vacuum flask) while aspirating ∼10 ml elution buffer or deionized water through the tip to rinse it. Dry and then count the tips on a scintillation counter (to ensure all radioactivity has been removed) before reusing them.
15. Lyophilize the eluates in a SpeedVac, then count them by Cerenkov counting. The counts here should be slightly higher than those of the liquid eluate. The number of cpm in this final sample of eluted peptide will often determine how it can be analyzed further.
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BASIC PROTOCOL 2
DETERMINATION OF THE POSITION OF THE PHOSPHORYLATED AMINO ACID IN THE PEPTIDE BY MANUAL EDMAN DEGRADATION If insufficient material is available for direct sequencing, a manual Edman degradation of the peptide can be performed to determine at which position the phosphorylated amino acid is present in the peptide. During each cycle of Edman degradation, the most amino-terminal amino acid residue is released from the peptide, and a sample from the reaction mixture is taken after each cycle. Phosphoserine or phosphothreonine is released as a derivative of serine or threonine and free phosphate; in contrast, phosphotyrosine is released as the anilinothiazolinone derivative of phosphotyrosine. Free phosphate and the PTH derivative of phosphotyrosine can be separated from the peptide by electrophoresis on a TLC plate. This approach indicates at which cycle the radioactivity, and thus the phosphorylated amino acid, is released from the peptide. Materials Eluted phosphopeptide (see Support Protocol 1) 5% (v/v) phenylisothiocyanate (PITC) in pyridine 10:1 (v/v) heptane/ethyl acetate—mix 10 parts heptane with 1 part ethyl acetate 2:1 (v/v) heptane/ethyl acetate—mix 2 parts heptane with 1 part ethyl acetate 100% (w/v) trifluoroacetic acid (TFA) Electrophoresis buffer, pH 1.9 (see recipe) 200 to 500 cpm 32P (prepared by diluting 32P orthophosphate with deionized water) or 2 mg/ml PTH-phosphotyrosine (see recipe) 1.5-ml microcentrifuge tubes, pretested for suitability (see Critical Parameters) 45°C water bath Scintillation counter appropriate for Cerenkov counting Glass-backed TLC plates (20 × 20 cm, 100 µm cellulose; EM Science) 65°C drying oven or fan Additional reagents and equipment for electrophoresis of peptides on a TLC plate (see Basic Protocol 1 and Figure 13.9.3) and autoradiography (UNIT 10.11) Determine experimental parameters 1. Decide the number of cycles to be run based on the list of candidate peptides. The number of cycles is designated as X. The starting volume for each cycle will be 20 ìl.
2. Dissolve the eluted peptide in 20 µl deionized water in a 1.5-ml microcentrifuge tube (henceforth called the reaction tube). 3. Remove a sample equal to 20/(X + 1) µl to a new tube; this is the starting material sample. Store this at 4°C. This sample will be lyophilized with the other cycle fractions at a later point.
Perform the Edman reactions 4. Add enough deionized water to the reaction tube to restore the volume to 20 µl. Count the sample at this point:
Phosphopeptide Mapping and Identification of Phosphorylation Sites
a. to insure that the expected number of cpm have in fact been removed from the initial sample (as the starting material sample); b. to check the cpm at the start of each given cycle. 5. Add 20 µl of 5% phenylisothiocyanate in pyridine to each reaction tube, vortex well, spin briefly in a microcentrifuge to collect the sample at the bottom, and incubate 30 min at 45°C.
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6. Add 200 µl of 10:1 heptane/ethyl acetate to each reaction tube and vortex for 15 sec. Microcentrifuge 1 min at full speed to separate the two phases. The pyridine will partition into the (upper) organic phase.
7. Carefully remove the upper organic phase using a plastic transfer pipet. Reextract the (bottom) aqueous phase a second time with 10:1 heptane/ethyl acetate as in step 6. 8. Extract the aqueous phase two more times as in step 6, this time using 2:1 heptane/ethyl acetate. 9. Freeze the aqueous phase on dry ice and lyophilize in a SpeedVac evaporator. 10. Dissolve the dried sample in 50 µl of 100% TFA and incubate this 10 min at 45°C. 11. Lyophilize the sample in a SpeedVac evaporator. 12. Count the sample by Cerenkov counting. There should be the same number of cpm as at the beginning of the cycle (i.e., at step 4 in this case).
13. Add 20 µl deionized water to the reaction tube, vortex, and microcentrifuge briefly. Remove a portion for analysis of the first-cycle products that is equal to 20/X. Store this at 4°C with the starting material sample. 14. Add deionized water to restore the sample volume to 20 µl to start the second cycle. Repeat steps 5 to 12. 15. After the second cycle, add 20 µl deionized water, resuspend the remaining sample, and remove 20/(X − 1) µl to a new tube for analysis of the second-cycle products. Repeat steps 4 to 12. 16. Continue repeating steps 4 to 12 until the desired number of cycles have been run. For each new cycle, the amount of the sample to be removed is 20/X − Y where Y equals the cycle number minus 1.
Analyze the Edman products 17. Lyophilize all samples to dryness in a SpeedVac evaporator. Count all final samples by Cerenkov counting, 18. If lyophilized, dissolve the samples in 5 µl of pH 1.9 electrophoresis buffer or deionized water. Microcentrifuge 2 min at maximum speed to bring down any insoluble material. Alternatively, if the sample volumes removed after each cycle are small enough, skip steps 17 and 18 and load the samples directly onto the TLC plate.
19. Spot all samples from the analysis of a given phosphopeptide at least 1 cm apart on a line of origins running vertically through the center of the TLC plate (Fig. 13.9.5). As a marker, depending on the phosphoamino acid content of the peptide under investigation, spot 50 to 200 cpm of [32P]phosphate or 1 to 2 µg PTH-phosphotyrosine (0.5 to 1.0 µl of 2 mg/ml PTH-phosphotyrosine) at an origin on that same vertical line. Load 1⁄3 to 1⁄2 µl of sample at a time, air drying the sample between applications (see Basic Protocol 1, step 25). 20. Wet the plate as described in Figure 13.9.5. 21. Prepare the HTLE 7000 apparatus and electrophorese the samples for 25 min at 1.0 kV in pH 1.9 electrophoresis buffer (see Basic Protocol 1 and Figure 13.9.3). 22. After drying the plate (either in a 65°C oven or with a fan), mark it appropriately with radioactive or fluorescent markers and expose it to presensitized film with an intensifying screen at −70°C (autoradiography; UNIT 10.11).
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A
B marker origin cycle 6 cycle 5 cycle 4
4.5 cm
2 cm
15 cm
cycle 3 cycle 2
15.5 cm
cycle 1 starting material 3 cm 10 cm
5.0 cm
10 cm 12.5 cm
12.5 cm
Figure 13.9.5 Sample and dye origins and blotter dimensions for analysis of manual Edman degradation products at pH 1.9. (A) Location of the sample and standard origins. To mark a TLC plate, the plate is placed on top of a life-sized template on top of a light box and the origins are marked on the cellulose side using a very blunt extra-soft pencil. (B) Dimensions of the blotter and the location of the slot that fits over multiple sample and marker origins. The blotter is soaked in electrophoresis buffer, blotted with a sheet of Whatman paper to remove most of the buffer, and placed on top of the TLC plate so that the origins are in the middle of the slot.
BASIC PROTOCOL 3
DIAGNOSTIC SECONDARY DIGESTS TO TEST FOR THE PRESENCE OF SPECIFIC AMINO ACIDS IN THE PHOSPHOPEPTIDE Further information about a phosphopeptide of interest can be obtained by digestion with sequence-specific proteases or cleavage by site-specific chemicals. After incubation with a protease or chemical, the peptide is analyzed by separation in two dimensions on a TLC plate. A change in mobility upon treatment with a particular reagent indicates that the peptide was susceptible to cleavage, and consequently that the amino acid or amino acid sequence that confers susceptibility to cleavage by this reagent must be present in the peptide. Materials Eluted phosphopeptide (see Support Protocol 1) Enzyme to be used for digestion and appropriate buffer (see Table 13.9.1) 2-Mercaptoethanol (2-ME) Electrophoresis buffer of appropriate pH (see recipe) Water bath at appropriate temperature for enzyme digestion Glass-backed TLC plates (20 × 20 cm, 100 µm cellulose; EM Science) Additional reagents and equipment for chromatography and electrophoresis of phosphopeptides (see Basic Protocol, steps 24 to 31)
Phosphopeptide Mapping and Identification of Phosphorylation Sites
1. Dissolve the eluted phosphopeptide in 50 µl of the appropriate buffer in a microcentrifuge tube and microcentrifuge briefly to collect all solution at the bottom of the tube. Check the pH of the peptide solution by spotting 1 µl on a piece of pH paper to be sure that this final pH will allow enzyme activity. If the buffer’s pH has been altered dramatically by addition of the peptide, adjust it before adding enzyme.
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Table 13.9.1
Specificities and Digestion Conditions for Enzymes and Other Cleavage Reagents
Enzyme or reagent Specificitya
Digestion conditions
Comments
TPCK-trypsin
K–X; R–X
pH 8.0-8.3
α-Chymotrypsin Thermolysin
F–X; W–X; Y–X X–L; X–I; X–V
pH 8.3 pH 8.0, 1 mM CaCl2, 55°C
Does not cut K/R-P; cuts inefficiently at K/R-XP.Ser/P.Thr and K/R-D/E; cuts wells at K/RP.Ser/P.Thr; cuts X-K/R-K/R-K/R incompletely Does not cleave F/W/Y-P or P.Tyr-X Will recognize most apolar residues to some extent; CaCl2 may affect the electrophoretic mobility —
Proline-specific P–X endopeptidase Cyanogen bromide M–X (CNBr)
pH 7.6
V8 protease
E–X
50 mg/ml CNBr in 70% formic acid, 90 min, 23°C pH 7.6
Endoproteinase Asp-N Formic acid
X–CSO3H; X–D
pH 7.6
D–P
70% formic acid, 37°C, 24-48 hr
Toxic; will cleave only unoxidized methionine
Will not cleave at every E in whole proteins; does give a consistent pattern Will cleave X-E at high enzyme concentrations —
aThe dash indicates the cleavage site. See APPENDIX 1A for definitions of the one-letter abbreviations for amino acids.
2. Remove a portion of the sample (usually 50%) to be run both as an undigested control and as a mix with a portion of the digested sample. 3. Add 1 to 2 µg enzyme to the portion of the sample to be digested, vortex, and concentrate the sample in the bottom of the tube by microcentrifuging briefly. 4. Incubate all tube(s) in a water bath at the appropriate temperature for at least 1 hr. 5. Add another aliquot of enzyme and continue the incubation step for an additional hour. 6. Add 1 µl 2-ME to each sample and boil 5 min to inactivate the enzyme Do this to all samples to ensure uniformity of sample preparation. It is necessary to completely inactivate the enzyme prior to loading the sample on the plate when analyzing a mix of digested and undigested peptide, since the undigested sample may be rapidly digested when the two samples are mixed.
7. Lyophilize the samples in a SpeedVac evaporator. 8. Resuspend the samples by vortexing vigorously in 6 µl of electrophoresis buffer of the appropriate pH. Microcentrifuge at full speed to bring down insoluble material. 9. Load half of the undigested sample on a single TLC plate. Load half of the digested sample on each of two TLC plates; on one of these load the other half of the corresponding undigested sample as a mix. 10. Perform electrophoresis and chromatography on the plate as described above (see Basic Protocol 1, steps 24 to 31). Based on the position where the particular phosphopeptide being analyzed ran in the original map, choose a pH and running time that will allow good separation of the peptide from its potential cleavage products but will ensure retention of the smaller cleavage products on the plate.
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SUPPORT PROTOCOL 2
PREPARATION OF PHOSPHOPEPTIDES FOR MICROSEQUENCE DETERMINATION OR MASS SPECTROMETRY The following is a general protocol and list of considerations for generating enough material for analysis by mass spectrometry or microsequencing starting with either intact cells or an in vitro system. 1. Optimize 32P labeling of the protein of interest. If the site of interest is seen only in stimulated cells, a time course of phosphorylation following stimulation may be helpful, as would determination of the optimal concentration of agonist. If an in vitro system is being employed, determine the optimum conditions (time, and ratio of kinase and substrate concentrations) for the kinase reaction. Include 1 mM cold ATP in the reactions to maximize the stoichiometry. UNIT 13.7, which deals with in vitro phosphorylation reactions, provides a detailed discussion of these parameters and how to manipulate them.
2. Calculate the number of cells or amount of substrate needed to isolate 10 pmol phosphorylated material. Even under optimal conditions, it is often not possible to achieve more than 25% stoichiometry of phosphorylation in vitro. In intact cells, the stoichiometry may be even less. It cannot hurt to overestimate the amount of starting material required, as the losses during the isolation procedures will always exceed expectation.
3. When calculating how to scale up the reactions, consider the following points.
Phosphopeptide Mapping and Identification of Phosphorylation Sites
a. The radioactivity of these samples is only used for visualization purposes—i.e., to determine which gel band to isolate, which phosphopeptide to isolate from the TLC plate, and which HPLC fraction(s) to use for final analysis. Thus, the majority of the material can be unlabeled, as only ∼1000 cpm per map spot are necessary for analysis at the time when the preparative HPLC is run. When isolating overexpressed protein from cells, labeling only 2 or 3 dishes of the 20 needed to generate enough material may be sufficient. When using an in vitro system, perform only one reaction using [γ-32P]ATP (include only the very minimum amount of unlabeled ATP necessary for kinase activity) to generate the labeled material. To generate enough material for further analysis, perform an additional kinase reaction with unlabeled ATP only. For visualization, mix the labeled and unlabeled samples before resolving them by SDS-PAGE. b. The efficiency of protein elution decreases as the amount of gel increases, so try to keep the number of lanes on the preparative gel(s) to a minimum. About four lanes/slices of acrylamide gel can be successfully extracted per tube. c. If ∼20 µg substrate protein is present in each elution sample, it will not be necessary to add carrier protein at the TCA precipitation step (see Basic Protocol 1, step 11). This will result in a cleaner sample, as the tryptic fragments of the carrier protein will be eliminated from the mix of fragments run on the TLC plate. d. The 20 µg trypsin used to digest map samples in Basic Protocol 1 is in vast excess. While it is important that the digestion go as far to completion as possible, it is probably not necessary to scale up the amount of trypsin used. Instead, consider pooling several like samples at the performic acid digestion step (at the end of the 60-min incubation, in order to give the protein the maximum time to dissolve). Adding 1 to 5 µg (total) trypsin to even 50 µg of protein for digestion is not unreasonable. Minimizing the amount of trypsin used will in turn minimize the amount of “extra” protein loaded on each TLC plate and ensure that the sample does not streak due to overloading.
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e. Determine the number of TLC plates to be run based on the amount of total protein to be analyzed—total protein includes the amount of trypsin and the amount (if any) of carrier protein used as well as the amount of the protein of interest. Even though the capacity of the TLC plates is ∼100 µg, to ensure good separation, no more than 60 µg of protein should be run on each plate. REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Chromatography buffers Phosphochromatography buffer 750 ml n-butyl alcohol 500 ml pyridine 150 ml glacial acetic acid 600 ml deionized water Store at room temperature Isobutyric acid buffer 1250 ml isobutyric acid 38 ml n-butyl alcohol 96 ml pyridine 58 ml acetic acid 558 ml deionized water Regular chromatography buffer 785 ml n-butyl alcohol 607 ml pyridine 122 ml glacial acetic acid 486 ml deionized water Store all of the above buffers up to 6 months at room temperature.
Electrophoresis buffers For each of these buffers, mix well and check the pH. Record the pH and the date on the bottle; if the pH is more than a tenth of a unit off, remake the solution. Do not adjust the pH. Store all buffers at room temperature. pH 1.9 buffer: 50 ml formic acid (88% w/v) 156 ml glacial acetic acid 1794 ml deionized water pH 3.5 buffer 100 ml glacial acetic acid 10 ml pyridine 1890 ml deionized water pH 4.72 buffer 100 ml n-butyl alcohol 50 ml pyridine 50 ml glacial acetic acid 1800 ml deionized water continued
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pH 6.5 buffer 8 ml glacial acetic acid 200 ml pyridine 1792 ml deionized water pH 8.9 buffer 20 g ammonium carbonate 2000 ml deionized water Green marker dye Prepare a solution containing 5 mg/ml ε-dinitrophenyllysine (yellow) and 1 mg/ml xylene cyanol FF (blue) in pH 4.72 electrophoresis buffer (see recipe) diluted 1:1 with deionized water. Store up to 1 year at room temperature. PTH-phosphotyrosine, 2 mg/ml Combine 20 µl of 100 mg/ml phosphotyrosine with 20 µl of 5% (v/v) phenylisothiocyanate in pyridine. Incubate 30 min at 45°C. Extract twice with 200 µl of 10:1 (v/v) heptane/ethyl acetate, then twice with 200 µl of 2:1 (v/v) heptane/ethyl acetate (see Basic Protocol 2, steps 6 to 8, for extraction technique). Freeze the aqueous phase and lyophilize in a SpeedVac evaporator. Dissolve the sample in 0.1 N HCl, incubate 20 min at 80°C, and lyophilize again in a SpeedVac evaporator. Dissolve in 1 ml pH 1.9 buffer (see recipe for electrophoresis buffers). PVP-360 in 100 mM acetic acid 0.5 g PVP-360 (Sigma) 575 µl glacial acetic acid 99.4 ml deionized water Store up to 1 year at room temperature COMMENTARY Background Information
Phosphopeptide Mapping and Identification of Phosphorylation Sites
Phosphopeptide mapping is a very sensitive technique that can help the investigator answer a variety of questions about a protein of interest. For some, phosphopeptide mapping is a tool to find out whether a particular protein is phosphorylated on one or more sites. This question can be answered by simply running a phosphopeptide map of the protein labeled in living cells. Other investigators want to know whether the increase in phosphorylation seen when cells are treated with a particular agent is restricted to one or more specific sites or whether it is evenly distributed over all phosphorylation sites present in the protein. Finally, detailed analysis of phosphopeptides isolated from a TLC plate can be used to identify the residues that are phosphorylated in a protein of interest. Several different strategies may be followed to identify the phosphorylation site represented by a particular spot on a phosphopeptide map. This is most definitively accomplished by eluting phosphopeptides from cellulose plates either for direct sequencing or for analysis by mass spectrometry (see Support Protocol 2; Fischer et al.,
1991; Wang et al., 1993; Mitchelhill et al., 1997; Mitchelhill and Kemp, 1999). While these two techniques require expensive instruments and expertise not found in most laboratories, such analysis can often be arranged by collaboration. However, sometimes it is not possible to take advantage of these techniques, since they require 1 to 10 pmol of material for analysis. For a 50-kDa protein one would need 0.1 to 1.0 mg starting material, assuming 50% recovery and 100% stoichiometry of phosphorylation at the site of interest. Use of an in vitro phosphorylation system that mimics the situation in intact cells will simplify matters greatly. Further considerations and strategies for preparation of samples for these two techniques are discussed in Support Protocol 2. Another approach to phosphorylation-site identification is to make an educated guess as to the identity of the site. Clues to a site’s identity include phosphoamino acid analysis of the individual phosphopeptide (UNIT 13.3); the result of manual Edman degradation of a phosphopeptide providing the cycle at which the phosphate is released and thus the position of
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the phosphorylated residue in the peptide (see Basic Protocol 2); and secondary enzymatic digests of the phosphopeptide, which can be used in a diagnostic sense to determine the presence of other specific amino acids in the peptide (see Basic Protocol 3). All three of these procedures are easily accomplished in a laboratory that is already set up for phosphopeptide mapping. The first step in all three is the isolation of the phosphopeptide from the cellulose plate (see Support Protocol 1). The validity of one’s guess can be tested by phosphopeptide mapping of a mutant protein lacking a phosphate acceptor at the site in question. Alternatively the guess can be substantiated by synthesizing the tryptic phosphopeptide and testing it for comigration with the phosphopeptide isolated from the peptide map.
Critical Parameters and Troubleshooting Generating phosphopeptide maps Keep in mind that the sort of analyses presented throughout this unit will give information regarding only the acid-stable forms of phosphoamino acids (i.e., phosphoserine, phosphothreonine, and phosphotyrosine) and will essentially ignore other forms such as phosphohistidine and phosphoaspartate, should they be present. Carrier protein. The authors prefer to use RNase as carrier protein during TCA precipitation, particularly when analyzing proteins labeled in intact cells, because it degrades 32P-labeled RNA species that may have copurified with the protein of interest. The nucleotides generated by the degradation of RNA do not precipitate in TCA. Cleaving the protein. In order to generate a phosphopeptide map, the 32P-labeled protein needs to be cleaved into smaller fragments that can be separated by electrophoresis and chromatography on TLC plates. To do this requires an enzyme or chemical agent that cleaves reproducibly and with a certain frequency. If not enough cleavage sites are present, the fragments generated will be too large and will not be separated easily by electrophoresis and chromatography on TLC plates. In addition, large fragments may contain multiple phosphorylation sites. This leads to maps that are less informative and more difficult to analyze. The authors routinely use trypsin and chymotrypsin. Other reagents are available (Table 13.9.1), but most of them cut much less frequently and some do not work very efficiently on full-length proteins.
Removing ammonium bicarbonate. Following digestion, repeated cycles of lyophilization are carried out to evaporate all ammonium bicarbonate. The presence of salts in the sample will interfere with the electrophoretic separation of the peptides. After lyophilization, the protein digest will be an invisible film at the bottom of the tube. The presence of any crystalline material indicates the presence of salts, most likely ammonium bicarbonate that can be removed by additional rounds of lyophilization. Controlling oxidation. Both cysteine and methionine can give rise to several oxidized derivatives. The oxidation state of these amino acids affects the mobility of peptides during chromatography, resulting in separation of oxidation-state isomers. This complicates the interpretation of the phosphopeptide map. To prevent this, the protein or peptides are oxidized by incubation in performic acid at 4°C. Incubation at higher temperatures may give rise to unwanted side reactions and should be avoided. Elution and TCA precipitation or transfer to a membrane? In Basic Protocol 1, the 32P-labeled protein is isolated from a small slice of a dried polyacrylamide gel by rehydrating and grinding up the gel and then eluting it in a buffer containing SDS and 2-mercaptoethanol. The protein is subsequently TCA precipitated, oxidized, and digested with trypsin. This is a timeconsuming and laborious procedure. Yields are variable and usually not better than 50%. The alternative is to transfer the protein to a PVDF membrane; any unoccupied protein-binding sites on the strips of membrane containing the protein of interest are blocked by incubation with PVP-360 in acetic acid before incubation with trypsin. Most peptides dislodge from the membrane during the digestion. This protocol is much faster and less laborious, and does not require the use of additional carrier proteins that may lead to overloading of the TLC plate and to streaky maps. Obviously this method is a poor choice for proteins that transfer poorly from the polyacrylamide gel to a membrane. In addition, it is possible that certain peptides that are generated during protease digestion retain a high affinity for the membrane and therefore fail to elute. If those peptides contain a phosphorylation site, this site will not be represented on the peptide map. This can lead to misinterpretations of the results. It is therefore advisable to first compare maps generated with Basic Protocol 1 and the Alternate Protocol. If these maps are identical, and if the protein transfers
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well from the gel to the membrane, the Alternate Protocol should be the protocol of choice. Amount of sample. The authors like to load at least 1000 cpm on a plate for a peptide map. If the final sample has many more than 1000 cpm and a “pretty-looking” map is desired, it may be better to load only a fraction of the sample. Remember that overloading can lead to streaky maps. If a preparative map from which a particular peptide will be isolated is being run, it may be best to run the entire sample on two (or more) separate plates. Theoretically, it should be possible to separate 100 µg of material on a single TLC plate; this is often not the case in practice. Check the rate at which the first drop spotted sinks into the cellulose; as more sample is spotted, this rate will decrease. If, while spotting, one reaches a point where the sample drop just sits on the origin and does not spread into the cellulose, it is time to stop loading. Peptide diffusion. Peptides diffuse during the electrophoresis and chromatography, and this leads to a reduction in resolution and sensitivity. To counteract this, the authors try to keep the area on the TLC plate onto which the sample is spotted as small as possible by spotting only a small amount at a time (i.e., <1 µl) and drying the sample origin between spottings. In addition, the sample is concentrated by wetting the TLC plates with electrophoresis buffer using a blotter with holes cut out around the origin (Figs. 13.9.1 and 13.9.3). Pressing the edges of the hole onto the plate makes buffer move from the blotter towards the center of the hole. This concentrates the sample on the origin. For this process to work well, the origin has to be precisely in the center of the hole. In addition, the buffer has to move with similar speed from the entire circumference towards the origin. The sample will inevitably streak if the buffer takes a long time to wet the spot, or moves unevenly through the spot. Electrophoresis system. In the authors’ laboratories the HTLE 7000 electrophoresis system (Fig. 13.9.2) is used. This system features water cooling and an inflatable airbag that presses the TLC plate against the cooling plate. Water cooling prevents overheating during electrophoresis. The inflatable airbag presses excess buffer from the TLC plate; this limits diffusion of the peptides and improves resolution. Buffers. Three different buffers are typically used for electrophoresis: pH 1.9, pH 4.72, and pH 8.9. Less often used are pH 3.5 and pH 6.5 electrophoresis buffer. To find out which buffer gives the best separation of peptides generated
from a particular protein, all three buffers should be tested. If possible, the authors prefer to work with pH 1.9 buffer. Most peptides dissolve well at this pH, and its use less often results in streaky maps. The authors usually spot the digest on the origins as marked in Figures 13.9.1 and 13.9.3. For optimal separation of the phosphopeptides generated from a particular protein, the position of the origin and the electrophoresis time may need to be changed. The authors prefer to change the time of electrophoresis rather than the voltage. Microcentrifuge tubes. Historically we have found that certain tubes are more apt than others to produce such unwanted side reactions; for this reason it is advisable to stock certain lots of tubes that do not produce such artifacts in the final maps. Similarly, brands and batches of tubes appear to differ in the extent to which peptides “stick” to them during the final steps of the protocol. The authors use tubes from Myriad Industries with success. Chromatographic process. Chromatography usually takes 12 to 15 hr, but the exact time may vary depending on the age of the chromatography buffer, the batch of plates, the buffer system, the quality of reagents used in the buffer, and the temperature in the room. Three different chromatography buffers are commonly used: isobutyric acid buffer, regular chromatography buffer, or phosphochromatography buffer (see Reagents and Solutions). Pyridine, which is present in all three chromatography buffers, oxidizes and turns yellow over time. Do not use oxidized pyridine to make up chromatography buffers. To find out which buffer gives the best separation of the peptides generated from a particular protein, all three chromatography buffers should be compared. Most investigators prefer not to use isobutyric acid buffer because it is particularly foul smelling. During the chromatography run, the air space in the tank saturates with buffer and this makes it possible for the volatile chromatography buffer to run all the way to the top of the TLC plate. When the chromatography tank is opened, most of the buffer-saturated air will escape from the tank. This makes it counterproductive to extend the run after the tank has been opened. Therefore, do not open the tank when chromatography is in progress, and take all plates out when the chromatography tank is opened. Separation of the yellow and blue dye functions as a control for successful electrophoresis and allows one to follow the progress of the chromatography as it proceeds. The dyes can
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also be used as standards relative to which the mobility of phosphopeptides of interest can be described, and can be used as markers for the comparison of one plate to another. The yellow compound ε-dinitrophenyllysine is neutral at pH 4.72 and pH 8.9 and defines the position to which neutral peptides migrate; at pH 1.9 this compound is positively charged. Phosphopeptide identification After running several phosphopeptide maps, it may become apparent that particular phosphopeptide spots present on the map change in intensity upon treatment of the cells with specific reagents. Such observations often lead to the next question—what is the identity of peptide “A” that becomes phosphorylated following treatment of the cells with factor “B”? If ∼1 to 10 pmol of phosphorylated peptide can be generated, the peptide is isolated from the TLC plate, purified by HPLC, and identified by mass spectrometry or microsequencing. In many cases, it is not possible to obtain a phosphorylated peptide in such quantities. The investigator is then forced to learn as much about the phosphopeptide as possible before making an educated guess. The authors find it useful to make a list of all possible candidate peptides including some of their properties. The next step is to eliminate as many candidates as possible using mobility predictions and the results of relatively simple experiments that can be performed on the minute amounts of labeled peptides isolated from TLC plates. Making a list of candidate peptides and eliminating the first candidates Make a list of all possible phosphopeptides that could be generated from the protein of interest given the enzyme used in the primary digest; be sure to include partial cleavage products on this list. This list of peptides should include the nature and position of amino acids that can be phosphorylated and the peptides’ susceptibility to further cleavage by proteases or chemicals (for an example, see van der Geer and Hunter, 1990). After making such a list, first calculate and then plot the predicted mobilities of all candidate phosphopeptides. See Table 13.9.2 for values that can be used to do this. To calculate electrophoretic mobility. The mobility of a peptide in the electrophoresis dimension is dependent on its charge (e) to mass (M) ratio, as Mr = keM−2/3. When calculating relative mobilities (Mr) the simplified
equation Mr = eM−1 can be used with good success. The net charge on a peptide is calculated by summing the charges of the N and C termini and those of the side chains of its amino acids at a given pH, and dividing by either the actual mass of the peptide or simply by the number of amino acids in it. Approximate charge values at the specific pHs commonly used for electrophoresis are given in Table 13.9.2. To calculate chromatographic mobility. A peptide’s mobility in the chromatographic dimension is dependent on its hydrophobicity, and thus on its amino acid sequence. The order of the amino acids will also change the peptide’s mobility; thus, two peptides of identical sequence that are phosphorylated at one or the other of two possible sites may migrate different distances in the second map dimension, although they migrate identically in the first dimension since their charge/mass ratio is the same. While it is not possible to predict chromatographic mobilities exactly, relative mobilities can be plotted with some success by calculating an average mobility for the peptide based on migratory values of its constituent amino acids. This is not ideal, since the calculated Rf values of individual amino acids are significantly influenced by the presence of their charged amino and carboxy termini, which of course are noncontributory in the context of a peptide. This accounts in part for the compression of the calculated maps compared with the observed peptide migrations. Values for chromatographic mobilities of amino acids have been published in Boyle et al. (1991); these were determined for each individual amino acid relative to the ε-dinitrophenyllysine (yellow) marker using cellulose plates available 20 years ago. The quality of the cellulose used in such plates has changed markedly over the years; contact Ned Lamb (CNRS, Paris) for values that have been empirically determined more recently (http://www.genestream.org). Bear in mind that these calculations can also be accomplished using a computer program. Lamb has constructed a Web site for analysis of phosphopeptide maps, which may be found at the same Web site. Phospepsort 4, the program that he has developed based on an earlier version which originated at the Salk Institute, gives the biophysical characteristics as well as the electrophoretic and chromatographic mobilities of each proteolytic fragment. Alternatively, predicted peptide mobilities can be visualized using the graphical interface to Phospepsort 4: Mobility. In addition, Lamb is working
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Table 13.9.2 Approximate Charge Values at Specific pHs Commonly Used for Electrophoresisa
Amino-terminal NH2 Carboxy-terminal COOH Arginine Aspartate Cysteine (oxidized) Histidine Glutamate Lysine Phosphoserine Phosphothreonine Phosphotyrosine
pH 1.9
pH 3.5
pH 4.7
pH 6.5
pH 8.9
+1 N +1 N −1 +1 N +1 −1 −1 −1
+1 −0.5 +1 N −1 +1 N +1 −1 −1 −1
+1 −1 +1 −0.7 −1 +1 −0.5 +1 −1 −1 −1
+1 −1 +1 −1 −1 +0.5 −1 +1 −1.3 −1.3 −1.3
+0.5 −1 +1 −1 −1 N −1 +1 −2 −2 −2
aN indicates neutral.
on a program called Resolve that fits the calculated mobility values to the actual values for peptides of known composition. The program then reads the position of a spot on the actual map and calculates which peptide(s) derived from the protein being mapped could have the mobility of that spot. It is imperative to note that to date there is no program that accurately predicts the mobility of all peptides of a protein. This may be explained by the fact that mobilities are calculated using values established for single amino acids rather than for peptides. Plotting predicted phosphopeptide mobilities on a graph using linear axes results in a greatly compressed map as compared to that observed in autoradiograms, especially in the chromatographic dimension. Therefore, do not despair if the predicted map of all phosphopeptides in the protein of interest looks nothing like the actual map that was generated experimentally. The value of such predictions comes from the fact that the relative positions of phosphopeptides are predicted with great accuracy by such programs/calculations. Thus, if the predicted mobility of a hypothetical phosphopeptide places it on the anode side of a cluster of phosphopeptide candidates, while the phosphopeptide of interest on the actual map is present on the cathode side of the cluster, the hypothetical phosphopeptide may be eliminated from further consideration. This example illustrates how careful use of predicted peptide mobility maps may lead to elimination of candidate peptides. Phosphopeptide Mapping and Identification of Phosphorylation Sites
Isolating peptides from TLC plates Phosphopeptides isolated by elution from cellulose, as described above, can be used without further purification for certain types of
analysis. What many people overlook, however, is that such a sample is by no means necessarily pure. It includes, in addition to the radioactive phosphopeptide in question, any unlabeled tryptic fragments that may have comigrated with it on the cellulose plate—these peptides are generated from trypsin itself and from the carrier protein used in the TCA precipitation during sample preparation. For this reason, the sample is usually further purified by HPLC to clean it up before analysis by mass spectrometry or microsequencing; the column fractions are counted in a scintillation counter to determine which ones to use for further analysis. However, for manual Edman degradation (see Basic Protocol 2), secondary cleavage (see Basic Protocol 3), and phosphoamino acid analysis (UNIT 13.3) no further purification is necessary, as the interpretation of the results relies solely on the visualization of the resultant 32P-containing reaction products. The presence of unlabeled contaminants does not interfere with the interpretation of results. Phosphoamino acid analysis Perhaps the most obvious step to take with an unidentified phosphopeptide in hand is to determine the phosphoamino acid content of the peptide of interest. This will eliminate many candidate peptides from consideration; this step is obviously not necessary if a long exposure of the phosphoamino acid analysis of the labeled protein in question indicates that only one species of phosphorylated amino acid is present. For phosphoamino acid analysis only 50 cpm of purified phosphopeptide is needed (though in peptide mapping and related protocols one can never have enough cpm). The details of such analysis have been discussed in
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detail in UNIT 13.3. Briefly, the phosphopeptide eluted from the TLC plate is hydrolyzed by incubation for 60 min at 110°C in 30 µl of 6 N HCl. The appearance of yellow to brown color in the sample during hydrolysis indicates that some cellulose remained despite efforts to clarify the phosphopeptide eluate. This sometimes causes the sample to streak. After the sample is lyophilized, it is resolved with stainable standards in two dimensions by electrophoresis as described (UNIT 13.3). The phosphoamino acid composition is determined by matching the resultant spot(s) on the autoradiogram with the ninhydrin-stained standards on the cellulose plate. Manual Edman degradation At pH 8 to 9, phenylisothiocyanate reacts with the free amino group(s) of a peptide to form a corresponding phenylthiocarbamyl peptide. Treatment of these PTC-peptides with acid (TFA) results in the cleavage of the derivatized amino-terminal amino acid and its release as an anilinothiazolinone molecule. This latter species is not stable, and will cyclize to yield the phenylthiohydantoin (PTH) derivative of the amino acid in aqueous acid. If a phosphoserine or phosphothreonine residue is present, a βelimination during the cyclization releases free phosphate. Phosphotyrosine, however, is simply released as the anilinothiazolinone derivative. This may be converted to the phenylhydantoin form for analysis by incubating it in 0.1 N HCl at 80°C for 20 min. PTH-phosphotyrosine to use as a marker is easily synthesized by reacting phosphotyrosine with phenylisothiocyanate and then heating it in acid (see Basic Protocol 2, step 5); it can be visualized as a dark spot using a hand-held UV light. While the protocol is relatively simple, each cycle takes ∼2 hr to complete and requires at least 100 cpm for an unambiguous result. It is important to run a portion of the starting material out on the TLC plate to show how much, if any, free phosphate is there, since some hydrolysis of the peptide may have occurred during its isolation. Because at each cycle the reaction may not go to completion, one should plan to do at least one more cycle than is predicted to be necessary to release the phosphate (i.e., if all the candidate peptides are phosphorylated at or before the third residue from the N terminus, at least 4 cycles should be run). Thus, how many cpm are in the eluted map spot may determine how many cycles are run. In any case, the authors generally do not attempt to perform more than 5 or 6 cycles. If no
phosphate is released during the course of these cycles, this conclusion in itself will usually greatly reduce the number of candidate sites. In addition to the position of the phosphorylated residue, clues to a peptide’s sequence may also be gleaned from Edman degradation. The way in which the residual peptide shifts its electrophoretic position on the plate after each cycle will indicate whether an acidic or basic amino acid has just been removed. If the tryptic peptide’s carboxy-terminal residue is a lysine, the lysine’s ε-amino group will be derivatized in the first cycle, and so a positive charge will be lost in that instance as well. If an automated peptide sequencer is available, 20 or more Edman cycles may be analyzed by coupling the phosphopeptide via carboxyl groups to a Sequelon membrane (Millipore) and letting the machine do the work. At the end, the fractions are counted to see where the radioactivity is released. This method obviously requires far fewer cpm in a phosphopeptide sample than does the manual Edman protocol, since one only analyzes the released material for radioactivity rather than a portion of the whole sample at the end of each cycle. A method for adapting an automated sequencer for such a purpose is discussed in Mitchelhill et al. (1997). An adaptation of the protocol presented above is found in Fischer et al. (1997). This protocol uses a volatile isothiocyanate (trifluoroethyl isothiocyanate) and volatile buffers; as a consequence the extraction steps can be eliminated and this results in shorter cycle times (∼45 min). Secondary digests In the past, secondary digests of tryptic phosphopeptides represented a large part of the further analysis of these peptides. The utility of these digests, however, is totally dependent on the sequence of the protein in question. A list of enzymes commonly used in such digests is found in Table 13.9.1, along with their cleavage site(s) and optimal pH and temperature. As is true for trypsin, these enzymes sometimes cleave inefficiently when encountering a cleavage site in a certain sequence context—some of these problematic sites are listed as well. In the authors’ experience, the enzymes/reagents that cleave only one amino acid (i.e., prolinespecific endopeptidase, V8, or cyanogen bromide) tend to be more useful and give less ambiguous results. Do not expect that the results of one enzyme digest will eliminate more than a few candidate
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Phosphopeptide Mapping and Identification of Phosphorylation Sites
sites. As it is most likely that the enzyme in question will not cleave the phosphopeptide, it is imperative that a positive control be included in the digest—i.e., a peptide whose sequence is known and which the enzyme will cleave. If no such peptide is available, a portion of the primary digest might be used as a positive control—with luck this will contain at least one peptide whose migration will be changed by this secondary digestion. Running out the mix of undigested and digested peptides is very important, since failure to comigrate with the original peptide will unequivocally demonstrate a change in the phosphopeptide’s mobility. A sample treated exactly like the digested sample, but without the cleavage reagent, should be analyzed in parallel. This is to ensure that changes in mobility seen in the digested peptide are truly due to the presence of the cleavage reagent. When deciding where to spot the samples and what conditions to use to run the plates, keep in mind that if the peptide did not migrate very far in the original map, it may be possible to load two sample on a single plate. Also keep in mind, however, that free phosphate may be released during the enzymatic digestion (due to elevated temperatures and resulting hydrolysis). Free 32P-phosphate originating from the sample loaded to the right may complicate the interpretation of the sample loaded on the left (anode) side of the plate. It should be noted that additional information about the other amino acids in a phosphopeptide may be gleaned just by running a tryptic digest in the electrophoretic dimension at another pH—identification of the different peptides by their mobility in the chromatography dimension will allow one to compare the mobility of peptides when run at different pHs and decide if a particular spot has changed its migration in the first dimension. Such a change in migration would be affected only if the phosphopeptide contained an amino acid whose charge was changed at the second pH used.
and that, while enough is loaded to be easily visualized, the plate is not overloaded (which would cause streaking and therefore render the comigration evidence ambiguous). If the peptide is synthesized as a phosphopeptide, ∼5 to 25 µg of pure peptide is needed for each comigration, as it will need to be ninhydrin stained for visualization. This is by far the easier approach—starting with an unphosphorylated peptide entails not only finding a kinase that will phosphorylate it, but also purifying the phosphopeptide first before running the comigration. In either case, synthesis of peptides, and in particular those with special residues such as phosphorylated amino acids, is not inexpensive, and so this type of experiment is usually attempted after one has accumulated several other clues regarding a site’s identity. The alternative is to mutate the remaining candidate phosphorylation sites. An epitopetagged version of the mutant protein can then be expressed in cells and mapped to check whether the spot in question disappears from the map. This approach of course assumes that the sequence of the protein in question is known and that one has a clone in hand for mutagenesis. While satisfying to many, this mutagenesis approach is not definitive, as it can be argued that the mutated form of the protein may not fold correctly and therefore may not be phosphorylated correctly, resulting in a misleading loss of map spots. Another potential problem is that, if the phosphorylation of the site in question is an ordered event, dependent on another site’s state of phosphorylation, mutation of this other site could lead to the erroneous conclusion that the disappearance of a particular spot is the direct result of the mutation introduced, rather than an event of secondary consequence. Mutagenesis then should be taken as a supporting argument for the presumed identity of a phosphorylation site, rather than as definitive proof. Taken in conjunction with other evidence, it can nevertheless be quite convincing.
Comigration of a synthesized phosphopeptide with a phosphopeptide isolated from a peptide map After all but a few of the candidate peptides have been eliminated, comigration of a synthetic phosphopeptide with the 32P-labeled peptide generated by digesting a protein from labeled cells may provide convincing evidence as to the latter’s identity. Obviously, care must be taken that the synthetic peptide be quite pure (generating only one spot on the cellulose plate)
Scaling up phosphopeptide mapping to isolate peptides for mass spectrometry and microsequencing The most definitive methods for determining the site(s) of phosphorylation in a protein are the determination of the amino acid sequence or mass of the phosphopeptides. While the sensitivity of the instruments used in both these techniques has improved over the past 5 years, both methods require on the order of 1 to 10 pmol of material for analysis. When
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analyzing the phosphorylation of a receptor protein-tyrosine kinase of which ∼10,000 molecules are present on the cell surface, one would need to grow 60 10-cm dishes of cells to isolate the 6 × 1011 molecules (1 pmol) required for a successful analysis. This calculation assumes 100% recovery and 100% stoichiometry of phosphorylation at the sites of interest. However, since it is very possible that not every molecule is phosphorylated at the sites in question, and that one is likely to take at least a 50% loss of material over the entire protocol, it would be best to start with at least 240 dishes of cells. The use of overexpressed protein in cells would obviously facilitate the accumulation of sufficient amounts of protein for analysis. Isolation of enough material for analysis becomes less difficult if an in vitro system can be used to generate the phosphopeptides in question. Protein kinases are notoriously promiscuous in vitro, so it may be possible to generate the sites of interest using a kinase that is actually not the one responsible for the phosphorylation in vivo. The protein of interest produced in bacteria makes a good substrate, as it is most likely not phosphorylated to start with. It is first necessary to show that the phosphopeptides generated by incubation of recombinant protein with a purified kinase in vitro comigrate exactly with those purified from labeled cells. If there is any doubt about this, it is reassuring to run the maps for a longer time than usual in the electrophoresis dimension, or at a different pH to further demonstrate comigration. Given the specificity of the two-dimensional separation on these maps, it is unlikely that two phosphopeptides that truly comigrate will not be identical.
Anticipated Results Upon developing the first autoradiogram of an initial peptide map, depending on the number of spots seen, the researcher will most likely be left wondering what it all means. There are several things to keep in mind when studying the pattern of spots on a peptide map. First, migration in the electrophoresis dimension is a function of the charge to mass ratio of a peptide. Migration in the chromatography dimension is related to the hydrophobicity of a peptide. The more hydrophobic a peptide, the further it migrates in the chromatographic dimension. In general, most phosphopeptides of the protein studied will be represented by one spot in the map. It is possible then to determine the relative stoichiometry of phosphorylation at different
sites by comparing the intensity of each spot on the autoradiogram. This, however, relies on the assumption that all tryptic peptides are recovered during the the entire protocol with similar efficiencies. There are several cases, however, in which this one phosphopeptide/one spot rule does not hold. One such case is when the enzyme used to digest the protein of interest has not worked to completion, yielding both partial and complete digestion products seen in the map. Multiple digestion products are also generated at sites where a run of basic residues is present. Trypsin works very efficiently as an endopeptidase, hydrolyzing peptide bonds following basic residues. In contrast, trypsin works poorly as an exopeptidase. Trypsin will cleave randomly within the run of basic residues and is unable to take off additional basic residues that may have been left. This results in a series of digestion products differing by the number of basic residues at their amino- or carboxy-terminus. Addition of such a basic residue to a phosphopeptide changes its migration in both the electrophoresis and the chromatography dimension (it is more positively charged and runs further towards the cathode; it is also more hydrophilic, and as a consequence runs less far relative to the buffer front in phosphochromatography buffer). Similarly, a given peptide will migrate differently if phosphorylated at two positions rather than one. Addition of another negative charge will again make the peptide more hydrophilic and thus it will migrate less far in the chromatography dimension. This time, however, it will migrate further towards the anode in the electrophoresis dimension, giving a diagonal pattern descending in the opposite direction to that seen with the addition of a lysine or an arginine. It is important to remember that the electrophoretic mobility of a peptide is dependent on its mass. For larger peptides, the slope of the diagonal seen with the addition of either positive or negative charges will be steeper, since the addition of another charge when divided by the mass will make less of a difference to the distance the peptide travels. While trypsin has traditionally been used in peptide mapping, it may not be the enzyme of choice for proteins phosphorylated by PKA, PKC, or other protein kinases whose recognition sequence involves multiple arginines or lysines, as trypsin often fails to cleave after all such residues when they are present in runs. In addition, trypsin cleaves inefficiently at argini-
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Figure 13.9.6 An example of a tryptic phosphopeptide map based on that of human Nck-alpha. For the first (horizontal) dimension, electrophoresis was run at pH 1.9 for 25 min at 1.0 kV; the anode is at the left. Ascending chromatography was run for 15 hr in phosphochromo buffer. The sites represented by spots 1 to 7, with the exception of spot 2, have been identified. Spot 1 is an 11-amino-acid, phosphotyrosine-containing peptide. While spot 2 also contains phosphotyrosine, and runs in a position likely to be the doubly phosphorylated version of this peptide, it turns out to be unrelated to spot 1. Spot 3 represents a 5-amino-acid, phosphoserine-containing peptide; this same peptide with an amino-terminal arginine runs as spot 4; thus it can be seen that in this case the tryptic cleavage is largely incomplete. Spot 5 represents a 20-amino-acid phosphoserine-containing peptide which, with an amino-terminal lysine, runs as spot 6. Spot 7 represents a peptide that is unrelated to 5 and 6. Spot 8 represents free phosphate, released during sample preparation.
Phosphopeptide Mapping and Identification of Phosphorylation Sites
nes or lysines two residues amino-terminal of a phosphoserine or phosphothreonine (i.e., R/K-X-P.Ser). Sometimes an individual peptide may appear to have an electrophoretic partner that migrates directly above or below it in the chromatographic dimension. This sort of pattern may be observed as the result of two different scenarios: (1) it may be the result of incomplete oxidation of the peptide if it contains a methionine residue (in which case the lower spot is the oxidized form), or (2) it may be the result of methylation of the peptide running in the lower position. Such a methylation may occur during the performic acid oxidation and is dependent
on the 1.5-ml microcentrifuge tubes being used (see Critical Parameters). An exemplary tryptic phosphopeptide map based on that of a real protein (Nck) is shown in Figure 13.9.6. This map illustrates the points mentioned above. Perhaps most importantly, it also illustrates the fact that just because two spots appear to be on a diagonal it is not a foregone conclusion that they are related. Although peptides 1 and 2 appear to represent the singly and doubly phosphorylated forms of a single tryptic peptide, in this case it turned out that they represent two completely different peptides. Peptides 3 and 4, and 5 and 6, respectively, represent two sets of peptides that are
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related and differ only by the addition of a basic residue. Spot 8 represents free phosphate, liberated by hydrolysis of phosphoester bonds that has occurred during sample preparation. It is useful both to compare the amount of free phosphate generated in different samples and to use the phosphate spot as another standard marker when comparing peptide mobilities on different plates.
Time Considerations To generate a two-dimensional phosphopeptide map, at least 9 days will elapse from the time the 32P label is added to the cells until the autoradiogram of the map is in hand. The typical researcher, intrigued by one or more particular spots that appear or disappear from such maps depending on how the cells or samples were treated, may rush to attempt to identify the phosphorylation site represented by such spot(s). Please be advised that this will take at least 4 months of hard work and effort, assuming that everything goes well. There are several different strategies to follow, which are outlined throughout the unit (especially see Background Information). Choice of a particular course will depend on the reagents and the equipment available for analysis.
(D.G. Hardie, ed.) pp. 127-151. Oxford University Press, Oxford. Mitchelhill, K.I., Michell, B.J., House, C.M., Stapleton, D., Dyck, J., Gamble, J., Ullrich, C., Witters, L.A., and Kemp, B.E. 1997. Posttranslational modifications of the 5′-AMP-activated protein kinase β1 subunit. J. Biol. Chem. 272:24475-24479. van der Geer, P. and Hunter, T. 1990. Identification of tyrosine 706 in the kinase insert as the major colony-stimulated factor 1 (CSF-1)–stimulated autophosphorylation site in the CSF-1 receptor in a murine macrophage cell line. Mol. Cell. Biol. 10:2991-3002. Wang, Y.K., Liao, P.-C., Allison, J., Gage, D.A., Andrews, P.C., Lubman, D.M., Hanash, S.M., and Strahler, J.R. 1993. Phorbol 12-myristate 13-acetate-induced phosphorylation of op18 in Jurkat T cells. J. Biol. Chem. 268:14269-14277.
Key References Boyle et al., 1991. See above. van der Geer, P., Luo, K. Sefton, B.M., and Hunter, T. 1993. Phosphopeptide mapping and phosphoamino acid analysis on cellulose thin-layer plates. In Protein Phosphorylation; a Practical Approach (D.G. Hardie, ed.) pp. 97-126. IRL Press, Oxford. Both of these papers discuss many of the protocols described in this unit.
Internet Resources Literature Cited
http://www.genestream.org
Boyle, W.J., van der Geer P., and Hunter, T. 1991. Phosphopeptide mapping and phosphoamino acid analysis by two-dimensional separation on thin-layer cellulose plates. Methods Enzymol. 201:110-148.
This Web site contains a program for calculating the mobility of a peptides of known composition and a program that reads the position of a spot on the actual map and calculates which peptide(s) derived from the protein being mapped could have the mobility of that spot.
Fischer, W.H., Karr, D., Jackson, B., Park, M., and Vale, W. 1991. Microsequence analysis of proteins purified by gel electrophoresis. Methods Neurosci. 6:69-84. Fischer, W.H., Hoeger, C.A., Meisenhelder, J. Hunter, T., and Craig, A.G. 1997. Determination of phosphorylation sites in peptides and proteins employing a volatile Edman reagent. J. Protein Chem. 16:329-333. Mitchelhill, K.I. and Kemp, B.E. 1999. Phosphorylation site analysis by mass spectrometry. In Protein Phosphorylation: A Practical Approach
Contributed by Jill Meisenhelder and Tony Hunter The Salk Institute for Biological Studies La Jolla, California Peter van der Geer University of California, San Diego La Jolla, California
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Use of Protein Phosphatase Inhibitors
UNIT 13.10
Reversible protein phosphorylation is recognized as a major mechanism regulating the physiology of plant and animal cells. Virtually every biochemical process within eukaryotic cells is controlled by the covalent modification of key regulatory proteins. This in turn dictates the cellular response to a variety of physiological and environmental stimuli; errors in signals transduced by phosphoproteins contribute to many human diseases. Thus, defining protein phosphorylation events, and specifically, the phosphoproteins involved, is crucial for obtaining a better understanding of the physiological events that distinguish normal and diseased states. In studying protein phosphorylation, two common experimental problems arise that mandate the use of protein phosphatase inhibitors. The first problem arises when the goal of the experiment is to decipher physiological events regulated by reversible protein phosphorylation but the hormonal stimuli or signaling pathways involved are not known. This situation is further complicated by the fact that many protein phosphorylations are rapid and transient, thereby eluding investigation. One solution is to employ cell-permeable compounds that inhibit cellular protein phosphatases in order to amplify the basal activity of protein kinases. Inhibition of protein phosphatases also prevents the turnover of protein-bound phosphate, thereby enhancing the detection of phosphoproteins by standard techniques such as SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11) or phosphorimaging. In other situations, the aim may be to analyze the impact of hormones and other physiological stimuli on the function of a specific phosphoprotein. Such detailed biochemical analysis is only made possible after the protein of interest has been separated from other cellular components that potentially interfere with these studies. In this circumstance, the cells or tissues exposed to hormones must be homogenized in the presence of protein phosphatase inhibitors to preserve the cellular phosphorylation events promoted by the hormones. The goal is to utilize the most effective inhibitors to abolish the activity of protein phosphatases that may dephosphorylate the proteins of interest during the lengthy protein isolation procedures. This unit describes the general or basic protocols for inhibiting the cellular PP1/PP2A activity with okadaic acid (see Basic Protocol 1) as well as in vitro with microcystin-LR (see Basic Protocol 2). Basic Protocols 3 and 4 describe approaches recommended for inhibiting cellular and in vitro activity, respectively, of PP2B/calcineurin. Finally, Basic Protocol 5 outlines a widely utilized strategy for inhibiting protein tyrosine phosphatases. The reader should be aware that these Basic Protocols are deliberately designed to cover the widest range of experimental situations and therefore represent prototypic procedures that can be readily used with many phosphatase inhibitors that are not specifically mentioned in this unit. A variety of very effective inhibitors, particularly those targeting protein serine/threonine phosphatases, are currently commercially available. Some of these compounds are cell-permeable and yet others are better utilized in vitro to suppress phosphatase activity in lysates and/or cell fractions. This unit describes methods for defining physiologically relevant protein phosphorylation events. Thus, in some cases, cells in culture are treated with cell-permeable phosphatase inhibitors to detect and establish the in vivo phosphorylation events. In other cases, the cells are stimulated with known physiological stimuli that modify cellular protein phosphorylation. In either situation, subsequent analyses require that the cells be homogenized (UNIT 4.3) in buffers containing phosphatase inhibitors that preserve the protein modifications through the steps of protein purification and analysis. This in turn facilitates the identification of the particular amino acids modified Contributed by Douglas C. Weiser and Shirish Shenolikar Current Protocols in Protein Science (2003) 13.10.1-13.10.13 Copyright © 2003 by John Wiley & Sons, Inc.
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and sets the stage for more defined molecular approaches, such as site-directed mutagenesis, that will allow the investigator to establish the functional role of individual protein phosphorylation events. PROTEIN SERINE/THREONINE PHOSPHATASES Serines and threonines constitute the major sites of protein phosphorylation in all eukaryotic cells. Varying estimates suggest that even in cells transformed using oncogenic viral protein tyrosine kinases, >97% of the protein-bound phosphate appears on serines and threonines. Thus, it is not surprising that numerous microorganisms produce toxins and natural products that elicit their deleterious effects by inhibiting cellular protein serine/threonine phosphatases (McCluskey et al., 2002). Many of these compounds show some degree of selectivity for subgroups of protein serine/threonine phosphatases and may provide insight into the cellular phosphatases that catalyze protein dephosphorylation events. The specificity of some of these compounds has been determined in vitro as differences in their concentration-response curves for the inhibition of purified protein phosphatases. When used in vivo, the chemical properties of the compounds, many of which are hydrophobic, do not allow accurate estimates of cellular concentration or predict their subcellular distribution. Thus, the in vitro specificity of the phosphatase inhibitors may be lost and other more direct molecular approaches may be required to define the protein phosphatases that regulate specific cellular events. The principal use of phosphatase inhibitors, as described in this unit, is to complement the in vitro use of purified phosphatases (UNIT 13.5) to define the regulatory importance of reversible protein phosphorylation, rather than to identify the physiologically relevant protein phosphatases. Early biochemical studies classified eukaryotic protein serine/threonine phosphatases into two major groups, termed type-1 (PP1) and type-2 protein phosphatases (Shenolikar and Nairn, 1991). The type-2 phosphatases are further separated into three distinct groups, PP2A, PP2B, and PP2C, showing very different profiles of inhibition by the available compounds (Table 13.10.1). Molecular cloning of mammalian phosphatase catalytic subunits further expanded this family of enzymes, which to date includes up to five additional phosphatase catalytic subunits numbered PP3 to PP7. Of these five, all but PP7 share a highly conserved catalytic site, and therefore are similarly inhibited by various compounds. Yet others, such as PP2C or PP7, show complete insensitivity to the known phosphatase inhibitors and there are no currently available tools to inhibit these phosphatases. Table 13.10.1
Commercially Available PP1/PP2A Inhibitorsa
Compound
Phosphatases
Recommended use
Okadaic acid Calyculin A Tautomycin Cantharidin A Microcystin-LR Nodularin Fostreicin
PP2A > PP1 PP1 ≥ PP2A PP1 > PP2A PP2A > PP1 PP1 = PP2A PP1 = PP2A PP2A>>PP1
In vivo In vivo In vivo In vivo In vitro In vitro In vitro
Supplier(s) Alexis Biochemicals, Sigma Alexis Biochemicals, Sigma Biomol Research Laboratories — Alexis Biochemicals, Sigma — A.B. Scientific, Sigma, Alexis Biochemicals
aAll compounds listed in this table also inhibit PP3, PP4, PP5, and PP6. Their shared sensitivity to these compounds
Use of Protein Phosphatase Inhibitors
results in their inclusion in the category of PP2A-like enzymes. In contrast, these compounds show no significant inhibitory activity towards PP2B, PP2C, and PP7. As the compounds possess little structural homology, several different inhibitors may be used to validate the physiological effects of inhibiting their shared targets, serine/threonine phosphatases.
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Table 13.10.2
Compound Cyclosporin Ab FK506 Cypermethrinc Deltamethrin
Commercially Available PP2B (Calcineurin)a Inhibitors
Phosphatases
Recommended use
PP2B PP2B PP2B PP2B
In vivo In vivo In vivo In vitro
Suppliers Alexis Biochemicals, Sigma Alexis Biochemicals Alexis Biochemicals Alexis Biochemicals
aPP2B (calcineurin) is a Ca2+/calmodulin-activated protein serine/threonine phosphatase and, like many Ca2+/calmodulin-activated enzymes, is inhibited by the calcium chelator, EGTA. Inhibition of PP2B activity using
EGTA is not a very viable approach in cells and the compounds listed in this table more effectively suppress cellular PP2B activity. bA number of inactive and partially active compounds related to cyclosporin are available. Rapamycin, which shares common target proteins with FK506, namely FK506-binding proteins or FKBPs, does not inhibit PP2B activity and is used as control for FK506. cCompounds related to cypermethrin, such as resmethrin, show weak or no activity as PP2B inhibitors and can function controls for the pyrethroid inhibitors.
The compounds listed in Table 13.10.1 inhibit several classes of protein serine/threonine phosphatases. Enzymes such as PP3, PP4, PP5, and PP6 share structural homology to PP2A and therefore show similar sensitivity to these compounds. In contrast, PP2B is inhibited by a distinct group of compounds (Table 13.10.2) that show no activity against either PP1 or the PP2A-like phosphatases. PP1/PP2A Classes of Protein Serine/Threonine Phosphatases PP1 and PP2A together account for >90% of protein serine/threonine phosphatase activity in most eukaryotic cells. With the exception of mammalian skeletal muscle, where PP1 predominates, most cells and tissues possess equivalent amounts of PP1 and PP2A. This has made okadaic acid and calyculin A, both potent cell-permeable PP1/PP2A inhibitors, the most widely utilized compounds for studying cellular protein phosphorylation. INHIBITION OF CELLULAR PP1/PP2A ACTIVITY WITH OKADAIC ACID Okadaic acid (OA), a polyether produced by marine dinoflagellates, is concentrated by shellfish and is a common cause of diarrhetic shellfish poisoning in humans. OA is commercially available as a free acid or as a potassium, sodium, or ammonium salt. Prevailing evidence favors the sodium salt of OA as the most readily permeable across cell membranes.
BASIC PROTOCOL 1
Biochemical studies demonstrate that OA inhibits the purified PP2A catalytic subunit with an IC50 value of 0.2 to 1.0 nM (Cohen et al., 1989). This concentration of OA, however, has little effect on PP2A activity in cell lysates, which typically exhibit 30- to 150-fold higher IC50 values than the purified enzyme. Extensive dilution of cell extracts enhances the efficacy of OA as a PP2A inhibitor. This may suggest that OA binds, albeit with low affinity, to other cellular components in cell extracts. This may be more of an issue in intact cells where significantly higher OA concentrations (10 to 100 nM) are required for effective PP2A inhibition. PP1, unlike PP2A, has a higher IC50 value for OA (10 to 100 nM; Cohen et al. 1989), whether assayed with purified enzyme or cell lysates, and the sensitivity of PP1 activity to OA is not modified by dilution of cell lysates. In some cellular studies, OA concentrations between 10 and 50 nM resulted in significant enhancement of protein phosphorylation, presumably reflecting the inhibition of PP2A and other PP2A-like enzymes. In contrast, OA concentrations >10 µM are needed to effectively inhibit cellular PP1 activity.
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This difference in affinity can provide an opportunity to discriminate between the physiological functions of the two major classes of cellular protein serine/threonine phosphatases, namely PP1 and PP2A. Numerous studies suggest that the predominant site of OA accumulation is the lipid bilayer of the plasma membrane (Suganuma et al., 1989; Nam et al., 1990; Namboodiripad and Jennings, 1996). OA transfers slowly from this site to other compartments and this may account for the slow onset of phosphatase inhibition by OA in most cells (up to 90 min) as well as the requirement of 10- to 100-fold higher concentrations of OA to inhibit PP2A activity in intact cells compared to cell lysates. Thus, it is surprising to find that OA can also be washed out of cells, albeit thorough washing can take hours. This reversible behavior may allow investigators to analyze the reversal of phosphorylation events triggered by OA. Duration of OA treatment to elicit maximal changes in cellular protein phosphorylation is largely empirically determined and must be defined for each individual cell type. In general, cells display a maximal response to micromolar concentrations of OA within 10 to 30 min. Lower OA concentrations require significantly longer exposure times (up to 2 hr). Due to their remarkable ability to activate multiple cellular signaling pathways, OA and other cell-permeable phosphatase inhibitors display some cytotoxicity. This is partially alleviated by lowering OA concentrations and reducing exposure times. Thus, OA treatments for several hours or days are not recommended. If longer times are necessary, calyculin A (CA), which inhibits PP1 and PP2A with near equal potency, may be a better substitute. While the literature on calyculin A is less extensive, this compound can be used at much lower concentrations (0.1 to 1.0 µM) than OA to yield similar physiological effects. OA induces errors in DNA repair that are not mediated by phosphatase inhibition, (Nakagama et al., 1997) which suggests the presence of additional targets for OA. This behavior mandates that appropriate controls be used in cellular studies. In this regard, the treatment of cells with vehicle (DMSO) is commonly used as a control for OA, CA, and other phosphatase inhibitors that are dissolved in this solvent. There are, however, a number of commercially available inactive OA analogues that may serve as better controls. The best of these is 1-norokadaone, whose structure closely matches that of OA and maintains some of the non-phosphatase-mediated effects elicited by OA. Structurefunction studies that established a critical role for the acidic side-chain of OA in phosphatase inhibition also generated a number of inactive OA analogues in which the critical carboxylate group is esterified (Nashiwaki et al., 1991). Such compounds, like methyl okadaate, show low or no activity in vitro as PP1/PP2A inhibitors. However, these compounds may not be effective negative controls, as they are slowly de-esterified in cells, yielding the active OA. Finally, OA, CA, and several other phosphatase inhibitors show significant activity as tumor promoters in mouse skin and gut (Fujuki et al., 1988; Suganuma et al., 1990). Thus, it is important that the investigator takes extreme care in handling and disposing of solutions containing these compounds.
Use of Protein Phosphatase Inhibitors
Materials Cultured mammalian cells grown to <50% confluence (APPENDIX 3C) 0.1 or 1.0 mM okadaic acid sodium salt (Alexis Biochemicals or Sigma), in DMSO (store at −20°C in a light-proof container under N2) 0.1 or 1.0 mM 1-norokadaone (Calbiochem or Sigma) in DMSO (store at −20°C in a light-proof container under N2)
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NOTE: Okadaic acid (OA), either as sodium, potassium, or ammonium salt, shows increased solubility in aqueous solutions compared to the free acid. The solutions of the OA salts are also more stable during storage. However, for reasons that are not entirely clear, aqueous solutions of OA lose their activity as phosphatase inhibitors over time. By comparison, the stock solutions of OA in DMSO (or ethanol or methanol) can be stored in a dark glass container under nitrogen at −20°C for many months without significant loss of activity. 1. Prepare cultured mammalian cells to ∼50% confluence (APPENDIX 3C). Okadaic acid activates a variety of cell signaling pathways and physiological events in mammalian cells (Schonthal, 1998). Thus, responses of different mammalian cells to OA vary widely. In general, changes in protein phosphorylation in quiescent or confluent cells induced by OA are less profound. Moreover, quiescent cells often initiate apoptosis following OA exposure (Kiguchi et al., 1994; Morimoto et al., 1997). In contrast, programmed cell death appears less of a problem when analyzing actively growing cultured cells.
2. Add okadaic acid (final concentration 10 µM) to the culture medium or exchange with fresh medium containing 10 µM okadaic acid. Expose control cells to medium containing vehicle (DMSO) or the inactive analogue, 1-norokadaone. Incubate cells for 30 min prior to harvesting (APPENDIX 3C) for analysis of physiological functions (Schonthal, 1998) and/or phosphoproteins (Chapter 13). The maximum (final) concentration of DMSO in the assay should not exceed 1.0% (v/v). Inactive analogues, such as 1-norokadaone, should also be analyzed at a final concentration of 10 ìM.
INHIBITION OF PP1/PP2A ACTIVITY IN VITRO WITH MICROCYSTIN-LR Microcystin-LR, a cyclic heptapeptide produced by the blooms of cyanobacteria in freshwater lakes, is a member of a family of cyclic peptide products that inhibit PP1 and PP2A at nanomolar or subnanomolar concentrations. Many members of this family, including the cyclic pentapeptides, nodularins, show similar phosphatase target specificity. Liver cells possess a unique uptake mechanism for these compounds. Hence, microcystins are potently hepatotoxic but in general, microcystins do not enter most tissues and cells.
BASIC PROTOCOL 2
The broad specificity of microcystin-LR for PP1, PP2A, and other PP2A-like phosphatases as well as its remarkable potency and stability in buffered solution (compared to OA or CA) makes this compound an ideal reagent for preservation of cellular protein phosphorylations in lysates and during lengthy chromatographic purification of phosphoproteins. Microcystin-LR is also significantly less expensive than CA or OA and thus well suited for inclusion in large volumes of buffers used for dialysis and protein chromatography. Moreover, in contrast to OA and CA, no special precautions are needed in the storage of microcystin-LR, although prolonged storage of stock solutions should be at −20°C. It is, however, necessary to take some steps to avoid ingestion of this compound as toxicological studies show that very small quantities of microcystins can lead to serious liver failure and extensive tissue damage. X-ray crystallography defined the binding site for microcystin-LR in the catalytic center of PP1α. Other studies suggest that microcystin-LR forms a stable adduct with a highly conserved cysteine adjacent to the PP1 catalytic site. This process is spontaneous and slow and plays no role in phosphatase inhibition. However, the possibility remains that prolonged exposure (days to weeks) of phosphatases to microcystins generates the enzyme-toxin adduct and limits subsequent studies. For example, following short-term
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exposure to microcystins, the observed phosphatase inhibition is readily reversed by dilution or dialysis. Significant adduct formation may prevent enzyme reactivation and reversal of covalent modifications. Materials Tissue or cells 0.1 or 1.0 mM microcystin-LR (Alexis Biochemicals or Sigma) in DMSO (store at −20°C) 1. Homogenize (UNIT 4.3) tissue or cells in buffers containing 1 µM microcystin-LR and process samples for functional or phosphoprotein analyses (Chapter 13). BASIC PROTOCOL 3
INHIBITION OF CELLULAR PP2B/CALCINEURIN ACTIVITY WITH CYCLOSPORIN A PP2B, also known as calcineurin, is a Ca2+/calmodulin-activated protein serine/threonine phosphatase. PP2B was thus the first phosphatase determined to be directly controlled by a second messenger. PP2B is the therapeutic target for the two major immunosuppressants, cyclosporin A and FK506. These drugs, however, do not directly interact with the PP2B catalytic subunit but function through intermediate protein receptors collectively known as immunophilins to inhibit enzyme activity. Distinct immunophilins bind cyclosporin A and FK506. However, both drug/receptor complexes are effective PP2B inhibitors. The unique activity of these compounds on T-lymphocytes is due to their ready access to T-cells, the low abundance of their target, PP2B, in T-lymphocytes, and the key role of PP2B in activating cytokine genes that promote T-cell proliferation. PP2B and the immunophilins are, however, widely expressed in mammalian tissues and PP2B also plays an important role in regulating the physiology of mammalian heart, brain, and other tissues. While the reduced access within various tissues may limit the utility of these compounds in animals, both cyclosporin A and FK506 are excellent inhibitors of PP2B functions in experimental systems such as tissue slices and cultured cells. Cyclosporin A has an apparent IC50 value for PP2B inhibition ranging from 10 to 100 nM. The use of cyclosporin A is limited by its poor solubility in aqueous solutions and its slow entry into mammalian cells. In contrast, FK506 is a ten-fold more potent PP2B inhibitor and enters cells rapidly. However, commercial availability of FK506 is still restricted, therefore, this protocol focuses on cyclosporin A, which is widely available from many different commercial sources. There are also numerous inactive analogues of cyclosporin available for use as controls. Many of these inactive analogues bind effectively to the target cyclophilin but fail to inhibit PP2B activity. In contrast, another immunosuppressant, rapamycin, binds to the same immunophilin as FK506 but does not inhibit PP2B and can be used as a control for FK506. Rapamycin is a potent inhibitor of the signaling pathway controlled by the protein kinase, TOR (target of rapamycin), and therefore, has its own effects on cellular protein phosphorylation. Materials 0.1 mM cyclosporin A (Alexis Biochemical or Sigma) in DMSO Cultured cells 1. Add 0.1 mM cyclosporin A (final concentration 0.1 to 1 µM) to culture medium or exchange with fresh medium containing cyclosporin A. Incubate cells for 60 min prior to harvesting for analysis.
Use of Protein Phosphatase Inhibitors
The slow entry of cyclosporin A into cells often requires incubating cells and tissue slices for periods >1 hr, as very few physiological events respond to this drug in periods under 30 min.
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IN VITRO INHIBITION OF PP2B/CALCINEURIN ACTIVITY WITH CYPERMETHRIN
BASIC PROTOCOL 4
Cypermethrin and other type II pyrethroids are synthetic insecticides that show activity as PP2B inhibitors and are available from many commercial sources. In addition, a number of pyrethroid analogues, resmethrin, allethrin, and permethrin, show reduced or no activity as PP2B inhibitors and can be used as controls. This is particularly important for cellular studies using cypermethrin, as many of the type-II pyrethroids have a wide range of physiological effects including oxidative stress, cytotoxicity, and tumorigenesis. Only a few of these are likely to be mediated by PP2B inhibition. Given the availability of cyclosporin A, a better characterized and more potent PP2B inhibitor, the value of using pyrethroids in mammalian cells is questionable. Materials 1.0 mM cypermethrin (Alexis Biochemicals) in DMSO and store in dark glass containers pretreated with 1% polyethylene glycol (PEG) Sample cell extracts or subcellular fractions NOTE: Cypermethrin and other type II pyrethroids are sold in both solid and liquid form. The liquid form represents cypermethrin dissolved in organic solvents, such as DMSO, acetone, or ethanol. Hydrophobicity of these compounds means that long-term storage in plastic or polyethylene containers should be avoided because these compounds adhere to such surfaces and may be slowly extracted from solution. Many pyrethroids are also light-sensitive. If stored in dark glass containers pretreated with 1% (w/v) PEG to prevent surface adherence, stock solutions of cypermethrin are stable for many months. For cellular studies using cypermethrin, the compound should be added directly to the culture medium as its PP2B inhibitory activity is reduced when made up in fresh medium. While the underlying basis for a time-dependent loss of cypermethrin activity is not clear, this provides a strong argument for using cypermethrin and the pyrethroid inhibitors solely for in vitro studies of PP2B function. 1. Add 1.0 mM cypermethrin (10 to 20 µM final concentration) to cell extracts or subcellular fractions and analyze phosphoproteins within a period not exceeding 60 min. Cypermethrin addition in vitro should result in rapid PP2B inhibition that is evident within minutes. Prolonged incubations with this compound are not recommended.
INHIBITION OF PROTEIN TYROSINE PHOSPHATASES WITH SODIUM ORTHOVANADATE
BASIC PROTOCOL 5
Protein tyrosine phosphatases (PTPases) represent a larger family of enzymes derived from a common ancestral gene and share no structural homology with the protein serine/threonine phosphatases (UNIT 13.1). Thus, PTPases are not sensitive to any of the compounds described above. PTPases share a common catalytic mechanism that involves the formation of an enzyme phospho-intermediate with the modification of a conserved cysteine within the catalytic cleft. While cysteine-modifying reagents, such as phenyl arsene oxide, and metals such as zinc chloride, potently inhibit PTPases, these reagents are non-specific and likely target many proteins other than PTPases and therefore, will not be further discussed here. Currently, the most useful reagent for inhibiting PTPases in vivo and in vitro is vanadate, a phosphate analogue. One might predict that vanadate would also be non-specific and inhibit many phosphotransferases and phosphohydrolases, including protein serine/threonine phosphatases. However, the remarkable chemistry of the vanadium metal
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ion (Gordon, 1991) can be exploited to produce a preferential and effective PTPase inhibitor. Materials Sodium orthovanadate (Sigma) 0.1 M NaOH Sample cells Boiling water bath or heating block Additional reagents and equipment for metabolic labeling (UNIT 13.2) or immunoblotting with an anti-phosphotyrosine antibody (UNIT 13.6) 1. Dissolve sodium orthovanadate (200 mM final concentration) in sterile, distilled water (often a yellowish solution) and adjust pH of the solution to 10.0 with 0.1 NaOH. Place solution in a boiling water bath until the solution clarifies or becomes translucent. Readjust pH to 10.0 and store the stock solution in a flint glass container at room temperature or dispense into aliquots in plastic containers and store at −20°C. Sodium orthovanadate, whether solid or in solution at neutral pH, slowly oxidizes and acquires a yellow color. The presence of thiols, often used to maintain protein structure and function following the homogenization of tissues or cells, promotes a rapid (<30 min) conversion to the oxycation, vanadyl. Finally, vanadate, particularly in solutions below 0.1 mM at neutral pH, also undergoes polymerization, which is also associated with increased yellow color. All of these factors result in reduced vanadate activity as a PTPase inhibitor. While vanadate slowly reverts to a colorless active monovalent form, especially on dilution, this process is greatly accelerated by increasing pH and temperature, as described above.
2. Incubate cells with 0.1 mM orthovanadate for periods not to exceed 2 to 3 hr before analyzing tyrosine-phosphorylated proteins by metabolic labeling (UNIT 13.2) or immunoblotting with an anti-phosphotyrosine antibody (UNIT 13.6). Maximal inhibition of cellular PTPase activity often requires up to 1.0 mM vanadate. This concentration of vanadate is also chosen to preserve protein tyrosine phosphorylation following cell disruption. However, much lower concentrations of vanadate are recommended for the treatment of intact cells to limit or avoid cytotoxicity. Even 0.1 mM vanadate is detrimental to cell function if the duration of treatment extends for several hours or days. This, however, makes the complete inhibition of PTPases achieved in cell lysates and other fractions by 1.0 mM vanadate easier to interpret than the results of cellular studies where changes in cell function cannot always be attributed to PTPase inhibition. Finally, some protein serine/threonine phosphatases display low but measurable PTPase activity in vitro and are not inhibited by 1.0 mM vanadate. These enzymes may account for slow dephosphorylation of tyrosine-phosphorylated proteins during lengthy isolation procedures.
COMMENTARY Background Information
Use of Protein Phosphatase Inhibitors
In contrast to the large number (300 to 500) of genes encoding mammalian protein kinases, many fewer genes encode the catalytic subunits of protein serine/threonine phosphatases (Cohen, 1997). Thus, the functional diversity of protein serine/threonine phosphatases arises through other mechanisms that include a rapidly expanding number of regulatory subunits that may dictate the substrate specificity and subcellular localization of the phosphatase catalytic subunits (Cohen, 2002). This means that the approaches outlined in this unit for
inhibiting protein serine/threonine phosphatases, while highly effective, may not distinguish between the functions of various intracellular pools of these enzymes. In addition, many phosphatase inhibitors are highly hydrophobic and may display significant differences in accessing intracellular pools of phosphatases. Hence, these protocols are most appropriately used for detecting and defining the physiological role of the phosphorylated protein substrate as opposed to making conclusions regarding the types or pools of phosphatases involved.
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While more genes encode protein tyrosine phosphatases than serine/threonine phosphatases, these too fall far short of the numbers representing the complimentary protein tyrosine kinases. Mammalian PTPases can be separated into two broad groups, transmembrane “ receptor” PTPases and soluble PTPases (Fischer et al., 1991). Many transmembrane PTPases contain two catalytic domains and emerging studies suggest that only one of these domains plays an active role in enzyme catalysis. The “ inactive” catalytic domain is thought to regulate the function of receptor PTPases. It is unclear whether the second catalytic site functions as a decoy for PTPase inhibitors or if other mechanisms account for the apparent reduced sensitivity of transmembrane PTPases to vanadate. Considerable anecdotal evidence points to the preferential inhibition of soluble PTPases in cells treated with vanadate. While the protocols discussed in this unit focus on a few pharmacological reagents, more potent and selective phosphatase inhibitors are likely to become available to investigators in the near future. The driving force behind the development of new phosphatase inhibitors is the desire by pharmaceutical companies to provide new therapies for human diseases associated with defects in signal transduction. The proof-of-principle for phosphatase inhibitors as potential medicines has already been established by the two PP2B/calcineurin inhibitors, cyclosporin A and FK506, which are the most effective immunosuppressants currently available. However, both compounds function by an indirect mechanism (i.e., via cellular receptors or immunophilins) to inhibit PP2B activity and both cause side-effects or toxicity in humans. Thus, there is a continued need for novel PP2B/calcineurin inhibitors as safe and improved immunosuppressants. With the availability of three-dimensional structures for PP2B/calcineurin and other phosphatases with bound inhibitors, improvements in structureactivity profiles for phosphatase inhibitors are inevitable. The current paucity of reagents to inhibit protein tyrosine phosphatases (PTPases) should change very rapidly in the coming years as many pharmaceutical companies are currently embarked on the search for PTPase inhibitors (Shenolikar and Brautigan, 2000). With growing evidence that a specific PTPase, PTP1B, antagonizes insulin signaling in many mammalian tissues, several compounds have been identified that selectively target this phosphatase (Goldstein, 2001). Whether these com-
pounds eventually emerge as therapies for diabetes is uncertain, but they will undoubtedly yield excellent tools for the investigation of signaling pathways regulated by PTP1B and other PTPases. Despite having a common catalytic mechanism, some phosphatases have been defined as “ dual-specificity” in that they dephosphorylate adjacent phosphotyrosines and phosphothreonines in their target substrates, many of which control gene expression and cell proliferation. Perhaps due to differences in the architecture of their catalytic centers, dualspecificity phosphatases also display a decreased sensitivity to vanadate. Recent studies show that some members of the PTPase family do not act on phosphoproteins but rather they dephosphorylate phospholipids (Maehama et al., 2001). One example of this is PTEN, a tumor-suppressor gene that is mutated in many human cancers. PTEN functions as a lipid phosphatase that antagonizes insulin signaling. Development of selective PTEN inhibitors may also yield new therapies for diabetes. The ability of vanadate to inhibit both PTP1B and PTEN may account for its ability to elicit physiological effects similar to insulin and other growth factors. The focus of this unit on a few selected protein serine/threonine phosphatase inhibitors should not minimize the value of a multitude of other protein serine/threonine phosphatase inhibitors currently available to researchers. Several hundred natural products appear to elicit their effects on mammalian cell physiology by inhibiting phosphatases. Such compounds include antibiotics such as tautomycin, insecticides such as cantharidins, and other structurally diverse compounds (McCluskey et al., 2002). Among these, one particular compound bears special mention, fostreicin, produced by a strain of Streptomyces, may be the most specific currently available inhibitor of the PP2A-like enzymes. Fostreicin demonstrates an IC50 value for PP2A of 1 to 3 nM while >100 µM fostreicin is needed to inhibit the structurally related serine/threonine phosphatase, PP1. Use of fostreicin is, however, limited by its propensity to be inactivated by oxidation and thus must be stored in the presence of a large excess of antioxidants. Commercially available fostreicin, therefore, contains numerous additives, making it very difficult to use as a specific reagent in cellular studies. Nevertheless, it is likely that selective PP1 and PP2A inhibitors can and will be developed (see Key References). Compared to most
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current serine/threonine phosphatase inhibitors, which display cytotoxicity and tumor promoter activity, clinical trials established fostreicin as a potential anti-neoplastic drug. This also hints at the possibility that more selective PP1 or PP2A inhibitors may be better experimental tools than currently available compounds whose cytotoxicity may be a result of inhibiting many different protein serine/threonine phosphatases. On the other hand, it is the remarkable chemistry of the vanadium metal ion that makes the current compounds such effective tools for analyzing highly labile and difficult-to-detect cellular phosphorylation events. Finally, a number of broad-acting or nonspecific reagents have been used to inhibit protein phosphatases. These include sodium fluoride (50 mM) and sodium pyrophosphate or glycerophosphate (10 mM). These compounds suppress protein dephosphorylations in cell lysates and fractions, although not as effectively as other inhibitors. They can serve as alternatives for inhibiting phosphatases in vitro but show no utility in phosphoprotein analysis in intact cells.
Critical Parameters
Use of Protein Phosphatase Inhibitors
Treatment of cells with the cell-permeable phosphatase inhibitors described in this unit is often so effective that it saturates and occludes the physiological signaling pathways, making studies that utilize combinations of hormones and phosphatase inhibitors difficult to interpret. Many mammalian proteins are regulated both positively and negatively by phosphorylation, generally occurring at distinct sites. Thus, the treatment of cells with phosphatase inhibitors may facilitate both types of modifications and yield a compound functional readout, which may bear little or no relationship to the physiological mechanisms that orchestrate the orderly control of protein function. Even when utilizing phosphatase inhibitors in vitro, artifactual or post-homogenization phosphorylations can result from inappropriate exposure of proteins to kinases normally located in distinct subcellular compartments and therefore are not normally physiological regulators of these proteins. Thus, this unit stresses the detection of cellular protein phosphorylation and events bearing on the functional role of such modifications. The pharmacological approach to studying cellular protein phosphorylation as described in this unit can be employed by investigators from many backgrounds. However, further op-
timization of the outlined experimental strategy may be necessary for individual cell types, where permeability of the compounds and the array of phosphatases inhibited may dictate unique physiological consequences. Thus, the investigator is encouraged to undertake pilot studies that determine the optimal concentrations of the compound and the duration of cell exposure required to obtain a measurable and meaningful cellular response. Also, as emphasized above, while the cell-permeable phosphatase inhibitors display some target specificity, they cannot unequivocally define the phosphatases that regulate specific phosphoproteins or physiological events. Finally, the investigator should be aware that many of the compounds described above display some capacity to bind proteins other than phosphatases albeit with much lower affinity. Thus, control experiments should utilize compounds, such as the inactive okadaic acid or cyclosporin analogues, that fail to inhibit the target phosphatases. These controls should provide a more compelling argument supporting the biological effects uniquely observed with the active compounds as reflecting the inhibition of the target protein phosphatases. To define the specific phosphatases, the investigator must turn to more molecular approaches that may involve the overexpression of phosphatases (currently a very challenging task), or the endogenous protein inhibitors that show exquisite selectivity for individual phosphatases (Oliver and Shenolikar, 1998). An alternate to overexpression may be the microinjection of phosphatases and the inhibitor proteins, an approach that has been successful in modulating cellular phosphatase activity (Alberts et al., 1993; Mulkey et al., 1994; Morishita et al., 2001). The difficulty of introducing protein inhibitors into cells, whether by overexpression or microinjection, is that most, if not all, of these proteins are tightly regulated by covalent modification. The functions of various inhibitors may be enhanced or dampened by reversible protein phosphorylation, making it difficult to predict their effects on phosphatases. Yet another approach is the use of antisense oligonucleotides or interference RNAs to “ knock-down” expression of specific protein phosphatases. While this approach harbors tremendous specificity, it is difficult to control the effectiveness of these inhibitors which lower phosphatase expression, and elicit appropriate physiological responses that cannot be readily predicted. The use of dominantnegative reagents, including inactive phos-
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phatase catalytic subunits or fragments of phosphatases, has shown some promise in suppressing phosphatase functions, but these reagents most likely rely on their competition with endogenous phosphatases for regulators rather than substrates, thereby making it difficult to predict the physiological outcome. However, catalytically inactive phosphatases have proven very useful in deciphering the functions of PTPases. These inactive phosphatases function as substrate traps and form stable complexes with cellular phosphoproteins, allowing identification of the potential physiological substrates of the PTPase. Even in this situation, there are serious concerns that overexpression of inactive PTPases may distort the natural specificity of these enzymes and yield spurious or unreliable data. Protein serine/threonine phosphatases themselves also occasionally form stable complexes with cellular substrates and such complexes can be isolated using affinity chromatography based on phosphatase inhibitors, such as microcystin-LR, immobilized on Sepharose or other dextran surfaces. Such affinity matrices are not commercially available but a number of publications describe the synthesis and use of such resins. The reader may refer to Campos et al. (1996) or Damer et al. (1998). However, this approach has thus far been more successful in identifying phosphatase regulators than substrates.
Troubleshooting The use of phosphatase inhibitors as described in this unit is associated with several problematic issues regarding compound stability and permeability. First and foremost, many of these compounds are highly hydrophobic. Some are light-sensitive and others difficult to store for long periods. These factors contribute to the problem of defining their precise concentration in culture media and buffers and predicting their effects on target phosphatases. In cellular studies, there are additional problems in that external inhibitor concentration is not an accurate predictor of the intracellular concentration of compounds, which are not evenly distributed throughout the cell. Thus, the literature reports widely varying IC50 values for inhibition of purified phosphatases and the sensitivity of known PP1 and/or PP2A-regulated events in cells. Thus, the protocols described in this unit deliberately err on the side of using excess phosphatase inhibitors. While this ensures effective phosphatase inhibition, additional problems may arise in terms of the cytotoxicity elicited by these reagents, which varies
widely among different cell types. Quiescent, confluent and highly differentiated cells appear more susceptible to programmed cell death in response to phosphatase inhibitors such as OA and CA, although the prolonged overactivation of multiple signaling pathways by these inhibitors may also contribute to the cytotoxicity observed in actively growing cells. Thus, investigators are encouraged to determine the lowest concentrations of inhibitors and shortest duration of cell exposure necessary for a consistent and measurable physiological response.
Anticipated Results Phosphatase inhibitors that inhibit the major mammalian protein serine/threonine or tyrosine phosphatases unmask an array of kinase cascades and thereby enhance the phosphorylation of a wide range of proteins. This in turn permits the investigator to detect and analyze novel phosphorylation events. By selecting specific phosphatase inhibitors and restricting their concentrations, the investigator can also inhibit a subset of cellular phosphatases yielding more defined results by not only limiting the panel of phosphoproteins visualized but also the sites modified on the given phosphoprotein. For an example of a specific phosphatase inhibition experiment, refer to Haystead et al. (1989). Functional effects associated with the rapid phosphorylation-dephosphorylation of cellular proteins are more readily visualized in the presence of phosphatase inhibitors. These experiments can establish the importance of protein phosphorylation in physiological events where the precise protein target or phosphorylation event has not been identified. However, readouts such as gene expression that result from increased phosphorylation of nuclear proteins often require long exposure of cells to phosphatase inhibitors and thereby risk the potentially confounding effects of cytotoxicity. Here, a compromise must be reached and inhibitor concentrations lowered to levels that maintain cell viability and still promote the transcription activation of target genes.
Time Considerations A common scenario for the experiments described above is that cells are metabolically labeled using 32P-orthophosphate for periods between 90 and 120 min (UNIT 13.2). Phosphatase inhibitors are then added to the media and the cells incubated for a further 30 min. Following homogenization, the phosphoproteins are analyzed using SDS-PAGE (UNIT 10.1) and autora-
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diography or phosphorimaging, which adds an additional 2 to 3 hr. Thus, the total time required may be minimally estimated at 6 hr. In the absence of metabolic labeling, SDSPAGE may be followed by electrophoretic transfer and analysis of changes in protein mobility reflecting increased phosphorylation or the modification of specific proteins (UNIT 13.9). Phosphorylation sites may be identified by western immunoblotting using phosphospecific antibodies (UNIT 13.4). These approaches provide different pieces of information but carry similar total time considerations.
Literature Cited Alberts, A.S., Thorburn, A.M., Shenolikar, S., Mumby, M.C., and Feramisco, J.R. 1993. Regulation of cell cycle progression and nuclear affinity of the retinoblastoma protein by protein phosphatases. Proc. Natl. Acad. Sci. U.S.A. 90:388392. Campos, M., Fadden, P., Alms, G., Qian, Z., and Haystead, T.A. 1996. Identification of protein phosphatase-1-binding proteins by microcystinbiotin affinity chromatography. J. Biol. Chem. 271:28478-28484. Cohen, P.T.W. 1997. Novel protein serine/threonine phosphatases: Variety is the spice of life. Trends Biochem. Sci. 22:245-251. Cohen, P.T. 2002. Protein phosphatase 1- targeted in many directions. J. Cell Sci. 115:241-256. Cohen, P., Klumpp, S., and Schelling, D.L. 1989. An improved procedure for identifying and quantitating protein phosphatases in mammalian tissues. FEBS Lett. 250:596-600. Damer, C.K., Partridge, J., Pearson, W.R., and Haystead, T.A. 1998. Rapid identification of protein phosphatase 1-binding proteins by mixed peptide sequencing and data base searching. Characterization of a novel holoenzymic form of protein phosphatase 1. J. Biol. Chem. 273:2439624405.
Use of Protein Phosphatase Inhibitors
Haystead, T.A., Sim, A.T., Carling, D., Honnor, R.C., Tsukitani, Y., Cohen, P., and Hardie, D.G. 1989. Effects of the tumour promoter okadaic acid on intracellular protein phosphorylation and metabolism. Nature 337:78-81. Kiguchi, K., Glesne, D., Chubb, C.H., Fujiki, H., and Huberman, E. 1994. Differential induction of apoptosis in human breast tumor cells by okadaic acid and related inhibitors of protein phosphatases 1 and 2A. Cell Growth & Differentiation 5: 995-1004. Maehama, T., Taylor, G.S., and Dixon, J.E. 2001. PTEN and myotubularin: Novel phosphoinositide phosphatases. Annu. Rev. Biochem. 70:24779. McCluskey, A., Sim, A.T.R., and Sakoff, J.A. 2002. Serine-threonine protein phosphatase inhibitors: Development of potential therapeutic strategies. J. Medicinal Chem. 45:1151-1175. Morimoto, Y., Ohba, T., Kobayashi, S., and Haneli, T. 1997. The protein phosphatase inhibitors okadaic acid and calyculin A induce apoptosis in human osteoblastic cells. Exp. Cell Res. 230:181-186. Morishita, W., Connor, J.H., Xia, H., Quinlan, E.M., Shenolikar, S., and Malenka, R.C. 2001. Regulation of synaptic strength by protein phosphatase 1. Neuron 32:1133-1148. Mulkey, R.M., Endo, S., Shenolikar, S., and Malenka, R.C. 1994. Involvement of a calcineurin/inhibitor-1 phosphatase cascade in hippocampal long-term depression. Nature 369:486-488. Nakagama, H., Kaneko, S., Shima, H., Fukuda, H., Kominami, R., Sugimura, T., and Nagao, M. 1997. Induction of minisatellite mutation in NIH3T3 cells by treatment with the tumor promoter okadaic acid. Proc. Natl. Acad. Sci. U.S.A. 94:10813-10816. Nam, K.Y., Hiro, M., Kimura, S., Fujiki, H., and Imanishi, Y. 1990. Permeability of a non-TPAtype tumor promoter, okadaic acid, through lipid bilayer memebrane. Carcinogenesis 11:11711174.
Fischer, E.H., Charbonneau, H., and Tonks, N.K. 1991. Protein tyrosine phosphatases: A diverse family of intracellular and transmembrane enzymes. Science 253:401-406.
Namboodiripad, A.N. and Jennings, M.L. 1996. Permeability characteristics of erythrocyte membrane to okadaic acid and calyculin A. Am. J. Physiol. 270:C449- C456.
Fujuki, H., Suganuma, M., Suguri, H., Yoshizawa, S., Takagi, K., Uda, N., Wakanatsu, K., Yamada, K., Murata, M., and Yasumoto, T. 1988. Diarrhetic shellfish toxin, dinophysistoxin-1, is a potent tumor promoter on mouse skin. Jap. J. Cancer Res. 79:1089-1093.
Nashiwaki, S., Fujiki, H., Suganuma, M., FuruyaSuguri, H., Matsushima, R., Iida, Y., Ojika, M., Yamada, K., Uemura, D., and Yasumoto, T. 1991. Structure-activity relationship within a series of okadaic ac id derivatives. Carcinogenesis 11:1837-1841.
Goldstein, B.J. 2001. Protein-tyrosine phosphatase 1B (PTP1B): A novel therapeutic target for type 2 diabetes mellitus, obesity and related states of insulin resistance. Curr. Drug Target–Immun. Endocrin. Metab. Disorders 1:265-275.
Oliver, C.J. and Shenolikar, S. 1998. Physiologic importance of protein phosphatase inhibitors. Frontiers in Biosci. 3:D961-D972.
Gordon, J.A. 1991. Use of vanadate as protein-phosphotyrosine phosphatase inhibitor. Methods Enzymol. 201:477-482.
Schonthal, A.H. 1998. Analyzing gene expression with the use of serine/threonine phosphatase inhibitors. Methods Mol. Biol. 93:35-40. Shenolikar, S. and Nairn, A.C. 1991. Protein phosphatases: Recent progress. Adv. Second Messenger Phosphoprotein Res. 23:1-121.
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Shenolikar, S. and Brautigan, D.L. 2000. Targeting protein phosphatases: Medicines for the new millenium. Science (STKE) Nov. 2 000. http://stke.sciencemag.org/. Suganuma, M., Suttajit, M., Suguri, H., Ojika, M., Yamada, K., and Fujiki, H. 1989. Specific binding of okadaic acid, a new tumor promoter in mouse skin. FEBS Lett. 250:615-618. Suganuma, M., Fujiki, H., Furuya-Suguri, H., Yoshizawa, S., Yasumoto, S., Kato, Y., Fusetani, N., and Sugimura, T. 1990. Calyculin A, an inhibitor of protein phosphatases, is a potent tumor promoter on CD-1 mouse skin. Cancer Res. 15:3521-3525.
Key References Current Medicinal Chemistry. November, 2002, vol. 9. A special “ hot topics” issue, which is devoted to the synthesis of serine/threonine phosphatase inhibitors.
Contributed by Douglas C. Weiser and Shirish Shenolikar Duke University Medical Center Durham, North Carolina
Post-Translational Modification: Phosphorylation and Phosphatases
13.10.13 Current Protocols in Protein Science
Supplement 31
CHAPTER 14 Post-Translational Modification: Specialized Applications INTRODUCTION
T
he amino acid sequence of a protein determines its structural and functional properties. Yet the primary structure of a polypeptide is not a complete description of its chemical constituents; modifications of the polypeptide backbone may alter the intracellular destination, as well as modify functional properties of the polypeptide in question. Some of these modifications occur as the polypeptide rolls off the ribosome; others take place on complete polypeptide chains; or both co- and post-translational modification may occur on the same polypeptide, as exemplified by protein-bound carbohydrates. A number of post-translational modifications may be inferred from the primary structure or from the open reading frame established by DNA sequencing. These include potential cleavage sites for signal peptidase in the case of secretory or type-I membrane proteins; cleavage sites for other proteases that process inactive precursors into active polypeptides, as exemplified by neuropeptide precursors; consensus sequences for N-linked glycosylation; and C-terminal modification by isoprenoid moieties in conjunction with carboxymethylation. Similarly, consensus sequences for certain types of phosphorylation events have been established. Although for many of these modifications consensus sequences indeed indicate their possible occurrence, it is not always safe to conclude that such modifications therefore necessarily take place. For that reason, methods to identify and characterize these modifications are essential. All of the naturally occurring amino acids can occur in proteins in a modified form. In this manual, separate chapters are devoted to the most intensively and possibly widespread protein modifications, glycosylation (Chapter 12) and phosphorylation (Chapter 13). In this chapter, methods for the study of other protein modifications will be addressed.
The presence of disulfide bonds stabilizes the structure of proteins and allows them to survive in the extracellular milieu. This type of modification cannot be inferred from primary structure without the aid of homologous proteins for which the disulfide bonding pattern has been established by chemical analysis, as described elsewhere in this manual (UNIT 7.3). Furthermore, in guiding protein folding and in allowing the formation of higher-order structures such as homo- or hetero-oligomers, disulfide bond formation is an important aspect of how the conformation of a mature, functional protein is reached. UNIT 14.1 describes the technology with which to study disulfide bond formation in living cells and in a completely cell-free system. Modification of proteins by lipid moieties allows proteins to interact with membranes. The functional properties of a number of proteins depend critically on this modification, and establishing whether or not acylation (UNIT 14.2) or isoprenylation (UNIT 14.3) have occurred is therefore important. There are other modifications that occur at distinct subcellular localizations, such as the modification of tyrosine with sulfate. Post-Translational Modification: Specialized Applications Contributed by Hidde L. Ploegh and Ben M. Dunn Current Protocols in Protein Science (2000) 14.0.1-14.0.2 Copyright © 2000 by John Wiley & Sons, Inc.
14.0.1 Supplement 20
The effect of oxygen on proteins can lead to modifications of cysteine, tyrosine, aspartic acid, and asparagine, as well as producing new reactive carbonyl groups. Methods for quantifying the products of such oxidative damage are given in UNIT 14.4. Future supplements will describe additional post-translational modifications to aid the investigator in achieving a more complete description of the protein under study. Hidde L. Ploegh and Ben M. Dunn
Introduction
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Current Protocols in Protein Science
Analysis of Disulfide Bond Formation
UNIT 14.1
Most proteins synthesized in the endoplasmic reticulum (ER) in eukaryotic cells and in the periplasmic space in prokaryotes are stabilized by disulfide bonds. Disulfide bonds are of two types: intrachain (within a polypeptide chain) and interchain (between separate chains). Intrachain disulfide bonds are formed during cotranslational and post-translational folding of a newly synthesized protein. Most interchain disulfide bonds are formed at a later stage in the maturation process and establish covalent links between subunits in oligomeric proteins. Disulfide bond formation can be followed in cultures of intact cells (Basic Protocol 1 and Alternate Protocols 1 and 2) or in an in vitro translation system containing isolated microsomes (Basic Protocol 2 and Alternate Protocol 3). First, the newly synthesized protein of interest is biosynthetically labeled with radioactive amino acids in a short pulse. The labeled protein is chased with unlabeled amino acids. At different times during the chase, a sample is collected, membranes are lysed with detergent, and the protein is isolated by immunoprecipitation (Support Protocol 1). The immunoprecipitates are analyzed for the presence of disulfide bonds by SDS-PAGE with and without prior reduction (Support Protocol 2). The difference in mobility observed between the gels with unreduced and reduced samples is due to disulfide bonds in the unreduced protein. CAUTION: Radioactive materials require special handling. See APPENDIX 2B concerning safe use of radioisotopes. ANALYSIS OF DISULFIDE BOND FORMATION IN INTACT MONOLAYER CELLS
BASIC PROTOCOL 1
In vivo, disulfide bond formation can be examined in cells growing in monolayers on tissue culture dishes (Braakman et al., 1991). This allows multiple wash steps in a short period of time and makes it easy to use short pulse and chase times. In addition, adherent cells are suitable for studies requiring frequent changes of medium with very different composition. When cells do not adhere or when low volumes of medium are desirable (e.g., when expensive additives are needed), the analysis can be done in suspension (see Alternate Protocol 1). This method detects cotranslational and post-translational disulfide bond formation; it is also possible to analyze post-translational disulfide bond formation (see Alternate Protocol 2). Materials Adherent cells Tissue culture medium containing methionine, 37°C Wash buffer (see recipe), 37°C Depletion medium (see recipe), 37°C Labeling medium (containing 125 to 250 µCi/ml [35S]methionine; see recipe), 37°C Chase medium (see recipe), 37°C Stop buffer (see recipe), 0°C Lysis buffer (see recipe), 0°C 60-mm tissue culture dishes, sterile 37°C humidified 5% CO2 incubator 37°C water bath with rack or insert to hold tissue culture dishes (e.g., Unwire racks for 15- and 50-ml tubes, Nalge) Aspiration flask for radioactive waste Flat, wide ice pan with fitted metal plate (e.g., VWR Scientific) Contributed by Ineke Braakman and Daniel N. Hebert Current Protocols in Protein Science (1996) 14.1.1-14.1.15 Copyright © 2000 by John Wiley & Sons, Inc.
Post-Translational Modification: Specialized Applications
14.1.1 Supplement 3
Cell scraper Additional reagents and equipment for immunoprecipitation (see Support Protocol 1) and nonreducing and reducing SDS-PAGE (see Support Protocol 2) NOTE: The volumes described here are for a 60-mm dish of cells. Volumes must be adjusted, based on the surface area of the dish, for other sizes. NOTE: All culture incubations are performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Perform pulse and chase 1. Set up cultures of adherent cells in tissue culture medium containing methionine in 60-mm tissue culture dishes so the cells will form a subconfluent monolayer on the day of the experiment. Incubate at 37°C. The experiment requires at least 1 dish per time point, and each dish should contain ≥106 cells.
2. Set up 37°C water bath with rack or insert for tissue culture dishes. Check that the water level is in contact with the bottom of a tissue culture dish but does not allow it to float if the lid is removed. Arrange the aspiration flask for radioactive waste so the pipet easily reaches dishes in the water bath. 3. Rinse cells twice with 2 ml wash buffer. Aspirate wash buffer and add 2 to 2.5 ml depletion medium. Incubate 15 to 30 min at 37°C in an incubator. This step depletes methionine in the cultured cells.
4. Remove dishes from the incubator and place on rack in 37°C water bath. 5. Pulse-label the cells, one dish at a time: aspirate depletion medium, add 400 µl labeling medium containing 50 to 100 µCi [35S]methionine, and incubate 1 to 2 min on rack in 37°C water bath. The pulse time should be equal to or shorter than the time required to synthesize the protein of interest (assuming a rate of 4 to 5 amino acid residues/sec) and long enough to detect the protein. If the protein of interest does not contain many methionines, or if it contains several cysteines, it may be worthwhile to deplete cysteine as well as methionine and label with [35S]methionine + [35S]cysteine (50 to 100 ìC; the total ratio depends on the protein) or with the same quantity of unpurified [35S]methionine, which usually contains ∼15% (v/v) [35S]cysteine as well. The stabilized form of unpurified methionine (e.g., [35S]in vitro labeling mix, Amersham) is less volatile and should be used for short pulses in a water bath to minimize radioactive contamination of air, pipets, and equipment.
For 0-min chase interval 6a. Add 2 ml chase medium to stop the pulse at precisely the end of the labeling interval. Rock gently to mix. 7a. Aspirate chase medium as quickly as possible. Transfer dish to aluminum plate on ice pan. Immediately add 2.5 ml cold stop buffer to end the chase. For all other chase intervals 6b. At precisely the end of the pulse labeling interval, add 2 ml chase medium to start the chase. Rock gently to mix. Aspirate medium and add 2 ml chase medium again. Incubate in a 37°C incubator or 37°C water bath for the desired chase intervals. Analysis of Disulfide Bond Formation
Choice of chase times is determined by the time it takes a protein to fold and form disulfide bonds. Generally, the first chase interval is equal to pulse time, the next approximately double that, and so on (e.g., 2, 5, 10, 20, and 40 min).
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Current Protocols in Protein Science
For short chase intervals, dishes can be incubated in a water bath, but for chase intervals >15 to 20 min, pH is best maintained in an incubator.
7b. At the end of the chase interval, aspirate chase medium and transfer dish to aluminum plate on ice pan. Add 2.5 ml cold stop buffer to end the chase. Cold stop buffer is used to stop all cellular processes. Cells may be left ≤30 min on ice in stop buffer.
Prepare the lysate 8. Remove stop buffer and add 2.5 ml cold stop buffer. 9. Aspirate dish as dry as possible. Add 600 µl cold lysis buffer. No incubation with lysis buffer is necessary when the buffer contains an alkylating agent. If alkylating agent is omitted from the lysis buffer, incubate 10 min with 20 mM N-ethylmaleimide (NEM) or longer with 20 mM iodoacetamide or 20 mM iodoacetic acid (typically 15 to 45 min).
10. Scrape dish and mix cell lysate with cell scraper. Transfer the lysate to a labeled 1.5-ml microcentrifuge tube. 11. Microcentrifuge 5 min at 12,000 rpm, 0°C, to pellet nuclei. Transfer supernatant (postnuclear lysate) to a clean 1.5-ml microcentrifuge tube. 12. Immediately analyze postnuclear lysate by immunoprecipitation (see Support Protocol 1 and UNIT 13.2) and nonreducing and reducing SDS-PAGE (see Support Protocol 2) or freeze it rapidly in liquid nitrogen and store at −80°C. Lysate with alkylating agent can be stored 1 to 2 hr on ice; lysate without alkylating agent should not be stored >5 min on ice.
ANALYSIS OF DISULFIDE BOND FORMATION IN CELLS IN SUSPENSION When cells do not adhere adequately to a solid support, or when it is desirable to use small volumes of reagents for analyses, cells in suspension may be used to analyze disulfide bond formation. One advantage to this approach is that samples at different chase intervals may be collected from a single tube of labeled cells. However, wash steps cannot be included during the pulse-chase incubations because the washes take too much time. Prior to starting the experiment it is necessary to determine (1) the minimum volume for incubating the cells (x µl), (2) the number of chase time points (y), and (3) the desired sample volume (z µl). Approximately 106 cells should be used for each time point.
ALTERNATE PROTOCOL 1
Additional Materials (also see Basic Protocol 1) Culture of suspension cells >1000 10 mCi/ml [35S]methionine (>1000 Ci/mmol; Amersham) 2× lysis buffer (see recipe) Concentrated chase medium (see recipe) 50-ml polystyrene tube with cap, sterile Beckman GPR cell centrifuge or equivalent Additional reagents and equipment for immunoprecipitation (see Support Protocol 1) and nonreducing SDS-PAGE (see Support Protocol 2) NOTE: All culture incubations are performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. NOTE: Keep cells in suspension during incubations by gently swirling the tube at regular intervals.
Post-Translational Modification: Specialized Applications
14.1.3 Current Protocols in Protein Science
Supplement 3
1. Transfer suspension cells for analysis to a sterile 50-ml polystyrene tube with cap. Use ∼106 cells per data point. If it is necessary to minimize the volumes of reagents used for the experiment, adherent cells can be removed from the dish by treating them with 0.25% (w/v) trypsin/0.2% (w/v) EDTA and resuspending them in a low-calcium medium.
2. Centrifuge cells 4 min at 500 × g (1500 rpm), 20° to 37°C. Resuspend pellet in 2y ml depletion medium and centrifuge again. Resuspend in 2y ml depletion medium. Incubate 15 to 30 min at 37°C. Pelleting conditions may vary with cell type.
3. Centrifuge cells 4 min at 500 × g, 20° to 37°C. Resuspend cells in x µl depletion medium in appropriate tube and place in water bath. 4. At the start of pulse, add 50 to 100 µCi undiluted [35S]methionine per time point to the cell suspension and mix by swirling. Incubate for the pulse period. 5. At the end of the pulse, add ≥4 vol (4 × x µl) concentrated chase medium. Mix by swirling. The total volume after addition of concentrated chase medium should be slightly more than y × z ìl to allow for loss due to evaporation. The final concentrations of chase medium components must be identical to those for the chase medium used in Basic Protocol 1 (see recipe for chase medium). Calculate the concentrations needed for the concentrated chase medium. A 1.25× solution is appropriate if 4 vol concentrated chase medium is added to the labeling mixture; for other added volumes, the concentrations must be adjusted accordingly.
6. Immediately collect the first sample of volume z. Add an equal volume of 2× lysis buffer, mix well, and place on ice. 7. At every chase time point, take a sample, add an equal volume of 2× lysis buffer, mix well, and place on ice. After the sample for the last chase time point is collected, the tube of cells should be almost empty.
8. Immediately analyze samples by immunoprecipitation (see Support Protocol 1) and nonreducing and reducing SDS-PAGE (see Support Protocol 2) or freeze rapidly in liquid nitrogen and store at −80°C. ALTERNATE PROTOCOL 2
ANALYSIS OF POST-TRANSLATIONAL DISULFIDE BOND FORMATION IN INTACT CELLS To determine the effect of certain conditions, such as ATP depletion, on folding, it may be necessary to separate the folding process from translation. This can be done by delaying folding until translation, glycosylation, and signal-peptide cleavage are complete. Folding of a disulfide-bonded protein may be prevented by incubating cells in reducing agent. Newly synthesized proteins cannot form disulfide bonds and often cannot fold when oxidation is prevented. When the reducing agent is removed after pulse-labeling, formation of disulfide bonds and folding may proceed (Braakman et al., 1992a,b; see Fig. 14.1.1).
Analysis of Disulfide Bond Formation
Wash the cells and deplete the methionine (see Basic Protocol 1, steps 2 and 3). Add dithiothreitol (DTT; APPENDIX 3A) to labeling medium to give a final concentration of 5 mM. Perform a pulse-chase experiment and prepare lysates (see Basic Protocol 1, steps 4 to 11).
14.1.4 Supplement 3
Current Protocols in Protein Science
It may be necessary to optimize the final concentration of DTT in labeling medium for each cell type. During long incubations, DTT may affect cellular ATP levels. Efficiency of incorporation may also be slightly diminished by the presence of DTT. The duration of pulse labeling may be increased to improve incorporation of label in protein. Before the effects of different conditions on folding are tested, the efficiency, rate, and outcome of post-translational folding should be compared to those of cotranslational folding. ANALYSIS OF DISULFIDE BOND FORMATION IN ROUGH ENDOPLASMIC RETICULUM–DERIVED MICROSOMES
BASIC PROTOCOL 2
Formation of disulfide bonds in proteins which possess signal sequences that target the protein to the endoplasmic reticulum can also be studied with an in vitro translation/rough endoplasmic reticulum–derived microsome translocation and folding system (Fig. 14.1.1). 35S-labeled proteins are generated by the translation of mRNA with a rabbit reticulocyte lysate in the presence of [35S]methionine. Translation is carried out in the presence of dog pancreas microsomes; this combination permits cotranslational insertion of 35S-labeled protein into microsomes. An oxidizing agent (i.e., oxidized glutathione, GSSG; APPENDIX 3A) is present during translation to allow monitoring of cotranslational or vectorial oxidation from the N- to the C-terminus of the protein, the mechanism by which proteins acquire their disulfide bonds under normal physiological conditions.
A
microsome membrane
ribosome
mRNA
GSSG/ DT T polypeptide
lumen
carbohydrate
B GSSG
DTT
lumen
Figure 14.1.1 In vitro translation/rough endoplasmic reticulum–derived microsome translocation and folding system. (A) For cotranslational folding, the protein is translocated into microsomes containing an oxidizing environment (GSSG). This provides an opportunity for the protein to fold vectorially during the translation and translocation processes. (B) For post-translational folding, oxidizing agent is added after translation and translocation, permitting synchronization and isolation of the folding process.
Post-Translational Modification: Specialized Applications
14.1.5 Current Protocols in Protein Science
Supplement 3
Oxidation of proteins in microsomes provides a system where the experimental conditions can be more extensively controlled and manipulated. The oxidation state of the protein can be trapped by alkylation with N-ethylmaleimide (NEM) and monitored by immunoprecipitation (see Support Protocol 1) and nonreducing and reducing SDS-PAGE (see Support Protocol 2). Materials 1 equivalent/µl nuclease-treated dog pancreas microsomes (Promega) Rabbit reticulocyte lysate treated with ATP-regenerating system and nucleases (Promega) 1 mM amino acid mixture lacking methionine (Promega) 10 mCi/ml [35S]methionine (1000 Ci/mmol, Amersham) 100 mM dithiothreitol (DTT) RNase-free H2O (e.g., DEPC-treated; see recipe) 23 U/ml RNase inhibitor (e.g., RNasin, Promega) 1 µg/µl mRNA for the protein of interest 100 mM oxidized glutathione (GSSG; APPENDIX 3A), titrated to neutrality with KOH 120 mM N-ethylmaleimide (NEM) in 100% ethanol (prepare from 1 M stock; see recipe) Lysis buffer (see recipe) 2× SDS sample buffer (UNIT 10.1; optional) 27°C water bath 1.5-ml microcentrifuge tubes, RNase-free Ice bath Additional reagents and equipment for immunoprecipitation (see Support Protocol 1) and nonreducing SDS-PAGE (see Support Protocol 2) NOTE: The major source of failure in the in vitro translation/translocation of proteins is contamination with ribonucleases (RNases) that degrade mRNA. To avoid contamination, water and salt solutions should be treated with diethylpyrocarbonate (DEPC) to chemically inactivate RNases; glass and plasticware should be treated with DEPC-treated water or otherwise treated to remove RNase activity (see recipe for DEPC treatment). Freshly opened plasticware that has not been touched by unprotected hands is also acceptable. It is also helpful to keep a set of solutions for RNA work alone to ensure that “dirty” pipets do not contaminate them. In addition, gloves should be worn at all times. 1. Thaw reagents and solutions rapidly with agitation in 27°C water bath. Rabbit reticulocyte lysate and dog pancreas microsomes should be stored in aliquots of working volumes (i.e., 6 ìl and multiples of 6 ìl). Repeated freeze-thaw cycles result in inefficient translation and translocation of protein. Freeze other reagents rapidly in liquid nitrogen after use.
2. Prepare reaction mix in an RNase-free 1.5-ml plastic microcentrifuge tube (96.7 µl total):
Analysis of Disulfide Bond Formation
6 µl 1 equivalent/µl nuclease-treated dog pancreas microsomes 52 µl treated rabbit reticulocyte lysate 2 µl 1 mM amino acid mixture lacking methionine 8 µl 10 µCi/µl [35S]methionine 1 µl 100 mM DTT 16 µl RNase-free H2O 4 µl 23 U/µl RNase inhibitor 4 µl 1µg/µl mRNA for the protein of interest 3.7 µl 100 mM GSSG.
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Current Protocols in Protein Science
This reaction mix is enough for 10 samples. Adjust the volumes proportionately for more or fewer samples. Each sample represents a different time point. mRNA for the protein of interest can be transcribed from cDNA for the protein of interest into a commercially available vector (e.g., pBluescript, Stratagene) which contains both T7 and T3 promoters for RNA polymerases.
3. Mix thoroughly with a pipet. Incubate at 27°C. Set pipettor to less than the total volume of reaction mixture to avoid introducing air bubbles. Mix by repeatedly taking the mixture up and gently expelling it with the pipettor.
4. At the appropriate time (e.g., 0.5, 1, 1.5, 2, 3,…n hr), transfer 9-µl samples to separate tubes and add 2.4 µl of 120 mM NEM in ethanol (25 mM final) to alkylate proteins. Incubate 10 min on ice. In vitro translation of proteins is 3- to 5-fold slower than in vivo translation, so extended time points are required.
5. Add 600 µl lysis buffer to each alkylated sample or add 30 µl of 2× sample buffer for direct analysis of total proteins translated. 6. Analyze lysate by immunoprecipitation (see Support Protocol 1) and nonreducing SDS-PAGE (see Support Protocol 2). ANALYSIS OF POST-TRANSLATIONAL DISULFIDE BOND FORMATION IN ROUGH ENDOPLASMIC RETICULUM–DERIVED MICROSOMES
ALTERNATE PROTOCOL 3
Post-translational oxidation is performed by translating the protein under reducing conditions in the presence of microsomes to allow accumulation of translocated and glycosylated proteins with the signal sequence removed and lacking disulfide bonds. Oxidizing agents are added post-translationally to initiate oxidation (see Fig. 14.1.1). Perform in vitro translation and analysis as for the previous method (see Basic Protocol 2) with the following exceptions in the indicated steps. 2. Omit oxidized glutathione (GSSG) from the reaction mixture (to make 93 µl). 3. After the initial incubation of the reaction, add 4.2 µl of 100 mM GSSG and incubate 1 hr more at 27°C before proceeding to alkylate and analyze proteins (steps 4 to 8). IMMUNOPRECIPITATION OF LYSATES The protein of interest is precipitated from lysates of cells or microsomes using a specific antibody bound to Staphylococcus aureus cells or Protein A–Sepharose beads (beads give less background but are twice as expensive as the cells). The immunoprecipitate is then analyzed by nonreducing and reducing SDS-PAGE (see Support Protocol 2). Materials 10% (w/v) killed, fixed Staphylococcus aureus cells (Zymed) Antibody against protein of interest Lysate from pulse-chase labeled cells or microsomes (see Basic Protocol 1 or 2 or Alternate Protocol 1, 2, or 3) Immunoprecipitation wash buffer (see recipe), 37°C TE buffer, pH 6.8 (see recipe) 2× nonreducing sample buffer (see recipe) 200 mM dithiothreitol (DTT) Rotator
SUPPORT PROTOCOL 1
Post-Translational Modification: Specialized Applications
14.1.7 Current Protocols in Protein Science
Supplement 3
Additional reagents and equipment for nonreducing and reducing SDS-PAGE (see Support Protocol 2) 1. Add 60 µl of 10% killed, fixed Staphylococcus aureus cells to 0.5 to 100 µl antibody. Rotate 1 hr at 4°C. The amount of antibody used depends on the antibody and its concentration. The time required to bind antibody to S. aureus cells may vary from 20 min to several hours. Alternatively, 60 ìl of 10% (w/v) Protein A–Sepharose beads, washed five times in an appropriate buffer, may be used instead of S. aureus cells.
2. Add 100 to 600 µl postnuclear lysate. Incubate 1 hr at 4°C, with shaking. If background is high, preincubate the lysate with S. aureus cells without antibody 1 hr at 4°C. Microcentrifuge the cells 4 min at 2000 × g, room temperature, and transfer the cleared lysate to antibody-coated S. aureus cells.
3. Pellet cells by microcentrifuging 4 min at 2000 × g (5000 rpm), room temperature. Remove supernatant and resuspend cells in 1 ml wash buffer. Shake 5 min. Repeat wash once and pellet cells. The temperature of the wash can affect the level of background; generally, the higher the temperature, the lower the background. Typical wash temperatures range from 10°C to room temperature. Protein A–Sepharose beads can be pelleted by centrifuging 1 min at 8000 × g (10,000 rpm), room temperature.
4. Aspirate the supernatant. Add 20 µl TE buffer. Shake 5 min or until cells are resuspended. 5. Add 20 µl of 2× nonreducing sample buffer without reducing agent and vortex. Boil 5 min at 95°C and vortex. 6. Pellet S. aureus cells by microcentrifuging 4 min at 12,000 × g (12,000 rpm) to give nonreduced sample in the supernatant. 7. Transfer 20 µl supernatant to a tube containing 2 µl of 200 mM DTT and vortex. Boil 5 min at 95°C. Microcentrifuge briefly at 12,000 × g to give reduced sample. 8. Analyze nonreduced and reduced samples by SDS-PAGE (see Support Protocol 2). Samples can be rapidly frozen in liquid nitrogen and stored at −80°C, but storage may increase background. SUPPORT PROTOCOL 2
NONREDUCING AND REDUCING SDS-PAGE Immunoprecipitates (see Support Protocol 1) are analyzed by nonreducing SDS-PAGE to detect changes in mobility due to disulfide bond formation. Materials Samples in 1× sample buffer (see Support Protocol 1) 2× nonreducing sample buffer (see recipe) PBS (APPENDIX 2E)/30% (v/v) methanol 1.5 M salicylate/30% (v/v) methanol pH paper Whatman 3MM filter paper
Analysis of Disulfide Bond Formation
Additional reagents and equipment for SDS-PAGE minigel with Laemmli buffers (UNIT 10.1) and Coomassie blue staining and destaining (UNIT 10.5)
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Current Protocols in Protein Science
1. Prepare a 1- or 0.75-mm thick polyacrylamide separating and stacking minigel. Percent acrylamide depends on molecular weight of the protein.
2. Load 8 µl of each sample. Load 1× nonreducing sample buffer in the two lanes next to the samples. When nonreduced and reduced samples are to be loaded on the same gel, leave two empty lanes between the two sample types, and load empty lanes with 1× nonreducing sample buffer. If nonreduced and reduced samples need to be loaded next to each other, cool the samples and add NEM to a final concentration of 100 mM to both sets of samples. Mix and pellet the cells before loading the samples on the gel.
3. Run gel ∼1 hr at 20 to 25 mA until the dye front is close to or at the bottom of the gel. 4. Stain the gel, including the stacking gel, with Coomassie blue stain and destain (see UNIT 10.5). Stacking gels often contain unreduced or aggregated protein.
5. Neutralize gel by incubating it in three changes of PBS/30% methanol, 5 min each, until pH >6. Check the pH with pH paper. 6. Treat gel 30 min in 1.5 M salicylate to enhance. Salicylate is used as a safer, less expensive, and faster way to enhance the signal 3- to 5-fold. Fluorography can also be enhanced using commercial solutions (e.g., Enhance, DuPont or Amplify, Amersham).
7. Dry gel onto Whatman 3MM filter paper. Autoradiograph/fluorograph at −80°C. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Chase medium Complete tissue culture medium appropriate for the cells containing: 20 mM HEPES (sodium salt), pH 7.3 (see recipe) 5 mM methionine (see recipe) 1 mM cycloheximide (optional; see recipe) Prepare fresh daily Add HEPES, methionine, and optional cycloheximide from concentrated stocks (see recipes). When labeling is done with cysteine, the chase medium should also contain 5 mM cysteine (from 1 M stock; see recipe). The chase medium may contain fetal bovine serum or another protein source, especially when chase times are long. Protein should be omitted when more defined conditions are needed. When the reducing agent dithiothreitol (DTT) is added, proteins present in the medium will partly quench the reductant.
Concentrated chase medium Adjust the concentration of chase medium components (see recipe) so the final concentrations are 20 mM HEPES, 5 mM methionine, and 5 mM cycloheximide after the concentrated chase medium has been added to the labeling medium for cells in suspension (see Alternate Protocol 1). Post-Translational Modification: Specialized Applications
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Cycloheximide, 500 mM stock 28.1 g cycloheximide H2O to 100 ml Store 1- to 2-ml aliquots ≤2 years at −20°C Thaw and warm to room temperature to completely dissolve before using. Do not freeze and thaw more than three times.
Cysteine, 1 M stock 12.12 g cysteine H2O to 100 ml Store 1- to 2-ml aliquots ≤2 years at −20°C Thaw and warm to room temperature to completely dissolve before using. Do not freeze and thaw more than three times.
Depletion medium Methionine-free tissue culture medium containing: 0.026 M sodium bicarbonate (2.2 g/liter) 20 mM HEPES, pH 7.3 (from 1 M stock; see recipe) Store ≤1 year at 4°C Diethylpyrocarbonate (DEPC) treatment of solutions and labware CAUTION: Wear gloves and use a fume hood when working with DEPC because it is a suspected carcinogen. Solutions: Add 0.2 ml DEPC per 100 ml of the solution to be treated. Shake vigorously to dissolve DEPC. Autoclave the solution to inactivate remaining DEPC. Any water or salt solutions used in RNA preparation should be treated with DEPC. Note that solutions containing Tris cannot be effectively treated with DEPC because Tris reacts with DEPC to inactivate it.
Labware: Rinse glass and plasticware thoroughly with DEPC solution. Alternatively, bake glassware 4 hr at 300°C; rinse plasticware with chloroform, or use fresh plasticware straight from a package that has not been touched by unprotected hands. Wear gloves for all manipulations. Note that autoclaving alone will not fully inactivate many RNases.
Ethylenediaminetetraacetic acid (EDTA), 200 mM stock 18.61 g EDTA 180 ml H2O 10 mM NaOH added dropwise with mixing just until EDTA dissolves Adjust pH to 7.3 H2O to 250 ml Store at 4°C For EDTA, pH 6.8, adjust the pH to 6.8.
HEPES (N-2-hydroxyethylpiperidine-N′-ethanesulfonic acid), 1 M stock 119.15 g HEPES (sodium salt) 400 ml H2O Adjust pH to 7.3 with 10 M NaOH H2O to 500 ml Store 100-ml aliquots at −20°C and thawed aliquots at 4°C
Analysis of Disulfide Bond Formation
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Immunoprecipitation wash buffer PBS (APPENDIX 2E)/0.5% (v/v) Triton X-100 or PBS/150 mM NaCl Store at 4°C or room temperature For every protein-antibody combination, the optimal wash buffer needs to be determined. The two buffers listed are particularly mild wash buffers that will maintain most antigen-antibody interactions but may lead to high background. To decrease background, other detergents or multiple detergents may be added, salt concentration may be increased, or a combination of the two may be used. SDS at a concentration ≥0.05% (w/v) may be especially helpful, with or without the quenching effect of added nondenaturing detergents.
Labeling medium Methionine-free tissue culture medium containing: 125 to 250 µCi [35S]methionine/ml Prepare fresh for each experiment (400 µl per chase time point) Addition of 50 to 100 ìCi in 400 ìl to a 60-mm-diameter dish (∼3–5 × 106 cells) should be sufficient to visualize a 1- to 2-min-pulse-labeled protein that is expressed to a high level in the cell. [35S]methionine + [35S]cysteine or unpurified [35S]methionine + [35S]cysteine can be used to label proteins that contain few methionines but several cysteines.
Lysis buffer PBS (APPENDIX 2E) or similar buffer containing: 0.5% (v/v) Triton X-100 1 mM EDTA, pH 6.8 (see recipe), added just before use 20 mM NEM (see recipe), added just before use 1 mM PMSF (see recipe), added just before use Small peptide mixture (10 µg/ml each final; see recipe), added just before use Use within 3 hr Add EDTA, NEM, and protease inhibitors (PMSF and small peptide mixture) from concentrated stocks (see recipes). For 2× lysis buffer, double the concentrations of reagents. Depending on the protein analyzed and on the purpose of the experiment, a variety of detergents may be used. Triton X-100 may disrupt noncovalent interactions between proteins. Milder detergents that can be used include CHAPS (3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate), deoxycholate, octylglucoside, or a mixture of detergent and lipid.
Methionine, 200 mM stock 2.984 g methionine H2O to 100 ml Store 1- to 2-ml aliquots ≤2 years at −20°C Thaw and warm to room temperature to completely dissolve before using. Do not freeze and thaw more than three times.
N-ethylmaleimide (NEM), 1 M stock 12.5 g NEM 100 ml 100% ethanol Store 1- to 2-ml aliquots ≤2 years at −20°C protected from light NEM is sensitive to light and to hydrolysis in water. If it is frozen and thawed too many times or exposed to too much light, the solution will turn yellow and the NEM will precipitate.
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Supplement 3
Nonreducing sample buffer, 2× 400 mM Tris⋅Cl, pH 6.8 6% (w/v) SDS 20% (v/v) glycerol 2 mM EDTA, pH 6.8 (from 200 mM stock; see recipe) 0.01% (w/v) bromphenol blue Phenylmethylsulfonyl fluoride (PMSF), 200 mM stock 3.484 g PMSF 10 ml anhydrous isopropanol Store in 500-µl aliquots ≤2 years at −20°C PMSF is highly unstable in water (the half life is ∼30 min at 37°C and a few hours on ice). Add this stock to solution immediately before use.
Small peptide mixture stock 10 mg/ml chymostatin in dimethyl sulfoxide (DMSO) 10 mg/ml leupeptin in DMSO 10 mg/ml antipain in DMSO 10 mg/ml pepstatin in DMSO Store in 10-µl aliquots ≤2 years at −20°C Use at a final concentration of 10 µg/ml each Stop buffer Wash buffer (see recipe) containing salts only 20 mM NEM (from 1 M stock; see recipe) Prepare fresh and store on ice <5 hr NEM may be replaced by iodoacetamide or iodoacetic acid, which react more slowly with free –SH groups. They are less specific than NEM and penetrate cells more slowly than NEM, but they irreversibly carboxymethylate the –SH group, whereas NEM may dissociate slowly. Deblocking of –SH groups after NEM treatment is not a problem if cells are stored ≤2 hr on ice.
TE buffer, pH 6.8 10 mM Tris⋅Cl, pH 6.8 1 mM EDTA, pH 6.8 (from 200 mM stock; see recipe) Store at 4°C Wash buffer PBS (APPENDIX 2E) containing 0.9 mM Ca2+/0.5 mM Mg2+, Earles balanced salt solution, or other isotonic buffer containing salts only. For most cells, calcium ions are crucial to maintain adherence to the plastic dishes, especially to withstand many washes. When cost is not an issue, all washes can be performed using depletion medium. However, depletion medium may not be used as a substitute in stop buffer.
COMMENTARY Background Information
Analysis of Disulfide Bond Formation
Folding of a newly synthesized protein can be followed by assaying for conformational changes in the molecule immediately after synthesis. For this type of analysis, a large number of antibodies that recognize specific epitopes are required to analyze the changes in the complete molecule. In a mature protein that is stabilized by intrachain disulfide bonds (which is
true of most proteins synthesized in the endoplasmic reticulum), the formation of these cross-links is indicative of folding (Creighton, 1986). A disulfide bond can be formed only when the conformation allows the two participating cysteines to be in close proximity. With only few antibodies available, the rate and extent of folding of a protein can be examined grossly through detection of disulfide bond
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formation. Of course, disulfide bond formation is not identical to folding; therefore, a comparison of disulfide bond formation and conformational changes is required for every protein studied. When a disulfide bond–containing protein is prepared for SDS-PAGE, it is routinely reduced. In nonreducing denaturing polyacrylamide gel electrophoresis, however, the protein is denatured with SDS, but the disulfide bonds remain intact. The consequence is a more compact conformation of the protein, resulting in a higher electrophoretic mobility; however, in some cases the protein may bind less SDS, resulting in a lower electrophoretic mobility. Most proteins will run faster in oxidized form than in reduced form. The choice of radioactive labeling is dictated by the desire to follow a protein from the moment of synthesis. No other method permits following the life of a protein in a cell in such detail. Any other method of detection would require bulk production of proteins, which would be very difficult and nonphysiological. The in vitro translation system coupled with dog pancreas microsomes has been traditionally employed to study the translocation of secretory, lysosomal, and many integral membrane proteins across the membrane of the endoplasmic reticulum (Blobel and Dobberstein, 1975). However, more recently it has been shown that microsomes possess a complete set of foldases and chaperones and thus provide an environment in which proteins can be oxidized rapidly and efficiently (Scheele and Jacoby, 1982; Marquardt et al., 1993; Hebert et al., 1995). Microsomes provide a system that can be controlled and manipulated to a much higher degree than intact cells. Direct access is provided to the lysate (equivalent to the cytosol in the cell), and intralumenal components can be manipulated with ionophores, detergents, toxins, or alkaline pH (Bulleid and Freedman, 1988; Nicchitta and Blobel, 1993; Hebert et al., 1995). Post-translational oxidation of proteins (see Alternate Protocol 2 and Alternate Protocol 3) can be used in both intact cells and microsomes to isolate the oxidation process from other biosynthetic events. Together, analysis of cellular and microsomal folding can be employed to dissect the folding pathway of a protein.
Critical Parameters Wash buffer for adherent cells should contain Ca2+ and Mg2+ to maintain adherence to
the plate, especially when there are several washes. If cost is not an issue, cells can be washed in depletion medium. Labeling medium should contain 50 to 100 µCi 35S-labeled methionine for ∼3–5 × 106 cells. If the protein of interest has few methionines, or if it contains several cysteines, a combination of [35S]methionine and [35S]cysteine, or unpurified labeled methionine and cysteine, may be used for labeling. When short pulses are performed in a water bath, the stabilized version of unpurified methionine (35S in vitro labeling mix, Amersham) should be used to minimize contamination of air, pipets, and incubators. Cycloheximide in the chase medium will stop elongation of unfinished nascent peptide chains. When the kinetics of disulfide bond formation are being studied, cycloheximide should be added to the chase medium; however, it should be omitted when a maximum amount of incorporated label is required (Braakman et al., 1991). An alkylating agent—e.g., N-ethylmaleimide, iodoacetamide, or iodoacetic acid— must be included in the stop buffer and lysis buffer to prevent artifactual formation of disulfide bonds. Initially, it is recommended to test at least two alkylating agents and compare the results with those obtained in the absence of alkylating agent. Ideally, there should be no difference in results obtained with the various alkylating agents, except possibly a change in electrophoretic mobility due to alkylation. SDS-PAGE should be performed under nonreducing conditions. Reducing agents should be completely absent. If a reducing environment does exist in (one of) the samples, all samples should be quenched with at least a two-fold concentration of N-ethylmaleimide or another alkylating agent. The translocation process is dependent upon free sulfhydryl groups, but oxidation requires an oxidizing environment. Thus, it would seem impossible for cotranslational oxidation to occur in the in vitro translation system with microsomes. However, it is possible to simultaneously create a reducing extralumenal compartm en t an d an o xidizing intralumenal compartment, probably due to the presence of a putative oxidized glutathione (GSSH) transporter in microsomal membranes. The redox range over which these two processes occur efficiently is small. In addition, the redox potential is directly related to the pH, so pH must be carefully controlled.
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Troubleshooting The presence of disulfide bonds in a protein does not guarantee an electrophoretic mobility difference between reduced and oxidized forms, especially when a protein is large and disulfide bonds form small peptide loops. The largest shift can be expected at a late chase time when all disulfide bonds have been formed. If the presence of disulfide bonds is not certain and needs to be tested, the radioactive cell lysate can be treated with GSSG, followed by immunoprecipitation and analysis by nonreducing and reducing SDS-PAGE as described (see Support Protocol 1 and Support Protocol 2). Alternatively, free sulfhydryl groups can be labeled with [14C]iodoacetamide before and after reduction. When the mobility difference between reduced and oxidized protein is minimal, resolution might be improved by using a lower percentage acrylamide gel and/or changing the alkylating agent. The treated reticulocyte lysate used here is optimized to efficiently translate a large range of mRNAs with an ATP-generating system, mixture of tRNAs, and salts. The concentration of salts greatly affects the translation efficiency. If the efficiency of translation is low, an untreated lysate should be used and optimized specifically for the mRNA for the protein of interest. If there is difficulty in differentiating between translocated and untranslocated proteins in microsomes, untranslocated protein can be removed by protease digestion or centrifugation. Translocation of protein across microsomal membrane protects the protein from digestion by added proteases. Furthermore, dense rough microsomes can be pelleted easily by centrifugation, thereby isolating translocated proteins from extralumenal untranslocated proteins. If the background is high in immunoprecipitation, try different immunoprecipitation wash buffers, using other detergents or multiple detergents, increased salt concentration, or a combination of both. SDS at concentrations ≥0.05% (w/v) may be especially helpful, with or without the quenching effect of added nondenaturing detergents.
Anticipated Results
Analysis of Disulfide Bond Formation
In most cases, formation of disulfide bonds creates a more compact structure that results in an increase in the mobility of a protein in nonreducing SDS-PAGE (for representative results, see Braakman et al., 1992 a,b). The num-
ber of disulfide bonds and the distance between the cysteines in a sulfhydryl pair influences the observed change in mobility. During the chase, the protein will move from the more reduced position in the gel to the more oxidized position. For some proteins distinct oxidative intermediates may be observed, but for others, a fuzzy band may precede formation of native protein. In reducing SDS-PAGE, ideally one band will be found in intact cells. Two bands will be found in the microsomal system—one for untranslocated protein and one for translocated protein. If the protein possesses a signal sequence (1.5 to 3.0 kDa) that is cleaved during translocation, a small mobility shift dependent upon the protein’s molecular weight may be observed. However, differentiation of translocated and untranslocated proteins in reducing SDS-PAGE is best seen for multiglycosylated proteins, because each glycosylation step adds 2.5 kDa to the translocated protein. Modifications of oligosaccharides and other post-translational modifications can change the electrophoretic mobility as well, but these changes can be distinguished from disulfide bond formation by comparing reducing with nonreducing gels.
Time Considerations Design and preparation of the experiment should take the most time. The very first time, this may take 1 or more days, depending on experience and available equipment. Preparation for routine experiments is around 1 hr. The pulse-chase portion requires 1 hr plus the maximum chase time. For rapidly folding proteins, folding can be over in 15 min; for others it may take hours. Cultures of cells should be started 1 or 2 days before the experiment so they will be subconfluent on the day of the experiment. Pulse-chase, immunoprecipitation, and SDS-PAGE are optimally done in 1 day. It is possible to rapid-freeze lysates or immunoprecipitates in liquid nitrogen and store them at −80°C until further use, but freezing may lead to higher background and less reproducibility.
Literature Cited Blobel, G. and Dobberstein, B. 1975. Transfer of proteins across membranes. II. Reconstitution of functional rough microsomes from heterologous components. J. Cell Biol. 67:852-862. Braakman, I., Hoover-Litty, H., Wagner, K.R., and Helenius, A. 1991. Folding of influenza hemagglutinin in the endoplasmic reticulum. J. Cell Biol. 114:401-411.
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Braakman, I., Helenius, J., and Helenius, A. 1992a. Manipulating disulfide bond formation and protein folding in the endoplasmic reticulum. EMBO J. 11:1717-1722. Braakman, I., Helenius, J., and Helenius, A. 1992b. Role of ATP and disulphide bonds during protein folding in the endoplasmic reticulum. Nature 356:260-262. Bulleid, N.J. and Freedman, R. 1988. Defective cotranslational formation of disulphide bonds in protein disulphide-isomerase-deficient microsomes. Nature 335:649-651. Creighton, T.E. 1986. Disulfide bonds as probes of protein folding pathways. Methods Enzymol. 131:83-106. Hebert, D.N., Foellmer, B., and Helenius, A. 1995. Glucose trimming and reglycosylation determine glycoprotein association with calnexin in the endoplasmic reticulum. Cell 81:425-433. Marquardt, T., Hebert, D.N., and Helenius, A. 1993. Post-translational folding of Influenza hemagglutinin in isolated endoplasmic reticulum–derived microsomes. J. Biol. Chem. 268:1961819625. Nicchitta, C.V. and Blobel, G. 1993. Lumenal proteins of the mammalian endoplasmic reticulum are required to complete protein translocation. Cell 73:989-998.
Scheele, G. and Jacoby, R. 1982. Conformational changes associated with proteolytic processing of presecretory proteins allow glutathione-catalyzed formation of native disulfide bonds. J. Biol. Chem. 257:12277-12282.
Key References Braakman et al., 1991. See above. Describes the protocol in intact cells, results with influenza hemagglutinin, and considerations for ultra-short pulse times. Hebert et al., 1995. See above. Marquardt et al., 1993. See above. Describes the protocol in microsomes.
Contributed by Ineke Braakman University of Amsterdam Academic Medical Centre The Netherlands Daniel N. Hebert Yale University School of Medicine New Haven, Connecticut
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Analysis of Protein Acylation
UNIT 14.2
Protein acylation is the covalent attachment of fatty acids to a protein; the most commonly added fatty acids are myristate (14:0) and palmitate (16:0). First, radiolabeled fatty acids are used to label eukaryotic cells in vitro (see Basic Protocol 1). The radiolabeled material produced can then be analyzed by various methods: the type of fatty acid linkage can be determined (see Basic Protocol 2), the nature of the protein-bound label can be determined to check for interconversion (see Basic Protocol 3), and the protein-bound fatty acid can be identified (see Basic Protocol 4). BIOSYNTHETIC LABELING WITH FATTY ACIDS To identify proteins that are modified with fatty acid groups, cultured cells are incubated first in medium containing sodium pyruvate, which acts as a source of acetyl-CoA and minimizes interconversion of the fatty acid to other metabolites, and then with [3H]fatty acids. Fatty acids tritiated at positions 9 and 10 provide the best combination of high specific activity and detectability for in vitro labeling, and because the tritium label is far removed from the carboxyl end where β-oxidation occurs, reincorporation of label is minimized.
BASIC PROTOCOL 1
Materials Cells for culture Complete tissue culture medium appropriate for cells Labeling medium: complete tissue culture medium containing the relevant dialyzed serum and 5 mM sodium pyruvate, 37°C 5 to 10 µCi/µl [9,10(n)-3H]fatty acid, e.g., [9,10(n)-3H]palmitic acid or [9,10(n)-3H]myristic acid (30 to 60 Ci/mmol; Amersham International, American Radiolabeled Chemicals, or NEN Research Products) in ethanol PBS, pH 7.2 (APPENDIX 2E), ice-cold 1% (w/v) SDS or SDS sample buffer (for SDS-PAGE, when using adherent or nonadherent cells respectively; UNIT 10.1) or RIPA lysis buffer (for immunoprecipitation; UNIT 13.2) 5× SDS sample buffer (see recipe) Cell scrapers Nitrogen gas Additional reagents and equipment for immunoprecipitation (UNIT 13.2), SDS-PAGE (UNIT 10.1), treating a gel with sodium salicylate (UNIT 14.3) or DMSO/PPO solution (UNIT 10.2), and fluorography (UNIT 10.2) NOTE: All reagents and equipment coming into contact with live cells must be sterile, and proper sterile technique should be used accordingly. NOTE: All culture incubations are performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. 1. On the day before the labeling experiment, split the cells into fresh complete tissue culture medium. Set up the cells at two split ratios; then choose the culture closest to 70% to 80% confluency for labeling.
2. The next day, replace the medium with a minimum volume of 37°C labeling medium. Incubate 1 hr.
Contributed by Caroline S. Jackson and Anthony I. Magee Current Protocols in Protein Science (1996) 14.2.1-14.2.9 Copyright © 2000 by John Wiley & Sons, Inc.
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Cells in suspension should be used at a cell density of 106 to 107 cells/ml. For adherent cells that are 70% to 80% confluent, the minimum amount of medium necessary to cover the dish—e.g., 1.5 ml for 60-mm dishes and 3 ml for 100-mm dishes—should be used.
3. Add 5 to 10 µCi/µl [9,10(n)-3H]fatty acid to a concentration of 50 to 500 µCi/ml. Incubate up to 24 hr. Cells vary in the rate and extent of incorporation (see Critical Parameters), so both the amount of label and the duration of incubation need to be optimized. Labeling cells overnight in the presence of 200 ìCi/ml [3H]fatty acid will maximize the chances of detecting labeled proteins. The amount of label and/or time of incubation can then be reduced if good incorporation of label is achieved, or increased if poor incorporation is attained. Short labeling times (e.g., pulses on the order of minutes up to 2 hr) require amounts of label at the higher end of the indicated range. In this case, uptake is relatively low and the medium plus label can be reused one or more times. The level of label in the medium can be monitored by scintillation counting. For longer incubations the interconversion of fatty acids becomes a greater problem, and the protein-bound fatty acid label should be analyzed (see Basic Protocols 3 and 4). If the [3H]fatty acid is not supplied in ethanol or if the concentration is too low, remove the solvent by blowing nitrogen over the solution in its original container until dry and redissolve the label in ethanol at a concentration of 5 to 10 ìCi/ìl. Do not transfer into another container or evaporate the solvent in a plastic container, as this will cause a significant loss of label that will adhere to the side of the container.
For adherent cells 4a. Place the dish on ice and aspirate the medium. Wash the cells twice with ice-cold PBS and lyse the cells by adding 1% SDS for SDS-PAGE (UNIT 10.1) or RIPA lysis buffer for immunoprecipitation (UNIT 13.2), using 100 µl of 1% SDS for a 60-mm dish or 300 µl for a 100-mm dish, or 1 ml RIPA lysis buffer. CAUTION: Radioactive medium and washes must be disposed of appropriately.
5a. Using a cell scraper, remove the lysed cells from the dish and transfer them to a 1.5-ml microcentrifuge tube. Add 20 µl lysate to 5 µl of 5× SDS-PAGE sample buffer. Use all of RIPA lysate for immunoprecipitation. Resuspend immunoprecipitate in 20 µl SDS sample buffer. For SDS-PAGE, use DTT at a concentration ≤20 mM, and do not boil the samples, but incubate them only 3 min at 80°C; this is necessary because the thioester linkage of the fatty acid is susceptible to cleavage.
For nonadherent cells 4b. Microcentrifuge the cell suspension 1 min at 6000 rpm, 4°C, to pellet the cells. Decant the supernatant and wash the cell pellet once by resuspending it in 1 ml ice-cold PBS and centrifuging again. 5b. Lyse the cells by resuspending the cell pellet in 100 µl SDS-PAGE sample buffer for discontinuous SDS-PAGE (UNIT 10.1) or 1 ml RIPA lysis buffer for immunoprecipitation (UNIT 13.2) for 106 to 107 cells. Resuspend immunoprecipitate in 20 µl SDS sample buffer. CAUTION: Radioactive medium and washes must be disposed of appropriately. For analysis of total protein-bound fatty acid label, lyse the cells in 100 ìl 1% SDS. For SDS-PAGE, use DTT at a concentration ≤20 mM, and do not boil the samples, but incubate them only 3 min at 80°C; this is necessary because the thioester linkage of the fatty acid is susceptible to cleavage. Analysis of Protein Acylation
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6. Analyze whole-cell lysate or immunoprecipitate on an SDS-PAGE minigel, using 20 µl lysate per lane. Store remaining lysate at −20°C. 7. Treat the gel with sodium salicylate (UNIT 14.3) or DMSO/PPO solution (UNIT 10.2). Using preflashed film, fluorograph the gel (UNIT 10.2) at −70°C. Typical exposure times are overnight to 1 month.
ANALYSIS OF FATTY ACID LINKAGE TO PROTEIN 3
To determine the type of linkage by which the [ H]fatty acid is attached to the protein (i.e., thioester, oxyester, or amide linkage), the fatty acid is selectively cleaved from the protein. The most convenient method is to run replicate lanes on an SDS-PAGE gel, cut the lanes apart, and analyze each lane separately.
BASIC PROTOCOL 2
Materials Lysate or immunoprecipitate from [3H]fatty acid–labeled cells (see Basic Protocol 1, step 6) 0.2 M potassium hydroxide (KOH) in methanol Methanol 1 M hydroxylamine⋅HCl, titrated to pH 7.5 with NaOH 1 M Tris⋅Cl, pH 7.5 (APPENDIX 2E) Additional reagents and equipment for SDS-PAGE (UNIT 10.1), treating a gel with sodium salicylate (UNIT 14.3) or DMSO/PPO solution (UNIT 10.2), and fluorography (UNIT 10.2) 1. Run an SDS-PAGE gel (UNIT 10.1) using 20 µl lysate or immunoprecipitate from [3H]fatty acid–labeled cells in each of four lanes. 2. Cut the four lanes apart and transfer each lane to a 15-ml tube containing one of the following solutions: 0.2 M KOH in methanol Methanol 1 M hydroxylamine⋅HCl 1 M Tris⋅Cl, pH 7.5. Incubate 1 hr at room temperature with shaking. The 0.2 M KOH in methanol will cleave thio- and oxyesters, but not amides; 1 M hydroxylamine⋅HCl will rapidly cleave thioesters but will cleave oxyesters only poorly, and will not cleave amides. Methanol and 1 M Tris⋅Cl serve as controls.
3. Wash each gel strip three times, 5 min each time, with water. Treat the strips with sodium salicylate (UNIT 14.3) or DMSO/PPO solution (UNIT 10.2), and fluorograph using preflashed film at −70°C. Typical exposure times are overnight to 1 month. Cleavage is measured as a reduction in the fluorographic signal compared to those for controls, and can be quantitated by densitometric scanning of the lane or scintillation counting of excised bands. Bands with fatty acids linked to the protein by thioesters will be missing or greatly reduced in lanes treated with 0.2 M KOH in methanol and 1 M hydroxylamine⋅HCl; oxyesters will be greatly reduced or missing in the lane treated with 0.2 M KOH in methanol and may be slightly reduced in the lane treated with 1 M hydoxylamine⋅HCl; and amide linkages will not be affected by any of these treatments, so that proteins with amide-linked fatty acids will appear in all four lanes.
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Supplement 5
BASIC PROTOCOL 3
ANALYSIS OF TOTAL PROTEIN-BOUND FATTY ACID LABEL IN CELL EXTRACT Due to problems of interconversion of fatty acids by β-oxidation and chain elongation and of reincorporation of label into other metabolic precursors, the protein-bound label derived from [3H]fatty acids should be analyzed, especially for experiments with long labeling incubations. This protocol is used to determine how much of the label has been converted into other fatty acids or metabolites during the incubation; a different procedure must be used to determine whether the fatty acid on the protein of interest is different from that added during labeling (see Basic Protocol 4). Materials 0.1 M HCl/acetone, −20°C Lysate from [3H]fatty acid–labeled cells in 1% SDS (see Basic Protocol 1, step 4a or 5b) 1% (w/v) SDS 2:1 (v/v) chloroform/methanol Diethyl ether 6 M HCl (concentrated HCl diluted 1:1 with H2O) Hexane 5 to 10 µCi/µl [9,10(n)-3H]fatty acid standards (30 to 60 Ci/mmol; Amersham International, American Radiolabeled Chemicals, or NEN Research Products) in ethanol 90:10 (v/v) acetonitrile/acetic acid EN3HANCE spray (NEN Research Products) 15-ml polypropylene centrifuge tubes Mistral 3000i benchtop centrifuge with swing-out four-bucket rotor or equivalent Nitrogen gas 30-ml thick-walled Teflon container with an air-tight screw top 110°C oven Thin-layer chromatography tank RP18 thin-layer chromatography plate (e.g., Merck) Kodak XAR-5 film, preflashed Precipitate protein 1. Add 5 vol of 0.1 M HCl/acetone to 100 µl lysate from [3H]fatty acid–labeled cells in 1% SDS in a 15-ml polypropylene tube. Incubate ≥1 hr at −20°C. This will precipitate the protein.
2. Centrifuge 10 min at 1500 × g (1000 rpm in Mistral 3000i swing-out rotor), 4°C, to pellet the precipitate. Remove the supernatant and allow the pellet to air dry gently. Remove free label 3. Dissolve the pellet in a minimum volume of 1% SDS and transfer to a 1.5-ml microcentrifuge tube. Add 5 vol of 0.1 M HCl/acetone. Incubate ≥1 hr at −20°C. 4. Repeat steps 2 and 3. These precipitation steps concentrate the protein and remove much of the SDS and free label.
Analysis of Protein Acylation
5. Add 500 µl of 2:1 chloroform/methanol and vortex. Centrifuge 10 min at 1000 rpm, 4°C, and remove the supernatant. Repeat this step at least three times until no more free label is extracted into the organic solvent, as determined by scintillation counting of the supernatant.
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6. Add 100 µl diethyl ether to the pellet and vortex. Centrifuge 10 min at 1000 rpm, 4°C, and decant the supernatant. Dry the pellet by placing the microcentrifuge tube under a gentle stream of nitrogen. 7. Place the tube into a 30-ml thick-walled Teflon container with a air-tight screw top containing 1 ml of 6 M HCl. Flush the tube and container with nitrogen. Close the lid tightly and incubate in an oven 16 hr at 110°C. This hydrolyzes the fatty acids from the protein.
Extract hydrolyzed fatty acids 8. Extract the contents of the tube twice with 0.5 ml hexane and pool the extracts. Dissolve the residue in 0.5 ml of 1% SDS. Determine the radioactivity in the hexane extracts and in the residue. Fatty acids will be extracted into hexane, while label incorporated into sugars and amino acids will be mainly in the hexane-insoluble residue.
9. Evaporate the hexane extracts just to dryness with a gentle stream of nitrogen. Dissolve in 2 to 5 µl of 2:1 chloroform/methanol. It is important not to overdry the sample because it may then be difficult to dissolve.
Identify fatty acids 10. Preequilibrate a thin-layer chromatography tank with 90:10 acetonitrile/acetic acid for 15 min. 11. Spot resuspended hexane extract onto an RP18 thin-layer chromatography plate. Dilute 1 µl [9,10(n)-3H]fatty acid standards in ethanol to give 1 µCi/µl and spot 0.5 µl in parallel lanes. Develop the plate in 90:10 acetonitrile/acetic acid. Air dry the plate.
solvent front
origin [3H]palmitate
1
2
[3H]myristate
Figure 14.2.1 Fluorogram of thin-layer chromatography plate showing analysis of acylated nerve growth factor (NGF) receptor. Outside lanes, migration of 0.5 µCi [3H]palmitate and [3H]myristate standards. Lane 1, NGF receptor immunoprecipitated from cells labeled with [3H]palmitic acid. Lane 2, NGF receptor immunoprecipitated from cells labeled with [3H]myristic acid. Although the cells were labeled with different fatty acids, the protein was labeled with palmitic acid due to chain elongation of [3H]myristic acid to [3H]palmitic acid by the cells. Exposure for standards, 1 week; exposure for lanes 1 and 2, 1 month.
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12. Detect the radioactivity by spraying the plate with En3hance spray and exposing it to preflashed Kodak XAR-5 film overnight at −70°C. Identify the fatty acids. See Figure 14.2.1 for an example of a typical fluorogram. BASIC PROTOCOL 4
ANALYSIS OF FATTY ACID LABEL IDENTITY This protocol is used to identify the labeled fatty acid(s) associated with a specific protein band on an SDS-PAGE gel. Following electrophoresis, the band of interest is located either by comparison with molecular weight standards or by fluorography of a sodium salicylate–treated gel (see UNIT 14.3). DMSO/PPO–treated gels cannot be used. The labeled material is analyzed by thin-layer chromatography. Materials SDS-PAGE gel of lysate from [3H]fatty acid–labeled cells Additional reagents and equipment for analysis of protein-bound label (see Basic Protocol 3) 1. Excise the band(s) of interest from a wet or dried (fluorographed) SDS-PAGE gel. Wash three times with shaking, 5 min each, with 0.5 ml water. If the gel is fluorographed it should be treated with sodium salicylate (UNIT 14.3), not DMSO/PPO solution. The dried gel piece will rehydrate and the salicylate will be washed out during the washes.
2. Place the gel piece in a 1.5-ml microcentrifuge tube and lyophilize. 3. Hydrolyze the fatty acids in the band and identify them by thin-layer chromatography (see Basic Protocol 3, steps 7 to 12). REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
SDS sample buffer (for discontinuous systems), 5× 3.125 ml 1 M Tris⋅Cl, pH 6.8 (0.313 M final) 1 g SDS (10% final) 5 mg bromphenol blue (0.05% final) 5 ml glycerol (50% final) H2O to 10 ml Store at room temperature Add DTT to appropriate concentration just before use Warm the solution before use because it tends to solidify.
COMMENTARY Background Information
Analysis of Protein Acylation
The two most common acyl groups that modify proteins are 14-carbon myristic acid and 16-carbon palmitic acid (Fig. 14.2.2), and they occur both on different and on overlapping sets of proteins. By increasing the hydrophobicity of the protein, these fatty acid moieties can play a role in localization of the protein to the membrane and sometimes to specific types of membrane structures—e.g., glycolipid-enriched domains or caveolae (Rodgers et al.,
1994). Identifying the type of acylation of a protein and determining whether the level of modification can be affected by stimuli can provide more information on the mechanisms of action of proteins involved in signaling pathways. Fatty acids are used in labeling cells in vitro because they will diffuse across the plasma membrane and then be converted to acyl-CoA by the action of the enzyme acyl-CoA synthetase. This activated form of the fatty acid is
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O OH myristic acid
O OH palmitic acid
Figure 14.2.2 Structures of myristic and palmitic acids.
the substrate for protein-acyl transferases that transfer the acyl group to the protein. Tritiated fatty acids are most commonly used in biosynthetic labeling of proteins, but fluorescent analogs of fatty acids have been used to study palmitoylation of rhodopsin (Moench et al., 1994a,b), and [ω-125I]iodo–fatty acids have been used to study myristoylation of v-src (Peseckis et al., 1993). A wide variety of proteins are myristoylated, including viral structural proteins and many proteins involved in cell signaling, such as the α subunits of trimeric G proteins and the Src family of tyrosine kinases (Resh, 1994; Wedegaertner et al., 1995). Myristoylation most commonly occurs cotranslationally via an amide linkage to an NH2-terminal glycine residue, and is an irreversible modification. Myristoylation is dependent on the removal of the initiator methionine and has been shown to occur by the time the nascent polypeptide is 100 amino acids long. Inhibitors of protein synthesis will therefore block myristoylation. The enzyme responsible for NH2-terminal myristoylation, N-myristoyl transferase (NMT), was first isolated from Saccharomyces cerevisiae; both the yeast and human homologs have been well characterized and show different protein substrate specificities (Gordon et al., 1991). Substrate specificity of yeast NMT is determined by recognition of a sterically unhindered NH2terminal glycine followed by an amino acid sequence that conforms to the following criteria: an uncharged residue or proline at position 2; any amino acid at positions 3 and 4; serine (more common), alanine, or threonine at position 5; no proline at position 6; and, preferably, basic residues at positions 7 and 8. Residues C-terminal to this region appear to be important in recognition of the α subunits of trimeric G
proteins and the Src family of tyrosine kinases (Glover et al., 1988; Gordon et al., 1991). The NMT enzymatic reaction proceeds by formation of a myristoyl-CoA-enzyme complex, subsequent binding of the peptide, transfer of the myristate moiety to the peptide, release of CoA, and release of the myristoylated peptide (Rudnick et al., 1991). Several assays have been developed for this enzyme (e.g., King and Sharma, 1991; Rudnick et al., 1992; French et al., 1994). Inhibitors of myristoylation include 2-hydroxymyristic acid, which is converted in vivo to 2-hydroxymyristoyl-CoA, a potent inhibitor of NMT. Other inhibitors of acylation are fatty acid analogs and other compounds that inhibit acyl CoA synthetase and therefore block the conversion of fatty acids to acyl CoA; the latter include α-bromopalmitate (DeGrella and Light, 1980) and the triacsin family of antibiotics (Tomoda et al., 1987). The endogenous pool of fatty acid can be depleted by incubation with cerulenin, an antibiotic that inhibits de novo fatty acid biosynthesis. Myristoylation of a protein can be necessary for its activity, e.g., the transforming activity of Src (Kamps et al., 1986). Myristoylation alone may not be sufficient for a protein to be localized to the membrane; for this further lipid modification, such as palmitoylation or cooperative interaction with protein sequences, is required (Resh, 1994). Palmitoylation (with and without myristoylation) occurs on many signaling molecules, including rhodopsin, α-subunits of G proteins, and Src-family tyrosine kinases. Palmitoylation is a post-translational event occurring via a thioester linkage to a cysteine residue, and where it occurs with NH2-terminal myristoylation, the myristoylation is usually a prerequisite for palmitoylation. This may be because the enzyme responsible for palmitoylation recog-
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Analysis of Protein Acylation
nizes the myristoylated protein or, more likely, because myristoylation brings the protein to the correct cell location for palmitoylation to occur. Palmitoylation is also found in conjunction with isoprenoid modification at the C-terminus of proteins belonging to the Ras superfamily; it is responsible for the localization of these proteins to the membrane (Newman and Magee, 1993). This palmitoylation is dependent on prior modification of the protein by isoprenoid. G protein subunits such as the α1 subfamily and members of the Src family of tyrosine kinases have an NH2-terminal amino acid sequence of Met-Gly-Cys, where the initiator methionine is removed and replaced by myristate and the cysteine is palmitoylated (Resh, 1994). Palmitoylation is a reversible modification and has been shown to be dynamic in vivo, with the level of palmitoylation changing in response to various stimuli such as receptor activation, insulin, and growth factors (James and Olsen, 1989; Jochen et al., 1991; Wedegaertner et al., 1995). This phenomenon is thought to play a role in switching on or off signaling molecules by altering either the localization of the molecules or their presentation to other signaling molecules with which they interact. Depalmitoylation of proteins is catalyzed by the activity of protein-palmitoyl thioesterase (Camp and Hoffman, 1993). Protein-palmitoyl transferase has been localized to the Golgi/endoplasmic reticulum, and attempts to purify it to homogeneity have so far been unsuccessful. Analysis of the type of fatty acid linkage present should always be performed. Posttranslational myristoylation via a thioester linkage has been found in platelets (Muszbek and Laposata, 1993). The term “palmitoylation” is not strictly accurate because other long-chain fatty acids, such as stearate (18:0) and oleate (18:1), can also be thioesterified to proteins; "S-acylation" is now becoming more commonly used to describe this modification. The acylating activity seems to be relatively unspecific for chain length and degree of unsaturation, and utilizes acyl-CoAs partly in proportion to their abundance in the cell—hence the predominance of palmitate. This and the potential for interconversion of labeled fatty acids necessitate analysis of the chain length of the attached label. Other more complicated methods can be used to identify the type of fatty acid attached to the protein; these include gas chromatography, reversed-phase HPLC, and mass spectroscopy (Aitken, 1992). Structural and
sequence analysis of the acylated protein can be performed using mass spectrometry (Aitken, 1992). Further information about the cellular localization of acylated proteins can be found by extraction of the cell lysate in the presence of either Triton X-114 or X-100. Extraction with Triton X-114, which has the property of phase separation at 30°C, can distinguish between hydrophobic and hydrophilic proteins (Aitken, 1992). Extraction of membranes with Triton X-100 at 4°C and 37°C can identify proteins that are associated with glycolipid-enriched membranes (Rodgers et al., 1994).
Critical Parameters and Troubleshooting Cells should be subconfluent; it is recommended that the cells be plated at two split ratios so the culture closest to 70% to 80% confluency can be selected to be used for labeling. The pH of all solutions should be ≤7.5 in order to avoid hydrolysis of labile thioesters; the high pH of SDS-PAGE buffers does not seem to be a problem when using minigels, where the running times are relatively short. Dithiothreitol (DTT) or 2-mercaptoethanol should be used with care, as these will also cleave thioester bonds; for SDS-PAGE a maximum concentration of 20 mM DTT should be used and samples should not be boiled but incubated only 3 min at 80°C. The use of short labeling times (especially for palmitoylation) will reduce reincorporation of the label. The ability to detect myristoylation and/or palmitoylation of a protein using these methods will depend on the ability of the cells to take up radiolabeled compounds and incorporate them into metabolic precursors; the pool sizes of endogenous fatty acids and fatty acyl–CoA esters; the expression level of the proteinmyristoyl transferase and supposed proteinpalmitoyl transferase; the abundance, rate of synthesis, and turnover of the protein(s) and modification(s) of interest; and the efficiency of antibodies for immunoprecipitation.
Anticipated Results Typically, it is possible to detect myristoylated proteins using fluorographic exposure times ranging from <1 week for high-level expression of protein in transformed cells (e.g., Lck in LSTRA cells) to 1 to 4 weeks for a well-expressed endogenous protein.
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Time Considerations In vitro labeling experiments require 2 to 3 days for growing and labeling the cells. Harvesting the cells, preparing the lysate for SDSPAGE (with or without prior immunoprecipitation) and SDS-PAGE require 1 to 2 days plus time for fluorography. Linkage analysis of the labeled proteins takes 1 to 2 hr after the gel has been run. Analysis of the label in cell lysates requires 1 day to precipitate the protein, remove free label, and hydrolyze the sample. A second day is required to extract the label and perform thinlayer chromatography, and fluorography requires an overnight exposure. Analysis of the protein-bound label in gel bands takes ∼3 days after fluorography (dried gel) and a similar amount of time from a wet gel, except the analysis begins after the gel has been run.
Literature Cited Aitken, A. 1992. Structure determination of acylated proteins. In Lipid Modification of Proteins, A Practical Approach (N.M. Hooper and A.J. Turner, eds.) pp. 63-88. Oxford University Press, Oxford. Camp, L.A. and Hoffmann, S.L. 1993. Purification and properties of a palmitoyl-pro tein thioesterase that cleaves palmitate from H-Ras. J. Biol. Chem. 268:22566-22574. DeGrella, R.F. and Light, R.J. 1980. Uptake and metabolism of fatty acids by dispersed adult rat heart myocytes. J. Biol. Chem. 255:9739-9745. French, S.A., Christakis, H., O’Neill, R.R., and Miller, S.P.F. 1994. An assay for myristoylCoA:protein N-myristoyltransferase activity based on ion-exchange exclusion of [3H]myristoyl peptide. Anal. Biochem. 220:115-121. Glover, C.J., Goddard, C., and Felsted, R.L. 1988. N-Myristoylation of p60src. Biochem. J. 250:485-491. Gordon, J.I., Duronio, R.J., Rudnick, D.A., Adams, S.P., and Gokel, G.W. 1991. Protein N-myristoylation. J. Biol. Chem. 266:8647-8650. James, G. and Olson, E.N. 1989. Identification of a novel fatty acylated protein that partitions between the plasma membrane and cytosol and is deacylated in response to serum and growth factor stimulation. J. Biol. Chem. 264:2098821006. Jochen, A., Hays, J., Lianos, E., and Hager, S. 1991. Insulin stimulates fatty acid acylation of adipocyte proteins. Biochem. Biophys. Res. Commun. 177:797-801. Kamps, M.P., Buss, J.E., and Sefton, B.M. 1986. Rous sarcoma virus transforming protein lacking myristic acid phosphorylates known polypeptide substrates without inducing transformation. Cell 45:105-112.
King, M.J. and Sharma, R.K. 1991. N-Myristoyl transferase assay using phosphocellulose paper binding. Anal. Biochem. 199:149-153. Moench, S.J., Terry, C.E., and Dewey, T.G. 1994a. Fluorescence labeling of the palmitoylation sites of rhodopsin. Biochemistry 33:5783-5790. Moench, S.J., Moreland, J., Stewart, D.H., and Dewey, T.G. 1994b. Fluorescence studies of the location and membrane accessibility of the palmitoylation sites of rhodopsin. Biochemistry 33:5791-5796. Muszbek, L. and Laposata, M. 1993. Myristoylation of proteins in platelets occurs predominantly through thioester linkages. J. Biol. Chem. 268:8251-8255. Newman, C.M.H. and Magee, A.I. 1993. Post-translational processing of the ras superfamily of small GTP-binding proteins. Biochim. Biophys. Acta 1155:79-96. Peseckis, S.M., Deichaite, I., and Resh, M.D. 1993. Iodinated fatty acids as probes for myristate processing and function. J. Biol. Chem. 268:5107-5114. Resh, M.D. 1994. Myristoylation and palmitoylation of Src family members: The fats of the matter. Cell 76:411-413. Rodgers, W., Crise, B., and Rose, J.K. 1994. Signals determining protein tyrosine kinase and glycosyl-phosphatidylinositol-anchored protein targeting to a glycolipid-enriched membrane fraction. Mol. Cell. Biol. 14:5384-5391. Rudnick, D.A., McWherter, C.A., Rocque, W.J., Lennon, P.J., Getman, D.P., and Gordon, J.I. 1991. Kinetic and structural evidence for a sequential ordered bi bi mechanism of catalysis by Saccharomyces cerevisiae myristoyl-CoA:protein N-myristoyltransferase. J. Biol. Chem. 266:9732-9739. Rudnick, D.A., Duronio, R.J., and Gordon, J.I. 1992. Methods for studying myristoyl-CoA:protein N-myristoyltransferase. In Lipid Modification of Proteins, A Practical Approach (N.M. Hooper and A.J. Turner, eds.) pp. 37-61. Oxford University Press, Oxford. Tomoda, H., Igarashi, K., and Ømura, S. 1987. Inhibition of acyl-CoA synthetase by triacsins. Biochim. Biophys. Acta 921:595-598. Wedegaertner, P.B., Wilson, P.T., and Bourne, H.B. 1995. Lipid modifications of trimeric G proteins. J. Biol. Chem. 270:503-506.
Key Reference Casey, P.J. and Buss, J.E. 1995. Lipid modification of proteins. Methods Enzymol., Vol. 250. A compilation of methods used in studying lipid modification of proteins.
Contributed by Caroline S. Jackson and Anthony I. Magee National Institute for Medical Research London, United Kingdom
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Analysis of Protein Prenylation and Carboxyl-Methylation
UNIT 14.3
In the last fifteen years or so, several post-translational modifications of proteins with lipid moieties have been discovered. Analysis of fatty acylation is covered in UNIT 14.2. This unit describes methods for analysis of prenylation and the carboxyl-methylation that often accompanies it. The two prenoid groups that have been found attached to proteins— farnesyl (C15) and geranylgeranyl (C20)—are both derived from intermediates in the isoprenoid biosynthetic pathway that utilizes mevalonic acid. In the protocols described here, radiolabeled mevalonate is used to label these intermediates in either intact cells (see Basic Protocol 1) or in vitro translations (see Basic Protocol 2); the labeled intermediates then become incorporated into proteins. Alternatively, the preformed radioactive prenyl diphosphates can be used for in vitro translations (see Basic Protocol 2). Carboxyl-methylation of C-terminal prenylated cysteine residues often occurs subsequent to prenylation. Methods are given for radiolabeling of the methyl group with [3H-methyl]methionine, that is converted intracellularly into S-adenosylmethionine (see Basic Protocol 3), and for radiolabeling with preformed S-adenosyl[3H]methionine (see Basic Protocol 4). Three recent publications (Casey, 1990; Hooper and Turner, 1992; Casey and Buss, 1995) contain a number of methods related to analysis of lipid modification of proteins. PRENYLATION OF PROTEINS IN CULTURED CELLS To radiolabel intermediates in the isoprenoid biosynthetic pathway, cultured cells are incubated with [3H]mevalonic acid, which is the 6-carbon precursor to isopentenyl diphosphate. Isopentenyl diphosphate (5-carbon) is polymerized in a stepwise fashion to yield farnesyl (15-carbon) and geranylgeranyl (20-carbon) diphosphates, which are the known precursors of protein-bound prenyl groups. To maximize incorporation of exogenous [3H]mevalonate, the endogenous pool of unlabeled mevalonate is usually depleted by preincubation with an inhibitor of hydroxymethyl glutaryl–coenzyme A (HMG-CoA) reductase, the enzyme responsible for production of mevalonate from hydroxymethyl glutarate. Medium containing dialyzed serum is also used to reduce the endogenous pool.
BASIC PROTOCOL 1
Materials Cells Complete tissue culture medium appropriate for the cells, 37°C Complete tissue culture medium containing dialyzed serum, 37°C 10 mM mevinolin (see recipe) 1 µCi/µl R-[5-3H]mevalonic acid (10 to 40 Ci/mmol; DuPont NEN or American Radiolabeled Chemicals) PBS, pH 7.2 (APPENDIX 2E) ice-cold 1× SDS-PAGE sample buffer (UNIT 10.1) or RIPA lysis buffer (UNIT 13.2) Nitrogen gas Additional reagents and equipment for immunoprecipitation (UNIT 13.2), SDS-PAGE (UNIT 10.1), treating a gel with sodium salicylate (see Basic Protocol 3) or DMSO/PPO solution (UNIT 10.2), and fluorography (UNIT 10.2) NOTE: All reagents and equipment coming into contact with live cells must be sterile, and proper sterile technique should be used accordingly. NOTE: All culture incubations are performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified.
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Contributed by Anthony I. Magee
14.3.1
Current Protocols in Protein Science (1996) 14.3.1-14.3.11 Copyright © 2000 by John Wiley & Sons, Inc.
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1. On the day before the labeling experiment, split the cells into complete tissue culture medium. Set up cells at two split ratios and choose the culture closest to 70% to 80% confluency for labeling. The cultures should contain 106 to 107 cells.
2. On the day of the experiment, replace the culture medium with a minimum volume of prewarmed complete tissue culture medium containing dialyzed serum. Add 10 mM mevinolin to a final concentration of 50 µM. Preincubate the cells in this medium ≥60 min. Mevinolin is an inhibitor of HMG-CoA reductase. Alternative HMG-CoA reductase inhibitors are compactin/mevastatin (Sigma) and pravastatin (Pravachol, Bristol-Meyers Squibb).
3. Transfer enough 1 µCi/µl R-[5-3H]mevalonic acid to label all samples at a concentration of 50 to 200 µCi/ml into a microcentrifuge tube. If necessary, remove organic solvent by exposing the tube to a gentle stream of nitrogen, but do not dry the mevalonic acid completely. The volume of the [3H]mevalonic acid can be made up with sterile water to facilitate accurate pipetting. Alternatively, the organic solvent may be removed in a Speedvac evaporator. The mevalonic acid must not be heated above room temperature because it is unstable.
4. Add the partially dried [3H]mevalonic acid at the desired concentration to the culture medium. Incubate up to 24 hr. The most appropriate concentration of label and duration of incubation should be experimentally determined for each cell type.
5. Place the cells on ice and aspirate the medium. Wash the cells twice with ice-cold PBS. CAUTION: Radioactive medium and washes must be disposed of appropriately.
6. Lyse cells in the appropriate solution for further processing—e.g., for a 50-mm dish containing 4 × 106 cells, add 0.2 ml 1× SDS-PAGE sample buffer (UNIT 10.1) or 1 ml RIPA lysis buffer for immunoprecipitation (UNIT 13.2). Use the entire lysate for immunoprecipitation.
7. Analyze the lysate or immunoprecipitate by SDS-PAGE (UNIT 10.1). Use 25 µl of the whole cell lysate for a minigel or 50 µl for a standard gel; load all of the immunoprecipitate after solubilizing it in 25 or 50 µl of 1× SDS-PAGE sample buffer.
8. After SDS-PAGE, treat the gel with sodium salicylate (see Basic Protocol 3, step 6) or DMSO/PPO solution (UNIT 10.2; Magee et al., 1995) and fluorograph the gels at −70°C using preflashed film. Typical exposure times are 1 week to 1 month, although exposures up to 3 months may sometimes be required (see Time Considerations).
Analysis of Protein Prenylation and CarboxylMethylation
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PRENYLATION OF PROTEINS DURING IN VITRO TRANSLATION Commercially available reticulocyte lysates contain protein:prenyl transferases, so proteins can be labeled during in vitro translation of mRNA using [3H]mevalonic acid (Hancock et al., 1991; Newman et al., 1992; Giannakouros et al., 1993), which is converted to prenyl diphosphates in situ. When [3H]mevalonic acid is used, the identity of the incorporated label as farnesyl or geranylgeranyl must be experimentally determined by cleavage of the label followed by chain-length analysis using the method described by Farnsworth et al. (1990). In the author’s experience, commercially available reticulocyte lysates contain little endogenous mevalonate, so prenylation is highly dependent on added precursors. Alternatively, presynthesized [3H]prenyl diphosphates can be used; this allows an assignment of the protein-bound prenoid by labeling duplicate translations with [3H]farnesyl diphosphate or [3H]geranylgeranyl diphosphate. Incorporation of only one precursor into the protein unambiguously identifies the prenoid substituent. Incorporation of both suggests either that farnesyl diphosphate has been converted to geranylgeranyl diphosphate in the translation mix, or that the protein under investigation genuinely contains both prenoids. This question must be resolved by chain-length analysis (Farnsworth et al., 1990).
BASIC PROTOCOL 2
Materials Nuclease-treated rabbit reticulocyte lysate (Promega) Methionine-free amino acid mix (Promega) 100 mM L-methionine 1 µCi/µl R-[5-3H]mevalonic acid (10 to 40 Ci/mmol; DuPont NEN or American Radiolabeled Chemicals) mRNA: e.g., in vitro transcribed RNA derived from cDNA cloned into an appropriate vector such as SP64T, pGEMp9Z(f-), or pBluescript 6× SDS sample buffer (UNIT 10.1) Trichloroacetic acid Nitrogen gas 30°C water bath Additional reagents and equipment for immunoprecipitation (UNIT 13.2; optional), SDS-PAGE (UNIT 10.1), and fluorography (UNIT 10.2) 1. Using nuclease-treated rabbit reticulocyte lysate and methionine-free amino acid mix and following manufacturer’s instructions, prepare a master mix containing 25 µl translation mix for each mRNA to be translated and a negative control tube on ice. 2. Add 0.25 µl of 100 mM L-methionine (final concentration 1 mM) per reaction. 3. For each sample, aliquot 20 µCi R-[5-3H]mevalonic acid into a microcentrifuge tube. Remove the solvent under a stream of nitrogen gas. Alternatively, use 2.5 µCi of [1-3H]farnesyl diphosphate or [1-3H]geranylgeranyl diphosphate (30 to 60 Ci/mmol; American Radiolabeled Chemicals or DuPont NEN) per sample and remove the solvent under a stream of nitrogen gas.
4. Add the translation master mix to the tube containing labeled precursor. Incubate 10 min on ice. 5. Aliquot 25 µl of the mix into separate microcentrifuge tubes for each sample. 6. Start the translation by adding mRNA (e.g., 1 µg of in vitro–transcribed RNA). Incubate 90 min in a 30°C water bath. The optimum RNA concentration and duration of translation must be determined for each batch of reticulocyte lysate and each mRNA.
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7. At the end of the translation reaction, stop the reaction by adding 5 µl of 6× SDS-PAGE sample buffer or 1 ml RIPA lysis buffer, and process the samples for SDS-PAGE (see Basic Protocol 1 and UNIT 10.1), or immunoprecipitation (see Basic Protocol 1 and UNIT 13.2). Load the entire sample on the gel. Alternatively, add trichloroacetic acid to a final concentration of 15% (w/v) and process for prenoid analysis (Farnsworth et al., 1990). BASIC PROTOCOL 3
CARBOXYL-METHYLATION OF PROTEINS IN CULTURED CELLS Prenylation at C-terminal CXXX and CXC motifs is followed by carboxyl-methylation of the α-carboxyl group. This is performed by a membrane-associated methyl transferase that uses the universal methyl group donor S-adenosylmethionine (SAM). The methyl group of SAM can be radiolabeled in vitro by [3H-methyl]methionine, and it is then transferred onto the α-carboxyl group as an alkali-labile ester. This can be distinguished from alkali-stable methylations such as lysyl N-methylation by treating the sample with sodium hydroxide, which releases [3H]methanol. The released volatile [3H]methanol is collected by distillation into scintillation fluid and counted (Chelsky et al., 1985; Gutierrez et al., 1989). Labeled methionine will also be incorporated into the peptide backbone and is alkali-stable, thus providing an internal standard that can be used to calculate the stoichiometry of carboxyl-methylation. The length of the labeling incubation required to get an acceptable signal depends inversely on the expression level of the protein under study. Materials Cells Complete tissue culture medium (CM) appropriate for the cells Methionine-free tissue culture medium containing dialyzed serum (MFM) or 5% (v/v) complete culture medium with dialyzed serum (MFM-5%CM) 1 µCi/µl L-[3H-methyl]methionine (∼80 Ci/mmol, Amersham) PBS, pH 7.2 (APPENDIX 2E), ice-cold 1× SDS-PAGE sample buffer (UNIT 10.1) or RIPA lysis buffer (UNIT 13.2) 1 M sodium salicylate, or other water-soluble fluor (e.g., Enlightening, DuPont NEN, or Amplify, Amersham) Radioactive ink (see recipe) 1 M NaOH 1 M HCl Warm room or incubator at 37°C Additional reagents and equipment for immunoprecipitation (UNIT 13.2), SDS-PAGE (UNIT 10.1), and fluorography (UNIT 10.2) NOTE: All reagents and equipment coming into contact with live cells must be sterile, and proper sterile technique should be used accordingly. NOTE: All culture incubations are performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Label the cells 1. On the day before the labeling experiment, split the cells into fresh complete tissue culture medium.
Analysis of Protein Prenylation and CarboxylMethylation
Set up cells at two split ratios and choose the culture closest to 70% to 80% confluency for labeling. The cultures should contain 106 to 107 cells.
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For labeling times ≤8 hr 2a. On the day of the experiment, replace the culture medium with the minimum volume MFM, e.g., 3 ml for a 100-mm tissue culture dish. Incubate 1 hr. 3a. Add 200 µCi [3H-methyl]methionine per milliliter MFM. Incubate up to 8 hr. For labeling times of 8 to 24 hr 2b. On the day of the experiment, replace the culture medium with the minimum volume MFM-5%CM, e.g., 3 ml for a a 100-mm tissue culture dish. Incubate 1 hr. 3b. Add 50 µCi [3H-methyl]methionine per milliliter MFM-5%CM. Incubate 8 to 24 hr. 4. After labeling is complete, aspirate the medium and wash the cells twice with ice-cold PBS. 5. Lyse in the appropriate solution for further processing—e.g., for a 50-mm dish containing 4 × 106 cells, add 0.2 ml 1× SDS-PAGE sample buffer (UNIT 10.1) or 1 ml RIPA lysis buffer for immunoprecipitation (UNIT 13.2). Separate and fluorograph the samples 6. Analyze the immunoprecipitate or lysate by SDS-PAGE (UNIT 10.1). Use 25 µl whole-cell lysate for a minigel or 50 µl for a standard gel. Use the whole immunoprecipitate solubilized in 25 or 50 µl of 1× SDS-PAGE sample buffer. The α-carboxyl-methyl esters are stable at the moderately alkaline pH of the standard Laemmli gel system, whereas methyl esters of the side-chain carboxyls of aspartate and glutamate are quite labile.
7. Soak the gel 20 min in 1 M sodium salicylate. Dry the gel immediately at 60°C on a backing of filter paper. The gel must not be dried at too high a temperature.
8. Using radioactive ink, mark the filter paper to note the alignment of the gel. Fluorograph the gel at −70°C, using preflashed film. Expose the film for 1 week to 1 month. An inexpensive radioactive ink can made by washing out old radiolabel vials with a small volume (e.g., 0.1 ml) of an aqueous solution containing 0.01% (w/v) bromphenol blue.
9. Using the alignment marks, align the film exactly with the gel. Excise the radioactive band(s) of interest with a scalpel. Release the alkali-labile label 10. Place each gel piece into a separate 1.5-ml microcentrifuge tube. Carefully lower the open tube into a scintillation vial that contains enough scintillation fluid to come about half way up the side of the tube. 11. Add enough 1 M NaOH into the tube to cover the gel piece (usually 100 to 200 µl). Immediately cap the scintillation vial, leaving the tube open inside. Incubate overnight in a 37°C warm room or 37°C incubator. During this incubation, radioactive methanol is hydrolytically released and distilled into the scintillation fluid.
12. The next day, uncap the vial and carefully remove the microcentrifuge tube, taking care not to spill any of the contents into the vial. Recap the vial. 3
This scintillation vial contains the alkali-labile [ H]methanol.
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Analyze the sample for alkali-stable label 13. Neutralize the NaOH in the microcentrifuge tube by adding an equal volume of 1 M HCl. Transfer the contents of the tube to a new scintillation vial. This scintillation vial contains the alkali-stable [3H]label which represents [3H]methionine incorporated into the peptide backbone. However, alkali-stable methyl groups cannot be distinguished from backbone methionine in this way, so it is essential to have good reason to believe that other methyl groups are not present before alkali-stable counts can be used to calculate stoichiometry of carboxyl-methylation.
14. Add a volume of scintillation fluid to this vial equal to that used for the alkali-labile sample (step 12). 15. Count both samples in a scintillation counter using the tritium channel. Calculate the methylation stoichiometry 16. Calculate the methylation stoichiometry as: stoichiometry =
alkali−labile cpm × number of methionine residues in the protein alkali−stable cpm.
This calculation can only be made if the methionine content of the protein is known. Also the possibility that the initiator methionine might be removed must be taken into account. This can be predicted from the primary sequence (Aitken, 1992). BASIC PROTOCOL 4
CARBOXYL-METHYLATION DURING OR AFTER IN VITRO TRANSLATION Carboxyl-methylation can also be carried out during or after in vitro translation of mRNA (Newman et al., 1992; Giannakouros et al., 1993). Unlabeled mevalonic acid must be included in the reaction because carboxyl-methylation obligatorily follows prenylation. A source of the membrane-bound methyltransferase must also be added—either a crude 100,000 × g membrane pellet from cultured cells or commercially available dog pancreatic microsomes will suffice. These membranes also contain the protease required to remove the XXX residues from the C-terminal CXXX motif of prenylated proteins, which must occur before carboxyl-methylation can take place. Translation can be performed in the presence of preformed S-adenosyl[3H-methyl]methionine and membranes, or alternatively, the radiolabel and membranes can be added after translation has been completed. Methionine must be added to support translation, because the manufacturer’s amino acid mix lacks it. Materials Nuclease-treated rabbit reticulocyte lysate (Promega) Methionine-free amino acid mix (Promega) 100 mM L-methionine 100 mM mevalonic acid (see recipe) 1 µCi/µl S-adenosyl[3H-methyl]methionine (5 to 15 Ci/mmol, Amersham or ICN Biomedicals) Microsomal membranes: 100,000 × g pellet from cultured cells or canine pancreatic microsomal membranes (Promega; store at −70°C and thaw on ice immediately before use) mRNA: e.g., in vitro transcribed RNA derived from cDNA cloned into an appropriate vector such as SP64T, pGEMp9Z(f-), or pBluescript
Analysis of Protein Prenylation and CarboxylMethylation
Nitrogen gas Water bath at 30°C Additional reagents and equipment for immunoprecipitation (UNIT 13.2), SDS-PAGE (UNIT 10.2), and fluorography (UNIT 10.2)
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1. Using nuclease-treated rabbit reticulocyte lysate and methionine-free amino acid mix and following manufacturer’s instructions, prepare a master mix containing 25 µl of translation mix for each mRNA to be translated and a negative control tube on ice. 2. Add 0.25 µl of 100 mM L-methionine (final concentration 1 mM) and 1.25 µl of 100 mM mevalonic acid (final concentration 5 mM) for each 25 µl of master mix. 3. For each reaction, aliquot 30 µCi S-adenosyl[3H-methyl]methionine into a microcentrifuge tube. Remove the solvent under a stream of nitrogen. Alternatively, the solvent can be removed in a Speedvac evaporator.
4. Add the translation master mix to the tube that contains labeled precursor. Incubate 10 min on ice. 5. Add 3.6 equivalents of microsomal membranes for each 25 µl of master mix to the tube. Either total microsomal membranes (the 100,000 × g pellet) from cultured cells or tissue or a commercial preparation of canine pancreatic microsomal membranes can be used. The optimum amount of membrane to use will have to be determined experimentally, but a good starting point is 3.6 equivalents (Promega) per 25 µl of translation mix. One equivalent of microsomal membranes is that amount of membranes required to cleave the signal sequence of preprolactin from 50% of the translation products, as measured by a shift in mobility on SDS-PAGE gels (from Promega Canine Pancreatic Microsomal Membranes information sheet).
6. Aliquot 25 µl of the master mix into separate microcentrifuge tubes for each sample. 7. Start the translation by adding mRNA (e.g., 1 µg of in vitro transcribed RNA). Incubate the tubes 90 min in a 30°C water bath. The optimum RNA concentration and duration of translation must be determined for each batch of reticulocyte lysate and each mRNA. Alternatively, in vitro translation can be performed in the absence of label and membranes. In that case, add 30 µCi label and 3.6 equivalents of microsomal membranes to each tube after translation and incubate an additional 2 hr at 30°C.
8. At the end of the translation reaction, process the samples for SDS-PAGE (see UNIT 10.1) or immunoprecipitation (see UNIT 13.2) and fluorography (UNIT 10.2). REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Mevalonic acid, 100 mM Hydrolyze mevalonic acid lactone (Sigma) to the free acid (Kita et al., 1980) by dissolving 30 mg mevalonic acid lactone in 1.8 ml ethanol at 55°C. Add 0.9 ml of 0.6 M NaOH and 18 ml water. Incubate 30 min at room temperature. Adjust to pH 8 with HCl and dilute to 100 mM. Store indefinitely at −20°C. Mevinolin, 10 mM Dissolve 90 mg mevinolin in 1.8 ml ethanol at 55°C. Add 0.9 ml of 0.6 M NaOH and 18 ml distilled water. Incubate 30 min at room temperature. Adjust to pH 8 with HCl and dilute to 10 mM. Store indefinitely at −20°C in 500-µl aliquots. Mevinolin, also called Lovastatin and Mevacore, is available from Merck & Co. Inc. or Biomol Research Labs (see SUPPLIERS APPENDIX; Alberts, 1988). It is supplied as the lactone and must be hydrolyzed to the free acid as described in this recipe (Kita et al., 1980).
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Radioactive ink An inexpensive radioactive ink can be made by washing out old radiolabel vials with a small volume (∼0.1 ml) of an aqueous solution containing 0.01% (w/v) bromphenol blue. COMMENTARY Background Information Prenylation of proteins is a widespread posttranslational modification that occurs at C-terminal motifs of the type CXXX, CXC, or CC (where C is cysteine and X is any amino acid). Recent reviews give a thorough description of this field (Clarke, 1992; Giannakouros et al., 1993). Either the 15-carbon isoprenoid farnesol or the 20-carbon isoprenoid geranylgeraniol (see Fig. 14.3.1) can be added, depending on the exact sequence at the C-terminus. CX1X2X3 motifs where X3 is a small residue (e.g., methionine, serine, or alanine), are farnesylated; whereas if X3 is large and hydrophobic (e.g., leucine or phenylalanine), they are geranylgeranylated. The nature of the X1 and X2 residues also influences the ability of the motif to become prenylated such that some apparent CXXX sequences are not modified. The CXC and CC motifs are also geranylgeranylated on both cysteines. The prenoid moiety is attached by a thioether linkage to the sulphydryl group of cysteine; this is an extremely stable linkage that is irreversible in vivo, but it can be chemically cleaved in vitro to allow analysis of the chain length of the attached isoprenoid (Farnsworth et al., 1990). Prenylation occurs on otherwise cytosolic soluble proteins and is catalysed by a ubiquitous family of at least three soluble heterodimeric enzymes: farnesyl transferase, geranyl-
geranyl transferase I (active on CXXX motifs) and geranylgeranyl transferase II (active on CXC and CC motifs). In addition, geranylgeranyl transferase II has a third associated subunit called Rab escort protein that is required for activity. These enzymes utilize the activated diphosphates of farnesol and geranylgeraniol that are normal intermediates in the isoprenoid biosynthetic pathway, that leads to many important end products such as steroids, ubiquinone, and dolichols (Goldstein and Brown, 1990). Mevalonic acid is the precursor of these polyisoprenyl diphosphates and is readily taken up by most cells in culture. Hence radioactive mevalonate can be used to radiolabel prenyl chains biosynthetically. To maximize incorporation, endogenous mevalonate is usually depleted by preincubation of cells with an inhibitor of hydroxymethylglutaryl coenzyme A (HMG-CoA) reductase, the enzyme responsible for mevalonate synthesis. Both the ability of cells to take up exogenous mevalonate and the sensitivity to HMG-CoA reductase inhibitors can vary enormously and needs to be experimentally tested (e.g., the yeasts Saccharomyces cerevisiae and Schizosaccharomyces pombe; Giannakouros et al., 1992). Subsequent to and dependent on prenylation, other modifications can take place. CXXX motifs are proteolyzed to remove the XXX residues by a membrane-associated en-
OH geranylgeraniol
OH farnesol
Analysis of Protein Prenylation and CarboxylMethylation
Figure 14.3.1 Structures of isoprenoid groups used in post-translational modification.
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doprotease; then they are carboxyl-methylated on the free C-terminal α-carboxyl by a membrane-bound carboxyl-methyl transferase. This methyl ester group is chemically relatively labile; it is sensitive to mild alkali, although it is not as labile as the side-chain methyl esters of aspartate and glutamate, which can hydrolyze even during electrophoresis in mildly alkaline buffers such as are used in the standard Laemmli SDS-PAGE system (Laemmli, 1970). This lability is used experimentally as a test for the C-terminal α-carboxyl-methyl ester (Chelsky et al., 1985; Gutierrez et al., 1989). CXC motifs become doubly geranylgeranylated and are therefore also carboxyl-methylated because the methyl transferase recognizes a C-terminal prenylated cysteine. CC motifs, on the other hand, are not carboxyl-methylated. Many prenylated proteins also become palmitoylated S-acylated on nearby upstream cysteines by a membrane-bound palmitoyl transferase that is poorly characterized. Palmitoylation is discussed in UNIT 14.2. The primary function of prenylation is in membrane localization of substrate proteins that would otherwise be cytosolic because they are usually lacking in hydrophobic peptide sequences (Parenti and Magee, 1995). To perform this localization function, prenyl groups generally act in cooperation with themselves (e.g., for the doubly geranylgeranylated CXC proteins), with other modifications such palmitate moieties, or with peptide sequences. For example, the prenylated CXXX sequences of Ras proteins must cooperate with palmitoylation sites (in the case of H-ras, N-ras and K[A]-ras) or a polybasic region (as in K[B]-ras), which probably interacts with acidic membrane phospholipids (Hancock et al., 1991; Newman and Magee, 1993). These cooperating signals specify not only general membrane binding, as might be expected for a hydrophobic lipid moiety, but also targeting to specific subcellular destinations. The mechanism of this intracellular targeting is at present obscure. Membrane association of prenylated proteins is frequently reversible in vivo. This is achieved in several ways (Newman et al., 1992). Both palmitoylation and carboxylmethylation can be dynamic in vivo, thus allowing modulation of the hydrophobicity and/or charge of the C-terminus. Phosphorylation of residues near the C-terminus will change the charge, reducing the interaction with acidic phospholipid head groups. Thirdly, many prenylated proteins are removed from membranes by sequestration of their prenylated do-
mains by binding proteins that mask the hydrophobic group and form soluble 1:1 complexes. These mechanisms allow prenylated proteins to move between subcellular sites, which is often an essential part of their function.
Critical Parameters and Troubleshooting All of the labeling protocols in this unit that use cultured cells perform optimally when subconfluent cells are used. Certainly cells that are fully confluent should not be used. To ensure that cells at an appropriate density are available, it is helpful set up cultures at two different split ratios and use the ones that are most suitable. Occasionally a cell line is encountered that labels very poorly. Increasing the amount of radiolabel and/or the duration of labeling may lead to a satisfactory result. In the case of [3H]mevalonic acid labeling, the amount of mevinolin used and the duration of pretreatment can be increased. [14C]mevalonic acid has been used by some workers for radiolabeling prenyl moieties. Although tritium has a lower intrinsic detectability than 14C due to its weaker emission, the author nevertheless finds 3H is more sensitive because tritiated compounds can generally be obtained at much higher specific activities than 14C derivatives, and that more than compensates for the lower emission. For [3H-methyl]methionine labeling, it may help to reduce the concentration of unlabeled methionine in the medium to zero and to increase the duration of pretreatment. It is always useful to include an aliquot of the total cell lysate on the SDS-PAGE gel to assess the efficiency of labeling. Low levels of tritium require the highest sensitivity of fluorography for most rapid detection. The author has found that treating the gel with 2,5-diphenyloxazole in dimethylsulfoxide (DMSO/PPO solution, UNIT 10.2) is the most sensitive method (Laskey, 1980; Magee et al., 1995). However, this method cannot be used if further processing of the gel for prenyl group analysis or alkaline release of carboxyl-methyl groups is to be performed. In these cases, fluorography is best performed after treating the gel with the water-soluble fluor salicylic acid (Chamberlain, 1979) or a commercial equivalent; this method is approximately half as sensitive. Mevalonic acid and mevinolin tend to cyclize to form the lactone. If this is suspected (e.g., as indicated by a fall-off in labeling efficiency), the lactone can be hydrolyzed by the method of Kita et al. (1980) as described in the
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recipes (see Reagents and Solutions). Lipid substrates should never be dried to completion in plastic tubes because they are often irreversibly adsorbed to the surface. The ability to detect prenylation and/or carboxyl-methylation of any given protein using these methods will depend on a number of factors: the ability of the cells to take up radiolabeled compounds and incorporate them into metabolic precursors; the sensitivity of the cells to hydroxymethylglutaryl coenzyme A (HMGCoA) reductase inhibitors; the pool sizes of endogenous mevalonate, prenyl diphosphates, and S-adenosylmethionine (SAM); the expression level of the protein prenyl transferases and carboxyl-methyl transferase; the abundance, rate of synthesis, and turnover of the protein(s) and modification(s) of interest; and for immunoprecipitation, the efficiency of the antibodies.
Anticipated Results Typically it is possible to detect prenylated proteins using fluorographic exposure times of one to a few days for transfected cDNAs expressed at high levels, e.g., in COS cells, or in vitro translated mRNAs, 2 or 3 weeks for wellexpressed endogenous proteins, or up to 3 months for endogenous proteins expressed at low levels or in cells that incorporate mevalonate poorly. Similar exposure times are required for carboxyl-methylated proteins. However, because much of the radiolabel incorporated into these proteins is in the form of backbone methionine, the exposure time to detect the protein by fluorography may not be a good guide to the sensitivity of detection for alkali-labile methyl groups. This can be predicted if the primary sequence of the protein and therefore, its methionine content is known; it clearly becomes more of a problem with larger proteins and those with many methionines.
Time Considerations
Analysis of Protein Prenylation and CarboxylMethylation
Labeling experiments using cultured cells typically require ∼3 days to grow and label the cells. Harvesting and analyzing the cells by SDS-PAGE (with or without immunoprecipitation) take an additional 2 days, plus the time required for fluorography. After the fluorogram is developed, it takes 2 to 4 hr plus an overnight incubation to perform the analysis for alkalilabile and alkali-stable label, and the samples can be counted the next day. The in vitro translation protocols typically take a single full day for translation and labeling
and an additional day if immunoprecipitation is performed. The time required to complete the analysis is the same as that for the experiments using cultured cells.
Literature Cited Aitken, A. 1992. Structure determination of acylated proteins. In Lipid Modifications of Proteins: A Practical Approach (N.M. Hooper and A.J. Turner. eds.) pp. 63-88. IRL Press, Oxford. Alberts, A.W. 1988. Discovery, biochemistry and biology of lovastatin. Am. J. Cardiol. 62:10J-15J. Casey, P. (ed.) 1990. Covalent Modification of Proteins by Lipids. Methods, Volume 1. Casey, P., and Buss, J. (eds.) 1995. Lipid Modifications of Proteins. Methods Enzymol., Volume 250. Chamberlain, J.P. 1979. Fluorographic detection of radioactivity in polyacrylamide gels with the water-soluble fluor, sodium salicylate. Anal. Biochem. 98:132-135. Chelsky, D., Ruskin, B., and Koshland, D.E. Jr. 1985. Methyl-esterified proteins in a mammalian cell line. Biochemistry 24:6651-6658. Clarke, S. 1992. Protein isoprenylation and methylation at carboxyl-terminal cysteine residues. Annu. Rev. Biochem. 61:355-386. Farnsworth, C.C., Casey, P.J., Howald, W.N., Glomset, J.A., and Gelb, M.H. 1990. Structural characterization of prenyl groups attached to proteins. Methods 1:231-240. Giannakouros, T., Armstrong, J., and Magee, A.I. 1992. Protein prenylation in Schizosaccharomyces pombe. FEBS Letts. 297:103-106. Giannakouros, T., Newman, C.M.H., Craighead, M.W., Armstrong, J., and Magee, A.I. 1993. Post-translational processing of S. pombe YPT5 protein: In vitro and in vivo analysis of processing mutants. J. Biol. Chem., 268:24467-24474. Goldstein, J.L. and Brown, M.S. 1990. Regulation of the mevalonate pathway. Nature 343:425-430. Gutierrez, L., Magee, A.I., Marshall, C.J., and Hancock, J.F. 1989. Post-translational processing of p21ras is two-step and involves carboxyl-methylation and carboxy-terminal proteolysis. EMBO J. 8:1093-1098. Hancock, J.F., Cadwallader, K., and Marshall, C.J. 1991. Methylation and proteolysis are essential for efficient membrane binding of prenylated p21K-ras(B). EMBO J. 10:641-646. Hooper, N.M. and Turner, A.J. (eds.) 1992. Lipid Modification of Proteins: A Practical Approach. IRL Press, Oxford. Kita, T., Brown, M.S., and Goldstein, J.L. 1980. Feedback regulation of 3-hydroxy-3-methylglutaryl coenzyme A reductase in livers of mice treated with mevinolin, a competitive inhibitor of the reductase. J. Clin. Invest. 66:1094-1107.
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Laemmli, U.K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680-685. Laskey, R.A. 1980. The use of intensifying screens or organic scintillators for visualizing radioactive molecules resolved by gel electrophoresis Methods Enzymol. 65:363-369. Magee, A.I., Wootton, J., and de Bony, J. 1995. Optimized methods for detecting radiolabeled lipid-modified proteins in polyacrylamide gels. Methods Enzymol. 250:330-336. Newman, C.M.H., Giannakouros, T., Hancock, J.F., Fawell, E.H., Armstrong, J., and Magee, A.I. 1992. Post-translational processing of Schizosaccharomyces pombe YPT proteins. J. Biol. Chem. 267:11329-11336.
Parenti, M. and Magee, A.I. 1995. Fatty acid- and isoprenoid-linked membrane proteins. In Biomembranes, Vol. 1 (A.G. Lee, ed.) pp. 79-105. JAI Press, Greenwich, Connecticut.
Key References Casey, P. (ed.) 1990. See above. Casey, P. and Buss, J. (eds.) 1995. See above. These references contain compilations of methods for studying lipid modifications of proteins.
Contributed by Anthony I. Magee National Institute for Medical Research London, United Kingdom
Post-Translational Modification: Specialized Applications
14.3.11 Current Protocols in Protein Science
Supplement 5
Analysis of Oxidative Modification of Proteins
UNIT 14.4
Reactions between protein molecules and reactive oxygen species (ROS) often lead to the modification of certain amino acid residues such as histidine, lysine, arginine, proline, and threonine, forming carbonyl derivatives. Carbonylation of proteins has thus often been employed for the quantification of generalized protein oxidation. Besides carbonylation, other types of oxidative damage that have been investigated in depth are the modifications of cysteine, tyrosine and aspartate, or asparagine residues. Except for cysteine residues, whose oxidation is often determined by the loss of protein thiol groups, quantification of oxidative damage to tyrosine and aspartate residues is usually carried out by the measurement of specific oxidation products such as dityrosine, nitrotyrosine (when nitrogen species are the oxidants), and isoaspartate. This unit provides a variety of protocols for the determination of the protein oxidation products indicated above. A method for the quantification of protein carbonyls using spectrophotometric determination with a carbonyl specific reagent, 2,4-dinitrophenylhydrazine (DNPH), is described (see Basic Protocol 1), as are the details of immunoblot detection using anti-DNP antibodies to detect the carbonylation of specific proteins (see Support Protocol 1). Radiolabeling detection of total and specific proteins using tritiated sodium borohydride, another quantification method, is also included (see Basic Protocol 2 and Support Protocol 2) in this unit. Procedures are outlined for the selective determination of protein thiol groups radiolabeled with [14C] iodoacetamide followed by gel electrophoresis (see Basic Protocol 3). Procedures for dityrosine measurement by GC/MS are described (see Basic Protocol 4), as are preparation of dityrosine standards and nitrotyrosine detection by competitive ELISA (see Support Protocols 3 and 4). Methods for the detection of isoaspartate formation in proteins/peptides, analyzed by a methyl transfer reaction catalyzed by protein-L-isoaspartyl methyltransferase (PIMT), using [3H]methyl-S-adenosyl-L-methionine (SAM) as the methyl donor are given (see Basic Protocol 5), and the details of the different buffers needed for the electrophoretic separation of proteins containing isoaspartate residues are described (see Support Protocol 5). IMPORTANT NOTE: Tissue collection and preparation must be carried out in buffers supplemented with antioxidants, such as 100 µM diethylenetriaminepentaacetic acid (DTPA) and 1 mM butylated hydroxytoluene (BHT). The use of antioxidant buffers may be especially critical for the detection of trace amounts of oxidized products formed during aging or under oxidative stress. In addition, all buffers should be bubbled with nitrogen before use. SPECTROPHOTOMETRIC QUANTITATION OF PROTEIN CARBONYLS USING 2,4-DINITROPHENYLHYDRAZINE
BASIC PROTOCOL 1
Protein carbonyl groups can specifically react with 2,4 dinitrophenylhydrazine (DNPH) to generate protein conjugated hydrazones (protein-DNP) which have a peak absorbance around 360 nm. The use of DNPH thus provides an index for the quantification of protein carbonyl content in protein mixtures or purified proteins. Materials DNPH solution (see recipe) Protein solution 2 M HCl Contributed by Liang-Jun Yan and Rajindar S. Sohal Current Protocols in Protein Science (2000) 14.4.1-14.4.24 Copyright © 2000 by John Wiley & Sons, Inc.
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20% (v/v) trichloroacetic acid solution, ice-cold (TCA; see recipe) 1:1 (v/v) ethanol:ethyl acetate 0.2% (w/v) SDS/20 mM Tris⋅Cl, pH 6.8 (APPENDIX 2E) Bicinchoninic acid protein assay kit (BCA; Pierce Co.) Bovine serum albumin (BSA) Benchtop centrifuge Branson 2200 sonicator 1. Add 200 µl of DNPH solution to 1 ml of protein solution. Incubate the mixture at room temperature for 60 min. Prepare a blank by adding 200 µl of 2 M HCl without DNPH to one sample. Incubate under the same conditions. For tissue samples, start with 0.5 to 1.0 mg/ml protein. If higher protein concentrations are used for DNPH treatment, it will be very hard to dissolve the pellet after washing off the free DNPH (see Critical Parameters and Troubleshooting).
2. Add 1.2 ml 20% trichloroacetic acid (TCA) solution to the DNPH-treated protein solution and blank, then incubate on ice 10 min. Centrifuge the sample in a benchtop centrifuge for 10 min at 750 to 1000 × g, room temperature. 3. Wash the pellet by adding 3 ml 1:1 (v/v) ethanol:ethyl acetate, followed by centrifugation in a benchtop centrifuge for 10 min at 750 to 1000 × g at room temperature. Sonicate at full power, room temperature, in a Branson 2200 sonicator until the pellet is completely broken up. 4. Repeat step 3 twice. Solubilize the final pellet in 1 ml 0.2% (w/v) SDS/20 mM Tris⋅Cl, pH 6.8. If the carbonyl content of lipoproteins is to be determined, use a denaturing buffer of 3% (v/v) SDS/150 mM sodium phosphate buffer, pH 6.8 (APPENDIX 2E), to dissolve the final pellet (see Critical Parameters and Troubleshooting).
5. After the pellet is completely dissolved, pipet 100 µl of the protein solution for protein assay using the BCA kit. Use bovine serum albumin (BSA) as the protein standard. Initially, 6 M guanidine⋅HCl was used to dissolve the protein pellet, but samples dissolved in such a high concentration of guanidine⋅HCl are not suitable for further analysis by other techniques such as polyacrylamide gel electrophoresis (SDS-PAGE).
6. Scan the sample in a spectrophotometer from 320 nm to 450 nm. Use the peak absorbency around 360 nm to calculate the carbonyl content. Use the protein samples treated with HCl, but not with DNPH, as blanks. The extinction coefficient (ε) for DNPH is 22,000 M−1cm−1. Protein carbonyl content (nmol/mg protein) = (absorbance × 106/22,000)/mg protein = absorbance × 45.45/mg protein.
7. Identify specific carbonylated proteins by SDS-PAGE and immunoblotting (see Support Protocol 1). SUPPORT PROTOCOL 1
Analysis of Oxidative Modification of Proteins
IMMUNOBLOT DETECTION OF PROTEIN CARBONYLS To identify specific proteins that are carbonylated, DNPH-treated proteins can be further separated by gel electrophoresis according to standard techniques. Once the proteins are electrotransferred from the gel to a PVDF membrane, protein-bound DNP in individual proteins can be probed by the use of anti-DNP antibodies (also see UNIT 10.10). Materials DNPH-treated proteins (Basic Protocol 1) 5% (w/v) nonfat dry milk in Tris-buffed saline with Tween-20 (TBST; see recipe) Primary antibody (anti-DNP antibody; Sigma)
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Secondary antibody: may be horseradish peroxidase–conjugated; select on the basis of nature of primary antibody Tris-buffered saline with and without Tween-20 (TBS and TBST; see recipe) ECL detection solution (Amersham) Minigel electrophoresis unit and transfer unit (Bio-Rad; also see recipe for minigel recipes) Immobilon-P membranes (Millipore) UV-transparent plastic wrap X-ray film Additional reagents and equipment for SDS-PAGE (UNIT 10.1), staining gels with Coomassie blue (UNIT 10.5), and electroblotting proteins onto membranes (UNIT 10.7) 1. Separate DNPH-treated proteins by SDS-PAGE in a minigel electrophoresis unit (UNIT 10.1). DNPH-treated protein samples dissolved in 0.2% SDS/20 mM Tris buffer, pH 6.8 can be directly mixed with SDS-PAGE loading buffer and analyzed by SDS-PAGE. Two gels are usually run simultaneously, one for protein staining by Coomassie blue (UNIT and the other for immunoblot detection.
10.5),
2. Electroblot proteins from polyacrylamide gel to Immobilon-P membrane using a transfer unit per manufacturer’s instructions. For an efficient transfer, perform tank transfer rather than semidry transfer (see UNIT 10.7 for additional details).
3. Block the membrane by immersing it in 5% (w/v) nonfat dry milk TBST for at least 1 hr at room temperature. 4. Wash the membrane with at least 50 ml TBST for 10 min. Repeat twice. 5. Immerse the membrane in primary antibody solution for at least 1 hr. Primary antibody solution is prepared by diluting the primary antibody in TBST solution containing 0.2% (w/v) BSA. Dilution of anti-DNP antibody is highly variable. A range of 1:1,000 to 1:10,000 is recommended (see Critical Parameters and Troubleshooting). Overnight incubation at 4°C usually gives better results. Do not reuse the antibody.
6. Wash the membrane with at least 50 ml TBST three times, each time for 10 min. 7. Incubate the membrane with secondary antibody at room temperature for at least 3 hr. Antibody is diluted in TBST solution containing 0.2% (w/v) BSA. Secondary antibody dilution usually ranges between 1:10,000 and 1:25,000 (see Critical Parameters and Troubleshooting). Do not reuse the antibody.
8. Wash the membrane three times with at least 50 ml TBST, each time for 10 min. 9. Wash the membrane twice with at least 50 ml TBS (no Tween 20), each time for 5 min. 10. Immerse the membrane with ECL detection solution for 1 min at room temperature. ECL detection solution comes with the enhanced chemiluminescence (ECL) detection kit from Amersham.
11. Drain off excess detection solution and wrap membrane in plastic wrap. Gently smooth out any air bubbles. Immediately expose the membrane to X-ray film. Film is usually exposed to the membrane for 0.5 to 2 min.
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During immunoblot detection of protein carbonyls, it is preferable to include both a positive and a negative control on the SDS-PAGE. A positive control can be BSA oxidized by an oxidant such as hypochlorite followed by DNPH treatment as described (see Basic Protocol 1). A negative control is usually the sample not treated with DNPH. The inclusion of a positive control will indicate whether the anti-DNP antibody reacts properly, while the inclusion of a negative control will identify any proteins that cross-react with anti-DNP antibody. Occurrence of a cross-reaction between a protein and the antibody indicates that immunoblotting is not DNP-dependent for this particular protein. In such a case, anti-DNP antibodies from a different source may have to be tried. BASIC PROTOCOL 2
QUANTITATION OF PROTEIN CARBONYLS DERIVATIZED WITH TRITIATED SODIUM BOROHYDRIDE When protein concentration is a limiting factor, or when the studied proteins, such as heme-containing proteins, have absorption maxima around 360 nm, protein carbonyls cannot be determined by the use of DNPH. In such cases, tritiated sodium borohydride, which is able to convert protein carbonyl groups into protein-bound ethanol groups, can be used. Because of the introduction of tritium onto oxidized proteins, the method provides a quantitative measurement of protein carbonyl content. Materials Protein solution 3 M Tris⋅Cl, pH 8.6 (APPENDIX 2E) 0.5 M EDTA, pH 8.0 (APPENDIX 2E) [3H]NaBH4 working solution (see recipe) 2 M HCl 20% (v/v) trichloroacetic acid solution, ice-cold (TCA; see recipe) 1:1 (v/v) ethanol:ethyl acetate 0.2% (w/v) SDS/20 mM Tris⋅Cl , pH 6.8 (APPENDIX 2E) 0.5% (w/v) SDS/0.1 M NaOH BCA protein assay kit (Pierce) Scintisafe Plus 50% cocktail (Fisher Scientific.) Benchtop centrifuge Scintillation vials Additional reagents and equipment for determination of protein concentration (UNIT 3.4) CAUTION: Perform all incubations in a hood as tritium gas may be released during the reaction. Follow standard guidelines for handling, storage, and disposal of radioactive materials (APPENDIX 2B). 1. Prepare a 1 ml reaction mixture containing 86 mM Tris⋅Cl, pH 8.6 (add from 3 M stock), 0.86 mM EDTA (add from 5 M stock), 50 mM tritiated sodium borohydride (add from working solution described in Reagents and Solutions), and 0.5 to 1.0 mg of protein. 86 mM Tris buffer, pH 8.6 is used as the incubation buffer, as the authors have found that if phosphate buffer is used, proteins can be degraded during incubation. EDTA is included in the reaction mixture so that any potential carbonyl-independent labeling can be prevented.
2. Incubate 30 min at 37°C. Add 200 µl of 2 M HCl to stop the reaction. Add an equal volume of 20% TCA (final concentration 10%) to precipitate the proteins. Analysis of Oxidative Modification of Proteins
HCl also serves to render the alkaline reaction mixture acidic; otherwise, 10% TCA (final concentration) would not be sufficient to precipitate all proteins.
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3. Keep the solution on ice for 10 min and then centrifuge 10 min at 750 to 1000 × g, 4°C, in a benchtop centrifuge. 4. Wash the protein pellet with 1:1 ethanol:ethyl acetate at least three times. 5. Dissolve the final pellet in 1 ml 0.5% SDS/0.1 M NaOH. Determine protein concentration for each sample using a BCA protein kit (UNIT 3.4). Determination of protein concentration is necessary because the recovery after washing with ethanol:ethyl acetate is rarely 100% (see Critical Parameters and Troubleshooting).
6. Count the radioactivity of each sample by adding 4 ml Scintisafe Plus 50% cocktail to the sample and counting on a scintillation counter, or quantitate by electrophoretic separation and excision of a specific protein band (see Support Protocol 2). For the determination of the carbonyl content in a protein mixture, the sample can be counted right after the addition of scintillation cocktail. For the quantitation of the carbonyl content of a specific individual protein, the protein has to be separated from other proteins by means of SDS-PAGE, then processed as described in Support Protocol 2.
GEL ELECTROPHORETIC QUANTITATION OF PROTEIN CARBONYLS DERIVATIZED WITH TRITIATED SODIUM BOROHYDRIDE
SUPPORT PROTOCOL 2
To quantitate the carbonyl content of a specific protein in a protein mixture, as identified by immunoblot technique, protein samples derivatized with tritiated sodium borohydride can be separated by SDS-polyacrylamide gel electrophoresis. The target protein band on the gel can then be excised and dissolved in 30% hydrogen peroxide, and incorporated radioactivity measured by the use of a scintillation counter. Additional Materials (also see Basic Protocol 2) Tritiated protein sample (see Basic Protocol 2) 30% (v/v) hydrogen peroxide Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and staining gels with Coomassie blue R-250 (UNIT 10.5). CAUTION: Radioactive materials require special handling. See APPENDIX 2B concerning safe use of radioisotopes. 1. Label protein samples with [3H]NaBH4 (see Basic Protocol 2). Mix the labeled protein sample with SDS-PAGE loading buffer. TCA precipitation and washing with organic solvent is not needed.
2. Heat the sample 5 min at 95°C. Separate the tritiated protein sample by SDS-PAGE. 3. Stain the gel with Coomassie blue R-250 (UNIT 10.5) and destain until the bands can be clearly visualized. 4. Excise each band with a razor blade. 5. Put the excised gel bands in scintillation vials; incubate for 2 hr at 60°C to dry the gel slices. 6. Add 1 ml 30% hydrogen peroxide to each vial and incubate for 24 hr at 60°C. 7. Add 4 ml Scintisafe Plus 50% cocktail and keep the vials at 4°C for at least 12 hr. Use Scintisafe Plus 50% cocktail because it is specifically designed for gel slices with solubilizers such as hydrogen peroxide. Other cocktails, such as Scintiverse BD, become emulsive after mixing with the solubilized gels, which can decrease counting efficiency.
8. Count radioactivity with a scintillation counter.
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BASIC PROTOCOL 3
GEL ELECTROPHORETIC ANALYSIS OF PROTEIN THIOL GROUPS LABELED WITH [14C] IODOACETAMIDE To identify specific proteins in a tissue sample or a protein mixture exhibiting loss of thiol groups due to oxidative damage, the thiols can be labeled with [14C] iodoacetamide and analyzed by gel electrophoresis. The gels, after being dried onto filter papers, can be autoradiographed. Loss of thiol groups is inversely correlated with radioactive intensity shown by autoradiography. Oxidized proteins thus identified can be further quantitated by the use of liquid scintillation counting technique, exactly as described in Support Protocol 2 of this unit. Materials Protein sample 1% (w/v) SDS/0.6 mM Tris⋅Cl buffer, pH 8.6 (APPENDIX 2E) 2-mercaptoethanol, neat Nitrogen gas 500 mM [14C] iodoacetamide, 1 µCi/ml (Amersham) 500 mM nonradiolabeled iodoacetamide SDS-PAGE gels for Bio-Rad Mini gel system (see recipe): 10% resolving gel 4% stacking gel 10% (v/v) trichloroacetic acid (see recipe) Whatman 3MM filter paper X-ray film Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and staining gels with Coomassie blue R-250 (UNIT 10.5). CAUTION: Radioactive materials require special handling. See APPENDIX 2B concerning safe use of radioisotopes. 1. Dissolve protein sample (up to 1 mg) in 1 ml 1% SDS/0.6 M Tris⋅Cl, pH 8.6. The inclusion of 1% SDS is meant to completely denature the protein and expose buried thiol groups.
2. Add 10 µl 2-mercaptoethanol and incubate under nitrogen gas for 3 hr at room temperature. 3. Add 100 µl colorless 500 mM [14C] iodoacetamide solution to a final concentration of 1 µCi/ml, using 500 mM freshly prepared, nonradiolabeled iodoacetamide, in distilled water, to dilute the [14C] iodoacetamide to the proper level of radioactivity. Incubate at 37°C for 30 min with constant gentle agitation. IMPORTANT NOTE: Iodoacetamide is intrinsically unstable in light, especially in solution; reactions should therefore be carried out in the dark. Iodoacetamide solutions must be colorless. A yellow color indicates the presence of iodine, which will quickly oxidize protein thiol groups and prevent labeling. Free iodine can also modify tyrosine residues. The use of nonradiolabeled iodoacetamide will ensure an excess of the reagent over total thiol groups and a complete labeling.
4. Resolve proteins by SDS-PAGE with a 10% separating gel and 4% stacking gel using the Laemmli gel system (UNIT 10.1). After proteins are adequately separated, fix the gel in 10% TCA for 60 min. Stain with Coomassie blue R-250 and destain (UNIT 10.5). The sample incubation prior to loading should be done at 95°C for 5 min. Analysis of Oxidative Modification of Proteins
With method described in this protocol, actual radioactivity counts can be obtained for any given samples, which can be used for statistical data analysis. A phosphor imager may not
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be appropriate in terms of quantitation, as it only gives the relative band intensity and thus may not be able to detect slight differences between samples.
5a. For unknown protein samples: Dry gels onto Whatman 3MM filter paper and expose to X-ray film for up to two weeks at −70°C. To identify target proteins, purify the protein using two-dimensional gel electrophoresis (UNIT 10.4) and perform N-terminal microsequencing (UNIT 11.11), followed by a computer database search (UNIT 2.1).
5b. Given target proteins: Proceed to Support Protocol 2, step 4. QUANTIFICATION OF PROTEIN DITYROSINE RESIDUES BY MASS SPECTROMETRY In this protocol, trace amounts of dityrosine residues formed in proteins due to oxidative damage are first released by acid hydrolysis. The released dityrosine residues are then analyzed by mass spectrometry following derivatization with heptaflurorbutyric anhydride/ethyl acetate. The assay entails the use of radiolabeled dityrosine as the internal standard in order to be quantitative.
BASIC PROTOCOL 4
Materials Tissue sample o,o′-dityrosine internal standards, labeled and unlabeled (see Support Protocol 3) Nitrogen gas 6 M HCl/1% (v/v) benzoic acid/1% (v/v) phenol Argon 10% and 0.1% (v/v) TCA solution (see recipe) 50 mM NaHPO4/100 µM diethylenetriamine pentaacetic acid (DTPA), pH 7.4 25% methanol 1:3 (v/v) HCl/n-propyl alcohol 1:4 (v/v) pentafluoropropionic anhydride/ethyl acetate Ethyl acetate n-propanol 0.1% (w/v) trifluoroacetic acid (TFA) Supelclean SPE reversed-phase C-18 column (Supelco) Hewlett Packard 5890 gas chromatography equipped with a 12-m DB-1 capillary column interfaced with Hewlett-Packard 5988A mass spectrometer (see Chapter 16 for details of mass spectrometry) CAUTION: Radioactive materials require special handling. See APPENDIX 2B concerning safe use of radioisotopes. 1. Dry tissue sample under nitrogen and add 50 pmol 13C-labeled o,o′-dityrosine as an internal standard. 2. Hydrolyze the sample in a tube flushed with an inert gas (i.e., Argon) at 110°C for 24 hr in 0.5 ml of 6 M HCl/1% benzoic acid/1% phenol (UNIT 11.9). 3. Supplement the sample with 50 µl 10% (v/v) TCA solution and pass over a reversedphase C-18 column that has been previously washed with 12 ml of 50 mM NaHPO4/100 µM diethylenetriamine pentaacetic acid (DTPA), pH 7.4, followed by 12 ml of 0.1% TFA. 4. Elute amino acids with 25% methanol and dry under vacuum for derivatization. Dityrosine is stable to acid hydrolysis and ∼80% of the internal standard should be recovered from the C-18 column using this procedure.
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5. Convert amino acids to carboxylic acid esters by the addition of 200 µl of 1:3 (v/v) concentrated HCl/n-propyl alcohol. Heat for 10 min at 65°C. 6. Add 50 µl of 1:4 (v/v) pentafluoropropionic anhydride/ethyl acetate to prepare pentafluoropropionyl derivatives of the amino acids. Heat at 65°C for 30 min. 7. Dry the derivatized samples under nitrogen and redissolve in 50 µl of ethyl acetate. 8. Analyze 1 µl of the sample using a gas chromatograph interfaced with a mass spectrometer with extended mass range. Set the injector and ion source temperature at 250°C and 150°C, respectively. Obtain full scan mass spectra and selected ion monitoring with the n-propyl heptafluorobutryl and the n-propyl pentafluoropropionyl derivatives of both authentic and isotopically labeled dityrosine in the negative-ion chemical ionization mode with methane as the reagent gas. The mass spectrum of the n-propyl heptafluorobutyryl derivative of dityrosine includes a small molecular ion at mass-to-charge (m/z) 1228 (M−) and prominent ions at m/z 1208 (M−-HF) and 1030 (M−-CF3(CF2)CHO). Use m/z 1208 to quantify dityrosine.
9. For the analysis of tyrosine, dilute one aliquot of the derivatized amino acid (from step 7) 1:100 (v/v) with ethyl acetate. Inject 1 µl of the sample into the gas chromatograph with a 1:100 split prior to mass analysis. Maintain the initial column temperature of 120°C for 1 min and then increase to 220°C at 10°C/min. The mass spectrum of the n-propyl heptafluorobutyryl derivative of tyrosine reveals prominent ions at m/z 595 (M−-HF) and 417 (M−-CF3(CF2)2CHO). Use m/z 417 to quantify tyrosine. To ensure that interfering ions are not coeluting with the analyte, monitor the ratio of ion currents of the two most abundant ions of tyrosine and dityrosine in all analyses. Baselineseparate authentic and radiolabeled standards to exhibit retention times identical to those of analytes derived from tissue samples. SUPPORT PROTOCOL 3
PREPARATION OF o,o′-DITYROSINE STANDARD Dityrosine can be formed by horseradish peroxidase-catalyzed oxidation of tyrosine in the presence of hydrogen peroxide. Dityrosine produced in the mixture can then be purified by chromatographic methods. If radiolabeled dityrosine is to be made, radiolabeled tyrosine should be used as a starting material. Materials Horseradish peroxidase (grade I; Boehringer Mannheim) 0.1 M borate buffer, pH 9.1 (see recipe) 5 mM L-tyrosine (Sigma) or [13C6] L-tyrosine (Cambridge Isotope Laboratories) in 0.1 M borate buffer, pH 9.1 30% (v/v) H2O2 2-mercaptoethanol 0.01 M NaOH (APPENDIX 2E) 200 µM borate buffer, pH 8.8: diluted from 0.2 M borate buffer (see recipe) with H2O 2.75 × 19.5–cm DEAE cellulose chromatography column (Bio-Rad) 20 µM NaHCO3, pH 8.8 (see recipe) Concentrated and 100 mM formic acid 100 mM NH4HCO3
Analysis of Oxidative Modification of Proteins
Benchtop centrifuge 4 × 34.5–cm BioGel P-2 column (200-4-mesh; Bio-Rad)
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CAUTION: Radioactive materials require special handling. See APPENDIX 2B concerning safe use of radioisotopes. 1. Mix 10 mg horseradish peroxidase with 500 ml of 5 mM tyrosine solution prepared in 0.1 M borate buffer, pH 9.1. Use [13C6]tyrosine to prepare o,o′-[13C12]dityrosine if radiolabeled dityrosine is needed. Follow guidelines on the use and disposal of radioactive materials.
2. Add 142 µl 30% H2O2 and swirl the solution briefly. After incubation at room temperature for 30 min, add 175 µl 2-mercaptoethanol to the reaction mixture. Immediately freeze the solution in liquid nitrogen and lyophilize to dryness. 3. Dissolve the lyophilized substance in 250 ml of distilled water and adjust the pH to 8.8 with a few drops of 0.01 M NaOH. Load the solution onto a 2.75 × 19.75-cm DEAE column that has been pre-equilibrated with 20 µM NaHCO3, pH 8.8. Elute the column with 200 µM borate buffer, pH 8.8. Under these conditions, tyrosine and dityrosine will elute in the breakthrough fractions.
4. Pool and lyophilize the dityrosine-containing solutions. Resuspend the lyophilized material in 20 ml of cold water and centrifuge for 15 min at 1000 × g in a benchtop centrifuge. Extract the pellet with 15 ml of water and combine the two supernatants. Adjust the pH to 7.0 with formic acid and incubate at 0°C overnight. 5. Remove any precipitate by centrifuging for 10 min at 1000 × g in a benchtop centrifuge, 4°C. Load the supernatant on a BioGel P-2 column equilibrated with 100 mM NH4HCO3. Elute the column with 100 mM NH4HCO3 with a flow rate 40 ml/hr. Monitor the elution of dityrosine at 370 nm. Under these conditions, dityrosine elutes earlier than tyrosine. Therefore, fractions constituting the first peak should be collected. Dityrosine elution can also be monitored by the use of a fluorometer (λex = 280 nm; λem = 300 nm); however, the collected fractions need to be diluted before measurement.
6. Collect dityrosine fractions and lyophilize. Dissolve the lyophilized dityrosine in 20 ml of 100 mM formic acid. Adjust the pH to 2.5 by adding concentrated formic acid. Remove any precipitate that is formed during the pH adjustment by microcentrifuging for 10 min at 1000 × g, room temperature, in a benchtop centrifuge. Load the solution to a BioGel P-2 column (same as above) equilibrated with 100 mM formic acid. Elute the column with 100 mM formic acid, lyophilize the dityrosine-containing solution, and store at −20°C. The yield of dityrosine should be around 20%.
ANALYSIS OF PROTEIN-BOUND NITROTYROSINE BY A COMPETITIVE ELISA METHOD The availability of anti-nitrotyrosine antibodies provides a very sensitive method for the detection of protein-bound nitrotyrosine in tissue samples, namely enzyme-linked immunosorbant assay (ELISA). In this protocol, nitro-BSA is coated onto ELISA plates, and nitrotyrosines are quantitated by the use of anti-nitrotyrosine antibodies. Competition is accomplished by adding either a potentially nitrated protein sample or a known amount of nitrotyrosine (in the form of nitro-BSA) as a standard. Each competes with the coated nitrated proteins for antibody binding. The amount of antibody that binds to the coated nitro-BSA is inversely proportional to the amount of nitrated protein (sample or standard) present in the solution added to the well of the plate.
SUPPORT PROTOCOL 4
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Materials 10 µg/ml nitro-bovine serum albumin (nitro-BSA; Alexis Biochemicals) in plate coating buffer ELISA buffers (see recipe): Plate coating buffer 1× phosphate-buffered saline/Tween 20 (PBST) Blocking buffer 1× diethanolamine (DEA) buffer Protein sample Primary antibody: mouse anti-nitrotyrosine antibodies (Upstate Biotechnology) Secondary antibody: rabbit anti-mouse IgG conjugated with alkaline phosphatase Tris-buffered saline/Tween-20 (TBST; see recipe) 1 mg/ml p-nitrophenyl phosphate (5-mg tablets; Sigma) in DEA buffer 96-well ELISA plates Plastic wrap Plate reader Prepare the ELISA plate 1. The day before the assay, coat each of the wells of a 96-well ELISA plate that are to be used to assay protein samples with 100 µl of 10 µg/ml nitro-BSA in plate coating buffer (1 µg nitro-BSA/well). Leave the wells to be used for blanks empty. Plate coating may have to be optimized (see Critical Parameters and Troubleshooting).
2. Wrap the plate with plastic wrap and incubate at 4°C overnight. 3. Add 100 µl of each protein sample (1 to 10 µg/ml) to be assayed to 100 µl primary antibody diluted 1:500 with blocking buffer, and incubate at 4°C overnight. Add 100 µl of this solution to the designated well. Prepare nitro-BSA competitor the same way for a standard curve. The nitrotyrosine in BSA and the protein sample will compete with the coating antigen (nitro-BSA in this case) for antibody binding. The standard curve range is determined with serial dilutions of nitro-BSA between 10 µg/ml to 1 µg/ml. This will give a curve whose middle part is linear and flanked by a plateau curve on either side. The points that signal the start and the end of the linear part should be taken as the range for the standard curve. A series of dilutions of the protein sample may need to be tried in order to have readings that will fall within the linear range of nitro-BSA competition curve (see Critical Parameters and Troubleshooting).
Perform ELISA 4. The day of the assay, rinse the plate three times with 1× PBST, add 300 µl blocking buffer to each well, including blanks, and incubate for 60 min at room temperature. 5. Remove the blocking buffer from the plate and discard. Add 100 µl of a serial dilution of each of the protein samples prepared in primary antibody solution (step 3) to each sample well. Add 100 µl of a serial dilution of nitro-BSA prepared with primary antibody to the designated wells on the same plate to generate a standard curve. Add 100 µl 1× PBST, without antibody, to blank wells. 6. Incubate at room temperature for 2 hr. Analysis of Oxidative Modification of Proteins
7. Rinse three times with TBST. Add 100 µl of secondary antibody diluted 1:1000 with blocking buffer to each sample and standard well. Add 100 µl PBST to blank wells. 8. Incubate at room temperature for 2 hr.
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Detect nitrotyrosine 9. Discard reaction solutions from the wells and wash three times with PBST. Add 100 µl 1 mg/ml p-nitrophenyl phosphate to all wells including the blanks. Remove any air bubbles with a clean needle. p-Nitrophenyl phosphate is a substrate for alkaline phosphatase and will provide a colorimetric group upon cleavage.
10. After 30 min, clean the bottom of the plate with a Kimwipe and read at 450 nm on a plate reader. Estimate nitrotyrosine concentration using the nitro-BSA standard curve. Express nitrotyrosine concentration in the protein sample as an equivalent of nitrotyrosine in nitro-BSA. ENZYMATIC ANALYSIS OF ISOASPARTATE FORMATION A novel technique for the determination of isoaspartate formation involves the use of protein-L-isoaspartyl methyltransferase (PIMT). PIMT catalyzes methyl transfer reactions using methyl-S-adenosyl-L-methionine (SAM) as the methyl donor. Because PIMT is highly specific for protein isoaspartate residues, the use of tritiated SAM, where the methyl group is radiolabeled, provides a quantitative method for the determination of isoaspartate residues, formed in proteins/peptides.
BASIC PROTOCOL 5
Materials 0.2 M Bis-Tris buffer, pH 6.0 (see recipe) 10 µM [3H]methyl-S-adenosyl-L-methionine (5 to 15 Ci/mmol; [3H] SAM; NEN) Protein-L-isoaspartyl methyltransferase (PIMT; Promega Corporation or purified from a known source) 0.2 M NaOH (APPENDIX 2E) Safety-Solve II counting fluor (Research Products International) Sponge plugs (Jaece Industries): cut into small pieces Scintillation vials with extra caps CAUTION: [3H]methanol is volatile at room temperature. Perform all reactions under a hood and follow guidelines on handling, storage, and disposal of radioactive materials (APPENDIX 2B). 1. Add [3H] SAM to a final concentration of 10 µM to a protein sample solution in 0.2 M Bis-Tris buffer, pH 6.0, containing 40 U of PIMT in a microcentrifuge tube. The sample protein amount is usually between 0.5 and 1 mg and the total reaction volume is 50 µl. Blanks that do not have the protein to be analyzed and standards that have known concentrations of isoaspartate should be carried out under the same conditions at the same time. The Promega version of the enzyme comes included in the ISOQUANT Isoaspartate Detection kit.
2. Incubate the reactions for 30 min at 30°C. 3. Place the tubes on ice for 15 min to stop the PIMT-catalyzed methyl transfer reaction. 4. Microcentrifuge the tubes at 4°C for 2 min to bring all of the liquid to the bottom of the tube. Place on ice. 5. Add 50 µl of 0.2 M NaOH to hydrolyze the methyl esters so that [3H] methanol can be released from the assayed proteins. Tubes may be vortexed briefly to collect liquid.
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6. Immediately transfer 50 µl of each reaction solution to a sponge insert in a scintillation vial cap and place the cap on a vial containing 4 ml Safety-Solve scintillation cocktail. 7. Incubate the capped vials at 37°C for 60 min. Diffusion of methanol into the scintillation liquid is time-dependent. To obtain reproducible results, it is critical to incubate all reactions for the same length of time.
8. Remove the caps and replace with new caps that do not contain sponge inserts. 9. Count the samples in a scintillation counter. SUPPORT PROTOCOL 5
GEL ELECTROPHORETIC ANALYSIS OF ISOASPARTATE FORMATION To identify which proteins in a tissue sample or a protein mixture contain isoaspartate residues, the protein sample, incubated with [3H]SAM and PIMT (see Basic Protocol 5), can be further analyzed by SDS-PAGE (UNIT 10.1) after mixing with acid SDS-PAGE sample buffer (see recipes for acid SDS-PAGE buffers). Protein bands can then be excised from the gel and solubilized in 30% hydrogen peroxide, exactly as described in Support Protocol 2 of this unit; however, for the measurement of isoaspartate formation, acid SDS-PAGE (pH 2.4), rather than the Laemmli gel system (pH 8.3), should be performed (see recipes for acid SDS-PAGE). This is necessary, as high pH running buffer will hydrolyze the base labile isoaspartate methyl esters, leading to the release of [3H]methanol into the running buffer. Do not use sodium hydroxide to stop the PIMT-catalyzed reaction when the samples are to be analyzed by acid SDS-PAGE. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Acid SDS-PAGE buffers (pH 2.4) 2× resolving gel buffer: 0.1 M H2NaPO4 2% (w/v) SDS 6 M urea Adjust pH to 2.4 with HCl Store up to several months at 4°C 1× running buffer: 0.03 M NaH2PO4 0.1% (w/v) SDS 0.2 M acetate Adjust pH to 2.4 with HCl Store up to 1 year at room temperature
Analysis of Oxidative Modification of Proteins
2× sample buffer: 2% (w/v) SDS 6 M urea 10% (v/v) glycerol 0.01% (w/v) pyronin Y dye 16.5 mM NaCl Adjust pH to 1.4 with HCl Store up to 1 year at −20°C
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2× stacking gel buffer: 2% (w/v) SDS 6 M urea 0.033 M NaCl Adjust pH to 1.4 with HCl Store up to 2 months at 4°C Bis-Tris buffer, 0.2 M, pH 6.0 41.84 g bis-Tris (Sigma) Adjust volume to 1 liter with H2O Adjust pH with concentrated HCl Borate buffer, 0.1 M, pH 9.1 6.184 g boric acid Adjust volume to 1 liter with H2O Adjust pH with 1 M NaOH Store up to 2 months at room temperature Borate buffer, 0.2 M, pH 8.8 12.37 g boric acid Adjust volume to 1 liter with H2O Adjust pH with 1 M NaOH Store up to 2 months at room temperature DNPH solution Dissolve 198 mg 2,4-dinitrophenylhydrazine (DNPH; mol. wt. 198.1) in 100 ml 2 M HCl (10 mM DNPH final). Store up to 1 year at room temperature in the dark since DNPH can be destroyed by light. DNPH may require 1 to 2 hrs of constant stirring for complete solubilization.
ELISA buffers Blocking buffer: Prepare 0.2% (w/v) gamma globulin (Sigma) in PBST (see recipe). Filter this buffer through 0.45 µm filter and store up to 1 week at 4°C. 1× diethanolamine (DEA) buffer: 24.25 ml 98% diethanolamine 200 ml Milli-Q H2O 0.25 ml 20% NaN3 25 mg MgCl2⋅6H2O Adjust pH to 9.8 with concentrated HCl Bring to 250 ml with Milli-Q water 10× phosphate-buffered saline/Tween 20 (PBST): 80 g NaC1 2 g KH2PO4 11.5 g Na2HPO4 2 g KC1 5 ml Tween 20 10 ml 20% (w/v) NaN3 Dilute to a total volume of 2 liters with Milli-Q water Store up to 2 months at room temperature Post-Translational Modification: Specialized Applications
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Plate coating buffer: 0.795 g Na2CO3 1.465 g NaHCO3 0.5 ml 20% (w/v) NaN3 Bring to 500 ml with Milli-Q water If necessary, adjust the pH with 1 M NaOH to ensure it is ∼9.6 [3H]Sodium borohydride ([3H]NaBH4) working solution Add 5 ml 1 M nonradiolabeled sodium borohydride dissolved in 0.1 M NaOH to the original bottle in which tritiated sodium borohydride (activity 222.3 mCi/mmol; NEN Life Science Products) was received. Divide the solution into 100-µl aliquots in microcentrifuge tubes (0.5 ml/tube) and store up to 2 years at −80°C. NaHCO3, 20 µM, pH 8.8 16.8 mg NaHCO3 Adjust volume to 1 liter with H2O Adjust pH with 1 M NaOH Dilute 1:10 just before use Store up to 2 months at room temperature Nitro-BSA standard Adjust the pH of a 5 mg/ml BSA solution (in water) to 3.5 with acetic acid. Add concentrated sodium nitrite (200 mM) to the BSA solution to obtain a final concentration of 1 mM. Incubate the reaction on a rotator at 37°C for 24 hr. The color change of the solution from colorless to yellow after the incubation usually indicates the presence of nitrotyrosine. Dialyze the nitrated BSA solution overnight against ELISA coating buffer. Determine nitrotyrosine concentration at 430 nm using a spectrophotometer (ε = 4100 M-1cm-1, pH > 8.5). Store up to 2 months at 4°C. SDS-PAGE gels (for Bio-Rad Mini gel system) 10% resolving gel: 6.1 ml 2× resolving buffer (Bio-Rad) 2.5 ml 40% 29:1 acrylamide/bisacrylamide 0.1 ml 0.06%, w/v FeSO4 0.1 ml 1%, w/v ascorbic acid 1.1 ml distilled water 0.1 ml 0.3% H2O2 Store up to 6 months at 4°C 4% stacking gel: 4.0 ml 2× stacking gel buffer (Bio-Rad) 0.5 ml 40% 29:1 acrylamide/bisacrylamide 0.065 ml 0.06%, w/v FeSO4 0.065 ml 1%, w/v ascorbic acid 0.305 ml distilled water 0.065 ml 0.3% H2O2 Store up to 6 months at 4°C IMPORTANT NOTE: FeSO4, ascorbic acid, and H2O2 are used as gel polymerization catalysts; prepare fresh.
Analysis of Oxidative Modification of Proteins
Trichloroacetic acid (TCA) solution, 20%, 10%, and 0.1% (v/v) Prepare TCA stock solution by adding 227 ml water to a 500-g bottle of TCA. Add 10 ml of TCA stock solution to 40 ml of water to prepare 20% (v/v) TCA. Dilute to appropriate percentage with water. Store up to 1 year at room temperature.
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Tris-buffered saline with and without Tween-20 (TBST and TBS) 100 mM Tris⋅, pH 7.5 (APPENDIX 2E) 0.9% (w/v) NaCl 0.1% (v/v) Tween-20 (omit for TBS) Store up to 2 months at 4°C. COMMENTARY Background Information Proteins are acknowledged to be one of the major targets of endogenously generated reactive oxygen species (ROS) (Stadtman, 1992). The study of protein oxidative modifications not only contributes to the elucidation of the mechanisms of ROS-mediated damage, but may also aid in understanding how protein oxidation is linked to losses in physiological functions under pathological processes or during aging. Furthermore, the identification of the protein targets of oxidative damage may potentially lead to the development of strategies that may ameliorate oxidative stress. Several factors seem to be involved in intracellular accumulation of oxidized proteins. One major factor is the cellular level of oxidative stress, determined by the balance between the rate of ROS generation and the efficiency of antioxidative defenses (Stadtman and Berlett, 1997). The other factor determining the steady-state amount of oxidized proteins is the activity of proteolytic enzymes (Stadtman and Berlett, 1997). It is now firmly established that mild oxidation of proteins facilitates their eventual degradation by proteases, while heavily oxidized proteins in which crosslinks usually occur are resistant to proteolysis (Davies et al., 1987). Oxidative damage to proteases themselves can further attenuate cellular ability to degrade oxidized proteins (Agarwal and Sohal, 1994). Analysis of protein carbonylation Some known mechanisms by which protein carbonylation occurs are shown in Figure 14.4.1. The reaction between DNPH and protein carbonyls, shown in Figure 14.4.2, was initially used for the measurement of metal catalyzed oxidative damage to glutamine synthase (Levine, 1983). The procedure, although very useful for the determination of protein carbonyl content in tissue samples, is not selective and subject to interference by proteins containing chromophores that have absorbance near 360 nm. In contrast, immunoblotting is highly selective and can detect carbonylation of individual proteins, but the method is not meant to provide quantitative information. The borohydride reduction method, when used in conjunction with gel electrophore-
sis, is both selective and quantitative (Yan and Sohal, 1998a). Using protein carbonyls as an index, oxidative damage to proteins has been found to be associated with aging and oxidative stress as well as a number of pathologies (Stadtman, 1992). Moreover, protein oxidation during aging has been recently established to be a highly selective phenomenon. Using the housefly as a model system, the authors’ laboratory has demonstrated that in flight-muscle mitochondria, only aconitase and adenine nucleotide translocase exhibited a discernible age-dependent increase in protein carbonylation and a corresponding loss in their activities (Yan et al., 1997; Yan and Sohal, 1998b). The four protocols for the measurement of protein carbonyls described in this unit are relatively simple and easy to perform, without the requirement of elaborate instrumentation. Analysis of protein carbonylation by methods other than those described here may also be used, should the relevant equipment be readily accessible. One such procedure requires HPLC separation, which has the advantage that the removal of free DNPH is unnecessary because the HPLC column can separate proteins from small molecules (Levine et al., 1994). HPLC requires a very small amount of proteins and can monitor both protein and protein-bound DNP absorption at two wavelengths simultaneously. The derivatization of protein carbonyls by DNPH, however, needs to be carried out in solutions that are compatible with HPLC columns. Another method, which also involves the use of anti-DNP antibodies, is the ELISA measurement of protein carbonyls (Buss et al., 1997). This method is quite sensitive and may be used for determination of whole tissue carbonyls, but it is only semiquantitative and lacks selectivity. Recently, a fairly new method, the MALDIMS technique (UNIT 16.2 and UNIT 16.3), has been applied for the determination of protein carbonyls. The method works very well for small peptides or proteins such as cytochrome c, but for large peptides, the proteins need to be digested with proteases. The resulting small peptides are then separated by HPLC. A potential problem for large proteins with this method is that if a large
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A CHO CH2
+
PNH2
CHO
CH=NP
CH-NHP
CH=NP
CH2
CH2
CH2
CHO
CHO
CHOH
P N
P-His
N R O
OH
B
S-P
R
polyunsaturated fatty acids
“oxygen”
O
P-HS
R O
OH OH NH-P P-NH2
R O
OH CHO
C P-NH2
C-NH-P
CHO +
(HCOH)n
O2,Me2+
C=O (HCOH)n –1
P-NH2
C=O (HCOH)n –1
R R
R
Figure 14.4.1 Generation of protein carbonyls by glycation and glycoxidation and by reactions with lipid peroxidation products of polyunsaturated fatty acids. (A) Reactions of protein amino groups (PNH2) with the lipid peroxidation product, malondialdehyde. (B) Michael addition of 4-hydroxy-2-nonenal to protein lysine (P-NH2), histidine (P-His), or cysteine (PSH) residues. (C) Reactions of sugars with protein lysyl amino groups (P-NH2). “Me” represents “metal ions.”
number of small peptides are generated after protease digestion, HPLC separation of these peptides can become very tedious.
Analysis of Oxidative Modification of Proteins
Analysis of protein thiol groups Protein thiol groups (mainly cysteine residues) not only stabilize and maintain the three-dimensional conformation of many proteins, but are also involved in biological redox couplings. Due to their high reactivity, protein thiol groups are very sensitive to oxidative damage induced by a variety of oxidation systems. Loss of protein thiol groups has been demonstrated during aging and in disease (Hughes et al., 1980; McKenzie et al., 1996). Most of the currently available methods for quantitative analysis of protein thiol groups de-
pend upon the formation of chemical derivatives. Due to the nucleophilic property of the –SH group, alkylation of cysteine thiol groups by alkylators such as iodoacetamide is widely used for either blocking or quantification of protein-bound thiols. There are two advantages when iodoacetamide is used. First, iodoacetamide does not create a carboxylate functional group; therefore, no new negative charges are introduced into the protein (Fig. 14.4.3). This may be necessary when the alkylated protein is to be analyzed by isoelectric focusing gel electrophoresis. Second, the bond formed from the reaction of iodoacetamide and a thiol group is a stable thioether linkage that is irreversible under normal gel electrophoretic
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NO2
NHNH2
protein—C=O + 2 ON
(1)
NO2
2ON
protein—C=O + NaB[3 H]4
NHN==C—protein
protein—CH2 O[3H]
(2)
Figure 14.4.2 Methods for labeling protein carbonyls: (1) Derivatization of protein carbonyls with 2,4- dinitrophenylhydrozine (DNPH), forming protein conjugated dinitrophenylhydrozones. (2) Derivatization of protein carbonyls with tritiated sodium borohydride.
protein—SH + ICH2[ 14C]ONH2
protein—SCH2[ 14C]ONH2
Figure 14.4.3 Radiolabeling of protein thiol groups with [14C]iodoacetamide.
conditions. Furthermore, when radiolabeled iodoacetamide is used, the reaction provides a highly quantitative method for the measurement of protein thiol groups. The use of iodoacetamide, together with gel electrophoresis, has led to a successful identification of glyceraldehyde-3-phosphate dehydrogenase as the enzyme that exhibits the most loss of protein thiol groups during inflammatory bowel disease (McKenzie et al., 1996). Another classical method for the determination of protein thiol groups involves the use of 5,5’-dithiobis (2-nitrobenzoic acid) also known as DTNB or Ellman’s reagent (Ellman, 1959). DTNB undergoes disulfide interchange with the release of 5-thio-2-nitrobenzoic anion (TNB), which has an absorption maximum at 412 nm and can be quantitated spectrophotometrically. The disadvantage of the DTNB method is that TNB is highly subject to reoxidation in an alkaline solution in the presence of oxygen and metal ions and its molar extinction coefficient changes slightly as a function of the solvent system used. In addition, DTNB itself can undergo hydrolysis
of its disulfide bond at a pH higher than 9. Moreover, without prior protein purification, DTNB cannot be used for the quantification of individual proteins that exist in tissue homogenates and may have undergone the most loss of thiol groups caused by oxidative damage during aging or in diseases. Analysis of tyrosine modification Dityrosine formation. o,o′-Dityrosine is formed either when a hydroxyl radical crosslinks tyrosines or by the reaction between protein bound tyrosine and a tyrosyl radical (Fig. 14.4.4). Dityrosine exhibits intense fluorescence at ∼400 nm and thus can be measured with an excitation wavelength of ∼315 nm in alkaline solutions or ∼284 nm in acidic solutions. Based on this property, dityrosine, formed in proteins oxidized in vitro, have been frequently measured by HPLC equipped with a fluorometer as the detection device (Giulivi and Davies, 1993). A disadvantage is that molecules that co-elute, but structurally differ from dityrosine may confound the procedure. Moreover, the HPLC
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OH
2
OH
O
OH
(4)
2
R tyrosine
R
R
R
o,o′-dityrosine
Figure 14.4.4 Formation of one molecule of o,o′-dityrosine from two molecules of tyrosine via tyrosyl radical intermediates.
OH
OH NO2 reactive nitrogen species
R tyrosine
(5)
R 3-nitrotyrosine
Figure 14.4.5 Formation of nitrotyrosine via reactive nitrogen species-mediated nitration of tyrosine.
Analysis of Oxidative Modification of Proteins
fluorometric method has not yet been successfully used for the detection of dityrosine formation during natural aging, suggesting that the amount of dityrosine in proteins, without challenge by severe oxidative stress, is below HPLC detection limit. In contrast, the GC-MS method described in this unit for dityrosine detection is highly sensitive and can structurally distinguish dityrosine from other tyrosine modified derivatives, thereby reducing potential confusions caused by compounds that coelute with dityrosine during chromatography. GC/MS analysis also permits the use of a stable, isotopically labeled internal standard which, other than its heavy isotope, is structurally identical to dityrosine, and therefore behaves identically during sample preparation and analysis. Inclusion of such a standard also corrects for the dityrosine loss during processing and thus increases the precision of quantitative measurements. Using the GC/MS method, dityrosine concentrations in various mouse tissues have been found to be elevated during aging and are signifi-
cantly decreased by caloric restriction (Leeuwenburgh et al., 1997). However, this method lacks selectivity when tissue samples are used as experimental materials, since no distinction can be made in the degree of dityrosine content among different protein species. Nitrotyrosine formation. Nitrotyrosine is formed by reactions between tyrosine and reactive nitrogen intermediates (Fig. 14.4.5). Its level is elevated, especially during inflammation. Peroxynitrite, produced by the reaction between the nitrogen monoxide radical and superoxide, has been proposed to be the major oxidant that causes nitrotyrosine formation in living systems (Ischiropoulos et al., 1992). Nitrotyrosine can be easily determined spectrophotometrically as its maximum absorbance ranges from 350 nm to 450 nm, shifting from 365 nm (colorless) at pH < 3 to 428 nm (yellow) at pH > 9. However, the amount contained in tissue samples without oxidative challenge is far below the detection limit of a spectrophotometer. The ELISA method described in this unit,
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HN
O
O
CNH2
COH
NH
C
HN
NH
C
O
O
asparaginyl peptide
aspartyl peptide
NH2
H2O O
C N HN
C O
succinimide intermediate
O
O
C
HN
CO–
NH
CO–
HN
C
NH
O
O
isoaspartyl peptide [70-85%]
aspartyl peptide [15-30%]
Figure 14.4.6 Formation of isoaspartate by the deamidation of asparagine or the isomerization of aspartate.
although only semiquantitative, is very sensitive for the measurement of nitrotyrosine in tissue samples. This method is especially useful when individual proteins in tissue samples need not be analyzed. For identification of specific proteins that might exhibit selective nitration during aging or under oxidative stress, immunoblot detection using anti-nitrotyrosine antibodies may be preferred. Analysis of isoaspartate formation Isoaspartate residues are usually formed from the deamidation of asparaginyl residues or isomerization of aspartyl residues (Fig. 14.4.6).
Formation of isoaspartate residues in a protein not only alters the protein’s structure, but may also cause the loss of the protein’s activity. Although isoaspartate formation was initially found in in vitro aging of proteins stored at neutral pH (Graf et al., 1971), the discovery of the enzyme protein-L-isoaspartyl methyltransferase (PIMT) suggests that isoaspartate formation in proteins is physiologically relevant (Clark, 1985). Indeed, it has been found that a variety of tissue samples are able to be tritiated when incubated in the presence of PIMT and [3H]S-adenosyl methionine (SAM; Aswad, 1995), demonstrating the occurrence of isoaspartate formation
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O C
HN
NH
CO– O isoaspartate
[3H] SAM PIMT
O HN
C
NH gel electrophoretic quantification (support protocol 5 )
CO—C[3 H]3 O [3H] methylated intermediate
O C N HN
C
O succinmide intermediate + [3H] methanol
quantification (basic protocol 5)
Figure 14.4.7 Pathway for the PIMT-catalyzed methylation of isoaspartate used for quantitation.
Analysis of Oxidative Modification of Proteins
in vivo. Therefore, methods for the detection and quantification of isoaspartate formation in proteins/peptides are of interest to investigators studying post-translational modification of proteins. The discovery of PIMT also provides a useful tool for the quantification of isoaspartate formation in proteins (Fig. 14.4.7), as described in this unit. The only limitation of the method, however, is that PIMT, as a single enzyme, is not commer-
cially available. Isoaspartate formation can also be measured by isoelectric focusing gel analysis or proteolytic digestion of the protein followed by mass spectral analysis of the peptides resolved by reversed-phase HPLC (UNIT 16.2). The isoelectric focusing method is based on the fact that isoaspartate formation in a protein results in a net change of the protein’s charge. However, the method cannot detect the formation of isoaspartate residues derived from aspartic acid
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which do not alter the net charge of the protein (i.e., aspartyl isomerization). In addition, changes in the isoelectric focusing pattern of a protein may also result from the modification of other amino acid residues, such as methionine or lysine oxidation; therefore, the isoelectric focusing method is not specific. Proteolytic digestion of a protein followed by HPLC and mass spectral analysis is capable of detecting the presence of isoaspartate residues in the protein; however, the HPLC method, based on the retention time of the digested peptides, may not be able to resolve subtle changes in the peptide that arise from the deamidation of a particular peptide. Moreover, many changes may affect the retention time of a particular peptide, which could coelute with the isoaspartate-containing peptide and therefore does not necessarily indicate the presence of isoaspartate residues.
Critical Parameters and Troubleshooting In all studies concerning oxidative damage to proteins, auto-oxidation of the tissues by ambient oxygen, following tissue isolation, should be minimized. Accordingly, tissue collection and preparation must be carried out in buffers supplemented with antioxidants, such as diethylenetriaminepentaacetic acid (DTPA) and butylated hydroxytoluene (BHT). The use of antioxidant buffers may be especially critical for the detection of trace amounts of oxidized products formed during aging or under oxidative stress. In addition, all buffers should be bubbled with nitrogen before use. Analysis of protein carbonylation Depending upon the method used, protein carbonyl content can be expressed as either nmol/mg protein or cpm/mg protein for the DNPH (see Basic Protocol 1) and NaBH4 (see Basic Protocol 2) method respectively. Therefore, the only critical parameter for protein carbonyl quantification is the protein concentration. As a rule, protein concentration after washing with ethanol:ethyl acetate, should always be determined. In addition, it should be noted that a high concentration of protein-conjugated DNP interferes with the BCA protein assay; therefore, for heavily oxidized proteins that have extremely high carbonyl content determined by DNPH, protein concentration should be assayed by UV absorption at 280 nm using BSA as a standard (UNIT 3.1). Two potential problems may arise when protein carbonyl content is measured spectro-
photometrically using DNPH. First, absorption of protein-bound DNP in some experiments may be lower than that of a blank, which is the same protein sample treated with HCl (2 M) rather than DNPH. This problem may be caused by chromophores of certain proteins in the sample. If this is indeed the case, protein carbonyl content may have to be determined by other methods, such as the use of tritiated sodium borohydride (see Basic Protocol 2). The problem can also be caused by the difference in protein concentrations between a blank and a sample due to loss of proteins during washing. Therefore, care should be taken to minimize protein loss during washing steps, and protein concentration should always be determined after the removal of free DNPH (see Basic Protocol 2, step 4). Second, for tissue samples, it may be very difficult to dissolve the final protein pellet after washing. This problem is most likely caused by the use of high concentration of proteins in the initial step where protein samples are treated with DNPH. We have found that starting with 1 mg/ml protein will render the final pellet readily soluble. The solubility of a protein pellet will also depend on the nature of the protein; for example, lipoproteins and collagen are always quite difficult to dissolve after treatment with DNPH. In such a case, a denaturing buffer containing 150 mM phosphate and 3% SDS, pH 6.8 should be used. For immunoblot detection of protein carbonyls, a major concern is the dilution of antibodies. If overdiluted, the immunostaining signal may be very weak; if underdiluted, the background may be too dark. Therefore, it is critical to optimize each antibody’s dilution so that the immunostain signal is maximized while background staining is minimized. In addition, the amount of protein loaded onto SDS-PAGE should not be too high. Otherwise, nonspecific immune reactions may occur. Generally, 5 µg of a protein mixture is enough and less may be used for a purified protein. When protein carbonyl content is determined by tritiated sodium borohydride (see Basic Protocol 2), protein concentrations should always be determined after the removal of free tritium because protein recovery after washing is rarely 100%. An 80% to 90% recovery for mitochondrial proteins has been routinely obtained in the authors’ laboratory. When tritiated proteins are analyzed by SDS-PAGE, solubilization of more than one band of the same protein in one vial may be necessary to obtain enough radioactivity counts.
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Analysis of protein thiol oxidation When protein thiol groups are determined by radiolabeling with [14C] iodoacetamide in conjunction with gel electrophoresis, it is necessary to include a final concentration of 1% SDS in the reaction mixture so that complete labeling can be ensured. For some proteins, the nucleophilicity of the thiolate (S−) anion may be lower than potentially competing alcohol and amino groups in the same proteins; therefore, the iodoacetamide reaction may not be absolutely specific. In such cases, experimental conditions may need to be optimized by varying temperature, pH, and duration of the reactions so that the labeling for cysteine residues will be specific. It should be noted that iodoacetamide is intrinsically unstable in light, especially in solution; reactions should therefore be carried out in the dark. Adding cysteine, glutathione, or mercaptosuccinic acid to the reaction mixture will quench thiol-reactive species, forming highly water-soluble adducts that are easily separated from proteins during gel electrophoresis.
Analysis of Oxidative Modification of Proteins
Analysis of tyrosine oxidative modification For GC/MS measurement of protein dityrosine residues, dityrosine always needs to be released by acid hydrolysis. Although it has been demonstrated by other investigators that the procedure of acid hydrolysis does not induce more oxidation of tyrosine, which can lead to additional dityrosine formation, care should be taken to ensure no further oxidation indeed takes place for a given sample under given experimental conditions by adding antioxidants to buffers. For such purposes, pure tyrosine should be used as a control during the acid hydrolysis process. In addition, for tissue samples (protein mixtures), the dityrosine level is usually above the detection limit of GC/MS method; however, this may not be the case for an individual protein. Therefore, it may be advisable to determine whether the protein of interest has a dityrosine level that is above the limit of detection. To achieve this, analyze the protein at two widely different concentrations, 50- to 100-fold. If no dityrosine is detectable in the concentrated sample, it is most likely that the protein does not contain a significant amount of dityrosine residues. For the measurement of protein nitrotyrosine by competitive ELISA, antibody dilution and the amount of nitro-BSA used for plate coating may have to be optimized. To achieve this, first titrate the amount of nitro-BSA needed to coat the plate versus a constant, high concentration of anti-nitrotyrosine antibody. Plot the absorbance values
and select the lowest level that yields a strong signal. Next, coat the plate with this amount of nitro-BSA and vary the concentration of antibodies. Plot the obtained values versus antibody dilution and select a level of the antibody that is within the linear portion of the curve. Optimal dilution for the secondary antibody can also be obtained by similar titration. In addition, in most of the competitive ELISA assays, the first antibody (here anti-nitrotyrosine) may bind preferentially to the coating antigen because the latter on the plate is highly localized. Therefore, to ensure a thorough competition, samples to be analyzed should always be premixed with the first antibody and incubated overnight before the actual ELISA measurement. Analysis of isoaspartate formation When protein isoaspartate is measured by PIMT-catalyzed methyl transfer reaction, the protein solution must not contain any detergent such as SDS or chaotropic reagents such as guanidineHCl, as these will denature the PIMT enzyme. Moreover, for an unknown protein or peptide, as a first step, it is useful to determine whether the protein has any isoaspartate residues. The protein can be diluted to several different concentrations and then analyzed. If the concentration of isoaspartic acid is not protein concentration-dependent, it is most likely that there is little or no isoaspartic acid present in the protein or peptide. Some proteins, when not denatured, may contain isoaspartate residues in domains that make them poor substrates for PIMT. In such cases, the PIMT methylation reaction may not be complete and isoaspartate concentration may be underestimated. If so, proteins may need to be fragmented by proteolysis before they are used for isoaspartate determination. To determine if proteolysis is necessary, process a sample of undigested protein along with a digested sample. If the amount of isoaspartate measured in the intact protein is similar to that in the digested protein, the protein need not be fragmented for isoaspartate quantification. If a protein should require digestion by protease, the sample buffer used for blank reactions should contain the same protease and buffer components existing in the digestion, but without the protein sample. This will exclude any isoaspartate residues that may be present in the protease.
Anticipated Results When performed correctly, protein carbonyl content of native proteins, such as BSA and human plasma, should be around 1 nmol/mg
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protein (usually 0.5 to 1.5 nmol/mg protein). Literature values for carbonyl content in normal tissues are usually between 1.5-2.0 nmol/mg protein (Reanick and Packer, 1994). If immunoblot is performed, immunostaining of protein carbonyls should be differential, which means protein oxidative damage does not occur equally in all proteins. Protein carbonylation analyzed by the use of tritiated borohydride is highly dependent on the source of tritiated borohydride, but for the same batch of the chemical with the same concentration, results are generally reproducible. If protein carbonyls derivatized with tritiated sodium borohydride are to be quantitated by a gel electrophoretic technique (see Support Protocol 2), radioactivity counts for each individual band in a protein mixture (<20 µg loaded), are usually less than 300 cpm if no more than one band is solubilized in a single vial. For protein thiol groups measured by gel electrophoresis of iodoacetamide-treated proteins (see Basic Protocol 3), results of autoradiographic intensities should show a highly differential pattern for each individual protein. If a band exhibits the highest intensity among all the proteins resolved by SDS-PAGE, the protein would have the least loss in its thiol groups. In contrast, if a protein band exhibits a very weak intensity following autoradiography, it should have a maximal loss in its thiol groups. Depending on what proteins are being studied, results of liquid scintillation counting of an excised individual band may range from 500 cpm to 2000 cpm. For mass spectrometric quantification of dityrosine (see Basic Protocol 4), the concentration of protein dityrosine is usually expressed as the ratio of dityrosine to tyrosine. In mouse, dityrosine concentration has been found to be around 0.3 to 0.6 mmol/mol for heart, 0.1 to 0.3 mmol/mol for skeletal muscle and brain, and 0.2 mmol/mol for liver (ter Steege et al., 1998). Nitrotyrosine, when determined by competitive ELISA and expressed as nitro-BSA equivalents, is about 0.12 µM in human plasma. Isoaspartate concentration in proteins, usually expressed as pmol/mg protein, is 200 pmol/mg protein in murine brain and heart, and < 50 pmol/mg protein in liver with no detectable isoaspartate residues in the plasma (Kim et al., 1997).
Time Considerations Without the consideration of preparation of protein samples, for a small number of samples (<20), spectrophotometric quantitation (see Basic Protocol 1) can be finished within one day after
the DNPH treatment. If a large number of samples are involved, the washing step may take a longer time, but one can always leave the protein pellet in the ethanol:ethyl acetate solution with the tubes capped lightly and stored under a hood. Immunoblot detection of protein carbonyls (see Support Protocol 1), starting from SDSPAGE, can be finished within 48 hr. The time can be shortened to 12 hr if the PVDF membrane is incubated for 1 hr each, with both primary and secondary antibodies. The quantitation of protein carbonyls by electrophoresis (see Support Protocol 2) may require one week. Most of the time is spent on gel solubilization and liquid scintillation counting. Gel electrophoretic analysis of protein thiol groups labeled with [14C]iodoacetamide (see Basic Protocol 3) will take at least two weeks. Generally, radiolabeling of the protein samples and gel analysis can be finished within two days. The rest of the time is used for the exposure of X-ray film to the Whatman 3MM filter paper onto which gels have been dried. Quantification of protein carbonyls by tritiated borohydride labeling (see Support Protocol 2) typical requires two days for ~20 samples. Labeling is generally accomplished in the first day, while the second day is left for scintillation counting. Quantitation of protein dityrosine by mass spectrometry (see Basic Protocol 4) usually takes up to one week. Sample preparation, including tissue hydrolysis, can take three days; the actual quantitation by mass spectrometry can be finished within two days. Preparation of dityrosine standard (see Support Protocol 3) may take up to one week and the time used for lyophilization steps are flexible. In addition, the lyophilized samples can be stored at −80°C for several months before further analysis. Competition ELISA measurement of nitrotyrosine (see Support Protocol 4) usually takes two days. Day one is allotted for sample preparation, plate coating and preincubation of the samples or nitro-BSA with anti-nitrotyrosine antibodies. Day two is used for antibody incubation, plate washing and data collection. Enzymatic analysis of isoaspartate formation (see Basic Protocol 5), up to the step of liquid scintillation counting, can be finished in one day. After the addition of scintillation cocktail, the samples can be counted at any time. Gel electrophoretic analysis of isoaspartate formation (see Support Protocol 5) may require a week with most of the time spent on gel solubilization and liquid scintillation counting.
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ACKNOWLEDGEMENTS The authors thank Dr. Christiaan Leeuwenburgh for his help on the procedure for dityrosine quantification by GC/MS.
LITERATURE CITED Agarwal, S., and Sohal, R.S. 1994. Aging and proteolysis of oxidized proteins. Arch. Biochem. Biophys. 309:24-28.
Reanick, A.Z. and Packer, L. 1994. Oxidative damage to proteins: Spectrophotometric method for carbonyl assay. Methods Enzymol. 233:357-363. Stadtman, E.R. 1992. Protein oxidation and aging. Science. 257:1220-1224.
Aswad, D.W. (ed.). 1995. Deamidation and isoaspartate formation in peptides and proteins. CRC, Boca Raton, Fla.
Stadtman, E.R., and Berlett, B.S. 1997. Reactive oxygen-mediated protein oxidation in aging and disease. Chem. Res. Toxicol. 10:485-494.
Buss, H., Chan, T.P., Sluis, K.B., Domigan, N.M., and Winterbourn, C.C. 1997. Protein carbonyl measurement by a sensitive ELISA method [published errata appear in Free Radic Biol Med 1998 May; 24:1352]. Free Radic. Biol. Med. 23: 361-366.
ter Steege, J.C., Koster-Kamphuis, L., van Straaten, E.A., Forget, P.P., and Buurman, W.A. 1998. Nitrotyrosine in plasma of celiac disease patients as detected by a new sandwich ELISA. Free Radic. Biol. Med. 25:953-963.
Clark, S. 1985. Protein carboxyl methyltransferase: Two distinct classes of enzymes. Annu. Rev. Biochem. 54:479-506.
Yan, L.J., Levine, R.L., and Sohal, R.S. 1997. Oxidative damage during aging targets mitochondrial aconitase. Proc. Natl. Acad. Sci. USA. 94:1116811172.
Davies, K.J., Lin, S.W., and Pacifici, R.E. 1987. Protein damage and degradation by oxygen radicals. IV. Degradation of denatured protein. J. Biol. Chem. 262:9914-9920. Ellman, G.L. 1959. Tissue sulfhydryl groups. Arch. Biochem. Biophys. 82:70-77. Giulivi, C., and Davies, K.J. 1993. Dityrosine and tyrosine oxidation products are endogenous markers for the selective proteolysis of oxidatively modified red blood cell hemoglobin by (the 19 S) proteasome. J. Biol. Chem. 268:8752-8759. Graf, L., Bajusz, S., Patthy, A., Barat, E., and Cseh, G. 1971. Revised amide location for porcine and human adrenocorticotropic hormone. Acta Biochim. Biophys. Acad. Sci. Hung. 6:415-418. Hughes, B.A., Roth, G.S., and Pitha, J. 1980. Age-related decrease in repair of oxidative damage to surface sulfhydryl groups on rat adipocytes. J. Cell. Physiol. 103:349-33. Ischiropoulos, H., Zhu, L., Chen, J., Tsai, M., Martin, J.C., Smith, C.D., and Beckman, J.S. 1992. Peroxynitrite-mediated tyrosine nitration catalyzed by superoxide dismutase. Arch. Biochem. Biophys. 298:431-437. Kim, E., Lowenson, J.D., MacLaren, D.C., Clarke, S., and Young, S.G. 1997. Deficiency of a protein-repair enzyme results in the accumulation of altered proteins, retardation of growth, and fatal seizures in mice. Proc. Natl. Acad. Sci. U.S.A. 94:6132-6137. Leeuwenburgh, C., Wagner, P., Holloszy, J.O., Sohal, R.S., and Heinecke, J.W. 1997. Caloric restriction attenuates dityrosine cross-linking of cardiac and skeletal muscle proteins in aging mice. Arch. Biochem. Biophys. 346:74-80. Levine, R.L. 1983. Oxidative modification of glutamine synthetase. J. Biol. Chem. 258:1182311827. Analysis of Oxidative Modification of Proteins
McKenzie, S.J., Baker, M.S., Buffinton, G.D., and Doe, W.F. 1996. Evidence of oxidant-induced injury to epithelial cells during inflammatory bowel disease. J. Clin. Invest. 98:136-141.
Levine, R.L., Williams, J.A., Stadtman, E.R., and Shacter, E. 1994. Carbonyl assays for determination of oxidatively modified proteins. Methods Enzymol. 233:346-357.
Yan, L.J., and Sohal, R.S. 1998a. Gel electrophoretic quantitation of protein carbonyls derivatized with tritiated sodium borohydride. Anal. Biochem. 265:176-82. Yan, L.J., and Sohal, R.S. 1998b. Mitochondrial adenine nucleotide translocase is modified oxidatively during aging. Proc. Natl. Acad. Sci U.S.A. 95:12896-12901.
Key References Berlett, B.S., and Stadtman, E.R. 1997. Protein oxidation in aging, disease, and oxidative stress. J. Biol. Chem. 272:20313-20316. Detailed biochemical mechanisms of protein oxidation, general principles of intracellular accumulation of oxidized proteins and implication of oxidized proteins in aging and disease. Heinecke, J.W, Hsu, F.F, Crowley, J.R, Hazen, S.L., Leeuwenburgh, C., Mueller, D.M., Rasmussen, J.E. and Turk, J. 1999. Detecting oxidative modification of biomolecules with isotope dilution mass spectrometry: Sensitive and quantitative assays for oxidized amino acids in proteins and tissues. Methods Enzymol. 300:124-144. Extensive description of tyrosine modified products, including dityrosine, nitrotyrosine, and chlorotyrosine, by GC/MS method. Aswad, 1995. See above. Extensive coverage of methods for the quantification of isoaspartate formation in proteins/peptides. A useful collection of examples of isoaspartate formation in individual proteins.
Contributed by Liang-Jun Yan and Rajindar S. Sohal Southern Methodist University Dallas, Texas
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Analysis of Protein Ubiquitination
UNIT 14.5
The covalent attachment of the polypeptide ubiquitin (Ub) to target proteins is probably best known for its role in directing modified substrates to the 26S proteasome for degradation. However, Ub modification of some substrates can have nonproteolytic functions (Hicke, 1999). Two methods for assaying protein ubiquitination are described in this unit (see Basic Protocols 1 and 2). Since cells also express a number of Ub-like (Ubl) molecules that can be conjugated to substrate proteins, these analytical methods can be adapted for studying the modification of proteins by Ubls as well. These methods are suitable for a variety of eukaryotic cells, but techniques are specifically described for use with yeast and mammalian cells. Attachment of Ub to a protein requires a complex of enzymes that recognize the substrate and promote Ub transfer. Sequence motifs present in these enzymes (e.g., a HECT domain, RING motif, PHD domain, or U box; Hatakeyama et al., 2001) may indicate that other uncharacterized proteins containing these motifs have a biochemical function in Ub-protein ligation, and several in vitro methods are described in this unit for determining if a protein has Ub-transferring activity. These include assaying for auto-ubiquitination of E3 (see Basic Protocol 3), and assaying ubiquitination of a model substrate protein (see Basic Protocol 4). IMMUNOPRECIPITATION OF TARGET PROTEIN FOLLOWED BY ANTI-Ub IMMUNOBLOTTING
BASIC PROTOCOL 1
This assay evaluates whether a protein is ubiquitinated by enriching the protein of interest from cell lysates using immunoprecipitation, fractionating this precipitated material by SDS-PAGE, and then immunoblotting with Ub-specific antibodies. Because deubiquitinating activity is often high in cell extracts, precautions must be taken to guard against substrate deubiquitination. Therefore, it is essential to include an inhibitor of deubiquitinating enzymes (Dubs) in all lysis buffers. Since these enzymes usually have a cysteine at their active site, the sulfhydryl alkylating agent N-ethylmaleimide (NEM) is commonly used to inactivate them. A highly specific Dub inhibitor, ubiquitin aldehyde, is available commercially (Affiniti Research Products) but is rather expensive. This inhibitor may be helpful in circumstances when the use of NEM is not practical (e.g., when analyzing a protein with an NEM-sensitive epitope). Antibodies capable of precipitating the protein of interest are required since this method relies on immunoprecipitation of the substrate. If substrate-specific antibodies are not available, a common alternate approach is epitope-tagging, where the protein of interest is engineered to carry one of a variety of different antigenic determinants. These epitopetagged proteins can then be monitored with commercially available monoclonal antibodies whose immunoprecipitation capabilities are thoroughly established. Materials Yeast or mammalian cells expressing the protein of interest and matching control cells PBS (APPENDIX 2E) RIPA buffer (see recipe) containing freshly dissolved 10 mM N-ethylmaleimide (NEM) and protease inhibitors (see recipe), ice-cold H2O, sterile 100% ethanol containing freshly dissolved 50 mM NEM, ice-cold SDS buffer (see recipe)
Contributed by Jeffrey D. Laney and Mark Hochstrasser Current Protocols in Protein Science (2002) 14.5.1-14.5.11 Copyright © 2002 by John Wiley & Sons, Inc.
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Triton lysis buffer (see recipe) containing freshly dissolved 10 mM NEM and protease inhibitors, ice-cold Protein A-agarose (Repligen) or protein G-beads (Amersham Biosciences) Triton lysis buffer (see recipe) supplemented with 0.05% (w/v) SDS 2× SDS-PAGE sample buffer (see recipe) Blocking buffer—e.g., 5% (w/v) nonfat dry milk Anti-Ub antibodies (Covance Research Products, Zymed Laboratories, Santa Cruz Biotechnology) Enhanced chemiluminescence (ECL), enhanced chemifluorescence (ECF), or 125 I-labeled reagents 425- to 600-µm acid-washed glass beads (Sigma) End-over-end rotator Additional reagents and equipment for SDS-PAGE (UNIT 10.1), electroblotting (UNIT 10.7), and immunoblot detection (UNIT 10.10) For mammalian cells 1a. Wash two 100-mm plates, one containing cultured mammalian cells expressing the protein of interest and the other matching control cells, once with PBS. Protein expression is often achieved by transient transfection with an appropriate expression vector. Control experiments with cells lacking the protein of interest should be conducted in parallel. With epitope-tagged substrates, the untagged protein is commonly used as the control.
2a. Lyse the cells by incubating on ice 30 min with 1 ml ice-cold RIPA buffer containing freshly dissolved 10 mM N-ethylmaleimide (NEM) and protease inhibitors. 3a. Collect the lysates in 1.5-ml microcentrifuge tubes and clear by centrifuging 15 min at 14,000 × g, 4°C. Alternative lysis procedures may be utilized (e.g., PBS containing 0.1% NP-40, 10 mM NEM and protease inhibitors) depending on the protein being studied.
For yeast cells 1b. Grow two 10-ml cultures, one of yeast cells expressing the protein of interest and the other matching control cells, to mid-log phase (OD600 = 0.5 to 1.0). Collect cells by centrifuging 5 min at 1200 × g, room temperature. Wash once with 10 ml sterile water. Repeat centrifugation. Resuspend cells in 1 ml sterile water and transfer to a 1.5-ml microcentrifuge tube. Collect cells by centrifuging 10 sec at 14,000 × g, room temperature. Control experiments with cells lacking the protein of interest should be conducted in parallel. With epitope-tagged substrates, the untagged protein is commonly used as the control.
2b. Discard the supernatant and resuspend the cell pellet in 500 µl ice-cold 100% ethanol containing freshly dissolved 50 mM NEM to lyse the cells. Add ∼300 µl of 425- to 600-µm acid-washed glass beads. Vortex samples continuously for 5 min at room temperature. Transfer liquid to a new tube. Wash the glass beads twice with 500 µl fresh ethanol/NEM and pool the liquid (total ∼1.5 ml). The transferred liquid should be cloudy from precipitated protein. Do not centrifuge the sample, as the glass beads will settle rapidly. Analysis of Protein Ubiquitination
3b. Dry the samples completely in a Speedvac evaporator without heating. Resuspend the precipitated protein in 100 µl SDS buffer by vortexing and heating in a boiling
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water bath 5 min. Dilute the sample with 1 ml ice-cold Triton lysis buffer containing freshly dissolved 10 mM NEM and protease inhibitors. Remove any remaining precipitate by centrifuging 15 min at 14,000 × g, 4°C. Immunoprecipitate the protein of interest 4. Normalize cell extract volumes. Transfer normalized volumes of extract to fresh tubes. Add an appropriate amount of antibody and incubate 1 to 4 hr at 4°C using an end-over-end rotator. Extracts are typically normalized according to cell number (mammalian cells) or culture OD600 (yeast), but normalization based upon total protein concentration may be preferable as it is probably more accurate.
5. After incubation, add an appropriate amount of protein A-agarose to quantitatively precipitate the antibody-antigen complex, and rotate end-over-end 1 hr at 4°C. Substitute protein G beads when using antibodies that are not bound effectively by protein A. 6. Collect immunoprecipitates by microcentrifuging <5 sec at top speed and wash the beads three times with 1 ml Triton lysis buffer supplemented with 0.05% (w/v) SDS. For epitope-tagged proteins, anti-tag antibodies covalently coupled to beads are available commercially and are convenient for precipitations.
7. Completely remove wash buffer from the beads with a micropipettor, add 1 vol of 2× SDS-PAGE sample buffer, and incubate in a boiling water bath 3 min. Analyze samples by electrophoresis on an SDS-PAGE gel (UNIT 10.1) and transfer to a PVDF or nitrocellulose membrane (UNIT 10.7). Immunoblot transfer of high-molecular-mass Ub-conjugates may be more efficient from a low percentage polyacrylamide gel.
8. Block nonspecific binding sites with an appropriate blocking buffer—e.g., 5% (w/v) nonfat dry milk—then incubate the membrane with an appropriate dilution of anti-Ub monoclonal antibody. 9. Develop the immunoblot signal with enhanced chemiluminescence (ECL), enhanced chemifluorescence (ECF), or 125I-labeled reagents according to the manufacturer’s instructions (also see UNIT 10.10). Use of 125I-labeled or chemifluorescent detection reagents allows for the ubiquitinated species to be analyzed quantitatively. The sensitivity of anti-Ub immunoblots may be improved by boiling or autoclaving the membrane after protein transfer, which presumably denatures and exposes buried epitopes in the easily refolded Ub molecule.
AFFINITY PURIFICATION OF UBIQUITINATED PROTEINS FROM CELLS EXPRESSING HIS6-Ub
BASIC PROTOCOL 2
This method to analyze protein ubiquitination utilizes hexahistidine (His6)-tagged Ub as a purification reagent for Ub-protein conjugates. Cell lysates are made under denaturing conditions (6 M guanidine⋅HCl), which effectively preserves the Ub-modified species without altering the ability of His6-modified protein to interact with the affinity resin (Trier et al., 1994; Kaiser et al., 2000).
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Materials Yeast or mammalian cells expressing the protein of interest and His6-tagged Ub (UNITS 5.6-5.8) and matching control cells PBS (APPENDIX 2E) 6 M guanidine⋅HCl/100 mM sodium phosphate buffer, pH 8.0 (APPENDIX 2E) with and without 5 mM imidazole 2+ Ni -NTA-agarose beads (Qiagen, Clontech) Disposable chromatography columns (e.g., Bio-Rad poly-prep) 6 M guanidine⋅HCl/100 mM sodium phosphate buffer, pH 5.8 (APPENDIX 2E) Protein buffer (see recipe) with and without 10 and 200 mM imidazole 10% (v/v) TCA 2× SDS-PAGE sample buffer (see recipe) Guanidine wash buffer (see recipe) with and without 20 mM imidazole 1 M imidazole Urea wash buffer, pH 8.0 (see recipe) containing 20 mM imidazole Urea wash buffer, pH 6.0 (see recipe) containing 20 mM imidazole 2× urea sample buffer (see recipe) Probe-type sonicator with microtip End-over-end rotator 425- to 600-µm acid-washed glass beads (Sigma) Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and immunoblot detection (UNIT 10.10) For mammalian cells 1a. Wash two 100-mM plates, one containing mammalian cells expressing the protein of interest and His6-tagged Ub and one containing matching control cells, once in PBS. Lyse in 2 ml of 6 M guanidine⋅HCl/100 mM sodium phosphate buffer, pH 8.0 with 5 mM imidazole. Collect the extract in a 1.5-ml microcentrifuge tube. Reduce the viscosity of the lysate by brief treatment with a probe-type sonicator set at the minimum setting required to achieve vigorous mixing without foaming. Clear by centrifuging 15 min at 14,000 × g, 4°C. Protein expression is often achieved by transient transfection of appropriate expression vectors (UNITS 5.9 & 5.10). Control experiments with cells lacking the protein of interest should be conducted in parallel. With epitope-tagged substrates, the untagged protein is commonly used as the control. Cells lacking His6-tagged Ub should also be examined.
2a. Combine the clarified extract with 75 µl (settled volume) Ni2+-NTA-agarose beads per 2 ml extract and incubate 4 hr at 4°C on an end-over-end rotator. 3a. Pour the slurry into a disposable chromatography column (e.g., Bio-Rad poly-prep) and wash successively with the following:
Analysis of Protein Ubiquitination
1 ml 6 M guanidine⋅HCl/100 mM sodium phosphate buffer, pH 8.0 without imidazole 2 ml 6 M guanidine⋅HCl/100 mM sodium phosphate buffer, pH 5.8 1 ml 6 M guanidine⋅HCl/100 mM sodium phosphate buffer, pH 8.0 without imidazole 2 ml 1:1 mix of 6 M guanidine⋅HCl/100 mM sodium phosphate buffer, pH 8.0 and protein buffer without imidazole 2 ml 1:3 mix of 6 M guanidine⋅HCl/100 mM sodium phosphate buffer, pH 8.0 and protein buffer without imidazole 2 ml protein buffer without imidazole 1 ml protein buffer with 10 mM imidazole.
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4a. Elute bound protein in 1 ml protein buffer containing 200 mM imidazole. Precipitate the eluate with 10% (v/v) TCA and resuspend by adding 1 vol of 2× SDS-PAGE sample buffer and incubating in a boiling water bath for 5 min. Analyze by SDSPAGE (UNIT 10.1) and immunoblot with antibodies directed against the target protein (UNIT 10.10). For yeast cells 1b. Wash two cultures of yeast cells, one expressing His6-tagged Ub and the other matching control cells, once with water. Add ∼300 µl of 425- to 600-µm acid-washed glass beads (Sigma) and lyse in guanidine wash buffer without imidazole by vortexing 5 min. Control experiments with cells lacking the protein of interest should be conducted in parallel. With epitope-tagged substrates, the untagged protein is commonly used as a control. Cells lacking His6-tagged Ub should also be examined.
2b. Centrifuge the lysate 15 min at 14,000 × g, 4°C. Combine the clarified extract with 50 µl Ni2+-NTA-agarose beads per 2.75 mg total protein in a fresh tube and add 1 M imidazole to a final concentration of 10 mM. Incubate 4 hr at 4°C using an end-over-end rotator. 3b. Wash the beads successively with the following: 1 ml guanidine wash buffer containing 20 mM imidazole 1 ml urea wash buffer, pH 8.0, containing 20 mM imidazole 1 ml urea wash buffer, pH 6.0, containing 20 mM imidazole 1 ml urea wash buffer, pH 6.0, containing 20 mM imidazole. Wash the beads well with the urea-containing buffers as it is critical to remove all traces of guanidinium that will precipitate in the presence of SDS.
4b. Elute the bound material from the beads by boiling in urea sample buffer. Analyze samples by SDS-PAGE (UNIT 10.1) and immunoblotting with antibodies directed against the target protein (UNIT 10.10). ASSAY FOR AUTO-UBIQUITINATION BY E3 Ub-PROTEIN LIGASES Many E3 Ub-protein ligases can catalyze the transfer of ubiquitin to themselves. This protocol allows determination of whether a protein of interest can promote E2-dependent (but substrate-independent) ubiquitin transfer.
BASIC PROTOCOL 3
Materials 5× ubiquitination buffer (see recipe) Purified GST-fusion protein: glutathione-S-transferase (GST) fused with protein of interest (UNIT 6.6) E1 ubiquitin-activating enzyme, E2 ubiquitin-conjugating enzyme, and ubiquitin (purified from recombinant sources or commercially available from Calbiochem, Boston Biochem, Sigma, A.G. Scientific) 2× SDS-PAGE sample buffer (see recipe) Anti-Ub antibodies (Covance Research Products, Zymed Laboratories, Santa Cruz Biotechnology) or anti-GST antibodies (Santa Cruz Biotechnology) Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and immunoblot detection (UNIT 10.10) Post-Translational Modification: Specialized Applications
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1. For each reaction prepare a mixture containing the following: 4 µl 5× ubiquitination buffer 1 to 10 µg purified GST-fusion protein 100 ng E1 ubiquitin-activating enzyme 500 ng E2 ubiquitin-conjugating enzyme 2.5 µg ubiquitin H2O to 20 µl. Also prepare control reactions singly omitting each of the enzyme components for assay in parallel. The GST-fusion protein is often a fusion of a specific protein domain to GST.
2. Incubate 1.5 hr or longer at room temperature. The duration of incubation depends on the protein being assayed.
3. Stop the reactions by adding 20 µl of 2× SDS-PAGE sample buffer and heat 5 min in a boiling water bath. Analyze by SDS-PAGE (UNIT 10.1) and immunoblot using anti-Ub or -GST antibodies (UNIT 10.10). BASIC PROTOCOL 4
ASSAY FOR TRANSFER OF UBIQUITIN TO A MODEL SUBSTRATE PROTEIN The function of an E3 Ub-protein ligase is to catalyze the transfer of ubiquitin from an E2 conjugating enzyme to a substrate protein. This transfer reaction usually requires that the presumptive E3 contact both the E2 Ub-conjugating enzyme and the substrate. The following method has been developed for situations in which the substrate is unknown or difficult to purify, or the substrate interaction domain in the putative E3 is uncharacterized. This assay determines if a protein of interest can promote the transfer of ubiquitin to the model substrate S protein by exploiting the high affinity binding of the ribonuclease A fragments called S protein and S peptide, thereby bringing an E2/E3 complex to the substrate (Figure 14.5.1). Materials 5× ubiquitination buffer (see recipe) Purified GST-S peptide fusion protein: glutathione-S-transferase (GST), ribonuclease S peptide and the protein of interest (UNIT 6.6) Ribonuclease S protein (Sigma) E1 ubiquitin-activating enzyme, E2 ubiquitin-conjugating enzyme, and ubiquitin (purified from recombinant sources or commercially available from Calbiochem, Boston Biochem, Sigma, A.G. Scientific)
Lys S
Ub
F
Ub
F
E2
E2 Lys S
Analysis of Protein Ubiquitination
Figure 14.5.1 Diagram of the ribonuclease S protein/S peptide-mediated Ub transfer reaction. Abbreviations: S, ribonuclease S protein; F, S peptide-protein of interest fusion protein; E2, Ub-conjugating enzyme.
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2× SDS-PAGE sample buffer (see recipe) Anti-Ub antibodies (Covance Research Products, Zymed Laboratories, Santa Cruz Biotechnology) Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and immunoblot detection (UNIT 10.10) 1. For each reaction, prepare a mixture containing the following: 4 µl 5× ubiquitination buffer 1.5 µg purified GST-S peptide fusion protein 1.5 µg ribonuclease S protein 100 ng E1 ubiquitin-activating enzyme 500 ng E2 ubiquitin-conjugating enzyme 2.5 µg ubiquitin H2O to 20 µl. Also prepare control reactions singly omitting each of the enzyme components to assay in parallel. The GST-S peptide fusion protein is often a fusion of a specific protein domain to GST-S peptide (using the pET41 or pET42 vectors from Novagen). A control reaction containing 1.5 ìg of the ribonuclease S peptide as a competitor should also be included.
2. Incubate 1.5 hr or longer at room temperature. 3. Stop the reactions by adding 20 µl of 2× SDS-PAGE sample buffer and heat for 5 min in a boiling water bath. Analyze by SDS-PAGE (UNIT 10.1) and anti-Ub immunoblotting (UNIT 10.10). REAGENTS AND SOLUTIONS Guanidine wash buffer 6 M guanidine⋅HCl 50 mM sodium phosphate buffer, pH 8.0 (APPENDIX 2E) 10 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 300 mM NaCl 5 mM NEM, freshly dissolved 2 µg/ml aprotinin, leupeptin, and pepstatin, freshly dissolved from 5 mg/ml stocks Store up to 6 months at 4°C without NEM or protease inhibitors Protease inhibitors For solutions requiring protease inhibitors, add protease inhibitor cocktail tablets (Roche Molecular Biochemicals) per manufacturer’s instructions and 5 mg/ml pepstatin in DMSO to 10 µg/ml on the day of use. Protein buffer 50 mM sodium phosphate buffer, pH 8.0 (APPENDIX 2E) 100 mM KCl 20% (v/v) glycerol 0.2% (v/v) NP-40 Store up to 6 months at 4°C
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RIPA buffer 0.15 M NaCl 1% (v/v) NP-40 0.5% (w/v) sodium deoxycholate 0.1% (w/v) SDS 0.05 M Tris⋅Cl, pH 8.0 Store up to 1 year at 4°C SDS buffer 1% (w/v) SDS 45 mM HEPES (sodium salt), pH 7.5 Store up to 1 year at room temperature SDS-PAGE sample buffer, 2× 20% (v/v) glycerol 4% (w/v) SDS 0.125 M Tris⋅Cl, pH 6.8 (APPENDIX 2E) 0.2 M DTT 0.01% bromphenol blue Store up to 6 months at −20°C Triton lysis buffer 1% Triton X-100 150 mM NaCl 50 mM HEPES (sodium salt), pH 7.5 5 mM disodium EDTA Store up to 1 year at room temperature Ubiquitination buffer, 5× 250 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 12.5 mM MgCl2 2.5 mM DTT 10 mM ATP Store in small aliquots up to 6 months at −20°C Urea sample buffer, 2× 8 M urea 4% (w/v) SDS 0.125 M Tris⋅Cl, pH 6.8 (APPENDIX 2E) 0.2 M DTT 0.01% (w/v) bromphenol blue Store up to 6 months at −20°C Urea wash buffer, pH 8.0 8 M urea (from solid) 50 mM sodium phosphate buffer, pH 8.0 (APPENDIX 2E) 10 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 300 mM NaCl 5 mM NEM, freshly dissolved 2 µg/ml aprotinin, leupeptin, and pepstatin, freshly dissolved from 5 mg/ml stocks Prepare fresh on day of use Urea wash buffer, pH 6.0 8 M urea (from solid) 50 mM sodium phosphate buffer, pH 6.0 (APPENDIX 2E) 10 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 300 mM NaCl 5 mM NEM, freshly dissolved 2 µg/ml aprotinin, leupeptin, and pepstatin, freshly dissolved from 5 mg/ml stocks Prepare fresh on day of use Analysis of Protein Ubiquitination
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COMMENTARY Background Information Evidence from a large number of different experimental systems has demonstrated a central role for the Ub system in cellular function. In addition to its housekeeping responsibilities in ridding the cell of misfolded, damaged, or misassembled proteins, the Ub pathway has important regulatory duties. Substrates that are modified by Ub attachment exist in nearly every cellular compartment and include cell surface receptors, signal transduction kinases, cell cycle regulators, transcription factors, and mediators of apoptosis (Hicke, 1999; Brodsky and McCracken, 1999; Koepp et al., 1999). The small highly conserved Ub polypeptide is conjugated to target proteins by a reversible isopeptide linkage between lysine side chains in the substrate and the carboxyl terminus of Ub. A well-characterized cascade of enzymatic reactions mediates the covalent attachment of Ub to substrates. First, the C-terminal carboxyl group of Ub is activated in an energy-dependent manner by the Ub-activating enzyme (E1), resulting in a high-energy thioester bond between Ub and E1. Ub is then passed to an E2 Ub-conjugating enzyme by a thioester transfer reaction. E2, together with a specificity factor (a Ub-protein ligase or E3), directly transfers Ub to the ε-amino group of lysine residues in the substrate. These E3 proteins are thought to be critical for the high degree of specificity in protein ubiquitination. The ubiquitination complex formed by E2 and E3 generally catalyzes the formation of a polyubiquitin chain on the substrate (Hershko and Ciechanover, 1998). Different topologies of Ub chains exist in cells. For efficient targeting to the 26S proteasome for degradation, a K48-linked chain is required (Chau et al., 1989). The individual Ub subunits are linked to one another in these chains through the carboxyl-terminal glycine of one Ub joined to the side chain of lysine 48 (K48) on another Ub molecule. Ub polymers with alternative amide linkages (through K29 or K63) are also found in cells, where they appear to function in a number of other cellular processes (Arnason and Ellison, 1994; Spence et al., 1995; Galan and Haguenauer-Tsapis, 1997). To address whether a particular protein is modified by a Ub chain and to determine the topology of the Ub subunits in these polymers, the in vivo and in vitro assays described in this unit can be utilized with a few minor adaptations. Mutant forms of the 76-residue Ub protein can be overexpressed in cells and the
ubiquitination pattern observed can be compared to that in cells overexpressing wild-type Ub. These mutations substitute an arginine for different lysine residues in the Ub sequence; these mutants inhibit the ability of Ub to form poly-Ub chains through particular lysine residues without affecting the ability of the mutant Ub to be attached to substrates. If overexpression of the chain-terminating Ub mutant inhibits the formation of a poly-Ub chain on the protein of interest, it will be visible as a shift from highly ubiquitinated forms to conjugates with fewer Ub molecules attached. The effect is sometimes more pronounced in cells overexpressing a doubly mutated form of Ub in which the chain-terminating K-to-R substitution is combined with a G76A mutation. This latter substitution inhibits deubiquitination, so that once a Ub chain is capped with the Ub double mutant, it is modified stably because the capped chain cannot be disassembled efficiently (Hodgins et al., 1992; Finley et al., 1994). For the in vitro assays, the reactions can be programmed with mutant forms of Ub instead of wild-type Ub to infer the topology of the assembled chains. Cell-free systems have been established for a number of substrate ubiquitination events and have aided in defining the requirements for these specific ubiquitin transfer reactions. For example, these in vitro reactions have been critical in determining whether a particular protein exhibits E3 ligase activity. In vitro ubiquitination of substrates has been observed in a wide array of cell extracts, including lysates made from yeast, mammalian cells, frog eggs, and plant cells (e.g., Hershko et al., 1983; Scheffner et al., 1993; King et al., 1995; Verma et al., 1997; Spencer et al., 1999). Reconstitution of substrate ubiquitination using purified components has also been accomplished, providing mechanistic details of the modification reaction (Feldman et al., 1997; Skowyra et al., 1999). Perhaps not surprisingly, these studies require considerable knowledge of the specific Ub transfer event, as these reactions are often tightly regulated by cell-cycle position, substrate modification, or interaction with other proteins.
Critical Parameters and Troubleshooting As mentioned in the protocols, to determine if a protein is ubiquitinated in vivo, negative controls with cells lacking the purification tag (either the antigen used to immunoprecipitate
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Analysis of Protein Ubiquitination
the target or the His6-tagged Ub used to enrich Ub-protein conjugates) should be assayed in parallel. With the immunoprecipitation/immunoblotting experiments, constructs that express epitope-tagged Ub in cells can be used and the signal can be developed with an anti-tag antibody. Use of these constructs generates another useful negative control: cells expressing untagged Ub should be free of any signal. Another control experiment can be performed with mutant cells in which Ub activation is impaired or Ub levels are depleted (e.g., the mammalian cell line ts85; Ciechanover et al., 1984) or yeast strains carrying the uba1-2 (Swanson and Hochstrasser, 2000) or doa4∆ mutations (Swaminathan et al., 1999). A decrease in any positive signal should be produced with these cells. With the in vitro experiments that assay Ub-transfer activity, control experiments utilizing a mutant form of the protein of interest should be performed to substantiate any positive results. Since these proteins often contain a sequence motif that suggests a role in protein ubiquitination, mutations in conserved residues of these domains can be constructed and assayed. For example, RING domain–containing proteins contain a cluster of structurally important cysteine and histidine residues that can be mutated and analyzed (Joazeiro et al., 1999).
ubiquitinated forms of the S protein should be distinguishable.
Anticipated Results
Ciechanover, A., Finley, D., and Varshavsky, A. 1984. Ubiquitin dependence of selective protein degradation demonstrated in the mammalian cell cycle mutant ts85. Cell 37:57-66.
Gels of in vivo ubiquitination assays often yield smears or ladders of high-molecular-mass species that react with anti-Ub antibodies. The intensity distribution of the signals will differ, depending on the method utilized. With the immunoprecipitation and immunoblotting approach, the higher molecular weight conjugates should give rise to stronger signals, since these presumably contain many Ub moieties that can be recognized by anti-Ub antibodies. On the other hand, the purification of ubiquitinated proteins with His6-Ub will produce signals that are visually quite different than those produced with the other assay. Relatively weaker signals of the most highly modified forms are expected, since these conjugates contain only one target protein molecule regardless of the number of Ub monomers attached. Similar smears of immunoreactivity are produced with the in vitro auto-ubiquitination assays. The Ub transfer promoted by the S peptide fusion protein to the S protein model substrate may be obscured by the auto-ubiquitination occurring on the fusion protein. Nevertheless, distinct bands representing mono- and di-
Time Considerations After the cells are grown, the in vivo experiments require 1 day to make lysates and perform the immunoprecipitation or affinity purification. Another day is required for the SDSPAGE and protein transfer. Probing of the blot with antibodies can be performed overnight, with the development of the immunoblot signal performed on the following day. Ubiquitination in vitro requires 1 day to perform the reaction, analyze by SDS-PAGE, and transfer the protein to a membrane. The immunoblot can be incubated with antibodies overnight.
Literature Cited Arnason, T. and Ellison, M.J. 1994. Stress resistance in Saccharomyces cerevisiae is strongly correlated with assembly of a novel type of multiubiquitin chain. Mol. Cell Biol. 14:7876-7883. Brodsky, J.L. and McCracken, A.A. 1999. ER protein quality control and proteasome-mediated protein degradation. Semin. Cell Dev. Biol. 10:507-513. Chau, V., Tobias, J.W., Bachmair, A., Marriott, D., Ecker, D.J., Gonda, D.K., and Varshavsky, A. 1989. A multiubiquitin chain is confined to specific lysine in a targeted short-lived protein. Science 243:1576-1583.
Feldman, R.M., Correll, C.C., Kaplan, K.B., and Deshaies, R.J. 1997. A complex of Cdc4p, Skp1p, and Cdc53p/cullin catalyzes ubiquitination of the phosphorylated CDK inhibitor Sic1p. Cell 91:221-230. Finley, D., Sadis, S., Monia, B., Boucher, P., Ecker, D., Crooke, S., and Chau, V. 1994. Inhibition of proteolysis and cell cycle progression in a multiubiquitination-deficient yeast mutant. Mol. Cell. Biol. 14:5501-5509. Galan, J. and Haguenauer-Tsapis, R. 1997. Ubiquitin lys63 is involved in ubiquitination of a yeast plasma membrane protein. EMBO J. 16:5847-5854. Hatakeyama, S., Yada, M., Matsumoto, M., Ishida, N., and Nakayama, K.I. 2001. U box proteins as a new family of ubiquitin-protein ligases. J. Biol. Chem. 276:33111-20. Hershko, A. and Ciechanover, A. 1998. The ubiquitin system. Annu. Rev. Biochem. 67:425479. Hershko, A., Heller, H., Elias, S., and Ciechanover, A. 1983. Components of ubiquitin-protein ligase system: Resolution, affinity purification, and
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role in protein breakdown. J. Biol. Chem. 258:8206-8214. Hicke, L. 1999. Gettin’ down with ubiquitin: Turning off cell-surface receptors, transporters and channels. Trends Cell Biol. 9:107-112. Hodgins, R.R.W., Ellison, K.S., and Ellison, M.J. 1992. Expression of a ubiquitin derivative that conjugates to protein irreversibly produces phenotypes consistent with a ubiquitin deficiency. J. Biol. Chem. 267:8807-8812. Joazeiro, C.A., Wing, S.S., Huang, H., Leverson, J.D., Hunter, T., and Liu, Y.C. 1999. The tyrosine kinase negative regulator c-Cbl as a RING-type, E2-dependent ubiquitin-protein ligase. Science 286:309-312.
Spencer, E., Jiang, J., and Chen, Z.J. 1999. Signalinduced ubiquitination of IκBα by the F-box protein Slimb/β-TrCP. Genes Dev. 13:284-294. Swaminathan, S., Amerik, A.Y., and Hochstrasser, M. 1999. The Doa4 deubiquitinating enzyme is required for ubiquitin homeostasis in yeast. Mol. Biol. Cell 10:2583-2594. Swanson, R. and Hochstrasser, M. 2000. A viable ubiquitin-activating enzyme mutant for evaluating ubiquitin system function in Saccharomyces cerevisiae. FEBS Lett. 477:193-198. Treier, M., Staszewski, L.M., and Bohmann, D. 1994. Ubiquitin-dependent c-Jun degradation in vivo is mediated by delta domain. Cell 78:787798.
Kaiser, P., Flick, K., Wittenberg, C., and Reed, S.I. 2000. Regulation of transcription by ubiquitination without proteolysis: Cdc34/SCF(Met30)mediated inactivation of the transcription factor Met4. J. Biol. Chem. 276:303-314.
Verma, R., Chi, Y., and Deshaies, R.J. 1997. Cellfree ubiquitination of cell cycle regulators in budding yeast extracts. Methods Enzymol. 283:366-376.
King, R.W., Peters, J.M., Tugendreich, S., Rolfe, M., Hieter, P., and Kirschner, M.W. 1995. A 20S complex containing CDC27 and CDC16 catalyzes the mitosis-specific conjugation of ubiquitin to cyclin B. Cell 81:279-288.
KEY REFERENCES
Koepp, D.M., Harper, J.W., and Elledge, S.J. 1999. How the cyclin became a cyclin: Regulated proteolysis in the cell cycle. Cell 97:431-434. Scheffner, M., Huibregste, J.M., Vierstra, R.D., and Howley, P.M. 1993. The HPV E6 and E6-AP complex functions as a ubiquitin-protein ligase in the ubiquitination of p53. Cell 75:495-505. Skowyra, D., Koepp, D.M., Kamura, T., Conrad, M.N., Conaway, R.C., Conaway, J.W., Elledge, S.J., and Harper, J.W. 1999. Reconstitution of G1 cyclin ubiquitination with complexes containing SCFGrr1 and Rbx1. Science 284:662-665. Spence, J., Sadis, S., Haas, A., and Finley, D. 1995. A ubiquitin mutant with specific defects in DNA repair and multiubiquitination. Mol. Cell. Biol. 15, 1265-1273.
Treier, 1994. See above. Describes the affinity purification of ubiquitinated proteins from tissue culture cells using His6-Ub and the mammalian expression vectors for the tagged forms of Ub. Laney, J.D. and Hochstrasser, M. 2002. Assaying protein ubiquitination in Saccharomyces cerevisiae. Methods Enzymol. 351:248-257. Describes the yeast methods and the yeast expression vectors for mutant and tagged forms of Ub.
Contributed by Jeffrey D. Laney and Mark Hochstrasser Yale University New Haven, Connecticut
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Analysis of Protein S-Nitrosylation
UNIT 14.6
S-nitrosylation is the binding of a NO group to a cysteine or other thiol. Peptides and proteins that have an NO group attached to a cysteine are called S-nitrosothiols (SNOs). In recent years it has become evident that cell signaling is regulated by S-nitrosylation of critical cysteine residues on proteins. However, analysis of protein S-nitrosylation is technically challenging because S-NO bonds are easily lost or gained during sample preparation. In addition, the levels of endogenously S-nitrosylated intracellular proteins approach the limits of detection of currently available technologies. Despite these difficulties, several useful methods have been developed to analyze protein S-nitrosylation, including the Saville assay (see Basic Protocol 1). However, all the currently available methods have potential disadvantages and none is considered the gold standard. Therefore, S-nitrosylation measurements should be confirmed using at least two different methods. Chemical reduction/chemiluminescence (see Basic Protocol 2) and DAN assays (see Alternate Protocol 1) are other useful methods for measuring S-nitrosylation of purified nitrosylated intracellular proteins. The biotin-switch assay (see Basic Protocol 3) and anti-SNO immunofluorescence assays (see Alternate Protocol 2) are useful techniques for detecting total protein S-nitrosylation in cells or cell lysates. S-NITROSYLATION ANALYSIS OF PURIFIED RECOMBINANT PROTEINS USING A SAVILLE ASSAY
BASIC PROTOCOL 1
The easiest method for studying protein S-nitrosylation is to S-nitrosylate a purified recombinant protein in vitro with an SNO donor compound. SNO donor compounds are S-nitrosylated peptides. The SNO donor transfers its NO group to a cysteine residue on a protein via a transnitrosation reaction. S-nitrosylation of the protein is then measured using a Saville assay. In the Saville assay (Saville, 1958), NO+ is displaced from S-NO bonds by mercuric ion (HgCl2). The NO+ then reacts under acidic conditions with sulfanilamide to form a diazonium salt. The diazonium salt forms an intensely colored azo dye when coupled with N-(1-naphthyl)-ethylenediamine. Levels of the azo dye are measured at 540 nm (Stamler and Feelisch, 1996). Materials Purified recombinant protein of interest (Chapter 6) HEN buffer (see recipe) 1 mM S-nitrosoglutathione (GSNO) in H2O (store up to 3 months in small aliquots at −70°C) Solution A: 1% (w/v) sulfanilamide in 0.5 M HCl (store up to 1 month at room temperature protected from light) Solution B: Solution A containing 0.2% (w/v) HgCl2 (store up to 1 month at room temperature protected from light) Solution C: 0.02% (w/v) N-(1-naphthyl)-ethylenediamine dihydrochloride in 0.5 M HCl (store up to 1 month wrapped in foil at –20°C; if solution turns purple, discard) Sephadex G-25 column (PD-10; Amersham Biosciences) Spectrophotometer
Post-Translational Modification: Specialized Applications Contributed by Joan B. Mannick and Christopher M. Schonhoff Current Protocols in Protein Science (2004) 14.6.1-14.6.20 Copyright ©2004 by John Wiley & Sons, Inc.
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Carry out in vitro S-nitrosylation of proteins 1. Resuspend purified recombinant protein in HEN buffer to a final concentration of 10 µM. 2. Add 1 mM GNSO to a final concentration of 100 µM to each sample. Incubate samples for 30 min in the dark at room temperature. As a control, incubate 100 µM GSNO alone in HEN buffer. 3. Remove GSNO by passing the samples through a Sephadex G-25 PD-10 column, pre-equilibrated with HEN buffer. The NO donor that produces the most efficient S-nitrosylation may vary between proteins. Therefore, if GSNO does not efficiently S-nitrosylate, then other NO donors should be tried such as S-nitroso-N-acetylpenicillamine (SNAP) or S-nitrosocysteine (SNOcys). The time required for optimal nitrosylation may vary depending on the protein and NO donor used. The NO donor should be protected from light and frozen at −80°C when not in use.
Analyze protein S-nitrosylation using a Saville assay 4. Add 50 µl of each sample to 50 µl of solution A in wells of a 96-well plate. In separate wells, add 50 µl of each sample to 50 µl of solution B. CAUTION: HgCl2 (in solution B) is toxic and appropriate precautions should be taken when handling and disposing of this compound. APPENDIX 2A includes a protocol for disposal of mercury compounds.
5. Incubate samples 5 min at room temperature, then add 100 µl of solution C to each sample. 6. After 5 min, measure the absorbance of the samples spectrophotometrically at 540 nm. 7. Determine the amount of SNO in the sample by subtracting the absorbance of samples in solution A (nitrite) from the absorbance of samples in solution B (nitrite + SNO). Generate standard curves by repeating steps 4 to 6 using 2-fold serial dilutions of GSNO ranging from 250 to 15 µM. Use the control samples that contain GSNO in the absence of protein to determine the background absorbance generated by residual GSNO in the samples. GSNO is used to generate standard curves because it contains one SNO group per molecule. The standard curve generated by plotting GSNO concentration versus absorbance in solution B minus absorbance in solution A is then used to calculate the SNO concentration in protein samples (see Table 14.6.1 and Fig. 14.6.1). Although the Saville assay is useful for measuring nitrosylation of in vitro nitrosylated proteins, in general it is not sensitive enough to detect S-nitrosylation of intracellular proteins, although some groups have had success in doing so (Haendeler et al., 2002). To confirm the physiologic relevance of in vitro S-nitrosylation assays, it is important to demonstrate that a protein of interest is S-nitrosylated intracellularly by endogenous sources of NO as described in later protocols.
Analysis of Protein S-Nitrosylation
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S-NITROSYLATION ANALYSIS OF PURIFIED INTRACELLULAR PROTEINS BY CHEMICAL REDUCTION/CHEMILUMINESCENCE It is much more difficult to analyze endogenous S-nitrosylation of intracellular proteins than in vitro S-nitrosylated recombinant proteins because the concentration of intracellular proteins endogenously nitrosylated by intracellular sources of NO is typically near the limit of detection of currently available assays. The concentration of intracellular S-nitrosylated proteins can be increased by exposing cells or cell lysates to NO donor compounds. However, the physiologic relevance of SNO measurements of samples treated with NO donors is less clear than the physiologic relevance of SNO levels in samples that are S-nitrosylated by endogenous intracellular sources of NO. At present, there is no gold standard for measuring intracellular protein S-nitrosylation, and each of the currently available methods has potential drawbacks. Therefore, S-nitrosylation measurements need to be confirmed using at least two different methods. The most
BASIC PROTOCOL 2
0.5
OD Difference
0.5 0.4 0.3 0.2 0.1 0 0
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100
150
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[SNO]
Figure 14.6.1 Plot of data from Saville assay (see Table 14.6.1).
Table 14.6.1
Representative Data Using the Saville Assaya,b
Sample 250 µM GSNO 125 µM GSNO 62.5 µM GSNO 31.25 µM GSNO (SNO) – (BSA) BSA
Solution A
Solution B
(Solution B) – (Solution A)
0.072 0.051 0.044 0.039 0.195 0.216
0.563 0.337 0.186 0.129 0.382 0.202
0.491 0.286 0.142 0.09 0.187 0
aKnown concentrations of S-nitrosoglutathione standards (GSNO) or 100 mM samples of control or in
vitro S-nitrosylated bovine serum albumin (BSA and SNO-BSA) were treated with solution A or solution B for 5 min at room temperature (see Basic Protocol 1 for recipes). Solution C was then added to each sample. The absorbance of the samples measured spectrophotometrically at 540 nm is shown. The difference in absorbance between the samples treated with solution B and the samples treated with solution A [(solution B) – (solution A)] is proportional to the SNO content of each sample. The concentration of SNO in the SNO-BSA sample (∼80 µM) was determined by generating a standard curve of [SNO] versus absorbance using the GSNO standards. bGraphic depiction of these data is shown in Figure 14.6.1.
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sensitive and specific method for detecting intracellular protein nitrosylation is to immunoprecipitate a protein of interest from cell lysates, then measure the NO content of the immunoprecipitate by chemical reduction/chemiluminescence. There are several different methods of chemical reduction that can be used in this assay (Fang et al., 1998; Ruiz et al., 1998; Samouilov and Zweier, 1998; Mannick et al., 1999). In this protocol, the chemical reduction/chemiluminescence method requires a nitric oxide analyzer, which is not available in many laboratories. Therefore, an alternative method is provided for detecting nitrosylation of immunoprecipitated proteins using the compound 2,3-diaminonaphthalene (DAN). DAN reacts with NO released from S-nitrosylated proteins to form fluorescent products (see Alternate Protocol 1). Immunoprecipitation experiments require the availability of antibodies capable of precipitating proteins of interest. If such antibodies are not available, then an alternative approach is to express an epitope-tagged protein in cells and immunoprecipitate using commercially available antibodies directed against the epitope. Reducing agents must be eliminated from buffers when immunoprecipitating S-nitrosylated proteins because reducing agents break S-NO bonds. In addition, metal chelators such as EDTA must be added to buffers to eliminate free metals that catalyze the loss and/or gain of S-NO bonds. Immunoprecipitation should be performed in the dark at 4°C, and pH shifts must be avoided, since UV light and pH shifts will disrupt S-NO bonds. In the authors’ experience, chemical reduction/chemiluminescence method is the most sensitive and specific method for detecting low levels of nitrosylated protein in samples. However, as mentioned above, it requires a nitric oxide analyzer, which is not available in many laboratories. In this method, NO is displaced from S-NO bonds by mercuric ion (HgCl2). The released NO rapidly reacts with dissolved oxygen in samples to form the stable end-product nitrite (NO2–). In the presence of potassium iodide (KI) in acetic acid, NO2– is reduced back to NO. I− + NO2− + 2H+ → NO + 1⁄2I2 + H2O Free NO is detected by chemiluminescence in a nitric oxide analyzer (NOA, Sievers). In the NOA, NO reacts with ozone, forming excited-state NO2 that emits light (Fang et al., 1998).
Analysis of Protein S-Nitrosylation
Materials Cells expressing protein of interest Phosphate-buffered saline (PBS; APPENDIX 2E) NP-40 lysis buffer (see recipe) with and without freshly dissolved protease inhibitors, ice-cold 2 mg/ml protein A– or protein G–Sepharose bead slurry (Amersham Biosciences) 0.4 µg/ml normal rabbit serum Antibody directed against the protein of interest Isotype-matched control antibody High-salt lysis buffer (see recipe), ice cold, containing freshly dissolved 1 mM N-ethylmaleimide (NEM; add from 100 mM stock in ethanol) and protease inhibitors PBS (APPENDIX 2E) containing 1 mM EDTA (store up to 1 year at 4°C) 2× SDS-PAGE sample buffer (see recipe) Glycine elution buffer (see recipe) Bovine serum albumin (BSA) Polyacrylamide gel (UNIT 10.1) Silver staining kit (Sigma) Glacial acetic acid Helium or argon gas
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KI solution (see recipe) Glacial acetic acid Antifoaming agent (Sievers), diluted 1:30 in cold H2O (stable at room temperature up to several years) 100 mM HgCl2 (store up to 1 year at room temperature) NaNO2 (Sigma-Aldrich) Refrigerated centrifuge (e.g., with Sorvall RTH-750 rotor) Platform rocker Boiling water bath Nitric oxide analyzer (Sievers NOA) and associated software Hamilton syringe capable of delivering 50-µl aliquot, airtight Additional reagents and equipment for SDS-PAGE (UNIT 10.1) Prepare cell lysates 1. Collect 1 × 108 cells. The number of cells required for the immunoprecipitation will depend on the intracellular abundance of the protein of interest. Use sufficient cells to immunoprecipitate at least 3 nmol of a protein of interest.
2. Centrifuge cells 10 min at 300 × g (1200 rpm in a Sorvall RTH-750 rotor), 4°C. Discard supernatant and resuspend cell pellet in 50 ml ice-cold PBS. Repeat centrifugation and again discard the supernatant. 3. Lyse cell pellet in 1 to 1.5 ml of NP-40 lysis buffer containing freshly dissolved protease inhibitors. Alternative lysis buffers may be utilized (e.g., high-salt buffer), depending on the protein being studied. The volume of buffer added will depend on the cell number used for the immunoprecipitation. In general, the authors lyse 60 × 106 cells in 1 ml of lysis buffer.
4. Incubate the cell lysates in the dark at 4°C for 30 min on a platform rocker. Prepare the “preclear” beads while this incubation is in progress (see step 7).
5. Remove insoluble debris from the lysates by centrifuging 10 min at 14,000 × g, 4°C. 6. Divide the supernatants into two equal aliquots in fresh ice-cold microcentrifuge tubes and keep in the dark at 4°C until preclear beads are ready. Do not leave on ice for >30 min. Preclear cell lysates It is important to obtain clean immunoprecipitates for nitrosylation measurements to ensure that the NO signal is derived from the protein of interest rather than from contaminating proteins in the immunoprecipitate. Therefore, to reduce the level of background proteins in the immunoprecipitates, cell lysates are “precleared” with irrelevant antibody bound to protein A–Sepharose beads. The beads remove proteins that nonspecifically bind to antibody or to protein A–Sepharose from the cell lysates. 7. While lysates are rocking for 30 min (step 4), prepare “preclear beads” as follows. a. Resuspend 0.5 ml of a 2 mg/ml slurry of protein A–Sepharose beads in 0.5 ml of NP-40 lysis buffer in a 1.5-ml microcentrifuge tube on ice. If protein G–Sepharose is used for immunoprecipitation (step 16), substitute protein G–Sepharose for protein A–Sepharose in this step.
b. Let the beads sink to the bottom of the tube, then remove the supernatant. c. Repeat steps 7a and 7b an additional three times. d. Resuspend the beads in volume of NP-40 lysis buffer equal to the volume of the beads.
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8. Incubate 50 µl of the washed bead suspension with 20 µl of a 0.4 µg/ml stock of rabbit serum in 300 µl of NP-40 lysis buffer on a platform rocker for 30 min at 4°C. 9. Pellet the beads by pulsing briefly in a microcentrifuge for 2 to 3 sec, remove supernatant, add 1 ml of NP-40 lysis buffer, microcentrifuge again with a 2- to 3-sec pulse, and remove supernatant. Repeat wash three more times. 10. After the last wash, resuspend the beads in NP-40 lysis buffer to a final volume of 50 µl. These beads are referred to as “preclear beads.”
11. Add 25 µl of the suspension of preclear beads prepared in step 10 to each of the two aliquots of cell lysate from step 6. 12. Incubate the cell lysates with the preclear beads on a platform rocker for 2 hr at 4°C in the dark. 13. Pellet the beads by pulsing briefly in a microcentrifuge for 2 to 3 sec. 14. Transfer the supernatants to fresh ice-cold microcentrifuge tubes and discard the beads. Immunoprecipitate protein of interest 15. Add an appropriate amount of antibody directed against the protein of interest to one of the precleared aliquots of cell lysate and add an equal concentration of isotypematched control antibody to the other, corresponding, precleared aliquot of cell lysate. Incubate overnight in the dark on a platform rocker at 4°C. 16. After incubation, add an appropriate amount (usually 25 to 50 µl) of protein A–Sepharose bead suspension (step 7d) to each tube. Incubate samples 2 hr in the dark on a platform rocker at 4°C. Substitute protein G–Sepharose beads when using antibodies that do not bind protein A.
17. Collect immunoprecipitates on the beads by pulsing briefly in a microcentrifuge for 2 to 3 sec. Remove supernatant and wash the beads four times, each time by adding 1 ml high-salt lysis buffer containing freshly dissolved protease inhibitors and 1 mM NEM, collecting the beads by microcentrifuging with a 2- to 3-sec pulse, and removing the supernatant. NEM is added to block free thiols and thereby prevent artifactual S-nitrosylation when samples are acidified prior to S-nitrosylation measurements.
18. After the fourth wash, resuspend the immunoprecipitates in 1 ml high-salt lysis buffer containing NEM and protease inhibitors. 19. Transfer 1⁄10 (100 µl) of each resuspended immunoprecipitate to a fresh tube, add 900 µl of PBS containing 1 mM EDTA, collect the beads by pulsing in a microcentrifuge for 2 to 3 sec, and discard the supernatant. 20. Add 1 vol (usually 25 µl) of 2× SDS-PAGE sample buffer to each tube from step 19, and incubate in a boiling water bath for 3 to 5 min. These samples will be used for silver stained gels (see steps 25 to 27 below).
21. Collect the remaining 9⁄10 of each immunoprecipitate (from step 18) by pulsing briefly in a microcentrifuge for 2 to 3 sec. Discard the supernatant. 22. Elute the antigen-antibody complexes from the beads by adding 70 µl of glycine elution buffer and incubating for 10 min in the dark on ice. Analysis of Protein S-Nitrosylation
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23. Collect the beads by pulsing in a microcentrifuge for 2 to 3 sec. Transfer the supernatants (containing eluted proteins) to a fresh ice-cold microcentrifuge tube and keep in the dark on ice. 24. Repeat the glycine incubation/elution (steps 22 and 23) two more times, pooling the supernatants containing eluted antigen/antibody complexes. Freeze at −80°C (stable up to 1 week) until ready to use for chemical reduction/chemiluminescence assays (step 33). Antigen/antibody complexes must be eluted from protein A–Sepharose beads because the beads interfere with subsequent chemical reduction/chemiluminescence measurements.
Analyze immunoprecipitates by silver staining 25. Prepare 2-fold serial dilutions (ranging from 500 ng to 4 ng) of a known concentration of the protein of interest in PBS. If purified preparations of the protein of interest are not available, serially dilute known concentrations of bovine serum albumin (BSA). Add an equal volume of 2× SDS-PAGE sample buffer to each dilution and heat in boiling water bath. 26. Load the samples containing 1/10 of each immunoprecipitate (from step 20) and the serial dilutions of protein standards (from step 25) on a polyacrylamide gel. Separate the proteins by SDS-PAGE (UNIT 10.1). 27. Silver stain the gels using a silver stain kit per the manufacturer’s instructions. Determine the quantity of protein in each immunoprecipitate by comparing the intensity of bands in the immunoprecipitate to the intensity of bands corresponding to known concentrations of protein standards. In addition, assess the presence of contaminating background proteins in the immunoprecipitates. It is important for subsequent nitrosylation measurements that the protein of interest be the only protein precipitated by its specific antibody, and not by control antibody.
Perform chemical reduction/chemiluminescence assay 28. Pipet 5 ml of glacial acetic acid into the purge vessel of the Sievers NOA nitric oxide analyzer. Purge the vessel with helium or argon gas for several minutes with the outlet stopcock closed and the screw cap off as described in the nitrite reduction section of the operations manual (Sievers, 1999). 29. Add 1 ml of KI solution to the purge vessel and let purge for several more minutes. 30. Add 100 ml of antifoaming agent to the purge vessel. 31. Install the screw cap and open the outlet stopcock of the purge vessel and allow the baseline reading to stabilize. 32. To generate standard curves, make two-fold serial dilutions of NaNO2 in water (1 mM to 15 nM). Introduce 50 µl of the first standard into the reaction chamber of the NOA using an airtight Hamilton syringe as described in the operations manual for the NOA (the NOA will plot the NO released from each sample as the NO chemiluminescence signal in arbitrary units on the y axis versus the time over which the signal is measured on the x axis as shown in Fig 14.6.2; the area under each curve is proportional to the amount of NO in the sample). Once the NO tracing has leveled off to baseline, inject the next standard. After all the standards have been injected, use the Create Calibration command on the NOA to generate a standard curve as explained in the operations manual. The standard curve will be used to calculate the concentration of NO in subsequent samples.
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– +
– +
– +
– +
30
60
125
250
500
[GSNO] (nM)
buffer
buffer
– +
30 60 125 250 500
[NO2–] (nm)
Figure 14.6.2 Chemical reduction/chemiluminescence assay. Varying concentrations of S-nitrosoglutathione (GSNO) were treated with (+) or without (–) mercuric chloride for 10 min at room temperature and 20 min on ice. The NO content of the samples was then determined by chemical reduction/chemiluminescence in a solution of potassium iodide in glacial acetic acid. The NO-derived chemiluminescence signal (arbitrary units) is plotted on the y-axis, and the time course over which the signal was measured is plotted on the x-axis. The area under each curve is proportional to the NO released from each sample. The NO detected in buffer alone (buffer), in nitrite standards (NO2−), and in GSNO samples not treated with mercuric chloride is due to background nitrite. The increase in NO signal detected in GSNO samples treated with mercuric chloride is the SNO-specific signal.
33. Divide each sample of eluted antigen/antibody complex (from step 24) into two equal aliquots. Add 100 mM HgCl2 to one aliquot for a final concentration of 2.2 mM, then incubate for 10 min at room temperature in the dark. HgCl2 displaces NO from S-NO bonds.
34. Introduce 50 µl of each sample into the reaction chamber using an airtight Hamilton syringe. The NOA will plot the NO chemiluminescence signal of each sample. The Calculate Concentrations command on the NOA is then used to calculate the concentration of NO in each sample as explained in the operations manual of the NOA. The NOA will plot the NO chemiluminescence signal of each sample. The Calculate Concentrations command on the NOA is then used to calculate the concentration of NO in each sample as explained in the operations manual of the NOA. The NO signal in the samples that have not been treated with HgCl2 is generated from background nitrite in the samples. The increase in NO signal above background in the HgCl2-treated samples is the SNO-specific signal (Fig. 14.6.2).
Analysis of Protein S-Nitrosylation
There are a variety of other chemical reduction methods that can be used to detect protein S-nitrosylation, including use of a saturated solution of CuCl in 100 mM cysteine (Fang et al., 1998). The optimal chemical reduction method for detecting protein S-nitrosylation will depend on the protein of interest, so if S-nitrosylation is not detected with one method, an alternate method should be employed.
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S-NITROSYLATION ANALYSIS OF PURIFIED INTRACELLULAR PROTEINS USING A DAN ASSAY
ALTERNATE PROTOCOL 1
If a nitric oxide analyzer is not available in a laboratory, then S-nitrosylation of purified proteins can be measured using a DAN assay. In the DAN assay, NO+ is released from S-NO bonds on S-nitrosylated proteins by HgCl2. The reagent 2,3-diaminonapthalene (DAN) reacts with the released NO+ to form a fluorescent product. The extent of DAN fluorescence is proportional to the amount of SNO in each sample (Park et al., 2000; Haendeler et al., 2002). The disadvantage of the DAN method is that it is not as sensitive as chemical reduction/chemiluminescence. Therefore endogenously S-nitrosylated proteins that are unstable or not abundant may not be detected by the DAN method. Additional Materials (also see Basic Protocol 2) 10 mM 2,3-diaminonapthalene (DAN) in dimethylformamide or dimethylsulfoxide (store up to 4 to 5 months at −20°C) 10 mM HgCl2 in H2O (store up to 1 year at room temperature) 1 N NaOH Fluorimeter capable of excitation at 375 nm and emission detection at 450 nm 1. Perform immunoprecipitation to obtain eluted antigen/antibody complexes in glycine elution buffer (see Basic Protocol 2, steps 1 to 24) and analyze the immunoprecipitates by silver staining (see Basic Protocol 2, steps 25 to 27). 2. Divide each sample of eluted antigen/antibody complex (see Basic Protocol 2, step 24) into two equal aliquots. To one aliquot, add 10 mM DAN to a final concentration of 100 µM; to the other, add 10 mM HgCl2 to a final concentration of 100 µM plus DAN at a final concentration of 100 µM. 3. Incubate samples in the dark for 30 min at room temperature on a platform rocker. 4. To each sample, add 1 N NaOH to a final concentration of 0.1 N. 5. Measure the amount of fluorescent product formed when NO+ released from S-nitrosylated proteins reacts with DAN, using a fluorimeter with an excitation wavelength of 375 nm and an emission wavelength of 450 nm. 6. Subtract the fluorescence intensity generated in the absence of HgCl2 (background) from the fluorescence intensity generated in the presence of HgCl2 to determine the SNO-specific fluorescence in each sample. Generate standard curves using known concentrations of in vitro nitrosylated proteins or GSNO (Fig. 14.6.3). GSNO can be used to provide an approximate estimate of the amount of S-nitrosylated protein in samples. However, the stability of S-NO bonds varies between proteins. Therefore, ideally, standard curves should be generated using in vitro S-nitrosylated preparations of the protein of interest.
S-NITROSYLATION ANALYSIS OF CELLS OR CELL LYSATES USING THE BIOTIN-SWITCH METHOD The previously described methods are useful for S-nitrosylation analysis of individual immunoprecipitated proteins. The methods are not as well suited for analyzing general protein S-nitrosylation in cell lysates because many components in lysates, including proteins, nitrite, oxidants, and antioxidants interfere with the assays (Fang et al., 1998; Nagata et al., 1999; Fernandez-Cancio et al., 2001; Jourd’heuil, 2002; Zhang et al., 2002). Therefore, to assess general protein nitrosylation in cell lysates, it is recommended that either a biotin-switch or immunofluorescence assay (Alternate Protocol 2) be used.
BASIC PROTOCOL 3
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500 450
Relative fluorescence
400 350 300 250 200 150 100 50 0 0
25
50
100
500
1000
nM of GSNO
Figure 14.6.3 DAN assay. Serial dilutions of GSNO in glycine elution buffer were incubated with DAN (100 µM) in the presence (+Hg; broken line) or absence (−Hg; solid line) of HgCl2. DAN fluorescence was then measured on a Perkin Elmer LS45 luminescence spectrometer with an excitation wavelength of 375 nm and an emission wavelength of 450 nm. The assay was performed in triplicate and data are presented as the mean standard deviation.
In the biotin-switch method, S-NO bonds are selectively labeled with biotin and then either purified on streptavidin agarose or detected on an anti-biotin immunoblot (Jaffrey et al., 2001; Figure 14.6.4). In the first part of the procedure, free thiols are blocked with the thiol-specific compound methyl methanethiosulfonate (MMTS, contained in the blocking buffer). In the second part, S-NO bonds are reduced with ascorbic acid to generate new free thiols. Finally, the new free thiols are biotinylated and detected on immunoblots or purified by streptavidin-affinity chromatography. The method has been used to detect both endogenously S-nitrosylated proteins in cell lysates and immunoprecipitates, and proteins that have been in vitro S-nitrosylated. A biotin-switch kit is also commercially available (NitroGlo, Perkin-Elmer).
Analysis of Protein S-Nitrosylation
Materials Tissue culture cells of interest Phosphate-buffered saline (PBS; APPENDIX 2E) Cell lysis buffer (with protease inhibitors; see recipe) Blocking buffer (see recipe) Acetone, –20°C HENS buffer: HEN buffer (see recipe) containing 1% (w/v) SDS [add 0.04 vol 25% (w/v) SDS]; stable up to 6 months at room temperature wrapped in foil to protect from light 50 mM stock solution of N-[6-(biotinamido)hexyl]-3′-(2′-pyridyldithio)propionamide (biotin-HPDP; Pierce) in dimethylformamide (DMF): store up to 1 month at –20°C Dimethylsulfoxide (DMSO)
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50 mM sodium ascorbate, freshly prepared in H2O 2× SDS sample buffer without reducing agents (see recipe) 2.5% (w/v) nonfat dry milk (store up to 1 week at 4°C) Anti-biotin mouse monoclonal primary antibody (Pierce) PBS (APPENDIX 2E) containing 0.1% (v/v) Tween 20 (store up to several years at 4°C) Horseradish peroxidase–conjugated anti-mouse secondary antibody (Amersham Biosciences) Enhanced chemiluminescence (ECL) detection kit (Amersham Biosciences) Neutralization buffer (see recipe) Streptavidin agarose (Sigma) or NeutrAvidin agarose (Pierce) Neutralization buffer (see recipe) containing 600 mM NaCl Elution buffer (see recipe) 2× SDS sample buffer with reducing agents (see recipe) Silver staining kit (optional; Sigma) Tissue homogenizer Refrigerated centrifuge (e.g., with Sorvall RTH-750 rotor) 50°C water bath Additional reagents and equipment for protein assay (UNIT 3.4), SDS-PAGE (UNIT 10.1), blotting onto nitrocellulose membranes (UNIT 10.7), and enhanced chemiluminescence (ECL) detection (UNIT 10.10)
O CH S S CH3
SH
S S
CH3
O S
S
NO
NO
MMTS
S
S
S
S MMTS removed by spin column or acetone
S
S
CH3
S
S
biotin
S N
S
CH3
S–S–biotin SH
biotin – HPDP S
S
S
S
Figure 14.6.4 Schematic diagram of the biotin switch assay. A theoretical protein is indicated with cysteines in either the free-thiol, disulfide, or nitrosothiol conformation. In the first step, free thiols are rendered unreactive by methylthiolation with MMTS. This modification can be reversed by reduction with 2-mercaptoethanol (2-ME). MMTS is removed in the next step by acetone precipitation. Nitrosothiols are selectively reduced with ascorbate to re-form the thiol, which is then reacted in the final step with the thiol-modifying reagent biotin-HPDP. The biotin-labeled proteins are then either detected on an anti-biotin immunoblot or are purified on streptavidin-agarose. Modified and reprinted with permission from Jaffrey et al. (2001).
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Prepare samples 1. Collect tissue culture cells (enough to obtain between 1 and 5 mg total protein) by centrifuging 5 min at 300 × g (1200 rpm in a Sorvall RTH-750 rotor), 4°C. Discard supernatant. Resuspend cells in 50 ml ice-cold PBS. Repeat centrifugation and again discard supernatant. 2. Lyse cells by resuspending pellet in 1 ml cell lysis buffer per 60 × 106 cells. Rock sample on a platform rocker for 30 min at 4°C in the dark. 3. Centrifuge insoluble debris for 10 min at 14,000 g at 4°C. Recover the supernatant and assay protein concentration using Bradford or Lowry assay (UNIT 3.4). 4. Adjust protein concentration to 0.8 µg/µl with cell lysis buffer. This assay has also been used to detect S-nitrosylated proteins in tissue samples (Jaffery et al., 2001) and in immunoprecipitates of proteins (Kim et al., 2004).
Block free thiols in protein samples 5. Add 4 vol blocking buffer and incubate 20 min at 50°C, vortexing frequently but gently to avoid foaming. 6. Remove the blocking buffer by acetone precipitation as follows. a. Add 10 vol of prechilled (–20°C) acetone to the sample. b. Incubate 20 min at −20°C, then centrifuge 10 min at 2000 × g, 4°C. c. Resuspend pellet in 0.1 ml HENS buffer per mg of protein in the starting sample. Biotinylate nitrosothiols 7. Just before use, dilute the 50 mM stock of biotin-HPDP in DMSO to a final concentration of 4 mM to prepare the labeling solution. Vortex before use to ensure even suspension of the biotin-HPDP. 8. To the blocked protein sample from step 6, add one-third vol labeling solution and 0.02 vol 50 mM sodium ascorbate. Incubate 1 hr at room temperature. As a control for endogenously biotinylated proteins in samples, treat some samples with one-third vol DMSO alone, instead of labeling solution. Prepare a fresh solution of sodium ascorbate for each experiment. After this step, samples do not need to be protected from light.
Analyze samples In addition to identifying biotin-labeled S-nitrosylated proteins on anti-biotin immunoblots (steps 9a to 13a), biotinylated proteins can be purified on streptavidin agarose (steps 9b to 16b). The purified S-nitrosylated proteins are then identified by mass spectrometry (Chapter 16) or analyzed on immunoblots for specific protein(s) of interest. To detect biotinylated proteins by immunoblotting 9a. Add an equal volume of SDS-PAGE sample buffer without reducing agents to samples. Do not boil samples or use reducing agents in the SDS-PAGE sample buffer. Sample buffer should be added directly to the samples and then immediately loaded onto the gel.
10a. Analyze samples by electrophoresis on a SDS-PAGE gel (UNIT 10.1) and transfer to nitrocellulose (UNIT 10.7). Analysis of Protein S-Nitrosylation
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11a. Block nonspecific binding sites on the blot by incubating with 2.5% nonfat dry milk for 30 min at room temperature Wash the blot 5 min at room temperature in PBS containing 0.1% Tween 20. Repeat wash two more times. 12a. Incubate the membrane with a 1:1000 dilution of an anti-biotin mouse monoclonal primary antibody in PBS containing 0.1% Tween 20 for 2 hr at room temperature. Decant the antibody and repeat the wash three times as in step 11a. 13a. Incubate the membrane with a 1:1000 dilution of horseradish peroxidase-conjugated anti-mouse secondary antibody in PBS containing 0.1% Tween 20. Wash the blot five times as in step 11a, then develop the immunoblot with enhanced chemiluminescence (ECL) according to the manufacturer’s instructions (also see UNIT 10.10). Purify biotinylated proteins 9b. Remove biotin-HPDP by adding 2 vol of prechilled (−20°C) acetone. Incubate 20 min at −20°C, then centrifuge 10 min at 2000 × g, 4°C. Discard supernatant and gently wash walls of tube, as well as surface of pellet, with −20°C acetone to remove traces of biotin-HPDP. 10b. Resuspend pellet in 0.1 ml HENS buffer per mg of protein in the initial protein sample. 11b. Add 2 vol neutralization buffer. 12b. Add 15 µl of packed streptavidin-agarose per mg of protein used in the initial protein sample and incubate 1 hr at room temperature. NOTE: NeutrAvidin-agarose (Pierce) has been used with identical results.
13b. Wash beads five times, each time by centrifuging 5 sec at 200 × g, room temperature, removing the supernatant, then adding 10 vol neutralization buffer containing 600 mM NaCl. Remove supernatant after final wash. 14b. Incubate beads with an equal volume of elution buffer to recover the bound proteins. 15b. Add an equal volume of 2× SDS-PAGE sample buffer with reducing agent to the eluted protein samples. Perform SDS-PAGE (UNIT 10.1) and transfer the electrophoresed samples to nitrocellulose (UNIT 10.10) for immunoblotting. 16b. Probe the blot using antibodies specific for a protein of interest as described in steps 11a to 13a, above. Alternatively, silver stain the gels using a kit according to the manufacturer’s instructions, excise desired bands, and identify proteins by mass spectrometry (Chapter 16). S-NITROSYLATION ANALYSIS OF CELLS OR CELL LYSATES USING ANTI-SNO IMMUNOFLUORESCENCE Recently, an antiserum has been produced that recognizes the S-NO moiety. The antiserum was generated against S-nitrosylated cysteine conjugated to BSA and detects S-nitrosylated proteins by immunohistochemical and immunofluorescence staining (Gow et al., 2002). This procedure can be used to detect proteins that are S-nitrosylated by endogenous sources of NO in cells. The limits of detection of the assay are not known and may vary from protein to protein. However, endogenously S-nitrosylated proteins have been detected using this assay in a variety of cell types (Gow et al., 2002).
ALTERNATE PROTOCOL 2
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Materials Cells of interest growing or plated on coverslips (APPENDIX 3C) 4% (w/v) paraformaldehyde in PBS (see APPENDIX 2E for PBS) Normal goat serum (Sigma) 0.2% (w/v) HgCl2 in PBS (see APPENDIX 2E for PBS) Primary antibody: rabbit polyclonal anti–SNO cysteine antibody (Alexis Biochemicals) Secondary antibody: FITC-conjugated F(ab′)2 fragment of anti-rabbit antibody Phosphate-buffered saline (PBS; APPENDIX 2E) 0.1% (w/v) sodium borohydride in PBS Humidified chamber: sealable container containing moistened paper towels Fluorescence microscope with HQ470140X EX filter, Q495LP BS filter, and HQ525/50m EM filter (Chroma Technology) 1. Fix cells on coverslips in PBS-buffered 4% paraformaldehyde for 30 min at room temperature. The optimal length of fixation is tissue and cell-type dependent. Paraffin-embedded tissue samples can also be stained with the anti-SNO antibody
2. Cover the cells on the coverslip with 100% normal goat serum and incubate 30 min at room temperature in a humidified chamber to block nonspecific protein binding. 3. For negative controls, cover the cells on the coverslip with 0.2% HgCl2 and incubate 30 min at room temperature in a humidified chamber. 4. Cover cells on the coverslip with the primary antibody (anti–SNO cysteine antibody) diluted 1:50 in normal goat serum, and incubate overnight at 4°C in a humidified chamber. 5. Cover cells on the coverslip with the secondary antibody [FITC-conjugated F(ab′)2 fragment] diluted 1:100 in PBS and incubate 1 to 2 hr at room temperature in a humidified chamber in the dark. 6. Cover cells on the coverslip with 0.1% sodium borohydride and incubate 30 min at room temperature in the dark in a humidified chamber to reduce autofluorescence. 7. Examine cells using a fluorescence microscope using HQ470140X EX filter, Q495LP BS filter, and HQ525/50m EM filter. Figure 14.6.5 shows an example of results obtained via this method.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Blocking buffer HEN buffer (see recipe) containing: 2.5% (w/v) SDS 20 mM methyl methanethiosulfonate (MMTS; Sigma) Prepare fresh for each experiment Cell lysis buffer HEN buffer (see recipe) containing: 0.5% (w/v) CHAPS 0.1% (w/v) SDS Analysis of Protein S-Nitrosylation
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Figure 14.6.5 Anti-SNO Immunofluorescence. Inducible nitric oxide synthase (iNOS) expression was induced in RAW 254.7 (macrophage) cells with bacterial lipopolysacharide (200 ng/ml) and γ-interferon (200 U/ml). Levels of intracellular SNOs in untreated control cells (basal) or cytokinetreated cells (cytokine) were then determined by immunofluorescent staining using an anti-SNO antibody. Reprinted with permission from Gow et al. (2002).
Just before, use add protease inhibitor stock solutions (see recipe) to the following final concentrations: 10 µg/ml leupeptin 5 µg/ml aprotinin 1 mM PMSF Elution buffer 20 mM HEPES pH 7.7 100 mM NaCl 1 mM EDTA 100 mM 2-mercaptoethanol Prepare fresh for each experiment Glycine elution buffer Prepare 100 mM glycine, pH 3, in water. Store up to 3 months at 4°C. Heat the solution in a sealed vessel at 95°C for 2 hr prior to use in order to decrease nitrite contamination, then cool solution to 4°C just prior to use. HEN buffer 250 mM HEPES pH 7.7 1 mM EDTA 0.1 mM neocuproine (Sigma) Store up to 1 year at room temperature wrapped in foil to protect from light High-salt lysis buffer 500 mM NaCl 1% (v/v) NP-40 50 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) continued
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Store up to 1 year at 4°C If indicated in protocol, just before use, add protease inhibitors (see recipe for stock solutions) as follows: 10 µg/ml leupeptin 5 µg/ml aprotinin 1 mM PMSF If indicated in protocol, just before use, add N-ethylmaleimide (NEM) to 1 mM final concentration (from 100 mM stock in ethanol)
KI solution Dissolve ∼50 mg KI in 1 ml of deionized water. Prepare fresh for each experiment. Accurate measurement is not necessary since this is an excess of the reagent.
Neutralization buffer 20 mM HEPES pH 7.7 100 mM NaCl 1 mM EDTA 0.5% (v/v) Triton X-100 Store up to 6 months at room temperature NP-40 lysis buffer 150 mM NaCl 1% (v/v) NP-40 50 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) Store up to 1 year at 4°C If indicated in protocol, just before use, add protease inhibitors (see recipe for stock solutions) as follows: 10 µg/ml leupeptin 5 µg/ml aprotinin 1 mM PMSF Protease inhibitor stock solutions Leupeptin: Prepare 5 mg/ml leupeptin stock solution in PBS (APPENDIX 2E). Store up to 6 months as single aliquots at −20°C. Aprotinin: Prepare 1.5 mg/ml aprotinin stock solution in 0.9% (w/v) NaCl/0.9% (v/v) benzyl alcohol (Sigma). Store up to 4 years at 4°C. PMSF: Prepare 200 mM PMSF stock solution in anhydrous isopropyl alcohol. Store up to 9 months at 4°C. SDS-PAGE sample buffer, 2× 125 mM Tris⋅Cl, pH 6.8 (APPENDIX 2E) 20% (v/v) glycerol 4% (w/v) SDS 0.004% (w/v) bromphenol blue Store up to 6 months at room temperature Just before use, add 2-mercaptoethanol (2-ME) to 10% (v/v). If the protocol indicates 2× SDS-PAGE sample buffer without reducing agents, do not add 2-ME. Other sample buffers may be appropriate depending on the gel system used but reducing agents should not be included.
Analysis of Protein S-Nitrosylation
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COMMENTARY Background Information Nitric oxide (NO) is a free radical gas that regulates a wide variety of biological responses including vasodilation, smooth muscle relaxation, neurotransmission, and apoptosis. One of the mechanisms by which NO regulates these processes is via S-nitrosylation of proteins. S-nitrosylation is the binding of a NO group to a cysteine (or other thiol). The function of many proteins has been shown to be regulated by S-nitrosylation in cell-free systems, and an increasing number of proteins have been shown to be endogenously S-nitrosylated in cells. Recent reviews give a thorough description of this field (Lane et al., 2001; Stamler et al., 2001; Mannick and Schonhoff, 2002). Like phosphorylation, S-nitrosylation is a rapidly reversible and precisely targeted posttranslational modification that serves as an on/off switch for protein function during cell signaling. However, unlike phosphorylation and dephosphorylation, S-nitrosylation and denitrosylation are not strictly enzyme-independent reactions. Although enzymatically mediated S-nitrosylation (Inoue et al., 1999) and denitrosylation (Liu et al., 2001) have been described, S-nitrosylation and denitrosylation are also mediated by enzyme-independent redox chemistry in cells. Instead, S-nitrosylation and denitrosylation are mediated, at least in part, by redox-regulated chemical reactions in cells. Alterations in pH, pO2, cellular reductants, transition metals, and UV light lead to the loss and/or gain of S-NO bonds. Thus, S-nitrosylation transduces intracellular redox shifts into alterations in protein function and cell signaling. The intracellular chemistry that results in S-NO bond formation is incompletely understood. In general, NO+ or an equivalent (such as N2O3), rather than native NO, reacts with thiols. In addition, NO groups are directly transferred from one S-nitrosylated peptide or protein to another via transnitrosation reactions.
Critical Parameters and Troubleshooting While S-nitrosylation of critical cysteine residues has been shown to regulate the function of many proteins in cell-free systems, it has been much more challenging to identify proteins that are endogenously nitrosylated in cells. This difficulty is due in part to the redoxsensitive nature of the S-NO bond, which often leads to the artifactual loss and/or gain of S-NO
bonds during sample preparation. A simple immunoblot method of detecting S-nitrosylated proteins has not been developed, because it has been very difficult to generate antibodies against the unstable S-NO bond. In addition, proteins often lose their S-NO bonds during electrophoresis. Despite these challenges, several methods have been developed to measure intracellular protein nitrosylation with reasonable accuracy. However, all currently available methods have drawbacks, and therefore S-nitrosylation measurements should be confirmed with at least two different methods. In each method, great care needs to be taken during sample preparation to prevent disruption of S-NO bond formation. For example, UV light exposure, reducing agents, and pH shifts must be avoided during sample preparation. In addition, metal chelators must be included in buffers to prevent artifactual metal-catalyzed SNO bond gain and loss. The precise conditions favoring S-NO bond stability vary between proteins. For instance, some SNO bonds are most stable at acidic pH, whereas others are most stable at neutral pH. Therefore, the method with the optimal sensitivity and specificity is likely to vary between S-nitrosylated proteins. It is erroneous to assume that, if a method detects an S-nitrosylated standard such as GSNO or SNO-BSA, then it will necessarily detect another S-nitrosylated protein of interest. Ideally, S-nitrosylation analysis should be performed using conditions optimized for each protein of interest. In vitro S-nitrosylation experiments The simplest method of assessing protein nitrosylation is to S-nitrosylate a purified recombinant protein in vitro with an SNO donor compound and to measure S-nitrosylation using a Saville assay (see Basic Protocol 1). An advantage of in vitro S-nitrosylation experiments is that sensitive detection methods are not required if relatively high (micromolar) concentrations of recombinant proteins are used. Site-directed mutagenesis can also identify the cysteine residue(s) that are the S-nitrosylation targets. As is the case with phosphorylation, consensus motifs target specific cysteine residues for S-nitrosylation (Stamler, 1994; Lane et al., 2001; Stamler et al., 2001; Mannick and Schonhoff, 2002). Functional assays can be performed to determine if in vitro S-nitrosylation alters protein activity. A major limitation of in vitro S-nitrosylation studies is that
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they do not determine the physiologic relevance of protein S-nitrosylation. To establish physiologic relevance, it is important to demonstrate that a protein is S-nitrosylated intracellularly by endogenous sources of NO. Analysis of intracellular protein S-nitrosylation Studies of intracellular nitrosylation are much more difficult than studies of in vitro nitrosylation because the levels of S-nitrosylated proteins in cells approach the limits of detection of currently available technology. In general, a protein of interest must be immunoprecipitated from cells, and its SNO content determined by chemical reduction/chemiluminescence (see Basic Protocol 2) or DAN assay (see Alternate Protocol 1). In the authors’ experience, a minimum of 30 nM of protein is required to reliably detect S-nitrosylation by chemical reduction/chemiluminescence, whereas higher concentrations (at least 100 nM) of S-nitrosylated protein are required for detection in DAN assays. If sufficient endogenous protein cannot be immunoprecipitated, then an alternative approach is to immunoprecipitate overexpressed proteins with the caveat that S-nitrosylation of overexpressed proteins may differ from S-nitrosylation of endogenous proteins.
Analysis of Protein S-Nitrosylation
Chemical reduction/chemiluminescence Although in the authors’ experience the chemical reduction/chemiluminescence method is currently the most sensitive and specific method for measuring protein S-nitrosylation, it also has several important limitations. One limitation is that it detects NO derived from nitrite as well as from SNOs. Detection of a SNO-specific NO signal is difficult if the concentration of nitrite in a sample is far higher than the concentration of SNOs. Some investigators add 5% sulfanilamide in 1 N HCl to samples prior to S-nitrosylation measurements in order to reduce nitrite background (Feelisch et al., 2002). However, addition of sulfanilamide also eliminates signal derived from some S-NO bonds (presumably due to HCl-induced protein denaturation and/or reaction of sulfanilamide with NO+ equivalents released from SNO bonds). Therefore, the authors do not generally recommend pretreating samples with sulfanilamide prior to SNO measurements. Another limitation of the chemical reduction/chemiluminescence method is that it does not distinguish between a NO signal derived from a protein of interest and a NO signal
derived from an irrelevant protein contaminating a sample. Therefore an immunoprecipitate must be scrupulously clean to ensure that the NO signal is derived from the protein of interest. If a clean immunoprecipitate is not obtainable, then an alternative approach is to determine if site-directed mutagenesis of a cysteine(s) on the protein of interest decreases the NO signal. A final limitation is that addition of mercuric chloride may acidify samples and thereby generate a SNO-independent NO signal. Thus, the pH of samples must be monitored after the addition of HgCl2 and buffers adjusted if necessary to avoid significant decreases in pH. DAN assay If a nitric oxide analyzer is not available, S-nitrosylation of immunoprecipitated proteins can be assessed using a DAN assay (see Alternate Protocol 1). In the authors’ experience, the DAN assay is less sensitive than chemical reduction/chemiluminescence, and a variety of substances in cell extracts, including proteins, may quench the fluorescence of DAN (Fernandez-Cancio et al., 2001). In addition, it is possible that DAN fluorescence increases via NO-independent mechanisms (Zhang et al., 2002). Therefore it is important to always include controls demonstrating that DAN fluorescence decreases when immunoprecipitates are obtained from cells that have been pretreated with NOS inhibitors. Detection of protein S-nitrosylation in cells or cell lysates Although the most accurate measures of protein nitrosylation are obtained using isolated proteins, it is often desirable to assess general protein nitrosylation in cells. However, as noted above, it is very difficult to analyze protein nitrosylation in cell lysates using either chemical reduction/chemiluminescence or DAN assays due to high nitrite backgrounds and competing reactions in cell lysates. The two methods that are most useful for measuring SNOs in cells or cell lysates are the biotin switch method (see Basic Protocol 3) or immunofluorescence microscopy (see Alternate Protocol 2). Biotin-switch assay In the biotin-switch method (see Basic Protocol 3), S-NO bonds are selectively labeled with biotin and then purified on streptavidinagarose or visualized on an anti-biotin immunoblot. This method has the advantage that
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it provides a general screen of nitrosylated proteins in cell lysates and is not hampered by nitrite background. In addition, individual Snitrosylated proteins labeled by the biotin switch assay can be identified by mass spectrometry (Jaffrey et al., 2001). The assay has also been used to detect S-nitrosylation of immunoprecipitated proteins (Kim et al., 2004). However, the sensitivity of the biotin switch assay is sometimes not as high as chemical reduction/chemiluminescence, due in part to loss of SNOs during sample preparation. In addition, false positive signals arise if free thiols are not completely blocked by MMTS prior to labeling with biotin. It is important to keep protein concentrations ≤0.8 µg/µl to maximize cysteine accessibility to MMTS. Finally, proteins that are endogenously biotinylated are a significant source of background signal in the assay. It is critical to include controls that are not labeled with biotinHPDP to control for endogenously biotinylated proteins. Immunofluorescence assay of SNOs Immunofluorescent staining of S-nitrosylated proteins using an anti-SNO antibody (Alternate Protocol 2) is relatively easy and provides data about the subcellular localization of S-nitrosylated proteins. However, the antibody also has been reported to detect non-nitrosylated thiols; therefore controls must be performed to verify the specificity of staining (Sun et al., 2001). In particular, it is important to demonstrate that NOS-inhibitor pretreatment of live cells, and HgCl2 pretreatment of fixed cells, decreases anti-SNO staining. In addition, it is not known if the anti-SNO antibody detects all S-nitrosylated proteins.
Anticipated Results Successful detection of protein S-nitrosylation is critically dependent on the concentration and stability of SNO in a sample. The optimal conditions for maintaining SNO stability vary between proteins. In general, it is anticipated that protein S-nitrosylation will be detectable by chemical reduction/chemiluminescence if 50 nM or more of a protein is cleanly immunoprecipitated. The DAN and biotin switch assays are generally less sensitive than the chemical reduction/chemiluminescence assay. Therefore, higher concentrations or more stable SNOs may be required for detection in these assays. The sensitivity of immunofluorescence is not known and specificity may be a problem, so controls must be performed to assure that a
fluorescent signal is derived from SNOs and not from non-nitrosylated thiols.
Time Considerations One day is required to S-nitrosylate proteins in vitro and measure their SNO content. Immunoprecipitation requires 1.5 days. Subsequent S-nitrosylation analysis of the immunoprecipitates by chemical reduction/chemiluminescence or DAN assays requires 0.5 to 1 day. Silver stain analysis of immunoprecipitated proteins requires 1.5 days. Biotin-switch assays require 1 day to prepare samples, a second day for SDS-PAGE, gel staining, and protein transfer, and a third day for immunoblotting. Immunofluorescence staining requires 1 day.
Literature Cited Fang, K., Ragsdale, N.V., Carey, R.M., MacDonald, T., and Gaston, B. 1998. Reductive assays for S-nitrosothiols: Implications for measurements in biological systems. Biochem. Biophys. Res. Commun. 252:535-540. Feelisch, M., Rassaf, T., Mnaimneh, S., Singh, N., Bryan, N.S., Jourd’Heuil, D., and Kelm, M. 2002. Concomitant S-, N-, and heme-nitros(yl)ation in biological tissues and fluids: Implications for the fate of NO in vivo. FASEB J. 16:1775-1785. Fernandez-Cancio, M., Fernandez-Vitos, E.M., Centelles, J.J., and Imperial, S. 2001. Sources of in te rf er ence in the u se of 2 ,3 -diaminonaphthalene for the fluorimetric determination of nitric oxide synthase activity in biological samples. Clin. Chim. Acta. 312:205-212. Gow, A.J., Chen, Q., Hess, D.T., Day, B.J., Ischiropoulos, H., and Stamler, J.S. 2002. Basal and stimulated protein S-nitrosylation in multiple cell types and tissues. J. Biol. Chem. 277:96379640. Haendeler, J., Hoffmann, J., Tischler, V., Berk, B.C., Zeiher, A.M., and Dimmeler, S. 2002. Redox regulatory and anti-apoptotic functions of thioredoxin depend on S-nitrosylation at cysteine 69. Nat. Cell Biol. 4:743-749. Jaffrey, S.R., Erdjument-Bromage, H., Ferris, C.D., Tempst, P., and Snyder, S.H. 2001. Protein S-nitrosylation: A physiological signal for neuronal nitric oxide. Nat. Cell Biol. 3:193-197. Jourd’heuil, D. 2002. Increased nitric oxide-dependent nitrosylation of 4,5-diaminofluorescein by oxidants: Implications for the measurement of intracellular nitric oxide. Free Radic. Biol. Med. 33: 676-684. Inoue, K., Akaike, T., Miyamoto, Y., Okamoto, T., Sawa, T., Otagiri, M., Suzuki, S., Yoshimura, T., and Maeda, H. 1999. Nitrosothiol formation catalyzed by ceruloplasmin: Implication for cytoprotective mechanism in vivo. J. Biol. Chem. 274:27069-27075.
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Kim, J.E. and Tannenbaum, S.R. 2004. S-Nitrosation regulates the activation of endogenous procaspase-9 in HT-29 human colon carcinoma cells. J. Biol. Chem. 279:9758-9764. Lane, P., Hao, G., and Gross, S.S. 2001. S-nitrosylation is emerging as a specific and fundamental posttranslational protein modification: Head-tohead comparison with O-phosphorylation. Sci. STKE 2001(86):RE1. Liu, L., Hausladen, A., Zeng, M., Que, L., Heitman, J., and Stamler, J.S. A metabolic enzyme for S-nitrosothiol conserved from bacteria to humans. Nature 410:490-494. Mannick, J.B. and Schonhoff, C.M. 2002. Nitrosylation: The next phosphorylation? Arch. Biochem. Biophys. 408:1-6. Nagata, N., Momose, K., and Ishida, Y. 1999. Inhibitory effects of catecholamines and anti-oxidants on the fluorescence reaction of 4,5-diaminofluorescein, DAF-2, a novel indicator of nitric oxide. J. Biochem. (Tokyo) 125:658-661. Ruiz, F., Corrales, F.J., Miqueo, C., and Mato, J.M. 1998. Nitric oxide inactivates rat hepatic methionine adenosyltransferase in vivo by S-nitrosylation. Hepatology 28:1051-1057. Samouilov, A. and Zweier, J.L. 1998. Development of chemiluminescence-based methods for specific quantitation of nitrosylated thiols. Anal. Biochem. 258:322-330. Saville, B. 1958. A scheme for the colorimetric determination of microgram amounts of thiols. Analyst 83:670-672.
Sievers. 1999. Sievers Nitric Oxide Analyzer NOATM280 Operation and Maintenance Manual. Sievers Instruments, Inc., Boulder, Co. Stamler, J.S. 1994. Redox signaling: Nitrosylation and related target interactions of nitric oxide. Cell 78:931-936. Stamler, J.S. and Feelisch, M. 1996. Preparation and detection of S-nitrosothiols. In Methods in Nitric Oxide Research (M. Feelisch and J.S. Stamler, eds.) p. 527. John Wiley and Sons. Chichester, U.K. Stamler, J.S., Lamas, S., and Fang, F.C. 2001. Nitrosylation: The prototypic redox-based signaling mechanism. Cell 106:675-683. Sun, J., Xin, C., Eu, J.P., Stamler, J.S., and Meissner, G. 2001. Cysteine-3635 is responsible for skeletal muscle ryanodine receptor modulation by NO. Proc. Natl. Acad. Sci. U.S.A. 98:1115811162. Zhang, X., Kim, W.S., Hatcher, N., Potgieter, K., Moroz, L.L., Gillette, R., and Sweedler, J.V. 2002. Interfering with nitric oxide measurements: 4,5-diaminofluorescein reacts with dehydroascorbic acid and ascorbic acid. J. Biol. Chem. 277:48472-48478.
Contributed by Joan B. Mannick and Christopher M. Schonhoff University of Massachusetts Medical School Worcester, Massachusetts
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Tyrosine O-Sulfation
UNIT 14.7
Tyrosine O-sulfation of proteins is a widespread post-translational modification that occurs in all animals. The sulfate transfer to tyrosine residues is catalyzed by one of two tyrosylprotein sulfotransferase (TPST) isoenzymes (EC 2.8.2.20; Fig. 14.7.1), which are integral membrane proteins of the trans-Golgi network (TGN), with their catalytic site facing the lumen. Consequently, all tyrosine-sulfated proteins must transit through the TGN. Moreover, the recognition by TPST of the tyrosine residue to be sulfated in a particular protein requires the presence of certain structural features (see Strategic Planning). Tyrosine sulfate in proteins can be identified by various methods, including metabolic labeling using inorganic [35 S]-sulfate (see Basic Protocol 1) followed by tyrosine sulfate analysis (see Basic Protocol 2). The [35 S]-labeling may be performed with whole animals, tissue explants, tissue slices, and cells in culture. This unit outlines procedures to determine whether a protein of interest contains a tyrosine-sulfated residue, using mammalian adherent cells.
STRATEGIC PLANNING Prediction of Tyrosine Sulfation Sites The first step to investigate whether the protein of interest contains a tyrosine-sulfated residue is to determine the presence of a potential tyrosine sulfation site. In order to be a substrate for TPST, a tyrosine residue in the protein must meet the following three criteria. 1. The tyrosine residue must be situated either on a secretory protein or on the lumenal domain of a transmembrane protein. This is a consequence of the fact that the catalytic site of TPST is oriented toward the TGN lumen. Practically, this means that a tyrosine can only be sulfated if the protein contains an N-terminal signal sequence or at least one transmembrane domain. For prediction of tyrosine sulfation sites in transmembrane proteins, the topology of the membrane insertion must be considered (UNIT 2.2). 2. The tyrosine residue must be situated in an acidic environment. Although no strict consensus sequence for tyrosine sulfation sites exists, all sequences exhibit the
Figure 14.7.1 The formation of a tyrosine-O4 -sulfate residue is catalyzed by tyrosylprotein sulfotransferase (TPST) using 3 -phosphoadenosine 5 -phosphosulfate (PAPS) as co-substrate and sulfate donor. This enzymatic reaction occurs in the lumen of the trans-Golgi network (TGN).
Contributed by Denis Corbeil, Christoph Thiele, and Wieland B. Huttner Current Protocols in Protein Science (2005) 14.7.1-14.7.14 C 2005 by John Wiley & Sons, Inc. Copyright
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presence of acidic amino acid residues in the vicinity, i.e., positions −5 (N-terminal) to +5 (C-terminal), of the sulfated tyrosine residue. Turn-inducing amino acids (P, G) are also frequently present. The few examples of tyrosine sulfation sites lacking proline and glycine are located near the N- or C-terminus of the protein and/or contain several of the three other amino acid residues (D, S, N) that have significant turn-conformational potential. 3. The tyrosine residue must be sterically accessible to TPST. This usually requires the absence of cysteine residues or potential N-glycosylation sites (NXS or NXT) in the vicinity (positions −7 to +7), which could give rise to disulfide bridges or bulky carbohydrate structures, respectively. The prediction of potential tyrosine sulfation sites can be supported by bioinformatics. The most recent software is called Sulfinator, developed by the SWISS-PROT group (Monigatti et al., 2002) and available at the ExPASy server (http://www.expasy.org/ tools/sulfinator/). Sulfinator compares the input sequence with a consensus pattern derived from a multiple sequence alignment of 68 stretches with known tyrosine-sulfated residues. Note that Sulfinator restricts itself to the above criterion 2, while criteria 1 and 3 are not covered. BASIC PROTOCOL 1
LONG-TERM [35 S]-SULFATE LABELING OF MAMMALIAN CELLS IN CULTURE AND IMMUNOPRECIPITATION To investigate whether a protein of interest contains a sulfated tyrosine residue, long-term [35 S]-sulfate labeling is used. The radiolabeled protein can be recovered by immunoprecipitation either from the cell lysate or, in the case of a secreted protein, the culture medium, and further analyzed by SDS-PAGE and autoradiography or fluorography (see Support Protocol 1). This assay is appropriate for a protein of interest endogenously expressed in a given cell line or upon its heterologous expression in host cells. If antibodies capable of precipitating the protein of interest are not available, an alternative approach is epitope tagging, whereby the protein of interest is engineered to carry an antigenic determinant to which a commercial antibody is available. It is important that the suitability of the antibody for immunoprecipitation is thoroughly established, and the target epitope does not contain a potential tyrosine sulfation site.
Materials
Tyrosine O-Sulfation
Mammalian adherent cells expressing protein of interest and appropriate control cells lacking the protein of interest or expressing the protein of interest containing a point mutation in the tyrosine sulfation site Complete tissue culture medium appropriate for cell line Labeling medium with and without label (see recipe), prewarmed PBS (APPENDIX 2E), cold Immunoprecipitation buffer (see recipe) with protease inhibitors, ice cold RIPA lysis buffer (see recipe) with protease inhibitors, ice cold Antibody Protein A-agarose beads Wash buffers A, B, and C (see recipes) 2× SDS-PAGE sample buffer (see recipe) Disposable pipets Centricon concentrator (Amicon) 2.0- and 1.5-ml microcentrifuge tubes Cell scraper
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15-ml centrifuge tubes End-over-end rotator Additional reagents and equipment for immunoprecipitation (UNIT 13.2) and SDS-PAGE (UNIT 10.1) NOTE: All reagents and equipment coming into contact with live cells must be sterile, and proper sterile technique should be used accordingly. NOTE: All culture incubations are performed in a 37◦ C, 5% CO2 humidified incubator unless specified otherwise. Some media (e.g., DMEM) may require altered levels of CO2 to maintain the pH at 7.4. NOTE: The size of the cultures to be labeled depends on the amount of the protein of interest present in the cells. Typically, 100- to 500-mm diameter culture dishes are required. CAUTION: Radioactive materials require special handling. See APPENDIX 2B concerning safe use of radioisotopes.
Prepare cells 1. Culture the mammalian adherent cells expressing the protein of interest and appropriate control cells to an appropriate stage of growth in complete tissue culture medium appropriate for the cell line (see Critical Parameters and Troubleshooting). For maximal 35 SO4 incorporation, cells should be subconfluent (60% to 80%). It is recommended to replace the medium with fresh growth medium 18 hr prior to labeling. Protein expression is often achieved by transient transfection with an appropriate expression vector. Several transfection methods are described in Current Protocols in Molecular Biology, UNITS 9.1-9.4. Control experiments with cells lacking the protein of interest or expressing the protein of interest containing a point mutation in the tyrosine sulfation site should be conducted in parallel.
2. Remove complete tissue culture medium from the cultured cells by aspiration. Wash away any residual serum-containing medium by adding prewarmed labeling medium without label (10 ml per 100-mm dish), and aspirate. It is important to hold the dishes at an angle and remove and add medium at one corner so as not to dislodge the cells from the plates.
3. Add 5 ml prewarmed labeling medium per 100-mm culture dish and incubate 7 to 16 hr. At the end of the labeling period, the medium can be discarded if the protein of interest is associated with cells. Preliminary experiments should be conducted to determine if the protein of interest is released into the medium.
For proteins released into the medium: 4a. Remove the labeling medium manually from the dish using a disposable pipet, centrifuge 5 min at 300 × g, 4◦ C, and carefully collect the supernatant. Discard the cell pellet and pipet as radioactive waste. 5a. Concentrate the supernatant in a Centricon unit to 500 µl by centrifugation (explicitly follow manufacturer’s instructions). Carefully choose the cut-off value of the Centricon to ensure retention of the protein of interest.
6a. Dilute to 1.5 ml with cold PBS and repeat the concentration and dilution steps 5a and 6a.
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7a. Concentrate to 200 µl as in step 5a and collect the concentrated protein solution. 8a. Dilute to 1.8 ml with ice-cold immunoprecipitation buffer containing protease inhibitors in a 2-ml microcentrifuge tube. Proceed to step 9. For solutions requiring protease inhibitors, add protease inhibitor cocktail tablets (Roche Molecular Biochemicals) following the manufacturer’s instructions.
For proteins associated with cells: 4b. Wash cells two times with 5 ml ice-cold PBS per 100-mm dish, and remove wash fluid by aspiration. CAUTION: The wash fluid is radioactive—follow applicable safety regulations for disposal.
5b. Add 5 ml ice-cold PBS per 100-mm dish and scrape cells carefully with a disposable plastic cell scraper. 6b. Transfer the suspension to a 15-ml centrifuge tube, centrifuge 5 min at 300 × g, 4◦ C, and discard supernatant. 7b. Add ice-cold RIPA lysis buffer containing protease inhibitors to cell pellet, using 0.3 to 0.5 ml per 100-mm dish, resuspend the pellet by gentle agitation, and transfer to a 1.5-ml microcentrifuge tube. Incubate 30 min at 4◦ C. 8b. Clarify the detergent lysate by centrifuging 10 min at 10,000 × g, 4◦ C, in a refrigerated microcentrifuge. Transfer the supernatant (total lysate) to a 2.0-ml microcentrifuge tube and dilute to 1.8 ml with ice-cold immunoprecipitation buffer containing protease inhibitors. Discard the 1.5-ml microcentrifuge tube and pellet as radioactive waste. It is worthwhile analyzing an aliquot of the clarified total lysate (step 8b) and/or the concentrated medium (step 7a) by SDS-PAGE (UNIT 10.1). The protein of interest may be highly sulfated and readily detectable (see Support Protocol 1) without prior enrichment, even if it is a relatively minor fraction of the total protein.
Immunoprecipitate 9. Carry out immunoprecipitation (UNIT 9.8), using the appropriate amount of antibody and incubate 16 hr at 4◦ C using an end-over-end rotator. 10. After the 16-hr incubation, add an appropriate amount of protein A-agarose beads to quantitatively precipitate the antibody-antigen complexes, and rotate end-over-end for 2 hr at 4◦ C. Substitute with protein G beads when using antibodies that are not bound effectively by protein A.
11. Collect immunoprecipitate by microcentrifuging for 10 sec at maximum speed and wash the beads at 4◦ C successively with the following: a. b. c. d.
one time with 1 ml immunoprecipitation buffer; three times with 1 ml wash buffer A; two times with 1 ml wash buffer B; one time with 1 ml wash buffer C.
12. Completely remove wash buffer from the beads with a micropipettor, add 1 vol of 2× SDS-PAGE sample buffer, and incubate 3 min in boiling water. Tyrosine O-Sulfation
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Analyze protein of interest 13. Analyze the protein of interest, potentially containing radiolabeled tyrosine sulfate, by SDS-PAGE (UNIT 10.1). After electrophoresis, detection of [35 S]-sulfate incorporation into the protein of interest is carried out as described in Support Protocol 1.
DETECTION OF [35 S]-SULFATED PROTEINS [35 S]-sulfated
To detect proteins after SDS-PAGE, several methods are available. The fixed gel, after being dried onto filter paper, can be exposed using a phosphorimager. Alternatively, the fixed, wet gel can be treated with sodium salicylate (UNIT 14.3) and fluorographed (UNIT 10.2). A rapid procedure to obtain an autoradiogram is to transfer labeled proteins from the gel to a nitrocellulose membrane using the blotting technique (UNIT 10.7), and to expose the nitrocellulose membrane to film.
SUPPORT PROTOCOL 1
Additional Materials (also see Basic Protocol 1) Gel containing protein of interest (see Basic Protocol 1) 1 M sodium salicylate, pH 5 to 7 20% PPO (2,5-diphenyloxazole) in toluene (optional) Whatman 3MM filter paper or nitrocellulose membrane X-ray film (e.g., Kodak XAR-5 or equivalent) or phosphorimager Additional reagents and equipment for staining and destaining gels (UNIT 10.5), fluorography (UNIT 10.2), and blotting (UNIT 10.7) CAUTION: Solutions that come into contact with the gel may become radioactive. Gloves should be worn at all times. Sodium salicylate can cause allergic reactions and is readily absorbed through the skin.
To detect [35 S]-sulfated proteins by phosphorimaging: 1a. After electrophoresis, fix the gel and (if necessary) stain and destain by standard methods (UNIT 10.5). Because the tyrosine sulfate ester is acid-labile, the use of acetic acid rather than trichloroacetic acid is recommended.
2a. Dry the fixed gel onto Whatman 3MM filter paper and then detect labeled proteins directly using a phosphorimager. Time of exposure will depend on the amount of radioactivity in the sample and, in most cases, will have to be determined empirically by making multiple exposures for different lengths of time.
To detect [35 S]-sulfated proteins by fluorography: 1b. After electrophoresis, fix the gel and (if necessary) stain and destain by standard methods (UNIT 10.5). Because the tyrosine sulfate ester is acid-labile, the use of acetic acid rather than trichloroacetic acid is recommended.
2b. Soak the fixed gel for 1 to 5 hr in 20 vol water to remove residual acid, which could cause precipitation of salicylic acid from sodium salicylate in step 3b.
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3b. Soak gel for 30 min in 10 vol of 1 M sodium salicylate, pH 5 to 7. Dry the gel and proceed with fluorography (UNIT 10.2) at −70◦ C. The sodium salicylate method is recommended for fluorography because salicylate is water-soluble and can, therefore, easily be removed from the gel after fluorography. This renders the proteins in the gel suitable for further biochemical analysis such as tyrosine sulfate analysis (see Basic Protocol 2). If the labeling of proteins is intense, salicylate treatment of the gel can be omitted and the dried gel can be subjected to autoradiography. To prevent cracking of gels that are >10% polyacrylamide or >1.5-mm thick, 2% (v/v) glycerol can be added to the sodium salicylate solution.
To detect [35 S]-sulfated proteins by membrane autoradiography: 1c. After electrophoresis, transfer labeled proteins from the gel to a nitrocellulose membrane using the blotting technique (UNIT 10.7). 2c. Air-dry the nitrocellulose membrane and autoradiograph, or dip the nitrocellulose membrane in 20% PPO (2,5-diphenyloxazole) in toluene, dry, and fluorograph at −70◦ C. This is an alternative, rapid procedure to obtain an autoradiogram after SDS-PAGE given that the time needed for autoradiography after transfer to nitrocellulose membrane is shorter than with dried polyacrylamide gels. BASIC PROTOCOL 2
TYROSINE SULFATE ANALYSIS—ALKALINE HYDROLYSIS METHOD Since incorporation of 35 SO4 into proteins may occur not only on tyrosine but also on carbohydrate residues, it is important to obtain positive evidence that the [35 S]-sulfated protein of interest indeed contains [35 S]-sulfated tyrosine. This protocol presents a simple chemical method based on exhaustive alkaline hydrolysis followed by thin-layer electrophoresis (see Support Protocol 2) to demonstrate the presence of radiolabeled tyrosine residues. The immunoprecipitate containing the protein of interest (see Basic Protocol 1) can be used as starting material.
Materials Immunoprecipitate containing the protein of interest (see Basic Protocol 1) Wash buffer D (see recipe) Pronase solution: 100 µg/ml Pronase (Roche) in 5 mM NH4 HCO3 , pH 8 0.2 and 0.02 M barium hydroxide (Ba(OH)2 ) N2 source 1 and 0.1 M sulfuric acid Uncapped 3-ml glass centrifuge tube 110◦ C oven CAUTION: Barium hydroxide may be fatal if swallowed, harmful if inhaled, and causes irritation to skin, eyes, and respiratory tract. It affects muscles and the central nervous system. Use gloves when handling and perform this experiment in a chemical fume hood. 1. Elute the [35 S]-sulfated protein of interest from antibody-protein A–agarose complexes (see Basic Protocol 1, step 11, with an additional final wash with 1 ml wash buffer D) by adding 200 µl Pronase solution. Incubate 12 to 24 hr at 37◦ C and microcentrifuge 10 sec at maximum speed. The Pronase solution should be preincubated according to the manufacturer’s instructions. Tyrosine O-Sulfation
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2. Transfer eluate into a disposable 3-ml glass centrifuge tube. Lyophilize eluate. The disposable glass tubes must tolerate the subsequent steps, i.e., centrifugation at 10,000 × g, and pH 14 at 110◦ C; they must also be suitable for gastight sealing. Uncapped 3-ml glass centrifuge tubes work well.
3. Prepare a fresh solution of 0.2 M barium hydroxide. Add 1 ml of 0.2 M Ba(OH)2 to lyophilized eluate and seal tube under N2 so that it is gas-tight. Use degassed water to reduce formation of BaCO3 . Usually, the barium hydroxide does not dissolve completely. While it is being stirred, take 1 ml of the barium hydroxide solution and add it to the residue of the lyophilized eluate. For uncapped glass tubes, freeze sample, evacuate, and seal by melting in flame.
4. Place the sealed tube in a 110◦ C oven for 20 to 24 hr. 5. Cool the sample to 4◦ C and break the seal manually by wrapping tube in a paper towel. Centrifuge for 10 min at 10,000 × g, 4◦ C, collect the supernatant and transfer it to another centrifuge tube. 6. (Optional) Add 0.4 ml water to pellet, vortex, centrifuge for 10 min at 10,000 × g, 4◦ C. Pool the second supernatant with first one. This step can be omitted if only qualitative results are required.
7. Discard the tube with the pellet as radioactive waste. 8. Neutralize the (pooled) supernatant by adding sulfuric acid, using the phenol red in the sample as indicator. Centrifuge sample for 10 min at 10,000 × g, 4◦ C. Carefully add 1 M sulfuric acid with frequent vortexing until the phenol red turns transiently yellow at the site of sulfuric acid addition. Then continue the neutralization with 0.1 M sulfuric acid until the sample is homogenously red-orange (pH ∼7). If excess sulfuric acid is added accidentally (yellow color), back-titrate with 0.02 M barium hydroxide.
9. Collect the supernatant, and transfer to a 2-ml microcentrifuge tube. 10. (Optional) Add 0.4 ml water to pellet, vortex, and centrifuge as in step 8. Pool second supernatant with first one. Discard the tube with the pellet as radioactive waste. The pooled supernatant is referred to as neutralized supernatant. This step can be omitted if only qualitative results are required.
11. Lyophilize the neutralized supernatant. The residue of the neutralized supernatant is used for the identification of tyrosine [35 S]sulfate by thin-layer electrophoresis (see Support Protocol 2).
THIN-LAYER ELECTROPHORESIS OF TYROSINE [35 S]-SULFATE Thin-layer electrophoresis of tyrosine sulfate is performed under the same conditions as that of tyrosine phosphate (UNIT 13.3). Given that the alkaline hydrolysis and neutralization conditions used to liberate tyrosine sulfate from proteins (see Basic Protocol 2) render tyrosine sulfate the major, if not the only, [35 S]-labeled product in the neutralized supernatant, 1-D thin-layer electrophoresis is sufficient in most instances to demonstrate the presence of tyrosine sulfate. Because electrophoresis equipment differs considerably in design, the details of the assembly and placement of the plate are not discussed here (see UNIT 13.3). It is assumed that a suitable apparatus is available for use by an experienced operator. For fluorography, EN3 HANCE (PerkinElmer Life Sciences) can be used.
SUPPORT PROTOCOL 2
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Materials Lyophilized sample (see Basic Protocol 2) Electrophoresis buffer, pH 3.5 (see recipe) Acetone Unlabeled L-tyrosine-O-sulfate (Bachem) Phenol red Unlabeled L-serine-O-sulfate (Bachem) Unlabeled L-tyrosine (Bachem) 0.2% (w/v) ninhydrin in acetone (as spray) Cellulose-coated plastic-backed TL chromatography sheet (20 cm × 20 cm × 100 µm) Whatman 3 MM filter paper 100◦ C oven Additional reagents and equipment for thin-layer electrophoresis (UNIT 13.3) 1. Dissolve the sample (residue after lyophilization of the neutralized supernatant, see Basic Protocol 2, step 11) in 20 µl electrophoresis buffer, pH 3.5. If the residue is small and dissolves readily, proceed to step 4 as the sample will run well in the thin-layer electrophoresis. However, if the residue appears to be too big to be loaded on a thin-layer sheet or does not dissolve readily, perform an acetone precipitation (steps 2 to 3). 2. Add 100 µl acetone to a 20-µl sample. 3. Sediment the precipitate by microcentrifuging 5 min. Collect the supernatant and evaporate the acetone. Repeat step 1. 4. Add 3 µg of unlabeled L-tyrosine-O-sulfate, and a trace of phenol red (100 to 300 ng) to the sample. Usually, the phenol red is already present from the alkaline hydrolysis.
5. Spot 25% to 100% of the sample, in 0.50-µl aliquots, on a 20-cm × 20-cm × 100-µm cellulose thin-layer chromatography sheet, 3 cm from the cathodic edge of the plate and adjacent to unlabeled L-tyrosine-O-sulfate, L-tyrosine, and L-serineO-sulfate standards (3 µg each, in electrophoresis buffer, pH 3.5; see Fig. 14.7.2 for arrangement of samples). Between each application, dry the sample spot with compressed air delivered through a Pasteur pipet plugged with cotton. Use long, thin plastic micropipet loading tips for sample application, and do not let the tip touch the plate. The entire hydrolysate can be spotted onto a single origin; however, streaking may be observed upon overloading.
6. Wet the cellulose sheet with electrophoresis buffer, pH 3.5. This can be done either by spraying (if the spot is small) or by using a “mask’ of Whatman 3 MM filter paper (if the spot is to be concentrated). Briefly, take a piece of the filter paper that is slightly larger than the cellulose sheet, cut circular holes slightly larger than the sample spots at the position corresponding to that of the sample and standards on the cellulose sheet. Wet the filter paper with electrophoresis buffer, pH 3.5 and place it on the cellulose sheet such that sample spots are not touching the filter paper. Allow the electrophoresis buffer to concentrically migrate toward the center of the spots, thereby concentrating the samples. The cellulose sheet should be homogeneously wet. Tyrosine O-Sulfation
7. Place wet cellulose sheet in the electrophoresis apparatus (follow manufacturer’s instructions).
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Figure 14.7.2 Thin-layer electrophoresis of tyrosine sulfate. Schematic diagram of a 1-D separation, showing the positions of sulfated and nonsulfated hydroxyamino acids. The sample (S) containing [35 S]-labeled tyrosine sulfate, and standards (tyrosine, T; tyrosine sulfate, TS; serine sulfate, SS) are analyzed by ninhydrin staining (A) and autoradiography/fluorography (B). The position of phenol red is indicated.
8. Electrophorese the samples in electrophoresis buffer, pH 3.5, until the phenol red marker has migrated 7 to 8 cm, i.e., just past the middle of the cellulose plate. Electrophoresis conditions will depend on the thin-layer electrophoresis apparatus used. Consequently, the appropriate duration and voltage of the electrophoresis need to be determined empirically by examining the rate of migration of unlabeled standards.
9. Remove the plate and quickly air-dry with a fan without heating. It takes ∼20 min to dry the plate.
10. Spray the dry plate with 0.2% ninhydrin in acetone. Develop the color in a 100◦ C oven for a few minutes to visualize the sulfated amino acid standards. 11. Perform autoradiography or fluorography of the plate to detect radioactive tyrosine sulfate.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Dialyzed serum Dialyze serum overnight against PBS (APPENDIX 2E) at 4◦ C. Use 100 times the serum volume for the dialysis, and change the solution once during dialysis. Sterile-filter after dialysis and store in 1- or 2-ml aliquots in sterile tubes for up to 1 year at −20◦ C. After thawing, store dialyzed serum for up to 1 month at 4◦ C. Electrophoresis buffer, pH 3.5 5% (v/v) acetic acid 0.5% (v/v) pyridine Store up to 1 year in a sealed bottle at room temperature
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Immunoprecipitation buffer 1% Triton X-100 0.1% SDS 150 mM NaCl 10 mM EDTA 50 mM Tris · Cl, pH 7.8 (APPENDIX 2E) Store up to 1 year at 4◦ C Labeling medium Sulfate-free tissue culture medium (see recipe) containing: MgCl2 instead of MgSO4 10% of the normal concentration of methionine and cysteine 1% dialyzed serum (see recipe) Carrier-free Na2 35 SO4 (1.0 mCi/ml; >900 Ci/mmol) (Amersham Biosciences) Prepare fresh See Critical Parameters for more information.
Low-Met/Cys amino acid mixture, 10x For 1 liter: 0.84 g L-arginine · HCl 0.063 g L-cystine 0.30 g glycine 0.42 g L-histidine · HCl 1.05 g L-isoleucine 1.05 g L-leucine 1.46 g L-lysine · HCl 0.03 g L-methionine 0.66 g L-phenylalanine 0.42 g L-serine 0.95 g L-threonine 0.16 g L-tryptophan 0.94 g L-valine Dissolve in sterile Milli-Q-purified water and adjust volume to 1 liter Store up to 6 months at −20◦ C Having low methionine and cysteine improves the [35 S]-sulfate incorporation by reducing the amount of unlabeled sulfate that may arise from the metabolism of sulfur-containing amino acids and that would reduce the specific activity of the radioactive sulfate. Methionine and cysteine at 10% the normal concentration is usually sufficient to maintain protein synthesis and keep the cells healthy. Tyrosine is omitted from the above 10× amino acid mixture and included directly in the sulfate-free tissue culture medium (see recipe) because tyrosine is insoluble at 10× the normal concentration. Do not add antibiotics, as some (e.g., streptomycin) are sulfate salts.
Tyrosine O-Sulfation
RIPA lysis buffer 0.15 M NaCl 1% (v/v) Nonidet P-40 (NP-40) 0.5% (w/v) sodium deoxycholate 0.1% (w/v) SDS 0.05 M Tris · Cl, pH 8.0 Store up to 1 year at 4◦ C
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SDS-PAGE sample buffer, 2× 20% (v/v) glycerol 4% (w/v) SDS 0.125 M Tris · Cl, pH 6.8 (APPENDIX 2E) 0.2 M DTT, added fresh 0.01% bromophenol blue Store up to 6 months at −20◦ C Sulfate-free tissue culture medium For 1 liter: 265 mg CaCl2 · 2H2 O 0.10 mg Fe(NO3 )3 · 9H2 O 400 mg KCl 168.75 mg MgCl2 · 6H2 O 6.4 g NaCl 125 mg NaH2 PO4 · H2 O 5 ml 20% (w/v) glucose 10 ml 11.0 mg/ml sodium pyruvate (added dropwise, see below) 49.3 ml 7.5% (w/v) NaHCO3 72.0 mg tyrosine 40 ml 100× MEM vitamins (Sigma) 100 ml 10× low-Met/Cys amino acid mixture (see recipe) 3 ml 0.5% (w/v) phenol red Mix the above ingredients (except phenol red) with 900 ml Milli-Q-purified water and stir for several hours at room temperature or overnight at 4◦ C to dissolve. Add phenol red and adjust pH to 7.2 with 1 M NaOH. Adjust volume to 1 liter with water. Filter sterilize solution through a 0.2-µm filter. Store up to 6 months at 4◦ C in the dark. Just before use, add L-glutamine (4 mM final). NOTE: Sodium pyruvate is added drop-wise to avoid precipitation. Wash buffer A 0.2% NP-40 150 mM NaCl 2 mM EDTA 10 mM Tris · Cl, pH 7.5 (APPENDIX 2E) Store up to 1 year at 4◦ C Wash buffer B 0.2% NP-40 500 mM NaCl 2 mM EDTA 10 mM Tris · Cl, pH 7.5 (APPENDIX 2E) Store up to 1 year at 4◦ C Wash buffer C 10 mM Tris · Cl, pH 7.5 (APPENDIX 2E) Store up to 1 year at 4◦ C Wash buffer D 5 mM NH4 HCO3 , pH 8 Store up to 1 year at 4◦ C
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COMMENTARY Background Information
Tyrosine O-Sulfation
Sulfation of tyrosine residues is a late post-translational modification that takes place in the trans-Golgi network and occurs in a wide spectrum of transmembrane and secretory proteins (Huttner, 1982). Tyrosinesulfated proteins are found in all animals, and also exist in the plant kingdom. While occurring throughout metazoan evolution, tyrosine-sulfated proteins appear to be absent in unicellular eukaryotes and prokaryotes, implicating this post-translational modification in some aspect of multicellularity (Huttner and Baeuerle, 1988; Moore, 2003; Corbeil and Huttner, 2004). For most tyrosine-sulfated proteins, the physiological role of this post-translational modification is presently unknown. A review with a comprehensive list of tyrosine-sulfated proteins has recently been published (Moore, 2003). With regard to the cases in which the physiological relevance of tyrosine sulfation of a particular protein has been elucidated, the common denominator has emerged that tyrosine sulfation promotes extracellular protein-protein interactions occurring in various biological processes, ranging from the receptor binding of regulatory peptides to the interaction of viral envelope proteins with the cell surface (Huttner et al., 1991; Farzan et al., 1999; Kehoe and Bertozzi, 2000; Costagliola et al., 2002). Moreover, tyrosine sulfation has been reported to modulate the secretion kinetics of certain secretory proteins (Friederich et al., 1988) and to promote proteolytic processing (Bundgaard et al., 1995). Thus, the demonstration that a given protein undergoes tyrosine sulfation and the identification of the sulfated tyrosine residue are important aspects of the characterization of its physiological activity. The sulfate linked to tyrosine is present as an O4 -sulfate ester (Fig. 14.7.1). Such a modified tyrosine residue was first identified in 1954 in bovine fibrinopeptide B (Bettelheim, 1954). Remarkably, up to 1% of all tyrosine residues of the total protein in an organism can be sulfated, making this the most common post-translational modification of this particular residue (Huttner and Baeuerle, 1988). The sulfate transfer reaction to tyrosine residue is catalyzed by one of two trans-Golgi-associated tyrosylprotein sulfotransferases (TPSTs; EC 2.8.2.20) (Niehrs et al., 1994; Ouyang et al., 1998a,b; Beisswanger et al., 1998), which use the universal sulfate donor 3 -phosphoadenosine
5 -phosphosulfate (PAPS) as a co-substrate (Robbins and Lippmann, 1956) (Fig. 14.7.1). Biologically, tyrosine sulfation appears to be an irreversible modification in vivo due to the lack of a sulfatase capable of catalyzing the desulfation of tyrosine-sulfated proteins under physiological conditions. Extending previous attempts to detect serine sulfate and threonine sulfate in protein hydrolysates (Huttner, 1984; Huttner and Baeuerle, 1988), it has recently been demonstrated that, other than tyrosine residues, serine and threonine residues can be subject to sulfation (Medzihradszky et al., 2004). Analysis of tyrosine sulfate The method described in this unit is based on alkaline hydrolysis. Because the tyrosine sulfate ester is alkali-stable, but acid-labile, alkaline hydrolysis is the method of choice to release tyrosine sulfate from a polypeptide chain. For such hydrolysis, barium hydroxide is used because the inorganic radioactive sulfate, generated by hydrolysis of alkali-labile–sulfated carbohydrate residues of glycoproteins, is precipitated as barium sulfate, whereas tyrosine sulfate remains in solution (Huttner, 1984). The alkaline hydrolysis of [35 S]-labeled protein with barium hydroxide is not restricted to proteins that have been isolated by immunoprecipitation, but is also applicable to proteins contained in polyacrylamide gels, and to purified proteins (for details, see Huttner, 1984). Several alternative analytical methods for studying tyrosine sulfation that are independent of radioisotope incorporation have been developed (Bundgaard et al., 2002). The sulfation of tyrosine residues in a protein of interest can be demonstrated using various chromatographic techniques, including reversed-phase high-performance liquid chromatography (UNIT 8.7). These methods are based on the chemical properties of the sulfate moiety, which introduces a highly acidic group into the protein of interest that alters its physical and chemical properties such as conformation and hydrophilicity. However, these analytical techniques require a pure protein or the use of specific tyrosine sulfate antisera, which are very difficult to generate. Nevertheless, an immunization protocol describing the generation of such antibody has been reported (Bundgaard et al., 2002). Tyrosine sulfate analysis can also be achieved using various techniques of mass spectrometry (MS), including matrix-assisted laser desorption/
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ionization time-of-flight (MALDI-TOF) MS (UNIT 16.5) and electrospray ionization (ESI) ¨ et al., 2004). MS (UNIT 16.1) (Onnerfjord Although these techniques are very sensitive and precise, they require sophisticated and expensive instrumentation. Moreover, it is important to note that the tyrosine sulfate ester bond, which is more labile than peptide bonds, might be broken during the mass spectrometry acquisition, resulting in false-negative data (Wolfender et al., 1999).
Critical Parameters and Troubleshooting A critical factor in the [35 S]-sulfate labeling of mammalian cells is the optimization of sulfate incorporation into the protein to be analyzed. The efficiency of sulfation is the most variable parameter and relies on maintaining the cells in a medium that is compatible with continued protein synthesis and with sulfate labeling conditions. The labeling medium that the authors use is a modified DMEM. Other media, such as RPMI, do not allow efficient sulfate incorporation (for additional information, see Current Protocols in Cell Biology, UNIT 7.3). The advantage of using the isotope at high specific activity should be balanced against the potential disadvantage that may result from the use of dialyzed serum. It is therefore recommended to ascertain that the use of sulfate-free medium and dialyzed serum during long-term [35 S]-sulfate labeling does not reduce the viability of the cells under investigation. To facilitate the detection of [35 S]-labeled protein, it is important that most, if not all, cells used express the protein of interest. If the efficiency of the cell transfection is low, establishing a permanent cell line expressing the protein of interest is strongly recommended. NIH 3T3 cells (Niehrs et al., 1992), CHO cells (Weigmann et al., 1997), and PC12 cells (Glombik et al., 1999) are excellent host cells. To determine if the protein of interest is sulfated on a tyrosine residue, control experiments utilizing a mutant form of the protein should be performed in parallel to substantiate any positive results. A phenylalanine residue can substitute the presumably sulfated tyrosine residue (Friederich et al., 1988). Negative controls with cells lacking the protein of interest should also be performed. To exclude the possibility that the incorporation of [35 S]-sulfate occurs on N- and Olinked carbohydrates, the protein of interest recovered after immunoprecipitation can be treated with N-glycosidase F, neuraminidase,
and/or O-glycosidase, prior to SDS-PAGE and detection of [35 S]-sulfated protein (Farzan et al., 1999). Likewise, it can be informative to perform [35 S]-labeling in the absence or presence of tunicamycin, an inhibitor of Nglycosylation. If, upon such treatments, the protein of interest displays a similar pattern of [35 S]-labeling, i.e., without any loss of signal intensity, it can be concluded that the protein contains sulfated tyrosine residues. Final proof that [35 S]-sulfate is indeed incorporated into tyrosine residues is provided by thin-layer electrophoresis of a Ba(OH)2 hydrolysate of the immunoprecipitated protein of interest.
Anticipated Results Typically, on thin-layer electrophoresis at pH 3.5, tyrosine sulfate migrates ∼1.5 times as fast toward the anode as phenol red. The non-sulfated tyrosine residue remains near the origin, whereas serine sulfate migrates ∼1.5 times as fast as tyrosine sulfate (see Fig. 14.7.2).
Time Considerations After the cells have grown to the desired density, the [35 S]-labeling experiment can be completed within 1 day, unless an overnight labeling is needed. Harvesting the cells, preparing the cell lysate and/or concentrating the culture medium as well as the immunoprecipitation require at least 2 days. An additional day is needed for SDS-PAGE and fixing the gel. If transfer of labeled proteins from the gel to nitrocellulose membrane is to be performed, several additional hours are required depending upon the blotting technique used. Exposure time to X-ray film or phosphorimaging must be determined on an experiment-to-experiment basis. Tyrosine sulfate analysis (alkaline hydrolysis, thin-layer electrophoresis) requires ∼1 day, and exposing the cellulose thin-layer sheet containing tyrosine [35 S]-sulfate another 2 to 5 days.
Literature Cited Beisswanger, R., Corbeil, D., Vannier, C., Thiele, C., Dohrmann, U., Kellner, R., Ashman, K., Niehrs, C., and Huttner, W.B. 1998. Existence of distinct tyrosylprotein sulfotransferase genes: Molecular characterization of tyrosylprotein sulfotransferase-2. Proc. Natl. Acad. Sci. U.S.A. 95:11134-11139. Bettelheim, F.R. 1954. Tyrosine-O-sulfate in a peptide from fibrinogen. J. Am. Chem. Soc. 76:28382839. Bundgaard, J.R., Johnsen, A.H., and Rehfeld, J.F. 2002. Analysis of tyrosine-O-sulfation. Methods Mol Biol. 194:223-239.
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Corbeil, D. and Huttner, W.B. 2004. Tyrosine sulfation. Encyclopedia Biol. Chem. 4:294-297. ( W. Lennarz and M.D. Lane, eds.) Elsevier, Oxford. Costagliola, S., Panneels, V., Bonomi, M., Koch, J., Many, M.C., Smits, G., and Vassart, G. 2002. Tyrosine sulfation is required for agonist recognition by glycoprotein hormone receptors. EMBO J. 21:504-513. Farzan, M., Mirzabekov, T., Kolchinsky, P., Wyatt, R., Cayabyab, M., Gerard, N.P., Gerard, C., Sodroski, J., and Choe, H. 1999. Tyrosine sulfation of the amino terminus of CCR5 facilitates HIV-1 entry. Cell 96:667-676. Friederich, E., Fritz, H.J., and Huttner, W.B. 1988. Inhibition of tyrosine sulfation in the trans-Golgi retards the transport of a constitutively secreted protein to the cell surface. J. Cell Biol. 107:16551667. Glombik, M.M., Kr¨omer, A., Salm, T., Huttner, W.B., and Gerdes, H.H. 1999. The disulfidebonded loop of chromogranin B mediates membrane binding and directs sorting from the transGolgi network to secretory granules. EMBO J. 18:1059-1070. Huttner, W.B. 1982. Sulphation of tyrosine residues—A widespread modification of proteins. Nature (London) 299:273-276. Huttner, W.B. 1984. Determination and occurrence of tyrosine O-sulfate in proteins. Methods Enzymol. 107:200-223. Huttner, W.B. and Baeuerle, P.A. 1988. Protein sulfation on tyrosine. Mod. Cell Biol. 6:97-140. Huttner, W.B., Niehrs, C., and Vannier, C. 1991. Bind or bleed. Curr. Biol. 1:309-310. Kehoe, J.W. and Bertozzi, C.R. 2000. Tyrosine sulfation: A modulator of extracellular proteinprotein interactions. Chem. Biol. 7:R57-R61. Medzihradszky, K.F., Darula, Z., Perlson, E., Fainzilber, M., Chalkley, R.J., Ball, H., Greenbaum, D., Bogyo, M., Tyson, D.R., Bradshaw, R.A., and Burlingame, A.L. 2004. OSulfonation of serine and threonine: Mass spectrometric detection and characterization of a new posttranslational modification in diverse proteins throughout the eukaryotes. Mol Cell Proteomics. 3:429-443. Monigatti, F., Gasteiger, E., Bairoch, A., and Jung, E. 2002. The Sulfinator: Predicting tyrosine sulfation sites in protein sequences. Bioinformatics 18:769-770. Moore, K.L. 2003. The biology and enzymology of protein tyrosine O-sulfation. J. Biol. Chem. 278:24243-24246. Niehrs, C., Huttner, W.B., and R¨uther, U. 1992. In vivo expression and stoichiometric sulfation of the artificial protein sulfophilin, a polymer of tyrosine sulfation sites. J. Biol. Chem. 267:1593815942.
Niehrs, C., Beiβwanger, R., and Huttner, W.B. 1994. Protein tyrosine sulfation, 1993—An update. Chem.-Biol. Interact. 92:257-271. ¨ Onnerfjord, P., Heathfield, T.F., and Heinegard, D. 2004. Identification of tyrosine sulfation in extracellular leucine-rich repeat proteins using mass spectrometry. J. Biol. Chem. 279:26-33. Ouyang, Y-B., Lane, W.S., and Moore, K.L. 1998a. Tyrosylprotein sulfotransferase: Purification and molecular cloning of an enzyme that catalyzes tyrosine O-sulfation, a common posttranslational modification of eukaryotic proteins. Proc. Natl. Acad. Sci. U.S.A. 95:28962901. Ouyang, Y-B. and Moore, K.L. 1998b. Molecular cloning and expression of human and mouse tyrosylprotein sulfotransferase-2 and a tyrosylprotein sulfotransferase homologue in Caenorhabditis elegans. J. Biol. Chem. 277:24770-24774. Robbins, P. and Lippmann, F. 1956. Identification of enzymatically active sulfate as adenosine3 -phosphate-5 -phospho-sulfate. J. Am. Chem. Soc. 78:2652-2653. Weigmann, A., Corbeil, D., Hellwig, A., and Huttner, W.B. 1997. Prominin, a novel microvillispecific polytopic membrane protein of the apical surface of epithelial cells, is targeted to plasmalemmal protrusions of nonepithelial cells. Proc. Natl. Acad. Sci. U.S.A. 94:12425-12430. Wolfender, J.L., Chu, F., Ball, H., Wolfender, F., Fainzilber, M., Baldwin, M.A., and Burlingame, A.L. 1999. Identification of tyrosine sulfation in Conus pennaceus conotoxins alpha-PnIA and alpha-PnIB: further investigation of labile sulfoand phosphopeptides by electrospray, matrixassisted laser desorption/ionization (MALDI) and atmospheric pressure MALDI mass spectrometry. J. Mass Spectrom. 34:447-454.
Key References Huttner, 1984. See above. Describes methods for sulfate labeling and various procedures to detect tyrosine sulfate in proteins. The determination of the stoichiometry of tyrosine sulfation of proteins is also discussed. Bundgaard et al., 2002. See above. Describes several analytical methods for tyrosine sulfate analysis.
Contributed by Denis Corbeil, Christoph Thiele, and Wieland B. Huttner Max-Planck-Institute of Molecular Cell Biology and Genetics Dresden, Germany
Tyrosine O-Sulfation
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Analysis of Protein Sumoylation
UNIT 14.8
Small ubiquitin-like modifier (SUMO) proteins are polypeptides that are covalently attached to specific lysine residues of other proteins, an event referred to as sumoylation. As the name indicates, SUMO proteins are structurally similar to ubiquitin. In contrast to ubiquitination, however, sumoylation does not promote degradation of proteins but instead regulates key functional properties of target proteins, including subcellular localization, protein partnering, transactivation functions of transcription factors, and other properties (Dohmen, 2004; Hilgarth et al., 2004; Johnson, 2004; Hay, 2005). In this unit two methods for determining whether a protein is a substrate for sumoylation are described: one uses immunoprecipitation followed by immunoblotting with anti-SUMO antibodies (Basic Protocol 1), while the other involves in vitro translation in the presence of 35 S-methionine followed by an enzymatic sumoylation reaction, SDS-PAGE, and autoradiography (Basic Protocol 2). In conjunction with point mutagenesis, these protocols can also be used to identify the lysine residue(s) within a protein where SUMO attachment occurs.
IMMUNOPRECIPITATION OF TARGET PROTEINS FOLLOWED BY ANTI-SUMO IMMUNOBLOTTING
BASIC PROTOCOL 1
This protocol describes methods for immunoprecipitating proteins to be tested for sumoylation and immunoblotting sumoylated proteins, using anti-SUMO antibodies (cell lysis buffers containing SDS or N-ethylmaleimide inhibit desumoylase enzymes). Although the theoretical molecular weight of the SUMO proteins is ∼11 kDa, the size increase of sumoylated proteins determined by SDS-PAGE typically falls in the range of 15 to 17 kDa for each SUMO protein added. In the case of a protein with multiple sumoylation sites, or where SUMO chains form on a lysine target site (see Commentary), multiples of this size increase are expected, sometimes yielding very large shifts in electrophoretic mobility. Multiple bands representing different sumoylation states of the protein are also possible. Immunoprecipitation and immunoblotting can be used to analyze sumoylation of endogenous proteins or transfected proteins containing an epitope tag (e.g., FLAG, myc; see UNIT 9.9). The transfection approach can also be used to determine the lysine residue(s) to which the SUMO group is attached by comparing the sumoylation of wild-type constructs to ones in which candidate lysine residues have been changed to nonsumoylatable arginines.
Materials 100-mm plates with cells expressing the protein of interest Phosphate-buffered saline (PBS; APPENDIX 2E) SDS lysis solution (see recipe) PBS (APPENDIX 2E) with 0.5% (v/v) nonylphenyl-polyethylene glycol (NP-40; Sigma-Aldrich) and 1× complete protease inhibitor (Roche) N-ethylmaleimide (optional) Antibodies to the protein of interest or to the epitope tag of a transfected, tagged protein IgG, species-matched, nonspecific control (Sigma-Aldrich) Protein G–Sepharose or protein A–Sepharose (Amersham) 2× SDS-PAGE sample buffer (see recipe) Blocking buffer: e.g., 5% (w/v) nonfat dry milk Antibodies to SUMO-1, SUMO-2, or SUMO-3 (Invitrogen) Contributed by Roland S. Hilgarth and Kevin D. Sarge Current Protocols in Protein Science (2006) 14.8.1-14.8.7 C 2006 by John Wiley & Sons, Inc. Copyright
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Enhanced chemiluminescence (ECL) or enhanced chemifluorescence (ECF) reagents (Pierce) Plate scraper 2-ml microcentrifuge tubes Refrigerated centrifuge Sonicator Dounce homogenizer (optional) Micropipettor End-over-end rotator Additional reagents and equipment for SDS-PAGE electrophoresis (UNIT 10.1), electroblotting (UNIT 10.7), and immunoblot detection (UNIT 10.10) 1. Wash two 100-mm plates of cells with PBS, harvest cells by scraping each plate in 1 ml PBS, transfer to a 2-ml microcentrifuge tube, and collect by centrifuging 30 sec at 13,000 × g, room temperature. More cells may be required for proteins expressed at low levels.
2. Add 150 µl SDS lysis solution and sonicate briefly. Add 1.5 ml PBS with 0.5% NP-40 and 1× complete protease inhibitor. Sonication helps lyse cells and also shears DNA to make the resulting solution less viscous. Optimal sonication times and settings will depend on the particular sonicator model, but in principle sonication can be stopped as soon as viscosity is noticeably reduced. Some researchers add N-ethylmaleimide (to 20 mM, freshly dissolved) to the 1.5 ml dilution solution (PBS with 0.5% NP-40 and 1× complete protease inhibitor) to enhance inactivation of desumoylases. Alternatively, the cells can be lysed in any of the standard immunoprecipitation lysis buffers, e.g., RIPA (see UNIT 14.5; for others examples see UNIT 9.8), known to extract the protein of interest, providing the lysis solution is supplemented with 20 mM Nethylmaleimide (freshly dissolved). If an alternative lysis buffer is used, add 1.5 ml to the cells; further dilution is unnecessary. It is important that lysis proceed quickly, perhaps aided by pipetting, Dounce homogenizing, or sonicating to insure that desumoylases are rapidly and uniformly inactivated.
3. Centrifuge 10 min at 16,000 × g, 4◦ C to clear the lysates. At this point a small aliquot (e.g., 30 µl) is usually taken out and frozen at −20◦ C to be later run as “input” in SDS-PAGE.
4. Divide the cleared lysate into two 1.5-ml microcentrifuge tubes. To one of the tubes add an antibody specific to the protein of interest (or to the epitope tag if analyzing transfected tagged protein), and to the other tube add a species-matched, nonspecific IgG. Add an appropriate amount of protein G–Sepharose to both tubes according to the supplier’s instructions. Rotate tubes for one hr at 4◦ C, using an end-over-end rotator. Substitute protein A–Sepharose when using antibodies that are not efficiently bound by protein G. It is best if the nonspecific IgG is species-matched, i.e., rabbit IgG if using a rabbit polyclonal primary antibody. Some researchers preclear the lysate before adding primary antibodies by incubating it with protein G–Sepharose, centrifuging, and then proceeding with the supernatant.
5. Collect the beads by centrifuging 10 sec at 16,000 × g, 4◦ C, and pipet off the supernatant. Wash beads three times, 5 min each, with 1 ml PBS containing 0.5% NP-40 and complete protease inhibitor. Analysis of Protein Sumoylation
If an alternative lysis buffer is used (step 2 annotation), use that lysis buffer for washing.
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6. Remove as much of the last wash as possible with a micropipettor. Add 20 µl 2× SDS-PAGE sample buffer to the beads and place in a boiling water bath for 4 min. Analyze samples by SDS-PAGE (UNIT 10.1) and transfer to a nitrocellulose or nylon membrane (UNIT 10.7). 7. Incubate the membrane for 60 min with blocking buffer (e.g., 5% (w/v) nonfat dry milk). Add an appropriate dilution of anti-SUMO-1, -SUMO-2, or -SUMO-3 antibodies. SUMO-1 antibodies are the most commonly used for initial experiments, but antibodies against the other SUMO proteins can be employed if results for SUMO-1 are negative.
8. Visualize the immunoblot signal using enhanced chemiluminescence (ECL) or enhanced chemifluorescence (ECF) reagents, following the manufacturer’s instructions (also see UNIT 10.10).
IN VITRO SUMOYLATION OF A CANDIDATE PROTEIN SUBSTRATE USING A RECONSTITUTED ENZYMATIC REACTION
BASIC PROTOCOL 2
In this protocol the protein of interest is translated in vitro (typically with 35 S-methionine incorporation) and then incubated in a reaction containing the SUMO E1 and E2 enzymes and the SUMO-1 protein (see Figure 14.8.1 and Background Information for more information on the functions of these proteins in the sumoylation reaction), followed by SDS-PAGE and autoradiography to determine whether a lower mobility band appears that would be consistent with a sumoylated form of the target protein. Performing this same reaction using mutants of the protein in which candidate sumoylation site lysine residues are changed to nonsumoylatable arginines can be used to determine the site(s) at which SUMO attachment is occurring.
Materials In vitro coupled transcription/translation kit (e.g., Promega TNT T7 Quick for PCR DNA kit) PCR product or plasmid containing an open reading frame (ORF) for the protein of interest 10× sumoylation buffer (see recipe) Creatine phosphokinase (Sigma-Aldrich) Creatine phosphate (Sigma-Aldrich) Inorganic pyrophosphatase (Sigma-Aldrich) SAE1/SAE2 heterodimer, purified (E1; LAE International, http://www.laebio.com) Ubc9, purified (E2; IAE International, http://www.laebio.com) SUMO-1, SUMO-2, or SUMO-3 (purified; LAE International, http://www.laebio.com) 2× SDS-PAGE sample buffer (see recipe) SDS-PAGE fixing solution (see recipe) Whatman filter paper Additional reagents and equipment for SDS-PAGE electrophoresis (UNIT 10.1), electroblotting (UNIT 10.7), and autoradiography (UNIT 10.11) 1. Translate the protein to be tested using a commercially available in vitro coupled transcription/translation kit using PCR product or plasmid containing an ORF for the protein of interest and following the kit manufacturer’s instructions. Use of the translation kit suggested in the materials list requires a PCR product containing the ORF of the protein that incorporates a T7 promoter sequence on the appropriate end of the construct. However, a plasmid in which the ORF with some 5 flanking sequence of the gene of interest is present downstream of a T7 promoter can also be used with related translation kits.
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Figure 14.8.1 The SUMO conjugation pathway. After they are translated, SUMO proteins must first be processed by a SUMO protease (e.g., Ulp1) which removes four C-terminal residues, resulting in the mature SUMO protein form, ending with a glycine residue. (SUMO proteases are also responsible for removing SUMO groups from proteins.) This mature form is then activated in an ATP-dependent reaction by forming a thioester bond with a cysteine residue in the Uba2 subunit of the heterodimeric E1-activating enzyme. The SUMO moiety is then transferred to the E2-conjugating enzyme Ubc9. In the final step SUMO is transferred in a ligation reaction from Ubc9 to substrate proteins, forming an isopeptide bond between the terminal glycine on SUMO and the ε-amino group of a lysine in the target protein. The efficiency of sumoylation of some proteins is enhanced by SUMO ligase E3 proteins because of their ability to bind both Ubc9 and the target protein, thereby increasing the kinetics of the SUMO transfer.
For general information about producing a PCR product or plasmid containing an ORF see APPENDIX 4C or APPENDIX 4J.
2. Prepare a sumoylation master mixture containing the following (on ice):
1 µl 10× sumoylation buffer 0.4 U creatine phosphokinase 10 mM creatine phosphate 0.6 U inorganic pyrophosphatase 100 ng purified SAE1/SAE2 heterodimer (E1) 400 ng purified Ubc9 (E2) 1 µg purified SUMO-1 Bring volume to 10 µl with H2 O Also prepare a negative control mix in which the SUMO protein is omitted. Purified SUMO E1 (SAE1/SAE2 subunits) and E2 (Ubc9) enzymes and SUMO proteins are commercially available (e.g., LAE International) The relatively high concentrations of E1 and E2 enzymes present in this reaction are sufficient to drive sumoylation of most substrates even in the absence of E3 proteins. If using this reaction to identify E3 proteins important for sumoylation, the amounts of E1 and E2 enzymes added can be reduced to make the reaction sensitive to E3 contributions. Purified SUMO-2 or SUMO-3 can be used in this reaction instead of SUMO-1, if desired, although in most cases SUMO-1 is tried initially. The choice is empirically determined.
3. Add 2 µl translated protein solution to 10 µl sumoylation master mix and incubate 1 hr at 37◦ C. 4. Add 12 µl 2× SDS-PAGE sample buffer and boil 5 min in a boiling water bath. Separate proteins by SDS-PAGE (see UNIT 10.1). Analysis of Protein Sumoylation
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5. Incubate gel 10 min in SDS-PAGE fixing solution, dry on Whatman paper, and expose to X-ray film. The exposure time will depend on the amount of labeled translation product and the efficiency of sumoylation, but exposures of at least 10 hr are typical. 35
S emissions are fairly low energy and it is advisable not to leave plastic wrap between the dried gel and the X-ray film if possible because this can increase required exposure times. If an epitope tag is present in the translated protein, immunoblot detection with antibodies against the tag could be used.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
SDS lysis solution 150 mM Tris·Cl, pH 6.8 (APPENDIX 2E) 5% (w/v) SDS 30% (v/v) glycerol Prepare fresh just before use SDS-PAGE fixing solution 50% (v/v) methanol 10% (v/v) glacial acetic acid Prepare fresh just before use SDS-PAGE sample buffer, 2× 125 mM Tris·Cl, pH 6.8 (APPENDIX 2E) 20% (v/v) glycerol 4% (w/v) SDS 0.2 M dithiothreitol (DTT) 0.01% bromophenol blue Store up to 6 months at −20◦ C Sumoylation buffer, 10× 500 mM Tris·Cl, pH 7.6 500 mM KCl 50 mM MgCl2 10 mM DTT 10 mM ATP Store in ∼100-µl aliquots up to 6 months at −20◦ C COMMENTARY Background Information Like ubiquitination, sumoylation involves a series of enzymatic steps (see Fig. 14.8.1), but the proteins involved are distinct from those in the ubiquitin conjugation pathway. First, SUMO proteins must undergo proteolytic processing, which is performed by SUMO proteases (Ulps), near the C-terminal end to form the mature protein. These proteases are dual-functional, and they are also responsible for removing SUMO groups from substrate proteins by cleaving the isopeptide bonds by which they are joined (Dohmen, 2004; Johnson, 2004; Hay, 2005). The mature, pro-
cessed SUMO protein is covalently attached via a thioester bond to the Uba2 subunit of the heterodimeric SUMO E1 activating enzyme in an ATP-dependent reaction (Johnson et al., 1997; Desterro et al., 1999; Gong et al., 1999; Okuma et al., 1999). The SUMO moiety is transferred from the E1 to Ubc9, the SUMO E2 enzyme, which then binds to the KXE consensus sequence ( represents a hydrophobic amino acid) found in target proteins and forms an isopeptide bond between the ε-amino group of the lysine within this sequence and the carboxyl group of the C-terminal glycine of the SUMO polypeptide (Desterro et al.,
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1997; Johnson and Blobel, 1997; Rodriguez et al., 2001; Sampson et al., 2001). SUMO E3 proteins have been identified that enhance the efficiency of SUMO attachment by interacting with both Ubc9 (the E2 enzyme) and the substrate, thereby acting as bridging factors (Dohmen, 2004; Hilgarth et al., 2004; Johnson, 2004; Hay, 2005). Vertebrate cells contain three SUMO paralogs. SUMO-2 and SUMO-3 are very similar to each other in sequence, and have ∼50% sequence identity with SUMO-1, which is the best characterized of the three vertebrate SUMO proteins.
Critical Parameters and Troubleshooting One critical issue affecting success in detecting sumoylation of cellular proteins is that desumoylating enzymes are very active in cell lysates and will rapidly remove SUMO groups from cellular proteins. For this reason it is very important for these analyses that cells be lysed using solutions containing agents that rapidly and efficiently inactivate desumoylases, e.g., SDS or N-ethylmaleimide. Another issue is that cells do not contain very large free pools of unconjugated SUMO proteins; thus, particularly for approaches where candidate sumoylation target proteins are overexpressed in cells by transfection, some researchers cotransfect expression constructs encoding one or more of the SUMO proteins to ensure that the free pools of these proteins do not become limiting. Regarding the in vitro sumoylation reaction, the amounts of SUMO E1 and E2 enzymes included in the reaction are sufficient to drive sumoylation of many proteins even in the absence of any E3 proteins. However, it is possible that a negative result from this approach could be due to the need for an E3 protein to enhance sumoylation of a particular protein to levels detectable by this assay. In this case, supplementing the in vitro sumoylation reaction with cell extracts or known E3 proteins might overcome this limitation.
Anticipated Results
Analysis of Protein Sumoylation
Immunoprecipitation analysis should show the presence of a SUMO antibody-reactive band in samples immunoprecipitated by antibodies against the protein of interest but not by the nonspecific species-matched IgG control antibodies. The expected size of putative sumoylated forms of a protein on SDS-PAGE will depend on how many different SUMO attachment sites the protein has. In addition, SUMO-2 and SUMO-3 have been reported to form polymeric chains reminiscent of ubiqui-
tin, which could result in large increases in size on a gel for a sumoylated protein compared to the nonsumoylated form (Tatham et al., 2001). Thus, it is possible to observe bands that are multiples of the ∼15 to 17 kDa size of each SUMO unit, as well as multiple bands representing different degrees of sumoylation of the protein. Results from in vitro sumoylation assays showing the appearance of a new, higher molecular weight band in reactions containing the reconstituted sumoylation enzyme machinery (E1, E2, SUMO), but not in reactions in which one of the components (often the SUMO donor) is left out, would suggest that the protein is a sumoylation substrate. However, it is important to note that such a result does not prove that the endogenous protein expressed in cells is sumoylated, which should be demonstrated using the immunoprecipitation approach (Basic Protocol 1).
Time Considerations For the immunoprecipitation approach (Basic Protocol 1), once cell pellets are obtained it should be possible to complete the steps up to the point of electrophoresing the immunoprecipitated proteins and transferring them to a membrane in one day. The anti-SUMO antibody immunodetection steps can then be performed the following day. For the in vitro sumoylation approach (Basic Protocol 2), if the template for the in vitro translation has been made (plasmid DNA or PCR product), it should be possible to complete all the steps in one day, including separating the protein products on SDS-PAGE and fixing and drying the gel, which can then be exposed overnight.
Literature Cited Dohmen, R.J. 2004. SUMO protein modification. Biochim. Biophys. Acta 1695:113-131. Desterro, J.M., Rodriguez, M.S., Kemp, G.D., and Hay, R.T. 1999. Identification of the enzyme required for activation of the small ubiquitin-like protein SUMO-1. J. Biol. Chem. 274:1061810624. Desterro, J.M., Thomson, J., and Hay, R.T. 1997. Ubch9 conjugates SUMO but not ubiquitin. FEBS Lett. 417:297-300. Gong, L., Li, B., Millas, S., and Yeh, E.T. 1999. Molecular cloning and characterization of human AOS1 and UBA2, components of the sentrin-activating enzyme complex. FEBS Lett. 448:185-189. Hay, R.T. 2005. SUMO: A history of modification. Mol. Cell 18:1-12. Hilgarth, R.S., Murphy, L.A., Skaggs, H.S., Wilkerson, D.C., Xing, H., and Sarge, K.D.
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2004. Regulation and function of SUMO modification. J. Biol. Chem. 279:53899-53902. Johnson, E.S. 2004. Protein modification by SUMO. Annu. Rev. Biochem. 73:355-382. Johnson, E.S. and Blobel, G. 1997. Ubc9p is the conjugating enzyme for the ubiquitin-like protein Smt3p. J. Biol. Chem. 272:26799-26802. Johnson, E.S., Schwienhorst, I., Dohmen, R.J., and Blobel, G. 1997. The ubiquitin-like protein Smt3p is activated for conjugation to other proteins by an Aos1p/Uba2p heterodimer. EMBO J. 16:5509-5519. Okuma, T., Honda, R., Ichikawa, G., Tsumagari, N., and Yasuda, H. 1999. In vitro SUMO-1 modification requires two enzymatic steps, E1 and E2. Biochem. Biophys. Res. Commun. 254:693-698. Rodriguez, M.S., Dargemont, C., and Hay, R.T. 2001. SUMO-1 conjugation in vivo requires both a consensus modification motif and nuclear targeting. J. Biol. Chem. 276:12654-12659.
Sampson, D.A., Wang, M., and Matunis, M.J. 2001. The small ubiquitin-like modifier-1 (SUMO-1) consensus sequence mediates Ubc9 binding and is essential for SUMO-1 modification. J. Biol. Chem. 276:21664-21669. Tatham, M.H., Jaffray, E., Vaughan, O.A., Desterro, J.M., Botting, C.H., Naismith, J.H., and Hay, R.T. 2001. Polymeric chains of SUMO-2 and SUMO-3 are conjugated to protein substrates by SAE1/SAE2 and Ubc9. J. Biol. Chem. 276:35368-35374.
Contributed by Roland S. Hilgarth and Kevin D. Sarge University of Kentucky Lexington, Kentucky
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Identifying and Quantifying Sites of Protein Methylation by Heavy Methyl SILAC
UNIT 14.9
Protein methylation occurs on arginine, lysine, proline, histidine, and carboxyl groups such as isoprenylated cysteine. Methylation primarily occurs on lysine and arginine residues, though the degree of modification can vary. Lysine can have one, two, or up to three methyl groups on its ε-nitrogen, while arginine may have a single methyl, or two methyl groups in a symmetric or asymmetric configuration. Much of the existing knowledge of methylation comes from in vitro radioactive labeling experiments with purified methyltransferases and substrates. Proteome-wide studies of protein methylation are less common due to the inherent difficulty in identification of this post-translational modification by current methodologies. This unit describes a metabolic labeling strategy using stable isotope labeling by amino acids in cell culture (SILAC) to label methylated proteins and facilitate their identification and quantification with mass spectrometry–based proteomics. In these protocols, cultured mammalian cells grow in media where methionine is substituted with a nonradioactive, stable isotope-labeled version of methionine (L-methionine-13 C, 2 H3 (methyl-13 C, 2 H3 )), which is 4 mass units heavier than the unlabeled version. Cells convert ‘heavy’ methionine to ‘heavy’ S-adenosylmethionine and then methylate cellular substrates such as protein and DNA. Methylated and methionine-containing peptides will be present in ‘light’ and ‘heavy’ forms, allowing identification and relative quantitation between two samples (see Fig. 14.9.1). SILAC is compatible with all MS-based analytical techniques, but, in this unit, the authors discuss the use of nano-scale liquid chromatography (LC) coupled to mass spectrometry (MS) for protein and peptide analysis. Preparation of SILAC media for metabolic labeling of mammalian cells using DMEM is described as an example (Basic Protocol 1). Cells are cultured in this medium and the degree of cellular incorporation of the SILAC amino acid is determined in Basic Protocol 2. Finally, a discussion of how heavy methyl SILAC is applied in the study of protein methylation is provided (Basic Protocol 3).
PostTranslational Modification: Specialized Applications Contributed by Shao-En Ong and Matthias Mann Current Protocols in Protein Science (2006) 14.9.1-14.9.12 C 2006 by John Wiley & Sons, Inc. Copyright
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Figure 14.9.1 Heavy methyl SILAC to study protein methylation. Cells are grown separately in ‘light’ and ‘heavy’ SILAC medium, containing L-methionine and L-methionine-13 C, 2 H3 , respectively. ‘Heavy’ methionine is converted into ‘heavy’ S-adenosylmethionine (SAM), intracellularly. This biological methyl donor labels all methylation substrates in the cell with a ‘heavy’ methyl group. A simple one-to-one mixing experiment is ideal for identifying protein methylation sites. Alternatively, differential treatment of the cells allows the quantitative profiling of protein methylation. The proteins from ‘light’ and ‘heavy’ cells are extracted and enriched for methylated proteins. The sample is then processed and analyzed by mass spectrometry in the usual manner. Protein Methylation by Heavy Methyl SILAC
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PREPARATION OF SILAC MEDIA SILAC is a metabolic labeling strategy for quantitative analysis of proteins by mass spectrometry. It replaces natural isotope abundance amino acids with nonradioactive, stable-isotope labeled amino acids in growth medium to introduce a residue-specific mass difference in cellular proteomes. In this protocol, the authors describe the preparation of SILAC media to be used for the analysis of protein methylation. A more general description of SILAC and its applications in quantitative proteomics is described in the Commentary.
BASIC PROTOCOL 1
Materials SILAC amino acid stock solutions (see recipes) Tissue culture medium appropriate for cells (e.g., Dulbecco’s Modified Eagle’s Medium, DMEM, APPENDIX 3C; Life Technologies Inc.) deficient in SILAC amino acid of choice (methionine, in this example; custom-synthesized “drop-out” amino acid medium is available as a special order product from most medium distributors) L-methionine, natural abundance form, tissue-culture-grade L-methionine-13 C,2 H3 (methyl-13 C, 2 H3 ), stable isotope enriched form, where stable isotopes are located in the methyl group on the side chain of methionine (Sigma-Isotec) Antibiotics for cell culture, e.g., penicillin and streptomycin Glutamine Dialyzed serum (Invitrogen) 0.2-µm pore size filter Tissue-culture-grade filter bottles, sterile NOTE: All reagents and equipment coming in contact with live cells must be sterile and standard sterile tissue culture technique should be used. NOTE: All culture incubations are performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. NOTE: Medium that is deficient in multiple amino acids should always be reconstituted to its original basal formulation as prescribed by manufacturer’s instructions. 1. Prepare concentrated amino acid stock solutions (see recipes) to reconstitute SILAC DMEM ‘light’ and ‘heavy’ media. This protocol uses a 1000× stock of methionine for basal DMEM but this can be adjusted to any convenient concentration for other medium formulations. Where possible, prepare highly concentrated stocks to avoid dilution of the medium upon reconstitution. It may be desirable to avoid filtering costly SILAC amino acid stock solutions, but a final filtration of the reconstituted medium must then be performed.
2. If the SILAC medium is depleted of several amino acids, reconstitute the original DMEM formulation of the culture medium with all but the amino acid of choice (e.g., methionine) using common, nonlabeled amino acids. Divide the SILAC medium depleted of the amino acid of choice into two aliquots. Place these into two separate media storage bottles and add ‘light’ methionine to one aliquot and ‘heavy’ methionine to the other. 3. Filter reconstituted media through a 0.2-µm filter into tissue-culture-grade filter bottles. This step is unnecessary if all amino acid stocks are sterile filtered.
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4. Add antibiotics, glutamine, and dialyzed serum to both ‘light’ and ‘heavy’ media. Although most immortalized cell lines behave similarly in SILAC media, some cell lines do not respond well to dialyzed serum. It may be possible to use a small percentage of normal serum together with dialyzed serum (Gehrmann et al., 2004). It is good practice to perform test SILAC experiments with primary cell cultures or slowly dividing cells, as incorporation rate will depend more on turnover than cell division. Although it is sufficient to analyze the ‘heavy’ sample by MS alone to assay for full incorporation, culturing a dish of ‘light’ cells in parallel provides a comparison of growth rate and cell morphology between the ‘light’ and ‘heavy’ cell states. The results of such an evaluation will hold for a set of frozen cell stocks, and parameters need not be re-determined if using the same set of cells.
5. Store media at 4◦ C, up to three months. BASIC PROTOCOL 2
TESTING FOR FULL INCORPORATION OF SILAC AMINO ACID In order to accurately compare the relative abundance of proteins and peptides from two cell states with SILAC directly, full incorporation of the SILAC amino acid is necessary. Cells are grown in SILAC media for at least five cell doublings, where the theoretical incorporation should be about 97% (1 − (1/2)5 ), before consideration of incorporation due to protein turnover. Additional passage of cells greatly accelerates the adaptation process (i.e., incorporation of heavy amino acids throughout the proteome of the cells). Proteins from adapted cells are extracted, proteolytically cleaved to peptides by trypsin, and analyzed by standard LC-MS techniques commonly used in proteomics (Ishihama, 2005). The degree of incorporation is determined by manually inspecting the raw MS data and determining that peptides containing the SILAC amino acid are present only in the ‘heavy’ form. The length of time required for full incorporation is obtained from this experiment, as well as confirmation of the ability of the cells to grow in SILAC labeling medium.
Materials Cells SILAC media from Basic Protocol 1 Tissue-culture-grade trypsin for lifting adherent cells Urea lysis buffer or a mass spectrometry-compatible lysis buffer for in-solution proteolytic digest of proteins. 1 M dithiothreitol 600 mM iodoacetamide Endoproteinase Lys-C, proteomics grade (Roche Diagnostics) 50 mM ammonium bicarbonate Trypsin, proteomics grade Formic acid C18 reversed phase micro-columns (StageTips; Proxeon Biosystems, http://www.proxeon.com, Rappsilber et al., 2003a) Additional reagents and equipment for standard tissue cell culture and cell harvesting (APPENDIX 3C) and mass spectrometry (Chapter 16) 1. To begin the labeling process, split a dish of cells into two separate dishes, each containing ‘light’ or ‘heavy’ SILAC medium, respectively.
Protein Methylation by Heavy Methyl SILAC
Split the cells into low density cultures of about 10% to 20% confluence, depending on the doubling rate of the cells. Cells can be cultured in small dishes and expanded later in the adaptation phase or after five doublings to save SILAC media.
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2. Passage cells as needed, taking care to keep cells in their respective SILAC media throughout the experiment. Culture the cells in the same manner as one would use with standard cell culture medium. Be sure to change the medium every 2 days, at the minimum.
3. After five doublings (5 days for HeLa), harvest cells grown in ‘heavy’ media into microcentrifuge tubes containing 750 µl 8 M urea, for each 15-cm dish with ∼107 cells. Vortex, then pellet cellular debris in a microcentrifuge for 10 min at 13,000 × g, 15◦ C and save the supernatant. The number of cells required for this assay is low as whole-cell lysates are digested to peptides with an in-solution digest. A few micrograms of peptide from lysates should be sufficient to generate good MS signals. Alternatively, proteins can be separated on a 1-D SDS-PAGE gel and bands excised for in-gel protein digestion to determine the degree of incorporation.
4. Reduce proteins with 10 mM dithiothreitol for 30 min at 37◦ C, then alkylate the proteins with 55 mM iodoacetamide for 20 min at room temperature, in the dark. Heating urea can result in extensive carbamylation of primary amines, complicating and reducing the sensitivity of MS analyses.
5. Add endoproteinase Lys-C (1 µg enzyme per 100 µg protein) and incubate for 2 hr at 37◦ C. 6. Dilute the digest with 3 vol of 50 mM ammonium bicarbonate until the concentration of urea is below 2 M and add trypsin (1 µg enzyme per 100 µg protein). Incubate for at least 4 hr or leave to digest overnight at 37◦ C. 7. Stop the digestion by acidifying the mixture to a final concentration of 1% formic acid. 8. Purify the peptide mixture on a C18 reversed phase micro-column (StageTip) and analyze sample using LC-MS. 9. To identify as many peptides as possible, collect MS and tandem MS/MS experiments in data-dependent mode. Many different MS instruments are used for analyses but most proteomics experiments in the literature rely on tandem MS/MS spectra and database search algorithms to identify peptides and proteins (Peng and Gygi, 2001). In this mode, sequence information is obtained for each precursor mass, and the mass of the peptide and its fragment masses are used to search a database of protein sequences (Steen and Mann, 2004).
10. Identify peptides with ‘heavy’ methionine by specifying this as a possible mass modification in the MS search algorithm. Methylated peptides should also be detectable only in their ‘heavy’ form, but the likelihood of detecting these peptides without any specific enrichment step is low, as they occur far less frequently than methionine-containing peptides. With full incorporation, cells would have incorporated the amino acid throughout their proteome and ‘heavy’ peptides should be observed without their ‘light’, natural abundance counterparts. This requires manual examination of the MS data as the analysis may already be complexity-limited and data-dependent experiments often fail to detect low-abundance species. Inspect the raw MS data to see if any peptide cluster can be detected at the mass corresponding to the unlabeled peptide for identified ‘heavy’ peptides. High mass accuracy MS instruments are desirable for quantitation in general and provide greater quantitative accuracy (Aebersold and Mann, 2003).
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BASIC PROTOCOL 3
STUDYING PROTEIN METHYLATION WITH HEAVY METHYL SILAC WITH ANTIBODIES AGAINST METHYLATED PROTEINS The basic SILAC experimental design requires that cells be adapted to ‘light’ and ‘heavy’ media before any differential treatment to the cells is applied. Proteins from the two states are of equal relative abundance in relation to one another at the start of the experiment. Upon differential treatment, protein levels and degree of methylation may change, and this is determined from the quantitative proteomics data. One set of cells can be treated with a drug or may be induced to differentiate into a different cell type in order to induce a change in methylation levels. Heavy methyl SILAC quantifies changes in the degree of methylation at a given site. If the goal of the proteomic experiment is a survey of protein methylation, the identities of modified proteins are mostly unknown a priori. Methylation changes the mass of the modified residue by multiples of a –CH2 group. Since so many peptide sequences exist in the database and their chemical compositions can differ by exactly n (–CH2 ) groups, database search algorithms often find it difficult to distinguish between the myriad of possible matches. Heavy methyl SILAC is ideally suited for identifying methylated peptides as it provides orthogonal lines of evidence that unambiguously confirm the peptide identification.
Materials Cells fully adapted to ‘light’ and ‘heavy’ SILAC medium as described in Basic Protocol 2 Lysis buffer (see recipe) compatible with co-immunoprecipitation experiments Trypsin, tissue-culture-grade Protein A or G agarose beads (Pierce or Sigma) Antibodies directed against a specific protein or a class of methylated residues, such as anti-methylarginine or anti-methyl-lysine antibodies 2× or 6× SDS-PAGE loading buffer (see recipe) 50 mM glycine, pH 2.4 Cell scraper Econo-Pac chromatography column (Bio-Rad) Additional reagents and equipment for adapting cells to SILAC media (Basic Protocol 2), Bradford protein assay (UNIT 3.4), 1-D SDS-PAGE (UNIT 10.1), staining of gels (UNIT 10.5), and mass spectrometry (Chapter 16) 1. Adapt sufficient numbers of ‘light’ and ‘heavy’ labeled cells in SILAC media as described in Basic Protocol 2. The number of cells required depends on the type of experiment to be performed. Identification of post-translationally modified proteins and, specifically, methylated peptides, is difficult without any specific enrichment (Mann and Jensen, 2003). While the performance of a particular reagent in specifically enriching for its target greatly varies, it is desirable to start with at least a few milligrams of protein from whole-cell lysates. Optimization of the co-precipitation should be performed in a separate experiment to determine the appropriate amounts of antibody, beads, and lysate to use. Ideally, distinct protein bands between the two experimental conditions would be observable in a colloidal Coomassie-stained gel.
Protein Methylation by Heavy Methyl SILAC
2. Harvest ‘light’ and ‘heavy’ cells separately by scraping in lysis buffer and placing on ice. Centrifuge the lysates for 10 min at 13,000 × g, 4◦ C to remove cellular debris. Determine the total protein concentrations of the two lysates with a protein assay (e.g., Bradford; UNIT 3.4).
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Alternatively, subcellular fractionation of cellular organelles can be performed by harvesting cells in a buffer that does not contain detergents. In this case, remove a small aliquot of the cells from each condition to lyse and determine the protein concentrations. Mix intact cells and perform subcellular fractionation (UNITS 4.2 & 4.3).
3. Mix proteins from ‘light’ and ‘heavy’ lysates in a known ratio, typically 1:1. 4. Preclear the lysates with Protein A or G agarose beads for 1 hr to remove abundant proteins. The lysates are “pre-cleared” to reduce background signal from proteins that interact nonspecifically with the solid phase or secondary antibodies. This is achieved by incubating the lysates with the same capture matrix, but without the primary antibody.
5. Centrifuge the beads 1 min at 600 × g, 4◦ C and save the supernatant. 6. Add the appropriate amounts of anti-methyl-antibody and Protein A or G beads as previously determined in the optimization phase. Incubate 6 hr at 4◦ C. 7. Centrifuge the beads 1 min at 600 × g, 4◦ C. Aspirate and save the supernatant, taking care not to disturb the beads. If the volume of lysate and beads is large (i.e., >500 µl beads, >4 ml lysate), it is convenient to perform washing and elution steps in a column format.
8. Wash beads two to three times, each time by adding at least 10× bead volume of lysis buffer (see recipe), centrifuging at 600 × g for 1 min at 4◦ C and removing the supernatant. 9a. If bead volume is not greater than 100 µl; Elute proteins by boiling the beads directly in an equal volume of 2× SDS-PAGE loading buffer directly. 9b. If bead volume is greater than 100 µl; Elute protein from beads in a column with 5 to 10 bead volumes of 50 mM glycine, pH 2.4. Neutralize the pH of the eluate immediately with 1M NaOH. Add 2× or 6× SDS-PAGE loading buffer to final concentration of 1×. To reduce the volume of sample for loading onto a 1-D gel, apply a concentration step with a spin column (e.g., Millipore Centricon spin columns) and further evaporation in vacuum to yield a working solution. Add 6× SDS-PAGE loading buffer (1× final).
10. Separate proteins on a 1-D SDS-PAGE gel (UNIT Coomassie blue stain (UNIT 10.5).
10.1).
Stain gel with colloidal
11. If a specific methylated protein was enriched by immunoprecipitation excise the protein band or cut the gel into multiple lanes for a GeLC-MS analysis. The protein gel isolates the heavy and light chains of the antibody to a particular molecular weight band and improves the dynamic range of MS analysis, as gel slices are analyzed separately.
12. Digest proteins with an in-gel digestion protocol and analyze peptide mixture with LC-MS (Chapter 16). Trypsin is a good protease to use in proteomics (Olsen et al., 2004), but note that it cleaves very inefficiently at methylated arginines and lysines. Despite that, trypsin does not appear to be a poor choice, as peptides still retain a basic residue at their C-termini and positively charged N-termini, generating good fragmentation spectra. An alternative enzyme is Glu-C (V8) from Staphylococcus aureus, which cuts C-terminal to aspartic and glutamic acid residues.
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SUPPORT PROTOCOL
DATA ANALYSIS TO IDENTIFY AND QUANTIFY METHYLATED PEPTIDES Proteomic identification of methylated peptides is challenging because of the dramatic increase in combinatorial search space. Some database search algorithms cannot accommodate several different modifications simultaneously to perform a complete analysis of the data. As a result, bona fide methylated peptides may appear as false negatives based solely on their probability scores. Heavy methyl SILAC addresses this issue because the labeling of a methylated peptide introduces a modification-specific mass shift that specifies the location of the modification and the type of methylation itself (each methyl group adds a 4-Da mass differential from the unlabeled peptide form; see Fig. 14.9.1). Furthermore, observing fragment ion masses generated by two separate MS/MS experiments greatly increases confidence in peptide identification. The ratio of signal intensities from the peptide pair is the relative abundance of the peptide from one sample to the other, allowing quantitation of protein methylation between two cellular states. For data analysis the following should be done: 1. Identify putative methylated peptide sequences using the MS database search algorithm (e.g., Mascot). 2. Manually inspect the data. Verify methylated peptide identifications based on the presence of two peptide clusters that are separated by the expected mass difference. The MS/MS spectra from the pair must support the peptide identification. 3. Use narrow mass widths (∼0.02 Da) to extract ion chromatograms for ‘light’ and ‘heavy’ peptides and use these values to determine relative abundance of the methylated peptides (this presupposes a high-resolution mass spectrometer).
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Lysis buffer 150 mM NaCl 50 mM Tris·Cl, pH 7.4 (APPENDIX 2E) 1% (v/v) NP-40 0.1% (w/v) sodium deoxycholate 1 mM EDTA Store at room temperature Add one Roche Complete protease inhibitor tablet and keep cold on ice before use SDS loading buffer, 2× 25 ml 4× Tris·Cl/SDS, pH 6.8 (see recipe) 20 ml glycerol 4 g SDS 2 ml 2-ME or 3.1 g DTT 1 mg bromphenol blue Add H2 O to 100 ml and mix Store in 1-ml aliquots at −70◦ C SDS loading buffer, 6×
Protein Methylation by Heavy Methyl SILAC
7 ml 4× Tris·Cl/SDS, pH 6.8 (see recipe) 3.0 ml glycerol 1 g SDS 0.93 g DTT continued
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1.2 mg bromphenol blue Add H2 O to 10 ml (if needed) Store in 0.5-ml aliquots at −70◦ C SILAC amino acid stock solutions Prepare 30 mg/ml stocks of L-methionine and 32.41 mg/ml L-methionine-13 C,2 H3 (methyl-13 C, 2 H3 ; Sigma Isotec) in PBS (APPENDIX 2E) and sterile filter using a 0.22-µm pore size syringe filter. Note that the difference in mass of the amino acid should be taken into account when preparing the stock solutions. The concentration of amino acid to use depends entirely on the medium formulation. Prepare concentrated stocks to avoid dilution of the medium when the SILAC amino acid is added.
Tris·Cl/SDS, pH 6.8 (0.5 M Tris·Cl containing 0.4% SDS), 4× Dissolve 6.05 g Tris base in 40 ml H2 O. Adjust to pH 6.8 with 1 N HCl. Add H2 O to 100 ml total volume. Filter the solution through a 0.45-µm filter, add 0.4 g SDS, and store at 4◦ C up to 1 month.
COMMENTARY Background Information SILAC is a metabolic labeling strategy for quantitative proteomics that has gained popularity very rapidly because it is simple to perform, is robust, and its versatility allows its inclusion into many existing proteomics workflows (Ong and Mann, 2005). SILAC is a general approach and has been successfully applied in yeast (Gruhler et al., 2005a), Arabidopsis thaliana (Gruhler et al., 2005b), Plasmodium falciparum (Nirmalan et al., 2004) and should be extensible to higher organisms. The current unit is limited to discussion of mammalian cell culture. SILAC relies on the replacement of specific normal, natural isotope abundance amino acids, such as L-arginine, with their stable isotopeenriched counterpart, L-arginine with six 13 C atoms (Ong et al., 2002; 2003). This introduces a 6-Da mass difference for each occurrence of arginine in the cellular proteome. Incorporation of the labeled amino acid is complete after six cell doublings. Cells grown in ‘light’ and ‘heavy’ media are harvested and protein or whole cells are mixed in a defined ratio (usually 1:1). Mixing cells at harvest allows subcellular fractionation, or other protein purification, to be performed without compromising quantitative accuracy. Upon digestion with trypsin, arginine-containing peptides from a mixture of labeled and unlabeled samples are present in a pair separated by a sequence-specific mass difference. The ratio of peak intensities of ‘light’ and ‘heavy’ peptides is the measure of relative abundance of proteins from which they are derived. The
choice of SILAC amino acid depends on the application. Using both lysine and arginine allows quantitation from every peptide, except the carboxyl-terminal peptide of the protein, because trypsin cleaves specifically after these residues (Olsen et al., 2004) and is excellent for protein profiling and analysis of post-translational modifications (Ibarrola et al., 2003). In heavy methyl SILAC, the cell’s conversion of methionine to the methyl donor, S-adenosylmethionine, is used to introduce the mass difference through the modifying substituting group itself (Ong et al., 2004). This is a critical advantage, since the method distinguishes between the different forms of methylation possible on a single residue. The SILAC approach for quantitative proteomics does not involve any chemical labeling steps. As a result, it is efficient, simple to perform, and uses only standard tissue cell culture techniques and equipment common in most laboratories. Although protein methylation has been recognized for some time (Ambler and Rees, 1959) and has been implicated in a number of important biological processes, including transcriptional regulation and protein-protein interactions, relatively little is known about this post-translational modification. Its role in histone biology is the most established, and the stability of the methylation mark was always its hallmark. However, the startling and important discoveries of arginine and lysine demethylases in 2004 (Cuthbert et al., 2004; Shi et al., 2004; Wang et al., 2004) changed that perspective dramatically and
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heralded new avenues of research (McBride and Silver, 2001; Bannister and Kouzarides, 2005; Bedford and Richard, 2005; Fackelmayer, 2005). Researchers have relied on in vitro radioactive isotope labeling experiments using radioisotope-labeled S-adenosylmethionine and purified, recombinant methyltransferases as their primary tools to study protein methylation. These experiments are cumbersome to perform and rarely characterize more than a single protein. Other mass spectrometry approaches have been used to analyze protein methylation, but these are either not general for all methylated residues (Brame et al., 2004; Rappsilber et al., 2003b; Zhang et al., 2004) or are not amenable to protein mixtures. By comparison, heavy methyl SILAC is simple to perform, is a general approach that can be used for all methylated residues, and greatly improves the confidence in identifying and quantifying methylation sites with conventional MS-based proteomics methods (Ong et al., 2004).
Critical Parameters and Troubleshooting The primary consideration in heavy methyl SILAC is the ability of the cells to grow in the SILAC medium. SILAC medium can be obtained as a custom synthesized product from medium manufacturers and, therefore, should be identical to regular cell culture medium, save for the dialyzed serum. If the dialyzed serum is preventing normal cell growth, using a small percentage of normal serum in combination with dialyzed serum may be a solution (Gehrmann et al., 2004). The level of incomplete incorporation from unlabeled amino acid in normal serum can then be accounted for in a separate analysis of the ‘heavy’ sample. Performing a labeling experiment to determine the amount of time required for complete labeling is well worth the effort. Incomplete labeling adversely affects quantitative accuracy and complicates data interpretation.
Anticipated Results
Protein Methylation by Heavy Methyl SILAC
Many proteomics approaches include a prefractionation of the sample either by strong cation exchange chromatography or separation on a one-dimensional SDS-PAGE gel. An example of a LC-MS analysis of a single gel slice from a 1-D SDS-PAGE gel (GeLCMS) is shown in Figure 14.9.2. The MS instrument
detects peptide species in MS spectra as they elute off the reversed phase column, selecting each of these in turn for tandem MS/MS experiments and generating MS/MS spectra. The rapid data acquisition capabilities of modern MS instruments allow the collection of thousands of MS and MS/MS spectra (Panels B and C in Fig. 14.9.2) in a single LC-MS run where tens or even hundreds of thousands of individual peptide species are detectable. In Figure 14.9.2 it is apparent, even from the base peak chromatogram (the extracted ion signal for the most intense peak in each MS spectrum), that the peptide mixture is extremely complex even after fractionation of the sample. The MS/MS spectral data (Figure 14.9.2C) are extracted from the data file and submitted to database search engines for peptide and protein identification. In a typical proteomics experiment with nano-scale LC-MS, one can expect to identify a few hundred proteins from an analysis of whole-cell lysate. The identification of posttranslationally modified peptides is far more difficult, particularly if no enrichment step is performed. It is not surprising, therefore, if the overwhelming majority of identified peptides are unmodified. Analysis of methylated peptides is made more challenging because most database search algorithms are not designed to handle the wide range of mass modifications that can occur. It may be necessary to search the data using a subset of possible mass modifications, and then further verification of the MS data can extract the complementary heavy methyl SILAC information from the pairs of methylated peptides.
Time Considerations Cells typically take about 5 to 6 days to adapt and grow in SILAC media. Expanding the number of cells necessary for a proteomicscale experiment may require additional time. Harvesting and affinity purification of proteins will require a day. Analysis of proteins by MS typically takes 2 to 3 days, including an overnight protein digestion step and analysis time on the MS instrument. Sample fractionation by orthogonal LC separation or a GeLCMS analysis will improve analytical coverage of the sample, but requires additional time. If necessary, adapted cells can be maintained in SILAC medium and a second round of experiments can be performed as soon as sufficient cell numbers are available.
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Figure 14.9.2 Mass spectrometric data acquisition and analysis to identify and quantify methylated peptides. (A) The base peak chromatogram from a typical 60-min gradient LC-MS. The arrow (marked B) indicates the 1-sec MS scan acquired and shown in Panel B. (B) The inset panel is a zoomed view of the light and heavy methyl labeled triply charged methylated peptide pair S(diMeR)GGGGGGFHR separated by 2.67 m/z (8 Da/3). The ratio of peak intensities of the pair indicate the relative abundance of the peptides and is used in quantitation. Tandem MS/MS spectra collected from both members of the methylated peptide pair are shown in Panel C. (C) MS/MS spectra from the light and heavy peptide are shown on the top and bottom panels, respectively. The dimethyl-arginine is located towards the beginning of the peptide, and is easily confirmed by the characteristic mass shift ( of 8 Da) of the b ions between the light and heavy MS/MS spectra, while the y ions are unchanged between the two MS/MS spectra.
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Literature Cited Aebersold, R. and Mann, M. 2003. Mass spectrometry-based proteomics. Nature 422:198-207. Ambler, R.P. and Rees, M.W. 1959. Epsilon-NMethyl-lysine in bacterial flagellar protein. Nature 184:56-57. Bannister, A.J. and Kouzarides, T. 2005. Reversing histone methylation. Nature 436:1103-1106. Bedford, M.T. and Richard, S. 2005. Arginine methylation an emerging regulator of protein function. Mol. Cell 18:263-272. Brame, C.J., Moran, M.F., and McBroomCerajewski, L.D. 2004. A mass spectrometry based method for distinguishing between symmetrically and asymmetrically dimethylated arginine residues. Rapid Commun. Mass Spectrom. 18:877-881. Cuthbert, G.L., Daujat, S., Snowden, A.W., Erdjument-Bromage, H., Hagiwara, T., Yamada, M., Schneider, R., Gregory, P.D., Tempst, P., Bannister, A.J., and Kouzarides, T. 2004. Histone deimination antagonizes arginine methylation. Cell 118:545-553. Fackelmayer, F.O. 2005. Protein arginine methyltransferases: Guardians of the Arg? Trends Biochem. Sci. 30:666-671.
Ong, S.E., Blagoev, B., Kratchmarova, I., Kristensen, D.B., Steen, H., Pandey, A., and Mann, M. 2002. Stable isotope labeling by amino acids in cell culture, SILAC, as a simple and accurate approach to expression proteomics. Mol. Cell. Proteomics 1:376-386. Ong, S.E., Kratchmarova, I., and Mann, M. 2003. Properties of 13C-substituted arginine in stable isotope labeling by amino acids in cell culture (SILAC). J. Proteome Res. 2:173-181. Ong, S.E. and Mann, M. 2005. Mass spectrometrybased proteomics turns quantitative. Nat. Chem. Biol. 1:252-262. Ong, S.E., Mittler, G., and Mann, M. 2004. Identifying and quantifying in vivo methylation sites by heavy methyl SILAC. Nat. Methods 1:119126. Peng, J. and Gygi, S.P. 2001. Proteomics: The move to mixtures. J. Mass Spectrom. 36:1083-1091. Rappsilber, J., Ishihama, Y., and Mann, M. 2003a. Stop and go extraction tips for matrix-assisted laser desorption/ionization, nanoelectrospray, and LC/MS sample pretreatment in proteomics. Anal. Chem. 75:663-670.
Gehrmann, M.L., Hathout, Y., and Fenselau, C. 2004. Evaluation of metabolic labeling for comparative proteomics in breast cancer cells. J. Proteome Res. 3:1063-1068.
Rappsilber, J., Friesen, W.J., Paushkin, S., Dreyfuss, G., and Mann, M. 2003b. Detection of arginine dimethylated peptides by parallel precursor ion scanning mass spectrometry in positive ion mode. Anal. Chem. 75:3107-3114.
Gruhler, A., Olsen, J.V., Mohammed, S., Mortensen, P., Faergeman, N.J., Mann, M., and Jensen, O.N. 2005a. Quantitative phosphoproteomics applied to the yeast pheromone signaling pathway. Mol. Cell. Proteomics 4:310-327.
Shi, Y., Lan, F., Matson, C., Mulligan, P., Whetstine, J.R., Cole, P.A., Casero, R.A., and Shi, Y. 2004. Histone demethylation mediated by the nuclear amine oxidase homolog LSD1. Cell 119:941953.
Gruhler, A., Schulze, W.X., Matthiesen, R., Mann, M., and Jensen, O.N. 2005b. Stable isotope labeling of Arabidopsis thaliana cells and quantitative proteomics by mass spectrometry. Mol. Cell. Proteomics 4:1697-1709.
Steen, H. and Mann, M. 2004. The ABC’s (and XYZ’s) of peptide sequencing. Nat. Rev. Mol. Cell. Biol. 5:699-711.
Ibarrola, N., Kalume, D.E., Gronborg, M., Iwahori, A., and Pandey, A. 2003. A proteomic approach for quantitation of phosphorylation using stable isotope labeling in cell culture. Anal. Chem. 75:6043-6049. Ishihama, Y. 2005. Proteomic LC-MS systems using nanoscale liquid chromatography with tandem mass spectrometry. J. Chromatogr. A. 1067:73-83. Mann, M. and Jensen, O.N. 2003. Proteomic analysis of post-translational modifications. Nat. Biotechnol. 21:255-261. McBride, A.E. and Silver, P.A. 2001. State of the arg: Protein methylation at arginine comes of age. Cell 106:5-8.
Protein Methylation by Heavy Methyl SILAC
lysine residues. Mol. Cell. Proteomics 3:608614.
Nirmalan, N., Sims, P.F., and Hyde, J.E. 2004. Quantitative proteomics of the human malaria parasite Plasmodium falciparum and its application to studies of development and inhibition. Mol. Microbiol. 52:1187-1199. Olsen, J.V., Ong, S.E., and Mann, M. 2004. Trypsin cleaves exclusively C-terminal to arginine and
Wang, Y., Wysocka, J., Sayegh, J., Lee, Y.H., Perlin, J.R., Leonelli, L., Sonbuchner, L.S., McDonald, C.H., Cook, R.G., Dou, Y., Roeder, R.G., Clarke, S., Stallcup, M.R., Allis, C.D., and Coonrod, S.A. 2004. Human PAD4 regulates histone arginine methylation levels via demethylimination. Science 306:279-283. Zhang, K., Yau, P.M., Chandrasekhar, B., New, R., Kondrat, R., Imai, B.S., and Bradbury, M.E. 2004. Differentiation between peptides containing acetylated or tri-methylated lysines by mass spectrometry: an application for determining lysine 9 acetylation and methylation of histone H3. Proteomics 4:1-10.
Contributed by Shao-En Ong The Broad Institute of MIT and Harvard Cambridge, Massachusetts Matthias Mann Max Planck Institute for Biochemistry Martinsried, Germany
14.9.12 Supplement 46
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CHAPTER 15 Chemical Modification of Proteins INTRODUCTION
C
hemical modifications of reactive amino acid side chains on proteins have many applications in modern protein science. Common applications include: fluorescent or radioactive tagging of macromolecules; identifying the subcellular localization of a specific protein, i.e., cell surface, bilayer imbedded, or internal; coupling the protein to another component including another protein, nucleic acids, or a solid support; and as a aid in structural analyses (Chapter 11) such as amino acid analysis, sequencing, peptide mapping and mass spectrometry. Another historical application included the use of chemical reagents for structure/function applications, such as identification of residues in enzyme active sites. Over the past decade many, but not all, of these latter approaches have been replaced by site-directed mutagenesis analyses. However, despite the impressive power of mutagenesis methods, it is often most productive to utilize both methods in concert—i.e., initial chemical modification studies can be used to identify likely residues involved in a particular biological function, and mutagenesis studies are then used to confirm and extend these initial observations. The most common chemical modifications target the most reactive amino acid side chains and are primarily oxidations, reductions, and nucleophilic or electrophilic substitutions. Chemical reagents that react with side chain amines or carboxyl groups will also react with the terminal amino and carboxyl groups of a protein, respectively, unless these groups were already modified in vivo. Terminal amino and carboxyl groups of a protein are, on average, more reactive than side-chain amino and carboxyl groups, and this difference can in theory be exploited to specifically label these terminal functional groups. Such specific, single-site labeling or attachment would be very useful for a range of applications, including solid phase sequencing and directional, defined attachment of proteins for affinity chromatography. In practice, selective reactivity of the terminal residue is rarely feasible. Achieving selective end-group modification requires empirical determination of appropriate reaction conditions, which must then be rigorously controlled. However, a more important limitation is that the microenvironments of individual reactive side chains can vary widely and affect reactivity to the point where some side chains are more reactive than the terminal residue.
One of the most reactive and most frequently modified amino acid side chains is the SH group of cysteine (UNIT 15.1). Cysteines occur in proteins either in the reduced state (cysteine or SH form) or in the oxidized state, where a disulfide bond is formed between two cysteines (cystine or -S-S- form). Disulfide bonds can be between two cysteines on the same polypeptide chain or between two different polypeptides. They are major stabilizing components of many folded proteins, especially proteins normally found in an oxidizing environment such as is typically found on the exterior of cells. In most proteins, cysteines occur relatively infrequently and are often useful sites to either tag a protein or to link it to a matrix, e.g., when crosslinking a protein to a chromatography matrix to produce an affinity column or to couple the protein to a biosensor chip for studying macromolecular interactions. Reactions using cysteine-specific reagents can also be used to distinguish between cysteine on the surface of a protein (no reduction, no denaturants), buried cysteines (denaturants, no reduction) and residues in disulfide bonds (reduction and denaturants). Finally, because cysteines are very reactive, it is necessary
Chemical Modification of Proteins
15.0.1 Copyright © 1996 by John Wiley & Sons, Inc.
Supplement 3 CPPS
to convert them to more stable derivatives prior to amino acid or sequence analyses if identification of cysteines is desired. Future supplements will provide protocols for derivatization of lysines and specific protocols for tagging proteins with biotin either through lysines or other reactive side chains. Labeling proteins with biotin has an especially wide range of applications in modern protein science. Later supplements will deal with reactions using carboxyl groups and other reactive side chains. David W. Speicher
Introduction
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Modification of Cysteine
UNIT 15.1
This unit describes a number of methods for modifying cysteine residues of proteins and peptides by reduction and alkylation procedures. These methods vary in duration, concentration, denaturant, and reductant, but they are basically similar in approach. They consist of four basic steps: (1) preparation of the sample (i.e., buffering, denaturation, and/or reduction); (2) reaction with excess alkylating reagent; (3) quenching or removing unreacted alkylating reagent; and (4) removal of salts and reaction by-products. When pH and duration of the reaction are controlled, the modification is normally very specific for cysteinyl residues. In solution, the reaction is usually carried out under denaturing conditions, in either 6 M guanidine⋅HCl or 8 M urea, to optimize accessibility of cysteine side chains that may be protected from reagent access by the structure of the native protein. When denaturation is desired, the denaturing reagents should be added to the sample prior to reduction and/or addition of modifying reagent. Finally, most protocols can be adjusted to accommodate any amount of protein from the low microgram to milligram range. A general procedure for alkylation of cysteine residues in a protein of known size and composition with haloacyl reagents or N-ethylmaleimide (NEM) is presented in Basic Protocol 1. Alternate Protocols 1 and 2 present similar procedures for use when the size and composition are not known and when only very small amounts of protein are available. Alkylations that introduce amino groups using bromopropylamine and N-(iodoethyl)-trifluoroacetamide are presented in Basic Protocols 2 and 3. Two procedures that are often used for subsequent sequence analysis of the protein, alkylation with 4-vinylpyridine and acrylamide, are described in Basic Protocols 4 and 5. In addition, a specialized procedure for 4-vinylpyridine alkylation of protein that has been adsorbed onto a sequencing membrane is presented in Basic Protocol 9. Reversible modification of cysteine residues by way of sulfitolysis is described in Basic Protocol 6, and oxidation with performic acid for amino acid compositional analysis is described in Basic Protocol 7. Gentle oxidation of cysteine residues to disulfides by exposure to air is described in Basic Protocol 8. Support Protocols are included for recrystallization of iodoacetic acid (Support Protocol 1), colorimetric detection of free sulfhydryls (Support Protocol 2), and desalting of modified samples (Support Protocol 3). STRATEGIC PLANNING Modification of cysteine side chains in proteins and peptides is usually done for one or more of three main reasons: (1) to break disulfide bonds and block their reformation, (2) to tag the cysteine residue with an adduct that will assist in its identification or quantitation in subsequent analytical techniques, and (3) to introduce a reporter group, such as a fluorescent label, into a protein or peptide. The reagents most commonly used for modification of cysteine are reagents susceptible to nucleophilic attack by the cysteine sulfur (including reactive halogen compounds and compounds containing reactive double bonds) and oxidizing agents. Strong oxidizers, such as performic acid, convert thiols to sulfonic acid groups; weak oxidizers, such as oxygen, convert thiols to disulfides. Cysteine residues can also form mixed disulfides with other thiol reagents (see Background Information). Mixed disulfide formation is, in fact, the basis for thiol reagent–induced reduction of disulfide bonds, which is often the first step in cysteine modification procedures. Reaction of cysteine with reactive halogen compounds and reactive double bonds requires the presence of the free thiolate anion. Thus, any disulfides originally present must first be reduced to free thiols unless the experiment is designed to restrict modification only to preexisting thiols. Contributed by Mark W. Crankshaw and Gregory A. Grant Current Protocols in Protein Science (1996) 15.1.1-15.1.18 Copyright © 2000 by John Wiley & Sons, Inc.
Chemical Modification of Proteins
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Table 15.1.1
Common Reagents for Modification of Cysteine
Reagent
Molecular Reaction product weight
Applicationsa
Iodoacetic acid
186
General
Iodoacetamide
185
5-I-AEDANSb
434
N-ethylmaleimide
125
3-Bromopropylamine 138
267
N-(iodoethyl)trifluoroacetamide 4-Vinylpyridine Acrylamide
105 71
Sodium sulfite
126
Performic acid
—
Air (oxygen)
—
Comments
Adduct not completely stable to acid hydrolysis; introduces negative charge Converted to S-carboxymethyl S-carboxamidomethyl General cysteine cysteine upon acid hydrolysis (see iodoacetic acid) Does not interfere with Edman S-AEDANS-cysteine Introduces fluorescent label sequence analysis Membrane permeable; can also react S-(ethylsuccinimido)- General cysteine with amino groups and histidine; adduct decomposes to S-succinyl cysteine and ethylamine upon acid hydrolysis Sequencing and Adduct stable and well resolved in S-aminopropyl cysteine amino acid analysis sequencing and amino acid analysis; introduces a positive charge Introduces trypsin Introduces a positive charge S-(2-aminoethyl)cysteine cleavage site Introduces a positive charge S-pyridylethyl cysteine Sequencing Cysteine-S-βSequencing PTH-derivative is well resolved propionamide Reversible Introduces negative charge; adduct S-sulfocysteine modification stable only at neutral and acidic pH Cysteine sulfonic acid Amino acid Quantitation of Cys in amino acid (cysteic acid) analysis analysis; destroys His, Tyr, Ser, Thr, Met, Trp; introduces negative charge Cystine Formation of Slow disulfide bonds S-carboxymethyl cysteine
aThe most frequent use of each is listed, but reagents are not necessarily restricted to this use alone. bN-iodoacetyl-N′-(5-sulfo-1-naphthyl)ethylenediamine.
The most common method for cleaving disulfide bonds is by reduction via a disulfide interchange between the protein cysteine disulfide and a sulfhydryl group of the reducing agent. Historically, 2-mercaptoethanol (2-ME) was used extensively, but its popularity has been eclipsed by the more efficient dithiothreitol (DTT). The efficiency of DTT is the result of the thermodynamically favored formation of an intramolecular disulfide ring structure within the reagent, which greatly shifts the equilibrium of the reaction toward complete reduction of the protein. Practically, this means that only slight excesses of DTT are necessary to achieve complete reduction; this can be meaningful for reactions using an expensive or radioactive alkylating agent. Aesthetically, even though DTT has a sulfurous odor, it does not compare with the volatile stench of 2-ME.
Modification of Cysteine
In the case of proteins, where secondary and tertiary structure may interfere with reagent access to the cysteine side chains, modification reactions are usually carried out in the presence of a protein denaturant such as 6 M guanidine⋅HCl or 8 M urea. In general, these denaturants can be used interchangeably and selection is often a matter of personal preference. However, urea solutions produce cyanate ions, which can carbamylate amino groups in proteins and peptides and thus block them to subsequent sequence analysis. If urea is used, it should always be treated with a mixed-bed ion-exchange resin to remove the cyanate just prior to its use. In addition, urea should be removed from the protein as
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soon as possible after the completion of the reaction. Of course, if carbamylation of free amino groups is not a concern, this precaution can be omitted. Also, the choice between urea and guanidine⋅HCl may be influenced by the choice of protease that may subsequently be used on the modified protein. This is particularly an issue when modifying very low quantities of protein with the intent to subsequently perform a proteolytic digestion. Certain proteases are active in lower concentrations of either urea or guanidine, so it may not be necessary to completely remove the denaturant. Instead, a simple dilution may be all that is necessary to proceed (see UNIT 11.1). Alternatively, cysteine modifications can be done with little or no denaturation to study free sulfhydryl content or the accessibility of free sulfhydryls in the native form of the protein. The choice of which reagent to use or which protocol to follow may depend to some extent on the purpose of the modification. For instance, the intent may be to modify cysteine to stabilize it specifically for amino acid compositional analysis or sequence analysis. Some cysteine adducts are more suitable than others for this purpose; Table 15.1.1 lists those that have been used most successfully. Alternatively, the intent may simply be to prevent disulfide bond reformation for peptide separation after proteolysis, in which case most reagents are suitable. When working with very low levels of protein, volume and sample manipulation must be kept to a minimum to avoid losses in handling. The protocols presented here are general procedures that can be adapted to a number of specific uses. All of them are very simple and easy to perform. Notes in the individual protocols indicate where one offers some advantage over another for a particular use (also see Critical Parameters and Troubleshooting and Table 15.1.1). A number of specialized protocols involving cysteine modification in conjunction with a specific application are presented elsewhere in this manual. Reduction and alkylation of peptides in digest mixtures is discussed in UNIT 11.1, and reduction and alkylation of proteins in acrylamide gels prior to digestion is presented in UNIT 11.3. ALKYLATION OF A PROTEIN OF KNOWN SIZE AND COMPOSITION WITH HALOACYL REAGENTS OR N-ETHYLMALEIMIDE
BASIC PROTOCOL 1
Reaction of the sulfhydryl group of cysteine with α-haloacids or their amide derivatives— iodoacetic acid, iodoacetamide, and N-iodoacetyl-N′-(5-sulfo-1-naphthyl)ethylenediamine (known as IAEDANS)—is historically the most common method used to alkylate cysteine. Use of iodo compounds is typical because iodide is a better leaving group (i.e., has faster reaction times) than the other halides. The choice between iodoacetic acid or iodoacetamide may be dictated by what subsequent studies are to be performed and, in some cases, the accessibility of the cysteine residues in the protein being modified (Gerwin, 1967). For instance, if the cysteine side chain is in a local charged environment on the surface of the protein, charge interactions with the acid form of the reagent may cause complications, and thus iodoacetamide may be a better choice. Modification using iodoacetic acid yields S-carboxymethyl cysteine, whereas iodoacetamide yields S-carboxamidomethyl cysteine. Both of these products are relatively stable in protein sequencing. The carboxymethyl group imparts a negative charge but the carboxamidomethyl group does not. Depending on the cysteine content of the protein of interest, the negative charge may alter the characteristics of the protein. Upon acid hydrolysis, S-carboxamidomethyl cysteine deamidates to S-carboxymethyl cysteine; this derivative is not as stable as some others to hydrolysis. Alkylation with IAEDANS introduces a fluorescent label that allows sensitive fluorometric detection following chromatography, SDS-PAGE (UNIT 10.1), or electroblotting from gels (UNIT 10.7).
Chemical Modification of Proteins
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N-ethylmaleimide (NEM) is a reagent containing a reactive double bond that has historically been used to modify cysteine residues and is still mentioned frequently in the literature. The advantages of NEM are that it is membrane permeable, it can be obtained in radioactive form, and the ethyl group can be replaced with other substituents, such as fluorescent adducts, to permit the introduction of reporter groups. Its disadvantages are that above pH 7 it is unstable and seems to have a higher propensity to react with amino groups and histidine. For these reasons, it is not recommended for routine use. If it is desirable for a particular application, however, NEM can be used in this protocol, but the reaction pH should be adjusted to 7.0. The adduct breaks down to S-succinylcysteine and ethylamine during acid hydrolysis. This protocol can be used to prepare material suitable for amino acid analysis, sequence analysis, and proteolytic digestion (UNITS 11.1 & 11.4). Materials Protein, lyophilized Tris/guanidine buffer, pH 8.0 (see recipe) or 0.1 M Tris⋅Cl, pH 8.0 50 mM dithiothreitol (DTT; 7.7 mg/ml) Nitrogen Alkylating agent (see recipe) 2-mercaptoethanol (2-ME; 14.4 M) 0.5- to 2.5-ml microcentrifuge tube Additional reagents and equipment for desalting modified protein (see Support Protocol 3) 1. Dissolve the protein in a minimum volume of Tris/guanidine buffer to give a protein concentration ≥100 µg/ml in a volume between 50 and 500 µl, if possible. Buffering capacity and pH are critical, so interference by other components in the protein sample must be considered and removed, if necessary, prior to modification. Guanidine⋅HCl may be omitted from the buffers (i.e., 0.1 M Tris⋅Cl may be used) if protein denaturation is not desired. Use Tris/guanidine buffer, pH 7.0, or 0.1 M Tris⋅Cl, pH 7.0, if NEM is used as an alkylating agent.
2. Add DTT to give a 10-fold molar excess over expected disulfides. To calculate the amount of DTT required, assume all cysteines are present as disulfides and use the following equation:
(µg protein) (nmol protein/µg protein) (nmol disulfide/nmol protein) = nmol protein disulfide For example, for 500 ìg of a 25,000-Da protein containing 8 cysteines the amount of disulfide is (500 ìg protein) (1 nmol protein/25 ìg) (4 nmol disulfide/nmol protein) = 80 nmol protein disulfide. A 10-fold molar excess of DTT requires
10 (nmol protein disulfide) = nmol DTT For the 25-kDa protein, this is 10 (80 nmol protein disulfide) = 800 nmol DTT. To calculate the amount of DTT solution required use the following equation:
(nmol DTT) (1 µl/nmol DTT in stock solution) = µl DTT For a 50 mM DTT stock solution, this becomes (800 nmol DTT) (1 ìl/50 nmol DTT) = 16 ìl of 50 mM DTT. Modification of Cysteine
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3. Flush nitrogen over the surface of the liquid and seal the microcentrifuge tube. Incubate 1 hr at 37°C. 4. Add alkylating agent to give a 5-fold molar excess of reagent over total thiols in solution. A 0.2 M stock solution of alkylating agent contains 200 nmol/ìl. Total thiols in solution includes both protein thiol and reductant thiol. DTT contributes 2 moles of thiol per mole of reagent. To obtain a 5-fold molar excess, the amount of alklyating agent required is
5 [(nmol DTT) (2 nmol SH/nmol DTT) + (nmol protein disulfide) (2 nmol SH/disulfide)] = nmol alkylating agent In the example here, the calculation is 5 [(800 nmol DTT) (2 nmol SH/nmol DTT) + (80 nmol protein disulfide) (2 nmol SH/disulfide)] = 8800 nmol. The volume of alkylating agent required is therefore
(nmol alkylating agent required) (1 µl/x nmol alkylating agent in stock solution) = µl alkylating agent For this example, with alkylating agent at 200 nmol/ìl, this is (8800 nmol) (1 ìl/200 nmol) = 44 ìl.
5. Flush nitrogen over the surface of the liquid and seal the microcentrifuge tube. Incubate 1 hr at 37°C in the dark. When iodinated reagents are used, exclusion of light is important to minimize the chance of formation of iodine from available iodide ions. As the reaction proceeds, it generates protons that will tend to make the solution more acidic if the buffer is not strong enough. If large amounts of protein (>5 mg) are being modified, it is advisable to monitor the pH and readjust to 8.0 as necessary.
6. Add ≥10-fold excess of 2-ME relative to the alkylating agent. A 10-fold excess of 2-ME requires 10 (8800 nmol alkylating agent) = 88 ìmol 2-ME, which corresponds to (88 ìmol) (1 ìl/14.4 ìmol) = 6 ìl. As long as the 2-ME is in molar excess relative to the modifying reagent, the volume of 2-ME added can be rounded up to the nearest microliter or most convenient pipetting volume. For instance, if 1.3 ìl is calculated, 2 to 5 ìl can be added without harm.
7. Desalt the alkylated protein (see Support Protocol 3). ALKYLATION OF A PROTEIN OF UNKNOWN SIZE AND COMPOSITION WITH HALOACYL REAGENTS OR N-ETHYLMALEIMIDE
ALTERNATE PROTOCOL 1
If the size and number of cysteines present in a protein are unknown follow procedure for a protein of known size and composition (see Basic Protocol 1), assuming that the protein is large (i.e., ∼100,000 Da) and contains a high amount of cysteine (i.e., 50 cysteines = 25 disulfides per molecule) for the calculations. ALKYLATION OF ≤50 ìg PROTEIN WITH HALOACYL REAGENTS OR N-ETHYLMALEIMIDE Reduction and alkylation of small amounts of protein requires the use of very small volumes of reagents based on the calculations in Basic Protocol 1. If <50 µg protein is being alkylated, whether or not the composition and size are known, excess reagents can be used at each step. Follow procedure as for larger amounts (see Basic Protocol 4) but use 2 µl of 50 mM DTT, 6 µl of 0.2 M alkylating reagent, and 2 µl of 2-ME.
ALTERNATE PROTOCOL 2
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BASIC PROTOCOL 2
ALKYLATION WITH 3-BROMOPROPYLAMINE Use of 3-bromopropylamine is especially well suited to the identification of cysteine in automated amino-terminal protein sequencing methods (Jue and Hale, 1993). The modified cysteine, as aminopropylcysteine, is stable and well resolved in both amino acid analysis and protein sequencing separations (Jue and Hale, 1994). Furthermore, 3-bromopropylamine does not cause preview in protein sequencing, which is a potential problem with 4-vinylpyridine (see Basic Protocol 4). Preview in protein sequencing is the appearance of an amino acid residue in a sequencing cycle just prior to the cycle in which it would normally be cleaved from the protein. Materials Protein, lyophilized 20 mM and 2 M dithiothreitol (DTT) in 0.5 M Tris⋅Cl (pH 8.2)/6 M guanidine⋅HCl 0.5 M 3-bromopropylamine (see recipe) Nitrogen 0.5- to 2.5-ml microcentrifuge tube Additional reagents and equipment for desalting modified protein (see Support Protocol 3) 1. Dissolve protein in a minimum volume of 20 mM DTT/0.5 M Tris⋅Cl/6 M guanidine⋅HCl. Enough reducing power for ∼200 nmol of –SH is provided by 100 ìl of 20 mM DTT/Tris⋅Cl/guanidine⋅HCl. This amount of –SH is comparable to 1 mg of a 50,000-Da protein containing 10 cysteines. Alternatively, the necessary amount can be calculated as described in the haloacid/NEM procedure (see Basic Protocol 1, step 2). If more reducing power is required, either use more volume or increase the concentration of DTT in the buffer.
2. Flush with nitrogen over the surface of the liquid and seal the microcentrifuge tube. Incubate 1 hr at 60°C. 3. Add 1⁄5 vol of 0.5 M 3-bromopropylamine. For example, if the protein is dissolved in a total of 100 ìl, add 20 ìl of 3-bromopropylamine.
4. Flush with nitrogen over the surface of the liquid and seal the microcentrifuge tube. Incubate 2 hr at 60°C in the dark. 5. Add 1⁄5 vol of 2 M DTT/0.5 M Tris⋅Cl/6 M guanidine⋅HCl. For example, if the protein is in a total of 120 ìl at this point, add 24 ìl of 2 M DTT/Tris⋅Cl/guanidine⋅HCl.
6. Desalt the modified protein (see Support Protocol 3).
Modification of Cysteine
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ALKYLATION WITH N-(IODOETHYL)-TRIFLUOROACETAMIDE Introduction of a primary amine by conversion of cysteine to S-(2-aminoethyl)-cysteine introduces a trypsin-sensitive cleavage site. Previously, this was accomplished using ethyleneimine, which is now restricted as a hazardous material. However, N-(iodoethyl)trifluoroacetamide has been shown to be a comparable substitute (Schwartz et al., 1980); this reagent is available from Pierce Chemical as Aminoethyl-8 Reagent.
BASIC PROTOCOL 3
Materials Protein, lyophilized 0.2 M N-ethylmorpholine acetate (pH 8.1)/6 M guanidine⋅HCl 50 mM dithiothreitol (DTT; 7.7 mg/ml) Nitrogen Aminoethyl-8 Reagent (Pierce Chemical) Methanol 2 N acetic acid 0.5- to 2.5-ml microcentrifuge tube Additional reagents and equipment for desalting modified protein (see Support Protocol 3) 1. Dissolve protein in a minimum volume of 0.2 M N-ethylmorpholine acetate (pH 8.1)/6 M guanidine⋅HCl. 2. Add DTT to a 10-fold molar excess over protein disulfide. Calculate the amount of 50 mM DTT required as described in the haloacid/NEM procedure (see Basic Protocol 1, step 2).
3. Flush with nitrogen and seal the microcentrifuge tube. Incubate 2 hr at room temperature. 4. Dissolve Aminoethyl-8 Reagent in a volume of methanol to give a 25-fold molar excess of reagent to total thiol groups and a final methanol concentration of 10% (v/v) relative to total reaction volume. To calculate the number of micrograms of reagent required use the following equation:
25[(µg protein)(nmol protein/µg protein)(nmol disulfide/nmol protein)(2 nmol SH/nmol disulfide) + (µl DTT added)(nmol DTT/µl)(2 nmol SH/nmol DTT)](0.267 µg reagent/nmol) = µg Aminoethyl-8 Reagent For example, for 500 ìg of a 25,000-Da protein containing 8 cysteines, the amount of reagent would be 25[(500 ìg protein)(1 nmol protein/25 ìg)(4 nmol disulfide/nmol protein)(2 nmol SH/nmol disulfide) + (16 ìl DTT added)(50 nmol DTT/ìl)(2 nmol SH/nmol DTT)](0.267 ìg reagent/nmol) = 11,748 ìg or 11.7 mg reagent. Dissolve the reagent in a volume of methanol equal to 0.11 × sample volume. If the volume of methanol required is too low to handle easily, increase the volume of the sample by adding 0.2 M N-ethylmorpholine acetate (pH 8.1)/6 M guanidine⋅HCl. Keep the final methanol concentration of the reaction at 10% (v/v). The solubility of Aminoethyl-8 Reagent in methanol is ∼450 mg/ml.
5. Add one-half of the Aminoethyl-8 Reagent/methanol solution. Incubate 1 hr at room temperature. 6. Add the remaining Aminoethyl-8 Reagent. Continue the incubation an additional 1 to 1.5 hr at room temperature. 7. Acidify reaction to ∼pH 5.5 with 2 N acetic acid. 8. Desalt modified protein (see Support Protocol 3).
Chemical Modification of Proteins
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BASIC PROTOCOL 4
ALKYLATION WITH 4-VINYLPYRIDINE Modification of cysteine with 4-vinylpyridine is simple and efficient. The resulting S-pyridylethyl cysteine derivative is stable to acid hydrolysis and also performs well in automated amino-terminal protein sequencing methods. However, there is evidence that some N-alkylation of the amino terminus of the protein may take place (Hempel et al., 1986). Upon reaction with phenylisothiocyanate (PITC), this adduct is unstable to the basic coupling conditions of the sequencing cycle, causing the alkylated amino acid to be released from the peptide and allowing the next amino acid to react with PITC. The result will be that the first cycle of sequencing can show some preview of the second residue and so on. The following protocol has been adapted from Applied Biosystems User Bulletin Number 28 (Hawke and Yuan, 1987). Materials 1 M Tris⋅Cl (pH 8.5)/4 mM EDTA 8 M guanidine⋅HCl Protein, lyophilized 10% (v/v) 2-mercaptoethanol (2-ME) Argon 4-vinylpyridine, fresh 0.5- to 2.5-ml microcentrifuge tube Additional reagents and equipment for desalting modified protein (see Support Protocol 3) 1. Mix 1 part 1 M Tris⋅Cl (pH 8.5)/4 mM EDTA to 3 parts 8 M guanidine⋅HCl to prepare the working buffer (0.25 M Tris⋅Cl/1 mM EDTA/6 M guanidine⋅HCl final). 2. Dissolve protein in <50 µl working buffer to a final concentration of 0.1 to 10 µg/µl and add 2.5 µl of 10% 2-ME. The procedure presented in this protocol will accommodate ≤500 ìg of protein, but it can be scaled up for larger amounts. The reaction volume should be kept to a minimum and the amounts of reductant, alkylating reagent, and quenching reagent should be increased proportionally to the amount of protein. For example, if 750 ìg of protein is used, reagent volumes should be increased by 50%.
3. Flush with argon and seal the microcentrifuge tube. Incubate 2 hr at room temperature. 4. Add 2 µl undiluted 4-vinylpyridine and mix. Only colorless 4-vinylpyridine that has been stored under argon at −20°C should be used.
5. Flush microcentrifuge tube with argon and seal the container. Incubate 2 hr at room temperature in the dark. 6. Either quench the reaction with 2.5 µl of 10% 2-ME and desalt, or desalt immediately with a rapid technique such as reversed-phase HPLC or membrane adsorption using a ProSorb cartridge (see Support Protocol 3).
Modification of Cysteine
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ALKYLATION WITH ACRYLAMIDE Although the use of acrylamide to alkylate cysteine is not widespread, it is a viable alternative. The cysteine-S-β-propionamide derivative is stable to automated amino-terminal sequencing and yields a readily identifiable phenylthiohydantoin (PTH)-cysteineS-β-propionamide. Use of acrylamide (Brune, 1992) may circumvent some problems associated with the use of 4-vinylpyridine (see Basic Protocol 4 and Commentary).
BASIC PROTOCOL 5
Materials Protein, lyophilized 0.3 M Tris⋅Cl, pH 8.3 50 mM dithiothreitol (DTT; 7.7 mg/ml; optional) 6 M acrylamide 10% (v/v) 2-mercaptoethanol (2-ME) 0.5- to 1.5-ml microcentrifuge tube Additional reagents and equipment for desalting modified protein (see Support Protocol 3) CAUTION: Acrylamide monomer is neurotoxic. A mask should be worn when weighing acrylamide powder. Gloves should be worn while handling the solution, and the solution should not be pipetted by mouth. 1. In a microcentrifuge tube, dissolve protein in a minimal volume (<100 µl, if possible) of 0.3 M Tris⋅Cl, pH 8.3, to give a protein concentration ≥100 µg/ml. 2. If reduction is desired, add DTT to yield a 10-fold molar excess relative to expected disulfides (see Basic Protocol 1, step 2). 3. Add acrylamide to a final concentration of 2 M. Incubate 2 hr at 37°C. Alternatively, 9.7 M dimethylacrylamide (available as a premade solution from Aldrich) can be added to a final concentration of 2 M. Dimethylacrylamide produces a more hydrophobic derivative that in turn produces a later-eluting adduct of PTH-Cys. Acrylamide and dimethylacrylamide can be used interchangeably; the choice is a matter of personal preference (Brune, 1992).
4. Either quench the reaction with 2.5 µl of 10% 2-ME and desalt, or desalt immediately with a rapid technique such as reversed-phase HPLC or membrane adsorption using a ProSorb cartridge (see Support Protocol 3). SULFITOLYSIS Sulfitolysis is a mild procedure for cleavage of protein disulfide bonds, and the resultant S-sulfocysteine is easily reduced back to native cysteine with sulfhydryl compounds, such as 2-mercaptoethanol (2-ME; Chan, 1968). Only disulfides are reactive, but cysteine in proteins can be modified by including catalytic levels of free cysteine (or other sulfhydryl compound), which will drive the reaction through formation of mixed disulfides. The S-sulfocysteine derivative is stable only at neutral or acidic pH. For these reasons, this procedure is useful only in a limited number of applications; it is suitable when subsequent removal of the modifying group is an issue. Materials Protein, lyophilized Sodium sulfite reagent buffer (see recipe) 4 M urea/1 M Tris⋅Cl, pH 7.5 2-mercaptoethanol (2-ME) 0.5- to 2.5-ml microcentrifuge tube
BASIC PROTOCOL 6
Chemical Modification of Proteins
15.1.9 Current Protocols in Protein Science
Supplement 3
Additional reagents and equipment for desalting modified protein (see Support Protocol 3) Cleave disulfide bonds 1. Dissolve protein in sodium sulfite reagent buffer to a final concentration of 2 mg/ml. Incubate 1 hr at 25°C in an open microcentrifuge tube. 2. Desalt modified protein (see Support Protocol 3). Reverse the modification 3. Dissolve the S-sulfocysteine protein in 4 M urea/1 M Tris⋅Cl, pH 7.5, to give a concentration of ∼1 mg/ml. 4. Add 2-ME to a final concentration of 0.7 M. Incubate 2 hr at 25°C. 5. Desalt the protein (see Support Protocol 3). BASIC PROTOCOL 7
OXIDATION WITH PERFORMIC ACID Modification of cysteine by performic acid oxidation is most commonly used in conjunction with amino acid compositional analysis to produce a derivative of cysteine (cysteic acid) that is stable to hydrolysis. The procedure is not recommended for modification of cysteine for purposes other than quantitation of cysteine by amino acid analysis. Conversion of cysteine to cysteic acid with this protocol is simple and quantitative. However, performic acid is a very strong oxidizer and several other amino acids such as histidine, tyrosine, tryptophan, methionine, serine, and threonine are adversely affected (Hirs, 1967). These other affected amino acids must therefore be quantitated by amino acid analysis of a separate aliquot. Analysis of the performic acid–treated sample can be normalized to the untreated sample by comparing the levels of stable amino acids such as aspartic acid (Asp + Asn) and glutamic acid (Glu + Gln). The following protocol is intended for samples to be used for amino acid analysis. CAUTION: This entire procedure should be performed in a fume hood. Materials Formic acid 30% hydrogen peroxide Protein, lyophilized and chilled 48% (v/v) hydrobromic acid Ice bath 1. Prepare fresh performic acid by adding 1 ml of 30% hydrogen peroxide to 9 ml formic acid. Mix and incubate 2 hr at room temperature. 2. Chill performic acid in an ice bath to 0°C. 3. To a chilled tube containing lyophilized protein, add sufficient chilled performic acid to thoroughly dissolve the protein. Incubate 1 hr on ice. 4. Add 0.2 ml hydrobromic acid. 5. Dry sample by lyophilization or Speedvac evaporation. The sample is now ready for hydrolysis for amino acid analysis.
Modification of Cysteine
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AIR OXIDATION TO A DISULFIDE Successful formation of the correct disulfide bonds in a protein or peptide is a critical step in achieving a biologically active product from chemical peptide synthesis or protein expression systems. The protocol presented here is very simplistic and the complexities of correct disulfide formation in proteins may require empirically derived combinations of denaturants and reducing agents (Noiva, 1994; Chau and Nelson, 1992; Zhang and Snyder, 1991). This protocol works best for small peptides such as those obtained from peptide synthesis procedures.
BASIC PROTOCOL 8
Materials Protein or peptide, lyophilized 0.1 M ammonium bicarbonate Additional reagents and equipment for reversed-phase HPLC (UNIT 11.6) and desalting oxidized protein (see Support Protocol 3) 1. Dissolve protein or peptide in 0.1 M ammonium bicarbonate. To reduce the likelihood of intermolecular disulfide formation, keep the protein or peptide concentration <0.25 to 0.5 mg/ml.
2. Gently stir several days with vessel open to air. 3. Monitor effects of oxidation by changes in elution position on reversed-phase HPLC relative to fully reduced starting product (more sensitive for smaller peptides). Alternatively, monitor the decrease in free sulfhydryl content colorimetrically with Ellman Reagent (see Support Protocol 2).
4. Desalt the oxidized protein (see Support Protocol 3). ALKYLATION OF CYSTEINE IN PROTEIN APPLIED TO SEQUENCER MEMBRANES
BASIC PROTOCOL 9
Modification of cysteine in samples applied directly to sequencer membranes can be employed as an alternative to modification in solution. This procedure minimizes sample loss by reducing the manipulation required for derivatization and sample cleanup. Although this procedure has been found to be generally applicable, there are instances where it has failed with certain proteins, for reasons that are not clear. If a negative result is obtained, alkylation in solution followed by application to a polyvinylidene difluoride (PVDF) membrane is a reasonable alternative (see Support Protocol 3). Tributylphosphine is used as a reductant in this procedure. Although its use is not widespread, this compound can be useful in cleavage of disulfide bonds for several reasons. First, only a slight excess (5% to 20%) over protein disulfide groups is necessary. Second, subsequent alkylation can take place in the presence of tributylphosphine without generating additional reaction by-products and with no allowance for thiols other than those associated with the protein. Finally, tributylphosphine is soluble in organic solvents, which makes it well suited to concomitant in situ reduction and alkylation immediately prior to protein microsequencing. The following protocol is adapted from a Perkin Elmer Applied Biosystems Division (PE-ABD) User Bulletin to prepare samples for sequencing with a PE-ABD sequencer (Hawke and Yuan, 1987). It can be modified for use with other sequencers by setting up a wash cycle similar to that used on the PE-ABD sequencer (described in step 4). CAUTION: Tributylphosphine is a hazardous reagent; proper precautions should be used when handling it.
Chemical Modification of Proteins
15.1.11 Current Protocols in Protein Science
Supplement 3
Materials Protein Reduction/alkylation cocktail (see recipe) Sequencer reagents (Perkin-Elmer or equivalent) Biobrene coated glass fiber disk, precycled PE-ABD sequencer, cartridge (Perkin-Elmer) or equivalent 1. Apply protein to a precycled Biobrene coated glass fiber disk in the usual manner for sequencing and dry. 2. Pipet 20 µl reduction/alkylation cocktail onto the glass fiber disk. 3. Immediately reassemble the cartridge. 4. Execute the 4-VP cycle: Deliver coupling buffer for 60 sec Pause 50 min Dry with argon 5 min Wash 5 min with butyl chloride Dry with argon 2 min Deliver coupling buffer for 1 min Dry with argon 2 min Wash with ethyl acetate for 5 min Dry with argon 2 min. The reaction takes place during the 50-min pause.
5. Start sequencing run.
SUPPORT PROTOCOL 1
RECRYSTALLIZATION OF IODOACETIC ACID Good quality iodoacetic acid (IAA) should be available from chemical companies such as Aldrich; IAA crystals should be white with no yellow tint. If the IAA has yellowed during storage, its use can result in iodination of the protein. The quality of the IAA can be significantly improved by recrystallizing it from petroleum ether until the yellow color is gone. CAUTION: Ether is very flammable. Use it in a fume hood away from flame. Materials Iodoacetic acid (IAA) Diethyl ether Petroleum ether, ice cold Buchner funnel Whatman No. 1 filter paper or equivalent Amber bottle with Teflon-lined lid 1. Dissolve 10 to 15 mg IAA in a minimal volume of diethyl ether (4 to 5 ml). Stir until the IAA dissolves. 2. Pour into a beaker containing 350 ml cold petroleum ether. 3. Collect crystals on a Buchner funnel with Whatman No. 1 filter paper.
Modification of Cysteine
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Current Protocols in Protein Science
4. Dry 24 hr under vacuum. If crystals are not snow-white, repeat steps 1 to 4 until no yellow tint remains.
5. Store crystals in a tightly sealed amber bottle with a Teflon-lined lid at −20°C. COLORIMETRIC QUANTITATION OF FREE SULFHYDRYLS Free sulfhydryl content can be quantitatively determined with Ellman reagent, 5,5′-dithiobis-(2-nitrobenzoic acid) from Pierce Chemical. Molar extinction coefficients of thionitrobenzoate, the colored species generated when the reagent reacts with a free thiol, are given for various solutions in Table 15.1.2.
SUPPORT PROTOCOL 2
The sensitivity of the reaction is in the low nmol/ml range for sulfhydryl groups. This method is often not sensitive enough for the amounts of protein available when proteins from natural sources are being analyzed, but it is well-suited to synthetic peptides or recombinant proteins that are usually available in larger quantities. Materials 0.1 M sodium phosphate, pH 8.0 (APPENDIX 2E) Reagent solution: 4 mg/ml Ellman reagent/0.1 M sodium phosphate, pH 8.0 Cysteine standard stock solution (see recipe) Unknown sample 13 × 100-mm clear test tubes 1. Prepare a cysteine standard curve by adding different amounts of cysteine standard stock solution to separate tubes—e.g., 25 µl, 50 µl, 100 µl, 150 µl, 200 µl, and 250 µl. Add ≤250 µl of each unknown to separate tubes. Bring the volume in each tube to a total of 250 µl with 0.1 M sodium phosphate, pH 8.0. Add 250 µl 0.1 M sodium phosphate, pH 8.0, to a blank tube. The cysteine content of an unknown should fall within the range of the standard curve (37.5 to 375 nmol). If a rough estimate of the cysteine content of an unknown cannot be made, at least two different amounts—e.g., 25 ìl and 250 ìl—should be tested.
2. Add 50 µl reagent solution and 2.5 ml of 0.1 M sodium phosphate, pH 8.0, to each tube. Mix and incubate 15 min at room temperature. 3. Measure absorbance at 412 nm. 4. Plot the A412 values obtained for the standards after subtracting the A412 value for the blank to derive a standard curve. 5. Use this curve to determine the free sulfhydryl content for the experimental samples.
Table 15.1.2 Molar Extinction Coefficients of Thionitrobenzoate in Various Solutionsa
Solvent
ε412
2% SDS 0.1 M phosphate (pH 7.27)/1 mM EDTA Buffered 6 M guanidine⋅HCl 6 M guanidine⋅HCl 8 M urea
12,500 14,150 13,700 13,880 14,290
aTaken from the Pierce Chemical technical bulletin for product number
22582.
Chemical Modification of Proteins
15.1.13 Current Protocols in Protein Science
Supplement 3
SUPPORT PROTOCOL 3
DESALTING THE SAMPLE AFTER CYSTEINE MODIFICATION Except for the in situ method (Basic Protocol 9), all of the protocols presented leave the protein in the presence of high salt and reaction by-products which are usually not compatible with subsequent analytical techniques. This problem may be remedied by one of the following desalting methods. The methods are presented in increasing order of technological complexity. Protein recovery is a concern because small quantities of protein may be lost in dialysis or gel filtration. Modern microanalytical techniques typically work with <100 pmol protein, which can easily disappear due to nonspecific binding to dialysis tubing, gel-filtration columns, and ultrafiltration devices. The most commonly applied desalting techniques currently used for picomole quantities of proteins are ProSpin and ProSorb cartridges (Perkin-Elmer) for direct sequence analysis, or reversed-phase HPLC (UNIT 11.6) using narrow or microbore columns. Dialysis. Dialysis is well suited to large-scale (milligram quantity) desalting needs. The protein is simply placed in dialysis tubing of an appropriate molecular weight cut-off (MWCO) such that the protein will be retained. The tubing is placed in a large vessel of appropriate buffer and gently stirred. After two or three changes of buffer over 24 to 48 hr, the composition of buffer in the tubing is now effectively the same as in the large vessel, due to the exchange of low-molecular-weight salts and other small molecules through the dialysis tubing. The volume inside the tubing will increase, so it is necessary to allow for this by leaving additional space in the dialysis tubing when the protein sample is added. Dialysis may be followed by lyophilization or Speedvac evaporation to concentrate the sample. Some brands of dialysis tubing require special preparation before use. For more information on dialysis, see UNIT 4.4 & APPENDIX 3B. Gel filtration (size-exclusion chromatography). Like dialysis, this method is better suited to large-scale desalting and is relatively inexpensive. It can be performed as a low-pressure bench-top technique using media such as Sephadex G-15 or G-25 resins or as a high-pressure liquid chromatography application (in which case it can be more readily done on a microscale). As with dialysis, buffer exchange may be accomplished with gel filtration, but subsequent concentration of the sample may be necessary. See UNIT 4.4 and UNIT 8.3 for additional details. Centrifugal filtration. Centrifugal filtration with membrane cartridges (Amicon; UNIT 4.4) combines the buffer exchange of dialysis with a concentration step by using centrifugation to force liquid through a membrane of appropriate MWCO. The protein is retained, may be repeatedly washed, and is ultimately recovered in a very small volume. When routinely desalting the same protein, a Centricon cartridge can be reused; this can potentially increase recovery of protein as nonspecific binding sites become saturated. Adsorption to a membrane. This method is well suited to samples that will subsequently be microsequenced because the resulting protein-containing polyvinylidene difluoride (PVDF) membrane can be placed directly in the reaction cartridge of a protein sequencer. ProSpin and ProSorb cartridges (Perkin-Elmer) are useful for this procedure. ProSpin uses centrifugation to force liquid through the PVDF membrane. ProSorb, a more recent development that has superseded ProSpin, uses an absorptive process to transfer protein to the membrane. Both cartridges require a homogeneous protein solution because the PVDF membrane will bind all protein present. Refer to manufacturer’s instructions for procedures with ProSpin and ProSorb cartridges.
Modification of Cysteine
Reversed-phase HPLC (RP-HPLC). This method is the most technologically complex, because it requires the use of a complete high-pressure liquid chromatography (HPLC)
15.1.14 Supplement 3
Current Protocols in Protein Science
system. For microscale work, the HPLC system should be configured for narrow-bore (2-mm) or micro-bore (1-mm) work because larger column diameters are better suited to milligram quantities of protein. The most common elution method uses a two-component system with 0.1% (v/v) trifluoroacetic acid in water (buffer A) and 0.1% (v/v) trifluoroacetic acid in acetonitrile (buffer B). Protein is detected by ultraviolet absorbance at 210 or 280 nm. The protein is acidified and injected onto a column that has been equilibrated with 100% buffer A, then eluted either isocratically with 80% buffer B or with an increasing gradient of buffer B. The column eluant is monitored and the appropriate fraction(s) are pooled, if necessary, and exposed to a jet of nitrogen or argon gas to evaporate the majority of the trifluoroacetic acid and acetonitrile phase. The remaining, predominantly aqueous, phase is lyophilized or vacuum concentrated. When working at narrow or microbore scale, the recovered fraction volumes are small enough (<100 µl) that they may be directly loaded onto a protein sequencer sample support. Ideally desalted samples should be lyophilized and stored in a freezer in the presence of a dessicant. Alternatively, they can be stored frozen in the dissolved state. REAGENTS AND SOLUTIONS Use Milli-Q water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Alkylating agent Prepare buffer for alkylating agent by adding 1 g guanidine⋅HCl to 1 ml 0.95 M Tris⋅Cl, pH 8.0 (0.5 M Tris⋅Cl and 6 M guanidine⋅HCl final). To this buffer add one of the following alkylating agents to a final concentration of 0.2 M: Iodoacetic acid (FW 186) Iodoacetamide (FW 185) N-iodoacetyl-N′-(5-sulfo-1-naphthyl)ethylenediamine (IAEDANS, FW 434) N-ethylmaleimide (FW 125) A concentration of 0.2 M provides 200 nmol alkylating agent/ìl. If N-ethylmaleimide is used as the alkylating agent, prepare the buffer with 0.95 M Tris⋅Cl, pH 7.0.
3-Bromopropylamine, 0.5 M 0.5 M 3-bromopropylamine 0.5 M Tris⋅Cl, pH 8.2 6 M guanidine⋅HCl Prepare fresh Cysteine standard stock solution, 1.5 mM Dissolve 26.34 mg cysteine hydrochloride monohydrate in 100 ml 0.1 M sodium phosphate, pH 8.0, immediately before use. Reduction/alkylation cocktail 2 µl tri-n-butylphosphine 5 µl 4-vinylpyridine 100 µl acetonitrile Prepare fresh Sodium sulfite reagent buffer 0.05 M sodium sulfite 0.2 mM cysteine 8 M urea or 6 M guanidine⋅HCl 0.1 M Tris⋅Cl, pH 8.4 Prepare fresh
Chemical Modification of Proteins
15.1.15 Current Protocols in Protein Science
Supplement 3
Tris/guanidine buffer 1 g guanidine⋅HCl (6 M final) 1 ml 0.19 M Tris⋅Cl, pH 8.0 (0.1 M final) Store at 25°C For N-ethylmaleimide, use Tris⋅Cl, pH 7.0.
COMMENTARY Jensen, 1989; Hoogerheide and Campbell, 1992), these methods are omitted here, for several reasons. Unlike alkylations, which are irreversible, mixed disulfide formation is freely reversible and the disulfides may thus be lost during subsequent manipulations. In addition, unless subsequent conditions are carefully controlled, disulfide scrambling may occur once the excess disulfide reagent is removed. Moreover, many of the reagents that have been reported in the literature for this purpose are not commercially available and must be synthesized prior to use. Thus, modification by mixed disulfide formation does not usually lend itself well to routine modification. However, the procedures for use of mixed disulfides in amino acid analysis (Barkholt and Jensen, 1989; Hoogerheide and Campbell, 1992) use commercially available reagents and appear to be quite accurate.
Background Information Cysteine is the most reactive residue commonly found in proteins and peptides. The thiol group, in its unprotonated state, is strongly nucleophilic and reacts with a wide variety of reagents. In addition to the free thiol, cysteine also occurs in disulfide linkage where it is relatively unreactive. The methods most commonly used to modify cysteines involve alkylating reagents, including active halogen compounds (Basic Protocols 1, 2, and 3) and compounds containing reactive double bonds (Basic Protocols 1, 4, 5, and 9). Cysteine thiols can also be oxidized either with stong oxidizing agents such as performic acid (resulting in conversion of the thiol to a sulfonic acid group; Basic Protocol 7) or with weak oxidizers such as oxygen (resulting in formation of disulfides; Basic Protocol 8; see Fig. 15.1.1). Finally, cysteine residues can form mixed disulfides with other thiol reagents. Although the formation of mixed disulfides has been used to modify protein thiols (Wynn and Richards, 1995) and to provide stabilizing adducts in amino acid analysis (Barkholt and
Na2SO3
S
Critical Parameters and Troubleshooting With few exceptions, disulfides generally need to be reduced to free thiols before they
S
(BP6) reducing agent
S
S
SO–3
SO 3–
O2
pH 8
strong oxidizer
(BP7)
(BP8) Na2SO3 +RSH S
–
S
SH
SH
+
+RSH alkylating agent (R-X) (BP1-5, 9)
Modification of Cysteine
SO 3–
–
–H
SR
SO 3–
+H
+
SR
Figure 15.1.1 Modification of cysteine. Interrelationship of the cysteine-modifying reactions described in this unit. Abbreviations: BP, Basic Protocol; R, alkyl group.
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can be derivatized. In addition to cysteine, proteins and peptides contain other reactive groups that, under the right conditions, can also react with many of the reagents used to derivatize cysteine. These groups include lysine ε-amino groups and the imidazole nitrogens of histidine, both of which are nucleophilic in the unprotonated state, and methionine, which is a pHindependent nucleophile. However, derivatization of cysteine can be made highly specific because of its pK and high reactivity. For instance, at pH 8, the thiol group, which has a pK of 8.3, has a significant population of unprotonated and thus nucleophilic species, but other potential nucleophiles are either still largely protonated (lysine ε-amino group) or react so slowly compared to the thiolate anion that they remain essentially unreacted in the time it takes for complete modification of cysteine (histidine and methionine). Thus, pH and duration of the reaction are critical factors in performing cysteine modifications. If residues other than cysteine become alkylated, repeat the experiment reducing the time of alkylation and check that the pH is correct and stable during the alkylation reaction. Any procedure that utilizes sulfhydryl-containing compounds to reduce disulfides prior to modification must take these sulfhydryl reagents into account when an alkylating reagent is added. The added sulfhydryl-containing compounds will also react with the alkylating reagent; in cases where the alkylating reagent is not in sufficient excess, they will consume the reagent and make it unavailable for the intended cysteine modification, resulting in only partial modification. Thus, it is essential to ensure that the alkylation reagent is in excess of the total thiol content of the sample. Once the alkylation reaction is completed, any excess alkylating reagent that remains unreacted should be destroyed before proceeding. Conditions can change significantly with subsequent sample manipulation, and this may allow remaining alkylating reagent to react in an unwanted manner with other components of the protein. Moreover, if halogen-containing reagents are used, halide ions are formed as a product of the alkylation reaction, and prolonged exposure to halide ions may result in halogenation of the protein. Occasionally specific cysteine residues will be resistant to alkylation with particular reagents. This is more apt to occur if the protein is not denatured prior to alkylation. For example, if the local environment of a protein thiol is charged, iodoacetic acid may not be as effec-
tive as an uncharged reagent such as iodoacetamide. If certain cysteine residues seem to be resistant to alkylation and a denaturing agent is being used, employing a different alkylating reagent may be helpful.
Anticipated Results Alkylation of cysteine residues is generally rapid and specific if the reaction conditions are controlled and care is taken to assure that correct ratios of reagents are employed. Thus, rapid and selective modification of cysteine residues in high yield (>90%) can generally be expected.
Time Consideratons
Allow ∼30 to 60 min for buffer and reagent preparation. With the exception of air oxidation of cysteines to disulfides (Basic Protocol 8), each protocol should take 2 to 4 hr. Production of disulfides by air oxidation is a slow process and will take from 1 to 7 days for completion. Recrystallization of iodoacetic acid (Support Protocol 1) takes 2 days. Assaying free sulfhydryls with Ellman reagent can be completed in 3 to 4 hr. The time required to desalt the sample after modification (Support Protocol 3) depends on which procedure is used and may also depend on the volume of sample: dialysis requires 24 to 48 hr; gel filtration requires 6 to 24 hr; centrifugal filtration requires 2 to 8 hr; adsorption to a membrane requires 1 to 6 hr; and HPLC requires 1 to 3 hr.
Literature Cited Barkholt, V. and Jensen, A.L. 1989. Amino acid analysis: Determination of cysteine plus halfcystine in proteins after hydrochloric acid hydrolysis with a disulfide compound as additive. Anal. Biochem. 177:318-322. Brune, D.C. 1992. Alkylation of cysteine with acrylamide for protein sequence analysis. Anal. Biochem. 207:285-290. Chan, W. 1968. A method for the complete S-sulfonation of cysteine residues in proteins. Biochemistry. 12:4247-4253. Chau, M.-H. and Nelson, J.W. 1992. Cooperative disulfide bond formation in apamin. Biochemistry. 31:4445-4450. Gerwin, B.I. 1967. Properties of the single sulfhydryl group of streptococcal proteinase. A comparison of the rates of alkylation by chloroacetic acid and chloroacetamide. J. Biol. Chem. 242:251. Hawke, D. and Yuan, P. 1987. S-Pyridylethylation of cystine residues. Applied Biosystems User Bulletin Issue No. 28.
Chemical Modification of Proteins
15.1.17 Current Protocols in Protein Science
Supplement 3
Hempel, J., Nilsson, K., Larsson, K., and Jörnvall, H. 1986. Internal chain cleavage and product heterogeneity during Edman degradation of isosteric peptide analogs lacking the carbonyl function. FEBS Lett. 194:333-337. Hirs, C.H.W. 1967. Performic acid oxidation. Methods Enzymol. 11:197-199. Hoogerheide, J.G. and Campbell, C.M. 1992. Determination of cysteine plus half-cystine in protein and peptide hydrolysates: Use of dithioglycolic acid and phenylisothiocyanate derivatization. Anal. Biochem. 201:146-151. Jue, R.A. and Hale, J.E. 1993. Identification of cysteine residues alkylated with 3-bromopropylamine by protein sequence analysis. Anal. Biochem. 210:39-44. Jue, R.A. and Hale, J.E. 1994. Alkylation of cysteine residues with 3-bromopropylamine: Identification by protein sequencing and quantitation by amino acid analysis. In Techniques in Protein Chemistry V (J.W. Crabb, ed.) pp. 179-188. Academic Press, San Diego.
Noiva, R. 1994. Enzymatic catalysis of disulfide formation. Protein Expression Purif. 5:1-13. Schwartz, W.E., Smith, P.U., and Garfield, P.R. 1980. N-(β-iodoethyl) trifluoroacetamide: A new reagent for the aminoethylation of thiol groups in proteins. Anal. Biochem. 106:43-48. Wynn, R. and Richards, F.M. 1995. Chemical modification of protein thiols: Formation of mixed disulfides. Methods Enzymol. 251:351-356. Zhang, R. and Snyder, G.H. 1991. Factors governing selective formation of specific disulfides in synthetic variants of α-Conotoxin. Biochemistry 30:11,343-11,348.
Contributed by Mark W. Crankshaw and Gregory A. Grant Washington University School of Medicine St. Louis, Missouri
Modification of Cysteine
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Modification of Amino Groups
UNIT 15.2
This unit describes group-specific modifications of amino groups. These reactions remain valid tools for early-stage evaluation of structure-function relationships, but are now valued even more for their applications in the preparation of bioconjugates, affinity columns, biosensors, and tagged macromolecules. Despite this shift in emphasis, a practical acquaintance with the classical amino group modifications (see Table 15.2.1) remains the key to a valuable toolbox of protein biochemistry methods. The amino groups of a polypeptide chain may include one N-terminal α-amino group, if it is not blocked, and any number of ε-amino groups of lysine. When the N-terminus is blocked by cell-mediated acetylation, cyclization of N-terminal Gln to pyroglutamate, or any other less common mechanism, it is inaccessible to chemical modification. Side-chain amino groups may also undergo post-translational modification, as in the ε-N,N,Ntrimethylation of Lys-115 in calmodulin, but they are unmodified and available for chemical modification in the great majority of cases. With the exception of N-terminal Pro, the free amino groups of proteins are primary amines. All of these groups have broadly similar reactivities, but there is a major difference in the typical pKa values for α-amino groups (pKa ∼8) and ε-amino groups (pKa ∼10). Selective modification of the N-terminus by careful selection of the reaction pH is an interesting route to site-specific tagging, but the usual numerical preponderance of ε-amino groups partly offsets the higher relative reactivity of the α-amino group. For reasons unconnected with pKa, amino-terminal residues of proteins also participate in several specialized reactions that are not available to lysine side chains (see Table 15.2.2). The most important reactions of amino groups are acylations, which yield amide (protein-NH-CO-R) or, less importantly, sulfonamide (protein-NH-SO2-R) products (where R indicates a group added to the protein by formation of a new bond). Other valuable modifications are based on nucleophilic substitution, nucleophilic addition, and reductive alkylation. In all cases, the amino group acts as a nucleophile that attacks an electrophilic center to acquire a modifying group of atoms. In particular, the reactions of amino groups with succinimidyl esters (see Basic Protocol 1) and isothiocyanates (see Basic Protocol 2) are broadly useful for the stable coupling to proteins of groups with useful, non-native functional properties. These include biotin for detection or recovery, fluorescent groups for biophysics or cytochemistry, cross-linking reagents for making bioconjugates, or metal-chelators that permit proteins to be loaded with radioisotopes for medical imaging or antitumor therapy. These applications require accurate product characterization, which preferably is performed by mass spectrometry (see Support Protocol). Succinic or acetic anhydrides may be used to change the charge state of protein amino groups (see Basic Protocol 3); reductive alkylations leave their charge unaltered but convert primary amines to secondary or tertiary amines (see Basic Protocol 4). STRATEGIC PLANNING Cysteine is most often modified for analytical purposes (UNIT 15.1), because its unmodified form is not reliably detected in amino acid analysis or Edman sequencing. The native structure of the protein is expendable in these applications, and denaturants are commonly used to ensure that sulfhydryls (which may otherwise be buried) are exposed to the reagents. Amino group modification has quite a different history. Because they are reactive and accessible, amino groups were and are often modified under native conditions to assess their contributions to protein function. Today, however, they are increasingly modified
Chemical Modification of Proteins
Contributed by Kieran F. Geoghegan
15.2.1
Current Protocols in Protein Science (1996) 15.2.1-15.2.18 Copyright © 2000 by John Wiley & Sons, Inc.
Supplement 4
Supplement 4
15.2.2
Current Protocols in Protein Science
Arylation; adds a chromophore and can be used to measure free amino groups
Addition by phenylthiocarbamylation of amino groups; couples a strongly fluorescent label to the protein
2,4,6-Trinitrobenzenesulfonic acid (TNBS)
Fluorescein isothiocyanate (FITC)
Alternative reagent for tagging with fluorescein; increasingly popular alternative to FITC
Succinylation; renders positively charged amino groups negative
Succinic anhydride
Fluorescein succinimidyl ester
Acetylation; renders positively charged amino groups neutral
Application
Structure of reagent
Some Common Reagents for Modification of Amino Groups
Acetic anhydride
Reagent
Table 15.2.1
Structure of modified amino groupa
2,4-Dinitrobenzenesulfonic acid (reacts very slowly); 2,4dinitrofluorobenzene
Rhodamine isothiocyanate
Hirs (1967); Fields (1972)
Banks and Paquette (1995)
continued
5-Carboxytetramethylrhodamine succinimidyl ester
Maleic anhydride (Butler and Hartley, 1972); citraconic anhydride (Atassi and Habeeb, 1972); both are reversible by low pH, but modification with citraconic anhydride is more easily reversible
Klotz (1967)
Banks and Paquette (1995)
Acetylimidazole (reacts primarily with phenolic groups of tyrosine, but has significant side reaction with amino groups); iodoacetic anhydride (see Table 15.2.2)
Related reagents
Means and Feeney (1971)
Reference(s)
15.2.3
Current Protocols in Protein Science
Supplement 4
An imidoester; modifies amino groups to a strongly basic form
Cross-linker; a homobifunctional imidoester
Methyl acetimidate hydrochloride
Dimethylsuberimidate dihydrochloride
Reacts with an amino group to form a sulfonamide that is stable even to acid hydrolysis conditions
Dimethylaminonaphthylsulfonyl chloride (dansyl chloride)
aP, protein molecule.
Alkylates the amino group once or twice; amino group remains capable of being protonated
Aldehyde/ NaBH3C0N
Disuccinimidyl suberate Cross-linker; a homobifunctional succinimidyl ester
Application
Structure of reagent
Structure of modified amino groupa
Some Common Reagents for Modification of Amino Groups, continued
Reagent
Table 15.2.1
Jentoft and Dearborn (1983)
Tae (1983)
Inman et al. (1983)
Reference(s)
Bis-(sulfosuccinimidyl)suberate is a more water-soluble alternative
Related cross-linkers differ by length of alkyl chain, e.g. dimethyladipimidate; spacer chain may be thiol-cleavable, e.g., dimethyl-3,3’-dithiobispropionimidate
Related reagents
Supplement 4
Selectively oxidizes N-terminal Ser or Thr to an α-N-glyoxylylresidue
Converts the N-terminal residue to an α-oxo-acyl moiety
Introduces a sulfhydryl-reactive iodoacetyl group at the amino terminus
NaIO4
Metal-catalyzed transamination
Iodoacetic anhydride used at pH 6
aR, side chain of the N-terminal residue.
Application
I-CH2-CO-HN-protein
(I-CH2-CO)2O
Wetzel et al. (1990)
Dixon and Fields (1972); Gaertner and Offord (1996)
O=CR-CO-HN-proteina
Metal [e.g., Ni(II)], Base (e.g., acetate), and amino acceptor (glyoxylate)
Reference Dixon and Fields (1972); Geoghegan and Stroh (1992); Alouani et al. (1995); Zhang and Tam (1996)
− 4
Structure of modified amino group O=CH-CO-protein
IO
Reagent
Modification Reactions Specific for the Amino Terminus
Reagent
Table 15.2.2
Modification of Amino Groups
15.2.4
Current Protocols in Protein Science
Creates a chemically unique site in the protein for conjugation with sulfhydryl-containing modifying groups; has applications for cyclizing peptides
Replaces the α-amino group with carbonyl oxygen; procedure considered prone to side reactions; when successful, can be used as the locus for further chemical modifications
Creates a chemically unique site in the protein for conjugation with modifying group; adducts described include hydrazones, reduced hydrazones (alkyl hydrazides), oximes, and thiazolidines
Comments
for preparative purposes, as they make convenient attachment points for the nonpeptide elements of molecular hybrids. Reactions done for either purpose are subject to the same issues at the planning stage. Key points to consider when preparing for such modifications are the reaction conditions, the extent of reaction desired, the method for adding reagents, and the technique for assessing results. ε-Amino groups are largely protonated at pH <9, but it is their unprotonated form that is chemically reactive: protein-NH3+
−H+
protein-NH 2
ZZZZZ X YZZZZ + Z protonated and unreactive
+H
unprotonated and reactive
Despite this, it is not necessary to set the reaction buffer at a very basic pH (e.g., pH 11) to ensure that all the amino groups in the protein are unprotonated. Doing so could be disastrous, in fact, because many modification reagents are labile to alkaline hydrolysis and proteins themselves undergo alkali-driven degradative reactions. It is only necessary to set and maintain a pH at which the amino groups are unprotonated to an extent that permits a reasonable reaction rate. This is usually pH 8 to 9 for reactions of protein amino groups with important reagents such as succinimidyl esters, sulfonyl halides, carboxylic acid anhydrides, and isothiocyanates. As unprotonated amino groups are consumed by the reaction, the original equilibrium distribution between protonated and unprotonated amino groups is continuously regained, so that all the amino groups can eventually be modified if desired. Elevated temperatures are hardly ever needed for the reactions considered here. A generous range of suitably reactive modifiers exists for amino groups, and most work can be done in the range of 0° to 22°C. The extent of modification required is a function of the application. When coupling biotin to a protein using a sulfosuccinimidyl ester, the most desirable result might be the minimum extent of modification consistent with biotinylating a large fraction of the protein molecules. Other applications call for saturation of all amino groups with the modifying group. Whenever possible, a pilot experiment should be done in which small amounts of protein are exposed to a range of treatments with varied levels of modifier to assess the outcome. Reagents must be added in a way that preserves their reactivity as well as respecting the sensitivity of the protein to elevated levels of organic solvent and pH. For example, ester reagents are base-labile and should be protected from exposure to alkaline conditions until they are added to the protein. As to the means by which the result is evaluated, fast electrophoretic methods are excellent tools for this purpose; optical spectroscopy may give instant information regarding the introduction of a chromophore; and mass spectrometry (see Chapter 16) offers the greatest accuracy at the cost of more complexity and the need for expensive equipment. AMIDATION USING A SUCCINIMIDYL ESTER It is often useful to add a non-native functional property to a protein by modifying it with biotin, a fluorescent group, a metal chelator, or other selected group. The most versatile and reliable methods for this purpose use succinimidyl ester reagents. When water-solubility of the reagent needs to be enhanced or ensured, a sulfosuccinimidyl ester may be used instead. N-hydroxysuccinimide or N-hydroxysulfosuccinimide, each a good leaving group, is displaced from the carboxylic reagent by the attack of a protein amino group. The bond formed between the modifying reagent and the protein is a stable amide (see Fig. 15.2.1).
BASIC PROTOCOL 1
Chemical Modification of Proteins
15.2.5 Current Protocols in Protein Science
Supplement 4
A O
O O
protein NH2
N O C "tag"
O protein NH C "tag"
O protein
N OH O
succinimidyl ester
modified protein N –hydroxysuccinimide
B –O S 3
protein NH2
O O N O C "tag" O
protein
sulfosuccinimidyl ester
O
–O S 3
protein NH C "tag"
O N OH O
modified protein N –hydroxysulfosuccinimide
Figure 15.2.1 General representation of the modification of a protein amino group by (A) a succinimidyl ester and (B) a sulfosuccinimidyl ester. Sulfosuccinimidyl esters are more water-soluble than the succinimidyl esters of the same acids. The “tag” denoted in the figure represents the group added to the protein and may take many forms.
Numerous reagents from this group are commercially available from vendors such as Pierce and Molecular Probes (see SUPPLIERS APPENDIX), whose catalogs are useful resources; a concise review of these reagents was provided by Brinkley (1993). Biotin succinimidyl esters that include spacer arms of varying lengths are popular reagents for the production of biotinylated proteins; fluorescent reagents such as 5-carboxytetramethylrhodamine succinimidyl ester and fluorescein succinimidyl ester are convenient for conferring visible-range fluorescence on a protein. Some reactive media designed for protein immobilization are based on succinimidyl esters—e.g., Affi-Gel 10 (Bio-Rad) and HiTrap NHS Sepharose (Pharmacia Biotech). Materials 0.1 to 10 mM succinimidyl ester reagent in acetonitrile, dimethylformamide (DMF), or dimethylsulfoxide (DMSO); or 0.1 to 10 mM sulfosuccinimidyl ester reagent in H2O Protein solution: 0.1 to 2 mM protein in 0.1 M sodium bicarbonate, pH not adjusted Additional reagents and equipment for gel-filtration chromatography (UNIT 8.3) or dialysis (UNIT 4.4 and APPENDIX 3B) and for mass spectrometry (see Chapter 16) 1. Add succinimidyl ester or sulfosuccinimidyl ester reagent to the protein solution ensuring a ≥2:1 molar excess of succinimidyl ester over the number of amino groups to be modified. Incubate ≥1 hr at room temperature. If the reagent has been dissolved in an organic solvent, before preparing the reaction mixture, use a small amount of the protein solution to test that the protein is soluble in the partly organic reaction mixture. Modification of Amino Groups
If made up in acetonitrile, DMF, or DMSO, these reagents are stable; if made up in water, they will be more stable than if made up in the basic reaction buffer.
15.2.6 Supplement 4
Current Protocols in Protein Science
2. Remove excess reagent and by-products from the protein by gel-filtration chromatography (UNIT 8.3), dialysis (UNIT 4.4 and APPENDIX 3B), or HPLC. See UNIT 15.1 for additional advice on a number of these methods. Microconcentrators (e.g., Centricon, Amicon) are useful for rapidly reducing sample volumes and exchanging buffer; however, these devices may need to be treated with polyethylene glycol to minimize protein adsorption (see manufacturer’s instructions).
3. Evaluate the extent of modification by mass spectrometry (see Chapter 16), UV-visible absorption spectroscopy (if the modifying group is chromophoric), or another suitable method as dictated by the nature of the modifying group. ADDITION OF FLUORESCEIN ISOTHIOCYANATE Fluorescence-activated cell sorting is a central technique of cell biology and frequently requires the use of antibodies that carry a fluorescent tag. Fluorescein isothiocyanate (FITC) is the reagent most commonly used to provide antibodies with the requisite fluorescence. The isothiocyanate functional group (–N=C=S, in which the carbon atom is the target for nucleophilic attack) readily reacts with amino groups at pH 9.0 to form phenylthiocarbamyl adducts that are stable at pH 7.0 (see Figure 15.2.2).
BASIC PROTOCOL 2
The following procedure is based on recommendations supplied by Molecular Probes for use with amine-reactive probes in general. Materials Protein, lyophilized or as a 5 to 20 mg/ml solution in 0.1 M sodium bicarbonate, pH 9.0 0.1 M sodium bicarbonate, pH 9.0 (pH adjusted with sodium hydroxide) Fluorescein isothiocyanate (FITC) Dimethylformamide (DMF) or dimethylsulfoxide (DMSO) 1.5 M hydroxylamine⋅HCl, pH 8.5, prepared fresh Gel-filtration column containing chromatography medium (e.g., PD-10 column containing Sephadex G-25; Pharmacia Biotech), equilibrated in PBS (APPENDIX 2E) 1. If necessary, dilute or dissolve protein to 5 to 20 mg/ml in 0.1 M sodium bicarbonate, pH 9.0. 2. Dissolve FITC at 10 mg/ml in DMF or DMSO, sonicating or vortexing briefly if necessary.
protein NH2
protein NH C S NH
S C N pH 9.0 COOH
HO
O
O
fluorescein isothiocyanate (FITC)
COOH
HO
O
O
fluoresceinated protein (may be modified at multiple amino groups)
Figure 15.2.2 Modification of a protein amino group with fluorescein isothiocyanate.
Chemical Modification of Proteins
15.2.7 Current Protocols in Protein Science
Supplement 4
3. Slowly add 50 to 100 µl FITC per milliliter of protein solution while gently shaking the tube to mix. Incubate 1 hr at room temperature. 4. Stop the reaction by adding 0.1 ml of 1.5 M hydroxylamine⋅HCl, pH 8.5, per milliliter of reaction mixture. This provides excess amino groups to quench residual FITC.
5. Pass the reaction mixture over a gel-filtration column equilibrated with PBS. Dye-conjugated protein should emerge before all low-mass components, and should be stored at −70°C or used immediately. BASIC PROTOCOL 3
SUCCINYLATION At acidic, neutral, or mildly basic pH (5. This modification can radically alter the solubility of a protein or its ability to participate in intermolecular associations. Klotz (1967) provides a practical guide to using succinic anhydride and related reagents (e.g., S-acetylmercaptosuccinic anhydride). The protein is dissolved in a buffer of pH 8.0 to 9.0 that contains no potentially reactive amine. Tris must not be used, because it is a primary amine that will compete with protein amino groups for the anhydride. Buffers made with HEPES (N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid), borate, or bicarbonate (see Table 15.2.3) are better choices. Succinic anhydride may be added as a solid, and its reactions with the protein take place in competition with its hydrolysis to succinic acid. The reaction with amino groups (see Fig. 15.2.3) predominates, but tyrosine side chains may also be modified at the phenolic oxygen to a derivative that is stable at pH <5, and therefore may be present if the product of the modification reaction is examined directly by HPLC performed with solvents containing 0.1% trifluoroacetic acid. If this occurs or is feared to have occurred, the protein may then be treated with 1 M hydroxylamine⋅HCl, pH 7.0, ≥1 hr at 20°C to remove the modifying group from tyrosine. Amino groups remain modified under these conditions. Side reactions of succinic acid also occur with other amino acid side chains (Klotz, 1967), but these are so readily reversible that they are of little general importance. Other types of modifications chemically related to succinylation are also important. This protocol may be used with acetic anhydride, a liquid reagent, in essentially the same fashion as with succinic anhydride. In general, succinylation of amino groups tends to promote the aqueous solubility of proteins, whereas acetylation tends to diminish it. Treating a protein with acetic anhydride causes amino groups to be acetylated: protein-NH2 + (CH3CO)2O → protein-NH-CO-CH3 + CH3COOH This reaction converts the positive charge of the amino group to no charge. Maleic anhydride (Butler and Hartley, 1972) and citraconic anhydride (Atassi and Habeeb, 1972) are other related amino-specific reagents. They add negative charge to the protein, but the modification can be reversed by exposing the modified protein to acidic pH. Reversing a modification provides a way to ensure that functional changes attributed to the modification really were caused by it and not by incidental factors. It is wise, however, to determine early on whether the protein is stable to the reversal conditions, which can be harsh.
Modification of Amino Groups
The following conditions are recommended as a starting point for succinylation. If sufficient protein is available, a range of conditions can be explored in parallel small-scale
15.2.8 Supplement 4
Current Protocols in Protein Science
Current Protocols in Protein Science
Weigh 0.1 mol boric acid per liter H2O and add it to ∼90% of the final intended volume. Adjust the pH to the desired value by adding 5 M NaOH (CAUTION: Wear eye protection and gloves) while stirring the mixture and monitoring the pH with a pH meter. All the boric acid will have dissolved when the pH reaches 9.0. Add H2O to final volume. Dissolve 0.1 mol NaHCO3 per liter of H2O. The solution has a pH of 8.3. Use it as prepared; no further adjustment is needed.
0.1 M sodium borate, pH 9.0
0.1 M NaHCO3 (not pH-adjusted), pH 8.3b
bThis solution is not a true buffer at pH 8.3, but provides an excess of a weak base (bicarbonate) to neutralize acid produced by the modification chemistry.
aAbbreviation: HEPES, N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid.
Weigh 0.1 mol solid HEPES (zwitterion form) per liter H2O and add it to ∼90% of the final intended volume of H2O. Adjust the pH to the desired value by adding suitable volumes of 5 M NaOH (CAUTION: Wear eye protection and gloves) while stirring the mixture and monitoring the pH with a pH meter. Add H2O to final volume.
Procedure
0.1 M HEPES, pH 8.0
Relevant equilibria
Selected Buffer or Base Solutions Useful in Chemical Modification of Amino Groups
Buffer or basea
Table 15.2.3
Chemical Modification of Proteins
15.2.9
Supplement 4
O O O succinic anhydride
A protein NH2
protein NH CO CH2 CH2 COOH major reaction
B protein
OH
protein
O CO CH2 CH2 COOH minor reaction
Figure 15.2.3 The principal reactions of succinic anhydride with proteins. (A) Succinylation of an amino group, which is normally the major result of treating a protein with succinic anhydride. (B) Succinylation of tyrosine, the minor reaction with a phenolic group of tyrosine; this reaction may be reversed by treating the protein with hydroxylamine⋅HCl at pH 7. Ionizable groups are drawn in their neutral state (amino as NH2 and carboxyl as COOH).
experiments. Details of the procedure may have to be revised based on experience with a specific protein. Materials Succinic anhydride 1 to 10 mg protein/ml in 0.1 M sodium bicarbonate, pH 8.3 1 M NaOH 1 M hydroxylamine⋅HCl, pH 7.0 pH 6 to 10 dye-indicator pH sticks Gel-filtration column containing chromatography medium (e.g., PD-10 column containing Sephadex G-25, Pharmacia Biotech) Additional reagents and equipment for gel-filtration chromatography (UNIT 8.3) or dialysis (UNIT 4.4 and APPENDIX 3B) and for mass spectrometry (see Chapter 16) or isoelectric focusing (UNIT 10.2) 1a. To succinylate some, but not all, amino groups in the protein: Calculate the amount of succinic anhydride that represents a 5-fold molar excess over ε-amino groups of Lys. To obtain a 5-fold molar excess, the amount of succinic anhydride required is: 5 (mol protein) × number Lys/mol protein × formula weight of succinic anhydride = grams of reagent
Modification of Amino Groups
For example, the mass of succinic anhydride giving a 5-fold excess over the Lys residues in 2 mg of a 20,000-Da protein that contains 10 Lys residues per mole of protein is determined as follows. (a) Determine the molar quantity of protein, 0.002 g/20,000 g mol−1 = 100 nmol protein; (b) multiply by 10 to determine the molar quantity of Lys residues in 100 nmol of this protein, 10 × 100 nmol = 1 ìmol; (c) multiply by 5 to obtain the molar quantity of succinic anhydride required to give a 5-fold excess over Lys residues, 5 × 1 ìmol = 5 ìmol succinic anhydride; (d) calculate the mass of succinic anhydride (FW 100.1) corresponding to 5 ìmol, (5 × 10−6 mol)× (100 g mol−1) = 0.5 mg succinic anhydride.
15.2.10 Supplement 4
Current Protocols in Protein Science
1b. To succinylate all amino groups: Calculate the amount of succinyl anhydride that represents a 20-fold molar excess over amino groups. 2. Add one-tenth the calculated amount of solid succinic anhydride to the protein with gentle shaking or stirring. This reaction is normally performed at room temperature, but it will proceed at 0°C. The succinic anhydride will dissolve within ∼1 min while reacting with water or the protein.
3. Check the pH of the reaction mixture by placing 1 µl on the appropriate part of a pH 6 to 10 dye-indicator pH stick. If the pH has dropped to <8, add small amounts of 1 M NaOH until it returns to near the starting value. Succinic anhydride is a neutral compound, but its reactions with the protein and water liberate acidic groups and suppress basic amino groups. Consequently, the pH in the reaction mixture tends to fall. To maintain optimal conditions for modifying the amino groups, the pH must be kept at 8.0 to 9.0.
4. Add succinic anhydride in small portions at intervals of a few minutes until all of the required reagent is added. Continue to monitor and maintain the pH. After the final addition, incubate an additional 15 min. 5. Pass the reaction mixture over the gel-filtration column to remove residual low-mass reaction components. Dialysis (UNIT 4.4 and APPENDIX 3B) or diafiltration could be used as alternatives. The modified form of the amino groups is stable to these procedures.
6. Evaluate the success of the reaction. Mass spectrometry (see Chapter 16) is the best technique when full succinylation is desired; each succinylation adds 100.1 Da. Isoelectric focusing (e.g., on a Pharmacia PhastSystem apparatus or as described in UNIT 10.2) may be helpful with incomplete succinylations, as each charge changed from positive to negative shifts the isoelectric point. A colorimetric assay of amino groups using trinitrobenzenesulfonic acid (see Table 15.2.1) is also useful.
7. Optional: Reverse modifications at tyrosines by incubating the modified protein with 1 M hydroxylamine⋅HCl, pH 7.0, for 1 hr at 20°C. REDUCTIVE METHYLATION Reductive dimethylation of an amino group amounts to replacing its two hydrogen atoms with methyl groups, which converts a primary amine into a tertiary amine. The monomethyl amino group is formed first and readily undergoes a second round of modification to yield the final product. The resulting tertiary amine retains the ability to become protonated, making this modification a subtle probe of steric constraints about amino groups in the molecule. As the modified amino group is no longer an effective nucleophile, reductive methylation can be used to muzzle the reactivity of protein amino groups without otherwise greatly changing their properties. The chemistry of each round of reductive methylation is itself a two-step process (see Fig. 15.2.4). First, reaction of formaldehyde with the protein results in reversible formation of a Schiff base (imine) adduct between the aldehyde and any protein amino group. Second, the modification is made permanent by the reducing (hydrogen-donating) action of sodium cyanoborohydride, which converts the imine to a carbon-nitrogen single bond. Unless the reagents are present in limiting supply, the resulting monomethylated protein amino group undergoes a further round of the same reaction to give the dimethylated form. The overall extent of substitution varies with the conditions, as described in detail
BASIC PROTOCOL 4
Chemical Modification of Proteins
15.2.11 Current Protocols in Protein Science
Supplement 4
BH3CN –
protein NH2
O CH2
protein N CH2
protein NH CH3 monomethylated amino group
BH3CN –
protein NH
CH3
O CH2
protein N(CH3) CH2
monomethylated amino group
protein N(CH3)2 dimethylated amino group
Figure 15.2.4 Reductive methylation of a protein amino group.
by Jentoft and Dearborn (1979). When consulting their excellent paper, note that the extent of modification in many of the examples approaches 1.0 methyl group per lysine, while the maximum possible extent of modification is 2.0 methyl groups per lysine. Higher concentrations of the reagents than those given by Jentoft and Dearborn (1979) are required when full dimethylation of all amino groups is sought, as may be the case when investigating structure-function relationships; for examples of such conditions, see Geoghegan et al. (1981). Comprehensive guidance for this procedure is available from the review by Jentoft and Dearborn (1983). ε-N-methyl-L-lysine and ε-N,N-dimethy-L-lysine are not normally present in acid hydrolyzates of proteins (although they do occur in rare cases), so standard chromatographic procedures for amino acid analysis require modification to permit determination of those amino acids (Elzinga and Alonzo, 1983). Useful variations of this procedure have been developed. For example, aldehydes or ketones larger than formaldehyde can be used; the technique is then called, generally, reductive alkylation. This should not be confused with the approaches to sulfhydryl modification collectively known as reduction and alkylation (see UNIT 15.1). Other possibilities include the selection of modifying groups that can later be removed (Geoghegan et al., 1979; for a review, see Feeney, 1987). Reductive alkylation chemistry has also been used to immobilize protein or amino acid ligands for affinity chromatography, as in the Actigel ALD line of media from Sterogene Bioseparations. Materials 1 to 10 mg protein/ml in 0.1 M sodium citrate, pH 6.0 37% formaldehyde (Aldrich) Sodium cyanoborohydride (NaBH3CN, Aldrich; see CAUTION below) Methanol Additional reagents and equipment for dialysis (UNIT 4.4 and APPENDIX 3B) or gel-filtration chromatography (UNIT 8.3) and for mass spectrometry (Chapter 16) or amino acid analysis (UNIT 3.2) CAUTION: Sodium cyanoborohydride is a highly toxic compound. Follow all precautions recommended in the Material Safety Data Sheet. Do not attempt to sniff the compound’s odor, and be aware that exposing it to strong acid will result in the release of cyanide gas. Modification of Amino Groups
15.2.12 Supplement 4
Current Protocols in Protein Science
1. Measure the volume of the protein/0.1 M sodium citrate solution. Calculate the volume of 37% formaldehyde required to give a final concentration of 20 mM and add to the solution. Formaldehyde has an Mr of 30, and a 37% solution therefore has a concentration of 12.3 M. A 1/100 dilution is 0.12 M; a further 1/6 dilution into the reaction mixture gives a concentration of 20 mM. It is important not to exceed the recommended level of formaldehyde in the reaction, as it is a cross-linking agent.
2. Calculate the volume of 1 M sodium cyanoborohydride solution that is required to give a final concentration of 10 mM in the reaction. Sodium cyanoborohydride has a formula weight of 62.84. A 1 M solution requires 63 mg/ml. As an alternative, a 1 M solution of reportedly highly purified NaBH3CN can be purchased from Sterogene Bioseparations (see SUPPLIERS APPENDIX) under the name of ALD Coupling Solution. If there is a particular desire to avoid the use of sodium cyanoborohydride, one of a number of amineboranes can be substituted (Geoghegan et al., 1981).
3. Weigh the hazardous sodium cyanoborohydride in the following manner. Tare a balance to zero with the tube to be used to prepare the solution. Working in a fume hood, transfer into the tube a quantity of sodium cyanoborohydride near the amount required. After closing the tube inside the hood, take the tube to the balance and weigh it again. Calculate the amount of methanol required to give a solution at the desired concentration, and carefully add it to the tube. Working in a fume hood, collect and rinse discarded spatulas and other equipment left over from the preparation of this solution. Use the solution immediately, as sodium cyanoborohydride begins to decompose as soon as it is dissolved.
4. Add methanolic sodium cyanoborohydride to the protein solution to a final concentration of 10 mM, i.e., a 1/100 dilution of the 1 M solution. Incubate 120 min at 22°C. 5. Separate the protein from low-mass reactants by dialysis (UNIT 4.4 and APPENDIX 3B) or HPLC. 6. Use mass spectrometry (see Chapter 16) or amino acid analysis (UNIT 3.2) to determine the extent to which the amino groups of the protein have been methylated. Monomethylation of an amino group increases the mass of the protein by 14 Da, and dimethylation increases its mass by 28 Da.
RAPID DESALTING OF PROTEIN SAMPLES FOR ELECTROSPRAY MASS SPECTROMETRY
SUPPORT PROTOCOL
Mass spectrometry (Chapter 16) has revolutionized protein chemistry by making full molecular characterization readily attainable. Under favorable conditions, it can be used to measure the progress of a modification experiment with unmatched analytical accuracy. However, to reap the potential benefits of this technique, it is necessary to have a convenient means of preparing samples for analysis. Mass spectrometry of biomolecules is an exercise with two major parts. The second is the mass analysis itself, but before that comes the requirement to generate a gas-phase ion of a large and labile analyte. Electrospray is one of a variety of methods for generating gas-phase ions. In this technique, a protein sample is sprayed as a population of droplets
Chemical Modification of Proteins
15.2.13 Current Protocols in Protein Science
Supplement 4
1.0
100
0.5
50
%solvent B (– – – –)
Absorbance (220nm)
protein peak
0
0.0 0
2
4
6 Time (min)
8
10
Figure 15.2.5 Rapid reversed-phase HPLC of a 61-kDa human recombinant intracellular enzyme (purified to homogeneity before this fractionation). The protein peak indicated by the arrow was collected, dried, and successfully analyzed by electrospray mass spectrometry.
into a compartment where it is exposed to high voltage. Evaporation of solvent from the sample droplets creates a destabilizing high charge density, and the droplets disintegrate in a process that makes multiply protonated gas-phase protein ions available for mass analysis. This can be achieved with 0.01% accuracy in a quadrupole-type spectrometer, i.e., to an accuracy of ± 4 Da in 40,000 Da. The electrospray mode of sample introduction is very refractory to the presence of nonvolatile salt and buffer components in the sample. A convenient way to remove them is to use an HPLC system, consisting of a gradient pump system running at 1 ml/min, an injection device, and a UV-detector set to 220 nm. The system is configured to allow hand collection of peak fractions for rapid reversed-phase HPLC fractionation. The protein is retained on a 2.1-mm-i.d. × 3-cm-long reversed-phase HPLC column (e.g., Poros R2/H 2.1/30, PerSeptive Biosystems) for a short period while buffers and low-mass reagents are washed out in the injection peak. Elution is with an acetonitrile gradient using solvent A (0.1% [v/v] trifluoroacetic acid in water) and solvent B (0.085% [v/v] trifluoroacetic acid in 80:20 acetonitrile/water). This places the protein in a water/acetonitrile/trifluoroacetic acid solvent that is removed by evaporation. If the dried protein residue is visible (although this is not essential), there is usually enough protein for mass spectrometry (see Fig. 15.2.5 for the HPLC profile for a 61-kDa protein). Although the HPLC column selected has an internal diameter of 2.1 mm (i.e., it is a narrow-bore column), the character of the packing allows it to run at flow rates >1 ml/min without excessive back-pressure. A flow rate of 1 ml/min is recommended. Gradient elution is performed over 6 min from 0 to 70% solvent B (i.e., 0% to 56% acetonitrile). First, a small portion of the protein is injected to determine that the protein can be recovered and to estimate the retention time; this will also give an impression of whether multiple protein peaks are to be expected. A quantity of the protein sample containing 1 Modification of Amino Groups
15.2.14 Supplement 4
Current Protocols in Protein Science
to 2 nmol is then injected, and the protein peak is collected by hand. Solvent is removed in a Speedvac evaporator. NOTE: One nanomole of any protein is calculated on the basis of one microgram per kilodalton of relative mass, i.e., 1 nmol of a 20-kDa protein is 20 µg, and 1 nmol of a 100-kDa protein is 100 µg. COMMENTARY Background Information
Critical Parameters
This unit presents only a limited selection of approaches to amino-group modification. In general, the methods currently of greatest interest are those adaptable to bioconjugate formation, protein immobilization, and other technological applications. There is less attention to methods that appear useful only as probes of protein function, but they are well documented in the literature (Means and Feeney, 1971). Reactions in this category include carbamylation of amino groups with cyanate (also still significant as an incidental reaction of proteins exposed to high concentrations of urea, from which cyanate can form; Stark, 1967); formation of imidoamides (amidines) by reaction with imidoesters (Inman et al., 1983); and arylation of amino groups with a selection of reagents such as 1-fluoro-2,4-dinitrobenzene (Sanger reagent; Hirs, 1967), trinitrobenzenesulfonic acid (Fields, 1972), and dinitrobenzenesulfonic acid. Of this last group, trinitrobenzenesulfonic acid is the easiest to work with and readily modifies amino groups in aqueous solution at room temperature. Dinitrobenzenesulfonic acid reacts much more sluggishly under comparable conditions, but it can be useful for tagging small, heat-stable peptides with a dinitrophenyl group. The classical modes of modification described in this unit now possess an importance rather different from that attached to them when they were new. Having once been among the most current tools for studying protein structure and function, they are now best viewed as prototypes for chemical modification schemes representing many ways to introduce new and useful groups into proteins. Succinimidyl esters, for example, are quite easy to prepare from carboxylic acids (see Brinkley, 1993), and they allow a growing range of non-native structures to be made part of proteins by reaction with the protein amino groups. Bifunctional reagents, a special category, are often directed toward amino groups at one or both of their two reactive sites (reviewed by Ji, 1983).
Proteins are subjected to chemical modification for a number of purposes, and the objective of the work defines the terms for all estimates of its success. In such a broad area, it is difficult to set practical guidelines for every case. The key discipline is to acknowledge that a chemical reaction is like a newly written computer program: it ruthlessly follows its own logic rather than the intentions of its creator. As a result, it is always important to consider whether the chemical modifications that have occurred (if any!) are actually those that were intended, and to ask whether any modifications that were not intended have also taken place. If the reaction has been driven to achieve quantitative and exclusive modification of the amino groups in the protein, the product should be homogeneous and mass spectrometry (see Chapter 16) will be the appropriate and preferred way to characterize it. Partial modification gives a mixture of products, and the resulting molecules are more likely to have a mixture of properties. This is the classical flaw in the chemical modification approach. A mass spectrometric technique, such as MALDI-TOF MS (matrix-assisted laser desorption and ionization time-of-flight mass spectrometry; UNIT 16.2), that accepts mixtures may still be useful for analyzing the results in these cases. Laboratories lacking mass spectrometers may find isoelectric focusing (UNIT 10.2) useful, and even SDS–polyacrylamide gel electrophoresis (UNIT 10.1) may be worth considering if the modification effects sufficient mass change in the protein to be detected. Finally, if a chromophore has been coupled to the protein molecule, absorption spectroscopy may furnish an estimate of the number of modifying groups introduced per molecule of protein.
Troubleshooting Whatever the purpose of a modification experiment, failure to achieve the desired reaction is the most obvious form of trouble (see Table 15.2.4). Several likely explanations must always be considered. At the top of the list is the
Chemical Modification of Proteins
15.2.15 Current Protocols in Protein Science
Supplement 4
Table 15.2.4
Troubleshooting Guide for Modification of Amino Groups
Problem
Possible cause(s)
Solution
Little or no modification is achieved
Reagent is already degraded when added to protein
Use fresh reagent. For reagents labile to hydrolysis, minimize or eliminate their exposure to aqueous solution before adding them to the protein.
Reaction pH is inappropriate
Check the reaction pH after reagent has been added; it should be pH 8-9 in most cases. Monitor the pH and adjust it if necessary.
The reagent is reacting with the buffer
If the buffer first chosen contains a primary or secondary amino group (e.g., Tris), change to a buffer that does not; e.g., HEPES, sodium bicarbonate, or boric acid/sodium borate.
Modification product is insoluble
Determine whether altered ionic strength or the addition of more or less organic solvent (e.g., acetonitrile) corrects the difficulty.
Protein becomes insoluble when a large fraction of its amino groups are modified
If a lower level of modification can still be useful, reduce the amount of reagent and/or the reaction time.
A solvent added with the reagent causes the precipitation
Test the effect of the solvent alone on protein solubility. If solvent causes the problem, minimize the amount added or use another solvent in its place.
Visible absorption spectrum of protein conjugate with a fluorescent reagent differs from that of free reagent
Multiple copies of strongly fluorescent groups (e.g., fluorescein) linked to a single protein molecule may achieve resonance energy transfer when irradiated
Distortion of the absorption spectrum is an artifact of placing multiple fluorescent groups in close proximity to each other. No unpredicted chemical change has occurred in the tagging group. Consult the literature.
Mass spectrometry of a heavily modified protein indicates that more equivalents of reagent were added than there are amino groups to accept them
Modification has occurred on sites other than amino groups, particularly tyrosyl residues
Incubate the modified protein with 1 M hydroxylamine⋅HCl for 1-5 hr at room temperature to reverse possible modifications at tyrosine, then recheck the product.
Protein comes out of solution during modification reaction
Modification of Amino Groups
possibility that the reagent is degraded and unsuitable for use. For example, succinimidyl esters are labile esters of carboxylic acids and can hydrolyze if exposed to water at neutral or basic pH before use. The risk of hydrolysis is not present when reagents are dissolved in dry aprotic solvents such as acetonitrile or dimethylformamide. Long-term exposure to trace levels of water may be effective in degrading reagents, so older or poorly stored supplies are always suspect. Another likely error is inclusion in the reaction medium of amino-containing compounds that compete with protein amino groups for the
modifying group. Apart from the hazards, already mentioned, of using amine-containing buffer components such as Tris or glycine, any compound with a primary or secondary amino group is a potential competitor with the protein amino groups. The reaction conditions must also be reconsidered when trouble occurs. Was the pH at or near the desired value? See Scopes (1994) for a superb discussion and practical advice on choosing and making buffers. Did the pH remain at this value after the time allowed for reaction has been completed? If pH is satisfactory, then temperature must also be considered.
15.2.16 Supplement 4
Current Protocols in Protein Science
The reactions described in this unit are fairly fast at room temperature (∼20°C), but some other procedures for amino-group modification may require gentle heating. In particular, dinitrophenylation of amino groups using dinitrobenzenesulfonic acid is a slow reaction that requires temperatures >40°C. This may make it unsuitable for use with many proteins, but it can be used successfully with peptides.
Anticipated Results The amino groups of proteins are easy to modify, and failure of these procedures is usually caused by overlooking some technical detail. A harder task is to predict accurately all the properties of a modified protein. Even a single hydrophobic fluorescent tag, for example, can seriously diminish the solubility of a protein that was easily handled in its native form. This type of uncertainty is part of any chemical experiment; it simply calls for vigilance and a guarded attitude by way of response, accepting that there may be unforeseen consequences when the charge distribution on a protein’s surface is varied or novel structures are introduced. As the native folded form of a protein typically enjoys only a marginal advantage in stability relative to many unfolded structures, it is not surprising that changing the structure sometimes has consequences that go beyond those first intended.
Time Considerations Most modification reactions should be completed in ≤2 hr, and fast methods such as gelfiltration chromatography (UNIT 8.3) can be used to avoid long delays in the follow-up. Microconcentrators such as the Centricon and related lines (Amicon) are also valuable tools for reducing sample volumes or achieving buffer exchanges in relatively short order, but note that their internal surfaces may need to be treated with polyethylene glycol to reduce adsorptive losses of protein (see manufacturer’s instructions and UNIT 4.4). The results of most chemical modification experiments should be well understood within 24 hr of their inception.
Literature Cited
Banks, P.R. and Paquette, D.M. 1995. Comparison of three common amine-reactive fluorescent probes used for conjugation to biomolecules by capillary zone electrophoresis. Bioconjugate Chem. 6:447-458. Brinkley. M. 1993. A brief survey of methods for preparing protein conjugates with dyes, haptens, and cross-linking reagents. In Perspectives in Bioconjugate Chemistry (C.F. Meares, ed.) pp. 59-70. American Chemical Society, Washington, DC. Butler, P.J.G. and Hartley, B.S. 1972. Maleylation of amino groups. Methods Enzymol. 25:191-199. Dixon, H.B.F. and Fields, R. 1972. Specific modification of NH2-terminal residues by transamination. Methods Enzymol. 25:409-419. Elzinga, M. and Alonzo, N. 1983. Analysis for methylated amino acids in proteins. Methods Enzymol. 91:8-13. Feeney, R.E. 1987. Chemical modification of proteins: Comments and perspectives. Int. J. Pept. Protein Res. 29:145-161. Fields, R. 1972. The rapid determination of amino groups with TNBS. Methods Enzymol. 25:464468. Geoghegan, K.F. and Stroh, J.G. 1992. Site-directed conjugation of nonpeptide groups to peptides and proteins via periodate oxidation of a 2-amino alcohol. Application to modification at N-terminal serine. Bioconjugate Chem. 3:138-146. Gaertner, H.F. and Offord, R.E. 1996. Site-specific attachment of functionalized poly(ethylene glycol) to the amino terminus of proteins. Bioconjugate Chem. 7:38-44. Geoghegan, K.F., Ybarra, D.M., and Feeney, R.E. 1979. Reversible reductive alkylation of amino groups in proteins. Biochemistry 18:5392-5399. Geoghegan, K.F., Cabacungan, J.C., Dixon, H.B.F., and Feeney, R.E. 1981. Alternative reducing agents for reductive alkylation of amino groups in proteins. Int. J. Pept. Protein Res. 17:345-352. Hirs, C.H.W. 1967. Reactions with reactive aryl halides. Methods Enzymol. 11:548-555. Inman, J.K., Perham, R.N., DuBois, G.C., and Appella, E. 1983. Amidination. Methods Enzymol. 91:559-569. Jentoft, N. and Dearborn, D.G. 1979. Labeling of proteins by reductive methylation using sodium cyanoborohydride. J. Biol. Chem. 254:43594365. Jentoft, N. and Dearborn. D.G. 1983. Protein labeling by reductive alkylation. Methods Enzymol. 91:570-579.
Alouani, S., Gaertner, H.F., Mermod, J.J., Power, C.A., Bacon, K.B., Wells, T.N., and Proudfoot, A.E. 1995. A fluorescent interleukin-8 receptor probe produced by targetted labelling at the amino terminus. Eur. J. Biochem. 227:328-334.
Ji, T.H. 1983. Bifunctional reagents. Methods Enzymol. 91:580-609.
Atassi, M.Z. and Habeeb, A.F.S.A. 1972. Reaction of proteins with citraconic anhydride. Methods Enzymol. 25:546-553.
Means, G.E. and Feeney, R.E. 1971. Chemical Modification of Proteins. Holden-Day, San Francisco.
Klotz, I.M. 1967. Succinylation. Methods Enzymol. 11:576-580. Chemical Modification of Proteins
15.2.17 Current Protocols in Protein Science
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Scopes, R.K. 1994. Control of pH: Buffers In Protein Purification: Principles and Practice, pp. 324-333. Springer-Verlag, New York. Stark, G.R. 1967. Modification of proteins with cyanate. Methods Enzymol. 11:590-594.
Key References Brinkley, M. 1993. See above. Surveys current techniques for preparing protein conjugates.
Tae, H.J. 1983. Bifunctional reagents. Methods Enzymol. 91:580-609.
Jentoft, N. and Dearborn, D.G. 1983. See above.
Wetzel, R., Halualani, R., Stults, J.T., and Quan, C. 1990. A general method for highly selective cross-linking of unprotected polypeptides via pH-controlled modification of N-terminal αamino groups. Bioconjugate Chem. 1:114-122.
Means, G.E. and Feeney, R.E. 1971. See above.
Zhang, L. and Tam, J.P. 1996. Thiazolidine formation as a general and site-specific conjugation method for synthetic peptides and proteins. Anal. Biochem. 233:87-93.
Tae, H.J. 1983. See above.
A comprehensive guide to reductive alkylation.
A classic book that remains invaluable for direct advice and fundamental information on chemical modification of proteins.
Comprehensive coverage of modifying agents.
Contributed by Kieran F. Geoghegan Pfizer, Inc. Groton, Connecticut
Modification of Amino Groups
15.2.18 Supplement 4
Current Protocols in Protein Science
CHAPTER 16 Mass Spectrometry INTRODUCTION
F
rom the point of view of protein scientists, in the course of the 1990s mass spectrometry (MS) has developed from a fairly exotic technique practiced in only a handful of laboratories into a routine, essential tool for protein characterization. MS is complementary to other protein and peptide analysis methods such as protein sequencing, amino acid analysis, and high-performance liquid chromatography (HPLC) peptide mapping (Chapter 11). It is particularly useful for identifying post-translational modifications (Chapters 12 to 14), which are often quite difficult to distinguish by other methods. To some extent, MS is beginning to replace alternative methods such as Edman sequencing as continuing advances in instruments, software, and user-friendliness improve the sensitivity and utility of MS methods.
A detailed discussion of peptide and protein MS analysis, including sample ionization methods, basic principles and parameters, and data interpretation, is presented in UNIT 16.1. The unit also lists numerous literature references that should be especially useful to scientists seeking more detailed information on specific MS-related issues. A key advance in the application of MS for routine analysis of peptides and proteins has been the development of commercial instruments that employ matrix-assisted laser desorption/ionization (MALDI) techniques to transfer peptides and proteins efficiently into the gas phase for time-of-flight based mass measurements. This technique has multiple advantages compared with older techniques, including the instruments’ relatively low cost and ease of operation, relatively good tolerance of nonvolatile buffer components, and ability to analyze heterogeneous samples as mixtures, since the ionization method predominantly produces a single charged species per sample component. A general discussion of MALDI-MS analysis of peptides is presented in UNIT 16.2, and UNIT 16.3 provides specific protocols for analysis of both peptides and proteins, with emphasis on sample preparation. A frequent research question in biology is the correlation of observed function with the proteins responsible. MS-based methods have emerged as the most sensitive, rapid, and economical method for identifying proteins. The simplest approach is MALDI-MS fingerprint mapping following in-gel protease digestion of the protein of interest (UNIT 16.4). The observed peptide masses are used to search sequence databases over the Internet (UNIT 16.5) using one of several alternative algorithms. This method works best when the protein of interest is expected to be found in an available sequence database—i.e., when the complete genome of the species from which the protein was isolated has been sequenced. When the database contains only related, but nonidentical, proteins, or when the results of the fingerprint mapping search are ambiguous, partial sequence information is generally required to identify homologous proteins or to search expressed-sequence tag (EST) databases. Such partial sequence information can be obtained on a MALDI spectrometer either using fragmentation induced within the mass analyzer (UNIT 16.6) or via exopeptidase digestion of samples on the MS sample target (UNIT 16.7). The other common mass analysis method for biomolecules is electrospray ionization (ESI)-MS (UNITS 16.1, 16.8, & 16.9). In recent years, the cost of ESI-based instruments has Contributed by David W. Speicher Current Protocols in Protein Science (2000) 16.0.1-16.0.2 Copyright © 2000 by John Wiley & Sons, Inc.
Mass Spectrometry
16.0.1 Supplement 27
decreased and they have become more sensitive and user friendly. Triple-quadrupole and ion-trap instruments with ESI interfaces are particularly well suited to tandem mass spectrometry (MS-MS) experiments, in which an initial mass measurement is followed by fragmentation of a target peptide and reanalysis of fragment mass spectra. ESI-MS is much less tolerant of salts and buffers than MALDI-MS. Hence, interfering components can be removed by trapping the peptides or proteins on a small amount (typically <2 µl) of reversed-phase resin. After removing polar components with water, the peptides or proteins are eluted with a few microliters of a high-purity organic solvent such as acetonitrile. The desalted samples are then introduced into an ESI mass spectrometer using nanoliter flow rates from a borosilicate nanospray glass needle (UNIT 16.8). This sample preparation/analysis is especially well suited for detailed structural studies of simple samples containing a small number of protein or peptide components, e.g., analysis of posttranslationally modified peptides. Alternatively, samples can be introduced into ESI mass spectrometers by on-line liquid chromatography (LC) using a microscale capillary reversed-phase column incorporated into the electrospray needle (UNIT 16.9). The on-line reversed phase column can concentrate the sample, remove contaminating buffers and salts, and partially separate peptide/protein components prior to MS analysis. This is the method of choice for analyzing complex mixtures such as tryptic peptides from in-gel digestion of proteins excised from polyacrylamide gels. When on-line nanocapillary LC columns are used without post-column stream splitting, very high sensitivities can be obtained. One of the most common uses of tandem MS (MS/MS) spectra is identification of proteins in polyacrylamide gels or purified macromolecular complexes using the SEQUEST software to search DNA and protein sequence databases (UNIT 16.10). However, database searches with SEQUEST or analogous MS/MS spectral matching algorithms are most reliable wehn the complete sequence of the genome for the species being studied is in the database that is searched. Even when a highly homologous protein from a closely related species is in the database, a definitive match may be missed because sequence coverage by the MS/MS data is rarely complete and nearly all substitutions change tryptic peptide masses. High-quality MS/MS spectra that do not match sequences in databases can be manually interpreted; that is, by de novo peptide sequencing (UNIT 16.11). De novo sequence interpretation is also used to locate specific residues that are posttranslationally or chemically modified, or to locate naturally occurring substitutions in a protein with a known sequence. David W. Speicher
Introduction
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Current Protocols in Protein Science
Overview of Peptide and Protein Analysis by Mass Spectrometry WHY IS MS AN ESSENTIAL TOOL IN PROTEIN STRUCTURE ANALYSIS? Mass spectrometry (MS) is now regarded as an indispensable tool for peptide and protein primary structure analysis. The increasingly widespread use of this technique is due in large part to its ability to solve structural problems not easily handled by conventional protein chemistry techniques. MS can provide accurate molecular weights for peptides and proteins with masses up to 500,000 Da using only a few picomoles (tens of femtomoles in favorable cases) of material. The accuracy achieved by MS is frequently better than 0.01% of the calculated mass (see Table 16.1.1). This means, for example, that the experimentally determined Mr of a 25-kDa protein will be ±2.5 Da of the actual mass. In contrast, molecular weight estimates obtained by SDS-PAGE have accuracies of 5% to 10%; moreover, the mobility of the protein in the gel can be grossly affected (up to 50%) by the presence of covalent modifications such as lipids and carbohydrates. The accuracy of protein molecular weight estimates obtained by other methods, such as sedimentation velocity and gel permeation, can also be affected by the presence of lipid or carbohydrate. For correction and interpretation of this data, a molecular weight derived by mass spectrometry is needed (Hensley et al., 1994). Unlike these other techniques, mass spectrometry provides molecular mass information independent of any modifications to the structure (such as glycosylation and phosphorylation). A mass difference between the observed and the expected mass of the protein or a peptide suggests the presence of post-translational modification or proteolytic processing, and the mass difference will be the starting point for defining the structure of the modification. A difference of +42 Da, for example, together with the lack of a free NH2-terminus, is indicative of the presence of an α-N-acetyl group. Similarly, a difference of +80 Da suggests the presence of a phosphate or sulfate moiety, and a cluster of molecular ion peaks separated by 162, 203, 291, or 146 Da indicates glycoforms differing by the presence of hexose, N-acetylhexosamine, N-acetylneuraminic acid, or deoxyhexose (see Tables A.1A.2 and A.1A.3 in
UNIT 16.1
APPENDIX 1A).
Thus molecular weight information provides a useful measure of the integrity, purity, and overall state of modification of a peptide or protein. When evaluated in the context of other available structural data, the molecular mass information alone can be used to solve a variety of peptide and protein structural problems. The ability of MS to determine the molecular weights of peptides in mixtures (such as result from a proteolytic digest of a protein) with better than 0.01% accuracy forms the basis of the mass spectrometric peptide mapping strategy, a rapid and highly efficient method to verify the fidelity of translation of protein sequences deduced from DNA or cDNA sequencing. This can be done either by direct analysis of unfractionated mixtures (Billeci and Stults, 1993) or on-line with liquid chromatography (Hess et al., 1993; Carr et al., 1993; Kassel et al., 1994) or capillary electrophoresis (Wahl et al., 1992; Smith et al., 1996; Foret et al., 1994). Mass spectrometry also has the ability to provide amino acid sequence information on peptides using methodology generically referred to as tandem mass spectrometry (MS/MS). Using MS/MS, complete or partial sequence information may be obtained at the femtomole to picomole level for peptides containing up to 25 amino acid residues. In favorable cases some internal sequence data can often be obtained for peptides up to 30 or 40 residues. Unlike Edman sequencing, MS can provide this type of sequence information even if the peptides are present in complex mixtures, and, more importantly, even if they are modified. Methodologies such as Edman degradation and amino acid analysis have serious limitations with regard to characterizing structurally modified proteins and peptides, because many commonly occurring modifications are lost or destroyed by the harsh cleavage and derivatization conditions employed (Uy and Wold, 1977; Wold, 1981; Wold and Moldave, 1984; Carr et al., 1991). Furthermore, the sensitivity and speed of MS-based sequencing now exceeds that of Edman-based sequence analysis (Hunt et al., 1992; Wilm et al., 1996). Vast amounts of cDNA sequence data, obtained by grand-scale sequencing of the human and other genomes, are presently flooding into Mass Spectrometry
Contributed by Steven A. Carr and Roland S. Annan Current Protocols in Protein Science (1996) 16.1.1-16.1.27 Copyright © 2000 by John Wiley & Sons, Inc.
16.1.1 Supplement 4
Table 16.1.1 Key Ionization Methods for Biomolecule Analysis
Ionization method
Tolerance for buffers, salts, and detergents
MW range
Mass assignment accuracy
Compatible with on-line LC/CE-MS
Electrospray ionization (ESI)
Low tolerance (≤20 mM) for nonvolatile buffers, alkali metal salts, detergents; on-line LC-MS used for contaminant removal
≤150 kDA
±0.005%–0.01% to 50 kDA ±0.02%–0.03% to 150 kDA
Yes
Matrix-assisted laser desorption/ionization (MALDI)
High tolerance (≥50 mM) for alkali metal salts, phosphate, urea, etc.; tolerant of some nonionic detergents; intolerant of SDS
≥350 kDA
±0.01%–0.05% to 25 kDA ±0.05%–0.3% to 300 kDA
No
Overview of Peptide and Protein Analysis by Mass Spectrometry
databases. However, little is known about the functions of most of the proteins the various genes encode, nor is much known about the processed states of the expressed, mature forms of the proteins. In contrast, the protein content of a cell, referred to as the proteome (Kahn, 1995; Wilkins et al., 1995), can be studied by protein differential display using two-dimensional gel electrophoresis (UNIT 10.4), immunoprecipitation, and related procedures, allowing physiologically relevant proteins to be recognized by changes in their expression levels in response to stimuli. Because of its speed and sensitivity, mass spectrometry, through the use of strategies that employ molecular weight and/or partial sequence information to search protein databases, has emerged as the primary technology for identifying these proteins (Henzel et al., 1993; Mann et al., 1993; Pappin et al., 1993; Wilm and Mann, 1994; Eng et al., 1994). This introductory overview has been designed with several goals in mind. The first and perhaps the most important one is to try and dispel the still widely held, but now mistaken, belief that mass spectrometry is beyond the technical resources of the typical laboratory or facility engaged in the structural characterization or synthesis of peptides and proteins. Over the last five years, mass spectrometers and associated data systems have become available that can be operated by anyone capable of running a modern amino acid analyzer or Edman sequencer, yet are powerful enough to handle the most demanding analyses that the peptide or protein investigator might require. As a result, it is no longer necessary to be a “card-carrying” mass spectroscopist to use the instruments and techniques of MS effectively and to apply it successfully to one’s own research program. The remaining goals of this
article are to familiarize peptide and protein chemists with the types of mass spectrometers that are appropriate for the majority of their analytical needs, to describe the kinds of experiments that can be performed with these instruments on a routine basis, and to discuss the kinds of information that these experiments provide. The emphasis here is on established tools and techniques that can realistically be used for problem solving. As a result, many useful instruments and techniques employed by the trained mass spectroscopist are not discussed here, because they do not satisfy these criteria or are not yet widely available. A few of these are briefly mentioned at the end of this introductory overview. The units that follow this overview will further discuss many of the principles introduced here and provide detailed protocols and applications of the methods. The overview concludes with a tutorial discussion on the meaning and practical importance of a number of fundamental experimental and performance-related issues that have important implications for interpretation and use of mass spectral data, such as mass accuracy and resolution. More comprehensive discussion of mass spectrometry of biological molecules may be found in several recent books (Burlingame and McCloskey, 1990; McCloskey, 1990; Burlingame and Carr, 1996) and review articles (Carr et al., 1991; Biemann, 1992; Chait and Kent, 1992; Aebersold, 1993; Wang and Chait, 1994; Mann and Wilm, 1995; Williams and Carr, 1995; Mann and Talbo, 1996). The biannual review of mass spectrometry in Analytical Chemistry by Burlingame and colleagues provides a particularly good annotated bibliography of the recent literature on techniques and application of MS in the biological sciences
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(Burlingame et al., 1994). Mass spectrometry information is also rapidly proliferating on the World Wide Web; Table 16.2.4 contains a useful listing of sites that provide information about MS.
WHAT IS MS? Mass spectrometry is a powerful analytical technique for forming gas-phase ions from intact, neutral molecules and subsequently determining their molecular masses. All mass spectrometers have three essential components: an ion source, a mass analyzer, and a detector. Ions are produced from the sample in the ion source using a specific ionization method (see discussion of Key Ionization Methods and Related Ancillary Techniques). The ions are separated in the mass analyzer based on their mass-tocharge (m/z) ratios and then detected, usually by an electron multiplier. The data system produces a mass spectrum, which is a plot of ion abundance versus m/z. Except in the case of electrospray mass spectrometry (where the ion source is at atmospheric pressure), the ion source, mass analyzer, and detector are usually situated inside a high-vacuum chamber (pressure between 10−8 and 10−4 Torr).
KEY IONIZATION METHODS AND RELATED ANCILLARY TECHNIQUES The decade of the 1980s marked a revolutionary period in the development of “soft ionization” techniques for the analysis by mass spectrometry of large, polar, nonvolatile molecules (Carr et al., 1991). These ionization techniques produced primarily intact protonated molecular ions of peptides and proteins. In the early 1980s, fast atom bombardment (FAB) and, to a lesser extent, plasma desorption (PD) mass spectrometry were used extensively for structural characterization of peptides and small proteins. In 1988, electrospray ionization (ESI) and matrix-assisted laser desorption/ionization (MALDI) mass spectrometry were first shown to be useful for the analysis of peptides, proteins, carbohydrates, and oligonucleotides. Within a few years of their introduction, ESI and MALDI techniques became the MS methods of choice for biopolymers analysis, largely supplanting FAB-MS and PD-MS. For this reason these techniques will be the focus of this overview. The capabilities and sample requirements of these two MS techniques are compared in Table 16.1.1. The performance specifications in the table are those that can be routinely achieved by trained opera-
tors and do not represent the ultimate performance of each technique. Several factors account for the widespread acceptance of ESI-MS and MALDI-MS. Both ionization methods (see detailed descriptions below) are optimally combined with relatively low-cost, simple-to-operate mass analyzers. A quadrupole mass filter is typically used for ESI-MS, whereas a time-of-flight (TOF) analyzer is commonly employed for MALDI-MS. The sensitivity of both methods is in the lowto subpicomole range, and both are capable of analyzing molecules with a wide range of molecular weights (Table 16.1.1). The methods have many analytical capabilities in common, but do differ in a number of respects that are of practical significance. Both techniques work best with clean (salt- and detergent-free) samples, but MALDI is capable of providing information directly on relatively dirty samples. Both methods can provide molecular weight information for large proteins, but the information is invariably easier to obtain and more sensitive with MALDI-MS. In the molecular weight range from 5 to 50 kDa, however, analysis by ESI-MS usually provides much more accurate molecular weight measurements. ESIMS in this molecular weight range also typically exhibits better mass resolution than MALDI, which permits detection of protein variants that differ only slightly in mass. These practical differences between the techniques, and their impact on the information that can be obtained about a protein structure, are discussed below. In addition to the development of the ionization approaches described above, a number of key enabling technologies that have significantly improved the performance and sensitivity of ESI-MS and MALDI-MS have been introduced in the last few years. These technologies have moved MS onto the center stage of protein discovery and characterization. The first such advance has been the development and refinement of microscale desalting procedures such as efficient liquid-liquid extraction for peptides produced by in-gel digestion (Shevchenko et al., 1996), micropacked glass capillaries for cleanup of 1 to 5 µl of sample (Houthaeve, et al., 1995), and microscale highperformance liquid chromatography (HPLC) employing columns with internal diameters ≤1mm (Zhang et al., 1995). Second has been the development of ultra-low-flow (<50 nl/min) electrospray sources that enable 1 to 2 µl of a dilute sample containing femtomole per microliter concentrations of peptides to be ana-
Mass Spectrometry
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lyzed for periods of >60 min (Wilm and Mann, 1996). The long duration of analysis permits many experiments, including conventional MS experiments and MS/MS, to be carried out on a single sample loading. A third development has been the introduction of novel scanning methods whereby MS can be used to selectively detect specific protein modifications (Carr et al., 1993; Huddleston et al., 1993a,b). Fourth is the development of the so-called post-source decay (PSD) experiment (Kaufmann et al., 1994), which provides the ability to mass select particular peptide molecular ions and to analyze the metastable fragment ions from this precursor that are formed during the MALDI process. Like the ESI tandem MS experiment, PSD can provide a substantial amount of peptide sequence information (see section entitled What Is Tandem MS?). Finally, a variety of computer-based searching strategies have been developed whereby known proteins can be identified from protein and DNA databases by searching MS-derived molecular weights with and without partial sequence information (Henzel et al., 1993; Mann et al., 1993; Pappin et al., 1993; Wilm and Mann, 1994; Eng et al., 1994; Hall et al., 1996).
WHAT IS TANDEM MS?
Overview of Peptide and Protein Analysis by Mass Spectrometry
In tandem mass spectrometry (commonly referred to as MS/MS), two consecutive stages of mass analysis are used to detect secondary fragment ions that are formed from a particular precursor ion. The first stage serves to isolate a peptide precursor ion of interest based on its m/z, and the second stage to mass analyze the product ions formed by spontaneous or induced fragmentation of the selected precursor ion. Interpretation of the product-ion spectrum provides sequence information for the peptide selected. The manner in which product ions are produced and detected is different in MALDI and ESI mass spectrometry, and therefore is described in the following sections that are devoted to these specific techniques. MS/MS provides information that is highly complementary to that provided by Edman degradation; in addition, MS/MS has several advantages relative to Edman degradation for determining peptide sequence. First, MS/MS can produce sequence information for blocked or otherwise modified peptides that are either difficult or impossible to sequence by Edman degradation (for example, see the MS/MS spectrum of the phosphopeptide in Fig. 16.1.6). Second, the ability to individually select and fragment peptides in mixtures eliminates the
need for extensive purification of peptides prior to sequence analysis. Third, MS-based peptide sequencing is faster than Edman sequencing because it is not a stepwise process. All of the structural information, in the form of fragment ions, is presented in a single mass spectrum acquired in a single experiment. Finally, MSbased approaches are now at least as sensitive as Edman degradation (e.g., 1 to 5 pmol of peptide or protein is required), and state-of-theart MS-based sequencing by either ESI-MS or MALDI-MS is capable of providing sequence information at the 50- to 250-fmol level (Hunt et al., 1992; Wilm et al., 1996). Peptide fragment ions are produced in the mass spectrometer primarily by cleavage of the amide bonds (—CO-NH—) that join pairs of amino acid residues. The most commonly observed fragment ions and their nomenclature are shown in Figure 16.1.1, and a list of commonly observed low-mass fragment ions indicative of the presence of particular amino acid residues is presented in Table 16.1.2. The fragmentation of peptides in MS has been well described (Hunt et al., 1986; Biemann, 1988, 1990; Falick et al., 1993). The interested reader is referred to these studies and references therein for more detailed discussions of fragmentation and its possible mechanisms. The difference in mass between adjacent sequence ions of the same type defines an amino acid. Of the twenty commonly occurring amino acids, sixteen have unique masses, Leu and Ile have exactly the same mass, and Lys and Gln have nearly the same mass (Table A.1A.1 in APPENDIX 1A). A mass difference between two adjacent sequence ions of the same fragment type that does not correspond to the mass of an amino acid residue may represent the presence of a post-translational modification. A list of common post-translational modifications and their masses is given in Table A.1A.3 in APPENDIX 1A. Two caveats to the above discussion on MS/MS should be noted. First, although it is generally very easy to obtain the internal sequence of a peptide by this procedure, it is often difficult to determine the order of the first two and sometimes the last few residues in the peptide without additional experiments. Several methods involving reanalysis by MS/MS after chemical derivatization have been developed to help sort out the sequence at the ends of the peptide when required (Sherman et al., 1995; Gulden et al., 1996). Alternatively, there are several sequence ladder approaches, using either chemical derivatives or peptidases, that can be useful for identifying the N- and C-ter-
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y3 +2H R1 H2N CH
CO
y2 +2H
y1 +2H
R2
R3
NH CH CO a2
NH CH CO a3
b2
b1
R4 NH CH
COOH
b3
sequence-specific ions an
H
(NH
CHR
+ CO)n –1 NH = CHRn
bn
H
(NH
CHR
+ CO)n –1 NH = CHRn C ≡ O
+H yn
H
(NH
CHR
CO)n
OH
non-sequence-specific ions
internal acyl ions
H2N CHR2
amino acid immonium ions
CO
NH CHR3
+ H2N = CHRn
+ C≡O
Figure 16.1.1 Nomenclature for peptide fragment ions. R represents the variable amino acid side chain (see APPENDIX 1A). The non-sequence-specific ions are designated by the single-letter amino acid codes (Table A.1A.1) for the R groups they include.
Table 16.1.2
Common Low-Mass Ions Characteristic of Amino Acidsa
m/z
Amino acid present
70, 87, 100, 112, 129 70 (w/o 87, 100, and 112) 72 84 (w/o 101) 86 101 (w/o 84) 102 104 106 110 120 133 134 136 147 159 216
Arginine Proline Valine Lysine Leucine or isoleucine Glutamine Glutamic acid Methionine Pyridylethyl cysteine Histidine Phenylalanine Carbamidomethylated cysteine Carboxymethylated cysteine Tyrosine Acrylamide-modified cysteine Tryptophan Phosphotyrosine
aUnderlining indicates more abundant ions; w/o, without.
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minal residues (Chait et al., 1993; Bartlet-Jones et al., 1994; Thiede et al., 1995; Patterson et al., 1995). The second caveat is that, unlike Edman degradation, MS cannot at present be used reliably to determine the NH2-terminal sequence of proteins directly, although several successful attempts have been made (Speir et al., 1995).
MALDI-MS
Overview of Peptide and Protein Analysis by Mass Spectrometry
Matrix-assisted laser desorption/ionization (MALDI)-MS as commonly practiced today was first described by Karas and Hillenkamp (1988). They demonstrated that by cocrystallizing analyte molecules with a large molar excess of a small, UV-absorbing organic acid, large proteins such as serum albumin could be desorbed and ionized by short, intense pulses from a UV laser and detected using a TOF mass spectrometer. Refinement of the technique was rapid. New laser wavelengths and correspondingly optimized matrices were introduced (Beavis et al., 1992). In less than two years, proteins >100 kDa were being routinely analyzed with subpicomole sensitivity (Hillenkamp et al., 1991) and molecular weights could be assigned with an accuracy of ∼0.1%. Beavis and Chait (1990) demonstrated that for small proteins (<30 kDa) mass accuracies on the order of 0.01% to 0.02% could be achieved by coadding peptides or proteins of known mass to the sample to serve as internal mass reference standards (Beavis and Chait, 1990). This is 2 to 3 orders of magnitude more accurate than SDSPAGE. MALDI-MS has also been shown to be extremely useful for the analysis of peptides (Billeci and Stults, 1993). Unlike ESI, MALDI produces predominately singly charged molecular ions from peptides and proteins (although larger proteins can produce multiply charged ions as well; see Fig. 16.1.8), making the analysis of mixtures very much more straightforward. The ease of spectral interpretation combined with femtomole sensitivity and a tolerance for many of the common biological buffers makes MALDI an obvious choice for the analysis of unfractionated enzyme digests. Recent developments in the use of MALDI-MS for the analysis of peptides and proteins have been reviewed (Mann and Talbo, 1996; Wang and Chait, 1994). The MALDI process can result in extensive fragmentation of the sample. Decomposition that occurs in the ion source, upstream from the field-free region of the flight tube, is referred to as “prompt fragmentation” and can be ob-
served in a linear spectrum. Ions formed by prompt fragmentation are usually of low abundance, and are formed when very high laser irradiance is used. For example, disulfide bonds have been demonstrated to undergo prompt fragmentation to yield the free peptides (Zhou et al., 1993; Patterson and Katta, 1994). Fragmentation that occurs after the source, in the flight tube, is referred to as metastable decay or post-source decay (Kaufmann et al., 1994). These fragment ions, which are often abundant, cannot be observed in a linear TOF instrument but can be observed with the use of a reflectron. As described below, they can be very useful for peptide sequencing. The mechanism by which MALDI operates is still a matter of some debate (Dreisewerd et al., 1995). The one point upon which everyone agrees, however, is that the matrix is critical to the success of the experiment. In its simplest form, sample preparation for MALDI involves mixing the sample with a large (104) molar excess of matrix and applying 0.5 to 2.0 µl of this solution to the target, where it air dries. Once in the mass spectrometer, the sample is bombarded with photons of UV light. The matrices, all compounds that are strongly UV-absorbing at the designated wavelength, serve three major functions in the experiment. First, the large excess of matrix separates the analyte molecules from one another, thereby reducing intramolecular interactions. Second, the matrix rapidly absorbs large amounts of energy from the incoming photons, resulting in an explosive breakdown of the matrix-analyte lattice. This explosion sends both matrix and analyte molecules into the gas phase. Third, the matrix is necessary for ionization of the analyte molecules. Transfer of protons from the matrix to the peptide and protein molecules is believed to occur via gas-phase reactions in the dense cloud that forms above the target during laser irradiance. The choice of matrix depends on the irradiance wavelength and the type of sample to be analyzed. A N2 laser operating in the UV at 337 nm is the most typical laser in use on commercial MALDI instruments. Matrices that have been shown to be routinely useful at 337 nm are 3,5-dimethoxy-4-hydroxy-trans-cinnamic acid (sinapinic acid), 2,5-dihydroxybenzoic acid, and α-cyano-4-hydroxy-trans-cinnamic acid (Hillenkamp et al., 1991; Beavis et al., 1992). Lasers that operate in the infrared at 2.94 µm have also been shown to be useful in MALDI applications, particularly for the analysis of proteins directly from gels or blots (Strupat et al., 1994). MALDI has been dem-
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A
laser light linear detector
precursor ions
ion source
B
charged fragments
laser light
linear detector
ion source reflectron (ion mirror)
reflecting detector
Figure 16.1.2 Schematic diagrams of linear (A) and reflectron (B) MALDI-TOF mass spectrometers. The flight paths of precursor and fragment ions in each instrument are shown.
onstrated on magnetic-sector (Annan et al., 1992; Medzihradszky et al., 1996), ion-trap (Qin and Chait, 1995), and Fourier-transform ion-cyclotron resonance (Wu et al., 1995) mass spectrometers, but by far the most common application of MALDI has been on TOF instruments. Typical TOF mass spectrometers can be classified into two broad categories, linear and reflector-type instruments. A simple linear instrument (Fig. 16.1.2, panel A) consists of an ion source, where the MALDI process takes place; a flight tube, where ions of different masses are separated from one another; and a detector. All of these components, with the exception of the laser, are housed in a high vacuum maintained at 10−5 to 10−8 Torr. After ions are formed by the MALDI process, they are accelerated out of the source under the influence of a strong electric field. All of the ions have essentially the same final kinetic energy upon entering the flight tube; thus, the arrival time of a given ion at the linear detector
will vary depending on its mass. These relationships are mathematically expressed by classical equation for kinetic energy (here expressed in two forms), where E is the kinetic energy of an ion, m is its mass, and v is its velocity: E = 1⁄2 mv2 v = (2E/m)0.5
These equations tell us that because all ions of the same charge have been accelerated to the same energy, their velocities (and therefore, their flight times) will be inversely proportional to the square root of their molecular masses. Thus ions of greater mass will travel slower than lighter ones and will therefore reach the detector later. Simple linear MALDI-TOF instruments are rather low-resolution mass spectrometers, operating at a resolving power of ∼450 to 600 for peptides and 50 to 400 for proteins. For peptides, this means that one cannot measure the distribution of the various naturally occurring isotopes for each peptide, or separate peptides
Mass Spectrometry
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Overview of Peptide and Protein Analysis by Mass Spectrometry
which differ in mass by only a few daltons. It also means that given a resolution (R) of 400, it will be difficult to resolve two proteins >30 kDa that differ only by an initiator Met residue (e.g., see Fig. 16.1.8). These issues are considered in more detail in the discussion of Fundamentals of Mass Measurement Accuracy and Mass Resolution. Several important considerations relating to the production of ions in the MALDI source limit the resolving power of linear TOF instruments. The most important of these is the initial kinetic energy spread of individual ion populations: as a result of this spread, ions of any particular mass will, after reaching final accelerating voltage, exhibit a spread in velocities. This means that various members of the same ion population will arrive at the detector at slightly different times, and that in turn gives rise to a broader peak for each ion than would be observed if there were no initial kinetic energy spread. Because the initial energy spread is mass dependent, peaks for highermass ions will be disproportionally broader than those for lower-mass ions. Resolution in a linear TOF instrument can be improved by increasing the length of the flight tube. This enhances the time dispersion of ions of different m/z; unfortunately, it also increases the spread of arrival times for ions of the same m/z due to the initial kinetic energy distribution. This energy spread is typically reduced in magnitude by increasing the final accelerating voltage. A more effective way to correct for energy distributions is through the use of an ion mirror, or reflectron. A reflectron TOF instrument (Fig. 16.1.2, panel B) corrects for the initial energy spread by acting as an energy-focusing device. The reflector works by slowing an ion down until it stops (the back of the reflector is at a voltage slightly higher than the source-accelerating voltage), turning it around, and then reaccelerating it back out to a second detector. Ions with an initial kinetic energy (and velocity) slightly lower than full accelerating potential will not penetrate the reflector as deeply, and will therefore turn around sooner, catching up to those ions with full kinetic energy. Ions with slightly greater energies (and thus higher velocities) will penetrate more deeply into the reflector, be turned around later, and have their flight times retarded, allowing the other ions to catch up. All of this causes ions of a given mass-to-charge ratio to be spatially focused into packets having flight times that are closer together, thus improving the resolution. This
improvement in resolution is illustrated in Figure 16.1.3, which shows the molecular ion region of the linear (panel A) and reflector (panel B) spectra of a synthetic tyrosine phosphorylated peptide TRDIYETDpYYRK. The isotopically resolved ion cluster in the reflector spectrum shows the contributions of all of the isotopes of C, H, N, O, and P that make up the elemental composition of this peptide. The improvement in resolution is most noticeable at masses of ∼3000 and lower. For larger molecules, where the resolution is no longer sufficient to provide separation of the isotopes, the natural width of the unresolved isotopic envelope has a large influence on the apparent resolution of the instrument, as explained in the discussion of Fundamentals of Mass Measurement Accuracy and Mass Resolution. In addition to resolution, mass accuracy is also improved when spectra are recorded in the reflector mode (Vorm and Mann, 1994). Tremendous improvement in the resolution achievable on MALDI-TOF instruments has recently been demonstrated using a focusing technique developed many years ago by Wiley and McLaren (1953). This method, referred to as time-lag focusing or “delayed extraction,” has been shown to provide a resolution of ∼2000 to 4000 (full-width, half-maximum; or FWHM) in linear mode for peptides and resolution of ∼3000 to 6000 FWHM in reflecting mode, on a 2-meter-long instrument (Brown and Lennon, 1995; Whittal and Li, 1995; King et al., 1995; Vestal et al., 1995). In a delayed extraction source, ions are created in a field-free region and allowed to spread out before an extraction voltage is applied and the ions are accelerated into the drift tube. The delayed extraction also probably limits the number of collisions the ions undergo on their way out of the source, thereby reducing additional peak broadening caused by metastable decomposition. Mass accuracy with the use of an internal reference standard may also be improved with the use of the delayed extraction source. Assigning an accurate m/z value to an ion whose flight time has been measured requires that the mass spectrometer be properly calibrated. This is usually accomplished by measuring the flight times of two compounds whose molecular weights are accurately known. In practice, it is usually best to select two compounds with molecular weights that bracket the mass range over which samples are to be examined. For molecules >10,000 Da, the [M+H]+ and the [M+2H]2+ ions of a well-
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A
1690 1695 1700 1705 1710 1715
Figure 16.1.3 Molecular ion region from MALDI spectra of the peptide TRDIYETDpYYRK (monoisotopic Mr = 1701.8) recorded in the (+) ion mode. Spectra were recorded in (A) the linear mode, where the average mass is measured at the centroid of the unresolved isotopic cluster, and (B) the reflector mode, where the monoisotopic mass is measured at the centroid of the first peak in the cluster. This peak contains the lightest isotopes of the elements present (see discussion of Fundamentals of Mass Measurement Accuracy and Mass Resolution).
B
1690 1695 1700 1705 1710 1715 m /z
characterized protein standard work very well for calibration (see UNIT 16.2). Application of an internal calibration, where the calibrant peaks are present in the same spectrum as the sample, can provide accuracies of 0.01% to 0.05% (e.g., 0.1 to 0.5 Da at m/z = 1000). Internal calibration is sometimes experimentally difficult owing to the need to adjust the amount of calibrant compounds added so that the calibrant peaks and the analyte peaks are simultaneously observed with reasonable intensity. MALDI mass spectra may also be calibrated externally; this approach is experimentally simpler. In this method calibrant compounds are measured using the same matrix and laser irradiance as were used for sample analysis. A mass accuracy of ∼0.1 to 0.2% (e.g., ±1 to 2 Da at m/z = 1000) can be achieved in linear mode using external calibration (Karas et al., 1989). In reflector mode, one can routinely expect mass accuracies on the order of 0.01% to 0.05% using external calibrations. In reflector mode it is also possible to use either the [M+H]+ ion or the [2M+H]+ ion (the protonated gas-phase dimer) of the matrix as one of the two calibrant
peaks, which means that only one calibrant compound needs to be added.
MALDI Post-Source Decay MS for Peptide Sequencing A peptide molecular ion that is sufficiently stable to be transported out of the ion source but insufficiently stable to survive the flight to the detector will decompose in the flight tube, giving rise to a series of fragment ions that are characteristic of the molecule’s original structure. This process is referred to as metastable decay or post-source decay. It is believed that molecular ions acquire excess internal energy necessary for fragmentation via multiple collisions with matrix ions in the source (Kaufmann et al., 1994). When a precursor ion decomposes in the flight tube, all of the fragment ions will have the same velocity as the precursor, but they will have only a fraction of its kinetic energy. This means that the precursor and metastable fragments will all strike the linear detector at the same time and will be detected at the same apparent mass. Thus metastable or post-sourcedecay fragment ions are never observed in a linear spectrum.
Mass Spectrometry
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Relative abundance (%)
A 100
2066
1284
50
804
1300
634
1705 1776 1550
1002 1250 1082
530
2168 2384
2705 2937
0
Relative abundance (%)
B
600
800 1000 1200 1400 1600 1800 2000 2200 2400 2600 2800
100 2066 y3 50
15 14 13 12 11 10 9
y4
8
7 6
5
4 3 2 1
I–F–Y–P–E–I–E–E–V–Q–A–L–D–D–T–E–R
yn
PEIE
y1 PEI y2
PEIEE y6 y5
y7
600
800
y8
y9 y10
y11
y12
y14
y15
0 200
400
1000 1200 m /z
1400
1600
1800
2000
Figure 16.1.4 MALDI-MS analysis of a tryptic digest of an 18-kDa protein (Ladner et al., 1996). A total of 30 pmol of protein was separated on an SDS-PAGE gel, the band of interest excised from the gel, and the protein digested in situ with trypsin; then 1⁄20 of the sample was analyzed by MALDI-MS. (A) MALDI mass spectrum of the tryptic peptides recorded in the linear mode; (B) MALDI-PSD spectrum of the [M+H]+ ion at m/z 2066. The peak was selected using an ion gate (R = 100). A nearly complete series of yn ions (see Fig. 16.1.1) makes it possible to deduce the sequence of residues 3 to 17.
Overview of Peptide and Protein Analysis by Mass Spectrometry
In contrast, a reflector TOF is capable of discriminating fragment ions by flight-time dispersion. Although all ions will have essentially the same velocity, the kinetic energy of a post-source-decay fragment ion will be determined by the ratio of its mass to the mass of the precursor. Thus the fragments, which have less kinetic energy than the precursor, will not penetrate as deeply into the ion mirror, and so will have their flight times decreased relative to the precursor. The fragments are therefore detected at a lower mass. By successively lowering the reflector voltage, thereby bringing lower- and lower-mass fragment ions into focus, a complete post-source decay fragment ion spectrum can be acquired in segments. The individual segments are calibrated and assembled afterward automatically by the data system. The energy-focusing features of reflectrontype mass spectrometers make it possible to use post-source decay for peptide sequencing (Kaufmann et al., 1994). Post-source decay
spectra resemble low-energy collision-induced dissociation (CID) spectra, similar to those commonly recorded on triple-quadrupole mass spectrometers (Hunt et al., 1986). The spectra contain mainly bn and yn ions, often with very abundant internal fragments and immonium ions (see Fig 16.1.1 and Table 16.1.2). The use of an ion gate makes it possible to perform a low-resolution (R = 100) selection of a precursor ion of interest from an unfractionated mixture, such as a protein digest. This approach is very powerful for providing internal sequence information on in situ–digested proteins that have been purified by SDS-PAGE (see Fig. 16.1.4 for an example).
ESI-MS In electrospray ionization (ESI)-MS, ions are formed from peptides and proteins by spraying a dilute solution of these analytes at atmospheric pressure from the tip of a fine metal capillary (Fig. 16.1.5). The spray pro-
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possible sample inlets: syringe pump sample injection loop HPLC
triple-quadrupole mass spectrometer
atmosphere (760 Torr)
vacuum (10 – 6 Torr)
liquid 1st quadrupole collision 2nd quadrupole mass analyzer cell mass analyzer
capillary electrophoresis
data system expansion of the ion formation and sampling regions
electrospray needle
nitrogen drying gas vacuum
atmosphere
3–5 kV
liquid nebulizing gas
droplets containing solvated ions
ions sampling orifice counter electrode
Figure 16.1.5 Schematic diagram of a triple-quadrupole mass spectrometer (top) and the electrospray interface (bottom).
cess, often assisted by pneumatic nebulization, creates a fine mist of droplets (Fenn et al., 1990; Kebarle and Tang, 1993). The droplets are formed in a very high electric field created by applying a high voltage (∼4 kV) to either the spray tip (Fig. 16.1.5) or the counter electrode; in this process, the droplets become highly charged. Solvent evaporation is rapid from these small droplets. As the droplets evaporate, the peptide and protein molecules in the droplets pick up one, two, or more protons from the solvent to form singly or, more frequently, multiply charged ions (e.g., [M+H]1+, [M+2H]2+, etc.). The number of charges acquired by a molecule is roughly equivalent to the number of possible sites of proton attachment. As the droplets continue to shrink, the charge density on the surface of each one increases to the point where charge repulsion overcomes the forces holding together the droplet and the solvated ions contained within it. Ions are then “emitted” or “evaporated” from the droplet surface. The ions are sampled into the high-vacuum region of the mass spectrometer for mass analysis and detection, most often using a quadrupole (or triple-quadrupole) mass analyzer (Fig. 16.1.5). Usually little or no fragmentation is observed in the normal ESI mass
spectra of peptides. The typical solvent for peptides and proteins is a mixture of water, an organic modifier such as CH3CN, and up to a few percent by volume of acetic, trifluoroacetic, or another volatile acid (the latter included to enhance ionization of sample constituents). Because the ions are produced at atmospheric pressure from flowing liquid streams, ESI is ideally suited for on-line coupling to high-performance liquid chromatography, making it possible to analyze mixtures of peptides and proteins rapidly (see below). As mentioned above, a key feature of the electrospray process is the formation of multiply charged molecular species from analytes that contain more than one possible site of proton attachment (Fenn et al., 1990; Smith et al., 1991). Proteins usually exhibit a characteristic series of multiply charged ions (e.g., see Fig. 16.1.8). As one proton (i.e., one charge) is typically attached for each 1000 Da of molecular mass, the ion series for a protein, even a large protein, will fall in the m/z range of 800 to 3000. This makes it possible for simple instruments like quadrupole mass spectrometers to be used for mass analysis of ions produced by ESI. The molecular mass of the protein can easily be calculated using the observed masses of any
Mass Spectrometry
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Relative abundance (%)
A
100 75
2+ 673.8
2+ (– H, +Na) 2+ (– H, +K)
50
1+ 805.4
25
1+ 1346.6 600
800
Relative abundance (%)
B
1000
b3 y2
100
y4 75 50
b2
y6
1200 2+
10 9 8 7 6 5 4 3 2
D G D G pY I S A A E L R
y7
b5
bn
2 3 4 5 6 7 8 9 10
y3 b7 b4
200
yn
y5
y1
25
1400
400
b9
b6
600
y8 b 10 y9
b8 800
y10 1000
1200
1400
m /z
Figure 16.1.6 Electrospray analysis of an HPLC fraction from the tryptic digest of a 17-kDa phosphoprotein. The protein was purified by SDS-PAGE, electroblotted onto PVDF membrane, eluted, digested with trypsin, and the peptides analyzed by on-line LC-ESI-MS using a phosphopeptide-selective scanning method (Huddleston et al., 1993b). The column flow was split after UV detection, with 10% going to the mass spectrometer and 90% to a fraction collector. (A) Nanoelectrospray mass spectrum of a putative phosphopeptide-containing fraction. Peaks labeled (+Na) and (+K) correspond to [M + H + Na]2+ and [M + H + K]2+, respectively. (B) Collision-induced decomposition (CID) spectrum of the doubly charged ion ([M+2H]2+) at m/z = 673.8. This spectrum confirms the sequence of the peptide and identifies the site of phosphorylation as Tyr5.
Overview of Peptide and Protein Analysis by Mass Spectrometry
two adjacent ions in the series (Covey et al., 1988; Mann et al., 1989; Reinhold and Reinhold, 1992). Normally these calculations are performed automatically by the data system. Because each multiply charged peak provides an independent measure of the molecular mass of the protein, an ion series from a single experiment provides a measure of the mass measurement precision. Most data systems also provide algorithms for transforming the mass-tocharge axis into a molecular mass axis for simplifying interpretation and presentation of the data. Proteins with molecular masses of up to 150,000 Da have been successfully analyzed by ESI using commercially available quadrupole instrumentation. Mass assignment accuracies of 0.01% are routine (see Table 16.1.1 and discussion of Fundamentals of Mass Measure-
ment Accuracy and Mass Resolution). ESI has also been successfully interfaced with ion-trap mass spectrometers (Korner et al., 1996), Fourier-transform ion-cyclotron resonance mass spectrometers (Winger et al., 1993; O’Conner et al., 1995; Wu et al., 1995; Smith et al., 1996), and TOF mass spectrometers (Verentchikov et al., 1994; Mirgorodskaya et al., 1994). In the case of peptides, the maximum number of charges observed often correlates reasonably well with the number of amino acid side chains that can readily accept a proton at the low pH of the analyte stream (i.e., Arg, Lys, and His, plus the free amino terminus). In the case of tryptic peptides, at least two charge sites are normally present, and tryptic peptides with molecular weights of 800 to 3000 typically
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exhibit a major, if not dominant, [M+2H]2+ ion. For example, the ESI mass spectrum of an HPLC fraction from a tryptic digest of a phosphoprotein shows a simple mixture of peptides, in which doubly charged forms of the major peptide predominate (Fig. 16.1.6). In this spectrum, in addition to proton attachment, doubly charged ions due to Na and K adduction are also abundant. In general, the smaller the peptide, the more likely it is that the singly charged [M+H]+ ion will also be observed. As the size of the peptide increases, triply and highercharged states of the peptide will increase in abundance in the spectra while the abundance of ions with lower charge states will decrease (Fig. 16.1.7). Electrospray mass spectra are keen reminders that mass spectrometers measure the ratio of mass to charge (m/z) and not molecular mass directly. The peak at m/z = 1346.6 in Figure 16.1.6 (panel A) is the monoisotopic [M+H]+ of a peptide of molecular weight 1345.6 (see section on Fundamentals of Mass Measurement Accuracy and Mass Resolution for a discussion of monoisotopic mass). Evidence that this ion carries a single charge (1+) as opposed to a higher number of charges is obtained from the spacing of the isotope peaks, which are one m/z unit apart. The doubly charged form of this same peptide is observed on the m/z scale at [1345.6 +2]/2 = 673.8. The isotope peaks in the [M+2H]2+ isotope cluster are spaced 0.5 m/z units apart, indicating that these ions are doubly charged. Quadrupole mass analyzers can routinely achieve resolution sufficient to distinguish singly from doubly charged ions up to m/z = 3000 (if sufficient sample is available and the instrument is scanned slowly). This capability is particularly relevant to tryptic digests of proteins where, as noted above, the doubly charged ion may be the only ion observed for a given peptide. In the absence of the peak spacing information, the analyst could have difficulty determining if such an ion is singly or doubly charged. For ions with three or more charges, a charge series (e.g., 2+ and 3+) is usually present that permits the analyst to determine the charge state and mass of any given ion in the spectrum without the need to determine the isotopic peak spacing.
LC-ESI-MS Electrospray ionization has made on-line liquid chromatography (LC)-MS a practical and robust method for sample introduction because gas-phase ions, as opposed to liquid droplets, are what is mainly being sampled by the
instrument. The ideal milieu for presenting a sample to any mass spectrometer, regardless of the ionization mechanism, is in mixtures of highly pure water and volatile organic solvents containing low levels of volatile acid or base. Fortunately, such mixtures (e.g., water, acetonitrile, and trifluoroacetic acid at 0.05% to 0.1% by volume) are also those that provide optimal separation of peptides and proteins by reversedphase liquid chromatography. From a practical standpoint, LC-MS reduces or eliminates the need to preparatively fractionate complex mixtures prior to MS, thereby saving valuable instrument time and preventing possible sample losses. LC-MS data may also be searched retrospectively for components present at very minor levels that may not have been considered worthy of collection based on a weak UV response. In an LC-MS experiment, mass spectra are recorded continuously as the components elute from the HPLC column. If the mass spectrometer is made to scan once every 4 sec, the LC-MS data file will contain ∼900 mass spectra. To identify regions of data that are likely to contain useful spectra, the mass spectrometer’s data system produces a plot of the total number of ions detected during each mass spectrum scan versus time (the scan number). This plot, which is called a mass chromatogram or a total ion current (TIC) trace, looks very much like the UV chromatogram (Fig. 16.1.7). If it is of interest to know whether a particular peptide is present in the digest and where it elutes, the analyst simply requests the data system to display a selected ion-current trace for the specific masses of the most likely charge states (e.g., 1+ to 4+). Figure 16.1.7 illustrates the type of data typically acquired during the LC-ESI-MS analysis of a protein digest. In this example, the protein is heterogeneously glycosylated at a single site, and the multiplicity of peaks evident is due to differing carbohydrates attached to a single tryptic peptide (Fig. 16.1.7, panel C). Because an ESI mass spectrometer behaves as a concentration-dependent detector, much like a UV detector, only a few percent of the total column effluent (∼1 to 5 µl/min) needs to be directed into the mass spectrometer. The remainder can be diverted, via a flow splitter, through an in-line UV detector to a fraction collector. This arrangement makes it possible to perform straightforward correlation of the UV trace with the total ion current trace and then analyze individual fractions further by MS/MS (see discussion of ESI MS/MS for Peptide Sequencing) or some other method.
Mass Spectrometry
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Relative intensity (%)
A
125 54.6
100 75 10.8
50
19.8
25 0 1 0.0
101 7.5
201 14.9
301 22.4
401 29.8
501 37.3
601 44.9
701 52.4
801 59.8
scan/time (min)
Relative intensity (%)
B
125
10.8 min (scans 317– 319)
1+
439.2
100
= Mr 438.2
75 50 [2M+H]+ 877.6
25 0
Relative intensity (%)
C
Relative intensity (%)
D
[M+2H]2+
125
2+
E292
100
75 50 25
1317.8 1398.8 x, y = 4 (Hex)x (HexNAc)y ( dHex) – N296 x = 3 y=4 204.0 366.2 R300 [M+3H]3+ x=5 933.0 y=4 1480.0
0 125
3+
2+
= Mr 2344.4
1173.2
= Mr 2846.8
4+
712.8
= Mr 2795.6
54.6 min (scans 731–74 4)
950.0
100
19.8 min (scans 261– 271)
2+
1424.4 1564.4
1+
2345.2
0 200
400
600
800 1000 1200 1400 1600 1800 2000 2200 m/z
Overview of Peptide and Protein Analysis by Mass Spectrometry
Figure 16.1.7 On-line LC-ESI-MS analysis of a tryptic digest of reduced and alkylated heavy chain (51 kDa) from a recombinant monoclonal antibody. (A) Total ion current trace showing the distribution of tryptic peptides. (B) ESI spectrum of the peak at 10.8 min showing a small singly charged peptide and its gas-phase dimer. (C) ESI spectrum of the peak at 19.8 min corresponding to a tryptic glycopeptide with heterogenous carbohydrate. At least five different carbohydrate structures are indicated. Abundant low-m/z ions at m/z = 204 and m/z = 366 are characteristic fragment ions produced from the carbohydrate (Roberts et al., 1995). (D) ESI spectrum of the peak at 54.6 min showing two large tryptic peptides, each represented by several different charge states.
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The UV response is also useful for estimating the amount of a peptide that has been injected on the column. This is generally not possible to do by MS without appropriate internal standards (see section entitled Is MS Data Quantitative?). LC-ESI-MS has become a key analytical method for the analysis of natural and recombinant proteins, and data from these experiments is now often included in the regulatory filings for new biopharmaceutical products (Carr et al., 1989; Palczewski et al., 1994; Watts et al., 1994; Stults et al., 1994; Rush et al., 1995; Schindler et al., 1995; Roberts et al., 1995; Resing et al., 1995; Hemling et al., 1996).
ESI MS/MS for Peptide Sequencing The principles of MS/MS using triple-quadrupole mass spectrometers have been reviewed (Yost and Boyd, 1990). Briefly, the first quadrupole (Fig. 16.1.5) acts as a mass filter to select specifically the [M+nH]n+ ion (where n is typically 1, 2, 3, or 4) of a peptide of interest and isolate it from other ions produced in the source. The selected precursor ion is then fragmented with a neutral gas such as nitrogen or argon in a quadrupole collision cell. This process is known as collision-induced decomposition. The fragment or product ions produced are then transmitted into the quadrupole second mass analyzer, which is being used as a scanning mass analyzer. The product ions are separated based on m/z and detected. The result is a product-ion mass spectrum of the selected precursor that is free of interferences from other components present in the mixture. An example of this process is shown in Figure 16.1.6. The fragmentation of this peptide reveals its sequence and the location of the phosphorylated residue. Fragmentation can also be induced in the ESI source by increasing the energetics of the ion sampling process. This is done by adjusting the lens potentials in the high-pressure region upstream from the first quadrupole mass filter. Ions with increased kinetic energy undergo decomposition via collisions with residual gas molecules (Katta et al., 1991). Structurally useful fragmentation can thus be obtained on a single-quadrupole instrument, with the caveat that because this method has no ability to preferentially select precursor ions, it requires a highly purified sample. Recently strategies have been developed that take advantage of this ability of the ESI interface to generate fragment ions prior to mass analysis. Selective detection of glycosylated peptides (Carr et al., 1993; Huddleston et al., 1993a), phosphorylated pep-
tides (Huddleston et al., 1993b; Till et al., 1994), and sulfated peptides (Bean et al., 1995) during on-line LC-ESI-MS analysis of protein digests can be accomplished by monitoring the total ion current trace for highly diagnostic low-m/z fragment ions produced in the ESI interface. These procedures are equally applicable to the detection of these modifications in unseparated mixtures.
IS MS DATA QUANTITATIVE? A question that frequently arises in the analysis of peptide or protein mixtures is whether the relative peak heights (or areas) are quantitatively representative of the relative solution concentrations of the sample components. Unfortunately, the answer to this question is no, unless appropriate internal reference standards are used. Due to differing ionization and desorption efficiencies, the absolute yields of different ions from one peptide in a mass spectrometer can vary by perhaps as much as 2 orders of magnitude, depending on the sequence, composition, and size of the peptides (Jesperson et al., 1995; Dunayevskiy et al., 1995; Cohen and Chait, 1996). Furthermore, strongly ionizing components can frequently suppress the ionization of more weakly ionizing components present in the same sample or chromatographic fraction. In MALDI, the matrix itself can strongly influence the detectability of specific sample constituents (see discussion in UNIT 16.2). Quantitation by mass spectrometry requires that the response of a specific sample component be determined relative to that of a reference compound added to the sample and analyzed under identical conditions. In certain cases, semiquantitative estimates of relative amounts of components may be made if the components are of nearly identical structure and their chemical differences do not significantly affect the overall charge or hydrophobic/hydrophilic character of the molecules. For example, the relative ratios of the variants of β-casein observed in the ESI mass spectrum (Fig. 16.1.8) are probably a good reflection of the relative solution concentrations of these variants, as they differ from one another by only one or two amino acid substitutions out of more than 200 amino acids. The smaller the molecule, the more likely it is that such estimates will be in error.
SAMPLE PREPARATION The ideal method for preparing samples for any type of MS analysis is reversed-phase highperformance liquid chromatography (RP-
Mass Spectrometry
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A
B 23,984 2
14+ 13+
Relative abundance (%)
[M +H]1+ 24,010 30
24,024 2
+
18+ 15 16+
24,093 2 12+
1400
1420
11+ 10+ 9+
20,000 22,000 24,000 26,000
800
1200
1600
8+
2000
2400
m /z Figure 16.1.8 Molecular weight determination of β-casein, a 24-kDa phosphoprotein with known sequence heterogeneity. (A) Linear MALDI-TOF spectrum. Molecular weight was determined from the centroid of the smoothed peak using an external calibration. (B) ESI spectrum. The inset, an expansion of the region around the 14+ charge state, reveals the presence of at least three variants. The molecular weights shown are the averages of determinations made from ten individual charge states.
Overview of Peptide and Protein Analysis by Mass Spectrometry
HPLC). If purification by RP-HPLC is not an option because of known problems with sample loss, then the cleanup protocol employed should produce a sample composition that resembles as closely as possible that of a sample which has been purified by RP-HPLC. That is, the sample should contain the least amount possible of buffers, salts, detergents, and anything else besides water, organic modifier, and volatile acid or base. The quality of the data obtained using ESIMS is very dependent on the type and concentration of excipients present in the sample. If buffers are necessary for protein solubility or enzymatic digestion, it is best to use a volatile buffer such as ammonium bicarbonate or ammonium acetate at a concentration of ≤30 mM. In general it is best to avoid using any ionic detergents (surfactants), even though it is possible to obtain spectra for some proteins when these are included as long as the surfactant concentration is kept <0.01%. ESI tolerates nonionic detergents better. Analytically useful data may be obtained for sample solutions containing certain nonionic detergents at concen-
trations between 0.01% and 0.1% (Ogorzalek Loo et al., 1994). Samples containing unacceptable amounts of buffers, salts, and detergents need to be purified prior to ESI-MS analysis. If the sample can be eluted from an RP column, some type of on-line procedure is both rapid and efficient. For water-soluble salts and buffers, a short, small-i.d. RP cartridge installed in place of the sample loop on a HPLC injector serves to both concentrate and purify samples (Stoney and Nugent, 1995). By placing the trapping cartridge in the injector loop, it is possible to wash the buffers and salts into waste rather than into the mass spectrometer. After loading the aqueous sample onto the trap (sample volume is no longer important except for very hydrophilic sample components), the trap is washed with 1% to 5% organic solvent and the salts and buffers are diverted to the waste line. The trap column is then is put in-line with the solvent delivery pump by switching the valve, and the sample is eluted into the mass spectrometer with the appropriate solvent. This is also a
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convenient way to analyze dilute solutions, as the trap serves to preconcentrate the sample. The same approach can be used to purify samples containing anionic and nonionic detergents (Stoney and Nugent, 1995). However, on-line use of detergent columns require that this step be done in tandem with an RP column, because either the sample will have to be eluted with a buffer and thus will need to be desalted prior to introduction into the mass spectrometer, or it will not be retained on the column and will need to be trapped on an RP column. An alternative approach for removing SDS from samples is to load the sample, dissolved in a high concentration of organic solvent, onto a polyhydroxyethyl aspartamide column (Jeno et al., 1993). Under these conditions, peptides and proteins are retained while the SDS passes through with the solvent front. The sample is then eluted using a gradient of decreasing organic strength. All of the purification strategies described above can, of course, also be performed offline. In either case, the use of small-bore HPLC columns and a low-flow HPLC system will provide better results with sample amounts in the picomole range or below. Because the ESI mass spectrometer acts as a concentration-dependent detector, the highest sensitivity will be achieved using the smallest-i.d. columns. For off-line cleanup, the use of the lower flow rates required for small-bore columns means that samples are collected in smaller volumes. This can be important for small amounts of sample, where losses associated with reducing solvent volumes prior to the next step of analysis often are great. The authors have found that 0.5- to 1-mm-i.d. columns provide a practical compromise between on-line sensitivity and easy collection of fractions using a post-column flow splitter. A number of groups have developed sample purification and fraction collection methods that use packed fused silica capillary columns and solvent flow rates of 0.1 to 5.0 µl/min (Zhang et al., 1995; Gulden et al., 1996) or micropipets filled with packing material (Houthaeve, et al., 1995). Other approaches to protein purification for MS include ultrafiltration (UNIT 4.4), TCA precipitation, size exclusion chromatography (UNIT 8.3), and microdialysis (UNIT 4.4). Among these alternatives, the authors find the easiest and most reliable methods for desalting samples or effecting a buffer change to be ultrafiltration using membranes of varying molecular weight cutoff (MWCO), and small-column (1-mmi.d.) size-exclusion chromatography. Final vol-
umes of 25 µl can be achieved using the ultrafiltration devices. If the protein is being purified for subsequent proteolytic digestion with the ultimate goal being mass spectrometry–based peptide mapping, a suitable approach is SDS-PAGE followed by in situ enzymatic digestion (UNIT 11.3; Williams and Stone, 1995; Wilm et al., 1996; Schevchenko et al., 1996). The broad applicability and high resolving power of SDSPAGE applied to protein mixtures makes this approach very useful for achieving difficult separations or for working with proteins that have poor solubility characteristics. Electrophoresing as little as a few micrograms of individual protein on an analytical gel will usually provide an informative peptide map. Following SDS-PAGE, the protein can also be blotted onto a membrane and enzymatically digested (UNIT 11.2; Patterson, 1995; Hess et al., 1993). Recovery of peptides from individual proteins appears to be similar regardless of whether they were produced from an in-gel digest or digestion off a membrane. If the peptide map is to be recorded using MALDI, the peptides can be sampled either directly from the digest or from the peptide extract. If the map is to be recorded using ES, then on-line LC-MS is usually the best approach. Although MALDI is very much more tolerant of the presence of salts, buffers, and some detergents, it too generally benefits from having the sample in the purest form possible, especially when dealing with subpicomole amounts of proteins or low femtomole amounts of peptides. All of the methods discussed above work equally well for preparing samples for MALDI-MS analysis.
FUNDAMENTALS OF MASS MEASUREMENT ACCURACY AND MASS RESOLUTION Monoisotopic Mass and Average Mass Most of the elements that comprise typical biological molecules have more than one naturally occurring isotope, each with a unique mass and natural abundance. For example, carbon has two principle isotopes, 12C and 13C (see Table 16.1.3). The lighter, more abundant isotope of carbon has, by definition, a mass of 12.000000 and a natural abundance of 98.9%, while the heavier isotope has a mass of 13.003355 and a natural abundance of 1.1%. In a mass spectrometer one measures a large, statistical sampling of the molecules present and, therefore, observes all of the isotopes for each
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Table 16.1.3
Isotope 1H 2H 12C 13C 14N 15N 16O 17O 18O 19F 23Na 28Si 29Si 30Si 31P 32S 33S 34S 36S 35Cl 37Cl 39K 40K 41K 79Br 81Br 127I
Isotopic Mass and Abundance Values
Natural abundance (%)b
Massa 1.007 825 035 2.014 101 779 12.000 000 000 13.003 354 826 14.003 074 002 15.000 108 97 15.994 914 63 16.999 131 2 17.999 160 3 18.998 403 22 22.989 767 7 27.976 927 1 28.976 494 9 29.973 770 1 30.973 762 0 31.972 070 698 32.971 458 428 33.967 866 650 35.967 080 620 34.968 852 728 36.965 902 619 38.963 707 4 39.963 999 2 40.961 825 4 78.918 336 1 80.916 289 126.904 473
99.985 0.015 98.90 1.10 99.634 0.366 99.762 0.038 0.200 100 100 92.23 4.67 3.10 100 95.02 0.75 4.21 0.02 75.77 24.23 93.2581 0.0117 6.7302 50.69 49.31 100
aData from Wapstra and Audi (1985). Standard errors are given in this
article but are omitted from the present table. bLide (1995).
Overview of Peptide and Protein Analysis by Mass Spectrometry
element present. The monoisotopic mass of a molecule is the sum of the accurate masses (including the decimal component, referred to as the mass defect) for the most abundant isotope of each element present (Yergey et al., 1983). As the number of atoms of any given element increases, the percentage of the population of molecules having one or more atoms of a heavier isotope of this element also increases. The most significant contributor to the isotopic peak pattern for biomolecules is the 13C isotope of carbon. For peptides with molecular masses <2000 Da, the first peak in the resolved isotopic cluster corresponding to the all-12C species will be the most abundant (Fig. 16.1.9, panel A). However, for peptides with molecular masses ≥2500 Da, the first peak in the isotopic cluster will no longer be the most
abundant (Fig. 16.1.9, panel B). In this example, the species containing one 13C atom is now the most abundant. For proteins, the heavy isotopes of nitrogen, sulfur, and oxygen (in addition to carbon) make significant contributions to the mass, and, as a result, the monoisotopic peak in the molecular ion cluster appears to vanish (Fig. 16.1.9, panels C and D). In this case a different definition for the mass of the molecule must be used. Whenever it is not possible to observe the all-12C peak for a molecular ion, the average mass should be used. The average mass of a molecule is the sum of the chemical average masses of the elements present (Fig. 16.1.9, peaks labeled A). The chemical average mass of an element is the sum of the abundanceweighted masses of all of its stable isotopes
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Mono.
Peak top
Relative abundance (%)
100
A
B
Ave.
Ave. Mono.
R = 500
75
R = 500 R = 1000
R = 1000
R = 5000
50
R = 5000 25
0 1268
1270
1272
1274
1276 2532 2534 2536 2538 2540 2542 2544
Ave.
C
Ave.
D
Relative abundance (%)
100
R = 500
R = 500
R = 1000
75
R = 1000 R = 5000
R = 5000 50
R = 20,000 25
Mono. Mono.
0 5720 5724 5728 5732 5736 5740 5744 23,940 23,960 23,980 24,000 24,020
m /z
Figure 16.1.9 Effect of resolution (10% valley definition) on the appearance and relative peak intensity of the isotopic envelopes for representative peptides and proteins in four different mass ranges. Abbreviations: R, resolution; M, monoisotopic mass (Mono.); A, average mass (Ave.); P, peak top. (A) C60H86N16O15: M = 1270.7, A = 1271.5, P = 1270.8. (B) C115H161N31N35: M = 2536.2, A = 2537.7. (C) C254H377N65O75S6: M = 5729.6, A = 5733.6. (D) C1080H1697N2680325P5S6: M = 23968.2, A = 23983.3.
(Yergey et al., 1983). For example, the isotopeweighted average mass of carbon is 12.0111, of hydrogen is 1.0079, of nitrogen is 14.0067, of oxygen is 15.9994, of sulfur is 32.0666, and of phosphorus is 30.9738. These masses are known with less precision than the monoisotopic masses of the elements because of variations in the natural abundances of the isotopes. The theoretical appearance of a molecule’s isotope envelope may be precisely modeled by solving a polynomial expression that calculates
the abundance-weighted sum of the isotopes of each element to the molecular ion cluster (Yergey, 1983). Nearly all modern mass spectrometer data systems come with programs that perform these calculations and can simulate the appearance of the resulting isotope pattern at different instrumental resolving powers.
Resolution Whether or not one observes the individual isotopes that make up the molecular ion cluster
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100
Relative abundance (%)
80
60 Max 40
∆m (50% Max = FWHM)
10% valley
20
∆m (5% of Max)
m1
m2 m /z
Figure 16.1.10 Schematic representation of the two common definitions of resolution used in mass spectrometry—10% valley and full-width, half-maximum (FWHM).
Overview of Peptide and Protein Analysis by Mass Spectrometry
for a given compound depends on the mass resolution of the instrument being used. The resolution of the mass analyzer is a measure of its ability to separate signals of adjacent molecular mass. Mass resolution is often expressed as the ratio m/∆m, where m and m+∆m are the masses (in atomic mass units or Da) of two adjacent peaks of approximately equal intensity in the mass spectrum (Fig. 16.1.10). There are presently two definitions of resolution in widespread use. In the “10% valley” definition, the two overlapping, adjacent peaks each contribute 5% to the resulting 10% valley between them (Fig. 16.1.10). The resolution of the mass analyzer may also be determined using a single peak by measuring the peak width (in m/z) at the 5% level and dividing this number into the m/z of the peak. The 10% valley definition is most frequently used for molecules with masses <5000 Da, where it is possible on certain analyzers to resolve the isotope peaks. For unresolved isotopic clusters the “fullwidth, half-maximum” (FWHM) definition is used. By this definition, the resolution is equal to the mass of the peak (in m/z) divided by the width (in m/z) of the isotopic envelope measured at the half-height of the peak (Fig. 16.1.10). An unfortunate consequence of the use of this definition is that, given an FWHM
resolving value, it is more difficult to visualize what the separation between two peaks of a given masses will be. For example, a resolution of 1000 by the FWHM definition means that a peak at mass 1001 will be unresolved from the peak at mass 1000. Creation of a detectable separation between these two peaks would require a FWHM resolution >1200. A useful rule of thumb is that the value for the resolution determined using the FWHM definition is approximately twice that obtained using the 10% valley definition. For example, a resolution of 1000 using the 10% valley definition is approximately equivalent to a resolution of 2000 using the FWHM definition (i.e., the measured degree of separation of the peaks in Da/z is identical).
Mass Accuracy Getting as accurate a molecular weight as possible is, of course, a central goal of any MS experiment. Here mass accuracy is taken to mean the difference in mass between the experimentally measured value and the theoretical value. By definition, the mass measurement accuracy can be directly determined only for compounds of known elemental composition and, therefore, known molecular mass. For unknown samples, the accuracy of the mass
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determination is approximated by measuring the mass of a peptide or protein of known elemental composition and similar mass to the unknown. It is important that these measurements be made under experimental conditions as close as possible to those used for analysis of the unknown. How useful this estimate of the mass measurement accuracy is depends on how well the instrument has been calibrated and on the reproducibility of the measurement. The reproducibility, or precision, of the measurement is best evaluated by making replicate measurements of the sample. However, in cases where the amount of sample is limited or the data is too noisy, a reasonable approach is to estimate the precision from replicate measurements of a known peak of mass and intensity similar to the unknown. It cannot be overemphasized that the mass spectrometer’s performance with respect to mass measurement accuracy and precision must be regularly evaluated using well-characterized standards covering the mass range of interest in order to provide confidence in the measurements of unknowns being made. It is desirable to achieve a mass measurement precision of ±0.5 Da or less for peptides and ±1 Da for proteins. In practice, it is relatively easy to achieve the stated value for peptides but more difficult for proteins (Table 16.1.1). Whether the monoisotopic mass or the average mass should be used when measuring and reporting molecular weights will depend on the mass of the substance and the resolving power of the mass spectrometer (see discussion of Interplay Between Resolution and Molecular Weight). The accuracy of the mass measurement may be stated in several different ways: as either an absolute mass error or a percentage of the measured mass (e.g., molecular mass = 1000 ±0.1 Da or ±0.01%), or as parts per million (e.g., molecular mass = 1000 ±100 ppm). As the mass being considered increases, the absolute mass error corresponding to the percent or ppm error will also increase proportionally (e.g., 0.01% or 100 ppm = 0.1 Da at m/z = 1000, 0.5 Da at m/z = 5000, or 5 Da at m/z = 50,000). It is important not to confuse mass accuracy with the term “accurate mass”; the latter is reserved for mass measurements accurate to ≤10 ppm.
Interplay Between Resolution and Molecular Weight
In the mass range up to ∼3000 Da it is recommended that, if possible, a resolution sufficient to partially resolve the isotope cluster
be used to facilitate mass assignment of the monoisotopic peak (Ingendoh et al., 1994). Resolution sufficient to separate single-massunit differences is also important in order to detect the presence of modifications such as deamidation of Asn to Asp and to make it possible to directly determine the charge state of an ESI ion by measuring the isotope peak spacing. For poorly resolved or unresolved isotope clusters, the mass accuracy can be dramatically affected by how the data is treated by the instrument’s data system. It is important to be aware of whether the data sytem is assigning masses to the centroid of the peak or to the peak top. In general, the centroid and the peak top mass will not be the same in this mass range because the isotopic distribution comprising the unresolved envelope of peaks is asymmetric in shape. Below mass ∼3000 it is sometimes difficult to accurately determine the centroid, and therefore the average mass, of an unresolved isotope cluster. The reason for this difficulty is that the bottom of the cluster, which must be included to get an accurate determination of the centroid for peaks in this mass range, is often distorted or poorly defined due to sample background (or, in MALDI, matrix) noise and/or poor ion statistics. These factors can easily contribute errors of ≥1 Da to the mass determination for compounds in the mass range of 1000 to 3000 Da. Furthermore, it should be noted that the peak top mass, which is the most easily determined value, corresponds to neither the monosiotopic mass or the average mass in this mass range. At ∼1200 and below, the mass at the peak top of the unresolved envelope will be closer to the monoisotopic mass, whereas at ∼3000 Da and above, the peak top will be closer to the average mass. For these reasons the use of resolved isotopes in this mass range is clearly preferable. Determining the monoisotopic mass of a protein is difficult or impossible due to the insignificant contribution that the monoisotopic peak makes to the isotopic envelope (Fig. 16.1.10, panel D). Furthermore, quadrupole and TOF mass analyzers cannot provide the resolving power necessary to separate the isotope peaks for molecules with masses in excess of ∼8000 Da. Fortunately, at masses above ∼10,000, the unresolved peak profiles are quite symmetrical and the mass at the peak top is nearly identical to the chemical average mass of the protein. For proteins, errors in measuring the centroid can be caused by the presence of unresolved adducts along with the protonated molecular ion (with, for example, Na+, K+, or
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photochemical adducts of MALDI matrices) or of other proteins of similar but not identical mass that are not resolved. The relative abundance of any of these adducts is unpredictable, and will shift the apparent centroid of the unresolved cluster to higher mass in an unpredictable fashion. In these cases, it will be impossible to obtain an accurate mass assignment unless the instrument resolving power can be increased or the contributing adduct ions can be removed by more stringent cleanup of the sample (Ingendoh et al., 1994; Zubarev et al., 1995). Thus it is still important even at higher mass to have a reasonable amount of resolving power. Resolution should be sufficient to enable detection of common protein variants that may be present in the sample. For example, β-casein exists as a family of isoforms differing by the substitution of one or more amino acids. The MALDI and ESI mass spectra of β-casein are compared in Figure 16.1.8. Detection of one of these variants, which differs by 40 Da in 24,000 Da, requires a resolution of ∼1200 FWHM (24,000/4010% valley def. × 2), which is easily achievable by ESI-MS on quadrupole mass analyzers. The resolution in MALDI-MS with a TOF analyzer is typically <1000 at this mass, and is insufficient to detect such small modifications. The minimum desirable resolution can be estimated from the natural width of the molecular ion envelope obtained from the computer programs mentioned above. For example, the envelope of isotopes for a 24-kDa protein has a width of ∼10 Da at the half-height of the peak (Fig. 16.1.9, panel D). This corresponds to an apparent resolution of 2400 (24,000/10). Having an instrumental resolution higher than 2400 but less than ∼20,000 (see Fig. 16.1.9) will not improve the apparent resolution because the minimum width of the unresolved protein molecular ion cluster is determined by the weighted average of the natural abundance of the isotopes present. It should be clearly stated, however, that having a higher resolution than is theoretically required is not detrimental (as long as it does not result in a decrease in sensitivity), and it may have some beneficial effect on the ability to precisely define the peak top and therefore the average mass.
OUTLOOK Overview of Peptide and Protein Analysis by Mass Spectrometry
Mass spectrometry and its application to biological problems are clearly enjoying a second childhood. MS-based methods for protein sequencing have sensitivity in the femtomole
range, exceeding that of Edman-based sequencing, and MS-based methods are uniquely capable of analyzing the ever-broadening spectrum of protein modifications. Mass spectrometry is even showing promise as a tool for studying protein folding (Englander, 1993; Smith and Zhang, 1994; Wagner and Anderegg, 1994) and noncovalent interaction (Smith and Light-Wahl, 1993; Cheng et al., 1995). Further increases in the understanding of the ionization/desorption processes in ESI and MALDI are likely to translate into even higher sensitivity through higher ion-sampling efficiency. It can also be expected that new methods for dissociating molecular ions of peptides and proteins (possibly involving surface-induced collisions or photoexcitation) will be developed that will be more efficient or more selective than collision-induced or post-souce decay. One can also anticipate that computer algorithms for the interpretation or “fingerprinting” of MS/MS data will continue to improve so that soon a minimum amount of human intervention will be required for identification (for an example, see Eng et al., 1994). Although MS-based approaches have sufficient raw sensitivity to analyze low femtomole amounts of peptides and proteins, purifying and handling samples at these levels remains a significant challenge for which there is no general strategy at present. With further improvements, gel- and membrane-based methods and microchromatography-based methods for immobilizing minute amounts of sample for biochemical manipulation and cleanup may meet this challenge. The mantra for MS now and into the next millenium is “smaller, cheaper, faster, simpler, more sensitive.” These criteria are based in large part on the need to make MS more accessible to the biological community. A variety of new types of mass spectrometers have recently been introduced that satisfy one or more of these criteria, and certain of these new instruments may provide unique benefits for molecular mass determination and sequencing of minute amounts of peptides and proteins. ESI and MALDI have shown particular promise when coupled to quadrupolar ion trap analyzers (Korner et al., 1996; Qin and Chait, 1995). These devices are capable of very high resolution and very rapid scanning, and can perform MSn experiments, in which fragment ions formed in the first MS/MS (MS2) experiment are further fragmented to provide additional structural information. TOF mass analyzers have been coupled to ESI sources (Verent-
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chikov et al., 1994; Mirgorodskaya, 1994) and offer the promise of improved resolution, very high sensitivity, and very fast sampling rates. An interesting variation on these two approaches couples an ion trap to a TOF analyzer, providing structural information as well as molecular mass data (Lee and Lubman, 1995). Additional variations on this theme of connecting two different, relatively inexpensive analyzers together to produce a more versatile instrument are under development. Although neither small or inexpensive, Fourier-transform ion-cyclotron resonance mass spectrometers equipped with MALDI and ESI are extremely versatile instruments with the potential for simultaneously providing very high mass accuracy, high resolution (routinely ≥20,000), and extremely high sensitivity for peptide and protein mass measurement (Winger et al., 1993; O’Conner et al., 1995; Wu et al., 1995; Smith et al., 1996). These devices are also capable of performing MSn experiments.
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Overview of Peptide and Protein Analysis by Mass Spectrometry
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Contributed by Steven A. Carr and Roland S. Annan SmithKline Beecham Pharmaceuticals King of Prussia, Pennsylvania
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Matrix-Assisted Laser Desorption/Ionization Time-of-Flight Mass Analysis of Peptides Matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS; Karas and Hillenkamp, 1988; Tanaka et al., 1988; Hillenkamp et al., 1991; Beavis, 1992; Bahr et al., 1994; Kaufmann, 1995; Stults, 1995) is one of the most useful techniques for determining the mass of biomolecules, with exceptional capabilities for mass analysis of peptides. Relative to other ionization techniques, MALDI provides high sensitivity and excellent tolerance of salt and other common buffer components. Routine detection limits for peptides are in the subpicomole range. The ions commonly observed are the protonated molecules (M+H+), which makes data analysis relatively easy. The use of MALDI-MS is becoming widespread, due in part to the availability of modern, commercially available instruments that are easier to operate and provide greater performance at lower cost than earlier versions. Peptide analysis is one of the most important applications of MALDI-MS. The ability to ionize many peptides in a mixture simultaneously makes MALDI-MS a popular choice for rapid analysis of protein digests. In fact, the high sensitivity and the high salt tolerance of MALDI have permitted the mass analysis of peptides directly from a single neuron without using purification (Li et al., 1994). MALDI-MS is frequently used for analysis of HPLC fractions from peptide maps. It has also been coupled with capillary electrophoresis (CE; see UNIT 10.9) for off-line analysis (Keough et al., 1992; vanVeelen et al., 1993; Walker et al., 1995). Many other specialized applications of MALDI-MS have been described in the literature, including protein identification by peptide mass fingerprinting (Henzel et al., 1993; Mann et al., 1993; Pappin et al., 1993; Mann, 1994; Patterson, 1995), C-terminal peptide sequencing (Patterson et al., 1995; Thiede et al., 1995; Woods et al., 1995), post-translational modification determinations (Huberty et al., 1993; Yip and Hutchens, 1993; Liao et al., 1994; Robertson et al., 1994; Stahl et al., 1994; Sutton et al., 1994; Talbo and Mann, 1994; Crimmins et al., 1995; Rüdiger et al., 1995), and protein epitope mapping (Papac et al., 1994; Zhao and Chait, 1994). Another popular application for MALDIMS of peptides is as a complement to Edman
UNIT 16.2
degradation (Elicone et al., 1994; Geromanos et al., 1994). A small aliquot (10 to 100 fmol) can be removed prior to Edman degradation to provide information on the number of cycles for which the sequencer should be programmed. The peptide mass can be used to confirm the sequence obtained by Edman degradation and may in some cases provide indirect information on a residue that was not observed in the sequence data. A difference in the mass calculated from the sequence and that measured by MALDI-MS may indicate the presence of a post-translational modification.
INSTRUMENT CONFIGURATIONS Most often the MALDI ion source is mated to a time-of-flight (TOF) mass analyzer from which the high mass range, resolution, and mass accuracy characteristics are derived (Cotter, 1992). The basic TOF mass spectrometer is a linear instrument, where the ions follow a straight, field-free path between the acceleration of the ion source and the detector. This configuration provides the highest sensitivity, yields few, if any, fragment ions, and requires the least manipulation of tuning parameters. The limitations of this instrument include relatively poor resolving power (m/∆m <2000) and poor mass accuracy (±0.15% with external calibration). Resolving power is the ability to distinguish peaks that are close in mass. For TOF instruments, it is defined by the mass of a peak divided by its width at half-height. Reflector instruments (or mode of operation) yield higher resolution (m/∆m ∼2000 to 5000) and better mass accuracy (±0.015% with external calibration). The reflectron is a focusing device that improves resolution by allowing ions of the same mass with different energies to arrive at the detector at the same time. The improved performance, however, is often obtained at the expense of slightly lower signal and the occasional appearance of undesired fragment ions. MALDI has been described as a “soft” ionization technique; however, significant fragmentation often occurs, but it is apparent only when operating in the reflector mode. Fragmentation is rarely observed in a linear TOF instrument because no electric field separates the fragments from their precursor ions. Although the fragment ions can be quite useful for sequence Mass Spectrometry
Contributed by William J. Henzel and John T. Stults Current Protocols in Protein Science (1996) 16.2.1-16.2.11 Copyright © 2000 by John Wiley & Sons, Inc.
16.2.1 Supplement 4
determination in post-source decay (PSD) spectra (Kaufmann et al., 1993, 1994; see UNIT 16.1), the fragment ions in the full-energy reflector spectrum do not appear at the proper masses and caution should be exercised so that they are not mistaken for molecular ions. Acquisition of an accompanying spectrum in the linear mode is the easiest method to ensure the proper identity of ions. Particularly when operating in reflector mode, it is necessary to determine whether the instrument reports average or monoisotopic peak values (see UNIT 16.1). Low-mass peptides (<1200 Da) display well-resolved isotope peaks that are assigned as monoisotopic peaks by most data systems. Higher-mass peaks (>2000 Da) in the same spectrum may show little or no isotope resolution and the data system will report the average mass value. Peaks that are intermediate in mass will have partially resolved isotope peaks, and the data system may report without an indication whether the mass is monoisotopic or average. A relatively new configuration, “delayed extraction,” delivers higher resolution by temporally uncoupling the ionization event from ion acceleration (Colby et al., 1994; Brown and Lennon, 1995; Vestal et al., 1995; Whittal and Li, 1995). It is useful in both linear (m/∆m = 2000 to 4000) and reflector modes (m/∆m = 3000 to 6000). When delayed extraction is used in the reflector mode, a smaller degree of fragmentation is observed; this allows operation at higher laser power and results in higher signal intensity of the molecular ion. The higher resolution and mass accuracy afforded by delayed extraction have helped to give TOF-MS more of the characteristics commonly expected of a “high performance” mass spectrometer.
SAMPLE PREPARATION GUIDELINES
MALDI-TOF Mass Analysis of Peptides
Sample preparation is straightforward and rapid (Lecchi and Caprioli, 1996). It is generally identical for all types of MALDI instruments. Peptide samples, either single components or mixtures, are cocrystallized with a large excess (∼104) of a UV-absorbing compound (the “matrix”). The peptide molecules are released into the gas phase and ionized by a pulse of UV radiation, typically from a nitrogen laser (λ = 337 nm). A wide range of peptide concentrations can be analyzed, from 10−4 M (100 pmol/µl) to 10−8 M (10 fmol/µl). Often simply adding the matrix to the sample on the sample stage will result in high-quality spectra, as described in UNIT 11.6. Samples may also be
prepared by mixing the sample solution and the matrix solution in a microcentrifuge tube. A total of 0.5 to 2 µl is applied to the target. A large number of different matrices have been tried for peptide analysis; however, only two matrices are in common use: α-cyano-4hydroxycinnamic acid (αCN; Beavis et al., 1992) and 2,5-dihydroxybenzoic acid (DHB; Strupat et al., 1992). A third matrix, less frequently used, is 3,5-dimethoxy-4-hydroxycinnamic acid (sinapinic acid, SA; Beavis and Chait, 1989), which has proven useful for highmass peptides (see Fig. 16.2.1 for chemistries and some guidelines for choosing a matrix). For a peptide with mass >1000 Da, αCN is preferred; DHB is a cleaner matrix for peptides of lower mass, and it is more tolerant of buffer salts at very low peptide concentrations. None of the matrices gives rise to significant photoadduct peaks with peptides, although photoadducts are often observed with proteins. The matrices are prepared as 10 mg/ml solutions in the solvents listed. Short-cuts, such as preparation of saturated solutions in these solvents, are occasionally used but yield less reproducible results. The matrix solutions are stable for weeks if they are kept in the dark and refrigerated when not in use. The matrices are normally used as obtained, without further purification. These compounds are photoreactive, and bad lots of material may be obtained on occasion. In addition, some commercial preparations contain salt contamination, particularly sodium, that will produce salt adducts and reduce the limits of detection. To avoid this, the matrices may be recrystallized and desalted by ionexchange chromatography (UNIT 8.2). Alternatively, highly purified matrix preparations are available from some instrument vendors. The detection limit for a peptide is dependent on the peptide’s ionization efficiency, its mass, its hydrophobicity, its solubility, the availability of a basic site for protonation, and other unknown factors. MALDI is relatively tolerant of salts and small amounts of some detergents when compared to most other forms of mass spectrometry ionization (Mock and Sutton, 1992; Rosinke et al., 1995; Cohen and Chait, 1996; Gharahdaghi et al., 1996). Table 16.2.1 lists salt and detergent concentrations that are compatible with MALDI. Some of these components interfere with ionization, and others can prevent crystal formation. As a general rule, any component present in a concentration higher than that of the matrix (∼50 mM) may be a problem. Detection limits are quite dependent on sample purity, especially at
16.2.2 Supplement 4
Current Protocols in Protein Science
Compound name
Suggested solvent Mass range (Da)
α-cyano-4-hydroxy-trans-cinnamic acid (αCN, CCA)
50% acetonitrile
400 -100,000
50% ethanol
200-5,000
COOH CN
HO
2,5-dihydroxybenzoic acid (DHB, gentisic acid)
COOH OH
HO 3,5-dimethoxy-4-hydroxy-trans-cinnamic-acid 50% acetonitrile (sinapinic acid, SA)
1,000 -100,000
COOH
MeO
HO MeO
Figure 16.2.1 Common matrix compounds for MALDI-MS.
Table 16.2.1
Tolerance for Common Buffer Components
Buffer componenta
Maximum concentrationb
Sodium chloride Phosphate Tris base Urea Guanidine Azide Glycerol PEG 2000 SDS Triton X-100, RTX-100, or NP-40 Tween CHAPS n-Octyl-β-glucopyranoside Zwittergent Lauryldimethylamine oxide (LDAO)
50 mM 10 mM 50 mM 1M 1M 0.1% (v/v) 1% (v/v) 0.1% (w/v) 0.01% (w/v) 0.1% (v/v) 0.1% (v/v) 0.01% (w/v) 1% (v/v) 0.1% (v/v) 1% (w/v)
aAbbreviations: CHAPS, 3-[(3-cholamidopropyl)-dimethylammonio]-1-propane
sulfonate; PEG, polyethylene glycol; SDS, sodium dodecyl sulfate; Tris, tris(hydroxymethyl)aminomethane. bThese values should be viewed as approximate. The maximum concentrations that can be tolerated depend on the matrix used, the method of sample preparation, the extent of rinsing, the amount of analyte, and the presence of mutiple buffer components.
Mass Spectrometry
16.2.3 Current Protocols in Protein Science
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MALDI-TOF Mass Analysis of Peptides
the subpicomole levels. The presence of buffer components and other peptides may lower the signal of the component of interest. In all cases, best results are obtained when salts and detergents are kept to as low a concentration as possible. A number of methods have been developed that enhance the tolerance for buffer components. The easiest method is simple dilution; due to the excellent sensitivity of MALDI-MS, samples may be diluted significantly in order to reduce the buffer concentration. If the buffer is volatile, it may be removed by drying with vacuum centrifugation. Unfortunately, this approach often leads to sample losses by adsorption to the tube walls. A strong solvent (e.g., 70% isopropanol/5% trifluoroacetic acid) may assist in recovery of the sample. Often salts will crystallize separately from the matrix and analyte, so a simple, quick rinse with cold dilute acid is often useful with αCN or SA (Beavis and Chait, 1990); DHB is too water-soluble to permit rinsing. Rinsing the sample with water not only may enhance the signal for some peptides but also, in some cases, may result in detection of ions that are completely suppressed by the salt. However, care should be exercised because some hydrophilic peptides may be lost by washing. Matrix additives such as 50 mM fucose (Billeci and Stults, 1993) or 10% 5-methoxysalicylic acid (Karas et al., 1993) have been developed; in some case these additives can enhance the quality of spectra when buffer salts are present. Adsorption to nylon (Zaluzec et al., 1994), polyethylene (Blackledge and Alexander, 1995), nitrocellulose (Mock et al., 1992; Preston et al., 1993; Bai et al., 1995), Nafion (Bai et al., 1994), or polyvinylidene difluoride (PVDF; Vestling and Fenselau, 1994) allows extensive rinsing before matrix is added. In addition, adsorption to a membrane provides a platform for chemical or enzymatic reactions (Fabris et al., 1995). An excellent alternative for desalting a sample utilizes a short fused-silica capillary that contains a small volume of reversed-phase HPLC packing material (Zhang et al., 1995). A sample is concentrated on this capillary column, the salts are rinsed away, and the analyte is eluted directly onto the MALDI sample stage with 1 µl of organic mobile phase. Another alternative method is attachment of affinity molecules (e.g., antibodies) to the sample stage; this permits selective concentration of targeted analytes from dilute solution and allows for subsequent washing before the matrix is added to disrupt the affinity binding (Nelson et al., 1995).
Using higher laser power can increase detection of larger peptide fragments as well as detection of some peptides at very low levels. Some ions may not appear until higher laser power is used. Excessive laser power, however, can result in a loss of resolution, and in reflector-TOF operation, increased fragment ion formation may result in a loss of signal for the molecular ion. In general the laser power should be set to the lowest level that will still produce a good-quality signal. Preparation of sample either in a microcentrifuge tube or on a sample stage with matrix is a rather straightforward task. Over recent years a number of alternative methods have been described. These include the thin-film method (Xiang and Beavis, 1994) and the rapid evaporation method (Vorm and Mann, 1994; Vorm et al., 1994). Advantages of these methods include easier rinsing of contaminants, lower detection limits, and/or more uniform crystal formation. A modification of the rapid evaporation method incorporates nitrocellulose for more reproducible results (Shevchenko et al., 1996). The exact method of sample preparation can have a significant effect on the results (Cohen and Chait, 1996), although no one method appears to be universally successful. These authors have tried these different techniques for many samples and find that similar results are often obtained. The lowest detection limits have been achieved when significantly smaller samples are used (<1 nl; Jespersen et al., 1994), but these require specialized sample-handling devices (picoliter vials).
INSTRUMENT CALIBRATION The mass axis of the time-of-flight mass spectrometer is normally calibrated with a twopoint calibration procedure. For peptides, these points are either a high- and a low-mass peptide or a single peptide plus a matrix ion for the low-mass point. The spectrum used for calibration should be acquired under the same conditions as the analyte spectra (same matrix, peptide concentration, and laser power). Table 16.2.2 is a list of common peptides used for calibration and their masses. Mass accuracy for external calibration is typically ±0.15% of the ion mass (e.g., ±1.5 Da at 1000 Da) for linear mode, and ±0.015% for reflector mode. Addition of mass standards to the analyte sample (internal calibration) can improve the accuracy to ±0.01%. The standards are chosen to bracket the analyte mass (above and below); they are added optimally at a concentration near that of the analyte. Peptides used for external calibra-
16.2.4 Supplement 4
Current Protocols in Protein Science
Table 16.2.2
Common Mass Calibration Compounds
Sample name
M+H+ (12C)
DHB matrix αCN matrix αCN matrix (dimer) desArg bradykinin Bradykinin Angiotensin I Substance P Glu-fibrinopeptide B Renin substrate ACTH-CLIP (18-39) Melittin Bovine insulin B-chain (oxidized) ACTH (7-38) Bovine insulin E. coli thioredoxin
155.0344 190.0504 379.0930 904.4681 1060.5692 1296.6853 1347.7360 1570.6774 1758.9331 2465.1989 2846.7459 3494.6513 3657.9294 5730.6087 —b
M+H+ (avg.)a 155.13 190.18 379.35 905.04 1061.23 1297.50 1348.65 1571.60 1760.04 2466.71 2848.49 3496.94 3660.17 5734.55 11676.44
aDue to variation in isotope abundance, the average mass is precise to only two decimal
places. bThe nonisotopic peak abundance is too small to be observed.
tion (see Table 16.2.2) can also be used as internal calibration standards. The authors find that addition of internal standards for analyte levels <100 fmol has limited utility because the standard can suppress ionization of the analyte, and mass accuracy is compromised by poor ion statistics at low levels. A matrix peak may be used for a one-point calibration adjustment (Vorm and Mann, 1994).
ANALYSIS OF PEPTIDE MIXTURES MALDI-MS is often the method of choice for rapid analysis of peptide mixtures. Ion suppression does occur in MALDI, but it is possible to analyze peptide mixtures containing dozens of components. Although the number of papers that describe applications of complex peptide mixtures is limited, spectra of unseparated tryptic digests have been published for human growth hormone and tissue plasminogen activator (Billeci and Stults, 1993), human interferon α and interleukin-4 (Tsarbopoulos et al., 1994), barley α-amylase (Andersen et al., 1994), and recombinase A (Lecchi and Caprioli, 1996). Complex peptide mixtures should, when possible, be analyzed using two different matrices. Each matrix may result in some differences in peak abundance (Lecchi and Caprioli, 1996). These differences may in some cases
result in the appearance of new peptide ions and the disappearance of other ions. Measurements made using more than one analyte concentration may also lead to greater numbers of peptides being observed. Finally, spectra should be obtained from a number of different spots or crystals on the target because the homogeneity of the analyte can vary from crystal to crystal. Detection of some ions may be limited by MALDI instruments with low resolution. For example, when a 100-kDa protein is digested with a protease such as trypsin, a large number of peptides with similar masses are typically generated. As the peptide masses approach one another (±1 to 2 Da), the ions may not be resolved. Instruments with higher resolution (m/∆m >3000) will be able to separate masses that are within 1 Da. Larger peptides (>3000 Da) can be more problematic to analyze due to low ionization efficiency and low solubility. Peptides produced by cyanogen bromide are often hydrophobic and can be refractory toward MALDI analysis. Methods have been developed that provide some success by extraction of cyanogen bromide–generated peptides from nitrocellulose (Bai et al., 1995), cationic (Zhang et al., 1994), and PVDF membranes (Henzel et al., 1994) after electroblotting from SDS-PAGE. Octyl-β-glucoside (Cohen and Chait, 1996; Gharahdaghi et al., 1996) has been
Mass Spectrometry
16.2.5 Current Protocols in Protein Science
Supplement 4
Table 16.2.3
Autolytic Fragments of Trypsin and Lysine-C
Trypsin (porcine) Peptide T2 T3 T7 T13 T10 T6 T9 T14 T8 T11 T4 T5 T12 T1
Avg.
massa
261.3 514.6 842.0 906.1 1006.1 1045.2 1469.7 1737.0 1769.0 2158.5 2211.4 2283.6 3014.3 4491.1
Lysine-C (Achromobacter) Peptide
Avg. massa
K2 K1 K5 K4 K3 K6
1922.1 3094.5 4875.3 5503.5 5917.5 6514.0
aMasses in bold print are the most frequently observed fragments. All masses are
isotopic averages.
MALDI-TOF Mass Analysis of Peptides
a useful additive to increase the solubility of hydrophobic peptides. Peptides can be generated in solution (UNIT 11.1), from in situ digestion on a PVDF or nitrocellulose membrane after electroblotting from a gel (UNIT 11.2), in an SDS-polyacrylamide gel (UNIT 11.3), in a silver-stained SDS-polyacrylamide gel (Shevchenko et al., 1996), or directly on the sample stage of a mass spectrometer (Fabris et al., 1995; Nelson et al., 1995). Most of these methods are directly compatible with MALDI analysis, with the exception of some of the in situ methods that utilize detergents. For example, hydrogenated Triton is a detergent that has been used for in-gel and on-membrane protein digestion. Although hydrogenated Triton is compatible with UV detection, its compatibility with MALDI-MS is more limited (Gharahdaghi et al., 1996). The highestquality spectra are obtained from peptides that are purified by HPLC using volatile buffers. Peptide mixtures that are derived from a proteolytic digestion should be analyzed along with a protease control. The protease control should consist of both the same amount of enzyme and the same conditions that were used to digest the sample. Analysis of the enzyme control allows identification of protease autolytic fragments that can interfere with sample analysis. These fragments may be difficult to identify without the use of a protease control because some enzymes are modified to increase
stability. Table 16.2.3 lists the commonly observed autolytic fragments for trypsin and lysine-C.
ANALYSIS OF SYNTHETIC PEPTIDES Mass spectrometry has taken a leading role in the verification of proper synthesis of peptides. In particular, it is sensitive to identification of deletions, improper composition, unremoved protecting groups, scavenger adducts, oxidation, deamidation, and other undesired synthetic products. Amino acid analysis, although the traditional method for verification, cannot identify many of these problems. For this reason, mass spectrometry has replaced amino acid analysis as the routine method for synthetic peptide analysis in many synthesis labs. Crude samples can be analyzed successfully by MALDI. Samples are typically prepared to yield a peptide concentration of 1 to 50 pmol/µl. If the quantity is not known, two or three 10-fold dilutions of the crude peptide sample will usually yield a suitable concentration. Care should be taken in interpreting the results when extremely acid-labile protecting groups are used because these may be removed by the acidic environment of the MALDI matrix. A number of studies of peptide synthesis have been carried out by the Association of Biomolecular Resource Facilities (ABRF); in these studies various ionization techniques
16.2.6 Supplement 4
Current Protocols in Protein Science
were evaluated for their utility in peptide analysis (Fields et al., 1994, 1995). In comparisons with electrospray ionization (ESI), MALDI gave nearly identical results for identification of improperly synthesized or improperly deprotected peptides. Approximate determination of the relative amount of properly synthesized material was also nearly the same for MALDI and ESI and correlated well with HPLC analysis. MALDI should not normally be relied upon for quantitation unless appropriate internal standards are used. Nonetheless, in the case of synthetic peptides, the approximate relative amounts of product/byproducts can often be determined successfully because the byproducts are very similar in sequence and properties to the product. When possible, the spectra should be obtained with sufficient resolution and mass accuracy to confirm that the correct peptides were prepared. In many cases, this requires resolving masses that differ by just 1 Da, with mass accuracy <0.5 Da, to eliminate the possibility of deamidation. When peptides are formed with disulfide bonds, the mass difference between the oxidized and reduced forms is only 2 Da. The mass measurement accuracy must distinguish between these two possibilities. Moreover, a mixture of reduced and oxidized forms often exists, and sufficient resolution is required to show the relative abundance of each form. On instruments in which the necessary accuracy or resolution is not possible to achieve, an alternative is to show reactivity of free sulfhydryls to alkylation, with the accompanying larger shifts in mass (Zaluzec et al., 1994; also see Analysis of Chemical Modifications).
ANALYSIS OF CHEMICAL MODIFICATIONS MALDI-MS can provide rapid identification of peptide modifications. One modification that is commonly observed with this technique is oxidation of tryptophan and methionine residues. Oxidation of a tryptophan results in an increase of 32 Da and oxidation of methionine results in an increase of 16 Da (methionine sulfoxide); oxidation of methionine to methionine sulfone (+32 Da) is generally not observed. The presence of a +16- or +32-Da adduct can be used as an indirect indication of the presence of an oxidized methionine or tryptophan residue. Oxidation can occur during peptide purification, storage, or mass analysis. Thorough degassing of the HPLC mobile
phases and use of nonmetal frits can minimize oxidation during purification. Storage at −5° or −80°C can minimize oxidation during sample storage. Because oxidation can also result when a peptide is stored with matrix on a sample stage, best results are obtained when the sample is analyzed immediately after the matrix has been added. Some peptides may be more susceptible to such oxidation than others. A second type of modification that may occur during MALDI analysis is disulfide bond cleavage (Zhou et al., 1993; Patterson and Katta, 1994; Crimmins et al., 1995). The extent of disulfide cleavage may be greater at higher laser power. A protocol has been developed for detection of disulfide bonds using on-the-probe reduction (Fischer et al., 1993). After a spectrum is acquired, reducing agent is added to the sample stage, and the spectrum is acquired again. The peak for each disulfide-linked peptide then disappears, and new, smaller peaks appear whose masses when summed equal the mass of the oxidized peptide. The reducing agent should be carefully titrated because a large excess can cause severe ion suppression. Another commonly observed modification is the formylation of amino groups (+28 Da) when formic acid is used as a solvent. Formic acid is a common solvent for cleavage at methionine residues using cyanogen bromide; however, other solvents—e.g., 0.1 N HCl or 70% trifluoroacetic acid—can be substituted. Amino-terminal glutamine is known to cyclize to pyroglutamic acid under certain aqueous conditions, and this reaction may also take place in the acidic environment of the matrix. Peptides containing cysteine that are derived from a protein separated by SDS-PAGE can be modified by free acrylic acid present in the gel. This adduct will add 71 Da to peptide mass. Proteins that are purified or refolded in the presence of urea may become carbamylated. Peptides can be carbamylated at the amino terminal and at lysine residues, resulting in a +43-Da increase for each carbamyl group added. Some of the more commonly observed chemical and post-translational modifications are listed in Table A.1A.3 (APPENDIX 1A). A more extensive listing of post-translational modifications and their corresponding mass shifts is available on the World Wide Web (see Table 16.2.4 for useful mass spectrometry sites). Future supplements will present detailed methods for MALDI-MS analysis of phosphorylation and glycosylation, two frequently observed post-translational modifications.
Mass Spectrometry
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Table 16.2.4
Useful Mass Spectrometry Web Sites
Site
Features
WWW URL
Delta Mass
Extensive list of mass shifts due to amino acid modifications
http://www.medstv.unimelb.edu.au/WWWDOCS/ SVIMRDocs/MassSpec/deltamassV2.html
EMBL Protein Core Facility home page
Extensive information on database searching
http://www.mann.embl-heidelberg.de
Rockefeller/NYU mass spectrometry home page
Many features including a library of matrices
http://chait-sgi.rockefeller.edu
UCSF mass—spectrometry home page
On-line database searching
http://rafael.ucsf.edu
Murray’s mass spectrometry home page
Best set of hyperlinks to other mass spectrometry web sites
http://tswww.cc.emory.edu/~kmurray/mslist.html
American Society for Mass Spectrometry home page
Information on ASMS meetings and short courses
http://www.trail.com/asms
Univ. of Washington (Ken Walsh’s laboratory) home page
Programs for peptide digest interpretation
http://128.95.12.16/WalshLab.html
Univ. of Washington (John Yates’ laboratory) home page
Information on database searching with fragmentation spectra
http://thompson.mbt.washington.edu
Pedro’s BioMolecular Research Tools
A large collection of useful hyperlinks to mass spectrometry, protein chemistry, and molecular biology sites
http://www.public.iastate.edu/~pedro/ research_tools.html
SUMMARY
MALDI-TOF Mass Analysis of Peptides
Peptides as a class of compounds are extremely amenable to analysis by MALDI-MS. The high sensitivity, tolerance of many buffer components, rapidity of measurement, and ease of use of the technique have made it an indispensible tool for routine analysis of peptides from a wide variety of natural and synthetic sources. In particular, the direct measurement of molecular mass gives insight into modifications that are frequently not observed by most other techniques. Advances in instrumentation for time-of-flight analyzers are giving MALDITOF instruments the performance characteristics (resolution, mass accuracy, and fragmentation measurement) found previously only in other types of mass spectrometers. The other major mass spectrometry technique for peptide analysis, electrospray ionization (see UNIT 16.1), has many characteristics similar to MALDI-MS. Electrospray offers the advantage of on-line coupling with chromatographic and electrophoretic techniques, but MALDI-MS displays a higher tolerance for buffer components, often results in higher sensitivity, and is operationally a bit easier to use.
Nonetheless, both techniques are equally applicable to a large variety of problems and they complement each other in many ways. The choice of technique frequently depends on the particular problem at hand.
LITERATURE CITED Andersen, J.S., Sogaard, M., Svensson, B., and Roepstorff, P. 1994. Localization of an O-glycosylated site in the recombinant barley α-amylase 1 produced in yeast and correction of the amino acid sequence using matrix-assisted laser desorption/ionization mass spectrometry of peptide mixtures. Biol. Mass Spectrom. 23:547-554. Bahr, U., Karas, M., and Hillenkamp, F. 1994. Analysis of biopolymers by matrix-assisted laser desorption/ionization (MALDI) mass spectrometry. Frensenius J. Anal. Chem. 348:783791. Bai, J., Liu, Y.-H., Cain, T.C., and Lubman, D.M. 1994. Matrix-assisted laser desorption/ionization using an active perfluorosulfonated ionomer film substrate. Anal. Chem. 66:3423-3430. Bai, J., Qian, M.G., Liu, Y., Liang, X., and Lubman, D.M. 1995. Peptide mapping by CNBr degradation on a nitrocellulose membrane with analysis by matrix-assisted laser desorption/ionization mass spectrometry. Anal. Chem. 67:1705-1710.
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Beavis, R.C. 1992. Matrix-assisted ultraviolet laser desorption: Evolution and principles. Org. Mass Spectrom. 27:653-659.
sis. In Techniques in Protein Chemistry V (J.W. Crabb, ed.) pp. 501-507. Academic Press, San Diego.
Beavis, R.C. and Chait, B.T. 1989. Cinnamic acid derivatives as matrices for ultraviolet laser desorption mass spectrometry of proteins. Rapid Commun. Mass Spectrom. 3:432-435.
Fields, G.B., Angeletti, R.H., Bonewald, L.F., Moore, W.T., Smith, A.J., Stults, J.T., and Williams, L.C. 1995. Correlation of cleavage techniques with side-reactions following solid-phase peptide synthesis. In Techniques in Protein Chemistry VI (J.W. Crabb, ed.) pp. 539-546. Academic Press, San Diego.
Beavis, R.C. and Chait, B.T. 1990. Rapid, sensitive analysis of protein mixtures by mass spectrometry. Proc. Natl. Acad. Sci. U.S.A. 87:6873-6877. Beavis, R.C., Chaudhary, T., and Chait, B.T. 1992. α-Cyana-4-hydroxy cinnamic acid as a matrix for matrix-assisted laser desorption mass spectrometry. Org. Mass Spectrom. 27:156-158. Billeci, T.M. and Stults, J.T. 1993. Tryptic mapping of recombinant proteins by matrix-assisted laser desorption/ionization mass spectrometry. Anal. Chem. 65:1709-1716. Blackledge, J.A. and Alexander, A.J. 1995. Polyethylene membrane as a sample support for direct matrix-assisted laser desorption/ionization mass spectrometric analysis of high mass proteins. Anal. Chem. 67:843-848. Brown, R.S. and Lennon, J.J. 1995. Mass resolution improvement by incorporation of pulsed ion extraction in a matrix-assisted laser desorption/ionization linear time-of-flight mass spectrometer. Anal. Chem. 67:1998-2003. Cohen, S.L. and Chait, B.T. 1996. Influence of matrix solution conditions on the MALDI-MS analysis of peptides and proteins. Anal. Chem. 68:31-37. Colby, S.M., King, T.B., and Reilly, J.P. 1994. Improving the resolution of matrix-assisted laser desorption/ionization time-of-flight mass spectrometry by exploiting the correlation between ion position and velocity. Rapid Commun. Mass Spectrom. 8:865-868. Cotter, R.J. 1992. Time-of-flight mass spectrometry for structural analysis of biological molecules. Anal. Chem. 64:1027A-1039A. Crimmins, D.L., Saylor, M., Rush, J., and Thoma, R.S. 1995. Facile, in situ matrix-assisted laser desorption ionization–mass spectrometry analysis and assignment of disulfide pairings in heteropeptide molecules. Anal. Biochem. 226:355361. Elicone, C., Lui, M., Geromanos, S., ErdjumentBromage, H., and Tempst, P. 1994. Microbore reversed-phase high-performance liquid chromatographic purification of peptides for combined chemical sequencing-laser-desorption mass spectrometric analysis. J. Chromatogr. 676:121-137. Fabris, D., Vestling, M.M., Cordero, M.M., Doroshenko, V.M., Cotter, R.J., and Fenselau, C. 1995. Sequencing electroblotted proteins by tandem mass spectrometry. Rapid Commun. Mass Spectrom. 9:1051-1055. Fields, G.B., Angeletti, R.H., Carr, S.A., Smith, A.J., Stults, J.T., Williams, L.C., and Young, J.D. 1994. Variable success of peptide-resin cleavage and deprotection following solid-phase synthe-
Fischer, W.H., Rivier, J.E., and Craig, A.G. 1993. In situ reduction suitable for matrix-assisted laser desorption/ionization and liquid secondary ionization using tris(2-carboxyethyl)phosphine. Rapid Commun. Mass Spectrom. 7:225-228. Geromanos, S., Casteels, P., Elicone, C., Powell, M., and Tempst, P. 1994. Combined Edman-chemical and laser-desorption mass spectrometric approaches to micropeptide sequencing: Optimization and applications. In Techniques in Protein Chemistry V (J.W. Crabb, ed.) pp. 143-150. Academic Press, San Diego. Gharahdaghi, F., Kirchner, M., Fernandez, J., and Mische, S.M. 1996. Peptide-mass profiles of polyvinylidene difluoride–bound proteins by matrix-assisted laser desorption/ionization timeof-flight mass spectrometry in the presence of nonionic detergents. Anal. Biochem. 233:94-99. Henzel, W.J., Billeci, T.M., Stults, J.T., Wong, S.C., Grimley, C., and Watanabe, C. 1993. Identifying proteins from two-dimensional gels by molecular mass searching of peptide fragments in protein sequence databases. Proc. Natl. Acad. Sci. U.S.A. 90:5011-5015. Henzel, W.J., Grimley, C., Bourell, J.H., Billeci, T.M., Wong, S.C., and Stults, J.T. 1994. Analysis of two-dimensional gel proteins by mass spectrometry and microsequencing. Methods Compan. 6:239-247. Hillenkamp, F., Karas, M., Beavis, R.C., and Chait, B.T. 1991. Matrix-assisted laser desorption/ionization mass spectrometry of biopolymers. Anal. Chem. 63:1193A-1203A. Huberty, M.C., Vath, J.E., Yu, W., and Martin, S.A. 1993. Site-specific carbohydrate identification in recombinant proteins using MALDI-TOF MS. Anal. Chem. 65:2791-2800. Jespersen, S., Niessen, W.M.A., Tjaden, U.R., van der Greef, J., Litborn, E., Lindberg, U., and Roeraade, J. 1994. Attomole detection of proteins by matrix-assisted laser desorption/ionization mass spectrometry with the use of picolitre vials. Rapid Commun. Mass Spectrom. 8:581584. Karas, M. and Hillenkamp, F. 1988. Laser desorption ionization of proteins with molecular masses exceeding 10,000 daltons. Anal. Chem. 60:2299-2301. Karas, M., Ehring, H., Nordoff, E., Stahl, B., Strupat, K., Hillenkamp, F., Grehl, M., and Krebs, B. 1993. Matrix-assisted laser desorption/ionization mass spectrometry with additives to 2,5-dihydrobenzoic acid. Org. Mass Spectrom. 28:1476-1481.
Mass Spectrometry
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Kaufmann, R. 1995. Matrix-assisted laser desorption ionization (MALDI) mass spectrometry: A novel analytical tool in molecular biology and biotechnology. J. Biotechnol. 41:155-175. Kaufmann, R., Spengler, B., and Luetzenkirchen, F. 1993. Mass spectrometric sequencing of linear peptides by product-ion analysis in a reflectron time-of-flight mass spectrometer using matrixassisted laser desorption ionization. Rapid Commun. Mass Spectrom. 7:902-910. Kaufmann, R., Kirsch, D., and Spengler, B. 1994. Sequencing of peptides in a time-of-flight mass spectrometer: Evaluation of postsource decay following matrix-assisted laser desorption ionization (MALDI). Int. J. Mass Spectrom. Ion Processes 131:355-385. Keough, T., Takigiku, R., Lacey, M.P., and Purdon, M. 1992. Matrix-assisted laser desorption mass spectrometry of protein isolated by capillary zone electrophoresis. Anal. Chem. 64:15941600. Lecchi, P. and Caprioli, R.M. 1996. Matrix-assisted laser desorption mass spectrometry for peptide mapping. In New Methods in Peptide Mapping for the Characterization of Proteins (W.S. Hancock, ed.) pp. 219-240. CRC Press, Boca Raton, Fla. Li, K.W., Hoek, R.M., Smith, F., Jimenez, C.R., van der Schors, R.C., Veelen, P.A.V., Chen, S., van der Greef, J., Parish, D.C., Benjamin, P.R., and Geraerts, W.P.M. 1994. Direct peptide profiling by mass spectrometry of single identified neurons reveals complex neuropeptide-processing pattern. J. Biol. Chem. 269:30288-30292. Liao, P.-C., Leykam, J., Andrews, P.C., Gage, D.A., and Allison, J. 1994. An approach to locate phosphorylation sites in a phosphoprotein: Mass mapping by combining specific enzymatic degradation with matrix-assisted laser desorption/ionization mass spectrometry. Anal. Biochem. 219:9-20. Mann, M. 1994. Sequence database searching by mass spectrometric data. In Microcharacterization of Proteins (R. Kellner, F. Lottspeich, and H.E. Meyer, eds.) pp. 223-245. VCH Publishers, New York. Mann, M., Hojrup, P., and Roepstorff, P. 1993. Use of mass spectrometric molecular weight information to identify proteins in sequence databases. Biol. Mass Spectrom. 22:338-345. Mock, K.K., Sutton, C.W., and Cottrell, J.S. 1992. Sample immobilization protocols for matrix-assisted laser desorption mass spectrometry. Rapid Commun. Mass Spectrom. 6:233-238. Nelson, R.W., Dogruel, D., Krone, F.R., and Williams, P. 1995. Peptide characterization using bioreactive mass spectrometer probe tips. Rapid Commun. Mass Spectrom. 9:1380-1385.
MALDI-TOF Mass Analysis of Peptides
Papac, D.I., Hoyes, J., and Tomer, K.B. 1994. Epitope mapping of the gastrin-releasing peptide/anti-bombesin monoclonal antibody complex by proteolysis followed by matrix-assisted laser desorption ionization mass spectrometry. Protein Sci. 3:1485-1492.
Pappin, D.J.C., Hojrup, P., and Bleasby, A.J. 1993. Rapid identification of proteins by peptide-mass fingerprinting. Curr. Biol. 3:327-332. Patterson, D.H., Tarr, G.E., Regnier, F.E., and Martin, S.A. 1995. C-terminal ladder sequencing via matrix-assisted laser desorption mass spectrometry coupled with carboxypeptidase Y timedependent and concentration-dependent digestions. Anal. Chem. 67:3971-3978. Patterson, S.D. 1995. Matrix-assisted laser-desorption/ionization mass spectrometric approaches for the identification of gel-separated proteins in the 5-50 pmol range. Electrophoresis 16:11041114. Patterson, S.D. and Katta, V. 1994. Prompt fragmentation of disulfide-linked peptides during matrixassisted laser desorption ionization mass spectrometry. Anal. Chem. 66:3727-3732. Preston, L.M., Murray, K.K., and Russell, D.H. 1993. Reproducibility and quantitation of matrix-assisted laser desorption ionization mass spectrometry: Effects of nitrocellulose on peptide ion yields. Biol. Mass Spectrom. 22:544550. Robertson, J.G., Adams, G.W., Medzihradszky, K.F., Burlingame, A.L., and Villafranca, J.J. 1994. Complete assignment of disulfide bonds in bovine dopamine β-hyroxylase. Biochemistry 33:11563-11575. Rosinke, B., Strupat, K., Hillenkamp, F., Rosenbusch, J., Dencher, N., Krüger, U., and Galla, H.J. 1995. Matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS) of membrane proteins and non-covalent complexes. J. Mass Spectrom. 30:1462-1468. Rüdiger, A., Rüdiger, M., Weber, K., and Schomburg, D. 1995. Characterization of post-translational modifications of brain tubulin by matrixassisted laser desorption/ionization mass spectrometry: Direct one-step analysis of a limited subtilisin digest. Anal. Biochem. 224:532-537. Shevchenko, A., Wilm, M., and Mann, M. 1996. Mass spectrometric sequencing of proteins from silver-stained polyacrylamide gels. Anal. Chem. 68:850-858. Stahl, B., Klabunde, T., Witzel, H., Krebs, B., Steup, M., Karas, M., and Hillenkamp, F. 1994. The oligosaccharides of the Fe(III)-Zn(II) purple acid phosphatase of the red kidney bean. Eur. J. Biochem. 220:321-330. Strupat, K., Karas, M., and Hillenkamp, F. 1992. 2,5-Dihydroxybenzoic acid: A new matrix for laser desorption–ionization mass spectrometry. Int. J. Mass Spectrom. Ion Processes. 111:89102. Stults, J.T. 1995. Matrix-assisted laser desorption ionization mass spectrometry (MALDI-MS). Curr. Opin. Struct. Biol. 5:691-698. Sutton, C.W., O’Neill, J.A., and Cottrell, J.S. 1994. Site-specific characterization of glycoprotein carbohydrates by exoglycosidase digestion and laser desorption mass spectrometry. Anal. Biochem. 218:34-46.
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Talbo, G. and Mann, M. 1994. Distinction between phosphorylated and sulfated peptides by matrixassisted laser desorption/ionization reflector mass spectrometry at the subpicomole level. In Techniques in Protein Chemistry V (J.W. Crabb, ed.) pp. 105-113. Academic Press, San Diego. Tanaka, K., Waki, H., Ido, Y., Akita, S., Yoshida, Y., and Yoshida, T. 1988. Protein and polymer analyses up to m/z 100,000 by laser ionization timeof-flight mass spectrometry. Rapid Commun. Mass Spectrom. 2:151-153. Thiede, B., Wittmann-Liebold, B., Bienert, M., and Krause, E. 1995. MALDI-MS for C-terminal seuqence determination of peptides and proteins degraded by carboxypeptidase Y and P. FEBS Lett. 357:65-69. Tsarbopoulos, A., Karas, M., Strupat, K., Pramanik, B.N., Nagabhushan, T.L., and Hillenkamp, F. 1994. Comparative mapping of recombinant proteins and glycoproteins by plasma desorption and matrix-assisted laser desorption/ionization mass spectrometry. Anal. Chem. 66:2062-2070. vanVeelen, P.A., Tjaden, U.R., van der Greef, J., Ingendoh, A., and Hillenkamp, F. 1993. Off-line coupling of capillary electrophoresis with matrix-assisted laser desorption mass spectrometry. J. Chromatogr. 647:367-374. Vestal, M.L., Juhasz, P., and Martin, S.A. 1995. Delayed extraction matrix-assisted laser desorption time-of-flight mass spectrometry. Rapid Commun. Mass Spectrom. 9:1044-1050. Vestling, M.M. and Fenselau, C. 1994. Poly(vinylidene difluoride) membranes as the interface between laser desorption mass spectrometry, gel electrophoresis, and in situ proteolysis. Anal. Chem. 66:471-477. Vorm, O. and Mann, M. 1994. Improved mass accuracy in matrix-assisted laser desorption/ionization time-of-flight mass spectrometry of peptides. J. Am. Soc. Mass Spectrom. 5:955-958. Vorm, O., Roepstorff, P., and Mann, M. 1994. Improved resolution and very high sensitivity in MALDI TOF of matrix surfaces made by fast evaporation. Anal. Chem. 66:3281-3287. Walker, K.L., Chiu, R.W., Monnig, C.A., and Wilkins, C.L. 1995. Off-line coupling of capillary electrophoresis and matrix-assisted laser desorption ionization time-of-flight mass spectrometry. Anal. Chem. 67:4197-4204. Whittal, R.M. and Li, L. 1995. High-resolution matrix-assisted laser desorption/ionization in a linear time-of-flight mass spectrometer. Anal. Chem. 67:1950-1954.
1995. Simplified high-sensitivity sequencing of a major histocompatibility complex class I–associated immunoreactive peptide using matrixassisted laser desorption/ionization mass spectrometry. Anal. Biochem. 226:15-25. Xiang, F. and Beavis, R.C. 1994. A method to increase contaminant tolerance in protein matrixassisted laser desorption/ionization by the fabrication of thin protein-doped polycrystalline films. Rapid Commun. Mass Spectrom. 8:199204. Yip, T. and Hutchens, T.W. 1993. Protein phosphorylation: Sequence-specific identification of in vivo phosphorylation sites by MALDI-TOF mass spectrometry. In Techniques in Protein Chemistry IV (R.H. Angeletti, ed.) pp. 201-210. Academic Press, San Diego. Zaluzec, E.J., Gage, D.A., and Watson, J.T. 1994. Quantitative assessment of cysteine and cystine in peptides and proteins following organomercurial derivatization and analysis by matrix-assisted laser desorption ionization mass spectrometry. J. Am. Soc. Mass Spectrom. 5:359-366. Zaluzec, E.J., Gage, D.A., Allison, J., and Watson, J.T. 1994. Direct matrix-assisted laser desorption ionization mass spectrometric analysis of proteins immobilized on nylon-based membranes. J. Am. Soc. Mass Spectrom. 5:230-237. Zhang, H.Y., Andrén, P.E., and Caprioli, R.M. 1995. Micro-preparation procedure for high-sensitivity matrix-assisted laser desorption ionization mass spectrometry. J. Mass Spectrom. 30:17681771. Zhang, W., Czernik, A.J., Yungwirth, T., Aebersold, R., and Chait, B.T. 1994. Matrix-assisted laser desorption mass spectrometric peptide mapping of proteins separated by two-dimensional gel electrophoresis: Determination of phosphorylation in synapsin I. Protein Sci. 3:677-686. Zhao, Y. and Chait, B.T. 1994. Protein epitope mapping by mass spectrometry. Anal. Chem. 66:3723-3726. Zhou, J., Ens, W., Poppe-Schriemer, N., Standing, K.G., and Westmore, J.B. 1993. Cleavage of interchain disulfide bonds following matrix-assisted laser desorption. Int. J. Mass Spectrom. Ion Processes 126:115-122.
Contributed by William J. Henzel and John T. Stults Genentech, Inc. South San Francisco, California
Woods, A.S., Huang, A.Y.C., Cotter, R.J., Pasternack, G.R., Pardoll, D.M., and Jaffee, E.M.
Mass Spectrometry
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Sample Preparation for MALDI Mass Analysis of Peptides and Proteins
UNIT 16.3
The discovery that polar biological substances may be ejected from solid and liquid matrices as protonated or deprotonated ions by deposition of laser pulses initiated the development of methodology and instrumentation for characterization of biomacromolecules at unprecedented sensitivities. This is generally known by the acronym, MALDI-MS, for matrix-assisted laser desorption/ionization mass spectrometry (also see UNITS 16.1 & 16.2). The physicochemical properties of matrices that work well analytically are diverse; some matrices are suited to UV/visual laser wavelengths and others to the infrared. Although originally thought to be “soft” in terms of minimal deposition of internal energy (in the case of UV/visual laser frequencies), most modulate the vibronic activation of the analyte, giving rise to a fraction of metastable ions that are formed by unimolecular dissociation of the [M + H]1+ or [M − H]1−. The recorded mass spectra of these metastable ions, known as post-source decay (PSD) mass spectra, are similar to those of low-energy collision-induced dissociation (CID) fragment ions. The preponderance of ions formed are singly charged. Thus, they are well suited to high-energy collisional activation using appropriate tandem experiments to give high-energy CID-like mass spectra. Sample preparation may be the most crucial step in mass spectrometric analysis of peptides and proteins by MALDI (also see sample preparation guidelines in UNIT 16.2). The free dipolar ionic peptides or proteins must be incorporated into a crystalline solid solution that is comprised of excess solvent (the matrix), which modulates desorption/ionization by preferentially absorbing the laser energy at the irradiation wavelength and laser pulse intensity being employed. Usually the cocrystalline sample and matrix are prepared by mixing solutions of each component and permitting the mixture to crystallize through evaporation of the solvents. Two methods employing this approach are presented in this unit: the dried drop method (see Basic Protocol 1) and the rapid evaporation method (see Basic Protocol 2); also see Background Information and Table 16.3.1). This process can be disturbed by the presence of contaminants such as detergents and salts (see Table 16.2.1). Matrices that are sufficiently water insoluble will permit a degree of on-target washing without dissolving and liberating the analyte. Such matrices are useful for removing some contaminants, and include α-cyano-4-hydroxy-trans-cinnamic acid (HCCA) and sinapinic acid. Alternatively, the water-soluble 2,5-dihydroxybenzoic acid (DHB) may be used, since this matrix excludes some impurities during crystallization. DRIED DROPLET METHOD Materials 100% methanol Peptide or protein sample of interest: ∼1 pmol/µl in 0.1% (v/v) trifluoroacetic acid (TFA) Peptide or protein matrix solution (see recipes) Peptide or protein standard mixture (see recipes) Matrix-assisted laser desorption/ionization (MALDI) mass spectrometer
BASIC PROTOCOL 1
1. Clean the sample target with the appropriate solvents. Wipe with a Kimwipe wetted with water followed by one wetted with 100% methanol, and repeat for several cycles. Mass Spectrometry Contributed by C.R. Jiménez, L. Huang, Y. Qiu, and A.L. Burlingame Current Protocols in Protein Science (1998) 16.3.1-16.3.6 Copyright © 1998 by John Wiley & Sons, Inc.
16.3.1 Supplement 14
This cleaning procedure is usually sufficient. Alternatively, wipe with 0.1% (v/v) TFA and/or sonicate the target in 100% methanol or 0.1% TFA. Detergents are not recommended as they are hard to wash away and tend to give background peaks.
2. Pipet 0.5 to 1 µl peptide or protein sample onto the target, add 0.5 to 1 µl peptide or protein matrix solution, and mix by pipetting up and down. Allow to air dry. Samples may also be prepared by mixing the sample and matrix solution in a microcentrifuge tube, from which 0.5 to 1 ìl is applied to the target.
3. For external calibration, apply 0.5 to 1.0 µl of an appropriate peptide or protein standard mixture and 0.5 to 1.0 µl peptide or protein matrix solution (as in step 2) to the target in the vicinity of the sample. For highest mass accuracy, the standard solution may be mixed with an aliquot of the sample for internal calibration. However, do not add an excess of internal standard, as this may result in ion suppression of the sample. The optimal ratio of sample/standard may have to be empirically determined, although starting with equimolar amounts is recommended. In the case of peptide analysis, internal calibration may be performed on matrix ions.
4. Insert sample target into the MALDI mass spectrometer and mass analyze as recommended by the manufacturer. The sample target with applied matrix/sample preparations may be stored in the dark at ambient temperature for days without significant loss of signal. For longer storage, the target should be kept at −20°C in a sealed container. Irradiate the sample just above the threshold for appearance of the ions. Generally, the ion signals (mass-to-charge ratio, m/z) of peptides correspond to [M + H]1+. Larger peptides and proteins yield doubly protonated ions [M + 2H]2+ that appear at half the peptide or protein mass since m/z is being measured. Due to microheterogeneity of the matrix crystals, spectra should be accumulated from several different crystals of the matrix/sample preparation to obtain a representative sum spectrum. Under certain circumstances, the relative peak abundances may give a semiquantitative indication of the relative concentrations of peptides in a mixture. BASIC PROTOCOL 2
RAPID EVAPORATION METHOD Materials 100% methanol Peptide matrix/nitrocellulose solution (see recipe) Peptide sample of interest: ∼1 pmol/µl in 0.1% (v/v) trifluoroacetic acid (TFA) 10% (v/v) formic acid, 0° to 4°C Peptide standard mixture (see recipe) Matrix-assisted laser desorption/ionization (MALDI) mass spectrometer 1. Clean the sample target with the appropriate solvents (see Basic Protocol 1, step 1). 2. Pipet ∼0.5 µl peptide matrix/nitrocellulose solution on the sample target. Pipet quickly because of the high volatility of the solvent. The matrix/nitrocellulose solution will spread out rapidly, allowing fast evaporation of the solvent and the formation of a uniform white layer.
3. Apply ∼0.5 µl peptide sample onto the matrix surface on the sample target. Allow the solvent to evaporate at ambient temperature. Sample Preparation for MALDI Mass Analysis
The sample solution should not completely dissolve the matrix layer. Therefore, the concentration of organic solvent in the sample should be kept <30% and the pH should be acidic. For samples with higher organic solvent concentrations or high pH, apply 0.5 ìl
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water or 0.5 ìl of 10% (v/v) formic acid, respectively, onto the matrix surface and then apply the sample.
4. Rinse the sample with ∼10 µl cold 10% formic acid. Leave the solution on the sample for 10 sec and then remove using pressurized air, vacuum, or careful pipetting. Repeat the rinse step with water. This step is optional and can be omitted for samples that are free of salts (e.g., reversedphase HPLC fractions). These rinse steps are strongly recommended for unseparated peptide digest mixtures.
5. Apply peptide standard mixture (for details on internal versus external standards, see Basic Protocol 1, step 3). 6. Insert sample target into the mass analyzer and analyze (see Basic Protocol 1, step 4). For best ion intensities, matrix/sample preparations made using the rapid evaporation method should be analyzed immediately.
PURIFICATION/RECRYSTALLIZATION OF HCCA α-Cyano-4-hydroxy-trans-cinnamic acid (HCCA) matrix that has been purified often yields better sensitivities than matrix used as supplied by the manufacturer. The HCCA cleanup protocol described below was developed by B.O. Keller (Department of Chemistry, University of Alberta).
SUPPORT PROTOCOL
Materials α-Cyano-4-hydroxy-trans-cinnamic acid (HCCA; Sigma or Aldrich) 95% (v/v) ethanol: 50°C, room temperature, and 4°C Class C and E Whatman glass filters 50°C water bath 1. Suspend 7 to 8 g HCCA in 100 ml of 95% ethanol at room temperature. Dissolve all brownish crumbs, so that only bright yellow crystals remain undissolved. 2. Filter suspension through a class C Whatman glass filter under a vacuum. Wash the remaining crystals with 50 ml of 95% ethanol. The yield of undissolved HCCA is ∼50%.
3. Dry the crystals and then pick out and discard any remaining black or brown crumbs in the dried residue. 4. Warm 50 ml of 95% ethanol in an Erlenmeyer flask or beaker in a 50°C water bath. To obtain a saturated solution, add dried HCCA crystals to the ethanol while swirling the flask, and continue until no more HCCA dissolves and yellow crystals remain undissolved at the bottom. The temperature should not exceed 50°C, as this may cause the HCCA to decompose.
5. Filter the hot (50°C) HCCA solution using a class C Whatman glass filter. Use a very clean glass flask to collect the filtrate. Cool to room temperature, cover flask, and leave at −20°C overnight. The next day, a precipitate of tiny crystals should be observed. If no precipitate is observed, the HCCA solution may be too dilute. Add more HCCA from step 3 to the solution in a 50°C water bath and filter again (step 4).
6. Separate the precipitate from the HCCA solution by filtration using a class E Whatman glass filter. Wash the precipitate moderately with cold (∼4°C) 95% ethanol.
Mass Spectrometry
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7. Cover the purified HCCA precipitate in the glass filter with foil. Dry 3 to 4 hr, preferably in a laminar-flow hood, and transfer to glass vials. Store below 0°C (stable for years, if repeated freezing/thawing is avoided). REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Peptide matrix/nitrocellulose solution Dissolve 10 mg nitrocellulose and 40 mg/ml α-cyano-4-hydroxy-trans-cinnamic acid (HCCA; Sigma or Aldrich) in 1 ml acetone. If necessary, vortex 1 to 2 min to dissolve. Add 1 ml isopropanol (final 20 mg/ml HCCA and 5 mg/ml nitrocellulose in acetone/isopropanol). Prepare fresh before each use. Peptide matrix solution Prepare 20 mg/ml α-cyano-4-hydroxy-trans-cinnamic acid (HCCA; Sigma or Aldrich) either in 0.1% (v/v) trifluoroacetic acid (TFA)/50% (v/v) acetonitrile or in 99:1 (v/v) acetone/water (also see Support Protocol 1). Alternatively, use 20 mg/ml 2,5-dihydroxybenzoic acid (DHB) in 0.1% (v/v) TFA/30% (v/v) acetonitrile. Divide into aliquots and store at −20°C (stable for months to years). With DHB, the percentage of acetonitrile may be varied depending on the hydrophobicity of the peptide(s) analyzed. The matrix additive 2-methoxy-5-benzoic acid may be added at 1 mg/ml to the DHB matrix solution for better detection of larger peptides (>3000 Da).
Peptide standard mixture Prepare a mixture of peptides with known masses (e.g., bradykinin, mellitin, and ACTH) at ∼1 pmol/µl in 0.1% (v/v) TFA. Divide into aliquots and store at −20° or −80°C (stable for years). Protein matrix solution Prepare 20 mg/ml sinapinic acid in 0.1% (v/v) trifluoroacetic acid (TFA)/50% (v/v) acetonitrile or in 99:1 (v/v) acetone/water. Divide into aliquots and store up at −20°C (stable for years). Protein standard mixture Prepare a mixture of insulin, myoglobulin, and BSA at ∼1 pmol/µl (v/v) in 0.1% TFA. Divide into aliquots and store at −20°C (stable for years). COMMENTARY Background Information
Sample Preparation for MALDI Mass Analysis
Different on-target sample preparation methods have been developed for MALDI mass spectrometry. The most frequently used is the dried droplet method, in which the matrix is simply added to the sample on the sample stage (see Basic Protocol 1 above; see UNIT 11.6, Basic Protocol 3). Alternatively, a thin layer of matrix material dissolved in a rapidly evaporating solvent may be prepared on the sample target holder, onto which a solution of the analyte sample is subsequently deposited to avoid complete dissolution of the preprepared matrix film. The latter method has provided improved sensitivity and uniform desorption properties
(Vorm et al., 1994). Moreover, the addition of nitrocellulose to the matrix solution may be employed to achieve better peptide binding to the target, allowing more extensive rinsing of the sample (see Basic Protocol 2; also Alternate Protocol of UNIT 11.6; Shevchenko et al., 1996).
Critical Parameters and Troubleshooting The method of sample preparation as well as the matrix solvent composition can significantly affect results (Table 16.3.1) due to differential mass discrimination and ion suppression effects (Cohen and Chait, 1996). This is especially true when complex peptide mixtures
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are analyzed, such as unseparated protein digests. For example, in tryptic digest mixtures, the C-terminal arginine-containing components are preferentially detected. Therefore, to obtain better detection of most of the components in such a mixture, spectra should be acquired using different matrix solution conditions and different matrices. Contaminants such as salts and detergents should be <50 mM. When crystallization is impaired due to the presence of contaminants,
the matrix/sample preparation should be rinsed 1 to 3 times with ice-cold 0.1% (v/v) TFA or 10% (v/v) formic acid, and the solvent should be removed after a few seconds by pipetting or with blown pressurized air. However, hydrophilic peptides may be lost in this washing procedure, and it is not suitable for DHB matrix solutions. Alternatively, the sample can be diluted and reapplied to the target. Volatile buffers may be removed by partial drying with vacuum centrifugation. Complete lyophilization is not
Table 16.3.1 Effects of Matrix Solution Composition, pH, Presence of Detergent, and Crystallization Rate on MALDI-MS of a Complex Mixture of Peptidesa
Relative peptide ion intensity
Composition of HCCA solution 0.1 N HCl/ACN (2:1) 1% TFA/ACN (2:1) Formic acid/water/ACN (1:3:2) Formic acid/water/IPA (1:3:2) 0.5% TFA/ACN (2:1) Formic acid/water/methanol (1:3:2) 0.01 N HCl/ACN (2:1) Formic acid/water/IPA (0.2:3:2) 0.1% TFA/ACN (2:1) 0.1% TFA/methanol (2:1) 0.1% TFA/IPA (2:1) 2 M acetic acid/ACN (2:1) Water/ACN (1:1) Water/ACN (2:1) Water/IPA (2:1) Water/ethanol (2:1) Water/methanol (2:1) N-Octyl-â-D-glucoside Water/ACN (2:1) alone 20 mM in water/ACN (2:1) Rate of crystallization Formic acid/water/IPA (1:3:2) Slow Dried droplet Rapid Water/ACN (2:1) Slow Dried droplet Rapid
pH
0.8-2 kDa
2-6 kDa
6-11 kDa
1.1 1.1 1.1 1.3 1.4 1.6 1.9 2.0 2.0 2.0 2.1 2.3 2.4 2.5 2.7 2.8 2.9
+ + + + + + ++ ++ ++ ++ ++ ++ +++ +++ +++ +++ +++
+++ +++ ++ ++ +++ ++ +++ +++ +++ +++ +++ +++ ++ ++ ++ ++ ++
++ ++ +++ +++ ++ +++ ++ ++ ++ ++ ++ ++ + + + + +
+++ +++
++ +++
+ +++
+ + +++
+++ +++ +
ND ND ND
+ ++ +++
+++ ++ +
ND ND ND
aPeptide mixture is a partial V8 protease digest of the 11 kDa protein Max (for details see Cohen and Chait, 1996). The
dried droplet method (see Basic Protocol 1) was used for the analysis of the effect of matrix composition/pH and N-octyl-β-D-glucoside on MALDI-MS performance. For each matrix condition, the three mass ranges are assigned a response level: +++ designates the mass range with the highest relative ion intensities; ++ indicates moderate ion intensities; + indicates weak or no peptide ions. Abbreviations: ACN, acetonitrile; IPA, isopropyl alcohol; ND; not determined; TFA, trifluoroacetic acid.
Mass Spectrometry
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recommended because of severe sample losses. A partially lyophilized sample can be diluted with 0.1% to 5% (v/v) TFA in 20% (v/v) acetonitrile to reduce adsorptive losses and to facilitate MALDI mass analysis.
Anticipated Results Using MALDI-MS, peptides generally yield singly protonated ions, and reasonable signal-to-noise ratios can be obtained, even in the low femtomole range. When salt is present in the sample, sodium (+22.99) and potassium (+39.10) adducts may be observed in the spectrum as well. Mass accuracy for linear time-of-flight (TOF) mode with external calibration is typically ±2 to 3 Da for peptides <3000 Da. Reflector-TOF instruments offer improved mass accuracy with external calibration (±0.5 to 1 Da), but a 5- to 10-fold loss in signal may be observed, especially for larger peptides that are more prone to metastable fragmentation. With delayed extraction in reflectron mode, the fullwidth, half-maximum mass resolution (M/∆M, FWHM) can reach >10,000 Da for analytes that have mass-to-charge ratios of 1000 to ∼6000 kDa, and mass accuracies in the low parts-permillion range can be achieved at low levels of gel-separated proteins (Clauser et al., 1996 and unpub. observ.). In addition, delayed extraction provides significant improvement in MALDI spectral quality, including a lower level of chemical noise arising from fragmentation in the desorption plume, reduction or elimination of matrix background, and significant reduction in the dependence of ion flight times on laser intensity (Vestal et al., 1995). Peak width and centroids are much less dependent on the absolute intensity of the ion signal. Therefore, high mass resolution and improved mass accuracy can be achieved even when the peaks are small.
Time Considerations One of the main advantages of MALDI-MS is its speed: MALDI sample preparation only takes a few minutes and a typical MALDI mass measurement may take less than a minute, or up to 10 min when very low amounts of sample or high levels of contaminants are present.
Literature Cited Clauser, K.R., Baker, P., and Burlingame, A.L. 1996. Peptide fragment-ion tags from MALDI/PSD for error-tolerant searching of genomic databases. In Proceedings of the 44th ASMS Conference on Mass Spectrometry and Allied Topics, p. 365. American Society for Mass Spectrometry, Portland, Oreg. Cohen, S.L. and Chait, B.T. 1996. Influence of matrix solution conditions on the MALDI-MS analysis of peptides and proteins. Anal. Chem. 68:31-37. Shevchenko, A., Wilm, M., Vorm, O., and Mann, M. 1996. Mass spectrometric sequencing of proteins from silver-stained polyacrylamide gels. Anal. Chem. 68:850-858. Vestal, M.L., Juhasz, P., and Martin, S.A. 1995. Delayed extraction matrix-assisted laser desorption time of flight mass spectrometry. Rapid Commun. Mass Spectrom. 9:1044-1050. Vorm, O., Roepstorff, P., and Mann, M. 1994. Improved resolution and very high sensitivity in MALDI TOF of matrix surfaces made by fast evaporation. Anal. Chem. 66:3281-3287.
Key References Burlingame, A.L. and Carr, S. (eds.) 1996. Mass Spectrometry in the Biological Sciences. Humana Press, Totawa, N.J. This book may serve as an introduction to the field of biological mass spectrometry. Burlingame, A.L., Boyd R.K., and Gaskell, S.G. 1998. Mass spectrometry. Anal. Chem. 70:647716. An extensive overview of the literature on the use of mass spectrometry for protein studies between 1997 and mid-1998. Mann, M. and Talbo, G. 1996. Developments in matrix-assisted laser desorption/ionization mass spectrometry. Curr. Opin. Biotechnol. 7:11-19. A review of the progress of MALDI-MS up to 1996. Roepstorff, P. 1997. Mass spectrometry in protein studies from genome to function. Curr. Opin. Biotechnol. 8:6-13. A comprehensive review on applications and recent developments in MS instrumentation.
Contributed by C.R. Jiménez, L. Huang, Y. Qiu, and A.L. Burlingame University of California, San Francisco San Francisco, California
Sample Preparation for MALDI Mass Analysis
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In-Gel Digestion of Proteins for MALDI-MS Fingerprint Mapping
UNIT 16.4
Polyacrylamide gel electrophoresis is a widely used technique for separating proteins from biological samples. Moreover, the development of two-dimensional gel electrophoresis (UNIT 10.4) has provided a tool for differential protein display, which allows for the quantitative analysis of thousands of proteins simultaneously. Complex biological questions can now be approached by analyzing differences in the two-dimensional gel patterns between control and experimental states (Fig. 16.4.1; Patterson and Aebersold, 1995; Arnott et al., 1998; Qiu et al., 1998). Although highly sensitive staining methods could visualize proteins at increasingly low levels, originally protein identification was limited by the sensitivity of Edman sequencing. However, in the last few years mass spectrometry (MS) has emerged as a more sensitive, versatile, and rapid alternative, following the advent of electrospray ionization mass spectrometry (ESI-MS) and matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS). The advantages of MALDI-MS over ESI-MS include its relatively high tolerance to contamination from biological matrices, its high sensitivity, the relative ease of interpreting spectra from mixtures, and the formation of singly protonated
untreated cells
treated cells
excise protein spot of interest and destain/wash
enzymatically digest protein in gel plug
extract peptides from the gel and mass analyze Figure 16.4.1 Protein in situ (in-gel) digestion. A spot from an SDS-PAGE gel is excised, cut into small particles (not shown), and placed in a silanized tube. The gel particles are destained, washed, dried, and rehydrated with a protease. The resulting peptides are extracted from the gel plug and subjected to mass measurement. Contributed by C.R. Jiménez, L. Huang, Y. Qiu, and A.L. Burlingame Current Protocols in Protein Science (1998) 16.4.1-16.4.5 Copyright © 1998 by John Wiley & Sons, Inc.
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tryptic peptides
protein sequence database theoretical digest
MALDI-MS peptide masses
theoretical peptide masses MS-Fit
PSD/ tandem MS
theoretical fragmentation protein match
peptide fragment masses
theoretical fragment masses
MS-Tag
Figure 16.4.2 Peptide fingerprint mass mapping and peptide fragmentation mapping, using the database search programs MS-Fit and MS-Tag (http://prospector.ucsf.edu), respectively. The masses of peptides generated by enzymatic cleavage of the protein of interest (left) are matched with peptides generated by theoretical cleavage of each protein in the database (right) using the search program MS-Fit (see UNIT 16.5 for further explanation). The masses of the post-source decay (PSD) ions or fragment ions (left) are matched with theoretical fragmentation products of each protein in the database (right) using the search program MS-Tag (see UNIT 16.6 for details). For a conclusive protein identification, MS-Tag search results should confirm MS-Fit search results. When no reasonable matches are obtained, the protein may be unknown, in which case MS/MS sequence information is needed to design oligonucleotides for cloning.
molecular ions for tandem analysis (see UNIT 16.6 for analysis of MALDI collision-induced dissociation, or MALDI-CID). Peptide fingerprint mass mapping (see UNIT 16.5) and partial peptide sequencing using post-source decay (PSD; see UNIT 16.6), and ladder sequencing by MALDI-MS in combination with algorithms for sequence database interrogation have the potential for identification and structural investigation of proteins (Fig. 16.4.2). This protocol describes the in-gel digestion procedure as it is routinely performed in the authors’ laboratory for peptide mass mapping of picomole to subpicomole quantities of protein derived from Coomassie- (UNIT 10.5) or silver-stained polyacrylamide gels (Shevchenko et al., 1996). First, SDS is removed from the excised gel fragments using several ammonium bicarbonate/acetonitrile washing steps. The gel pieces are subsequently dried and rehydrated with the digestion enzyme (trypsin) in buffer. Alternatively, the washed gel pieces may be reduced and S-alkylated prior to rehydration with enzyme. After digestion, the peptides are extracted from the gel and mass analyzed. BASIC PROTOCOL
In-Gel Digestion for MALDI-MS Fingerprint Mapping
IN-GEL DIGESTION PROCEDURE Materials Stained polyacrylamide gel containing protein band of interest (UNITS 10.1, 10.4 & 10.5) 25 mM ammonium bicarbonate/50% (v/v) acetonitrile 10 mM dithiothreitol (DTT) in 25 mM ammonium bicarbonate 55 mM iodoacetamide in 25 mM ammonium bicarbonate 25 mM ammonium bicarbonate, pH 8
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0.05 to 0.1 mg/ml trypsin (sequence grade; Promega) in 25 mM ammonium bicarbonate, pH 8 5% (v/v) trifluoroacetic acid (TFA)/50% (v/v) acetonitrile 0.65-ml tubes (e.g., PGC Scientific), silanized (APPENDIX 3E) Vacuum centrifuge 56°C water bath Sonication bath NOTE: To avoid or reduce contamination with human keratins, one should wear gloves and preferably work in a laminar-flow hood. Excise and wash gel fragments 1. Excise protein bands/spots of interest from a stained polyacrylamide gel. Cut each gel piece into small particles (∼1 mm2) using a scalpel, and place into a 0.65-ml siliconized tube. Also cut out and dice a gel piece from a protein-free region of the gel, for a parallel control digestion to identify trypsin autoproteolysis products. The amount of protein loaded on the gel must be determined empirically, as it depends on properties of the specific protein (for example, degree of hydrophobicity and size). In general, picomole levels are preferred to lower levels, although some reports claim that femtomole sensitivities are attainable with in-gel digestion and mass spectrometry. The small gel particle size facilitates the removal of SDS (and Coomassie) during the washes, and improves enzyme access to the gel. For Coomassie-stained proteins, one gel particle per tube is probably sufficient protein. With faint silver-stained spots, particles can be pooled, but no more than three should be pooled as this will lead to excessive levels of contaminants. For a silver-staining protocol compatible with mass spectrometry, see Shevchenko et al. (1996).
2. Add ∼100 µl of 25 mM ammonium bicarbonate/50% acetonitrile (or enough to immerse the gel particles) and vortex for 10 min. Use gel-loading pipet tips to remove the solution (pale blue in the case of Coomassie staining) and discard. Repeat this wash/dehydration step up to ∼3 times. At this point, the gel slices shrink and become white. This visual criterion should be used to determine whether or not additional washes should be performed.
3. Dry gel particles for ∼30 min in a vacuum centrifuge. Perform reduction and alkylation (optional) 4. Add enough 10 mM DTT solution to cover the gel pieces, and reduce for 1 hr at 56°C. Steps 4 to 7 may be included when a maximum of protein coverage is required or when digesting a band from a one-dimensional gel. Proteins separated by two-dimensional gel electrophoresis are already reduced and alkylated, so their steps can be omitted.
5. Cool to room temperature and replace the DTT solution with roughly the same volume of 55 mM iodoacetamide solution. Incubate for 45 min at room temperature in the dark with occasional vortexing. 6. Wash gel pieces (rehydrate) with ∼100 µl of 25 mM ammonium bicarbonate, pH 8, for 10 min while vortexing, and dehydrate with ∼100 µl of 25 mM ammonium bicarbonate/50% acetonitrile. Repeat rehydration and dehydration. 7. Remove the liquid phase and dry the gel pieces in a vacuum centrifuge. Digest protein sample 8. Rehydrate gel particles in 1 vol of 0.05 to 0.1 mg/ml trypsin solution by vortexing for 5 min. Do not add more solution than can be absorbed by the gel particles. Mass Spectrometry
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The volume needed can be estimated by calculating the total gel volume excised (e.g., 2 mm × 8 mm × 1 mm = 16 mm3 = 16 µl). The enzyme-to-substrate ratio employed for in-gel digestions (>1:10) is greater than for in-solution digestions due to hindered enzyme access to the protein substrate in the gel. Moreover, the relatively low salt concentration (25 mM) is used to reduce the possibility of subsequent salt interference with ionization in the mass spectrometer.
9. If necessary, overlay the rehydrated gel particles with a minimum amount of 25 mM ammonium bicarbonate to keep them immersed throughout digestion. 10. Incubate 12 to 16 hr at 37°C. Recover peptides 11. Add 2 vol water, vortex for 5 min, and then sonicate for 5 min. Use a gel-loading tip to remove the peptide solution and transfer it to a silanized tube. 12. Perform two additional extractions using 2 vol of 5% TFA/50% acetonitrile. The silanized microcentrifuge tubes and the high TFA concentration are used to minimize adsorptive sample loss. Formic acid (5%) may be used as an alternative to TFA.
13. Concentrate recovered peptides by reducing the final volume of the extracts to ∼10 µl in a vacuum centrifuge. Bring to 25 µl with 5% TFA/50% acetonitrile. Store the recovered peptides at −20°C until MALDI-MS is performed. For MALDI-MS analysis, mix 0.5 µl of the unseparated digest on the sample target with 0.5 µl of 20 mg/ml α-cyano-4-hydroxycinnamic acid in 0.1% TFA/50% acetonitrile and let air dry (for additional details, see UNIT 16.3, Basic Protocols 1 and 2). Insert sample target into the mass spectrometer and mass analyze as recommended by the manufacturer.
14. Optional: Decrease the amount of volatile salts by adding a few cycles of water (100 to 200 µl) and subsequently reducing the volume in a vacuum centrifuge. Salt reduction may improve mass signals, particularly at subpicomole levels.
COMMENTARY Background Information
In-Gel Digestion for MALDI-MS Fingerprint Mapping
Generally, in-gel tryptic digests of proteins yield higher recovery of peptides than proteins digested after transblotting to polyvinylidene difluoride (PVDF) membranes (UNIT 11.2). Trypsin is the protease of choice for peptide mass fingerprinting because of its reliability and its substrate specificity, yielding peptides with Cterminal basic residues (Arg and Lys), which facilitates ionization and subsequent mass spectrometric sequencing. However, autolysis of the protease contributes contaminating peptides that can interfere with the analysis. For trypsin, this problem is partially overcome by using modified porcine enzyme. A few labile sites are often cleaved nonetheless, resulting in peptides with monoisotopic ions of 842.51, 2211.1045, and 2284.6, which may be used for internal accurate mass calibration of the spectrum. Endoprotease Lys-C is a good alternative to trypsin, because it produces less autolysis and it is more stable under denaturing condi-
tions. Unfortunately, endo-Lys-C yields fewer, larger fragments that may be released from the gel particles less efficiently. However, larger average lengths provide greater specificity for the search algorithm (UNIT 16.5).
Critical Parameters and Troubleshooting Low concentrations of ammonium bicarbonate in the digestion buffer give the best results for subsequent mass measurement by MALDI-MS. Adsorption of the released peptides to the sample tube and other surfaces is a problem when working in the femtomole range. To prevent sample losses, digestion is performed in silanized tubes. Moreover, the presence of organic solvent (10% acetonitrile) in the digestion buffer aids in solubility without degrading enzyme activity. The use of low amounts of pure, single-component detergents (e.g., N-octyl-β-D-glucoside) during digestion may also improve peptide recovery.
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If a higher peptide coverage of the protein is required, the amount of organic solvent and the acidity of the matrix composition can be varied (see Cohen and Chait, 1996), as can the matrix. For example, α-cyano-4-hydroxycinnamic acid is the matrix of choice for best detection limits and post-source decay analysis. However, 2,5-dihydroxybenzoic acid (DHB), or DHB with fucose, yields spectra of peptide mixtures with the least suppression characteristics. If sequence coverage is still insufficient, use a microbore liquid chromatography/ESI-MS (LC-ESI-MS) system.
Anticipated Results MALDI-MS analysis of unseparated tryptic digests generally yields 40% to 50% sequence coverage, although peptide recovery may be lower for hydrophobic proteins. Typically, with sufficient mass accuracy, four tryptic peptides may already be enough to identify a protein from the database using the search program MS-Tag (see UNIT 16.6).
Time Considerations Generally, the in-gel digestion procedure is performed overnight. Shorter digestion times (e.g., 6 hr) are possible, but at the expense of less complete digestion.
Literature Cited Arnott, D., O’Connell, K.L., King, K.L., and Stults, J.T. 1998. An integrated approach to proteome analysis: Identification of proteins associated with cardiac hypertrophy. Anal. Biochem. 258:118.
Cohen, S.L. and Chait, B.T. 1996. Influence of matrix solution conditions on the MALDI-MS analysis of peptides and proteins. Anal. Chem. 68:31-37. Patterson, S.D. and Aebersold, R. 1995. Mass spectrometric approaches for the identification of gel-separated pr oteins. Electrophoresis 16:1791-1814. Qiu, Y., Benet, L.Z., and Burlingame, A.L. 1998. Identification of the hepatic protein targets of reactive metabolites of acetaminophen in vivo in mice using two-dimensional gel electrophoresis and mass spectrometry. J. Biol. Chem. 273:17940-17953. Shevchenko, A., Wilm, M., Vorm, O., and Mann, M. 1996. Mass spectrometric sequencing of proteins from silver-stained polyacrylamide gels. Anal. Chem. 68:850-858.
Key References Jungblut, P. and Thiede, B. 1997. Protein identification from 2-DE gels by MALDI mass spectrometry. Mass Spectrom. Rev. 16:145-162. This extensive review article provides a comprehensive introduction to the analysis of gel-separated proteins. Fenselau, C. 1997. MALDI-MS and strategies for protein analysis. Anal. Chem. 69:661A-665A. A tutorial-type review on the use of MALDI-MS for protein analysis.
Contributed by C.R. Jiménez, L. Huang, Y. Qiu, and A.L. Burlingame University of California, San Francisco San Francisco, California
Mass Spectrometry
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Searching Sequence Databases Over the Internet: Protein Identification Using MS-Fit Peptide fingerprint mass mapping is widely used for initial identification of proteins separated by gel electrophoresis (Cottrell, 1994; Jungblut and Thiede, 1997). This method involves the enzymatic in-gel digestion of proteins to generate peptides (UNIT 16.4), followed by mass measurement of the cleaved peptides. The experimental mass values are then compared with theoretical values from protein databases, calculated using the cleavage specificity of the enzyme. This protocol describes the use of the search program MS-Fit, which can be accessed through ProteinProspector at http://prospector.ucsf.edu. By using an appropriate scoring algorithm, the closest match or matches can be identified (Cottrell, 1994).
UNIT 16.5
BASIC PROTOCOL
Peptide masses can be determined using either a matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) or liquid chromatography/electrospray ionization (LC-ESI) mass spectrometer. MALDI-TOF mass spectrometry has been preferred for direct analysis of unseparated digest mixtures because of its tolerance to low levels of contaminants such as buffer salts. Because of the inherently high throughput, sensitivity, and simplicity of this technique, MALDI peptide mapping is now a powerful method for identification of proteins present at greater than or equal to ∼0.5-pmol on a gel (Shevchenko et al., 1996). Input data and set search parameters 1. Input the peptide masses obtained by mass spectrometry into a spreadsheet, and then copy and paste the values into the space for masses (m/z) in the MS-Fit program. Monoisotopic peptide masses from MALDI-TOF are preferred. Additional peptide mass values obtained in parallel from multiply charged ions (m/z) using electrospray ionization mass spectrometry along with charge state (z) are submitted at the same time. Average masses may also be used for the search; however, more false positive results will be obtained due to the lower mass accuracy.
2. Set mass tolerance, which should be no less than the mass accuracy obtained experimentally. Typical mass accuracy values for external calibration should be ∼100 to 300 ppm for delayed-extraction MALDI-TOF, and ∼100 ppm for electrospray orthogonal TOF. For internal calibration, ∼5 to ∼20 ppm should be used for both methods. Typical mass accuracy of ∼ 0.5 Da is appropriate when regular MALDI-TOF (i.e., without delayed extraction) is used with internal calibration.
3. Choose a minimum number of peptides required for matching a particular protein in the database to generate a hit. The default value is four, which can be changed depending on the search output and the total number of peptides submitted. If the number of search results is too large (i.e., >25), the minimum number of peptides required to match can be increased gradually to eliminate the number of false positives. In general, if the search results give more than five different proteins, and the number of hits exceeds 25 (except all the hits referring to one protein in different species), a false positive is present.
4. Choose a protein database to use for the search. Choose any database listed in the program, such as NCBInr, Genpept, Swiss Prot, Owl, and dbEST. NCBInr is recommended as the first choice for the search, because it combines most of the public domain protein databases (though dbEST is not included) to generate the largest nonredundant protein database, and because it is updated most frequently. Swiss Prot may also be chosen because it is the best annotated database. Contributed by C.R. Jiménez, L. Huang, Y. Qiu, and A.L. Burlingame Current Protocols in Protein Science (1998) 16.5.1-16.5.6 Copyright © 1998 by John Wiley & Sons, Inc.
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5. If the source of the protein of interest is known, select the species to perform a species-limited search. Alternatively, designate “all” for the species. In general, it is useful to designate “all,” because it allows proteins occurring in different species with high homology to be found. Moreover, species prefiltering is imperfect because of poor usage of taxonomy (standard species naming conventions) in the databases. If the Latin taxonomic name for the species of interest is not known, see the NCBI Taxonomy Browser at http://www.ncbi.nlm.nih.gov/Taxonomy/tax.html.
6. Select the enzyme specificity and the number of “missed cleavage sites” to be employed. MS-Fit will search the peptide mass values submitted against calculated theoretical values of peptides cleaved by the specified enzyme. The termini of the matched peptides will be consistent with the cleavage specificity of the enzyme used to generate the peptide. Increasing the maximum number of allowed missed cleavages, enables matching to sequences with uncleaved sites internal to the peptide. If the digestion time employed is fairly long (>6 hr), it is possible (even likely) to find peptides present in the digest that are derived from nonspecific proteolytic cleavage. The option for employing the fictive enzyme “slymotrypsin” allows the occurrence of chymotryptic-like cleavages in tryptic digests. When using this choice it is important to increase the amount of missed cleavages allowed to three or more. It is also possible to combine the rules for two or more enzymes by adding options to the Enzyme item on the HTML form.
7. Optional: Select the molecular weight and pI range. Perform the first search with no molecular weight or pI restriction. Then adjust the range according to the search output. Because molecular weights from SDS-PAGE are not generally accurate, the range needs to be set at ±10 kDa. If the protein being analyzed is a processing or degradation product of a larger precursor, the molecular weight estimated from the gel would not be useful for the search.
8. Choose the desired types of modified residues to be searched. a. Specify modifications desired for cysteine residues to generate theoretical masses to match those cysteine-containing peptides that have been reduced and then alkylated with iodoacetic acid (carboxymethylation), iodoacetamide (carbamidomethylation), or 4-vinylpyridene (pyridylethylation). b. Specify additional chemical modifications of proteins that can be generated during PAGE, such as oxidation of methionine residues and acrylamide modification of cysteine residues. c. Specify peptide mass changes due to known posttranslational modifications, such as conversion of N-terminal glutamine to pyroglutamic acid, and phosphorylation of serine, threonine, and tyrosine. d. For any database entry with a methionine at the N terminus, specify whether the N-terminal peptide is to be considered in its original form or in a form where the methionine is removed and the penultimate amino acid is acetylated. If the database curators have removed the N-terminal methionine from the sequence, MS-Fit will not apply the acetylation modification.
Searching Sequence Databases Over the Internet: MS-Fit
Perform the search 9. Initiate the search. Two scoring systems are automatically displayed in search results: a modified MOWSE score (Pappin et al., 1993) and an MS-Fit rank. The higher the score, the higher number of peptide masses matched and the more likely the hit is of relevance. The output indicates the number of peptide masses matched and their corresponding sequences, and the matching protein’s molecular weight, pI, accession number, species, and protein name. Click on the MS-digest index number to display
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the coverage of the matched peptides in the entire protein sequence. Click on the accession number to link to the NCBI Entrez. A detailed description of the MOWSE and MS-Fit scoring system can be found by clicking on this parameter in the program. Currently, MS-Fit employs only a simple ranking system. The results are sorted so that if multiple database entries are matched, the more likely matches are listed higher in the rank. All database entries matching the input data and parameters are ranked on the following basis: (1) database entries with the least number of unmatched masses are ranked higher, and (2) among equivalent matches (those with the same rank), the results are sorted in order of increasing index number.
10. To perform an alternative search in homology mode, select the box that allows for a single amino acid substitution per peptide. In this mode, the MS-Fit option for other possible modifications is by-passed. In practice, the homology mode should only be used when one or more of the following conditions applies: the peptide mass data have excellent mass accuracy (±10 ppm or better), a narrow intact protein molecular weight filter is used, and/or the hits will be saved and searched using MS-Tag (UNIT 16.6). The minimum amount of peptides with no amino acid substitution (usually set at one or two) controls the protein entry pulled from the database. The output displays the necessary single amino acid substitution and the corresponding sequence consistent with the experimental peptide mass data (not the sequence present in the database). These predictions need to be confirmed by MALDI post-source decay (MALDIPSD) or tandem mass spectrometry.
11. Optional: To use one ProteinProspector search program as a prefilter for another search program, save the hits (index numbers for matching database entries) from the first program to a user-specified file. Retrieve this file using the second program, so that only those matching database entries are searched by the second program. Saved hits from MS-Fit can be used by other programs such as MS-Tag (UNIT 16.6) and MS-Edman.
COMMENTARY Background Information Initially, protein identification using peptide mass mapping was hampered mainly by the poor mass accuracy obtained using MALDI mass spectrometers, making unambiguous identification difficult. Recently, the development of MALDI-TOF equipped with delayed extraction has dramatically improved the mass resolution and mass accuracy (Vestal et al., 1995; Edmondson and Russell, 1996) at the subpicomole to femtomole protein levels. Moreover, the rapid growth of gene and protein databases resulting from large-scale genome sequencing projects of different organisms now offers significant power for mass spectrometric identification and characterization of proteins isolated from gels. The advantage of the improved mass accuracy using delayed extraction is increased search specificity of each peptide, leading to a better discrimination of true hits versus falsepositive search results (Clauser et al., unpub. observ.). Finally, peptide mass mapping is not
only useful for identification of proteins existing in databases, but also for cross-species identification. This possibility relies on sequence conservation between two proteolytic cleavage sites (a stretch of ∼10 to 15 amino acids) in homologous proteins of different organisms (Cordwell et al., 1997; Shevchenko et al., 1997).
Critical Parameters and Troubleshooting Measuring masses as accurately as possible is the single most important factor in achieving the highest certainty of protein identification in a peptide mass fingerprinting experiment (K.R. Clauser, unpub. observ.). The more accurate the masses measured, the lower the mass tolerance is set, and the more unambiguous the results obtained. Several steps must be followed to obtain reliable protein identification using peptide mass mapping, especially when dealing with cases yielding ambiguous search results. First,
Mass Spectrometry
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sort out all of the unassigned experimental peptide masses by attributing them to posttranslationally modified peptides (such as modification of cysteine, methionine, and glutamine; see Basic Protocol, step 8), peptides with nonspecific cleavage sites, enzyme autolysis products, adducts (sodium, potassium, and matrix adducts), matrix peaks, and contaminants (keratins from human and other species). Series of peaks with constant mass differences can be attributed to organic polymers, which may come from tubes or may have been added during the isolation or preparation procedures (Jungblut and Thiede, 1997). If there are still quite a few abundant peaks left unassigned by such means, perform a second database search using the set of unmatched mass values excluded from the first match hit, because more than one protein may be present in the gel spot due to comigration during gel electrophoresis (Jensen et al., 1996). Second, try another digest using a different enzyme, search both sets of data, and select the results that are common to both lists. The chance of obtaining a protein with a high score in both searches is extremely small (Jensen et al., 1996). When no match is found in the protein databases, this may suggest that the gene encoding the protein of interest has not yet been sequenced or cloned. However, it may be known as an expressed sequence tag (EST) that represents part of the protein. Therefore, it may be useful to screen a dbEST database. Search times will typically be longer because of the
multiframe translations and because the dbEST file is >3 times larger than the NCBInr file. When an EST database is selected, the number for DNA frame translation needs to be set. A user should select frame mode 6 unless it is known that the database being searched contains sequences exclusively cloned in one direction or contains known genes with sequences already in frame. Of course, a longer search time will be required when using frame mode 6 instead of 3. Users can also generate their own database for searching; the instructions are in the ProteinProspector home page. Finally, to obtain a protein identification unambiguously, in the authors’ opinion, it is always prudent and in fact may be necessary to obtain a partial peptide sequence to choose correctly among possible MS-Fit search result possibilities, unless information from other experiments indicates what the protein could possibly be.
Anticipated Results As an example, the use of MS-Fit for unambiguous protein identification is illustrated here. Figure 16.5.1 shows a spectrum obtained by delayed-extraction MALDI-reflectron-TOF mass spectrometry of an in-gel-digested protein spot from a two-dimensional gel. The peptide masses were externally calibrated using peptide standards. Twelve monoisotopic (Burlingame and Carr, 1996) peptide masses (Fig. 16.5.1) were submitted into the MS-Fit program. In this case, four modifications were considered: conversion of peptide N-terminal
2273.2222 2289.2285
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1744.9784
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1130.5549 1152.6232 1217.6711 1334.8252 1337.7146 1380.7812 1396.7758 1471.8635
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0 800
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1000
1200
1400
1600
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Figure 16.5.1 Mass spectrum obtained by delayed-extraction MALDI-reflectron-TOF mass spectrometry of an in-gel-digested protein spot from a two-dimensional gel.
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MS-Fit Search Results Press stop on your browser if you wish to abort this MS-Fit search prematurely. Sample ID (comment): Spot #3, MALDI mass mapping Database searched: NCBInr.06.09.98 Molecular weight search (1000-30000 Da) selects 180441 entries. Full pI range: 309620 entries. Species search (RATTUS NORVEGICUS) selects 6834 entries. Combined molecular weight, pI and species searches select 3032 entries. MS-Fit search selects 1 entry. Considered modifications: | Peptide N-terminal Gln to pyroGlu | Oxidation of M | Protein N-terminus Acetylated | Min. # Peptides Peptide Mass Peptide Masses Digest Max. # Missed Cysteines to Match Used Tolerance ( ) are Cleavages Modified by 100.000 ppm monoisotopic Trypsin acrylamide 4 1
Peptide N terminus Hydrogen (H)
Peptide C terminus Free Acid (O H)
Input # Peptide Masses 12
Result Summary
Rank 1
MOWSE Score 9.71e+003
# (%) Masses Matched 11/12 (91%)
Protein MW (Da)/pI 28419.4 / 5.29
NCBInr.06.09.98 Accession # Protein Name Species 130860 RATTUS RATTUS (D90258) proteasome subunit C8
Detailed Results 1. 11/12 matches (91%). 28419.4 Da, pI = 5.29. Acc. # 130860. RATTUS RATTUS. (D90258) proteasome subunit C8
Data submitted 763.3923 935.5383 1130.5549 1152.6232 1217.6711 1380.7812 1396.7758 1471.8635 1578.8493 1744.9784 1960.0006
MH+ . matched 763.4103 935.5427 1130.5556 1152.6053 1217.6490 1380.7422 1396.7371 1471.8161 1578.7876 1744.9135 1959.8936
Delta start ppm –23.5303 67 –4.6758 223 –0.6253 21 15.5002 101 18.1578 30 28.2609 73 27.7120 73 32.2353 197 39.0750 87 37.2063 101 54.5998 1
end 72 230 29 110 41 86 86 208 100 115 20
Peptide Sequence Modifications (Click for Fragment Ions) (R)LFNVDR(H) (K)GRHEIVPK(D) (R)VFQVEYAMK(A) 1 Met-ox (R)SNFGYNIPLK(H) (K)AVENSSTAIGIR(C) (R)HVGMAVAGLLADAR(S) 1 Met-ox (R)HVGMAVAGLLADAR(S) (K)IIYIVHDEVKDK(A) (R)SLADIAREEASNFR(S) (R)SNFGYNIPLKHLADR(V) Acet N (-)SSIGTGYDLSASTFSPDGR(V)
1 unmatched mass: 1337.7 The matched peptides cover 43% (110/255 AA’s) of the protein. Coverage Map for This Hit (MS-Digest index #): 79534
Figure 16.5.2 MS-Fit search result. A database search was performed using the tryptic masses shown in Figure 16.5.1. A single protein was identified after restriction of the molecular weight range to 20 to 40 kDa and restriction of the species to rat. Reprinted with permission of the Regents of the University of California.
Mass Spectrometry
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glutamine to pyroglutamate, oxidation of methionine, acetylation of protein N-terminus, and carboxymethylation or acrylamide modification of cysteine. The peptide mass tolerance was chosen as 100 ppm because external calibration was used. The default values were used for maximum number of missed cleavage sites and minimum number for peptides to match (both set at 4). A first search was performed without selection of a particular species (enter “all”) or molecular weight range (1000 to ∼100,000 Da). In this unrestricted search, 182 entries were obtained in the search results, of which the three highest ranking proteins all indicated proteasome subunit C8 in human and rat, due to the high interspecies homology. To reduce the number of false positives, the molecular weight range was set at 20 to ∼30 kDa, as determined by SDS-PAGE. Alternatively, the minimum number of peptides matched may be increased for this purpose. Thirteen entries were displayed in the second search result, and the high-ranking proteins were the same as in the first search. At this point, it is fairly certain that the protein of interest is proteasome subunit C8. This conclusion was further supported in the third search result, in which the species searched was limited to rat. As shown in Figure 16.5.2, 91% of the peaks submitted were matched. Two molecular ions matched modified peptides containing oxidized methionine (1130.5549, 1396.7758) and N-terminal acetylation (1960.0006). Subsequently, the latter peptide with [M + H]1+ at 1960.006 was analyzed further using MALDIPSD. This 42-Da shift in spectrum established the presence of N-terminal acetylation by observation of the appropriate ion sequence. When operating in reflectron mode, peptides containing oxidized methionine residues usually eliminate 64 u (mass units; −SOCH4) as a neutral moiety, and the resulting metastable fragment ion is not focused well and thus typically shows a 62-Da loss from the parent ion. The peak at 1334.8252 represents a fragment ion from peptide of [M + H]1+ at 1396.7758. In conclusion, this protein can be successfully identified as rat proteasome subunit C8 using the MS-Fit program. However, in general, to ascertain reliable protein identification, it is always advisable to confirm such MS-Fit results with independent peptide sequence data using either MALDI-PSD or MALDI collision-induced dissociation (MALDI-CID).
Time Considerations Search times depend on the database se-
lected (also see Critical Parameters and Troubleshooting) and on the quality of data submitted (e.g., the amount of tryptic peptides, the mass accuracy). They can vary from less than 1 min to 15 min or more.
Literature Cited Burlingame, A.L. and Carr, S.A. 1996. The meaning and usage of the terms monoisotopic mass, average mass, mass resolution, and mass accuracy for measurements of biomolecules. In Mass Spectrometry in the Biological Sciences (A.L. Burlingame and S.A. Carr, eds.), pp. 546-553. Humana Press, Totowa, N.J. Cordwell, S.J., Wasinger, V.C., Cerpa-Poljak, A., Duncan, M.W., and Humphrey-Smith, I. 1997. Conserved motifs as the basis for recognition of homologous proteins across species boundaries using peptide-mass fingerprinting. J. Mass Spectrom. 32:370-378. Cottrell, J.S. 1994. Protein identification by peptide mass fingerprinting. Pept. Res. 7:115-123. Edmondson, R. and Russell, D.H. 1996. Evaluation of matrix-assisted laser desorption ionizationtime-of-flight mass measurement accuracy by using delayed extraction. J. Am. Soc. Mass Spectrom. 7:995-1001. Jensen, O.N., Podtelejnikov, A., and Mann, M. 1996. Delayed extraction improves specificity in database searches by matrix-assisted laser desorption/ionization peptide maps. Rapid Commun. Mass Spectrom. 10:1371-1378. Jungblut, P. and Thiede, B. 1997. Protein identification from 2-DE gels by MALDI mass spectrometry. Mass Spectrom. Rev. 16:145-162. Pappin, D.J.C., Hojrup, P., and Bleasby, A.J. 1993. Rapid identification of proteins by peptide-mass finger-printing. Curr. Biol. 3:327-332. Shevchenko, A., Jesen, O.N., Podtelejnikov, A.V., Sagliocco, F., Wilm, M., Vorm, O., Mortensen, P., Boucherie, H., and Mann, M. 1996. Linking genome and proteome by mass spectrometry: Large scale identification of yeast proteins from two dimensional gels. Proc. Natl. Acad. Sci. U.S.A. 93:14440-14445. Shevchenko, A, Wilm, M., and Mann, M. 1997. Peptide sequencing by mass spectrometry for homology searches and cloning of genes. J. Prot. Chem. 16:481-490. Vestal, M.L., Juhasz, P., and Martin, S.A. 1995. Delayed extraction matrix-assisted laser desorption time of flight mass spectrometry. Rapid Commun. Mass Spectrom. 9:1044-1050.
Internet Resources http://prospector.ucsf.edu University of California’s ProteinProspector site.
Contributed by C.R. Jiménez, L. Huang, Y. Qiu, and A.L. Burlingame University of California, San Francisco San Francisco, California
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UNIT 16.6
Although peptide mass mapping is fast and simple, its success can be compromised by the purity of the protein in one gel spot, errors in the sequence database, and the detection of too few peptides in the MS map of a given protein to permit reliable database matching. In those cases, partial amino acid sequence that is determined using post-source decay (PSD) analysis or tandem mass spectrometry is required to establish the correct protein identification among possibilities suggested by MS-Fit (UNIT 16.5). Post-source decay (Kaufmann et al., 1993, 1994) involves the detection of the fragmentation product ions (metastable ions) of a selected mass value window containing the precursor ion chosen, that occur in the field-free region between the ion source and the reflectron. For a discussion of the basic principle of MALDI-PSD-MS for peptide sequencing, refer to UNIT 16.1. This unit covers some of the practical issues in obtaining and analyzing a typical PSD spectrum. The Basic Protocol introduces MS-Tag (see http:// prospector.ucsf.edu), a database interrogation program that enables the identification of proteins with a limited number of peptide fragment ions by matching existing database sequences. Although a typical PSD spectrum will usually not provide enough sequence ions for de novo manual sequence interpretation (particularly in the higher mass range, e.g., >500 to 800 Da), these ions together with the parent ion are statistically much better identifiers in database searching than only the molecular weights of tryptic peptides. Unlike peptide fingerprinting, database searching based on fragment ion information is often able to identify a single peptide sequence in the database that matches all the input information. Several computer programs employing fragment ion information to search databases are available on the internet (see http://donatello.ucsf.edu/onramp.html and sites listed therein). Among them, MS-Tag is a user-friendly program that simultaneously considers a variety of ion types, but still allows single-mismatch-tolerant searching. However, the authors suggest that a frequent user of MS-Tag should be able to interpret sequences from those spectra that do contain sufficient information to enable de novo interpretation. Although MALDI-PSD database searching is a widely used tool, an alternative method is analysis by MALDI collision-induced dissociation (MALDI-CID). This is presented in the Alternate Protocol. DATABASE SEARCHING WITH MS-TAG The basic principles of MS-Tag are detailed in the instruction section of the Web site (http://prospector.ucsf.edu). Users can find explanations for every input parameter by just clicking on the highlighted terms.
BASIC PROTOCOL
1. Type in the mass and charge state of the parent ion (masses detected in MS mode are preferred). 2. Define the parent ion mass tolerance. The tolerance range will depend on the instrument.
3. Input all fragment ions and define their mass tolerance, which is usually higher than the parent ion mass tolerance. Ions with higher intensities are preferred if a number of ions are available. Contributed by C.R. Jiménez, L. Huang, Y. Qiu, and A.L. Burlingame Current Protocols in Protein Science (1998) 16.6.1-16.6.7 Copyright © 1998 by John Wiley & Sons, Inc.
Mass Spectrometry
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1500
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b7 828.902
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Figure 16.6.1 PSD spectrum of a tryptic peptide of an unknown protein isolated from mouse liver. This peptide has sequence homology to bovine inorganic pyrophosphatase (DVFMVVEVPR), where one mutation is found (from E1 to D1).
4. Specify the type of input masses (e.g., all monoisotopic, all average, or parent ion is monoisotopic and fragment ions are average). 5. Specify the database searched and the type of frame translation if a nucleotide database is selected. The ability to search against MS-Fit hits is particularly useful when the target protein is not in current protein databases but is included in the expressed sequence tag (EST) database. The EST databases are currently much larger than other databases. Therefore, direct MS-Tag searching against an EST database is statistically much more demanding and may fail due to the much longer search time.
6. Specify the species from which the protein of interest is isolated, the protein molecular weight range, the type of enzyme used for protein digestion, the type of cysteine modification, and the maximum unmatched fragment ions. Default parameters may be used for other choices listed on the Web page. 7. Specify the search mode as identity or homology. Search in identity mode first; if not successful, then search in homology mode. Begin search. Typical results are depicted in Figure 16.6.1 and Figure 16.6.2, and are discussed in Anticipated Results.
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MS-Tag Search Results Press stop on your browser if you wish to abort this MS-Tag search prematurely. Sample ID (comment): Apo A-1 1040 AKPVLEDLR Database searched: NCBInr.05.04.98 Molecular weight search (1000 - 100000 Da) selects 287683 entries. Number of sequences passing through parent mass filter: 150221 MS-Tag search selects 1 entries. Ion Types Considered: a b B y n h l Search Max. # Peptide Masses Digest Max. # Missed Unmatched Ions are Used Cleavages Mode monoisotopic homology Trypsin 0 1
Cysteines Modified by acrylamide
Peptide N terminus Hydrogen (H)
Peptide C terminus Free Acid (O H)
Parent mass: 1327.6000 (± 0.6000 Da) Fragment Ions used in search: 174.9, 255.0, 268.9, 271.9, 284.9, 371.0, 415.9, 471.1, 499.0, 629.8, 729.2, 828.9 ( ±1.20 Da) Parent mass shift: ± 15.0 Da Composition Ions present: HF Result Summary
Rank 1
MS-Digest Index # 101341
Modification E1->D
NCBInr.05.04.98 Protein Accession # MW (Da) 585322 32844.4
Species BOVIN
Calculated MH+ (Da) 1341.6989
MH+ Error (Da) –14.0989
Sequence (K)DVFHMVVEVPR(W)
Protein Name INORGANIC PYROPHOSPHATASE (PYROPHOPHATE PHOSPHO-HYDROLASE) (PPASE) gil539751IpirllA45153 inorganic pyrophosphatase (EC 3.6.1.1) - bovine Detailed Results Rank
1
MS-Digest Index # 101341
Modification E1->D
NCBInr.05.04.98 Protein MW (Da) Accession # 585322 32844.4
Species BOVIN
Calculated MH+ (Da) 1341.6989
MH +Error (Da) –14.0989
Sequence (K)DVFHMVVEVPR(W)
Protein Name INORGANIC PYROPHOSPHATASE (PYROPHOPHATE PHOSPHO-HYDROLASE) (PPASE) gil539751IpirllA45153 inorganic pyrophosphatase (EC 3.6.1.1) - bovine Fragment-Ion 174.9 255.0 268.9 271.9 284.9 371.0 415.9 471.1 499.0 629.8 729.2 828.9 (m/z) y1 y2-NH3 HM y2 FH y3 FMH a4 Ion-type b4 b5 b6 b7 Delta Da –0.22 –0.15 –0.21 –0.27 –0.24 –0.24 –0.28 –0.14 –0.23 –0.47 –0.14 –0.49 y7 –0.56
Figure 16.6.2 MS-Tag search result. An MS-Tag search of the NCBInr database in homology mode with the PSD ions of the tryptic peptide shown in Figure 16.6.1 yields one single peptide sequence (DVFHMVVEVPR). Reprinted with permission from the University of California, San Francisco, Mass Spectrometry Facility.
Mass Spectrometry
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ALTERNATE PROTOCOL
SEQUENCE ANALYSIS USING MALDI-CID Although MALDI-PSD combined with database searching (see Basic Protocol) has emerged as the primary approach for identifying gel-separated proteins, limitations exist. For example, due to the nature of the gating device, the resolution of a precursor ion selection cannot exceed ∼100 (M/∆M) without serious loss of sensitivity. This value would barely enable the distinction between a parent ion at m/z 2000 and its water-loss ion. More importantly, the performance of PSD tends to be unpredictably dependent on the amino acid composition of individual peptides. This may be avoided by relying on high-energy MALDI collision-induced dissociation (MALDI-CID), because the use of collision gas with current reflectron instruments usually increases the intensities of product ions in the low mass region only (mainly immonium ions; Stimson et al., 1997). Often complete sequence ion information is needed to determine the sequence of peptides de novo (particularly important if the protein of interest is not represented in any current databases) or in studies of the structure of posttranslational and xenobiotic modifications. In many cases, these problems may be addressed by high-energy MALDI-CID. High-energy MALDI-CID is often performed on hybrid instruments, such as a tandem double-focusing magnetic orthogonal-acceleration time-of-flight (TOF) mass spectrometer (Bateman et al., 1995; Medzihradszky et al., 1996, 1997). The use of a double–focusing sector mass analyzer (such as MS1, a mass analyzer for separating parent ions using appropriate electric fields, and in combination with magnetic fields) enables the selection of a monoisotopic precursor ion with mass up to m/z 2000. The ions selected by MS1 then collide with xenon gas to generate a variety of ion types, which are then accelerated into a linear TOF analyzer orthogonal to the original direction of travel. The sampling frequency for this mode of operation is 100%. The signal-to-noise ratio in a MALDI-CID spectrum is far superior to that of a PSD spectrum. More importantly, many ion types that are unique to the high-energy process (e.g., d, v, and w ions) can be observed in a high-energy MALDI-CID spectrum but not in a PSD spectrum (which is very similar to a low-energy CID process). These high-energy ions have considerable value in the de novo determination of complete and unambiguous amino acid sequence (Medzihradszky and Burlingame, 1994), such as discrimination of leucine and isoleucine residues (Biemann, 1990). Therefore, MALDI-CID holds the promise to characterize unknown proteins, particularly those not represented in any current database, and can more explicitly determine the sites of posttranslational or xenobiotic modifications on target proteins. For example, Figure 16.6.3 shows a MALDI-CID spectrum of a tryptic peptide of human serum albumin (HPYFYAPELLFFAK*R, m/z 2360.5) that is modified by benoxaprofen glucuronide (Qiu et al., 1998), as indicated by the asterisk in the sequence above. As benoxaprofen contains a chlorine atom, the 12C35Cl isobar for the precursor mass was selected by MS1 for CID analysis. In this spectrum, all C-terminal sequence ions (y ions) together with their corresponding high-energy ions (v and w ions) allow the unambiguous determination of both the sequence and the site of modification of this peptide. The 461-unit mass shift of all y ions except y1 indicates the benoxaprofen glucuronide modification on the lysine residue near the C terminus. The presence of w6 (m/z 1183.5) and w7 (m/z 1296.6) distinguishes leucines from isoleucines (for isoleucines, w6 and w7 should be at m/z 1197.5 and 1310.6, respectively). The N-terminal sequence ions (b ions), internal fragments, and fragments from the drug moiety also aid or confirm the final data interpretation.
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×30.00 b
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100 ×60.00 90 w7 80 w9 –(B+H2 O) w8 70 y9 y14 60 y7 v11 b y v13 13 14 50 40 y10 30 y11 v y6 12 y12 b13 20 10 0 1200 1300 1400 1500 1600 1700 1800 1900 2000 2100 2200 2300 Mass (m /z ) 1296.6
Relative abundance (%)
100 90 80 70 60 50 40 30 20 10 0
Figure 16.6.3 MALDI-CID spectrum of a tryptic peptide of human serum albumin (amino acids 146 to 160; HPYFYAPELLFFAK*R; m/z 2360.5). Asterisk indicates modification with benoxaprofen glucuronide.
COMMENTARY Background Information In MALDI-TOF-MS spectra obtained in a reflectron mode, most of the fragment ions (metastable) formed in the field-free region cannot be observed because they are not in energy focus. However, fragment ions with masses close to the mass value of the parent ion can still be observed, albeit with both lower resolution and mass accuracy. To focus these metastable fragment ions throughout the entire mass range, the reflectron voltage needs to be successively stepped down to record approximately nine to fourteen consecutive mass scale segments, depending on the mass of the parent ion. These partially overlapping mass segments are then mass resolved to an accuracy of ±0.3 Da, and can be calibrated and stitched together to form a full PSD mass spectrum. One may still find that fragment ion peaks with different masses recorded in one segment have different mass accuracy (sometimes the difference can be as high as 0.3 Da within a segment). Typically, ions with higher masses will have better
mass accuracy than those with lower masses because they are better focused by the reflectron. More recently introduced curved-field reflectrons (Cordero et al., 1995) focus all the ions into a small region near the exit of the reflectron. This new feature enables approximately ten times faster acquisition of the product ion spectrum, because it can simultaneously collect the entire PSD mass spectrum without changing the reflectron voltage.
Critical Parameters and Troubleshooting After submitting a search request, oftentimes no match is retrieved for a given set of input parameters, and the MS-Tag program suggests decreasing the specificity of the input parameters. Because there are many restrictions that can be changed, it may be difficult for a new user to pinpoint the critical parameters that prevent the search from retrieving hits. In this situation, the following troubleshooting suggestions may save time.
Mass Spectrometry
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1. Verify that the mass calibration of the data is correct. Mass accuracy of the parent ion is very important for minimizing false positives. 2. Do not specify the species in the first search, because some species may have different names in different databases. Specify species only when too many hits have been identified. 3. Do not limit molecular weight range. 4. Provide a broad enough mass tolerance window for fragment ions. Fragment ions are differentially focused by the reflectron, so their mass accuracy varies significantly. 5. Deselect enzyme specificity. Nonspecific enzymatic cleavages often occur, particularly after overnight incubations. 6. Increase the number of ions allowed to be unmatched when no hit is found. 7. Deselect immonium ions. 8. Use the NCBInr database as a first choice. If nothing is found, search other databases. If none of these changes results in search hits, switch to homology searching. In homology search mode, it is very important to maintain the maximum specificity of input data, or too many false positives may be generated. Additionally, only peptides with one amino acid difference from a peptide sequence existing in the database can be retrieved. If two mutations occur in your peptide, no hit will be found. For peptides containing oxidized methionine, homology mode must be used. In this case, the parent mass shift can be limited to ±17 Da.
database for protein identification. Figure 16.6.1 shows the PSD spectrum of a tryptic peptide of an unknown protein isolated from mouse liver. An MS-Tag search of the NCBInr database in homology mode yields one single peptide sequence (DVFHMVVEVPR) with a single amino acid mismatch, compared to the sequence of a tryptic peptide from bovine inorganic pyrophosphatase (Fig. 16.6.2).
Anticipated Results
Kaufmann, R., Kirsch, D., and Spengler, B. 1994. Sequencing of peptides in a time-of-flight mass spectrometer—evaluation of post-source decay following matrix-assisted laser desorption ionisation (MALDI). Int. J. Mass Spectrom. Ion Process. 355-385.
PSD spectra are usually acquired at a laser power well above the threshhold required for observation of ions corresponding to the intact analytes (Riahi et al., 1994). Whether this enhancement of internal (vibronic) energy is due to the desorption of parent ions with deposition of higher internal energy or to the increased collision number resulting from a larger matrix plume volume is currently not known. The particular results obtained during a PSD experiment are also dependent on the properties of the particular matrices used (see MALDI sample preparation, UNIT 16.3; Stimson et al., 1997). For PSD analysis of peptides, α-cyano4-hydroxy-trans-cinnamic acid (HCCA) is the matrix of choice because it seems to promote ion fragmentation more than other matrices. The following example illustrates the use of PSD combined with MS-Tag searching of the
Time Considerations Search times depend on the database selected and on the quality of data submitted (e.g., the amount of tryptic peptides, the mass accuracy). They can vary from less than 1 min to 15 min or more.
Literature Cited Bateman, R.H., Green, M.R., Scott, G., and Clayton, E. 1995. A combined magnetic sector time-offlight mass spectrometer for structural determination studies by tandem mass spectrometry. Rapid Commun. Mass Spectrom. 9:1227-1233. Biemann, K. 1990. Sequencing of peptides by tandem mass spectrometry and high-energy collision-induced dissociation. Methods Enzymol. 193:455-479. Cordero, M.M., Cornish, T.J., Cotter, R.J., and Lys, I.A. 1995. Sequencing peptides without scanning the reflectron—post source decay with a curved field reflectron time-of-flight mass spectrometer. Rapid Commun. Mass Spectrom. 9:1356-1361. Kaufmann, R., Spengler, B., and Lützenkirchen, F. 1993. Mass spectrometric sequencing of linear peptides by product-ion analysis in a reflectron time-of-flight mass spectrometer using matrixassisted laser desorption ionization. Rapid Commun. Mass Spectrom. 7:902-910.
Medzihradszky, K.F. and Burlingame, A.L. 1994. The advantages and versatility of a high-energy collision-induced dissociation-based strategy for the sequence and structural determination of proteins. Methods Comp. Methods Enzymol. 6:284303. Medzihradszky, K.F., Adams, G.W., Burlingame, A.L., Green, M., and Bateman, R. 1996. Peptide sequence determination by matrix-assisted laser desorption ionization employing a tandem double focusing magnetic orthogonal acceleration time-of-flight mass spectrometer. J. Am. Soc. Mass Spectrom. 7:1-10. Medzihradszky, K.F., Maltby, D.A., Qiu, Y., Yu, Z., and Burlingame, A.L. 1997. Protein sequence and structural studies employing matrix-assisted laser desorption ionization—high energy colli-
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sion-induced dissociation. Int. J. Mass Spectrom. Ion Process. 160:357-369. Qiu, Y., Burlingame, A.L., and Benet, L.Z. 1998. Mechanisms for covalent binding of benoxaprofen glucuronide to human serum albumin—studies by tandem mass spectrometry. Drug Metab. Dispos. 26:246-256. Riahi, K., Bolbach, G., Brunot, A., Breton, F., Spiro, M., and Blais, J.C. 1994. Influence of laser focusing in matrix-assisted laser desorption ionization. Rapid Commun. Mass Spectrom. 8:242247. Stimson, E., Truong, O., Richter, W.J., Waterfield, M.D., and Burlingame, A.L. 1997. Enhancement of charge remote fragmentation in protonated peptides by high energy CID MALDI-TOF-MS using “cold” matrices. Int. J. Mass Spectrom. Ion Process. 169:231-240.
Internet Resources http://prospector.ucsf.edu University of California’s ProteinProspector site. http://donatello.ucsf.edu/onramp.html University of California site for programs employing fragment ion information to search databases (also see sites listed therein).
Contributed by C.R. Jiménez, L. Huang, Y. Qiu, and A.L. Burlingame University of California, San Francisco San Francisco, California
Mass Spectrometry
16.6.7 Current Protocols in Protein Science
Supplement 14
Enzymatic Approaches for Obtaining Amino Acid Sequence: On-Target Ladder Sequencing
UNIT 16.7
Peptide sequencing by mass spectrometry (MS) is usually based on detecting mass differences associated with various amino acids in the polymer chain. Post-source decay (PSD) and MS/MS spectra may yield internal peptide sequences. However, determination of the order of the first two, and sometimes the last few, amino acids in the peptide is often problematic without additional experiments. Several enzymatic approaches have proven useful for identifying the N- and C-terminal residues (Patterson et al., 1995; Thiede et al., 1995; Woods et al., 1995). They involve the use of carboxypeptidases and aminopeptidases to produce peptide ladders for rapid analysis by matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS). The ladder sequence method described in the Basic Protocol may generate sequence information from low- to subpicomole quantities of peptides and can be performed directly on the sample stage, thus minimizing potential sample losses to vials and through sample transfer. It should be pointed out that success is not predictable because of the substrate specificity of the enzymes. However, the experiments are easy to perform and the resulting mass spectra easy to interpret.
Materials Carboxypeptidase P, Y, and A 50 mM sodium citrate buffer, pH 6.0 Aminopeptidase M 50 mM ammonium carbonate buffer, pH 7.0 ∼1 pmol/µl peptide substrate in 0.1% (v/v) trifluoroacetic acid (TFA) Matrix solution (UNIT 16.3, Basic Protocol 1) 10% (v/v) formic acid, cold Matrix-assisted laser desorption/ionization (MALDI) mass spectrometer
BASIC PROTOCOL
1. Prepare C-terminal sequencing enzyme in sequencing buffer immediately before use: 0.1 to 0.2 µg/µl carboxypeptidase P in 50 mM sodium citrate buffer, pH 6.0 0.1 to 0.2 µg/µl carboxypeptidase Y in 50 mM sodium citrate buffer, pH 6.0 0.1 to 0.2 µg/µl carboxypeptidase A in 50 mM sodium citrate buffer, pH 6.0. Carboxypeptidases have different preferences for each amino acid, causing the kinetics of enzymatic release of different amino acids to vary considerably. Therefore, it is advisable to use different carboxypeptidases in parallel in a time-monitored fashion to release all amino acids in an interpretable manner. When the amount of sample is limited and C-terminal sequence information is needed, start with carboxypeptidase Y (as this enzyme can cleave all C-terminal residues) or use a mixture of all three enzymes.
2. Prepare N-terminal sequencing enzyme in sequencing buffer immediately before use: 2.0 µg/µl aminopeptidase M in 50 mM ammonium carbonate buffer, pH 7.0 Mass Spectrometry Contributed by C.R. Jiménez, L. Huang, Y. Qiu, and A.L. Burlingame Current Protocols in Protein Science (1999) 16.7.1-16.7.3 Copyright © 1999 by John Wiley & Sons, Inc.
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3. Apply 0.3 µl of ∼1 pmol/µl peptide substrate in 0.1% TFA to a clean sample target for each time point and enzyme to be used. Add an equal volume of sequencing enzyme to each. At relatively high peptide concentrations (≥1 pmol/ìl) the incubation may also be performed in a microcentrifuge tube from which aliquots are subsequently taken and applied to the target at different time points. For six time points and four enzymes, a total of twenty-four spots is needed.
4. Incubate sample target for 9, 15, 30, 60, 90, and 600 sec at room temperature. Lower concentrations of substrate generally require shorter incubation periods with enzymes to sequentially cleave the same number of residues from both the C- and N-terminal ends. The above time points are suitable for ∼1 pmol/ìl of peptide substrate. It is advisable to perform the digestion time course in a humidified chamber, as spots of the longer incubations may dry up.
5. To end the digestion, add 0.6 µl matrix solution to the digest mixture, mix by pipetting up and down, and allow to air dry. 6. Rinse the matrix/sample preparations with 10 µl cold 10% formic acid (UNIT 16.3, Basic Protocol 2, step 4). 7. Insert the sample target into the mass spectrometer and perform mass analysis according to the instrument manufacturer’s instructions (UNIT 16.3, Basic Protocol 1). COMMENTARY Background Information
On-Target Ladder Sequencing
Enzymatic sequencing is an alternative approach to chemical sequencing or fragmentation induced in the mass spectrometer. Carboxypeptidases offer a simple method by which amino acids can be sequentially cleaved from the C terminus of a protein or a peptide. Carboxypeptidase Y is a particularly attractive enzyme as it nonspecifically cleaves all residues from the C terminus, proline included. Aminopeptidase M is a metalloprotease completely hydrolyzing peptides and proteins with a free α amino group and L amino acids (X-Pro bonds are not cleaved). Enzymatic sequencing was historically performed by analysis of the released amino acids, which was complicated by amino acid contaminants and required relatively large amounts of sample. Using mass spectrometric techniques such as MALDI-MS, however, it is possible to perform direct mass analysis of the peptide fragments resulting from carboxypeptidase digestion. In this ladder sequencing approach, a sequence can be read, in the correct order, by calculating the differences in mass of adjacent peptide peaks. Enzymatic sequencing has been performed mostly in a time-dependent approach (see Basic Protocol), although concentration-dependent carboxypeptidase digestion has been successfully used for enzymatic sequencing as well (Patter-
son et al., 1995). Both digestion approaches can be conveniently performed on the MALDI sample target, minimizing the amounts of sample and enzyme consumed and losses of sample to vials.
Critical Parameters Carboxypeptidase digestions often need to be optimized to obtain the maximum sequence information in the shortest amount of time. For example, when the enzyme concentration is too high, sequence gaps may occur for adjacent amino acids that are readily cleavable. Lower concentrations of enzyme may allow for all amino acids to be sequenced, but often require long periods of time (up to days) for total digestion. Mass spectrometry cannot differentiate between leucine and isoleucine or between lysine and glutamine because they have identical molecular weight. Moreover, the confidence with which amino acid sequences are determined depends on the instrumental mass accuracy. For MALDI mass analysis without delayed extraction, depending on the amino acids, multiple measurements may be needed for reliable assignments. For example, more analyses are required for mass differences of 113 to 115 Da (Ile/Leu, Asn, and Asp) and 128 to 129 Da (Glu/Lys/Glu) than for mass differences around
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163 (Tyr) and 57 (Gly), when excluding all but one assignment as statistically unlikely. See Patterson et al. (1995) for statistical considerations.
Anticipated Results Using MALDI-MS analysis of on-target carboxypeptidase digestions, peptides of various amino acid composition, size, charge, and polarity have been successfully C-terminally sequenced at low picomole levels (Patterson et al., 1995; Thiede et al., 1995; Woods et al., 1995). Due to the salt content of the enzyme/buffer solution, prominent MNa+ ions may be detected in addition to MH+ ions when the carboxypeptidases are used at 0.2 µg/µl.
Time Considerations The amount of digestion time needed to obtain C-terminal or N-terminal sequence information of a peptide or protein may vary from a few minutes to many hours.
Literature Cited
Thiede, B., Wittman-Liebold, B., Bienert, M., and Krause, E. 1995. MALDI-MS for C-terminal sequence determination of peptides and proteins degraded by carboxypeptidase Y and P. FEBS Lett. 357:65-69. Woods, A.S., Huang, A.Y.C., Cotter, R.J., Pasternack, G.R., Pardoll, D.M., and Jaffee, E.M. 1995. Simplified high-sensitivity sequencing of a major histocompatibility complex class I-associated immunoreactive peptide using matrix-assisted laser desorption ionization mass spectrometry. Anal. Biochem. 226:15-25.
Key Reference Patterson et al., 1995. See above. This article provides an elaborate description of both time-dependent and concentration-dependent digestion strategies. In addition, it provides a protocol for statistical analyses of the resulting peptide mass ladder.
Contributed by C.R. Jiménez, L. Huang, Y. Qiu, and A.L. Burlingame University of California, San Francisco San Francisco, California
Patterson, D.H., Tarr, G.E., Regnier, F.E., and Martin, S.A. 1995. C-terminal ladder sequencing via matrix-assisted laser desorption ionization mass spectrometry coupled with carboxypeptidase Y time-dependent and concentration-dependent digestions. Anal. Chem. 67:3971-3978.
Mass Spectrometry
16.7.3 Current Protocols in Protein Science
Supplement 15
Introducing Samples Directly into Electrospray Ionization Mass Spectrometers Using a Nanospray Interface
UNIT 16.8
The development of electrospray (ES) ionization provided the first practical means of directly analyzing masses of peptide and protein samples in solution. As originally conceived, stable electrospray emission was most readily achieved at flow rates of 1 to 5 µl/min (Whitehouse et al., 1985). There was a successful early effort by mass spectrometer manufacturers to develop ES ion sources capable of stable emission at much higher flow rates in order to couple directly to existing liquid chromatography (LC) systems. However, to a first approximation, ES is a concentration-dependent phenomenon, and operation at high flow rates results in a corresponding high rate of sample consumption. Sensitivity increases are most readily achieved by decreasing the flow rate of sample solution into the electrospray source. In recent years, a number of methodologies have been developed to achieve ES analyses at sub-µl/min flow rates. These methods have made it possible to greatly expand the ways that mass spectrometry can be applied to analysis of complex biological systems. The large number of microscale ES devices currently in use make a comprehensive description of current practices impractical. This unit and UNIT 16.9 will describe in detail two micro-scale interfaces, one for direct liquid introduction without separation (i.e., nanospray; described here), and the second coupling to capillary liquid chromatography (cLC; UNIT 16.9). Both protocols assume that the mass spectrometer is equipped with a three-dimensional (x-y-z) translational stage for aligning the electrospray emitter with the orifice. The alignment is easier if a magnifying video camera is used to image the orifice and electrospray emitter. Such devices are available commercially (e.g., from Protana or New Objective). Detailed plans for modifying a Thermoquest LCQ ion trap mass spectrometer are available over the Internet (http://www.cityofhope.org/microseq/ download.html). NANOSPRAY ANALYSIS OF PEPTIDE AND PROTEIN SAMPLES This procedure describes the design of the nanospray interface and the method for loading and analyzing samples. It is suitable for samples that do not contain impurities such as buffer salts that suppress sample ionization. Procedures for removing such impurities on a small scale are described (see Commentary and Wilm and Mann, 1996; Jin et al., 1999).
BASIC PROTOCOL
Materials 50% (v/v) aqueous acetonitrile, 2% (v/v) acetic acid sample solvent Suitably prepared protein sample (free from buffer salts and strong acids) Nanospray needle holder, constructed from: Delrin metric webbed union (LC union; Upchurch) 1⁄ -in. o.d. × 0.020-in. i.d. Teflon tubing (Upchurch) 16 0.015-in. diameter platinum wire (Alfa Aesar) Delrin metric flangeless nut (Upchurch) Flangeless ferrule for 1⁄16-in. o.d. tubing (Upchurch) 1-mm o.d. × 0.75-mm i.d. × 2-cm long, tip i.d. 1 ± 0.5-µm pulled borosilicate glass needles (e.g., Protana, New Objective, or World Precision Instruments) Pipet and 0.25-mm o.d. gel loading pipette tips (Fisher) High-voltage clip Electrospray mass spectrometer Contributed by Terry D. Lee, Roger E. Moore, and Mary K. Young Current Protocols in Protein Science (2000) 16.8.1-16.8.5 Copyright © 2000 by John Wiley & Sons, Inc.
Mass Spectrometry
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1. Construct a nanospray needle holder device from an LC union, a length of platinum (Pt) wire, and a length of Teflon tubing (see Fig. 16.8.1). Enlarge the hole in the center web of the union to 1⁄16 in., so that the Teflon tube will pass completely through. Thread the Pt wire through the Teflon tube and hold in place by tightening the nut and ferrule. Be sure the wire extends beyond the Teflon tube ∼2.5 cm on one end and 1 cm on the other. The union and Teflon tube are used to support the wire electrode and provide a point of attachment to a clamp mounted on the x-y-z positioner. The wire in turn supports the nanospray needle. The other end of the wire is used to connect with the electrospray high voltage lead, typically with an alligator clip.
2. Clean the Pt wire of the mounting device with 50% aqueous acetonitrile, 2% acetic acid sample solvent and a laboratory tissue to remove any contamination from the previous sample. 3. Cut the pulled borosilicate glass needle to length so that when placed on the wire it does not reach to the Teflon tube. This ensures that there is good electrical contact as the sample solution is consumed during the analysis.
4. Take the suitably prepared protein sample solution (i.e., 1 to 10 µl) up in a gel loader pipet tip. Insert the tip into the electrospray needle until it encounters the narrow part of the taper. Expel the fluid slowly while withdrawing the pipet to reduce the probability that air will be trapped at the bottom of the column of fluid. Small amounts of air in the tapered section can be expelled by shaking the needle like a fever thermometer.
5. Slip the ES needle over the end of the Pt wire, mount the holder in the clamp on the x-y-z positioner, and attach the high-voltage clip. 6. Center the tip of the ES needle in front of the mass spectrometer opening ∼1 mm from the plane of the orifice. Turn on the ES voltage (i.e., 500 to 700 V) and optimize the position and voltage by monitoring the intensity of a sample ion in the real time display of the mass spectrometer. 7. After the analysis is complete, turn off the ES voltage, remove the clip, and remove the holder from the x-y-z positioner. Remove the ES needle from the Pt wire and discard it (proper disposal in a sharp object container is essential).
teflon tubing (1.6-mm-o.d. × 0.5-mm-i.d.)
0.015-in.-o.d. Pt wire
pulled glass needle (1-mm-o.d., 0.75-mm-i.d.)
1-µm tip 1/16-in. plastic union ESI-MS Using a Nanospray Interface
Figure 16.8.1 Nanospray needle and holder assembly.
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COMMENTARY Background Information The use of pulled-glass capillaries with micron-sized openings for electrospray emitters reported by Wilm and Mann (1994, 1996) was a seminal event in the evolution of microscale electrospray interfaces. The work showed that stable ion emission is possible at flow rates on the order of 20 nl/min and that the signal intensities of sample ions at a given concentration were similar to those obtained at higher flow rates. The methodology, which became known as “nanospray,” was simple and robust and is used routinely in most protein labs that have electrospray mass spectrometers. All nanospray setups have two things in common, a pulled glass or fused silica electrospray needle and a three-dimensional micropositioner. The electrospray needle must be positioned in close proximity to the orifice of the mass spectrometer for optimal performance. Although not essential, one or two video cameras equipped with macro-lenses make it easier to position the needle. Commercial setups for mounting and positioning nanospray needles are available from mass spectrometer manufacturers or third-party sources, or can be readily assembled from off the shelf components (http://www.cityofhope.org/immunology/ download.html). Most nanospray systems use needles constructed from borosilicate glass tubing (1.0- to 1.2-mm o.d. × 0.6- to 0.75-mm i.d.). A micropipet puller is used to heat the center of a length of tubing (8 to 15 cm) and pull it to separation. If the settings of the puller are correct, each piece will have a tapered tip with a hole ∼1 µm across. The technology for creating these needles originates from neuroscience studies (Rae and Levis, 1997) where they are used as microelectrodes that can be placed inside living cells. Consequently, micropipet pullers are commonly available in neuroscience departments. Pullers range in cost from <$2,000 for the simplest heated filament models to ∼$12,000 for one with a laser heat source and multiple program capability. Heated filaments are sufficient for pulling glass capillaries, but a CO2 laser source is needed for fused silica tubing. Nanospray needles are available commercially at a cost of several dollars each. Needle tips are fragile and should be handled very carefully. They should be stored in a covered container, otherwise they collect dust and moisture, inhibiting proper function.
Electrical contact for the spray potential can be made through the bulk liquid or directly at the tip via a metal coating on the outside of the glass capillary. Commercial emitter needles generally come with a sputtered metal coating on one side. If electrical contact is made via the outside of the needle, it is necessary to pressurize the fluid to ensure flow to the end of the tip and contact with the metal surface. Once the spray has started, flow will generally continue in the absence of any applied pressure. If the potential is applied to the bulk fluid, surface tension forces and the field gradient are sufficient to induce flow. The simple device described in the protocol will have a single set flow rate (typically 20 to 100 nl/min) defined by the viscosity of the fluid, the dimension of the needle orifice, and the voltage applied. The spray parameters can be controlled more precisely using a more complex device that continuously feeds sample solution to the emitter via a transfer line instead of placing the solution directly in the needle (Geromanos et al., 2000). The sample reservoir is pressurized to push liquid through the transfer line to the needle. Flow rates in the range of 1 to 500 nl/min can be achieved, depending on the tip i.d. and the pressure. Needles are constructed from 75-µm fused silica capillary tubing in order to reduce the dead volume. Even so, an hour or more may be needed to push the sample solution to the tip at the lower flow rates. Such precise control of the experimental parameters is unnecessary for most applications, but may be essential to obtain the most information from very small sample amounts. At the lowest flow rates, a few microliters of sample can be analyzed for several hours if necessary.
Critical Parameters The most common problem with respect to obtaining good results using nanospray is sample purity. Electrospray is very sensitive to impurities, particularly salts that interfere with sample ionization. These can be effectively removed by trapping peptides or proteins on a reversed-phase support, washing away the very polar components with water, and then eluting the sample with a solution containing a high percentage of organic solvent. Techniques for doing this using a few microliters of solution have been published (Wilm and Mann, 1996; Jin et al., 1999). Sensitivity to impurities also makes it imperative to use the highest available grade of solvents and reagents.
Mass Spectrometry
16.8.3 Current Protocols in Protein Science
Supplement 22
ESI-MS Using a Nanospray Interface
Final optimization of the voltage and position of the electrospray needle should be done by monitoring a sample ion. If the applied potential is too low, there is no electrospray and no signal will be observed. If the potential is too high, the spray is unstable, resulting in intermittent ion formation and an unstable signal on the monitor. Because of the close proximity of the needle to the grounded opening of the mass spectrometer, it is possible to get an arc between the two if the voltage is too high (i.e., >1500 V). Such an arc may damage the electrospray needle tip. There is generally a range of acceptable positions and voltages and very little time (<2 min) is needed to adjust these parameters. Strongly acidic solvents such as those containing trifluoroacetic acid (TFA) will extract Na and K ions from borosilicate glass, resulting in high levels of adducts of these ions with sample molecules. The result is a significant increase in the complexity of the spectrum and a decrease in sensitivity. The ion-pairing effect of TFA also tends to lower signal intensity. Weak organic acids, such as acetic and formic acids, do not present these problems. Although the analysis of 100% aqueous solutions is possible using nanospray, loading the needle is more difficult. The higher surface tension of water makes it more prone to trap air at the bottom of the needle, and air bubbles are more difficult to displace. Needles with an internal filament greatly reduce the risk of trapping air at the needle tip.
Samples left for prolonged periods in electrospray needles will slowly evaporate, leaving a precipitate in the tip that will block the opening. Highly concentrated samples may exhibit similar clogging problems after spraying for a few moments; sudden cessation of spray after a brief period of strong signal is almost always an indication that the sample is too concentrated. It is not practical to clean ES needles between use, and a new one should be used for each sample. Systems that deliver flow to a nanospray needle via a transfer line experience decreased flow over time as a result of particles that collect at the tip. Simply increasing the pressure to overcome the increased resistance to flow only seems to pack the particles more tightly into the tip, completely blocking the opening. It is usually better to readjust the voltage and position to accommodate the lower flow rate. For the protocol described here, trouble with particles clogging the needle are rarely encountered, but would be handled the same way.
Troubleshooting
Time Considerations
With pressurized setups, fluid can be observed to collect on the needle if no voltage is applied. The fluid should disappear when the voltage is applied. If this is observed, then ions are being formed. With the nanospray protocol given here, there are no visual clues to indicate flow. If no ions are observed over the normal range of applied voltage, then it is likely that the tip is either clogged or was completely closed during the pulling operation. It is handy to have a low-power microscope to examine the tip to see if fluid completely fills the tapered section. A break in the fluid that cannot be expelled by gentle shaking of the ES needle is an indication that the opening is closed. Closed tips can be opened by gently touching the tip to a hard surface. However, the fractured tip will now have a much larger opening and a correspondingly higher flow rate. Unless increased sample consumption is not an issue, it is better to transfer the solution to a new needle.
Assembly of the nanospray holder requires 10 to 15 min. Once assembled, all that needs to be done is to clean the Pt wire between samples. Loading and mounting the sample and optimizing parameters require only a few minutes each.
Anticipated Results Ion signal should be observed in the mass spectrometer the moment the voltage is turned on, if set in the proper range. Sample consumption is very low (i.e., 20 to 100 nl/min), so analysis times can be extended more than an hour if needed. Some sample does adhere to the Pt wire electrode and sides of the ES needle; therefore it is not possible to completely consume the full amount loaded.
Literature Cited Geromanos, S., Freckleton, G., and Tempst, P. 2000. Tuning of an electrospray ionization source for maximum peptide-ion transmission into a mass spectrometer. Anal. Chem. 72:777-790. Jin, X., Chen, Y., Lubman, D.M., Misek, D., and Hanash, S.M. 1999. Capillary electrophoresis/tandem mass spectrometry for analysis of proteins from two-dimensional sodium dodecyl sulfate polyacrylamide gel electrophoresis. Rapid Commun. Mass Spectrom. 13:2327-2334. Rae, J.L. and Levis, R.A. 1997. Fabrication of patch pipets. In Current Protocols in Neuroscience (J.N. Crawley, C.E. Gerfen, M.A. Rogawski, D.R. Sibley, P. Skolnick, and S. Wray, eds.) pp. 6.3.31. John Wiley & Sons, New York.
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Whitehouse, C.M., Dreyer, R.N., Yamashita, M., and Fenn, J.B. 1985. Electrospray interface for liquid chromatographs and mass spectrometers. Anal. Chem. 57:675-679. Wilm, M.S. and Mann, M. 1994. Electrospray and Taylor-Cone theory, Dole’s beam of macromolecules at last? Int. J. Mass Spectrom. Ion Proc. 136:167-180. Wilm, M. and Mann, M. 1996. Analytical properties of the nanoelectrospray ion source. Anal Chem. 68:1-8.
Internet Resources http://www.cityofhope.org/microseq/download. html Web site for downloading information on custombuilt electrospray interfaces and low-flow-rate gradient pumping systems.
Contributed by Terry D. Lee, Roger E. Moore, and Mary K. Young Beckman Research Institute of the City of Hope Duarte, California
Mass Spectrometry
16.8.5 Current Protocols in Protein Science
Supplement 22
Introducing Samples Directly into Electrospray Ionization Mass Spectrometers Using Microscale Capillary Liquid Chromatography
UNIT 16.9
This unit describes the design and operation of a microscale electrospray (ES) interface suitable for the on-line liquid chromatography (LC) separation and mass spectrometry (MS) analysis of mixtures of peptides and proteins. The interface utilizes an ES needle packed with reversed-phase support. Such a design has the advantage of minimizing any void volume between the end of the column and point of electrospray ionization, thus maintaining the integrity of the LC separation and maximizing sensitivity. Recently, electrospray needles packed with supports have become commercially available. Consequently, researchers have the option of building everything from scratch (see Basic Protocol 1), buying fritted needles that can be packed (see Basic Protocol 2), or buying needles already packed with a variety of supports. The protocols are divided into sections to adequately cover all of these options. The protocols assume access to a solvent delivery system capable of delivering gradients at flow rates <200 nl/min. Various options for low-flow solvent delivery systems are also discussed (see Commentary and Chervet et al., 1992, 1996; Davis et al., 1995; Ducret et al., 1998). BUILDING AN INTEGRATED LC COLUMN ES NEEDLE This protocol describes building an empty column ES needle assembly from scratch.
BASIC PROTOCOL 1
Materials 360-µm-o.d. × 150-µm-i.d. fused silica capillary (ES needle; Polymicro Technology) Laser micopipet puller (e.g., Sutter Instruments) 140-µm-o.d. × 25-µm-i.d. fused silica capillary (Polymicro Technology) Tool for cleaving fused silica capillaries Low-power dissecting microscope GF/A filter paper (Whatman) 1. Pull a fused silica needle with a 5-µm orifice from a 15-cm length of 360-µm-o.d. × 150-µm-i.d. fused silica capillary using a laser micropipet puller. Each pulling operation produces two needles. An effective program uses a two-stage approach to generate needles with a long tapered section that is easy to align with the mass spectrometer inlet. The first step of the program uses the following parameters: Heat 550, Filament 0, Velocity 30, Delay 200, Pull 0. For the second and succeeding step, the Heat is decreased to 500, the Velocity is decreased to 20, and the Filament is increased to 5.
2. To create the support for the filter, push a length of 140-µm-o.d. × 25-µm-i.d. fused silica capillary into the 150-µm-i.d. ES-emitter needle to the point where the taper begins. Withdraw the capillary until ∼5 mm is left in the needle. Cut the excess length of capillary and push the cut piece back down into the tapered end. Use a low-power dissecting microscope to view the process. 3. Place a sheet of GF/A glass fiber filter on a hard surface. Press the large end of the column into the filter to cut out a circular piece. Push the circularly cut filter into the column using a length of 140-µm-o.d. capillary. Repeat the process two more times. Mass Spectrometry Contributed by Terry D. Lee, Roger E. Moore, and Mary K. Young Current Protocols in Protein Science (2000) 16.9.1-16.9.7 Copyright © 2000 by John Wiley & Sons, Inc.
16.9.1 Supplement 22
fused silica capillary (360-µm-o.d. × 150-µm i.d.)
C18 column packing
fused silica capillary (140-µm-o.d. × 25-µm-i.d.)
GF/A filter
5-µm orifice
Figure 16.9.1 Column/ES needle assembly.
4. Use the 140-µm-o.d. capillary to compress the three pieces of filter together so that they appear to be a single piece under the microscope. See Figure 16.9.1 for a diagram of the finished product. BASIC PROTOCOL 2
PACKING THE ES NEEDLE This protocol describes how to pack the column ES needle assembly. Materials 5- to 10-µm particle size silica-based reversed-phase chromatography medium Acetonitrile Pulled fused silica capillary needle with frit (column/ES needle, see Basic Protocol 1) Empty columns with fritted tips, 360-µm-o.d. × 75-µm-i.d., 8-µm orifice (optional; can be obtained commercially from New Objective) 1⁄ - × 1⁄ -in. stainless steel reducing union 16 32 1⁄ -in. to 0.4-mm graphite reducing ferrule 32 1⁄ -in. gauge plug 16 Packing reservoir: 1⁄16-in.-o.d. × 0.02-in.-i.d. × 10-cm long stainless steel tubing with a compression screw and metal ferrule permanently attached to each end 1⁄ -in. stainless steel union 16 1-ml disposable syringe and 16-G blunt-ended needle Compression screws 1.5-ml microcentrifuge tubes High pressure pump 1. Attach the ES needle to the small end of a 1⁄16- × 1⁄32-in. stainless steel reducing union using a 1⁄32-in. to 0.4-mm graphite reducing ferrule. Use a 1⁄16-in. gauge plug to ensure that the needle does not extend too far past the central web of the union. 2. Attach the packing reservoir to a 1⁄16-in. stainless steel union. Attach a 1-ml syringe equipped with a 16-G blunt-ended needle and compression screw to the other side of the union. 3. Prepare a 1:10 slurry of 5- to 10-µm particle size silica-based reversed-phase packing:acetonitrile in a 1.5-ml microcentrifuge tube. Agitate the slurry vigorously to ensure an even suspension. Draw the packing slurry into the reservoir using the 1-ml syringe (Figure 16.9.2A).
ESI-MS Using Microscale Capillary Liquid Chromatography
4. Attach the reducing union with the needle to the open end of the packing reservoir. Be sure to clean the fitting of any adhering slurry, which may cause the fitting to bind.
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Remove the syringe and fitting from the union on the other end of the reservoir and attach the reservoir to the high-pressure pump at this end (Figure 16.9.2B). 5. Ramp the pump pressure up to 4000 psi over a period of 1 min while tapping gently on the side of the reservoir with a wrench or similar tool to ensure even flow of the slurry. After reaching 4000 psi, turn off the pump and allow the pressure to bleed off slowly through the column. After an initial period of gentle flow into the column, the slurry will often become compressed at the bottom of the reservoir and stop flowing until a higher pressure is reached, at which point it flows down in a mass.
6. Remove the reservoir from the pump and reattach the syringe. Remove the reducing union and needle from the reservoir. Remove the needle and store it until use. Expel any unused slurry from the packing reservoir back into the centrifuge tube for reuse. Thoroughly clean all fittings before reassembling or storing the apparatus.
A
B to pump
syringe
reservoir
reservoir
ES needle
Figure 16.9.2 Packing the column/ES needle assembly. (A) Draw the packing slurry into the reservoir using the 1-ml syringe. (B) Remove the syringe and fitting from the union on the other end of the reservoir and attach the reservoir to the high-pressure pump at this end.
Mass Spectrometry
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BASIC PROTOCOL 3
MOUNTING AND USING THE MICROSCALE ES LC/MS INTERFACE ASSEMBLY This protocol describes the assembly and performance testing of the ES interface, consisting of a plastic HPLC micro Tee, a gold wire electrode, and a packed fused silica ES needle (Figure 16.9.3). Materials Buffer A Cytochrome c tryptic digest mixture test sample Liquid chromatography (LC) pump PEEK micro Tee (Upchurch) PEEK sleeve (Upchurch) Packed fused silica ES needle (see Basic Protocol 2; also available commercially from New Objective) 0.025-in.-diameter gold wire (Alfa Aesar) ES mass spectrometer and alligator clip 1. Connect the transfer line from the LC pump to the micro Tee using a PEEK sleeve to make up the difference between the diameter of the line and the ferrule hole size. Connect the ES needle the same way on the opposite arm. Connect the 0.025-in.-diameter gold wire electrode through the third (top) arm of the Tee and attach the ES voltage lead to it with an alligator clip. Mount the Tee directly to a clamp on the x-y-z positioner. 2. Equilibrate the column to initial conditions with the aqueous component of the gradient system (buffer A). Fluid will collect on the needle tip with the electrospray voltage off.
3. Center the tip in front of the MS orifice ∼1 mm back from the plane of the opening. 4. Increase the ES voltage until the threshold for achieving spray is crossed (i.e., typically 800 to 1200 V). Stable emission of normal background ions should be observed.
5. Run several short blank gradients through the system to remove contaminants and ensure that the column packing is thoroughly wetted.
gold wire (0.02-in.-diameter)
fused silica transfer line (360-µm-o.d. × 50-µm-i.d.)
ESI-MS Using Microscale Capillary Liquid Chromatography
PEEK sleeve (0.025-in.-o.d. × 0.015-in.-i.d.) ES needle/column assembly
PEEK micro tee
Figure 16.9.3 Microscale ES LC/MS interface assembly.
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6. Run a blank gradient through the system at the normal running flow rate (i.e., 150 to 200 nl/min) to determine levels of normal background ions and to be certain that the spray is stable over the course of the entire gradient. 7. Next, analyze the standard cytochrome c tryptic digest mixture to determine if the chromatographic resolution is adequate and that stable ion emission is achieved for all of the components. COMMENTARY Background Information An efficient microscale liquid chromatography (LC) system can serve three purposes. It can effectively remove impurities that interfere with sample ionization. It can concentrate samples, thus significantly increasing the lower limits of detection. Finally, it can separate the components of a complex mixture and present them one or a few at a time for analysis by the mass spectrometer. In theory, all the advantages of nanospray also apply to on-line LC separations. However, there are a number of additional technical issues that define the practical limits. For example, with nanospray a new emitter is typically used for each sample. With on-line LC, emitters are more difficult to make and integrate into the system and are often expected to last for days and even weeks. With nanospray, the solvent composition is constant for the entire analysis. With gradient elution, the parameters that affect spray stability are constantly changing. Although microscale LC/MS is certainly more difficult than nanospray, it is not significantly more difficult than normal-scale LC/MS. Systems and components are available commercially that are adequate for chromatographic separations at flow rates <200 nl/min. It is no longer necessary for researchers to build their own systems, although they may want to in order to reduce costs or adapt to particular needs or applications. Almost without exception, microscale (≤150-µm-i.d.) capillary columns are constructed from fused silica tubing. The interface between the column and the ES needle is critical. Any void volume will result in peak broadening, degrading both the chromatographic resolution and sensitivity. The borosilicate needles typically used for nanospray are not suitable for coupling to capillary chromatography because of the large inner and outer diameters. Packing the column in the ES needle makes it easy to eliminate any void volumes (Davis and Lee, 1998; Gatlin et al., 1998). Needles used for this purpose will have larger orifices than are typically used for nanospray needles in order to facilitate the column packing opera-
tion. Although needles with orifices as large as 25 µm can be used, smaller tip dimensions make it easier to maintain stable ion emission over the broad range of solvent compositions associated with gradient elutions. For the lowflow systems described here, emitters with tips <10-µm o.d. are recommended, and tips <5-µm o.d. are needed for special techniques such as peak parking where flow rates are similar to those obtained with nanospray. Tips with small holes are easily plugged, and special techniques are needed to avoid clogging during the packing operation. Needles with long narrow tapers can be successfully packed with polymeric supports such as the 10-µm Poros R2 media. Silica-based supports generally have a broader size distribution, and it is more difficult to make freely flowing columns without protecting the tip with some kind of filter. In the protocol given here, the filter is a glass fiber membrane supported by a small piece of fused silica capillary at the base of the needle. Once assembled, the structure is very robust and column plugging nearly always occurs at the inlet end, where it can be removed by trimming off that end. In many ways, microscale columns are more robust than conventional HPLC columns. They are essentially immune to thermal shocking from rapid changes in solvent composition, and they can be allowed to dry out completely and be rehydrated without special precautions. It is also possible to polymerize supports directly in the needle to create single monolithic structures (Moore et al., 1998). However, it is not convenient to make such structures one or a few at a time. Electrospray needles are available commercially that have preformed frits in the tip that very effectively eliminate any post column void volumes. The choice of using commercial columns or constructing them in-house is largely one of economics. If a laser puller is available, then the cost of building one’s own is much less even when the cost of labor is factored in. There is less reluctance to discard a column that has become contaminated or has marginal perform-
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ance. On the other hand, laser pullers are expensive ($12,000) and would not be justified unless hundreds of columns are needed or the puller is also used for other purposes. The delivery of gradients at flow rates <200 nl/min remains problematic. No commercial pumping systems exist that can do it efficiently by making the gradient in real time. The solution that most researchers have adopted is to use pumps operating at much higher flow rates, and then to split the flow, sending the large majority to waste. This is a very workable solution although it is important to be aware of the problems that can occur during split flow operation. Any change in the resistance of the LC column and ES interface will affect the split ratio and may have a serious effect on the separation. This can be overcome to a certain extent by designing the system such that most of the resistance to flow is within the splitting device (Chervet et al., 1992, 1996). Alternative solvent delivery systems that do not use split flows have been described in the literature (Davis et al., 1995; Ducret et al., 1998). The gradient-loop HPLC described by Davis and co-workers utilizes preformed gradients and has proven very effective for efficient smallscale separations coupled to mass spectrometry. A detailed parts list and assembly instructions for this system can be downloaded from the Internet (http://www.cityofhope.org/microseq/ download.html).
Critical Parameters
ESI-MS Using Microscale Capillary Liquid Chromatography
When assembling the electrospray emitter with an internal filter, it is essential to work in a clean environment. Dirt trapped on the wrong side of the filter can lead to immediate column failure if it becomes wedged in the opening during the packing operation. It is often possible to unplug the tip of a plugged column assembly by etching it with hydrofluoric acid (HF), but careful precautions must be taken when working with HF because it is highly toxic and causes insidious burns. Once made, columns should be stored in a covered container. If used and then stored, it is important to thoroughly flush the needle to prevent clogging due to formation of precipitates in the tip. The same procedure can be used to pack polymeric supports. However, the maximum pressure is set at 1000 instead of 4000 psi. Because of the lower pressure requirements, many experimenters have been able to use a pressure bomb to pack columns with polymeric supports, instead of a high-pressure pump.
Troubleshooting The most useful tool for determining the source of problems with a microscale ES LC/MS system is an imaging system that provides a magnified image of the ES needle tip as it is being used. With the pump on and the spray voltage off, there should be a steady flow of liquid from the tip as evidenced by a drop on the end of the needle that steadily increases in size. If there is no flow and the pump is on, there is an obstruction somewhere in the line, usually in the column. If there is intermittent flow and gas bubbles are observed at the needle tip or in the transfer line, this is an indication of a leak in some part of the plumbing or inadequately degassed HPLC buffers. Connections to fused silica capillaries can fail either because they are too loose or because the capillary has cracked due to over-tightening of the ferrule. With the ES voltage turned on, the drop on the tip should disappear unless it has flowed too far back on the needle. If fluid does not collect on the needle with the voltage on, the ES process is working and it is certain that ions are being formed. Failure to observe ions in the mass spectrometer under these conditions can only happen if the needle is grossly misaligned or the mass analyzer has failed or is out of tune. A number of possible problems can cause liquid to collect on the tip with the voltage applied. The tip may be broken, resulting in a larger orifice and higher spray threshold. The orifice to the mass spectrometer may not be properly grounded. This can happen as the result of a loose connection or from sample contamination on the surfaces surrounding the entrance into the mass spectrometer. Ion sources utilizing heated metal capillaries need to be cleaned at regular intervals, depending on use and the nature of the samples being analyzed. To minimize contamination of the interface, the ES needle should be moved back from the MS entrance and the ES voltage turned off when cleaning and equilibrating the column between runs and while loading sample onto the column. Material will drip off the end of the column rather than being sprayed onto the MS entrance. Finally, there may be a problem with the ES high voltage connection. By checking all of the possibilities in a logical fashion, the source of the problem can generally be quickly determined. It is also important to check for bubbles in the tip of the electrospray needle with the ES voltage on. Gas bubbles can be generated at the electrode as a result of electrolysis. The degree that electrolysis occurs is dependent on the
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current. There are two pathways to be considered. The first is the result of ions spraying from the end of the needle. The second is leakage from the electrode back through the transfer line to the grounded pump. The resistance of the transfer line is a function of the solution composition, the length and diameter of the transfer line. The transfer line should be at least half a meter long and have a diameter no larger than 50 µm. Many of the problems with microscale capillary HPLC are the same as those that plague normal-scale HPLC. Samples that contain detergents will have higher levels of spectral background that will limit sensitivity. Contamination of the sample injector is a common occurrence. Off-line loading of the column via a pressurized reservoir can eliminate this problem although at the cost of frequent making and breaking of the connection to the pumping system. For novice users, there is a tendency to overload the column, significantly altering the back pressure or compromising the separation efficiency. Problems with the chromatography are best diagnosed by analyzing a standard peptide mixture.
Anticipated Results The system described in this protocol should yield high quality mass spectral data for mixture components in quantities as low as 100 fmol. Below 100 fmol, peptide molecular ions become increasingly more difficult to detect within the context of the spectral background, and MS/MS spectra may be too weak to be useful. Even so, it is usually possible to make protein identifications using only a few tens of fmol.
Time Considerations Placing a frit in an ES needle requires a certain amount of dexterity and practice. An experienced assembler may place the frit in 5 to 10 min, but it will take several times longer for a novice. Making a packing reservoir takes only a few minutes, particularly if a precut length of stainless steel tubing is used. Packing a needle that already contains a frit takes at least a few minutes, and the time required for the pressure to bleed off can vary considerably
depending on the internal volume of the highpressure pump. Mounting the ES interface assembly and initiating spray takes only a few minutes.
Literature Cited Chervet, J.P., Meijvogel, C.J., Ursem, M., and Salzmann, J.P. 1992. Recent advances in capillary liquid chromatography. LC-GC 10:140-148. Chervet, J.P., Ursem, M., and Salzmann, J.B. 1996. Instrumental requirements for nanoscale liquid chromatography. Anal. Chem. 68:1507-1512. Davis, M.T. and Lee, T.D. 1998. Rapid protein identification using a microscale electrospray LC/MS system on an ion trap mass spectrometer. J. Am. Soc. Mass Spectrom. 9:194-201. Davis, M.T., Stahl, D.C., and Lee, T.D. 1995. Low flow high-performance liquid chromatography solvent delivery system designed for tandem capillary liquid chromatography mass spectrometry. J. Am. Soc. Mass Spectrom. 6:571-577. Ducret, A., Bartone, N., Haynes, P.A., Blanchard, A., and Aebersold, R. 1998. A simplified gradient solvent delivery system for capillary liquid chromatography electrospray ionization mass spectrometry. Anal. Biochem. 265:129-138. Gatlin, C.L., Kleemann, G.R., Hays, L.G., Link, A.J., and Yates, J.R., III. 1998. Protein identification at the low femtomole level from silver stained gels using a new fritless electrospray interface for liquid chromatography-microspray and nanospray mass spectrometry. Anal. Biochem. 263:93-101. Moore, R.E., Licklider, L., Schumann, D., and Lee, T.D. 1998. A micro-scale electrospray interface incorporating a monolithic, polystyrene-divinylbenzene support for on-line liquid chromatography-tandem mass spectrometry analysis of peptides and proteins. Anal. Chem. 70:4879-4884.
Internet Resources http://www.cityofhope.org/microseq/download. html Web site for downloading information on custom built electrospray interfaces and low flow rate gradient pumping systems.
Contributed by Terry D. Lee, Roger E. Moore, and Mary K. Young Beckman Research Institute of the City of Hope Duarte, California
Mass Spectrometry
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Protein Identification Using a Quadrupole Ion Trap Mass Spectrometer and SEQUEST Database Matching
UNIT 16.10
The constantly expanding wealth of protein sequence information contained in various publicly available databases has made the rapid identification of proteins a practical reality. Most methodologies rely on some form of mass spectrometry to generate the necessary sequence information on a sample. Ion trap mass spectrometers equipped with electrospray ion sources are relatively low-cost systems that yield high quality tandem MS (MS/MS) spectra on very low amounts of sample. Because they are readily coupled to liquid chromatography (LC) separations, they can be used to analyze very complex mixtures of peptides. SEQUEST is a powerful suite of programs that can take the information in the peptide MS/MS spectra and correlate it to a database of known DNA or protein sequences. COLLECTION OF PEPTIDE MS/MS SPECTRA AND ANALYSIS BY SEQUEST
BASIC PROTOCOL
This procedure describes the use of a Quadrupole Ion Trap Mass Spectrometer (QIT-MS) and the commercially available SEQUEST (Eng et al., 1994) program to identify proteins. The proteins are digested and the digests are analyzed by LC/MS/MS. The MS/MS spectra are then used to query a sequence database to identify the proteins from which the peptides are derived. SEQUEST is a highly automated approach to database searching suitable for high-throughput analysis, but requires considerable computer power. Materials Protein sample digested into peptides with an appropriate enzyme (UNITS 11.1-11.3) Capillary HPLC system (UNIT 16.9) Quadrupole Ion Trap Mass Spectrometer (QIT-MS) with up-to-date software (Thermoquest LCQ with Excaliber version 1.1) Fast computer for database searching (Microsoft Windows NT 4.0 with Service Pack 5 or higher) SEQUEST and SEQUEST Browser software (Version 26 or higher; Eng et al., 1994) Protein or nucleotide sequence database in FASTA format (also see UNIT 2.1) Additional reagents and equipment for desalting by gel filtration chromatography (UNIT 8.3) Gather data 1. Analyze protein digests using a capillary HPLC system (UNIT considerable advantage in sensitivity.
16.9),
which gives
For maximum sensitivity, thoroughly desalt the digest before analysis either by extended on-line column washing or an off-line technique such as employing a zip tip or hydrophilic interaction column (UNIT 8.3).
2. Carry out the analysis using automated data analysis on a QIT-MS. Use a “triple-play” style data analysis with dynamic exclusion. In “triple-play” analysis, the instrument gathers full mass range scans until the base peak exceeds a threshold, which automatically triggers a high-resolution, narrow mass range scan (“zoom scan”), and then a MS/MS scan of the peak of interest. Dynamic exclusion prevents reanalysis of already analyzed precursor ions. Contributed by Roger E. Moore, Mary K. Young, and Terry D. Lee Current Protocols in Protein Science (2000) 16.10.1-16.10.9 Copyright © 2000 by John Wiley & Sons, Inc.
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Prepare data for SEQUEST analysis 3. Prepare raw data for SEQUEST analysis by transferring the data file from the QIT-MS instrument computer to a separate fast database searching computer. It is possible to run SEQUEST on the same computer that is used to control the mass spectrometer, but the computational needs of SEQUEST make this inadvisable. SEQUEST does not read mass spectrometer data files directly. Instead, the data are translated into the intermediate .dta format, with each .dta file containing the data from a single MS/MS scan or a group of scans from the same precursor ion.
4. Use the Setup Directory and Create DTA routine in SEQUEST Browser to convert the raw data file into .dta files in a separate directory for analysis. Screen .dta files 5. Screen .dta files by using one of the automated screening procedures from SEQUEST Browser, either a simple ion count cutoff or IONQUEST (Yates et al., 1998), or another program, such as winnow.pl (Moore et al., 2000; available at http://www.cityofhope.org/microseq/download.html), to screen the spectra. If there are only a few spectra to screen, use the View DTA program to check spectral quality and the correctness of charge state assignments manually. The process of generating .dta files is imperfect. The automated generation programs can assign the wrong precursor ion charge state or incorrectly group spectra, and include many low-quality spectra that are not worth searching. The screening procedures described in this step should overcome these concerns.
Run SEQUEST 6. Perform the SEQUEST analysis. The Run SEQUEST routine in SEQUEST Browser automates the process of running SEQUEST and setting up parameters. Significant parameters include the database to search, enzyme used, precursor ion and fragment mass tolerances, amino acid modifications to consider, ion series expected, and parameters influencing the output format (see Critical Parameters for additional detail). Use the Inspector program to monitor the progress of searches in progress.
Work up the data 7. Work up the data. Use the SEQUEST Summary routine to group results for work-up and report generation. SEQUEST will always generate at least some matches provided that there are any possible peptides having a mass within the precursor ion tolerance, making it necessary to check the results to ensure that the matches generated are actually significant.
COMMENTARY Background Information
SEQUEST Database Matching
Protein identification utilizing endoproteolytic digestion followed by mass spectrometry is now a standard approach. It has all but replaced approaches based on chemical sequence determination, and appears to be generally accepted as an essential technique for proteomics. The first approach to protein database searching was based on determining accurate masses for proteolytic peptides (Henzel et al., 1993; James et al., 1993; Pappin et al., 1993). This peptide mass map approach de-
pends heavily on high mass accuracy, which is not characteristic of quadrupole and quadrupole ion trap mass spectrometers. Triple quadrupole and quadrupole ion trap mass spectrometers are capable of generating fragment ion MS/MS spectra which reveal structural information about peptides being analyzed. To take advantage of the data available from MS/MS spectra, the sequence tag approach to database searching was developed (Mann and Wilm, 1994). In the sequence tag method, a MS/MS spectrum is interpreted to give a partial
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sequence that is used to search the database along with the intact mass and the masses of the fragments on either side of the interpreted region. Like the sequence tag approach, SEQUEST uses the intact mass and fragment ion data from individual peptides to query the database, meaning that each peptide is analyzed independently. Unlike the sequence tag approach, SEQUEST does not attempt to interpret spectra directly. Instead, the actual fragmentation pattern is compared to predicted fragmentation patterns for all peptides in the database having the same molecular weight as the precursor peptide. SEQUEST thus uses the maximum amount of information available in the MS/MS spectra, requires little human intervention, and is very effective in extracting data from noisy spectra. The disadvantage of SEQUEST is that it uses much more computer time than other search programs.
Critical Parameters Equipment Capillary HPLC integration with mass spectrometry is discussed in more detail in UNIT 16.9. Be sure to use the most up-to-date versions of all relevant software (i.e., Mass Spectrometer, SEQUEST, and SEQUEST Browser) that are known to be free of serious bugs. The SEQUEST Browser package is not absolutely essential, but it provides a vastly improved interface for SEQUEST and its associated routines as well as a number of new programs for data manipulation and reporting. It is very important to have a fast computer for SEQUEST searching. A 500-MHz Pentium III processor with 128-MB RAM and a 4-GB hard drive is a practical minimum standard, and a faster processor will improve performance. The latest released version (version 26) of SEQUEST is not capable of using multiple processors directly, but it is possible to run multiple instances of SEQUEST on a multiprocessor computer without loss of speed. Procedure Gathering the data. Trypsin is the enzyme of choice for MS/MS identification of proteins because the presence of a basic amino acid at the carboxy terminus helps to give strong series for both b and y ions and hence the most sequence information. Use sequencing grade trypsin that has been reductively methylated to reduce autolysis, such as a modified trypsin (e.g., Promega). For extremely basic proteins it
may be preferable to use sequencing grade endoprotease Lys-C, which will generate fewer, larger peptides. Avoid using other proteases unless there is a strong reason not to use trypsin and Lys-C. Digests can undergo degradation on prolonged storage. Minimize the time between digestion and analysis and store digests at −20°C until they are analyzed. The two critical experimental choices with regard to the MS analysis are how to set up dynamic exclusion parameters and whether to gather zoom scans. There are four critical parameters for dynamic exclusion: exclusion width, exclusion duration, repeat count, and repeat duration. The exclusion width is simply the range on either side of the analyzed peak that will not be further analyzed. Set the exclusion width wide enough to include common isotopic peaks from a +1 parent ion, as lowerabundance isotopic peaks can trigger an automated scan. The exclusion duration is the time that a peak remains excluded from reanalysis. Set the duration to approximately the typical peak width. This maximizes the ability to analyze later isobaric (same mass) peptides. The repeat count and repeat duration define the process by which ions are placed on the list of excluded ions. Ions are excluded if they are analyzed the number of times set by the repeat count within the time specified by the repeat duration. A single scan is usually enough to identify a peptide, so a repeat count of 1 (which makes the repeat duration unimportant) is generally acceptable. Zoom scans should almost always be used. The only case where zoom scans should be avoided is with samples of overwhelming complexity, where every possible scan must be devoted to generating MS/MS spectra. This is only true for digests of very complex multiprotein mixtures. Preparing raw data for SEQUEST. Create DTA will attempt to group spectra from the same precursor ion into a single .dta file if possible. Three parameters control grouping of scans. The grouped scan parameter is the number of scans required before a .dta file will be generated, and should be set to 1. The precursor mass tolerance is the maximum mass difference between two ions that will still allow them to be grouped; a value of 1 to 2 works well. The final parameter is the intermediate scan count, which is the maximum number of unrelated MS/MS scans that can fall between two related scans and still have them grouped. The value chosen should be small enough that scans from isobaric ions are not grouped. This value can be obtained by dividing the exclusion duration
Mass Spectrometry
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SEQUEST Database Matching
by three times the average scan time (since only one scan in three is a MS/MS scan). When setting the intermediate scan count, it is safer to be too low than too high, because incorrectly grouped scans present a much greater problem than incorrectly separated ones. The user can also specify a range of scans and a range of allowed precursor masses. Very low and very high molecular weight peptides rarely give strong matches by SEQUEST, so it is safe to consider only peptides in the range of 700 to 3500 Da. The program will also assign a charge state for each scan by (in order of preference) using a user-supplied charge state, using a charge state obtained from a zoom scan, or generating two .dta files, one assuming +2 and one assuming +3 precursor charge. The zoom scan–derived charge state is usually the best choice, which is one reason that zoom scans are so desirable. Screening .dta files. Due to imperfections in the .dta generation process, screening the files before searching is necessary. A very simple approach, which is actually used during .dta file generation, is to eliminate all spectra with fewer than a specified number of ions. Very few spectra with <35 to 65 ions will give a strong SEQUEST match. Eliminating all such spectra can save 1⁄4 to 1⁄3 of the total search time. More sophisticated routines, such as IONQUEST and winnow.pl, can have a larger impact. IONQUEST automatically searches for and eliminates spectra that match common contaminants, such as trypsin and keratin. Winnow.pl uses a more sophisticated set of heuristics than ion count to eliminate spectra that are unlikely to generate positive search results. Both IONQUEST and winnow.pl can run in much less than the time needed to search a single spectrum against the National Center for Biotechnology Information (NCBI) nonredundant database, while eliminating large numbers of uninteresting spectra. Running SEQUEST. Choice of parameters for running SEQUEST can have a large impact of the quality of the search results and on the amount of time taken to generate them. Some important factors include: a. Database used: databases are available for download from the National Center for Biotechnology Information (ftp://ncbi.nlm. nih.gov/blast/db/). SEQUEST can use both protein and nucleotide databases; it can translate nucleotide databases in 1, 3, or 6 reading frames on the fly, although translating in more than one frame multiplies the effective size of the database and the search time. Index the
database using the Database Indexer routine in SEQUEST Browser to speed access to results in SEQUEST Summary. Search time scales roughly with the size of the database plus a small constant. This means that searching large databases takes much longer than searching smaller databases. This would seem to make it desirable to search against species-specific databases as much as possible. In practice it is best to search against the largest available database if enough processor time is available. Identifying a protein from the correct species from a large database can often increase confidence in the quality of the match, and searching a complete database will help to show any contaminants or minor components present in the sample. Using a large database rather than a species-specific one is also very helpful when the complete genome of the species in question is not available. It is sometimes possible to identify homologous proteins from related species when the analyzed protein is not in the database. b. Enzyme used: the SEQUEST algorithm starts by finding peptides that are within a specified tolerance of the known precursor ion mass by performing a theoretical digestion on all proteins in the database. At a minimum, the enzyme used for the theoretical digestion must cleave at all possible sites cut by the enzyme used in step 1. SEQUEST may match a few additional peptides if the theoretical digestion uses a less specific enzyme, such as including possible chymotryptic cleavage sites in addition to tryptic cleavages or allowing cleavage at all peptide bonds. Allowing more cleavage sites increases the time taken for the search. c. Amino acid modifications: SEQUEST allows the operator to specify both quantitative and nonquantitative amino acid modifications. Quantitative modifications do not greatly affect search time, and are well suited for cases, such as cysteine reduction and alkylation, where the protein has been deliberately modified. Nonquantitative modifications can be used for either of two cases: incidental chemical modification and site-specific biological modification. Incidental modifications, such as methionine oxidation, occur essentially randomly though the protein, and searching for them rarely results in identification of additional peptides. Instead some peptides are identified twice, once with and once without the modification. In the case of site-specific biological modifications, such as phosphorylation, the potentially modified amino acid is completely unmodified at most sequence positions and quantitatively modified at other positions.
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Figure 16.10.1 Sample SEQUEST Summary output. Numbers on the left side of the figure indicate different regions of the output that are described as follows: (1) Header that contains the page name and hyperlinks to the other SEQUEST Browser programs. (2) Sample data file information and parameters used for the search. Db: Sequence database used for the search. Mass: indicates whether mass values given are monoisotopic or average. Dir: Directory containing the datafile. Enz: indicates what enzyme parameters were used to limit the search. Tot: indicates number of .dta files searched | number of spectra searched (may be smaller than number of .dta files because of charge state ambiguity) | summed zoom scan base peak intensity. (3) Set of control buttons that can be used to alter the format of the Main Results section and access further levels of information. A brief description of each button is included in Figure 16.10.3. (4) Main Results section that contains information for each consensus protein match. The first two lines of an entry contain information about the protein and a summary of the search results that led to the match. A description of the format of these lines is given in Figure 16.10.2. Following the first two lines are the results for each peptide of the consensus protein match, given as a single line of text. The format of this line is described in Figure 16.10.3. Protein matches based on only one peptide are designated as — No Consensus. (5) Complete consensus protein list. The format is nearly the same as the Main Results section (see Figure 16.10.2).
Peptides containing modification sites can only be identified using nonquantitative modification. Avoid searching for site-specific modifications unless that modification is known to be present in the sample being analyzed. d. Mass tolerances: SEQUEST allows setting tolerances for both precursor and fragment ion masses. Only peptides that fall within the precursor ion tolerance of the measured precursor molecular weight are further compared to the actual spectrum. The precursor ion tolerance should be set at a value similar to the observed mass measurement error. The fragment ion tolerance is only used in the preliminary stage of SEQUEST scoring. Missing one
or a small number of fragment ions will not compromise the entire search, so the fragment ion tolerance can be set low, i.e., ≤1 mass unit, without bad results. e. Ion series: SEQUEST is able to look at all of the common peptide ion series. The low energy CID characteristic of quadrupole ion trap MS/MS generates predominantly b and y ions, so those are the only series that should be used. f. Output parameters: other available parameters affect the format of the SEQUEST output files. They include: the number of top matches shown, the number of protein descriptions shown, whether to show references for all
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Name
Value shown in figure 1 Meaning A Consensus letter Each consensus match is given a letter corresponding to its rank by consensus score. Protein A is the top scorer. Each letter in the main list is given a unique color. Protein reference owl||CYC_HORSE The protein reference from the searched database. Also a hyperlink to the protein sequence with matched peptides highlighted (sequence alignment). 48 Consensus score Score based on number of distinct peptides matched. Score is 10 points per best match, 8 points per second best match, etc. and 1 point per match below fifth best. 5|7.4e6 Spectra | zoom Number of spectra contributing to the consensus and their base peak summed zoom scan base peak intensities. {4,1,0,0,0,0} Peptides by rank Number of peptides rated as best match, second best, etc. Each distinct peptide sequence contributes only once, so a peptide that generates two best matches and one third best match counts as only as one best match. (1 3 7 8, 4, x, The scans that contributed to this consensus match, grouped Matched .dtas x, x, x) by commas into best matches, second best matches, etc. If some of the scans have been displayed for a higher scoring protein, they will not be shown again for this protein. If all of these scans have been shown for other proteins, this protein will be displayed only in the complete consensus list at the end of the summary. Protein description CYTOCHROME C. - Description of the protein from the database. EQUUS CABALLUS (HORSE) Additional proteins owl||CYC_EQUAS If several proteins contain all of the same matched peptides, only one is shown in the main summary. The others are shown, without descriptions, in the complete consensus list. Also a hyperlink to a sequence alignment. C owl||CYC_BOVIN If all peptides for a protein are also in higher scoring Lower matches proteins, it is shown only in the complete consensus list. The letter for such proteins is the same color as the higher scoring protein that contained the same peptides. Also a hyperlink to a sequence alignment.
Figure 16.10.2 Description of SEQUEST Summary output for consensus protein matches. Consensus matched proteins are ones identified on the basis of at least two distinct peptide sequence matches.
SEQUEST Database Matching
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Name Scan
Sort Button
Value shown 1
zBP
2.3e6
Scan range
0003
Parent charge
2
Mass error
0.7
Mass
1495.0
Xcorr
4.04
deltaCn
0.20
Sp
2124
RSp
3
Ion matches
19/22
DB reference
Extra proteins
Preceding AA Peptide
Explanation Input files are numbered consecutively by scan number or scan range. The number for scans that contribute to a consensus match is colored the same as the Consensus letter. Base peak intensity for precursor ion zoom scan. Bolded if greater than the median value. Scan or scan range from data file that contributed to this .dta file. Hyperlink to actual Sequest output file. Charge state of precursor ion. Difference between predicted and measured mass for matched peptide. Measured precursor ion mass as monoprotonated ion. Cross Correlation score. Best measure of match between actual and predicted spectrum. Bolded if above a threshold. Delta Correlation. Difference between best and second best Xcorr as fraction of best Xcorr. Displayed as ---for matches below first place. Bolded if above a threshold. Preliminary Score. Score from preliminary matching algorithm that pares down list of peptides before cross correlation scoring. Bolded if above a threshold. Rank by preliminary score. Bolded if 1 or 2.
Number of ions matched / predicted for this sequence. Hyperlink to the spectrum with matched ions marked. owl||cyc_horse Database reference to the protein that the matched peptide is from. Hyperlink to display of protein sequence with peptide highlighted (sequence alignment). +1 Number of additional proteins also containing this peptide. Hyperlink to display of all proteins containing the peptide with sequence alignments. (K) Amino acid amino-terminal to matched peptide. EETLMEYLENPK Actual peptide sequence matched. Hyperlink to NCBI BLAST search using this sequence. Sort button only. When sorted by consensus, peptides that contribute to a consensus match are grouped under a heading for the consensus protein. Sort button only. Causes a reconstructed chromatogram to be displayed, with scans contributing to consensus matches labeled and colored, instead of the normal main section. Control button only. Causes additional controls for data manipulation and report generation to be shown. Control button only. Causes display of second or lower place matches in place of top match if they are relevant to a consensus protein.
Figure 16.10.3 Description of individual match lines and display controls in SEQUEST Summary. Each match line summarizes the results from searching a single .dta file. Unless otherwise noted, sort buttons cause SEQUEST Summary to display the lines sorted in the order of the value referred to by that button.
Mass Spectrometry
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proteins containing a matched peptide or only one, and whether to show fragment information. The number of matches and descriptions shown is largely a matter of personal preference. Matches below the top five are almost never significant for protein identification. It is very important to know every protein that contains a peptide of interest, so duplicate protein references should always be shown. Fragment information is available using SEQUEST Summary and need not be shown. Working up data. SEQUEST Summary automatically correlates and summarizes the results from a SEQUEST search (Figure 16.10.1), but it is still necessary to validate the individual matches in the summary. Different authors have suggested different approaches to validating sequence matches based on numerical criteria for match quality (Ducret et al., 1998; Link et al., 1999). SEQUEST Summary also makes it simple to compare actual and predicted spectra manually and validate only those matches that are subjectively judged to be good. Protein identification is based on matches of individual peptides. A common approach is to consider a protein identified if more than one peptide from the protein is validated by numerical criteria or a single peptide is validated manually. A more conservative approach does not allow identifications based on a single peptide match. The format of the output from SEQUEST Summary is rather cryptic and does not come with a good explanation. As an aid to the reader, a typical output from SEQUEST Summary is shown in Figure 16.10.1. A brief description of each part is given in the figure legend. Figure 16.10.2 describes the format of the two line description of each consensus protein match and Figure 16.10.3 describes the format of the single line description of each matched peptide spectrum.
Anticipated Results
SEQUEST Database Matching
The number and quality of sequence matches varies dramatically depending on the quantity of protein digested, the protein sequence, and the quality of the chromatographic separation. With abundant sample and by using a very long, low flow rate HPLC gradient, it is possible to identify dozens of proteins from a complex digest, or to obtain fifty or more peptide matches for a large protein. Even with comparatively low sample levels, such as the digest from a faint Coomassie stained gel band, it is quite possible to identify five or more peptides from a single protein routinely. Pro-
teins from light silver stained gel bands can be identified, but obtaining such results requires meticulous technique and success is not assured.
Troubleshooting A common cause of poor search results is low-quality mass spectrometry data. The HPLC and mass spectrometry methods should be carefully validated using standard digests before analyzing experimental samples. Check the quality of input data, either by manually spot-checking spectra or using a program such as winnow.pl, before modifying search parameters in an attempt to improve search results. If the spectra submitted to SEQUEST are of high quality and still do not produce positive matches, the peptides that generated the spectra are probably not being searched. Try a more general search that gives SEQUEST a larger range of peptides to match against the spectra. Steps to try, in rough order of preference are: 1. Search a larger database. A number of databases are available from the National Center for Biotechnology Information (ftp://ncbi.nlm.nih.gov/blast/db/). Use the NCBI non-redundant database if possible. 2. Expand the enzyme specificity for the search to include more possible cleavage sites or eliminate the enzyme restriction entirely. 3. Look for nonquantitative modifications of amino acids. 4. Examine spectra eliminated using programs such as IONQUEST and winnow.pl. 5. Expand mass tolerances. 6. Reanalyze the sample.
Time Considerations The time required for digestion and mass spectral analysis are described in more detail in sections devoted to those topics. Generating and automatically filtering .dta files takes a few minutes. Manually filtering .dta files requires ~10 to 20 sec per file. The computer time needed for the SEQUEST search proper is almost always the most time-consuming step of the process. SEQUEST can search a single .dta file against ~100,000 to 500,000 proteins/min, depending on the parameters chosen, when using the minimum standard computer system described above. Validating SEQUEST results using numerical criteria requires only a 1 or 2 sec per result and validating results manually is about as fast as manually filtering .dta files.
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Current Protocols in Protein Science
Literature Cited Ducret, A., Van Oostveen, I., Eng, J.K., Yates, J.R. III, and Aebersold, R. 1998. High throughput protein characterization by automated reversedphase chromatography/electrospray tandem mass spectrometry. Protein Sci. 7:706-719. Eng, J.K., Mccormack, A.L., and Yates, J.R. III. 1994. An approach to correlate tandem mass spectral data of peptides with amino acid sequences in a protein database. J. Am. Soc. Mass Spectrom. 5:976-989. Henzel, W.J., Billeci, T.M., Stults, J.T., Wong, S.C., Grimley, C., and Watanabe, C. 1993. Identifying proteins from two-dimensional gels by molecular mass searching of peptide fragments in protein sequence databases. Proc. Natl. Acad. Sci. USA 90:5011-5015. James, P., Quadroni, M., Carafoli, E., and Gonnet, G. 1993. Protein identification by mass profile fingerprinting. Biochem. Biophys. Res. Commun. 195:58-64. Link, A.J., Eng, J., Schieltz, D.M., Carmack, E., Mize, G.J., Morris, D.R., Garvik, B.M., and Yates, J.R. III. 1999. Direct analysis of protein complexes using mass spectrometry. Nat. Biotechnol. 17:676-682. Mann, M. and Wilm, M. 1994. Error-tolerant identification of peptides in sequence databases by peptide sequence tags. Anal. Chem. 66:43904399.
Moore, R.E., Young, M.K., and Lee, T.D. 2000. Method for screening peptide fragment ion mass spectra prior to database searching. J. Am. Soc. Mass Spectrom. 11:422-426. Pappin, D.J.C., Hojrup, P., and Bleasby, A.J. 1993. Rapid identification of proteins by peptide-mass fingerprinting. Curr. Biol. 3:327-332. Yates, J.R. III, Morgan, S.F., Gatlin, C.L., Griffin, P.R., and Eng, J.K. 1998. Method to compare collision-induced dissociation spectra of peptides: Potential for library searching and subtractive analysis. Anal. Chem. 70:3557-3565.
Internet Resources ftp://ncbi.nlm.nih.gov/blast/db/ Web site for downloading a number of databases available from the National Center for Biotechnology Information. http://www.cityofhope.org/microseq/download. html Web site for downloading the winnow.pl program to screen spectra prior to a SEQUEST search.
Contributed by Roger E. Moore, Mary K. Young, and Terry D. Lee Beckman Research Institute of the City of Hope Duarte, California
Mass Spectrometry
16.10.9 Current Protocols in Protein Science
Supplement 22
De Novo Peptide Sequencing via Manual Interpretation of MS/MS Spectra
UNIT 16.11
In a matter of a few seconds, a mass spectrometer is capable of ionizing a mixture of peptides with different sequences and measuring their respective mass-to-charge ratios. Under low-energy collisions in a tandem mass spectrometer (MS/MS; UNIT 16.1), such as a quadrupole ion trap (UNIT 16.10), a triple quadrupole (UNIT 16.1), or a quadrupole/time-offlight (UNIT 16.10), sequence information can be generated by selectively fragmenting each peptide along its amide backbone, and measuring the mass-to-charge ratios (m/z) of the fragment ions, thus producing a different MS/MS spectrum for peptides of varying amino acid composition and sequence. Because databases have been filling up rapidly with complete genome sequences, the recent trend in interpretation of MS/MS spectra of peptides has been database matching (UNITS 16.5 & 16.6). A caveat, however, to database matching is that it can only be successful if the database being searched contains the identical amino acid sequence, or one with high homology, to the peptide being investigated. Often times, high-quality MS/MS spectra are obtained for which there is no corresponding database match. This may be due to the fact that the peptide of interest is derived from a novel protein from an organism whose genome is not known, has amino acids in its sequence that may be post-translationally modified, or has multiple amino acid substitutions as compared to the homologous sequence from a different species. In these instances, de novo (or manual) interpretation of the MS/MS spectrum is required (see Basic Protocol). The results of the de novo analysis can be verified through chemical modification (see Support Protocols 1 and 2) and limited manual Edman degradation (see Support Protocol 3) of the peptide. MANUAL INTERPRETATION OF MS/MS SPECTRA The objective of a de novo sequencing exercise is to determine the sequence of a peptide from its MS/MS spectrum (UNIT 16.1), without the use of database matching programs (see Commentary; UNITS 16.5 & 16.6). Low-energy collision-induced dissociation (CID) of peptides generally results in a limited number of sequence-specific fragment ions (Fig. 16.11.1). Ions of type y include the C terminus of the peptide, while b-type ions include the N terminus of the peptide. Described here is a step-by-step explanation of manual interpretation of an MS/MS spectrum. The annotations to the steps provide a detailed example. The only items needed other than those listed in the materials section below are a table of molecular weights for the residue masses of the twenty most common amino acids (Table 16.11.1; also refer to APPENDIX 1A), pencil, paper, and a standard calculator.
b1 a1 H2N
CH R1
b3
b2 a2
O C
NH
CH
BASIC PROTOCOL
a3
O C
NH
R2
O
CH
C
R3
nH
O NH
CH
C
OH
R4
Figure 16.11.1 Generation of sequence-specific fragment ions. Mass Spectrometry Contributed by Terri Addona and Karl Clauser Current Protocols in Protein Science (2002) 16.11.1-16.11.19 Copyright © 2002 by John Wiley & Sons, Inc.
16.11.1 Supplement 27
Materials Peptides generated from a tryptic digest of a reduced and alkylated single protein or protein mixture Liquid chromatography (LC) apparatus coupled in-line to a microelectrospray ionization source on a tandem mass spectrometer (quadrupole ion trap, triple quadrupole, or hybrid quadrupole/time-of-flight) with <200 nl/ml flow rates and 75-µm-i.d. capillary reversed-phase columns Additional equipment and reagents for confirming sequence information via chemical modification (see Support Protocol 1 and 2) and manual Edman degradation (see Support Protocol 3) Generate MS/MS spectrum 1. For best results, analyze tryptic digests of a single protein or protein mixture by liquid chromatography (LC) apparatus coupled in-line to a microelectrospray ionization source on a tandem mass spectrometer with <200 nl/min flow rates and 75-µm-i.d. capillary reversed-phase columns. 2a. For quadrupole ion-trap mass spectrometers (UNIT 16.10): Acquire MS/MS spectra in automatic data-dependent mode whereby every MS experiment is followed by a user-defined number of MS/MS experiments (e.g., MS/MS of the top four most intense ions per MS scan). The number of MS/MS scans per MS scan is dependent upon the chromatographic peak width of the LC/MS/MS system. Dynamic exclusion should also be enabled to minimize the number of times a MS/MS spectrum is acquired on any given ion. Global collision energy can be used throughout the LC/MS/MS experiment and will still yield high quality, “interpretable” MS/MS spectra. For the analysis of single protein digests, the charge of Table 16.11.1
De Novo Peptide Sequencing via Manual Interpretation of MS/MS Spectra
Residue Masses of Amino Acidsa
Name
Symbol
Monoisotopic Average massb mass
Immonium ion mass
Glycine Alanine Serine Proline Valine Threonine Cysteine Leucine Isoleucine Asparagine Aspartic acid Glutamine Lysine Glutamic acid Methionine Histidine Phenylalanine Arginine Carbamidomethyl cysteine Tyrosine Tryptophan
Gly or G Ala or A Ser or S Pro or P Val or V Thr or T Cys or C Leu or L Ile or I Asn or N Asp or D Gln or Q Lys or K Glu or E Met or M His or H Phe or F Arg or R Camc Tyr or Y Trp or W
57.02 71.04 87.03 97.05 99.07 101.05 103.01 113.08 113.08 114.04 115.03 128.06 128.09 129.04 131.04 137.06 147.07 156.10 160.03 163.03 186.08
— — — 70 72 — — 86 86 — — 101 84 102 104 110 120 70, 112 133 136 159
57.05 71.08 87.08 97.12 99.13 101.11 103.14 113.16 113.16 114.10 115.09 128.13 128.17 129.12 131.20 137.14 147.18 156.19 — 163.18 186.21
aA dash indicates the immonium ion is not typically observed. bEntries are ordered by monoisotopic mass.
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the precursor m/z can be determined through “zoom scans,” which are slow, narrow, mass-range scans that yield isotopic resolution. For the analysis of complex protein digests, zoom scans should be eliminated in order to maximize the number of MS/MS spectra acquired.
2b. For triple quadrupole (UNIT 16.1) or hybrid quadrupole/TOF mass spectrometers (UNIT 16.2): Acquire the MS/MS spectrum with sufficient collision energy to yield a MS/MS spectrum in which the precursor m/z is ∼50% of the total ion current. On instruments that have automatic, data-dependent acquisition, a range of collision energies is typically used.
Interpret MS/MS spectrum Figure 16.11.2 is an MS/MS spectrum recorded on the doubly charged precursor ion at m/z 531.5 (MH+ = 1062.5 = 531.5 × 2 − 2H + 1) using an ion trap mass spectrometer (LCQ; Thermo Finnigan). The following steps are designed based on the peptide depicted in this figure. Peptides of different length will require greater or fewer iterations of steps 6 to 12. 3. Assume that a peptide resulting from a trypsin digest has either arginine or lysine present at the C terminus. Calculate y1 for both lysine: 128.09 (for lysine) + 18.01 (for H2O) + 1.01 (for added proton) = 147.11 and arginine: 156.10 (for arginine) + 18.01 (for H2O) + 1.01 (for added proton) = 175.21. For triple quadrupole and hybrid quadrupole/TOF mass spectrometers, these low-mass y ions are usually observed; however, in data generated on an ion-trap mass spectrometer (as is the data in Fig. 16.11.2), low-mass y ions may not be observed due to the low-mass cutoff, typically 1⁄3 precursor m/z.
4. If low-mass ions are not observed, calculate the high-mass b ion that would be present for C-terminal lysine and arginine, respectively. a. If the C-terminal residue is lysine, search for the corresponding b ion at MH+ − 18.01 (for H2O) − 128.09 (residue mass for lysine) In Figure 16.11.2, this would be: 1062.5 − 18.01 − 128.09 = 916.4.
b. If the C-terminal residue is arginine, search for the corresponding b ion at MH+ − 18.01 (for H2O) − 156.1 (residue mass for arginine). In Figure 16.11.2, this would be 1062.5 − 18.01 − 156.1 = 888.4. Looking at the spectrum, it is clear that the C-terminal residue is lysine, since an ion at m/z 916.3 is observed. If high-mass ions consistent with both lysine and arginine are present, then extension of both ion series should be attempted in subsequent steps. The incorrect one should not extend very far.
916.3 1062.5
1062.5 Lys 147
Note that there are ions 18 and 36 Da lower than m/z 916.3. These ions correspond to single- and double-neutral losses of water from this b ion, indicating that the peptide sequence will contain residues that are capable of losing water, such as serine, threonine, aspartic acid, and glutamic acid.
Mass Spectrometry
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Supplement 27
848.2
1.1
963.2
Abundance ( x 10 6)
619.2
733.2
215.0 490.2 169.1 187.0
916.3
702.2 361.1
684.3
573.1 443.8
330.2
785.0
666.9
555.0
881.1 898.1
945.0
803.1
260.1
0 200
400
600 m/z
800
1000
1 2 3 4 5 6 7 8 v / D/ D/ N/ E / E / T / I / K 8 7 6 5 4 3 2 1
T 101
E 129
E 129
N 114
D 115
1.1 D 115
N 114
E 129
E 129
T 101
D 115
y7 848.2 I 113
y8 963.2 y5 619.2
Abundance (× 10 6 )
b2 215.0 169.1 a2 187.0
y6 733.2
y4 490.2 y3 361.1
b3 y2 330.2 260.1
b4 443.8
b8 - H2O 898.1
702.2 b8 b6 - H2O b8 - 2H2O 916.3 b5 684.3 b7 - H2O 881.1 y8 - H2O 573.1 945.0 785.0 666.9 b6 b7 555.0 803.1
0 200
400
600 m/z
800
1000
Figure 16.11.2 (A) Unannotated and (B) annotated MS/MS spectrum recorded on the doubly charged precursor ion at m/z 531.5 (LCQ). (Leu/Ile are assigned due to database match.)
De Novo Peptide Sequencing via Manual Interpretation of MS/MS Spectra
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5. Examine the high-mass end of the spectrum and find the y-type ion formed by the loss of the N-terminal residue. The m/z value of this ion will be found between 57 and 186 Da below the MH+, the residue masses of the smallest (glycine) and the largest (tryptophan) amino acids, respectively (Table 16.11.1; also see APPENDIX 1A). In Figure 16.11.2, the strong signal at m/z 963.2 is the most likely candidate for this y ion. The mass difference between 1062.5 and 963.2 is 99.3, close to the residue mass of valine; therefore, the N-terminal residue is valine, and the predicted b1 ion can be calculated as 99.07 (residue mass of valine) + 1.01 (proton).
100.1 Val 1062.5
916.3
1062.5 Lys 147
963.2
A b1 ion will almost never be observed without chemical derivatization. Again note the neutral loss of water from m/z 963.2.
6. Continuing with the y-type ion series at high mass, subtract the next adjacent m/z that has yet to identified and is between 56 and 186 Da from the m/z for the N-terminal residue. Continue until a value roughly corresponding to that expected by an amino acid (Table 11.16.1; also refer to Table A.1A.1 in APPENDIX 1A) is identified. This is the second residue in the peptide. In Figure 16.11.2, the mass difference between m/z 963.2 (N-terminal residue) and 848.2 is 115, the residue mass of aspartic acid (Table 16.11.1; also see APPENDIX 1A). No other mass difference that corresponds to a residue mass can be calculated.
7. Calculate the corresponding b2 ion for a y-type ion at the second position by the following formula: MH+ − putative y ion + proton = corresponding b-type ion. In Figure 16.11.2 this is 1062.5 − 848.2 + 1.01 = 215.3, and an ion at m/z 215.0 can be found in the spectrum. Ions of type a are formed by neutral loss of carbon monoxide (CO) from b-type ions, and are often observed 28 Da less than b ions. An ion at m/z 187.0 is observed in the spectrum in Figure 16.11.2, which is 28 Da lower than the calculated b2 ion above. The presence of both b2 at m/z 215.0 and y7 at m/z 848.2 supports aspartic acid as the second residue from the N terminus.
100.1
215.0
Val 1062.5
Asp 963.2
916.3
1062.5 Lys 147
848.2
Low-mass b ions, particularly b2 and b3, are often easily identified by the presence of an a-b ion pair because they always differ in mass by 28 Da.
8. Continue building the sequence from the N terminus by calculating the next residue mass from the y7 ion. A mass difference of 115 (aspartic acid) is observed between m/z 848.2 (y7) and m/z 733.2. If this is correct, then the corresponding b3 ion can be calculated (1062.5 − 733.2 + 1.01 = 330.3). Note a weak ion at m/z 330.2 is observed in the spectrum.
100.1
215.0
330.2
Val 1062.5
Asp 963.2
Asp 848.2
916.3 733.2
1062.5 Lys 147 Mass Spectrometry
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Supplement 27
9. Continue building the sequence from the N terminus by calculating the next residue mass from the y6 ion. A mass difference of 114 (asparagine) is observed between m/z 733.2 (y6) and m/z 619.2. The corresponding b4 ion can be calculated (1062.5 − 619.2 + 1.01 = 444.3). Note a weak ion at m/z 443.8 is observed.
100.1
215.0
330.2
444.2
Val 1062.5
Asp 963.2
Asp 848.2
Asn 733.2
916.3
1062.5 Lys 147
619.2
10. Continue building the sequence from the N terminus by calculating the next residue mass from the y5 ion. A nominal mass difference of 129 (glutamic acid) is observed between m/z 619.2 (y5) and m/z 490.2. The corresponding b5 ion can be calculated (1062.5 − 490.2 + 1.01 = 573.3). Note an ion at m/z 573.1 is observed.
100.1
215.0
330.2
444.2
573.1
Val 1062.5
Asp 963.2
Asp 848.2
Asn 733.2
Glu 619.2
916.3
1062.5 Lys 147
490.2
Note the single and double neutral water losses from the b5 ion (i.e., m/z 555.1 and m/z 537.1, respectively) due to the aspartic acid and glutamic acid residues in this fragment.
11. Continue building the sequence from the N terminus by calculating the next residue mass from the y4 ion. A nominal mass difference of 129 (glutamic acid) is observed between m/z 490.2 (y4) and m/z 361.1. The corresponding b6 ion can be calculated (1062.5 − 361.1 + 1.01 = 702.4). Note an ion at m/z 702.2 can be observed.
100.1
215.0
330.2
444.2
573.1
702.2
Val 1062.5
Asp 963.2
Asp 848.2
Asn 733.2
Glu 619.2
Glu 490.2
916.3
1062.5 Lys 147
361.1
12. Continue the y-ion series by calculating the next residue mass from the y3 ion. A nominal mass difference of 101 (threonine) is observed between m/z 361.1 and m/z 260.1. The corresponding b7 ion can be calculated (1062.5 − 260.1 + 1.01 = 803.4). Note an ion at m/z 803.1 can be observed.
13. The remaining residue (leucine/isoleucine, or Lxx) can be determined by the mass difference using either the b- or y-type ions. In Figure 16.11.2, this residue is either leucine or isoleucine (Lxx). Because leucine and isoleucine have the same residue mass of 113, these amino acids cannot be differentiated under low-energy collision-induced dissociation of peptides in the ion trap.
De Novo Peptide Sequencing via Manual Interpretation of MS/MS Spectra
100.1
215.0
330.2
444.2
573.1
702.2
803.1
916.3
1062.5
Val 1062.5
Asp 963.2
Asp 848.2
Asn 733.2
Glu 619.2
Glu 490.2
Thr 361.1
Lxx 260.1
Lys 147
14. Confirm the sequence through chemical derivatization of the peptide (see Support Protocols 1 and 2). Determine the first and second amino acids in the sequence, which may be ambiguous in MS/MS analysis, through manual Edman degradation (see Support Protocol 3).
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Current Protocols in Protein Science
CONFIRMATION OF THE DE NOVO SEQUENCE THROUGH METHYL ESTERIFICATION
SUPPORT PROTOCOL 1
Chemical derivatization of peptides is a useful means of facilitating de novo interpretation. The modifications described here often aid in the assignment of y-type and b-type ions. Conversion of the C-terminal carboxylic acid group and the carboxylic acid groups on aspartate and glutamate to the corresponding methyl esters is used to determine the number of acidic groups in the peptide. It is also used to assign y-type ions since a mass shift of +14 Da occurs for every methyl ester formed. Materials Sample Acetyl chloride Anhydrous methanol, ice cold Additional reagents and equipment for LC/MS/MS (see Basic Protocol) 1. Lyophilize appropriate amount of sample to dryness. 2. Prepare a solution of 2 N methanolic HCl by adding 160 ml acetyl chloride dropwise with stirring, to 1 ml ice-cooled anhydrous methanol. Warm to room temperature before use. 3. Add 30 to 40 µl of above reagent to the sample and allow to react at room temperature for 90 min. 4. Remove solvent via lyophilization or vacuum centrifugation using a SpeedVac evaporator. 5. Resuspend esterified material and analyze by LC/MS/MS (see Basic Protocol). In the above example, the mass of the intact peptide would shift by 70 Da (5 × 14) following methyl esterification. This is indicative of four acidic residues plus the C terminus of the peptide. Ions of type b and y would shift by multiples of fourteen as acidic residues are assigned in the sequence (e.g., b2 and y1 shift by 14, b3 and y4 shifts by 28, etc.)
CONFIRMATION OF THE DE NOVO SEQUENCE THROUGH ACETYLATION
SUPPORT PROTOCOL 2
Acetylation of the N-terminal amino group and the ε-amino group of lysine is used to determine the number of lysine residues in a peptide. Conversion of a peptide to its acetylated form results in a mass shift of 42 Da per primary amino group, and as a result, aids in the assignment of b-type ions in a MS/MS spectrum. Acetylation can also result in less extensive charging on the parent, which in turn, can significantly reduce sensitivity. NOTE: This on-column method is best applied when low amounts of sample need to be modified. The same derivatization can be performed in a microcentrifuge tube. In addition, partial acetylation of histidine residues can often be observed. Additional Materials (also see Basic Protocol) Sample Acetic anhydride 200 mM ammonium bicarbonate, pH 8.5 Additional reagents and equipment for LC/MS/MS (see Basic Protocol) 1. Load appropriate amount of sample onto a capillary column (see Basic Protocol). 2. Wash sample with 3 to 4 column volumes water. Mass Spectrometry
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Supplement 27
3. Prepare 1% (v/v) acetic anhydride in 200 mM ammonium bicarbonate, pH 8.5, just prior to use and wash sample with acetylating reagent for 5 to 10 column volumes. 4. Analyze derivatized samples by LC/MS/MS (see Basic Protocol). In the above example, the mass of the intact peptide would shift by 84 Da following acetylation. This is indicative of one lysine and the N terminal of the peptide becoming modified. All b- and y-type ions would shift by 42 Da. SUPPORT PROTOCOL 3
MANUAL EDMAN DEGRADATION One cycle of Edman degradation for the removal of the N-terminal amino acid of a peptide aids in the assignment of the first two amino acids in a sequence, which are often ambiguous in the MS/MS interpretation. Materials Sample Phenylisothiocyanate Pyridine Trifluoroacetic acid, concentrated Butyl acetate Additional reagents and equipment for LC/MS/MS (see Basic Protocol) 1. Reduce sample volume to 2 to 3 µl by vacuum centrifugation in a SpeedVac evaporator, but do not allow to dry completely. 2. Prepare 5% (v/v) phenylisothiocyanate in pyridine, add 5 to 10 µl to the sample, and allow to react 45 min at 45°C. Remove reagent by lyophilization or vacuum centrifugation. 3. Release the phenylthiohydantoin (PTH) derivative of the N-terminal amino acid by adding 15 µl concentrated trifluoroacetic acid. Allow to react 10 min at 37°C. Remove the trifluoroacetic acid by lyophilization or vacuum centrifugation. 4. Separate the PTH-N-terminal amino acid from the truncated peptide by extraction with 15 µl water/15 µl butyl acetate followed by a second extraction with 15 µl butyl acetate. 5. Bring aqueous phase to dryness by lyophilization or vacuum centrifugation in a SpeedVac evaporator. 6. Resuspend material and analyze by LC/MS/MS (see Basic Protocol). Truncated peptides containing the amino acid lysine anywhere in their sequence will also be modified by the above procedure. In these cases, a mass shift of +135 Da per lysine is observed.
De Novo Peptide Sequencing via Manual Interpretation of MS/MS Spectra
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COMMENTARY Background Information Peptide sequencing via tandem mass spectrometry (MS/MS) is one of the most powerful tools in analytical biochemistry for identifying proteins (UNIT 16.1). Until recently, this approach has been limited to experts in the field both for instrument operation and data interpretation; however, commercial instruments today are capable of automated, “on-the-fly” MS/MS with little operator intervention. By coupling reversed-phase liquid chromatography (LC) to mass spectrometry, a single, automated LC/MS/MS experiment can generate hundreds to thousands of MS/MS spectra, enabling the analysis of highly complex mixtures of peptides. Because complete genome sequences have been accumulating rapidly, the recent trend in interpretation of MS/MS spectra has been database matching (Mann and Wilm, 1994; Eng et al., 1994; Clauser et al., 1996; Fenyo et al., 1998; UNITS 16.5 & 16.6). A caveat, however, to database matching is the fact that it can only be successful if the database being searched contains the identical amino acid sequence, or one with high homology, to the peptide being investigated. Even after extensive searching against protein and EST databases, quite often one is left with high-quality tandem mass spectra of peptides for which no exact database match can be made and de novo interpretation is required. The primary difficulty in deriving a complete peptide sequence from a single MS/MS spectrum is the extent of fragmentation that a peptide undergoes during an MS/MS experiment. Ideally, in order to derive complete sequence, fragmentation of peptides in the mass spectrometer must occur at every peptide bond, and the ions resulting from this fragmentation must be detected. In practice, complete fragmentation along the peptide backbone rarely occurs. As a result, the interpretation of most MS/MS spectra yields sequences with varying levels of completeness and ambiguity.
Critical Parameters and Troubleshooting Acquisition of the data When it comes to de novo interpretation, doubly charged tryptic peptides are a mass spectrometrist’s favorite. Since trypsin cleaves on the C-terminal side of arginine and lysine, tryptic peptides contain these basic residues at the C terminus. Under low-energy CID, these
peptides tend to fragment more predictably at amide bonds throughout the length of the peptide, and their MS/MS spectra are comprised mostly of singly charged fragment ions and few doubly charged fragments (Figs. 16.11.3 and 16.11.4). As a result, MS/MS spectra of doubly charged tryptic peptides have the potential to yield nearly complete sequence information. The fragmentation of peptides with internal basic residues is not nearly as complete, due to the absence of fragmentation at peptide bonds adjacent to the basic site. The interpretation of such MS/MS spectra (usually recorded on triply charged precursors and higher) typically results in incomplete sequence information. In addition, MS/MS spectra recorded on triply charged precursors and above are often complicated by the presence of multiply charged fragment ions (Fig. 16.11.5). Oftentimes, the de novo interpretation only yields partial sequence information using a multiply-charged ion series. Instruments, such as hybrid quadrupole/TOF (QTOF, Micromass), that have the resolving power to determine charge state on fragment ions because of isotope spacing, can significantly aid interpretation, although complete sequence is still not guaranteed (Fig. 16.11.6). De novo interpretation of MS/MS spectra The only way to learn de novo interpretation of MS/MS spectra is by reviewing a substantial number of MS/MS spectra of known amino acid sequence. The figures in this unit are a good representation of common observations made by looking at larger numbers of MS/MS spectra. By doing so, one becomes familiar with general observations and trends that may aid de novo interpretation of MS/MS spectra generated on peptides of unknown sequence. It is important to note that none of these observations should become hard and fast rules. Each peptide will fragment differently depending upon its amino acid composition; therefore, what is observed for one peptide may not be observed for another with similar amino acids. Sequence-specific fragment ions. Whether or not a complete peptide sequence is attainable for any given MS/MS spectrum depends upon the extent of fragmentation and detection of sequence-specific fragment ions (b- and y-type ions). Often the fragmentation is incomplete, and the spectrum is complicated by nonsequence-specific fragment ions (see below) and neutral losses from b- and y-type ions (water
Mass Spectrometry
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Supplement 27
1 2 3 4 5 6 7 8 9 L/ S/ S / P/ A/ T / L/ N S R 9 8 7 6 5 4 3 2 1 y8++ - H2O 414.3
Relative abundance (× 10 6 )
4.1
L
T
113
101
A
P
S
S
71
97
87
87
y5 590.1
y8++ 423.2 y7++
y3 b2 200.8
y7 758.2
y8++ - 2H2O
y4 489.4 b6 558.2
PAT 269.8 376.2
y8 845.2
y6 661.2
y9 932.5
0 200
400
600 m/z
800
1000
Figure 16.11.3 MS/MS spectrum recorded on the doubly charged precursor ion at m/z 523.4 (LCQ), showing near complete y-type ion series. (Leu/Ile are assigned due to database match.)
1 2 3 4 5 6 7 8 A/ A/ E / D/ Y/ G V/ I / K 8 7 6 5 4 3 2 1 I 113
(GV) 156
Y 163
D 115
E 129
(GV) 156
I 113
4.1 E 129
D 115
Y 163
A 71
y 7 823.2
Abundance (× 10 5 )
y6 694.1
b4 - H2O b5 b2 y5 b2 y4 ++ y b 579.3 416.1 8 3 y2 y1 447.8 142.9 260.1 271.3 387.1 549.3 147.0
b8 b7 819.3 706.0
y
8
894.2
0 200
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16.11.10 Supplement 27
400
600 m/z
800
1000
Figure 16.11.4 MS/MS spectrum recorded on the doubly charged precursor ion at m/z 483.2 (LCQ), showing nearly complete b- and y-type ion series. (Leu/Ile are assigned due to database match.) Current Protocols in Protein Science
1 2 3 4 5 6 7 8 9 10 11 R H/ P E/ Y/ A/ V/ S/ V / L/ L R 11 10 9 8 7 6 5 4 3 2 1 L
V
S
V
YA
113 b6++ 377.7
99
87
99
234
1.2
PE
Y
A
226
163
71
V 99
Abundance (× 10 6 )
y5 y3 587.4 401.4 b7++ 427.3 b6 754.4
b8++ 470.3
b5++
y10++ y4 574.1 y6 500.3 686.5 b5 b4 683.4 520.3
b2 342.3 294.2 y2 288.3
b7 y8 920.5 853.0
0 200
400
600
800 m/z
1000
1200
1400
Figure 16.11.5 MS/MS spectrum recorded on the triply charged precursor ion at m/z 480.8 (LCQ) with extensive doubly charged fragments. Resolution on the ion trap precludes charge state determination on these fragment ions. (Leu/Ile are assigned due to database match.)
1 2 3 4 5 6 7 8 9 10 11 R H/ P E/ Y/ A/ V/ S/ V / L/ L R 11 10 9 8 7 6 5 4 3 2 1 L
L
V
S
V
YA
113
113
99
87
99
234
b6++ 377.7
100
377.73
100 [M+2H]++ y5 480.7 587.5
Relative abundance
b5++ 342.2 y2 PE 288.2 260.7 b2 294.2 V 72.1 Y y1 136.1175.1 I/L 86.1
378.23
PE
Y
A
V
226
163
71
99
y3 401.4
378.74 0
377
378
379
b7++ 427.3 y4 500.4 y6 686.6 b6 b4 y ++ 754.5 10 520.3 573.9 b5 b7 683.4 853.6
y8 920.7
0 200
400
600
800 m/z
1000
1200
1400
Figure 16.11.6 MS/MS spectrum recorded on the triply charged precursor ion at m/z 480.7 (QTOF), with several doubly charged fragments. TOF resolution enable charge state determination on the fragment ions from the isotope spacing (inset). (Leu/Ile are assigned due to database match.)
Mass Spectrometry
16.11.11 Current Protocols in Protein Science
Supplement 27
detected) between the first two N-terminal amino acids. Subsequently, b1 ions are almost never observed, and this results in ambiguity at the N terminus of a peptide (Fig. 16.11.10). Quite often, one knows the mass of possible dipeptides (Table 16.11.2) that could complete the sequence, but the order of the amino acids cannot be determined. Sequence gaps are not limited to the N terminus, can involve more than just a dipeptide, and can occur anywhere in the putative sequence (Figs. 16.11.11 & 16.11.12). If fragmentation is sufficiently poor, an MS/MS spectrum may only contain fragment ions that are related to the same part of the peptide. In this case, the interpretation yields redundant and overlapping information despite the abundance of b- and y-type ions (Fig. 16.11.13). Another complication is the observation that certain amino acids, such as proline, can influence the fragmentation process. In the case of proline, the peptide bond on the N-terminal side of this amino acid appears to be particularly labile. For peptides containing proline, the resulting MS/MS spectrum is usually dominated by the y-type ion ending in proline (Fig. 16.11.14).
and ammonia). Loss of water from y- and/or b-type ions usually occurs with peptides that contain serine, threonine, or glutamic and aspartic acid. Loss of ammonia from y- and/or b-type ions usually occurs with peptides that contain arginine, lysine, and to a lesser extent, asparagine and glutamine. Neutral-loss ions can add complimentary information while interpreting a spectrum. If loss of water or ammonia is observed from a y-type or b-type ion, then the appropriate amino acid should be present in the putative sequence; however, neutralloss ions can also complicate MS/MS spectra when these spectra are generated on peptides that contain an abundance of residues that can yield them (Figs. 16.11.7 & 16.11.8). In these spectra, ions resulting from loss of ammonia or water can be higher in abundance than the corresponding b- or y-ion, complicating the interpretation. Furthermore, neutral-loss ions resulting from loss of phosphoric acid (98 Da) in MS/MS spectra of phosphopeptides are extremely useful in indicating the presence of a phosphorylated residue in the peptide and aid in determining the site of phosphorylation in these sequences (Fig. 16.11.9). Perhaps the most common observation for de novo sequencing is the lack of fragmentation (and subsequently the lack of fragment ions
T 101 3.0
1 2 3 4 5 6 7 8 9 D E/ D/ N/ N/ L/ L / T/ E/ K 9 8 7 6 5 4 3 2 1 L L N N D 113 113 114 114 115
y3 377.1 D N 115
114
y8 946.2
N
L
L
TE
114
113
113
230
y4 490.2
Abundance (× 10 5 )
b7 - NH3 b6 - NH3 797.0 b2 b7 y6 244.9 814.0 y5 717.2 603.2 b4 - H2O y7 b O H 5 2 b3 - H2O 831.1 779.0 b7 - NH3 - H2O b8 - NH3 a2 b6 569.8 217.0 y2 b8 - NH3 - H2O b3 b4 684.2 701.2 898.1 276.0 359.9 454.9 b 5 b9 472.8 880.1 341.7 587.8 1044.2 0
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16.11.12 Supplement 27
200
400
600
800
1000
1200
m/z
Figure 16.11.7 MS/MS spectrum recorded on the doubly charged precursor ion at m/z 595.9 (LCQ), with extensive losses of water and ammonia from side chains. (Leu/Ile are assigned due to Current Protocols in Protein Science
1 2 3 4 5 6 7 L S/ S/ P A T / L/ N 7 6 5 4 3 2 1 S 87 y5 515.2
1.7
b7 - 2H2O b7 - H2O 634.2 652.1 L 113
b6 - 2H2O
b7 670.1
Abundance (× 10 6 )
b6 - H2O 521.1 539.1
y6 602.1
[M+H]+ - 2H2O [M+H]+ - H2O
[M+H]+ PATL 383.2
PAT 269.9
766.2 784.2
b6 557.1
802.3
0 400
600
800
m/z
Figure 16.11.8 MS/MS spectrum recorded on the singly charged precursor ion at m/z 802.3 (LCQ), with extensive neutral loss of water from the side chains of serine and threonine. (Leu/Ile are assigned due to database match.)
1 2 3 4 5 6 7 N Y/ M/ s/ E H L L / K 8 7 6 5 4 3 2 1 [M+2H - H3 PO4]++ 559.1
9.9
s 167
Abundance (× 10 5 )
M 131
M 131
y7 - H3PO4 839.4
[M+2H - H3 PO4 - H2O] ++ 550.5
a2 250.1 b2 278.0 b3 409.3 0 200
400
y6 806.3 y6 - H3PO4 y5 708.3 639.4 600
800
y7 937.3 b8 1068.2 1000
1000
m/z
Figure 16.11.9 MS/MS spectrum recorded on the doubly charged precursor ion at m/z 608.1 (LCQ), with neutral loss ions resulting from the loss of phosphate. Note that these ions aid in the definitive assignment of the phosphorylated serine residue. (Leu/Ile are assigned due to database match.)
Mass Spectrometry
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Supplement 27
16.11.13
1 2 3 4 5 6 7 8 G F/ L / I / D/ G/ Y/ P/ R 8 7 6 5 4 3 2 1 Y G D I L 163 57 115 113 113 y5 590.1
2.8 L 113
I 113
D 115
G 57
Y 163
P 97
Abundance (× 10 7 )
y6 720.3
b3 317.9 y4 492.2 b5 546.1 431.0 b6 b4
b2 y2 204.9 272.2 a2 177.0
y3
a4
y7 b7 833.2 766.1 b8 864.2
603.2
0 200
400
600
800
1000
m/z
Figure 16.11.10 MS/MS spectrum recorded on the doubly charged precursor ion at m/z 519.4 (LCQ), showing near complete series of both b- and y-type ions, but the order of the first two residues (GF and FG) cannot be determined from the spectrum. (Leu/Ile are assigned due to database match.)
1 L
2 3 4 5 6 7 8 9 10 11 12 13 Y/ T / L / V / L / T / D P / D / A /P S / R 13 12 11 10 9 8 7 6 5 4 3 2 1
A 71
(PD) 212
Abundance (× 10 6 )
1.1
D 115
T 101
L 113
y7 y9 757.1 y6 971.2 y8 642.2 858.2
(LV)
L
212
113
b5 590.1
359.2
y4 430.1 b3 378.3
L 113
T 101
T
D
(PD)
(APS)
101
115
212
255
b6 919.2
b6 703.1 y3
V 99
y10 1070.1
b7 833.2
y11
1184.4 y12 b10 1284.6 b13 1130.8 1386.3
0 400
De Novo Peptide Sequencing via Manual Interpretation of MS/MS Spectra
600
800
1000
1200
1400
m/z
Figure 16.11.11 MS/MS spectrum recorded on the doubly charged precursor ion at m/z 781.0 (LCQ). Note the order of the residues PD/DP and PS/SP cannot be determined due to the lack of fragmentation between these residues. (Leu/Ile are assigned due to database match.)
16.11.14 Supplement 27
Current Protocols in Protein Science
Mass Spectrometry
16.11.15
Current Protocols in Protein Science
Supplement 27
Gly (57)
115 129 145 155 157 159 161 171 172 173 186 187 189 195 205 214 218 221 244
Amino acid (mass)
Gly (57) Ala (71) Ser (87) Pro (97) Val (99) Thr (101) Cys (103) Leu/Ile (113) Asn (114) Asp (115) Qln/Lys (128) Glu (129) Met (131) His (137) Phe (147) Arg (156) Camc (160) Tyr (163) Trp (186) 143 159 169 171 173 175 185 186 187 200 201 203 209 219 228 232 235 258
Ala (71)
175 185 187 189 191 201 202 203 216 217 219 225 235 244 248 251 274
Ser (87)
Table 16.11.2 Dipeptide Masses (MH+)
195 197 199 201 211 212 213 226 227 229 235 245 254 258 261 284
Pro (97)
199 201 203 213 214 215 228 229 231 237 247 256 260 263 286
Val (99)
203 205 215 216 217 230 231 233 239 249 258 262 265 288 207 217 218 219 232 233 235 241 251 260 264 267 290 227 228 229 242 243 245 251 261 270 274 277 300 229 230 243 244 246 252 262 271 275 278 301 213 244 245 247 253 263 272 276 279 302 257 258 260 266 276 285 289 292 315
259 261 267 277 286 290 293 316
263 269 279 288 292 295 318
275 285 294 298 301 324
295 304 308 311 324
313 317 320 343
321 324 347
327 350
373
Thr Cys Leu/Ile Asn Asp Gln/Lys Glu Met His Phe Arg Camc Tyr Trp (101) (103) (113) (114) (115) (128) (129) (131) (137) (147) (156) (160) (163) (186)
1 2 3 4 5 6 7 8 9 10 I/ F/ G /V / T/ T L / D/ I V R 10 9 8 7 6 5 4 3 2 1 TL T V G 214 101 99 57 y7 817.3
Abundance (× 10 6 )
1.9
G
V
T
TL
D
57
99
101
214
115
y9 916.3
y6
b2 260.9
a4 y4 389.1 b4 502.2 b3 318.0 417.1
a2
F 147
233.0
716.3 b7
b5
b8
y8 916.3 732.0 846.9
517.5
y10 1120.6
0 200
400
600
800
1000
1200
m/z
Figure 16.11.12 MS/MS spectrum recorded on the doubly charged precursor ion at m/z 617.3 (LCQ). Note the order of the residues TL/LT cannot be determined because of the lack of fragmentation between them. (Leu/Ile are assigned due to database match.)
1 2 3 4 5 6 7 8 9 10 N V / V / I / A /A D G V L 10 9 8 7 6 5 4 3 2 1 A 71
K
I 113
V 99
8.5 V 99
I 113
A 71
y9 885.4
y8 786.4
Abundance (× 10 6 )
b5 497.0 b3 313.1 b2 213.9 a2 185.9
y7 673.3
b4 426.0
y6 602.0
0 200
De Novo Peptide Sequencing via Manual Interpretation of MS/MS Spectra
400
600 m/z
800
1000
Figure 16.11.13 MS/MS spectrum recorded on the doubly charged precursor ion at m/z 550.0 (LCQ), with redundant fragmentation limited to one portion of the peptide. (Leu/Ile are assigned due to database match.)
16.11.16 Supplement 27
Current Protocols in Protein Science
monium ions are almost always observed on triple quadrupole and hybrid quadrupole/TOF mass spectrometers, but are not generally observed on ion traps due to the low mass restriction employed on these instruments. High-energy CID instruments produce a richer variety of immonium and related ions (Falick et al., 1993). Isobaric amino acids. Isobaric amino acids, such as leucine/isoleucine and lysine/glutamine, add ambiguity to de novo interpretation. Under low-energy CID in a trap, triple quadrupole, or hybrid quadrupole/TOF instrument, these residues cannot be differentiated by mass, with the possible exception of K/Q in a hybrid quadrupole/TOF. The difference in mass between lysine and glutamine is 0.03 Da, which is within the mass-accuracy range of this instrument, provided that the instrument is properly calibrated. Lysine and glutamine can also be differentiated by chemical derivatization, such as acetylation, which modifies the ε-amino group of lysine and the N terminus; however, blocking these basic sites has the potential of decreasing the ionization efficiency for the acetylated peptide, especially if there are no other basic sites in the sequence with which to carry a charge.
Nonsequence-specific fragment ions There are two types of nonsequence-specific fragment ions: internal fragment ions and immonium ions. Internal ions result when fragmentation occurs at two peptide bonds simultaneously. In MS/MS spectra generated on ion trap mass spectrometers, internal fragment ions are rarely observed; however, they can be quite common in MS/MS spectra generated on hybrid quadrupole/TOF or triple quadrupole instruments. Typically, the more abundant internal fragment ions contain proline at the N terminus of the internal fragment (Fig. 16.11.8). Immonium ions are low-mass ions that are indicative of amino acid composition (Fig. 16.11.15; Falick et al., 1993). They are not indicative of where in the sequence the specific amino acid may be located nor are they indicative of the number of occurrences of that particular amino acid. Not all immonium ions for every amino acid are typically observed, and therefore, the absence of a particular immonium ion does not necessarily mean the absence of its corresponding amino acid in the sequence; however, the presence of an immonium ion of somewhat significant abundance typically indicates that its corresponding amino acid is present in the sequence. Im-
1 2 3 4 5 6 7 L/ P/ L/ Q D/ V Y K 7 6 5 4 3 2 1 y7++ 431.9
3.6
Abundance (× 10 6 )
(QD) 243
L 113
P 97
y6 765.3
b2 210.9 a2
y3 409.1
200
400
y5 b5 652.2 567.2
y7 862.2
0 600 m/z
800
1000
Figure 16.11.14 MS/MS spectrum recorded on the doubly charged precursor ion at m/z 488.1 (LCQ). Fragmentation is dominated by the proline residue.
Mass Spectrometry
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Supplement 27
ine, which has the same nominal residue mass as phenylalanine. Oxidation of methionine is observed oftentimes in the analysis of in-gel digestions. Similarly, the mass of certain dipeptides is equal (or nearly equal) to the residue mass of a single amino acid (Table 16.11.3).
A common pitfall of de novo interpretation is the fact that some modified amino acids may have the same nominal residue mass as unmodified amino acids (Table 16.11.3), so that distinguishing them without ambiguity may require a high-mass accuracy instrument. The most common example of this is oxidized methion-
Table 16.11.3 Residue Mass Combinations and Modifications that Equal the Mass of Single Residues
Dipeptide or modified residue
Nominal mass
Single residue
Gly-Gly Gly-Ala Val-Gly Gly-Glu Ala-Asp Ser-Val AcGly AcAla AcSer Metox AcAsn
114 128 156 186 186 186 99 113 129 147 156
Asn Qln/Lys Arg Trp Trp Trp Val Leu/Ile Glu Phe Arg
1 2 3 4 5 6 7 8 9 10 11 12 L G/ E / Y/ G/ F/ Q/ N/ A/ L / I/ V/ R 12 11 10 9 8 7 6 5 4 3 2 1 E 129
Y 163
b3 300.2
100 V b2 99 171.1
I
L
A
113 113 71
N
Q
F
G
Y
114
128
147
57
163
Abundance
y2 274.2
175.1 y1 I/L 86.1 F Y 120.1136.1
y3 387.3
[M+2H]++ 740.4
b4 463.2
GE GEY 187.1 350.2
y10++ 590.9y
6 y7 y4 685.5 813.6 500.4 571.4 y 5
y9 1017.7 y8 960.7
y10 1180.7
0 200
400
600
800
1000
1200
1400
m/z
De Novo Peptide Sequencing via Manual Interpretation of MS/MS Spectra
Figure 16.11.15 MS/MS spectrum recorded on the triply charged precursor ion at m/z 740.4 (QTOF). Note that the N-terminal LGEY sequence is difficult to discern as the b-type ions and internal ions complicate the interpretation.
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Current Protocols in Protein Science
Time Considerations Although quick arithmetic skills and memorized amino acid masses facilitate the process, manual interpretation of MS/MS spectra is extremely variable and depends primarily on the quality and complexity of the MS/MS spectrum. Some interpretations require only a few minutes, whereas other spectra require extensive manipulation and chemical derivatization to complete the interpretation.
Literature Cited Clauser, K.R., Baker, P.R., and Burlingame, A.L. 1996. Peptide Fragment-Ion Tags from MALDI/PSD for Error Tolerant Searching of Genomic Databases. In Proceedings of the 44th ASMS Conference Mass Spectrom. p. 365. Allied Topics, Portland, Oreg.
Falick, A.M., Hines, W.M., Medzihradszky, K.F., Baldwin, M.A., and Gibson, B.W. 1993. Lowmass ions produced from peptides by high-energy collision-induced dissociation in tandem mass spectrometry J. Am. Soc. Mass Spectrom. 4:882-893. Fenyo, D., Qin, J., and Chait, B.T. 1998. Protein identification using mass spectrometric information. Electrophoresis 19:998-1005. Mann, M. and Wilm, M. 1994. Error tolerant identification of peptides in sequence databases by peptide sequence tags. Anal. Chem. 66:43904399.
Contributed by Terri Addona and Karl Clauser Millennium Pharmaceuticals Cambridge, Massachusetts
Eng, J.K., McCormick, A.L., and Yates, J.R. 1994. An approach to correlate tandem mass spectral data of peptides with amino acid sequences in a protein database. J. Am. Soc. Mass Spectrom. 5:976-989.
Mass Spectrometry
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CHAPTER 17 Structural Biology INTRODUCTION
P
roteins have evolved to perform specific functions which depend on their three-dimensional structure. These three-dimensional structures are directly related to amino acid sequences (primary structure) of the proteins. Different segments of the primary structure form units of secondary structure, such as α helices or β pleated sheets. The tertiary structure refers to the three-dimensional arrangement of all the atoms that constitute a protein molecule. It is defined by the precise spatial coordination of the component secondary structural elements. The tertiary structure of proteins can usually be divided into domains which are comprised of distinct compact folding units usually containing about 100 to 200 amino acid residues. Small proteins may contain only a single domain, whereas most proteins contain multiple domains. A fold refers to the characteristic spatial assembly of secondary structure elements that comprise a domain; they are usually common to many proteins, e.g., the immunoglobulin fold. In UNIT 17.1, the authors give an overview of protein structural characteristics and provide an extensive survey of common structural folds that form the building blocks of the three-dimensional structures of proteins. For each fold described, the salient structural features are described, usually accompanied by illustration. Several technologies are employed to study the three-dimensional structures of proteins. Electron microscopy (EM) can be utilized in protein structural analyses (UNIT 17.2). Although EM has long been employed to visualize the substructure of cells and tissues and the spatial array of proteins within them, it has not been extensively utilized for the study of proteins in general. The goal of this unit is to give a general overview of current EM procedures for observing proteins and to outline the kind of information that may be acquired by illustrating this with specific examples. The premise is that EM is complementary to—not in competition with—more established techniques such as X-ray crystallography and nuclear magnetic resonance (NMR) for the study of three-dimensional structure. Another means of imaging biological samples with molecular resolution is by atomic force microscopy (AFM), as described in UNIT 17.7. In addition to providing topographical images of surfaces with nanometer- to angstrom-scale resolution, forces between single molecules and mechanical properties of biological samples can be investigated. Importantly, unlike EM, the biological samples are studied within a physiological-like environment, permitting measurement of temporal changes in structure. The major limitations of AFM are that the specimen must be firmly attached to a solid support for measurement and the information gained is localized to surface structures. UNIT 17.3 deals with the principles of macromolecular X-ray crystallography. The first part
of the unit provides an overview of the underlying principles upon which this technology is based. It discusses the generation of the diffraction pattern, data collection, phase determination, resolution, refinement, and the Ramachandran plot, which is used by crystallographers to assess the quality of their structure determinations and to detect errors in the tracing of the polypeptide chain. Appreciation of these parameters should allow one to critically assess the quality of reported crystal structures. In the second part of the unit, the methods and concerns for preparing a protein sample for crystallographic studies are described. Included are details about preparation, assessment of quality and complexity, and storage of the protein samples. Procedures for setting up initial crystallization Contributed by John E. Coligan Current Protocols in Protein Science (2003) 17.0.1-17.0.2 Copyright © 2003 by John Wiley & Sons, Inc.
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trials are described with suggestions for modifying these trials should they be unsuccessful. Finally, for cases where all else has failed to gain useful crystals, protein engineering strategies are discussed using HIV-1 integrase as a model protein for which these strategies have proven successful. The major bottleneck in structural determination by X-ray crystallography is no longer a lack of sufficiently pure material—this problem has largely been eliminated by producing proteins with recombinant technology—but instead the preparation of suitable crystalline samples. Advances in procedures for macromolecular crystallization, combined with the availability of commercial screening kits, have made it possible to set up crystallization screening trials in a conventional biochemistry laboratory. This is particularly desirable given that crystallization of the protein will generally enhance an investigator’s chances of attracting a crystallographic collaborator. UNIT 17.4 outlines the steps for accomplishing this starting with a purified sample. Protocols are given for concentrating the sample, for preparing and assessing the screening trials, and finally for optimizing the crystallization conditions. Other than X-ray crystallography, the only technique currently available that can provide a complete description of macromolecules at the atomic level is nuclear magnetic resonance (NMR) spectroscopy. An advantage of NMR measurements over crystallography is that they can be carried out in solution under potentially physiological conditions. Recent advances in NMR technology, including the use of triple-resonance methods and deuterium labeling, have dramatically increased the molecular weight limitations inherent to this methodology. It is now feasible to consider using this approach for proteins approaching 100 kDa in size. UNIT 17.5 aims to provide an overview of the application of NMR spectroscopy to protein structure determination. It is not meant to be a how-to guide, as the technique itself requires extensive expertise. It does, however, provide information about sample preparation and the type of information that can be gleaned from the technology. The unit starts with a description of the basic principles and instrumentation of NMR, followed by spectral assignment methods for various size proteins and the utility of 13C and 15N to aid in structural assignments for larger proteins. Employment of multiple technologies other than those described above can provide valuable information about protein structure and dynamics. Among these, the determination of hydrogen exchange rates at backbone peptide amide linkages of proteins has been used for many years as a sensitive probe for detecting changes in protein structure in response to changing conditions, such as pH and temperature. The rates at which amide hydrogens in protein exchange with deuterium or tritium in the bulk solvent are highly dependent on protein conformation. Mass spectrometry is a sensitive and accurate method for detecting such interchanges. After fragmentation, hydrogen exchange-mass spectrometry (HX-MS), as described in UNIT 17.6, can be used to determine deuterium levels in short segments of protein. UNIT 17.6 discusses the interplay between protein dynamics and hydrogen exchange and provides details for designing and performing typical HX-MS experiments, as well as common procedures for processing and interpreting HX-MS data. Raman spectroscopy, described in UNIT 17.8, is a method for probing structures and interactions of molecules by measuring the energies (or frequencies) of molecular vibrations. UNIT 17.8 surveys the general nature, underlying theoretical basis and experimental requirements of protein Raman spectroscopy. Emphasis is given to applications that probe protein conformation, intra- and inter-molecular protein interactions, subunit folding and assembly pathways, and protein-nucleic acid interactions. John E. Coligan Introduction
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Overview of Protein Structural and Functional Folds INTRODUCTION TO PROTEIN STRUCTURE Proteins fold into stable three-dimensional shapes, or conformations, that are determined by their amino acid sequence. The complete structure of a protein can be described at four different levels of complexity: primary, secondary, tertiary, and quaternary structure. As a multitude of protein structures are rapidly being determined by X-ray crystallography and by nuclear magnetic resonance (NMR), it is becoming clear that the number of unique folds is far less than the total number of protein structures. Not only do functionally related proteins generally have similar tertiary structures (see below), but even proteins with very different functions are often found to share the same tertiary folds. As a consequence, structural conservation at the tertiary level is perhaps more profound than it is at the primary. The identification of the fold of a protein has therefore become an invaluable tool since it can potentially provide a direct extrapolation to function, and may allow one to map functionally important regions in the amino acid sequence. Several groups have already attempted to classify protein structures into fold families and superfamiles without focusing on function (Orengo et al., 1993; Murzin et al., 1995). The scope of this unit is not to enumerate all the existing folds and tertiary structures determined to date, but rather to provide a comprehensive overview of some commonly observed protein fold families and commonly observed structural motifs which have functional significance. Likewise, the PDB-entry tables given in this unit provide some examples of various folds, but are not comprehensive lists. The unit is organized into sections based on both structural and functional relations.
Primary Structure Primary structure is defined as the linear amino acid sequence of a protein’s polypeptide chain. In fact, the term protein sequence is often used interchangeably with primary structure. In 1973, Chris Anfinsen demonstrated that the primary amino sequence of a protein
UNIT 17.1
uniquely determines the higher orders of structure for a protein and is thus of fundamental importance (Anfinsen, 1973). It is noteworthy, however, that changes in the local biological environment of a protein molecule can sometimes perturb its three-dimensional structure. For example, interactions with ligands, substrates, or other proteins can bring about controlled conformational changes producing potentially profound effects. Furthermore, a few proteins have been found to have intrinsically unstructured regions (Wright and Dyson, 1999; Tompa, 2002). Hence, although structural uniqueness associated with a protein sequence is a powerful principle, it can sometimes depend on the local environment and is not rigidly followed in every case.
Secondary Structure Secondary structure is defined as the local spatial conformation of the polypeptide backbone excluding the side chains. Regular secondary structures (also referred to as secondary structure elements) common to many proteins include α-helices, β-sheets, and turns (see below). They can vary widely in length, from as few as three to five residues in short helices and sheets, to over fifty residues in some coiled-coil helices (see Frequently Observed Secondary Structure Assemblies or Structural Motifs). Such structures are generally defined both by characteristic main chain φ and ψ dihedral angles (i.e., the torsion angles between backbone atoms Ci-1–Ni–Ciα–Ci and Ni–Ciα–Ci–Ni+1, respectively; Fig. 17.1.1A) and by regular main chain hydrogen bonding patterns. When discussing the structure of a protein, the term topology is often used to refer to the connectivity of secondary structure elements. For example, a portion of a protein containing a β-strand connected to an α-helix and then another βstrand is said to have a βαβ topology. The complete secondary structure of a polypeptide chain is often represented in two dimensions by a topology diagram (e.g., Fig. 17.1.1H) which shows both the connectivity and the relative orientation of neighboring secondary structure elements. Such diagrams are particularly useful in classifying β-sheets.
Structural Biology Contributed by Peter D. Sun, Christine E. Foster, and Jeffrey C. Boyington Current Protocols in Protein Science (2004) 17.1.1-17.1.189 Copyright © 2004 by John Wiley & Sons, Inc.
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α-helices An α-helix is formed when a protein backbone adopts a right-handed helical conformation with 3.6 residues per turn and a set of hydrogen bonds formed between the main chain carbonyl (CO) of the ith residue and the main chain NH of the (i+4)th residue (Fig. 17.1.1B). Occasionally, hydrogen bonds are observed between residues i and i+3, resulting in a 310 helix. Compared to the more common α-helices, 310 helices are much shorter, usually comprising only one to three turns. The optimal backbone φ and ψ dihedral angles for a righthanded α-helix are −57° and −47°, while for a 310 helix they and −49° and −26°, respectively. β-sheets β-sheets (also referred to as β-pleated sheets) make up another major secondary structural element in proteins. A β-sheet consists of at least two β-strands, each approaching an extended backbone conformation with dihedral angles confined to the region where the φ torsion angle is between −60° and −180°, and the ψ torsion angle is between 30° and 180° (i.e., −180°<φ<−60°, 30°<ψ<180°). All main-chain CO-to-NH hydrogen bonds lie between adjacent strands (Fig 17.1.1C). A parallel β-sheet is formed when all the sheet-forming strands run parallel to each other and in the same direction from N to C termini, whereas an antiparallel β-sheet is formed when the strands are still parallel to each other, but run in opposite directions (Fig. 17.1.1C); a β-sheet formed from both parallel and antiparallel strands in referred to as a mixed β-sheet. A characteristic feature of β-sheets is the right handed twist which is visible when the sheet is viewed edgeon (Fig. 17.1.1D).
Overview of Protein Structural and Functional Folds
Turns and loops Loops and turns connect helices and βsheets in protein structures. Most turns and loops assume irregular secondary structures, but important exceptions are the type I and II reverse turns (also referred to as β-turns or β-bends), which are tight turns connecting adjacent, antiparallel β-strands. Shown in Figure 17.1.1E, type I and II turns each have a hydrogen bond between the CO of the first residue in a turn and the NH of the fourth residue (also the last residue) in the turn. The difference between the type I and II turn is a 180° flip of the peptide plane in the middle of the turn. The type I turn is more frequently observed than the type II. The terms loops and coils generally
refer to secondary structure elements that display less regular hydrogen bonding patterns than those observed in α-helices, β-sheets, and reverse turns.
Tertiary Structure Tertiary structure refers to the three-dimensional arrangement of all the atoms that constitute a protein molecule. It relates the precise spatial coordination of secondary structure elements and the location of all functional groups of a single polypeptide chain.
Domains, Folds, and Motifs The tertiary structure of a protein can often be divided into domains—i.e., distinct compact folding units usually comprising 100 to 200 residues. Small proteins may contain a single domain, whereas larger proteins often contain multiple domains. A fold refers to a characteristic spatial assembly of secondary structure elements into a domain-like structure that is common to many different proteins. Particular folds are often, though not always, related to a certain function (e.g., nucleotide-binding folds). A structural motif is similar to a fold, but is generally smaller, tending to form the building blocks of folds. Some structural motifs (e.g., β-barrels) are observed in a vast array of unrelated proteins with many variations, while others, especially those related to a unique biochemical function (e.g., zinc fingers; see Zinc-Containing DNA-Binding Motifs), are highly conserved, generally occurring in protein domains of similar function. Highly conserved structural motifs often display a characteristic signature at their amino acid sequence level, the so-called sequence motif. Very often the term motif is used alone to refer to either a structural motif or a sequence motif. This can lead to confusion since structural motifs do not always have a unique amino acid signature. It is important to note that the terms domain, fold, and motif are often used interchangeably with blurred distinctions. This happens especially for larger, highly conserved structural motifs which may be as equally well known as a particular type of domain or fold.
Quaternary Structure The structure of many proteins, especially those >100 kDa in mass, are oligomers consisting of more than one polypeptide chain. The precise spatial arrangement of the subunits within a protein is referred to as the quaternary structure.
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Frequently Observed Secondary Structure Assemblies Or Structural Motifs
separate subunits but reside in a topologically equivalent part of a protein.
This section describes some structural motifs commonly observed in a wide variety of proteins. Other structural motifs of particular functional significance, such as DNA- and RNA-binding motifs, are described in later sections of this unit.
β-hairpin When an antiparallel β-sheet contains only two strands connected by a single tight turn, it is termed a β-hairpin (Fig. 17.1.1G). A β-hairpin is usually highly twisted and less regular compared to a more extended β-sheet.
α-helix bundles Helix bundles (or helical bundles) usually refer to the packing of several helices in a nearly parallel or antiparallel orientation. In some cases the term also applies to the general packing of helices regardless of their orientation to each other. There are many examples of protein structures that contain three-, four-, or multiple-helix bundles, such as cyclin (three-helix bundle), hematopoietic cytokines (four-helix bundle), globins (six-helix bundle), and cytochrome c oxidase (22-helix bundle). Helix bundles are often distinguished by the number of helices in the bundle, the relative direction of the helices, and the characteristic twist of each helix bundle. In describing the direction of the helices in a bundle, both parallel/antiparallel and up/down terminologies are used in the literature, although the latter terms give a more precise description (Fig. 17.1.1F). In the first convention, two helices are said to be parallel if their N- to C-terminal vectors, or peptide directions, are parallel to each other (i.e., pointing in the same direction). The opposite is referred to as antiparallel. In the up/down notation, the first helix in a bundle is defined as the ‘up’ helix, and all subsequent helices are assigned either ‘up’ or ‘down’ depending on whether they are parallel or antiparallel, respectively, to the first helix. In most helical bundles, the axes of individual helices are inclined to the representative axis of the entire bundle in a systematic manner, forming a twist with defined handedness that resembles a spiral. Most helix bundles carry a left-handed twist.
Greek key and jelly-roll motifs In an antiparallel β-sheet assembly, four adjacent strands are sometimes arranged in a pattern similar to the repeating unit of one of the ornamental patterns used in ancient Greece. Thus, this type of strand assembly is termed a Greek key motif (Fig. 17.1.1H). The immunoglobulin fold is one example of a fold that contains the Greek key motif. A certain arrangement of two Greek key motifs form the so-called jelly roll (Fig. 17.1.1I). In this case, strands 1, 2, 7, and 8 form one Greek key, while strands 3, 4, 5, and 6 form the other. The eight strands fold into a nearly closed barrel shape. A characteristic feature in the topology of any β-sheet is the arrangement of all the strands in the sheet, namely the strand order within the hydrogen bonding of the sheet. This hydrogen bonding-directed strand order is also called strand connectivity. In the case of a jelly roll motif, this strand connectivity has an order of 1-2-7-4-5-6-3-8.
Coiled-coil As a special case of helix bundles, coiledcoil refers to two or more helices intertwined with each other to form a supercoiled helical structure. The helices involved in a coiled-coil configuration are normally long, with certain characteristic repeats in their amino acid sequences, and are usually parallel rather than antiparallel to each other. In most cases, the participating helices of a coiled-coil are from
β-sandwich β-sandwich refers to a set of β-strands arranged into two β-sheets packed face-to-face against each other (Fig. 17.1.1J). The two sheets can be made out of parallel, antiparallel, or mixed strands. In many β-sandwiches, the strands of the two sheets are orientated either parallel/antiparallel or approximately orthogonal to each other. A common feature in some types of β-sandwiches is strand switching, which involves an edge strand that hydrogen bonds to one sheet at its N-terminal end and the other at its C-terminal end. Mixed α/β sandwich Aside from the β-sandwich, other layered secondary structure assemblies include α/β, α/β/α, and β/β/α sandwiches. For example, an α/β/α-sandwich is defined as structures with one layer of β-sheet (middle layer) sandwiched in between two layers of α-helix. Both layers of helix pack closely against the middle β-sheet layer. The α//β/α sandwich is commonly ob-
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served in nucleotide-binding proteins (see DNA-Binding Motifs and Domains; also see RNA-Binding Structural Motifs and Domains). β-barrel β-barrels are assemblies of β-strands folded into closed barrel-like structures when viewed from the ends of the strands (Fig. 17.1.1K). The size and shape of a barrel vary widely in protein structures. A given barrel can have as few as five strands, such as in the src homology SH3 domain (see Modular Domains Involved in Signal Transduction), or as many as 22 strands, as observed in the structures of bacterial outer membrane proteins FepA and FhuA (see β-Barrel Membrane Proteins). In addition, their strands can be arranged in parallel, antiparallel, or mixed orientations. α/β-barrel In contrast to a β-barrel, which consists entirely of β-strands, an α/β-barrel is made out of alternating α and β structures. A typical example of an α/β-barrel fold is found in the structure of triosephosphate isomerase (TIM; see The TIM-Barrel Fold). The structure has eight repeats of an alternating unit consisting of a β-strand followed by an α-helix (Fig. 17.1.1L). It is organized such that the eight β-strands form an inner β-barrel which is then surrounded by eight α-helices. α/β-barrels are frequently observed in enzyme structures.
Overview of Protein Structural and Functional Folds
β-propeller β-propellers consist of a disk-shaped circular arrangement of four to eight small β-sheets around a central tunnel (for reviews see Fulop and Jones, 1999; Jawad and Paoli, 2002). Each blade of the propeller comprises a twisted, four-stranded antiparallel sheet with β-meander topology (i.e., one strand directly follows another, forming an array of β-hairpins; also see Lipocalins) that packs face to face with neighboring blades, commonly through hydrophobic interactions (Fig. 17.1.1M). β-propeller blades generally align with 40- to 60-residue sequence repeats that start with either the second, third, or fourth β-strand, and continue into the next sheet for a total of four strands. Ring closure usually occurs with the N- and C-terminal β-strands coming together within the final propeller blade in an association sometimes referred to as ‘molecular Velcro’. β-propeller functions vary widely and include enzymatic catalysis (e.g., collagenase, galactose oxidase, sialidase), ligand binding (e.g.,
tachylectin-2, haemopexin), signaling (e.g., seven-bladed G protein β-subunit), proteinprotein interaction (e.g., integrin extracellular segment), and scaffolding functions (e.g., constitutive photomorphogenic 1 (COP1) protein). Well known repeating sequence motifs observed in β-propeller proteins include the WD motif (also called WD40), the kelch motif, the RCC1 repeat, the TolB repeat, the YWTD repeat, the aspartate box, the tachylectin-2 repeat, and the tryptophan docking motif (Fulop and Jones, 1999).
WEB-BASED STRUCTURAL BIOINFORMATICS The rapid growth of protein structural data along with the simultaneous growth of the World Wide Web is facilitating the assembly and posting of numerous structural biology databases for researchers throughout the world. Concurrently, web servers that provide interactive computations such as protein structure alignments, graphics-based structural analysis, and pattern matching are also being developed at a swift pace. This section summarizes several helpful bioinformatics websites that relate to protein structure, classification, and structural folds; however, due to the large number of sites available, some excellent ones are not discussed in this unit. Instead, compiled here are some representative web sites that appear suited for a broad range of researchers interested in structural biology; sites focused on more narrow topics or technical issues are not discussed. The databases are arranged into the following categories: 1. Identification and annotation of protein domains and/or folds. Describes websites that serve as databases of protein folds and motifs and also allows new protein sequences submitted by the user to be scanned for domain/motif profiles. 2. Protein structure databases. Discusses worldwide databases for protein structures and their services. 3. Molecular graphics programs for PC and Macintosh personal computers. Brief descriptions and listings of freely available graphics programs for displaying and manipulating macromolecular structures on PCs and Macintosh (Mac) computers. 4. Classification of protein structures. Databases that classify protein folds and domains by various protocols. 5. Structural alignments. Databases that provide structural alignments of proteins as well
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as servers that align user-submitted structures interactively. For more detailed descriptions of the databases as well as instructions on how to use them, the reader is referred to Current Protocols in Bioinformatics (Baxevanis et al., 2004).
Identification and Annotation of Protein Domains and/or Folds There are now numerous database projects with interactive web sites that serve to categorize proteins, or portions of proteins, according to families, domains, motifs, or even functional sites using various types of sequence clustering methods or profile matching, usually using Hidden Markov Models (HMMs). These database web sites enable the user to upload sequences to be scanned interactively for real time diagnostic classification and annotation. Most sites also allow the user to browse the site’s database to view annotation on a particular type of domain or motif, including a short descriptive paragraph and key literature references, or to view the complete annotation of a particular protein in the database. Below are brief descriptions of some of the more well known databases. PROSITE This database (Bairoch and Bucher, 1994; Falquet et al., 2002) is part of the ExPASy (Expert Protein Analysis System) proteomics project (Appel et al., 1994) operated by the Swiss Institute for Bioinformatics (SIB). The PROSITE web server (http://us.expasy.org/ prosite) uses protein-sequence-profile matching to identify a wide variety of functional regions in protein sequences. In addition to some large domains, PROSITE can also scan for very short stretches of less than five residues (e.g., regions of posttranslational modifications). At the time of this writing, PROSITE included 1178 documentation entries, and 1614 different patterns, rules, and profiles/matrices. SMART The Simple Modular Architecture Research Tool (SMART; Schultz et al., 1998; Letunic et al., 2002) specializes in identifying and annotating mobile eukaryotic protein domains. The general focus of this database is on regulatory domains and extracellular modules with little emphasis on enzymatic domains. Over 650 different domains (HMMs) are currently included in the SMART database. SMART can detect signal peptides, transmembrane regions, coiled-coils, and internal repeats as well. The SMART web site
(http://smart.embl-heidelberg.de) provides numerous annotation hyperlinks for each type of domain, including the current number of proteins identified with the domain, lists of proteins containing the domain, a comprehensive sequence alignment of domain family members, a consensus sequence for the domain family, the species distribution of proteins containing the domain, the PDB entry codes (hyperlinked to PDBsum) for solved structures containing the domain, and links to the InterPro database (see below). Pfam The Pfam database is a manually curated (for the most part) collection of multiple sequence alignments and HMMs that describe various protein families (Bateman et al., 2002). It is maintained by the Wellcome Trust Sanger Institute. Approximately one-quarter of Pfam is automatically generated from the PRODOM database and is referred to as Pfam-B. Pfam is built solely from proteins within the SwissProt/TrEMBL protein sequence database releases. Over 5200 groups of homologous sequences are included in Pfam. They are each classified as either a family, domain, repeat, or motif. So-called nondomain regions, such as signal sequences, transmembrane regions, and coiled-coils, are also predicted through third p ar ty sof tware. The Pfam web site (http://www.sanger.ac.uk/Software/Pfam) and its four mirror sites provide an interface for uploading sequences and searching the database. Annotations include an InterPro abstract (when available), the protein family sequence alignment, hyperlinks to the HMMs that were used, the known species distribution, and various hyperlinks (if available) to entries in other databases such as PROSITE, RCSB PDB, SCOP, HOMSTRAD, PDBsum, COGS, SYSTERS, and InterPro. When structures are available, they can be viewed interactively on graphics programs (if installed on the user’s PC or Mac) by selecting them and clicking on the Chime or RasMol web page buttons (see below for databases and software). InterPro The InterPro database (http://www.ebi.ac. uk/interpro) is a consortium of several different database projects for annotating protein families, domains, and motifs (Apweiler et al., 2001). It is maintained by the European Bioinformatics Institute (EBI) which is part of the European Molecular Biology Laboratory (EMBL). Member databases include Pfam,
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SMART, PROSITE, TIGRFAM, PRINTS, and ProDom, in addition to the protein sequence database SwissProt/TrEMBL. InterPro has combined data from these seven member databases to form a much larger, coherent database. Redundant, overlapping entries are merged into single unique InterPro entries and hierarchical family relationships are modified accordingly. Through the InterPro web site, text searches of the InterPro database can be made or sequences can be uploaded and scanned against the database. The web page for each InterPro entry provides a brief descriptive abstract (frequently used by or linked to other databases), links to member database entries, and links to related InterPro domain entries, in addition to literature reference links. Databases not discussed Other similar databases not discussed in this section include TIGRFAM (http:// www.tigr.org/ TIGRFAMs/index.shtml), PRINTS (http:// www.bioinf.man.ac.uk/dbbrowser/ PRINTS), ProDom (http://prodes.toulouse. inra.fr/prodom/ doc/prodom.html), Blocks (http://www. blocks.fhcrc.org), and CDD (http://www.ncbi. nlm.nih.gov/Structure/cdd/ cdd.shtml).
Protein Structure Databases
Overview of Protein Structural and Functional Folds
RCSB PDB The Research Collaboratory for Structural Bioinformatics (RCSB) Protein Data Bank (PDB) is a comprehensive database of three-dimensional protein and nucleic acid structures determined by X-ray crystallography, NMR, cryoelectron microscopy, and theoretical modeling (Berman et al., 2000). It is maintained by Rutgers, The State University of New Jersey; the San Diego Supercomputer Center (SDSC) at the University of California, San Diego; and the National Institute of Standards and Technology (NIST). The PDB was established in 1971 at Brookhaven National Laboratories (BNL) with the deposition of seven structures. Since then, management of the PDB has been transferred to the RCSB (fall, 1998) and well over 20,000 structures have been deposited. The PDB has become an international resource for macromolecular structural coordinates and most peer-reviewed journals now require the deposition of coordinates in this database prior to publishing structural data. PDB structures can be easily accessed through the main web site (http://www. rcsb.org/pdb) or six other international mirror sites by searching key words or PDB entry
codes. An advanced search form is also available at http://www.rcsb.org/pdb/cgi/queryForm.cgi. From the resulting list, entries can be chosen by clicking on the EXPLORE hyperlink on the right-hand side of the page. At this point, a wealth of information is available; the following is a list of the hyperlinks available from this page (also refer to the documentation available from the web site): 1. Download/Display File. Allows downloading or displaying the coordinate file. 2. Medline. Provides the MedLine abstract of the primary publication describing the structure. 3. View Structure. Permits interactive viewing of the structure. On this page, links are available for coordinates that are formatted specifically to be parsed into interactive graphics programs such as RasMol, Chime, Swiss PdbViewer, VRML, or MICE (see below) by simply clicking on the name of the program. Links are provided on the same page that explain the downloading and installation of these programs. If there are no graphics programs installed on the user’s computer, a simple Java applet called QuickPDB is also available requiring no installation. Clicking on the QuickPDB button provides a red colored wire α-carbon trace of the structure that can be rotated, translated, or magnified (zoomed) with a mouse. The protein sequence(s) is displayed as well, enabling residues to be selected by highlighting regions of the sequence. As of early 2003, QuickPDB can display only protein structures; DNA, RNA, and ligand structure display is not supported. Additionally, prearranged static images of the molecule created by RASTER3D are available in jpg and tiff formats from the View Structure web page. 4. Structural Neighbors. Clicking on this will provide links for that particular structure within the CATH, CE, FSSP, SCOP, or VAST databases (see below). 5. Geometry. This provides considerable structural analysis of the structure including outputs from WHAT CHECK and PROCHECK (comprehensive geometric analyses), PROMOTIF (secondary structure analysis), castP (pocket and cavity analysis), LPC (ligand contact analysis), and CSU (contact analysis), in addition to a Ramachandran plot, and tabulated analyses of average bond length, bond angles, and dihedral angles. 6. Sequence Details. This link includes sequence(s) from the coordinate file in FASTA format and links to sequence databases.
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Other Existing Structural Databases MSD. The Macromolecular Structure Database (MSD; http://www.ebi.ac.uk/msd/index. html) at the European Bioinformatics Institute (EBI) manages and distributes macromolecular structural data. It collaborates closely with the RCSB PDB (see above), providing services for the deposition (AutoDep; http://autodep. ebi.ac.uk/, retrieval (OCA browser; http://oca.ebi.ac.uk/oca-bin/ocamain) and analysis (Biotech validation server; http:// biotech.ebi.ac.uk:8400) of PDB data. Other MSD services are listed below. 1. SSM. Through the Secondary Structure Matching (SSM) server (http://www.ebi. ac.uk/msd-srv/ssm/ssmstart. html) one can pick a structure from the PDB or upload a coordinate file and then scan the PDB or SCOP databases for the best Cα three-dimensional superpositions. These superpositions can then be viewed immediately using RasMol or RasTop. 2. The Ligand Chemistry Search Service. This site provides access to the Ligands and Small Molecules Dictionary of the MDS database encompassing all chemically distinct components found within the PDB (http://www.ebi. ac.uk/msd-srv/chempdb/cgi-bin/cgi.pl). Included are ligands, small molecules, hetero groups, and standard and nonstandard amino acids. Various chemical details such as formula, atom connectivity, and charge can be retrieved in addition to the coordinates of real or idealized structures. 3. Sequences of structural genomics projects. A list of structural genomics targets from The National Institute of General Medical Sciences (NIGMS)–supported research centers has been compiled, converted into FASTA format, and made accessible at this site (http://www.ebi. ac.uk/msd/projects/SGtargets.html). 4. PQS. The Protein Quaternary Structure (PQS) file server (http://pqs.ebi.ac.uk) uses an automated procedure (Henrick and Thornton, 1998) to apply crystallographic symmetry to PDB crystallographic coordinates. The resulting oligomers are analyzed by multiple criteria to predict which interchain contacts are nonspecific crystal contacts and which specifically contribute to an oligomeric complex. These results are then used to predict the most likely quaternary state of the macromolecule. Advanced or simple search forms are available to look up the results for current PDB entries; however, currently, users cannot upload files for analysis. 5. The 3D-EM Macromolecular Structure Database. This is a newly launched database for
macromolecular electron microscopy data such as volume maps, sections, structure factor files, layer-line data, figures, and textual descriptors (http://www.ebi.ac.uk/msd/iims/3D_EMdep .html). MMBD. The Molecular Modeling Database (MMDB; http://www.ncbi.nlm.nih.gov/ Structure/MMDB/mmdb.shtml) is maintained by the National Center for Biotechnology Information (NCBI), part of the National Institutes of Health (NIH). It is a monthly updated database containing experimentally determined macromolecular structure coordinates downloaded form the RCSB PDB (see above). Theoretical models are excluded. Using NCBI’s Entrez data retrieval system, one can run key word searches to find protein structures. Searching results in a list of hyperlinked PDB entry codes with brief one- or two-line descriptions. Upon choosing a particular hyperlink, an MMDB Structural Summary is presented. From this page, several hyperlinked options are available. These include: 1. The PDB web page for that particular entry. 2. A list of related PubMed references. 3. NCBI’s Taxonomy browser. 4. A three-dimensional view of the protein using NCBI’s own molecular graphics program, Cn3D (obtained by clicking View 3D Structure; program must be installed locally). 5. A list of structurally homologous proteins (referred to as structural neighbors) generated by the program VAST (Vector Alignment Search Tool; obtained by clicking on the colored bar representing the protein chain or domain of interest). Superpositions of these structures with the original structure can be viewed using the Cn3D or MAGE programs. 6. Protein sequence alignments between various domains in the structure and other sequences within the same domain family from NCBI’s Conserved Domain Database (CDD).
Molecular Graphics Programs for PC and Macintosh Platforms High-end graphics work stations are no longer needed to view three-dimensional models of macromolecular structures or perform simple manipulations in real time. This can now be accomplished with PC or Mac personal computers running any of a variety of operating systems. Below is a sample listing from the growing inventory of personal computerfriendly molecular graphics programs that are freely available from the World Wide Web. Unless otherwise noted, each program listed runs on both PC and Mac platforms. These
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programs enable the user to input PDB-formatted coordinates and then view ribbon, CPK, wireframe, or combination models of macromolecules which can be rotated, translated, or magnified. The user can then select certain residues or atoms for manipulations and output static images for future reference. Most programs have several additional features, such as selecting atom-atom distances, displaying hydrogen bonds, and viewing in a stereodisplay or dot-surface format. Despite the growing popularity of these small graphics programs, many structural biologists often use more sophisticated programs (e.g., O, Ribbons, MolScript, Raster3D, GRASP) on UNIX or LINUX platforms to model, build, and analyze structures, as well as generate figures for publication. These programs are not discussed here. RasMol RasMol is probably the most popular program for viewing macromolecular structures on PC and Mac computers (Sayle and MilnerWhite, 1995; Bernstein, 2000; http://www. bernstein-plus-sons.com/software/ rasmol). Web sites that provide access to macromolecular coordinates frequently have hyperlinks that will activate a local copy of RasMol and parse coordinates into the viewer.
Overview of Protein Structural and Functional Folds
Chime This program, currently available from MDL Information Systems (http://www. mdlchime.com/chime), was built partly from RasMol code and runs as a web browser plug-in using Netscape or Internet Explorer as the viewing platform. Like RasMol, database web pages providing macromolecular coordinates often have hyperlinks specifically for running Chime; however, unlike RasMol, Chime only reads in the coordinates supplied by the sponsoring website. There are several Chime-interfacing programs such as Protein Explorer (http://www.proteinexplorer.org) , ST ING (http://honiglab.cpmc.columbia.edu/STING/ help), and No ncovalent Bo nd Finder (http://www.umass.edu/microbio/chime/find-ncb/ index.htm) that provide enhanced or additional features to Chime, including different interfaces for program control, simple animation, automation of tasks, highlighting atomic interactions, or enabling users to upload their own PDB files for viewing. Further documentation and Chime resources are available at http://www.umass.edu/microbio/chime.
Mage This program is particularly well suited for use as a teaching or presentation tool and can run simple animation. It is also the oldest of the macromolecular graphics freeware available for personal computers (Richardson and Richardson, 1992; http://kinemage.biochem. duke.edu/kinemage/kinemage.php). Deep View/Swiss-PdbViewer This program was formerly known as simply the Swiss-PdbViewer. Deep View is one of the more sophisticated programs freely available f or use on personal computers (http://us.expasy.org/spdbv). Features include performing torsion manipulations and mutations of residues, displaying electron density maps, energy minimization, computing and displaying electrostatic potential maps, displaying molecular surfaces, and generating POV-ray scenes. It is also closely linked with the molecular modeling server, SWISSMODEL (http://www.expasy.org/swissmod). RasTop This program is adapted from RasMol; however, it provides a menu-driven graphics interface rather than the command-line interface that RasMol uses (http://www.geneinfinity. org/ rastop). As of early 2003, RasTop runs on Windows but not Mac platforms (except within environments such as virtual PC). Cn3D This program is specifically designed for NCBI’s MMDB system and reads only ASN.1formatted coordinate files (Wang et al., 2000b; http://www.ncbi.nlm.nih.gov/Structure/CN3D/ cn3d.shtml). In addition to being a moleculargraphics viewer, Cn3D also displays the protein sequence in a second window. As regions of the sequence are highlighted with a mouse, the corresponding residues are also highlighted in the structure. Furthermore, homologous sequences can be imported for alignment, enabling one to map sequence differences directly onto the three-dimensional structure. By accessing NCBI’s VAST database, the user can also view superpositions of structurally related proteins in Cn3D. MolMol This software has been developed with a special emphasis on displaying protein or nucleic acid structures determined by NMR (Koradi et al., 1996; (http://www.mol. biol.ethz.
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ch/wuthrich/software/molmol). The program runs only on UNIX or Windows platforms. PocketMol This software runs on Pocket PCs running Windows CE 3.0 or higher, using a stylus for m olecu le m an ipu lations (http://birg.cs. wright.edu/pocketmol/pocketmol.html). MolView This is a general purpose molecular viewer that runs on Mac platforms only (Smith, 1995; http://www.danforthcenter.org/smith/MolView/ molview.html). MolView can also create QuickTime movies. Qmol This viewer generates molecular surfaces and AVI movies (Gans and Shalloway, 2001; http://www.mbg.cornell.edu/shalloway/jason/ qmol.html. WebMol A JAVA based viewer that can run as an applet or a stand-alone application (Walther, 1997; http://www.cmpharm.ucsf.edu/~walther/ webmol.html).
Classification of Protein Structures Several web-accessible databases have been developed that classify the domains of known three-dimensional structures of protein molecules in a hierarchical manner. Three of the most well known databases, SCOP, CATH, and FSSP, are briefly described below. SCOP The Structural Classification of Proteins (SCOP) database classifies protein domains and multidomain regions with known structure according to evolutionary and structural relationships (Murzin et al., 1995; http://scop.mrclmb.cam.ac.uk/scop/index.html). For most entries, hyperlinks are available to run RasMol or Chime (see above) or to display a static jpg image of the molecule. This database is periodically curated/modified in a manual fashion with the aid of various automatic programs (Lo et al., 2002). SCOP release 1.61 (November 2002) includes 44,327 domains from 17,406 proteins. Structures are classified at four major levels (i.e., class, folds, superfamily, family). 1. SCOP class. Includes, alpha, beta, alpha/beta (mostly parallel β-sheets), alpha/beta (mostly antiparallel β-sheets), multidomain alpha/beta, membrane, cell surface proteins (ex-
cluding proteins of the immune system), and small proteins. 2. Folds. Classified according to the arrangement of secondary structure elements and topological connections. 3. Superfamilies. Proteins or domains with low sequence identity are grouped into superfamilies according to structural and functional similarities to reflect potential evolutionary relationships. 3. Families. Structures are clustered into families if they have sequence identities ≥30% or if they display compelling functional and structural similarity. CATH The Class, Architecture, Topology, Homologous (CATH) Superfamily database is assembled from crystallographically determined protein structures with resolution better than 3.0 Å and from NMR structures, using a largely automated set of procedures (Orengo et al., 1997, 2002; http://www.biochem.ucl.ac.uk/ bsm/cath_new/index.html). After delineating domain boundaries, protein domains are systematically classified at the five different levels listed below, forming a hierarchical tree. Domain entries can be viewed via RasMol hyperlinks (see above) or static gif images. CATH database release 2.4 (January 2002) contains 39,480 domains. 1. Class. Each domain is designated a specific class, namely alpha, beta, alpha/beta, or structures with few secondary elements. 2. Architecture. Domains are grouped by architecture according to the general three-dimensional orientations of their secondary structure elements. 3. Topology. Domains are categorized into topology or fold group by the connectivity of their secondary structure elements. 4. Homologous superfamily. Topology groups are further divided into homologous superfamiles by homology utilizing sequence, and structural and functional data. 5. Family. Final classification is by sequence families according to sequence identity. FSSP The Families of Structurally Similar Proteins (FSSP) database comprises all protein chains from the PDB (see above) that are longer than 30 residues (Holm and Sander, 1996, 1 99 8; http://www.ebi.ac.uk/dali/fssp) . Sequence redundancy is removed by generating a representative set of sequence-unique proteins, none of which have >25% sequence iden-
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tity with any others in the set. Each member of the representative set represents a group of sequence homologs with >25% sequence identity. Pairwise structural alignments are made (using the Dali program) between all members of the representative set and between each representative protein and its sequence homologs. The individual alignments, complete with RasMol links (see above) to view three-dimensional superpositions, color-coded sequence alignments, and various alignment statistics such as the root mean square deviation (RMSD), Z-scores, and percent sequence identity, are available on the web. These structural comparisons are used in a hierarchical clustering procedure to create a fold tree within FSSP for the classification of protein structures. A Z-score cutoff of ≥2 is used to define individual folds at the top of the hierarchy. Subsequent Z-score cutoffs of 4, 8, 16, 32, and 64 are used to designate further classification levels with increasing structural similarity. Moreover, another database, referred to as a supplement to FSSP and called the Dali Domain Dictionary (http://www.ebi.ac.uk/dali/domain), has recently been built in a very similar manner but focuses on protein domains rather than whole proteins (Dietmann and Holm, 2001).
Structural Alignments Several databases accessible through the World Wide Web offer comprehensive collections of structural alignments between known protein structures. Some servers also allow the user to upload coordinate files for interactive structural alignments. Proper cutoff values applied to alignment parameters result in lists of structural neighbors for each entry. Three of these database servers are briefly described below. HOMSTRAD The Homologous Structure Alignment Database (HOMSTRAD) groups protein structures determined by crystallography or NMR into several hundred families according to sequence identity and provides structural alignments within the families that can be downloaded or viewed via RasMol (see above) hyp er link s (Mizug uchi et al., 1998; http://www-cryst.bioc.cam.ac.uk/~homstrad). As of early 2003, 1033 multimember families and 3187 single member families were included in the database. Overview of Protein Structural and Functional Folds
VAST The Vector Alignment Search Tool (VAST) provides a systematic algorithm for aligning and comparing the three-dimensional structures of proteins (Madej et al., 1995; Gibrat et al., 1996; http://www. ncbi.nlm.nih.gov/ Structure/VAST/vast.shtml). More than 18,000 protein domains within the NCBI’s Molecular Modeling Database (MMD) have been compared with each other in a pairwise fashion using VAST and are stored in an NCBI database. From these results a list of ‘Structural Neighbors’ has been tabulated for each domain within the MMD. The alignments of structural neighbors can be viewed using NCBI’s Cn3D or the MAGE program (see above). Furthermore, the user can upload new structures using the VAST Search web page (http://www.ncbi. nlm.nih.gov/Structure/VAST/ vastsearch.html) to run VAST interactively and obtain a list of structural neighbors. CE The Combinatorial Extension (CE) web server uses a structural alignment method that involves aligning fragments of proteins and then searching for continuous paths of aligned fragment pairs (AFPs; Shindyalov and Bourne, 1998; http://cl.sdsc.edu/ce.html). Several options available for the user are listed below. 1. Choose a protein from the PDB (see above) and scan the entire PDB or representative proteins from the PDB for structural neighbors. 2. Upload a coordinate file and scan against the PDB 3. Calculate the structural alignment between two proteins (PDB entries or user-uploaded). 4. Perform multiple sequence alignment of proteins in the PDB using a Monte Carlo–based method. The aligned structures generated can be downloaded as a PDB file or viewed interactively using RasMol (see above), Protein Explorer (requires Chime; see above), or the web site’s own Java applet called Compare3D that requires no locally installed program.
PROTEINS INVOLVED IN THE FUNCTION OF IMMUNE SYSTEMS Immunoglobulins and Immunoglobulin-Like Superfamilies Traditionally, the term immunoglobulin (Ig) fold refers to the homologous domain structures present in the variable and constant regions of all immunoglobulins. In recent years, however, with the rapid growth of sequence and
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structure knowledge, many other proteins whose functions are not related to immunoglobulins have nonetheless been found to contain one or more Ig-like domains in their tertiary structures. Examples of such proteins include the cell surface glycoprotein receptors such as growth hormone receptor, CD4 and CD8, adhesion molecules such as cadherins and type III fibronectins, class I and class II major histocompatibility proteins, chaperone protein PapD, the transcriptional factor NF-κB, and even some enzymes such as Cu and Zn superoxide dismutase and β-galactosidase. The discovery of the Ig-like domains in many proteins not only broadens the definition of superfamily, but more importantly, demonstrates the versatility of the fold and the ability of these domains to function as modular units in many diverse systems. The immunoglobulin fold is defined as a domain that is formed from seven to nine βstrands folded into a Greek key β-sandwich (Fig. 17.1.2; also see above). The topology consists of one β-sheet formed by strands A, B, E, and D, while the opposing sheet is formed by strands C, C′, and occasionally C′′, as well as F and G. Based on the strand hydrogen-bonding pattern and the number of strands in each domain, the superfamily is further divided into five commonly observed subclasses: V, C1, C2, I, and E (Fig. 17.1.2). The V-type domain has the same topology as immunoglobulin variable (V) regions. It has nine strands with a typical strand switch occurring in the first strand, which usually breaks into two halves (A and A′) with the first half (strand A) hydrogen bonding to strand B of the B-E-D β-sheet and the second half (strand A′) hydrogen bonding to strand G of the opposing β-sheet. Strands C′ and C′′ are usually shorter than the other strands. In immunoglobulins, the antigen binding site is composed primarily of three loop regions termed complementary determining regions (CDRs). They are located between strands B and C, C′ and C′′, and F and G. The C1 type of immunoglobulin fold refers to the topology observed in the structure of constant regions of immunoglobulins. It has seven strands, of which strands A-B-E-D form one β-sheet and strands C-F-G form the other. Most members of the C2-type subclass have seven strands and differ from the C1 type by a simple switch whereby strand D hydrogen bonds with strand C in the C-F-G sheet rather than strand E in the A-B-E sheet. Hence strand D is labeled C′ in the C2-type subclass. Some members of the C2 family also have additional strands at
either the N or C termini. Based on tertiary structure similarities, there are two frequently adopted tertiary folds in the C2 class; one is similar to the C2 domain of T-lymphocyte cell surface glycoproteins CD2 and CD4, while the other is similar to the fibronectin type III domain, and hence is also referred to as the fibronectin type III-like fold, shown as C2(Fn) in Figure 17.1.2C. The I-type domains closely resemble the topology of immunoglobulinvariable domains except that they lack strands C′ and C′′. The members of the I-type subclass also have the characteristic strand switching phenomena in the first strand, as do the immunoglobulin variable domains. The E-type immunoglobulin domains also present a topology similar to those of the V type with the difference being the lack of C′′ and D strands. Table 17.1.1 summarizes all the observed immunoglobulin subclasses. Among the members of the immunoglobulin superfamily, occasionally, additional strands are observed at the N and C termini and sometimes insertion strands which lie outside the core immunoglobulin fold can be found connecting the adjacent immunoglobulin strands.
The MHC Peptide Binding Fold The three-dimensional structures of class I and class II major histocompatibility complex (MHC) molecules reveal a distinct fold associated with the peptide-binding domain. In addition to MHC molecules, the structures of neonatal Fc receptor (FcRn), hereditary haemochromatosis protein HFE, MHC-I chain homolog (MIC), UL16 binding protein (ULBP), and Rae-1 have all been shown to contain an MHC-like fold (Radaev et al., 2001b; Boyington and Sun, 2002). MHC molecules are heterodimers consisting of an α/β peptide-binding domain and two Ig C1-type domains (Fig. 17.1.3). In class I MHC molecules, the peptide-binding α1α2 domain and the α3 Ig domain reside on one chain (the heavy chain), while the second Ig domain is contributed by an associated β2m molecule (Fig. 17.1.3A). Class II MHC molecules consist of an α and β chain (Fig. 17.1.3B). The N-terminal part of each chain contributes to half of the peptide-binding domain and the C-terminal half of each chain provides a single Ig domain. The MHC Ig domains all have Ig C1-type folds (see Immunoglobulins and ImmunoglobulinLike Superfamilies). In class I MHC molecules, the peptide-binding domain is formed by the α1 and α2 regions of the heavy chain whereas
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in class II molecules it is formed by the α1 and β1 regions of both α and β chains. The peptidebinding domain consists of two α-helices (one from each region), forming the two sides of the groove and an eight-stranded antiparallel βsheet (four strands from each region), forming the floor of the groove (Fig. 17.1.3C). The size of the groove is ∼30 Å long and 12 Å wide in the middle. Both class I and II MHC molecules bind peptides in extended conformations with anchor residues buried inside pockets of the groove. The distribution of these pockets along the binding site determines the peptide selectivity of a particular allele of class I or class II molecules. Class I MHC molecules are restricted to binding peptides of 8 to 11 residues, whereas class II MHC molecules bind longer peptides of 15 to 24 residues. In contrast, none of the MHC-like proteins, such as FcRn, HFE, MICs, ULBPs, and Rae-1s associate with peptides. Their putative peptide-binding grooves are narrower or even closed compared to the MHC peptide-binding groove (Table 17.1.1).
The C-Type Lectin-Like Receptor Fold In addition to immunoglobulin-like superfamilies, many recently identified immunoreceptors are members of the C-type lectin-like superfamily. Examples include mannose-binding protein, FcεRII (CD23), CD94, members of NKG2 family receptors, Ly49 receptors, and DC-SIGN. Their structures resemble that of C-type lectin, a calcium-dependent carbohydrate-binding protein (see C-type Lectins). What is unique to many of these C-type lectinlike receptors is the absence of a functional calcium-binding site and their ability to form a common disulfide-bonded dimer (Boyington et al., 1999). Table 17.1.1 lists the known structures of C-type lectin-like receptors.
Structure of Immunoreceptor-Ligand Complexes
Overview of Protein Structural and Functional Folds
The goal of structural immunology in recent years has been to define molecular structures of immunoreceptor-ligand complexes rather than revealing new protein folds. The notable milestones include the publications of several T cell receptors and their cognate MHC-peptide complexes (Garboczi et al., 1996; Garcia et al., 1998; Reinherz et al., 1999)—i.e., T cell coreceptor CD8 or CD4 and their class I or II MHC complexes (Gao et al., 1997; Wang et al., 2001), T cell coreceptor CTLA-4 and ligand B7 complex (Stamper et al., 2001; Zhang et al., 2001), NK cell killer immunoglobulin-like receptor (KIR) and its MHC ligand complexes (Fan et
al., 2001; Boyington et al., 2000), NK cell activating receptor NKG2D and its ligand complexes (Li et al., 2001; Radaev et al., 2001b; Li et al., 2002b), and Fc receptors in complex with their Fc ligands (Garman et al., 2000; Radaev et al., 2001a; Sondermann et al., 2000).
Proteins in the Complement System The complement system is composed of a large number of plasma proteins which participate in an innate immune response to bacteria. Antibodies, mannose-binding lectins, or other complement components binding to the bacterial cell surface can initiate a cascade of complement reactions ultimately leading to recruitment of inflammatory cells, and the opsonization and killing of bacterial pathogens. The following section reviews the structural folds of fragments of several complement components (Table 17.1.2). C3 fragment C3d Complement protein C3 is a central component in the opsonization of bacteria. One of its natural cleavage products, C3dg can act as a ligand for the B cell receptor CR2 (CD21) when covalently attached to the pathogen surface. The three-dimensional structure of C3d, a protease-resistant fragment (residues 996 to 1303) of C3dg, reveals an α/α barrel composed of twelve α helices (Fig. 17.1.4; Nagar et al., 1998). The barrel is composed of two concentric rings of six α helices each. Each ring is made of parallel helices inclined slightly from the main axis of the barrel. The topology of the molecule causes consecutive helices to alternate from outside to inside with the orientation of the outer ring helices antiparallel to those of the inner ring. This same fold is also observed in enzymes such as glucoamylase, endoglucanase, and the β subunit of protein farnesyltransferase. The three residues essential for covalent attachment to the pathogen, Cys17 (mutated to Ala in the crystal structure to avoid incorrect disulfide bond formation), Gln20, and His133, are situated at the rim of one end of the barrel (right panel of Fig. 17.1.4A). C5a C5a is a 74-residue glycoprotein derived from C5 and is very potent in mediating local inflammatory responses within the complement cascade. The solution NMR structures show a five α helix antiparallel bundle (Fig. 17.1.4B; Zhang et al., 1997). Mutational analysis indicates that the C-terminal twelve residues, including the C-terminal helix, contribute
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to agonist activity and receptor binding (Zhang et al., 1997). Complement factor D Complement factor D is a serine protease which cleaves complement factor B only when complexed with C3b. The crystal structure shows a typical trypsin fold (see Protease Folds) consisting of two nearly identical sixstranded antiparallel β barrels with Greek key topology (see above), two flanking α helices, and an active site situated between the two barrels (Fig. 17.1.4C; Narayana et al., 1994). The active site is very unusual in that the Asp and His residues of the canonical Asp-His-Ser triad are misoriented from the conformation that is favorable for catalytic activity of a serine protease. It is proposed that complement factor D activity is regulated by a reversible inducedfit mechanism which activates the enzyme only in the presence of its natural substrate, factor B/C3b. Mutation and X-ray studies indicate that three residues, Ser 94, Thr 214, and Ser 215 are responsible for the altered conformation of the active site relative to other serine proteases (Kim et al., 1995a). Complement regulatory protein CD59 CD59 (also known as MEM-43, MIRL, H19, MACIF, HRF20, and protectin) is a glycophosphatidylinositol (GPI)-anchored glycoprotein that functions as a potent inhibitor of complement-mediated cell lysis and may also play a role in T-cell activation. CD59 belongs to a group of cysteine-rich cell surface molecules known as the Ly6 superfamily which includes the urokinase plasminogen activator receptor, mouse thymocyte antigen ThB, and murine Ly6 antigens. Two NMR solution structures of the extracellular portion (residues 1 to 70) of the protein reveal a disk-shaped molecule with five disulfide bridges comprising a threeand two-stranded antiparallel β-sheet abutting each other, and a two-turn α-helix packed against one side of the larger β-sheet (Fig. 17.1.4D; Fletcher et al., 1994; Kieffer et al., 1994). This structure is also very similar to those of snake venom toxins (see Small SingleSubunit Toxins) with which the Ly6 superfamily shares sequence homology. Complement control protein modules Complement control protein (CCP) modules, also known as complement protein (CP) modules, short consensus repeats (SCRs), or sushi repeats, are small domains of ∼60 amino acids containing four invariant cysteines form-
ing intramodule disulfide bridges and several other highly conserved residues. These modules are present in tandem arrays in many complement regulatory proteins in addition to a number of cell surface proteins. Three-dimensional structures have been determined for the fifth, fifteenth, and sixteenth CCP domains of factor H, CCP modules 3 and 4 of a Vaccinia virus complement control protein mimic, and CCP modules 1 and 2 from the complement cofactor CD46 (Table 17.1.2; Norman et al., 1991; Barlow et al., 1992, 1993; Wiles et al., 1997; Casasnovas et al., 1999). The domain structure consists of an elongated, twisted βsandwich formed from a three- or four-stranded antiparallel β-sheet and a two-stranded antiparallel β-sheet (Fig. 17.1.4E). Although the known structures of SCR modules all have essentially the same backbone fold, differences in the number of β strands between various modules arise from differences in hydrogen bonding patterns. Structures containing two tandem CCPs show an elongated head to tail arrangement with elbow angles that vary significantly from structure to structure.
PROTEINS INVOLVED IN SIGNAL TRANSDUCTION PATHWAYS Cytokines Four-helix bundles Members of the hematopoietic cytokines possess a common four-helix bundle topology in their structures. The four helices (labeled a, b, c, d) always pack in a left-handed up-updown-down manner, with helices a and b, and c and d forming two pairs of parallel helices juxtaposed in opposite directions (Sprang and Bazan, 1993; Wlodawer et al., 1993). The position of the four helices and the packing angles between them are generally well conserved. The superfamily is further divided into longchain, short-chain, and interferon (IFN) γ–like families of cytokines. Long-chain family. Members in this family include growth hormone (GH), granulocyte colony-stimulating factor (G-CSF), erythropoietin (EPO), leukemia inhibiting factor (LIF), IL-6, IL-12 α-chain, leptin, and ciliary neurotrophic factor. The four helices of this family are each ∼25-residues long (Fig. 17.1.5A and D). Short-chain family. The short-chain family includes most of the interleukins (i.e., IL-2, -3, -4, -5, -7, -9, and -13), granulocyte/macrophage-colony-stimulating factor (GM-CSF),
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macrophage-colony-stimulating factor (MCSF), and stem-cell factor (SCF). The helices are shorter and uneven in length (eight to fifteen amino acids long) compared to members of the long-chain family. Another signature characteristic of short-chain hematopoietic cytokines is the presence of a pair of short, twisted antiparallel β-strands, one in the loop connecting helices a and b, and the other in the loop connecting helices c and d (Fig. 17.1.5B and D). Many of them also have disulfide bridges in their structures; however, the locations of these disulfide bonds are not well conserved. Interferon γ–like. This family includes IFNγ and IL-10. Both contain six helices (a, b, c, d, e, and f) and function as homodimers (Ealick et al., 1991; Zdanov et al., 1995). The dimerization is particularly interesting in this family. The two domains of the dimer are formed by swapping the last two helices (e and f) between the two monomers. The four-helix bundle forming helices in each domain are made out of helices a, c, and d from one monomer, and helix f from the other (Fig. 17.1.5C and E). β-Trefoil Members of the β-trefoil family include not only cytokines such as interleukin-1α, interleukin-1β, and fibroblast growth factor, but also proteins such as soybean trypsin inhibitor, ricin B, amaranthin, and hisactophilin, which display the same fold (Priestle et al., 1989; Graves et al., 1990; Clore and Gronenborn, 1991; Eriksson et al., 1991; Zhang et al., 1991; Zhu et al., 1991). The overall structure is a sixstranded closed β-barrel (A-D-E-H-I-L) sitting atop three β-hairpins (B-C, F-G and J-K). The β-trefoil has internal pseudo three-fold symmetry (Fig. 17.1.6A) relating three four-stranded antiparallel β-sheets. Each β-sheet is made of four consecutive strands. For example, the three β-sheets of IL-1β are comprised of strands L-A-B-C, D-E-F-G, and H-I-J-K, respectively.
Overview of Protein Structural and Functional Folds
The cystine-knot growth factor superfamily The recently solved crystal structures of four different growth factors, nerve growth factor (NGF), transforming growth factor-β (TGF-β), platelet-derived growth factor (PDGF), and human chorionic gonadotropin (hCG), have revealed a common overall topology among these functionally diverse protein families (Daopin et al., 1993; Murray-Rust et al., 1993; Murzin et al., 1995; Sun and Davies, 1995). These proteins share very little sequence homology. Nevertheless, they all have an unusual arrangement of six cysteines that are disulfide linked
to form a knotted conformation and this core structure, the cystine-knot, is flanked by four strands of an antiparallel β-sheet (McDonald et al., 1991; Oefner et al., 1992; Daopin et al., 1993; Schlunegger and Grutter, 1993; Lapthorn et al., 1994; Wu et al., 1994). Fig. 17.1.6B shows the crystal structure of human TGF-β2 with the common flanking β-strands (1, 2, 3, and 4) shown in green and the six disulfidelinked cysteines shown in ball-and-stick models. Outside the four structurally conserved β-strands, the helices and loops observed in the structure of TGF-β2 are not conserved in NGF, PDGF, and hCG. The three disulfide bonds are formed between the six cysteines of TGF-β2 as follows: Cys 15–Cys 78, Cys 44–Cys 109, and Cys 48–Cys 111, creating a linkage pattern of I–IV, II–V, and III–VI among the six cysteines. A topological knot is formed by threading disulfide I–IV through a ring formed with disulfides II–V and III–VI as its two sides, and the peptides in between cystine II and III, and V and VI as its other two sides. Another common structural feature among the cystine-knot cytokines is that the active forms of these proteins are always dimers, either homo- or heterodimers; however, despite the overall topological similarity between the monomers, each family of growth factors form their own distinct dimers with a unique dimer interface that is characteristic to each family of growth factors. EGF-like domain EGF-like domains are often observed in tandem repeats in many proteins involved with a receptor-related signal transduction component (Groenen et al., 1994). Members of this family include epidermal growth factor, transforming growth factor α, the EGF-domain of E-selectin, Factor X, Factor IX, plasminogen activator, prostaglandin H2 synthase-1, and thrombomodulin. An EGF module has ∼50 amino acids and is stabilized by three disulfide bonds (Graves et al., 1994; Kohda and Inagaki, 1992; Rao et al., 1995). Its structure elements consist of two small β-sheets: a three-stranded antiparallel β-sheet close to the N terminus and a two-stranded antiparallel β-sheet near the C-terminus (Fig. 17.1.6C). The disulfide bonds are formed between cysteines I–III, II–IV, and V–VI in sequence. Chemokines There are two subfamilies of chemokines, the CXC type and the CC type (also known as the α and β type, respectively), referring to the spacing between the first two conserved cyste-
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ines in sequence. They are potent chemotactic activators for neutrophils (CXC type), monocytes, and lymphocytes (CC type). The structures of several of these chemokines are available, such as interleukin 8 (IL-8; Baldwin et al., 1990; Clore et al., 1990) of the CXC type and neutrophil-activating peptide-2 (NAP-2) of the CXC type (Malkowski et al., 1995), as well as the macrophage inflammatory protein-1β (MIP-1β), RANTES, and monocyte and chemoattractant proteins (MCP-1 and -3) of the CC type (Table 17.1.3; Lodi et al., 1994; Skelton et al., 1995; Handel and Domaille, 1996; Lubkowski et al., 1997). The two disulfide bonds are formed between the four conserved cysteines in a pattern of I–III and II–IV. Both subfamilies of chemokine are composed of dimeric proteins which share the same structural fold at the monomer level, but they differ drastically in their dimerization (Fig. 17.1.6D). The basic fold consists of a C-terminal α-helix packing against a three-stranded antiparallel β-sheet. The CXC chemokines dimerize on the edge of the first β-strand, forming an extended six stranded antiparallel β-sheet with the two C-terminal helices packing on one side of the sheet. In contrast, the CC chemokines form the dimer using their N-terminal loop region, resulting an elongated dimer with the β-sheets of the monomers opposing each other (Fig. 17.1.6E).
Receptors TNF receptor family and its associated proteins The structure of a human 55 kDa tumor necrosis factor (TNF) receptor shows a unique fold consisting of four homologous cysteine repeat domains (Banner et al., 1993). There are no helices and very few regular β-sheets in its secondary structure (Fig. 17.1.7A). Instead, the fold is composed mostly of loops and β-hairpins. Upon complex formation with TNF, there are three receptor molecules symmetrically bound to a TNF trimer. In addition to TNF, the structure of a TNFassociated protein, TRAF2 (Park et al., 1999), and the structure of a TNF receptor superfamily member, TRAIL, as well as a TRAIL-DR5 complex (Cha et al., 1999; Mongkolsapaya et al., 1999) have also been determined. Receptors for cystine-knot growth factors The structures of the ligand-binding domains for several cystine-knot growth factor receptors have been solved to date. They in-
clude a type II murine activin receptor (ActRII; Greenwald et al., 1999), a type I human bone morphogenic protein receptor (BRIA; Kirsch et al., 2000), a type II human transforming growth factor receptor (TBRII; Fig. 17.1.7B; Hart et al., 2002), the TrkA subunit of a human nerve growth factor (Wiesmann et al., 1999), and domain 2 of a vascular endothelial growth factor (VEGF) receptor, Flt-1 (Wiesmann et al., 1997; Table 17.1.3). Two of these, TrkA and Flt-1, are tyrosine kinase receptors, whereas the other three are serine/threonine kinase receptors. Structurally, the ligand-binding domains of TrkA and Flt-1 are members of the immunoglobulin superfamily whereas the ligandbinding domains of ActRII, BRIA, and TBRII display a characteristic three-finger toxin fold. The latter three are also part of the so-called TGF-β receptor superfamily. Nuclear and nuclear transport receptors The nuclear receptor superfamily is composed of steroid receptors, the thyroid hormone receptor, retinoic-acid receptor, and a number of orphan receptors (Weatherman et al., 1999). All have two domains, an N-terminal DNAbinding domain that functions as a transcription activator, and a C-terminal ligand-binding domain that is specific for different ligands. The DNA-binding domain, rather conserved among the members of the nuclear-receptor superfamily, uses two Zn finger motifs to recognize short DNA repeats (see DNA-Binding Motifs and Domains). The ligand-binding domain is more variable than the DNA-binding domain, but nevertheless displays a similar fold of mostly α-helical segments among its members. To date, the ligand-binding domains of thyroid hormone receptor (TR; Fig. 17.1.7C; Darimont et al., 1998), retinoic-acid receptor (RAR; Renaud et al., 1995), estrogen receptor (ER; Shiau et al., 1998), progesterone receptor (PR; Williams and Sigler, 1998), and peroxisome proliferator-activated receptor (PPAR; Xu et al., 2002) have been determined (Table 17.1.3). Importin-α and -β, also known as karyopherin-α and -β, are nuclear import receptors that recognize nuclear localization signals and bind and transport proteins from the cytoplasm to the nucleus through the nuclear pore complex. The structures of importin-α and -β consist of homologous ARM and HEAT repeats (Table 17.1.3; Conti et al., 1998; Chook and Blobel, 1999; Cingolani et al., 1999; Kobe, 1999). The structural fold of ARM and HEAT repeats are reviewed in the section below (see Structural Biology
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Modular Domains Involved in Signal Transduction). Integrins and adhesion-related proteins The structures of the I-domain from CD11b and CD11a integrin α-chains show that these domains possess a flavodoxin-like fold similar to members of the dinucleotide-binding family of proteins (Lee et al., 1995; Qu and Leahy, 1995). The tertiary structure consists of a sixstranded mostly parallel β-sheet with only one antiparallel strand on one edge of the sheet (Fig. 17.1.7D; Table 17.1.3). The connectivity of the strands follows a 3-2-1-4-5-6 order. Flanking the central β-sheet are several α-helices on each side. A bound divalent metal ion at the end of the β-sheet is postulated to play an important role in its activity. The structure of an intact αVβ3 integrin exhibits a multidomain organization of the molecule (Xiong et al., 2001). The adhesion domain consists of a β-propeller domain from the αV-chain and the I-domain from the β3-chain. Following the adhesion domain, the αV-chain has three immunoglobulin-like domains, whereas the β3-chain possess four EGF-like modules. β-catenins are cytosolic proteins essential for cell adhesion and signal transduction. The structure of β-catenin consists of a central domain of twelve armadillo (ARM) repeats (see Modular Domains Involved in Signal Transduction), which mediate binding to cadherins and Tcf family transcription factors (Graham et al., 2000; Huber and Weis, 2001). Scavenger receptor cysteine-rich domain The cysteine-rich domain of the scavenger receptor is a conserved domain of 110 amino acids. It is found widely in cell surface receptors and secreted proteins that participate in innate immune functions. The structure of a scavenger receptor cysteine-rich domain (SRCR) from the Mac-2 binding protein has been determined (Hohenester et al., 1999). The fold consists of a curved six-stranded β-sheet cradling an α-helix. There are three conserved disulfide bonds (Fig. 17.1.7E; Table 17.1.3): one bridging between strand C and the EF loop (Cys 31-Cys 95), one between the α-helix and strand F (Cys 44-Cys 105), and one within the EF loop (Cys 75-Cys 85).
Overview of Protein Structural and Functional Folds
Glutamate receptor Glutamate is an important neurotransmitter of mammalian central nervous systems mediating long-term potentiation/depression, learning, and memory. There are two forms of glu-
tamate receptors, the metabotropic (mGluR) and ionotropic (iGluR) glutamate receptors. While metabotropic glutamate receptors are members of G-protein-coupled seven transmembrane receptors, ionotropic glutamate receptors contain ligand-gated channels. The structures of the extracellular ligand-binding domains of mGluR and iGluR are similar (Armstrong et al., 1998; Kunishima et al., 2000), with two domains each assuming an α/β fold topology (Fig. 17.1.7F; Table 17.1.3). Glutamate is bound in an interdomain crevice. Transferrin receptor Transferrin receptors (TfR) facilitate iron uptake in vertebrates by binding to iron-loaded transferrins on the cell surface, and then internalizing and releasing the iron in an endosome. The TfR-bound apo-transferrin then recycles to the cell surface where it is exchanged with iron-bound transferrin. The crystal structure of a human TfR shows a receptor dimer with each monomer consisting of three domains (Fig. 17.1.7G; Lawrence et al., 1999). The first protease-like domain has a fold similar to that of the carboxy- or aminopeptidases with a central seven-stranded β-sheet flanked by α-helices. The second apical domain folds into a β-sandwich similar to domain four in aconitase. The third domain is all α-helical and forms an irregular four-helical bundle (Table 17.1.3).
Modular Domains Involved in Signal Transduction Domains that bind phosphotyrosine SH2 domains. SH2 domains were initially identified in the oncoprotein v-Fps, and subsequently in the Src and Abl tyrosine kinases (Pawson et al., 2001). The name is derived from the designation Src homology domain 2. The function of this module is to bind phosphotyrosine peptides (Table 17.1.4). Through this very specific phosphorylation-dependent interaction, various types of cellular signaling processes can be controlled, such as enzyme regulation, sequestering or recruiting of specific proteins, linking proteins together through adaptor molecules, and protein multimerization. Most SH2 domains are involved in some aspect of the cytoplasmic tyrosine kinase signaling pathways (for reviews of the SH2 domain, see Kuriyan and Cowburn, 1997; Pawson et al., 2001). Examples of proteins with SH2 domains include protein tyrosine kinases (PTKs), phosphatases (PTPases), a subunit of phospholipase C (PLC-γ1), the p85 subunit of
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phosphatidylinositol-3OH kinase, the cytoplasmic tail of cell surface receptor proteins, and adaptor proteins like Grb2; however, as of this writing, the total number of SH2-containing proteins is well over 1100 according to the SMART database (see Web-Based Structural Bioinformatics and http://smart.embl-heidelberg.de/ browse.shtml). The fold of the ∼100 residue SH2 domain is characterized by an α/β barrel-like sandwich with a central core formed by a four-stranded antiparallel β-sheet followed by a smaller three-stranded antiparallel β-sheet (Fig. 17.1.8A). The β-barrel is flanked by two α-helices, one on each side of the larger β-sheet (Booker et al., 1992; Overduin et al., 1992; Waksman et al., 1992). A phosphopeptidebinding site is located on a surface formed by the fourth strand of the central β-sheet, the first helix, the loop between strands 5 and 6, and the loop at the end of helix 2. The phosphate moiety of the phosphotyrosine forms an ion pair with an arginine residue from the second strand of the central β-sheet. Another arginine from the first helix and a lysine from strand 4 appear to have favorable interaction with the phosphotyrosine ring. The peptide binding specificity of SH2 domains are defined largely by amino acids at the +1, +2, and +3 site of the phosphotyrosine peptide. PTBs. Phosphotyrosine-binding domains (PTBs), also known as protein-interaction domains (PIs), function as signaling domains which bind to proteins containing the consensus sequence Asn-Pro-X-(pTyr/Tyr). For reviews, see Yaffe (2002b) and Yan et al. (2002). Most PTBs target sequences with a phosphorylated tyrosine; however, some recognize nonphosphorylated sequences. Furthermore, some PTB domains are reported to recognize peptide ligands containing no tyrosine residue or even phospholipid ligands (Yaffe, 2002b; Yan et al., 2002). Despite functional diversity within this family, structural homology is strongly conserved and many members display very high ligand specificity. The PTB fold consists of a seven-stranded antiparallel β-sandwich flanked by an α-helix (Fig. 17.1.8B; Zhou et al., 1995, 1996; Eck et al., 1996). Remarkably, this is the same fold observed for the pleckstrin homology (PH) domain and the EVH1 domain (see Polyproline Binding Domains). In the case of the PTB domain, the peptide ligand generally forms an additional β-strand between β-strand 5 and the α-helix, essentially extending the second βsheet (Yan et al., 2002). First identified in the
adaptor protein SCH (Blaikie et al., 1994; Kavanaugh and Williams, 1994; van der Geer et al., 1995) and the insulin receptor substrate 1 (IRS-1; Gustafson et al., 1995), over 430 PTB domains are now listed in the SMART database (see Web-Based Structural Bioinformatics and http://smart.embl-heidelberg.de/browse.shtml). Phosphoserine- and phosphothreonine-binding domains Forkhead-associated domains (FHA). The forkhead-associated domain (FHA) is a small modular phosphothreonine-binding domain widely found, from prokaryotes to eukaryotes, in numerous types of multidomain proteins, including transcription factors, kinases, phosphatases, kinesins, RNA-binding proteins, and some metabolic enzymes (Li et al., 2000; Durocher and Jackson, 2002). Initially identified by Hofmann and Bucher (1995) in the family of forkhead-associated transcription factors, this domain has since been found on over 400 different proteins (http://smart.embl-heidelberg. de/browse.shtml). In addition to recognizing phosphothreonine, some FHA domains are reported to bind phosphotyrosine (Liao et al., 1999; Wang et al., 2000a) or even unphosphorylated peptides (Li et al., 2002a). The structure of the FHA domain (Fig. 17.1.8C) comprises a ∼100 residue eleven-stranded β-sandwich (Liao et al., 1999; Durocher et al., 2000; Wang et al., 2000a). Some FHA domains have small helices within the loop regions as well. Structurally, this domain is most closely related to the MH2 domain of the SMAD family that also functions as a protein interaction module. Interestingly, the FHA fold includes nearly twice as many residues as the 55– to 75–amino acid FHA homology region initially studied. This FHA homology region covers only β-strands 3 to 10 on one face and includes the ligand recognition site (Durocher and Jackson, 2002). The phosphothreonine-containing peptide ligand binds in an extended conformation across three loops (β3/4, β4/5 and β6/7) along the end of one face of the β-sandwich. 14-3-3 proteins. 14-3-3 proteins constitute a large array of multifunctional phosphoserine/phosphothreonine-binding proteins that play significant roles in signal transduction for numerous eukaryotic-cell processes, including regulation of gene expression, cell-growth control, apoptosis, and neuronal development (Fu et al., 2000b; Yaffe, 2002a). Many different types of proteins are targeted by 14-3-3 proteins, including kinases, acetylases, acetyl transferases, hydroxylases, cell surface recep-
Structural Biology
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tors, cytoskeletal and scaffolding proteins, small G-proteins, and transcription factors. Indeed over 100 different proteins have been reported to interact with 14-3-3 proteins (Yaffe, 2002a). The name 14-3-3 refers to the elution position of these proteins from a DEAE cellulose column and the subsequent gel electrophoresis migration pattern during isolation from bovine brain by Moore and Perez in 1967. 14-3-3 proteins are relatively abundant, acidic proteins found in eukaryotic organisms from fungi to plants and mammals. In mammals, seven different isoforms have been characterized, referred to as β, γ, ε, η, σ, τ, and ζ. They function as homo- and heterodimers with a monomeric molecular weight of ∼30 kDa. The first three-dimensional structures of 14-3-3 determined in 1995 revealed a crescent shaped dimer (Fig. 17.1.8D) composed of a bundle of antiparallel α-helices (nine per monomer; Liu et al., 1995; Xiao et al., 1995). The inner concave surface (or central channel) is more negatively charged and more highly conserved than the outer surface. Subsequent crystal structures of peptide-ligand complexes revealed an amphipathic binding groove in each monomer within the central channel that binds peptides in an extended conformation (Yaffe, 2002a). The recent crystal structure of 14-3-3 ζ complexed with serotonin N-acetyltransferase bound to a bisubstrate analog in addition to isothermal titration calorimetry (ITC) and activity studies provide intriguing evidence that 14-3-3 binding structurally modulates serotonin N-acetyltransferase such that substrate binding affinity and activity increase (Obsil et al., 2001). Thus, 14-3-3 proteins are not simply protein interaction domains, but can also regulate enzymatic activity.
Overview of Protein Structural and Functional Folds
Polyproline-binding domains SH3 domains. SH3 (src homology 3) domains are protein interaction modules of 50 to 75 residues that recognize proline-rich sequences. Like SH2 domains, SH3 domains are perhaps best known for their role in the regulation of tyrosine kinases; however, they are also found in numerous other proteins, including the adaptor proteins Grb2 and Crk, as well as PTPases such as the γl subunit of phopholipase C (PLC-γl), certain cytoskeletal proteins such as myosin-1B and fodrin, and the yeast actinbinding protein ABP-1. The SMART database now lists over 2200 proteins containing SH3 domains (see Web-Based Structural Bioinform atics and http://smart.embl-heidelberg. de/browse.shtml).
The core of an SH3 domain consists of two perpendicular antiparallel β-sheets folded into a β-barrel like structure. One sheet is formed with strands B, C, and D, the other with A and E (Fig. 17.1.9A; Musacchio et al., 1992; Yu et al., 1992). The relatively flat, nonpolar recognition site is formed primarily by two loops of variable length, one between strand A and B (the so-called RT loop), and one between B and C (the so-called n-Src loop). The core binding motif is Φ-Pro-X-Φ-Pro, where Φ is a hydrophobic residue and x is any residue. Each of the ΦP dipeptides occupies a conserved hydrophobic binding pocket. An interesting feature of the SH3 domain is that it recognizes its ligands in either of two orientations, depending on the placement of positively charged specificity residues flanking the Φ-Pro-X-Φ-Pro sequence. For reviews detailing the structural and functional aspects of SH3 domains see Cesareni et al. (2002), Kuriyan and Cowburn (1997), and Mayer (2001). WW domains. WW domains (also known as WWP or rsp5 domains) are very small modules consisting of only 35 to 40 residues folded into a three-stranded antiparallel β-sheet (Fig. 17.1.9B; Macias et al., 1996). As its name suggests, this domain contains a pair of signature tryptophan residues spaced ∼20 to 22 residues apart. Additionally, there is a highly conserved proline three residues beyond the second tryptophan. WW domains have been observed throughout eukaryotes, including fungi, plants, nematodes, and various vertebrates. They function specifically in binding proline-rich sequences and have been classified into four groups based on ligand specificity. Group I domains bind to PPxY-containing sequences, group II bind PPLP sequences, group III bind polyproline sequences flanked by Arg or Lys, and group IV domains bind to ligands containing phospho-(S/T)P. Structural solutions of several ligand complexes show that the polyproline ligand interacts with the convex face of the β-sheet. For reviews, see Macias et al. (2002) and Sudol and Hunter (2000). EVH1 domains. Enabled/vasodilator-stimulated phosphoprotein homology 1 (EVH1) domains (also known as WH1 domains) are protein-interaction modules critical for a diverse array of signal transduction pathways related to actin cytoskeleton remodeling (Ball et al., 2002). EVH1 domains of ∼115 residues are widely present in eukaryotic species from fungi to mammals and are generally part of large multidomain proteins. The EVH1 fold comprises a small seven-stranded antiparallel β-
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sandwich flanked by a C-terminal α-helix (Fig. 17.1.9C; Fedorov et al., 1999; Prehoda et al., 1999). Interestingly, this fold closely resembles both the pleckstrin homology (PH) and phosphotyrosine binding (PTB) domains. EVH1 domains function by recognizing and targeting ligands containing the polyproline consensus sequence Phe-Pro-Pro-Pro-Pro; however, based on ligand specificity, EVH1 domains can be divided into class I proteins that recognize the sequence Phe-Pro-X-Φ-Pro, (X represents any residue and Φ represents a hydrophobic residue) and class II proteins that recognize the sequence Pro-Pro-X-X-Phe (Ball et al., 2002). The peptide ligand binds across the concave surface formed by β-strands 1, 5, 6, and 7. PH and PTB domains also bind their ligands on this same face of the β-sandwich, albeit in different regions (Fedorov et al., 1999). GYF domains. The GYF domain was first identified in CD2BP2, a signaling protein that binds to a proline-rich region of the cytoplasmic tail of the T cell adhesion protein CD2 (Nishizawa et al., 1998). The SMART database now lists 54 occurrences of GYF domains including sequences from fungi, plants, and metazoans (see Web-Based Structural Bioinformatics and http://smart.embl-heidelberg.de/browse.shtml) . The NMR solution structure of the C-terminal 62 residues of Drosophila CD2BP2 reveals a small four-stranded antiparallel β-sheet adjacent to a small α-helix (Fig. 17.1.9D; Freund et al., 1999). NMR and mutational data suggest that the binding site is centered around the highly conserved triplet Gly-Tyr-Phe situated in the loop between the α-helix and the third β-strand (Freund et al., 1999). Hence, this domain has been named the GYF domain. Phospholipid-binding domains PH domains. Pleckstrin homology (PH) domains are modules of ∼100 amino acids that are present in a wide range of signaling proteins, including many kinases, isoforms of phospholipase C, GTPases, GTPase-activating proteins, and nucleotide-exchange factors. The majority of characterized PH domains recognize and bind phosphoinositides or inositol phosphates. Indeed, most PH domains are part of membrane-associated proteins and it has been suggested that PH domains function as membrane adaptors, targeting proteins to specific membranes (Hurley and Misra, 2000; Maffucci and Falasca, 2001); however, recent evidence suggests that some PH domains, such as those of the β-adrenergic receptor kinase (β-ARK), the TecII tyrosine kinase, insulin
receptor substrate 1 (IRS-1), and the Etk tyrosine kinase, may bind protein ligands (see Maffucci and Falasca, 2001, and references therein). Each PH domain folds into a seven-stranded antiparallel β-sandwich flanked by a C-terminal long α-helix at one edge of the sandwich (Downing et al., 1994; Ferguson et al., 1994; Macias et al., 1994; Yoon et al., 1994). Strands A, B, C, and D form one sheet of the β-sandwich while strands E, F, and G form the other (Fig. 17.1.10A). The identified IP3 binding site is located at the two loop regions, loop AB (the loop between strand A and B ) and CD (the loop between strand C and D), opposite of the long helix. The basic PH fold has also been observed in PTB and EVH1 domains and the Pan-binding domain, despite the fact that these domains have no sequence homology with the PH domain. This has led to its reference by some as the PH superfold (Blomberg et al., 1999). C1 domains. The C1 domain is a cysteinerich compact structure of ∼50 residues. It was first identified as a conserved region of protein kinase C (PKC) isozymes that binds diacylglycerol (DAG) and phorbol esters and participates in allosteric regulation. Since then, several hundred C1 domains have been found in other proteins. Not all of these bind DAG but may instead bind proteins or other lipids. For reviews, see Cho (2001) and Hurley and Misra (2000). Structurally, the C1 domain (Fig. 17.1.10B) comprises a two-stranded β-sheet, a threestranded β-sheet, a small α-helix and two bound Zn2+ ions (Hommel et al., 1994; Zhang et al., 1995). Each Zn2+ ion is coordinated by three Cys and one His side chain. The DAG/phorbol ester binding site is situated at one end of the molecule in a region surrounded by hydrophobic residues. A belt of basic residues abuts this region, encircling the middle part of the molecule. C2 domains. C2 domains were initially identified as a Ca2+-binding conserved homology region of protein kinase C (PKC) isozymes. To date, however, C2 domains have been observed in over 1600 different proteins according to the SMART database (see Web-Based Structural Bioinformatics and http://smart. embl-heidelberg.de/browse.shtml), most of which function in signal transduction or membrane trafficking (see Cho, 2001, and Hurley and Misra, 2000 for reviews). The majority bind membrane phospholipids in a Ca2+-dependent manner; however, not all require Ca2+ and some domains bind proteins as well. The
Structural Biology
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Overview of Protein Structural and Functional Folds
structure of C2 domains consists of ∼130 residues folded into an eight-stranded β-sandwich with topologies resembling an Ig fold (Fig.17.1.10C and Fig. 17.1.2, respectively; Sutton et al., 1995; Grobler and Hurley, 1997). Although the β-strands are consistently in the same location, two different circularly permuted C2 domain topologies are observed. The first β-strand (strand a) in synaptotagmin-like C2 domains becomes the last β-strand in the phospholipase C–like C2 domains. The rest of the topology is identical between various C2 domains. Three variable loops at one end of the domain, referred to as Ca2+-binding regions (CBRs), are intimately involved in the binding of two to three Ca2+ ions and in phospholipid binding. Interestingly, the CBRs also correspond topologically to the complementaritydetermining regions (CDRs) of the variable domains of antibodies. Structure-function studies of C2 domains from PKCα and cytosolic phospholipase A2 (cPLA2) have shown that the two bound Ca2+ ions in each C2 domain play distinct roles: one provides a bridge between the C2 domain and anionic phospholipids, whereas the second Ca2+ induces a conformational change that enhances membrane-protein interactions (Cho, 2001; Stahelin and Cho, 2001). FYVE domains. FYVE fingers are modular domains of ∼60 residues that specifically bind to phosphatidylinositol 3-phosphate (PI3P). They are observed in numerous eukaryotic proteins from yeast to human, and function primarily in the regulation of endocytic membrane trafficking (for a review, see Stenmark et al., 2002). The name originates from the first four FYVE finger-containing proteins identified: Fab1p, YOTB, Vac1p, and EEA1. The three-dimensional structure of the FYVE finger consists of two double-stranded antiparallel βsheets, a C-terminal helix and two long loops (Fig. 17.1.10D; Misra and Hurley, 1999; Mao et al., 2000). The core structure is further stabilized by two Zn2+ ions that are coordinated by four conserved Cys-X-X-Cys motifs. A 2.2Å crystal structure of the homodimeric EEA1 FYVE finger bound to inositol 1,3-bisphosphate (a soluble analogue of PI3P) reveals the probable PI3P binding site, which includes residues from the following conserved FYVE signature motifs: the N-terminal Trp-X-X-Asp, the C-terminal RCV, and the Arg-(Arg/Lys)His-His-Cys-Arg motif (Dumas et al., 2001). A lower resolution NMR structure of EEA1 bound to di-C4-PI3P shows a similar binding site, but with different atomic contacts (Ku-
tateladze and Overduin, 2001). Furthermore, a hydrophobic N-terminal loop (referred to as the turret loop) adjacent to the PI3P binding site is proposed to partially penetrate and interact with the membrane surface. Protein interaction domains PDZ domains. PDZ domains are compact globular domains of ∼90 residues that serve as protein-protein interaction modules in the assembly of large complexes. For reviews, see Hung and Sheng (2002) and Ponting et al. (1997). These relatively abundant domains are widely observed in bacteria, plants, and higher eukaryotes, including humans. Several of these domains (up to 13) are often found within one protein. Formerly known as Disc-large homology regions (DHRs) or GLGF repeats, PDZ modules derived their current nomenclature from the first three PDZ-containing proteins identified: PSD-95/SAP90, Disc-large, and ZO-1. The first PDZ structures determined were the third PDZ of rat PSD-95 (Doyle et al., 1996) and the third PDZ domain of human Dlg (Morais Cabral et al., 1996). Structurally, the domain comprises a five- or six-stranded βsandwich and two α-helices that specifically bind the C-terminal peptide of certain proteins (Fig. 17.1.11A). The peptide ligand binds as an antiparallel β-strand between the β-strand and the αB-helix with up to nanomolar affinity. PDZ domains have also been reported to interact with internal peptide sequences in addition to C-terminal peptides, as observed in the crystal structure of nNOS and syntropin (Hillier et al., 1999). Three classes of PDZ domains have been loosely defined based on their specificity for the last four residues of the bound peptide: class I binds the sequence -X-Ser/Thr-X-Φ, class II binds -X-Φ-X-Φ, and class III binds -X-Asp/Glu-X-Φ, where X represents any amino acid and Φ represents a hydrophobic residue. However, a more comprehensive and complicated classification has also been proposed based on the identity of two residues within the PDZ fold (Bezprozvanny and Maximov, 2001). VHS domains. VHS domains comprise ∼140 residues and reside in the N-termini of certain proteins that play key roles in vesicular trafficking and protein sorting. The name VHS originates from Vps27, Hrs, and STAM, three proteins that harbor this domain; however, to date, well over 100 proteins are known to contain VHS domains. These can be divided into four categories based on their relative placement
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with other modular domains (Lohi et al., 2002). Group 1 consists of the STAM/EAST/Hbp family, each of which have the VHS-SH3-ITAM domain organization (see above for SH3). Group 2 proteins have a VHS-FYVE arrangement which includes the Vps27 and Hrs proteins (see above for FYVE). Group 3 contain Golgi-localizing, γ-adaptin ear homology domain, ARF-interacting (GGA) proteins with a VHS-GAT-ear arrangement. Finally, group 4 are VHS-containing proteins that do not fit into groups 1 to 3. Crystal structures of the VHS domains of Drosophila Hrs and human TOM1 (Mao et al., 2000; Misra et al., 2000) revealed a righthanded superhelix of eight α-helices (Fig. 17.1.11B) with striking structural homology to the eight-helix ENTH domain of epsin-1. The first seven helices of the VHS and ENTH domains superimpose with rms deviations of ∼1.8 Å (Lohi et al., 2002). Furthermore, the VHS domain displays structural similarities to HEAT and ARM repeats. More recently, crystal structures of the VHS domains of GGA1 and GGA3 complexed with target peptides showed that signal peptides containing an acidic-cluster dileucine (ACLL) motif, Asn-X-X-Leu-Leu, bind to a groove between α-helices 6 and 8 (Fig. 17.1.11B; Misra et al., 2002; Shiba et al., 2002). SNAREs. Soluble, n-ethylmaleimide-sensitive factor attachment-protein receptors (SNAREs) comprise a superfamily of conserved eukaryotic proteins that function in membrane fusion. They are characterized by ∼60 residue SNARE domains consisting of a long amphipathic α-helix. Most SNAREs are type II transmembrane proteins with an N-terminal cytoplasmic SNARE domain and a very short extracytoplasmic region; however, some are fastened to the membrane by palmitoylation or prenylation rather than by a transmembrane domain (Hay, 2001). Upon complex formation, SNARE domains anchored to opposite membranes come together to form a helical bundle that promotes membrane fusion. For reviews, see Hay (2001) and Misura et al. (2000a). First observed to atomic resolution in the crystal structure of the syntaxin-1A/synaptobrevin-II/SNAP-25B complex (Sutton et al., 1998), this complex comprises a structurally conserved coiled-coil of four parallel α-helices (Fig. 17.1.11C). Superficially, it resembles the tetrameric GCN4 four-helix-bundle with a largely hydrophobic core; however, deviations in the heptad repeat pattern and composition, as well as local deviations in helical curvature,
set this complex apart from known leucine-zipper or coiled-coil proteins. SNARE complexes consist of a single SNARE from one membrane termed an RSNARE and three Q-SNARES associated with the opposing membrane. In some cases, two Q-SNARES are on the same polypeptide (e.g., SNAP-25) while in other cases all three QSNARES are on separate molecules. The “R” and “Q” designations are based on the presence of conserved Arg or Gln residues in the center (or 0 layer) of each respective SNARE domain. The Arg and three Gln residues interact with each other at the core of the complex to form an intriguing hydrogen bond network completely isolated from solvent. The Q-SNAREs have been further classified by protein profiling (Bock et al., 2001). Group Qa are homologous to syntaxins, group Qb are homologous to the N-terminal helix of SNAP-25 and the group Qc SNAREs are homologous to the C-terminal helix of SNAP-25. Furthermore, the R-, Qa-, Qb-, and Qc-SNAREs each appear to occupy conserved positions within the complex. In isolation, SNARE domains are unstructured and are referred to as existing in an open conformation that favors SNARE complex formation; however, syntaxin SNAREs have an Habc domain N-terminal to the SNARE domain that has an antiparallel triple-helix structure (Fernandez et al., 1998). The Habc domain can complex with the SNARE domain (referred to as H3 in this context), induce helical structure, and form a four-helix bundle (Munson et al., 2000). This is referred to as a closed SNARE conformation and inhibits complex formation with other SNAREs. Syntaxin appears to exist in equilibrium between open and closed forms. Regulatory proteins such as nSec1 can perturb this equilibrium by binding to the closed conformation, thereby stabilizing it and preventing SNARE complex formation. The crystal structure of yeast syntaxin Sso1p (H3Habc; Munson et al., 2000) shows the H3 helix is very similar in structure to the comparative helix in a SNARE complex, although its orientations with neighboring helices is strikingly different. Curiously, the crystal structure of the nSec1/syntaxin 1a complex (Misura et al., 2000b) reveals the H3 domain adopting an irregular bent helical structure distinct from that of the SNARE complex. Thus, the H3 region of syntaxin can have at least three different conformations depending on its local environment or binding partners. Structural Biology
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Overview of Protein Structural and Functional Folds
SAM domains. The sterile alpha motif (SAM) is a modular protein interaction domain present in a wide variety of signaling proteins, including receptor tyrosine kinases, Ser/Thr kinases, diacylglycerol kinases, transcription factors, scaffolding and adaptor proteins, polyhomeotic proteins, and GTPases. The SMART database (see Web-Based Structural Bioinformatics and http://smart.embl-heidelberg.de/ browse.shtml) lists over 700 proteins containing SAM domains. SAM domains are ∼70 residues in size and generally reside in the Nor C-terminal ends of proteins. The basic fold consists of an N-terminal extended region followed by four short α-helices and one long C-terminal α-helix (Fig. 17.1.11D). Several modes of SAM domain interaction have been observed crystallographically, including dimer formation as well as polymerization (Smalla et al., 1999; Stapleton et al., 1999; Thanos et al., 1999; Kim et al., 2002). Furthermore, there are reports of SAM domains binding other nonSAM containing proteins. MH2 domains (C-terminal domain of Smad proteins). Smad proteins are an integral part of the receptor signal transduction apparatus for the transforming growth factor-β (TGF-β) family of cytokines such as TGF-β, bone morphogenetic protein (BMP), and activins. Smads are divided into three classes: comediator Smads (Co-Smads), receptor regulated Smads (RSmads), and inhibitory Smads (I-Smads). Both the N- and C-terminal domains of Smads are conserved across species and connected by a variable linker region (L-domain). The N-terminal domain (MH1) interacts with DNA, whereas the C-terminal MH2 domain is involved in phosphorylation-regulated homoand hetero-oligomerization among Smads as well as interaction with other proteins in the signaling pathway. The fold of the MH2 domain comprises a twisted antiparallel β sandwich made from five- and six-stranded β-sheets capped at one end by a three-helix bundle pointing away from the sandwich, and capped at the other end by a three large loops and a helix (Fig. 17.1.11E; Shi et al., 1997). Signal-dependent phosphorylation of two serines at the C-terminal Ser-Ser-X-Ser sequence of R-Smads is believed to trigger the formation of a heterotrimer with a Co-Smad. Homotrimeric crystal structures of R-Smads 1, 2, and 4 MH2 domains (psuedophosphorylated, phosphorylated, and unphosphorylated, respectively) have provided models for this interaction and indicate a phosphoserine binding site at the L3 loop. (Fig. 17.1.11F; Shi et al., 1997; Qin et al., 2001; Wu
et al., 2001). There are also crystal structures of complexes of the MH2 domains of both Smad2 and Smad3 with extended structures of SARA (Smad anchor for receptor activation; Fig 17.1.11G; Wu et al., 2000; Qin et al., 2002). Structural repeat motifs In addition to the numerous modular domains that form independently folded functional units, a number of much smaller structural motifs of 20 to 40 residues are observed that exist as tandem arrays of 5 to 25 repeats (Groves and Barford, 1999). In proteins containing these arrays, the functional unit is a set of repeats rather than the individual motifs. The stacking of adjacent motifs often results in curving superhelical structures, many of which function in the context of macromolecular binding interactions. Four of the more well known of these structural repeat motifs are discussed below. Leucine-rich repeats. The leucine-rich repeat (LRR) was first identified in 1985 in the sequence of α2-glycoprotein (Takahashi et al., 1985). Now, however, over 2900 proteins are reported to contain LRRs according to the SMART database (see Web-Based Structural Bioinformatics and http://smart.embl-heidelberg.de/browse.shtml). Most of these are eukaryotic proteins, but >100 are in prokaryotic sequences. Although these proteins have diverse functions, a common element that most LRRs share is mediating protein-protein interaction. LRRs comprise repeats of 20 to 30 amino acids with the following eleven-residue repeat consensus sequence LeuX-X-Leu-X-Leu-X-X-(Asn/Cys)-X-Leu, where X represents any residue and some L positions can be occupied by Val, Ile, or Phe (Kajava, 1998; Kobe and Kajava, 2001). The three-dimensional structure of the repeat consists of an α-helix followed by a single β-strand (Kobe and Deisenhofer, 1993). Subsequent repeats stack directly on top of each other such that the β-strands form a curved parallel β-sheet and the helices line up parallel to a common axis (Fig. 17.1.12A). LRR structures are generally crescent shaped with the β-sheet along the inner concave surface and the helices on the outside convex surface with very little twist along the central axis. So far, the β-sheet region and adjacent loops of LRRs appear to be the most frequent areas for protein-protein interaction (Kobe and Kajava, 2001). HEAT repeats. The HEAT repeat derives its name from four of the first proteins identified to contain it: Huntingtin, elongation factor 3, the PR65/A subunit of protein phosphatase 2A,
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and lipid kinase TOR (Andrade and Bork, 1995). Andrade and coworkers (2001) reported the detection of 71 HEAT repeat-containing eukaryotic proteins in the Swissprot database. HEAT repeats are 37 to 43 residues long and consist of two antiparallel α-helices designated A and B. Between 3 and 22 repeats stack directly on top of each other and thus form two layers of parallel helices with an overall superhelical curvature (Fig. 17.1.12B; Groves et al., 1999). B helices line the concave surface and A helices the convex surface. The crystal structures of importin β1 and β2 bound with different protein ligands each reveal protein recognition sites among the inner B helices (Chook and Blobel, 1999; Cingolani et al., 1999; Vetter et al., 1999). For recent reviews about HEAT repeats, see Andrade et al. (2001), Groves and Barford, (1999), and Kobe et al. (1999). ARM repeats. ARM repeats are named after the armadillo protein, the Drosophila segment polarity gene product in which they were first identified. These repeats are very similar to HEAT repeats and are ∼40 residues in length; however, instead of the two helices observed in each HEAT repeat, ARM repeats consist of three helices, designated H1, H2, and H3 (Groves and Barford, 1999; Andrade et al., 2001). The SMART database reports nearly 400 proteins containing ARM repeats (see Web-Based Structural Bioinformatics and http://smart.embl-heidelberg.de/browse.shtml). The crystal structures of β-catenin and karyopherin α (importin α) revealed stacking of repeats to form three ladders of parallel helices arranged in a superhelical structure (Fig. 17.1.12C; Conti et al., 1998; Huber et al., 1997). The superhelical twist is formed by an ∼30° rotation of each repeat from its neighbors. Within individual repeats, the antiparallel helices H2 and H3 have orientations similar to helices A and B of HEAT repeats. Helix H1, however, is smaller and lies perpendicular to the H2/H3 pair. H3 and its flanking loops are observed to form a groove lined with conserved residues along the entire length of the ARM repeat structure. The structure of β-catenin complexed with the cytoplasmic domain of E-cadherin provides a striking example of how this groove can be used for protein recognition (Huber and Weis, 2001). In the complex, the C-terminal 100 residues of cadherin bind in a predominantly extended conformation along the entire length of the H3 groove of β-catenin. A variant of the ARM motif is also observed to bind to specific RNA sequences in the Pumilio
family of mRNA translation regulators (Wang et al., 2002). Ankyrin repeats. The ∼33-residue ankyrin (ANK) repeat is found in numerous proteins with varied functions, such as cyclin-dependent protein kinase inhibitors, the notch membrane receptor, the inhibitory subunit (IκBα) of nuclear factor κB (NF-κB), the transcriptional regulator GABPβ, the p53-binding protein 53BP2, certain cytoskeletal organizing proteins, and toxins (Groves and Barford, 1999; Sedgwick and Smerdon, 1999). The SMART database reports over 2700 proteins containing ankyrin repeats (see Web-Based Structural Bioinformatics and http://smart.embl-heidelberg. de/browse.shtml). The ankyrin repeat receives its name from the cytoskeletal protein ankyrin in which 24 repeats were found (Lux et al., 1990); however, it was first identified three years earlier in several cell cycle and developmental regulators (Breeden and Nasmyth, 1987). Amidst the diverse assortment of proteins containing ankyrin repeats, the common functional theme that has developed is that of protein-protein interaction. The ankyrin fold was first observed in the crystal structure of 53BP2 bound to the p53 tumor suppressor (Gorina and Pavletich, 1996). The structural repeating unit consists of a βhairpin loop followed by a pair of antiparallel α-helices that are roughly perpendicular to the β-hairpin (Fig. 17.1.12D). However, the repeating amino acid sequence actually begins with the second β-strand in one unit and ends with the first β-strand in the next. Stacking of repeats results in a curving structure with a left-handed twist, consisting of two rows of parallel helices and a narrow extended β-sheet. Ankyrin repeat proteins recognize their target proteins in varied ways, but a strikingly common theme involves interaction through the ends of the βhairpin fingers.
Protein Kinase Fold Eukaryotic protein kinases (PK) comprise an enormous superfamily of homologous proteins that catalyze the transfer of the γ phosphate of ATP to specific hydroxyamino acids on a protein substrate. These proteins play numerous roles directly regulating and participating in a diverse array of signal transduction cascades. PKs can be broadly categorized as tyrosine kinases and serine/threonine kinases; however, a comprehensive classification system was published in 1995 based primarily on sequence homology (Hanks and Hunter, 1995). This classification has recently been expanded
Structural Biology
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Overview of Protein Structural and Functional Folds
to include a total of seven major groups of PK families (Manning et al., 2002a, 2002b). Tyrosine kinases (TK) comprise one group and include both membrane-spanning receptors, such as the insulin receptor, and cytosolic proteins, such as Src kinases. The Ser/Thr kinases comprise six main groups. The AGC group includes cyclic nucleotide-regulated PKs and diacylglyceride-regulated/phospholipid-depe ndent PKs (PKC), among others. The CaMK group consists of calcium/calmodulin-dependent PKs. The CMGC group includes cyclin-dependent PKs, mitogen activated protein (MAP) kinases, glycogen synthase kinase-3, and casein kinase-II. The STE grouping includes several MAPK cascade families. The CK1 group comprises CK1, tau tubulin kinase (TTBK), and vaccinia related kinase (VRK) families. The sixth group of serine/threonine kinases comprises tyrosine kinase-like (TKL) families. These include the mixed lineage kinase (MLK), interleukin-1 receptor-associated kinase (IRAK), LIMK/TESK, Raf, receptor interacting protein kinase (RIPK), and the activin and TGF-β receptor kinases. At least 50 proteins have been identified that have sequence similarity with eukaryotic protein kinases and yet are either reported or predicted to lack catalytic activity (Manning et al., 2002b). Finally, more than a dozen atypical protein kinases (aPKs) have been identified that are reported to have protein kinase activity, but have no sequence homology with the eukaryotic protein kinase superfamily (Manning et al., 2002b). Protein kinase catalytic domain. Despite the diversity of this superfamily, structural studies have consistently revealed a structurally conserved core fold for the catalytic domain of eukaryotic protein kinases. Other modular domains (e.g., SH2, SH3, PH, SAM, C2, and PDZ domains, to name a few) are sometimes fused to the catalytic domain for regulatory or localizing purposes. First elucidated in the crystal structure of cyclic AMP-dependent protein kinase (also known as PKA; Knighton et al., 1991), the bilobal protein kinase fold consists of an N-terminal lobe (N-lobe) comprising a five-stranded antiparallel β-sheet with a flanking α-helix and a larger C-terminal, primarily α-helical lobe (C-lobe) of eight to ten α helices and some short β-strands (Fig. 17.1.13A). The active site is located between the lobes with the N-lobe contributing an ATP-binding glycinerich loop and the C-lobe providing most of the protein-substrate-binding site. An activation loop in the C-lobe contains one or more phosphorylation sites that help activate the enzyme
when phosphorylated. Although the topology and core catalytic domain structures are well conserved across this large superfamily, there are some noticeable differences among various members. These include additions and deletions of secondary structure elements, changes in the orientation of secondary structure elements, N- and C-terminal extensions, and variations in the relative orientation between the Nand C-lobes. Some of these structural differences appear to correspond to the sequencebased classifications of PKs discussed above. Noteworthy variations include a 25- to 30-residue insertion between helices G and H (PKA nomenclature) in the CMGC protein kinases (e.g., cyclin-dependent and MAP kinases) and the extra helix C observed in the cAMP-dependent protein kinases. Two atypical protein kinases, the transient receptor potential (TRP) Ca2+-channel kinase (also referred to as ChaK or TRP-PLIK) domain (Yamaguchi et al., 2001) and the actin-fragmin kinase (Steinbacher et al., 1999) have been found to share significant structural homology with the eukaryotic PK superfamily members despite a lack of sequence similarity; however, the structurally homologous region in these enzymes is smaller, comprising most of the N-lobe but only the N-terminal portion—i.e., helices D and E (PKA nomenclature) and the following β-strands—of the C-lobe. The rest of the C-lobe consists largely of α-helices that do not correspond to those in the PK superfamily. Furthermore, actin-fragmin kinase has an extra two-helix insertion in the N-lobe, and the C-lobe of ChaK has a bound zinc atom that is sequestered from solvent. Src family kinases. The structure of src tyrosine kinases is briefly discussed here as a special case to illustrate how SH2 and SH3 domains are used in cis to regulate and localize protein kinase activity. Most src family tyrosine kinases share a common architecture comprising four Src homology (SH) regions in addition to a variable region. The N-terminal SH4 region contains a myristylation signal required for attachment to the membrane, followed by a variable region that may localize the enzyme more specifically. The subsequent SH3 and SH2 domains bind proline-rich targets and certain phosphotyrosine motifs respectively (for more information, see SH2 Domains and SH3 Domains, respectively). These two domains are involved in both regulation of the enzymatic activity and subcellular localization of the kinase. The catalytic activity of the kinase is found in the C-terminal SH1 region. A short
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segment C-terminal to the SH1 domain contains a tyrosine residue that inactivates the kinase when phosphorylated. Similar to other protein kinases, phosphorylation of a tyrosine in the activation loop increases activity of the kinase. The crystal structure of inactive haematopoietic cell kinase (Hck; Sicheri et al., 1997) includes the SH2, SH3, and catalytic domain of the enzyme, while the structures of the catalytic domains alone have been determined for lymphoid cell kinase (Lck) in its activated form (Yamaguchi and Hendrickson, 1996) and C-terminal Src kinase (Csk) in its inactive form (Lamers et al., 1999). The Hck crystal structure shows the SH2 domain interacting with a C-terminal phosphotyrosine and the SH3 domain bound to the linker region between the SH2 and kinase domains, effectively locking the kinase in an inactive state (Fig. 17.1.13B). Despite the fact that both the SH3 and SH2 domains are located distal to the active site of the kinase, their intramolecular interactions in the inactive form results in positioning helix C of the N-lobe such that the orientation of a key residue in the active site is disrupted.
Phosphatases Protein phosphatases Eukaryotic protein phosphatases comprise three major families of enzymes, each of which have conserved core folds (Barford et al., 1998). The protein tyrosine phosphatase (PTP) family dephosphorylates phosphotyrosine residues, while the phosphoprotein phosphatase (PPP) and protein phosphatase M/Mg2+-dependent phosphatase (PPM) families consist of enzymes that dephosphorylate phosphorylated serine and threonine residues. PTPs. Protein tyrosine phosphatases (PTPs) include cytosolic and membrane-spanning, receptor-like, tyrosine-specific phosphatases in addition to dual-specificity phosphatases (DSPs) that can dephosphorylate phosphorylated tyrosine, serine, or threonine. Receptorlike PTPs generally have two tandem PTP domains whereas other PTPs contain one PTP domain. Structurally, there are three subtypes of PTP catalytic domains, all of which share a threelayer α/β/α sandwich architecture. The lowmolecular-weight (subtype I) PTPs are composed of a single domain containing a central four-stranded parallel β-sheet flanked by six helices on both sides (Fig. 17.1.13C). The βsheet has a strand order of 2-1-3-4. Subtype II
PTPs have a central highly curved five- or ten-stranded mixed β-sheet sandwiched between four to seven helices on one side and one or two on the other (Fig. 17.1.13D). Unlike subtype I, the strand order for the central parallel strands of this β-sheet is 1-4-2-3. Subtype II PTPs include the so-called high-molecularweight PTPs such as PTB1, Yersinia PTP, and SHP-1 and -2, in addition to the smaller DSPs such as VHR, MAP kinase phosphatase, kinase-associated phosphatase (KAP), and tumor suppressor PTEN (Table 17.1.5). The high-molecular-weight PTPs have a nine-stranded βsheet whereas the DSPs have a smaller fivestranded β-sheet composed of the same four central parallel strands and one preceding antiparallel strand, and have an additional one or two extra helices. Although superficially quite similar to other DSP structures, the catalytic domain of CDC25 has a rather different topology and the strand order of its five-stranded parallel β-sheet is 1-5-4-2-3 (Fig. 17.1.13E). Therefore, it may be referred to as a third subtype of protein phosphatase. Interestingly, CDC25 has the same fold as the sulfurtransferases rhodanase, GlpE, and the ERK2-binding domain of MAPK phosphatase-3. Although the topologies of these protein phosphatase subtypes are different, the active site strandloop-helix regions show a striking structural similarity, and they all feature an essential nucleophilic cysteine residue located in a cleft between the β-sheet and the larger helical bundle. Protein serine/threonine phosphatases. The PPP and PPM families both comprise metaldependent serine/threonine protein phosphatases. The fold of the PPP family catalytic domain is a mixed β sandwich flanked by seven α helices on one side, an α/β structure of two to three α helices, and a couple of β-strands on the other side (Fig. 17.1.13F). The active site, including the essential Zn2+ and Fe3+ cations, is located among the loops at one end of the β sandwich. Enzymes in this family include PP1, calcineurin (PP2B), and the bacteriophage λ protein phosphatase. This fold is also shared by at least three other phosphoryltransfer metalloenzymes that are not protein phosphatases: purple acid phosphatase, the N-terminal domain of UDP-sugar hydrolase, and the N-terminal domain of the DNA double-strand-break repair enzyme MRE11 nuclease (Table 17.1.5). The PPM family includes both eukaryotic and prokaryotic protein phosphatases and has been defined by protein phosphatase 2C (PP2C). Although PP2C has no sequence ho-
Structural Biology
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DNA-BINDING STRUCTURAL MOTIFS AND DOMAINS
Alkaline phosphatases Alkaline phosphatases (AP) are dimeric enzymes that catalyze the hydrolysis of phosphomonoesters and are common to all organisms. The overall structure of an AP monomer comprises two domains, both of which are α/β/α sandwiches (Fig. 17.1.14A). The smaller domain consists of a three- or four-stranded antiparallel β-sheet sandwiched between two αhelices, one on each side. The larger domain has a central ten-stranded β-sheet in the middle flanked by thirteen helices of various lengths. Except for the eighth strand, this β-sheet is composed of parallel strands. Two other enzymes that share the core AP fold include arylsulfatase and prokaryotic cofactor-independent phosphoglycerate mutase (Table 17.1.5).
Numerous DNA-binding domains are observed to utilize a small number of conserved substructures or structural motifs for binding DNA. In most cases a portion of the structural motif protrudes from the domain and interacts with the major and/or minor groove of DNA and also with the phosphate backbone through hydrogen bonds, salt bridges, and nonpolar van der Waals interactions. By far the most prevalent interactions involve an α-helix binding in the major groove. Other less common interactions include β-strands or loops binding in either the major or minor groove of the DNA. In a few cases, considerable distortion of Bform DNA is induced upon binding. This section summarizes some commonly observed DNA-binding structural motifs and DNA-binding domains.
Sugar phosphatases The sugar phosphatase fold is shared among the families of fructose-1,6-bisphosphatase, inositol monophosphatase, and inositol polyphosphate 1-phosphatase. The molecule folds into a five-layered α/β/α/β/α sandwich (Fig. 17.1.14B and Table 17.1.5). The two β-sheets are five and eight-stranded, respectively, and mostly antiparallel. The location of the AMP binding site is at the end of the N-terminal α-helix. This fold also remotely resembles the actin fold.
The Cyclin Fold
Overview of Protein Structural and Functional Folds
IIB and retinoblastoma tumor suppressor domains. Based on the structure of a cyclinACDK2 complex (De Bondt et al., 1993), the activation of CDK has been postulated to occur through a cyclin-induced conformational change in the PSTAIRE helix (C helix) and the T-loop of the kinase domain.
mology with PPP phosphatases, it is also a metalloenzyme with a binuclear metal (in this case Mn2+) center at the active site and has a fold that resembles that of PPP phosphatases. The N-terminal domain consists of an elevenstranded antiparallel β sandwich surrounded by α helices. As in the PPP family, the active site is located at one end of the β sandwich. The C-terminal domain consists of three antiparallel helices.
The eukaryotic cell cycle is regulated by a series of cyclin-dependent kinases (CDKs) that are activated by association with cyclins. Several different forms of cyclins (i.e., cyclins A, B, C, D, E, and H) are known to regulate separate CDKs. Conserved among these cyclins is a region of ∼100 amino acids, termed the cyclin box. The basic cyclin box fold contains five α-helices with the central helix, α3, being surrounded by the remaining four (Jeffrey et al., 1995; Noble et al., 1997). Cyclin A consists of two cyclin box domains in tandem (Fig. 17.1.14C and Table 17.1.5). The cyclin box fold is also observed in transcription factor
Helix-Turn-Helix Motifs The helix-turn-helix (HTH) structural motif, first observed in the crystal structure of the λ Cro protein (Anderson et al., 1981), is one of the most common and well characterized DNAbinding motifs, and is found in a very diverse group of proteins (Table 17.1.6). The HTH motif is part of a three-helix bundle in which the second and third helices (α2 and α3) of the bundle make direct contacts with DNA. The third helix, the recognition helix, protrudes from the rest of the protein into the major groove of DNA, making the majority (but not all) of the base-specific contacts, while the second helix sits above the recognition helix at an ∼120° angle to it, forming a bridge across the two phosphate backbones on either side of the major groove (Fig. 17.1.15). The N-terminal helix (α1) lies across the second helix opposite the DNA, completing the helical bundle where it forms interactions with the rest of the protein and may or may not interact with the DNA. There are now tens of permutations of the DNA-binding HTH motif listed as various superfamilies in the online SCOP database (see Web-Based Structural Bioinformatics). For the purposes of this overview, four of these permu-
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tations are discussed here: prokaryotic or phage repressor, eukaryotic homeodomain, winged helix, and PurR repressor, with some structures falling into more than one category (Table 17.1.6; Fig. 17.1.15). Prokaryotic repressor. The structures of prokaryotic repressor HTHs have served as the prototypical model for the overall HTH motif. In this particular class, the HTH motif by itself is not stable enough to fold or bind DNA, but requires the scaffolding of the rest of the protein. Furthermore, monomers often lack DNAbinding ability and thus dimerization is required for binding to DNA. Interestingly, outside the HTH motif of prokaryotic repressors there is little structural consensus among these DNA-binding proteins. Their structures have been observed to vary from α/β folds to all helical folds. Eukaryotic homeodomain. The eukaryotic homeodomain HTHs all use an N-terminal arm from helix 1 to bind DNA in the adjacent minor groove (Li et al., 1995). Some also use a C-terminal arm for minor groove binding as well. The additional interactions of the arm(s) enable homeodomains to bind DNA as monomers, although most homeodomains are observed to function as homo- or heterodimers in vivo. Also, some proteins in this class (e.g., yeast RAP1 telomere-binding protein, proto-oncogene product Myb, the Paired class homeodomains, the Oct-1 POU domain) contain two HTH motifs connected in tandem by a polypeptide linker. Winged-helix motif. Another class of HTH proteins is referred to as the winged-helix motif (Fig. 17.1.15D). First characterized as a motif in the eukaryotic HNF-3/fork head transcription factor (Clark et al., 1993), this motif uses one or two flexible loops (or wings) that protrude from the HTH on either side at approximately right angles to the recognition helix to provide additional DNA contacts. The more prominent wing is usually part of a small twoor three-stranded antiparallel β-sheet, a distinctive feature of the winged-helix motif. For a detailed review of the winged helix motif see Gajiwala and Burley (2000). PurR repressor. The PurR repressor of the LacI family represents a more extreme variant of the HTH motif with three major differences from the canonical HTH (Schumacher et al., 1994). First, although the spatial orientation is consistent with the HTH, the connectivity of the three helices is completely different so that helices 1 and 2 interact with the DNA instead of helices 2 and 3 (Fig. 17.1.15E). Second, the
orientation of the HTH with respect to the DNA dyad is in the reverse direction from what is observed in the classical HTH. Third, in addition to the HTH motif a fourth helix (the hinge helix) is present to bind the DNA in the adjacent minor groove. In a physiological dimer, the two symmetry-related hinge helices interact with each other in an antiparallel manner to pry open the minor groove with a set of leucine levers which forces the DNA axis to bend 45°.
Zinc-Containing DNA-Binding Motifs Six of the many classes of DNA-binding proteins that use zinc coordination to form the stable core or folding template for DNA-binding will be discussed in this section. These include the TFIIIA type zinc finger, the GATA1 Cys4 type zinc finger, the Zn2Cys6 binuclear cluster, the nuclear hormone receptor DNAbinding domains, the transcription elongation factor TFIIS, and the zinc binding loop of the p53 tumor suppressor. Zinc fingers The Cys2His2 zinc finger structural motif is the most common DNA-binding motif among eukaryotic transcription factors, with several thousand protein sequences identified (Berg and Shi, 1996). A canonical zinc finger motif contains the consensus sequence (Tyr/Phe)-XCys-X2–4-Cys-X3-Phe-X5-Leu-X2-His-X3–5His where X represents any amino acid, and the Cys and His residues serve as zinc ligands (Berg and Shi, 1996). The structure of the motif is comprised of two short antiparallel β-strands followed by an α-helix, which are stabilized by a small hydrophobic core and a tetrahedrally coordinated zinc atom (Fig. 17.1.16A). The N-terminal end of the α-helix rests in the major groove where it makes base-specific contacts while the β-sheet lies on the other side of the helix opposite the DNA (Fig. 17.1.17A) to form limited contact with the phosphate backbone. Zinc finger-containing proteins generally use this motif in tandem arrays within the DNAbinding domain. Since the first three-dimensional structure determination of a zinc fingerDNA complex (Zif268 from mouse intermediate protein; Pavletich and Pabo, 1991), the structures of several other protein-DNA complexes have been elucidated, including the DNA-binding domains from human oncogene GLI, the Drosophila regulator protein tramtrack, and TFIIIa, as well as numerous structures of uncomplexed zinc fingers (Fairall et al., 1993; Pavletich and Pabo, 1993; Foster et al., 1997).
Structural Biology
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The zinc-ribbon motif First observed in the eukaryotic transcription elongation factor TFIIS (Qian et al., 1993), the zinc ribbon motif consists of a small threeor sometimes four- or five-stranded antiparallel β-sheet that coordinates a zinc atom within the loop region at one end of the sheet (Fig 17.1.16B; Pan and Wigley, 2000). The zinc is coordinated by three Cys and one His residue (or four Cys side chains in the case of TFIIS). The mechanism of DNA binding for this motif is controversial and awaits further clarification from future structure determinations of zinc ribbon-DNA complexes.
Overview of Protein Structural and Functional Folds
Zn2Cys6 binuclear cluster A DNA-binding structural motif containing a Zn2Cys6 binuclear cluster has been characterized in numerous fragments of fungal transcription factors. The three-dimensional structure of fragments of these proteins—e.g., GAL4 and pyrimidine pathway regulator 1 (PPR1)—complexed with DNA have been determined (Marmorstein et al., 1992; Marmorstein and Harrison, 1994), revealing homodimers each containing three regions: two N-terminal zinc-coordinated DNA-binding modules (one in each monomer), a linker region, and a short coiled-coil made from two C-terminal helices which protrude away from the DNA. The DNA-binding modules are nearly identical in both proteins, consisting of two short helices, each followed by an extended loop (Fig. 17.1.16C). Six cysteines from each module tetrahedrally coordinate two zinc atoms. An internal two-fold symmetry relates the two helix-loop zinc-coordinated motifs within each module. In both GAL4 and PPR1, the C-terminal end of the first helix from each module points into the major groove, mediating basespecific DNA contacts to a CGG triplet (Fig. 17.1.17C and D, respectively). Apart from the DNA-binding motifs, the structures of GAL4 and PPR1 are rather different from each other. The linker region in each monomer of GAL4 is completely extended, making practically no contacts with the rest of the protein, whereas this same region in PPR1 folds into a more compact antiparallel β-ribbon. The GAL4 homodimer is highly symmetric, with the coiledcoil dimerization region projecting perpendicularly from the DNA. The PPR1 homodimer, however, becomes very asymmetric beyond the N-terminal DNA-binding modules, leaning to one side to form a hydrophobic core. Several other complexes of Zn2Cys6 motif-containing proteins with DNA show similar per-
turbations of domain orientation while retaining the basic fold (Table 17.1.6). GATA-binding motif The NMR solution structure of the chicken erythroid transcription factor GATA-1 complexed with its target DNA reveals a DNAbinding structural motif distinctly different from the canonical zinc finger structure (Omichinski et al., 1993). The consensus sequence Cys-X2-Cys-X17-Cys-X2-Cys characterizes an entire family of regulatory proteins expressed in numerous cell types, each of which recognize a GATA DNA sequence. The DNA-binding domain of GATA-1 consists of two short two-stranded irregular antiparallel β-sheets followed by an α-helix and a long C-terminal loop (Fig. 17.1.16D). The tetrahedrally coordinated zinc is ligated by cysteines from the first β-sheet and helix. The α-helix and loop which connects the two β-sheets are both located in the DNA major groove, and the long C-terminal tail crosses the phosphate backbone to sit in the minor groove (Fig. 17.1.17B). DNA-binding domains of nuclear hormone receptors Eukaryotic nuclear hormone receptors form a superfamily of ligand-activated transcription factors that regulate numerous cellular processes. These proteins generally consist of an N-terminal DNA-binding domain and a C-terminal hormone-binding domain responsible for transactivation. The minimal core of the N-terminal DNA-binding domain (60 to 70 residues) consists of two helices separated by a long loop packed against each other at approximately right angles (Fig. 17.1.16E). Two zinc atoms, one at the N-terminus of each helix, are each coordinated by four cysteine side chains. The first helix, also termed the recognition helix, lies in the major groove of bound DNA, making sequence-specific contacts while other portions of the core domain make extensive contacts with the phosphate backbone on either side of the major groove (Fig. 17.1.17E). Nuclear receptors are observed to bind cooperatively to both direct and inverted DNA repeats as either homo- or heterodimers. The three-dimensional structures of the DNAbinding domains of several nuclear receptors complexed with DNA have been elucidated (Table 17.1.6). These include the glucocorticoid receptor (Luisi et al., 1991), estrogen receptor (Schwabe et al., 1993), 9-cis retinoic
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acid receptor, and thyroid hormone receptor (Rastinejad et al., 1995).
MADS Box Domain The MADS box (also known as the SRFlike) family of proteins comprises many eukaryotic regulatory proteins, including human serum response factor (SRF), Drosophila trachea development factor (DSRF), the METF2 myocyte-specific enhancer factors, and the MCM1 gene regulator from yeast, all of which contain a highly conserved 60-residue core consensus sequence (Table 17.1.6; Pellegrini et al., 1995). SRF was the first of these proteins to have its three-dimensional structure determined (Pellegrini et al., 1995). This protein is homodimeric and consists of three layers (Fig. 17.1.18A). The first layer is a left-handed antiparallel coiled-coil formed from an amphipathic helix from each monomer. The coiled-coil sits above the minor groove near the DNA dyad (the binding site) in an approximate parallel orientation to the minor groove and makes contacts with the phosphate backbone. The DNA is bent by 72° along its helical axis at this point, wrapping around the coiled-coil and enabling the N-terminal portions of the two helices to fit into the major groove of the DNA. In addition, N-terminal arms from each helix extend across the phosphate backbone and bind in the minor groove of the DNA. The coiled-coil is supported by a four-stranded antiparallel βsheet that lies flat across the helices opposite the DNA. The third layer consists of a small two-helix subdomain above the sheet. Similar to other dimeric DNA-binding proteins, the two-fold symmetry axis of the dimer often coincides with the dyad axis of the DNA binding site. The crystal structure of the MCM1 MADS-box protein bound to DNA and the MATα2 homeodomain core (Tan and Richmond, 1998) reveals an intriguing ternary complex in which a β-hairpin from MATα2 extends the MCM1 β-sheet, and both the MATα2 HTH motif and the MCM1 MADS-box dimer bind DNA.
Basic Region Leucine Zippers The fold of basic region leucine-zipper (bZIP) domains in eukaryotic transcription factors is relatively simple. First observed in the structure of the GCN4 homodimer (Ellenberger et al., 1992) and the c-Fos/c-Jun heterodimer (Glover and Harrison, 1995), it consists of two, long parallel α-helices that dimerize via a coiled-coil leucine zipper motif at their C-terminal ends and spread apart like partially
opened scissors at the N-terminal end, gripping the DNA between them (Fig. 17.1.18B; Table 17.1.6). The primary sequence of leucine zippers contains heptad repeats (a-b-c-d-e-f-g) with hydrophobic residues at position a, leucine at position d, and charged or hydrophilic residues at the other positions (Ellenberger, 1994). Residues at positions a and d from each helix form extensive hydrophobic interactions at the core of the dimer interface, while charged side chains at positions e and g form inter- and intrahelical salt bridges at the edge of the interface. The N-terminal primarily basic ends of the helices lie in the major groove on either side of the DNA contacting base-pair edges and the phosphate backbone.
The Helix-Loop-Helix Motif The helix-loop-helix (HLH) structural motif is similar to the bZIP domain in both its DNAbinding and dimerization strategies. As with bZIP proteins, this structural motif binds DNA as a homo- or heterodimer. Each monomer consists of two long α-helices connected by a loop of variable length (5 to 23 residues) and composition forming a left-handed, parallel, four-helix bundle in the dimer (Fig. 17.1.18C). The first helix in each monomer binds DNA with its N-terminal basic region sitting in the major groove on opposite sides of the DNA helix. Dimer contacts are made through the C-terminal portion of the first helix and a coiled-coil formed by the second helix of each monomer. In some members of the HLH family, the coiled-coil contacts are formed by a leucine zipper; in other HLH proteins the second helix is shorter and the coiled-coil is stabilized primarily through less regular hydrophobic contacts. In both cases, the dimer is required for DNA binding. DNA bound to this motif is primarily B-form with very little distortion observed. Examples of three-dimensional structures of HLH proteins include the eukaryotic transcription factor Max (Ferre-D’Amare et al., 1993), the MyoD transcription activator (Ma et al., 1994), the HLH region of the upstream stimulatory factor (USF; Ferre-D’Amare et al., 1994), and the E47 regulatory protein (Ellenberger et al., 1994; Table 17.1.6).
HMG Box Domains DNA-binding proteins containing the highmobility-group (HMG) box comprise a diverse superfamily of eukaryotic regulatory proteins including various transcription factors (e.g., LEF-1), components of chromatin, products of fungal mating type genes, and the mammalian
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sex-determining gene product SRY (Table 17.1.6; for reviews see Laudet et al., 1993; Travers, 2000). The HMG consensus box consists of ∼80 residues with an average conservation of 25% amino acid identity between boxes. HMG box proteins fall into two subfamilies. The first is observed in all cell types, contains multiple HMG box domains, and tends to bind DNA with little specificity, while the second subfamily is found in fewer cell types, contains only one HMG box domain, and binds DNA specifically. In the mid-1990s, the NMR solution structures of the HMG domains in two proteins of the second subfamily complexed with DNA (Love et al., 1995; Werner et al., 1995) and the solution structures of uncomplexed HMG domains from the first subfamily revealed a three-helix L-shaped structure which binds DNA exclusively in the minor groove and induces profound structural changes in bound DNA (Fig. 17.1.18D). Helices 1 and 2 are antiparallel to each other (with an acute angle between them) forming the base of the L. Helix 3, the longest helix, is almost perpendicular to helix 2 and forms the vertical arm of the L. All three helices participate in forming a concave surface for binding the minor groove and phosphate backbone of the target DNA, causing severe DNA unwinding and bending away from the major groove.
DNA-Binding Proteins that use β-Sheet Motifs
The Histone Fold
TATA-box binding protein A second β-sheet motif for binding DNA is observed in the TATA box–binding protein (TBP), a component of the class II general transcription initiation factor TFIID that is required for preinitiation-complex formation with RNA polymerase II (Table 17.1.6). TBP consists of a C-terminal region of ∼180 residues which is highly conserved across species, and a variable length N-terminal region. The crystal structure of TBP from Arabidopsis thaliana bound to the TATA-box DNA consensus sequence reveals a quasi-symmetric α/β structure composed of a curved ten-stranded β-sheet with four α-helices sitting on the convex side of the sheet with an overall shape resembling a saddle (Fig. 17.1.19B; Burley, 1996). Two distinct loops project away from the protein on each end of the β-sheet forming stirrups for the saddle. DNA containing the TATA box consensus sequence binds to the concave surface formed by the eight central strands of the βsheet with an induced fit mechanism resulting in remarkably distorted DNA that is partially unwound and bent almost 90°. TBP has marked intramolecular pseudosymmetry with an ap-
A unique fold shared by the H2A, H2B, H3, and H4 subunits of the octameric histone core of the nucleosome was first described by Arents et al. (1991) as the histone fold. Since then, the histone fold has been observed in the archeael histones, the TBP-associated factor (TAF) family of transcription factors (e.g., TAFII42, TAFII62, TAFII18, TAFII28; Xie et al., 1996; Birck et al., 1998), and the TBP-associated negative cofactors (Kamada et al., 2001). The individual polypeptides display a helix-loophelix structure, with a central long α-helix flanked on each side by a short α-helix (Fig. 17.1.18E). The histone core subunits and the TAF fragments form similar heterodimeric and heterotetrameric structures. The function of histones in eukaryotic cells is to provide a scaffold for nuclear DNA to wrap around into a highly compacted nucleosome. It has been postulated in TAFs that a similar TAF octameric organization is involved in transcription activation. Overview of Protein Structural and Functional Folds
The vast majority of DNA-binding proteins structurally characterized to date are observed to interact with DNA through helical structures. However, three different families of DNAbinding proteins have been characterized in which β-sheet structures are used as the primary contacts for protein-DNA interactions. Ribbon-helix-helix motif The ribbon-helix-helix motif is found in prokaryotic and phage repressor proteins such as Arc (Raumann et al., 1994), MetJ (Somers and Phillips, 1992) and Mnt (Burgering et al., 1994; Table 17.1.6). These proteins bind DNA as dimers of dimers (homotetramers) and all have homologous structures in which the homodimer consists of a small helical bundle with a symmetric double-stranded antiparallel βsheet (β-ribbon) on the surface that can fit specifically into the major groove of DNA (Fig. 17.1.19A). In the three-dimensional structures of complexes with DNA, additional contacts are also made through helix and loop residues, but base-specific contacts to DNA are mediated exclusively through side chains of residues in the β-ribbon. Binding of the tetramers to the two tandem operator sites is cooperative and occurs with limited distortion of B-form DNA.
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proximate two-fold axis passing through the center of the protein perpendicular to the βsheet. The primary sequence of TBP can be divided into two long repeats (89 to 90 residues each) related by 30% sequence identity. The three-dimensional structure of each half is topologically identical with a root-meansquare (rms) deviation between equivalent Cα positions of 1.1 Å. Several other protein structures are observed to contain a fold similar to one TBP repeat (as opposed to two); however, none of them use the β-sheet to bind DNA. The histone-like HU family A third family of proteins that use β-ribbons to bind DNA is the highly conserved histonelike HU protein family (Table 17.1.6). These are prokaryotic dimeric proteins that bind DNA nonspecifically to form nucleosome-like particles containing negatively supercoiled DNA. The crystal structure of a 19-kDa uncomplexed homodimeric HU protein (also known as DNAbinding protein II) from Bacillus stearothermophilus reveals a V-shaped protein consisting of a six-helix bundle with two three-stranded, curved, antiparallel β-sheets extending out from the surface to form an open cradle (Fig. 17.1.19C; Tanaka et al., 1984). The crystal structure of a similar protein, heterodimeric integration host factor (IHF) from E. coli, complexed with DNA (Rice et al., 1996), reveals that the two β-sheets wrap around the DNA along the minor groove inducing a 160° bend in the DNA (Fig 17.19D).
The Ig-Like Family of Transcription Factors The structures of several families of Ig-like transcription factors have been elucidated (Table 17.1.6). Each of them uses an Ig-like fold with C2-type (also known as s-type) topology to bind DNA through loop regions at one end of the β sandwich (see Immunoglobulins and Immunoglobulin-Like Superfamilies for a discussion of Ig folds). The SCOP database classifies these proteins as the superfamily of p53like transcription factors (Murzin et al., 1995; see Web-Based Structural Bioinformatics for SCOP database). Although the individual structures and binding modes vary somewhat among these families, three common loops are used by all six families to contact DNA (Rudolph and Gergen, 2001). The loop between strands A and B (AB loop, sometimes referred to as the recognition loop) interacts with the major groove, making sequence-specific contacts. The loop between strands E and F (EF loop) contacts the
DNA backbone, in some cases making basespecific contacts in the minor groove, and the C-terminal tail emanating from strand G sits in the major groove, making base-specific contacts. The structural details of these families are described below. p53 tumor suppressors The first of these families to be examined structurally, p53 tumor supressors (Table 17.1.6), can bind DNA as a monomer (Cho et al., 1994). p53 tumor suppressors are key proteins facilitating protection against many types of cancer in animals. It is estimated that up to one-half of all cancers involve genetic inactivation of p53 (Vogelstein, 1990). In this structure, the major groove-binding C-terminal tail is an α-helix and the EF loop is stabilized by a zinc atom (Fig. 17.1.19E). NF-κB The NF-κB transcription factors are part of the larger Rel family of transcription factors containing a Rel homology region (RHR). They function as homo- and heterodimers; each monomer being composed of two Ig-like domains. The first domain is a DNA-binding C2type fold with an extension of strand E (designated E′) and an additional insert of two antiparallel α-helices between strands E and F. The second domain has an E-type Ig fold that provides the dimerization contacts. The NF-κB dimer/DNA complex has been described as resembling a butterfly with a DNA torso and protein wings (Fig. 17.1.19F; Ghosh et al., 1995; Muller et al., 1995). NFAT The NFAT family of transcription factors is similar to NF-κB in that its members possess a DNA-binding N-terminal C2-type Ig-like fold and a C-terminal E-type Ig fold. Furthermore, the DNA-binding domain has an additional helix inserted between strands E and F, two extra β-strands between C and C′, and an additional β-strand after strand E (E′). It is not surprising that the TonEBP (NFAT5) transcription factor forms the same type of dimeric butterfly complex with DNA as the NF-κB proteins with the added feature that its DNAbinding domains make dimer contacts as well, thus completely encircling the DNA (Stroud et al., 2002); however, other NFAT proteins bind DNA as monomers (Zhou et al., 1998) or bind DNA cooperatively with AP-1 transcription factors. The structure of a tetrameric complex of NFAT1/Fos/Jun/DNA (Chen et al., 1998b)
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shows the N-terminal domain interacting with DNA through the AB and EF loops and interdomain linker, as observed in other Ig-like transcription factors (Fig. 17.1.19G). However, the C-terminal domain forms an acute angle with the N-terminal domain and maintains only minimal contact with the N-terminal domain, the bound DNA, and the neighboring Fos helix. The Fos and Jun helices interact extensively with the NFAT1 N-terminal domain resulting in a 20° DNA bend and a 15° bend in the middle of the Fos/Jun heterodimer (relative to the Fos/Jun/DNA complex (see Basic Region Leucine Zippers; Chen et al., 1998b). CBF The core binding factors (CBF) comprise a family of heterodimeric transcription factors. The α subunit (CBFα) has the C2-type Ig-like fold (Berardi et al., 1999; Nagata et al., 1999) for binding DNA and is encoded by several genes. The invariant β subunit (CBFβ) has an α/β fold consisting of a partly open sixstranded β-barrel with four α-helices packed against the top and bottom (Fig. 17.1.19H; Goger et al., 1999; Huang et al., 1999). Although CBFβ does not interact with DNA itself, CBFα-DNA binding is apparently enhanced significantly by dimerization with CBFβ. Unlike the two Ig -like do mains in the NFAT1/DNA complex, the CBF α and β subunits interact extensively with each other either alone or in complex with DNA (Fig. 17.1.19H; Warren et al., 2000; Bravo et al., 2001; Tahirov et al., 2001). T-domain The T-domain (also known as T-box) transcription factors comprise a single C2-type Iglike domain with an additional two C-terminal helices (H3 and H4) that are capped at the DNA-binding end by several additional small twisted β-strands. The crystal structure of a Brachyury T-domain/DNA complex reveals a triangle-shaped dimer with the two T-domains forming the sides and the DNA forming the bottom of the triangle (Fig. 17.1.19I; Muller and Herrmann, 1997). The C-terminal α-helix (H4) has a rather unusual DNA interaction, being embedded deeply in the minor groove.
Overview of Protein Structural and Functional Folds
STAT proteins The signal transducers and activators of transcription (STAT) family of eukaryotic transcription factors includes large proteins (750 to 850 residues) that dimerize (homo- and heterodimers) upon phosphorylation and translo-
cate to the nucleus. Crystal structures of STAT-1 and -3β homodimers bound to DNA reveal elongated, four-domain monomers forming an arc-shaped ternary complex with DNA (Fig. 17.1.19J; Becker et al., 1998; Chen et al., 1998c). The N-terminal domain is a coiled-coil bundle of four antiparallel α helices. These helices project away from the DNA, have no interactions with DNA or the other monomer, and possess a primarily hydrophilic surface. The second domain is the DNA-binding domain. Like the other members of the Ig-like transcription factor superfamily, this domain is a modified C2-type Ig fold. The AB loop has an extra β-strand (A′), the CC′ loop contains an inserted α-helix (α5) and β-strand (X), the EF loop has an extra β-strand (E′), and β-strand G ends with an additional β-strand (G′). All three of the modified loops and the C-terminal tail participate in DNA binding. Following the Ig domain and prior to the SH2 domain is a helical connector or linker domain consisting of a bundle of four α helices and a small twostranded β-sheet. The SH2 domain (see Modular Domains Involved in Signal Transduction) binds a phosphorylated tyrosine from the C-terminal tail of the other monomer in the complex, effectively clamping the two monomers together.
RNA-BINDING STRUCTURAL MOTIFS AND DOMAINS Relative to DNA, RNA is capable of folding into a much more diverse array of structural motifs. In addition to the standard A-form double helix, irregularities such as bulges, hairpin loops, helix unwinding, noncanonical basepairing, and unstacked bases are all common in RNA structures. This results in RNA-binding protein structural motifs designed to interact with various perturbations of the RNA A-helix rather than a focus on major- and minor-groove interactions as observed in DNA-binding proteins. Although numerous three-dimensional structures have been determined for RNAbinding proteins, many of these use either unique or less common folds to recognize RNA; however, a small group of RNA-binding modules with known structures are noted for their more widespread use in RNA-protein interactions. These include RNP, OB-type domains, dsRBP, KH domains, and zinc fingers. This group of folds, along with a handful of larger, more unique structures, are briefly described below and in Table 17.1.7. For a review of some of these folds, see Perez-Canadillas and Varani, (2001).
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The RNP Domain
OB-Fold
The RNP domain, also known as the RNA recognition motif (RRM) or consensus sequence RNA-binding domain (cs-RBD), is one of the most widely observed and well characterized RNA-binding domains (Burd and Dreyfuss, 1994). It is identified in amino acid sequences by two highly conserved consensus sequences referred to as RNP1 and -2, within a more weakly conserved region of ∼80 residues. The SMART database lists over 3400 proteins containing RNP (see Web-Based Structural Bioinformatics and http://smart. embl-heidelberg.de/browse.shtml ). The threedimensional structure of RNP domains determined in the early 1990s (U1A splicosomal protein; Oubridge et al., 1994) and the RNAbinding domain of hnRNP C (Wittekind et al., 1992) revealed a four-stranded antiparallel βsheet with two α-helices laying across one face with a βαββαβ topology. The crystal structure of the RNA-binding fragment of the U1A splicosomal protein complexed with a 21-nucleotide RNA hairpin showed that the RNA binds as an open structure to the exposed β-sheet surface opposite the helices (Fig. 17.1.20A; Oubridge et al., 1994). Two loops from one end of the β-sheet protrude through the middle of the RNA loop, positioning and stabilizing the RNA structure on the sheet, and preventing base pairing. This enables sequence-specific interactions to occur between the bases and both main- and side-chain atoms on the β-sheet. Although it has no sequence homology with RNP proteins, the ribosomal protein S6 has been observed to have the same fold with a topology identical to the U1A protein (Fig. 17.1.20B; Lindahl et al., 1994). Similar α/β folding patterns have also been observed in the structures of other ribosomal proteins such as L6, L7/L12, and L30 (Lindahl et al., 1994). The bacteriophage T4 regA translational regulator protein also contains two regions that have sequence homology to the RNP1- and -2 sequence motifs. This protein specifically represses the translation of 35 early T4 mRNAs. The 1.9-Å crystal structure of T4 regA consists of two antiparallel β-sheets (three and four strands, respectively) and four helices (Fig. 17.1.20C; Kang et al., 1995). The larger β-sheet contains the RNP1 and -2 sequence motifs in two adjacent β-strands exposed to solvent, and superficially resembles the RNA-binding βsheet in the U1A protein.
The cold-shock proteins CspA and -B, from E. coli and Bacillus subtilis, respectively, are implicated in binding both RNA and singlestranded DNA, and each contain a region that has sequence homology with a canonical RNP domain. The structures of these proteins (Schindelin et al., 1993, 1994; Schnuchel et al., 1993; Newkirk et al., 1994), however, consist of a five-stranded antiparallel β-barrel known as the oligonucleotide/oligosaccharide (OB) fold (Fig. 17.1.21A and B; Murzin, 1993). The RNP sequence motifs reside in two adjacent β-strands suggesting a possible RNA-binding site. A similar β-barrel structure is also observed in the ribosomal proteins S17 (Golden et al., 1993) and L14, and in the N-terminal anticodon-binding domains of both lysyl- and aspartyl-tRNA synthetases (Fig. 17.1.21C-E; Cavarelli et al., 1993). Since the early 1990s, nearly 100 unique protein structures with OB folds have been deposited in the PDB, revealing a remarkably versatile fold used to bind not only RNA, but also ssDNA, oligosaccarides, and proteins (Arcus, 2002).
KH and dsRBP Domains Two other RNA-binding domains, the K homology (KH) and double-stranded RNAbinding domains (dsRBD), have structures that are similar in terms of general orientation of secondary structures (although not in topology) to the structure of the RNP domain. As with the RNP domain, these domains can be identified by a short consensus sequence. The three-dimensional structures for two dsRBD-containing proteins, E. coli RNase III (Kharrat et al., 1995) and Drosophila staufen protein (Fig. 17.1.22A; Bycroft et al., 1995), and one KH motif-containing protein, human vigilin (Fig. 17.1.22C; Castiglone Morelli et al., 1995), each contain a three-stranded antiparallel β-sheet with two or three helices packed against one face to form an α/β structure, albeit with different topology. Furthermore, a topological variant of the canonical KH domain has been observed in the ribosomal protein S3 from T. thermophilus (Carter et al., 2001; Grishin, 2001). Despite the superficial similarities in structure, the RNP, dsRBD, and KH domain RNA-binding structural motifs all appear to bind RNA by distinctly different modes. The dsRBD motif is proposed to interact with double-stranded RNA through the end loops of the β-sheet, the N-terminal end of the last helix, and a portion of one face of the β-sheet (Bycroft et al., 1995; Kharrat et al., 1995). The KH
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domain has been suggested to bind singlestranded RNA via the helical side of the domain (Gibson et al., 1993). The N-terminal domain of ribosomal protein S5 (Fig. 17.1.22B) from Bacillus stearothermophilus shares a number of conserved residues with dsRBD of staufen protein in addition to a similar topology, suggesting it may bind RNA in the same way as a dsRBD motif (Bycroft et al., 1995).
Zinc Finger Motif The zinc finger structural motif already described in this unit as a DNA-binding motif (see Zinc-Containing DNA-Binding Motifs), has also been implicated in binding RNA. A well characterized example is the TFIIIA protein which binds both 5S ribosomal RNA and the DNA encoding the 5S rRNA gene (Burd and Dreyfuss, 1994).
Four-Helix Bundle RNA-Binding Protein The RNA-binding protein Rop (repressor of primer) from an E. coli plasmid helps regulate plasmid copy number by facilitating sense-antisense RNA pairing. This protein functions as a homodimeric four-helix bundle structure comprised of two helix-turn-helix monomers (Fig. 17.1.23A; see above). Rop has no structural or sequence homology to any other known RNA-binding proteins (Predki et al., 1995). The results of mutation studies combined with binding assays provide strong evidence that RNA binding occurs along one face of the bundle formed by helices 1 and 1′ (Predki et al., 1995).
Large RNA-Binding Structures
Overview of Protein Structural and Functional Folds
MS2 phage coat protein One hundred and eighty copies of MS2 phage coat protein assemble to form the icosahedral viral capsid of the RNA bacteriophage MS2. Although comprised of the same amino acid sequence, the 129-residue MS2 subunits are present in the viral capsid in three structurally similar, but not identical, conformations referred to as A, B, and C (Valegard et al., 1994). Icosahedral symmetry results in the formation of AB and CC dimers arranged in the capsid as trimers of dimers (Fig. 17.1.23B). The dimers each consist of a ten-stranded antiparallel βsheet with two small β-hairpins and four helices packed against one face of the sheet. In addition to its structural role, this protein also prevents translation of viral replicase by binding to replicase mRNA. A nineteen-nucleotide RNA
hairpin of the Shine-Dalgarno sequence for viral replicase diffused into crystals of empty phage capsid was observed crystallographically to bind specifically to AB dimers of MS2 coat protein (Valegard et al., 1994). Crystal symmetry precluded crystallographic analysis of any RNA bound to the CC dimer. The RNA bound to AB interacts as a hairpin structure laying as a rigid crescent against the face of the β-sheet, opposite the helices, causing very little change in the protein structure relative to the unbound form of MS2. Trp RNA-binding attenuation protein Trp RNA-binding attenuation protein (TRAP) from Bacillus Subtilis binds to mRNA for the tryptophan (trpEDCFBA) operon in the presence of tryptophan, inducing the formation of a transcription terminator structure, thereby halting expression of the operon as tryptophan concentrations increase (Antson et al., 1995). The 1.8-Å crystal structure of TRAP in its activated form bound to tryptophan reveals a doughnut-shaped eleven-subunit oligomer containing 56% β-sheet (Fig. 17.1.23C). Each 75-residue subunit is composed of both a threeand four-stranded β-sheet packed face to face against each other. Tryptophan binds at one end of the interface between the two sheets in each subunit towards the center of the wheel. Adjacent three- and four-stranded β-sheets from neighboring subunits hydrogen bond to each other to form eleven seven-stranded β-sheets, each tilted in a way that resembles the blades of a turbine. Although the RNA binding site is not known, mutation analysis of both RNA and TRAP suggest that RNA containing eleven (G/U)AG elements forms a matching circle that binds to one side of the β-sheet wheel (Antson et al., 1995). Elongation factor Tu Elongation factor Tu (EF-Tu) is one of several elongation factors that participate in the translation of mRNA into a protein sequence on the ribosome. The crystal structure of EF-Tu from Thermus aquaticus in a ternary complex with Phe-tRNAPhe and a GTP analogue shows a three-domain structure in which all three domains interact with tRNA (Nissen et al., 1995). The N-terminal domain 1 is a structurally conserved G-protein domain which binds guanine nucleotide. Domains 2 and 3 are sevenand six-stranded β-barrels respectively. The 3′ CCA-Phe end of the tRNA binds in a cleft at the interface between domains 1 and 2 whereas the 5′ end of the tRNA binds at the interface of
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all three domains. The T-stem of the tRNA interacts primarily with the β-barrel in domain 3. Except for the terminal Phe side chain and terminal adenine, both at the 3′ end, EF-Tu interacts primarily with the ribose moieties and the phosphate backbone of the tRNA. Aminoacyl-tRNA synthetases Aminoacyl-tRNA synthetases (aaRS) are ATP-requiring enzymes that ligate amino acids to their cognate tRNAs during translation of RNA. The aaRS family of enzymes is remarkably diverse in terms of sequence, size, and three-dimensional structure. On the basis of sequence, structural analysis, and catalytic properties, aaRS can be divided into two classes, each comprising ten members (Eriani et al., 1990; Carter, 1993). As of early 2003, there are over 100 three-dimensional structures of aaRS and various complexes in the PDB (Table 17.1.8). These include bacterial and yeast aaRS for nineteen different amino acids (excluding alanine). Class I aaRS all share a classical dinucleotide-binding or Rossman fold in the catalytic domain (Fig. 17.1.24A) and generally acylate the 2′ hydroxyl of the terminal ribose (see The Classical Dinucleotide-Binding Fold for a discussion of the Rossman fold). The two sequence motifs, His-Ile-Gly-His and LysMet-Ser-Lys-Ser (more generally known as Met-Ser-Lys) are both present in all class I enzymes in regions that bind ATP. Class I enzymes are generally single-chain polypeptides, except for MetRS, TyrRS, and TrpRS which are homodimers. The ten class I aaRS are further divided into three subclasses: Ia, -b, and -c (Cusack, 1995; Ibba and Soll, 2000). Class II aaRS have a seven-stranded antiparallel β-sheet instead of a dinucleotide-binding fold in the catalytic domain (Fig. 17.1.24B) and primarily acylate the 3′ hydroxyl of the terminal ribose (except for PheRS which acylates the 2′ hydroxyl). The amino acid sequences of class II enzymes are identified by three different sequence motifs within the catalytic domain (Fig. 17.1.24B). The first sequence motif is involved in homodimerization, while the other two participate in catalysis. Eight class II enzymes are homodimers, two are α2β2 heterotetramers (GlyRS and PheRS) and one is a homotetramer (AlaRS). Note that GlyRS comes in two varieties, a homodimer (E. coli) and a heterodimer. It is also noteworthy that there are two types of LysRS: class I and II. Like their class I counterparts, the class II enzymes are also divided into three subclasses:
IIa, -b, and -c (Table 17.1.8; Cusack, 1995; Ibba and Soll, 2000). Despite the strong conservation of catalytic domains observed within each aaRS class, the structures and arrangements of the other domains involved in RNA binding and dimerization are remarkably variable, even within certain subclasses. AaRS domains include α-helical bundles, β-barrels, α/β f olds, and β-coiled-coils, among others. The sizes of single chains range from 300 to over 950 residues. Due to this variability, it is beyond the scope of this unit to describe each one or to develop structural classifications.
Signal Recognition Particle Nascent polypeptide chains synthesized by ribosome machinery are targeted to different cellular compartments. All proteins with signal peptides are sorted for secretion or membrane location via signal recognition particles (SRPs), a ubiquitous machinery throughout all kingdoms of life. Although the structure of a holo-SRP is yet to be determined, fragments of SRPs with or without their bound RNA, and a fragment of SRP receptor have been solved (Table 17.1.7). SRP domains Eukaryotic SRPs consist of six proteins organized into two domains, the Alu domain and the S domain. The function of the Alu domain is to retard the ribosome elongation of a nascent polypeptide chain for targeting to endoplasmic reticulum. The S domain forms the core of the SRP by binding to the signal peptide and targets it to the SRP receptor-mediated translocation machinery. Both domains are organized around SRP RNA and are connected by RNA. The Alu domain contains a heterodimer of SRP9 and -14 bound to the 5′ and 3′ terminal SRP RNA. The crystal structure of a mammalian SRP9/14 dimer in complex with the 5′ RNA domain, and a construct containing both the 5′ and 3′ RNA domains has been solved (Fig. 17.1.24C; Weichenrieder et al., 2000). SRP9 and -14 share a common fold of (αβββα topology with the two helices packed on the convex side of the three-stranded β-sheet. The SRP9/14 heterodimer is formed with the β-sheets aligning against each other forming a six-stranded antiparallel β-sheet with the four helices packed on one side of the β-sheet. The RNA domain that is recognized by SRP9/14 consists of the first 47 nucleotides of the 5′ RNA and 39 nucleotides from the 3′ RNA domain. The RNA assumes a helical conformation with a 180° re-
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verse bend in the middle at the SRP9/14 binding site. The S domain is formed by an SRP19, an SRP54 that contains a GTPase domain, an SRP68/72 heterodimer, and the core part of SRP RNA. The structure of a human SRP19 in complex with its cognate RNA has been determined (Fig. 17.1.24D; Wild et al., 2001). SRP19 folds into a KH-like (K-homology) fold (see KH and dsRBP Domains) with a central three-stranded antiparallel β-sheet packed against two helices on one side of the sheet. It is different from the fold observed in SRP9 and -14, even though both have two α-helices packed against a three-stranded β-sheet. The core SRP54 comprises three domains, the amino-terminal N domain, the GTPase domain G, and the methionine-rich M domain that contains binding sites for signal peptide and SRP RNA. The structure of a Thermus aquaticus Ffh, a homolog of mammalian SRP54, showed that the N domain forms a four-helical bundle, and the G domain assumes a Ras-like GTPase fold with an insertion that is unique to the SRP family of GTPases (Keenan et al., 1998). The M domain of Ffh in complex with domain IV of 4.5S RNA revealed that the five α-helices containing M domain recognizes the minor groove of SRP RNA by a helix-turn-helix motif (Fig. 17.1.24E; Batey et al., 2000). SRP receptor SRP receptor mediates the ER translocation of signal peptide containing proteins through binding to the signal peptide complexed SRP. The translocation depends on the hydrolysis of GTP through a GTPase domain of the receptor, the so-called NG domain. The structure of the GTPase-containing portion of FtsY, a functional homologue of the SRP receptor of E. coli, revealed a GTPase fold that is unique to SRPtype GTPases, including Ffh of SRP. Unlike the Ras-related GTPases, the SRP-type GTPases contain a small insertion domain of ∼29 amino acids called the I box, within the H-Ras fold. The I box comprises one β-strand flanked by two α-helices. Preceding the GTPase domain is a four helical bundle domain, called the N domain (Fig. 17.1.24F; Montoya et al., 1997).
Overview of Protein Structural and Functional Folds
binding monosaccharides, many lectins function as oligomeric proteins to increase the valency and thus, the affinity of binding toward carbohydrate ligands. Despite their functional similarity, neither the mode of oligomerization (dimers, tetramers, and pentamers have all been observed) nor the mode of carbohydrate recognition are strictly conserved in members of this family. Some members require Ca2+ or Mn2+ for saccharide binding, while others are thiodependent (Sharon, 1993; Drickamer, 1995; Rini, 1995a,b). Lectins can be divided into subtypes according to their structural homology. For the purpose of this overview, four more frequently observed subtypes are discussed: the legume and cereal lectins in plants, and the S-type and C-type lectins in animals (Table 17.1.9). β-trefoil lectins are discussed above (see Secondary Structure). Generally, members within each subtype of lectin share much higher degrees of structural conservation and conservation in carbohydrate recognition than the members of different subtypes.
Legume Lectins Lectins from various leguminous plants have very homologous three-dimensional structures and share similar locations for the carbohydrate recognition site. They often form dimers or tetramers and require both Ca2+ and Mn2+ ligated to each domain for carbohydrate binding. The general fold consists of a thirteenstranded β-sandwich formed from a sixstranded flat β-sheet packed against the convex face of a seven-stranded curled β-sheet. The carbohydrate-binding site is located on the outside concave portion of the seven-stranded βsheet (Fig. 17.1.25A; Dessen et al., 1995). Although the general topology (i.e., connectivity and strand order) is preserved in legume lectins, some members of the family do have deleted or inserted β-strands, or permutated termini. Lectins usually have their N and C termini close to each other in structure. This makes termini permutation particularly simple without disrupting the overall structure. For example, the N- and C-termini for ConA from jack bean are located at the respective N-terminal end of strand F and the C-terminal end of strand E of soybean agglutinin (Fig. 17.1.25A).
CARBOHYDRATE-BINDING PROTEINS
Cereal Lectins
Lectins represent a structurally diverse group of proteins that bind carbohydrate specifically. They are widely distributed throughout plants, animals, and even viruses, as well as bacterial toxins. Although often weak in
Wheat germ agglutinin (WGA) represents a member of plant chitin-binding lectins that are specific for both N-acetylglucosamine and Nacetylneuraminic acid (sialic acid). It functions as a homodimer of 17-kDa subunits. The X-ray
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crystal structure of WGA shows that each monomer contains four domains, each with four disulfides, and has a tertiary structure consisting mostly of loops and a few one-turn α-helices with no regular β-structure (Fig. 17.1.25B; Wright and Jaeger, 1993). Other structurally homologous cereal lectins include hevein (Rodriguez-Romero et al., 1991) and Urtica dioica agglutinin (UDA; Saul et al., 2000).
C-Type Lectins This family of animal lectins is characterized by a conserved ∼120 amino acid Ca2+dependent carbohydrate-recognition domain (CRD) that is often associated with other functional domains to mediate cell adhesion and other cellular signaling events. Members of this group include endocytic glycoprotein receptors, macrophage receptors, selectins, and the soluble collectins. Each CRD consists of a set of five to seven short β-strands forming two abutting β-sheets that are sandwiched between two α-helices. The carbohydrate-binding site is located in loop regions at one end of the β-sheet (I-G-H; Fig. 17.1.25C; Weis et al., 1992; Graves et al., 1994; Boyington et al., 1999). Calcium ions are directly involved in the binding of saccharide. A much larger superfamily of proteins referred to as C-type lectin-like proteins is now being structurally characterized. Many of these proteins do not have calcium or carbohydrate binding sites even though they share the same fold. These include numerous cell surface receptors, particularly in the immune system.
S-Type Lectins S-lectins are a family of thio-dependent, β-galactoside-binding proteins that have been proposed to play important roles in modulating cell-cell and cell-matrix interactions (Hirabayashi and Kasai, 1993). Galectins are the best known examples of this family. They often form dimers or associate with other accessory domains. Crystal structures of galectin-1 and -2 show a striking structural homology with legume lectins with the exception of a few inserted and deleted strands (Liao et al., 1994; Lobsanov et al., 1993). Each monomer folds into a curled β-sandwich with the flat sheet consisting of five β-strands and the curled sheet consisting of six β-strands (Fig. 17.1.25D). Although the carbohydrate binds on the concave portion of the curled sheet, similar to those in legume lectins, the specific mode of saccha-
ride recognition is not conserved between legume lectins and galectins.
CALCIUM-BINDING PROTEINS EF-Hand Calcium-Binding Motif The EF-hand is the most commonly observed Ca2+-binding motif in proteins (Satyshur et al., 1988; Nakayama and Kretsinger, 1994). As of early 2003, the SMART database listed over 2000 proteins with this domain (see Web-Based Structural Bioinformatics and http://smart.embl-heidelberg.de/browse.shtml) . Examples of EF-hand-containing proteins include calmodulin, troponin C, essential lightchain and regulatory light-chain myosin, parvalbumin, intestinal and sarcoplasm calciumbinding proteins, calcineurin B, recoverin, and extracellular matrix protein BM-40 (Table 17.1.10). There are two kinds of Ca2+-binding sites among the EF-hand motif proteins: high and low affinity. The high affinity Ca2+-binding sites appear to primarily provide structural stability to proteins, whereas the low affinity sites often function to regulate the active protein conformations, and hence are also termed regulatory sites. For example, calmodulin undergoes a conformational change upon binding of Ca2+ ions to the regulatory sites (Meador et al., 1992). Both crystal structures and NMR structures of a number of EF-hand proteins have been solved (Herzberg and James, 1988; Satyshur et al., 1988; Meador et al., 1992; Svensson et al., 1992; Flaherty et al., 1993; Rayment et al., 1993; Findlay et al., 1994; Xie et al., 1994; Hohenester et al., 1996). A typical EF-hand consists of a helix-loop-helix arrangement with the two helices oriented ∼90° to each other and the Ca2+-binding site located in the loop region (Fig. 17.1.26A). In most situations, they form pairs with the two EF-hands in a pair related by approximately two-fold symmetry. The Ca2+ coordination is usually described as a pentagon bipyramid geometry with a total of six to seven ligands which are from the carboxylate groups of Asp, Glu, and main chain carbonyl oxygens in the loop region. Water molecules are often found to provide one of the apical coordinations.
Annexins Annexins are a family of calcium-dependent phospholipid-binding proteins. Unlike the classical EF-hand calcium-binding proteins, the annexins chelate calcium in an entirely different way (Huber et al., 1990a,b, 1992; Bewley et al., 1993). Each calcium-binding
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domain comprises five alpha helices, A through E (Fig. 17.1.26B). The coordination geometry is that of a pentagonal bipyramid with three ligands from the backbone carbonyl oxygen groups of the residues in the AB loop (the loop between helices A and B). The three ligands are each separated by one amino acid in the sequence. Two other ligands are provided by the side chain carboxylate of a Asp or Glu residue, located at the end of helix D, 44 amino acids downstream from the first carbonyl ligand of the AB loop. The six and seventh ligands are less conserved and can be replaced with water molecules.
NUCLEOTIDE-BINDING DOMAINS Nucleotide-binding folds first began to be characterized in the early 1970s as the structures of several NAD cofactor-containing dehydrogenases were elucidated (Rossmann et al., 1975). It was observed that single domains with conserved folding patterns were responsible for nucleotide-binding in many proteins. With the structural determination and analysis of many more nucleotide-binding proteins, this particular fold has been divided into three primary classes: dinucleotide-binding domains, the mononucleotide-binding fold, and the actin fold; however, not all nucleotide-binding proteins have these folding patterns. Proteins such as the class II aaRS (containing an antiparallel β-sheet for binding ATP) and some FMN-binding proteins (TIM-barrel structures), among others, have very different folds for interacting with single nucleotides. To avoid confusion, the di- and mononucleotide-binding folds discussed in this section (Table 17.1.11) are usually referred to in the literature as classical or canonical nucleotide-binding folds and the dinucleotide-binding fold is sometimes referred to simply as the Rossman fold.
The Classical Dinucleotide-Binding Fold
Overview of Protein Structural and Functional Folds
In general, classical (canonical) dinucleotide-binding proteins fall into one of three groups: NAD(P)-dependent dehydrogenases and oxidoreductases, FAD-linked oxidoreductases, and FAD/NAD-linked oxidoreductases. Although many of these proteins are multidomain α/β structures, their dinucleotide-binding domains typically have a common three-dimensional fold. These binding domains contain a central five- or six-stranded parallel sheet, strand order 3-2-1-4-5-6, connected to helices on each face through right-handed loops (Table 17.1.11; Fig. 17.1.27A). The conserved topol-
ogy for this α/β/α sandwich motif as compiled by G.E. Schultz (1995) and A.M. Lesk (1995) is illustrated in Figure 17.1.27B. In each case, the dinucleotide binds across the C-terminal end of the sheet with the ADP portion interacting specifically with the central βαβ unit of this fold (β1-α1-β2). The core sequence motif (also called the fingerprint sequence) for this βαβ fold is Gly-X-Gly-X2-Gly (where X is any amino acid) and is located in a loop between β-strand 1 and helix 1. This sequence permits unusual main chain dihedral angles in the binding loop and close contacts with ADP. The pyrophosphate moiety sits at the N-terminal end of the helix following the loop, a favorable environment for the negative charges (Wierenga et al., 1985). Another conserved feature is an Asp or Glu residue at the C-terminal end of the second β-strand of the βαβ unit for binding the ribose moieties of NAD and FAD, respectively. NADP-binding domains contain an Arg at this position for binding the 2′-phosphate (Schultz, 1992). One additional eleven-residue sequence motif has been located in β-strand 5 of the canonical dinucleotide binding fold in FAD-binding domains which includes a C-terminal aspartic acid that interacts with the ribitol moiety of FAD (Eggink et al., 1990).
The Classical Mononucleotide-Binding Fold First noted in the structure of adenylate kinase (Fig. 17.1.27C; Schulz et al., 1974), the mononucleotide-binding fold is superficially very similar to the dinucleotide-binding fold (above), but has a different topology. This fold also consists of an α/β/α sandwich with a central five-stranded parallel β-sheet and righthanded connections to helices (Table 17.1.11, Fig. 17.1.27C and D; Schultz, 1992). The strand order is 2-3-1-4-5. As in the case of the dinucleotide-binding fold, mononucleotide binds to a critical glycine-rich loop at the C-terminal end of the β-sheet between strand 1 and helix 1. This loop (also referred to as the P-loop) is much longer than the shorter dinucleotidebinding loop, enabling it to form a giant positively charged anion hole for interacting with the mononucleotide triphosphate moiety (Dreusicke and Schulz, 1986). The loop’s core sequ en ce mo tif, Gly-X2-Gly-X-Gly-Lys (where X is any amino acid) has been used to identify many more mononucleotide-binding proteins including some from the more distantly related GTP-binding proteins (also known as G proteins) which also have a P-loop (Saraste et al., 1990; Schultz, 1992; Traut,
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1994). Slight variations of the mononucleotidebinding fold include the flavodoxin-like and the resolvase-like proteins. In these families, the first β-strand is displaced to create a strand order of 2-1-3-4-5 instead of 2-3-1-4-5. Furthermore, in the resolvase-like fold, the fifth β-strand is antiparallel to the rest. G proteins have an extra sixth strand in the β-sheet with strand order 2-3-1-4-5-6 and the second βstrand is antiparallel to the rest. Other variations include protein structures that do not contain the canonical P-loop fingerprint sequence— e.g., CheY chemotactic protein; Volz and Matsumura, 1991), γδ-resolvase (Sanderson et al., 1990), flavodoxin (Watenpaugh et al., 1972). Further perturbations of the mononucleotidebinding fold are listed under the heading Fold: P-loop Containing Nucleotide Triphosphate Hydrolases in the SCOP database (see WebB ased Stru ctu ral Bio informatics and http://scop.mrc-lmb.cam.ac.uk/scop/index. html). It should also be noted that some mononucleotide-binding proteins, such as adenylate kinase, also bind monophosphate nucleotides in a distinctly different binding site, albeit close in proximity to the classical site discussed in this section.
ENZYMES INVOLVED IN DNA/RNA REPLICATION AND DNA TRANSCRIPTION DNA/RNA Polymerases Polymerases are fundamentally important enzymes responsible for replication, repair, and transcription of DNA and RNA in all living organisms. The structures of these enzymes vary widely from small single-subunit proteins of only a few tens of kilodaltons in size to large multisubunit complexes approaching 1000 kDa (Joyce and Steitz, 1994). DNA polymerases and small RNA polymerases DNA polymerase and some small RNA polymerases have a central catalytic domain responsible for DNA binding, nucleotide binding, and phosphoryl transfer. The three-dimensional structures of the single-chain catalytic domains of several different nucleic acid polymerases from eukaryotic, prokaryotic, and viral organisms have been solved (Table 17.1.12). These include DNA polymerase structures from the Pol I (A family), Pol α (B family), Pol β (X family), lesion-bypass polymerase (Pol Y family), and reverse transcriptase (RT) families. In addition, there are structurally
homologous eukaryotic poly(A) RNA polymerases and viral RNA polymerase structures from the T7 RNA polymerase family, the viral RNA-dependent RNA polymerase family, and the double-stranded (ds)phage RNA–dependent RNA polymerase family. Including mutants and structures of polymerases complexed with DNA, nucleotides, drugs, and various salts, the number of PDB entries is now in the hundreds (see Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb). Despite numerous structural differences, the single-chain catalytic domains of the abovementioned polymerases each possess a common arrangement of subdomains centered around a large nucleotide-binding cleft which can be loosely described as a U-shaped structure resembling a person’s right hand. Using nomenclature introduced by Ollis et al. (1985), the central base of the structure is referred to as the palm subdomain, and the two vertical walls of the U are called the fingers and thumb subdomains (Fig. 17.1.28A). It is noteworthy that bovine and yeast poly(A) RNA polymerases do not have a thumb subdomain (Bard et al., 2000; Martin et al., 2000). The palm catalyzes the phosphoryl transfer reaction, the fingers are involved in template positioning, and the thumb may be important in DNA binding and processivity (Hubscher et al., 2002). Structures of polymerase/DNA and polymerase/DNA/dNTP complexes suggest that the thumb and finger subdomains in some cases make dramatic movements during different stages of the polymerization reaction (Fig. 17.1.28B; Beese et al., 1993; Pelletier et al., 1994; Franklin et al., 2001). Associated catalytic functions distinct from polymerase activity, such as the 3′-5′ exonuclease activity in the Klenow fragment (KF) of E. coli DNA polymerase I, RNase activity in RT, 5′-nuclease activity of pol-β and Taq polymerase, and nascent RNA binding by T7 RNA polymerase, are located in separate N- or C-terminal domains. Still more related functions reside in associated subunits. The central palm is the most structurally conserved subdomain among polymerases. It consists of a threeto six-stranded antiparallel β-sheet with two helices packed against one face of the sheet (Fig 17.1.28C), a structure similar to the RNP motif found in many RNA-binding proteins (see KH and dsRBP Domains). The exposed side of the sheet displays the conserved active site Asp residues. With regard to this subdomain, however, the eukaryotic β polymerase and eukaryotic poly(A) RNA polymerases are unique (Davies et al., 1994; Sawaya et al., 1994; Bard
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et al., 2000; Martin et al., 2000). Although the palm subdomain has a similar α/β sandwich fold, the overall topology is quite different (Fig 17.1.28D). In contrast to the palm subdomain, the fingers and thumb subdomains appear to be functionally related rather than structurally analogous between polymerase domains. For example, the thumb subdomains of KF, Taq polymerase, and T7 RNA polymerase all consist of a helical bundle with one or two long helices; however, the thumb subdomains of pol-β and RT are α/β structures each having three α-helices and one or two β-strands, respectively. Finger subdomains are similarly divergent. The Y family polymerases have an additional conserved α/β domain sometimes referred to as the little finger (Ling et al., 2001). The eukaryotic poly(A) RNA polymerases have an additional C-terminal domain with an RRM-like fold (see The RNP Domain) that is known to interact with the RNA primer (Bard et al., 2000; Martin et al., 2000). Not only are the folds of the subdomains variable, but the order of the subdomains within the primary sequence differs as well in various polymerases (Sawaya et al., 1994). Moreover, RT functions as a heterodimer (p66/p51) while other homologous polymerase domains function as monomers. Interestingly, the p51 subunit of RT results from the cleavage of the C-terminal RNase H domain from p66. Thus, both p51 and p66 have the structurally equivalent thumb, fingers, and palm subdomains. In addition, both RT subunits also contain another α/β subdomain termed the connection subdomain, which bridges the RNase H and polymerase domains in the larger subunit (Jacobo-Molina et al., 1993). P66 and p51 differ dramatically in the spatial arrangement of their subdomains (Fig. 17.1.28E), resulting in two morphologically and functionally asymmetric subunits, with the polymerase activity residing only in the larger p66 subunit (Jacobo-Molina et al., 1993; Joyce and Steitz, 1994).
Overview of Protein Structural and Functional Folds
RNAP The bacterial, archeal, and eukaryotic DNAdependent RNA polymerases (RNAPs) are responsible for the transcription of DNA into RNA during gene expression. All known RNAPs including eukaryotic RNAP I, II, and III are members of a conserved protein family also known as the multisubunit RNAP family (Ebright, 2000). Structurally, RNAPs superficially resemble the DNA and viral RNA polymerases described above in that they possess a U-shaped structure; however, RNAPs are much
larger, more complicated multisubunit enzymes that have a rather different architecture. The crystal structures of the prokaryotic core RNAP and holoenzyme RNAP from Thermos aquaticus, and eukaryotic RNAP II from Saccharomyces cerevisiae each reveal a large crabclaw-shaped molecule with a molecular weight of ∼400 to 500 kDa (Fig. 17.1.28F and H; Zhang et al., 1999; Cramer et al., 2000, 2001; Murakami et al., 2002a; Vassylyev et al., 2002). The active site is found in the bottom of a deep channel between the two pincers of the claw. For clarity, most of following description of RNAPs is based on the simpler structure of the bacterial form rather than the structure from yeast. The bacterial core RNAP is composed of five subunits (α2ββ′ω). The holoenzyme additionally incorporates a σ initiation factor that is essential for transcription initiation. In general terms, the β and β′ββ′ω subunits make up the two arms of the claw, respectively, and the two identical α subunits cap the end of the molecule opposite the claw opening (Fig. 17.1.28F). The ω subunit sits on the outer surface of the β′ subunit and the σ70 subunit straddles the ends of both pincers on one side of the claw, interacting with β and β′. The α subunit N-terminal domain (NTD) consists of two α/β subdomains (Fig. 17.1.28G; Zhang and Darst, 1998; Zhang et al., 1999). Subdomain 1 is a four-stranded antiparallel β-sheet packed against two nearly orthogonal α-helices along one face. The second subdomain comprises a three-stranded antiparallel β-sheet flanked on one face by two twostranded β-sheets and one α-helix. The two α-subunits dimerize through antiparallel helix interactions of the two helices in subdomain 1. The C-terminal domain (CTD) of the α subunit is not visible in the RNAP complex, but NMR studies have revealed a compact fold consisting of four small helices. The first three helices are nearly perpendicular to each other, while the fourth is antiparallel to the third (Jeon et al., 1995). The ω subunit consists of a small four-helix bundle and a β-hairpin. The σ70 initiation factor of the T. aquaticus holoenzyme is an elongated all-helical protein comprising three domains (Fig. 17.1.28F, dark blue). The second and third domains are separated by a long linker region (Murakami et al., 2002a; Vassylyev et al., 2002). Crystal structures of E. coli σ70 and T. aquaticus σA factors reveal similar all-helical subunits (Malhotra et al., 1996; Campbell et al., 2002). A helix-turn helix motif (see Helix-
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Turn-Helix Motif) in σA is observed to make critical interactions with the promoter DNA (Campbell et al., 2002). The β and β′ subunits are rather complicated α/β polypeptides with eight and six structural domains, respectively. Due to their complexity, these two subunits are not described in this overview. For a detailed analysis of these two subunits, see Cramer et al. (2001) for the structurally homologous yeast RNAP II in which subunits RBP1 and -2 are equivalent to β′ and β subunits, respectively. The RNAP II complex from yeast consists of ten subunits (RBP1 to -10; Fig. 17.1.28H). Five of the yeast subunits (RBP1, -2, -3, -11, and -6) correspond in structure and function to the five bacterial subunits (β′, β, αI, αII, and σ, respectively). In fact, R. Ebright (2000) has detailed remarkably extensive structural similarity between the bacterial and yeast RNAP complexes ranging from the general subunit locations to the folding topologies of individual domains. The yeast RNAP II structure has an additional five subunits, most of which decorate the outer surface of the ‘claw’, away from the internal channel (Fig. 17.1.28H; Cramer et al., 2001).
DNA Polymerase Processivity Factors Certain DNA polymerases, such as those involved in replication, require a highly processive polymerase complex. The characteristic of processivity is conferred upon polymerases by associated processivity factors that tether the polymerase to DNA in an ATP-dependent manner in order to prevent dissociation (Kong et al., 1992). One type of processivity factor known as a sliding clamp is a simple ring-shaped structure that both encircles DNA and interacts with the respective polymerase. The three-dimensional structures of several of these sliding clamps, including the β-subunit homodimer of pol III, the RB69 and T4 bacteriophage processivity factors, and PCNA, are each starshaped six-fold symmetric closed circular rings consisting of a circle of twelve α-helices perpendicular to the ring plane surrounded by six antiparallel β-sheets (Kong et al., 1992; Krishna et al., 1994; Shamoo and Steitz, 1999; Moarefi et al., 2000). The inside diameter of both rings is 30 to 35 Å and the outer diameters are each ∼80 Å. Each ring is composed of six nearly identical domains, each of which is twofold symmetric, consisting of two fourstranded antiparallel β-sheets packed edge-toedge at a ∼45° angle with two antiparallel helices packed against the concave side of the
domain (Fig. 17.1.29A). The β-sheets of adjacent domains hydrogen bond to each other forming six curved eight-stranded antiparallel β-sheets that line the outer surface of the ring. The complete ring structure results from the 12-fold repeat of the simple topological structural motif βαβββ round the entire circle (Fig. 17.1.29B). Despite the striking structural similarity, there is only 15% sequence identity when individual domains from each protein are aligned (Krishna et al., 1994). Sequence alignments of individual domains within the same protein give similar results. A head-to-tail dimer of the β-subunit of pol III forms the entire six-fold circle; each subunit comprises three domains. The bacteriophage RB69 and T4 rings, and the PCNA ring consist of trimers with two domains per subunit. ATP and auxiliary proteins are required for assembly around duplex DNA and dissociation does not occur easily, unless the DNA is linearized. In each case the ring has a substantial net negative charge and positively charged side chains are concentrated on the twelve α-helices lining the central channel, which should allow favorable interaction with the phosphate backbone of DNA passing through the channel (Kong et al., 1992; Krishna et al., 1994). In addition, the orientation of helices facing the channel is such that they are nearly perpendicular to the phosphate backbone of B-form DNA, precluding specific interaction with the grooves. These physical characteristics, as well as the size of the inner diameter (30 to 35 Å), make both proteins well suited for encircling DNA in a manner that allows for unrestricted movement along a duplex DNA strand during polymerization. Finally, direct interaction has been observed between the T4 and RB69 sliding clamps and the corresponding DNA polymerases (Shamoo and Steitz, 1999). In other systems, intermediate proteins facilitate this polymerase linkage.
Topoisomerases Topoisomerases are essential enzymes of eukaryotes, prokaryotes, and viruses that alter the topology of double-stranded DNA. They function by transiently breaking one or two strands of DNA, passing DNA through the opening, and then resealing the break (for reviews, see Wigley, 1995; Champoux, 2001). Topoisomerases can be divided into two primary classes: type I enzymes which transiently cleave only one strand of DNA, and type II enzymes which cleave both DNA strands during catalysis. Structural Biology
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Overview of Protein Structural and Functional Folds
Type I topoisomerases Type I enzymes are monomeric proteins which relax supercoiled DNA in an ATP-independent manner. They can also catenate and decatenate nicked double-stranded circles and form a double-stranded DNA circle from two complementary single-stranded circles. Subtype IA enzymes form a link to the 5′ phosphate of the DNA and subtype IB enzymes link to the 3′ phosphate. Subtype IA. The three-dimensional structures of the N-terminal catalytic fragments of two subtype IA enzymes from E. coli, topoisomerases I and III, reveal a common structural architecture (Lima et al., 1994; Mondragon and DiGate, 1999). The N-terminal fragment is made of four distinct domains arranged around a large central channel with a diameter of 27.5 Å (Fig. 17.1.30A). Domain I is a four-stranded parallel β-sheet sandwiched between two pairs of α-helices, a fold similar to the classical dinucleotide-binding fold (see above). Domain II is an unusual looking β-sandwich composed of two long-curving three-stranded antiparallel β-sheets that cross each other orthogonally at one end of each sheet. A single helix lies between the sheets at one end of the sandwich. The β-strands of this domain line the inside of the top half of the central channel, bearing a striking resemblance to the TATA-box binding protein (see DNA-Binding Proteins that use β-Sheet Motifs; Burley, 1996). Domains III and IV are each primarily helical except for a small β-hairpin in domain IV. Domain III contains the active-site tyrosine, Tyr319, and closes one side of the central channel through an extensive interface with domains I and IV (Fig. 17.1.30A). This interface effectively buries Tyr319, suggesting that large conformational changes are required during catalysis. It has been proposed that domains II and III may move together as a rigid body away from domains I and IV, using a protease-sensitive region between domains II and IV as a hinge (Lima et al., 1994). Such a movement would open a gate to the central channel and expose the active site Tyr319. The size of the closed channel is large enough to accommodate double-stranded B-form DNA without steric hindrance. The inner surface of the channel has a strong positive potential suitable for interaction with DNA. Subtype IB. Eukaryotic subtype IB topoisomerases share no structural or sequence similarity with prokaryotic subtype IA enzymes. Structural studies of an N-terminally truncated form of human topoisomerase IB (excluding an
unstructured, highly charged ∼200 residue Nterminal domain) in complex with a 22-base pair DNA duplex reveal a five-domain protein structure that wraps around the DNA (Redinbo et al., 1998; Stewart et al., 1998; Fig 17.1.30B and C). Core subdomains I, II, and III are each α/β folds, the linker domain is a two-helix antiparallel coiled-coil, and the C-terminal domain is composed of five short helices. Subdomain II is noted to resemble the fold of the homeodomain family despite only 11% sequence identity and no evidence from the topoisomerase structure that it interacts with DNA (Redinbo et al., 1998). Additionally, subdomain III shows striking structural similarity to bacteriophage HP1 integrase (Redinbo et al., 1998). Finally, the coiled-coil of the linker domain superimposes with the Rop RNA-binding protein (see Four-Helix Bundle RNA-Binding Protein) with a root-mean square (rms) distance of 0.8 Å for 60 Cα atoms (Stewart et al., 1998). The catalytic Tyr723 resides in the C-terminal domain within the positively charged channel region. Like prokaryotic topoisomerase IA, the central pore is lined with positively charged side chains for interaction with DNA, and concerted movement of subdomains I and II is proposed to take place during the catalytic mechanism, allowing DNA to enter and exit the complex; however the proposed mechanisms for the two enzymes are quite different (Redinbo et al., 1998; Stewart et al., 1998; Lima et al., 1994). Type II topoismerases Type II enzymes are multimeric proteins with homologous amino acid sequences. The eukaryotic type II topoisomerases are homodimers, while the prokaryotic enzymes are A2B2 tetramers where the A and B subunits correspond to the N- and C-terminal halves of the eukaryotic monomer, respectively; viral enzymes can have either subunit composition. Type II topoisomerases require ATP for catalysis and are capable of either relaxing DNA or introducing supercoils. Like the type I enzymes, they can also catenate or decatenate circular DNA. Crystal structures of the C-terminal half of yeast topoisomerase II, the homologous E. coli gyrA (gyrase subunit A), and the ATPase-containing subunit (gyrB) of E. coli gyrase complement each other and together provide a model for the structure of an intact type IIA topoisomerase. A crystal structure of the A subunit from Methanococcus jannaschii DNA topoisomerase VI provides insight regarding type IIB topoisomerases.
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Subtype IIA. The monomer structure of the C-terminal 92-kDa yeast topoisomerse II fragment reveals a modular crescent-shaped protein with five domains (Fig. 17.1.30D). Berger et al (1996) divided the polypeptide chain into two subfragments, A′ and B′, based on their relation to the A and B subunits of the homologous bacterial enzymes. The N-terminal subfragment B′ has two domains, a Rossman fold–like five-stranded mostly parallel β-sheet (see The Classical Dinucleotide-Binding Fold for a discussion of the Rossman fold) sandwiched between four helices and a three-stranded antiparallel β-sheet packed against one helix. The C-terminal subfragment A′ has three domains. Domain I consists of a winged-helix motif (see above) of three helices and a three-stranded antiparallel β-sheet embedded between an Nterminal α-helix, two short C-terminal α-helices, and a β-hairpin. The active site tyrosine residue, Tyr783 (Fig. 17.1.30E), is located in a loop of the winged-helix at the bottom of a short tunnel leading to the protein surface. Domain II comprises a four-stranded antiparallel βsheet packed against two helices on one face and an abutting three-stranded antiparallel βsheet. The third domain of subfragment A′ consists of a six-helix bundle and a small βhairpin. Three of the helices are unusually long, causing this domain to project out from the rest of the protein. In the initial crystal structure, the two monomers come together to form a flat heart-shaped dimer with a large pear-shaped central cavity 60 Å high and 55 Å wide towards the base (Fig. 17.1.30E). Although the primary dimer contacts occur at the A′-A′ interface, the B′-B′ interface involves extensive overlap of domains such that the B′ subfragment from one monomer interacts with the A′ subfragment of the other. At the interface between the first two domains in subfragment A′, a 20- to 25-Å groove runs across the protein surface including the region containing Tyr783. B-form DNA can be modeled into this groove with the recognition helix of the winged-helix motif lying in the DNA major groove (see Helix-Turn-Helix Motifs). A second crystal form of yeast topoisomerase II (Fass et al., 1999) and the structure of E. coli GyrA (Morais Cabral et al., 1997) reveal similar shaped dimers, but significant alterations in subdomain orientations between the three structures suggest a flexible complex that undergoes conformational changes during catalysis. The crystal structures of yeast topoisomerase II lack the N-terminal ATPase domain that is proposed to lie on top of the B′ subfragments; however, two crystal
structures of the ATPase domain of the homologous E. coli gyrase B have been determined, enabling the modeling of the entire topoisomerase II structure (Wigley et al., 1991; Brino et al., 2000). The 43-kDa fragment of DNA gyrase B is an elongated molecule consisting of two α/β domains each with a unique fold (Fig. 17.1.30F). The first domain, which contains the ATPase site, is an eight-stranded mostly antiparallel β-sheet with five α-helices packed against the convex face. It binds ADPNP at the center of one face of the β-sheet with the phosphates coordinated by a glycine-rich loop at the N-terminus of a helix. The second domain (also referred to as the DNA capture domain) is a four-stranded mixed β-sheet surrounded by four helices, including one long C-terminal helix that protrudes away from the protein. In the gyrase B dimer, the most extensive interaction is through the N-terminal domains (Fig. 17.1.30F). A long 15-residue N-terminal arm from each monomer wraps around the other monomer contributing to the ATP-binding site. It is interesting to note that the binding of ATP is reported to induce dimerization (Berger et al., 1996). The C-terminal domain forms dimer contacts only through one end of the protruding helix from each monomer. This dimer structure creates a large 20 Å-wide hole between the two C-terminal domains (Fig. 17.1.30F). The C-terminal half of this hole is observed to be rich in Arg residues, suggesting possible interaction with DNA. Subtype IIB. The crystal structure of a truncated form (missing residues 1 to 68) of the A subunit from Methanococcus jannaschii DNA topoisomerase VI reveals a U-shaped homodimer (Nichols et al., 1999; Fig. 17.1.30G). Each monomer consists of two domains linked by a short tether. The N-terminal domain has five α helices packed against a two-stranded β-sheet with a fold similar to the winged helix motif observed in catabolite activator protein (CAP; see Helix-Turn-Helix Motifs). The second domain consists of three subdomains. Subdomain I is a five-stranded antiparallel β-sandwich. Subdomain II has a Rossman fold consisting of a four-stranded parallel β-sheet flanked on each face by a pair of α helices ribose (see The Classical Dinucleotide-Binding Fold for a discussion of the Rossman fold). The last subdomain is an α/β structure comprising five α helices and a small two-stranded β-sheet. Apart from the winged helix motif, the Rossman fold, and a large central channel, this dimer is quite different from the known topoisomerase type I Structural Biology
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and II structures (see above). The only noted similarity is in the Rossman fold.
Polynucleotidyl Transferase RNase H–Like Fold The structures of a number of enzymes involved in polynucleotide transfer have been observed to have the same basic fold as ribonuclease H (RNaseH; Katayanagi et al., 1990, 1992; Yang et al., 1990). Members of this family include RNase H, retroviral integrase, Holiday junction RuvC resolvase, TN5 transposase, the RNase H domain of retroviral reverse transcriptase, a core fragment of bacteriaphage Mu transposase, and the 3′-5′ exonuclease domains from several DNA polymerase structures, including the Klenow fragment (Ollis et al., 1985; Davies et al., 1991, 2000; Ariyoshi et al., 1994; Dyda et al., 1994; Rice and Mizuuchi, 1995). The sequence identity among family members is generally <15%. At the center of the structure is a five-stranded β-sheet, aligned with a strand order of 3-2-1-4-5. The second strand is antiparallel to the other four strands (Fig. 17.1.31). The number and location of the helices are less conserved than the central β-sheet. Helices A and E are topologically more conserved than the other helices. They are located between strands 3 and 4, and at the C-terminal of strand 5 in sequence, primarily interacting with the central β-sheet. The number of helices between strands 4 and 5 varies, as do the helices after helix E (Fig. 17.1.31). The family can be further divided into RNase- (e.g., HIV-1 reverse transcriptase RNase H, E. coli RNase H, Archeon Class II RNase H) and integrase-like structures (e.g., integrase, Mu A transposase, RuvC) with greater structure resemblance observed between the members of each subclass. Limited structural homology, particularly among the central five-stranded β-sheet, is also observed with Mut S domain II, actin-like ATPase domains, and ribosomal proteins L18 and S11.
PROTEINS INVOLVED IN CYTOSKELETON AND MUSCLE MOVEMENTS
to as the actin fold, despite the absence of any significant sequence homology (Table 17.1.13). This fold consists of two α/β domains connected by a hinge region with the ATP binding site between them (Fig. 17.1.32A-D; Kabsch and Holmes, 1995). Each domain can be further divided into two subdomains. Two of the four subdomains (subdomains 1 and 3 according to actin notation) share a conserved core, a five-stranded β-sheet (with only one antiparallel strand) that is assembled as a β-meander surrounded by three helices. ATP binds at the interface between the β-sheets of subdomains 1 and 3. These two β-sheets are connected by two conserved helices and have identical topology, suggesting a gene duplication event. The other subdomains (2 and 4) are not as well conserved among these proteins, and may carry out specialized functions in different proteins sharing this fold. In actin, hsc70, and glycerol kinase, subdomains 2 and 4 are each α/β regions positioned at the C-terminal end of the sheets of the actin fold. Subdomain 4 has identical topology in the above three proteins, but a different tertiary structure in glycerol kinase (Hurley et al., 1993). Hexokinase only has subdomain 4 (subdomain 2 is absent), which is an α-helical bundle.
Actin-Binding Proteins The assembly of actin to form filament, where it is the major component, and cortex is regulated by several actin-binding proteins, including profilin, gelsolin, severin, and villin. These molecules, through their association with actin, not only regulate cellular locomotion and shape, but also function as signaling molecules and keep actin filament in a dynamic balance between the free monomeric and polymerized forms. For example, profilin inhibits the assembly of F-actin by sequestering monomeric actin molecules, whereas gelsolin controls the actin cortex formation through fragmentation of F-actin filaments. The threedimensional structures of several of these actin-binding proteins has revealed their folds and helped to elucidate their functions in filament formation.
The Actin Fold
Overview of Protein Structural and Functional Folds
The three-dimensional structures of several ATP-binding proteins, including actin (Kabsch et al., 1990), the N-terminal domain of heatshock cognate protein (hsc70; Flaherty et al., 1990), hexokinase (Anderson et al., 1978), glycerol kinase (Hurley et al., 1993), and acetate kinase (Buss et al., 2001), have been found to contain a conserved structural motif referred
Profilin Both the X-ray crystal and NMR solution structure of profilin determined to date show a single domain fold with a central five-stranded antiparallel β-sheet pack against four α-helices, two on each side of the sheet (Fig. 17.1.32E; Schutt et al., 1993). The β-sheet is curled toward the N-terminal α-helices on one side and
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the five strands are connected in the order of 1-2-3-4-5. Profilin, when complexed with βactin, forms two major contacts with its third helix, as well as the amino-terminal portion of the fourth helix and strands 4, 5, and 6. Gelsolin The members of the gelsolin superfamily include gelsolin, villin, and severin. The structure of the segment 1 domain of gelsolin shows the basic fold, which consists of a central fivestranded β-sheet flanked by two α-helices, one on each side of the sheet (Fig. 17.1.32F; McLaughlin et al., 1993). The central β-sheet contains mostly antiparallel strands with the exception of the last strand of the sheet, which is in parallel configuration. Spectrin repeats Spectrins are actin cross-linking proteins. Their associations with actin generate meshworks to support the plasma membrane and facilitate cellular interactions. The so-called spectrin repeat defines a segment of 100 to 120 amino acid and folds into a three-helix bundle (Fig. 17.1.32G; Yan et al., 1993). The observed repeats form a stable domain-swapped dimer in which the third helix of each monomer is exchanged in the helix bundle.
ENZYMES The α/β Hydrolase Fold A distinct protein fold, observed among several hydrolytic enzymes, has emerged in the last few years. These enzymes, including serine carboxypeptidases, microbial lipases, esterases, dehalogenases, peroxidases, and various small molecule-hydrolyzing enzymes, such as dienelactone hydrolase, share little or no detectable sequence homologies among them and differ widely in their catalytic functions (Table 17.1.14; Ollis et al., 1992; Nardini and Dijkstra, 1999). Nonetheless, they share the same structural topology and furthermore a conserved catalytic triad consisting of a nucleophile (Ser, Cys, or Asp) at the end of β-strand 5, an absolutely conserved His after β-strand 8, and an Asp or Glu residue at the end of β-strand 7 (Liao and Remington, 1990; Franken et al., 1991; Sussman et al., 1991; Noble et al., 1993; Nardini and Dijkstra, 1999). The prototype of this fold consists of an eight-stranded mostly parallel β-sheet connected by one or more helices between adjacent strands (Fig. 17.1.33A and B). The strands are in the order 1-2-4-3-56-7-8 with strand 2 antiparallel to the rest.
While the central eight-stranded β-sheet is well conserved among these enzymes, the length and number of helices spaced between the adjacent strands are less conserved and their positions vary significantly from enzyme to enzyme. The length and location of the loops connecting the secondary structure elements are also highly variable among members of this family.
Sialidases Sialidases and influenza neuraminidases form a family of glycohydrolases that cleave terminal α-ketosidically-linked sialic acids of many glycoproteins, glycolipids, and oligosaccharides. Structurally, they define a prototype with a β-propeller fold composed of six fourstranded antiparallel β-sheets arranged similar to a six-blade propeller (Fig. 17.1.33C; Varghese and Colman, 1991; Burmeister et al., 1992; Crennell et al., 1993; White et al., 1995). There is a six-fold pseudo-symmetry through the center of the subunit relating each of the six blades. The topological connections between the strands within each sheet and among the adjacent sheets are identical from blade to blade. Each β-sheet displays a typical righthanded twist with strands 1 to 4 running antiparallel from the center of the propeller to the edge. The connection between each adjacent blade is always through the fourth strand of the preceding sheet to the first strand of the connecting sheet. The active site is located near the center of the propeller (see Frequently Observed Secondary Structure Assemblies of Structural Motifs for further discussion).
The TIM-Barrel Fold The TIM-barrel fold, also referred to as an α/β-barrel or (β/α)8 fold, is one of the most frequently observed catalytic folds in protein structures. This fold is used by a multitude of different enzymes with very diverse functions and received its name from triose phosphate isomerase (TIM), the protein in which it was first discovered (Banner et al., 1975). The threedimensional structure of the TIM-barrel is a rather elegant looking eight-stranded parallel β-barrel surrounded by a ring of helices that are essentially antiparallel to the β-strands (Fig. 17.1.34A and B). Both the β-strands and the α-helices are inclined to the axis of the barrel. The classical topology of this structure is an array of eight tandem up-down β-α motifs which are aligned side-by-side in a parallel manner with consecutive strand order, eventually wrapping around to form a β-barrel (Fig.
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17.1.34C). In some TIM-barrel structures, this array is not completely regular, containing instead small insertions in loop regions or, as in the cases of enolase and muconate lactonizing enzyme, missing a helix between the last two β-strands. To date all of the crystal structures in which the TIM-barrel has been found have been enzymes and in almost every case the active site is located within the loop regions at the C-terminal end of the barrel.
Protease Folds The various proteases that have been examined structurally can be grouped into four primary classes based on characteristics of the active sites. First, serine proteases use an essential Ser as the active site nucleophile. Second, cysteine proteases require an active site Cys nucleophile. Third, aspartic proteases work primarily at low pH, utilizing two essential Asp side chains for catalysis. Finally, metalloproteases all contain a critical zinc atom coordinated at the active site. These four classes can be further subdivided into two or more different protein folds despite the fact that members of a particular class share nearly the same reaction mechanism. A fifth class that has been observed in proteasome complexes and which uses a Thr as the active site residue is also discussed in this section.
Overview of Protein Structural and Functional Folds
Serine proteases Nearly all serine proteases contain the well known catalytic triad Ser-His-Asp in their active sites. The three side chains are positioned close together through tertiary structure and are hydrogen bonded to each other in a manner which causes the Ser hydroxyl oxygen to be a strong catalytic nucleophile. An exception is the cytomegalovirus protease which uses a similar Ser-His-His catalytic triad for the same purpose. Four completely different protein folds have been observed to date in this class of proteases. Each fold is described below. Trypsin-like serine proteases. Trypsin-like proteases are folded into two nearly identical domains, each consisting of a six-stranded antiparallel β-barrel with a Greek key topology (Fig. 17.1.34D; Fischer et al., 1994). In many cases, an α-helix is associated with each domain. The active site is formed from loop regions in a cleft between the two domains. Numerous eukaryotic, prokaryotic, and viral serine proteases have been observed to utilize this fold. Some of the more well known trypsin-like proteases include thrombin, elastase, chymotrypsin, plasmin(ogen), urokinase, and
complement C1R and C1S proteases. At least three types of viral cysteine proteases also have the trypsin-like fold (see below). Subtilisns. The three-dimensional structures of subtilisns from prokaryotic and fungal organisms (Fig. 17.1.34E) show a common core folding pattern made up of a seven-stranded parallel β-sheet (strand order 2-3-1-4-5-6-7) sandwiched between two layers of α-helices (Murzin et al., 1995). The active site is located at the C-terminal ends of the central β-sheet strands with the essential Ser residue situated at the N-terminus of an α-helix. Additions to this core fold in some subtilisns include a small two- to four-stranded antiparallel sheet positioned at the C-terminal edge of the large βsheet and a C-terminal β-ribbon hydrogen bonded to the main β-sheet (extending it by two strands). Serine carboxypeptidases. Ser ine carboxypeptidases are members of a larger family of enzymes known as the α/β-hydrolases (see The α/β Hydrolase Fold). These serine proteases contain a central, eleven-stranded, mixed, mostly parallel β-sheet (three more strands than the canonical α/β-hydrolase fold) surrounded by fifteen helices on either side of the sheet (Fig. 17.1.34F). The β-sheet has a pronounced twist of almost 180°. As in the subtilisn class of proteases (see above), the active site is located at the C-terminal end of the β-sheet with the catalytic serine at the Nterminal end of a helix (Liao and Remington, 1990). Herpes virus–like serine proteases. First observed in the structure of a serine protease from human cytomegalovirus (hCMV), the three-dimensional structures of these single-domain proteases consist of a seven-stranded mostly antiparallel β-barrel with one Greek key motif and seven helices surrounding approximately one half of the circumference of the barrel as well as both ends of the barrel (Qiu et al., 1996; Shieh et al., 1996; Tong et al., 1996). This fold has subsequently been observed in Herpes simplex virus type II serine protease (Hoog et al., 1997), Karposi’s sarcoma-associated herpes virus protease (Reiling et al., 2000), and Varicella-Zoster viral protease (Qiu et al., 1997). The active site Ser-His-His triad is positioned on the outside of the barrel on the face without any helices. This group of proteases is unique among serine proteases in that the tertiary structure of its catalytic triad contains a conserved His residue in the same position that the Asp residue is normally found in the classical Ser-His-Asp triad. These proteins func-
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tion physiologically as dimers with the active sites situated distally to each other. Cysteine proteases To date, three different subfamilies of cysteine proteases have been characterized, each of which utilize the sulfhydryl of a Cys residue as the active site nucleophile. The most well known and studied are the papain-like cysteine proteases which are sometimes confused with the entire cysteine protease family. Papain-like cysteine proteases. The proteases of this subfamily can be recognized by the conserved catalytic triad Cys-His-Asn, in addition to a unique fold. Analogous to the Ser-His-Asp triad of serine proteases, this hydrogen bonded triad is also formed through tertiary interactions resulting in a nucleophilic sulfhydryl on the Cys. The basic papain-like fold consists of two distinct domains designated L and R (papain nomenclature; Fig. 17.1.35A). The L domain is a small three- to four-helix bundle. The R domain is comprised of a highly curved and twisted six-stranded antiparallel β-sheet flanked by one or two helices (Musil et al., 1991). The active site is in a V-shaped crevice between the two domains. Similar to the subtilisns and serine carboxypeptidases, the nucleophilic cysteine residue is located at the N-terminal end of an α-helix. Although small structural variations of the described fold are observed in several papainlike proteases, bleomycin hydrolase from yeast has extensive structural modifications (JoshuaTor et al., 1995): a small helical domain is inserted into loops in the L domain and an additional N-terminal helical domain is also present. This cysteine protease, which cleaves the anticancer peptide bleomycin, functions as a hexamer with a large central channel containing the six active sites. It also binds to doubleand single-stranded DNA, and represses genes of the GAL system. Caspases. Cysteine-dependent aspartatespecific proteases (caspases) are a family of structurally homologous cysteine endopeptidases that are involved in processing proinflammatory cytokines and in apoptotic signaling (Alnemri et al., 1996). The prototypic caspase structure, that of interleukin-1β-converting enzyme (ICE or caspase-1) is a tetramer made from two heterodimers related by a two-fold rotation axis. Each heterodimer (p10/p20) contains an independent active site and is derived from a 45-kDa precursor that has been cleaved in three locations. The fold of the heterodimer consists of a six-stranded mostly parallel β-
sheet sandwiched between two helices on one face and four helices on the other (Fig. 17.1.35B; Walker et al., 1994; Wilson et al., 1994). The active site formed by loops at the C-terminal end of the β-sheet contains a catalytic Cys-His diad. The active site Cys is situated at the end of a β-strand instead of the N-terminus of an α-helix as in the papain subfamily. The crystal structures of several other caspases (i.e., caspase-3, -7, -8, and -9) have confirmed this unique fold and quaternary structure except for differences in substrate binding sites (see Wei et al., 2000; Chai et al., 2001; Renatus et al., 2001, and references therein). Although caspases are mammalian enzymes, the crystal structure of the catalytic domain of a bacterial cysteine protease, gingipain R from Porphyromonas gingivalis, has been observed to have a very similar fold and a Cys-His diad at the active site (Eichinger et al., 1999). The two subdomains within the single chain catalytic domain correspond to the small and large subunits of the caspase fold. Viral trypsin-like cysteine proteases. Another set of cysteine proteases that have been characterized structurally include the 3C and 2A viral cysteine proteases (Allaire et al., 1994; Matthews et al., 1994; Petersen et al., 1999). They use a Cys-His-Glu/Asp catalytic triad and surprisingly have the same fold as the trypsinlike serine proteases (see above). The coronavirus cysteine protease also shares this fold but has an extra helical domain and a Val instead of Asp or Glu in the third position of the canonical catalytic triad (Anand et al., 2002). Aspartic proteases The aspartic proteases (also known as acid proteases) contain primarily β-sheets and utilize two Asp residues to catalyze proteolysis at low pH. They are divided into two groups, the two-domain pepsin-like proteases found in eukaryotes and the dimeric retroviral proteases. Both groups contain a conserved catalytic triad sequence Asp-Thr/Ser-Gly at the active site and limited regions of sequence conservation are shared between them. Pepsin-like proteases. The pepsin-like proteases are comprised of two pseudosymmetric domains, each containing a partially opened, six-stranded, antiparallel, distorted β-barrel structure with a few short helices within the loop regions (Fig. 17.1.35C; Davies, 1990). Between the domains and away from the center of the structure, a twisted six-stranded antiparallel β-sheet lays against the surface of the protein as if to form a base on which to set the
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molecule. Three strands from each domain contribute to the sheet. On the side of the protein opposite the six-stranded sheet, a large cleft runs across the domain interface forming the site for the polypeptide substrate to bind. The two catalytic Asp residues (one from each domain) are located at the bottom of the active site cleft (Fig. 17.1.35C). A long hairpin loop from the N-terminal domain forms a flap which projects over the top of the active site shielding it from solvent. When pepsin-like proteases bind an inhibitor, the flap also participates in binding by clamping down over the inhibitor. Retroviral proteases. Retroviral proteases are homodimers that cleave newly synthesized viral polyproteins into individual functional units during viral maturation. The two identical subunits are structurally very analogous to the two domains of the pepsin-like proteases. Each is comprised of a six-stranded antiparallel βbarrel, a single helix located at one end of the barrel, and a two-stranded β-sheet at the C-terminal end of the helix (Fig. 17.1.35D; Navia et al., 1989; Davies, 1990). In the dimer, the two small β-sheets hydrogen bond to form a fourstranded β-sheet analogous to the six-stranded sheet observed in pepsin-like proteases. Two conserved, catalytic Asp residues (one from each monomer) are positioned at the bottom of the active site cleft at the subunits interface. A flap from each subunit extends over the top of the active site in a manner structurally and functionally resembling the flap of the pepsinlike enzymes.
Overview of Protein Structural and Functional Folds
Metalloproteases There are two main superfamilies of zinccontaining metalloproteases, each of which extend across both prokaryotes and eukaryotes: the zinc-dependent endopeptidases, also referred to as zincins (Bode et al., 1993), and the zinc-dependent exopeptidases. Zincins. The conserved tertiary structure of the catalytic domain of zincins consists of a five-stranded mostly parallel β-sheet with two helices packed against the concave side Fig.17.1.35E; Bode et al., 1993; Gomis-Ruth et al., 1994). This fold is generally observed in the N-terminal domains of zincins. The conserved His-Glu-X2-His sequence motif identifies three critical active site residues. These three residues are in a helix situated at the edge of the β-sheet with the two histidines coordinating the catalytic zinc and Glu, polarizing a zinc-bound water that acts as a nucleophile during catalysis. Several zincins also bind multiple Ca2+ ions.
Zincins comprise two distinct families known as the thermolysin-like proteases and the metzincins. The known tertiary structures of thermolysin-like proteases have a small irregular twisted β-sheet structure at one end of the central β-sheet and a C-terminal helical domain with the active site in a crevice between the two domains. A conserved glutamate residue provides the fourth zinc ligand. The metzincins, which include astacins, adamalysins, matrix metalloproteases, and serralysins have an extended consensus sequence (His-Glu-X2His-X2-Gly-X2-His), including a third zinc-coordinated histidine. The name metzincins comes in part from a unique structural characteristic of this family referred to as a Met-turn, which is a methionine-containing β-turn located just beneath the zinc atom (Bode et al., 1993). It is noteworthy that one metzincin, an alkaline protease of the serralysins, has a particularly unusual C-terminal domain consisting of a calcium-binding 21-strand β-sandwich, also referred to as a right-handed parallel β-roll (Fig. 17.1.35G; Baumann et al., 1993). Zinc-dependent exopeptidases. Zinc-dependent exopeptidases share a common catalytic fold consisting of an α/β/α sandwich. The central eight-stranded β-sheet (strand order 12-4-3-8-5-7-6) is mostly parallel (only strands 2 and 8 are antiparallel) and hosts the active site at the middle of its C-terminal end (Fig. 17.1.35F; Burley et al., 1992). Carboxypeptidases A, B, and T each have one active site zinc atom pentacoordinated to two His, a Glu (both oxygens are ligands), and a water molecule. The aminopeptidases in this family, however, each have two zinc atoms coordinated less than 3.5 Å apart at the active site. As with endopeptidases, zinc-bound water in this exopeptidase family is considered a possible nucleophilic catalyst (Strater et al., 1995). Ntn hydrolase fold The N-terminal nucleophile (Ntn) hydrolase fold is used by eukaryotic and archael proteaso mes o r pr oteaso me h om ologues in prokaryotes (e.g., Hs1V; Table 17.1.14; Bochtler et al., 1999). This fold has an αββα core structure consisting of a ten-stranded antiparallel β-sandwich flanked on each face by helices (Fig. 17.1.35H; Brannigan et al., 1995). The strand order for one sheet is 3-4-5-6-7 and the other is 8-1-2-9-10. A catalytic triad of Thr, Glu/Asp, and Lys forms at least part of the active site at one end of the β-sandwich and is highlighted by an essential N-terminal threonine nucleophile (Groll et al., 1997). Variations
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of this fold (α- and β-subunits) are used to make large cylinder-shaped multisubunit complexes known as proteasomes or proteasome homologues that are responsible for protein degradation (see Proteasome for further discussion). Only certain β-subunits are catalytic and none of the α-subunits are. This same fold is also found in several other enzymes, such as glutamine PRPP aminotransferase, glucosamine 6-phosphate synthase, and various penicillin acylases, to name a few (Oinonen and Rouvinen, 2000). Sequence homology is generally not conserved between these enzymes, nor is the identity of the catalytic nucleophile.
Enzymes Involved in Ubiquitination Ubiquitin-targeted protein degradation plays an important role in protein degradation by proteasomes, in DNA repairs, and in selected signal transduction pathways (Hershko and Ciechanover, 1998). A ubiquitin pathway involves three distinct steps. (1) E1-catalyzed ubiquitin activation which attaches the C-terminal Gly residue of ubiquitin to a Cys residue on E1 through a thioester linkage. (2) E2s-mediated ubiquitin conjugation that transfers the activated ubiquitin from E1 to the active site Cys residue of E2s. (3) Ligation of ubiquitin to substrate proteins by E3 ubiquitin ligases. Ubiquitin, a conserved protein of 76 amino acids, and all ubiquitin-like proteins fold into small αα/β structure with a five-stranded antiparallel β-sheet wrapped around an α-helix (sometimes referred to as a β-grasp; Fig. 17.1.36A; Vijay-Kumar et al., 1987; Rao-Naik et al., 1998). While there is usually one E1, there are multiple and diverse E2 and E3 enzymes, each specific for different protein substrates. Ubiquitin-conjugating enzymes The structures of ubiquitin-conjugating E2 enzymes solved to date include UbcH7, Ubc9, and a Mms2-Ubc13 heterodimer complex (Bernier-Villamor et al., 2002; Moraes et al., 2001; VanDemark et al., 2001; Zheng et al., 2000). The catalytic core of E2 enzymes consists of ∼160 amino acids that fold into an α/β structure with its central four- to seven-stranded antiparallel β-sheet flanked by four α-helices (Fig. 17.1.36B and C). The active site cysteine, which catalyzes the transfer of ubiquitin from E1 through a thioester linkage to E2, is located in a loop between the fourth β-strand and the second α-helix.
Ubiquitin ligases The ubiquitin ligases (E3s) are a very diverse family of proteins whose function is to bring protein substrates into proximity with ubiquitin-conjugated E2 enzymes. The structural diversity of ubiquitin-ligases presumably reflects the need for specificity among diverse protein substrates. Common among E3s is their ability to bind E2 and specific protein substrates. There are two types of E3 enzymes, which differ in their ubiquitin transfer mechanisms. The HECT-type (homology to E6AP binding protein C terminus) ubiquitin ligases catalyze the transfer of ubiquitin by first forming an E3-ubiquitin thioester intermediate. The E6AP-Ubc7 complex is an example of an HECT-type E3 enzyme. In contrast, the RINGfamily ubiquitin ligases do not form an E3ubiquitin intermediate. They all have a RING domain that interacts with the E2-ubiquitin conjugate. Examples of RING-type ubiquitin ligases include c-Cbl (Fig. 17.1.36B; Zheng et al., 2000), members of the seven in absentia homolog (Siah) family (Fig. 17.1.36D; Polekhina et al., 2002), and the SCF (Skp1-CullinF-box) family of E3 ligases (Fig. 17.1.36E; Zheng et al., 2002). The SCF members constitute the largest and most complex family of protein ligases and ubiquitinate a rather diverse range of proteins. The SCF complex consists of a protein-substrate-interacting subunit, Skp2, (Schulman et al., 2000), an elongated two-domain cullin subunit (Cul1), a connecting unit, Skp1, that links the Skp2 F-box–containing protein with cullin, and a RING domain nested on cullin (Fig. 17.1.36E). Human Skp2 has an ∼100 residue N-terminal domain of unknown structure, followed by an F-box, ten leucine-rich repeats (LRR; see Leucine-Rich Repeats for a discussion of the LRR fold), and a 30-residue tail with little secondary structure (Schulman et al., 2000). The F-box is an ∼40residue domain comprising three helices, with one helix packing orthogonally against a pair of antiparallel helices. The N-terminal domain of Skp1 has an α/β fold consisting of a threestranded β-sheet packed on one face against a bundle of α-helices (also known as a BTB/POZ fold) and an additional two-helix extension (Schulman et al., 2000). The N-terminal domain of Cul1 consists of three cullin repeats, each comprising a bundle of five α-helices (Fig. 17.1.36F). The C-terminal domain of Cul1 is made of two winged helix motifs (see HelixTurn-Helix Motifs), a four-helix bundle, and an α/β subdomain. The zinc-containing Rbx1 RING domain interacts closely with the C-ter-
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minal domain of Cul1 and contributes one βstrand to the five-stranded β-sheet of the α/β subdomain. Small ubiquitin-like modifiers In addition to the more well known E2-E3 system, there is a family of small ubiquitin-like modifiers (SUMO), that regulate nuclear transport, stress response, and signal transduction controlling cell-cycle progression. The E2 enzymes for SUMO, as shown in the structure of a Ubc9-RanGAP1 complex (Bernier-Villamor et al., 2002), can directly recognize and modify Lys residues on their protein targets without additional E3s.
ELECTRON TRANSFER PROTEINS There are several superfamilies of proteins involved in processes of electron transfer. Most of them have a heme prosthetic group or porphyrin ring central to electron transfer coordinated in a hydrophobic pocket; however, their tertiary folds vary from the four-helix-bundle fold of ferritin to the mostly helical (multiple α-helices and some β-strands) structure of cytochrome P450, to the typical cytochrome c fold, with the polypeptide chain organized into a series of α-helices and reverse turns. Below are summaries of a few commonly observed folds in this category (Table 17.1.15). Another group of electron transfer proteins, integral membrane respiratory proteins, is discussed below (see Respiratory Enzymes).
Cytochrome P450
Overview of Protein Structural and Functional Folds
Cytochrome P450 catalyzes the mono-oxygenation of a number of organic molecules in a heme-dependent reaction mechanism. There are now at least eight different types of P450 that have been examined structurally. These include P450cam, P450eryF, P450BM-3 , P450terp, P450NOR, P4502C5, P450 sterol 14α-demethylase, and CYP119, each utilizing very different substrates. The sequence homology remains relatively high within each group but relatively weak between them. Despite low sequence homologies (<20%) and very diverse substrate specificities, a common fold is maintained among all the P450 cytochromes. The molecule folds into two domains, α and β (Fig. 17.1.37A). The α domain consists of a righthanded four-helix bundle sandwiched between two clusters of helices, a top cluster of five helices, and a bottom cluster of three helices with two β-strands. The β domain consists of a β-barrel surrounded by three helices. The βbarrel is formed by two β-sheets, a two-
stranded β-sheet, and a five-stranded β-sheet, folded against each other (Raag et al., 1993). The heme molecule is embedded between the two domains.
Cytochrome c Domain Cytochrome c is often observed as an electron transfer domain in many redox enzymes. Members of this family include cytochromes c, c2, c5, c551, c553, and c555. Other proteins, such as the N-terminal domain of cytochrome cd1-nitrite reductase, flavocytochrome c sulfide dehydrogenase (FCSD), and di-haem cytochrome c peroxidase, also contain cytochrome c–type domains in their structures. Most cytochrome c proteins contain one cytochrome c domain, but some enzymes, including FCSD and di-haem cytochrome c peroxidase, contain two. The typical cytochrome c fold consists of three long α-helices and a pair of β-strands packed in the core region, with two additional short helices surrounding the core (Fig. 17.1.37B). The length of the helices, strands, and loop conformations vary considerably among the members of the family. The heme group is coordinated by residues from both helices and reverse turns in the core region (Louie and Brayer, 1990).
Cytochrome c3 The structure of cytochrome c3 from Desulfovibrio desulfuricans consists of three to four helices (often one long helix amidst other shorter helices), a pair of β-hairpins, and three to four long irregular loops (Fig. 17.1.37C). There are four heme molecules ligated to the ∼110 residues of cytochrome c3. The heme coordination frame is formed by packing the long helix against the β-hairpins and the loop regions.
Cytochrome b5 In addition to cytochrome b5, the N-terminal domain of flavocytochrome b2 also belongs to this fold family. The structure folds into a α/β/α sandwich with a central five-stranded mixed β-sheet surrounded by six short α helices, two on the concave side of the β-sheet and four on the other (Fig. 17.1.37D; Mathews et al., 1972). The heme is coordinated by the four helices and the central two strands of the βsheet. The five strands are hydrogen-bonded in a 1-5-3-2-4 order in cytochrome b5 and 5-3-12-4 order in the cytochrome b5–type domain of flavocytochrome b2.
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Four-Helix Bundle Cytochromes There are at least three families of cytochromes known to assume the fold of four-helix bundles: cytochrome b562, cytochrome c′, and ferritin. Among them, cytochrome b562 and cytochrome c′ share a close resemblance, both forming up-down-up-down helical bundles with a left-handed twist (Fig. 17.1.37E). The heme is inserted between helices 1 and 4, with helix 3 forming the floor. The structure of ferritin, on the other hand, forms an up-downdown-up four-helix bundle with left-handed twist. There is a short one-turn helix at the C-terminal of the four-helix bundle. Unlike cytochrome c′ or b562, ferritin forms a dimer with only one heme molecule inserted at the interface of the two helical bundle subunits (Fig. 17.1.37F; Frolow et al., 1994).
GLOBIN-LIKE PROTEINS The helical globin-like proteins encompass three different families: globins, phycocyanins, and colicin. The core fold for all of these proteins consists of an α-helical sandwich which is simply two layers of α-helices stacked against each other. Using the globin nomenclature, helices A, E, and F make up one side of the sandwich, while helices B, G, and H make up the other, with helices on each side essentially orthogonal to those on the other (Table 17.1.16, Fig. 17.1.38A). Various members of these families are all completely α-helical (with the exception of flavohemoglobin), with minor differences in helix packing due to the binding of associated cofactors and insertions of helices to the basic fold (Holm and Sander, 1993; Ermler et al., 1995).
bound in a cooperative manner, making it well suited for oxygen transport. Other nonmammalian hemoglobins are found to have very different oligomeric states. Pig roundworm hemoglobin, for example, is an octamer of a two-domain globin fold (Kloek et al., 1993), while sea cucumber hemoglobin can be present in both monomeric and dimeric states under physiological conditions. Flavohemoglobin is a rather unusual globin. Unlike the other known globin structures, flavohemoglobin is an α/β three-domain protein containing an FAD-binding domain and an NAD-binding domain as well as an N-terminal globin domain (Ermler et al., 1995). The middle FAD-binding domain is a six-stranded antiparallel β-barrel with Greek key topology and the C-terminal NADbinding domain is a classical dinucleotide binding domain with a five-stranded parallel βsheet. Both redox domains share significant structural homology with both domains of ferridoxin reductase (Karplus et al., 1991). The physiological function of flavohemoglobin remains unknown.
Phycocyanins Phycocyanins make up part of the light-harvesting antennae in cyanobacteria and red algae. The three-dimensional structure of this protein has the six helices of the globin fold with two additional antiparallel α-helices protruding from the N-terminus (Fig. 17.1.38C; Schirmer et al., 1987). A phycocyanobilin cofactor is located in the same place as the heme group of the globin structures (between helices E and F).
Colicins Globins The globin family is made up of several heme-containing helical proteins used specifically for oxygen storage and transport within bacteria, insects, vertebrates, and plants. Myoglobin is the smallest of these proteins, consisting of the basic globin fold with the addition of two more short helices (C and D; Fig. 17.1.38A; Phillips, 1980). This monomeric protein binds oxygen for storage in muscle using a heme group located in a cleft between helices E and F. Mammalian hemoglobin is composed of four separate globin chains (α2β2) in an approximately tetrahedral arrangement (Fig. 17.1.38B; Fermi et al., 1984). Each chain binds a heme group in the same location and orientation as in myoglobin. The binding of oxygen induces changes in the quaternary structure of hemoglobin such that four oxygen molecules are
Colicins are a class of multidomain antibacterial toxins produced by E. coli that participate in killing target cells by binding to surface receptors and forming pores in the cytoplasmic membrane. The crystal structure of a C-terminal fragment of colicin A is observed to have a canonical globin fold with three additional Nterminal helices and an extra C-terminal helix (Fig. 17.1.38D; Parker et al., 1989).
TOXINS The structures of toxins represent a collection of a diverse family of folds. Some toxins, such as scorpion and snake toxins, are relatively small, contain few well defined secondary structure elements, and are rich in disulfides, while others, such as diphtheria toxin and heatlabile toxin, have well defined secondary structures and often form complicated mutidomain
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oligomeric organizations. Numerous toxin structures have been determined by X-ray crystallography or NMR, and most belong to one of several distinct structural folds summarized in this section (Table 17.1.17).
Ribosome-Inactivating Toxins The A chains of ricin and abrin as well as the single-chain toxins gelonin, α-momorcharin, α-trichosanthin, and shiga toxin are members of the ribosome-inactivating protein (RIP) fold family, named for their ability to block protein synthesis. Figure 17.1.39D illustrates the structure of the ricin A chain (Katzin et al., 1991). It contains mixed β-sheets and several α-helices. The B chain of ricin and abrin each form a β-trefoil type structure (see β-Trefoils and Table 17.1.9). There are two domains of such trefoils in the B chain of each toxin (Rutenber and Robertus, 1991).
ADP Ribosylation Toxins Diphtheria toxin (DT), Pseudomonas aeruginosa exotoxin A (ETA), cholera toxin, heat-labile enterotoxin, and pertussis toxin all contain ADP ribosylation domains which enable these toxins to inhibit protein synthesis by catalyzing the ADP-ribosylation (and thereby the inactivation) of essential cellular proteins. The basic fold of an ADP ribosylation domain, shown in Fig. 17.1.39A, consists of a mixed structure of seven or eight β-strands and several short helices. The β-strands are grouped into two β-sheets that are packed against each other forming the core of this domain.
Overview of Protein Structural and Functional Folds
a pentameric receptor-binding B domain (Sixma et al., 1991; Swaminathan et al., 1992; Stein et al., 1994). The topology of the B domain, shown in Fig. 17.1.39B, contains a pentameric ring structure of layered helices-sheetshelices. The pentameric repeat unit is an α/β/α sandwich that consists of a one-turn α-helix, six-stranded antiparallel β-sheet and long α-helix. Superantigen toxins The three-dimensional structures of staphylococcal enterotoxins A (SEA), B (SEB), C2 (SEC2), and a toxic shock syndrome toxin 1 (TSST-1) define a family of toxins that function as superantigens to T cells (Swaminathan et al., 1992, 1995; Prasad et al., 1993). The common fold of these toxins is a two-domain structure (Fig. 17.1.39C). The N-terminal domain folds with a Greek key topology and consists primarily of a five-stranded β barrel with strands in the order 1-2-3-5-4. It is also a member of the oligonucleotide/oligosaccaride binding (OB) fold (see The OB Fold; Murzin, 1993). The C-terminal domain of these toxins folds into a modified β-grasp motif. The primary feature of this domain is a long α-helix packed against a five-stranded mixed β-sheet.
Small Single-Subunit Toxins
Diphtheria toxin Diphtheria toxin (DT) and Pseudomonas exotoxin A (ETA) structures are each organized into three domains: catalytic ADP ribosylation, transmembrane, and receptor-binding (Allured et al., 1986; Choe et al., 1992). The sequential order of the three domains is from N- to C-terminal in DT and from C- to N- terminal in ETA. The transmembrane domain in both DT and ETA is all α-helical. The receptor-binding domains are both immunoglobulin-like; however, the receptor-binding domain of DT resembles an immunoglobulin V domain (see above) whereas that of ETA resembles a Con A lectin domain (see above).
Scorpion toxins Members of scorpion toxins are usually small proteins of ∼50 amino acids strengthened by disulfide bonds. They include scorpion margatoxin, noxiustoxin, charybdotoxin, scyllatoxin, agitoxin, chlorootoxin, toxin I5a, kaliotoxin, and others. The basic topology consists of a N-terminal α-helix packed against a threestranded antiparallel β-sheet (Fig. 17.1.39E; Johnson et al., 1994). The fold is stabilized by three sets of disulfide bonds connected in the sequential order of Cys I–IV, II–V, and III–VI. Cys II and III usually follow the CxxxC pattern whereas Cys V and VI usually follow the CxC pattern. Some members of the family, referred to as long-chain scorpion toxins, have an additional loop structure at the C-terminal end and an additional disulfide bond (Housset et al., 1994). The rest of the family is known as the short-chain scorpion toxins.
Bacterial AB5 toxins Cholera toxin, pertussis toxin, and heatlabile enterotoxin belong to a family of bacterial AB5 toxins. All of them contain an ADP ribosylation domain (domain A; see above) and
Snake toxins Most snake toxins have similar structures, known as the snake-toxin fold. Members with known structures include sea snake erabutoxin A and B, neurotoxin I and II, cobratoxin I and
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II, cardiotoxin I, II, IIB, III and V, and bungarotoxin. Their structure has two antiparallel βsheets, an N-terminal two-stranded β-sheet, and a C-terminal three-stranded β-sheet (Fig. 17.1.39F; Tsernoglou et al., 1978). There are four disulfides with the sequential bonding of Cys I–III, II–IV, V–VI, and VII–VIII in each structure. Spider toxins The family of spider toxins include ω-conotoxins from sea snail and agatoxins from web spider. These are very small proteins, <40 residues in sequence, and are stabilized by Cys bridges. The structure consists of largely irregular loops with only two short β-strands forming an antiparallel β-sheet at the center of the fold (Fig. 17.1.39G; Reily et al., 1995). The three common disulfides are connected with the sequence order of Cys I–IV, II–V, and III–VI. Members of agatoxins have one more disulfide near the C-terminal end.
Anthrax Toxin Anthrax toxin is the major virulent factor of Bacillus anthracis. Owing to its high mortality when the infection is untreated, and its ability to exist in an aerosol form, it poses a threat to public health as a potential reagent for biological warfare and terrorism. The toxin consists of three proteins, lethal factor (LF),edema factor (EF), and protective antigen (PA). Among them, the lethal factor appears to be a protease that targets the mitogen-activated protein kinase kinase (MAPKK) family of proteins.edema factor is an adenylate cyclase and PA forms a membrane pore-like transporter once activated by furin-like cellular proteases. The crystal structures of all three factors have been solved (Table 17.1.17; Petosa et al., 1997; Pannifer et al., 2001; Drum et al., 2002; Shen et al., 2002). Lethal factor Lethal factor (LF), with a molecular weight of 90 kDa, consists of four domains. Domains I and IV are structurally similar, each with a set of nine to twelve helices packed against a fourstranded β-sheet (Fig. 17.1.40A). The function of domain I appears to be to bind protective antigen (PA; see below) whereas domain IV appears to be the catalytic domain of LF. The α/β part of domain IV loosely resembles a thermolysin-like zincin (see Protease Folds) with a Zn2+ coordinated within a His-Glu-X2His motif. Domain I has no zinc-binding site. Domain II has an α/β fold similar to ADP
ribosylation toxins (see ADP Ribosylation Toxins), however, a critical Glu is not conserved, suggesting this domain does not have ADP-ribosylating activity (Pannifer et al., 2001). Domain III is a small α-helical bundle. The N-terminal peptide of MAPKK is bound at the interface between domains III and IV. Oedema factor Oedema factor (EF) is a calmodulin (CaM)dependent adenylate cyclase, whose activity increases 1000-fold upon activation by CaM. The structures of both the catalytic portion of EF alone and EF in complex with CaM have been determined (Drum et al., 2002; Shen et al., 2002). The fold of EF consists of a largely α-helical domain, three switch regions, and two α/β catalytic domains, CA and CB (Fig. 17.1.40B). Domains CA and CB have folds similar to the palm domains of DNA polymerase β and RNA poly(A) polymerase (see DNA Polymerases and Small RNA Polymerases). The binding of CaM induces large conformational changes in the association of the helical domain with respect to the switch and catalytic domains (Fig. 17.1.40C). Protective antigen Protective antigen (PA), an 83-kDa protein, consists of four domains, I to IV (Fig. 17.1.40D). Domain I contains a jelly-roll-like β-sandwich and two calcium ions coordinated by residues in an EF-hand motif. Domain I has the furin cleavage site and, once cleaved, PA forms a heptameric membrane pore transporter. Domain II folds into a modified Greek-key-like β-barrel. Domain III adopts the same topology as ferredoxins and the domain A of the toxicshock-syndrome toxin-1 (TSST-1). Domain IV contains an immunoglobulin-like fold (see above) and is presumably responsible for host cell binding of PA.
LIPID-BINDING PROTEINS There are numerous different protein folds involved in binding to lipids or lipid-like molecules, most with β-sheet structures. Three well known families of lipid-binding proteins are discussed in this section: lipocalins, fatty acidbinding proteins, and serum albumins. See Modular Domains Involved in Signal Transduction for a discussion of other phospholipid-binding proteins.
Lipocalins Lipocalins (or retinol-binding protein-like proteins) comprise a family of mostly extracel-
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lular proteins 160 to 180 residues in length, typically involved in the transport of small hydrophobic molecules (Flower et al., 2000). Examples include retinol-binding protein, βlactoglobulin, bilin-binding protein, and ordorant-binding protein (Table 17.1.18). The crystal structure of retinol binding protein (Cowan et al., 1990) and subsequent structures reveals an eight-stranded antiparallel β-barrel with a β-meander topology (Fig. 17.1.41A; also see Frequently Observed Secondary Strucutre Assemblies or Structural Motifs). A short helix is sometimes located before the first β-strand and after the last β-strand of the β-barrel. Ligands bind inside the barrel.
Fatty Acid–Binding Proteins Fatty acid binding proteins (FABPs) constitute a family of small, predominantly cytoplasmic lipid-binding proteins. They include myelin P2 protein, adipocytes lipid-binding protein, cellular retinol-binding protein (CRBP), and numerous tissue-specific forms of FABP. The fold comprises a ten-stranded antiparallel βsheet with a pair of α-helices between the first and second β-strands. Like lipocalins, the topology is that of a β-meander. The β-sheet folds in half to form what is often referred to as a β-clam (Fig 17.1.41B). In the resulting structure the β-strands in each half of the clam shell are nearly orthogonal to each other and a ligand-binding cavity is created between them.
Serum Albumin
Overview of Protein Structural and Functional Folds
Serum albumin is a remarkably abundant fatty acid transport protein in the circulatory system. In humans, concentrations of 40 to 50 mg/ml are maintained in the blood. Despite extensive characterization, the atomic three-dimensional structure of serum albumin remained elusive until the early 1990s. Crystal structures of human and horse forms reveal a mostly helical heart-shaped protein composed of three structurally homologous domains (I to III; Fig. 17.1.41C; He and Carter, 1992; Ho et al., 1993; Curry et al., 1998; Sugio et al., 1999). Each domain has ten helices and is divided into two subdomains (a and b) with six and four helices, respectively. Furthermore, the first four helices in subdomain a are structurally homologous to the four helices in subdomain b. It is noteworthy that serum albumin contains 35 cysteine residues resulting in 17 disulfide bridges and one free cysteine. Despite the striking symmetry within the structure of serum albumin, its five fatty acid binding sites are located asymmetrically around the molecule
(Curry et al., 1998): domain I has two binding sites, domain II has one binding site (shared with domain I), and β domain III has three binding sites (Fig. 17.1.41C).
LARGE MULTISUBUNIT PROTEINS Chaperonin Proteins: GroES and GroEL Chaperonins are a class of proteins that assist in catalyzing the folding of other proteins. The GroEL/GroES complex represents one of the best characterized chaperonin systems and both components are required for proper protein folding in bacteria (Xu et al., 1997b). Crystal structures of GroEL, GroES, an asymmetric GroEL/GroES complex, and other homologous chaperonin complexes have been determined by several groups (Table 17.1.19). GroEL (L for large) forms the cylindrical core of the GroEL/GroES complex and consists of fourteen identical subunits arranged as two heptameric rings stacked back to back (Fig 17.1.42A). Each individual protomer is comprised of three distinct domains: equatorial, intermediate, and apical (Fig 17.1.42B and C). The equatorial domain consists primarily of six long α-helices, two short helices, and three pairs of short β-strands. This is also the ATPase domain of GroEL. The intermediate domain contains a three-stranded β-sheet and three αhelices. The apical domain consists of a mixture of five α-helices and a seven-stranded β-sandwich. Structurally, the equatorial domain provides most of the intersubunit contacts within each heptameric ring and all the contacts between the two back-to-back stacked heptameric rings. The intermediate domain appears to bridge between the equatorial and the apical domains. Most of the contacts between GroEL and GroES are mediated through the apical domain. GroES (S for small; also known as chaperonin-10) is a single ring of seven identical subunits that forms a dome over one end of the GroEL cylinder (Fig 17.1.42A). Each individual subunit folds into a five-stranded antiparallel β-barrel with four more associated βstrands and a long mobile loop (Fig. 17.1.42D). The end of the barrel near the long loop interacts with the GroEL subunits. In the GroEL complex alone, both heptameric rings are essentially identical; however, when complexed with GroES and ADP at one end, the cis ring (i.e., the one closest to GroES and bound to ADP) of GroEL has dramatically
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different domain orientations than the trans ring (see Fig. 17.1.42B,C). During the cycles of protein folding, the GroEL/GroES complex is proposed to undergo significant conformational changes, including the dissociation and reassociation of the GroES cap (Keskin et al., 2002).
Myosin Subfragment-1 The crystal structure of myosin subfragment-1 (Table 17.1.19) illustrates the overall arrangement of myosin heavy chain, regulatory light chain, and essential light chain (Houdusse et al., 1999; Rayment et al., 1993). The heavy chain forms much of the myosin head as well as an ∼85-Å-long coiled-coil α-helix (Fig. 17.1.42E). The myosin head consists of mostly α-helices with the actin-binding site located near the tip. Both the regulatory and essential light chain subunits bind to the long coiled-coil helical tail of the heavy chain. The structure of the regulatory light chain resembles that of calmodulin with a divalent cation binding site in its first EF-hand region (see above). The essential light chain consists mostly helical elements that wrap around the heavy chain helical tail.
G Proteins and Regulators of G-Proteins Small G proteins and their activators Ras and RasGAP. As a prototype of small G proteins, the structure of p21Ras (Pai et al., 1990) reveals a variation of the mononucleotide binding fold (Fig. 17.1.43A; Table 17.1.19; also see The Classical Mononucleotide-Binding Fold). Like many small G proteins, the intrinsic GTPase activity of Ras is quite low and requires GTPase activation protein (GAP) to increase the enzymatic rate of GTP hydrolysis. The structure of p120GAP is all α-helical (Fig. 17.1.43B; Scheffzek et al., 1996). The structure of a complex between Ras and the Ras guaninenucleotide-exchange-factor region of the son of sevenless (Sos) reveals a tight association between them that buries 3600 Å2 of surface area. This results in the dissociation of the nucleotide from Ras due to alteration of the conformation of the switch 1 and 2 regions (Boriack-Sjodin et al., 1998). The Ras-exchange-factor region of Sos consists of N- and C-terminal α-helical domains, with the C-terminal α-helical domain interacting with Ras (Fig. 17.1.43C). Another group of Ras-like GTP-binding proteins is the Ran family that are involved in nuclear transport. The structure of
Ran-GppNHp has also been determined (Vetter et al., 1999). Rho and RhoGAP. The Rho family of small G proteins includes Rho, Rac, and Cdc42Hs. They regulate the phosphorylation rate of a variety of proteins involved in cell proliferation and cytoskeleton formation. The intrinsic GTPase activity of the Rho family of G proteins (i.e., the rate of hydrolysis of GTP to GDP) is slow but can be increased by 105-fold upon interacting with Rho family of G protein activators. Several crystal structures of Rho family G proteins and their complexes with RhoGAPs have been determined, thus revealing the molecular mechanism of the activation of Rho by its activator proteins (Fig. 17.1.43D). Among them, the structures of Rac1 (Hirshberg et al., 1997) and p50rhoGAP (Barrett et al., 1997), as well as the complexes between RhoA and RhoGAP (Rittinger et al., 1997b), Cdc42Hs and p50rhoGAP, and Rac1 and Tiam 1 have been published (Rittinger et al., 1997a; Worthylake et al., 2000; Table 17.1.19). The structure of Rho assumes a typical Ras-like G protein fold and RhoGAP adopts an α-helical fold similar to the p120GAP. In the structure of the complexes, an Arg residue on RhoGAP interacts with the P-loop of Rho, presumably to stabilize the transition state of the GTP hydrolysis. Large GTP-binding proteins Unlike the Ras-like small G-proteins, the family of large GTP-binding proteins is characterized by higher molecular weight and higher intrinsic GTPase activity. Members of this family include guanylate-binding proteins 1 and 2 (GBP1 and 2), Mx, and dynamin. The crystal structure of an intact human GBP1 with a molecular weight of 67,000 reveals a two-domain fold (Fig. 17.1.43E; Prakash et al., 2000). The N-terminal domain resembles a modified Ras-like G-protein fold with several insertions compared to the canonical Ras structure. The C-terminal domain is elongated and consists of seven α-helices including one that is 118-Å long. GBP1 lacks the conserved nucleotide binding motif (Asn/Thr)-Lys-X-Glu as observed in all Ras-like G-proteins. GBP1 is stable without the bound nucleotide, whereas Ras is not. Heterotrimeric G proteins and their regulators G protein trimer. G protein-coupled transmembrane receptor signaling cascades are ubiquitous throughout eukaryotic cells and util-
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Overview of Protein Structural and Functional Folds
ize heterotrimeric G protein complexes. Crystal structures of two heterotrimeric Gtα/iαGDP Gtβγ complexes reveal the molecular architecture and provide mechanistic insight (Table 17.1.19; Wall et al., 1995; Lambright et al., 1996). The Gtα subunit (Fig. 17.1.43F and G) is organized into three structural components: (1) a Ras-like GTPase domain that contains a mononucleotide-binding fold (see The Classical Mononucleotide-Binding Fold), (2) an all α-helical domain with a central long helix surrounded by five shorter helices, and (3) an N-terminal helix that projects away from the remainder of the Gtα. The Gtβ subunit has an N-terminal helix followed by seven repeating WD motifs that are organized into a β-propeller structure (see Frequently Observed Secondary Structure Assemblies or Structural Motifs). Each WD-structural motif (also known as the Trp-Asp or WD40 motifs) consists of four antiparallel β-strands that form one propeller blade; however, the so-called WD repeat sequence corresponds to the three inner strands of a blade together with the outside (fourth) strand of the preceding blade. The Gtγ subunit consists of two helices that pack against the β subunit. The RGS fold. Regulators of G-protein signaling (RGS) modulate the GTPase activity of heterotrimeric G proteins. Their function is analogous to the GAPs that activate the GTPase activity of Ras-like G proteins (see above). The mechanism of RGS modulation is proposed to be stabilization of the transition state of catalytic Gα subunits, since RGS possesses no affinity for the Gα-GDP complex, modest affinity for the Gα-GTP state, and high affinity for the transition analog Gα-GDP-AlF4− complex. The crystal structures for members of RGS include the structure of RGS4 in complex with GiαGDP-AlF4− (Tesmer et al., 1997), and the structure of the rgRGS domain of p115RhoGEF (Chen et al., 2001). The RGS domain consists of ∼130 amino acids that fold into an all α-helical structure with nine helices divided into two subdomains (Fig. 17.1.43H; Tesmer, 1997). The larger domain, comprised of helices α4 to α7, assumes a classical right-handed antiparallel four-helix bundle fold, while the small domain consists of helices α1 to α3, α8, and α9. Instead of binding to the catalytic residues of the GTPase domain, RGS4 interacts with the three switch regions of Giα, thereby stabilizing their conformation. The GoLoco motif. The nineteen–amino acid GoLoco motif regulates the function of heterotrimeric G proteins by binding to the
GDP-bound form of the Gα subunit, thereby preventing reassociation of Gβγ with the GDPbound Gα. By doing so, GoLoco enables a continued presence of Gβγ which can interact with effector proteins without activating Gα. This motif is present in RGS12, RGS14, LOCO, Purkinje-cell protein-2 (Pcp2), and Rap1GAP isoforms. The crystal structure of a Gαi-GDP bound to the GoLoco region of RGS14 revealed key interactions between the GoLoco motif and the Gαi 1 subunit (Fig. 17.1.43I; Kimple et al., 2002). In particular, the conserved (N/E)QR triad of GoLoco interacts with the nucleotide binding pocket of Gα to make direct contacts with the α- and β-phosphates of GDP. The binding site of GoLoco on the Gαi1 partially overlap with that of Gβγ, thus preventing reassociation of the G protein heterotrimer.
The F1-ATPase (ATP synthase) The F1F0ATPase found in the membranes of mitochondria, chloroplasts, and bacteria is responsible for synthesizing ATP from ADP and inorganic phosphate by utilizing a proton gradient that has been generated across the membrane. The large multisubunit ATPase complex is composed of an F1 globular head catalytic domain linked by a central stalk to an integral membrane F0 proton translocating domain. The F1 domain, including the stalk, consists of five subunits with the stoichiometry α3β3γδε. The F0 domain is composed of a, b, and c subunits with varying stoichiometry. Various species have additional F0 subunits. Crystal structures of the F1 domain and stalk regions from mitochondria, chloroplasts, and bacteria (Table 17.1.19) reveal that the α and β subunits are arranged in an alternating fashion around a pseudosymmetric three-fold axis to make a structure shaped like a slightly squashed pear (Fig. 17.1.44A and B). The α and β subunits are structurally very homologous even though there is marginal sequence identity (20%) between them. They each consist of three domains: an N-terminal six-stranded antiparallel β-barrel, a central nine-stranded mostly parallel β-sheet surrounded on both faces by nine helices, and a C-terminal six- to seven-helix bundle (Fig. 17.1.44B). The central domain of both α and β subunits bind mononucleotides, although only the β subunit is catalytic. Both subunits have a P-loop with a conserved mononucleotide-binding sequence motif at the C-terminal end of the sheet. The γ, δ, and ε subunits fold together intimately to form an α/β foot located just below the α3β3 complex, con-
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nected to it by a long 90-Å α-helix from the γ subunit that runs roughly parallel to the F1 pseudo-three-fold axis (Fig. 17.1.44B; Gibbons et al., 2000). The γ subunit comprises a Rossman-like fold (see The Classical Dinucleotide-Binding Fold) with a five-stranded mostly parallel β-sheet (strand order 3-2-1-4-5) surrounded by six helices. The long, curving C-terminal helix and shorter N-terminal helix of this fold form the stalk region. The eukaryotic δ subunit (bacterial ε subunit) is a ten-stranded β-sandwich followed by a pair of antiparallel helices that project away from one end of the sandwich. The 50-residue mitochondrial ε subunit comprising a helix-loop-helix structure apparently has no counterpart in chloroplast or bacterial ATP synthases (Gibbons et al., 2000). The structure of the entire complex has been more elusive. A 3.9-Å resolution unrefined Cα model of F1F0ATPase from yeast shows ten c subunits from the F0 domain arranged in a circle (Stock et al., 1999). Each c subunit is a pair of antiparallel helices 47- and 58-Å long, respectively. This results in two concentric rings of helices with their helical axes running approximately perpendicular to the plane of the circle (Fig. 17.1.44C). The outer helices are kinked in the middle, giving the barrel a slight hourglass appearance.
Proteasome The proteasome is the central nonlysosomal protein degradation enzyme observed in archeabacterium and eukaryotes. In most prokaryotes, a simpler proteasome homolog such as the ATP-dependent protease Hs1V plays a similar role. In addition to a host of cellular functions, including the degradation of abnormal or misfolded proteins, and the removal of various transient regulatory proteins, the proteasome also plays a major role in generating antigenic peptides for presentation by MHC class I molecules in mammals. The proteasome is composed of a 20S core proteolytic chamber and a 11S or 19S regulatory particle at the ends of the chamber. The crystal structures of archeabacterium Thermoplasma acidophilum (Lowe et al., 1995), yeast (Groll et al., 1997), and bovine (Unno et al., 2002) 20S proteasomes reveal structures consisting of four heptameric rings, α7β7β7α7, stacked on top of each other to form a barrel-shaped complex with an approximate length of 150 Å and a diameter of 110 Å (Fig. 17.1.45, red and green regions). The α and β subunits share the same fold (known as the Ntn
hydrolase fold; see Ntn Hydrolase Fold, consisting of a central β-sandwich flanked by two α-helices on one side, and two or three on the other (Fig. 17.1.35H). The eukaryotic forms consist of seven distinct types of β-subunits and seven distinct α-subunits, whereas the archael proteasome has one type of α- and one type of β-subunit. Only three of the eukaryotic βsubunits (β1, β2, and β5) are fully post-translationally processed to form mature catalytic domains with an N-terminal threonine nucleophile. None of the α-subunits are catalytic. Jawed vertebrates express an additional set of three catalytic β-subunits (β1i, β2i, and β5i) that are incorporated into the immunoproteasome that is responsible for generating MHC ligands for antigen presentation (Unno et al., 2002). The Escherichia coli ATP-dependent protease HslVU forms a dimer of two hexamers (Bochtler et al., 1997). The single HslV subunit is homologous to the β-subunit of proteasomes and has an N-terminal threonine nucleophile. Crystal structures of the 11S regulator alone (human Regα) and in complex with the 20S proteasome (T. brucei 11S and yeast 20S) reveal a barrel-shaped heptamer with outer dimensions of ∼90 Å in diameter and 70 Å in length (Knowlton et al., 1997; Whitby et al., 2000). Each monomer is a bundle of four long α-helices. This barrel is situated at both ends of the 20S particle essentially extending the proteasome cylinder (Fig 17.1.45). The inner diameter is 33 Å at the proteasome-binding end and 24 Å at the opposite end.
Viral Coat Proteins of Spherical Viruses The capsid proteins of a virus provide a protective shell around the enclosed RNA or DNA. These viral coats are made from an assembly of identical or nearly identical small protein subunits. The protein capsids of all known spherical viruses are built using icosahedral symmetry. In its simplest form, an icosahedron is a 20-sided polygon with two-, three-, and five-fold symmetry, made from an arrangement of identical equilateral triangles (Fig. 17.1.46A). Because constructing a protein subunit with equilateral three-fold symmetry is difficult, the simplest known viruses contain three asymmetric subunits per triangle or 60 subunits per virus. Larger and more complex viral coats can be made by further subdividing these asymmetric units and using more protein subunits, all the while maintaining icosahedral symmetry. Viruses can be classified by their triangulation (T) number which indicates the
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number of capsid subunits in multiples of 60. For example, a T = 1 virus has 60 subunits and a T = 3 virus has 3 × 60 = 180 subunits. The three-dimensional structures of numerous capsid proteins for spherical viruses that infect bacteria, plants, insects, and animals have revealed a common fold for most of these structures, namely an eight-stranded jelly-roll β sandwich (see Elements of Protein Structure). Strands 1-8-3-6 form one sheet and strands 2-7-4-5 form the other sheet of the β sandwich (Fig. 17.1.46B). Despite the lack of sequence homology between various classes of viral-coat proteins, the jelly-roll β sandwich topology is well conserved in >50 known capsid structures according to SCOP classification of the PDB (Murzin et al., 1995). It forms a wedge-shaped β sandwich, generally with short loops at one end and longer loops at the other. A striking exception to this fold is observed in the coat protein of bacteriophage MS2, which is a T = 3 virus (see MS2 Phage Coat Protein).
INTEGRAL MEMBRANE PROTEINS
Overview of Protein Structural and Functional Folds
High-resolution structural information is available for only ∼50 integral membrane proteins (Table 17.1.20), compared with over 10,000 soluble proteins, although an estimated 15% to 30% of genes encode membrane proteins. Difficulties in protein expression, solubilization, and the growth of well-ordered crystals have plagued structural studies. Nonetheless, remarkable progress has been made in solving structures of integral membrane proteins owing to advances in overexpression, availability of detergents, and the use of antibody fragments in crystallization. Complementary methodological approaches, including Xray diffraction, cryo-electron microscopy, NMR spectroscopy, and electron paramagnetic resonance spectroscopy promise even faster future progress. Updated compendiums of structures of membrane proteins can be found at http://blanco.biomol.uci.edu/Membrane_ Proteins_xtal.html and http://www. mpibpfrankfurt.mpg.de/michel/public/memprotstru ct.html. The diversity of integral membrane protein folds appears to be much less than soluble proteins (Schulz, 2002). Due to the thermodynamic cost of burying charges in the hydrophobic environment of the membrane, integral membrane structures must minimize exposed charges and unsatisfied hydrogen bonds. Two classes of structures are observed satisfying these requirements: the α-helix and the closed
β-barrel. The α-helical structures are most prevalent and are found in cytoplasmic membrane proteins. They include both transmembrane α-helices and α-helical structures in monotopic integral membrane proteins which do not cross the bilayer. β-barrel structures are characteristic of proteins of the outer membranes of Gram-negative bacteria.
Photosynthetic Proteins The first high-resolution structure determination of an integral membrane protein was a photosynthetic reaction center from the purple bacterium Rhodopseudomonas virdis, for which Johann Deisenhofer, Robert Huber, and Hartmut Michel were awarded the Nobel Prize in Chemistry in 1988 (Deisenhofer et al., 1984, 1985, 1995; Deisenhofer and Michel, 1989). This large complex includes four protein subunits and fourteen cofactors. The transmembrane core of the complex has pseudo two-fold symmetry, formed by the structurally related chains L and M; each of which contains five membrane-spanning α-helices. The chlorophyll, quinone, carotenoid, and nonheme iron cofactors associate with the L and M subunits. The H subunit forms an additional transmembrane α-helix and also contains β structures on the cytoplasmic face. The eleventransmembrane helices are arranged approximately parallel to each other (Fig. 17.1.47A). The cytochrome subunit, bound to four heme groups, is on the periplasmic face and does not span the membrane. More recently, additional photosynthetic complexes have been solved, including the reaction center of Rhodobacter sphaeroides, the two photosystems of chloroplasts, and antenna light-harvesting complexes (Axelrod et al., 2002; Chirino et al., 1994; Jordan et al., 2001; Koepke et al., 1996; Prince et al., 1997; Zouni et al., 2001).
Seven-Helix Proteins A number of families of integral membrane proteins contain seven membrane-spanning αhelices. Among them, the microbial rhodopsins are the best studied. The crystallographic studies of bacteriorhodopsin from Halobacterium salinarum have provided insight into the mechanism of this light-driven proton pump (Lanyi, 1999; Subramaniam, 1999). The helices pack around a covalently linked retinal chromophore, forming an internal channel lined by hydrophilic residues (Fig. 17.1.47B). Conformational changes in the transmembrane regions have been observed during the course of the photocycle (Kataoka and Kamikubo,
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2000). More recently, structures of two related haloarchaeon proteins, halorhodopsin (Kolbe et al., 2000) and sensory rhodopsin II (Luecke et al., 2001; Royant et al., 2001; Gordeliy et al., 2002), have also been determined. The versatility of the seven transmembrane helix bundle is apparent. For example, bacteriorhodopsin and halorhodopsin are ion transporters, while sensory rhodopsin participates in photosensory signaling and interacts with transducer proteins. Structural differences, particularly in the retinal binding pocket and conformation of bound retinal, reflect the distinct functional role of sensory rhodopsin (Luecke et al., 2001; Royant et al., 2001). In higher organisms, the seven-helix fold is seen in receptors coupled to G proteins, a large family of integral membrane proteins. The crystal structure of the mammalian retinal photoreceptor rhodopsin represents the first highresolution structure of a G protein-coupled receptor (Palczewski et al., 2000). Although the overall topology of the seven-helix bundle of bovine rhodopsin resembles that of microbial rhodopsins, its structure appears unique. Unlike the parallel arrangement of the helices in bacteriorhodopsin, some of the bovine rhodopsin helices are tilted or strongly bent at proline residues (Fig. 17.1.47C). Moreover, bovine rhodopsin contains more extensive secondary structures in the intra- and extracellular regions, consistent with the disparate receptor function. There is also an additional short helix that lies parallel to the membrane surface on the cytoplasmic face. This amphipathic eighth helix is of particular interest because of its location in the G protein binding region. On the extracellular face, the three loops associate with portions of the N-terminus to form a compact structure centered around two twisted β hairpins. The second extracellular loop, which contains the inner hairpin, is linked through a conserved disulfide bond to transmembrane helix 3, and forms a plug covering the chromophore binding pocket (Bourne and Meng, 2000).
Ion Channels Ion channels allow movement of selected ions along electrochemical gradients, and can open and close (i.e., gate) in response to various stimuli. A successful approach to their structural study has involved use of bacterial homologs. Two bacterial potassium channel structures have been determined: the KcsA H+gated channel and the Ca2+-gated MthK channel (Doyle et al., 1998; Jiang et al., 2002a,b).
The bacterial mechanosensitive channels MscL and MscS, which respond to mechanical stretching of the cell membrane, have also been solved (Bass et al., 2002; Chang et al., 1998). Structures of two ClC chloride channels were recently determined from E. coli and Salmonella typhimurium (Dutzler et al., 2002). The ClC chloride channel is dimeric, with each subunit forming its own pore to make up a double-barreled structure, as predicted by biophysical studies (Fig. 17.1.47D). The secondary structure is all α-helical, with each subunit containing seventeen membrane-inserted αhelices. Many of the membrane-inserted helices are tilted relative to the plane of the membrane, and not all pass entirely through the bilayer. The crystal structure revealed an internal structural repeat, wherein the N-terminal portion of each subunit is structurally related to the C-terminal portion. This unanticipated internal symmetry results in an antiparallel arrangement of the subunit halves. A selectivity filter is formed by helix dipoles and main chain atoms that interact with bound Cl−. The partial positive charge generated by the helix dipoles likely supports electrostatic stabilization of Cl− but does not result in tight binding (and thus does not restrict Cl− movement through the pore). The carboxylate of a conserved glutamic acid near the chloride-binding site protrudes into the pore, suggesting a gating function to open and close the channel through electrostatic repulsion.
ABC Transporters The large family of ATP binding cassette (ABC) transporters are involved in ATP-driven import and export of a variety of substrates. The functional transporters contain two transmembrane domains and two cytoplasmic nucleotide-binding domains. The structures of MsbA lipid transporter (flippase) and BtuCD vitamin B12 transporter are two members of this family solved to date (Chang and Roth, 2001; Locher et al., 2002). The membrane-spanning portion of the E. coli BtuCD transporter, made up of two BtuC subunits, has a total of 20 transmembrane α-helices that display a complex packing arrangement (Fig. 17.1.47E). The translocation channel seems to be located at the BtuC dimer interface. The two BtuD cassettes interact with each other and are attached to the BtuC module on the cytoplasmic face. A large, water-filled vestibule located in the cytoplasm between the four subunits may allow vitamin B12 to exit the transporter; two cytoplasmic loops of BtuC appear to serve as the channel gate. Comparison
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of BtuCD with MsbA shows that the arrangement and packing of the transmembrane domains differ significantly. While in BtuCD, both the cytoplasmic and the transmembrane domains of each subunit remain in close contact, in MsbA they are oriented away from each other.
P-type ATPases The structure of a calcium ATPase from skeletal-muscle sarcoplasmic reticulum has been determined in both E1 (Ca2+-bound) and E2 (Ca2+-free) states, providing details of the mechanism of this P-type ATPase (Toyoshima et al., 2000; Toyoshima and Nomura, 2002). The Ca2+ ATPase has ten transmembrane αhelices and three cytoplasmic domains. Two calcium ions are bound at the center of four transmembrane helices in the E1 state, and two of these helices are in unwound conformations to efficiently coordinate the ions. Four transmembrane helices, one of which is 41-residues long, appear to extend through the bilayer into the cytoplasm, forming a stalk region that connects the transmembrane portion to the cytoplasmic domains. The crystal structure of the E2 state revealed that large movements of the cytoplasmic domains and rearrangements of the transmembrane helices take place during the reaction cycle (Fig. 17.1.47F; Toyoshima and Nomura, 2002).
Other Channels and Transporters
Overview of Protein Structural and Functional Folds
The structure of E. coli efflux transporter AcrB, a major multidrug exporter which cooperates with the β-barrel membrane protein TolC (see below), was recently determined by X-ray crystallography (Murakami et al., 2002b). The first structure of a mammalian channel protein was that of a bovine aquaporin water channel, AQP1, which was studied by electron crystallography, electron microscopy, and X-ray crystallography (Murata et al., 2000; Ren et al., 2001; Sui et al., 2001). AQP1 shares the same overall topology as its bacterial homolog GlpF, a glycerol channel (Fu et al., 2000). AQP1 contains a quadruple-barreled channel, with individual pores formed by each of its four subunits. Each monomer has six transmembrane and two membrane-inserted α-helices (Fig. 17.1.47G). Severe tilting of the transmembrane helices relative to the membrane normal, as was seen in the ClC chloride channel, allows for formation of the dumbbell-shaped pore in AQP1. The structure of the pore reveals that water selectivity is based on a narrow constric-
tion region that imposes a 2.8-Å diameter steric limit on the channel.
Respiratory Enzymes Several structures of respiratory enzymes have been solved, including both bacterial and mammalian cytochrome c oxidase (Iwata et al., 1995; Tsukihara et al., 1996; Ostermeier et al., 1997; Soulimane et al., 2000), fumarate reductase (Iverson et al., 1999; Lancaster et al., 1999), and formate reductase-N (Jormakka et al., 2002). Crystal structures of the bovine and chicken cytochrome bc1 complex (complex III) as well as its Saccharomyces cerevisiae homolog provide intriguing insights into the function of the respiratory chain (Xia et al., 1997; Iwata et al., 1998; Zhang et al., 1998; Hunte et al., 2000). The complex functions as a dimer of multisubunit monomers. In the mammalian bc1 complex, seven of the eleven subunits have membrane-spanning components, forming a total of 26 transmembrane helices in the dimer (Fig. 17.1.47H). Conformations of the Rieske iron-sulfur protein in different crystal forms have led to the suggestion that the soluble portion of the subunit may move during catalysis to facilitate the transfer of electrons from ubiquinol to cytochrome c1 (Iwata et al., 1998; Zhang et al., 1998).
β-Barrel Membrane Proteins A distinct class of membrane proteins contain membrane-spanning β-strands arranged into closed barrels. Transmembrane β-barrel structures are composed of 8 to 22 antiparallel strands (Schulz, 2002). This class is best illustrated by porins, which are channels of the outer membrane of Gram-negative bacteria (Fig. 17.1.1K). Related to the porins are the iron transporters FepA and FhuA, which form 22stranded barrels housing globular plug domains (Fig. 17.1.47I; Buchanan et al., 1999; Ferguson et al., 2002). Both porins and maltoporins appear as trimers with each subunit forming one β-barrel channel. A more unusual structure is TolC, an E. coli outer membrane protein involved in export of a range of molecules. Three protein subunits each contribute a fourstranded β sheets to form a single twelvestranded outer membrane-spanning β-barrel (Fig. 17.1.47J). In addition, TolC also has a large, left-handed α-helical barrel stabilized by coiled-coil interactions. The α-helical barrel is contiguous with the β-barrel domain, and extends 100 Å into the periplasm. This results in the formation of a 140-Å long channel-tunnel that spans both the outer membrane and the
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periplasmic space, allowing interaction with inner membrane proteins.
Andrade, M.A. and Bork, P. 1995. HEAT repeats in the Huntington’s disease protein. Nat. Genet. 11:115-116.
ACKNOWLEDGEMENTS
Andrade, M.A., Petosa, C., O’Donoghue, S.I., Muller, C.W., and Bork, P. 2001. Comparison of ARM and HEAT protein repeats. J. Mol. Biol. 309:1-18.
The authors would like to thank L. Birnbaumer, T. Darden, and T. Transue for the insightful comments and suggestions they contributed to this manuscript.
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Overview of Protein Structural and Functional Folds
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Contributed by Peter D. Sun and Christine E. Foster National Institute of Allergy and Infectious Diseases National Institutes of Health Rockville, Maryland Jeffrey C. Boyington National Institute of Environmental Health Science National Institutes of Health Research Triangle Park, North Carolina
APPENDIX The following section lists all of the Tables and Figures mentioned throughout the text (figures follow tables). All ball-and-stick models and ribbon diagrams appearing in this unit were created from Protein Data Bank (PDB) coordinates (Berman et al., 2000; also see Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb) using the program MolScript (Kraulis, 1991) or Raster3D (Merritt and Bacon, 1997). Unless otherwise noted, α-helices and β-strands are colored red and green, respectively. For reasons of clarity, certain proteins are colored by domain or by subunit as designated in the figure legends. Ball-and-stick models of amino acid side chains or ligands are colored by atom type: carbon, black; oxygen, red; nitrogen, blue; and sulfur, yellow. DNA and RNA are represented by blue ball-and-stick models. Metal ions are shown as colored spheres. Note that the black and white facsimile of the figures throughout the printed version of this unit are intended only as placeholders; for the full-color version of figures go to the Current Protocols website at Wiley Interscience (http://www.interscience.wiley.com/ c_p/colorfigures.htm).
Overview of Protein Structural and Functional Folds
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Table 17.1.1
Immunoglobulin and Other Immunoreceptor Folds
Protein/complex
Domain/region
Folda
PDBb
References
1igt
Harris et al. (1997)
2clr
(Bjorkman et al., 1987; Collins et al., 1994b)
1aqd
Brown et al. (1993)
1ao7
Garboczi et al. (1996)
1cd8 1fru
Leahy et al. (1992a) Burmeister et al. (1994)
C2(Fn) C2(Fn)
3hhr 1ern
de Vos et al. (1992) Livnah et al. (1999)
C2(Fn) C2(Fn) I-C2 C2(Fn) C2(Fn) C2(Fn) C2(Fn)
1bp3 1fg9 1e0o 1cd9 1j7v 1iar 1i1r 3cd4
Somers et al. (1994) Walter et al. (1995) Pellegrini et al. (2000) Aritomi et al. (1999) Josephson et al. (2001) Hage et al. (1999) Chow et al. (2001) Ryu et al. (1990), Wang et al. (1990)
1fnl 1fcg 1f2q
Zhang et al. (2000) Maxwell et al. (1999) Garman et al. (1998)
Immunoglobulins Antibodies Variable region Constant region Major histocompatibility complexes MHC class I
V- type C1-type
Heavy chain α1+α2 domain MHC Heavy chain α3 domain C1-type β2-microglobulin C1-type MHC class II Heavy chain α1+β1 domain α chain, C-terminal domain β chain, C-terminal domain Cell surface immunoglobulin-like receptors T-cell receptor, αβ type Vα, Vβ domains Cβ domain CD8, T-cell coreceptor Neonatal Fcγ receptor Heavy chain α3 domain β chain Growth hormone receptor Domains 1 and 2 EPO receptor Prolactin receptor Domains 1 and 2 IFN-γ receptor FGFR2 GCSFR IL-10R IL-4R Gp130 receptor CD4, T-cell surface glycoprotein Domains 1 and 3 Domains 2 and 4 Fcγ receptor, type III Fcγ receptor, types IIA and IIB Fcε receptor, type I
MHC C1-type C1-type
V-type C1-type V-type C1-type C1-type
V-typec C2-like C2-like C2-like C2-like
continued
Structural Biology
17.1.91 Current Protocols in Protein Science
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Table 17.1.1
Immunoglobulin and Other Immunoreceptor Folds, continued
Protein/complex
Domain/region
Coagulation factor Tissue factor Ectodomain Cell matrix and adhesion proteins Fibronectin Type III domain Tenascin, repeats CD2 Adhesion domain C-terminal domain VCAM-1 Domain 1 Domain 2 d Cadherin Muscle proteins Telokin Titin Transcription factorse NF-κB, p50 subunit
Folda
PDBb
C2(Fn)
2hft
Muller et al. (1996)
C2(Fn) C2(Fn)
1fna 1ten 1hnf
Main et al. (1992) Leahy et al. (1992b) Jones et al. (1992)
V-type C2 like 1vca
Jones et al. (1995)
I-type C2 like C1 type
1nci
Shapiro et al. (1995)
I-type I-type
1tlk 1tnm
Holden et al. (1992) Pfuhl and Pastore (1995)
1nfk
Ghosh et al. (1995), Muller et al. (1995) Muller et al. (1995)
First Rel homology domainf E-type Second Rel homology E-type domain Enzymes with immunoglobulin-like domains Chaperone protein, papD
Oxidoreductase Cu, Zn superoxide dismutasei Galactose oxidase Transferase Cyclodextrin glycosyltransferase Lyase Chitinase A C-type lectin-like receptors Mannose binding protein (MBP) CD94 Ly48A NKG2D DC-SIGN
References
1svc
3dpa
Holmgren and Branden (1989)
Domain 1g Domain 2h
C2 C2
C domain
C2 E-type
2sod 1gof
Richardson et al. (1976) Ito et al. (1991)
E-type
1cdg
Lawson et al. (1994)
E-type
1ctn
Perrakis et al. (1994)
CTLR
1msb
Weis et al. (1991)
CTLR CTLR CTLR CTLR
1b6e 1qo3 1hq8 1k9j
Boyington et al. (1999) Tormo et al. (1999) Wolan et al. (2001) Feinberg et al. (2001)
N-terminal domain
continued
Overview of Protein Structural and Functional Folds
17.1.92 Supplement 35
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Table 17.1.1
Immunoglobulin and Other Immunoreceptor Folds, continued
Folda
PDBb
References
Immunoreceptor-ligand complexes A6 TCR/HLA-A2(Tax) 2C TCR/H-2Kb(dEV8) CD8/HLA-A2 CD4/I-A(k) B7-1/CTLA-4 KIR2DL2/HLA-Cw3 KIR2DL1/HLA-Cw4 NKG2D/MICA NKG2D/ULBP3 NKG2D/Rae-1b FcεRI/Fc FcγRIII/Fc
TCR-MHC TCR-MHC CD8/MHC-I CD4/MHC-II B7/CTLA-4 KIR/HLA KIR/HLA NKG2D/ligand NKG2D/ligand NKG2D/ligand FcR/Fc FcR/Fc
1ao7 2ckb 1akj 1jl4 1i84 1efx 1im9 1hyr 1kcg 1jsk 1f6a 1iis,1iix, 1e4k
Garboczi et al. (1996) Garcia et al. (1998) Gao et al. (1997) Wang et al. (2001) Stamper et al. (2001) Boyington et al. (2000) Fan et al. (2001) Li et al. (2001) Radaev et al. (2001b) Li et al. (2002b) Garman et al. (2000) Radaev et al. (2001a), Sondermann et al. (2000)
MHC-like molecules HLA-A2 HLA-Aw6B HLA-B27 H-2Kb
MHC MHC MHC MHC
2clr 1hsb 1hsa 1vac
HLA-DR1
MHC
1aqd
Neonatal Fc receptor HLA-DM HLA-E CD1 Zinc α-2-glycoprotein (Zag) MIC-A T22
MHC MHC MHC MHC MHC MHC MHC
1fru 1k8i 1mhe 1cd1 1zag 1b3j 1c16
Bjorkman et al. (1987) Garrett et al. (1989) Madden et al. (1992) Fremont et al. (1992), Zhang et al. (1992) Brown et al. (1993), Stern et al. (1994) Burmeister et al. (1994) Fremont et al. (1998) O’Callaghan et al. (1998) Zeng et al. (1997) Sanchez et al. (1999) Li et al. (1999) Wingren et al. (2000)
Protein/complex
Domain/region
aSee Proteins Involved in the Function of Immune Systems for explanations of folds. bSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb. cStrand A pairs with strand G only. dStrand A pairs with strand G instead of B. eSee also Table 17.1.6. fInsertion strands between strands C and C′, and E and F. gStrand A pairs with both B and G. hOne additional strand at C terminus. iOne additional strand at N terminus.
Structural Biology
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Table 17.1.2
Protein
Complement System Protein Folds
PDB entrya
C3 fragment C3d 1c3d C5a anaphylatoxin 1kjs Complement factor D 1dsu CD59 (1-70) 1cdq,1erg Factor H CCP module 16 1hcc Factor H CCP modules 15 and 16 1hfh 1vvc CCP modules 3 and 4b SCR modules 1 and 2c 1ckl
Reference Nagar et al. (1998) Zhang et al. (1997) Narayana et al. (1994) Fletcher et al. (1994), Kieffer et al. (1994) Norman et al. (1991), Barlow et al. (1992) Barlow et al. (1993) Wiles et al. (1997) Casasnovas et al. (1999)
aSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb. bFrom Vaccinia virus complement control protein mimic. cFrom CD46.
Overview of Protein Structural and Functional Folds
17.1.94 Supplement 35
Current Protocols in Protein Science
Table 17.1.3
Cytokines and Receptors
Protein Four-helix bundle cytokines Long chain, G-CSF Short chain, IL-4 Interferon-γ-like IL-10 β-trefoil cytokines IL-1α IL-1β Fibroblast growth factor
Cystine-knot cytokines TGF-β
PDB entrya
References
1rhg 1rcb 1rfb 1ilk
Hill et al. (1993) Rozwarski et al. (1994), Smith et al. (1994) Ealick et al. (1991) Zdanov et al. (1995)
2ila 1hib
Graves et al. (1990) Clore and Gronenborn (1991), Priestle et al. (1989) Eriksson et al. (1991) Zhang et al. (1991) Zhu et al. (1991)
4fgf 2fgf 1bas, 1bar 2tgi
Nerve growth factor 1bet Platelet-derived growth factor 1pdg Bb Human chorionic gonadotropin 1hrp, 1hcn Chemokines IL-8 1il8 MIP-1β 1hum RANTES 1rto Neutrophil activating peptide-2 1nap TNF receptor superfamily TNFR 1tnr TRAIL 1d4v Receptors for cystine-knot growth factors ActRII 1bte BRIA 1es7 TBRII 1ktz TrkA 1www Domain 2 of Flt-1 1flt Nuclear receptors TR 1bsx RAR 2lbd ER 2ert PR 1a28 PPAR 1kkq Importin-α 1ial Importin-β 1qgr Adhesion receptors Integrin I domain (LFA-1) 1lfa Integrin I domain (CR3) 1ido Integrin (intact) 1jv2 β-catenin/E-cadherin 1i7x β-catenin/Tcf 1g3j Cysteine-rich scavenger receptor SRCR domain of Mac-2 1by2 binding protein Glutamate receptors iGluR 1gr2 mGluR 1ewt, 1ewv Transferrin receptor Human transferrin receptor 1cx8
Daopin et al. (1992, 1993), Schlunegger and Grutter (1993) McDonald et al. (1991) Oefner et al. (1992) Lapthorn et al. (1994), Wu et al. (1994) Clore et al. (1990) Lodi et al., 1994 Skelton et al. (1995) Malkowski et al. (1995) Banner et al. (1993) Mongkolsapaya et al. (1999) Greenwald et al. (1999) Kirsch et al. (2000) Hart et al. (2002) Wiesmann et al. (1999) Wiesmann et al. (1997) Darimont et al. (1998) Renaud et al. (1995) Shiau et al. (1998) Williams and Sigler (1998) Xu et al. (2002) Kobe (1999) Cingolani et al. (1999) Qu and Leahy (1995) Lee et al. (1995) Xiong et al. (2001) Huber and Weis (2001) Graham et al. (2000) Hohenester et al. (1999)
Armstrong et al. (1998) Kunishima et al. (2000) Lawrence et al. (1999)
aSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb.
Structural Biology
17.1.95 Current Protocols in Protein Science
Supplement 35
Table 17.1.4
Modular Domains Involved In Signal Transduction
Protein Domains that bind phosphotyrosine SH2 domains v-src tyrosine kinase SH2 domain p85 α subunit of PI 3-kinase SH2 C-abl SH2 domain PTB domains Shc PTB domain IRS-1 PTB domain
PDB entrya
References
1sha 2pnb 1ab2
Waksman et al. (1992) Booker et al. (1992) Overduin et al. (1992)
1shc 1irs
Zhou et al. (1995) Eck et al. (1996), Zhou et al. (1996)
Phosphoserine and phosphothreonine binding domains FHA domains RAD53 FHA domain 1g6g, 1dmz 14-3-3 domains 14-3-3 ζ 14-3-3 τ Polyproline binding domains SH3 domains α Spectrin SH3 domain Chicken c-src SH3 domain p85 α subunit of PI 3-kinase SH3 ww domains Kinase associated protein WW Distrophin WW domain EVH1 domains Murine enabled EVH1 domain Murine ena/vasp-like EVH1 domain GYF domains Drosophila CD2BP2 GYF domain Phospholipid binding domains PH domains Dynamin PH domain β-spectrin PH domain Pleckstrin PH domain C1 domains Rat protein kinase C-γ C1 domain Rat protein kinase C-δ C1 domain C2 domains Synaptotagmin I C2 domain Phospholipase C-δ C2 domain Protein kinase C-α C2 domain FYVE domains Hrs FYVE domain Vps27p FYVE domain Eea1 FYVE domain Protein interaction domains PDZ domains PSD-95 third PDZ domain Dlg third PDZ domain VHS domains Hrs VHS domain Tom1 VHS domain GGA1 VHS domain Overview of Protein Structural and Functional Folds
Liao et al. (1999), Durocher et al. (2000)
1a40 —
Liu et al. (1995) Xiao et al. (1995)
1shg 1rlp 2pni
Musacchio et al. (1992) Yu et al. (1992) Booker et al. (1993)
— 1eg4
Macias et al. (1996) Huang et al. (2000)
1evh 1qc6
Prehoda et al. (1999) Fedorov et al. (1999)
1gyf
Freund et al. (1999)
1dyn 1btn 1pls
Downing et al. (1994), Ferguson et al. (1994) Macias et al. (1994) Yoon et al. (1994)
— 1ptq
Hommel et al. (1994) Zhang et al. (1995)
I1rsy 1qas 1dsy
Sutton et al. (1995) Grobler et al. (1996) Verdaguer et al. (1999)
1dvp 1vfy 1joc
Mao et al. (2000) Misra and Hurley (1999) Dumas et al. (2001)
1be9, 1bfe 1pdr
Doyle et al. (1996) Morais Cabral et al. (1996)
1dvp 1elk 1jwg, 1jwf
Mao et al. (2000) Misra et al. (2000) Shiba et al. (2002) continued
17.1.96 Supplement 35
Current Protocols in Protein Science
Table 17.1.4
Modular Domains Involved In Signal Transduction, continued
Protein
PDB entrya
References
GGA3 VHS domain SNARES Syntaxin-1A/synaptobrevin-II/SNAP-25B SAM domains EphB2 receptor SAM domain
1juq, 1jpl
Misra et al. (2002)
1sfc
Sutton et al. (1998)
1b4f, 1sgg
EphA4 receptor SAM domain Polyhomeotic SAM domain MH2 domains Smad2 MH2 domain Smad3 MH2 domain Structural repeat motifs Leucine rich repeats (LRRs) Ribonuclease inhibitor
1b0x 1kw4
Smalla et al. (1999), Thanos et al. (1999) Stapleton et al. (1999) Kim et al. (2002)
1dev 1mk2
Wu et al. (2000) Qin et al. (2002)
2bnh
Ribonuclease inhibitor/RNAse A
1dfj
Internalin B RanGAP protein rna1p HEAT repeats Protein phosphatase 2A PR65/A subunit Karyopherin 2/Ran-GppNHp complex Armadillo (ARM) repeats Karyopherin α β-catenin β-catenin bound to E-cadherin Ankyrin (ANK) repeats GABPβ p16INK4a bound to Cdk6 IκBα/NF-κB complex
1d0b 1yrg
Kobe and Deisenhofer (1993) Kobe and Deisenhofer (1995) Marino et al. (1999) Hillig et al. (1999)
1b3u 1qbk
Groves et al. (1999) Chook and Blobel (1999)
1bk5, 1bk6 3bct 1i7w
Conti et al. (1998) Huber et al. (1997) Huber and Weis (2001)
1awc 1bi7 1ikn
Gorina and Pavletich (1996) Russo et al. (1998) Huxford et al. (1998)
aSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb. For proteins with no entry listed, coordinates
have not been deposited in the Protein Data Bank.
Structural Biology
17.1.97 Current Protocols in Protein Science
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Table 17.1.5
Kinases and Phosphatases
Protein
PDB entrya
References
Protein kinases
Overview of Protein Structural and Functional Folds
Protein tyrosine kinases Tyrosine kinase C-src Lymphocyte cell kinase (lck) Haematopoietic cell kinase (hck) C-terminal src kinase Tie2 protein tyrosine kinase Human insulin receptor kinase Insulin-like growth factor-1 receptor FGF receptor-1 EPHB2 receptor Vascular growth factor receptor-2 Protein serine/threonine kinases Cyclic AMP kinases
1fmk 3lck 1ad5 1byg 1fvr 1irk 1k3a 1fgk 1jpa 1vr2
Xu et al. (1997a) Yamaguchi and Hendrickson (1996) Sicheri et al. (1997) Lamers et al. (1999) Shewchuk et al. (2000) Hubbard et al. (1994) Favelyukis et al. (2001) Mohammadi et al. (1996) Wybenga-Groot et al. (2001) McTigue et al. (1999)
1atp
Cyclin-dependent kinase 2
1bi7
Cyclin A-CDK Cell cycle protein CksHs1 MAP kinase ERK2 MAP kinase p38 Phosphorylase kinase Calcium/calmodulin-dependent kinase Twitchin kinase Casein kinase I Death associated protein kinase
1fin 1dks 1gol 1p38 1phk 1ao6 1koa 1cki 1jks
Knighton et al. (1991), Olah et al. (1993) De Bondt et al. (1993), Jeffrey et al. (1995) Jeffrey et al. (1995) Arvai et al. (1995) Zhang et al. (1994) Wang et al. (1997) Owen et al. (1995) Goldberg et al. (1996) Hu et al. (1994) Xu et al. (1995) Tereshko et al. (2001)
1phr
Su et al. (1994)
2hnq 1gwz 2shp 1ypt 1g4u 1lar
Barford et al. (1994) Yang et al. (1998) Hof et al. (1998) Stuckey et al. (1994) Stebbins and Galan (2000) Nam et al. (1999)
1vhr 1fpz 1mkp 1d5r
Yuvaniyama et al. (1996) Song et al. (2001) Stewart et al. (1999) Lee et al. (1999)
1c25 1hzm
Fauman et al. (1998) Farooq et al. (2001)
1fjm 1aui 1g5b 1kbp
Goldberg et al. (1995) Kissinger et al. (1995) Voegtli et al. (2000) Strater et al. (1995)
Phosphatases Protein tyrosine phosphatases Subtype I Low molecular weight Subtype II High molecular weight PTB1 SHP-1 SHP-2 Yersinia PTP SptP tyrosine phosphatase domain RPTP LAR tandem phosphatase Dual specificity VHR Kinase associated phosphatase (kap) MAP kinase phosphatase PTEN tumor suppressor Third subtype CDC25A ERK2 binding domain of MKP-3 Protein serine/threonine phosphatases PPP family enzymes PP1 Calcineurin Bacteriophage λ phosphatase Purple acid phosphatase
continued
17.1.98 Supplement 35
Current Protocols in Protein Science
Table 17.1.5
Kinases and Phosphatases, continued
Protein
PDB entrya
References
UDP sugar hydrolase MRE11 nuclease PPM family Protein phosphatase 2C Alkaline phosphatase fold Alkaline phosphatase Arylsulfatase Prokaryotic cofactor-independent phosphoglycerate mutase
1ush 1ii7
Knofel and Strater (1999) Hopfner et al. (2001)
1a6q
Das et al. (1996)
1ali 1auk 1eqj
Kim and Wyckoff (1991) Lukatela et al. (1998) Jedrzejas et al. (2000)
5fbp 2hhm 1inp 1qgx 1jp4
Ke et al. (1991) Bone et al. (1992) York et al. (1994) Albert et al. (2000) Patel et al. (2002)
1fin 1vol 1gux
Jeffrey et al. (1995) Nikolov et al. (1995) Lee et al. (1998)
Sugar phosphatases Fructose 1,6-biosphastase Inositol monophosphatase Inositol polyphosphate 1-phosphatase 3,5-adenosine bisphosphatase PIPase The cyclin fold Cyclin A Transcription factor IIB Retinoblastoma tumor suppressor protein
aSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb.
Structural Biology
17.1.99 Current Protocols in Protein Science
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Table 17.1.6
DNA-Binding Proteins
Protein Helix-turn-helix motifs Prokaryotic and phage repressors λ Cro protein λ-repressor 434 repressor 434 cro protein Homeodomains or homeodomain-like Engrailed Antennapedia Oct-1 POU Proto onco gene product Myb Hin-related prokaryotic DNA recombinase Winged helix HNF-3/fork head transcription factor Globular domain of histone H5 ETS transcription factor E. coli CAP protein Biotin repressor PurR repressor PurR repressor Zinc-containing DNA-binding motifs Cys2 His2 zinc fingers ZIF286 GLI oncogene product Tramtrack Zinc ribbon motif Transcription elongation factor TFIIS Transcription initiation factor TFIIB RNA Polymerase II RPB9 fragment DNA primase fragment GATA-binding motif GATA-1 Zn2Cys6 binuclear cluster GAL-4 Pyrimidine pathway regulator 1(PPR1) PUT3 CD2-LAC9 Nuclear hormone receptor DNA-binding domains Glucocorticoid receptor Estrogen receptor Orphan nuclear receptor NGFI-B Thyroid hormone receptor MADS box domain Serum response factor MCM1 transcriptional regulator MEF2A core Basic leucine zipper GCN4 homodimer c-Fos:c-Jun heterodimer
PDB entrya
Reference
1cro 1lmb 2or1 2cro
Anderson et al. (1981) Jordan and Pabo (1988) Aggarwal et al. (1988) Wolberger et al. (1988)
1hdd 1hom 1oct 1mse 1hcr
Kissinger et al. (1990) Qian et al. (1989) Klemm et al. (1994) Ogata et al. (1994) Feng et al. (1994)
2hfh 1hst 1etc 1cgp 1bia
Clark et al. (1993), Marsden et al. (1998) Ramakrishnan et al. (1993) Donaldson et al. (1996) Schultz et al. (1991) Wilson et al. (1992)
1pru
Schumacher et al. (1994)
1zaa 2gli 2drp
Pavletich and Pabo (1991) Pavletich and Pabo (1993) Fairall et al. (1993)
1tfi 1pft 1qyp 1d0q
Qian et al. (1993) Zhu et al. (1996) Wang et al. (1998) Pan and Wigley (2000)
1gat
Omichinski et al. (1993)
1d66 1pyi 1zme 1cld
Marmorstein et al. (1992) Marmorstein and Harrison (1994) Swaminathan et al. (1997) Gardner et al. (1995)
1gdc 1hcq 1cit 2nll
Luisi et al. (1991) Schwabe et al. (1993) Meinke and Sigler (1999) Rastinejad et al. (1995)
1srs 1mnm 1egw
Pellegrini et al. (1995) Tan and Richmond (1998) Santelli and Richmond (2000)
1ysa 1fcs
Ellenberger et al. (1992) Glover and Harrison (1995) continued
Overview of Protein Structural and Functional Folds
17.1.100 Supplement 35
Current Protocols in Protein Science
Table 17.1.6
DNA-Binding Proteins, continued
Protein
PDB entrya
Helix-loop-helix MAX transcription factor MyoD transcription activator Upstream stimulatory factor Transcription factor E47 HMG box domain Human SRY LEF-1 transcription factor HMG1 B domain Histone-fold proteins Nucleosomal core octamer (H2A, H2B, H3, H4) Drosophila TAFII42 and TAFII62 Human TAFII28 and TAFII18 Negative cofactor 2 HMFA and HMFB Archeon histones β-sheet motifs Ribbon-helix-helix Arc repressor Met repressorb Mnt repressor Histone-like HU family Histone-like protein (HU) Integration host factor (IHF)/DNA complex DNA-binding protein TF1 TBP family TATA box–binding protein/DNA complex Ig-like transcription factors p53 tumor supressors Human p53/DNA complex Mouse p53 DNA-binding domain NF-κB p50/p50/DNA complex p52/p52/DNA complex p65/p65/DNA complex p65/p50/DNA complex NFAT NFAT1/Fos/Jun/DNA complex NFATC1/DNA complex TonEBP/DNA complex CBF Human PEBP2(CBFα) runt domain Human CBFα runt domain Human Runx-1(CBFα)/CBFβ/DNA complex Mouse Runx-1(CBFα)/CBFβ/DNA Mouse Runx-1(CBFα)/CBFβ/c-EBPβ/DNA T-domain Xenopus laevis Brachyuri T-domain/DNA complex Human TBX3 T-domain/DNA complex STAT proteins STAT-1/STAT-1/DNA complex STAT3β/STAT3β/DNA complex
Reference
2an2 1mdy 1an4 —
Ferre-D’Amare et al. (1993) Ma et al. (1994) Ferre-D’Amare et al. (1994) Ellenberger et al. (1994)
1hry 1lef 1hme
Werner et al. (1995) Love et al. (1995) Weir et al. (1993)
1hio 1taf 1bh9 1jfi 1b67
Arents et al. (1991) Xie et al. (1996) Birck et al. (1998) Kamada et al. (2001) Decanniere et al. (2000)
1par 1cma 1mnt
Raumann et al. (1994) Somers and Phillips (1992) Burgering et al. (1994)
1hue
Tanaka et al. (1984), Vis et al. (1995) Rice et al. (1996) Jia et al. (1996)
1ihf 1wtu
1ytb/1tgh Kim et al. (1993a,b)
1tsr, 1tup 1hu8
Cho et al. (1994) Zhao et al. (2001)
1nfk, 1svc Ghosh et al. (1995), Muller et al. (1995) 1a3q Cramer et al. (1997) 1ram Chen et al. (1998d) 1vkx Chen et al. (1998a) 1a02 1a66 1imh
Chen et al. (1998b) Zhou et al. (1998) Stroud et al. (2002)
1cmo 1co1 1h9d 1hjb 1io4
Nagata et al. (1999) Berardi et al. (1999) Bravo et al. (2001) Tahirov et al. (2001) Tahirov et al. (2001)
1xbr 1h6f
Muller and Herrmann (1997) Coll et al. (2002)
1bf5 1bg1
Chen et al. (1998c) Becker et al. (1998)
aSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb. For proteins with no entry listed, coodinates
have not been deposited in the Protein Data Bank.
bThis protein is also known as MetJ.
Structural Biology
17.1.101 Current Protocols in Protein Science
Supplement 35
Table 17.1.7
RNA-Binding Proteins
Protein RNP domain U1A splicosome hnRNP C Ribosomal protein S6 T4 regA translational regulator OB-fold B. subtilis major cold-shock protein (CspB) E. coli major cold-shock protein (CspA) Ribosomal protein L14 Ribosomal protein S17 N-terminal domain of lysyl and aspartyl tRNA synthetases
PDB entrya
Reference
1urn 1sxl 1ris 1reg
Oubridge et al. (1994) Wittekind et al. (1992) Lindahl et al. (1994) Kang et al. (1995)
1csp, 1nmg 1mef, 1mjc
Schindelin et al. (1993) Newkirk et al. (1994), Schindelin et al. (1994) Davies et al. (1996) Golden et al. (1993) Cavarelli et al. (1993), Onesti et al. (1995)
1whi 1rip 1asz, 1lyl
KH domain Vigilin
1vih
Castiglone Morelli et al. (1995)
dsRNA-binding domain E. coli RNase III Drosophila staufen protein N-terminal domain of ribosomal protein s5
1stu 1pkp
Kharrat et al. (1995) Bycroft et al. (1995) Ramakrishnan and White (1992)
1aaf 1a6b
South and Summers (1993) Demene et al. (1994)
1rop
Predki et al. (1995)
1mst 1wap 1ttt
Valegard et al. (1994) Antson et al. (1995) Nissen et al. (1995)
1e8o, 1e8s 1jid 2ffh 1dul 1fts
Weichenrieder et al. (2000) Wild et al. (2001) Keenan et al. (1998) Batey et al. (2000) Montoya et al. (1997)
Retroviral CCHC zinc fingers NCp7 complex with dACGCC Ncp10 Four-helix bundle RNA-binding protein E. coli ROP Other RNA-binding proteins MS2 phage coat protein Trp RNA-binding attenuation protein (TRAP) Elongation factor-Tu (EF-Tu) SRP domains SRP9/14 in complex with RNA SRP19 in complex with RNA Ffh M domain and RNA complex SRP receptor NG domain
aSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb.
Overview of Protein Structural and Functional Folds
17.1.102 Supplement 35
Current Protocols in Protein Science
Table 17.1.8
Aminoacyl-tRNA Synthetases
Protein Class I Subclass Ia ArgRS CysRS IleRS LeuRS MetRS ValRS LysRS Ib Subclass Ib GlnRS GluRS Subclass Ic TrpRS TyrRS Class II Subclass IIa GlyRSc HisRS ProRS SerRS ThrRS Subclass IIb AspRS AsnRS LysRS IIb Subclass IIc PheRS AlaRSc GlyRSd
Subunits
PDB entriesa
α α α α α, α2 α α
1bs2, 1f7u, 1f7v, 1iq0 li5, 1li7 1ile, 1ffy, 1jzq, 1jzs, 1qu2, 1qu3 1h3n 1a8h, 1f4l, 1qqt 1gax 1irx
α α
1euq, 1euy, 1exd, 1gsg, 1gtr, 1gts, 1qrs, 1qrt, 1qru, 1qtq 1g59, 1gln
α2 α2
1d2r, 1i6k, 1i6l, 1i6m, 1m83, 1mau, 1maw, 1mb2 1tya, 1tyb, 1tyc, 1tyd, 1h3f, 1jii, 1jij, 1jik, 1jil, 2ts1, 3ts1, 4ts1, 1h3e
α2 α2 α2 α2 α2
1ati, 1b76, 1ggm 1h4v, 1qe0, 1adj, 1ady, 1htt, 1kmm, 1kmn 1hc7, 1h4q, 1h4s, 1h4t 1sry, 1ser, 1ses, 1set 1evk, 1evl, 1fyf, 1kog, 1qf6
α2 α2 α2
1l0w, 1b8a, 1eov, 1eqr, 1g51, 1asy, 1asz, 1coa, 1efw, 1il2 11as, 12as 1bbw, 1bbu, 1e1o, 1e1t, 1e22, 1e24, 1lyl
(αβ)2 α4, α (αβ)2
1ehz, 1pys, 1b70, 1b7y, 1eiy, 1jjc –c –c
aSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb. bThere are both class I and class II LysRS cAs of 2003, structures have not been determined. dGlyRS is found in two subclasses: IIa and IIc.
Table 17.1.9
Carbohydrate-Binding Proteins
Protein Legume lectins Soybean agglutinin Cereal lectins Wheat germ agglutinin C-type Mannose-binding E-selectin S-type S-lectin S-lac lectin β-trefoil lectins Ricin B Amaranthus agglutinin
PDB entrya
Reference
2sba
Dessen et al. (1995)
2wgc
Wright and Jaeger (1993)
2msb 1esl
Weis et al. (1992) Graves et al. (1994)
1slt 1hlc
Liao et al. (1994) Lobsanov et al. (1993)
1aai 1jlx, 1jly
Rutenber and Robertus (1991) Transue et al. (1997)
aSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb.
Structural Biology
17.1.103 Current Protocols in Protein Science
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Table 17.1.10
Calcium-Binding Proteins
PDB entrya Reference
Protein EF-hand motif Calmodulin Chick skeletal muscle troponin C Turkey skeletal muscle troponin C Calbindin Myosin subfragment Scallop myosin Recoverin BM-40 Tr1c fragment of sketetal muscle troponin C Annexins Annexin V
1cdl 1top 5tnc 4icb 1mys 1scm 1rec 1bmo 1trf
Meador et al. (1992) Satyshur et al. (1988) Herzberg and James (1988) Svensson et al. (1992) Rayment et al. (1993) Xie et al. (1994) Flaherty et al. (1993) Hohenester et al. (1996) Findlay et al. (1994)
1ala
Huber et al. (1990a,b), Bewley et al. (1993)
aSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb.
Table 17.1.11
Protein
Nucleotide-Binding Proteins
PDB entrya
Dinucleotide-binding fold Lactate dehydrogenase 8ldh Alcohol dehydrogenase 6adh Carbonyl reductase 1cyd Glyceraldehyde 3-phosphate 1gpd dehydrogenase Glutaminyl tRNA synthetase 1gtr Tyrosyl-tRNA synthetase 1tyb Mononucleotide-binding fold Adenylate kinase 3adk Guanylate kinase 1gky Uridylate kinase 1uky Deoxyribonucleoside kinase 1j90 Estrogen sulfotransferase 1aqu Flavodoxin-like mononucleotide-binding fold Flavodoxin 1fxl CheY chemotactic protein 3chy Acetylhydrolase 1es9 Methionine synthase 1bmt Ribosomal protein S2 1j5e Resolvase-like mononucleotide-binding fold γδ-resolvase 2rsl 5′-3′ exonuclase from Taq DNA 1taq polymerase G-protein mononucleotide-binding fold H-ras p21 121p Elongation factor Tu (EF-Tu) 1ttt Elongation factor G (EF-G) 1efg CDC42 1grn Transducin α 1tnd
Reference Adams et al. (1970) Eklund et al. (1981) Tanaka et al. (1996) Moras et al. (1975) Rould et al. (1991) Irwin et al. (1976) Schulz et al. (1974) Stehle and Schulz (1990) Muller-Dieckmann and Schulz (1994) Johansson et al. (2001) Kakuta et al. (1997) Watenpaugh et al. (1972) Volz and Matsumura (1991) Ho et al. (1997) Drennan et al. (1994) Wimberly et al. (2000) Sanderson et al. (1990) Kim et al. (1995b)
Pai et al. (1990) Nissen et al. (1995) Czworkowski et al. (1994) Nassar et al. (1998) Noel et al. (1993)
aSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb.
Overview of Protein Structural and Functional Folds
17.1.104 Supplement 35
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Table 17.1.12
Replication and Transcription Enzymes
Protein
PDB entrya,b Reference
DNA polymerases Pol I family (A family) Klenow fragment from E. coli
1kln
Beese et al. (1993)
Pol α family (B family) Bacteriophage RB69 DNA polymerase
1ih7, 1ig9
Franklin et al. (2001)
Pol β family (X family) Polymerase β from rat
1bpd
Sawaya et al. (1994)
Lesion bypass polymerase family (Y family) DNA polymerase IV from S. solfataricus
1jx4, 1jxl
Ling et al. (2001)
Reverse transcriptase family HIV-1 reverse transcriptase
2hmi
Jacobo-Molina et al. (1993)
Small RNA polymerases T7 polymerase family Bacteriophage T7 polymerase 4rnp Viral RNA-dependent RNA polymerase family Poliovirus 3D RNA polymerase 1rdr dsRNA phage RNA–dependent RNA polymerase family Bacteriophage Φ6 RNA polymerase 1hhs Eukaryotic poly(A) RNA polymerases Yeast poly(A) RNA polymerase 1fa0 Bovine poly(A) RNA polymerase 1f5a DNA-dependent RNA polymerases (RNAPs) T. aquaticus core RNAP (α2ββ′ω)
1hqm, 1i6v
T. aquaticus holoenzyme RNAP (α2ββ′ωσ70) T. thermophilus holoenzyme RNAP (α2ββ′ωσ70) E. coli RNAP α subunit N-terminal domain E. coli RNAP α subunit C-terminal domain E. coli RNAP σ70 initiation factor T. aquaticus RNAP σA factor
1l9u 1iw7 1bdf 1coo 1sig 1ku2, 1ku3, 1ku7 Saccharomyces cerevisiae RNAP II (10 subunits) 1en0, 1I50 DNA polymerase processivity factors Dimeric processivity factors E. coli β subunit of polymerase III Trimeric processivity factors Proliferating cell nuclear antigen (PCNA) Topoisomereases
Sousa et al. (1993) Hansen et al. (1997) Butcher et al. (2001) Bard et al. (2000) Martin et al. (2000) Zhang et al. (1999), Campbell et al. (2001) Murakami et al. (2002a) Vassylyev et al. (2002) Zhang and Darst (1998) Jeon et al. (1995) Malhotra et al. (1996) Campbell et al. (2002) Cramer et al. (2000, 2001)
2pol
Kong et al. (1992)
1plq
Krishna et al. (1994)
Subtype IA E. coli topoisomerase I 67-kDa fragment
1ecl
Lima et al. (1994)
Subtype IB Human topoisomerase I
1a36
Redinbo et al. (1998), Stewart et al. (1998)
Subtype IIA Yeast topoisomerase II 92-kDa fragment
1bgw
Berger et al. (1996) continued Structural Biology
17.1.105 Current Protocols in Protein Science
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Table 17.1.12
Replication and Transcription Enzymes, continued
Protein
PDB entrya,b Reference
E coli GyrB 43-kDa fragment
1ei1
Wigley et al. (1991), Brino et al. (2000)
1d3y
Nichols et al. (1999)
2m2
Katayanagi et al. (1990), Yang et al. (1990), Katayanagi et al. (1992) Davies et al. (1991) Ariyoshi et al. (1994) Dyda et al. (1994) Rice and Mizuuchi (1995) Davies et al. (2000) Beese et al. (1993)
Subtype IIB M. jannaschii topoisomerase VI-A′ RNAse H-like polynucleotide transferases E. coli RNase H
HIV-I reverse transcriptase RNase H RuvC resolvase HIV-I integrase Mu transposase TN5 transposase Klenow fragment 3′-5′ exonuclease
Table 17.1.13
1hrh 1hjr 1itg 1beo 1f3i 1kln
Proteins Involved in Cytoskeleton and Muscle Movements
Protein
PDB entrya
Reference
Actin fold Actin 70-kDa heat-shock cognate protein Hexokinase Glycerol kinase
1atn 1nga 2yhx 2gla
Kabsch et al. (1990) Flaherty et al. (1990) Anderson et al. (1978) Hurley et al. (1993)
Actin-binding protein Profilin
2btf
Schutt et al. (1993)
Actin depolymerizing proteins Gelsolin segment I
—
McLaughlin et al. (1993)
Actin cross-linking proteins Spectrin
1spc
Yan et al. (1993)
aSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb. For proteins with no entry listed,
coordinates have not been deposited in the Protein Data Bank.
Overview of Protein Structural and Functional Folds
17.1.106 Supplement 35
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Table 17.1.14
Enzymes
Protein α/β Hydrolases Haloalkane dehaologenase Torpedo californica acetylcholinesterase Pseudomonas glumae triacylglycerol lipase Sialidases Influenza A neuraminidase Influenza B neuraminidase Salmonella typhimurium LT2 sialidase TIM barrel fold Chicken muscle triose phosphate isomerase Serine proteases Trypsin-like α-Chymotrypsin Subtilisins Subtilisin BPN′ Serine carboxypeptidases Wheat carboxypeptidase II Herpes-like serine protease Cytomegalovirus protease Cysteine proteases Papain-like Papain Caspases ICE (Caspase-1) Viral trypsin-like Hepatitus A virus 3C proteinase Aspartic proteases Pepsin-like Human rennin Retroviral HIV-1 Aspartyl protease Metalloproteases Zincins Thermolysin Astacin Zinc-dependent exopeptidases Leucine aminopeptidase Bovine carboxypeptidase A Proteasome (Ntn hydrolase fold) Archeon T. acidophilum 20S proteasome Yeast 20S proteasome Bovine 20S proteasome E. coli HslV Proteins in the ubiquitin pathway Ubiquitin c-Cbl-UbcH7 Ubc9-RanGAP1 Siah hMms2-Ubc13 Mms2-Ubc13 Skp1-Skp2 Cul1-Rbx1-Skp1-F-box-Skp2
PDB entrya
Reference
2had 1ack 1tah
Franken et al. (1991) Sussman et al. (1991) Noble et al. (1993)
1nn2 1nsb 1dil
Varghese and Colman (1991) Burmeister et al. (1992) Crennell et al. (1993)
1tim
Banner et al. (1975)
2cha
Birktoft and Blow (1972)
1sbt
Alden et al. (1971)
1wht
Liao and Remington (1990)
1cmv, 1wpo, 1lay
Qiu et al. (1996), Shieh et al. (1996), Tong et al. (1996)
9pap
Kamphuis et al. (1984)
1ice
Wilson et al. (1994)
1hav
Allaire et al. (1994)
2ren
Sielecki et al. (1989)
2hvp
Navia et al. (1989)
4tln 1ast
Holmes and Matthews (1982) Bode et al. (1992)
1lap 5cpa
Burley et al. (1990) Rees et al. (1983)
1pma 1ryp 1iru 1ned
Lowe et al. (1995) Groll et al. (1997) Unno et al. (2002) Bochtler et al. (1997)
1ubq 1fbv 1kps 1k2f 1j74,1j7d 1jat 1fs2,1fqv 1ldd,1ldj,1ldk
Vijay-Kumar et al. (1987) Zheng et al. (2000) Bernier-Villamor et al. (2002) Polekhina et al. (2002) Moraes et al. (2001) VanDemark et al. (2001) Schulman et al. (2000) Zheng et al. (2002)
aSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb.
Structural Biology
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Table 17.1.15
Electron-Transfer Proteins
Protein
PDB entrya
Reference
Cytochrome P450CAM Yeast iso-I-cytochrome c Cytochrome b5 Ferritin
1phc 1ycc 1cyo 1bcf
Raag et al. (1993) Louie and Brayer (1990) Mathews et al. (1972) Frolow et al. (1994)
aSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb.
Table 17.1.16
Globin-Like Proteins
Protein
PDBa entry
Reference
Globins Oxymyoglobin Human deoxyhemoglobin Sea cucumber hemoglobin
1mbd 1hhb 1hlm, 1hlb
Phillips (1980) Fermi et al. (1984) Mitchell et al. (1995)
1cpc
Schirmer et al. (1987)
1col
Parker et al. (1989)
Phycocyanins Cyanobacterial C-phycocyanin Colicins Colicin A
aSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb.
Table 17.1.17
Toxins
Protein Small single-subunit toxins Margatoxin Androctonus australis hector toxin II Snake venom neurotoxin Agelenopsis aperta venom Ribosome-inactivating toxins Ricin A ADP ribosylation toxins Diphtheria toxin Pseudomonas aeruginosa exotoxin A E. coli enterotoxin Pertussis toxin Cholera toxin Superantigen toxins Staphylococcus enterotoxin C2 Staphylococcus enterotoxin B Toxic shock syndrome toxin I Anthrax toxin Leathal factor (LF) Oedema factor (EF)
Overview of Protein Structural and Functional Folds
Edema factor (EF) Protective antigen (PA)
PDB entrya
Reference
1mtx 1ptx 1nxb 1agg
Johnson et al. (1994) Housset et al. (1994) Tsernoglou et al. (1978) Reily et al. (1995)
1rtc
Katzin et al. (1991)
1ddt 1ikq 1ltt 1prt 1chp
Choe et al. (1992) Wedekind et al. (2001) Sixma et al. (1991) Stein et al. (1994) Merritt et al. (1995)
1se2 — 3tss
Swaminathan et al. (1995) Swaminathan et al. (1992) Prasad et al. (1997)
1j7n,1jky Pannifer et al. (2001) 1k8t,1k93,1k9 Drum et al. (2002) 0 1lvc Shen et al. (2002) 1acc Petosa et al. (1997)
aSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb. For proteins with no entry listed,
coodinates have not been deposited in the Protein Data Bank.
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Table 17.1.18
Lipid-Binding Proteins
Protein
PDB entrya
Reference
Lipocalins Pieris brassicae bilin-binding protein Human retinol binding protein Bovine odorant-binding protein Bovine β-lactoglobulin
1bbp 1rbp 1obp 3blg, 2blg
Huber et al. (1987) Cowan et al. (1990) Bianchet et al. (1996) Qin et al. (1998)
Fatty acid-binding protein like Rat intestinal FABP Human muscle FABP Rat liver FABP Rat cellular retinol-binding protein II Human retinoic acid-binding protein II Bovine P2 myelin protein Murine adipocyte lipid-binding protein
2ifb 2hmb 1lfo 1opa, 1opb 1cbs 1pmp 1lid
Sacchettini et al. (1989) Zanotti et al. (1992) Thompson et al. (1997) Winter et al. (1993) Kleywegt et al. (1994) Cowan et al. (1993) Xu et al. (1993)
Serum albumin Human serum albumin
1uor, 1ao6
Human serum albumin liganded Horse serum albumin
1bj5, 1bke —
He and Carter (1992), Sugio et al. (1999) Curry et al. (1998) Ho et al. (1993)
Structural Biology
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Table 17.1.19
Large Multisubunit Proteins
Protein
PDB entrya
Reference
1oel, 1aon 1iok
Braig et al. (1994), Braig et al. (1995), Xu et al. (1997b) Fukami et al. (2001)
1aon 1lep 1hx5 1g31
Xu et al. (1997b) Hunt et al. (1996), Mande et al. (1996) Taneja and Mande (2002) Hunt et al. (1997)
1b7t 2mys 1br2 1lkx
Houdusse et al. (1999) Rayment et al. (1993) Dominguez et al. (1998) Kollmar et al. (2002)
1gp2, 1got
Lambright et al. (1996), Wall et al. (1995) Pai et al. (1990) Scheffzek et al. (1996) Boriack-Sjodin et al. (1998) Vetter et al. (1999) Rittinger et al. (1997a) Prakash et al. (2000) Tesmer et al. (1997) Kimple et al. (2002)
Chaperonin complexes GroEL-like E. coli GroEL P. denitrificans chaperonin-60 GroES-like E. coli GroES M. leprae chaperonin-10 M. tuberculosis chaperonin-10 T4 co-chaperonin gp31 Myosin Scallop myosin subfragment S1 Chicken myosin S1 Chicken myosin motor domain Slime mold myosin motor domain G-proteins and associated regulators G-protein αβγ heterotrimer p21Ras p120GAP Ras-Sos complex Ran RhoA-RhoGAP GBP1 RGS4 GoLoco
1gnr 1wer 1bkd 1k5d,1ibr 1tx4 1f5n 1agr 1kjy
F1 ATPases Bovine mitochondrial α3β3γδε
1cow, 1e79
Rat liver mitochodrial α3β3γ Spinach chloroplast α3β3γε Thermophilic bacillus α3β3 E. coli γ-ε
1mab 1fx0 1sky 1fs0
Abrahams et al. (1994), Gibbons et al. (2000) Bianchet et al. (1998) Groth and Pohl (2001) Shirakihara et al. (1997) Rodgers and Wilce (2000)
F1F0 ATPase Yeast mitochondrial α3β3γδc10
1qo1
Stock et al. (1999)
1pma
Lowe et al. (1995)
1ryp 1fnt
Groll et al. (1997) Whitby et al. (2000)
1iru 1ned 1avo
Unno et al. (2002) Bochtler et al. (1997) Knowlton et al. (1997)
Proteasome Archeon T. acidophilum 20S proteasome Yeast 20S proteasome Yeast 20S proteasome and T. brucei 11S regulator Bovine 20S proteasome E. coli HslV Human Regα 11S regulator
aSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb.
Overview of Protein Structural and Functional Folds
17.1.110 Supplement 35
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Table 17.1.20
Integral Membrane Proteins of Known Structure
Protein Photosynthetic proteins Photosynthetic reaction center Photosystem I Photosystem II Light-harvesting complex Seven-helix proteins Microbial rhodopsins Bacteriorhodopsin
Halorhodopsin Sensory rhodopsin II
PDB entrya
Reference
1prc 1pss 1l9j 1jb0 1fe1 1kzu 1lgh
Deisenhofer et al. (1995) Chirino et al. (1994) Axelrod et al. (2002) Jordan et al. (2001) Zouni et al. (2001) Prince et al. (1997) Koepke et al. (1996)
1c3w 1ap9 2brd 1at9 1e12 1jgj 1h68
Luecke et al. (1999) Pebay-Peyroula et al. (1997) Grigorieff et al. (1996) Kimura et al. (1997) Kolbe et al. (2000) Luecke et al. (2001) Royant et al. (2001)
G protein-coupled receptors Rhodopsin 1f88 Transporters and channels AcrB multidrug efflux transporter 1iwg Aquaporin water channel 1fqy 1ih5 1j4n GlpF glycerol channel 1fx8 Ion channels KcsA potassium channel 1bl8 MthK potassium channel 1lnq MscL mechanosensitive channel 1msl MscS voltage-modulated 1mxm mechanosensitive channel ClC chloride channel 1kpk, 1kpl P-type ATPases Calcium ATPase 1eul 1iwo ATP-binding-cassette transporters MsbA lipid flippase 1jsq 1l7v BtuCD vitamin B12 transporter Respiratory enzymes Fumarate reductase 1l0v 1qla, 1qlb Cytochrome c oxidases 1occ 1ar1 1ehk Cytochrome bc1 complex 1qcr 1bcc 1bgy 1ezv Formate dehydrogenase-N 1kqf, 1kqg Monotopic membrane proteins Cyclooxygenase-1 1prh
Palczewski et al. (2000) Murakami et al. (2002b) Murata et al. (2000) Ren et al. (2001) Sui et al. (2001) Fu et al. (2000) Doyle et al. (1998) Jiang et al. (2002a,b) Chang et al. (1998) Bass et al. (2002) Dutzler et al. (2002) Toyoshima et al. (2000) Toyoshima and Nomura (2002) Chang and Roth (2001) Locher et al. (2002) Iverson et al. (1999) Lancaster et al. (1999) Tsukihara et al. (1996) Ostermeier et al. (1997) Soulimane et al. (2000) Xia et al. (1997) Zhang et al. (1998) Iwata et al. (1998) Hunte et al. (2000) Jormakka et al. (2002) Picot et al. (1994) continued
Structural Biology
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Table 17.1.20
Integral Membrane Proteins of Known Structure, continued
Protein
PDB Entrya
Reference
Cyclooxygenase-2 Squalene-hopene cyclase Fatty acid amine hydrolase Monoamine oxidase B β-barrel membrane proteins Porin OmpF porin PhoE phosphoporin Maltoporin
1cx2 2sqc, 3sqc 1mt5 1gos
Kurumbail et al. (1996) Wendt et al. (1999) Bracey et al. (2002) Binda et al. (2002)
2por 2omf 1pho 1mal 2mpr 1bxw 1qjp 1g90 1qj8 1qd5, 1qd6 1ek9 1fcp, 2fcp 1by3, 1by5 1fep 1kmo, 1kmp 7ahl
Weiss and Schulz (1992) Cowan et al. (1992) Cowan et al. (1992) Schirmer et al. (1995) Meyer et al. (1997) Pautsch and Schulz (1998) Pautsch and Schulz (2000) Arora et al. (2001) Vogt and Schulz (1999) Snijder et al. (1999) Koronakis et al. (2000) Ferguson et al. (1998) Locher et al. (1998) Buchanan et al. (1999) Ferguson et al. (2002) Song et al. (1996)
OmpA OmpX OmpLA TolC FhuA FepA FecA α-hemolysin
aSee Web-Based Structural Bioinformatics and http://www.rcsb.org/pdb.
Overview of Protein Structural and Functional Folds
17.1.112 Supplement 35
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A
Hi
α
Ci – 1
α
Oi Ci–1
φi
Ni
Ci
α Ci
ψi
Oi – 1 Ri
C i+1 N i+1
H i+1
α
Hi
B CA
N
C O
N
C
CA C N O C
O N
CA
antiparallel β sheet
parallel β sheet
D
Figure 17.1.1 (continues on next three pages) (A) Drawing of an L-polypeptide chain using a ball-and-stick model to illustrate torsion angles φ and ψ for residue i. Torsion angle φ defines the angle between the planes specified by atoms Ci-1–Ni–Ciα and Ni–Ciα–Ci, respectively. Torsion angle ψ defines the angle between the plane specified by atoms Ni–Ciα–Ci and Ciα–Ci–Ni+1 respectively. Also shown are both ball-and-stick and ribbon representations of an (B) α-helix and (C) β-sheet. The latter is shown in both anti- and parallel orientations. (D) Illustration of the characteristic right-handed twist of a β-sheet as observed in flavodoxin (PDB entry 1flv). (E) Types I and II tight turns. Examples of commonly observed secondary structure assemblies: (F) four-helix bundle (top and side view; PDB entry 1bcf), (G) β-hairpin structure (PDB entry 1bpi), (H) β-sheet with Greek key topology (topology diagram), (I) jelly-roll motif (PDB entry 1pgs); (J) β-sandwich (PDB entry 4gcr), (K) 16-stranded β-barrel (PDB entry 2por), (L) α/β-barrel (PDB entry 1btm), and (M) seven-bladed β-propeller (PDB entry 1got). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.) Structural Biology
17.1.113 Current Protocols in Protein Science
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E O
2
C
N
ON
3 2
3
C
1 4 1 4
type I
type II
F
b d
a
b c
d a c
G
H
4
1
2
3
Figure 17.1.1 (continued)
Overview of Protein Structural and Functional Folds
17.1.114 Supplement 35
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I
J
8 7
4
5
4
5
2
7 6
1
1 3
8
K
3
6
2
L
Figure 17.1.1 (continued)
Structural Biology
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Figure 17.1.1 (continued)
Overview of Protein Structural and Functional Folds
17.1.116 Supplement 35
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A
A′ F
G
C
A′
C′ C′′ C′ C F G
B
B E D
E A
A D
C′′
V type
B
G
F
B
A
C
C F G
E
A
B E D
D
C1 type
Figure 17.1.2 (continues on next page) Tertiary and secondary structures of immunoglobulin fold. The coordinates used for the ribbon diagrams are taken from the PDB entries (A) 3hfl (V type), (B) 1hnf (C1 type), (C) 1fna (C2 type), (D) 1 + tlk (I type), and (E) 1gof (E type). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
Structural Biology
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C
G G
F C
F
A
E
C E
C′
C′ C F G
A
A B E
C′ B
B
C2 type
C2(Fn)
C2
D
A′ C F
B E D
G A
I type
E C G
F
A
A′ C′
C′ C F G
E
B E A
B
E type Figure 17.1.2 (continued)
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Figure 17.1.3 MHC structures. (A) Class I MHC HLA-A2 complex (PDB entry 2clr) and (B) class II MHC complex HLA-DR1 (PDB entry 1dlh), each with an antigenic peptide. The α and β chains in (B) are colored blue and green respectively. The peptide is shown as a ball-and-stick model. (C) Close-up view of the α1α2 peptide-binding domain of HLA-A2 (PDB entry 2clr). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
Structural Biology
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Figure 17.1.4 Protein folds in the complement system. (A) C3d (PDB entry 1c3d; residues 996 to 1287). Left side shows the view down the barrel axis; the right side shows the side view of the barrel. The α helices are numbered 1 to 12 and the N-terminal 310 helix is labeled T1. The residues critical for covalent attachment to the pathogen surface, His 133, Gln 20, and Ala 17 (Cys residue in the wild type protein), are represented by a ball-and-stick model. (B) C5a (PDB entry 1kjs).The α helices are numbered from 1 to 5 (C) Complement factor D serine protease (PDB entry 1dsu). Catalytic residues His 57, Asp 102, and Ser 195 are represented by a ball-and-stick model. (D) Complement regulatory protein CD59 (PDB entry 1cdq; residues 1 to 70). The β strands are numbered according to their order in the protein sequence. (E) CCP modules 15 and 16 from complement factor H (PDB entry 1hfh). The β strands for each separate CCP module are numbered according to their order in the protein sequence. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
Overview of Protein Structural and Functional Folds
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Figure 17.1.5 Long and short-chain cytokines. The structure of (A) a long-chain helical cytokine, GCSF (PDB entry 1rhg), (B) a short-chain helical cytokine, IL-4 (PDB entry 1rcb), and (C) interferon-γ (PDB entry 1rfb). The two monomers of interferon-γ are shown in red and purple. (D) Connectivity between helices in four-helix bundle cytokines. The up helices are drawn in white and the down helices are in black. (E) Connectivity between helices of IFN-γ. The block shaded regions correspond to the four-helix bundles. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
Structural Biology
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Figure 17.1.6 Cytokines and chemokines. (A) Human IL-1β, a member of the β-trefoil fold. (PDB entry 1hib). (B) Human transforming growth factor-β2 (PDB entry 2tgi). The four strands that define the cysteine-knot fold are shown in green (Anderson et al., 1978). The six knotted cysteines are shown in ball-and-stick model (with yellow sulfur atoms). (C) Murine EGF (PDB entry 1epj). (D) CXC chemokine (PDB entry 1il8), (E) CC chemokine (PDB entry 1rto). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.7 (continues on next page) The structures of (A) tumor necrosis factor receptor TNFR (PDB entry 1tnr), (B) type II TGF-β receptor (PDB entry 1ktz), (C) thyroid hormone receptor (PDB entry 1bsx), (D) integrin I domain (PDB entry 1lfa), (E) scavenger receptor (PDB entry 1by2), (F) glutamate receptor (PDB entry 1gr2) bound to the neurotoxin kainate (ball-and-stick model), and (G) transferrin receptor (PDB entry 1cx8). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.7 (continued)
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Figure 17.1.8 Folds that bind phosphopeptides. The phosphopeptides are shown as magenta colored worms. The phosphorylated amino acid is represented by a ball-and-stick model. (A) The SH2 domain from v-src tyrosine kinase bound to a five-residue phosphotyrosine peptide (PDB entry 1sha). (B) PTB domain from shc complexed with a twelve-residue phosphotyrosine peptide (PDB entry 1shc). (C) FHA domain from protein kinase RAD53 complexed to a twelve-residue phosphothreonine peptide (PDB entry 1g6g). (D) Homodimer of 14-3-3 protein ζ bound to eight-residue phosphoserine peptides (PDB entry 1qja). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.9 Folds that bind to polyproline peptides. Bound polyproline peptides are represented by ball-and-stick models. (A) An SH3 domain from the Abl tyrosine kinase complexed with the ten-residue synthetic peptide 3Bp-1 (PDB entry 1abo). (B) A WW domain from dystrophin in complex with a seven-residue β-dystroglycan peptide (PDB entry 1eg4). (C) An EVH1 domain from Enabled, bound to the Acta peptide (PDB entry 1evh). (D) GYF domain from CD2Bp2 (PDB entry 1gyf). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.10 Phospholipid-binding domains. Lipid ligands are displayed as ball-and-stick models and metal cations are represented by magenta spheres. (A) Pleckstrin homology (PH) domain from Dappl/Phish complexed with inositol 1,3,4,5-tetrakisphosphate (PDB entry 1fao). (B) C1 domain from protein kinase Cδ complexed with phorbol-13-acetate (PDB entry 1ptr). (C) C2 domain from protein kinase C(α) complexed with Ca2+ and phosphatidylserine (PDB entry 1dsy). (D) A FYVE domain from EEA1 bound to inositol 1,3-diphosphate (PDB entry 1joc). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.11 (continues on next page) Protein interaction domains. Bound peptides are represented by magenta worms. (A) The syntrophin PDZ domain bound to the peptide GVKESLV (PDB entry 2pdz). (B) VHS domain of GGA1 complexed with cation-independent mannose-6-phosphate receptor C-terminal peptide (PDB entry 1jwg). (C) SNARE fusion complex containing syntaxin-1A (green), synaptobrevin-II (red), and SNAP-25B (blue; PDB entry 1sfc). (D) SAM domain from human EPHB2 receptor (PDB entry 1b4f). (E) MH2 domain from human Smad2 (PDB entry 1khx). (F) Homotrimer of a phosphorylated MH2 domain from human Smad2 (PDB entry 1khx). Phosphorylated Ser465 and -467 are represented by ball-and-stick models. The phosphoserine binding loop L3 is shown in magenta. (G) Complex of the Smad2 MH2 domain (cyan) with the SARA complex (magenta; PDB entry 1dev). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.11 (continued)
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Figure 17.1.12 (continues on next page) Structural repeat motifs. (A) A single Leu-rich repeat (left) from ribonuclease inhibitor (right; PDB entry 2bnh). (B) A HEAT repeat (left) from the PR65/A subunit of protein phosphatase 2A (right; PDB entry 1b3u). (C) An ARM repeat (left) from β-catenin (right; PDB entry 2bct). (D) An ankyrin repeat (left) from GABPβ (right; PDB entry 1awc). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.12 (continued)
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Figure 17.1.13 (continues on next page) Protein kinase and phosphatase structures. (A) The structure of phosphorylated cyclic AMP-dependent protein kinase complexed with ATP, Mn, and inhibitor (PDB entry 1atp). The peptide inhibitor is represented by a magenta worm. The ATP, Mn, and phosphorylated serine and threonine are shown as ball-and-stick models. (B) The structure of the inactive form of hematopoietic cell kinase of the Src family of protein kinases (PDB entry 1ad5). The SH3 and SH2 domains are colored blue-green and magenta, respectively, and the catalytic domain is colored green and red. The phosphorylated tyrosine 572 is represented by a blue ball-and-stick model. The following protein phosphatases are all shown in approximately the same orientation with the catalytic cysteine displayed as a ball-and-stick model. (C) Low-molecular-weight protein tyrosine phosphatase (PDB entry 1phr). (D) High-molecular-weight protein tyrosine phosphatase-1B (PDB entry 2hnq). (E) Dual-specificity protein phosphatase CDC25A (PDB entry 1c25). (F) Protein serine/threonine phosphatase-1 of the PPP family (PDB entry 1fjm). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.14 Phosphatase, kinase, and related structures. (A) Alkaline phosphatase (PDB entry 1ali). (B) Fructose-1,6bisphosphatase (PDB entry 5fbp) as a representative of the sugar phosphatase fold. (C) Cyclin A (PDB entry 1fin). The N-terminal cyclin box domain is colored magenta. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.15 Variations of the helix-turn-helix (HTH) structural motif and the MADS box. The recognition helix is oriented horizontally in all but panel A. (A) Homodimer of the DNA-binding domain of λ repressor bound to DNA (PDB entry 1lmb). The three helices of the HTH motif are labeled α1, α2, and α3 in each monomer. Both recognition helices (α3) sit in the major groove. (B) λ repressor (prokaryotic HTH; PDB entry 1lmb). (C) Engrailed homeodomain (PDB entry 1hdd). (D) Globular domain of histone H5 (PDB entry 1hst). This example of the winged helix motif has only one wing (the β hairpin). (E) Purine repressor (purR; PDB entry 1pru). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.16 Zinc-binding motifs within DNA-binding domains. The zinc atom and side-chain ligands to the zinc are represented by ball-and-stick models. (A) Second zinc finger from ZIF268 mouse intermediate protein (PDB entry 1zaa). (B) Elongation factor TFIIS (PDB entry 1tfi) (C) Zn2Cys6 binuclear cluster from GAL4 (PDB entry 1d66). (D) GATA-1 chicken erythroid transcription factor (PDB entry 1gat). (E) DNA-binding module from the glucocorticoid receptor (PDB entry 1gdc). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.17 (continues on next page) Binding of zinc-containing modules to DNA. The zinc atoms are represented by spheres. (A) The three zinc fingers of ZIF268 (PDB entry 1zaa). (B) GATA-1 transcription factor (PDB entry 1gat). Zn2Cys6 binuclear clusters (C) Gal4 (PDB entry 1d66) and (D) pyrimidine pathway regulator 1 (PPR1) DNA-binding fragment (PDB entry 1pyi) bound to DNA. (E) Glucocorticoid receptor complexed with DNA (PDB entry 1glu). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.17 (continued)
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Figure 17.1.18 (continues on next page) Helical DNA binding domains (A) MADS box of serum response factor bound to DNA (PDB entry 1srs). (B) Basic-region leucine-zipper (bZIP) c-Fos/c-Jun heterodimer complexed with DNA (PDB entry 1fos). Monomers within the dimer are shaded differently. (C) Basic helix-loop-helix (bHLH) MyoD homodimeric transcription activator complexed with DNA (PDB entry 1mdy). Monomers within the dimer are colored differently. (D) High-mobility group (HMG) fragment B from rat (PDB entry 1hme). (E) Structure of a heterodimer of dTAF42 and dTAF62 (PDB entry 1taf). Each monomer contains the histone fold. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.18 (continued)
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Figure 17.1.19 (continues on next two pages) β-sheet DNA-binding motifs. (A) Arc-repressor tetramer complexed with DNA (PDB entry 1par). (B) TATA-box-binding protein (TBP) complexed with DNA (PDB entry 1ytb). (C) Histone-like HU protein (PDB entry 1hue). (D) HU-like IHF complexed with DNA (PDB entry 1ihf). (E) The p53 tumor-suppressor monomer bound to DNA (PDB entry 1tsr). Regions involved in DNA binding are labeled and colored red. The zinc atoms and side-chain ligands to the zinc are represented by ball-and-stick models. (F) Structure of the p50/p50 homodimer complexed with DNA (PDB entry 1svc). The two insertion regions (magenta) may play a role in binding other transcription factors. (G) Tetrameric complex of NFAT1 (magenta), Fos/Jun (AP-1), and DNA (PDB entry 1a02). (H) Ternary complex of CBFα (green), CBFβ (magenta), and DNA (blue) (PDB entry 1h9d). (I) Brachyuri T-domain homodimer bound to DNA (PDB entry 1h6f). (J) STAT-1 homodimer bound to DNA (PDB entry 1bf5). The coiled-coil domain, DNA-binding domain, linker domain, and SH2 domain are colored blue, red, green, and magenta, respectively. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.19 (continued)
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Figure 17.1.19 (continued)
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Figure 17.1.20 RNP domains. β strands containing the RNP1 or -2 consensus sequences are labeled (arrows). (A) U1A splicosomal protein (PDB entry 1urn). (B) Ribosomal protein S6 (PDB entry 1ris). (C) Bacteriophage T4 regA protein (PDB entry 1reg). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.21 The OB fold. (A) Cold-shock protein A (PDB entry 1mjc). β strands containing RNP1 or -2 consensus sequences are colored purple. (B) Cold-shock protein B (PDB entry 1csp). β strands containing RNP1 or -2 consensus sequences are labeled. (C) Ribosomal protein S17 (PDB entry 1rip). (D) Ribosomal protein L14 (PDB entry 1whi). (E) Anticodon-binding domain of aspartyl-tRNA synthetase (PDB entry 1asz). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.22 Double-stranded RNA-binding domain (dsRBD) and KH domain. (A) Drosophila staufen protein (PDB entry 1stu), an example of dsRBD fold. (B) N-terminal domain of ribosomal protein S5 (PDB entry 1pkp). (C) Human vigilin (PDB entry 1vih), an example of a KH domain. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.23 (A) The RNA-binding protein Rop from E. coli. (PDB entry 1rop). Helices 1 and 1′, postulated to bind to RNA, are labeled. (B) Hexamer of the MS2 phage-coat protein (PDB entry 1mst). Two AB dimers and a CC dimer are arranged around a quasi-3-fold rotation axis located in the center of the figure (labeled q3). (C) Eleven-subunit oligomer of Trp RNA-binding attenuation protein (TRAP; PDB entry 1wap). Bound tryptophan molecules are shown by a ball-and-stick model. Alternating monomers are shaded differently for clarity. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.24 (continues on next page) Aminoacyl tRNA transferase catalytic domains (A to B) and signal recognition particle (SRP) domains (C to E). (A) Example of a class I aaRS catalytic domain (GluRS, PDB entry 1gln). The MSK and HIGH motifs are highlighted by green and red respectively. An insertion domain (common in class I catalytic domains) is colored magenta. (B) Example of a class II catalytic domain (HisRS, PDB entry 1htt). Motifs 1 to 3 are colored green, red, and magenta, respectively. (C) Structure of SRP9/14 heterodimer of SRP Alu domain bound to RNA (red coil). SRP9 and -14 are colored in green and cyan, respectively (PDB entry 1e8o). (D) Structure of SRP19 in complex with RNA (cyan; PDB entry 1jid). (E) Structure of ffh-M domain (green) in complex with RNA (red; PDB entry 1dul). (F) Structure of a SRP receptor (PDB entry 1fts). The N domain and I box are both colored magenta. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.24 (continued)
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Figure 17.1.25 Lectin folds. (A) Legume lectin soybean agglutinin (PDB entry 1sba). The β strands are labeled A through M. The bound carbohydrate molecules are represented by ball-and-stick models. (B) Wheat-germ agglutinin monomer with bound carbohydrate. Disulfide bonds are shown as yellow sticks. (PDB entry 2wgc). (C) The C-type lectin mannose-binding protein A (PDB entry 2msb) with bound carbohydrate. Calcium ions are shown as magenta spheres. (D) S-lectin with bound carbohydrate (PDB entry 1slt). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.26 Calcium-binding folds. Calcium ions are drawn as magenta spheres. (A) A pair of calcium-binding EF-hands (red and orange respectively) complexed with Ca2+ from bovine calbindin D9K. The corresponding helices are labeled E, F, E′, and F′ (PDB entry 4icb). (B) Calcium-binding domain of annexin V (PDB entry 1ala). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.27 The classical dinucleotide- and mononucleotide-binding folds shown by ribbon drawings and topology diagrams. In the topology diagrams, the β-sheets are viewed from the C-terminal edge with β-strands represented by green triangles and α-helices by red circles. Numbers indicate strand and helix order within the structure. Circles containing an “X” represent possible domain insertions. Bound nucleotides are colored purple. (A) Ribbon drawing of the NAD-binding domain of dogfish lactate dehydrogenase with bound NAD represented as a ball-and-stick model (PDB entry 1ldm). (B) Topology diagram for the common core of the classical dinucleotide-binding fold. (C) Ribbon drawing of adenylate kinase as an example of a mononucleotide-binding protein (PDB entry 1aky). The portion of the structure that is not part of the nucleotide-binding fold is colored purple. The bound inhibitor bis(adenosine)-5′-pentaphosphate (AP5) is shown using a ball-and-stick model. (D) Topology diagram for the common core of the classical mononucleotide-binding fold. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.28 (continues on next two pages) Structures of DNA and RNA polymerases. (A) The 39-kDa catalytic domain of rat DNA polymerase β (PDB entry: 1bpd). (B) The Klenow fragment of E. coli DNA polymerase I bound to an 11-bp duplex DNA in an editing complex (PDB entry: 1kln). The DNA is represented by a yellow and purple phosphate backbone trace. (C) The conserved palm subdomain from Klenow fragment (PDB entry 1kln). (D) The palm subdomain from rat DNA polymerase β (PDB entry 1bpd). (E) Structure of the HIV-1 reverse transcriptase heterodimer complexed with a 19/18-base duplex DNA (PDB entry 1hmi). The p66 subunit is colored by subdomain and the p51 subunit is dark blue. The bound DNA is represented by a yellow and purple phosphate backbone trace. (F) Thermus thermophilus RNA polymerase holoenzyme (PDB entry 1iw7). The β′, β, α′, α′′, ω, and σ70 subunits are labeled and colored green, cyan, red, tan, magenta, and dark blue, respectively. (G) The N-terminal domain of the RNAP α subunit from E. coli (PDB entry 1bdf). (H) RNAP II from yeast (PDB entry 1i50). Visible subunits are labeled and the five subunits in common with bacterial RNAP follow the same coloring scheme as in (F). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.28 (continued)
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Figure 17.1.28 (continued)
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Figure 17.1.29 Structure of the β-subunit of pol III (PDB entry: 2pol), an example of a sliding-clamp DNA polymerase processivity factor. (A) A single domain of the β-subunit. (B) View of the entire ring structure of the homodimer looking down the six-fold symmetry axis. The two monomers are colored red and green, respectively. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.30 (continues on next two pages) Topoisomerase structures. (A) Structure of the N-terminal 67 kDa fragment of E. coli topoisomerase I (PDB entry 1ecl). The four domains are labeled and the side chain of the active-site Tyr319 is represented by a ball-and-stick model. (B) Complex of human topoisomerase IB with 22-bp DNA duplex (PDB entry 1a36). Each domain is color coded and labeled. DNA is a light-blue ball-and-stick model. (C) Same as (B), but rotated 90° around the vertical axis. (D) Structure of the 92-kDa fragment of yeast topoisomerase IIA showing the organization of domains. (PDB entry 1bgw). (E) Homodimeric form of the 92-kDa fragment of yeast topoisomerase IIA. For clarity, the two monomers are colored red and green, respectively. (F) Homodimer of the 43-kDa fragment of E. coli GyrB (PDB entry 1ei1). One monomer is blue-green, the other is red-orange. ADPNP bound to each monomer is represented by a ball-and-stick model. (G) Homodimer of Methanococcus jannaschii topoisomerase VI subunit A (PDB entry 1d3y). One monomer is colored according to domain and subdomain, the other is gray (program color “white”). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.30 (continued)
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Figure 17.1.30 (continued)
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Figure 17.1.31 The polynucleotidyl transferase RNase H-like family of folds. (A) Structure of RNase H from E. coli (PDB entry 2rn2) and (B) Corresponding secondary structure topology. (C) The secondary structural topology of HIV integrase. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.32 (continues on next page) Structures containing the actin fold (A to D) and structures that bind to actin (E to G). (A) Actin complexed with ATP (PDB entry 1atn). Domains 1 and 2 are colored purple and blue, respectively. Subdomains 1 to 4 are labeled. ATP is shown as a ball-and-stick model. (B) The N-terminal fragment of heat-shock cognate protein (hsc70) complexed with ADP (PDB entry 1nga). ADP is shown as a ball-and-stick model. (C) Hexokinase B in its open form (PDB entry 2yhx). Structures of (D) glycerol kinase (PDB entry 1gla), (E) profilin (PDB entry 2btf), and (F) severin (NMR structure, PDB entry 1svq). (G) The spectrin repeat is shown as a dimer (PDB entry 1spc), with the two monomers colored red and blue, respectively. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.32 (continued)
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Figure 17.1.33 (continues on next page) (A) Structure of acetylcholinesterase from Torpedo Californica. The central eight-stranded β-sheet is shown in green. (PDB entry 1ack). (B) Diagram of the secondary structure topology common to all α/β hydrolases. The strands are numbered from one to eight and helices are labeled A through F. (C) Structure of an influenza virus neuraminidase complexed with an inhibitor (PDB entry 1nnc). The active site-bound inhibitor is shown as ball-andstick model. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.33 (continued)
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Figure 17.1.34 (continues on next page) The TIM-barrel (A to C) and serine protease fold (D to F). The side chains of residues in the active site catalytic triad are shown as ball-and-stick models in panels (D) and (F). (A) Side view of a ribbon drawing of triose phosphate isomerase as an example of a TIM-barrel. (B) Ribbon drawing of triose phosphate isomerase viewed from the top. (C) Secondary structure schematic of the classical TIM-barrel fold. β-strands are represented by green arrows and α-helices by red rectangles. (D) Structure of the trypsin-like serine protease, collagenase (PDB entry 1hyl). (E) Structure of the subtilisn serine protease, subtilisn BPN′ (PDB entry 1sup). (F) The structure of the serine carboxypeptidase, wheat serine carboxypeptidase II (PDB entry 1wht). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.34 (continued)
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Figure 17.1.35 (continues on next two pages) Examples of cysteine proteases (A and B), aspartic proteases (C and D), and metalloproteases (E to G). The side chains of critical active site residues mentioned in the text are shown as ball-and-stick models. Zinc atoms are shown as magenta balls. (A)The papain-like cysteine protease, human cathepsin B (PDB entry: 1huc). (B) Interleukin-1b converting enzyme (ICE; PDB entry: 1ice). (C) Pepsin-like protease, human renin (PDB entry 1bbs). (D) The retroviral protease, HIV-1 protease (PDB 1hpx). (E) Structure of a zinc-dependent endopeptidase, a metzincin, snake venom adamalysin II (PDB entry 1iag). (F) Structure of a zinc-dependent exopeptidase, aminopeptidase from Aeromonas proteolytica (PDB entry 1amp). (G) Structure of alkaline protease, a metzincin from Pseudomonas aeruginosa (PDB entry 1akl). The active site zinc and coordinated side chains are shown as ball-and-stick models in the N-terminal catalytic domain (left side). Bound calcium ions are shown as black balls in the C-terminal parallel β-roll domain (right side). (H) A catalytic β-subunit from the 20S yeast proteasome as a representative of the Ntn fold (PDB entry 1ryp). The nucleophilic threonine is shown as a ball-and-stick model. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.35 (continued)
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Figure 17.1.35 (continued)
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Figure 17.1.36 (continues on next page) Enzymes of the ubiquitin pathway. (A) The structure of ubiquitin (PDB entry 1ubq). (B) Complex between the ubiquitin-conjugating E3 enzyme c-Cbl (cyan) and the E2 enzyme UbcH7 (green; PDB code 1fbv). The c-Cbl RING domain is colored magenta with Zn shown as yellow balls and the c-Cbl-bound ZAP-70 peptide is shown in red. The active site Cys86 of UbcH7 is shown as a yellow ball-and-stick model. (C) SUMO E2 enzyme Ubc9 with active site Cys93 shown as a yellow ball-and-stick model (PDB entry 1kps). (D) Siah homodimer (PDB entry 1k2f). The monomers are colored blue and green, respectively. The RING domains are in orange. Zinc atoms are magenta spheres. (E) Cul1-Rbx1-Skp1-F-box-Skp2 complex. The Cul1, Ring-Box, SKP1, and Skp2-F-box are colored in cyan, red, orange, and magenta, respectively (PDB code 1ldk). Zn ions are shown as yellow spheres. (F) The second cullin repeat from cul1 (PDB entry 1ldk). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.36 (continued)
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Figure 17.1.37 (continues on next page) Structures showing the fold of (A) cytochrome P450-CAM (PDB entry 1phc), (B) cytochrome c (PDB entry 1ycc), (C) cytochrome c3 (PDB entry 2cy3) with the four heme molecules colored differently, (D) cytochrome b5 (PDB entry 1cyo), (E) cytochrome b562 (PDB entry 1cgn), and (F) a dimer of bacterioferritin (also known as cytochrome b1; PDB entry 1bcf). The heme molecules are shown by ball-and-stick models. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.37 (continued)
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Figure 17.1.38 Structures of globin-like proteins from each of the three globin-like families. All three proteins are shown in the same orientation with respect to the globin fold. Helices that are not considered part of the core globin fold are colored green. (A) Sperm whale myoglobin (PDB entry 1mbd). Helices are labeled A to H according to traditional globin nomenclature. Helices A, E, and F are colored red, while helices B, G, and H are tan. The heme group is represented by a ball-and-stick model. (B) Structure of human deoxyhemoglobin showing all four subunits (PDB entry 1hhb). The heme groups are represented by ball-and-stick models. The α subunits are colored red and the β subunits are colored yellow. (C) C-phycocyanin from cyanobacteria (PDB entry 1cpc). The phycocyanobilin cofactor is represented by a ball-and-stick model. (D) Colicin A from E. coli (PDB entry 1col). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.39 (continues on next page) Tertiary folds of toxins. (A) ADP ribosylation domain of diphtheria toxin (PDB entry 1ddt). (B) Pentameric B domain of bacteria AB5 cholera toxin with each domain colored differently (PDB entry 1chp). (C) Superantigen toxin (PDB entry 1se2). (D) Ricin A chain (PDB entry 1rtc) The structure of (E) scorpion toxin (PDB entry 1mtx), (F) snake toxin (PDB entry 1nxb), and (G) spider toxin (PDB entry 1eit). Disulfide bonds are represented by yellow stick models. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.39 (continued)
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Figure 17.1.40 Structure of anthrax toxin. (A) Domains I through IV of the lethal factor (LF) are colored in blue, green, yellow, and orange, respectively (PDB entry 1jky). The bound MAPKK peptide is shown in red. This PDB entry does not include coordinates for the Zn atom. (B) The structure ofedema factor (EF; PDB entry 1k8t). The catalytic domains CA and CB are shown in blue and cyan, respectively. The helical domain and the switch A and C regions are shown in green, yellow, and magenta, respectively. The location of switch B is shown in the next panel. (C) CaM complexededema factor (PDB entry 1k90). The coloring scheme is the same as (B) with CaM in red and switch B in orange. (D) Domains I to IV of the protective antigen (PA) are shown in blue, green, yellow, and red, respectively (PDB entry 1acc). Ca2+ ions are shown as magenta spheres. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.41 Lipid-binding proteins with respective ligands shown as ball-and-stick models. (A) Human retinol-binding protein bound to retinol (PDB entry 1rbp), an example of a lipocalin. (B) Rat intestinal fatty acid-binding protein bound to palmitate (PDB entry 2ifb). (C) Human serum albumin with five bound myristate molecules (PDB entry 1bj5). Note that a helix spans both the I/II and II/III domain boundaries. This results in 28 instead of 30 helices in the structure. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.42 (continues on next page) Chaperonin and myosin structures. (A) The GroEL/GroES complex viewed from the side (left) and top (right; PDB entry 1aon). The GroEL trans ring is green, the cis ring is blue, and the GroES ring is magenta. The domain structure of GroEL in the (B) trans and (C) cis rings of the GroEL/GroES complex (PDB entry 1aon). The equatorial domains are blue, the intermediate domains are green, and the apical domains are red. ADP is shown as a ball-and-stick model. (D) The GroES subunit (PDB entry 1aon). (E) The structure of the scallop myosin subfragment S1 (PDB entry 1b7t). Bound Ca2+ is shown as a small purple sphere. The heavy-chain domain is shown in green. The essential and regulatory light chains are colored blue and red, respectively. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.42 (continued)
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Figure 17.1.43 (continues on next two pages) G-protein and its regulators. The bound Mg2+ and GDP are shown as a magenta sphere and ball-and-stick model. (A) Structure of p21Ras (PDB entry 1gnr). A nonhydrolyzable GTP analog is shown as a ball-and-stick model. (B) The structure of the Ras GTPase activation domain of a human p120GAP (PDB entry 1wer) . (C) The structure of Ras in complex with the Ras guanine-nucleotide-exchange-factor region of Sos (PDB entry 1bkd). Ras and Sos are colored in green and blue, respectively. (D) The structure of RhoA (green) in complex with RhoGAP (blue; PDB entry 1tx4). GDP and AIF4 are shown as ball-and-stick models. (E) The structure of GBP1 (PDB entry 1f5n). (F) Side and (G) top view of the structure of a heterotrimeric G-protein complex (PDB entry 1got). The Gtα-Giα chimera is in red, Gτβ is green, and Gτγ subunit is blue. (H) The structure of RGS4 (blue) in complex with Giα-GDP-AlF4− (green; PDB entry 1agr). (I) The crystal structure of a Gαi-GDP (green and yellow) bound to the GoLoco region of RGS14 (blue; PDB entry 1kjy). GDP is shown as a ball-and-stick model. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.43 (continued)
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Figure 17.1.43 (continued)
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Figure 17.1.44 Structure of the F1ATPase. (A) Top view of bovine F1ATPase α (yellow), β (red), and γ (magenta, N- and C-terminal helices only) subunits (PDB entry 1e76). (B) Side view of bovine mitochondrial F1ATPase showing one α subunit (left), one β subunit (right), and the γ (magenta, center), δ (cyan, bottom), and ε (green, bottom) subunits. The α and β subunits are each color coded by domain: N-terminal β-barrel is cyan, the central nucleotide-binding domain is green, and the C-terminal α-helical bundle is red. (C) Cα model of ten c subunits from a yeast F0ATPase membrane domain (PDB entry 1qo1). Each subunit is a long α-helical hairpin. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.45 The 20S proteasome from yeast complexed with the 11S regulator from T. brucei (PDB entry 1fnt). The α, β, and regulator subunits are colored red, green, and blue, respectively. (A) Side view of the barrel-shaped complex. (B) View of the complex looking down the axis of the barrel. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.46 Viral capsid proteins. (A) Illustration of an icosahedron showing two-, three-, and five-fold symmetry. (B) Structure of the jelly-roll β-sandwich fold of satellite tobacco necrosis virus-coat protein (PDB entry 2stv). (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.47 (continues on next two pages) Structures of integral membrane proteins. (A) Photosynthetic reaction center (PDB entry 1prc). The four protein subunits are shown: cytochrome (gray), M (green), L (red), and H (yellow). (B) The structure of Halobacterium salinarum bacteriorhodopsin (PDB entry 1c3w), with the cytoplasmic face at the top. The retinal chromophore is shown as a blue ball-and-stick model. (C) Rhodopsin from the outer segment of bovine rod photoreceptor cells (PDB entry 1f88). The receptor is shown with the C-terminal (cytoplasmic) domain at the top, and the N-terminal (extracellular) domain at the bottom of the diagram. The covalently linked eleven-cis-retinal ligand is shown as a blue ball-and-stick model. The additional eighth helix is colored in yellow. (D) The ClC chloride channel from Salmonella typhimurium (PDB entry 1kpl). The double barrel of the homodimer is shown with Cl− ions, visible in the pores, displayed as green spheres. View is perpendicular to the plane of the membrane with the two subunits colored red and blue. (E) The E. coli vitamin B12 transporter BtuCD. The dimer is depicted, with BtuC subunits in red and BtuD subunits in blue (PDB entry 1l7v). (F) Calcium ATPase from skeletal-muscle sarcoplasmic reticulum in the E1 state with bound calcium ions displayed as orange spheres. The three cytoplasmic domains are at the top (PDB entry 1eul). (G) Bovine AQP1 water channel monomer (PDB entry 1j4n). View is from the plane of the membrane with the extracellular face at the top. The two membrane-inserted helices that do not span the membrane are colored in blue. (H) Bovine cytochrome bc1complex (PDB entry 1bgy). The subunits with membrane-spanning portions are colored in the following scheme. Cytochrome b is red, cytochrome c1 is aqua, Rieske iron-sulfur protein is blue, subunit 7 is orange, subunit 10 is green, and subunit 11 is yellow. (I) The E. coli ferric enterobactin receptor FepA (PDB entry 1fep). The β-barrel domain is shown in green and the N-terminal plug domain is shown in red. (J) TolC outer membrane protein of E. coli (PDB entry 1ek9). The trimer is shown with the subunits colored separately. (For full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.)
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Figure 17.1.47 (continued)
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Overview of Macromolecular Electron Microscopy: An Essential Tool in Protein Structural Analysis Although the majority of biological studies employing the electron microscope (EM) aim to visualize the substructure of cells and tissues or the spatial distributions of specific protein antigens in them, EM also has a long and distinguished record in the study of isolated protein molecules. Today, its potentialities for incisive observations of proteins and interactions between them are unprecedentedly high. These exciting prospects stem from innovations in specimen-preparation techniques, image-recording procedures, and importantly, advances in digital image processing (Baumeister and Zeitler, 1992; Frank, 1996). Of particular note, EM lends itself well to the examination of proteins or protein complexes that are refractory to other experimental approaches—e.g., proteins that are exceptionally large, flexible, filamentous or otherwise asymmetric, membrane-bound, or irremediably heterodisperse. EM is an underutilized resource for the study of proteins in general, and one goal of this unit is to illustrate why EM is a powerful tool at the protein scientist’s disposal. A second objective is to illustrate the potential of EM for structural analysis of large, multicomponent protein complexes. The significance of this area is heightened by the consideration that many, if not most, important biological processes are carried out by such complexes, not by simple enzymes that communicate only via random diffusion of their reaction products. DNA replication, transcription, RNA splicing, protein folding, protein synthesis, regulatory proteolysis, membrane trafficking, and signal transduction represent a few examples of such processes. EM is complementary to—not in competition with—other methods for studying the structures of proteins. In that context, this unit will try to identify situations in which EM may offer the best, if not the only, prospect of progress—as well as situations where a combined approach in which EM is integrated cohesively with other techniques may be advantageous. Although a few atomic models of proteins have now been worked out by EM (see discussion of High-Resolution Studies) and more are on the way, it seems inevitable that the majority of new high-resolution structures will continue to
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emerge from X-ray crystallographic and NMR spectroscopic studies. One of the major opportunities that EM offers is in providing a means to capitalize on this hard-won information on the structures of individual subunits. In other words, EM has the potential to visualize how such subunits interact in higher-order, functional complexes. The goals of this chapter are to give a general overview of current EM procedures for observing proteins, to outline the kind of information that may be acquired, and to illustrate their potential with some specific examples. The references represent an eclectic mix of reviews, early influential papers on given topics, accounts of methodology, and examples of recent or notably successful applications.
ELECTRON MICROSCOPY AND DIGITAL IMAGE PROCESSING: A PERFECT PARTNERSHIP Electron micrographs of proteins tend to be noisy images, even when recorded with the most advanced instruments. Moreover, they provide only two-dimensional representations, usually projections, of specimens whose threedimensional structures are the objects of primary interest. Both of these drawbacks have been largely eliminated by the emergence of digital image processing techniques specifically designed for analyzing micrographs of biological specimens (Misell, 1978; Frank, 1996). These techniques clarify and quantify the information recorded in the original micrographs. For instance, one may average many noisy images to obtain a better defined and more readily interpretable image of a protein, synthesize multiple views into a three-dimensional density map, or apply a variety of visualization tools to explore these maps and averaged two-dimensional images. In consequence, the scope of EM for studies of proteins has been immensely enhanced. In contrast to X-ray crystallography, in which most practitioners tend to use the same highly developed and professionally supported programs, EM-related image-processing software is still at an earlier stage of development, and research in this domain is actively pursued in many laboratories. Nevertheless, the proStructural Biology
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grams already available—in many cases at little or no expense—amount to an extensive body of well-tested and user-friendly software, capable of a wide range of operations (Carragher and Smith, 1996).
IN THE EM, PROTEINS ARE VISUALIZED ONE MOLECULE AT A TIME
Overview of Macromolecular Electron Microscopy
The primary attribute of the EM as an analytical tool is that its images relate to individual molecules, not vast ensembles. This is the case whether the molecules are ordered as in X-ray crystallography or dispersed as in other biophysical techniques such as ultracentrifugation or NMR spectroscopy. The physical basis of this capability is the charge of the electron, which makes it possible to accelerate electrons from an emitter, collimate them, and focus the resulting beam through electromagnetic lenses (Misell, 1978; Spence, 1980). A second important consequence of the electron’s charge is that the beam interacts with the specimen strongly enough for a single protein molecule to yield a discernible, albeit noisy, image. However, the strength of this interaction is a two-edged sword. To avoid degradation of the coherence and monochromaticity of the electron beam—and hence of its imaging qualities—by interactions with gas molecules, the EM column must be maintained under a high vacuum. Until recently, this vacuum has required that specimens be dehydrated before entering the EM column. Preservation of native protein conformations despite complete dehydration poses a severe, possibly unattainable, technical challenge. To date this problem has not been solved, and the various procedures whereby dehydration is accomplished in practice—which are useful and productive, at least to limited resolution—represent damage-control measures. However, with the recent innovation of cryo-EM (see discussion of Frozen-Hydrated Specimens), specimens remain fully hydrated, immobilized in vitreous ice (Stewart and Vigers, 1986; Dubochet et al., 1988; Booy, 1993; Chiu, 1993). Accordingly, this approach has unique potential for visualizing native protein structures in that it circumvents the fundamental problem of dehydration-induced denaturation. Another obstacle posed by the electron charge, particularly for studies that aspire to higher resolutions, is radiation damage. The energy imparted to a protein molecule in a single inelastic scattering event is sufficient to rupture many bonds (Zeitler, 1990), so molecu-
lar structures degrade rapidly during observation. However, some workable solutions have been devised. For specimens prepared by negative staining or metal shadowing (see discussion of Specimen Preparation), the electrons that dominate the image are scattered not from the protein molecules themselves but from metallic replicas. Although not immune to the effects of electron irradiation, such replicas are much more resistant than unstained proteins. Nevertheless, even with unstained specimens, the adverse effects of electron irradiation may be minimized by employing imaging strategies in which the electron dose is minimized—i.e., virtually all of the electrons that pass through the protein are used to form its image (Williams and Fisher, 1970; Unwin and Henderson, 1975). Typically, a promising area of the specimen is identified at very low magnification (e.g., 2000×) at a negligibly low electron dose. Next, focusing, stigmation, and other initial adjustments are performed on a neighboring area. Finally the beam is switched on to the targeted area just long enough to record the micrograph. Hence, the observer shoots blindly, but not randomly. It also helps to ensure that (nearly) every electron counts by developing the negatives under conditions that maximize the detection efficiency of the film, or by using sensitive digital recorders—e.g., a charge-coupled device (CCD) camera (Brink and Chiu, 1994). By operating the microscope at relatively low magnifications (30,000× to 50,000×), the total electron dose needed to achieve the required minimum darkening of the negative is correspondingly reduced (by the inversesquare law). However, it becomes counterproductive to reduce the magnification much further because of the limit set by the finite grain size of the recording medium (Downing and Grano, 1982), and because the lenses tend to work less well at lower magnifications—i.e., they are more susceptible to external perturbations.
THE NICHE OF EM IN PROTEIN STRUCTURE DETERMINATION In general, the larger and more asymmetric a complex, the more difficult it is to solve its structure by X-ray crystallography. NMR spectroscopy, also capable of high resolution, is restricted in applicability to proteins up to about 25 to 30 kDa. In the past few years, EM has graduated into the elite class of experimental techniques that yield atomic models of proteins (see discussion of High-Resolution Studies).
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To date, the specimens that have been imaged in such detail have been highly ordered crystalline monolayers of intrinsic membrane proteins embedded in lipid bilayers, and, consequently, in their natural environment. However, even at lower resolution, many questions concerning the structures and interactions of proteins may be addressed readily and rewardingly by EM, and less easily if at all by other techniques. Some of these applications are outlined below: in several cases, a more detailed account follows later in the unit.
Assessing Purity and Determining Stoichiometry EM provides a rapid and informative assay with respect to the purity and monodispersity of a preparation, particularly if the protein in question has a high molecular weight and/or is oligomeric. It is also economical: 10 ng of protein in a 2-µl droplet of buffer may suffice for an observation by negative staining (see discussion of Negative Staining). Electron micrographs are also a direct source of information concerning the shape, dimensions, symmetry, and architecture of protein molecules. The r otational symmetry and thence the stoichiometry of ring-like oligomers may often be determined by simple appraisal of micrographs. In ambiguous cases—e.g., “donuts” (see Fig. 17.2.1A and C)—the order of symmetry may be determined with greater confidence and objectivity by applying symmetry-detection algorithms (Crowther and Amos, 1971; Kocsis et al., 1995) or classification procedures (Dube et al., 1993) to sets of digitized images. If preliminary observations prove promising, the specimen may be suitable for a systematic analysis of its three-dimensional structure (see discussion of Architecture and Interactions of Protein Complexes). Moreover, if the shapes of two or more interacting components have been determined individually, EM of the complex may serve to define their docking geometry (see Fig. 17.2.1C).
Molecular-Weight Measurements of Individual Proteins Scanning transmission electron microscopy (STEM) of unstained proteins provides a direct and unambiguous method of determining their molecular weights (Wall and Hainfeld, 1986; Müller et al., 1992; Thomas et al., 1994). This method is based on the principle that the number of electrons scattered by a molecule should be directly proportional to its molecular weight. In STEM imaging, this condition is met. The
electron beam is focused down to a tiny probe only a few angstrom units (Å) across, to scan across the specimen. For each point sampled by this probe, the scattered electrons are collected in an annular detector situated behind the specimen and recorded as a digital signal. This imaging system provides an efficient dark-field mode (most electrons scattered are recorded), whose signal is linear over a wide range. As such, it is ideally suited for molecular-mass measurements. These measurements are effected by integrating the image density associated with an object of interest in a digital micrograph, subject to an appropriate background subtraction to discount scattering from the support film. For this paradigm to hold, the protein should be unstained; also, the imaging mode should be linear (as it is, to a good approximation, in dark-field). Although these conditions cease to hold for very thick specimens (e.g., >500 Å) or when comparing molecules of quite different atomic compositions, the method works remarkably well for most proteins. Structural preservation of the unstained molecules is enhanced by freeze-drying (Wall, 1979). Such preservation is not an absolute requirement for mass measurement of the whole molecule, but it may allow multiple domains to be visualized and consequently their masses to be determined on an individual basis (e.g., Cerritelli et al., 1996; Fig. 17.2.2). To date, most EM-based mass analyses have been performed in the few laboratories that are equipped with a dedicated STEM, as distinct from a conventional TEM (transmission electron microscope) adapted with a STEM imaging mode. Notwithstanding their small number, the few biological STEMs in operation have been exceptionally productive. In particular, the STEMs operated at Brookhaven National Laboratory are accessible to user-defined projects (see discussion of Resources and User Facilities). Recent developments in energy-filtering electron microscopy promise to make EM-based molecular mass determinations more widely available in individual laboratories (Feja et al., 1997). Although the remarkable progress of mass spectrometry has provided another analytical tool capable of operating in the elusive-highmolecular-weight range (see Chapter 16 for mass spectrometry techniques useful in this range), STEM retains some unique capabilities in this domain. For example, it is particularly well suited for large, asymmetric molecules; for the analysis of preparations that are het-
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erodisperse but in which the molecules of interest are recognizable; and for complexes such as nucleoproteins in which spatial information pertaining to the positions of protein-binding sites relative to the ends of a nucleic acid molecule provides a valuable complement to mass data (Thomas et al., 1994).
Architecture and Interactions of Protein Complexes Higher-order structures constitute the class of proteins for which computer-enhanced EM holds the greatest potential. These complexes are typically composed of many subunits, and their masses range from several hundreds kilodaltons to many megadaltons. Their subunits may be arranged on a symmetric lattice—as in a viral capsid (see Fig. 17.2.3)—or they may be without symmetry, as in a ribosome. In either case, image averaging, applied in one form or another (Steven et al., 1991), is the foundation on which detailed structural analysis is based. Initially, such averaging was effected by Fourier filtration of regular two-dimensional arrays and thus was applicable only to this small class of specimens. Since then, the methodology has gradually generalized and diversified to the point where it may now be applied to virtually any kind of protein molecule, whether freestanding or polymerized (Frank, 1996; also see Fig. 17.2.4 for an illustrative analysis of freestanding molecules). Although a detailed account of this methodology goes beyond the scope of this article, the principles are easily outlined. Analysis starts with collection of a suitably large set of molecular images (two-dimensional projections) of the highest possible quality. The required size of this data set depends on the dimensions of the molecule in question, the resolution aspired to, and whether analysis in two or three dimensions is envisaged. Typically, the data set
ranges from a hundred or so to several thousand images (Henderson, 1995). For a two-dimensional analysis, the images are mutually aligned and sorted into intrinsically similar classes, and these are independently averaged. Structural conclusions are then drawn from these averaged images (see, e.g., Fig. 17.2.1). For a full three-dimensional analysis, it is necessary to determine which view of the molecule is given by each image before a density map can be calculated. There are several strategies that may be followed. If a plausible, even if crude, three-dimensional model is available a priori, it may be projected computationally in all possible directions to generate a set of reference images that may be used to sort the data in a “supervised classification” (Penczek et al., 1994; Baker and Cheng, 1996). A more detailed model is then calculated from the sorted data, and the process iterated until no further improvement is obtained. Clearly, the end product of such a calculation has to be insensitive to the details of the starting model if it is to truly reflect the information content of the data and not just replicate the initial assumptions. If an appropriate starting model is not available, several strategies may be pursued to obtain one. For instance, internal symmetries may be exploited (Crowther, 1971) to determine the orientations of enough particles to calculate a first-generation model. More generally, the mutual orientations of averaged two-dimensional projections may be estimated by matching their common one-dimensional projections (a “common-lines” approach to “angular reconstitution”; van Heel, 1987). Alternatively, data from tilting experiments may be used to obtain a model—from either tilt pairs, as in the “random conical tilt” approach (Radermacher, 1988), or a full tilt series in the case of tomographic reconstruction (Dierksen et al., 1995).
Figure 17.2.1 (at left) Components of the energy-dependent protease, ClpAP, visualized in negative stain. The bar at the top of panel B is equal to 500 Å. Each panel shows the molecule as originally imaged (large field) and as enhanced by averaging, at higher magnification (inserts in bottom right of each panel). (A) ClpP (21-kDa monomer), the protease, is a ring-shaped molecule, ∼115 Å in diameter. ClpP orients preferentially on EM grids to present its axial view, whereby the 7-fold rotational symmetry is revealed by rotational-symmetry analysis. ClpP consists of two 7-fold rings stacked to form a round, hollow complex that looks annular in projection from any direction. (B) ClpA (81-kDa monomer), the ATP-hydrolyzing, chaperonin-like component, forms hexamers in the presence of nucleotide which present two preferred views. The rarer top view discloses ClpA’s 6-fold symmetry. The more prevalent side view presents two parallel striations, corresponding to the two tiers of domains (the subunit consists of two ATP-binding domains of approximately equal size). (C) Functional complexes of ClpP with ClpA are bipolar. They contain two molecules of ClpA, seen here in side view, one on either side of a ClpP molecule, also in side view. Adapted from Kessel et al. (1995) courtesy of Dr. M. Kessel.
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Figure 17.2.2 Dark-field STEM microscopy of bacteriophage T4 tail fibers, unstained and prepared by freeze-drying. Panels A, B, and C show micrographs alongside histograms of mass measurements made from data; the bar at the bottom of panel C is equal to 500 Å. (A) Proximal half-fibers; (B) Distal half-fibers; (C) Whole long tail-fibers. (D) Averaged images of the long tail fiber give consistent accounts of its domainal structure: top, unstained, freeze-dried; bottom, negatively stained with vanadate. (E) Molecular model of a short segment of the tail-fiber, as a three-stranded β helix. Adapted from Cerritelli et al. (1996) courtesy of Dr. M.E. Cerritelli. Overview of Macromolecular Electron Microscopy
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Figure 17.2.3 Cryo–electron micrographs of capsids of bacteriophage HK97. Original micrographs are shown in the top row of each panel (with the bar in this section of panel C equal to 200 Å on the scale of those micrographs); the outside surfaces of reconstructions at ∼25-Å resolution are shown in the middle row; and their inner surfaces in the bottom row (with the bar in the bottom row of panel C equal to 200 Å on the scale of the reconstructions). (A) Prohead I, the earliest precursor, is composed of 420 copies of the 42-kDa capsid protein. (B) Prohead II, produced by proteolytic removal of 101 residues from the N terminus, which resides on the inner surface of Prohead I. (C) Head II is the end product of a large-scale conformational change in Prohead II that produces Head I, which then becomes covalently cross-linked together to yield Head II. Adapted from Conway et al. (1995) courtesy of Dr. J.F. Conway.
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Figure 17.2.4 Electron micrography and reconstructions of calcium release channel. (A) Cryo– electron micrograph of calcium release channel. The bar is equal to 1000 Å. (B) Stereo pairs of a three-dimensional model of this tetrameric molecule (monomer mass = 170 kDa), as reconstructed to 25-Å resolution from cryo–electron micrographs by the random conical tilt method (Radermacher et al., 1994). The molecule is portrayed by surface rendering, as viewed along three mutually orthogonal axes. The bar is equal to 100 Å. Adapted from Radermacher et al. (1994) courtesy of Dr. M. Radermacher.
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As noted above, a rich variety of reconstruction strategies has been developed. However, one key condition applies: the raw images must contain enough information to permit reliable determination of the view to which each micrograph corresponds. Tomography is an exception to this rule because the relative orientations of the various tilts are specified by the observer. Thus, the larger and more asymmetric a complex, the more marked is the differentiation between its various views, and the more precisely one may hope to distinguish among them. The other side of this coin is that if the specimen size is doubled, one must then halve the uncertainty in its orientation angles to achieve the same resolution. In this context, small globular proteins will look essentially the same from any direction, even with micrographs of the highest possible quality, and thus will remain difficult if not impossible to tackle by single-particle methods. Nevertheless, such molecules are quite amenable to study by EM if organized into regular arrays (two-dimensional crystals or helices) in which the subunits are held in well-defined orientations (Jap et al., 1992).
Membrane Proteins To facilitate the handling of intrinsic membrane proteins, such molecules are often solubilized in detergents, or else studied in genetically truncated form with their major hydrophobic elements excised or modified. For expediency, some investigations focus on expressing single domains or even peptide fragments for study, a strategy that is by no means confined to membrane proteins. While such provisions may provide material that is more amenable to in vitro biochemistry, it is worth bearing in mind that these fragments may no longer be membrane proteins and that isolated fragments may fail to adopt the conformations that they assume in the intact protein. Moreover, detergent-solubilized proteins may be significantly altered in structure by binding the surfactant. In this context the opportunity to visualize the intact proteins, even at limited resolution, is of considerable value. For instance, cryo-EM of lipoprotein vesicles and of natural or reconstituted membranes provides unique opportunities to visualize membrane proteins in their lipid-embedded environment (Brisson and Unwin, 1984).
Protein Filaments Proteins that polymerize into helices are readily studied by EM. Although the first three-
dimensional studies of helical polymers were performed almost 30 years ago (DeRosier and Klug, 1968), the methodology has continued to advance in applicability and resolving power (Stewart, 1988). Proteins that have been studied by this approach include cytoskeletal filaments as well as motility elements—either intracellular (actomyosin or microtubule-kinesin) or extracellular (e.g., flagellae; see discussion of Attainable Goals, below). Other examples include microbial adhesins (e.g., fimbriae; Heck et al., 1996) and nucleic acid–binding proteins such as the genetic recombination factor, recA (Stasiak and Egelman, 1994). In addition to naturally occurring and functionally active filaments, there are many cases on record of proteins polymerizing adventitiously into ordered tubes or filaments in vitro. Examples of proteins that have exhibited such behavior include the acetylcholine receptor (Brisson and Unwin, 1984) and the rev transactivator protein of HIV (Wingfield et al., 1991). In such cases, the ordered packing of molecules may be of dubious biological significance, but their helical symmetry is convenient for structural analysis in that they hold the subunits in well-defined orientations—helical filaments are one-dimensional crystals. A tentative generalization concerning proteins that have a tendency or a functional requirement to polymerize into filaments is that they are refractory to forming well-ordered three-dimensional crystals. Under conditions that cause such proteins to precipitate, the interactions that lead to filament formation may be expected to assert themselves. Thus, filament formation is likely to preempt three-dimensional crystallization, except in the rare cases in which the helical symmetry allows filaments to pack together to generate a threedimensional crystal of some repeating unit. More generally, disordered lateral packing of filaments into tactoids or paracrystals will occur. This obstacle to crystal formation is circumvented in the case of filament-forming proteins that bind a ligand masking their self-interacting surface, thus providing a binary complex that may be amenable to co-crystallization. In the event that a high-resolution structure of the subunit can be obtained, either directly (e.g., for the gonococcal pilin; Parge et al., 1995) or from cocrystals (e.g., of actin complexed with DNase I; Kabsch et al., 1990), three-dimensional EM may be a source of information on the intersubunit interactions that govern their packing in the filament (Holmes et al., 1993; Aebi et al., 1996).
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Fibrous Oligomers
Overview of Macromolecular Electron Microscopy
Many proteins consist of one or more highmolecular-weight subunits that have extended, often flexible, conformations. Such molecules are particularly prevalent in “connectivity” interactions in such settings as the extracellular matrix, receptor recognition, and cytoskeletonassociated phenomena. To a first approximation, they may be considered as one-dimensional proteins, since their folds often employ one or more of the standard fibrous-protein motifs—coiled-coil α helices or collagen-like 310 helices (Fraser and MacRae, 1973) or the recently discovered β helix (Yoder et al., 1993)—or else they consist of multiple globular domains connected by fibrous linkers. The former two motifs may be readily detected by analyzing the protein’s amino acid sequence— i.e., coiled-coil α helices consist of multiple tandem copies of canonical heptads (Cohen and Parry, 1990) and collagen-like sequences tend to have glycine in every third position. Algorithms have been developed to detect the presence of heptads in a sequence (Lupas et al., 1991) and reasonable predictions may be made with respect to the packing of the α helices in coiled coils (i.e., the number of strands and their mutual orientation) on the basis of empirical rules concerning the occupancy of key sites in the heptads (Woolfson and Alber, 1995). For oligomeric fibrous proteins, EM can serve as an important source of structural information that complements sequence-based predictions. The first order of business is to acquire an image that is sufficiently detailed to divulge the molecule’s domainal structure, which may then be correlated with inferences based on the primary sequence, as noted above. The information required for such a correlation—i.e., the number of distinct domains, their lengths and widths (in the case of rod domains), and their sizes and spacing (for globular domains)—may be inferred from averaged electron micrographs in the moderate-resolution range of 20 to 30 Å. For such analyses, fibrous proteins have the advantage that they are almost always present in a definite view—the side view. Direct imaging can be very effective with such proteins (e.g., myosin; Walker et al., 1985) and image averaging adds a valuable refinement. Intrinsic flexibility may result in variations from particle to particle that pose a problem for averaging, but this may be rectified by computational straightening (Egelman, 1986; Steven et al., 1991). By combining EM with sequence-based predictions as outlined above, it may be possi-
ble to develop a model for the overall path of the polypeptide chains through the molecule. A number of fibrous proteins have been analyzed in this way, including the tail fibers of bacteriophages T7 (Steven et al., 1988) and T4 (Cerritelli et al., 1996; see Fig. 17.2.2) and the filamentous hemagglutinin of Bordetella pertussis (Makhov et al., 1994). This form of analysis appears to have considerable potential for the study of other fibrous proteins. An alternative approach to the study of large filamentous proteins is via high-resolution studies of domains by crystallography (Strelkov et al., 1996) or NMR spectroscopy (Baron et al., 1991; Pascual et al., 1996). As a prelude to such studies, the domain of interest must be identified, expressed in bulk, and purified. The decision as to what to express is, of course, pivotal; the number of candidates is large and the number for which in-depth study is practical is small. EM has the potential to reveal the full domainal organization of such proteins. EM may also help clarify whether an expressed domain of a fibrous protein has folded correctly.
Visualizing Conformational Changes and Switches Conformational changes control the assembly and activation of many protein complexes. The scale of these structural revisions varies widely. At one extreme, subtle local changes may suffice for a substantial alteration in the affinity of a binding site. At the other extreme, movements of structural elements over distances of 25 Å or more (Jontes et al., 1995) and extensive refolding of polypeptide chains (e.g., in bacteriophage T4 capsid maturation; Black et al., 1994) accompany some of the more dramatic conformational changes on record. Other examples of this phenomenon include activation of reovirus to a transcriptionally competent state (Dryden et al., 1993), herpesvirus capsid maturation (Trus et al., 1996), and chaperonin switching upon binding nucleotides (Roseman et al., 1996). EM provides a powerful tool to study conformational transitions, particularly those in which large-scale structural changes take place (see Fig. 17.2.3). The suitability of EM for this application is enhanced by the consideration that differential changes may be detected with considerably greater sensitivity than the nominal resolution of the analysis implies (see discussion of Immuno-EM). A further consideration is that the distinct states in a functional cycle or sequence may vary considerably in stability, with some
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states being relatively short-lived. Instability is not conducive to the success of crystallography in that the proteins may well switch into a lower-energy state prior to or during crystallization. However, EM requires only that a conformation be sustained long enough to allow observation. By judicious use of rapid freezing techniques to capture short-lived states, the time resolution achievable in cryoEM studies is now of the order of a few milliseconds (Berriman and Unwin, 1994; Walker et al., 1995).
CONCEPTS AND MISCONCEPTIONS To some, electron microscopy has the dubious reputation of having little to offer for the study of protein structure. According to this point of view, first of all, it is prone to artifacts— i.e., “you can see anything you want.” Second, too small a fraction of molecules in a bulk population are observed for the results to be statistically meaningful. Third, the resolution is too low to yield worthwhile information.
Artifacts and How to Avoid Them In principle, artifacts can arise from two major sources. The first relates to failure of the observer to identify the molecule of interest correctly thus basing conclusions on some other component present on the grid. With the second, the molecule of interest is correctly identified, but the acts of specimen preparation and observation (e.g., radiation damage) have caused changes, random and/or systematic, in its native structure that are sufficiently large to seriously mislead the observer. In practice, the risk of both kinds of artifacts may be largely eliminated by observing a few straightforward precautions. One essential measure is to avoid contamination of the EM grid. It should be standard operating procedure to use double-distilled water and chemicals of the highest available purity in making specimen buffers and gridwashing solutions. It should be recalled that sterility is not the same thing as cleanliness. The specimen substrate, usually a thin carbon film, should be clean; glow-discharging or casting the film immediately before use can be helpful provisions. The risk of cross-contamination from tweezers used to prepare grids of other specimens should be borne in mind. Cellulose fibers from filter paper used to blot grids should not be mistaken for protein filaments, and vice versa. One distinguishing criterion is that protein filaments often exhibit distinctive sub-
structure such as striations or serrations in the size range typical of individual subunits or domains (20 to 50 Å) that are not seen on cellulose fibers.
The Cardinal Importance of Reproducibility When examining a new protein by EM, the first priority is to establish conditions under which the appearance of the particles is fully reproducible. It is also desirable to optimize their distribution on the grid by systematically varying the sample concentration and other experimental conditions (such as grid-washing procedure). Optimal concentrations tend to vary from protein to protein, but typically, 10 to 50 µg/ml may be appropriate for specimens adsorbed to a carbon film for negative staining or nebulized onto a mica surface for shadowing; 0.5 to 2.5 mg/ml may be appropriate when preparing thin films of frozen suspensions for cryo-EM (see discussion of Specimen Preparation Techniques for more information on these procedures). Once an adequate distribution of particles has been achieved, it should be determined if they are uniform in size and appearance and if there are any indications of substructure. For example, featureless globular particles of randomly varying size seen in a negative-staining experiment should be viewed with suspicion. They may simply represent an emulsion of grease droplets or bubbles somehow formed in the stain layer. If the observed particles show distinctive substructure but fall into several size classes, there are several testable possibilities. Particles that are oligomeric in solution may be breaking down when prepared for EM. This hypothesis may be tested by cross-linking the preparation with glutaraldehyde, which should suppress this tendency to break down, or by repeating the procedure using cryo-EM, in which specimens are not subjected to such extreme forces. Alternatively, the specimen may be impure or inhomogeneous. On occasion, protein preparations that seem to be quite pure according to SDS-PAGE (UNITS 10.1 & 10.4) may look considerably less pure by EM. For instance, there may be contaminants that contribute only faint gel bands, but cumulatively amount to a significant fraction of the total molecular population. Also, some of these intruders may have higher binding affinities for the support film than the majority protein of interest and thus be overrepresented in micrographs. A further important reproducibility check, particularly if doubt persists, is consistency of
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appearance of the same specimens when prepared for EM in quite different ways (e.g., staining and shadowing or cryo-EM). Finally, in identifying the molecule of interest in micrographs, it is important to take into consideration all other available information. If the protein has been purified by gel-filtration chromatography (UNIT 8.3) or sucrose-gradient centrifugation, an estimate of its molecular weight should be available. Within the limited scope of such a calculation, that molecular weight should be consistent with a particle of the size seen in the EM images. A typical value for the partial specific volume of protein should be compared with the estimated particle volume. Assuming that the particle is correctly identified, it should be determined whether the observed features make sense in the light of what else is known.
and whose dimensions or spacings in solution are known, such as the plate-like crystals of catalase (Wrigley, 1968) or tobacco mosaic virus (Wall, 1979) with its distinctive 23-Å axial repeat. A second problem is that proteins may spread when they are dried down on a support film, thus exaggerating their size, or they may shrink upon drying and appear smaller. The advent of cryo-EM has greatly reduced uncertainties associated with the state of preservation of the specimen. Thus, it is usually possible to determine the dimensions of protein molecules to within a few percent for freestanding specimens; the main source of uncertainty arises from imprecise edge detection with images of limited resolution. With periodic specimens, edge detection is less of an issue because spacings may be determined from diffraction patterns to within ≤1%, and in favorable cases to even higher precision.
Variability in Appearance If the particles seen in electron micrographs are approximately uniform in size but distinctively variable in appearance, several explanations are possible. It may be that the protein of interest assumes several different orientations on the EM grid, thus presenting distinct views. If that is so, this property may be exploited to yield a three-dimensional representation of the protein by synthesizing the various two-dimensional projections (Frank, 1996). One check is to perform tilting experiments: suitable rotation of the specimen in the EM may convert one view into another. Alternatively, the preparation may contain two or more similarly sized proteins. To evaluate this possibility, it is essential to make a careful review of the biochemical evidence pertaining to purity. In cases of ambiguity, specific antibodies to the protein of interest, if available, can be used to advantage. The question here is: do the antibodies label all particles or only a specific subset?
Measuring the Dimensions of Protein Molecules and Polymers
Overview of Macromolecular Electron Microscopy
In principle, systematic errors of several kinds may affect size measurements made by EM. In practice, awareness of such effects makes it possible to minimize their impact. For instance, the magnification may diverge by up to 5% or so from its nominal value because of lens hysteresis and the state of alignment of the microscope column. This limitation may be overcome by employing an internal calibration—i.e., a specimen that is easily recognized
Reproducibility Again: Statistically Meaningful Results This section concludes with a reiteration of the importance of reproducibility. The sample should be reproducible from one biochemical preparation to another and should yield consistent results in independent EM experiments. For studies involving the use of image processing, the reproducibility of the end-product images should be assessed in quantitative terms. Finally, consider statistical representativity. Suppose that a sample consists of 0.1 ml of a solution containing a 250-kDa protein at 0.5 mg/ml—a total of ∼0.3 × 1014 molecules. If an EM study is based on, say, 3000 mutually consistent images, these molecules amount only to an infinitesimal fraction (∼10−10) of the total population. Can they be taken as statistically representative? Consider the following proposition: suppose the population consists of two kinds of microscopic particles: the protein of interest (fraction p), and contaminants (fraction 1−p). Then, in an unbiased sampling experiment of 3000 trials, the probability of picking out only contaminants is (1−p)3000—an extremely small number for any reasonable value of p. Of course, this proposition is idealized, particularly with respect to the assumed lack of bias in the trial, which in practice may be affected by selective adsorption of contaminants to the EM grid and subjectivity on the part of the observer. It does, however, make a point with respect to the issue of sample sizes and statistical representativity.
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RESOLUTION Instrumental Factors The resolution achieved in an EM study of proteins is constrained by many factors. Among these, it is possible to distinguish between instrumental factors and specimen-dependent factors. In purely optical terms, the wavelength of the electrons is not a resolution-limiting factor, as it is in light optics. At the commonly used accelerating voltage of 80 kV, the wavelength of the electrons is ∼0.04 Å. Nor, in principle, is the outcome of most experiments limited by the intrinsic resolving power of the microscope. Commercially available instruments routinely attain resolutions of 2.5 to 3.0 Å when correctly aligned and operated close to focus and with the lenses fully excited—i.e., at very high magnification (for example, 400,000×)—for imaging of radiation-resistant inorganic specimens. Structural information at this resolution should allow the tracing of polypeptide chains. Why is this resolving power not generally realized with biological material? The answer is, at best, complex and incomplete. Many factors are involved. One major consideration is the need to minimize radiation damage when imaging biological macromolecules. In consequence, the EM is operated at lower magnifications, where it tends to perform less well— e.g., at these magnifications EM performance is more susceptible to impairment by stray magnetic fields. Another consideration relates to the size of proteins and the complexity of their substructure, and the large numbers of precisely defined views that are needed to determine their three-dimensional structures to high resolution (see discussion of Quantitative Resolution Criteria in Two and Three Dimensions). The instrumental resolution limit of a given micrograph may be assessed from its diffraction pattern (obtained optically or by digital Fourier transform; see Fig. 17.2.5C). The spacing of the outer limit of intensity in this pattern represents the maximum resolution that the image is potentially capable of yielding.
Specimen-Dependent Factors Because of specimen-related factors, the resolution limit of biologically meaningful information is usually considerably lower than the instrumentation-imposed limit. One such specimen-related factor is the possibility of random distortions in native structure induced by specimen preparation. Another is radiation
damage. “Shot noise” in images that were recorded at very low dose to reduce radiation damage and in consequence, are statistically ill-defined, as well as metal-grain size (if shadowing or staining are used) are additional specimen-related constraints on the resolution limit. Since resolution may be constrained by so many factors, any of which may be limiting in a given case, a pragmatic measure is to define the effective resolution of an EM experiment in terms of the limit to which structure is consistently represented over a substantial population of molecules (see Quantitative Resolution in Two and Three Dimensions).
Resolution and Signal-to-Noise Ratio As noted above, electron micrographs of proteins are noisy images. The sources of noise are many (Frank, 1996), and a full accounting will not be attempted here. In general, however, the finer a detail, the more likely it is to be obscured by noise. This problem may be alleviated by image averaging. In its simplest form, this involves visual appraisal of an array of like images and identifying common features. On a more systematic basis, averaging is performed by digital summation of many individual images, appropriately aligned. Each image is assumed to consist of a signal overlaid with additive noise; if it is random, the noise is suppressed by averaging; if it is consistent, the signal is reinforced. In this context, the resolution criteria outlined below apply only to averaged images. The same information is, of course, latently present in individual images; the problem is that it cannot be inferred unambiguously without image averaging.
Quantitative Resolution Criteria in Two and Three Dimensions For periodic specimens—e.g., crystals or helices—the resolution limit may readily be calculated in terms of the spatial frequency of the outermost periodic reflections that are visible above background in the diffraction patterns (e.g., Fig. 17.2.5B). For EM data (images, not electron-diffraction patterns) this limit is a conservative one and may be an underestimate because of lattice disorders. The adverse effects of disorder, which result in smearing of the averaged image with a consequent reduction in resolution, may be mitigated by computational correction of the lattice (van Heel and Frank, 1981; Steven et al., 1991). This procedure may yield a significant improvement in the resolution of the averaged image (see Fig. 17.2.5). Structural Biology
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For nonperiodic specimens, corresponding measures of resolution have been expressed in terms of mathematically defined criteria applicable to sets of digitized images (each representing a distinct copy of the same protein). Several such criteria are widely used—e.g., the differential phase residual (Frank et al., 1981), the Fourier ring correlation coefficient (Saxton and Baumeister, 1982), and the spectral signalto-noise ratio (Unser et al., 1987). The numbers that these criteria give for the resolution of a given data set tend to vary slightly because different thresholds are used to define the limit of resolution. Although such variations are small, studies seeking incremental improve-
A
Overview of Macromolecular Electron Microscopy
ments in resolution on a given specimen should apply the same criterion consistently. Similarly, when comparing several studies of the same protein, investigators should consider whether any small differences in reported resolution may not simply reflect the choice of resolution criterion. The resolution of two-dimensional images imposes an upper bound on the resolution of the three-dimensional density map that may be calculated from them. In three dimensions, however, an additional resolution-limiting factor comes into play—i.e., the question of how fully and how uniformly the underlying set of two-dimensional views covers three-dimen-
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Figure 17.2.5 (A) Negatively stained polyhead (tubular capsid variant; 750-Å wide) of bacteriophage T4. (B) Optical diffraction pattern showing double hexagonal reciprocal lattice (from the top and bottom surfaces of the flattened tube). Periodic reflections extend to a resolution of 30 Å. (C) Optical diffraction pattern of same micrograph, overexposed to illustrate spectral limit of potentially accessible information (∼12 Å). (D,E) Correlation averaged surface lattice images. The actual resolution achieved is 22 Å, indicating both that computational correction for lattice disorders has improved the resolution from 30 to 22 Å, and that the “signal” generated by the stain beyond that limit (i.e., between 22 Å and 12 Å) is not reproducible from capsomer to capsomer. The hexagonal surface lattices are shown for the “cleaved, expanded” state (panel E) and the “uncleaved, expanded” state (panel D). Upon maturation, the T4 surface lattice undergoes a major conformational change in which its lattice constant increases from 118 to 140 Å in the “expanded” state. Normally, this transition is preceded by proteolytic cleavage of the capsid protein, gp23, from its initial molecular weight of 56,000 Da to 49,000 Da, yielding the “cleaved, expanded” state (see Black et al., 1994). However, expansion may be induced in vitro in the absence of cleavage by treatment with low concentrations of guandine hydrochloride or urea, giving the “uncleaved, expanded” state. Comparison of these images allows the 7-kDa “∆-domain” (which is removed by the cleavage reaction) to be localized: additional densities are visible around the rim of the proteomers of the uncleaved lattice (panel D). The center-to-center spacing between hexameters in the surface lattice is 140 Å. Adapted from Kocsis et al. (1997) courtesy of Dr. E. Kocsis.
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sional space. For a three-dimensional map calculated from N equally spaced projections of a particle of diameter D, an estimate of the resolution (r) is given by r ≅ π × (D/N) (Klug and Crowther, 1972). With fewer views or irregularly spaced views, the resolution deteriorates. One specific case—anisotropic resolution— encountered because of the limited angular range accessible in a tilting experiment, is considered later in the unit (see discussion of Mapping Specific Components in Large Protein Complexes).
Attainable Goals: The Art of the Possible The following discussion briefly summarizes goals that are realistically achievable with current methodology for several classes of protein specimens. Two-dimensional crystals With highly ordered molecular monolayers, electron crystallography can achieve resolutions of ∼3.0 Å in the plane (see discussion of High-Resolution Studies). With such three-dimensional density maps, it is possible to trace polypeptide chains and render a detailed account of the molecular conformation, including the locations and dispositions of prosthetic groups (Kühlbrandt et al., 1994). For less-wellordered two-dimensional crystals embedded in ice or sucrose, a more realistic goal may be a density map at 6- to 8-Å resolution. This is sufficient to describe the configuration of a bundle of α helices, a capability that is valuable for the study of intrinsic membrane proteins, many of which have transmembrane segments that are α-helical. As a prelude to detailed analysis of unstained specimens, two-dimensional and three-dimensional analyses of stained or shadowed specimens in the resolution range of 15 to 30 Å are eminently worthwhile. In general, surface details (e.g., loops connecting α helices) are visualized less well in such maps, even at high resolution, because of the limited tilt range covered (see discussion of High-Resolution Studies) and possibly also because of interactions with the support film or air interface. To overcome this limitation, at least in part, high-resolution shadowing (Fuchs et al., 1995) or atomic force microscopy (Shao and Yang, 1995; Engel and Gaub, 1997) offer important sources of complementary information on surface detail.
Helical polymers The three-dimensional structures of filaments may readily be calculated to resolutions of 20 to 30 Å for negatively stained or ice-embedded specimens. These resolutions are sufficient to define the shapes and packings of their subunits. The binding of ancillary proteins to filaments may be studied in detail by difference mapping. If an atomic model is available for the subunit, EM may be used to determine how they associate in the filament and to interpret induced structural changes (Stasiak and Egelman, 1994; Aebi et al., 1996). With bacterial flagellae, it has been possible to extend the resolution to ∼10 Å by combining many images, thereby yielding information on the packing of α helices in the filament backbone (Mimori et al., 1995; Morgan et al., 1995). The highest resolution to date with helical specimens, ∼8 Å, has been achieved with acetylcholine receptors packed in crystalline membrane tubes, again revealing the locations and dispositions of α helices (Unwin, 1996). As a class, filaments have some liabilities as compared with planar specimens, but they have the advantage of providing a full range of angular views (after combining multiple particles), so their density maps are not affected by anisotropic resolution. Oligomeric fibrous proteins When adsorbed to a thin carbon film for visualization by EM, fibrous oligomers usually present a side view. Their domainal structures may be inferred directly from micrographs, or more reliably, after image averaging. So far, negative staining has been the method of choice because of its high signal-to-noise ratio, which is necessary for precise computational straightening of these thin, curved molecules. Resolutions of 20 Å or so are usually attainable. If integrated with information from other sources—primarily sequence-based predictions, which are relatively reliable in the case of fibrous-protein motifs—it may be possible to sketch out the overall path followed by the polypeptide chain through the molecule (Steven et al., 1988; also see Fig. 17.2.2). Icosahedral viruses As a geometrically defined class of specimen, spherical viruses occupy a special niche. On one hand, they are single particles and imaged as such; on the other hand, they possess a high degree of internal symmetry that may be exploited for purposes of structural analysis. A Structural Biology
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formalism for calculating the three-dimensional structures of spherical (i.e., icosahedrally symmetric) virus capsids was developed long ago (Crowther, 1971). However, this approach came to fruition only after the advent of cryo-EM (Adrian et al., 1984; Fuller, 1987; Baker et al., 1988), whereupon it has been applied with great effect to many capsids and derivatives thereof (reviewed by Steven et al., 1997). The resolutions attained, initially in the range of ∼35 Å to ∼45 Å, have improved steadily. Recent developments have pushed the resolution down below 10 Å for favorable specimens, revealing elements of secondary structure (i.e., α helices; Conway et al., 1997; Trus et al., 1997). Isolated protein complexes There are also some large multienzyme complexes that conform to icosahedral symmetry, and accordingly may be analyzed by the same procedures as are used to study virus capsids (Stoops et al., 1992). However, protein or nucleoprotein complexes that observe lower symmetry or no symmetry constitute a much broader class of specimens, and several general approaches have been developed for calculating their three-dimensional structures from sets of images of freestanding particles (reviewed by Frank, 1996). Such complexes do not necessarily possess any symmetry, but if they do, it may be exploited. Most studies start with negative staining (see discussion of Specimen Preparation). Distinctive two-dimensional projections of the complex are recognized and averaged to yield images that are typically in the resolution range of 20 to 30 Å, and these may be synthesized into a three-dimensional density map. These experiments may then be followed up with cryo-EM, with its improved preservation and direct representation of native structure (as distinguished from the indirect representation afforded by a stain distribution). Currently, resolutions in the range of 20 to 25 Å are attained in such three-dimensional maps (Frank et al., 1995; Stark et al., 1995) and there is every reason to anticipate progress to higher resolutions in the near future.
SPECIMEN PREPARATION: YOU GET OUT WHAT YOU PUT IN Negative Staining Overview of Macromolecular Electron Microscopy
The principle of negative staining (Hayat and Miller, 1990) is to enhance the low intrinsic contrast of a protein by embedding it in a dense medium (Fig. 17.2.6B). The protein molds the
surrounding stain, and its shape is ultimately deduced from that of the stain distribution, which scatters most of the electrons. In principle, the mode of dehydration employed in negative staining—air drying—should be severely disruptive to native protein conformations. The high concentrations of heavy-metal salts and abrupt changes in pH imposed during staining are also causes for concern. Notwithstanding, several negative-staining studies of model proteins have documented excellent correspondence between experimental images and images calculated from the known, native, structure (Steven and Navia, 1980). Moreover, the technique has been found to work surprisingly well on a wide range of specimens (Bremer et al., 1992). To an increasing extent nowadays, negative staining serves as a prelude to more detailed investigation by cryo-EM. The main advantages of negative staining are that it is quick and easy to perform and has a generally sound record as attested to by studies with model proteins and with specimens subsequently analyzed by cryo-EM. Ease of use and the capability for screening many samples rapidly and inexpensively are likely to guarantee the continuing popularity of this technique. Its limitations relate mainly to structural preservation, especially where fine details are concerned, and the fact that the stain depicts only surface features, yielding little information about internal structure other than the presence of stain-penetrable cavities (see Fig. 17.2.1A and C).
Shadowing Shadowing experiments also use heavy metals to amplify the low intrinsic contrast of proteins. In this case, the metal is applied by evaporation to form a thin coating on predried specimens (Fig. 17.2.6C). Dehydration is usually effected by air drying, which is technically the simplest drying method but which usually provides poor preservation, or freeze-drying, which generally gives superior preservation (Kistler et al., 1977; Nermut, 1977; Heuser, 1983). Critical-point drying represents a third option (Newcomb and Brown, 1989). Nebulizing droplets of a protein-containing glycerol solution onto a mica surface prior to drying under vacuum at room temperature has been found to mitigate the adverse effects of this form of air drying, presumably because of the conformation-preserving effects of glycerol. This technique has been particularly efficacious as applied to filamentous proteins (Tyler
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and Branton, 1980; Milam and Erickson, 1982; Fig. 17.2.7). The metals most commonly used for shadowing are platinum and tungsten/tantalum. Apart from the state of preservation of the specimen, the major resolution-limiting factor is the grain size of the deposited metal, which in the case of platinum depends on the temperature of the specimen during shadowing. Low temperatures (e.g., <−80°C) limit the surface mobility of incident metal atoms and are con-
ducive to small grains. Room-temperature shadowing is accompanied by greater surface mobility of the atoms (larger, fewer grains). The latter property may be exploited to localize nucleation hot spots on proteins in a technique called “decoration” (Weinkauf et al., 1991). Metal may be evaporated either unidirectionally onto a static specimen or by rotating the specimen while the shadowing is taking place (Fig. 17.2.6C). Unidirectionally shadowed images allow specimen heights to be
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low-angle rotary shadowing
unidirectional shadowing
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Figure 17.2.6 Preparation of protein molecules for EM. (A) Schematic drawing of protein molecules in solution. (B) Protein molecules prepared for EM by negative staining. The molecules are adsorbed to a thin carbon film substrate and embedded in negative stain. In drying, they have undergone variable amounts flattening. (C) Protein molecules prepared for EM by heavy-metal shadowing. The molecules are supported on a thin carbon film. The top diagram depicts rotary shadowing with platinum from a low angle; note the grain-free zone adjacent to the molecule. The bottom diagram depicts unidirectional shadowing from a 45° elevation angle. Note the smaller grain size and the grain-free shadow zone. (D) Protein molecules prepared for observation by cryo-EM. The molecules are embedded in a thin film of vitreous ice maintained at a temperature of approximately −180°C.
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Figure 17.2.7 Transmission electron micrographs of filamentous proteins prepared by glycerol-spraying/low-angle rotary metal shadowing. The bar at the bottom of panel A is equal to 1000 Å. (A) At their first level of structural organization, nuclear lamin polypeptides form “myosin-like” parallel, unstaggered dimers (arrows). (B) At their second level of structural organization, lamin dimers assemble into linear head-to-tail polymers. Adapted from Heitlinger et al. (1991) courtesy of Dr. U. Aebi.
measured from the lengths of the shadows that they cast, and these micrographs may also be analyzed digitally to calculate topographic maps of molecular surfaces (Smith and Kistler, 1977; Thomas et al., 1991; Fuchs et al., 1995). On the other hand, images of rotary-shadowed specimens tend to be easier to interpret because they contain fewer lacunae where no metal grains reach. Low-angle rotary shadowing has proved particularly effective for the study of filamentous proteins (see discussion of Protein Filaments and Fig. 17.2.7).
Frozen-Hydrated (Vitrified) Specimens: Cryo-EM
Overview of Macromolecular Electron Microscopy
In cryo-EM, the specimen is not dehydrated, but instead is preserved and observed in a thin film of ice (Fig. 17.2.6D). Provided that this film is thin enough and the freezing is fast enough, the water molecules are immobilized in an amorphous vitreous state, thus avoiding dehydration and/or physical disruption of the specimen by the formation of ice crystals. Once frozen, the specimen is inserted into a specially cooled specimen holder, or cryoholder, for transfer into the EM. Below about −110°C, ice
does not sublime at a significant rate, even in the high vacuum of the EM column. Provided that the specimen remains below about −150°C, vitreous ice will not undergo a phase transition to a crystalline state. Commercially available cryoholders cooled by liquid nitrogen can maintain specimen temperatures of about −180°C, preventing both sublimation and crystallization. A few EMs are equipped to operate at liquid helium temperatures (approximately −269°C; Fujiyoshi et al., 1991), which confers some additional radiation-hardiness. Because cryospecimens remain fully hydrated, there is good reason to suppose that native protein conformations are preserved. This expectation is supported by high-resolution electron-diffraction patterns recorded from ice-embedded catalase crystals (Taylor and Glaeser, 1974). A number of technical problems arise in connection with observing vitrified specimens. Heat transfer between the cryoholder and the microscope may cause the specimen to drift. Frozen specimens are liable to deposition of hoar-frost or other contaminants while in transit into the EM and even within the column, although additional cryoshielding
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protein molecules in solution
Figure 17.2.8 Formation of two-dimensional membrane crystals. (A) Intrinsic membrane proteins, initially solubilized in detergent, are reinserted into lipid bilayers as the detergent is removed by dialysis. (B) Extrinsic membrane protein with a specific affinity for a lipid monolayer (LangmuirBlodgett film).
helps to minimize this effect. Because ice is not a good conductor, electric charge may build up on the specimen and distort the image. Also, vitrified specimens are acutely sensitive to electron irradiation (Conway et al., 1993) and, as noted previously, every precaution must be made to minimize the electron dose. These matters are still the focus of active developmental work; nevertheless, progress to date has made possible a wide range of experiments at resolutions that are already good and that continue to improve.
Crystallization of Proteins in Two Dimensions Because monolayer crystals are so important for EM studies of proteins at high resolution (see discussion of High-Resolution Studies), a good deal of effort has been invested in their production. Occasionally, conditions that promote the formation of such arrays have been found serendipitously; e.g., Zn2+ ions promote monolayer crystal formation for soluble tubulin (Larsson et al., 1976). Two systematic approaches have been pursued, both based on the principle of confining the crystallands to a two-dimensional surface while maintaining their hydration (Fig. 17.2.8). Intrinsic membrane proteins are an attractive class of molecules for study by EM (also
see discussion of Membrane Proteins). The insolubility of these proteins adds to the list of parameters to be varied in a crystal screen. The sequence of steps that leads to crystallization— purification, concentration, and precipitation—must be expanded to include reinsertion into a lipid bilayer (Fig. 17.2.8A). Thus, the choice and concentration of detergent used to maintain solubility, as well as the choice and concentration of exogenous lipids for the reinsertion step, are also important. The temperature dependence of lipid mobility offers a possible mechanism for causing intramembranous proteins to undergo ordered aggregation. As the temperature falls below the threshold for a gel/liquid crystal phase transition, proteins may be “frozen out” of the ordered lipid phase. Based on this idea, a temperature-ramp method has been developed (Engel et al., 1992). A comprehensive overview of membrane-protein crystallization methods has been given by Kühlbrandt (1992). An alternative crystallization strategy, applicable to extrinsic membrane proteins, involves the use of lipid monolayers formed at a (flat) air/water interface (Brisson et al., 1994; Fig. 17.2.8B). Such a monolayer presents a suitable surface for ordered aggregation of the proteins adhering from the underlying bulk phase. The protein of interest should have an
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affinity for the polar surface of such a monolayer. This affinity may either be natural or induced by appropriate chemical modification of the protein or the lipid head group (Kubalek et al., 1994; Celia et al., 1994). It should cause the protein to attach to the monolayer at a defined angle, leaving each molecule’s azimuthal orientation about this axis as its the sole remaining degree of freedom, apart from the packing density. This crystallization technique has already been applied successfully in a number of systems, and excellent ordering and impressive resolutions have been obtained—e.g., annexins (Olofsson et al., 1994), α-actinin (Taylor and Taylor, 1994), and streptavidin (Avila-Sakar and Chiu, 1996).
MAPPING SPECIFIC COMPONENTS IN LARGE PROTEIN COMPLEXES Once the three-dimensional structure of a large multiprotein complex has been determined to a resolution of 20 to 30 Å, it remains a major task to localize its various components within this frame of reference. A variety of approaches have been devised for different biological systems. They have in common the principle of seeking to modify the complex in some chemically defined way (e.g., the extraction of a loosely bound subunit or the binding of an antibody) and then visualizing the accompanying structural change. One serendipitous finding has been that when two well-defined— i.e., essentially noise-free—two-dimensional images or three-dimensional maps are compared, it is possible to detect structural differences that are smaller than the nominal resolution of the data. The explanation for this seeming paradox is that detectability is not the same as resolvability, but is a considerably more achievable goal. For instance, a point scatterer (infinitely thin) may be visualized by imaging techniques of modest resolution, although its representation is correspondingly blurred.
Difference Imaging: Depletion, Limited Proteolysis, and Complementation
Overview of Macromolecular Electron Microscopy
One powerful approach to mapping subunits or domains is based on seeking to determine conditions under which one (or more) subunit or domain is selectively removed, while the remainder of the complex is left unchanged. Quantitative comparison of the modified complex with the original complex (“difference imaging”) allows the vacated site to be localized. Two methods of accomplishing such re-
movals are proteolytic dissection and selective extraction. Treatment with a specific protease may excise a domain or subunit (Carrascosa and Steven, 1978). Alternatively, extraction may be achieved by treating the specimen with a critical concentration of some denaturant such as urea or guanidine hydrochloride (Newcomb et al., 1993) or an extreme of pH (Yeager et al., 1994). Alternatively, alteration of surface charge by citraconylation (Aebi et al., 1977a) or similar reactions such as succinylation may prise particular subunits out of a complex by electrostatic repulsion. In any case, the biochemical groundwork has to be established before structural analysis becomes worthwhile. A third variation on this theme is to study the formation of complexes between the protein of interest and another protein that is known to bind to a particular subunit (Olson et al., 1993). This ligand then serves as a bulk marker for that subunit. For any of the above treatments to achieve its intended purpose, it is important that the induced change should be strictly local. Thus, quantitative structural comparisons between images of treated and control (untreated) complexes are essential to guard against generalized structural changes, which potentially compromise the interpretability of such an experiment.
Immuno-EM: Labeling with Antibodies and Fab Fragments Perhaps the most widely used variant of the marker-binding approach is visualization of complexes labeled with antibodies. It is essential that the specificity of the antibodies be clearly established by immunochemical methods. Immuno-EM labeling experiments may be performed either with intact antibodies or with monovalent Fab fragments (Aebi et al., 1977b; Wang et al., 1992; Porta et al., 1994). Both have their advantages. Whole antibodies may cause sizable immunoprecipitates to form if the protein happens to contain more than one copy of the epitope, and these precipitates may result in complicated images that are difficult to interpret. This problem does not arise with Fab fragments. On the other hand, precipitation may be advantageous in some situations. For instance, if the specimen is undesirably dilute and too fragile to survive centrifugation, it may be concentrated by immunoprecipitation (Newcomb et al., 1996). With three-dimensional density maps calculated from cryomicrographs of antibody-labeled specimens, it turns out to be possible to localize the “footprint” of the antibody-binding
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site to within a few angstrom units, even if the map is limited to, say, 25-Å resolution (Steven et al., 1992; Smith et al., 1993). Similarly, the interaction between antibody and antigen can be modeled in detail by drawing on a priori information on the (conserved) structures of Fab moieties. The characteristic bilobed shape of an Fab may be readily detected in a cryo-EM density map at 25- to 30-Å resolution. Moreover, its orientation may be ascertained with high precision on the reasonable assumption that its point of contact with the protein marks the epitope.
Heavy-Metal Clusters: Small Site-Specific Labels An important innovation capable of localizing sites on a protein complex has been the development of heavy-metal cluster compounds with organic moieties of defined functionality. Thiol-reactive clusters are particularly useful for labeling free cysteine residues, and site-specific mutagenesis techniques make it possible to insert cysteines into promising sites in a protein of interest. Single heavy atoms are hard to detect in the EM when attached to protein molecules, although they may be visualized when dispersed over a thin carbon film (Crewe et al., 1968). However, clusters of heavy atoms have enough scattering power to be visible, even when attached to proteins. The 11atom gold cluster, undecagold, has been exploited successfully in several studies (Hainfeld, 1987; Milligan et al., 1990; Yang et al., 1994; Wagenknecht et al., 1995; Fig. 17.2.9), as well as a larger cluster of about 40 gold atoms called Nanogold (Gregori et al., 1997; Fig. 17.2.10). Nanogold may be visualized against a background of low-density negative stains (Wetzel and Baumeister, 1995), a property that expands its ease of use. Cluster compounds are now commercially available (see discussion of Resources and User Facilities), and as experience with them accumulates, this powerful labeling technique should become broadly applicable.
HIGH-RESOLUTION STUDIES In the past few years, two EM-based approaches have progressed to the point of describing protein structures in near-atomic detail and revealing the detailed folds of polypeptide chains. The first is electron crystallography of highly ordered crystalline plates. These arrays are typically several microns or more in lateral extent, but very thin—ideally, only one molecule thick. To date, the two systems in which electron crystallography has advanced fur-
Figure 17.2.9 Dark-field STEM micrograph of a single ferritin molecule surrounded by goldlabeled Fab′ fragments of anti-ferritin antibodies. The specimen was unstained and supported on a thin carbon film. The small white dots are undecagold clusters and the large central white spot is the iron core of the ferritin molecule. Fab′ antibody fragments were covalently linked to undecagold at their hinge sulfhydryl, then incubated with ferritin and column purified. Courtesy of Dr. J. Hainfeld, Brookhaven National Laboratory.
thest—purple membrane (Grigorieff et al., 1996) and the light-harvesting complex (Kühlbrandt et al., 1994)—are both intrinsic membrane proteins, and encouraging progress has also been made in a number of other systems. This work is not confined to membrane proteins—a resolution of ∼6 Å has been achieved with sheets of tubulin (Nogales et al., 1995), a cytoskeletal protein of great importance that has so far resisted attempts to produce three-dimensional crystals suitable for X-ray diffraction analysis. In the second approach, three-dimensional density maps calculated by cryo-EM to resolutions of the order of 20 to 30 Å have been found to contain enough information to reveal the precise docking of subunits whose three-dimensional structures were separately solved to high resolution by other methods (Rayment et al., 1993; Stewart et al., 1993; Baker and Johnson, 1996). Thus, cryo-EM at moderate resolution serves as a bridge that allows the high-resolution information on individual subunits to be extended to the study of higherorder structures.
Electron Crystallography of Highly Ordered Two-Dimensional Arrays Much of the electron crystallography methodology now available was pioneered in a series
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Figure 17.2.10 (A) STEM micrograph of proteasomes after binding Nanogold-labeled amyloid β protein, thus indicating its binding location. Labeling and binding were confirmed by electrophoresis, blots, and activity studies (Gregori et al., 1997). The 14-Å Nanogold clusters (arrows) are clearly visible with the low-density stain, methylamine vanadate (NanoVan; Nanoprobes), which enhances visualization of subunit detail. The bar is equal to 100 Å. (B) An array of top views of labeled proteasomes. (C) Side-views of corresponding labeled proteasomes. (D) Maps of the label sites recorded on many micrographs of the two views in panels B and C. Reprinted from Gregori et al. (1997), with permission, courtesy of Dr. J. Hainfeld, Brookhaven National Laboratory.
Overview of Macromolecular Electron Microscopy
of studies of the purple membrane of Halobacter halobium (see Grigorieff et al., 1996, and the references in that paper). Under anerobic conditions, this bacterium expresses a pigmented protein called bacteriorhodopsin (bR), which functions as a light-driven proton pump to generate an alternative source of energy. In vivo, bR assembles into a crystalline monolayer that is exceptionally well ordered and thus ideally suited for study by crystallographic methods. The procedure relies on combining Fourier amplitudes (structure factors) collected from electron-diffraction patterns of single crystalline patches, with phases obtained from calculated Fourier transforms of low-dose electron micrographs (Unwin and Henderson, 1975). A single-projection view thus synthesized provides a central plane through the three-dimensional Fourier transform of the repeating unit
(in this case, a trimer of bR). The remainder of the transform is filled in by tilting specimens to progressively higher angles and collecting diffraction patterns and images at each step. Each such view contributes another plane of the transform. Finally, the three-dimensional density map is calculated by inverse Fourier transformation. Because the accessible tilt range is limited and because data collection becomes progressively more difficult at higher tilt angles, the resolution of the density map is lower in the dimension perpendicular to the membrane. Nevertheless, electron-crystallographic analysis of bR has already been carried to a resolution of 3.5 Å in plane and ∼5.5 Å in the third dimension (Grigorieff et al., 1996), which is sufficient to define the fold of the polypeptide chain and to locate the prosthetic group (retinal).
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The number of naturally crystalline protein monolayers that are as well ordered as bacteriorhodopsin is very small. To extend the range of specimens eligible for electron-crystallographic analysis, it is necessary to form artificial monolayer crystals (see discussion of Crystallization of Proteins in Two Dimensions). So far, the greatest success has been achieved with photosynthetic light-harvesting proteins (Kühlbrandt et al., 1994) and porins, a class of bacterial channels (Jap et al., 1991). However, promising results have also been obtained with several other proteins, including members of the aquaporin class of hydration channels (Walz et al., 1995) and gap junction connexins (Unger et al., 1997; see Fig. 17.2.11). The latter study (of connexins) represents an important new departure in that the connexin arrays studied were not extracted from a source tissue but were expressed from a recombinant baculovirus in baby hamster kidney (BHK) cells, a cultured cell line that does not normally deploy gap junctions. Just as overexpression of soluble proteins has become a central element for structural studies, the expression and assembly of membrane proteins has become a vital objective for high-resolution studies of membrane proteins. As an added element of challenge, the requirement that expressed proteins should be inserted into membranes imposes a constraint on the amounts that can be expressed. For example, overloading the plasma membrane with foreign proteins may weaken the cell and impair its synthetic competence. If expressed membrane proteins precipitate into inclusion bodies, extraction and reconstitution may be difficult; perhaps incorporation into cytoplasmic storage membranes, if this can be contrived, may be a practically useful solution.
Hybrid Approaches: EM and X-Ray Crystallography The complementarity between EM and Xray crystallography (or any other technique capable of yielding subunit structures at high resolution) has been demonstrated in recent hybrid studies that combine the best of both approaches (Whittaker et al., 1995; Baker and Johnson, 1996). These studies serve as compelling role models for investigations of interactions that underlie protein-protein recognition phenomena and the assembly and functioning of protein complexes. In such studies, a highresolution technique (to date, X-ray crystallography) is used to define the structures of individual molecules or subunits, and three-di-
mensional cryo-EM at 20- to 30-Å resolution is used to define the placement of these subunits in larger complexes. The pivotal point, which may not be evident, is that the cryo-EM density maps, despite their modest nominal resolution, contain sufficient information to allow the subunits to be docked to within a few angstrom units in their relative positions and orientations. To come to terms with this seemingly fanciful proposition, it should be borne in mind that interpreting the cryo-EM map is not a structure determination per se, but rather is an exercise in pattern recognition for which much a priori information is available. In this context, it is worth bearing in mind that interpreting a crystallographic map is also in part an exercise in pattern recognition. The assumptions made include the following: the density distribution should represent a known number of polypeptide chains; these chains are continuous and unbranched; their amino acid sequences are (usually) known, which assists chain tracing; and familiar elements of secondary structure, such as α-helices, may be recognized at an early stage, thereby reducing the complexity of the problem. In this vein, interpreting a cryo-EM map of a complex of subunits of known structure hinges on whether there is enough information for the locations and orientations of these subunits to be identified correctly. It is sometimes argued that one only needs to determine six parameters per subunit—three Cartesian coordinates for its center of mass and three orientation angles. For this purpose, a limited amount of structural information should suffice. The only situation in which such correlation of crystallography with cryo-EM would be compromised would be if large-scale conformational changes were to accompany complex formation. The likelihood of such an eventuality has to be assessed on a case-by-case basis.
RESOURCES AND USER FACILITIES The scanning transmission electron microscopy (STEM) facility at Brookhaven National Laboratory, Upton, Long Island, New York, is supported by the National Center for Research Resources (NCRR) of the National Institutes of Health (NIH). This facility operates two high-performance scanning transmission electron microscopes and specializes in STEMbased mass determinations and mass-mapping analyses. Its resources are available to appropriate user-initiated projects. The Brookhaven facility also has considerable experience in la-
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Figure 17.2.11 Projection of gap junction connexons, as visualized at 7-Å resolution by electron crystallography of ice-embedded specimens (Unger et al., 1997). The hexagonal unit cell contains two layers of hexameric rings of a rat connexin protein, truncated at its C terminus. This construct was expressed in BHK cells, where it spontaneously formed connexon plaques. There are four transmembrane α helices per subunit; two are oriented perpendicular to the membrane and are visible as the 10-Å disks of densities that lie, respectively, outside and inside the ring. The other two helices adopt a slanting orientation. As seen in projection, they overlap to contribute the features seen in the ring. (A) Contour plot. The side of the hexagonal unit cell is 95 Å. (B) Gray-scale representation. High densities are shown as white. Courtesy of Dr. M. Yeager.
Overview of Macromolecular Electron Microscopy
beling molecules with heavy-metal clusters and in visualizing cluster-labeled molecules embedded in low-density stains. For more information see Internet Resources. The National Center for Macromolecular Imaging at Baylor College of Medicine, also supported by the NCRR, has a 400-keV electron cryomicroscope and computer processing equipment to assist or collaborate with biological users in studying high-resolution three-dimensional structures of macromolecules. For more information see Internet Resources. Heavy-metal cluster compounds for protein labeling experiments may be purchased from Nanoprobes (see SUPPLIERS APPENDIX for contact information; for technical questions contact Dr. Richard Powell, Research Director).
Image-processing software for the analysis of biological electron micrographs of proteins and complexes has been summarized comprehensively in a recent special issue of the Journal of Structural Biology (Carragher and Smith, 1996).
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Overview of Macromolecular Electron Microscopy
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Kocsis, E., Cerritelli, M.E., Trus, B.L., Cheng, N., and Steven, A.C. 1995. Improved methods for determination of rotational symmetries in macromolecules. Ultramicroscopy 60:219-228.
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Roseman, A.M., Chen, S., White, H., Braig, K., and Saibil, H.R. 1996. The chaperonin ATPase cycle: Mechanism of allosteric switching and movements of substrate-binding domains in GroEL. Cell 87:241-252.
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INTERNET RESOURCES http://www.bnl.gov (click on User Facilities and Scanning Transmission Electron Microscope) Brookhaven National Laboratory, Upton, New York [email protected] E-mail for Dr. Martha Simon, Manager, User Projects, Brookhaven National Laboratory [email protected] E-mail for Dr. J. Wall, Director, Bookhaven National Laboratory http://ncmi.bioch.bcm.tmc.edu National Center for Macromolecular Imaging, Baylor College of Medicine, Houston, Texas [email protected] E-mail for Dr. Wah Chiu of the National Center for Macromolecular Imaging
Contributed by Alasdair C. Steven National Institute of Arthritis and Musculoskeletal and Skin Diseases Bethesda, Maryland
Structural Biology
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Principles of Macromolecular X-Ray Crystallography In the last 10 years there has been an astonishing increase in the number of protein structures determined by X-ray crystallography. As of September, 1996, there were some 4700 such structures in the Protein Data Bank, and the number is increasing exponentially. There is almost no limit to the complexity of the structures that can presently be determined by X-ray diffraction, provided that adequate ordered crystals are available. These advances have had a corresponding effect on concepts of modern biochemistry, where the profusion of three-dimensional structures has introduced a new degree of chemical sophistication into the discussion of the functions and interactions of biological macromolecules. However, the technical nature of crystallography presents a formidable barrier to the biochemist who wishes to assess a crystallographic paper. How can the quality of the information coming from this technique involving crystals, computers, and X rays be critically evaluated? This unit does not attempt to explain all the details of crystal-structure determination, which are more adequately described in textbooks. It does, however, provide a brief overview of the general principles and methodology, as well as a more detailed discussion of the methods used to prepare a protein sample for crystallization trials and the various stratagems that are sometimes necessary to achieve this goal.
CRYSTAL STRUCTURE DETERMINATION BY X-RAY DIFFRACTION There are a number of excellent reference works on macromolecular crystallography. These include early classic texts such as Blundell and Johnson (1976) and the more recent, comprehensive description of the principles of X-ray diffraction by Drenth (1994). There are several volumes of Methods in Enzymology (Wyckoff et al., 1985; Sweet, 1997) that provide more detail about advanced techniques. Consistent with the growing accessibility of protein crystallography to a broader community of scientists, there are also texts such as Jones et al. (1996) and Rhodes (1993) that are meant more for the biochemist or molecular biologist. Another recent text is by McRee (1993).
X-ray diffraction analysis of crystal structures is quite analogous to using a very highpower microscope that can see molecular shapes. The wavelength of the light used is usually in the vicinity of 1.5 Å, about the length of a carbon-carbon single bond, and the use of X rays with this wavelength permits, in theory, resolution of individual atoms. In a microscope, a light beam strikes a specimen and the scattered light is brought to a focused image by a lens. In principle, X-ray diffraction functions in much the same way except that there is no X-ray lens to focus the X rays that are scattered by the electrons within the crystal. Therefore, it is not possible to directly obtain an image that can be interpreted. Instead the crystallographer measures the intensities of the scattered X rays from the crystal and uses these, together with some additional data described below, to compute an image. Crystallographers use a crystal because the signal from a single molecule would be too weak to detect. The crystal acts as a three-dimensional diffraction grating and scatters the incident beam of X rays only in certain directions. But all the molecules in the crystal scatter together in these directions, and since there are >1015 molecules, at least in most crystals, this enables the scattered signal to be recorded before the specimen is destroyed by radiation damage. It can be demonstrated that this diffraction is the vector sum of the diffraction by the individual atoms of the unit cell, each having an amplitude that is related to the number of electrons and the size of the atom as well as a phase that is derived from the position of the atom within the unit cell. The sum of these individual contributions is a vector with an amplitude |F(h)| and a phase α(h) where h is equal to (hkl) and h, k, and l are the Miller indices that define the diffraction maxima. F(h), called the structure factor (because it depends only on the structure of the scatterer), can be expressed in terms of the fj, the individual scattering factors for the j atoms in the unit cell, and the atomic coordinates xj. F ( h) = Σ j f j exp 2πi( h ⋅ x j ) = F ( h ) exp iα ( h)
By rotating the crystal relative to the X-ray beam, it is possible to record and measure the
Contributed by Alison B. Hickman and David R. Davies Current Protocols in Protein Science (1997) 17.3.1-17.3.15 Copyright © 1997 by John Wiley & Sons, Inc.
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complete diffraction pattern of the crystal. The crystallographer can measure |F(h)|, which is directly related to the intensities of the diffraction maxima, but cannot experimentally measure the phase angle, α(h). These lost phases have to be recovered by some other method. When both the amplitudes and the phases of the scattered X rays are known, they can be combined to produce an electron-density map of the contents of the unit cell, the repeating unit of the crystal. The electron density (ρ) at any position xyz in the unit cell of the crystal is represented by the following equation (with i equal to the square root of −1). ρ( x , y, z ) = Σ hkl F ( h, k , l ) exp iα ( hkl ) exp −2 πi( hx + ky + lz )
Data Collection
Principles of Macromolecular X-Ray Crystallography
There are many methods available for measuring the intensities of the scattered X rays in X-ray crystallography. The two early methods involved film, which could measure an array of peaks simultaneously, and proportional or scintillation counters, which measured the data one spot at a time. Film has been replaced by imaging plates, which can be exposed to the scattered X rays, then rapidly read, erased, and reexposed. To reduce dead time, commercial imaging-plate detectors use two imaging plates so that one can be exposed while the other is being read. Multiwire proportional detectors have replaced single detectors, again permitting the measurement of many reflections simultaneously. A modern laboratory might employ either or both of these methods of data collection; for the future there is the emerging technology of charge-coupled devices (CCDs). The laboratory X-ray source has traditionally been a rotating anode X-ray generator, usually with a copper target, which gives a peak of CuKα radiation of wavelength 1.54 Å. However, increasingly data are being collected using synchrotron X-ray sources at a few centers specifically designed for this purpose. Synchrotrons can provide X-ray intensities many hundreds or even thousands of times more intense than those attainable in the laboratory. They also have the advantage of providing a continuous band of radiation wavelengths from which a monochromatic beam of predefined wavelength can be selected for anomalous dispersion experiments (see discussion of Phase Determination). Until recently, data for biological macromolecules were measured from crystals at am-
bient temperatures. However, lower temperatures reduce radiation damage to the crystal and much current data collection is carried out using so-called “cryo” techniques at temperatures close to that of liquid nitrogen. In this way, a single crystal will suffice to measure a complete data set.
Phase Determination Having obtained crystals of a protein that diffract X rays to a reasonable resolution, the next step is to obtain an interpretable electrondensity map, and for this one needs a reasonably accurate set of phase angles, α(hkl). Protein crystallographers generally use three methods to obtain phase information, two of which (multiple isomorphous replacement and anomalous scattering) involve heavy atoms, while the third uses a procedure called molecular replacement. Multiple isomorphous replacement The isomorphous heavy-atom replacement method, which was originally applied to the determination of phases for hemoglobin by Max Perutz and colleagues (Green et al., 1954), requires the attachment of a heavy atom to the protein in the crystal. There are enough sites of weak attachment on protein surfaces that it is usually sufficient simply to soak the crystals in a solution containing the heavy atom. An alternative procedure involves cocrystallizing the protein in the presence of the heavy-atom compound. The heavy atom will then also contribute to the X-ray scattering, and because it is electron-dense (with a large fj), there will be a measurable difference in the intensities of the diffraction pattern. These differences can be used to locate the site of binding of the heavy atom. If several such crystals can be prepared with different heavy atoms binding to different sites on the protein, the results can be combined to calculate the phases of the protein structure factors. From these, an electron-density map can be computed. This map will contain errors resulting from the difficulties associated with the accurate measurement of small intensity differences. It will contain additional errors that are due to the imprecision of the phasing process, which in turn result from the lack of isomorphism between the native and heavyatom crystals. Nevertheless, it is usually possible to identify regions that will fit the known sequence and to proceed to construct a model that will be consistent with the polypeptide backbone and side chains of the protein.
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Anomalous-scattering methods A second method of phase determination also uses heavy atoms but takes advantage of the fact that atoms at wavelengths near their absorption edges will scatter X rays anomalously with a change in amplitude and phase. This anomalous scattering can be used for a single heavy-atom derivative, first to identify the binding sites for the heavy atom, then to compute the phases for the crystal as a whole. Although this procedure can be carried out at a single wavelength, the effects are much greater at wavelengths near or at the absorption edge of the anomalous scatterer. By measuring the diffraction intensities at wavelengths near and distant from the absorption edge, optimal intensity differences can be observed and used to solve the structure. The availability of tunable sources of X rays from synchrotron radiation has provided an effective means of doing this and has led to the development of the multiwavelength anomalous dispersion (MAD) procedures by Wayne Hendrickson and his colleagues, who used selenomethionine as a replacement for methionine to provide a suitable anomalous scatterer (Yang et al., 1990). The big advantage of the anomalous-scattering procedure is that the structure can be determined with data collected from a single crystal containing a suitable heavy atom. Molecular replacement When the structure of the protein is expected to be closely related to another known protein structure—e.g., in the case of a mutant protein that has crystallized in a different crystal form, or when the unknown protein has high sequence homology with another protein of known structure—this similarity may be used to obtain preliminary phase information about the unknown structure, which can then be refined. This method, known as the molecular-replacement method, is used in two stages—first to determine the rotation function and then to determine the translation function. The rotation function determines the rotation necessary to position the known molecule in the unit cell with an orientation corresponding to the unknown molecule. The translation function is then used to determine the position of the correctly oriented search molecule in the cell. In general this consists of stepping the search molecule through the unit cell, avoiding bad stereochemical contacts and looking for a location that will provide a reasonable agreement between the observed and calculated structure factors. A detailed description of the applica-
tion of this method is beyond the scope of this chapter; interested readers may consult texts such as Drenth (1994) or the chapter by Tickle and Driessen (1996) for more information. Molecular replacement can be very powerful when the two molecules are not too dissimilar, and for molecules belonging to some superfamilies, such as antibody Fabs, it is now the method of choice. Several software packages exist for the molecular-replacement method. These include AMORE (Navaza, 1994), MERLOT (Fitzgerald, 1988), and CCP-4 (Collaborative Computer Project No. 4, 1994).
Resolution Once the phases are known, an electron-density map can be calculated. The resolution of this map depends on the angular extent to which the intensity data have been measured. This is related to the spacing, d, of lattice planes within the unit cell according to Bragg’s Law:
λ = 2d sin θ where λ is the wavelength of the diffracted X rays and θ is the diffraction angle. Resolution in a protein structure is usually defined as the value of the dmin corresponding to the θmax of the data set. For highly ordered small-molecule crystals that can diffract to the limit allowed by the wavelength of the X rays used—i.e., to θmax = 90°—the resolution would be λ/2 Å. Proteins have not yet been observed to attain this resolution, and crystallographers usually have to settle for something less. Figure 17.3.1 shows the effect of resolution on an electron-density map for an α helix. It can be seen that at low resolution (e.g., 6 Å) there is no detail, but the helix takes the form of a rod of density. As the resolution increases (i.e., d decreases between 3.0 and 2.0 Å) the details of the main-chain and side-chain structures begin to appear. Individual atoms cannot be seen until the resolution is 1.0 Å or better, but based on the knowledge of peptide and amino acid shapes from small-molecule crystal structures it is possible to fit the map with a model based on the known sequence of the protein. The ultimate resolution of protein data is determined by the degree of order in the crystal. Small-molecule crystals that do not contain large amounts of water usually can diffract to a resolution of better than 0.5 Å. However, protein crystals are inherently less ordered than this, partly because the protein structure itself has flexibility as a result of being a long polypeptide chain held together by weak forces— e.g., van der Waals’ forces and hydrogen bonds.
Structural Biology
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G
H
Figure 17.3.1 Calculated electron density for an α helix of myohemerythrin at resolutions of (A) 6.0 Å; (B) 5.0 Å; (C) 4.5 Å; (D) 4.0 Å; (E) 3.0 Å; (F) 2.5 Å; (G) 2.0 Å; and (H) 1.5 Å. Courtesy of Dr. Steven Sheriff, Bristol-Myers Squibb, Princeton, N.J. Principles of Macromolecular X-Ray Crystallography
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Also, the protein molecules are not packed identically in the crystal because there tend to be relatively few interactions between adjacent protein molecules; these are mostly weak and often they occur through intermediate water molecules. Finally, the spaces between the molecules are filled with large amounts of solvent, and these solvent channels tend to be quite disordered. Typically, a protein crystal will contain 50% solvent, the bulk of which is disordered. As a result, protein crystals do not often diffract beyond 1.8 Å, and the majority of protein crystals used for structural studies diffract in the range of 2.0 to 2.8 Å. It should be noted that since a protein crystal diffraction pattern does not just abruptly stop but gradually fades away with diffraction angle, the assignment of a resolution limit for a given crystal is to some extent subjective. These limitations on resolution place limits on the accuracy of the structure that is being determined. For instance, water molecules are difficult to locate with precision in crystal structures where the resolution is worse than 2.5 Å. Also the lower resolution of the electron-density maps in protein crystals means that individual bonded atoms cannot be resolved. However, the knowledge from small-molecule crystallography about the dimensions of the amino acid components enables the crystallographer to fit the map and make reasonable assumptions about the positions of the atoms. Let us assume that the heavy-atom phasing (see discussion of Phase Determination) has produced an electron-density map to a resolution of 2.5 Å. The next step is to fit a polypeptide chain with the correct sequence to this map. This can be done by first using a software package such as O (Jones et al., 1991) that will fit the polypeptide backbone automatically wherever the map quality is good enough to permit such interpretation. Once this is done, the next step is to identify sequence markers— i.e., those amino acids with large side chains such as Cys or Trp that can be recognized easily. The sequences between these markers can then be filled in. In this way a preliminary model for most of the structure can be assembled, although there are usually a few segments of polypeptide chain for which the map quality does not permit interpretation as a result of phasing errors or disorder in the protein structure. This preliminary model of the protein is now ready for the process of refinement.
significant errors resulting from the imperfect phasing of the experimental electron-density map. This model can be improved by a procedure that crystallographers call refinement. The aim of refinement is to minimize differences between the calculated and experimental intensities while at the same time optimizing the stereochemistry. The initial model can be used to compute the scattering amplitudes from the equation for structure factors (see discussion of Crystal Structure Determination by X-ray Diffraction). The calculated amplitudes |Fc(h)| are compared with the observed |Fo(h)|, and the value of
Refinement
and can vary from 0.4 to 0.5 (or from 40% to 50%, if expressed as a percentage) for an es-
The initial model of the protein will contain
∑ Fo ( h ) − Fc ( h )
2
h
is minimized, subject to some chemical constraints. There are a variety of software packages available for refinement, listed in Westhof and Dumas (1996). Since the number of diffraction data measured is usually less than the sum of the atomic occupancy and positional parameters to be determined, it is necessary to provide some additional information—e.g., about the allowed bond lengths, angles, and planarity of groups of atoms—in order to effect a meaningful refinement of the structure. This can be done either by placing constraints on the dimensions of groups of atoms or by restraining the dimensions to be within an ideal set of distances and angles. Reference dimensions are taken from the crystal structures of chemically similar groups determined by small-molecule crystallography where the resolution of the X-ray data is generally much higher than for proteins. The residual ∆F will ultimately approach a limiting low value, which cannot be reduced by further refinement. This ultimate residual is due partly to errors in the experimental measurements of the values of F, partly to the failure to locate all of the ordered waters, and partly to the fact that the refined structure, with the constraints on stereochemistry, does not necessarily correspond exactly to the protein structure in the crystal. Also, it is difficult to model the dynamic nature of the protein molecule. The progress of the refinement can be indicated partly by the value of the so-called R factor, given by: R=
∑ Fo ( h) − Fc ( h) ∑ Fo ( h) Structural Biology
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sentially correct but unrefined structure, to values as low as 0.12 (12%) for an exceptionally well-refined structure. Most well-refined structures produce R values <0.2 (20%). A low R value does not necessarily indicate that the structure is correct, and there have been several examples of partially incorrect structures that have yielded apparently low R values. Brunger (1992) has proposed a now widely used crossvalidation procedure that involves omitting a certain percentage (10% is suggested) of the reflections in the refinement but using them to calculate an Rfree, which will then provide an independent check of the validity of the structure. Rfree is calculated in the same way as the standard R value, except that it is limited to these unrefined reflections.
The Ramachandran Plot
Principles of Macromolecular X-Ray Crystallography
Another criterion used by crystallographers to assess their structures and to detect errors in the tracing of the polypeptide chain is known as the Ramachandran plot (Sasisekharan, 1962; Ramachandran et al., 1963). These authors demonstrated that the polypeptide backbone is not free to assume any conformation and proceeded to define systematically which regions of conformational space were allowed. The unallowed regions result from unacceptable contacts between the main-chain atoms, including the β carbon atom. The Ramachandran plot provides a simple way of displaying the conformational data for a protein. The conformation at each α carbon atom can be defined by the dihedral angles φ and ψ, which correspond to rotations about the single bonds to the α carbon from the amide nitrogen and the carbonyl carbon, respectively. Varying φ and ψ will cause various atoms to make stereochemically unacceptable contacts. These combinations of φ and ψ will then represent high-energy conformations, and will for the most part be forbidden. This can be plotted on a planar diagram, the Ramachandran plot. This plot defines the parts of conformational space that are available to a polypeptide chain. The various secondary structures, of course, lie within the allowed regions—e.g., the α helix, the β sheet, and the collagen fold (see UNIT 17.1). The Ramachandran plot is used by crystallographers to demonstrate that the structure they have determined is stereochemically acceptable, and therefore appears in many structure papers. These days, most refinement programs have stereochemistry restraints built into them, so that it is rare to see a Ramachandran plot with a number of outliers.
PROCEDURES FOR CRYSTALLIZATION Growing protein crystals suitable for structure determination has frequently been referred to as an art rather than a science. Although certain aspects of the method are empirical, there are many factors that can be controlled to maximize the probability of success. The overall approach is an iterative one. An initial set of crystallization conditions is tried, the outcome is observed, and if crystals do not immediately form, the conditions are then modified to take into account what was learned during the previous attempt. The following sections describe a general approach to obtaining protein crystals and include considerations to keep in mind throughout; examples from the recent literature and the authors’ own experiences with HIV-1 integrase are included where appropriate. Many excellent reference texts and reviews cover specific aspects of protein crystallization in more detail than will be presented here. These include McPherson (1982, 1990), Weber (1991), Carter (1992), Ducruix and Giegé (1992), and Gilliland and Ladner (1996). Valuable information can also be obtained from a number of Internet Web sites. The reader is referred to the Hampton Research Home Page (see Internet Resources) and the other Web sites described therein. Another useful reference is the Biological Macromolecule Crystallization Database (Gilliland et al., 1994; http://ibm4. carb.nist.gov:4400/bmcd/bmcd.html) where information is compiled on biological macromolecules that have been crystallized and the methods used.
Sample Preparation The decision to attempt protein crystallization often begins with the observation that one has in hand a protein that is purifiable to relatively high homogeneity and shows promise in terms of solubility. Unfortunately, there are no absolute requirements for either homogeneity or solubility—what works for one protein may not be sufficient for another. Nevertheless, “the higher the better” is a good rule of thumb. Assessing sample homogeneity Protein homogeneity is usually assessed using denaturing gel electrophoresis—e.g., sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE; UNIT 10.1). It is important to examine the protein over a range of concentrations—a concentrated sample (to the point of overloading the gel) will provide information on any contaminating proteins that
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are present in low amounts. A more dilute sample, on the other hand, may reveal that the protein is actually two or more proteins of very similar molecular weight arising, for example, from proteolysis. A good target range for purity is ∼95%, although—as in organic chemistry— crystallization itself can be a method of protein purification. For example, the final step in the purification of cytochrome c peroxidase is repeated crystallization by dialysis against water (Fishel et al., 1987) and crystallization of the molybdenum-iron protein is part of the purification protocol for nitrogenase (Burgess et al., 1980). It is often useful to determine the nature of any contaminating proteins. If the contaminants are unrelated proteins, alternate or additional purification steps may be required. However, if the contaminating proteins are smaller fragments of the protein of interest (which may provide useful information regarding domain boundaries), the relative amounts of these can sometimes be manipulated. For example, in the case of recombinant proteins expressed in Escherichia coli that are susceptible to proteolysis during expression, the extent of protein degradation can sometimes be reduced by decreasing the cell density at which cells are induced, changing the concentration of the inducer, or dropping the temperature after induction (e.g., from 37°C to 20°C; also see UNIT 6.1). Although SDS-PAGE is the method of choice to initially characterize protein homogeneity, a single band on a SDS-PAGE gel can be misleading. Heterogeneity can arise from a number of sources other than contaminating proteins, including post-translational modification, deamidation, amino acid side-chain oxidation, or multiple cross-linked species formed by intermolecular or intramolecular disulfide bonds. Advantage should be taken of the several analytical techniques that exist for evaluating protein heterogeneity. Attempts should be made, for example, to analyze the protein by nondenaturing (native) gel electrophoresis (UNIT 10.3) and by isoelectric focusing (IEF; UNIT 10.2). The critical role of information obtained by IEF was recently demonstrated for the complex between the human interferon γ receptor and interferon γ; only those protein fractions selected according to IEF gel criteria of homogeneity ultimately yielded crystals (Windsor et al., 1996). N-terminal sequencing (UNIT 11.10) and mass spectrometry analysis (Chapter 16) are also becoming routine as initial screens of homogeneity. An important consideration to keep in mind throughout crystallization attempts is the redox
state of any cysteine residues in the protein. If cysteine residues are present, it may be necessary to include dithiothreitol (DTT) or 2-mercaptoethanol in the protein buffer because the protein may become heterogeneous as a function of time as the protective reducing agents become oxidized. For example, analytical methods indicated that growth factor receptor– bound protein 2 (Grb2) formed covalent dimers as a result of cysteine oxidation, and it was necessary to include 5 mM DTT in the crystallization buffer to prevent their formation (Guilloteau et al., 1996). It is often useful to perform the initial protein characterization (e.g., SDSPAGE and native gel electrophoresis) under both reducing and nonreducing conditions to determine if there is an effect on heterogeneity. Another form of protein heterogeneity can arise from multiple conformations of the protein in solution. This may be a particular problem for flexible multidomain proteins where the domains may not be held in fixed positions relative to one another. Amount of sample Once it is clear that it is possible to obtain a protein of sufficient purity, the next factor to consider is quantity—i.e., how much protein is needed for initial crystallization trials? An initial screen of 50 different crystallization conditions is often the place to begin, and each condition requires ∼3 µl of protein solution at a reasonable concentration (e.g., 10 mg/ml). This corresponds to 1.5 mg of purified protein and should be considered a bare minimum. A few milligrams of material allow a preliminary look at the protein’s behavior in the presence of precipitating agents, but the first screening attempt is unlikely to produce large regular crystals. Enough material (10 to 20 mg) should be available for a few iteration cycles. The second half of the “quantity” equation is that the 10 or 20 mg of protein must be in concentrated form. Most crystals are obtained using solutions with starting protein concentrations of 10 to 20 mg/ml, although crystals have been grown from solutions as low as 2 to 3 mg/ml (Kühlbrandt, 1988; Gilliland and Bickman, 1990). At the other end of the spectrum, crystals of the hemoprotein domain of P450BM-3 were reportedly grown starting with a protein solution at 275 mg/ml (Boddupalli et al., 1992). It is difficult to know if 10 to 20 mg/ml reflects the optimal condition or simply the most commonly tried condition. Nevertheless, protein solutions in this concentration range appear likely to yield the best results.
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Sample concentration Several methods exist for concentrating protein, most of which are familiar to the biochemist. Cases have even been reported where protein crystals have formed during concentration—for example, tRNA-guanine transglycosylase crystallized during concentration after the final purification step (Romier et al., 1996). A common technique is to remove excess protein buffer by filtration through a membrane (e.g., using ultraconcentrators under gas pressure or small concentration units that operate by centrifugal force; UNIT 4.4 and APPENDIX 3B). However, not all proteins are amenable to this method, as some stick to or aggregate on the membrane surface. The authors have observed that this effect can sometimes be mitigated by changing the temperature at which the concentration is done; for example, certain deletion derivatives of HIV-1 integrase do not concentrate well at 4°C but behave nicely at room temperature. It should be noted that buffer components such as glycerol and detergents often also concentrate above these membranes. To ensure reproducibility, it is best to extensively dialyze a concentrated protein solution containing such additives against a standard buffer, remembering that it may take several days to reach equilibrium. Another useful method of concentrating protein is ammonium sulfate precipitation (UNIT 4.5). A concentrated solution of ammonium sulfate or the salt in solid form is added to the protein solution and the resulting protein precipitate collected by centrifugation. The precipitated protein is then resuspended in a smaller volume of buffer, and it is a matter of taste whether the protein is used “as is” or after removal of the coprecipitating ammonium sulfate. One drawback of this method is that protein recovery may be low if the initial protein concentration is less than ∼1 mg/ml (Scopes, 1982). It may become apparent at this stage that it is not possible to concentrate the protein of interest beyond a certain threshold point or that when the protein is concentrated, it only transiently remains in solution before precipitating out. Protein engineering approaches may help circumvent these problems (see discussion of When All Else Has Failed).
Principles of Macromolecular X-Ray Crystallography
Sample storage The input of the biochemist is particularly important when the method of protein storage is selected because the decision involves not only knowledge of the protein’s biophysical
properties as a function of time but also its catalytic properties if it is an enzyme. Does the protein tolerate freezing at −80°C and subsequent thawing? Alternatively, is it stable for extended periods of time at 4°C? If the protein permits, it is convenient from the standpoint of reproducibility to freeze small aliquots of a concentrated sample that can be thawed as needed. This may require that glycerol or some other cryoprotectant be included in the protein buffer. Glycerol can subsequently be removed by dialysis prior to crystallization attempts, although this may not be necessary as glycerol is sometimes used as an additive in crystallization trials (Sousa, 1995). Storing the protein at 4°C avoids the perhaps traumatic step of freezing, but the possibility of sample variability as a function of time is then introduced. Caution: Affinity-tagged proteins It has become routine to engineer recombinant proteins to possess affinity tags attached to one of the termini; these tags permit rapid and selective purification on affinity columns. Such tags can take the form of another protein such as glutathione-S-transferase (GST; UNIT 6.6) or the maltose-binding protein (MBP), or can be as simple as a stretch of histidine residues that confers specific binding to metal affinity columns (UNIT 9.4). It cannot be assumed that these tags are innocuous. For example, the authors have observed that the presence of a 20–amino acid histidine-containing tag on the catalytic core domain of HIV-1 integrase negatively affects both its solubility and its ability to form crystals (A.B. Hickman and D.R. Davies, unpub. observ.). Similarly, Grishin et al. (1996) report that ornithine decarboxylase only forms crystals when the six histidine residues introduced at the N terminus are removed. Others (e.g., Zhang et al., 1994; Mol et al., 1996; Prodromou et al., 1996) have successfully crystallized proteins that retain tags, but in general these tags are short peptides of six to nine amino acids. As the inclusion of a protease cleavage site between the tag and the protein allows for tag removal during or after purification (Craigie et al., 1995; Grishin et al., 1996; Guilloteau et al., 1996), this precaution should be seriously considered. Exposure of the protein of interest to a protease may bring with it new problems such as proteolysis at internal sites (Guilloteau et al., 1996) or heterogeneity as a result of incomplete removal of the tag. Nevertheless, it may be worth the time invested to surmount these hurdles prior to undertaking crystallization attempts.
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Physical Characterization of Protein Sample
instruments often sell prepacked columns capable of high-resolution separation. Small-bore columns are also available to allow the analysis of very small samples—e.g., 10 to 50 µl. If a sample does contain aggregated material, gel filtration on a preparative scale can be a valuable technique for removing the high-molecular-weight material prior to crystallization trials.
If it has been possible to concentrate the protein to crystallographically useful levels, the next points to address are the multimeric or aggregation state of the protein and the minimal buffer components required to keep the protein soluble. Although these questions will be considered separately, it should be apparent that the results are intertwined; when the protein buffer is modified, the biophysical properties of the protein may also change. Information on the relationship between these two considerations is often invaluable in understanding and interpreting the results of initial crystallization trials. A protein’s multimeric or aggregation state in solution is an important predictor of success in forming crystals. The tendency of a protein to form aggregates or high-molecular-weight clumps is easy to evaluate when these aggregates are insoluble and precipitate out of solution. However, aggregated protein can also remain in solution as soluble high-molecularweight material. If this material is heterogeneous—i.e., the aggregates are irregular in shape and size—it is unlikely that a regular crystal lattice containing reproducible individual units will form. It is crucial to determine if such aggregates are present and, if so, to devise methods to avoid their formation or to remove them prior to crystallization. Useful methods for determining the multimeric state of a protein in solution include gel-filtration chromatography (UNIT 8.3), dynamic light scattering, and analytical ultracentrifugation (UNIT 7.5).
Dynamic light scattering Dynamic light scattering is another method that yields information about protein multimeric state. This technique affords the advantage of doing the analysis at the protein concentration of interest—unlike gel filtration, in which sample dilution is de rigeur (for review see Ferré-d’Anaré and Burley, 1994). In this method, a laser beam scattered by the sample is observed at a 90° scattering angle for a period of several seconds. The effective hydrodynamic radius of the scattering protein molecules in solution can be determined via the autocorrelation function of the scattered light intensity variation during the observation period. If the sample is monodisperse, the observed data can be interpreted in terms of a single hydrodynamic radius and the molecular mass of the scattering species thereby calculated. As an example of the usefulness of light-scattering data, Bentsen et al. (1996) reported that lightscattering measurements on phosphoribosylpyrophosphate synthetase and its complexes showed a direct correlation between polydispersity, the time required for crystals to appear, and ultimate crystal quality.
Gel filtration chromatography Gel filtration is a commonly used chromatographic step during protein purification; it separates biomolecules according to molecular size (for review see Stellwagen, 1990, and UNIT 8.3). The chromatographic support is comprised of porous beads and the diameter of these pores dictates the range of molecular weights over which proteins of different sizes are resolved. Information can be obtained about the protein multimeric state(s) if the column is calibrated using a set of appropriate molecular weight standards, although nonspecific adsorption and protein shape can have unpredictable effects on these results. The extent of aggregation in a sample intended for crystallography experiments can also be readily determined by measuring the proportion of material eluting in the column void volume. Companies that market fast protein liquid chromatography (FPLC)
Analytical ultracentrifugation Analytical ultracentrifugation can be used to derive information about the shape of molecules, molecular masses of species in solution, and thermodynamic parameters relating to their association properties (for reviews see Hansen et al., 1994, and Hensley, 1996). In sedimentation-equilibrium experiments, a protein sample is centrifuged at low speeds such that the rate of transport of molecules by sedimentation is balanced by the rate resulting from diffusion; under these conditions, the distribution of molecules at equilibrium can be measured and molecular masses calculated. Thus, for a wellbehaved protein, it is possible to determine if the protein is present in solution as a monomer or a higher-order multimer and the appropriate association constants can be derived. For example, in the case of the catalytic core domain of HIV-1 integrase with an introduced point
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mutation (F185K), sedimentation-equilibrium studies demonstrated that the protein was a monodisperse dimer over a wide concentration range and therefore a suitable crystallization target (Jenkins et al., 1995). Since the method involves fitting experimental data to an assumed mathematical model, meaningful results can only be obtained when the analysis is performed over a range of conditions to obtain a unique fit to the possible variables. Unfortunately, this has the drawback of being both sample- and time-consuming. Choice of buffer components As indicated earlier, the biophysical state of a protein in solution may be dependent on the properties of the buffer used. As crystallization attempts involve changing the protein environment in a controlled manner, often by the addition of precipitating agents, keeping the concentrations and number of buffer components to a minimum allows the precipitant solution to dominate the properties of the resulting mixture. As most proteins are not soluble or stable in water alone, the usual buffer components that should be evaluated include: the buffering agent itself and the pH to which it is adjusted; protective compounds such as dithiothreitol, 2-mercaptoethanol, or EDTA; salts, if these are needed to keep the protein in solution; and any other agents required to stabilize the protein. A series of experiments to evaluate the effect of removing these components or decreasing their concentration can be performed using a concentrated protein solution with a full complement of buffer components (usually remnants of the last protein purification step), and then removing each by slow dialysis. It should be determined if changes in the buffer cause corresponding changes in the biophysical state of the protein or its homogeneity as defined by the previously described battery of techniques. In such a way, it is possible to define the minimal set of buffer components necessary to keep the protein in a homogeneous state for the expected duration of a crystallographic study.
First Crystallization Attempts
Principles of Macromolecular X-Ray Crystallography
In essence, methods commonly used to coax protein crystals to form involve the very gentle approach to conditions of supersaturation (McPherson, 1985, 1990). In theory, once supersaturation is reached, if conditions are just right, the protein will gradually crystallize out of solution rather than fall out in the undesired form of an amorphous protein precipitate. Slow physical processes that can be exploited to drive
a protein solution to supersaturation include evaporation, vapor diffusion, and dialysis. The vapor-diffusion technique is probably the most widely used method for growing protein crystals. It involves the suspension of a drop of protein solution above a larger volume of a solution of the precipitant in a sealed chamber. Customarily, the droplet, ∼3 µl in volume, is pipetted onto a siliconized coverslip, an equal volume of the well solution added and mixed, and the coverslip inverted and sealed with vacuum grease or oil over the well solution. Equilibration by vapor diffusion of the volatile species results in equalizing the vapor pressure over the well and the drop. This can produce conditions of protein supersaturation in the drop, which sometimes result in crystallization. Typically, the experiment is performed in 24-well tissue culture plates (“crystal trays”); the clear coverslips permit the visualization of the drop contents at regular intervals using a microscope. More detailed descriptions of this method can be found in Ducruix and Giegé (1992) and McPherson (1990). A complete investigation of all the possible variables—e.g., different precipitants, temperatures, pHs, and concentrations—would involve unachievable numbers of experiments. Attempts to reduce this number by factorial procedures have been made (Carter and Carter, 1979), but they have been only partly successful. However, the empirical data obtained from thousands of proteins and protein-nucleic acid complexes have indicated the most popular and successful methods (Gilliland et al., 1994). The commercial availability of screening kits containing likely precipitants for protein crystallization (e.g., from Hampton Research; see SUPPLIERS APPENDIX) significantly reduces the number of solutions to be prepared. Thus, a reasonable place to begin is with a concentrated protein sample and a screening kit containing ∼50 precipitant solutions. Other methods that can be used to screen crystallization conditions (e.g., sitting drops, dialysis, and batch methods) are reviewed in McPherson (1982); gel methods are described in Robert et al. (1992).
Modifying Screening Conditions As it is impossible to predict the rate at which crystals might form, it is important to monitor the processes occurring in the protein drops as equilibrium conditions are approached. The first set of observations should be obtained as soon as the crystal trays are set up. The drops should be carefully examined
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under the microscope. Are the drops clear or have precipitates formed? Is there phase separation of the protein in the form of oil-like droplets? Is there “extraneous material” in the drops that should not be mistaken for crystals at a later date? The crystal trays should be reexamined within the next 24 hr and periodically thereafter. Each drop should be thoroughly examined at the bottom where heavy precipitate (or crystals) may settle, around the edges where equilibration may be the fastest, and against the surface of the coverslip, which can provide nucleation sites for crystals. If crystals do not form under the initial conditions, then careful observations can guide the rational modification of subsequent conditions. What is the relative proportion of drops remaining clear to those with precipitates? A large number of clear drops may suggest that the protein was not concentrated enough or that the amounts of precipitants should be increased. Alternatively, if most conditions give rise to heavy precipitates, the protein concentration should probably be decreased or the well solutions diluted. Extensive protein precipitation can also be caused by the initial two-fold dilution of buffer components upon mixing the protein with the precipitant solutions, if protein solubility is particularly sensitive to these. It should also be considered whether there are general trends across the range of precipitating agents that suggest certain compounds lead to protein precipitation while others tend to keep the protein soluble. Similarly, one should ask whether there is an apparent effect caused by pH or a particular salt. Remembering that the goal is to slowly approach supersaturation and induce the formation of crystals rather than precipitate, it is often clear which precipiting agents will be useful and how their concentration should be adjusted in subsequent crystallization trials. It should be noted, however, that crystals sometimes grow in a drop in which an amorphous protein precipitate initially formed. Furthermore, initial precipitates may disappear over time as equilibrium conditions are reached in the drop. Thus, precipitates are not necessarily a cause for undue concern. Perhaps one of the most difficult assessments in the initial stages of crystallization trials is to judge the nature or promise inherent in a particular precipitate—there are “good” precipitates and then there are “bad” precipitates. Particular attention should be paid to how heavy the precipitate is and its color. It should also be determined if a precipitate is shiny or
appears to have distinct edges, which may be indicative of tiny crystals rather than amorphous precipitate. Unfortunately, not much replaces experience—the investigator’s own, and if available, that of a seasoned crystallographer. In subsequent crystallization experiments in which the amounts or the nature of the components of the precipitant solutions are changed, another possibility is to supplement the well solution with various additives. The composition of the well solution, by virtue of its larger volume, will control the approach to vaporpressure equilibrium. If salts are present in the protein buffer, they can be added to the well solution to control the ultimate volume of the drop. Under certain conditions, salts may also affect the rate of approach to equilibrium (Luft and DeTitta, 1995). It may be desirable to supplement the well with a reducing agent such as dithiothreitol (DTT) or 2-mercaptoethanol if previous experiments have demonstrated that a reducing environment is required to keep the protein in a homogeneous state. Inclusion of such reducing agents can have marked effects on crystallization. For example, the authors have observed an absolute requirement for 5 mM DTT in the crystallization of the core domain of HIV-1 integrase (Dyda et al., 1994); others have reported similar results for other proteins (Guilloteau et al., 1996). Once microcrystals or small crystals are observed, it is usually necessary to optimize conditions to obtain crystals of sufficient size and quality for data collection. This is done by varying such parameters as the concentrations of the precipitant components, the types of components (e.g., the nature of the salts or the molecular weights of polyethylene glycols), the protein concentration, and the pH. Obtaining suitable crystals may also require resorting to other methods such as sitting drops (McPherson, 1990), microseeding, or macroseeding (Stura and Wilson, 1992).
When All Else Has Failed: The Use of Protein Engineering At some point during the isolation or characterization of a particular protein, it may become clear that the protein persists in being insoluble and polydisperse. On the other hand, a protein that appears to fulfill all the criteria for being well-behaved in solution may adamantly refuse to form crystals. One method that has enjoyed some success in circumventing these problems is the use of antibody complexes (for reviews see Laver, 1990, and Kovari et al., 1995). Alternatively, instead of modify-
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ing the protein’s environment, it is possible to modify the protein itself. The authors’ experience with HIV-1 integrase, an essential enzyme in the retrovirus infection cycle, illustrates a number of protein-engineering approaches to modifying a protein in order to obtain a version amenable to concentration and crystallization.
Principles of Macromolecular X-Ray Crystallography
Crystallizing protein domains Wild-type HIV-1 integrase as recombinantly expressed in E. coli is relatively insoluble (∼1 mg/ml in buffers containing 1 M NaCl) and tends to aggregate, forming high-molecularweight material that either elutes in the void volume of a gel-filtration column or precipitates out of solution. A series of deletion mutants, or smaller versions of the protein, was examined to determine if any of these was more soluble that the full-length protein. Limited proteolysis experiments demonstrated that when integrase was exposed to trypsin, chymotrypsin, or V8 protease, it was converted to a series of smaller products as visualized by SDS-PAGE (Engelman and Craigie, 1992). Notably, a prominent protein of about half the molecular weight of the full-length protein was formed, which was resistant to further degradation. When the N-terminal sequence of this smaller protein was determined, it was clear that it represented the middle portion, or core, of the protein from which the two termini had been removed. Such protease-resistant protein fragments may be useful crystallographic targets as they are likely to represent distinct folded domains of the protein that are relatively compact (and hence relatively inaccessible to proteases). The approach of using limited proteolysis and then defining distinct domains by N-terminal sequencing or, more recently, mass spectrometry (reviewed in Cohen, 1996; also see Chapter 16) has been successful in many cases. In the case of HIV-1 integrase, limited protease digestion experiments defined three protein domains: N-terminal, C-terminal, and central domains. All three domains were then separately expressed in E. coli and their solubility examined. While the central domain was no more soluble than the full-length protein, the C-terminal domain by itself was extremely soluble (Engelman et al., 1994) and could be readily concentrated to 20 mg/ml. Thus, the solubility of a full-length protein is not a good predictor of the solubility of its distinct domains. The structure of the C-terminal domain of integrase, which possesses the nonspecific DNA-binding properties of the intact enzyme,
was recently determined by NMR spectroscopy (Eijkelenboom et al., 1995; Lodi et al., 1995). Although probably not generalizable, an interesting example of the proteolysis approach to crystallizable domains was recently reported (Prodromou et al., 1996) in which hanging drops containing yeast Hsp82 chaperone protein were observed to be contaminated with fungus. Crystals of full-length Hsp82 that normally would have been obtained under these conditions did not form. However, a few days later, a new crystal form was observed. The crystals were dissolved, and N-terminal sequencing indicated that proteolysis, presumably by fungal enzymes, had generated a truncated form of the protein consisting of the N-terminal domain, which readily crystallized. The importance of being compact Many proteins that are of interest from a structural point of view are composed of multiple domains. It is becoming clear that when attempting crystallization of an isolated protein domain, ultimate success can be crucially dependent on the residues chosen to define the domain boundaries. Thus, it is fairly common to generate a series of protein constructs of slightly different lengths and to investigate the crystallizability of each protein. The need for such an approach was demonstrated in the cases of the N-terminal region of cartilage oligomeric matrix protein (Efimov et al., 1996), and the transcriptional coactivators dTAF42 and dTAF62 (Xie et al., 1996). Truncation may also be necessary for other reasons. For example, the N terminus of the extracellular domain of the human interferon γ receptor is particularly susceptible to proteolysis (and hence heterogeneity); crystals were only obtained when eight amino acids were removed from the N terminus (Windsor et al., 1996). Mutation of nonessential cysteine residues During isolation of the separately expressed C-terminal domain of HIV-1 integrase, it was observed that concentrated protein precipitated out of solution, an effect readily reversible by the addition of dithiothreitol (DTT). Examination of the amino acid sequence revealed a single cysteine residue, suggesting that intermolecular cross-linking might be responsible for aggregation and subsequent precipitation. Accordingly, the cysteine residue was mutated to serine; the mutated protein remained soluble and no longer aggregated in solution. It was verified that the Cys-to-Ser mutation did not affect the DNA-binding properties of integrase.
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This approach of mutating nonessential cysteine residues to prevent aggregation is generally applicable. For example, the N-terminal domain of the Mu phage transposase contains a cysteine residue that was mutated to leucine prior to structure determination by NMR spectroscopy (Clubb et al., 1996). Site-specific mutagenesis Integrase activity assays demonstrated that the region of structural interest (the enzyme active site) was likely to reside in its central core domain (Bushman et al., 1993). Unfortunately, this domain proved to be as insoluble and as susceptible to aggregation as the fulllength protein. Thus, a novel approach to improving solubility was undertaken (Jenkins et al., 1995). The hydrophobic residues in the central core domain were selected for site-specific mutation according to two criteria—(1) where an isolated hydrophobic residue occurred by itself within the amino acid sequence, it was mutated to a charged lysine residue, and (2) where two or more hydrophobic residues clustered in the primary sequence, they were collectively mutated to alanine residues. Altogether, 29 sets of mutations were introduced by PCR mutagenesis (see Cormack, 1997), the proteins expressed in E. coli, and cell-lysis experiments devised to rapidly screen for improved solubility. Three mutated proteins were identified that were more soluble than the unmutated core domain as assessed by initial solubility after cell lysis, and these were studied further. One of these proteins, in which a single phenylalanine residue was mutated to lysine (F185K), was subsequently demonstrated to be soluble to at least 20 mg/ml, dimeric and monodisperse under all conditions examined, and fully active (Jenkins et al., 1995). The introduction of this mutation was directly responsible for the subsequent crystallization and structure determination of the catalytic core domain of integrase (Dyda et al., 1994). Thus, surprising as it may seem, a single amino acid change can be sufficient to convert a protein that is completely unsuitable for crystallographic attempts to one that readily crystallizes. The problem, of course, is identifying such a mutation. In other successful structure-determination efforts, “magic” mutations have been introduced by accident, as in the case of polymerase chain reaction (PCR) artifacts during cloning of the bacterial chaperonin GroEL (Braig et al., 1994). In other cases, sets of site-directed mutants have been generated for different reasons than those used for HIV-1 integrase. For exam-
ple, to increase protein yield during purification, Staphylococcus aureus dihydrofolate reductase was deliberately engineered to be more soluble; the protein containing two introduced point mutations readily crystallized (Dale et al., 1994). To summarize, protein engineering can be applied to problematic crystallographic targets in a number of ways. Compact domains can be defined by proteolysis and subsequent identification by N-terminal sequencing (UNIT 11.10) or mass spectrometry (Chapter 16). Further trimming of the domain boundaries may be required to yield a tractable protein. Cysteine residues that may cause intermolecular cross-links can be removed. Furthermore, site-specific mutations can be introduced that may dramatically affect a protein’s solubility, multimeric properties, and crystallizability.
LITERATURE CITED Bentsen, A.-K., Larsen, T.A., Kadziola, A., Larsen, S., and Harlow, K.W. 1996. Overexpression of Bacillus subtilis phosphoribosylpyrophosphate synthetase and crystallization and preliminary x-ray characterization of the free enzyme and its substrate-effector complexes. Proteins 24:238246. Blundell, T.L. and Johnson, L.N. 1976. Protein Crystallography. Academic Press, San Diego. Boddupalli, S.S., Hasemann, C.A., Ravichandran, K.G., Lu, J.-Y., Goldsmith, E.J., Deisenhofer, J., and Peterson, J.A. 1992. Crystallization and preliminary x-ray diffraction analysis of P450terp and the hemoprotein domain of P450BM-3, enzymes belonging to two distinct classes of the cytochrome P450 superfamily. Proc. Natl. Acad. Sci. U.S.A. 89:5567-5571. Braig, K., Otwinowski, Z., Hegde, R., Boisvert, D.C., Joachimiak, A., Horwich, A.L., and Sigler, P.B. 1994. The crystal structure of the bacterial chaperonin GroEL at 2.8 Å. Nature 371:578586. Brunger, A.T. 1992. The free R value: A novel statistical quantity for assessing the accuracy of crystal structures. Nature 355:472-474. Burgess, B.K., Jacobs, D.B., and Stiefel, E.I. 1980. Large-scale purification of high activity Azotobacter vinelandii nitrogenase. Biochim. Biophys. Acta 614:196-209. Bushman, F.D., Engelman, A., Palmer, I., Wingfield, P., and Craigie, R. 1993. Domains of the integrase protein of human immunodeficiency virus type 1 responsible for polynucleotidyl transfer and zinc binding. Proc. Natl. Acad. Sci. U.S.A. 90:3428-3432. Carter, C.W. Jr. 1992. Design of crystallization experiments and protocols. In Crystallization of Nucleic Acids and Proteins: A Practical Approach (A. Ducruix and R. Giegé, eds.) pp. 4771. Oxford University Press, New York.
Structural Biology
17.3.13 Current Protocols in Protein Science
Supplement 10
Principles of Macromolecular X-Ray Crystallography
Carter, C.W. Jr., and Carter, C.W. 1979. Protein crystallization using incomplete factorial experiments. J. Biol. Chem. 50:760-763. Clubb, R.T., Mizuuchi, M., Huth, J.R., Omichinski, J.G., Savilahti, H., Mizuuchi, K., Clore, G.M., and Gronenborn, A.M. 1996. The wing of the enhancer-binding domain of Mu phage transposase is flexible and is essential for efficient transposition. Proc. Natl. Acad. Sci. U.S.A. 93:1146-1150. Cohen, S.L. 1996. Domain elucidation by mass spectrometry. Structure 4:1013-1016. Collaborative Computer Project No. 4. (CCP-4). 1994. Acta Cryst. Sect. A 50:760-763. Cormack, B. 1997. Directed mutagenesis using the polymerase chain reaction. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 8.5.1-8.5.10. John Wiley & Sons, New York. Craigie, R., Hickman, A.B., and Engelman, A. 1995. Integrase. In HIV: A Practical Approach, Vol. 2 (J. Karn, ed.) pp. 53-71. Oxford University Press, Oxford. Dale, G.E., Broger, C., Langen, H., D’Arcy, A., and Stüber, D. 1994. Improving protein solubility through rationally designed amino acid replacements: Solubilization of the trimethoprim-resistant type S1 dihydrofolate reductase. Protein Eng. 7:933-939. Drenth, J. 1994. Principles of Protein X-Ray Crystallography. Springer-Verlag, New York. Ducruix, A. and Giegé, R. 1992. Methods of crystallization. In Crystallization of Nucleic Acids and Proteins: A Practical Approach (A. Ducruix and R. Giegé, eds.) pp. 73-98. Oxford University Press, New York. Dyda, F., Hickman, A.B., Jenkins, T.M., Engelman, A., Craigie, R., and Davies, D.R. 1994. Crystal structure of the catalytic domain of HIV-1 integrase: Similarity to other polynucleotidyl transferases. Science 266:1981-1986. Efimov, V.P., Engel, J., and Malashkevich, V.N. 1996. Crystallization and preliminary crystallographic study of the pentamerizing domain from cartilage oligomeric matrix protein: A fivestranded α-helical bundle. Proteins 24:259-262. Eijkelenboom, A.P.A.M., Puras-Lutzke, R.A., Boelens, R., Plasterk, R.H.A., Kaptein, R., and Hard, K. 1995. The DNA-binding domain of HIV-1 integrase has an SH3-like fold. Nature Struct. Biol. 2:807-810. Engelman, A. and Craigie, R. 1992. Identification of conserved amino acid residues critical for human immunodeficiency virus type 1 integrase function in vitro. J. Virol. 66:6361-6369. Engelman, A., Hickman, A.B., and Craigie, R. 1994. The core and carboxyl-terminal domains of the integrase protein of human immunodeficiency virus type 1 each contribute to nonspecific DNA binding. J. Virol. 68:5911-5917. Ferré-d’Anaré, A.R. and Burley, S. 1994. Use of dynamic light scattering to assess crystalliz-
ability of macromolecules and macromolecular assemblies. Structure 2:357-359. Fishel, L.A., Villafranca, J.E., Mauro, J.M., and Kraut, J. 1987. Yeast cytochrome c peroxidase: Mutagenesis and expression in Escherichia coli show tryptophan-51 is not the radical site in compound I. Biochemistry 26:351-360. Fitzgerald, P.M.D. 1988. MERLOT, an integrated package of computer programs for the determination of crystal structures by molecular replacement. J. Appl. Crystallogr. 2:273-278. Gilliland, G.L. and Bickman, D.M. 1990. The Biological Macromolecule Crystallization Database: A tool for developing crystallization strategies. Methods 1:6-11. Gilliland, G.L. and Ladner, J.E. 1996. Crystallization of biological macromolecules for X-ray diffraction studies. Curr. Opin. Struct. Biol. 6:595-603. Gilliland, G.L., Tung, M., Blakeslee, D.M., and Ladner, J.E. 1994. Biological Macromolecule Crystallization Database, Version 3.0: New features, data and the NASA archive for protein crystal growth data. Acta Crystallogr. Sect. D 50:408-413. Green, D.W., Ingram, V.M., and Perutz, M.F. 1954. The structure determination of haemoglobin. IV. Sign determination by the isomorphous replacement method. Proc. R. Soc. Lond. A225:287307. Grishin, N.V., Osterman, A.L., Goldsmith, E.J., and Phillips, M.A. 1996. Crystallization and preliminary X-ray studies of ornithine decarboxylase from Trypanosoma brucei. Proteins 24:272-273. Guilloteau, J.P., Fromage, N., Ries-Kautt, M., Reboul, S., Bocquet, D., Dubois, H., Faucher, D., Colonna, C., Ducruix, A., and Becquart, J. 1996. Purification, stabilization, and crystallization of a modular protein: Grb2. Proteins 25:112-119. Hansen, J.C., Lebowitz, J., and Demeler, B. 1994. Analytical ultracentrifugation of complex m ac ro m o l ec u l ar s y ste ms. Biochemistry 33:13155-13163. Hensley, P. 1996. Defining the structure and stability of macromolecular assemblies in solution: The re-emergence of analytical ultracentrifugation as a practical tool. Structure 4:367-373. Jenkins, T.M., Hickman, A.B., Dyda, F., Ghirlando, R., Davies, D.R., and Craigie, R. 1995. Catalytic domain of human immunodeficiency virus type 1 integrase: Identification of a soluble mutant by systematic replacement of hydrophobic residues. Proc. Natl. Acad. Sci. U.S.A. 92:60576061. Jones, T.A., Zhou, J.Y., Cowan, S.W., and Kjeldgaard, M. 1991. Improved methods for the building of protein models in electron density maps and the location of errors in these models. Acta Crystallogr. Sect. A 47:110-119. Jones, C., Mulloy, B., and Sanderson, M.R. (eds.) 1996. Crystallographic Methods and Protocols. In Methods in Molecular Biology, Vol. 56. Humana Press, Totowa, N.J.
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Current Protocols in Protein Science
Kovari, L.C., Momany, C., and Rossman, M.G. 1995. The use of antibody fragments for crystallization and structure determinations. Structure 3:1291-1293. Kühlbrandt, W. 1988. Three-dimensional crystallization of membrane proteins. Q. Rev. Biophys. 21:429-477. Laver, W.G. 1990. Crystallization of antibody-protein complexes. Methods 1:70-74. Lodi, P.J., Ernst, J.A., Kuszewski, J., Hickman, A.B., Engelman, A., Craigie, R., Clore, G.M., and Gronenborn, A.M. 1995. Solution structure of the DNA binding domain of HIV-1 integrase. Biochemistry 34:9826-9833. Luft, J.R. and DeTitta, G.T. 1995. Chaperone salts, polyethylene glycol and rates of equilibration in vapor-diffusion crystallization. Acta Crystallogr. Sect. D 51:780-785. McPherson, A. 1982. Preparation and Analysis of Protein Crystals. John Wiley & Sons, New York. McPherson, A. 1985. Crystallization of macromolecules: General principles. Methods Enzymol. 114:112-120. McPherson, A. 1990. Current approaches to macromolecular crystallization. Eur. J. Biochem. 189:1-23. McRee, D.E. 1993. Practical Protein Crystallography. Academic Press, New York. Mol, C.D., Harris, J.M., McIntosh, E.M., and Tainer, J.A. 1996. Human dUTP pyrophosphatase: Uracil recognition by a β hairpin and active sites formed by three separate subunits. Structure 4:1077-1092. Navaza, J., 1994. AMORE: An automated package for molecular replacement. Acta Crystallogr. Sect. A 50:157-163. Prodromou, C., Piper, P.W., and Pearl, L.H. 1996. Expression and crystallization of the yeast Hsp82 chaperone, and preliminary X-ray diffraction studies of the amino-terminal domain. Proteins 25:517-522. Ramachandran, G.N., Ramakrishnan, C., and Sasisekjaran, V. 1963. Stereochemistry of polypeptide chain configuration. J. Mol. Biol. 7:9599. Rhodes, C. 1993. Crystallography Made Crystal Clear: A Guide For Users Of Macromolecular Models. Academic Press, New York. Robert, M.C., Provost, K., and Lefaucheux, F. 1992. Crystallization in gels and related methods. In Crystallization of Nucleic Acids and Proteins: A Practical Approach (A. Ducruix and R. Giegé, eds.) pp.127-143. Oxford University Press, New York. Romier, C., Ficner, R., Reuter, K., and Suck, D. 1996. Purification, crystallization, and preliminary X-ray diffraction studies of tRNA-guanine transglycosylase from Zymomonas mobilis. Proteins 24:516-519. Sasisekharan, V. 1962. Stereochemical criteria for polypeptide and protein structures. In Collagen (N. Ramanathan, ed.) pp. 39-78. John Wiley & Sons, New York.
Scopes, R. 1982. Protein Purification: Principles and Practice, p. 52. Springer-Verlag, New York. Sousa, R. 1995. The use of glycerol, polyols, and other protein structure stabilizing agents in protein crystallization. Acta Crystallogr. Sect. D 51:271-277. Stellwagen, E. 1990. Gel filtration. Methods Enzymol. 182:317-328. Stura, E.A. and Wilson, I.A. 1992. Seeding techniques. In Crystallization of Nucleic Acids and Proteins: A Practical Approach (A. Ducruix and R. Giegé, eds.) pp. 99-126. Oxford University Press, New York. Sweet, R.M. (ed.) 1997. Macromolecular Crystallography. Methods Enzymol. Vol. 276. Tickle, I.J. and Driessen, H.P. 1996. Molecular replacement using known structural information. In Methods in Molecular Biology (J.M. Walker, ed.), Vol. 56, pp. 173-203. Humana Press, Totowa, N.J. Weber, P.C. 1991. Physical principles of protein crystallization. Adv. Prot. Chem. 41:1-36. Westhof, E. and Dumas, P. 1996. Refinement of protein and nucleic acid structures. In Methods in Molecular Biology (J.M. Walker, ed.), Vol. 56, pp. 227-244. Humana Press, Totowa, N.J. Windsor, W.T., Walter, L.J., Syto, R., Fossetta, J., Cook, W.J., Nagabhushan, T.L., and Walter, M.R. 1996. Purification and crystallization of a complex between human interferon γ receptor (extracellular domain) and human interferon γ. Proteins 26:108-114. Wyckoff, H.W., Hirs, C.H.W., and Timasheff, S.G. (eds.) 1985. Diffraction methods for biological molecules (Parts A and B). Methods Enzymol. Vols. 114 and 115. Xie, X., Kokubo, T., Cohen, S.L., Mirza, U.A., Hoffmann, A., Chait, B.T., Roeder, R.G., Nakatani, Y., and Burley, S.K. 1996. Structural similarity between TAFs and the heterotetrameric core of the histone octamer. Nature 380:316-322. Yang, W., Hendrickson,W.A., Crouch, R.J., and Satow, Y. 1990. Structure of ribonuclease H phased at 2 Å by MAD analysis of the selenomethionine protein. Science 249:1398-1403. Zhang, F., Strand, A., Robbins, D., Cobb, M.H., and Goldsmith, E.J. 1994. Atomic structure of the MAP kinase ERK2 at 2.3 Å resolution. Nature 367:704-711.
INTERNET RESOURCES www.hamptonresearch.com Hampton Research Home Page; discusses aspects of protein crystallography; contains links to other Web sites.
Contributed by Alison B. Hickman and David R. Davies National Institute of Diabetes and Digestive and Kidney Diseases Bethesda, Maryland Structural Biology
17.3.15 Current Protocols in Protein Science
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Crystallization of Macromolecules
UNIT 17.4
X-ray crystallography has evolved into a very powerful tool for determining the three-dimensional structure of macromolecules and macromolecular complexes. Developments in computational methodologies, crystal data-collection procedures, and recombinant DNA technology have resulted in a near exponential pace for structure determinations over the last century. The major bottleneck in structure determination by X-ray crystallography is the preparation of suitable crystalline samples. Advances in procedures for macromolecular crystallization, together with the availability of commercial screening kits, have made it relatively straightforward to set up screens for crystallization in a conventional molecular biology or biochemistry laboratory. The following unit outlines the steps required for the crystallization of a macromolecule starting with a purified sample. The starting macromolecule must be purified and homogeneous (≥95% pure by SDS-PAGE; UNIT 10.1) but can be at a fairly dilute concentration (≤1 µg/ml). The macromolecule should be in a standard nonphosphate buffer between pH 5.5 and 8.0, with a fairly low salt concentration (preferably <100 mM NaCl, and no higher than 250 mM). A reducing agent, either 2-mercaptoethanol (2-ME) or dithiothreitol (DTT), should also be present at a concentration between 1 and 10 mM. The total amount of macromolecule is critical, and should be at least 2 mg, preferably >5 mg. The first protocols describe the preparation of the macromolecular sample—i.e., proteins (see Basic Protocol 1), nucleic acids (see Alternate Protocol 1), macromolecular complexes (see Alternate Protocol 2), and membrane proteins (see Alternate Protocol 3). The preparation and assessment of crystallization trials is then described (see Basic Protocol 2), along with a protocol for determining whether the crystals obtained are composed of the macromolecule of interest or just salt (see Support Protocols 1 and 2). Next, the optimization of crystallization conditions is presented (see Basic Protocol 3 and Alternate Protocol 4). Finally, protocols that facilitate the growth of larger crystals through seeding are described (see Alternate Protocols 5 and 6). PREPARATION OF PROTEIN FOR CRYSTALLIZATION Before a protein can be crystallized, it needs to be concentrated to at least 5 mg/ml. An ideal concentration is 20 mg/ml; however, somewhere between 5 and 20 mg/ml is usually suitable.
BASIC PROTOCOL 1
Materials Soluble protein sample 15 ml to 500 µl and 2 ml to ∼50 µl Centriprep and Centricon microconcentrators (Amicon) 1.5-ml screw-cap microcentrifuge tubes Additional reagents and equipment for measurement of dynamic light scattering (UNIT 7.8; optional) 1. Place 15 ml soluble protein sample in a Centriprep microconcentrator that concentrates 15 ml to ∼500 µl. Centrifuge according to manufacturer’s instructions. 2. Transfer the sample to a Centricon microconcentrator that concentrates 2 ml to ∼50 µl, and continue centrifugation according to manufacturer’s instructions. Structural Biology Contributed by Troy Messick and Ronen Marmorstein Current Protocols in Protein Science (2003) 17.4.1-17.4.25 Copyright © 2003 by John Wiley & Sons, Inc.
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3. Monitor concentration C (in M) of an appropriately diluted sample (i.e., that produces an absorbance between 0.1 and 1.0) by determining A280 with a UV spectrophotometer and applying the Beer-Lambert law: C = A280/ εl where l is the path length (cm) and ε , the molar absorption coefficient (M−1cm −1), is calculated as follows: ε = (5600 × no. Trp) + (1420 × no. Tyr) + (197 × no. Phe). For proteins that do not contain Trp or Tyr residues, a colorimetric assay such as the Bradford (UNIT 3.4) can be used.
4. Continue to concentrate the protein until it is 5 mg/ml. If this is not attainable, concentrate the protein until visible precipitate is observed. If concentration of the protein sample leads to formation of precipitate, the remaining soluble portion should be isolated by centrifugation. Reconcentration of the soluble portion should then be attempted under different salt concentrations, pHs, or concentrations of other appropriate additives.
5. If a dynamic light scattering instrument is available (e.g., Protein Solutions model DynaProMS/X, DynaPro-801), check the protein solution for monodispersity (UNIT 7.8). Generally, proteins that are monodisperse at high concentrations (typically a polydispersity value ≤15% of the Stokes radius) have a strong tendency to crystallize, and those that are polydisperse have an equally strong tendency not to crystallize (Ferre-D’Amare and Burley, 1994). However, there are exceptions to this rule, and crystallization attempts should not be abandoned solely because of poor dynamic light scattering measurements.
6. Divide into 20- to 50-µl aliquots in 1.5-ml screw-cap microcentrifuge tubes, and flash freeze in liquid nitrogen. Store frozen (preferably at −70°C) until use for crystallization (up to 1 month in most cases). If possible, subject part of the concentrated protein sample to crystallization trials (see Basic Protocol 2) prior to freezing, as some proteins do not respond well to freeze-thawing. If there is an activity assay available for the protein, it is recommended that the effect of freeze-thawing on this activity be established before freezing the protein sample. If the protein responds adversely, try to freeze the protein sample in the presence of a small amount of glycerol (5% to 15%) or another cryoprotectant. In such cases, it is usually necessary to remove the cryoprotectant (preferably with a microconcentrator) prior to crystallization. ALTERNATE PROTOCOL 1
PREPARATION OF NUCLEIC ACID FRAGMENTS FOR CRYSTALLIZATION Before performing this protocol, the oligonucleotide(s) of interest (i.e., DNA or RNA) should be purified (preferably by reversed-phase HPLC; UNIT 11.6), and usually diluted (i.e., 1 mg/ml). The total amount of nucleic acid should be at least 0.5 mg, preferably >1.5 mg. This usually requires solid-phase synthesis at a phosphoramidite scale of 1 µmol or greater. Materials DNA or RNA sample, purified (e.g., UNIT 11.6) 10 mM sodium cacodylate (pH 6.8)/50 mM NaCl
Crystallization of Macromolecules
2000-MWCO dialysis tubing (i.e., Spectrum Spectra/Por 7) 50-ml polypropylene centrifuge tubes (Falcon or Corning) Hypodermic needle 500-ml lyophilization vessel
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Current Protocols in Protein Science
1.5-ml screw-cap microcentrifuge tubes Additional reagents and equipment for dialysis (UNIT 4.4 & APPENDIX 3B) 1. Transfer a volume of purified nucleic acid sample containing ≥0.5 mg oligonucleotide (preferably ≥1.5 mg) to 2000-MWCO dialysis tubing and dialyze against a total of 200 vol. water at least 4 hr at 4°C (UNIT 4.4 & APPENDIX 3B). 2. Transfer dialyzed material to a 50-ml polypropylene centrifuge tube, cap, and shell freeze the sample (i.e., freeze it onto the sides of the tube) by immersing in crushed dry ice. 3. Punch holes in the tube cap using a hypodermic needle, place the tube in a 500-ml lyophilization vessel, and lyophilize the sample to dryness. 4. Dissolve the lyophilized material with 1 ml water and transfer to a 1.5-ml screw-cap microcentrifuge tube. 5. Refreeze the oligonucleotide solution on crushed dry ice. Punch small holes in the top of the microcentrifuge tube using a hypodermic needle, place the tube in a 500-ml lyophilization vessel, and lyophilize to dryness. 6. Dissolve the dried oligonucleotide in 30 µl of 10 mM sodium cacodylate (pH 6.8)/50 mM NaCl. 7. Determine the nucleic acid concentration C (M) by measuring A260 and converting using the Beer-Lambert equation: C = A260/ εl where l is the path length (cm), and ε, the molar absorption coefficient (M−1cm−1), is calculated as follows: ε = (15,200 × no. dATPs) + (9300 × no. dCTPs) + (13,700 × no. dGTPs) + (9600 × no. dTTP) + (9600 × no. dUTPs). For an oligonucleotide that has been synthesized on a 1-ìmol phosphoramidite scale, this procedure should yield an oligonucleotide concentration of ∼0.8 to 1.5 mM from a minimum of 0.5 mg starting material (step 1).
Form duplexes (optional) 8. Mix complementary strands in stoichiometric molar amounts in a 1.5-ml screw-cap microcentrifuge tube. Tightly seal the microcentrifuge tube. 9. Place the tube in a room-temperature water bath (i.e., beaker) and heat the sample and bath to 75°C on a hot plate for 5 min. 10. Remove the water bath from the hot plate and cool to 4°C over ∼2 hr by allowing the water bath to come to room temperature gradually. 11. Slowly add ice to the water bath until the temperature reaches 10°C. Store the renatured duplex at 4°C until use for crystallization (up to 2 weeks). For prolonged storage (>2 weeks), store the sample frozen (preferable at −70°C) and renature again prior to use. The sample may be stored for up to 1 yr.
Structural Biology
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ALTERNATE PROTOCOL 2
PREPARATION OF MACROMOLECULAR COMPLEXES FOR CRYSTALLIZATION There are two types of macromolecular complexes that are generally prepared for crystallization: protein–nucleic acid and protein-protein complexes. For protein–nucleic acid complexes, the concentrated macromolecules (see Basic Protocol 1 and Alternate Protocol 1) are mixed in near stoichiometric amounts. It has been empirically determined that a slight excess of nucleic acid over protein (∼20%; Aggarwal, 1990) is best. In general, one should aim for 0.8 mM complex. Some protein-nucleic acid complexes form precipitate when mixed at the high concentrations required for crystallization. There are two common solutions to this problem: (1) lower the concentration of complex 2- to 3-fold, or (2) add monovalent or divalent metal ions at increasing concentrations. Generally, one should first try adding monovalent ions at a concentration of 25 mM, and increase slowly to a maximum of 200 mM if the precipitate persists. For divalent ions, one should start at 10 mM increasing to a maximum of 75 mM. NaCl is the most popular monovalent ion, while MgCl2 and CaCl2 are the most commonly used divalent ions. Although divalent ions are often effective in solubilizing some protein-nucleic acid complexes, they should only be used as a last resort, because they may give rise to salt crystals (this typically occurs under crystallization conditions that contain sulfate or phosphate counterions), which may be mistaken for macromolecular crystals (to determine if this is the case, see Support Protocols 1 and 2). Homogeneous protein-protein complexes are generally prepared by isolating the desired complex by gel filtration prior to concentration for crystallization. Multiprotein-nucleic acid complexes can be prepared either by mixing the preformed concentrated protein complex with the concentrated nucleic acid sample in appropriate stoichiometric amounts, or by isolating the multiprotein-nucleic acid complex by gel filtration (e.g., UNIT 8.3) prior to concentration for crystallization.
ALTERNATE PROTOCOL 3
PREPARATION OF MEMBRANE PROTEINS FOR CRYSTALLIZATION Membrane proteins represent by far the most underrepresented class of high-resolution structures currently available in the protein database (PDB). This is attributable to the increased difficulties in the overexpression, solubilization, and crystallization of membrane proteins compared to those that are water-soluble. Overexpression It is particularly difficult to obtain milligram quantities of membrane proteins due to several factors, including a dearth of protein expression systems, and a propensity for inclusion body formation and proteolysis. The most effective host for expression is still E. coli , although the strain selection—i.e., BL21(DE3) pLysS or C43(DE3)—can have a significant impact on the amount of protein expression. Also, an expression vector should be chosen which prevents leaky expression during cell growth and allows manipulation of protein expression levels in an inducer concentration–dependent manner; leaky expression of these often-toxic proteins may result in proteolysis or cell death and high expression levels may result in inclusion body formation. The pBAD-based system (Invitrogen), which uses the tightly regulated ara BAD promoter and allows expression levels to be modulated by using between 0.00002% to 0.2% of the arabinose inducer, satisfies these concerns. Growth and expression conditions that need to be optimized include type of culture medium, OD at induction, growth and induction temperatures, and induction time.
Crystallization of Macromolecules
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Solubilization The main obstacle in solubilizing a membrane protein is releasing the extensive hydrophobic surfaces of the protein from its interaction with the surrounding lipids. A membrane-spanning protein, for example, must be released from the encompassing lipid bilayer using a detergent. The hydrophobic tail of the detergent molecule binds to the hydrophobic portion of the protein surfaces, while the hydrophilic head of the detergent interacts with the aqueous solvent. During the protein solubilization process, however, many of the hydrophobic surfaces of the protein can be exposed to other protein monomers, leading to aggregation. Thus, detergents must readily bind the surfaces of monomers, preventing aggregation without simultaneously denaturing the protein. Another complication commonly encountered during the preparation of membrane proteins is the heterogeneity of the sample, which is antithetical to forming uniform crystallization contacts. Specifically, the protein sample may contain pure detergent micelles, with a variety in the number and types of interactions of the detergent molecules with the hydrophobic surfaces. It is desirable to purify one species of protein monomers having specific interactions with detergent molecules in order to obtain a homogeneous monodispersed protein solution. A variety of detergents have been employed to facilitate the solubilization of membrane proteins. As with crystallization conditions, it is difficult to determine a priori which detergents will provide better protein solubilization, and a trial-and-error approach is often employed. However, OG (octyl-β-D-glucopyranoside) and LDAO (N,N-dimethyldodecylamine-N-oxide) appear to be the most successful detergents used. (For an updated list of solved membrane protein structures and the crystallization conditions and references, see http://www.mpibp-frankfurt.mpg.de/michel/public/memprotstruct.html). As with water-soluble proteins, care must be taken to prevent the aggregation of the protein sample. Dynamic Light Scattering (DLS) provides a method for ascertaining the aggregation state of a protein (UNIT 7.8). However, the DLS measurements are very sensitive to large contaminants, such as large micelles, and may overwhelm the signal from monomer proteins. In the case of membrane proteins, analytical ultracentrifugation (UNIT 7.5) and electron microscopy (UNIT 17.2) may provide better measures of the homogeneity and monodispersity than DLS. Ultracentrifugation, varying salt concentration, and changing the detergent can sometimes improve the monodispersity of the sample. The target concentration of membrane proteins for crystallization trials are typically higher (10 to 30 mg/ml) than for water soluble proteins (5 to 20 mg/ml). Normally, the same detergent used in solubilizing the protein may be added in the crystallization trials, however, a second detergent has been useful in a number of cases. Again, the ideal concentration of detergent must be empirically determined, however, a good starting concentration is one to three times the critical micelle concentration (CMC). Also, it is often useful to add a small molecule amphiphile for crystallization. Amphiphiles that have been the most successful include heptane-1,2,3-triol and benzamidine. Refer to http://www.mpibp-frankfurt.mpg.de/michel/public/memprotstruct.html for a list of second detergents and amphiphiles. PREPARATION AND ANALYSIS OF CRYSTALLIZATION TRIALS UNIT 17.3 discusses parameters that effect the crystallization of soluble macromolecules. Overall, the most important parameter in obtaining well-ordered macromolecular crystals is the actual macromolecular sample (see Critical Parameters and Troubleshooting). Purity, protein monodispersity, and stability (i.e., remains intact) are the most critical factors in the ability of a protein sample to crystallize. An overloaded PAGE gel (UNIT 10.1) should be run to check for contaminants. Ideally, the protein sample should be >99% pure. Gel filtration (UNIT 8.3) or dynamic light scattering (DLS; UNIT 7.8) are suitable methods
BASIC PROTOCOL 2
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for assessing protein monodispersity. In the case of protein expression in eukaryotic expression systems, mass spectrometry (Chapter 16) of the sample can be used to check for post-translational modification of the protein, which may adversely effect crystallization properties. Primarily three components make up any crystallization solution: salt, buffer, and precipitating agent. Because there is an infinite number of combinations of these parameters (McPherson, 1982), a complete, methodical search for the crystallization conditions that give rise to macromolecular crystals would take an enormous amount of time and material. There are basically two approaches for pursuing crystallization conditions of a macromolecule which has never previously been crystallized: screening matrix and a systematic approach. The most popular approach is the screening matrix, which is employed to quickly determine which variables influence the behavior of the protein solution. This so-called shotgun approach utilizes a rather large number of wide-ranging conditions unrelated to one another. The goal of this procedure is to narrow the conditions that yield macromolecular crystals. One would typically refine the components of the successful crystallization condition(s) by separately varying the concentration of the components. (see Basic Protocol 3.) The first widely popular screen, described by Jancarik and Kim in 1991, was heavily biased towards previously published crystallization conditions (Table 17.4.1). In 1992, the Jancarik and Kim screen became commercially available as “Crystal Screen” from Hampton Research. Since then, there has been a proliferation in the number of companies that provide sparse matrix solutions for performing initial screenings (Table 17.4.2). There has also been an expansion in the number of different crystallization screens, many dedicated to certain types of macromolecules. For instance, Natrix (Hampton) is designed for nucleic acids or nucleic acid–protein complexes and MemStart (Molecular Dimensions) screens are suitable for membrane proteins. It should be remembered that a sparse matrix screen which does not yield promising crystals does not mean that a particular macromolecule will not crystallize. For example, lysozyme, canavalin, and many immunoglobulins will not crystallize using the commercially available kits.
Table 17.4.1 Crystallization Matrix Components Used in Crystal Screen 1 (Hampton Research) to Crystallize Proteins
Crystallization of Macromolecules
Precipitant
Salt
Buffer
Ammonium phosphate Ammonium sulfate Lithium sulfate Magnesium formate MPDa PEGb (400, 1500, 4000, 8000) Potassium phosphate Potassium tartrate 2-Propanol Sodium acetate Sodium citrate Sodium formate Sodium phosphate Sodium tartrate
Ammonium acetate Ammonium sulfate Calcium acetate Calcium chloride Lithium sulfate Magnesium acetate Magnesium chloride Sodium acetate Sodium citrate Zinc acetate
Imidazole, pH 6.5 Sodium acetate, pH 4.6 Sodium citrate, pH 5.6 Sodium cacodylate, pH 6.5 Sodium HEPES, pH 7.5 Tris⋅Cl, pH 8.5
a MPD, 2-methyl-2,4-pentanediol. b PEG, polyethylene glycol.
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Table 17.4.2
Commercially Available Crystallization Screens
Screens DeCODE
Comment Geneticsa
Wizard I Wizard II Cryo I Cryo II Peg/Ion Kinase screen
Wim Hol screen Wim Hol screen Wizard I with cryoprotectant Wizard II with cryoprotectant Various PEGs/various salts Screen designed for kinases
Hampton Researchb Crystal Screen Crystal Screen 2 Crystal Screen Cryo Crystal Screen Lite Natrix Nucleic Acid Mini Screen MembFac Index Salt Rx Grid Screen ammonium sulfate Grid Screen PEG 6000 Grid Screen MPD Grid Screen PEG/LiCl Grid Screen sodium chloride Quick Screen Low Ionic Strength Screen
Original Jancarik & Kim Screen Extension of original Jancarik & Kim screen Crystal Screen with glycerol added Crystal Screen with lower precipitating agent Screen for nucleic acids or protein–nucleic acid complexes Screen for nucleic acids, Various salts/pH screen Screen for membrane proteins Combines classic, contemporary, new conditions Various salts/pH screen Ammonium sulfate/pH screen PEG 6000/pH screen MPD/pH screen PEG 6000/pH screen Sodium chloride/pH screen Sodium potassium phosphate/pH screen PEG 3350/pH screen, special protocol is required
Jena Bioscienced JB Screen 1 JB Screen 2 JB Screen 3 JB Screen 4 JB Screen 5 JB Screen 6 JB Screen 7 JB Screen 8 JB Screen 9 JB Screen 10
PEG 400 to 3000 based PEG 4000 based PEG 4000 plus based PEG 5000 MME to 8000 based PEG 8000 to 20000 based Ammonium sulfate based MPD based MPD and alcohol based Alcohol and salt based Salt based
Molecular Dimensionsc Structure Screen 1 Structure Screen 2 3D Structure Screen Clear Strategy Screen I Clear Strategy Screen II ZetaSol Stura Footprint Screens MacroSol
Same as Hampton Crystal Screen 2, different order plus 2 conditions Same as Hampton Crystal Screen, different order 24 nucleation conditions, 24 crystal growth conditions Brzozowski and Walton screen; various PEGs/various salts screen; pH buffer provided by user Extension of Clear Strategy Screen 1 (same principles) Riès-Kautt & Ducruix physico-chemical approach Enrico Stura Footprint Screen intended for glycoproteins or protein complexes continued
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Table 17.4.2
Commercially Available Crystallization Screens, continued
Screens
Comment
MemStart NR-LBD
Some similarities to Hampton MembFac Screen Screen based on successful crystallization of nuclear receptor-ligand binding domains
Sigma-Aldriche Basic kit Extension kit Cryo kit Membrane Proteins kit Low Ionic kit
Same as Hampton Research Crystal Screen Same as Hampton Research Crystal Screen 2 Same as Hampton Research Crystal Screen Cryo Same as Hampton Research MembFac Screen PEG 3350/pH screen
Nextal Biotechnologiesf The Classics
Same as Hampton Research Crystal Screen and Crystal Screen 2
aRefer to http://decode.com/emeraldbiostructures for more details. Note that this supplier was formerly called
Emerald Biostructures. bRefer to http://www.hamptonresearch.com for more details. cRefer to http://www.moleculardimensions.com for more details. dRefer to http://www.jenabioscience.com for more details. eRefer to http://www.sigmaaldrich.com for more details. fRefer to http://www.nextalbiotech.com for more details.
The second method for macromolecular crystallization involves systematically obtaining information about the solubility of the particular macromolecule (Stura, 1994). This is done by using a matrix containing increasing concentrations of precipitating agent along one axis versus pH along the other. In this way, the solubility curve of the protein can be determined, along with the optimum pH or type or concentration of precipitating agent. The observation of dull or fibrous precipitate may permit the elimination of a particular pH or precipitating agent. On the other hand, observing a shiny precipitate or one which appears colorful (birefringence) when viewed through a polarizer may warrant further investigation (see Basic Protocol 3). Subsequent crystallization trials may then focus on the introduction of additives such as salt, alcohol (ethanol), or MPD additives to promote crystallization of the protein. This approach involves repeated and methodical probing of the solubility curve of the macromolecule until crystals are obtained. A summary of the different screens and their applications is given in Table 17.4.2. Provided that the macromolecular sample is able to be crystallized, these screens will generally yield directions in which to proceed in order to obtain suitable crystals for structural studies, if not diffraction-quality crystals themselves. Although each of the commercially available premixed solutions can be prepared in the laboratory, it is convenient, at least initially, to obtain the premixed solutions. Any reagents that are not obtained from vendors should generally be of the highest-quality grade. This is especially true for PEG. This protocol focuses only on screens that are used most often for crystallizing soluble proteins, nucleic acids, macromolecular complexes, and membrane proteins.
Crystallization of Macromolecules
There are several different crystallization techniques that are used for macromolecules (Weber, 1997). Only the one most common, the vapor-diffusion technique using hanging drops (Fig. 17.4.1), is discussed here. Steps 1 to 9 should be carried out both at room temperature and 4°C. Both the Jancarik and Kim sparse crystallization screen (Crystal Screen) and the extended sparse crystallization screen (Crystal Screen 2 or its equivalent
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macromolecule solution coverslip vacuum grease crystallization chamber
reservoir solution
Figure 17.4.1 Typical vapor-diffusion crystallization setup.
from another vendor) should be employed for optimal results with proteins, although at a minimum the sparse matrix screen can be used alone. The nucleic acid sparse matrix (Natrix) should be used for nucleic acids or protein–nucleic acid complexes. Crystallization droplets will typically show one of three results: clear drops, crystals, or precipitate/phase separation (see Anticipated Results). Materials Vacuum grease (Hampton Research) Crystallization screening kit(s) (Hampton Research): Proteins and multiprotein complexes: Crystal Screens 1 (Table 17.4.3) and 2 (latter is optional) Nucleic acids and protein–nucleic acid complexes: Natrix (Table 17.4.4) Concentrated macromolecule sample: protein (see Basic Protocol 1), nucleic acid (see Alternate Protocol 1), or macromolecular complex (see Alternate Protocol 2) 10-ml polypropylene syringe 24-well crystallization trays (VDX Plate, Hampton Research) 22-mm2 plastic or siliconized coverslips (Hampton Research) Self-closing tweezers Microscope with ≥40× magnification, light source, and viewing platform (at least 3 × 4 in.), preferably with polarizer (e.g., Leica MZ16, Zeiss STEMI SV11, Nikon SMZ1000) NOTE: Preparation and crystallization (steps 1 to 9) should be done both at 4°C and room temperature (±3°C). Prepare crystallization trays 1. Fill a 10-ml polypropylene syringe with vacuum grease. 2. Apply a uniform layer of vacuum grease to 50 rims of three 24-well crystallization trays using the 10-ml syringe without a needle. Do not leave any gaps in the grease. Label the 50 wells, numbering across. Fifty rims require two entire plates plus two additional wells from a third plate.
3. Mix the solutions (reservoir mixes) in the appropriate crystallization screening kit according to manufacturer’s instructions. 4. Add 500 µl of each reservoir mix to a separate well, using a new pipet tip for each transfer.
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Table 17.4.3
Components of Crystal Screen 1 (Hampton Research) for Soluble Proteinsa
Condition
Formulationb
1 2 3 4 5 6 7 8 9 10 11 12
30% MPD, 0.1 M sodium acetate, pH 4.6, 0.02 M calcium chloride 0.4 M potassium/sodium tartrate 0.4 M ammonium phosphate 2.0 M ammonium sulfate, 0.1 M Tris⋅Cl, pH 8.5 30% MPD, 0.1 M sodium HEPES, pH 7.5, 0.2 M sodium citrate 30% PEG 4000, 0.1 Tris⋅Cl, pH 8.5, 0.2 M magnesium chloride 1.4 M sodium acetate, 0.1 M sodium cacodylate, pH 6.5 30% 2-propanol, 0.1 M sodium cacodylate, pH 6.5, 0.2 M sodium citrate 30% PEG 4000, 0.1 M sodium citrate, pH 5.6, 0.2 M ammonium acetate 30% PEG 4000, 0.1 M sodium acetate, pH 4.6, 0.2 M ammonium acetate 1.0 M ammonium phosphate, 0.1 M sodium citrate, pH 5.6 30% 2-propanol, 0.1 M sodium HEPES, pH 7.5, 0.2 M magnesium chloride 30% PEG 400, 0.1 M Tris⋅Cl, pH 8.5, 0.2 M sodium citrate 28% PEG 400, 0.1 M sodium HEPES, pH 7.5, 0.2 M calcium chloride 30% PEG 8000, 0.1 M sodium cacodylate, pH 6.5, 0.2 M ammonium sulfate 1.5 M lithium sulfate, 0.1 M sodium HEPES, pH 7.5 30% PEG 4000, 0.1 M Tris⋅Cl pH 8.5, 0.2 M lithium sulfate 20% PEG 8000, 0.1 M sodium cacodylate, pH 6.5, 0.2 M magnesium acetate 30% 2-propanol, 0.1 M Tris⋅Cl, pH 8.5, 0.2 M ammonium acetate 25% PEG 4000 30% MPD, 0.1 M sodium cacodylate, pH 6.5, 0.2 M magnesium acetate 30% PEG 4000, 0.1 M Tris⋅Cl, pH 8.5, 0.2 M sodium acetate 30% PEG 400, 0.1 M sodium HEPES, pH 7.5, 0.2 M magnesium chloride 20% 2-propanol, 0.1 M sodium acetate, pH 4.6, 0.2 M calcium chloride 1.0 M sodium acetate, 0.1 M imidazole, pH 6.5 30% MPD, 0.1 M sodium citrate, pH 5.6, 0.2 M ammonium acetate 20% 2-propanol, 0.1 M sodium HEPES, pH 7.5, 0.2 M sodium citrate 30% PEG 8000, 0.1 M sodium cacodylate, pH 6.5, 0.2 M sodium acetate 0.8 M potassium/sodium tartrate, 0.1 M sodium HEPES, pH 7.5 30% PEG 8000, 0.2 M ammonium sulfate 30% PEG 4000, 0.2 M ammonium sulfate 2.0 M ammonium sulfate 4.0 M sodium formate 2.0 M sodium formate, 0.1 M sodium acetate, pH 4.6 1.6 M sodium/potassium phosphate, 0.1 M sodium HEPES, pH 7.5 8% PEG 8000, 0.1 M Tris⋅Cl, pH 8.5 8% PEG 4000, 0.1 M sodium acetate, pH 4.6 1.4 M sodium citrate, 0.1 M sodium HEPES, pH 7.5 2% PEG 400, 2.0 M ammonium sulfate, 0.1 M sodium HEPES, pH 7.5 20% 2-propanol, 20% PEG 4000, 0.1 M sodium citrate, pH 5.6 10% 2-propanol, 20% PEG 4000, 0.1 M sodium HEPES, pH 7.5 20% PEG 8000, 0.05 M potassium phosphate 30% PEG 1500 0.2 M magnesium formate
13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44
continued Crystallization of Macromolecules
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Table 17.4.3 continued
Components of Crystal Screen 1 (Hampton Research) for Soluble Proteinsa,
Condition
Formulationb
45 46 47 48 49 50
18% PEG 8000, 0.1 M sodium cacodylate, pH 6.5, 0.2 M zinc acetate 18% PEG 8000, 0.1 M sodium cacodylate, pH 6.5, 0.2 M calcium acetate 2.0 M ammonium sulfate, 0.1 M sodium acetate, pH 4.6 2.0 M ammonium phosphate, 0.1 M Tris⋅Cl, pH 8.5 2% PEG 8000, 1.0 M lithium sulfate 15% PEG 8000, 0.5 M lithium sulfate
a Jancarik and Kim (1991). b MPD, 2-methyl-2,4-pentanediol; PEG, polyethylene glycol.
Set up hanging drops 5. Place 1 µl concentrated macromolecule sample onto the center of a 22-mm2 plastic or siliconized coverslip. Keep stock macromolecule sample on ice. 6. With a new tip, withdraw 1 µl from the reservoir mix in the first well. Add this to the 1-µl sample aliquot and pipet up and down several times to mix, taking care not to introduce air bubbles into the droplet. 7. Flip the coverslip over with self-closing tweezers so that the drop is hanging down from the bottom of the coverslip. Place the coverslip over the appropriate well solution. To seal the coverslip-well interface, gently tap the edges of the coverslip into the grease using the bottom of the tweezers. Figure 17.4.1 illustrates the components of a completed vapor-diffusion setup.
8. Repeat steps 5 to 7 for the rest of the well solutions. Incubate 9. After all of the hanging drops have been prepared, tape together the top and bottom of each tray. Transfer the trays to a location that is free from vibrations and is maintained at a fairly constant temperature (±3°C). Leave the trays undisturbed for at least 2 days. In general, crystallization trays should not be discarded for at least 6 months. Some proteins have been known to undergo time-dependent modifications that promote crystallization.
Analyze results 10. Use a microscope with ≥40× magnification, light source, and viewing platform (preferably with polarizer) to view the crystallization drops. Note whether the drops are clear (0), show crystals (C), or show precipitate or phase separation (P). 11. For those drops that produce crystals, note the crystallization conditions and see if there is a common parameter (i.e., precipitant, pH, or ionic strength). Also note the crystal size, shape, and ability to polarize light. In general, large (≥0.4-mm3) crystals that have sharp edges and that polarize light are preferable.
12. Continue to observe the crystallization trials every 2 days until no further change is noted in the drops.
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Table 17.4.4 Components of Natrix (Hampton Research) for Nucleic Acid and Protein-Nucleic Acid Complexes Screena
Condition Formulationb 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 Crystallization of Macromolecules
0.01 M magnesium chloride, 0.05 M MES, pH 5.6, 2.0 M lithium sulfate 0.01 M magnesium acetate, 0.05 M MES, pH 5.6, 2.5 M ammonium sulfate 0.1 M magnesium acetate, 0.05 M MES, pH 5.6, 20% MPD 0.2 M KCl, 0.01 M magnesium sulfate, 0.05 M MES, pH 5.6, 10% PEG 400 0.2 M KCl, 0.01 M magnesium chloride, 0.05 M MES, pH 5.6, 5% PEG 8000 0.1 M ammonium sulfate, 0.01 M magnesium chloride, 0.05 M MES, pH 5.6, 20% PEG 8000 0.02 M magnesium chloride, 0.05 M MES, pH 6.0, 15% isopropanol 0.005 M magnesium sulfate, 0.1 M ammonium acetate, 0.05 M MES, pH 6.0, 0.6 M NaCl 0.1 M KCl, 0.01 M magnesium chloride, 0.05 M MES, pH 6.0, 10% PEG 400 0.005 M magnesium sulfate, 0.05 M MES, pH 6.0, 5% PEG 4000 0.01 M magnesium chloride, 0.05 M sodium cacodylate, pH 6.0, 1.0 M lithium sulfate 0.01 M magnesium sulfate, 0.05 M sodium cacodylate, pH 6.0, 1.8 M lithium sulfate 0.015 M magnesium acetate, 0.05 M sodium cacodylate, pH 6.0, 1.7 M ammonium sulfate 0.1 M KCl, 0.025 M magnesium chloride, 0.05 M sodium cacodylate, pH 6.0, 15% isopropanol 0.04 M magnesium chloride, 0.05 M sodium cacodylate, pH 6.0, 5% MPD 0.04 M magnesium acetate, 0.05 M sodium cacodylate, pH 6.0, 30% MPD 0.2 M KCl, 0.01 M calcium chloride, 0.05 M sodium cacodylate, pH 6.0, 10% PEG 4000 0.01 M magnesium acetate, 0.05 M sodium cacodylate, pH 6.5, 1.3 M lithium sulfate 0.01 M magnesium sulfate, 0.05 M sodium cacodylate, pH 6.5, 2.0 M ammonium sulfate 0.1 M ammonium acetate, 0.015 M magnesium acetate, 0.05 M sodium cacodylate, pH 6.5, 10% isopropanol 0.2 M KCl, 0.005 M magnesium chloride, 0.05 M sodium cacodylate, pH 6.5, 10% 1,6-hexanediol 0.08 M magnesium acetate, 0.05 M sodium cacodylate, pH 6.5 0.2 M KCl, 0.01 M magnesium chloride, 0.05 M sodium cacodylate, pH 6.5, 10% PEG 4000 0.2 M ammonium acetate, 0.01 M calcium chloride, 0.05 M sodium cacodylate, pH 6.5, 10% PEG 4000 0.08 M magnesium acetate, 0.05 M sodium cacodylate, pH 6.5, 30% PEG 4000 0.2 M KCl, 0.1 M magnesium acetate, 0.05 M sodium cacodylate, pH 6.5, 10% PEG 8000 0.2 M ammonium acetate, 0.01 M magnesium acetate, 0.05 M sodium cacodylate, pH 6.5, 30% PEG 8000 0.05 M magnesium sulfate, 0.05 M sodium HEPES, pH 7.0, 1.6 M lithium sulfate 0.01 M magnesium chloride, 0.05 M sodium HEPES, pH 7.0, 4.0 M lithium chloride 0.01 M magnesium chloride, 0.05 M sodium HEPES, pH 7.0 0.005 M magnesium chloride, 0.05 M sodium HEPES, pH 7.0, 25% PEG monomethyl ether 550 continued
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Table 17.4.4 Components of Natrix (Hampton Research) for Nucleic Acid and Protein-Nucleic Acid Complexes Screena, continued
Condition Formulationb 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48
0.2 M KCl, 0.01 M magnesium chloride, 0.05 M sodium HEPES, pH 7.0, 20% 1,6-hexanediol 0.2 M ammonium chloride, 0.01 M magnesium chloride, 0.05 M sodium HEPES, pH 7.0, 30% 1,6-hexanediol 0.1 M KCl, 0.005 M magnesium sulfate, 0.05 M sodium HEPES, pH 7.0, 15% MPD 0.1 M KCl, 0.01 M magnesium chloride, 0.05 M sodium HEPES, pH 7.0, 5% PEG 400 0.1 M KCl, 0.01 M calcium chloride, 0.05 M sodium HEPES, pH 7.0, 10% PEG 400 0.2 M KCl, 0.025 M magnesium sulfate, 0.05 M sodium HEPES, pH 7.0, 20% PEG 200 0.2 M ammonium acetate, 0.15 M magnesium acetate, 0.05 M sodium HEPES, pH 7.0, 5% PEG 4000 0.1 M ammonium acetate, 0.02 M magnesium chloride, 0.05 M sodium HEPES, pH 7.0, 5% PEG 8000 0.01 M magnesium chloride, 0.05 M Tris⋅Cl, pH 7.5, 1.6 M ammonium sulfate 0.1 M KCl, 0.015 M magnesium chloride, 0.05 M Tris⋅Cl, pH 7.5, 10% PEG monomethyl ether 550 0.01 M magnesium chloride, 0.05 M Tris⋅Cl, pH 7.5, 5% isopropanol 0.01 M magnesium chloride, 0.05 M ammonium acetate, 0.05 M Tris⋅Cl, pH 7.5, 10% MPD 0.2 M KCl, 0.05 M magnesium chloride, 0.05 M Tris⋅Cl, pH 7.5, 10% PEG 4000 0.025 M magnesium sulfate, 0.05 M Tris⋅Cl, pH 8.5, 1.8 M ammonium sulfate 0.005 M magnesium sulfate, 0.05 M Tris⋅Cl, pH 8.5, 35% 1,6-hexanediol 0.1 M KCl, 0.01 M magnesium chloride, 0.05 M Tris⋅Cl, pH 8.5, 30% PEG 400 0.01 M calcium chloride, 0.2 M ammonium chloride, 0.05 M Tris⋅Cl, pH 8.5, 30% PEG 4000
a Scott et al. (1995). b MES, 2-(N-morpholino)ethanesulfonic acid; MPD, 2-methyl-2,4-pentanediol; PEG, polyethylene glycol.
ESTABLISHING WHETHER CRYSTALS ARE MACROMOLECULE OR SALT USING A CRYOLOOP Often one obtains crystals that are salt rather than macromolecule. This occurs notoriously when the crystallization conditions contain divalent ions such as magnesium or calcium with phosphate or sulfate counterions. If crystals are suspected to be salt, prepare a control crystallization trial in which everything but the macromolecule is included. For example, after concentrating in a microconcentrator, the flow-through can be used instead of the protein sample. In preparing this control, substitute the macromolecule sample with the storage buffer that is identical to that used for the macromolecule in the drops. If this test does not show crystals, one can analyze a washed crystal directly by SDS-PAGE (UNIT 10.1). The procedure for preparing an SDS-PAGE sample from a crystal is described here. A crystal that is at least 0.2 mm3 is usually large enough to wash with a cryoloop and dissolve for visualization by SDS-PAGE and silver staining (UNIT 10.5; both proteins and nucleic acids are stained with silver reagent). The procedure involves extensively washing the crystal surface, so that it is free of macromolecule from the hanging drop, before dissolving the crystal for gel analysis. For microcrystals, see Support Protocol 2.
SUPPORT PROTOCOL 1
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Materials Crystallization tray(s) containing 0.2-mm3 (or larger) crystal(s) and reservoir solutions (see Basic Protocol 2) Macromolecule storage solution (i.e., solution in which the macromolecule is soluble) Purified macromolecule 2× SDS sample buffer (UNIT 10.1) Microscope with ≥40× magnification, light source, and viewing platform (at least 3 × 4 in.), preferably with polarizer (e.g., Leica MZ16, Zeiss STEMI SV11, Nikon SMZ1000) Cryoloops (Hampton Research) Plastic petri dish Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and silver staining (UNIT 10.5) 1. Invert the coverslip harboring 0.2-mm3 crystals from the crystallization tray and place on a flat surface. Pipet 100 µl of the respective reservoir solution directly onto the drop. 2. View the drop under the microscope with ≥40× magnification, use a cryoloop to mix the solution in the drop, and gently move the crystal to make sure it is detached from the coverslip surface. 3. Pipet three separate 100-µl drops of reservoir solution into a plastic petri dish. 4. Place the cryoloop under the crystal such that the crystal is hanging half inside and half outside the circle, then lift the loop swiftly upward. Transfer the crystal to the first 100-µl drop, shake slightly to pry the crystal off the loop, and mix the solution slightly. When the crystal is “lassoed” in this manner, it is held in the loop by the surface tension of the liquid that is attached to the rim of the loop.
5. Transfer the crystal to the second drop, mix slightly by swirling the loop in the reservoir solution, and finally transfer the crystal to the third drop. 6. Use the cryoloop to transfer the crystal to a microcentrifuge tube containing 5 µl macromolecule storage solution, and shake the cryoloop to release the crystal. Vortex the solution to fully dissolve the crystal. 7. Set up the following controls: a. For a negative control, transfer 5 µl from the third washing drop to a microcentrifuge tube containing 5 µl of the macromolecule storage solution. b. For a positive control, add 1 µg purified macromolecule in a final volume of 5 µl macromolecule storage solution to another microcentrifuge tube. The negative control ensures that all of the free macromolecule in solution has been washed from the crystal surface.
8. Add 5 µl of 2× SDS sample buffer to each tube and analyze the samples by SDS-PAGE (UNIT 10.1). Visualize macromolecule(s) by silver staining the gel (UNIT 10.5).
Crystallization of Macromolecules
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ESTABLISHING WHETHER MICROCRYSTALS ARE MACROMOLECULE OR SALT BY FILTRATION Often one obtains a shower of small microcrystals that are difficult to manipulate by the procedure outlined above (see Support Protocol 1). In such cases, the procedure outlined below can be used to establish whether the microcrystals contain macromolecules.
SUPPORT PROTOCOL 2
Materials Crystallization tray(s) containing 0.2-mm3 (or larger) crystal(s) and reservoir solutions (see Basic Protocol 2) Macromolecule storage solution (i.e., solution in which the macromolecule is soluble) 2× SDS sample buffer (UNIT 10.1) Purified macromolecule Centricon-100 microconcentrator (MWCO 100; Amicon) 1.5-ml screw-cap microcentrifuge tube Hypodermic needle 500-ml lyophilization vessel Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and silver staining (UNIT 10.5) 1. Invert the coverslip harboring the crystals from the crystallization tray and use a 10-µl micropipettor to transfer the drop to a Centricon-100 microconcentrator. 2. Wash the coverslip with 50 µl of the respective reservoir solution by pipetting up and down several times with a 200-µl micropipettor. Use the 200-µl micropipettor to transfer this wash solution to the microconcentrator containing the drop. 3. Add an additional 500 µl reservoir solution to the microconcentrator and concentrate the entire sample by centrifugation to dead volume per manufacturer’s instructions. 4. Add 2 ml additional reservoir solution to the microconcentrator and reconcentrate by centrifugation to dead volume. 5. Add 100 µl macromolecule storage solution to the microconcentrator and mix by pipetting up and down several times with a 200-µl micropipettor. 6. Invert the microconcentrator and centrifuge to recover sample, which should now contain the dissolved crystals. 7. Transfer the sample to a 1.5-ml screw-cap microcentrifuge tube, punch holes in the top using a hypodermic needle, and freeze over crushed dry ice. Place in a 500-ml lyophilization vessel and lyophilize the sample to dryness. 8. Redissolve the lyophilized sample with 5 µl macromolecule storage solution and 5 µl of 2× SDS-PAGE sample buffer. 9. Set up the following controls: a. For a negative control, transfer 5 µl from the third washing drop to a microcentrifuge tube containing 5 µl of the macromolecule storage solution. b. For a positive control, add 1 µg purified macromolecule in a final volume of 5 µl macromolecule storage solution to another microcentrifuge tube. The negative control ensures that all of the free macromolecule in solution has been washed from the crystal surface.
10. Analyze the samples by SDS-PAGE (UNIT 10.1). Visualize macromolecule(s) by silver staining the gel (UNIT 10.5). Structural Biology
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BASIC PROTOCOL 3
OPTIMIZATION OF CRYSTALLIZATION PARAMETERS Although crystals that are obtained from sparse matrix screens are sometimes already suitable for structural studies, more often the crystals need to be optimized prior to crystallographic analysis. Common problems include crystals that are too small, too thin, clustered, or intergrown. In these cases, the optimization of precipitant concentration, pH, and ionic strength often improve crystal quality. There are different ways of optimizing crystallization parameters. The procedure generally used in the author’s laboratory is presented here. Figure 17.4.2 illustrates a typical optimization procedure, which is discussed further in annotations to the steps below. Materials Vacuum grease (Hampton Research) Crystallization screening kit(s) (Hampton Research): Proteins and multiprotein complexes: Crystal Screens 1 (Table 17.4.3) and 2 (latter is optional) Nucleic acids and protein–nucleic acid complexes: Natrix (Table 17.4.4) Concentrated macromolecule sample: protein (see Basic Protocol 1), nucleic acid (see Alternate Protocol 1), or macromolecular complex (see Alternate Protocol 2) 10-ml polypropylene syringe 24-well crystallization trays (VDX Plate, Hampton Research) 22-mm2 plastic or siliconized coverslips (Hampton Research) Self-closing tweezers Microscope with ≥40× magnification, light source, and viewing platform (at least 3 × 4 in.), preferably with polarizer (e.g., Leica MZ16, Zeiss STEMI SV11, Nikon SMZ1000) Additional reagents and equipment for preparation of crystallization trials (see Basic Protocol 2) 1. For each crystallization condition that yields crystals, prepare a crystallization tray (see Basic Protocol 2, steps 1 and 2) that contains an initial one-dimensional grid to optimize the concentration of precipitant. Using the crystallization screening kit, design precipitant concentrations that range in 10% and 15% increments from zero to ∼20% higher than that used in the screen condition. As an example (Fig. 17.4.2A), protein crystallization trials (see Basic Protocol 2) using Crystal Screen 1 (Table 17.4.3) yielded a shower of nicely shaped but small crystals under condition 9, which contains 30% polyethylene glycol (PEG) 4000, 0.2 M ammonium acetate, 0.1 M sodium citrate, pH 5.6. This condition was used as a starting point for further optimization. The starting protein concentration in the crystallization drop was 10 mg/ml. Figure 17.4.2A shows the first grid screen that was prepared to optimize the precipitant concentration.
2. Prepare reservoirs by adding the appropriate amounts of stock precipitant, buffer, and salt directly to the well and diluting with water to a final volume of 1 ml (see Basic Protocol 2, steps 3 and 4). Mix by pipetting up and down several times with a 1-ml micropipettor. 3. In each well, mix 1 µl concentrated macromolecule with 1 µl reservoir solution and equilibrate several days over the appropriate reservoir as described (see Basic Protocol 2, steps 5 to 9). Trays should be incubated at the same temperature used to obtain the initial crystals.
Crystallization of Macromolecules
4. Observe crystallization trays under a microscope with ≥40× magnification, light source, and viewing platform, and note the conditions that yield the best crystals (i.e., those with the sharpest edges, largest size, and most invariant form).
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Figure 17.4.2 Optimization of crystallization parameters (see Basic Protocol 3 for discussion).
5. Continue to observe crystallization trays for at least 1 week. In the example, after one week it was determined that crystals prepared from reservoir 3 were bigger and larger than those prepared from reservoirs 4 and 5. The condition for this reservoir (15% PEG 4000, 0.2 M ammonium acetate, 0.1 M sodium citrate, pH 5.6) was used as a starting point to optimize pH and ionic strength.
6. Prepare a two-dimensional grid to optimize pH and ionic strength conditions as a function of precipitant concentration (e.g., Fig. 17.4.2B) in the following manner: a. Vary the precipitant concentration more narrowly, based on the results of steps 1 to 5. b. Vary the ionic strength in 20% to 25% increments. c. Vary the pH in 1-unit increments both above and below the values used in the initial screen. Structural Biology
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Set up and observe trials as in steps 2 to 5 above. Note that different buffers will have to be used to establish different pH values. Generally, use the buffers that are employed in the sparse matrix screens. In the example, observations made after 1 week determined that crystals from reservoirs 10 and 13 looked the longest and most nicely shaped (Fig. 17.4.2B). Crystals in reservoir 18 looked striated with blunt edges. Therefore, the conditions from reservoirs 10 and 13 were used to further optimize the conditions at pH 4.6 and 0.05 M ammonium acetate.
7. To more finely optimize crystal growth conditions, use smaller changes of precipitating agent, ionic strength, and pH with 2-dimensional grids. Set up and observe trials as described in steps 2 to 5 above. In the example, observations made after 1 week determined that crystals grown from reservoirs 9 (16% PEG 4000, 50 mM ammonium acetate, 0.1 M sodium acetate, pH 4.6) and 14 (13% PEG 4000, 25 mM ammonium acetate, 0.1 M sodium acetate, pH 4.6) looked best (Fig. 17.4.2C); the crystals in reservoir 14 looked marginally better.
8. Use these conditions to prepare the next experimental grid. Use the results of this grid as the optimal crystallization parameters. In the example, observation of this grid shows that crystals in reservoirs 2 and 3 are the best (Fig. 17.4.2D). Therefore, the best crystallization parameters were 11% PEG 4000, 35 mM ammonium acetate, and 0.1 M sodium acetate, pH 4.6. ALTERNATE PROTOCOL 4
OPTIMIZATION OF CRYSTAL EQUILIBRATION CONDITIONS It is often the case that optimization of the chemical parameters for crystallization (see Basic Protocol 3) still yields crystals that are too small for crystallographic analysis. In such cases, it may be helpful to use a different crystallization procedure such as the sitting-drop method (Weber, 1997). Alternatively, the macromolecule concentration, the ratio of macromolecule solution to reservoir solution, and the drop size can be varied to obtain larger crystals using the vapor-diffusion technique. This latter approach is described here. Crystallization trials are performed as described previously (see Basic Protocols 2 and 3). Figure 17.4.3 illustrates a typical procedure for optimization of equilibration conditions. See Basic Protocol 3 for materials. 1. Using the optimized ionic strength, pH, and precipitant concentration (see Basic Protocol 3, steps 1 and 2), design a set of two-dimensional grids varying the parameters as follows: a. In each of these grids, vary the precipitant concentration finely along the 4-well axis, using four different precipitant concentrations. b. Duplicate this parameter in each of the columns in the other crystal trays. 2. Vary three different parameters in the other dimension of the grid screen (Fig. 17.4.3A). a. To decrease macromolecule concentration in constant drop volume: Dilute the macromolecule 2- and 4-fold relative to the concentration used in the initial screen, prior to mixing with the reservoir solution (1 µl dilute macromolecule to 1 µl reservoir solution). b. To increase drop volume at constant concentration: Vary the drop size by increasing the volume of both macromolecule solution and reservoir solution (i.e., 1 µl macromolecule to 1 µl reservoir, 2 to 2, and 3 to 3). c. To shift the equilibration process: Vary the reservoir solution/macromolecule solution ratio (i.e., 1:2, 2:1, 2:3, 3:2) in a constant drop volume (2 µl).
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Figure 17.4.3 Optimization of crystal equilibration parameters (see Alternate Protocol 4 for discussion).
Using the conditions determined (see Basic Protocol 3 and Fig. 17.4.2D), a screen to optimize crystallization equilibration conditions was prepared by the authors (Fig. 17.4.3A).
3. Set up trays as described (see Basic Protocol 3, steps 3 to 5) and observe them after several days. In the example (Fig. 17.4.3A), observations made after 1 week determined that reservoirs 14, 18, and 31 gave the largest crystals. Thus, the best parameters are 5 mg/ml protein mixed at a 2:3 protein/reservoir ratio in a final drop volume of 6 ìl.
4. Optional: If necessary, refine drops with an additional optimization step. In the example, one last round of optimization testing provided several crystals large enough for X-ray crystallographic analysis (Fig. 17.4.3B). Structural Biology
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OPTIMIZATION BY CRYSTAL SEEDING Many times the crystals obtained from initial crystallization trials are too small or unsuitable for X-ray diffraction studies. Along with the previously described optimization strategies, seeding techniques can yield larger, better diffracting crystals. When seeding, it is important to nucleate a crystallization drop just prior to crystal nucleation (microseeding) or crystal growth (macroseeding). More specifically, it is desirable to determine the time for introducing the seed. Since one does not typically know this ahead of time, crystal seeds are introduced into the crystallization drops at different time points (e.g., 2, 4, 8, 12, 18, and 24 hr). Ideally, one of these will result in the growth of large stable crystals. Procedures for crystal macroseeding and microseeding are described below (see Alternate Protocols 5 and 6). ALTERNATE PROTOCOL 5
Macroseeding Macroseeding involves the transfer of an existing crystal through a number of stabilizing solutions into a “fresh” pre-equilibrated drop. A crystal may not grow to a larger size for a number of reasons, including macromolecule depletion and surface poisoning. During the growth phase, monomers are added to the crystal and the concentration of the macromolecule decreases. Consequently, it may be desirable to transfer the crystal to a macromolecule solution where the concentration of macromolecule is higher and the crystal can grow again. Another reason a crystal fails to continue to grow is that the surfaces have been poisoned by contaminants. As such, it is desirable to “wash” the surfaces of the contaminants by transferring the crystal through multiple stabilizing solutions. Crystal drops can all be set up at the same time and seeded at different times after equilibration (e.g., 2, 4, 8, 12, 18 and 24 hr). The drop is created the same way as described above (see Basic Protocol 2). A typical seeding procedure is described below. Additional Materials (also see Basic Protocol 2) Microscope with ≥40× magnification, light source, and viewing platform (at least 3 × 4 in.), preferably with polarizer (e.g., Leica MZ16, Zeiss STEMI SV11, Nikon SMZ1000) Tweezers Glass fibers (see recipe) or crystal probe manipulators (Hampton Research) Capillary-tipped syringe (optional; see recipe) 1. Place four plastic or siliconized coverslips on top of the tray cover. Pipet 50 to 100 µl wash solutions (typically the same solution as the reservoir solution) onto the middle of each coverslip. Placing the coverslips on the tray cover permits one to quickly perform the manipulations without having to make drastic adjustments to the focus of the microscope. The top of a 15-ml conical screw-cap tube can be placed over the wash solutions to prevent dehydration. The wash solutions may also need to be optimized. If the crystals dissolve too quickly or visibly crack, then adjustments should be made to the precipitating agent. A typical wash solution contains the crystallization reservoir conditions with a 20% increase in the concentration of the precipitating agent. It may be necessary to omit any volatile alcohol solution (i.e., ethanol) in the wash solutions. Finally, some procedures suggest adding 0.5% to 2.0% (w/v) of a detergent, typically OG (octyl-β-D-glucopyranoside), to the wash solution.
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2. Using a microscope with ≥40× magnification, light source, and viewing platform, select a crystal without visible defects in the tray to macroseed. It is important to select a crystal without visible defects. If the underlying crystal has defects, the crystal may grow to be an accumulation of multiple crystals.
3. Being careful not to break the coverslip, gently break the vacuum grease seal using the tweezers. Carefully lift the coverslip, flip it over and place it on the tray cover next to the wash solutions. 4. Add ∼20 to 50 µl wash solution to the drop to generate sufficient volume with which to work. 5. Under the microscope, isolate a single crystal from other crystals or precipitate with a glass fiber or crystal probe manipulators. For crystal clusters, carefully use the probe to gently touch the cluster to break it apart. For a crystal embedded in precipitate, starting at the edge of the crystal, sweep the precipitate away from all sides of the crystal. A single crystal should be selected and swept to an edge of the drop free of other crystals or precipitate, so that the crystal can easily be picked up by a capillary-tipped syringe or cryoloop.
6a. Using a capillary-tipped syringe: Under the microscope, gently use the capillarytipped syringe to draw up the crystal with as little liquid as possible. Watching through the microscope, expel the crystal into the first wash solution. 6b. Using a cryoloop: Perform crystal transfers with a cryoloop (Support Protocol 1). It is often advantageous to wet the capillary before this step using wash solution. Also, before drawing up the crystal, it is useful to pull the plunger out half-way to be able to expel the crystal. The length of time of incubating the crystal in each wash step can be from 15 sec to 5 min. If cracks appear or the crystal starts to dissolve, shorter washes may be necessary.
7. Using a new capillary, draw up the crystal and expel into the next wash solution as in step 4. Repeat for a total of 4 washes. 8. Remove the equilibrated drop from above the reservoir, expel the crystal in the pre-equilibrated drop and replace this equilibrated drop (now containing a macroseeded crystal) over the reservoir. Make sure that vacuum grease forms an airtight seal with the coverslip.
Microseeding: Streak Seeding
ALTERNATE PROTOCOL 6
Microseeding involves providing a nucleation center for crystals to grow. The trick is to present only a few minute crystals per pre-equilibrated drop around which suitable crystals will form. The most popular form of microseeding, streak seeding, is described below. This and other seeding techniques are also described by Stura and Wilson (1990). Materials (also see Basic Protocol 2) Stabilization solution (e.g., reservoir solution with 20% higher precipitating reagent) Glass fibers or crystal probe manipulators (Hampton Research) Capillary-tipped syringe (see recipe) Glass minipestle (see recipe) Rabbit hair streaker (see recipe) Structural Biology
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1. Prepare a pre-equilibrated drop the day before the experiment as described (see Basic Protocol 2). Lower the concentration of the precipitating agent so as to be conducive for the growth of crystals, but not nucleation. 2. Using a microscope with ≥40× magnification, light source, and viewing platform, select which crystal in the tray to microseed. It is important to select a well-formed crystal without visible defects.
3. Being careful not to break the coverslip, gently break the vacuum grease seal using tweezers. Carefully lift the coverslip, flip it over, and place it on the tray cover. 4. Under the microscope, isolate the crystal from other crystals or precipitate with a glass fiber or crystal probe. 5. Under the microscope, use a capillary-tipped syringe gently to draw up the crystal with as little liquid as possible. Expel the crystal into a microcentrifuge tube containing the stabilization solution. 6. Using a minipestle, break the crystal apart in the stabilization solution. 7. Centrifuge the microcentrifuge tube containing the solution of crushed crystal 10 min at low speed (100 × g), at the temperature used to grow the crystals. 8. Transfer 10 µl crushed crystal solution into a new microcentrifuge tube containing 90 µl stabilization solution. Repeat to obtain a serial dilution of 10−4 to 10−8. 9. Gently remove the coverslip from the pre-equilibrated drop, submerge the rabbit hair streaker into various dilutions of the microcrystal solution and streak through the pre-equilibrated drop. 10. Replace the coverslip over the reservoir solution, making sure that the vacuum grease forms an airtight seal with the coverslip. Typically a grid of dilution versus time of equilibration is used to optimize the choice of adding just a few nucleation sites at the appropriate time for nucleation.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Capillary-tipped syringe A capillary-tipped syringe is used to draw up the crystal in the liquid. To make this useful instrument, place silicone tubing over the tip of a syringe. Next obtain a glass capillary. Note that glass capillaries have two different ends: one side is sealed off, the other opens up to a larger diameter. Score the glass capillary with a diamond pencil near the tip of the sealed end and carefully break it off. Place the capillary with the large end snugly into the silicone tubing (Fig. 17.4.4). Small amounts of vacuum grease may be applied to the inside of the tubing to provide a sealed fit. Glass fibers Glass fibers are useful for manipulating crystals, breaking up crystal clusters and separating crystals from precipitate (Fig. 17.4.4). To prepare, hold both ends of a Pasteur pipet with each hand and place the middle part of the pipet in the flame of a Bunsen burner. Rotate the pipet in the flame until the glass becomes orange. After the pipet becomes orange, remove it from the flame, and with one quick motion pull the two ends apart. Wait for Pasteur pipet to cool and separate into 10- to 20-cm sizes. It is important to not allow holes in the glass fiber or it will draw the liquid up through capillary action. Crystallization of Macromolecules
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Figure 17.4.4 Typical home-made crystal manipulation tools.
Minipestle The minipestle is made from a Pasteur pipet using a Bunsen burner. Using a diamond pen (Hampton Research), score the circumference of the pipet on the small diameter side, ∼1 in. away from the neck, and break it off. Put the smaller side gradually in the flame and rotate the pipet until a ball is formed. Then let the minipestle cool. Make sure the minipestle will fit in the size microcentrifuge tube containing the stabilization solution. Rabbit-hair streaker Many crystallographers avow that only rabbit hair works to seed crystals. However, human hair or mouse whiskers have also proven successful. For easier manipulation, make the streaker by gluing the hair onto the end of an unused pen. COMMENTARY Background Information X-ray crystallography has undergone a renaissance over the last decade. This change has been due to a combination of several factors. Among the most important of these are advances in data collection techniques, computa-
tional resources, and crystallographic refinement procedures. In addition, advances in recombinant DNA technology have opened up structural biology to many more types and varieties of proteins and macromolecular complexes. Because of these advances, X-ray crys-
Structural Biology
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tallography has moved from being a technique that was only applied to available protein samples to one that is open to any biological question. In other words, crystallography no longer drives the questions; biological questions drive the crystallography. A significant result of this renaissance is that the procedures of crystallography are no longer restricted to a dedicated crystallography laboratory. Crystallography is now a tool used by many laboratories that do not necessarily specialize in crystallographic analysis. In fact, it is quite straightforward for a molecular biology laboratory to prepare crystals that may be suitable for structure determination using X-ray crystallography. Macromolecular crystallization is the process by which macromolecules are coaxed out of solution in the form of crystals. The induction of crystal formation is facilitated by a precipitating agent that functions to drive equilibration between the small macromolecule-containing solution and large reservoir solution. A volatile precipitating agent, such as 2-methyl-2,4-pentanediol (MPD) or isopropanol, transfers by vapor-diffusion from the reservoir to the drop, thus increasing the precipitant concentration in the drop and driving the macromolecule out of solution (in the form of precipitate or crystals). Alternatively, a nonvolatile precipitating agent, such as salt or large polyethylene glycol (PEG), functions to promote the diffusion of water from the drop to the reservoir, causing the precipitant concentration to increase in the drop, thus driving the macromolecule out of solution. A more detailed discussion underlying the theory of crystal formation from a supersaturated macromolecular solution can be found in McPherson (1982). Here the authors present a method for macromolecular crystallization by the vapor-diffusion technique using hanging drops. It is the most convenient and commonly used technique, and often yields crystals from suitable macromolecular samples; however, more advanced techniques (e.g., sitting drops, microdialysis; UNIT 17.3 ) should be used if these procedures are unsuccessful.
Critical Parameters and Troubleshooting
Crystallization of Macromolecules
The most critical parameter for obtaining macromolecular crystals that are suitable for structure determination using X-ray crystallography is the macromolecular sample itself. In general, homogeneous, compact, monodisperse macromolecular samples are straightfor-
ward to crystallize, while nonhomogeneous, polydisperse samples with flexible domains, or samples that tend to aggregate nonspecifically, are very difficult to crystallize (D’Arcy, 1994; Ferre-D’Amare and Burley, 1994). Generally, if crystals do not form with the appropriate sparse matrix crystallization screens at either room temperature or 4°C, it is likely that the macromolecule needs to be modified in some way to obtain well-ordered crystals. One modification that can be performed is to remove flexible domains of the protein. In general, flexible domains are most easily identified by limited proteolysis, followed by protein sequencing of the remaining globular product. Recombinant DNA technology can then be used to genetically engineer the desired protein fragment. Another modification is to prepare a macromolecular complex rather then a nascent protein. Formation of a complex sometimes serves to bury otherwise hydrophobic regions that can nucleate protein aggregation and thus hamper crystallization. In this regard, proteinantibody complexes sometimes crystallize more readily than free proteins. Aside from the macromolecular sample, physiologically relevant cofactors often play major roles in crystallization. For example, enzymes may show entirely different crystallization behavior when they are bound to various nonhydrolyzable small-molecule or peptide substrates. The most common problem associated with macromolecular crystallization is that the protein precipitates either upon concentration or in a large majority of crystallization conditions. This often indicates that the protein has a strong tendency to aggregate at high concentrations. If this is the case, one of the two modification strategies described above should be considered. Diluting the macromolecule 2- to 4-fold may also help. Alternatively, crystallization trays may show predominantly clear drops. This indicates that the protein concentration is too low and needs to be raised (between 2- and 4-fold), or that the protein is overly soluble. In the latter case, one might try crystallization screens at or very near the pI of the macromolecule in order to reduce its charge. Another common problem in crystallizing macromolecules is that crystals of salt rather than of macromolecule are obtained. To determine if this is the case, set up a control drop containing everything except macromolecule. Salt crystals are commonly obtained with divalent ions (particularly with phosphate or sulfate counterions).
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Anticipated Results Crystallization trials generally show one of three types of results: clear drops, crystals, or precipitate or phase separation. Most often, one obtains a mixture of all of these results depending on the well conditions. If crystals are obtained under several different screen conditions, it is usually straightforward to proceed from small crystals to well-ordered, large crystals suitable for structure analysis by X-ray crystallography. Conditions that result in precipitate or phase separation generally are incompatible with crystal formation. If all, or nearly all, of the drops show precipitate, this is a bad sign and suggests that a modification of the macromolecule is required (see Critical Parameters and Troubleshooting).
Time Considerations Once a purified macromolecule is on hand, it can usually be concentrated and set up in crystallization trials in a matter of a few days. Assessment and refinement of crystallization parameters can take between a week to several months, depending on the behavior of the macromolecular sample.
Jancarik, J. and Kim, S.-H. 1991. Sparse matrix sampling: A screening method for crystallization of proteins. J. Appl. Cryst. 24:409-411. McPherson, A. 1982. The Preparation and Analysis of Protein Crystals. John Wiley & Sons, New York. McPherson, A. 1999. Crystalization of Biological Macromolecules. Cold Spring Harbor Press, Cold Spring Harbor, New York. Scott, W.G., Finch, J.T., Grenfell, R., Fogg, J., Smith, T., Gait, M.J., and Klug, A. 1995. Rapid crystallization of chemically synthesized hammerhead RNAs using a double screening procedure. J. Mol. Biol. 250:327-332. Stura, E.A., Satterthwait, A.C., Calvo, J.C., Kaslow, D.C., and Wilson, I.A. 1994. Reverse screening. Acta Cryst. 50: 448-455. Stura, E.A. and Wilson, I.A. 1990. Analytical and production seeding techniques. In. Methods: A Companion to Methods in Enzymology. 1:3849. Weber, P. 1997. Overview of protein crystallization methods. Methods Enzymol. 276:13-22.
Key References Stura, E.A. and Wilson, I.A. 1991. Seeding Techniques. In: Crystallation of Nucleic Acids and Proteins: A Practical Approach (A. Ducruix and G. Giegé, eds.) IRL Press, Oxford. pp. 241-254.
Literature Cited Aggarwal, A.K. 1990. Crystallization of DNA binding proteins with oligonucleotides. Methods Companion 1:83-90. D’Arcy, A. 1994. Crystallizing proteins: A rational approach? Acta Cryst. D50:469-471.
Contributed by Troy Messick and Ronen Marmorstein The Wistar Institute Philadelphia, Pennsylvania
Ferre-D’Amare, A. and Burley, S. 1994. Use of dynamic light scattering to assess crystallizability of macromolecules and macromolecular assemblies. Structure 2:357-359.
Structural Biology
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Introduction to NMR of Proteins Nuclear magnetic resonance (NMR) is an extremely important technique for structural analysis that bridges chemistry, physics, biochemistry, and medicine. From the characterization of a small organic (or inorganic) molecule to the full conformational determination of a biomolecule, NMR is at the forefront of structural analysis. This has largely been the result of major advances in the field of NMR. The advent of two-dimensional spectroscopy in the early 1980s sparked a revolution in the applicability of NMR to complex molecules with overlapping resonances. Isotope labeling, enabling the extension of NMR into heteronuclear chemical shift dimensions, has further increased the utility of NMR for protein analysis. The molecular weight limit for which NMR methods are applicable has been increased dramatically by using triple-resonance methods and deuterium labeling. Recent advances have enabled more interactions between atoms to be mapped and has led to the exciting possibility of NMR structural determination of very high-molecular-weight proteins (>100 kDa). This unit aims to overview the application of NMR spectroscopy to proteins. It is not intended to provide an exhaustive “how to” guide, but rather to give a flavor of both well-established and emerging NMR techniques used in the elucidation of protein structure. It starts with a brief introduction to the basic principles of NMR and the information provided by this technique, and goes on to discuss the instrumentation involved, spectral assignment methods for small and large proteins, and the utility of other spin active nuclei (e.g., 13C and 15N) to aid assignment of the latter.
PRINCIPLES OF NMR The physical mechanism of the NMR technique can be summarized briefly by its name: nuclear (a physical phenomenon involving the nuclei of atoms), magnetic (the effect requires a magnetic field), resonance (the absorption of energy at a well-defined energy, i.e., frequency). Most elements in the periodic table have at least one nuclear isotope that will interact with a magnetic field and exhibit the phenomenon of NMR. The property of a nucleus that allows this interaction is called “spin,” and is expressed in terms of a nuclear spin quantum number (I). Table 17.5.1 shows some NMR properties of several nuclei present in proteins. The hydrogen nucleus (the proton, 1H) has a half-integer spin,
UNIT 17.5
is close to 100% natural abundance, and is the most important nucleus in the study of proteins. The isotopes 13C and 15N also have half-integer spin, but their natural abundance is somewhat lower than for the proton, making these nuclei difficult to observe directly. 17O and 33S are the most common NMR active isotopes of oxygen and sulfur, respectively, but are not generally studied due to their insensitivity. The “spinning” nucleus can be thought of as a rotating positive charge, generating a minute magnetic field. In the absence of an external magnetic field, the direction of the axis of spin is random, but in the presence of such a field, each “nuclear magnet” will adopt a specific orientation. For spin-1⁄2 nuclei, two orientations are adopted: a low-energy (or α) state and a high-energy (or β) state. The α state is aligned in the same direction as the applied magnetic field, whereas the β state is aligned in the opposite direction. When a sample (an ensemble of spins) is placed in a magnetic field, a bulk magnetization is induced, as there are fractionally more spins in the α state than in the β state. The slight excess of α spins makes NMR an inherently insensitive spectroscopic technique. The induced nuclear magnetization traces out a circular path about the field axis, a motion termed precession. The rate of precession (called the Larmor frequency, vL) is proportional to both the experimentally determined magnetogyric ratio (γ in Table 17.5.1) and the magnetic field (represented by B0).
vL =
γB0 2π
The absorption of energy by the nuclear spins results in an excited state, in which the populations of the α and β energy levels are perturbed from their equilibrium values. Transitions between the α and β energy levels (the NMR phenomenon) occur when an appropriate amount of energy (resonance) is supplied to the system. The energy is supplied by a burst (or pulse) of electromagnetic radiation at a frequency of νL. For common nuclei (i.e., those in Table 17.5.1) in commercially available magnets, νL corresponds to radiofrequency radiation (100 to 800 MHz). The higher the external magnetic field, the greater the Larmor frequency of a given nucleus, the greater the difference in energy between the α and β states, and the more sensitive the NMR experiment. Structural Biology
Contributed by Andrew J. Edwards and David Reid Current Protocols in Protein Science (2000) 17.5.1-17.5.39 Copyright © 2000 by John Wiley & Sons, Inc.
17.5.1 Supplement 19
Table 17.5.1
NMR Properties of Selected Nuclei
Nucleus (isotope)
Spin (I)
Natural Magnetogyric Relative abundance (%) sensitivityb ratio (γ)a
NMR frequency (νL in MHz) at 9.4 Tc
Hydrogen (1H) Deuterium (2H or D)
1/2 1/2
99.98 0.015
26.8 4.1
1 1.5 × 10−6
400 61.4
Carbon (13C) Carbon (12C)
1/2 0
1.1 98.9
6.7 0
1.8 × 10−4 0
100.6 0
Nitrogen (14N) Nitrogen (15N)
1 1/2
99.63 0.37
1.9 −2.7
10−3 3.9 × 10−6
28.9 40.5
Oxygen (17O) Phosphorous (31P)
5/2 1/2
0.04 100
−3.6 10.8
10−5 6.6 × 10−2
54.2 162
aUnits are 107 rad s−1 T−1. bAt natural abundance. cThe strength of the magnetic field is measured in units of tesla (T).
INSTRUMENTATION Several types of magnet produce sufficiently high magnetic fields for NMR spectroscopy. Permanent iron magnets and electromagnets can generate fields of ≤2.35 T (100 MHz), but are little used in protein NMR due to insufficient resolution. The majority of magnets used for protein NMR are of the superconducting or “cryo” type, generating stable homogeneous fields up to 18.8 T (800 MHz). The magnetic field is produced by an electric current (up to 100 amperes) flowing through a superconducting wire wound into a solenoid. The wire is composed of a niobium/tin alloy, having the property of zero electrical resistance at very low temperatures. A current will flow in such a superconducting wire without loss, as long as the coil is kept below a critical temperature, achieved by immersion in a bath of liquid helium at a temperature of 4.2K (−269°C).
The console is the spectrometer control center. It houses the frequency synthesizers that produce the radiofrequency radiation, the control systems for probe heating/cooling, a shim interface that controls the homogeneity of the magnetic field on a sample-to-sample basis, and electronic circuits to amplify and digitize the signal from the sample. The computer hardware and associated software are designed to enable an operator to control the workings of the console and ultimately what happens at the probe. Data are written to a local hard disk at the end of an NMR experiment, and are processed using either the spectrometer vendor’s own software or an ever-expanding range of third-party NMR processing and analysis packages. The specialized hardware required for NMR spectroscopy of proteins is expensive to purchase and maintain. Thus, NMR spectroscopy is a technique that is usually found in centers of excellence or specialized departments.
Probe
PRACTICALITIES
Magnet
Introduction to NMR of Proteins
Console and Computer
The probe is the interface between the spectrometer and the sample, and is designed to place the sample in the region of strongest and most homogeneous field. Most importantly, it contains a resonant coil that surrounds the sample and transmits the radiofrequency energy required to promote the NMR transition. Modern sophisticated probe designs allow coils tuned to frequencies of several nuclei to be in proximity to the sample. The probe also houses an electrical heating system that, with the use of cooled gas, allows the sample in the magnet to be maintained at a given constant temperature.
Sample Preparation NMR is an inherently insensitive technique and requires relatively high sample concentrations to generate spectra with sufficiently high signal-to-noise ratio in a reasonable length of time. As such, most NMR studies of proteins are carried out at concentrations of 1 mM or above (e.g., 5 mg of a 10 kDa protein in 0.5 ml solvent = 1 mM). This requirement for unusually concentrated samples is rather limiting, as the protein may aggregate at this concentration or may not even dissolve fully. It may be possible to
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reverse or limit aggregation by altering the pH or temperature or by adding salts or detergents, the selection of which may be aided by prior knowledge of the properties of the protein. The most important sample preparation/analysis parameters for NMR are solvent, pH, temperature, concentration, and purity. By far the most usual solvent for the NMR analysis of proteins is water. Water is used for experiments that require the observation of amide protons, usually with the addition of ∼10% 2H O (heavy water). The latter is added to pro2 vide a field/frequency lock, which is a method used on most spectrometers to provide stable conditions for extended NMR experiments. Proton NMR spectroscopy of a sample dissolved in H2O requires that the intense solvent resonance be eliminated; otherwise, dynamic range problems will be encountered, leading to poor spectra. Many methods have been devised to achieve this, from simple presaturation to sophisticated techniques using pulsed magnetic field gradients in conjunction with selective radiofrequency pulses (e.g., Piotto et al., 1992). The use of 2H2O as solvent may be preferable in some cases. The residual resonance of partially deuterated water (HOD) is far less intense than the H2O resonance, and excellent quality spectra can usually be obtained with the minimum of effort. Amide NH and other exchangeable resonances will not be observed if sufficient time is left between dissolution and spectral acquisition. It is possible to use solvents other than H2O or 2H2O; for instance, dimethylsulfoxide-2H6 (DMSO-2H6) or trifluoroethanol (TFE) may be used for small peptides. It is a wise precaution to include a preservative in the NMR sample, especially for samples dissolved in H2O at neutral pH. This is commonly achieved by adding ∼1 mM sodium azide. It is common to record protein NMR spectra at acidic pH (∼3 to 5), as the rate of exchange of backbone amide protons with water attains a minimum within this range. This is a key consideration for experiments that rely on the observation of the amide proton resonance (e.g., see Three-Dimensional NMR and 15N-Edited Experiments, and see Spectral Assignment Using Triple-Resonance Multidimensional Heteronuclear Methods), and especially for regions of protein with little or no “structure” or for regions that are highly mobile. The actual pH chosen for the NMR study of a particular protein may be influenced more by solubility or propensity to aggregate at a particular pH than by the amide exchange rate. In some cases, it may be desirable to record spectra in 2H2O, so that the
amide exchange rate is no longer as important a consideration (since all amide protons will eventually exchange in 2H2O, given adequate time). Fully deuterated formic and acetic acids are useful as buffers in the acidic pH region, whereas phosphate can be used around neutral pH, if compatible with the protein under investigation. When using nonprotic solvents (e.g., DMSO), it is important to remember to use proton-containing acids or bases to alter pH, as the use of 2HCl may exchange all amide protons. The choice of temperature will be largely dictated by the thermal stability of the protein under analysis (both short- and long-term stability should be kept in mind). If the protein is thermally stable, it may be useful to record spectra at elevated temperature, as the increased thermal mobility of the protein will lead to a shorter correlation time (τc; see Relaxation Times: T1, T2, τc, and the nOe), and hence to sharper resonances. The sample must be as chemically pure as possible. The familiar techniques of protein purification are not incompatible with NMR, but due to the large amounts of protein required, a concentration step is usually needed. Freeze drying (lyophilization), although not compatible with all proteins, is an excellent method of concentration, but will also concentrate any inorganic salts present. If applicable, a desalting procedure using (for instance) ionic interaction chromatography (UNIT 8.4) may be used prior to lyophilization, where volatile salts such as ammonium bicarbonate can be used very effectively as the mobile phase. Ultrafiltration (UNIT 4.4) is another widely used method to bring pure protein solutions to NMR concentrations.
Titration of Metal Ions or Organic Molecules It is often desirable to monitor the NMR spectrum of a protein as a function of the concentration of a ligand (see Protein-Ligand Interactions), in order to determine, for instance, the location of a binding site or the effect of binding on the overall structure of a protein. In most cases, NMR structural information will have already been obtained on the non-ligand-bound protein; this is required, of course, to compare the apo- and the ligand-bound structures. In order to minimize bulk solution effects (e.g., dilution, change in ionic strength, or solvent composition), it is desirable to add the ligand in small aliquots taken from a concentrated stock. Small organic molecules are usually dissolved to high concentration in deuterated solvents such as DMSO-2H6 or methanol-2H4, whereas
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metal ions are generally introduced as solutions in H2O or 2H2O. As a minimum, one-dimensional proton spectra are recorded at intermediate titer points (e.g., 0.1, 0.5 molar equivalents of ligand) and the effect of the ligand on the spectra is assessed. Small variations of pH may occur as the ligand is titrated into the sample, so correction may be necessary at each titer point.
Isotopic Labeling
Introduction to NMR of Proteins
Labeled peptides and small proteins may be synthesized using solid-phase techniques with appropriately labeled commercially available precursor amino acids (UNITS 18.1 & 18.2). Medium to large proteins with non-native isotopic composition must be prepared from organisms that express the protein naturally or in which overexpression has been genetically engineered (see Chapter 5; Markley and Kainosho, 1993; Stockman, 1996; Mossakowska and Smith, 1997). Many expression hosts may be used (e.g., yeasts, UNITS 5.6-5.8; photosynthetic algae; and mammalian cell lines, UNITS 5.9 & 5.10), but by far the most common is the bacterium Escherichia coli (E. coli), due in part to its well-understood genetic makeup and its widespread use in molecular biology (UNITS 5.1-5.3). Often the expression methodology is in place before NMR studies are required. The choice of isotopic label and its distribution in the protein depends markedly on the protein and the information required from the NMR study. Examples of the use of uniform and specific isotopic labeling are discussed in other sections of this unit (see Spectral Assignment Using Double-Resonance Heteronuclear Methods; see Spectral Assignment Using TripleResonance Multidimensional Heteronuclear Methods; see Small Molecule Interactions; and see Other Protein NMR Techniques and Future Directions). The choice of growth media depends on the pattern of enrichment required. For instance, specific labeling requires the use of a minimal medium to which the isotopically labeled feedstocks are added. This is the method of choice for labeling specific amino acids in specific ways (e.g., labeling each leucine with 15N or substituting all phenylalanines with Phe2H ), but the possibility of isotopic scrambling 5 through the host’s metabolism must be considered. Sources of nitrogen and carbon in minimal media are usually inorganic salts (such as ammonium chloride) or small organics (like glucose). Uniform enrichment may also be accomplished this way, by substituting isotopically enriched sources of nitrogen and carbon.
For uniformly labeled proteins it is also possible to use a “complex medium” containing isotopically enriched algal hydrolysates. Growth in such a rich medium may produce the protein in higher yield, but this must be decided on a case-by-case basis. Random fractional deuteration can be accomplished using either minimal medium supplemented with an appropriate proportion of 2H2O or complex medium in which deuterium is incorporated into the feedstocks.
INFORMATION NMR CAN PROVIDE Chemical Shift The molecular framework of a molecule (the chemical bonds between atoms) induces slight changes in the magnetic field experienced by the nuclei in that molecule. This is because the electrons that constitute chemical bonds also interact with the magnetic field, and act to “shield” the nuclei from the field. The more electron density associated with a particular nucleus, the greater the shielding from the static B0 field, and the lower the resonance frequency. Conversely, a nucleus bonded to an electronegative atom has a reduced electron density. Such a nucleus is said to be deshielded and experiences a greater proportion of the static B0 field; consequently, its NMR transition is of higher energy. A universal convention in NMR spectroscopy is to express resonance frequencies in such a way as to be independent of B0. This is the chemical shift scale (δ), and is defined as follows:
δ=
(vobs − v ref ) v0
× 10 6
where νobs is the frequency of a resonance of interest, v0 is the spectrometer frequency, and νref is the frequency of a signal from a chosen reference compound. Chemical shifts are expressed in dimensionless “units” of parts per million (ppm). The reference substance used and its frequency depend on the nucleus under observation (this question will be addressed later for nuclei other than protons). For proton magnetic resonance, the reference substances generally used are water-soluble alkyl silane derivatives: 5,5-dimethylsilapentanesulfonate (DSS) and 3-(trimethylsilyl)propionic-2,2,3,32H acid sodium salt (TSP). The resonance 4 frequency of the trimethylsilyl protons of these compounds is assigned as 0 ppm. The chemical
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A Met ε
acetate
HOD
TSP
Ala
Gly α Met δ Met β
7
6
5
4
3
2
1
0
ppm
B
C Phe δ Phe ζ
Tyr δ
D
Tyr ε
Tyr β
Phe ε
3J 2
Met α Tyr α Ala α
Phe α
αβ
2J
J ββ′
ββ′ 3J
αβ′
Phe β 7.5 7.4 7.3 7.2 7.17.0 6.9 6.8 6.7 ppm
4.6 4.54.4. 4.3 4.2 4.1 ppm
3.25 3.15 3.05 2.95 2.85 ppm
ε
E
S β
O
ε
δ
O α
γ
O
NH α
NH2
β
β
O α
α
NH
NH
α β
O
H
NH
O
OH
δ
ζ
ε
Figure 17.5.1 (A) Anisotropic shielding effect of carbonyl and aromatic rings. Regions of shielding are marked “+” and regions of deshielding are marked “−”. Nuclei within the shielding region are shifted upfield. (B) The effect of secondary structure on chemical shifts. In α helices, both the α and carbonyl carbons are shifted downfield (dark shaded ovals) compared to the “random coil shift,” whereas the β carbon and α proton are shifted upfield (light shaded ovals). In β sheets, shifts in the opposite sense are observed. These empirical observations are useful indicators of the presence of secondary structure in a protein. However, more definitive methods like the observation of characteristic nuclear Overhauser interactions are required to confirm such observations.
shift scale is convenient, as NMR spectra obtained on spectrometers of different operating frequencies can be directly compared. By convention, the ppm scale is shown with zero at the right-hand side, with chemical shift increasing to the left. By convention, an "upfield shift" refers to an increase in nuclear shielding and a lower resonance frequency. Chemical shifts in proteins are also influenced by noncovalent interactions, specifically hydrogen bonding and the proximity and relative orientation of carbonyl groups and aromatic
rings (Fig. 17.5.1A). These influences on the chemical shift reflect the local environment of a nucleus in a protein and are thus a function of the secondary and tertiary folded structures. Table 17.5.2 lists the “random coil” (i.e., the chemical shift expected in the absence of noncovalent interactions) proton and carbon-13 chemical shifts of all 20 amino acids. Corresponding R groups of the amino acids are given for reference in Figure 17.5.2. The deviation of observed chemical shifts from their random coil values can give an indi-
Structural Biology
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Table 17.5.2
Proton and Carbon Chemical Shifts of Amino Acid Residues in Proteinsa
Residueb
α
β
Ala
4.35 50.8 3.97 43.9 4.18 60.7 4.38 53.8 4.22
1.39 17.7
59.6
37.5
4.50 56.6 4.35 60.1 4.69 53.9 4.52 54.0 4.76 52.7 4.29 55.4 4.75 51.5 4.37 54.1 4.36 54.6 4.38 54.6 4.63 53.6 4.44 61.9 4.66 56.2 4.60 56.3 4.70
3.88 62.3 4.22 68.4 3.28, 2.96 40.0 2.15, 2.01 31.7 2.84, 2.75 39.8 2.9. 1.97 28.7 2.83, 2.75 37.7 2.13, 2.01 28.1 1.85, 1.76 31.9 1.89, 1.79 29.6 3.26, 3.20 28.1 2.28, 2.02 30.6 3.22, 2.99 38.4 3.13, 2.92 37.5 3.22, 3.19
2.38 32.2 1.45 23.2 1.70 25.5 — 129.5 2.03 25.6 — 137.8 — 129.5 —
1.70 27.6 3.32 41.8 7.14 (1) 119.0 3.68, 3.65 48.3 7.30 130.4 7.15 131.8 7.24 (1)
55.7
28.1
110.3
125.9
Gly Val Leu Ile
Ser Thr Cys Met Asp Glu Asn Gln Lys Arg His Pro trans Phe Tyr Trp
2.13 31.4 1.65 41.2 1.90
γ
0.97, 0.94 19.6, 18.6 1.64 25.7 0.95 (2), 1.48, 1.19 (1) 15.8 (2), 25.5 (1)
δ
ε
ζ
η
0.94, 0.90 23.6, 22.1 0.89
11.1
1.23 20.0
2.64 30.6
2.13 15.5
2.31, 2.28 34.9
3.02 40.4
8.12 (1) 135.2
7.39 129.9 6.86 116.7 7.65 (3)
7.34 128.3 — 155.5 7.17 (3), 7.24 (2) 7.50 (2) 137.6 (2), 113.2 (2), 119.8 (2) 120.7 (3) 123.2 (3)
aNomenclature as in IUPAC-IUB Commission on Biochemical Nomenclature (1970). 1H shifts taken from Wüthrich (1986) and shown in normal text. 13C shifts from Richarz and Wüthrich (1978) and shown in bold. bSpecific atoms (α, β, γ...) of the amino acid substituent (R) groups are shown in Figure 17.5.2. For full amino acid names and one-letter codes, see APPENDIX 1A.
Introduction to NMR of Proteins
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β
γ 1
CH3
β
γ 2
γ
γ 2
δ 1
β
δ
β
γ 1
δ 2
Ala
Val
Leu
Ile
γ β β
OH Ser γ β
γ
Glu
γ
δ
δ
NHη
Lys O
β
1 η NH22
NH ζ ε
γ
δ N 1
ε
NH ε2
δ 2
β
His β ε
γ
δ
ζ
N
α
1
Arg δ
NH2
ε
γ
Gln
δ
γ
β CONH2
β
CO2H δ
β
Asp
Asn
β
β CO2H
CONH2
γ
Cys
ε
β
SH
Thr
SCH3
Met
β
β
OH
δ γ
Pro ε β
ε ζ OH
3
γ δ1
δ2 NH ε2 ε
ζ
3
η2 ζ
2
1
Phe
Tyr
Trp
Figure 17.5.2 Substituent (R) groups of amino acids. For Gly (not shown), R = H.
cation of the local secondary structure. For instance, upfield shifts (0.1 to 0.3 ppm) are generally observed for Hα protons in α helices, whereas these protons in β-sheet structures are shifted by similar amounts to lower field. This is more statistically relevant when groups of sequential residues exhibit shift deviations in the same direction, and is the basis of a method of predicting the presence of secondary structure called the chemical shift index (Wishart et al., 1996). The effect of secondary structure is also manifest by deviations of Cα and Cβ shift, as summarized in Figure 17.5.1B.
Scalar Coupling
The nuclear spin state (i.e., α or β) of one nucleus can be transmitted through the intervening bonding electrons to another nucleus up to
four chemical bonds away. The energy levels of each of the interacting nuclei are mutually perturbed (split), resulting in a splitting in the NMR resonance of each of the interacting nuclei. The effect is referred to as J, scalar, spinspin, or through-bond coupling, and its magnitude is called the coupling constant and given the symbol J. The symbolism nJAX is widely used, where n is the number of bonds separating the coupled nuclei, and AX shows that the coupling is between nucleus A and nucleus X. It can be shown that a nucleus coupling to n identical spin-1⁄2 nuclei is split into n + 1 lines, each separated by J and having intensities given by the binomial coefficients. J is always measured and quoted in Hz, not ppm. This interaction between spins in a molecule is of great use in spectral assignment, forming the basis for the
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0 - 18 H′β Hβ 0 - 12
χ
Cγ 2
140
30 Cβ H
30 χ
O
1
Hα ψ
Cα
7 N
55
11
15
φ
ω
N
Cα
Cα
95 O
H
average value of J in Hz
definition of dihedral angles
Figure 17.5.3 Average coupling constants (J in Hz) between nuclei in the peptide backbone and side chain are shown using bold numerals on the bond between the nuclei (e.g., 1JN-Cα = 11 Hz) or on the arrows connecting two nuclei (e.g., 2JN-Cα = 7 Hz); see text for nomenclature. The accepted nomenclature for dihedral angles is shown using light Greek characters (e.g., the angle between NH and Hα is given by φ).
majority of modern NMR experiments. Scalar coupling can be observed between any two nuclei if they are within several chemical bonds of each other and have a spin >0. The magnitude of the coupling will depend on the number of intervening bonds and the isotopes that are coupled. Figure 17.5.3 shows approximate values of the couplings possible in the peptide backbone. The three-bond (vicinal) proton-proton coupling constant is a function of the dihedral angle between them, obeying the well-known Karplus relationship (see Macomber, 1998; Friebolin, 1993), with the largest value being observed for a dihedral angle of 180°. Thus, if scalar coupling constants between protons in a protein can be measured, useful information about conformations can be obtained. For instance, angles φ and χ1 could be estimated by measuring 3JNH,Hα and 3J Hβ,Hα, respectively (Fig. 17.5.3).
Relaxation Times: T1, T2, τc, and the nOe
Introduction to NMR of Proteins
The excited spin state responsible for the NMR signal eventually returns to the equilibrium ground state by a process called relaxation. At first sight this may seem such a trivial occurrence that it should be ignored. However, the
relaxation process and, more importantly, its rate greatly influence NMR spectroscopy of proteins. There are two key relaxation mechanisms. Longitudinal relaxation, also called spin-lattice relaxation, is a measure of how easily an excited spin can exchange its excess energy with the surroundings (lattice). Transverse relaxation, also called spin-spin relaxation, is a measure of the efficiency of the exchange of energy between spins. Each of these relaxation processes is generally characterized by the reciprocal of the appropriate relaxation rate—i.e., the longitudinal relaxation time, T1, and the transverse relaxation time, T2. In complete isolation, a nuclear spin would have very long relaxation times, but relaxation within molecules can be promoted by exchange of energy with the lattice or other spins in the same molecule. If a nucleus experiences a fluctuating magnetic field at the Larmor frequency, then relaxation by the T1 mechanism is efficient. The relaxation rates are specifically related to molecular motion. Frequencies slower or faster than the Larmor frequency are not as efficient in promoting T1 relaxation, but motions slower than the Larmor frequency do contribute to a decrease in T2 (i.e., promote T2 relaxation). These motions are related to the reorientation
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Fast motion (small molecules) τc short, ω2τc2 << 1
Slow motion (e.g. proteins) τc long
in NOESY, cross peaks are opposite phase to diagonal; nOes positive
in NOESY, cross peaks are same phase as diagonal; nOes negative
nOe
ωτc
extreme narrowing limit
spin diffusion limit
Figure 17.5.4 The variation in the maximum theoretical nOe as a function of correlation time (τc) and spectrometer frequency (ω). The curve passes through zero, separating the regimes of small (left) and large (right) molecules. Proteins generally have long correlation times and exist in the “spin diffusion limit”.
time of the molecule, also called the correlation time, τc. A way of thinking about the correlation time of a molecule is the time taken by the molecule to rotate through an angle of one radian. For small molecules, this can be picoseconds, whereas for proteins the timescale is nanoseconds. A large molecule like a protein tumbles slowly in solution, and T2 relaxation is efficient. Short T2 values mean that NMR resonance lines become broad (Heisenberg broadening) and that transverse magnetization quickly decays. The magnetic fields produced by two spinning nuclei can mutually interact if they are close enough in space. This effect is called dipolar coupling, and should not be confused with J coupling described above (see Scalar Coupling), which was an effect solely transmitted through bonding electrons. The nuclear Overhauser effect (nOe) is a through-space interaction mediated by dipolar coupling (i.e., a mutual interaction of magnetic fields) and is the basis of one of the most important homonuclear NMR techniques (Neuhaus and Williamson, 1989). Two protons will yield a mutual nOe if they are close enough to experience each other’s magnetic dipole moment (in practice, ≤5 Å apart in space). If a nuclear spin is perturbed from its equilibrium Boltzmann population, the spin will recover to equilibrium by relaxation. The rate of relaxation may be enhanced by the proximity of a magnetic field fluctuating at the correct frequency (i.e., the NMR transition frequency), as in the case of a nearby nucleus undergoing an NMR transition. The increase in relaxation
rate by one nucleus due to the presence of another nucleus close in space is called the nuclear Overhauser effect. A later section discusses how these nOes can be used to aid spectral assignment, and how they can act as indicators of secondary structure in proteins; see Nuclear Overhauser Effect Spectroscopy (NOESY) and Rotating-Frame Nuclear Overhauser Effect Spectroscopy (ROESY). The nOe is dependent on the correlation time (τc) of the molecule under investigation. Figure 17.5.4 shows the variation in the maximum theoretical nOe between two protons as a function of correlation time multiplied by the spectrometer frequency. For small molecules nOes are positive, with a maximum value of 0.5 (or 50% enhancement, although this is very rarely achieved in practice). Large molecules, such as most proteins, exhibit negative nOes. Notice that the curve in Figure 17.5.4 goes through zero (“zero crossing”). For molecules in this unfortunate regime, no nOes would be observed between two protons, no matter how close in space they are. Notice also in Figure 17.5.4 the nomenclature commonly used when discussing nOes—e.g., large molecules are said to be in the “spin diffusion limit.” The reader interested in a more detailed and in-depth discussion of NMR theory is referred to some of the fine texts currently available (Derome, 1987; Friebolin, 1993; Macomber, 1998). Additionally several excellent texts discuss the application of NMR to biomolecules (in particular, Roberts, 1993; Evans, 1995). Structural Biology
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SINGLE-DIMENSION PROTON SPECTRA
Introduction to NMR of Proteins
To illustrate the above principles, it is useful to consider some real proton NMR spectra. Figure 17.5.5A shows a proton NMR spectrum of a synthetic pentapeptide Tyr-Ala-Gly-PheMet (YAGFM; Sigma; Fig. 17.5.5E) dissolved in 2H2O. The chemical shift reference is TSP, the singlet resonance at 0 ppm. The NMR resonances are very sharp, owing to the long T2 values characteristic of such small molecules. The lowest field resonances (shown expanded in Fig. 17.5.5B) are due to the aromatic residues Tyr and Phe, each distinct and nonoverlapped. The doublet at high field (1.33 ppm) is due to the three Ala β-methyl protons. The α proton region is expanded in Figure 17.5.5C, showing the near overlap of Tyr and Met α protons. Of particular note in this spectrum are the Phe β protons (Fig. 17.5.5D), which are not equivalent and which resonate at two distinct frequencies (3.17 and 3.01 ppm). This magnetic inequivalence is caused by the fact that these protons are proximate to a chiral center, the α carbon atom, such that each of the protons experience different magnetic environments for all possible conformations (Fig. 17.5.6). The double-doublet nature of these resonances is due to the three scalar couplings 2Jββ′ (13.9 Hz), 3Jβα (8.6 Hz), and 3Jβ′α (5.9 Hz), as indicated. The Phe β protons are said to be diastereotopic. In a similar manner, the two α protons of glycine are diastereotopic, resonating at 3.8 and 3.9 ppm, and coupling together with a large (17 Hz) two-bond (geminal) coupling constant (Fig. 17.5.6). Because the spectrum in Figure 17.5.5 was recorded in 2H2O, no exchangeable protons (OH, NH) are observed, as these exchange with the deuterons in the solvent. The scalar coupling that would be present between the NH protons and the α protons of all amino acids is thus not present. In contrast, Figure 17.5.7 shows the one-dimensional proton NMR spectrum of a protein considered to be of medium size for NMR, the calcium binding protein calmodulin (148 residues; 17kDa), at a concentration of ∼1.5 mM, obtained at a temperature of 310K at 500 MHz. The spectrum in Figure 17.5.7 was recorded in H2O, so that the exchangeable protons, especially the amides, remain visible. The resonance from the large excess of water protons (present at 56 M) is partially suppressed, leaving a residual signal at ∼4.8 ppm. The annotations on Figure 17.5.7 show approximate chemical shift ranges for typical functional groups or environments.
Resonance widths of ∼15 Hz typify a protein of this size having a correlation time of ∼9 nsec. Homonuclear (i.e., proton-proton) scalar couplings are not directly observed, as the linewidth is greater than nJHH. The resonances at the lowest field represent individual amide protons, which are deshielded due to their nonbonded environment. Noncovalent interactions and hydrogen bonding present in the tertiary structure of the protein result in the observed dispersion of chemical shifts. As calmodulin is mostly α-helical, the majority of its α protons resonate in a narrow band (∼130 α protons resonate between 3.1 and 4.8 ppm). Small regions of β sheet result in the deshielding of several α protons, which resonate downfield of the water signal.
SPECTRAL ASSIGNMENT OF PEPTIDES AND SMALLER PROTEINS—HOMONUCLEAR METHODS The first step in any NMR determination of structure is resonance assignment, in which NMR signals are correlated to specific nuclei in the protein. All subsequent structure calculations rely on these assignments, so the more complete and (especially) accurate this procedure is the better. Even if a full structure determination is not the goal of the study, when armed with resonance assignments, it is possible to (1) interpret structural changes in the protein following binding of small molecules or metal ions (i.e., characterize binding events), (2) classify secondary structural elements in the protein, and (3) gain information about solution dynamics. For smaller proteins and peptides, homonuclear (i.e., proton only) NMR assignment methods are usually adequate. The cutoff point for which homonuclear assignment is applicable cannot be clearly defined, but depends on a number of factors other than molecular weight. For instance, if the protein is mainly composed of α helix or β sheet (which may be determined, for instance, from circular dichroism or infrared measurements), then the chemical shift dispersion of the alpha proton signals may not be sufficient to allow unambiguous assignment. Additionally, if the protein has a large number of the same residue type, or it is only partially folded, then degeneracies in chemical shift may preclude full assignment. The strategy often employed is that developed by Wüthrich (1986). The steps in this “sequence-specific” assignment procedure can be summarized as (1) assignment of an NMR resonance at a particular frequency to a specific proton in an amino acid (e.g., Hβ of a serine
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A Met ε
acetate
HOD
TSP
Ala
Gly α Met δ Met β
7
6
5
4
3
2
1
0
ppm
B
C Phe δ Phe ζ
Tyr δ
D
Tyr ε
Tyr β
Phe ε
3J
Phe α
2
Met α Tyr α Ala α
αβ
2J
J ββ′
ββ′ 3J
αβ′
Phe β 7.5 7.4 7.3 7.2 7.17.0 6.9 6.8 6.7 ppm
4.6 4.54.4. 4.3 4.2 4.1 ppm
3.25 3.15 3.05 2.95 2.85 ppm
ε
E
S β
O
ε
δ
O α NH2
γ
O
NH α β
β
O α
α
NH
NH
α β
O
H
NH
O
OH
δ
ζ
ε
Figure 17.5.5 One-dimensional 400.13 MHz NMR spectrum of the synthetic peptide YAGFM (2 mg) dissolved in 2H2O, recorded at a temperature of 300K. The full spectrum is shown in (A). The aromatic region is expanded in (B) and the α-proton region in (C). Expansion (D) details the coupling constants responsible for the observed splitting of the diastereotopic β-methylene protons of Phe-4. The structure of YAGFM (including the designation of α, β, γ, etc. protons) is shown in (E).
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17.5.11
Hβ′
phenyl CO –
NH –
CO –
Hβ′
Hβ
Hβ
Hβ CO –
NH –
phenyl
NH –
Hβ′
phenyl
Hα
Hα
Hα
I
II
III
Figure 17.5.6 Newman projections drawn along the Cβ–Cα bond of phenylalanine. The β protons in conformations I to III are never equivalent, and thus resonate at different chemical shifts. These protons are diastereotopic.
methyl protons in Val, Ala, Thr, Ile, Leu
deshielded amides
Met ε methyl
10.5
10.0 ppm
α protons in α helix and random coil β protons
amide & aromatic TSP
α protons in β sheet
10
9
8
7
6
5
4
3
2
1
ppm
Figure 17.5.7 One-dimensional 500.13 MHz NMR spectrum of calcium-saturated calmodulin (∼12 mg) in H2O, recorded at 310K. The annotations indicate approximate chemical shift ranges encountered in proteins. Introduction to NMR of Proteins
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residue), and (2) assignment of that amino acid to a specific position in the primary sequence (e.g., serine-25). Once this level of assignment has been achieved, it is then necessary to determine the location of each residue in space, relative to other residues in the sequence. The assignment techniques used encompass a range of two-dimensional NMR experiments, which are discussed below in terms of what information is available and how the data can be interpreted. As this technique may be unfamiliar to some, a brief discussion of two-dimensional NMR principles is presented. A detailed discussion of the mechanism of magnetization transfer and the pulse sequences used for two-dimensional NMR are beyond the scope of the present work; the interested reader is referred to some of the many articles on this topic (Friebolin, 1993; Croasmun and Carlson, 1994). In addition, for a more in-depth discussion of homonuclear assignment than is possible here, see Basus (1989) and Redfield (1993).
Two-Dimensional NMR A one-dimensional NMR spectrum is obtained by the sequential process: preparation, detection. For the simplest one-dimensional NMR experiment, the preparation period would consist of a single radiofrequency pulse, used to excite the nuclear spins as described above (see Principles of NMR). During the detection period, the excited magnetization evolves with respect to time and induces a signal in the receiver, called the free induction decay (FID). A two-dimensional experiment would have, in addition, an evolution period, during which the magnetization generated by the preparation sequence is “frequency labeled” by evolution of transverse magnetization with respect to a time variable (t1). Because two time variables are involved in two-dimensional NMR spectroscopy, it is conventional to describe the evolution time as t1 and the acquisition period as t2. In many two-dimensional experiments, a mixing period forms part of the overall sequence, which becomes: preparation, evolution (t1), mixing, detection (t2). The preparation period may simply be a single radiofrequency pulse, depending on the particular experiment, but transverse magnetization will always exist at the end of the preparation period. The mixing period is often the characteristic of a particular type of experiment (e.g., COSY, NOESY; see below), and the detection period is identical to that in one-dimensional NMR.
In the process of generating a two-dimensional spectrum, a series of one-dimensional data is generated in which each spectrum is associated with a particular value of the time variable t1. The t1 period is systematically incremented by an amount n∆t1, where ∆t1 is the increment and n is the number of the experiment. At the end of the experiment, a series of n FIDs is stored to disk. Fourier transformation with respect to t2 yields a series of n one-dimensional NMR spectra, whose amplitude (or phase) will vary as a function of the t1 value used. A further Fourier transformation with respect to t1 results in a two-dimensional matrix, consisting of two independent frequency axes (f1 and f2) and one intensity axis. It is conventional to display the data such that the f2 axis is horizontal and the f1 axis is vertical, with intensity information encoded by contours that join points of equal amplitude (e.g., Fig. 17.5.8). The f2 axis is the result of Fourier transformation of signals induced in the receiver, and is called the directly acquired dimension; those dimensions derived from the incrementation of a time variable (e.g., f1) are referred to as indirect dimensions. Most homonuclear two-dimensional spectra have a diagonal (about which they are symmetrical), in which the f1 frequency coordinate is identical to that in f2. This is the result of “selfcorrelations,” for which no mixing (or coherence transfer) has occurred. The most important features of two-dimensional NMR spectra are the off-diagonal elements. These represent interactions (called correlations or cross peaks) between nonidentical protons. The actual interaction mapped in this way will depend on the experiment chosen, as discussed below.
Correlation Spectroscopy (COSY) A COSY experiment correlates protons that are mutually J (or scalar) coupled. The coordinates of the off-diagonal elements of the COSY matrix represent the chemical shifts of J-coupled protons. A proton may, of course, be coupled to n chemically distinct protons, in which case there will be n cross peaks in the COSY spectrum originating from the frequency coordinate of that proton. The COSY spectrum detects J couplings over, generally, two to four bonds. There are several methods of recording COSY spectra. These range from the simple COSY-90, which yields little more than qualitative connectivity information, to the more complex phase-sensitive methods, such as double quantum filtered phase-sensitive COSY (DQF-COSY), which can yield values of coupling constants (Neuhaus et al., 1985).
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a c b 1.5 M(α–β) 2.0
2.5
M(β–γ) F(β–β′)
ppm
F(α–β′) 3.0
Y(α–β) F(α–β) G(α–α′)
3.5
4.0
4.5
5.0 5.0
4.5
4.0
3.5
3.0
2.5
2.0
1.5
ppm
Figure 17.5.8 Two-dimensional 400 MHz COSY-90 spectrum of YAGFM (2 mg) dissolved in 2H2O, recorded at a temperature of 300K. The spectrum is presented as a contour plot. A “high-resolution” one-dimensional spectrum of the peptide is plotted along the upper f2 (horizontal) axis to aid interpretation. Annotations of most correlations are indicated; for a discussion of a through c, see Correlation Spectroscopy (COSY).
Introduction to NMR of Proteins
As a simple illustration of the interpretation of a two-dimensional spectrum, Figure 17.5.8 shows a COSY-90 spectrum of the synthetic pentapeptide YAGFM discussed above (see Single-Dimension Proton Spectra). The analysis starts by looking at the correlations from the alanine methyl doublet at 1.31 ppm on the top spectrum. This resonance is located in f2, and a line (a) is drawn down to the diagonal. A line (b) drawn parallel to f2 that intersects the diagonal at the f1 coordinate of this peak also meets an off-diagonal peak (c). This peak represents
the three-bond coupling of the Ala β methyl and its α proton; its coordinates are the frequency of Ala-α(f1) and Ala-β(f2). These two resonances have thus been correlated (i.e., a connection has been established between them). There are no further correlations evident (or indeed expected) for this residue. All other correlations are annotated on Figure 17.5.8. There are no J couplings evident between the Met γ and Met ε protons, or between the β and δ protons of the aromatic residues Tyr or Phe.
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3.5 3.6 I-28
3.7 3.8
I-31
3.9 R-25 4.0
T-32
Q-34
4.1
R-35 A-23
ppm
R-33
4.2
L-30
4.3
Y-21
S-22
Y-27
Y-20
4.4
L-24
N-29 H-26
4.5 4.6 4.7 Y-36 4.8 4.9 5.0
8.8
8.7
8.6
8.5
8.4
8.3
8.2
8.1
8.0
7.9
7.8
7.7
7.6
7.5
7.4
7.3
ppm
Figure 17.5.9 Expansion of the fingerprint region of a DQF-COSY spectrum recorded on the C-terminal domain of NPY (residues 18 to 36) at 400 MHz. Each cross peak represents the correlation between an amide NH proton in f2 (horizontal) and its α proton in f1 (vertical). The bold boxes are drawn around those correlations that are partially overlapped. Each correlation shows a pattern of four peaks, with the separation between these peaks being representative of the coupling constant. This is a property of the DQF-COSY experiment that is not exemplified in the “simple” COSY spectrum shown in Figure 17.5.8.
For a more demanding example, a discussion is given for spectra of the 19-residue C-terminal domain of neuropeptide Y (NPY), a 36-residue neurotransmitter present in large quantities in the brain. This 19-residue domain, referred to as NPY[18-36], is where much of the pharamacological activity of NPY resides. Its sequence is: A18-R19-Y20-Y21-S22-A23-L24-R25-H26-Y27I28-N29-L30-I31-T32-R33-Q34-R35-Y36. Part of a DQF-COSY spectrum of NPY[1836] is shown in Figure 17.5.9. This region, which shows NH protons in the f2 dimension and α protons in f1, is generally referred to as the fingerprint region (obviously this experiment must be carried out on a sample of the
protein dissolved in H2O). The correlations observed are due to 3
J
NH i H i α
and are between the NH proton of residue i and the alpha proton of residue i (i.e., they are intraresidue). There are no proton-proton interresidue J couplings possible. Seventeen out of a possible nineteen correlations are observed for NPY[18-36]; the amide proton resonances of the N-terminal residues Ala-18 and Arg-19 are not observed, presumably due to rapid exchange with solvent. Although not present in this example, glycine residues often yield two Structural Biology
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γ
A
δ
2
β
γ2 γ1
1
β
β
2
β
β,β '
α
β α
α α α
β
β
α
α
3
4
5
*
6
H-26
δ−ε
7
I-31
Y-21
Y-20
S-22 A-23 Y-36 Y-27
8
9
I-28
10 8.7
B
γ
8.6
8.5
8.4
8.3
8.2
8.1
8.0
7.9
7.8
γ2 γ
2
7.6
7.5
7.4
1
β
1
β
β
7.7
2
β α
α
α
β
β
α
α
3
4
5
*
6
7
dNN(i,i-1)
dNN(i,i+1)
Y-20
dNN(i,i-1) I-28 8.8
8.7
A-23 I-31
8.6
8.5
Y-21
9
dNN(i,i+1) Y-27
8.4
8.3
8.2
8
8.1
8.0
7.9
7.8
7.7
7.6
7.5
7.4
Figure 17.5.10 (A) A 100-msec TOCSY (400 MHz) spectrum of the C-terminal domain of NPY[18-36]. The NH-Hα correlations are identified from the COSY experiment in Figure 17.5.9. For each NH resonance in f2 (horizontal), correlations are observed to Hα, Hβ, Hγ (as appropriate). A correlation between the δ and ε protons of the single histidine residue is also seen in this region of the spectrum. (B) A 400-msec NOESY spectrum of the C-terminal domain of NPY[18-36]. Cross peaks that are observed in addition to those in the TOCSY spectrum (A) are due to the spatial proximity of protons in different residues: for instance, the sequential nOes between Ile-28 NH and the amide protons of Asn-29 and Tyr-27 (marked dNN(i, i + 1) and dNN(i, i − 1), respectively) and the nonsequential nOes between NH’s and aliphatic protons. Star (right-hand side of panels A and B) indicates correlations from Tyr δ to ε protons; all others originate from amide NH’s.
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dβN(i,i +1)
Hβ Cβ
dαN(i,i +1)
H O
O
Hα N
Cα N
dNN(i,i +1)
O
H
residue i
Hα
residue i + 1
Figure 17.5.11 Sequential nOes (curved arrows) expected in a peptide containing residues i and i+1.
correlations in the fingerprint region: from the NH to the diastereotopic methylene protons Hα and Hα′. Prolines lack an NH and are therefore always absent from the fingerprint region.
protons in addition, classified as type U (i.e., Arg, Glx, Lys, Met, and Pro).
Total Correlation Spectroscopy (TOCSY)
Nuclear Overhauser Effect Spectroscopy (NOESY) and Rotating-Frame Nuclear Overhauser Effect Spectroscopy (ROESY)
A TOCSY experiment has many similarities to a COSY experiment: for instance, correlations depend on J coupling (Braunschweiler and Ernst, 1983). However, in a TOCSY experiment, a correlation may be observed between proton “a” and proton “d” even though there is no direct coupling between them (nJad = 0), as long as the coupling between each successive member of the spin system is not zero (i.e., Jab ≠ 0, Jbc ≠ 0, Jcd ≠ 0). The experimenter uses the mixing time to control how far down the spincoupling chain the correlations can be observed. For instance, a TOCSY with a mixing time of ∼10 msec yields only correlations from directly coupled spins (as in COSY); however, in a 100-msec TOCSY, it is possible that all protons in the spin system may be correlated (i.e., all aliphatic protons and the amide proton may be mutually correlated). Figure 17.5.10A shows a TOCSY experiment recorded on the NPY[1836] peptide. TOCSY data recorded with long mixing times yield unique correlation patterns for Ala, Gly, Ile, Leu, Thr, and Val. The remainder of residue types can be put into two categories: those with a four-spin system (NH, Hα, Hβ, Hβ′), classified as type J (i.e., Asx, Cys, His, Phe, Ser, Trp, and Tyr), and those that have γ (or more)
A brief introduction to the nOe was given above (see Relaxation Times: T1, T2, τc, and the nOe). NOESY (Kumar et al., 1980) is a two-dimensional experiment correlating protons ≤5 Å apart in space. The experiment is thus a probe of local structure, potentially revealing information about conformation and protein folding. nOe forms an important part of the sequential assignment procedure, making the link between protons across the peptide bond. Figure 17.5.11 shows the nOes expected between the sequential residues i and i+1. The accepted nomenclature for nOes in proteins is dAB(i,j), in which the nOe is between protons A and B, which are located within residues i and j, respectively. nOes can be of great value in resonance assignment. For instance, nOes are frequently observed in His, Phe, Trp, and Tyr residues between a β-methylene proton and an aromatic proton. Additionally, the primary amido protons of Asn and Gln often yield nOe correlations to the β- or γ-methylene protons, respectively, of their own side chains. nOes can also be of great utility in the determination of secondary structure. For instance, Figure 17.5.12A and 17.5.12B show nOes typical in α helices and β sheets, respectively. In the
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O
A
to C terminus
O N
O
H
N H dNN(i,i+1)
H N
dαβ(i,i+3)
O
dαN(i,i+3)
O
H
N H
N
to N terminus
H
B
H
dαN(i,i+1)
H
O
N N
N
H
H
O dNN(i, j)
O
dαα(i,j)
H
H dαN(i,j)
H
O
N N
N
H
H
O
H
Figure 17.5.12 Summary of longer-range interresidue nOes (curved arrows) expected between residues in (A) α-helical and (B) β-sheet structures. Dashed arrows in (B) show antiparallel directions of the two peptide chains.
Introduction to NMR of Proteins
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case of β-sheet structures in particular, nOes can be observed between protons separated by many residues in the primary sequence. Similarly, the tertiary fold of a protein may well result in the spatial proximity of protons, even though they are far apart in the linear sequence. The mixing time (τm) is an important parameter in a NOESY experiment, as it is during this time that protons undergo cross-relaxation (i.e., exhibit the nOe phenomenon). Too short a mixing time results not only in small cross peaks, but also in artifacts that distort cross-peak intensity. Too long a mixing time can result in spin diffusion, an undesirable “relay” of the nOe, resulting in cross peaks between protons that could potentially be >>5 Å apart in space. Spin diffusion reduces cross-peak intensity and precludes quantitative geometric interpretation. Cross-peak intensity (or volume) increases roughly linearly with mixing time up to a point, and then starts to decay. The initial nOe build up is proportional to r−6, where r is the internuclear distance. In practice, a series of NOESY spectra are acquired with increasing mixing times, and the cross-peak volumes are plotted against the mixing time. Cross-peak volume ratios can be used to give quantitative relative distances if they are obtained from an experiment with a τm in the linear regime. Especially useful in this respect are the δ and ε aromatic protons of Tyr residues, whose mutual nOe cross-peak volume yields a reference distance for the calculation of other interproton distances. Figure 17.5.10B shows a 400-msec NOESY experiment recorded on the NPY[18-36] peptide. Intraresidue through-space interactions can be identified by comparison with the TOCSY data (Fig. 17.5.10A). Additional correlations yield the most useful information about the relative distances of different parts of the peptide. Sequential nOes are observed, for instance, from Ile-28 NH to the amide resonances of Tyr-27 and Asn-29, as marked on the spectrum. The two-dimensional ROESY experiment (Bothner-By et al., 1984; Bax and Davis, 1985) is analogous to NOESY in that correlations occur between protons that are close in space. However, in the ROESY method, all molecules behave as if they were in the extreme narrowing limit, despite molecular size or correlation time, and thus there is no zero crossing point. True ROESY correlations (i.e., through-space interactions) are always positive (opposite in phase to the diagonal), but of lower absolute intensity than the analogous NOESY correlations. How-
ever, negative correlations may be observed in a ROESY spectrum, and arise from TOCSYtype transfer or from chemical exchange. The latter process is a consequence of the exchange of (chemical) environment, for instance, during the mixing time of the experiment. The phenomenon of chemical exchange can be of great utility in the determination of dynamic processes of proteins in solution. For examples, see Fejzo et al. (1991) and Lian and Roberts (1993).
NMR Characteristics of Secondary Structural Elements The identification of secondary structure in a protein can be achieved by several complementary methods. The chemical shift perturbations of the alpha proton (and of Cα and Cβ, if available) of an amino acid can be characteristic, as seen above (see Chemical Shift). Amide NH protons involved in hydrogen bonding (the interaction that stabilizes α helices and β sheets) usually have much slower proton exchange rates than those in regions of little structure; these rates of exchange are usually measured by dissolving the protein in 2H2O and rapidly recording spectra as a function of time. Hydrogenbonded amide protons show a lower temperature dependence of chemical shift than those that are not hydrogen bonded. The NH-Hα coupling constant is a function of the dihedral angle between them (φ); this coupling is small (3 to 5 Hz) in α-helical structures, whereas it is usually much larger (8 to 12 Hz) in β sheets. As seen in the above section and in Figure 17.5.12, certain short-range nOes are characteristic of α helices and β sheets, and it is the observation of these correlations that gives the most definitive method of characterizing the location and boundaries of secondary structure. It is common practice to summarize the NMR data (as it defines secondary structure) as shown in Figure 17.5.13.
Toward the Structure: Putting It All Together Each resonance assignment experiment can also yield information about the three-dimensional structure of the protein itself. For instance, coupling constants revealed by one-dimensional experiments or two-dimensional methods based on COSY (e.g., DQF-COSY) constrain the dihedral angle between the coupled protons. The main structural constraints revealed by NMR are nOes, J coupling, and chemical shift. nOes reveal much about the overall protein folding (as well as secondary structure; see above discussion of NOESY and
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sequence A D F Q R RS T A R E K R E A I S V Q R A QD D S E E N S F Q Y R H E D A N 3J u 69445u 4544799 7 u 989uu99 8 7 8u9 445 444 489 8 (Hz) NH,α NH exchange
dαN(i,i+3)
dαβ(i,i+3)
dαN(i,i +1) dNN(i,i +1) dβN(i,i+ 1) chemical shift index secondary structure
1 –1 α helix
β sheet
u
α helix
slow exchange fast exchange strong medium weak unobservable
Figure 17.5.13 Usual presentation of NMR information regarding secondary structure, illustrated using a fictitious peptide and data. Coupling constants between amide NH and Hα (intraresidue) are shown in Hz. Fast or slow amide exchange rates are shown with open or closed circles, respectively. Long-range (i,i+3) nOes between two residues are shown using horizontal bars connecting those residues, with line thickness used to indicate the relative size of these correlations. Shorter-range (i,i+1) nOes are shown as blocks representing large, medium, and small magnitudes. The chemical shift index represents the deviation of Hα shift from that found in random coil structures; a sequence of +1 values (representing an actual deviation of +0.3 and above) is representative of β-sheet structure, whereas the converse is representative of α helices.
Introduction to NMR of Proteins
ROESY), and are generally not used to determine accurate interatomic distances, but rather to provide upper distance bounds. Powerful software programs are available that calculate three-dimensional structures based on the satisfaction of NMR constraints. Usually, after all available constraints have been employed in the calculation, a “family” of structures is obtained. Each member of this family will have a slightly different structure, but will satisfy the NMR constraints within a certain tolerance. The quality of structures generated by these methods is generally quoted in terms of the maximum deviation within the family from the “average” structure (actually given as the root mean square deviation, RMSD). “Good” quality structures will have backbone RMSDs on the order of 0.5 Å. The interested reader is referred to the following articles for a more detailed
discussion of this topic: Sutcliffe (1993), Weber (1996), and Güntert (1997).
SPECTRAL ASSIGNMENT USING DOUBLE-RESONANCE HETERONUCLEAR METHODS Assignment of larger (>10 kDa) proteins is not well suited to homonuclear methods. The number of resonances increases monotonically with the number of residues, and even correlations observed in two-dimensional NMR spectra frequently overlap, precluding assignment. The overlap problem is exacerbated for proteins with a high degree of α-helical content, in which the dispersion of the Hα proton resonances is generally low. The widths of the proton signals increase with the size of the protein, due to the shortening of T2 relaxation times, as the rotational correlation time (τc) increases, leading to
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reduced sensitivity for homonuclear correlation experiments, particularly COSY. Incorporation of NMR-active stable isotopes allows the use of large heteronuclear couplings to transfer magnetization (see Fig. 17.5.3), leading to experiments with high sensitivity. Generally, NMR correlation experiments that rely on a coupling of J Hz require a delay of (2J)−1 sec to fully transfer magnetization. Small couplings (i.e., nJHH) result in large (2J)−1 times, during which magnetization (i.e., signal) will be attenuated by T2 relaxation. Two-dimensional NMR techniques can be extended into three or more dimensions, any of which could be a heteronuclear frequency dimension. Spreading correlations into the heteronuclear dimension partially circumvents the resonance overlap problem, as the chemical shift dispersion of heteronuclei is generally much greater than that of protons. NMR of proteins uniformly and completely labeled with the nitrogen-15 isotope (U-15N) can exploit the large one-bond 15N-1H coupling of ∼90 Hz to establish correlations. Applications include isotope-edited correlation experiments, such as three-dimensional 15N/1H-TOCSYHMQC and NOESY-HMQC, which will be discussed below (see Isotope Editing: Two-Dimensional Heteronuclear Chemical Shift Correlations). These three-dimensional experiments have been successfully used in favorable cases—i.e., for smaller proteins (∼100 residues) or for larger proteins rich in β sheets (Marion et al., 1989)—to obtain nearly complete assignments of the protein backbone 15N and 1H resonances (Gronenborn et al., 1989). NMR of proteins labeled with both 13C and 15N (U-13C/15N protein) allows many additional heteronuclear couplings to potentially contribute to magnetization transfer (Fig. 17.5.3). The one-bond couplings are generally much larger than multiple bond 1H-1H couplings and greater than or comparable to 13C and 15N linewidths for most medium-sized proteins (i.e., <30 kDa). Unlike the techniques of TOCSY and NOESY, these experiments yield correlations that are independent of protein conformation.
Dispersion of 13C and 15N Spectra of Proteins The backbone amide protons of folded proteins generally resonate between ∼6 and 10 ppm, although strong noncovalent interactions can increase this limit by deshielding or shielding (e.g., the one-dimensional spectrum of calmodulin in Fig. 17.5.7 shows hydrogen-bonded amide protons at nearly 11 ppm). In contrast,
the nitrogen nuclei of backbone amides generally resonate between 105 and 130 ppm, a dispersion of nearly six times that of amide protons. Nitrogen NMR spectra are generally referenced with respect to liquid ammonia (defined as 0 ppm), using indirect means. Figure 17.5.14 shows a one-dimensional 15N NMR spectrum of U-15N Trypanosome calmodulin recorded at a field strength of 9.4 T (40.5 MHz for 15N observation). Carbon-13 chemical shifts cover a very large range (∼10 to 190 ppm). However, the dispersion of carbon atoms of similar type is much smaller (e.g., backbone carbonyl carbons generally resonate between 170 and 176 ppm). Figure 17.5.15 shows the one-dimensional proton-decoupled 13C spectrum of U-13C/15N Trypanosome calmodulin recorded at a field strength of 11.7 T (125.7 MHz for 13C observation). Multiplicities in this spectrum are caused by 13C-13C and 15N-13C coupling. Typical chemical shift ranges of different carbon types are indicated. As seen above (see Chemical Shift), 13C chemical shifts are sensitive to secondary structure. 13C chemical shifts are usually referenced internally to TSP or DSS at 0 ppm.
Isotope Editing: Two-Dimensional Heteronuclear Chemical Shift Correlations
The presence of a spin-1⁄2 heteronucleus at near 100% incorporation opens new doors to the spectroscopist. For instance, it is possible to selectively observe only those protons directly coupled to the heteronucleus (i.e., 1JXH ≠ 0, where X = 13C or 15N). It is equally possible to select all protons that do not have a one-bond heteronuclear coupling. These techniques are called isotope editing and filtering, respectively, and examples are shown in Figure 17.5.16 using a sample of U-15N Trypanosome calmodulin. Figure 17.5.16A shows the low-field region of the isotopically normal protein (also shown in Fig. 17.5.7), in which amide and aromatic protons overlap. The 15N-isotope-edited spectrum is shown in Figure 17.5.16B, in which only NH resonances are selected, by virtue of 1JNH. Figure 17.5.16C shows only those protons from aromatic residues (one His, one Tyr, nine Phe), as the NH protons have been filtered out. A similar spectrum to that shown in Figure 17.5.16C would be observed in the one-dimensional proton spectrum of the protein dissolved in 2H2O. It is possible to record isotope-edited spectra as two-dimensional matrices, in which the f1 dimension contains the heteronuclear chemical shift, e.g., the heteronuclear multiple and single
Structural Biology
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135
130
125
120 ppm
115
110
105
Figure 17.5.14 One-dimensional directly observed 15N spectrum of calcium-saturated U-15N-calmodulin, recorded at 40.54 MHz at 310K. Amide 15N resonances cover a range of ∼30 ppm. Generally, 15 N resonances are observed in “inverse” experiments such as the HSQC (Fig. 17.5.17), due to the low sensitivity of the directly observed experiment. This experiment took four times longer to acquire than the experiment shown in Figure 17.5.17.
C γ,δ Cβ backbone CO
aromatic
Cα
aliphatic methyl
1 amide CO Arg ζ
180
160
140
120
100 ppm
80
60
40
20
Figure 17.5.15 One-dimensional directly observed 13C spectrum of calcium-saturated U-13C/15N-calmodulin, recorded at 100.61 MHz at 310K with 1H decoupling. 13C-13C coupling is evident on all resonances. The chemical shift ranges of selected carbon environments in proteins are marked. Introduction to NMR of Proteins
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A
B
Y-138
H-107ε1
C
11.5
11.0
10.5
10.0
9.5
9.0
8.5
8.0
7.5
7.0
6.5
ppm
Figure 17.5.16 An illustration of isotope editing and filtering using calcium-saturated U-15N-calmodulin. Spectra were recorded at a proton frequency of 400.13 MHz at 310K; only the low-field regions of the spectra are shown. The spectrum shown in (A) is the 15N-decoupled proton spectrum (1H{15N}), in which all amide NH and aromatic resonances appear and overlap. The spectrum in (B) is the 15N-edited spectrum, in which only those protons directly attached to 15N are observed. The spectrum in (C) is the 15 N-filtered spectrum, in which all 15N-attached protons are filtered out or removed. Thus, in (C) only aromatic resonances remain; this is equivalent to recording the spectrum in 2H2O. Addition of spectra (B) and (C) would provide spectrum (A).
Structural Biology
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quantum coherence experiments (HMQC and HSQC, respectively). These powerful methods spread overlapped proton resonances into the heteronuclear chemical shift dimension, which is generally more disperse. These one-bond correlation experiments are the first step in the heteronuclear chemical shift assignment procedure. Figure 17.5.17 shows the assigned 1H-15N HSQC matrix acquired on a sample of calciumsaturated U-15N Trypanosome calmodulin at 9.4 T and a temperature of 310K. Even though a field strength of 9.4 T is considered to be at the low field end for protein assignment, each 1H/15N amide correlation is almost completely resolved, due to the fact that calmodulin is a highly structured protein. The boxed region in Figure 17.5.17A is expanded in Figure 17.5.17B.
Three-Dimensional NMR and 15 N-Edited Experiments
Introduction to NMR of Proteins
Two-dimensional heteronuclear correlation experiments can be combined with traditional homonuclear methods to yield extremely useful three-dimensional experiments, such as NOESY-HSQC and TOCSY-HSQC. Figure 17.5.18 shows such a three-dimensional experiment in schematic form. There are two proton dimensions and one heteronuclear dimension. To make the discussion less abstract, consider a 15N/1H NOESY-HSQC. A homonuclear two-dimensional NOESY experiment would map nOes from all protons in the protein, and would be severely overlapped (Fig. 17.5.18A). If the two-dimensional NOESY experiment also included an isotope editing sequence, then the resulting spectrum would be simplified, as only nOes from NH (i.e., amide) resonances would appear (Fig. 17.5.18B). If the HSQC part of the experiment is included by introducing the 15N chemical shift, this simplified two-dimensional spectrum is spread into a third dimension, according to the 15N chemical shift of those amides giving rise to the signal. Although the data can be viewed as a “cube” (Fig. 17.5.18C), it is more common (and actually easier) to display the data as two-dimensional slices or planes, each containing the proton-proton NOESY information (Fig. 17.5.18E). Each slice through the cube can be regarded as a slice through the two-dimensional HSQC spectrum (Fig. 17.5.18D); indeed, the projection of the three-dimensional cube onto the 1H/15N axis yields essentially the 15N1H HSQC data. Figure 17.5.19 shows one slice (taken at an 15N shift of 113 ppm) from a three-dimensional 15N/1H NOESY-HMQC experiment recorded
on U-15N Trypanosome calmodulin at 11.7 T and a temperature of 310K, with a NOESY mixing time of 100 msec. The diagonal in this spectrum contains those amide proton resonances whose directly bonded 15N atoms resonate at the 15N chemical shift shown. It is thus possible to “map” the local environment of the backbone amide protons in terms of their proximity to other groups in the protein. The nOes observed in this way can also be used as constraints to calculate the three-dimensional protein structure (see Toward the Structure: Putting It All Together). In a similar way, the threedimensional TOCSY-HMQC yields spin system correlations between the amide proton and the side chain. An excellent description of the methodology used can be found in Marion et al. (1989), showing the application of three-dimensional TOCSY- and NOESY-HMQC methods to the β-sheet protein interleukin 1β (153 residues).
SPECTRAL ASSIGNMENT USING TRIPLE-RESONANCE MULTIDIMENSIONAL HETERONUCLEAR METHODS When a protein is fully labeled with 13C in addition to 15N (U-13C/15N), many more techniques are available to the spectroscopist. All couplings in Figure 17.5.3 can effect magnetization transfer, and thus contribute to the resonance assignment process. The term “tripleresonance NMR” is applied to those experiments that use mainly one-bond heteronuclear scalar couplings for magnetization transfer (Ikura et al., 1990). The large values of heteronuclear scalar couplings (Fig. 17.5.3) ensure that the transfer of magnetization is particularly efficient, even for large molecules where linewidths can exceed even the largest homonuclear proton couplings. Resonance assignment strategies based on triple-resonance methods usually begin with the correlation of 15N and NH resonances (e.g., 15N-1H HSQC), as most triple-resonance experiments detect the NH signal and at least one other dimension encompasses the 15N chemical shift. These amide pairs are then correlated to nuclei in residues i and i−1, which provides the basis of the sequential assignment. The names given to triple-resonance experiments generally reflect the nuclei correlated in the experiment, using the following abbreviations: H and N for amide 1H and its directly bonded 15N, respectively; CO for carbonyl carbon; and CA and CB for Cα and Cβ, respectively. Figure 17.5.20 shows a graphical representation
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A 105.00
G-40
G-33
G-113 T-62 G-59
G-23
Q-135
V-55
G-96
110.00
G-132
G-98
T-44
T-29
G-134
T-26
S-17
G-25 T-117
115.00
G-61
120.00 K-13 A-102 S-101 I-63 I-27
I-100
F-68
L-116 K-21
K-115
F-141 I-136
125.00
K-94
Q-57 K-148
V-130
N-137 D-64
11.00
10.00
9.00
8.00
7.00
6.00
B T-117
D-22
D-95
114.00
T-110 S-10
F-19
Q-135
E-127
R-90
F-99
T-70
S-147 M-142 N-97 M-109 M-145 E-67 E-54 F-92 S-79 T-28 N-53 D-131 D-93 R-74 T-34 M-36 Q49 D-129 R-106 K-143 I-52 D-20 E-139 S-5 V-91 I-125 A-103 Q-41 I-108 F-89 F-16 Y-138 E-87 L-69 E-47 M-144 S-38 H-107 M-76 R-37 A-128 E-119 V-121 E-83 E-104 D-122 D-50 E-140 K-75 F-12 E-123 V-142 N-97 E-120 L-112 Q-3 E-7 Q-8 E-14 N-6 L-39 L-48 K-30 L-32 D-24 E-114 D-133 L-18 E-45 A-46 A-73 D-78 D-118 I-85 E-31 K-86 A-15 I-9 L-105 V-35 A-88 D-80 K-13 N-111 D-56 F-68 L-71 E-82 L-4 N-42
S-60
S-81
M-72
116.00
D-58
F-65
A-102
S-101
K-115 F-141
9.00
L-116
8.00
120.00
122.00
124.00
K-21
8.50
118.00
7.50
Figure 17.5.17 15N-1H HSQC recorded on calcium-saturated U-15N-calmodulin at a field of 9.4 T at 310K. The full spectrum is shown in (A), with annotations of the more disperse amide correlations. The region marked with a dotted box in (A) is expanded in (B). The assignments were accomplished using a U-13C/15N-calmodulin sample and a suite of triple-resonance experiments. Dashed lines in (A) and (B) connect the in-equivalent primary amido proton resonances of Gln-135 and Asn-97, respectively.
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B
A
D
E
15N
C
1H
1H
1H
1H
1H
1H
15N
1H
1H
1H
Figure 17.5.18 A schematic representation of three-dimensional 15N-edited NOESY experiment. The nonedited NOESY spectrum is represented in (A), showing correlations between all protons. Panel (B) shows the effect of 15N editing. The two-dimensional matrix is simplified (leading to less overlap), as only nOes originating from amide protons are visible. If the 15N-edited spectrum of (B) is combined with a third (15N) chemical shift dimension, a three-dimensional experiment results. This can be viewed as a cube (C) in which each slice (plane) represents a 1H/1H NOESY, containing as diagonal resonances only those amide protons having a particular 15N chemical shift. Panel (E) shows selected slices from the cube. Comparing each slice in (E) with the two-dimensional edited spectrum in (B) shows the spectral simplification afforded by this technique. Looking down on the cube (D; i.e., taking an 15N-1H projection of the data) yields essentially the 15N/1H HSQC data.
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of the nuclei that are correlated in five important triple-resonance experiments. Examples are taken from a study of the binding of calmidazolium (Fig. 17.5.21) to U-13C/15N Trypanosome calmodulin, in which sequential assignments were obtained using combinations of the complementary HNCO and HN(CA)CO experiments, and CBCA(CO)NH and HNCACB
experiments. An HBHA(CBCACO)NH experiment was used to derive the Hα and Hβ chemical shifts. Spectra are shown as planes taken through the three-dimensional data at the 15N shift specified. All data were acquired at a field strength of 11.7 T at a temperature of 310K. The HNCO experiment correlates the resonances Hi, Ni, and COi−1. The 15N-1H plane of
Nγi
Nγi
Nγi 2.00
{
Nβi –1
NH2βi Nαi –1 Nαi
{
}
Nβi
Nαi
Nαi
4.00 Nαi
Nαi Nαi
N-111
G-98 & G-25 6.00 S-17 T-29
G-98/ F-99δ
T-44
T-26
G-134 NNi + 1 L-18
NNi + 1 F-99
NNi + 1 Q-135
NNi – 1 D-24
NNi – 1 D-133
8.00
NNi – 1 F-16
NN G-25/T-26
10.00
10.50
10.00
9.00
9.50
8.50
8.00
ppm
Figure 17.5.19 A slice taken at 113 ppm from an 15N-edited NOESY-HMQC spectrum of calcium-saturated U-15N-calmodulin at 500.13 MHz at 310K. In this slice, eight amide protons appear (G-98, G-25, G-134, T-29, T-44, T-26, S-17, and N-111). The mutual correlations between the amide protons of Gly-25 and Thr-26 (solid box) appear symmetrically about the diagonal (dotted line), due to the fact that both of these amide protons appear in the same slice. The resonances of Gly-25 and Gly-98 are overlapped at the resolution obtained in the 15N dimension.
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the three-dimensional matrix is equivalent to a two-dimensional HSQC, whereas the 13C-1H plane contains the interresidue carbonyl resonances. The spectra are usually displayed as 13CO-1H planes, with successive planes having different 15N chemical shifts (cf. HMQC-
O
H
Hα
N
N Hα
H
Hα
N H
Hα i-1
H
N
N
O HNCA i
O
H
Hα
N
H
Hα
O HN(CA)CO i
Key stronger correlations
N
O HNCACB
i-1
N
N
H
Hα
H
Hα
H
Cα
i-1 H
H
Hα
N
i-1
Cα
N
Cα
O
Hα
O CBCA(CO)NH
Cβ
N Cβ
N
i
O
H
N
i
N Cβ
H
O HNCO
O Cα
H
Cα
i-1 H
TOCSY). Figure 17.5.22A shows a typical slice through the three-dimensional HNCO experiment recorded on a sample of U-13C/15N Trypanosome calmodulin, taken at an 15N chemical shift of 113 ppm. In this slice, the horizontal axis represents the 1H chemical shift of those NH
weaker correlations
i
Figure 17.5.20 Correlated nuclei in selected triple-resonance experiments discussed in the text (see Spectral Assignment Using Triple-Resonance Multidimensional Heteronuclear Methods). Each experiment results in detection of the amide NH resonance. Note particularly the complementarity of the HNCO/HN(CA)CO and the CBCA(CO)NH/HNCACB pairs of experiments.
24′
Cl
23′ 24
24′
23a′
Cl
H 23
21 23a
N 5
2
+ N
4
24a
20 H
6,6′
19
7
Cl
R 8,8′
O
Figure 17.5.21 The chemical structure of calmidazolium. Note the presence of the chiral center at C7, which is drawn with absolute configuration R. The optical rotation of R-calmidazolium is negative, and throughout the text this molecule is referred to as R-(−)-calmidazolium.
17 Cl
14
Cl
13 11 Cl
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protons having an 15N shift of ∼113 ppm, with the vertical axis being the 13C shift of the carbonyl carbon of the preceding residue. The complementary experiment to the HNCO is the HN(CA)CO, which correlates the same 15N-1H pair with the intraresidue carbonyl shift. Because this experiment relies on the small 15N13C coupling, its sensitivity is low. Actually the α HN(CA)CO experiment can yield both interand intraresidue correlations, but the former are generally much weaker in intensity, and are
often not observed. A slice (δ 15N = 113 ppm) taken from the HN(CA)CO experiment recorded on the same sample is shown in Figure 17.5.22B. The horizontal axis is the same as the HNCO, but the vertical axis (displayed on the same scale in both panels) contains the intraresidue carbonyl carbon shift. With the data in Figure 17.5.22, the 13CO chemical shifts of the carbonyl carbons of residue i and residue i−1 can be obtained for the 15N-1H pair of residue i. Because this information is available for most
HNCO
A
HN(CA)CO
B
172 G-98 i G-134 i 174
13C
(ppm)
G-25 i
T-44 i 176 G-98 i–1
G-98 i–1
T-29 i–1
G-25 i–1
178
G-25 i–1
10.5
10.0
T-44 i–1
T-44 i–1
G-134 i–1
11.0
G-134 i–1
9.5
9.0
8.5
11.0
10.5
10.0
9.5
9.0
8.5
1H(ppm)
Figure 17.5.22 Slices taken at an 15N chemical shift of 113 ppm through (A) HNCO and (B) HN(CA)CO three-dimensional matrices. Data were acquired at 500.13 MHz at a temperature of 310K on a sample of calcium-saturated U-13C/15N Trypanosome calmodulin. The sensitive HNCO triple-resonance experiment provides correlations between the amide proton of residue i (horizontal axis), the amide nitrogen of residue i (chemical shift axis, perpendicular to the plane of the figure), and the carbonyl carbon of the preceding residue (i−1). The HN(CA)CO experiment is far less sensitive; however, it provides the interresidue correlation as in the HNCO, as well as the intraresidue correlation (i.e., NH pair of residue i with the 13C of residue i). These two experiments yield complementary data.
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NH pairs, it is possible to link sequential residues via the CO resonance. In practice, due to the weakness of the HN(CA)CO data and degeneracies in the CO shift, it is not possible to complete the assignment process. Another source of sequential information is required. Often two complementary three-dimensional experiments are used to complete or extend the sequential assignment. In the case of Trypanosome calmodulin, the CBCA(CO)NH and HNCACB experiments were used. The former (highly sensitive) experiment correlates the amide pair of residue i with the Cα and Cβ chemical shifts of residue i−1; the latter corre-
A 30
B
HNCACB
Q-135β
Q-57β
lates the same amide pair with the Cα and Cβ chemical shifts of residue i. In addition, the HNCACB experiment may also yield correlations to the i−1 carbons, although these are generally weaker than the intraresidue cross peaks. The usefulness of this approach is that these experiments contain both the Cα and the Cβ chemical shift, which can facilitate the identification of residue type and also gives two frequencies to match between successive residues rather than just one. Figure 17.5.23 shows planes taken at an 15N chemical shift of 125 ppm from the CBCA(CO)NH and the HNCACB experiments, performed on U-13C/15N Trypano-
CBCA(CO)NH E-114β
E-114β K-115β
Q-135β
K-115β
K-115β
I-136β
40
D-56β
13C
(ppm)
D-56β
L-116β
50
Q-135α Q-135α
D-56α Q-57α
I-136
9.5
L-116α
D-56α
K-115α
E-114α
K-115α
K-115α
I-136α
60
E-114α
Q-57 K-115
9.0
8.5
L-116
I-136
8.0 1H
9.5 (ppm)
Q-57 K-115
9.0
8.5
L-116
8.0
Figure 17.5.23 Slices taken at an 15N chemical shift of 125 ppm through (A) HNCACB and (B) CBCA(CO)NH three-dimensional matrices. Data were acquired at 500.13 MHz at a temperature of 310K on a sample of calcium-saturated U-13C/15N Trypanosome calmodulin complexed with R-(−)-calmidazolium. The CBCA(CO)NH experiment correlates the amide 15N-1H pair of residue i with the Cα and Cβ carbons of residue i−1. These correlations also appear in the HNCACB data. Additionally, the HNCACB experiment provides intraresidue correlation information (i.e., correlations are observed from the “ith” 15N-1H pair to the Cα and Cβ carbons of residue i). Correlations that arise from intraresidue magnetization transfer are labeled in bold font. Notice also in the HNCACB experiment that Cα correlations are 180° out of phase with respect to those from Cβ. This is indicated on the figure by boxed contours for negative levels (β carbons).
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I-52
N-53
E-54
V-55
D-56
Q-57
D-58
G-59
S-60
G-61
T-62
I-63
D-64
F-65
45.00 i i
50.00 i
i
i
C (ppm)
55.00
i i i
i i i
60.00
i
i 65.00
i
70.00 117.2 117.3 116.8 108.0 121.9 126.4 115.9 7.73 8.52 7.71 7.26 7.89 8.97 8.17
108.9 116.2 115.7 107.8 125.2 128.8 119.1 N (ppm) 7.88 8.42 10.8 7.78 9.37 9.04 9.04 H (ppm)
Figure 17.5.24 Series of “strips” taken from a single HNCA experiment performed on calcium-saturated U-13C/15N Trypanosome calmodulin at 500.13 MHz at a temperature of 310K. For each strip, the pertinent 15N chemical shift and the 1H shift of the marked correlation are shown at the bottom, with the residue assignment indicated at the top. The vertical lines connect the Cαi (marked i) and Cαi−1 for each given amide proton within a strip. The horizontal lines connect those 13C resonances that are common to consecutive residues. In one strip, this peak is the strong intraresidue Cαi correlation, whereas in the succeeding strip it represents the weaker Cαi−1 shift.
some calmodulin complexed with calmidazolium. An additional benefit of the HNCACB experiment is that correlations from β carbons are of opposite phase to those from the α carbons (i.e., they are inverted, and are shown boxed in Fig. 17.5.23A). In principle, it may be possible to use a single three-dimensional experiment to achieve sequential assignment using the Cα chemical shift alone. Such an experiment is the HNCA, which correlates the “ith” amide pair with the Cα carbons of both residues i and i−1. The experiment is not very sensitive, but the intraresidue correlation (i.e., Ni to Cαi) is generally stronger than the interresidue correlation (Ni to Cαi−1). However, in practice, it is not often possible to complete the sequential assignment process using this experiment alone due to resonance overlap. Figure 17.5.24 shows a series of “strips” of sequential residues in the HNCA spectrum of calmodulin (assignments of these residues were achieved using the HNCACB/CBCA(CO)NH
methodology described above). The connectivity between successive residues, evidenced by the observation of a common carbon resonance in each strip, is marked by horizontal lines. The vertical lines connect the Cαi and Cαi−1 observed for the given amide proton. The intraresidue α carbon resonance is marked with an i. Many other triple-resonance experiments are possible, some of which incorporate a fourth frequency dimension (four-dimensional NMR). The fourth dimension further alleviates resonance overlap. In principle, a single four-dimensional experiment could yield the same information content as two three-dimensional experiments, although digital resolution in the indirect dimensions is likely to be reduced compared to the three-dimensional analogs. Analysis of the data generated from a suite of triple-resonance experiments is a time-consuming process that may become the bottleneck in the assignment process. Chemical shift matching is ideally suited to computing techStructural Biology
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niques, and several automated resonance assignment programs have been introduced that analyze correlation data generated by a variety of three- and four-dimensional triple-resonance experiments (e.g., Lukin et al., 1997). There are many examples of the use of triple-resonance NMR for protein structure determination (e.g., Ikura et al., 1992; Bagby et al., 1994; Jerala et al., 1996; Wang et al., 1996).
PROTEIN-LIGAND INTERACTIONS
Introduction to NMR of Proteins
Interactions of proteins with metal ions, peptides, other proteins, DNA/RNA hormones, neurotransmitters, enzyme substrates, and pharmacologically active small molecules (drugs) are of fundamental importance to biological processes. As such, an understanding of the molecular basis of these interactions is an important goal of biochemistry. This section concentrates on the interactions of proteins with small organic molecules (drugs) and metal ions. NMR is ideally suited to the study of protein-ligand complexes, as chemical shifts are sensitive markers of local structure. By analyzing perturbations in the NMR spectra of either or both interacting species, it may be possible to deduce key features of the complex, such as stoichiometry, affinity, kinetics, and interaction site(s). An important objective is separate observation of the resonances of each interacting molecule. In the case of metal ion binding, it is possible to observe the NMR signals of the protein in the absence and presence of the ion. Additionally, if the metal ion itself has a nonzero spin, its resonance can be observed under the same conditions. In the case of small molecule– protein interactions, however, it is likely that the resonances of the ligand will overlap (at least partially) with those of the protein, making both resonance assignment and characterization of the interaction extremely difficult. One way of alleviating this problem is to isotopically label either the protein or the ligand; it is often easier, and more common, to label the protein (see Isotopic Labeling). Isotope editing and filtering techniques may be used to select or remove the proton resonances of the labeled protein (see Isotope Editing: Two-Dimensional Heteronuclear Chemical Shift Correlations). In practice, the ligand is titrated (see Titration of Metal Ions or Organic Molecules) into a solution of the protein, and spectra of the protein and/or ligand are recorded at intermediate points. In this way, the dynamics and
stoichiometry of the interaction can be more accurately assessed. To illustrate some of the main principles involved, the following discussion returns to the small protein calmodulin to study its interactions with Ca2+ and a small molecule, calmidazolium (Fig. 17.5.21; Edwards, 1998).
Metal Ion Interactions Figure 17.5.25 shows the low-field region of proton spectra obtained during the course of titrating calmodulin with Ca2+ (Sweeney, 1990). On a qualitative level, the spectra of Ca2+-free and fully Ca2+-ligated calmodulin (bottom and top spectra in Fig. 17.5.25, respectively) are very different. The significant chemical shift changes observed represent a major protein conformational change upon Ca2+ binding. For instance, the very high-field resonance in the Ca2+-free protein changes shift dramatically on the addition of Ca2+. The methyl group giving rise to this resonance is removed from the anisotropic shielding region of an aromatic ring by a change in conformation of the protein upon binding Ca2+. Although calmodulin contains >900 nonexchangeable protons, significant spectral changes can be observed. However, at this stage interpretation on the atomic or structural level is not possible; this would require characterization of both Ca2+-free and Ca2+-ligated forms of the protein. Nevertheless, several aspects of the Ca2+binding process can be understood in a qualitative way. Ca2+ binding occurs in two distinct phases. The first phase, characterized by the addition of up to two equivalents of Ca2+, causes resonances characteristic of apocalmodulin (Ca2+-free) to disappear, while a distinct set of “new” resonances appear. This additional set of resonances belongs to (Ca2+)2-calmodulin, as there is no distinct (Ca2+)1 species. This is well illustrated by the His-107 Hε1 resonance and by the Tyr-138 Hδ protons marked on Figure 17.5.25. This well-characterized behavior is indicative of slow exchange kinetics—that is, strong binding between protein and ligand. Changes accompanying the second phase of Ca2+ binding, that is between two and four equivalents bound to calmodulin, are more subtle. Resonances change their position gradually from being characteristic of the (Ca2+)2- to the (Ca2+)4-calmodulin complex, as again there is no distinct (Ca2+)3 form. This can be clearly seen for the Phe-65 Hδ protons, marked on Figure 17.5.25. This behavior, in contrast to that observed in the first phase of Ca2+ binding, is called
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approximate equivalents of calcium
F-65δ
H-107ε1
ε
Y-138 δ 4.00
D-64,T-26α
3.50 3.00
2.50 2.00 1.75 1.50
1.25 1.00 0.75 0.50 0.25 0.00
8.0
7.5
7.0
6.5
6.0
5.5
ppm Figure 17.5.25 Low-field region of proton spectra obtained during the course of a titration of Trypanosome calmodulin with calcium at 360.13 MHz and 310K. Each spectrum was obtained after the addition of an aliquot of a 200 mM CaCl2 solution in 2H2O, corresponding to 0.25 calcium equivalents. The bottom-most spectrum represents calcium-free calmodulin and the top-most spectrum the fully calcium-ligated protein. The calculated molar ratio of calcium to calmodulin is shown on the figure. Notice the two resonances of His-107 Hε1 in the spectra obtained with 0.75 to 1 equivalent of calcium. This is due to slow exchange between calcium-ligated and calcium-free calmodulin, and means that the binding of calcium to calmodulin is very strong. The His-107 Hε1 resonance characteristic of calcium-free calmodulin (8.25 ppm) has completely disappeared after the addition of 2 equivalents of calcium. No further changes to the spectra were observed following addition of >4 equivalents of calcium. The appearance and disappearance of resonances is represented by up and down arrows, respectively; the dotted line indicates a resonance that changes shift.
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approximate equivalents Y-138 calmidazolium ε1,2 δ1,2 D-64α 2.0
1.7 1.5 1.3 1.0 0.6 0.3 ε1 8.4
H-107
δ2
0.0
8.2 8.0 7.8 7.6 7.4 7.2 7.0 6.8 6.6 6.4 6.2 6.0 5.8 5.6 5.4 5.2 ppm
Figure 17.5.26 Low-field region of proton spectra obtained during the course of a titration of calciumsaturated Phe-2H5 Trypanosome calmodulin with R-(−)-calmidazolium at 500.13 MHz and 310K. Each spectrum was obtained after the addition of an aliquot of 25 mM R-(−)-calmidazolium in methanol-2H4. It can easily be seen that the His-107 Hε1 proton acts as a marker for the titration, due to the very tight binding of calmidazolium to calmodulin, showing distinct chemical shifts for the apo- and calmidazoliumsaturated forms of the protein. After the addition of one molar equivalent of calmidazolium, the His-107 Hε1 resonance is split into two signals of equal area. This represents an equimolar mixture of calmodulin and the calmodulin-[calmidazolium]2 complex. The Tyr-138 ε1,2 protons exhibit similar behavior. All resonances (apart from the single His-107 δ2 proton) between 6.65 and 7.80 ppm are due to the aromatic protons of calmidazolium.
fast exchange kinetics. Although not shown in Figure 17.5.25, no further changes in chemical shift are observed following the addition of more than four Ca2+ equivalents. Calmodulin consists of two domains, each containing two Ca2+ binding sites. The C-terminal domain contains the higher-affinity Ca2+ binding sites, which are occupied first. This accounts for the observed slow exchange kinetics involving primarily C-terminal residues. The binding is cooperative (the binding of the first equivalent increases the affinity of the unoccupied site for Ca2+), explaining the absence of a (Ca2+)1 intermediate species. The lower-affinity Ca2+ binding sites that are located in the N-terminal domain are filled only after the C-terminal sites are occupied. The lower affinity of these sites for Ca2+ is reflected in the fast exchange kinetics observed.
Small Molecule Interactions
Introduction to NMR of Proteins
Using the techniques of isotope editing and filtering, it may be possible to observe, at will, the NMR resonances of both the protein and the ligand if one is isotopically normal but the other
is, for example, labeled with 13C. As part of a study to investigate the interaction between calmodulin from Trypanosoma brucei rhodiesiense and the very potent antagonist calmidazolium (Fig 17.5.21), a different, but nevertheless illustrative, labeling strategy was adopted. The only aromatic amino acids in Trypanosome calmodulin are nine Phe residues and single Tyr and His residues. Specific labeling with Phe-2H5 calmodulin effectively removes 90% of the protein aromatic resonances from the 1H NMR spectrum. The antagonist itself is mostly aromatic, so that the majority of its protons will resonate in the 7 to 8 ppm (aromatic) region of the 1H spectrum. Thus, the spectrum of a complex of calmidazolium with Phe-2H5 calmodulin will contain mostly antagonist signals in the aromatic region. Additionally, by using 2H2O as solvent, the NH signals, which would also resonate in the aromatic region, are absent. Figure 17.5.26 shows the 500 MHz one-dimensional proton spectrum of Phe-2H5 calmodulin in 2H2O during the course of titration with R-(−)-calmidazolium. The bottom spectrum in Figure 17.5.26 shows the protein in the
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absence of ligand, clearly demonstrating the spectral simplification obtained using the Phe2H label (compare with the top-most spectrum 5 of Fig. 17.5.25 at 360 MHz or with Fig. 17.5.16C at 500 MHz) with only His-107 ε1, δ2 and Tyr-138 ε1,2, δ1,2 protons visible in the aromatic region. The number of calmidazolium equivalents is shown in the figure. Calmidazolium shows slow exchange binding kinetics, exemplified by the behavior of the His-107 ε1 proton and, to a more limited extent, by the Tyr-138 ε1,2 protons. Distinct changes in protein chemical shifts over the entire spectrum are observed throughout the titration. It can be deduced from Figure 17.5.26 that calmodulin binds two equivalents of calmidazolium, and that the spectrum obtained after the addition of one calmidazolium equivalent is indicative of a 1:1 mixture of antagonist-free calmodulin and the calmodulin-[calmidazolium]2 complex. Two-dimensional homonuclear methods (e.g., TOCSY and NOESY) can be used to assign the resonances of the bound calmidazolium molecule. The NOESY spectrum will also show intermolecular nOes between the bound calmidazolium molecule and the protein. These nOes will be observed from calmidazolium protons to those protons of calmodulin that are involved in or around the drug binding site. As an example of the use of these methods, Figure 17.5.27 shows three regions taken from a 200-msec NOESY experiment on the calmodulin-calmidazolium complex. Figure 17.5.27A shows intermolecular nOes between calmidazolium and calmodulin, the majority of which occur to the methyl groups of hydrophobic residues. Figure 17.5.27B shows nOes between calmidazolium aromatic and aliphatic protons. Figure 17.5.27C shows intramolecular nOes between calmidazolium aromatic protons. The nOe data not only aid in the assignment of calmidazolium resonances, but also yield information about the conformation of the molecule when bound to the protein. However, it is not adequate to completely assign those protein residues involved in complex formation. To accomplish this, a drug-protein complex obtained from doubly labeled calmodulin is required, which can be subjected to the very powerful isotope-editing NMR techniques introduced above (see Isotope Editing: Two-Dimensional Heteronuclear Chemical Shift Correlations). For a more detailed discussion of the use of NMR in the analysis of protein-ligand interactions, the reader is referred to the following articles and references therein: Feeney and Bird-
sall (1993), Cooke (1996), and Craik and Wilce (1997).
OTHER PROTEIN NMR TECHNIQUES AND FUTURE DIRECTIONS As the molecular weight of a protein increases, it becomes more and more challenging to generate data amenable to interpretation and assignment. The most obvious effect of increasing molecular weight is the increased number of resonances in discrete spectral regions, yielding chemical shift degeneracy. In addition, the correlation time (τc) increases with molecular weight, resulting in more efficient T2 relaxation and hence broader lines. As a result, correlation experiments that rely on small J couplings become extremely inefficient. The dipolar interaction between protons also contributes significantly to increased rates of relaxation and to spin diffusion in nOe experiments. The dipolar interaction between protons and 13C and 15N also becomes more efficient, resulting in lower sensitivity for double- and triple-resonance experiments. The dipole-dipole interaction between protons can be reduced by decreasing the “concentration” of protons in a protein. This may be achieved by substituting a certain proportion of protons with deuterons. Fractional deuteration is an isotope labeling technique producing a heterogeneous mixture of isotopomers in which each position within the protein has the same probability of being deuterated (Sattler and Fesik, 1996). Although the number of protons giving a signal is reduced, the concomitant decrease in linewidth more than compensates, resulting in an overall increased signal-to-noise ratio. This labeling technique is particularly powerful when combined with U-13C and/or 15N labeling. In this case, triple-resonance methods become far more efficient as the T2 values of the heteronuclei, which are dominated by one-bond dipolar interaction with 1H, become much longer. The mutual interaction between two dipoles is proportional to the distance and angle between them. The angular dependence normally averages to zero in isotropic liquids. If molecules have preferred orientations in space relative to B0, this produces new signal splittings in the case of uncoupled nuclei, or changes in splitting of coupled nuclei, because the dipolar interaction no longer averages to zero. Adding the protein to a low concentration of self-orienting particles such as phospholipids or even viruses, which can form oriented bicelles in solution, a degree
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A
1.0
2.0
B
ppm
4.5
5.0
5.5
C 6.8
7.0
7.2
7.4
7.6
7.6
7.4
7.2
7.0
6.8
ppm
Figure 17.5.27 Expansions of a 200-msec NOESY spectrum obtained on the Phe-2H5 Trypanosome calmodulin-[calmidazolium]2 complex recorded at 500.13 MHz at a temperature of 310K. Panels A through C are each drawn with the same horizontal scale. (A) Intermolecular nOes between the aromatic protons of calmidazolium and calmodulin. The majority of such nOes are observed to hydrophobic residues of calmodulin, e.g., terminal methyl groups of Val, Ala, Ile, and Leu, and the ε methyl groups of Met residues. The assignment of these intermolecular nOes can potentially reveal the drug-binding sites on the protein. (B) Intramolecular nOes between the aromatic and aliphatic protons (i.e., H6, 7, and 8) of calmidazolium, again yielding both assignment and conformational information about the bound drug. (C) Mutual intramolecular nOes are observed between the aromatic protons of calmidazolium. These correlations contribute to the resonance assignment of these signals, and also give an indication of the conformation of the bound drug.
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of alignment is imposed on the protein (Bax and Tjandra, 1997). A comparison of J couplings observed in the absence and presence of such additives yields the magnitude and sign of the dipolar coupling (Jobs = J + D, where D is the dipolar coupling). The contribution of the dipolar coupling to the observed splitting is a function of the relative orientation of the bond with respect to the B0 field. This information yields constraints that can be used in addition to nOes and torsion angles to further refine protein structure (Tjandra et al., 1997). A further source of nuclear relaxation is chemical shift anisotropy (CSA), an effect that is proportional to the correlation time of the protein and to the square of the applied magnetic field. This relaxation mechanism becomes particularly prominent for the 15N-1H amide group in high magnetic fields. It is possible to record the 15N-1H HSQC experiment such that the heteronuclear spin coupling is retained in f1 and f2. This results in four correlations for each 15N-1H pair, each of which relax at a different rate. For one of these multiplet components, the dipole-dipole (DD) and the CSA mechanisms contribute to relaxation in an opposing sense, and balance exactly at high enough magnetic field (the DD relaxation is field independent), so that one of the four multiplet lines becomes sharp. It is calculated for 15N-1H that the reduction in linewidth will be a maximum at a magnetic field of ∼1 GHz, 23.5 T. An experiment called transverse relaxation optimized spectroscopy (TROSY; Pervushin et al., 1997) selects only the sharp multiplet component. Using this as a building block for triple-resonance experiments (Salzmann et al., 1999), it may be possible to determine the structure of very high-molecular-weight proteins. Thus, the quest for magnets with increasingly high field strength is not only desirable for increased resolution and sensitivity, but also for its potential to produce highquality narrow-line spectra for very high-molecular-weight proteins. Further general discussion of the topics covered here can be found in Dötsch and Wagner (1998) and Wüthrich (1998).
LITERATURE CITED Bagby, S., Harvey, T.S., Kay, L.E., Eagle, S.G., Inouye, S., and Ikura, M. 1994. Unusual helixcontaining Greek keys in development-specific Ca2+-binding protein S. 1H, 15N, and 13C assignments and secondary structure determined with the use of multidimensional double and triple resonance heteronuclear NMR spectroscopy. Biochemistry 33:2409-2421.
Basus, V.L. 1989. Proton nuclear magnetic resonance assignment. In Nuclear Magnetic Resonance, Part B (T.L. James and N.J. Oppenheimer, eds.) pp. 132-149. Academic Press, New York. Bax, A. and Davis, D.G. 1985. Practical aspects of two-dimensional transverse NOE spectroscopy. J. Magn. Reson. 63:207-213. Bax, A. and Tjandra, N. 1997. High-resolution heteronuclear NMR of human ubiquitin in an aqueous liquid crystalline medium. J. Biomol. NMR 10:289-292. Bothner-By, A.A., Stephens, R.L., Lee, J., Warren, C.D., and Jeanloz, R.W. 1984. Structure determination of a tetrasaccharide: Transient nuclear Overhauser effects in the rotating frame. J. Am. Chem. Soc. 106:811-813. Braunschweiler, L. and Ernst, R.R. 1983. Coherence transfer by isotropic mixing: Application to proton correlation spectroscopy. J. Magn. Reson. 53:521-528. Cooke, R.M. 1996. Protein-ligand interactions: Examples in drug design. In NMR in Drug Design (D.J. Craik, ed.) pp. 245-274. CRC Press, Boca Raton, Fla. Craik, D.J. and Wilce, A. 1997. Studies of proteinligand interactions by NMR. In Protein NMR Techniques (D.G. Reid, ed.) pp. 195-232. Humana Press, Totowa, N.J. Croasmun, W.R. and Carlson, R.M.K. 1994. TwoDimensional NMR Spectroscopy. Applications for Chemists and Biochemists. VCH Publishers, New York. Derome, A.E. 1987. Modern NMR Techniques for Chemistry Research. Pergamon Press, Oxford. Dötsch, V. and Wagner, G. 1998. New approaches to structure determination by NMR spectroscopy. Curr. Opin. Struct. Biol. 8:619-623. Edwards, A.J. 1998. An NMR isotope labelling analysis of calmodulin interactions with high affinity chiral inhibitors. Ph.D. thesis, University of Hertfordshire, U.K. Evans, J.N.S. 1995. Biomolecular NMR Spectroscopy. Oxford University Press, Oxford. Feeney, J. and Birdsall, B. 1993. NMR studies of protein-ligand interactions. In NMR of Macromolecules: A Practical Approach (G.C.K. Roberts, ed.) pp. 183-215. Oxford University Press, Oxford. Fejzo, J., Westler, W.M., Macura, S., and Markley, J.L. 1991. Strategies for eliminating unwanted cross-relaxation and coherence-transfer effects from two-dimensional chemical-exchange spectra. J. Magn. Reson. 92:20-29. Friebolin, H. 1993. Basic One- and Two-Dimensional NMR Spectroscopy. VCH Publishers, New York. Gronenborn, A.M., Bax, A., Wingfield, P.T., and Clore, G.M. 1989. A powerful method of sequential proton resonance assignment in proteins using 15N-1H multiple quantum coherence spectroscopy. FEBS Lett. 243:93-98. Structural Biology
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Güntert, P. 1997. Calculating protein structures from NMR data. In Protein NMR Techniques (D.G. Reid, ed.) pp. 157-194. Humana Press, Totowa, N.J.
Neuhaus, D. and Williamson, M.P. 1989. The Nuclear Overhauser Effect in Structural and Conformational Analysis. VCH Publishers, New York.
Ikura, M., Kay, L.E., and Bax, A. 1990. A novel approach for sequential assignment of 1H, 13C, and 15N spectra of larger proteins: Heteronuclear triple-resonance three-dimensional NMR spectroscopy. Application to calmodulin. Biochemistry 29:4659-4667.
Neuhaus, D., Wagner, G., Vasák, M., Kägi, J.H.R., and Wüthrich, K. 1985. Systematic application of high-resolution, phase-sensitive two-dimensional 1H-NMR techniques for the identification of the amino-acid-proton spin systems in proteins. Eur. J. Biochem. 151:257-273.
Ikura, M., Clore, G.M., Gronenborn, A.M., Zhu, G., Klee, C.B., and Bax, A. 1992. Solution structure of a calmodulin–target peptide complex by multidimensional NMR. Science 256:632-638.
Pervushin, K., Riek, R., Wider, G., and Wüthrich, K. 1997. Attenuated T2 relaxation by mutual cancellation of dipole-dipole coupling and chemical shift anisotropy indicates an avenue to NMR structures of very large biological macromolecules in solution. Proc. Natl. Acad. Sci. U.S.A. 94:12366-12371.
IUPAC-IUB Commission on Biochemical Nomenclature. 1970. Abbreviations and symbols for the description of the conformation of polypeptide chains. J. Mol. Biol. 52:1-17. Jerala, R., Almeida, P.F.F., Ye, Q., Biltonen, R.L., and Rule, G.S. 1996. 1H,15N and 13C resonance assignments and secondary structure of group II phospholipase A2 from Agkistrodon piscivorus: Presence of an amino-terminal helix in solution. J. Biomol. NMR 7:107-120. Kumar, A., Ernst, R.R., and Wüthrich, K. 1980. A two-dimensional nuclear Overhauser enhancement (2D NOE) experiment for the elucidation of complete proton-proton cross-relaxation networks in biological macromolecules. Biochem. Biophys. Res. Commun. 95:1-6. Lian, L. and Roberts, G.C.K. 1993. Effects of chemical exchange on NMR spectra. In NMR of Macromolecules: A Practical Approach (G.C.K. Roberts, ed.) pp. 153-182. Oxford University Press, Oxford. Lukin, J.A., Gove, A.P., Talukdar, S.N., and Ho, C. 1997. Automated probabilistic method for assigning backbone resonances of (13C,15N)-labeled proteins. J. Biomol. NMR 9:151-166. Macomber, R.S. 1998. A Complete Introduction to Modern NMR Spectroscopy. John Wiley & Sons, New York. Marion, D., Driscoll, P.C., Kay, L.E., Wingfield, P.T., Gronenborn, A.M., and Clore, G.M. 1989. Overcoming the overlap problem in the assignment of 1H NMR spectra of larger proteins by use of three-dimensional heteronuclear 1H-15N Hartmann-Hahn-multiple quantum coherence and nuclear Overhauser-multiple quantum coherence spectroscopy: Application to interleukin 1b. Biochemistry 28:6150-6156. Markley, J.L. and Kainosho, M. 1993. Stable isotope labelling and resonance assignments in larger proteins. In NMR of Macromolecules: A Practical Approach (G.C.K. Roberts, ed.) pp. 101-152. Oxford University Press, Oxford. Mossakowska, D.E. and Smith, R.A.G. 1997. Production and characterisation of recombinant proteins for NMR structural studies. In Protein NMR Techniques (D.G. Reid, ed.) pp. 325-335. Humana Press, Totowa, N.J. Introduction to NMR of Proteins
Piotto, M., Saudek, V., and Sklenár, V. 1992. Gradient-tailored excitation for single-quantum NMR spectroscopy of aqueous solutions. J. Biomol. NMR 2:661-665. Redfield, C. 1993. Resonance assignment strategies for small proteins. In NMR of Macromolecules: A Practical Approach (G.C.K. Roberts, ed.) pp. 71-99. Oxford University Press, Oxford. Richarz, R. and Wüthrich, K. 1978. Carbon-13 NMR chemical shifts of the common amino acid residues measured in aqueous solutions of the linear tetrapeptides H-Gly-Gly-C-L-Ala-OH. Biopolymers 17:2133-2141. Roberts, G.C.K. 1993. NMR of Macromolecules: A Practical Approach. Oxford University Press, Oxford. Salzmann, M., Wider, G., Pervushin, K., Senn, H., and Wüthrich, K. 1999. TROSY-type triple resonance experiments for sequential NMR assignments of large proteins. J. Am. Chem. Soc. 121:844-848. Sattler, M. and Fesik, S.W. 1996. Use of deuterium labeling in NMR: Overcoming a sizeable problem. Structure 4:1245-1249. Stockman, B.J. 1996. Preparation of 2H, 13C and 15N isotopically enriched proteins for NMR spectroscopic investigations. In NMR Spectroscopy and its Application to Biomedical Research (S.K. Sarkar, ed.) pp. 159-185. Elsevier/NorthHolland, Amsterdam. Sutcliffe, M.J. 1993. Structure determination from NMR data II. Computational approaches. In NMR of Macromolecules: A Practical Approach (G.C.K. Roberts, ed.) pp. 359-390. Oxford University Press, Oxford. Sweeney, P.J. 1990. NMR analysis of drug interactions with isotopically labelled calmodulin. Ph.D. thesis, Hatfield Polytechnic, Hertfordshire, U.K. Tjandra, N., Omichinski, J.G., Gronenborn, A.M., Clore, G.M., and Bax, A. 1997. Use of dipolar 1H-15N and 1H-13C couplings in the structure determination of magnetically oriented macromolecules in solution. Nat. Struct. Biol. 4:732738.
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Wang, Y., Frederick, A.F., Senior, M.M., Lyons, B.A., Black, S., Kirschmeier, P., Perkins, L.M., and Wilson, O. 1996. Chemical shift assignments and secondary structure of the Grb2 SH2 domain by heteronuclear NMR spectrsocoopy. J. Biomol. NMR 7:89-98. Weber, P.L. 1996. Protein structure determination from NMR data. In NMR Spectroscopy and Its Application to Biomedical Research (S.K. Sarkar, ed.) pp. 187-239. Elsevier/North-Holland, Amsterdam. Wishart, D.S., Sykes, B.D., and Richards, F.M. 1996. The chemical shift index. A fast and simple method for the assignment of protein secondary structure through NMR spectroscopy. Biochemistry 31:1647-1651.
Wüthrich, K. 1986. NMR of Proteins and Nucleic Acids. John Wiley & Sons, New York. Wüthrich, K. 1998. The second decade—into the third millenium. Nat. Struct. Biol. 5(NMR Suppl.):492-495.
Contributed by Andrew J. Edwards and David Reid SB Pharmaceuticals Welwyn, Herts, United Kingdom
The authors would like to thank their colleagues at SmithKline Beecham Pharmaceuticals for their assistance, especially Drs. Lesley K. MacLachlan and Julia Hubbard for providing the spectra of NPY. In addition, the authors would like to thank Prof. John Walker of the University of Hertfordshire for his collaboration on the Trypanosome calmodulin project.
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Probing Protein Structure and Dynamics by Hydrogen Exchange–Mass Spectrometry
UNIT 17.6
Determining protein 3-D structure to a resolution of 2 to 3 Å provides a good starting point in understanding protein structure-function relationships. However, protein molecules are highly dynamic, which may be essential for many of their natural functions. For example, the structures of viral coat proteins may change extensively when viruses bind to cellular receptors and disassemble, releasing their genetic material into the cell. To understand structural biology at the molecular level requires a variety of analytical methods for detecting changes in the structures and dynamics of proteins. Low resolution experimental methods, such as circular dichroism (CD; UNIT 7.6) and fluorescence (UNIT 7.7), have been used to detect protein structural changes induced by changing temperature and pH or by adding denaturants. Although the exact locations of structural changes cannot be determined by these methods, they do provide convenient means for showing that changes have occurred. High resolution methods, such as NMR (UNIT 17.5) and X-ray crystallography (UNIT 17.3), are used to determine the location of every atom in a molecule. Hydrogen exchange rates at backbone peptide amide linkages of proteins have been used for many years as a sensitive probe for detecting changes in protein structure and dynamics (Hvidt and Nielsen, 1966; Woodward et al., 1982; Englander and Kallenbach, 1984). This approach is based on the fact that the rate constants at which amide hydrogens in protein exchange with deuterium or tritium in the bulk solvent are highly dependent on protein conformation. Hydrogen exchange rates are correlated with the solvent accessibility of amide hydrogens and their intramolecular hydrogen bonding. Even minor changes in protein conformation may cause rearrangement in the hydrogen-bonding network and alter the solvent accessibility of certain amide hydrogens. These changes often alter the hydrogen exchange rates at some amide linkages. Experimental procedures discussed in this unit are relevant only to hydrogens located at peptide amide linkages in polypeptides. Exchangeable hydrogen located in sidechains, as well as at the N- and C-termini, exchange too fast to be measured by these methods (Englander et al., 1985; Bai et al., 1993). Nuclear magnetic resonance (NMR; UNIT 17.5) has been particularly useful for hydrogen exchange measurements because exchange rates at individual peptide amide linkages can be determined. However, this approach is limited by requirements for large quantities of sample, high protein solubility, and relatively low molecular weight proteins. Numerous studies have demonstrated that mass spectrometry can also be used to detect deuterium levels at peptide amide linkages in proteins labeled under a variety of conditions (Katta and Chait, 1991; Thévenon-Emeric et al., 1992; Miranker et al., 1993; Zhang and Smith, 1993). When proteins are fragmented by acid proteases or collision-induced dissociation (CID MS/MS; UNTI 16.6), hydrogen exchange–mass spectrometry (HX-MS) can be used to determine deuterium levels in short segments of proteins. Particularly important advantages of mass spectrometry for detecting hydrogen exchange include the ability to study large proteins with high sensitivity and at very low concentrations; and the ability to determine the intermolecular distribution of deuterium. This unit provides a brief discussion of the interplay between protein dynamics and hydrogen exchange, a detailed prescription for designing and performing typical HX-MS experiments, and common procedures for processing and interpreting HX-MS data.
Structural Biology Contributed by Lintao Wang and David L. Smith Current Protocols in Protein Science (2002) 17.6.1-17.6.18 Copyright © 2002 by John Wiley & Sons, Inc.
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SOME BACKGROUND ON HYDROGEN EXCHANGE The details of hydrogen exchange have been reviewed extensively elsewhere (Woodward et al., 1982; Englander and Kallenbach, 1984; Englander et al., 1985; Bai et al., 1993; Miller and Dill, 1995; Englander et al., 1997; Smith et al., 1997). The following discussion provides the basic hydrogen exchange background required to understand the general HX-MS protocols and to interpret HX-MS data. Hydrogen Exchange in an Unstructured Polypeptide Isotopic hydrogen exchange at each peptide amide linkage in an unstructured peptide is catalyzed by acid and base, as illustrated by the following equation: k int = k H [H + ] + kOH [OH - ] Equation 17.6.1
where kint is the intrinsic exchange rate constant, and kH and kOH are acid- and base-catalyzed rate constants, respectively. The linear dependence of kint on H+ and OH− shows that hydrogen exchange rates are highly dependent on pH. The pH dependence of amide hydrogen exchange rate constant calculated for polyalanine (see Fig. 17.6.1) is slowest at pH 2.4 to 3.0. Above this range, the exchange rate changes by one order of magnitude for each unit of pH. This high sensitivity to pH is an important consideration in all HX-MS experiments. For example, labeling may be performed under physiological conditions, then quenched by decreasing the pH to 2.5 prior to analysis. Temperature is another important factor that affects amide hydrogen exchange rates. The temperature dependence of exchange rates can be predicted using the effective activation energies for kH and kOH, which are 14 and 17 kcal/mol, respectively (Bai et al., 1993). Aside from the effects of pH and temperature, the exchange rate at each individual amide linkage is also dependent on the side chains of their neighboring amino acid residues. These effects, which are generally inductive and steric in nature, appear to be additive. Exchange results from model peptides can be used to predict hydrogen exchange rates in unstructured peptides at any pH and temperature (Molday et al., 1972; Bai et al., 1993). These calculated rate constants for exchange from an unstructured polypeptide are often
Probing Protein Structure and Dynamics by HX-MS
17.6.2 Supplement 28
Figure 17.6.1 Dependence on pH of the rate constant for amide hydrogen exchange in polyalanine (Bai et al. 1993). Current Protocols in Protein Science
compared with measured rate constants for exchange from a folded polypeptide to determine the extent the folded structure decreases isotope exchange. Although side-chain effects in unstructured polypeptides alter the amide hydrogen exchange rates by as much as 30-fold, secondary, tertiary, and quaternary structural features of folded proteins may decrease amide exchange rates by as much as 108. It is this large reduction that makes hydrogen exchange a sensitive probe of the folded structures of proteins. Hydrogen Exchange in Folded Proteins Hydrogen exchange under conditions where a protein is generally folded may be described using a two-process model (Kim and Woodward, 1993; Bai et al., 1994; Li and Woodward, 1999), as illustrated in the following equations: kex f
, → f (D) f (H)
Equation 17.6.2
k
k
k−1
k1
−1 kint 1 → u(D) → f (D) → u(D) ← f (H) ←
Equation 17.6.3
where f and u refer to folded and unfolded forms, respectively, and H and D refer to hydrogen and deuterium, respectively. Exchange of amide hydrogens directly from the folded form of a protein is described by Equation 17.6.2. The rate constant for this form of isotope exchange can be described by Equation 17.6.4: kex,f = βk int Equation 17.6.4
where β is the probability for exchange from folded forms and kint is the rate constant for isotope exchange from the totally unfolded polypeptide. It has been suggested that β represents highly localized, low amplitude structural changes (Elber and Karplus, 1987; Kim and Woodward, 1993). In the process illustrated by Equation 17.6.3, hydrogen exchange occurs only after a segment of protein unfolds, exposing several amide hydrogens to the bulk solvent. When exposed to solvent, the amide hydrogens exchange as if they were in an unfolded polypeptide. The unfolding and refolding rate constants are designated here by k1 and k-1, respectively. For most proteins at neutral pH and in the absence of denaturants, k-1 >> kint. In this case, the rate constant for exchange through momentary unfolding of a protein is expressed as kex,u =
k1 kint = K unf kint k-1
Equation 17.6.5
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where Kunf is the equilibrium constant for momentary unfolding. When k-1 >> kint, a segment of any molecule must unfold and refold many times before exchange within it is complete. This unfolding may involve a short segment or the entire polypeptide backbone. Exchange at individual amide hydrogens may occur through a combination of these two models. Hence, the rate constant for hydrogen exchange, kex can be expressed as the sum of two rate constants, as illustrated in Equation 17.6.6. kex = kex,f + kex,u = (β + K unf )kint
Equation 17.6.6
This expression is particularly important because it links the experimentally measured deuterium exchange rate constant, kex, to important parameters describing protein dynamics, β and Kunf. The variation of kex with low levels of denaturants has been used to separate these two processes (Bai et al., 1993; Kim et al., 1993). Exchange by both processes leads to a random intermolecular distribution of isotope peaks and a single envelope of isotope peaks in mass spectra (Gross et al., 1996; Zhang et al., 1996). It may be noted that other protein dynamics, such as those found when k-1 << k2 (Arrington et al., 1999) or in protein folding studies (Miranker et al., 1993; Deng and Smith, 1999), may give multiple envelopes of isotope peaks. BASIC PROTOCOLS FOR HX-MS EXPERIMENTS The general HX-MS procedure to be discussed here starts by exposing the protein to D2O for various times to effect partial exchange at peptide amide linkages (see Fig. 17.6.2). This strategy, which may be viewed as continuous labeling, is appropriate for studying the structures and dynamics of proteins where the labeling time is the primary variable. In an alternative labeling strategy called pulse labeling, the labeling time is fixed while another experimental parameter, such as folding time, is varied. Continuous- and pulselabeling strategies have been discussed elsewhere (Deng et al., 1999). According to the procedure illustrated in Fig. 17.6.2, a protein sample is diluted into D2O exchange buffer to initiate isotope exchange. After various labeling times, the exchange solution is acidified and cooled to quench isotope exchange. Two basic approaches for analysis giving two different types of information may then follow. In the first approach, the deuterium level incorporated into the whole protein is determined by analyzing the intact, labeled protein by directly-coupled HPLC/electrospray ionization mass spectrometry (ESI-MS). These results provide the global exchange profile of a protein, that is, the exchange properties averaged over all peptide amide linkages. In an alternative approach, the protein fragmentation/MS method, the labeled protein is proteolytically fragmented into peptides whose deuterium levels are determined by HPLC/ESI-MS. This approach provides the local exchange profile of a protein, facilitating detection of localized structural changes. The following discussion focuses on experimental details of the protein fragmentation approach. Details for performing the simpler, but nevertheless useful procedure used to follow global exchange will be evident. Initiation of Protein Hydrogen Exchange Probing Protein Structure and Dynamics by HX-MS
Prior to labeling, the protein should be in a well-defined state. For example, in studies of enzymes, it is important to verify that the protein is active. Other sources of complication include misfolded, insoluble, or aggregated forms of a protein. Because the structure of
17.6.4 Supplement 28
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protein D2O exchange buffer labeled protein quench buffer, pH 2.4, 0°C labeling quenched
local exchange profile
pepsin digest
global exchange profile
HPLC/ESIMS
pH 2.4, 0°C HPLC/ESIMS
deuterium content of protein
deuterium content of peptic fragments
Figure 17.6.2 A general experimental procedure of hydrogen exchange/mass spectrometry.
a lyophilized protein may be different from its solution structure, proteins should be allowed to equilibrate in H2O prior to exposure to D2O. Dilution of the protein stock solution with an excess of deuterated exchange buffer is the simplest approach for initiating hydrogen exchange. A 20-fold dilution gives a labeling solution that is 95% D, which sets the highest deuterium content achievable in the protein. The use of large dilution factors leads to low protein concentrations, which adversely affects the detection limits of ESI-MS. Although effective procedures for concentrating the labeled protein have been described (Ehring, 1999; Wang et al., 2001), they require more time and result in greater losses of deuterium. Exchange has also been initiated using a spinning Sephadex G-10 column (Jeng et al., 1990; Zhang et al., 1996). This approach avoids significant dilution of the sample, but is somewhat more tedious. Although control of pH (i.e., pD) during labeling is critical, pH is a relatively flexible parameter and is usually chosen to satisfy the particular needs of a study. Because HPLC will be used to remove buffers and other salts just prior to MS analysis, the choice of buffer will depend on the desired pH. Phosphate buffers are often used because phosphate has a pK1 of 2.15 and pK2 of 6.82, providing excellent buffering capacity at both the exchange step (neutral pH) and the quench step (pH 2.4). Because rate constants for hydrogen exchange in folded proteins typically span a range of 108, choosing an appropriate exchange time is important. The shortest and longest time periods determine the range of isotope exchange rate constants that can be detected. For example, only those rate constants in the range 0.001 to 4 min−1 could be determined for exchange times from 10 sec to 8 hr. The shortest exchange time is often chosen to be just
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adequate to effect complete exchange in an unfolded polypeptide. In this case, labeling for 5 sec is adequate at pH 7 and 20°C. Shorter labeling times, which may be implemented using a high-speed mixing apparatus, may be useful for pH >7 (Yang and Smith, 1997). Although labeling may be continued for days or weeks, as required for detectable H/D exchange at the slowest exchanging amide linkages, protein aggregation and bacterial growth may present problems. If rate constants are to be derived from exchange results, samples should be analyzed at 4 to 5 time points for each decade of time. When performing continuous-labeling experiments, a “master” solution of protein is usually prepared. This solution contains enough protein for LCMS analysis of all samples. For example, if analysis required 10 time points with 300 pmol consumed for each LCMS analysis (typical for a microbore column, 1 × 50–mm), a total of 3 nmol of protein is required. To minimize sources of random error, it is prudent to analyze 2 to 3 samples for each time point. Quenching Exchange Reaction Once hydrogen exchange is initiated, a timer is started to indicate the exchange time. At various time points (10 sec, 30 sec, 1 min, and so on), an aliquot of sample containing enough protein for a single LCMS analysis is removed from the labeling solution. Isotope exchange in this aliquot is quenched by decreasing the pH to 2 to 3 and the temperature to 0°C. If the solution already contains phosphate buffer, the pH can be decreased by simply adding the appropriate quantity of acid. Hydrochloric, phosphoric, and trifluoroacetic (TFA) acids work well. If a buffer effective at pH 2.5 is not present, one must be added. The volume of quench solution to be added depends on several factors, which include volume, pH, and buffer concentration of the labeling solution, as well as the pH and buffer concentration of the quench solution. To minimize dilution of the sample, experiments should be designed so the volume of quench solution is <10% of the labeling solution. Whether H2O or D2O is used to prepare the quench solution is not important for most experiments. Decreasing the sample to pH 2 to 3 and the temperature to 0°C is critical for efficient quenching of isotope exchange. Figure 17.6.1 shows that exchange is slowest at pH 2 to 3. If protein is labeled at pH 7, decreasing the pH to 2 to 3 will decrease the intrinsic exchange rate at amide linkages by ∼10−4. Decreasing the temperature from 20°C to 0°C lowers the exchange rate another ten fold. Although hydrogen exchange is relatively slow under quench conditions, some back exchange during analysis is inevitable. Precise control of quench conditions (i.e., pH and temperature) is essential for obtaining highly reproducible results. It is noted that retention of folded structure in the protein to reduce exchange during analysis, as used in many hydrogen exchange NMR studies, is not necessary for the HX-MS protocol described here. Proteolytic Fragmentation of Labeled Protein
Probing Protein Structure and Dynamics by HX-MS
To study the structure and dynamics within specific regions of a protein, the labeled protein may be fragmented with an acid protease under conditions where isotope exchange remains quenched (Rosa and Richards, 1979; Englander et al., 1985; Zhang and Smith, 1993). Pepsin is ideal because it has maximum activity at pH 2 to 3 where the exchange rates are slowest. Due to its low specificity, pepsin usually gives many overlapping peptides. The spatial resolution may be improved considerably by considering the deuterium levels determined from differences in levels in overlapping peptic fragments (Zhang and Smith, 1993). Deuterium levels at specific amide linkages may be determined by collision-induced decomposition (CID) MS/MS analysis of peptic fragments (Deng
17.6.6 Supplement 28
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et al., 1999; also see UNIT 16.6). However, this procedure may lead to erroneous results because some MS instrument conditions may lead to intramolecular H/D scrambling. The simplest approach for digesting labeled protein is to add pepsin to the quenched solution. Because digestion is performed at a low temperature, a large amount of pepsin (e.g., substrate/enzyme ratio of 1:1) is usually required to fragment the protein rapidly. For example, 2 to 10 µl of a concentrated pepsin solution (1 to 5 mg/ml) may be added to each sample (200 to 500 pmol protein in 50 to 100 µl). Addition of larger volumes will contribute to the dilution factor. Despite use of high concentrations of pepsin, there is generally little evidence for peptic fragments of pepsin. Digestion for 5 min often gives many peptides with 5 to 20 residues. Immobilized pepsin has been used to reduce the digestion time considerably by increasing the effective concentration of pepsin (Ehring, 1999; Halgand et al., 1999; Wang et al., 2001). If these procedures do not lead to sufficient digestion, denaturing agents, such as organic solvents, urea and guanidine hydrochloride may be used (Wang and Smith, 1999; Wang et al., 2001). Although the abundance of specific peptic fragments may be sensitive to the experimental conditions, careful control of these conditions will usually lead to reproducible abundances of fragments. Because the specificity of pepsin is low (Smith et al., 1997), digestion usually gives a large number of peptic fragments. For example, digestion of a 50-kDa protein may generate >200 peptides. Because pepsin cleavage sites cannot be predicted with certainty, the peptides cannot be identified reliably by nominal molecular mass alone. However, these peptides may be readily identified by a combination of exact molecular mass, CID MS/MS or C-terminal sequencing using various carboxy-peptidases (McCloskey, 1990). HPLC/ESI-MS Analysis of Deuterated Peptides Although various strategies for measuring deuterium levels in polypeptides by mass spectrometry without the aid of HPLC have been reported, use of directly coupled HPLC offers many advantages. Because the response of ESI to specific peptides often depends on whether other peptides are present, fractionation of peptides prior to analysis by ESI-MS (UNITS 16.8 & 16.9) can substantially increase the number of peptides detected. In addition, separation of the peptides reduces the probability that peaks for peptides with similar molecular masses will overlap. Use of HPLC also allows considerable freedom in the choice of buffers, salts, and denaturants during sample preparation. Because gradient HPLC also serves as a solid phase extractor, it allows analysis of very dilute solutions (Wang et al., 2001). Finally, it is important to note that HPLC is performed with protic solvents that effectively remove deuterium from the side chains (N- and C-termini) via back exchange. As a result, the measured deuterium level reflects the number of deuteriums located at peptide amide linkages. In addition, removal of deuterium from the rapidly exchanging side chains may be important for minimizing deuterium losses during the electrospray ionization process. Although a wide variety of reversed-phase HPLC approaches has been used for HX-MS experiments, minimizing deuterium loss from the peptide-amide linkages is usually the principal concern. When choosing HPLC conditions, it is important to note that the half-life for deuterium loss from peptide amide linkages under typical quench conditions is 20 to 500 min (Bai et al., 1993). In addition to maintaining hydrogen exchange quench conditions (pH 2 to 3, 0°C), one normally uses short gradients and high flow rates to minimize the analysis time. The low temperature is maintained by submerging the injector and column in an ice bath, while the appropriate pH may be maintained using 0.05% TFA or 0.25% formic acid in both mobile phases. Small columns are generally preferred because they minimize sample dilution, thereby increasing sensitivity. However, the low flow rates used with small columns may create delays in formation of the gradient if the
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HPLC system was not designed for such flow rates. In addition, low flow rates may substantially increase the residence time of peptides in the region following the column. To avoid excessive loss of deuterium in this region, which is not cooled, the residence should be less than ∼1 sec. A typical HPLC protocol suitable for most HX-MS experiments uses a 1 × 100–mm C18 column (e.g., Microtech/Vydac). Columns packed with materials that are not designed for large molecules should be avoided because they are easily plugged by undigested sample or pepsin. Mobile phases consisting of water and acetonitrile, both with 0.05% TFA, are used to elute the peptides within 10 min. A flow rate of 40 to 50 µl/min is a good compromise between separation and speed. Injection of 400 pmol is usually adequate for modern, high-sensitivity mass spectrometers. When the required sample volume is large (e.g., greater than ∼50 µl), the flow rate can be increased to shorten the sample loading time. The upper pressure limit of the column should not be exceeded. To decrease deuterium loss, the loading syringe should be cooled before sample injection. Depending on the salt content of the sample, 1 to 3 min of desalting on the column prior to elution may be advisable. Mass spectra of peptides eluting from the HPLC are recorded continuously. Several different types of ESI mass spectrometers have been used successfully for HX-MS measurements. The authors’ experience is based on four instruments, a Micromass Autospec magnetic sector, a Finnigan LCQ ion-trap, a Micromass VG platform singlequadruple, and a Micromass QTOF. To minimize deuterium loss in the electrospray source, the source tuning parameters, especially the source temperature, must be optimized. For example, significant deuterium loss in the source was detected in the Autospec when the source was operated at temperatures >60°C. Whether significant deuterium loss occurs in any part of the procedure, including inside the ESI source, the instrument should be monitored routinely using completely exchanged peptides. Processing of LCMS Data The next step following data acquisition is the identification of scans that were acquired during elution of each peptide of interest. Spectra from these scans are combined and analyzed to give the average molecular mass of the peptide, from which the number of deuteriums present can be determined. The ions representative of a peptide will have a range of molecular masses depending on the number of heavy atoms (e.g., 13C, 18O, 15N, and D) present. For most studies where the hydrogen exchange processes described by Equations 17.6.2 and 17.6.3 dominate, the intermolecular distribution of deuterium is random, which gives a single envelope of isotope peaks. Linkage between the width of this distribution and the structure/dynamics of the protein has been discussed (Gross et al., 1996; Zhang et al., 1996). Mass spectra of proteins with multiple, long-lived populations may have multiple envelopes of isotope peaks (Miranker et al., 1993; Zhang et al., 1996; Deng and Smith, 1999). The average mass of a peptide, which takes into account both the abundance (Ii) and the mass (mi) of each isotopic form of the peptide, is defined by Equation 17.6.7:
∑ I i mi average mass = i ∑ Ii i
Probing Protein Structure and Dynamics by HX-MS
Equation 17.6.7
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The software provided with most mass spectrometers can be used to determine the average molecular mass of polypeptides whose isotope peaks are not resolved, as is usually the case for proteins. Additional software, such as MagTran (Zhang) or Excel (Microsoft) may be required to determine the average molecular masses of peptides whose isotope peaks are resolved. Typical HX-MS results obtained using a Micromass Autospec mass spectrometer equipped with an ESI source and array detector are given in Fig. 17.6.3. Approximately 400 pmol of brome mosaic virus (BMV) coat protein (mol. wt. 20 kD) peptic digest was loaded on a C18 reversed-phase column (1 × 50–mm, flow rate 40 µl/min, 2% to 60% acetonitrile in 6 min, 0.05% TFA). The total ion current (TIC), which is similar to a UV chromatogram, is shown in Fig. 17.6.3A. This chromatogram shows that most of the peptides eluted within 6 to 7 min. The selected ion plot for m/z 391-393, representing the peptic fragment 183-188 of BMV capsid protein, shows that this peptide eluted in a 30-sec time interval with an average retention time of 3.7 min (Fig. 17.6.3B). Although many
A
Relative intensity
100
50
0 2:24
B
total ion current (TIC)
3:36
4:48 min
6:00
7:12
100 Relative intensity
TIC of m/z 391-393
0 2:24
100 Relative intensity
C
50
3:36
4:48 min
6:00
7:12
391.7
mass spectrum of scans from 3:30-3:58
50 639.5
430.4 0 320
782.8
720
520 m/z
100 Relative intensity
D
50
391.7
mass spectrum of m/z 391.7
392.2
centroid = 391.85 charge state = 2
392.7 0 390
394
392
396
m/z region selected to determine the average mass of this peak
Figure 17.6.3 Representative HPLC/ESI-MS spectra in a typical HX-MS analysis showing data processing steps involved in determining the average mass of a particular peptide.
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other peptides eluted in the same time interval (see Fig. 17.6.3C), their ions were well resolved in the mass spectra. The average molecular mass of this peptide was determined from the centroid of the entire envelope of isotope peaks multiplied by the charge-state (Fig. 17.6.3D). The deuterium level in this peptide was determined by subtracting the average mass of the non-deuterated form of the same peptide, which was determined in a separate experiment. Adjustment for Artifactual Exchange Although quench conditions may decrease isotope exchange by a factor of 105, a significant amount of exchange usually occurs during digestion and HPLC. To compare differences in the structure and dynamics of two closely related forms of a protein, one may simply compare the deuterium levels found for the two forms. That is, it is not necessary to adjust measured deuterium levels for exchange that occurred during digestion and analysis. However, more detailed analysis, which may include quantitative comparison of found and calculated deuterium levels, requires such adjustments. The amount of deuterium lost from a partially deuterated peptide may be estimated from the amount of deuterium lost from a totally deuterated peptide (i.e., totally exchanged in D2O), which can be determined by analyzing a sample of the totally deuterated protein (Zhang and Smith, 1993). This sample, the 100% reference, may be prepared most easily
A
120 100% control <m100%> = 393.46 D = 3.22
Relative intensity
100 80 60 40 20 0 390
B
391
392
393
394
labeled peptide <m> = 392.84 D = 1.98
Relative intensity
80 60 40 20 0 390
391
392
393
394
Relative intensity
395
396
120 0% control <m0% > = 391.98 D = 0.26
100 80 60 40 20 0 390
Probing Protein Structure and Dynamics by HX-MS
396
120 100
C
395
391
392
393
394
395
396
m/z
Figure 17.6.4 Mass spectra of a peptide fragment from BMV capsid protein that are (A) fully deuterated, (B) partially deuterated, and (C) nondeuterated.
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by incubating the protein in D2O for 24 hr at pD 2 to 3 and elevated temperature. Denaturants may be required to achieve complete exchange. The mass spectrum of the segment including residues 183-188 of BMV capsid protein completely exchanged in D2O is given in Figure 17.6.4A. The centroid of this envelope of isotope peaks shows that this segment had 3.22 deuteriums when it arrived at the mass spectrometer. This segment has 4 exchangeable amide hydrogens (6 residues, one of which is Pro). Finding 3.22 deuteriums shows that 80.5% of the deuterium at peptide-amide linkages was retained during digestion and analysis. The deuterium recovery, 80.5%, may be used to adjust the deuterium level found in the same peptide when it is only partially deuterated at the time exchange was quenched. The mass spectrum of the 183-188 segment derived from partially deuterated BMV capsid protein (Fig. 17.6.4B) indicates that this segment had 1.98 deuteriums when it arrived at the mass spectrometer. Adjusting for the 80.5% recovery indicates that this segment actually had 2.46 deuteriums at the time exchange was quenched. Because deuterium loss is sequence specific (Bai et al., 1993), the deuterium recovery for each peptide must be determined. Furthermore, because the recovery is based on the sum of deuterium levels found at all positions, the adjustment is only an approximation of the recovery in partially deuterated forms of the same peptide. When the recoveries are relatively high, adjustments for artifactual deuterium losses are small. In such cases, errors in the adjustment may be insignificant. For optimized experimental conditions, one may expect deuterium recoveries of 65% to 95% for analysis of peptic fragments and 85% to 95% for intact proteins. Failure to consider deuterium exchange-in during digestion is another source of error in adjustments based only on deuterium recovery. A more accurate method, which adjusts for deuterium exchange-in during digestion, has been described (Zhang and Smith, 1993). This approach requires analysis of a 0% reference sample that was digested in a solution under the same conditions (including H/D ratio) as used for all other samples. The mass spectrum of the 183-188 segment from a 0% reference sample of BMV capsid protein (Fig. 17.6.4C) indicates that this segment had 0.26 deuteriums. These deuteriums exchanged-in during quench and digestion. The deuterium level in this segment at the time H/D exchange was quenched can be estimated from Equation 17.6.8: D=
< m > − < m0% > ×N < m100% > − < m0% > Equation 17.6.8
where D is the adjusted number of deuteriums in a peptide, N is the total number of amide linkages, and <m>, <m0%>, and <m100%> are the average masses of a peptide from a partially deuterated sample, the 0% control, and the 100% control, respectively. The application of Equation 17.6.8 to segment 183-188 indicates that this segment had 2.32 deuteriums at the time exchange was quenched. Errors in such adjustments due to different exchange rates at different linkages have been discussed (Zhang and Smith, 1993). Experimental Error The accuracy with which deuterium levels can be determined is of critical importance because it sets the threshold for detecting structural changes in a protein. For example, significant structural changes may alter deuterium levels by as little as 0.1 deuteriums. Experimental errors come from two major sources: sample preparation and mass measurement. The most important sources of error in sample preparation include time, pH, and temperature at all steps (e.g., exchange-in, digestion, and HPLC). It is helpful to note that the exchange rate changes ten fold for each pH unit and approximately three fold for
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each 10°C change in temperature (Bai et al., 1993). The accuracy of the mass measurement depends on the signal-to-noise ratio and the particular instrument. However, one may expect the uncertainty in mass measurement to be 10 to 500 ppm. Samples to be compared with one another should be prepared in the identical buffer and analyzed under identical conditions. Deuterium in solvents and buffers may impose isotope effects on measured hydrogen exchange behavior (Bai et al., 1993; Connelly et al., 1993). The isotope effects on amide hydrogen exchange rates in peptides are generally small. However, the solvent isotope effect on the glass electrode used to measure pH (pD) may be important for some types of analyses. The authors’ laboratory, as well as many others, normally reports the value actually read from the pH meter. HX-MS DATA INTERPRETATION The deuterium levels in proteins or protein fragments may be used to detect changes or differences in protein structure and dynamics. For the simplest interpretation of hydrogen exchange results, finding different deuterium levels for the same labeling time is strong evidence that a change has occurred. For a more detailed interpretation, time-course exchange-in data can be fitted into a sum of first-order rate expressions to obtain the distribution of rate constants for exchange at amide linkage in the protein or peptic fragment. Changes in the exchange rate distribution are used to characterize changes in protein structure and dynamics. The models for H/D exchange (Equations 17.6.2 and 17.6.3) provide physical models for relating H/D exchange data and protein structure/dynamics. The discussion below illustrates both approaches. Detecting Structural Changes in Protein by Comparing Deuterium Levels Changes in the non-covalent structures of proteins often involve rearrangement of the intra-molecular hydrogen bonding network and alteration of the solvent accessibility of peptide amide hydrogens. Such structural changes are reflected by changes in the parameters β and Kunf defined in Equation 17.6.6. Furthermore, such structural changes may be reflected by changes in the deuterium levels found in peptides if the measurements are performed under conditions where kint is unchanged. Many applications of HX-MS have followed this approach (i.e., conditions where the pH and temperature are constant). Because the dependence of kint on pH and temperature is known, H/D exchange can also be used to detect changes in structure/dynamics that are induced by changes in pH and temperature. To cancel the effect of pH on the intrinsic rate of H/D exchange requires consideration of the labeling time. The deuterium level at any peptide-amide linkage (D) is given in Equation 17.6.9:
D = 1 − e− kex t Equation 17.6.9
where t is the time the protein was exposed to D2O. According to Equations 17.6.1 and 17.6.6, kex can be expressed as in Equation 17.6.10:
Probing Protein Structure and Dynamics by HX-MS
kex = (β + K unf ) kOH [OH − ] Equation 17.6.10
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Combining Equations 17.6.9 and 17.6.10 results in Equation 17.6.11:
D =1− e
−(β+ Kunf ) kOH [OH − ]t
Equation 17.6.11
which indicates that deuterium levels may be used to detect pH-induced structural changes in folded proteins if different exposure times are used for different pH such that the product [OH−]t is constant. This general approach of using different exposure times to compensate for changes in intrinsic rates can also be used to compare hydrogen exchange results obtained at different temperatures (Zhang and Smith, 1993; Liu and Smith, 1994). This approach has been used to study the pH-induced swelling of the BMV capsid. Table 17.6.1 lists the exposure times used at different pD (D2O buffer), to determine structural changes in the virus capsid when the pH was increased from 5 to 7. These labeling times were chosen to keep the product [OH–]t constant. To satisfy this requirement, the labeling time for pD 5.43 was 74.1 times greater than the labeling time at pH 7.30. The general experimental procedure used for this study is illustrated in Figure 17.6.2. The differences in the deuterium levels found in the peptic fragments from BMV capsid protein–labeled at pD 5.43 and 7.30 are shown in Figure 17.6.5. A set of 16-peptide fragments were reported to cover nearly the entire backbone of the BMV capsid protein (20 kD, 187 residues). The change in deuterium level found for each fragment is expressed in both relative units (the percentage of the total number of amide linkages, given as the y-axis) and absolute units (the number of deuteriums, given above each bar). Both presentation modes have specific merits. Expressing the deuterium difference as the average difference per amide linkage facilitates comparing deuterium changes in fragments of different sizes. On the other hand, showing the difference in the absolute number of deuteriums facilitates comparing these changes with the error of the measurement. The uncertainty of measurement for each segment, expressed as percent, is indicated by the error bars in Figure 17.6.5. These results indicate that H/D exchange was greater in most segments of the capsid protein when the pD was 7.3, suggesting increased flexibility with increasing pH. The largest increases in deuterium level were found in segments including residues 70-90 and 118-151 (shaded dark gray), suggesting that these regions are most susceptible to destabilization with increasing pH. The X-ray crystal structure of cowpea chlorotic mottle virus (CCMV; Speir et al., 1995), a virus closely related to BMV, indicates that these structural changes likely occur at the center of the icosahedral asymmetric trimer.
Table 17.6.1 Exchange Times Used to Label Intact Brome Mosaic Virus in D2O at pD 5.43 and 7.30a,b (Wang and Smith 1999)
pD 7.30 (min) 0.33 1.00 5.50 30.00 90.00
pD 5.43 (min) 24.5 74.1 407.6 2223.0 7113.6
aThe labeling time at pD 5.43 was 74.1 times longer than the labeling time at pD 7.30. bpD values were taken directly from the pH meter without correction.
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50 40 30
4.0
0.9
3.2 4.1
20
1.0
6.1
10
–1.1 –0.8
–0.5 166-170 156-165
–30
183-188
171-182
deuterium change at pD 7 _ + 0
152-155 149-151
136-148
123-135
109-118
118-121
97-108
91-96
70-90
48-61
62-69
–20
0.2
0.3
0 –10
1.1
1.0
0.5
1-47
Deuterium difference (%)
0.9
1.1
Protein fragment
Figure 17.6.5 Changes in deuterium levels found in segments of the BMV capsid protein following labeling of the intact virus particles at pD 5.43 or 7.30. The changes are plotted as rectangular boxes, where the height indicates the change expressed as a percentage of the total number of NHs in the peptide and the width indicates the length of the backbone segment. The change in the absolute number of deuteriums is given on the top of each bar. Shading indicates the relative degree of change found in each region. Adapted from Wang and Smith (2001).
Probing Protein Dynamics by Comparing Hydrogen Exchange Rate Distribution More detailed analysis of time-course data leads to the distribution of exchange rates within a particular segment and information on localized dynamics. The deuterium level measured in a peptic fragment is the sum of the deuterium levels at each peptide amide linkage. The levels at these linkages may span a range of 106 to 108. Although exchange rates at specific linkages cannot be determined from time-course data, the distribution of exchange rates within a peptic fragment can be estimated by fitting the data to a simple model. For example, the four-compartment model illustrated by Equation 17.6.12 has been used to determine the number of amide hydrogens with very fast, fast, slow, and very slow exchange rates (Engen et al., 1999). D = N − [( n1 (e − mi t ) + n2 (e − m2 t ) + n3 (e − m3t ) + n4 (e − m4 t ) Equation 17.6.12
Probing Protein Structure and Dynamics by HX-MS
Variable parameters n1, n2, n3, and n4 indicate the number of linkages where the H/D exchange rates are fast, intermediate, or slow (m1, m2, m3, and m4, respectively). Although the model used here is a major simplification of reality, it does allow one to distinguish between conformations where all linkages exchange at the same rate and conformations where there is a wide distribution of exchange rates. Other useful approaches for determining the distribution of exchange rates within peptic fragments of proteins have been described (Zhang and Smith, 1993; Zhang et al., 1997).
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16.0
Deuterium level
12.0
8.0
4.0
0.0 0.01
0.1
1 10 Exchange time (min)
100
1000
Figure 17.6.6 Time-course exchange-in plot of segment 7-27 of free SH2 (solid triangles) or SH2 bound with a short peptide (open circles). Adapted from Engen et al. (1999).
Application of Equation 17.6.12 is illustrated using exchange-in data for a peptic fragment of the SH2 domain of hematopoietic cell kinase (Hck). The SH2 domain was labeled for 10 sec to 8 hr in D2O buffer (pH 6.9) with and without a 12-residue peptide known to bind to SH2. Deuterium levels found in the peptic fragment including residues 7-27 are presented in Figure 17.6.6 for both free and bound SH2. These results show that the bound form always has less deuterium than the free form. Application of Equation 17.6.12 to these data (see Table 17.6.2) shows that rate constants for H/D exchange within this short segment span a range >0.001 to 4 min−1. For the free form of SH2, this analysis shows that the hydrogens at 9 amide linkages exchange with rate constants >4 min−1 while three exchange with rate constants <0.001 min−1. Similar analysis of exchange-in data for SH2 bound to a peptide shows that the number of linkages where exchange occurred with kex >4 min−1 has decreased from 9 to 8, and the number of linkages where exchange occurred with kex <0.001 min−1 has increased by 3 to 4. A convenient means for illustrating how binding to the SH2 domain alters the distribution of H/D exchange rates within the segment including residues 7-27 is presented in Fig. 17.6.7. In this presentation, the number of amide linkages with a certain average rate of exchange is plotted versus the
Table 17.6.2 Distribution of Rate Constants for Isotope Exchange at Peptide Amide Linkages in a Peptic Fragment (Residues 7-27) of Hck SH2–Free and Bound to 12-Residue Peptide (Engen, et al., 1999)
Segmenta 7-27 7-27 bound
Categoryb Very fast
Fast
Slow
Very slowc
9.0 (>4.0) 8.0 (>4.0)
2.5 (2.1) 3.0 (1.6)
4.5 (0.004) 1.5 (0.006)
3.0 (<0.001) 6.5 (<0.001)
aPeptic segment 7-27 and other segments of Hck SH2 were identified in preliminary experiments by techniques
described previously. bThe number of peptide amide linkages in each category are given outside the parentheses, while the isotope exchange rates (min−1) are indicated inside the parentheses. cThe number of very slow NHs was determined by subtracting the number of NHs exchanged in 8 hr from the total number of NHs in the segment.
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10
Number of NHs
8 6 4 2 0 0
0.01
0.10
1.0
Exchange rate (min–1)
Figure 17.6.7 Distribution of amide hydrogen exchange rates in segment 7-27 derived from free SH2 (solid circles) and SH2 bound with the peptide (open circles). Adapted from Engen et al. (1999).
exchange rate, which is plotted as a log scale because it spans a range of several orders of magnitude. The ability to resolve deuterium exchange-in data into a distribution of exchange rates opens the possibility for determining whether exchange occurred from the folded or unfolded forms of a protein (i.e., models described by Equations 17.6.2 and 17.6.3). As illustrated by Equation 17.6.6, amide-hydrogen exchange can be viewed as the sum of contributions of exchange from folded (kex,f) and unfolded (kex,u) forms of the protein. It has been proposed that amide hydrogens exchanging very fast (i.e., little protection) must do so from the folded form of a protein (Engen et al., 1999). However, highly protected hydrogens may exchange from either unfolded or folded forms of a protein. Furthermore, structural changes required for hydrogen exchange from folded and unfolded forms of a protein differ in the magnitude of atomic displacements at individual peptide amide linkages. Exchange in a folded and compact protein is believed to occur through small atomic movements, probably ∼1 Å, but sufficient to allow diffusion of OD− and D2O to the exchange sites (Elber and Karplus, 1987; Kim and Woodward, 1993). Exchange in momentarily unfolded forms of a protein occurs only after many or all residues have unfolded. It follows that exchange from unfolded forms of a protein requires large movements of many or all of the atoms in a molecule. The reduction in the number of amide hydrogens exchanging very fast with ligand binding (see Table 17.6.2 and Figure 17.6.7) was interpreted as a reduction in exchange from the folded form of SH2 (Engen et al., 1999). SUMMARY Hydrogen exchange/mass spectrometry is playing a more and more important role in studies of protein structure and dynamics. Using this technique, protein can be studied at a wide range of conditions, such as different pHs and temperatures, high concentration of denaturants, or as a component of a large heterogeneous system. In addition, both global and localized structure and dynamics features of protein can be characterized. ACKNOWLEDGEMENT Probing Protein Structure and Dynamics by HX-MS
This work was supported by a grant from the National Institutes of Health (GM 40384) and the Nebraska Center for Mass Spectrometry.
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LITERATURE CITED Arrington, C.B., Teesch, L.M., and Robertson, A.D. 1999. Defining protein ensembles with native-state NH exchange: Kinetics of interconversion and cooperative units from combined NMR and MS analysis. J. Mol. Biol. 285:1265-1275. Bai, Y., Milne, J.S., Mayne, L., and Englander, S.W. 1993. Primary structure effects on peptide group hydrogen exchange. Proteins Struct. Funct. Genet. 17:75-86. Bai, Y.W., Milne, J.S., Mayne, L., and Englander, S.W. 1994. Protein stability parameters measured by hydrogen exchange. Proteins Struct. Funct. Genet. 20:4-14. Connelly, G.P., Bai, Y., Jeng, M.-F., and Englander, S.W. 1993. Isotope effects in peptide group hydrogen exchange. Proteins Struct. Funct. Genet. 17:87-92. Deng, Y., Pan, H., and Smith, D.L. 1999. Selective isotope labeling demonstrates that hydrogen exchange at individual peptide amide linkages can be determined by collision-induced dissociation mass spectrometry. J. Am. Chem. Soc. 121:1966-1967. Deng, Y. and Smith, D.L. 1999. Rate and equilibrium constants for protein unfolding and refolding determined by hydrogen exchange-mass spectrometry. Anal. Biochem. 276:150-160. Deng, Y., Zhang, Z., and Smith, D.L. 1999. Comparison of continuous and pulsed labeling amide hydrogen exchange/mass spectrometry for studies of protein dynamics. J. Am. Soc. Mass Spectrom. 10:675-684. Ehring, H. 1999. Hydrogen exchange/electrospray ionization mass spectrometry studies of structural features of proteins and protein/protein interactions. Anal. Biochem. 267:252-259. Elber, R. and Karplus, M. 1987. Multiple conformational states of proteins: A molecular dynamics analysis of myoglobin. Science 235:318-321. Engen, J.R., Gmeiner, W.H., Smithgall, T.E., and Smith, D.L. 1999. Hydrogen exchange shows peptide binding stabilizes motions in Hck SH2. Biochemistry 38:8926-8935. Englander, J.J., Rogero, J.R., and Englander, S.W. 1985. Protein hydrogen exchange studied by the fragment separation method. Anal. Biochem. 147:234-244. Englander, S.W. and Kallenbach, N.R. 1984. Hydrogen exchange and structural dynamics of proteins and nucleic acids. Q. Rev. Biophys. 16:521-655. Englander, S.W., Mayne, L., Bai, Y., and Sosnick, T.R. 1997. Hydrogen exchange: The modern legacy of Linderstrom-Lang. Protein Sci. 6:1101-1109. Gross, M., Robinson, C.V., Mayhew, M., Hartl, F.U., and Radford, S.E. 1996. Significant hydrogen exchange protection in GroEL-bound DHFR is maintained during iterative rounds of substrate cycling. Protein Sci. 5:2506-2513. Halgand, F., Dumas, R., Biou, V., Andrieu, J.P., Thomazeau, K., Gagnon, J., Douce, R., and Forest, E. 1999. Characterization of the conformational changes of acetohydroxy acid isomeroreductase induced by the binding of Mg2+ ions, NADPH, and a competitive inhibitor. Biochemistry 38:6025-6034. Hvidt, A. and Nielsen, S.O. 1966. Hydrogen exchange in proteins. Adv. Protein Chem. 21:287-385. Jeng, M.-F., Englander, S.W., Elöve, G.A., Wand, A.J., and Roder, H. 1990. Structural description of acid-denatured cytochrome c by hydrogen exchange and 2D NMR. Biochemistry 29:10433-10437. Katta, V. and Chait, B.T. 1991. Conformational changes in proteins probed by hydrogen-exchange electrospray-ionization mass spectrometry. Rapid Commun. Mass Spectrom. 5:214-217. Kim, K.-S., Fuchs, J.A., and Woodward, C.K. 1993. Hydrogen exchange identifies native-state motional domains important in protein folding. Biochemistry 32:9600-9608. Kim, K.-S. and Woodward, C. 1993. Protein internal flexibility and global stability: Effect of urea on hydrogen exchange rates of bovine pancreatic trypsin inhibitor. Biochemistry 32:9609-9613. Li, R. and Woodward, C. 1999. The hydrogen exchange core and protein folding. Protein Sci. 8:1571-1590. Liu, Y. and Smith, D.L. 1994. Probing high order structure of proteins by fast-atom bombardment mass spectrometry. J. Am. Soc. Mass Spectrom. 5:19-28. McCloskey, J.A. 1990. Mass spectrometry. Methods Enzymol. 193. Miller, D.W. and Dill, K.A. 1995. A statistical mechanical model for hydrogen exchange in globular proteins. Protein Sci. 4:1860-1873. Miranker, A., Robinson, C.V., Radford, S.E., Aplin, R.T., and Dobson, C.M. 1993. Detection of transient protein folding populations by mass spectrometry. Science 262:896-900. Molday, R.S., Englander, S.W., and Kallen, R.G. 1972. Primary structure effects on peptide group hydrogen exchange. Biochemistry 11:150-158. Rosa, J.J. and Richards, F.M. 1979. An experimental procedure for increasing the structural resolution of chemical hydrogen-exchange measurements on proteins: Application to ribonuclease S peptide. J. Mol. Biol. 133:399-416.
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Smith, D.L., Deng, Y., and Zhang, Z. 1997. Probing the non-covalent structure of proteins by amide hydrogen exchange and mass spectrometry. J. Mass Spectrom. 32:135-146. Speir, J.A., Munshi, S., Wang, G., Baker, T.S., and Johnson, J.E. 1995. Structures of the native and swollen forms of cowpea chlorotic mottle virus determined by X-ray crystallography and cryo-electron microscopy. Structure 3:63-78. Thévenon-Emeric, G., Kozlowski, J., Zhang, Z., and Smith, D.L. 1992. Determination of amide hydrogen exchange rates in peptides by mass spectrometry. Anal. Chem. 64:2456-2458. Wang, L., Pan, H., and Smith, D.L. 2002. Hydrogen exchange/mass spectormetry: Optimization of digestion conditions. Mol. Cell. Proteom. 1:132-138. Wan, L., Lane, L., and Smith, D.L. 2001. Detecting structural changes in viral capsids by hydrogen exchange and mass spectrometry. Protein Sci. 10:1234-1243. Woodward, C., Simon, I., and Tuchsen, E. 1982. Hydrogen exchange and the dynamic structure of proteins. Mol. Cell. Biochem. 48:135-160. Yang, H. and Smith, D.L. 1997. Kinetics of cytochrome c folding examined by hydrogen exchange and mass spectrometry. Biochemistry 36:14992-14999. Zhang, Z., Li, W., Logan, T.M., Li, M., and Marshall, A.G. 1997. Human recombinant [C22A] FK506-binding protein amide hydrogen exchange rates from mass spectrometry match and extend those from NMR. Protein Sci. 6:2203-2217. Zhang, Z., Post, C.B., and Smith, D.L. 1996. Amide hydrogen exchange determined by mass spectrometry: Application to rabbit muscle aldolase. Biochemistry 35:779-791. Zhang, Z. and Smith, D.L. 1993. Determination of amide hydrogen exchange by mass spectrometry: A new tool for protein structure elucidation. Protein Sci. 2:522-531.
Lintao Wang and David L. Smith University of Nebraska Lincoln, Nebraska
Probing Protein Structure and Dynamics by HX-MS
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Introduction to Atomic Force Microscopy (AFM) in Biology The atomic force microscope has the unique capability of imaging biological samples with molecular resolution in buffer solution. In addition to providing topographical images of surfaces with nanometer- to angstrom-scale resolution, forces between single molecules and mechanical properties of biological samples can be investigated. Importantly, the measurements are made in buffer solutions, allowing biological samples to “stay alive” within a physiological-like environment while temporal changes in structure are measured—e.g., before and after addition of chemical reagents to the fluid cell. These qualities distinguish AFM from conventional imaging techniques of comparable resolution—i.e., electron microscopy (EM). However, limitations of AFM are that the specimen must be firmly attached to a solid support for measurement and the information gained is generally restricted to regions of the specimen that are directly exposed to the scanning tip—i.e., its surface structures. This unit provides an introduction to AFM on biological systems and describes specific examples of AFM on proteins. The physical principles of the technique and methodological aspects of its practical use and applications are also described. The principle of AFM as a member of the class of scanning probe microscopies (SPM) was first described by Binnig et al. (1986). Subsequent development of the technique for biological applications has enabled investigations of a range of different samples; individual protein or nucleic acid molecules, membranes, living cells, and tissue samples. AFM imaging is fundamentally different from conventional microscopies: a fine tip attached to a cantilever “spring” is scanned over the sample surface, ideally in a physiological buffer solution (Figure 17.7.1). The interaction between sample and tip causes the cantilever to bend and the bending of the cantilever is detected by the deflection of a laser beam focused on the end of the cantilever (other detection methods also exist, however, the laser-beam method is most commonly used in commercial AFMs). From the cantilever bending, a force between the sample and tip can be calculated (typical range: pN to nN). A piezoelectric (polycrystalline) ceramic (piezo) is used to scan in the x and y direction and to adjust the vertical distance
between sample and tip. In “contact mode” the feedback loop between the laser detector and the piezo adjusts the vertical distance from tip to sample such that the cantilever is held at constant deflection. In “dynamic mode,” the feedback loop keeps the amplitude of a vibrating cantilever constant by correcting the vertical distance. Thus, in both modes, surface topographies are obtained and sample heights can be measured within a sub-nanometer resolution range. Additionally, the bending of the cantilever can be recorded as a function of its distance from the surface in the form of force-distance curves (Figure 17.7.2). Forces felt by the cantilever as the tip approaches and retracts from the sample can reveal long-range attractive or repulsive forces between the tip and sample surfaces, allowing measurements of electrostatic, chemical, or mechanical properties such as adhesion or elasticity (if the tip is used to indent the sample) on a nanometer scale. “Force mapping” was a technique developed to determine the elastic modulus at multiple points on a sample surface and thus correlate elasticity with topography, e.g., over the structure of a cell surface. Furthermore, by immobilizing single molecules on the tip and the surface, AFM can be used to unfold proteins or to pull single receptor-ligand complexes apart, allowing binding and rupture forces to be determined. The following reviews provide comprehensive discussions of the various applications of AFM: Lal et al. (1994), Bustamante et al. (1997), Hansma et al. (1997), Engel et al. (1999), Heinz and Hoh (1999), Allison et al. (2002), ClausenSchaumann et al. (2000), Engel and Müller (2000), Fisher et al. (2000), Stolz et al. (2000).
EXPERIMENTAL SETUP Many factors influence the quality of AFM data. Namely the characteristics of the sensing tip, the choice of application mode (“contact” or “dynamic” mode), the quality of the support to which the sample is adsorbed, and of course, the sample preparation itself. These points are discussed in more detail below.
The Tip and Cantilever The tip and the cantilever play important roles in the quality of the data and resolution of the AFM. As biological samples are soft and delicate, tips attached to cantilevers with low
Contributed by Claire Goldsbury and Simon Scheuring Current Protocols in Protein Science (2002) 17.7.1-17.7.17 Copyright © 2002 by John Wiley & Sons, Inc.
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can then be selected depending on the particular application mode. Standard tips have a curvature radius of ∼5 nm. For high resolution imaging, sharper tips such as oxide sharpened or contact etched are also commercially available. An increase in resolution can also be attained in some cases by attaching a carbon nanotube to the end of the tip (Cheung et al., 2000). Tips may be further modified by chemical or adsorptive functionalization, e.g., for the measurement of specific binding and rupturing forces, a ligand may be attached to the tip and its corresponding receptor immobilized to the surface (see also Preparation of the Support and Specimen Attachment; Dammer et al., 1996; Fritz et al., 1999; Raab et al., 1999). For some
spring constants (<1 N/m) are preferable (the lower the spring constant of the cantilever, the less force exerted between tip and sample during imaging allowing finer adjustment of force). Commonly used commercial cantilevers are made of silicon nitride and have an integrated sharp tip at their end. Typically, cantilevers for biological applications are 100 to 200 µm in length and have spring constants of 0.06 to 0.6 N/m. For dynamic mode, cantilevers are excited close to their resonance peak, typically between 8 and 9 kHz in aqueous solution. In most cases, there are up to four cantilevers with different spring constants attached to a single substrate (Figure 17.7.3A). This substrate is mounted in the AFM and the cantilever
A
B laser diode F
detector cantilever fluid cell
buffer solution
z y
sample piezotube
x
C
Introduction to Atomic Force Microscopy (AFM) in Biology
Figure 17.7.1 Scheme showing how AFM produces images of surfaces. (A) The sample surface is scanned with a fine tip. The interaction force between the tip and the surface causes the cantilever to bend. (B) The sample is mounted in the AFM under a transparent cantilever holder (fluid cell). A laser beam is focused on the end of the cantilever. The deflection of the laser spot, induced by the bending of the cantilever, is measured by a photodiode detector and a feed-back loop corrects the vertical distance by adjusting the piezo position. (C) The AFM thereby produces a topographical image of the sample (surface of “constant force”). This figure was obtained from the M.E.MuellerInstitute for Structural Biology, Basel, Switzerland, publication “Imaging, measuring and manipulating native biomolecular systems with the atomic force microscope” prepared by Daniel J. Müller, Ueli Aebi, and Andreas Engel.
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contact
no contact
3
Force
2
1
0 contact
4
0
Distance
Figure 17.7.2 Force versus distance curve. The deflection (y-axis) of the free end of the cantilever is measured as the fixed end of the cantilever is brought vertically towards and then away from the sample surface. (1) The tip is initially not touching the surface—no cantilever deflection occurs. (2) As the tip approaches the surface, attractive forces cause it to “jump” into contact with the sample (note: this adhesive force is less pronounced when acquiring images under physiological conditions, i.e., in liquid, as compared to air). (3) Cantilever deflection becomes maximal when the tip is in contact with the sample (relates linearly to further approach to the sample as the tip is already in contact). If the tip indents the surface, elasticity forces related to the sample can be measured from the force curve. (4) As the tip retracts from the sample, adhesion or specific receptor-ligand binding forces can cause the cantilever to stay attached to the sample some distance past the original point of contact. The point on the force curve where the tip eventually jumps off the surface can thus provide information about bond rupturing forces. Adapted from Digital Instruments.
of these applications, precise determination of the cantilever spring constant is required to calibrate the measured forces. Several publications address this problem (e.g., Hutter and Bechhoefer, 1993; Neumeister and Ducker, 1994). It may be desirable to re-use a particular tip for measuring another sample. In this case, tips may be cleaned, e.g., in 1% SDS followed by three washes in distilled water (Scheuring et al., 2001), in helmanex (Hörber et al., 1995), or by ultraviolet radiation (Thomson et al., 1996). Further useful information on tips, including modification of and cleaning, can be found at the Digital Instruments Web site: http://www.di.com/AppNotes_PDFs/AN44Bi otipsApps.pdf. Also see Tip Shape for a discussion of experimental artifacts related to the tip and Table 17.7.1 for a list of tip manufacturers.
The Piezo Piezos are used to either scan the cantilever over a fixed sample or actuate the sample relative to a fixed cantilever. Thus, depending on the design of the AFM, either the tip or the sample undergoes corrective vertical movement, but the principle of the feedback control is the same in both cases. Piezos are commercially available in various sizes and differ mainly in their maximum scan range and in their precision of x, y, and z movements. Usually, the larger the maximal scan range, the lower the precision. The accuracy of the piezo in x, y and z dimensions is typically given by the manufacturer but has to be recalibrated periodically. The precision can be tested relatively easily before each experiment by scanning a 10-µm2 or a 1-µm2 calibration grid with 1-µm grooves. Atomic-scale calibration in x and y direction is easily performed with a mica
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A
B
Figure 17.7.3 (A) Substrate with two cantilevers attached. (B) Integration of a fine tip on the end of a cantilever. Images are from Digital Instruments: http://www.di.com/products2/NewProbeGuide/ ContactModeProbes.html.
Table 17.7.1
Introduction to Atomic Force Microscopy (AFM) in Biology
AFM and Tip Manufacturers
AFM manufacturers
Web sites
Digital Instruments JEOL Molecular Imaging Omicron Pacific Scanning Park Scientific Quesant Instrument Corporation Thermo Microscopes (Merger of Park Scientific and Topometrix) Tip Manufacturers Molecular Devices and Tools (NT-MDT) Olympus cantilevers
http://www.di.com/ http://www.jeol.com/spm/spm.html http://www.molec.com/ http://www.omicron-instruments.com/ http://www.pacificnanotech.com/ http://www.park.com/ http://www.quesant.com/ http://www.thermomicro.com/
http://www.siliconmdt.com/ http://www.olympus.co.jp/probe/index.html
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surface (hexagonal lattice). Atomic precision in the z dimension is more difficult to determine, but it can be tested by scanning MoTe2, a layered dihalcogenide crystal that reveals terraces of atomic layers with a height step-size of 7Å (Müller and Engel, 1997). Since the scan ranges of piezos are fairly small (typically only up to 200 µm2) compared to the sample area, e.g., of an electron microscopy grid, it is not always easy to find nicely adsorbed objects within this region. Hence, the choice of the appropriate piezo is important. Usually, it is of interest to start investigations with piezos that have a scan range >100 µm to initially find the adsorbed objects and to improve the preparation. If the sample can be adsorbed with high surface density, one can then change to a smaller piezo, which might be advantageous for higher resolution imaging.
Measurement Modes Contact mode In contact mode, the tip is permanently in contact with the sample. Contact mode can be used throughout a large scan and resolution range, from imaging of whole cells at a lateral resolution of ∼10 nm to single molecules and 2-D crystalline protein arrays at a resolution of ∼5 Å. Although temporal resolution is possible, contact mode imaging is restricted to samples that are firmly attached to the support and exhibit strong lateral stability. Dynamic mode Not all specimens can be firmly attached to a solid support. Single molecules or filamentous structures have less surface area for adhering to the support (i.e., compared to crystalline protein arrays), and are often pushed away by the tip during scanning. In some cases, reduced surface adherence is even preferred in order to preserve or increase temporal resolution of protein movement and activity (Chen et al., 1998; Bustamante et al., 1999). Dynamic mode, also called “MAC Mode” or “Tapping-mode” (MAC Mode, Molecular Imaging; TappingMode, Digital Instruments) was developed to address these issues. In dynamic mode, the tip is oscillated in fluid at a fixed resonance frequency and a given amplitude. The reduced contact time of the oscillating tip with the sample reduces lateral frictional forces compared to contact mode, and sample molecules are therefore less likely to be pushed away. For biological samples, the same tips used for contact mode can be used in dynamic mode. Nor-
mally, dynamic mode has a lower resolution compared to contact mode. Nevertheless, recent advances have demonstrated that with optimal operating conditions, dynamic mode is capable of attaining image resolution comparable to that of contact mode (Möller et al., 1999). A drawback of dynamic mode is the slow scan rates required, typically a factor of four slower than contact mode. New innovations in microscope and tip design (Viani et al., 1999, 2000) are needed to circumvent this shortcoming and ultimately allow higher temporal resolution. More information on dynamic mode can be found at: http://www.molec.com/products/PicoSPM/ MAC/MACMode.html and http://www.di.com/ AppNotes/FluidTap/FluidMain.html. Force curve measurement The ability of AFM to measure molecular interaction forces has found wide applications. First, the tip can be used to indent the specimen and, from the resulting surface deformation, one can determine the mechanical properties of the sample (Radmacher et al., 1994). Such experiments allow calculation of the elastic moduli of a wide range of biological samples from tissues and cells to protein layers. Second, the shape of the force curve during retraction of the tip from a surface can be used to elucidate attractive forces of nonspecific nature, as well as highly specific binding between biological molecules (e.g., antigen-antibody). Third, stretching and unfolding experiments on single molecules can be conducted. For the latter two techniques, the term “force-spectroscopy” has been used.
Preparation of the Support and Specimen Attachment The sample can either be physically adsorbed or covalently bound to a flat and stable support. The quality of this support is one of the most critical factors for obtaining optimal results in AFM. Poorly prepared supports result in vibrational oscillations, so that only poor image resolution can be achieved. A reliable method for preparing AFM supports has been described by Schabert and Engel (1994). In this publication, supports are 5-mm discs of muscovite mica (Mica) attached to 10-mm teflon discs using a two-component araldite glue (Ciba-Geigy, or Devcon Epoxy, ITW Brands). The hydrophobic teflon repels the fluid droplet and thus contains it on the mica surface and protects the piezo from contact with the buffer solution. Before attaching the mica, the teflon discs are glued to metal stubs using superglue.
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Introduction to Atomic Force Microscopy (AFM) in Biology
The mica and teflon discs are prepared by punching them out of thin sheets of the respective materials. Key factors in support preparation to minimize subsequent vibrational oscillations in the AFM are (1) ensuring that the surfaces to be glued are clean and (2) ensuring that the surfaces are sealed, i.e., there are no bubbles under the glued surfaces. Other material with different properties, e.g., hydrophobic highly oriented pyrolitic graphite (HOPG) or glass, can accordingly replace mica as a support attached to the teflon disc. For adsorption of the specimen, mica, HOPG, or other surfaces should be as clean as possible. For this purpose, mica and HOPG can easily be cleaved to reveal a fresh and atomically flat surface. Glass can be washed, e.g., with concentrated HCl/HNO3 (3:1) and rinsed with ultrapure water in a sonicating bath to obtain a clean flat surface (Karrasch et al., 1993). Not all biological preparations will adhere to standard AFM specimen supports such as mica, HOPG or glass. In these cases, various alternatives must be explored. As one example, native Xenopus oocyte nuclear envelopes could not be immobilized on mica or glass, but were successfully attached to 400-mesh copper electron microscopy (EM) grids (Plano) coated with parlodion and evaporated carbon (also see Time-Lapse imaging, Figure 17.7.7b; Stoffler et al., 1999). The EM grids were glued onto 12-cm diameter plastic petri dishes using 5-min araldite glue. Depending on how an isolated oocyte nuclear envelope (NE) was opened and the charge of the EM grid, the NE preferentially adhered by its cytoplasmic or nuclear face to the carbon support. Non-covalent attachment of additional material, e.g., lipid layers to the support can also facilitate adsorption of the protein of interest. This was achieved in the case of actin filaments which bound to lipid bilayers attached to mica using the Langmuir-Blodgett technique (Stolz et al., 2000; Shi et al., 2001). Specific protein-lipid interactions have also been studied using this technique (Czajkowsky et al., 1999; Rinia et al., 2000). Lipids may also be adsorbed to mica using the vesicle fusion technique. Highly specialized supports for covalently binding the sample may be prepared by chemical modification of the support, e.g., by silanization of glass, functionalizing gold surfaces with thiol groups (for Cys residue binding) or by deposition of functionalized lipid bilayers onto the support (Karasch et al., 1993; Müller et al., 1997a; Reviakine et al., 1998; Wagner, 1998). In order to measure specific molecular interactions (using force spec-
troscopy), both the surface and the tip have to be functionalized. This can be achieved through functionalization of the respective surfaces with linker molecules and subsequent cross linking with the interacting molecules (Rief et al., 1997). Often, the tip and support are coated with gold, or the strong interaction of streptavidin-biotin is exploited to attach biotinylated molecules. Sometimes, simple physical adsorption of the molecules to the two respective surfaces is sufficient. Adsorption of the specimen to mica or HOPG Firm adsorption of a specimen to the support depends on buffer conditions (Figure 17.7.4). In the case of a hydrophilic negatively charged specimen adsorbing to mica, adsorption buffers must be of sufficient ionic strength to shield surface charges and consequently shield longrange electrostatic repulsive force interactions (Müller et al., 1997c). A good starting point for adsorption is a buffer at pH 7.3 with an ionic strength of ∼300 mM monovalent ions or up to ∼50 mM divalent ions; however, adsorption buffer conditions must be optimized for each new sample. In the case of a partially hydrophobic sample, a hydrophobic support such as HOPG might be preferable to mica. In this case, hydrophobic interactions will mediate sample immobilization, e.g., immobilization of lipid monolayers (Scheuring et al., 1999b).
APPLICATIONS AFM can be used to measure an abundance of molecular properties—from structure to unfolding forces. In this section, various applications are outlined and illustrated with specific examples under the headings of High-Resolution Imaging, Time-Lapse Imaging, and Force Spectroscopy. Methodological aspects of the experimental set up for each application are summarized, however, it should be noted that these conditions often have to be determined empirically for each new sample.
High-Resolution Imaging Biological samples are soft and fragile. In order to preserve the sample and not distort its conformation, loading forces between the AFM tip and specimen must be minimized. The forces between tip and specimen are a composite of electrostatic forces (at long distances: up to 20 nm), van der Waals forces (at short distances: in the range of several Angstroms), and Pauli-repulsion forces (∼1Å). The AFM tip and proteins both expose charged surfaces in aque-
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ous solution. Consequently, these surfaces attract counter-ions which shield the charges and form a diffuse electric layer. When two equally charged surfaces approach one another, the counter-ion distribution of both surfaces are disturbed resulting in repulsive forces. The thickness of the counter-ion layer is influenced by the total amount of ions in the buffer solution; consequently, the distance from the surface where repulsive force interactions appear can be altered. Taken together, the pH and/or ions in the buffer solution influence the charges of the interacting surfaces and hence the interacting forces. Adjustment of the buffer conditions can minimize the loading force between AFM tip and sample (Figure 17.7.5). A good starting point for scanning buffer screening is
A
LiCl NaCl
400 Adsorption density [PM / 1000 µm2]
pH 7.3 with an ionic strength of ∼100 mM monovalent ions. Some biological samples might expose a hydrophobic surface. Often, force curves between the hydrophilic AFM tip and a hydrophobic sample show attractive forces. Such interactions can damage the specimen. In these cases, the rationale outlined in the above paragraph is not valid. Buffers with low ionic strength (e.g., 20 mM monovalent ions) are preferable in these cases, possibly because smaller hydrophilic surface regions generate repulsive forces at such an ionic strength, which together with the attractive interaction regions average out to slightly favorable repulsive force interactions.
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Figure 17.7.4 Adsorption density of purple membrane to mica as a function of the electrolyte concentration. Two micrograms of protein was incubated for 5 min at room temperature on 6-mm-diameter supports in buffer containing different amounts of (A) monovalent or (B) divalent ions. Adapted from Müller et al., 1997c.
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400 pN
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Figure 17.7.5 Force-distance curves recorded on the extracellular surface of purple membrane. Force-distance curves were recorded during the approach of sample and AFM tip under different electrolyte concentrations at constant pH (7.6). The dotted lines represent force-distance curves recorded on purple membrane without electrostatic repulsion. Conditions: scan frequency: 1.97 Hz, scan range: 50 nm (512 pixels). Arrows (1) mark the onset of measurable electrostatic repulsion, whereas arrows (2) indicate the point of contact between tip and sample. Adapted from Müller et al. (1999c).
Introduction to Atomic Force Microscopy (AFM) in Biology
High-resolution imaging has so far been mainly restricted to crystalline samples or proteins densely packed within a membrane (Karrasch et al., 1994, Schabert et al., 1995; Müller et al., 1997b, 1999c; Scheuring et al., 1999a, 2001b; Fotiadis et al., 2000), not because the signal-to-noise ratio of the AFM is insufficient to detect widely dispersed individual proteins, but for reasons relating to the lateral flexibility of individual molecules adsorbed to a support. Single isolated molecules are normally very flexible and are easily pushed away by the scanning AFM tip, which makes their highresolution imaging impossible. Without the stabilizing element of a two-dimensional network, single molecules often must be crosslinked to the support to increase their stability. Dynamic mode (see discussion on Dynamic Mode) can
sometimes circumvent these problems by reducing the applied lateral force loaded on to the sample. Tip convolution effects also limit resolution of single molecules (see Tip Shape). This is due to broadening of the image in the x-, y-dimension, which decreases the lateral resolution to as low as 30Å. Nevertheless, it has been shown for membrane proteins densely packed in lipid bilayers that imaging of noncrystalline regions can yield sub-nanometer resolution (Scheuring et al., 1999a, 2001a; Fotiadis et al., 2000; Seelert et al., 2000; Stahlberg et al., 2001). From such topographs, averages could be calculated using single-particle alignment routines (Table 17.7.2). Similarly for soluble proteins, the highest resolution topographs of non-crystalline molecules have been achieved on densely packed proteins that
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Table 17.7.2
Data Processing Softwarea
Averaging of proteins ordered in a 2-D-lattice: MRC CRISP
ftp://ftp.mrc-lmb.cam.ac.uk/ http://www.calidris-em.com/
Averaging of proteins ordered in a 2-D-lattice and single particle averaging: http://www.synoptics.co.uk/ SEMPER http://www.ImageScience.de/ Imagic-5 Single particle averaging: http://www.wadsworth.org/spider_doc/spider/docs/spider.html SPIDER General image treatment: http://www.adobe.com/ Photoshop ftp://rsbweb.nih.gov/pub/nih-image/ NIH Image (can be powered for AFM image analysis by Image SXM by St. Barrett, http://reg.ssci.liv.ac.uk/) Software for analyzing force curves: http://www.nanotec.es/ WSxM http://www.cp.umist.ac.uk/Analysis/spmcon.html SPMcon http://www.wavemetrics.com/ Igor (can be powered with routines written by users; see the forum for AFM users on the di-page) http://www.imagemet.com/ SPIP http://www.forceview.de/ Forceview2000 aThis list of image processing software is by no means comprehensive. Some of the programs are downloadable
freeware while some are fairly expensive. Most of them are used for analysis of electron micrographs and must be adapted to the special needs for analyzing atomic-force microscopy images.
Figure 17.7.6 High-resolution topograph of a two-dimensional crystal of the membrane protein aquaporin Z (AqpZ) using contact-mode AFM recorded in buffer solution. Adapted from Scheuring et al., 1999a.
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adsorbed to the support unidirectionally (Mou et al., 1995, 1996a,b; Czajkowsky et al., 1998; Dorn et al., 1999). High-resolution AFM imaging has been shown to directly yield information about protein flexibility (Müller et al., 1998; Scheuring et al., 2002b; see Figure 17.7.6). Topographical image processing yields both average height and standard deviation values of protein surfaces. High standard deviations in surface averages indicate high regional protein flexibility. The flexibility of distinct protein regions can be related to functionally flexible peptide domains. See Table 17.7.2 for image treatment software programs.
Time-Lapse Imaging: Proteins at Work
Introduction to Atomic Force Microscopy (AFM) in Biology
With the introduction of dynamic mode AFM (see discussion on Dynamic Mode), researchers were given the possibility of repetitive imaging of proteins only weakly attached to the support. An early example of this application was the measurement of movement of a single DNA molecule through an RNA polymerase enzyme (Figure 17.7.7A; Kasas et al., 1997; Bustamante et al., 1999). As outlined in the High-Resolution Imaging section, it is important to minimize the force exerted on the sample during scanning and to optimize buffer conditions. The force in dynamic mode may be modulated by altering the amplitude of oscillation and/or the distance of the tip from the sample. Nonetheless, due to lower forces involved between the tip and the sample, the buffer can be exchanged while measurements are taking place, thus allowing responses to chemical effectors to be measured directly. Dynamic mode is not required for timelapse imaging if the sample can be attached firmly enough to the support. The structural changes induced in native nuclear pore complexes upon addition of calcium (Figure 17.7.7B, Stoffler et al., 1999) have been measured in contact mode as have cellular dynamics in cultured cells (Hoh and Schoenenberger, 1994; Schoenenberger and Hoh, 1994; Hofmann et al., 1997). AFM with temporal resolution has also found an application in the field of protein crystallography. Crystallization can be monitored in situ at high resolution, allowing insight into the mechanisms and kinetics of the growth process (Reviakine et al., 1998; McPherson et al., 2000, 2001). Similarly, protein aggregation/amyloid fibril formation can be measured in vitro at the level of individual fibrils growing on a supporting surface (Figure
17.7.7C). This has been achieved in the case of human amylin—the peptide forming pancreatic amyloid deposits in type 2 diabetes mellitus, as well as the amyloid beta protein (Aβ; Goldsbury et al., 1999, 2001). Clearly, all of these applications require the specimen to be attached to a support, so surface effects may play a role in the processes being measured.
Force Spectroscopy: Single Molecule Unfolding and Bonding Interactions As well as being used for imaging, the AFM tip can be used to perform high precision forcespectroscopy experiments (see Figure 17.7.2, Introduction to unit and discussion on Force Curve Measurement) on biological samples under physiologically relevant conditions. Indeed, imaging and force spectroscopy can be elegantly combined. AFM force spectroscopy has wide applications including (1) determination of the elastic modulus of organic films, protein layers, cells, and tissues (Radmacher et al., 1995; Radmacher, 1997; Domke et al., 1999); (2) determining the forces involved in antibody-antigen and other ligand-receptor interactions (Florin et al., 1994; Allen et al., 1996; Dammer et al., 1996; Fritz et al., 1998; Raab et al., 1999)—force mapping may be applied to determine, e.g., the receptor distribution on a surface; (3) molecule stretching (Rief et al.,1997a); (4) measuring forces associated with protein unfolding (Mitsui et al., 1996; Rief et al., 1997b, 2000; Carrion-Vazquez et al., 1999; Fisher et al., 1999a,b; Oesterhelt et al., 2000); (5) measuring electrostatic charges (Xu et al., 1995; Levadny et al., 1996; Müller and Engel 1997); (6) determination of protein layer stability (Müller et al., 1999a; Scheuring et al., 2002a); and (7) measuring enzyme activity (Radmacher et al., 1994). Such measurements have provided insight into both intermolecular and intramolecular forces. Interactions between hydrophobic regions of the support-adsorbed protein and the tip can be strong enough to “unzip” molecules (Figure 17.7.8). In cases where the tip need not to be functionalized, comparative topography image acquisition is possible (Müller et al., 1999; Scheuring et al., 2002a), allowing force measurements to be correlated with topographical structure. In all cases, many force-distance curves (Figure 17.7.2) must be acquired and carefully analyzed. Instrumental effects must also be taken into account, e.g., rupture forces (correlating with thermodynamic “off-rates” of a ligand-receptor complex) measured between bonded
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A
B
C
Figure 17.7.7 Time-lapse imaging: watching proteins at work. (A) Movement of a single DNA molecule through an RNA polymerase complex (adapted from Kasas et al., 1997). (B) Structural change induced in the nuclear basket of individual native nuclear pore complexes (NPCs) with addition of calcium ions. The same specimen area was imaged in two distinct conformational states. Three corresponding NPCs are marked by arrowheads. For a more quantitative comparison of the “closed” and “open” states, 30 corresponding NPCs were aligned and averaged, and their average radial height profiles computed. Scale bar, 100 nm (adapted from Stoffler et al., 1999), (C) Growth of an individual Aβ amyloid fibril adhering to a mica support (Goldsbury et al., 2001).
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A
B
C
100pN
10nm
Figure 17.7.8 Image before a) and after b) unzipping of a full hexameric core of the Corynebacterium glutamicum S-layer. c) Force-distance curve corresponding to the change of the surface structure as shown in a) and b). The hexameric core is unzipped in a sequence of three dimer-like rupture events (Scheuring et al., 2002).
molecular pairs depend on the speed of withdrawal of the AFM tip (Fritz et al., 1998).
IMAGE INTERPRETATION: TROUBLESHOOTING AND INSTRUMENTAL EFFECTS
Introduction to Atomic Force Microscopy (AFM) in Biology
AFM is now an established technique for imaging of biological materials. The instrumentation has greatly improved, so that high clarity, high-resolution images can be achieved (Fotiadis et al., 1998; Müller et al., 1999c; Scheuring et al., 2001b). Instrumental effects that were previously poorly understood are beginning to unravel, leading to better control of the imaging process. As outlined in the previous sections, major factors that can affect the quality of the image include applied force, ionic strength and composition of the scanning buffer (see High Resolution Imaging), and the quality of the support to which the sample is attached (see Preparation of the Support and Specimen
Attachment). The shape and characteristics of the scanning tip used for imaging (see The Tip and Cantilever) also plays a major role in the resolution of AFM images. Furthermore, as AFM is a surface-imaging technique, it is important to fully characterize and understand the features of the supporting surface to which the sample is attached.
Tip Shape The shape of the scanning tip affects the shape of the features in the image. The most common tip effect is convolution of the imaging tip’s shape with the actual shape of the object. In such convolutions, the height of the object (z-dimension) is not affected by tip geometry, but the width (x-, y-dimension) becomes broadened due to convolution with the radius of curvature of the interacting tip. Broadening of the width measurement becomes predominant with specimens isolated on the support, e.g.,
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filamentous structures or single antibody molecules (Allen et al., 1992; Willemsen et al., 1998). The broadening effect is less of an issue with closely packed objects, e.g., crystalline protein arrays or filament rafts. Furthermore, the interaction between the surface and the tip sometimes involves more than one region of the tip, creating an effect of “multiple tips” scanning the surface simultaneously. This effect can be dramatic and is easily recognized as a clear double picture, or it can be more subtle, requiring a careful eye. Generally, if isolated objects on the surface appear to all have the same shape and orientation, the tip is probably being imaged rather than the sample. Also, if cross sections measured on different parts of the specimen (e.g., along a filamentous structure) exhibit almost identical contours, this is strongly indicative that the shape of the crosssection is dominated by, and represents the shape of the tip rather than sub-structure within the specimen. The multiple-tip effect may be eliminated in some cases by slightly changing the angle of the cantilever relative to the scanned support such that a slightly different region of the end of the tip contacts the sample. Particular care must be taken when investigating helical structures by AFM. Markiewicz ( 19 88; http://www.weizmann.ac.il/surflab/ peter/phfpaper/index.html) demonstrated apparent existence of both right- and left-handed filaments in preparations of Alzheimer paired helical filaments (PHFs). In computational simulations using different shaped tips, they showed that this directionality could be an artifact due to a slightly oblong shaped tip. In the simulations, apparent beaded structures often appeared, masking the filament substructure, and in some instances, further distortion occurred resulting in reversal of the apparent helical twist direction. It is recommended, wherever possible, to compare the structural features seen by AFM with what is seen using conventional electron microscopy (i.e., after negative staining or unidirectional metal shadowing the specimen). In some cases, changes in tip shape may be due to contamination resulting from organic molecules adsorbing to the very end of the stylus. Scanning with such tips result in smears at the left edge of the “trace-image” and at the right edge of the “retrace-image.” The more material that is adsorbed on the end of the tip, the stronger the effect. In severe cases, a completely smeared image occurs without any resolvable surface features. To try to clear the tip of contamination, one can scan a sample-free
support region with increased force for a few seconds, or let the tip scan in a non-contact region above the surface for a few minutes. Normally, the best solution is to exchange the tip.
Specimen Support Artifacts Image artifacts can occasionally be caused by the surface of the support to which the sample is adsorbed. It is therefore very important to characterize this surface under identical scanning conditions with and without adsorption of the specimen. Mica rarely produces image artifacts. If it is carefully cleaved with tape before every experiment, the only effects that might be observed are mica-layer edges, which are usually very sharp, or large hill-like protrusions, probably originating from air inclusions between the mica layers. On the other hand, HOPG is more prone to surface effects when imaged under buffer solution, e.g., parallel stripes in various directions (usually oriented in line with the 60°-angles of the HOPG carbon lattice) are sometimes observed with a lateral spacing of ∼5 to 10 nm and a height of 0.5 to 1 nm. These features may pose a problem if they closely resemble features of the specimen.
SUMMARY The atomic-force microscope provides image resolution comparable to the electron microscope, while additionally allowing specimens to be viewed “alive” in a dynamic state within an aqueous environment (Bustamante et al., 1997; Engel and Müller, 2000). To achieve this, a sharp tip at the end of a flexible cantilever scans and senses the surface contours of the sample. Over the past decade, AFM has been refined to the point of achieving sub-nanometer resolution on densely packed proteins, e.g., membrane proteins. Importantly, these samples are completely hydrated and can be imaged in a physiologically relevant buffer in their native state without the requirement for further fixation or contrasting. Other samples range from single molecules to live cells, and although the resolution is lower (on the order of ∼3 nm for single proteins, complexes, and filaments, and ∼10 nm for cells), the specimen remains in an aqueous environment, so repetitive imaging is possible allowing temporal resolution, i.e., regarding conformational changes, polymerization kinetics, and dynamic movement. In most cases, the specimen must be firmly attached to a solid support. In dynamic-mode AFM, due to smaller lateral forces, it is possible to measure
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samples that are less well attached to the support, hence, dynamic movement of the specimen on the support may be visualized. The fine sensitivity of the scanning tip to contact forces has also been exploited to measure both intermolecular and intramolecular force interactions in the pico-Newton (pN) range as well as mechanical properties (elastic modulus) of the specimen (Radmacher et al., 1997; Rief et al., 1997a,b, 2000; Allison et al., 2002). Intermolecular forces include the binding and rupturing of receptor-ligand complexes, finding applications in studies on cell adhesion molecules and antibody-antigen interactions. Protein unfolding and membrane protein “unzipping” experiments monitoring intramolecular forces have opened doors for gaining new insight into protein folding dynamics at the level of single molecules. New emerging applications of AFM include using the tip to “write” and later “read” nano-arrays of proteins (Lee et al., 2002). These arrays have been used to screen mixtures of potential ligands for molecular recognition of the protein of interest, perhaps pointing towards applications in the fast growing field of proteomics. Other applications use AFM cantilevers as free-standing biosensors that bend or change their resonance frequency when molecules bind to their surface (Fritz et al., 2000; Ilic et al., 2000; Wu et al., 2001). However, the major limitations of conventional AFM remain the scanning time and the requirement for the sample to be adhered to a supporting surface. Future development of faster scanning microscopes/tips will enable higher temporal resolution to be gained both in imaging and in force measurements.
Acknowledgements The authors thank Dr. Jürgen Fritz for a critical reading and editing of the manuscript. Drs. Claire Goldsbury and Simon Scheuring were postdoctoral fellows working on atomicforce microscopy in the laboratories of Prof. Ueli Aebi and Prof. Andreas Engel at the M.E. Müller Institute for Structural Biology, Biozentrum, University of Basel, Switzerland. We thank Andreas Engel, Ueli Aebi, Daniel Müller, and Paul Hansma for providing contributions to the figures.
LITERATURE CITED Introduction to Atomic Force Microscopy (AFM) in Biology
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Engel, A., Gaub, H.E., and Müller, D.J. 1999. Atomic force microscopy: A forceful way with single molecules. Curr. Biol. 9:R133-R136. Fisher, T.E., Marszalek, P.E., Oberhauser, A.F., Carrion-Vazquez, M., and Fernandez, J.M. 1999a. The micro-mechanics of single molecules studied with atomic force microscopy. J. Physiol. (Lond). 520:5-14. Fisher, T.E., Oberhauser, A.F., Carrion-Vazquez, M., Marszalek, P.E., and Fernandez, J.M. 1999b. The study of protein mechanics with the atomic force microscope. Trends Biochem. Sci. 24:379-384. Fisher, T.E., Marszalek, P.E, and Fernandez, J.M. 2000. Stretching single molecules into novel conformations using the atomic force microscope. Nat. Struct. Biol. 7:719-724. Florin, E.-L., Moy, V.T., and Gaub, H.E. 1994. Adhesion forces between individual ligand-receptor pairs. Science 264:415-417. Fotiadis, D., Müller, D.J., Tsiotis, G., Hasler, L., Tittmann, P., Mini, T., Jeno, P., Gross, H., and Engel, A. 1998. Surface analysis of the photosystem I complex by electron and atomic force microscopy. J. Mol. Biol. 283:83-94. Fotiadis, D., Hasler, L., Müller, D.J., Stahlberg, H., Kistler, J., and Engel, A. 2000. Surface tongueand-groove contours on lens MIP facilitate cellto-cell adherence. J. Mol. Biol. 300:779-789. Fritz, J., Katopodis, A.G., Kolbinger, F., and Anselmetti, D. 1998. Force-mediated kinetics of single P-selectin/ligand complexes observed by atomic force microscopy. Proc. Natl. Acad. Sci. U.S.A. 95:12283-12288. Fritz, J., Baller, M.K., Lang, H.P., Rothuizen, H., Vettiger, P., Meyer, E., Güntherodt, H.-J., Gerber, C., and Gimzewski, J.K. 2000. Translating biomolecular recognition into nanomechanics. Science 288:316-318. Goldsbury, C., Aebi, U., and Frey, P. 2001. Visualizing the growth of Alzheimer’s Åβ amyloid-like fibrils. Trends Mol. Med. 7:582. Goldsbury, C., Kistler, J. Aebi, U., Arvinte, T., and Cooper, G.J. 1999. Watching amyloid fibrils grow by time-lapse atomic force microscopy. J. Mol. Biol. 285:33-39. Hansma, H.G., Kim, K.J., Laney, D.E., Garcia, R.A., Argaman, M., Allen, M.J., and Parsons, S.M. 1997. Properties of biomolecules measured from atomic force microscope images: A review. J. Struct. Biol. 119:99-108. Heinz, W.F. and Hoh, J.H. 1999. Spatially resolved force spectroscopy of biological surfaces using the atomic force microscope. Trends Biotechnol. 17:143-150. Hofmann, U.G., Rotsch, C., Parak, W.J., and Radmacher, M. 1997. Investigating the cytoskeleton of chicken cardiocytes with the atomic force microscope. J. Struct. Biol. 119:84-91. Hoh, J.H. and Schoenenberger, C.A. 1994. Surface morphology and mechanical properties of MDCK monolayers by atomic force microscopy. J. Cell Sci. 107:1105-1114.
Hörber, J.K., Mosbacher, J., Haberle, W., Ruppersberg, J.P. and Sakmann, B. 1995. A look at membrane patches with a scanning force microscope. Biophys. J. 68:1687-1693. Hutter, J.L. and Bechhoefer, J. 1993. Calibration of atomic force microscope tips. Rev. Sci. Instr. 64:1868-1873 Ilic, B., Czaplewski, D., Craighead, H.G., Neuzil, P., Campagnolo, C., and Batt, C. 2000. Mechanical resonant immunospecific biological detector. Appl. Phys. Lett. 77:450-452. Karrasch, S., Dolder, M., Schabert, F., Ramsden, J., and Engel, A. 1993. Covalent binding of biological samples to solid supports for scanning probe microscopy in buffer solution. Biophys. J. 65:2437-2446. Karrasch, S., Hegerl, R., Hoh, J., Baumeister, W., and Engel, A. 1994. Atomic force microscopy produces faithful high-resolution images of protein surfaces in an aqueous environment. Proc. Natl. Acad. Sci. U.S.A. 91:836-838. Kasas, S., Thomson, N.H., Smith, B.L., Hansma, H.G., Zhu, X., Guthold, M., Bustamante, C., Kool, E.T., Kashlev, M., and Hansma P.K. 1997. Escherichia coli RNA polymerase activity observed using atomic force microscopy. Biochemistry 36:461-468. Lal, R. and John, S.A. 1994. Biological applications of atomic force microscopy. Am. J. Physiol. 266:C1-C21. Lee, K.-B., Park, S.-J., Mirkin, C., Smith, J., and Mrksich, M. 2002. Protein nanoarrays generated by dip-pen nanolithography. Science 295:17021705 Levadny, V.G., Belaya, M.L., Pink, D.A., and Jericho, M.H. 1996. Theory of electrostatic effects in soft biological interfaces using atomic force microscopy. Biophys. J. 70:1745-1752. Markiewicz, P. 1988. Orientational dependency of AFM images revealed by Alzheimer paired helical filaments. Ph.D. thesis. Department of Chemistry, University of Toronto. McPherson, A., Malkin, A.J., and Kuznetsov, Y.G. 2000. Atomic force microscopy in the study of macromolecular crystal growth. Annu. Rev. Biophys. Biomol. Struct. 29:361-410. McPherson, A., Malkin, A.J., Kuznetsov, Y.G., and Plomp, M. 2001. Atomic force microscopy applications in macromolecular crystallography. Acta Crystallogr. D. Biol. Crystallogr. 57:10531060. Mitsui, K., Hara, M., and Ikai, A. 1996. Mechanical unfolding of alpha(2)-macroglobulin molecules with atomic force microscope. FEBS Lett. 385:29-33. Möller, C., Allen, M., Elings, V., Engel, A., and Müller, D.J. 1999. Tapping-mode atomic force microscopy produces faithful high-resolution images of protein surfaces. Biophys. J. 77:11501158.
Structural Biology
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Mou, J.X., Yang, J., and Shao, Z.F. 1995. Atomic force microscopy of cholera toxin B-oligomers bound to bilayers of biologically relevant lipids. J. Mol. Biol. 248:507-512.
Radmacher, M., Fritz, M., Hansma, H.G., and Hansma, P.K. 1994. Direct observation of enzyme activity with the atomic force microscopy. Science 265:1577-1579.
Mou, J., Sheng, S., Ho, R., and Shao, Z. 1996a. Chaperonins GroEL and GroES: Views from atomic force microscopy. Biophys. J. 71:22132221.
Radmacher, M., Fritz, M. and Hansma, P.K. 1995. Imaging soft samples with the atomic force microscope: Gelatin in water and propanol. Biophys. J. 69:264-270.
Mou, J., Czajkowsky, D.M., Sheng, S., Ho, R., and Shao, Z. 1996b. High resolution surface structure of E. coli GroES oligomer by atomic force microscopy. FEBS Lett. 381:161-164.
Reviakine, I., Bergsma-Schutter, W., and Brisson, A. 1998. Growth of protein 2-D con supported planar lipid bilayers imaged in situ by AFM. J. Struct. Biol. 121:356-361.
Müller, D.J. and Engel, A. 1997. The height of biomolecules measured with the atomic force microscope depends on electrostatic interactions. Biophys. J. 73:1633-1644.
Rief, M., Oesterhelt, F., Heymann, B., and Gaub, H.E. 1997a. Single molecule force spectroscopy on polysaccharides by AFM. Science 275:12951298.
Müller, D.J., Engel, A., and Amrein, M. 1997a. Preparation techniques for the observation of native biological systems with the atomic force microscope. Biosens. Bioelectron. 12:867-877.
Rief, M., Gautel, M., Oesterhelt, F., Fernandez, J.M., and Gaub, H.E. 1997b. Reversible unfolding of individual titin immunoglobulin domains by AFM. Science 276:1109-1112.
Müller, D.J., Engel, A., Carrascosa, J., and Veléz, M. 1997b. The bacteriophage ø29 head-tail connector imaged at high resolution with atomic force microscopy in buffer solution. EMBO J. 16:101107.
Rief, M., Gautel, M., and Gaub, H.E. 2000. Unfolding forces of titin and fibronectin domains directly measured by AFM. Adv. Exp. Med. Biol. 481:129-136.
Müller, D.J., Amrein, M., and Engel, A. 1997c. Adsorption of biological molecules to a solid support for scanning probe microscopy. J. Struct. Biol. 119:172-188. Müller, D.J., Fotiadis, D., and Engel, A. 1998. Mapping flexible protein domains at subnanometer resolution with the atomic force microscope. FEBS Lett. 430:105-111.
Introduction to Atomic Force Microscopy (AFM) in Biology
Rinia, H.A., Kik, R.A., Demel, R.A., Snel, M.M., Killian, J.A., van Der Eerden, J.P., and de Kruijff, B. 2000. Visualization of highly ordered striated domains induced by transmembrane peptides in supported phosphatidylcholine bilayers. Biochemistry 39:5852-5858. Schabert, F.A. and Engel, A. 1994. Reproducible acquisition of Escherichia coli porin surface topographs by atomic force microscopy. Biophys. J. 67:2394-2403.
Müller, D.J., Baumeister, W., and Engel, A. 1999a. Controlled unzipping of a bacterial surface layer with atomic force microscopy. Proc. Natl. Acad. Sci. U.S.A. 96:13170-13174.
Schabert, F.A., Henn, C., and Engel, A. 1995. Native Escherichia coli OmpF porin surfaces probed by atomic force microscopy. Science 268:92-94.
Müller, D.J., Sass, H.-J., Müller, S., Büldt, G., and Engel, A. 1999b. Surface structures of native bacteriorhodopsin depend on the molecular packing arrangement in the membrane. J. Mol. Biol. 285:1903-1909.
Scheuring, S., Ringler, P., Borgina, M., Stahlberg, H., Müller, D.J., Agre, P., and Engel, A. 1999a. High resolution topographs of the Escherichia coli waterchannel aquaporin Z. EMBO J. 18:4981-4987.
Müller, D.J., Fotiadis, D., Scheuring, S., Müller, S.A., and Engel, A. 1999c. Electrostatically balanced subnanometer imaging of biological specimens by atomic force microscopy. Biophys. J. 76:1101-1111.
Scheuring, S., Müller, D.J., Ringler, P., Heymann, J.B., and Engel, A. 1999b. Imaging streptavidin 2D crystals on biotinylated lipid monolayers at high resolution with the atomic force microscopy. J. Microsc. 193:28-35.
Neumeister, J.M. and Ducker, W.A. 1994. Lateral, normal and longitudinal spring constants of atomic force microscopy cantilevers. Rev. Sci. Instr. 65:2527-2531.
Scheuring, S., Fotiadis, D., Möller, C., Müller, S.A., Engel, A., and Müller, D.J. 2001a. Single proteins observed by atomic force microscopy. Single Mol. 2:59-67.
Oesterhelt, F., Oesterhelt, D., Pfeiffer, M., Engel, A., Gaub, H.E., and Müller, D.J. 2000. Unfolding pathways of individual bacteriorhodopsins. Science 288:143-146.
Scheuring, S., Freiss-Husson, F., Engel, A., Rigaud, J., and Ranck, J. 2001b. High resolution topographs of the Rubrivivax gelatinosus light-harvesting complex 2. EMBO J. 20:3029-3035.
Raab, A., Han, W., Badt, D., Smith-Gill, S., Lindsay, S., Schindler, H., and Hinterdorfer, P. 1999. Antibody recognition imaging by atomic force microscopy. Nature Biotech. 17:902-905.
Scheuring, S., Stahlberg, H., Chami, M., Houssin, C., Rigaud, J., and Engel, A. 2002a. Charting and unzipping the surface-layer of Corynebacterium glutamicum with the atomic force microscope. Mol. Microbiol. 44:675-684.
Radmacher, M. 1997. Measuring the elastic properties of biological samples with the atomic force microscope. IEEE Medicine and Engineering Biology 16:47-57.
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Current Protocols in Protein Science
Scheuring, S., Müller, D.J., Stahlberg, H., Engel, H.-A., and Engel, A. 2002b. Sampling the conformational space of membrane protein surfaces with the AFM. Eur. Biophys. J. 31:172-178. Schoenenberger, C.A. and Hoh, J.H. 1994. Slow cellular dynamics in MDCK and R5 cells monitored by time-lapse atomic force microscopy. Biophys. J. 67:929-936. Seelert, H., Poetsch, A., Dencher, N.A., Engel, A., Stahlberg, H., and Müller, D.J. 2000. Proton powered turbine of a plant motor. Nature 405:418-419. Shi, D., Somlyo, A.V., Somlyo, A.P., and Shao, Z. 2001. Visualizing filamentous actin on lipid bilayers by atomic force microscopy in solution. J. Microsc. 201:377-382. Stahlberg, H., Müller, D.J., Suda, K., Fotiadis, D., Engel, A., Matthey, U., Meier, T., and Dimroth, P. 2001. Bacterial ATP synthase has an undacemeric rotor. EMBO Reports 2:229-235. Stolz, M., Stoffler, D., Aebi, U., and Goldsbury, C. 2000. Monitoring biomolecular interactions by time-lapse atomic force microscopy. J. Struct. Biol. 131:171-180. Thomson, N.H., Fritz, M., Radmacher, M., Cleveland, J.P., Schmidt, C.F., and Hansma, P.K. 1996. Protein tracking and detection of protein motion using atomic force microscopy. Biophys. J. 70:2421-2431.
Wagner P. 1998. Immobilization strategies for biological scanning probe microscopy. FEBS Lett. 430:112-115. Willemsen, O.H., Snel, M.M., van der Werf, K.O., de Grooth, B.G., Greve, J., Hinterdorfer, P., Gruber, H.J., Schindler, H., van Kooyk, Y., and Figdor, C.G. 1998. Simultaneous height and adhesion imaging of antibody-antigen interactions by atomic force microscopy. Biophys. J. 75:2220-2228. Wu, G., Datar, R.H., Hansen, K.M., Thundat, T., Cote, R.J., and Majumdar, A. 2001. Bioassay of prostate-specific antigen (PSA) using microcantilevers. Nature Biotech. 19:856-860. Xu, S.H. and Arnsdorf, M.F. 1995. Electrostatic force microscope for probing surface charges in aqueous solutions. Proc. Natl. Acad. Sci. U.S.A. 92:10384-10388.
INTERNET RESOURCES http://www.di.com/SPM%20Forum/SPMForum MailingList.html Digital Instruments hosts a free internet mailing list for troubleshooting, discussing, and exchanging technical information, views, issues, and applications of AFM.
Viani, M.B., Schäfer, T.E., Chand, A., Rief, M., Gaub, H.E., and Hansma, P.K., 1999. Small cantilevers for force spectroscopy of single molecules. J. Appl. Phys. 86:2258-2262.
Contributed by Claire Goldsbury Cytoskeleton Group Max Planck Unit for Structural Molecular Biology Hamburg, Germany
Viani, M.B., Pietrasanta, L.I., Thompson, J.B., Chand, A., Gebeshuber, I.C., Kindt, J.H., Richter, M., Hansma, H.G., and Hansma, P.K. 2000. Probing protein-protein interactions in real time. Nat. Struct. Biol. 7:644-647.
Simon Scheuring Institut Curie Paris, France
Structural Biology
17.7.17 Current Protocols in Protein Science
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Raman Spectroscopy of Proteins
UNIT 17.8
Raman spectroscopy, like infrared (IR) spectroscopy, is a method for probing structures and interactions of molecules by measuring the energies (or frequencies) of molecular vibrations. Although the two methods differ fundamentally in the mechanism of interaction between radiation and matter, one obtains, in both cases, a vibrational spectrum consisting of a number of discrete bands, the frequencies and intensities of which are determined by the nuclear masses in motion, the equilibrium molecular geometry, and the molecular force field. An important advantage of Raman over IR spectroscopy for applications to proteins and their complexes is the virtual transparency of liquid water (both H2O and D2O) in the Raman effect. This greatly simplifies the analysis of spectra of aqueous solutions of biomolecules and also facilitates the investigation of hydrogenisotope-exchange phenomena. Other notable advantages of Raman spectroscopy are ease of implementation for samples in different physical states (solutions, suspensions, gels, precipitates, fibers, single crystals, amorphous solids, etc.), applicability to large supramolecular assemblies, a spectroscopic time scale (∼10−14 sec) that is fast in comparison to both biomolecular structure transformations and protium/deuterium (H/D) exchanges, non-destructiveness of data collection protocols, relatively low mass (<1 mg) and volume (∼1 µl) requirements for most samples, and freedom from the need for chemical labels or probes. The changes in molecular geometry or conformation that characterize many molecular biological phenomena can produce relatively large changes in Raman band positions, which are usually referred to as frequency shifts. Such shifts in the Raman band frequencies are the basis for applications of the technique to analyze protein secondary structures, determine side chain conformations, and detect intramolecular interactions. Because the molecular geometry and force field are also sensitive to interactions between molecules, the Raman method can be effectively applied to probe protein intermolecular interactions, including the formation of specific complexes and large assemblies. Currently, there exists a large database of Raman spectra of model compounds for which reliable band assignments and detailed spectra-structure correlations have been developed. Many of these correlations are directly transferable to proteins and their complexes, thus facilitating structural interpretations of protein Raman spectra. Raman spectroscopy continues to gain increasing use as a method for probing protein structure, dynamics, assembly, and recognition. This unit surveys the general nature, underlying theoretical basis and experimental requirements of protein Raman spectroscopy. Specific molecular structural properties that can be elucidated from protein Raman spectra are described and recent applications are discussed to illustrate the scope of the method. Emphasis is given to applications from the authors’ laboratory, which probe protein conformations, intra- and inter-molecular protein interactions, subunit folding and assembly pathways, and protein-nucleic acid recognition. These examples employ conventional (off-resonance) Raman spectroscopy, resonance Raman spectroscopy and polarized Raman microspectroscopy. This unit does not include discussions of more specialized Raman techniques, such as Raman imaging, Raman optical activity (ROA), and surface-enhanced Raman spectroscopy (SERS), which may also find use in protein science. More comprehensive treatments of Raman, resonance-Raman and related spectroscopic methods in life science research have been given elsewhere (Carey, 1982; Barron et al., 2000; Chalmers and Griffiths, 2002).
Structural Biology Contributed by James M. Benevides, Stacy A. Overman, and George J. Thomas, Jr. Current Protocols in Protein Science (2003) 17.8.1-17.8.35
17.8.1
Copyright © 2003 by John Wiley & Sons, Inc.
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NATURE OF THE DATA The Raman effect is an inelastic light scattering process in which quanta of energy are transferred from a beam of intense monochromatic radiation (laser photons) to the target molecule with which the photons interact. The transferred energy is typically a small fraction of the total energy of the incident photon. The quanta are gained by the molecule in the form of discrete vibrational energies, which constitute the Raman frequencies. These frequencies, along with their relative scattering probabilities (Raman intensities) and tensor characteristics (Raman polarizations), constitute the Raman spectrum of the molecule. The spectrum is defined by the intramolecular geometry and intra- and intermolecular force fields, therefore, representing a unique signature of the molecular structure and its environment. The Raman spectrum of a protein is obtained by using a highly sensitive spectrophotometer to measure the energies and relative numbers of photons scattered by the target protein. Scattered photons exhibiting frequencies less than that of the impingent laser photons are a measure of the vibrational quanta transferred to the protein. The intensity of a Raman protein band is a function of the efficiency of the energy exchange between the incident laser photon and the vibrational mode in question, as well as of the number of subgroups in the protein that exhibit that particular vibrational mode. With respect to the former, the following discussion is helpful in understanding the mechanism that determines the intrinsic Raman scattering intensity. When monochromatic light of frequency (v0) irradiates a molecule, the oscillating electric field (E) promotes a small discrepancy between the centers of negative (electronic) and positive (nuclear) charge, and an oscillating dipole moment is introduced into the molecule. This phenomenon is referred to as polarization. To a good approximation, the induced dipole moment is proportional to the incident electric field strength E: P = αE Equation 17.8.1
where the proportionality factor, α, is the molecular electric polarizability tensor. Each component of P (Pi, where i = x, y, z) is related to the three components of the electric field strength by Equation 17.8.2: Pi = ∑ αij E j Equation 17.8.2
where the sum is over i and j (= x, y, z). The coefficients αij are the elements of the polarizability tensor and the subscripts i and j represent, respectively, the components of P and E to which it relates. Since both α and E are time-dependent (α is dependent on the internuclear separations that vary with the frequency vv of molecular vibration, while E oscillates at frequency v0), it can be shown that P has terms that depend upon v0, v0 + vv and v0 − vv as Equation 17.8.3:
( )
1 Pi (t ) = ∑ αij 0 E × cos 2π v0 t + αij′ qE × cos 2π (v0 ± vv ) t 2 Raman Spectroscopy of Proteins
Equation 17.8.3
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Current Protocols in Protein Science
where αij0 is the component ij of the molecular polarizability at the equilibrium configuration, αij′ is the first derivative of the polarizability component taken with respect to the vibrational coordinate q, and |E| is the amplitude of E. The first term on the right of Equation 17.8.3 is a component of Pi oscillating with frequency ν0 that, according to classic electromagnetic theory, represents energy radiated (scattered) with the same frequency as the incident light. This is called Rayleigh scattering. The second term on the right of Equation 17.8.3 contains components of Pi oscillating at frequencies ν0 ± νv, i.e., representing radiation scattered with frequencies that are either a vibrational quantum larger (anti-stokes Raman scattering) or smaller (Stokes Raman scattering) than the incident frequency. Although both anti-Stokes and Stokes Raman scattering can be observed experimentally, the latter is favored by the Boltzmann distribution law and in practice is easier to detect. Equation 17.8.3 indicates that the Raman effect is dependent upon a change in polarizability with the vibration, i.e., αij′ ≠ 0. If the polarizability does not change during a molecular vibration, there is no Raman scattering generated and hence the vibration is said to be Raman inactive. On the basis of the above considerations, the following generalizations can be made with respect to Raman intensities associated with vibrations of protein subgroups. Stretching vibrations involving peptide C=O and C–N bonds are expected to yield high Raman intensities because of the large polarizability changes associated with their stretching vibrations. In-plane vibrations of the rings of aromatic side chains (Trp, Tyr, Phe) are also expected to produce Raman bands of high intensity. Stretching vibrations of C–C, C–N, and C–O bonds also produce intense Raman bands, especially if they involve the concerted symmetrical displacements of side chain skeletons. Conversely, out-of-plane ring deformations and bending motions of exocyclic substituents are usually weak in Raman spectra of proteins. The Raman bands associated with the bending and stretching modes of individual hydrogenic substituents (e.g., C–H, N–H, and O–H) are generally weak, but their collective spectral intensities, which result from large numbers of such groupings in a protein, can be high. Vibrations that involve the displacement of heavy atoms such as sulfur in C–S stretching modes of Met and Cys, and S–S stretching of cystine, S–H stretching of cysteine, and Zn–S stretching in zinc metalloproteins are expected to be relatively intense. Additional factors govern the intensities of bands appearing in the ultraviolet-resonance Raman (UVRR) spectra of proteins. Accordingly, spectral intensities may differ greatly between conventional (off-resonance) Raman and UVRR spectra of the same protein molecule. Further discussion of UVRR spectra of proteins is given by Austin and co-authors (1993). Polarization of the Raman scattered light is a feature that can be exploited to provide additional useful information on the nature of a given normal mode of vibration. Although the incident laser beam is plane polarized, Equation 17.8.2 indicates that the Raman scattered light is not necessarily polarized in the same plane. The change of the plane of polarization is defined quantitatively by the depolarization ratio ρ (≡ I⊥/I||), which is the ratio of intensities of light scattered along directions perpendicular (I⊥) and parallel (I||) to the direction of polarization of the incident light. In isotropic ensembles of molecules, measurements of ρ can help distinguish normal modes of vibration that are totally or locally symmetric from those that are non-totally symmetric. In anisotropic ensembles of molecules, such as single crystals or oriented fibers, detailed analysis of the Raman polarizations can yield information on the orientations of molecules or their subgroups (Tsuboi et al., 1996b; Tsuboi and Thomas, 1997; Tsuboi et al., 2001a). Further discussion is given below (see Data Analysis).
Structural Biology
17.8.3 Current Protocols in Protein Science
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EXPERIMENTAL DESIGN Instrumentation Instrumentation for Raman spectroscopy has changed dramatically in recent years, reflecting improved design of laser light sources, optical filters and photon detectors. Powerful and stable continuous-wave lasers are now commercially available for Raman excitations extending from near infrared to ultraviolet. Diode-pumped solid-state lasers are generally replacing their ion laser equivalents. Such lasers have smaller dimensions and require neither a high line voltage nor a circulating coolant. Long wavelength Raman excitations in the red and near-infrared offer the significant advantage over shorter wavelength visible excitations of greater freedom from potentially interfering fluorescence phenomena. Recently, several solid-state lasers operating between 780 and 830 nm have become available commercially. For ultraviolet (UV) wavelengths, continuous-wave laser technology has improved to the point that collection of UV resonance Raman (UVRR) spectra of proteins and nucleoprotein complexes, including viruses, is now feasible without significant photodecomposition of the sample. Advent of the holographic notch filter to more effectively reject elastically scattered (Rayleigh) light, introduction of the volume-phase holographic transmissive grating to maximize photon throughput, and implementation of efficient charge-coupled-device (CCD) detectors have also contributed to dramatic increases in spectrophotometric sensitivity and selectivity. In addition, transmissive optics provide an appreciably more stable Raman platform that preserves spectrometer calibration and alignment (Slater et al., 2001). This is of particular importance in the application of Raman digital difference spectroscopy, a very useful and powerful approach for characterizing conformational changes in proteins and other biomolecules (Thomas, 1986). State-of-the-art Raman spectrometer systems have been described for biological applications of UVRR and near-infrared Raman (NIR) spectroscopies (Asher et al., 1993; Hashimoto et al., 1993; Russell et al., 1995; Dong et al., 1998). Sample Requirements and Handling Although protein concentrations in the range of 5 to 40 µg/µl are generally employed for Raman spectroscopy, concentrations as low as 1 µg/µl have been reported (Dong et al., 1998). Sample volume requirements are also modest (∼1 µl). For most purposes, the sample is sealed within a glass capillary. When NIR excitation wavelengths >752 nm are used to excite the Raman spectrum, quartz capillaries are preferred to circumvent fluorescence interference from glass. Quartz capillaries are also required for UVRR spectroscopy. Salts, buffers, and chelating agents appropriate for specific Raman spectroscopy applications to proteins are described in the general literature. Monatomic cationic and anionic species should be employed whenever possible, because polyatomic species generate Raman bands that may interfere with those of the protein. DATA ANALYSIS
Raman Spectroscopy of Proteins
Definitive band assignments are required for the successful application of Raman spectroscopy in protein analysis. These can be achieved from a combination of the following experimental and theoretical approaches. (1) Determine vibrational frequency shifts that accompany stable isotope substitutions, the most useful generally being 1H → 2H(D) (H/D) exchange. (2) Conduct detailed analyses of the Raman spectra of appropriate model compounds (smaller, more symmetrical molecules that are structurally related to the protein constituents). (3) Characterize effects of pH, temperature, and other environmental factors on the spectra. (4) Use ab initio molecular orbital calculation of intramolecular force fields and normal coordinate calculations based upon them to assist empirical assignments. (5) Measure depolarization ratios of the Raman bands of isotropic solutions. (6) Measure polarized Raman spectra of oriented samples. (7) Examine Raman
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excitation profiles for laser wavelengths in the visible and ultraviolet regions. Because selection rules governing UVRR and off-resonance transitions differ fundamentally from one another, the intensities of Raman bands observed in the two types of spectra can be quite different (Nishimura et al., 1978; Tsuboi et al., 1987; Austin et al., 1993; Miura and Thomas, 1995a,b). As an example Figure 17.8.1 compares Raman spectra of the filamentous virus fd (600 to 1800 cm−1 region) obtained using off-resonance (514.5 nm) and UV-resonance (257 and 229 nm) excitations. Thus, while the 514.5-nm spectrum exhibits a rich pattern of Raman bands from all viral components, the 229-nm excitation produces Raman signatures only from the tryptophan (Trp 26) and tyrosine (Tyr 21 and Tyr 24) residues of the coat protein subunit, and the 257-nm excitation produces a spectrum dominated by Raman signatures of the DNA bases of the packaged single-stranded (ss) DNA genome. Further discussion of UVRR excitation profiles is given elsewhere (Hudson and Mayne, 1987; Tsuboi et al., 1987). For both proteins and nucleic acids, many group frequencies have been identified and their assignments supported by normal coordinate calculations (Tsuboi et al., 1987; Thomas and Wang, 1988; Austin et al., 1993; Thomas and Tsuboi, 1993; Peticolas, 1995; Miura and Thomas, 1995a,b). For example, Figure 17.8.2 illustrates many such assignments for the Raman spectrum of a large nucleoprotein complex, the icosahedral doublestranded (ds) DNA bacteriophage P22. The legend identifies representative Raman bands that are diagnostic group frequencies of the viral coat protein subunits and packaged dsDNA genome and indicates details of sample conditions. Digitally computed Raman difference spectra can facilitate visualization of changes in Raman protein bands that accompany changes in sample temperature, molecular environment, and interactions with ligands. Raman digital difference spectroscopy was developed independently in several laboratories, and many interesting applications of Raman difference spectroscopy in studies of biomolecular structure have been reviewed (Thomas, 1986; Callender and Deng, 1994). The technique is particularly powerful when combined with residue-specific isotope substitutions (Aubrey and Thomas, 1991; Overman and Thomas, 1999) and site-specific mutagenesis (Overman et al., 1994; Tsuboi et al., 2001a). Information on three-dimensional structure may be obtained from polarized Raman spectra of proteins if two key conditions are met. First, the target proteins must be of uniform and known orientation with respect to the electric vector of the exciting radiation, and second, Raman tensors for the diagnostic vibrational bands must be known. The first criterion is satisfied for appropriately oriented single crystals or fibers; the second requires tedious polarized Raman measurements on suitable model structures (Benevides et al., 1993; Thomas et al., 1995; Tsuboi et al., 1996a; Benevides et al., 1997). Once both criteria have been met, the oriented protein—e.g., in a fibrous specimen—is placed on the sample holder of a Raman microscope with its internal axis system (abc) positioned such that two axes, say c (the fiber axis) and b (perpendicular to c), are in the horizontal plane (Fig. 17.8.3). The exciting radiation with electric vector polarized along c is directed onto the fiber through an objective. The Raman scattered radiation is collected with the same objective and directed through a polarizer to the monochromator. The polarizer transmits only the Raman scattering that is polarized along the same direction as that of the exciting radiation. By initially orienting c parallel to the electric vector of the exciting radiation (also parallel to the electric vector of the scattered radiation), the cc-polarized Raman spectrum (Icc) is obtained. Subsequent 90° rotation of the fiber axis yields the bb-polarized spectrum (Ibb). As illustrated in Figure 17.8.4, the Eulerian angles θ and χ relate the orientation of the axis system abc of the protein specimen to the orientation of the Raman tensor coordinate system (xyz) of a given normal mode of vibration (Tsuboi et al., 1996b). To each normal mode (Raman band), a Raman tensor α′ is assigned, which is defined as the first derivative
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1646 T,C 1651
257 nm
1575 A,G
1335 A
Raman intensity
853 Y
229 nm
1483 A,G
fd virus
981 SO42– 1010 W
of the molecular polarizability with respect to the vibrational normal coordinate (see Equation 17.8.3; hereafter, the prime symbol is omitted for convenience). The tensor component αij represents the change of polarizability for radiation with incident and scattered electric vectors polarized, respectively, in the i and j directions. As previously noted (Tsuboi and Thomas, 1997), only the relative magnitudes of the diagonal tensor components, namely αxx/αzz ≡ r1 and αyy/αzz ≡ r2, need be considered. Highly anisotropic Raman tensors are characterized by values of r1 and/or r2 differing greatly from unity, whereas isotropic tensors have r1 = r2 = 1. For Raman tensors that are axially symmetric with respect to z, r1 = r2 ≡ r (Overman et al., 1996). Tensor components associated with
600
amide I
668 G
514.5 nm
800
1000
1200
1400
1600
1800
cm–1
Raman Spectroscopy of Proteins
Figure 17.8.1 Raman spectra (600-1800 cm−1) of fd virus excited at 514.5 nm (bottom; 50 µg/µl), 257 nm (middle; 0.5 µg/µl), and 229 nm (top; 0.5 µg/µl). Data are from various sources (Aubrey and Thomas, 1991; Overman et al., 1994; Overman and Thomas, 1995; Overman and Thomas, 1998a) and illustrate the complementary nature of off-resonance (514.5 nm) and UVRR (257 and 229 nm) excitations. Off-resonance excitation produces a rich pattern of Raman bands, primarily from viral coat subunits, which constitute ∼88% of the virion mass. The strongest band, amide I at 1651 cm−1, is diagnostic of subunit α-helix; the weak band at 668 cm−1 identifies C3′-endo/anti dG in packaged DNA; neither band appears in UVRR spectra. The spectrum excited at 257 nm is primarily the Raman signature of the packaged ssDNA, with assignments to specific bases, as labeled. The spectrum excited at 229 nm is the Raman signature of the capsid subunit Trp and Tyr residues, as labeled. Reproduced with permission from Tuma and Thomas (2002).
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1666
1579
1490
2573
1093
1003 830
681
Raman intensity
1618
878
1000
1500
2565 2575
H2O 1000
1500
2500 2000 –1 cm
3000
3500
Figure 17.8.2 Raman spectrum (514.5 nm excitation) of P22 virus at 80 µg/µl in H2O buffer (10 mM Tris, 200 mM NaCl, 10 mM MgCl2, pH 7.5) at 20°C; sample volume 2 µl; no smoothing of data. Bottom trace shows the complete spectrum, including H2O solvent (3000-3700 cm−1) and aliphatic C–H stretch modes of viral protein and DNA (ca. 2900 cm−1). Most of the structurally informative bands occur in the 600-1800 cm−1 interval, expanded in the upper trace. Labels indicate representative Raman markers of dG (681 cm−1), DNA backbone (830 and 1093 cm−1), Trp (878 cm−1), Phe (1003 cm−1), dA and dG (1490 and 1579 cm−1), Tyr and Trp (1618 cm−1), and protein amide I (1666 cm−1). Also labeled is the S–H stretch (2573 cm−1), informative of the environment and dynamics of Cys 405 of the capsid subunit. Note the prominence of the 2573 cm−1 band, even though it represents only one of ∼20,000 chemical bonds in the virion. Reproduced with permission from Tuma and Thomas (2002).
any vibration can be calculated from knowledge of the appropriate polarized Raman band intensities Ikl, where k and l ( = a, b, c) are the directions of the incident and scattered electric vectors, respectively (Benevides et al., 1993). The polarized Raman intensity ratio Icc/Ibb of a spectral band is given in terms of θ, χ, r1 and r2 by Equation 17.8.4 (Tsuboi et al., 1996b).
I cc / I bb =
4[sin 2 θ (r1 cos2 χ + r2 sin 2 χ) + cos 2 θ]2 cos 2 θ (r1 cos 2 χ + r2 sin 2 χ) + r1 sin 2 χ + r2 cos 2 χ + sin 2 θ
2
Equation 17.8.4
Use of Equation 17.8.4 in conjunction with polarized Raman measurements to determine protein side chain and main chain orientations has been described for a number of specific cases (Overman et al., 1996; Tsuboi et al., 1996b; Tsuboi et al., 2001a,b; Tsuboi et al., 2003a,b). Structural Biology
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polarizer
mirror
scattered light to spectrometer
mirror exciting laser beam
beam splitter
mirror
a objective c salt solution
b
Raman cell fiber
Figure 17.8.3 Schematic diagram of a Raman microscope and sampling stage suitable for polarized Raman spectroscopy of oriented single crystals and fibers of biological molecules.
Analysis of Protein Secondary Structure Most useful for protein secondary structure analysis by Raman spectroscopy are amide I, which is primarily a carbonyl stretching mode, and amide III, which combines both in-plane N–H bending and C–N stretching motions. Amide I and amide III bands are expected in the spectral intervals of 1640 to 1680 cm−1 and 1230 to 1310 cm−1, respectively, and the band shapes and peak positions are sensitive to the protein secondary structure. Accordingly, these Raman amide bands are often used as indicators of protein secondary structure. Table 17.8.1 lists the assigned Raman amide I and amide III bands of representative polypeptides and proteins of differing secondary structures. In addition to amide I and amide III, other conformation-sensitive bands have been identified in Raman and UVRR spectra of peptide model compounds and proteins. For example, amide II (1500 to 1600 cm−1 interval), which involves substantial C–N stretching is promoted to prominence in UVRR spectra obtained with 200-nm excitation. Raman amide II intensity depends sensitively on protein α-helical content (Wang et al., 1991; Copeland and Spiro, 1986). Polarized Raman spectra of α-helical proteins also generate a useful secondary structure marker near 1340 to 1345 cm−1, which can be exploited for α-helix orientation analyses (Tsuboi et al., 2000). Additional amide-related modes near 1390 cm−1 have been proposed in UVRR spectra. The UVRR 1390 cm−1 band has been assigned to the overtone of amide V and appears to be sensitive to the local conformation of the peptide linkage (Song and Asher, 1988, 1989; Wang et al., 1991; Austin et al., 1993). Raman Spectroscopy of Proteins
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Table 17.8.1 Raman Amide I and Amide III Bands of Representative Polypeptide and Protein Secondary Structuresa
Molecule
Amide I
Amide III
Secondary structure
α-poly-L-alanine α-poly-L-glutamate α-poly-L-lysine β-poly-L-alanine β-poly-L-glutamate β-poly-L-lysine poly-L-lysine, pH 4 poly-L-glutamate, pH 11 fd subunit Pf1 subunit cI repressor (1-102)
1655 1652 1645 1669 1672 1670 1665 1656 1650 1650 1650 1675 1653 1655 1668 >1668 1659
1265-1348 1290 1295-1311 1226-1243 1236 1240 1243-1248 1249 1270-1300 1270-1300 1286-1297 1245 1273 1235 1231 1249 1236
α-helix α-helix α-helix β-strand β-strand β-strand irregular irregular α-helix α-helix α-helix turns/irregular α-helix β-strand/turns β-strand turns/irregular β-strand
P22 subunit P22 tailspike BPMV subunitb
aData are from the following sources: Carey, 1982; Thomas et al., 1986; Thomas et al., 1987; Sargent et al., 1988;
Li et al., 1990; Prevelige et al., 1990; Bandekar, 1992; Prevelige et al., 1993. bAbbreviation: BPMV, bean pod mosaic virus.
c
z
θ
a O
χ b
y
N
x
Figure 17.8.4 Definitions of the Eulerian angles θ and χ, which designate the orientation of the principal axes (x, y, z) of a tyrosine Raman tensor with respect to the axes (a, b, c) of the oriented protein. Here, θ is the angle by which the Raman tensor principal axis z (perpendicular to the plane of the phenolic ring) is tilted from axis c of the oriented protein, and χ is the angle between the Raman tensor principal axis y (line O-y) and the line of intersection (O-N) between the plane of the phenolic ring and the plane normal to axis c of the oriented protein. An increase in χ corresponds to a counterclockwise rotation of the ring plane about the positive z-axis.
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Raman amide modes typically undergo significant and well-characterized perturbations with peptide deuteration. The resulting H/D shifts serve as probes of both conformational structure and dynamics (Tuma and Thomas, 1997; Tuma et al., 1998a; Tuma et al., 2001). Representative examples are considered in the applications discussed below. Side Chain Conformations and Local Environments A number of correlations have been established between Raman spectral data (band intensities and/or frequencies) and the local environments or structures of specific side chains. Employing such relationships, the side chain conformations, hydropathicities, hydrogen-bonding strengths and similar properties can be determined from the Raman spectrum. In this section, Raman indicator bands of cysteine, tyrosine, tryptophan, and histidine are briefly considered. Cysteine The Raman band resulting from the cysteine sulfhydryl bond (S–H) stretching vibration is a unique probe of local SH structure and dynamics (Li and Thomas, 1991; Raso et al., 2001). The sulfhydryl Raman marker occurs in a region of the spectrum (2500 to 2600 cm−1) that is virtually devoid of interference from other Raman bands of the protein and its sensitivity to hydrogen-bonding interactions of the S–H group is well documented. Correlations established between sulfhydryl hydrogen bonding and the S–H stretching frequency are listed in Table 17.8.2. Because the band is specific to the SH group and its intensity is a direct measure of the concentration of cysteinyl SH sites, it may also be exploited to measure the pKa of thiolate titration (Li et al., 1993; Tuma et al., 1993). Tyrosine It has long been recognized that the para-substituted phenyl ring of tyrosine can generate a pair of relatively intense Raman bands at 850 and 830 cm−1 (Lord and Yu, 1970). A correlation between the ratio of intensities of the tyrosyl doublet (I850/I830) and the hydrogen-bonding state of the phenolic OH group was first recognized by Siamwiza and co-workers (1975). The doublet results from the phenomenon of Fermi resonance involving the in-plane fundamental mode expected near 840 cm−1 (designated Y1) and the first overtone of the out-of-plane fundamental near 420 cm−1 (Y16a) (Siamwiza et al., 1975; Harada and Takeuchi, 1986). Both members of the Fermi doublet, i.e., Y1 and 2Y16a, share the intensity of the uncoupled Y1 band. The hydrogen-bonding state of the phenolic OH group perturbs the Y1 and Y16a modes and thus affects the strength of coupling and the degree of shared intensity. Siamwiza and co-workers demonstrated that the ratio I850/I830 reflects the OH hydrogen-bonding state as follows: (1) when the phenolic OH Table 17.8.2 Bondinga
Raman Spectroscopy of Proteins
Dependence of the Raman S–H Frequency and Band Width on Hydrogen
Hydrogen bonding state of S–H group
S–H band frequency (cm−1)
S–H band Examples width (cm−1)
no hydrogen bond S acceptor weak S–H donor
2581-2589 2590-2595 2575-2580
12-17 12-17 20-25
moderate S–H donor
2560-2575
25-30
strong S–H donor
2525-2560
35-60
S–H donor and S acceptor
2565-2575
30-40
thiols in CCl4 (dilute solution) thiols in CHCl3 thiol neat liquids; thiols in thioesters thiols in acetone; crystal structures thiols in dimethylacetamide; crystal structures thiols in H2O
aResults from Li and Thomas, 1991 and Li et al., 1992.
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group is a strong hydrogen-bond donor, I850/I830 = 0.3; (2) when the phenolic OH group is a strong hydrogen-bond acceptor, I850/I830 = 2.5; and (3) when the phenolic OH group functions as both a donor and an acceptor, I850/I830 = 1.25 (Siamwiza et al., 1975). For a protein containing a single tyrosine, the I850/I830 ratio is an unambiguous marker of the hydrogen-bonding state. If the protein contains multiple tyrosines, I850/I830 reflects the average of tyrosyl hydrogen-bonding states. Recent studies have shown that the tyrosyl Fermi doublet intensity ratio may lie outside the range 0.3 < I850/I830 < 2.5, notably I850/I830 = 6.7 for a non-hydrogen-bonded phenoxyl group (Arp et al., 2001). This state, while unlikely for globular proteins, is encountered for the tyrosine side chains in coat-protein subunits of filamentous virus capsids and may also occur for certain tyrosines within transmembrane α-helices. Further discussion is given in the applications section below. Tryptophan The indolyl moiety of tryptophan generates many prominent Raman bands (Takeuchi and Harada, 1986), several of which have been correlated with the local environment and geometry of the tryptophan side chain in proteins. Notable among these are the following. (1) Normal mode W3 generates an intense and sharp Raman band in the interval 1540 to 1560 cm−1, the peak position of which is indicative of the absolute value of the side chain torsion angle χ2,1 (Cδ1–Cγ–Cβ–Cα) (Miura et al., 1989). This relationship is illustrated in Figure 17.8.5. For proteins containing more than one tryptophan residue, the Raman peak position, bandwidth and band shape provide information on the |χ2,1 | distribution among all tryptophan side chains. (2) The tryptophan residue also generates a Fermi doublet with components at 1360 and 1340 cm−1. The Fermi doublet intensity ratio (I1360/I1340) increases with increasing hydrophobicity of the indolyl ring environment and thus serves as an indicator of local hydropathy (Harada et al., 1986). This correlation extends to UVRR spectra (Miura et al., 1988). (3) Normal mode W17 produces a Raman band near 880 cm−1, the frequency of which is sensitive to indolyl N–H hydrogen-bond donation (Takeuchi and Harada, 1986). W17 exhibits a relatively high value (∼883 cm−1) when the indolyl moiety is located in a highly hydrophobic environment, such as the hydrophobic core of a globular protein (Kitagawa et al., 1979; Miura et al., 1991). The band center decreases with increasing strength of N–H⋅⋅⋅X hydrogen bonding and values as low as 871 cm−1 have been reported for strongly hydrogen-bonded indoles (Takeuchi and Harada, 1986). N1 deuteration causes the band to shift to a lower wave number by ∼20 cm−1. (4) Normal mode W18, which is an indole ring-breathing vibration (Takeuchi and Harada, 1986), generates one of the most prominent bands (near 755 cm−1) in Raman spectra of proteins. Typically, the band intensity increases with decreasing hydrophobicity of the indolyl ring environment (Miura et al., 1991), as judged by normalization of the W18 band intensity to that of the W16 mode near 1010 cm−1 (Takeuchi and Harada, 1986). Histidine In H2O solution spectra of proteins, Raman markers of histidine (including the histidinium ion) are relatively weak in comparison to those of tyrosine and tryptophan. In D2O solution, however, histidine generates a moderately intense Raman band near 1408 cm−1, which is assigned to an in-plane vibration of the N-deuterated imidazolium ring (Tasumi et al., 1982). The band is particularly useful for monitoring histidine H/D exchange dynamics as well as histidine-histidinium equilibria (in D2O) (Harada et al., 1982; Okada et al., 2003). A linear correlation between the frequency of the 1408 cm−1 Raman band and the histidine side chain conformation has been proposed (Takeuchi et al., 1991). Structural Biology
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W3 frequency(cm–1)
1555
1550
C χ1 Cα χ2,1 Cβ N 3 2 N1 H
1545
60°
90°
120°
χ 2,1 : glycyl-L -tryptophan dihydrate : N-acetyl-L -tryptophan methyl ester : DL-tryptophan formate : N-acetyl-DL-tryptophan methylamide : DL-tryptophan ethyl ester : N-acetyl-L -tryptophan : L-tryptophan HCI
Figure 17.8.5 Relationship between the frequency of tryptophan normal mode W3 and the absolute magnitude of the side chain torsion χ2,1 (abscissa), as established from the examination of Raman spectra of the following model compounds of known structure: glycyl-L-tryptophan dihydrate; N-acetyl-L-tryptophan methyl ester; DL-tryptophan formate; N-acetyl-DL-tryptophan methylamide; DL-tryptophan ethyl ester; N-acetyl-L-tryptophan; L-tryptophan HCl (Miura et al., 1989; reproduced with permission).
APPLICATIONS TO PROTEIN STRUCTURES AND ASSEMBLIES In this section, recent representative Raman applications are discussed to illustrate the scope of the method and to delineate many of the principles and practical considerations discussed in the preceding sections. Proteins of Icosahedral Viruses
Raman Spectroscopy of Proteins
Raman spectroscopy provides a useful means of investigating protein folding pathways in the context of supramolecular assembly. An example is the assembly of the Salmonella bacteriophage P22, the Raman spectrum of which is shown in Figure 17.8.2. The icosahedral capsid lattice of P22 is constructed from 420 copies of a 429-residue coat subunit (gp5, 47 kDa), 12 copies of a 725-residue portal subunit (gp1, 84 kDa), plus a small number of minor proteins (Prevelige and King, 1993). In the initial assembly step, a procapsid or immature shell is formed with the help of ∼300 subunits of a 303-residue scaffolding protein (gp8, 34 kDa). The twelve portal subunits form a ring, which is located at a unique five-fold vertex of the procapsid, where it serves as the channel for DNA translocation at the time of genome packaging. DNA packaging, which is under the control of virally encoded terminase and ATP hydrolysis, results in expulsion of scaffold-
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ing subunits through orifices of the procapsid lattice with concurrent expansion of the procapsid to the mature capsid. The coat protein subunits of the mature capsid lattice are conformationally altered in this maturation process, a phenomenon that is common to many dsDNA viruses. The portal is subsequently closed by the binding of additional minor proteins. Attachment of up to six trimeric tailspikes (gp9, 71 kDa) results in the infectious phage particle (Fig. 17.8.6). Raman spectroscopy has been used to monitor many properties of P22 structural proteins, including key events related to the assembly of the mature virion. Here, the following representative examples are considered: (1) formation of the P22 procapsid shell from the coat protein monomer (the initial assembly step) and subsequent transformation of the procapsid shell into the mature capsid; (2) formation of the dodecameric portal from the portal monomer; and (3) analysis of the sulfhydryl hydrogen-bonding states of tailspike cysteines. Bacteriophage P22 capsid The assembly of an icosahedral viral shell from identical protein subunits requires highly specific interactions to sequester hydrophobic residues at subunit interfaces and avert unproductive aggregations. A plausible assembly mechanism is one involving several discrete steps, each with attendant changes in subunit structure. In this scheme, an assembly intermediate (precursor shell or procapsid) would consist of subunits in incompletely folded states, while the mature shell (capsid) would contain the most stable subunit fold. Because assembly intermediates are likely to possess subunits of well-defined secondary and tertiary structures, resembling the late-folding intermediates observed for small globular proteins, they should be distinguishable by the extent of protection of their peptide NH groups against deuterium exchange. Although NMR measurement of H/D exchanges is feasible in small globular proteins (Bai et al., 1995), icosahedral viral shells typically contain hundreds of large protein subunits and are well beyond the size limitations of the NMR method. Raman spectroscopy is not hindered by this size limitation. Raman spectroscopic investigations of the P22 procapsid-to-capsid maturation (Prevelige et al., 1993; Tuma et al., 1998a) have provided molecular details of the lattice rearrangement (reviewed previously by Thomas, 1999). Here, focus is on a recent structural characterization of the initial assembly step, i.e., the formation of the P22 procapsid shell from 420 copies of the coat protein monomer (Tuma et al., 2001). This study was made possible by the greatly improved sensitivity of Raman instrumentation, which allows examination of the coat protein in its monomeric state at very low concentration. Relevant data are shown in Figure 17.8.7 (Tuma et al., 2001). Although Raman spectra of the monomeric and procapsid shell states of the P22 coat protein share many similar features, it is clear from Figure 17.8.7 that procapsid assembly perturbs significantly the structure of the coat protein monomer. The computed difference spectrum shown in the bottom trace of Figure 17.8.7 highlights the spectral changes accompanying assembly. Importantly, the center of the Raman amide I marker is shifted from 1667 cm−1 in the monomer to 1663 cm−1 in the procapsid shell. This shift gives rise to Raman difference peaks at 1655 cm−1 (assigned to α-helix) and 1673 cm−1 (β-strand) and corresponding troughs at lower and higher wavenumber values (disordered main chain). The difference band pattern of Figure 17.8.7 reflects a narrower amide I profile in the procapsid shell spectrum, i.e., the more ordered secondary structure of the shell subunit occurs at the expense of the disordered chain segments of the monomer. The more highly ordered secondary structure of the subunit in the assembled shell is further demonstrated in the difference spectrum of Figure 17.8.7 by the diminished intensity of the Raman amide III band at 1255 cm−1, assigned to disordered structure and by the enhanced intensity of the Raman amide III marker at 1230 cm−1 (β-strand). These results show that the polypeptide backbone of the coat protein monomer undergoes refolding during procapsid assembly. Overall, ∼4% ±
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coat scaffolding portal gp5(420) gp8(~300) gp1(12)
procapsid assembly
procapsid pilot proteins gp7, gp16, gp20
DNA packaging and expansion gp9 gp4, gp10, gp26
mature virion
portal closure
tailspike attachment
capsid
maturation
Figure 17.8.6 Assembly pathway of bacteriophage P22 (Casjens and King, 1975; King et al., 1976) proceeding clockwise from upper left. Co-assembly of 420 copies of the coat subunit (gp5), 12 copies of the portal subunit (gp1) and ∼300 copies of the scaffolding subunit (gp8), together with a few copies of minor pilot proteins (gp7, gp16, gp20), leads to formation of the procapsid, which subsequently matures to the icosahedral capsid at the time of DNA packaging and scaffold ejection. The ATP-driven DNA packaging event is accomplished with the aid of a virally encoded phosphatase and additional minor proteins (terminase subunits gp2 and gp3; not shown). Portal closure by minor proteins (gp4, gp10, gp26) and attachment of six trimeric tailspikes (gp9) lead to the mature virion. Adapted with permission from Rodriguez-Casado et al. (2001).
1% of the backbone, or 17 of 428 peptide bonds, are estimated to undergo refolding based on the normalized area of the amide I band differences. The likely candidates for refolding are residues within a flexible hinge. Thus, assembly of the P22 procapsid leads to local reordering of a small but significant percentage of the peptide main chain. In addition to the changes in subunit secondary structure noted above, the Raman spectrum also provides information on the effect of procapsid assembly on subunit tertiary structure. For example, the shift of the tryptophan (Trp) marker band (normal mode W18) at 759 cm−1 in the monomer to 756 cm−1 in the procapsid shell with a concomitantly large decrease in intensity (Fig. 17.8.7) signifies a change in the hydropathic environment of the average Trp side chain. Broadening of the 1360 cm−1 component of the Trp Fermi doublet indicates that this change is due to a more hydrophilic indolyl environment in the procapsid shell subunit. Overall, the changes in Trp side chain environment are small. Further, cysteine (Cys) sulfhydryl and tyrosine (Tyr) phenoxyl hydrogen-bonding environments are essentially invariant to the assembly process. Thus, despite measurable refolding of the coat protein backbone with procapsid assembly, the tertiary structure of the subunit is not greatly affected. Raman Spectroscopy of Proteins
The folding state of the coat protein monomer has also been characterized by H/D exchange (Tuma et al., 2001). Figure 17.8.8A compares Raman markers of the exchanged
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1242
1341
1251
1341 1360
1400
1655 1673
1551 1605 1618
1667
1449
1551
1200
1470*
1000
1341 1360
800
1230 1255
965
1042*
760
difference
958
759 828 855 878
621 643
monomer
600
1663
1003 877
756
procaspid shell
1600
1800
cm–1 Figure 17.8.7 Raman spectra in the region 600-1800 cm−1 (514.5 nm excitation) of the bacteriophage P22 procapsid shell at 18 µg/µl and 10°C in aqueous buffer (top) and unassembled coat protein monomer at 16 µg/µl and 2°C in aqueous buffer (middle). The difference spectrum (bottom) was computed by subtraction of the middle spectrum from the top spectrum. Spectra were corrected for Raman scattering of the buffer and intensities were normalized using the 1003 cm-1 band of phenylalanine (Tuma et al., 2001; reproduced with permission).
ND groups (amide III′ region, 900 to 1000 cm−1) in monomer and procapsid shell states, following exposure of each sample to D2O (pD 7.8) under native conditions (2°C for monomer and 10°C for procapsid shell). Also shown is the amide III′ profile of a coat protein control in which all NH groups have been exchanged by deuterium under denaturing conditions prior to refolding of the protein in D2O. These data show that H/D exchange is more extensive in the monomer than in the procapsid shell subunits. In particular, greater amide III′ intensity is observed at 930 to 945 cm−1, assignable to deuterated α-helix (Yu et al., 1973; Overman and Thomas, 1998a), at 960 cm−1, assignable to deuterated disordered chain segments (Chen and Lord, 1974; Chen et al., 1974; Tuma et al., 1998a), and at 980 to 990 cm−1, assignable to deuterated β-strand (Yu et al., 1973; Chen and Lord, 1974; Chen et al., 1974; Tuma et al., 1998a). The amide III′ exchange profile of the monomer indicates that the protein fold comprises unprotected and protected residues in regions of α-helix, β-strand, and disordered secondary structure. Figure 17.8.8A also shows that neither the procapsid shell subunit nor the monomer undergoes complete exchange of all the peptide residues. The data suggest further that the most protected residues of the monomer are those located in regions of β-strand structure. Raman-monitored H/D exchange can also be used to identify specific side chains of the protein folding core (Miura and Thomas, 1995a; Thomas, 1999). For example, the degree of protection of the S–H group of the single cysteine (Cys 405) in the coat protein is a measure of the accessibility of this residue and of the adjoining C-terminal region of the protein to H/D exchange. Figure 17.8.8B shows Raman spectra in the region 1840 to 1890 cm−1, where the stretching vibration of the deuterated sulfhydryl group (S–D) is expected
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A
850
900
950
B 100% D control monomer 120 hr 2°C D2O monomer 90 hr 2°C D2O
1840
985
1000
1050
1100
cm–1
1850
1866
800
960
930
100% D control monomer procapsid
1860
1870
1880
1890
cm–1
Figure 17.8.8 Normalized Raman signatures of native-state H/D exchanges of the P22 coat protein monomer (2°C) and procapsid shell (10°C), following 12 hr exposure to D2O buffer. (A) amide III′ profiles; included as a control is the S–D band profile of the fully exchanged procapsid subunit incubated at 40°C. (B) S–D stretching band profiles, following prolonged exposure to D2O (Tuma et al., 2001; reproduced with permission).
Raman Spectroscopy of Proteins
to occur (Tuma et al., 2001). The absence of an S–D band in the spectrum of the monomer shows that the Cys 405 sulfhydryl is fully protected against H/D exchange. Protection persists for samples maintained at native conditions (pD 7.8, 2°C) for up to 6 days. The S–H group can be exchanged only at conditions that favor global unfolding (40°C). Thus, the Cys 405 sulfhydryl is buried within the hydrophobic folding core of the subunit monomer. A similar degree of sulfhydryl exchange protection is observed when the subunit is assembled into the procapsid shell. Conversely, such protection is not observed for the mature capsid shell (Tuma et al., 1998a). This shows that the procapsid → capsid lattice transformation is accompanied by loss of sequestration of the subunit C-terminal region from H/D exchange. In summary, during P22 shell assembly, ∼20 peptide linkages undergo local refolding without affecting the global structure of the monomer. The procapsid subunit achieves enhanced protection against H/D exchange vis-à-vis the monomer. On the other hand, the monomer possesses a sizable protected folding core that
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2585
2565
2550 2530
Am I CH def
Am III
F
cm–1
2600
1000
1667
1238
W
W
Y W
W
Y F
Raman intensity
2500
1500
2000
2500
3000
Wavenumber (cm–1)
Figure 17.8.9 Raman spectrum (600-2800 cm−1) of the P22 trimeric tailspike protein at 10°C. The tailspike was dissolved at 92 mg/ml in 50 mM Tris, 2 mM EDTA, pH 7.0. The inset at upper right, which shows an amplification of the spectral interval 2480-2620 cm−1, exhibits the composite S–H stretching profile (band components are resolved at 2530, 2550, 2565, and 2585 cm−1) of the eight cysteines per subunit. The data are uncorrected for contributions of the solvent and scattering background (Raso et al., 2001; reproduced with permission).
encompasses Cys 405. Monomeric and procapsid states of the subunit represent distinct and compact late-folding intermediates, which ultimately mature to the fully folded capsid state. Cysteinyl SH interactions of the P22 tailspike protein Attachment of a virion to its host cell receptor requires a specialized apparatus called a tailspike. In the case of P22, six tailspikes are arranged symmetrically at the unique icosahedral vertex that also incorporates a portal apparatus for channeling cellular entry of viral DNA (Fig. 17.8.6). The tailspike of bacteriophage P22 is a trimeric assembly of a 666-residue subunit that contains eight cysteine residues widely distributed through the sequence: Cys 169, Cys 267, Cys 287, Cys 290, Cys 458, Cys 496, Cys 613, and Cys 635 (Sauer et al., 1982). Each cysteine contributes to the prolific S–H stretching band observed in the 2500 to 2600 cm−1 region of the Raman spectrum (Fig. 17.8.9) (Sargent et al., 1988; Thomas et al., 1990; Raso et al., 2001). The X-ray crystal structure of the trimeric tailspike (Steinbacher et al., 1994; Steinbacher et al., 1997a,b) locates subunit cysteines in the context of the protein fold—a parallel β-helix—as illustrated in Figure 17.8.10. However, the X-ray structure does not locate sulfhydryl protons and provides no information on the hydrogen-bonding states of these groups. Sulfhydryl interactions are important not only for their contributions to the stability of the native structure but also because the folding and assembly of the trimeric tailspike requires selective and transient oxidation of a subset of cysteine sulfhydryl (SH) groups to the disulfide (SS) form (Robinson and King, 1997; Haase-Pettingell et al., 2001). Identification of the specific Cys residues participating in sulfur redox chemistry has the potential to provide important insights into the protein folding and assembly pathway.
Structural Biology
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A
N-terminal head binding domain
B
Cys 169 Cys 290 β-coil domain
C
Cys 267 Cys 287
Cys 290
Cys 458 Cys 496 β-prism region
Met 448 Cys 496
Cys 613 Cys 635
Figure 17.8.10 (A) Ribbon diagram of the X-ray crystal structure of the trimeric P22 tailspike protein, showing the N-terminal head-binding domain, the central parallel β-helix (or β-coil) domain and the C-terminal β-prism domain (Steinbacher et al., 1994; Steinbacher et al., 1997a; Steinbacher et al., 1997b). (B) The β-helix and β-prism domains of a single subunit, showing the locations of the eight cysteines with space-filling atoms. (C) Locations of Cys 290 and Cys 496 within the predominantly hydrophobic stack of the parallel β-helix domain. Met 448 is also labeled to avoid confusion with cysteine (Raso et al., 2001; reproduced with permission).
Although all cysteines of the mature tailspike are in the sulfhydryl form, the complexity of the Raman S–H band envelope (Fig. 17.8.9) suggests a diverse range of hydrogenbonding strengths. To correlate S–H⋅⋅⋅X strengths with specific Cys sites, eight single-site Cys → Ser mutants were engineered genetically and investigated by Raman spectroscopy (Raso et al., 2001). Raman signatures of the individual Cys S–H groups, obtained by difference spectroscopy, are shown in Figure 17.8.11. On the basis of previously established correlations between the state of sulfhydryl hydrogen bonding and the Raman S–H frequency (Li and Thomas, 1991; Li et al., 1992), the tailspike cysteines can be ranked according to the strength of S–H⋅⋅⋅X hydrogen-bonding as in Figure 17.8.12. Of particular interest is Cys 613, which not only forms the strongest S–H⋅⋅⋅X hydrogen bond so far encountered in a protein, but also is implicated in transient disulfide bond formation (Haase-Pettingell et al., 2001). Other structural and dynamic characteristics of the Cys S–H groups determined from the Raman spectra have been discussed (Raso et al., 2001). These studies provide a basis for future Raman characterization of Cys SH interactions in folding intermediates of the P22 tailspike, and demonstrate a general approach for probing cysteine environments in protein assemblies.
Raman Spectroscopy of Proteins
P22 portal protein Portal proteins participate in many stages of virion assembly—including formation of a topologically correct ring for procapsid incorporation, retention of the ring during shell maturation, participation of the assembly in ATPase-catalyzed DNA translocation, and interactions with proteins that subsequently cap the translocation channel. This suggests
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A
B 2565
wild type C169S
2550
C267S
2585
C169S
2550
C287S C287S
Raman intensity
2585
C267S
2585
C290S C290S 2550
C458S C458S
2585
C496S
C613S C613S
2530
C496S
2576
C635S C635S
2500
2540 cm –1
2580
2620
2500
2540
2580
2620
cm –1
Figure 17.8.11 S—H Raman signatures (2480-2620 cm−1) of tailspike cysteines. (A) The Raman S—H profiles observed for the wild-type tailspike and for each of the eight Cys → Ser mutants, as labeled. (B) Raman difference spectra, computed as mutant minus wild-type, for each of the eight Cys → Ser mutants; in each trace, the S—H Raman signature of the mutated Cys site is revealed as a negative spectrum (Raso et al., 2001; reproduced with permission).
strongest H-bond
weakest H-bond
Cys 613 > Cys 458 > Cys 267 > Cys 287 > Cys 169 > Cys 635 > Cys 290 = Cys 496
Figure 17.8.12 Tailspike cysteines ranked according to the strengths of S–H⋅⋅⋅X hydrogen bonding. Structural Biology
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a highly versatile portal subunit structure capable of undergoing path-dependent switching between different conformational states (Bazinet and King, 1985; Valpuesta and Carrascosa, 1994). Crystallography and imaging methods reveal many architectural features of isolated portals (Valpuesta et al., 1999; Simpson et al., 2000; Orlova et al., 1999; Droge and Tavares, 2000), but provide little insight into molecular mechanisms of subunit folding, assembly, or stability. Recent biochemical (Moore and Prevelige, 2001) and Raman studies (Rodriguez-Casado et al., 2001; Rodriguez-Casado and Thomas, 2003) of the P22 portal protein provide a starting point for elucidating the complex structural and functional roles of the portal subunit. Raman spectra of unassembled (monomeric) and assembled (dodecameric) forms of the P22 portal protein are shown in Figure 17.8.13. The data indicate an α/β fold that is not appreciably altered by subunit assembly. Conversely, portal assembly generates major changes in the Raman markers that are diagnostic of side chain local environments. Most striking are changes observed in the 2520 to 2600 cm−1 region (Fig. 17.8.13, right panel), which indicate alterations in cysteine S–H hydrogen-bonding environments. In the case of the monomer, the Raman S–H profile is relatively broad with a central peak at 2562 cm−1 and a prominent shoulder of nearly equal intensity at 2557 cm−1. The positions, intensities, and widths of these overlapping bands suggest that although all four cysteines (Cys 153, Cys 173, Cys 283, Cys 516) of the monomer engage in relatively robust S–H⋅⋅⋅X hydrogen bonds, they partition into two sub-populations characterized by markers near 2557 and 2562 cm−1, respectively. For the dodecamer, the Raman S–H bands are collectively shifted to higher wavenumber values, indicative of general weakening of S–H⋅⋅⋅X hydrogen-bond strengths with assembly (Rodriguez-Casado et al., 2001). To further characterize the roles of portal cysteines in subunit folding and dodecamer assembly pathways, Raman spectroscopy was employed in combination with site-directed mutagenesis to measure Raman S–H signatures of the four single-site Cys → Ser mutant portals (Fig. 17.8.14; Rodriguez-Casado and Thomas, 2003). The data of Figure 17.8.14 resolve the residue-specific Raman sulfhydryl bands and provide an internally consistent S–H⋅⋅⋅X classification scheme for the portal monomers, as given in Table 17.8.3. On the basis of the previously established correlations (Li and Thomas, 1991; Li et al., 1992), the results show that the composite S–H Raman band envelope of the WT monomer comprises the following: (1) moderate S–H⋅⋅⋅X interactions from Cys 173 and Cys 283, and (2) strong S–H⋅⋅⋅X interactions from Cys 153 and Cys 516. Corresponding analysis of the data from mutant portal assemblies leads to the following additional conclusions: (3) Each portal variant exhibits a Raman S–H signature that is greatly perturbed by assembly; (4) all assemblies exhibit a Raman peak at 2570 cm−1 or higher, despite the absence of such a marker in the monomeric form; (5) with the exception of C516S, no mutant exhibits an S–H group that is promoted to a stronger hydrogen-bonding state with assembly.
Raman Spectroscopy of Proteins
The difference spectra computed between mutant and WT portal assemblies are complex. In previous work on Cys → Ser mutant tailspikes of phage P22, Raso and co-workers (2001) were able to deduce the unique S–H signatures of all eight cysteines of the WT tailspike. The tailspike Raman data indicated that Cys signatures are uncoupled to one another, i.e., mutation of one Cys site did not perturb the S–H⋅⋅⋅X hydrogen-bonding state of any other Cys site (Raso et al., 2001). While Figure 17.8.14A shows this to be also true for the portal monomers, Figure 17.8.14B indicates that cysteine signatures of the assemblies are not similarly conserved. A tabulation of plausible assignments and S–H⋅⋅⋅X hydrogen-bonding schemes for the mutant portal assemblies is given in the right column of Table 17.8.3.
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A
800
1200 cm–1
1600
2587 2559
1648 1656 1678 1669
1548 1558
1469
1400
2587
2557 2562
1653 1672
2565
1000
1211 1232 1259 1278 1305 1354 1334
1080
1006
1064
852 892
759
difference
600
2566
1606 1617 1655 1672
1551
1554 1606 1617
1031
1080
827 852 901 934 957
757 756
828 852 901 935 959
monomer 620 642
Raman intensity
620 642
dodecamer
1002 1031 1079 1103 1102 1126 1126 1173 1174 1207 1207 1252 1244 1268 1269 1318 1320 1340 1340 1400 1400 1423 1423 1448 1448 1460 1459
B
1800 2520 2560 cm–1
2600
Figure 17.8.13 (A) Raman spectra (600-1800 cm−1, 532 nm excitation) of the P22 portal protein monomer (middle trace) and dodecamer (top trace) and the digital difference spectrum (second trace from bottom) obtained by subtracting the spectrum of the monomer from that of the ring. Also shown (bottom trace) is an approximate three-fold amplification of the difference spectrum. Proteins were dissolved to ∼40 mg/ml in 20 mM Tris⋅Cl (pH 7.5)/100 mM NaCl/2 mM EDTA. Spectra were corrected for contributions of the buffer solution and the gently sloping background typical of protein Raman spectra. Identical spectra were obtained for protein concentrations in the range 25 to 100 mg/ml. (B) Raman spectra (2520-2600 cm−1, 532 nm excitation) of the P22 portal protein monomer (middle trace) and dodecamer (top trace) and the digital difference spectrum (bottom trace) obtained by subtracting the spectrum of the monomer from that of the dodecamer. Data are adapted with permission from Rodriguez-Casado et al., 2001.
In addition to assembly-related changes to the hydrogen-bonding state of the portal cysteines, assembly-related perturbations are observed for portal tyrosines and tryptophans. The portal subunit contains 26 tyrosines (Eppler et al., 1991). Most occur within the central 50% of the sequence (residues 223 to 571) and none occurs in the C-terminal segment (572 to 725). The bottom trace of Figure 17.8.13 shows that the tyrosine marker at 852 cm−1 undergoes the largest relative intensity change in the Raman spectrum. This implicates the central region of the sequence in the intersubunit interface. A similar conclusion is reached from consideration of perturbed Trp markers (Rodriguez-Casado and Thomas, 2003). In contrast to the considerable perturbations to side chain environments accompanying portal assembly, changes in secondary structure are small. In the case of amide I (1600 to 1750 cm−1, Fig. 17.8.13), only small intensity changes (∼2%) in α-helix (1656 cm−1) and β-strand (1669 cm−1) markers are observed. The difference spectrum is consistent
Structural Biology
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Raman intensity
C516S C283S C173S
differences 2555 2563
monomer profiles 2558 2563
A
C516S-WT
C283S-WT
C173S-WT
C153S C153S-WT
WT
2555
assembly profiles 2563 2574 2586
B
2590
2555
2590
differences
C516S-WT
Raman intensity
C516S C283S C173S
C283S-WT
C173S-WT
C153S
2555
2590
2555 2590
C516S
2563
2586
assembly-monomer differences 2559 2563
C
2586
WT
2563
C153S-WT
2587
Raman intensity
C283S
C173S C153S
2563
WT
2555
2590
cm–1
Raman Spectroscopy of Proteins
2555
2590
cm–1
Figure 17.8.14 Raman profiles of WT and mutant portal proteins in the region of cysteine S–H stretching vibrations (2520-2620 cm−1). Conditions are as given in Figure 17.8.13. (A) The left panel compares S–H profiles of the monomeric proteins (WT and mutants, as labeled). The right panel shows the S–H signature specific to each cysteine of the WT monomer, obtained by subtracting the WT spectrum from the corresponding mutant spectrum. (B) The left panel compares S–H profiles of the assembled proteins (WT and mutants, as labeled). The right panel shows the S–H signature specific to each cysteine of the WT assembly, obtained by subtracting the WT spectrum from the corresponding mutant spectrum. (C) The left panel shows the computed assembly-minusmonomer difference spectrum for each variant, as labeled. The right panel demonstrates the non-additivity of cysteine signatures for the assembled proteins. Thus, the Raman signature of C516 that is obtained by subtracting the C516S mutant spectrum from the WT spectrum (B, top right) is not identical to the C516 signature (dashed-line trace) generated by subtracting the sum of signatures of C153, C173 and C283 (solid-line trace) from the WT spectrum (dotted-line trace). Data are adapted with permission from Rodriguez-Casado and Thomas, 2003.
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Table 17.8.3 Cysteine Sulfhydryl Raman Markers and S–H⋅⋅⋅X Hydrogen-Bond Strengths of the P22 Portal Proteina
Cysteineb
Monomeric form
Assembled form
cm−1
H-bond
cm−1
H-bond
Cys 153 Cys 173
2558c 2563c
strong moderate
Cys 283
2563c
moderate
Cys 516
2558c
strong
2563c 2563d (50%) 2586d (50%) 2563d (27%) 2586d (73%) 2563 (50%) 2586 (50%)
moderate moderate very weak moderate very weak moderate very weak
aHydrogen-bond strengths are based upon model compound studies of Li and Thomas (1991). bAmino acid residue in wild-type portal protein. cPeak of Raman S-H stretching band. dComponent of closely spaced doublet. The percentage of the total Raman intensity contributed by each
component of the doublet is given in parentheses.
with curve fitting results (Berjot et al., 1987), which suggest only a marginal change in portal subunit secondary structure with assembly. It is interesting that this type of Raman difference fingerprint parallels that observed with P22 procapsid maturation (Prevelige et al., 1993). Such a different signature may be diagnostic of relatively strong intersubunit interactions. In summary, Raman studies of the P22 portal protein show that assembly leads to only a small perturbation of subunit secondary structure but large changes in the local environments of subunit side chains. In particular, portal assembly results in a major reorganization of cysteine S–H hydrogen-bonding states, substantial alteration of tyrosine O–H hydrogen-bonding states and large changes in tryptophan hydropathic environments. The Raman-monitored changes in side chains indicate that the N-terminal third and central third of the portal sequence are involved in intersubunit recognition. Other applications to icosahedral viruses In other recent Raman applications to icosahedral viruses, the method has been employed to probe the coat protein recognition domain of the P22 scaffolding protein (Tuma et al., 1998b), identify secondary and tertiary structures and enzymatic activity of the RNAtranslocating enzyme (P4) of the φ6 nucleocapsid (Juuti et al., 1998; Jenkins et al., 1999), investigate capsid assembly in PRD1 (Tuma et al., 1996a), identify subunit-specific interactions of the φ6 procapsid (Benevides et al., 2002), and characterize conformations and interactions of the packaged viral DNA genomes within capsids of bacteriophages T7, PRD1, and P22 (Aubrey et al., 1992; Tuma et al., 1996b; Overman et al., 1998). Proteins of Filamentous Viruses Bacteriophages fd, f1, and M13 are structurally identical members of the Ff group of class I filamentous viruses, which infect strains of Escherichia coli. The cylindrical Ff filament (∼880-nm length, ∼6-nm diameter) contains a covalently-closed ssDNA genome of 6410 nucleotides, sheathed by ∼2750 copies of a 50-residue α-helical capsid subunit (protein pVIII) plus a few copies of minor proteins at the filament ends. The capsid subunit sequence (1AEGDDPAKAA 11FDSLQASATE 21YIGYAWAMVV 31VIVGATIGIK 41 LFKKFTSKAS) is essentially identical among members of the Ff class (Day et al., 1988). The structure of the native Ff assembly has been studied by methods of solution
Structural Biology
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and fiber spectroscopy and fiber X-ray diffraction, which collectively provide many details of capsid structure and filament architecture (Day et al., 1988; Marvin, 1998). Overall, the results show that the ssDNA core of Ff is coated by a superhelical array of pVIII subunits arranged with five-fold rotational symmetry and an approximately twofold screw axis (C5S2 symmetry), and that the capsid subunit is a continuous α-helix tilted by a small angle from the virion axis. The class II filamentous viruses, represented below by Pseudomonas phages Pf1 and Pf3, differ in filament symmetry (C1S5.4) and other details from the class I particles (Welsh et al., 1998; Welsh et al., 2000). Although a complete three-dimensional structure is not known for any filamentous virus, assembly models have been proposed for Ff, Pf1, and Pf3 on the basis of available data. Because ssDNA constitutes only a small percentage (<15%) of the virion mass, the mode of DNA interaction with capsid subunits has been difficult to assess experimentally. Fiber X-ray diffraction provides no direct evidence on the structure of the packaged genome, or on the nature of DNA base environments or interactions between DNA and capsid subunits. Information on the orientations and interactions of subunit side chains is also absent from fiber diffraction maps. However, these questions can be addressed by Raman and UVRR methods and numerous studies have been reported (Thomas and Murphy, 1975; Thomas et al., 1983; Thomas et al., 1988; Aubrey and Thomas, 1991; Overman et al., 1994; Overman and Thomas, 1995; Overman et al., 1996; Takeuchi et al., 1996; Tsuboi et al., 1996b; Wen et al., 1997; Overman and Thomas, 1998a; Matsuno et al., 1998; Overman and Thomas, 1999; Wen et al., 1999; Wen and Thomas, 2000; Tsuboi et al., 2000; Arp et al., 2001; Tsuboi et al., 2001a; Wen et al., 2001; Tsuboi et al., 2003a, b). To illustrate the power of Raman spectroscopy in conjunction with polarization and isotopeediting methods, recent applications that have yielded information on the orientations of subunit helices and key side chains in native filamentous virus assemblies are reviewed. These applications have also provided new empirical correlations between the data of Raman spectroscopy and the structures of protein moieties. Orientations of α-helical capsid subunits in Ff (fd), Pf1, and Pf3 virions The filamentous virus capsid comprises several thousand identical α-helical subunits, arranged in accordance with the helical symmetry of the capsid lattice. Knowledge of this symmetry and of appropriate Raman tensors permits determination of the orientations of specific molecular subgroups with respect to the virion axis. This approach was first applied to fd by Overman and co-workers, who assessed the average tilt angle of the subunit α-helix axis with respect to the virion axis (Overman et al., 1996). The polarized Raman amide I band was interpreted using the Raman tensor transferred from crystalline aspartame to reveal that the α-helix axis of the capsid subunit makes an average angle of 16° ± 4° with the virion axis. The same protocol was also applied to the polarized Raman data of Pf1 and Pf3 to demonstrate that the α-helix axis of the capsid subunit of each of these class II phages also makes an average angle of 16° ± 4° with the virion axis (Tsuboi et al., 2003a,b). Alternative Raman approaches to determine subunit α-helix orientation have also been described (Takeuchi et al., 1996; Tsuboi et al., 2000). For example, the novel Raman assignment of a prominent band at 1340 to 1345 cm−1 in the spectrum of Pf1 to a coupled C–Cα–H valence angle bending and C–Cα bond stretching mode of the polypeptide main chain (Overman and Thomas, 1998b) was exploited to determine the α-helix tilt angle in Pf1 (Tsuboi et al., 2000). It is interesting to note that this relatively simple method circumvents the somewhat tedious determination of Raman tensors. Raman Spectroscopy of Proteins
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Orientations of aromatic side chains in the Ff (fd, f1) and Pf3 virions Polarized Raman microspectroscopy of oriented filamentous viruses may also be applied to determine subunit side chain orientations, provided that the Raman tensors are known for the diagnostic Raman bands of the target side chain. Using tensors for key Raman markers of tryptophan (Tsuboi et al., 1996a), the precise orientation of the Trp 26 side chain in the coat protein subunit of the native fd assembly has been determined (Tsuboi et al., 1996b). The same protocol has also been used to determine the orientation of the Trp 38 side chain in the coat protein subunit of the native Pf3 assembly. The indolyl ring orientations of Trp 26 in fd and Trp 38 in Pf3, as determined by polarized Raman microspectroscopy, provide important constraints for improved modeling of subunit structures in the viral capsids (Marvin, 1998). It is also of interest to extend such determinations to aqueous solutions of the filamentous viruses. For this purpose, Takeuchi and co-workers (1996) developed the Raman linear intensity difference (RLID) method, which utilizes Raman polarizations measured in UVRR spectra of flow oriented virions in conjunction with knowledge of the transition moments of excited electronic states to deduce side chain orientations. Importantly, the RLID method applied to UVRR bands of the Trp 26 residue in the capsid subunit of aqueous fd yields the same indolyl ring orientation determined by polarized Raman spectroscopy for oriented fd fibers. The polarized Raman and polarized UVRR (RLID) methodologies have been applied most recently to determinations of the orientations of the two tyrosine side chains (Tyr 21 and Tyr 24) in the capsid subunit of Ff (Matsuno et al., 1998; Tsuboi et al., 2001a). In both experimental approaches, single-site mutations of Tyr 21 and Tyr 24 to methionine were exploited in the f1 virion. These mutant viruses are designated f1(Y21M) and f1(Y24M). In the case of the polarized Raman study (Tsuboi et al., 2001a), the mutants were also subjected to deuterium isotope-editing of the non-mutated tyrosyl ring (Overman and Thomas, 1995) and were designated f1(Y21M; Y24d4) and f1(Y24M;Y21d4). This introduces tyrosyl Raman markers (viz. C–D stretching bands in the 2200 to 2400 cm−1 region) that are amenable to Raman tensor determination in the model compound L-tyrosine-2,3,4,5-d4 and well suited to polarized Raman measurement in the labeled f1 virion (Tsuboi et al., 2001a). The C–D stretching bands conveniently occur in a spectral region that is free of interference from other virion constituents; also, the highly localized nature of the carbon-deuterium stretching vibrations is considered favorable to transferability of the Raman tensors. The procedure, results, and interpretation are summarized in Figure 17.8.15, which illustrates the coordinate system defined for the virion (Fig. 17.8.15A), the nature of the experimental data obtained (Fig. 17.8.15B) and the molecular model derived for the capsid subunit (Fig. 17.8.15C; Tsuboi et al., 2001a). These results complement and extend earlier structural studies (Aubrey and Thomas, 1991; Marvin et al., 1994; Symmons et al., 1995; Overman et al., 1996; Tsuboi et al., 1996b). APPLICATION TO DNA-BINDING PROTEINS The DNA-binding domains of gene-regulatory proteins exhibit considerable diversity in structural motif, mechanism of nucleotide recognition, and extent of induced fit at the protein-nucleic acid interface (Steitz, 1990; Allemann and Egli, 1997; Patikoglou and Burley, 1997). High-resolution structures of specific DNA target sites in both proteinbound and protein-free states are unavailable. Barriers are posed by the difficulty of obtaining well-ordered crystals of B-DNA, issues of structural interpretation with respect to the influence of the crystal lattice and a continuing debate regarding the accuracy and precision of NMR approaches to nucleic acid structure determination (Nilges, 1996; Neidle, 1998). Raman difference spectroscopy has been employed as an alternative. In this approach, the spectrum of the complex (DNA with bound protein) is compared digitally with the sum of spectra of the isolated components to yield a difference spectrum
Structural Biology
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A
c filament axis
B f1(Y24M;Y21d 4)
b
θ
z y
Raman intensity
α - helix axis
I cc 2282
1651
a
I bb
f1(Y21M;Y24d 4)
I cc 2282
1651
x
I bb
1600
1800
2200
2400
cm–1
C
Raman Spectroscopy of Proteins
Figure 17.8.15 (A) The coordinate system (abc) for the uniaxially oriented Ff fiber in relation to the α-helical subunit and a tyrosine side chain. The drawing (not to scale) represents a small segment of the ∼880 nm Ff virion and a few α-helical turns of one subunit. The orientation of abc with respect to a tyrosine Raman tensor system (xyz, Figure 17.8.4) is defined by the Eulerian coordinates, θ and χ. (B) Polarized Raman spectra (Icc, Ibb) of oriented fibers of the deuterium-labeled mutant viruses f1(Y21M;Y24d4) and f1(Y24M;Y21d4) in the spectral region 1500-2400 cm−1. (C) Stereo diagram of the energy-minimized structure of neighboring subunits in the assembly model proposed for Ff on the basis of coordinates for Tyr 21 and Tyr 24 developed from the polarized Raman spectra of B and coordinates for other residues as previously described (Aubrey and Thomas, 1991; Marvin et al., 1994; Symmons et al., 1995; Overman et al., 1996; Tsuboi et al., 1996b; Tsuboi et al., 2001a). The virion axis runs vertically and subunit N-termini are at the top. The subunits shown correspond to index numbers k = 0 and k = 5, as defined in the Protein Databank Accession Code 1IFJ. The molecular graphics were obtained using MOLSCRIPT (Kraulis, 1991). Adapted with permission from Tuma and Thomas (2002).
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identifying the DNA and protein sites perturbed by interaction. This approach is well suited for the analysis of protein/DNA complexes in solution. It provides the capability of rapidly screening many gene-regulatory complexes, although with limited 3-D-structural resolution. To date, Raman spectra have been obtained for a number of protein/DNA complexes, which encompass a broad spectrum of DNA-binding motifs and mechanisms of DNA recognition (Benevides et al., 1991a,b; Benevides et al., 1994a,b; Benevides et al., 1996; Benevides et al., 2000a,b). Included are proteins that bind DNA specifically through recognition of DNA bases in either the major groove (phage λ cI repressor; yeast transcriptional activator, GCN4) or minor groove (SRY-HMG box encoded by the human male-determining region of the Y chromosome), as well as proteins that bind DNA nonspecifically (gVp, the ssDNA binding protein of M13 phage). Figure 17.8.16 compares Raman difference spectra for several models of protein/DNA recognition and demonstrates the dramatic differences between spectroscopic signatures of the various binding modes. The spectra of Figure 17.8.16 provide an empirical database proposed as diagnostic of different protein/DNA binding mechanisms. Such a database can be exploited to identify the mechanism of protein/DNA recognition in gene-regulatory complexes that are refractory to high-resolution structural methods. A case in point is the specific complex between phage D108 Ner repressor and a 61-bp operator site that has been structurally characterized by its Raman difference signature (Benevides et al., 1994a). The empirical database can also be used to compare and evaluate binding mechanisms of closely related biological complexes, such as wild-type and mutant variants of transcription factors or repressors with cognate DNA (Benevides et al., 1994b; Benevides et al., 2000a). DEVELOPMENT OF NOVEL SPECTRA-STRUCTURE CORRELATIONS FOR PROTEINS Although viruses are among the largest and most complex supramolecular assemblies so far subjected to Raman spectroscopic investigation, they have served fortuitously as excellent “model” systems, by facilitating the development of new and interesting vibrational spectra-structure correlations (Overman and Thomas, 1998a,b; Tsuboi et al., 2000; Wen and Thomas, 2000; Arp et al., 2001). Of particular interest is the structural significance of the tyrosyl Fermi doublet at 850 and 830 cm−1, which is prominent in Raman spectra of tyrosine-containing globular proteins (Siamwiza et al., 1975). For the filamentous viruses fd and Pf1, the tyrosines of the capsid subunit give rise to an apparent singlet near 853 cm−1 in lieu of the doublet that is found in globular proteins (Fig. 17.8.17). These data illustrate the power of the combined usage of isotope-editing and Raman digital difference spectroscopy as a means of identifying specific contributions of molecular subgroups to the Raman spectrum of a complex nucleoprotein assembly. Through a detailed analysis of the Raman spectra of Ff filamentous viruses (Overman et al., 1994; Overman and Thomas, 1995) and a comprehensive vibrational assessment of the phenolic moiety (Arp et al., 2001), it has now been established that the “singlet-like” tyrosyl signature in Raman spectra of filamentous viruses is diagnostic of the non-hydrogenbonded state of the phenoxyl group. Thus, in the absence of hydrogen bonding, the Raman intensity of the higher wavenumber component of the canonical Fermi doublet is greatly enhanced, such that I2/I1 = 6.8. The feeble Raman intensity of the lower wavenumber member of the Fermi doublet (I1) gives the appearance of an intense singlet at the spectral position of the higher wavenumber member (I2). This finding extends and refines the earlier correlation of Siamwiza and co-workers (1975) and profoundly impacts the use of tyrosyl Raman signatures as diagnostic of phenoxyl group hydrogen bonding in proteins. Structural Biology
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hydrophobic bonding/DNA base stacking
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Figure 17.8.16 Raman difference spectra reflecting DNA reorganizations due to different DNA binding proteins. From top-to-bottom, bZIP protein GCN4 binding to an AP-1 target site; ssDNA binding protein of M13 phage (gVp) binding to the ssDNA analogue, poly(dA); cI repressor of phage λ binding to its OL1 target site; human sex-determining factor hSRY-HMG box binding to its DNA target site. Ordinates of the difference spectra have been scaled to reflect the same 1090 cm−1 intensity in the parent complex. Labeling of boxed segments refers to bands of DNA. Data are from Benevides et al., 2000b.
Raman Spectroscopy of Proteins
Raman spectroscopy of viruses and viral assemblies has also suggested new spectra-structure correlations for the indolyl moiety of tryptophan (Wen and Thomas, 2000) and provided a new understanding of the extraordinary range of hydrogen-bonding interactions accessible to the sulfhydryl group of cysteine (Raso et al., 2001). In addition, Raman spectroscopy of isotope-edited viruses has furnished novel approaches to assessing α-helical secondary structure in proteins (Tsuboi et al., 2000) and identifying specific spectral contributions of the peptide main chain (Overman and Thomas, 1998a), aromatic side chains (Aubrey and Thomas, 1991; Overman and Thomas, 1995) and a variety of nonaromatic side chains (Overman and Thomas, 1999).
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E fd(3Fd5)
Raman intensity
D fd(2Yd4)
C fl(Y24M)
B fl(Y21M)
A fd cm–1
Figure 17.8.17 Raman spectra in the region 600-900 cm−1 of isotopic and mutant variants of the Ff filamentous virus demonstrating assignment of the prominent bands near 853 cm−1 to tyrosine and those near 827 cm−1 to phenylalanine. From bottom to top, wild-type unlabeled fd virus (A), mutant f1 virus in which Met replaces Tyr 21 (B), mutant f1 virus in which Met replaces Tyr 24 (C), wild-type fd in which phenolic rings of Tyr 21 and Tyr 24 are deuterated (D), and wild-type fd in which phenyl rings of Phe 11, Phe 42, and Phe 45 are deuterated (E). Note that the Raman bands of the two mutants, at 855 (B) and 851 cm−1 (C), respectively, are about half as intense as their composite at 853 cm−1 in the wild-type virus (A). Because the 827 cm−1 band is virtually completely removed by phenylalanine deuteration (E), it is assigned to Phe not Tyr. The 853 → 817 cm−1 deuteration shift and the absence of an 827 cm−1 deuteration shift (D) establish the singlet-like tyrosine markers for Tyr 21 and Tyr 24. Further details are given elsewhere (Overman et al., 1994; Overman and Thomas, 1995; Arp et al., 2001). Adapted with permission from Arp et al., (2001).
LITERATURE CITED Allemann, R.K. and Egli, M. 1997. DNA recognition and bending. Chem. Biol. 4:643-650. Arp, Z., Autrey, D., Laane, J., Overman, S.A., and Thomas, G.J. Jr. 2001. Tyrosine Raman signatures of the filamentous virus Ff are diagnostic of non-hydrogen-bonded phenoxyls: Demonstration by Raman and infrared spectroscopy of p-cresol vapor. Biochemistry 40:2522-2529. Asher, S.A., Bormett, R.W., Chen, X.G., Lemmon, D.H., Cho, N., Peterson, P., Arrigoni, M., Spinelli, L., and Cannon, J. 1993. UV resonance Raman spectroscopy using a new CW laser source: Convenience and experimental simplicity. Appl. Spectrosc. 47:628-633.
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Thomas, G.J., Jr., Prescott, B., Opella, S.J., and Day, L.A. 1988. Sugar pucker and phosphodiester conformations in viral genomes of filamentous bacteriophages: fd, If1, IKe, Pf1, Xf and Pf3. Biochemistry 27:4350-4357. Thomas, G.J., Jr., Becka, R., Sargent, D., Yu, M.-H., and King, J. 1990. Conformational stability of P22 tailspike proteins carrying temperature-sensitive-folding mutations. Biochemistry 29:4181-4187. Thomas, G.J., Jr., Benevides, J.M., Overman, S.A., Ueda, T., Ushizawa, K., Saitoh, M., and Tsuboi, M. 1995. Polarized Raman spectra of oriented fibers of A DNA and B DNA: Anisotropic and isotropic local Raman tensors of base and backbone vibrations. Biophys. J. 68:1073-1088. Tsuboi, M. and Thomas, G.J. Jr. 1997. Raman scattering tensors in biological molecules and their assemblies. Appl. Spectrosc. Revs. 32:263-299. Tsuboi, M., Nishimura, Y., and Hirakawa, A.Y. 1987. Resonance Raman spectroscopy and normal modes of the nucleic acid bases. In Biological Applications of Raman Spectroscopy. Vol. 2: Resonance Raman Spectra of Polyenes and Aromatics (T.G. Spiro, ed.) pp. 109-179. John Wiley & Sons. New York. Tsuboi, M., Ueda, T., Ushizawa, K., Ezaki, Y., Overman, S.A., and Thomas, G.J. Jr. 1996a. Raman tensors for the tryptophan side chain in proteins determined by polarized Raman microspectroscopy of oriented N-acetyl-L-tryptophan crystals. J. Mol. Struct. 379:43-50. Tsuboi, M., Overman, S.A., and Thomas, G.J. Jr. 1996b. Orientation of tryptophan-26 in coat protein subunits of the filamentous virus Ff by polarized Raman microspectroscopy. Biochemistry 35:10403-10410. Tsuboi, M., Suzuki, M., Overman, S.A., and Thomas, G.J. Jr. 2000. Intensity of the polarized Raman band at 1340-1345 cm−1 as an indicator of protein α-helix orientation: Application to Pf1 filamentous virus. Biochemistry 39:2677-2684. Tsuboi, M., Ushizawa, K., Nakamura, K., Benevides, J.M., Overman, S.A., and Thomas, G.J. Jr. 2001a. Orientations of Tyr 21 and Tyr 24 in the capsid of filamentous virus Ff determined by polarized Raman spectroscopy. Biochemistry 40:1238-1247. Tsuboi, M., Aida, M., Nakamura, K., Bondre, P., Overman, S.A., Benevides, J.M., and Thomas, G.J. Jr. 2001b. Raman tensors of arginine: A basis for determining orientations of arginine side chains in filamentous viruses (Pf1 and Pf3) by polarized Raman spectroscopy. Biophys. J. 80:397a. Tsuboi, M., Kubo, Y., Ikeda, T., Overman, S.A., Osman, O., and Thomas, G.J. Jr. 2003a. Protein and DNA residue orientations in the filamentous virus Pf1 determined by polarized Raman and polarized FTIR spectroscopy. Biochemistry 42:940-950. Tsuboi, M., Overman, S.A., Nakamura, K., Rodriguez-Casado, A., and Thomas, G.J. Jr. 2003b. Orientation and interactions of an essential tryptophan (Trp-38) in the capsid subunit of Pf3 filamentous virus. Biophys. J. 34:1-8. Tuma, R. and Thomas, G.J. Jr. 1997. Mechanisms of virus assembly probed by Raman spectroscopy: The icosahedral bacteriophage P22. Biophys. Chem. 68:17-31. Tuma, R. and Thomas, G.J. Jr. 2002. Raman spectroscopy of viruses. In handbook of vibrational spectroscopy. Vol. 5. Applications in Life, Pharmaceutical and Natural Sciences (J.M. Chalmers and P.R. Griffith, eds.) pp. 3519-3535. John Wiley & Sons, Chichester, U.K. Tuma, R., Vohník, S., Li, H., and Thomas, G.J. Jr. 1993. Cysteine conformation and sulfhydryl interactions in proteins and viruses. 3. Quantitative measurement of the Raman S-H band intensity and frequency. Biophys. J. 65:1066-1072. Tuma, R., Bamford, J.K.H., Bamford, D.H., Russell, M.P., and Thomas, G.J. Jr. 1996a. Structure, interactions and dynamics of PRD1 virus. I. Coupling of subunit folding and capsid assembly. J. Mol. Biol. 257:87-101. Tuma, R., Bamford, J.K.H., Bamford, D.H., and Thomas, G.J. Jr. 1996b. Structure, interactions and dynamics of PRD1 virus. II. Organization of the viral membrane and DNA. J. Mol. Biol. 257:102-115. Tuma, R., Prevelige, P.E., Jr., and Thomas, G.J. Jr. 1998a. Mechanism of capsid maturation in a doublestranded DNA virus. Proc. Natl. Acad. Sci. U.S.A. 95:9885-9890. Tuma, R., Parker, M.H., Weigele, P., Sampson, L., Sun, Y., Rama Krishna, N., Casjens, S.R., Thomas, G.J., Jr., and Prevelige, P.E. Jr. 1998b. A helical coat protein recognition domain of the bacteriophage P22 scaffolding protein. J. Mol. Biol. 281:81-94. Tuma, R., Tsuruta, H., Benevides, J.M., Prevelige, P.E., Jr., and Thomas, G.J., Jr. 2001. Characterization of subunit structural changes accompanying assembly of the bacteriophage P22 procapsid. Biochemistry 40:665-674. Valpuesta, J.M. and Carrascosa, J.L. 1994. Structure of viral connectors and their function in bacteriophage assembly and DNA packaging. Q. Rev. Biophys 27:107-155. Valpuesta, J.M., Fernandez, J.J., Carazo, J.M., and Carrascosa, J.L. 1999. The three-dimensional structure of a DNA translocating machine at 10 Å resolution. Structure Fold Des. 7:289-96. Raman Spectroscopy of Proteins
Wang, Y., Purrello, R., Jordan, T., and Spiro, T.G. 1991. UVRR spectroscopy of the peptide bond. J. Am. Chem. Soc. 113:6359-6368.
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Welsh, L.C., Symmons, M.F., Sturtevant, J.M., Marvin, D.A., and Perham, R.N. 1998. Structure of the capsid of Pf3 filamentous phage determined from X-Ray fibre diffraction data at 3.1 Å resolution. J. Mol. Biol. 283:155-177. Welsh, L.C., Symmons, M.F., and Marvin, D.A. 2000. The molecular structure and structural transition of the alpha-helical capsid in filamentous bacteriophage Pf1. Acta Crystallogr. D. Biol. Crystallogr. 56:137150. Wen, Z.Q. and Thomas, G.J., Jr. 2000. Ultraviolet-resonance Raman spectroscopy of the filamentous virus Pf3: Interactions of Trp 38 specific to the assembled virion subunit. Biochemistry 39:146-152. Wen, Z.Q., Overman, S.A., and Thomas, G.J., Jr. 1997. Structure and interactions of the single-stranded DNA genome of filamentous virus fd: Investigation by ultraviolet resonance Raman spectroscopy. Biochemistry 36:7810-7820. Wen, Z.Q., Armstrong, A., and Thomas, G.J., Jr. 1999. Demonstration by ultraviolet resonance Raman spectroscopy of differences in DNA organization and interactions in filamentous viruses Pf1 and fd. Biochemistry 38:3148-3156. Wen, Z.Q., Overman, S.A., Bondre, P., and Thomas, G.J., Jr. 2001. Structure and organization of bacteriophage Pf3 probed by Raman and ultraviolet resonance Raman spectroscopy. Biochemistry 40:449-458. Yu, T.-J., Lippert, J.L., and Peticolas, W.L. 1973. Laser Raman studies of conformational variations of poly-L-lysine. Biopolymers 12:2161-2176.
James M. Benevides, Stacy A. Overman, and George J. Thomas, Jr. School of Biological Sciences University of Missouri-Kansas City Kansas City, Missouri
Structural Biology
17.8.35 Current Protocols in Protein Science
Supplement 33
Crystallization of Integral Membrane Proteins
UNIT 17.9
X-ray crystallography has become a very powerful tool for determining the structures of integral membrane proteins, with >120 unique membrane protein structures solved to date (http://blanco.biomol.uci.edu/Membrane Proteins xtal.html). The major bottlenecks in this field are the expression of sufficient quantities of functional membrane proteins and the growth of well ordered, three-dimensional crystals for X-ray analysis. Although it is necessary to screen more parameters to obtain good membrane protein crystals than for soluble protein crystals, the approach is systematic and benefits from the availability of a wide variety of homogeneous detergents, the implementation of nanoliter pipetting robots, and the use of numerous 96-condition screens. This unit illustrates the steps required for crystallization of a bacterial outer membrane protein starting from a purified sample in detergent. For a description of expression and purification procedures used to isolate an E. coli outer membrane protein, see Yue et al. (2003). Crystallization procedures will vary depending on the membrane protein of interest, but the procedures described in this unit are generally applicable to inner membrane proteins as well. Support Protocol 1 describes the exchange of the purified membrane protein into a detergent solution suitable for crystallization, using size-exclusion chromatography. Support Protocol 2 describes preparation of the sample for crystallization, while Basic Protocols 1 to 3 describe crystallization procedures to identify and optimize conditions for the production of highly ordered membrane protein crystals.
ROBOTIC SCREENING OF CRYSTALLIZATION CONDITIONS For robotic screening, the hanging drop method is preferred over the sitting drop method because small (0.4 µl), detergent-containing sitting drops tend to adhere to the side of the sitting drop well, making visualization much more difficult.
BASIC PROTOCOL 1
Materials 96-condition screen (Table 17.9.1) 96-well U-bottom microplates (Greiner Bio-One) 8- or 12-channel multi-channel pipet Nanoliter pipetting robot (e.g., Mosquito, TTP LabTech) 96-well format crystallization sheets (e.g., CrystalClene Sheets HT, Molecular Dimensions) Constant temperature incubators (e.g., Rumed, Molecular Dimensions) 1. Prepare the screens by transferring 50 µl of each well solution from a 96-condition commercial screen to a 96-well plate using a multi-channel pipet. To prevent dust from entering the wells and to minimize evaporation, stack the finished plates, placing an empty plate on top. These plates should be used on the day they are prepared.
2. Initialize and calibrate the nanoliter pipetting robot according to the manufacturer’s instructions. The authors use a dispensing program that pipets 0.2 µl of protein onto all 96 positions of a CrystalClene sheet, and then pipets 0.2 µl of well solution onto the corresponding protein drops. Structural Biology Contributed by Travis J. Barnard, Jeremy L. Wally, and Susan K. Buchanan Current Protocols in Protein Science (2007) 17.9.1-17.9.15 C 2007 by John Wiley & Sons, Inc. Copyright
17.9.1 Supplement 47
Table 17.9.1 Suggested 96-Condition Screens for Initial Crystallization Trials of Membrane Proteinsa
Hampton Research Crystal Screen HT Index HT Salt Rx HT Nextal Biotechnologies The Classics Lite The PEGs The AmSO4 The Anions The Cations The MPDs The pHClear The MbClass Suite The MbClass II Suite Molecular Dimensions Memstart/MemSys Emerald Biosystems Wizard Bloc (I&II) Cryo Bloc (I&II) OZMA 1-4 OZMA 8-10 a These screens are listed by manufacturer in random order.
Many robots do not have the capability to pipet hanging drops as described above, but instead are designed to pipet sitting drops. In this case, sitting drops can be used with alternative microplates and coversheets described in the Alternate Protocol, using a robot to pipet 0.2 µl protein plus 0.2 µl well solution.
3. Once the robot has finished the pipetting program, carefully apply the CrystalClene sheet to the 96-well microplate so the drops are centered over the wells. Firmly smooth the coversheet over the plate to seal the wells. Remove the protective backing from the cover sheet so the drops can be visualized. The Mosquito robot comes with a convenient adaptor that facilitates centering the coversheet over the plate. The plate is now complete.
4. Repeat the pipetting program and step 3 for the remaining plates. Incubate the plates at the desired temperature(s), usually 21◦ C and/or 4◦ C. If the membrane protein is known to be unstable at room temperature, it may be advisable to incubate the plates at 4◦ C; however, crystallization reagents sometimes stabilize membrane proteins, such that samples requiring purification at 4◦ C may still form crystals at 21◦ C. These two temperatures have yielded the most crystals of all types, but other temperatures may also be tried.
Crystallization of Integral Membrane Proteins
With either Basic Protocol 1 or the Alternate Protocol, the plates should be evaluated after 2 days, again after 1 week, and finally after 2 weeks. The drops usually reach equilibrium within 1 week, therefore if no crystals are observed after 2 weeks, it is advisable to discard the plates. If crystals are obtained, the conditions should then be scaled up and optimized in 24-well plates using larger volumes, as described in Basic Protocols 2 and 3. If no crystals are obtained, the following modifications can be made to the initial 96-condition screens. (1) Adjust the protein concentration. If the majority of
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the drops are still clear after 2 weeks, try a higher protein concentration. If the majority have precipitated (characterized by diffuse, amorphous material in the drop), then try lowering the protein concentration. (2) Adjust the incubation temperature. If no crystals are obtained, try 15◦ C or 10◦ C. Also try higher temperatures, such as 25◦ C or 30◦ C, as it is unclear how temperature will affect the solubility of a protein and/or its propensity to aggregate. Note that the critical micelle concentration (cmc) of the detergent will also change with temperature, affecting crystallization results. (3) Change the detergent. The authors routinely screen new membrane proteins in every detergent in which they are stable (with reference to Table 17.9.4), and this large-scale screening is done early in a project. All 17 kits are screened for a single membrane protein in each detergent being tested to identify detergents that may yield useful membrane protein crystals. (4) If all else fails, try multiple homologs (Chang et al., 1998), a different construct (Locher et al., 2002), or complexing the membrane protein to a soluble protein (such as a natural ligand or protein partner, or an antibody fragment; Dutzler et al., 2003). The large detergent micelle surrounding a membrane protein may make crystallization difficult, so having a soluble protein bound to it may provide additional crystal contacts by expanding the hydrophilic surface.
MANUAL SCREENING OF CRYSTALLIZATION CONDITIONS If a nanoliter pipetting robot is not available, the same 96-condition kits can be screened manually, using 96-well sitting-drop plates. This method is very fast but requires significantly more protein because 1-µl drops of membrane protein are mixed with 1-µl drops of well solution. To screen all 17 kits as described in Basic Protocol 1, at least 1700 µl of concentrated (10 mg/ml) membrane protein sample is required.
ALTERNATE PROTOCOL
Materials 96-condition commercial screens (Table 17.9.1) 96-well conical sitting-drop microplates (Corning/Hampton Research) 8- or 12-channel multi-channel pipet (50-µl volume) 8- or 12-channel multi-channel pipet (1-µl volume) 1-µl repeater pipet ClearSeal Film (Hampton Research) Microplate centrifuge Constant temperature incubators (e.g., Rumed, Molecular Dimensions) 1. Prepare the screens by transferring 50 µl of each well solution from a 96-condition commercial screen to a 96-well sitting drop plate using a 50-µl volume 8- or 12-channel multi-channel pipet. 2. Using a 1-µl multichannel pipet, pipet 1 µl from each well into each sitting drop sub-well (small depression where the sample drop will reside). 3. Using the 1-µl repeater pipet, quickly pipet 1 µl of concentrated membrane protein sample into each of the 96 sitting-drop sub-wells. To avoid cross-contamination, do not mix the droplets at this point; centrifugation of the microplates will ensure mixing.
4. Cover the microplate with a ClearSeal film, using light pressure to seal it. 5. Centrifuge the microplate according to manufacturer’s recommendations for 2 min to merge precipitant and protein droplets. 6. Repeat steps 1 to 5 for all other 96-condition kits, as appropriate. 7. Incubate the plates at the desired temperature(s), usually 21◦ C and/or 4◦ C. See annotation of Basic Protocol 1, step 4, for guidance on optimization of conditions.
Structural Biology
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BASIC PROTOCOL 2
PREPARATION OF 24-WELL PLATES FOR OPTIMIZATION OF CRYSTALLIZATION CONDITIONS Once promising crystallization conditions have been identified using the screens described in Basic Protocol 1 or the Alternate Protocol, these conditions are repeated using larger drop volumes and expanded into finely sampled grid screens, usually varying two parameters (such as precipitant concentration and pH) on a given 24-well plate. This protocol describes how to set up 24-well plates in the context of working with membrane proteins in detergent.
Materials 10× salt stock solutions 10× buffer stock solution Precipitant stock solutions (concentrations will vary) Cryoprotectants (e.g., glycerol), optional 24-well plates (e.g., VDX plates with sealant, Hampton Research) Repeater pipet (optional) Orbital platform shaker (optional) 22-mm siliconized square cover slides (Hampton Research) Constant temperature incubators (e.g., Rumed, Molecular Dimensions) 1. Prepare the well solutions. Sequentially add the 10× salt solution, 10× buffer solution, and precipitant stock solutions to each well of a 24-well plate to attain a final volume of 1.0 ml in each well. If a cryoprotectant, such as glycerol, is to be added, include it in the precipitant solution. It is not necessary to add detergent to the well solutions, because the protein contains detergent at a concentration of at least twice the cmc so that dilution with an equal amount of precipitant will still result in sufficient detergent to maintain the protein in solution. All solutions or their components should be made with the highest quality water available and passed through 0.22-µm filters to remove particulates. Hampton Research (as well as many other companies) provides many salt, buffer, and precipitant stock solutions that are ready-to-use. Plates should be set up in a clean area as free of dust as possible. Normally, 0.1 ml of the salt and buffer stock solutions is added per well, which has a total volume of 1 ml. The precipitant stock solutions (a separate solution is made for each precipitant concentration) are made such that 0.8 ml can be added per well to yield the correct final concentration. If necessary, cryoprotectant is added to the precipitant stock solutions to help prevent damage to the crystals during flash-freezing in liquid nitrogen or liquid nitrogen–cooled propane. The authors normally use glycerol as a cryoprotectant at a final well concentration of 5% to 15% (v/v). Since these components are often very viscous, quantities are measured by weight according to density and not by volumetric pipetting. A repeater pipet will greatly reduce the time necessary to dispense the stock solutions into individual wells. In addition, these pipets operate using positive displacement, handling viscous solutions better than traditional pipets. It is very important that the preparation procedure for the well solutions be reproducible, because if crystals grow, more wells of the same condition will need to be prepared.
2. Mix the well solutions on an orbital platform shaker for at least 1 hr.
Crystallization of Integral Membrane Proteins
Shake the well solutions as vigorously as possible without spillage. It is important that the well solutions are thoroughly mixed for reproducible results. This can be difficult for highly concentrated PEG solutions. Check for proper mixing by stirring a well containing the highest precipitant concentration. If no Schlieren effects are observed, the plates are ready. The plates can be wrapped in plastic wrap and stored overnight at 4◦ C or used immediately. If a suitable orbital shaker is not available, each well can be mixed by stirring with a pipet tip.
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3. Set up the 22-mm siliconized square cover slides on a clean, lint-free surface. Place 1.0 µl of protein on a siliconized cover slide, then add 1.0 µl of well solution to each protein drop, pipetting up and down four times for each drop to ensure proper mixing. Avoid introducing air bubbles and try to make the drops as hemispherical as possible. Change tips between each well. Place the cover slide over the appropriate well and press down to seal. If only one drop per slide is planned, six slides (one row of a VDX plate) can be prepared at a time. Care should be taken to work as quickly as possible to avoid evaporation. The detergent (some more than others) will cause the drop to spread on the slide. As a result, cover slides should always be siliconized to keep the surface area of the drop minimized. In addition, it is not uncommon to find batches of poorly siliconized cover slides as evidenced by drop spreading. These slides should not be used for membrane proteins. Place the slides over the appropriate wells and gently push them down to seal. Care should be taken not to get grease near the center of the slide as this will complicate visualization of the drop.
4. Once the plate is complete, incubate it at a constant temperature in an environment as free of vibrations as possible.
REFINEMENT OF INITIAL CRYSTALLIZATION CONDITIONS IN 24-WELL FORMAT
BASIC PROTOCOL 3
This protocol describes a 24-well format to refine initial crystallization conditions from commercial screens. In this protocol, precipitant concentration and pH are optimized because these parameters greatly affect crystal formation and growth and are good starting points for refinement. However, there are many other parameters that affect crystal growth (Table 17.9.2) and these often need to be optimized as well. Conditions should be refined for each crystal type observed because the type that diffracts best will not be known until diffraction data are collected. Thus, during initial optimization, crystals are judged by their appearance. Once conditions that produce crystals are identified, the Hampton Additive Screen (Table 17.9.3) can be used to identify small molecules that may improve crystal growth. Finally, for the best conditions and additives, multiple drops are prepared and the plates are left undisturbed until the crystals have reached their optimum size. Table 17.9.2 Variables Affecting Membrane Protein Crystal Growtha
Protein primary sequence and conformational flexibility Protein purity, stability, and concentration Temperature and pH Precipitant type and concentration Detergents, additives, and lipids Specific ions and ionic strength Presence or absence of ligand Presence or absence of affinity tags a All variables may substantially affect crystallization, so equal consideration should be
given to each item.
Structural Biology
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Table 17.9.3 Additive Screen Reagent Formulation (Hampton Research)
Crystallization of Integral Membrane Proteins
1
0.1 M barium chloride
42
0.1 M trimethylamine HCl
2
0.1 M cadmium chloride
43
1 M guanidine HCl
3
0.1 M calcium chloride
44
0.1 M urea
4
0.1 M cobaltous chloride
45
0.1 M nicotinamide adenine dinucleotide
5
0.1 M cupric chloride
46
0.1 M adenosine-5-triphosphate disodium salt
6
0.1 M magnesium chloride
47
0.1 M TCEP hydrochloride
7
0.1 M manganese (II) chloride
48
0.1 M L-cysteine
8
0.1 M strontium chloride
49
0.1 M EDTA sodium salt
9
0.1 M yttrium chloride
50
5% (w/v) polyvinylpyrrolidone K15
10
0.1 M zinc chloride
51
30% (w/v) dextran sulfate
11
0.1 M ferric (III) chloride
52
40% (v/v) pentaerythritol ethoxylate
12.
0.1 M nickel (II) chloride
53
10% (w/v) polyethylene glycol 3350
13
0.1 M chromium (III) chloride
54
30% (w/v) D(+)-glucose
14
0.1 M praseodymium (III) acetate
55
30% (w/v) D (+)-sucrose
15
1 M ammonium sulfate
56
30% (w/v) xylitol
16
1 M potassium chloride
57
30% (w/v) sorbitol
17
1 M lithium chloride
58
12% (w/v) inositol
18
2 M sodium chloride
59
30% (w/v) D-trehalose
19
0.5 M sodium fluoride
60.
30% (w/v) D-galactose
20
1 M sodium iodide
61
30% (v/v) ethylene glycol
21
2 M sodium thiocyanate
62
30% (v/v) glycerol
22
1 M potassium sodium tartrate
63
3 M NDSB-195
23
1 M tri-sodium citrate
64
2 M NDSB-201
24
1 M cesium chloride
65
2 M NDSB-211
25
1 M sodium malonate pH 7.0
66
2 M NDSB-221
26
0.1 M L-proline
67
2 M NDSB-256
27
0.1 M phenol
68
15% (w/v) 1,2,3-heptanetriol
28
30% (v/v) dimethyl sulfoxide
69
20% (w/v) benzamidine HCl
29
0.1 M sodium bromide
70
5% (w/v) LDAO
30
30% (w/v) 6-aminocaproic acid
71
5% (w/v) n-octyl-b-D-glucoside
31
30% (w/v) 1,5-diaminopentane di-HCl
72
5% (w/v) n-dodecyl-b-D-maltoside
32
30% (w/v) 1,6-diaminohexane
73
30% (w/v) trimethylamine N-oxide
33
30% (w/v) 1,8-diaminooctane
74
30% (w/v) 1,6 hexanediol
34
1 M glycine
75
30% (v/v) 2-methyl-2,4-pentanediol
35
0.3 M glycyl-glycyl-glycine
76
50% (w/v) polyethylene glycol 400
36.
0.1 M taurine
77
50% (v/v) jeffamine M-600 pH 7.0
37
0.1 M betaine hydrochloride
78
40% (v/v) 2,5 hexanediol
38
0.1 M spermidine
79
40% (v/v) (+)-1,3 butanediol
39
0.1 M spermine tetra-HCl
80
40% (v/v) polypropylene glycol P 400
40
0.1 M hexamine cobalt (III) chloride
81
30% (v/v) dioxane
41
0.1 M sarcosine
82
30% (v/v) ethanol
17.9.6
continued
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Table 17.9.3 Additive Screen Reagent Formulation (Hampton Research), continued
83
30% (v/v) isopropanol
90
40% (v/v) n-propanol
84
30% (v/v) methanol
91
5% (v/v) ethyl acetate
85
40% (v/v) 1,4 butanediol
92
40% (v/v) acetone
86
40% (v/v) tert-butanol
93
0.25% (v/v) dichloromethane
87
40% (v/v) 1,3 propanediol
94
7% (v/v) n-butanol
88
40% (v/v) acetonitrile
95
40% (v/v) 2,2,2-trifluoroethanol
89
40% (v/v) g-butyrolactone
96
40% (v/v) hexafluoro-2-propanol
Materials Commercial screens with crystal formation (see Basic Protocol 1 or Alternate Protocol) 10× salt stock solutions 10× buffer stock solution Precipitant stock solutions (concentrations will vary) 24-well plates (e.g., VDX plates with sealant, Hampton Research) Light microscope (4× to 20× magnification) Hampton Additive Screen (optional) 22-mm siliconized square cover slides (Hampton Research) Constant temperature incubators (e.g., Rumed, Molecular Dimensions) 1. From the results obtained using Basic Protocol 1 or Alternate Protocol, select the conditions that produced crystals or crystalline precipitate as a guide for setting up the 24-well plates. If no promising conditions were observed, it may be necessary to modify the protein through mutagenesis or to try homologs. If small needles or plates were observed, switching the detergent may produce high quality three-dimensional crystals. For one outer membrane protein, the authors observed needles in N,N-dimethyldodecylamine-N-oxide, thin plates in octyl tetraoxyethylene, and three-dimensional crystals in octyl-β-D-glucoside. Thus, detergent screening is a good approach to try early in the process, especially if initial optimization attempts only yield needles or thin plates.
2. From the chosen conditions, set up plates that screen a range of precipitant concentration and pH. Observe each plate directly after it is set up to ensure that all the drops are not precipitated. If all are precipitated, set the plate up again using a lower precipitant concentration range. The goal is to have clear drops at the lowest precipitant concentration and fully precipitated drops at the highest concentration, once the plate has reached equilibrium. For example, in a commercial screen, small crystals were observed in 30% PEG 1000, 200 mM NaCl, and 0.1 M Bis-Tris, pH 6.5. Using these conditions as a guide, the plates shown in Figure 17.9.1A were set up with the same protein concentration and incubated at the same temperature as the commercial screen. For the conditions at pH 6.5, two drops were set up per slide: control drops containing buffer (including detergent and any additives) without protein and drops containing buffer with protein. If no crystals appear in the control drops, this is a good indication, but not absolute proof, that the crystals obtained in sample drops are protein. In this example, no cryoprotectant is present (PEG 1000 is not a good cryoprotectant, whereas PEG 400, glycerol, and some other reagents offer significant cryoprotection; Garman and Doublie, 2003). Thus, if there is sufficient protein, the plates in Figure 17.9.1A
Structural Biology
17.9.7 Current Protocols in Protein Science
Supplement 47
Figure 17.9.1 Optimization of pH and precipitant concentration. (A) Initial crystal optimization using three 24-well plates, with the top plate containing pH 3.5 to 5.0, the middle plate containing pH 5.5 to 7.0, and the bottom plate containing pH 7.5 to 9.0. Each plate uses the same six precipitant concentrations. The best condition in this example is marked ‘X’. (B) Two 24-well plates are designed using the best conditions found in A.
would be set up in duplicate, adding glycerol to all of the well solutions of one set to a final concentration of 10% (v/v). If the presence of glycerol did not adversely affect crystal formation, it would be included in all well solutions throughout the optimization process and, in subsequent purifications, the buffer used for size-exclusion chromatography in Support Protocol 1. Including glycerol in the size-exclusion buffer simplifies setting up plates that have equal concentrations of glycerol in the drop and the well. This can be important because glycerol gradients can significantly affect the rate of equilibration. Crystallization of Integral Membrane Proteins
Sometimes, crystals obtained from commercial screens are not reproducible in 24-well plates. This is likely due to different physical and chemical conditions between the 96-well and 24-well formats. Samples in 96-well plates equilibrate much more rapidly than in 24-well plates, due to the smaller volumes used.
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3. Score plates from step 2 and refine the best conditions. Examine the plates under a microscope every 24 hr for the first few days after they are set up and then every other week for 2 to 3 months. Scoring plates is described on the Hampton Research Web site (http://www.hamptonresearch.com/) and by McPherson (1999). Briefly, indicate whether a drop is clear, contains some sort of precipitate, or contains crystals. If a precipitate is discrete and granular, it should be examined under high magnification for the presence of microcrystals. For drops containing crystals, indicate their general shapes and sizes. Often, different crystal morphologies are seen at different pH values. Optimize conditions for each crystal type because the best looking crystals do not always diffract strongly or to high resolution.
4. Further refine precipitant concentration and pH (or other variables) to optimize crystal growth. In the example illustrated in Figure 17.9.1, only one crystal type was found for the plates set up in Figure 17.9.1A and the best condition contained 28% PEG 1000 and 10% glycerol at pH 5.0. Figure 17.9.1B shows how these conditions would be further refined by more finely sampling precipitant and pH conditions.
5. (Optional) Select a single (reproducible) crystallization condition and multiply the precipitant and salt concentrations by 1.1 (to account for subsequent dilution with the additive). Set up four 24-well plates with this modified condition using the 96-condition Hampton Research Additive Screen according to manufacturer’s instructions. Pipet 0.9 µl protein onto a siliconized cover slide, add 0.2 µl additive, then add 0.9 µl well solution. Repeat the Additive Screen plates for other optimized crystallization conditions as needed. This screen contains a variety of small molecules that can affect membrane protein crystallization (Table 17.9.3). Additives are often used in combination with one or more detergents to optimize the size of the detergent micelle for crystallization (Table 17.9.4). If well-formed, single crystals are obtained using the additive screen, re-optimize, beginning at step 2 and include the additive(s) for all conditions.
6. Score the optimized plates from steps 4 and 5. Set up the best conditions multiple times to maximize the number of large, single crystals. For conditions that give large, well-formed crystals, set up at least four wells (one column of a VDX plate) with three drops per cover slide. Not all of the drops will behave identically due to random pipetting errors, but a large number of crystals suitable for X-ray diffraction should grow. In addition, leave these optimized plates undisturbed in the incubator for the amount of time it takes for crystals to grow to their optimum size. This time frame can be approximated from the behavior of previous plates. At this stage, excessive handling of the plates should be avoided because sudden temperature changes, shocks, or vibrations can induce nucleation, giving rise to showers of small crystals. The goal now is to limit nucleation and maximize crystal growth. Finally, in this example, the path to macroscopic membrane protein crystals required only optimizing the precipitant concentration and the pH. While these are important parameters, there are many more conditions that can and usually must be optimized (Table 17.9.2). See McPherson (1991), Michel (1991), McPherson (1999), and Iwata (2003) for a more comprehensive list of parameters that affect both soluble and membrane protein crystal growth, as well as a more complete discussion of crystal growth refinement.
DETERGENT EXCHANGE BY SIZE-EXCLUSION CHROMATOGRAPHY The starting membrane protein must be purified in a detergent in which it is stable and active, such as dodecyl maltoside (DDM). If possible, the protein should show no contaminants as judged by SDS-PAGE (UNIT 10.1). Because affinity tags used for purification could either help or hinder crystallization, they can be removed prior to detergent exchange if desired. The starting protein concentration is not important as long as the
SUPPORT PROTOCOL 1
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total volume contains ≥1 mg of protein, but the sample must be concentrated to ≤5 ml for the chromatography column described in this protocol. The sample should be eluted from the size-exclusion column at a low flow rate for optimal detergent exchange to occur. An analysis of the peak shape gives an indication of the homogeneity of the sample, as well as separating different oligomeric states and removing large aggregates. A homogeneous sample will exhibit a symmetrical, Gaussian peak shape, whereas an unhomogeneous sample may display tailing on the leading or trailing side of the peak (making the peak asymmetric) or multiple peaks, reflecting the presence of higher molecular weight aggregates or multiple oligomeric states, such as monomers, dimers, and tetramers.
Materials Column buffer (see recipe) Purified membrane protein sample (≥1 mg) Sephacryl S300 HR 16/60 size exclusion column (GE Healthcare) ¨ Akta prime chromatography system (GE Healthcare) or equivalent 15-ml Centriprep concentrators (Millipore) 0.22-µm syringe filter 1. Attach a Sephacryl S300 HR 16/60 size exclusion column to the chromatography system according to the manufacturer’s instructions. Equilibrate the column at a flow rate of 0.5 ml/min with 240 ml (two column volumes) of column buffer. 2. Place up to 15 ml purified membrane protein in a 15-ml Centriprep concentrator, choosing the molecular weight cut-off of the ultrafiltration membrane according to the size of the protein. Centrifuge at 4◦ C according to manufacturer’s instructions. Refill the concentrator if necessary and repeat until the sample volume measures 3 to 5 ml. Filter the concentrated sample using a 0.22-µm syringe filter. It may be necessary to use a smaller molecular weight cut-off Centriprep membrane to avoid loss of sample. Additional information on choosing ultrafiltration devices of the appropriate molecular weight cut-off can be found in Critical Parameters and Troubleshooting.
3. Apply the membrane protein sample to the equilibrated Sephacryl column and elute the sample at a rate of 0.3 ml/min. If the sample elutes as a single, Gaussian peak, determine the protein concentration of the pooled peak fractions (UNITS 3.1 & 3.4). If the peak shape is not Gaussian, a decision must be made to either take fractions from the major species (avoiding leading or trailing peak shoulders) or to discard the sample entirely.
4. Repeat steps 1 to 3 for new protein samples in each detergent that may be useful for crystallization. Any of the detergents in Table 17.9.4 (and possibly other homogenous, monodisperse detergents) yielding a Gaussian peak shape for the protein of interest is a candidate for crystallization (Lemieux et al., 2003).
Crystallization of Integral Membrane Proteins
While dynamic light scattering (UNIT 7.8) is sometimes used to assess the monodispersity of soluble protein samples, this technique is difficult to use for membrane proteins because the detergent micelle (in which the membrane protein is incorporated) contributes to light scattering and may itself be polydisperse. For this reason, the authors assess approximate monodispersity by size exclusion peak shape. Analytical ultracentrifugation (UNIT 7.5) can also be used in some cases, if care is taken to match the buoyant density of the detergent (Lebowitz et al., 2002).
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PREPARATION OF MEMBRANE PROTEIN SAMPLE FOR CRYSTALLIZATION
SUPPORT PROTOCOL 2
Once a purified protein solution containing the desired detergent has been obtained, further preparation of the sample is necessary for crystallization trials. The sample needs to be concentrated to a concentration suitable for crystal trials and then filtered to remove particulate matter and aggregates. After concentration and filtration, the sample can be stored prior to setting up crystallization trials (see Basic Protocols 1 through 3).
Materials Detergent-exchanged membrane protein sample (see Support Protocol 1) 15-ml Centriprep concentrators (Millipore) 500-µl Microcon concentrators (Millipore) 500-µl Ultrafree 0.22-µm filtration devices (Millipore) 1. Concentrate the membrane protein sample to 10 mg/ml using the largest molecular weight cut-off membrane possible without losing sample. First concentrate the sample in a 15-ml Centriprep concentrator according to manufacturer’s instructions, and then transfer the sample to 500-µl Microcon concentrator(s) for further concentration, if necessary. It is important to use the highest molecular weight cut-off membranes possible to avoid concentrating the detergent more than necessary. Detergents with low cmc values, such as DDM, have large micellar molecular weights, often exceeding the molecular weight of the protein of interest. Ideally, the final concentrated sample will have a detergent concentration close to that of the size-exclusion buffer.
2. If one or more additives are being used in combination with the detergent chosen for crystallization, these should be added to the concentrated sample, taking care to mix gently to prevent the formation of excess air bubbles. Incubate the sample for 1 hr on ice. A list of additives successfully used to crystallize membrane proteins is found in Table 17.9.4. Screening for useful additives is described in Basic Protocol 3.
3. Transfer the sample to one or more Ultrafree filtration devices and centrifuge according to the manufacturer’s instructions. This step removes aggregates that might inhibit crystallization. An alternative method for removing aggregated protein is to spin the sample in an ultracentrifuge for 1 hr at ≥100,000 × g to pellet aggregated protein.
4. Store the concentrated, filtered sample on ice or at 4◦ C. The amount of time a concentrated membrane protein sample can be stored depends on the identity of the membrane protein, and must be determined empirically. Some membrane proteins lose activity within hours, while others are stable for several weeks. Do not freeze the sample, because many membrane proteins may not crystallize if they have been frozen, even if they retain full activity. Once the protein has been exchanged into the detergent of choice and concentrated, a broad screen of crystallization conditions is recommended. This is most easily done using a nanoliter pipetting robot (see Basic Protocol 1). A survey of the crystallization conditions that have led to published structures of membrane proteins (see http://www.mpibpfrankfurt.mpg.de/michel/public/memprotstruct.html) indicates that most have included polyethylene glycol (PEG) as the precipitant. Table 17.9.4 lists precipitants that have successfully yielded membrane protein crystals. Some recommended crystallization screens for membrane proteins are listed in Table 17.9.1. These screens have been chosen because Structural Biology
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they sample a wide range of conditions while retaining enough overlap to indicate the consistency of the results. The choice of which of these screens to use depends simply upon the volume of concentrated protein available; if enough protein is available, all 17 screens can be set up for each membrane protein sample. For a robot screen, a suggested drop volume of 0.2 µl protein solution plus 0.2 µl well solution minimizes losses due to evaporation (seen in smaller drops) but also conserves the protein sample, allowing a large number of experiments to be carried out for a given amount of concentrated membrane protein. Using the method described in Basic Protocol 1 and allowing for sample losses, each 96-condition screen requires 30 µl of a 10 mg/ml sample. The procedure for screening using a crystallization robot is given in Basic Protocol 1. Table 17.9.4 Detergents, Additives, and Precipitants that Have Been Used to Solve Crystal Structures of Outer and Inner Membrane Proteinsa
Outer membrane proteins
Inner membrane proteins
Detergents Octyl tetraoxyethylene
Octyl-β-D-glucoside
Octyl-β-D-glucoside
Dodecyl-β-D-maltoside
N,N-Dimethyldodecylamine-N-oxide
N,N-Dimethyldodecylamine-N-oxide
N,N-Dimethyldecylamine-N-oxide
Nonyl-β-D-glucoside
Decyl pentaoxyethylene
Undecyl-β-D-maltoside
Octyl-2-hydroxy ethylsulfoxide
Decyl-β-D-maltoside
Octyl polyoxyethylene
Dodecyl nonaoxyethylene
Octyl pentaoxyethylene
Dodecyl octaoxyethylene
Decyl-β-D-maltoside
Octyl-β-D-maltoside
Cyclohexylpentyl-β-D-maltoside
Dodecyl-α-D-maltoside
Additives Heptane-1,2,3-triol
Heptane-1,2,3-triol
N,N-Dimethylhexylamine-N-oxide
Benzamidine
Heptyl-β-D-glucoside
CHAPS
2-Propanol
HECAMEG
Dioxane
Dodecyl octaoxyethylene
Inositol
Dioxane Ethanol 2-Propanol 1,6-Hexanediol Dioleylphosphatidylcholine
Precipitants
Crystallization of Integral Membrane Proteins
Polyethylene glycol 2000
Polyethylene glycol 4000
2-Methyl-2,4-pentandiol
Polyethylene glycol 400
Polyethylene glycol 2000 MME
Polyethylene glycol 2000 MME
Polyethylene glycol 4000
Various phosphates
Polyethylene glycol 1000
Polyethylene glycol 350 MME
Polyethylene glycol 600
Ammonium sulfate
Ammonium sulfate
Polyethylene glycol 1000
a In each category, items are listed from most to least successful (adapted from http://www.mpibp-frankfurt.mpg.de/
michel/public/memprotstruct.html).
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Column buffer 20 mM Tris·Cl, pH 7.5 150 mM NaCl 0.5 mM EDTA 0.02% (w/v) NaN3 Detergent of choice at a concentration two times its cmc (le Maire et al., 2000) Prepare fresh A list of detergents successfully used to crystallize membrane proteins is found in Table 17.9.4. Properties of detergents, such as cmc, aggregation number, formula weight, and others, can be found in the Anatrace catalogue (http://www.anatrace.com). General properties of detergents can be found in Zulauf (1991).
COMMENTARY Background Information X-ray crystallography has been used to determine the structures of a large number of proteins. As of 2005, there were >32,000 structures deposited in the Protein Data Bank (http://www.rcsb.org/pdb/), yet only 0.5% of these represent integral membrane proteins. Membrane protein structures are underrepresented for several reasons. Many membrane proteins, both prokaryotic and eukaryotic, are difficult to express in functional form in sufficient quantities for structure determination (Grisshammer and Tate, 1995). Extraction of membrane proteins from their native lipid environment can lead to inactivation, as can subsequent purification using detergents. Crystallization is accomplished in the presence of detergents and/or lipids, adding more parameters that must be screened to obtain well-ordered 3-D crystals. Despite these difficulties, the number of membrane protein structures solved each year is increasing exponentially (White, 2004) due to the availability of new detergents, as well as advances in small-scale crystallization equipment, X-ray data collection at synchrotrons, and software used to more rapidly compute new structures.
Critical Parameters and Troubleshooting The most critical parameter for obtaining well-ordered membrane protein crystals is a natively folded, fully functional membrane protein sample of high purity. Care must be taken to analyze the activity of the sample in the desired detergent(s). Stability, defined as the length of time that a membrane protein retains full activity in a detergent environment, is also crucial to the crystallization process.
While determination of residual lipid and/or number of detergent molecules bound would be useful for establishing uniformity of protein preparations, these parameters are difficult to assay in a standard biochemistry laboratory. As with soluble proteins, compact, homogeneous molecules are more likely to crystallize than flexible molecules of limited purity. In addition, some ectodomains of membrane proteins remain folded and soluble without detergent when the transmembrane sequences are removed. These domains may be regarded as soluble proteins and crystallized as described in Chapter 17. Care should be taken when choosing molecular weight sizes for ultrafiltration devices. When a protein must be concentrated in earlier stages of the purification protocol, the filtration membrane pore size should be selected that minimizes protein losses. Membrane proteins are often observed to require smaller molecular weight cut-off membranes than their actual masses would predict, and we try to avoid sample losses during concentration wherever possible. The final detergent concentration is not critical at these stages. However, detergent micelles can have very large molecular weights, and many detergents are concentrated along with the protein sample when using molecular weight cut-off membranes of 10,000, 30,000, and sometimes even 50,000. This means that a ten-fold concentration of protein may also concentrate the detergent tenfold. Increased detergent concentrations can negatively influence crystallization, resulting in poorly ordered crystals, or no crystals at all. Therefore, once detergent exchange has been accomplished as described in Basic Protocol 1, the largest cut-off membranes possible are
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used to maximize protein retention while minimizing concentration of detergent. Ideally, the final detergent concentration of the concentrated protein sample will be approximately twice the critical micelle concentration of the detergent. One parameter that has a greater influence on membrane protein crystals than soluble protein crystals is temperature. Since membrane protein crystals have large amounts of detergent in the crystal lattice, small fluctuations in effective detergent concentration can easily destroy growing crystals. These fluctuations occur when the temperature increases or decreases, due to the dependence of the cmc of the detergent on temperature. It, therefore, makes sense to buy the best quality crystallization incubators available, with the most constant temperature control. One final difference between crystallizing membrane and soluble proteins lies in the strategies for improvement of crystal quality. For soluble proteins, if no crystals are obtained from commercial screens, one usually modifies the construct or tries a homolog. Therefore, the screening process for a given protein or protein fragment is rather brief. For membrane proteins, it is often not possible (or desirable) to remove part of the protein, and much more optimization is done using different detergents and additives for a single membrane protein construct. It is common to obtain membrane protein crystals that diffract to medium ◦ resolution (∼10 A), but refinement of conditions to higher resolution takes more time, more protein, sampling of more conditions, and more patience.
Anticipated Results
Crystallization of Integral Membrane Proteins
Crystallization trials generally show one (or more) types of behavior: clear drops, amorphous precipitate, granular precipitate, phase separation, microscopic crystals, or macroscopic crystals. Due to the increasing detergent concentration over time in a vapor-diffusion experiment (as described in this unit), phase separation is very common, and occasionally leads to crystal growth. Such a system is unstable, however, because areas of detergentrich and detergent-poor solution coexist in the same drop, and growing crystals can be destabilized by these differences. The best crystals are obtained in drops where no phase separation occurs, with slow growth of a few macroscopic crystals. For an example of how various parameters affect the growth of well-ordered crystals for a bacterial inner membrane pro-
tein, see Lemieux et al. (2003). Several reviews on the crystallization of integral membrane proteins are also available (Garavito et al., 1996; Loll, 2003; Wiener, 2004).
Time Considerations Once a purified membrane protein is available, it can usually be concentrated and screened with a robot in just a few days. Refinement of crystallization conditions may take weeks to months or even years, depending on the number of parameters that must be optimized. It is therefore important to produce enough protein routinely to set up numerous crystallization experiments so that traditional parameters such as precipitant and pH, as well as membrane-protein specific parameters such as detergent type and concentration, additives, and possibly lipids, can be evaluated for their influence on crystalline order. In the authors’ experience, the best ordered membrane protein crystals grow to completion in a time frame of 1 week to 1 month, with more rapidly growing crystals displaying poorer order.
Acknowledgement The work of T.J.B., J.L.W., and S.K.B. is supported by the Intramural Research Program of the NIH, National Institute of Diabetes and Digestive and Kidney Diseases.
Literature Cited Chang, G., Spencer, R.H., Lee, A.T., Barclay, M.T., and Rees, D.C. 1998. Structure of the MscL homolog from Mycobacterium tuberculosis: A gated mechanosensitive ion channel. Science 282:2220-2226. Dutzler, R., Campbell, E.B., and MacKinnon, R. 2003. Gating the selectivity filter in ClC chloride channels. Science 300:108-112. Garavito, R.M., Picot, D., and Loll, P.J. 1996. Strategies for crystallizing membrane proteins. J. Bioenerg. Biomembr. 28:13-27. Garman, E.F. and Doublie, S. 2003. Cryocooling of macromolecular crystals: Optimization methods. Methods Enzymol. 368:188-216. Grisshammer, R. and Tate, C.G. 1995. Overexpression of integral membrane proteins for structural studies. Q. Rev. Biophys. 28:315-422. Iwata, S. ed. 2003. Methods and Results in Crystallization of Membrane Proteins. La Jolla, International University Line. le Maire, M., Champeil, P., and Møller, J.V. 2000. Interaction of membrane proteins and lipids with solubilizing detergents. Biochim. Biophys. Acta 1508:86-111. Lebowitz, J., Lewis, M.S., and Schuck, P. 2002. Modern analytical ultracentrifugation in protein science: A tutorial review. Protein Sci. 11:20672079.
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Lemieux, M.J., Song, J., Kim, M.J., Huang, Y., Villa, A., Auer, M., Li, X.D., and Wang, D.N. 2003. Three-dimensional crystallization of the Escherichia coli glycerol-3-phosphate transporter: A member of the major facilitator superfamily. Protein Sci. 12:2748-2756.
Internet Resources
Locher, K.P., Lee, A.T., and Rees, D.C. 2002. The E. coli BtuCD structure: A framework for ABC transporter architecture and mechanism. Science 296:1091-1098.
http://www.mpibp-frankfurt.mpg.de/michel/ public/memprotstruct.html Hartmut Michel’s listing of crystallized membrane proteins, including a statistical analysis of crystallization conditions and structure references.
Loll, P.J. 2003. Membrane protein structural biology: The high throughput challenge. J. Struct. Biol. 142:144-153. McPherson, A. 1991. Useful principles for the crystallization of proteins. In Crystallization of Membrane Proteins. (H. Michel, ed.) pp. 1-51. CRC Press, Boca Raton, Fla. McPherson, A. 1999. Crystallization of Biological Macromolecules. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Michel, H. 1991. General and practical aspects of membrane protein crystallization. In Crystallization of Membrane Proteins. (H. Michel, ed.) pp. 73-88. CRC Press, Boca Raton, Fla. White, S.H. 2004. The progress of membrane protein structure determination. Protein Sci. 13:1948-1949. Wiener, M.C. 2004. A pedestrian guide to membrane protein crystallization. Methods 34:364372. Yue, W.W., Grizot, S., and Buchanan, S.K. 2003. Structural evidence for iron-free citrate and ferric citrate binding to the TonB-dependent outer membrane transporter FecA. J. Mol. Biol. 332:353-368.
http://www.rcsb.org/pdb Protein data base—the site where all protein structures (determined by X-ray crystallography, electron microscopy, and NMR) are deposited and made publicly available.
http://blanco.biomol.uci.edu/ Membrane Proteins xtal.html Stephen White’s listing of all solved membrane protein structures. This list is continuously updated and contains references to the individual structures. http://www.emeraldbiosystems.com/OnlineStore/ home.php http://www.hamptonresearch.com/ http://www.moleculardimensions.com/us/ index.ihtml http://www.nextalbiotech.com/ Web sites for crystallization instrumentation and reagents. http://www.anatrace.com/default.htm The Anatrace Web site for the widest variety of high purity detergents for membrane protein purification and crystallization.
Contributed by Travis J. Barnard, Jeremy L. Wally, and Susan K. Buchanan National Institutes of Health Bethesda, Maryland
Zulauf, M. 1991. Detergent phenomena in membrane protein crystallization. In Crystallization of Membrane Proteins. (H. Michel, ed.) pp. 5372. CRC Press, Boca Raton, Fla.
Structural Biology
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CHAPTER 18 Preparation and Handling of Peptides INTRODUCTION
A
n important component of the armamentarium of modern protein scientists is the ability to design and construct peptides of almost any sequence. Peptides may be employed for a wide range of applications, including as substrates for kinases, proteases, or glycosidases, or as antigens to produce antisera that may recognize a protein containing that sequence. Peptides also have many biological functions on their own, interacting with receptors to stimulate changes in cellular function, or altering the interactions between cells, for example. Chemists developed the means to synthesize peptides early in this century (UNIT 18.1). The ability to create a molecule that possessed biological activity by organic synthesis is certainly a milestone in our science. For many years, however, peptide synthesis was practiced only by highly skilled and dedicated individuals, and the lengthy time required to obtain even simple peptides limited their use as reagents for biological experiments. Difficulties in purification effectively eliminated the study of complex peptides of more than 5 to 10 amino acids.
Several developments have revolutionized this field, fortunately, and they have brought peptide science into the realm of standard biological techniques available to many scientists. First was the development of solid-phase methods by Bruce Merrifield and subsequent improvements in the reagents available for synthesis that are detailed in UNIT 18.1. Second was the design and construction of automated instruments to perform the repetitive steps of the solid-phase methodology, culminating in the 1980s with instruments that provided nearly global accessibility to synthetic peptides of ever-increasing length. Another advance was the optimization of reversed-phase HPLC as a separation method for the rapid purification and analysis of synthetic peptides (UNIT 11.6). A future unit in this chapter will focus on medium-to-large scale peptide purification by HPLC methods. In addition, recent advances in mass spectroscopy (Chapter 16) have provided another critical analytical tool to verify the products of synthesis. A convenient, low-cost method for creating large numbers of related peptides or for assembly of a set of peptides that together span an entire protein to search for the epitope recognized by an antibody that binds to the full-length protein is provided by the synthesis of peptides on a multipin apparatus (UNIT 18.2). Frequently, genomic studies lead to the discovery of an open reading frame (ORF), predicting a protein sequence that might have an activity of interest. One step that could follow would be the generation of an antibody to use as a reagent to identify the protein product of the gene in a sample isolated from the organism under study. A small peptide of 10 to 15 amino acids can frequently be used as an antigen to stimulate antiserum production. This requires picking the best sequence out of an ORF of hundreds of amino acids and preparing the conjugate for injection into an animal for generation of the antiserum (UNIT 18.3). As the procedures employed in solid-phase peptide synthesis have improved, researchers’ desire to attempt the preparation of longer sequences has grown. Methods to ligate Contributed by Ben M. Dunn Current Protocols in Protein Science (2001) 18.0.1-18.0.2 Copyright © 2001 by John Wiley & Sons, Inc.
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medium- to large-sized peptides together through defined chemistry are presented in UNIT 18.4. Success in this approach has opened the door to a variety of new synthetic objectives.
The chemistry and strategy utilized to prepare peptide dendrimers (UNIT 18.5) for use as immunogens, designed protein mimics, new reagents for drug discovery, or new biomaterials has been an outgrowth of solid-phase peptide synthesis. Methods for both Boc and Fmoc syntheses are described here. In addition, several procedures for the formation of cyclic peptides are presented, including derivatization of Lys and Cys residues. Preparing bioactive peptides or peptides with a fixed conformation often involves creating disulfide bridges (UNIT 18.6). The strategy and reactions necessary to achieve selective linkages in multi-Cys-containing peptides is described here; the analytical techniques required to characterize the peptides at each stage of the process are highlighted as well. Units planned for future expansion of this chapter will cover synthesis of derivatives such as phosphopeptides or other modification that can be done at the resin-bound stage, the creation of derivatives in which selective peptide bonds have been replaced to create inhibitors or to stabilize against degradation in biological systems, methods for storage and handling of peptides, specialized methods for different types of resins for solid-phase synthesis, the creation of peptide libraries by chemical or biological approaches, the chemistry necessary to achieve the coupling of difficult or hindered peptides, and methods for the assembly of peptides of 100 amino acid residues in a single, long synthesis. Ben M. Dunn
Introduction
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Introduction to Peptide Synthesis DEVELOPMENT OF SOLIDPHASE PEPTIDE-SYNTHESIS METHODOLOGY A number of synthetic peptides are significant commercial or pharmaceutical products, ranging from the dipeptide sugar substitute aspartame to clinically used hormones such as oxytocin, adrenocorticotropic hormone, and calcitonin. Rapid, efficient, and reliable methodology for the chemical synthesis of these molecules is of utmost interest. The stepwise assembly of peptides from amino acid precursors has been described for nearly a century. The concept is a straightforward one, whereby peptide elongation proceeds via a coupling reaction between amino acids, followed by removal of a reversible protecting group. The first peptide synthesis, as well as the creation of the term “peptide,” was reported by Fischer and Fourneau (1901). Bergmann and Zervas (1932) created the first reversible Nα-protecting group for peptide synthesis, the carbobenzoxy (Cbz) group. DuVigneaud successfully applied early “classical” strategies to construct a peptide with oxytocin-like activity (duVigneaud et al., 1953). Classical, or solution-phase methods for peptide synthesis have an elegant history and have been well chronicled. Solution synthesis continues to be especially valuable for largescale manufacturing and for specialized laboratory applications. Peptide synthesis became a more practical part of present-day scientific research following the advent of solid-phase techniques. The concept of solid-phase peptide synthesis (SPPS) is to retain chemistry that has been proven in solution but to add a covalent attachment step that links the nascent peptide chain to an insoluble polymeric support (resin). Subsequently, the anchored peptide is extended by a series of addition cycles (Fig. 18.1.1). It is the essence of the solid-phase approach that reactions are driven to completion by the use of excess soluble reagents, which can be removed by simple filtration and washing without manipulative losses. Once chain elongation has been completed, the crude peptide is released from the support. In the early 1960s, Merrifield proposed the use of a polystyrene-based solid support for peptide synthesis. Peptides could be assembled stepwise from the C to N terminus using Nαprotected amino acids. SPPS of a tetrapeptide
Contributed by Gregg B. Fields Current Protocols in Protein Science (2001) 18.1.1-18.1.9 Copyright © 2001 by John Wiley & Sons, Inc.
was achieved by using Cbz as an α-amino-protecting group, coupling with N,N′-dicyclohexylcarbodiimide (DCC), and liberating the peptide from the support by saponification or by use of HBr (Merrifield, 1963). SPPS was later modified to use the t-butyloxycarbonyl (Boc) group for Nα protection (Merrifield, 1967) and hydrogen fluoride (HF) as the reagent for removal of the peptide from the resin (Sakakibara et al., 1967). SPPS was thus based on “relative acidolysis,” where the Nα-protecting group (Boc) was labile in the presence of moderate acid (trifluoroacetic acid; TFA), while side-chain-protecting benzyl (Bzl)– based groups and the peptide/resin linkage were stable in the presence of moderate acid and labile in the presence of strong acid (HF). The first instrument for automated synthesis of peptides, based on Boc SPPS, was built by Merrifield, Stewart, and Jernberg (Merrifield et al., 1966). From the 1960s through the 1980s, Boc-based SPPS was fine-tuned (Merrifield, 1986). This strategy has been utilized for synthesis of proteins such as interleukin-3 and active enzymes including ribonuclease A and all-L and all-D forms of HIV-1 aspartyl protease. In 1972, Carpino introduced the 9-fluorenylmethoxycarbonyl (Fmoc) group for Nα protection (Carpino and Han, 1972). The Fmoc group requires moderate base for removal, and thus offered a chemically mild alternative to the acid-labile Boc group. In the late 1970s, the Fmoc group was adopted for solid-phase applications. Fmoc-based strategies utilized t-butyl (tBu)–based side-chain protection and hydroxymethylphenoxy-based linkers for peptide attachment to the resin. This was thus an “orthogonal” scheme requiring base for removal of the Nα-protecting group and acid for removal of the side-chain protecting groups and liberation of the peptide from the resin. The milder conditions of Fmoc chemistry as compared to Boc chemistry—which include elimination of repetitive moderate acidolysis steps and the final strong acidolysis step—were envisioned as being more compatible with the synthesis of peptides that are susceptible to acid-catalyzed side reactions. In particular, the modification of the indole ring of Trp was viewed as a particular problem during Boc-based peptide synthesis (Barany and Merrifield, 1979), which could be alleviated using Fmoc chemistry. One example of the potential advantage of Fmoc chemistry
UNIT 18.1
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18.1.1 Supplement 26
(King et al., 1990). Thus, the mild conditions of Fmoc chemistry appeared to be advantageous for certain peptides, as compared with Boc chemistry. One of the subsequent challenges for practitioners of Fmoc chemistry was to refine the technique to allow for construction of proteins, in similar fashion to that which had been achieved with Boc chemistry. Fmoc chemistry had its own set of unique problems, including suboptimum solvation of the peptide/resin,
for the synthesis of multiple-Trp-containing peptides was in the synthesis of gramicidin A. Gramicidin A, a pentadecapeptide containing four Trp residues, had been synthesized previously in low yields (5% to 24%) using Boc chemistry. The mild conditions of Fmoc chemistry dramatically improved the yields of gramicidin A, in some cases up to 87% (Fields et al., 1989, 1990). A second multiple-Trp-containing peptide, indolicidin, was successfully assembled in high yield by Fmoc chemistry
X
A
NH
O
CH C
+
OH
linker
resin
anchoring X
repetitive cycle
A
NH
O
CH C
linker
resin N α -deprotection
Y
NH
A
O
CH C
X
+
OH
H2 N
O
CH C
linker
resin
coupling
Y NH
A
O
CH C
X
NH
O
CH C
linker
resin α
(1) N -deprotection (2) cleavage (3) side-chain deprotection
Z H2 N
Introduction to Peptide Synthesis
O
CH C
R NH
Y
O
CH C
NH n
O
CH C
X
NH
O
CH C
NH2 OH
Figure 18.1.1 Generalized approach to solid-phase peptide synthesis. Symbols: A, Nα-protecting group; circle, side-chain protecting groups; R, X, Y, and Z, side-chain functionalities.
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slow coupling kinetics, and base-catalyzed side reactions. Improvements in these areas of Fmoc chemistry (Atherton and Sheppard, 1987; Fields and Noble, 1990; Fields et al., 2001) allowed for the synthesis of proteins such as bovine pancreatic trypsin inhibitor analogs, ubiquitin, yeast actin-binding protein 539-588, human β-chorionic gonadotropin 1-74, minicollagens, HIV-1 Tat protein, HIV-1 nucleocapsid protein Ncp7, and active HIV-1 protease. The milder conditions of Fmoc chemistry, along with improvements in the basic chemistry, have led to a shift in the chemistry employed by peptide laboratories. This trend is best exemplified by a series of studies (Angeletti et al., 1997) carried out by the Peptide Synthesis Research Committee (PSRC) of the Association of Biomolecular Resource Facilities (ABRF). The PSRC was formed to evaluate the quality of the synthetic methods utilized in its member laboratories for peptide synthesis. The PSRC designed a series of studies from 1991 to 1996 to examine synthetic methods and analytical techniques. A strong shift in the chemistry utilized in core facilities was observed during this time period—i.e., the more senior Boc methodology was replaced by Fmoc chemistry. For example, in 1991 50% of the participating laboratories used Fmoc chemistry, while 50% used Boc-based methods. By 1994, 98% of participating laboratories were using Fmoc chemistry. This percentage remained constant in 1995 and 1996. In addition, the overall quality of the peptides synthesized improved greatly from 1991 to 1994. Possible reasons for the improved results were any combination of the following (Angeletti et al., 1997): 1. The greater percentage of peptides synthesized by Fmoc chemistry, where cleavage conditions are less harsh; 2. The use of different side-chain protecting group strategies that help reduce side reactions during cleavage; 3. The use of cleavage protocols designed to minimize side reactions; 4. More rigor and care in laboratory techniques. The present level of refinement of solidphase methodology has led to numerous, commercially available instruments for peptide synthesis (Table 18.1.1). The next step in the development of solidphase techniques includes applications for peptides containing non-native amino acids, posttranslationally modified amino acids, and pseudoamino acids, as well as for organic molecules in general. Several areas of solid-phase
synthesis need to be refined to allow for the successful construction of this next generation of biomolecules. The solid support must be versatile so that a great variety of solvents can be used, particularly for organic-molecule applications. Coupling reagents must be sufficiently rapid so that sterically hindered amino acids can be incorporated. Construction of peptides that contain amino acids bearing posttranslational modifications should take advantage of the solid-phase approach. Finally, appropriate analytical techniques are needed to assure the proper composition of products.
THE SOLID SUPPORT Effective solvation of the peptide/resin is perhaps the most crucial condition for efficient chain assembly during solid-phase synthesis. Swollen resin beads may be reacted and washed batch-wise with agitation, then filtered either with suction or under positive nitrogen pressure. Alternatively, they may be packed in columns and utilized in a continuous-flow mode by pumping reagents and solvents through the resin. 1H, 2H, 13C, and 19F nuclear magnetic resonance (NMR) experiments have shown that, under proper solvation conditions, the linear polystyrene chains of copoly(styrene1%-divinylbenzene) resin (PS) are nearly as accessible to reagents as if free in solution. 13C and 19F NMR studies of Pepsyn (copolymerized dimethylacrylamide, N,N′-bisacryloylethylenediamine, and acryloylsarcosine methyl ester) have shown similar mobilities at resin-reactive sites as PS. Additional supports created by grafting polyethylene glycol (polyoxyethylene) onto PS—either by controlled anionic polymerization of ethylene oxide on tetraethylene glycol–PS (POE-PS) or by coupling Nω-Boc– or Fmoc–polyethylene glycol acid or –polyethylene glycol diacid to amino-functionalized PS (PEG-PS)—combine the advantages of liquid-phase synthesis (i.e., a homogeneous reaction environment) and solid-phase synthesis (an insoluble support). 13C NMR measurements of POE-PS showed the polyoxyethylene chains to be more mobile than the PS matrix, with the highest T1 spin-lattice relaxation times observed with POE of molecular weight 2000 to 3000. Other supports that have been developed that show improved solvation properties and/or are applicable to organic synthesis include polyethylene glycol polyacrylamide (PEGA), cross-linked acrylate ethoxylate resin (CLEAR), and augmented surface polyethylene prepared by chemical transformation (ASPECT). As the solid-phase
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Table 18.1.1
Instruments for Solid-Phase Synthesis
Suppliera
Instrument model
Peptide synthesis systems Advanced ChemTech 90 Apex 396 348 357 Bachem Bioscience SP4000-LAB SP4000-PRO Gilson/Abimed AMS422 Perkin-Elmer ABI433A Pioneer-MPS Rainin PS3 Sonata/Pilot Symphony/Multiplex CS Bio CS100 CS336 036 136 CS536 CS936S CS936 Intavis AG AutoSpot Multiple organic synthesis units Advanced ChemTech 384 Vantage aFor
Fmoc Boc
Batch Flow Monitoring
Scale (mmol)
No. of peptides
Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes
Yes Yes Yes Yes Yes No No Yes No Yes Yes No Yes Yes Yes Yes Yes Yes Yes No
Yes Yes Yes Yes Yes Yes Yes Yes No Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes
No No No No No No No No Yes No No No No No No No No No No No
No No No No No No Yes Yes Yes No No No No No No No No No No No
0.1-12 0.005-1 0.005-0.15 0.005-0.25 0.25-5 5-50 0.005-1 0.05-1 0.005-0.1 0.1-0.25 0.1-50 0.005-0.35 0.05-1 0.05-0.25 0.1-2.5 0.1-2.5 0.2-25 ≤600 ≤12,500 3-4 nmol/mm2
1-2 96 48 42 1 1 1 1 16 3 1 12 1 3 1 1 1 1 1 384
Yes Yes
Yes Yes
Yes Yes
No No
No No
— —
4 × 96 96
contact information, see SUPPLIERS APPENDIX.
method has expanded to include organic-molecule and library syntheses, the diversity of supports will enhance the efficiency of these new applications. Successful syntheses of problematic sequences can be achieved by manipulation of the solid support. In general, the longer the synthesis, the more polar the peptide/resin will become (Sarin et al., 1980). One can alter the solvent environment and enhance coupling efficiencies by adding polar solvents and/or chaotropic agents (Fields and Fields, 1994). Also, using a lower substitution level of resin to avoid interchain crowding can improve the synthesis (Tam and Lu, 1995). During difficult syntheses, deprotection of the Fmoc group can proceed slowly. By spectrophotometrically monitoring deprotection as the synthesis proceeds, one can detect problems and extend base-deprotection times and/or alter solvation conditions as necessary. Introduction to Peptide Synthesis
COUPLING REAGENTS The classical examples of in situ coupling reagents are N,N′-dicyclohexylcarbodiimide (DCC) and the related N,N′-diisopropylcarbodiimide (Rich and Singh, 1979). The generality of carbodiimide-mediated couplings is extended significantly by the use of either 1hydroxybenzotriazole (HOBt) or 1-hydroxy-7azabenzotriazole (HOAt) as an additive, either of which accelerates carbodiimide-mediated couplings, suppresses racemization, and inhibits dehydration of the carboxamide side chains of Asn and Gln to the corresponding nitriles. Protocols involving benzotriazol-1-ylo xy -tr is(d imethy lam ino )p hosphonium hexafluorophosphate (BOP), benzotriazol-1y l - o x y - t r i s ( p y r r o li d in o ) p h o s p h o n iu m hexafluorophosphate (PyBOP), 7 -azabenzotriazol-1-yl-oxytris(pyrrolidino)phosphoni um hexafluorophosphate (PyAOP), O-benzotriazol-1-yl-N,N,N′,N′-tetramethyluronium hexafluorophosphate (HBTU), O-(7-azaben-
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Table 18.1.2
Coupling Reagents and Additives Used in Solid-Phase Peptide Synthesis and Suppliers
Reagent
Abbreviation
Supplier(s)a
N,N′-dicyclohexylcarbodiimide N,N′-diisopropylcarbodiimide
DCC DIPCDI
A, ACT, AO, CI, CN, F, PI, PL, Q, S A, ACT, AO, CI, F, PE, Q, S
O-benzotriazol-1-yl-N,N,N′,N′-tetramethyluronium hexafluorophosphate O-benzotriazol-1-yl-N,N,N′,N′-tetramethyluronium tetrafluoroborate O-(7-azabenzotriazol-1-yl)-N,N,N′,N′tetramethyluronium hexafluorophosphate
HBTU
A, ACT, AS, CI, CN, F, NS, PI, PL, Q, S
TBTU
A, ACT, B, CI, CN, F, NS, PE, PI, PL, Q, S
HATU
PE
Benzotriazol-1-yl-oxy-tris(dimethylamino) phosphonium hexafluorophosphate Benzotriazol-1-yl-oxy-tris(pyrrolidino)phosphonium hexafluorophosphate 7-azabenzotriazole-1-yl-oxy-tris(pyrrolidino)phophonium hexafluorophosphate
BOP
A, ACT, AO, B, CI, CN, F, NS, PL, PI, Q, S
PyBOP
A, ACT, AO, CI, CN, F, S
PyAOP
PE
Tetramethylfluoroformamidinium hexafluorophosphate 1-hydroxybenzotriazole
TFFH
ACT, PE
HOBt
A, ACT, AO, AS, CI, CN, NS, PE, PI, Q, S
1-hydroxy-7-azabenzotriazole N,N-diisopropylethylamine
HOAt DIEA
PE A, ACT, AO, CI, F, PE, Q, S
N-methylmorpholine
NMM
A, AO, CI, F, S
aAbbreviations: A, Aldrich; ACT, Advanced ChemTech; AO, Acros Organics; AS, AnaSpec; B, Bachem; CI, Chem-Impex; CN, Calbiochem-Novabiochem; F, Fluka; NS, Neosystem/SNPE; PE, Perkin-Elmer; PI, Peptides International; PL, Peninsula Laboratories; Q, Quantum Biotechnologies; S, Sigma. For contact information, see SUPPLIERS APPENDIX.
zotriazol-1-yl)-N,N,N′,N′-tetramethyluronium hexafluorophosphate (HATU), and O-benzotriazol-1-yl-N,N,N′,N′-tetramethyluronium tetrafluoroborate (TBTU) result in coupling kinetics even more rapid than that obtained with carbodiimides. Amino acid halides have also been applied to solid-phase peptide synthesis (SPPS). Nα-protected amino acid chlorides have a long history of use in solution synthesis. Fmoc–amino acid chlorides and fluorides react rapidly under SPPS conditions in the presence of HOBt/N,N-diisopropylethylamine (DIEA) and DIEA, respectively, with very low levels of racemization. For convenience, tetramethylfluoroformamidinium hexafluorophosphate (TFFH) can be used for automated preparation of Fmoc–amino acid fluorides. Amino acid fluorides have been found to be especially useful for the preparation of peptides containing sterically hindered amino acids, such as peptaibols. All of the coupling reagents and additives discussed here are commercially available (see Table 18.1.2).
SYNTHESIS OF MODIFIED RESIDUES AND STRUCTURES Peptides of biological interest often include structural elements beyond the 20 genetically encoded amino acids. Particular emphasis has been placed on peptides containing phosphorylated or glycosylated residues or disulfide bridges. Incorporation of side-chain-phosphorylated Ser and Thr by solid-phase peptide synthesis (SPPS) is especially challenging, as the phosphate group is decomposed by strong acid and lost with base in a β-elimination process. Boc-Ser(PO3phenyl2) and Boc-Thr(PO3phenyl2) have been found to be useful derivatives, where hydrogen fluoride (HF) or hydrogenolysis cleaves the peptide/resin and hydrogenolysis removes the phenyl groups. Fmoc-Ser(PO3Bzl,H) and Fmoc-Thr(PO3Bzl,H) can be used in conjunction with Fmoc chemistry with some care. Alternatively, peptide/resins that were built up by Fmoc chemistry to include unprotected Ser or Thr side chains may be subject to “global” or post-assembly phosphorylation. Side-chain-phosphorylated Tyr is less susceptible to strong-acid decomposition and is not at all base-labile. Thus, SPPS
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Introduction to Peptide Synthesis
has been used to incorporate directly FmocTyr(PO3methyl2), Fmoc-Tyr(PO3tBu2), FmocTyr(PO3H2), and Boc-Tyr(PO3H2). Phosphorylation may also be accomplished on-line, directly after incorporation of the Tyr, Ser, or Thr residue but prior to assembly of the whole peptide. Methodology for site-specific incorporation of carbohydrates during chemical synthesis of peptides has developed rapidly. The mild conditions of Fmoc chemistry are more suited for glycopeptide syntheses than Boc chemistry, as repetitive acid treatments can be detrimental to sugar linkages. Fmoc-Ser, -Thr, -5-hydroxylysine (-Hyl), -4-hydroxyproline (-Hyp), and -Asn have all been incorporated successfully with glycosylated side chains. The side-chain glycosyl is usually hydroxyl-protected by either benzoyl or acetyl groups, although some SPPSs have been successful with no protection of glycosyl hydroxyl groups. Deacetylation and debenzylation are performed with hydrazine/methanol prior to glycopeptide/resin cleavage or in solution with catalytic methoxide in methanol. Disulfide-bond formation has been achieved on the solid-phase by air, K3Fe(CN)6, dithiobis(2-nitrobenzoic acid), or diiodoethane oxidation of free sulfhydryls, by direct deprotection/oxidation of Cys(acetamidomethyl) residues using thallium trifluoroacetate or I2, by direct conversion of Cys(9-fluorenylmethyl) residues using piperidine, and by nucleophilic attack by a free sulfhydryl on either Cys(3-nitro-2-pyridinesulfenyl) or Cys(S-carboxymethylsulfenyl). The most generally applicable and efficient of these methods is direct conversion of Cys(acetamidomethyl) residues by thallium trifluoroacetate. Intra-chain lactams are formed between the side-chains of Lys or Orn and Asp or Glu to conformationally restrain synthetic peptides, with the goal of increasing biological potency and/or specificity. Lactams can also be formed via side-chain-to-head, side-chain-to-tail, or head-to-tail cyclization (Kates et al., 1994). The residues used to form intra-chain lactams must be selectively side-chain deprotected, while all side-chain protecting groups of other residues remain intact. Selective deprotection is best achieved by using orthogonal side-chain protection, such as allyloxycarbonyl or 1-(4,4-dimethyl-2,6-dioxocyclohex-1-ylidene)ethyl protection for Lys and allyl or N-[1-(4,4,-dimethyl-2,6-dioxocyclohexylidene)-3-methyl butyl]aminobenzyl protection for Asp/Glu in combination with an Fmoc/tBu strategy. Cyclization is carried out most efficiently with BOP
in the presence of DIEA while the peptide is still attached to the resin. The three-dimensional orthogonal protection scheme of Fmoc/tBu/allyl protecting groups is the strategy of choice for head-to-tail cyclizations. An amide linker is used for sidechain attachment of a C-terminal Asp/Glu (which are converted to Asn/Gln) and the αcarboxyl group is protected as an allyl ester. For side-chain-to-head cyclizations, the N-terminal amino acid (head) can simply be introduced as an Nα-Fmoc derivative while the peptide-resin linkage and the other side-chain protecting groups are stable to dilute acid or carry a third dimension of orthogonality.
PROTEIN SYNTHESIS There are three general chemical approaches for constructing proteins. First is stepwise synthesis, in which the entire protein is synthesized one amino acid at a time. Second is “fragment assembly,” in which individual peptide strands are initially constructed stepwise, purified, and finally covalently linked to create the desired protein. Fragment assembly can be divided into two distinct approaches: (1) convergent synthesis of fully protected fragments, and (2) chemoselective ligation of unprotected fragments. Third is “directed assembly,” in which individual peptide strands are constructed stepwise, purified, and then noncovalently driven to associate into protein-like structures. Combinations of the three general chemical approaches may also be employed for protein construction. Convergent synthesis utilizes protected peptide fragments for protein construction (Albericio et al., 1997). The advantage of convergent protein synthesis is that fragments of the desired protein are first synthesized, purified, and characterized, ensuring that each fragment is of high integrity; these fragments are then assembled into the complete protein. Thus, cumulative effects of stepwise synthetic errors are minimized. Convergent synthesis requires ready access to pure, partially protected peptide segments, which are needed as building blocks. The application of solid-phase synthesis to prepare the requisite intermediates depends on several levels of selectively cleavable protecting groups and linkers. Methods for subsequent solubilization and purification of the protected segments are nontrivial. Individual rates for coupling segments are substantially lower then for activated amino acid species by stepwise synthesis, and there is always a risk of racemization at the C-terminus of each segment. Care-
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ful attention to synthetic design and execution may minimize these problems. As an alternative to the segment condensation approach, methods have been developed by which unprotected peptide fragments may be linked. “Native chemical ligation” results in an amide bond being generated between peptide fragments (Muir et al., 1997). A peptide bearing a C-terminal thioacid is converted to a 5-thio-2-nitrobenzoic acid ester and then reacted with a peptide bearing an N-terminal Cys residue (Dawson et al., 1994). The initial thioester ligation product undergoes spontaneous rearrangement, leading to an amide bond and regeneration of the free sulfhydryl on Cys. The method was later refined so that a relatively unreactive thioester can be used in the ligation reaction (Dawson et al., 1997; Ayers et al., 1999). Safety-catch linkers are used in conjunction with Fmoc chemistry to produce the necessary peptide thioester (Shin et al., 1999).
SIDE-REACTIONS
The free Nα-amino group of an anchored dipeptide is poised for a base-catalyzed intramolecular attack of the C-terminal carbonyl. Base deprotection of the Fmoc group can thus release a cyclic diketopiperazine while a hydroxymethyl-handle leaving group remains on the resin. With residues that can form cis peptide bonds, e.g., Gly, Pro, N-methylamino acids, or D-amino acids, in either the first or second position of the (C → N) synthesis, diketopiperazine formation can be substantial. The steric hindrance of the 2-chlorotrityl linker may minimize diketopiperazine formation of susceptible sequences during Fmoc chemistry. The conversion of side-chain protected Asp residues to aspartimide residues can occur by repetitive base treatments. The cyclic aspartimide can then react with piperidine to form the α- or β-piperidide or α- or β-peptide. Aspartimide formation can be rapid, and is dependent upon the Asp side-chain protecting group. Sequence dependence studies of Asp(OtBu)-X peptides revealed that piperidine could induce aspartimide formation when X = Arg(2,2,5,7,8pentamethylchroman-6-sulfonyl; Pmc), Asn(triphenylmethyl; Trt), Asp(OtBu), Cys(Acm), Gly, Ser, Thr, and Thr(tBu) (Lauer et al., 1995). Aspartimide formation can also be conformation-dependent. This side-reaction can be minimized by including 0.1 M HOBt in the piperidine solution (Lauer et al., 1995), or by using an amide backbone protecting group (i.e., 2-hydroxy-4-methoxybenzyl) for the resi-
due in the X position of an Asp-X sequence (Quibell et al., 1994). Cys residues are racemized by repeated piperidine deprotection treatments during Fmoc SPPS. Racemization of esterified (C-terminal) Cys can be reduced by using 1% 1,8diazabicyclo[5.4.0]undec-7-ene in N,N-dimethylformamide (DMF). Additionally, the steric hindrance of the 2-chlorotrityl linker minimizes racemization of C-terminal Cys residues. When applying protocols for Cys internal (not C-terminal) incorporation which include phosphonium and aminium salts as coupling agents, as well as preactivation in the presence of suitable additives and tertiary amine bases, significant racemization is observed. Racemization is generally reduced by avoiding preactivation, using a weaker base (such as collidine), and switching to the solvent mixture DMF-dichloromethane (DCM) (1:1). Alternatively, the pentafluorophenyl ester of a suitable Fmoc-Cys derivative can be used. The combination of side-chain protecting groups and anchoring linkages commonly used in Fmoc chemistry are simultaneously deprotected and cleaved by TFA. Cleavage of these groups and linkers results in liberation of reactive species that can modify susceptible residues, such as Trp, Tyr, and Met. Modifications can be minimized during TFA cleavage by utilizing effective scavengers. Three efficient cleavage “cocktails” quenching reactive species and preserving amino acid integrity, are TFA-phenol-thioanisole-1,2-ethanedithiol-H2O (82.5:5:5:2.5:5) (reagent K) (King et al., 1990), TFA-thioanisole-1,2-ethanedithiol-anisole (90:5:3:2) (reagent R) (Albericio et al., 1990), and TFA-phenol-H2O-triisopropylsilane (88:5:5:2) (reagent B) (Solé and Barany, 1992). The use of Boc side-chain protection of Trp also significantly reduces alkylation by Pmc or 2,2,4,6,7-pentamethyldihydro-benzofuran-5sulfonyl (Pbf) groups.
PURIFICATION AND ANALYSIS OF SYNTHETIC PEPTIDES Each synthetic procedure has limitations, and even in the hands of highly experienced workers, certain sequences defy facile preparation. The maturation of high-performance liquid chromatography (HPLC) has been a major boon to modern peptide synthesis, because the resolving power of this technique facilitates removal of many of the systematic low-level by-products that accrue during chain assembly and upon cleavage. Peptide purification is most commonly achieved by reversed-phase HPLC
Preparation and Handling of Peptides
18.1.7 Current Protocols in Protein Science
Supplement 26
(RP-HPLC; UNIT 11.6). Either alternatively to or in tandem with RP-HPLC, ion-exchange HPLC (UNIT 8.2) and gel-filtration HPLC (UNIT 8.3) can be used for isolation of desired peptide products. The progress of peptide purification can be monitored rapidly by matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF-MS; UNITS 16.2 & 16.3) or ion-trap electrospray MS (UNIT 16.8). The homogeneity of synthetic materials should be checked by at least two chromatographic or electrophoretic techniques, e.g., RP-HPLC (UNIT 11.6), ion-exchange HPLC (UNIT 8.2), and capillary zone electrophoresis (UNIT 10.9). Also, determination of a molecular ion by MS (see Chapter 16) using a mild ionization method is important for proof of structure. Synthetic peptides must be checked routinely for the proper amino acid composition, and in some cases sequencing data are helpful. The PSRC studies (see discussion of Development of Solid-Phase Peptide Synthesis Methodology) have allowed for a side-by-side comparison of a variety of analytical techniques. Efficient characterization of synthetic peptides best been obtained by a combination of RP-HPLC and MS, with sequencing by either Edman degradation sequence analysis or tandem MS (UNIT 16.1) being used to identify the positions of modifications and deletions. Proper peptide characterization by multiple techniques is essential.
LITERATURE CITED Albericio, F., Kneib-Cordonier, N., Biancalana, S., Gera, L., Masada, R.I., Hudson, D., and Barany, G. 1990. Preparation and application of the 5-(4(9-fluorenylmethyloxycarbonyl)aminomethyl3,5-dimethoxyphenoxy)valeric acid (PAL) handle for the solid-phase synthesis of C-terminal peptide amides under mild conditions. J. Org. Chem. 55:3730-3743. Albericio, F., Lloyd-Williams, P., and Giralt, E. 1997. Convergent solid-phase peptide synthesis. Methods Enzymol. 289:313-336. Angeletti, R.H., Bonewald, L.F., and Fields, G.B. 1997. Six year study of peptide synthesis. Methods Enzymol. 289:697-717. Atherton, E. and Sheppard, R.C. 1987. The fluorenylmethoxycarbonyl amino protecting group. In The Peptides, Vol. 9 (S. Udenfriend and J. Meienhofer, eds.) pp. 1-38. Academic Press, New York. Ayers, B., Blaschke, U.K., Camarero, J.A., Cotton, G.J., Holford, M., and Muir, T.W. 1999. Introduction of unnatural amino acids into proteins using expressed protein ligation. Biopolymers (Peptide Sci.) 51:343-354. Introduction to Peptide Synthesis
Barany, G. and Merrifield, R.B. 1979. Solid-phase peptide synthesis. In The Peptides, Vol. 2 (E. Gross and J. Meienhofer, eds.) pp. 1-284. Academic Press, New York. Bergmann, M. and Zervas, L. 1932. Über ein allgemeines Verfahren der Peptidsynthese. Ber. Dtsch. Chem. Ges. 65:1192-1201. Carpino, L.A. and Han, G.Y. 1972. The 9-fluorenylmethoxycarbonyl amino-protecting group. J. Org. Chem. 37:3404-3409. Dawson, P.E., Muir, T.W., Clark-Lewis, I., and Kent, S.B.H. 1994. Synthesis of proteins by native chemical ligation. Science 266:776-779. Dawson, P.E., Churchill, M.J., Ghadiri, M.R., and Kent, S.B.H. 1997. Modulation of reactivity in native chemical ligation through the use of thiol additives. J. Am. Chem. Soc. 119:4325-4329. duVigneaud, V., Ressler, C., Swan, J.M., Roberts, C.W., Katsoyannis, P.G., and Gordon, S. 1953. The synthesis of an octapeptide amide with the hormonal activity of oxytocin. J. Am. Chem. Soc. 75:4879-4880. Fields, C.G. and Fields, G.B. 1994. Solvents for solid-phase peptide synthesis. In Methods in Molecular Biology, Vol. 35: Peptide Synthesis Protocols (M.W. Pennington and B.M. Dunn, eds.) pp. 29-40. Humana Press, Totowa, N.J. Fields, G.B. and Noble, R.L. 1990. Solid phase peptide synthesis utilizing 9-fluorenylmethoxycarbonyl amino acids. Int. J. Peptide Protein Res. 35:161-214. Fields, C.G., Fields, G.B., Noble, R.L., and Cross, T.A. 1989. Solid phase peptide synthesis of 15Ngramicidins A, B, and C and high performance liquid chromatographic purification. Int. J. Peptide Protein Res. 33:298-303. Fields, G.B., Otteson, K.M., Fields, C.G., and Noble, R.L. 1990. The versatility of solid phase peptide synthesis. In Innovation and Perspectives in Solid Phase Synthesis: Peptides, Polypeptides and Oligonucleotides, Macro-organic Reagents and Catalysts (R. Epton, ed.) pp. 241-260. Solid Phase Conference Coordination, Ltd., Birmingham, U.K. Fields, G.B., Lauer-Fields, J.L., Liu, R.-q., and Barany, G. 2001. Principles and practice of solidphase peptide synthesis. In Synthetic Peptides: A User’s Guide, 2nd ed. (G.A. Grant, ed.) in press. W.H. Freeman, New York. Fischer, E. and Fourneau, E. 1901. Über einige Derivate des Glykocoils. Ber. Dtsch. Chem. Ges. 34:2868-2877. Kates, S.A., Solé, N.A., Albericio, F., and Barany, G. 1994. Solid-phase synthesis of cyclic peptides. In Peptides: Design, Synthesis and Biological Activity (C. Basava and G.M. Anantharamaiah, eds.) pp. 39-57. Birkhaeuser, Boston. King, D.S., Fields, C.G., and Fields, G.B. 1990. A cleavage method which minimizes side reactions following Fmoc solid phase peptide synthesis. Int. J. Peptide Protein Res. 36:255-266.
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Lauer, J.L., Fields, C.G., and Fields, G.B. 1995. Sequence dependence of aspartimide formation during 9-fluorenylmethoxycarbonyl solid-phase peptide synthesis. Lett. Peptide Sci. 1:197-205. Merrifield, R.B. 1963. Solid phase peptide synthesis I: The synthesis of a tetrapeptide. J. Am. Chem. Soc. 85:2149-2154. Merrifield, R.B. 1967. New approaches to the chemical synthesis of peptides. Recent Prog. Hormone Res. 23:451-482. Merrifield, R.B. 1986. Solid phase synthesis. Science 232:341-347. Merrifield, R.B., Stewart, J.M., and Jernberg, N. 1966. Instrument for automated synthesis of peptides. Anal. Chem. 38:1905-1914. Muir, T.W., Dawson, P.E., and Kent, S.B.H. 1997. Protein synthesis by chemical ligation of unprotected peptides in aqueous solution. Methods Enzymol. 289:266-298. Quibell, M., Owen, D., Packman, L.C., and Johnson, T. 1994. Suppression of piperidine-mediated side product formation for Asp(OBut)containing peptides by the use of N-(2-hydroxy4-methoxybenzyl) (Hmb) backbone amide protection. J. Chem. Soc. Chem. Commun. 2343-2344. Rich, D.H. and Singh, J. 1979. The carbodiimide method. In The Peptides, Vol. 1 (E. Gross and J. Meienhofer, eds.) pp. 241-314. Academic Press, New York. Sakakibara, S., Shimonishi, Y., Kishida, Y., Okada, M., and Sugihara, H. 1967. Use of anhydrous HF in peptide synthesis I: Behavior of various protective groups in anhydrous HF. Bull. Chem. Soc. Jpn. 40:2164-2167.
Sarin, V.K., Kent, S.B.H., and Merrifield, R.B. 1980. Properties of swollen polymer networks: Solvation and swelling of peptide-containing resins in solid-phase peptide synthesis. J. Am. Chem. Soc. 102:5463-5470. Shin, Y., Winans, K.A., Backes, B.J., Kent, S.B.H., Ellman, J.A., and Bertozzi, C.R. 1999. Fmocbased synthesis of peptide-αthioesters: Application to the total chemical synthesis of a glycoprotein by native chemical ligation. J. Am. Chem. Soc. 121:11684-11689. Solé, N.A. and Barany, G. 1992. Optimization of solid-phase synthesis of [Ala8]-dynorphin A. J. Org. Chem. 57:5399-5403. Tam, J.P. and Lu, Y.-A. 1995. Coupling difficulty associated with interchain clustering and phase transition in solid phase peptide synthesis. J. Am. Chem. Soc. 117:12058-12063.
KEY REFERENCES Atherton, E. and Sheppard, R.C. 1989. Solid Phase Peptide Synthesis: A Practical Approach. IRL Press, Oxford. An extensive collection of Fmoc-based synthetic methods and techniques. Barany and Merrifield, 1979. See above. The definitive, comprehensive overview of the solidphase method. Fields, G.B. 1997. Solid-phase peptide synthesis. Methods Enzymol. Vol. 289. A contemporary collection of SPPS techniques and applications.
Contributed by Gregg B. Fields Florida Atlantic University Boca Raton, Florida
Preparation and Handling of Peptides
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Synthesis of Multiple Peptides on Plastic Pins
UNIT 18.2
Scanning protein sequences by bioassay for smaller bioactive peptide sequences requires a source of many peptides homologous with the parent protein sequence. This unit deals with one of the synthetic methods for making such sets of peptides (see Fig. 18.2.1). The key to preparing large numbers (hundreds to thousands) of synthetic peptides in a short time and at minimal cost is to use a parallel synthesis technique which is efficient and can be done on a small scale. The multipin technology is suitable because it can be performed without expensive synthesizers and it uses equipment available to most laboratories. Prior experience with organic synthesis techniques or peptide chemistry is useful but not essential. The products of synthesis by multipin technology are unpurified peptides which are useful as screening reagents and may also be used to prepare purified peptide on a small scale. Most multipin techniques exploit the conventional 8 × 12 matrix layout of common microtiter equipment which simplifies handling of the synthesis, the products (peptides), and the test results. Computer assistance with synthesis and data analysis also speeds the cycle from designing the experiment through analyzing the results. With multipin technology, peptides are synthesized in parallel on plastic “pins” (Fig. 18.2.2) to give sets of peptides suitable not only for B and T cell epitope scanning but also for other bioassays. Peptides can be either permanently bound to the surface of the plastic for direct binding assays, or they can be released into solution. There is a choice of N- and C-terminal peptide endings. For solution-phase peptides, the synthesis scale can be 1 or 5 µmol (for a 10-mer, ∼1 mg or 5 mg, respectively). The preferred coupling/deprotection chemistry used is the milder 9-fluorenylmethyloxycarbonyl (Fmoc) protection scheme rather than the older t-butyloxycarbonyl (t-Boc) protection scheme (see UNIT 18.1), thus reducing the level of chemical safety risk arising from synthetic peptide chemistry. This unit covers the strategy of the multiple peptide approach to biological scanning, the synthetic protocols, and the handling of peptides after synthesis—cleavage, preliminary purification, storage, and analysis (see Basic Protocol). It is specific for the multipin technique using equipment obtained from Chiron Technologies, although some of the approaches are applicable to other multiple synthesis techniques. Procedures for multipin equipment obtained from other suppliers may differ from the procedures described here, and the manufacturer’s literature should be consulted. This unit also includes protocols for preparing Fmoc–amino acid solutions (see Support Protocol 1) and for acetylating (see Support Protocol 2) or biotinylating (see Support Protocol 3) synthesized peptides. STRATEGIC PLANNING For a protein whose primary structure is known, the conceptually simplest method of locating all the bioactive linear peptide sequences is to make all possible peptide subsets of the protein sequence and test them. If only selected parts of the sequence are synthesized, or only the predicted active parts, bioactive sequences could be missed. The use of a set of highly overlapping peptides likewise reduces the possibility that the most bioactive sequences might be missed because they are absent from the set. A set of all overlapping 20-mers offset along the sequence by one residue at a time should capture the entire set of, for example, helper T cell epitopes, and this is a much more reliable approach than trying to predict motifs. In reality, a synthetic peptide scan through a protein is a compromise between the cost and effort in making and screening all peptides and the Contributed by Stuart J. Rodda Current Protocols in Protein Science (1997) 18.2.1-18.2.19 Copyright © 1997 by John Wiley & Sons, Inc.
Preparation and Handling of Peptides
18.2.1 Supplement 9
need for completeness. Thus, one worker may choose to make all overlapping 8-mers to find the linear (continuous) B cell epitopes, and another may make 12-mers offset along the sequence by five residues for the same purpose. In each case, all sequences of eight residues from the protein are present in at least one peptide, but the latter approach requires only one-fifth the number of peptides.
decide on type of peptides to be made: e.g., noncleavable, cleaved, biotinylated, acetylated (see Table 18.2.1)
obtain kit with the required pin type and install software
Fmoc deprotect the pins
(n cycles) generate synthesis schedule using Pepmaker software based on protein sequence or individual peptide sequences
pipet activated amino acids (Support Protocol 1) into the wells of the reaction trays and place freshly deprotected blocks of pins into the appropriate trays; incubate
position the required number of pins in the pinholder, following the information on the synthesis schedule
deprotect the side chains of peptides; for the MPS kit, cleave the peptides from the pins
wash the peptides, either as a precipitate in a tube (MPS kit) or still on the pins (NCP, GAP, and DKP kits)
wash pins
Fmoc deprotect the pins
acetylate (Support Protocol 2 ) or biotinylate (Support Protocol 3) the N-terminus of the peptides before side chain deprotection or cleavage as required
test pin-bound peptides with conjugate alone, prior to testing with specific antiserum
carry out assays with pin-bound peptides Synthesis of Multiple Peptides on Plastic Pins
cleave the peptides (GAP or DKP kit)
carry out assays with cleaved peptides
Figure 18.2.1 Flow chart for multipin peptide synthesis.
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A
stem
gear
pin with gear
macrocrown
pin with macrocrown
leg
B
Figure 18.2.2 Apparatus for multipin peptide synthesis. (A) Assembled synthesis block with 96 gears (left) or 96 macrocrowns (right). (B) Components of the pin assembly. Components are either push-fitted together (e.g., legs or stems into the pin holder) or clipped on (gears or macrocrowns onto stems). All components are solvent-resistant plastic, either polyethylene, polypropylene, or copolymers of these two monomer types.
Planning the Synthesis Synthetic peptides are assembled by solid-phase synthesis one amino acid at a time, commencing with the C-terminal end of the peptide on the solid phase (see UNIT 18.1). The assembly process, or coupling, requires activation of the α-carboxyl group of each incoming amino acid to make it reactive with the α-amino group of the growing peptide chain. To prevent unwanted polymerization or side reaction, reactive groups in each amino acid must be temporarily protected, and the protecting group removed before further reaction can be carried out. The protecting group on the α-amino function of the most recently added amino acid must be removed before another amino acid can be coupled to it, so the α-amino protection must be labile under conditions that do not remove side-chain protection. Later, the side-chain-protecting groups must be removable under conditions that do not attack the peptide bonds. The two common protecting group “schemes” are known as t-butoxycarbonyl (t-Boc) or 9-fluorenylmethyloxycarbonyl (Fmoc). The protecting group scheme currently recommended for multipin peptide synthesis is the milder Fmoc scheme, which is the only scheme described in this chapter. Before beginning to plan the actual synthesis in detail, a choice needs to be made regarding how the peptides will eventually be presented in the bioassay. The options available to investigators are listed in Table 18.2.1. For noncleavable peptide (NCP) kits, peptides are permanently bound on the solid phase (pin surface) and can be used for direct binding assays but not for interaction with living cells or other complex (e.g., multicomponent) systems. In this case, the peptides must be
Preparation and Handling of Peptides
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Table 18.2.1
Types of Pins for Multipin Peptide Synthesisa
Name
Linkerb
Physical formatc Loading
Final form of peptide
NCP MPS MPS DKP GAP
Noncleavable AA ester Rink amide DKP-forming Glycine ester
Gear Macrocrown Macrocrown Gear Gear
(N-capping)-PEPTIDE-linker-pin (N-capping)-PEPTIDE-acid (N-capping)-PEPTIDE-amide (N-capping)-PEPTIDE-DKP (N-capping)-PEPTIDE-glycine-acid
50 nmol 5 µmol 5 µmol 1 µmol 1 µmol
aAbbreviations: DKP, diketopiperazine; GAP, glycine acid peptide; MPS, multiple peptide synthesis; NCP, noncleavable
peptide; (N-capping), a free amine, acetyl group, or biotin; PEPTIDE, the sequence of the peptide being made. bNature of linker between peptide and graft polymer on the pin: noncleavable linker, β-alanine-hexamethylenediamine;
DKP, diketopiperazine; AA ester, amino acid ester; Rink amide, Rink amide–forming linker. cSee Figure 18.2.2B.
“regenerated” between repeat assays by disrupting the peptide-ligand interaction without damaging the peptide. The quantity of peptide made is very small (50 nmol), but it is sufficient to provide a high surface density of peptide for direct binding assays. In the other options, peptides are synthesized on pins and then released into solution. The mechanism of peptide release into solution affects the postsynthesis handling and thus the suitability of peptides produced by each cleavage method for various assay systems. For multiple peptide synthesis (MPS) kits, the released peptides have a “native” free acid or an amide carboxy terminus. To make free acid C-termini, it is necessary to use macrocrowns that already have the first (C-terminal) amino acid on them because the chemistry of forming the first (ester) link is too difficult for the inexperienced user. In contrast, the Rink amide linker allows formation of a peptide with a C-terminal amide of any amino acid by adding the C-terminal amino acid to the Rink handle macrocrown using the standard amino acid coupling protocol. A Rink amide linker is a linker that can accept an amino acid but then can be cleaved in trifluoroacetic acid (TFA) to release the amide form of that amino acid (Rink, 1987). Although acid or amine endings are often the most desirable peptide format to have, they are also the most complex to produce because the cleavage of the peptides from the pin is into neat TFA plus scavengers which needs to be evaporated to recover the peptide. The scale of peptide synthesis for MPS kits is 5 µmol (∼5 mg of a decamer). For glycine acid peptide (GAP) kits, peptides with a glycine at the carboxy terminus are cleaved as the free acid, so that the C-terminal residue is a natural amino acid (glycine) and is not blocked. The peptides are also relatively simple to release from the pin and require little postsynthesis handling. However, the presence of glycine at the C-terminus may be undesirable where the C-terminus plays an important role in peptide bioactivity. The scale of synthesis for GAP kits is 1 µmol (∼1 mg of a decamer). In diketopiperazine (DKP) kits, peptides are synthesized with a DKP group at the C terminus. The DKP group is a cyclic dipeptide formed from C-terminal lysine and proline residues during the facile cleavage of the peptide under the mildest possible conditions: neutral aqueous buffer. In applications where the presence of the DKP group is acceptable, this type of peptide can make the downstream processing of synthetic peptides very simple and fast. The peptides can be placed into a bioassay system immediately after completing the cleavage. The scale of synthesis for DKP kits is 1 µmol (∼1 mg of a decamer). Synthesis of Multiple Peptides on Plastic Pins
For these five kit options, it is also possible to choose a variety of N-terminal endings on the peptides. For example, it may be desirable to acetylate pin-bound peptides (see Support Protocol 2) to eliminate the positive charge that would otherwise be present on
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the α-amino group of the N-terminal residue, or to enhance the activity of a peptide in a T helper assay (Mutch et al., 1991). A handy option for cleaved peptides is to place a biotin group on the N-terminus (see Support Protocol 3) so the peptide can be captured using avidin or streptavidin. These additions must be made prior to side-chain deprotection of the peptides. There are other configurations for multiple peptide synthesis—e.g., the SPOTS or “peptides on paper” system (Zeneca/CRB), the RaMPS system (DuPont), and multi-synthesizer machines (e.g., Advanced ChemTech). Assessing Peptide Sequences Peptides differ so much in properties that it is important to assess the likely properties of the peptides before attempting to synthesize them. Peptide length and hydrophobicity are the two main attributes affecting successful synthesis. The longer the peptide, the lower will be the purity of the product, as each amino acid coupling cycle is never 100% efficient. Synthesis of peptides longer than 20 residues should be avoided unless special attention can be given to each sequence. Hydrophobic peptides may be difficult to synthesize, but more significantly they may be poorly soluble in aqueous buffers, restricting their ultimate usefulness in bioassays. Prior to beginning synthesis of a set of peptides, it is sensible to assess them all for hydrophobicity (Fauchere and Pliska, 1983; UNIT 2.2) and decide if all should be attempted as they stand. In many cases, it is possible to choose slightly different peptides (longer, shorter, or using a different starting and finishing point in the homologous protein sequence) that will have more user-friendly properties. As well as these general factors affecting peptides, particular peptide sequences may have characteristics that make them difficult to synthesize, or they may be problematic after synthesis. It is not feasible to discuss all the common problems here. To help assessment of peptide sequences, a software application called Pinsoft is available free from Chiron Technologies. This allows any sequence to be typed in, and an assessment is automatically reported. Generating Peptide Sequences Computer software (Pepmaker) supplied with synthesis kits allows sets of overlapping peptide sequences to be generated from a protein sequence computer file using the single-letter amino acid code. Alternatively, sequences can be created using a word processor and the resulting computer text file can then be used by Pepmaker to guide synthesis. The use of this software simplifies the otherwise complex and tedious task of adding the right amino acids to each reaction plate on each synthesis cycle. MULTIPIN SYNTHESIS OF PEPTIDES Derivatized pins with macrocrowns or gears are attached to a pin holder. Each peptide is built up on the reactive surface of one pin by successive cycles of amino acid coupling, followed by washing and removal of the 9-fluorenylmethyloxycarbonyl (Fmoc) aminoprotecting group to prepare for the next amino acid coupling cycle. A critical step is properly dispensing activated amino acid solutions into the appropriate wells of each reaction tray. A list of the well locations for dispensing of each amino acid is generated by the Pepmaker software for this purpose. When the peptides are complete, trifluoroacetic acid (TFA) that contains scavengers is used to remove the side-chain-protecting groups, and for MPS kits, to cleave the peptides from the pins. The manual provided with each type of kit (see Table 18.2.1) includes instructions and hints for kit-specific procedures. NOTE: All reagents should be of the highest grade possible, preferably peptide synthesis or analytical reagent grade.
BASIC PROTOCOL
Preparation and Handling of Peptides
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Materials 20% (v/v) piperidine/dimethylformamide (DMF; see recipe) DMF, analytical reagent grade Methanol, analytical reagent grade 100 mM activated 9-fluorenylmethyloxycarbonyl (Fmoc)–protected amino acid solutions (see Support Protocol 1) Side chain deprotecting (SCD) solution (see recipe) Acidified methanol: 0.5% (v/v) glacial acetic acid/methanol 1:2:0.003 (v/v/v) ether/petroleum ether/2-mercaptoethanol (2-ME) 1:2 (v/v) ether/petroleum ether 0.1 M NaOH 0.1 M acetic acid 0.1 M sodium phosphate buffer, pH 8.0 (APPENDIX 2E) Sonication buffer (see recipe) Peptide Synthesis Starter Kit (e.g., Chiron Technologies) of the desired type, containing: Pepmaker computer program and ELISA reading and plotting programs Manual Pins with gears or macrocrowns Storage boxes or sealable bags, polyethylene or polypropylene (ICN Biomedicals) Pipettor tips, polyethylene or polypropylene (ICN Biomedicals) 0.3- or 1.5-ml reaction trays, polyethylene or polypropylene (Chiron Technologies, Nunc, or Beckman) Sonicator with power output of ∼500 W Dry nitrogen Rack containing 96 1-ml polypropylene tubes (Bio-Rad) 10-ml capped conical polypropylene centrifuge tubes Additional reagents and equipment for N-terminal acetylation (see Support Protocol 2; optional) or biotinylation (see Support Protocol 3; optional) CAUTION: Perform all chemistry steps in a well-functioning chemical fume hood. Wear solvent-resistant gloves, safety glasses, and protective clothing. The reagents can be flammable, toxic, and/or carcinogenic. Avoid sources of contamination which may affect the pins, including direct contact with the bench surface or exposure to vapors. The reagents for multipin synthesis can be handled in unsealed systems, but the amount of time that these reagents are left exposed to the open air should be minimized by using capped containers for liquids or polyethylene bags for pins wherever practical. Local regulations for safe disposal of solvents and used reagents must be followed. Prepare synthesis schedule and equipment 1. Use the Pepmaker computer program according to the instructions to generate the required set of peptide sequences (Fig. 18.2.3). Generate the printouts, which show for each coupling cycle how much of each amino acid solution, catalyst, and activating agent needs to be prepared (see Fig. 18.2.4) and where each amino acid solution is to be added to the reaction tray (see Fig. 18.2.5).
Synthesis of Multiple Peptides on Plastic Pins
The standard microtiter plate layout is an 8 × 12 matrix, in which the eight rows are identified as A through H and the twelve columns are identified as 1 through 12. However, the Pepmaker software uses a designation in which the column number is given first followed by a number designation for the row, beginning with row H, given in parentheses—i.e., 1(1) for well H1, 1(2) for well G1, 2(1) for well H2, and 12(8) for well A12 (see Fig. 18.2.6).
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2. Label each pin holder block indelibly on the top (e.g., Synthesis #1, Block A, Synthesis #1, Block B, and so forth), preferably by scratching into the plastic with a sharp tool. Place the label where it will help orient the block so that the pins are not accidentally placed into amino acid solutions in an inverted orientation. For example, keep pin H1 and well H1 at the lower left corner of the block (Fig. 18.2.6). Ink labels will run or disappear with exposure to solvents. The multipin system is based on the standard microtiter plate layout. The block is the complete unit and consists of the pin holder, which is the support that holds 96 pins (in an 8 × 12 array with standard ELISA microtiter plate spacing), and five legs to support the device and correctly position the active surfaces. A pin consists of an inert stem that supports either a gear or a macrocrown, both of which have an active surface on which the peptide is synthesized (see Figure 18.2.2). A gear is a detachable gear-shaped unit that fits on the thin end of a stem. A macrocrown is a detachable, vaned tip that fits on the thin end of a stem. It is made of high-density polyethylene and the surface is derivatized to give a solvent-compatible polymer matrix. Macrocrowns are provided in two forms: one has a linker that cleaves to give peptides with an amide at the carboxy terminus; the other has a linker that cleaves to give the free acid at the carboxy terminus and is supplied with an amino acid already attached to the linker. The reaction tray used for the synthesis is a polyethylene or polypropylene tray consisting of 96 wells in the standard microtiter plate 8 × 12 matrix. Shallow reaction trays (0.3-ml) are used with gears and deep trays (1.5-ml) are used with macrocrowns.
3. Remove any pins that are not required for the first cycle of amino acid coupling and store them dry in a plastic bag in the refrigerator until needed. Some pins need to be removed when the peptides in the synthesis are of various lengths because the software is designed to arrange all peptides to complete their synthesis on the same (final) coupling cycle. This approach eliminates unnecessary Fmoc–deprotection cycles for pins that are designated to carry the shorter peptides. The synthesis printout from the Pepmaker software shows which pins need to be added for each cycle of amino acid addition (Fig. 18.2.5). Pins (stems) can be pushed out from the top side of the pin holder. In the case of the MPS kit, where the first amino acid is already on the macrocrown as supplied, choose and mount the correct macrocrown for each position on the block.
Deprotect α-amino groups 4. Add 20% piperidine/DMF to a bath and place the pins in the bath so that the tips (macrocrowns or gears) are covered. Cover and let stand for 20 min at room temperature. CAUTION: Piperidine is flammable. The volume of reagent needed for all the bath steps depends on the shape of the bath, the critical factor being that all surfaces of the pins (i.e., the gears or macrocrowns) bearing the peptide need to be totally covered. A small bath suitable for gears is the upturned polypropylene lid of a pipettor tip box. For macrocrowns, deeper baths or deep-well polypropylene trays as supplied with the kit can be used.
5. Remove the block from the bath, shake off the excess liquid, and then wash the pins in a DMF bath for 2 min at room temperature. Again, the DMF must fully cover the tips.
6. Shake off the excess DMF and immerse the block completely in a deep bath of methanol for 2 min so that all surfaces of the block are washed. CAUTION: Methanol is flammable and toxic. In a shallower bath the block can be turned over so that the pin holder part is washed as well.
Preparation and Handling of Peptides
18.2.7 Current Protocols in Protein Science
Supplement 9
A
GENERAL NET SYNTHESIS SCHEDULE NUMBER : 1
Page 1
Description: Example of a scan through Sperm Whale Myoglobin 8-mer peptides based on the sequence MBN-SW Peptide spacing increment is 1 Segment 1: 146 peptides starting at residue 1 First peptide: [VLSEGEWQ] Last peptide: [YKELGYQG] Protein sequence: MBN-SW (153 residues) 1: VLSEGEWQLV LHVWAKVEAD VAGHGQDILI RLFKSHPETL EKFDRFKHLK 51: TEAEMKASED LKKHGVTVLT ALGAILKKKG HHEAELKPLA QSHATKHKIP 101: IKYLEFISEA IIHVLHSRHP GNFGADAQGA MNKALELFRK DIAAKYKELG 151: YQG Amino Acid set to be used - AASET1 aaset 1:Free acid L-Fmoc amino acids - DIC/HOBt chemistry Number of copies of each peptide 1 Schedule based on a 250 microliter fill/well (Well concentration is 100 mM)
Figure 18.2.3 (above and at right) A portion of the synthesis schedule worksheets generated by Pepmaker software for Schedule no. 1 for synthesis of a set of all possible overlapping octamers of sperm whale myoglobin. (A) This page of the synthesis schedule is a summary of the features of the protein including its sequence. (B) This page of the synthesis schedule shows the sequences of the first 96 peptides that will be synthesized, the first two of which are the controls, PLAQGGGG and GLAQGGGG. Peptide sequences are shown in the conventional amino-to-carboxy-terminal direction (from left to right), with a “<” sign indicating the end attached to the solid phase during synthesis. Because amino acid couplings are carried out in the carboxy-to-amino direction, the first amino acids coupled are at the right-hand end of each sequence, adjacent to the “<”.
7. Place the block in a second methanol bath to fully cover the tips. Wash for 2 min. Repeat this washing step again with a fresh methanol bath for a total of three methanol washes. 8. Remove the block and allow it to air dry in an acid-free fume hood for a minimum of 30 min. Avoid exposure to acidic fumes as this could prevent efficient coupling in the next step. The block can be conveniently left to dry while the amino acid solutions are being dispensed.
Dispense activated amino acid solutions 9. Dispense the required volume of each activated amino acid solution (see Support Protocol 1; 150 µl for gears or 450 µl for macrocrowns) into the appropriate wells of the reaction tray as specified by the synthesis schedule for the coupling cycle (e.g., Fig. 18.2.5).
Synthesis of Multiple Peptides on Plastic Pins
Perform the amino acid coupling 10. Place the pins in the activated amino acid solutions in the reaction tray, ensuring that the block is correctly oriented before actually lowering the pins into the solution. Incubate ≥2 hr at 20° to 25°C in a polyethylene box with a lid or in a sealable polyethylene bag. Coupling begins immediately and is irreversible.
18.2.8 Supplement 9
Current Protocols in Protein Science
Figure 18.2.3 (continued) Preparation and Handling of Peptides
18.2.9 Current Protocols in Protein Science
Supplement 9
Wash the pins 11. Remove the block of pins from the amino acid solutions and, if the next cycle is to start immediately, place the block in a methanol bath in which the pins are immersed to half their height (tips are fully immersed). Wash with agitation for 5 min. Flick off excess methanol and allow to air dry for 2 min. 12. Place the block in a DMF bath in which the pins are immersed to half their height (tips are fully immersed). Wash with agitation for 5 min. If extra pins are to be added as synthesis progresses (if the peptides being made differ in length), add the pins to the appropriate spaces (see Fig. 18.2.5).
13. Repeat steps 4 through 12 for each cycle of amino acid addition. Follow the synthesis schedule for preparing Fmoc–protected amino acid solutions and for depositing
Synthesis of Multiple Peptides on Plastic Pins
Figure 18.2.4 This page of the synthesis schedule is used for the preparation of activated amino acid solutions. It shows the amounts of each amino acid (represented by the single letter code, A through Y, along the left-hand margin), activator (diisopropylcarbodiimide [DIC] in dimethylformamide [DMF]), and catalyst (additive; 1-hydroxybenzotriazole [HOBt] in DMF) needed for the first amino acid coupling cycle. (In this example the amounts are calculated for a 250-µl reaction volume.) The amino acid powder is dissolved in the HOBt/DMF solution (right-hand column) before activation with DIC/DMF solution.
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GENERAL NET SYNTHESIS SCHEDULE NUMBER : 1 PIN POSITIONS for Synthesis coupling NEW PIN POSITIONS A 1(1) TO B 6(5)
Page 4 1
Well positions for amino acid dispensing A
A10(1) A 2(2) A 5(2) A12(4) A 4(5) A 6(6) A 9(6) A 7(7) A 1(8) A 5(8) B11(1) B 2(3) B 4(3) B 7(3) B11(3) B 8(4) TO B 9(4)
D
A 3(2)
E
A 1(2) A 9(3) A12(3) A11(4) A 1(5) A 6(5) A 8(7) B 6(1) B10(1) B 1(4) B 1(5)
F
A 4(3)
G
A 1(1) TO A 2(1) A 6(2) A 8(2) A12(5) A 8(6) A 3(7) B 1(1) TO B 2(1) B10(2) B 1(3) B 6(3) B 3(5) B 6(5)
H
A 7(1) A 7(2) A 7(3) A 7(4) A11(5) A 4(7) TO A 5(7) A 4(8) A 8(8) B 2(2) B 5(2) B 8(2)
I
A11(2) A 1(3) A10(6) TO B 1(2) B 7(4)
K
A11(1) A 5(3) A 1(4) A 6(4) A 9(4) A 3(5) A 9(5) TO A10(5) A12(6) TO A 2(7) A10(7) A 7(8) A 9(8) B 3(1) B10(3) B 5(4) B10(4) B12(4)
L
A 4(1) A 6(1) A12(2) A 3(3) A11(3) A 8(4) A 8(5) A 4(6) A 7(6) A11(6) A 9(7) A12(7) B 5(1) B 4(2) B12(3) B 2(4) B 2(5)
M
A 2(5)
B 8(3)
N
B11(2)
B 9(3)
P
A 8(3)
A11(7)
A11(8)
B 9(2)
Q
A 3(1)
A 9(2)
A 2(8)
B 5(3)
R
A 2(3)
A 4(4)
B 7(2)
B 4(4)
S
A 6(3)
A 5(5)
A 3(8)
B 9(1)
B 6(2)
T
A10(3)
A10(4)
A 2(6)
A 5(6)
A 6(8)
V
A 5(1)
A 8(1)
A12(1)
A 4(2)
A 1(6)
W
A 9(1)
Y
B 4(1)
B11(4)
B 4(5)
A10(2)
A 2(4)
A 3(4)
A 5(4)
A 7(5)
B 7(1)
A10(8)
B 3(3)
B12(2)
A12(8)
B 6(4) A 6(7)
B 3(4)
B 8(1)
B12(1)
B 5(5)
A 3(6)
B 3(2)
Figure 18.2.5 This page of the synthesis schedule identifies which wells of the reaction tray receive which activated amino acid solution. Each amino acid solution is identified along the left-hand margin using the single-letter amino acid code. Individual reaction trays and pin holders (A or B in this case) are identified by letters of the alphabet, and the paired numbers—e.g., 10(1) for well 10H—identify individual wells within reaction trays according to the numbering system illustrated in Figure 18.2.6.
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aliquots of the activated amino acids to the appropriate wells of the reaction tray for each cycle. Two coupling cycles can be carried out during a normal working day, and a third coupling can be carried out overnight, so a total of three amino acids can be added to each pin during a 24-hr period.
14. Deprotect the final Fmoc amino acid by repeating steps 4 through 8. Then proceed with step 15 or step 16. 15. Optional: For B cell epitope scanning or for T helper cell epitope scanning, the N-terminus of the peptide can be capped by acetylation (see Support Protocol 2). To allow later recapture onto avidin, the N-terminus of the peptide can be capped with biotin or long-chain biotin (see Support Protocol 3). Deprotect the side chains 16. Dispense the required volume of SCD solution into a bath or tubes (1.5 ml/tube) and fully immerse the peptide-bearing portion of the pins. Cover bath or cap tubes and incubate 2.5 hr at 20° to 25°C. CAUTION: SCD solution is a toxic, corrosive liquid. For blocks where the peptide is to remain attached to the pin during side chain deprotection (i.e., NCP, GAP, and DKP kits), side chain deprotection is carried out in a bath of reagent. For pins where the peptide is simultaneously side chain deprotected and cleaved from the pin (MPS kits), the process is carried out in individual 10-ml capped conical polypropylene centrifuge tubes that become the containers for the recovered peptide.
17a. For NCP, GAP, or DKP kits: Wash the pins 3 times in acidified methanol to remove the SCD solution prior to further treatment. 17b. For MPS kit: In a good chemical fume hood, reduce the volume of SCD solution containing cleaved peptide to ∼0.1 ml either with a gentle stream of dry nitrogen gas or in a centrifugal vacuum drier (e.g., Speedvac) equipped to handle corrosive fumes. Precipitate the peptide in the remaining solution with 8 ml of 1:2:0.003 ether/petroleum ether/2-ME. Decant and discard the supernatant, and wash the
1
Synthesis of Multiple Peptides on Plastic Pins
(8)
A
(7)
B
(6)
C
(5)
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(4)
E
(3)
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Figure 18.2.6 The numbering system for pins and reaction trays for Chiron Technologies’ multipin synthesis system. Each well is identified by a pair of numbers rather than a number and a letter, e.g, 1(1). The first number identifies the column number; the second (in parentheses) identifies the lettered row, beginning with H as (1) and ending with A as (8).
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precipitated peptide with 4 ml of 1:2 ether/petroleum ether. Dry the precipitate with a gentle stream of dry nitrogen. CAUTION: Ether/petroleum ether/2-ME and ether/petroleum ether solutions are highly flammable. These dry peptides can now be redissolved for assay purposes or may be further purified, e.g., by HPLC (see UNIT 11.6).
Prepare the peptides 18a. For GAP kits: Add 0.7 ml of 0.1 M NaOH to each tube of a rack of 96 1-ml polypropylene tubes. Place the pins into the solution and incubate ∼1.5 hr to cleave the peptides from the pins. Immediately after cleavage, neutralize with one equivalent of 0.1 M acetic acid. Alternatively, the 0.1 M NaOH solution can contain 40% (v/v) acetonitrile to facilitate solubilization of the more hydrophobic peptides. The incubation time can be shorter if the tubes are sonicated during cleavage.
18b. For DKP kits: Cleave the peptide from the pin in a suitable, reasonably well-buffered solution with a pH >7 overnight (16 hr). The solution for cleavage—e.g., 0.1 M sodium phosphate, pH 8, or 0.1 M HEPES, pH 8—can be chosen to be compatible with the assay for which the peptides will be used; the solution should have a buffering capacity of ∼0.05 M. Again, an organic modifier such as acetonitrile can be added to the cleavage solution to facilitate solubilization of the more hydrophobic peptides.
18c. For NCP kits: Prepare the pins for binding assays by floating the block, pin-side down, in sonication buffer and sonicating 10 min at ∼60°C. Rinse the pins first in water, then in 20° to 45°C methanol for immediate use in an assay or air dry the pins for storage until they are used in an assay. PREPARING ACTIVATED Fmoc–PROTECTED AMINO ACID SOLUTIONS Activated 9-fluorenylmethyloxycarbonyl (Fmoc)–protected amino acid solutions are prepared in two steps. First, the protected amino acid is dissolved in a solution of the catalyst, 1-hydroxybenzotriazole (HOBt) in dimethylformamide (DMF). Then, just before dispensing, the amino acid is activated by adding an activating agent. The following procedure illustrates the use of diisopropylcarbodiimide (DIC) as the activating reagent. DIC is a liquid and can be measured by volume.
SUPPORT PROTOCOL 1
Additional Materials (also see Basic Protocol) 9-fluorenylmethyloxycarbonyl (Fmoc)–protected amino acids with side-chain-protecting groups (Sigma, Bachem, Novachem, or Chiron Technologies), stored at 4°C Catalyst: 1-hydroxybenzotriazole (HOBt) Activating agent: diisopropylcarbodiimide (DIC) Dimethylformamide (DMF), amine-free Ethanol, analytical reagent grade 5- or 10-ml glass, polyethylene, or polypropylene bottles with inert (e.g., polythylene, Teflon) lids and liners 1. Remove the Fmoc–protected amino acids from the refrigerator and allow them to come to room temperature before weighing them out. Warming the containers to room temperature avoids the possibility of uptake of moisture from the air onto the cold solids.
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The side chains of the amino acids must also be protected during peptide synthesis: t-butyl ether is used for serine, threonine, and tyrosine; t-butyl ester is used for aspartic acid and glutamic acid; t-butoxycarbonyl (t-Boc) is used for lysine, histidine, and tryptophan; 2,2,5,7,8-pentamethylchroman-6-sulfonyl (Pmc) is used for arginine; and trityl (Trt) is used for cysteine. If benzotriazolyl-N-oxytris(dimethylamino)phosphonium hexafluorophosphate (BOP) or benzotriazol-1-yl-oxytripyrrolidinophosphonium hexafluorophosphate (PyBOP) activation is to be used, trityl protection should be used for asparagine and glutamine. BOP and PyBOP require greater care in handling and different protection should be used for some amino acids.
2. Weigh individual amino acids and HOBt using the quantities specified for the current synthesis cycle (e.g., see Fig. 18.2.4) into separate appropriately sized clean and dry glass, polyethylene, or polypropylene bottles. Rinse the spatula with ethanol and dry it after weighing each reagent. The bottle caps and inserts must be inert to any of the reagents or solvents used in making up the activated solutions (they must be Teflon, not rubber). Care should be taken to avoid cross-contamination of amino acids by rinsing the spatula in ethanol between weighings (the spatula must be dry before each use) and by making sure that all lids, as well as the containers, are labeled so they can be replaced on the correct bottles after weighing has been completed.
3. Measure the appropriate amount of DIC (see “Activator” as in Fig. 18.2.4). DIC is a liquid so it is more convenient to measure it by volume rather than by weight. Multiply the indicated weight by 1.23 (based on the density of DIC, 0.815 g/ml) as a conversion factor from the calculated weight to get the required volume of DIC in microliters.
4. Prepare HOBt and DIC solutions, by pipetting the appropriate volumes of purified (amine-free) DMF as shown on the synthesis schedule (e.g., see Fig. 18.2.4). Both reagents should be fully dissolved in the DMF before using them to prepare the activated amino acid solutions.
5. Add the specified volume of HOBt/DMF solution (e.g., Fig. 18.2.4, column headed “HOBt”) to the individual amino acids. Make sure the amino acids are completely dissolved before adding the activator solution. Unactivated amino acid solutions may be stored a few days at 4°C.
6. Activate the individual amino acid solutions by adding the specified volume of DIC/DMF solution to each amino acid solution (e.g., Fig. 18.2.4, column headed “DIC”). Mix thoroughly and use immediately for peptide synthesis. Activated amino acids should be prepared immediately before use and any excess should be discarded. SUPPORT PROTOCOL 2
Synthesis of Multiple Peptides on Plastic Pins
N-TERMINAL ACETYLATION OF PEPTIDES N-terminal capping of the peptides is carried out after a final 9-fluorenylmethyloxycarbonyl (Fmoc)–deprotection cycle and prior to side chain deprotection. The process is similar to coupling of amino acids, except that in the case of acetylation the active reagent can be acetic anhydride rather than acetic acid. Acetic anhydride does not require activation. If acetic anhydride is not available, simply use acetic acid as if it were an amino acid (see Support Protocol 3). Additional Materials (also see Basic Protocol) Acetylation solution (see recipe), prepared just before use Pins with completed peptides (see Basic Protocol)
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1. Add freshly prepared acetylation solution to appropriate bath container. 2. Immerse the pins with completed peptides in acetylation solution and incubate 90 min at room temperature. 3. Wash the pins in a methanol bath and air dry. The pins can now be used for side chain deprotection (Basic Protocol, step 16).
N-TERMINAL BIOTINYLATION OF PEPTIDES Biotin can also be coupled to the N-terminus of peptides after N-terminal deprotection and before side chain deprotection. The reagent is used as if it were an amino acid, using the same solvent, activating agent, and catalyst.
SUPPORT PROTOCOL 3
Additional Materials (also see Basic Protocol) Biotin or long-chain biotin Dimethylformamide (DMF), amine-free Diisopropylcarbodiimide (DIC) Pins with completed peptides (see Basic Protocol) 1. Dissolve biotin in amine-free DMF to a concentration of 125 mM. 2. Prepare a 10× solution of DIC (activating agent) by dissolving 158 mg DIC in 1 ml DMF and prepare a 10× solution of HOBt (catalyst) by dissolving 192 mg HOBt in 1 ml DMF. 3. Activate the biotin with the 10× concentrate solutions of activation agent and catalyst (80:10:10 [v/v/v]). 4. Dispense 150 µl/well (for gears) or 450 µl/well (for macrocrowns) into reaction trays. 5. Immerse the pins with completed peptides in the reaction tray and incubate ≥2 hr. 6. Wash the pins in methanol. The pins can now be used for side chain deprotection (see Basic Protocol, step 16).
REAGENTS AND SOLUTIONS Use Milli-Q water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Acetylation solution For 200 ml: 193 ml dimethylformamide (DMF) 6 ml acetic anhydride 1 ml N-ethyldiisopropylamine Prepare immediately before use and discard after use DO NOT expose pins to acetic anhydride at any other time except during acetylation. Also, do not store acetic anhydride anywhere near where peptide synthesis is performed. The DMF does not need to be amine-free.
20% piperidine/DMF Prepare a 20% (v/v) solution of the best quality piperidine available in analytical reagent–grade dimethylformamide (DMF). Prepare a fresh solution for each synthesis (solution can be reused several times within a synthesis). Store at room temperature in an amber bottle containing activated molecular sieves to remove moisture. CAUTION: This solution is highly flammable and toxic. continued
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If high-quality piperidine is not available, it may have to be treated with solid sodium hydroxide and redistilled. DMF need not be amine-free.
Side chain deprotecting (SCD) solution 33 parts (v/v) trifluoroacetic acid 1 part (v/v) ethanedithiol 2 parts (v/v) anisole 2 parts (v/v) thioanisole 2 parts (v/v) H2O Prepare immediately before use and do not store or reuse CAUTION: This solution is corrosive and extremely malodorous. Contamination of the laboratory, especially with ethanedithiol, should be avoided. Wipe the outside of ethanedithiol-contaminated equipment or containers with dilute, 0.1% aqueous hydrogen peroxide to oxidize ethanedithiol to a nonodorous compound before removing the container from the fume hood. DO NOT allow hydrogen peroxide to contact other readily oxidizable materials or reagents.
Sonication buffer 1% (w/v) SDS 0.1 M sodium phosphate buffer, pH 7.2 (APPENDIX 2E) 0.1% (v/v) 2-mercaptoethanol (2-ME) Store at room temperature up to 1 week CAUTION: Before discarding sonication buffer, destroy remaining 2-ME by adding 2 ml 30% hydrogen peroxide per liter of buffer.
COMMENTARY Background Information The multipin method was developed by Dr. H.M. Geysen and coworkers (Geysen et al., 1984, 1987) as a scanning method for linear antibody-defined epitopes. Eventually in the late 1980s, the method was adapted to parallel synthesis of cleaved (soluble) peptides (Maeji et al., 1990), opening the way for systematic scanning of T helper (Reece et al., 1993) and cytotoxic epitopes (Burrows et al., 1994). Initially only suitable for synthesis of short peptides (up to 10 amino acid residues), the method can now routinely produce peptides of up to 20 residues of acceptable quality for initial screening experiments (Valerio et al., 1993).
Critical Parameters
Synthesis of Multiple Peptides on Plastic Pins
Successful peptide synthesis requires reagents of a quality appropriate to the particular step, and the careful application of those reagents. For example, the protected amino acids need to be free of reactive counterions such as dicyclohexylamine (DCHA), contaminating unprotected amino acid, isomers such as the D-amino acid, and water. Check carefully that the amino acid as supplied is EXACTLY the same as specified in the manual or on the software. Apart from quality testing each amino
acid, the best assurance of quality is to buy only from reputable suppliers. Dimethylformamide (DMF) is the primary solvent for carrying out reactions (couplings) on pins. Its low volatility and moderate polarity make it suitable for dissolving the amino acids and solvating the graft polymer/growing peptide on the pin surface. Purity is not critical for some (washing) steps, but is critical for the DMF used just before and during amino acid coupling. Presence of excessive amine in the DMF results in loss of activated amino acid because the amino acid couples to the amine rather than to the peptide on the pin. Fortunately, the pin system allows use of substantial molar excesses of incoming amino acid (typically 6- to 1000-fold), so loss of some amino acid is not disastrous. Fresh DMF of the best available grade should be used for the coupling, and it is recommended that the amine level be tested using the FDNB test (Stewart and Young, 1984). Liberal use is made of methanol as a washing solvent. Analytical reagent grade methanol is readily available at low cost in large containers (20 or 200 liters) and is relatively easy to dispose of. It is possible to reduce the use of methanol by reusing it for washes: the last wash
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bath in any series should be in fresh (pure) methanol. In the next round of washes, the former last bath is then reassigned as the second-to-last wash, the previously second-to-last bath becomes the third-to-last, and so on. For each synthesis cycle, the first wash bath in the series is the one which is discarded. The presence of methanol is undesirable during reactions on the pins, but as it evaporates readily it can be easily removed by standing the block in a moving stream of air, such as the opening of an operating chemical fume hood. Methanol will dry more rapidly and the methanol-washed pins will take up less moisture from the air if the methanol is warm (e.g., prewarmed to 45°C in a closed bottle in a water bath). Other solvents (e.g., ether, petroleum ether, acetonitrile) should be the best available grade. Carrying out the correct synthesis of the peptides requires that all steps are performed with a very high level of attention to detail. All cyclically repeated steps (washes and deprotections) must be performed, and the activation and dispensing of the amino acids for each coupling cycle must be carried out exactly, or the peptides made may have the incorrect sequence, may be missing an amino acid, or may be truncated. Computerized equipment is available for assisting with the accurate dispensing of amino acids to the wells in a reaction tray (e.g., “Pin-Aid,” Chiron Technologies; Carter et al., 1992). The growing peptides must not be subjected to conditions that would prematurely block or deprotect the side chains (for example, from premature exposure to acetic anhydride or trifluoroacetic acid, which should be stored well away from where peptide synthesis is being performed). As a spot test for correct completion of all the steps of synthesis, it is wise to synthesize controls on each block of 96 pins. For noncleavable peptides, these controls can be peptide sequences that can be probed with an antibody known to react with the peptide. In this case, one of the two peptides should be a negative control, such as a randomized sequence. For cleavable peptides, the quantity and quality of the controls can be monitored by the usual techniques of HPLC (UNIT 11.6), amino acid analysis (UNIT 11.9), and mass spectrometry (Chapter 16). Ultimately, proof that an assay result is a function of the particular peptide made has to rely on a confirmatory experiment carried out with more highly-characterized
peptide or on analysis of a sample of the particular peptide used in the experiment. Once peptides have been made, they need to be handled and stored carefully to prevent degradation. Noncleavable peptides (pins) should be stored dry in a refrigerator after removal of any bound protein. If stored with desiccant they should be stable for months to years. Cleaved peptides can be stored frozen or as dry powder. After a long period of storage, it is wise to reassay controls or confirm the quality of the stored peptide by analysis. Another parameter critical to data from large numbers of peptides is to ensure that the identity of each peptide is properly tracked and that activity is not ascribed to the wrong peptide. Consistent use of the 8 × 12 microtiter plate format for synthesis, storage, assay, and use of computerized records for tracking all three processes can help avoid mistakes. Tracking and control is particularly easy if the assay data is read directly from a microtiter plate reader to a computer that is programmed with the peptide information because this method avoids manual data transcription.
Anticipated Results For a noncleavable pin-peptide synthesis, two control peptides, one of which is reactive with a monoclonal antibody in ELISA and the other serving as a nonbinding peptide control, should show the specific binding expected based on past data. For cleaved peptides, the yield of control peptide should be in the range expected from the stated pin loading (substitution level), e.g., 1 µmol for GAP and DKP kits or 5 µmol for the MPS kit. Purity of the cleaved controls should be consistent with the results of previous batches and should be of an acceptable standard. Testing of a systematic set of peptides in a bioassay can give data that is interpretable without recourse to additional controls, because a systematic set of peptides through a protein includes many sequences that are unlikely to be reactive sequences, i.e., they act as internal negative controls. Figure 18.2.7 shows one set of ELISA data from scanning noncleaved peptides with a monoclonal antibody. In screening for T helper cell responsiveness it is critical to include many control cultures, not only controls with no peptide added but also controls with nonstimulatory peptide. Systematic sets of peptides automatically include such controls (Reece et al., 1994).
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Time Considerations If amino acid coupling is carried out at 3 cycles/day, which can fit into a conventional working day, then it will take up to 2 weeks to make a set of 15-mers, as there is extra time required for side chain deprotection and drying down (depending on the peptide format). Although this may seem slow, the fact that hundreds or thousands of peptides can be made
simultaneously means that a project requiring large numbers of peptides is completed in a very short time. Indeed, the rate-limiting step may be the time it takes to carry out the assays on the large number of peptides when they become available. From this perspective, biotinylated peptides produced on glycine acid peptide (GAP), diketopiperazine (DKP), or multiple peptide syn-
+2-261,-
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Amino acid residue number
Synthesis of Multiple Peptides on Plastic Pins
Figure 18.2.7 Multipin capture ELISA. (A) Setup for multipin capture ELISA. Pins (gears) with peptides covalently attached are incubated in primary antibody, secondary antibody, and substrate developer in ELISA plates. The absorbance is measured and the resulting absorbance values are graphed versus peptide number, corresponding to the N-terminal residue number of the peptide in the protein sequence. (B) Peptide pin capture ELISA results with a monoclonal antibody against pins bearing octamer peptides of gonococcal pilin protein. All the peptides that show high readings contain a significant portion of the epitope. (Diagram courtesy of Dr. Fred Cassels, Walter Reed Army Institute of Research, Washington, D.C.)
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thesis (MPS) pins have a great advantage over the noncleavable peptide (NCP) pin-bound peptides, as the latter can only be assayed once a day, whereas hundreds of parallel assays can be carried out on all biotinylated peptides at once. Reading data directly into a computer enables the massive amounts of data to be stored efficiently for later analysis. Dispensing amino acids can be carried out efficiently by two people, one reading out the position into which the amino acid is to be dispensed and the other doing the actual dispensing. The passive partner (reader) can also act as a cross-checker to ensure no mistakes are made. If a computer-controlled pointing device is used, accuracy is improved and dispensing becomes a one-person operation. For large syntheses (>200 peptides), it is important that the dispensing be fast and accurate so that three couplings can be carried out per day.
Literature Cited Burrows, S.R., Gardner, J., Khanna, R., Steward, T., Moss, D.J., Rodda, S., and Suhrbier, A. 1994. Five new cytotoxic T cell epitopes identified within Epstein-Barr virus nuclear antigen 3. J. Gen. Virol. 75:2489-2493. Carter, J.M., VanAlbert, S., Lee, J., Lyons, J., and Deal, C. 1992. Shedding light on peptide synthesis. Bio/Technology 10:509-513. Fauchere, J.L. and Pliska, V. 1983. Hydrophobic parameters of amino acid side chains from the partitioning of N-acetyl-amino-acid amides. Eur. J. Med. Chem. 18:369-375. Geysen, H.M., Meloen, R.H., and Barteling, S.J. 1984. Use of peptide synthesis to probe viral antigens for epitopes to a resolution of a single amino acid. Proc. Natl. Acad. Sci. U.S.A. 81:3998-4002.
Geysen, H.M., Rodda, S.J., Mason, T.J., Tribbick, G., and Schoofs, P.G. 1987. Strategies for epitope analysis using peptide synthesis. J. Immunol. Methods 102:259-274. Maeji, N.J., Bray, A.M., and Geysen, H.M. 1990. Multi-pin peptide synthesis strategy for T cell determinant analysis. J. Immunol. Methods 134:23-33. Mutch, D.A., Rodda, S.J., Benstead, M., Valerio, R.M., and Geysen, H.M. 1991. Effects of end groups on the stimulatory capacity of minimal length T cell determinant peptides. Pept. Res. 4:132-137. Reece, J.C., Geysen, H.M., and Rodda, S.J. 1993. Mapping the major human T helper epitopes of tetanus toxin: The emerging picture. J. Immunol. 151:6175-6184. Reece, J.C., McGregor, D.L., Geysen, H.M., and Rodda, S.J. 1994. Scanning for T helper epitopes with human PBMC using pools of short synthetic peptides. J. Immunol. Methods 172:241254. Rink, H. 1987. Solid-phase synthesis of protected peptide fragments using a trialkoxydiphenylmethylester resin. Tetrahedron Lett. 28:37873790. Stewart, J.M. and Young, J.D. 1984. Solid Phase Peptide Synthesis, 2nd ed. Pierce Chemical Co., Rockford, Ill. Valerio, R.M., Bray, A.M., Campbell, R.A., Dipasquale, A., Margellis, C., Rodda, S.J., Geysen, H.M., and Maeji, N.J. 1993. Multipin peptide synthesis at the micromole scale using 2-hydroxyethyl methacrylate grafted polyethylene supports. Int. J. Pept. Protein Res. 42:1-9.
Contributed by Stuart J. Rodda Chiron Technologies Pty. Ltd. Victoria, Australia
Preparation and Handling of Peptides
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Synthetic Peptides for Production of Antibodies that Recognize Intact Proteins
UNIT 18.3
Antibodies that recognize intact proteins can be produced through the use of synthetic peptides based on short stretches of the protein sequence, without first having to isolate the protein. The procedure for selecting stretches of protein sequence likely to be antigenic is relatively straightforward. However, no procedure will identify a single sequence guaranteed to be effective, nor will it usually identify the best single sequence to use. Rather, several sequences will be identified that have a higher-than-average probability of producing an effective antigen. The steps to produce an effective antibody include: (1) designing the peptide sequence based on the sequence of the protein; (2) synthesizing the peptide; (3) preparing the immunogen either by coupling the synthetic peptide to a carrier protein or through the use of a multiple antigenic peptide (MAP); (4) immunizing the host animal; (5) assaying antibody titer in the host animal’s serum; and (6) obtaining the antiserum and/or isolating the antibody. This unit covers steps 1 and 3; step 2 requires a laboratory with expertise in peptide synthesis. Peptide synthesis services are widely available both academically and commercially. The best method to select potentially effective sequences is via a computer-assisted strategy (see Basic Protocol 1). An alternative manual method is also described (see Alternate Protocol 1) but is not recommended to replace the use of algorithms if there is a choice. A small synthetic peptide is usually insufficiently immunogenic on its own, and two methods have been developed to solve this problem. The first (see Basic Protocol 2) involves chemically coupling the synthetic peptide to a carrier protein to boost the immune response. The second method (see Alternate Protocol 2) entails direct synthesis of a MAP covalent multimer of the simple peptide sequence. Both methods have proven effective and it is a matter of personal preference which to use. Coupling to a carrier protein requires additional chemical manipulations after synthesis of the peptide, while the MAP is complete and ready for immunization at the conclusion of the synthetic protocol. Disadvantages of MAPs are that they are more difficult to produce homogeneously and to analyze postsynthetically. They also may be more prone to insolubility problems. A carrier protein is a relatively large molecule capable of stimulating an immune response independently. A synthetic peptide coupled to a carrier protein acts as a hapten and produces antibodies specific for the hapten (antibodies against the carrier protein are also produced). The most commonly used carrier proteins are keyhole limpet hemocyanin (KLH) and bovine or rabbit serum albumin (BSA or RSA). KLH is usually preferred, because it tends to elicit a stronger immune response and is evolutionarily more remote from mammalian proteins. A common problem with KLH, however, has been its solubility. Pierce Chemical Company sells a preparation of KLH purported to have better solubility properties (see below). Alternatively, peptides can be coupled to carrier proteins through either their amino (see Alternate Protocol 3) or carboxyl groups (see Alternate Protocol 4). These two alternate protocols are not recommended as a first choice for coupling, but are included because they have been used successfully and may be advantageous for certain special applications discussed in the Commentary. Also presented are methods for assaying free sulfhydryl content and for reducing disulfide bonds in synthetic peptides (see Support Protocols 1 and 2).
Contributed by Gregory A. Grant Current Protocols in Protein Science (2002) 18.3.1-18.3.19 Copyright © 2002 by John Wiley & Sons, Inc.
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Once the coupling procedure has been performed, it is possible to determine the approximate degree of coupling by amino acid analysis (see Support Protocol 3). However, in most instances this is unnecessary and the product can be used directly. BASIC PROTOCOL 1
COMPUTER-ASSISTED SELECTION OF APPROPRIATE ANTIGENIC PEPTIDE SEQUENCES An antibody produced in response to a simple linear peptide will most likely recognize a linear epitope in a protein. Furthermore, that epitope must be solvent-exposed to be accessible to the antibody. The general features of protein structure that correspond to these criteria are turns or loop structures, which are generally found on the protein surface connecting other elements of secondary structure, and areas of high hydrophilicity, especially those containing charged residues. As a consequence, computer algorithms that predict protein hydrophilicity and tendency to form turns are very useful. Several analytic programs or algorithms that attempt to do this have been developed. Although the choice of method may rely on availability or personal preference, there tends to be a high level of agreement among them. As stated earlier, none of the methods will identify the one single sequence guaranteed to produce an effective antibody against any given protein. Rather, the methods will offer several good candidates, one or several of which can be used. Many of these algorithms may already be available on a local computer system. They are included in many commercial software packages such as GCG (Genetics Computer Group; see SUPPLIERS APPENDIX). The ExPASy Web site of the University of Geneva offers free access to a variety of different programs over the Internet at http://expasy.org/tools. The following protocol utilizes the hydropathy index developed by Kyte and Doolittle (1982) and the secondary structure prediction method for β turns developed by Chou and Fasman (1974) found in the tool “Protscale” at the ExPASy Internet address. 1. Using the selected algorithms, compute the hydropathy index and the tendency for β-turns of the protein sequence. Use a window size of 7 or 9 and give equal weight to each amino acid. Record the results in either graphical or numerical form, or both. As an example, the graphical representation of these results for the protein sequence shown in Figure 18.3.1 is presented in Figure 18.3.2. A window size determines the number of amino acids to be used in computing a value for the amino acid at the center of the window. For example, a window size of 9 includes 4
10
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MAKVSLEKDK IKFLLVEGVH QKALESLRAA GYTNIEFHKG ALDDEQLKES IRDAHFIGLR SRTHLTEDVI NAAEKLVAIG CFCIGTNQVD LDAAAKRGIP VFNAPFSNTR SVAELVIGEL 120 LLLLRGVPEA NAKAHRGVWN KLAAGSFEAR GKKLGIIGYG HIGTQLGILA ESLGMYVYFY 180 DIENKLPLGN ATQVQHLSDL LNMSDVVSLH VPENPSTKNM MGAKEISLMK PGSLLINASR 240 GTVVDIPALC DALASKHLAG AAIDVFPTEP ATNSDPFTSP LCEFDNVLLT PHIGGSTQEA 300 QENIGLEVAG KLIKYSDNGS TLSAVNFPEV SLPLHGGRRL MHIHENRPGV LTALNKIFAE 360 QGVNIAAQYL QTSAQMGYVV IDIEADEDVA EKALQAMKAI PGTIRARLLY
Selecting Synthetic Peptides for Production of Antibodies
Figure 18.3.1 The amino acid sequence of a 410-residue protein analyzed by the method presented in Basic Protocol 1. The results are shown in Figure 18.3.2.
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amino acids on each side of the central amino acid. The value computed for the central amino acid is the simple average of the values for each amino acid in the window.
2. Compare the results of the two analyses and look for areas of sequence that are high in turn tendency and high in hydrophilicity (low in hydrophobicity). In Figure 18.3.2, these areas correspond to positive peaks in the Chou-Fasman analysis and negative peaks in the Kyte-Doolittle analysis. The three best areas in terms of amplitude and correlation are shaded. These correspond to the sequences underlined in Figure 18.3.1. (Note the alignment of these peak optima as compared to the peaks around residue 300.)
A
1.3 1.2
Score
1.1 1.0 0.9 0.8 0.7 0.6
B
3
2
Score
1
0
–1
–2 50
100
150
200
250
300
350
400
Position
Figure 18.3.2 Graphical representation of the results generated by a computer algorithm for the sequence in Figure 18.3.1, analyzed by the method presented in Basic Protocol 1. The shaded areas represent three regions in the sequence meeting criteria for selection as potential immunogens. (A) Analysis for β turns (Chou and Fasman, 1974). (B) Analysis for hydrophobicity (Kyte and Doolittle, 1982).
Preparation and Handling of Peptides
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3. Examine the sequences for glycosylation site motifs and discard any sequences that contain them unless it is known that the protein is not glycosylated. Amino acids in glycosylated regions may be shielded from presentation to an antibody by masking carbohydrates. Amino-linked carbohydrate chains can occur at Asn-X-Ser or Asn-X-Thr sequences. Hydroxyl-linked carbohydrate chains do not appear to have a set motif. A program to assist in the prediction of mucin-type GalNAc O-glycosylation sites in mammalian lipoproteins is found in the tool “NetOGlc” at the Expasy site (http://expasy.org/tools). However, before using read the documentation carefully and keep in mind that such prediction methods cannot always be successful.
4. Select the best sequences resulting from this analysis to use as antigenic peptides. These are sequences where the largest positive values (peaks with positive deflection) for turn propensity correspond in position to the largest negative values (peaks with negative deflection) for hydrophobicity. The values obtained in these analyses are relative and dependent on the individual protein’s composition, so it is not possible to set an arbitrary minimum value as a cutoff for rejecting a particular peak. Rather, always select the peaks of greatest magnitude in any given sequence. In addition, the immediate amino-terminal and carboxyl-terminal regions of proteins are often exposed to solvent. If these areas appear to be hydrophilic in nature, they are also acceptable candidates. Thus each analysis may provide several potential sequences. How many peptides to make (see Anticipated Results) is a matter of individual choice. ALTERNATE PROTOCOL 1
MANUAL INSPECTION TO SELECT APPROPRIATE PEPTIDE SEQUENCES If computer algorithms are not available, it is possible to select potential sequences by manual inspection. Although there is no evidence that a manual method is any less effective than the use of computer algorithms, there is a greater probability of overlooking potentially important areas of sequence. It is therefore recommended that computer analysis be used whenever it is available. Although it can be done, it would be very time consuming and labor intensive to manually calculate values for every overlapping peptide offset by a single amino acid in the same way that the algorithms do. For this reason, areas rich in polar residues are selected for manual calculation of hydrophilicity and turn propensity. 1. Visually inspect the protein sequence and select areas that contain at least two to three charged residues (Lys, Arg, His, Asp, Glu) within a 10- to 15-residue span. If this criterion cannot be met, select sequences with the greatest number of charged residues.
2. From the sequences identified in step 1, select a subset of sequences that are the highest in Ser, Thr, Asn, Gln, Pro, and Tyr content. 3. Calculate average hydrophilicity and turn propensity for each amino acid in the selected sequences using the values given in Table 18.3.1 and a window of 9 residues (see Basic Protocol 1, step 1). Be sure to include the residues flanking the selected sequence for calculation of values for the residues at the ends of the selected sequence. In other words, do not use different size windows.
4. Plot the values for each amino acid of a chosen sequence. Selecting Synthetic Peptides for Production of Antibodies
Sequences whose optimal values for hydrophilicity and turn propensity correspond (as in Fig. 18.3.2) are considered good candidates.
5. Inspect sequences for glycosylation motifs and discard these candidates (see Basic Protocol 1, step 3).
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Table 18.3.1
Hydrophobic and β-Turn Indices of Amino Acids
Amino acid
Symbols
Arginine Lysine Aspartic acid Glutamic acid Asparagine Glutamine Histidine Proline Tyrosine Tryptophan Serine Threonine Glycine Alanine Methionine Cysteine Phenylalanine Leucine Valine Isoleucine
Arg (R) Lys (K) Asp (D) Glu (E) Asn (N) Gln (Q) His (H) Pro (P) Tyr (Y) Trp (W) Ser (S) Thr (T) Gly (G) Ala (A) Met (M) Cys (C) Phe (F) Leu (L) Val (V) Ile (I)
aKyte
Hydrophobicity β-turn valuea propensityb −4.5 −3.9 −3.5 −3.5 −3.5 −3.5 −3.2 −1.6 −1.3 −0.9 −0.8 −0.7 −0.4 1.8 1.9 2.5 2.8 3.8 4.2 4.5
0.95 1.01 1.46 0.74 1.56 0.98 0.95 1.52 1.14 0.96 1.43 0.96 1.56 0.66 0.60 1.19 0.60 0.59 0.50 0.47
and Doolittle (1982). and Fasman (1974).
bChou
Amino acids of glycosylated regions may be masked in native proteins, so an antibody raised against them would be ineffective.
6. Select the best sequences (see Basic Protocol 1, step 4 for criteria), choosing a high turn-propensity-to-hydrophobicity ratio. DESIGNING A SYNTHETIC PEPTIDE FOR COUPLING TO A CARRIER PROTEIN
BASIC PROTOCOL 2
Although there is no direct evidence to show that the state of the termini of the peptide affects its ability to produce antibodies that will react with the protein, most procedures suggest that the termini of the peptide should mimic their native state. Thus, sequences whose terminal residues normally are in peptide linkage in the protein can have their amino-terminal and carboxyl-terminal groups modified by acetylation and amidation, respectively, during synthesis. Modification of the amino or carboxyl termini will decrease the polarity of the peptide in solution and could have a significant effect on the peptide’s solubility. If the peptide lacks sufficient protonatable side chains, modification of the termini can be omitted. A general rule to predict solubility is that the total number of charges at a given pH should be at least 20% of the number of residues in the peptide. 1. Choose a sequence of 10 to 15 amino acid residues for the synthetic peptide. Longer peptides are more difficult and expensive to make, and they are usually unnecessary.
Preparation and Handling of Peptides
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Try to choose a stretch of sequence that contains some charged residues such as Arg, Lys, His, Glu, and Asp. In addition to the high likelihood of these amino acids being located on the surface of the protein, they aid handling of the synthetic product by promoting solubility.
2a. If the selected sequence does not contain an internal cysteine: Place a cysteine on either the amino or carboxyl terminus for use in coupling to a carrier protein with a heterobifunctional cross-linking reagent such as MBS, m-maleimidobenzoyl-N-hydroxysuccinimide ester (see Basic Protocol 3). Cross-linking with heterobifunctional reagents is the recommended procedure for most peptides (see Basic Protocol 3). As an alternative to using a chemical cross-linking reagent, any peptide, regardless of amino acid content, can also be photochemically linked to a carrier protein if p-benzoyl benzoic acid is added to the peptide during synthesis (see Alternate Protocol 5). Although photochemical cross-linking is effective, it is not widely used. If the sequence includes the immediate amino or carboxyl terminal sequence of the protein, the cysteine should be placed on the end that would normally be engaged in the internal peptide bond. For sequences internal to the protein, the cysteine may be placed at either end according to the preference of the synthetic chemist. However, if amino-terminal capping (acetylation) is used after the coupling of each amino acid during synthesis, it is preferable to place the cysteine on the amino-terminal end of the peptide since then only the full-length peptide will contain the cysteine residue. In this way, if synthetic difficulties are encountered, only the full-length peptide will couple to the carrier. If placed on the carboxyl terminal end, the cysteine residue tends to racemize during synthesis unless a chlorotrityl resin is used. However, this should not have an effect on the rest of the peptide or the generation of antibodies.
2b. If the sequence contains an internal cysteine residue: Do not add a terminal cysteine for MBS cross-linking. Rather, use an alternative coupling procedure (see Alternate Protocols 3, 4, or 5) or synthesize a multiple antigenic peptide (MAP; see Alternate Protocol 2). Internal cysteine sulfhydryl groups will also cross-link to the carrier protein, and multiple cysteines will result in a peptide attached at multiple points. If the sulfhydryl will eventually be important for antibody recognition of the protein, the immunization may not produce effective antibodies. Furthermore, the additional constraint produced by the existence of multiple points of coupling may affect the ultimate ability of the antibody to recognize the protein.
3a. For sequences whose terminal residues are in peptide linkage within the protein: If the peptide is coupled using a heterobifunctional reagent such as MBS (see Basic Protocol 3), modify the amino and carboxyl ends by acetylation and amidation, respectively, during the synthetic procedure. If coupling is performed with a homobifunctional reagent that reacts with amino groups such as glutaraldehyde (see Alternate Protocol 3) or by the photochemical method (see Alternate Protocol 5), only amidate the carboxyl terminus. If coupling is performed with EDC (see Alternate Protocol 4), only acetylate the amino terminus. 3b. For sequences that are amino or carboxyl terminal to the protein:
Selecting Synthetic Peptides for Production of Antibodies
i. If the sequence is the immediate amino or carboxyl terminal sequence of the protein and the peptide will be coupled with a heterobifunctional reagent such as MBS (see Basic Protocol 3), leave the end that is not in peptide linkage (and the end that does not contain the additional cysteine residue for MBS coupling) as the free amino or carboxyl group unless it is known that they are normally blocked. ii. If the sequence is the immediate amino terminal sequence of the protein and the peptide will be coupled with a homobifunctional reagent that reacts with amino
18.3.6 Supplement 28
Current Protocols in Protein Science
NH 2
aan C
O NH
H N
aa3 O
HO
O
C
O C
C
NH
NH β-Ala
aa2 O C N
O C
N
O
ε
Lys
H C N
H
C O H
O aa3
O
C
H N
C
Lys H Nε
aa2
O
NH
H
aa1
aan
NH 2
aa3
C
H2N
aan
O
H N α
aa1
O
O
H
C
Nα
C N
C
Lys
NαH
NεH
C O
C O
aa1
aa1
NH
NH
C O
C O
aa2
aa2
H N
O C
aa3
H N
C
aan
NH2
O
Figure 18.3.3 Representation of a four-branched multiple antigenic peptide (MAP). The MAP core is usually synthesized with a β-alanine residue attached to the solid phase support followed by a scaffold of lysine residues. The first lysine residue, attached through its α-carboxyl group to β-alanine, provides two primary amino groups (α and ε) for chain elongation. The next level of the lysine scaffold consists of two lysine residues which in turn provide four primary amino groups for chain elongation. MAPs with eight branches are formed by providing one additional layer of lysine to that depicted here.
groups such as glutaraldehyde (see Alternate Protocol 3) or photochemically (see Alternate Protocol 5), amidate the C-terminus. iii. If the sequence is the immediate carboxyl terminal sequence of the protein and the peptide will be coupled with glutaraldehyde (see Alternate Protocol 3) or photochemically (see Alternate Protocol 5), leave both termini free. 4. If the peptide will be coupled with 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC; see Alternate Protocol 4), block the amino terminus by acetylation. This should be done even if the sequence is at the immediate amino terminus of the protein, since treatment with EDC can result in the production of covalent multimers of the peptide through reaction with the N-terminal amino group. Some procedures recommend using citraconylation to temporarily protect the amino group. However, this also adds a free carboxyl group that could result in multiple site attachment of the peptide to the carrier.
Preparation and Handling of Peptides
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ALTERNATE PROTOCOL 2
DESIGNING A SYNTHETIC MULTIPLE ANTIGENIC PEPTIDE A multiple antigenic peptide (MAP; Posnett et al., 1988; Tam, 1988) is an effective alternative to coupling a simple linear peptide to a carrier protein. MAPs are covalent constructs consisting of a simple peptide sequence synthesized on a branched core (Fig. 18.3.3) with one copy of the peptide sequence on each of four or eight branches. One advantage of a MAP is that it is suitable for use as an immunogen at the conclusion of the synthetic process. On occasion, MAPs have produced effective protein antibodies when the conventional peptide coupled to carrier protein has not. Thus, MAPs represent an effective alternative approach for antiserum production. 1. Select a sequence between 10 and 15 residues in length. Longer sequences are unnecessary and increase the probability of synthetic problems. The presence of internal cysteine residues are not a concern with MAPs, but if present, take precautions to keep them reduced.
2. Synthesize the MAPs utilizing a four-branch core. Synthesize the MAP core de novo, or purchase resins for solid-phase peptide synthesis with four- or eight-branched cores (available commercially from Advanced ChemTech, Novabiochem, Applied Biosystems, AnaSpec, and Bachem Bioscience). Eight-branched cores are suitable if the peptide is no more than 12 to 14 residues and has a high degree of hydrophilicity. There are more synthetic problems with eight-branched MAPs, presumably due to the higher density of structure during synthesis: they are more difficult to characterize and probably raise a diverse antibody population against some synthetic artifacts (see Mints et al., 1997).
3. Optional. If the selected sequence was not the amino terminus of the protein, acetylate the new amino terminus. In the case of a MAP, the carboxyl terminus will remain in covalent linkage to the branched core.
4. Use the MAP directly as an immunogen. Coupling to a carrier protein as described in Basic Protocol 3 is usually not necessary. BASIC PROTOCOL 3
COUPLING SYNTHETIC PEPTIDES TO A CARRIER PROTEIN USING A HETEROBIFUNCTIONAL REAGENT If the synthetic peptide was designed with a cysteine residue at one terminus (see Basic Protocol 2, step 2a), the following procedure should be followed for coupling to keyhole limpet hemocyanin (KLH) or other carrier proteins. Care must be taken to assure that the cysteine sulfhydryl group has remained reduced. Under normal synthetic conditions, if the peptide was lyophilized and stored dry immediately after synthesis, the sulfhydryl usually remains in the reduced state. The presence of free sulfhydryl groups in the peptide can be determined with Ellman’s reagent (see Support Protocol 1) just prior to use; alternatively, high-resolution mass spectrometry can be used. If reduction is needed, follow the cysteine reduction procedure (see Support Protocol 2) before starting the coupling process.
Selecting Synthetic Peptides for Production of Antibodies
The reagent most commonly used for this purpose is m-maleimidobenzoyl-N-hydroxysuccinimide ester (MBS). However, several related reagents (all available from Pierce) offer some additional features. Sulfo-MBS (Pierce) is a water-soluble alternative to MBS. Succinimidyl 4-(N-maleimidomethyl)-cyclohexane-1-carboxylate (SMCC) and its sulfonated analog sulfo-SMCC provide the same chemistry with a more pH-stable maleimide (see step 1). The MBS and SMCC reagents can be used interchangeably in this protocol (the sulfo reagents can be dissolved in aqueous solution, while the others must be dissolved in an organic solvent; the concentrations listed are appropriate for all four).
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Materials Keyhole limpet hemocyanin (KLH; Pierce, Sigma, Calbiochem, or Boehringer Mannheim) 0.01 M sodium phosphate buffer, pH 7.5 (APPENDIX 2E) 10 mg/ml MBS in fresh N,N-dimethylformamide (DMF) 0.05 M and 0.1 M sodium phosphate buffer, pH 7.0 (APPENDIX 2E) Synthetic peptide with a reduced cysteine residue at either the N- or C-terminus 6 M guanidine⋅HCl (see recipe) Small glass vial with flat bottom ∼0.9 × 15–cm gel filtration column with Sephadex G-25 or G-50 (Pharmacia Biotech) or Bio-Gel P2 or P4 (Bio-Rad) resin; or prepacked PD-10 column (Pharmacia Biotech) Additional reagents and equipment for gel filtration chromatography (UNIT 8.3) NOTE: Do not use Tris or other buffers with primary amino groups in this procedure. CAUTION: MBS is a moisture-sensitive irritant. Read the Material Safety Data Sheet before use. 1. Dissolve 5 mg KLH in ∼0.5 ml of 0.01 M sodium phosphate buffer, pH 7.5, in a small, flat-bottomed vial. A pH range of 7.0 to 7.5 offsets competing reactions. Although the unprotonated form of the amine reacts with the N-hydroxysuccinimide ester and would be optimal at pH >8.0, hydrolysis of the ester bond and reaction of the maleimide group with amines is enhanced at higher pH.
2. Add 100 µl of 10 mg/ml MBS/DMF solution and stir gently with a micro stir-bar 30 min at room temperature. A small amount of precipitate may form during this procedure and is acceptable. However, if the precipitate is large, perform the procedure again with fresh components. As an alternative to performing this coupling procedure from scratch, it is possible to purchase MBS-activated KLH (Pierce; Boehringer Mannheim) or kits containing MBS-activated KLH and an alternate MBS-activated protein for use in ELISA assays (Pierce).
3. Separate MBS-activated KLH from free MBS on a ∼0.9 × 15–cm gel filtration column, equilibrating and eluting the column with 0.05 M sodium phosphate buffer, pH 7.0. Collect 0.5-ml fractions and read their absorbance at 280 nm (UNIT 8.3). The first peak to elute is the KLH-MBS conjugate. These fractions may appear cloudy. The second peak is uncoupled MBS.
4. Pool the KLH-MBS conjugate fractions in a separate tube. 5. Dissolve 5 mg of the synthetic peptide in 0.01 M sodium phosphate buffer, pH 7.0, immediately prior to use. If the peptide is poorly soluble, use 6 M guanidine⋅HCl. Maleimide groups react specifically with sulfhydryls at slightly acid to neutral pH.
6. Add the peptide solution to the KLH-MBS conjugate. Stir gently with a micro stir-bar 3 hr at room temperature. The coupling may be continued overnight.
7. Dialyze against 4 liters distilled water overnight at 4°C. Use for immunizations within 24 hr. Preparation and Handling of Peptides
18.3.9 Current Protocols in Protein Science
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Selecting Synthetic Peptides for Production of Antibodies
18.3.10
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Current Protocols in Protein Science
O
O
N O C
O
NO2
OH (DTT)
OH
SH
COOH (Ellman's Reagent)
S S
O
HS CH2 CH CH CH2 SH
HOOC
O2N
C OH
O
N
O
R1 NH
OH
HOOC
O2N
R2
N
C O C
O
KLH
N OH
O
O
N
O
KLH
S S
NH2
OH
SH
S CH2 H C C H OH
S H2C OH
HOOC
O N C H
SH
KLH
KLH
(A = 412 nm)
C N H
O
O2N
SH
C R1 N N R2 H H
O
N C (CH2)3 C N H H
O
N C H
S SCH2 CH CH CH2 SH
O
O
KLH
O
N
O
O
S
Figure 18.3.4 Representations of the chemistries described in this unit. (A) Cross-linking with MBS; (B) cross-linking with glutaraldehyde; (C) cross-linking with EDC; (D) reaction of free sulfhydryls with Ellman’s reagent to produce a colorimetric product; (E) reduction of disulfides with DTT. Abbreviations: KLH, keyhole limpet hemocyanin; MBS, m-maleimidobenzoyl-N-hydroxysuccinimide ester; EDC, 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide; DTT, N,N-dithiothreitol.
S
S
E
SH
D
O
O
NH2 HC (CH2)3 CH H2N
NH2
R1 N C N R2 (EDC)
C
KLH
B
KLH
A
ASSAY OF FREE SULFHYDRYLS WITH ELLMAN’S REAGENT Free sulfhydryl content of a peptide can be quantitatively determined with Ellman’s reagent, 5,5′-dithio-bis(2-nitrobenzoic acid). The molar extinction coefficient at 412 nm of thionitrobenzoate, the colored species generated when the reagent reacts with a free thiol, is 14,150 in 0.1 M sodium phosphate buffer (see Fig. 18.3.4). The sensitivity of the reaction is in the low nmol/ml range for sulfhydryl groups, making it well suited for synthetic peptides. By dry weight most synthetic peptides are only ∼60% to 75% peptide, the remainder consisting of counterions and water of hydration. Amino acid analysis (see UNIT 11.9) is needed to establish the actual peptide content unambiguously, but such precise measurement is not usually necessary for qualitative evaluation of the free sulfhydryl content of a peptide sample.
SUPPORT PROTOCOL 1
Materials Cysteine standard stock solution (see recipe) 0.1 M sodium phosphate, pH 8.0 (APPENDIX 2E) Peptide to be assayed Ellman’s reagent solution (see recipe) 13 × 100–mm glass test tubes 1. Prepare a cysteine standard curve by adding 25 µl, 50 µl, 100 µl, 150 µl, 200 µl, and 250 µl of cysteine standard stock solution to separate 13 × 100–mm tubes. Add ≤250 µl of each peptide to be tested to separate tubes. Bring the volume in each tube to 250 µl with 0.1 M sodium phosphate, pH 8.0. Add 250 µl of 0.1 M sodium phosphate, pH 8.0, to a blank tube. The cysteine content of the peptide to be assayed should fall within the range of the standard curve (37.5 to 375 nmol).
2. Add 50 µl Ellman’s reagent solution and 2.5 ml of 0.1 M sodium phosphate, pH 8.0, to each tube. Mix and incubate 15 min at room temperature. 3. Measure absorbance at 412 nm (A412). 4. Plot the A412 values of the standards after subtracting the value for the blank to produce a standard curve. Use this curve to determine the free sulfhydryl content of the peptides. REDUCING CYSTEINE GROUPS IN PEPTIDES When a peptide is synthesized with a terminal cysteine residue to be used for coupling with MBS (see Basic Protocol 3), the cysteine must be in the reduced state (present as free -SH rather than as a disulfide) in order to participate in the reaction with the coupling reagent. If peptides are lyophilized immediately after extraction from the resin cleavage cocktail or reversed-phase HPLC and used immediately after reconstitution, oxidation of cysteine side chains is usually not a problem. However, if oxidation to disulfides has occurred, the peptide can be reduced prior to use with the protocol presented here.
SUPPORT PROTOCOL 2
Dithiothreitol (DTT) is preferred to 2-mercaptoethanol (2-ME) as a reducing agent because its lower redox potential allows it to be effective at lower concentrations, and the reaction goes to completion because formation of the six-membered ring containing an internal disulfide is energetically favorable (see Fig. 18.3.4; also see APPENDIX 3A). To determine if reduction is necessary, quantitate the level of free sulfhydryl groups with Ellman’s reagent (see Support Protocol 1). Additional methods for reducing disulfides include using sodium borohydride (Gailit, 1993) and Tris(2-carboxyethyl)phosphine (TCEP; Getz et al., 1999).
Preparation and Handling of Peptides
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Materials Synthetic peptide 0.1 M sodium phosphate, pH 8.0 (APPENDIX 2E) 1 M aqueous dithiothreitol (DTT) 1 N HCl 100- or 250-µl polypropylene tubes Nitrogen gas source Additional reagents and equipment for reversed-phase HPLC of peptides (see UNIT 11.6) 1. Dissolve 5 to 10 mg of peptide in 0.1 M sodium phosphate, pH 8.0. 2. Add 100 µl of 1 M DTT. 3. Flush nitrogen over the surface of the liquid, seal the tube, and incubate 1 hr at 37°C. 4. Acidify with 1 N HCl and desalt by reversed-phase HPLC (UNIT 11.6). 5. Pool peptide fractions and lyophilize. Store lyophilized at 4°C until ready to use (up to several days). The oxidation state of the peptide can usually be followed by analytical monitoring of its elution position on reversed-phase HPLC. Disulfide-linked dimers of peptides generally elute later than the monomeric peptide. ALTERNATE PROTOCOL 3
COUPLING SYNTHETIC PEPTIDES TO A CARRIER PROTEIN USING A HOMOBIFUNCTIONAL REAGENT The available homobifunctional reagents couple compounds through primary amino groups. Therefore, peptides with internal lysine residues should not be used in this procedure. The reagent most commonly used for this procedure is glutaraldehyde, but it should not be used with peptides containing internal Cys, Tyr, or His residues. Other homobifunctional cross-linking reagents that can be used in the same way as glutaraldehyde, but do not cross react with Cys, Tyr, or His residues, are also available: disuccinimidyl suberate (DSS), disuccinimidyl glutarate (DSG), and bis(sulfosuccinimidyl) suberate (BS3; all available from Pierce). However, these reagents are not widely employed for coupling peptides to proteins and are not considered the method of choice, and methods for their use have not been formalized. This is probably because coupling of the synthetic peptide to itself and aggregation of the carrier protein can occur with homobifunctional reagents such as these. Glutaraldehyde, on the other hand, although also subject to this limitation, has generally been used successfully. Additional Materials (also see Basic Protocol 3) 50 mM sodium borate buffer, pH 8.0 (pH adjusted with HCl) Glutaraldehyde solution (see recipe) 1 M glycine in 50 mM sodium borate buffer, pH 8.0 NOTE: Do not use Tris or other buffers with primary amino groups in this procedure. 1. Dissolve 5 mg KLH in ∼1.0 ml of 50 mM sodium borate buffer, pH 8.0. 2. Add 5 mg synthetic peptide.
Selecting Synthetic Peptides for Production of Antibodies
3. Slowly add 1 ml fresh glutaraldehyde solution with gentle mixing at room temperature. Allow to react for an additional 2 hr with gentle mixing. Formation of a yellowish color or milkiness is normal and does not affect the sample.
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4. Add 0.25 ml of 1 M glycine to bind unreacted glutaraldehyde. A darker yellow to brown color may develop.
5. Dialyze the reaction mixture overnight at 4°C against 4 liters of 50 mM sodium borate buffer, pH 8.0, and then overnight against water. Use immediately. COUPLING SYNTHETIC PEPTIDES TO A CARRIER PROTEIN USING A CARBODIIMIDE
ALTERNATE PROTOCOL 4
This procedure couples amino groups to carboxyl groups by way of activation of the carboxyl group with a water-soluble carbodiimide. Since the procedure is most easily performed in one step, peptides containing internal Asp, Glu, Lys, Tyr, or Cys residues should not be used. Also, in order to avoid making polymers of the peptide, the amino terminus should be blocked by acetylation during synthesis (see Basic Protocol 2). While this method has been used successfully, it is not considered to be the method of choice except in special situations. Additional Materials (also see Basic Protocol 3) 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC; Pierce), used fresh or stored desiccated and frozen 0.1 N HCl NOTE: Buffers containing amino or carboxyl groups should not be used in this procedure. According to some reports, buffers containing phosphate groups should also be avoided. Water is the safest choice as a solvent. 1. Dissolve 5 mg synthetic peptide in 1 ml water. 2. Add 25 mg EDC and carefully adjust pH to 4.0 to 5.0 by adding small amounts of 0.1 N HCl. Allow to react for 5 to 10 min at room temperature with gentle mixing. pH paper suffices to monitor this adjustment.
3. Dissolve 5 mg KLH in 0.5 ml water and add to solution from step 2. React 2 hr at room temperature with gentle mixing. 4. Dialyze against 4 liters of water overnight at 4°C. Use immediately. COUPLING SYNTHETIC PEPTIDES TO A CARRIER PROTEIN PHOTOCHEMICALLY
ALTERNATE PROTOCOL 5
Most synthetic peptides today are made with 9-fluorenylmethyloxycarbonyl (Fmoc) chemistry (see Fig. 18.1.1), and therefore the following protocol links the photoactive group to the free amino terminus of the peptide. An alternative approach if t-butyloxycarbonyl (Boc) chemistry is used is to link it to an ε-amino group of a terminal lysine (Gorka et al., 1989). Additional Materials (see also Basic Protocol 3) 4-benzoyl benzoic acid (Sigma or Aldrich) Quartz spectrophotometry cuvettes 1. Have the synthetic chemist who is making the peptide attach a benzoyl benzoic acid group to the amino terminus simply by treating the reagent as the terminal residue during normal synthesis. This blocks the amino terminus.
Preparation and Handling of Peptides
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2. Dissolve 5 mg KLH in ∼0.5 ml of 0.01 M sodium phosphate buffer, pH 7.5. 3. Add 2 mg of synthetic peptide containing the benzoyl benzoate adduct. 4. Place in a 1-cm quartz cuvette and irradiate with 366-nm light for 3 hr at a distance of 0.5 cm. Use immediately. The peptide can be used directly for immunization. SUPPORT PROTOCOL 3
CALCULATION OF THE MOLAR RATIO OF PEPTIDE TO CARRIER PROTEIN The molar ratio of peptide to carrier protein coupling efficiency can be calculated to determine the level of substitution achieved by the coupling procedure. This information can be obtained using the results of amino acid compositional analysis. By performing the calculations presented in this protocol, the molecules of peptide in the conjugate per molecule of carrier protein in the conjugate can be determined. 1. Obtain the amino acid composition of the carrier protein, the peptide, and the peptide/carrier conjugate. Amino acid compositional analysis (of these hydrolysates) is usually available at sources that provide automated peptide synthesis (see UNITS 18.1 & 18.2). Be sure that the conjugate is free of unconjugated peptide (i.e., it should be well dialyzed). 2. Determine a scaling factor (SF) that relates the moles of protein in the unconjugated carrier protein to the moles of protein in the peptide/carrier conjugate. This is done by comparing the molar ratio of ≥3 amino acids present in the carrier protein and peptide/carrier conjugate but not present in the peptide. For example, if the peptide TGLRDSC (Table 18.3.2) is coupled to a carrier protein, choose A, K, and I. The calculation is done as: Table 18.3.2 Sample Calculation of the Extent of Coupling of the Peptide TGLRDSC to Carrier Proteina
Amino acid
Selecting Synthetic Peptides for Production of Antibodies
D E G S T H P A M V F L I C Y K R Total pmol amino acid
Composition of carrier protein
Composition of peptide/carrier conjugate
Amount of carrier protein amino acids in conjugate
Amount of peptide amino acids in conjugate
80 110 95 65 70 10 25 103 5 60 22 55 65 7 13 65 75
185 222 215 150 163 19 51 206 11 118 45 133 135 22 25 125 177
160 220 190 130 140 20 50 206 10 120 44 110 130 14 26 130 150
25 — 25 20 23 — — — — — — 23 — 8 — — 27
925
2002
1850
151
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SF = [(pmol A in conjugate/pmol A in carrier) + (pmol K in conjugate/pmol K in carrier) + (pmol I in conjugate/pmol I in carrier)]/3. For these amino acids, the carrier protein yields are as follows: A = 103 pmol, K = 65 pmol, and I = 65 pmol. For these same amino acids, the peptide/carrier conjugate yields: A = 206 pmol, K = 125 pmol, and I = 135 pmol. From these values, the relative amount of carrier protein in the conjugate versus the unconjugated carrier protein (SF) can be calculated as follows: (206/103 + 125/65 + 135/65)/3 = 2.0, indicating that there is twice as much carrier protein in the peptide/carrier conjugate hydrolysate as in the carrier-protein hydrolysate. 3. Calculate the moles of peptide present in the conjugate by subtracting the moles of amino acid present in the carrier from the moles of amino acid present in the conjugate. Choose ≥3 amino acids present in the peptide. The relative amount (SF) of protein present in the carrier protein versus the amount in the conjugate as calculated in step 2, must also be considered as follows: pmol peptide in conjugate = {[pmol G in conjugate − (SF × pmol G in carrier)]+ [pmol L in conjugate − (SF × pmol L in carrier)]+ [pmol R in conjugate − (SF × pmol R in carrier)]}/3
Therefore, the amount of peptide in the conjugate hydrolysate for the example shown in Table 9.4.1, calculated using the amino acids G, L, and R, is {[215 − (2 × 95)] + [133 − (2 × 55)] + [177 − (2 × 75)]}/3 = 25 pmol. 4. Calculate the number of moles of protein in the conjugate hydrolysate as follows:
pmol carrier protein in conjugate =
(total pmol carrier protein amino acids) × 110 molecular weight of carrier protein
where total pmol carrier protein amino acids = SF × (total amino acid composition of carrier in pmol) and 110 is the average molecular weight of an amino acid. In this example, there are 1850 pmol of carrier protein amino acids in the conjugate; therefore, 1850 pmol × (110/100,000) = 2.04 pmol carrier protein in conjugate. 5. Determine the ratio of peptide to carrier protein as follows: molecules peptide in conjugate/molecules carrier protein in conjugate = pmol peptide in conjugate/pmol carrier protein in conjugate. Using the values calculated in steps 3 and 4, the result is: 25 pmol peptide in conjugate/2.04 pmol carrier protein in conjugate = 12.2 molecules peptide in conjugate per molecule carrier protein in conjugate.
Preparation and Handling of Peptides
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Supplement 28
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Cysteine standard stock solution Dissolve 26.3 mg cysteine hydrochloride monohydrate in 100 ml of 0.1 M sodium phosphate, pH 8.0 (APPENDIX 2E). Prepare immediately before use. Ellman’s reagent solution Dissolve 4 mg Ellman’s reagent, 5,5′-dithio-bis-(2-nitrobenzoic acid) (Pierce), in 1 ml of 0.1 M sodium phosphate, pH 8.0 (APPENDIX 2E). Prepare immediately before use. Glutaraldehyde solution, 0.15% Add 30 µl of 25% aqueous glutaraldehyde solution to 5 ml of 50 mM sodium borate buffer, pH 8.0 (pH adjusted with HCl). Prepare fresh and use immediately. If the glutaraldehyde precipitates, check the pH. It should not be above 8.0; a slightly lower pH can be used (pH 7 to 8). CAUTION: Glutaraldehyde is a sensitizing agent that should be handled in a hood and only according to the recommendations in the Material Safety Data Sheet. When mixing solutions or performing reactions, keep the container covered to prevent vapors from escaping into the atmosphere.
Guanidine⋅HCl, 6 M Dissolve 1 g guanidine⋅HCl in 1 ml of 0.05 M sodium phosphate, pH 7.0 (APPENDIX 2E). Store up to several weeks at room temperature. The resulting 1.8-ml solution should be ∼0.025 M phosphate/6 M guanidine⋅HCl at pH 7.0.
COMMENTARY Background Information
Selecting Synthetic Peptides for Production of Antibodies
Synthetic peptides are linear arrays of amino acids that in most instances possess a random structure in solution. While it is not difficult to produce antipeptide antibodies, it does not necessarily follow that the antibodies will recognize a protein containing the same stretch of sequence found in the peptide. In order for this to occur, the amino acids in the protein must be oriented to the antibody in a way similar to that of the synthetic peptide. This generally requires three basic features of the protein: (1) that the stretch of sequence be exposed to solvent; (2) that the sequence be a continuous stretch of amino acids; and (3) that it not possess a higherorder structure that renders it unrecognizable by the antibody population. The large number of model protein structures now available indicate that almost all of the ionized groups in water-soluble proteins are on the protein surface. Asp, Glu, Lys, and Arg residues, on the average, comprise 27% of the protein surface and only ∼4% of the protein interior. The fraction of residues that are at least 95% buried range from 0.36 to 0.60 for nonpolar residues and 0.01 to 0.23 for polar residues. Only 1% of Arg and 3% of Lys residues fall
into the 95% buried range (Creighton, 1993). Therefore, it is reasonable to expect solvent-exposed areas of proteins to display relatively high levels of polar and charged residues, particularly Arg and Lys. Proteins display three kinds of secondary structure: α-helices, β-sheets, and turns or loops (see UNIT 17.1). Turns or loops generally connect elements of α-helices and β-sheets, and can either fit one of several rather strict motifs with recognizable hydrogen bonding patterns or be of a more extended, random nature. These turn or loop structures appear to be most useful for antibody production because they tend to be found on the surface of proteins connecting larger arrays of helices and sheets, and they consist of continuous stretches of amino acids. Although many amino acid residues in helices and sheets are also exposed at the surface, the regular geometry of amino acids contained within them makes them less suitable for this purpose. For instance, in β-sheet structures the side chain of each successive amino acid in the β-sheet strand points in the opposite direction to the ones immediately preceding and following it. Thus, even if the amino acid side chains are not predominantly buried in the interior of
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the protein, only every other side chain is exposed on the same surface of the sheet. This can hinder recognition by an antibody produced with a linear peptide capable of assuming a more random structure. A similar situation exists for α-helices. Although the change in direction of the side chains of successive amino acids is perhaps not as abrupt as in β-sheets, only approximately every third or fourth side chain is found on the same surface of the helix. Epitopes in proteins have been identified in amphipathic helices, but unless the synthetic peptide assumes a similar helical structure in solution, recognition by the antibody may be problematic. These considerations have led to more useful methods for predicting sequences that will produce antibodies recognizing intact proteins. A variety of different indices that predict hydrophilicity or hydrophobicity and secondary structure are available (see UNITS 2.2 & 2.3, respectively). In addition, predictive methods based on segmental mobility, side chain accessibility, and sequence variability (see Van Regenmortel et al., 1988) have also been proposed. All of these methods generally tend to yield similar results, but it must be noted that these procedures were developed for (and work best with) water-soluble proteins composed of a single globular structure. Additional complications can arise with multisubunit proteins, where normally exposed structures may be shielded by subunit interactions, or membrane proteins with large sections shielded from the solvent. The method presented in this unit utilizes the correlation between the hydrophilic character of a peptide sequence (Kyte and Doolittle, 1982) and its propensity to form β-turn structures (Chou and Fasman, 1974). Free access to these and many other algorithms is provided at the ExPASy Web site of the University of Geneva at http://expasy.org.tools. After selection of the peptide sequence, an effective immunogen is generally produced by coupling the peptide to a carrier protein or by synthesizing a multiple antigenic peptide (MAP), with four or eight identical peptides assembled simultaneously on the α and ε amines of the terminal lysines of a branched core (see Fig. 18.3.3).
Critical Parameters Analyzing protein sequences with algorithms or tables of assigned values for amino acids is a well-established procedure, but evaluating these results and selecting the candidate sequences requires some consideration. To take
full advantage of the results, choose areas of sequence that give the maximum values for the properties being evaluated and that also show the highest degree of residue-by-residue correlation. In other words, choose areas of maximum amplitude where the centers of the peaks correspond to the same sequence with a divergence of no more than two to three residues. Examples of this are given in Figure 18.3.2, which shows results from the method presented in Basic Protocol 1 for the sequence shown in Figure 18.3.1. The top panel in Figure 18.3.2 predicts β-turns as calculated by the method of Chou and Fasman (1974). The bottom panel is a prediction of hydrophobicity using the parameters of Kyte and Doolittle (1982). The data are analyzed by looking for areas of high turn propensity (maximum positive deflection in the top panel) and high hydrophilicity (maximum negative deflection in the bottom panel). The shaded areas in Figure 18.3.2 designate three segments that meet these criteria. Note that the maximum and minimum values of these three stretches of protein sequence correlate very well. Additional areas of high hydrophilicity (bottom panel) are found near residues 64, 132, 137, 149, and 345, although the β-turn values of these secondary candidates are not as high as those of the three shaded areas. Two equally hydrophilic areas at residues 49 and 299 correspond to downward deflections in the β-turn profile and are thus not good candidates based on this analysis. Many different chemistries are available for coupling synthetic peptides to carrier proteins to produce effective immunogens (Van Regenmortel et al., 1988). In many cases, however, side reactions or incompatibilities in chemistry between the coupling agent and the residues present in the peptide can be problematic. In order to simplify the process and present the greatest probability of success in most cases, only a few coupling methods are presented in this unit. In this regard, the recommended coupling procedure is cross-linking of the peptide via cysteine residues to keyhole limpet hemocyanin (KLH) with the heterobifunctional reagent m-maleimidobenzoyl-N-hydroxysuccinimide ester (MBS; see Basic Protocol 3). This effective method has enjoyed great success and can be used for virtually any peptide. The one caveat is that it is not recommended for peptides with internal cysteine residues, since they will also link to the carrier. It is also critical, when coupling with MBS through an added terminal cysteine residue, that the sulfhydryl group of
Preparation and Handling of Peptides
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Selecting Synthetic Peptides for Production of Antibodies
the peptide be present in the free or reduced form (see Support Protocols 1 and 2). In addition to MBS coupling, other procedures commonly used (see Alternate Protocols 3 and 4) are included as alternatives for use in special situations, but these are not recommended as a general alternative to MBS because they are more restrictive and have the potential for undesirable side reactions. Glutaraldehyde coupling (see Alternate Protocol 3) should not be used with peptides containing internal Lys, Cys, Tyr, or His residues and, since it is a homobifunctional reagent, cross-linking of the peptide to itself and the carrier to itself can occur. The latter lowers antigenicity and can result in extensive aggregation and precipitation of the carrier. 1-ethyl-3(3-dimethylaminopropyl) carbodiimide (EDC; see Alternate Protocol 4) is a water-soluble carbodiimide and should not be used with peptides containing internal Lys, Glu, Asp, Tyr, or Cys residues. Alternate Protocol 5 describes a simple photochemical coupling strategy (Gorka et al., 1989). Another good alternative for most peptides is the production of a multiple antigenic peptide (MAP; see Alternate Protocol 2). With this method the composition of the peptide is not a concern beyond its potential solubility properties. In most cases, since hydrophilic sequences are selected, this also is not a major problem. Both four- and eight-branched MAPs have been found to be effective. However, four-branched MAPs are recommended because they are less prone to synthesis problems and are easier to characterize. As with any synthetic peptide, the product must be well characterized before use. If the peptide is not what it was intended to be, this decreases the probability of generating antibodies that will recognize the protein. At the very least, check synthetic peptides for homogeneity by analytical HPLC and correct mass by mass spectrometry (see Chapter 16). Characterization of MAP can be more problematic due to their multibranched nature (Mints et al., 1997): HPLC and mass spectrometric analysis can be compromised by the presence of four to eight peptide chains per molecule, each of which may have only a small percentage of modification at any particular residue but which in the aggregate contribute to broad spectra. However, this feature of MAPs usually does not tend to compromise their ability to form antigens of the proper peptide since the correct sequence is usually present in high enough concentration that a significant amount of specific antibody is produced among the polyclonal population. Amino acid analysis (UNIT 11.9), which is less sensitive to
multiple small differences, tends to give a reasonable assessment of the MAP integrity.
Anticipated Results The methods outlined in this unit produce an effective polyclonal antiserum against an intact protein from a single peptide sequence ∼50% to 70% of the time. Therefore, it is advisable to prepare two or three different peptides from a given protein to increase the probability of at least one of them being effective.
Time Considerations Computer-assisted analysis of a protein sequence and inspection of the data to select several candidate sequences takes from 5 to 30 min. Manual analysis of a protein sequence can take several hours but can certainly be accomplished in <1 day. Selection of peptide design and manner of synthesis as well as selection of a coupling method will take <1 hr. Actual preparation of the peptide can be accomplished in 3 to 4 days, but this may vary depending on the turnaround time of the synthetic laboratory. Coupling a synthetic peptide to a carrier protein takes from 1 to 2 days. Although not covered in this unit, production of the antisera will vary with the animal and protocol used, but generally requires 2 to 3 months. It is therefore advisable to inject several animals with different peptides at one time.
Literature Cited Chou, P.Y. and Fasman, G.D. 1974. Prediction of protein conformation. Biochemistry 13:222-245. Creighton, T.E. 1993. Proteins: Structure and Molecular Properties, 2nd ed. W.H. Freeman, New York. Gailit, J. 1993. Restoring free sulfhydryl groups in synthetic peptides. Anal. Biochem. 214:334-335. Getz, E.B., Xiao, M. Chakrabarty, T., Cooke, R., and Selvin, P.R. 1999. A comparison between the sulfhydryl reductants Tris(2-carboxyethyl) phosphine and dithiothreitol for use in protein biochemistry. Anal. Biochem. 272:73-80. Gorka, J., McCourt, D.W., and Schwartz, B.D. 1989. Automated synthesis of a C-terminal photoprobe using combined Fmoc and t-Boc synthesis strategies on a single automated peptide synthesizer. Peptide Res. 2:376-380. Kyte, J. and Doolittle, R.F. 1982. A simple method for displaying the hydropathic character of a protein. J. Mol. Biol. 157:105-132. Mints, L., Hogue Angeletti, R., and Nieves, E. 1997. Analysis of MAPS peptides. ABRF News 8:22-26. Posnett, D.N., McGrath, H., and Tam, J.P. 1988. A novel method for producing anti-peptide antibodies. Production of site-specific antibodies to the T-cell antigen beta-chain. J. Biol. Chem. 263:1719-1725.
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Tam, J.P. 1988. Synthetic peptide vaccine design: Synthesis and properties of a high density multiple antigenic peptide system. Proc. Natl. Acad. Sci. U.S.A. 85:5409-5413. Van Regenmortel, M.H.V., Briand, J.P., Muller, S., and Plaué, S. 1988. Synthetic polypeptides as antigens. In Laboratory Techniques in Biochemistry and Molecular Biology, Vol. 19 (R.H. Burdon and P.H. van Knippenberg, eds.). Elsevier/North-Holland, Amsterdam.
Key References Van Regenmortel et al., 1988. See above. Comprehensive treatment of theory and method. Tam, J.P. 1988. High density multiple antigen peptide system for preparation of anti-peptide antibodies. Methods Enzymol. 168:7-15. Original methods article for MAPs.
Internet Resource http://expasy.org/tools.html Web site for programs to analyze protein sequences.
Contributed by Gregory A. Grant Washington University School of Medicine St. Louis, Missouri
Preparation and Handling of Peptides
18.3.19 Current Protocols in Protein Science
Supplement 28
Native Chemical Ligation of Polypeptides
UNIT 18.4
The total synthesis and semisynthesis of proteins allows the site-specific incorporation of unnatural amino acids, post-translational modifications, and biophysical/biochemical probes into the target molecule. Among the various chemical and enzymatic approaches available for the synthesis/semisynthesis of proteins, the native chemical ligation technique has proven especially useful and will be the exclusive focus of this unit. Native chemical ligation allows native backbone proteins to be assembled from fully unprotected polypeptide building blocks. To facilitate the ligation reaction, the α-carboxylate group of the N-terminal polypeptide fragment must be mildly activated as an aryl thioester, while the C-terminal polypeptide fragment must contain an amino-terminal cysteine residue. The reaction is normally carried out in aqueous buffers at around neutral pH and can be performed with complete regioselectivity in the presence of all the functionalities commonly found in proteins. Even the presence of additional cysteine sulfhydryl groups in one or both peptide fragments does not affect the reaction regioselectivity. The unit first discusses how to choose the ligation site(s) in the target protein and then outlines how to obtain the necessary polypeptide building blocks using either chemical synthesis or recombinant DNA expression (see Strategic Planning). Next, the synthesis of a protein by native chemical ligation of two polypeptide fragments is described (see Basic Protocol 1). The synthesis of a protein from three polypeptide fragments using a sequential native chemical ligation strategy is also described (see Basic Protocol 2). The support protocols describe how to obtain the necessary polypeptide fragments using either chemical synthesis or recombinant DNA expression. NOTE: All the operations involving volatile chemicals should be performed in a well ventilated fume hood. See APPENDIX 2A for general safety guidelines. STRATEGIC PLANNING The first step in synthesizing a protein by native chemical ligation is to choose the ligation site(s), thereby defining the polypeptide fragments to be used in the ligation reactions. The position of the ligation site depends upon several factors: the primary sequence of the protein; the secondary and tertiary structures of the protein (if known); and the type of strategy being used to generate the fragments, i.e., chemical synthesis or recombinant DNA expression. Ideally the target protein should be divided up into synthetically accessible fragments the size of which depends upon whether chemical synthesis or biosynthesis is being used for their generation; optimized solid-phase peptide synthesis (SPPS; also see UNIT 18.1) allows polypeptides of up to ∼50 residues to be efficiently prepared, whereas recombinant DNA expression permits the preparation of much larger fragments. For example, a hypothetical 100-amino-acid protein could be assembled either from two ∼50-residue synthetic peptides or from a 20-residue synthetic fragment and an 80-residue recombinant fragment. Note that in the latter strategy (i.e., semisynthesis) it is important to design the synthesis in such a way that the chemically synthesized fragment encompasses the region of the protein to be engineered. Where possible, naturally occurring X-Cys motifs in the native sequence should be chosen as the ligation sites, where X can be any residue except Asp, Asn, Glu, Gln, and Pro. In the absence of an endogenous Cys at the appropriate position in the primary sequence, it will be necessary to mutate a native residue in order to facilitate ligation. The effect of this Cys mutation on the structure and function of the protein can usually be minimized by following a few simple rules: (1) If the tertiary structure of the protein is known, choose a region remote from the active site of the protein, preferably in a flexible surface loop; (2) choose the mutation to be as conservative as possible, e.g., Ser→Cys or Ala→Cys; (3) try to avoid Contributed by Julio A. Camarero and Tom W. Muir Current Protocols in Protein Science (1999) 18.4.1-18.4.21 Copyright © 1999 by John Wiley & Sons, Inc.
Preparation and Handling of Peptides
18.4.1 Supplement 15
Table 18.4.1 Resins and Amino Acid Derivatives Usually Employed in the Synthesis of N-Terminal Cys and α-Thioester Polypeptidesa
Polypeptide N-terminal Cys
Nα-protecting group strategy Boc
Fmoc
α-thioester
Boc
Side-chain protecting groups Arg(Tos), Asn(Xan), Asp/Glu(OcHx), Cys(Meb), Gln(no protection), His(Dnp), Lys(2-Cl-Z), Met(no protection), Ser/Thr(Bzl), Trp(For), Tyr(2-Br-Z) Asn(Trt), Arg(Pmc), Asp/Glu(OtBu), Cys(Trt), Gln(Trt), His(Trt), Lys(Boc), Met(no protection), Ser/Thr/Tyr(tBu), Trp(Boc) As in the Boc strategy for obtaining Nterminal Cys polypeptides, except for His(Bom) and Trp(no protection or Hoc)
Resin(s)
C-terminal functionality after cleavage
MBHA PAM
—CONH2 —COOH
Rink amide Wang
—CONH2 —COOH
3-mercaptopropionamide -MBHA
—COSCH2CH2CONH2
aAbbreviations:
Boc, t-butyloxycarbonyl; Bom, benzyloxymethyl; 2-Br-Z, 2-bromobenzyloxycarbonyl; tBu, t-butyl; 2-Cl-Z, 2-chlorobenzyloxycarbonyl; Dnp, 2-4-dinitrophenyl; Fmoc, 9-fluorenyloxycarbonyl; For, formyl; Meb, p-methylbenzyl, OtBu, t-butyl ester; OcHx, cyclohexyl ester; Pmc, 2,2,5,7,8-pentamethylchroman-6-sulfonyl; Tos, tosyl (p-toluenesulfonyl); Trt, trityl; Hoc, cyclohexyloxycarbonyl; PAM, 4-hydroxymethylphenylacetamidomethyl; Xan, xanthyl.
mutating residues that are conserved across a gene family; and (4) take advantage of any known mutational data on the system since the effect on structure/function of mutating a particular residue may already be known. Chemical synthesis and recombinant DNA expression each allow the generation of polypeptides containing an amino-terminal cysteine residue (C-terminal fragment) and the generation of polypeptides containing an α-thioester functionality (N-terminal fragment). For polypeptides of ∼50 residues or less, chemical synthesis is usually the method of choice since it is relatively quick and efficient and allows the incorporation of unnatural amino acids. Amino-terminal cysteine–containing peptides can be chemically synthesized via established t-butyloxycarbonyl (Boc) or 9-fluorenyloxycarbonyl (Fmoc) SPPS protocols using commercially available resins (e.g., from NovaBiochem, Bachem, Peptides International, Applied Biosystems, or Neosystems) and amino acid derivatives (see Table 18.4.1). The synthesis of peptide α-thioesters can currently only be achieved using Boc chemistry, due to the sensitivity of the thioester functionality to the base-deprotection conditions associated with the Fmoc-SPPS strategy. Note, the resin required for the synthesis of peptide α-thioesters is not commercial available, but can be prepared in the laboratory in a single day.
Native Chemical Ligation of Polypeptides
For polypeptide fragments larger than ∼50 amino acids in length, a biosynthetic strategy should be employed. It must be noted that this assumes the gene has been cloned. Polypeptides containing amino-terminal Cys residues can be obtained using a mutagenesis/proteolysis strategy. This involves constructing an expression vector in which a DNA sequence encoding the peptide Ile-Glu-Gly-Arg-Cys is inserted between a upstream affinity-purification handle (e.g., GST or His tag) and the appropriate gene fragment of interest (for cloning strategies see UNIT 6.6). Following expression and affinity purification (UNIT 6.6), the fusion protein is treated with Factor Xa to give the desired recombinant Cys-polypeptide. Polypeptides containing α-thioester functionalities can be obtained by expressing the corresponding polypeptide sequence using the IMPACT expression system (New England Biolabs). This results in the generation of a fusion protein in which the polypeptide of interest is attached through its C-terminus to an intein-CBD, where the intein is an engineered protein-splicing element and CBD (chitin binding domain) is an
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+ H2N
+ NH3 + H3N
+ H2N
NH2 NH
+ NH3
HS
OH O
Polypeptide 2
CO2–
SR
CO2–
CO2– + H2N
+ NH3
H2N O S
spontaneous S to N acyl transfer + H2N NH2 + NH3 OH NH O + Polypeptide 2 H3N NH – SH CO
NH2 OH
NH
+ H2N
CONH
+ NH3
CO2–
SH
CO2–
SH
OH
Polypeptide 1
CONH
SH
2
NH
Polypeptide 1
H2N CONH
SH
reversible transthioesterification + H2N NH2 + OH NH3 NH + Polypeptide 2 H3N
NH2
CO2–
SH
NH2 OH
NH
Polypeptide 1
CO2–
CO2–
SH
Figure 18.4.1 The mechanism of native chemical ligation. The initial step is a reversible transthioesterification reaction involving the thiol group of the N-terminal Cys-polypeptide (C-terminal fragment) and the α-thioester moiety of the N-terminal polypeptide fragment. This intermediate undergoes a spontaneous rearrangement to form a natural peptide bond at the ligation site.
affinity handle. The desired recombinant α-thioester polypeptide can be obtained by simply treating the immobilized fusion protein with an appropriate thiol; the intein-CBD construct remains attached to the affinity matrix. NATIVE CHEMICAL LIGATION OF TWO POLYPEPTIDES This protocol describes the chemical ligation of two polypeptide segments to afford a target protein containing both fragments linked through a native peptide bond. The reaction is initiated by dissolving both peptides in aqueous buffer at pH 7.5 in the presence of thiol cofactors. The reaction usually takes between 1 and 3 days to go to completion, at which point the product is purified using reversed-phase high-performance liquid chromatography (RP-HPLC) and subsequently folded. A schematic of the chemistry behind the procedure is presented in Figure 18.4.1. Materials 6 M guanidine⋅HCl buffer (see recipe) Nitrogen source (ultrapure) Benzyl mercaptan (Aldrich) Thiophenol (Aldrich) Polypeptide with N-terminal Cys residue (see Support Protocols 1 and 2) Polypeptide with α-thioester functionality (see Support Protocols 3 and 4) 1 M NaOH Tris(2-carboxyethyl)phosphine hydrochloride (TCEP, 99% pure; Strem Chemicals) continued
BASIC PROTOCOL 1
Preparation and Handling of Peptides
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Buffers for analytical C18 reversed-phase HPLC Buffer A: H2O containing 0.1% trifluoroacetic acid (TFA) Buffer B: 1 part H2O/9 parts acetonitrile/0.1% TFA Appropriate final buffer in which target protein will be dissolved (e.g., for long-term storage, activity studies, or structural studies), containing 6 M guanidine⋅HCl and 5 mM DTT Appropriate final buffer in which target protein will be dissolved, containing 4 M, 2 M, and 0 M guanidine⋅HCl (APPENDIX 3A) and 1 mM DTT Dialysis membrane with MWCO substantially smaller than target protein Centricon filter (Amicon; optional) Tabletop centrifuge (Sorvall RT-7 or equivalent) Additional reagents and equipment for analytical and semipreparative or preparative C18 reversed-phase HPLC, electrospray ionization (ESI) or matrix-assisted laser desorption ionization time-of-flight (MALDI-TOF) mass spectrometry (MS; UNITS 16.1 & 16.2), dialysis (UNIT 4.4 & APPENDIX 3B), and spectrophotometric determination of protein concentration (UNIT 3.1) Prepare the ligation buffer 1. Degas the 6 M guanidine⋅HCl buffer by purging with ultrapure nitrogen for at least 20 min. 2. Add benzyl mercaptan and thiophenol, each at a concentration of 0.5% (v/v), to the degassed buffer. Mix by vortexing to produce the ligation buffer. IMPORTANT NOTE: This ligation buffer should be prepared just prior to use and should not be stored for long periods of time. Thiols are, in general, toxic and have an unpleasant odor; therefore thiol-containing solutions should be treated with 5% (v/v) commercial bleach in 1:1 (v/v) methanol/water before being disposed of. Since benzyl mercaptan and thiophenol are barely soluble in aqueous buffers, vortexing is required to form a homogeneous thiol suspension in the ligation buffer.
Ligate the N-terminal and C-terminal polypeptide fragments 3. Dissolve equimolar amounts of the polypeptides to be ligated (i.e., one with an N-terminal Cys residue and the other with an α-thioester functionality) in the ligation buffer to a final concentration of at least 0.5 mM with respect to each peptide. Reactions can be run in volumes of 50 to 100 µl for analytical purposes and up to 0.5 to 5 ml for preparative purposes. Native chemical ligation reactions are usually carried out in the presence of chaotropic agents (guanidium⋅HCl or urea) in order to maximize the concentration of the two reactants and to increase the availability of reacting moieties by alleviating any steric hindrance that might be associated with folded polypeptides. The use of chemical denaturants, of course, presumes that the final product can be properly folded in vitro. In cases where the fragments being ligated must be kept in a folded state, the ligation reaction can be carried out in the absence of chemical denaturants.
4. Check the pH of the ligation mixture by spotting a 2- to 3-µl aliquot onto a strip of pH paper. If it is above or below 7.0, adjust the pH by titrating in small increments with 1 M HCl or NaOH, respectively. Repeat this process until a pH of ∼7.5 is reached.
Native Chemical Ligation of Polypeptides
Dissolving lyophilized peptides at high concentrations can cause the pH of the ligation mixture to drop; this is particularly so when HPLC-purified materials are being used (i.e., from acidic conditions). It is extremely important to ensure that the pH of the reaction mixture is kept between 7 and 8, since the efficiency of the native chemical ligation is strongly dependent on the pH. As the pH drops below pH 7.0, the reaction slows down, and as it increases above pH 8.0 the chemoselectivity of the process is lost.
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5. Keep the reaction at room temperature with gentle stirring for 24 hr. 6. After 24 hr, take a 5-µl aliquot of the reaction mixture (allowing the rest of the mixture to continue reacting as in step 5) and dilute the aliquot in a polypropylene microcentrifuge tube with 50 µl of 6 M guanidine⋅HCl buffer in the presence of one crystal of tris-(2-carboxyethyl)phosphine hydrochloride (TCEP, used as a reducing agent). Shake the sample until the TCEP is completely dissolved, wait 10 min, then microcentrifuge 5 min at 10,000 rpm. 7. Analyze 25 µl of the supernatant by analytical C18 reversed-phase HPLC, initially using a linear gradient from 0% buffer B to 73% buffer B over 60 min at a flow rate of 1 ml/min and refining as necessary based on the elution times of the reactants and product. Monitor the run via UV detection at 214 nm (and 280 nm if aromatic residues are present in the target protein). Manually collect every peak (except the solvent peak) into a separate polypropylene microcentrifuge tube and analyze each by ESI-MS (UNITS 16.1 & 16.2), which will allow the retention times of the reactants and the product to be determined Occasionally the peak corresponding to the ligation product is buried beneath the thiophenol or benzyl mercaptan peaks. This can usually be resolved by optimizing the HPLC gradient or, alternatively, by lyophilizing the reduced sample a few times and then reanalyzing it by HPLC.
8. After 36 hr take another 5-µl aliquot of the reaction mixture and analyze it by HPLC as in step 7. If significant amounts of both reactants still remain and the product peak area is still increasing, then leave the reaction for another 12 hr and repeat the analysis. If, on the contrary, both reactants have been consumed or if the product peak area has not increased appreciably, then terminate the reaction as in step 9. Ligation reactions normally take 24 to 36 hr to go to completion, although on rare occasions longer periods (3 to 4 days) or shorter periods (a few hours) may be required.
9. Dilute the crude reaction mixture into 5 to 10 vol of 6 M guanidine⋅HCl buffer and reduce by adding 15 molar equivalents of TCEP (i.e., based on the quantities of reactants used; 15 times the number of moles of either one of the polypeptides added). Shake the solution until all the TCEP is dissolved and wait for 20 min. 10. Centrifuge the sample 20 min at 3000 rpm in a tabletop centrifuge (Sorvall RT-7 or equivalent), room temperature. Retain the supernatant. A white cloudiness often appears in the reaction medium during the course of the reaction. This is due to the slow formation of insoluble benzyl mercaptan and thiophenol disulfides. The reactants and ligation product will generally remain in solution, allowing the unwanted disulfides to be removed by centrifugation. The crude ligation mixture can be stored at −80°C for 2 to 3 days, although it is advisable to purify it as soon as possible.
Purify the target protein 11. Purify the crude reaction mixture by semipreparative or preparative reversed-phase HPLC using an optimized gradient system. Analyze each HPLC fraction by mass spectrometry (UNITS 16.1 & 16.2) and analytical reversed-phase HPLC in order to identify those fractions containing the target protein. Pool the fractions of highest purity (>95%) and lyophilize. Reversed-phase HPLC purification will denature the majority of proteins, meaning that a subsequent refolding step is required. In cases where the ligation reaction has been carried out under more physiological conditions (i.e., without chemical denaturants in the ligation buffer), the final purification step can be performed using, e.g., gel-filtration chromatography (UNIT 8.3) or ion-exchange chromatography (UNIT 8.4) under conditions compatible with protein folding.
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Fold the purified protein 12. Dissolve the lyophilized target protein to a final concentration of ≤100 µM in the appropriate freshly degassed buffer (e.g., for long-term storage, activity studies, or structural studies), containing 6 M guanidine⋅HCl and 5 mM DTT. Although the present protocol involves the stepwise dialysis from 6 M guanidium⋅HCl, it should be stressed that this approach may not work for all proteins and that a certain amount of optimization will be required for every system. It should also be noted that for disulfide-containing proteins, an additional oxidation step will have to be performed. In general, the lower the final concentration of protein the better, since this will tend to minimize aggregation during the refolding step.
13. Dialyze the solution at 4°C (UNIT 4.4 & APPENDIX 3B) against the same buffer as in step 12, this time containing 1 mM DTT and decreasing amounts of guanidine⋅HCl (i.e., first against 4 M guanidine⋅HCl, then against 2 M guanidine⋅HCl, then against final storage buffer). In each step perform dialysis for at least 12 hr and change the buffer 2 to 3 times during this period. The MWCO of the dialysis membrane should be chosen to be substantially smaller than the purified protein.
14. Remove sample from dialysis bag, centrifuge to remove any precipitate, and quantify the supernatant by UV spectrophotometry at 280 nm (UNIT 3.1). If necessary concentrate the protein solution using a Centricon filter (UNIT 4.4; Amicon), then divide into aliquots and freeze at −70°C. BASIC PROTOCOL 2
SEQUENTIAL CHEMICAL LIGATION OF THREE POLYPEPTIDES If the region of interest in the protein is >50 residues from the N- or C-termini, then its chemical modification will be extremely difficult using a two-fragment ligation strategy, since this would require the chemical synthesis of a peptide >50 residues in length. This problem can be overcome by assembling the target protein from three polypeptide fragments using a sequential native ligation strategy. The basic method for sequential native ligation uses a N-terminal Cys polypeptide (C-terminal fragment), an α-thioester polypeptide (N-terminal fragment), and an Nα(methylsulfony)ethyloxycarbonyl-Cys, α-thioester polypeptide (central fragment). The procedure starts by ligating the middle fragment and the C-terminal fragment at pH 7.5 in the presence of thiol cofactors. The Nα(methylsulfony)ethyloxycarbonyl (Msc) group is then removed and the resulting polypeptide (central plus C-terminal fragment) is ligated with the N-terminal fragment to give the target protein, which is purified and refolded. The general approach is depicted in Figure 18.4.2. Materials Three polypeptide fragments to be ligated: Polypeptide with N-terminal Cys residue (see Support Protocols 1 and 2) Polypeptide with Nα(Msc)-Cys, α-thioester functionality (see Support Protocol 5) Polypeptide with α-thioester functionality (see Support Protocols 3 and 4) 1 M HCl Additional reagents and equipment for native chemical ligation of two polypeptides (see Basic Protocol 1)
Native Chemical Ligation of Polypeptides
1. Prepare the ligation buffer (see Basic Protocol 1, steps 1 and 2). Ligate the central [i.e., Nα(Msc)-Cys, α-thioester] and C-terminal (N-terminal Cys) polypeptide fragments [see Basic Protocol 1, steps 3 to 8, but use the Nα(Msc)-Cys, α-thioester polypeptide in the reaction mix in place of the α-thioester polypeptide].
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2. When the ligation reaction is complete, remove the Nα-Msc protecting group by raising the pH of the crude ligation mixture to 13 with 1 M NaOH solution (see Basic Protocol 1, step 4). After 1 min, lower the pH to 5.0 to 7.0 with 1 M HCl (use ∼1.1 to 1.2 times the volume of 1 M NaOH required to raise the pH to 13). Usually, for a 1-ml ligation mixture, the pH can be raised by adding 75 µl of 1 M NaOH, then dropped by adding 95 µl of 1 M HCl. It is recommended that the pH be dropped by adding the acid all at once, instead of by titration, for reasons of speed. The presence of thiols improves the yield in the deprotection step, since they trap the still reactive methyl ethylenyl sulfone side product.
3. Analyze the crude reaction mixture (see Basic Protocol 1, steps 6 and 7) and examine to see if the Msc deprotection is complete. Purify the polypeptide fragment (see Basic Protocol 1, steps 9 to 11), which will consist of the central fragment ligated to the C-terminal fragment. 4. Starting with the lyophilized (central plus C-terminal) polypeptide fragment prepared in steps 1 to 3 above and the N-terminal polypeptide fragment (containing an α-thioester functionality), perform the ligation reaction to generate the target protein (see Basic Protocol 1, steps 3 to 10). 5. Purify and refold the target protein (see Basic Protocol 1, steps 11 to 15).
HS
HS
Msc-HN-Cys
peptide 2
CO-SR
+ H2N-Cys
– peptide 1 CO2
aqueous buffer, pH 7.5 thiophenol 0.5% benzyl mercaptan
ligation 1
HS
HS
Msc-HN-Cys peptide 2
CO-NH- Cys
removal of Msc group
HS
peptide 1
CO2–
pH 13 HS
H2N-HN-Cys peptide 2
CO-NH- Cys
peptide 1
CO2–
aqueous buffer, pH 7.5 thiophenol 0.5% benzyl mercaptan
ligation 2
+ H3N peptide peptide 33 HS + H3N-Cys
peptide 3
CO-NH-Cys peptide 22
CO-SR
HS CO-NH-Cys
peptide 1
CO2–
Figure 18.4.2 The principle of sequential native chemical ligation. The key to this approach is the reversible protection of the α-amino group of the central peptide fragment, thereby preventing self-reaction with the α-thioester moiety present in the same molecule. This can be accomplished with the base-labile Msc [Nα(methylsulfony)ethyloxycarbonyl] group, which can be easily removed by brief treatment with base after the first ligation step. The newly deprotected fragment is now ready for the next ligation step.
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SUPPORT PROTOCOL 1
CHEMICAL SYNTHESIS OF N-TERMINAL CYS-POLYPEPTIDES The chemical synthesis of N-terminal Cys-polypeptides can be easily carried out using standard SPPS with Boc or Fmoc Nα-protected amino acids and the appropriate resins (see Table 18.4.1 and UNIT 18.1). In both cases, the synthesis can be performed on automated solid-phase peptide synthesizers, which are available in the core facilities of many institutions. NOTE: Extreme care must be taken to avoid exposing N-terminal Cys-polypeptides to even trace amounts of carbonyl-containing compounds (e.g., acetone or formaldehyde). These chemicals react rapidly with the α-amino and thiol groups of the N-terminal Cys to give a very stable thiazolidine derivative which prevents the peptide from participating in subsequent native chemical ligation reactions. For the same reason, the groups benzyloxymethyl (Bom) and t-butyloxymethyl (Bum), used to protect the side-chain of His in Boc and Fmoc SPPS, respectively, should be avoided during the synthesis of these polypeptides since they release formaldehyde during the cleavage/deprotection step.
SUPPORT PROTOCOL 2
SUPPORT PROTOCOL 3
Native Chemical Ligation of Polypeptides
BIOSYNTHESIS OF N-TERMINAL CYS POLYPEPTIDES Biosynthesis represents a complementary approach to chemical synthesis (see Support Protocol 1) for obtaining N-terminal-Cys polypeptides, and is especially useful when the target polypeptide fragment is somewhat greater than 50 residues in length (i.e., too large to be chemically synthesized). Assuming that the cDNA for the gene is available, N-terminal Cys polypeptides can be obtained using a mutagenesis/proteolysis strategy (Erlandson et al., 1996). This involves constructing an expression vector in which a DNA sequence encoding the peptide Ile-Glu-Gly-Arg-Cys is inserted between a upstream affinity-purification handle (e.g., MBP, GST, or His tag) and the appropriate gene fragment of interest (for cloning and mutagenesis see UNIT 6.6). Following expression and affinity purification, the fusion protein is treated with Factor Xa to give the desired recombinant Cys-polypeptide, which can then be further purified if necessary. These recombinant Cys-polypeptides can be used in native chemical ligation reactions as per their synthetic counterparts. CHEMICAL SYNTHESIS OF α-THIOESTER POLYPEPTIDES This protocol describes the chemical synthesis of a 3-mercaptopropionamide α-thioester polypeptide on a 3-mercaptopropionamide-MBHA resin. This resin can be easily prepared from commercially available MBHA resin using a three-step solid-phase procedure (Fig. 18.4.3). The solid-phase synthesis of the polypeptide is achieved using Boc amino acid derivatives employing in situ neutralization/HBTU [2-(1H-benzotriazolyl)-1,1,3,3tetramethyluronium hexafluorophosphate] activation protocols for Boc-SPPS (Schnölzer et al., 1992). The corresponding α-thioester polypeptide, suitable for native chemical ligation, is obtained after cleavage with hydrogen fluoride and purification. Materials Methylbenzhydrylamine (MBHA) resin (Peptides International) Dimethylformamide (DMF, spectrophotometric grade; Fisher) 5% (v/v) diisopropylethylamine (DIEA, peptide synthesis grade; Perkin-Elmer Applied Biosystems) in DMF (store up to 1 month at room temperature) 97% 3-bromopropionic acid Dichloromethane (DCM, spectrophotometric grade; Fisher) 99% diisopropyl carbodiimide (DIPC, 99%; Aldrich) Diisopropylethylamine (DIEA, peptide synthesis grade; Perkin-Elmer Applied Biosystems)
continued
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Br MBHA
NH2
CO2H, DIPC
O MBHA
NH
DCM
O
CH3-COSH, DIEA MBHA Br DMF
NH
NS
O MBHA
NH
S
CH3
OH, DIEA, DMF
O
Boc-AA-OSu MBHA
S-AA-Boc
O
NH
SH
DMF
Figure 18.4.3 Scheme showing the preparation of the Boc-aminoacyl-3-mercaptopropionamide-MBHA resin using solidphase transformations. Abbreviations: AA, aminoacyl; Boc, t-butyloxycarbonyl; CH3-COSH, thioacetic acid; DCM, dichloromethane; DIEA, diisopropylethylamine; DIPC, diisopropyl carbodiimide; DMF, dimethylformamide; MBHA, methylbenzhydrylamine; OSu, hydroxysuccinimide ester.
Ac2O/DIEA/DMF solution (see recipe) AcSH/DIEA/DMF solution (see recipe) BME/DIEA/DMF solution (see recipe) Boc–amino acyl–N-hydroxysuccinimide ester (Boc-AA-OSu; where AA is the first amino acid to be incorporated into the polypeptide; Bachem) Trifluoroacetic acid (TFA, BioGrade) Ninhydrin test reagents: monitor 1, monitor 2, and monitor 3 (Perkin-Elmer Applied Biosystems) HF/p-cresol solution (see recipe) Diethyl ether, cold 10% to 50% acetonitrile in H2O containing 0.1% TFA 15-ml manual peptide synthesis vessel (Peptides International) Black rubber tubing (1/4 in. i.d. × 5/8 in. o.d. × 3/16 in. wall thickness; Fisher), resistant to acids and organic solvents 2-liter side-arm flasks with rubber stoppers and glass tubing to fit Pasteur pipet containing glass wool for filtration 2-ml polypropylene column (Microcolumn X from Isolab) with Teflon stopcock 13 × 100–mm glass test tube 110°C heating block 60% ethanol HF cleavage apparatus (Peptides International) Additional material and equipment for solid-phase peptide synthesis (Schnölzer et al., 1992; UNIT 18.1), preparative C18 reversed-phase HPLC (see Basic Protocol 1, step 11), and ESI-MS (Chapter 16) Prepare the MBHA resin 1. Build a manual solid-phase peptide synthesis system (Fig. 18.4.4) consisting of a 15-ml manual peptide synthesis vessel attached by 12 in. of black rubber tubing through a rubber stopper to a 2-liter side-arm flask, which is in turn connected to a vacuum source.
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2. Place 1 g of MBHA resin (1.0 mmol/g) into the peptide synthesis vessel, add enough DMF to cover the dry resin (∼5 ml), and wait 30 min. This preswelling step is crucial for the success of any reaction carried out on a solid support, since it renders the functional groups on the polymer available for reaction.
3. Drain the resin. Cover the resin with 5% DIEA/DMF, leave for 1 min, then drain the resin. Repeat this process two more times, then perform three 20-sec flow washes with DMF—i.e., wash the resin with a continuous flow of DMF for 20 sec while keeping a constant volume of solvent above the resin bed (usually 3 to 4 mm), then drain the resin. A continuous flow wash is an extremely efficient way of exchanging solvent in a swollen polymer—much more so than the commonly used bulk wash (i.e., by adding solvent, mixing, and draining).
Prepare the symmetrical anhydride of 3-bromopropionic acid 4. Dissolve 1.222 g 3-bromopropionic acid (8 mmol) in a minimal volume of DCM (∼1 to 2 ml), then add 630 µl of 99% DIPC (4 mmol). Shake the solution vigorously and wait 10 min. A white precipitate corresponding to the diisopropyl urea will appear during the activation reaction. If no such precipitate forms, the reaction has not taken place and should be repeated. It is best to prepare the symmetrical anhydride fresh for each coupling reaction.
5. Filter the solution through a Pasteur pipet containing glass wool. It is very important to remove the diisopropyl urea by filtration before adding the symmetrical anhydride solution to the resin. The authors have found that if this step is not carried out the acylation reaction does not go to completion.
Couple the symmetrical anhydride of 3-bromopropionic acid to the MBHA resin 6. Add the filtered solution from step 5 to the MBHA resin in the peptide synthesis vessel. Without applying any suction, add 800 µl DIEA (4.5 mmol) and the minimal amount of DMF (1 to 2 ml) required to give a good slurry. 7. Leave the coupling reaction for 30 min with occasional stirring using a glass stirring rod.
solid-phase peptide reaction vessel vacuum resin stopcock
waste container
Native Chemical Ligation of Polypeptides
rubber tubing resistant to acids and organic solvents Figure 18.4.4 Suggested apparatus for the manual solid-phase synthesis of thioester peptides.
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8. Drain the resin and wash thoroughly with three 20-sec DMF flow washes as in step 3. 9. Repeat the coupling process (steps 4 to 8) two more times. 10. Add 5 ml Ac2O/DIEA/DMF solution to the resin and wait 10 min. Drain the resin and wash three times with DMF as in step 3. This acetylation step ensures that the small amount of unreacted amine groups remaining on the MBHA resin cannot participate in any subsequent acylation steps.
Prepare the 3-mercaptopropionamide-MBHA resin 11. Add 5 ml AcSH/DIEA/DMF solution to the resin and wait 20 min. Drain the resin and wash three times with DMF as in step 3. Repeat this entire process (treatment with AcSH/DIEA/DMF solution, along with the washings) two more times. 12. Add 5 ml BME/DIEA/DMF solution to the resin and wait 20 min. Drain the resin and wash three times with DMF as in step 3. Repeat this entire process (treatment with BME/DIEA/DMF solution, along with the washings) two more times. The 3-mercaptopropionamide-MBHA resin which is the product of this step is susceptible to oxidation upon long-term storage. The authors therefore recommend that it be immediately acylated with the appropriate Boc amino acid derivative.
Couple the first Boc amino acid to the 3-mercaptopropionamide-MBHA resin 13. Dissolve 3 molar equivalents of the appropriate Boc-AA-OSu (where AA is the first amino acid to be incorporated in the synthesis) in ∼6 ml DMF, then add this solution to the 3-mercaptopropionamide-MBHA resin. Add 714 µl DIEA (4 molar equivalents, 4 mmol) and leave the coupling reaction for 3 to 4 hr with occasional stirring. Most of the Boc amino acid derivatives are commercially available as N-hydroxysuccinimide esters. If the required Boc amino acid derivative is not commercially available it can be readily prepared manually (Bodanszky and Bodanszky, 1994).
14. Drain the resin and wash three times with DMF as in step 3. Add 5 ml Ac2O/DIEA/DMF solution to the resin and wait 10 min, then wash the resin three times with dichloromethane (DCM) using 20-sec flow washes as described in step 3. Dry the resin under vacuum and store at −20°C. Determine the final substitution of the Boc-amino acyl-3-mercaptopropionamideMBHA resin 15. Place ∼5 mg of the dry Boc–amino acyl–3-mercaptopropionamide–MBHA resin from step 14 in a 2-ml polypropylene Microcolumn X column with a Teflon stopcock, connected to a vacuum source. 16. Add 1 ml TFA to the resin and wait 2 min. Drain the column, then flow wash first with DMF and then with DCM. Drain again and dry under vacuum. 17. Weigh an exact amount of the dried resin (∼3 to 5 mg) and place in a 13 × 100–mm glass test tube. 18. Add 2 drops of ninhydrin test monitor 1 reagent, 4 drops of ninhydrin test monitor 2 reagent, and 2 drops of ninhydrin test monitor 3 reagent, then incubate 5 min at 110°C in a heating block. 19. Dilute the blue solution with 60% ethanol to 25 ml and mix well. Measure the absorbance at 570 nm using 1-cm path-length cuvette. Calculate the substitution of the thioester resin (in mmol/g) as 1.67 × (A570/amount of resin in mg).
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For a resin with an initial loading of ∼1 mmol/g the final substitution for the corresponding acylated thioester resin is usually ∼0.2 mmol/g.
Perform solid-phase synthesis of the α-thioester polypeptide 20. Synthesize the polypeptide sequence using the in situ neutralization/HBTU activation protocols for Boc-SPPS (Schnölzer et al., 1992; UNIT 18.1). Solid-phase deprotection of the 2,6-dinitrophenyl (Dnp) and formyl (For) protecting groups (commonly used for the protection of the side chains of His and Trp residues, respectively) results in premature cleavage of α-thioester polypeptides from the resin. Therefore, these residues should either be deprotected after the ligation reaction is complete or incorporated as Boc-Trp-OH (no side-chain protection) or Boc-Trp(Hoc)-OH and Boc-His(Bom)-OH during SPPS .
21. Once the synthesis is complete, flow wash with DMF and DCM as in step 14. Dry the resin under vacuum and store at −20°C. Cleave and purify the α-thioester polypeptide 22. Using 200 mg of peptide α-thioester resin from step 21, cleave the peptide-resin by treating with 5 ml of HF:p-cresol solution for 1 hr at 4°C in an HF cleavage apparatus. CAUTION: Anhydrous HF is a highly toxic and corrosive gas and should only be manipulated in a fume hood using the commercially available specialized apparatus.
23. Remove the HF under vacuum and resuspend both peptide and resin in ∼40 ml cold diethyl ether with gentle stirring for 10 min. Filter the suspension on a glass Buchner funnel under vacuum, without letting air pass through the filter, as this could oxidize the cleaved peptide. 24. Wash the material in the filter (containing the cleaved peptide and the resin) three times, each time with 10 ml cold diethyl ether. Wash an additional three times, each time with 10 ml dichloromethane, then wash with 10 ml cold diethyl ether again. Discard the wash liquid. These washes will remove most of the scavenger byproducts.
25. Add 10 ml freshly degassed 50% acetonitrile in water containing 0.1% TFA to the filter, wait 10 min (to dissolve the cleaved peptide), then filter. Repeat this process three more times, then recover and lyophilize the filtrates. 26. Analyze a 50-µl aliquot by reversed-phase HPLC and ESI-MS (Chapter 16; also see Basic Protocol 1, step 7). 27. Purify the lyophilized propionamide α-thioester peptide by preparative C18 RPHPLC (see Basic Protocol 1, step 11). SUPPORT PROTOCOL 4
Native Chemical Ligation of Polypeptides
BACTERIAL EXPRESSION OF α-THIOESTER POLYPEPTIDES This protocol describes the preparation of α-thioester polypeptides using bacterial expression in E. coli. The desired polypeptide-encoding gene fragment is first cloned into a commercially available pTYB vector (Fig. 18.4.5). Following E. coli transformation, soluble expression of the polypeptide-intein-CBD (chitin binding domain) fusion protein is induced and the cells are harvested and lysed. The lysate is then loaded onto a chitin column and the fusion protein affinity purified. Finally, the target α-thioester polypeptide is cleaved from the column and eluted using a buffer containing ethanethiol. NOTE: Initial studies using an MBP-intein-CBD system indicate that the majority of amino acid residues, when located immediately before the intein N-terminal cysteine, allow both purification of fusion proteins and efficient cleavage with thiols. However, Pro,
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Figure 18.4.5 pTYB1, one of the series of pTYB vectors useful for the cloning, expression and purification of recombinant α-thioesters polypeptides. (A) pTYB1 vector; (B) sequence of pTYB1 cloning/expression region. This vector uses a bacteriophage T7 promoter-driven system (see UNIT 5.1 and Ausubel et al., 1999). The target gene is cloned into the multiple cloning site (MCS) polylinker region to create an in-frame fusion between the C-terminus of the target gene and the N-terminus of the gene encoding a modified intein. The DNA encoding a small chitin binding domain (CBD) has been also added to the C-terminus of the intein gene to allow the resulting fusion protein to be purified by affinity chromatography. Following purification, the target α-thioester polypeptide is obtained through a transthioesterification reaction by treating the fusion protein with ethanethiol. The pTYB vectors are available from New England Biolabs. Preparation and Handling of Peptides
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Cys, and Asn residues were found to inhibit the in vitro cleavage with thiols. Note that partial cleavage during bacterial expression can sometimes be observed in certain systems, resulting in a decrease in the yield of fusion precursors. For example, in the MBP system, Asp, Arg, His, and Glu residues gave rise to in vivo cleavage (>50%). Materials Gene construct encoding target polypeptide pTYB1 vector (New England Biolabs) E. coli BL21 (or any other suitable E. coli strain) LB medium containing 100 µg/ml ampicillin (APPENDIX 4A) 1 M isopropyl-1-thio-β-D-galactopyranoside (IPTG), filter sterilized Chitin beads slurry: 50% (w/v) suspension of chitin beads (New England Biolabs) in 40% ethanol Column buffer (see recipe) Lysis buffer (see recipe) Cleavage buffer (see recipe) Refrigerated centrifuge with Beckman JA-10 rotor and centrifuge buckets to hold 1 liter of culture 1.5 × 10–cm glass or polypropylene column Refrigerated centrifuge with Beckman JA-17 rotor and appropriate centrifuge tubes Shaker Additional reagents and equipment for introducing plasmid vectors into bacterial cells (APPENDIX 4D), growth of bacteria in liquid medium (APPENDIX 4A), lysis of bacterial cells using a French press (UNIT 6.3), ESI-MS (UNITS 16.1 & 16.2) Express the fusion protein 1. Insert the gene construct encoding the target polypeptide into the pTYB1 vector (see manufacturer’s instruction for vector and Ausubel et al., 1999) so as to express the target protein as an intein-CBD fusion. Introduce the vector into the E. coli BL21 cells (see APPENDIX 4D and Ausubel et al., 1999). Inoculate 15 ml LB/ampicillin with E. coli BL21 containing a pTYB vector expressing the desired protein fusion. Grow overnight with shaking at 37°C (APPENDIX 4A). 2. Inoculate 1000 ml LB/ampicillin with 10 ml of the overnight culture and grow with shaking at 37°C to an OD600 of 0.5 to 0.6 (APPENDIX 4A). 3. Add 1 ml of 1 M IPTG (final concentration of 1 mM) to the culture and continue incubation for 1 to 6 hr at 37°C. The optimal induction conditions (i.e., incubation time, temperature, and final IPTG concentration) for soluble expression will depend on the in vivo properties of the overexpressed protein, and should be optimized for every system.
4. Centrifuge the cell culture 10 min at 8700 × g (7000 rpm in Beckman JA-10 rotor), 4°C. Discard supernatant. Proceed to step 5 immediately or freeze pellet at −70°C indefinitely. If extract preparation is to be be carried out immediately, the chitin column may be prepared during the centrifugation step.
Prepare the affinity column 5. Add ∼10 ml of chitin bead slurry to a 1.5 × 10–cm column and allow liquid to drain just to the top of the packed resin bed (∼5 ml). Native Chemical Ligation of Polypeptides
6. Wash the column with 100 ml column buffer at a flow rate of 1 ml/min.
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Prepare the cell extract 7. If cell pellet was frozen, (step 4), thaw on ice. Resuspend pellet in 30 ml ice-cold lysis buffer. Lyse cells using a French press (UNIT 6.3). IMPORTANT NOTE: From this step on, all procedures should be carried out at 4°C.
8. Centrifuge lysate for 30 min (or until supernatant is clear) at 25,000 × g (14,000 rpm in Beckman JA-17 rotor), 4°C. Decant supernatant into a clean container on ice and discard pellet. The supernatant can be frozen and stored at −70°C indefinitely before continuing with the procedure. In order to monitor the expression, extraction, and purification steps, it is convenient to take small aliquots at every step and analyze them by SDS-PAGE (UNIT 10.1).
Purify the fusion protein 9. If extract is frozen, thaw on ice. Load onto the chitin column (at a rate no faster than 0.5 ml/min). Collect flowthrough and reapply to column, then repeat this process one more time. Chitin beads have a capacity of ∼2 mg of CBD-tagged protein per ml packed beads. Therefore, the amount of extract that can be loaded on the column will depend on the amount of soluble fusion protein in the extract.
10. Wash the column with 100 ml column buffer at a flow rate of 1 ml/min. Discard flowthrough. Be sure that all traces of crude extract have been washed off the sides of the column. Obtain the α-thioester polypeptide 11. Add 5 ml cleavage buffer to the column and gently shake the reaction slurry overnight at room temperature. 12. Drain the beads, then wash with 15 ml cleavage buffer (three times, each time with 5 ml). Pool all of the fractions. IMPORTANT NOTE: Thioesters are susceptible to hydrolysis under alkaline conditions; consequently purification and storage of α-thioester polypeptides should be performed at pH 6.0 or below.
Analyze and purify the α-thioester polypeptide 13. Analyze a 50-µl aliquot by reversed-phase analytical HPLC and ESI-MS (see UNITS 16.1 & 16.2; also see Basic Protocol 1, step 7). 14. Purify the target α-thioester polypeptide by reversed-phase preparative HPLC see Basic Protocol 1, step 11). CHEMICAL SYNTHESIS OF Nα(Msc)-CYS, α-THIOESTER POLYPEPTIDES This protocol describes how to introduce the Nα(methylsulfony)ethyloxycarbonyl (Msc) protecting group onto the α-amino group of an N-terminal Cys polypeptide synthesized on a 3-mercaptopropionamide-MBHA resin (see Support Protocol 3). The resulting Nα(Msc)-Cys, α-thioester polypeptides are used in sequential native chemical ligation reactions (see Basic Protocol 2). NOTE: Boc-SPPS of α-thioester polypeptides requires the use of the Bom side-chain protecting group for His. The Bom group releases fomaldehyde during its deprotection with HF; however the formation of the thiazolidine adduct with the N-terminal Cys cannot take place due to the presence of the NαMsc group.
SUPPORT PROTOCOL 5
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Additional Materials (also see Support Protocol 3) Fully protected Boc-polypeptide-3-mercaptopropionamide-MBHA resin (see Support Protocol 3, step 21), dried (Methylsulfonyl)-ethyl 4-nitrophenyl carbonate (Msc-ONp; Fluka) Prepare the Boc-polypeptide-3-mercaptopropionamide-MBHA resin 1. Place ∼0.5 g of fully protected Boc-polypeptide-3-mercaptopropionamide-MBHA resin in a peptide synthesis vessel attached to a vacuum source and swell with DMF (see Support Protocol 3, steps 1 and 2). 2. Deprotect the Boc group by adding ∼5 ml of TFA to the resin, waiting 1 min, then draining the resin. Repeat this process one additional time, then drain the resin and perform three 20-sec flow washes with DMF (see Support Protocol 3, step 3). 3. Neutralize the polypeptide-3-mercaptopropionamide-MBHA resin by treating with 5% DIAE/DMF and flow washing with DMF (see Support Protocol 3, step 3). Introduce the Msc group on the α-amino group of the polypeptide-3-mercaptopropionamide-MBHA resin 4. Dissolve 560 mg (2 mmol) of Msc-ONp in 4 to 5 ml DMF and add this mixture to the peptide thioester resin, then add 340 µl (2 mmol) of DIEA. Leave the coupling reaction for 3 hr, with occasional stirring using a glass stirring rod, then drain the resin and perform three 20-sec flow washes with DMF. 5. Place ∼1 mg of resin from step 4, in a column (see Support Protocol 3, step 15). Flow wash first with DMF and then with DCM, drain again, and dry under vacuum. Place the aliquot in a glass test tube and perform the ninhydrin test (see Support Protocol 3, step 18). If the color of the solution is strongly blue, repeat the coupling reaction (step 4) and test again; if not, continue with step 6, below. Sometimes the ninhydrin test can give false positives (i.e., strong blue coloration) even although the acylation reaction is complete. This is especially true when the α-amino group is protected with a base-labile group (i.e., Fmoc or Msc). Therefore, if the ninhydrin test is still positive after the third coupling, proceed to step 6.
6. Perform three 20-sec flow washes with DMF and then with DCM. Dry the resin under vacuum and store at −20°C until further use. 7. Cleave and purify 200 mg of peptide-thioester resin (see Support Protocol 3, steps 22 to 26). REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Ac2O/DIEA/DMF solution 15 parts (v/v) 99% acetic anhydride (Ac2O) 15 parts (v/v) diisopropylethylamine (DIEA, peptide synthesis grade; Perkin-Elmer Applied Biosystems) 70 parts (v/v) dimethylformamide (DMF, spectrophotometric grade; Fisher) Prepare fresh
Native Chemical Ligation of Polypeptides
AcSH/DIEA/DMF solution 1 part (v/v) 96% thiolacetic acid (AcSH; Fluka) 1 part (v/v) diisopropylethylamine (DIEA, peptide synthesis grade; Perkin-Elmer Applied Biosystems) 8 parts (v/v) dimethylformamide (DMF, spectrophotometric grade; Fisher) Prepare fresh
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BME/DIEA/DMF solution 1 part (v/v) 98% 2-mercaptoethanol (BME) 1 part (v/v) diisopropylethylamine (DIEA, peptide synthesis grade; Perkin-Elmer Applied Biosystems) 8 parts (v/v) dimethylformamide (DMF, spectrophotometric grade; Fisher) Prepare fresh Cleavage buffer 0.1 mM EDTA 200 mM sodium phosphate, pH 6.0 250 mM NaCl 3% (v/v) ethanethiol 0.1% (v/v) Triton X-100 Adjust pH to 6.0 with 1 M NaOH or HCl Store up to 6 months at 4°C Column buffer 0.1 mM EDTA 20 mM sodium phosphate, pH 7.2 250 mM NaCl 0.1% (v/v) Triton X-100 Adjust pH to 7.2 with 1 M NaOH or HCl Store up to 6 months at 4°C Guanidine⋅HCl buffer, 6 M 6 M guanidine⋅HCl (APPENDIX 3A) 0.1 M sodium phosphate 1 mM EDTA Adjust pH to 7.5 with 1 M NaOH Store up to 6 months at room temperature HF/p-cresol solution 96 parts (v/v) hydrogen fluoride (HF; anhydrous) 4 parts (v/v) p-cresol Prepare fresh CAUTION: Anhydrous HF is a highly toxic and corrosive gas and should be manipulated only in a fume hood, using the commercially available specialized apparatus (HF cleavage apparatus from Peptides International).
Lysis buffer 0.1 mM EDTA 1 mM PMSF 25 mM HEPES, pH 8.0 250 mM NaCl 5% (v/v) glycerol Adjust pH to 8.0 with 1 M NaOH Store up to 6 months at 4°C COMMENTARY Background Information The introduction of solid-phase peptidesynthesis (SPPS) by Bruce Merrifield revolutionized the chemical synthesis of peptides (Merrifield, 1963; also see UNIT 18.1). Despite the enormous impact of SPPS in the generation
and study of small bioactive peptides, it is now clear that the combination of incomplete acylation/deprotection reactions and other well documented side reactions places an intrinsic limit on the size of peptides accessible by efficient stepwise SPPS. Thus, polypeptides of up
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Native Chemical Ligation of Polypeptides
to ∼50 residues in length can be prepared by stepwise SPPS with reasonable confidence, but beyond this, the chances of success fall off precipitously. In order to overcome this size limitation, recent years have seen renewed interest in the use of convergent synthetic strategies—i.e., the synthesis of large polypeptides from smaller peptide building blocks which are themselves accessible via the SPPS approach. It should be noted that fragment condensation has a long and illustrious history in the peptide chemistry field; however, there have always been serious problems associated with the manipulation of the fully protected peptides in these classical convergent strategies. Although the use of minimal protection strategies represented a step in the right direction (see LloydWilliams et al., 1993 for an extensive review), it was not until the early 1990s that a truly practical way of performing fragment condensations was developed—i.e., chemical ligation (for reviews see Muir, 1995; Wallace, 1995). The original chemical ligation strategies were all based on the premise that an unnatural moiety could be used to covalently join two fully unprotected (and hence water-soluble) polypeptides, each bearing unique and mutually reactive groups. A number of different chemistries have been developed for this purpose, all of which give rise to an unnatural covalent structure at the ligation site (Muir, 1995; Wallace, 1995). In 1994, a second-generation ligation chemistry was introduced, known as “native chemical ligation,” which allows the preparation of proteins with native backbone structures from fully unprotected peptide building blocks (Dawson et al., 1994). This important extension of the chemical ligation concept makes use of the mild acylating power of the α-thioester functionality. The principle of native chemical ligation is depicted in Figure 18.4.1. The first step involves the chemoselective reaction which occurs between the free thiol group of an unprotected N-terminal Cys-polypeptide and a second, unprotected polypeptide containing an α-thioester group. This transthioesterification reaction gives rise to a thioester-linked intermediate which spontaneously rearranges to form a native peptide bond at the ligation site. The target full-length polypeptide is thus obtained without any further manipulation. Native chemical ligation reactions are performed in aqueous buffers at pH 7 to 7.5 in the presence of thiol cofactors. At this pH, the regioselectivity of the reaction is such that the reaction can to be performed in the presence of all the
functionalities commonly found in proteins. Even the presence of additional Cys residues in one or both fragments does not affect the regioselectivity of the ligation (Hackeng et al., 1997). Small proteins or protein domains ∼100 to 120 amino acids in length can be reliably generated from two peptide building blocks in a single chemical ligation step (for a few examples see Dawson et al., 1994, 1997; Lu et al., 1996; Hackeng et al., 1997; Camarero et al., 1998). The total chemical synthesis of larger protein targets (>100 residues) via the ligation of just two fragments becomes more and more problematic as the size increases. This is due to the difficulties associated with the direct stepwise SPPS of polypeptide segments bigger than 50 residues. This difficulty can be overcome by performing multiple ligation reactions using three or more synthetic peptides (e.g., Canne et al., 1995). Native chemical ligation has been extended to allow multiple ligation steps to be performed sequentially in a controlled and directed way (Muir et al., 1997; Camarero et al., 1998). The general approach is depicted in Figure 18.4.2. Key to this strategy is the temporary protection of the α-amino group of the central peptide with the base-labile 2-(methylsulfonyl)ethyloxycarbonyl (Msc) group (Tesser and Balvert-Geers, 1975). The presence of this protecting group prevents the N-terminal Cys from reacting in an intramolecular or intermolecular fashion with the α-thioester functionality present in the same polypeptide fragment. Once the first ligation reaction is finished, the Nα-Msc group can be efficiently removed, allowing the next ligation step to be performed (Fig. 18.4.2). The native chemical ligation approach can also be used in the semisynthesis of proteins (Muir et al., 1998; Severinov and Muir, 1998; Erlandson et al., 1996; Evans et al., 1998) from recombinant and synthetic polypeptide fragments. As described above, native chemical ligation of two polypeptides requires that one of the fragments possess an N-terminal Cys and that the other contain an α-thioester moiety. Polypeptides containing an N-terminal cysteine residue for use in litigation can be obtained using standard recombinant DNA expression methods (see Chapter 5). Importantly, biosynthetic methods are also now available for the generation of α-thioester polypeptides (Muir et al., 1998; Severinov and Muir, 1998; Evans et al., 1998). This is made possible using the IMPACT expression system, commercially available from New England Biolabs (Chong
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A
HS intein
NH
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B
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clone protein gene in IMPACT system
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+
CBD
i) 3% ethanethiol at pH 6 overnight ii) filtration
C-extein SH
recombinant protein
CO-SCH2 CH3
O intein NH O
Figure 18.4.6 Principles of the biosynthetic preparation of α-thioester polypeptides by recombinant techniques. (A) Scheme representing the proposed mechanism of protein splicing involving the intein from Saccharomyces cerevisiae VMA1 gene (Xu and Perler, 1996). (B) Expression, purification, and cleavage of polypeptide-intein*-CBD fusion protein (where the asterisk refers to the mutation Asn454→Ala in the intein element and CBD refers to the chitin binding domain) with an appropriate thiol to give the α-thioester polypeptide.
et al., 1997). This system utilizes a protein splicing element, an intein from the Saccharomyces cerevisiae VMA1 gene, in conjunction with a chitin binding domain (CBD) which allows purification by affinity chromatography (Fig. 18.4.6). The natural intein has been modified in the expression system (Asn454→Ala) in order to block the normal protein splicing reaction in midstream. This results in the formation of a thioester linkage between the polypeptide of interest and the intein (Fig. 18.4.6). Cleavage of the thioester can thus be induced by treatment of the expressed fusion protein with the appropriate
thiol to give the target α-thioester polypeptide (Muir et al., 1998; Severinov and Muir, 1998; Evans et al., 1998).
Critical Parameters and Troubleshooting As mentioned in Basic Protocol 1, pH is the most critical parameter in the native chemical ligation of two fully unprotected peptides in aqueous buffers. The efficiency of the ligation reaction is strongly dependent on the pH; above pH 8.0 the reaction loses its regioselectivity and below pH 6.0 the reaction is usually very slow.
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Another crucial factor is the concentration of the two reactants. The bimolecular nature of native chemical ligation (and ligations in general) means that the concentration of both reactants should be as high as possible for efficient reaction. Generally the use of chemical denaturants (GdmCl or urea) will allow high concentrations of both reactants to be achieved. Furthermore, the use of denaturing conditions helps to alleviate potential steric problems that may be associated with the use of folded polypeptides. Another important parameter for the success of the ligation reaction is the availability of the thiol and/or α-amino groups of the N-terminal Cys polypeptide during the ligation reaction. If one or both groups are chemically blocked, the ligation will not take place. It is well known that N-terminal Cys peptides can react rapidly with carbonyl-containing compounds (e.g., acetone and formaldehyde) to give the corresponding N-terminal thiazolidine adducts, which are totally unreactive in the native ligation process. It is thus crucial to avoid the use of these substances while handling all peptides and buffers. Note that the authors have found acetone, a commonly used solvent for washing glassware in many laboratories, to be particularly problematic in this regard.
Anticipated Results Native chemical ligation has been applied to the synthesis and semisynthesis of a large number of proteins and protein domains. These studies indicate that very high yields (80% or better) are typically obtained for the ligation step. Following purification, the total yield usually drops to 50% to 60%. In sequential ligations where the Nα-Msc group has to be deprotected in situ after the first ligation step, the typical total yield after purification is ∼30% to 40%. It is also important to note that the protein-folding step may further decrease the final yield of product.
Time Considerations
Native Chemical Ligation of Polypeptides
In Basic Protocols 1 and 2, the chemical ligation step typically requires 2 days, although in some cases the ligation reaction can proceed very rapidly (in a few hours) or somewhat more slowly (in 4 days). The purification step (RPHPLC or other liquid chromatographies) can be performed in half a day. Processing of the purified samples (lyophilization or concentration) can take 1 to 2 days (depending on the volume). Finally, folding of the protein (if necessary) can be achieved in 3 days.
The thioester resin can be prepared in 1 day. The time required for the chemical synthesis of an α-thioester polypeptide (see Support Protocol 3) will depend on its size. Typically, for peptides ∼50 residues in length, the solid-phase chain assembly can be carried out in 3 days manually or in 1 day using an automated synthesizer. The deprotection-cleavage and purification can be performed in 1 day, and lyophilization of peptide fractions can take 1 to 2 days (depending on the volume). Bacterial expression of a peptide-inteinCBD fusion protein and preparation of the crude cell extract requires 2 days for the system described here (see Support Protocol 4). Column preparation requires ∼2 to 3 hr. Loading and washing the column can be done in 3 hr. The cleavage of the α-thioester polypeptide from the affinity column requires 10 to 15 hr (overnight). Purification and processing of the polypeptide fractions requires 1 to 2 days.
Literature Cited Ausubel, F.A., Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G., Smith, J.A., and Struhl, K. (eds.). 1999. Current Protocols in Molecular Biology. John Wiley & Sons, New York. Bodanszky, M. and Bodanszky, A. 1994. The Practice of Peptide Synthesis, 2nd ed. Springer-Verlag, Berlin. Camarero, J.A., Cotton, G.J., Adeva, A., and Muir, T.W. 1998. Chemical ligation of unprotected peptides directly from a solid support. J. Pept. Res. 51:303-316. Canne, L.E., Ferre-D’Amare, A.R., Burley, S.K., and Kent, S.B.H. 1995. Total chemical synthesis of a unique transcription factor-related protein: cMyc-Max. J. Am. Chem. Soc. 117:2998-3007. Chong, S., Mersha, F.B., Comb, D.G., Scott, M.E., Landry, D., Vence, L.M., Perler, F.B., Benner, J., Kucera, R.B., Hirvonen, C.A., Pelletier, J.J., Paulus, H., and Xu, M.Q. 1997. Single-column purification of free recombinant proteins using a self-cleavable affinity tag derived from a protein splicing element. Gene 192:271-281. Dawson, P.E., Muir, T.W., Clark-Lewis, I., and Kent, S.B.H. 1994. Synthesis of proteins by native chemical ligation. Science 266:776-779. Dawson, P.E., Churchill, M.J., Ghadiri, M.R., and Kent, S.B.H. 1997. Modulation in native chemical ligation through the use of thiol additives. J. Am. Chem. Soc. 119:4325-4329. Erlandson, D.A., Chytill, M., and Verdine, G.L. 1996. The leucine zipper domain controls the orientation of AP-1 in the NFAT AP-1 DNA complex. Chem. Biol. 3(12):981-991. Evans, T.C., Benner, J.J., and Xu, M.-Q. 1998. Semisynthesis of cytotoxic proteins using a modified protein splicing element. Protein Sci. 7:22562264.
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Hackeng, T.M., Mounier, C.M., Bon, C., Dawson, P.E., Griffin, J.H., and Kent, S.B. 1997. Total chemical synthesis of enzymatically active human type II secretory phospholipase A2. Proc. Natl. Acad. Sci. U.S.A. 94:7845-7850.
Schnölzer, M., Alewood, P., Jones, A., Alewood, D., and Kent, S.B.H. 1992. In situ neutralization in Boc-chemistry solid phase peptide synthesis: Rapid, high yield assembly of difficult sequences. Int. J. Pept. Protein Res. 40:180-193.
LLoyd-Williams, P., Albericio, F., and Giralt, E. 1993. Convergent solid-phase peptide synthesis. Tetrahedron 49:11065-11133.
Severinov, K. and Muir, T.W. 1998. Expressed protein ligation: A novel method for studying protein-protein interactions in transcription. J. Biol. Chem. 273:16205-16209.
Lu, W., Qasim, M.A., and Kent, S.B.H. 1996. Comparative total synthesis of turkey ovomucoid third domain by both stepwise solid phase synthesis and native chemical ligation. J. Am. Chem. Soc. 118:8518-8523. Merrifield, R.B. 1963. Solid phase peptide synthesis. I. The synthesis of a tetrapeptide. J. Am. Chem. Soc. 85:2149-2154. Muir, T.W. 1995. A chemical approach to the construction of multimeric protein assemblies. Structure 3:649-652. Muir, T.W., Dawson, P.E., and Kent, S.B.H. 1997. Protein synthesis by chemical ligation of unprotected peptides in aqueous solution. Methods Enzymol. 289:266-298. Muir, T.W., Sondhi, D., and Cole, P.A. 1998. Expressed protein ligation: A general method for protein engineering. Proc. Natl. Acad. Sci. U.S.A. 95:6705-6710.
Tesser, G.I. and Balvert-Geers, I.C. 1975. The methylsulfonylethyloxycarbonyl group: A new and versatile amino protective function. Int. J. Pept. Prot. Res. 7:295-305. Wallace, C.J.A. 1995. Peptide ligation and semisynthesis. Curr. Opin. Biotechnol. 6:403-410. Xu, M.-Q. and Perler, F.B. 1996. The mechanism of protein splicing and its modulation by mutation. EMBO J. 15:5146-5153.
Contributed by Julio A. Camarero and Tom W. Muir The Rockefeller University New York, New York
Preparation and Handling of Peptides
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Synthesis and Application of Peptide Dendrimers As Protein Mimetics
UNIT 18.5
Peptide dendrimers are branched polymers with peptides attached centrally to a template or core matrix. They are synthesized as defined dendritic structures. Their molecular weights increase geometrically as a function of generation branching of monomers. Usually, 2 to 16 peptidyl branches of the same or different sequences are used to form a peptide dendrimer. Architecturally, peptide dendrimers belong to the multichained, cascade-shaped polymer family. Thus, they contain multiple N- or C-terminal ends that can be exploited as a platform for multitasks. Such a molecular architecture is different from conventional polypeptides or proteins that contain a single unbranched contiguous structure with two terminal ends. Peptide dendrimers are also different from organic star-burst dendrimers, which are spherically shaped and often consist of >16 hyperbranched small organic monomers. However, the fundamental dendrimeric design consisting of monomers radiated from a center core is similar, conceptually, to the organic dendrimers. Peptide dendrimers come in different forms and sizes. Their forms are determined by the templates and peptide structures while their sizes are determined by the numbers and lengths of peptide monomers. Although their sizes vary over a wide range from 3 to >100 kDa, most peptide dendrimers reported to date are in the range of 6 to 20 kDa. A driving force in developing peptide dendrimers is the effort to mimic forms and functions of proteins. Thus far, peptide dendrimers have found applications in four active areas: (1) immunological applications as immunogens and antigens; (2) de novo design of artificial proteins; (3) protein mimetics as agonists and antagonists in drug discovery; and (4) new biopolymers and biomaterials. However, this unit will focus on the use of peptide dendrimers as protein mimetics. The use of peptides to mimic a portion of a protein structure is a challenging and powerful tool in the discovery of new drugs. In native proteins, discontinuous bioactive peptide surfaces are held together in a particular conformation by the structural rigidity of the protein. One approach to mimicking the structural surface on the protein is to bring the potential peptide sequences together by assembling the peptide chains on a template as a peptide dendrimer. Recently, small peptides have been demonstrated to adopt well defined conformations when they are attached on a template (Schneider and Kelly, 1995). In the past 15 years, templates suitable for protein engineering have been developed. These templates are flexible dendrimeric peptides (Tam, 1988; Wrighton et al., 1997) and cyclic peptides (Mutter and Vuilleumier, 1989), as well as more rigid organic molecules (Unson et al., 1984; Sasaki and Kaiser, 1989). In 1988, the multiple antigenic peptide (MAP) system was introduced in the authors’ laboratory (Tam, 1988; Tam and Lu, 1989) as a novel approach to preparing peptide immunogens. The MAP consists of an inner core matrix built up of a large layer of Lys residues and a surface of peptide chains attached to the core matrix (Fig. 18.5.1). Because of its dendrimeric structure, MAP can be very useful as a template for assembling potential peptide surfaces. Furthermore, the effect of MAP on secondary structures has also been studied (Shao and Tam, 1995). These study results showed that the peptide dendrimer has an increased ordered helical structure, while the peptide monomer and the tetravalent lysinyl core matrix do not have any ordered structure. The synthetic schemes for preparing MAPs suitable for protein mimetics are the same as those for generating MAPs as peptide immunogens. In addition to the applications in immunology, examples in the literature Contributed by James P. Tam and Jane C. Spetzler Current Protocols in Protein Science (1999) 18.5.1-18.5.35 Copyright © 1999 by John Wiley & Sons, Inc.
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B
A
NH NH NH
NH NH
NH Ala-OH
Ala-OH
NH NH
NH NH
NH
NH
Figure 18.5.1 Schematic drawing of the MAP system. (A) Tetravalent MAP core matrix. (B) Octavalent MAP core matrix. The solid dots represent Lys and the ribbons with arrows represent the peptide.
have applied MAPs in areas such as inhibitors, artificial proteins, affinity purifications, and intracellular transport. The MAP, which is chemically defined and homogeneous, was first intended as an approach to overcoming the limitations of the conventional method for producing antipeptide antibodies. In the conventional approach, the peptide antigen is conjugated to a known large protein, or synthetic polymer carrier, to form a peptide-carrier conjugate. Although this approach has been used successfully in eliciting animal antibodies, it has several inherent limitations. First, only a small portion of peptide antigen is represented in the whole conjugate. Second, there is chemical ambiguity in the antigen composition and structure. Third, irrelevant epitopes and antibodies may be produced, and finally, carrier toxicity and carrier-induced epitope suppression may occur. The MAP systems have been used successfully to produce both polyclonal and monoclonal antibodies that specifically recognize native proteins. They have also been used to produce sera that have a significantly higher titer of antibodies than sera with antibodies against the same peptides conjugated to keyhole limpet hemocyanin (KLH) as a carrier protein (Tam, 1988).
Synthesis and Application of Peptide Dendrimers As Protein Mimetics
Cyclic peptides are a powerful approach in the design of hormones and bioactive peptides, since favorable conformational constraints associated with biological activity can be fixed and conformational mobility diminished. In addition, the stability against biodegradation is increased. Cyclic peptides are also beneficial as peptide antigens, since they can elicit higher-quality antibodies than their linear counterparts. Based on these demands, the cyclic multiple antigen peptide (cMAP) approach (Spetzler and Tam, 1996; Zhang and Tam, 1997), which has branched multiple closed-chain architectures, was developed. The cMAP system is a superior approach for protein mimetics because the multiple constrained peptides can mimic bioactive conformations. Whether to select this approach over MAP depends on the properties of the peptides, but usually if the peptides are too small to adopt a stable conformation on their own, incorporation of a cyclic structure may be necessary. Two methods for preparing MAP systems are described in this unit. The direct approach for preparing MAP systems is presented in the first two protocols, including the procedure
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direct functionalized peptide
peptide MAP core matrix
functionalized MAP core
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functionalized functionalized X Y free MAP core free peptide
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Figure 18.5.2 The direct and indirect approaches for preparing MAPs. Spheres represent the solid support; the MAP core matrix is Lys2-Lys-Ala-OH or Lys4-Lys2-Ala-OH; X represents nucleophile; Y represents electrophile.
for t-butyloxycarbonyl (Boc) chemistry (see Basic Protocol 1) and the procedure for 9-fluorenylmethyloxycarbonyl (Fmoc) chemistry (see Alternate Protocol). An indirect approach for preparing MAP systems, in which peptide and core matrix are synthesized separately and conjugated by several ligation methods, is then described (see Basic Protocols 2 and 3). The cMAP approach is also executed using either the direct or indirect approach, but requires an additional cyclization step to constrain the peptides after synthesis. The synthesis of cMAP is described (see Basic Protocols 4, 5, and 6), and the preparation of cyclic peptides is illustrated (see Support Protocols 4, 5, and 6). Support Protocol 1 describes the ninhydrin assay to assess the completeness of the coupling reaction. In most cases, MAP systems can be used directly after simple dialysis or desalting. Some immunological studies, however, require purified MAPs. Support Protocols 2 and 3 describe MAP system purification by dialysis and high-performance gel-filtration chromatography. STRATEGIC PLANNING Realizing the distinction between the direct and indirect approach is essential (Fig. 18.5.2) when synthesizing MAP systems. In the direct approach, the MAP core is formed by attaching several layers of lysine to the resin, which contains either Gly or Ala, using solid-phase peptide synthesis (Merrifield, 1963; UNIT 18.1). The peptide is then assembled onto this MAP core matrix (Basic Protocol 1 and Alternate Protocol). After the synthesis, the entire MAP is deprotected and cleaved from the resin. In the indirect approach, the MAP core and peptide are prepared and purified separately (Basic Protocols 2 to 3). Then, they are linked together in the absence of any side-chain protecting groups, after purification, to form a completed MAP. For preparing MAPs by either the direct or indirect approach, both the peptide synthesis and the ligation chemistry are easy to perform. Other features of the direct and indirect approach are discussed in the Commentary (see Background Information). However, the considerations for choosing a direct or indirect approach involve requirements for purity, properties of the peptide antigens, and time factors.
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-COCHOf NH2-OCH2CO-
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x CO2H s-s -CO- CH=N- OCH2 CO-
H2N x H2N
CO2H s-s
s-s
-COCHOf NH2-OCH2CO- Pallin and Tam, 1996 Cys
Cys
Tam et al., 1991
Figure 18.5.3 Available methods for preparing cyclic peptides from unprotected linear peptides. Notes: (a) arrow indicates the direction of the amide bond; (b) types of cyclic peptides are designated as e-e, end-to-end; e-s, end-to-side chain; and s-s, side-chain-to-side-chain; (c) an electrophile/nucleophile pair is placed at the ends or side chain of the linear peptide (e.g., for formation of e-e cyclic peptide, the amino nucleophile is Cys and the carbonyl electrophile can be a thioester); (d) R = (CH2)2-CONH2; (e) from periodate oxidation of a C-terminal glyceric ester; (f) from periodate oxidation of Ser and Thr attached to side chain of Lys.
For generating cMAPS, peptides on the core matrix need to be constrained as an additional step. A key tactic in the authors’ strategy for preparing cMAP is the use of unprotected peptides as precursors. In the past few years, the authors’ laboratory has developed a plethora of methods to form cyclic peptides from unprotected linear peptides in aqueous solution. Figure 18.5.3 lists those methods that are useful in preparing cMAPs. In planning of the cMAP synthesis, two questions need to be addressed: (1) how should the peptide antigen be constrained and (2) what are the suitable methods? The following are simple guidelines in the strategic planning for preparing cMAPs. There are three general methods for constraining peptides in the cMAP system: end-toend, end-to-side chain, and side chain-to-side chain (Figure 18.5.3). In one example (Basic Protocol 4) using the direct approach, cMAP is self-assembled via thiazolidine linkages after its unprotected precursor is synthesized by solid-phase synthesis. This procedure produces cyclic peptides that are linked in an end-to-side chain manner. A similar strategy can be applied for end-to-end or side chain-to-side chain methods by modifying the synthetic strategy accordingly. Peptides with a motif that favors a cyclic structure are highly recommended in this approach. Two other procedures (Basic Protocols 5 and 6) describe the use of an indirect approach for the synthesis of purified cMAP in two distinct steps. These protocols involve the synthesis of side chain-to-side chain and end-to-end cyclic peptides containing a free thiol, which are then linked to a MAP core matrix Synthesis and Application of Peptide Dendrimers As Protein Mimetics
A critical point in the preparation of both MAP and cMAP is the selection of the linear peptide sequences. Some of the available methods for identifying potential bioactive sequences include peptide library approaches (Tam et al., 1991; Houghten et al., 1991),
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simultaneous synthesis of a high number of peptides (UNIT 18.2), and computer-assisted modeling (UNIT 18.3). The procedures described in this unit involve use of an automated peptide synthesizer (e.g., Perkin-Elmer Applied Biosystems). For Boc syntheses, a special hydrogen fluoride cleavage apparatus (Peptide Institute or Peninsula Laboratories) and hood are required. Manual procedures can also be used (Stewart and Young, 1984). A detailed discussion of peptide synthesizers and reagents is presented in UNIT 18.1. CAUTION: Some chemicals used in these procedures are harmful. N-N′-dicyclohexylcarbodiimide (DCC) can cause severe allergy in sensitive persons. Avoid any skin contact with DCC! Benzotriazolyl N-oxytrisdimethylaminophosphonium hexafluorophosphate (BOP) and 2-(1H-benzotriazol-1-yl)-1,1,3,3-tetramethyl uronium hexafluorophosphate (BBTU) can be used as alternatives to DCC. Trifluoroacetic acid (TFA) is a toxic, volatile organic acid and should only be used under a hood. Besides being toxic, hydrogen fluoride (HF) is also highly corrosive to glass instruments. Never use glass instruments or containers in HF cleavage. Thick polyvinyl gloves and a protective face mask should be worn. More details about these chemicals can be found in the literature (Stewart and Young, 1984). DIRECT Boc SYNTHESIS OF MAP SYSTEMS In the direct approach for preparing MAP systems, the MAP core is first formed by attaching several levels of lysine onto an amino acid coupled to a resin matrix. The peptide sequence is then assembled stepwise on this MAP core resin. After peptide assembly is complete, the MAP system is cleaved from the resin support (Tam, 1988). The procedure for preparing MAP systems by the direct approach using Boc chemistry is presented in Figure 18.5.4 and Table 18.5.1. Solid-phase peptide synthesis (Merrifield, 1963) using
BASIC PROTOCOL 1
X-Ala-O
synthesis of MAP core matrix
X-HN X-HN-Lys
Lys2 -Lys-Ala-O 4
peptide synthesis Peptide Peptide
Lys
Lys2 -Lys-Ala-O 4
cleavage
Peptide Peptide
Lys
Lys2 -Lys-Ala-OH 4
MAPs
Figure 18.5.4 Direct approach of preparing MAPs with an octabranched core matrix. Sphere represents solid support; X represents Boc or Fmoc.
Preparation and Handling of Peptides
18.5.5 Current Protocols in Protein Science
Supplement 17
Table 18.5.1
Schedule for Boc Solid-Phase Peptide Synthesis
Reaction Description 1 2 3 4 5 6 7 8a 9 10 11 12 13c
DCM wash 50% (v/v) TFA/DCM prewash 50% (v/v) TFA/DCM deprotection DCM wash (5×) 5% (v/v) DIEA/DCM prewash 5% (v/v) DIEA/DCM neutralization DCM wash (5×) Boc-AA (3 eq) in DCM DCC (3 eq) in DCM Addition of DMF DMF wash (2 times) DCM wash (2 times) Ninhydrin test
Volume Repetitions (ml/g resin) and time 15 15 15 15 15 15 15 5 1 15 15 15 —
3 × 1 min 1 × 1 min 1 × 20 min 5 × 1 min 1 × 1 min 1 × 5 min 5×1 1 × 1 minb 1 × 30 minb 1 × 30 min 2 × 1 min 2 × 1 min 5 min
aIn case of coupling of Boc-Asn-OH, and Boc-Gln-OH, the DCC/HOBt coupling method should be
used to avoid dehydration of unprotected side chain amide groups. Alternatively use Boc-Asn(Trt)OH and Boc-Gln(Trt)-OH to minimize degradation side reactions. Boc-Arg(Tos)-OH should also be coupled with the DCC/HOBt method. bDo not drain until the completion of reaction 10. cIf ninhydrin test does not show completion of coupling reaction (>99.5%), repeat reactions 7 to 13.
Boc chemistry involves prewashing the resin, removing the Boc protecting group, neutralizing the resin, coupling the next Boc–amino acid residue, and then repeating this cycle until the complete peptide sequence is assembled. Finally, the peptide is cleaved from the resin support. Materials Synthesis reagents (Table 18.5.1; prepared as described in Stewart and Young, 1984): Dichloromethane (methylene chloride, DCM) 50% (v/v) trifluoroacetic acid (TFA) in DCM (TFA/DCM) 5% (v/v) diisopropylethylamine (DIEA) in DCM (DIEA/DCM) 1.0 M N,N′-dicyclohexylcarbodiimide (DCC) in DCM (DCC/DCM) N,N′-dimethylformamide (DMF) 0.1 mmol/g Boc-β-Ala-OCH2-PAM-resin (Bachem California) Boc-Lys(Boc) (Bachem California) Boc-amino acids (Boc-AA), with protected side-chain groups (Bachem California, Peninsula Laboratories) appropriate for synthesis of the desired peptide antigen, dissolved in dichloromethane (DCM). 10% (v/v) thiophenol in DMF or 10% (v/v) mercaptoethanol (2-ME)/5% (v/v) DIEA in DMF p-cresol p-thiocresol Dimethyl sulfide (DMS) Liquid hydrogen fluoride (HF), –78°C 99:1 (v/v) cold diethyl ether/2-mercaptoethanol (2-ME) 10% (v/v) acetic acid in water Synthesis and Application of Peptide Dendrimers As Protein Mimetics
Automated peptide synthesizer (e.g., Perkin-Elmer Applied Biosystems #ABI430A) Manual reaction vessel (Pierce) Hydrogen fluoride cleavage apparatus (Peptide Institute or Peninsula Laboratories)
18.5.6 Supplement 17
Current Protocols in Protein Science
50-ml round-bottom flask Coarse- and fine-porosity fritted glass funnels 50°C water bath Prepare the core matrix 1. Place 0.5 g of 0.1 mmol/g Boc-β-Ala-OCH2-PAM-resin in a reaction vessel in an automated peptide synthesizer. Begin the synthesis, using the synthesis reagents and directions outlined in Table 18.5.1, making the substitutions in reaction 8 of the table as described in step 2a, b, or c. Reactions 1 through 13 (Table 18.5.1) constitute one complete cycle; one cycle is required for each amino acid in the sequence. Some automated synthesizers may include the test to evaluate the coupling reaction in their cycle; if not, the ninhydrin test (reaction 13) will have to be performed manually (Support Protocol 1, Sarin et al., 1981). β-alanine is used as an internal standard for amino acid analysis which provides the molar ratios of other amino acids against β-alanine. Instead of β-alanine, other simple amino acids such as alanine can also be used.
2a. To prepare a MAP core matrix with two lysine branches: In reaction 8 of the first cycle, couple 0.15 mmol (52 mg) Boc-Lys(Boc) to the resin added in step 1. Proceed to step 3. 2b. To prepare a MAP core matrix with four lysine branches: Complete step 2a, then, in reaction 8 of the second cycle, couple 0.3 mmol (104 mg) Boc-Lys(Boc) to the resin from step 2a. Proceed to step 3. The substitution level doubles with each coupling of Boc-Lys(Boc).
2c. To prepare a MAP core matrix with eight lysine branches: Complete steps 2a and 2b, then, in reaction 8 of the third cycle, couple 0.6 mmol (208 mg) Boc-Lys(Boc) to the resin from step 2b. Proceed to step 3. Preformed MAP core matrix is also commercially available from Applied Biosystems, Bachem California, and Calbiochem-Novabiochem.
Synthesize the peptide on the MAP core matrix 3. Perform synthesis of the peptide on the MAP core matrix prepared in step 2a, b, or c by carrying out the reactions in Table 18.5.1 (one cycle of 13 reactions per amino acid), using, in reaction 8 of the table, 0.15 mmol of Boc-amino acids for a MAP core matrix with two branches, 0.3 mmol of Boc-amino acids for a MAP core matrix with four branches, or 0.6 mmol of Boc-amino acids for a MAP core matrix with eight branches, based on 0.05 mmol of starting Boc-β-Ala-OCH2-PAM resin. Select the appropriate Boc-amino acids for the synthesis of the peptide antigen. For descriptive purposes, the peptide antigen IEDNEYTARQG (IG-11) is used in this protocol. For this peptide, the Boc-amino acids would be Boc-Gly, Boc-Gln, Boc-Arg (Tos), Boc-Ala, Boc-Thr(Bz1), Boc-Tyr(BrZ), Boc-G1u(OBzl), Boc-Asn, Boc-Asp (OBzl), and Boc-Ile for the synthesis of IG-11-MAP. The peptide chain would be assembled in sequence G, Q, R, A, T, Y, E, N, D, E, I. The most commonly used side-chain-protecting groups for Boc amino acids are: Ser and Thr, benzyl ether (Bzl); Asp and Glu, benzyl ester (OBzl); Asn, and Gln, unprotected or trityl (Trt) amide; Met, unprotected; Tyr, 2-bromobenzyloxycarbonyl (BrZ); Arg, toluenesulfonyl (Tos); Lys, 2-chlorobenzyloxycarbonyl (CIZ); His, Nim-2,4-dinitrophenyl (Dnp); Trp, Nin-formyl, Cys, p-methylbenzyl (MeBzl), or acetamidomethyl (Acm). Some special side-chain-protecting groups can also be found in the literature listed at the end of this unit (see Literature Cited).
4. After synthesis of the peptide is complete, transfer the resin to a manual reaction vessel and remove the Dnp from peptide residues with Nim-Dnp protecting groups using 10% thiophenol in DMF (three 8-hr treatments, 5 ml each), or 10% 2-ME/5% DIEA in DMF (two 30-min treatments, 5 ml each).
Preparation and Handling of Peptides
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5. Remove N-terminal Boc groups by treating for 20 min with 10 ml of 50% TFA/DCM, neutralize the resin with 10 ml of 5% DIEA/DCM, and then wash the resin five times, each time with 10 ml DCM (procedure similar to “reactions 1 to 4” in Table 18.5.1). Place the reaction vessel in a desiccator, and dry the resin under a vacuum. Boc groups would be removed in HF cleavage. It is advisable, however, to remove these t-butyl protecting groups prior to HF cleavage to avoid generating long-lived, reactive t-butyl cations during HF cleavage.
Cleave the peptides 6. Place 300 to 500 mg of MAP resin in the reaction vessel of the hydrogen fluoride cleavage apparatus. Melt p-cresol and p-thiocresol by placing bottles in 50°C water bath. Add 0.75 ml of the p-cresol and 0.25 ml of the p-thiocresol to the reaction vessel. Once this mixture has cooled to room temperature and solidified, add a magnetic stir-bar and 6.5 ml DMS to the reaction vessel and mix by stirring for 2 min at low speed without splattering resin on the reaction vessel. 7. For low HF cleavage: Add liquid HF at −78°C to give a final volume of 10 ml 7.5/2.5/65/25 (v/v/v/v) p-cresol/p-thiocresol/DMS/HF. Equilibrate the mixture to 0°C by immersing the reaction vessel in an ice bath and stirring. Stir the mixture vigorously for 2 hr at 0°C, and remove HF and DMS under a vacuum. Proceed to step 8 for high HF cleavage, or work up reaction as in step 9. CAUTION: HF cleavage must be performed in a well ventilated chemical fume hood. Thick polyvinyl chloride gloves and a protective face mask must be used. The outside wall of the vessel should have a predetermined mark for 10 ml.
8. For high HF cleavage: Recharge the mixture with 14 ml liquid HF at −78°C to give a final total volume of 15 ml HF/p-cresol/p-thiocresol. Stir the reaction mixture 1 hr at 0°C. Remove HF under a vacuum using a water aspirator at 0°C. See precautions for working with HF (step 7 annotation)
9. Work up reaction: Wash the residue by adding 30 ml of cold 99:1 diethyl ether/2-ME to the reaction vessel to remove p-thiocresol and p-cresol. Filter the resin and the precipitated peptide with a fritted-glass funnel in a 50-ml round-bottom flask. Discard the ether wash (the peptide will be distributed partially in the reaction vessel and partially in the fritted-glass funnel). Dissolve the crude peptide on both the reaction vessel and the fritted-glass funnel with 100 ml of 10% acetic acid, collect the filtrates, and lyophilize the solution to dryness. Store at −70°C. ALTERNATE PROTOCOL
Synthesis and Application of Peptide Dendrimers As Protein Mimetics
DIRECT Fmoc SOLID-PHASE SYNTHESIS OF MAP SYSTEMS Fmoc synthetic chemistry is another widely used method for solid-phase peptide synthesis. The procedure for preparing MAP systems (Defoort et al., 1992) by the direct approach using Fmoc chemistry is presented in Figure 18.5.4 and Table 18.5.2. The basic procedure in Fmoc solid-phase peptide synthesis involves prewashing the resin, removing Fmoc protecting groups, and coupling the next Fmoc-amino acid residue. This cycle is repeated until peptide elongation is completed, and the peptide has been cleaved from the resin support. Materials Fmoc-Ala-Wang-resin (p-alkoxybenzyl alcohol resin preloaded with Fmoc-alanine at 0.3 to 0.5 mmol alanine/g resin; Bachem California) Fmoc synthesis reagents N,N′-dimethylformamide (DMF) 20% (v/v) piperidine in DMF (Pip/DMF) 0.5% (w/v) 1-hydroxybenzotriazole (HOBt) in DMF (HOBt/DMF)
18.5.8 Supplement 17
Current Protocols in Protein Science
Table 18.5.2
Schedule for Fmoc Solid-Phase Peptide Synthesis
Reaction Description 1 2 3 4 5 6a 7 8 9 10c
DMF wash 20% (v/v) Pip/DMF prewash 20% (v/v) Pip/DMF deprotection DCM wash 0.5% HOBt/DMF wash Fmoc-AA (3 eq) in DMF HOBt (3 eq) in DMF DCC (3 eq) in DCM DMF wash Ninhydrin test
Volume Repetitions and (ml/g resin) time 15 15 15
2 × 1 min 1 × 3 min 1 × 20 min
15 12 12 1 1 15 —
6 × 1 min 2 × 1 min 1 × 1 minb 1 × 1 minb 1 × 60 min 5 × 1 min 5 min
aCoupling procedure for all amino acids. bDo not drain until the completion of reaction 7. cIf ninhydrin test does not show completion of coupling reaction (>99.5%), repeat reactions 5 to 7.
Fmoc-Lys(Fmoc) (Bachem California; Peninsula Laboratories) Acetic anhydride Fmoc-amino acids (Fmoc-AA) with protected side-chain groups (Bachem California; Peninsula Laboratories) appropriate for synthesis of the desired peptide antigen, dissolved in DMT Cleavage solution (see recipe), prepare fresh and prechill to 0°C Anhydrous methyl t-butyl ether, cold Anhydrous diethyl ether 10% (v/v) acetic acid in water Automated peptide synthesizer (e.g., Perkin-Elmer Applied Biosystems #ABI430A) Manual reaction vessel (Pierce) Coarse- and fine-porosity fritted glass funnels Prepare the resin 1. Place 0.5 g of 0.3 to 0.5 mmol/g Fmoc-Ala-Wang-resin in a reaction vessel in an automated peptide synthesizer. Couple 0.055 mmol (0.32 mg) Fmoc-Lys(Fmoc) onto the resin by carrying out reactions 1 to 9 in Table 18.5.2, adding the Fmoc-Lys(Fmoc) in reaction 6 of the table. Omit the ninhydrin test step. Reactions 1 through 10 (Table 18.5.2) constitute one complete cycle; one cycle is required for each amino acid in the sequence. Some automated synthesizers may include the test to evaluate the coupling reaction in their cycle; if not, the ninhydrin test will have to be performed manually (Support Protocol 1, Sarin et al., 1981). The commercially available Fmoc-Ala-Wang-resin usually has a loading of 0.3 to 0.5 mmol Fmoc-Ala/g resin. This content, however, is too high for synthesis of AMP (see Background Information). To prepare a resin with 0.1 mmol lysine/g resin at the first level in MAP core, only 0.055 mmol of Fmoc-Lys (Fmoc) should be used. This incomplete coupling will result in the loading of ∼0.1 mmol lysine/g resin.
2. Add 15 ml DCM and 0.75 ml acetic anhydride to the reaction vessel and incubate 1 hr to terminate unreacted amines. Wash the resin five times with 10 ml DMF (1 min each) as in Table 18.5.2, reaction 9. Preparation and Handling of Peptides
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Supplement 17
As a result of the previous incomplete coupling step, an excess of free amino groups will be on the resin. This step will acylate (cap) free amino groups prior to coupling the next level of lysine.
Prepare the core matrix 3a. To prepare a MAP core matrix with four or eight branches: In reaction 6, couple 0.3 mmol (177 mg) Fmoc-Lys(Fmoc) to the resin from step 2 by following the reaction steps in Table 18.5.2. Proceed to step 3b (for core matrix with eight branches) or step 4 (for core matrix with four branches). 3b. To prepare a MAP core matrix with eight branches: In reaction 6, couple 0.6 mmol (354 mg) Fmoc-Lys(Fmoc) to the resin from step 3a by following the reaction steps in Table 18.5.2, adding the Fmoc-Lys(Fmoc) at reaction 6 in the table. Proceed to step 4. Preformed MAP core matrix can be purchased from Applied Biosystems or CalbiochemNovabiochem.
Synthesize the peptide on the MAP core matrix 4. Perform synthesis of the peptide on the MAP core matrix prepared in step 3a or b by carrying out the reactions in Table 18.5.2 (one cycle of 10 reactions per amino acid to be added) using, in reaction 6 of the table, 0.6 mmol of Fmoc-amino acids for a MAP core matrix with four branches or 1.2 mmol of Fmoc amino acids for a MAP core with eight branches. The most commonly recommended side-chain-protecting groups for Fmoc-amino acids are: Ser, Thr, and Tyr, t-butyl ether (tBu); Asp and Glu, t-butyl ester (OtBu); Asn, Gln, trityl (Trt) amide; Arg, 2,2,5,7,8-pentamethylchroman-6-sulfonyl (PMC) or 4-methoxy-2,3,6trimethylhenzenesulfonyl (Mtr); Lys, tert-butyloxycarbonyl (Boc); His, Nim-trityl (TrT) Trp, and Met, unprotected; Cys, trityl (Trt) thioether or acetamidomethyl (Acm). All are commercially available.
5. After completing the synthesis, perform reactions 1 to 4 in Table 18.5.2, to remove the Fmoc group on the amino group of N-termini (last coupling) with 20% pip/DMF. Wash the resin with 10 ml DMF (three times, 1 min each) and 10 ml DCM (three times, 1 min each). Cleave the peptides 6. Transfer peptide/resin mixture to a manual reaction vessel. Add prechilled cleavage solution to the peptide/resin at a ratio of 10 ml cleavage solution per 0.2 to 0.4 g peptide/resin. Stir mixture 1.5 to 2 hr at room temperature. Other cleavage solutions can be used. Reagent K [82.5:5:5:2.5:5 (v/v/v/v/v) TFA/thioanisole/phenol/1,2-ethanedithiol (EDT)/H2O] can be used for peptides that contain multiple Arg and Trp residues. For simple peptides, a cleavage solution of 5% H2O/95% (v/v) TFA can be used.
7. Filter the peptide/resin mixture through a coarse-porosity fritted glass funnel, and wash the resin three times, each time with 5 ml TFA. Concentrate the filtrate to a small volume (∼2 ml) under a vacuum. 8. Precipitate peptide with 30 to 50 ml cold anhydrous methyl t-butyl ether.
Synthesis and Application of Peptide Dendrimers As Protein Mimetics
9. Filter the peptide precipitate through a fine-porosity fritted glass funnel, and wash the peptide three times, each time with 10 ml anhydrous diethyl ether. Dry the crude peptide (in the glass funnel) under a vacuum. 10. Dissolve the crude peptide in 10% acetic acid and lyophilize to dryness. Store at −70°C.
18.5.10 Supplement 17
Current Protocols in Protein Science
INDIRECT Boc AND Fmoc SYNTHESIS OF MAP SYSTEMS USING THIOL CHEMISTRY
BASIC PROTOCOL 2
MAP systems have been successfully prepared by the direct approach, but they can also be synthesized by an indirect approach. In this method, the peptide antigen and MAP core matrix are prepared and purified separately and then linked together through functional groups on both elements to form a MAP system (Fig. 18.5.2 and Fig. 18.5.5; Lu et al., 1991). The indirect approach combines the advantages of solid-phase peptide synthesis with those of conjugation in a solution phase. An unprotected, purified peptide can be used in ligation. The orientation of peptides, particularly those derived from sequences at or near carboxyl termini, can be arranged as in native protein by linkage through the N-terminus to the core matrix. A variety of functional groups have been used for ligation to guarantee site-specific, chemically unambiguous MAP systems (also see Background Information). This protocol describes ligation through a sulfhydryl group of Cys in the peptide and a chloroacetyl group on the MAP core matrix. The same ligation chemistry is used to attach cyclic peptides to the MAP core matrix (see Basic Protocol 6). Materials Chloroacetic acid (Aldrich) 1.0 M N,N′-dicyclohexylcarbodiimide (DCC) in DCM (DCC/DCM) Sulfhydryl reducing solution (see recipe) Nitrogen source 1 M Tris⋅Cl 1 M NaOH 0.1 M NH4HCO3 or 10% acetic acid Additional reagents and equipment for solid-phase peptide synthesis using Boc (see Basic Protocol 1) or Fmoc chemistry (see Alternate Protocol), purification
Reactive groups Peptide
directiona
C N
N C
N
C
N
C
N
N
Ligation site
Electrophileb
O S-CH2 C-
S
Nucleophilec References O X-CH2 CX=Cl or Br
Lu et al., 1991
OO HC-C-d
Shao and Tam, 1995
O O S C C N H O O -C-CH2 ON=CH-C-
O -C
O -C-CH2 ONH2
OO HC-C-d
Shao and Tam, 1995
C
O O -C-NHN=CH-C-
O -C-NHNH2
OO HC-C-d
Shao and Tam, 1995
C
O -C
OO HC-C-d
Spetzler and Tam, 1995
O O N=N-CH2 -C- -C
SH NH2
NHNH2
Figure 18.5.5 Ligation methods for preparing MAP or cMAP by the indirect approach. Notes: (a) The direction in which the peptide is attached to the core matrix; (b) the functional group is placed at the end of the peptide; (c) the functional group is placed at the end of the branched Lys residues; (d) Generated for NalO4 oxidation of Ser of Thr.
Preparation and Handling of Peptides
18.5.11 Current Protocols in Protein Science
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of peptides via RP-HPLC (UNIT 11.6), dialysis (see Support Protocol 2), and high performance gel-filtration chromatography (see Support Protocol 3) Prepare peptide antigen by solid-phase peptide synthesis 1. Synthesize the peptide using either Boc or Fmoc chemistry as in Basic Protocol 1 or the Alternate Protocol. For the ligation of peptide to a chloroacetylated MAP core matrix, an additional Cys should be attached to either C-termini (for peptide antigens derived from internal or N-terminal regions) or N-termini (for peptide antigens derived from the C-terminal region) of the peptide.
2. Purify the peptide using the methods in UNIT 11.6 or in Support Protocols 2 and 3 of this unit. Prepare chloroacetylated MAP core matrix 3. Prepare a MAP core matrix with four or eight lysine branches (see Basic Protocol 1, step 2), or Alternate Protocol, step 3). 4. Remove the Nα-terminal protecting group (see Basic Protocol 1, step 5, or Alternate Protocol, step 5). 5. Couple chloroacetic acid (4 molar equivalents) to the free amino groups of core matrix using 4 molar equivalents of DCC/DCM as a common amino acid coupling. This is done as in reactions 4 to 12 of Table 18.5.1 for Boc chemistry or reactions 4 to 9 of Table 18.5.2 for Fmoc chemistry, except that chloroacetic acid is being used instead of a protected amino acid.
6. Cleave the MAP core matrix from resin (see Basic Protocol 1, steps 6 to 9, for Boc-amino acids—using high HF cleavage—or Alternate Protocol, steps 6 to 8, for Fmoc-amino acids—using 95% (v/v) TFA in water as the cleavage solution). 7. Purify MAP core matrix using the methods in UNIT 11.6 or in Support Protocols 2 and 3 in this unit. Conjugate peptide antigen with chloroacetylated MAP core matrix 8. Dissolve 28 µmol peptide (from step 2) in 5 ml sulfhydryl reducing solution. Keep this solution under nitrogen and adjust pH to 6.0 to 6.5 with 1 M Tris⋅Cl. Flush the solution with nitrogen for 30 min, then seal the solution (no air contact) for 2 hr. Before the peptide is conjugated to the chloroacetylated MAP core matrix, the disulfide bond of the terminal Cys should be reduced to a sulfhydryl group by treatment with tributylphosphine (Bu3P). Dithiothreitol (DTT) cannot be used for this purpose because DTT may react with the chloroacetylated core matrix. Note that this reaction is performed off-resin and in solution phase. On-resin conjugation can be performed by modifying the sequence of these reactions and the use of an aqueous-compatible resin support, e.g., PEG-modified resin, which is available commercially.
9. Add 4 µmol of the tetrabranched chloroacetylated MAP core matrix (from step 7) to the peptide solution (from step 8). Adjust the pH to 8.5 to 9.0 with 1 M NaOH. Stir the mixture 45 min at room temperature. Synthesis and Application of Peptide Dendrimers As Protein Mimetics
10. Dialyze the MAP system conjugate sequentially against 2 liters of 0.1 M NH4HCO3 or 10% acetic acid and then deionized water at room temperature, 8 hr each change. Lyophilize to dryness and store at −70°C.
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Current Protocols in Protein Science
This dialysis procedure is different from that described in Support Protocol 2 because the peptides have already been purified.
11. Remove any byproducts by purifying the MAP conjugate by high-performance gel-filtration chromatography (Support Protocol 3). INDIRECT Boc AND Fmoc SYNTHESIS OF MAP SYSTEMS USING CARBONYL CHEMISTRY
BASIC PROTOCOL 3
Another method for preparing the MAP system by the indirect approach is to exploit the selectivity of other weak bases, particularly in the condensation reaction with aldehydes (Fig. 18.5.5; Shao and Tam, 1995). Two types of weak bases can be used. The first type consists of conjugated amines whose basicities are lowered by neighboring electron-withdrawing groups such as hydroxylamine and substituted hydrazines. These react with aldehyde to form oxime from hydroxylamine and hydrazones from hydrazines. The second type contains the 1,2-disubstituted patterns such as 1,2-aminoethanethiol derivatives of N-terminal cysteine and 1,2-aminoethanol derivatives of N-terminal threonine. Proline-like rings such as thiazolidine or oxazolidine are formed from the reaction between an aldehyde and cysteine or threonine, respectively. An aldehyde moiety on a peptide or MAP core matrix can be obtained by NaIO4 oxidation of N-terminal Ser, Thr, or Cys under neutral conditions to give an α-oxoacyl group. The additional weak base available for conjugation is attached to the N-termini of the peptide so the orientation of the peptide is C to N. However, the weak base-aldehyde chemistry has also been used to ligate constrained peptides to a MAP core matrix in the preparation of cMAP (see Basic Protocol 5). Materials Boc-aminooxyacetic acid (see recipe) or Boc-succinic acid hydrazide (see recipe) Benzotriazolyl N-oxytrisdimethylaminophosphonium hexafluorophosphate (BOP) (Richelieu Biotechnologies) Diisopropylethylamine (DIEA; Aldrich) 0.01 M sodium phosphate buffer, pH 7 (APPENDIX 2E) Sodium periodate (Aldrich) Ethylene glycol (Aldrich) 0.4 M sodium acetate buffer; mix equal volumes 0.4 M sodium acetate and 0.4 M acetic acid Dimethyl sulfoxide (DMSO; Aldrich) Dimethyl formamide (DMF) 0.1 M NaOH or glacial acetic acid Argon source C8 RP-HPLC column (see UNIT 11.6) Additional reagents and equipment for solid-phase peptide synthesis using Boc chemistry (Basic Protocol 1) or Fmoc chemistry (Alternate Protocol), and purification of peptides by semipreparative RP-HPLC (UNIT 11.6) Prepare the peptide by solid-phase peptide synthesis 1. Synthesize the peptide using Boc or Fmoc chemistry as in Basic Protocol 1 or the Alternate Protocol. 2. Remove the N-terminal protecting group (see Basic Protocol 1, step 5, or Alternate Protocol, step 5). 3. Couple (on-resin) the Cys residue (for thiazolidine ligation; 4 molar equivalents) to the free amino group of the peptide using 4 molar equivalents of DCC/DCM as a
Preparation and Handling of Peptides
18.5.13 Current Protocols in Protein Science
Supplement 17
common amino acid coupling (see Table 18.5.1, steps 4 to 12, for Boc chemistry, or Table 18.5.2, steps 4 to 9, for Fmoc chemistry). Couple Boc-aminooxyacetic acid (for oxime ligation) or Boc-succinic acid hydrazide (for hydrazone ligation; 4 molar equivalents) to the amino terminus of the peptide using 4 molar equivalents of 1:1 (molar:molar) BOP/DIEA, dissolved in DMF, as coupling reagent in reaction 9 of Table 18.5.1 (in place of the DCC). For thiazolidine ring formation, an additional Cys residue is attached to the N-terminus of the peptide. For coupling through an oxime or hydrazone bond, the weak base is also placed at the N-terminus of the peptide.
Prepare the glyoxyl-MAP core matrix 4. Prepare the tetravalent MAP core matrix (Lys2-Lys-Ala) by the stepwise solid-phase synthesis method using either Boc chemistry (see Basic Protocol 1 and Table 18.5.1) or Fmoc chemistry (see Alternate Protocol and Table 18.5.2). Normal resin loading, ranging from 0.3 to 0.9 mmol/g, is recommended since only four coupling steps are required.
5. Remove the Boc group with 50% TFA/DCM (see Basic Protocol 1) or the Fmoc group with 20% pip/DMF (see Alternate Protocol). 6. Couple the fully protected Ser residue to the MAP core matrix using DCC and HOBt as described in step 3. 7. Cleave the Ser4-MAP core matrix from the resin by the high HF procedure, HF/pcresol (9:1, v/v), as described for Boc chemistry (see Basic Protocol 1, steps 6 to 9) or 95% (v/v) TFA in water as described for Fmoc chemistry (see Alternate Protocol, steps 6 to 8), and lyophilize the intermediate without purification. 8. Dissolve 5.86 µmol (5 mg) Ser4-MAP core matrix (from step 7) in 2 ml 0.01 M sodium phosphate buffer, pH 7, and add 46.9 µmol (10 mg) sodium periodate in a 2-ml microcentrifuge tube. 9. Stir the mixture for 5 min at room temperature, then quench the reaction by adding 93.8 µmol ethylene glycol (5.3 µl). 10. Isolate the product by semipreparative RP-HPLC as described in UNIT 11.6. Conjugate peptide with glyoxyl-MAP core matrix 11. Prepare a 5 mM peptide stock solution in water using the peptide from step 3 and a 5 mM glyoxy-MAP stock solution in water using the purified MAP from step 10. 12. Add 400 µl of 0.4 M sodium acetate buffer and 40 µl of the 5 mM glyoxy-MAP stock solution (0.2 µmol glyoxy-MAP) to 400 µl of the 5 mM peptide stock solution (2 µmol peptide). Then, add 800 µl DMSO in the case of oxime and hydrazone ligation and 800 µl DMF in the case of thiazolidine ligation. 13. Adjust pH of the solution to 5.7 (for oxime and hydrazone ligation) or 4.5 (for thiazolidine ligation) using 0.1 M NaOH or glacial acetic acid. 14. Stir the mixture under argon for 8 hr at room temperature, and monitor the completion of the ligation on a C8 RP-HPLC column (UNIT 11.6). Synthesis and Application of Peptide Dendrimers As Protein Mimetics
15. Purify the MAP conjugate by semipreparative HPLC (UNIT 11.6). Lyophilize to dryness and store at −70°C.
18.5.14 Supplement 17
Current Protocols in Protein Science
NINHYDRIN TEST The ninhydrin assay is used to check for completeness of the coupling reaction (Sarin et al., 1981). It is performed at the end of each synthesis cycle, usually as described here.
SUPPORT PROTOCOL 1
Materials Ninhydrin test reagents: Solution A/B mixture and Solution C (see recipe) 60% (v/v) ethanol in H2O 0.5 M tetraethylammonium chloride in dichloromethane (DCM) 10 × 75–mm test tubes 100°C heating block Pasteur pipet containing glass wool plug 1. At the end of each synthesis cycle (see Table 18.5.1 and Table 18.5.2), remove ∼10 mg wet resin from the reaction vessel. Dry the resin 1 hr under vacuum in a desiccator. 2. Weigh a 3- to 5-mg sample of dry resin into a 10 × 75–mm test tube. Add 100 µl of the solution A/B mixture and 25 µl solution C. To another tube add only the ninhydrin test reagents without resin. Mix. 3. Heat both tubes 10 min in a heating block at 100°C. Cool tubes in cold water. 4. Add 1 ml of 60% ethanol to the tubes. Mix thoroughly and filter through a Pasteur pipet containing a glass wool plug. 5. Rinse the resin twice with 0.20 ml of 0.5 M tetraethylammonium chloride in DCM. Combine the filtrates. 6. Dilute the combined filtrate to 2 ml with 60% ethanol. 7. Measure the A570 of the sample filtrate against the reagent blank. For routine monitoring, an effective extinction coefficient of 1.5 × 104 may be used.
PURIFICATION OF MAP SYSTEM BY DIALYSIS The chemistry of solid-phase peptide synthesis has been significantly improved since it was first introduced in 1963. Now synthesis of peptides of 100 or more residues with satisfactory yield and purity is possible. In general, crude synthetic MAP systems (after the cleavage step) are pure enough for many purposes (e.g., for production of antisera). When highly purified MAP systems are required (e.g., for vaccines for humans), the crude synthetic product must be purified to near homogeneity. MAP systems can be purified by dialysis, gel-filtration chromatography, RP-HPLC (UNIT 11.6), and high-performance gel-filtration chromatography (Support Protocol 3). This protocol details the special procedure required for the sequential dialysis of MAP systems.
SUPPORT PROTOCOL 2
Materials Crude MAP system (see Basic Protocol 1, 2, or 3, or Alternate Protocol) 0.1 M NH4HCO3/(NH4)2CO3 (pH 8.0) in 8 M and 2 M urea 1 M acetic acid Dialysis tubing (e.g., Spectra/Por 6, MWCO 1000, Spectrum) 1. Dissolve crude MAP system in 100 ml of 0.1 M NH4HCO3/(NH4)2CO3 (pH 8.0) in 8 M urea. 2. Load the peptide solution into dialysis tubing.
Preparation and Handling of Peptides
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3. Dialyze the peptide solution sequentially against 2 liters of each of the following solutions, for 8 hr each, at room temperature: 0.1 M NH4HCO3/(NH4)2CO3 (pH 8.0) in 8 M urea 0.1 M NH4HCO3/(NH4)2CO3 (pH 8.0) in 2 M urea H2O 1 M acetic acid. 4. Remove peptide solution from dialysis tubing. Lyophilize to dryness. Store at −70°C. SUPPORT PROTOCOL 3
PURIFICATION OF MAP USING HIGH-PERFORMANCE GEL-FILTRATION CHROMATOGRAPHY In this protocol, a crude MAP system is dissolved in potassium phosphate and purified over a specialized gel-filtration column by HPLC. Materials 10 mg crude MAP system (Basic Protocol 1, 2, or 3, or Alternate Protocol) 0.1 M potassium phosphate, pH 7.0 (APPENDIX 2E) Column for high-performance gel-filtration chromatography (prepacked Bio-Sil TSK 250, 300-mm length × 7.5-mm i.d.; mol. wt. range for separation: 10 to 300 kDa; Bio-Rad #125-0062) Guard column (75-mm length × 7.5-mm i.d.; Bio-Rad #125-0061) Additional reagents and equipment for HPLC (UNIT 11.6) 1. Dissolve 10 mg crude MAP system in 1 ml of 0.1 M potassium phosphate, pH 7.0. 2. Set up HPLC system with the Bio-Sil TSK column linked to the guard column. Set UV detector at 225 nm. Equilibrate the gel-filtration column with 0.1 M potassium phosphate, pH 7.0, at a flow rate of 0.5 ml/min. 3. Inject 200 µl of the crude MAP solution (from step 1). Elute peptide at a flow rate of 0.5 ml/min. Collect the purified MAP product. 4. Repeat injection (step 3) four times to purify all of the material. Pool the purified MAP product. 5. Lyophilize MAP fraction to dryness. Store MAP at −70°C.
BASIC PROTOCOL 4
Synthesis and Application of Peptide Dendrimers As Protein Mimetics
DIRECT SYNTHESIS OF END-TO-SIDE CHAIN cMAP This approach is suitable for preparing cMAP with multiple end-to-side chain cyclic peptides. It combines solid-phase (on-resin synthesis) with solution-phase synthesis (off-resin cyclization; Spetzler and Tam, 1996). First, stepwise solid phase synthesis is used to assemble the peptides linked to the MAP core matrix on resin. The cMAP precursor is then cleaved from the resin, and its protecting groups removed. The peptide antigens are cyclized in cMAP in aqueous solution. The chosen sequences were peptide antigens derived from the V3 loop of the surface protein gp120 of human immunodeficiency virus-1 (HIV-1), MN strain. Two peptides with the sequence GPGRAFYTTKNIIG and KRKRIHIGPGRAFYTTKNIIG were used. The essential motif GPGR was placed in the middle of the antigen sequence. The cMAP precursor contains a tripartite design (Fig. 18.5.6). The first two parts are constant regions. Part 1 at the C-terminus contains a short peptide which can be used for identification, delivery to give an MHC class I molecule or a T-helper epitope, followed by a tetralysyl core matrix. Part 2 contains a specific chemical cleavage site AspPro that gives a monomeric cyclic peptide to verify the yield of the intrachain cyclization. This part may not be necessary if verification is
18.5.16 Supplement 17
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Self-assembly of multiple cyclic antigen peptide
-Lys -Lys
Lys-Ala-
stepwise synthesis
Ser(tBu) Cys(StBu)
Lys-(Asp-Pro)-MPC-
MPC TFA
Ser Lys-(Asp-Pro)-MPC
Cys(StBu) H
O
O NaIO4
Lys-(Asp-Pro)-MPC
Cys(StBu) H
R3P
O
SH NH2-Cys S N H
pH 5.6
O
Lys-(Asp-Pro)-MPC
O Lys-(Asp-Pro)-MPC
= HOCH2
OCH2
= peptide
R = CH2 CH2CO2H
Figure 18.5.6 The scheme for preparing cMAP using the direct method.
not needed. The third part is the variable region and contains a peptide framed by a lysine at the COOH-terminus (ε-Lys) and a Cys at the amino terminus to give the end-to-side chain cyclic structure. The two mutually reactive functions are the α-Cys and a masked aldehyde attached to the side chain of the C-terminal lysine. Conversion to the aldehyde under mild aqueous conditions will permit an end-to-side chain cyclization via thiazolidine (Fig. 18.5.6). The protecting strategy for the two reactive moieties, Cys and aldehyde, is designed to be compatible with Fmoc chemistry (Alternate Protocol and Table 18.5.2) and the removal of these moieties under aqueous conditions. Sulfenyl tertbutyl is chosen for Cys as Cys(StBu) because it can be removed under aqueous conditions by trialkylphosphine and is stable to oxidation by periodate. The masked form of the aldehyde is the 1,2-aminoalcohol of a Ser residue anchored on the lysyl side chain. It can be converted to an aldehyde by periodate oxidation. The use of trialkylphosphine in the scheme has two advantages. First, it inhibits intermolecular disulfide formation as a side reaction, and second, it allows a one-pot reaction for unmasking the thiol protecting group and cyclization. Materials Fmoc-Lys[methyltrityl(Mtt)] (Calbiochem-Novabiochem) Fmoc-Lys(Fmoc) (Bachem California) Fmoc-Cys(StBu) (Calbiochem-Novabiochem) 1% (v/v) TFA/5% (v/v) triisopropylsilane (TIS; Aldrich) in DCM Cleavage solution: 92.5/2.5/2.5/2.5 (v/v) trifluoroacetic acid (TFA)/triisopropyl silane (TIS) (Aldrich)/thioanisole (Aldrich)/H2O, prepared fresh 0.01 M sodium phosphate buffer, pH 6.8 (APPENDIX 2E)
Preparation and Handling of Peptides
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Sodium periodate (Aldrich) Tris(2-carboxyethyl)phosphine hydrochloride (TCEP; Calbiochem-Novabiochem) 10 mM sodium phosphate, pH 6.8 (APPENDIX 2E) 10 mM sodium acetate buffer, pH 4.2 (APPENDIX 2E) 70% (v/v) formic acid in H2O Additional reagents and equipment for peptide synthesis (Alternate Protocol and Table 18.5.2), semipreparative HPLC (UNIT 11.6), and assay of free sulfhydryls with Ellman’s reagent (UNIT 18.3) Perform on-resin synthesis of the cMAP precursor 1. Synthesize the cMAP precursor starting from Fmoc-Ala-Wang resin (0.5 g, 0.1 mmol/g) manually or via an automated peptide synthesizer using the synthesis reagents and directions outlined in Table 18.5.2 and described in Alternate Protocol. The order of assembly of the precursor is: the MAP core matrix, the cleavage site Asp-Pro, Fmoc-Lys(Mtt), peptide sequence, and Fmoc-Cys(S-tBu). See Figure 18.5.6.
2. Remove (on-resin) the side-chain-protecting 4-methyltrityl (Mtt) group of Lys using 1% TFA/5% TIS in DCM using the procedure described in step 6 of the Alternate Protocol. Discard the TFA/TIS/DCM solution and neutralize the TFA-amine salt on-resin by performing reactions 4 to 7 of Table 18.5.1. 3. Couple Fmoc-Ser(tBu) to the lysyl side chain by DCC and HOBt procedure as a common amino acid coupling as in reactions 6 to 9 of Table 18.5.2. 4. Deprotect and cleave the precursor of cMAP from the resin by 92.5/2.5/2.5/2.5 (v/v) TFA/TIS/thioanisole/H2O as outlined in the Alternate Protocol, steps 5 to 10. Lyophilize the peptide. Avoid thiol scavengers due to the presence of Cys(S-tBu).
Perform off-resin cyclization to form cMAP 5. Dissolve 0.44 µmol cMAP precursor (from step 4) in 2 ml 0.01 M sodium phosphate buffer, pH 6.8, and add 3.52 µmol (0.75 mg) sodium periodate in 27 µl water. 6. Stir the solution for 2 min, then purify the peptide by semipreparative HPLC as described in UNIT 11.6. Lyophilize the oxidized cMAP precursor. 7. Add 17.6 µmol (1.5 mg) TCEP in 47 µl of 10 mM sodium acetate buffer, pH 4.2, to the solution of oxidized cMAP in 1.5 ml of 10 mM sodium acetate buffer, pH 4.2. 8. Stir the mixture at 22°C for 48 hr, and monitor the progress of the cyclization by assay of free sulfhydryls with Ellman’s reagent (UNIT 18.3) until all the thiols are consumed. 9. Purify cMAP by semipreparative HPLC as described in UNIT 11.6. 10. For determining the intrachain yield, add 0.5 ml 70% formic acid to 0.05 µmol of the lyophilized cMAP in a glass test tube. Incubate the solution either at 37°C for 48 hr or 60°C for 24 hr. Add 0.5 ml water and lyophilize the sample. This procedure will cleave the cyclic peptide at the AspPro site as a monomer.
11. Separate the monomeric sample by RP-HPLC (UNIT 11.6).
Synthesis and Application of Peptide Dendrimers As Protein Mimetics
The intrachain yield is calculated from the amino acid hydrolysis of the cMAP and the cyclic monomer.
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Current Protocols in Protein Science
INDIRECT Fmoc SYNTHESIS OF SIDE CHAIN-TO-SIDE CHAIN cMAP USING CARBONYL CHEMISTRY
BASIC PROTOCOL 5
In this approach, a side chain-to-side chain cyclized peptide is attached to the core matrix through a lysyl side chain (Pallin and Tam, 1996). The sequence used is a peptide antigen chosen from the neutralizing determinant of the V3 loop of gpl20. The target amino acid sequence is IGPGP. The precursors, which contain three Lys residues, are introduced as Fmoc-Lys(Mtt) and Boc-Lys(Fmoc), respectively, for attaching three functional groups (hydroxylamine, Ser, and Cys(StBu) through the side chains (Fig. 18.5.7). An O-alkylhydroxylamine group is attached to the side chain of the first Lys before the peptide sequence. The second Lys is introduced after the peptide sequence, and a Ser moiety is coupled to the side chain as a masked aldehyde, which is oxidized to an aldehyde by NaIO4. A cyclic peptide is formed by using the intramolecular oxime formation from the reaction between O-alkylhydroxylamine and the aldehyde. A Cys(StBu) residue on the
A
ONHBoc stepwise synthesis
Ser(tBu)
O=C
Cys(StBu) ONH2 O=C TFA
-Lys-Ala-
Boc-Lys-Ala-Lys
1 Ser
Lys-Ala-Lys
Lys-Ala-OH
Cys(StBu) 2 ONH2 O=C NaIO4
Lys-Ala-Lys
H
O
O Lys-Ala-OH
Cys(StBu)
3 Lys-Lla-Lys O=C Cys ON=CHC=O
pH 5.5 TCEP
Lys-Ala-OH
4
B SH NH2
HCOCO HCOCO-Lys HCOCO-Lys
Lys-Ala-OH
4
Y Y-Lys Y-Lys
HCOCO
= HOCH2
Lys-Ala-OH
Y
OCH2
Y=
S HN O
SH NH2 4
Lys-Ala-Lys
=
O=C Cys ON=CHC=O Lys-Ala-OH
Figure 18.5.7 The scheme for preparing cMAP utilizing carbonyl chemistry and using the indirect method. (A) Oxime approach for preparing unprotected cyclic peptide antigen. (B) Attachment to MAP via thiazolidine linkage.
Preparation and Handling of Peptides
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side chain of the third Lys moiety is used to ligate the cyclic peptides to the aldehyde moieties of the MAP core. Materials Fmoc-Lys[methyltrityl(Mtt)] (Calbiochem-Novabiochem) 1% (v/v) TFA/5% (v/v) triisopropylsilane (TIS; Aldrich) in DCM 20% piperidine in DMF (pip/DMF) Boc-Ser(tBu; Bachem California) Boc-(aminooxy)acetic acid (Bachem California) Boc-Lys(Fmoc; Bachem California) 0.01 M sodium acetate buffer, pH 7 (APPENDIX 2E) Sodium periodate (NaIO4;Aldrich) Ethylene glycol 0.1 M NaOH Tris(2-carboxyethyl)phosphine hydrochloride (TCEP; Calbiochem-Novabiochem) 0.05 M sodium acetate buffer, pH 6 (APPENDIX 2E) Tetravalent MAP core containing aldehyde groups (see Basic Protocol 3) Acetic acid Additional reagents and equipment for peptide synthesis (see Alternate Protocol and Table 18.5.2), semipreparative and preparative HPLC (UNIT 11.6), and mass spectrometry (Chapter 16), Perform on-resin synthesis of the cyclic precursor 1. Synthesize the linear unprotected peptide by the stepwise solid phase strategy using the Fmoc chemistry protocol (see Alternate Protocol and Table 18.5.2) and couple the first Lys residue as Fmoc-Lys(Mtt). 2. Remove the Mtt protecting group using 1% TFA/5% TIS in DCM and couple Boc-Ser(tBu) to the side chain of Lys as described in step 2 of Basic Protocol 4. 3. Synthesize the peptide sequence using the Fmoc strategy as described in the Alternate Protocol. 4. Incorporate the second Lys residue as the Fmoc-Lys(Mtt) derivative. 5. Couple Boc-(aminooxy)acetic acid to its side chain. 6. Couple the third Lys as a Boc-Lys(Fmoc) derivative. 7. Remove the Fmoc group by 20% pip/DMF (see Alternate Protocol). 8. Couple Fmoc-Cys(StBu) to the side chain of lysine. 9. Cleave and purify the peptide precursor as described for Fmoc chemistry (Alternate Protocol). Perform off-resin cyclization to form side chain-to-side chain cyclic peptides via an oxime linkage 10. Dissolve the peptide in 0.01 M sodium acetate buffer, pH 7, to give a final concentration of 0.8 M.
Synthesis and Application of Peptide Dendrimers As Protein Mimetics
11. Oxidize the 1,2-amino alcohol moiety to the glyoxyl derivative by addition of a 2 M excess of NaIO4 for 2 min at room temperature. Quench reaction with a >20-fold excess of ethylene glycol. 12. Purify the peptide by semipreparative HPLC (UNIT 11.6) to remove the formaldehyde byproduct.
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13. Collect the product from semipreparative HPLC and adjust pH to 5.5 with 0.1 M NaOH, to effect the oxime cyclization. 14. Monitor the progress of the intramolecular cyclization by RP-HPLC (UNIT 11.6). The reaction is usually completed within 12 hr.
15. Characterize the cyclic peptide by mass spectrometry (Chapter 16). Attach cyclic peptides to the MAP core matrix via a thiazoline linkage 16. Add 1.5 µmol (0.13 mg) TCEP to a solution of 0.3 µmol cyclic peptide in 0.5 ml of 0.05 M sodium acetate buffer, pH 6. Stir the mixture for 1 hr at room temperature. 17. Add 0.036 µmol tetravalent MAP core containing aldehyde groups (Basic Protocol 3) to the above mixture and adjust pH to 5 with acetic acid. 18. Stir the solution for 36 hr at 18°C, and purify the product by preparative HPLC as described in UNIT 11.6. INDIRECT PREPARATION OF END-TO-END cMAP USING THIOL CHEMISTRY
BASIC PROTOCOL 6
In this protocol, the cyclic peptide is attached through its side chain to the core matrix (Zhang and Tam, 1997). The selected sequences were YGGFL and AVEIQFMHNLGK. First, the precursor of the unprotected cyclic peptide and the functionalized core matrix are synthesized by SPPS (Fig. 18.5.8). The cyclic peptide is generated by an intramolecular transthioesterification reaction in which the cyclization occurs under equilibration of ring chain tautomerization in aqueous solution. Sequentially, the end-to-end cyclic peptide is formed via an S- to N-acyl transfer reaction, and a Cys residue is generated. In this method, racemization is minimized, and the cyclization can occur in high concentration. The generated free thiol group of the Cys residue is then available for ligation. Preparation of the (chloroacetyl)lysyl MAP core matrix is described in Basic Protocol 2. In the second step, the thioether bond is formed between the cyclic peptide and the core matrix at slightly basic pH. Materials 3-thiopropionic acid (Aldrich) MBHA (p-methyl benzhydrylamine) or Gly-PAM resin (Calbiochem-Novabiochem) 1 mM dithiothreitol (DTT; Aldrich) in distilled water 20 mM Na2HPO4/10mM citric acid buffer, pH 7.5 Tris(2-carboxyethyl)phosphine, hydrochloride (TCEP; Calbiochem-Novabiochem) 0.2 M sodium phosphate buffer, pH 7.4 (APPENDIX 2E)/10 M ethylenediaminetetraacetic acid (EDTA; Aldrich) Argon source Tetravalent chloroacetyl(lysinyl) core matrix (see Basic Protocol 2) Dimethylformamide (DMF) Additional reagents and equipment for peptide synthesis (see Basic Protocol 1), preparative RP-HPLC (UNIT 11.6), and mass spectrometry (Chapter 16) Perform on-resin synthesis of the cyclic precursor 1. Couple 3-thiopropionic acid to 0.5 g MBHA or Gly-PAM resin at 1.1 mmol/g by DCC/DCM in a reaction vessel (see Basic Protocol 1). Boc-Gly-PAM resin must be deprotected to give a free α-amine. The idea of this step is to form a reversible linker containing a thioester at the C-terminus after HF cleavage. For
Preparation and Handling of Peptides
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A stepwise synthesis 4-MeBzl-S Boc-NH
O HF
HS H2N
1
O NH2
SR pH 7.2 O
S
O 3
2
O
O
SH NH O 4
B SH
ClCH2 CO ClCH2 CO-Lys Lys-Ala-OH ClCH2 CO-Lys ClCH2 CO
SCH2 CO SCH2 CO-Lys
4 pH 8.5
SCH2 CO-Lys SCH2 CO
= HS-(CH2)2 CO-MBHA resin
Lys-Ala-OH
5
O
SH
R = (CH2)2 CONH2
SH
= 4
NH O
Figure 18.5.8 The scheme for preparing cMAP utilizing thiol chemistry and using the indirect method. (A) Thioester approach for preparing unprotected cyclic peptide antigen. (B) Attachment to MAP via thioalkylation.
an alternative method to prepare this resin, see recipe for thioester resin (HS-CH2CH2COMBHA-resin 1) for C-terminal thioester.
2. Add 1 mM DTT to the resin for 1 hr. Disulfide bonds may have been formed during the acylation reaction.
3. Synthesize the peptide sequence by stepwise solid phase synthesis (also see Fig. 18.5.7) either with an automated peptide synthesizer or manually using the synthesis reagents and directions outlined in Table 18.5.1 and Basic Protocol 1. Avoid the use of base-mediated coupling reagents such as benzotriazol-1-yl-oxy-tris(dimethylamino)phosphonium hexafluorophosphate (BOB), 2-(IH-benzotriazol-1-yl)-1, 1, 3,3-tetramethyluronium tetrafluorohorate (TBTU), and 2-(IH-benzotriazol-1-yl)1,1, 3,3-tetramethyluronium hexafluorophosphate (HBTU), since the thioester is not stable to base. Synthesis and Application of Peptide Dendrimers As Protein Mimetics
4. Cleave the peptide thioester by the HF protocol as described in Basic Protocol 1. Purify the product by preparative RP-HPLC (UNIT 11.6). Lyophilize and characterize the product by mass spectrometry (Chapter 16).
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If the His residue is present in the sequence, use Boc-His(Tos) instead of Boc-His(DNP) which requires thiolysis to release the DNP protecting group.
Perform off-resin cyclization to form an end-to-end cyclic peptide containing an XCys ligation site 5. Dissolve 3.5 µmol linear peptide thioester (from step 4) in a solution of 2 ml of 20 mM Na2HPO4/10mM citric acid buffer, pH 7.5, containing 3.5 µmol (1.04 mg) TCEP. 6. Stir the solution for 6 hr, and monitor the progress of the intramolecular cyclization by RP-HPLC (UNIT 11.6). 7. Purify the cyclic peptide by preparative RP-HPLC as described in UNIT 11.6. Characterize and lyophilize the product. Attach cyclic peptides to the MAP core via a thioether linkage 8. Dissolve 28 µmol cyclic peptide (from step 7) in 1.25 ml of 0.2 M sodium phosphate buffer containing 10 µM EDTA, pH 7.4. 9. Purge the solution with argon for 10 min. 10. Add 1.4 µmol (1.1 mg) tetravalent (chloroacetyl)lysinyl core matrix (prepared as described in Basic Protocol 2) predissolved in 0.75 ml DMF. 11. Stir the mixture for 24 hr at room temperature, then purify the cMAP by preparative RP-HPLC as described in UNIT 11.6. 12. Characterize cMAP by mass spectrometry (Chapter 16). PREPARING END-TO-END CYCLIC PEPTIDES VIA AN X-CYS LIGATION SITE
SUPPORT PROTOCOL 4
In this protocol, an unprotected cyclic peptide containing an X-Cys ligation site is prepared (Liu and Tam, 1997). The sequence is a peptide antigen derived from the V3 loop of the gp120 of IIIB. The peptide precursor is synthesized on a thioester benzhydryl resin which, after HF cleavage, generates Cα-thiocarboxylic acid. A Cys residue activated as Cys(Npys) is incorporated at the amino-terminus. First, the end-to-end peptide is cyclized from formation of a mixed acyl disulfide which undergoes an intramolecular S,N-acyl transfer reaction. The generated hydrodisulfide (S-SH) can be reduced by thiolysis to give the native Cys-residue. Materials Thiobenzhydryl resin for C-terminal thioacid (see recipe) Boc-Cys(Npys) (Bachem California) 25% acetonitrile/0.05% TFA in H2O, pH 2 Sodium acetate, solid Dithiothreitol (DTT; Aldrich) Additional reagents and equipment for peptide synthesis (see Basic Protocol 1 and Table 18.5.1) and RP-HPLC (UNIT 11.6) 1. Perform on-resin synthesis of the cyclic precursor containing an X-Cys ligation site. Synthesize the precursor of the cyclic peptide on a thiobenzhydryl resin using the Boc-chemistry protocol as described in Basic Protocol 1 and outlined in Table 18.5.1. Cap the amino-terminal with Boc-Cys(Npys), and cleave and deprotect the peptide by HF. The resulting precursor is a peptide with a Cys(Npys) residue at the N-terminal and a thiocarboxylic acid at the C-terminal. The other side chains are unprotected.
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2. Initiate the cyclization after HF cleavage (see Basic Protocol 1) in 25% acetonitrile in H2O containing 0.05% TFA, pH 2, giving a final concentration of 1.75 µM with respect to the peptide solution. Monitor the progress of the cyclization by RP-HPLC (UNIT 11.6). During the cyclization, a sulfur-sulfur exchange occurs first and a yellow color appears due to the release of Npys-H.
3. Adjust the solution to pH 6 by using solid sodium acetate to complete the cyclization reaction. 4. After 10 min, add 1 eq DTT to the solution to reduce the cysteinyl hydrodisulfide to cysteine, and monitor the reaction by RP-HPLC (UNIT 11.6). 5. Purify the cyclic peptide by preparative RP-HPLC as described in UNIT 11.6. SUPPORT PROTOCOL 5
PREPARING END-TO-END CYCLIC PEPTIDES VIA AN X-THIAPROLINE LIGATION SITE In this example, the unprotected peptide precursor is synthesized on an Fmoc-amino acid-O-CH2-cyclic acetal resin (Botti et al., 1996). The peptide sequence is as follows: GRAFVTIGKIG derived from the V3 loop of gpl20, HIV-1. During the TFA cleavage, a glyceric ester is generated at the C-terminal. The diol of the glyceric ester is converted to glycoaldehyde by oxidation with NaIO4. The precursor contains a Cys(StBu) residue at the N-terminus while all the other side chains are unprotected. Two steps are involved to form thiaproline. First, the thiazolidine ring is generated from the reaction between an aldehyde moiety and the 1,2-aminothiol of the N-terminal Cys. Then, an O- to N-acyl transfer reaction occurs through a tricyclic ring contraction to give a peptide bond. The thiazolidine ring formation is mediated after NaIO4 oxidation and removal of the StBu protecting group, and the intramolecular O- to N-acyl transfer reaction occurs upon readjusting the pH to 5.9. Materials Fmoc-Gly-O-CH2-cyclic-acetal resin (see recipe for acetal resin) Cleavage solution: 91:3:3:3 (v/v) TFA/H2O/thioanisole/anisole NaIO4 0.01 M sodium phosphate buffer, pH 5.5, 5.7 and 6 (APPENDIX 2E) Tributyl phosphine (Bu3P) Isopropyl alcohol Additional reagents and equipment for peptide synthesis (see Alternate Protocol 2 and Table 18.5.2) and RP-HPLC (UNIT 11.6) 1. Synthesize end-to-end cyclic peptides via a thiaproline ligation site. Assemble the peptide sequence on Fmoc-Gly-O-CH2-cyclic-acetal resin by Fmoc chemistry using stepwise phase synthesis as described in the Alternate Protocol and Table 18.5.2. Cap the amino-terminal with Boc-Cys(StBu). 2. Cleave and deprotect the precursor as in the Alternate Protocol, steps 6 to 10, using a cleavage solution containing 91:3:3:3 TFA/H2O/thioanisole/anisole and allowing the reaction procedure to proceed for 2.5 hr. Lyophilize the product (Alternate Protocol, step 10).
Synthesis and Application of Peptide Dendrimers As Protein Mimetics
3. Add 30 µmol NaIO4 in 0.3 ml 0.01 M sodium phosphate buffer, pH 5.7, to 7 µmol peptide glyceric ester also dissolved in 1 ml 0.01 M sodium phosphate buffer, pH 5.7. Stir the solution for 30 min, and purify the product by semipreparative RP-HPLC (UNIT 11.6).
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4. Dissolve 1.18 µmol peptide-glycoaldehyde (from step 3) in 0.2 ml 0.01 M sodium phosphate buffer, pH 5.5, and add 10.9 µmol tributyl phosphine (Bu3P) in 30 µl isopropyl alcohol (1:10 v/v dilution of Bu3P in isopropyl alcohol). Stir and monitor the progress of the cyclization by analytical RP-HPLC (UNIT 11.6). Purify and lyophilize the product. The starting product forms two products corresponding to the cis and trans isomers of the cyclic peptide.
5. Redissolve the purified peptide in 0.2 ml 0.01 M sodium phosphate buffer, pH 6, and monitor the amide formation by analytical RP-HPLC (the O- to N-acyl transfer can take up to several days). If the reaction is slow, warm up the solution to 40°C. Purify the product and analyze by HPLC. PREPARING END-TO-SIDE CHAIN CYCLIC PEPTIDES VIA A THIAZOLIDINE OR OXIME LINKAGE
SUPPORT PROTOCOL 6
In these two examples (Pallin and Tam, 1995; Botti et al., 1996), the precursor contains a Lys residue which is introduced as an Fmoc-Lys(Mtt) for attaching Ser through its side chain. The sequences are peptide antigens ranging from 5 to 26 amino acids in length and are derived from the V3 loop of gp120, HIV-1. The 1,2-amino alcohol function of Ser acts as a masked aldehyde and can be converted to glyoxyaldehyde after treatment with NaIO4. A Cys(StBu) moiety is incorporated at the N-terminus to generate a thiazolidine ring while Boc-(aminooxy)acetic acid is incorporated to form an oxime bond. All the other side chains are unprotected. The thiazolidine ring is formed by first oxidizing the amino-terminal Ser to an aldehyde moiety and then removing the StBu protection group by TCEP. In generating cyclic oxime peptides, cyclization occurs after oxidation to the aldehyde. Materials Fmoc-Lys(Mtt) 1% (v/v) TFA/5% (v/v) triisopropylsilane (TIS; Aldrich) in DCM Boc-Ser(tBu) Cleavage solution: 92.5/2.5/2.5/2.5 (v/v) TFA/TIS/thioanisol/H2O Sodium periodate 0.01 M sodium phosphate buffer, pH 6.8 (APPENDIX 2E) 0.01 M sodium acetate buffer, pH 4.2 (APPENDIX 2E) Tris(2-carboxyethyl)phosphine, hydrochloride (TCEP; Calbiochem-Novabiochem) Additional reagents and equipment for peptide synthesis (see Alternate Protocol and Table 18.5.2), RP-HPLC (UNIT 11.6), and mass spectrometry (Chapter 16) 1. Synthesize end-to-side chain cyclic peptides via a thiazolidine or oxime linkage. Assemble the peptide segment on a Wang resin by the Fmoc-chemistry protocol using stepwise peptide synthesis as described in the Alternate Protocol and outlined in Table 18.5.2. The order is as follows: X-protected peptide sequence-Lys[Boc-Ser(tBu)]βAla, where X = Boc-Cys(StBu) or Boc-NH-O-CH2-CO-.
2. Couple the Lys residue as Fmoc-Lys(Mtt) and remove the Mtt protection group with 1% TFA,/5% TIS in DCM. Couple Boc-Ser(tBu) to the side chain of Lys (Alternate Protocol). 3. Cleave and deprotect the peptide (see Alternate Protocol, steps 6 to 10) using 92.5/2.5/2.5/2.5 TFA/TIS/thioanisol/H2O and allowing the reaction to proceed for 1 hr. Lyophilize the crude product (Alternate Protocol, step 10).
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4. Add 1.55 µmol sodium periodate (0.33 mg) in 3.5 µl H2O to 0.76 µmol peptide precursor dissolved in 1 ml 0.01 M sodium phosphate buffer, pH 6.8. Shake the solution for 2 min, and purify the product by semipreparative RP-HPLC (UNIT 11.6). For oxime formation, the intramolecular cyclization occurs after oxidation to the aldehyde. Purify and characterize the product.
5. Dissolve 0.94 µmol peptide in 1 ml 0.01 M sodium acetate buffer, pH 4.2, and add 9.3 µmol TCEP (2.3 mg) dissolved in 23 µl 0.01 M sodium acetate buffer, pH 4.2. Stir the solution, and monitor the cyclization by analytical RP-HPLC (UNIT 11.6). After purification by semipreparative RP-HPLC (UNIT 11.6), characterize the product by mass spectrometry (Chapter 16). REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Acetal resin Add 0.2 g sodium bicarbonate to a suspension of chloromethylated polystyrene (1% divinylbenzene) copolymer (Merrifield resin, purchased, e.g., from Bachem California; 0.76 mmol/g, 0.5 g) in 3 ml DMSO. After stirring for 7 hr at 155°C, wash the resin with water, methanol, DMF, DCM, and diethyl ether and then dry under vacuum. To confirm the conversion of the chloromethyl resin to the aldehyde resin, use FT-IR analysis, which should show the typical carbonyl absorption at 1700.5 cm–1, and microanalysis, which should show that the resin does not contain any chlorine. The benzaldehyde resin (0.59 mmol/g, 1.0 g) is suspended in 10 ml anhydrous dimethoxyethane with 1 g glycerol and 10 mg p-toluenesulfonic acid. After 24 hr at 90°C, wash the resin with a 1:1 (v/v) mixture of 3% sodium bicarbonate and 1,4-dioxane, followed by water, dioxane, DMF, methanol, DCM, and diethyl ether. Dry under vacuum. Use FT-IR analysis to monitor the disappearance of the carbonyl absorbance at 1700.5 cm–1. To functionalize the acetal resin with a Fmoc-amino acid, activate the Fmoc-amino acid, e.g., Fmoc-Gly-OH (5.58 mmol, 1.66 g) using DCC (3 mmol, 0.62 g), dimethylaminopyridine (0.28 mmol, 35 mg), and HOBt (0.28 mmol, 44 mg) for 20 min in 10 ml DMF. Insoluble dicyclohexylurea is then removed, and the solution is added to a suspension of the glycerol-acetal resin (1.0 g) in 5 ml DMF. After 18 hr, wash the resin three times each with dioxane, DMF, methanol, DCM, diethyl ether and then dry under vacuum. Microanalysis will indicate 0.71% of nitrogen, corresponding to a substitution of 0.5 mmol/g. UV absorbance of the cleaved Fmoc (wavelength 301 nm, I = 7840) will indicate a substitution of 0.5 mmol/g.
Boc-aminooxyacetic acid Add 10 mmol carboxymethylamine hemihydrochloride (Aldrich) to a mixture of 20 ml dioxane, 10 ml water, and 10 ml 1 N NaOH on ice. Add 11 mmol di-t-butyl pyrocarbonate (Aldrich) to the above solution at 0°C and stir the solution overnight at room temperature. Adjust pH to 4 using diluted KHSO4 and extract the product twice, each time for 30 min, with ethyl acetate. Dry the organic phase over MgSO4. Evaporate the organic phase under vacuum, and recrystallize Boc-(aminooxy) acetic acid (Boc-NHNH2) from ethyl acetate–hexane. Synthesis and Application of Peptide Dendrimers As Protein Mimetics
Boc-succinic acid hydrazide On ice, dissolve 42 mmol Boc-NHNH2 (Aldrich) in 20 ml dichloromethane (DCM) and add 40 mmol succinic anhydride. Then add 10 mmol diisopropylethylamine (DIEA) to the above solution at 0°C, and stir the solution for 6 hr at room
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temperature. Remove DCM under vacuum, and dissolve the residue in 60 ml ethyl acetate. Wash the solution twice with 15 ml 5% KHSO4, twice with 10 ml saturated NaCl solution, and finally with 10 ml water. Dry the organic phase over MgSO4, and evaporate the organic phase under vacuum. Wash the oily residue three times, each time with 20 ml ether-hexane and dry under vacuum. Cleavage solution for Fmoc protecting group 90 parts trifluoroacetic acid (TFA) 6 parts thioanisole 3 parts 1,2-ethanedithiol (EDT) 1 part anisole Prepare fresh and chill to 0°C The above reagents are available from Aldrich
Ninhydrin test reagents Solution A: Dissolve 40 g reagent grade phenol in 10 ml absolute ethanol. Solution B: Dissolve 65 mg potassium cyanide in 100 ml water. Dilute 2 ml of this solution to 100 ml with freshly distilled pyridine. Mix solution A and solution B, and store indefinitely at 4°C. Solution C: Dissolve 2.5 g ninhydrin in 50 ml absolute ethanol. Store solution C indefinitely, in the dark. Pyridine from any commercial source may be used to prepare solution B and must be distilled before use. Reflux 500 ml pyridine with 2 g ninhydrin for 2 hr, then distill the pyridine and store several months at 4°C, in a brown bottle (away from light).
Sulfhydryl reducing solution 4.25 ml 6 M guanidine⋅HCl 0. 25 ml 5% (w/v) EDTA (50 mg EDTA/ml) 0.5 ml 2% (w/v) tributylphosphine (Bu3P) in propanol Prepare just before use CAUTION: Tributylphosphine (Bu3P) is a highly toxic, volatile reducing agent and should only be handled in a chemical hood.
Thiobenzhydryl resin 2 for C-terminal thioacid Add thiolacetic acid (CH3COSH, 30 mmol; Aldrich Chemical) and triethylamine (30 mmol; Aldrich Chemical) to a suspension of 4-methylbenzhydryl chloride resin (6 g, 3 mmol) in 60 ml DMF. React the mixture at 60°C for 12 to 18 hr until a negative silver chloride test (see below) of the resin indicates that no free chloride groups are present. Wash the resin three times each with DMF, DCM, CH3OH, DCM, and DMF. Treat resulting resin with a mixture of cysteine methyl ester hydrochloride (9 mmol), triphenylphosphine (9 mmol), and DIEA (9 mmol) in DMF/DCM (3:1, v/v, 60 ml). After 2 hr, wash the resin thoroughly three times each with DMF, DCM, and CH3OH and dry under vacuum. The yield from this procedure should be 6.3 g. For the silver chloride test, remove ∼20 mg of resin to a small glass test tube, add 0.5 ml of pyridine, and heat at 100°C in a heating block for 15 min in a well-ventilated hood. Cool and dilute with an equal volume of 0.1 N nitric acid. Add a few drops of 0.1 N silver nitrate solution. The presence of silver chloride precipitate indicates an incomplete reaction.
Thioester resin (HS-CH2CH2CO-MBHA-resin 1) for C-terminal thioester Add 3-thiopropionic acid (1.2 ml, 13 mmol), HOBt (1.8 g, 13 mmol), and DIC (2 ml, 13 mmol) sequentially to a suspension of 4-methylbenzhydrylamine (MBHA) resin (6 g, 3.2 mmol) in 60 ml DMF. Shake the mixture at room temperature for 30 min until a ninhydrin test (see Support Protocol 1) of the resin indicates that no free
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amino groups are present. Wash the resin three times each with DMF, DCM, CH3OH, DCM, and DMF. Treat resulting resin with a mixture of cysteine methyl ester hydrochloride (0.55 g, 3.2 mmol), triphenylphosphine (0.85 g, 3.2 mmol), and DIEA (0.68 ml, 3.9 mmol) in DMF/DCM (3:1, V/V, 60 ml). After 2 hr, wash the resin thoroughly three times each with DMF, DCM, and CH3OH and dry under vacuum. The yield from this procedure should be 6.3 g.
COMMENTARY Background Information
Synthesis and Application of Peptide Dendrimers As Protein Mimetics
The multiple antigen peptide (MAP) system was originally designed to replace the protein carrier and to overcome the limitations of conventional conjugation approaches (Tam, 1988, 1996; Posnett et al., 1988). The MAP system consists of three structural elements. A simple amino acid (e.g., alanine) is bound through a proper linkage to a solid-phase polymer to initiate the synthesis and serve as an internal standard. An inner core matrix is constructed from several levels of lysine to generate multiple reactive ends (two to eight ends) to which peptides can be linked. The surface layer is composed of either multiple copies of linear or cyclic synthetic peptides attached to the lysine inner-core matrix (Fig. 18.5.1). The MAP makes use of a limited sequential propagation of a trifunctional amino acid to form a low-molecular-weight core matrix. Lysine is suitable for this purpose because both Nα- and Nε-amino groups are available as reactive ends. The sequential propagation of lysine generates 2n reactive ends, each of which can serve as an attachment site for peptides. The complete MAP has been shown to be large enough to stimulate immune responses. The lysine core matrix is small; the bulk of the complex is formed by up to 16 peptides layered around the inner core matrix. As an example, a peptide dendrimer with an octabranched core matrix and eight copies of the peptide, each containing 14 amino acid residues, has a total molecular weight of 13 to 14 kDa. Only 7% of the mass is due to lysine residues of the core matrix; 93% of the mass is due to peptide sequences. The core matrix is oligomeric and contains noncationic peptidyl and isopeptidyl lysine amide on both Nα- and Nε-termini of lysine. As a result, the MAP core matrix is totally bioinactive and functions only as a template to keep the multiple peptides together. It is the dendritic peptide chains on the core matrix that are likely to be mobile and contribute to the biological activity of the MAP system. Recently, Wrighton and co-workers (Wrighton et al., 1997) have used β-Ala-Lys as
a pseudosymmetrical dendrimeric template and have attached two copies of the erythropoietin mimetic peptide (EMP) to the ε-amino group and the α-amino moiety, respectively. The two intramolecular disulfide bridges in EMP were formed by using an orthogonal protecting group scheme. As a result, the dimer has 100-times higher affinity for the erythropoietin receptor (EPOR) and 10-fold better antagonistic activity in vivo than the monomer. The MAP system is chemically unambiguous, and the method of synthesis provides accurate knowledge of conformation and quantity of the peptide. The MAP system is also versatile. If half of the reactive lysine amino chains (i.e., ε-amino groups) are chemically blocked during synthesis, a peptide dendrimer containing two different peptide sequences can be constructed. For example, Nα- and Nε-amino groups of lysine at the last level are orthogonally protected with t-butyloxycarbonyl (Boc) and 9-fluorenylmethyloxycarbonyl (Fmoc), respectively. The first peptide chain can be assembled on the Nα-termini of this special lysine core matrix using Boc chemistry. The Fmoc groups of Nε-termini are not affected during the Boc synthesis. After synthesis of the first peptide is complete, the second peptide chain can be synthesized on the Nε-termini using Fmoc chemistry. The entire MAP system with two different peptide sequences is then cleaved from resin by hydrogen fluoride (HF) cleavage. One such MAP system was used as the model for synthetic hepatitis B vaccines (Tam and Lu, 1989). The simplicity and flexibility of the design and synthesis of MAP also permit incorporation of a hydrophobic built-in adjuvant such as tripalmitoyl-S-glyceryl cysteine (P3C; Defoort et al., 1992). Constrained peptides that mimic native protein conformations are highly desirable because of the advantages in improving their biological activity. To achieve such constrained peptides, linear peptide sequences need to be circularized and then attached to a lysyl MAP core matrix. This approach requires two types of synthetic manipulations: a cyclization strat-
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egy of a linear precursor and a chemoselective method to attach to the template. The conventional approach to achieving two synthetic components is usually cumbersome and requires a scheme of multi-tiered protecting groups. For example, conventional cyclization to form cyclic peptides requires a protection scheme for selected functional side chains. The reactions are carried out in organic solvent (off-resin) or solid phase (on-resin). After cyclization, the side-chain protecting groups are removed. This unit describes two simplified approaches (the direct approach and the indirect approach; also see Fig. 18.5.2) to form cMAPs through unprotected constrained peptides generated from their linear unprotected peptides under aqueous conditions. The authors’ two approaches do not use protecting groups and produce cMAPs directly for a variety of purposes, including antigen scanning and generation of constrained peptide libraries. The direct and indirect methods for preparing cMAP are the same as for MAP, and include an extra cyclization step after the linear unprotected precursors are synthesized. Synthetic peptides have been used extensively for production of site-specific antibodies for confirming the identity of proteins derived from recombinant DNA, for exploring biosynthetic pathways, and for defining precursorproduct relationships (e.g., proenzyme and preproenzyme) and structural domains of proteins (Lerner, 1982). So far, the MAP approach has been used in the preparation of experimental vaccines (Amon and Horwitz, 1992) against hepatitis (Tam and Lu, 1989), malaria (Tam et al., 1990; Pessi et al., 1991), foot-and-mouth disease (Francis et al., 1991), and HIV (Defoort et al., 1992; Nardelli et al., 1992). Various templates derived from peptide and organic compounds containing several peptide chains have been able to induce protein structural motifs like α-helix, β-sheet, β-turn, and loop conformations (Schneider and Kelly, 1995). Since the MAP system also has an increased ordered helical structure (Shao and Tam, 1995) and its synthesis is well established, MAP is suitable as a template to mimic proteins. Many important questions, such as how a certain structural motif can be induced and how the secondary structures interact with each other, can be addressed by using the authors’ MAP system. Artificial enzymes can also be designed and generated because certain tertiary structures associated with a specific enzymatic activity can be mimicked on the surface of the MAP core matrix.
Critical Parameters Chemistry of synthesis The MAP system can be effectively synthesized by both Boc and Fmoc chemistry (Tam and Spetzler, 1997). The selection of the synthetic method depends on several factors, including peptide sequence, instrumentation requirements, cost, and chemical wastes. The most important factor is probably peptide sequence. For sequences very rich in amino acids that are susceptible to strong acid modifications [e.g., alkylation (Cys, Met, Trp, and Tyr), acylation (Glu), and dehydration (Asp-Gly, AspSer)], Fmoc chemistry may provide better results than Boc chemistry. However, use of the low-high hydrogen fluoride cleavage technique has greatly reduced these types of side reactions. Sequences rich in Arg, Trp, Tyr, and Met, as well as peptides with proline as one of the first two residues from the C-terminus, may do poorly in Fmoc chemistry. Occasionally, a peptide cannot be successfully prepared by either of these two methods. A failed synthesis may be caused by sequence-dependent side reactions. The second factor that affects the choice of synthetic method is the instrumentation requirement. In general, Fmoc chemistry is easier in operation and requires less equipment. Boc chemistry normally needs special equipment for the final hydrogen fluoride cleavage of peptide from resin. If cost is a consideration, Boc chemistry is preferable because Bocamino acid derivatives are less expensive than Fmoc-amino acid derivatives. The nature and quantity of chemical wastes produced during synthesis should also be considered. The solvent, coupling reagent, and cleavage reagent wastes generated in Boc chemistry are acidic (due to TFA) and contain halogens (TFA, DCM). The waste generated by Fmoc chemistry is basic (due to piperidine) and is not halogen-containing; the DCM used in reaction 8 of the procedure (Table 18.5.2) is negligible in this regard. For the synthesis of cMAPs, the preparation of their linear precursors are often limited to either Boc or Fmoc chemistry depending on the cyclization method used (see discussion of Method of Synthesis, below). However, the same precautions have to be taken as for the synthesis of MAP. Method of synthesis Both direct and indirect approaches for preparing MAPs are presented in this unit. In the direct approach, the linear peptide sequence
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Synthesis and Application of Peptide Dendrimers As Protein Mimetics
and MAP core matrix are synthesized in a single operation, directly on the resin matrix, by stepwise solid-phase peptide synthesis (Basic Protocol 1 and the Alternate Protocol). The core matrix (two or three levels of lysine) is formed first by coupling lysine to a small amino acid attached to a resin support. The peptide sequence is then synthesized stepwise on this branched-lysine scaffolding. After peptide synthesis, the MAP is cleaved from the resin support. In the indirect approach, the MAP core matrix and peptide are synthesized separately, purified, and then combined to form the complete MAP by one of a number of different ligation methods illustrated in Figure 18.5.5 (Basic Protocols 2 to 3). One advantage of the indirect approach is that the components— MAP core matrix and peptide—are purified before conjugation. Any impurity generated during solid-phase peptide synthesis can be removed; this may be difficult in the direct approach because of the unique structural characteristic of MAP. Preparation of cMAP using the direct approach can be complicated by competing intramolecular versus intermolecular cyclization. The competing reactions can be minimized for single-chain peptides under dilute conditions, but this is not possible for cMAPs because of simultaneous multiple peptide copies on the branched structure. Furthermore, extensive intermolecular cyclization among the precursors can also give polymers. The authors’ solution for avoiding side products and for simultaneously obtaining a self-assembly process to give multiple cyclic peptide is based on ring-chain tautomerization. Ring-chain tautomerization is an entropy process and is conformation-driven as well as concentration-independent, to favor intra- over intermolecular reactions. The initial bond formation is reversible in ring-chain tautomerism. Thermodynamically stable products are formed under equilibrating conditions suitable for self-assembly of the multiple peptide chains on the dendrimer to form end-toside chain cyclic peptides. In the synthetic scheme of cMAP, the ring-chain tautomerization for intramolecular cyclization is a thiazolidine ring generated from an N-terminal Cys and an aldehyde attached to the side chain of Lys (Fig. 18.5.6). Peptides with a motif that favor a cyclic structure are highly recommended in this approach. The authors’ schemes for preparing cyclic peptides by the indirect approach result in ligation sites of peptide bonds like X-Cys and X-SPro and surrogate peptide bonds such as
thiazolidine and oxime that both are metabolically stable (Fig. 18.5.3). These schemes also produce peptides which are cyclized through end-to-end, end-to-side chain, or side chain to side chain (Pallin and Tam, 1995, 1996; Botti et al., 1996; Zhang and Tam, 1997), depending on the strategic placement of the two intended sites that must react with each other. Although peptide and nonpeptide bonds are described, the chemistries governing their syntheses are rather similar. To generate unprotected cyclic peptides with peptide bonds such as X-Cys and X-SPro from linear unprotected precursors, the reaction must be regioselective (Liu and Tam, 1994; Tam, 1995). A common strategy for obtaining regioselectivity is entropic activation achieved by capturing the Nα- and Cα-termini at close proximity as a covalent bond allowing an intramolecular acyl shift and forming a peptide bond. In cyclization reactions, the capturing reaction between an Nα-amine and a weakly activated Cα-acyl moiety has to be specific in the presence of many other side-chain functional groups, including Nε-amine. Two methods are available to form a Cys bond in the ligation site. In one method, cyclization is achieved via an intramolecular transthioesterification of the linear precursor containing an N-terminal Cys and a C-terminal thioester. The thioester intermediate subsequently undergoes a proximity-driven S- to N-acyl transfer to give the desired amide bond, and introduces a thiol moiety (Basic Protocol 5). In the other method, the cyclization occurs via an acyl disulfide–mediated intramolecular acylation. The linear precursor consists of an N-terminal Cys protected and activated as Cys(Npys) and a C-terminal thiocarboxylic acid. The mixed disulfide undergoes an S to N-acyl transfer through a six-member ring and a native Cys residue is obtained after thiolytic reduction (Support Protocol 4). Cyclic peptides with a thiol moiety are useful as building blocks in the indirect approach for preparing cMAPs (see discussion of Method of Ligation, below). Also, cyclic peptides containing an SPro residue to mimic Pro can be generated (Support Protocol 2). The cyclization is mediated by an amine-aldehyde condensation. The linear precursor contains an N-terminal Cys and a C-terminal glycoaldehyde generated from NaIO4 oxidation of the corresponding glyceric ester. The Cys residue is protected as Cys(StBu) because of its sensitivity to NaIO4 oxidation. First, a thiazolidine ring is formed when the StBu protecting group is removed by phosphine. Then a proximity-driven O to Nacyl transfer reaction occurs through a tricyclic
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ring contraction to afford the peptide bond. Cyclic peptides through side-chain-to-end and end-to-end can be achieved. In the preparation of cyclic peptides with peptide bond surrogates from totally unprotected linear precursors, chemoselective ligation based on carbonyl chemistry is used. Carbonyl chemistry exploits the selectivity of weak bases in the condensation with an aldehyde under acidic conditions where the basic sidechain nucleophiles are protected by protonation. For the purposes here, weak bases such as thiols, hydrazides, and amino-oxy groups are available, since they are easy to incorporate into the peptide during solid-phase synthesis. An aldehyde is obtained by NaIO4 oxidation of N-terminal Ser, Thr, or Cys under neutral conditions to give an α-oxoacyl group. A thiazolidine ring is formed from the reaction between the aldehyde and the 1,2-aminoethanethiol of N-terminal Cys. Under the same conditions, an amino group will form a reversible Schiff base with the aldehyde. The Cys residue, which is incorporated as Cys(StBu), can play a dual function in the synthetic scheme. First, it can be used for cyclization where the cyclization is mediated after deprotection of the thiol of Cys, as already mentioned above. Second, it can be applied for ligating unprotected cyclic peptides to a core matrix when the Cys residue is deprotected after the cyclic peptide is formed. Another weak base is aminooxy, which affords oxime at low pH. In the authors’ scheme, the aldehyde group is attached to the side chain of Lys while the nucleophilic aminooxyl or cysteinyl moiety is incorporated to either the amino terminal of the peptide or through the side chain of Lys to afford end-to-side chain or side chain-to-side chain cyclic peptides. Method of ligation The advantage of the indirect approach is that conjugation between the unprotected peptide and MAP core is chemically unambiguous. The functional groups on the peptide chain and core matrix can be designed to avoid reacting with other functional groups on the peptide. A variety of conjugation methods and cross-linking reagents may be used for conjugation to avoid modification of the peptide. The methods described in this unit are some of many variations (Fig. 18.5.5). A pair of mutually reactive functional groups such as a nucleophile and an electrophile are introduced on the monomeric peptide and MAP core matrix, respectively. The two functional groups only react specifically with each other. The methods that use thiol and
carbonyl chemistries are suitable for the preparation of both MAP and cMAP with nonpeptide bonds. A thiol group can be selectively reacted in the presence of amine and other side-chain groups because it is a stronger nucleophile, especially in neutral or acidic conditions where the amine group is protonated. Haloacetyl (Fig. 18.5.5) is a functional group used for selectively reacting with a thiol group under neutral to slightly basic pH conditions. However, under these conditions, the thiol tends to be oxidized to disulfide. Both unprotected linear (Basic Protocol 2) and cyclic (Basic Protocol 6) peptides contain a free thiol that has been ligated to chloroacetyl groups incorporated on the lysine core matrix. The indirect approach using thiol chemistry also allows the peptides to be attached to the MAP core matrix through either the amino or the carboxyl terminus, increasing the flexibility of the design. However, other functional groups such as maleimide or aromatic thiols like 2-thiopyridyl, nitropyridyl sulfenyl, or 4-dithiopyridyl, which also react specifically with a thiol, can be applied to ligate unprotected peptides to the MAP core matrix. Carbonyl chemistry that exploits the selectivity of weak bases in condensation with an aldehyde has already been mentioned (see discussion of Method of Synthesis, above) and is shown in Figure 18.5.5. Thiazolidine ring formation has been used to ligate both linear (Basic Protocol 3) and constrained peptides (Basic Protocol 5) to a MAP core matrix containing α-oxoacyl groups. Other types of weak bases are hydroxylamine and hydrazine, which afford oxime and hydrazone bonds with an aldehyde moiety. Oxime and hydrazone bonds in the ligation site have been suitable for linear unprotected peptides (Basic Protocol 3). Both types of weak bases are added to the N-terminus, so the peptide sequence will be oriented with the amino terminal residues proximal to the core matrix and the carboxyl terminal residues distal to core matrix. The conjugation reactions listed in Figure 18.5.5 have some limitations. The haloacetyl derivative may react with amine or other nucleophilic groups in peptides in basic medium. This type of reaction will lead to chemical ambiguity of the conjugate. To prevent this reaction, the pH of the conjugating solution should not be above 9.0. Aldehyde groups reacting with N-terminal cysteine rapidly form a stable thiazolidine derivative at pH 4 to 6. The resulting thiazolidine is quite stable under normal manipulating conditions, but the aldehyde
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itself is unstable. However, this can be circumvented by inert gas protection. Hydroxylamine and hydrazine react with aldehydes, forming an oxime and a hydrazone linkage, respectively, at pH ∼5. Both linkages are stable at physiological pH (6 to 8), but the hydrazone derivative will be hydrolyzed at a lower or higher pH. The rate of hydrolysis is pH dependent. Stability of the linkage can be increased by reducing the hydrazone bond to hydrazine with sodium cyanoborohydride (NaBH3CN).
Troubleshooting
Synthesis and Application of Peptide Dendrimers As Protein Mimetics
After the peptide sequence is selected, MAP or cMAP with this special sequence can be synthesized by solid-phase synthesis. As with solid-phase peptide synthesis, the reactions of each cycle (including deblocking and coupling reactions) must be very efficient and should be driven to completion in a short time. The following calculations demonstrate the importance of high yield from each cycle. If the mean yield of each cycle of synthesis is 95%, the final yield for a peptide with 20 amino acid residues would be only 35.8%. However, if the yield of each cycle is 98%, 99%, 99.5%, or 99.9%, the total yields for the synthesis of a peptide with 20 residues would be 66.8%, 81.8%, 90.5%, or 98.0%, respectively. Furthermore, the byproducts produced in peptide synthesis usually closely resemble the desired peptide product in sequence and structure and are very difficult to remove by normal separation methods. Because our MAP system is a dendrimer, the degree of heterogeneity of peptide products will be much higher than that of solid-phase peptide synthesis. For example, in the synthesis of a MAP system with four branches, a 99% yield from each cycle results in 96% of MAP with the correct structure. If the mean yield of each cycle of synthesis is 99%, the synthesis of a four-branched MAP system with a peptide antigen of 20 amino acid residues will result in final yield of 44.2% with the correct sequence, instead of 81.8% obtained in usual solid-phase peptide synthesis. The major side reactions during solid-phase peptide synthesis are: deletion, where one or more amino acid residues are skipped during synthesis; termination, where chain elongation stops at a point because of acylation or other side reaction; and racemization, where the amino acid is racemized during peptide elongation. All these side reactions result in heterogeneity of peptide product, and decrease the total yield of synthesis. To overcome these side reactions, excess amounts of reagents should
be used in the synthesis to force reactions to completion and to shorten the reaction time. Double coupling or even coupling at elevated temperatures (50° to 60°C) should be used if a single coupling fails to complete the reaction. Efficiency of synthesis should be closely monitored by the ninhydrin test (Support Protocol 1) or quantitative ninhydrin test (Sarin et al., 1981). The unusual properties of MAP systems (e.g., their multibranched structure and tendency to aggregate) should be considered to achieve a high-quality product. The synthesis should be started with a lower level of amino acid loading on the resin than is customary with usual solid-phase peptide synthesis, because peptide content will increase geometrically with each additional layer of lysine. Peptide synthesis will be difficult if the conventional loading of 0.3 to 0.8 mmol amino acid/g resin is used. In the Alternate Protocol for Fmoc chemistry, because resin with 0.1 mmol/g loading is not commercially available, synthesis starts with a resin of 0.3 to 0.5 mmol alanine/g resin, but only some of the amino groups are used for preparation of the dendrimer. Dimethyl formamide (DMF) is a more suitable solvent than dichloromethane (DCM) for avoiding potential aggregation of peptides. The peptide resin should not be dried at any stage of synthesis because resolvation of dried resin that contains MAP scaffolding is difficult. During HF cleavage in Boc synthesis, several precautions should be taken to obtain satisfactory results. The HF line must be kept clean. The line should be checked before each cleavage. Very often, heterogeneous and complex peptide products are caused by a dirty, contaminated HF line. The HF line should also be checked for leaks before each cleavage. A leaky HF line can result in the loss of HF and dimethyl sulfide (DMS) during the cleavage reaction; this can lead to unsatisfactory results. A leaky HF line may also allow water and carbon dioxide into the reaction vessel, diluting the HF cleavage mixture. The cleavage mixture should be mixed completely during the reaction. Sometimes the magnetic stir-bar is frozen by the p-cresol or p-thiocresol mixture during the introduction of the reagent. This can be avoided by adding the reagents in the following order: (1) peptide resin, (2) p-cresol and pthiocresol in melted form, then, after cooling and when the mixture has solidified, (3) the magnetic stirbar, and (4) DMS. In Fmoc synthesis, trifluoroacetic acid (TFA) cleavage should be prolonged if the peptide contains Arg(Pmc). Thioanisole can
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accelerate the cleavage of the Pmc group. The deprotection of trityl (Trt) from cysteine residues is reversible, and scavengers such as thiophenol and 1,2-ethanedithiol (EDT) should be added to the cleavage mixture to prevent Trt reattachment to Cys. EDT also minimizes the alkylation reaction of Trp. The optimal volume of cleavage solution should be determined experimentally. In general, larger volumes of cleavage solution will, to some extent, suppress side reactions because most of the side reactions in the cleavage stage are bimolecular reactions. However, the more cleavage solution used, the larger the volume of TFA that must be removed, and the larger the amount of scavenger that will remain, making purification and analysis of the peptide more difficult. TFA can be reduced to a smaller volume under a vacuum by means of rotary evaporation, but the mixture should never be dried. A small-scale cleavage, using 20 to 30 mg of peptide resin, should be performed to determine optimal cleavage reaction conditions such as the combination and the ratio of scavengers, reaction time, and the volume of cleavage mixture. HPLC can be used to evaluate the results. Reaction conditions should be modified if the crude cleavage product is not as homogeneous as expected. The desired degree of purity should be ∼60% or a major peak in the HPLC. MAP systems prepared by the direct approach have been found to have an unusual tendency to aggregate after cleavage from the resin. The entire MAP should be dialyzed under basic, denaturing conditions using 8 M urea to remove unwanted additives from the cleavage reaction. Treatment with base under these conditions will also convert the possible strong acid–catalyzed O-acyl rearrangement product of serinyl and threonyl peptides back to N-acyl peptides. After dialysis, MAP systems can be further purified by either RP-HPLC or highperformance gel-filtration chromatography. In the additional cyclization step for preparing cyclic unprotected peptide building blocks and the entire cMAP by the direct approach, some of the protocols involve oxidation with NaIO4. The order of the synthetic scheme to generate aldehydes from the oxidation has to be followed carefully, since not all the functional groups in their unprotected form, suitable for cyclization, are stable under the condition of NaIO4. Perform the NaIO4 oxidation at neutral pH in the presence of a large excess of free methionine as a scavenger, to prevent oxidation of Met to Met(O). The oxidation rate increases
as pH decreases, but side reactions occur at lower pH. The aldehyde may decompose, and the absence of air is necessary. Flushing with nitrogen or argon for 30 min results in an oxygen-free solution. For cyclizations that include the presence of free sulfhydryl groups, the reactions must also be carried out in a solution devoid of oxygen to avoid dimerization. Also, addition of thiols or trialkylphosphine to the solution prevents the formation of dimers. The cyclization reactions can be carried out in a final concentration of up to 20 mM without affecting the yield of the desired product, since the reactions usually are concentration-independent. In the indirect approach where either linear or cyclic unprotected peptides are ligated to a MAP core matrix using carbonyl or thiol chemistry, the ligation reactions are also performed in an oxygen-free solution. Usually, the ligation is performed with a peptide concentration at 1.2- to 2-fold excess relative to core matrix. For the thiol alkylation reaction in thiol chemistry, iodoacetylated and bromoacetylated derivatives are known to be more reactive than the chloroacetylated derivative. However, the chloroacetylated derivative is more stable during the preparation of core matrix, especially during HF cleavage. The peptide and nonpeptide bonds formed in MAP and cMAP are stable at physiological pH (pH 6 to 8) as well as at low pH during analytical and preparative HPLC. However, for long-term storage, the peptides must be stored as a dry powder at a maximum temperature of −20°C.
Anticipated Results If synthesis of an eight-branched MAP with ∼15 amino acid residues starts from 0.5 g resin with a resin loading of 0.1 mmol/g, then >0.5 g of crude MAP can be obtained. Its purity depends on both the peptide sequence and the method used for synthesis. Usually the purity of crude products will be >60%, and after purification ∼40% to 50% can be recovered as purified MAP. This amount is usually more than sufficient for biological purposes. In general, the reaction should start with 0.3 to 0.5 g resin. For preparing cMAP using the direct approach, the overall yield can be up to 60% for cyclic peptides containing 17 and 24 amino acid residues, also starting with 0.5 g resin (loading is 0.1 mmol/g). In the direct approach, the average yield for MAP is between 50% and 60% using the ligation methods in Figure
Preparation and Handling of Peptides
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18.5.5. The yield of the pure cMAP system is ∼30% when cyclic peptides ranging from 6 to 14 amino acids are used. However, the site and structure, as well as the excess and concentration of the peptides influence the obtained yield. Both methods for generating MAP and cMAP systems are applicable for mimetic proteins. If HPLC of a crude MAP prepared by the direct approach shows multiple peaks without a major product, the complex may be difficult to purify, and the yield of MAP may not be satisfactory. In this case, the synthetic procedure (synthetic chemistry, protecting combination, coupling procedure, and/or cleavage conditions) should be modified. Also, the progress of the ligation reactions in the indirect approach is conveniently monitored by RP-HPLC (UNIT 11.6). If no product is formed, the peptides should be reanalyzed by mass spectrometry (Chapter 16) for the presence of nucleophilic and electrophilic moieties that have to react with each other. The peptide may even have to be resynthesized. Also, be aware of the formation of byproducts that can be generated from hydrolysis of the starting material at a given pH. Therefore, recheck the pH during the cyclization and assembling reactions.
Time Considerations
Synthesis and Application of Peptide Dendrimers As Protein Mimetics
An automatic peptide synthesizer can assemble ∼15 residues in one day by a rapid procedure using a 15-min cycle or in 2 days with the the normal cycle of ∼1 hr. The synthesis of a MAP system with its peptide chain of 10 to 20 residues will usually take 1 to 2 days using the direct approach. Using a manual shaker for solid-phase peptide synthesis, assembly of as many as 6 to 12 amino acid residues onto the resin in one day is possible. The same MAP system can be prepared within 4 days by manual synthesis. Cleavage and subsequent purification can take from 2 days to 1 week, depending on the purity of the crude product as well as the required purity of the isolated MAP system. In preparing cMAP using the direct approach, 5 days should be added for the extra steps of cyclization, purification and characterization. In the indirect approach for preparing MAP systems, both the peptide and core matrix are prepared and purified before ligation. Ligation of the peptide with the core matrix requires an additional reaction step and adds 2 to 3 days to the process. For cyclization of the linear unprotected precursors, 3 extra days are needed.
Literature Cited Amon, R. and Horwitz, R.J. 1992. Synthetic peptides as vaccines. Curr. Opin. Immunol. 4:449453. Botti, P., Pallin, T.D., and Tam, J.P. 1996. Cyclic peptides from linear unprotected peptide precursors through thiazolidine formation. J. Am. Chem. Soc. 118:10018-10024. Defoort, J.-P., Nardelli, B., Huang, W., Ho, D.D., and Tam, J.P. 1992. Macromolecular assemblage in the design of a synthetic AIDS vaccine. Proc. Natl. Acad. Sci. U.S.A. 89:3879-3883. Francis, M.J., Hastings, G.Z., Brown, F., McDermed, J., Lu, Y.A., and Tam, J.P. 1991. Immunological evaluation of the multiple antigen peptide (MAP) system using the major immunogenic site of foot-and-mouth disease virus. Immunology 73:249-254. Houghten, R.A., Pinilla, C., Blondelle, S.E., Appel, J.R., Dooley, C.T., and Cuervo, J.H. 1991. Generation and use of synthetic peptide combinatorial libraries for basic research and drug discovery. Nature 354(6348):84-6 Lerner, R.A. 1982. Tapping the immunological repertoire to produce antibodies of predetermined specificity. Nature (Lond.) 299:592-596. Liu, C.-F. and Tam, J.P. 1994. Peptide segment ligation strategy without use of protecting groups. Proc. Natl. Acad. Sci. U.S.A. 91:6584-6588. Liu, C.-F. and Tam, J.P. 1997. Synthesis of a symmetric branched peptide: Assembly of a cyclic peptide on a small tetraacetate template. Chem. Commun. 1619-1620. Lu, Y.A., Clavijo, P., Galantino, M., Shen, Z.-Y., Liu, W., and Tam, J.P. 1991. Chemically unambiguous peptide immunogen: Preparation, orientation and antigenicity of purified peptide conjugated to the multiple antigen peptide system. Mol. Immunol. 28(6):623-630. Merrifield, R.B. 1963. Solid phase synthesis. I. J. Am. Chem. Soc. 85:2149-2154. Mutter, M. and Vuilleumier, S. 1989. A chemical approach to protein design-template-assembled synthetic proteins (TASP). Angew. Chem. Int. Ed. Engl. 28:535-554. Nardelli, B., Lu, Y.A., Shim, D.R., Delpierre-Defoort, C., Profy, A.T., and Tam, J.P. 1992. A chemically defined synthetic vaccine model for HIV-1. J. Immunol. 148:914-920. Pallin, D.T. and Tam, J.P. 1995. Cyclisation of totally unprotected peptides in aqueous solution by oxime formation. Chem. Commun. 2021-2022. Pallin, T.D. and Tam, J.P. 1996. Assembly of cyclic peptide dendrimers from linear building blocks in aqueous solution. Chem. Commun. 11:13451346. Pessi, A., Valmori, D., Migliorni, P., Tougne, C., Bianchi, E., Lambert, P.-H., Corradin, G., and Giudice, del G. 1991. Lack of H-2 restriction of Plasmodium falciparum (NANP) sequence as multiple antigen peptide. Eur. J. Immunol. 21:2273-2276.
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Posnett, D.N., McGrath, H., and Tam, J.P. 1988. A novel method for producing antipeptide antibodies. J. Biol. Chem. 263:1719-1725. Sarin, V.K., Kent, S.B.H., Tam, J.P., and Merrifield, R.B. 1981. Quantitative monitoring of solidphase peptide synthesis by the ninhydrin reaction. Anal Chem. 117:147-157. Sasaki, T. and Kaiser, E.T. 1989. Helicrome: Synthesis and enzymatic activity of a designed heme protein. J. Am. Chem. Soc. 111:380-381. Schneider, J.P. and Kelly, J.W. 1995. Templates that induce α-helical, β-sheet, and loop conformations. Chem. Rev. 95:2169-2187. Shao, J. and Tam. J.P. 1995. Unprotected peptides as building blocks for the synthesis of peptide dendrimers with oxime, hydrazone and thiazolidine linkages. J. Am. Chem. Soc. 117:38933899. Spetzler, J.C. and Tam, J.P. 1996. Self-assembly of cyclic peptides on a dendrimer: Multiple cyclic antigen peptides. Pept. Res. 9:290-296. Stewart, J.M. and Young, J.D. 1984. Solid Phase Peptide Synthesis, 2nd ed. Pierce Chemical Co., Rockford, Ill. Tam, J.P. 1988. Synthetic peptide vaccine design: Synthesis and properties of a high density multiple antigenic peptide system. Proc. Natl. Acad. Sci. U.S.A. 85:5409-5413. Tam, J.P. 1995. Synthesis and applications of branched peptides in immunological methods and vaccines. In Peptides: Synthesis, Structures and Applications (ed. B. Gutte), pp. 455-501. Academic Press, San Diego. Tam, J.P. 1996. Recent advances in multiple antigen peptides. J. Immunol. Methods 196:17-32. Tam, J.P. and Lu, Y.-A. 1989. Vaccine engineering: Enhancement of immunogenicity of synthetic peptide vaccine related to hepatitis in chemically defined models consisting of T and B cell epitopes. Proc. Natl. Acad. Sci. U.S.A. 86:90849088. Tam, J.P. and Spetzler, J.C. 1997. Multiple antigen peptide system. Methods Enzymol. 289:612-637. Tam, J.P., Clavijo, P., Lu, Y.A., Nussenzweig, R.S., Nussenzweig, V., and Zavala, F. 1990. Incorporation of T and B epitopes of the circumsporozoite protein in a chemically defined synthetic vaccine against malaria. J. Exp. Med. 171:299306.
Tam, J.P., Wu, C.-R., Liu, W., and Zhang, J.-W. 1991. Disulfide bond formation in peptides by dimethyl sulfoxide: Scope and applications. J. Am. Chem. Soc. 113:6657-6662. Unson, C.G., Erickson, B.W., Richardson, D.G., and Richardson, J.S. 1984. Protein Engineering: Design and synthesis of a protein. Fed. Proc. 43:1837. Wrighton, N.C., Balasubramanian, P., Barbone, F.P., Kashyap, A.K., Farrell, F.X., Jolliffe, L.K., Barrett, R., and Dower, W.J. 1997. Increased potency of an erythropoietin peptide mimetic through covalent dimerization. Nat. Biotechnol. 15:1261-1265. Zhang, L. and Tam, J.P. 1997. Synthesis and application of unprotected cyclic peptides as building blocks for peptide dendrimers. J. Am. Chem. Soc. 119:2363-2370.
Key References Fields, G.B. and Noble, R. 1990. Solid phase peptide synthesis utilizing 9-fluorenylmethoxycarbonyl amino acids. Int. J. Pept. Protein Res. 35:161214. An extensive and useful literature review of Fmoc solid-phase peptide synthesis. Grant, G.A. (ed.) 1992. Synthetic Peptides—A User’s Guide. W.H. Freeman, New York. A practical book for solid-phase peptide synthesis which covers synthesis, purification, analysis, and applications of synthetic peptides. References include papers published in 1992. Stewart and Young, 1984. See above. An excellent laboratory book for solid-phase peptide synthesis. Tam and Spetzler, 1997. See above. A practical protocol for preparing MAPs and cMAP using direct and indirect approaches. Zhang and Tam, 1997. See above. A useful research paper discussing preparation and analysis of cyclic peptides and cMAP.
Contributed by James P. Tam and Jane C. Spetzler Vanderbilt University School of Medicine Nashville, Tennessee
Preparation and Handling of Peptides
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Disulfide Bond Formation in Peptides
UNIT 18.6
The formation of disulfide bridges is often a crucial final stage in peptide synthesis. There is compelling evidence that the disulfide pattern can be critical in the folding and structural stabilization of many natural peptide and protein sequences, while the artificial introduction of disulfide bridges into natural or designed peptides may often improve biological activities/specificities and stabilities. This unit will provide the experimentalist with a highly selective, albeit state-of-the-art, menu of procedures that can be tried in order to establish intramolecular or intermolecular disulfide bridges in targets of varying complexities. In all, this unit consists of 15 procedures, divided into two major classes: (1) formation of disulfides from free thiol precursors (see Basic Protocols 1 through 5 and Alternate Protocols 1 through 5), and (2) symmetrical formation of disulfides from S-protected precursors (see Basic Protocols 6 through 8 and Alternate Protocols 6 and 7). See Strategic Planning for further discussion of this breakdown. Directed methods for the unsymmetrical formation of disulfides, which offer control when two different chains are to be linked, use relatively sophisticated and specialized chemistry; examples are not provided herein. Reactions to form disulfides can be carried out with peptides either in solution or while they remain anchored to a polymeric support—in the latter case, taking advantage of the pseudo-dilution phenomenon that favors intramolecular cyclization over intermolecular side reactions. Other possible advantages of solid-phase methods relate to the circumvention of peptide solubility issues, and the chance to readily remove excess oxidizing agents as well as to exchange the solvent milieu by simple filtration and washing steps. The authors assume throughout this unit that the linear precursors (free or S-protected), which are to be converted to the corresponding disulfide species, can be prepared in a reasonable state of chemical homogeneity. When the plan is to carry out the disulfide-forming steps in solution, it is often advantageous to first purify the linear precursor. Solution and solid-phase methods of peptide synthesis, and purification of synthetic products both before or after the chemical steps to form disulfide bonds, are beyond the scope of the unit, but have been amply documented in the primary and review literature. STRATEGIC PLANNING Formation of Disulfides from Free Thiol Precursors The most straightforward approach for the preparation of disulfide-containing peptides involves initial assembly of the linear chain, using the same (invariably) acid-labile protecting group for all Cys residues. Figure 18.6.1 shows the structures of some such protecting groups that are compatible with solid-phase peptide synthesis (SPPS) methods: S-Xan, S-Tmob, S-Mmt, or S-Trt for Fmoc SPPS, and S-Mob or S-Meb for Boc SPPS. Subsequently, the polypeptide is released from the support, concomitant with removal of acid-labile protecting groups for the side chains of all (or almost all) other amino acid residues. This may be followed by treatment with reducing agents to ensure the sole presence of the linear monomeric chain, and then purification under acidic conditions that minimize the possibility of inadvertent premature oxidation. The resultant precursor, in which all Cys residues should be in the free sulfhydryl form, can then be subjected to a variety of solution oxidation procedures, keeping in mind that the precise experimental conditions, e.g., pH, ionic strength, organic cosolvent, temperature, time, and concentration, can often make a substantial difference in terms of the quality of disulfide product obtained. High dilution is recommended in order to minimize physical aggregation, and also to minimize the chemical formation of dimers, oligomers, or intractable polymers. Contributed by Lin Chen, Ioana Annis, and George Barany Current Protocols in Protein Science (2001) 18.6.1-18.6.19 Copyright © 2001 by John Wiley & Sons, Inc.
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In a variation of the general approach, free sulfhydryl precursors can be converted to the corresponding disulfides while the peptide is still retained on the solid support; this requires cysteine protection that is selectively removable in an earlier on-resin step. Practical examples of this approach are given in Basic Protocols 1 to 5 and Alternate Protocols 1 to 5. Symmetrical Formation of Disulfides from S-Protected Precursors Several oxidizing reagents can be used for the formation of disulfide bridges directly from certain S-protected cysteine residues, without the intermediacy of free bis(thiol) intermediates. This approach is used for intramolecular or intermolecular pairing of two Cys residues that originally have the same protecting group, and it can be generalized in experiments aimed at the construction of multiple disulfides. Most of the S-protecting groups already shown in Figure 18.6.1 can be manipulated in this way, as can the S-Acm group, the structure of which is presented in Figure 18.6.2. Typical oxidative reagents include iodine, thallium (III) trifluoroacetate [Tl(tfa)3], and a variety of alkyltrichlorosilane-sulfoxide combinations, and both solution and on-resin deprotection/oxidation methodologies have been described. In most cases, caution is required when peptide substrates contain sensitive residues, especially Met and Trp. Practical examples of this approach are given in Basic Protocols 6 through 8 and Alternate Protocols 6 and 7.
Typically used with Boc SPPS
– S – CH2
X = CH3
X
p-methylbenzyl (Meb)
X = CH3O p-methoxybenzyl (Mob)
Typically used with Fmoc SPPS
–S
9H-xanthen-9-yl (Xan) O
H
CH3O – S – CH2
OCH3
2,4,6-trimethoxybenzyl (Tmob)
OCH3
–S –C
X
X = CH3O 4-methoxytriphenylmethyl (Mmt) X = H triphenylmethyl (trityl; Trt)
Disulfide Bond Formation in Peptides
Figure 18.6.1 Acid-labile Cys protecting groups. The protecting group structure is drawn to include the sulfur (shown in bold) of the cysteine that is being protected.
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Typically used with Boc SPPS
– S – CH2
9 - fluorenylmethyl
H
Typically used with either Boc or Fmoc SPPS O – S – CH2 – NH – C – CH3 acetamidomethyl (Acm)
CH3 – S – C – CH3
tert-butyl (tBu)
CH3 CH3 – S – S – C – CH3
tert-butylmercapto (StBu)
CH3
Figure 18.6.2 Essentially acid-stable Cys protecting groups that are removed by orthogonal methods. The protecting group structure is drawn to include the sulfur (shown in bold) of the cysteine that is being protected.
AIR OXIDATION Molecular oxygen can promote disulfide formation under slightly alkaline conditions, usually pH 7.5 to 8.5. Oxidation is carried out by simple aeration under gentle stirring, or by bubbling oxygen through a dilute solution of the peptide. Reactions are accelerated at higher pH, so long as materials are soluble. This widely used approach may be subject to one or more of the following limitations: (1) oligomerization, despite precautions to carry out reactions at low concentrations of substrate; (2) inadequate solubility of some basic or hydrophobic peptides; (3) long reaction times (1 to 5 days); (4) difficulty in optimizing the oxidation, because the rate depends on trace amounts of metal ions; and (5) accumulation of side products due to oxidation of Met residues.
BASIC PROTOCOL 1
Materials Poly(thiol) peptide, previously purified (see UNIT 8.7 for purification techniques) Buffer, selected from among: 0.1 to 0.2 M Tris⋅Cl, pH 7.7 to 8.7 (APPENDIX 2E) Tris⋅acetate, pH 7.7 to 8.7 0.01 M phosphate buffers, pH 7 to 8 (APPENDIX 2E) 0.01 M ammonium bicarbonate, pH 8 Air or oxygen Additional reagents and equipment for dialysis (UNIT 4.4 and APPENDIX 3B), HPLC (UNIT 8.7), and gel-filtration (UNIT 8.3) or ion-exchange (UNIT 8.2) chromatography 1. Dissolve the peptide substrate in the appropriate buffer, at a typical concentration of 0.01 to 0.10 mg/ml (∼0.01 to 0.1 mM). 2. Stir the solution in open atmosphere, or while oxygen is bubbling through it.
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3. Monitor progress of the reaction by analytical HPLC (UNIT 8.7); withdraw small aliquots (e.g., 20 µl to fill an HPLC injection loop of the same size) to carry out the analysis. 4. Upon completion of the oxidation reaction, concentrate the peptide material by direct lyophilization. Alternatively, first quench the reaction by acidification, and perform dialysis (UNIT 4.4 or APPENDIX 3B) to exchange out salts. Some disulfide-containing peptides are labile when concentrated at alkaline pH, whereas the presence of acid minimizes disulfide interchange.
5. Purify the product by standard methods, e.g., preparative HPLC (UNIT 8.7), gel filtration (UNIT 8.3), or ion-exchange (UNIT 8.2) chromatography, as appropriate. Peptide concentrations have been reported that are as as low as 1 ìM and as high as 1 mM, and the reaction time can vary from a few hours to a few days. As a starting point, try one of the buffers listed in Materials. For the oxidation of certain peptides, the use of organic cosolvents [e.g., methanol, acetonitrile, dioxane, and 2,2,2-trifluoroethanol (TFE)] and the addition of tertiary amines [e.g., N-methylmorpholine (NMM), triethylamine, or N,N-diisopropylethylamine (DIEA)], has been recommended. Addition of moderate concentrations of denaturants such as 0.5 to 3 M urea or 0.1 to 1.5 M guanidinium hydrochloride may help avoid aggregation and facilitate the renaturation process. Addition of 0.1 to 1 ìM CuCl2 has also been reported to improve certain oxidations. ALTERNATE PROTOCOL 1
ON-RESIN AIR OXIDATION In favorable cases, it is possible to use air to oxidize the resin-bound product of a linear sequence assembled by SPPS. According to this plan, the protecting groups for Cys must be removed selectively while negligible or no cleavage occurs of the anchoring linkage; whether other side-chain protecting groups are retained or removed at the same time is usually not as paramount a consideration. With acid-labile Cys protecting groups (Fig. 18.6.1) this means that the anchor should be acid-stable, or more commonly, that the Cys protecting group chosen is one that can be removed by short treatment with dilute acid. Alternatively, the usual acidolyzable anchors can be used together with acid-stable, orthogonally removable Cys protecting groups (Fig. 18.6.2). Once the on-resin deprotection step has been completed, the resin is washed, suspended in a polar solvent in the presence of a tertiary amine base, and exposed to air for the required oxidation. Subsequent cleavage from the support may provide, in a respectable yield, the wanted monomeric (if intramolecular cyclization) or homodimeric (if intermolecular co-oxidation) disulfide product; peptidic material not readily accounted for is invariably oligomeric and serves to diminish the overall yield. Materials Protected peptide-resin, prepared by optimized linear SPPS chain assembly Deprotection reagents and solvents Appropriate wash solvents 0.02 to 0.175 M triethylamine in N-methylpyrrolidone (NMP) Air or oxygen Appropriate peptide cleavage cocktail (UNIT 18.5), and materials for workup 2-ml plastic syringe fitted with a polypropylene frit (use larger syringe for larger amounts of resin)
Disulfide Bond Formation in Peptides
Additional reagents and equipment for HPLC (UNIT 8.7) and gel-filtration (UNIT 8.3) or ion-exchange (UNIT 8.2) chromatography
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1. Place protected peptide-resin into a 2-ml plastic syringe fitted with a polypropylene frit, and carry out a selective deprotection step that removes protection on Cys while not affecting the anchoring linkage. Upon completion of the deprotection, wash thoroughly with appropriate solvents. For example, S-Tmob or S-Xan can be removed selectively by brief treatment with dilute acid (Munson and Barany, 1993; Munson et al., 1993; Hargittai and Barany, 1999). S-StBu can be removed by thiolysis (Eritja et al., 1987). S-Acm can be removed by treatment with Hg2+ salts (Tam and Shen, 1992). S-Fm can be removed by piperidine-promoted β-elimination (Albericio et al., 1991). Precise protocols to do this are in the aforementioned literature references.
2. Incubate the peptide-resin (containing free thiol moieties) with 0.02 to 0.175 M (2 to 10 equivalents) triethylamine in NMP at 25°C. Other polar solvents (e.g., DMF) that swell the resin can be used also.
3. Gently bubble air or oxygen through the suspension. 4. Optional: Monitor progress of the reaction by taking small aliquots of the peptideresin (e.g., 10 to 20 mg) for final cleavage, and evaluate the cleaved material by HPLC analysis. The reaction time can vary from 5 to 36 hr. This step is optional, but is particularly valuable as part of exploratory studies.
5. Cleave the peptide from the resin with appropriate (thiol-free) peptide cleavage cocktail (UNIT 18.5). Purify the product by standard methods, e.g., preparative HPLC (UNIT 8.7), gel filtration (UNIT 8.2), or ion-exchange (UNIT 8.3) chromatography, as appropriate. These methods have been applied to make intramolecular disulfides, and also to make symmetrical intermolecular disulfides.
CHARCOAL/AIR-MEDIATED INTRAMOLECULAR DISULFIDE FORMATION
ALTERNATE PROTOCOL 2
Oxygen adsorbed onto charcoal surfaces has proven efficient in mediating disulfide bond formation in a variety of peptides under basic conditions. The reactions were significantly faster than DMSO- or air-mediated cyclizations of the same substrates. Thermodynamic studies suggest that cyclization is accelerated by reduction of entropy of the peptides, upon transient adsorption to the charcoal surface, resulting in a lower activation energy (Volkmer-Engert et al., 1998). Materials Bis(thiol) peptide, previously purified 5% (v/v) aqueous NH4OH Granulated charcoal Ellman’s reagent (UNIT 18.3) Additional reagents and equipment for assaying free sulfhydryls with Ellman’s reagent (UNIT 18.3) 1. Dissolve the bis(thiol) peptide in water to a final concentration of 1 mg/ml (∼1 mM). 2. Adjust the pH to 7.5 to 8.0 with 5% aqueous NH4OH. 3. Add granulated charcoal to the peptide solution, using up to a 1:1 (w/w) ratio of charcoal to peptide.
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4. Gently shake the heterogeneous reaction mixture at 25°C. 5. Monitor the progress of the reaction by Ellman’s assay for disappearance of free sulfhydryls. Take 70-µl aliquots of the reaction mixture, dilute each with 0.7 ml of H2O and 70 µl of Ellman’s reagent, and measure the absorbance of 2-nitro-5-thiobenzoic acid anion (NTB−) at 420 nm. In general, the reaction is complete in 2 to 6 hr. See Figure 18.6.3 and UNIT 18.3 for details on assaying free sulfhydryls with Ellman’s reagent.
6. Upon completion of the reaction, filter the reaction mixture, and lyophilize the filtrate. BASIC PROTOCOL 2
INTRAMOLECULAR DISULFIDE FORMATION BY POTASSIUM FERRICYANIDE OXIDATION Efficient oxidation rates have been reported, especially in the oxytocin and somatostatin families, when using potassium ferricyanide, a relatively mild inorganic oxidizing reagent. Because K3Fe(CN)6 is slightly light sensitive, reactions are best conducted in the dark. The indicated order of addition, i.e., peptide to oxidizing agent, is designed to favor intramolecular cyclization because the concentration of free thiol species is kept minimal. Oxidation side products are possible when Met or Trp residues are present in the substrate. Materials Poly(thiol) peptide, previously purified 0.01 M phosphate buffer, pH ∼7 0.01 M aqueous K3Fe(CN)6 solution 10% (v/v) aqueous NH4OH 50% (v/v) aqueous acetic acid Celite resin (Aldrich) AG-3 anion-exchange resin Additional reagents and equipment for HPLC (UNIT 8.7) and gel-filtration (UNIT 8.3) or ion-exchange (UNIT 8.2) chromatography 1. Dissolve the poly(thiol) peptide in 0.01 M phosphate buffer, pH ∼7, to achieve a final concentration of 0.1 to 1 mg/ml (∼0.1 to 1 mM). 2. At 25°C under nitrogen, add the peptide solution slowly to a 0.01 M aqueous K3Fe(CN)6 solution (the amount of oxidant used should be in 20% excess over the calculated theoretical amount). Addition times vary between 6 and 24 hr, with purer products noted upon slower addition.
3. Keep the reaction mixture constant at pH 6.8 to 7.0, by addition of 10% aqueous NH4OH. 4. After completion of the reaction, adjust the pH to 5 with 50% aqueous acetic acid. 5. Remove ferrocyanide and ferricyanide ions by filtering first through Celite resin and then, under mild suction, through a bed of AG-3 weakly basic anion-exchange resin. 6. Purify the product by standard methods, e.g., preparative HPLC (UNIT 8.7), gel filtration (UNIT 8.2), or ion-exchange (UNIT 8.3) chromatography, as appropriate.
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INTERMOLECULAR OR INTRAMOLECULAR DISULFIDE FORMATION BY POTASSIUM FERRICYANIDE OXIDATION
ALTERNATE PROTOCOL 3
The protocol that follows differs from the preceding one principally with respect to order of addition, and it is carried out on a shorter time frame. General considerations are the same. The extent of intermolecular homodimer formation is favored by increased peptide concentration, but intramolecular cyclization is also possible if conformationally favored. For Materials see Basic Protocol 2. 1. Dissolve the poly(thiol) peptide in 0.01 M phosphate buffer, pH ∼7, to achieve a final concentration of 0.1 to 1 mg/ml. 2. At 25°C and over a 30-min period, titrate the peptide solution with 0.01 M aqueous K3Fe(CN)6 solution, until a slight yellow color persists. 3. Follow the oxidation by Ellman analysis (optional; UNIT 18.3). 4. Adjust the pH to 5 with 50% aqueous acetic acid, and remove the oxidant with AG-3 anion-exchange resin. 5. Purify the product by standard methods, e.g., preparative HPLC (UNIT 8.7), gel filtration (UNIT 8.2), or ion-exchange (UNIT 8.3) chromatography, as appropriate. ON-RESIN DISULFIDE FORMATION BY POTASSIUM FERRICYANIDE OXIDATION
ALTERNATE PROTOCOL 4
Following the same general motivation explained in Alternate Protocol 1, resin-bound thiols can, under favorable circumstances, be oxidized by reagents other than air, e.g., K3Fe(CN)6. Materials Protected peptide-resin, prepared by optimized linear SPPS chain assembly Dimethylformamide (DMF) 0.1 to 0.5 M K3Fe(CN)6 solution Dichloromethane (CH2Cl2) Appropriate (thiol-free) peptide cleavage cocktail (UNIT 18.5), and materials for workup 2-ml plastic syringe fitted with a polypropylene frit (use larger syringe for larger amounts of resin) Additional reagents and equipment for HPLC (UNIT 8.7) and gel-filtration (UNIT 8.3) or ion-exchange (UNIT 8.2) chromatography 1. Place the protected peptide-resin into a 2-ml plastic syringe fitted with a polypropylene frit, and carry out a selective deprotection step that removes protection on Cys while not affecting the anchoring linkage. For more details, see the corresponding section of Alternate Protocol 1. At the end of the process, swell the peptide-resin in DMF, and then drain the mixture. 2. Incubate the swollen peptide-resin with a homogeneous mixture of 0.1 to 0.5 M aqueous K3Fe(CN)6 with DMF (1:1 to 1:10 v/v ratio), at 25°C in the dark. The overall amount of K3Fe(CN)6 used is calculated to be 10 to 20 equivalents with respect to the level of SH groups on the peptide-resin, and the higher concentration of oxidant in the aqueous solution requires correspondingly less of the DMF co-solvent.
3. Upon completion of the incubation (12 to 24 hr reaction), wash the peptide-resin with water, DMF, and CH2Cl2.
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4. Cleave the peptide from the resin with the appropriate (thiol-free) peptide cleavage cocktail (UNIT 18.5) and materials. 5. Purify the product by standard methods, e.g., preparative HPLC (UNIT 8.7), gel filtration (UNIT 8.2), or ion-exchange (UNIT 8.3) chromatography, as appropriate. BASIC PROTOCOL 3
OXIDATION UNDER SLIGHTLY ACIDIC pH CONDITIONS BY DMSO Dimethylsulfoxide (DMSO)-promoted oxidation of thiols is efficient over an extended pH range, i.e., pH 3 to 8. The oxidant is miscible with water, so concentrations of up to 20% (v/v) can be achieved and the higher concentrations in turn promote faster reactions. Additional advantages cited for this approach (Tam et al., 1991) are (1) improved peptide solubility; (2) the effect of DMSO as a denaturing co-solvent; and (3) compatibility with side-chains susceptible to oxidation (Met, Trp, or Tyr). A limiting factor, however, is the difficulty in removal of DMSO upon completion of the oxidation. Materials Poly(thiol) peptide, previously purified 5% (v/v) aqueous acetic acid Ammonium carbonate, (NH4)2CO3 Dimethylsulfoxide (DMSO) 0.05% (v/v) trifluoroacetic acid (TFA)/5% (v/v) aqueous acetonitrile Additional reagents and equipment for HPLC (UNIT 8.7) 1. Dissolve the poly(thiol) peptide in 5% aqueous acetic acid to a final concentration of 0.5 to 1.6 mg/ml (∼0.5 to 1.6 mM). 2. Adjust pH to 6 with (NH4)2CO3. 3. Add DMSO (10% to 20% by volume) to the peptide solution. 4. Monitor progress of the reaction by analytical HPLC (UNIT 8.7); withdraw small aliquots (e.g., 20 µl to fill an HPLC injection loop of the same size) to carry out the analysis. The reaction time can vary from 5 to 24 hr.
5. Upon completion of the reaction, dilute the reaction mixture two-fold with 0.05% TFA/5% aqueous acetonitrile. 6. Load onto a preparative reversed-phase HPLC column (UNIT 8.7) for DMSO removal and product purification. ALTERNATE PROTOCOL 5
OXIDATION UNDER SLIGHTLY BASIC pH CONDITIONS BY DMSO Assuming that the peptide is soluble, it is often advantageous to carry out oxidation by DMSO at higher pH, because the intrinsically higher rate means less reagent is required and the final purification becomes correspondingly simpler. Materials Poly(thiol) peptide, previously purified 0.01 M phosphate buffer, pH 7.5 Dimethylsulfoxide (DMSO) Additional reagents and equipment for HPLC (UNIT 8.7)
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1. Dissolve the poly(thiol) peptide to a final concentration of ∼1 mg/ml (∼1 mM) in 0.01 M phosphate buffer, pH 7.5. 2. Add DMSO (1% by volume), at 25°C. 3. Monitor progress of the reaction by analytical HPLC (UNIT 8.7); withdraw small aliquots (e.g., 20 µl to fill an HPLC injection loop of the same size) to carry out the analysis. The reaction time can vary from 3 to 7 hr.
4. Quench the reaction by lyophilization. OXIDATION BY REDOX BUFFERS Peptides and proteins that contain multiple disulfide bonds can be oxidized in the presence of mixtures of low molecular weight disulfides and the corresponding free thiols. This approach may provide better yields and regioselectivities than straightforward air oxidations covered earlier (see Basic Protocol 1), because the mechanism changes from direct oxidation (free radical intermediates) to thiol-disulfide exchange (thiolate intermediate), which facilitates the reshuffling of incorrectly paired disulfides to the natural arrangements. The most commonly used systems are mixtures of oxidized and reduced glutathione, cysteine, cysteamine, or 2-mercaptoethanol (also see APPENDIX 3A). As with all solution oxidations, yields are improved under high dilution conditions that reduce the potential levels of oligomerization.
BASIC PROTOCOL 4
Materials Poly(thiol) peptide, previously purified Glutathione redox buffer (see recipe) EDTA Sephadex G-10 or G-25 column Additional reagents and equipment for dialysis (UNIT 4.4 and APPENDIX 3B), HPLC (UNIT 8.7), and gel-filtration (UNIT 8.3) or ion-exchange (UNIT 8.2) chromatography 1. Dissolve the poly(thiol) peptide in glutathione redox buffer, which contains 1 mM EDTA, to a final concentration of 0.05 to 0.1 mg/ml (∼0.05 to 0.1 mM). 2. Incubate the solution at 25°C to 35°C. 3. Monitor progress of the reaction by analytical HPLC (UNIT 8.7); withdraw small aliquots (e.g., 20 µl to fill an HPLC injection loop of the same size) to carry out the analysis. The reaction time can vary from 16 hr to 2 days.
4. Upon completion of the reaction, concentrate the peptide material by direct lyophilization. Alternatively, first quench the reaction by acidification, and perform dialysis to exchange out salts.
5. Purify the product by gel filtration on a Sephadex G-10 or G-25 column or by other standard methods, e.g., preparative HPLC (UNIT 8.7) or ion-exchange (UNIT 8.3) chromatography, as appropriate. In cases where formation of physical aggregates competes with the oxidation process, addition of a nondenaturing chaotropic agent is recommended (1 to 2 M guanidine⋅HCl or urea; also see APPENDIX 3A). If intractable precipitates form during this procedure, an alternative folding/oxidation is recommended in which the peptide is oxidized against a series of redox buffers with a slow pH gradient from 2.2 to 8.0.
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BASIC PROTOCOL 5
OXIDATION MEDIATED BY SOLID-PHASE ELLMAN’S REAGENT Ellman’s reagent, 5,5′-dithiobis(2-nitrobenzoic acid), when bound through two sites to a suitable solid-support (PEG-PS or modified Sephadex), is an effective and mild oxidizing reagent that promotes the formation of intramolecular disulfide bridges (Annis et al., 1998; Figure 18.6.3). The likely mechanism for this process involves a key resin-bound intermediate (Fig. 18.6.3, middle structure) to which pseudo-dilution applies; dimer formation should therefore be reduced, and indeed this is borne out experimentally. Oxidations are efficient under a wide pH range, from 2.7 to 6.6, with yields and rates increasing with higher pHs. The main side reaction involves covalent adsorption of the substrate to the support through intermolecular disulfide bridges; this diminishes the yield but does not affect the purity of the desired intramolecular disulfide that is isolated. Moreover the thiol substrate can be recovered by reduction, and recycled through the oxidation process. Materials Poly(thiol) peptide, previously purified 2:1:1 (v/v/v) suitable buffer (pH 2.7-7.0)/acetonitrile/CH3OH Solid-phase Ellman’s reagent (Annis et al., 1998) Dichloromethane (CH2Cl2) Dimethylformamide (DMF) 50-ml plastic syringe fitted with a polypropylene frit (use larger syringe for larger amounts of resin) Septum or plastic lock cap Additional reagents and equipment for HPLC (UNIT 8.7) 1. Dissolve the poly(thiol) peptide to a final concentration of ∼1 mg/ml (∼1 mM) in a 2:1:1 mixture of buffer (pH 2.7 to 7.0)/acetonitrile/CH3OH. 2. Weigh the solid-phase Ellman’s reagent (e.g., ∼50 mg resin/ml peptide solution, at 0.2 mmol/g, which corresponds to a 15-fold excess of DTNB functions over thiol groups) into a 50-ml plastic syringe fitted with a polypropylene frit. 3. Swell the resin in CH2Cl2, wash with DMF, wash again with CH2Cl2, and drain. The above steps apply when the parent support is PEG-PS. In the case of Sephadex as the parent support, only DMF washes are necessary prior to draining.
4. Plug the syringe tip using a small septum or a plastic lock cap. 5. Add the solution of peptide substrate to the syringe already containing the solid-phase Ellman’s reagent. 6. Gently stir the reaction mixture magnetically, or gently agitate on a rotary shaker, at 25°C. 7. Monitor progress of the reaction by taking aliquots (e.g., 20 µl to fill an HPLC injection loop) from the liquid phase for HPLC analysis (UNIT 8.7). Reaction times vary from 0.5 to 30 hr depending on the buffer pH, solid support, and ease of oxidation, with significantly faster rates at more basic pH, and when PEG-PS is the solid support.
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NO2 C NH SH
S
SH
+
S
O R O C NH
“capture” NO2
NO2
C NH SH
S
O
HS
O
S
R C NH NO2
“cyclization”
NO2 C NH
S
S
+
HS HS
O R O C NH NO2
Figure 18.6.3 Proposed mechanism for intramolecular disulfide formation mediated by solidphase Ellman’s reagents. R = solid supports. Reprinted with the permission of the American Chemical Society (Annis et al., 1998).
8. Upon completion of oxidation, drain the liquid phase into a vial, using positive air pressure. 9. Wash the resin with the 2:1:1 mixture of buffer (pH 2.7 to 7.0)/acetonitrile/CH3OH, and combine the washes with the drained liquid phase from the previous step. 10. Lyophilize the reaction mixture to obtain the oxidized peptide. No additional purification of the oxidized peptide is required, if the original poly(thiol)peptide was pure.
SIMULTANEOUS DEPROTECTION/OXIDATION WITH IODINE Iodine-mediated oxidation has been applied in conjunction with S-Xan, S-Tmob, S-Mmt, S-Trt, and S-Acm protection. The choice of solvent is critical, and optimal solvents include 80% (v/v) acetic acid, 80% (v/v) methanol, or 80% (v/v) DMF in water. It is advisable to monitor reactions (e.g., by HPLC); once oxidation is complete, quenching should be carried out to lessen the extent of overoxidation of the thiol functionality to the corresponding sulfonic acid, as well as to prevent or minimize modification of other sensitive amino acid side-chains (Tyr, Met, Trp).
BASIC PROTOCOL 6
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Materials S-protected peptide (protected by S-Acm, S-Xan, S-Tmob, or S-Trt), previously purified 80% (v/v) acetic acid Iodine Carbon tetrachloride (CCl4) Additional reagents and equipment for HPLC (UNIT 8.7) and gel-filtration (UNIT 8.3) or ion-exchange (UNIT 8.2) chromatography 1. Dissolve the S-protected peptide in 80% acetic acid to a final concentration of ∼2 mg/ml (∼2 mM). 2. Add solid iodine (5 equivalents with respect to S-protected Cys) in one portion to the solution. Mix the solution vigorously at 25°C. 3. Monitor progress of the reaction by analytical HPLC (UNIT 8.7); withdraw small aliquots (e.g., 20 µl to fill an HPLC injection loop of the same size) to carry out the analysis. The reaction time can vary from 10 min to 24 hr.
4. Upon completion of oxidation, quench the reaction by diluting to two times the volume with water. 5. Extract with CCl4 (4 to 5 times, equal volume each time) to remove the iodine, and retain the upper, aqueous phase. Upon completion of the extractions, lyophilize the aqueous phase. 6. Purify the product by standard methods, e.g., preparative HPLC (UNIT 8.7), gel filtration (UNIT 8.2), or ion-exchange (UNIT 8.3) chromatography, as appropriate. ALTERNATE PROTOCOL 6
SIMULTANEOUS ON-RESIN DEPROTECTION/OXIDATION OF S-ACM WITH IODINE The direct iodine-mediated oxidation of S-protected cysteine residues can sometimes be carried out effectively while the peptide, with its essentially full complement of protecting groups, remains on the support. A wide range of anchoring linkages and non-cysteine side-chain protecting groups are compatible in the sense that they are not cleaved prematurely under the relatively mild reaction conditions. The disulfide formation chemistry is subject to the same advantages and caveats of other solid-phase procedures covered in this unit; ultimate cleavage/deprotection from the support will provide a mixture of monomeric (desired), dimeric, and oligomeric products. Materials S-protected peptide resin (protected by S-Acm, S-Xan, S-Tmob, or S-Trt, prepared by optimized linear SPPS chain assembly) Dimethylformamide (DMF) Iodine Suitable peptide cleavage cocktail Additional reagents and equipment for for dialysis (UNIT 4.4 and APPENDIX 3B), HPLC (UNIT 8.7), and gel-filtration (UNIT 8.3) or ion-exchange (UNIT 8.2) chromatography
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1. Swell the S-protected peptide-resin in DMF. 2. Add solid iodine (5 to 10 equivalents with respect to S-protected Cys). Stir gently for 1 to 4 hr at 25°C. 3. Drain, and wash the peptide-resin with DMF and CH2Cl2. 4. Cleave the peptide with suitable peptide cleavage cocktail, avoiding the use of thiol scavengers. 5. Purify the product by standard methods, e.g., preparative HPLC (UNIT 8.7), gel filtration (UNIT 8.2), or ion-exchange (UNIT 8.3) chromatography, as appropriate. SIMULTANEOUS DEPROTECTION/OXIDATION OF S-ACM WITH Tl(III) Thallium (III) trifluoroacetate [Tl(tfa)3] is a mild oxidant that can be used in conjunction with S-Acm and S-Tmob protection. For certain specific applications, this approach has provided better yields and purities of desired disulfide products with respect to methods using I2. However the high toxicity of Tl(tfa)3, and its difficult removal from sulfur-containing peptides, represent limitations. The optimal solvent for reasons both of solubilization and chemistry is TFA, but DMF is an acceptable solvent for on-resin oxidations in conjunction with TFA-labile anchoring linkages. Anisole should be added as a scavenger for the alkyl cations generated during the deprotection/oxidation. His and Tyr survive exposure to Tl(tfa)3, but Met and Trp are compatible only if protected.
BASIC PROTOCOL 7
Materials S-protected peptide (protected by S-Acm, S-Xan, S-Tmob, or S-Trt), previously purified 19:1 (v/v) trifluoroacetic acid (TFA)/anisole Thallium (III) trifluoroacetate [Tl(tfa)3] Ethyl ether 10 to 20-ml screw-cap centrifuge tube Additional reagents and equipment for HPLC (UNIT 8.7) 1. Dissolve the S-protected peptide in 19:1 TFA/anisole to a final concentration of ∼1 mg/ml (∼1 mM), in a 10 to 20-ml screw-cap centrifuge tube. Chill to 4°C. 2. Add solid Tl(tfa)3 (0.6 equivalents per S-protected Cys). Stir the reaction for 5 to 18 hr at 4°C. 3. Optional: Monitor progress of the reaction by analytical HPLC (UNIT 8.7); withdraw small aliquots (e.g., 20 µl to fill an HPLC injection loop of the same size) to carry out the analysis. 4. Evaporate as much of the TFA as possible, under positive nitrogen flow, and add ethyl ether (∼3.5 ml per 1 µmol peptide) to precipitate the peptide. 5. Triturate for 2 min. Collect the peptide by centrifuging for 2 min at 4000 rpm in a standard benchtop clinical centrifuge, room temperature. 6. Decant the ethyl ether. Repeat the trituration/centrifugation cycle two more times to ensure complete removal of the toxic Tl salts. CAUTION: Thallium is very toxic. Handle it in the hood; use of gloves is strongly recommended. Solutions must be disposed of according to institutional guidelines. Preparation and Handling of Peptides
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ALTERNATE PROTOCOL 7
ON-RESIN SIMULTANEOUS DEPROTECTION/OXIDATION OF S-ACM WITH Tl(III) The on-resin Tl(tfa)3 procedure represents an efficient and often preferred method for creating disulfide bonds. The general considerations from the standpoint of the reagent have already been covered (see Basic Protocol 7), and relevent discussions of solid-phase methodologies are found in Alternate Protocols 1 and 6. Materials S-protected peptide-resin (protected by S-Acm, S-Tmob, prepared by optimized linear SPPS chain assembly) Thallium (III) trifluoroacetate (Tl(tfa)3) Dimethylformamide (DMF) Anisole Dichloromethane (CH2Cl2) Suitable (thiol-free) peptide cleavage cocktail (UNIT 18.5) 2-ml plastic syringe fitted with a polypropylene frit Additional reagents and equipment for for dialysis (UNIT 4.4 and APPENDIX 3B), HPLC (UNIT 8.7), and gel-filtration (UNIT 8.3) or ion-exchange (UNIT 8.2) chromatography 1. Swell the S-protected peptide-resin in DMF in a 2-ml plastic syringe fitted with a polypropylene frit. 2. Add Tl(tfa)3 (1.5 to 2 equivalents per S-protected group) in 19:1 DMF/anisole (∼0.35 ml/25 mg resin). Gently stir the heterogeneous mixture at 4°C. 3. Optional: Monitor progress of the reaction by cleavage of small portion of the peptide-resin and subjecting to HPLC analysis (UNIT 8.7). The reaction time can vary from 1 to 18 hr.
4. Drain, and wash the peptide-resin with DMF and CH2Cl2 to remove the excess Tl reagent. CAUTION: Thallium is very toxic, handle it in the hood. Use of gloves is strongly recommended. Solutions must be disposed of according to institutional guidelines.
5. Cleave the peptide with suitable thiol-free peptide cleavage cocktail (UNIT 18.5). Avoid the use of thiol scavengers.
6. Purify the product by standard methods, e.g., preparative HPLC (UNIT 8.7), gel filtration (UNIT 8.2), or ion-exchange (UNIT 8.3) chromatography, as appropriate. BASIC PROTOCOL 8
ALKYLTRICHLOROSILANE-SULFOXIDE OXIDATION A harsher oxidizing milieu uses mixtures of alkyltrichlorosilanes and sulfoxides to form disulfide bridges directly from the linear substrates protected with a variety of S-protecting groups (e.g., S-Acm, S-tBu, S-Mob, S-Meb). The reactions are fast, and are reported to be compatible with pre-existing disulfides formed in an orthogonal scheme. The protecting groups present in the substrate determine the optimal chlorosilane-sulfoxide combinations, but the most generally effective milieus apply diphenylsulfoxide [Ph(SO)Ph] with either CH3SiCl3 or SiCl4. The method is compatible with most amino acid side-chains, although Trp must be protected as its Nin-formyl derivative to prevent chlorination of the side-chain indole.
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Materials S-protected peptide (protected by S-Acm, S-tBu, S-Mob, or S-Meb), previously purified Trifluoroacetic acid (TFA) Diphenylsulfoxide (Ph(SO)Ph) Trichloromethylsilane (CH3SiCl3) Anisole Ammonium fluoride (NH4F) Diethyl ether Sephadex G-15 column 4 N aqueous acetic acid Additional reagents and equipment for HPLC (UNIT 8.7) and gel-filtration (UNIT 8.3) or ion-exchange (UNIT 8.2) chromatography 1. Dissolve the S-protected peptide in TFA to a final concentration of 1 to 10 µg/ml (∼1 to 10 µM). 2. Add Ph(SO)Ph (10 equiv), CH3SiCl3 (100 to 250 equiv), and anisole (100 equiv) to the solution. Allow the reaction to proceed for 10 to 30 min at 25°C. 3. Quench the reaction by addition of solid NH4F (300 equiv). 4. Precipitate the crude product with a large excess of dry diethyl ether. 5. Isolate the crude product by gel filtration on Sephadex G-15 column eluting with 4 N aqueous acetic acid. 6. Lyophilize the peptide fraction, and purify the product by standard methods, e.g., preparative HPLC (UNIT 8.7) or ion-exchange (UNIT 8.3) chromatography, as appropriate. REAGENTS AND SOLUTIONS Glutathione redox buffer Prepare a 0.1 M Tris⋅Cl or Tris⋅acetate buffer at pH 7.7 to 8.7. Add both reduced (1 to 10 mM) and oxidized (0.1 to 1.0 mM) glutathione to the buffer, with the molar ratio of reduced to oxidized glutathione typically 10:1. Alternative redox systems include: cysteine/cystine, cysteamine/cystamine, 2-mercaptoethanol/2-hydroxyethyl disulfide, dithiothreitol (DTT)/1,2-dithiane-4,5-diol (oxidized DTT). The same general procedure should be used.
COMMENTARY Background Information In natural or artificial peptides and proteins, the ease of intramolecular disulfide formation evidently depends on conformational factors, and is tied to questions of what are the driving forces (kinetic versus thermodynamic) and mechanisms (direct oxidation versus disulfide exchange) of tertiary structure acquisition in vivo and in vitro. Whereas the oxidation of simple ω-dithiols to monomeric cyclic disulfides is optimal when six-membered rings can form, the heterodetic disulfide rings found in a range of biological substances are of size 3n +
8, where n is the number of residues between a given pair of half-cystines and can start at 0 (relatively rare, because of conformational strain), and continues to 1, 2 (e.g., thioredoxin active-site loop), 3, 4, and higher. For n = 4, the conformationally flexible 20-membered ring occurs in oxytocin, prothrombin, and various toxin families. The smaller “local” loops are typically associated with β- and γ-turns; assumption of these secondary structures is possible only with certain intervening residues (e.g., Gly, Pro, Ser, Asn). The proper pairing of two particular cysteines that are relatively far
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apart in the linear sequence becomes possible if the corresponding folded conformation is energetically favorable, somewhat flexible, and readily accessible. The environment (steric; neighboring charges; buried versus surface) about the cysteine residues affects pKa values, and consequently the ease with which they oxidize. The rates, yields, and specificities of oxidations can be influenced as well by the reagent used, the solvent for the reaction, the presence of denaturants, and trace metals that act as catalysts. There exists a substantial empirical base of knowledge on a number of these issues, but as of yet, few clear correlations with predictive power have been drawn. Intermolecular disulfides can be established by careful co-oxidation, an approach that is often limited by the formation of homodimers along with the desired heterodimers. “Directed” methods are more general, but they are also more demanding in terms of requirements for a range of selectively removable sulfhydryl protecting groups, and the need to avoid conditions that give rise to disproportionation of unsymmetrical disulfides to the symmetric species. The issues just raised increase in complexity when more than two half-cystine residues are involved, since in principle mispairing can occur. In vitro renaturation studies are clearly predicated on the assumption that the “correct” structure (as judged by proper pairing of halfcystines) will either form preferentially or be the major species after equilibration (perhaps, as promoted in thiol-disulfide “redox” buffers or by the presence of protein factors such as disulfide isomerases or thioredoxin). While this ideal can often be achieved in practice with careful attention to experimental conditions, e.g., pH, ionic strength, temperature, time, and protein concentration, it is also common (particularly with mutant materials prepared by recombinant techniques) for substantial amounts of isomers with incorrect disulfide pairing to form. Furthermore, oxidation/folding of purified synthetic materials, even carried out under high dilution, is frequently plagued by the production of significant levels of dimeric, oligomeric, and intractable polymeric byproducts. This problem is exacerbated by the difficulty in “recycling” nonmonomeric species through alternating reduction and reoxidation steps. A central conceit of this unit is that disulfide formation can be controlled to some extent by astute choices of chemical procedures. Sulfursulfur bonds are created most often by oxidation (using air or stronger reagents) of precur-
sors with free or protected sulfhydryls. Chemistries for unsymmetrical “directed” formation of sulfur-sulfur bonds can be applied as well (the latter is not specifically covered in this unit). In general, the more complex peptide targets have been obtained in relatively low yields, only after extensive optimization of experimental protocols for synthesis, purification, and oxidation/folding. The details of how to protect cysteine, and the corresponding deblocking conditions, are central issues. Unlike the case with deprotection from most other side-chains, where the parent functionality is recovered, removal of protecting groups from cysteine can yield the corresponding free thiol, a thiolate metal salt (mercaptide), or a disulfide, depending on the reagent(s) applied and the precise reaction conditions. A significant avenue for current research is to achieve some measure of regioselectivity in cysteine pairing by applying appropriate pairwise combinations of orthogonally removable Cys protecting groups.
Critical Parameters and Troubleshooting Experimentalists seeking to obtain disulfide-containing peptides are faced with a set of options that might be considered both daunting and reassuring. Certainly, the conformational forces related to the desired covalent structures need to be favorable. Beyond that, reactions can be conducted in solution (e.g., Basic Protocols 1 to 4, Basic Protocols 6 to 8, Alternate Protocol 3, and Alternate Protocol 5) or in the solidphase (e.g., Alternate Protocols 1, 4, 6, and 7) mode, or by a hybrid approach wherein the substrate is in solution but the oxidizing reagent is resin-bound (e.g., Basic Protocol 5 and Alternate Protocol 3); the reaction milieus can range from primarily aqueous to aqueous with organic cosolvent to fully organic (even anhydrous); the linear precursor can be the free poly(thiol) or have its Cys residues appropriately protected; and the strategy to generate the disulfide array can involve a single procedure or sequential steps. A minimum prerequisite for solution strategies is that the materials are indeed soluble throughout the processes (i.e., as linear substrate, as intermediates, and as final disulfide product), and the value of high dilution has been noted earlier. Solid-phase reactions are based on the pseudo-dilution principle and, when successful, have the added advantage that they are operationally much simpler than the corresponding solution reactions.
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Even before attacking the disulfide-forming steps, a plan must be made for construction of the linear sequence(s). This entails selection of the repetitively removable Nα-amino protecting group, choices of all of the corresponding side-chain protecting groups with particular emphasis on Cys, consideration of how to manage certain susceptible residues (e.g., Met, Tyr, Trp) that could potentially offer compatibility problems, and (in the case of SPPS), decisions on what resin support and anchoring handle to use, and the appropriate cleavage conditions. Reasonable guidelines and precedents for all of this can either be inferred from the text in this unit, or are covered in some of the Key References as well as the Literature Cited. Some of the initial synthetic work can even be contracted out. The next phase of the work depends on whether or not any of the disulfides will be formed by solid-phase reactions. If yes, parameters that may impact the outcome include: solvent; acidity/basicity; reaction temperature and time; and the loading, morphology, and swelling of the solid support. Later, the anchoring linkage is cleaved to release the oxidized material into solution for evaluation, additional chemical steps (sometimes), and eventual purification (under all circumstances). When disulfide-forming steps are to be carried out in solution, a myriad of experimental details may be critical, as has been described throughout this unit. Starting with purified precursors and intermediates can often make a significant difference, and in the cases that rely on oxidations of the corresponding poly(thiols), it is often imperative that such species have been pretreated with suitable reducing agents in order to ensure the absence of interfering species that contain prematurely formed disulfides (UNIT 18.3). In aqueous solution, oxidations generally proceed more rapidly with increased pH (e.g., pH 7.5 to 8.5) because of the greater level of ionization to the thiolate, but these conditions can also promote disulfide scrambling. Methods to form disulfides that are effective and reliable at lower pH (e.g., pH 2.5 to 6.0) are therefore of considerable interest. Again, because the facility of disulfide formation hinges on polypeptide conformation, changes in ionic strength and addition of denaturants can sometimes have a salutary effect. Most of the general principles raised throughout here can be extrapolated to protocols that rely on oxidizing agents other than oxygen, that are conducted in media other than water, and/or that do not involve free thiol intermediates.
As the reactions to form disulfides are underway, particularly as part of exploratory studies, conversion of precursors to products should be monitored chromatographically in order to determine the endpoints. This is most straightforward for solution processes that start with poly(thiol) precursors, and the information gleaned can be reinforced by following the disappearance of sulfhydryl titer, e.g., by Ellman’s assay. Monitoring solid-phase oxidations is somewhat more difficult, since generally the peptide-resin must be sampled, then subjected to cleavage, and finally analyzed to determine the distribution of released materials (corresponding to starting, intermediates, and product that were originally resin-bound). When working with synthetic disulfidecontaining peptides, it is often helpful to provide a full accounting of the molecular species formed. This general objective covers the isolation and structural characterization of both the target and any significant byproducts (e.g., parallel or antiparallel dimers, oligomers, and variants with misaligned disulfides). At a minimum, one should quantify components of crude synthetic products by comparison of HPLC peak areas with those of purified standards (if available) of known concentration. Also, analytical gel filtration gives information about the relative levels of monomers versus species of greater size. In order to assign and/or confirm structures, standard analytical methods for characterizing peptides, e.g., amino acid analysis, analytical HPLC, and (most informative) any of a number of mild ionization mass spectrometric methods, are supplemented by techniques adapted directly to study the disulfide arrangements such as proteolysis/peptide mapping or partial reduction/peralkylation/sequencing procedures. If after following the aforementioned guidelines for synthesis, purification, and characterization, the desired disulfide-containing structures are not obtained in sufficiently satisfactory overall yields and purities, an iterative troubleshooting process can be followed. This can range from further optimization of specific reaction conditions to trying alternative protocols to a partial or complete overhaul of the essential strategy. Literature precedents suggest that most synthetic peptides with a single or two disulfide bonds can eventually be obtained by effective routes, but the outcome for more complicated targets is by no means assured and quite dependent on the experimental art of the investigator.
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Anticipated Results As indicated in earlier sections of this unit, there are no definitive rules that guarantee success, and several of the described protocols may need to be evaluated individually or in tandem in order to converge upon optimal results. In calculations based on 100% being a homogeneous linear precursor, either resin-bound or in solution, many of the disulfide-forming reactions under review can proceed in yields of ≥80% (turnover of starting material can generally be taken to completion, but the required disulfide will not necessarily be the exclusive product). After an efficient purification step, the overall isolated yield usually drops to the 40% to 60% range. Clearly, when multiple reactions and purifications are carried out, the overall yields will be lower.
Time Considerations
Disulfide Bond Formation in Peptides
Formation of the disulfide bond(s) in synthetic peptides is integrated with other steps, starting from linear chain assembly and including one or more purification steps. The following description assumes that pilot experiments and optimizations specific to the desired target have already been completed. Corresponding to those cases where a particular chemical protocol is especially efficient, valuable time can be saved by skipping an intermediate purification. Solid-phase synthesis of the linear sequences can be completed within several days, and is accelerated somewhat when using an automated synthesizer. Cleavage and workup procedures require several hours, and half a day of effort should be budgeted for each purification. Processing of purified samples by lyophilization can take 1 to 2 days, depending on the volume. The focus of this unit is on disulfide formation, and the individual protocols should be referred to in order to learn the recommended times that often suffice to achieve complete reaction. Potentially the most time-consuming procedures are those in which a linear poly(thiol) precursor is oxidized in solution, using aeration, a redox buffer, or the solidphase Ellman’s reagent. For these cases, reaction times of 1 to 3 days are typical. Accelerated rates may be achieved when applying specific oxidizing agents such as ferricyanide or DMSO. Methods that start from S-protected precursors and involve disulfide formation concurrent with deprotection are invariably more rapid, and are often complete in ≤4 hr (sometimes, as short as 10 min). The majority of
solid-phase reactions under review are relatively fast as well.
Literature Cited Albericio, F., Hammer, R.P., García-Echeverría, C., Molins, M.A., Chang, J.L., Munson, M.C., Pons, M., Giralt, E., and Barany, G. 1991. Cyclization of disulfide-containing peptides in solid-phase synthesis. Int. J. Peptide Protein Res. 37:402413. Annis, I., Chen, L., and Barany, G. 1998. Novel solid-phase reagents for facile formation of intramolecular disulfide bridges in peptides under mild conditions. J. Am. Chem. Soc. 120:72267238. Eritja, R., Ziehler-Martin, J.P., Walker, P.A., Lee, T.D., Legesse, K., Albericio, F., and Kaplan, B.E. 1987. On the use of S-t-butylsulphenyl group for protection of cysteine in solid-phase peptide synthesis using Fmoc-amino acids. Tetrahedron 43:2675-2680. Hargittai, B. and Barany, G. 1999. Controlled syntheses of natural and disulfide-mispaired regioisomers of α-conotoxin SI. J. Peptide Res. 54:468-479. Munson, M.C. and Barany, G. 1993. Synthesis of α-conotoxin SI, a bicyclic tridecapeptide amide with two disulfide bridges: Illustration of novel protection schemes and oxidation strategies. J. Am. Chem. Soc. 115:10203-10210. Munson, M.C., Lebl, M., Slaninová, J., and Barany, G. 1993. Solid-phase synthesis and biological activity of the parallel dimer of deamino-oxytocin. Peptide Res. 6:155-159. Tam, J.P. and Shen, Z.-Y. 1992. Efficient approach to synthesis of two-chain asymmetric cysteine analogs of receptor-binding region of transforming growth factor-α. Int. J. Peptide Protein Res. 39:464-471. Tam, J.P., Wu, C.-R., Liu, W., and Zhang, J.-W. 1991. Disulfide bond formation in peptides by dimethyl sulfoxide. Scope and applications. J. Am. Chem. Soc. 113:6657-6662. Volkmer-Engert, R., Landgraf, C., and SchneiderMergener, J. 1998. Charcoal surface-assisted catalysis of intramolecular disulfide bond formation in peptides. J. Peptide Res. 51:365-369.
Key References Albericio, F., Annis, I., Royo, M., and Barany, G. 2000. Preparation and handling of peptides containing methionine and cysteine. In Fmoc Solid Phase Peptide Synthesis: A Practical Approach (W.C. Chan and P.D. White, eds.) pp. 77-114. Oxford University Press, Oxford. Relatively recent coverage of cysteine protection and disulfide formation, in the context of a 14-chapter monograph on Fmoc solid-phase methodology.
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Andreu, D., Albericio, F., Solé, N.A., Munson, M.C., Ferrer, M., and Barany, G. 1994. Formation of disulfide bonds in synthetic peptides and proteins. In Methods in Molecular Biology, Vol. 35: Peptide Synthesis Protocols (M.W. Pennington and B.M. Dunn, eds.) pp. 91-169. Humana Press, Totowa, N.J. State-of-the-art review at the time that it was written. Some experimental information and 358 literature citations. Annis, I., Hargittai, B., and Barany, G. 1997. Disulfide bond formation in peptides. Methods Enzymol. 289:198-221. Another book chapter, more recent but less comprehensive than the Andreu et al. reference above. The parent volume, edited by G.B. Fields, is a highly useful 32-chapter compendium of methods of solidphase peptide synthesis.
Contributed by Lin Chen AxCell Biosciences Corporation Newtown, Pennsylvania Ioana Annis Union Carbide Corporation Bound Brook, New Jersey George Barany University of Minnesota Minneapolis, Minnesota
Preparation and Handling of Peptides
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Guide for Resin and Linker Selection in Solid-Phase Peptide Synthesis
UNIT 18.7
The past decade has seen the chemistry of solid-phase peptide synthesis (SPPS) mature considerably. What was once a technique solely accessible to specialized laboratories has now become routine practice for chemists and biologists alike who have no specific training in peptide chemistry per se. For the first ∼25 years (beginning with Merrifield, 1963), the majority of SPPS efforts employed Boc/benzyl chemistry, which is based on a system of graduated acid lability. In Boc/benzyl-based syntheses, trifluoroacetic acid (TFA) is used for iterative removal of the t-butyloxycarbonyl (Boc) protecting group from the α-amino (Nα ) functionality, while side chain protecting groups and the peptide-resin anchorage are simultaneously cleaved using a much stronger acid, anhydrous hydrogen fluoride (HF). As investigators sought milder methods for SPPS that avoided the use of HF for the final deprotection step, the Fmoc/t-butyl approach rose in popularity. This methodology differs from Boc/benzyl chemistry in that Nα protection and side-chain protection are based on a system of orthogonality—i.e., the selective removal of each type of protection occurs by a mechanism that leaves the other type of protection entirely intact. In Fmoc/t-butyl chemistry, piperidine is used for iterative removal of the Nα 9-fluorenylmethoxycarbonyl (Fmoc) protecting group, while side chain protecting groups and the peptide-resin anchorage are simultaneously cleaved using TFA. While many monographs have been written on the theory and practice of SPPS, these discussions have almost invariably splintered into camps centered around the Boc/benzyl methodology or the Fmoc/t-butyl methodology, with little to no interplay between the two. While the chemical minutiae of these two approaches are ostensibly at odds, the development of linkers and resins for Boc-based SPPS and Fmoc-based SPPS have followed the same chemical trends. It is noteworthy that some of the most intriguing applications of SPPS have bridged the gap between these two strategies. For instance, peptide nucleic acids (PNAs), which hold promise in drug delivery and antisense technologies, have brought the concept of orthogonality between side-chain protection and Nα protection, hitherto a hallmark of the Fmoc approach, to Boc chemistry. Conversely, the synthesis of glycopeptides and fluorescence resonance energy transfer (FRET) substrates using Fmoc chemistry requires the application of side-chain protection that is removable under conditions of graduated acid lability, which is the basis of Boc chemistry. To both novice and more experienced practitioners of SPPS, the vast selection of commercially available linkers and resins has become something of a babel. The goal of this unit is to clarify the situation, which is best understood by distillation to first principles, through an appreciation of chemical trends and consequences, as well as practical considerations. It must be emphasized that some of the philosophies and procedures described in this unit are naturally skewed in accordance with the author’s own experience and preferences. For example, there are many satisfactory ways to couple a linker to a resin via an amide bond (although some are better than others), but only some are presented here. As the development of SPPS has involved a long series of (usually incremental) improvements upon existing methodologies, it is hoped that this unit and others will provide food for thought that might lead to further improvements in the practice of SPPS.
Peptides Contributed by Jason A. Moss Current Protocols in Protein Science (2005) 18.7.1-18.7.19 C 2005 by John Wiley & Sons, Inc. Copyright
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Figure 18.7.1 resin.
Linkage of the target peptide, linker, internal reference amino acid (IRAA), and
Linker Selection In the majority of cases, synthetic peptide targets are designed to possess C-terminal carboxylic acids or primary amides. For applications in which more exotic unnatural functionalities are desired at the C-terminus, a variety of procedures, each with different degrees of complexity, generality, and effectiveness, have been devised. Although the literature is awash with technologies that allow the preparation of most any peptide-derived target, many of these reports have been only sparingly validated and applied. Furthermore, in many cases, such techniques require significant experience in synthetic and solid-phase chemistry, as well as access to equipment not normally found in most peptide synthesis laboratories. This discussion will focus on established state-of-the-art methodologies that fulfill the needs of most peptide synthesis applications. Whenever possible, the use of preformed linkers—sometimes termed handles—prepared by solution-phase organic synthesis is preferable to the traditional strategy of functionalizing linkers onto resins via direct modification of native polystyrene. One attractive feature of preformed linkers is their compatibility with the use of an internal reference amino acid (IRAA) between the linker and the resin (Fig. 18.7.1). While perhaps unnecessary for the synthesis of smaller, more routine peptides, an IRAA allows cleavage of the scissile peptide-resin linkage to be quantitated. This is particularly useful for newcomers to Fmoc-based SPPS; if, for example, an unexplainably poor yield of isolated peptide is obtained, solid-phase amino acid analysis (AAA) of the IRAA will allow the quantitation of cleavage efficiency. In such cases, the ability to accurately troubleshoot problems may allow the cleavage to be repeated and/or may make it possible to improve cleavage protocols for use in future syntheses. A second benefit of preformed linkers is that, almost without exception, their use in conjunction with high-quality amino-functionalized resins leads to significantly improved synthetic efficiency.
Linkers yielding C-terminal carboxylic acids
Resin and Linker Selection in SPPS
Peptides bearing C-terminal carboxylic acids are almost exclusively prepared using linkers derived from substituted benzyl esters (Fig. 18.7.2A). Whereas an unsubstituted benzyl ester is quite stable to acidolysis—and therefore ideally suited to use as a peptide-resin anchorage in Boc-based SPPS—the increasing prevalence of Fmocbased SPPS has spurred the development of more acid-labile peptide-resin linkages. Benzyl esters can be decorated with various alkoxy substituents and/or aryl rings to afford increasingly acid-labile peptide-resin linkages. Because of the variation in electronic properties as a function of the location, number, and type of such substituents, a wide range of linkers with predictable, graduated acid stabilities have been developed. Although only some of these linkers have found widespread application, the elegance with which graduated acid lability can be applied to linker design has been one of the key factors leading to the widespread use of solid-phase synthesis in biomedical research involving peptides, peptide-derived natural products, proteins, and other biopolymers.
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Figure 18.7.2 Acidolytic bond scission in the synthesis of peptides bearing (A) a C-terminal carboxylic acid (C-O bond scission) and (B) a C-terminal primary amide (C-N bond scission). Examples shown are the p-alkylbenzyl ester linkage (panel A; Fig. 18.7.3, entry no. 1) and the p-methylbenzhydrylamide (MBHA) linkage (panel B; Fig. 18.7.4, entry no. 1).
The classical peptide-resin linkage employed in Boc-based SPPS is the p-alkylbenzyl ester (Fig. 18.7.3, entry no. 1; Merrifield, 1964). It is important to note that this is only formally a benzyl ester; in polystyrene resins, the 4-vinyl substituent of styrene (which is saturated upon polymerization) donates electron density to the benzyl ester linkage through the σ bond, thereby increasing the acid lability of the linkage. Nonetheless, this linkage is sufficiently stable to the repetitive TFA treatments necessary for subsequent Nα Boc deprotection cycles, but quantitatively cleaved by anhydrous HF during the final deprotection/cleavage step. Still, the unacceptable cumulative loss of peptide chains through repeated TFA-mediated Nα Boc deprotection cycles motivated the development of a more acid-stable alternative, the phenylacetamidomethyl (PAM) linker (Fig. 18.7.3, entry no. 2; Mitchell et al., 1976; Mitchell et al., 1978). The PAM linker is first prepared as a derivative of the desired C-terminal residue using a two-step solution-phase synthesis, and this preformed amino acid–PAM handle is then coupled to an aminomethyl-derivatized resin. The electron-withdrawing nature of the p-acetyl substituent on the PAM benzyl ester linkage renders the PAM linker ∼100 times more stable to TFA acidolysis than a traditional p-alkylbenzyl ester (obtained by direct esterification of a chloromethylderivatized resin). The result is a peptide-resin linkage that is sufficiently stable for Boc-based synthesis of large peptides (>50 residues), with negligible chain loss in each TFA-mediated Nα deprotection step. For the routine synthesis of peptides by Boc chemistry, the PAM linker is recommended in all cases in place of the classic p-alkylbenzyl ester obtained by direct esterification of chloromethyl-derivatized polystyrene. This recommendation stems not only from the greater acid stability of the PAM linker but also from chemical considerations regarding the alternative; a variety of deleterious side reactions in Boc-based SPPS have been ascribed to incomplete esterification reactions involving chloromethyl-derivatized polystyrene resins, and all of these side reactions are avoided by using preformed PAM linkers. It must be emphasized that the peptide bond connecting the PAM linker to the resin is not cleaved during HF deprotection/cleavage; only the ester linkage between the C-terminal amino acid residue and the PAM linker is cleaved, yielding the target peptide as a C-terminal acid. Preloaded Boc-Xaa-PAM resins are available from several commercial sources (e.g., Peptides International, Midwest Biotech), eliminating the need for the user to prepare the PAM-linked derivative of the C-terminal amino acid. Preformed Boc-Xaa-PAM-COOH derivatives are also commercially available (NeoMPS, Peptides International), allowing the user to select a preferred aminomethyl-derivatized resin for use in SPPS.
Peptides
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Figure 18.7.3 Linkers yielding peptides that bear a C-terminal carboxylic acid. Typical minimal acidolytic deprotection conditions are noted; acidolysis should be carried out at 25◦ C whenever not otherwise specified. Entries marked with an asterisk have the greatest general utility in routine peptide synthesis applications. TFA, trifluoroacetic acid; TFE, trifluoroethanol; HFIP, hexafluoroisopropanol. Resin and Linker Selection in SPPS
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Despite their incompatibility with Boc-based SPPS, TFA-labile peptide-resin linkages were found early on to be ideally suited to Fmoc-based SPPS. In Fmoc-based SPPS, the use of an orthogonal, base-labile Nα protecting group eliminates the need to use HF for the final deprotection/cleavage step. Fortunately, an appropriate linker was already in hand at the time the Fmoc approach was being developed, as a TFA-labile p-alkoxybenzyl ester had previously been developed for a somewhat sparsely applied variant of Boc chemistry that relied on greater acid lability. This substituted benzyl ester, known as the Wang linker, was prepared by a base-mediated ether coupling of 4-hydroxymethylphenol and chloromethyl-functionalized polystyrene resin (Fig. 18.7.3, entry no. 3; Wang, 1973). It is important to emphasize that the commonly used term Wang resin is a misnomer, clouding the demarcation between linker and resin. Wang-functionalized polystyrene resins are commercially available from many sources, both in free alcohol form and preloaded with Fmoc-protected amino acid residues. One significant problem inherent in the synthesis of Wang-functionalized polystyrene resins is that it is difficult to achieve clean, quantitative functionalization of the resin at all chloromethyl sites with minimal polymerization of 4-hydroxybenzyl alcohol, one of the starting materials in the process. To address this issue, the preformed 2-(4-hydroxymethyl)phenoxyacetic acid (HMPA) linker was developed (Sheppard and Williams, 1982). This linker (Fig. 18.7.3, entry no. 4) can be rapidly and quantitatively coupled to any aminomethyl-derivatized resin using standard amide coupling protocols and affords roughly the same electronic characteristics as the parent Wang linker. As the p-alkoxybenzyl alcohol is substantially more difficult to acylate than an amine, autoacylation of the unprotected benzylic alcohol during linker coupling is not a significant concern, provided that an appropriately weak combination of activation chemistry and solvent is employed (see Basic Protocol 1). An additional benefit of the HMPA linker is that TFA acidolysis of the peptide-resin anchorage cleaves only the peptide-HMPA ester linkage, such that the linker remains attached to the resin. This contrasts with the traditional Wang linker, as the Wang linker–resin ether bond, in addition to the peptide–Wang linker ester bond, is cleaved during TFA acidolysis, leading to unnecessary contamination of the cleaved peptide product. Among the TFA-labile carboxylic acid linkers, the Wang linker and HMPA handle provide the most stable peptide-resin anchorages and are therefore the most useful for chemoselective side-chain modification of the resin-bound peptide chain. Modification possibilities include the coupling of biotin, a fluorophore, or a fatty acid to a side-chain amine or alcohol, as well as the phosphorylation or glycosylation of a serine side chain. In such cases, pseudo-orthogonal trityl-based protection or truly orthogonal allyl- or reductionbased protection can be used to prevent the target functionality from reacting during chain assembly, and these protecting groups can be selectively removed in the presence of more stable t-butyl esters, ethers, and carbamates following chain assembly. (For clarification, truly orthogonal side-chain protection strategies employ deprotection mechanisms that are entirely harmless to the usual Fmoc/t-butyl side-chain protection scheme, whereas pseudo-orthogonal side chain protecting groups are removed using mild acidolytic mechanisms that often give rise to varying degrees of undesired deprotection side products.) For a peptide linked to the solid support via a p-alkoxybenzyl ester, the brief TFA (1% to 5%) treatment necessary to remove trityl protecting groups, for example, leads to negligible premature chain loss due to acidolysis of the peptide-resin anchorage. (TFA concentrations here and throughout are expressed as percent v/v in an inert solvent, usually methylene chloride.) Only at the final stage of the synthesis, after all desired side-chain modifications are incorporated, is the peptide cleaved from the resin, via treatment for 1 to 2 hr with a ∼95% TFA cleavage cocktail. Peptides
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In some cases, a cleaved peptide bearing a free C-terminal acid but having protected side chains and/or a protected N-terminus is desired. For example, peptide-derived natural products and proteins may be prepared by the controlled, iterative condensation of partially or fully protected peptide segments. Such reactions are referred to as fragment condensations and are usually employed when it would be difficult or impossible to prepare the entire target molecule using standard stepwise SPPS. To allow cleavage of the peptide-resin anchorage while preserving the integrity of side-chain and N-terminal protection, linkers with greater acid lability, which are labile to low concentrations of TFA or other acids, have been developed. Brief treatments (<5 min) with 1% to 5% TFA allow cleavage of highly acid-labile peptide-resin linkages while leaving the standard, comparatively acid-stable t-butyl-based side chain protecting groups mostly or entirely intact. The linkers shown in Figure 18.7.3, entry no. 5 to entry no. 10, are all suitable for the synthesis of protected peptides. The increased acid lability of these linkers is derived from the presence of alkoxy substituents in the key ortho- and para- position(s) in the benzyl ester linkage, or from the presence of conjugated aryl rings in the linker. Entry no. 5 depicts a polystyrene resin functionalized with 4-hydroxymethyl-3-methoxyphenol, which serves as a more acid-labile version of a Wang-functionalized polystyrene resin. Developed by Bachem for the synthesis of protected peptides, this resin is marketed under the trade name SASRIN (super-acid-sensitive resin; Mergler et al., 1988). The 2,4-bisalkoxybenzyl ester formed using this linker is ∼100 times more labile to acidolysis compared with the p-alkoxybenzyl ester linkage and therefore allows the isolation of protected peptides by cleavage with 5% TFA. Analogous to the HMPA handle, which was devised as an amide-linked surrogate for the ether-linked Wang linker, 4-(4hydroxymethyl-3-methoxyphenoxy)butyric acid (Fig. 18.7.3, entry no. 6; Floersheimer and Riniker, 1991) was developed to address the problems encountered during the functionalization of chloromethyl-derivatized polystyrene resins with 4-hydroxymethyl-3methoxyphenol. The benefits of using this linker (as a preformed handle coupled to an aminomethyl-derivatized resin) over the parent 4-hydroxymethyl-3-methoxyphenol linker (in the SASRIN resin) are identical to those that favor the use of the HMPA handle in place of the Wang linker. For applications in which an extremely high degree of acid lability is needed, the electronic properties of the 2,4-bisalkoxybenzyl ester–type linker were further fine-tuned via the incorporation of an additional methoxy substituent in the ortho- position (Fig. 18.7.3, entry no. 7). The resulting linker, 4-(4-hydroxymethyl-3,5-dimethoxyphenoxy)butyric acid, also known as the HAL (hyper-acid-labile) linker, can be coupled to aminomethylderivatized resins using standard coupling protocols, as in the case of the preformed 2,4-bisalkoxybenzyl ester handle (Albericio and Barany, 1991). Equipped with electrondonating substituents in both key ortho- positions and in the lone key para- position, this linker provides a peptide-resin linkage that is at least an order of magnitude more acidlabile than the 2,4-bisalkoxybenzyl esters, allowing fully protected peptides to be cleaved under conditions as mild as 0.1% TFA. Similarly acid-labile peptide-resin linkages can be found in Figure 18.7.3, entry no. 8 to entry no. 10. The 2-chlorotrityl linker (entry no. 8; Barlos et al., 1991a,b), the corresponding handle (Bayer et al., 1993), and the Rink acid linker (entry no. 9; Rink, 1987) are functionally interchangeable with the HAL linker. While these linkers may be slightly different in terms of their acid lability, they are functionally equivalent to the extent that peptides may be cleaved from any of them using mild organic acids such as 0.1% TFA, acetic acid, or fluorinated alcohols. (Of these three options, fluorinated alcohols are preferred, due to their volatility and their excellent solvation of protected peptides.) Resin and Linker Selection in SPPS
While the linkers depicted in Figure 18.7.3, entry no. 5 to entry no. 10, all suffice for the synthesis of protected peptide acids, cost, time, and purity considerations have elevated
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entry no. 8, the 2-chlorotrityl linker, to a role of primary importance and widespread application in SPPS. The popularity of this linker stems in part from the convenience of 2-chlorotrityl-modified polystyrene resins, which are available at relatively low cost, preloaded with side-chain-protected amino acid residues, from many commercial sources. This widespread commercial availability, in turn, is a function of the chemical superiority and general utility of the linker, not only due to its appropriate acid lability but also due to its ability to preserve the chiral integrity of the C-terminal residue. The linkers shown in Figure 18.7.3, entries no. 3 to 7 and entry no. 10, all require the use of potent activation chemistries, which are accompanied by highly undesirable side reactions (including dipeptide formation and racemization), for esterification of the relatively unreactive resin-bound benzylic alcohol. The 2-chlorotrityl linker, however, is chemically unique in that linkage of the C-terminal amino acid is performed by direct esterification of the resin-bound trityl chloride analog. Because the C-terminal carboxylic acid is used as the nucleophile in this reaction—and therefore is not activated by any of the potent methods typically used to acylate resin-bound alcohols—the danger of racemization of the C-terminal amino acid during coupling to the solid support is negligible. In light of the drastically reduced probability of racemization during the anchorage of the C-terminal amino acid to the 2-chlorotrityl-functionalized resin, peptides bearing particularly racemization-prone C-terminal residues (e.g., Cys, His) are best prepared using this linker. Additionally, the enormous steric hindrance afforded by the 2chlorotrityl ester peptide-resin anchorage minimizes the potential for premature peptide chain loss due to diketopiperazine (DKP) formation following Nα Fmoc deprotection of the resin-bound dipeptide. Prevention of this deleterious side reaction is of particular importance when a peptide target is synthesized bearing a proline residue in the C-terminal or penultimate (relative to the C-terminus) position. While the 2-chlorotrityl peptide-resin linkage is highly acid-labile and, in fact, originally designed for the synthesis of protected peptides, it may also be used to prepare unprotected peptides using a ∼95% TFA cleavage cocktail. Indeed, for the synthesis of peptides bearing racemizationprone C-terminal residues, or whenever DKP formation is foreseen to be a problem, the 2-chlorotrityl-derivatized polystyrene resin is preferred over the Wang-functionalized resin regardless of whether the preservation of side-chain and/or N-terminal protection is desired. The 2-chlorotrityl-functionalized resin (Fig. 18.7.3, entry no. 8) does not employ an amide-linked preformed handle. While the Wang linker can easily be transformed for use in handle format, it is much less straightforward to do so using the 2-chlorotrityl linker. However, this is somewhat less of a concern for several reasons. Most significantly, the recent commercial application of the 2-chlorotrityl resin for very large-scale peptide syntheses has led to significant optimization of chlorotrityl-derivatized resin production; consequently, this resin (native or preloaded) can now be obtained in a high-quality form with reliable batch-to-batch reproducibility. Nevertheless, the development of new amino-derivatized solid supports has spurred the creation of a preformed-handle version of the 2-chlorotrityl linker (Fig. 18.7.3, entry no. 9; Bayer et al., 1993). Although it possesses nearly the same chemical properties as the parent linker (Fig. 18.7.3, entry no. 8), this linker is contraindicated on the basis of cost, availability, and practicality. Whereas high-quality preloaded 2-chlorotrityl resins are widely available at relatively low cost, the corresponding linker for use in preformed handles is expensive and difficult to obtain; furthermore, the linker’s marginal stability as a trityl chloride derivative has led to its commercialization as a free alcohol, requiring the user to activate it by transforming it into a chloride using highly water-sensitive and corrosive halogenating reagents (e.g., SOCl2 , PBr3 ) that will react to completion only under absolutely optimal conditions. Once the linker has been activated in this way, direct esterification is not as straightforward to perform as might be expected, because of the lability of trityl halides to
Peptides
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trace impurities (e.g., water, dimethylamine in dimethylformamide, secondary amines in N,N-diisopropylethylamine) and the impossibility of monitoring the esterification reaction prior to an Fmoc release assay. In short, when the synthetic target demands the use of a 2-chlorotrityl resin, it is recommended that the user purchase a preloaded 2-chlorotrityl resin from a reputable commercial source rather than prepare the analogous handle-derivatized resin in the laboratory. To summarize, the synthesis of peptides bearing C-terminal acids can be understood by simple electronic principles, with the key factor usually being the additive electrondonating character of the substituents surrounding the benzyl ester peptide-resin anchorage. For Boc-based SPPS, the PAM linker is recommended in all cases, owing to its ideal level of acid stability and the chemical side reactions associated with the use of the classic alternative, the p-alkylbenzyl ester. For Fmoc-based SPPS, the choice is slightly more complex, but not as much as would be predicted given the rather bewildering array of commercially available linkers. In the vast majority of cases, the p-alkoxybenzyl ester peptide-resin anchorage, whether made using a Wang or HMPA linker (although the latter is preferred), will suffice. A significant benefit of this linkage type is its pseudo-orthogonality with respect to the graduated acid lability schemes used to make chemoselective side-chain modifications to the resin-bound peptide prior to the final deprotection/cleavage step. In cases in which (1) some or all of the side chain and/or N-terminal protecting groups are to be left intact on the cleaved peptide product, (2) the C-terminal residue is particularly prone to racemization, or (3) a proline residue is located at the C-terminus or at the penultimate (relative to the C-terminus) position, the 2-chlorotrityl linker is recommended, preferably in a commercially obtained preloaded resin.
Linkers yielding C-terminal primary amides Synthetic peptide targets are often desired bearing C-terminal primary amides. Whereas C-terminal carboxylic acids are recognized and rapidly degraded by carboxypeptidases in vivo, peptides bearing C-terminal primary amides are substantially more stable to resident carboxypeptidase activity. It is perhaps for this reason that most naturally occurring peptide hormones, such as oxytocin, neuropeptide Y, and substance P (among many others), are expressed in vivo as C-terminal primary amides. Interest in the pharmacology of these and other peptide hormones stimulated much of the early work on a general, straightforward methodology for the synthesis of peptide amides. In addition, the synthesis of peptide segments from proteins for the elucidation of structure-activity relationships has long been (and continues to be) a significant motivating force behind the synthesis of peptide amides. In such cases, a C-terminal primary amide is much more desirable than a C-terminal carboxylic acid, as the former more closely mimics the stereoelectronic characteristics of a secondary amide in the backbone of a parent protein, whereas the latter is ionized at physiological pH.
Resin and Linker Selection in SPPS
Early methodologies for the synthesis of peptides as C-terminal primary amides were irreproducible, inefficient, and yielded more side products than intended products. These early routes were all variations on the theme of direct installation of the C-terminal nitrogen by a nitrogen-containing nucleophile under strongly basic conditions. A quantum leap in the safety, practicality, and efficiency of the solid-phase synthesis of peptide amides occurred when the chemistry of primary amide protection was adapted to a peptideresin anchorage. This technology—the application of acid-labile, electron-rich monoor dibenzylic secondary amides as peptide-resin linkages—continues to be the standard method for the synthesis of peptide amides. As described for linkages used in the synthesis of C-terminal peptide acids, these linkages have been fine-tuned with respect to their electronic characteristics to afford a wide spectrum of acid lability.
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Entry no.
Peptide-linker-resin structure
Necessary linker or derivatized resin
Typical minimal acidolytic cleavage conditions
O
peptide
NH
H2N
1*
Anhydrous HF, 0°C, 1 hr
O peptide
NH
O
2
FmocNH
O
OMe MeO
MeO
O peptide
O
N H
OMe
4
O
OH
OMe
95% (v/v) TFA, 1 to 2 hr
MeO
MeO O
O
FmocNH
O
NH
3*
peptide
95% (v/v) TFA, 1 to 2 hr
OMe
MeO NH
MeO FmocNH
O O
N H
MeO
O O
OH
95% (v/v) TFA, 2 to 3 hr
MeO
O peptide
NHFmoc
NH
5
1% (v/v) TFA, 10 min O
O
O peptide
O
O
NHFmoc NH
6 O
H N
O
O
OH
O
1% (v/v) TFA, 10 min
O
O
Figure 18.7.4 Linkers yielding peptides that bear a C-terminal amide. Typical minimal acidolytic deprotection conditions are noted; acidolysis should be carried out at 25◦ C whenever not otherwise specified. Entries marked with an asterisk have the greatest general utility in routine peptide synthesis applications. TFA, trifluoroacetic acid.
The first linker to gain widespread use in the synthesis of C-terminal primary amides was the p-methylbenzhydryl amide (MBHA) linker (Fig. 18.7.4, entry no. 1), introduced by modification of a native, nonderivatized polystyrene resin (Matsueda and Stewart, 1981). The MBHA linker is well suited for use in Boc-based syntheses of peptide amides, as the bis-p-alkylbenzhydrylamide anchorage made by this linker is stable to repetitive TFA deblocking treatments but is quantitatively cleaved by HF during the final deprotection/cleavage step. The sole shortcoming of MBHA-functionalized resins is the high degree of batch-to-batch variation in the quality of commercially available material, which can be ascribed to the incidence of side reactions during the multistep synthetic route used to prepare this material. That problem notwithstanding, the MBHA linker continues to be the standard tool for Boc-based synthesis of peptide amides, largely for lack of a better alternative.
Peptides
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Much like in the development of linkers for the solid-phase synthesis of peptide acids, the benzhydryl core of the MBHA linker was functionalized with various electron-donating substituents to furnish linkers suitable for TFA-mediated cleavage following Fmoc-based SPPS. The first linker widely used for this purpose was the trialkoxybenzhydryl Rink linker (Fig. 18.7.4, entry no. 2; Rink, 1987). This linker, introduced by modification of a chloromethyl-derivatized polystyrene resin using ether coupling conditions similar to those used for the Wang linker, bears three electron-donating alkoxy substituents in place of the MBHA linker’s two alkyl substituents. The scissile C-N amide-forming bond is therefore labile to substantially milder acidolytic conditions, being cleaved quantitatively by ∼95% TFA in 1 to 2 hr. Again, like the Wang linker, the Rink linker was further refined to yield a preformed handle, the carboxymethyloxy-derivatized Rink handle (Fig. 18.7.4, entry no. 3; Bernatowicz et al., 1989). As is the case for handles used in the synthesis of peptide acids, the use of the Rink handle coupled to an aminomethyl-derivatized resin (which serves as an electronic surrogate for the ether-functionalized resin) is recommended over the use of the originally developed Rink-functionalized resin. Again, the possibility of TFA-mediated cleavage of the linker from the resin is essentially abrogated by the presence of an amide bond between the linker and the resin. It should be noted that the Rink linker’s benzyl 4carboxamidomethyl-3,5-dimethoxybenzyl ether is substantially more electron-rich than the Wang linker’s benzyl 4-hydroxymethylbenzyl ether. Accordingly, the potential for contamination of the cleaved peptide product by linker degradation byproducts is significantly reduced when the preformed Rink handle is coupled to an aminomethyl-derivatized resin rather than a Rink-functionalized polystyrene resin. From a practical standpoint, perhaps the prime motivation for using a preformed Rink handle is the often poor quality of commercially available Rink-functionalized polystyrene resins, which can be grossly discolored, ground, and incompletely and/or densely functionalized, leading to problems during chain assembly. The widely commercially available Rink handle (Senn Chemicals, Midwest Biotech) is colorless and may be coupled to similarly high-quality aminomethylderivatized resins to afford final products that are of vastly higher quality compared with Rink-functionalized polystyrene resins and that can be obtained at a lower cost. Since the development of the Rink linker, the PAL (primary amide linker) handle (Fig. 18.7.4, entry no. 4) has been devised as an improved TFA-labile linker for peptide amide synthesis (Albericio et al., 1990). Equipped with the same number of electron-donating alkoxy substituents as the Rink linker but one fewer aryl ring, this linker affords a slightly more stable peptide-resin anchorage compared with the Rink linker. One theoretical benefit of the PAL linker (commercially available from Advanced ChemTech) is that whereas the benzhydryl core of the Rink linker may decompose during TFA cleavage to liberate a carbocation that can alkylate the cleaved peptide, the PAL linker is chemically inert, yielding no soluble byproducts upon acidolysis. This potential benefit notwithstanding, the PAL handle offers little to no significant benefit over the Rink handle with regard to product purity, and it is substantially more expensive (Bernatowicz et al., 1989). As is the case with p-alkoxybenzyl ester linkages, the PAL and Rink linkers depicted in Figure 18.7.4, entry no. 2 to entry no. 4, are all suitable for side-chain modification of the resin-bound peptide. These linkages require long-term exposure to high concentrations of TFA for C-N bond scission and consequent peptide release as a C-terminal amide. Highly acid-labile trityl-based protecting groups on Lys, Ser, and Thr side chains, as well as monomethoxytrityl (Mmt) and xanthenyl (Xan) protecting groups on Cys side chains, may therefore be chemoselectively removed from the resin-bound peptide without any danger of significant chain loss due to acidolysis of the peptide-resin anchorage. Resin and Linker Selection in SPPS
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In much the same way that increasingly electron-rich linkers are used to allow the solidphase synthesis of peptide acids bearing either partial or complete side chain and/or N-terminal protecting groups, electron-rich linkers may also be used for the synthesis of protected peptide amides. Whereas o- and p-alkoxy substituents can readily be appended to the benzyl ester linkage to furnish highly acid-labile ester linkers, the trialkoxybenzhydryl core of the Rink linker leaves little room for further modification. A different aryl core is therefore used for this purpose—one that is substantially more electron-rich compared with the benzhydryl system. This core, known as the xanthenyl core, is used in the Sieber amide linker (Fig. 18.7.4, entry no. 5), which, like the Wang and Rink linkers, was first introduced via functionalization of a chloromethyl-derivatized polystyrene resin (Han et al., 1996). The high resonance stability of the xanthenyl cation allows acidolytic cleavage of the C-N bond using TFA concentrations as low as 1%, allowing t-butyl-based side chain protecting groups to remain intact in the cleaved peptide amide. This linker has been adapted for use in preformed-handle format (commercially available from Advanced ChemTech) by carboxymethylation of the xanthenyl alcohol to allow coupling to an aminomethyl-derivatized resin (Fig. 18.7.4, entry no. 6; Han et al., 1996). Again, it is advisable to use the Sieber handle coupled to a solid support via an amide linkage, as the benzyl xanthenyl ether formed by functionalization of a chloromethyl-derivatized polystyrene resin is highly acid-labile and will lead to significant and unnecessary contamination of crude peptide cleavage products. A common thread can be observed throughout the evolution of linkers that afford cleaved peptide products via acidolytic mechanisms. Specifically, whenever possible, the use of preformed handles is recommended over the installation of linkers onto polystyrene resins using ether bonds. As described above, this recommendation is founded on two principal concerns. The first is that resins for peptide synthesis can be much more cleanly and quantitatively prepared by coupling preformed, purified handles to a solid support via amide bonds, rather than coupling linker precursors to a solid support using comparatively labile ether bonds, with multiple subsequent solid-phase reactions needed to complete the synthesis of the linker. The second consideration is that inert, unactivated amide bonds are much more stable than are highly electron-rich ethers; this increased stability generally ensures that the linker remains attached to the resin during the final peptide deprotection/cleavage step, thereby limiting the creation of soluble carbocations that can alkylate the peptide product. A further benefit of using preformed handles for coupling to aminomethyl-derivatized resins is that these handles allow resins other than poly(styreneco-divinylbenzene) to be employed for SPPS. One practical caveat is that it is the author’s standard practice, whenever possible, to purchase a single large batch of high-quality aminomethyl-derivatized resin and functionalize this resin with a linker and a C-terminal amino acid as needed; this ensures reproducibility and high quality from the functionalized resin that is to be used. Commercial sources may certainly be used to obtain preloaded resins, although appropriate caution must be exercised when making any such purchase; in the case of Wang- or HMPA-functionalized resins in particular, it is important for the buyer to verify that the commercial source employs optimal chemistry (preferably the Sieber method; Sieber, 1987; see Basic Protocol 2) to anchor the C-terminal amino acid. Furthermore, it is recommended that the buyer be loyal to a single, reliable commercial source for preloaded resins, so as to ensure batch-to-batch reproducibility. This may be an overly cautious recommendation, but it is useful to be able to ascribe the failure of a peptide synthesis to problems with chain assembly or cleavage, while being able to rule out resin quality as a possible causative factor. Peptides
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Resin Selection Having selected the desired linkage chemistry to be employed, the decision regarding which insoluble polymer support to use must be made. It cannot be overemphasized that in all but a few cases, the linkage and resin selections are independent of one another; unfortunately, the prevalence of misnamed products (e.g., “Wang resin,” “Rink AmideMBHA Resin”) and the vast number of commercially available linker/resin combinations have somewhat clouded this point. For the sake of generality, the discussion that follows will focus exclusively on commercially available resins, particularly those whose merits have been widely validated in published studies conducted in both commercial and academic laboratories.
Hydrophobic resins (polystyrene backbone) The solid support employed by Merrifield in his landmark synthesis of a tetrapeptide continues to be the mainstay of SPPS (Merrifield, 1963). This resin, a 98:2 binary copolymer of polystyrene and divinylbenzene (PS-DVB)—commonly referred to as polystyrene, 2% cross-linked with divinylbenzene—possesses many qualities that substantiate its continued popularity in manual and automated batchwise SPPS. Polystyrene resins are inexpensive and widely available, and while the quality of commercially available resins is varied, there are several sources from which clean polystyrene resins can be reliably obtained (notably, Polymer Laboratories). From a chemical standpoint, polystyrene resins are ideal because of their chemical inertness to all of the reaction conditions encountered in SPPS, owing to the absence of any potentially labile functionalities in either the polymer backbone or the cross-linker (usually 1,4-divinylbenzene). PS-DVB resins are also superior to other resins on the basis of practical handling considerations, because of their free-flowing, powderlike consistency when dry, facile swelling in most organic solvents, and amenability to repeated shrink/swell and dry/wet cycles. These favorable qualities notwithstanding, traditional PS-DVB resins are marked by one shortfall in the context of manual batchwise SPPS. While these resins swell readily in nonpolar aprotic solvents (e.g., CH2 Cl2 , the original solvent used for peptide synthesis as conceived by Merrifield), their swelling in the polar aprotic solvents typically used today (e.g., dimethylformamide, 1-methyl-2-pyrrolidinone) is substantially diminished. The impact of this poor swelling is most significant during the first coupling step, which is noticeably sluggish. Thus, if possible, this step is best performed in mixed solvents, such as CH2 Cl2 /DMF or toluene/DMF; the nonpolar component (CH2 Cl2 or toluene) in such mixed solvent systems ensures adequate solvation of the polymer backbone, while the polar component allows high concentrations of amino acid derivatives and coupling reagents to be used.
Resin and Linker Selection in SPPS
Recent years have seen the introduction of the JandaJel resins, which were developed for solid-phase organic synthesis; however, these resins have since proven useful for addressing the swelling issue in SPPS, as they incorporate a 1,4-bis(4-vinylphenoxy)butane cross-linker that is more flexible and hydrophilic than the 1,4-divinylbenzene cross-linker. The reasoning behind the use of JandaJel resins for solid-phase organic synthesis (SPOS) is that the poor swelling of PS-DVB is often detrimental to reaction efficiencies when polar solvents are required. Not surprisingly, then, these resins have afforded significant advantages when applied to SPPS, as their swelling in DMF is ∼2.5 times that of traditional PS-DVB. In the author’s experience, one of the most telling instances in which improved resin swelling yielded a benefit in terms of coupling efficiency was during the attachment of the pentafluorophenyl ester of the HMPA handle (Fig. 18.7.3, entry no. 4) to aminomethyl-derivatized JandaJel, as compared with aminomethyl-derivatized
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PS-DVB. Whereas aminomethyl-derivatized PS-DVB required overnight reaction to give a ninhydrin-negative (see Basic Protocol 3) HMPA-functionalized resin, aminomethylderivatized JandaJel required only 30 min (J. A. Moss, unpub. observ.). Aminomethylderivatized JandaJel resin is commercially available from Sigma and has been functionalized with a variety of linker handles (e.g., HMPA, Rink, PAM) for a variety of solid-phase peptide and organic synthesis applications (Clapham et al., 2002; Moss et al., 2002).
Hydrophilic resins (polyethylene glycol backbone) One of the significant benefits of solid-phase peptide synthesis over solution-phase peptide synthesis is its amenability to automation. While the early automated peptide synthesizers operated in a batchwise manner, using glass or polychlorotrifluoroethylene vessels equipped with frits composed of sintered glass or woven Teflon fibers to retain the resin while solvents flowed freely to waste, continuous-flow synthesizers subsequently gained popularity in parallel with the development of the Fmoc/t-butyl method. Continuousflow synthesizers operate in much the same way as ion-exchange, gel-filtration, or highperformance liquid chromatography apparatuses, using rigid columns to isolate the resin, which remains fully solvated while reagents and solvents are washed through it during repetitive coupling, deprotection, and washing cycles. In the continuous-flow method, resin swelling must obviously remain constant during the course of the synthesis; should resin swelling increase, the column end caps may burst, leading to failure of the synthesis and instrument damage. Hydrophilic resins were designed to circumvent this problem; the swelling of these resins changes negligibly over the course of a synthesis, as the high polarity of the resin in its initial form ensures that its overall character does not change from uniformly nonpolar to highly polar as the grafted peptide chain grows in length. This contrasts starkly with PS-DVB resins (and JandaJel resins, albeit to a lesser extent), which easily swell to three times their initial swollen bed volume during syntheses of large peptides (>30 residues). PEGA (polyethylene glycol–polyacrylamide) resin is a cross-linked, aminefunctionalized ternary copolymer commercially available from Sigma and Polymer Laboratories (Meldal, 1992). This resin contains only secondary amide and ether bonds and is therefore highly hydrophilic and inert to most peptide synthesis conditions. While intended for automated Fmoc-based SPPS under continuous-flow conditions, this resin is applicable to batchwise synthesis as well, provided that only mild rotary shaking (so as to minimize resin fracture) is performed. The handling properties of PEGA resins leave much to be desired, as they have poor mechanical stability and must always be maintained in a moist slurry to avoid irreversible collapse of the polymer matrix. A compromise between chemical and mechanical stability can be found in a solid support known as CLEAR (cross-linked ethoxyacrylate resin), which is commercially available from Peptides International (Kempe and Barany, 1996). Like PEGA, CLEAR is a ternary copolymer of hydrophilic monomers; however, CLEAR is composed partially of alkyl ester linkages, rather than the secondary amide linkages found in PEGA. While stable to all of the standard conditions encountered in Boc- and Fmoc-based SPPS, CLEAR is not stable to harsh alkali treatments. Nevertheless, the rarity of such reaction conditions in SPPS mitigates the significance of this drawback, as the instability of many peptide side chain protecting groups to base, as well as the known base-mediated side reactions involving protected peptides, has virtually abolished the need for base treatments at any stage in SPPS. This shortcoming notwithstanding, CLEAR is a useful solid support by virtue of its good mechanical stability, which is intermediate between that of PEGA and polystyrene resins.
Peptides
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“Hybrid” resins (polystyrene/polyethylene glycol composites) An alternative strategy for improving resin swelling in polar solvents, as well as for maintaining consistent swelling throughout peptide chain assembly, entails the grafting of polyethylene glycol (PEG) chains onto a PS-DVB resin. These hydrophilic chains are usually terminated with pendant amino groups that are ready for functionalization with an IRAA or a linker handle. This modification provides increased swelling relative to PS-DVB in most organic solvents, and particularly in the polar aprotic solvents (e.g., dimethylformamide, 1-methyl-2-pyrrolidinone, acetonitrile), in which PS-DVB exhibits comparatively poor swelling. The length and the sterically unhindered nature of the PEG chain, which serves as a bridge between the peptide chain and the polystyrene resin backbone, are thought to result in a more “solvent-like” environment for the resinbound peptide. Both resins described below are amenable to synthesis under batchwise or continuous-flow conditions, as they exhibit good mechanical stability and consistent swelling. The first chemistry used to attach PEG grafts to polystyrene resins involved an amide linkage to an MBHA-functionalized polystyrene resin (Fig. 18.7.3, entry no. 5; Zalipsky et al., 1994). The resulting resin, appropriately termed PEG-PS, is suitable only for Fmoc-based SPPS, due to the lability of the MBHA linkage to strong acids such as HF. A key feature of the PEG-PS resin is the incorporation of a norleucine residue between the PEG graft and the MBHA linker as an IRAA for solid-phase AAA. As in any other amine-derivatized resin, the free amine–functionalized PEG terminus must be acylated with a linker handle before peptide chain assembly can be initiated; the final TFA deprotection/cleavage step cleaves this linkage without detriment to the PEG-resin MBHA linkage. PEG-PS resin is commercially available from Applied Biosystems in native, linker-functionalized, and preloaded forms. An alternative approach for the attachment of PEG chains to polystyrene resins involves the generation of PEG in situ by polymerization of ethylene oxide onto a PSDVB resin. The resulting family of resins is marketed under the trade name TentaGel and is commercially available from Rapp Polymere. TentaGel resins are available in a wider selection compared with PEG-PS resins, and they can also be obtained in native, linker-functionalized, and preloaded forms. Whereas the early variants of TentaGel were marked by exceedingly low loading (∼0.1 to 0.2 mmol/g) and acid-labile benzylic ether linkages connecting the pendant PEG chains to the polystyrene backbone, significant improvements have broadened the applicability of TentaGel resins for SPPS. Notably, the high-capacity TentaGel HL resin is commercially available, and offers satisfactory loading capacities of 0.4 to 0.5 mmol/g. In addition, TentaGel resins are now made almost exclusively with an additional methylene spacer between the PEG graft and the polystyrene backbone. This feature is of the utmost importance, as it essentially eliminates the specter of acidolytic cleavage of the PEG graft from the PS-DVB core (with resulting contamination of the crude peptide product).
Resin and Linker Selection in SPPS
The choice of resin for a given synthesis is dictated principally by the manner in which the synthesis is to be performed. For manual batchwise SPPS using either Boc/benzyl or Fmoc/t-butyl chemistry, any resin may in principle be used. However, certain practical considerations must be considered prior to commencement of any synthesis. The inferior mechanical stability of PEG-based resins—principally PEGA resin and CLEAR— contraindicates their use in manual batchwise SPPS, in which mechanical agitation is likely to lead to resin fracture and frit clogging. Conventional polystyrene resins and hybrid resins, such as TentaGel and PEG-PS, are therefore recommended, as the mechanical rigidity imparted by the polystyrene core of such resins ensures the integrity of the resin throughout the course of synthesis. For automated SPPS, the mode of
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operation of the synthesizer is of prime importance. Automated synthesizers that operate in a batchwise fashion (e.g., ABI 431; ABI 433; all CSBio, Rainin, and Advanced ChemTech synthesizers) are amenable to nearly all resin types, although for longer peptides, the mechanical stability of polystyrene resins is desirable. For the few synthesizers that employ continuous-flow methodology (e.g., ABI Pioneer, MilliGen/Bioresearch Crystal), resins that exhibit minimal changes in swelling throughout the course of synthesis (i.e., PEG-based and hybrid resins, as opposed to conventional polystyrene resins) are required so that instrument failure resulting from increases in resin swelling can be avoided. While automated Fmoc-based SPPS under continuous-flow conditions is most often performed using hybrid PEG-grafted polystyrene resins, it is the author’s experience that a high-quality polystyrene resin may be used for automated or manual batchwise SPPS using Fmoc/t-butyl or Boc/benzyl chemistry. Because they can be functionalized with nearly any linker for a given application, aminomethyl-derivatized polystyrene resins are recommended above all others, being of unmatched chemical and mechanical stability, versatility, and cost-effectiveness. The last of those three qualities is a key consideration for most users, as the cost of PEG-grafted polystyrene resins and PEG-based resins is in the range of two to five times the cost of aminomethyl-derivatized polystyrene resins. Furthermore, this cost comparison is on a per-weight basis and therefore does not take into account the fact that PEG-grafted resins typically have one-fourth to one-half the amine loading capacity (per gram) of standard polystyrene resins. This disparity in resin loading is of critical economic importance, as loading capacity is the ultimate limiting factor determining the total quantity of synthetic peptide product that can be prepared in a given synthesis.
Conclusions The most important point to be taken from this unit is that the linker and resin are separate entities and can be selected independently. The selection of the linker depends primarily on three factors: (1) the synthetic chemistry to be employed (Fmoc-based, Boc-based, or a combination thereof); (2) whether any protecting groups are to be left intact on the cleaved peptide product; and (3) the desired C-terminal functionality (primary amide or carboxylic acid). While usually separable, these issues are sometimes interdependent, such as when chemoselective postassembly modification of the resin-bound peptide is desired, or when partially or fully protected peptides are desired upon cleavage of the peptide-resin anchorage. It is hoped that this distillation of the somewhat daunting selection of commercially available linkers down to a simple spectrum of carbocation stabilities will allow the newcomer to peptide chemistry to make an educated decision prior to starting a synthesis. The selection of the resin is simple by comparison; here, the overriding question is whether the user plans to perform automated Fmoc-based SPPS under continuous-flow conditions. If this is the case, only a PEG-based or PEG-grafted polystyrene resin is appropriate, with the majority of published and anecdotal practical experience dealing with the use of the latter, although the former is acceptable as well. For automated or manual batchwise SPPS using either Boc-based or Fmoc-based chemistry, polystyrene resins are the most appropriate. This stems principally from their unrivaled mechanical stability throughout repetitive vortexing, stirring, sparging, and/or inversion to aid in the diffusion of reactants throughout the polymeric matrix during coupling and/or washing. From an economic standpoint, low cost, ready availability, and the wide range of amine loading capacities also serve to elevate aminomethyl-derivatized polystyrene resins to their current status as the mainstay of modern SPPS. Peptides
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BASIC PROTOCOL 1
FUNCTIONALIZATION OF AN AMINE-DERIVATIZED RESIN USING PREFORMED HANDLES This protocol provides three alternatives for functionalizing an amine-derivatized resin using a hydroxyl linker as a preformed handle. Although any of these three methods will work, the preferred method is the one involving a 2,4,5-trichlorophenyl (Tcp) or pentafluorophenyl (Pfp) ester linker. Preactivated Tcp esters are commercially available from Peptides International or may be prepared according to published procedures (Atherton and Sheppard, 1989).
Materials 5% (v/v) triethylamine in dimethylformamide (DMF) or 5% (v/v) triethylamine in 1:1 DMF/toluene Linker (free carboxylic acid, Tcp ester, or Pfp ester) to be used in peptide synthesis N,N -diisopropylcarbodiimide (DIC; optional) N-hydroxybenzotriazole (HOBt; optional) Benzotriazol-l-yloxytris(dimethylamino)phosphonium hexafluorophosphate (BOP; optional) DMF Amine-derivatized resin to be used in peptide synthesis Glass or polypropylene fritted vessel with Teflon stopcock Rotary shaker Wash bottle 1. Measure out the desired amount of amine-derivatized resin and transfer to a fritted vessel. Set the vessel’s stopcock to the closed position, and then preswell the resin by adding DMF containing ∼5% triethylamine or 1:1 DMF/toluene containing ∼5% triethylamine (10 ml DMF/triethylamine or DMF/toluene/triethylamine per g resin), capping the vessel, and incubating at room temperature with rotary shaking for a minimum of 1 hr, but preferably overnight. The inclusion of a base such as triethylamine or DIEA (N,N-diisopropylethylamine) in the preswelling solution is recommended, as this will neutralize any resin-bound salts.
2. Open the stopcock to drain all liquid from the vessel. Thoroughly wash the resin twice, each time by using a wash bottle to apply a constant flow of DMF for 20 sec and then allowing all liquid to drain from the vessel. After the second wash, set the stopcock in the closed position. 3a. If using a free carboxylic acid linker and DIC/HOBt catalysis: Dissolve 3 equivalents of the linker and 3 equivalents of HOBt to a concentration of 0.5 M in DMF. Use a pipet filler and pipet to apply this solution to the resin, and then apply 3 equivalents of DIC to the resin. 3b. If using a free carboxylic acid linker and BOP catalysis: Dissolve 3 equivalents of the linker and 3 equivalents of BOP to a concentration of 0.5 M in DMF. Use a pipet filler and pipet to apply this solution to the resin, and then apply 3 equivalents of DIEA to the resin. 3c. If using a Pfp or Tcp ester linker and HOBt catalysis: Dissolve 3 equivalents of the (already activated) linker and 3 equivalents of HOBt to a concentration of 0.5 M in DMF. Use a pipet filler and pipet to apply this solution to the resin. 4. To ensure complete mobility of the resin slurry during coupling, fill the vessel to between two-thirds and three-quarters of its capacity with DMF. Cap the vessel, invert and vent by opening the stopcock, and then place on a rotary shaker for several hours. Resin and Linker Selection in SPPS
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5. Perform the qualitative ninhydrin test (see Basic Protocol 3) to assess the completeness of the resin functionalization reaction. Current Protocols in Protein Science
ACYLATION OF HYDROXYL LINKERS VIA REACTION WITH A MIXED ANHYDRIDE
BASIC PROTOCOL 2
This protocol describes the preferred method for acylation of a hydroxyl linker with the C-terminal amino acid of the target peptide. While there are many methods that can be used to accomplish this acylation, most carry a risk of undesirable side reactions (e.g., racemization, premature Fmoc deprotection and resultant dipeptide formation) or are exceedingly complicated, requiring expensive reagents and rigorously anhydrous reaction conditions. The method presented here represents a compromise, providing an acceptably low level of racemization (<3%) and negligible Fmoc deprotection, albeit with subquantitative acylation efficiency (usually ∼70%). However, the reader should note that quantitative acylation efficiency is not critical in this setting, as (1) most unreacted hydroxyl groups are capped by 2,6-dichlorobenzoylation, (2) any uncapped hydroxyl groups will remain unreactive throughout the synthesis, and (3) the user can compensate for suboptimal efficiency by using 50% more hydroxyl-functionalized resin to afford the desired product yield.
Materials Glass or polypropylene fritted vessel (with Teflon stopcock) containing hydroxyl-functionalized resin (see Basic Protocol 1) Dimethylformamide (DMF) Fmoc-protected amino acid (to serve as the C-terminal amino acid in the synthesized peptide) Anhydrous pyridine (packaged under nitrogen in a septum-equipped bottle; Aldrich or Acros) 2,6-Dichlorobenzoyl chloride Wash bottle NOTE: 2,6-Dichlorobenzoyl chloride is highly reactive. Therefore, it should be purchased in small quantities and exposed to air for as short a period as possible. The use of decomposed 2,6-dichlorobenzoyl chloride will lead to greatly decreased acylation efficiency and additional undesirable side reactions. 1. Set the stopcock on the resin-containing vessel to the open position. To ensure that all residual water, base, and other reagents from preceding reaction steps are fully removed, wash the resin twice, each time by using a wash bottle to apply a constant flow of DMF for 20 sec and then allowing all liquid to drain from the vessel, leaving a wet resin cake. After the second wash, set the stopcock to the closed position. If the resin has been dried, it should be allowed to swell in DMF or 1:1 DMF/toluene containing ∼5% (v/v) triethylamine (as in Basic Protocol 1, step 1) for at least several hours before washing.
2. Dissolve 5 equivalents of the Fmoc-protected amino acid and 6 equivalents of anhydrous pyridine in DMF such that the concentration of the amino acid is 0.5 M. Use a pipet filler and pipet to apply this solution to the drained resin, and then apply 5 equivalents of neat 2,6-dichlorobenzoyl chloride to the resin. 3. To ensure complete mobility of the resin slurry during acylation, fill the vessel to between two-thirds and three-quarters of its capacity with DMF. Cap the vessel, invert and vent by opening the stopcock, and then place on a rotary shaker for 12 to 24 hr. 4. Optional: Perform an Fmoc release assay to assess the completeness of the linker acylation reaction (Chan and White, 2000). 5. Proceed to peptide synthesis using the acylated material in the vessel.
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BASIC PROTOCOL 3
QUALITATIVE NINHYDRIN TEST When functionalizing an amino-derivatized resin using a preformed linker, a qualitative ninhydrin test should be performed to assay the completeness of the functionalization reaction. The protocol provided here can also be used to assess the completeness of subsequent peptide coupling steps.
Materials Resin to be tested (see Basic Protocol 1) Dichloromethane Unreacted aminomethyl-derivatized resin (with loading capacity similar to that of test resin) 4% (w/v) ninhydrin in ethanol 4:1 (v/v) phenol/ethanol Anhydrous pyridine (packaged under nitrogen in a septum-equipped bottle; Aldrich or Acros) 3-ml polypropylene fritted syringe (Applied Separations) Wash bottle Tygon tubing Vacuum source 10 × 75–mm glass culture tubes 1. Transfer 10 to 20 mg of putatively functionalized resin to a 3-ml polypropylene fritted syringe. To ensure that all residual water, base, and other reagents from previous reaction steps are fully removed, wash the resin once by using a wash bottle to apply a constant flow of dichloromethane for 20 sec and then allowing all liquid to drain from the syringe. Attach a piece of Tygon tubing to the bottom of the syringe and apply vacuum until the resin becomes a dry, free-flowing powder (∼20 sec). It is important that this wash step be performed thoroughly, as any residual DMF will decompose in the ninhydrin test reaction, leading to a false-positive result.
2. Transfer the dried resin (i.e., the test sample) to a 10 × 75–mm glass culture tube. To a separate tube, as a positive control, add a few beads of unreacted aminomethylderivatized resin with a loading capacity similar to that of the test resin. 3. Add three drops of 4% ninhydrin in ethanol, three drops of a 4:1 mixture of phenol and ethanol, and three drops of pyridine to each of the tubes prepared in step 2. As a negative control, aliquot three drops of each of those same three solutions into an empty glass culture tube. 4. Place all three tubes in a heating block set to 100◦ C. After 2 min, check the color of the solution in each tube. The positive control solution should have a strong indigo blue color, and the negative control solution should be faintly yellow. If there are no blue beads in the test sample, then acylation is complete, even if the solution above the resin has a faint blue or gray coloration.
LITERATURE CITED Albericio, F. and Barany, G. 1991. Hypersensitive acid-labile (HAL) tris(alkoxy)benzyl ester anchoring for solid-phase synthesis of protected peptide segments. Tetrahedron Lett. 32:10151018. Resin and Linker Selection in SPPS
Albericio, F., Kneib-Cordonier, N., Biancalana, S., Gera, L., Masada, R.I., Hudson, D., and Barany,
G. 1990. Preparation and application of the 5-(4(9-fluorenylmethyloxycarbonyl)aminomethyl3,5-dimethoxyphenoxy)valeric acid (PAL) handle for the solid-phase synthesis of Cterminal peptide amides under mild conditions. J. Org. Chem. 55:3730-3743. Atherton, E. and Sheppard, R.C. 1989. Solid Phase Peptide Synthesis: A Practical Approach. IRL Press, Oxford.
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Barlos, K., Chatzi, O., Gatos, D., and Stavropoulos, G. 1991a. 2-Chlorotrityl chloride resin: Studies on anchoring of Fmoc-amino acids and peptide cleavage. Int. J. Pept. Protein Res. 37:513-520. Barlos, K., Gatos, D., and Schafer, W. 1991b. Synthesis of prothymosin alpha (pro T-alpha), a protein consisting of 109 amino acid residues. Angew. Chem. Int. Edit. Engl. 30:590-593. Bayer, E., Goldammer, C., and Zhang, L. 1993. A new acid labile anchor system for the synthesis of fully protected peptide fragments. Chem. Pept. Proteins 5/6(Pt. A):3-8. Bernatowicz, M.S., Daniels, S.B., and K¨oster, H. 1989. A comparison of acid labile linkage agents for the synthesis of peptide C-terminal amides. Tetrahedron Lett. 30:4645-4648. Chan, W.C. and White, P.D. 2000. Fmoc Solid Phase Synthesis: A Practical Approach. Oxford University Press, Oxford. Clapham, B., Lee, S.H., Koch, G., Zimmermann, J., and Janda, K.D. 2002. The preparation of polymer bound beta-ketoesters and their conversion into an array of oxazoles. Tetrahedron Lett. 43:5407-5410. Floersheimer, A. and Riniker, B. 1991. Solidphase synthesis of peptides with the highly acidsensitive HMPB linker. In Peptides 1990: Proceedings of the Twenty-First European Peptide Symposium (E. Giralt and D. Andreu, eds.) pp. 131-133. Escom, Leiden, The Netherlands. Han, Y., Bontems, S.L., Hegges, P., Munson, M.C., Minor, C.A., Kates, S.A., Albericio, F., and Barany, G. 1996. Preparation and applications of xanthenylamide (XAL) handles for solid-phase synthesis of C-terminal peptide amides under particularly mild conditions. J. Org. Chem. 61:6326-6339. Kempe, M. and Barany, G. 1996. CLEAR: A novel family of highly cross-linked polymeric supports for solid-phase peptide synthesis. J. Am. Chem. Soc. 118:7083-7093. Matsueda, G.R. and Stewart, J.M. 1981. A pmethylbenzhydrylamine resin for improved solid-phase synthesis of peptide amides. Peptides 2:45-50. Meldal, M. 1992. PEGA: A flow-stable polyethylene glycol-dimethylacrylamide copolymer for solid-phase synthesis. Tetrahedron Lett. 33:3077-3080.
Merrifield, R.B. 1963. Solid phase peptide synthesis: I: The synthesis of a tetrapeptide. J. Am. Chem. Soc. 85:2149-2153. Merrifield, R.B. 1964. Solid phase peptide synthesis: II: The synthesis of bradykinin. J. Am. Chem. Soc. 86:304-305. Mitchell, A.R., Erickson, B.W., Ryabtsev, M.N., Hodges, R.S., and Merrifield, R.B. 1976. Tert-butoxycarbonylaminoacyl-4(oxymethyl)phenylacetamidomethyl-resin, a more acid-resistant support for solid-phase peptide synthesis. J. Am. Chem. Soc. 98:7357-7362. Mitchell, A.R., Kent, S.B., Engelhard, M., and Merrifield, R.B. 1978. A new synthetic route to tert-butyloxycarbonylaminoacyl4-(oxymethyl)phenylacetamidomethyl-resin, an improved support for solid-phase peptide synthesis. J. Org. Chem. 43:2845-2852. Moss, J.A., Dickerson, T.J., and Janda, K.D. 2002. Solid phase peptide synthesis on JandaJel resin. Tetrahedron Lett. 43:37-40. Rink, H. 1987. Solid-phase synthesis of protected peptide fragments using a trialkoxy-diphenylmethylester resin. Tetrahedron Lett. 28:37873790. Sheppard, R.C. and Williams, B.J. 1982. Acid-labile resin linkage agents for use in solid phase peptide synthesis. Int. J. Pept. Protein Res. 20:451-454. Sieber, P. 1987. A new acid-labile anchor group for the solid-phase synthesis of C-terminal peptide amides by the Fmoc method. Tetrahedron Lett. 28:2107-2110. Wang, S.S. 1973. Para-alkoxybenzyl alcohol resin and para- alkoxybenzyloxycarbonylhydrazide resin for solid-phase synthesis of protected peptide fragments. J. Am. Chem. Soc. 95:13281333. Zalipsky, S., Chang, J.L., Albericio, F., and Barany, G. 1994. Preparation and applications of polyethylene glycol-polystyrene graft resin supports for solid-phase peptide synthesis. React. Polym. 22:243-258.
Contributed by Jason A. Moss The Scripps Research Institute La Jolla, California
Mergler, N., Tanner, R., Gosteli, J., and Grogg, P. 1988. Peptide synthesis by a combination of solid-phase and solution methods I: A new very acid-labile anchor group for the solid phase synthesis of fully protected fragments. Tetrahedron Lett. 29:4005-4008.
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CHAPTER 19 Identification of Protein Interactions INTRODUCTION
P
rotein interactions are intrinsic and vital to many cellular processes. In recent years many new techniques have been developed for the study of this important topic; the review by Phizicky and Fields (1995) provides an excellent general overview of this subject. This chapter describes biochemical, physical, and genetic methods that are used to detect specific protein interactions. The companion Chapter 20 presents a series of complementary methods that are used to provide quantitative information about protein interactions, such as the strength and kinetics of binding.
This chapter begins with a brief overview of the analysis of protein interactions in which some of the basic kinetic and thermodynamic concepts are described (UNIT 19.1). This is followed by a presentation of one of the most widely used approaches for detecting protein interactions, the yeast two-hybrid/interaction trap system (UNIT 19.2). In this genetic method transcriptional activity is used as a measure of protein-protein interaction. The high-throughput screening using yeast two-hybrid arrays is described in UNIT 19.6. In this approach, a protein array composed of living yeast cells is used in a functional screen for protein interactions using the two-hybrid assay. In the commonly used technique of co-immunoprecipitation (UNIT 19.4), cell-free extracts are incubated with an antibody to a protein or an epitope tag in order to coprecipitate associated proteins. This approach is used to isolate multiprotein complexes that presumably were present in the intact cell. An advanced microscopic technique that detects the transfer of excited state energy (fluorescence resonance energy) between matching donor and acceptor fluorophores is known as FRET microscopy. As energy transfer is nanometer-scale distance dependent, proximity information between interacting labeled proteins can be obtained. Practically, protein-protein interactions can be observed and localized within a single cell. In the method described here (UNIT 19.5) a Green Fluorescence Protein (GFP)-tagged protein is used as the donor and a Cy3 dye-labeled antibody as the acceptor. FRET microscopy monitors the interaction of the proteins due to the quenching of the donor protein after contact with labeled antibody. Support protocols describe microinjection of proteins into the nucleus and cytoplasm and the labeling of antibody with Cy3. The probing of proteins immobilized on a solid support membrane following separation by SDS-PAGE by antibodies is termed western blot analysis (UNIT 10.10). In UNIT 19.7, a variant of this method, termed far western analysis is described. In this method, proteins resolved by SDS-PAGE are probed by non-antibody proteins in order to identify specific protein-protein interactions. This method may be useful for studying protein interactions between proteins difficult to analyze by other methods due to, for example, solubility problems. LITERATURE CITED Phizicky, E.M. and Fields, S. 1995. Protein-protein interactions: Methods for detection and analysis. Microbiol. Rev. 59:94-123.
Paul T. Wingfield Contributed by Paul T. Wingfield Current Protocols in Protein Science (2001) 19.0.1 Copyright © 2001 by John Wiley & Sons, Inc.
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Analysis of Protein-Protein Interactions
UNIT 19.1
BASIC CONCEPTS Most of the work of living organisms is performed by proteins. Proteins do their work by acting on other macromolecules: nucleic acids, carbohydrates, lipids, and, especially, other proteins. Inside cells, protein-protein interactions are instrumental in enzymatic actions upon protein substrates and in the fleeting and lasting protein assemblies that govern signal transduction, cell division, DNA replication, and transcription initiation. Outside cells, protein interactions allow cells to talk with each other: ligands expressed on the cell surface often bind protein receptors expressed on adjacent cells, and secreted protein ligands bind receptors on distant cells. Techniques to detect and study proteinprotein interactions have become steadily more easy to use in recent years. This unit, and those that follow, lay them out. Qualitative statements about protein-protein interactions inform much of the current biological literature. This unit presents the basics for understanding such qualitative statements—i.e., the quantitative concepts by which these can be evaluated. Colloquially, protein interactions are often referred to as either stable or transient. The statement that a certain interaction is stable is typically used when the interacting proteins form a complex. Membership in a protein complex is most commonly defined operationally— based on detection of the protein after its coprecipitation with a reagent specific for another member of the complex—but is sometimes defined based on a strong and biologically plausible interaction in a two-hybrid experiment. Proteins that form such complexes with one another are often referred to as partners. By contrast, the term transient is used to denote, e.g., interactions between an enzyme and a protein substrate. As the preceding example suggests, there is no particular correlation between the stability of a protein interaction and its biological significance. It is also worth noting that there are exceptions to commonly held assumptions about stable and transient interactions: some interactions observed in coprecipitation experiments have no biological validity, whereas some enzyme-substrate interactions last for minutes. For these reasons, it is often useful to think of protein interactions in more formal terms. The strength, or affinity, of an interaction between two proteins is described by equilibrium parameters. The speed at which an interaction occurs, and at which interacting proteins come apart, is given by kinetic parameters. EQUILIBRIUM PARAMETERS The strength of an interaction can be given as the equilibrium dissociation constant, Kd, or by its reciprocal, Ka (discussed below). When two proteins A and B associate, the Kd is given by
Kd =
[A][B] [AB]
where [AB] is the concentration of the complexed species and [A] and [B] are the concentrations of the noncomplexed species. Concentrations are given in molar terms, as is the Kd. There are two useful special cases to keep in mind. First, when A and B are present in equal concentration, the concentration at which half of each protein is present in the form of AB complex is the Kd. Second, when the concentration of A vastly exceeds the Contributed by Roger Brent Current Protocols in Protein Science (1998) 19.1.1-19.1.4 Copyright © 1998 by John Wiley & Sons, Inc.
Identification of Protein Interactions
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Table 19.1.1 Interactions
Dissociation Constants for Some Biologically Significant
Interaction
Kd
Streptavidin-biotin binding Antibody-antigen interaction with good antibody DNA binding protein with specific site Antibody-antigen interaction with weak antibody Enzyme-substrate interations Cooperative interaction between DNA-bound phage repressors Cooperative interaction between phage repressor and E. coli RNA polymerase
10−14 M 10−8 to 10−10 M 10−8 to 10−10 M 10−6 M 10−4 to 10−10 M 10−4 M 10−2 M
concentration of B (say, by 100-fold)—typically indicated [A] >> [B]—the concentration of A at which half of the B is found in the AB complex approximates Kd. Typical Kd values for biologically significant interactions are given in Table 19.1.1. Note that many significant interactions are weak. Alternatively, affinity may be given as an equilibrium association constant, Ka. The Ka is simply Ka =
[ AB] [ A ][ B]
In other words, it is the reciprocal of the equilibrium dissociation constant: Ka = 1/Kd. Association constants are used less often than Kd, but do appear in the literature of some subfields—for example, in descriptions of antibody-antigen interactions. The strength of an interaction is directly proportional to the change in Gibbs free energy (∆G) when A and B interact, which is given by ∆G = ∆H − T∆S
where T is the temperature in degrees Kelvin, ∆S is the change in entropy (S), and ∆H is the change in enthalpy (H). ∆G is given in units of kilocalories/mole (kcal mol−1). An increase in the free energy of an interaction by 1 kcal decreases the Kd by a factor of ∼7. Note that the temperature affects the entropy term of the equation. As the temperature decreases, reactions driven by enthalpy are less affected by losses in entropy, and are usually favored. By contrast, reactions whose free energy is derived largely from favorable changes in entropy are disfavored at lower temperatures. This means that the dependence of a protein association on temperature can often provide a clue to which term predominately contributes to the free energy change involved in that association. The relationship between Ka and ∆G is defined as follows: ∆G 0 = − RT ln
Analysis of Protein-Protein Interactions
[ AB] [ A ][ B]
∆G 0 = − RT ln K a = − RT ln
1 = − RT ln K Kd d
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where ∆G0 is the free energy change under standard conditions (25oC); R is the universal gas constant (1.9872 calmole−1K–1); and T is the temperature in degrees Kelvin (25°C is 289.1 K). Therefore, ∆G 0 = 0.588 ln K d
and, since ln x = 2.303 log10 x, ∆G0 = 1.36 log10 Kd. For example:
(a) Kd = 1 × 10−14 , then ∆G0 = −19.04 kcal mol −1 (b) Kd = 1 × 10−2 , then ∆G0 = −2.72 kcal mol −1 The strepavidin-biotin reaction (a) is intrinsically more favorable in the direction of binding, [AB] formation, than the low-affinity interaction (b) involving phage repressor (see Table 19.1.1). It should be pointed out that the DG values calculated above assume that the molar ratio of reactants [A] [B] and product [AB] is 1 M (standard state). Many (but by no means all) biologically important protein interactions seem to be largely driven by ∆H, or changes in enthalpy. That is fortunate, in that changes in entropy on binding are very hard to quantitate, or even think about precisely. For example, proteins in solution are surrounded by water molecules that form hydrogen bonds with surface residues. When two proteins interact, the ordered arrangement of the water molecules that surrounded the interacting surfaces of the proteins is often disrupted, and this loss of order provides an entropically favorable term to the free energy of the interaction. Such changes in the free energy due to entropy are very hard to predict. By contrast, enthalpic changes are easier to understand. If formation of one hydrogen bond liberates about −1 kcal M−1, and formation of a particular ionic contact liberates about −2 kcal M−1, then the energies of these changes are additive such that the formation of both bonds usually liberates −3 kcal M−1—which means that the Kd is decreased >100-fold by the enthalpic changes. Kinetic Parameters The above descriptions of protein interactions make no reference to the speed at which association or dissociation occurs. These speeds are given by kinetic parameters. The dissociation rate constant, kdissoc, gives the speed at which the AB complex dissociates into A and B (AB → A + B). kdissoc is a first-order rate constant—i.e., one that is dependent on the concentration of one species, in this case the AB complex—and is given by the rate of decrease in the concentration of AB: kdissoc [A][B] =
− d[ AB] dt
Its units are those of reciprocal time (t in the equation), usually given in sec−1. For example, a dissociation rate constant of 10−4 sec−1 means that one in 104 of the AB complexes present comes apart each second. Similarly, the association rate constant, kassoc, gives the speed at which A and B associate to form an AB complex (A + B → AB). This is a second-order reaction—i.e., its speed depends on the concentrations of both A and B—and its rate constant is is given by
kassoc [A][B] =
+ d[ AB] dt
Its units are those of reciprocal concentration × reciprocal time, typically given in M−1 sec−1.
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For example, suppose that protein A is present at a concentration of 10−6 M in a cell, that protein B is injected to a nuclear concentration of 10−5 M, and that the rate constant for this antibody-antigen association is 10−4 M−1 sec−1. After injection, the concentration of AB will be [AB] = [A] × [B] × K assoc = (10 −6 M)(10 −5 M)(10 −4 M ) = 10 −7 M sec −1
That is, in the first second after mixing, 10−7 M of AB complex will form. Every 10-fold increase in the concentration of either reactant increases the rate of product formation 10-fold. Note that, at equilibrium, by definition,
d[AB] =0 dt and, therefore, for the AB interaction, k K d = dissoc kassoc
The fact that the strength of equilibrium interactions reflects the speed of associations and dissociations has important consequences. To understand this, imagine two pairs of proteins that interact with the same Kd. Proteins A and B come together slowly, but, once the AB complex forms, it takes a long time to come apart. By contrast, proteins C and D come together rapidly, and the CD complex dissociates rapidly. There are two common cases in which these kinetic differences in the AB and CD associations would be significant. One is in measurement. Many techniques, such as the “pulldown” and immunological coprecipitation techniques described in this chapter, rely on the the fact that proteins remain associated during some sequence of steps, while they are being separated from other proteins in a mixture and while the isolated complex is being rinsed. No matter how tightly the proteins associate, if they come apart before their complex can be separated and rinsed, the complex will not be detected. Moreover, if the AB and CD interactions have the same Kd, but CD comes apart more rapidly, then a coprecipitation experiment can falsely suggest that the CD association is weaker. The second case concerns the biological effects. Many biological phenomena, such as the transcription phenotypes resulting from protein-protein interactions in two-hybrid experiments (UNIT 19.3), seem to be well-described by consideration of equilibrium measurements. However, it is worth keeping in mind that any biological process that results from the association of two proteins requires a minimum time to occur. For some enzyme-substrate interactions, the minimum time may be on the order of microseconds, but for others, such as the initiation of DNA replication, it may be measured in seconds. If the complex dissociates faster than the minimum time, then, no matter how tight the interaction, the process will not occur. If AB and CD have the same Kd, but CD dissociates faster, the association may not produce a biological effect.
Analysis of Protein-Protein Interactions
Contributed by Roger Brent The Molecular Sciences Institute Berkeley, California
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Interaction Trap/Two-Hybrid System to Identify Interacting Proteins
UNIT 19.2
To understand the function of a particular protein, it is often useful to identify other proteins with which it associates. This can be done by a selection or screen in which novel proteins that specifically interact with a target protein of interest are isolated from a library. One particularly useful approach to detect novel interacting proteins—the twohybrid system or interaction trap (see Figs. 19.2.1 and 19.2.2)—uses yeast as a “test tube” and transcriptional activation of a reporter system to identify associating proteins (see Background Information). This approach can also be used specifically to test complex formation between two proteins for which there is a prior reason to expect an interaction. In the basic version of this method (see Fig. 19.2.2), the plasmid pEG202 or a related vector (see Fig. 19.2.3 and Table 19.2.1) is used to express the probe or “bait” protein as a fusion to the heterologous DNA-binding protein LexA. Many proteins, including transcription factors, kinases, and phosphatases, have been successfully used as bait proteins. The major requirements for the bait protein are that it should not be actively excluded from the yeast nucleus, and it should not possess an intrinsic ability to strongly activate transcription. The plasmid expressing the LexA-fused bait protein (see Table 19.2.1) is used to transform yeast possessing a dual reporter system responsive to transcriptional activation through the LexA operator. In one such example, the yeast strain EGY48 (see Table 19.2.2) contains the reporter plasmid pSH18-34. In this case, binding sites for LexA are located upstream of two reporter genes. In the EGY48 strain, the upstream activating sequences of the chromosomal LEU2 gene—required in the biosynthetic pathway for leucine (Leu)—are replaced with LexA operators (DNA binding sites). pSH18-34 contains a LexA operator–lacZ fusion gene. These two reporters allow selection for transcriptional activation by permitting selection for viability when cells are plated on medium lacking Leu, and discrimination based on color when the yeast is grown on medium containing Xgal (APPENDIX 4A). In Basic Protocol 1, EGY48/pSH18-34 transformed with a bait is characterized for its ability to express protein (Support Protocol 1), growth on medium lacking Leu, and for the level of transcriptional activation of lacZ (see Fig. 19.2.2A). A number of alternative strains, plasmids, and strategies are presented which can be employed if a bait proves to have an unacceptably high level of background transcriptional activation. In an interactor hunt (Basic Protocol 2), the strain EGY48/pSH18-34 containing the bait expression plasmid is transformed (along with carrier DNA made as described in Support Protocol 2) with a conditionally expressed library made in the vector pJG4-5 (see Fig. 19.2.6 and Table 19.2.3). This library uses the inducible yeast GAL1 promoter to express proteins as fusions to an acidic domain (“acid blob”) that functions as a portable transcriptional activation motif (act) and to other useful moieties. Expression of libraryencoded proteins is induced by plating transformants on medium containing galactose (Gal), so yeast cells containing library proteins that do not interact specifically with the bait protein will fail to grow in the absence of Leu (see Fig. 19.2.2B). Yeast cells containing library proteins that interact with the bait protein will form colonies within 2 to 5 days, and the colonies will turn blue when the cells are streaked on medium containing Xgal (see Fig. 19.2.2C). The DNA from interaction trap positive colonies can be analyzed by polymerase chain reaction (PCR) to streamline screening and detect redundant clones in cases where many positives are obtained in screening (see Alternate Protocol 1). The plasmids are isolated and characterized by a series of tests to confirm specificity of the interaction with the initial bait protein (Support Protocols 3 to 5). Those found to be specific are ready for further analysis (e.g., sequencing).
Identification of Protein Interactions
Contributed by Erica A. Golemis, Ilya Serebriiskii, Russell L. Finley, Jr., Mikhail G. Kolonin, Jeno Gyuris, and Roger Brent
19.2.1
Current Protocols in Protein Science (1998) 19.2.1-19.2.40 Copyright © 1998 by John Wiley & Sons, Inc.
Supplement 14
prepare bait strain(s) for interaction mating (Alternate Protocol 2, steps 1-3)
construct bait protein plasmid and transform yeast (Basic Protocol 1, step 1)
characterize bait protein expression and activity
transform cDNA library into lexA operatorLEU2 strain to make pretransformed library strain (Alternate Protocol 2, steps 4-11)
introduce cDNA library into bait strain(s) by interaction mating (Alternate Protocol 2, steps 12-20)
obtain cDNA library in pJG4-5
transform cDNA library into lexA-operator-LEU2/lexA-operator-lacZ/pBait yeast (Basic Protocol 2, steps 1-7 )
select for library plasmid (Basic Protocol 2, step 8)
assess transcriptional activity (Basic Protocol 1, steps 4-7 ) assess repressor activity (Basic Protocol 1, steps 8 -11) test for Leu requirement (Basic Protocol 1, steps 12 -13) assess protein synthesis (Support Protocol 1)
freeze and replate transformants (Basic Protocol 2, steps 9 -15)
select for interacting proteins (Basic Protocol 2, steps 16 -19)
transform E. coli Basic Protocol 2, steps 20-22)
test for specificity (Basic Protocol 2, steps 23 -27, and Support Protocol 5 )
analyze and sequence positive isolates (Basic Protocol 2, step 28, and Support Protocol 5 )
assess whether clones are independent by restriction mapping or by filter hybridization (Support Protocol 3)
or obtain profile of independent interactors by microplate plasmid rescue (Support Protocol 4) or
warehouse clones and repeat screen with less sensitive strain analyze positive clones by PCR and restriction endonuclease digestion (Alternate Protocol 1)
Figure 19.2.1 Flow chart for performing an interaction trap.
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A
B
C act act bait
bait
bait LEU2
LEU2
LEU2
act
act bait
bait lacZ
bait lacZ
lacZ
Figure 19.2.2 The interaction trap. (A) An EGY48 yeast cell containing two LexA operator–responsive reporters, one a chromosomally integrated copy of the LEU2 gene (required for growth on −Leu medium), the second a plasmid bearing a GAL1 promoter–lacZ fusion gene (causing yeast to turn blue on medium containing Xgal). The cell also contains a constitutively expressed chimeric protein, consisting of the DNA-binding domain of LexA fused to the probe or bait protein, shown as being unable to activate either of the two reporters. (B) and (C), EGY48/pSH18-34/pbait-containing yeast have been additionally transformed with an activation domain (act)–fused cDNA library in pJG4-5, and the library has been induced. In (B), the encoded protein does not interact specifically with the bait protein and the two reporters are not activated. In (C), a positive interaction is shown in which the library-encoded protein interacts with bait protein, resulting in activation of the two reporters (arrow), thus causing growth on medium lacking Leu and blue color on medium containing Xgal. Symbols: black rectangle, LexA operator sequence; open circle, LexA protein; open pentagon, bait protein; open rectangle, library protein; shaded box, activator protein (acid blob in Fig. 19.2.6).
When more than one bait will be used to screen a single library, significant time and resources can be saved by performing the interactor hunt by interaction mating (see Alternate Protocol 2). In this protocol, EGY48 is transformed with library DNA and the transformants are collected and frozen in aliquots. For each interactor hunt, an aliquot of the pretransformed EGY48/library strain is thawed and mixed with an aliquot of a bait strain transformed with the bait expression plasmid and pSH18-34. Overnight incubation of the mixture on a YPD plate results in fusion of the two strains to form diploids. The diploids are then exposed to galactose to induce expression of the library-encoded proteins, and interactors are selected in the same manner as in Basic Protocol 2. The advantage to this approach is that it requires only one high-efficiency library transformation for multiple hunts with different baits. It is also useful for bait proteins that are somewhat toxic to yeast; yeast expressing toxic baits can be difficult to transform with the library DNA. CHARACTERIZING A BAIT PROTEIN The first step in an interactor hunt is to construct a plasmid that expresses LexA fused to the protein of interest. This construct is transformed into reporter yeast strains containing LEU2 and lacZ reporter genes, and a series of control experiments is performed to establish whether the construct is suitable as is or must be modified, and whether alternative yeast reporter conditions should be used. These controls establish that the bait protein is made as a stable protein in yeast, that it is capable of entering the nucleus and binding LexA operator sites, and that it does not appreciably activate transcription of the LexA operator–based reporter genes. This last is the most important constraint on use of this system. The LexA-fused bait protein must not activate transcription of either re-
BASIC PROTOCOL 1
Identification of Protein Interactions
19.2.3 Current Protocols in Protein Science
Supplement 14
Table 19.2.1
Interaction Trap Componentsa,b
Plasmid name/source
Selection In yeast
Comment/description
In E. coli
LexA fusion plasmids HIS3 pEG202c,d,e HIS3 pJK202
Apr Apr
pNLexAe
HIS3
Apr
pGildad
HIS3
Apr
pEE202I
HIS3
Apr
pRFHM1e,f (control)
HIS3
Apr
pSH17-4e,f (control) pMW101f
HIS3
Apr
HIS3
Cmr
pMW103f
HIS3
Kmr
pHybLex/Zeof,g Zeor
Zeor
Activation domain fusion plasmids pJG4-5c,d,e,f TRP1 Apr
pJG4-5I
TRP1
Apr
pYESTrpg
TRP1
Apr
pMW102f
TRP1
Kmr
pMW104f
TRP1
Cmr
LacZ reporter plasmids URA3 pSH18-34d,e,f
Apr
pJK103e
URA3
Apr
pRB1840e
URA3
Apr
pMW112f pMW109f
URA3 URA3
Kmr Kmr
Contains an ADH promoter that expresses LexA followed by polylinker Like pEG202, but incorporates nuclear localization sequences between LexA and polylinker; used to enhance translocation of bait to nucleus Contains an ADH promoter that expresses polylinker followed by LexA; for use with baits where amino-terminal residues must remain unblocked Contains a GAL1 promoter that expresses same LexA and polylinker cassette as pEG202; for use with baits whose continuous presence is toxic to yeast An integrating form of pEG202 that can be targeted into HIS3 following digestion with KpnI; for use where physiological screen requires lower levels of bait to be expressed Contains an ADH promoter that expresses LexA fused to the homeodomain of bicoid to produce nonactivating fusion; used as positive control for repression assay, negative control for activation and interaction assays ADH promoter expresses LexA fused to GAL4 activation domain; used as a positive control for transcriptional activation Same as pEG202, but with altered antibiotic resistance markers; basic plasmid used for cloning bait Same as pEG202, but with altered antibiotic resistance markers; basic plasmid used for cloning bait Bait cloning vector compatible with interaction trap and all other two-hybrid systems; minimal ADH promotor expresses LexA followed by extended polylinker Contains a GAL1 promoter that expresses nuclear localization domain, transcriptional activation domain, HA epitope tag, cloning sites; used to express cDNA libraries An integrating form of pJG4-5 that can be targeted into TRP1 by digestion with Bsu36I (New England Biolabs); to be used with pEE202I to study interactions that occur physiologically at low protein concentrations Contains a GAL1 promoter that expresses nuclear localization domain, transcriptional activation domain, V5 epitope tag, multiple cloning sites; contains f1 ori and T7 promoter/flanking site; used to express cDNA libraries (Invitrogen) Same as pJG4-5, but with altered antibiotic resistance markers; no libraries yet available Same as pJG4-5, but with altered antibiotic resistance markers; no libraries yet available Contains 8 LexA operators that direct transcription of the lacZ gene; one of the most sensitive indicator plasmids for transcriptional activation Contains two LexA operators that direct transcription of the lacZ gene; an intermediate reporter plasmid for transcriptional activation Contains 1 LexA operator that directs transcription of the lacZ gene; one of the most stringent reporters for transcriptional activation Same as pSH18-34, but with altered antibiotic resistance marker Same as pJK103, but with altered antibiotic resistance marker continued
19.2.4 Supplement 14
Current Protocols in Protein Science
Table 19.2.1
Plasmid name/source pMW111f pMW107f pMW108f pMW110f pJK101e,f (control)
Interaction Trap Componentsa,b, continued
Selection In yeast
In E. coli
URA3 URA3 URA3 URA3 URA3
Kmr Cmr Cmr Cmr Apr
Comment/description Same as pRB1840, but with altered antibiotic resistance marker Same as pSH18-34, but with altered antibiotic resistance marker Same as pJK103, but with altered antibiotic resistance marker Same as pRB1840, but with altered antibiotic resistance marker Contains a GAL1 upstream activating sequence followed by two lexA operators followed by lacZ gene; used in repression assay to assess bait binding to operator sequences
aAll plasmids contain a 2µm origin for maintenance in yeast, as well as a bacterial origin of replication, except where noted (pEE202I, pJG4.5I). bInteraction Trap reagents represent the work of many contributors: the original basic reagents were developed in the Brent laboratory (Gyuris et al.,
1993). Plasmids with altered antibiotic resistance markers (all pMW plasmids) were constructed at Glaxo in Research Triangle Park, N.C. (Watson et al., 1996). Plasmids and strains for specialized applications have been developed by the following individuals: E. Golemis, Fox Chase Cancer Center, Philadelphia, Pa. (pEG202); J. Kamens, BASF, Worcester, Mass. (pJK202); cumulative efforts of I. York, Dana-Farber Cancer Center, Boston, Mass. and M. Sainz and S. Nottwehr, U. Oregon (pNLexA); D.A. Shaywitz, MIT Center for Cancer Research, Cambridge, Mass. (pGilda); R. Buckholz, Glaxo, Research Triangle Park, N.C. (pEE2021, pJG4-51); J. Gyuris, Mitotix, Cambridge, Mass. (pJG4-5); S. Hanes, Wadsworth Institute, Albany, N.Y. (pSH17-4); R.L. Finley, Wayne State University School of Medicine, Detroit, Mich. (pRFHM1); S. Hanes, Wadsworth Institute, Albany, N.Y. (pSH18-34); J. Kamens, BASF, Worcester, Mass. (pJK101, pJK103); R. Brent, The Molecular Sciences Institute, Berkeley, California (pRB1840). Specialized plasmids not yet commercially available can be obtained by contacting the Brent laboratory at (510) 647-0690 or [email protected] or the Golemis laboratory, (215) 728-2860 or [email protected]. cSequence data are available for pEG202 (pLexA) accession number pending. dPlasmids commercially available from Clontech and OriGene; for Clontech pEG202 is listed as pLexA, pJG4-5 as pB42AD, and pSH18-34 as p8op-LacZ. ePlasmids and strains available from OriGene. fIn pMW plasmids the ampicillin resistance gene (Apr) is replaced with the chloramphenicol resistance gene (Cmr) and the kanamycin resistance gene (Kmr) from pBC SK(+) and pBK-CMV (Stratagene), respectively. The choice between Kmr and Cmr or Apr plasmids is a matter of personal taste; use of basic Apr plasmids is described in the basic protocols. Use of the more recently developed reagents would facilitate the purification of library plasmid in later steps by eliminating the need for passage through KC8 bacteria, with substantial saving of time and effort. Apr has been maintained as marker of choice for the library plasmid because of the existence of multiple libraries already possessing this marker. These plasmids are the basic set of plasmids recommended for use. gPlasmids commercially available from Invitrogen as components of a Hybrid Hunter kit; this kit also includes all necessary positive and negative controls (not listed in this table). See Background Information for further details on commercially available reagents.
porter—the EGY48 strain (or related strain EGY191) that expresses the LexA fusion protein should not grow on medium lacking Leu, and the colonies should be white on medium containing Xgal. The characterized bait protein plasmid is used for Basic Protocol 2 to screen a library for interacting proteins. Materials DNA encoding the protein of interest Plasmids (see Table 19.2.1): pEG202 (see Fig. 19.2.3), pSH18-34 (see Fig. 19.2.4), pSH17-4, pRFHM1, and pJK101 for basic characterization; other plasmids for specific circumstances as described (Clontech, Invitrogen, OriGene, or R. Brent) Yeast strain EGY48 (ura3 trp1 his3 3LexA-operator-LEU2), or EGY191 (ura3 trp1 his3 1LexA-operator-LEU2; Table 19.2.2) Complete minimal (CM) medium dropout plates (APPENDIX 4L), supplemented with 2% (w/v) of the indicated sugars (glucose or galactose), in 100-mm plates: Glu/CM, −Ura, −His Gal/CM, −Ura, −His Gal/CM, −Ura, −His, −Leu Z buffer (APPENDIX 4L) with 1 mg/ml 5-bromo-4-chloro-3-indolyl-β-D-galactosidase (Xgal)
Identification of Protein Interactions
19.2.5 Current Protocols in Protein Science
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Pst I 8672
Aat II 7995 Stu I 7835 Nru I 7765 Pst I 7671 Pst I 7590
Tth 111I 10059 Nar I 180 Nae I 246 SphI 1110 Hin dIII 1514 Mlu I 1616 Pme I 2056 pBR backbone ADHpro EcoRI 2144 Bam HI r Sal I * Ap Nco I lexA Not I Xho I Sal I *2182 ADH ter Pst I 2188 pEG202 SphI 2396 10166 bp
HIS3 Hin dIII 6464 Bst XI 6380 HindIII 6277 Pst I 6089 Bss HII 5960
Sac I 5113
2 µm ori
Xba I 3487
Pst I 4780 Avr II 5004
Polylinker sequence
Sal I* Not I Sal I* EcoRI BamHI NcoI XhoI GAA TTC CCG GGG ATC CGT CGA CCA TGG CGG CCG CTC GAG TCG AC
Figure 19.2.3 LexA-fusion plasmids: pEG202. The strong constitutive ADH promoter is used to express bait proteins as fusions to the DNA-binding protein LexA. Restriction sites shown in this map are based on recently compiled pEG202 sequence data and include selected sites suitable for diagnostic restriction endonuclease digests. A number of restriction sites are available for insertion of coding sequences to produce protein fusions with LexA; the polylinker sequence and reading frame relative to LexA are shown below the map with unique sites marked in bold type. The sequence 5′-CGT CAG CAG AGC TTC ACC ATT G-3′ can be used to design a primer to confirm correct reading frame for LexA fusions. Plasmids contain the HIS3 selectable marker and the 2µm origin of replication to allow propagation in yeast, and the Apr antibiotic resistance gene and the pBR origin of replication to allow propagation in E. coli. In the recently developed LexA-expression plasmids pMW101 and pMW103, the ampicillin resistance gene (Apr) has been replaced with the chloramphenicol resistance gene (Cmr) and the kanamycin resistance gene (Kmr), respectively (see Table 19.2.1 for details).
Gal/CM dropout liquid medium (APPENDIX 4L) supplemented with 2% Gal Antibody to LexA or fusion domain: monoclonal antibody to LexA (Clontech, Invitrogen) or polyclonal antibody to LexA (available by request from R. Brent or E. Golemis) H2O, sterile 30°C incubator Nylon membrane Whatman 3MM filter paper Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins
Additional reagents and equipment for subcloning DNA fragments (Struhl, 1987), lithium acetate transformation of yeast (APPENDIX 4L), liquid assay for β-galactosidase (APPENDIX 4L), preparation of protein extracts for immunoblot analysis (see Support Protocol 1), and immunoblotting and immunodetection (UNIT 10.10)
19.2.6 Supplement 15
Current Protocols in Protein Science
Smal 0.00 Bam HI 0.01 EcoRI 0.01 XhoI 0.28
PstI 9.42 Hind III 9.22 EcoRI 9.19
EcoRI 0.60 Hind III 0.61 Bam HI 0.62
PstI 8.95 URA3
lexA op GAL1pro
2µm
lacZ LacZ reporter 10.3 kb
SacI 2.63
Apr
Hind III 7.05 Eco RI 6.95
pBR ori
Eco RI 3.70
Pst I 6.20
Figure 19.2.4 LacZ reporter plasmid. pRB1840, pJK103, and pSH18-34 are all derivatives of LR1∆1 (West et al., 1984) containing eight, two, or one operator for LexA (LexAop) binding inserted into the unique XhoI site located in the minimal GAL1 promoter (GAL1pro; 0.28 on map). The plasmid contains the URA3 selectable marker, the 2µm origin to allow propagation in yeast, the ampicillin resistance (Apr) gene, and the pBR322 origin (ori) to allow propagation in E. coli. Numbers indicate relative map positions. In the recently developed derivatives, the ampicillin resistance gene (Apr) has been replaced with the chloramphenicol or kanamycin resistance genes (see Table 19.2.1 for details).
NOTE: All solutions and equipment coming into contact with cells must be sterile, and proper sterile technique should be used accordingly. Transform yeast with the bait protein plasmid 1. Using standard subcloning techniques (e.g., Struhl, 1987), insert the DNA encoding the protein of interest into the polylinker of pEG202 (see Fig. 19.2.3) or other LexA fusion plasmid to make an in-frame protein fusion. The LexA fusion protein is expressed from the strong alcohol dehydrogenase (ADH) promoter. pEG202 also contains a HIS3 selectable marker and a 2ìm origin for propagation in yeast. pEG202 with the DNA encoding the protein of interest inserted is designated pBait. Uses of alternative LexA fusion plasmids are described in Background Information.
2. Perform three separate lithium acetate transformations (APPENDIX 4L) of EGY48 using the following combinations of plasmids: pBait + pSH18-34 (test) pSH17-4 + pSH18-34 (positive control for activation) pRFHM1 + pSH18-34 (negative control for activation). Use of the two LexA fusions as positive and negative controls allows a rough assessment of the transcriptional activation profile of LexA bait proteins. pEG202 itself is not a good negative control because the peptide encoded by the uninterrupted polylinker sequences is itself capable of very weakly activating transcription.
Identification of Protein Interactions
19.2.7 Current Protocols in Protein Science
Supplement 17
Table 19.2.2
Interaction Trap Yeast Selection Strainsa
Strain
Relevant genotype
EGY48b,c,d
MATα trp1, his3, ura3, lexAops-LEU2
6
EGY191
MATα trp1, his3, ura3, lexAops-LEU2
2
L40c
MATα trpl, leu2, ade2, GAL4, lexAops-HIS34, lexAops-lacZ8
Number of operators
Comments/description lexA operators direct transcription from the LEU2 gene; EGY48 is a basic strain used to select for interacting clones from a cDNA library EGY191 provides a more stringent selection than EGY48, producing lower background with baits with instrinsic ability to activate transcription Expression driven from GAL1 promoter is constitutive in L40 (inducible in EGY strains); selection is for HIS prototrophy. Integrated lacZ reporter is considerably less sensitive than pSH18-34 maintained in EGY strains
aInteraction Trap reagents represent the work of many contributors: the original basic reagents were developed in the Brent laboratory
(Gyuris et al., 1993). Strains for specialized applications have been developed by the following individuals: E. Golemis, Fox Chase Cancer Center, Philadelphia, Pa. (EGY48, EGY191); A.B. Vojtek and S.M. Hollenberg, Fred Hutchinson Cancer Research Center, Seattle, Wash. (L40). Specialized strains not yet commercially available can be obtained by contacting the Brent laboratory at The Molecular Sciences Institute, Berkeley, (510) 647-0690 or [email protected], or the Golemis laboratory, (215) 728-2860 or [email protected]. bStrains commercially available from Clontech. cStrains commercially available from Invitrogen as components of a Hybrid Hunter kit; the kit also includes all necessary positive and negative controls (not listed in this table). See Background Information for further details on commercially available reagents. dStrains commercially available from OriGene.
pSH18-34 contains a 2ìm origin and a URA3 selectable marker for maintenance in yeast, as well as a bacterial origin of replication and ampicillin-resistance gene. It is the most sensitive lacZ reporter available and will detect any potential ability to activate lacZ transcription. pSH17-4 is a HIS3 2ìm plasmid encoding LexA fused to the activation domain of the yeast activator protein GAL4. This fusion protein strongly activates transcription. pRFHM1 is a HIS3 2ìm plasmid encoding LexA fused to the N-terminus of the Drosophila protein bicoid. This fusion protein has no ability to activate transcription.
3. Plate each transformation mixture on Glu/CM −Ura, −His dropout plates. Incubate 2 days at 30°C to select for yeast that contain both plasmids. Colonies obtained can be used simultaneously in tests for the activation of lacZ (steps 4 to 7) and LEU2 (steps 12 to 13) reporters.
Assay lacZ gene activation by β-galactosidase assay 4. Streak a Glu/CM −Ura, −His master dropout plate with at least five or six independent colonies obtained from each of the three transformations in step 3 (test, positive control, and negative control) and incubate overnight at 30°C. The filter assay described in Steps 5a to 7a (based on Breeden and Nasmyth, 1985) provides a rapid assay for β-galactosidase transcription. Alternatively, a liquid assay (APPENDIX 4L) or a plate assay (described in Steps 5b to 7b) may be used.
Perform filter assay for β-galactosidase activity: 5a. Lift colonies by gently placing a nylon membrane on the yeast plate and allowing it to become wet through. Remove the membrane and air dry 5 min. Chill the membrane, colony side up, 10 min at −70°C. Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins
Whatman 3MM filters can be cut to the size of the yeast plate as a more economical alternative to nylon membranes for performing lifts. In addition, two or three 5-min temperature cycles (−70°C to room temperature) can be used instead of a single cycle to promote better lysis; this may be worth doing if there is difficulty visualizing blue color.
19.2.8 Supplement 17
Current Protocols in Protein Science
6a. Cut a piece of Whatman 3MM filter paper slightly larger than the colony membrane and soak it in Z buffer containing 1 mg/ml Xgal. Place colony membrane, colony side up, on Whatman 3MM paper, or float it in the lid of a petri dish containing ∼2 ml Z buffer with 1 mg/ml Xgal. Acceptable results may be obtained using as little as 300 ìg/ml Xgal.
7a. Incubate at 30°C and monitor for color changes. It is generally useful to check the membrane after 20 min, and again after 2 to 3 hr. Strong activators will produce a blue color in 5 to 10 min, and a bait protein (LexA fusion protein) that does so is unsuitable for use in an interactor hunt using this lacZ reporter plasmid. Weak activators will produce a blue color in 1 to 6 hr (compare versus negative control pRFHMI which will itself produce a faint blue color with time) and may or may not be suitable. Weak activators should be tested using the repressor assay described in steps 8 to 11.
Perform Xgal plate assay for lacZ activation: 5b. Prepare Z buffer Xgal plates as described in APPENDIX 4L. For activation assays, plates should be prepared with glucose as a sugar source. For repression assays (steps 8 to 11), galactose should be used as a sugar source. In our experience, when patching from a master plate to Xgal plates, sufficient yeast are transferred that plasmid loss is not a major problem even in the absence of selection; this is balanced by the desire to assay sets of constructs on the same plate to eliminate batch variation in Xgal potency. Hence, plates should be made either with complete minimal amino acid mix, or by dropping out only uracil (−Ura), to make the plates universally useful.
A +++ endogenous
GAL4 GALUAS
ops
lacZ
plasmid JK101
B
P1
endogenous
L e x A ops
GAL4 GALUAS
+ lacZ
plasmid JK101
Figure 19.2.5 Repression assay for DNA binding. (A) The plasmid JK101 contains the upstream activating sequence (UAS) from the GAL1 gene followed by LexA operators upstream of the lacZ coding sequence. Thus, yeast containing pJK101 will have significant β-galactosidase activity when grown on medium in which galactose is the sole carbon source because of binding of endogenous yeast GAL4 to the GALUAS (B). LexA-fused proteins (P1-LexA) that are made, enter the nucleus, and bind the LexA operator sequences (ops) will block activation from the GALUAS, repressing β-galactosidase activity (+) 3- to 5-fold. On glucose/Xgal medium, yeast containing pJK101 should be white because GALUAS transcription is repressed.
Identification of Protein Interactions
19.2.9 Current Protocols in Protein Science
Supplement 14
6b. Streak yeast from master plate to Xgal plate and incubate at 30°C. 7b. Examine plates for color development at intervals over the next 2 to 3 days. Strongly activating fusions should be visibly blue on the plate within 12 to 24 hr; moderate activators will be visibly blue after ∼2 days. When a bait protein appreciably activates transcription under these conditions, there are several recourses. The first and simplest is to switch to a less sensitive lacZ reporter plasmid; use of pJK103 and pRB1840 may be sufficient to reduce background to manageable levels. If this fails to work, it is frequently possible to generate a truncated LexA fusion that does not activate transcription.
Confirm fusion-protein synthesis by repression assay For LexA fusions that do not activate transcription, confirm by performing a repression assay (Brent and Ptashne, 1984) that the LexA fusion protein is being synthesized in yeast (some proteins are not) and that it is capable of binding LexA operator sequences (Fig. 19.2.5). The following steps can be performed concurrently with the activation assay. 8. Transform EGY48 yeast with the following combinations of plasmids (three transformations): pBait + pJK101 (test) pRFHM1 + pJK101 (positive control for repression) pJK101 alone (negative control for repression). 9. Plate each transformation mix on Glu/CM −Ura, −His dropout plates or Glu/CM −Ura dropout plates as appropriate to select yeast cells that contain the indicated plasmids. Incubate 2 to 3 days at 30°C until colonies appear. 10. Streak colonies to a Glu/CM −Ura, −His or Glu/CM −Ura dropout master plate and incubate overnight at 30°C. 11. Assay β-galactosidase activity of the three transformed strains (test, positive control, and negative control) by liquid assay (using Gal/CM dropout liquid medium), filter assay (steps 5a to 7a, first restreaking to Gal/CM plates to grow overnight), or plate assay (steps 5b to 7b, using Gal/CM −Ura XGal plates). This assay should not be run for more than 1 to 2 hr for membranes, or 36 hr for Xgal plates, as the high basal lacZ activity will make differential activation of pJK101 impossible to see with longer incubations. Use of Xgal plates, and inspection 12 to 24 hr after streaking, is generally most effective. The plasmid pJK101 contains the galactose upstream activating sequence (UAS) followed by LexA operators upstream of the lacZ coding sequence. Thus, yeast containing pJK101 will have significant β-galactosidase activity when grown on medium in which galactose is the sole carbon source because of binding of endogenous yeast GAL4 to the GALUAS. LexA-fused proteins that are made, enter the nucleus, and bind the LexA operator sequences block activation from the GALUAS, repressing β-galactosidase activity 3- to 20-fold. Note that on Glu/Xgal medium, yeast containing pJK101 should be white, because GALUAS transcription is repressed.
12. If a bait protein neither activates nor represses transcription, perform immunoblot analysis by probing an immunoblot of a crude lysate with antibodies against LexA or the fusion domain to test for protein synthesis (see Support Protocol 1).
Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins
Even if a bait protein represses transcription, it is generally a good idea to assay for the production of full-length LexA fusions, as occasionally some fusion proteins will be proteolytically cleaved by endogenous yeast proteases. If the protein is made but does not repress, it may be necessary to clone the sequence into a LexA fusion vector that contains a nuclear localization motif, e.g., pJK202 (see Table 19.2.1), or to modify or truncate the fusion domain to remove motifs that target it to other cellular compartments (e.g., myristoylation signals).
19.2.10 Supplement 14
Current Protocols in Protein Science
Test for Leu requirement These steps can be performed concurrently with the lacZ activation and repression assays. 13. Disperse a colony of EGY48 containing pBait and pSH18-34 reporter plasmids into 500 µl sterile water. Dilute 100 µl of suspension into 1 ml sterile water. Make a series of 1/10 dilutions in sterile water to cover a 1000-fold concentration range. 14. Plate 100 µl from each tube (undiluted, 1/10, 1/100, and 1/1000) on Gal/CM −Ura, −His dropout plates and on Gal/CM −Ura, −His, −Leu dropout plates. Incubate overnight at 30°C. There will be a total of eight plates. Gal/CM −Ura, −His dropout plates should show a concentration range from 10 to 10,000 colonies and Gal/CM −Ura, −His, −Leu dropout plates should have no colonies. Actual selection in the interactor hunt is based on the ability of the bait protein and acid-fusion pair, but not the bait protein alone, to activate transcription of the LexA operator-LEU2 gene and allow growth on medium lacking Leu. Thus, the test for the Leu requirement is the most important test of whether the bait protein is likely to have an unworkably high background. The LEU2 reporter in EGY48 is more sensitive than the pSH18-34 reporter for some baits, so it is possible that a bait protein that gives little or no signal in a β-galactosidase assay would nevertheless permit some level of growth on −Leu medium. If this occurs, there are several options for proceeding, the most immediate of which is to substitute EGY191 (see Table 19.2.2), a less sensitive screening strain, and repeat the assay. As outlined in this protocol, the authors recommend the strategy of performing the initial screening using the most sensitive reporters and then, if activation is detected, screening with increasingly less sensitive reporters (see Critical Parameters for further discussion).
PERFORMING AN INTERACTOR HUNT An interactor hunt involves two successive large platings of yeast containing LexA-fused probes and reporters and libraries in pJG4-5 (Fig. 19.2.6, Table 19.2.3) with a cDNA expression cassette under control of the GAL promoter. In the first plating, yeast are plated on complete minimal (CM) medium −Ura, −His, −Trp dropout plates with glucose (Glu) as a sugar source to select for the library plasmid. In the second plating, which selects for yeast that contain interacting proteins, a slurry of primary transformants is plated on CM −Ura, −His, −Trp, −Leu dropout plates with galactose/raffinose (Gal/Raff) as the sugar source. This two-step selection is encouraged for two reasons. First, a number of interesting cDNA-encoded proteins may be deleterious to the growth of yeast that bear them; these would be competed out in an initial mass plating. Second, it seems likely that immediately after simultaneous transformation and Gal induction, yeast bearing particular interacting proteins may not be able to initially express sufficient levels of these proteins to support growth on medium lacking Leu. Library plasmids from colonies identified in the second plating are purified by bacterial transformation and used to transform yeast cells for the final specificity screen.
BASIC PROTOCOL 2
A list of libraries currently available for use with this system is provided in Table 19.2.3. The protocol outlined below describes the steps used to perform a single-step screen that should saturate a library derived from a mammalian cell. For screens with libraries derived from lower eukaryotes with less complex genomes, fewer plates will be required. Occasionally, baits that seemed well-behaved during preliminary tests produce unworkably high backgrounds of “positives” during an actual screen (see Background Information and Critical Parameters). To forestall the waste of time and materials performing a screen with such a bait would entail, an alternative approach is to perform a scaled-back
Identification of Protein Interactions
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Supplement 14
screen when working with a new bait (e.g., 5 rather than 30 plates of primary transformants). The results can be assessed before doing a full screen; it is then possible to switch to lower-sensitivity reporter strains and plasmids, if appropriate. Although individual baits will vary, the authors’ current default preference is to use the lacZ reporter pJK103 in conjunction with either EGY48 or EGY191. Polymerase chain reaction (PCR) can also be used in a rapid screening approach that may be preferable if a large number of positions are obtained in a library screen (see Alternate Protocol 1).
SacI 6440 PvuII 6253 Afl III 6075
KpnI 6446
HindIII 528 EcoRI 849 XhoI 861 GAL pro Alw NI 5661 HindIII 867 fusion SphI 1191 cassette BamHI 1330 pUC backbone XbaI 1336 ADH ter SalI 1342 NotI 1350 PstI 1364 Ap HindIII 1474 pJG4-5 6449 bp ScaI 4704
2µm ori
AatII 4264
XbaI 2072
TRP1 XbaI 4002 HindIII 3573
PstI 3365
Fusion cassette NLS
HA Tag
B42 domain
EcoRI
XhoI
ATG GGT GCT CCT CCA AAA AAG AAG ... CCC GAA TTC GGC CGA CTC GAG AAG CTT ... M G A P P K K K ... P E F G R L E K L ...
Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins
Figure 19.2.6 Library plasmids: pJG4-5. Library plasmids express cDNAs or other coding sequences inserted into unique EcoRI and XhoI sites as a translational fusion to a cassette consisting of the SV40 nuclear localization sequence (NLS; PPKKKRKVA), the acid blob B42 domain (Ruden et al, 1991), and the hemagglutinin (HA) epitope tag (YPYDVPDYA). Expression of cassette sequences is under the control of the GAL1 galactose-inducible promoter. This map is based on the sequence data available for pJG4-5, and includes selected sites suitable for diagnostic restriction digests (shown in bold). The sequence 5′-CTG AGT GGA GAT GCC TCC-3′ can be used as a primer to identify inserts or to confirm correct reading frame. The pJG4-5 plasmid contains the TRP1 selectable marker and the 2µm origin to allow propagation in yeast, and the antibiotic resistance gene and the pUC origin to allow propagation in E. coli. In the recently developed pJG4-5 derivative plasmids pMW104 and pMW102, the ampicillin resistance gene (Apr) has been replaced with the chloramphenicol resistance gene (Cmr) and the kanamycin resistance gene (Kmr), respectively (see Table 19.2.2 for details). Currently existing libraries are all made in the pJG4-5 plasmid (Gyuris et al., 1993) shown on this figure. Unique sites are marked in bold type.
19.2.12 Supplement 14
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Materials Yeast containing appropriate combinations of plasmids (see Table 19.2.1 and Table 19.2.2): EGY48 containing LexA-operator-lacZ reporter and pBait (see Basic Protocol 1) EGY48 containing LexA-operator-lacZ reporter and pRFHM-1 EGY48 containing LexA-operator-lacZ reporter and any nonspecific bait Complete minimal (CM) dropout liquid medium (APPENDIX 4L) supplemented with sugars (glucose, galactose, and/or raffinose) as indicated [2% (w/v) Glu, or 2% (w/v) Gal + 1% (w/v) Raff]: Glu/CM −Ura, −His Glu/CM −Trp Gal/Raff/CM −Ura, −His, −Trp H2O, sterile TE buffer (pH 7.5; APPENDIX 2E)/0.1 M lithium acetate Library DNA in pJG4-5 (Table 19.2.3 and Fig. 19.2.6) High-quality sheared salmon sperm DNA (see Support Protocol 2) 40% (w/v) polyethylene glycol 4000 (PEG 4000; filter sterilized)/0.1 M lithium acetate/TE buffer (pH 7.5) Dimethyl sulfoxide (DMSO) Complete minimal (CM) medium dropout plates (APPENDIX 4L) supplemented with sugars and Xgal (20 µg/ml) as indicated [2% (w/v) Glu, and 2% (w/v) Gal + 1% (w/v) Raff]: Glu/CM −Ura, −His, −Trp, 24 × 24–cm (Nunc) and 100-mm Gal/Raff/CM −Ura, −His, −Trp, 100-mm Gal/Raff/CM −Ura, −His, −Trp, −Leu, 100-mm Glu/Xgal/CM −Ura, −His, −Trp, 100-mm Gal/Raff/Xgal/CM −Ura, −His, −Trp, 100-mm Glu/CM −Ura, −His, −Trp, −Leu, 100-mm Glu/CM −Ura, −His, 100-mm Gal/CM −Ura, −His, −Trp, −Leu, 100-mm TE buffer (pH 7.5), sterile (optional) Glycerol solution (see recipe) E. coli KC8 (pyrF leuB600 trpC hisB463; constructed by K. Struhl and available from R. Brent) LB/ampicillin plates (APPENDIX 4A) E. coli DH5α or other strain suitable for preparation of DNA for sequencing Bacterial defined minimal A medium plates: 1× A medium plates containing 0.5 µg/ml vitamin B1 (APPENDIX 4A) and supplemented with 40 µg/ml each Ura, His, and Leu 30°C incubator, with and without shaking Low-speed centrifuge and rotor 50-ml conical tubes, sterile 1.5-ml microcentrifuge tubes, sterile 42°C heating block Glass microscope slides, sterile Additional reagents and equipment for rapid miniprep isolation of yeast DNA (APPENDIX 4L), transformation of bacteria by electroporation (UNIT 5.10), miniprep isolation of bacterial DNA (APPENDIX 4C), restriction endonuclease digestion (APPENDIX 4I; optional), and agarose gel electrophoresis (APPENDIX 4F; optional) NOTE: All solutions and equipment coming into contact with cells must be sterile, and proper sterile technique should be used accordingly.
Identification of Protein Interactions
19.2.13 Current Protocols in Protein Science
Supplement 14
Table 19.2.3 Libraries Compatible with the Interaction Trap Systema
Source of RNA/DNA Cell lines HeLa cells (human cervical carcinoma)
Independent clones
Insert size (average)b Contact information
JG
9.6 × 106
0.3-3.5 kb (1.5 kb)
Y JG
3.7 × 106 5.7 × 106
0.3-1.2 kb 0.3-3.5 kb (1.5 kb)
JG
4.0 × 106
0.7-2.8 kb (1.5 kb)
R. Brent, Clontech, Invitrogen, OriGene Invitrogen R. Brent, Clontech, OriGene R. Brent
Y Y JG JG Y
3.2 × 106 3.0 × 106 5.7 × 106 2 × 106 5.4 × 106
0.3-1.2 kb 0.5-4.0 kb (1.8 kb) (>1.3) 0.7-3.5 kb (1.2 kb) 0.3-0.8 kb
Invitrogen Clontech OriGene S. Witte Invitrogen
JG JG JG
4.0 × 106 >106 1.5 × 106
0.4-4.0 kb (2.0 kb) 0.3-2.5 kb (>0.5 kb) 0.3-3.5 kb
Clontech R. Brent R. Brent
Vector
HeLa cells (human cervical carcinoma) WI-38 cells (human lung fibroblasts), serum-starved, cDNA Jurkat cells (human T cell leukemia), exponentially growing, cDNA Jurkat cells (human T cell leukemia) Jurkat cells (human T cell leukemia) Jurkat cells (human T cell leukemia) Jurkat cells (human T cell leukemia) Be Wo cells (human fetal placental choriocarcinoma) Human lymphocyte CD4+ T cell, murine, cDNA Chinese hamster ovary (CHO) cells, exponentially growing, cDNA A20 cells (mouse B cell lymphoma) Human B cell lymphoma Human 293 adenovirus–infected (early and late stages) SKOV3 human Y ovarian cancer MDBK cell, bovine kidney MDCK cells HepG2 cell line cDNA MCF7 breast cancer cells, untreated MCF7 breast cancer cells, estrogen-treated MCF7 cells, serum-grown LNCAP prostate cell line, untreated LNCAP prostate cell line, androgen-treated Mouse pachytene spermatocytes
Y JG JG
3.11 × 106 — —
0.3-1.2 kb — —
Invitrogen H. Niu K. Gustin
Y JG JG JG JG JG JG JG JG JG
5.0 × 106 5.8 × 106 — 2 × 106 1.0 × 107 1.0 × 107 1.0 ×107 2.9 × 106 4.6 × 106 —
(>1.4 kb) (>1.2 kb) — — (>1.5 kb) (>1.1 kb) 0.4-3.5 kb (>0.8 kb) (>0.9 kb) —
OriGene OriGene D. Chen M. Melegari OriGene OriGene OriGene OriGene OriGene C. Hoog
Tissues Human breast Human breast tumor Human liver Human liver
Y Y JG Y
9 × 106 8.84 × 106 >106 2.2 × 106
Invitrogen Invitrogen R. Brent Clontech
Human liver Human liver Human lung Human lung tumor Human brain Human brain Human testis Human testis Human ovary
JG JG Y Y JG Y Y JG Y
3.2 × 106 1.1 × 107 5.9 × 106 1.9 × 106 3.5 × 106 8.9 × 106 6.4 × 106 3.5 × 106 4.6 × 106
0.4-1.2 kb 0.4-1.2 kb 0.6-4.0 kb (>1 kb) 0.5-4 kb (1.3 kb) 0.3-1.2 kb (> 1 kb) 0.4-1.2 kb 0.4-1.2 0.5-4.5 kb (1.4 kb) 0.3-1.2 kb 0.3-1.2 kb 0.4-4.5 kb (1.6 kb) 0.3-1.2 kb
Invitrogen OriGene Invitrogen Invitrogen Clontech Invitrogen Invitrogen Clontech Invitrogen continued
19.2.14 Supplement 14
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Table 19.2.3 Libraries Compatible with the Interaction Trap Systema, continued
Source of RNA/DNA
Vector
Independent clones
Insert size (average)b Contact information
Human ovary Human ovary Human heart Human placenta Human placenta Human mammary gland Human peripheral blood leucocyte Human kidney Human fetal kidney Human spleen Human prostate Human normal prostate Human prostate Human prostate cancer Human fetal prostate Human fetal liver Human fetal liver Human fetal liver Human fetal brain
JG JG JG Y JG JG JG JG JG Y Y JG JG JG JG JG Y JG JG
4.6 × 106 3.5 × 106 3.0 × 106 4.8 × 106 3.5 × 106 3.5 × 106 1.0 × 107 3.5 × 106 3.0 × 106 1.14 × 107 5.5 × 106 1.4 × 106 1.4 × 106 1.1 × 106 — 3.5 × 106 2.37 × 106 8.6 × 106 3.5 × 106
(>1.3 kb) 0.5-4.0 kb (1.8 kb) 0.3-3.5 kb (1.3 kb) 0.3-1.2 kb 0.3-4.0 kb (1.2 kb) 0.5-5 kb (1.6 kb) (>1.3 kb) 0.4-4.5 kb (1.6 kb) (>1 kb) 0.4-1.2 kb 0.4-1.2 kb 0.4-4.5 kb (1.7 kb) (>1 kb) (>0.9 kb) — 0.3-4.5 kb (1.3 kb) 0.3-1.2 kb (>1 kb) 0.5-1.2 kb (1.5 kb)
Mouse brain Mouse brain Mouse breast, lactating Mouse breast, involuting Mouse breast, virgin Mouse breast, 12 days pregnant Mouse skeletal muscle Rat adipocyte, 9-week-old Zucker rat Rat brain Rat brain (day 18) Rat testis Rat thymus Mouse liver Mouse spleen Mouse ovary Mouse prostate Mouse embryo, whole (19-day) Mouse embryo Drosophila melanogaster, adult, cDNA D. melanogaster, embryo, cDNA D. melanogaster, 0-12 hr embryos, cDNA D. melanogaster, ovary, cDNA D. melanogaster, disc, cDNA D. melanogaster, head
JG JG JG JG JG JG JG JG JG JG JG JG JG JG JG JG JG JG JG JG JG
6.1 × 106 4.5 × 106 1.0 × 107 1.0 × 107 1.0 × 107 6.3 × 106 7.2 × 106 1.0 × 107 4.5 ×106 — 8.0 × 106 8.2 × 106 9.5 × 106 1.0 × 107 4.0 × 106 — 1.0 × 105 3.6 × 106 1.8 × 106 3.0 × 106 4.2 × 106
(>1 kb) 0.4-4.5 kb (1.2 kb) 0.4-3.1 kb 0.4-7.0 kb 0.4-5.5 kb 0.4-5.3 kb 0.4-3.5 kb 0.4-5.0 kb 0.3-3.4 kb — (>1.2 kb) (>1.3 kb) (>1.4 kb) (>1 kb) (>1.2 kb) — 0.2-2.5 kb 0.5-5 kb (1.7 kb) (>1.0 kb) 0.5-3.0 kb (1.4 kb) 0.5-2.5 kb (1.0 kb)
OriGene Clontech Clontech Invitrogen Clontech Clontech OriGene Clontech OriGene Invitrogen Invitrogen Clontech OriGene OriGene OriGene Clontech Invitrogen OriGene R. Brent, Clontech, Invitrogen, OriGene OriGene Clontech OriGene OriGene OriGene OriGene OriGene OriGene OriGene H. Niu OriGene OriGene OriGene OriGene OriGene OriGene OriGene Clontech OriGene Clontech R. Brent
JG JG JG
3.2 × 106 4.0 × 106 —
0.3-1.5 kb (800 bp) 0.3-2.1 kb (900 bp) —
R. Brent R. Brent M. Rosbash continued
19.2.15 Current Protocols in Protein Science
Supplement 15
Table 19.2.3 Libraries Compatible with the Interaction Trap Systema, continued
Source of RNA/DNA
Vector
Miscellaneous Synthetic aptamers Saccharomyces cerevisiae, S288C, genomic S. cerevisiae, S288C, genomic Sea urchin ovary Caenorhabditis elegans Agrobacterium tumefaciens Arabidopsis thaliana, 7-day-old seedlings Tomato (Lycopersicon esculentum) Xenopus laevis embryo
PJM-1 JG JG JG JG JG JG JG JG
Independent clones
>1× 109 >3 × 106 4.0 × 106 3.5 × 106 3.8 × 106 — — 8 × 106 2.2 × 106
Insert size (average)b Contact information
60 bp 0.8-4.0 kb 0.5-4.0 kb (1.7 kb) (>1.2 kb) — — — 0.3-4 kb (1.0 kb)
R. Brent R. Brent OriGene Clontech OriGene — H.M. Goodman G.B. Martin Clontech
aMost libraries are constructed in either the pJG4-5 vector or the pYESTrp vector (JG or Y in the Vector column); the peptide aptamer library is made in
the pJM-1 vector. Libraries available from the public domain were constructed by the following individuals: (1) J. Gyuris; (3) C. Sardet and J. Gyuris; (4) W. Kolanus, J. Gyuris, and B. Seed; (39) D. Krainc; (50-52) R. Finley; (55) P. Watt; (54) P. Colas, B. Cohen, T. Jessen, I. Grishina, J. McCoy, and R. Brent (Colas et al., 1996). All libraries mentioned above were constructed in conjunction with and are available from the laboratory of Roger Brent, (510) 647-0690 or [email protected]. The following individual investigators must be contacted directly: (18) J. Pugh, Fox Chase Cancer Center, Philadelphia, Pa.; (8,9) Vinyaka Prasad, Albert Einstein Medical Center New York, N.Y.; (57, 58) Gregory B. Martin, [email protected]; (11) Huifeng Niu, [email protected]; (16) Christer Hoog, [email protected]; (12) Kurt Gustin, [email protected]; (6) Stephan Witte, [email protected]. bInsert size ranges for pJG4-5 based libraries originally constructed in the Brent laboratory, which are now commercially available from Clontech, were reestimated by the company.
Transform the library 1. Grow an ∼20-ml culture of EGY48 or EGY191 containing a LexA-operator-lacZ reporter plasmid and pBait in Glu/CM −Ura, −His liquid dropout medium overnight at 30°C. For best results, the pBait and lacZ reporter plasmids should have been transformed into the yeast within ∼7 to 10 days of commencing a screen.
2. In the morning, dilute culture into 300 ml Glu/CM −Ura, −His liquid dropout medium to 2 × 106 cell/ml (OD600 = ∼0.10). Incubate at 30°C until the culture contains ∼1 × 107 cells/ml (OD600 = ∼0.50). 3. Centrifuge 5 min at 1000 to 1500 × g in a low-speed centrifuge at room temperature to harvest cells. Resuspend in 30 ml sterile water and transfer to 50-ml conical tube. 4. Centrifuge 5 min at 1000 to 1500 × g. Decant supernatant and resuspend cells in 1.5 ml TE buffer/0.1 M lithium acetate. 5. Add 1 µg library DNA in pJG4-5 and 50 µg high-quality sheared salmon sperm carrier DNA to each of 30 sterile 1.5-ml microcentrifuge tubes. Add 50 µl of the resuspended yeast solution from step 4 to each tube. The total volume of library and salmon sperm DNA added should be <20 ìl and preferably <10 ìl.
Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins
A typical library transformation will result in 2 to 3 × 106 primary transformants. Assuming a transformation efficiency of 105/ìg library DNA, this transformation requires a total of 20 to 30 ìg library DNA and 1 to 2 mg carrier DNA. Doing transformations in small aliquots helps reduce the likelihood of contamination, and for reasons that are not clear, provides significantly better transformation efficiency than scaled-up versions. Do not use excess transforming library DNA per aliquot of competent yeast cells because each competent cell may take up multiple library plasmids, complicating subsequent analysis.
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6. Add 300 µl of sterile 40% PEG 4000/0.1 M lithium acetate/TE buffer, pH 7.5, and invert to mix thoroughly. Incubate 30 min at 30°C. 7. Add DMSO to 10% (∼40 µl per tube) and invert to mix. Heat shock 10 min in 42°C heating block. 8a. For 28 tubes: Plate the complete contents of one tube per 24 × 24–cm Glu/CM −Ura, −His, −Trp dropout plate and incubate at 30°C. 8b. For two remaining tubes: Plate 360 µl of each tube on 24 × 24–cm Glu/CM −Ura, −His, −Trp dropout plate. Use the remaining 40 µl from each tube to make a series of 1/10 dilutions in sterile water. Plate dilutions on 100-mm Glu/CM −Ura, −His, −Trp dropout plates. Incubate all plates 2 to 3 days at 30°C until colonies appear. The dilution series gives an idea of the transformation efficiency and allows an accurate estimation of the number of transformants obtained.
Collect primary transformant cells Conventional replica plating (Treco and Winston, 1992) does not work well in the selection process because so many cells are transferred to new plates that very high background levels inevitably occur. Instead, the procedure described below creates a slurry in which cells derived from >106 primary transformants are homogeneously dispersed. A precalculated number of these cells is plated for each primary transformant. 9. Cool all of the 24 × 24–cm plates containing transformants for several hours at 4°C to harden agar. 10. Wearing gloves and using a sterile glass microscope slide, gently scrape yeast cells off the plate. Pool cells from the 30 plates into one or two sterile 50-ml conical tubes. This is the step where contamination is most likely to occur. Be careful.
11. Wash cells by adding a volume of sterile TE buffer or water at least equal to the volume of the transferred cells. Centrifuge ∼5 min at 1000 to 1500 × g, room temperature, and discard supernatant. Repeat wash. After the second wash, pellet volume should be ∼25 ml cells derived from 1.5 × 106 transformants.
12. Resuspend pellet in 1 vol glycerol solution, mix well, and store up to 1 year in 1-ml aliquots at −70°C. Determine replating efficiency 13. Remove an aliquot of frozen transformed yeast and dilute 1/10 with Gal/Raff/CM −Ura, −His, −Trp dropout medium. Incubate with shaking 4 hr at 30°C to induce the GAL promoter on the library. Raffinose (Raff) aids in growth without diminishing transcription from the GAL1 promoter.
14. Make serial dilutions of the yeast cells using the Gal/Raff/CM −Ura, −His, −Trp dropout medium. Plate on 100-mm Gal/Raff/CM −Ura, −His, −Trp dropout plates and incubate 2 to 3 days at 30°C until colonies are visible. 15. Count colonies and determine the number of colony-forming units (cfu) per aliquot of transformed yeast. In calculating yeast concentrations, it is useful to remember that 1 OD600 unit = ∼2.0 × 107 yeast cells. In general, if the harvest is done carefully, viability will be greater than 90%. Some intrepid investigators perform this step simultaneously with plating out on Leu selective medium (steps 16 and 17).
Identification of Protein Interactions
19.2.17 Current Protocols in Protein Science
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Screen for interacting proteins 16. Thaw the appropriate quantity of transformed yeast based on the plating efficiency, dilute, and incubate as in step 13. Dilute cultures in Gal/Raff/CM −Ura, −His, −Trp, −Leu medium as necessary to obtain a concentration of 107 cells/ml (OD600 = ∼0.5), and plate 100 µl on each of as many 100-mm Gal/Raff/CM −Ura, −His, −Trp, −Leu dropout plates as are necessary for full representation of transformants. Incubate 2 to 3 days at 30°C until colonies appear. Because not all cells that contain interacting proteins plate at 100% efficiency on −Leu medium (Estojak et al., 1995), it is desirable that for actual selection, each primary colony obtained from the transformation be represented on the selection plate by three to ten individual yeast cells. This will in some cases lead to multiple isolations of the same cDNA; however, because the slurry is not perfectly homogenous, it will increase the likelihood that all primary transformants are represented by at least one cell on the selective plate. It is easiest to visually scan for Leu+ colonies using cells plated at ∼106 cfu per 100-mm plate. Plating at higher density can contribute to cross-feeding between yeast, resulting in spurious background growth. Thus, for a transformation in which 3 × 106 colonies are obtained, plate ∼2 × 107 cells on a total of 20 selective plates.
17. Carefully pick appropriate colonies to a new Gal/Raff/CM −Ura, −His, −Trp, −Leu master dropout plate. Incubate 2 to 7 days at 30°C until colonies appear. A good strategy is to pick a master plate with colonies obtained on day 2, a second master plate (or set of plates) with colonies obtained on day 3, and a third with colonies obtained on day 4. Colonies from day 2 and 3 master plates should generally be characterized further. If many apparent positives are obtained, it may be worth making master plates of the much larger number of colonies likely to be obtained at day 4 (and after). See Critical Parameters and annotation to step 19 for additional information about appropriate colony selection for the master plate. If no colonies appear within a week, those arising at later time points are likely to be artifactual. Contamination that has occurred at an earlier step (e.g., during plate scraping) is generally reflected by the growth of a very large number of colonies (>500/plate) within 24 to 48 hr after plating on selective medium. Some investigators omit use of a Gal/Raff/CM −Ura, −His, −Trp, −Leu master plate, restreaking directly to a Glu/CM −Ura, −His, −Trp master plate as in step 19.
Test for Gal dependence The following steps test for Gal dependence of the Leu+ insert and lacZ phenotypes to confirm that they are attributable to expression of the library-encoded proteins. The GAL1 promoter is turned off and −Leu selection eliminated before reinducing. 18. Restreak from the Gal/Raff/CM −Ura, −His, −Trp, −Leu master dropout plate to a 100-mm Glu/CM −Ura, −His, −Trp master dropout plate. Incubate overnight at 30°C until colonies form. 19. Restreak or replica plate from this plate to the following plates: Glu/Xgal/CM −Ura, −His, −Trp Gal/Raff/Xgal/CM −Ura, −His, −Trp Glu/CM −Ura, −His, −Trp, −Leu Gal/Raff/CM −Ura, −His, −Trp, −Leu. Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins
At this juncture, colonies and the library plasmids they contain are tentatively considered positive if they are blue on Gal/Raff/Xgal plates but not blue or only faintly blue on Glu/Xgal plates, and if they grow on Gal/Raff/CM −Leu plates but not on Glu/CM −Leu plates. The number of positives obtained will vary drastically from bait to bait. How they are processed subsequently will depend on the number initially obtained and on the preference
19.2.18 Supplement 14
Current Protocols in Protein Science
of the individual investigator. If none are obtained using EGY48 as reporter strain, it may be worth attempting to screen a library from an additional tissue source. If a relatively small number (≤30) are obtained, proceed to step 20. However, sometimes searches will yield large numbers of colonies (>30 to 300, or more). In this case, there are several options. The first option is to warehouse the majority of the positives and work up the first 30 that arise; those growing fastest are frequently the strongest interactors. These can be checked for specificity, and restriction digests can be used to establish whether they are all independent cDNAs or represent multiple isolates of the same, or a small number, of cDNAs. If the former is true, it may be advisable to repeat the screen in a less sensitive strain background, as obtaining many different interactors can be a sign of low-affinity nonspecific background. Alternatively, if initial indications are that a few cDNAs are dominating the positives obtained, it may be useful to perform a filter hybridization with yeast (see Support Protocol 3) using these cDNAs as a probe to establish the frequency of their identification and exclude future reisolation of these plasmids. The second major option is to work up large numbers of positives to get a complete profile of isolated interactors (see Support Protocol 4). A third option is to temporarily warehouse the entire results of this first screen, and repeat the screen with a less sensitive strain such as EGY191, on the theory that it is most important to get stronger interactors first and a complete profile of interactors later. Finally, some investigators prefer to work up the entire set of positives initially obtained, even if such positives number in the hundreds. Particularly in this latter case, it is most effective to use Alternate Protocol 1 as a means to identify unique versus common positives.
Isolate plasmid from positive colonies by transfer into E. coli 20a. Transfer yeast plasmids directly into E. coli by following the protocol for direct electroporation (UNIT 5.10). Proceed to step 22. 20b. Isolate plasmid DNA from yeast by the rapid miniprep protocol (APPENDIX 4C) with the following alteration: after obtaining aqueous phase, precipitate by adding sodium acetate to 0.3 M final and 2 vol ethanol, incubate 20 min on ice, microcentrifuge 15 min at maximum speed, wash pellet with 70% ethanol, dry, and resuspend in 5 µl TE buffer. Cultures can be grown prior to the miniprep using Glu/CM −Trp to select only for the library plasmid; this may increase the proportion of bacterial colonies that contain the desired plasmid.
21. Use 1 µl DNA to electroporate (UNIT 5.10) into competent KC8 bacteria, and plate on LB/ampicillin plates. Incubate overnight at 37°C. Electroporation must be used to obtain transformants with KC8 because the strain is generally refractory to transformation.
22. Restreak or replica plate colonies arising on LB/ampicillin plates to bacterial defined minimal A medium plates containing vitamin B1 and supplemented with Ura, His, and Leu but lacking Trp. Incubate overnight at 37°C. Colonies that grow under these conditions contain the library plasmid. The yeast TRP1 gene can successfully complement the bacterial trpC-9830 mutation, allowing the library plasmid to be easily distinguished from the other two plasmids contained in the yeast. It is helpful to first plate transformations on LB/ampicillin plates, which provides a less stringent selection, followed by restreaking to bacterial minimal medium to maximize the number of colonies obtained (E.G., unpub. observ.).
23. Purify library-containing plasmids using a bacterial miniprep procedure (APPENDIX 4C). Some investigators are tempted to immediately sequence DNAs obtained at this stage. At this point, it is still possible that none of the isolated clones will express bona fide
Identification of Protein Interactions
19.2.19 Current Protocols in Protein Science
Supplement 14
interactors, and it is suggested that the following specificity tests be completed before committing the effort to sequencing (also see annotation to step 28). Because multiple 2ìm plasmids with the same marker can be simultaneously tolerated in yeast, it sometimes happens that a single yeast will contain two or more different library plasmids, only one of which encodes an interacting protein. The frequency of this occurrence varies in the hands of different investigators and may in some cases account for disappearing positives if the wrong cDNA is picked. When choosing colonies to miniprep, it is generally useful to work up at least two individual bacterial transformants for each yeast positive. These minipreps can then be restriction digested (APPENDIX 4I) with EcoRI + XhoI to release cDNA inserts, and the size of inserts determined on an agarose minigel (APPENDIX 4F) to confirm that both plasmids contain the same insert. An additional benefit of analyzing insert size is that it may provide some indication as to whether repeated isolation of the same cDNA is occurring, generally a good indication concerning the biological relevance of the interactor. See Background Information for further discussion.
Assess positive colonies with specificity tests Much spurious background will have been removed by the previous series of controls. Other classes of false positives can be eliminated by retransforming purified plasmids into “virgin” LexA-operator-LEU2/LexA-operator-lacZ/pBait–containing strains that have not been subjected to Leu selection and verifying that interaction-dependent phenotypes are still observed. Such false positives could include mutations in the initial EGY48 yeast that favor growth on Gal medium, library-encoded cDNAs that interact with the LexA DNA-binding domain, or proteins that are sticky and interact with multiple biologically unrelated fusion domains. 24. In separate transformations, use purified plasmids from step 23 to transform yeast that already contain the following plasmids and are growing on Glu/CM −Ura, −His plates: EGY48 containing pSH18-34 and pBait EGY48 containing pSH18-34 and pRFHM-1 EGY48 containing pSH18-34 and a nonspecific bait (optional). 25. Plate each transformation mix on Glu/CM −Ura, −His, −Trp dropout plates and incubate 2 to 3 days at 30°C until colonies appear. 26. Create a Glu/CM −Ura, −His, −Trp master dropout plate for each library plasmid being tested. Streak adjacently five or six independent colonies derived from each of the transformation plates. Incubate overnight at 30°C. 27. Restreak or replica plate from this master dropout plate to the same series of test plates used for the actual screen: Glu/Xgal/CM −Ura, −His, −Trp Gal/Raff/Xgal/CM −Ura, −His, −Trp Glu/CM −Ura, −His, −Trp, −Leu Gal/CM −Ura, −His, −Trp, −Leu. True positive cDNAs should make cells blue on Gal/Raff/Xgal but not on Glu/Xgal plates, and should make them grow on Gal/Raff/CM −Leu but not Glu/CM −Leu dropout plates only if the cells contain LexA-bait. cDNAs that meet such criteria are ready to be sequenced (see legend to Fig. 19.2.3 for primer sequence) or otherwise characterized. Those cDNAs that also encode proteins that interact with either RFHM-1 or another nonspecific bait should be discarded. Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins
It may be helpful to cross-check the isolated cDNAs with a database of cDNAs thought to be false positives. This database is available on the World Wide Web as a work in progress at http://www.fccc.edu:80/research/labs/golemis/InteractionTrapInWork.html. cDNAs reported to this database are generally those isolated only once in a screen in which obviously
19.2.20 Supplement 14
Current Protocols in Protein Science
true interactive partners were isolated multiple times, cDNAs that may interact with more than one bait, or cDNAs for which the interaction does not appear to make biological sense in the context of the starting bait. Although some proteins in this database may ultimately turn out in fact to associate with the bait that isolated them, they are by default unlikely to possess a unique and interesting function in the context of that bait if they are well represented in the database.
28. If appropriate, conduct additional specificity tests (see Support Protocol 5). Analyze and sequence positive isolates. The primer sequence for use with pJG4-5 is provided in the legend to Figure 19.2.4. DNA prepared from KC8 is generally unsuitable for dideoxy or automated sequencing even after use of Qiagen columns and/or cesium chloride gradients. Library plasmids to be sequenced should be retransformed from the KC8 miniprep stock (step 23) to a more amenable strain, such as DH5α, before sequencing is attempted.
RAPID SCREEN FOR INTERACTION TRAP POSITIVES Under some circumstances, it may be desirable to attempt the analysis of a large number of positives resulting from a two-hybrid screen. One such hypothetical example would be a bait with a leucine zipper or coiled coil known to dimerize with partner “A” that is highly expressed. In order to identify the rare novel partner “B”, it is necessary to work through the high background of “A” reisolates. This protocol uses the polymerase chain reaction (PCR) in a strategy to sort positives into redundant (multiple isolates) and unique classes prior to plasmid rescue from yeast, thus greatly reducing the number of plasmid isolations that must be performed. An additional benefit is that this protocol preidentifies positive clones containing one or multiple library plasmids; for those containing only one library plasmid, only a single colony needs to be prepared through KC8/DH5α.
ALTERNATE PROTOCOL 1
Additional Materials (also see Basic Protocol 2) Yeast plated on Glu/CM −Ura, −His, −Trp master plate (see Basic Protocol 2, step 19) Lysis solution (see recipe) 10 µM forward primer (FP1): 5′-CGT AGT GGA GAT GCC TCC-3′ 10 µM reverse primer (FP2): 5′-CTG GCA AGG TAG ACA AGC CG-3′ Toothpicks or bacterial inoculating loops (APPENDIX 4A), sterile 96-well microtiter plate Sealing tape, e.g., wide transparent tape 150- to 212-µm glass beads, acid-washed (UNIT 5.8) Vortexer with flat plate Additional reagents and equipment for performing an interactor hunt (see Basic Protocol 2), PCR amplification of DNA (APPENDIX 4J), agarose gel electrophoresis (APPENDIX 4F), restriction endonuclease digestion (APPENDIX 4I), electroporation (UNIT 5.10), and miniprep isolation of bacterial DNA (APPENDIX 4C) 1. Perform an interactor hunt (see Basic Protocol 2, steps 1 to 19). 2. Use a sterile toothpick or bacterial inoculating loop to transfer yeast from the Glu/CM, −Ura, −His, −Trp master plate into 25 µl lysis solution in a 96-well microtiter plate. Seal the wells of the microtiter plate with sealing tape and incubate 1.5 to 3.5 hr at 37°C with shaking. The volume of yeast transferred should not exceed ∼2 to 3 ìl of packed pellet; larger quantities of yeast will reduce quality of the DNA. DNA can be efficiently recovered from master plates that have been stored up to 1 week at 4°C. If yeast have been previously
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gridded on master plates, transfer to microtiter plates can be facilitated by using a multicolony replicator.
3. Remove tape from the plate, add ∼25 µl acid-washed glass beads to each well, and reseal with the same tape. Firmly attach the microtiter plate to a flat-top vortexer, and vortex 5 min at medium-high power. The microtiter plate can be attached to the vortexer using 0.25-in (0.64-cm) rubber bands.
4. Remove the tape and add ∼100 µl sterile water to each well. Swirl gently to mix, then remove sample for step 5. Press the tape back firmly to seal the microtiter plate and place in the freezer at −20°C for storage. 5. Amplify 0.8 to 2.0 µl of sample by standard PCR (APPENDIX 4J) in a ∼30-µl volume using 3 µl each of the forward primer FP1 and the reverse primer FP2. Perform PCR using the following cycles: Initial step: 31 cycles:
2 min 45 sec 45 sec 45 sec
94°C 94°C 56°C 72°C.
These conditions have been used successfully to amplify fragments up to 1.8 kb in length; some modifications, such as extension of elongation time, are also effective.
6. Load 20 µl of the PCR reaction product on a 0.7% low melting temperature agarose gel (APPENDIX 4F) to resolve PCR products. Based on insert sizes, group the obtained interactors in families, i.e., potential multiple independent isolates of identical cDNAs. Reserve gel until results of step 7 are obtained. No special precautions are needed for storing the gel. Since HaeIII digests typically yield rather small DNA fragments, running the second gel does not take a lot of time. Usually, the delay does not exceed 45 to 60 min, during which time the first gel may be stored in a gel box at room temperature or wrapped in plastic wrap at 4°C.
7. While the gel is running, use the remaining 10 µl of PCR reaction product for a restriction endonuclease digestion with HaeIII in a digestion volume of ∼20 µl (APPENDIX 4I). Based on analysis of the sizes of undigested PCR products in the gel (step 6), rearrange the tubes with HaeIII digest samples so that those thought to represent a family are side by side. Resolve the digests on a 2% to 2.5% agarose gel (APPENDIX 4F). Most restriction fragments will be in the 200-bp to 1.0-kb size range so using a long gel run is advisable. This analysis should produce a distinct fingerprint of insert sizes and allow definition of library cDNAs as unique isolates or related groups. A single positive yeast will sometimes contain multiple library plasmids. An advantage of this protocol is the ready detection of multiple library plasmids in PCR reactions; thus, following subsequent bacterial transformations, only a single TRP1 colony would need to be analyzed unless multiple plasmids were already known to be present.
8. Isolate DNA fragments from the low melting temperature agarose gel (step 6). If inspection of the banding pattern on the two gels suggests that a great many reisolates of a small number of cDNAs are present, it may be worthwhile to immediately sequence PCR products representative of these clusters, but it is generally still advisable to continue through specificity tests before doing so. If the PCR products are sequenced, the FP1 forward primer works well in automated sequencing of PCR fragments, but the FP2 primer is only effective in sequencing from purified plasmid. Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins
In general, priming from the AT-rich ADH terminator downstream of the polylinker/cDNA in library plasmid is less efficient than from upstream of the cDNA, and it is hard to design effective primers in this region.
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9. Remove the microtiter plate of lysates from the freezer, thaw it, and remove 2 to 4 µl of lysed yeast for each desired positive. Electroporate DNA into either DH5α or KC8 E. coli as appropriate, depending on the choice of bait and reporter plasmids (see Table 19.2.1 and see Background Information for further information). Refreeze the plate as a DNA reserve in case bacteria fail to transform on the first pass. KC8 E. coli should be used for electroporation when the original reagents pEG202/pJG45/pJK101 are used for the interaction trap. An additional strength of this protocol is that it identifies redundant clones before transfer of plasmids to bacteria, thus reducing the amount of work required in cases where plasmid identity can be unambiguously assigned. However, although restriction endonuclease digestion and PCR analysis are generally highly predictive, they are not 100% certain methods for estimating cDNA identity. Thus, if there is any doubt about whether two cDNAs are the same, investigators are urged to err on the side of caution.
10. Prepare a miniprep of plasmid DNA from the transformed bacteria (APPENDIX 4C) and perform yeast transformation and specificity assessment (see Basic Protocol 2, steps 24 to 28). PERFORMING A HUNT BY INTERACTION MATING An alternative way of conducting an interactor hunt is to mate a strain that expresses the bait protein with a strain that has been pretransformed with the library DNA, and screen the resulting diploid cells for interactors (Bendixen et al., 1994; Finley and Brent, 1994). This “interaction mating” approach can be used for any interactor hunt, and is particularly useful in three special cases. The first case is when more than one bait will be used to screen a single library. Interaction mating allows several interactor hunts with different baits to be conducted using a single high-efficiency yeast transformation with library DNA. This can be a considerable savings, since the library transformation is one of the most challenging tasks in an interactor hunt. The second case is when a constitutively expressed bait interferes with yeast viability. For such baits, performing a hunt by interaction mating avoids the difficulty associated with achieving a high-efficiency library transformation of a strain expressing a toxic bait. Moreover, the actual selection for interactors will be conducted in diploid yeast, which are more vigorous than haploid yeast and can better tolerate expression of toxic proteins. The third case is when a bait cannot be used in a traditional interactor hunt using haploid yeast strains (see Basic Protocol 2) because it activates transcription of even the least sensitive reporters. In diploids the reporters are less sensitive to transcription activation than they are in haploids. Thus, the interaction mating hunt provides an additional method to reduce background from transactivating baits.
ALTERNATE PROTOCOL 2
In the protocol described below, the library DNA is used to transform a strain with a LEU2 reporter (e.g., EGY48). This pretransformed library strain is then frozen in many aliquots, which can be thawed and used for individual interactor hunts. The bait is expressed in a strain of mating type opposite to that of the pretransformed library strain, and also bearing the lacZ reporter. A hunt is conducted by mixing the pretransformed library strain with the bait strain and allowing diploids to form on YPD medium overnight. The diploids are then induced for expression of the library-encoded proteins and screened for interactors as in Basic Protocol 2. NOTE: Strain combinations other than those described below can also be used in an interaction-mating hunt. The key to choosing the strains is to ensure that the bait and prey strains are of opposite mating types and that both have auxotrophies to allow selection for the appropriate plasmids and reporter genes. Also, once the bait plasmid and lacZ reporter plasmid have been introduced into the bait strain, and the library plasmids have
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been introduced into the library strain, the resulting bait strain and library strain must each have auxotrophies that can be complemented by the other, so that diploids can be selected. Additional Materials (also see Basic Protocols 1 and 2) Yeast strains: either RFY206 (Finley and Brent, 1994), YPH499 (Sikorski and Hieter, 1989; ATCC #6625), or an equivalent MATa strain with auxotrophic markers ura3, trp1, his3, and leu2 YPD liquid medium (APPENDIX 4L) Glu/CM –Trp plates: CM dropout plates −Trp (APPENDIX 4L) supplemented with 2% glucose pJG4-5 library vector (Fig. 19.2.6), empty 100-mm YPD plates (APPENDIX 4L) Additional reagents and equipment for lithium acetate transformation of yeast (APPENDIX 4L) Construct the bait strain The bait strain will be a MATa yeast strain (mating type opposite of EGY48) containing a lacZ reporter plasmid like pSH18-34 and the bait-expressing plasmid, pBait. 1. Perform construction of the bait plasmid (pBait; see Basic Protocol 1, step 1). 2. Cotransform the MATa yeast strain (e.g., either RFY206 or YPH499) with pBait and pSH18-34 using the lithium acetate method (APPENDIX 4L). Select transformants on Glu/CM –Ura,–His plates by incubating plates at 30°C for 3 to 4 days until colonies form. Combine 3 colonies for all future tests and for the mating hunt. The bait strain (RFY206/pSH18-34/pBait or YPH499/pSH18-34/pBait) can be tested by immunoblotting to ensure that the bait protein is expressed (see Support Protocol 1). Synthesis and nuclear localization of the bait protein can also be tested by the repression assay (see Basic Protocol 1, steps 8 to 12).
3. Optional: Assay lacZ gene activation in the bait strain (see Basic Protocol 1, steps 4 to 7). If the bait activates the lacZ reporter, a less sensitive lacZ reporter plasmid (Table 19.2.1), or an integrated version of the lacZ reporter should be tried. A bait that strongly activates the lacZ reporters usually cannot be used in a hunt based on selection of interactors with the LEU2 reporter, because the LEU2 reporters are more sensitive than the lacZ reporters. However, both reporters are less sensitive to activation by a bait in diploid cells, as compared to haploid cells. Thus, a more important test of the transactivation potential of a bait is to test the leucine requirement of diploid cells expressing it, as described in steps 6 to 20, below.
Prepare the pretransformed library strain (EGY48 + library plasmids) 4. Perform a large-scale transformation of EGY48 with library DNA using the lithium acetate method (see Basic Protocol 2, steps 1 to 8, except start with EGY48 bearing no other plasmids). To prepare for transformation, grow EGY48 in YPD liquid medium. Select library transformants on Glu/CM –Trp plates by incubating 3 days at 30°C. 5. Collect primary transformants by scraping plates, washing yeast, and resuspending in 1 pellet vol glycerol solution (see Basic Protocol 2, steps 9 to 12). Freeze 0.2 to 1.0 ml aliquots at −70° to −80°C. Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins
The cells will be stable for at least 1 year. Refreezing a thawed aliquot will result in loss of viability. Thus, many frozen aliquots should be made, so that each thawed aliquot can be discarded after use.
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Prepare the pretransformed control strain (EGY48 + pJG4-5) 6. Transform EGY48 grown in YPD liquid medium with the empty library vector, pJG4-5, using the lithium acetate method (APPENDIX 4L). Select transformants on Glu/CM –Trp plates by incubating 3 days at 30°C. 7. Pick and combine three transformant colonies and use them to inoculate 30 ml of Glu/CM –Trp medium. Incubate 15 to 24 hr at 30°C (to OD600 >3). 8. Centrifuge 5 min at 1000 to 1500 × g, room temperature, and remove supernatant. Resuspend in 10 ml sterile water to wash cells. 9. Centrifuge 5 min at 1000 to 1500 × g, room temperature, and remove supernatant. Resuspend in 1 pellet vol glycerol solution and freeze 100-µl aliquots at −70° to −80°C. Determine plating efficiency of pretransformed library and pretransformed control strains 10. After freezing (at least 1 hr) thaw an aliquot of each pretransformed strain (from step 5 and step 9) at room temperature. Make several serial dilutions in sterile water, including aliquots diluted 105-fold, 106-fold, and 107-fold. Plate 100 µl of each dilution on 100-mm Glu/CM –Trp plates and incubate 2 to 3 days at 30°C. 11. Count the colonies and determine the number of colony-forming units (cfu) per aliquot of transformed yeast. The plating efficiency for a typical library transformation and for the control strain will be ∼1 × 108 cfu per 100 ìl.
Mate the bait strain with the pretransformed library strain and the pretransformed control strain In steps 12 through 20, an interactor hunt is conducted concurrently with testing LEU2 reporter activation by the bait itself. For most baits, this approach will be the quickest way to isolate interactors. However, for some baits, such as those that have a high transactivation potential, or those that affect yeast mating or growth, steps 12 through 20 will serve as a pilot experiment to determine the optimal parameters for a subsequent hunt. 12. Grow a 30-ml culture of the bait strain in Glu/CM –Ura,–His liquid dropout medium to mid to late log phase (OD600 = 1.0 to 2.0, or 2 to 4 × 107cells/ml). A convenient way to grow the bait strain is to inoculate a 5-ml culture with approximately three colonies from a plate and grow it overnight at 30°C with shaking. In the morning, measure the OD600, dilute into a 30-ml culture to a final OD600 = 0.2, and grow at 30°C with shaking. The culture should reach mid to late log phase before the end of the day.
13. Centrifuge the culture 5 min at 1000 to 1500 × g, room temperature, to harvest cells. Resuspend the cell pellet in sterile water to make a final volume of 1 ml. This should correspond to ∼1 × 109 cells/ml.
14. Set up two matings. In one sterile microcentrifuge tube mix 200 µl of the bait strain with 200 µl of a thawed aliquot of the pretransformed control strain from step 9. In a second microcentrifuge tube mix 200 µl of the bait strain with ∼1 × 108 cfu (∼0.1 to 1 ml) of the pretransformed library strain from step 5. The library mating should be set up so that it contains a ∼2-fold excess of bait strain cfu over pretransformed library strain cfu. Because the bait strain was harvested in log phase, most of the cells will be viable (i.e., cells/ml = ∼cfu/ml), and the number of cfu can be sufficiently estimated from optical density (1 OD600 = ∼2 × 107 cells/ml). Under these conditions, ∼10% of the cfu in the pretransformed library strain will mate with the bait
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strain. Thus, a complete screen of 107 library transformants will require a single mating with at least 108 cfu of the pretransformed library strain and at least 2 × 108 cfu of the bait strain. To screen more library transformants, set up additional matings. The number of pretransformed library transformants to screen depends on the size of the library and the number of primary transformants obtained in step 5. If the size of the library is larger than the number of transformants obtained in step 5, the goal will be to screen all of the yeast transformants. In this case, complete screening of the library will require additional transformations of EGY48 and additional interactor hunts. If the size of the library is smaller than the number of transformants obtained in step 5, the goal will be to screen at least a number of transformants equivalent to the size of the library.
15. Centrifuge each cell mixture for 5 min at 1000 to 1500 × g, pour off medium, and resuspend cells in 200 µl YPD medium. Plate each suspension on a 100-mm YPD plate. Incubate 12 to 15 hr at 30°C. 16. Add ∼1 ml of Gal/Raff/CM –Ura, –His, –Trp to the lawns of mated yeast on each plate. Mix the cells into the medium using a sterile applicator stick. 17. Transfer each slurry of mated cells to a 500-ml flask containing 100 ml of Gal/Raff/CM –Ura, –His, –Trp dropout medium. Incubate with shaking 6 hr at room temperature to induce the GAL1 promoter, which drives expression of the cDNA library. 18. Centrifuge the cell suspensions 5 min at 1000 to 1500 × g, room temperature, to harvest the cells. Wash by resuspending in 30 ml of sterile water and centrifuging again. Resuspend each pellet in 5 ml sterile water. Measure OD600 and, if necessary, dilute to a final concentration of ∼1 × 108 cells/ml. This is a mixture consisting of haploid cells that have not mated and diploid cells. Under a microscope, the two cell types can be distinguished by size (diploids are ∼1.7× bigger than haploids) and shape (diploids are slightly oblong and haploids are spherical). Because diploids grow faster than haploids, this mixture will contain ∼10% to 50% diploid cells. The actual number of diploids will be determined by plating dilutions on –Ura, –His, –Trp medium, which will not support the growth of the parental haploids.
19. For each mating make a series of 1⁄10 dilutions in sterile water, at least 200 µl each, to cover a 106-fold concentration range. Plate 100 µl from each tube (undiluted, 10−1, 10−2, 10−3, 10−4, 10−5, and 10−6 dilution) on 100-mm Gal/Raff/CM –Ura, –His, –Trp, –Leu plates. Plate 100 µl from the 10−4, 10−5, and 10−6 tubes on 100-mm Gal/Raff/CM –Ura, –His, –Trp plates. Incubate plates at 30°C. Count the colonies on each plate after 2 to 5 days. 20. For the mating with the pretransformed library, prepare an additional 3 ml of a 10−1 dilution. Plate 100 µl of the 10−1 dilution on each of 20 100-mm Gal/Raff/CM –Ura, –His, –Trp, –Leu plates. Also plate 100 µl of the undiluted cells on each of 20 100-mm Gal/Raff/CM –Ura, –His, –Trp, –Leu plates. Incubate at 30°C. Pick Leu+ colonies after 2 to 5 days and characterize them beginning with step 17 of Basic Protocol 2.
Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins
The number of Leu+ colonies to pick to ensure that all of the pretransformed library has been screened depends on the transactivation potential of the bait protein itself. The transactivation potential is expressed as the number of Leu+ colonies that grow per cfu (Leu+/cfu) of the bait strain mated with the control strain, as determined in step 19 of this protocol. It can be calculated as the ratio of the number of colonies that grow on Gal/Raff/CM –Ura,–His, –Trp, –Leu to the number of colonies that grow on Gal/Raff/CM –Ura, –His, –Trp for a given dilution of the mating between the bait strain and the control strain. A bait with essentially no transactivation potential will produce less than 10−6 Leu+/cfu. For a bait to be useful in an interactor hunt it should not transactivate more than 10−4 Leu+/cfu.
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To screen all of the pretransformed library, it will be necessary to pick a sufficient number of Leu+ colonies in addition to background colonies produced by the transactivation potential of the bait itself. Thus, the minimum number of Leu+ colonies that should be picked in step 20 of this protocol is given by: (transactivation potential, Leu+/cfu) × (# library transformants screened). For example, if 107 library transformants were obtained in step 2 (and at least 108 cfu of these transformants were mated with the bait strain in step 14, since only ∼10% will form diploids), and the transactivation potential of the bait is 10−4 Leu+/cfu, then at least 1000 Leu+ colonies must be picked and characterized. In other words, if the rarest interactor is present in the pretransformed library at a frequency of 10−7, to find it one needs to screen through at least 107 diploids from a mating of the library strain. However, at least 1000 of these 107 diploids would be expected to be Leu+ due to the bait background if the transactivation potential of the bait is 10−4. The true positives will be distinguished from the bait background in the next step by the galactose dependence of their Leu+ and lacZ+ phenotypes.
PREPARATION OF PROTEIN EXTRACTS FOR IMMUNOBLOT ANALYSIS To confirm that the bait fusion protein constructed in Basic Protocol 1 is synthesized properly, a crude lysate is prepared for SDS-PAGE and immunoblot analysis (UNITS 10.1 & 10.10). The presence of the target protein is detected by antibody to LexA or the fusion domain.
SUPPORT PROTOCOL 1
Materials Master plates with pBait-containing positive and control yeast on Glu/CM −Ura, −His dropout medium (see Basic Protocol 1, step 4) Glu/CM −Ura, −His dropout liquid medium: CM dropout plates −Ura, −His (APPENDIX 4L) supplemented with 2% glucose 2× Laemmli sample buffer (see recipe) Antibody to fusion domain or LexA: monoclonal antibody to LexA (Clontech, Invitrogen) or polyclonal antibody to LexA (available by request from R. Brent or E. Golemis) 30°C incubator 100°C water bath Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and immunoblotting and immunodetection (UNIT 10.10) 1. From the master plates, start a 5-ml culture in Glu/CM −Ura, −His liquid medium for each bait being tested and for a positive control for protein expression (i.e., RFHMI or SH17-4). Incubate overnight at 30°C. For each construct assayed, it is a good idea to grow colonies from at least two primary transformants, as levels of bait expression are sometimes heterogenous.
2. From each overnight culture, start a fresh 5-ml culture in Glu/CM −Ura, −His at OD600 = ∼0.15. Incubate again at 30°C. 3. When the culture has reached OD600 = 0.45 to 0.7 (∼4 to 6 hr), remove 1.5 ml to a microcentrifuge tube. For some LexA fusion proteins, levels of the protein drop off rapidly in cultures approaching stationary phase. This is due to a combination of the diminishing activity of the ADH1 promoter in late growth phases and the relative instability of particular fusion domains. Thus, it is not a good idea to let cultures become saturated in the hopes of obtaining a higher yield of protein.
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4. Microcentrifuge cells 3 min at 13,000 × g, room temperature. When the pellet is visible, remove the supernatant. Inspection of the tube should reveal a pellet ∼1 to 3 ìl in volume. If the pellet is not visible, microcentrifuge another 3 min.
5. Working rapidly, add 50 µl of 2× Laemmli sample buffer to the visible pellet in the tube, vortex, and place the tube on dry ice. Samples may be frozen at −70°C.
6. Transfer frozen sample directly to a boiling water bath or a PCR machine set to cycle at 100°C. Boil 5 min. 7. Microcentrifuge 5 sec at maximum speed to pellet large cellular debris. 8. Perform SDS-PAGE (UNIT 10.1) using 20 to 50 µl sample per lane. 9. To detect the protein, immunoblot and analyze (UNIT 10.10) using antibody to the fusion domain or LexA. SUPPORT PROTOCOL 2
PREPARATION OF SHEARED SALMON SPERM CARRIER DNA This protocol generates high-quality sheared salmon sperm DNA (sssDNA) for use as carrier in transformation (Basic Protocol 2). This DNA is also suitable for other applications where high-quality carrier DNA is needed (e.g., hybridization). This protocol is based on Schiestl and Gietz (1989). For more details of phenol extraction or other DNA purification methods, consult APPENDIX 4E. Materials High-quality salmon sperm DNA (e.g., sodium salt from salmon testes, Sigma or Boehringer Mannheim), desiccated TE buffer, pH 7.5 (APPENDIX 2E), sterile TE-saturated buffered phenol (APPENDIX 2E) 1:1 (v/v) buffered phenol/chloroform Chloroform 3 M sodium acetate, pH 5.2 (APPENDIX 2E) 100% and 70% ethanol, ice cold Magnetic stirring apparatus and stir-bar, 4°C Sonicator with probe 50-ml conical centrifuge tube High-speed centrifuge and appropriate tube 100°C and ice-water baths 1. Dissolve desiccated high-quality salmon sperm DNA in TE buffer, pH 7.5, at a concentration of 5 to 10 mg/ml by pipetting up and down in a 10-ml glass pipet. Place in a beaker with a stir-bar and stir overnight at 4°C to obtain a homogenous viscous solution. It is important to use high-quality salmon sperm DNA. Sigma Type III sodium salt from salmon testes has worked well, as has a comparable grade from Boehringer Mannheim. Generally it is convenient to prepare 20- to 40-ml batches at a time.
2. Shear the DNA by sonicating briefly using a large probe inserted into the beaker. Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins
The goal of this step is to generate sheared salmon sperm DNA (sssDNA) with an average size of 7 kb, but ranging from 2 to 15 kb. Oversonication (such that the average size is closer to 2 kb) drastically decreases the efficacy of carrier in enhancing transformation. The original version of this protocol (Schiestl and Gietz, 1989) called for two 30-sec pulses at
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Table 19.2.4
System
Two-hybrid System Variantsa
DNA-binding Activation Selection domain domain
Two-hybrid Interaction trap “Improved two-hybrid” Modified two-hybrid KISS Contingent replication
GAL4 LexA GAL4 LexA GAL4 GAL4
GAL4 B42 GAL4 VP16 VP16 VP16
Activation of lacZ, HIS3 Activation of LEU2, lacZ Activation of HIS3, lacZ Activation of HIS3, lacZ Activation of CAT, hygr Activation of T-Ag, replication of plasmids
Reference Chien et al., 1991 Gyuris et al., 1993 Durfee at al., 1993 Vojtek at al., 1993 Fearon et al., 1992 Vasavada et al., 1991
aAbbreviations: CAT, chloramphenicol transferase gene; hygr, hygromycin resistance gene; T-Ag, viral large T antigen.
three-quarter power, but optimal conditions vary between sonicators. The first time this protocol is performed, it is worthwhile to sonicate briefly, then test the size of the DNA by running out a small aliquot alongside molecular weight markers on an agarose gel containing ethidium bromide. The DNA can be sonicated further if needed.
3. Once DNA of the appropriate size range has been obtained, extract the sssDNA solution with an equal volume of TE-saturated buffered phenol in a 50-ml conical tube, shaking vigorously to mix. 4. Centrifuge 5 to 10 min at 3000 × g, room temperature, or until clear separation of phases is obtained. Transfer the upper phase containing the DNA to a clean tube. 5. Repeat extraction using 1:1 (v/v) buffered phenol/chloroform, then chloroform alone. Transfer the DNA into a tube suitable for high-speed centrifugation. 6. Precipitate the DNA by adding 1⁄10 vol of 3 M sodium acetate and 2.5 vol of ice-cold 100% ethanol. Mix by inversion. Centrifuge 15 min at ∼12,000 × g, room temperature. 7. Wash the pellet with 70% ethanol. Briefly dry either by air drying, or by covering one end of the tube with Parafilm with a few holes poked in and placing the tube under vacuum. Resuspend the DNA in sterile TE buffer at 5 to 10 mg/ml. Do not overdry the pellet or it will be very difficult to resuspend.
8. Denature the DNA by boiling 20 min in a 100°C water bath. Then immediately transfer the tube to an ice-water bath. 9. Place aliquots of the DNA in microcentrifuge tubes and store frozen at −20°C. Thaw as needed. DNA should be boiled again briefly (5 min) immediately before addition to transformations. Before using a new batch of sssDNA in a large-scale library transformation, it is a good idea to perform a small-scale transformation using suitable plasmids to determine the transformation efficiency. Optimally, use of sssDNA prepared in the manner described will yield transformation frequencies of >105 colonies/ìg input plasmid DNA.
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SUPPORT PROTOCOL 3
YEAST COLONY HYBRIDIZATION This protocol is adapted from a modification of the classic protocol of Grunstein and Hogness (1975; Kaiser et al., 1994). It is primarily useful when a large number of putative interactors has been obtained, and initial minipreps and restriction digests have indicated that many of them derive from a small number of cDNAs; these cDNAs can then be used as probes to screen and eliminate identical cDNAs from the pool. Materials Glu/CM −Trp plates: CM dropout plates −Trp (APPENDIX 4L) supplemental with 2% glucose Master dropout plate of yeast positive for Gal dependence (see Basic Protocol 2, step 18) 1 M sorbitol/20 mM EDTA/50 mM DTT (prepare fresh) 1 M sorbitol/20 mM EDTA 0.5 M NaOH 0.5 M Tris⋅Cl (pH 7.5)/6× SSC (APPENDIX 2E) 2× SSC (APPENDIX 2E) 100,000 U/ml β-glucuronidase (type HP-2 crude solution from Helix pomatia; Sigma) 82-mm circular nylon membrane, sterile Whatman 3MM paper 80°C vacuum oven or UV cross-linker Additional reagents and equipment for bacterial filter hybridization (Strauss, 1993; Duby et al., 1988) 1. Place a sterile nylon membrane onto a Glu/CM −Trp dropout plate. From the master dropout plate of Gal-dependent positives, gently restreak positives to be screened onto the membrane and mark the membrane to facilitate future identification of hybridizing colonies. Grow overnight (∼12 hr) at 30°C. Growth for extended periods of time (i.e., 24 hr) may result in difficulty in obtaining good lysis. It is a good idea to streak positive and negative controls for the cDNAs to be hybridized on the membrane.
2. Remove membrane from plate. Air dry briefly. Incubate ∼30 min on a sheet of Whatman 3MM paper saturated with 1 M sorbitol/20 mM EDTA/50 mM DTT. Optionally, before commencing chemical lysis, membranes can be placed at −70°C for 5 min, then thawed at room temperature for one or more cycles to enhance cell wall breakage.
3. Cut a piece of Whatman 3MM paper to fit inside a 100-mm petri dish. Place the paper disc in the dish and saturate with 100,000 U/ml β-glucuronidase diluted 1:500 in 1 M sorbitol/20 mM EDTA (2 µl glucuronidase per ml of sorbitol/EDTA to give 200 U/ml final). Layer nylon membrane on dish, cover dish, and incubate up to 6 hr at 37°C until >80% of the cells lack a cell wall. The extent of cell wall removal can be determined by removing a small quantity of cells from the filter to a drop of 1 M sorbitol/20 mM EDTA on a microscope slide and observing directly with a phase-contrast microscope at ≥60× magnification. Cells lacking cell wall are nonrefractile.
Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins
4. Place membrane on Whatman 3MM paper saturated with 0.5 M NaOH for ∼8 to 10 min. 5. Place membrane on Whatman 3MM paper saturated with 0.5 M Tris⋅Cl (pH 7.5)/6× SSC for 5 min. Repeat with a second sheet of Whatman 3MM paper.
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6. Place membrane on Whatman 3MM paper saturated with 2× SSC for 5 min. Then place membrane on dry Whatman paper to air dry for 10 min. 7. Bake membrane 90 min at 80°C in vacuum oven or UV cross-link. 8. Process as for bacterial filter hybridization (Strauss, 1993; Duby et al., 1988), hybridizing the membrane with probes complementary to previously isolated cDNAs. When selecting probes, either random-primed cDNAs or oligonucleotides complementary to the cDNA sequence may be used. If the cDNA is a member of a protein family, it may be advantageous to use oligonucleotides to avoid inadvertently excluding genes related but not identical to those initially obtained.
MICROPLATE PLASMID RESCUE In some cases, it is desirable to isolate plasmids from a large number of positive colonies (Basic Protocol 2, steps 18 and 19). The protocol described below is a batch DNA preparation protocol developed by Steve Kron (University of Chicago, Chicago, Ill.) as a scale-up of a basic method developed by Manuel Claros (Laboratoire de Génétique Moleculaire, Paris, France).
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Materials 2× Glu/CM −Trp liquid medium: 2× CM −Trp liquid medium (APPENDIX 4L) supplemented with 4% glucose Master plate of Gal-dependent yeast colonies (see Basic Protocol 2, step 18) Rescue buffer: 50 mM Tris⋅Cl (pH 7.5)/10 mM EDTA/0.3% (v/v) 2-mercaptoethanol (prepare fresh) Lysis solution: 2 to 5 mg/ml Zymolyase 100T/rescue buffer or 100,000 U/ml β-glucuronidase (type HP-2 crude solution from Helix pomatia; Sigma) diluted 1:50 in rescue buffer 10% (w/v) SDS 7.5 M ammonium acetate Isopropanol 70% ethanol TE buffer, pH 8.0 (APPENDIX 2E) 24-well microtiter plates Centrifuge with microplate holders, refrigerated Repeating micropipettor 37°C rotary shaker Grow yeast cultures 1. Aliquot 2 ml of 2× Glu/CM −Trp medium into each well of a 24-well microtiter plate. Into each well, pick a putative positive colony. Grow overnight with shaking at 30°C. The 2× minimal medium is used to maximize the yield of yeast. Four plates can generally be handled conveniently at once, based on the number that can be centrifuged simultaneously.
2. Centrifuge 5 min at 1500 × g, 4°C. Shake off supernatant with a snap and return the plate to upright. 3. Swirl or lightly vortex the plate to resuspend cell pellets in remaining liquid. Add 1 ml water to each well and swirl lightly. Cell pellets can most easily be resuspended in residual liquid before adding new solutions. Addition of liquid can be accomplished using a repeating pipettor.
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4. Centrifuge 5 min at 1500 × g, 4°C. Shake off supernatant and resuspend pellet. Add 1 ml rescue buffer. 5. Centrifuge 5 min at 1500 × g, 4°C. Shake off supernatant and resuspend pellet in the small volume of liquid remaining in the plate. Lyse cells 6. To each well, add 25 µl lysis solution. Swirl or vortex to mix. Incubate (with cover on) on a rotary shaker ∼1 hr at 37°C. Lysis solution need not be completely dissolved before use. By 1 hr, lysis should be obvious as coagulation of yeast into a white precipitate. Susceptibility of yeast strains to lytic enzymes varies. If lysis occurs rapidly, then less lytic enzyme should be used. If the lysis step is allowed to go too far, too much of the partially dissolved cell wall may contaminate the final material. Lysis can be judged by examining cells with a phase-contrast microscope. Living cells are white with a dark halo and dead cells are uniformly gray. Lysis leads to release of granular cell contents into the medium. Once cells are mostly gray and many are disrupted, much of the plasmid should have been released.
7. To each well, add 25 µl of 10% SDS. Mix gently by swirling to completely disperse the precipitates. Allow plates to sit 1 min at room temperature. At this point, the wells should contain a clear, somewhat viscous solution.
Purify plasmid 8. To each well, add 100 µl of 7.5 M ammonium acetate. Swirl gently, then incubate 15 min at −70°C or −20°C until frozen. Addition of acetate should result in the formation of a massive white precipitate of cell debris and SDS. The freezing step appears to improve removal of inhibitors of E. coli transformation.
9. Remove plate from freezer. Once it begins to thaw, centrifuge 15 min at 3000 × g, 4°C. Transfer 100 to 150 µl of the resulting clear supernatants to clean 24-well plates. In general, some contamination of the supernatant with pelleted material cannot be avoided. However, it is better to sacrifice yield in order to maintain purity.
10. To each well, add ∼0.7 vol isopropanol. Mix by swirling and allow to precipitate 2 min at room temperature. A cloudy fine precipitate should form immediately after isopropanol is added.
11. Centrifuge 15 min at 3000 × g, 4°C. Shake off supernatant with a snap. 12. To each well, add ∼1 ml cold 70% ethanol. Mix by swirling, centrifuge 5 min at 3000 × g, 4°C. Shake off supernatant with a snap, invert plates and blot well onto paper towel. Allow plates to air dry. 13. To each well, add 100 µl TE buffer. Swirl well and allow to rest on bench several minutes, until the pellets appear fully dissolved. Transfer preps to microcentrifuge tubes or 96-well plates for storage at −20°C.
Interaction Trap/ Two-Hybrid System to Identify Interacting Proteins
One to five microliters of each of the resulting preparations can be used to transform competent E. coli: for KC8, electroporation should be used (see Basic Protocol 2, step 21). Sometimes, the yield of transformants is low if E. coli carrying plasmids are not permitted time to increase the plasmid copy number above a critical threshold before the cells are placed on selective medium. Allow plenty of time for cells to express antibiotic resistance or the TRP1 gene before plating. If insufficient numbers of colonies are obtained by this approach, the final plasmid preparation can be resuspended in 20 ìl instead of 100 ìl TE buffer to concentrate the DNA stock.
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ADDITIONAL SPECIFICITY SCREENING The three test plasmids outlined (pSH18-34, pRFHM1, and pEG202; see Basic Protocol 2, step 24) represent a minimal test series. If other LexA-bait proteins that are related to the bait protein used in the initial library screen are available, substantial amounts of information can be gathered by additional specificity tests. For example, if the initial bait protein was LexA fused to the leucine zipper of c-Fos, specificity screening of interactor-hunt positives against the leucine zippers of c-Jun or GCN4 in addition to that of c-Fos might allow discrimination between proteins that are specific for fos versus those that generically associate with leucine zippers.
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REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Glycerol solution 65% (v/v) glycerol, sterile 0.1 M MgSO4 25 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) Store up to 1 year at room temperature Laemmli sample buffer, 2× 10% (v/v) 2-mercaptoethanol (2-ME) 6% (w/v) SDS 20% (v/v) glycerol 0.2 mg/ml bromphenol blue 0.025× Laemmli stacking buffer (see recipe; optional) Store up to 2 months at room temperature This reagent can conveniently be prepared 10 ml at a time.
Laemmli stacking buffer, 2.5× 0.3 M Tris⋅Cl, pH 6.8 0.25% (w/v) SDS Store up to 1 month at 4°C Lysis solution 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 10 mM EDTA 0.3% (v/v) 2-mercaptoethanol (2-ME), added just before use 2% (v/v) β-glucuronidase from Helix pomatia (Type HP-2; Sigma), added just before use COMMENTARY Background Information Interaction-based cloning is derived from three experimental observations. In the first, Brent and Ptashne (1985) demonstrated that it was possible to assemble a novel, functional transcriptional activator by fusing the DNAbinding domain from one protein, LexA, to the activation domain from a second protein, GAL4. This allowed the use of a single reporter system containing a single DNA-binding motif, the LexA operator, to study transcriptional ac-
tivation by any protein of interest. In the second, Ma and Ptashne (1988) built on this work to demonstrate that the activation domain could be brought to DNA by interaction with a DNAbinding domain. In the third, Fields and Song (1989), working independently of Ma and Ptashne, used two yeast proteins, SNF1 and SNF4, to make an SNF1 fusion to the DNAbinding domain of GAL4 and an SNF4 fusion to the GAL4 activation domain. They demonstrated that the strength of the SNF1-SNF4
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interaction was sufficient to allow activation through a GAL4 DNA-binding site. From this, they suggested the feasibility of selecting interacting proteins by performing screens of cDNA libraries made so that library-encoded proteins carried activating domains. Several groups have developed cDNA library strategies along these lines, with some systems using LexA and others using GAL4 as the DNA-binding domain (Table 19.2.4). LexA and GAL4 each have different properties that should be considered when selecting a system. LexA is derived from a heterologous organism, has no known effect on the growth of yeast, possesses no residual transcriptional activity, can be used in GAL4+ yeast, and can be used with a Gal-inducible promoter. Because GAL4 is an important yeast transcriptional activator, it has the disadvantage that experiments must be performed in gal4− yeast strains to avoid background due to activation of the reporter system by endogenous GAL4. Such gal4− strains are frequently less healthy and more difficult to transform than wild-type strains, and either libraries must be constitutively expressed or alternate inducible systems must be used. By contrast, the GAL4 DNA-binding domain may be more efficiently localized to the nucleus and may be preferred for some proteins (for a review of GAL4-based systems, see Bartel et al., 1993). Whichever system is used, it is important to remember that the bait protein constitutes a novel fusion protein whose properties may not exactly parallel those of the original unfused protein of interest. Although systems using the two-hybrid paradigm have been developed in mammalian cells (see Table 19.2.4), these have not been used effectively in library screens. It seems likely that the organism of choice for two-hybrid identification of novel partner proteins will remain yeast. cDNAs that pass specificity tests are referred to as positives, or “true positives.” In interactor hunts conducted to date, anywhere from zero to practically all isolated plasmids passed the final specificity test. If no positives are obtained, the tissue source for the library originally used may not be appropriate, and a different library may produce better results. However, there are some proteins for which no positives are found. Various explanations for this are provided below. Conversely, some library-encoded proteins are known to be isolated repeatedly using a series of unrelated baits, and these proteins demonstrate at least some specificity. One of these, heat shock protein 70, might be explained by positing that it
assists the folding of some LexA-fused bait proteins, or alternatively, that these bait proteins are not normally folded. This example illustrates the point that the physiological relevance of even quite specific interactions may sometimes be obscure. Because the screen involves plating multiple cells to Gal/CM −Ura, −His, −Trp, −Leu dropout medium for each primary transformant obtained, multiple reisolates of true positive cDNAs are frequently obtained. If a large number of specific positives are obtained, it is generally a good idea to attempt to sort them into classes—for example, digesting minipreps of positives with EcoRI, XhoI, and HaeIII will generate a fingerprint of sufficient resolution to determine whether multiple reisolates of a small number of clones or single isolates of many different clones have been obtained. The former situation is a good indication that the system is working well. An important issue that arises in an interactor hunt is the question of how biologically relevant interacting proteins that are isolated are likely to be. This leads directly to the question of what Kd of association two molecules must have to be detected by an interactor hunt. In fact, this is not at all a simple issue. For the system described here, most fusion proteins appear to be expressed at levels ranging from 50 nM to 1 µM (Golemis and Brent, 1992). Given the strength of the GAL promoter, it is likely that many library-encoded proteins are expressed at similarly high levels, ≥1 µM in the nucleus (Golemis and Brent, 1992). At this concentration, which is in considerable excess over the nuclear concentration of operatorbound bait protein, a cDNA library–encoded protein should half-maximally occupy the DNA-bound bait protein if it possesses a Kd of 10−6 M, making it theoretically possible that very-low-affinity interactions could be detected. Such interactions have been observed in some cases. In contrast, some interactions that have been previously established using other methods and are predicted by known Kd to be easily detected by these means, either are not detected or are detected only weakly (Finley and Brent, 1994; Estojak et al., 1995). Because of the conservation of many proteins between lower and higher eukaryotes, one explanation for this observation is that either one or both of the partners being tested is being sequestered from the desired interaction by fortuitous association with an endogenous yeast protein. A reasonably complete investigation of the degree of correlation between in vitro determina-
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tions of interaction affinity and apparent strength of interaction in the interaction trap is included in Estojak et al. (1995). The result of this investigation suggests it is important to measure the affinity of detected interactions under different conditions, using a second assay system, rather than to draw conclusions about affinity based on detection in the interaction trap. A number of different plasmids can be used for conducting an interactor hunt. Their properties are summarized in Tables 19.2.1 and 19.2.2. Because of the generous and open scientific exchange between investigators using the system, the number of available plasmids and other components has greatly expanded since the appearance of the initial two-hybrid reagents, facilitating the study of proteins inaccessible by the original system. The original parent plasmid for generating LexA fusions, pEG202 is a derivative of 202 + PL (Ruden et al., 1991; see Fig. 19.2.3) that contains an expanded polylinker region. The available cloning sites in pEG202 include EcoRI, BamHI, SalI, NcoI, NotI, and XhoI, with the reading frame as described in the legend to Figure 19.2.3. Since the original presentation of this system, a number of groups have developed variants of this plasmid that address specialized research needs. Those currently available, as well as purposes for which they are suited, are listed in Table 19.2.1. pGilda, created by David A. Shaywitz, places the LexAfusion cassette under the control of the inducible GAL1 promoter, allowing expression of the bait protein for limited times during library screening, reducing the exposure of yeast to toxic baits. pJK202, created by Joanne Kamens, adds nuclear localization sequences to pEG202, facilitating assay of the function of proteins lacking internal nuclear localization sequences. pNLexA, created by Ian York, places LexA carboxy-terminal in the fusion domain, allowing assay of interactions that require an unblocked amino-terminus on the bait protein. pEE202I, created by Mike Watson and Rich Buckholz, allows chromosomal integration of a pEG202-like bait, thus reducing expression levels so they are more physiological for bait proteins normally present at low levels intracellularly. All of these have been extensively tested by numerous researchers. pGilda, pJK202, and pEE202I work with complete reliability. pNLexA works effectively with ∼50% of the fusion domains tried, but synthesizes only very low levels of protein (relative to expression of the same fusion domain as a
pEG202 fusion) with the remaining 50%. Attachment of fusion domains amino-terminal either to LexA or GAL4 has been generally problematic in the hands of many investigators; it may be that appending additional protein sequences to the amino termini of these proteins is destabilizing, although the problem has not been rigorously investigated. A series of lacZ reporters of differing sensitivity to transcriptional activation can be used to detect interactions of varying affinity (see Table 19.2.2). These plasmids are LexA operator–containing derivatives of the plasmid LR1∆1 (West et al., 1984). In LR1∆1, a minimal GAL1 promoter lacking the GAL1 upstream activating sequences (GALUAS) is located upstream of the bacterial lacZ gene. In pSH18-34, eight LexA operators have been cloned into an XhoI site located 167 bp upstream of the lacZ gene (S. Hanes, unpub. observ.). pJK103 and pRB1840 contain two and one operator, respectively. pJK101 is similar to pSH18-34, except that it contains the GAL1 upstream activating sequences (GALUAS) upstream of two LexA operator sites. A derivative of del20B (West et al., 1984), it is used in the repression assay (Brent and Ptashne, 1984; see Fig. 19.2.5) to assess LexA fusion binding to operator. pSH17-4 is a HIS3 2µm plasmid that encodes LexA fused to the activation domain of the yeast activator GAL4. EGY48 cells bearing this plasmid will produce colonies in overnight growth on medium lacking Leu, and yeast that additionally contain pSH18-34 will turn deep blue on plates containing Xgal. This plasmid serves as a positive control for the activation of transcription. pRFHM1 is a HIS3 2µm plasmid that encodes LexA fused to the N-terminus of the Drosophila protein bicoid. The plasmid has no ability to activate transcription, so EGY48 cells that contain pRFHM1 and pSH18-34 do not grow on −Leu medium and remain white on plates containing Xgal. pRFHM1 is a good control for specificity testing, because it has been demonstrated to be sticky—that is, to associate with a number of library-encoded proteins that are clearly nonphysiological interactors (R. Finley, Wayne State University, Detroit, Mich., unpub. observ.). This protocol uses interaction libraries (Table 19.2.3) made in pJG4-5 or its derivatives (see Fig. 19.2.6). pJG4-5 was developed to facilitate isolation and characterization of novel proteins in interactor hunts (Gyuris et al., 1993). The pJG4-5 cDNA library expression
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cassette is under control of the GAL1 promoter, so library proteins are expressed in the presence of galactose (Gal) but not glucose (Glu). This conditional expression has a number of advantages, the most important of which is that many false-positives obtained in screens can be easily eliminated because they do not demonstrate a Gal-dependent phenotype. The expression cassette consists of an ATG to start translation, a nuclear localization signal to extend the interaction trap’s range to include proteins that are normally predominantly localized in the cytoplasm, an activation domain (acid blob; Ma and Ptashne, 1987), the hemagglutinin epitope tag to permit rapid assessment of the size of encoded proteins, EcoRI-XhoI sites designed to receive directionally synthesized cDNAs, and the alcohol dehydrogenase (ADH) termination sequences to enhance the production of high levels of library protein. The plasmid also contains the TRP1 auxotrophy marker and 2µm origin for propagation in yeast. A derivative plasmid, pJG4-5I, was created by Mike Watson and Richard Buckholz to facilitate chromosomal integration of the activation domain fusion expression plasmid. A series of recently developed derivatives of pEG202, pJG4-5, and lacZ reporter plasmids (MW101 to MW112) alter the antibiotic resistance markers on these plasmids from ampicillin (Apr) to either kanamycin (Kmr) or chloramphenicol (Cmr; Watson et al., 1996). Judiciously mixing and matching these plasmids in conjunction with Apr libraries would considerably reduce work subsequent to library screening, because the KC8 transformation, which involves trpC complementation in bacteria, could be omitted. EGY48 and EGY191 (see Table 19.2.2) are both derivatives of the strain U457 (a gift of Rodney Rothstein, Columbia University, New York, N.Y.) in which the endogenous LEU2 gene has been replaced by homologous recombination with LEU2 reporters carrying varying numbers of LexA operators, using a procedure detailed in Estojak et al. (1995). Interaction Trap–compatible reagents have recently become commercially available; Clontech and Invitrogen were the first to market such reagents and have recently been joined by OriGene. All suppliers use systems with the most sensitive reporters (EGY48 and pSH1834), and provide their own positive and negative controls for testing activation or interaction between defined proteins. For expression of bait and library proteins, the Clontech Matchmaker LexA two-hybrid system and the
OriGene Duplex-A system use some of the basic set of plasmids described here (see Table 19.2.1 for availability). Forward sequencing primers for bait and library plasmids are included in the Clontech kit, and Insert Screening Amplimer Sets for both plasmids can be acquired separately. Additional related products from Clontech include KC8 competent cells, anti-LexA monoclonal antibodies, a yeast transformation system, a yeast plasmid isolation kit, and an EGY48 partner strain for yeast mating to facilitate the analysis of interaction specificity. OriGene has a generally similar product line to Clontech. In contrast, Invitrogen has substantially modified the Interaction Trap core reagents to develop its own bait and library plasmids. pHybLex/Zeo, a novel bait plasmid, is ∼50% smaller than the original pEG202 (making it easier to clone into), and it has an enriched polylinker. Significantly, it replaces both the Apr and HIS3 genes with a novel gene that confers resistance to the antibiotic Zeocin (supplied with the kit), which provides selection in both bacteria and yeast. This elimination of auxotrophic selection for the bait plasmid renders the LexA-fusion construct usable with libraries and strains from all existing two-hybrid systems and additionally facilitates the direct selection of library plasmid in strains other than KC8. Some changes, which are designed to make the vector easier to use, have also been introduced in the library vector pYESTrp (e.g., it uses a V5 epitope tag for protein detection). The Invitrogen kit, termed Hybrid Hunter, includes the bait/library/reporter plasmids and EGY48 yeast strain as noted, and additionally includes primer sets for bait and library plasmids and the L40 yeast strain, should an investigator wish to use a HIS3 auxotrophy selection. Additional related products from Invitrogen include antibodies for detection of bait and prey fusion proteins (antiLexA and anti-V5), pJG4-5 library vector primers, and a Transformation Kit. A significant advantage of the entry of commercial entities into the Interaction Trap field is the rapid increase in the number of compatible cDNA libraries. A list of currently available premade libraries available from these companies is presented in Table 19.2.3, and custommade libraries are also available upon request. Because new libraries and other related reagents are being constantly added to the line of two-hybrid related products, it is advisable to contact the companies or visit their Web sites (www.clontech.com, www.invitrogen.com, and www.origene.com) for the latest information.
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Finally, over the last several years, a number of groups have adapted basic two-hybrid strategies to more specialized applications, and they have devised strategies to broaden their basic functionality. Interaction Mating (Finley and Brent, 1994) has been used to establish extended networks of targeted protein-protein interaction. In this approach, a panel of LexAfused proteins are transformed into a MATa haploid selective strain (such as RFY206), a panel of activation-domain fused proteins are transformed into a suitable MATα haploid (such as EG448), and the two panels are crossgridded against each other for mating. Selected diploids are then screened by replica plating to selective medium. This approach complements library screening in large-scale applications, such as proposed definition of interaction maps for entire genomes (Bartel et al., 1996). Interaction mating has also provided the basis for an alternative two-hybrid hunt protocol (see Alternate Protocol 2), useful in cases when a single library will be screened with different baits. In this approach (Bendixen et al., 1994; Finley and Brent, 1994: Kolonin and Finley, 1998), a library is introduced into a single strain, like EGY48, and aliquots are stored frozen. To conduct a hunt, an aliquot is thawed and mated with a strain expressing a bait. This allows one to avoid repeated high-efficiency transformations, since a single library transformation can provide enough pretransformed yeast to conduct dozens of interactor hunts. Moreover, some yeast strains pretransformed with libraries are becoming commercially available, which may eliminate altogether the need to conduct a high-efficiency library transformation for some researchers. Two-hybrid approaches have been shown to be effective in identifying small peptides with biological activities on selected baits (Yang et al., 1995; Colas et al., 1996), which may prove to be useful as a guide to targeted drug design. Rapid screening protocols have been devised using custom-synthesized libraries expressing sheared plasmid DNA to facilitate rapid mapping of interaction interfaces (Stagljar et al., 1996). Osborne and coworkers have demonstrated the effectiveness of a tribrid (or tri-hybrid) approach, in which an additional plasmid expresses a tyrosine kinase to specifically modify a bait protein, allowing detection of SH2domain-containing partner proteins that recognize specific phosphotyrosine residues (Osborne et al., 1995). A variety of more elaborate tribrid approaches, in which a DNA-binding domain fused protein is used to present an
intermediate nonprotein compound for interaction with a library, have been developed and proven effective. These approaches have allowed the identification of proteins binding specific drug ligands (Chiu et al., 1994; Licitra and Liu, 1996), as well as the identification of proteins binding to RNA sequences (SenGupta et al., 1996; Wang et al., 1996). It is expected that the range of utility of these systems will continue to expand.
Critical Parameters and Troubleshooting To maximize chances of a successful interactor hunt, a number of parameters should be taken into account. Before attempting a screen, bait proteins should be carefully tested to ensure that they have little or no intrinsic ability to activate transcription. Bait proteins must be expressed at reasonably high levels and must be able to enter the yeast nucleus and bind DNA (as confirmed by the repression assay). Optimally, integrity and levels of bait proteins should be confirmed by immunoblot analysis, using an antibody to either LexA or the fused domain. In particular, at this time, bait proteins that have extensive transmembrane domains or are normally excluded from the nucleus are not likely to be productively used in a library screen. Proteins that are moderate to strong activators will need to be truncated to remove activating domains before they can be used. If a protein neither activates nor represses, the most likely reason is that it is not being made. This can be determined by immunoblot analysis of a crude lysate protein extract of EGY48 (UNIT 10.10; Samson et al., 1989) containing the plasmid, using anti-LexA antibodies as primary antiserum. If the full protein is not made, it may be possible to express truncated derivatives of the protein. If the protein is made, but still does not repress, it may not enter the yeast nucleus effectively, although this appears to be a relatively rare problem. In this case, introducing the coding sequence for the fused moieity into a LexA fusion vector containing a nuclear localization motif (e.g., pJK202; J. Kamens, BASF, Worcester, Mass., unpub. observ.) may solve the problem. The test for the leucine (Leu) requirement is extremely important to determine whether the bait protein is likely to yield an unworkably high background. The LEU2 reporter in EGY48 is more sensitive than the pSH18-34 reporter for some baits (Estojak et al., 1995). Therefore, it is possible that a bait protein demonstrating little or no signal in a β-galac-
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tosidase assay may nevertheless permit some growth on −Leu medium. If this occurs, there are several options. First, a less sensitive strain can be used, as described in the text. Second, background can sometimes be reduced further by making the EGY strain diploid (e.g., D. Krainc, Harvard Medical School, Boston, Mass.; R. Finley and R. Brent, unpub. observ.) or by performing the hunt by interaction mating as described in Alternate Protocol 2. A third option is to attempt to truncate the bait protein to remove activating function. In general, it is useful to extrapolate from the number of cells that grow on −Leu medium to the number that would be obtained in an actual library screen, and determine if this is a background level that can be tolerated. For example, if two colonies arise from 100,000 plated cells on −Leu medium, 200 to 400 would be expected in an actual screen of 106 cDNAs. Although this is a high initial number of positives, the vast majority should be eliminated immediately through easily performed controls. This is a judgment call. Finally, very rarely it happens that a bait that appears to be well behaved and negative for transcriptional activation through all characterization steps will suddenly develop a very high background of transcriptional activation following library transformation. The reason for this is currently obscure, and no means of addressing this problem has as yet been found: such baits are hence inappropriate for use in screens. The protocols described in this unit use initial screening with the most sensitive reporters followed by substitution with less sensitive reporters if activation is detected. An obvious question is, why not start out working with extremely stringent reporters and know immediately whether the system is workable? In fact, some researchers routinely use a combination of pJK103 or pRB1840 with EGY191, and obtain proteins that to date appear to be biologically relevant partners from library screens. However, extensive comparison studies using interactors of defined in vitro affinity with different combinations of LacZ and LEU2 reporters (Estojak et al., 1995) have indicated that although the most sensitive reporters (pSH1834) may in some cases be prone to background problems, the most stringent reporters (EGY191, pRB1840) may miss some interactions that certainly are biologically relevant and occur inside cells. In the end, the choice of reporters devolves to the preference of individual investigators: the bias of the authors is to cast a broad net in the early stages of a screen,
and hence to use more sensitive reporters when practicable. It is important to move expeditiously through characterization steps and to handle yeast transformed with bait plasmids with care. In cases where yeasts have been maintained on plates for extended periods (e.g., 4 days at room temperature or >2 to 3 weeks at 4°C), unexpected problems may crop up in subsequent library screens. The transformation protocol is a version of the lithium acetate transformation protocol described by Schiestl and Gietz (1989) and Gietz et al. (1992; see APPENDIX 4L) that maximizes transformation efficiency in Saccharomyces cerevisiae and produces up to 105 colonies/µg plasmid DNA. In contrast to Escherichia coli, the maximum efficiency of transformation for S. cerevisiae is ∼104 to 105/µg input DNA. It is extremely important to optimize transformation conditions before attempting an interactor hunt. Perform small-scale pilot transformations to ensure this efficiency is attained and to avoid having to use prohibitive quantities of library DNA. In addition, as for any effort of this type, it is a good idea to obtain or construct a library from a tissue source in which the bait protein is known to be biologically relevant. In practice, the majority of proteins isolated by interaction with a LexA fusion turn out to be specific for the fused domain; a smaller number are nonspecifically sticky, and to date there appears to have been only one isolation from a eukaryotic library of a protein specific for LexA. However, it is generally informative to retest positive clones on more than one LexA bait protein; ideally, library-derived clones should be tested against the LexA fusion used for their isolation, several LexA fusions to proteins that are clearly unrelated to the original fusion, and if possible, several LexA fusions that there is reason to believe are related to the initial protein (e.g., if the initial probe was LexA-Fos, a good related set would include LexA-Jun and LexA-GCN4). Colony selection for master plate production is one of the more variable parts of the procedure. For strong interactors, colonies will grow up in 2 days. However, if plates are left at 30°C, new colonies will continue to appear every day. Those that appear rapidly are most likely to reflect interactors that are biologically relevant to the bait protein. Those that appear later may or may not be relevant. However, many parameters can delay the time of colony formation of cells that contain valid interactions, including the strength of the interaction
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and the level of expression of the library-encoded protein.
Anticipated Results Depending on the protein used as bait, anywhere from zero to hundreds of specific interactors will be obtained from 106 primary transformants.
Time Considerations If all goes well, once the required constructions have been made it will take ∼1 week to perform yeast transformations, obtain colonies, and determine whether bait proteins are appropriate. It will take a second week to perform library transformations, replate to selective medium, and obtain putative positives. A third week will be required to rescue the plasmid from the yeast, passage it through E. coli, transform fresh yeast, and confirm specificity.
Literature Cited Bartel, P.L., Chien, C.-T., Sternglanz, R., and Fields, S. 1993. Using the two-hybrid system to detect protein-protein interactions. In Cellular Interactions in Development: A Practical Approach (D.A. Hartley, ed.) pp. 153-179. Oxford University Press, Oxford. Bartel, P.L., Roecklein, J.A., SenGupta, D., and Fields, S. 1996. A protein linkage map of Escherichia coli bacteriophage T7. Nature Genet. 12:72-77. Bendixen, C., Gangloff, S., and Rothstein, R. 1994. A yeast mating-selection scheme for detection of protein-protein interactions. Nucl. Acids Res. 22:1778-1779. Breeden, L. and Nasmyth, K. 1985. Regulation of the yeast HO gene. Cold Spring Harbor Symp. Quant. Biol. 50:643-650. Brent, R. and Ptashne, M. 1984. A bacterial repressor protein or a yeast transcriptional terminator can block upstream activation of a yeast gene. Nature 312:612-615. Brent, R. and Ptashne, M. 1985. A eukaryotic transcriptional activator bearing the DNA specificity of a prokaryotic repressor. Cell 43:729-736. Chien, C.-T., Bartel, P.L., Sternglanz, R., and Fields, S. 1991. The two-hybrid system: A method to identify and clone genes for proteins that interact with a protein of interest. Proc. Natl. Acad. Sci. U.S.A. 88:9578-9582. Chiu, M.I., Katz, H., and Berlin, V. 1994. RAPT1, a mammalian homolog of yeast Tor, interacts with the FKBP12/rapamycin complex. Proc. Nat. Acad. Sci. U.S.A. 91:12574-12578. Colas, P., Cohen, B., Jessen, T., Grishina, I., McCoy, J., and Brent, R. 1996. Genetic selection of peptide aptamers that recognize and inhibit cyclindependent kinase 2. Nature 380:548-550. Duby, A., Jacobs, K.A., and Celeste, A. Using synthetic oligonucleotides as probes. In Current
Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 6.4.16.4.10. John Wiley & Sons, New York. Durfee, T., Becherer, K., Chen, P.L., Yeh, S.H., Yang, Y., Kilburn, A.E., Lee, W.H., and Elledge, S.J. 1993. The retinoblastoma protein associates with the protein phosphatase type 1 catalytic subunit. Genes & Dev. 7:555-569. Estojak, J., Brent, R., and Golemis, E.A. 1995. Correlation of two-hybrid affinity data with in vitro measurements. Mol. Cell. Biol. 15:58205829. Fearon, E.R., Finkel, T., Gillison, M.L., Kennedy, S.P., Casella, J.F., Tomaselli, G.F., Morrow, J.S., and Dang, C.V. 1992. Karyoplasmic interaction selection strategy: A general strategy to detect protein-protein interactions in mammalian cells. Proc. Nat. Acad. Sci. U.S.A. 89:7958-7962. Fields, S. and Song, O. 1989. A novel genetic system to detect protein-protein interaction. Nature 340:245-246. Finley, R.L., Jr., and Brent, R. 1994. Interaction mating reveals binary and ternary connections between Drosophila cell cycle regulators. Proc. Natl. Acad. Sci. U.S.A. 91:12980-12984. Gietz, D., St. Jean, A., Woods, R.A., and Schiestl, R.H. 1992. Improved method for high-efficiency transformation of intact yeast cells. Nucl. Acids Res. 20:1425. Golemis, E.A. and Brent, R. 1992. Fused protein domains inhibit DNA binding by LexA. Mol. Cell Biol. 12:3006-3014. Grunstein, M., and Hogness, D.S. 1975. Colony hybridization: A method for the isolation of cloned DNAs that contain a specific gene. Proc. Natl. Acad. Sci. U.S.A. 72:3961-3965. Gyuris, J., Golemis, E.A., Chertkov, H., and Brent, R. 1993. Cdi1, a human G1- and S-phase protein phosphatase that associates with Cdk2. Cell 75:791-803. Kaiser, C., Michaelis, S., and Mitchell, A. 1994. Methods in Yeast Genetics, a Cold Spring Harbor Laboratory Course Manual, pp.135-136. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Kolonin, M.G. and Finley, R.L., Jr. 1998. Targeting cyclin-dependent kinases in Drosophilia with peptide aptamers. Proc. Natl. Acad. Sci. U.S.A. In press. Licitra, E.J. and Liu, J.O. 1996. A three-hybrid system for detecting small ligand-protein receptor interactions. Proc. Nat. Acad. Sci. U.S.A. 93:12817-12821. Ma, J. and Ptashne, M. 1987. A new class of yeast transcriptional activators. Cell 51:113-119. Ma, J. and Ptashne, M. 1988. Converting an eukaryotic transcriptional inhibitor into an activator. Cell 55:443-446. Osborne, M., Dalton, S., and Kochan, J.P. 1995. The yeast tribrid system: Genetic detection of transphosphorylated ITAM-SH2 interactions. Bio/Technology 13:1474-1478.
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Ruden, D.M., Ma, J., Li, Y., Wood, K., and Ptashne, M. 1991. Generating yeast transcriptional activators containing no yeast protein sequences. Nature 350:426-430. Samson, M.-L., Jackson-Grusby, L., and Brent, R. 1989. Gene activation and DNA binding by Drosophila Ubx and abd-A proteins. Cell 57:1045-1052. Schiestl, R.H. and Gietz, R.D. 1989. High-efficiency transformation of intact yeast cells using single-stranded nucleic acids as a carrier. Curr. Genet. 16:339-346. SenGupta, D.J., Zhang, B., Kraemer, B., Pochart, P., Fields, S., and Wickens, M. 1996. A three-hybrid system to detect RNA-protein interactions in vivo. Proc. Nat. Acad. Sci. U.S.A. 93:8496-8501. Sikorski, R.S. and Hieter, P. 1989. A system of shuttle vectors and yeast host strains designed for efficient manipulation of DNA in Saccharomyces cerevisiae. Genetics 122:19-27. Stagljar, I., Bourquin, J.-P., and Schaffner, W. 1996. Use of the two-hybrid system and random sonicated DNA to identify the interaction domain of a protein. BioTechniques 21:430-432. Strauss, W.M. 1993. Using DNA fragments as probes. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 6.3.1-6.3.6. John Wiley & Sons, New York. Struhl, K. 1987. Subcloning of DNA fragments. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 3.16.1-3.16.11. John Wiley & Sons, New York. Treco, D.A. and Winston, F. 1992. Growth and manipulation of yeast. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 13.2.1-13.2.12. John Wiley & Sons, New York. Vasavada, H.A., Ganguly, S., Germino, F.J., Wang, Z.X., and Weissman, S.M. 1991. A contingent replication assay for the detection of protein-protein interactions in animal cells. Proc. Nat. Acad. Sci. U.S.A. 88:10686-10690. Vojtek, A.B., Hollenberg, S.M., and Cooper, J.A. 1993. Mammalian Ras interacts directly with the serine/threonine kinase Raf. Cell 74:205-214. Wang, Z.F., Whitfield, M.L., Ingledue, T.C.3, Dominski, A., and Marzluff, W.F. 1996. The protein that binds the 3′ end of histone mRNA: A novel RNA-binding protein required for histone pre-mRNA processing. Genes & Dev. 10:3028-3040. Watson, M.A., Buckholz, R., and Weiner, M.P. 1996. Vectors encoding alternative antibiotic resistance for use in the yeast two-hybrid system. BioTechniques 21:255-259.
West, R.W.J., Yocum, R.R., and Ptashne, M. 1984. Saccharomyces cerevisiae GAL1-GAL10 divergent promoter region: Location and function of the upstream activator sequence UASG. Mol. Cell Biol. 4:2467-2478. Yang, M., Wu, Z., and Fields, S. 1995. Protein-peptide interactions analyzed with the yeast two-hybrid system. Nucl. Acids Res. 23:1152-1156.
Key Reference Gyuris et al., 1993. See above. Initial description of interaction trap system.
Internet Resources http://www.clontech.com http://cmmg.biosci.wayne.edu/rfinley/lab.html Source of two-hybrid information, protocols, and links. http://www.invitrogen.com http://www.origene.com Commercial sources for basic plasmids, strains, and libraries for interaction trap experiments. [email protected] [email protected] Sources of interaction trap plasmids for specialized interactions. http://www.fccc.edu:80/research/labs/golemis/ InteractionTrapInWork/html Database for false postive proteins detected in interaction trap experiments; analysis of two-hybrid usage. http://xanadu.mgh.harvard.edu/brentlabhome page4.html Database of interaction trap protocols and related issues.
Contributed by Erica A. Golemis and Ilya Serebriiskii Fox Chase Cancer Center Philadelphia, Pennsylvania Russell L. Finley, Jr. and Mikhail G. Kolonin (hunt by interaction mating) Wayne State University School of Medicine Detroit, Michigan Jeno Gyuris Mitotix, Inc. Cambridge, Massachusetts Roger Brent The Molecular Sciences Institute Berkeley, California
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Phage-Based Expression Cloning to Identify Interacting Proteins
UNIT 19.3
Interaction cloning (also known as expression cloning) is a technique to identify and clone genes which encode proteins that interact with a protein of interest, or “bait” protein. Phage-based interaction cloning requires a gene encoding the bait protein and an appropriate expression library constructed in a bacteriophage expression vector, such as λgt11. The gene encoding the bait protein is used to produce recombinant fusion protein in E. coli. The cDNA is radioactively labeled with 32P. A recognition site for cyclic adenosine 3′, 5′-phosphate (cAMP)–dependent protein kinase (protein kinase A; PKA) is introduced into the recombinant fusion protein to allow its enzymatic phosphorylation by PKA and [γ-32P]ATP. The procedure presented here (see Basic Protocol) involves a fusion protein consisting of bait protein and glutathione-S-transferase (GST) with a PKA site at the junction between them (the protocol can, however, be adapted to use other PKA-containing recombinant proteins). The labeled protein is subsequently used as a probe to screen a λ bacteriophage-derived cDNA expression library, which expresses β-galactosidase fusion proteins that contain in-frame gene fusions. The phages lyse cells, form plaques, and release fusion proteins that are adsorbed onto nitrocellulose membrane filters. The filters are blocked with excess nonspecific protein to eliminate nonspecific binding and probed with the radiolabeled bait protein (see Fig. 19.3.1). This procedure leads directly to the isolation of genes encoding the interacting protein, bypassing the need for purification and microsequencing or for antibody production. NOTE: Radioactive label is used in this protocol, and appropriate precautions and shielding should be used (APPENDIX 2B). STRATEGIC PLANNING There are two important choices one must make to begin this procedure: (1) how to design the bait protein and (2) how to construct or acquire an appropriate phage-derived expression library. Vectors for recombinant fusion protein expression that contain a PKA recognition site can be obtained commercially. Several companies now sell these vectors with various affinity tags such as GST (Pharmacia Biotech), histidine (Novagen), or calmodulin-binding protein (Stratagene). Alternatively, one can engineer the PKA recognition site (the five–amino acid sequence RRASV) into existing vectors by using synthetic DNA that encodes it. Lambda-derived expression libraries that direct the expression of cDNAs made from many different mRNA sources are widely available. Alternatively, a library may be constructed for a particular experimental purpose; when designing a library to be used in expression cloning, several points should be considered. Libraries made from cDNA synthesized with random primers or an oligo dT-primer during first-strand synthesis (Klickstein et al., 1995; Klickstein and Neve, 1991) can often be advantageous in that multiple clones representing different portions of the same protein can be identified in the screening. Analysis of coding regions present in these multiple clones could provide useful information about what region of the protein is responsible for the observed interaction. Full-length cDNA clones can subsequently be obtained from other available libraries. Many suitable λ vectors are available for constructing cDNA expression libraries. The most widely used cDNA expression vector has been λgt11 (Huynh et al., 1985). There are modifications of these λ vectors, however, that facilitate the recovery of the cDNA Contributed by Julie M. Stone Current Protocols in Protein Science (1999) 19.3.1-19.3.9 Copyright © 1999 by John Wiley & Sons, Inc.
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inserts by avoiding the necessity of time-consuming λ phage DNA preparations. These modified vectors include those that make use of the Cre-lox recombination for in vivo conversion of recombinant phages into plasmid DNA in Cre recombinase-expressing host strains (e.g., λZipLox, Life Technologies) as well as those that employ helper phage for in vivo excision of a phagemid vector (e.g., the λZAP series of vectors, Stratagene). Most λ vectors available produce fusions of the cDNA inserts to β-galactosidase, because they are cloned into the lacZ gene. However, some λ-derived expression vectors (e.g., λSCREEN-1, Novagen) direct expression of proteins fused to the T7 DNA polymerase promoter (gene 10 under control of the T7 promoter) and may yield higher expression of the library proteins (Margolis et al., 1992). BASIC PROTOCOL
INTERACTION CLONING Phage-based interaction expression cloning is a simple, rapid, and powerful technique to identify interacting proteins. A protein of interest is expressed as a recombinant fusion protein and labeled with 32P at a serine residue in an engineered PKA recognition site to facilitate detection. β-galactosidase proteins that are fused in-frame to cDNA inserts in a bacteriophage λ-derived expression library are produced by the phage and adsorbed onto nitrocellulose filters. The filters are then screened with the radiolabeled protein probe to identify phage clones that express an interacting protein.
A G ST
PKA ba it
B probed with GST-bait probed with GST
Phage-Based Expression Cloning
Figure 19.3.1 Schematic representation of the interaction cloning technique used to identify proteins that associate with a protein of interest (bait protein), and the expected results of a successful screen. (A) Expression of β-galactosidase fusion proteins from in-frame cDNA inserts of a λgt11 Arabidopsis library is induced with IPTG-impregnated nitrocellulose membrane filters (indicated by an oval). Filters are probed with GST-bait fusion protein labeled with 32P (•) at the PKA recognition site located at the junction of the fusion. Interacting clones are detected by autoradiography. (B) Representative autoradiogram of a tertiary screen of a positive plaque after the control experiment to determine whether the interaction is specific for the bait portion of the probe. The top half of the filter is probed with GST-bait while the bottom half is probed with GST alone as a control.
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Materials cAMP-dependent protein kinase (PKA; e.g., 250-U lots from Sigma) 40 mM DTT, prepared fresh 10× PKA buffer (see recipe) 10 mCi/ml [γ-32P]ATP (6000 mCi/mmol) Purified glutathione-S-transferase (GST)–bait protein fusion protein with a PKA recognition site (UNIT 6.6), at ∼0.1 to 1 µg/µl concentration Z′-KCl (see recipe), ice cold Sephadex G-50 equilibrated in Z′-KCl E. coli Y1090r− or other appropriate host strain LB medium containing appropriate selective antibiotic (APPENDIX 4B), 10 mM MgSO4, and 0.2% maltose 10 mM MgSO4 10 mM IPTG (Table 1.4.2) 150- or 100-mm LB plates (with antibiotic, if necessary; APPENDIX 4B) 0.7% top agarose (APPENDIX 4B), 47°C Tris-buffered saline with Triton X-100 (TBS-T; see recipe) India ink HEPES blocking buffer (HBB; see recipe) Binding buffer (BB; see recipe) Suspension medium (SM; see recipe) Chloroform 3-ml disposable plastic columns or disposable syringe and glass wool Scintillation counter Tabletop centrifuge or equivalent Nitrocellulose membrane filters (137- and 82-mm disks) 22-G needle Additional reagents and equipment for preparation and purification of recombinant glutathione-S-transferase fusion protein (UNIT 6.6), SDS-PAGE (optional; UNIT 10.1), autoradiography (UNIT 10.11), titering and plating λ phage to generate plaques (Lech and Brent, 1988; Quertermous, 1996), and purification of bacteriophage clones (Quertermous, 1987) Prepare the 32P-labeled protein probe 1. Resuspend 250 U PKA in 25 µl freshly prepared 40 mM DTT. Let the reconstituted enzyme stand at room temperature ∼10 min before use. It is important to use freshly prepared 40 mM DTT. PKA is extremely unstable after reconstitution; the enzyme stock solution can be stored temporarily at 4°C but retains activity for only 2 to 3 days.
2. Prepare a phosphorylation reaction mixture containing: 1 µl 10 U/µl PKA (from step 1) 3 µl 10× PKA buffer 5 µl 10 mCi/ml (6000 mCi/mmol) [γ-32P]ATP 1 to 10 µl (∼1 µg) purified GST-bait fusion protein H2O to 30 µl. Incubate 1 hr at room temperature. A fusion protein unrelated to the bait protein, but containing the PKA recognition site, should also be expressed for use as a control to determine whether the observed interaction is specific for the bait moiety. Due to the instability of PKA, it is advisable to label both the bait protein and the unrelated control fusion protein simultaneously (in separate reactions).
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3. Add 170 µl ice-cold Z′-KCl (to stop reaction) and store on ice until use. 4. Prepare a gel filtration column by pouring Sephadex G-50 equilibrated in Z′-KCl into a 3-ml disposable plastic column (or a 3-ml syringe with a glass wool plug) for a final bed volume of ∼3 ml. Allow the column to drain until the level of the buffer is at the top of the column bed. The Sephadex G-50 can be swelled in water or buffer and stored at 4°C for months. In this case, five to ten column volumes of Z′-KCl can be used to equilibrate the column after it is poured.
5. Load the entire phosphorylation reaction (from step 3) onto the column and collect the effluent in a 1.5-ml microcentrifuge tube. Place a second tube under the column, add a 200-µl aliquot of Z′-KCl, and collect the effluent again. Repeat the above loading and collecting steps an additional ten times to collect a total of twelve 200-µl fractions. 6. Measure the Cerenkov counts with a scintillation counter to identify the fractions with the highest specific activities, and calculate cpm/µl. Typically two peaks of radioactivity are observed. The first peak elutes in fractions five to nine and corresponds to labeled protein. The second peak, usually found in the last few fractions collected, corresponds to unincorporated ATP and should be discarded. The hottest fraction(s), which elute first, should be used. Fractions can be stored at 4°C for several weeks.
7. Optional: Analyze a small amount (1 to 2 µl) of the 32P-labeled protein probe by SDS-PAGE and autoradiography (UNITS 10.1 & 10.11). Typically two strong signals are observed: one at the predicted size of the GST-bait fusion protein and the other at the predicted molecular weight of the GST alone (28 kDa). This is due to the fusion protein’s inherent susceptibility to protease cleavage at the GST-bait protein junction. The presence of labeled GST protein in the probe should not interfere with the screen; a control experiment with labeled GST will be performed at the later stages of screening.
Prepare host strain cells and dilution of bacteriophage cDNA library 8. Using serial dilution, determine the titer of the bacteriophage cDNA library. See Lech and Brent (1988) for serial dilution procedure.
9. Grow an overnight 50-ml culture of E. coli Y1090r− (or other appropriate host strain; APPENDIX 4B) in LB medium containing an appropriate selective antibiotic, 10 mM MgSO4, and 0.2% maltose. 10. Centrifuge cells 10 min at 2000 × g, room temperature, and resuspend in 25 ml of 10 mM MgSO4. The resuspended cells can be stored up to 1 week at 4°C before use.
Prepare the filters to screen the bacteriophage cDNA expression library 11. Soak eight 137-mm nitrocellulose filters in 10 mM IPTG for 15 min at room temperature and air dry. IPTG-impregnated filters can be stored in a petri plate until use.
Phage-Based Expression Cloning
12. Prepare eight 1.5-ml microcentrifuge tubes each containing 0.6 ml Y1090r− cells (from step 10) and ∼40,000 pfu bacteriophage (from step 8), and incubate 15 min at 37°C.
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13. Add contents of each tube to 7 ml of 0.7% top agarose at 47°C and pour onto 150-mm LB plates (with antibiotic, if necessary). Incubate plates ∼3 hr at 42°C, until small plaques are visible. 14. Overlay the plates with IPTG-impregnated filters and incubate an additional 6 to 8 hr at 37°C. Formation of plaques in the absence of induction of the lacZ gene promoter ensures that any library-encoded proteins that are deleterious to phage growth will not be expressed while the phage are forming a plaque. Incubation in the presence of IPTG-impregnated filters is usually performed for a 6- to 8-hr period, but can proceed overnight for convenience.
15. Chill plates 15 min (or overnight) at 4°C. 16. Pierce each filter in several locations with a 22-G needle dipped in India ink to mark orientation. Remove filters from plates and wash 15 min in TBS-T at room temperature with shaking. Screen bacteriophage cDNA expression library 17. Incubate filters 1 to 4 hr with rocking at 4°C in 100 ml HBB. This blocking step (to reduce nonspecific binding) can also be performed overnight for convenience.
18. Incubate overnight with rocking at 4°C in BB containing 2.5–5 × 105 cpm/ml of radiolabeled fusion protein (from step 6). All eight filters can be placed in one 150-mm plate and probed with 30 to 40 ml of solution. The plate is wrapped in Parafilm and placed in a Plexiglas box for shielding. Alternatively, the filters can be placed with solution in a heat-sealable bag. The probe solution can be stored at 4°C and used for the subsequent secondary and tertiary screenings.
19. Wash membrane filters three times, 10 min each, in 100 ml BB with shaking at room temperature. Air dry and expose to film (see UNIT 10.11). 20. Using the large end of a Pasteur pipet, take an agarose plug at the position of the positive clone. Place into a 1.5-ml microcentrifuge tube and add 1 ml SM and 1 drop of chloroform. Agarose plugs can be stored at 4°C for months. Therefore, if many putative positive clones are identified in the primary screen, they can all be stored for future analysis if necessary.
21. Determine the titer by serial dilution and perform successive screening procedures to obtain purified clones. See Lech and Brent (1988) for serial dilution procedure and Quertermous (1987) for screening methods. Subsequent screens are performed on 100-mm LB plates with ∼2000 pfu/plate for secondary screens and ∼300 to 500 pfu/plate for tertiary screens. Before purified clones are obtained (usually during the tertiary screen), a control to eliminate clones that might interact with the GST fusion portion of the probe should be performed. To do this, cut the filters in half and probe one half with the labeled GST fusion to the protein of interest and the other with labeled GST alone or an unrelated control GST fusion protein (see Fig. 19.3.1).
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REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. All solutions should be prepared from sterile, autoclaved stock solutions, except Z′-KCl, which should be filter sterilized. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Binding buffer (BB) 20 mM HEPES⋅OH, pH 7.4 7.5 mM KCl 0.1 mM EDTA 2.5 mM MgCl2 1% (w/v) nonfat dry milk The solution can be prepared without milk and stored indefinitely at room temperature (add milk prior to use).
HEPES blocking buffer (HBB) 20 mM HEPES⋅OH, pH 7.4 5 mM MgCl2 1 mM KCl 5% (w/v) nonfat dry milk The solution can be prepared without milk and stored indefinitely at room temperature (add milk prior to use).
PKA buffer, 10× 200 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 10 mM DTT 1 M NaCl 120 mM MgCl2 Store up to 6 months at room temperature Suspension medium (SM) 5.8 g NaCl 2 g MgSO4⋅7H2O 50 ml 1 M Tris⋅Cl, pH 7.5 (APPENDIX 2E) 5 ml 2% (w/v) gelatin H2O to 1 liter Sterilize by autoclaving. Store up to several months at 4°C Gelatin is prepared by adding 2 g gelatin to 100 ml H2O, then autoclaving to dissolve when needed.
Tris-buffered saline with Triton X-100 (TBS-T) 10 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 150 mM NaCl 0.05% (v/v) Triton X-100 Store up to 6 months at room temperature
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Z′-KCl 25 mM HEPES⋅OH, pH 7.4 12.5 mM MgCl2 20% (w/v) glycerol 100 mM KCl 1 mg/ml BSA 1 mM DTT Filter sterilize Store up to 6 months at 4°C
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COMMENTARY Background Information Historically, interacting proteins have been isolated by biochemical approaches, which required purification of the interacting protein for antibody production or microsequencing before a clone encoding the protein could be identified. Two of the molecular biological approaches described in this chapter, the yeast two-hybrid system (UNIT 19.2) and phage-based interaction expression cloning (this unit), however, directly yield a clone encoding the interacting protein. Bacteriophage cDNA expression libraries are commonly screened using antibodies (see St. John, 1990 and references therein) or radiolabeled DNA probes to identify DNA-binding proteins (see Singh, 1991 and references therein). Modifications to identify interacting proteins include screening with the protein of interest (the bait protein) and detecting that protein with antibodies (Chapline et al., 1993). However, by screening with a radiolabeled protein probe, one avoids the additional incubations and washes which are necessary for immunodetection but that increase the likelihood of disrupting weak protein-protein interactions. 125I-labeled protein probes have been used successfully to screen for interacting proteins (Hoeffler et al., 1991), although the technique described in this unit avoids the need to label the proteins with 125I, and the complications of handling this isotope. Protein probes autophosphorylated with 32P were originally used to screen cDNA expression libraries in the isolation of proteins that interact with receptor protein kinases, a technique referred to as CORT (cloning of receptor targets; Skolnik et al., 1991; Lowenstein et al., 1992; Margolis et al., 1992). By introducing a PKA recognition site into the protein probe, the technique was made suitable for proteins that were not themselves protein kinases (Blanar and Rutter, 1992; Kaelin et al., 1992). Phage-based interaction expression cloning as described in this unit has been used successfully to identify many interacting proteins, but may not be successful for all types of interactions. For example, many related proteins (known as A-kinase anchoring proteins, or AKAPs) have been identified by their ability to interact with the type II cAMP-dependent kinase regulatory subunit, RII (Carr and Scott, 1992). All the AKAPs can bind RII under denaturing conditions (Lester et al., 1996), illustrating that the success of interaction cloning is often dependent on the nature of the interactions and that interactions less dependent on three-di-
mensional structure may be favored. However, there are many examples of proteins identified by this technique that cannot interact under the denaturing conditions often used to confirm the binding (commonly referred to as overlay assays or Far Western analysis). For example, a protein domain identified by its interaction with a plant receptor–like protein kinase is capable of interacting with a denatured form of the bait protein, but is unable to interact when the interacting protein is denatured (Stone et al., 1994). This fact is consistent with the idea that immobilization on membranes denatures some proteins but not others.
Critical Parameters and Troubleshooting The success of phage-based interaction expression cloning is inherently dependent on the quality of the protein probe used and the extent of representation of the bacteriophage cDNA library. GST fusion proteins are often obtained in high quantity in a soluble form, avoiding the necessity of solubilizing and refolding during protein purification from E. coli extracts. In most cases the PKA recognition site is readily accessible, and allows production of radiolabeled protein with a high specific activity. The strength of the protein-protein interaction is also critical. Depending on the nature of the interaction, a filter-binding technique might not be suitable. This technique may not detect weak or transient interactions, such as an enzyme/substrate interaction, and techniques such as the yeast two-hybrid system (UNIT 19.2) might be more appropriate. If no positive clones are identified or if background is high, it may be helpful to add reducing agents or detergents or to alter the salt conditions of the binding and wash solutions (Vinson et al., 1988). Moreover, denaturation and renaturation of the proteins on the filters using 6 M guanidine⋅HCl (Singh, 1991) may facilitate the recovery of clones expressing interacting proteins, because adsorption of the β-galactosidase fusion proteins to nitrocellulose filters may alter the conformation of the proteins. For a discussion of common problems with this procedure and their diagnosis and possible solutions, see Table 19.3.1.
Anticipated Results Depending on the nature of the interaction being sought by this technique, many interact-
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Table 19.3.1
General Troubleshooting Guide for Interaction Cloning
Problem
Possible cause
Solution
High background
Insufficient washing
Use more BB wash solution
Poor choice of nitrocellulose membrane filters Insufficient blocking
Try a different supplier for membranes Block overnight in HBB
Poor-quality protein probe
Check the probe by SDS-PAGE and autoradiography
Wash conditions too stringent
Reduce wash times Vary salt and/or detergent concentration
Poorly folded library fusion proteins
Perform 6 M guanidine⋅HCl denaturation/renaturation (Singh, 1991)
Wash and binding conditions insufficiently stringent Library biased to proteins that interact with the affinity tag
Increase salt and/or detergent in BB
No positive plaques
Too many positive plaques
ing clones, none, or just a few may be identified. However, the technique is as simple as screening a library by hybridization and often well worth the effort. In any case, observed interactions should be confirmed by other means.
Include unlabeled affinity tag in blocking, binding, and wash solutions
take an additional week. There are a number of steps that can be performed either rapidly or overnight, providing a great deal of convenience and flexibility.
Literature Cited Time Considerations
Phage-Based Expression Cloning
The interaction cloning technique described in this unit is extremely rapid in comparison with other techniques to identify interacting proteins, such as the yeast two-hybrid system (UNIT 19.2). The technique does not require any unusual reagents that would not be readily available in any laboratory routinely engaged in molecular biology, other than the cAMP-dependent protein kinase. Once the appropriate recombinant fusion protein and cDNA expression library are obtained, the primary screen can be completed in a few days. Once the appropriate construct for expression of the GST-bait is obtained, purification of the recombinant GST-bait protein can be achieved in 1 day. 32P labeling of the bait protein requires 2 hr (several additional hours if the optional SDSPAGE and autoradiography are performed). The initial screening of the bacteriophage cDNA library takes 3 days; day 1 to prepare the filters and induce expression of library-encoded proteins, day 2 for blocking and probing overnight with 32 P-labeled bait protein, and day 3 for washing and autoradiography. Subsequent purification of the cDNA clones should only
Blanar, M.A. and Rutter, W.J. 1992. Interaction cloning: Identification of a helix-loop-helix zipper protein that interacts with c-Fos. Science 256:1014-1018. Carr, D.W. and Scott, J.D. 1992. Blotting and bandshifting: Techniques for studying protein-protein interactions. Trends Biochem. Sci. 17:246-249. Chapline, C., Ramsay, K., Klauck, T., and Jaken, S. 1993. Interaction cloning of protein kinase C substrates. J. Biol. Chem. 268:6858-6861. Hoeffler, J.B., Lustbader, J.W., and Chen, C.Y. 1991. Identification of multiple nuclear factors that interact with cyclic AMP response element-binding protein and activation transcription factor-2 by protein interactions. Mol. Endocrinol. 5:256266. Huynh, T.V., Young, R.A., and Davis, R.W. 1985. Constructing and screening cDNA libraries in λgt10 and λgt11. In DNA Cloning: A Practical Approach (D.M. Glover, ed.) pp. 49-78. IRL Press, Oxford. Kaelin, W.G.J., Krek, W., Sellers, W.R., DeCaprio, J.A., Ajchenbaum, F., Fuchs, C.S., Chittenden, T., Li, Y., Farnham, P.J., Blanar, M.A., Livingston, D.M., and Flemington, E.K. 1992. Expression cloning of a cDNA encoding a retinoblastoma-binding protein with E2F-like properties. Cell 70:351-364.
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Klickstein, L.B. and Neve, R. L. 1991. Ligation of linkers or adapters to double-stranded cDNA. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 5.6.1-5.6.10. John Wiley & Sons, New York.
St. John, T. P. 1990. Immunoscreening of fusion proteins produced in lambda plaques. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 6.7.16.7.6. John Wiley & Sons, New York.
Klickstein, L.B., Neve, R.L., Golemis, E.A., and Gyuris, J. 1995. Conversion of mRNA into double-stranded cDNA. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 5.5.1-5.5.14. John Wiley & Sons, New York.
Singh, H. 1991. Detection, purification, and characterization of cDNA clones encoding DNA-binding proteins. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 12.7.1-12.7.10. John Wiley & Sons, New York.
Lech, K. and Brent, R. 1988. Plating lambda phage to generate plaques. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 1.11.1-1.11.4 John Wiley & Sons, New York.
Skolnik, E.Y., Margolis, B., Mohammadi, M., Lowenstein, E., Fischer, R., Drepps, A., Ullrich, A., and Schlessinger, J. 1991. Cloning of PI3 kinase–associated p85 utilizing a novel method of expression/cloning of target proteins for receptor tyrosine kinases. Cell 65:83-90.
Lester, L.B., Coghlan, V.M., Nauert, B., and Scott, J.D. 1996. Cloning and characterization of a novel A-kinase anchoring protein: AKAP220, association with testicular peroxisomes. J. Biol. Chem. 271:9460-9465.
Stone, J.M., Collinge, M.A., Smith, R.D., Horn, M.A. and Walker, J.C. 1994. Interaction of a protein phosphatase with an Arabidopsis serinethreonine receptor kinase. Science 266:793-795.
Lowenstein, E.J., Daly, R.J., Batzer, A.G., Li, W., Margolis, B., Lammers, R., Ullrich, A., Skolnik, E.Y., Bar-Sagi, D., and Schlessinger, J. 1992. The SH2 and SH3 domain-containing protein GRB2 links receptor tyrosine kinases to ras signaling. Cell 70:431-442. Margolis, B., Silvennoinen, O., Comoglio, F., Roonprapunt, C., Skolnik, E., Ullrich, A., and Schlessinger, J. 1992. High-efficiency expression/cloning of epidermal growth factor–receptor-binding proteins with src homology 2 domains. Proc. Natl. Acad. Sci. U.S.A 89:8894-8898. Quertermous, T. 1987. Purification of bacteriophage clones. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 6.5.1-6.5.2. John Wiley & Sons, New York. Quertermous, T. 1996. Plating and transferring bacteriophage libraries. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 6.1.1-6.1.4. John Wiley & Sons, New York.
Vinson, C.R., LaMarco, K.L., Johnson, P.F., Landschulz, W.H., and McKnight, S.L. 1988. In situ detection of sequence-specific DNA binding activity specified by a recombinant bacteriophage. Genes & Dev. 2:801-806.
Key References Blanar and Rutter, 1992. See above. The basic protocol described in this unit is modified directly from the Blanar and Rutter protocol. Huynh et al., 1985. See above. Provides an excellent description of constructing and screening λgt11 cDNA expression libraries.
Contributed by Julie M. Stone Massachusetts General Hospital Boston, Massachusetts
Identification of Protein Interactions
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Detection of Protein-Protein Interactions by Coprecipitation
UNIT 19.4
Coprecipitation of proteins from whole-cell extracts is a valuable approach to test for physical interactions between proteins of interest. When a precipitating antibody is used, this method is referred to as co-immunoprecipitation. Coprecipitation can be used to study interactions between known proteins under a variety of conditions and as a means of identifying components of a complex. Coprecipitation may be the single method of choice, or may be used in combination with other methods that detect protein-protein interactions, such as two-hybrid analysis and copurification schemes (UNIT 19.2), and tests of physical associations using purified proteins. This unit describes basic approaches to immunoprecipitating tagged proteins from whole-cell extracts. The approaches described can be adapted for other systems. In a typical experiment, as described here, cells are lysed and a whole-cell extract is prepared under nondenaturing conditions (see Strategic Planning). The protein is precipitated from the lysate with a solid-phase affinity matrix and the precipitate is tested for the presence of a second specifically associated protein (see Basic Protocol and Alternate Protocol). The approach can be used for native or epitope-tagged proteins for which antibodies are available, or for recombinant proteins that have been engineered to bind with high affinity to a molecule that can be coupled to a solid-phase matrix (see Strategic Planning). The presence of an associated protein is detected by separating the precipitated proteins by SDS-PAGE (UNIT 10.1) and then immunoblotting (UNIT 10.10) with a second antibody that recognizes the putative associated protein. Controls to test specificity of interaction are crucial (see Strategic Planning). For additional background reading, the user should consult UNIT 3.8 for a theoretical discussion of immunoprecipitation and Coligan et al. (1999) for principles of antibody production and immunoassays; see UNIT 19.3 for approaches to tagging proteins; and see APPENDIX 4L and UNITS 5.6-5.8 for transformation and propagation of yeast. For an in-depth review of immunoprecipitation techniques, see Chapter 11 of Harlow and Lane (1988). For an in-depth review of coprecipitation and other approaches to detect protein-protein interactions, see Phizicky and Fields (1995). STRATEGIC PLANNING Detecting the Proteins in Question The first step is to generate reagents that detect the two proteins in the coprecipitate under nondenaturing conditions. If antibodies are available that can immunoprecipitate the proteins under nondenaturing conditions, then these can be used. Alternatively, the proteins can be differentially tagged in a variety of ways to allow their detection with commercially available antibodies or other affinity reagents. The tagged proteins are then introduced into the host organism using expression vectors. All tagged proteins must be assessed for function in vivo. A frequently used option is to add a short peptide or epitope that is recognized by a commercially available high-affinity monoclonal antibody (mAb). The epitope is typically added at the amino or carboxyl terminus, although internal positions that do not disrupt function can also be used. Two frequently used epitopes are derived from influenza hemagglutinin protein (HA) and human c-Myc and are recognized by high-affinity mAbs (12CA5 and 9E10, respectively; Kolodziej and Young, 1991). Others, such as FLAG, are Contributed by Elaine A. Elion Current Protocols in Protein Science (1999) 19.4.1-19.4.9 Copyright © 1999 by John Wiley & Sons, Inc.
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also available (BioSupplyNet Source Book, 1999). The choice of epitope may be dictated by its amino acid composition. It is often useful to insert tandem copies of the epitope in order to increase sensitivity. The number of additional tandem copies can range widely, from one (Field et al., 1988) to several (e.g., three; Tyers et al., 1993) to many (e.g., nine; Feng et al., 1998). Proteins can also be fused to small proteins or peptides that have high affinity to small molecules that can be attached to a solid support. This is a particularly valuable approach when the protein to be precipitated comigrates with immunoglobulin heavy or light chains in an SDS-polyacrylamide gel. Such alternative tagging methods include fusion to glutathione-S-transferase (to allow purification by a glutathione affinity matrix; UNIT 6.6) or maltose-binding protein (to allow purification by a maltose affinity matrix; UNIT 5.1). An excellent reference for identifying sources of commercially available antibodies and approaches to tagging proteins is the BioSupplyNet Source Book (1999). See Coligan et al. (1999) and Harlow and Lane (1988) for the generation and purification of specific antibodies. See Chapters 5, 6, and 7 of this manual for a discussion of tagging and expressing proteins. Preparing Whole-Cell Extracts The second step in a successful coprecipitation is generating whole-cell extracts that optimize the yield and activity of the proteins to be analyzed, using lysis buffer conditions that permit recognition of the proteins by the affinity matrix. The yield of total protein in a whole-cell extract is not always a reliable indicator of the relative yield and activity of specific proteins, so it is wise to verify both parameters at the onset of an experiment before proceeding with the coprecipitation. Yield and activity can be affected by a number of factors (see Chapter 10; see Harlow and Lane, 1988). Small variations in the relative amounts of salt and detergents in the lysis buffer can have large effects on yield and activity, as can the speed and efficiency of cell breakage. Both factors are particularly important for less soluble proteins that associate with macromolecular structures such as membranes or cytoskeleton. In addition, global inhibition of proteolysis through the inclusion of multiple classes of protease inhibitors may be essential. Methods for preparing whole-cell extracts from yeast (UNITS 5.6-5.8), Escherichia coli (UNIT insect cells (UNIT 5.12), and mammalian cells (UNITS 5.9 & 5.10) can be found elsewhere in this manual, and specifics will not be discussed here. In general, the lysis buffer conditions are not very different from the coprecipitation conditions. It is recommended that the investigator begin by comparing small-scale extract preparations that vary the amount of salt and nonionic detergent. As a starting point, a basic lysis buffer might contain the following components.
6.2),
Basic components. Basic components include a buffering agent (such as 50 mM Tris⋅Cl, pH 7.5), a small amount of nonionic detergent (such as 0.1% [v/v] Triton X-100), salt (such as 100 mM NaCl), a reducing agent (such as 1 mM DTT), and 10% (v/v) glycerol as stabilizer. Protease inhibitors. Protease inhibitor cocktails are described here in UNIT 5.8 and are also commercially available. A reasonable starting point would be to include 5 µg/ml each of chymostatin, pepstatin A, leupeptin, and antipain, as well as 1 mM phenylmethysulfonyl fluoride and 1 mM benzamidine. Detection of Protein-Protein Interactions by Coprecipitation
Chelating agents. EGTA (∼15 mM) is commonly included to chelate divalent metal ions that are essential for metalloprotease activity. Because EGTA also inhibits other metaldependent enzymes, it may be omitted, combined with the addition of a needed metal ion, and/or substituted with EDTA.
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Phosphatase inhibitors. If the phosphorylation state of the proteins in question is important, a mixture of phosphatase inhibitors should also be included in the lysis buffer. A starting mixture could contain 2.5 mM each meta- and ortho-vanadate, 10 mM NaF, and 10 mM β-glycerol phosphate. Simple modifications of this initial buffer include varying the amount of NaCl (from 0 to 500 mM) and of Triton X-100 (from 0% to 1%). The investigator may choose to compare different means of breaking the cells (for example, glass-bead breakage versus liquid nitrogen/grinding methods for yeast cells; UNITS 5.6-5.8).
1. Generate antibodies to protein 1 and protein 2; or differentially tag them ( ) and introduce genes encoding tagged proteins into host cell
protein 1
protein 2
2. Prepare whole-cell extract
3. Incubate extract with antibody to protein 1 4. Incubate extract with protein A–Sepharose, which binds antibody
5. Collect Sepharose beads by centrifugation
discard
P2 P1
supernatant
pellet
P2-NT P2 P1 P1-NT
protein 2 protein 1 Ig h Ig I
6. Wash pellet several times to remove proteins not bound to protein A–Sepharose. 7. Dissociate proteins from protein A – Sepharose. Separate proteins by SDS - PAGE. Immunoblot with antibody to protein 2 (P2). Reprobe with antibody to protein 1 (P1).
Figure 19.4.1 Flow chart for the coprecipitation of two proteins that have been differentially tagged and introduced into the host organism. Ig h and Ig l, immunoglobulin heavy and light chains; NT, no tag.
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Total protein concentration in the whole-cell extract is generally assayed by using the Bio-Rad protein assay and calculating protein concentration (UNIT 3.4). Extracts should be tested for the amount of each specific protein by immunoblot analysis (UNIT 10.10), analyzing 25 to 75 µg of total protein. In general, it is best to test for the presence of a second established protein (such as a housekeeping enzyme, cytoskeletal or ribosomal protein, or a previously defined component in the pathway being studied) as an internal control for normalization and as a positive control for the immunoblot. The amount of specific protein in the whole-cell extract is compared to the amount that is recovered by precipitation with an affinity matrix. Control Tests for Specificity of Interaction Controls are essential to verify that the antibodies and protein-protein interactions are specific. Proper controls are simplest to set up when the proteins are differentially tagged. In this instance, two parallel extracts are prepared from strains that contain each protein lacking the tag in the presence of the second tagged protein. An example is shown in the idealized gel in Figure 19.4.1, which includes lanes containing untagged protein 1 + tagged protein 2 and tagged protein 1 + untagged protein 2. If the antibodies are specific, untagged protein 1 will not immunoprecipitate. The presence of untagged protein 1 in the immunoprecipitate will indicate that it binds the affinity matrix nonspecifically. If the interaction between proteins 1 and 2 is specific, then tagged protein 2 will be present in the immunoprecipitate of tagged protein 1, but not in its absence. If antibodies to native proteins are used, it is necessary to compare extracts made from strains harboring deletions of the proteins in question to test for the specificity of the antibody and the interaction. However, this is obviously possible only if the deletions do not cause inviability. If deletion mutations cannot be used, a common approach is to show that the preimmune serum or an antibody not known to be specific to either of the proteins in question does not coprecipitate them in a parallel experiment. However, the latter two controls do not rule out the possibility that the antibody is precipitating the protein in question through an indirect association. It is also essential to compare the amount of coprecipitated protein with the amounts of the two proteins in question in the whole-cell extract. This allows one to determine whether apparent differences in the ability of the two proteins to coprecipitate are a secondary consequence of the relative abundance of the proteins. This control is particularly important when an interaction has been established and the investigator wishes to search for regulatory changes in association apart from changes in abundance. BASIC PROTOCOL
Detection of Protein-Protein Interactions by Coprecipitation
COPRECIPITATING PROTEINS WITH PROTEIN A/G–SEPHAROSE Once the conditions of extract preparation have been established (see Strategic Planning), the next step is to test for coprecipitation of the specific proteins. This protocol describes a standard coprecipitation procedure that uses an antibody coupled to protein A– Sepharose or protein G–Sepharose. An alternative coprecipitation method that uses GST coupled to glutathione-agarose is also provided (see Alternate Protocol). It is essential to keep all buffers and tubes cold by using an ice bath and a refrigerated centrifuge. The conditions of coprecipitation match the conditions of the lysis buffer described above. Materials Whole-cell extract (see Strategic Planning) Antibody Co-immunoprecipitation buffer (see recipe) 5 M NaCl (APPENDIX 2E)
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Protein A/G–Sepharose slurry (see recipe) 2× sample buffer for SDS-PAGE (UNIT 10.1) Test tube rotator 20-ml syringe and 18-G needle Hamilton syringe Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and immunoblotting (UNIT 10.10) 1. Prepare duplicate samples in microcentrifuge tubes on ice: 0.5 to 1 mg whole-cell extract 1 µg antibody 5 M NaCl to equalize at 100 mM NaCl Co-immunoprecipitation buffer to 0.5 ml final volume. Adjust buffer if necessary for activity of the protein in question (see Strategic Planning, discussion of chelating agents).
2. Invert tube gently several times and incubate on ice for 90 min with occasional tube inversion. It is recommended that the investigator begin with a 90-min incubation. However, this incubation step can be shortened or lengthened.
3. Microcentrifuge 10 min at maximum speed, 4°C, to pellet nonspecific aggregates. Transfer supernatant to a new microcentrifuge tube. 4. Add 50 µl of protein A– or protein G–Sepharose slurry (25 to 30 µl bead volume). Be sure to evenly suspend the slurry before distributing it to the samples. Protein A has been used more frequently for historical reasons; however, protein G binds a broader range of Ig subtypes at higher efficiency.
5. Rotate tube gently at 4°C for 30 to 60 min. Rocking is much less efficient and should be avoided.
6. Gently pellet protein A/G–Sepharose by centrifuging 30 sec at 1000 rpm in a tabletop centrifuge, 4°C. 7. Wash pellet three times with 1 ml co-immunoprecipitation buffer. For each wash, gently invert tube three times before pelleting. After each pelleting, use a 20-ml syringe with an 18-G needle to aspirate and remove supernatant. It may be possible to omit the costly protease inhibitors from the buffer at this stage, but this has not been attempted to date.
8. Aspirate as much liquid as possible from the final without touching the beads and add 25 µl of 2× sample buffer. If desired, samples containing sample buffer can be frozen up to several months at −80°C prior to SDS-PAGE. In this case the buffer should be prepared with sterile stock solutions and made with 1 mM sodium azide included.
9. Prepare for SDS-PAGE analysis by boiling for 5 min, vortexing, and microcentrifuging briefly to pellet beads. Use a Hamilton syringe to load eluates onto an SDS-polyacrylamide gel, arranging duplicate samples to allow preparation of duplicate blots. Separate by electrophoresis (UNIT 10.1). A Hamilton syringe works well to remove the eluate from the beads during loading.
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10. Immunoblot duplicate samples separately with antibodies for each of the two proteins (UNIT 10.10). Be sure to include aliquots of the whole-cell extract for comparison and as a positive control for the immunoblot. Each immunoblot can be reprobed with the antibody to the other protein. ALTERNATE PROTOCOL
COPRECIPITATING A GST FUSION PROTEIN GST fusion proteins may be coprecipitated by following the co-immunoprecipitation procedure (see Basic Protocol) with the modifications outlined below. This procedure might be used when the protein in question comigrates with immunoglobulin heavy or light chain in an SDS-PAGE gel or if the antibodies being used precipitate too many cross-reacting proteins, such as the protein being tested for association. Furthermore, it is possible to dissociate the purified GST fusion from the solid-state glutathione resin under gentle conditions through the use of imidazole. The approach is also useful in that it will increase the size of the protein sufficiently that this size increase can be used as a diagnostic feature in analyzing complexes. Additional Materials (also see Basic Protocol) Glutathione-agarose or glutathione-Sepharose slurry (see recipe) 1. Prepare duplicate samples as described for protein A/G–Sepharose (see Basic Protocol, step 1), omitting antibody. 2. Microcentrifuge 10 min at maximum speed, 4°C, to pellet nonspecific aggregates. Transfer supernatant to new microcentrifuge tube. 3. Add 30 µl glutathione-agarose or glutathione-Sepharose slurry (25- to 30-µl bead volume). Be sure to evenly suspend the slurry before distributing it to the samples. 4. Rotate the sample, pellet and wash glutathione-agarose/Sepharose, and perform SDS-PAGE and immunoblot analysis (see Basic Protocol, steps 5 to 10). REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Co-immunoprecipitation buffer 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 15 mM EGTA 100 mM NaCl 0.1% (w/v) Triton X-100 Store at 4°C Immediately before use add: 1× protease inhibitor mix (see recipe) 1 mM dithiothreitol (DTT) 1 mM phenylmethylsulfonyl fluoride (PMSF; from fresh 250 mM solution in 95% ethanol) If necessary for activity of the protein of interest, a divalent cation may need to be included in the buffer and EGTA omitted (as for extraction buffer; see Strategic Planning, discussion of chelating agents). Detection of Protein-Protein Interactions by Coprecipitation
The protease inhibitor mix, PMSF, and DTT should be added fresh at the time of experimentation. The mixture without those components can be stored for months at 4°C with the addition of 1 mM sodium azide. PMSF is labile in aqueous buffer and should be added at the last minute.
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Glutathione-agarose or glutathione-Sepharose slurry Swell 1.5 g glutathione-agarose or glutathione-Sepharose beads (e.g., Pierce, Sigma) in 30 ml of 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E), for 1 to 2 hr on ice. Pellet beads by gravity or very gentle centrifugation (1 min at 1000 rpm in a tabletop centrifuge) and then wash four times with co-immunoprecipitation buffer (see recipe) that lacks protease inhibitor mix and contains 1 mM sodium azide. Resuspend beads in 15 ml of this buffer to yield a final slurry concentration of ∼100 mg/ml. Store at 4°C (stable for months). This recipe can be scaled up or down.
Protease inhibitor mix, 1000× Dissolve in DMSO: 5 mg/ml chymostatin 5 mg/ml pepstatin A 5 mg/ml leupeptin 5 mg/ml antipain Store in aliquots up to 1 year at −20°C Protein A/G–Sepharose slurry Swell 1.5 g protein A– or protein G–Sepharose beads (e.g., Pierce, Sigma) in 30 ml of 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E), for 1 to 2 hr on ice. Pellet beads by gravity or very gentle centrifugation (1 min at 1000 rpm in a tabletop centrifuge) and then wash four times with co-immunoprecipitation buffer (see recipe) that lacks protease inhibitor mix and contains 1 mM sodium azide. Resuspend beads in 15 ml of this buffer to yield a final slurry concentration of ∼100 mg/ml. Store at 4°C (stable for months). This recipe can be scaled up or down.
COMMENTARY Background Information Coprecipitation is a powerful and simple approach to test for a physical interaction between proteins. There are many reasons to incorporate coprecipitation into a study. First, as a form of protein affinity chromatography, the method may be sensitive enough to detect weak associations that do not withstand the rigors of standard purification methods involving substantial dilution of the initial cell extract. Second, coprecipitation tests for associations between proteins within the milieu of a whole-cell extract, where the proteins are present at native concentration in a complex mixture of other cellular components. This feature makes it an important partner to two-hybrid methods and direct tests of interactions using purified proteins, because it provides a way to verify that a positive interaction reflects a true in vivo association. For example, nonphysiological interactions can be detected when purified proteins are present at too elevated a concentration. A falsely positive interaction between two proteins can also arise in a two-hybrid test when protein domains are inappropriately exposed
due to altered folding. In addition, not all proteins are amenable to two-hybrid analysis; a negative result may mask a true association. Nevertheless, a word of caution is in order. The ability to coprecipitate two proteins from a cellular extract is not proof that a particular interaction normally takes place in vivo. Additional experiments are needed to argue that a given interaction is not the result of mixing cell contents during extract preparation. Such evidence could include colocalization of the proteins or demonstration of functional relatedness. When performing coprecipitation, it is important to precipitate from both directions (i.e., individually precipitating protein 1 and protein 2, and testing for the presence of protein 2 and protein 1, respectively). This is important in that it can provide further verification of an interaction between the proteins. It is also important because it is possible the interaction will only be detected in one direction. An inability to detect an interaction in one direction could be due to a variety of factors including obstruction of an interaction by the binding of
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the antibody or other affinity agent, or differences in pool size representation of each protein. For example, protein 1 may bind to many proteins besides protein 2, while most of protein 2 binds to protein 1. In this scenario, it would be anticipated that detection of their association will be most efficient when protein 2 is precipitated.
Critical Parameters and Troubleshooting
Detection of Protein-Protein Interactions by Coprecipitation
It is important to vary conditions of both the extract preparation and the coprecipitation to determine what is optimal. When starting from scratch, it is most prudent to use a range of lysis and precipitation conditions from less to more stringent in terms of the amount of salt and nonionic detergent. When no interaction is detected, it is worthwhile to use less stringent conditions (reduced salt with little or no nonionic detergent). In addition, it may be necessary to avoid any dilution of the whole-cell extract. This can be done by adding the protein A/G–Sepharose directly to the extract after an initial clarification centrifugation and by using smaller wash volumes. Depending on the strength and nature of the interaction, the precipitation can be done in the presence of a mixture of detergents that includes ionic detergents (for example, 1% Triton X-100, 0.5% deoxycholate, 0.1% SDS), similar to that described in RIPA buffer (UNIT 3.8). In addition, it may be necessary to increase the expression levels of the proteins in question to be able to readily detect them by coprecipitation. A range of expression levels is recommended, because a level that is too high can lead to unregulated interactions (Feng et al., 1998). Alternatively, one can scale up the coprecipitation and use more than 0.5 to 1 mg of whole-cell extract (Feng et al., 1998). Here, the limiting factor is the concentration of the extracts, which must be high enough to allow the volume of the coprecipitation mixture to remain low. Larger-scale extract preparations may be necessary to generate more concentrated extracts. Finally, in cases of failure due to low abundance of the proteins in the host organism, one can overexpress a tagged version of one of the two proteins in the same or another host (such as E. coli), concentrate this protein by preimmobilization on an appropriate affinity matrix, and then incubate the affixed protein with extracts from the host organism. The most important objective in these experiments is to generate as great a signal-tonoise ratio as possible and avoid problems of
background. A variety of parameters can be changed to enhance the co-immunoprecipitation. Optimization of the precipitating antibody is one possibility. Protein A–Sepharose and protein G–Sepharose should give results comparable to anti-Ig serum. However, direct coupling of the antibody to Sepharose may lead to lower background and more quantitative precipitation. In addition, varying the ratio of antibody to whole-cell extract and the total amount of whole-cell extract is strongly suggested to determine the optimal amount of antibody that gives the most precipitation with the least amount of background. Affinity purification of the antibody may be necessary if the antibody immunoprecipitates additional crossreacting proteins. Additional approaches can be taken to minimize background. First, better clarification of the cell extract can be achieved by precentrifugation at 100,000 × g. These extracts can be directly used for coprecipitation without an intervening freezing step, which can increase the amount of protein precipitation. Second, both the lysis buffer and the coprecipitation buffer can be supplemented with 1% BSA to reduce nonspecific binding to the affinity matrix. Third, the whole-cell extract can be preincubated with protein A/G–Sepharose to remove nonspecific proteins that bind to the solid support. Fourth, the amount of salt and detergent can be increased in both the coprecipitation and the washes to reduce nonspecific binding. Fifth, increasing the number of washes may also help, although it may reduce the amount of specific protein that remains associated. Sixth, the expression levels of the proteins in question can be increased to generate a stronger signal that is above background binding. Alternatively, it may be possible to produce a whole-cell extract that is enriched for the proteins in question (e.g., by preparing a nuclear extract if the proteins are known to be in the nucleus). In instances where one of the proteins binds nonspecifically to Sepharose, the substitution of an agarose-based affinity matrix may help solve the problem. Finally, it may be necessary to generate a different set of reagents to precipitate the proteins in question (i.e., different antibodies and/or protein tag).
Anticipated Results Provided suitable antibodies are available to the proteins in question and the physical interaction is stable under the coprecipitation conditions, it should be possible to detect an interaction between two proteins.
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Time Considerations Once the extracts are prepared, coprecipitation can be done within 3 to 4 hr, yielding samples ready to load on a gel for SDS-PAGE and immunoblot analysis.
Literature Cited BioSupplyNet Source Book. 1999. BioSupplyNet, Plainview, N.Y., and Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Coligan, J.E., Kruisbeck, A.M., Margulies, D.H., Shevach, E.H., and Strober, W. 1999. Current Protocols in Immunology. John Wiley & Sons, New York. Feng, Y., Song, L.-Y., Kincaid, E., Mahanty, S.K., and Elion, E.A. 1998. Functional binding between Gβ and the LIM domain of Ste5 is required to activate the MEKK Ste11. Curr. Biol. 8:267-278. Field, J., Nikawa, J., Broek, D., MacDonald, B., Rodgers, L., Wilson, I.A., Lerner, R.A., and Wigler, M. 1988. Purification of RAS-responsive adenylyl cyclase complex from Saccharomyces cerevisiae by use of an epitope addition method. Mol. Cell. Biol. 8:2159-2165. Harlow, E. and Lane, D. 1988. Antibodies: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
Kolodziej, P.A. and Young, R.A. 1991. Epitope tagging and protein surveillance. Methods Enzymol. 194:508-519. Phizicky, E.M. and Fields, S. 1995. Protein-protein interactions: Methods for detection and analysis. Microbiol. Rev. 59:94-123. Tyers, M., Tokiwa, G., and Futcher, A.B. 1993. Comparison of the Saccharomyces cerevisiae G1 cyclins: Cln3 may be an upstream activator of Cln1, Cln2, and other cyclins. EMBO J. 11:17731784.
Key References BioSupplyNet Source Book. 1999. See above. Published yearly. Instant access is available on the WWW, at http://www.biosupplynet.com; hard copy may be requested by fax at (609) 786-4415. Information on its contents may be obtained by telephone at (516) 349-5595, fax at (516) 349-5598, or email at [email protected]. Phizicky and Fields, 1995. See above. General discussion of methodologies for detecting protein-protein interactions as well as their merits and drawbacks.
Contributed by Elaine A. Elion Harvard Medical School Boston, Massachusetts
Identification of Protein Interactions
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Imaging Protein-Protein Interactions by Fluorescence Resonance Energy Transfer (FRET) Microscopy
UNIT 19.5
Specific protein-protein interactions, whether induced by covalent protein modifications or not, are generally considered to mediate cellular signaling and function. Detection of these processes has long been restricted to bulk biochemical methods such as immunoprecipitation (UNIT 9.8) and immunoblotting (UNIT 10.10). These approaches have proven invaluable, e.g., in uncovering the major signal transduction pathways by delineating the hierarchy of protein-protein interactions and kinase-substrate relationships in the different phosphorylation cascades. However, immunoblotting techniques lack spatial information and the interactions that are detected depend on the stability of the complex during the experimental conditions that exist outside the cell in homogenates. Detection of proteins using immunofluorescence provides spatial information on the micrometer scale, and it is therefore not possible to infer protein-protein interactions. Limited information on the phosphorylation status of proteins can be obtained by the application of phospho-specific antibodies against specific phosphorylated residues in a given protein (see UNIT 13.4). The use of these antibodies is, however, restricted by availability and specificity. Only a small number of antibodies against phosphoproteins exist and phosphotyrosine residues have proven to produce the most specific antibodies thus far, leaving a large number of modifications undetectable by microscopy. This unit describes the preparation and execution of a typical fluorescence resonance energy transfer (FRET) experiment. The following procedures will specifically refer to the use of a GFP-tagged protein as donor and Cy3-labeled antibodies as acceptor. As an example, covalent modification by tyrosine phosphorylation of the epidermal growth factor receptor (EGFR) is determined by FRET between C-terminal GFP-tagged EGFR and Cy3-labeled anti-phosphotyrosine antibodies (Wouters and Bastiaens, 1999). FRET is measured by release of donor quenching through acceptor photobleaching. An advantage of this method is the convenient setup; it does not require specialized equipment but can be performed using common microscopy equipment, preferably a confocal laser scanning microscope (Bastiaens and Jovin, 1996; Bastiaens et al., 1996; Wouters et al., 1998). The method is quantitative, and by simple relationships, a FRET efficiency is obtained. The sample preparation described in this unit is identical for other FRET microscopy techniques or other donor-acceptor pairs. As an illustration, the result of a more advanced FRET measurement using fluorescence lifetime imaging microscopy (FLIM), which is based on the change in GFP fluorescence lifetime (see Background Information), is also included (Bastiaens and Squire, 1999; Ng et al., 1999; Wouters and Bastiaens, 1999). This unit describes FRET microscopy based on release of quenched donor fluorescence after acceptor photobleaching (see Basic Protocol), microinjection of reagents into the nucleus or cytosol (see Support Protocol 1), and labeling antibodies with Cy3 (see Support Protocol 2). FRET MICROSCOPY OF FIXED CELLS A number of quantitative FRET microscopy techniques are available at this time. A technique based on the release of quenched donor fluorescence after acceptor photobleaching will be described in this unit. This technique requires a pixel-by-pixel reference image to be created by photobleaching of the acceptor through continuous illumination at the absorption maximum of the acceptor. The time scale of the phoContributed by Fred S. Wouters and Philippe I.H. Bastiaens Current Protocols in Protein Science (2001) 19.5.1-19.5.15 Copyright © 2001 by John Wiley & Sons, Inc.
BASIC PROTOCOL
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tobleaching process is on the order of minutes. In a lifetime-based FRET approach, the photobleaching step is used at the end of a time-lapse sequence of measurements to introduce an intracellular reference in live cells. In the donor intensity approach (unquenching), described here in detail, the time scale of photobleaching restricts the technique to fixed cells but has the advantages of being relatively simple and quantitative. Materials Cells of interest Plasmid for GFP-tagged protein Transfection reagent (e.g., Fugene 5 from Boehringer Mannheim, Lipofectin from Life Technologies, Effectene from Qiagen, or Superfect from Qiagen) Serum-free medium Low-background fluorescence CO2-independent medium (Life Technologies, or see recipe) Phosphate-buffered saline (PBS; see recipe), pH 7.4 4% (w/v) formaldehyde fixative solution (see recipe) Quench solution: 50 mM Tris⋅Cl (pH 8.0)/100 mM NaCl 0.1% (v/v) Triton X-100 in PBS Antibody (e.g., PY72 monoclonal anti-phosphotyrosine antibody) labeled with Cy3 (see Support Protocol 2) 1% (w/v) bovine serum albumin (BSA, fraction V) in PBS Mowiol mounting medium (see recipe) 6- and 12-well tissue culture plates Coverslips Microscope slides Confocal laser scanning microscope (e.g., Zeiss LSM 510), equipped with argon (488 nm) and He/Ne (543 nm) lasers selected by the HFT 488/543 double dichroic filter, GFP fluorescence selected by the NFT 545 dichroic and BP 505-530 emission filter, and Cy3 fluorescence selected by the LP560 emission filter Imaging software package (e.g., NIH-image or IPLab Spectrum from Scanalytics) Additional reagents and equipment for transfection of mammalian cells (APPENDIX 3C) NOTE: All solutions and equipment coming into contact with cells must be sterile, and aseptic technique should be used accordingly. NOTE: All incubations are performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4. Prepare cells 1. Seed the cells onto culture dishes containing coverslips. For live-cell FRET experiments, seed the cells onto glass-bottom MatTek culture dishes. The cells should be adherent and grow in a monolayer.
2. Transfect cells with the plasmid for GFP-tagged protein of interest using calcium phosphate precipitation, lipofection, or the activated dendrimer reagent, Superfect.
Imaging Protein-Protein Interactions by FRET Microscopy
For cells that are difficult to transfect, due to resistance to transfection (e.g., primary cell lines), or due to cytotoxicity of the transfection reagent, the DNA for the GFP-tagged protein can be introduced by nuclear microinjection (see Support Protocol 1). The latter method is also particularly useful when the expression levels of the protein of interest have to be controlled accurately.
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3. Grow the cells, typically 1 to 2 days post transfection, until expression levels of the GFP-tagged protein are high enough to detect with fluorescence microscopy. When needed, starve the cells overnight with serum-free medium to make them semi-quiescent. For cells that undergo apoptosis upon serum deprivation, lower the serum concentration in the medium to 0.5%.
4. On the day of the experiment, transfer the coverslips to a 12-well tissue culture plate. Subject the cells to the desired experimental conditions (e.g., growth factor/hormone stimulation, or incubation with drugs or inhibitors).
Fix cells 5. Wash the cells with ice-cold PBS, aspirate, and add 1 ml 4% formaldehyde fixative solution. Allow cells to fix at room temperature for 10 min. The common alternative fixation protocol, where cells are incubated in −20°C methanol for 5 min, is not advised. Strictly speaking, this treatment does not fix the tissue, but precipitates the cellular proteins. Protein-protein interactions are therefore likely to be affected.
6. Quench excess fixative with quench solution by briefly washing the cells, then changing to fresh quench solution and incubating 5 min at room temperature. In a first short wash, the excess fixative is removed. In a second 5-min incubation, the remaining aldehyde groups are allowed to react with the primary amino group on the Tris molecule. Alternative, equally effective, quench solutions are 0.1 M hydroxylamine or 0.1 M glycine.
7. Permeabilize cellular membranes by incubating 5 min with 0.1% Triton X-100 to allow penetration of acceptor-labeled molecules into fixed cells. In this treatment, a compromise is made between optimal morphology and permeabilization. A milder detergent treatment with 0.1% saponin in PBS or a harsher permeabilization with −20°C methanol for 5 min can be considered if morphology or permeabilization, respectively, is compromised.
8. Wash the cells with PBS to remove the permeabilization solution. Add antibody 9. Dilute Cy3-labeled antibodies appropriately in 1% BSA/PBS. Incubate the coverslips with appropriately diluted Cy3-labeled antibodies for 1 hr, at room temperature. Typical concentrations are 0.1 to 10 ìg/ml, but it is advised that a titration series be performed to determine the antibody concentration where epitope binding saturates. To minimize the amount of antibody needed, press a sheet of Parafilm to the bench with some water, place 25-ìl drops of antibody solution on the Parafilm, place the coverslips cells-down on these drops, and incubate 1 hr. To facilitate handling of the coverslips at the end of the 1-hr incubation, place a pipet to the edge of a coverslip and add ∼100 ìl PBS underneath it. This will lift the coverslip so that it can be easily picked up with jeweler’s forceps.
10. Transfer the coverslips to 6-well tissue culture plates and wash four times, each time with 3 ml PBS, to remove excess antibody. 11. Blot off excess PBS with tissue and mount the coverslips on slides with ∼10 µl Mowiol mounting solution. Allow Mowiol to harden overnight at 4°C before imaging. Identification of Protein Interactions
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If necessary, cells can be imaged after a short drying period of 1 hr if the coverslip is attached to the slide by painting the edge of the coverslip with molten agarose or rubber cement at four points. Do not use nail polish since this has been shown to quench GFP fluorescence.
Examine cells and calculate results 12. View the specimen on a confocal microscope using a 63× or 100× oil-immersion objective. Acquire an image in the GFP channel (excitation, 488 nm; emission, NFT 545, BP505-530). Either take an image of the entire field of vision or select a region of interest containing the cell that is to be imaged. GFP fluorescence is selected by the NFT 545 dichroic and BP 505-530 emission filter. Do not use the full dynamic range of the detector, since unquenching will result in added fluorescence. Depending on the make of the confocal microscope, make sure that a second image can be made at exactly the same location. Do not adjust the settings (i.e., pinhole size, contrast, brightness, laser power, or averaging) for the GFP channel. This acquisition provides the FRET-quenched donor image (FDA), since the acceptor is present throughout the cell, causing FRET with the GFP donor. Minimize GFP photobleaching at this point by limiting the illumination.
13. Change to the Cy3 channel (excitation, 543 nm; emission, LP560) and take an image of the cell, minimizing Cy3 photobleaching. Select a portion of the cell in which the FRET efficiency is to be determined. Cy3 fluorescence is selected by the LP560 emission filter. This region is where the acceptor will be photobleached, thereby revealing the unquenched donor intensities enabling FRET calculation. Consequently, any portion of the cell where the acceptor is not photobleached serves as control [i.e., 1 − FDA/FDA = 0]. This area in the same cell will provide an essential control to judge the effects of photobleaching GFP, lateral movement, or focal mismatch between the two consecutive donor images.
14. Photobleach the selected acceptor region by repeated scanning with the 543-nm He/Ne laser line at full power. Follow the progress of photobleaching by monitoring the intensities of the respective images (emission: NFT 545, BP505-530). Continue until there is no more discernible Cy3 intensity. Depending on the staining intensity and the area that is bleached, this will typically take 1 to 20 min. When selecting the photobleaching region on the Zeiss LSM 510 confocal microscope, crop the acceptor image to this region (and note the zoom settings to return to the original image after bleaching) instead of selecting a region of interest (ROI). In the latter option, the entire area will be scanned but the laser will only switch on in the ROI. Consequently, the duty cycle is low and bleaching will take considerably longer.
15. Return to the GFP channel and make the second acquisition, using identical settings and location of the prebleached image. This provides the FD reference in the region where the acceptor was photobleached.
Imaging Protein-Protein Interactions by FRET Microscopy
16. To calculate the FRET efficiency in the bleached area, use an appropriate imaging software package. Subtract the prebleach donor image from the post-bleach donor image (i.e., FD − FDA). Divide this image by the postbleach donor image: (FD − FDA)/FD; this is identical to 1 − (FDA/FD) = E, the image of FRET efficiencies. A software package capable of performing the abovementioned image processing is NIH Image. This package is freely available from http://rsb.info.nih.gov/nih-image/ and ver-
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sions are available for Apple Macintosh, Windows (Scion image) and even a Java version is available that runs on any platform (Image J). A more versatile commercial package is IPLab Spectrum. Image arithmetic for calculating the FRET efficiency should be performed on a region of interest obtained by thresholding the images in order to prevent the amplification of background noise. Therefore, thresholds should be chosen in such a way that the background is omitted in the calculation. Accuracy can be improved by subtracting the average background intensity from the images before calculating the FRET efficiency.
NUCLEAR AND CYTOSOLIC MICROINJECTION The following is a generic protocol for introduction of DNA or labeled proteins into a cell. DNA is microinjected directly into the nucleus to achieve controlled and high expression of the protein of interest. Labeled proteins are injected into the cytosolic region next to the nucleus since the cell is at its highest here, facilitating injection.
SUPPORT PROTOCOL 1
Materials Cells of interest, cultured in MatTek glass-bottom 35-mm dishes (MatTek) DNA (e.g., human EGFR cDNA in the Clontech pEGFP-N1 expression vector; APPENDICES 4C & 4D) HPLC-grade water Millex-GV4 0.22-µm filtration unit GELloader tips (Eppendorf) Needles for microinjection (e.g., Femtotip from Eppendorf) Microinjector (e.g., Eppendorf model 5244) Micromanipulator (e.g., Eppendorf model 5170) Inverted microscope with 10× and 40× air objectives Additional reagents and equipment for preparation of DNA (APPENDIX 4C) 1. Prepare DNA, to be microinjected, of the highest possible quality. In the authors’ experience, double cesium chloride–banded DNA (Wilson, 1994) and DNA purified by Qiagen ion-exchange resin (Qiaprep midi/maxiprep columns; Budelier and Schorr, 1998) perform equally well. Cy3-labeled protein (e.g., antibody against protein of interest; see Support Protocol 2) may also be microinjected.
2a. For optimal expression a few hours after microinjection: Dilute DNA in HPLC-grade water to 100 µg/ml. 2b. For expression overnight: Dilute DNA in HPLC-grade water to 1 µg/ml. 3. Clear the DNA solution to prevent blocking of the glass needle during microinjection as follows. Place a 0.22 µm Millex filtration unit in a 0.5-ml microcentrifuge tube and place the entire unit in a 1.5-ml microcentrifugation tube to enable centrifugation. Filter 10 µl of the DNA solution by microcentrifugating 1 min at maximum speed, room temperature. Since these membranes have low-protein-binding characteristics, they can also be used for clearing Cy3-labeled proteins (see Support Protocol 2). The recovery of Cy3-labeled antibodies when cleared this way is also generally very high. Alternatively, microcentrifuge the Cy3-labeled protein for 20 min at maximum speed. Sacrifice a small amount of solution to prevent disturbing the pellet of aggregated protein. Identification of Protein Interactions
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4. Load 2 µl of DNA solution (or Cy3-protein solution) using GELloader tips into the capillary glass needle of the microinjector. Commercially available needles (Femtotip from Eppendorf) can be used. These needles fit directly into the needle holder of the Eppendorf microinjection device. Carefully remove the pipet tip that protects the needle. In the authors’ experience, this is most easily performed by holding the needle pointing downwards and loosening the tip by rotation until it falls to the ground. If access is available to a needle-pulling device, make sure that the diameter of the needle opening is ∼0.25 ìm. The diameter of the needle opening can be estimated using a simple syringe-operated micropipet bubble meter (Clark Electronic Instruments) by measuring the air pressure required to expel air bubbles from the pipet into a liquid (Mittman et al., 1987).
5. On an inverted microscope, microinject the DNA solution (or Cy3-protein solution) into the nucleus (or perinuclear cytosol) of cells grown in MatTek culture dishes. Typical settings are: 0.3 sec, 150 to 400 hPa injection pressure with 20-hPa back-pressure to prevent medium from entering the needle. The injection pressure may be varied according to the needle opening and cell type. No major movement of the nucleus (or cell organelles) should be observed. A visual indication for excessive pressure is the separation of the nucleus from the surrounding cellular material (i.e., light ring around the nucleus) and leakage into the cytosol, visible by movement of the cellular organelles. Restrict the microinjection procedure to a maximum of 10 min. In the authors’ laboratory, microinjection is performed at room temperature in normal CO2-dependent medium. After 10 min, the medium starts to acidify significantly (i.e., purple medium). CO2-independent (or HEPES-buffered) media can be used for longer periods. SUPPORT PROTOCOL 2
Imaging Protein-Protein Interactions by FRET Microscopy
PROTEIN LABELING WITH Cy3 Proteins are labeled on unprotonated free amino groups (i.e., α-amino terminus or ε-amino groups on lysine side chains) by the succinimide esters of the fluorescent sulfoindocyanine (Cy) dyes (also see Haugland, 2000). Alkaline labeling conditions ensure deprotonation of amino groups. Cy-dyes are water soluble and have high extinction coefficients, making them particularly useful for sensitive detection of proteins with minimal disturbance of protein function. The following protocol describes the labeling of antibodies with Cy3, a suitable donor for GFP in a FRET experiment, and also explains how to remove stabilizing compounds (e.g., gelatin, BSA), which are often added to prolong shelf life, as these contain amino groups that would compete with the labeling reaction. Materials Antibody (PY72 monoclonal anti-phosphotyrosine antibody) 1 M Tris⋅Cl, pH 8.0 (APPENDIX 2E) 10 mM and 100 mM Bicine/NaOH, pH 8.0 100 ml citric acid/NaOH, pH 2.8 1 M Bicine/NaOH, pH 9.0 1 M NaCl (APPENDIX 2E) Labeling buffer: 100 mM Bicine/NaOH (pH 8.0)/100 mM NaCl Cy3.29–OSu monofunctional sulfoindocyanine succinimide ester (Amersham Pharmacia Biotech) Dimethylformamide (DMF) dried by addition of 10 to 20 mesh 3-Å pore diameter molecular sieve dehydrate (Fluka) 1-ml Protein G HiTrap columns (Amersham Pharmacia Biotech) Centricon YM30 concentrators (Amicon)
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Biogel P6DG Econopac prepacked size-exclusion columns (5.5 × 1.5–cm, ∼10 ml; Bio-Rad) 1-ml and 10-ml syringes with HPLC Luer-Lok fitted tubing Additional reagents and equipment for spectrophotometric protein determination (UNIT 3.1) and SDS-PAGE (UNIT 10.1) Prepare antibody solution 1. Resuspend antibody to 1 mg/ml in PBS. Excess stabilizing agents containing free amino groups (e.g., BSA and gelatin) in commercially available antibody preparations compete for the labeling reagent and have to be removed by Protein A or Protein G affinity chromatography (UNIT 9.8). If the protein solution to be labeled is free of additional amino groups, proceed directly to step 10. A number of suppliers of antibodies, e.g., Transduction Laboratories and New England Biolabs, can provide their products free of these compounds. Request that the antibodies be provided at 1 mg/ml concentration in PBS.
2. Prepare a syringe-operated 1-ml protein G HiTrap column. Equilibrate column with 10 ml PBS at a maximum flow of 4 ml/min. All fluid handling is performed manually using appropriately sized syringes. These columns are easy to use and result in minimal loss of protein. A number of subclasses of IgG molecules do not bind to protein A. When using protein G or A chromatography, make sure that the antibodies are compatible (see Bonifacino and Dell’Angelica, 1998).
3. Add 0.1 vol 1 M Tris⋅Cl, pH 8.0, to the antibody solution. Commercial antibody solution is typically 0.5 ml at 0.1 mg/ml IgG.
4. Load antibody solution onto the column and wash column with 10 ml of 100 mM Bicine/NaOH, pH 8.0. Collect run-through. 5. Wash column with 10 ml of 10 mM Bicine/NaOH, pH 8.0. Collect run-through. 6. Elute column with 5 ml of 100 mM citric acid/NaOH, pH 2.8, and collect 0.5-ml fractions (i.e., 8 drops) in 1.5-ml microcentrifuge tubes containing 100 µl of 1 M Bicine/NaOH, pH 9.0, to neutralize the pH. Mix immediately and store on ice. 7. Determine the A280 of the eluted fractions using a spectrophotometer (UNIT 3.1) and pool the fractions that contain protein. Under the given conditions, the protein will typically elute in the first four fractions.
8. Add 0.1 vol of 1 M NaCl. Concentrate the solution in a YM30 Centricon to ∼200 µl by centrifuging at 5000 × g, 4°C. 9. Redilute to 2 ml with labeling buffer and repeat the concentration and redilution steps (steps 8 and 9). 10. Concentrate to ∼50 to 100 µl as in step 8 and collect the concentrated protein solution. When labeling proteins from solutions containing relatively high concentrations (i.e., millimolar) of compounds that contain free amino groups (e.g., Tris, glycine, glutathione), repeat the concentration-redilution cycle more often. Each cycle dilutes the compound ∼10 fold. Allow a maximum of 10% of contaminating primary amino groups, compared to the protein to be labeled. Identification of Protein Interactions
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For example, the protein concentration in a 0.5 mg/ml antibody solution is 3.3 ìM, assuming a molecular mass of 150 kDa. If this solution contains 50 mM Tris buffer (a 1.5 × 104 fold excess of free amino groups), a 1.5 × 105-fold dilution, corresponding to 5.2 concentration cycles, is needed to reach the 10% contamination level. When labeling proteins other than antibodies, a Bicine concentration of 50 mM is recommended in the labeling buffer. Additional compounds that are needed to maintain the function of the protein to be labeled should be included. At this point, no compounds containing free amino groups should be added. High concentrations of reducing agents are also known to inhibit the labeling reaction. It is recommended that these be included in the chromatography step following labeling to prevent these compounds from interfering with the labeling reaction. Choose the cut-off value of the Centricon carefully to ensure retention of the protein of interest.
Prepare dye 11. Reconstitute Cy3.29–OSu in 20 µl dry DMF to give a ∼10 mM Cy3 solution. Determine the exact concentration by measuring the absorption of a 104-fold diluted solution in PBS. From the ε550 of 150 mM−1cm−1, calculate the concentration. Cy3.29–OSu is supplied as a desiccated pellet in microcentrifuge tubes. DMF is dried by addition of hygroscopic beads to the container. The ε650 of Cy5 is 250 mM−1cm−1.
Perform labeling reaction 12. Determine the protein concentration of the antibody (or protein) solution to be labeled based on A280 reading (UNIT 3.1). Antibody concentration at A280 = 1.0 is typically 1 mg/ml. At low protein concentrations (i.e., <0.1 mg/ml), or for smaller proteins (where the A280 = 1.0 = 1 mg/ml protein relation does not hold true), the protein concentration can be determined using the Bio-Rad Coomassie-based protein determination assay. Antibodies are known to give a lower apparent concentration when BSA is used as a standard in this assay. Multiply the concentration found by a factor of two or use an IgG standard.
13. Slowly add a 10- to 20-fold molar excess of dye to the protein solution while simultaneously stirring the solution with the pipet tip. The added volume of Cy3/DMF should not exceed 10% of the total volume, to prevent protein denaturation by DMF.
14. Incubate 30 min at room temperature, shaking the tube gently every 10 min. Remove unconjugated dye 15. Remove the unconjugated dye by size-exclusion chromatography on a Bio-Rad P6DG Econopac (6-kDa exclusion size) column. Equilibrate the column with 30 ml PBS (or a buffer that is specifically formulated for the protein to be labeled). 16. Load the labeling reaction mixture onto the column and wash with a small amount (i.e., ∼200 µl) of PBS (or equilibration buffer). Discard the first 2.5 ml (void volume ∼3.3 ml).
Imaging Protein-Protein Interactions by FRET Microscopy
17. Collect the colored protein fraction (or 2 ml of eluate if the staining is too weak to be visible) in a YM30 Centricon concentrator and concentrate to approximately the volume of the labeling reaction.
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Determine labeling ratio 18. Estimate the labeling ratio by either of the following formulas: A554 × M/[A280 − (0.05 × A554) × 150] for Cy3 A650 × M/[A280 − (0.05 × A650) × 250] for Cy5 where Ax is the absorption at wavelength x and M is the molecular weight of the protein in kDa. The extinction coefficients (in units of mM−1cm−1) 150 and 250 are for Cy3 and Cy5, respectively. These formulas assume the A280 = 1.0 = 1 mg/ml relationship as found for larger proteins (i.e., >50 kDa) and correct for the 5% absorption of Cy3 at 280 nm. For smaller proteins or lower protein concentrations (i.e., <0.1 mg/ml) that cannot be reliably estimated from the A280, determine the protein concentration by the Bio-Rad protein assay and estimate the labeling ratio from: A554 × M/(P × 150) for Cy3 A650 × M/(P × 250) for Cy5 where P is the protein concentration in mg/ml as determined by Bio-Rad assay. As previously mentioned, IgG concentration obtained with BSA as standard has to be multiplied by 2. Using mass spectrometry, the authors have observed that ∼50% of the Cy3 molecules in the preparation provided by Amersham are reactive. The molar excess is thus 7.5-fold for a 15-fold excess and, the typical labeling ratio of 2 to 3 that is reached under these conditions is therefore reasonably efficient.
19. Verify covalent labeling of the antibody by SDS-PAGE of the labeled product (UNIT 10.1). Use transillumination with a UV source (302 nm) to visualize labeled protein. Ensure that there is no fluorescence at the running front of the gel; this indicates contamination with unconjugated dye. Where applicable, verify the specific activity of the protein and compare to the specific activity before labeling to ensure that labeling did not interfere with the biological function of the protein.
REAGENTS AND SOLUTIONS Use deionized or distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Formaldehyde fixative, 4% (w/v) Dissolve 4 g of paraformaldehyde (Sigma) in 50 ml water. Add 1 ml 1 M NaOH solution and stir gently on a heating block (>60°C) until the paraformaldehyde is dissolved. Add 10 ml of 10× PBS (see recipe) and allow to cool to room temperature. Adjust the pH to 7.4 using 1 M HCl (∼1 ml). Adjust to 100 ml with water and filter through a Millipore 0.45-µM filter using a syringe to remove traces of undissolved paraformaldehyde. Store up to several months at −20°C. Low-background-fluorescence CO2-independent medium Adjust the formulation of the standard medium by omitting the pH indicator phenol red, the antibiotics penicillin and streptomycin, folic acid, and riboflavin. Before use, supplement the medium with 50 mM HEPES/NaOH, pH 7.4. Store 1 to 2 months at 4°C.
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Mowiol mounting medium Mix 6 ml glycerol, 2.4 g Mowiol 4-88 (Calbiochem), and 6 ml water. Shake for 2 hr. Add 12 ml of 200 mM Tris⋅Cl, pH 8.5 (APPENDIX 2E), and incubate at 50°C with occasional mixing until the Mowiol dissolves (i.e., ∼3 hr). Filter through 0.45-µM Millipore filtration unit and store in aliquots up to several weeks at 4°C or up to several months at −20°C. Phosphate-buffered saline, 10× 68 g NaCl 18.8 g Na2HPO4 2 g KH2PO4 H2O to 1 liter COMMENTARY Background Information
Imaging Protein-Protein Interactions by FRET Microscopy
Fluorescence resonance energy transfer (FRET) is a photophysical process that can be exploited to obtain information on protein-protein interactions and protein modification in addition to location (Clegg, 1996). Its usefulness lies in the ability to sense the presence of acceptor fluorophores in the extreme vicinity of a donor fluorophore with a maximum separation distance that is in the order of magnitude of single protein molecules. FRET is a radiationless process whereby an excited donor fluorophore transfers energy to an acceptor by dipole-dipole coupling (Förster, 1948). Due to the 6th-order distance dependence of the FRET efficiency, two discrete states of the fluorophores can be discriminated: exhibition of efficient FRET when donor and acceptor are in close proximity or no occurrence of FRET due to distance. The transfer of energy (FRET) from an excited donor fluorophore to a nearby acceptor fluorophore has a number of consequences with respect to the fluorescent properties of both fluorophores and this can be exploited to measure the efficiency of this process. In the methods presented in this unit, FRET is determined by measuring the donor fluorescence emission exclusively (Bastiaens and Squire, 1999). The acceptor fluorophore is excluded from the measurement by the choice of optical filters. As a consequence, a large excess of acceptor can be used and, when using antibodies, specificity can be sacrificed to gain higher occupancy. In photophysical terms, FRET provides an extra channel of nonradiative decay by which the excited state of the donor fluorophore is depopulated. This results in a reduced quantum yield (Q), the ratio of emitted photons over
absorbed photons. The reduction in quantum yield can be determined in two ways: (1) by the decrease in steady-state fluorescence emission; or (2) the decrease in fluorescence lifetime (τ), which characterizes the duration of the excited state of the fluorophore. The measurement of the change in quantum yield by steady-state emission has to be calibrated due to its dependence on concentration and light path. In contrast, the fluorescence lifetime is proportional to Q and is independent of concentration and light path. Fluorescence lifetimes are measured by fluorescence lifetime imaging microscopy (FLIM; Lakowicz and Berndt, 1991; Gadella et al., 1993; Gadella and Jovin, 1995) and can be performed sufficiently fast to enable realtime live cell experiments. FLIM requires a specialized microscopy setup enabling highfrequency modulation of the excitation light and the gain on the detector. This technique can be combined with acceptor photobleaching to provide an internal lifetime reference, e.g., at the end of a live-cell time-lapse sequence. A detailed description of a frequency domain FLIM set-up is given by Squire and Bastiaens (1999). This type of imaging system is gradually being made commercially available by companies such as LaVision or Lambert Instruments (see SUPPLIERS APPENDIX). Calibration in the intensity-based FRET method is achieved by photobleaching the acceptor to provide the unquenched donor (Bastiaens et al., 1996; Bastiaens and Jovin, 1998; Wouters et al., 1998). In this method, an exposure of the donor fluorescence emission is made (FDA, fluorescence emission of donor in presence of the acceptor). Removal of the acceptor fluorophore from the sample by photobleaching enables the acquisition of an unquenched donor emission image when the ex-
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posure is taken with settings identical to those used for the original image (FD, fluorescence emission of the donor in absence of the acceptor). This essentially recovers that part of the fluorescence intensity that is lost to FRET. A pixel-by-pixel FRET efficiency map is obtained by simple image arithmetic: E = 1 − (FDA/FD). The specificity of the acceptor photobleaching techniques lies in the steep edge at the long-wavelength (i.e., red edge) of the absorption spectrum of the donor, enabling exclusive photobleaching of the acceptor. Essential to the success of this technique is that the photoproduct of the acceptor does not exhibit residual absorption and does not fluoresce at donor emission wavelengths. Cy3 meets both criteria to act as a proper acceptor for GFP; the same is the case for Cy5 acting as acceptor for Cy3. Since photobleaching of the acceptor occurs on a minute time scale, this type of FRET determination is restricted to fixed cells. FRET microscopy can be used to detect protein-protein interactions in single cells as if one were performing an immunoprecipitation experiment. There is, however, no need to isolate the complex or remove it from its physiological environment prior to investigation, and the experiment can be performed in living cells. A number of approaches can be followed: (1) the donor molecule can be purified, labeled with Cy3, and introduced into cells by microinjection (Bastiaens and Jovin, 1996); (2) the donor molecule can be detected by a Cy3-labeled antibody or Fab fragment (provided the antibody is highly specific) on fixed cells (Ng et al., 1999); (3) the donor protein can be fused to one of the mutants of the intrinsically fluorescent green fluorescent protein (GFP) from the jellyfish Aequoria victoria (Tsien, 1998), genetically encoded, and expressed; (4) acceptor proteins (mostly antibodies or Fab fragments) can be labeled with Cy5 (for Cy3 donors), or labeled with Cy3 (for GFP donors) and introduced by microinjection or incubation in the case of antibodies; and (5) the recently discovered and commercial available red fluorescent protein from the Anthozoa sp. (Matz et al., 1999) can be employed. The latter opens up a wide array of possibilities since an ideal acceptor for GFP can be coexpressed in the same cell, obviating the need for exogenous labeling and microinjection. Alternative methods for measuring FRET on the basis of fluorescence intensities are available. In these methods FRET is not determined from the donor fluorescence exclusively, but also from the sensitized emission of the
acceptor, excited by receiving energy from the donor. These approaches fall into two categories (1) those based on changes in donor/acceptor emission ratio (Adams et al., 1991; Miyawaki et al., 1997); and (2) those based on acceptor sensitized emission (Day, 1998; Gordon et al., 1998; Mahajan et al., 1998). These approaches are attractively simple to perform but may suffer from a number of drawbacks that cannot always be corrected for. In the ratiometric approach, the decrease in donor emission and concomitant increase in acceptor emission are imaged by calculating the ratio of these intensities at each pixel. Since this is a relative measure, a change in FRET can only be observed in live cells where the ratio changes over time. A decrease in the donor/acceptor emission ratio can be taken as an indication for the occurrence of FRET. However, this ratio is also dependent on the local concentrations of the donor and acceptor molecules measured in each pixel, which can complicate the interpretation by differential translocation of the biomolecules conjugated to donor and/or acceptor fluorophores. This problem does not occur when the donor and acceptor fluorophores are present on the same molecule (e.g., the “chameleon” Ca2+ biosensors; Miyawaki et al., 1997), since the relative concentrations are identical at each location in the cell. Measuring FRET from the sensitized emission intensity alone (by using a “FRET filter set”) is the most widely used method. Emission from the acceptor would be an exclusive readout for FRET when contamination with donor fluorescence (i.e., donor bleed-through) and direct excitation of the acceptor did not complicate the approach. Correction for these effects by extensive control measurements, some of which have to be performed in different samples where the acceptor is absent, is described in Gordon et al. (1998). Here, both donor-quenching and acceptor-emission information are used to derive the sensitized emission contribution, which is proportional to the FRET efficiency. However, sensitized emission is also proportional to the concentration of the acceptor. Therefore, relative populations of associated molecules cannot be determined. Molecules that are not participating in FRET are not detected. In contrast to donor-based measurements, no estimation can be made of the bound/unbound fractions from the sensitized emission alone. Since excitation spectra generally exhibit an extensive tail at the lower wavelength (blue-edge) the acceptor can easily be excited at the wavelength used for excitation
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of the donor. Direct excitation will be especially problematic when the recently discovered red fluorescent protein from the Anthozoa sp. is used as an acceptor, since its absorption spectrum shows multiple absorption peaks at the excitation optima for the green and yellow fluorescent protein.
Critical Parameters and Troubleshooting
Imaging Protein-Protein Interactions by FRET Microscopy
FRET measurements are prone to false negative results. While the finding of FRET by unquenched fluorescence is highly unlikely to be caused by artifactual processes or nonspecific proximity of donor and acceptor, the absence of FRET does not provide proof for the absence of an interaction for a number of reasons. The presence of a GFP moiety on the protein of interest might interfere with its function or targeting, thus affecting the interaction with the target protein. Cloning vectors for the construction of GFP fusion proteins routinely contain a multiple cloning site that has been optimized for a maximum number of restriction sites to facilitate cloning. As a consequence, the random amino acid residues that they translate into can adopt an unfortunate secondary structure that hinders the proper behavior of the fused protein. In the case of the EGFR-GFP, the amino acid residues encoded by the cloning site were replaced by a flexible six-glycine linker between the GFP moiety and the EGFR to prevent the GFP from affecting EGFR function (Wouters and Bastiaens, 1999). Another problem might arise when the separation distance between the GFP moiety on the donor protein and the Cy3 groups on the antibody is too large (i.e., >10 nm) for energy to be transferred efficiently. This might be overcome by using antibodies raised against a different epitope, or the entire protein, thus increasing the chance of a favorable orientation of the antibody to the donor fluorophore. The labeling ratio of the antibodies is a common reason for failure to detect FRET. The labeling ratio should exceed 1 to prevent unlabeled antibody from competing with the labeled antibodies. A higher labeling ratio is beneficial for FRET detection since the R0 distance (i.e., characteristic value for a given donor-acceptor pair, the distance at which 50% of the excited state energy is transferred by FRET) is effectively increased by a larger number of acceptor molecules per donor molecule. In the authors’ experience, labeling ratios up to 5 do not significantly affect the specificity of the
antibody. However, this should be verified by using the Cy3-labeled antibody in an immunofluorescence experiment and judging the staining pattern as compared to unlabeled antibody. Remember that absolute specificity is not a requirement in the FRET assay, because only the donor photophysical properties are used. However, in extreme cases, the highly labeled antibody could become nonspecific to a point where significant amounts bind to the donor. It is essential that the donor images before and after acceptor photobleaching be taken under identical conditions. Any change in parameters that influence the collection efficiency of donor fluorescence (i.e., brightness, contrast, laser power, and averaging) will influence the calculation of the FRET efficiency. This will affect the entire image, and the control region in the E map will no longer be distributed around 0 but will be uniformly shifted. This also occurs with donor photobleaching. When the laser is used at high intensity, or when too many scans are made for averaging, a substantial amount of the GFP becomes photobleached. This will lead to a shift to negative E values in the control region. Furthermore, it is essential to verify that the sample has not moved and that the same focal plane is imaged in both donor images. Movement causes structures in the control region in the E map with positive values at the leading edge and negative values at trailing edges. Movement is most often caused by temperature differences between the slide and the objective but can also be caused by drift in the slide holder. To correct for translation of the image after acceptor photobleaching, the maximum in the correlation function between pre- and post-bleaching images gives the shift between the two images (Bastiaens and Jovin, 1998). The correlation function of the two images is obtained by Fourier transformation on both images, followed by multiplication of the conjugate of the Fourier transform of one image by the Fourier transform of the other, and performing an inverse Fourier transform on the resulting image. Another source of structured contrast in the control region in the E map can be caused by focal mismatch between the two images, originating from the apparent appearance (positive E) or disappearance (negative E) of structures. This problem can be prevented by observing the z position of the stage between exposures. A larger pinhole can reduce these problems by decreasing the depth of focus but at the cost of z resolution. When using pre- and post-z sections rather than single-focus plane images, the
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Figure 19.5.1 (A) Cy3-PY72 photobleaching releases FRET-quenched EGFR-GFP emission. The histogram shows the distribution of calculated FRET efficiencies in the cell. (B) FRET measured by fluorescence lifetime imaging microscopy. The histogram shows the distribution of measured lifetime (nsec) and corresponding calculated FRET efficiencies prior and after photobleaching of the acceptor. This black and white facsimile of the figure is intended only as a placeholder; for full-color version of figure go to http://www.currentprotocols.com/colorfigures.
maximum in a three-dimensional correlation function can be used for registration of the two data sets and a correct three-dimensional E map can be obtained (Bastiaens et al., 1996).
Anticipated Results After photobleaching of the acceptor, an increase in donor emission intensity can be observed when substantial FRET is present.
Image arithmetic should produce FRET efficiencies that are closely distributed around zero in the portion of the cell that was not photobleached. Deviations from this condition indicate donor bleaching, lateral movement, or focal mismatch. With the occurrence of FRET, a separate efficiency distribution is expected in the photobleached area. The heterogeneity in this distribution contains information about
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variability in the formation of complexes of the proteins carrying the donor and acceptor fluorophores. Figure 19.5.1 shows an MCF-7 mammary carcinoma cell expressing EGFR-GFP that was stimulated with 100 ng/ml EGF for 5 min. This cell was incubated with Cy3-labeled anti-phosphotyrosine antibodies (e.g., 10 µg/ml PY72 monoclonal) and EGFR-GFP tyrosine phosphorylation was measured by FRET using the unquenching of GFP fluorescence by acceptor photobleaching (Fig. 19.5.1A) and FLIM (Fig. 19.5.1B). The EGFR-GFP distribution is shown in the D1 image. Most EGFR-GFP is localized at the plasma membrane in addition to internalized (punctate) EGFR-GFP. The corresponding antibody-staining A1 shows phosphorylation at the plasma membrane. A rectangular region was subjected to acceptor photobleaching (white box). The GFP fluorescence in this region increased after photobleaching as shown in the difference image (D2 − D1), demonstrating positive FRET efficiencies [(D2 − D1)/D2]. Highest FRET efficiencies can be observed in the plasma membrane corresponding to fully phosphorylated receptor at this location. The large peak in the energy-transfer efficiency histogram that is distributed around zero originates from the area outside the photobleached region and indicates proper image registration. The additional population, ranging from 15% to 35% efficiency, corresponds to the photobleached region. Figure 19.5.1B shows the analogous result of a fluorescence lifetime measurement. The steady-state fluorescence distribution of EGFR-GFP is shown in (D) and the corresponding lifetime map in (τ1). The anti-phosphotyrosine Cy3 immunofluorescence (A) is photobleached to obtain an intracellular reference lifetime in absence of acceptor (τ2). As can be seen in the fluorescencelif etim e h istog ram, these values are homogeneously distributed around an average of ∼2.0 nsec. The decrease in lifetime due to FRET is shown in the difference image (τ2 − τ1). The energy-transfer efficiency is given by the normalization of this difference to the reference EGFR-GFP lifetime [E = (τ2 − τ1)/τ2] and again shows highly phosphorylated receptor in the plasma membrane.
Imaging Protein-Protein Interactions by FRET Microscopy
incubation of cells; 1/2 day of data acquisition; and 1/2 day for data analysis. Removal of gelatin/BSA from commercial antibody takes ∼3 hr. The most time-consuming step in the procedure for labeling antibodies is the buffer exchange using a Microcon concentration device. Depending on the amount of contaminating free amino groups in the original buffer, this can take between 2 and 5 hr. The labeling reaction and subsequent gel filtration chromatography takes ∼1 hr. There are a number of points where the procedure can be interrupted: (1) optimal expression after overnight incubation rather than a 3- to 4-hr incubation post microinjection can be achieved by lowering the concentration of DNA; (2) after fixation and permeabilization, the cells can be stored overnight in PBS at 4°C before antibody incubation; (3) after mounting, the cells can be viewed immediately rather than the following day, by application of rubber cement instead of Mowiol; and (4) the samples can be stored at −20°C for weeks without appreciable loss of antibody staining or FRET.
Literature Cited Adams, S.R., Harootunian, A.T., Buechler, J., Taylor S.S., and Tsien, R.Y. 1991. Fluorescence ratio imaging of cyclic AMP in single cells. Nature 349:694-697. Bastiaens, P.I.H. and Jovin, T.M. 1996. Microspectroscopic imaging tracks the intracellular processing of a signal-transduction protein: Fluorescent labeled protein kinase C beta I. Proc. Natl. Acad. Sci. U.S.A. 93:8407-8412. Bastiaens, P.I.H. and Jovin, T.M. 1998. Fluorescence resonance energy transfer microscopy. In Cell Biology a Laboratory Handbook, Vol. 3 (J.E. Celis, ed.), pp. 136-146. Academic Press, New York. Bastiaens, P.I.H. and Squire, A. 1999. Fluorescence lifetime imaging microscopy: Spatial resolution of biochemical processes in the cell. Trends Cell Biol. 9:48-52. Bastiaens, P.I.H., Majoul, I.V., Verveer, P.J., Soling, H.D., and Jovin, T.M. 1996. Imaging the intracellular trafficking and state of the AB(5) quaternary structure of cholera-toxin. EMBO J. 15:4246-4253.
Time Considerations
Bonifacino, J.S. and Dell’Angelica, E.S. 1998. Immunoprecipitation. In Current Protocols in Cell Biology (J.S. Bonifacino, M. Dasso, J.B. Hartfort, J. Lippincott-Schwartz, and K.M. Yamada, eds.), pp. 7.2.1-7.2.21. John Wiley & Sons, New York.
From cell seeding, the entire procedure can be performed in 3 to 4 days. This period includes 1 to 2 days for expression of the GFP construct after transfection or microinjection; 1/2 day for treatment, fixation, and antibody
Budelier, K. and Schorr, J. 1998. Purification of DNA by anion-exchange chromatography. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.), pp. 2.1.11-2.1.18. John Wiley & Sons, New York.
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Clegg, R.M. 1996. Fluorescence resonance energy transfer spectroscopy and microscopy. In Fluorescence Imaging Spectroscopy and Microscopy (X.F. Wang and B. Herman eds.), pp. 179-251. John Wiley & Sons, New York. Day, R.N. 1998. Visualization of Pit-1 transcription factor interactions in the living cell nucleus by fluorescence resonance energy transfer microscopy. Mol. Endocrinol. 12:1410-1419. Förster, T. 1948. Zwischenmolekulare Energiewanderung und Fluoreszenz. Ann. Phys. 2:55-75. Gadella, T.W.J. and Jovin, T.M. 1995. Oligomerization of epidermal growth-factor receptors on A431 cells studied by time-resolved fluorescence imaging microscopy—a stereochemical model for tyrosine kinase receptor activation. J. Cell Biol. 129:1543-1558. Gadella, T.W.J., Jovin, T.M., and Clegg, R.M. 1993. Fluorescence lifetime imaging microscopy (FLIM)—spatial resolution of microstructures on the nanosecond time-scale. Biophys. Chem. 48:221-239. Gordon, G.W., Berry, G., Huan Liang, X., Levine, B., and Herman, B. 1998. Quantitative fluorescence resonance energy transfer measurements using fluorescence microscopy. Biophys. J. 74:2702-2713. Haugland, R.P. 2000. Antibody conjugates for cell biology. In Current Protocols in Cell Biology (J.S. Bonifacino, M. Dasso, J.B. Hartford, J. Lippincott-Schwartz, and K.M. Yamada, eds.), pp. 16.5.1-16.5.22. John Wiley & Sons, New York. Lakowicz, J.R. and Berndt, K. 1991. Lifetime-selective fluorescence imaging using an rf phase-sensitive camera. Rev. Sci. Instrum. 62:1727-1734. Mahajan, N.P., Linder, K., Berry, G., Gordon, G.W., Heim, R., and Herman, B. 1998. Bcl-2 and bax interactions in mitochondria probed with green fluorescent protein and fluorescence energy transfer. Nature Biotechnol. 16:547-552. Matz, M.V., Fradkov, A.F., Labas, Y.A., Savitsky, A.P., Zaraisky, A.G., Markelov, M.L., and Lukyanov, S.A. 1999. Fluorescent proteins from nonbioluminescent Anthozoa species. Nature Biotechnol. 17:969-973.
Mittman, S., Flaming, D.G., Copenhagen, D.R., and Belgum, J.H. 1987. Bubble pressure measurement of micropipette tip outer diameter. J. Neurosci. Methods 22:161-166. Miyawaki, A., Llopis, J., Heim, R., McCaffery, J.M., Adams, J.A., Ikura, M., and Tsien, R.Y. 1997. Fluorescent indicators for Ca2+ based on green fluorescent proteins and calmodulin. Nature 388:882-887. Ng, T., Squire, A., Hansra, G., Bornancin, F., Prevostel, C., Hanby, A., Harris, W., Barnes, D., Schmidt, S., Mellor, H., Bastiaens, P.I.H., and Parker, P.J. 1999. Imaging protein kinase C alpha activation in cells. Science 283:2085-2089. Squire, A. and Bastiaens, P.I.H. 1999. Three dimensional image restoration in fluorescence lifetime imaging microscopy. J. Microsc. 193:36-49. Tsien, R.Y. 1998. The green fluorescent protein. Annu. Rev. Biochem. 76:509-538. Wilson, K. 1994. Preparation of genomic DNA from bacteria. In Current Protocols in Molecular Biology (F.A. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds), pp. 2.4.1-2.4.5. John Wiley & Sons, Nwe York. Wouters, F.S. and Bastiaens, P.I.H. 1999. Fluorescence lifetime imaging of receptor tyrosine kinase activity in cells. Curr. Biol. 9:1127-1130. Wouters, F.S., Bastiaens, P.I.H., Wirtz, K.W.A., and Jovin, T.M. 1998. FRET microscopy demonstrates molecular association of non-specific lipid transfer protein (nsL-TP) with fatty acid oxidation enzymes in peroxisomes. EMBO J. 17:7179-7189.
Contributed by Fred S. Wouters Imperial Cancer Research Fund London, United Kingdom Philippe I.H. Bastiaens European Molecular Biology Laboratory Heidelberg, Germany
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High-Throughput Screening for Protein-Protein Interactions Using Yeast Two-Hybrid Arrays
UNIT 19.6
Arrays are used for parallel assays of large numbers of proteins in a single experiment. They provide an alternative to libraries for protein screening purposes. The use of arrays was fostered by genomics and proteomics with the sequencing of complete genomes, which can now be represented completely on an array. At the end of an array experiment, the positives can be identified immediately because each element, or protein, has a unique address or position in the array. Although many important studies have been carried out using DNA arrays, proteins are more chemically heterogeneous than oligonucleotides, so the construction of large-scale protein arrays that can be screened under uniform conditions is a challenge. Two types of protein arrays have been constructed: living arrays, where the proteins are expressed in cells, and nonliving arrays, where the proteins are purified and tested in vitro. Here the authors describe a protein array composed of living yeast (Saccharomyces cerevisiae) cells that can be used in a functional screen for protein interactions, the two-hybrid assay. In this assay (also see UNIT 19.2), two proteins, A and B, are expressed as fusions with, respectively, a DNA-binding domain (DBD) and a transcriptional activating domain (AD). If A and B interact in the yeast nucleus, the resulting complex will reconstitute a transcription factor, Gal4, that activates one or more reporter genes. Transcription of the reporter gene allows the cells to grow on selective media so that they can be easily scored on a selective plate. Yeast colonies expressing proteins for two-hybrid screening are arrayed in a high-density format, and proteins are screened using automated robotic procedures. The procedure can be modified for manual use or for use with alternative screening strategies such as synthetic lethal screens. With minor modifications, the array can be used to screen for protein interactions with DNA, RNA, or even small-molecule inhibitors of the two-hybrid interaction (see Background Information). The protocols described here use the yeast proteome as an example, but they can be applied to any other genome or subset thereof. The steps of the process involve the construction of the array (see Basic Protocol 1), including the cloning of the arrayed proteins (AD fusions or preys) and the DBD fusions (baits) in yeast; and the screening of the array by either manual (see Basic Protocol 2) or robotic (see Alternate Protocol) manipulation, including the selection of positives and scoring of results. NOTE: To prevent contamination, plate pouring and the transfer steps should be carried out in a sterile hood, or the work surface should be treated regularly with UV light. STRATEGIC PLANNING Many projects involving protein arrays are of a large scale and may be difficult to modify at later stages. Specifically, the size and character of the array must be designed while bearing in mind the initial construction of the array as well as the ultimate aims of the experiment. Many types of arrays are possible. Factors that may be varied include the form of protein arrayed (e.g., wild type or mutant, full length or single domain, ligand or drug bound, epitope tagged). Similarly, the arrayed proteins may be related (e.g., splice variants or alanine-substituted variants of a single protein, a family or pathway of related proteins, orthologs of a protein from different species, the entire protein complement of a model organism). Artificial peptides generated using combinatorial chemical synthesis Contributed by Gerard Cagney and Peter Uetz Current Protocols in Protein Science (2001) 19.6.1-19.6.12 Copyright © 2001 by John Wiley & Sons, Inc.
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may even comprise an array. Different types of arrays will require different construction strategies, and it may be best to carry out a small-scale pilot study, incorporating positive and negative controls, before committing to a full-scale project. Although high-throughput screening projects can be performed manually, automation is strongly recommended. Highly repetitive tasks are not only boring and straining but also error prone when done manually. High-throughput screening therefore should utilize a robotic device. Robots are expensive and require programming and maintenance. Before buying a robot, one should consider whether the robot can also be used for other procedures that are not directly related to the actual screen (e.g., for follow-up studies). BASIC PROTOCOL 1
CONSTRUCTION OF A PROTEOME-SCALE PROTEIN ARRAY FOR YEAST This procedure describes the cloning and expression of yeast proteins for a high-density living array. Because the application involves a large number of proteins (>6000), polymerase chain reaction (PCR) products encoding full-length yeast proteins (as predicted by the published genome sequence) are directly cloned into yeast hosts by transformation and subsequent homologous recombination into the vectors. The recombination event takes place inside the cell following the cotransformation of linearized vector and insert DNA using a modification of the lithium acetate/polyethylene glycol technique (Orr-Weaver et al., 1981). An aliquot of each transformed yeast clone is stored individually, and the entire set of clones is grown on agar so that it can be handled conveniently for screening purposes. To perform the interaction screen, the DNAs encoding the proteins of interest are inserted into two different vectors in two different yeast strains that are identical except at the MAT locus. To clone the activating domain (AD) fusion constructs, the pOAD vector is used in a strain of mating type a (PJ69-4A). These transformants are used to create the array. To clone the DNA-binding domain (DBD) fusion constructs, the pOBD2 vector is used in a strain of mating type α (PJ69-4α). The interactions occur when the DBD and AD strains are allowed to mate (see Basic Protocol 2). The array is made from an ordered set of AD-containing strains, rather than BD-containing strains, because the former do not generally result in self-activation of transcription.
Two-Hybrid Screening with Protein Arrays
Materials DNAs encoding proteins of interest (e.g., genomic DNA, cDNA library) Appropriate primers for amplification of the protein-coding sequences Two-hybrid plasmids: pOAD and pOBD2 (Hudson et al., 1997; Cagney et al., 2000; also see Internet Resources) NcoI and PvuII or other appropriate restriction enzymes, with buffers Liquid YEPD medium (see recipe) S. cerevisiae host strains: PJ69-4A (MATa trp1-901 leu2-3,112 ura3-52 his3-200 gal4∆ gal80∆ LYS2::GAL1-HIS3 GAL2-ADE2 met2::GAL7-lacZ; James et al., 1996) PJ69-4α (MATα trp1-901 leu2-3,112 ura3-52 his3-200 gal4∆ gal80∆ LYS2::GAL1-HIS3 GAL2-ADE2 met2::GAL7-lacZ; Uetz et al., 2000) 0.1 M lithium acetate 7.75 mg/ml salmon sperm DNA (Sigma), autoclaved (15 min at 121°C) and stored at −20°C 96PEG solution (see recipe) Dimethyl sulfoxide (DMSO) 35-mm plates containing solid −Leu (for pOAD) and −Trp (for pOBD2) dropout medium (see recipe) Liquid −Leu dropout medium (see recipe)
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Single-well microtiter plates (e.g., OmniTray, Nalge Nunc International) containing solid −Leu dropout medium (see recipe) 40% (v/v) glycerol 250-ml conical flasks 30°C shaking incubator 8-channel pipettor (to deliver 200 µl) and sterile pipetting troughs 96- and 384-well microtiter plates (e.g., Nalge Nunc International) Plastic adhesive tape for sealing plates 42°C incubator Centrifuge with insert for 96-well plates Toothpicks, sterile 384-Pin Replicator (Nalge Nunc International) 2-ml cryotubes (e.g., Nalge Nunc International) Additional reagents and equipment for PCR amplification (APPENDIX 4J), agarose gel electrophoresis (APPENDIX 4F), digestion of DNA with restriction endonucleases (APPENDIX 4I), and gel purification (APPENDIX 4F) and quantification (APPENDIX 4K) of DNA Amplify protein-coding DNA 1. Individually amplify the DNAs encoding the proteins of interest using appropriate primers and standard PCR conditions (APPENDIX 4J). A two-step PCR protocol may be used, where the first-round reaction uses primers specifying the 5′ and 3′ terminal ends of the DNA encoding the protein to be cloned. These primers are each flanked with ∼20-nucleotide tails that are homologous to sequences in the two-hybrid vectors. In order to increase the efficiency of homologous recombination, a second-round PCR step using ∼70-mer primers can be carried out, where the primers have ∼50 nucleotides that are homologous to the vectors (see Internet Resources; Hudson et al., 1997; Cagney et al., 2000). For the first round of PCR, specific primers must be used for each reaction; for the second round, a common set of primers is used. Appropriate primers for the cloning vector used in this protocol are: First-round forward primer: 5′-AATTCCAGCTGACCACCATGXXX20-30-3′, where ATG represents the start codon of the yeast ORF and XXX20-30 represents ORF-specific sequences of 20 to 30 bases. First-round reverse primer: 5′-GATCCCCGGGAATTGCCATG***XXX20-30-3′, where *** represents the reverse complement of one of the three stop codons and XXX20-30 represents 20 to 30 bases of the reverse complement of ORF-specific sequences at the terminus of the reading frame. Second-round forward primer: 5′-CTATCTATTCGATGATGAA GATACCCCACCAAACCCAAAAAAAGAGATCGAATTCCAGCTGACCACCATG-3′. Seco nd -ro un d reverse p rimer: 5′-CTTGCGGGGTTTTTCAGTATCTACGATTCATA GATCTCTGCAGGTCGACGGATCCCCGGGAATTGCCATG-3′. Other primers can be used when the cloning vectors are modified accordingly. It is recommended that primers be purified to the highest level that is practical (i.e., PAGE, or HPLC, UNIT 8.7). Insertions or deletions in the primers can result in the expression of fusion proteins that are out of frame. UNIT 10.1,
Suggested PCR conditions are: (first round) 30 ng template, 1.5 mM MgCl2, 5 U Taq polymerase (PE Biosystems), 0.02 U Pfu polymerase (Stratagene), 20 pmol each primer; (second round) 5 ng template, 1.5 mM MgCl2, 0.6 U Taq polymerase, 0.003 U Pfu polymerase, 2.5 pmol each primer. It is helpful to amplify the DNAs with approximately equal product sizes grouped together so that the extension times are similar and the products can be checked on the same agarose gel. Many full-length ORFs can now be purchased from commercial suppliers as PCR products or plasmids. Identification of Protein Interactions
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2. Examine the amplified DNAs by agarose gel electrophoresis (APPENDIX 4F) to confirm that they are the correct size. If resources permit, the DNAs can be partially or completely sequenced.
Prepare plasmids and yeast strains 3. Linearize the two-hybrid plasmids (pOAD and pOBD2 vector DNA) by digesting with NcoI and PvuII or other appropriate restriction enzymes (APPENDIX 4I). Gel purify (APPENDIX 4F) and quantify (APPENDIX 4K) the linearized plasmids. pOAD is used for the AD fusion constructs to create the array. pOBD2 is used for the DBD constructs, which will be used to screen the array. UNIT 19.2 describes alternative vector systems. The following steps can be scaled down to perform a smaller number of transformations.
4. For each multiple of 96 PCR products to be cloned, add 50 ml liquid YEPD medium to each of two 250-ml conical flasks. Inoculate each flask with one S. cerevisiae host strain (PJ69-4A and PJ69-4α) from an isolated colony using an inoculating loop and grow these cultures overnight in a 30°C shaking incubator. PJ69-4A is used to clone the AD fusion constructs, and PJ46-4α is used to clone the DBD fusion constructs.
5. Transfer each culture to a 50-ml conical centrifuge tube and centrifuge 3 min at 3000 × g, room temperature, to harvest the yeast cells. Remove the supernatants and resuspend each pellet in 2 ml of 0.1 M lithium acetate. Clone PCR products 6. Boil 1 ml of 7.75 mg/ml salmon sperm DNA in a microcentrifuge tube for 5 min and plunge it into ice water. 7. Add the following reagents (in order) to each of two 50-ml conical tubes: 20 ml 96PEG solution 0.5 ml salmon sperm DNA (step 6) 200 ng linearized pOAD or pOBD2 vector DNA (step 3) 2.5 ml DMSO. Shake vigorously for 30 sec followed by 1 min of vortexing. Add 2 ml yeast suspension (step 5) and mix well by hand for 1 min. 8. Pour each mixture into a sterile pipetting trough and use an 8-channel pipettor to deliver 200 µl into each well of a 96-well microtiter plate. Set up one plate for the pOAD vector and a second for the pOBD2 vector. Wells for control transformations, where no PCR product will be added, should be included.
9. Add 3 µl of each amplified DNA (insert DNA; step 2) to a separate vector-containing well and seal the wells with a secure plastic adhesive tape. Vortex 4 min and incubate 30 min in a 42°C incubator. A vector DNA to which no PCR product was added should be used as a negative control for transformation.
10. Centrifuge the 96-well plates 7 min at 3000 × g, room temperature.
Two-Hybrid Screening with Protein Arrays
11. Remove the supernatant by aspiration with the 8-channel pipettor. Add 200 µl sterile water to each well and resuspend the yeast with the pipettor.
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12. Spread the pOAD and pOBD2 yeast suspensions onto 35-mm −Leu and −Trp plates, respectively, and incubate 2 to 3 days at 30°C. The authors suggest testing the clones by colony PCR or sequencing before assembling the array. Ideally, each colony or clone should be tested for expression of its cognate fusion protein using immunoblotting (UNIT 10.10). This may be prohibitively difficult or expensive to carry out for thousands of proteins, especially with the low-copy plasmids described here.
Construct array 13. Pick individual AD colonies from the −Leu plates using sterile toothpicks and transfer to the wells of 384-well microtiter plates containing liquid −Leu dropout medium for inclusion in the array. Note the address of each colony, preferably in a searchable database. One colony is picked from each plate if it can be tested by PCR or sequencing. If there are too many clones to test individually, two or more clones from a single AD plate may be pooled at this stage in order to avoid occasional PCR mutants or empty vectors. A larger number of pooled clones may be used, but this increases the risk of fast-growing variants out-competing the desired ones. The authors have found that recovery of array clones is best following storage in YEPD. Therefore, a duplicate 384-well microtiter plate containing YEPD should be set up with the array clones, grown overnight at 30°C, and frozen indefinitely in 20% (v/v) glycerol (i.e., after adding an equal volume of 40% glycerol) at −80°C.
14. Incubate overnight at 30°C without shaking. 15. Use a 384-Pin Replicator to transfer the clones to single-well microtiter plates containing solid −Leu dropout medium, according to manufacturer’s instructions. Duplicate each element on the array (i.e., place two colonies that express the same protein in adjacent positions on the array). Alternatively, duplicate copies of the array can be made. Using duplicate elements or arrays assures reproducibility (see Anticipated Results, discussion of false positives). When arranging the colonies on the array, standard formats should be used so that the colonies can be manipulated by robots and other standardized labware. When the array is used for screening, this format can be condensed 4-, 8-, or 16-fold to include 384, 768, or 1536 colonies per microtiter-plate footprint. However, scoring positives becomes more difficult at higher densities. For day-to-day use, the array can be stored on solid medium lacking leucine (−Leu) for up to 3 months at 4°C. Working copies of the array grown on complete medium (YEPD) can be generated from the −Leu copy every 4 weeks (or as often as needed) for use in screens (see Basic Protocol 2).
Prepare bait 16. Pick individual DBD colonies from −Trp plates and grow overnight in 20 ml YEPD at 30°C. Dilute culture 1:1 with 40% glycerol and transfer aliquots to 2-ml cryotubes. Freeze indefinitely at −80°C. MANUAL SCREENING FOR PROTEIN INTERACTIONS USING A YEAST PROTEIN ARRAY The living array (see Basic Protocol 1) can be screened for protein interactions by a mating procedure that can be carried out manually (described here) or using a robot (see Alternate Protocol). A strain expressing a single candidate protein as a DBD fusion is mated to all the colonies in the array, which contains yeast colonies expressing individual AD fusion
BASIC PROTOCOL 2
Identification of Protein Interactions
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proteins. After mating, the colonies are transferred to diploid-specific medium, and then to two-hybrid selective medium. To manually screen with more than one bait, replicate copies of the array are used. For large numbers of baits, robotic screening is recommended. Materials 20% (v/v) bleach (∼1% sodium hypochlorite) 95% (v/v) ethanol Yeast protein array (see Basic Protocol 1) Single-well microtiter plates (e.g., OmniTray; Nalge Nunc International) containing solid YEPD +Ade medium (see recipe), YEPD medium (see recipe), −Leu −Trp dropout medium (see recipe), and −His −Leu −Trp +3AT dropout medium (see recipe) Liquid YEPD medium (see recipe) DBD fusion–expressing yeast strain (see Basic Protocol 1) 384-Pin Replicator (Nalge Nunc International) 30°C incubator 250-ml conical flask 1. Sterilize a 384-Pin Replicator by dipping the pins into 20% bleach for 20 sec, sterile water for 1 sec, 95% ethanol for 20 sec, and sterile water again for 1 sec. Repeat this sterilization before each transfer. 2. Use the sterile pin replicator to transfer a yeast protein array to single-well microtiter plates containing solid YEPD +Ade medium and grow the array overnight in a 30°C incubator. If duplicate colonies were not used to construct the array (see Basic Protocol 1, step 15), the entire experiment should be done using duplicate arrays. YEPD medium with supplemental adenine increases the efficiency of the mating step.
3. Inoculate 20 ml liquid YEPD medium in a 250-ml conical flask with a DBD fusion–expressing yeast strain and grow overnight at 30°C. Ideally, the frozen strain (see Basic Protocol 1, step 16) is streaked out on a plate containing solid YEPD medium and grown overnight at 30°C. A colony from this plate is then used to inoculate the liquid medium. This volume is sufficient for mating to the entire array.
4. Dip the pins of the pin replicator into the DBD fusion–expressing culture and place directly onto a fresh single-well microtiter plate containing solid YEPD medium. Repeat with the required number of plates. 5. Pick up the array (i.e., AD) yeast colonies with sterilized pins and transfer them directly onto the DBD strain, so that each of the 384 DBD yeast spots per plate receives different AD yeast cells (i.e., a different AD fusion). Incubate 1 to 2 days at 30°C to allow mating. A scaffold of some kind should be used to ensure that the DBD and array strains contact each other. The use of a robot for this procedure is discussed (see Alternate Protocol). Mating will take place in <24 hr, but a longer period is recommended because some bait (DBD) strains show poor mating efficiency.
Two-Hybrid Screening with Protein Arrays
6. For selection, transfer the colonies to single-well microtiter plates containing solid −Leu −Trp dropout medium using the sterilized pinning tool. Grow for 2 days at 30°C until the colonies are >1 mm in diameter.
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This is an essential control step because only diploid cells that contain Leu and Trp markers on pOAD and pOBD2, respectively, will grow on this medium. This step also helps recovery of the colonies and increases the efficiency of the next selection step.
7. Transfer the colonies to a single-well microtiter plate containing solid −His −Leu −Trp +3AT dropout medium using the sterilized pinning tool and grow at 30°C for up to 10 days (or longer if there is little or no background growth). The stringency of the screen can be varied by adding different amounts of 3AT, an inhibitor of the His3 gene product (imidazoleglycerolphosphate dehydratase). For standard screening purposes, 3 mM is suggested. In many cases (10% to 20% of full-length yeast proteins tested by the authors), the haploid strain expressing the DBD fusion has transcriptional self-activation properties. These haploid strains can be titrated on −His plates containing increasing amounts of 3AT. The highest level of 3AT tolerated should be added to the −His −Leu −Trp plates for selection of two-hybrid positive diploids. In many cases, however, the transcriptional activity is very strong (>200 mM 3AT) and alternative approaches must be considered, such as reengineering the construct to express smaller protein fragments. The vectors and strains described here are also suitable for two-hybrid screening on −Ade −Leu −Trp plates. The use of two alternative two-hybrid reporters (His and Ade) can be useful for reducing the levels of false positives, but the Ade selection is very stringent (equivalent to ∼200 mM 3AT) and the interactions of many proteins may be of insufficient affinity to permit growth on these plates.
8. Score the interactions by looking for signals (i.e., growing colonies) that are significantly above background (by size) and that are present for both duplicate colonies (see Basic Protocol 1, step 15). The plates should be examined every day. Most two-hybrid positive colonies appear within 3 to 4 days, but occasionally positive interactions can be observed later. Very small colonies are usually designated as background; however, there is no absolute measure to distinguish between background and real positives. When there are many (i.e., >30) large colonies per array of 6000 positions, the authors consider the baits used as random activators. In this case the screen should be repeated or should not be analyzed. Scoring can be done manually or using automated image analysis procedures. When using image analysis, care must be taken not to score contaminant colonies as positives. Aspects of the scoring process are discussed more fully (see Anticipated Results, discussion of analysis of screens).
ROBOTIC SCREENING FOR PROTEIN INTERACTIONS USING A YEAST PROTEIN ARRAY
ALTERNATE PROTOCOL
In many cases a hand-held 384-pin replicating tool can be used for routine transfer of colonies for screening. For larger projects, however, a robotic workstation (e.g., Biomek 2000; Beckman Coulter) may be used to speed up the screening procedures and to maximize reproducibility. A 384- or 768-pin stainless steel replicating tool (e.g., HighDensity Replication Tool; Beckman Coulter) can be used to transfer the colonies from one plate to another. Between transfer steps, the tool should be sterilized by sequential immersion into a 20% (v/v) bleach solution (20 sec), sterile water (1 sec), 95% (v/v) ethanol (20 sec), and sterile water (1 sec). The level of these liquids should be 2 to 4 mm from the base of the pin and care must be taken that the ethanol does not evaporate. It is important to ensure that plasticware is compatible with the movements of the robot. In the procedure described above, the array is gridded on eight 86 × 128–mm single-well microtiter plates (e.g., OmniTray, Nalge Nunc International) in 768-colony format. Identification of Protein Interactions
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
C dropout powder 1 g histidine 1 g methionine 1 g arginine 2.5 g phenylalanine 3 g lysine 3 g tyrosine 4 g tryptophan 4 g leucine 4 g isoleucine 5 g glutamic acid 5 g aspartic acid 7.5 g valine 10 g threonine 20 g serine 1 g adenine 1 g uracil Store up to 2 years at room temperature For leucine-, tryptophan-, and histidine-free media (−Leu, −Trp, and −His), leave out these amino acids.
Dropout medium For liquid medium: To 800 ml H2O add: 1.7 g yeast nitrogen base without amino acids 5 g ammonium sulfate 20 g dextrose 1.4 g C dropout powder (see recipe) Add H2O to 1 liter Autoclave and store up to 10 weeks at 4°C For +3AT dropout medium, add 3 mM 3-aminotriazole (final concentration).
For solid medium: Add 16 g agar (e.g., Difco, Becton Dickinson) before autoclaving. Cool to 45°C and pour into appropriate plates. Wrap plates and store up to 6 months at 4°C. 96PEG solution 45.6 g polyethylene glycol (avg. mol. wt. 3350; e.g., Sigma) 6.1 ml 2 M lithium acetate 1.14 ml 1 M Tris⋅Cl, pH 7.5 (APPENDIX 2E) 232 µl 0.5 M EDTA, pH 8.0 (APPENDIX 2E) H2O to 100 ml Store up to 1 year at room temperature
Two-Hybrid Screening with Protein Arrays
YEPD medium For liquid medium: To 800 ml H2O add: 10 g yeast extract (e.g., Sigma; Difco, Becton Dickinson) 20 g peptone (e.g., USBiological) 20 g dextrose (e.g., Fisher) continued
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Add H2O to 1 liter Autoclave and store up to 3 months at room temperature For solid medium: Add 14 g agar (e.g., Difco, Becton Dickinson) before autoclaving. Cool to 45°C and pour into appropriate plates. Wrap plates and store up to 6 months at 4°C. YEPD +Ade medium Prepare solid YEPD medium (see recipe) but add 10 ml 0.2% (w/v) adenine before autoclaving. COMMENTARY Background Information The two-hybrid system was originally invented to detect interactions between characterized proteins (Fields and Song, 1989). It quickly became clear that it could be used to identify new interactions when expression libraries were screened. With the advent of complete genome sequences, the original idea of testing known protein pairs was revived again: instead of using random libraries, all possible pairwise protein combinations were tested in order to identify interacting partners. Such systematic, genome-wide screens were first reported in studies using DNA chips but could also be applied to proteins. Arrays have the advantage that many tests can be performed under identical conditions at the same time. The results of these individual tests can be compared directly. This is important because baits vary in background levels. Preys with a tendency to cause false positives therefore can also be easily identified. In addition, because arrays allow rapid identification of positives by their position, it is not necessary to sequence positive clones. However, two-hybrid arrays as described here have disadvantages. Because all elements in the array are generated individually, the production of arrays is time and labor intensive. Furthermore, full-length proteins may not interact because steric factors prevent productive reconstitution of the transcription factor Gal4 on the promoter of the reporter gene. Other groups have recommended the use of random libraries that are screened conventionally (Fromont-Racine et al., 1997; Rain et al., 2001). Indeed, random libraries can detect interactions that have not been found using full-length proteins (Flajolet et al., 2000). On the other hand, large-scale screens of random libraries require significant sequencing capacities. For instance, the screening of 261 Helicobacter baits required the sequencing of >13,000 PCR products (Rain et al., 2001). Screens of random
libraries yield more interactions than screens of full-length libraries, but they also require highquality libraries that are representative of weakly expressed transcripts (when cDNA libraries are used). In addition to screening for protein-protein interactions, living protein arrays can be used to study DNA- and RNA-protein interactions (Kraemer et al., 2000) or genetic interactions. These might include screens of mutant proteins against arrays of strains deleted for nonessential proteins, or strains where expression of an essential gene is disrupted using promoter manipulation or drugs. In the reverse-two-hybrid screen, candidate compounds are examined for their ability to disrupt a positive two-hybrid interaction (Vidal and Legrain, 1999).
Critical Parameters Following successful cloning, the mating reaction between the bait and prey yeast strains is critical. Strains vary considerably in their mating efficiency. The strains must be in contact for ≥24 hr. Robots used for transfer functions should be calibrated regularly, in order to prevent misalignment and inefficient transfer of yeast cells. Mold contamination can be a problem, especially for plates growing for >1 week. To prevent contamination of plates, plate pouring and the transfer steps should be carried out in a sterile hood, or the work surface should be treated with UV light regularly.
Troubleshooting About 15% of all baits activate transcription on their own and show strong background growth. Weak activators can usually be screened by elevating the level of 3-aminotriazole up to 150 mM. Another approach is to use additional reporter genes, although multiple reporter genes, and therefore higher stringency, may result in the loss of weak interactions.
Identification of Protein Interactions
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A
B
Two-Hybrid Screening with Protein Arrays
Figure 19.6.1 Two subsequent two-hybrid screens of a full-length activation domain–open reading frame library with full-length PHO85 protein (a cyclin-dependent kinase) as bait. (A) First screen. (B) Second screen. Reproducible positives are indicated by arrows. Some reproducible positives are either nonyeast contaminants (c) or false positives (f). Nonyeast contaminants can be identified by their unusual shape, size, or color. False positives are either nonreproducible or reproducible positives, but the latter are usually found in many independent screens with unrelated baits (hence they are nonspecific). Given these criteria, the following preys were identified as reproducible positives (plates are numbered in columns): PCL10, PCL6, CLG1 (plate 4), SOR1 (plate 5), CDC36 (plate 11), PCL9, PCL2, YPL229W (plate 12), YDL246C (plate 13), PCL8 (plate 14). PCLs are cyclins, and most of them are known PHO85 interactors; CLG1 is a cyclin-like protein; CDC36 is a transcription factor; YPL229W is a protein of unknown function; SOR1 is sorbitol dehydrogenase; and YDL246C is a SOR1-related protein. Most of these preys are either known or highly plausible interactors of PHO85. The biological role of an interaction between SOR1 and PHO85 remains unclear but is supported by YDL246C, which is a relative of SOR1. Note that these two screens have an unusually high number of contaminants on plates 9 and 10. The false positives have been found in more than 10% of all screens.
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When arrays are kept over months or even years, it is important to avoid cross-contamination. These events can accumulate over time and lead to misidentification of colonies. Yeast arrays can also be maintained as frozen stocks in 20% glycerol at −80°C that are used to reestablish the working arrays every couple of months.
Anticipated Results A fairly typical screen is shown in Figure 19.6.1. Ideally it should have no background growth and only a few positives. In most cases a small number of reproducible positives is found (i.e., <10 per screen of ∼6000 preys). It is important to make sure that the positives are reproducible because ∼80% of all positives are caused by random mutations and are therefore not reproducible. A few baits have large numbers of such random positives (random activators). It remains unclear how such random positives arise, but they seem to be dependent on bait properties. About 15% of all baits activate transcription on their own, causing growth of all prey colonies (see Troubleshooting). Analysis of screens The authors analyze their screens using a computer program based on the FileMaker Pro (FileMaker) database application (available for both Windows and Macintosh). The software allows one to hold a two-hybrid selective plate (−His −Leu − Trp) onto a computer screen with an equally sized 384 grid and click on all positive colonies (agar plates are transparent). The software then retrieves all positions and prey identities from a linked database. False positives During the past 10 years, the two-hybrid system has become the foremost method for the detection of protein-protein interactions. In fact, about two thirds of all protein-protein interactions have been identified by this procedure (Uetz et al., 1998; see Internet Resources). Nevertheless, the two-hybrid system is often criticized because of its tendency to produce false positives. Whereas few false positives have been characterized on the molecular level, two main classes can be distinguished: reproducible and nonreproducible false positives. Reproducible false positives are based on nonphysiological two-hybrid interactions (i.e., interactions that take place in the two-hybrid
assay but not in vivo). Further investigations are required to distinguish between physiological and nonphysiological interactions. Nonreproducible false positives are caused by mutations in the reporter gene(s) or the two-hybrid plasmids. These can be identified by repeating the screen at least once. The probability of independent mutations in two different clones generating duplicate two-hybrid positives by chance is less than (x/n)2, where x is the maximum number of positives accepted in the screen, and n is the number of proteins in the array. For the yeast proteome, this probability is (30/6000)2 or 1/40,000. An alternative to two independent screens is to perform duplicate screens on the same plate (e.g., by pinning two prey colonies next to each other). This way, reproducible positives are easily scored by identifying double colonies on the selective (−His −Leu −Trp) plates. Another alternative is to test all the positives from an array screen in manual two-hybrid assays. This may be more labor intensive than duplicate screens, but the retests can be done using larger colonies, which are much easier to score. False negatives False negatives refer to protein interactions that occur in vivo but are not found in the two-hybrid screen. This may result from: (1) structural effects, where the fusion proteins fail to interact because of steric impedance, instability, or improper folding; (2) failure of one or both fusion proteins to localize to the nucleus; or (3) assay insensitivity, where the binding affinity of the proteins is low. Two-hybrid arrays using full-length proteins suffer from a high number of false negatives, and therefore protein fragments (or domains) may be used if possible (e.g., Flajolet et al., 2000).
Time Considerations Building a cell-based array is time consuming and can take several months for genomescale arrays. The yeast prey array described here required two to three full-time workers to assemble. Random libraries (UNIT 19.2) can be made more quickly, but the screens take longer because false positives can be identified only with additional tests. An array screen takes ∼2 weeks to complete, from the initial mating reaction to analysis of the results. Using a laboratory robot, 20 or more screens can be initiated per week. Identification of Protein Interactions
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Literature Cited
Key Reference
Cagney, G., Uetz, P., and Fields, S. 2000. Highthroughput screening for protein-protein interactions using two-hybrid assay. Methods Enzymol. 328:3-14.
Uetz et al., 2000. See above.
Fields, S. and Song, O. 1989. A novel genetic system to detect protein-protein interactions. Nature 340:245-246.
This paper, the first to describe two-hybrid arrays, provides the results of 192 screens and compares them to screens that used a library of preys.
Internet Resources depts.washington.edu/sfields/
Flajolet, M., Rotondo, G., Daviet, L., Bergametti, F., Inchauspe, G., Tiollais, P., Transy, C., and Legrain, P. 2000. A genomic approach of the hepatitis C virus generates a protein interaction map. Gene 242:369-379.
The Fields laboratory Web site contains more details on primer design, vectors, and array screening as well as results from several hundred screens.
Fromont-Racine, M., Rain, J.C., and Legrain, P. 1997. Toward a functional analysis of the yeast genome through exhaustive two-hybrid screens. Nature Genet. 16:277-282.
The Laboratory Robotics Interest Group Web site provides a guide to robotics manufacturers and product information.
Hudson, J.R., Dawson, E.P., Rushing, K.L., Jackson, C.H., Lockshon, D., Conover, D., Lanciault, C., Harris, J.R., Simmons, S.J., Rothstein, R., and Fields, S. 1997. The complete set of predicted genes from Saccharomyces cerevisiae in a readily usable form. Genome Res. 7:11691173.
www.proteome.com
James, P., Halladay, J., and Craig, E.A. 1996. Genomic libraries and a host strain designed for highly efficient two-hybrid selection in yeast. Genetics 144:1425-1436.
www.lab-robotics.org/
Proteome’s Web site includes a Yeast Protein Database. mips.gsf.de/proj/yeast/ The Munich Information Center for Protein Sequences Web site provides yeast protein databases and information on other genomes and proteomes. www.fccc.edu/research/labs/golemis/ InteractionTrapInWork.html
Kraemer, B., Zhang, B., SenGupta, D., Fields, S., and Wickens, M. 2000. Using the yeast three-hybrid system to detect and analyze RNA-protein interactions. Methods Enzymol. 328:297-321.
The Golemis laboratory Web site offers general information on two-hybrid screens and false positives.
Orr-Weaver, T.L., Szostak, J.W., and Rothstein, R.J. 1981. Yeast transformation: A model system for the study of recombination. Proc. Natl. Acad. Sci. U.S.A. 78:6354-6358.
Contributed by Gerard Cagney University of Washington Seattle, Washington and Banting and Best Institute of Medical Research University of Toronto Toronto, Ontario, Canada
Rain, J.C., Selig, L., De Reuse, H., Battaglia, V., Reverdy, C., Simon, S., Lenzen, G., Petel, F., Wojcik, J., Schachter, V., Chemama, Y., Labigne, A., and Legrain, P. 2001. The protein-protein interaction map of Helicobacter pylori. Nature 409:211-215. Uetz, P., Hughes, R.E., and Fields, S. 1998. The two-hybrid system: Finding likely partners for lonely proteins. Focus (Life Technologies) 20:6265.
Peter Uetz University of Washington Seattle, Washington and Research Center Karlsruhe Karlsruhe, Germany
Uetz, P., Giot, L., Cagney, G., Mansfield, T., Judson, R., Knight, J., Lockshon, D., Narayan, V., Srinivasan, M., Pochart, P., Qureshi-Emili, A., Li, Y., Godwin, B., Conover, D., Kalbfleisch, T., Vijayadamodar, G., Yang, M., Johnston, M., Fields, S., and Rothberg, J. 2000. A comprehensive analysis of protein-protein interactions in Saccharomyces cerevisiae. Nature 403:623-627. Vidal, M. and Legrain, P. 1999. Yeast forward and reverse “n”-hybrid systems. Nucl. Acids Res. 27:919-929.
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Identification of Protein Interactions by Far Western Analysis
UNIT 19.7
This unit describes far western blotting, a method of identifying protein-protein interactions. In a far western blot, one protein of interest is immobilized on a solid support membrane, then probed with a non-antibody protein. Far western blots can be used to identify specific interacting proteins in a complex mixture of proteins (see Basic Protocol). They are particularly useful for examining interactions between proteins that are difficult to analyze by other methods due to solubility problems or because they are difficult to express in cells. This method is performed totally in vitro, and the proteins of interest can be prepared in a variety of ways. Peptides can be used to determine the effects of specific residues or post-translational modifications on protein-protein interactions (see Alternate Protocol 2). In addition, many different detection techniques, either radioactive or nonradioactive, can be used. For example, the protein probe may be detected indirectly with an antibody, rather than being labeled radioactively (see Alternate Protocol 1). Thus, techniques and reagents already in hand can frequently be adapted for use with this assay. CAUTION: Appropriate safety precautions must be taken when working with radioactive materials. Information on proper handling and disposal of radioactive compounds can be found in APPENDIX 2B and may be obtained from local radiation safety officials. Specific information on handling 35S-labeled compounds can be found in UNIT 3.7 and in APPENDIX 2B. FAR WESTERN ANALYSIS OF A PROTEIN MIXTURE The following is a basic method for detecting protein-protein interactions by far western blotting when one protein is contained within a simple or complex mixture of proteins. First, the protein sample is fractionated on an SDS-PAGE gel (UNIT 10.1). After electrophoresis, the proteins are transferred from the gels onto a solid support membrane by electroblotting (UNIT 10.7). Transferred membranes may be stained with Ponceau S to facilitate location and identification of specific proteins. Nonspecific sites on the membranes are blocked with standard blocking reagents, and the membranes are then incubated with a radiolabeled non-antibody protein probe. After washing, proteins that bind to the probe are detected by autoradiography (UNIT 10.11).
BASIC PROTOCOL
Materials Samples to be analyzed 1× SDS sample buffer (UNIT 10.1) Ponceau S staining solution (see recipe) Blocking buffer I: 0.05% (w/v) Tween 20 in 1× PBS (see recipe for PBS); prepare fresh Blocking buffer II: dissolve 1 g bovine serum albumin (BSA; fraction V) in 100 ml 1× PBS (see recipe for PBS); prepare fresh Phosphate-buffered saline (PBS; see recipe), pH 7.9 cDNA encoding protein of interest cloned into an in vitro expression vector In vitro transcription/translation kit (Promega) 10 mCi/ml 35S-methionine (1000 Ci/mmol) Probe purification buffer (see recipe) Probe dilution buffer (see recipe) Identification of Protein Interactions Contributed by Diane G. Edmondson and Sharon Y. R. Dent Current Protocols in Protein Science (2001) 19.7.1-19.7.10 Copyright © 2001 by John Wiley & Sons, Inc.
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Polyvinyldifluoridine (PVDF) or nitrocellulose membrane for protein transfer Microfiltration centrifuge columns (e.g., Gelman Nanosep, Pall Filtron, or Millipore Microcon) Additional reagents and equipment for SDS-PAGE (UNIT 10.1), electrophoretic transfer of proteins to a support membrane (UNIT 10.7), and autoradiography (UNIT 10.11) NOTE: Always handle support membranes with gloves or membrane forceps. Prepare protein blot 1. Prepare the protein sample to be analyzed by resuspending it in 1× SDS sample buffer. UNIT 10.1 gives instructions on preparation of samples and amount of samples to load. In general, ∼50 to 100 ìg can be loaded in each lane for a complex mixture of proteins. A smaller amount, i.e., 10 to 20 ìg, is loaded for less complex protein samples. The amount loaded may also need to be adjusted for the size of gel. (Usually 30 ìg/mm2 loading surface can be resolved without smearing.)
2. Separate the samples on an SDS-polyacrylamide gel (UNIT 10.1). 3. Transfer the proteins from the gel to a solid support membrane (e.g., PVDF or nitrocellulose) by semidry electroblotting (UNIT 10.7). Either nitrocellulose or PVDF membranes can be used with good results. PVDF membranes are easier to handle and tend to give a slightly higher signal-to-noise ratio, probably due to increased protein retention by the membrane.
Stain with Ponceau S 4. After transfer, stain the membrane for 5 min in ∼100 ml freshly diluted 1× Ponceau S staining solution. Stain the membrane in a plastic container large enough to hold the blot and use sufficient Ponceau S to cover the membrane completely. This step is optional. When the protein samples consist of a few proteins, or when there are clearly visible bands that facilitate orientation of the blot, staining with Ponceau S can provide helpful landmarks. One can unequivocally identify interacting bands, mark the position of molecular weight standards, and trim away excess membrane more exactly.
5. Destain the membrane washing in several changes of deionized water until the proteins are clearly visible. Place light pencil marks adjacent to important protein bands to mark them for future reference. Trim away excess membrane. The stain fades quickly so the marks must be placed immediately.
6. Destain an additional 5 min in water until the red staining fades. Block membrane 7. Block blot for 2 hr in 200 ml blocking buffer I at room temperature with gentle agitation. 8. Decant and add 200 ml blocking buffer II. Incubate as in step 7. 9. Decant blocking buffer II and rinse the membrane briefly in 100 ml of 1× PBS. At this point, the blot may be probed immediately or may be wrapped in plastic wrap and stored for up to 2 weeks at 4°C. Identification of Protein Interactions by Far Western Analysis
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Prepare the probe 10. Following manufacturer’s procedures, prepare a radiolabeled in vitro–translated probe of the protein of interest using 35S-methionine. The probe can be conveniently prepared during the blocking steps. The authors routinely use the Promega TnT quick-coupled transcription/translation system for producing probes. For a small blot (e.g., ≤9 × 9–cm), one half of a standard in vitro transcription/translation reaction (i.e., 25 ìl) is sufficient for the probe. For larger blots, an entire 50-ìl reaction may be used. In the authors’ laboratory it is considered essential that the transcription/translation lysate not be repeatedly frozen and thawed.
11. After translation, dilute the probe with 500 µl probe purification buffer, and purify by microcentrifuging 15 to 30 min at 10,000 × g, room temperature, in a microfiltration column. Save aliquots of the purified probe for analysis by SDS-PAGE (i.e., 2 µl), and for scintillation counting (i.e., 2 to 5 µl). Check microfiltration column manufacturer’s procedure for exact centrifugation times required by different columns. In practice, it is not always necessary to purify the probe through a microfiltration column; many probes give good signals without purification. However, if signal-to-noise ratio is low, probe purification may improve results. In addition, it is possible to quantitate the proportion of probe bound if the probe has been purified.
Bind probe 12. Preincubate blot for 10 min in 50 ml of 1× probe dilution buffer (without probe) by gently agitating at room temperature. 13. Dilute the translated probe with 1× probe dilution buffer in a volume sufficient to cover the membrane to be probed (typically 3 ml). For small blots, i.e., ≤9 × 9–cm, a 50-ml conical tube makes a convenient incubation chamber. A volume of 3 ml is enough solution to cover the blot and tubes can be rotated on a mechanical rotator. In addition, the conical tube makes the radiolabeled probe easy to contain and dispose of. Larger blots can be incubated on a Nutator or orbital shaker in a heat-sealable bag, or rotated in a hybridization oven adjusted to room temperature.
14. Add the probe to the membrane and incubate 2 hr at room temperature. Rotate the tubes or agitate bags throughout the binding reaction. Wash the membrane 15. Transfer the membrane to a plastic dish and wash the membrane with 200 ml 1× PBS for 5 min, room temperature. Repeat for a total of four washes. Background is generally quite low and extended washing does not substantially reduce background.
16. Air dry the membrane and expose to X-ray film (autoradiography) or phosphor imager screen (see UNIT 10.11 for both techniques). Do not cover the blots with plastic wrap as this will quench the 35S signal. Overnight exposure to X-ray film is usually sufficient to detect positive interactions.
DETECTING INTERACTING PROTEINS BY IMMUNOBLOTTING In vitro–translated probes have the advantages of being quickly produced, easily detected, and quantitated to give an estimate of relative binding. In addition, mutations in the protein probe can be generated by simple cloning procedures and can provide information on binding domains and their critical residues. A disadvantage of in vitro–translated probes is the need for multiple methionine or cysteine residues to obtain a well labeled probe. For the same reasons, small peptide fragments are often not suitable for use as in
ALTERNATE PROTOCOL 1
Identification of Protein Interactions
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vitro–translated probes. [14C]leucine and [3H]leucine can also be used for in vitro translation of proteins; however, in the authors’ laboratory [14C]leucine has not yielded probes suitable for use as far western probes. There are many other ways to generate probes for far western blots. The protein probe may be labeled in vitro with 125I (UNIT 3.3) or enzymatically with 32P (Kimball, 1998). Biotin-labeled probes may be detected with strepavidin-biotin detection schemes (UNIT 3.6; Grulich-Henn et al., 1998; Kimball et al., 1998). Protein binding may be detected indirectly as well. If an antibody to the interacting protein is available, then an unlabeled protein probe can be bound to the blots as usual and then detected by western (immunoblot) analysis. This is especially useful when a tagged recombinant protein and antibody to the tag are available. The following procedure describes detection of an unlabeled protein probe with specific antibody. Many variations of immunoblotting exist; additional information and procedures may be found in UNIT 10.7. Additional Materials (also see Basic Protocol) Recombinant protein or unlabeled in vitro–translated protein for probe 5% (w/v) non-fat instant dry milk in 1× TBST (see recipe for TBST) Primary antibody specific for protein probe TBST (see recipe) Alkaline phosphatase (AP)–conjugated secondary antibody against Ig of species from which specific antibody was obtained Alkaline phosphatase buffer (see recipe) Developing solution (see recipe) 100 mM EDTA, pH 8.0 (APPENDIX 2E) Prepare blot and probe 1. Prepare and block the blot (see Basic Protocol, steps 1 to 9). 2. Prepare the probe protein by diluting in vitro–translated or recombinant protein in 3 ml of 1× probe dilution buffer. The amount of recombinant protein must be empirically determined for each protein. Various researchers have used from 0.5 to 20 ìg recombinant protein/ml of probe dilution buffer.
Expose blot to probe 3. Bind the probe to blot and wash (see Basic Protocol, steps 12 and 14 to 15). Do not dry the membrane after washing. 4. Incubate blot in 200 ml of 5% non-fat milk in 1× TBST for 1 hr, room temperature, with gentle rotation on an orbital shaker. Expose to antibodies 5. Dilute the primary antibody in 5% milk in 1× TBST. Incubate blot in 5 to 10 ml diluted antibody at room temperature with gentle agitation to ensure blot is evenly covered with the antibody solution. Incubations are usually carried out in heat-sealed plastic bags or hybridization bottles to minimize the volume necessary to completely cover the blot. A volume of 5 to 10 ml of diluted antibody is sufficient to cover most blots. Appropriate antibody concentrations vary for each antibody and must be determined empirically. Identification of Protein Interactions by Far Western Analysis
6. Wash for 10 min in ≥200 ml 1× TBST by agitating on an orbital shaker. Repeat an additional two times.
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7. Dilute the AP-conjugated secondary antibody in 5 to 10 ml of 5% milk in 1× TBST and incubate blot for 1 hr as in step 4. Suppliers generally provide an estimate of appropriate dilution for the secondary antibody.
8. Wash blot six times for 5 min each in ≥200 ml TBST, with agitation. Detect antibodies 9. Briefly, rinse blot in 50 ml alkaline phosphatase buffer. 10. Incubate blot in 20 ml developing solution for 1 to 15 min and rinse blot with 100 ml water. 11. Wash blot for 5 min with 100 ml of 100 mM EDTA, pH 8.0, to stop the development reaction. Rinse with 100 ml water, dry, and photograph.
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An example of a far western blot of proteins is shown in Figure 19.7.1. Lane 1 shows a Coomassie blue stain of a protein sample enriched for histone proteins separated on a 22% SDS-PAGE gel. Lane 2 shows a far western autoradiogram of a parallel lane probed with the yeast Tup1 protein according to this protocol. Lane 3 shows a parallel lane probed with an unlabeled Tup1 protein and detected with antibody specific to Tup1 as in Alternate Protocol 2. Both protocols yield the same result—Tup1p interacts with H3 and H4 but not with H2A or H2B. Lanes 4 and 5 show immunoblots of parallel lanes using anti-H3 and anti-H4 antibodies to identify histories H3 and H4 unequivocally.
H3 H2B H2A H3* H4
3
4
5
Figure 19.7.1 Far western of blotted SDS-PAGE gel. Lane 1, Coomassie blue–stained gel showing locations of histone bands. Lane 2, far western of a parallel lane using radiolabeled in vitro–translated probe. Lane 3, far western using unlabeled probe detected with probe-specific antibody. Lane 4, western blot using anti-histone H3 specific antibody. Lane 5, western blot using anti-histone H4 specific antibody. Identification of Protein Interactions
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ALTERNATE PROTOCOL 2
USING PEPTIDES TO IDENTIFY SPECIFIC INTERACTING SEQUENCES IN A FAR WESTERN BLOT Synthetic peptides can also be used in far western analyses. The use of peptides enables the identification of specific interacting sequences. Specific post-translational modifications can be examined for their effect on protein-protein interactions. Peptides as small as 11 amino acids have been used successfully as far western targets. Peptide far westerns differ from other far westerns only in the preparation of the blots. Peptide dilutions are prepared, then dot or slot blotted onto the support membrane. Blocking and probing of peptide blots are identical to procedures used for traditional far westerns. Duplicate blots are stained to verify that comparable quantities of different peptides have been loaded. Because Ponceau S staining is temporary, staining of duplicate blots with India ink is used to provide a permanent record for peptide blots. In the authors’ experience, only peptides that have been synthesized on MAP resins have worked well for peptide blots. MAP resins consist of branched lysine chains whose chemically active groups have been blocked. Although peptides prepared in other ways do give reproducible results, the peptide concentrations required are several orders of magnitude higher than those required for MAP peptides, making these blots very costly to perform. The reason why increased peptide is needed is unclear, but perhaps the MAP resin “presents” the peptide in such a way that it is more accessible for interaction. Additional Materials (also see Basic Protocol) Peptides 0.4% Tween 20/PBS (see recipe) India ink solution (see recipe) Slot or dot blot apparatus (e.g., Bio-Rad Bio-Dot SF or Schleicher & Schuell Minifold II) 1. Make dilutions of peptides between 5 ng and 5 µg in a final volume of 100 to 200 µl of distilled water. 2. Prepare slot or dot blotter and support membrane (PVDF or nitrocellulose) as described by the manufacturer. Load the peptide dilutions into wells. Prepare duplicate blots, one for far western and one for India ink staining. After all samples are loaded, apply vacuum to draw the peptide samples through the manifold device and onto the support membrane. 3. Block, bind, wash, and autoradiograph one blot for far western (see Basic Protocol, steps 7 to 16). 4. Incubate the second blot in 100 ml of 0.4% Tween 20/PBS for 5 min at room temperature with gentle agitation. Repeat incubation. 5. Stain blot by incubating 15 min to overnight with 100 ml India ink solution at room temperature. 6. Wash the filter for 2 hr in 4 changes of 1× PBS. Dry and store the membrane. This stain is permanent.
Identification of Protein Interactions by Far Western Analysis
Figure 19.7.2 shows an example of results from this alternate protocol, using peptides as a substrate for a far western. The right-hand panel is a blot stained with India ink verifying that comparable quantities of peptide were loaded on the blot. The left-hand panel is a far western of a duplicate blot demonstrating the effect of acetylation of lysine residues of histone peptides on Tup1p/histone interaction.
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5 µg 1 µg 0.2 µg 0.04 µg 0.008 µg far western blot
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Figure 19.7.2 Far western blot of peptides (A) dot blotted onto PVDF membrane and (B) stained with India ink. Figure reproduced with permission of Cold Spring Harbor Laboratory Press.
REAGENTS AND SOLUTIONS Use deionized or distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Alkaline phosphatase buffer 100 ml 1 M Tris⋅Cl, pH 9.5 (APPENDIX 2E) 20 ml 5 M NaCl (APPENDIX 2E) 5 ml 1 M MgCl2 (APPENDIX 2E) Add H2O to 1 liter Store up to 1 year at room temperature 5-Bromo-4-chloro-3-indolyl phosphate (BCIP) stock solution Dissolve 0.5 g of BCIP in 10 ml of 100% dimethylformamide. Store at 4°C or in small aliquots at −20°C. Discard when solution turns color. Developing solution Add 66 µl of NBT stock (see recipe) to 10 ml of alkaline phosphatase buffer (see recipe). Mix well. Add 33 µl of BCIP stock solution (see recipe) and mix again. Prepare fresh. India ink solution Add 100 µl India ink (Pelikan or Higgans) to 100 ml 0.4% Tween 20/PBS (see recipe). Prepare fresh. Nitroblue tetrazolium chloride (NBT) stock solution Dissolve 0.5 g of NBT in 10 ml of 70% dimethylformamide. Store at 4°C or in small aliquots up to 6 months at −20°C. Phosphate-buffered saline (PBS), 10× 80 g NaCl 2.2 g KCl 9.9 g Na2HPO4 2.0 g K2HPO4 Add H2O to 1 liter
Identification of Protein Interactions continued
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Adjust pH of the 10× solution to 7.4 Store indefinitely at room temperature Prior to use, dilute to 1× by mixing 1 part 10× PBS with 9 parts water Leftover 1× PBS should be stored at 4°C to discourage bacterial growth.
Ponceau S staining solution, 10× 2 g Ponceau S 30 g trichloroacetic acid 30 g sulfosalicylic acid Add H2O to 100 ml Store indefinitely at room temperature Just prior to use dilute to 1× by mixing 1 part 10× Ponceau S with 9 parts water Probe dilution buffer, 10× 3.0 g bovine serum albumin (BSA) 10 ml normal goat serum 10 ml 10× PBS (see recipe) H2O to 100 ml Store indefinitely at −20°C Just prior to use, dilute to 1× by mixing 1 part 10× stock with 9 parts 1× PBS Probe purification buffer 400 µl 1 M HEPES, pH 7.4 400 µl 1 M dithiothreitol (DTT) 9.2 ml H2O Prepare fresh TBST, 10× 90 g NaCl 100 ml 1 M Tris⋅Cl, pH 7.5 (APPENDIX 2E) 10 g Tween 20 Add H2O to 1 liter For 1× TBST dilute 1 part 10× TBST with 9 parts water prior to use Store indefinitely at room temperature Tween 20/PBS, 0.4% (w/v) Dissolve 0.4 g Tween 20 in 100 ml 1× PBS (see recipe). Store up to 1 week at room temperature. COMMENTARY Background Information
Identification of Protein Interactions by Far Western Analysis
The far western blot (also called a west western and a ligand blot) has been widely used to examine the interactions of many diverse proteins. For example, it has been used to examine the interactions between the subunits of eukaryotic initiation factors (Kimball et al., 1998), to look at interactions between basic helix-loop-helix DNA-binding proteins (Chaudhary et al., 1997), and to examine the interactions of keratin intermediate filaments with desmosomal proteins (Kouklis et al., 1994). Far westerns have been particularly useful in examining interactions of histones with regulatory proteins. Far westerns have been
used to look at interactions of WD repeat proteins with histones (Edmondson et al., 1996; Palaparti et al., 1997), the interaction of Epstein-Barr virus nuclear antigen 2 with histone H1 (Grasser et al., 1993), and the interaction of histones with the Xenopus oocyte protein N1 (Kleinschmidt and Seiter, 1988). In addition, far westerns have been used to study receptorligand interactions and to screen libraries for interacting proteins (Grulich-Henn et al., 1998; Hsiao and Chang, 1999). Sometimes, the nature of the proteins being examined is such that standard methods of studying protein-protein interactions are not possible. For example, some proteins are diffi-
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cult to solubilize or to extract from cells except under conditions that disrupt protein-protein interactions and therefore, are difficult to assay by immunoprecipitation. Other proteins cannot be expressed in bacteria or yeast due to toxicity problems, thus making the production of recombinant proteins or the use of two-hybrid assays impossible. Far westerns are particularly useful in such cases. Since far westerns are performed totally in vitro, they circumvent these types of problems. Another advantage of the far western blot is its flexibility. Proteins prepared in a variety of ways can be used for the assay. Cell extracts, recombinant proteins, and peptides can all be used as both probe and target proteins. For example, Palaparti et al. (1997) used cell extracts to probe a semipurified histone sample and detected bound proteins of interest with antibodies. Hsiao and Chang (1999) used phage-expressed proteins immobilized on filter lifts for a far western library screen. Unpurified E. coli extracts containing recombinant protein have been successfully used as probes (Fischer et al., 1997). In addition, many different detection techniques, either radioactive or nonradioactive, can be used. Kimball and co-workers used recombinant protein probes that were labeled radioactively with kinases and recombinant proteins labeled with biotin and subsequently detected with a strepavidin-biotin detection scheme (Grulich-Henn et al., 1998; Kimball et al., 1998). Thus, techniques and reagents already in hand can frequently be adapted for use with this assay. Finally, the far western can be modified to define the protein domains and amino acid residues that are important in protein-protein interactions. Mutagenized clones can be used to produce variants of protein probes. A single SDS-PAGE gel can be run with identical lanes and cut into strips, and a different in vitro–translated probe can be used for each strip. In this way, multiple variants of a protein can be tested simultaneously for their ability to interact with a target protein. Non-SDS polyacrylamide gels can also be used to separate proteins for far westerns. For example, acid urea gels, which separate on the basis of both size and charge, have been successfully employed (Edmondson et al., 1996). Finally, peptides corresponding to specific interacting sequences can be synthesized with specific post-translational modifications to test their effects on protein-protein interactions.
Critical Parameters Blocking nonspecific binding sites on the membranes is critical to achieving good results with far westerns. Too little blocking results in high background, while extended time in blocking solutions results in weakened or lost signal. The reason for the diminished signal is unclear, but protein renaturation apparently takes place during the blocking step, so an optimal renaturation may require limited blocking. The best time may well be different for each protein and require empirical optimization. In addition, different lots of BSA appear to result in diminished signal. Therefore, it is important to purchase high-quality BSA from a reputable manufacturer. An important consideration is the inclusion of appropriate controls to rule out nonspecific interactions that might result in false positives. Suitable controls should be furnished for both the target proteins and the protein probe. When using a complex mixture of proteins, such as cell extracts, as target, “negative” control proteins are already present. However, when using a mixture of only a few proteins, it is important to provide a protein that does not interact to serve as a negative control for nonspecific binding. The ideal negative control should be similar to the protein of interest in charge and size. Appropriate controls should be subjected to SDS-PAGE and blotted in parallel with the samples of interest. Another important control is the use of an unprogrammed translation lysate as a probe. Translation of an unrelated protein as a control probe is also often helpful.
Troubleshooting Precise conditions for far westerns vary from procedure to procedure and probably reflect the nature of the individual proteins being examined. Optimal conditions for each protein may need to be determined empirically. If background staining is too high, there are several possible remedies. The probe may be diluted or the sample concentration lowered. Other “blocking” reagents may be tested.The blocking reagent, nonfat dry milk, ranging in concentration from 1% to 5%, has been used successfully. Other detergents such as NP-40 and Triton X-100 are commonly employed for this procedure and may help to decrease background. Also, increasing the length of the blocking step may aid in background reduction. Most procedures that call for extended blocking times suggest incubation at 4°C. If no specific staining is observed, confirm the quality of the radiolabeled probe by SDS-
Identification of Protein Interactions
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PAGE and autoradiography. Make sure the reticulocyte lysate has not been repeatedly frozen and thawed. If using a recombinant protein/antibody scheme, confirm the affinity of that interaction by immunoblotting. Sometimes a decrease in blocking time or use of a different blocking reagent will increase signal. Some proteins may interact more readily following a denaturation/renaturation cycle. In this case, the membrane is incubated for 1 hr in PBS-buffered 7 M guanidine or 8 M urea and renatured overnight. Renature in blocking buffer I (see Basic Protocol) without agitation at 4°C with several changes of buffer. Finally, the length of the binding reaction can be increased.
Anticipated Results Sensitivity of the far western blot is dependent on the affinity of the protein-protein interactions being investigated and on the quality of the probe. Thus, extra attention given to the preparation of a high-quality probe is almost invariably worthwhile. When using a radiolabeled probe, a positive interaction can typically be visualized after overnight exposure to X-ray film.
Time Considerations The basic protocol can be performed in 2 days. An SDS-PAGE gel can be set up and run in 4 to 6 hr. The rest of the procedure can be performed in 8 to 10 hr. It is often convenient to set up the SDS-PAGE gels on one day and run them slowly overnight. The rest of the Basic Protocol can then be performed the next day. Semidry transfer requires ∼2 hr, the blocking steps take ∼4 hr, binding of the probe takes ∼2 hr, washing, drying, and setting up the autoradiography cassette takes ∼1 hr. Detection of interactions using antibodies to detect the probe protein requires an additional day. Peptide blots can be performed in one day. It is possible to store the blots after the blocking steps for up to 2 weeks at 4°C. The blots should be stored in airtight containers or wrappings so they do not dry out. In addition, in vitro–translated probes may be produced ahead of time and stored unpurified at −20°C, although the efficiency may be reduced for some probes.
Literature Cited Chaudhary, J., Cupp, A.S., and Skinner, M.K. 1997. Role of basic-helix-loop-helix transcription factors in Sertoli cell differentiation: Identification of an E-box response element in the transferrin promoter. Endocrinology 138:667-675. Edmondson, D.G., Smith, M.M., and Roth, S.Y. 1996. Repression domain of the yeast global repressor Tup1 interacts directly with histones H3 and H4. Genes & Dev. 10:1247-1259. Fischer, N., Kremmer, E., Lautscham, G., MuellerLantzsch, N., and Grasser, F.A. 1997. EpsteinBarr virus nuclear antigen 1 forms a complex with the nuclear transporter karyopherin alpha2. J. Biol. Chem. 272:3999-4005. Grasser, F.A., Sauder, C., Haiss, P., Hille, A., Konig, S., Gottel, S., Kremmer, E., Leinenbach, H.P., Zeppezauer, M., and Mueller-Lantzsch, N. 1993. Immunological detection of proteins associated with the Epstein-Barr virus nuclear antigen 2A. Virology 195:550-560. Grulich-Henn, J., Spiess, S., Heinrich, U., Schonberg, D., and Bettendorf, M. 1998. Ligand blot analysis of insulin-like growth factor-binding proteins using biotinylated insulin-like growth factor-I. Horm. Res. 49:1-7. Hsiao, P.W. and Chang, C. 1999. Isolation and characterization of ARA160 as the first androgen receptor N-terminal-associated coactivator in human prostate cells. J. Biol. Chem. 274:2237322379. Kimball, S.R., Heinzinger, N.K., Horetsky, R.L., and Jefferson, L.S. 1998. Identification of interprotein interactions between the subunits of eukaryotic initiation factors eIF2 and eIF2B. J. Biol. Chem. 273:3039-3044. Kleinschmidt, J.A. and Seiter, A. 1988. Identification of domains involved in nuclear uptake and histone binding of protein N1 of Xenopus laevis. EMBO J. 7:1605-1614. Kouklis, P.D., Hutton, E., and Fuchs, E. 1994. Making a connection: Direct binding between keratin intermediate filaments and desmosomal proteins. J. Cell Biol. 127:1049-1060. Palaparti, A., Baratz, A., and Stifani, S. 1997. The Groucho/transducin-like enhancer of split transcriptional repressors interact with the genetically defined amino-terminal silencing domain of histone H3. J. Biol. Chem. 272:26604-26610.
Contributed by Diane G. Edmondson and Sharon Y. R. Dent M.D. Anderson Cancer Center Houston, Texas
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Scintillation Proximity Assay (SPA) Technology to Study Biomolecular Interactions
UNIT 19.8
Scintillation proximity assay (SPA) is a versatile homogeneous technique for radioactive assays which eliminates the need for separation steps. It has been applied to radioimmunoassays (RIA), receptor-binding assays, enzyme assays, and molecular-interaction assays, such as protein/protein-binding studies (Cook, 1996). When a radioactive atom decays, it releases subatomic particles (e.g., β-particles), and depending upon the isotope, various forms of electromagnetic radiation (e.g., γ-rays). The distance these particles travel through aqueous solutions is limited and is dependent upon the energy of the particle; SPA exploits this limitation. For example, when a tritium atom (3H) decays it releases a β-particle. If the 3H atom is within 1.5 µm of a suitable scintillant molecule in aqueous solution, the energy of the β-particle will be sufficient to reach the scintillant and excite it to emit light. If the distance between the scintillant and the 3H atom is greater than 1.5 µm, then the β-particles will not have sufficient energy to travel the required distance (see Figure 19.8.1). In an aqueous solution, collisions with water molecules dissipate the β-particle energy and it therefore cannot stimulate the scintillant. Normally, the addition of a scintillation cocktail to radioactive samples ensures that the majority of 3H emissions are captured and converted to light. In SPA, the scintillant is incorporated into small fluomicrospheres. These microspheres or “beads” are constructed in such a way as to bind specific molecules. SPA beads have been developed from inorganic scintillants such as yttrium silicate (YSi) and hydrophobic polymers such as polyvinyltoluene (PVT). An optimized microsphere has been developed, consisting of a solid scintillant-containing PVT core coated with a polyhydroxy film that renders the bead more compatible with aqueous buffers. Coupling molecules, such as antibodies and streptavidin, are covalently attached to the coating, allowing generic links for assay design. Table 9.18.1 lists the range of coatings available on both PVT and YSi SPA beads. These coupling formats are high-affinity biologicalmolecule-mediated linkages. No complex chemistry is required to achieve attachment of assay components to the SPA particles.
Light
SPA Bead β-particle
β-particle
radiogland
Figure 19.8.1 Diagrammatic representation of SPA (not to scale). On the left, the bound radioligand is in close proximity, stimulating the bead to emit light. On the right, the unbound radioligand is not in sufficient proximity to stimulate light.
Identification of Protein Interaction
Contributed by Neil Cook, Alison Harris, Alison Hopkins, and Kelvin Hughes
19.8.1
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Table 19.8.1
Range of SPA Bead Types
Bead coating
Comments
Anti-mouse (PVT and YSi)
Used for radioimmunoassays, receptor binding assays, and enzyme assays in conjunction with an appropriate antibody. Anti-rabbit (PVT and YSi) Used for radioimmunoassays, receptor binding assays, and enzyme assays in conjunction with an appropriate antibody. Anti-sheep (PVT and YSi) Used for radioimmunoassays, receptor binding assays, and enzyme assays in conjunction with an appropriate antibody. Copper His-tag (PVT and YSi) Used to capture His-tagged proteins. Glutathione (PVT and YSi) Binds glutathione-S-transferase (GST) tagged proteins. Polyethylenimine (PEI) Wheat germ Binds membranes. Used in receptor agglutinin (PVT only) binding assays. Poly-L-lysine (PL) (YSi only) Binds negatively charged cell membranes. Protein A (PVT and YSi) Used for radioimmunoassays, receptor binding assays, and enzyme assays in conjunction with an appropriate antibody. Streptavidin (PVT and YSi) Used to capture biotinylated components. Can be used for receptor assays in conjunction with biotinylated lectins. Wheat germ agglutinin (WGA; PVT and YSi) Binds glycosylated proteins and membrane fractions for receptor binding assays.
If a radioactive molecule is bound to the bead, it is brought into close enough proximity that it can stimulate the scintillant contained within to emit light. Otherwise, the unbound radioactivity is too distant, the energy released is dissipated before reaching the bead, and these disintegrations are not detected. Several isotopes are suitable for SPA. Tritium is ideally suited for SPA as its β-particle has an extremely short path length of only 1.5 µm through aqueous solutions. This means that the background obtained from unbound tritium molecules is normally low, even when relatively large amounts of activity are used. 125I is another isotope which displays excellent properties for use in SPA. One way in which the 125I atom decays is by a process termed “electron capture.” This type of decay gives rise to two electrons which may be detected by SPA. These electrons have path lengths of ∼1 and 17.5 µm.
SPA Technology to Study Biomolecular Interactions
More energetic β-emitters such as 33P and 35S can also be used with SPA (DeLapp et al, 1999; Beveridge et al., 2000; Slater, 2001). The path lengths of the electrons emitted from these isotopes are longer in aqueous medium, which can result in stimulation of beads even by unbound radioisotope. This is termed “nonproximity effect” or NPE. In addition, bound isotope emissions will pass through the bead, resulting in a less efficient harvesting of energy from the bound radioisotope. In these circumstances, in order to discriminate the bound isotope from free, the NPE can be reduced by introducing a relatively simple centrifugation or settling step. If the SPA beads are allowed to settle out of solution, the distance between free radioisotope and beads is increased, such that electrons emitted from free radioisotope can no longer travel the distance required to interact with the scintillant in the bead. In addition, more energetic electrons released by bound radioisotope, although passing through their bead of origin, can now interact with neighboring
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beads. The time required for beads to settle is at most 15 min for the dense YSi beads and 4 to 6 hr for PVT beads. Gentle centrifugation of assay tubes or plates, typically ∼1000 × g for 5 min, can also be employed. In this unit, the application of SPA technology to measuring protein-protein interactions (see Basic Protocol 1), Src Homology 2 (SH2) and Src Homology 3 (SH3) domain binding to specific peptide sequences (see Basic Protocol 2), and receptor-ligand interactions (Basic Protocol 3) are described. Three other basic protocols discuss the application of SPA technology to cell-adhesion-molecule interactions (see Basic Protocol 4), proteinDNA interactions (see Basic Protocol 5), and radioimmunoassays (see Basic Protocol 6). In addition, protocols are given for the preparation of SK-N-MC cells (see Support Protocol 1) and cell membranes (see Support Protocol 2). NOTE: SPA is a patented technology that was initially made available for use in high-throughput screening applications through Technology Transfer/Access programs. Both SPA beads and kits are now fully available. Refer to the current Amersham Pharmacia Biotech catalog or http://amershambiosciences.com. THE APPLICATION OF SPA TECHNOLOGY TO THE MEASUREMENT OF PROTEIN-PROTEIN INTERACTIONS
BASIC PROTOCOL 1
Interactions between proteins are key features of many biochemical processes (e.g., cell signaling); however, in the absence of any enzymatic activity, the measurement of protein-protein interactions has presented problems. SPA technology circumvents this problem by permitting the direct measurement of binding of one protein to another. In addition, in cases where the dissociation rate (kD) of the interaction is high, SPA provides a means of equilibrium counting in an homogeneous assay format. This protocol describes the interaction of the GTPase activating protein, neurofibromin-1 (NF1), to a radiolabeled oncogenic Ras protein (RasL61; Skinner et al., 1994) by SPA. The procedure employed in configuring this assay will be used to illustrate the key features to be considered when designing a protein-protein interaction SPA. See Commentary for steps that need to be modified for different systems. Materials GST-NF1 protein (Eccleston et al., 1993) NF1-Ras SPA buffer (see recipe) RasL61⋅[3H]GTP (Lowe et al., 1991; also see Background Information) Anti-GST antibody (Molecular Probes) 500 mg/vial protein A PVT SPA beads, lyophilized (Amersham Biosciences; Table 19.8.1) Test compound and appropriate solvent Siliconized glass vial (APPENDIX 3E) 96-well microtiter plate and adhesive sealer or assay tubes and caps 22°C water bath or microtiter plate incubator Prepare reagents 1. In a siliconized glass vial, prepare a 2.5 nmol/ml dilution of GST-NF1 protein in NF1-Ras SPA buffer. The catalytic domain of NF1 is expressed as a fusion protein in E. coli and purified by chromatography on glutathione-agarose by elution with 5 mM glutathione in 10 mM Tris buffer, pH 7.5, containing 50 mM NaCl, 0.1 mM EDTA, and 1 mM DTT (Eccleston et al., 1993). All proteins are diluted in siliconized glass vials to reduce losses of proteins by adsorption.
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2. In a second siliconized glass vial, prepare a 1 nmol/ml solution of RasL61⋅[3H]GTP in NF1-Ras SPA buffer. RasL61 is expressed in E. coli and purified as described for other Ras proteins by Lowe et al. (1991). Purification is by chromatography on glutathione-agarose with elution using 5 mM glutathione in 10 mM Tris⋅Cl buffer, pH 7.5, containing 50 mM NaCl, 0.1 mM MgCl2, 50 ìM GDP, and 1 mM DTT. RasL61 is labeled with [3H]GTP as described (see Background Information). The relative amounts of both GST-NF1 and RasL61⋅[3H]GTP proteins are determined in optimization experiments performed to maximize the signal obtained in the assay. See Background Information for the key factors involved in the assay optimization process.
3. Prepare a 75 µg/ml dilution of anti-GST antibody in NF1-Ras SPA buffer. For all assays using an antibody, capture format, the specificity, purity, and affinity of the antibody will greatly influence the amount of antibody, and hence the amount of beads, required for maximal signal. It is essential that the antibody-protein interaction does not interfere with the protein-protein interaction. See Background Information for alternative capture formats.
4. Carefully add 50 ml NF1-Ras SPA buffer to the vial provided by the supplier containing 500 mg lyophilized protein A PVT SPA beads, to produce a 10 mg/ml protein A PVT bead solution and replace the stopper. Mix the beads by gentle inversion and swirl, taking care to avoid foaming. Store reconstituted beads up to 4 weeks at 2° to 8°C. The NF1-Ras SPA format exploits the fact that the NF1 protein is expressed as a fusion protein (i.e., GST-NF1). The GST-NF1 fusion protein is coupled to protein A PVT SPA beads via the commercially available anti-GST antibody. An obvious alternative bead for this type of format would be one of the anti-species antibody–binding beads (e.g., anti-rabbit, -mouse, or -sheep), according to the source of the capture antibody. The specificity, affinity, and purity of the antibody will effect the maximum level of signal achievable and the choice of bead; it is not possible to predict whether the anti-species antibody–binding or protein A beads would perform better. Both should be evaluated as an initial experiment in the development process. This format offers a generic design for the capture of any protein when a suitable antibody exists. Regardless of the bead type used, it is essential that the capture protein is present in excess with respect to the radiolabeled protein, and that beads are present in excess to capture all the complex to maximize the signal obtained (see Background Information).
5. Prepare test compound at appropriate dilution in appropriate solvent. Test compounds are used when screening a panel of drugs or monitoring inhibition (i.e., determining IC50). In general, any unlabeled drugs or compounds for testing should be added in aqueous solution (i.e., NF1-Ras buffer). If samples are insoluble in aqueous solution then they should be dissolved in an organic solvent. In this event, it is advisable to set up a solvent control tube to test whether the solvent alone has any effect on the assay.
Conduct assay Figure 19.8.2 presents a flowchart for optimizing this assay. 6. Label appropriate 96-well microtiter plates or assay tubes in duplicate. Pipet 20 µl NF1-Ras SPA buffer into all test wells or tubes, and 40 µl buffer into all control wells or tubes. SPA Technology to Study Biomolecular Interactions
It is advisable to include control wells in the absence of GST-NF1 fusion protein. Additional controls could include the absence of anti-GST antibody.
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source reagents: recombinant proteins antibodies
assay conditions: buffer system co-factor requirements protease inhibitors time temperature agitation required?
reagent concentrations: NSB NPE
sample preparation color quench correction
design radiolabel and capture mechanism
choice of SPA bead: streptavidin protein A/second antibody
optimize signal:noise
assay protocol: precoupled To delayed addition
validate assay: controls/inhibitors competition curves saturation binding
Figure 19.8.2 Flow chart for the development of SPA protein-protein interaction assays.
7. Pipet 50 µl appropriately diluted test compound into the sample wells or tubes, and 50 µl NF1-Ras SPA buffer into control wells or tubes. 8. Pipet 50 µl appropriate solvent into solvent control wells or tubes, if applicable. This is only required if the samples to be tested are insoluble in aqueous solution.
9. Pipet 20 µl of 2.5 nmol/ml GST-NF1 fusion protein (50 pmol; step 1) into all test wells or tubes. 10. Into every well or tube, add 10 µl of 1 nmol/ml RasL61⋅[3H]GTP (10 pmol; step 2), 50 µl of 75 µg/ml anti-GST antibody (3.75 µg; step 3), and 50 µl of 10 mg/ml protein A PVT SPA beads (0.5 mg; step 4). Ensure the SPA beads are an homogeneous suspension during pipetting by either recapping and shaking the bottle at regular intervals, or by stirring gently with a magnetic stirrer. All the tubes or wells should now contain a total volume of 200 ìl.
11. Cap the tubes or seal the microtiter plate with an adhesive sealer and incubate 1 hr in a 22°C water bath or microtiter plate incubator. 12. Count 1 min per well or tube in a β-scintillation counter. See Background Information section regarding counting instrumentation details and procedures. Identification of Protein Interaction
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BASIC PROTOCOL 2
THE APPLICATION OF SPA TECHNOLOGY TO Src HOMOLOGY 2 (SH2) AND Src HOMOLOGY 3 (SH3) DOMAIN BINDING TO SPECIFIC PEPTIDE SEQUENCES SH2 and SH3 domains are small independent domains of ∼70 to 120 amino acid residues. They are found in a variety of proteins, and can occur together or separately. SH2 domains are thought to be involved in signal transduction mechanisms (Cohen et al., 1995). This unit describes the interaction of the tandem SH2 domains within the growth factor receptor binding-2 (Grb2) protein, to a selected radiolabeled phosphopeptide (Pai et al., 1999) by SPA. The procedure employed in configuring this assay will be used to illustrate the key features to be considered when designing a SH2/SH3 interaction SPA. Materials Grb2-SH2/SH3 SPA buffer (see recipe) Biotin/streptavidin capture buffer (see recipe) 500 mg/vial freeze-dried protein A or streptavidin PVT SPA beads (Amersham Biosciences) Test compound and appropriate solvent (e.g., DMSO; optional) GST-SH2/SH3 fusion protein: express in E. coli, purify by chromatography on glutathione-Sepharose 4B, and filter centrifuge using a Centriprep 10 microconcentration device (Amicon) Anti-GST antibody (Molecular Probes) 96-well microtiter plates with adhesive sealer or assay tubes with caps Additional reagents and equipment for synthesizing peptides by solid-phase Fmoc chemistry (UNIT 18.5), radiolabeling peptides using either [125I]Bolton and Hunter reagent (UNIT 3.3), succinimidyl-N-[propionate-2,3-3H] (Tang et al., 1983), or catalytic reduction under tritium gas (Evans, 1966), and purifying peptides by reversed-phase HPLC (UNIT 11.6) NOTE: This example is an optimized protocol containing amounts of the various assay components that give an appropriate counting window. In order to optimize each component, assays should be performed by varying the final concentration of the component under investigation, while keeping the concentration of the others fixed. For example, Figure 19.8.3 shows the effect of varying the amount of the SPA beads in a Grb2 SH2 assay. Synthesize and radiolabel peptides 1. Synthesize peptides using standard methods of solid-phase peptide Fmoc chemistry (UNIT 18.5), and then radiolabel using either [125I]Bolton and Hunter reagent (UNIT 3.3), succinimidyl-N-[propionate-2,3-3H] (Tang et al., 1983), or catalytic reduction under tritium gas (Evans, 1966). If necessary, subsequently purify peptides by reversedphase HPLC (UNIT 11.6). Phosphopeptide ligands employed in such applications are designed to contain essential elements of SH2/SH3 domain recognition motifs. Either 3H- or 125I-labeled peptides can be used for measuring SH2/SH3 interactions by SPA. The signal and background are significantly lower for 3H compared to 125I. Ultimately, the choice of label will be determined by a combination of signal, signal-to-noise ratio required, acceptable half-life, and whether there is any steric hindrance of binding caused by labeling the peptide of choice. (see Pai et al., 1999 for details of the design and radiolabeling of the Grb2 SH2 phosphopeptide).
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12 NSB B0
SPA (cpm × 10 − 3 )
10
8
6 4
2
0 0
1
2 3 4 Streptavidin-coated SPA bead (mg)
5
6
Figure 19.8.3 The effect of varying the bead concentration on signal (Bo) and NSB in the GRB2 assay. Assays were performed using 20 mM MOPS buffer, pH 7.5, containing 0.1% (w/v) BSA, 10 mM DTT, and varying concentrations of streptavidin coated SPA beads. The beads were settled and the plate was counted at 4°C. Results are means (n = 3).
Prepare the SPA bead suspension and reaction 2. Carefully add 250 ml Grb2-SH2/SH3 SPA buffer to the vial provided by the supplier containing 500 mg freeze-dried protein A or 250 ml biotin/streptavidin capture buffer to streptavidin SPA beads (2 mg/ml final), and replace the stopper. Mix the beads by gentle inversion and swirling, taking care to avoid foaming. Store reconstituted beads up to 4 weeks at 2° to 8°C. For the detailed protocol given below, protein A beads are employed.
Conduct assay via antibody capture approach 3. Label appropriate 96-well microtiter plates or assay tubes in duplicate. Pipet 20 µl Grb2-SH2/SH3 SPA or biotin/streptavidin capture buffer into all test wells and 40 µl into all control wells. It is advisable to include control wells in the absence of GST-SH2/SH3 fusion protein. Additional controls could include the absence of anti-GST antibody.
4. Prepare test compound at appropriate dilution in appropriate solvent. In general, any unlabeled drugs/compounds for testing should be added in aqueous solution. If samples are insoluble in aqueous solution then they should be dissolved in an organic solvent. In this event, it is advisable to set up a solvent control tube to test whether the solvent alone has any effect on the assay.
5. Pipet 50 µl appropriately diluted test compound into the sample wells or tubes, or 50 µl Grb2-SH2/SH3 SPA or biotin/streptavidin capture buffer into control wells or tubes. 6. Pipet 50 µl appropriate solvent (e.g., DMSO) into solvent control wells or tubes if applicable. This is only required if the samples to be tested are insoluble in aqueous solution.
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For protein A beads: 7a. Pipet 20 µl (6 pmol) GST-SH2/SH3 fusion protein into all test wells or tubes. 8a. Pipet 10 µl of 0.04 µCi/0.6 pmol [3H]peptide, 50 µl (6 µg/34 pmol) anti-GST antibody, and 50 µl (0.1 mg) protein A PVT SPA beads into every well or tube. Ensure the SPA beads are an homogeneous suspension during pipetting by either recapping and shaking the bottle at regular intervals, or by gently stirring with a magnetic stirrer. Each tube or well should now contain 0.6 pmol (0.04 ìCi) 3H-radiolabeled binding peptide (i.e., [3H]Grb2; step 1), 6 pmol (0.25 ìg) GST-SH2/SH3 domain fusion protein, 34 pmol (6 ìg) anti-GST antibody, and 0.1 mg protein A-coated SPA beads in a suitable buffer (e.g., Grb2-SH2/SH3 SPA buffer) at a final volume of 200 ìl/well to capture the peptide-protein complex. Assay conditions taken from Pai et al. (1999).
For streptavidin beads: 7b. Pipet 20 µl (300 nmol) biotinylated GST-SH2 fusion protein into all test wells or tubes. 8b. Pipet 10 µl of 0.1 µCi pmol [125I]peptide and 100 µl (0.2 mg) of streptavidin SPA beads into every well or tube. Ensure the SPA beads are a homogeneous suspension during pipetting by either recapping and shaking the bottle at regular intervals, or by gently stirring with a magnetic stirrer. Each tube or well should contain 350 pM (0.1 ìCi) radiolabeled binding peptide, ∼300 nM biotinylated GST-SH2/SH3 domain fusion protein, and 200 ìg streptavidin-coated SPA beads in a suitable buffer (e.g., biotin/streptavidin capture buffer) at a final volume of 200 ìl to capture the peptide-SH2/SH3 binding protein complex. See Sheets et al. (1998) for details of a ZAP-70/protein tyrosine kinase SH2 domain assay employing this approach.
9. Cap the tubes or seal the microtiter plate with an adhesive sealer and incubate with shaking for 2 to 8 hr (depending on when equilibrium is reached) at room temperature. 10. Count for 1 min per tube/well in a β-scintillation counter. See Background Information for details of appropriate scintillation counting instrumentation. BASIC PROTOCOL 3
THE APPLICATION OF SPA TECHNOLOGY TO STUDY RECEPTOR-LIGAND INTERACTIONS Traditionally, techniques such as filtration have been used to study the interaction of radiolabeled ligands with cell-surface receptors. How SPA technology can be applied to study receptor-ligand interactions is briefly described here.
SPA Technology to Study Biomolecular Interactions
Receptors from sources ranging from crude tissue preparations to soluble purified proteins can be immobilized directly onto SPA beads via a number of coupling methods (see Table 19.8.1 for different bead types). For instance, WGA PVT beads are often used for coupling cellular membranes prepared from either homogenized tissues or cultured cells. Once immobilized, the receptor is close enough to the bead so that should a suitably radiolabeled ligand bind to the receptor, it will be in close enough proximity to stimulate the bead to emit light (see Figure 19.8.4). Any unbound radioligand is too distant from the bead to transfer energy and remains undetected. This discrimination of binding by proximity means that no labor-intensive separation of bound and free ligand is required, as in traditional methods. In addition, as SPA is a homogeneous system, it is possible to monitor the rate of association or dissociation of a radiolabeled ligand from the receptor, in real time, in a minimum of assay tubes using repeat counting.
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“cold” ligand or drug
radioligand
Figure 19.8.4 Diagrammatic representation of an SPA receptor binding assay (not to scale). Immobilized receptors on SPA beads are stimulated to emit light when a radioligand binds to the receptor. Any unbound radioligand is too distant from the bead to generate a signal.
In this protocol a competitive binding assay for the Y1-type neuropeptide Y-peptide YY receptor by SPA is described. Competition experiments are routinely used to analyze the pharmacological nature of the radioactively-labeled receptor and determine the binding characteristics of unlabeled drugs. The assay described below utilizes cell preparations from a human neuroblastoma cell line, which expresses Y1 receptors. Materials YY SPA buffer, 2° to 8°C (see recipe) 500 mg/vial lyophilized WGA PVT SPA beads (Amersham Biosciences) [125I]Tyr-labeled peptide YY (Amersham Biosciences) 500 µg/ml unlabeled porcine peptide YY (Bachem): prepare as recommended by the manufacturer; store up to at least two weeks at −15° to −30°C Test compound and appropriate solvent SK-N-MC cell (see Support Protocol 1) or membrane preparation (see Support Protocol 2) 96-well microtiter plate with adhesive seal or assay tubes with caps NOTE: SPA is an homogeneous technique, and as such it is important that the assay be allowed to reach equilibrium before counting is performed. This normally requires a more prolonged incubation period than traditional filtration techniques. As the assay incubation is longer, it is therefore essential to ensure that the assay components are themselves stable. This may involve the addition of stabilizing agents or protease inhibitors to protect the ligand and/or receptor from the effect of degrading enzymes present in the receptor preparation. In addition, certain other cofactors such as divalent cations may be required for the binding of the radioligand to its receptor. Prepare reagents 1. Carefully add 25 ml cold YY SPA buffer to the vial containing 500 mg lyophilized WGA PVT beads (20 mg/ml final), as provided by the supplier, and replace stopper. Mix the beads by gentle inversion and swirling, taking care to avoid foaming. Store reconstituted beads up to 4 weeks at 2° to 8°C. There are a variety of bead types available for SPA receptor binding assays (see Table 19.8.1) and the choice is dependent on both the receptor source and the radioligand. WGA PVT beads have been widely used for receptor assays. WGA PVT beads bind to N-acetylβ-D-glucosamine residues in membranes and select for membrane glycolipids and glycoproteins (Nagata and Burger, 1974). The binding capacity of these beads is in the range of
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10 to 40 ìg membrane protein per milligram beads; however, if a glycosylated ligand is used then high nonspecific binding (NSB) may be observed due to binding of the ligand to WGA PVT beads. Also, some ligands may interact with the PVT core, in which case poly-L-lysine (PL) YSi beads may be more appropriate. Here, there is an electrostatic interaction between negatively charged membranes and the positively charged beads (Kalish et al., 1978). A different approach was used by Zhuang et al. (1999). In this case, streptavidin beads were used to capture a membrane preparation from a baculovirus-infected insect cell line (Spodoptera frugiperda Sf9) expressing human interleukin-5 receptors using a variety of biotinylated lectins, many of which are now available commercially (Sigma). Interestingly, they found that some of the biotinylated lectins (e.g., Wisteria floribunda lectin) were more effective than WGA at capturing insect cell membranes (Zhang et al., 1999). Regardless of the bead type used, the amount of bead added should be carefully optimized during the assay development process in order to maximize the signal obtained (see Background Information).
2. Reconstitute 25 µCi (925 kBq) [125I]Tyr-labeled peptide YY in 0.5 ml distilled water. Dilute an aliquot of the 50 µCi/ml solution in YY SPA buffer to give the required volume for the assay such that a 50-µl aliquot contains 40,000 cpm (γ counts). Maintain at 2° to 8°C. Aliquot and store any remaining 50 µCi/ml stock solution up to 4 weeks at −15° to −30°C. It is recommended that silanized glass tubes be used to store the frozen stock solution to ensure good recovery when thawed. The amount of ligand added is dependent on the affinity of the ligand for the receptor but is normally at a concentration less than KD.
3. Prepare a 0.4 nmol/ml solution of unlabeled porcine peptide YY from a 500 µg/ml stock and maintain at 2° to 8°C. Prepare a range of standard dilutions from the 0.4 nmol/ml stock to add to the assay tubes to produce the competition binding curve. The full range of concentrations required should be determined by the researcher. As a guideline use 0.0012 to 10 nM. Standards should be freshly prepared before each assay and not reused. NSB tubes are set up in the assay by adding 50 ìl of the 0.4 nmol/ml stock solution directly to the assay, to give a final concentration of 100 nM in the assay tube.
4. Prepare test compounds. In general, any unlabeled drugs/compounds for testing should be added in aqueous solution. If samples are insoluble in aqueous solution then they should be dissolved in an organic solvent. In this event, an organic solvent should be used to dissolve the compound and it is advisable to set up a solvent control tube, to test whether the solvent alone has any effect on the assay.
5. Thaw the required amount of SK-N-MC cell or appropriate membrane preparation (see Support Protocol 1 or 2) and maintain at 2° to 8°C. Dilute cells 1:4 with cold YY SPA buffer. Mix gently and maintain at 2° to 8°C during assay preparation.
SPA Technology to Study Biomolecular Interactions
The amount of SK-N-MC cell preparation added to the assay should be carefully optimized, so that all membrane protein present is bound by SPA beads in the assay. This is vital to produce an optimum signal in the assay, as there may be a reduction in the signal obtained if there is an excess of receptor protein present that is not bound to the SPA beads. The optimization of protein/bead ratios is a critical factor in the assay development process for SPA and will be discussed further (see Background Information; also see Critical Parameters). If a cell line is selected as the receptor source, the density of receptors is important particularly if the ligand is labeled with 3H and is of low specific activity (i.e., 20 to 80 Ci/mmol). For 3H-labeled ligands, expression levels in the region of 500,000 receptors per cell are often required, corresponding to receptor densities greater than 2 pmol/mg membrane protein; however, in this example using an [125I]ligand (∼2,000 Ci/mmol), lower receptor densities are acceptable, in the region of 200 fmol/mg membrane protein (50,000 receptors per cell).
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receptor preparation grow and harvest cells and prepare membranes (see SUPPORT PROTOCOLS 1 AND 2 )
prepare reagents and label tubes/assay plate
pipet 50 µl assay buffer into B 0 wells
pipet 50 µl 0.4 nmol/ml peptide YY into NSB wells
pipet 50 µl standard dilution peptide YY into appropriate wells
pipet 50 µl [ 125 Ι] peptide YY into all wells
pipet 50 µl diluted cell preparation into all wells
pipet 50 µl of SPA beads into all wells
cap tubes/seal plate and shake at room temperature overnight
leave tubes to stand for 1 hr then count for 1 min/well in a β-scintillation counter (see Background Information )
Figure 19.8.5 Flow chart for the development of SPA receptor binding assays.
Conduct assay Figure 19.8.5 presents a flowchart outlining the assay. 6. Label appropriate 96-well microtiter plates or assay tubes in duplicate. Pipet 50 µl YY SPA buffer into the B0 (zero standard) wells or tubes and 50 µl of 0.4 nmol/ml unlabeled peptide YY into the NSB wells or tubes. Pipet 50 µl each standard dilution of peptide YY into appropriately labeled wells or tubes. Replicate tubes/wells should be used for each data point. B0 is equal to the total counts bound minus nonspecific binding.
7. Pipet 50 µl appropriate solvent into solvent control tubes if applicable. This is only required if the samples to be tested are insoluble in aqueous solution.
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8. Pipet 50 µl [125I]Tyr-labeled peptide YY solution, 50 µl diluted cell preparation, and 50 µl WGA SPA beads into all tubes. Ensure WGA SPA beads are an homogeneous suspension by mixing while pipetting into the assay tubes, by either recapping and shaking the bottle at regular intervals, or by stirring gently with a magnetic stirrer. All the tubes or wells should now contain a total volume of 200 ìl.
9. Cap the tubes or seal the microtiter plate with an adhesive seal and shake at room temperature (15° to 30°C) overnight (16 to 24 hr) on a suitable shaker. Shaking should be sufficient to maintain the assay components in suspension. There are a variety of suitable plate/tube shakers available including the DPC MicroMix. Time-course experiments should be performed to establish both the incubation time required for the attainment of equilibrium, and the stability of the SPA signal at equilibrium. The incubation time and signal stability will be dependent on the receptor and radiolabeled ligand.
10. Incubate the tubes or microtiter plates for 1 hr without shaking at room temperature. 11. Count 1 min/tube or well in a β-scintillation counter. See Background Information for details of appropriate scintillation counting instrumentation.
12. Process the assay data collected as follows. a. Calculate the average counts per minute (cpm) or dpm for each set of replicate tubes. b. Calculate the normalized percent bound (NPB) for each standard using the following equation: NPB =
B standard − NSB × 100 (%) = B0 B0 − NSB
NPY PYY
100
B/B 0 (%)
[Leu31Pro34]NPY NPY (13-36) 75
50
25
0 –13
–12
–11
–10
–9
–8
–7
–6
–5
Log10 [competitor] M
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Figure 19.8.6 Competition binding curve showing the inhibition of [125I]Tyr-labeled PYY binding by unlabeled NPY(squares), PYY(upright triangles), [Leu31Pro34]NPY (upside-down triangles), and NPY(13-36) (diamonds), expressed as percent B/B0. Results are means ± SEM (n = 3). Assay was performed using the conditions outlined (see Basic Protocol 3). Current Protocols in Protein Science
where B0 = zero standard, NSB = nonspecific binding, and all measurements are in cpm. This equation can be used for any SPA binding assay (e.g., receptor binding) where an expression of normalized (as opposed to absolute) results is required for comparison.
c. Generate a standard competition curve by plotting the normalized percent bound as a function of the peptide YY concentration. (See Figure 19.8.6.). SK-N-MC CELL CULTURE AND CELL HARVESTING This protocol offers a reliable method for culturing and harvesting SK-N-MC cells, a stable human neuroblastoma cell line expressing Y1 type receptors (Fuhlendorff et al., 1990). This SK-N-MC cell preparation may be stored at −70°C prior to use in the assay. Before freezing, add 5% (v/v) dimethyl sulfoxide (DMSO) and dispense into suitablesized aliquots. Under these conditions frozen cells should remain stable for at least 12 weeks.
SUPPORT PROTOCOL 1
Materials SK-N-MC human neuroblastoma cells (ATCC# HTB-10): store as recommended by ATCC Complete Dulbecco’s modified Eagle medium containing 10% (v/v) FBS (DMEM-10; e.g., Sigma; also see APPENDIX 3C) Dulbecco’s PBS, sterile (Life Technologies) Trypsin/EDTA (Life Technologies) Harvesting buffer, cold (see recipe) Cell scraper 168-cm2 flasks 50-ml centrifuge tube NOTE: SK-N-MC cells do not grow well when cultured in roller bottles. The use of large tissue culture flasks is therefore recommended so that the cells remain permanently immersed in culture medium. Routine subculture 1. On thawing, culture SK-N-MC human neuroblastoma cells in complete DMEM-10 per ATCC guidelines. 2. To subculture the cell lines, remove the culture medium and gently wash the cell monolayer with sterile Dulbecco’s PBS. 3. To the flask containing the cells, add 1 ml trypsin/EDTA per 25 cm2 to ensure that the surface is covered, then remove most of the solution leaving just enough to wet the cells. Incubate at room temperature until the cells begin to detach. Detachment occurs within 3 to 6 min for SK-N-MC cells. The time required for other cell lines should be determined empirically.
4. Add fresh culture medium at 37°C, and dispense into new flasks at a subcultivation ratio of 1:6 (recommended for SK-N-MC). Change the media every 2 to 3 days. As a guide, fifteen 168-cm2 flasks should provide sufficient cells for 2500 assay tubes. At this stage, cell density and growth can be monitored if required. See APPENDIX 4B for details.
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Cell harvesting 5. Harvest cells when confluent. The maximum cell and receptor yield is obtained if the cells are harvested when confluent, with fresh medium added 24 hr prior to harvesting.
6. Wash the confluent cell monolayers with ∼20 ml cold Dulbecco’s PBS per 168 cm2 flask. Remove PBS. 7. Add 5 ml cold harvesting buffer and gently detach the cells using a cell scraper. Aspirate the cell suspension and transfer to a 50-ml centrifuge tube, which should be kept on ice. 8. Wash the culture flask with a further 5 ml cold harvesting buffer. Add to the centrifuge tube. Repeat this process with each flask until all cells have been removed. 9. Centrifuge the cells 10 min at 600 × g, 4°C. Discard the supernatant and resuspend the cell pellet in 2.5 ml harvesting buffer/flask. To store, add 5% (v/v) DMSO, disperse into aliquots, and freeze at −70°C. Frozen cells should remain stable at least 12 weeks. SUPPORT PROTOCOL 2
PREPARATION OF CELL MEMBRANES The method described above for SK-N-MC cells (see Basic Protocol 3), using whole cells removed from culture flasks, is a very convenient route to develop receptor-binding assays; however, the cells rapidly die once removed from optimal growing conditions and in some cases autolysis may effect both assay performance and stability. Stabilization may be improved by allowing the assay to equilibrate at 2° to 8°C. In general, it is preferable to prepare membranes from the cells and to stabilize these appropriately with protease-inhibitor cocktails. The method below describes the preparation of membranes from Chinese hamster ovary (CHO) cells expressing the M1 muscarinic receptor, but could be adapted to any CHO cell line transfected with membrane-bound receptors. NOTE: CHO cells grow well in 168-cm2 flasks or the larger 250-ml roller bottles. Materials Confluent monolayers of Chinese hamster ovary (CHO) cells genetically modified to express M1 muscarinic acetylcholine receptors (mAChR1; ATTC# CRL-1985) Dulbecco’s PBS, cold (Life Technologies) Hypotonic buffer, ice-cold (see recipe) Dilution buffer (see recipe) Trypsin/EDTA (Life Technologies) Cell scraper 50-ml centrifuge tube Hand-held Dounce homogenizer and tight-fitting pestle with 0.07 mm clearance (Bellco Glass) Cell harvesting 1. Wash confluent monolayers of Chinese hamster ovary (CHO) cells genetically modified to express M1 muscarinic acetylcholine receptors (mAChR1) with cold PBS. Remove wash. The maximum cell and receptor yield is obtained if the cells are harvested when confluent.
SPA Technology to Study Biomolecular Interactions
2. Add 5 ml cold Dulbecco’s PBS (15 ml if using roller bottles) and gently detach the cells using a cell scraper. Aspirate and transfer the cell suspension to a 50-ml centrifuge tube, which should be kept on ice.
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3. Wash the flask with a further 5 ml PBS (10 ml if using roller bottles), and add to the centrifuge tube. Repeat this process with each flask (or roller bottle) until all cells have been removed. 4. Centrifuge 10 min at 600 × g, 4°C. Discard the supernatant. Homogenization 5. Resuspend the cell pellets in 2.5 ml/flask ice-cold hypotonic buffer (5 ml/roller bottle) and leave to stand on ice 10 min. Disrupt the cells using a hand-held Dounce homogenizer and tight-fitting pestle with 0.07 mm clearance. A range of methods are available for the disruption of cultured cells (Findlay and Evans, 1987).
6. Centrifuge the cells 10 min at 600 × g, 4°C. Discard the pellets, unless the pellet is large, in which case repeat steps 5 and 6. 7. Centrifuge the supernatant 30 min at 1000 × g, 4°C. Discard the supernatant and resuspend the pellets in cold dilution buffer, homogenizing with a hand-held Dounce homogenizer. 8. Centrifuge again 30 min at 1000 × g, 4°C. Discard the supernatant and resuspend the pellets in a small volume (5 to 10 ml) cold dilution buffer. Homogenize using the Dounce homogenizer. 9. Dispense the membrane preparation into suitable sized aliquots and store at −70°C. THE APPLICATION OF SPA TECHNOLOGY TO CELL ADHESION MOLECULE (CAM) INTERACTION ASSAYS
BASIC PROTOCOL 4
Cell adhesion molecules (CAMs) mediate a range of processes from embryonic development and differentiation to organization of organ structure and generation of an immune response. Changes in expression or function can result in disease states. For example, abnormal adhesion, molecular structure, function, or expression are associated with lung disease (Mackay and Imhof, 1993; Hamachter and Schaberg, 1994), cancer (Juliano and Varner, 1993), thrombosis, and stroke (Okada et al., 1994), and continue to be the focus of much research. Interactions between CAMs and their respective ligands are often of low affinity and SPA is ideal for studying this due to its homogeneous nature. Heterogenous techniques could cause dissociation of ligand binding due to weak binding interaction (Amersham Biosciences, 1997; http://www.amershambiosciences.com/application/drug_screening). SPA systems have been developed for a variety of CAM activities including selectins, integrins, and fibronectin (Pachter et al., 1995; Game et al., 1998). Here the key points for consideration when designing SPA for CAM interactions will be described and guidelines for assay optimization will be offered (see the flow diagram in Figure 19.8.7). Each interaction should be considered on an individual basis, as many features will be specific to and governed by the molecules involved. Particular attention should also be paid to the selection of an appropriate radiolabeled “counter-ligand.” Capture methods A variety of SPA capture methods exist. One format involves biotinylation of the CAM molecule immobilized onto streptavidin-coated SPA beads; however, if using this technique, biotinylation conditions have to be optimized to achieve functional conjugates. CAMs expressed as ZZ-fusion proteins can also be biotinylated (ZZ- is a synthetic dimer of the Staphylococcus aureus Z domain of protein A that binds to IgG).
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source reagents CAM “Counter-ligand”- whether radiolabelled cells or neoglycoconjugate
assay conditions buffer system co-factors for binding protease inhibitors time temperature agitiation required
reagent concentrations non-specific binding (NSB)
select capture mechanism biotinylate CAM if required
selection of SPA bead type streptavidin protein A/second antibody
optimize signal:noise ratio
assay protocol pre-coupled bead T 0 addition delayed addition validate assay controls inhibition profiles competition binding
color quench correction
performance criteria assay drift signal stability
Figure 19.8.7 Flow chart for the development of CAM interaction SPA.
Antibody capture is an alternate format requiring an antibody, raised against one partner in the adhesion event, to be captured by SPA beads coated with either protein A or an anti-species antibody. Protein A SPA beads can be used by binding the IgG domains of the fusion protein, or antibody binding beads can be used to capture ZZ-cell adhesion molecule fusion proteins. However, these capture systems are relatively inefficient compared to the streptavidin-biotin format already mentioned, and the experimenter must ensure that the binding of the anti-CAM antibody does not inhibit the CAM-ligand interaction. To improve capture efficiency, bridging antibodies are used—i.e., rabbit IgG is used as a bridge between anti-rabbit coated beads and the ZZ-fusion protein of E-selectin (Game et al., 1998). Cells metabolically labeled with [3H]amino acids provide the appropriate “counter-ligand” for both of these formats during the interaction. Cultured cells labeled with [3H]amino acids, expressing the appropriate counter-ligands on their cell surface with the necessary multivalency for high-avidity binding are used. HL60 cells express the correct glycoprotein ligand for E-selectin, and JY cells, which express lymphocyte function associated molecule-1 (LFA), are employed for an ICAM-1 SPA (Game et al., 1998).
SPA Technology to Study Biomolecular Interactions
The use of whole cells may introduce variable expression of counter-ligands due to downor up-regulation, which is of particular importance for CAMs that require activation or that are not constitutively expressed. The E-selectin SPA can be configured using either membranes or whole cells (see Support Protocols 1 and 2). This format depends upon both the constitutive expression of counter-ligands on the cell surface and that they also remain in binding conformation after cell disruption. In some cases cell membranes cannot be used—e.g., ICAM/LFA-1 interaction requires activation of the integrin LFA-1 for ICAM binding, but the cytoplasmic integrin-activating process appears to be disrupted when JY cell integrity is lost (Edwards, 1995; Pachter et al., 1995).
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Cell-free methods In another approach Pachter et al. (1995), using a cell-free format, employed an anti-β1 integrin monoclonal antibody and anti-mouse coated SPA beads to measure the binding of soluble [125I]fibronectin to α5β1 integrin. This avoided the use of [3H]amino acid-labeled cells as the “counter-ligand,” which can cause problems. If the cells lyse during assay incubation, they will release radiolabled cellular components that can give rise to higher background counts in the assay. Also, the use of metabolically labeled cells may give variable results due to the up- or down-regulation of counter-ligand expression. The anti-β1 integrin antibody captures the α5β1 integrin complex from crude CHO cell lysates stably over-expressing human α5 integrin. Another cell-free selectin adhesion assay option involves the use of soluble radiolabeled neoglycoconjugates. The carbohydrate recognition motifs for selectin binding, sialyl LewisX (sLeX) and sialyl LewisA (sLeA) are coupled to a poly-[N-(2-hydroxyethyl)acrylamide] (PAA) backbone (Bovin, 1993). A commercially available glycoconjugate precursor is then radiolabeled with [3H]leucine sLeX (20% molar ratio) [3H]polyacrylamide conjugate ligand available from Amersham Biosciences. The carbohydrate tetrasaccharide density connected to the PAA background can be controlled, and the attachment of multiple ligand copies (10% or 20% molar ratio) ensures presentation in the multivalent form necessary for avid binding. The assay is inhibition sensitive, simple, and convenient to use (i.e., benefits from “off-the-shelf” reagents) and is suitable for high-throughput screening. Optimizing the assay Following the selection of an appropriate “counter-ligand,” the optimal ratio of SPA bead to adhesion molecule can be determined from matrix experiments using increasing bead (typically 0.1 to 0.4 mg/well) and CAM concentrations (50 to 500 ng/well).
6000
SPA cpm bound
5000
4000
3000
total binding 10mM EDTA
2000
1000
0 0
250
300
750
1000
1250
5000
Incubation time (min)
Figure 19.8.8 Time course for the E-selectin SPA. Assay contained 0.1 mg/well SPA beads, 100 ng/well E-selectin-ZZ fusion protein, and 2 × 105 cpm sLex (20% molar ratio) [3H] glycoconjugate. Assays were counted using a MicroBeta 1450 scintillation counter after incubation for 8 hr at 22°C. Data points are means ± range of duplicate wells.
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Several factors can influence the quantity of SPA beads required. These include CAM quality and purity, biotinylation efficiency, and CAM affinity for its counter-ligand. In addition, the time for the assay to reach equilibrium should be determined. A typical time course for the E-selectin SPA is shown in Figure 19.8.8. Most CAM interactions are cation dependent, so an EDTA blank can be used to determine the level of nonspecific binding (NSB) to the SPA beads in the presence of assay components. Binding of [3H]-labeled cells or neoglycoconjugate to the bead without the biotinylated component should also be examined. Alternatively the CAM interaction specificity can be determined using appropriate antibodies such as anti-ICAM-1, or a structural analogue of the counter-ligand. Optimized CAM SPA can be evaluated by profiling with competing monomeric or polymeric carbohydrates. Game et al. (1998) described such an experiment for E-selectin interaction using metabolically labeled HL60 cells and labeled neoglycoconjugate as selectin “ligands.” Inhibition profiles of both formats were similar, the glycoconjugate assay being slightly more sensitive to monomeric carbohydrate inhibition. In both formats, specific binding was completely abolished by blocking anti-E-selectin antibodies. The application of SPA technology to the investigation of CAM interactions has been demonstrated. When designing CAM SPA each interaction must be considered individually as many features will be specific to that molecule, in particular the selection of an appropriate “counter-ligand” is important. Metabolically labeled cells are an option, but may give variable results due to the up- or down- regulation of the counter-ligand expression. Membranes prepared from labeled cells are another possibility. Finally, the introduction of validated, synthetic radiolabeled neoglycoconjugates mimicking the counter-ligands or recognition motifs for CAMs, have resulted in the development of cell-free, homogeneous SPA for cell adhesion molecule interactions. BASIC PROTOCOL 5
THE APPLICATION OF SPA TECHNOLOGY TO STUDY PROTEIN/DNA INTERACTIONS Transcription is a highly regulated cellular event, requiring the interaction of many proteins in multimeric complexes at the transcription initiation point (Buratowski, 1994). Some of these proteins are generic transcription factors, but large numbers of more specific factors that associate with particular DNA sequences have also been identified (Buratowski, 1994; Cowell, 1994; Hill and Treisman, 1995). Binding of transcription factors to DNA sites has primarily been studied by gel electrophoretic mobility shift assays or filter-binding assays using 32P-labeled DNA (UNIT 9.7; Dahlquist, 1978; Zabel et al., 1991); however, gel-shift assays are not readily quantifiable and filter assays are fairly labor-intensive, making SPA an attractive alternative method. Advantages SPA has several advantages over other techniques; namely speed, convenience, and the use of isotopes other than 32P. In addition, SPA is also useful for studying low-affinity binding interactions, requiring none of the separation steps that usually risk disturbing them. Another significant advantage is the potential for assaying binding in real time and being able to study association and dissociation rates. A number of SPA assay formats can be applied to the study of these interactions (e.g., streptavidin-biotin formats and antibody capture). The choice depends upon the radiolabel location and capture system chosen.
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An example for nuclear factor-κB Nuclear factor-κB (NF-κB) was chosen to develop an SPA protein-DNA interaction assay. This is an important factor due to its central role in regulating the transcription of a large number of genes involved in cellular defense (Baeuerle, 1991; Liou and Baltimore, 1993). NF-κB is a family of proteins forming a range of homo- and heterodimers. The affinities of these various dimers for different DNA consensus sites varies, leading to an additional mechanism of transcriptional control (Siebenlist et al., 1994; Thanos and Maniatis, 1995). This SPA used an NF-κB-GST-p65 fusion protein to quantify binding of the p65 subunit to DNA. The authors found that the best approach was to use radiolabeled consensus DNA, NF-κB-GST-p65 fusion protein subunit, anti-GST antibody, and protein A SPA beads to capture the complex. A double-stranded DNA (dsDNA) sequence, based on the NF-κB consensus site in the HIV-1 long terminal repeat, was synthesized and labeled with methyl 1′,2′-[3H]thymidine 5′ triphosphate ([3H]TTP; Amersham Biosciences) using Sequenase. Specific activities obtained were 1 to 2 Ci/µmol. Determining optimized assay conditions Optimized conditions for the NF-κB-GST-p65 SPA involved the incubation of the fusion protein with the [3H]DNA at room temperature for 30 min in a buffer containing: 10 mM HEPES, pH 7.0, 20 mM sodium acetate, 0.2 mM EDTA, 0.05% (v/v) Nonidet P40 (NP40), 5 mM DTT, and 1 mg/ml BSA. The anti-GST antibody and protein A-coated SPA beads (1 mg) were then added to a final assay volume of 100 µl. After a further incubation at room temperature for 60 min, the samples were counted using an appropriate scintillation counter. To obtain a stable and optimized SPA signal requires optimization of reagent concentration, buffering, and incubation conditions via a series of experiments and should be established for each individual assay. The amount of SPA beads and capture antibody used is particularly important, as all fusion protein present in the assay mixture should be associated with SPA beads. The order of addition can also have an impact, and titration experiments varying the concentration of radiolabeled DNA and fusion protein should be performed.
[NF-κ B:p65-GST/DNA complex] nM
0.18 0.16 0.14 0.12 0.10 0.08 0.06 0.04 0.02 0.00 0.0
0.1
0.2
0.3 [DNA] nM
0.4
0.5
0.6
Figure 19.8.9 Titration of consensus DNA against NF-κB-GST-p65. 25 nM GST-p65, was titrated with up to 0.5 nM [3H]DNA as described in the text. Results are means ± SD (n = 2) and are representative of three experiments.
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Typical results from such experiments are shown in Figure 19.8.9, illustrating DNA titration versus a constant concentration of NF-κB-GST-p65. SPA counts correlated with DNA concentration, and complex formation was saturated at 0.3 nM DNA for 25 nM GST-p65. (The concentration of complex formed is calculated after allowing for the approximate 50% efficiency of 3H SPA counting, compared with liquid scintillation counting). The specificity of the protein-DNA interaction was validated using further DNA titration and competition experiments (e.g., incubating the GST fusion protein in the presence of [3H]DNA with increasing amounts of either unlabeled DNA consensus sequence or a mutant DNA sequence). The authors showed that consensus sequence binding was not inhibited by up to 40-fold excess of mutant over consensus DNA. To summarize, the application of SPA for the study of protein-DNA binding has been demonstrated using NF-κB-GST-p65 as a model system. The binding detected is specific and can be assayed in real time. Preliminary data generated suggests it could be used for kinetic studies of GST-p65/DNA binding, although for some determinations a higher specific activity DNA substrate may be required. The existing format of room temperature incubation generating a stable signal for overnight counting would make it suitable for use as a high throughput-screening assay. In addition, the assay can distinguish specific competition between cold consensus DNA and mutated DNA, suggesting it would be suitable for NF-κB-GST-p65/DNA inhibitor screening. BASIC PROTOCOL 6
THE APPLICATION OF SPA TECHNOLOGY TO RADIOIMMUNOASSAYS (RIAs) In common with conventional heterogeneous radioimmunoassay (RIA) systems, SPA RIAs are based on the competition between unlabeled ligand and a fixed quantity of radiolabeled ligand for a limited number of binding sites on a specific antibody. With fixed amounts of antibody and radioactive ligand, the amount of radioactive ligand bound by the antibody will be inversely proportional to the concentration of added unlabeled ligand. SPA RIAs have been widely utilized with a broad spectrum of applications, including mass measurement in research, pharmacological studies, and cellular function screening for analytes such as prostaglandins (Baxendale et al., 1990; Sugatani et al., 1994), steroids (Fiet et al., 1991, 1994), drugs (Fenwick et al., 1994), and second messengers such as cyclic AMP and GMP (Horton et al., 1998). Assay Design SPA beads are available coupled to anti-rabbit, anti-mouse, and anti-sheep IgG as well as protein A (Amersham Biosciences), making this technology suitable for use in a large range of assays, by either converting existing RIAs to an SPA format or for developing new assays using this technology. Table 19.8.2 illustrates which SPA bead is suitable for use with different sources of primary antibody. Figure 19.8.10 shows in flow diagram format the key steps to be considered when developing an SPA radioimmunoassay.
SPA Technology to Study Biomolecular Interactions
Choice of SPA bead and tracer The choice of either PVT or YSi beads will depend primarily on the nature of the ligand and whether it interacts with the bead. Assays using YSi beads will require agitation to enable equilibrium to be attained but will have higher SPA counts than PVT beads. With the exception of higher signal output, YSi beads and PVT beads should produce similar assay characteristics. SPA technology is suitable for the detection of both 3H- and 125IRIA tracers.
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Table 19.8.2
Choice of SPA Bead for RIA
Source of primary antibody
Compatible SPA bead
Goat Guinea pig Mouse
Anti-sheep Protein A Anti-mouse Protein A (depending on the IgG subtype) Anti-rabbit Protein A (depending on the IgG subtype) Anti-sheep
Rabbit Sheep
select primary antibody: specificity affinity
choice of SPA bead: antibody coating PVT/YSi
assay protocol: T0 addition pre-equilibration of antibody and ligand
tracer concentration: NSB NPE
bead concentration: total capture of primary antibody
sample preparation color quench correction
reagent concentrations
assay conditions: time temperature
validate assay: standard curves
Figure 19.8.10 Flow chart for the development of SPA RIAs.
Optimization of SPA bead and antibody The most common assay format for SPA RIA systems is T0 addition (i.e., sequential addition of all reagents). This requires the presence of excess SPA beads to ensure complete capture of the primary antibody. Potential consequences of using large amounts of beads include an increase in NSB and NPE, contributing to higher background counts. It is therefore recommended that a matrix experiment be performed with increasing concentrations of beads and primary antibody to optimize the signal-to-noise ratio and obtain an acceptable level of SPA counts. It is also important that assays are counted at equilibrium, as the signal will vary until this point. Agitation may be required during the incubation period and will be essential if using
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YSi SPA beads. It is also recommended that incubations be carried out at a similar temperature to that at which the assay will be counted. Incubation time It is important that assays are incubated until equilibrium is reached, otherwise assay drift may occur. Agitation of the assay tube/well contents may be required during the incubation period and a variety of shakers are available commercially for this purpose. Incubation temperature Standard assay incubation temperatures are suitable for SPA; however, it is recommended where possible that incubations be carried out at a temperature similar to that at which the assay tubes/plate will be counted. Matrix effects Many matrices (e.g., plasma and serum) contain many compounds that may interfere nonspecifically with the assay reaction and effect the signal in some way. While not specific to SPA, these matrix effects should be considered and accounted for by preparing the assay standards in the same medium or matrix as the samples. Detection Instrumentation SPA can readily be counted using standard single-tube liquid scintillation counters such as the Wallac MicroBeta (PerkinElmer), provided appropriate care is taken to set the counting windows as recommended by the manufacturers. In situations where a higher level of throughput is required, as is the case when many thousands of samples are screened in the early stages of drug discovery within the pharmaceutical industry, SPA assays are conducted in 96- or, increasingly, 384-well microplates. Detection instrumentation that allows measurement of such high-density assay formats has been developed. Both the Packard Topcount and Wallac MicroBeta instruments have been developed for this purpose, and are now available from PerkinElmer. As such, it is important to appreciate the capabilities of these two instruments and how their design addresses the particular challenges of the HTS environment. Wallac MicroBeta The MicroBeta is a dual photomultiplier tube (PMT)–based microplate scintillation counter. Each well of the microplate being measured is provided with a photomultiplier tube both above and below. Each pair of tubes operates via coincidence detection circuitry which acts to eliminate unwanted background count events arising due to either thermal noise in the PMT, cosmic rays, or chemiluminescence in the sample. Models are available with 1, 2, 3, 6, and 12 detector pairs, which count in parallel to increase throughput. Plate handling for the manual versions is based on a cassette adapter system, which allows for great adaptability in the type of sample handling. Small vials, tubes, filter mats, and various designs of 24-, 96-, and 384-well plate formats can be accommodated. Operationally, one disadvantage is that plates have to be fitted into the appropriate adapter before counting, but this does then provide a greater degree of uniformity in the size of the assemblies being counted. This pays dividends in the reliability of the whole system. The MicroBeta can hold up to 12,288 samples (32 × 384 format). A robot loadable version is also available. SPA Technology to Study Biomolecular Interactions
Packard TopCount The TopCount, as the name suggests, relies on a single PMT arranged above the well being counted. This machine is optimized to make measurements on opaque plate formats
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and so is incapable of the dual PMT coincidence technique for reducing background noise. Instead, the TopCount uses a patented electronic method of discriminating “true” scintillation events from unwanted interfering events. The TopCount is available in both 6- and 12-detector versions. Sample loading is achieved with a stack mechanism which in the current version (TopCount NXT) allows for 25 plates to be loaded. Direct, robotic loading of plates is also possible. Plates with 96- and 384-wells can be counted and these are loaded directly without the use of adapters. The noise discrimination technique of the TopCount is well matched to the characteristics of inorganic solid scintillator materials. Such materials, for example calcium fluoride or YSi, doped with low levels of elements such as europium or cerium, respond directly to β particles by emitting light. These materials have been incorporated into 96-well plates for counting without the use of scintillation cocktail (LumaPlates) and used for some versions of the scintillation proximity assay technique. Conventional scintillation cocktail with slow decay characteristics is required to obtain the best counting efficiency for low energy isotopes in this instrument.
Figure 19.8.11 The LEADseeker Components. (1 and 2) Light-tight chambers containing CCD camera and microplate focusing mechanism. (3) Microplate stacker carousel. (4) Bar code reader. (5) Microplate shuttle mechanism. (6) Camera cooling unit.
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Color quench correction When samples are colored, a color quench correction program needs to be installed on the counting instrument. For SPA assays using both PVT and YSi beads, color quench correction kits for either 3H or 125I can be used (Amersham Biosciences). It is recommended that the user follow the instructions provided by the instrument suppliers when constructing color quench curves on the TopCount or MicroBeta. Imaging of SPA assays The move towards ultra-high throughput SPA assays within the pharmaceutical industry has increased the pressure on conventional plate-counting technology and reagents. As plates have increased in well density, counting times have also increased. Typically, scintillation counters can only read up to 12 wells of a 384-well plate at any one time, giving count times of up to 40 min per plate. Automated imaging devices such as LEADseeker (see Figure 19.8.11), developed jointly by Amersham Biosciences and Imaging Research, comprise a charge-coupled device (CCD) camera and a set of new bead reagents (LEADseeker beads; Amersham Biosciences) compatible with CCD-based imaging (see below). This system is capable of measuring an entire plate in a single image and thus reduces count times. In addition, the new bead types have emission characteristics that greatly reduce the effect of color quenching on signal. LEADseeker bead types As standard PMTs are most sensitive to light in the blue region of the emission spectrum (i.e., 400 to 450 nm), SPA beads were developed to emit light in this region. CCD chips are more sensitive to light in the red region of the spectrum; therefore, Amersham Biosciences, using proprietary technology, has developed new bead types which emit light in the ∼600 nm region, optimized for use with LEADseeker. An advantage of these beads is that the quenching effect of yellow and red compounds is eliminated. There are two core bead types, one based on polystyrene (PS), the other on yttrium oxide (YOx). Both types have an emission spectrum with a peak at 615 nm and exhibit a much higher light output than SPA beads. Each may be derivatized by covalently linking molecules such as streptavidin in the same way as SPA beads. REAGENTS AND SOLUTIONS Biotin/streptavidin capture buffer 20 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) 10 mM MgCl2 10 mM DTT 0.1% (w/v) BSA Store up to 4 weeks at 2° to 8°C Dilution buffer 50 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) 5 mM MgCl2 Appropriate protease inhibitor combination, for example: 80 µg/ml bacitracin 0.25 mg/ml Pefabloc SC (Roche Molecular Biochemicals) 4 µg/ml leupeptin 4 µg/ml chymostatin Store up to 4 weeks at 2° to 8°C SPA Technology to Study Biomolecular Interactions
Sigma now supplies ready prepared protease inhibitor cocktails specifically for mammalian cell preparations.
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Grb2-SH2/SH3 SPA buffer 20 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) 250 mM NaCl 0.1% (w/v) BSA Store up to 4 weeks at 2° to 8°C Harvesting buffer 50 mM HEPES, pH 7.4 145 mM NaCl2 25 mM CaCl2 1 mM MgCl2 10 mM glucose 0.1% (w/v) BSA 0.25 mg/ml bacitracin 0.25 mg/ml Pefabloc SC (Roche Molecular Diagnostics) 25µg/ml leupeptin 25µg/ml aprotinin Store up to 4 weeks at 2° to 8°C Hypotonic buffer 5 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) 5 mM MgCl2 Appropriate protease inhibitor combination, for example: 80 µg/ml bacitracin 0.25 mg/ml Pefabloc SC (Roche Molecular Biochemicals) 4 µg/ml leupeptin 4 µg/ml chymostatin Store up to 4 weeks at 2° to 8°C NF1-Ras SPA buffer 50 mM Tris⋅Cl buffer, pH 7.5 (APPENDIX 2E) 2 mM DTT (APPENDIX 2E) This buffer must be prepared fresh YY SPA buffer 50 mM HEPES, pH 7.4 1 mM MgCl2 2.5 mM CaCl2 0.1% (w/v) BSA 0.025% (w/v) bacitracin 0.025% (w/v) sodium azide Store up to 4 weeks at 2° to 8°C COMMENTARY Background Information Protein-protein interactions The interaction of one protein with another can be studied by SPA, provided one of the proteins is radiolabeled. In order to achieve a good signal in the assay, a chemically pure protein labeled to a high specific activity is desirable. Kahl et al. (1996) compared both 125I and 3H for labeling Ras directly using conventional labeling methods and for loading Ras
with radiolabeled GTPγS, which permits the use of wild-type Ras proteins. For the NF1-Ras SPA described in this unit, an oncogenic Ras protein (RasL61) was used. This mutant does not hydrolyze GTP even upon binding of a GTPase activating protein. This enabled RasL61 to be loaded with [3H]GTP using an exchange reaction performed in the presence of a GTP regenerating system (phosphoenol pyruvate and pyruvate kinase) in which GDP and GTP rapidly exchange from
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SPA Technology to Study Biomolecular Interactions
the protein into solution and any GDP in solution is converted into GTP, under optimized conditions (Skinner et al., 1994). The GST-NF1-Ras assay described (see Basic Protocol 1) involves the coupling of the GST-NF1-Ras complex via anti-GST antibodies to the protein A SPA beads; however, alternative coupling mechanisms can be applied to study protein/protein interactions using SPA technology. As described previously, anti-species antibody beads (e.g., anti-rabbit, antisheep, or anti-mouse) may be substituted for the protein A beads. Indeed it is advisable to test both anti-species antibody and protein A beads as part of the assay development process. Alternatively, if one of the proteins can be biotinylated without loss of biological activity, then the use of streptavidin beads becomes an option. GST-fusion proteins have been biotinylated for other assays without interfering with the functional activity of the protein. This format has been used for an SPA system designed to measure the interaction between Ras/Raf and the binding of HIV-1 Nef protein to the Src homology region 3 (SH3) domain of Hck (Kahl et al., 1996; Jones et al., 1998, respectively). In the Ras/Raf assay, a GST-Raf fusion protein was biotinylated using sulfosuccinimidyl-6(biotinamido) hexanoate. If biotinylation is considered, it is necessary to optimize the ratio of biotinylation reagent to protein. This must be done empirically for the system under study. There is currently no method for the precise determination of the number of biotin molecules that become attached to a protein. A suggested protocol would be to biotinylate the protein using increasing molar ratios of biotinylation reagent to protein from 1:1 up to 50:1. Following biotinylation of the protein and separation from the biotinylation reagent, the biotinylated protein can then be tested in a simple binding experiment using 1 mg streptavidin beads and a constant amount of radiolabeled protein. Once the optimal ratio of biotinylation reagent/protein is established, then further optimization experiments are necessary to determine amounts of both biotinylated protein and streptavidin beads required for the assay. Finally, alternative assay formats that employ GST- or His-tagged fusion proteins can also be considered, using the appropriate glutathione or copper-chelating coated beads, respectively. An important point to remember with all the SPA formats described in this section is that it is essential that the capture protein is present in
excess with respect to the radiolabeled protein in the assay, and that beads are present in excess to capture all the complex in order to maximize the signal obtained. To optimize the concentration of the GST-NF1 fusion protein, increasing concentrations of GST-NF1 were added to each assay in the presence of 0.5 mg protein A SPA beads and a constant amount of Ras [3H]GTP. The titration of GST-NF1 was performed using four concentrations of Ras⋅[3H]GTP (Figure 19.8.12). The signal increased with the concentration of GST-NF1 until all the available binding sites (i.e., anti-GST antibody bound to the protein A beads) were saturated. Further increases in the GST-NF1 concentration resulted in a reduction in signal as competition was established for anti-GST antibody by GST-NF1 and a complex of GST-NF1 bound to Ras⋅[3H]GTP. Improved signal was also observed by increasing the concentration of Ras⋅[3H]GTP. The specificity of the NF1-Ras interaction was determined using unlabeled GTP, ATP, and GDP over a range of concentrations in an SPA competition binding assay (see Figure 19.8.13). GTP appeared to have an inhibitory effect on the NF1-Ras interaction under normal assay conditions, with IC50 values of 0.1 to 1 mM being observed. Skinner et al. (1994) used a neutralizing anti-Ras monoclonal antibody Y13-259 and a detergent, n-dodecyl maltoside (a specific inhibitor of NF1 catalytic activity), to demonstrate specific binding. Both reagents abolished the signal from the NF1/Ras SPA, but neither affected the signal from a control SPA in which a [3H]GTP containing GST-Ras fusion protein was bound to protein A beads (unpub. observ.). The interaction between Ras and NF1 is known to show a high degree of specificity for the GTP-bound form of Ras. This was demonstrated using both Ras containing [3H]GTP and [3H]GDP, where no binding of Ras⋅[3H]GTP to GST-NF1 was detectable. The kinetics of both the NF1-Ras interaction and the Ras-Raf interaction have been studied (Skinner et al., 1994; Gorman et al., 1996; Sermon et al., 1996). The homogeneous nature of SPA technology permits the kinetic analysis of such interactions, despite very fast dissociation rates. Skinner et al. (1994) calculated the affinity of the NF1-Ras interaction, giving a Kd of ∼40 nM. SH2 and SH3 domain binding to specific peptide sequences SH2 domains control enzyme activity by binding membrane-bound receptors to regulate
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Specific SPA cpm bound(× 10 –3 )
10.0
7.5
5.0
2.5
0.0 0
10
20
30
70 40 50 60 GST-NF1(pmol/well)
80
90
100
110
Figure 19.8.12 Effect of increasing GST-NF1 and H-RasL61⋅[3H]GTP concentration on specific SPA cpm bound. 10 pmol/well H-RasL61⋅[3H]GTP (squares), 20 pmol/well H-RasL61⋅[3H]GTP (upward-facing triangles), 30 pmol/well H-RasL61⋅[3H]GTP (downward-facing triangles), 40 pmol/well H-RasL61⋅[3H]GTP (diamonds). Assay conditions: 3.75 µg anti-GST antibody (Molecular Probes), 0.5 mg protein A beads in 50 mM Tris⋅Cl, pH 7.5, 2 mM DTT. Results are means ± SEM (n = 3).
120
% Inhibition of binding
100
80
60
40
20
10 10 –6
10 –5
10 –4
10 –3
10 –2
10 –1
10 0
10 1
10 2
10 3
Concentration of competitor ( µM)
Figure 19.8.13 Inhibition of H-RasL61⋅[3H]GTP binding to GST-NF1 by GTP (square), ATP (downward-facing triangle), and GDP (upward-facing triangle). Assay conditions as for Figure 19.8.12 using 10 pmol H-RasL61⋅[3H]GTP and 40 pmol GST-NF1. Results are means ± SEM (n = 2). Identification of Protein Interaction
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downstream events such as kinase cascades; however, transcription factors—e.g., signal transducers and activators of transcription (STATs)—recently have been shown to contain SH2 domains (Rosen et al., 1995). SH3 domains are also thought to be a feature of signal transduction mechanisms, or to have a role in cytoskeletal organization (Musacchio et al., 1994). SH2 domains have been well characterized by in vitro mutagenesis studies, and are known to specifically bind phosphotyrosine residues with affinities in the low nanomolar range. Examples of proteins containing SH2 domains include enzymes—e.g., GTPase activating protein (GAP), Src—and nonenzymatic adapters—e.g., growth factor receptor binding-2 (GRB2), p85 domain of phosphatidylinositol 3-kinase. SH3 domains bind proline-rich sequences with a much lower affinity. Recently, the use of peptides as opposed to intact proteins for binding studies has received significant attention. Examples of proteins containing SH3 domains also include enzymes (Abl, GAP) and adapters (Crk and p85). Nonhydrolysable peptide mimetics might represent an attractive therapeutic target. SPA has been used successfully to quantitate SH2 and SH3 domain binding to radiolabeled peptides. Recent examples include the successful configuration of an assay that monitors the ability of compounds to inhibit the binding interaction of a selected phosphopeptide with the tandem SH2 domains of the ZAP-70 protein tyrosine kinase, an important enzyme involved in T-cell function (Sheets et al., 1998). Pai et al.
Table 19.8.3
(1999) developed an SPA to screen inhibitors of phosphopeptide binding to the SH2 domains of the proteins Grb2 and Syk, both of which play important roles in cell activation. Choice of N- or C-terminal phosphotyrosines. An investigation has been carried out into the relative affinities of the N- and C-terminal phosphotyrosines for the SH2 domain. The ability of various peptides to compete with [125I]ppY604–635 for binding to the p85 SH2 domain was tested. The results showed that the N-terminal pY bound the SH2 domain with a slightly higher affinity than the C-terminal pY. In addition, a preliminary experiment indicated that a mixture of the N- and C-terminal pY sequences did not compete with the radiolabel as efficiently as the intact sequence. This indicated that the sequence between the phosphotyrosines is important for binding, either directly or by providing conformational restraint. The effect of temperature on binding with reference to GAP (SH2-SH3-SH2). Three peptides sequences derived from the platelet derived growth factor receptor (PDGFr; see Table 19.8.3) were evaluated for binding to the GAP SH2 domain over a range of temperatures (5° to 30°C). The results revealed that the shorter sequence was more susceptible to temperature changes than the longer sequence, which in turn was more susceptible than the cyclic sequence (unpub. observ.). Low affinity SH3 domain binding: the Crk/Abl interaction. The biotin-streptavidin system was selected for this assay as the reagents were supplied prebiotinylated. This assay involved incubating biotinylated Crk SH3
Peptide Sequences Used for Testing Binding to the GAP SH2 Domain
Peptide
Sequence
98P 117P Cyclic 117P
SSNpYMAPYDNY GDIKYESSNpYMAPYDNYVPSA G D I K Y E S S N p Y M A P Y D N Y V P S A-cyclica
aIndicates covalent attachment of the N and C termini.
Table 19.8.4
SPA Technology to Study Biomolecular Interactions
Peptide Sequences Used for Testing Binding to the Crk SH3 Domaina
Source
Sequence
Code
Binds to
Abl-1 Abl-2 Btk-1
QAPELPTKTRT SEPAVSPLLPRKER TKKPLPPTPEED
108 109 110
Crk SH-3 Crk SH3 Hck SH3
aSpecificity was demonstrated by the Btk-1 peptide, known to bind the Hck SH3 domain, but not the Crk SH3
domain. Figure 19.8.14 shows competition between unlabeled peptides and peptide [3H]108.
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120 Peptide 108, B 0 =5830cpm Peptide 108, B 0 =3348cpm
100
B/B (%)
80
60
40
20
10 10 –4
10 –3
10 –2
10 –1
100
10 1
10 2
10 3
Concentration of unlabelled competing peptide (µ M)
Figure 19.8.14 Unlabeled peptides competing with peptide [3H]108 for binding to the Crk SH3 domain. Assays were performed using 20 mM MOPS buffer, pH 7.5, containing 10 mM DTT, 1% (v/v) Triton X-100, 0.1% (v/v) SDS, and 1 mg streptavidin SPA beads. Following overnight incubation at 2° to 8°C, the assays were counted using a TopCount (PerkinElmer). Results are means ± SEM (n = 3).
domain with radiolabeled binding peptides derived from the Abl tyrosine kinase (Table 19.8.4). Because of the extremely low affinity of this interaction (micromolar), a relatively large amount of radiolabeled peptide had to be added to produce an acceptable signal. 3H- and 125I-labeled peptides were compared, and it was found that only the 3H-labeled peptide produced a signal. The probable reason for this is steric hindrance caused by the 125I atom. Figure 19.8.14 shows an example of a competition experiment using peptide 108. The assay exhibited a severe temperature dependency, which meant that the assay had to be incubated at 4°C, and counted before the plate reached room temperature in order to produce a signal to noise ratio of 6:1 (data not shown). Using this approach, SPA has enabled quantification of binding events with micromolar affinities. Further work focused on protein/protein SH3 interactions and results suggested that these have higher affinities than the equivalent protein/peptide interaction. Receptor-ligand interactions In this unit, a competitive binding assay for the Y1-type neuropeptide Y/peptide YY receptor by SPA (see Basic Protocol 3) has been briefly described. It was emphasized in this protocol that optimization of the membrane
receptor/bead ratio is a critical factor in the successful development of receptor SPAs. This optimization process involves performing matrix experiments varying the quantities of both membrane-bound receptor and SPA beads added to the assay. As a guideline, set up B0/NSB tubes for different ratios of bead and membrane (or cells), with constant levels of ligand (concentration is dependent on the ligand affinity but should not be above KD levels), and assay volume (e.g., 200 µl). Select the membrane/bead ratio that gives the optimum specific signal in the assay. The following range of bead and membrane levels could form a useful starting point for assay optimization: [125I]ligands: 1 to 100 µg membrane protein (or 1 × 105 to 5 × 106 cells) and 0.5, 1.0, and 2.0 mg beads. [3H]ligands: 10 to 200 µg membrane protein (or 1 × 106 to 1 × 107 cells) and 1.0, 2.0, and 4.0 mg beads. In addition, it is important to confirm that the radioligand does not bind nonspecifically to the selected bead type by including controls performed in the absence of membrane protein. The assay described for receptor-ligand interactions (see Basic Protocol 3) is termed a T0 (time = 0) addition assay as it involves the sequential addition of radioligand, test samples
Identification of Protein Interaction
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(if required), membrane, and beads as separate additions. In this format, the coupling of membrane to beads occurs simultaneously with ligand-receptor binding. This is the most widely used SPA format and, as discussed previously, it is important to ensure that sufficient beads are added in order to capture all the membrane present. Certain assays require the precoupling of the membrane preparation to the SPA bead. This is particularly true if the experiment involves the investigation of receptor binding kinetics (e.g., measurement of ligand association and dissociation). For these studies, the use of a precoupled bead format is necessary to ensure that the rate of association detected is that of receptor/ligand complex formation, and to avoid interference from membrane/bead association. The precoupled bead format involves coupling membranes to the beads prior to assay. Again the quantities of both membrane-bound receptor and SPA bead are varied and, after coupling, excess unbound membrane is separated by either allowing settling of the beads or by a short centrifugation step. Figure 19.8.15 shows an example of this assay format used to determine the association/dissociation kinetics of [3H]scopolamine binding to the M1 muscarinic acetylcholine receptor (mAChR1) by SPA
[ 3H]-Scopolamine specific binding (dpm)
3000
(O’Beirne et al., 1994). Accurate measurements may be made by simple, repeat counting using a minimum of assay wells. This does not disrupt the equilibrium and therefore a study on the kinetics of the drug candidate can be completed with minimal use of reagents. The absence of a separation step with SPA is also an advantage when assaying receptors of weak affinity (e.g., soluble epidermal growth factor receptor; Hurwitz et al., 1991), where filtration or washing steps may shift the apparent equilibrium of the binding event under study.
Critical Parameters Assay optimization Bead amounts. Using 0.5 to 1 mg of beads is a good starting point for most assays. Varying the amount of beads between 0.1 and 1.5 mg as part of a matrix experiment will illustrate the concentration required to achieve a balance between signal, noise, and sensitivity. Bead mixing and settling. YSi beads are difficult to maintain in suspension and, if used, will require agitation to reach equilibrium. A variety of mixers are commercially available. PVT beads have a much lower density and can
QNB
2500
2000
1500
1000
500
0 0
SPA Technology to Study Biomolecular Interactions
50
100
150 200 Time (min)
250
300
350
Figure 19.8.15 Association/dissociation of [3H]copolamine binding to mAChR1 measured by SPA. Stock solutions of mAChR1 membranes were rolled for 2 hr with WGA PVT SPA beads at a ratio of 50 µg membranes/4 mg beads. [3H]Scopolamine (100 pM final) was added and the signal monitored by counting 1 min at 5-min intervals using the MicroBeta 1450 scintillation counter (PerkinElmer). At equilibrium (after 2.5 hr), (−)-quinuclidinyl benzilate (QNB) (2 µM final) was added and the decreasing signal measured as above. The data was fitted and kinetic constants estimated using EBDA (McPhearson, 1985).
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be counted while still in suspension within 4 to 6 hr. Biotinylation. It is essential to optimize biotinylation conditions by using increasing amounts of biotinylation reagent to protein (e.g., from 1:1 up to 50:1). There is no method of measuring the number of biotins that become attached. Test the biotinylated protein using a simple SPA binding experiment, 1 mg beads, and a constant amount of radiolabeled protein. Order of addition. This can have an impact on the SPA signal so it is worth experimenting with this during optimization. The most widely used SPA format is termed T0 (time = 0), which involves the sequential addition of radioligand, test samples, membrane/antibody, and beads. When using this format it is important to ensure that there are sufficient beads for capture. In receptor studies, the precoupled approach where membranes and beads are incubated together can also be successful, particularly for kinetic studies. Delayed addition, where the receptor is pre-equilibrated with the ligand before adding the SPA bead, is another useful approach when the bead-coupling mechanism may interfere with the receptor binding site, or where a shorter assay time is required. Assay conditions Buffer cofactors. Consult current publications for suitable buffering systems, including any cofactors such as divalent cations that may be essential for the assay. Choice of SPA bead (e.g., for protein-protein interactions). The specificity, affinity, and purity of the antibody will effect the maximum level of signal achievable and the choice of bead. Both anti-species antibody binding and protein A beads should be evaluated as it is impossible to predict which type will give the best results. Regardless of bead type, the capture protein must be present in excess with respect to radiolabeled protein. To ensure the entire complex is captured, and therefore ensure the signal is maximized, an excess of beads must also be present. Equilibrium. The SPA signal will vary until all the interactions reach equilibrium. Counting repeatedly at timed intervals will monitor this progress. The authors recommend the assay be agitated as this reduces the time taken to reach equilibrium. Interfering substances. As SPA is a homogeneous assay technology, the effects of potentially interfering substances should be thoroughly tested as part of the assay optimization. Substances that could affect SPA results in-
clude detergents (e.g., SDS, CHAPS, deoxycholate), organic solvents (e.g., DMSO, methanol), and compounds contained within plasma and serum. NSB effects. It is important to confirm that there is little interaction between ligand and bead in absence of, for example, receptors. Appropriate no-membrane controls must be included to measure the extent of the binding. The careful use of reagents such as BSA, detergents such as PEI or Triton X-100, or varying the buffer ionic strength may also help to reduce NSB. Protease inhibitor cocktails. During long assay incubations these may be required to guard against degradation of proteinaceous assay components by endogenous enzymes. This will help ensure that stable SPA results are generated. Sigma supplies ready prepared cocktails, which include Bacitracin, Pefabloc SC (Roche Molecular Biochemicals), leupeptin, and chymostatin. Temperature dependency. It is recommend that the temperature used for the incubation be the same as that used for counting the assay; however, some assays exhibit severe temperature dependency and this must be taken into account. Instrumentation Color quench correction. Colored samples that absorb light near 400 nm (yellow) may decrease the SPA signal. To correct for these effects a color quench curve must be set up for each assay and counter. Amersham Biosciences supplies color quench correction kits based on tartrazine for use with both 3H and 125I. Counting windows. These must be adjusted as recommended by the instrument suppliers to the appropriate settings to enable SPA counting with different isotopes.
Troubleshooting Poor signal The following is a checklist which should be performed if poor signal is a problem: 1. Perform a matrix experiment (e.g., bead titration versus assay components). 2. Check to ensure the amounts of critical components used are sufficient (e.g., enzyme, beads). 3. Perform a time-course experiment to determine the stable counting window for the assay. 4. Check the specific activity of the ligand and that it has been diluted correctly.
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5. Check for poor binding caused by interfering substances. 6. Ensure any required cofactors (e.g., Mg2+, Mn2+) are present in the assay. 7. Check the incubation time and temperature. 8. Ensure the instrument window settings are correct for the bead and label type. Poor reproducibility The following is a checklist which should be performed if poor reproducibility is a problem: 1. Ensure that a homogeneous suspension of beads is maintained during pipetting steps by gentle mixing with a magnetic stirrer. 2. Ensure that there is not uneven heat distribution in tubes or wells during incubation. 3. Ensure that all reagents have been pipetted into the bottom of the tube or well. 4. Check the assay precision by including more replicates. 5. Check to ensure that pipettes are calibrated and functioning reliably.
Anticipated Results Protein-protein interactions The measurement of protein/protein interactions in the absence of enzymatic activity is feasible using SPA technology. The NF1-Ras SPA format demonstrates that interactions of micromolar affinity and fast dissociation rates can be measured using SPA, permitting both kinetic analysis and the screening of potential inhibitors. Src homology domains Different approaches can be used for studying SH2/SH3 binding by SPA: the biotin/streptavidin system or the antibody capture method. The latter may be more suitable if biotinylation is known to affect binding of the protein to the peptide. A temperature dependency effect has been observed in some cases, so it is recommended that counting be carried out in a temperature-controlled counter. Assay conditions are relatively consistent with only minor alterations required between the systems tested to date. SPA has therefore been shown to be suitable for measuring SH2 and SH3 interactions with specific peptide sequences in the high-, intermediate-, and low-affinity range.
SPA Technology to Study Biomolecular Interactions
Receptor-ligand interactions SPA has been successfully applied to an extensive range of receptor binding assays (e.g., Hoffman and Cameron, 1992; Kienhuis
et al., 1992; Reinscheid et al., 1995; Dayton et al., 1996; Patel et al., 1996; Dairaghi et al., 1997; Mellor et al., 1997; Robinson et al., 1997; Fitzgerald et al., 1998; Nichols et al., 1998; Elbrecht et al., 1999). SPA technology overcomes the disadvantages associated with traditional filtration techniques for receptor assays. No separation step is required and so radioactive handling and waste generation are minimized. Moreover, the procedure can also be applied to study the rate of association and/or dissociation of a radiolabeled ligand from its receptor in real time by simple repeat counting using a limited number of sample wells.
Time Considerations The overall time it takes to develop and conduct an SPA protein interaction assay depends very much on the nature of the application under investigation. When considering configuring an assay using this homogeneous technology, as with other assay approaches, there are a number of defined phases of work that need to be addressed. These include the design, selection and preparation of key assay components, the evaluation of these components followed by assay development, the optimization of assay conditions and format, and finally, assay validation. In terms of the time taken to prepare and evaluate key biological components, this will vary considerably depending on the nature of the protein (or other component) being isolated. For example, it may take anywhere from 1 to 6 months to clone, express, and purify a particular receptor protein of interest that exhibits the appropriate binding characteristics. This assumes of course that the receptor is not available commercially. In light of this, it is not possible to assign typical time scales to these exploratory phases of the process; however, once the key biological components have been successfully sourced, predicting the time taken to optimize an assay becomes more straightforward. Again, using a receptor-ligand binding SPA assay as an example, configuration of a working assay with the desired signal/background ratio typically takes anything from a few weeks in a straightforward case, up to several months for a more difficult assay. As far as actually conducting an SPA is concerned, as detailed in the basic protocols given, this also can be broken down into a number of defined steps as follows.
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Assay set-up Manually pipetting volumes of each component (typically four) into each well of a typical 96-well microtiter plate or the equivalent number of individual reaction tubes can be completed in a matter of 10 to 15 min. Obviously, if more than one 96-well plate full of samples needs to be analyzed, then the time taken increases proportionately. Where high sample throughput is required, many laboratories now have fully automated dispensing instruments that allow many microplates/tubes to be processed at any one time. Generally, the use of such automated devices can speed up the time taken to dispense assay components into microplates considerably. Incubation of assay components For certain SPA configured assays measuring interactions involving proteins, incubation times (typically at ambient temperature) can be as short as 30 min for reactions that attain equilibrium quickly; however, for some low affinity binding assays, extended incubation times of up to 20 hr are often required and employed. The appropriate time of incubation needs to be determined empirically. As SPA is a homogeneous technique, determining the optimum incubation time for each particular assay is relatively straightforward. As no separation step is required, the wells or tubes can be repeatedly counted at various time points during the incubation to ascertain when equilibrium (and hence the optimum scintillation counts) is reached. Scintillation counting To count all wells of a single 96-well microplate using existing eight-detector microplate scintillation counters typically takes anywhere from ∼15 to 30 min. The time taken will depend on the count time per well preset on the instrument, which is largely determined by the scintillation counts obtained (i.e., the SPA signal) in the assay under investigation. The counting of an equivalent number of individual assay tubes in a more traditional (single detector) scintillation counter will take much longer. For example, at 1 min per tube counting time, 96 samples will be counted in ∼2 hr. Data analysis This will vary depending on the degree of data analysis required. Software packages exist, either as integral parts of counting instrumentation, or as off-the-shelf products that en-
able data to be analyzed quickly and reproducibly. Based on the above, in terms of the overall time it takes to conduct a 96-well/tube SPA (excluding the design and isolation of key biological component phases), this can vary from 1 to 2 hr in the best case scenario and up to 24 hr where longer incubation times are required.
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SPA Technology to Study Biomolecular Interactions
Game, S.M., Rajapurohit, P.K., Clifford, M., Bird, M.I., Priest, R., Bovin, N.V., Nifant’ev, N.E., O’Beirne, G., and Cook, N.D. 1998. Characterisation of selectin mediated adhesion using synthetic glycoconjugate ligands and scintillation proximity assay (SPA). Anal. Biochem. 258:127-135.
Gorman, C., Skinner, R.H., Skelly, J.V., Neidle, S., and Lowe, P.N. 1996. Equilibrium and kinetic measurements reveal rapidly reversible binding of Ras to Raf. J. Biol. Chem. 271:6713-6719. Hamachter, J. and Schaberg, T. 1994. Adhesion molecules in lung diseases. Lung 172:189-213. Hill, C.S. and Treisman, R. 1995. Transcriptional regulation by extracellular signals: Mechanisms and specificity. Cell 80:199-211. Hoffman, R. and Cameron, L. 1992. Characterization of a scintillation proximity assay to detect modulators of transforming growth factor α (TGFα ) binding. Anal. Biochem. 203:70-75. Horton, J.K., Seddon, L.S., Williams, A.S., and Baxendale, P.M. 1998. A method for measuring intracellular cyclic AMP levels with immunoassay technology and novel, cellular lysis reagents. Br. J. Pharmacol. 123:Suppl. 31P. Hurwitz, D.R., Emanuel, S.L., Nathan, M.H., Sarver, N., Ullrich, A., Felder, S., Lax, I., and Schlessinger, J. 1991. EGF induces increased ligand binding affinity and dimerization of soluble epidermal growth factor (EGF) receptor extracellular domain. J. Biol. Chem. 266:2203522043. Jones, A.E., Saksela, K., Game, S.M., O’Beirne, G.O., and Cook, N.D. 1998. Screening assay for the detection of the protein-protein interaction between HIV-1 Nef protein and the SH3 domain of Hck. J. Biomol. Screen. 3:37-41. Juliano, R.L. and Varner, J.A. 1993. Adhesion molecules in cancer: The roles of integrins. Curr. Opin. Cell Biol. 5:812-818. Kahl, S.D., Gygi, T., Eichelberger, L.E., and Manetta, J.V. 1996. A multiple-approach scintillation proximity assay to measure the association between Ras and Raf. Anal. Biochem. 243:282283. Kalish, D.I., Cohen, C.M., Jacobsen, B.S., and Brandon, D. 1978. Membrane isolation on polylysine-coated glass beads. Asymmetry of bound membrane. Biochim. Biophys. Acta 506:97-110. Kienhuis, C.B.M., Geurts-Moespot, A., Ross, A.H., Foekens, J.A., Swinkels, M.J.W.L., Koenders, P.G., Ireson, J.C., and Benraad, T.J. 1992. Scintillation proximity assay to study the interaction of epidermal growth factor with its receptor. J. Recept. Res. 12:389-399. Liou, H.-C. and Baltimore, D. 1993. Regulation of the NF-κB/rel transcription factor and IκB inhibitor system. Curr. Opin. Cell Biol. 5:477-487. Lowe, P.N., Page, M.J., Bradley, S., Rhodes, S., Sydenham, M., Paterson, H., and Skinner, R.H. 1991. Characterization of recombinant human Kirsten-ras (4B) p21 produced at high levels in Escherichia coli and insect baculovirus expression systems. J. Biol. Chem. 226:1672-1678. Mackay, C.R. and Imhof, B. 1993. Cell adhesion in the immune system. Immunol. Today 14:99-102. McPherson, G.A. 1983. A practical computer-based approach to the analysis of radioligand binding experiments. Comput. Programs Biomed. 17:107-113.
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Mellor, G.W., Fogarty, S.J., Shane O’Brien, M., Congreve, M., Banks, M.N., Mills, K.M., Jefferies, B., and Houston, J.G. 1997. Searching for chemokine receptor binding antagonists by high throughput screening. J. Bimol. Screen. 2:153157. Musacchio, A., Wilmanns, M., and Saraste, M. 1994. Structure and function of the SH3 domain. Prog. Biophys. Mol. Biol. 61:283-297. Nagata, Y. and Burger, M.M. 1974. Wheat germ agglutinin. Molecular characteristics and specificity for sugar binding. J. Biol. Chem. 249:31163122. Nichols, J.S., Parks, D.J., Consler, T.G., and Blanchard, S.G. 1998. Development of a scintillation proximity assay for peroxisome proliferator-activated receptor γ ligand binding domain. Anal. Biochem. 257:112-119. O’Beirne, G., Ireson, J.C., Lee, D.Y., and Cook, N.D. 1994. Kinetic analysis of receptor binding by scintillation proximity assay. Can. J. Physiol. Pharmacol. 72:23.1.18. Okada, Y., Copeland, B.R., Mori, E., Tung, M.M., Thomas, W.S., and del Zoppo, G.J. 1994. P-selectin and intracellular adhesion molecule-1 expression after focal brain ischemia and reperfusion. Stroke 25:202-211. Pachter, J.A., Zhang, R., and Mayer-Ezell, R. 1995. Scintillation proximity assay to measure binding of soluble fibronectin to antibody captured α5β1 integrin. Anal. Biochem. 230:101-107. Pai, J.J.-K., Kirkup, M.P., Frank, E.A., Pachter, J.A., and Bryant, R.W. 1999. Compounds capable of generating singlet oxygen represent a source of artifactual data in scintillation proximity assays measuring phosphopeptide binding to SH2 domains. Anal. Biochem. 270:33-40. Patel, S., Harris, A., O’Beirne, G., Cook, N.D., and Taylor, C.W. 1996. Kinetic analysis of inositol trisphosphate binding to pure inositol trisphosphate receptors using scintillation proximity assay. Biochem. Biophys. Res. Commun. 221:821825 Reinscheid, R.K., Nothacker, H.P., Bourson, A., Ardati, A., Henningsen, R..A., Bunzow, J.R., Grandy, D.K., Langen, H., Monsma, F.J. Jr., and Civelli, O. 1995. Orphanin FQ: A neuropeptide that activates an opioidlike G protein-coupled receptor. Science 270:792-794. Robinson, N., Gibson, T.M., Chicarelli-Robinson, M.I., Cameron, L., Hylands, P.J., Wilkinson, D., and Simpson, T.J. 1997. Cochliobolic acid, a novel metabolite produced by Cochliobolus lu-
natus, inhibits binding of TGF-alpha to the EGF receptor in an SPA assay. J. Nat. Prod. 60:6-8. Rosen, M.K., Yamazaki, T., Gish, G.D., Kay, C.M., Pawson, T., and Kay, L.E. 1995. Modular binding domains in signal transduction proteins. Nature 374:477-479. Sermon, B.A., Eccleston, J.F. ,Skinner, R.H., and Lowe, P.N. 1996. Mechanism of inhibition of arachidonic acid of the catalytic activity of Ras GTP-ase-activating proteins. J. Biol. Chem. 271:1566-1572. Sheets, M.P., Warrior, U.P., Yoon, H., Mollinson, K.W., Djuric, S.W., and Trevillyan, J.M. 1998. A high-capacity scintillation proximity assay for the discovery and evaluation of ZAP-70 tandem domain antagonists. J. Biomol. Screen. 3:139144. Siebenlist, U., Franzoso, G., and Brown, K. 1994. Structure, regulation and function of NF-κB. Annu. Rev. Cell Biol. 10:405-455. Skinner, R.H., Picardo, M., Gane, N.M., Cook, N.D., Morgan, L., Rowedder, J., and Lowe, P.N. 1994. Direct measurement of the binding of RAS to neurofibromin using a scintillation proximity assay. Anal. Biochem. 223:259-265. Slater, R. 2001. Radioisotopes in Biology: A Practical Approach. Second Edition. Oxford University Press, Oxford. Sugatani, J., Hughes, K.T., and Saito, K. 1994. Radioimmunoassay for quantitation of PAF in biological samples. J. Lipid Mediators 10:9-10. Tang, Y.S., Davis, A.M., and Kitcher, J.P. 1983. N-succinimidyl propionate characterisation and optimum conditions for use as a tritium labeling reagent for proteins. J. Labeled Compounds and Radiopharm. 20:277. Thanos, D. and Maniatis, T. 1995. NF-κB: A lesson in family values. Cell 80:529-532. Zabel, U., Schreck, R., and Baeuerle, P.A. 1991. DNA binding of purified transcription factor NFkB. J. Biol. Chem. 266:252-260. Zhang, J., Wu, P., Kuvelkar, R., Schwartz, J.L., Egan, R.W., Motasim Billah, M., and Wang, P. 1999. A scintillation proximity assay for human interleukin-5 (hIL-5) high-affinity binding in insect cells coexpressing hIL-5 receptor α and β subunits. Anal. Biochem. 268:134-142.
Contributed by Neil Cook, Alison Harris, Alison Hopkins, and Kelvin Hughes Amersham Biosciences Ltd. Cardiff, United Kingdom
Identification of Protein Interaction
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Identifying Protein Interactions by Hydroxyl-Radical Protein Footprinting
UNIT 19.9
Hydroxyl-radical protein footprinting is a direct method to map areas of a protein that interact with a ligand. Areas of interaction are those whose accessibility to a peptide cleavage reagent (i.e., diffusible hydroxyl radicals) changes when in a molecular complex. Accessibility can be altered as a result of either direct binding or conformational changes induced by the molecular partner (indirect). The procedure for the hydroxyl-radical footprinting of a protein:ligand interaction requires preparation of an end-labeled active test protein (TP; see Basic Protocol 1), subjecting it alone and in complex with a ligand to hydroxyl-radical cleavage (see Basic Protocol 2), resolving the cleaved products, and quantitatively analyzing the gel data (see Basic Protocol 3) against molecular weight standards prepared from the test protein (see Support Protocol). Figure 19.9.1 presents a schematic of this procedure.
GENERATION OF AN ACTIVE RECOMBINANT PROTEIN THAT IS END-LABELED
BASIC PROTOCOL 1
This protocol describes generating recombinant forms of the test protein that can be radioactively end-labeled at either the N- or C-terminus. End-labeling a test protein (TP) is achieved by introducing the recognition sequence (LRRASV) for a heart muscle kinase (HMK) to the N- or C-terminus, followed by phosphorylating it with radioactive
Figure 19.9.1
Flow chart of hydroxyl-radical protein footprinting protocol.
Contributed by Nick Loizos Current Protocols in Protein Science (2004) 19.9.1-19.9.11 C 2004 by John Wiley & Sons, Inc. Copyright
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Figure 19.9.2 Schematic drawings of (A) PK-pET16b and (B) pET1529-PK expression vectors. Features of these vectors include the ampicillin (AMP) resistance gene, lacIq repressor gene, pBR322 origin of DNA replication, and T7 promoter and terminator. PK-pET16b was kindly provided by Dr. O’Donnell (Kelman et al., 1995). Plasmid pET1529-PK was constructed to contain a pET15b background and pET29a multiple cloning site—i.e., the multiple cloning site of pET29a (sequences between the XbaI to Bpu1102I sites) was exchanged with the multiple cloning site of pET15b (sequences between the XbaI to Bpu1102I sites). The HMK coding sequence was then inserted by recombinant PCR directly downstream of and in frame with the His tag.
phosphate (Li et al., 1989). The pET vectors 16b and 29a (Novagen) were modified to code for HMK sites at the N- and C- termini of expressed open reading frames, respectively (Fig. 19.9.2).
Materials
HydroxyRadical Protein Footprinting
DNA insert containing test protein’s open reading frame Modified vectors (available from Nick Loizos on request) of the pET expression system: PK-pET-16b pET1529-pK Heart muscle kinase, bovine (HMK; Sigma): reconstitute in 40 mM dithiothreitol 6000 Ci/mmol [γ-32 P]ATP or 2000 Ci/mmol [γ-33 P]ATP (Perkin Elmer) 1 M Tris·Cl, pH 7.9 (APPENDIX 2E) 5.0 M NaCl (APPENDIX 2E) 1 M MgCl2 (APPENDIX 2E)
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Ni2+ -NTA agarose beads (Qiagen), preequilibrated in binding buffer Binding buffer (see recipe) Elution buffer (see recipe) 1.5-ml centrifuge tube 30◦ C water bath or heatblock Microcon 10 microconcentrator (Amicon) Additional reagents and equipment for restriction digestion (APPENDIX 4I), transformation of E. coli (APPENDIX 4D), metal chelate affinity chromatography (MCAC; UNIT 9.4), and appropriate activity assay for the protein of interest Construct vectors 1. Insert the test protein’s open reading frame (ORF) between the NdeI and BamHI restriction sites in PK-pET16b to generate a vector that directs production of the protein with an N-terminal HMK site and His6 tag, with the ORF in frame with the His6 tag (HMK-His6 -TP; APPENDIX 4I). 2. Also insert the ORF between the NdeI site and an appropriate restriction site(s) in pET1529-PK to generate a vector that directs the production of the protein with a C-terminal HMK site and His6 tag (TP-His6 -HMK; APPENDIX 4I), with the ORF in frame with the His6 tag.
Check activity of the recombinant forms of the test protein 3. Prepare and perform separate transformations of E. coli with each of the vectors (APPENDIX 4D). Induce expression as per instructions from Novagen. 4. Purify the proteins by Ni2+ column chromatography (MCAC; UNIT 9.4). 5. Check the activity of both HMK-His6 -TP and TP-His6 -HMK using an appropriate activity assay. Purified HMK-His6 -TP and/or TP-His6 -HMK must behave as the wild-type parent during in vitro assays for the protein. It must be shown that the terminal modifications to the test protein do not disrupt folding or interactions with the ligand. How this is shown will depend on the binding and/or activity assays available in each specific case. The presence of aggregates or possibly a fraction of misfolded proteins in a preparation of the derivative could affect cleavage susceptibility to hydroxyl radicals and thereby complicate the analysis of cleavage profiles. It is best to use refolded proteins in footprinting experiments only when an activity-based purification step is available (e.g., an affinity column using the ligand to be tested in the footprinting experiment).
End-label protein 6. In a 1.5-ml centrifuge tube, end-label 40 µg of the test protein (HMK-His6 -TP or TPHis6 -HMK) with 32 P using the following 400-µl reaction (5 µM final concentration): 100 U reconstituted HMK 0.02 µM [γ-32 P]ATP (6000 Ci/mmol) or [γ-33 P]ATP (2000 Ci/mmol) 20 mM Tris·Cl, pH 7.9 (dilute from 1 M stock) 0.5 M NaCl (dilute from 5.0 M stock) 12 mM MgCl2 (dilute from 1 M stock). Incubate 15 min at 30◦ C. HMK is also known as the protein kinase A catalytic subunit (bovine heart). Either 33 P or 32 P can be used. The lower energy beta particles arising from 33 P decay (0.248 MeV) compared with 32 P (1.71 MeV) do not penetrate and scatter as far into the detecting medium, resulting in sharper bands; however, 32 P gives a stronger signal, should a low amount of protein be used in the footprinting reaction.
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7. To remove free label, add 20 µl of Ni2+ -NTA agarose beads, preequilibrated in binding buffer, to the reaction. Mix gently and incubate 15 min on ice. Pellet beads by microcentrifuging 3 sec at maximum speed. 8. Wash by adding 1 ml binding buffer and inverting the tube three times. Centrifuge 2 to 3 sec (e.g., in a tabletop microcentrifuge). Decant supernatant. Repeat once. 9. Resuspend beads in 200 µl elution buffer and incubate 10 min on ice. Microcentrifuge 10 sec at maximum speed. Discard pellet. 10. Add 300 µl binding buffer to the supernatant and apply to a Microcon 10 microconcentrator. Concentrate sample to 100 µl according to manufacturer’s directions (also see UNIT 4.4). Add 400 µl binding buffer and reconcentrate to reduce imidazole concentration to <20 mM. The yield should be ∼6 µg end-labeled protein in 100 µl. BASIC PROTOCOL 2
HYDROXYL RADICAL CLEAVAGE OF PROTEIN IN ABSENCE AND PRESENCE OF LIGAND This protocol describes generating hydroxyl radicals by the Fenton reaction (Kim et al., 1985) and using them to cleave the peptide backbone. The free test protein as well as test protein in complex with a ligand are subjected to cleavage under single-hit conditions (Brenowitz et al., 1986).
Materials End-labeled protein (see Basic Protocol 1) Ligand Footprinting reaction buffer (see recipe) Iron-EDTA solution (see recipe) 0.2 M sodium ascorbate in footprinting reaction buffer 10 mM hydrogen peroxide 3× loading buffer (see recipe) 1. Add 25 pmol end-labeled protein alone or with 25 pmol ligand to appropriately labeled tubes. Adjust total volume to 14 µl per tube with footprinting reaction buffer. Incubate 5 min at 25◦ C to allow for complex formation. Alternative buffers can be used but should not contain radical scavengers (e.g., Tris, glycerol; Heyduk et al., 1996). MgCl2 can be omitted if necessary to prevent Mg2+ dependent activation of proteins during the reaction (Loizos and Darst, 1999).
2. Add 2 µl iron-EDTA solution, 2 µl of 0.2 M sodium ascorbate in footprinting reaction buffer, and 2 µl of 10 mM hydrogen peroxide. Initiate reactions simultaneously by spotting cleavage reagents on the inside lid of the test tubes followed by briefly centrifuging. Incubate reactions 20 min at 25◦ C. The final concentrations of cleavage and protein components are: 1 mM (NH4 )2 Fe(SO4 )2 , 2 mM EDTA, 20 mM ascorbate, 1 mM H2 O2 , and 1.25 to 2.5 µM test protein and ligand. The concentration of the components must leave a large fraction of the molecules uncut, thus satisfying the need for single-hit conditions during footprinting (Brenowitz et al., 1986).
HydroxyRadical Protein Footprinting
3. Terminate by adding 10 µl of 3× loading buffer. To end reactions simultaneously, spot 3× loading buffer on the inside lid of the test tubes followed by briefly centrifuging.
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ANALYZE GEL DATA Hydroxyl-radical cleaved products and molecular weight standards are subjected to polyacrylamide gel electrophoresis. Digital images of the footprinting gels are then quantitatively analyzed and the results presented in the form of a difference plot. This protocol describes how to (1) make a high percentage tricine-polyacrylamide gel, (2) generate intensity plots for each gel lane, (3) align them to each other to correct for lane-to-lane distortions, (4) subtract intensity values for free TP from the TP/ligand complex, and (5) graph the differences versus amino acid number.
BASIC PROTOCOL 3
Materials 33%T, 3%C polyacrylamide, ultrapure (ICN) 2 M Tris·Cl, pH 7.9 (APPENDIX 2E)/0.3% (w/v) SDS: store up to 6 months at room temperature 80% glycerol 10% (w/v) ammonium persulfate TEMED Anode buffer (see recipe) Cathode buffer (see recipe) Cleaved test protein (TP) or test protein:ligand (TP:ligand) complex (see Basic Protocol 2) Molecular weight standards (see Support Protocol) Vertical electrophoresis apparatus PhosphorImager (Storm model from Molecular Dynamics) or equivalent IMAGEQUANT software (Molecular Dynamics) ALIGN software (available upon request from T. Heyduk; [email protected]) Excel software (Microsoft) or equivalent spreadsheet program Set up tricine SDS-PAGE system 1. Prepare a Tricine/SDS 16.5%T, 3%C polyacrylamide separating gel by combining the following components: 17.5 ml 33% T, 3%C polyacrylamide 11.69 ml 2 M Tris·Cl(pH 7.9)/0.3% (w/v) SDS 1.19 ml H2 O 4.62 ml 80% glycerol 300 µl of 10% (w/v) ammonium persulfate 20 µl TEMED. 2. Prepare a 6% T, 3% C stacking gel by combining the following components:
4.5 ml of 33%T, 3%C polyacrylamide 6.2 ml 2 M Tris·Cl(pH 7.9)/0.3% (w/v) SDS 14.3 ml H2 O 100 µl of 10% (w/v) ammonium persulfate 10 µl TEMED. The gel dimensions are 14 cm × 16 cm × 0.8 mm.
3. Place the gel in a vertical electrophoresis apparatus and add anode and cathode buffers. 4. Load cleaved TP and TP:ligand reactions, and molecular weight standards onto the gel. Run 20 min at 14 mAmp. Run an additional 16 hr at 20 mAmps. Identification of Protein Interactions
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The hydroxyl-radical cleavage products are run on the same gel as a set of molecular weight markers and radical cleavage sites assigned by interpolation between the known cleavage sites of the markers.
5. Dry gel 2 hr with a slow ramp up speed to 80◦ C. 6. Obtain a digital image of the gel using a phosphorimager or equivalent instrument.
Align intensity plots 7. To generate intensity plots, use the program IMAGEQUANT to make a line down the center of each gel lane. Widen the line using the Object Attributes menu item until the entire lane is bracketed. 8. Use the Create Graph command to integrate phosphorimager intensities across full lane widths and plot them versus electrophoretic mobilities (pixels). 9. To correct for lane-to-lane distortions, align intensity plots using the program ALIGN. ALIGN was written in Basic by G. Lubecki and T. Heyduk, and uses a linear interpolation to adjust peaks of one plot to homologous peaks in another. Choose one plot (gel lane) as a reference to which all others will be aligned. The lanes containing molecular weight standards should only be aligned with the band corresponding to the uncleaved protein.
Generate difference plots 10. Import the aligned intensity scans, excluding portions corresponding to the uncleaved full-length protein, into a spreadsheet program (e.g., Excel). Place pixel numbers in column A of the spreadsheet and the corresponding aligned intensity values for cleaved TP and TP:ligand complex into columns B and C, respectively. In addition to obtaining the difference between TP:ligand and TP plots, the following steps correct for gel loading and cleavage efficiencies between the samples.
11. Calculate the ratios of the intensity values for each pixel in columns B and C (i.e., TP/TP:ligand) and report the values in column D of the spreadsheet. 12. Round the values in column D (TP/TP:ligand ratios) to a single digit and place in column E. Calculate the mode of these rounded values. 13. Multiply the values in column C (TP:ligand intensities) by the mode of the values in column E, then subtract the values in column B (TP intensities) and divide the difference by the additive values of columns B and C (TP and TP:ligand intensities, respectively) multiplied by the mode of the values in the column E:
Place these values in column G. 14. Assign each of the bands of standards in the gel an amino acid number, based on where the endoproteinase or CNBr cuts, and a pixel number. Pixel number is defined as the distance from the beginning of the line made to generate the intensity plot. Pixel numbers start from lowest to highest and amino acid numbers highest (full-length) to lowest available marker.
15. Place the standard pixel and amino acid numbers into columns I and J, respectively. The next series of steps convert pixel number to amino acid number. Knowledge of the residue specificity for each endoproteinase, as well as the primary amino acid sequence HydroxyRadical Protein Footprinting
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for TP, allows one to determine how many bands (i.e., molecular weight standards) would be generated under limited cleavage conditions and their mobilities relative to each other.
16. Determine the log of the values in the column J (standard amino acid numbers) and place in column K. 17. Use the data in column I (standard pixel numbers) and K (log standard amino acid numbers) columns to prepare a standard curve of log(Mr ) versus relative mobility. 18. Find the slopes and intercepts for the lines between the data points of the standard curve and place in columns L and M, respectively. It is necessary to use multiple slopes and intercepts given that the relationship between gel migration and peptide molecular weight is not linear.
19. The pixel values in column A can then be converted to amino acid number using the following formula: Interpolated amino acid no. = [(slope × pixel no.) + intercept]10 . Place interpolated amino acid values in column N. 20. Use the values in columns N (interpolated amino acid no.) and G (normalized difference) to generate a difference plot. 21. Average difference plots from different gel lanes and from different gels (interpolated to a reference gel). 22. Test the statistical significance of the differences between the (TP:ligand complex – TP) experiments using a Student’s t test for small samples with unequal variances.
PREPARATION OF MOLECULAR WEIGHT STANDARDS This protocol presents instructions on the generation of molecular weight markers derived from test protein cleaved with endoproteinases and chemical cleavage reagents. These molecular weight markers are used to convert gel mobilities of the hydroxyl-radical cleaved products into amino acid cleavage sites (see Basic Protocol 3; Heyduk et al., 1996).
SUPPORT PROTOCOL
Materials HMK-His6 -TP or TP-His6 -HMK (see Basic Protocol 2) Urea 1 M Tris·Cl, pH 9.0 (APPENDIX 2E) Endoproteinase Lys-C, Glu-C, or Asp-N (Sigma) 1× and 3× loading buffer (see recipe) CNBr (Aldrich) 20% SDS (APPENDIX 2E) 1 M HCl 1. Prepare endoproteinase reaction mixtures (30 µl total volume) containing the following:
1.25 µM HMK-His6 -TP or TP-His6 -HMK 8 M urea 50 mM Tris·Cl, pH 9.0 (dilute from 1 M stock) 0.25 µg Lys-C, 0.5 µg Glu-C, or 0.5 µg Asp-N. Incubate 15 min at 25◦ C. Identification of Protein Interactions
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2. Terminate reactions by adding 10 µl of 3× loading buffer. 3. Prepare methionine-specific cleavage reactions (20 µl total):
1.25 µM HMK-His6 -TP or TP-His6 -HMK 400 mM CNBr 0.4% SDS (dilute from 20% stock). Adjust to pH 2 with 1 M HCl. Incubate 20 min at 25◦ C. 4. Terminate by lyophilization and addition of 30 µl of 1× loading buffer. Endoproteinases Lys-C, Glu-C, or Asp-N are specific for lysine, glutamate, and aspartate residues, respectively. The chemical cleavage reagent cyanogen bromide is specific for methionine. Depending on the sequence of the test protein, other chemical cleavage reagents, such as BNPS skatole and 2-nitro-5-thiocyanobenzoic acid (specific for tryptophan and cysteine, respectively), may be used to generate molecular weight markers (Borukhov et al., 1991). The cleavage reactions can be stored at −20◦ C and loaded onto the gel at a later time; however, waiting longer than a few days could lead to radioactive decay of the peptide fragments.
REAGENTS AND SOLUTIONS Anode buffer Dissolve 24.22 g Tris base in 100 ml water. Adjust to pH 8.9 with HCl and then dilute to 1 liter with H2 O. Store up to 6 months at room temperature. Binding buffer 10 mM MOPS, pH 7.2 (adjust pH with NaOH) 500 mM NaCl 10 mM MgCl2 Store up to 1 month at 4◦ C Cathode buffer Dissolve the following in 1 liter H2 O: 12.11 g Tris base 17.92 g Tricine 1 g SDS Store up to 6 months at room temperature Elution buffer Prepare binding buffer (see recipe) containing 250 mM imidazole added from a 1 M stock solution (adjust stock to pH 7.5 with concentrated HCl). Store up to 1 month at 4◦ C. Footprinting reaction buffer 10 mM MOPS, pH 7.2 (adjust pH with NaOH) 200 mM NaCl 10 mM MgCl2 Store up to 1 months at 4◦ C
HydroxyRadical Protein Footprinting
Iron-EDTA solution Combine 25 µl of 40 mM ammonium iron (II) sulfate hexahydride in water, 25 µl of 80 mM EDTA in footprinting reaction buffer, and 50 µl footprinting buffer (see recipe). Prepare just before use.
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Loading buffer, 3× 150 mM Tris·Cl, pH 7.9 (APPENDIX 2E) 36% glycerol 12% SDS 6% 2-mercaptoethanol 0.01% bromophenol blue Store up to 1 year at −20◦ C COMMENTARY Background Information The hydroxyl-radical protein footprinting procedure presented here was developed by Heyduk & Heyduk (Heyduk and Heyduk, 1994; Heyduk et al., 1996) and has been the subject of recent reviews (Loizos and Darst, 1998; Heyduk et al., 2001). Using this procedure, data has been obtained for numerous protein:ligand complexes (Heyduk and Heyduk, 1994; Nagai and Shimamoto, 1997; Wang et al., 1997; Colland et al., 1998; Mossessova et al., 1998; Casaz and Buck, 1999; Loizos and Darst, 1999). Footprinting involves (1) endlabeling a protein with radioactivity (Li et al., 1989), (2) using hydroxyl radicals generated by the Fenton reaction (Kim et al., 1985) as a general, peptide backbone cleavage reagent to cleave the protein and protein:ligand complex, (3) resolving and identifying the cleavage products in a tricine SDS-PAGE system developed for the resolution of small polypeptides (Schagger and von Jagow, 1987), and (4) quantitatively analyzing the gel data (also see Fig. 19.9.1). Researchers often map sites involved in protein interactions by making point mutations or deletions and then testing these recombinants in binding assays. This approach, however, can result in misfolding or distal effects of the recombinants due to alterations in amino acid sequence. Hydroxyl-radical footprinting offers the advantage of using a recombinant protein containing the native sequence in solution as opposed to mutant or truncated forms for forming complexes. Hydroxyl-radical footprinting also offers the advantage of detecting conformational changes in a protein when bound by a ligand (e.g., see Baichoo and Heyduk, 1999). Theoretically, mass spectrometry could be an alternative to gel electrophoresis as a read-out for the cleavage products; however, experiments using synchrotron X-ray– generated radicals show that amino acid side chain modifications occur at a faster rate (greater than two orders of magnitude) than backbone cleavage (Maleknia et al., 1999). Modification to side chains would make mass
spectrometric analysis of hydroxyl-radical cleavage products very difficult.
Critical Parameters Before beginning any footprinting project, it is important to show that the wild-type test protein is not phosphorylated by the kinase at an internal cryptic site. If a cryptic site is present, phosphorylation conditions may be found that eliminate or reduce labeling to acceptably low levels compared to the usually robust labeling of the cognate site for the kinase. It has been found that cleavage products shorter than ∼25 amino acids in length are lost during electrophoresis (Heyduk et al., 1996); therefore, adding approximately twenty nonnative amino acids between the kinase site and the beginning of the protein can help resolve footprints at the termini of test proteins. Such was the case with footprinting GreB on a transcription elongation complex (Loizos and Darst, 1999).
Troubleshooting Unlike the situation for DNA footprinting, the footprint on the test protein resulting from the bound ligand may not be immediately apparent from observation of the imaged gel. The reason for this is mainly because single amino acid resolution is not achieved by the SDS-PAGE analysis. Thus, careful quantitative analysis of the phosphorimager intensities is required for the footprint to be fully revealed. Gel mobility of proteins is not related in a simple way to the molecular weight; thus, small protein fragments may show atypical mobility in SDS-PAGE. Therefore, calibration of gel mobility using molecular weight markers generated from the test protein itself (see Support Protocol) is essential to accurately convert the gel mobilities into amino acid cleavage sites. Usually, a cleavage site can be assigned to plus or minus three to five amino acids using the gel dimensions given (Heyduk et al., 2001).
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Given that nucleic acids are more susceptible than proteins to hydroxyl radical cleavage, care must be taken that the ligand and/or complex is not altered during the footprinting reaction. This can be done by testing for the functionality/integrity of the ligand under the cleavage conditions (e.g., see Loizos and Darst, 1999).
Anticipated Results A footprinting gel image and difference plot is shown in Figure 19.9.3. This experimental data is for the interaction between transcription factor GreB and E. coli RNA polymerase (Loizos and Darst, 1999). Regions of statis-
HydroxyRadical Protein Footprinting
tically significant protection (negative values) and enhancement (positive values) of radical cleavage of the transcription factor GreB by the binding of core RNA polymerase are observed. Protection of a site on the test protein from hydroxyl radical cleavage by the presence of a ligand can be most simply interpreted in two ways: from a direct association of the ligand with that site (blocking access by hydroxylradicals) or from a conformational change that altered the local environment, making cleavage less likely. Enhancement of cleavage in the presence of the ligand would be most likely to result from a conformational change that made cleavage more likely. It is tempting to
Figure 19.9.3 Hydroxyl radical footprint of core RNAP (ligand) on GreB (test protein). (A) phosphorimaged gel of hydroxyl-radical cleaved GreB and GreB-Core complex. Enhanced and protected areas are marked with arrows and brackets, respectively. (B) Top part of phosphorimaged gel from experiment in which gel run time was extended and GreB was labeled with 33 P. Asterisk marks additional protected area not seen in (A) due to masking by the intensity of the uncleaved product. (C) Difference plot showing normalized intensity difference, (Icomplex − IGreB )/(Icomplex + IGreB ), plotted against residue number (Icomplex and IGreB are the corrected intensities for the GreBcore complex and GreB, respectively). Statistically significant differences according to a Student’s t test (confidence level of 0.99999) are denoted by black bars.
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interpret protection from cleavage as being due to a direct interaction with the ligand. The access to information on conformational changes is a powerful feature of the protein footprinting method, but the effects of conformational changes on hydroxyl-radical cleavage make this direct interpretation less secure. Additional information, such as point substitutions within a protected region that affect the binding of the ligand, can bolster this interpretation. Alternatively, the results of the protein footprinting analysis can direct the generation of point substitutions to regions found to be affected by ligand binding.
Time Considerations The most time consuming part of this procedure is obtaining a test protein that can be radioactively end-labeled. That cloning, expression, and functional testing can take 1 to 2 months. The footprinting procedure itself should only take 2 weeks and 2 additional weeks for the quantitative analysis.
Literature Cited Baichoo, N. and Heyduk, T. 1999. DNA-induced conformational changes in cyclic AMP receptor protein: Detection and mapping by a protein footprinting technique using multiple chemical proteases. J. Mol. Biol. 290:37-48. Borukhov, S., Lee, J., and Goldfarb, A. 1991. Mapping of a contact for the RNA 3 terminus in the largest subunit of RNA polymerase. J. Biol. Chem. 266:23932-23935. Brenowitz, M., Senear, D.F., Shea, M.A., and Ackers, G.K. 1986. Quantitative DNase footprint titration: A method for studying protein-DNA interactions. Methods Enzymol. 130:132-181. Casaz, P. and Buck, M. 1999. Region I modifies DNA-binding domain conformation of sigma 54 within the holoenzyme. J. Mol. Biol. 285:507514. Colland, F., Orsini, G., Brody, E.N., Buc, H., and Kolb, A. 1998. The bacteriophage T4 AsiA protein: A molecular switch for sigma 70-dependent promoters. Mol. Microbiol. 27:819-829. Heyduk, E. and Heyduk, T. 1994. Mapping protein domains involved in macromolecular interactions: A novel protein footprinting approach. Biochemistry 33:9643-9650.
Heyduk, T., Baichoo, N., and Heyduk, E. 2001. Hydroxyl radical footprinting of proteins using metal ion complexes. Met. Ions Biol. Syst. 38:255-287. Kelman, Z., Yao, N., and O’Donnell, M. 1995. Escherichia coli expression vectors containing a protein kinase recognition motif, His6-tag and hemagglutinin epitope. Gene 166:177-178. Kim, K., Rhee, S.G., and Stadtman, E.R. 1985. Nonenzymatic cleavage of proteins by reactive oxygen species generated by dithiothreitol and iron. J. Biol. Chem. 260:15394-15397. Li, B.L., Langer, J.A., Schwartz, B., and Pestka, S. 1989. Creation of phosphorylation sites in proteins: Construction of a phosphorylatable human interferon alpha. Proc. Natl. Acad. Sci. U.S.A. 86:558-562. Loizos, N. and Darst, S.A. 1998. Mapping proteinligand interactions by footprinting, a radical idea. Structure 6:691-695. Loizos, N. and Darst, S.A. 1999. Mapping interactions of Escherichia coli GreB with RNA polymerase and ternary elongation complexes. J. Biol. Chem. 274:23378-23386. Maleknia, S.D., Brenowitz, M., and Chance, M.R. 1999. Millisecond radiolytic modification of peptides by synchrotron X-rays identified by mass spectrometry. Anal. Chem. 71:3965-3973. Mossessova, E., Gulbis, J.M., and Goldberg, J. 1998. Structure of the guanine nucleotide exchange factor Sec7 domain of human arno and analysis of the interaction with ARF GTPase. Cell 92:415-423. Nagai, H. and Shimamoto, N. 1997. Regions of the Escherichia coli primary sigma factor sigma70 that are involved in interaction with RNA polymerase core enzyme. Genes Cells 2:725-734. Schagger, H. and von Jagow, G. 1987. Tricinesodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa. Anal. Biochem. 166:368-379. Wang, Y., Severinov, K., Loizos, N., Fenyo, D., Heyduk, E., Heyduk, T., Chait, B.T., and Darst, S.A. 1997. Determinants for Escherichia coli RNA polymerase assembly within the beta subunit. J. Mol. Biol. 270:648-662.
Contributed by Nick Loizos ImClone Systems Incorporated New York, New York
Heyduk, T., Heyduk, E., Severinov, K., Tang, H., and Ebright, R.H. 1996. Determinants of RNA polymerase alpha subunit for interaction with beta, beta , and sigma subunits: Hydroxylradical protein footprinting. Proc. Natl. Acad. Sci. U.S.A. 93:10162-10166.
Identification of Protein Interactions
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Visualization of Protein Interactions in Living Cells Using Bimolecular Fluorescence Complementation (BiFC) Analysis
UNIT 19.10
Protein interactions control many cellular functions. The functional characteristics of many proteins are determined by the proteins that they interact with in each cell. Interactions with different partners and in response to different extracellular signals can occur in different subcellular locations. Determination of the locations of protein complexes and their regulation can provide insight into the functional roles of the protein interactions. This unit describes the bimolecular fluorescence complementation (BiFC) assay for the visualization of protein interactions in living cells (Hu et al., 2002). This approach is based on complementation between nonfluorescent fragments of the yellow fluorescent protein (YFP) when they are brought together by an interaction between proteins fused to each fragment (Fig. 19.10.1). The BiFC assay allows direct visualization of the subcellular locations of protein interactions in living cells (see Basic Protocol). This approach can be used for the analysis of many protein interactions and does not require information about the structures of the interaction partners. Most proteins have a large number of alternative interaction partners in each cell. Many of these interactions are mutually exclusive, resulting in competition for shared interaction partners in the cell. To study the competition for interactions between alternative interaction partners in living cells, the authors of this unit have developed a multicolor BiFC assay by fusing fragments of different fluorescent proteins to the interaction partners (see Alternate Protocol 1). The multicolor BiFC assay is based on the use of fragments of fluorescent proteins with distinct spectral characteristics. Bimolecular complexes formed between these fragments can be visualized using different excitation and emission wavelengths, enabling parallel visualization of multiple interactions in the same cell. The
Figure 19.10.1 Schematic representation of the principle of the BiFC assay Two fragments (YN and YC) of the yellow fluorescent protein (YFP) are fused to putative interaction partners (A and B). The association of the interaction partners allows formation of a bimolecular fluorescent complex. This black-and-white facsimile of the figure is intended only as a placeholder; for a full-color version of the figure go to http://www.interscience.wiley.com/c p/colorfigures.htm.
Contributed by Chang-Deng Hu, Asya V. Grinberg, and Tom K. Kerppola Current Protocols in Protein Science (2005) 19.10.1-19.10.21 C 2005 by John Wiley & Sons, Inc. Copyright
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multicolor BiFC assay allows comparison of the subcellular locations of different protein interactions in the same cell, as well as analysis of the competition among mutually exclusive interaction partners for complex formation with a common partner (Alternate Protocol 2). NOTE: Cells are cultured in a humidified 37◦ C, 5% CO2 incubator. Cells can be incubated in a 30◦ C, 5% CO2 incubator to promote maturation of the fluorophore and thereby increase the signal (see Commentary). Cells are maintained under conditions required for each line. BASIC PROTOCOL
VISUALIZATION OF PROTEIN INTERACTIONS IN LIVING CELLS The BiFC assay allows direct visualization of protein interactions in living cells based on complementation between nonfluorescent fragments of YFP or other fluorescent proteins.
Materials Plasmid vectors for expression of proteins of interest (also see Critical Parameter): e.g., pFlag-CMV2 (Sigma) or pHA-CMV (Clontech) DNA encoding amino acid residues 1 to 154 of yellow fluorescent protein (YN fragment; also see Critical Parameters and Table 19.10.1) DNA encoding amino acid residues 155 to 238 of yellow fluorescent protein (YC fragment; also see Critical Parameters and Table 19.10.1) Linkers (see Critical Parameters) DNA encoding proteins (interaction partners, wild-type) of interest DNA encoding mutated, single amino acid–substitution variants of the protein of interest that do not interact with each other (controls) Cells that can be transfected using plasmid DNA (preferably adherent, monolayer cell line) Appropriate culture medium FuGENE 6 (Roche Diagnostics) or other transfection reagent Tissue culture vessels appropriate for experiment: e.g., cluster plates, slide chambers, or glass coverslips Inverted fluorescence microscope equipped with: Sensitive CCD camera 20× to 100× objectives Filters for visualization of YFP (excitation 500 ± 10 nm; emission 535 ± 15 nm) Software for instrument control and data analysis Additional reagents and equipment for expressing proteins in mammalian cells (UNIT 5.10), mammalian cell culture (APPENDIX 3C), and immunoblotting (UNIT 10.10)
Table 19.10.1 Selection of Fluorescent Protein Fragments for BiFC and Multicolor BiFC Analyses
Bimolecular Fluorescence Complementation
Fluorescent protein fragments
Applications
Filters
YN155+YC155
BiFC analysis
YFP
YN173+YC173
BiFC analysis
YFP
CN155+CC155
BiFC analysis
CFP
YN155+CN155+CC155
Multicolor BiFC analysis
CFP YFP
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Construct plasmid vectors 1. Using the appropriate plasmid vectors, construct expression vectors by fusing the sequences encoding amino acid residues 1 to 154 (YN) and residues 155 to 238 (YC) of the yellow fluorescent protein to sequences encoding the proteins of interest. Whenever possible, test fusions to both the N- and C-terminal ends of the proteins to be investigated. As controls, construct plasmids encoding mutated variants of the proteins that do not interact with each other. For additional information about the design of fusion constructs, see Critical Parameters. The chimeric coding regions can be expressed in mammalian cells using vectors such as pFlag-CMV2 (Sigma) or pHA-CMV (Clontech). As an alternative, fragments of the cyan fluorescent protein (CFP) may be used (see Table 19.10.1).
Prepare cells 2. Seed cells the day before transfection at a density that will allow for cell proliferation over the course of the experiment and, at the same time, allow for the effects of cell growth and density on the interaction under investigation. The BiFC assay can be used for the analysis of protein interactions in many mammalian cell lines. The authors have studied protein interactions using the BiFC assay in COS-1, HEK293, HeLa, Hep3B, αTN4, and NIH3T3 cells (Hu et al., 2002; Grinberg et al., 2004; Rajaram and Kerppola, 2004).
Figure 19.10.2 Multiple combinations of fusion proteins should be tested for bimolecular fluorescence complementation. Amino- and carboxyl-terminal fusions can be used to test eight distinct combinations (A to H). This black-and-white facsimile of the figure is intended only as a placeholder; for a full-color version of the figure go to http://www.interscience.wiley.com/c p/colorfigures.htm.
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Cluster plates are convenient for processing multiple transfections in parallel. If shortworking-distance objectives will be used to visualize the interaction, the cells should be grown in slide chambers or on glass coverslips. APPENDIX 3C
describes basic mammalian cell culture techniques.
3. When cells reach appropriate confluence (e.g., ∼50%), transfect cells with appropriate amounts (e.g., 0.25 µg) of the BiFC plasmid (from step 1) encoding the fusion proteins. As controls, transfect cells in parallel with the plasmids encoding mutated variants of the proteins that do not interact with each other. FuGENE 6 from Roche Diagnostics is used according to the manufacturer’s instructions to introduce plasmid DNA into cells, as it has low background fluorescence and requires minimal manipulation of the cells. Users need to follow specific protocols and optimize the conditions of transfection if other transfection reagents are used.
4. Grow cells until fluorescence is detected (12 to 36 hr). If necessary, incubate the cells at 30◦ C with a 5% CO2 atmosphere to promote maturation of the fluorophore and to increase the signal (however, interpret results obtained under such conditions with care, as incubation at a lower temperature could alter protein interactions). Compare the levels of fusion protein expression with those of the endogenous proteins by immunoblot analysis (see UNIT 10.10).
Examine cells 5. Wash the cells with PBS to remove dead cells and debris, then add fresh medium. 6. View the cells using an inverted fluorescent microscope with a 20× long-workingdistance objective and excitation at 500 ± 10 nm with emission detection at 535 ± 15 nm. Confirm that fluorescent cells are alive and exhibit normal morphology compared to nontransfected cells. Higher-numerical-aperture objectives may be necessary to visualize weak signals and higher magnification may be helpful to determine the subcellular localization. Cells grown on coverslips can be fixed and individual proteins can be visualized by indirect immunofluorescence analysis. If CFP is used as the reporter, the excitation wavelength is 436 ± 5 nm and the emission wavelength is 470 ± 15 nm.
Assess results 7. Compare results obtained with cells transfected with the wild-type interaction proteins and those transfected with the control, mutant constructs with single amino acid substitutions that prevent interactions. Compare the number of fluorescent cells observed in cultures expressing fluorescent protein fragment fusions with that observed in cultures expressing intact fluorescent proteins. Fluorescence in a small subpopulation of cells is difficult to interpret, since it may represent complementation due to nonspecific interactions, or it may reflect a regulated interaction that occurs only in response to signaling in a subset of cells. The following conclusions may be drawn: Specific interaction: If fluorescence is detected when wild-type proteins are expressed, and this signal is eliminated or significantly reduced by single amino acid substitutions that prevent the interaction, it is likely that the bimolecular fluorescence complementation represents a specific interaction between the proteins fused to the fragments of fluorescent proteins. Bimolecular Fluorescence Complementation
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Nonspecific interaction: If mutations that are known to eliminate the interaction do not reduce or eliminate the fluorescence, then the bimolecular complementation is due to nonspecific interactions between the fusion proteins. If this is the case, the BiFC assay may not be an appropriate assay for the study of the two proteins. Alternatively, a different combination of fusion proteins or linkers should be tested. No fluorescence complementation detected: The lack of fluorescence complementation in the BiFC assay does not prove the absence of an interaction, even if coexpression of the same fusion proteins with other interaction partners results in bimolecular fluorescence complementation. Factors that may contribute to this are discussed in the Commentary. Different combinations of BiFC constructs or alternative approaches should be employed.
QUANTIFICATION OF FLUORESCENCE COMPLEMENTATION EFFICIENCY
SUPPORT PROTOCOL 1
Quantification of the efficiency of fluorescence complementation is useful for comparison of interactions involving closely related proteins. It is particularly important for determining whether fluorescence complementation represents a specific interaction by comparison of wild-type and mutated proteins. The relative efficiencies of bimolecular fluorescence complementation by structurally unrelated proteins cannot be used to determine the efficiencies of complex formation, since many factors unrelated to the efficiency of complex formation influence the efficiency of bimolecular fluorescence complementation.
Additional Materials (also see Basic Protocol) Plasmid encoding full-length CFP (assuming YFP fragments are used in the Basic Protocol) or other spectrally distinguishable fluorescent protein Filters for visualization of YFP (excitation 500 ± 10 nm; emission 535 ± 15 nm) and CFP (excitation 436 ± 5 nm; emission 470 ± 15 nm) fluorescence 1. Follow steps 1 to 5 of the Basic Protocol, but add 50 ng of a plasmid encoding full-length CFP or another spectrally distinguishable fluorescent protein into each transfection. 2. View cells with a fluorescence microscope using filters optimized for separation of CFP and YFP signals. Collect images of at least 100 cells that show detectable fluorescence using both filter sets. 3. Quantify the fluorescence intensities using both YFP and CFP filters integrated over the area of each cell. Subtract the background in an area of the field containing no cells. 4. Calculate the ratio of YFP/CFP fluorescence for each cell. This represents the complementation efficiency for the combination of proteins expressed in individual cells.
5. Plot the data in a histogram by binning cells with similar ratios as shown in Fig. 19.10.3. 6. Assess result. If a mutation significantly reduces the YFP/CFP ratio compared to the wild-type protein (but the levels of expression and localization of the proteins are not altered), the observed fluorescence is likely to represent a specific interaction between the proteins. If the ratios for wild-type and mutated proteins are similar, the observed fluorescence signal may represent nonspecific fluorescence complementation, or the mutation may not affect the interaction between the proteins.
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Figure 19.10.3 The effects of mutations that eliminate the interaction interface should be tested on the efficiency of bimolecular fluorescence complementation (data adapted from Hu et al., 2002). Plasmids encoding either wild-type (upper panel) or mutated (lower panel) interaction partners were transfected into cells together with an internal reference (CFP). The fluorescence intensities produced by bimolecular fluorescence complementation (YN-YC) and the internal reference (CFP) were measured in individual cells. The distribution of ratios between the fluorescence intensities in individual cells is plotted in each histogram. The dashed line indicates the maximal ratio produced by nonspecific interactions.
ALTERNATE PROTOCOL 1
SIMULTANEOUS VISUALIZATION OF MULTIPLE PROTEIN INTERACTIONS IN THE SAME CELL Comparison of the subcellular localization of different protein-protein complexes can provide information about similarities and differences in their functions based on colocalization or nonoverlapping distributions of the complexes. The locations of different complexes can be compared in different cells by using markers for various cellular structures to define the localization of each complex separately. However, it is often difficult to find markers that precisely match the localization of a particular complex. It is therefore preferable to compare the locations of protein complexes in the same cell. This protocol describes a multicolor BiFC approach that enables simultaneous visualization of several interactions in the same cell (Hu and Kerppola, 2003; Grinberg et al., 2004). The multicolor BiFC assay is based on complementation between fragments of different fluorescent proteins that produce bimolecular fluorescent complexes with distinct spectra. These fragments are fused to alternative interaction partners such that complexes formed with different partners can be visualized independently in the same cell using different excitation and emission wavelengths. This approach provides a unique method for comparison of the distributions of different protein complexes in the same cell and for analysis of the relative efficiencies of interactions with mutually exclusive interaction partners (Alternate Protocol 2).
Additional Materials (also see Basic Protocol)
Bimolecular Fluorescence Complementation
DNA encoding amino acid residues 155 to 238 of cyan fluorescent protein (also see Critical Parameters and Table 19.10.1) DNA encoding several alternative interaction partners
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DNA encoding amino acid residues 1 to 154 of the cyan and yellow fluorescent proteins (also see Critical Parameters and Table 19.10.1) Interference filters for fluorescence microscope designed to minimize cross-talk between the fluorescence signals from bimolecular fluorescent complexes formed by fragments of different fluorescent proteins (see below) Construct plasmid vectors 1. Using the appropriate plasmid vectors, construct mammalian expression vectors by fusing the sequences encoding amino acid residues 155 to 238 of the cyan fluorescent protein (CFP) to the protein whose interactions are to be investigated (this protein is designated Z-CC155), and fuse sequences encoding amino acid residues 1 to 154 of YFP and residues 1 to 154 of CFP to two alternative interaction partners (these proteins are designated A-YN155 and B-CN155, respectively). As controls, construct plasmids encoding these fluorescent protein fragments fused to the same interaction partners. For additional information about the design of fusion constructs, see Critical Parameters. Other fluorescent proteins whose fragments produce bimolecular fluorescent complexes with distinct spectra may be used (Hu and Kerppola, 2003; also see Critical Parameters).
Prepare cells 2. Seed cells the day before transfection at a density that will allow for cell proliferation over the course of the experiment and, at the same time, allow for the effects of cell growth and density on the interaction under investigation. If short-working-distance objectives will be used to visualize the interaction, the cells should be grown in slide chambers or on glass coverslips.
3. When cells reach appropriate confluence (e.g., ∼50%), transfect cells with an appropriate amounts (e.g., 0.25 µg) of the plasmids encoding A-YN155, B-CN155, and Z-CC155. As a control, transfect each pair of expression vectors (A-YN155 and Z-CC155 as well as B-CN155 and Z-CC155) into separate cells. 4. Grow cells until fluorescence from both complexes (i.e., A-YN155—Z-CC155 and B-CN155—Z-CC155) is detected using filters optimized for the detection of YFP and CFP fluorescence, respectively. If necessary, incubate the cells at 30◦ C with 5% CO2 to promote maturation of the fluorophores and to increase the signal (however, interpret results obtained under such conditions with care, as incubation at a lower temperature could alter protein interactions). 5. Compare the levels of expression of the proteins by immunoblot analysis and adjust the amounts of plasmids transfected if necessary.
Examine cells 6. Wash the cells with PBS to remove dead cells and debris, then add fresh medium. 7. View the cells using an inverted fluorescence microscope with a 20× long-workingdistance objective. To view the fluorescence emissions of A-YN155—Z-CC155 and B-CN155—Z-CC155 complexes use two filter sets with excitation at 500 ± 10 nm and emission at 535 ± 15 nm (to detect YN—CC complexes), and excitation at 436 ± 5 nm and emission at 470 ± 15 nm (to detect CN—CC complexes). The emission and excitation wavelengths are separated using a dichroic mirror with transmission windows at 450 to 490 nm and 520 to 590 nm. The fluorescence emissions of YN155—CC155 and CN155—CC155 bimolecular fluorescent complexes can be resolved with less than 2% cross-talk between the signals using the filters optimized for YFP and CFP detection.
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ALTERNATE PROTOCOL 2
ANALYSIS OF THE COMPETITION BETWEEN ALTERNATIVE INTERACTION PARTNERS IN LIVING CELLS The selectivity of protein interactions is determined by the relative binding affinities of alternative interaction partners and by the local concentrations of each protein. It is difficult to predict the selectivity of protein interaction in cells based on in vitro studies, since many factors, including covalent modifications, differences in subcellular distributions, interactions with additional proteins, and binding affinity, can influence the selectivity of interactions. The following protocol describes a quantitative multicolor BiFC assay that can provide information about the relative efficiencies of complex formation by mutually exclusive interaction partners in living cells (Hu and Kerppola, 2003; Grinberg et al., 2004).
Principles of competition experiment To investigate the competition between two alternative interaction partners (i.e., A and B) for a shared partner (i.e., Z), the proteins are fused to fragments of different fluorescent proteins that can form spectrally distinct bimolecular fluorescent complexes (i.e., AYN155, B-CN155, and Z-CC155). Equal concentrations of the two competing interaction partners (i.e., A-YN155 and B-CN155) are expressed with a limiting concentration of the shared partner (i.e., Z-CC155). The fluorescence intensities of bimolecular fluorescent complexes formed by the alternative interaction partners (i.e., A-YN155–Z-CC155 and B-CN155–Z-CC155) are measured in the same cells. For purposes of this protocol, the phrase “fluorescence intensities of complexes” and variations thereof will be used to refer to the fluorescence intensities of individual cells with specific spectral characteristics.
Additional Materials (also see Basic Protocol) Constructs prepared in Alternate Protocol 1, step 1 Prepare cells 1. Seed cells the day before transfection at a density that will allow for cell proliferation over the course of the experiment and, at the same time, allow for the effects of cell growth and density on the interaction under investigation. 2. When cells reach appropriate confluence (e.g., ∼50%), transfect cells with an appropriate amount (e.g., 0.05 µg) of the plasmid encoding Z-CC155 to produce a limiting concentration of the protein (established by immunoblot analysis) and appropriate amounts (i.e., 0.25 µg) of the plasmids encoding A-YN155 and B-CN155 to produce equal amounts of these proteins that are in molar excess relative to Z-CC155. For quantitation using absolute fluorescence intensities (see Support Protocol 2), also transfect each pair of expression vectors (A-YN155 and Z-CC155; as well as B-CN155 and Z-CC155) in separate cells. For quantitation using relative fluorescence intensities (see Support Protocol 3), also transfect expression vectors encoding fusions with the interaction partners fused to different combinations of fluorescent protein fragments (i.e. A-YN155 + A-CN155 + Z-CC155; as well as B-YN155 + A-CN155 + Z-CC155) into separate cells. 3. Grow cells until fluorescence from both complexes (i.e., A-YN155–Z-CC155 and B-CN155–Z-CC155) is detected. If necessary, incubate the cells at 30◦ C with a 5% CO2 atmosphere to promote maturation of the fluorophore and to increase the signal (however, interpret results obtained under such conditions with care, as incubation at a lower temperature could alter protein interactions). Bimolecular Fluorescence Complementation
4. Compare the levels of expression of the proteins and adjust the amounts of plasmids transfected if necessary.
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Examine cells 5. Wash the cells with PBS to remove dead cells and debris and add fresh medium. 6. View the cells using an inverted fluorescence microscope with a 20× long working distance objective. Identify cells that are healthy and that have a morphology similar to nontransfected cells. Interactions between competing partners will result in the formation of two BiFC complexes, A-YN155–Z-CC155 and B-CN155–Z-CC155, with distinct spectral characteristics. To image the fluorescence emissions of A-YN155–Z-CC155 and B-CN155–Z-CC155 complexes, use filters with excitation at 500 ± 10 nm and emission at 535 ± 15 nm to detect YN–CC complexes and filters with excitation at 436 ± 5 nm and emission at 470 ± 15 nm to detect CN–CC complexes. The emission and excitation light are separated using a dichroic mirror with transmission windows at 450 to 490 nm and 520 to 590 nm.
QUANTITATION OF MULTICOLOR BiFC ANALYSIS The authors of this unit have developed two methods for quantification of data from the multicolor BiFC assay that can provide information about the relative efficiencies of complex formation by mutually exclusive interaction partners in living cells (Hu and Kerppola, 2003; Grinberg et al., 2004). The first method, designated absolute competition (Support Protocol 2), is based on comparison of the absolute fluorescence intensities of bimolecular fluorescent complexes produced when a pair of interaction partners is expressed in the presence and absence of an alternative interaction partner in separate cells. The second method, designated relative competition (Support Protocol 3), is based on comparison of the relative fluorescence intensities of bimolecular fluorescent complexes produced when a protein is expressed together with two alternative interaction partners in the same cells. It is best to use both methods in combination to eliminate potential caveats inherent in each approach.
Comparison of the Absolute Fluorescence Intensities of Bimolecular Fluorescent Complexes Produced When a Pair of Interaction Partners Is Expressed in the Presence and Absence of a Competitor in Separate Cells
SUPPORT PROTOCOL 2
The absolute fluorescence intensities of bimolecular fluorescent complexes are compared when two fusion proteins are expressed alone (i.e. A-YN155+Z-CC155) and when they are expressed in the presence of a competitor (i.e. A-YN155+B-CN155+Z-CC155) in separate cells. This approach is designated absolute competition. This method is well suited for analysis of large differences in the efficiencies of complex formation between alternative interaction partners. Since it involves comparison of different cell populations, it is subject to error due to variations between the populations. Since the absolute fluorescence intensities vary between individual cells, this requires comparison of the fluorescence intensities of the populations or representative subsets of the populations. Using this approach, it is generally possible to detect only large differences in the efficiencies of complex formation. This approach compares the fluorescence intensities of bimolecular fluorescent complexes formed by the same interacting pair, in the absence and presence of competitor. This circumvents the effects of differences in the steric arrangement of the fluorescent protein fragments between different interaction partners that prevent comparison of the efficiencies of complex formation based on the absolute fluorescence intensities of different bimolecular fluorescent complexes. Identification of Protein Interactions
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Carry out Alternate Protocol 2, steps 1 to 6, then continue with the following steps. 1. Measure the fluorescence intensities of YN-CC complexes in 100 to 300 cells expressing A-YN155 + Z-CC155. 2. Measure the fluorescence intensities of YN-CC complexes in 100 to 300 cells expressing A-YN155 + B-CN155 + Z-CC155. 3. Measure the fluorescence intensities of CN-CC complexes in 100 to 300 cells expressing B-CN155 + Z-CC155. 4. Measure the fluorescence intensities of CN-CC complexes in 100 to 300 cells expressing A-YN155 + B-CN155 + Z-CC155. 5. Compare the fluorescence intensities of YN–CC complexes measured in steps 1 and 2 as well as the CN–CC complexes measured in steps 3 and 4. 6. Assess results. Case i: If the fluorescence intensities of the two bimolecular fluorescent complexes (i.e., A-YN155–Z-CC155 and B-CN155–Z-CC155) expressed in the same cells are ∼50% of the fluorescence intensities of the complexes when expressed separately, then the two complexes form with comparable efficiencies. Case ii: If the fluorescence intensity of one complex is similar whether it is expressed separately or in combination, whereas the fluorescence intensity of the other complex is reduced by >50% when expressed in combination, then the former complex forms with higher efficiency than the latter. Case iii: If the fluorescence intensities of both complexes expressed together are equal or higher than those observed when they are expressed separately, then the interaction partners do not compete for complex formation. The shared interaction partner may not be present at limiting concentration, and the relative efficiencies of complex formation cannot be determined based on the data. Case iv: If the fluorescence intensities of both complexes are reduced by significantly more than 50% when they are expressed together, or if the changes in fluorescence intensities are not reciprocal, then the interaction partners may affect the expression or functions of each other, and the relative efficiencies of complex formation cannot be determined based on the data.
SUPPORT PROTOCOL 3
Bimolecular Fluorescence Complementation
Comparison of the Relative Fluorescence Intensities of Bimolecular Fluorescent Complexes Produced when a Protein Is Expressed with Two Alternative Interaction Partners in the Same Cells The relative fluorescence intensities of bimolecular fluorescent complexes are compared when a fusion protein is expressed with two alternative interaction partners fused to fragments of different fluorescent proteins (i.e., A-YN155 + B-CN155 + Z-CC155). This approach is designated relative competition. This approach is suitable for detection of relatively modest differences in the efficiencies of complex formation between structurally related alternative interaction partners. Since this approach compares complexes in the same cells, it is less sensitive to variations between cells. However, since this approach compares the relative fluorescence intensitites of bimolecular fluorescent complexes produced by different interaction partners, it is subject to variations in the efficiency of association of the fluorescent protein fragments when they are fused to different interaction partners. Therefore, this approach can be used only when there are no large differences in the steric constraints to association of the fluorescent protein fragments between different complexes.
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Carry out Alternate Protocol 2, steps 1 to 6, then continue with the following steps. 1. Measure the fluorescence intensities of 100 to 300 cells expressing A-YN155 + A-CN155 + Z-CC155 to use as a reference. 2. Measure the fluorescence intensities of 100 to 300 cells expressing A-YN155 + B-CN155 + Z-CC155. 3. Measure the fluorescence intensities of 100 to 300 cells expressing B-YN155 + A-CN155 + Z-CC155. 4. Plot the fluorescence intensities of YN–CC versus CN–CC complexes for each individual cell as a scatter plot as shown in Figure 19.10.4 (middle row). 5. Compare the slopes produced by different combinations of interaction partners. The relative slopes of these plots provide information about the relative efficiencies of complex formation between alternative interaction partners. Alternatively, calculate the ratio YN-CC/(YN-CC + CN-CC) and plot the data as a histogram as shown in Figure 19.10.4 (bottom row). The distribution of ratios in the population can provide information about the relative efficiencies of complex formation. 6. Assess results. Case i: If the relative fluorescence intensities of all combinations of complexes display a linear relationship, and the slopes of the relationships are not significantly different, then the two interaction partners compete with indistinguishable efficiencies for the shared partner. Case ii: If the fluorescence intensities of all combinations of complexes display a linear relationship, and the slopes of the relationships are significantly different, then the two interaction partners compete with different efficiencies for the shared partner. Case iii: If the fluorescence intensities of one or all combinations of complexes do not display a linear relationship, or if the slopes cannot be determined with sufficient confidence, then the relative efficiencies of competition cannot be determined based on the data.
Comparison of Results from the Absolute and Relative Competition Approaches The relative competition approach (Support Protocol 3) is generally more accurate for complexes with similar structures. However, this approach can only be used when there are no differences in the steric constraints to association of the fluorescent protein fragments. To confirm that the identities of the fluorescent protein fragments fused to the alternative interaction partners do not influence the relative efficiencies of complex formation, it is necessary to exchange the fragments between the interaction partners (i.e., compare A-YN155 + B-CN155 + Z-CC155 with B-YN155 + A-CN155 + Z-CC155). In the absolute competition approach, exchange of the fragments should not affect the difference in fluorescence intensity in the presence and absence of the competitor. In the relative competition approach, the slope of the plot of YN-CC versus CN-CC intensities should shift in the opposite direction relative to the slope observed for the reference complexes (A-YN155 + A-CN155 + Z-CC155; compare the slopes of the plots on the right and left of the middle row of Fig. 19.10.4 with that of the plot in the center). If exchange of the fluorescent protein fragments between the interaction partners does not produce the predicted results, it is possible that the fluorescent protein fragments influence the specificity of the interaction. In this case, the relative efficiencies of complex formation cannot be determined using multicolor bimolecular fluorescence complementation analysis.
Limitations of the Method The multicolor BiFC assay for measurement of the efficiencies of protein interactions in cells enables determination of the relative efficiencies of competition between alternative interaction partners for a shared partner, but this does not necessarily reflect their relative
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Figure 19.10.4
Legend at right.
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binding affinities, since the complexes are not at equilibrium for the entire time of the experiment. The association of the fluorescent protein fragments is relatively slow (t1/2 ≈ 60 sec) and essentially irreversible in vitro (Hu et al., 2002). The relative efficiencies of complex formation reflect the competition between alternative interaction partners during the time before association of the fluorescent protein fragments. This is predicted to give a valid estimate of the relative efficiencies of complex formation by proteins with rapid exchange rates. However, interaction partners with very slow rates of association may not compete efficiently for complex formation in the assay. The multicolor BiFC approach is based on comparison of the relative amounts of bimolecular fluorescent complexes with different spectral characteristics. Differences between the rates of degradation of proteins in different complexes could alter the steady-state ratio between the complexes and thereby affect the apparent efficiencies of complex formation. The relative fluorescence intensities should therefore be measured at several time points, and the steady-state levels of proteins should be compared by western analysis. The rates of degradation of the complexes can be compared by monitoring the decrease in fluorescence in cells treated with protein-synthesis inhibitors (e.g., cycloheximide). The rates of degradation of the fusion proteins can also be measured directly using pulse-chase analysis.
Example of the Application of Quantitative Multicolor BiFC Analysis for Determination of the Relative Efficiencies of Complex Formation To examine the relative efficiencies of complex formation by protein Z with two alternative interaction partners, A and B, the authors of this unit performed an experiment in which the proteins were fused to fluorescent protein fragments (A-YN, B-CN, and Z-CC) and were expressed separately and in combination (Grinberg et al., 2004). The fluorescence intensities of the alternative complexes (i.e., A-YN–Z-CC and B-CN–ZCC) were measured when the complexes were expressed in separate cells and when they were expressed together in the same cell (Fig. 19.10.4A). As a reference, the fluorescent proteins were fused to the same interaction partners (B-YN, B-CN and Z-CC), and their fluorescence intensities were analyzed in parallel (Fig. 19.10.4B). The fluorescence intensities of the complexes were plotted in scatter plots (middle panels) and histograms
Figure 19.10.4 (at Left) Multicolor fluorescence complementation analysis of the competition for dimerization between alternative interaction partners. The relative efficiencies of complex formation between protein Z and two alternative interaction partners, A and B, can be quantified by fusing them to fragments of different fluorescent proteins and expressing them in different combinations. (A) BiFC analysis of A-YN + Z-CC (green), B-CN + Z-CC (red) and A-YN + B-CN + Z-CC (yellow). (B) BiFC analysis of B-YN + Z-CC (green), B-CN + Z-CC (red) and B-YN + B-CN + Z-CC (yellow). (C) BiFC analysis of B-YN + Z-CC (green), A-CN + Z-CC (red) and B-YN + ACN + Z-CC (yellow). The diagrams at the top indicate the combinations of proteins expressed either pairwise or in three-way competition in each experiment. The images show cells expressing the proteins indicated below each image. The cells were imaged using filters (described in the instrumentation section) that distinguish the fluorescence emissions of YN–CC (green) and CN– CC (red) complexes, and the images were superimposed. The fluorescence intensities of YN–CC complexes were plotted as a function of the fluorescence intensities of CN–CC complexes in individual cells (middle row of graphs). The fluorescence intensities are shown in green and red for cells that express the interaction partners pairwise, and in yellow for cells that express two alternative interaction partners in direct competition. The best fit of a linear function to the data from cells coexpressing three proteins is shown. The slope of this function reflects the relative efficiencies of complex formation between the alternative interaction partners. The ratio between the fluorescence emissions from each complex [Y/(Y + C) where Y represents YN–CC and C represents CN–CC] was determined in individual cells in each population, and was plotted in histograms (lower row of graphs) using the same colors as in the middle row. This black-and-white facsimile of the figure is intended only as a placeholder; for a full-color version of the figure go to http://www.interscience.wiley.com/c p/colorfigures.htm.
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(lower panels) when the complexes were expressed separately (red and green) and when they were expressed together in the same cells (yellow). The ratio between the fluorescence intensities of the complexes reflects the relative efficiencies of complex formation between the alternative interaction partners. When A-YN and B-CN were coexpressed with Z-CC, the fluorescence intensities produced by A-YN–Z-CC and B-CN–Z-CC exhibited a linear relationship with a slope of 1.3 and a 95% confidence interval of 0.08 (Fig. 19.10.4A, middle row). In comparison, when B-YN and B-CN were coexpressed with Z-CC, the fluorescence intensities produced by B-YN–Z-CC and B-CN–Z-CC exhibited a linear relationship with a slope of 0.98 and a 95% confidence interval of 0.05 (Fig. 19.10.4B, middle row). Likewise, the distribution of ratios between the fluorescence intensities [Y/(Y+C)] was shifted toward higher values for complexes formed by A-YN and B-CN with Z-CC compared to complexes formed by B-YN and B-CN with Z-CC (compare lower row of graphs for Fig. 19.10.4, panels A and B). These results are consistent with the interpretation that Z favors complex formation with A over complex formation with B. The absolute fluorescence intensities of A-YN–Z-CC and B-YN–Z-CC were comparable when expressed separately, suggesting that the efficiencies of association of the fluorescent protein fragments were comparable under conditions where no competitors for complex formation were present. To confirm that the fluorescent protein fragments fused to the alternative interaction partners did not influence the relative efficiencies of complex formation, the authors exchanged the fragments between the interaction partners (i.e., B-YN, A-CN and Z-CC) and compared the fluorescence intensities of bimolecular fluorescent complexes formed by these proteins (Fig. 19.10.4C). The fluorescence intensities of these complexes exhibited a linear relationship with a slope of 0.7 and a 95% confidence interval of 0.04 (Fig. 19.10.4C, middle row). Likewise, the distribution of fluorescence intensity ratios was shifted in the opposite directions compared to the proteins in which the fragments were fused to the opposite interaction partners (see lower row of graphs for Fig. 19.10.4; compare panel C with panels A and B). These data are consistent with the interpretation that the difference between the efficiencies of bimolecular fluorescent complex formation by A and B fusions is not due to a difference between the fluorescent protein fragments fused to these proteins, but is caused by a difference between the efficiencies of complex formation between A and Z versus B and Z (Grinberg et al., 2004).
Controls for Quantitative Multicolor BiFC Analysis The use of both absolute and relative competition methods for the quantification of relative efficiencies of complex formation is designed to correct for many of the factors that influence the fluorescence intensities of bimolecular fluorescent complexes formed by fragments of different fluorescent proteins. However, there are additional factors that need to be considered and examined as follows. 1. The relative levels of A-YN155 and B-CN155 expression affect the relative amounts of complexes formed. The alternative interaction partners must therefore be expressed at equal concentrations and the shared interaction partner must be expressed at a lower, limiting concentration. All protein concentrations should preferably be within the physiological range to observe the competition under normal cellular conditions. The levels of protein expression can be measured by immunoblot analysis using antibodies directed against the same epitope tag on the alternative interaction partners. Bimolecular Fluorescence Complementation
2. The efficiencies of association of different fluorescent protein fragments as well as the fluorescence intensities of the resulting bimolecular complexes can differ. It is therefore essential to determine the relative fluorescence intensities of the bimolecular fluorescent complexes when they are brought together by the same interaction
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partners (i.e., A-YN155, A-CN155, and Z-CC155). The relative fluorescence intensities of the two bimolecular fluorescent complexes (i.e., A-YN155–Z-CC155 and A-CN155–Z-CC155) reflect both differences between the efficiencies of association of the fluorescent protein fragments and differences between their fluorescence intensities when they are brought together by the same interaction partners. 3. The relative efficiencies of bimolecular fluorescence complementation can be affected by steric constraints imposed by the interaction partners on the association of the fluorescent protein fragments. To investigate this possibility, the fluorescent protein fragments can be fused to the interaction partners using several different extended linkers. If fusion proteins containing different linkers produce identical results, then the efficiencies of bimolecular complex formation are unlikely to be affected by steric constraints. 4. Different fluorescent protein fragments could have differential effects on the efficiencies of competition for the shared interaction partner. It is therefore essential to exchange the fluorescent protein fragments between the interaction partners to determine if differences between the fluorescent protein fragments affect the relative efficiencies of competition. This can also be verified in vitro by measuring the relative amounts of bimolecular fluorescent complexes formed when different ratios of the fluorescent protein fragments fused to the same interaction partners are mixed (Hu and Kerppola, 2003). If the amounts of complexes formed are proportional to the relative concentrations of fusion proteins in the reaction, then differences between the fluorescent protein fragments do not influence the relative efficiencies of complex formation.
COMMENTARY Background Information The biological functions of proteins are determined in large part by their interactions with other proteins in the physiological environment. Numerous protein interactions have been identified using genetic screens and in vitro binding assays. It is important to develop experimental approaches that allow determination of the subcellular locations of these interactions. The BiFC assay has been used to visualize interactions between many structurally unrelated proteins (Hu et al., 2002; Atmakuri et al., 2003; Deppmann et al., 2003; von der Lehr et al., 2003; Yu et al., 2003; Bracha-Drori et al., 2004; de Virgilio et al., 2004; Farina et al., 2004; Grinberg et al., 2004; Hynes et al., 2004a,b; Kanno et al., 2004; Rackham and Brown, 2004; Rajaram and Kerppola, 2004; Remy and Michnick 2004a,b; Remy et al., 2004; Tsuchisaka and Theologis, 2004a,b; Tzfira et al., 2004; Walter et al., 2004; Wei et al., 2004; Zal and Gascoigne, 2004; Zhang et al., 2004). The authors of this unit and other investigators have determined the localization of complexes formed by Fos, Jun, and ATF-2 (Hu et al., 2002); c-Maf and Sox (Rajaram and Kerppola, 2004); BATF and Jun (Deppmann et al., 2003); p65, p50, and IkB
(Hu et al., 2002); Myc, Max, Mxi1, Mad3, and Mad4 (Grinberg et al., 2004); Skp2 and c-Myc (von der Lehr et al., 2003); and Brd2 and histone H4 (Kanno et al., 2004). Protein interactions are crucial for the control of most cellular functions. Several methods have been developed to study protein interactions in living cells. One of the most commonly employed methods is FRET (UNIT 19.5). The FRET assay is based on the use of two fluorophores either chemically linked or genetically fused to two proteins whose interaction is to be examined. Compared to the BiFC assay, FRET analysis generally requires higher levels of protein expression to detect energy transfer. Also, structural information or a great deal of luck is required to place the two fluorophores within 100 Å of each other, which is the maximal distance over which any significant energy transfer between fluorescent proteins can be detected. The fraction of proteins that form complexes must also be high to produce a sufficient change in the donor and acceptor fluorescence intensities. To exclude alternative interpretations of the results, numerous controls must be performed and the fluorescence intensities must be measured with high quantitative accuracy. Despite these limitations, FRET has been successfully used for
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Bimolecular Fluorescence Complementation
the analysis of many protein interactions in living cells (Sorkin et al., 2000; Li et al., 2001; Majoul et al., 2002; Hink et al., 2003; Larson et al., 2003; Miyawaki, 2003; Tsien, 2003). A great advantage of FRET over BiFC analysis is the reversibility of complex formation, which potentially allows analysis of the interaction under equilibrium conditions. FRET is therefore superior for studies of the kinetics of protein association and dissociation. Bimolecular complementation between fragments of a variety of proteins has been used to investigate protein interactions in cells (Johnsson and Varshavsky, 1994; Rossi et al., 1997; Pelletier et al., 1998; Rossi et al., 1998; Ghosh, 2000; Galarneau et al., 2002; Wehrman et al., 2002; Paulmurugan and Gambhir, 2003). The basic principle is to fuse fragments of a reporter protein to the interaction partners. Selected fragments of many proteins are able to associate with each other to produce a functional complex. This complex can be detected by the use of chromogenic or fluorogenic substrates or ligands. Reporter proteins that have been used to date include ubiquitin (Johnsson and Varshavsky, 1994), β-galactosidase (Rossi et al., 1997), dihydrofolate reductase (Pelletier et al., 1998), luciferase (Paulmurugan and Gambhir, 2003), and β-lactamase (Galarneau et al., 2002; Wehrman et al., 2002). BiFC employs the classical principle of complementation between protein fragments (Ullmann et al., 1967, 1968). The advantage of the BiFC approach over other complementation approaches is that the reconstituted protein has strong intrinsic fluorescence that allows direct visualization of the protein complex. The complex can therefore be visualized with minimal perturbation of the cells. Using the BiFC approach, living cells can be observed over long periods, and the possibility of experimental manipulations altering the result is minimized. Potential problems with unequal distributions of the chromogenic or fluorogenic substrates or ligands are also avoided. Moreover, as described in Alternate Protocols 1 and 2, multiple protein interactions can be visualized in parallel using spectrally distinct bimolecular fluorescent complexes. One limitation of the BiFC approach is the time required for fluorophore maturation. This prevents real-time detection of rapid changes in interactions using the BiFC assay. In addition, bimolecular fluorescent complex formation is irreversible in vitro. These characteristics prevent the use of the BiFC assay for studies of the dissociation and subunit exchange of protein complexes in cells.
Characteristics of the BiFC assay The BiFC assay has several features that make it particularly useful for the study of protein interactions. First, it enables direct visualization of protein interactions and does not rely on their secondary effects. Second, the interactions can be visualized in living cells, eliminating potential artifacts associated with cell lysis or fixation. Third, the proteins are expressed in a relevant biological context, ideally at levels comparable to their endogenous counterparts. This increases the likelihood that they will reflect the properties of native proteins, including any post-translational modifications. Fourth, the BiFC assay does not require stoichiometric complex formation but can detect interactions between subpopulations of each protein. Fifth, multicolor BiFC analysis allows simultaneous visualization of multiple protein complexes in the same cell and enables analysis of the competition between alternative interaction partners for complex formation. Finally, BiFC does not require specialized equipment, apart from an inverted fluorescence microscope equipped with objectives that allow imaging of fluorescence in cells. The simple detection of bimolecular complex formation requires no post-acquisition image processing for interpretation of the data.
Critical Parameters Design of BiFC fusion constructs The fusion proteins for BiFC analysis must be designed with the specific proteins to be investigated as well as the purpose of the experiment in mind. Schematic diagrams of the different permutations of fusion proteins that can be used for BiFC analysis are shown in Figure 19.10.2. Some general guidelines for the design of fusion proteins for BiFC analysis are discussed below: Fragments of fluorescent proteins Several combinations of fluorescent protein fragments support bimolecular fluorescence complementation (Hu and Kerppola, 2003). The combinations of fluorescent protein fragments recommended for BiFC analysis are listed in Table 19.10.1. For most purposes, fragments of YFP truncated at residue 155 (designated YN155, containing N-terminal residues 1 to 154, and YC155, containing of C-terminal residues 155 to 238) are recommended, as they exhibit a relatively high complementation efficiency when fused to many interaction partners, yet produce low fluorescence when fused to proteins that do not
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interact with each other. Fragments of YFP truncated at residue 173 (designated YN173 containing residues 1 to 172, and YC173, containing residues 172 to 238) can also be used. Other combinations of fluorescent protein fragments can be also used (Hu and Kerppola, 2003) as described in the multicolor BiFC assay (see Alternate Protocols 1 and 2). Positions of fusions The positions of the fusions must be determined empirically to fulfill three essential criteria: 1. The fusions must allow the fragments of the fluorescent proteins to associate with each other if the fusion proteins interact. Information about the structure and location of the interaction interface may be useful for this purpose. However, this information is not essential since BiFC vectors can be designed by screening multiple combinations of fusion proteins for fluorescence complementation. A simple strategy for the identification of fusion proteins that allow bimolecular fluorescence complementation is to fuse each of the fluorescent protein fragments to the N- and C-terminal end of each interaction partner, and to test the fusion proteins for complementation in all eight combinations that contain both fragments of the fluorescent protein (Fig. 19.10.2). 2. The fusions must not affect the localization or the stabilities of the proteins. This should be confirmed by comparing the localization and the level of expression of the fusion proteins with those of wild-type proteins lacking the fusions using indirect immunofluorescence and immunoblot (UNIT 10.10) analyses. 3. The fusions must not affect the functions of the proteins under investigation. Ideally, the protein functions should be validated using assays that evaluate all of the known functions of the endogenous proteins, and they should be performed under the conditions used to visualize the protein interactions. Linkers The linkers connect the fragments of the fluorescent proteins to the proteins of interest in the fusion proteins. The linkers must provide flexibility for independent motion of the fluorescent protein fragments and the interaction partners, allowing the fragments to associate when the proteins interact. The authors have used the RSIAT and RPACKIPNDLKQKVMNH linker sequences in many fusion constructs used for
BiFC analysis (Hu et al., 2002; Hu and Kerppola, 2003). These linkers have been used for the visualization of interactions between many structurally unrelated proteins. The sequence AAANSSIDLISVPVDSR encoded by the multiple cloning sites of the pCMV-FLAG vector (Sigma) has also been successfully used as a linker in many BiFC experiments. Although these linker sequences worked well in the proteins examined, it is possible that the length or the sequence of the linker affects the complementation efficiency between other proteins. Design of constructs for multicolor BiFC analysis The basic principles for the design of BiFC constructs also apply to the design of multicolor BiFC constructs. The main difference is the use of fluorescent protein fragments that provide maximal spectral separation of the fluorescence signals from different bimolecular complexes. There are several combinations of fragments that can be used for multicolor BiFC analysis (Table 19.10.1). The use of CC155 (Cterminal fragment of CFP) paired with YN155 versus CN155 (N-terminal fragment of CFP) provides good spectral separation and high complementation efficiency. These combinations are therefore appropriate for the simultaneous analysis of two protein interactions. For the simultaneous analysis of more than two interactions, more selective interference filters and more complex spectral separation algorithms are required. Expression system The selection of an expression system should be based on the purpose of the experiment. To determine whether a pair of proteins can interact in cells and to determine the subcellular location of the complex, a transient overexpression system can be used. However, overexpression of proteins in cells can result in mislocalization of the proteins and formation of non-native complexes. To ensure that the observed fluorescence signal reflects native interactions, the fusion proteins should be expressed at levels comparable to the endogenous proteins. This can be done by the use of plasmids with weak promoters and plasmid vectors that do not replicate in mammalian cells. Also, cells can be transfected using small amounts of plasmid DNA, and they can be observed soon after transfection. More consistent results can be obtained by using inducible expression vectors integrated into the genomes of stable cell lines (e.g.,
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pIND). This allows for the control of protein expression at relatively uniform levels in the entire cell population. Controls In order to determine if any fluorescence observed reflects a specific protein interaction, it is essential to include negative controls in each experiment. This is especially important because the fluorescent protein fragments are able to form fluorescent complexes with a low efficiency even in the absence of a specific interaction. This spontaneous complementation is generally reduced when the fragments are fused to proteins that do not interact with each other. The validity of bimolecular fluorescence complementation results must therefore be confirmed using fusions to proteins in which the interaction interface has been mutated (Hu et al., 2002; Grinberg et al., 2004). The mutant proteins must be fused to the fluorescent protein fragments in a manner identical to the wild-type protein. The mutation must not affect the stability or the subcellular location of the fusion protein. The level of expression and the localization of the mutant protein should be compared with the wild-type fusion protein by immunoblot (UNIT 10.10) and indirect immunofluorescence analyses. The efficiencies of fluorescence complementation by the wild-type and mutant proteins should be quantified and compared. If the interaction interface has not been previously characterized, it is possible to screen for mutations that alter the efficiency of bimolecular fluorescence complementation, and thereby to determine if the complementation reflects a specific interaction. The BiFC assay can therefore be used to determine whether two proteins interact in cells without prior knowledge of the location or the structural nature of the interaction interface.
Bimolecular Fluorescence Complementation
Protein concentrations The levels of expression of fusion proteins used for BiFC analysis can have a profound influence on the experimental results and on interpretation of the data. A high level of expression may produce non-native protein interactions, and may result in bimolecular complementation independent of specific protein interactions. It may also alter the subcellular distributions of the fusion proteins and alter cell functions. These may lead to the misinterpretation of the observed interactions. It is therefore important to control the levels of expression of the fusion proteins and to compare them with those of the endogenous proteins.
This can be accomplished by using regulated expression vectors and by making stably transfected cell lines. Differences between the levels of expression of the fusion proteins and their endogenous counterparts should also be considered when interpreting the results from competition studies using the multicolor BiFC assay. Time and temperature The time required for the detection of fluorescence varies depending on the complementation efficiency. For most productive interactions, transiently transfected cells exhibit fluorescence at 12 to 36 hr after transfection. In the case of some interactions, fluorescence can be detected as early as 8 hr after transfection (Hu et al., 2002; Grinberg et al., 2004). Long incubation times should be avoided since this may result in higher expression of fusion proteins and complementation due to nonspecific interaction. Since fluorophore maturation is sensitive to high temperature, incubation at 30◦ C for 30 min to several hours can increase the signal. Keep in mind that results obtained at 30◦ C should be carefully interpreted, as lowtemperature incubation may alter protein localization and/or interactions. Interpretation of lack of fluorescence complementation The lack of fluorescence complementation does not prove the absence of an interaction because fusion of nonfluorescent fragments to the proteins of interest can alter their structures, which could interfere with an interaction between the proteins. Moreover, the spatial arrangement of the fragments of fluorescent proteins may not allow bimolecular complex formation. Only in cases where fluorescence complementation can be induced by an extracellular signal can the lack of fluorescence complementation in the absence of the signal be tentatively interpreted to reflect the absence of an interaction or a change in complex architecture. If there is strong evidence to indicate that the proteins interact in cells, fluorescence complementation could be examined in different cell types or in the presence of different extracellular stimuli. Alternatively, additional fusion proteins containing different linker sequences could be tested. Quantification of the efficiency of bimolecular fluorescence complementation The efficiency of fluorescence complementation is defined as the fluorescence intensity produced by bimolecular fluorescent complex formation relative to the levels of fusion
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proteins present in the cell. The efficiencies of bimolecular fluorescence complementation by structurally unrelated proteins cannot be used to determine their efficiencies of complex formation, since many factors unrelated to the efficiency of complex formation influence the efficiency of bimolecular fluorescence complementation. However, in situations where all of these factors are expected to be identical, differences in the efficiency of bimolecular fluorescence complementation can provide information about the relative efficiencies of complex formation. Thus, the effects of single amino acid substitutions that do not alter the level of protein expression or its localization can be examined by quantifying the efficiencies of fluorescence complementation by the wild-type and mutated proteins (Hu et al., 2002; Hu and Kerppola, 2003). To quantify the efficiency of fluorescence complementation, it is necessary to include an internal control in the experiment to normalize for differences in the efficiencies of transfection and protein expression in individual cells. For this purpose, cells are cotransfected with plasmids encoding the two fusion proteins together with a plasmid encoding a full-length fluorescent protein with distinct spectral characteristics (e.g., CFP). The fluorescence intensities derived from both bimolecular fluorescence complementation (i.e., YFP) and the internal control (i.e., CFP) are measured in individual cells. The ratio of YFP to CFP emission is calculated after subtraction of background signal (Fig. 19.10.3). The ratio of YFP to CFP fluorescence is a measure of the efficiency of bimolecular fluorescence complementation. For structurally related proteins under carefully controlled conditions, the efficiency of complementation can provide information about the efficiency of the protein interaction.
Anticipated Results If the fusion proteins under investigation support fluorescence complementation within 12 to 36 hr after transfection, it is likely that the complementation reflects a specific interaction. This possibility must be verified by the analysis of mutated proteins. For proteins that display complementation after longer times, it is increasingly probable that the complementation results from a nonspecific association. If eight different combinations of aminoand carboxy-terminal fusions are tested and no complementation is observed, it is unlikely that additional combinations will produce fluorescence complementation, although this does not
exclude the possibility that the proteins may interact.
Time Considerations The construction of plasmid vectors for BiFC and multicolor BiFC analysis can be accomplished in 1 to 2 weeks. For transfection, preparation of the cells takes 1 day and transfection takes 1 hr using FuGENE 6. Next, 1 to 2 days incubation at 37◦ C is required. To promote chromophore maturation, incubation at 30◦ C for 30 min to several hours or overnight may be necessary. Imaging of the cells may take a few hours depending on how the data will be analyzed. Quantitative analysis requires about half a day to 1 day for each experiment. If everything goes smoothly, 3 to 4 weeks should be enough to obtain preliminary results. However, it takes much longer to validate the results by performing all the necessary controls. Confirmation of the interaction using cells stably transfected with inducible expression vectors also takes longer, since it requires the establishment of stable cell lines expressing the fusion proteins.
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Grinberg, A.V., Hu, C.D., and Kerppola, T.K. 2004. Visualization of Myc/Max/Mad family dimers and the competition for dimerization in living cells. Mol. Cell. Biol. 24:4294-4308. Hink, M.A., Borst, J.W., and Visser, A.J. 2003. Fluorescence correlation spectroscopy of GFP fusion proteins in living plant cells. Methods Enzymol. 361:93-112. Hu, C.D. and Kerppola, T.K. 2003. Simultaneous visualization of multiple protein interactions in living cells using multicolor fluorescence complementation analysis. Nat. Biotechnol. 21:539545. Hu, C.D., Chinenov, Y., and Kerppola, T.K. 2002. Visualization of interactions among bZIP and Rel family proteins in living cells using bimolecular fluorescence complementation. Mol. Cell 9:789-798. Hynes, T.R., Mervine, S.M., Yost, E.A., Sabo, J.L., and Berlot, C.H. 2004a. Live cell imaging of G(s) and the beta(2)-adrenergic receptor demonstrates that both alpha(s) and beta(1)gamma(7) internalize upon stimulation and exhibit similar trafficking patterns that differ from that of the beta(2)-adrenergic receptor. J. Biol. Chem. 279:44101-44112. Hynes, T.R., Tang, L.N., Mervine, S.M., Sabo, J.L., Yost, E.A., Devreotes, P.N., and Berlot, C.H. 2004b. Visualization of G protein beta gamma dimers using bimolecular fluorescence complementation demonstrates roles for both beta and gamma in subcellular targeting. J. Biol. Chem. 279:30279-30286. Johnsson, N. and Varshavsky, A. 1994. Split ubiquitin as a sensor of protein interactions in vivo. Proc. Natl. Acad. Sci. U.S.A. 91:10340-10344. Kanno, T., Kanno, Y., Siegel, R.M., Jang, M.K., Lenardo, M.J., and Ozato, K. 2004. Selective recognition of acetylated histones by bromodomain proteins visualized in living cells. Mol. Cell 13:33-43. Larson, D.R., Ma, Y.M., Vogt, V.M., and Webb, W.W. 2003. Direct measurement of Gag-Gag interaction during retrovirus assembly with FRET and fluorescence correlation spectroscopy. J. Cell Biol. 162:1233-1244. Li, H.Y., Ng, E.K., Lee, S.M., Kotaka, M., Tsui, S.K., Lee, C.Y., Fung, K.P., and Waye, M.M. 2001. Protein-protein interaction of FHL3 with FHL2 and visualization of their interaction by green fluorescent proteins (GFP) two-fusion fluorescence resonance energy transfer (FRET). J. Cell. Biochem. 80:293-303. Majoul, I., Straub, M., Duden, R., Hell, S.W., and Soling, H.D. 2002. Fluorescence resonance energy transfer analysis of protein-protein interactions in single living cells by multifocal multiphoton microscopy. J. Biotechnol. 82:267-277. Miyawaki, A. 2003. Visualization of the spatial and temporal dynamics of intracellular signaling. Dev. Cell 4:295-305. Bimolecular Fluorescence Complementation
Nagai, T., Sawano, A., Park, E.S., and Miyawaki, A. 2001. Circularly permuted green fluorescent
proteins engineered to sense Ca2+ . Proc. Natl. Acad. Sci. U.S.A. 98:3197-202. Paulmurugan, R. and Gambhir, S.S. 2003. Monitoring protein-protein interactions using split synthetic renilla luciferase protein-fragmentassisted complementation. Anal. Chem. 75:1584-1589. Pelletier, J.N., Campbell-Valois, F.X., and Michnick, S.W. 1998. Oligomerization domaindirected reassembly of active dihydrofolate reductase from rationally designed fragments. Proc. Natl. Acad. Sci. U.S.A. 95:12141-12146. Rackham, O. and Brown, C.M. 2004. Visualization of RNA-protein interactions in living cells: FMRP and IMP1 interact on mRNAs. EMBO J. 23:3346-3355. Rajaram, N. and Kerppola, T. 2004. Transcription activation by Maf and Sox, and their subnuclear localization are disrupted by a mutation in Maf that causes cataract. Mol. Cell. Biol. 24:56945709. Remy, I. and Michnick, S.W. 2004a. A cDNA library functional screening strategy based on fluorescent protein complementation assays to identify novel components of signaling pathways. Methods 32:381-388. Remy, I. and Michnick, S.W. 2004b. Regulation of apoptosis by the Ft1 protein, a new modulator of protein kinase B/Akt. Mol. Cell. Biol. 24:14931504. Remy, I., Montmarquette, A., and Michnick, S.W. 2004. PKB/Akt modulates TGF-beta signalling through a direct interaction with Smad3. Nature Cell Biol. 6:358-365. Rossi, F., Charlton, C.A., and Blau, H.M. 1997. Monitoring protein-protein interactions in intact eukaryotic cells by beta-galactosidase complementation. Proc. Natl. Acad. Sci. U.S.A. 94:8405-8410. Rossi, F.M., Guicherit, O.M., Spicher, A., Kringstein, A.M., Fatyol, K., Blakely, B.T., and Blau, H.M. 1998. Tetracycline-regulatable factors with distinct dimerization domains allow reversible growth inhibition by p16. Nat. Genet. 20:389-393. Sorkin, A., McClure, M., Huang, F., and Carter, R. 2000. Interaction of EGF receptor and grb2 in living cells visualized by fluorescence resonance energy transfer (FRET) microscopy. Curr. Biol. 10:1395-1398. Tsien, R.Y. 2003. Imagining imaging’s future. Nat. Rev. Mol. Cell. Biol. Suppl. SS16-21. Tsuchisaka, A. and Theologis, A. 2004a. Heterodimeric interactions among the 1amino-cyclopropane-1-carboxylate synthase polypeptides encoded by the Arabidopsis gene family. Proc. Natl. Acad. Sci. U.S.A. 101:22752280. Tsuchisaka, A. and Theologis, A. 2004b. Unique and overlapping expression patterns among the arabidopsis 1-amino-cyclopropane-1carboxylate synthase gene family members. Plant Physiol. 136:2982-3000.
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Tzfira, T., Vaidya, M., and Citovsky, V. 2004. Involvement of targeted proteolysis in plant genetic transformation by Agrobacterium. Nature 431:87-92. Ullmann, A., Jacob, F., and Monod, J. 1967. Characterization by in vitro complementation of a peptide corresponding to an operator-proximal segment of the beta-galactosidase structural gene of Escherichia coli. J. Mol. Biol. 24:339343. Ullmann, A., Jacob, F., and Monod, J. 1968. On the subunit structure of wild-type versus complemented beta-galactosidase of Escherichia coli. J. Mol. Biol. 32:1-13. von der Lehr, N., Johansson, S., Wu, S., Bahram, F., Castell, A., Cetinkaya, C., Hydbring, P., Weidung, I., Nakayama, K., Nakayama, K.I., Soderberg, O., Kerppola, T.K., and Larsson, L.G. 2003. The F-box protein Skp2 participates in c-Myc proteosomal degradation and acts as a cofactor for c-Myc-regulated transcription. Mol. Cell 11:1189-1200. Walter, M., Chaban, C., Schutze, K., Batistic, O., Weckermann, K., Nake, C., Blazevic, D., Grefen, C., Schumacher, K., Oecking, C., Harter, K., and Kudla, J. 2004. Visualization of protein interactions in living plant cells using bimolecular fluorescence complementation. Plant J. 40:428-438.
Wehrman, T., Kleaveland, B., Her, J.H., Balint, R.F., and Blau, H.M. 2002. Protein-protein interactions monitored in mammalian cells via complementation of beta-lactamase enzyme fragments. Proc. Natl. Acad. Sci. U.S.A. 99:3469-3474. Wei, G.H., Liu, D.P., and Liang, C.C. 2004. Charting gene regulatory networks: Strategies, challenges and perspectives. Biochem. J. 381:1-12. Yu, H., West, M., Keon, B.H., Bilter, G.K., Owens, S., Lamerdin, J., and Westwick, J.K. 2003. Measuring drug action in the cellular context using protein-fragment complementation assays. Assay Drug Dev.Technol. 1:811-822. Zal, T. and Gascoigne, N.R. 2004. Using live FRET imaging to reveal early protein-protein interactions during T cell activation. Curr. Opin. Immunol. 16:418-427. Zhang, S.F., Ma, C., and Chalfie, M. 2004. Combinatorial marking of cells and organelles with reconstituted fluorescent proteins. Cell 119:137144.
Contributed by Chang-Deng Hu, Asya V. Grinberg, and Tom K. Kerppola Howard Hughes Medical Institute and University of Michigan Medical School Ann Arbor, Michigan
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Production and Use of Trimeric Isoleucine Zipper Fusion Proteins to Study Surface Receptor Ligand Interactions
UNIT 19.11
Cellular signaling is often initiated upon interaction of cell surface receptors with ligands. Analysis of such interactions is sometimes possible using standard techniques such as coimmunoprecipitation (UNIT 19.4) or phage display (UNIT 19.3). However, in many cases, the detection of such interactions on the surface of intact cells is desirable. Soluble forms of surface receptors can bind to their respective ligands on intact cells, and this binding can then be detected by methods such as flow cytometry, which enables the analysis of single cells. One way to produce soluble forms of a cell surface protein is to express only its extracellular portion as a recombinant protein. However, since the affinities of receptor-ligand interactions are usually low, it is sometimes difficult to detect the binding of a single receptor to its ligand. Creating multimerized receptors increases the avidity of the binding and facilitates detection of weak-affinity binding events. The approach in this unit describes the fusion of such extracellular portions of surface receptors with an isoleucine zipper (ILZ) sequence, resulting in recombinant, soluble, trimerized receptor fusion proteins. This unit provides advice on how to construct an ILZ fusion protein of a gene of interest (see Strategic Planning) and then focuses on the production and purification of such fusion proteins using a human cell line (see Basic Protocol 1). A second protocol describes the use of ILZ fusion proteins to detect receptor-ligand interactions by flow cytometry (see Basic Protocol 2).
STRATEGIC PLANNING Constructing or choosing an expression vector is the first step in producing ILZ fusion proteins. In many cases, it will be necessary to produce the ILZ fusion protein in a human cell line to ensure proper glycosylation of the protein. This can sometimes be essential for the correct binding of the recombinant protein to its cellular ligand. In these cases, it is necessary to choose a vector containing a strong mammalian promotor, such as the cytomegalovirus (CMV) or the elongation factor 1α (EF-1α) promotor. However, it should also be possible to produce ILZ fusion proteins in other expression systems described in Chapter 5 by introducing the ILZ sequence in appropriate expression vectors. For purification of the recombinant protein, it is advantageous to have the protein secreted into the cell culture medium. This can be achieved in mammalian cells by using a leader sequence of a cell surface protein, such as the Igκ leader sequence often used in expression vectors. Alternatively, the leader sequence of the surface protein of interest can be used. Both methods have produced successful outcomes. Without secretion, the protein will be more difficult to purify and, in addition, will not be properly glycosylated. The ILZ sequence is derived from the leucine-zipper dimerization domain of the yeast transcription factor GCN4 (Harbury et al., 1993). Leucine zipper motifs consist of a characteristic 7-amino-acid-residue repeat with hydrophobic residues at positions 1 and 4 (Landschulz et al., 1988). The ILZ sequence was generated by substituting those hydrophobic residues with isoleucine (Harbury et al., 1993; Fig. 19.11.1A). The 32-aminoacid-long ILZ peptides form a parallel three-stranded, α-helical coiled-coil (Harbury et al., 1994) and thereby lead to the trimerization of ILZ fusion proteins. Thus far, the ILZ fusion protein technique has only been successfully used with type I transmembrane
Contributed by Carsten Watzl Current Protocols in Protein Science (2006) 19.11.1-19.11.13 C 2006 by John Wiley & Sons, Inc. Copyright
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Figure 19.11.1 (A) Amino acid sequence (single amino acid code) and schematic structure of the isoleucine zipper helix. Isoleucine residues are marked in bold. (B) Schematic representation of the ILZ fusion protein expression vector pZipH. Abbreviations: EF1α, elongation factor 1α promotor; MCS, multiple cloning site; ILZ, isoleucine zipper sequence; BGH, bovine growth hormone. The backbone is derived from the pEF1 expression vector (Invitrogen). Modified from (Stark et al., 2005) with permission from Elsevier.
proteins (i.e., those with the amino-terminal end of the protein on the outside of the cell), with the ILZ sequence situated at the carboxyl-terminus of the extracellular domain of the gene of interest. However, it should also be possible to make ILZ fusion proteins from type II transmembrane proteins (i.e., those with the carboxy-terminal end of the protein on the outside of the cell). In this case, the ILZ sequence should be fused to the amino-terminus of the extracellular domain of the protein of interest. For easy purification, a His6 (6×His) tag is added to the carboxyl terminus of the ILZ sequence (for type I proteins). This enables a simple one-step purification of the secreted ILZ fusion protein from the culture supernatant. Other affinity tags such as FLAG or HA tags should also be compatible with this system (see UNIT 9.9 for more information on commonly used affinity tags).
Trimeric Isoleucine Zipper Fusion Proteins
A typical ILZ expression vector for type I transmembrane proteins is shown in Figure 19.11.1B. The strong EF-1α promotor drives expression of the construct. The Igκ leader sequence is followed by a multiple cloning site for easy subcloning of any extracellular domains. The ILZ sequence replaces the transmembrane region of the surface receptor, and is followed by the His6 tag and a stop codon.
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The human embryonic kidney cell line 293T has been used successfully as a producer cell line for ILZ fusion proteins; these cells are easily transfectable and can produce large amounts of protein. In addition, 293T cells express the large T antigen of SV40, which enables the amplification of plasmids carrying the SV40 origin of replication during transient transfection procedures. A transient transfection system is employed in this unit based on the very cost-efficient calcium phosphate method for transfecting 293T cells. Other transfection methods, such as lipid-based systems, can also be used for 293T cells. These methods might achieve higher transfection efficiencies, but are much more expensive. A stable producer cell line (UNIT 5.10) could also be generated, but this will require more time and effort. One of the benefits of a stable production system is having a consistent source of ILZ fusion proteins, which might be desirable if large and consistent amounts of ILZ fusion proteins are needed.
GENERATION AND PURIFICATION OF ILZ FUSION PROTEINS This protocol describes the expression of secreted ILZ fusion proteins using HEK 293T cells and the purification of the fusion protein using the His6 tag. An expression vector must be constructed containing the gene of interest fused to the ILZ sequence and a His6 tag, as described in Strategic Planning. The procedure described in this protocol will yield between 100 and 1000 µg of ILZ fusion protein, depending on the construct used. The protocol can be scaled accordingly, e.g., to produce smaller amounts for a pilot experiment. The time frame for the production and purification of an ILZ fusion protein is 9 to 10 days, starting with the plating of the HEK 293T cells.
BASIC PROTOCOL 1
Materials HEK 293T cells (ATCC no. CRL-11268) Dulbecco’s modified Eagle medium (DMEM) containing 10% fetal bovine serum (FBS) and 1× penicillin/streptomycin (pen/strep) Expression plasmid for the ILZ fusion protein of interest Expression vector for GFP (optional) 2 M CaCl2 (see recipe), sterilized using 0.22-µm filter (store up to 2 months at room temperature) 2× HBS solution (see recipe) 1 M imidazole solution (see recipe) 1 M NaOH Ni2+ -NTA agarose beads (e.g., Qiagen) Wash buffer (see recipe) Elution buffer (see recipe) 5× reducing SDS sample buffer (see recipe) SDS polyacrylamide gel (e.g., 10% Nu-PAGE Bis-Tris gel, Invitrogen; also see UNIT 10.1) Coomassie staining solution (e.g., SimplyBlue stain, Invitrogen; also see UNIT 10.5) 1× phosphate-buffered saline, pH 7.4 (PBS; see recipe for 10×) BCA protein assay kit (e.g., Pierce; also see UNIT 3.4) 175-cm2 tissue culture flasks 15- and 50-ml conical centrifuge tubes End-over-end rotator Poly-Prep disposable chromatography columns (Bio-Rad) Boiling water bath UV spectrophotometer Dialysis tubing, MWCO 8,000 to 10,000 (also see APPENDIX 3B) Disposable ultrafiltration device, MWCO 10,000 (e.g., Vivaspin 6-ml concentrator, Vivascience; http://www.vivascience.com/)
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Additional reagents and equipment for mammalian cell culture (APPENDIX 3C), SDS-PAGE (UNIT 10.1), staining of protein gels (UNIT 10.5), dialysis (APPENDIX 3B), and protein assay (UNIT 3.4) NOTE: All culture incubations should be performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified. NOTE: All solutions and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly.
Transfect HEK 293T cells Day 1 1. Dilute 2 × 108 trypsinized HEK 293T cells in 100 ml DMEM/10% FBS supplemented with pen/strep. Place a 20-ml aliquot of this suspension into each of five 175-cm2 tissue culture flasks. Grow cells overnight. APPENDIX 3C
describes basic mammalian cell culture protocols.
HEK 293T cells grow quickly and can usually be split 1:10 every 2 or 3 days. They should not be incubated for an extended period of time after reaching full confluency, in order to retain their transfectability. Thaw fresh cells every 3 months to ensure proper transfection and production capabilities. The amount of cells needed for this protocol can usually be obtained from one confluent 175-cm2 flask.
Day 2 The transfection reagents should be at room temperature at time of use. 2. On the next day (after ∼20 hr), combine 125 µg of ILZ expression plasmid and sterile water to a total volume of 11 ml in a sterile 50-ml conical tube, and add 1.5 ml of 2 M CaCl2 . Optionally, include an additional 10 µg of a GFP expression plasmid in the mix. This way, the transfection efficiency can be monitored by analyzing GFP expression of the transfected cells using an inverted fluorescence microscope with fluorescence excitation at 488 nm and emission at 545 nm. The GFP will not negatively influence the expression or purification of the ILZ fusion protein.
3. Add 12.5 ml of 2× HBS to the DNA/CaCl2 solution and bubble vigorously with automatic pipettor by keeping the dispense button pressed for 30 sec. Move the pipet up and down through the solution during bubbling. The bubbling ensures proper mixing of the DNA and HBS solutions and helps to form the DNA/CaPO4 precipitate.
4. Add 5 ml of the DNA/HBS solution dropwise to each 175-cm2 flask of 293T cells. Move the pipet across the entire surface of the flask to cover all cells. Adding the DNA/HBS solution to the medium will shift its color to orange. This makes it easy to verify that the entire flask is covered.
5. Rock the flasks gently back and forth a few times to evenly distribute the DNA/CaPO4 precipitate, and return the flasks to the incubator. The DNA/CaPO4 precipitate can be observed under the microscope. It will appear as very small particles and may sometimes resemble bacterial contamination.
Trimeric Isoleucine Zipper Fusion Proteins
Day 3 6. The next day (16 hr after transfection) aspirate the medium from the transfected HEK 293T cells and add 20 ml of fresh DMEM/10% FBS supplemented with pen/strep to each 175-cm2 flask.
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HEK 293T cells are only loosely adherent. Be careful not to detach the cells during the medium change. Stand the flasks upright for 1 min and aspirate the old medium. Add the new medium to the side opposite of the cells.
Harvest the supernatant Day 5 7. Approximately 48 hr after the medium change, harvest the supernatant containing the ILZ-fusion protein as follows. First, stand each flask upright and pipet the medium (100 ml total) into two 50-ml conical tubes. Next, carefully add 20 ml fresh DMEM/10% FBS supplemented with pen/strep to each 175-cm2 flask, return the flasks to the incubator, and continue incubation to day 8 (see step 9). At this stage the 293T cells are very fragile and detach easily. Some cells may already have detached from the flask, but most of the cells should still be adherent at this time. The medium color will be orange to yellow.
8. Centrifuge the 50-ml conical tubes containing the medium 5 min at 480 × g, 25◦ C, to remove any cells from the medium. Transfer the supernatant to two fresh 50-ml conical tubes. Proceed directly to the purification (step 10) or store the supernatant up to 3 days at 4◦ C at this stage. For some applications, it may be sufficient to use the supernatant directly without purification of the ILZ fusion protein.
Day 8 Many cells will probably have detached from the flask at this point. 9. Harvest the supernatant into two 50-ml conical tubes and centrifuge as described in step 8 to remove any cells from the medium. Transfer the clarified supernatant to two fresh 50-ml conical tubes. This second harvest usually contains less ILZ fusion protein than the first harvest, but it is usually still worthwhile to purify the ILZ fusion protein from the second harvest. If the supernatant from step 8 was not purified, the two samples can now be combined and purified together.
Purify the ILZ fusion protein 10. Add 1 ml of 1 M imidazole solution per 50 ml of supernatant for a final concentration of 20 mM imidazole (pH of the supernatant should now be ∼8.0). If pH is <7.5, adjust to ∼8.0 using 1 M NaOH. The histidine residues of the fusion protein tag need to be deprotonated for efficient binding to the Ni2+ agarose. The low concentration of imidazole is used to reduce nonspecific binding to the column.
11. Place 500 µl of packed Ni2+ agarose beads in a 15-ml conical centrifuge tube, add 10 ml wash buffer, and centrifuge 2 min at 480 × g, 25◦ C. Remove supernatant and resuspend beads with 500 µl of wash buffer to obtain 1 ml of a 50% slurry of Ni2+ agarose beads. This amount of beads has sufficient binding capacity for the purification of ILZ fusion protein from a 175-cm2 flask culture, even if the supernatants from steps 8 and 9 are combined.
12. Add the Ni2+ agarose beads to the supernatant from step 10. Use either 500 µl of bead slurry per 50 ml of supernatant when purifying 100 ml total, or only 250 µl of bead slurry per 50 ml of supernatant when purifying the combined supernatants from steps 8 and 9.
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13. Rotate the tubes containing the supernatant/beads on an end-over-end rotator for 2 hr at room temperature. 14. Centrifuge tubes 5 min at 480 × g, 25◦ C, to pellet the beads. Aspirate the supernatant. The ILZ fusion protein should now be bound to the Ni2+ agarose beads.
15. Resuspend beads in each tube with 20 ml wash buffer, rotate tubes on an end-overend rotator for 5 min at room temperature, and centrifuge tubes 5 min at 480 × g, 25◦ C. Aspirate supernatant. 16. Repeat step 15 once more for a second wash. 17. Resuspend all beads in a total of 5 ml wash buffer and transfer to a disposable Poly-Prep column containing a bed support to retain the Ni2+ beads. Rinse the 50-ml conical tubes with 5 ml of wash buffer to collect all beads, and transfer to the column. Make sure that the column is never allowed to run dry and that the beads settle without trapping air bubbles. The beads can be resuspended with more wash buffer inside the column using a fine pipet to release any trapped air.
18. Wash column with 10 ml wash buffer. 19. To elute the ILZ fusion protein, apply 1 ml of elution buffer to the column and collect the eluate in a 1.5-ml microcentrifuge tube. Repeat up to 10 times, collecting each fraction of the eluate in an individual tube. Keep the fractions on ice until and during electrophoresis. The ILZ fusion protein will typically elute in fractions 1 to 4.
20a. Evaluate purification by electrophoresis: Add 5 µl of 5× reducing SDS sample buffer to 20 µl of each fraction. Heat samples in a boiling water bath for 5 min,
Trimeric Isoleucine Zipper Fusion Proteins
Figure 19.11.2 Samples equivalent to 2 µg of the indicated ILZ fusion proteins were analyzed by 10% Nu-PAGE gel electrophoresis and stained using colloidal Coomassie. Adapted from (Stark et al., 2005) with permission from Elsevier.
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then apply to an SDS polyacrylamide gel (UNIT 10.1) and perform electrophoresis. After the gel run is complete, stain the gel with Coomassie blue (UNIT 10.5). Check for a single band corresponding to the ILZ fusion protein, which should appear in several of the early fractions. Combine the fractions containing the ILZ fusion protein and proceed to step 21. For a typical SDS-PAGE analysis of ILZ fusion proteins see Figure 19.11.2.
20b. If it has been established (e.g., in past experiments) that the column-purified ILZ fusion protein is without contaminating proteins: Analyze by measuring the absorbance at 280 nm. Combine the peak fractions and proceed to step 21. 21. Dialyze the purified ILZ fusion protein against several changes of 1× PBS using dialysis tubing with a MWCO of 8,000 to 10,000 (also see APPENDIX 3B). 22. Concentrate the ILZ fusion protein to ∼500 µl to 1 ml using an ultrafiltration device with a MWCO of 10,000. 23. Quantify the concentration of the ILZ fusion protein using a BCA protein quantification assay (UNIT 3.4). The authors use gel-filtration chromatography to confirm trimerization.
24. Adjust the concentration of the ILZ fusion protein to 200 to 1000 µg/ml with 1× PBS and store the protein in small aliquots at −20◦ C. Bovine serum albumin (1 mg/ml final) can also be added prior to freezing to stabilize the protein.
USE OF ILZ FUSION PROTEINS TO DETECT RECEPTOR-LIGAND INTERACTIONS ON LIVING CELLS
BASIC PROTOCOL 2
This protocol describes the use of ILZ fusion proteins to determine if cells express a binding partner or ligand for the respective ILZ fusion protein. The binding of the ILZ fusion protein to the surface of cells is detected by fluorescently labeled antibodies and analyzed by flow cytometry. The staining procedure is described for a 96-well plate, which will simplify the processing of multiple samples because 8- or 12-channel pipettors can be used.
Materials Cells of interest FACS buffer (see recipe) Purified ILZ fusion protein of interest (Basic Protocol 1, step 24) or HEK 293T cell supernatant containing the ILZ fusion protein of interest (Basic Protocol 1, step 8 or 9) FACS buffer (see recipe), ice cold 5 µg/ml anti-ILZ mouse monoclonal antibody (e.g., anti-ILZ-11; Stark et al., 2005); prepare in ice-cold FACS buffer just before use Phycoerythrin (R-PE)–labeled goat anti–mouse IgG antibody (Jackson ImmunoResearch); dilute 1:200 in ice-cold FACS buffer just before use 2% (v/v) formaldehyde in FACS buffer (optional) 96-well V-bottom microtiter plate Refrigerated centrifuge with adapter for 96-well plates Flow cytometer (e.g., Becton Dickinson FACScan) NOTE: All solutions should be at 4◦ C prior to use and all incubations must be done on ice.
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1. Centrifuge the cells of interest 5 min at 480 × g, 4◦ C, remove the supernatant, and resuspend the cell pellet in cold FACS buffer at a density of 2–4 × 106 cells/ml (see APPENDIX 3C for determining cell number). Place a 50-µl aliquot of this cell suspension into each designated well of a 96-well V-bottom plate, on ice. For each cell line and fusion protein a minimum of three cell-containing wells will be required: two for controls and one for specific staining of the ILZ fusion protein.
2. Add 50 ng of the ILZ fusion protein of interest to the well designated for specific staining of the fusion protein, to obtain a final concentration of 1 µg/ml. Do not add anything to the two wells reserved for controls. Incubate 20 min on ice. Most ILZ fusion proteins will give specific staining at a concentration of 1 µg/ml. In initial experiments, it may be necessary to titrate the concentration from 0.5 to 5 µg/ml to determine the optimal staining concentration. Use a negative control cell line to determine the concentration that produces nonspecific binding of the ILZ fusion protein. In most cases, 5 to 10 µg/ml will result in nonspecific binding. If using nonpurified ILZ fusion protein for staining, it will be necessary to determine the optimal staining dilution in a similar setup.
3. Add 150 µl cold FACS buffer to each well. Centrifuge the plate 5 min at 480 × g, 4◦ C. Carefully aspirate supernatant from each well without touching the cell pellet. This simplified wash step is usually sufficient. However, if nonspecific binding is observed, perform an additional wash step by resuspending the cell pellet in 150 µl FACS buffer and repeating the centrifugation step.
4. Resuspend cells in the well reserved for specific staining of the fusion protein, and in one control well (which will serve as a staining control), with 50 µl of 5 µg/ml anti-ILZ antibody solution (e.g., anti-ILZ-11). Resuspend cells in the other (negative) control well with 50 µl cold FACS buffer. Incubate 20 min on ice. If an antibody against the ILZ portion of the protein is not available, a commercial anti-6×His antibody (Qiagen) can be used for this step.
5. Add 150 µl cold FACS buffer to each well. Centrifuge the plate 5 min at 480 × g, 4◦ C. Carefully aspirate supernatant from each well without touching the cell pellet. 6. Resuspend cells in the well reserved for specific staining of the fusion protein, and in the staining control well, with 50 µl of cold PE-conjugated goat anti–mouse IgG antibody diluted 1:200 in FACS buffer. Resuspend cells in the other (negative) control well with 50 µl cold FACS buffer. Incubate 20 min in the dark on ice. After adding the fluorescently labeled antibody, the samples should be kept in the dark to limit bleaching of the fluorophore.
7. Add 150 µl cold FACS buffer to each well. Centrifuge the plate 5 min at 480 × g, 4◦ C. Carefully aspirate supernatant without touching the cell pellet. 8. Resuspend cells in 150 µl of cold FACS buffer and analyze immediately by flow cytometry. Alternatively, fix the cells by resuspending in 150 µl FACS buffer containing 2% formaldehyde at 4◦ C and store samples up to 1 week at 4◦ C for later analysis. The staining control was incubated with the secondary and tertiary antibody reagents but without fusion protein, and should not show any increased fluorescence compared to the negative control which was incubated with FACS buffer alone. If nonspecific binding of the secondary or tertiary reagent is observed in the staining control, include an additional wash in steps 5 and 7, or reduce the concentration of the reagents. Trimeric Isoleucine Zipper Fusion Proteins
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Elution buffer 50 mM NaH2 PO4 300 mM NaCl 250 mM imidazole 0.05% (v/v) Tween 20 Adjust pH to 8.0 using NaOH Store up to 6 months at room temperature FACS buffer Add fetal bovine serum (FBS) to sterile phosphate-buffered saline (PBS; see recipe) for a final concentration of 2% (v/v). Store up to 3 months at 4◦ C. HBS solution, 2× 8.0 g NaCl 6.5 g HEPES 10 ml 75 mM Na2 HPO4 300 ml H2 O Adjust pH to 7.0 using NaOH or HCl Adjust volume to 500 ml Check pH again and adjust if necessary Sterilize by filtering through 0.22-µm filter Store up to 2 months at room temperature Imidazole, 1 M Dissolve 6.8 g imidazole in water and adjust volume to 100 ml. Do not adjust pH. Store up to 6 months at room temperature. Phosphate-buffered saline (PBS), 10× 80 g NaCl 2 g KCl 35.8 g Na2 HPO4 ·12 H2 O or 17.8 g Na2 HPO4 ·2H2 O 2.4 g KH2 PO4 800 ml H2 O Adjust pH to 7.4 with NaOH Add H2 O to 1 liter Sterilize by autoclaving Store up to 1 year at room temperature Reducing SDS sample buffer, 5× Combine on a magnetic stirrer under a fume hood: 50 ml 99% glycerol 25 ml 2-mercaptoethanol 21 ml 1.5 M Tris·Cl, pH 6.8 (APPENDIX 2A) Add 10 g SDS and stir until dissolved. Add 10 mg bromphenol blue and stir until dissolved. The final volume should be 100 ml and usually does not need to be adjusted. Store at −20◦ C for up to 2 years. Working aliquots can be stored at room temperature for up to 3 months. Identification of Protein Interactions
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Wash buffer 50 mM NaH2 PO4 300 mM NaCl 20 mM imidazole 0.05% Tween 20 Adjust pH to 8.0 using NaOH Store up to 6 months at room temperature COMMENTARY Background Information
Trimeric Isoleucine Zipper Fusion Proteins
Soluble recombinant surface receptor proteins are very useful for studying cellular interactions and functions. They have been successfully used to investigate and identify the ligands of many cell surface receptors. In the case of an unknown ligand, they are the only tools available to study and demonstrate the expression of such a ligand. One way to obtain a soluble recombinant protein for a cell surface receptor is to express only the extracellular portion using any of the expression systems described in Chapter 5. However, bacterially expressed recombinant surface receptor proteins are not glycosylated, and glycosylation may be critical for the binding of a receptor to its physiological ligand. It is therefore desirable to produce such proteins in mammalian cells as described in UNIT 5.9 and in this unit. The affinities of receptor-ligand interactions are usually low, making it difficult to detect the binding of a single receptor to its ligand. Creating multimerized receptors increases the avidity of the binding and facilitates detection of weak-affinity binding events (Altman et al., 1996). One classical way to create receptor fusion proteins is to fuse the extracellular portion of a surface receptor to the constant region of the immunoglobulin heavy chain (Zettlmeissl et al., 1990). These so-called Fc fusion proteins form disulfidelinked dimers and show higher binding avidity than monomeric constructs. The ILZ fusion proteins described in this unit form trimers in solution, and therefore produce an improved avidity over Fc fusion proteins (Stark et al., 2005). ILZ fusion proteins can bind to ligands on the surface of cells at concentrations as low as 0.15 µg/ml, and usually show optimal binding at ∼1 µg/ml, which is 10 times less than the optimal staining concentration for Fc fusion proteins (Stark et al., 2005). The binding of soluble fusion proteins to their receptors on the surface of living cells can either stimulate the receptor or block receptor function by inhibiting the binding of the nat-
ural ligand. In the case of ILZ fusion proteins of members of the signaling lymphocyte activation molecule (SLAM)–related receptors, binding of ILZ ligands was found to block the function of these receptors (Flaig et al., 2004; Stark et al., 2005). However, members of the TNF family stimulate their respective receptors when binding in trimeric form. ILZ fusion proteins of the TNF family can therefore effectively stimulate cells by inducing receptor trimerization (Walczak et al., 1999). The outcome of ILZ fusion protein binding to a receptor can therefore differ depending on the nature of the receptor. The ability to block or to mimic the natural ligand makes ILZ fusion proteins attractive tools for both in vitro and in vivo experiments. In such applications, ILZ fusion proteins have another advantage over Fc fusion proteins. The Fc portions of antibodies interact with Fc receptors on many different cells to induce cellular responses. Also, Fc fusion proteins can bind to Fc receptors. This binding may create unwanted effects when using Fc fusion proteins in in vitro or in vivo experiments. As the ILZ portion does not have a physiological binding partner, the effects of ILZ fusion proteins are dependent only on the nature of the fusion partner. When using ILZ fusion proteins to block receptor-ligand interactions, it is important to note that the affinity between the soluble receptor and the ligand on the cell is likely to be the same as for the membrane-bound receptor. Therefore, a large excess of soluble receptor is needed for effective blocking. The ILZ part, including the His6 tag, will add only 5 kDa to the molecular weight of the monomers, and therefore 15 kDa to the total molecular weight of the trimeric fusion protein. In contrast, the Fc portion adds 50 kDa and thereby makes the Fc fusion protein much bulkier. This can negatively affect the interaction of the fusion protein with its ligand, but can also create a problem when using the recombinant fusion protein as an antigen to immunize animals. ILZ and Fc fusion proteins display the receptor in a native
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conformation, including the proper glycosylation. This makes ILZ and Fc fusion proteins attractive antigens to generate antibodies that can recognize the native receptor on the surface on living cells. Several different ILZ fusion proteins have been used successfully to generate monoclonal antibodies that recognize the native receptor (Flaig et al., 2004; Stark et al., 2005). When using Fc fusion proteins as antigen, many unwanted antibodies will form against the large Fc part. This is a problem, especially when generating polyclonal antisera. The smaller ILZ portion contains fewer epitopes for the generation of such antibodies. One monoclonal antibody against the ILZ portion, which was created as a byproduct (Stark et al., 2005), is now a valuable reagent for detecting the binding of ILZ fusion proteins to the surface of living cells in the FACS analysis described in Basic Protocol 2. The ILZ segment will form trimers in solution (Harbury et al., 1994). Therefore, ILZ fusion proteins will also form trimeric molecules. However, higher-order multimers have been observed when analyzing different ILZ fusion proteins by gel filtration (Stark et al., 2005). Some of this multimerization may be mediated by the fusion partner, e.g., by forming additional disulfide links (Stark et al., 2005), and therefore the multimerization can differ between ILZ fusion proteins. However, the ILZ part also seems to have a tendency to form such higher-order multimers. It is not clear how such multimers influence the activity of ILZ fusion proteins. They may further enhance the avidity of the ILZ fusion proteins, which could contribute to their efficient binding activity at low concentrations, but they may also reduce binding activity by making receptor molecules unavailable for ligand binding when they are buried inside the multimers. Future modifications of the ILZ sequence may be able to limit the amount of multimerization. Besides the use of ILZ fusion proteins as antigens or in in vitro or in vivo assays as described above, the detection of ligands on the surface of cells as described in Basic Protocol 2 is one of their most popular applications. In the case of several different ligands for one receptor, an ILZ receptor fusion protein can be especially useful for determining the expression of the different ligands with a single reagent. When the ligand for a receptor is unknown, an ILZ fusion protein can be a very valuable reagent for the investigation and identification of such a ligand (Flaig et al., 2004).
Critical Parameters and Troubleshooting Optimal transfection of HEK 293T cells is critical for the efficient production of ILZ fusion proteins. Critical parameters influencing transfection efficiency are the quality and density of cells and the preparation of the transfection reagents. The cells should be freshly thawed every 3 months and never allowed to overgrow. The optimal seeding density for the transfection may depend on the growth kinetics of HEK 293T cells and can vary between laboratories. These parameters may need to be adjusted if Basic Protocol 1 yields transfection efficiencies below 50%. Thaw a fresh vial of HEK 293T cells and seed the cells at different concentrations. Monitor transfection efficiency by transfecting the cells with a GFP expression plasmid and analyzing by flow cytometry with excitation at 488 nm and detection in FL1. Also prepare fresh transfection reagents and vary the pH of the 2× HBS solution slightly between 6.9 and 7.1. Choose the seeding density and pH that yield the highest transfection efficiency. Typically the transfection efficiency should exceed 80%. Even at optimal transfection efficiency, different ILZ fusion protein expression constructs may produce very different protein yields. In one case, the authors determined that an extremely low protein yield was connected to the use of the Igκ leader sequence. For unknown reasons, the resulting ILZ fusion protein was not properly secreted by the HEK 293T cells and accumulated intracellularly, as detected by anti-ILZ immunoblot analysis of cellular lysate and supernatant. Using the protein’s own leader sequence resulted in enhanced secretion and improved protein yields. Another potential factor negatively influencing protein yield is the presence of a binding partner for the respective ILZ fusion protein on the surface of the HEK 293T cells. The author of this unit successfully produced an ILZ fusion protein in HEK 293T cells even though the ligand for this fusion protein was present on the surface of the cells (Flaig et al., 2004; Stark et al., 2005). In this case, production of the ILZ fusion protein exceeded the binding capacity of the HEK 293T cells. However, in other cases, the production efficiency may be substantially reduced and may only be improved by using a producing cell line that does not contain the ligand. The purified ILZ fusion proteins are stable for several weeks at 4◦ C, at sufficiently high protein concentrations. Although substantial
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degradation of ILZ fusion proteins has not been observed when they are stored at 4◦ C for extended periods of time, the binding capacity of such ILZ fusion proteins can be greatly impaired, possibly due to partial denaturation. Therefore, ILZ fusion proteins should be stored in small aliquots at −20◦ C, where they are stable for many months. The aliquots should be thawed only once, stored at 4◦ C, and used within a few days. For optimal staining results, the condition of the cells and the concentration and quality of the ILZ fusion proteins is critical. Some protein ligands may be negatively affected by the use of trypsin to detach cells. Compare trypsinized versus nontrypsinized cells to determine if the binding of the ILZ fusion protein is affected by such treatment. As described above, the optimal concentration for the specific binding of an ILZ fusion protein may vary. To determine the optimal concentration, perform a pilot binding assay using a range of concentrations of the ILZ fusion protein. To distinguish specific from nonspecific bind-
Trimeric Isoleucine Zipper Fusion Proteins
ing, use a cell line lacking a binding partner for the specific ILZ fusion protein, or compare the binding of the ILZ fusion protein to that of an unrelated ILZ fusion protein. This will allow easy determination of the optimal staining concentration.
Anticipated Results The yield of ILZ fusion proteins obtained from five 175-cm2 flasks of cells using Basic Protocol 1 should range from 0.1 to 1 mg. The purity of ILZ fusion proteins is usually >90% as judged by Coomassie staining (Fig. 19.11.2). The authors regularly observe a contaminating protein of about 80 kDa, which copurifies by Ni2+ chromatography. The amount of this contaminating protein is greater in preparations with lower yields. While this protein has not yet been identified, there is also no indication that it negatively affects subsequent assays. The apparent size of the ILZ fusion proteins produced is greater than the calculated molecular weight of the recombinant fusion protein (Stark et al., 2005), and
Figure 19.11.3 Baf3 cells were transfected with a 2B4 (Baf3-2B4) or CD48 (Baf3-CD48) expression plasmid. Stable transfectants were either left unstained (control), stained with ILZ-11 followed by PE-conjugated goat anti-mouse antibody (GαM), or stained with the indicated ILZ fusion proteins. Cells were analyzed by flow cytometry. These results demonstrate the interaction between 2B4 and CD48.
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the most likely explanation for this when dealing with surface proteins is that the proteins are glycosylated (although this may differ depending on the gene of interest). Glycosylation is also indicated by the rather diffuse migration of the ILZ fusion proteins in SDS-PAGE (Fig. 19.11.2). A typical staining result demonstrating the interaction between the two surface receptors 2B4 (CD244) and CD48 is shown in Figure 19.11.3 (Brown et al., 1998). Note that the ILZ-11 and the PE-conjugated goat antimouse antibodies do not bind nonspecifically to the cells, as the fluorescence of this control staining is similar to the nonstained cells. The binding of ILZ-2B4 is only observed on cells transfected with CD48, and binding of ILZ-CD48 is only detected on cells expressing 2B4. This demonstrates the specificity of binding of the ILZ fusion proteins.
Time Considerations The production and purification of ILZ fusion proteins described in Basic Protocol 1 will usually take 9 to 10 days. In the author’s laboratory, the HEK 293T cells are typically seeded on a Monday evening (step 1) and the transfection is performed on Tuesday afternoon (steps 2 to 5). Medium is then changed on Wednesday morning (step 6) and the first supernatant is harvested on Friday (steps 7 to 8). The first purification (steps 10 to 20) is then performed directly, or the second supernatant is collected on Monday (step 9) and both harvests are purified together as in steps 10 to 20. Dialysis is typically performed overnight (step 21), and, on Tuesday, the purified ILZ fusion proteins are concentrated, quantified, and divided into aliquots (steps 22 to 24). The procedure should only be interrupted after harvesting the supernatant, which is stable for several days when stored under sterile conditions at 4◦ C. The staining procedure described in Basic Protocol 2 will only take about 2 hr, with relatively little hands-on time. Incubation periods can be shortened to 15 min without excessively compromising staining efficiency. They can also be extended to 1 or 2 hr without increasing nonspecific staining. As described in the protocol, the stained cells can either be analyzed directly or fixed and stored at 4◦ C in the dark. Fixed cells are stable for several days under these conditions.
Literature Cited Altman, J.D., Moss, P.A., Goulder, P.J., Barouch, D.H., McHeyzer-Williams, M.G., Bell, J.I., McMichael, A.J., and Davis, M.M. 1996. Phenotypic analysis of antigen-specific T lymphocytes. Science 274:94-96. Brown, M.H., Boles, K., van der Merwe, P.A., Kumar, V., Mathew, P.A., and Barclay, A.N. 1998. 2B4, the natural killer and T cell immunoglobulin superfamily surface protein, is a ligand for CD48. J. Exp. Med. 188:2083-2090. Flaig, R.M., Stark, S., and Watzl, C. 2004. Cutting edge: NTB-A activates NK cells via homophilic interaction. J. Immunol. 172:6524-6527. Harbury, P.B., Kim, P.S., and Alber, T. 1994. Crystal structure of an isoleucine-zipper trimer. Nature 371:80-83. Harbury, P.B., Zhang, T., Kim, P.S., and Alber, T. 1993. A switch between two-, three-, and fourstranded coiled coils in GCN4 leucine zipper mutants. Science 262:1401-1407. Landschulz, W.H., Johnson, P.F., and McKnight, S.L. 1988. The leucine zipper: A hypothetical structure common to a new class of DNA binding proteins. Science 240:1759-1764. Stark, S., Flaig, R.M., Sandusky, M., and Watzl, C. 2005. The use of trimeric isoleucine-zipper fusion proteins to study surface-receptor-ligand interactions in natural killer cells. J. Immunol. Methods 296:149-158. Walczak, H., Miller, R.E., Ariail, K., Gliniak, B., Griffith, T.S., Kubin, M., Chin, W., Jones, J., Woodward, A., Le, T., Smith, C., Smolak, P., Goodwin, R.G., Rauch, C.T., Schuh, J.C., and Lynch, D.H. 1999. Tumoricidal activity of tumor necrosis factor-related apoptosis-inducing ligand in vivo. Nat. Med. 5:157-163. Zettlmeissl, G., Gregersen, J.P., Duport, J.M., Mehdi, S., Reiner, G., and Seed, B. 1990. Expression and characterization of human CD4: Immunoglobulin fusion proteins. DNA Cell Biol. 9:347-353.
Contributed by Carsten Watzl University of Heidelberg Heidelberg, Germany
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Fluorescence Quenching Methods to Study Lipid-Protein Interactions
UNIT 19.12
This unit describes how fluorescence quenching methods can be used to determine the binding constants of lipids to intrinsic membrane proteins. The composition of the lipid shell, or annulus, around a membrane protein in a membrane will depend on the relative binding constants of the protein for all the species of lipids present in the membrane. The lipid annulus will be enriched in lipids that bind strongly to the protein and will be depleted of lipids that bind weakly to the protein. This is important because the activities of membrane proteins are sensitive to the structures of the lipid molecules that surround them in the membrane and, indeed, some membrane proteins are only active in the presence of particular classes of lipid (Lee, 2004). In those cases where a high-resolution structure is available for the membrane protein, it is possible to relate the measured binding constants to particular binding sites on the protein. For example, it has been possible to detect specific binding of anionic lipids to hot spots on the mechanosensitive channel MscL, corresponding to a cluster of positively charged residues (Powl et al., 2005). However, it is possible to use spectroscopic methods such as fluorescence quenching to determine lipid binding constants, even when the structure of the protein is not known. Other techniques, such as electron spin resonance (ESR), are also available for studying lipid-protein interactions (East et al., 1985; Marsh and Horvath, 1998), but fluorescence methods have the advantage of sensitivity and simplicity, and the bromine-containing lipid molecules required for the fluorescence methods are easy to synthesize by addition of bromine across cis carbon-carbon double bonds in unsaturated lipid molecules. For example, bromination of dioleoylphosphatidylcholine (DOPC) yields the brominated lipid molecule di(9,10-dibromostearoyl)phosphatidylcholine (BrPC). Phospholipids containing brominated fatty acyl chains behave much like conventional phospholipids with unsaturated fatty acyl chains because the bulky bromine atoms affect lipid packing in a manner similar to a cis double bond (East and Lee, 1982). The analysis of lipid-protein interactions assumes that the membrane-spanning region of a membrane protein is covered by a number of “sites” to which lipid molecules can bind (Fig. 19.12.1). The aim is to measure binding constants for lipids at these sites. The experimental approach makes use of the fact that the fluorescence of a Trp residue is quenched when the Trp residue is close to a bromine atom (East and Lee, 1982). Therefore, if a membrane protein contains a Trp residue close to a lipid-binding site on the protein, binding of a brominated lipid molecule to the site will result in quenching of the fluorescence of that Trp residue. Furthermore, if the membrane protein is reconstituted into a lipid bilayer containing a mixture of bromine-containing and bromine-free lipid molecules, the level of quenching will depend on the probability that a bromine-containing lipid molecule occupies the lipid binding site. The higher the binding constant for the bromine-containing lipid compared to that for the bromine-free lipid molecule, the more marked will be the fluorescence quenching; this is the basis for the measurement of binding constants. The method requires that the membrane protein being studied contain at least one Trp residue within the transmembrane region of the protein. Most membrane proteins in fact contain several Trp residues in the transmembrane region; while this aids sensitivity, the disadvantage is that analyzing the data generally requires the assumption that lipid binding constants at sites close to all the Trp residues are the same, and this may not be true. An ideal membrane protein for study is therefore a protein lacking Trp residues, Identification of Protein Interactions Contributed by Joanne Carney, J. Malcolm East, Sanjay Mall, Phedra Marius, Andrew M. Powl, J. Neville Wright, and Anthony G. Lee
19.12.1
Current Protocols in Protein Science (2006) 19.12.1-19.12.17 C 2006 by John Wiley & Sons, Inc. Copyright
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Figure 19.12.1 Lipid binding to an integral membrane protein. The figure shows lipid molecules (small circles) binding to part of the hydrophobic surface of a membrane protein. The lipid-binding sites are referred to as annular or boundary sites. For a protein reconstituted into a mixture of two types of lipid, A and B, each of the lipid-binding sites is occupied by either lipid A or by lipid B, The equilibrium is described by an equilibrium constant K given by: K = [Protein·A][B]/([Protein·B][A]). Here [A] and [B] are, respectively, the mole fractions of lipids A and B not bound to the protein and [Protein·A] and [Protein·B] are the fractions of the lipid-binding site occupied by A and B, respectively. In this example, two of the sites occupied by lipid molecules, shown by the dark circles, are sufficiently close to the Trp residue (W) that binding of brominated lipid to either of the sites will quench the fluorescence of the Trp residue. In this case, therefore, n = 2.
into which Trp residues can be introduced using site-directed mutagenesis, allowing the Trp residue to be placed at different positions, thereby allowing the determination of lipid binding constants at a variety of positions around the circumference of the protein (Powl et al., 2005). Measuring lipid binding constants for an intrinsic membrane protein requires purification of the protein followed by its reconstitution into lipid bilayers containing a brominated phospholipid. Ideally, the membrane protein will be one that can be purified in milligram quantities. A number of techniques are available for reconstitution of membrane proteins into bilayers containing brominated lipid molecules. The method described in this unit has the advantage of simplicity. It has been shown to work well with a wide range of membrane proteins, including ion channels (Williamson et al., 2002; Powl et al., 2003), ion pumps (East and Lee, 1982), enzymes (Pilot et al., 2001), and β-barrel porins (O’Keeffe et al., 2000). It is important that the membrane protein not purify with lipid molecules that remain tightly bound to the protein after reconstitution; since such lipid molecules would not exchange with brominated lipid molecules in the membrane, the fluorescence quenching method described here could not be used to determine the binding constant for a lipid of this type. In practice, this does not seem to be a major problem since lipid binding constants appear not to be so high that lipid binding to a protein is irreversible. The structures of any lipids purifying with a membrane protein can be characterized using standard techniques after extraction into organic solvent, as described by Bergelson (1980).
Fluorescence Quenching to Study Lipid-Protein Interactions
The protocols in this unit describe the preparation of brominated lipid, BrPC, from DOPC (Support Protocol 1) and its analysis by mass spectrometry (Support Protocol 2). Basic Protocol 1 describes the reconstitution of a membrane protein into a lipid bilayer formed from DOPC and BrPC and measurement of the fluorescence quenching efficiency. Basic Protocol 2 describes the measurement of the relative lipid-binding constant of a membrane protein using fluorescence quenching. Instructions are also given in Support Protocol 3 for the purification of commercial potassium cholate, which is required for membrane reconstitution.
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RECONSTITUTION OF A MEMBRANE PROTEIN INTO DOPC AND BrPC AND MEASUREMENT OF FLUORESCENCE QUENCHING EFFICIENCY
BASIC PROTOCOL 1
Membrane proteins are usually purified in detergents such as octyl glucoside or dodecyl maltoside. Such proteins can easily be reconstituted into lipid bilayers by mixing the protein in detergent with a sample of lipid dispersed in detergent, and then diluting to regenerate membranes in the form of unsealed vesicles. The detergent in which the membrane protein is purified will depend on the protein, but the authors of this unit have reconstituted proteins purified in Mega-9, octylglucoside, dodecylmaltoside, cholate, and octyl-POE. An important step in the reconstitution procedure is the efficient dispersal of the lipid into detergent. The authors have used a variety of detergents to solubilize the lipid, including octyl glucoside, octyl-POE, and cholate. Cholate was found to be a very convenient detergent for reconstitution because of its high critical micelle concentration (∼10 mM) and because it is inexpensive. Before embarking on a full set of experiments, it is recommended that a significant level of quenching be ascertained empirically for the particular membrane protein of interest. This protocol describes how a membrane protein can be reconstituted into bilayers of dioleoylphosphatidylcholine (DOPC) and its brominated analog di(9,10dibromostearoyl)phosphatidylcholine (BrPC); the ratio of the Trp fluorescence intensity in the two systems provides the level of quenching. The quantity of protein required for the measurements will depend on the number of Trp residues in the protein and on the sensitivity of the fluorimeter. The example given below describes the reconstitution for the specific case of the potassium channel KcsA purified as a His-tagged protein in dodecylmaltoside as detergent (Doyle et al., 1998); KcsA is a homotetrameric protein, with each monomer containing 5 Trp residues. KcsA can be replaced in this protocol (also see Basic Protocol 2) with any membrane protein of interest. NOTE: All reagents should be the purest available to minimize any fluorescent impurities.
Materials 25 mg/ml dioleoylphosphatidylcholine (DOPC; Avanti Polar Lipids) in chloroform 25 mg/ml di(9,10-dibromostearoyl)phosphatidylcholine (BrPC; Support Protocol 1) in chloroform Oxygen-free nitrogen Reconstitution buffer: HEPES/EGTA buffer (see recipe) containing 15 mM potassium cholate purified as in Support Protocol 3 KcsA (purified as His-tagged protein in Mega-9 or dodecylmaltoside, eluting from metal-chelate column with imidazole; Doyle et al., 1998; Williamson et al., 2002) at 2.5 mg protein/ml HEPES/EGTA buffer (see recipe) 10-ml screw-cap glass scintillation vials, 1-cm diameter Vacuum desiccator connected to vacuum pump 50◦ C water bath Retort stand and clamp Ultrasonication bath (model FB11010, Fisher Scientific, or equivalent) 4-ml quartz fluorescence cell Fluorimeter Reconstitute membrane protein 1. Add 16 µl of 25 mg/ml DOPC (0.5 µmol DOPC) to each of two glass scintillation vials. One vial will be used for reconstitution with the membrane protein; the other will serve as a fluorescence blank.
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The efficiency of the sonication process in the subsequent steps depends on the glass vial being used; in some vials the liquid within the vial is seen to “fountain” when the sonicator bath is switched on, whereas in other vials the surface of the liquid hardly moves; in general, the more violent the disturbance of the liquid, the better. Glass scintillation vials work well.
2. Add 22 µl of 25 mg/ml BrPC (0.5 µmol BrPC) to each of two new glass scintillation vials. Again, one vial will be used for reconstitution and the other for a fluorescence blank.
3. Remove most of the solvent in the four vials by evaporating in a stream of oxygenfree nitrogen, then place the four vials in a vacuum desiccator attached to a vacuum pump for ∼2 hr or overnight, to lyophilize the lipid onto the sides of the vials. 4. Add 400 µl reconstitution buffer to each vial. Flush the vials with nitrogen, screw the caps on tightly, and wrap Parafilm around the caps for a complete seal. Briefly warm the vials under hot running water or in a 50◦ C water bath and vortex for about 5 min to help disperse the lipid. 5. Using the retort stand and clamp, place the vials one by one in the ultrasonication bath and sonicate until the sample is optically clear; this will generally take ∼5 min. The efficiency of the sonication process depends very markedly on the ultrasonication bath used. A desirable sonication bath is one in which the liquid in the glass vial is agitated violently when the bath is switched on. Most traditional ultrasonication baths are not powerful enough for dispersing lipids in detergent. The Fisher model FB11010 is recommended. A cup-horn ultrasonicator such as the Misonix Programmable Ultrasonic Processor with a 2.5-in (6.25-cm) cup-horn probe, although considerably more expensive, also works well. A good test of optical clarity is to place the vial on top of a sheet of printed paper; if the sample is clear, the writing on the paper should be easy to read through the vial. If a cup-horn sonicator is being used, the horn should be filled with an ice-water mix and the vial should be clamped so that the bottom of the vial is just above the surface of the probe. The sample is then sonicated at full power, checking every few minutes that the temperature of the surrounding water has not become too high.
6. Add 34 µl of 2.5 mg/ml KcsA (equivalent to 5 nmol KcsA monomer) to one of the two vials containing DOPC. The molar ratio of lipid:KcsA monomer in the sample is 100:1.
7. Add 34 µl of 2.5 mg/ml KcsA (equivalent to 5 nmol of KcsA monomer) to one of the two vials containing BrPC. KcsA is purified as a His-tagged protein in Mega-9 or dodecylmaltoside, eluting from a metal-chelate column with imidazole (Doyle et al., 1998; Williamson et al., 2002). It is important to use the purest available grade of imidazole, since other grades contain fluorescent impurities.
8. Leave samples at room temperature for 15 min, then keep on ice for up to 1 hr before use.
Fluorescence Quenching to Study Lipid-Protein Interactions
Obtain fluorescence measurements 9. Dilute 63 µl of the sample of KcsA in DOPC into 3 ml of HEPES/EGTA buffer in a 4-ml quartz fluorescence cell and record the Trp fluorescence emission spectrum using a fluorimeter. Typically, Trp is excited at 290 nm and the fluorescence emission spectrum is recorded from 300 to 400 nm. The 50-fold dilution of the sample decreases the detergent concentration below its critical micelle concentration, allowing the membranes to reform.
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Reconstituted membrane samples tend to be cloudy, so light scatter can be a problem in fluorescence studies of membranes. Light scatter is minimized in a fluorimeter with good monochromators in the excitation and emission light paths and with variable slits. High sensitivity is also a considerable advantage. It should be possible to stir the sample to prevent photobleaching. Also, ideally, the fluorimeter is connected to a computer to allow rapid data manipulation.
10. Repeat step 9 for the sample of KcsA in BrPC, again measuring the Trp emission spectrum. 11. Repeat step 9 for the samples of DOPC and BrPC lacking protein. These spectra are blanks to account for light scatter from the lipid and any fluorescent impurities in the buffers.
12. Subtract the fluorescence blanks from the fluorescence spectra in DOPC and BrPC and plot the spectra. Fluorescence spectra for KcsA in DOPC and BrPC are shown in Figure 19.12.2. The positions of the fluorescence maxima can be different on different fluorimeters because fluorimeter detection systems have different sensitivities to different wavelengths of light, distorting the emission spectra.
13. Measure the fluorescence emission intensity at a wavelength such as 330 nm where the intensity is high. 14. Calculate the fractional fluorescence quenching as [(F1 – F3 )-(F2 – F4 )]/(F1 – F3 ), where F1 and F2 are the fluorescence emission intensities for KcsA in DOPC and BrPC, respectively, and F3 and F4 are the measured intensities for the DOPC and BrPC blank samples, respectively. For KcsA, the fractional quenching in BrPC is ∼0.39 (Williamson et al., 2002).
Figure 19.12.2 Fluorescence emission spectra for KcsA in buffer (a, dotted line) and after reconstitution into DOPC (b) and BrPC (c). The lower fluorescence intensity in BrPC shows fluorescence quenching. The protein concentration was 0.75 µM and the molar ratio of lipid:KcsA was 100:1.
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BASIC PROTOCOL 2
MEASUREMENT OF THE RELATIVE LIPID-BINDING CONSTANT OF A MEMBRANE PROTEIN USING FLUORESCENCE QUENCHING If the Trp fluorescence of the protein of interest is quenched significantly by brominated lipid, then it is possible to measure lipid-binding constants for the protein. This involves first determining the number of lipid-binding sites, n, from which the fluorescence of the Trp residues can be quenched (see Commentary). This determination requires measurements of the fluorescence intensity of the protein of interest (KcsA in this example) reconstituted into a series of mixtures of DOPC and BrPC with different amounts of BrPC. Having determined n, the relative binding constant for the lipid of interest is then determined from measurements of fluorescence intensity in mixtures of that lipid (lipid X) with BrPC. Having determined a relative binding constant from measurements of fluorescence quenching in mixtures of lipid X with BrPC, it is important to repeat the measurements in mixtures of brominated lipid X and DOPC. If all the lipid-binding sites on the protein have the same relative binding constants for lipid X, then the relative binding constant for lipid X determined from experiments with the lipid X/BrPC mixtures will be the same as that determined with the brominated lipid X/DOPC mixtures. However, if the relative lipid binding constants for lipid X determined from the two sets of experiments are different, this would suggest that lipid binding to the protein is heterogeneous, with different sites showing different relative affinities for lipid X. In particular cases, the data can be analyzed to give binding constants at each type of site (Powl et al., 2005). A membrane protein can be reconstituted into bilayers containing mixtures of two lipids by first mixing the two lipids in chloroform solution in the desired amounts, then lyophilizing the lipids in scintillation vials and reconstituting as described in Basic Protocol 1. The procedure does, however, involve a large number of sonications, and it is therefore more convenient to produce separate batches of the two lipids sonicated in cholate and then mix the two separate batches in the required proportions, as described here. The steps below are for KcsA (also see Basic Protocol 1), but KcsA can be replaced by any membrane protein of interest.
Materials
Fluorescence Quenching to Study Lipid-Protein Interactions
25 mg/ml dioleoylphosphatidylcholine (DOPC; Avanti Polar Lipids) in chloroform 25 mg/ml di(9,10-dibromostearoyl)phosphatidylcholine (BrPC; Support Protocol 1) in chloroform Oxygen-free nitrogen Reconstitution buffer: HEPES/EGTA buffer (see recipe) containing 15 mM potassium cholate purified as in Support Protocol 3 KcsA (purified as His-tagged protein in Mega-9 or dodecylmaltoside, eluting from metal-chelate column with imidazole; Doyle et al., 1998; Williamson et al., 2002) at 2.5 mg protein/ml HEPES/EGTA buffer (see recipe) 25 mg/ml dioleoylphosphatidylethanolamine (DOPE; Avanti Polar Lipids) in chloroform 10-ml screw-cap glass scintillation vials, 1-cm diameter Vacuum desiccator connected to vacuum pump 50◦ C shaking water bath Retort stand and clamp Ultrasonication bath (model FB11010, Fisher Scientific, or equivalent) 4-ml quartz fluorescence cell Fluorimeter SigmaPlot (Sigma) or equivalent data-fitting program
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Measure fluorescence quenching in mixtures of DOPC and BrPC 1. Add 140 µl of 25 mg/ml DOPC (4.4 µmol DOPC) to one glass scintillation vial. 2. Add 195 µl of 25 mg/ml BrPC in chloroform (4.4 µmol BrPC) to a second glass scintillation vial. 3. Remove most of the solvent in the vials by evaporating in a stream of oxygen-free nitrogen, then place the vials in a vacuum desiccator attached to a vacuum pump for ∼2 hr or overnight, to lyophilize the lipid onto the sides of the vials. 4. Add 2 ml reconstitution buffer to each vial. Flush the vials with nitrogen, seal the screw caps tightly, and wrap Parafilm around the caps for a complete seal. Briefly warm the vials under hot running water or in a 50◦ C water bath and vortex for about 5 min to help disperse the lipid. 5. Using the retort stand and clamp, place the vials one by one in the ultrasonication bath and sonicate until the sample is optically clear; this will generally take ∼5 min. The efficiency of the sonication process depends very markedly on the ultrasonication bath used. A desirable sonication bath is one in which the liquid in the glass vial is agitated violently when the bath is switched on. Most traditional ultrasonication baths are not powerful enough for dispersing lipids in detergent. The Fisher model FB11010 is recommended. A cup-horn ultrasonicator such as the Misonix Programmable Ultrasonic Processor with a 2.5-in (6.25-cm) cup-horn probe, although considerably more expensive, also works well. A good test of optical clarity is to place the vial on top of a sheet of printed paper; if the sample is clear then the writing on the paper should be easy to read through the vial. If a cup-horn sonicator is being used, the horn should be filled with an ice-water mix and the vial should be clamped so that the bottom of the vial is just above the surface of the probe. The sample is then sonicated at full power, checking every few minutes that the temperature of the surrounding water has not become too high.
6. Add 1.74 ml of the reconstitution buffer to each vial to yield a final volume of 3.74 ml in each. Sonicating the lipid at a higher lipid concentration and then diluting the sample is advantageous because it reduces the sample volume being sonicated at any one time. The efficiency of sonication decreases with increasing sample volume.
7. Prepare three sets of mixtures of DOPC and BrPC, each containing mole fractions 0.0, 0.2, 0.4, 0.6, 0.8, and 1.0 of BrPC by placing the volumes of the sonicated samples of DOPC and BrPC listed in Table 19.12.1 into 18 1.5-ml microcentrifuge tubes. The mole fraction of BrPC in the membrane, xBr , is calculated from the following equation: xBr = [BrPC]/([BrPC] + [DOPC]) where [BrPC] and [DOPC] are, respectively, the concentrations of BrPC and DOPC in the sample, expressed in the usual units of moles per liter.
8. Place the microcentrifuge tubes in a 50◦ C shaking water bath and shake the samples for 30 min to ensure good mixing of the lipids. 9. Remove the tubes from the water bath and allow them to cool to room temperature. 10. For two sets of the tubes, add 34 µl of the 2.5 mg/ml sample of KcsA to each tube, yielding 5 nmol KcsA monomer (final) in each tube at a molar ratio of 100:1 lipid:KcsA monomer. The two sets of tubes containing KcsA will allow duplicate measurements of fluorescence intensity; the set of tubes containing no KcsA will provide the sample blanks.
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Table 19.12.1 Required Volumes of Sonicated Samples of DOPC and BrPC
Tubes
Mole fraction BrPC
Volume DOPC sample (µl)
Volume BrPC sample (µl)
1, 2, 3
0
400
0
4, 5, 6
0.2
320
80
7, 8, 9
0.4
240
160
10, 11, 12
0.6
160
240
13, 14, 15
0.8
80
320
16, 17, 18
1.0
0
400
11. Leave the three sets of tubes at room temperature for 15 min, and then place on ice for up to 1 hr before use. 12. Take 100 µl of sample from each protein-containing tube and add it to 2.9 ml of HEPES/EGTA buffer in a 4-ml quartz fluorescence cell. Measure the fluorescence emission intensity at the fluorescence maximum (about 330 nm; see Basic Protocol 1, step 9). Perform three determinations of fluorescence intensity from the material in each microcentrifuge tube and calculate the average fluorescence intensity from the three measurements. 13. Similarly, add 100 µl of sample from each protein-free tube to 2.9 ml of HEPES/EGTA buffer in a fluorescence cuvette and measure the fluorescence intensity at the same wavelength used in step 12; this is the sample blank. Perform three determinations of the blank intensity from the material in each microcentrifuge tube and calculate the average blank intensity from the three measurements. 14. For each protein-containing measurement, subtract the sample blank to determine the average corrected fluorescence intensity. The average corrected fluorescence intensities should be the same for each pair of measurements at a given BrPC content.
15. Plot average corrected fluorescence intensities as a function of the mole fraction of BrPC in the membrane. 16. Use the SigmaPlot nonlinear least squares option with the routine given below to calculate the value of n, as described in the Commentary: K=1.0 fmax=1.0 fbra=x/(x+K*(1-x)) f=fmin+(fmax-fmin)*((1-bra)ˆn) fit f to y
Fluorescence Quenching to Study Lipid-Protein Interactions
Measure fluorescence quenching in mixtures of DOPE and BrPC 17. Repeat steps 1 to 15, replacing DOPC with DOPE. Because of the difference in molecular weights of DOPE and DOPC, in step 1 use 133 µl of a 25 mg/ml solution of DOPE. 18. Having plotted the fluorescence intensities for KcsA reconstituted in mixtures of BrPC and DOPE, use the SigmaPlot nonlinear least squares option with the routine shown below to calculate the relative binding constant for DOPE relative to DOPC,
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with the value for n calculated in step 16 to calculate the relative binding constant for DOPE relative to DOPC. n = {insert value of n here} fmax=1.0 fbra=x/(x+K*(1-x)) f=fmin+(fmax-fmin)*((1-bra)ˆn) fit f to y The binding constant for DOPE relative to BrPC should be ∼0.57 (Alvis et al., 2003).
PREPARATION OF BROMINATED LIPID This protocol presents a general method for bromination of lipid molecules. Brominated phospholipids are synthesized by addition of bromine across the cis-double bonds of phospholipids containing monounsaturated fatty acyl chains. The protocol describes the bromination of dioleoylphosphatidylcholine to give di(9,10dibromostearoyl)phosphatidylcholine, but the same procedure has been used to brominate phospholipids with a variety of different fatty acyl chain lengths, and with a variety of different headgroups including phosphatidylethanolamines, phosphatidic acid, phosphatidylglycerol, phosphatidylserine, and cardiolipin. The method can also be used to brominate cholesterol (Simmonds et al., 1982) and molecules such as oleic acid, oleoylamine and oleoyl alcohol (Froud et al., 1986).
SUPPORT PROTOCOL 1
Materials 25 mg/ml dioleoylphosphatidylcholine (DOPC; Avanti Polar Lipids) in chloroform Bromine, liquid (Br2 ) 95:35:6 (v/v/v) chloroform/methanol/water or 14:6:1 (v/v/v) chloroform/methanol/25% (w/v) ammonia 10-ml round-bottom flask with ground-glass stopper Retort stand Glass tank with cover, large enough to hold 3 × 7 – cm TLC plate and second chromatography tank of the same size containing a few crystals of solid iodine (I2 ) 3 × 7–cm thin-layer chromatography (TLC) plate (plastic-backed silica gel G plate without fluorophore; Whatman 4410-221) cut from a 20 × 20–cm sheet Hot air dryer Synthesize di(9,10-dibromostearoyl)phosphatidylcholine 1. Weigh an empty, clean, dry 10-ml round-bottom flask. Add 2 ml of 25 mg/ml solution of DOPC in chloroform to the flask and, in a fume hood, support the flask in an ice-bucket using a retort stand. 2. Working in the fume hood and using a 5-µl repeating pipettor, add bromine dropwise to the flask in 0.5-µl increments (initially, after each addition of bromine the solution will become colorless as the bromine reacts with the lipid). Continue adding the bromine until the solution remains slightly yellow. CAUTION: Handle the bromine with care, making sure not to spill any on the skin. The total volume of bromine required will be about 7 µl.
3. Insert the stopper into the flask, cover the flask with aluminum foil to protect from light, and leave on ice ∼30 min.
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4. Remove the chloroform and excess bromine using a rotary evaporator. Depending on the vacuum used, this will take about 30 min. If the sample contains more excess bromine than can easily be removed by rotary evaporation, it can be purified by gel-filtration chromatography, applying the sample dissolved in chloroform to the column packed with silica gel, eluting the column first with chloroform to remove the bromine and then with 3:7 (v/v) chloroform/methanol to elute the phospholipid, as described in detail in Bergelson (1980).
5. Making sure that the flask is dry, weigh the flask and check that the weight of lipid is correct. There should be ∼70 mg of BrPC, starting from 50 mg of DOPC.
6. Add chloroform to the flask to make an ∼25 mg/ml solution of BrPC and store up to several weeks at −20◦ C, wrapped in foil to protect from light.
Check the purity of the brominated lipid sample The purity of the brominated lipid can be checked by thin-layer chromatography as described here, or by mass spectrometry as described in Support Protocol 2 7. Pour solvent (95:35:6 chloroform/methanol/water or 14:6:1 chloroform/ methanol/ammonia) into a glass chromatography tank to a depth of ∼3 mm. 8. Spot the BrPC onto the thin-layer chromatography plate together with a spot of DOPC. The spots should be located about 1 cm from the bottom of the plate so that they will be located above the solvent layer when placed in the chromatography tank.
9. Dry the spots, using a hot-air dryer if necessary, and carefully place the plate in the tank, making sure that the lipid spots are above the solvent layer in the tank. 10. Allow the solvent to run to the top of the plate. Remove the plate from the tank and allow it to dry in air. 11. Place the plate in another tank containing a few crystals of iodine and leave it a few minutes to allow the iodine to stain the plate. A single brown-staining spot for the BrPC sample should be observed at the same position as for the DOPC sample. SUPPORT PROTOCOL 2
ANALYSIS OF BROMINATED PHOSPHOLIPID BY MASS SPECTROMETRY The mass spectra of zwitterionic phospholipids such as phosphatidylcholines can be recorded using electrospray ionization mass spectrometry (UNIT 16.8) in either positiveor negative-ion mode with the addition of sodium or chloride ions, respectively. The corresponding brominated phospholipids are best analyzed in the negative-ion mode because the major species observed in positive ion spectra is one that has gained positive charge by losing Br− rather than by association with Na+ . The conditions given below are for recording spectra using a Quattro II triple quadrupole mass spectrometer with a standard stainless steel capillary electrospray source.
Materials
Fluorescence Quenching to Study Lipid-Protein Interactions
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1 mg/ml di(9,10-dibromostearoyl)phosphatidylcholine (BrPC; Support Protocol 1) in chloroform 7:2:1 (v/v/v) methanol/chloroform/water 2 M HCl 2 mg/ml NaI in 1:1 propan-2-ol/H2 O ∼2-ml glass test tubes 1-ml gastight syringe with Teflon luer-lock connector (Hamilton P/N 81322) Current Protocols in Protein Science
Quattro II triple quadrupole mass spectrometer with electrospray probe (Waters) PEEK tubing (0.5 m length, 1.6 mm o.d., 0.12 mm i.d.) PEEK Fingertight HPLC fittings (Upchurch) PEEK Quick Connect luer adapter (Upchurch, P/N P-659) Additional reagents and equipment for electrospray ionization mass spectrometry (UNIT 16.8) Prepare the brominated lipid solution 1. Add 50 µl of 1 mg/ml di(9,10-dibromostearoyl)phosphatidylcholine to 1 ml of 7:2:1 methanol/chloroform/water in an ∼2-ml glass test tube. 2. Add 1 µl of 2 M HCl to the tube to acidify the solution. 3. Fill a 1 ml gastight luer-lock Hamilton syringe with the solution.
Set up the mass spectrometer 4. Connect the syringe to the electrospray probe of the Quattro II mass spectrometer using PEEK tubing, PEEK Fingertight connectors, and a PEEK Quick Connect luer adapter. If using a syringe with a metal fitting or needle, these must be grounded to prevent electric shocks when connected to the electrospray source.
5. Mount the syringe in a syringe pump set to deliver 10 µl/min. 6. Select negative ion mode on the Tune page. 7. Turn on nebulizing and desolvation gas. 8. Switch into “operate” to apply source voltages. 9. Display the molecular ion using the first quadrupole analyzer (MS 1) and adjust the tuning parameters to optimize signal intensity and resolution. Typical values are:
Capillary, 2.7 kV HV lens, 0.35 kV Cone, 55 V Source temperature 80◦ C. Acquire data 10. Acquire data from 400 to 1500 m/z with a scan duration of 4.5 sec and an interscan delay of 0.1 sec. 11. Halt acquisition after 2 min and combine 25 to 30 scans to obtain a spectrum with an enhanced signal/noise ratio.
Obtain calibration spectrum and plot mass spectra 12. Record a calibration spectrum using the NaI solution (2 mg/ml in propan-2-ol/water [1:1]) using settings identical to those used for the brominated phospholipid. 13. Use the NaI spectrum to make a calibration file and apply it to the sample spectrum to correct the m/z scale. 14. Plot the mass spectrum. This should show a major peak corresponding to di(9,10-dibromostearoyl) phosphatidylcholine with a single bound chloride ion, predicted to give a peak at 1140.2 (C44 H84 Br4 ClNO8 P). The peaks are illustrated in Figure 19.12.3. The isotope pattern is complex due to the two isotopes of bromine of equal abundance, differing by 2 mass units, and the two isotopes of chlorine, of mass units 35 and 37 and relative abundances of 76% and 24%, respectively (Fig. 19.12.3). Current Protocols in Protein Science
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Figure 19.12.3 Negative ion electrospray mass spectrum of di(9,10-dibromostearoyl)phosphatidylcholine in the presence of HCl, showing the peak at 1140.4 corresponding to the (M+Cl)− species. Insert: expanded mass scale showing the isotope cluster for (M+Cl)− .
SUPPORT PROTOCOL 3
PURIFICATION OF POTASSIUM CHOLATE Commercial samples of cholic acid should be recrystallized to remove impurities as described here.
Materials Cholic acid Methanol, pure-grade Potassium hydroxide Diethyl ether 1-liter Erlenmeyer flask 40◦ C water bath Large sintered-glass filter funnel Glass beaker 1. Weigh out 20 g (48.9 mmol) cholic acid into a 1-liter conical flask. 2. Gradually add ∼150 ml of methanol to the flask and warm to about 40◦ C in a water bath until all the cholic acid has dissolved. 3. Slowly add an equimolar amount of potassium hydroxide (2.75 g), swirling the flask until all has dissolved. 4. Fill the flask with diethyl ether. The potassium cholate should immediately form a precipitate. Fluorescence Quenching to Study Lipid-Protein Interactions
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5. Leave the potassium cholate to settle, then filter under vacuum using a large sinteredglass filter funnel. 6. Spread the potassium cholate as a thin layer in a glass beaker and leave in a fume hood under a gentle stream of air to evaporate off the diethyl ether. When free of ether, store up to several months at 5 ◦ C. Current Protocols in Protein Science
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
HEPES/EGTA buffer 40 mM HEPES 100 mM KCl 1 mM EGTA Adjust pH to pH 7.2 with KOH Store up to 2 weeks at 5◦ C All reagents should be the purest available to minimize any fluorescent impurities. EGTA is slow to dissolve; therefore, add it as a small aliquot of 100 mM EGTA dissolved in concentrated KOH. The buffer contains EGTA to complex any divalent metal ions present.
COMMENTARY Background Information Quenching of Trp fluorescence by brominated phospholipids is fitted to a lattice model for quenching (Caffrey and Feigenson, 1981; London and Feigenson, 1981; Mall et al., 1998; O’Keeffe et al., 2000). Because the time for two lipid molecules to exchange between the bulk lipid phase and the annular shell of lipids around a membrane protein is greater than the fluorescence lifetime (London and Feigenson, 1981; East et al., 1985), the level of fluorescence quenching observed in a mixture of a normal and a quenching lipid is proportional to the fraction of annular sites occupied by the quenching lipid, and is thus dependent on the binding constant of the quenching lipid.
Analysis of fluorescence results The probability that any particular Trp residue will give rise to fluorescence is proportional to the probability that none of the n lattice sites close enough to the residue to cause quenching are occupied by a molecule of brominated lipid (Fig. 19.12.1). The value of n for phosphatidylcholines is determined from experiments measuring fluorescence quenching in mixtures of DOPC and BrPC. It is convenient to express the fraction of BrPC in the membrane by its mole fraction xBr in the membrane, calculated from the equation:
Equation 19.12.1
where [BrPC] and [DOPC] are, respectively, the concentrations of BrPC and DOPC in the sample, expressed in the usual units of moles per liter. The mole fraction xDOPC of DOPC in
the membrane is given by:
Equation 19.12.2
A Trp residue in the protein will be fluorescent if none of the n lipid binding sites close enough to it to cause quenching are occupied by BrPC. The Trp fluorescence intensity in the mixture of DOPC and BrPC can therefore be written as:
Equation 19.12.3
where Fo and Fmin are the fluorescence intensities for protein in DOPC and in BrPC lipid, respectively, and F is the fluorescence intensity in the phospholipid mixture when the mole fraction of BrPC lipid is xBr . This equation assumes that BrPC and DOPC bind equally well to the protein. For convenience, if the fluorescence intensities are scaled to a value of 1 in DOPC, then Equation 19.12.3 becomes:
Equation 19.12.4
Figure 19.12.4 shows the meaning of Equation 19.12.4. In that figure, the value of the fluorescence intensity F calculated from Equation 19.12.4 for a membrane protein in a mixture of DOPC and BrPC varies with BrPC content for various assumed values of n; as the value of n increases and the Trp fluorescence is quenched from more lipid-binding sites, the quenching curves become steeper. Fitting a set of experimental quenching data for a mixture of a brominated lipid with its nonbrominated equivalent to Equation 19.12.4 gives the value of n for that lipid and protein. Typical values
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Figure 19.12.4 Effect of n, the number of lipid-binding sites on the protein from which a bound BrPC molecule can quench the fluorescence of a given Trp residue in a protein. The figure shows a set of theoretical quenching curves for a membrane protein reconstituted into bilayers containing mixtures of DOPC and BrPC, showing values of F/Fo calculated from Equation 19.12.4 with the given values of n, for a value of Fmin of 0.25.
for n are in the range 1.8 to 3.0 (Powl et al., 2005). To determine lipid-binding constants relative to DOPC, the membrane protein is reconstituted into mixtures of BrPC and a nonbrominated lipid X. The equations given above are readily extended to this case. Binding of lipid X and BrPC at each site around the protein can be described by a series of displacement reactions:
The mole fraction xX of lipid X can be expressed as:
Equation 19.12.8
The fractional occupancy of the site by BrPC, fBr , is given by:
Equation 19.12.9 Equation 19.12.5
This equilibrium is described by an equilibrium constant K, which is the binding constant at the site for lipid X relative to the binding constant at the site for BrPC, given by:
Equation 19.12.6
As long as the molar ratio of lipid:protein is sufficiently high that binding of lipid to the protein does not significantly deplete the concentration of lipid in the membrane, this can be rewritten as: Fluorescence Quenching to Study Lipid-Protein Interactions
From Equation 19.12.6, fBr can be expressed as:
Equation 19.12.10
If it is assumed that all the lipid binding sites on the membrane protein are equivalent, so that binding at each site can be described by the same equilibrium constant K, the fluorescence intensity for the protein will be described by an equation equivalent to Equation 19.12.3:
Equation 19.12.11 Equation 19.12.7
where xBr and xX are the mole fractions of BrPC and lipid X in the mixture, respectively.
with fBr given by Equation 19.12.10. Figure 19.12.5 shows how fluorescence intensity in a mixture of lipid X and BrPC varies with the mole fraction of BrPC in the mixture, for
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Figure 19.12.5 The figure shows a set of theoretical quenching curves for a membrane protein reconstituted into bilayers containing mixtures of lipid X and BrPC, showing values of F/Fo calculated from Equation 19.12.11 with n = 1.69, for a value of Fmin of 0.25, for the given values of the binding constant of lipid X relative to BrPC.
various values of the relative binding constant K. The stronger the binding of lipid X relative to BrPC (higher K values), the greater the fluorescence intensity observed at intermediate mole fractions of BrPC, because strong binding of lipid X means that little BrPC will be bound to the protein and there will be little fluorescence quenching. Conversely, weak binding of lipid X relative to BrPC will result in low fluorescence intensities at intermediate mole fractions of BrPC, because weak binding of lipid X means that many of the lipid-binding sites on the protein will be occupied by BrPC and that there will therefore be extensive fluorescence quenching. Fitting a set of quenching data for a mixture of BrPC with lipid X to Equation 19.12.11 yields the relative binding constant for lipid X compared to that for BrPC. Having determined a relative binding constant from measurements of fluorescence quenching in mixtures of lipid X with BrPC, it is important to repeat the measurements in mixtures of brominated lipid X and DOPC. The relative binding constants determined in the two sets of experiments will be the same if the relative binding constant is the same at all the lipid-binding sites on the protein from which fluorescence can be quenched. This is found to be the case, for example, for phosphatidylethanolamine binding to KcsA (Alvis et al., 2003). However, it is not the case for anionic lipid binding to KcsA, where higher rel-
ative binding constants for anionic lipids are obtained from experiments with brominated anionic lipids than from experiments with the equivalent nonbrominated anionic lipid mixed with BrPC (Alvis et al., 2003). The explanation is that all the lipid-binding sites on KcsA are not equivalent. Some show a high affinity for anionic lipid, explaining the marked quenching observed with brominated anionic lipid, and some show similar binding constants for anionic lipids and phosphatidylcholine. Binding to these latter sites is most important for the observed fluorescence quenching in experiments with BrPC (Alvis et al., 2003). Although these experiments indicate heterogeneous binding to KcsA and show that anionic lipids can bind with high affinity, derivation of actual binding constants requires a knowledge of the distribution of the Trp residues in the protein relative to the location of the various classes of lipid-binding site, and this is generally not known. An especially simple case, however, is a membrane protein containing a single Trp residue and, in this case, it is possible to derive individual binding constants for the various classes of lipid-binding site (Powl et al., 2005).
Critical Parameters The experiments described in this unit require the purified membrane protein of interest in milligram quantities, and this is clearly
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a significant limitation. Although it would be possible to study a membrane protein available in smaller amounts, the experiments would be technically more demanding. The measurements involve reconstitution of the purified protein into lipid bilayers of defined composition; this is relatively easy as long as the protein is stable and soluble in a detergent of high critical micelle concentration (CMC; see APPENDIX 1B), such as dodecylmaltoside or octylglucoside. The protein to be studied must contain Trp residues, with greater than ∼25% of the Trp residues being located in the transmembrane region where they can be quenched by brominated lipid molecules. Before embarking on a full set of experiments with a new membrane protein, it is therefore recommended that it be ascertained that the fluores-
Fluorescence Quenching to Study Lipid-Protein Interactions
cence is quenched significantly in brominated lipid, as described in Basic Protocol 1. Finally, the fluorescence measurements require the use of a fluorimeter of high sensitivity, with good monochromators to reduce the effects of light scatter from cloudy membrane samples.
Troubleshooting A critical step in the reconstitution procedure described above is the dispersal of the lipid into detergent micelles by ultrasonication. This requires careful selection of an ultrasonication bath, as described in the protocols above. It is important to remove all organic solvent when the lipid sample is lyophilized, since any residual solvent could partition into the final reconstituted membrane sample to a relatively high concentration within the
Figure 19.12.6 Experimental fluorescence quenching plots. KcsA was reconstituted into mixtures of DOPC and BrPC (A) and DOPE and BrPC (B). Fluorescence intensities are expressed as F/Fo where Fo is the fluorescence intensity in the nonbrominated lipid. The solid line in (A) shows a fit to Equation 19.12.3, giving a value for n of 1.66 ± 0.12. The solid line in (B) shows a fit to Equation 19.12.11 giving a value for the relative binding constant K for DOPE of 0.57 ± 0.09.
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membrane, leading to potential denaturation of the protein. As in all fluorescence measurements, it is necessary to use materials of the highest available purity to minimize the presence of fluorescent impurities. The possibility of fluorescence bleaching during the experiments can be minimized by stirring the sample during the fluorescence measurements. Fluorescence artifacts can also arise from adsorption of protein onto the walls of the fluorescence cuvette; the chance of this happening can be minimized by keeping the cell clean and soaking it occasionally in concentrated hydrochloric acid. It is best to avoid washing the cell in any detergentcontaining cleaner.
Anticipated Results The experiments described in this unit will generate two quenching curves, one for the membrane protein in mixtures of DOPC and BrPC and one for the membrane protein in mixtures of DOPE and BrPC (Fig. 19.12.6). Fitting the first to Equation 19.12.4 gives the value of n required to fit the second set of data to equation 19.12.11 to obtain the relative binding constant K for DOPE relative to BrPC.
Time Considerations It is convenient to lyophilize the lipid samples from organic solvent overnight. Determination of a value for n and for a relative binding constant each take about half a workday. Preparing the set of tubes containing the required mixtures of brominated and nonbrominated lipids (Basic Protocol 2, steps 1 to 6) takes about 3 hr, and the fluorescence measurements themselves take about 1 hr.
Literature Cited Alvis, S.J., Williamson, I.M., East, J.M., and Lee, A.G. 2003. Interactions of anionic phospholipids and phosphatidylethanolamines with the potassium channel KcsA. Biophys. J. 85:1-11. Bergelson, L.D. 1980. Lipid Biochemical Preparations. Elsevier/North Holland, Amsterdam. Caffrey, M. and Feigenson, G.W. 1981. Fluorescence quenching in model membranes. 3. Relationship between calcium adenosinetriphosphatase enzyme activity and the affinity of the protein for phosphatidylcholines with different acyl chain characteristics. Biochemistry 20:1949-1961. Doyle, D.A., Cabral, J.M., Pfuetzner, R.A., Kuo, A., Gulbis, J.M., Cohen, S.L., Chait, B.T., and Mackinnon, R. 1998. The structure of the potassium channel: Molecular basis of K+ conduction and selectivity. Science 280:69-77. East, J.M. and Lee, A.G. 1982. Lipid selectivity of the calcium and magnesium ion dependent adenosinetriphosphatase, studied with fluores-
cence quenching by a brominated phospholipid. Biochemistry 21:4144-4151. East, J.M., Melville, D., and Lee, A.G. 1985. Exchange rates and numbers of annular lipids for the calcium and magnesium ion dependent adenosinetriphosphatase. Biochemistry 24:2615-2623. Froud, R.J., East, J.M., Rooney, E.K., and Lee, A.G. 1986. Binding of long-chain alkyl derivatives to lipid bilayers and to (Ca2+ −Mg2+ )-ATPase. Biochemistry 25:7535-7544. Lee, A.G. 2004. How lipids affect the activities of integral membrane proteins. Biochim. Biophys. Acta 1666:62-87. London, E. and Feigenson, G.W. 1981. Fluorescence quenching in model membranes. 2. Determination of local lipid environment of the calcium adenosinetriphosphatase from sarcoplasmic reticulum. Biochemistry 20:1939-1948. Mall, S., Sharma, R.P., East, J.M., and Lee, A.G. 1998. Lipid-protein interactions in the membrane: Studies with model peptides. Faraday Discuss. 111:127-136. Marsh, D. and Horvath, L.I. 1998. Structure, dynamics and composition of the lipid-protein interface: Perspectives from spin-labelling. Biochim. Biophys. Acta 1376:267-296. O’Keeffe, A.H., East, J.M., and Lee, A.G. 2000. Selectivity in lipid binding to the bacterial outer membrane protein OmpF. Biophys. J. 79:20662074. Pilot, J.D., East, J.M., and Lee, A.G. 2001. Effects of bilayer thickness on the activity of diacylglycerol kinase of Escherichia coli. Biochemistry 40:8188-8195. Powl, A.M., East, J.M., and Lee, A.G. 2003. Lipidprotein interactions studied by introduction of a tryptophan residue: The mechanosensitive channel MscL. Biochemistry 42:14306-14317. Powl, A.M., East, J.M., and Lee, A.G. 2005. Heterogeneity in the binding of lipid molecules to the surface of a membrane protein: Hot-spots for anionic lipids on the mechanosensitive channel of large conductance MscL and effects on conformation. Biochemistry 44:5873-5883. Simmonds, A.C., East, J.M., Jones, O.T., Rooney, E.K., McWhirter, J., and Lee, A.G. 1982. Annular and non-annular binding sites on the (Ca2+ + Mg2+ )-ATPase. Biochim. Biophys. Acta 693:398-406. Williamson, I.M., Alvis, S.J., East, J.M., and Lee, A.G. 2002. Interactions of phospholipids with the potassium channel KcsA. Biophys. J. 83:2026-2038.
Contributed by Joanne Carney, J. Malcolm East, Sanjay Mall, Phedra Marius, Andrew M. Powl, J. Neville Wright, and Anthony G. Lee University of Southampton Southampton, United Kingdom
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Overview of Biacore Systems and Their Applications BIACORE SYSTEM OVERVIEW Introduction Surface plasmon resonance (SPR) was first shown to be amenable for the label-free study of interactions between biomolecules over 20 years ago (Lieberg et al., 1983). Biacore pioneered commercial SPR biosensors offering a unique technology for collecting highquality, information-rich data from biomolecular binding events. Since the release of the first instrument in 1990, researchers around the world have used Biacore’s optical biosensors to characterize binding events with samples ranging from proteins, nucleic acids, small molecules to complex mixtures, lipid vesicles, viruses, bacteria, and eukaryotic cells. Typical questions answered with Biacore instruments include:
r r r r r
How specific is an interaction? How strong is an interaction? What is the affinity? How fast is an interaction? What are the association and dissociation rate constants? Why is the interaction that strong or that fast? What are the thermodynamic parameters for an interaction? What is the biologically active concentration of a specific molecule in a sample?
This unit describes the various instrument platforms available and their primary uses.
System Features Biacore’s optical biosensors are designed around three core technologies: 1. An optical detector system that monitors the changes in SPR signal brought about by binding events in real time. 2. An exchangeable sensor chip upon which one of the interacting biomolecules is immobilized or captured. The resulting biospecific surface is the site where biomolecular interactions occur. Sensor chips are discussed in greater detail in Surface Preparation. 3. A microfluidic and liquid handling system that precisely controls the flow of buffer and sample over the sensor surface. These systems are contained within the processing unit that communicates with a com-
puter equipped with control and data evaluation software.
The SPR detection system Surface plasmon resonance (SPR) is a phenomenon that occurs in thin conducting films placed at the interface between two media of different refractive indices. In Biacore systems, a 50-nm layer of gold on the sensor chip is sandwiched between the glass layer of a sensor chip and the sample solution flowing through the microfluidic cartridge. Planepolarized light from a near-infrared LED is focused on the back of the sensor chip under conditions of total internal reflection and a diode array detector monitors the intensity of the reflected light. Under these conditions, the light leaks an electromagnetic component called an evanescent wave across the gold interface into the sample/buffer solution. At a certain angle of incident light, the evanescent wave field excites electrons in the gold film resulting in the formation of surface plasmons (electron charge density waves) within the gold film with a concomitant drop in the intensity of the reflected light at this angle (SPR angle). When a change in mass occurs near the sensor chip surface, e.g., as a result of a binding event, the angle of light at which SPR occurs shifts due to a change in refractive index near the sensor chip surface. A sensorgram depicts changes in SPR angle in real time, with responses measured in resonance units (RU). One RU corresponds to 0.0001◦ shift in SPR angle. Since only the evanescent wave penetrates the sample, measurements can be carried out with colored, turbid, or even opaque solutions without interference from conventional light absorption or light scattering. Flexchip instruments use a different configuration for SPR detection called gratingcoupled SPR. In the grating-coupled configuration, incident light is focused through the biomolecular layer onto the gold layer. A fine grating on the chip surface provides optical coupling and allows imaging of the entire surface at once, enabling simultaneous real-time detection of binding interactions taking place on up to 400 spots on the surface.
Contributed by Laure Jason-Moller, Michael Murphy, and JoAnne Bruno Current Protocols in Protein Science (2006) 19.13.1-19.13.14 C 2006 by John Wiley & Sons, Inc. Copyright
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The microfluidic system Biacore systems (except Flexchip) make use of an integrated microfluidic cartridge (IFC) to deliver samples and buffer to the sensor surface. The IFC consists of a series of micro-channels encased in a plastic housing. In automated systems, samples are transferred through a needle into the IFC. Flow cells are created through pressure contact between molded channels in the IFC and the sensor chip surface. A series of computer-controlled pneumatic valves meters precise access to the channels and the flow cells. The design of the IFC, as well as the number and configuration of flow cells, is specific to each instrument platform. Pulse-free syringe pumps ensure a continuous flow of buffer over the sensor surface whenever samples are not being injected. In Flexchip, one large flow cell with an inlet and outlet is created on the sensor surface by installing an adhesive-gasketted window over the sensor surface following external immobilization of ligands.
Available Instrument Platforms Life science research Biacore X Biacore X is a highly sensitive, semiautomated system for label-free protein interaction studies. There are two flow cells in the Biacore X instrument. Two different molecules can be immobilized on one sensor chip or one flow cell can be used as a reference and the other immobilized with a molecule of interest. Dual-channel detection of the SPR signal and serial flow allow simultaneous monitoring of both flow cells. The instrument control software can be set to automatic in-line reference subtraction of the control surface signal from the same sample injection. Sample bound to the sensor chip surface can be recovered for complementary analysis and/or identification, e.g., using mass spectrometry. Biacore X is well suited for laboratory environments with low-sample throughput where many users handle different types of samples.
Biacore 2000 and 3000
Overview of Biacore Systems
Biacore 2000 and 3000 are currently the most widely used Biacore instruments. They are equipped with a robotic autosampler and can be programmed through the use of wizards or methods in the control software to automatically inject samples for up to two 96well microtiter plates. The fluidic system in these instruments creates four serially linked
flow cell areas on one sensor chip, thereby increasing the potential number of simultaneous measurements. For example, it is possible to simultaneously collect data from a reference surface as well as three other surfaces with different immobilized ligands as well as reference-subtracted data for each surface. Flow cells can also be addressed individually, which allows for the creation of different surfaces on one sensor chip. The dynamic range of the SPR detector is greater in Biacore 3000 than in Biacore 2000. This expands the range of interactions that can be studied in Biacore 3000 to high refractive index solutions or high density surfaces. In Biacore 3000, a microrecovery protocol is available to recover material bound to the sensor chip surface for delivery either to a vial or a MALDI target plate for further analysis. The U.S. Food and Drug Administration’s Title 21 Code of Federal Regulations (21CFR Part 11) compliant software and instrument validation are available for Biacore 3000 to meet FDA regulations for GxP (Good Laboratory Practice, Good Manufacturing Practice, Good Clinical Practice, etc.) environments. GxP software includes, for example, restricted user level access, audit trail, and method publishing. GxP instrument validation includes, for example, equipment qualification services, and Part 11 assessment reports. The open platforms of Biacore 2000 and 3000 suggest that they are well suited for use in versatile environments ranging from research to preclinical studies.
Biacore T100 Biacore T100, released in 2005, is fully automated and is the latest generation of the fourflow cell SPR biosensors for protein interaction analysis. New hardware features include 384-well microtiter plate capacity, a built-in buffer degasser that eliminates the requirement for daily degassing of running buffers, and allows for up to 48 hr of unattended operation. New software functionality and a buffer selector valve enables users to carry out automatic analysis of up to four different buffer conditions or at different temperatures ranging from 4◦ to 45◦ C. The software has a number of built-in wizards to guide users in operations ranging from the initial immobilization of ligands to widely used assays such as concentration analysis, kinetic analysis, determination of solution affinity, and determination of thermodynamic parameters. In addition to equilibrium thermodynamic parameters, transition-state thermodynamic parameters can be derived from the rate constants
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obtained over a range of temperatures. The fast liquid exchange rates resulting from a redesigned microfluidic system enable the characterization of interactions with off rates as fast as 0.5 sec−1 . Biacore T100 is well suited to all types of protein interaction analysis with molecules as small as 100 Da or as large as phages. Validated (21 CFR Part 11 compliant) hardware and software options are available for use in regulated environments.
ment of samples is required since SPR measurements can be carried out in opaque or turbid samples. Wizard-based Biacore Q control software facilitates assay development, optimization, and routine testing. The wizards also provide automatic and user-specific result evaluation.
Drug discovery and development
Flexchip
Biacore C Biacore C is a dedicated SPR instrument for the measurement of biologically active sample concentration in a regulated environment. Biacore C is used in applications such as in-process control, batch release, and quality control. The ability of SPR systems to detect binding events in complex mixtures such as fermentation media, cell culture supernatants, and serum allows users to make rapid decisions based on the active concentration of sample, e.g., in determining the optimal time to harvest a bioreactor. Biacore C software is validated to support GxP-regulated working processes and includes technical controls for 21 CFR Part 11 compliance.
Biacore S51 High-throughput screening of drug candidates often produces many false-positive hits. Biacore S51 offers a unique platform designed for secondary screening applications and kinetic characterization of low-molecularweight compound leads. The unique design of the microfluidic cartridge allows for the detection of fast dissociating interactions (dissociation rate constants kd , up to 1 sec−1 ). There are two independent flow cells in Biacore S51, each of which contains three detection spots. The central spot is used as a reference surface, while different molecules can be immobilized on the outer two spots using hydrodynamic addressing. Biacore S51 software supports automatic calibration procedures used when analyzing samples that require up to 10% DMSO to maintain solubility.
Food analysis Biacore Q, together with Qflex kits, provides rapid, reliable solutions for determination of vitamin content and detection of veterinary drug residues in a wide variety of food-related products. Minimal pretreat-
Arrays
The Flexchip array platform features a single flow cell on a disposable sensor chip with the capacity for 400 target spots together with 400 adjacent reference spots. Immobilization on the sensor chip surface is performed outside the instrument with an external spotter. The detection system uses grating-coupled SPR. The system is well suited for multiplex analysis, ranking based on kinetic rate constants or affinity constants and epitope mapping. The minimum molecular weight of analyte for analysis with Flexchip is 5000 Da.
Biacore A100 Biacore A100 is Biacore’s most advanced SPR array system. A unique microfluidic system defines four independent flow cells, each with five detection spots. This means that assays can be multiplexed with up to 20 different immobilized targets per chip. The sensor chips are reusable, enabling multiplexing of samples as well. Binding interactions can be analyzed simultaneously in up to four different buffer conditions. Several hundred samples can be analyzed in a run due to an onboard degasser and a rack hotel that automatically exchanges 96- or 384-well sample plates. Biacore A100 is sensitive enough to detect the binding of low molecular weight analytes > 100 Da therefore it can be used in drug discovery applications to screen or characterize the interaction between small molecules and a panel of protein targets. Biacore A100 is well suited for information-rich, high-throughput kinetic characterization enabling selection of antibodies at early stages of development (e.g., hybridoma supernatants) and large-scale proteomics studies. Biacore A100 is also suitable for use in regulated environments.
SURFACE PREPARATION Sensor Chip Options Although SPR can be generated in thin films made from conducting metals, all Biacore sensor chips are coated with a thin,
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uniform gold layer. Gold has a number of advantages in that it results in a well-defined reflectance minimum when a visible light source is used to generate the SPR signal, gold is amenable to covalent attachment of surface matrix layers and, in physiological buffer conditions, gold is mostly inert. Sensor chip surfaces, except Sensor Chip Au, are covered with a covalently bound alkanethiol monolayer to prevent nonspecific adsorption of proteins to the gold surface. Sensor chips are reusable since noncovalently bound material can be removed from the Sensor Chip surface with an injection of a suitable regeneration solution.
Nonderivatized dextran-containing sensor chips The surface of Sensor Chips CM5, CM4, and CM3 is covered with a hydrogel matrix of flexible, unbranched carboxymethylated dextran covalently attached to the surface alkanethiol monolayer. The dextran layer ranges in thickness from ∼25 to 100 nm, depending on the sensor chip type. The dextran layer provides a hydrophilic, solution-like environment favorable for most interactions between proteins or other biomolecules. The carboxyl moieties can be derivatized using a wide range of well-defined chemistries for the covalent attachment of biomolecules. The flexibility of the unbranched dextran polymer allows covalently attached molecules to move with relative freedom within the surface layer. The dextran layer also increases the surface capacity in comparison with a two-dimensional surface.
Derivatized dextran-containing sensor chips
Overview of Biacore Systems
Three pre-derivatized chips are currently available for Biacore instruments, namely Sensor Chips SA, NTA, and L1. Sensor Chip SA is pre-immobilized with streptavidin and is suitable for the capture of biotinylated molecules (e.g., nucleic acids, peptides, proteins, liposomes, glycosaminoglycans). The capture of biotinylated molecules is essentially irreversible due to the high affinity of the biotinstreptavidin interaction (KD = 10−15 M). Histidine (His)-tagged recombinant proteins can be reversibly captured on NiCl2 -activated nitrilotriacetic acid (NTA) groups that are covalently coupled to the dextran matrix of Sensor Chip NTA. Captured molecules can easily be removed from the sensor chip surface with a pulse of EDTA. The affinity of the His-tag for the activated NTA surface is dependent on the microenvironment of the tag and, therefore,
the baseline stability following the capture step can vary from protein to protein. Sensor Chip L1 is derivatized with lipophilic anchors (alkane chains) designed to intercalate into lipid bilayers enabling the stable capture of intact membrane structures and liposomes.
Non-dextran sensor chips Sensor Chip C1 has a flat carboxylated surface without a dextran matrix. The carboxyl groups can be derivatized using the same immobilization chemistries used for covalent coupling to the dextran surface of Sensor Chips CM5, CM4, and CM3; however, the reduction in available surface area results in a significant decrease in the immobilization capacity (typically ∼10% of that on Sensor Chip CM5 under comparable conditions). Sensor Chip C1 is useful for monitoring interactions with large interactants such as cells and viruses. It also provides a suitable surface for studies where the flexibility of the dextran layer is not wanted (e.g., homodimerization studies, reducing avidity effects). Sensor Chip HPA lacks a carboxymethylated dextran matrix so that lipid monolayers can be formed on the hydrophobic thioalkane surface layer covering the gold film. Sensor Chip HPA is suitable for measuring interactions with a lipid monolayer or interactions with ligands associated with a membrane surface or partially inserted into the membrane. Sensor Chip L1 is more suitable for deeply inserted or transmembrane proteins. Sensor Chips Au and SIA Kit Au consist of a bare gold surface, which lends itself to the design of customized surface chemistries using self-assembled monolayers (SAM) or to study interactions between surface materials and biomolecules. Sensor Chip SIA Kit Au contains unmounted gold surfaces and a separate chip carrier for easy assembly after surface coating. This allows for the use of a wide variety of coating techniques, including those using harsh conditions (such as organic solvents) that the chip carrier would not usually withstand.
Flexchip sensor chips A number of sensor surfaces are currently available for Flexchip including bare gold, streptavidin, neutravidin, and protein A/G– coated chips. Proteins can be spotted on the bare gold surface that can subsequently be blocked with blocking solution to minimize non-specific interactions. Antibodies or Fc
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fusion proteins can be spotted onto protein A/G chips, thereby resulting in a more oriented surface than that created by spotting on a bare gold surface. Streptavidin and neutravidin surfaces can be used with biotinylated peptides, nucleic acids, and proteins.
Immobilization Chemistries A critical step in the development of reliable SPR assays is the selection of the most suitable immobilization technique such that ligand activity is maintained and binding sites are available to interacting partners. Commonly utilized immobilization strategies are outlined here. Guidelines and a detailed review of immobilization chemistries have been published recently (Karlsson and Larsson, 2004).
Direct immobilization chemistries A number of covalent immobilization techniques are available to covalently attach proteins or other biomolecules to the dextran on the sensor chip surface. All types of immobilization can be performed directly in the Biacore system, and a typical immobilization reaction usually takes <30 min to complete. Immobilization is usually carried out at low flow rates (5 to 10 µl/min) and the concentration of ligand used for immobilization is typically in the range of 1 to 100 µg/ml. To ensure specificity, it is recommended that the ligand purity exceed 95%.
Amine coupling Amine coupling is the most generally applicable covalent coupling chemistry used to immobilize protein ligands. Immobilization is via free primary amine groups such as lysine residues that are abundant in most proteins or the N-terminus of proteins and peptides. The dextran matrix on the sensor chip surface is first activated with a 1:1 mixture of 0.4 M 1-ethyl-3-(3dimethylaminopropyl) carbodiimide (EDC) and 0.1 M N-hydroxysuccinimide (NHS) to create reactive succinimide esters. The standard activation time is 7 min; this can be varied from 1 to 10 min to create fewer or more, respectively, reactive groups on the sensor chip surface depending on the immobilization level required. The ligand is then injected in low salt buffer lacking primary amines at a pH below the isoelectric point (pI) of the protein. Under these conditions, the ligand acquires a positive charge and is effectively preconcentrated into the negatively charged carboxymethyl dextran
matrix. High local concentrations of ligand obtained through electrostatic preconcentration maximize the efficiency of amine coupling. Finally, unreacted esters are blocked with ethanolamine. The volume or concentration of ligand injected may be varied to adjust the immobilization level. The ligand may be prevented from attaching to the surface via sites important to the interaction under study by having analyte present during immobilization. Stabilization of ligand activity during immobilization was recently reported by Casper et al. (2004), where conformationally sensitive protein kinases, p38α and GSK3β, were immobilized on Sensor Chip CM5 by amine coupling in the presence of a specific reversible inhibitor. This treatment resulted in a more stable surface with much higher specific binding activity. In cases where amine coupling interferes with the binding site on the ligand or in the case of very acidic proteins, the ligand can be attached using alternative coupling chemistries or a high-affinity capture approach.
Thiol coupling Thiol coupling is based on exchange reactions between thiol and active disulfide groups. In the case where the ligand has a free thiol group (typically cysteine residues), a surface thiol approach can be used where a disulfide group is introduced on the sensor chip dextran by first activating the surface with NHS and EDC to amine couple 2-(2-pyridinyldithio) ethaneamine (PDEA). Injection of the ligand results in thiol-disulfide exchange and excess PDEA groups are inactivated with cysteineHCl. If the ligand lacks a free thiol group, a reactive disulfide (PDEA) can be linked to carboxyl groups on the ligand. The pyridyldisulfide groups linked to the ligand are then coupled to thiol groups on the sensor surface that has been derivatized through injection of NHS and EDC, cystamine and subsequent reduction with DTE or DTT. Modification of carboxyl groups results in an increase in the isoelectric point of the proteins, which is of additional benefit in the case of acidic proteins.
Maleimide coupling Maleimide coupling is an alternative form of thiol coupling, which involves the formation of a stable thioether bond between reactive maleimido groups on the sensor surface and the thiol groups of the ligand molecule. Such surfaces have the capacity to withstand basic pH (>9.5) as well as reducing agents such as
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β-mercaptoethanol and dithiothreitol. Several heterobifunctional reagents are available for introduction of reactive maleimido groups to the sensor surface, including sulfoMBS (m-maleimidobenzoyl-N-hydroxysulfosuccinimide ester), sulfo-SMCC (sulfosuccinimidyl-4-(N-maleimidomethyl)cyclohexane-l-carboxylate), and GMBS (N(g-maleimidobutyryloxy)sulfosuccinimide ester).
Aldehyde coupling Aldehyde coupling involves the formation of a hydrazone bond via condensation of hydrazide groups on the sensor surface with aldehyde groups on the ligand molecule. These aldehyde moieties may be native to the protein or introduced through mild oxidation of cis-diols present in the ligand molecule. Aldehyde coupling is particularly useful for sitedirected immobilization of glyco-conjugates, glyco-proteins, and polysaccharides, and may also be useful for orientation-specific immobilization of proteins containing functional groups that may be converted to aldehyde moieties.
Indirect (capture) immobilization General capture methods Capture approaches provide an alternative to covalent immobilization and take advantage of tags commonly used for ligand purification. This technique involves high-affinity capture of the ligand onto a capturing molecule that has been covalently immobilized using one of the techniques described earlier. The requirement for ligand purity is less stringent for capture approaches than for covalent immobilization since the capture step can also provide a ligand purification step. Another benefit of capture approaches is the creation of a homogenous surface since all ligands are similarly oriented through a common site on the ligand (the tag). The affinity of the ligand for the capture molecule should be sufficiently high to ensure that little or no ligand dissociates from the surface for the duration of an analysis cycle. Monoclonal antibodies are frequently used as capture molecules, e.g., antiGST antibodies can be immobilized and used to capture GST-tagged molecules. In general, regeneration of the surface removes both the ligand and the analyte at the end of an assay cycle such that fresh ligand must be captured for a new cycle.
Specialized capture methods The high-affinity interaction between streptavidin and biotin (KD 10−15 M) makes it an extremely useful system for the capture of biotinylated ligands. Sensor Chip SA provides a robust, user-friendly streptavidin capture surface. Owing to the high affinity of the interaction, biotinylated ligands are immobilized irreversibly and, unlike most other capture approaches, cannot be removed from the sensor chip surface. Histidine (His)-tagged recombinant proteins can be captured on Sensor Chip NTA following activation of the surface with a pulse of NiCl2 . Ligands can be removed easily from the surface by stripping the nickel ions with a pulse of EDTA. An alternative to the NTA surface is to capture His-tagged ligands onto covalently immobilized anti-His antibodies. Large baseline drifts caused by unstable ligands or poor capture may be overcome by using EDC/NHS as a post-capture cross-linking step, i.e., crosslinking the tagged protein to the anti-tag antibody. This step may compromise ligand activity if active sites of the ligand are involved in the cross-linking; therefore, it must be empirically tested for each ligand-analyte system. In general, crosslinking should be as brief as possible; 15 sec is often sufficient to achieve acceptable baseline stability without compromising ligand activity. Lipid monolayers can be adsorbed onto the hydrophobic surface of Sensor Chip HPA while liposomes can be maintained in a membrane-like lipid bilayer structure on the surface of Sensor Chip L1. On-surface reconstitution (OSR) is a recently developed methodology for handling membrane proteins on Sensor Chip L1. In this process, the membrane protein is solubilized initially in detergent and then either captured through a tag or immobilized on the chip surface using amine coupling. The next step is to inject mixed micelles, which bind to the lipophilic tails on the sensor chip surface and to the ligand. Finally, the detergent is removed, inducing the lipids to form a plasma membrane-like bilayer, linked to the immobilized membrane protein via the natural affinity of lipids for the hydrophobic regions of the proteins. In this way, it is possible to present a uniform, oriented field of plasma membrane proteins within a lipid bilayer on a sensor chip and to study how they interact with binding partners (Karlsson and L¨of˚as, 2002; Stenlund et al., 2003; Navratilova et al., 2005).
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ASSAY TYPES Binding Specificity Biacore is well suited to carry out qualitative studies to confirm the specificity of interactions as well as quantitative measurements for affinity, kinetics, and concentration determination. A small volume of analyte can be tested easily for selective binding to 2 to 400 targets simultaneously, depending on the instrument platform chosen. Furthermore, analyte activity is not compromised since interactants do not need to be labeled. It is possible to monitor a number of sequential binding events since each yields a concomitant increase in mass on the sensor chip surface and all stages in the binding process can thus be monitored. This approach has been used to confirm the results of conventional immunoprecipitation assays and reduce assay time (Young et al., 2002). Epitope mapping of monoclonal antibodies can be carried out at much earlier stages in development using Biacore since little sample is required and samples do not need purification or labeling. Unfractionated hybridoma culture supernatants have been used successfully for epitope mapping studies (F¨agerstam et al., 1990). The technique is well suited to unattended instrument operation, and many samples can be processed without user intervention. This feature is valuable in epitope specificity determination of a large panel of MAbs, where the pair-wise combination matrix requires a large number of assay cycles. Other examples of specificity assays include identification of binding sites (Jokiranta et al., 2000), monitoring steps involved in complex formation (Schuster et al., 1993; Clark et al., 2001; Thai and Ogata, 2005), and assessing cofactor requirements for an interaction to occur (e.g., Ca2+ ; Schlattner et al., 2001). SPR technology is increasingly being used to monitor immune responses either to an immunotherapeutic protein, vaccine, or even whole virus in research and preclinical environments (Alaedini and Latov, 2001; Abad et al., 2002; Swanson, 2003; Rini et al., 2005; Thorpe and Swanson, 2005). One simple assay requiring small sample volumes can provide information regarding antibody isotype, and active or relative concentration in serum. Another advantage of using Biacore for immunogenicity studies is the ability of the technique to detect both high- and lowaffinity antibodies, whereas traditional endpoint assays (e.g., ELISA) often fail to detect
fast-dissociating antibodies (Swanson, 2003; Thorpe and Swanson, 2005).
Concentration Analysis Most chemical and spectroscopic methods used to quantify proteins measure total protein content, do not distinguish active from inactive molecules, and cannot be used with unpurified samples. Since SPR is a noninvasive technology (i.e., no light penetrates the sample), it is possible to measure sub-femtomole amounts of analyte bound to the sensor chip surface from complex matrices such as food products, serum, and cell extracts, to name a few (Nelson et al., 2000). Instrument automation decreases operator involvement thereby leading to highly reproducible measurements. Various assay formats are possible. For analytes >5000 Da, a direct binding assay format can be used with the optional response enhancement from a secondary detecting molecule. Enhancement not only increases the dynamic range of the assay but can also improve assay specificity. An enhancement step can also be used to determine the isotype of antibodies in serum that are generated in response to a protein therapeutic or vaccine. Unlike many other immunoassays, concentration analysis with Biacore requires no separation and washing steps and, since binding responses are monitored continuously, it is possible to quantify fast-dissociating, low-affinity interactants. The point at which analyte concentration is measured can be chosen, giving flexibility in the assay design, which is not available with standard end-point assays. Inhibition or competition assay formats are well suited for quantification of low-molecular-weight molecules (<5000 Da). Inhibition assays rely on mixing the sample with a known concentration of a molecule that binds with high affinity, typically an antibody, then injecting the mixture over the target molecule immobilized on the sensor surface to determine the concentration of the high-molecular-weight binder in the mixture. Biacore evaluation software provides a direct readout of analyte concentration from a calibration curve generated using standard analyte concentrations. In a competition assay, the low-molecular-weight analyte competes with a fixed concentration of a high-molecular-weight molecule that shares the same binding site. Generally, concentration analysis is carried out at high ligand densities and slow flow rates. Efficient regeneration of the surface while maintaining ligand activity is paramount for successful concentration analysis measurements.
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Kinetics and Affinity Analysis
Overview of Biacore Systems
A hallmark of Biacore’s SPR biosensors is the ability to derive kinetic association and dissociation rate constants from real-time measurement of binding interactions, thereby providing valuable information regarding complex formation and complex stability that is not revealed through affinity measurements. Rate constants can provide a link between protein function and structure, e.g., through the evaluation of the impact of amino acid substitutions on an interaction. Affinity values can be derived either from interactions that have reached equilibrium or from the ratio of the dissociation and association rate constants in cases where the system does not reach steady state during the time frame of the experiment. The typical working range for affinity measurements with Biacore is picomolar to high micromotar for KD (M). Association rate constants can be measured ranging from 103 to 108 M−1 sec−1 and dissociation rate constants from 10−5 to 1 sec−1 (Karlsson, 1999). To measure rate constants accurately, proper experimental design is critical. As with all surface-based analysis methods, the phenomenon of mass transport should be considered (see Fig. 20.2.5 for mass transport effects on the surface binding process). For analyte molecules to bind to a ligand on the sensor surface, they must be transported from the bulk analyte solution to the surface. Under laminar flow conditions used in Biacore, the rate of transport of the analyte to the surface is proportional to the cube root of the flow rate, and is also influenced by the dimensions of the flow cell and the diffusion properties of the analyte. If the rate of transport of analyte to the surface is much faster than the rate of analyte association with the ligand, the observed binding will be determined by the kinetic rate constants. However, if mass transport is much slower than association, the binding interaction will be limited by the rate of analyte transport and there will be partial or no kinetic information in the binding data (mass transport limitation). Optimal assay conditions to minimize mass transport limitations to measure rate constants are a combination of high flow rates and low surface binding capacity. High flow rates minimize the diffusion distance from the bulk flow to the surface, while low ligand densities reduce analyte consumption in the surface layer. In practice, this translates to using ligand densities that result in a maximum analyte binding response no greater than 50 to 150 RU and flow rates >30 µl/min. It is important to note that mass
transport is a well-understood physical property of the system and partial mass transport limitations can be accounted for during data analysis (Myszka et al., 1998; Karlsson, 1999). Reliable detailed kinetic analysis requires data from four to six analyte concentrations, spanning the range of 0.1 to 10 times KD . Analyte concentrations must be accurately known to determine correct association rate constants. Analytes should be in the same buffer as the continuous flow buffer to minimize bulk refractive index differences that can lead to low signal-to-noise ratios. This is often most easily achieved through dilution of a concentrated analyte stock into running buffer. Samples containing high refractive index solutions, such as high salt, glycerol, or DMSO, should either be exchanged into the running buffer or the concentration of the high refractive index component should be matched precisely in the continuous flow running buffer. Kinetic assays should include a series of start-up cycles using buffer as analyte to equilibrate the surface as well as cycles with zero concentration of analytes as part of the concentration series for the purposes of double-referencing (Myszka, 1999) during data analysis. Although it is not necessary to reach equilibrium, it is recommended that the association times used be sufficient for at least one analyte concentration to reach steady state. To accurately determine dissociation rate constants, a measurable decrease in signal should occur during the dissociation period. If possible, kinetic experiments should be designed such that the data are described by the simplest interaction model. For example, in the case of an antibody-antigen interaction, the antibody should be immobilized or captured on the surface and the antigen used as analyte to avoid avidity effects resulting from the bivalent nature of the antibody. Avidity refers to the ability of an antibody to bind to two antigen molecules simultaneously, thus, the antibody may not dissociate from the antigen immobilized on the chip surface before binding another antigen molecule. Avidity effects will slow down the dissociation rate yielding enhanced affinity values compared to those measured from a 1:1 interaction. It is also important that both the ligand and analyte be as homogeneous as possible. Impurities from partially purified material can complicate the results by affecting the accurate determination of analyte concentration or introducing nonspecific binding. Lastly, analyte should be injected over both a reference surface and an active ligand surface. Reference surfaces are necessary to subtract bulk
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refractive index responses from the specific binding signal as well as to ensure that there is no nonspecific interaction with the sensor chip surface. Several excellent reviews on the topic provide detailed guidelines on experimental setup and interpretation of results (Karlsson and F¨alt, 1997; Myszka, 1999; Myszka, 2000; Rich and Myszka, 2001; Van Regenmortel, 2003; Karlsson and Larsson, 2004).
Thermodynamics By studying temperature dependence of rate and affinity constants it is possible to determine thermodynamic parameters for a binding interaction. Not only can the equilibrium values for changes in enthalpy (H) and entropy (S) associated with complex formation be determined, but transition state energetics can also be evaluated (Roos et al., 1998). The van’t Hoff equation relates affinity (KD ) to the free energy difference (G◦ ) between the associated and unassociated states of the interacting molecules where G◦ = RT ln KD , where R is the gas constant and T is the absolute temperature. Therefore, thermodynamic parameters can be determined by measuring KD at several temperatures and plotting ln KD against inverse temperature. H◦ and S◦ may then be calculated from the slope and y-axis intercept of the resulting straight line, respectively, since ln KD = H◦ /RT – S◦ /R. In some cases, entropy may be determined more accurately by subtracting H◦ from G◦ . This approach provides complementary information to thermodynamic parameters determined through microcalorimetry since in Biacore only the direct binding is measured, whereas microcalorimetry measures all components, including heats of mixing, dilution, and hydration effects. ln KD will be a linear function of (1/T) if both H and S are independent of temperature (T); however, H and S may not always be independent of temperature. In such cases, taking changes in heat capacity (Cp◦ ) into account and nonlinear fitting of the data using the integrated van’t Hoff equation may be used to determine thermodynamic parameters (Shuman et al., 2004; Dhalluin et al., 2005). The Eyring equation can be used with either the association rate constant (ka ) or the dissociation rate constant (kd ) to calculate the transition state energetics (H◦‡ and S◦‡ ) for a given complex. Thus, a plot of ln ka or ln kd against (1/T) will have a slope of –H◦‡ /R and an intercept of S◦‡ /R – ln(h/kB ), where h is the Planck constant and kB is the Boltzmann constant.
Biacore has been used to investigate the thermodynamics of proteins interacting with small molecules (Deinum et al., 2002; Day and Myszka, 2003; Shuman et al., 2004) and with other proteins (Zeder-Lutz et al., 1997; McFarland and Strong, 2003; Baerga-Ortiz et al., 2004; Dhalluin et al., 2005). Some benefits of using Biacore to determine thermodynamic parameters include: low sample concentration and consumption; affinities up to 1011 M−1 can be measured; there is no need to wait for equilibrium since KD can be determined from kd ka . Transition state analysis can provide additional mechanistic information about an interaction, revealing if the interaction is driven by entropic or enthalpic forces.
PROTEIN INTERACTION ANALYSIS This section highlights examples of studies where Biacore technology has been used to investigate interactions between proteins and a variety of biomolecules, whole cells, and viruses. This summary represents a small sample of the over 4000 peer-reviewed publications that have made use of Biacore for investigating many disciplines within the life sciences. A searchable database containing the abstracts of these publications is available on the Biacore Website (http://www.biacore. com/lifesciences/index.html).
Protein-Protein Interactions The largest number of publications citing Biacore involve characterization of proteinprotein interactions. An example of the versatility of Biacore in protein studies is illustrated in the work of Seet and co-workers (2003). In this study, the functionality and selectivity of a putative chemokine binding protein (CBP) from a poxvirus was assayed by immobilizing the recombinant protein and screening a panel of cytokines and chemokines. The affinity and kinetic rate constants for the interaction between the CBP and several chemokines were measured. Lastly, residues important for the interaction between the chemokine MCP1 and the CBP were identified by comparing the binding profiles of MCP-1 substitution mutants to that of the wild-type protein (Seet et al., 2003). Modifications of proteins, e.g., with biotin, fluorescent labels, or polyethyleneglycol, may introduce steric hindrance and alter their binding characteristics toward their interaction partners (Peter et al., 2003). This can
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be especially detrimental in the case of therapeutic proteins. Since detection in Biacore is based on mass changes at the sensor chip surface, it is often used to compare the binding of native and modified proteins. The kinetic rate and affinity constants for pegylated forms of interferonα 2a to the extracellular domain of a cell membrane receptor were compared to those of the unmodified protein (Dhalluin et al., 2005). It was found that increased degrees of pegylation resulted in lower affinities mainly due to reduction in association rate. In vivo, the decreased affinity was compensated for by an increased half-life of the protein due to increased protection from proteolytic degradation and a reduction in renal clearance. A recent development is the combination of SPR with mass spectrometry (SPR-MS). While Biacore systems provide information regarding the kinetics of an interaction and the analyte concentration, they do not permit direct identification of an unknown binding partner. Thus, the possibility of combining both technologies to provide protein identification on the basis of functional binding criteria is quite powerful. This method has been implemented to identify proteins captured from cytosolic extract (Lopez et al., 2003), cell lysates (Gilligan et al., 2002; Natsume et al., 2002), tissue extracts (Kikuchi et al., 2003; Zhukov et al., 2004), and plant extracts (Borch and Roepstorff, 2004). The small sample volume required and the high sensitivity of SPR results in a small amount of material that is recovered from a single SPR experiment for MS analysis. This can be addressed through the use of MS systems with high sensitivity, multiple binding and recovery cycles or by the use of an external flow cell with an expanded area available as part of the Biacore 3000 analyte recovery package. Biacore 3000 has been updated to recover bound analyte automatically in a 2-µl volume that can be deposited directly onto MALDI targets or delivered to a recovery vial for enzymatic digestion.
Protein Interactions with Peptides and Small Molecules
Overview of Biacore Systems
Improvements in instrument sensitivity, experimental protocols, and data analysis software have allowed for routine study of the affinity and kinetics of interactions between proteins and small analytes (<1000 Da) with Biacore A100, S51, T100, 3000, and 2000. Owing to their higher sensitivity and improved fluidic systems, Biacore A100, S51, and T100 are best suited for analysis of very low-molecular-weight analytes (>100 Da) in
direct binding assays. Not only have such characterizations provided valuable information regarding the interaction of drug candidates with protein targets, but also details on their interaction with circulating plasma proteins such as human serum albumin (HSA) and α1-acid glycoprotein (AGP) (Frostell-Karlsson et al., 2000; Day et al., 2002). In the case of enzymes, the data for a direct binding assay of a potential inhibitor to the enzyme may be easier to interpret than that from activity assays since interactions can be studied in the absence of substrate and without restricting the assay conditions to those best suited for catalysis of a specific substrate (Backman and Danielson, 2003). Different experimental conditions can be used to monitor their effect on the kinetics and affinity of binding interactions, e.g., the influence of pH or temperature. Binding interactions between small molecules or peptides and targets ranging from kinases (Casper et al., 2004; Nordin et al., 2005) to proteases (Karlsson et al., 2000; Backman and Danielson, 2003; Gossas and Danielson, 2003) and other enzymes (Boehm et al., 2000; Day et al., 2002), hormone receptors (Rich et al., 2002), antibiotics (Tseng and Chu, 2005), and proteins (Fisher et al., 2003; Huber, 2005; Yoshitani et al., 2005) have been characterized.
Protein Interactions with Lipids, Membranes, and Membrane-Bound Proteins The impact of Biacore technology on studies with lipids, membranes, and integral membrane proteins has been growing steadily since the introduction of Sensor Chips HPA and L1, which were designed for the creation of lipid monolayers and bilayers, respectively, on the sensor surface. In a well-designed study, de Haro et al. (2004) investigated the mechanism whereby the v-SNARE synaptobrevin, VAMP, interacts with lipids and Ca2+ / calmodulin as part of the fusion of secretory granules in the plasma membrane of neurosecretory cells. The binding of recombinant VAMP to a bilayer of acidic phopholipids created on the surface of Sensor Chip L1 was abrogated in the presence of Ca2+ / calmodulin or mutations in the calmodulin binding domain, suggesting that this region of the protein is involved in interactions with lipids and calmodulin. Antibodies raised against the putative lipid/calmodulin binding domain were able to bind to oriented VAMP proteoliposomes reconstituted with VAMP containing mutations in this domain but not to proteoliposomes reconstituted with wild-type VAMP.
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These results, together with those of control experiments, demonstrated that membraneanchored VAMP is able to undergo cis-lipid interactions with acidic lipid headgroups in the outer leaflet of the vesicle membrane that may be altered by Ca2+ / calmodulin (de Haro et al., 2004). An area currently in development is the design of experimental conditions to study interactions with G-protein-coupled receptors and integral membrane proteins. The concentration of these proteins within membranes may be too low to be able to detect interactions between low-molecular-weight analytes and immobilized membrane preparations. Enrichment of membrane proteins through appropriate detergent solubilization followed by immobilization on the sensor chip surface and by on-surface reconstitution of lipid bilayers is a promising approach that has been reported by several authors (Karlsson and L¨of˚as, 2002; Stenlund et al., 2003; Navratilova et al., 2005).
eric repeats was developed by Maesawa et al. (2003). Addition of nucleotides to the 3 end of biotinylated oligonucleotides captured on sensor chip SA by telomerase, present in extracts from cancer cell lines and tumor samples, results in a post-injection baseline increase due to the increased mass on the surface that is proportional to the amount of telomerase present in the extract (Maesawa et al., 2003). This Biacore assay has significant advantages over current PCR-based methods in that the direct on-surface measurement of elongation rates avoids post-PCR procedures and potential PCR artifacts. Additionally, the method is label-free and automated, with a one-step analysis that provides a rapid, reproducible readout with real-time monitoring of the process. In contrast to PCR-based methods, the telomeric repeat elongation assay enables quantitative evaluation of telomerase activity from a range of sources, including both cancer and normal somatic cells.
Protein Interactions with Nucleic Acids Interactions between proteins and nucleic acids play an extensive role in biological regulation. Although affinity data obtained under equilibrium conditions (such as electrophoretic mobility shift assays) is valuable in many instances, it fails to address the dynamic nature of protein-nucleic acid interactions. The most common approach utilized to study these interactions with Biacore is to analyze the interaction between proteins in solution and biotinylated nucleic acids captured on the streptavidin surface of sensor chip SA. DNA and RNA oligonucleotides can be synthesized with a terminal biotin group with relative ease, thus enabling the creation of a homogeneous oriented surface. Katsamba et al. (2002) used Biacore technology to elucidate the binding mechanisms of the neuronspecific RNA-binding protein, HuD, and the spliceosomal protein, U1A, with their target RNAs. A review of this work, which also provides recommendations for RNA-protein interactions, was recently published (Katsamba et al., 2002). Tsoi and Yang (2002) used Biacore to investigate the kinetics of interaction between DNA polymerase β and a series of DNA substrates. Results indicated that the polymerase-bound, single-stranded and fully matched DNA-primer templates with similar affinity, but with different rates of association and dissociation, while mismatches in the primer sequence, significantly lowered the affinity. A novel assay to monitor telomerase activity by detecting on-surface elongation of oligonucleotides containing tandem telom-
Protein Interactions with Whole Cells and Viruses On the high end of the analyte molecular weight spectrum are whole cells and viruses. A number of SPR assays to detect and identify bacteria in a wide range of contaminated samples have been developed as an alternative to conventional microbiological and biochemical methods that are much lengthier. These assays have shown very good reproducibility and equal or better sensitivity than ELISA and PCR (Bokken et al., 2003; Leonard et al., 2005). Bacterial cells and viruses, unlike eukaryotic cells, are generally robust enough to withstand immobilization and the shear forces within the microfluidic system (Dubs et al., 1991; Abad et al., 2002; Medina, 2004). By measuring the ability of various compounds to reduce the binding of cell membrane proteins to an immobilized Salmonella typhimurium surface, Medina developed an SPR-based method for the rapid evaluation of compounds that may prevent bacterial attachment to poultry skin and carcasses with a vision to develop treatments to reduce pathogen contamination of poultry foods. A number of publications have reported the use of Biacore to detect interactions between eukaryotic cells and proteins or peptides (Quinn et al., 1997; Aggarwal et al., 2005). For example, Aggarwal and co-workers designed peptides to efficiently and selectively capture metastatic epithelial cancer cells from flowing blood. More detailed kinetic information provided by Biacore analysis of phage-displayed
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antibodies binding to the target antigen has also proved to be useful in guiding the selection of the highest affinity antibodies (Schier and Marks, 1996). For more than 15 years, Biacore systems have been helping scientists unravel the intricacies of protein interactions. Herein, only a small portion of this exciting technology has been highlighted and the breadth of applications it addresses. The release of the new SPR array platforms (Flexchip and Biacore A100) will add new dimensions to future developments within label-free protein interaction analysis research.
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F¨agerstam, L.G., Frostell, A., Karlsson, R., Kullman, M., Larsson, A., Malmqvist, M., and Butt, H. 1990. Detection of antigen-antibody interactions by surface plasmon resonance. Application to epitope mapping. J. Mol. Recognit. 3:208-214. Fisher, R.D., Wang, B., Alam, S.L., Higginson, D.S., Robinson, H., Sundquist, W.I., and Hill, C.P. 2003. Structure and ubiquitin binding of the ubiquitin-interacting motif. J. Biol. Chem. 278:28976-28984. Frostell-Karlsson, A., Remaeus, A., Roos, H., Anderson, K., Borg, P., H¨am¨al¨ainen, M., and Karlsson, R. 2000. Biosensor analysis of the interaction between immobilized human serum albumin and drug compounds for prediction of human serum albumin binding levels. J. Med. Chem. 43:1986-1992.
Borch, J. and Roepstorff, P. 2004. Screening for enzyme inhibitors by surface plasmon resonance combined with mass spectrometry. Anal. Chem. 76:5243-5248.
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Huber W. 2005 A new strategy for improved secondary screening and lead optimization using high-resolution SPR characterization of compound-target interactions. J. Mol. Recognit. 18:273-281. Jokiranta, T.S., Hellwage, J., Koistinen, V., Zipfel, P.F., and Meri, S. 2000. Each of the three binding sites on complement factor H interacts with a distinct site on C3b. J. Biol. Chem. 275:2765727662. Karlsson, R. 1999. Affinity analysis of non-steadystate data obtained under mass transport limited conditions using Biacore technology. J. Mol. Recognit. 12:285-292. Karlsson, R. and F¨alt, A. 1997. Experimental design for kinetic analysis of protein protein interactions with surface plasmon resonance biosensors. J. Immunol. Methods 200:121-133. Karlsson, R., Kullman-Magnusson, M., H¨am¨al¨ainen, M.D., Remaeus, A., Andersson, K., Borg, P., Gyzander, E., and Deinum, J. 2000. Biosensor analysis of drug-target interactions: Direct and competitive binding assays for investigation of interactions between thrombin and thrombin inhibitors. Anal. Biochem. 278:1-13. Karlsson, R. and Larsson A. 2004. Affinity measurement using surface plasmon resonance. Methods Mol. Biol. 248:389-415. Karlsson, O.P. and L¨of˚as, S. 2002. Flow-mediated on-surface reconstitution of G-protein coupled receptors for applications in surface plasmon resonance biosensors. Anal. Biochem. 300:132138. Katsamba, P.S., Park, S., and Laird-Offringa, I.A. 2002. Kinetic studies of RNA-protein interactions using surface plasmon resonance. Methods 26:95-104. Kikuchi, J., Furukawa, Y., and Hayashi, N. 2003. Identification of novel p53-binding proteins by biomolecular interaction analysis combined with tandem mass spectrometry. Mol. Biotechnol. 23:203-212. Lieberg, B., Nylander, C., and Lundstr¨om, I. 1983. Surface plasmon resonance for gas detection and biosensing. Sensors Actuators 4:299-304. Leonard, P., Hearty, S., Wyatt, G., Quinn, J., and O’Kennedy, R. 2005. Development of a surface plasmon resonance-based immunoassay for Listeria monocytogenes. J. Food Prot. 68:728735. Lopez, F., Pichereaux, C., Burlet-Schiltz, O., Pradayrol, L., Monsarrat, B., and Esteve, J.P. 2003. Improved sensitivity of biomolecular interaction analysis mass spectrometry for the identification of interacting molecules. Proteomics 3:402-412. Maesawa, C., Inaba, T., Sato, H., Iijima, S., Ishida, K., Terashima, M., Sato, R., Suzuki, M., Yashima, A., Ogasawara, S., Oikawa, H., Sato, N., Saito, K., and Masuda, T. 2003. A rapid biosensor chip assay for measuring of telomerase activity using surface plasmon resonance. Nucl. Acids Res. 31:E4-4.
McFarland, B.J. and Strong, R.K. 2003. Thermodynamic analysis of degenerate recognition by the NKG2D immunoreceptor: Not induced fit but rigid adaptation. Immunity 19:803-812. Medina, M.B. 2004. Binding interaction studies of the immobilized Salmonella typhimurium with extracellular matrix and muscle proteins, and polysaccharides. Int. J. Food Microbiol. 93:6372. Myszka, D.G. 1999. Improving biosensor analysis. J. Mol. Recognit. 12:1-6. Myszka, D.G. 2000. Kinetic, equilibrium, and thermodynamic analysis of macromolecular interactions with Biacore. Methods Enzymol. 323:325340. Myszka, D.G., He, X., Dembo, M., and Morton, T.A. 1998. Extending the range of rate constants available from Biacore: Interpreting mass transport-influenced binding data. Biophys. J. 75:583-594. Natsume, T., Taoka, M., Manki, H., Kume, S., Isobe, T., and Mikoshiba, K. 2002. Rapid analysis of protein interactions: On-chip micropurification of recombinant protein expressed in Esherichia coli. Proteomics 2:12471253. Navratilova, I., Sodroski, J., and Myszka, D.G. 2005. Solubilization, stabilization, and purification of chemokine receptors using biosensor technology. Anal. Biochem. 339:71-81. Nelson, R.W., Nedelkov, D., and Tubbs, K.A. 2000. Biomolecular interaction analysis mass spectometry. BIA/MS can detect and characterize protiens in complex biological fluids at the lowto subfemtomole level. Anal. Chem. 72:404A411A. Nordin, H., Jungnelius, M., Karlsson, R., and Karlsson, O.P. 2005. Kinetic studies of small molecule interactions with protein kinases using biosensor technology. Anal. Biochem. 340:359368. Peter, J.C., Briand, J.P., and Hoebeke, J. 2003. How biotinylation can interfere with recognition: A surface plasmon resonance study of peptideantibody interactions. J. Immunol. Methods 274:149-158. Quinn, J.G., O’Kennedy, R., Smyth, M., Moulds, J., and Frame, T. 1997. Detection of blood group antigens utilising immobilised antibodies and surface plasmon resonance. J. Immunol. Methods 206:87-96. Rich, R.L., Hoth, L.R., Geoghegan, K.F., Brown, T.A., LeMotte, P.K., Simons, S.P., Hensley, P., and Myszka, D.G. 2002. Kinetic analysis of estrogen receptor/ligand interactions. Proc. Natl. Acad. Sci. U.S.A. 99:8562-8567. Rich, R.L. and Myszka, D.G. 2001. Survey of the year 2000 commercial optical biosensor literature. J. Mol. Recognit. 14:273-294. Rini, B., Wadhwa, M., Bird, C., Small, E., GainesDas, R., and Thorpe, R. 2005. Kinetics of development and characteristics of antibodies induced in cancer patients against yeast expressed
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rDNA derived granulocyte macrophage colony stimulating factor (GM-CSF). Cytokine 29:5666. Roos, H., Karlsson, R., Nilshans, H., and Persson, A. 1998. Thermodynamic analysis of protein interactions with biosensor technology. J. Mol. Recognit. 11:204-210. Schier, R. and Marks, J.D. 1996. Efficient in vitro affinity maturation of phage antibodies using Biacore guided selections. Hum. Antibody Hybridomas 7:97-105. Schlattner, U., Dolder, M., Wallimann, T., and Tokarska-Schlattner, M. 2001. Mitochondrial creatine kinase and mitochondrial outer membrane porin show a direct interaction that is modulated by calcium. J. Biol. Chem. 276:4802748030. Schuster, S.C., Swanson, R.V., Alex, L.A., Bourret, R.B., and Simon, M.I. 1993. Assembly and function of a quaternary signal transduction complex monitored by surface plasmon resonance. Nature 365:343-346. Seet, B.T., McCaughan, C.A., Handel, T.M., Mercer, A., Brunetti, C., McFadden, G., and Fleming, S.B. 2003. Analysis of an orf virus chemokine-binding protein: Shifting ligand specificities among a family of poxvirus viroceptors. Proc. Natl. Acad. Sci. U.S.A 100:1513715142. Shuman, F.S., H¨am¨al¨ainen, M.D., and Danielson, U.H. 2004. Kinetic and thermodynamic characterization of HIV-1 protease inhibitors. J. Mol. Recognit. 17:106-119. Stenlund, P., Babcock, G.J., Sodroski, J., and Myszka, D.G. 2003. Capture and reconstitution of G protein–coupled receptors on a biosensor surface. Anal. Biochem. 316:243-250. Swanson, S.J. 2003. New technologies for the detection of antibodies to therapeutic proteins. Dev Biol (Basel). 112:127-133. Thai, C.T. and Ogata, R.T. 2005. Recombinant C345C and factor I modules of complement components C5 and C7 inhibit C7 incorporation into the complement membrane attack complex. J. Immunol. 174:6227-6232. Thorpe, R. and Swanson, S.J. 2005. Assays for detecting and diagnosing antibody-mediated pure red cell aplasia (PRCA): An assessment of available procedures. Nephrol. Dial. Transplant. 20(Suppl 4):iv16-22. Tseng, M.C. and Chu, Y.H. 2005. Using surface plasmon resonance to directly identify molecules in a tripeptide library that bind tightly to a vancomycin chip. Anal. Biochem. 336:172177. Tsoi, P.Y. and Yang, M. 2002. Kinetic study of various binding modes between human DNA polymerase beta and different DNA substrates by surface-plasmon-resonance biosensor. Biochem. J. 361:317-325.
Overview of Biacore Systems
Yoshitani, N., Satou, K., Saito, K., Suzuki, S., Hatanaka, H., Seki, M., Shinozaki, K., Hirota, H., and Yokoyama, S. 2005. A structure-based strategy for discovery of small ligands binding to functionally unknown proteins: Combination of in silico screening and surface plasmon resonance measurements. Proteomics 5:1472-1480. Young, P.J., Day, P.M., Zhou, J., Androphy, E.J., Morris, G.E., and Lorson, C.L. 2002. A direct interaction between the survival motor neuron protein and p53 and its relationship to spinal muscular atrophy. J. Biol. Chem. 277:2852-2859. Zeder-Lutz, G., Zuber, E., Witz, J., and Van Regenmortel, M.H.V. 1997. Thermodynamic analysis of antigen-antibody binding using biosensor measurements at different temperatures. Anal. Biochem. 246:123-132. Zhukov, A., Schurenberg, M., Jansson, O., Areskoug, D., and Buijs, J. 2004. Integration of surface plasmon resonance with mass spectrometry: Automated ligand fishing and sample preparation for MALDI MS Using a Biacore 3000 biosensor. J. Biomol. Tech. 15:112-119.
KEY REFERENCES Dhalluin et al., 2005. See above. Excellent example of the importance of kinetic and thermodynamic information in the development of a biotherapeutic. Karlsson and Larsson. 2004. See above. A practical guide to proper experimental design when using Biacore for affinity measurements. Nordin et al., 2005. See above. Excellent example of the kinetic characterization of small molecule inhibitors of a protein kinase. Shuman et al., 2004. See above. Excellent example of kinetic and thermodynamic characterization of small molecule inhibitors of a protease.
INTERNET RESOURCES http://www.biacore.com/lifesciences/index.html Details and animations of SPR technology, Biacore instruments, application notes, a reference database, guidelines to common immobilization chemistries, an immobilization and regeneration database, self-training tools, and course listings are available. http://www.piercenet.com Maleimide coupling reagents are available from Pierce Chemical.
Contributed by Laure Jason-Moller, Michael Murphy, and JoAnne Bruno Biacore, Inc. Piscataway, New Jersey
Van Regenmortel, M.H. 2003. Improving the quality of Biacore-based affinity measurements. Dev Biol (Basel) 112:141-151.
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Using Biacore to Measure the Binding Kinetics of an Antibody-Antigen Interaction
UNIT 19.14
The optical phenomenon of surface plasmon resonance (SPR) used by Biacore systems enables the detection and measurement of protein-protein interactions in real time, without the use of labels. Biacore systems are widely used for characterizing the interactions of proteins with other proteins, peptides, nucleic acids, lipids, and small molecules. Over 4000 references using Biacore to characterize protein interactions can be found in the literature. When Biacore is used to measure protein interactions, one of the interactants is immobilized onto a sensor chip surface and the other interactant is passed over that surface in solution via an integrated microfluidic flow system. The immobilized interactant is referred to as the ligand, and the injected interactant in solution is referred to as the analyte. Binding responses are measured in resonance units (RU) and are proportional to the molecular mass on the sensor chip surface and, consequently, to the number of molecules on the surface. A sensorgram is a plot of the binding response in resonance units versus time in seconds, which is displayed and recorded as a change in mass occurs on the sensor chip surface (Fig. 19.14.1). The sensorgram provides essentially two kinds of information that are relevant to different types of applications: (1) the rate of interaction (association, dissociation, or both), which provides information on kinetic rate constants and analyte concentration; and (2) the binding level, which provides information on affinity constants and can be used for qualitative or semi-quantitative applications.
Figure 19.14.1 The sensorgram is a plot of response in resonance units (RU) versus time in seconds, which is presented continuously in real time. Upon injection of an analyte, if a binding interaction occurs, then an increase in mass occurs on the sensor chip surface and association is measured. At the end of the analyte injection, as the complex decays, a decrease in mass occurs and dissociation is measured. Association and dissociation are measured as changes in response. Contributed by Michael Murphy, Laure Jason-Moller, and JoAnne Bruno Current Protocols in Protein Science (2006) 19.14.1-19.14.17 C 2006 by John Wiley & Sons, Inc. Copyright
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In this unit, the application of Biacore technology to measuring a protein-protein interaction is described using an antibody and its antigen as an example. The affinity of the antibody for its antigen is determined by measuring the binding kinetics of the interaction. The protocols are divided into three major steps that are required for measuring binding kinetics using Biacore: (1) surface preparation (see Basic Protocols 1 and 2); (2) assay development (see Basic Protocol 3); and (3) kinetic analysis (see Basic Protocol 4). Note that the materials used in these protocols are available from Biacore and that the protocols are intended for use with Biacore 3000, Biacore 2000, or Biacore T100 systems. The principles described here apply to use of other Biacore systems, however, the software interface and system configuration may differ from the provided protocols. BASIC PROTOCOL 1
IDENTIFICATION OF OPTIMAL IMMOBILIZATION pH (pH SCOUTING) For most chemical coupling methods, a mechanism for attraction of the ligand to the carboxymethylated dextran surface matrix on the sensor chip is important for the efficient immobilization of proteins. The mechanism employed is an electrostatic attraction between negative charges on the carboxymethylated dextran matrix and positive charges on the protein ligand, which is referred to as preconcentration. The preconcentration strategy enables efficient immobilization from relatively dilute solutions of ligand (10 to 100 µg/ml). The protein ligand is diluted into a low-ionic-strength immobilization buffer at a pH below its isoelectric point (pI) to generate a net positive charge on the protein. The carboxymethylated dextran matrix carries a net negative charge at pH values >3.5. The diluted ligand is injected over the sensor chip surface where it will accumulate if positively charged and, in the next step (see Basic Protocol 2), undergo chemical coupling to the carboxymethylated dextran. The goal of immobilization pH scouting is to identify an appropriate pH for the immobilization buffer to establish a set of controlled conditions that provide usable and reproducible immobilization levels so that both sample and sensor chip consumption is minimized.
Materials 200 ml HBS-P+ (see recipe) 1 mg/ml anti-β2µ-globulin antibody (Biacore) 10 mM sodium acetate, pH 4.0, 4.5, 5.0, and 5.5 50 mM NaOH Biacore 3000, 2000, or T100 system Sensor Chip CM5 0.8- or 1.5-ml polypropylene tubes with rubber caps (Biacore) Set up instrument 1. Undock the maintenance chip or previously used sensor chip and remove from the instrument by clicking on the Command tab and then selecting the Undock command. When undocked, the sensor chip should easily slide out of the instrument. Note that the sensor chip indicator light on the front of the instrument is lit steadily when a chip is docked, flashing when a chip is inserted but undocked, and unlit when no chip is inserted.
2. Insert and dock the new Sensor Chip CM5. Click the Command tab and then select the Dock command. 3. Insert the appropriate instrument buffer tubing into a container containing 200 ml HBS-P+. Biacore to Measure Binding Kinetics
The buffer should be filtered (using a 0.22-µm filter) and degassed prior to use.
4. Under the Tools tab in the control software, choose Working Tools and select Prime to exchange all of the solutions in the system with fresh HBS-P+ buffer.
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5. Keep the instrument in Standby (or Continue) mode when not in use. Under the Tools tab in the control software, choose Start Standby. The Standby mode maintains a slow flow rate for up to 4 days. When not performing experiments, the instrument should be kept in Standby using water with the maintenance chip docked.
Set up sample and method 6. Dilute 1 mg/ml anti-β2µ-globulin antibody to 10 µg/ml in each of the four immobilization buffers using 0.8- or 1.5-ml tubes and cap the tubes with rubber caps. Add 2 µl of 1 mg/ml anti-β2µ-globulin to 198 µl of each of the following: 10 mM sodium acetate, pH 4.0 10 mM sodium acetate, pH 4.5 10 mM sodium acetate, pH 5.0 10 mM sodium acetate, pH 5.5 7. In the instrument control software under the File tab, open the Application Wizards, select Surface Preparation, and then choose Immobilization pH Scouting. 8. Enter the names of the solutions to be used for pH scouting by selecting from the list in the wizard. Select or enter:
acetate pH 4.0 acetate pH 4.5 acetate pH 5.0 acetate pH 5.5 If the needed solution is not already a selection in the list, then additional solutions can be added.
9. Select flow path 2. 10. Provide a name for the ligand (e.g., antiB2micro), and enter values for the injection parameters: 120 sec contact time (injection) and 10 µl/min flow rate. 11. Enter 50 mM NaOH for the wash solution that is used to remove the ligand from the surface after the pH scouting experiment. Enter values for the injection parameters for the wash: 30 sec contact time and 50 µl/min flow rate. Dispense 100 µl of 50 mM NaOH into a 1.5-ml tube. 12. Next, a diagram of the reagent rack is shown and colored spots in various positions indicate samples (the samples will include the tubes of diluted ligand and one tube of NaOH). Move the samples on the diagram via clicking, dragging, and dropping or by entering the position in the table. Place the solutions in the appropriate reagent rack, taking care to match the sample content with the indicated rack positions. 13. Save the wizard template and the result file, and start the method.
Evaluate pH scouting results 14. When the method has finished, view the overlay plot of the sensorgrams for each immobilization buffer, presented automatically by the control software (or the evaluation software, if using T100). If the ligand accumulates on the sensor chip surface, then a response is generated due to the increased mass on the surface. The magnitude and slope of the preconcentration response provide information about the ligand density that can be achieved. Compare the preconcentration response for each immobilization buffer and identify the mildest pH solution that will achieve the targeted level for immobilization (1200 RU, described also in Critical Parameters).
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Figure 19.14.2 The effect of pH on preconcentration of anti-β2µ-globulin antibody onto the surface of Sensor Chip CM5. Electrostatic attraction of anti-β2µ-globulin antibody to the chip surface occurs for each pH tested, therefore, any pH solution that was tested can be used. The pH 5.5 sodium acetate solution was chosen because amine coupling occurs more efficiently at a higher pH.
Although the preconcentration levels were higher and achieved more quickly for pH 4.0 and pH 4.5, the preconcentration levels for pH 5.0 and pH 5.5 were acceptable. Based on the pH scouting results, the pH 5.5 solution is optimal for preconcentration of the anti-β2µ-globulin antibody (Fig. 19.14.2). BASIC PROTOCOL 2
IMMOBILIZATION OF LIGAND TO SENSOR CHIP USING AMINE COUPLING Several coupling chemistries are available for covalent immobilization of the ligand to the sensor chip surface, such as amine, aldehyde, and thiol (see UNIT 19.13). Amine coupling is the most widely applicable approach for immobilization of ligands. There are three steps in amine coupling: (1) activation of the surface, (2) contact of the ligand with the activated surface, and (3) blocking of unreacted sites. After identification of the optimal pH for preconcentration of the ligand (see Basic Protocol 1), the active and reference surfaces are prepared on Sensor Chip CM5 using, in this example, amine-coupling chemistry. The immobilization surface preparation application wizard is used to create active and reference surfaces automatically. The active surface will contain the anti-β2µ-globulin antibody ligand, and the reference surface will contain no ligand, but is activated and blocked with the same chemistry as the active surface. Generally, 75% surface activity can be expected for ligands immobilized directly by amine coupling. Note that when using the Aim for Immobilized Level option in the application wizard, the control software performs an injection of ligand to test for preconcentration prior to activating the surface to ensure that the desired immobilization level can be achieved during amine coupling. If the preconcentration response is too low to achieve the target immobilization level, the run will be stopped.
Materials
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100 mM N-hydroxysuccinimide (NHS) 400 mM N-ethyl-N-(3-dimethylaminopropyl)-carbodiimide hydrochloride (EDC) 10 µg/ml anti-β2µ-globulin antibody in 10 mM sodium acetate, pH 5.5 1 M ethanolamine 50 mM NaOH Current Protocols in Protein Science
Biacore 3000, 2000, or T100 system 0.8-ml tubes Sensor chip CM5 Set up sample and method 1. Under the File tab in the instrument control software, open the Application Wizards, choose Surface Preparation and then select Immobilization. Enter CM5 for Sensor Chip and Amine Coupling for the Immobilization Method. 2. Select Aim for Immobilized Level in the immobilization procedure or set-up window. 3. Select flow cell 1 as blank. 4. Enter the ligand name (e.g., antiB2micro) for flow cell 2 and enter a target level of 1200 RU. Target levels for ligand immobilization must be calculated using the rearranged Rmax equation, which is described in Critical Parameters under Immobilization levels.
5. Enter 50 mM NaOH for the wash solution. 6. Click on Next and the wizard will recommend solution volumes and vial positions. Dispense the indicated volume of each solution into 0.8-ml tubes. Place the solutions in the appropriate reagent rack positions taking care to match the sample content with the indicated rack positions. Check the sample positions and edit if necessary. 7. Save the immobilization wizard template and result files and start the immobilization.
Evaluate immobilization results 8. When the method has finished, view the table summarizing the results of the immobilization, which is presented automatically by the control software. Note the surface
Figure 19.14.3 Immobilization of anti-β2µ-globulin antibody to the chip surface. Each step and corresponding response is labeled at the top of the sensorgram. Each X on the response represents a report point, which provides a value of the response (averaged over a 5-sec window) at a given point. Report point values are assigned a name (such as baseline or binding level) and are recorded in a table. During the preconcentration test, the ligand is injected over the unmodified sensor chip surface to determine if the desired immobilization level can be achieved. During the activation step, the carboxymethyl groups on the dextran are converted into reactive esters, and then the ligand is injected in short pulses until the desired immobilization level is achieved. During the blocking step, uncrosslinked reactive esters are blocked with ethanolamine. The immobilization level or ligand density is the final response difference from baseline, denoted by RL .
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density (RL ), which is the amount of immobilized ligand given by the final response in resonance units (RU). View the sensorgrams from each flow cell. The sensorgram from the reference flow cell (Fc 1) shows a bulk refractive index response for the injection of EDC/NHS and ethanolamine. The sensorgram from the active flow cell (Fc 2) shows the preconcentration injection and wash, bulk refractive index change for injection of EDC/NHS, the response from the injection of ligand, and bulk refractive index change for ethanolamine (Fig. 19.14.3). BASIC PROTOCOL 3
REGENERATION SCOUTING: IDENTIFYING A SUITABLE BUFFER FOR SURFACE REGENERATION After the active and reference surfaces have been created (see Basic Protocols 1 and 2), the binding of analyte is tested using a concentration of analyte well above the equilibrium dissociation constant (KD ) to ensure that the interaction will be observed if it occurs. If the interaction is characterized by a slow dissociation rate (<10−3 /sec), then surface regeneration is necessary to remove the bound analyte for subsequent interaction analysis within a reasonable time frame. The procedure for this is described here. The aim of regeneration scouting is to identify a solution that removes the analyte from the immobilized ligand without affecting the ligand activity. In general, start with a mild reagent (e.g., buffered solutions of low or high pH, high-salt solutions) and then test stronger reagents (e.g., HCl, NaOH). Inject regeneration buffers at high flow rates for short contact times (≤1 min). When regeneration scouting, follow the regeneration pulse with another injection of the same concentration of analyte to confirm surface activity.
Materials 32 nM β2µ-globulin in HBS-P+ (see recipe for HBS-P+) 10 mM glycine, pH 1.5, 2.0, 2.5, and 3.0 Biacore 3000, 2000, or T100 system 1. In the instrument control software under the Run tab, select Run Sensorgram. 2. Choose Multi Detection, Fc1,2; 2-1. Fc 1 is the reference surface and Fc 2 is the active surface.
3. Set flow rate to 10 µl/min. 4. Inject 32 nM β2µ-globulin over Fc 1,2 for 2 min using the Inject command under the Command tab. 5. Use the reference line to insert a baseline report point ∼15 sec before the injection start, and measure the amount bound by placing a report point 15 sec before the injection end. 6. Under the Command tab, select Flow and change the flow rate to 50 µl/min. 7. Test the different 10 mM glycine regeneration buffers working from high to low pH (gentle to harsh) using the Quickinject command (include an extra cleanup). Place a report point 15 sec after the end of the regeneration injection to measure the amount of analyte still bound to the surface. 8. After each regeneration step, change the flow rate back to 10 µl/min and reinject the 32 nM β2µ-globulin to assess ligand-binding activity. Insert report points, as done in steps 5 and 7 and use these values to check the effectiveness of the regeneration. Biacore to Measure Binding Kinetics
9. When a suitable regeneration buffer has been identified, stop and save the sensorgram.
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SURFACE PERFORMANCE TEST TO ASSESS LIGAND STABILITY UNDER REGENERATION CONDITIONS
BASIC PROTOCOL 4
Once a regeneration buffer has been identified (see Basic Protocol 3), the stability of the ligand under that condition should be tested for several binding and regeneration cycles. The surface performance test is used to determine if the regeneration solution will remove the bound analyte while retaining the binding activity of the ligand.
Materials 32 nM β2µ-globulin 10 mM glycine, pH 2.5 Biacore 3000, 2000, or T100 system Set up sample and method 1. Under the File tab in the instrument control software, open the Application Wizards, choose Surface Preparation, and then select Surface Performance Test. 2. Enter the analyte name (e.g., 32 nM B2micro), and enter values for the injection parameters: 2 min for the injection time and 10 µl/min for the flow rate. 3. Run the test for 10 cycles over flow cell 2, using flow cell 1 as a reference. 4. Select Single Injection, and enter 10 mM glycine, pH 2.5, for the solution. 5. Enter values for the injection parameters: 30 sec for the injection time and 50 µl/min for the flow rate. 6. Include a 1-min stabilization time after the regeneration injection. 7. Place the solutions in the appropriate reagent rack positions matching the sample content and recommended volumes with the indicated rack positions. 8. Provide a name for the surface performance test wizard template and result files and start the run.
Evaluate surface performance results 9. View the results of the Surface Performance Test that the control software presents. Plots of the baseline level and the analyte response level for each cycle are shown
Figure 19.14.4 Results of the surface performance tests of four regeneration solutions. The responses for the baseline (solid diamonds) and for the binding response (open triangles) are plotted for each cycle. Cycle 2 corresponds to the initial baseline and binding response. The baseline and binding responses using glycine pH 3.0 (cycles 3 to 7), glycine pH 2.5 (cycles 8 to 12), glycine pH 2.0 (cycles 13 to 17), and glycine pH 1.5 (cycles 18 to 22).
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for each flow cell. Examine the plot from the active flow cell. The responses for the baseline and ligand-binding response may require several cycles before stabilizing. For illustrative purposes, the results of surface performance tests using four solutions of glycine with decreasing pH are shown in Figure 19.14.4. Ideal regeneration conditions will exhibit baseline and analyte responses that are consistent for each cycle. For glycine, pH 3.0, the β2µ-globulin response decreases as the baseline increases, which suggests that this regeneration condition is too mild. For glycine, pH 2.5, the β2µ-globulin response and the baseline are stable after the second cycle, which suggests that this regeneration condition is good and should be used. For glycine, pH 2.0, the baseline response drops initially, but the β2µ-globulin response is stable, which suggests that the regeneration condition could be used. For glycine, pH 1.5, the analyte response decreases and the baseline remains constant, which suggests that the regeneration condition is too harsh and should be avoided to prevent damaging the ligand. BASIC PROTOCOL 5
KINETIC ANALYSIS FOR LIGAND IMMOBILIZED DIRECTLY TO SURFACE Once the optimal regeneration solution has been identified and assessed in a surface performance test, kinetic analysis of the antibody-antigen interaction is performed. A dilution series of the antigen (analyte) is prepared in running buffer. Each concentration of analyte will be injected over the surface and the binding response will be measured. The bound analyte will be removed from the ligand using regeneration. This process will be repeated for each concentration of analyte in a separate cycle, i.e., the binding response for each concentration is recorded as a new sensorgram. The binding response from each sensorgram will be overlaid and curve fitting of the entire concentration series (global fitting) will be used to determine the kinetic rate constants and affinity of the interaction. Additionally, a mass transfer control test can be performed as part of the Kinetic Analysis wizard (B2000, B3000) or as an independent experiment (T100). The mass transfer control test will measure the binding response of a single concentration of analyte injected sequentially at three flow rates (5, 15, and 75 µl/min). Ideally, the magnitude of the binding responses will not be influenced by the flow rate. If the binding responses are higher at faster flow rates, then mass transfer limitations exist for the interaction (described in Critical Parameters and UNIT 19.13).
Materials 100 µg/ml (8.5 µM) β2µ-globulin HBS-P+ (see recipe) 10 mM glycine, pH 2.5 Biacore 3000, 2000, or T100 system Set up sample and method 1. Dilute 5.6 µl of 8.5 µM stock β2µ-globulin (100 µg/ml) into 1494.4 µl HBS-P+ to prepare the 32 nM sample. 2. Prepare two-fold serial dilutions of 32 nM β2µ-globulin down to 2 nM. Also include a buffer-only sample (0 nM) that will be used for double-reference subtraction during data analysis. 3. In the instrument control software Application Wizards, choose Kinetic Analysis, check the box for Concentration Series, and include the control for Mass Transfer, and then click Next. Choose Direct Binding and then click Next. Biacore to Measure Binding Kinetics
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4. Enter values for the analyte injection parameters: flow path 2, using 1 as a reference, 30 µl/min for the flow rate, 2 min for analyte injection time, and 5 min for dissociation time. Current Protocols in Protein Science
5. Enter the analyte names (e.g., 32n B2micro, 16n B2micro, etc.), molecular mass (11,800 Da), and number of replicates (enter 2 for one concentration in the series), and each concentration for β2µ-globulin in the series. Run at least one analyte sample and the buffer-only sample in duplicate. 6. Choose the Run Order as Random or Sorted low to high. 7. Enter the concentration 2 nM for the Mass Transfer control test. 8. Provide the regeneration solution name (Glycine pH 2.5) and enter values for the regeneration injection parameters: 50 µl/min for the flow rate, 30 sec for injection time, and 1 min stabilization time. 9. Place the solutions in the appropriate reagent rack positions, matching the sample content and recommended volumes with the indicated rack positions. The 2 nM sample requires more volume for the mass transfer test.
10. Save the kinetic wizard template and result files and start the kinetic analysis. When the kinetic analysis is finished, the application wizard will present a results window automatically (only in Biacore 2000 or 3000). The kinetic rate constants that are presented by the application wizard in the result window should not be reported, instead curve fitting must be performed using the BIAevaluation software as described below. Tabs that are found in the application wizard results window should be used to view the baseline stability for the active and reference surfaces, and result of the mass transfer control. A mass transport limitation exists for the interaction if the binding response rate increases in a flow rate–dependent manner. Ideally, the binding response should not be dependent on flow rate. See Critical Parameters for more information.
Evaluate data from kinetic analysis 11. Open the BIAevaluation software from the start menu, BIA programs. 12. Open the Result file (.blr) that was saved for the Kinetic Analysis wizard. 13. Select Curves and Report Point Table, and then select the reference-subtracted (Fc2 – Fc1) curves that will be used in the kinetic analysis evaluation. The curves can be sorted by the keyword Fc and the value 2-1, and selected or deselected by holding down the Ctrl button and clicking on the curve.
14. Create an overlay plot of the curves for each concentration for β2µ-globulin by choosing View, and select Plot Overlay. The last three cycles represent the curves from the mass transfer control are should not be included.
15. Next, begin curve fitting by choosing Fit, and select Kinetics Simultaneous ka/kd to start the kinetic evaluation wizard, which will guide the user through the fitting procedure. 16. Remove unnecessary data, such as a regeneration injection. Zoom in on the area of interest by dragging a box around the data with the left mouse button held. Select the unwanted data by dragging a box around the data with the right mouse button. Click on Cut in the wizard panel to remove the data. 17. Adjust the baseline to zero. Zoom in on the baseline prior to the analyte injection and select the region to be used as the baseline using the right mouse button, then click on Y-transform in the wizard panel. Choose Zero at Average of Selection, make sure that Apply to Entire Working Set is checked, and choose Replace Original. 18. Adjust the curves by blank-run subtraction or double-referencing (i.e., subtracting the buffer-only curve for each of the other curves). Click on Y-transform in the wizard panel, and choose Curve-Curve. Select the 0 nM analyte curve from the drop-down
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menu. To recall which curve is the 0 nM, cancel the Y-transform and Fit Kinetics windows and under View, choose Curve History for review, then re-open Kinetics Simultaneous ka /kd under Fit to perform the blank-run subtraction. Click Next. 19. Set the start and stop times for the injection and select the data range for the fitting for all of the curves in the data set at once (see Fig. 19.14.1). Use the black markers in the top bar to mark the start and end of the analyte injection. Use the grey markers in the top bar to select the portion of the curves that correspond with the analyte injection (association) and after the end of the injection (dissociation). Click on Next once the selections have been made. 20. Select 1:1 (Langmuir) Binding with mass transfer from the pull-down menu. Click on Fit to initiate the curve fitting (Fig. 19.14.5). 21. View the Report and examine the values for ka , kd , kt , KA , KD , Rmax , and chi2 . Assess the fit quality by viewing a plot of the relative Residuals. Assess the fit quality by clicking on Parameters. Note the standard error (SE). Curve fitting is an iterative, mathematical procedure to determine parameter values that best describe the experimental responses. Parameter values are determined by least square fitting of residuals using the Marquardt-Levenberg algorithm. Residuals are the differences between the experimental and fitted curves. The best fit of the data is defined as the minimum sum of the squared residuals, and is referred to as chi2 . Thus, the value of chi2 describes the goodness of fit and should ideally be a value that is <1% of Rmax . The association rate constant (ka ) has a unit of M−1 sec−1 , and the dissociation rate constant (kd ) has a unit of sec−1 . The affinity of an interaction is often described by the equilibrium dissociation rate constant (KD ), which has a unit of M and equals kd /ka . The Rmax value is the experimental maximum binding capacity of the immobilized ligand. The mass transfer rate constant (kt ) describes the degree of mass transport limitations for the interaction. Values for kt <109 indicate partial mass transport limitations, which can be adjusted for mathematically. The standard error for each parameter indicates the significance of that parameter. Standard error values that are 1000-fold less than the parameter suggest that there is high confidence in the value of that parameter. Association rate constants ranging from 103 to 107 M−1 sec−1 and dissociation rate constants from 10−5 to 1 sec−1 can typically be measured accurately. Note that the curve fitting procedure for T100 is mostly automated.
Biacore to Measure Binding Kinetics
Figure 19.14.5 Representative sensorgram of fitted kinetic data from the kinetic analysis of β2µglobulin binding to anti-β2µ-globulin antibody that has been directly immobilized to Sensor Chip CM5 using amine coupling as described in the text. The solid black line that is super-imposed on the responses is the fitted curve. The analyte concentrations from the top curve are 32 nM, 16 nM, 8 nM, 4 nM, and 2 nM. The 8 nM β2µ-globulin sample (middle curve) was run in duplicate. The values of the calculated parameters for the kinetic analysis are listed in the text.
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IMMOBILIZATION OF LIGAND TO SENSOR CHIP USING LIGAND CAPTURE METHOD
ALTERNATE PROTOCOL 1
An alternative approach to direct immobilization of the ligand to the sensor chip surface (see Basic Protocol 2) is capture of the ligand onto the sensor chip surface using a highaffinity binding molecule that has been coupled directly to the chip. In the ligand-capture approach described here, regeneration of the surface usually removes both the bound analyte and the captured ligand for each cycle of analyte binding. Some examples of ligand capture include: use of a streptavidin surface to capture biotinylated ligands; use of an anti-tag antibody surface to capture a tagged recombinant protein ligand; and use of an anti-IgG antibody surface to capture an antibody ligand. The use of an anti-mouse IgG surface to capture the mouse monoclonal anti-β2µ-globulin antibody ligand is described here. Amine coupling is used to prepare an anti-mouse IgG surface on the reference and active flow cells. The volume of anti-β2µ-globulin required to reach the desired ligand level must be determined. Regeneration conditions for the anti-mouse IgG surface are provided. Lastly, experimental set-up for kinetic analysis using ligand capture is described. The capture protocol as written assumes that flow cells 1 and 2 have been used for the direct immobilization protocol.
Materials 100 mM N-hydroxysuccinimide (NHS) 400 mM N-ethyl-N-(3-dimethylaminopropyl)-carbodiimide hydrochloride (EDC) 25 µg/ml anti-mouse IgG antibody in 10 mM sodium acetate, pH 5.0 (Biacore) 1 M ethanolamine 5 µg/ml solution of anti-β2µ-globulin in HBS-P+ 10 mM glycine, pH 1.7 Biacore 3000, 2000, or T100 system 0.8-ml tubes Sensor Chip CM5 Perform amine coupling of anti-mouse IgG to surface 1. Under the File tab in the instrument control software, open the Application Wizards, choose Surface Preparation, and then select Immobilization. Enter CM5 for Sensor Chip and Amine Coupling for the Immobilization Method. 2. Select Specify Flow Rate and Injection Time under the immobilization procedure or set-up window. 3. Enter the capture molecule name (e.g., anti IgG) for the ligand name for flow cells 3 and 4. Enter values for the injection parameters for both flow cells: 7 min injection time and 10 µl/min for the flow rate. 4. Click on Next and the wizard will recommend solution volumes and vial positions. Place the solutions in the appropriate reagent rack positions taking care to match the sample content with the indicated rack positions. 5. Save the immobilization wizard template and result files and start the immobilization.
Determine volume of ligand to capture 6. In the instrument control software under the Run tab, select Run Sensorgram. Start sensorgram in flow cell 4. Do not capture the anti-β2µ-globulin ligand in flow cell 3. Fc 3 will be the reference surface and Fc 4 will be the active surface. Identification of Protein Interactions
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7. Set the flow rate to 5 µl/min (this flow rate will be used for the capture step of the kinetic analysis). 8. Under Command Inject, select Manual Inject. Select Load loop, enter 50 µl, and the rack position for the 5 µg/ml anti-β2µ-globulin. 9. Enter 5 µl for the injection volume and Start. 10. Check the binding level. Use the reference line to insert a baseline report point ∼15 sec before the injection start, and measure the amount bound by placing a report point 15 sec before the injection end. 11. Repeat step 4 until the target level of ∼1200 RU is reached. For this experimental system, ∼15 µl will be required to reach the target level of anti-β2µ-globulin. 12. Regenerate the surface after determining the volume of 5 µg/ml anti-β2µ-globulin required for reaching the target level of ∼1200 RU. Adjust the Flow rate to 50 µl/min. Use the Quickinject command (include Extraclean) to inject 10 mM glycine-HCl, pH 1.7, for 2 min. Re-adjust the flow rate to 5 µl/min. 13. Inject the total volume that was determined to capture 1200 RU of anti-β2µ-globulin to confirm that the accumulated volume of anti-β2µ-globulin that was injected will produce the same response as a single amount. Regenerate the surface as in step 7 to remove the anti-β2µ-globulin from the anti-mouse IgG surface before starting Kinetic Analysis. 14. Stop and save the sensorgram. Note the volume required to reach the target level, which will be entered into the Kinetic Analysis wizard. ALTERNATE PROTOCOL 2
KINETIC ANALYSIS FOR LIGAND CAPTURE METHOD OF BINDING LIGAND TO SURFACE This section describes an alternative method to the direct kinetic analysis in Basic Protocol 5. A dilution series of the antigen (analyte), as well as a single dilution of the antibody (ligand), is prepared in running buffer. During each analysis cycle, the ligand is injected over the capture surface using the conditions identified in Alternate Protocol 1 and subsequently one concentration of analyte is injected over the captured ligand, and the binding responses of both are measured. The bound analyte and the captured ligand will be removed from the anti-mouse capture surface using a recommended regeneration. This process will be repeated for each concentration of analyte as a separate cycle, such that the binding response for the capture and each concentration of analyte is recorded in a new sensorgram. The binding responses of the analyte will be overlaid and curve fitting of the entire concentration series (global fitting) will be used to determine the kinetic rate constants and affinity of the interaction.
Materials 8.5 µM stock β2µ-globulin (100 µg/ml) HBS-P+ (see recipe) 10 mM glycine, pH 1.7 5 µg/ml solution of anti-β2µ-globulin in HBS-P+ Set up sample and method 1. Dilute 5.6 µl of 8.5 µM stock β2µ-globulin (100 µg/ml) into 1494.4 µl HBS-P+ to prepare the 32 nM sample. Biacore to Measure Binding Kinetics
2. Prepare 250 µl of a two-fold serial dilution of 32 nM β2µ-globulin (2 nM final).
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3. In the instrument control software Application Wizards, choose Kinetic Analysis. Check the box for Concentration Series and include the control for Mass Transfer, and then click Next. Select Binding Using Capturing Molecule and then click Next. 4. Enter the ligand name and values for the Capture parameters that were identified and used in the previous section: 5 µl/min for the flow rate, 3 min for the injection time, and 1 min stabilization time. 5. Next, enter values for the analyte injection parameters: flow path 4, using 3 as a reference, 30 µl/min for the flow rate, 2 min for analyte injection time, and 5 min for dissociation time. 6. Enter the analyte names (e.g., 32n B2micro, 16n B2micro, etc.), molecular mass (11,800 Da), and number of replicates for each β2µ-globulin concentration in the series. Run at least one analyte sample and the buffer-only sample (0 nM) in duplicate. Also include a buffer-only sample that will be used for double-reference subtraction during data analysis.
7. Choose the Run Order Sorted low to high. 8. Enter the concentration 2 nM for the Mass Transfer control test. 9. Next, provide the regeneration solution name (glycine pH 1.7) and enter values for the regeneration injection parameters: 50 µl/min for the flow rate, 120 sec for injection time, and 1 min stabilization time. 10. Place the solutions in the appropriate reagent rack positions, matching the sample content and recommended volumes with the indicated rack positions. 11. Save the kinetic wizard template and result files and start kinetic analysis.
Evaluate data from kinetic analysis using ligand capture 12. Follow the procedure for kinetic analysis using the direct ligand immobilization method (see Basic Protocol 5, steps 11 to 21) with one difference of removing unwanted data, including cutting the responses from the regeneration injection as well as the responses from ligand capture. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
HBS-P+ 10 mM HEPES, pH 7.4 150 mM NaCl 0.05% polysorbate 20 Store up to 1 week at 4◦ C Dilute 10× HBS-P+ to 1×, filter through a 0.2-µm filter, and degas before use with the Biacore 3000 or 2000 system. Equilibrate to room temperature before use. COMMENTARY Background Information provides descriptions of Biacore systems, sensor chips, immobilization chemistries, and general applications. UNIT 20.2 UNIT 19.13
provides a thorough discussion of background information on optical biosensors, which is pertinent to the protocols described in this unit.
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Critical Parameters and Troubleshooting Experimental design Some experimental design questions should be considered when using Biacore to measure the binding kinetics of protein interactions. Which interactant should be immobilized as the ligand? Often protein-protein interactions can be studied using Biacore with either interactant immobilized as the ligand. General properties such as purity, quantity, mass, stability, valency, isoelectric point, and available tag should be considered when deciding which interactant to immobilize. The purity of the ligand is very important to ensure binding specificity as well as binding capacity. Impurities in the ligand preparation can be immobilized as well as the ligand, which will complicate the determination of ligand density. Impurities in the analyte material can complicate results by introducing nonspecific binding or by affecting the accurate determination of analyte concentration. If an interactant is of limited quantity, it could be used as the ligand, because immobilization, either direct or via capture, usually requires very small amounts of material (2 to 10 µg). Biacore systems can detect molecules as small as 200 Da, therefore, the size of the interactants is not usually a limiting factor. However, the Rmax equation (see Eq. 19.14.1) should be used to calculate whether the ratio of the molecular masses of the interactants would limit the response of the interaction. If one of the interactants has a protein purification tag (e.g., 6His), then that protein could be captured as the ligand using an anti-tag antibody surface. When using Biacore to measure the binding kinetics of an antibody-antigen interaction, the antibody should be immobilized as the ligand to avoid binding avidity effects that result from the bivalency of the antibody. Immobilization of the bivalent protein enables determination of the kinetic rate constants by fitting the responses to a simple 1:1 (Langmuir) binding model.
Biacore to Measure Binding Kinetics
How should the ligand be immobilized? The two general techniques for immobilization of the ligand are: (1) direct coupling to the sensor chip surface using one of several surface chemistries; and (2) capture onto a sur-
face that has been derivatized with a binding molecule (e.g., via a purification tag). The various surface chemistries for direct coupling are described in UNIT 19.13. Ligands that are directly coupled to the surface will have a random orientation; therefore, some binding sites may not be accessible. However, captured ligands often exhibit higher binding activity, which can be critical for some proteins. The higher binding activity of the captured ligand can be attributed to a homogeneous presentation of the ligand, availability of the binding site, and use of fresh ligand for each binding cycle (no damage due to regeneration solution). The ligand capture approach can thus offer advantages over direct immobilization. Several examples of ligand capture strategies are described in UNIT 19.13. What is a good control surface? Usually, a surface that is treated with the same coupling chemistries used to immobilize the ligand serves as a suitable control for the reference surface. When using the ligand capture approach, the capture molecule serves as a suitable control for the reference surface. For some interactions, a nonspecific ligand may act as an appropriate control, such as a scrambled peptide sequence for a protein-peptide interaction. The use of bovine serum albumin as a control ligand is not recommended due to its tendency to nonspecifically bind to many proteins. How much of the ligand should be immobilized? A low ligand density surface is optimal for accurate measurement of the binding kinetics of an interaction. The maximum binding capacity (Rmax ) of the immobilized ligand should be in the range of 50 to 150 RU. Use the rearranged Rmax equation (Eq. 19.14.2) to determine an appropriate immobilization level that will generate an Rmax of ∼100 RU. For the direct immobilization of the anti-β2µ-globulin described in this protocol, 1200 RU was targeted, which results in a calculated Rmax of 189 RU. However, the experimental Rmax for the anti-β2µ-globulin surface is ∼40 RU due to a lower-than-expected surface activity. Is the immobilized ligand active? Surface activity should be calculated using Equation 19.14.3. The BIAevaluation software generates an experimental Rmax value after curve fitting as described in the Evaluation of the Kinetic Analysis section. Generally, 75% surface activity can be expected for ligands
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immobilized directly by amine coupling. Poor surface activity (<50%) can result from denaturation of the ligand due to the preconcentration solution, or inactivation of the binding site due to the immobilization chemistry. In this case, employing an alternative surface chemistry should be considered to improve the surface activity. Impurities in the ligand preparation can also be immobilized when using a direct coupling approach, which could affect the binding capacity of the surface, thereby lowering the apparent surface activity. In this case, employing the ligand capture approach can add an on-chip purification step that will usually result in improved surface activity. Is binding of the analyte specific to the ligand? Inspect the analyte response on the reference flow cell to identify nonspecific binding due to electrostatic or hydrophobic interactions with the sensor chip surface. The bulk refractive index has a square-shaped response, while nonspecific binding will typically have an increasing response on the reference. Electrostatic nonspecific binding can be minimized by the addition of NaCl to the sample and running buffer or by using a sensor chip with less carboxymethylation, such as CM4 (see UNIT 19.13). Nonspecific binding to the CM-dextran can be minimized by addition of soluble CMdextran to the sample at 1 mg/ml. Hydrophobic nonspecific binding often can be minimized by the addition of a detergent, such as 0.05% polysorbate 20 or 10 mM CHAPS, to the sample and running buffers. Nonspecific binding to a capture surface can be resolved by switching to another capture molecule. For example, when capturing biotinylated ligands, nonspecific binding to streptavidin can often be resolved by use of neutravidin (Pierce) as the capture molecule. How should the surface be regenerated? If the analyte does not dissociate from the ligand within a reasonable time (e.g., 20 min), then the surface should be regenerated. Identify a solution that removes the analyte from the immobilized ligand without affecting the ligand activity. An acidic or basic solution, or chaotropic salt solution injected at a high flow rate (50 µl/min) for a short time (1 min) will usually be effective in removal of the analyte from the ligand without damaging the binding activity of the ligand. Determination of ideal regeneration conditions may require optimization. Combining solutions together into “regeneration cocktails” can of-
ten produce efficient regeneration conditions (Andersson, 1999). For example, a combination of low pH and high ionic strength solutions such as 10 mM glycine, pH 2.0, and 1 M sodium chloride; a combination of basic and chaotropic solutions such as 10 mM sodium hydroxide and 500 mM sodium thiocyanate; a combination of a low pH solution and detergent such as 10 mm glycine, pH 2.0, and 0.05% polysorbate 20. An immobilization and regeneration database that provides examples and suggestions for a wide range of interactants is available at the Biacore Website (see Internet Resources). Bulk refractive index The SPR detection technology used in Biacore systems measures changes in the refractive index that are related to changes in mass at the sensor chip surface. Differences in the refractive index between the running buffer and injected sample buffer will give rise to changes in responses that are known as bulk refractive index changes. Bulk refractive index effects are common and do not affect the measurement of the binding interaction when a reference surface is used to subtract the bulk refractive index from the binding response. It is recommended that differences in the bulk refractive index between the analyte and running buffer are minimized by either dialysis or dilution of the analyte into the running buffer. Samples containing high refractive index solutions, such as high–salt solutions, glycerol, or DMSO, should either be buffer-exchanged into the running buffer or the solutions should be added to the running buffer to match the composition of the samples. Immobilization pH scouting Immobilization by covalent coupling cannot be repeated on the same sensor chip surface. Therefore, preconcentration is an important strategy to control the immobilization level for experimental optimization and efficiency. Typically, the optimal pH for preconcentration will be a 0.5- to 1-pH unit below the pI of the protein. Many proteins exhibit limited stability in the low-ionic-strength, low-pH solutions used for preconcentration, therefore, dilution of the ligand just prior to use is recommended. Always perform pH scouting on the flow cell that will be used for the immobilization, not on the reference flow cell. When using standard amine coupling (see Basic Protocol 2), ∼40% of the carboxymethyl groups are converted to reactive esters after a 7-min activation. Thus, the activated
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sensor chip surface maintains a net negative charge during the immobilization procedure, which enables preconcentration. Note that prepared surfaces can have a negative charge during experiments, depending on sensor chip type and blocking procedure. Amine coupling Buffer components that contain primary amines, such as Tris or sodium azide, must be avoided in amine coupling to prevent competition with the ligand for coupling. Ligand contact should be completed within 15 min after activation of the surface with EDC/NHS to ensure coupling before the reactive esters on the surface hydrolyze. Immobilization levels The binding capacity of the surface will depend on the level and activity of immobilized ligand. The SPR response correlates with change in mass concentration on the sensor chip surface, and therefore, depends on the molecular weight (mass) of the analyte in relation to the number of ligand sites on the surface. The term Rmax describes the maximum binding capacity of the surface ligand for analyte in RU. The theoretical Rmax is calculated from Equation 19.14.1:
Rmax = (analyte MW/ligand MW) × RL × Sm where, MW is the molecular weight (mass) of the ligand and analyte, RL (ligand response) is the amount of immobilized ligand in RU, and Sm is the stoichiometry as defined by the number of binding sites on the ligand. Rearrangement of the Rmax equation provides a means of calculating an appropriate ligand density to aim for when performing an immobilization for kinetic analysis as shown by the following Equation 19.14.2:
RL = (ligand MW/analyte MW) × Rmax × (1/Sm )
Biacore to Measure Binding Kinetics
The maximum binding capacity (Rmax ) of the immobilized ligand should be in the range of 50 to 150 RU for measurement of the binding kinetics of an interaction. Use of a low ligand density will minimize mass transfer limitations during the interaction. Mass transfer limitation refers to an experimental situation in which the supply of analyte to the sensor chip surface is a limiting factor for the interaction due to the demand for analyte by a high ligand density. Mass transfer limitations can result in calculated binding ki-
netics that are slower than the true binding kinetics. Partial mass transport limitations can be accounted for in data analysis using a binding model that includes a mass transfer rate constant. An experimental Rmax must be determined to assess the percent activity of the immobilized ligand. The quality of the binding data that can be obtained is reflected in the ligand activity as shown in the following Equation 19.14.3:
%ligand activity = experimental Rmax /theoretical Rmax × 100% Kinetic analysis Accurate determination of kinetic rate constants requires an optimal assay design. Ligand densities that result in a maximum analyte binding response of 50 to 150 RU and fast flow rates of at least 30 µl/min are recommended. Low ligand densities reduce analyte consumption at the surface and fast flow rates minimize the diffusion distance from the bulk flow to the surface, resulting in conditions that reduce the influence of mass transport to accurately measure binding rate constants. Check the surface activity and binding specificity. Always use a reference surface to monitor nonspecific binding and to subtract the bulk refractive index. Inject five analyte concentrations, spanning the range of 0.1 to 10 times KD . Adjust the analyte injection time to collect enough associationphase data to observe curvature in the response. Adjust the dissociation time to collect enough dissociation-phase data to observe decay (>3 RU). For very stable interactions (kd < 10-5 ), the required dissociation time may be as long as 1 to 2 hr. When interactions exhibit fast kinetics, short contact times and dissociation times can be used. If the binding response has reached equilibrium, e.g., interactions that exhibit fast kinetics, then steadystate affinity can be determined. Steady-state affinity is determined by plotting the response at equilibrium (flat response) versus the analyte concentration. The equilibrium dissociation constant (KD ) is determined by fitting this curve to the steady-state affinity model. Include buffer injections as part of the analyte concentration series for blank-run subtraction. Evaluate reference subtracted, blank-run subtracted data by globally (all curves together) fitting the kinetic rate constants simultaneously with the simplest model that describes the interaction. Assess the fit by inspecting the superimposed curves, residuals, and standard error for the calculated parameters. Refer to
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UNIT 19.13, Kinetics and Affinity Analysis section and key references for more information.
Routine maintenance Refer to the instrument’s handbook for comprehensive routine maintenance procedures. After each use or at least once per week, a Desorb routine (under the Tools tab, Working tools) should be used to keep the needle, tubing, and microfluidics clean. The reagents and protocol for the desorb routine are described by the software. When not in use, the Biacore instrument should be left with a maintenance chip docked, running in Standby mode using water as the running buffer.
Anticipated Results The values of the calculated parameters for the kinetic analysis of β2µ-globulin binding to a directly immobilized anti-β2µ-globulin antibody from three independent experiments are provided:
ka = (1.11 ± 0.04) × 106 M−1 sec−1 kd = (2.35 ± 0.25) × 10−3 sec−1 KD = 2.12 ± 0.15 nM Rmax = 32.8 ± 6 RU Chi2 = 0.17 ± 0.04 RU kt = 5.5 × 108 – 1.6 × 1012 Time Considerations The overall time that it takes to develop and conduct kinetic analysis of a protein interaction using Biacore will depend on the nature and behavior of the interactants. In terms of the time required to complete the assays as described here, one 8-hr day should be sufficient to accomplish the protocols for surface preparation and assay development, as well as set-up of the kinetic analysis, which can be run overnight. Data analysis requires ∼1 hr. Derivatized sensor chips can be stored at 4◦ C in a humid environment, however, the activity of the immobilized ligand after storage should be evaluated.
Literature Cited Andersson, K., Hamalainen, M., and Malmqvist, M. 1999. Identification and optimization of regeneration conditions for affinity-based biosensor assays. A multivariate cocktail approach. Anal. Chem. 71:2475-2481.
Key References Andersson et al., 1999. See above. This report describes a systematic approach to producing regeneration solutions and identifying regeneration conditions. Karlsson, R. and F¨alt, A. 1997. Experimental design for kinetic analysis of protein-protein interactions with surface plasmon resonance biosensors. J. Immunol. Methods 200:121-133. This classic paper describes the suitability of using a dextran hydrogel on the sensor chip surface, describes the importance of using a reference, and explores the use of control experiments to gain understanding of reaction mechanisms. Myszka, D.G. 1999. Improving biosensor analysis. J. Mol. Recognit. 12:1-6. This paper underscores the importance of careful experimental design when measuring binding kinetics with biosensors. ¨ Onell, A. and Andersson, K. 2005. Kinetic determinations of molecular interactions using Biacore: Minimum data requirements for efficient experimental design. J. Mol. Recognit. 18:307-317. This work provides recommendations for the design of cost-effective assays that deliver reliable kinetic measurements. Rich, R.L. and Myszka, D.G. 2005. Survey of the year 2003 commercial optical biosensor literature. J. Mol. Recognit. 18:1-39. Rich, R.L. and Myszka, D.G. 2005. Survey of the year 2004 commercial optical biosensor literature. J. Mol. Recognit. 18:431-478. These literature surveys are most highly recommended as guides to careful experimental design, data analysis and presentation, and as excellent resources for outstanding biosensor publications.
Internet Resources http://www.biacore.com/lifesciences/products/ catalog/index.html Biacore catalog. https://www.biacore.com/lifesciences/service/ online support/irdb/index.html Immobilization and regeneration database. http://www.biacore.com/lifesciences/service/ training/courses/usa canada/biacorebasics/ index.html Training courses.
Contributed by Michael Murphy, Laure Jason-Moller, and JoAnne Bruno Biacore, Inc. Piscataway, New Jersey
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Identifying Small-Molecule Modulators of Protein-Protein Interactions
UNIT 19.15
Protein-protein interactions are an essential aspect of all living organisms. These interactions are critical for enzymes that contain multiple subunits and for the regulation of enzymatic activity through allosteric interactions. On a larger scale, protein-protein interactions form the backbone of intricate protein networks and signaling cascades, which are crucial for controlling and regulating many biological processes. The essential nature of these interactions and their involvement in many medically relevant processes has made them attractive as potential drug targets (Cochran, 2000; Toogood, 2002). This unit describes a high-throughput strategy for identifying small molecules that can modulate protein-protein interactions. The basis for this system is the genetic encoding of both the interacting proteins and the small-molecule library inside a host cell, allowing for inhibitor discovery through genetic selection. The interaction of protein targets is monitored using the two-hybrid approach (UNIT 19.2); the primary difference with the approach described in this protocol is that it incorporates a bacterial two-hybrid system, based on the bacteriophage repressor concept (Ladant and Karimova, 2000; Di Lallo et al., 2001). As outlined herein, the two-hybrid system is engineered to operate in reverse, allowing for the identification of molecules that disrupt protein-protein interactions (Vidal and Endoh, 1999). In order to maximize the potential for this approach, the system was designed to interface with genetically encoded small-molecule libraries. The small-molecule libraries are produced using split intein-mediated circular ligation of peptides and proteins (SICLOPPS) technology for the intracellular synthesis of cyclic
Figure 19.15.1 Engineering the DnaE trans-intein of Synechocystis sp. PCC6803 to cyclize an intervening sequence. This strategy is utilized in the SICLOPPS method to make libraries of cyclic peptides (Scott et al., 1999). Identification of Protein Interactions Contributed by Alexander R. Horswill and Stephen J. Benkovic Current Protocols in Protein Science (2006) 19.15.1-19.15.19 C 2006 by John Wiley & Sons, Inc. Copyright
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peptides, developed in the Benkovic laboratory (see Scott et al., 1999). An intein is a segment of a protein that possesses a naturally occurring enzyme activity, which is able to excise itself and splice together the flanking protein fragments of the remaining portions (the exteins) with a peptide bond. A specific trans-intein from Synechocystis, engineered to perform the chemistry on an internal sequence instead of the flanking regions, allows for the cyclization of any sequence of choice (Fig. 19.15.1). This construct, when placed on a plasmid under control of an arabinose-inducible promoter, allows large libraries of cyclic peptides to be generated through transformation into host cells. The benefit of cyclic peptides is that the cyclization renders them resistant to cellular proteases and improves the binding to target proteins. They can also be synthesized in vitro using straightforward methods, enabling detailed biochemical studies on inhibitor function (Horswill et al., 2004). Using this protocol, both the target proteins and elements of a small-molecule library can be compartmentalized in a host cell, accelerating the ability to discover rare protein complex disruptors. Briefly, genes encoding the proteins of interest are fused to repressor DNA-binding domains, and expression is controlled with an IPTG-inducible promoter (Fig. 19.15.2). The formation of a functional repressor blocks the expression-selectable markers, and with defined culture medium a positive selection is created that can be
Small-Molecule Modulators of Protein Interactions
Figure 19.15.2 Scheme for genetic selection for cyclic peptide inhibitors of protein-protein interactions (Horswill et al., 2004). A protein of interest is fused to a repressor DNA-binding domain, which associates to block expression of three genes encoding HIS3 (IGPD, histidine biosynthesis), kan (aminoglycoside 3 -phosphotransferase, kanamycin resistance), and lacZ (β-galactosidase). Background expression can be chemically tuned with 3-aminotriazole (3-AT, competitive inhibitor of IGPD) and kanamycin. By placing E. coli host cells on minimal medium lacking histidine, a genetic selection is created that can be used to identify cyclic peptides that inhibit protein-protein interactions. Reproduced from Horswill et al. (2004) with permission, Copyright 2004 by National Academy of Sciences, U.S.A.
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prepare media for genetic selection
Support Protocol 1 prepare competent cells of selection strain
construct strain for genetic selections Support Protocol 2
transform plasmid library into selection strain
construct cyclic peptide plasmid library
plate on selective media, incubate
process candidates to identify best hits
DNA sequence best candidates
characterize best candidates
Figure 19.15.3 Steps involved in the Basic Protocol. The points at which Support Protocols 1 and 2 become relevant are indicated.
exploited to identify small-molecule inhibitors. By using a plasmid library encoding potential inhibitors to transform the host, cyclic peptides can be identified that disrupt the targeted protein-protein interactions and allow expression of the selectable markers. Additionally, since the cyclic peptides are genetically encoded, the identity of the inhibitor sequences can be rapidly determined, accelerating further rounds of optimization and advancement to biochemical studies. The Basic Protocol describes the genetic selection procedures for identification of cyclic peptide inhibitors of protein-protein interactions. Support Protocol 1 describes the preparation of a strain for the inhibitor selections, and Support Protocol 2 provides methods for the construction of cyclic peptide libraries. A flow chart outlines the steps comprising the various protocols (Fig. 19.15.3; also see the Commentary section and Horswill et al., 2004).
IDENTIFICATION OF SMALL-MOLECULE INHIBITORS OF IN VIVO PROTEIN-PROTEIN INTERACTIONS BY GENETIC SELECTION The combination of a reverse two-hybrid system with cyclic peptide libraries is a powerful approach to identifying inhibitors of protein-protein interactions. By merging these methods into a cell-based assay, large libraries can be quickly analyzed through genetic selection. This protocol presents an example of the use of this system in identifying cyclic peptide inhibitors of the murine ribonucleotide reductase heterodimer (Horswill et al., 2004).
BASIC PROTOCOL
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Materials Selection medium (see recipe) Luria-Bertani (LB) agar plates (APPENDIX 4A) supplemented with 25 µg/ml chloramphenicol E. coli strain SNS-126 containing the two-hybrid reporter and protein domains of interest (or control protein) fused to the bacteriophage repressor DNA-binding domains (Support Protocol 1) 50 to 200 ng cyclic peptide library (Support Protocol 2) SOC broth (see recipe) 1× MMA: diluted from 5× minimal medium A (APPENDIX 4A) LB broth (APPENDIX 4A) with and without 25 µg/ml chloramphenicol 200 mM IPTG Oligonucleotide that hybridizes to the intein fragment (in this case, pAR-NdeI, 5 GGAATTCCATATGGTTAAAGTTATCGGTCGTCGT-3 ) 245 × 245–mm culture plates (Nunc) 100 × 15–mm petri dishes Electroporator and cuvettes 18 × 100–mm culture tubes Toothpicks, sterile Inoculating loops, sterile Plasmid preparation kit, optional (e.g., QIAquick Spin Miniprep Kit; Qiagen) 96-well microtiter plates Multichannel pipettor DNA sequencing facility Additional reagents and equipment for growing bacteria (APPENDIX 4B), preparing electrocompetent cells for electroporation (Seidman et al., 2002), and purifying plasmid DNA (APPENDIX 4C) NOTE: All reagents used with E. coli cells must be sterile.
Prepare media and cells 1. Prepare selection medium with arabinose and optimized levels of 3-amino-1,2,5triazole (3-AT), isopropyl-β-D-thiogalactopyranoside (IPTG) and kanamycin for genetic selection and pour into 245 × 245–mm culture plates. Typically one or two plates are used for each target protein complex, with additional plates for repeats, if necessary. The 3-AT, IPTG, and kanamycin levels are optimized for each target protein complex. For example, for identifying inhibitors of the ribonucleotide reductase protein complex the following levels were used: 2.5 mM 3-AT, 200 µM IPTG, and 50 µg/ml kanamycin (Horswill et al., 2004). Other target protein complexes will require different levels of these tuning reagents to obtain the best results (see Support Protocol 1). If desired, 40 µg/ml Xgal can be added to the medium to improve the visualization of colonies. Xgal is usually dissolved in a small volume of dimethyl formamide prior to adding to culture media. Xgal is a substrate for β-galactosidase (lacZ) enzyme, and the cleavage of this substrate turns E. coli colonies blue. The expression of the lacZ reporter will be enhanced with protein-protein interaction inhibitors.
2. For processing potential inhibitors, prepare additional batches of this selection medium with and without arabinose in 100 × 15–mm petri dishes. If desired, add 40 µg/ml Xgal to the media to enhance the visual observation of growth. Small-Molecule Modulators of Protein Interactions
The number of plates prepared depends on the experiment. The authors typically process eight to twelve candidates per plate. Additional plates without IPTG may be prepared if the optional step 14 is to be carried out.
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3. Prepare LB agar plates with 25 µg/ml chloramphenicol in 100 × 15–mm petri dishes. This media will be used for estimation of the cyclic peptide library size.
4. Prepare high-efficiency electrocompetent cells (see Seidman et al., 2002) of E. coli strain SNS-126 expressing the ribonucleotide reductase fusions.
Transform and plate library 5. Electroporate 50 to 200 ng of a cyclic peptide library into 50 µl of prepared competent cells. Follow the instructions provided by the electroporator manufacturer For E. coli, usually 50 µl of competent cells are used with a 0.2-cm electroporation cuvette, and typical machine conditions are 2.5 kV, 25 µF, and 200 ohms.
6. After the pulse is applied, immediately recover the cells with 950 µl of SOC broth in 18 × 100–mm culture tubes and grow 1 hr at 37◦ C with shaking. The best results are obtained by directly transforming a desalted ligation of a cyclic peptide library. See Support Protocol 2 for the preparation of this ligation. To prepare larger libraries, perform this transformation multiple times and pool the cells.
7. Transfer the culture to a microcentrifuge tube and centrifuge 1 min at 12,000 × g, room temperature. Wash the cells by adding 1 ml of 1× MMA and vortexing. Centrifuge 1 min at 12,000 × g, room temperature and discard the supernatant. Repeat wash and resuspend cells in 1 ml of 1× MMA. 8. Remove 10 µl of resuspended cells and dilute into 990 µl of LB in a microcentrifuge tube. Mix thoroughly, remove 100 µl, and add to 900 µl of LB in a new microcentrifuge tube. Repeat this serial dilution three more times to obtain five tubes at different cell concentrations. 9. Plate 100 µl in duplicate of each dilution onto LB agar plates supplemented with 25 µg/ml chloramphenicol. Incubate overnight at 37◦ C. Only count plates with 30 to 300 colonies to obtain accurate numbers. Chloramphenicol is necessary to maintain plasmid selection.
10. Determine the library size by performing the following calculation:
average colony count × initial dilution factor × 10 = library size. The initial dilution factor ranges from 100 to 1 million depending on the tube used for plating. The additional factor of 10 is included in the calculation to account for plating only 100 µl of each dilution. Library sizes should be in the range of 1 × 107 to 1 × 109 members per ml. Additional information on dilution plating is provided in APPENDIX 4B.
11. Plate the rest of the cell resuspension on the 245 × 245–mm selection medium plate with arabinose and optimal levels of 3-AT, IPTG, and kanamycin prepared in step 1. Incubate the plate at 37◦ C for 3 to 5 days. Individual colonies will begin to appear in this time frame. The use of minimal medium and the addition of kanamycin and 3-AT makes the E. coli cells grow considerably more slowly than on rich media.
12. Count the number of colonies that grew on the 245 × 245–mm selection plate. Knowing the library size, perform the following calculation:
number of candidates/library size = frequency of positives. In a typical selection, 50 to 200 colonies will appear on the minimal medium. The frequency of putative hits arising in the genetic selection is usually in the range of
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1 × 10−5 to 1 × 10−7 positive candidates per library. The number will vary depending on the library size, target protein complex, and stringency of the selection conditions.
Identify and confirm best inhibitor candidates 13. Sector (usually 8 to 12 sectors per plate) and number 100 × 15–mm agar plates of selection medium with and without arabinose (prepared in step 2). Using sterile toothpicks, pick individual colonies from the plate in step 12, streak for isolation with a sterile loop in a sector on both types of plates, and incubate for 3 to 5 days at 37◦ C. A positive candidate should show significant growth only in the presence of arabinose. This processing step is critical to identify the candidates that respond to arabinose induction. Arabinose is required to express the cyclic peptide, and if the peptide is effective at disrupting protein-protein interactions, a growth enhancement should be observed in the presence of the inducer. The inclusion of positive and negative controls on these plates also is helpful in distinguishing the best hits. This processing step will uncover a significant number of false positives, sometimes as high as 90%, which can be removed from the pool of potential inhibitors.
14. Optional: Perform a second processing step in parallel to confirm that the target protein complex is properly expressed in each candidate by restreaking the candidates on selective medium plates lacking both the IPTG and arabinose supplements. Spot 2 µl drops of 200 mM IPTG within each restreak, and incubate the plates 3 to 5 days at 37◦ C. The presence of IPTG induces expression of the target protein complex, blocking growth on the plate. A false positive will grow in the presence of IPTG on selection medium without arabinose, and the candidate can be removed from further screening steps. A true positive will not grow in the presence of IPTG on selection medium without arabinose. A picture of the plates for both the arabinose and IPTG processing steps can be found in Horswill et al. (2004).
15. Purify plasmid DNA from each positive candidate (APPENDIX 4C). Alternatively, use a plasmid prep kit. 16. Reconstruct each strain by retransforming the purified plasmid into the parent E. coli strain. This step is important for identifying additional false positives that passed the arabinose response test.
Evaluate inhibitor candidates’ effectiveness 17. Identify additional false positives and rank the candidates from step 16 for effectiveness as follows:
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a. Grow each strain in culture tubes with LB broth supplemented with 25 µg/ml chloramphenicol to an OD600 of 1.0. b. Wash 1 ml of cells with 1× MMA and resuspend in 1 ml of 1× MMA as in step 6. c. Dispense 90 µl of 1× MMA into each of 6 wells of a microtiter plate per strain to be tested. d. Add 10 µl of resuspended cells into the first well, mix thoroughly, and transfer 10 µl to the next well. e. Repeat this process until serial dilutions have been created in all 6 wells. f. Using a multichannel pipettor, remove 2 µl from each well and spot rows onto the selection medium plates with and without arabinose (prepared in step 2). g. Grow plates at 37◦ C for 3 to 5 days until adequate growth is observed.
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This is a convenient and powerful approach to removing additional false positives and simultaneously rank ordering the best inhibitor candidates. By spotting serial dilutions of cells expressing the peptides on selective media, the growth trends can be compared at each dilution level. The most effective inhibitors should display growth at high dilutions of cells on the selective medium containing arabinose. False positives will not respond to the presence of arabinose. For more details and a visual example of the results of this procedure see Horswill et al. (2004).
18. Optional: Compare the best candidates to a control protein complex to determine the specificity of the cyclic peptide inhibitor. Repeat the process in Steps 15 and 16 to estimate the effectiveness of the identified inhibitors against the control protein complex. The easiest way to perform this test is to have another strain with control proteins (e.g., leucine zipper) fused to the DNA-binding domains. Take the purified plasmid DNA of each candidate inhibitor and transform this strain, and repeat the spotting of serial dilutions. Some cyclic peptide inhibitors identified in this protocol will be very hydrophobic and display nonspecific inhibitory effects that will allow disruption of other protein complexes. These types of inhibitors can be identified using this simple processing step. It is helpful to have a known inhibitor for the control protein complex to allow better comparison with candidate inhibitors.
Sequence and biochemically characterize candidates 19. Select the best candidates from the previous step and sequence the plasmid insert to determine the residues of the cyclic peptide inhibitor. Use an oligonucleotide that hybridizes to the intein fragment for automated DNA sequencing, and perform sequencing according the DNA facility instructions. The sequence of the encoded inhibitors can be compared for selection of the most interesting candidates for further characterization. See Scott et al. (1999) and Abel-Santos et al. (2003) for information about the SICLOPPS plasmids required to analyze sequencing data. At this stage it is important to take the best inhibitors identified through the genetic selection and characterize them biochemically.
20. Synthesize the cyclic peptides and biochemically characterize their ability to disrupt protein-protein interactions using, e.g., kinetic assays, ELISA, or fluorescence quenching methods. Often it is useful to test the inhibitor in linear form before making the cyclic form. Linear peptides can be ordered through many commercial suppliers. Cyclic peptides can be generated biologically for biochemical experiments (Scott et al., 1999), but it is difficult or expensive to obtain the amounts necessary for most assays. Straightforward methods for synthesizing cyclic peptides have been described elsewhere (Horswill et al., 2004; Naumann et al., 2005). Also see Chapter 18 for further information on cyclizing peptides. The choice of biochemical technique for analyzing the cyclic peptide potency will depend entirely on the target. For some proteins, kinetic assays are sufficient to determine inhibitor effectiveness (Tavassoli and Benkovic, 2005). For ribonucleotide reductase, competition and peptide ELISA assays were developed and tryptophan fluorescence quenching was performed (Horswill et al., 2004). Chapter 19 is dedicated to methods for investigating protein-protein interactions, and these protocols should serve as a good starting point for analyzing candidates.
CONSTRUCTION AND OPTIMIZATION OF A STRAIN FOR INHIBITOR SELECTION In order to identify inhibitors of protein-protein interactions, an E. coli strain for the reverse two-hybrid selection needs to be constructed, and media conditions need to be optimized. Any homodimeric or heterodimeric protein complex of interest can be incorporated into this selection system. For disrupting homodimeric interactions, E. coli strain SNS-118 is used, which possesses a triple reporter (depicted in Fig. 19.15.4)
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Figure 19.15.4 Graphic map of the RTHS plasmids used for making repressor fusions and their respective promoter sequences: (A) Plasmid pTHCP16 for constructing homodimeric fusions requires a strain containing the phage 434 wild-type promoter (E. coli strain SNS118 derivatives). (B) The wild-type 434 promoter. (C) Plasmid pTHCP14 for constructing heterodimeric fusions requires a strain containing the chimeric phage 434·P22 promoter (E. coli strain SNS126 derivatives). (D)The chimeric 434·P22 promoter.
under control of the homodimer-dependent promoter (Fig. 19.15.4B). For disrupting heterodimeric interactions, E. coli strain SNS-126 is used, which expresses the reporters from a heterodimer-dependent promoter (Fig. 19.15.4D). The level of protein-protein interactions will be determined by β-galactosidase assays (Miller, 1992). See Horswill et al. (2004) for further background information on these strains and constructs and additional details on optimizing media conditions.
Materials
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Oligonucleotides (specifically designed) Plasmid pTHCP16 (Fig. 19.15.4A, for making homodimeric fusions) or pTHCP14 (Fig. 19.15.4C, for making heterodimeric fusions) DNA template (gene of interest) 2.5 U/ml proofreading PCR polymerase (e.g., Deep Vent, New England Biolabs; Pfu, Stratagene) 0.1 M EDTA, pH 8.0 10% SDS 2.5 mg/ml proteinase K 3 M sodium acetate, pH 5.2 70% (v/v) and 100% ethanol PCR purification kit (e.g., QIAquick, Qiagen; optional) DNA purification kit (e.g., QIAquick Spin Miniprep Kit, Qiagen; optional) Restriction enzymes (specific to the cloning procedure) Gel extraction kit (e.g., QIAquick gel extraction kit, Qiagen; optional) T4 DNA ligase (New England Biolabs) E. coli competent cells (e.g., DH5α): prepared (APPENDIX 4D) or purchased from commercial suppliers LB agar plates (APPENDIX 4A) supplemented with 100 µg/ml ampicillin and with and without IPTG (0 to 1 mM)
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E. coli strain SNS-118 (homodimeric reporter) or SNS-126 (heterodimeric reporter) CRIM plasmids (E. coli Genetic Stock Center, Yale), optional Z buffer (see recipe) Chloroform 0.1% (w/v) SDS 4 mg/ml o-nitrophenyl-β-D-galactopyranoside (ONPG): prepared fresh 1 M sodium bicarbonate (Na2 CO3 ) 3 /4 in. sterile, optical glass culture tubes, optional 13 × 100–mm glass tubes (disposable) Additional reagents and equipment for growing bacteria (APPENDIX 4B), performing PCR (APPENDIX 4J), separating DNA by agarose gel electrophoresis (APPENDIX 4F), purifying plasmid DNA (APPENDIX 4C), quantitating DNA (APPENDIX 4K) NOTE: For more information about preparing, analyzing, and cloning DNA see APPENDIX 4.
Clone target gene 1. Design the oligonucleotide primer for cloning the target gene, incorporating restriction sites that are compatible with the appropriate vector for generating homodimeric (pTHCP16; Fig. 19.15.4A) or heterodimeric (pTHCP14; Fig. 19.15.4C) repressor fusions to the 434 repressor DNA-binding domain. Note that it is critical for the reading frame to be maintained, allowing the creation of a translational fusion. The restriction sites shown in Figure 19.15.4 are unique to plasmids pTHCP14 and pTHCP16. Before selecting a restriction site, check that the enzyme does not cut the target gene of interest. An example of an oligonucleotide design for cloning into the BamHI site would be: 5 -GTTGTTGGATCC-target sequence (20 to 25 bp)-3 .
2a. For a homodimeric interaction: Set up a PCR reaction with the designed oligonucleotide primers, DNA template, and proofreading PCR polymerase (see APPENDIX 4J for more details on PCR). 2b. For a heterodimeric interaction: Perform two different PCR reactions for the separate genes. 3. Run an analytical agarose gel (APPENDIX 4F) to confirm the size and quality of the PCR product(s).
Purify target gene PCR product and vector DNA 4. Optional, but recommended: To completely remove the DNA polymerase, add the following (assuming 50 µl of PCR mix): 2.5 µl of 0.1 M EDTA (pH 8.0), 2.5 µl of 10% SDS, 1 µl of 2.5 mg/ml proteinase K. Incubate 30 min at 55◦ C. DNA polymerase can reduce the cloning efficiency.
5. Purify the PCR product by adding one-tenth vol 3 M sodium acetate, pH 5.2 and two vol ice-cold 100% ethanol. Incubate 30 min on ice or at −20◦ C. 6. Centrifuge 10 min at 12,000 × g, 4◦ C. Wash the pellet with 70% ethanol and centrifuge 5 min at 12,000 × g. Alternatively, use a commercial PCR purification kit. 7. Purify plasmid DNA of the selected construct, either pTHCP14 or pTHCP16 (see APPENDIX 4C or other standard DNA purification procedure) or use a DNA purification kit.
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Insert target gene DNA into vector 8. Digest the plasmid DNA and purified PCR product using the selected restriction enzymes based on the cloning design. Follow the manufacturers’ instructions for setting up the restriction digestion. Restriction enzymes and buffers from New England Biolabs work well for cloning into these plasmids. Enzymes from other manufacturers should also work.
9. Electrophorese the digested DNA on an agarose gel (APPENDIX 4F). 10. Excise the portions of the gels containing the desired DNA fragments, and purify the DNA from the agarose using a standard procedure or a gel extraction kit. Quantitate the DNA using a spectrophotometer (see APPENDIX 4K). 11. Prepare a T4 DNA ligation reaction using pTHCP14/16 vector and PCR product from step 10. Follow the recommendations for DNA concentrations, incubation time, and incubation temperature provided by the manufacturer of the T4 DNA ligase. 12. Transform the ligation product by electroporation into E. coli competent cells. See steps 4 and 5 in the Basic Protocol for more details on DNA transformation. 13. Plate the transformation mixture onto LB agar plates supplemented with ampicillin. 14. Grow up individual transformants and purify plasmid DNA (see APPENDIX 4C). Verify that the plasmids were constructed correctly through restriction digests and DNA sequencing. Perform the DNA sequencing according to the DNA facility instructions.
Construct two-hybrid reporter strain 15. Transform the finished plasmids into E. coli strain SNS-118 or SNS-126, depending on the target protein complex. Plate the transformation mixture onto LB agar plates supplemented with ampicillin. The pTHCP14/16 plasmids are derivatives of pMAL-c2x (New England Biolabs), which are IPTG-inducible, medium-copy, ampicillin-resistant plasmids (Horswill et al., 2004). It is recommended that control strains be constructed with the pTHCP14 or 16 plasmid without inserts.
16. Optional: Integrate the constructs onto the chromosome to improve the genetic selection performance, using CRIM plasmids. Identify restriction enzyme cut sites and clone the construct into a suitable CRIM plasmid. Alternatively, perform the cloning with PCR. Integrate the plasmid into the chromosome and verify as described (Haldimann and Wanner, 2001). In the case of ribonucleotide reductase, the construct was removed from pTHCP14 using BsaBI and SacI and cloned into pAH68 cut with HincII and SacI (Horswill et al., 2004). Although this is an additional cloning step, it generally results in a significant improvement in the genetic selection of inhibitors.
Quantitate protein-protein interaction using the β-galactosidase reporter 17. Prepare overnight cultures of constructed strains in LB broth with ampicillin (100 µg/ml). 18. Subculture 50 µl of the overnight cultures into tubes containing 5 ml of LB with ampicillin and IPTG ranging from 0 to 1 mM. If cells need to placed in a cuvette for absorbance readings, the culture may be scaled up as necessary. Some spectrophotometers, such as a Spectronic 20D+, have adaptors that allow direct measurements on 3/4 -in. glass culture tubes. Small-Molecule Modulators of Protein Interactions
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19. Grow the stains at 37◦ C with shaking (∼250 rpm) until the OD600 reaches 0.3 to 0.7. Once the culture reaches this cell density, place the tube on ice. For best results, stop growth of all the cultures at similar OD readings. Current Protocols in Protein Science
20. Vortex cell cultures, remove 0.1 ml, and microcentrifuge 1 min at 10,000 × g, room temperature. Resuspend pellet in 1 ml of Z buffer and transfer to 13 × 100–mm glass tubes. It is important to wash the cells to remove excess IPTG. IPTG will competitively inhibit ONPG cleavage by β-galactosidase. Perform the assays in triplicate to obtain accurate numbers.
21. Add 2 drops of chloroform and 1 drop of 0.1% SDS to each tube. Vortex 10 sec and place in water bath at 28◦ C. The chloroform and SDS disrupt the E. coli membranes to allow ONPG to diffuse into the cells.
22. Start the β-galactosidase reaction by adding 0.2 ml of ONPG. Shake tubes briefly to mix. Record the time of ONPG addition. Staggering the start time for each reaction will make it easier to record the end point.
23. Watch the tubes carefully for the development of yellow color. Stop the reaction with 0.5 ml of 1 M Na2 CO3 , once sufficient yellowness has appeared. Note the time. 24. Transfer samples to microcentrifuge tubes and centrifuge 1 min at 10,000 × g. Determine the OD at both 420 and 550 nm for the supernatant of each sample. 25. Calculate Miller units of β-galactosidase activity (see APPENDIX 4L or Miller, 1992). The levels of repression will vary widely for different targets. Using the wild-type repressor 434 as a control, repression levels of over 10-fold are usually observed. With most protein fusions, the amount repression often is in the range of 3- to 6-fold, compared to vector only (pTHCP14/16). Although this level of repression seems small, it is adequate to perform the genetic selection. At high levels of IPTG (≥1 mM), even the DNA-binding domains expressed without fusions (pTHCP14/16 vector only) will show signs of repression. Additionally, it is important to note that a low background level of β-galactosidase is present due to secondary promoters within the HIS3-Kan region (Fig. 19.15.2). This additional transcription through the lacZ gene has no effect on the genetic selection, but it will make the repression range appear slightly smaller through β-galactosidase assays. For selecting the optimal level of inducer for genetic selection, it is recommended that the lowest amount of IPTG required to achieve maximal repression is chosen. Usually it is helpful to plot units of activity versus IPTG concentration, and the point at which the plot begins to plateau is selected for the media conditions. There is no defined method for determining the optimal 3-AT and kanamycin concentrations for genetic selection. The simplest approach is to prepare media with a range of 3-AT and kanamycin levels, and test the media with the constructed selection strain. For example, prepare minimal media (1× MMA) with 0.5% glycerol, 1 mM MgSO4 , 13 µM arabinose (2 × 10−4 %), 25 µg/ml chloramphenicol, IPTG (concentration determined above), and vary the 3-AT (5 to 15 mM) and kanamycin (25 to 100 µg/ml) levels. Following the Basic Protocol, transform a test library into the selection strain, plate on the prepared media, and incubate the plates. Wait until colonies arise on the plates, and visually determine which media conditions offer the best separation of background from the positive candidates. Use these conditions for inhibitor discovery in subsequent experiments.
CONSTRUCTION OF CYCLIC PEPTIDE LIBRARIES Cyclic peptide libraries are generated through a PCR-based technique using oligonucleotides containing randomized residues. The oligonucleotides described in this protocol will produce cyclic peptides containing six-membered rings. These libraries are termed C+5 to indicate the presence of one nucleophilic cysteine and five random residues. To make other types of libraries, the randomized region can be altered to include random or fixed residues as needed. Additionally, the nucleophilic residue can be changed to serine, although the cyclization step will occur at a slower rate (Scott et al., 1999). Current Protocols in Protein Science
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In order to facilitate library construction, a second zipper PCR is performed to ensure that random nucleotides properly anneal to a complementary strand. Under optimal conditions, this protocol can achieve libraries with >108 members. For additional information see Abel-Santos et al. (2003).
Materials LB broth (APPENDIX 4A) supplemented with 30 µg/ml chloramphenicol E. coli culture containing plasmid pARCBD-p (Scott et al., 1999), or another compatible plasmid DNA purification kit (e.g., QIAquick Spin Miniprep Kit, Qiagen; optional) 2.5 U/µl proofreading PCR polymerase (e.g., Deep Vent; New England Biolabs) and 10× buffer 2 mM (each) dNTP mix Oligonucleotide primers C+5 (5 -GGAATTCGCCAATGGGGCGATCGCCCACAATTGTNNSNNSNN SNNSNNSTGCTTAAGTTTTGGC-3 ) CBD-reverse (5 -GGAATTCAAGCTTTCATTGAAGCTGCCACAAGG-3 ) zipper (5 -GGAATTCGCCAATGGGGCGATCGCC-3 ) PCR purification kit, optional (e.g., QIAquick,Qiagen) 100 mM EDTA, pH 8 10% (w/v) SDS 2.5 mg/ml proteinase K Restriction enzymes: BglI and HindIII (e.g., New England Biolabs) Phosphatase (e.g., shrimp alkaline phosphatase; United States Biochemical) Gel extraction kit (e.g., QIAquick gel extraction kit, Qiagen) T4 DNA ligase (e.g., Promega) 3 M sodium acetate, pH 5.2 100% ethanol Thermal cycler 0.05 micron VM nitrocellulose membranes (Millipore), optional 250-ml beaker, optional Additional reagents and equipment for purifying plasmid DNA (APPENDIX 4C), purifying DNA (APPENDIX 4E), performing agarose gel electrophoresis (APPENDIX 4F), and quantitating DNA by spectroscopy (APPENDIX 4K) Purify plasmid pARCBD-p 1. Start a culture in 5 ml LB broth supplemented with 30 µg/ml chloramphenicol of an E. coli strain containing pARCBD-p Plasmid pARCBD-p contains an arabinose-inducible promoter, the engineered intein construct for SICLOPPS, and a C-terminal chitin-binding domain. This particular version generates the eight-residue cyclic peptide pseudostellarin F (Scott et al., 1999). A detailed protocol for preparing other constructs similar to pARCBD is described elsewhere (AbelSantos et al., 2003).
2. Purify plasmid DNA from the E. coli strain using any established procedure (see APPENDIX 4C) or DNA purification kit. 3. Prepare mix for PCR amplification as follows (final volume 50 µl):
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5 µl 10× DNA polymerase buffer (1×) 5 µl 2 mM (each) dNTP mix (200 µM) 5 µl 10 µM primer C+5 (1 µM) 5 µl 10 µM primer CBD-reverse (1 µM) 10 to 50 ng pARCBD-p template DNA (from step 2) 1 µl 2.5 U/µl proofreading PCR polymerase (0.1 U/µl) H2 O to a final volume of 50 µl. Current Protocols in Protein Science
4. Perform PCR using the following amplification cycles (typically): 30 cycles
30 sec 30 sec 1 min
94◦ C 53◦ C 72◦ C
(denaturation) (annealing) (extension)
See APPENDIX 4J for more information about performing PCR. The C+5 oligo contains five codons with the NNS reduced genetic code, with “N” being any one of the four bases and “S” being only G and C. This randomized region encodes for all 20 amino acids using only 32 codons and reduces the frequency of stop codons. To prepare different types of cyclic peptide libraries, oligonucleotides can be synthesized with altered randomized regions.
5. Check the PCR product using agarose gel electrophoresis (APPENDIX 4F). The expected size of the fragment is 0.6 kb.
6. Purify the PCR product through ethanol precipitation (APPENDIX 4C) or alternatively, use a purification kit. 7. Set up a second PCR as in step 4 using the oligonucleotides zipper and CBD-reverse. Use 10 ng of the purified PCR product from the previous step as template, and use the proofreading PCR polymerase according to manufacturer’s instructions. 8. As with the first PCR, run an agarose gel to confirm the second reaction. 9. To remove the DNA polymerase from the sample, add the following (assuming a 50-µl PCR reaction): 2.5 µl of 100 mM EDTA (pH 8.0), 2.5 µl of 10% SDS), 1 µl of 2.5 µg/ml proteinase K. Incubate 30 min at 55◦ C. 10. Purify the PCR product using ethanol precipitation (APPENDIX 4E) or with a purification kit. 11. Digest the pARCBD-p plasmid and second PCR product with the BglI and HindIII restriction enzymes according to manufacturer’s instructions. Dephosphorylate the vector to reduce background with a commercially available phosphatase. 12. Size-separate the entire restriction digest of the vector and PCR product on an agarose gel, and cut out the appropriate bands corresponding to the pARCBD vector (3.4 kb) and PCR product (0.6 kb). 13. Purify the DNA from the agarose fragments using a standard procedure or a gel extraction kit. Quantitate the DNA using a spectrophotometer (APPENDIX 4K). 14. Prepare a T4 DNA ligation reaction using purified pARCBD vector and PCR product. Follow the recommendations for DNA concentrations, incubation time, and incubation temperature provided by the manufacturer of the T4 DNA ligase. 15. Remove salt from the ligation reaction by ethanol precipitation using 3 M sodium acetate and 100% ethanol as in steps 5 and 6 of Support Protocol 1. An alternative and more convenient method of desalting ligations is through dialysis (Goryshin et al., 2000). For dialysis, place 0.05 micron VM nitrocellulose membranes (Millipore) onto a beaker containing 10,000-fold excess of sterile water. Spot the ligation onto the membrane, dialyze for one hr, and remove the sample from the membrane. Some volume change may occur during the dialysis. Identification of Protein Interactions
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Selection medium Autoclave 4 g of granulated agar in 200 ml of distilled water in a 500-ml flask. Once the molten agar has cooled, quickly add 50 ml of 5× minimal medium A (APPENDIX 4A) and the following supplements to the indicated final concentrations.
0.5% (v/v) glycerol 1 mM MgSO4 25 µg/ml chloramphenicol 13 µM arabinose (0.0002%; if required) 3-Amino-1,2,5-triazole (3-AT; see below) Isopropyl-β-D-thiogalactopyranoside (IPTG; see below; if required) Kanamycin (see below) 40 µg/ml 5-bromo-4-chloro-3-indolyl-β-D-galactoside (Xgal; optional). Mix thoroughly and pour into 245 × 245–mm culture plates Optimize 3-AT, IPTG, and kanamycin levels for each target protein complex. For example, for identifying inhibitors of the ribonucleotide reductase protein complex the following levels were used: 2.5 mM 3-AT, 200 µM IPTG, 50 µg/ml kanamycin (Horswill et al., 2004). Other target protein complexes will require different levels of these tuning reagents to obtain the best results (see Support Protocol 1).
SOC broth 20 g tryptone 5 g yeast extract 0.5 g NaCl 0.19 g KCl per liter Mix components with 900 ml water, adjust pH to 7.0 with NaOH, bring volume to 1 liter with water, and autoclave. After cooling add the following to the final concentrations indicated in parentheses: MgCl2 (10 mM), MgSO4 (10 mM), and glucose (0.5%).
Z buffer Combine the following ingredients to the final concentrations indicated in parentheses:
Na2 HPO4 ·7H2 O (60 mM) NaH2 PO4 ·H2 O (40 mM) KCl (10 mM) MgSO4 ·7H2 O (1 mM) 2-mercaptoethanol (50 mM) Adjust pH to 7.0 COMMENTARY Background Information
Small-Molecule Modulators of Protein Interactions
Disrupting protein-protein interactions The ability to discover small-molecule inhibitors that disrupt protein-protein interactions holds tremendous promise for drug development. The completion of many genome sequences and improvements in mapping pro-
tein networks have provided estimates of 6800 interacting pairs in E. coli (Marcotte et al., 1999), 16,000 to 26,000 in yeast (Grigoriev, 2003), and potentially over 375,000 in humans (Ramani et al., 2005). Considering that many of these protein-protein interactions are quite specific, those involved in medically relevant
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processes may ultimately become pharmaceutical targets. Identifying molecules that disrupt proteinprotein interactions has proven to be a challenging task. These interfaces are often large, flat, and lacking in discrete binding pockets that facilitate inhibitor binding, making rational design of candidate compounds very difficult. Additionally, there are few methods available for systematically screening for protein complex inhibitors (Toogood, 2002). However, pioneering work on the study of protein complexes suggests that a small portion of the interface residues constitute the majority of the free energy of binding (Clackson and Wells, 1995). These so called “hot spots” are a select group of clustered residues that could be targeted with inhibitors, allowing a small molecule to break apart a large protein-protein interface. The disruption of protein-protein interactions with reverse two-hybrid systems (RTHS) is a well established method for identifying potential dissociative inhibitors. The concept has been explored extensively in yeast, relying on the availability of reporters that are conditionally toxic to the cell, e.g., CYH2 and URA3, which confer cycloheximide and 5fluoroorotic acid (5-FOA) sensitivity, respectively (Leanna and Hannink, 1996; Vidal et al., 1996). In a RTHS with these markers, the formation of a functional protein complex activates transcription of the CYH2 or URA3 reporters, which blocks cell growth and creates a situation allowing positive selection of interface inhibitors. Despite the attractiveness of the RTHS, the full capability of this approach has been underutilized (Toogood, 2002). Typically an RTHS is coupled to a lowthroughput, phenotype-based screen to identify protein-protein disruptors from chemical libraries (Huang and Schreiber, 1997). Although this approach has been successful, it offers little improvement over high-throughput enzymatic assays, generates numerous false positives, and is limited by the size of a synthetic chemical library. Advances in combinatorial chemistry have improved and increased library sizes, but for many laboratories it is simpler and more cost effective to produce genetically encoded libraries, which are typically orders of magnitude larger and perfectly suited for interfacing with the RTHS. A number of approaches for preparing genetically encoded libraries have been pursued and some have even been coupled to a RTHS. Methods for making linear peptide
libraries have been developed, and screening one of those libraries identified candidate peptides with antibacterial properties (Walker et al., 2001). Although potentially useful, linear peptides have the inherent problem of being unstable in most living organisms, due to the presence of numerous proteases, and thus likely unsuitable to disrupting most protein complexes. Constraining peptides in protein scaffolds, often termed peptide aptamers, provided a significant breakthrough and allowed researchers to stabilize the presentation of a bioactive peptide and improve binding to the target, facilitating in vivo screening of libraries (Hoppe-Seyler et al., 2001). The attractive properties of peptide aptamers allowed researchers to exploit them as genetic tools, uncovering new players in complex regulatory networks (Geyer et al., 1999; Norman et al., 1999). Not surprisingly, peptide aptamers have also been integrated with the RTHS, with a goal of identifying constrained peptides that disrupt protein-protein complexes (Serebriiskii et al., 2002). Unfortunately, except for some rare examples (Park and Raines, 2000), few attempts have been made to draw on the power of this approach to discover and biochemically characterize those peptide sequences that can inhibit proteinprotein interactions. Comparison of reverse two-hybrid systems There are advantages and disadvantages to the various RTHS that have been developed. Most RTHS that have been described in the literature are designed for yeast (Vidal and Endoh, 1999), although some have been developed for bacteria, such as the system described in this unit. There are a number of advantages to the yeast systems: 1) the two-hybrid concept and the most commonly utilized systems are all developed for yeast; 2) the yeast systems are quite robust and generally give less background than bacterial systems; and 3) post-translational modifications are performed in yeast, which is critical for some mammalian proteins. However, there are disadvantages as well, such as lower transformation efficiency, compared to E. coli. Additionally, protein-protein interactions that are essential in yeast cannot be inhibited, but these complexes can be studied and disrupted in bacteria. When selecting an RTHS for discovering small-molecule inhibitors, care must be taken to determine the most appropriate system for the protein target. The RTHS described
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and utilized in this unit is a bacterial version, based on the bacteriophage repressor 434 (Horswill et al., 2004). This system offers a number of notable advantages compared to others: 1) the system is based in E. coli, allowing for very large libraries (>108 members) to be screened against a target; 2) it possesses two selectable markers, amplifying the power of the genetic selection; 3) the selectable markers are tunable, enabling the level of background to be controlled; and 4) it has a quantitative reporter, allowing the strength of protein-protein interactions to be determined.
Small-Molecule Modulators of Protein Interactions
Additional features of RTHS systems One of the most important features of the outlined RTHS system is its ability to couple with genetically encoded cyclic peptide libraries. Every aspect of the system was designed to allow its various components to integrate and work collectively as one convenient cell-based assay for inhibitor selection (Horswill et al., 2004). It starts with the E. coli strain, which serves as the host for all the whole-cell assays. The genetic background of the strain is a modified version of BW27786, a strain that was developed to improve gene expression from arabinose-inducible promoters (Khlebnikov et al., 2001). Since the intein construct of the SICLOPPS plasmids is under control of an arabinose promoter (Scott et al., 1999; 2001), the use of this E. coli strain allows tight control over the production of cyclic peptides. The next important feature of this RTHS is the design of the two-hybrid strategy (shown in Fig. 19.15.2). Available transcriptional repression schemes, based on the bacteriophage life cycle, allow the implementation of RTHS in a positive selection format. Briefly, proteins of interest are spliced to a phage 434 repressor (cI protein), and the interacting fusions associate to form a functional complex that blocks transcription of conditionally essential genes (Fig. 19.15.2). The release of the repression through complex disruption allows cells to grow under selective conditions and enables the identification of desired library candidates. To monitor homodimeric interactions, the gene encoding the protein of interest is cloned as a translation fusion to the 434 repressor DNA-binding domain on plasmid pTHCP16 (Fig. 19.15.4A). To monitor true heterodimeric interactions, an approach was adapted for creating heterodimeric repressors that specifically recog-
nize chimeric (434 and P22) bacteriophage operator sites (Di Lallo et al., 2001), thereby bypassing problems encountered using proteins that form higher ordered oligomers (Ladant and Karimova, 2000). Two genes of interest are cloned as translational fusions to the 434 and mutated 434 DNA-binding domains on plasmid pTHCP14 (Fig. 19.15.4C). Both plasmid pTHCP14 and pTHCP16 possess an IPTG-inducible pTac promoter, enabling control of the level of fusion protein produced in the cell. Additionally, if necessary, the entire construct can be integrated onto the chromosome of the E. coli host strain (Haldimann and Wanner, 2001), bypassing the need to maintain selection of the plasmid. The RTHS works through the formation of functional repressors, which block transcription from a compatible phage promoter. There are two promoters for this system, one for the native 434 repressor for examining homodimeric interactions (Fig. 19.15.4B), and a second promoter with a chimeric operator site for examining heterodimeric interactions (Fig. 19.15.4D). These promoters drive expression of two conditional markers, enabling coupled selections based on growth advantage. The last feature of the system is the triple reporter that contains the selectable markers (Fig. 19.15.2). The first reporter is the Saccharomyces cerevisiae HIS3 gene, encoding imidazole glycerol phosphate dehydratase (IGPD) in the histidine biosynthetic pathway, which is known to complement a corresponding E. coli histidine auxotrophy (Joung et al., 2000). Importantly, the choice of the HIS3 gene product as a selectable marker is expected to yield few false positives, an inherent problem of antibiotic-dependent selections. To enable this selectable marker, the gene encoding the corresponding E. coli enzyme, hisB, was deleted from the BW27786 strain (Horswill et al., 2004). Incorporation of a second auxiliary reporter gene encoding aminoglycoside 3 -phosphotransferase for kanamycin resistance was employed to amplify selection sensitivity. Further, the HIS3 enzyme is competitively inhibited by 3-amino-1,2,5triazole (3-AT) and levels of kanamycin resistance correlate well with the phosphotransferase expression, allowing chemical tunability (Brennan and Struhl, 1980). Finally, the lacZ gene was added to allow quantitative measurements of protein-protein interactions, and the entire triple reporter construct was integrated into an E. coli BW27786 strain containing the hisB deletion.
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Advantages of genetic selection One of the primary advantages of the method described in this unit is the capacity for genetic selection, which allows very large libraries (>108 members) to be screened against a variety of targets quite rapidly. By having everything genetically encoded, the entire method can be performed at low cost in most laboratory settings with minimal equipment, accelerating the identification of potential lead inhibitors. In contrast, synthetic chemical libraries have to be purchased or prepared at great expense and have to be screened, usually through low-throughput approaches. These features make it very difficult for many laboratories to exploit the benefits of synthetic libraries for inhibitor identification. Furthermore, genetic selection has other benefits, such as the ability to translate affinity to function. Using this protocol to identify ribonucleotide reductase protein-protein interaction inhibitors, candidate inhibitors were identified that unexpectedly bound to the smaller mR2 subunit (Horswill et al., 2004). If the target was preselected for an affinity-based method, such as phage display, these inhibitors may not have been discovered. Benefits of cyclic peptide libraries The use of cyclic peptide libraries offers additional advantages over other types of genetically encoded libraries. Cyclic peptides, like peptide aptamers, are constrained into a fixed conformation, improving the binding of a candidate inhibitor over its linear counterpart. Additionally, during biochemical characterization, it might be challenging to purify significant quantities of a peptide aptamer, and there is the constant concern over the contribution of the protein scaffold to the inhibitory effect. With cyclic peptides, the scaffold is unnecessary, and large quantities can be synthesized in a straightforward manner with chemical methods (Horswill et al., 2004; Naumann et al., 2005), facilitating biochemical studies. Moreover, the functional epitope on a cyclic peptide is presented in a structural state amenable to further synthetic optimization (Hirschmann et al., 1998), yet another benefit that can aid their development as lead compounds for pharmaceutical studies.
Critical Parameters and Troubleshooting The success of identifying small-molecule inhibitors is entirely dependent on the proteinprotein interaction in question and the ability of the fusion protein to form a complex that
behaves as a repressor to inhibit transcription of the reporter genes. If the protein target is eukaryotic in origin, post-translational modifications will not be performed in E. coli, and this could affect the level of interactions. Additionally, some proteins are deleterious to E. coli, and this toxicity can mask repression. Clearly, considerations should be made as to whether this protocol is the best suited to the target in question. One of the most common problems is poor repression of the reporter construct by the DNA-binding fusions to the target gene(s). Often this occurs with proteins that have a weak interaction or are poorly expressed in E. coli. If this is known to be an issue, it might suggest the proteins in question are not fully compatible with this system. For those not experienced with bacterial two-hybrid systems, the ones based on bacteriophage repressors tend to have smaller fold differences compared to yeast systems (Ladant and Karimova, 2000). Even if the repression level is only 3- to 4-fold, based on β-galactosidase assays, inhibitors can be identified with this protocol. If expected repression of β-galactosidase is quite poor or not observed at all, several factors should be considered. Sometimes it can be helpful to construct additional fusion constructs with a longer fragment or with the addition of a flexible peptide linker; in the case of heterodimers, the fragments can be swapped between the two DNA-binding partners (Fig. 19.15.4). Make sure a full range of IPTG concentrations is explored to produce adequate levels of fusion proteins. Although it takes time to build the constructs, integration of the gene(s) onto the chromosome can often bypass repressionrelated problems (Horswill et al., 2004). Some protein fusions perform better as singlecopy integrants rather than from multi-copy plasmids. For the genetic selection conditions, the most important step is the identification of media conditions where protein-protein interactions inhibitors will allow growth over background. If conditions are too stringent, positive candidates could be missed in the selection. Additionally, high-level expression of the fusion proteins can mask the effect of potent candidates by titrating out the inhibitor population. Using the minimum level of IPTG for fusion protein expression should help avoid this potential problem. The integration of the fusion constructs also helps maintain control over the expression level. For challenging targets, sometimes it can be helpful to perform the genetic selection at both low stringency
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(low 3-AT and kanamycin levels) and high stringency (high 3-AT and kanamycin levels), which can increase the pool of positive candidates available for screening.
Anticipated Results In a successful genetic selection, anywhere from 50 to >300 colonies will arise on library plates for each set of 108 transformants. Of these candidates, usually 5% to 20% pass the arabinose response and retransformation tests, while the rest are probably false positives. Further filtering through growth ranking will weed out the candidate pool, often to only 1% to 5% of the original number. For example, in a case study on disrupting the ribonucleotide reductase complex, 108 total transformants of a C+5 cyclic peptide library were subjected to genetic selection as described in this unit. Of 262 candidate inhibitors processed, 24 passed the arabinose response and retransformation tests. After growth ranking, 8 candidates were selected for biochemical studies. Under these optimal conditions, a library of 108 inhibitors was pared down to 8 promising candidates through a simple series of genetic selection and processing steps (Horswill et al., 2004). Similar results have been obtained with other homodimeric and heterodimeric complexes, suggesting this example is representative.
Time Considerations The amount of time it takes to proceed from targets of interest to isolating protein-protein interaction inhibitors can vary widely when using the methods outlined in this unit. The entire procedure, going from choice of protein target to inhibitor identification, should be expected to take at least two months. Although this is a lengthy protocol, most of the individual steps are not time or labor intensive, giving the researcher flexibility to perform the project in parallel with other work. Additionally, the bulk of the time is spent on the construction of the selection strain and the determination of media conditions. Once all the conditions have been optimized, additional rounds of inhibitor selections can proceed quite quickly, allowing rapid panning of new or focused inhibitor libraries.
Literature Cited Small-Molecule Modulators of Protein Interactions
Abel-Santos, E., Scott, C.P. and Benkovic, S.J. 2003. Use of inteins for the in vivo production of stable cyclic peptide libraries in E. coli. Methods Mol. Biol. 205:281-294.
Brennan, M.B. and Struhl, K. 1980. Mechanisms of increasing expression of a yeast gene in Escherichia coli. J. Mol. Biol. 136:333-338. Clackson, T. and Wells, J.A. 1995. A hot spot of binding energy in a hormone-receptor interface. Science 267:383-386. Cochran, A.G. 2000. Antagonists of protein-protein interactions. Chem. Biol. 7:R85-94. Di Lallo, G., Castagnoli, L., Ghelardini, P., and Paolozzi, L. 2001. A two-hybrid system based on chimeric operator recognition for studying protein homo/heterodimerization in Escherichia coli. Microbiology 147:1651-1656. Geyer, C.R., Colman-Lerner, A., and Brent, R. 1999. “Mutagenesis” by peptide aptamers identifies genetic network members and pathway connections. Proc. Natl. Acad. Sci. U.S.A. 96:8567-8572. Goryshin, I.Y., Jendrisak, J., Hoffman, L.M., Meis, R., and Reznikoff, W.S. 2000. Insertional transposon mutagenesis by electroporation of released Tn5 transposition complexes. Nat. Biotechnol. 18:97-100. Grigoriev, A. 2003. On the number of proteinprotein interactions in the yeast proteome. Nucleic Acids Res. 31:4157-4161. Haldimann, A. and Wanner, B.L. 2001. Conditional-replication, integration, excision, and retrieval plasmid-host systems for gene structure-function studies of bacteria. J. Bacteriol. 183:6384-6393. Hirschmann, R., Hynes, J., Jr., Cichy-Knight, M.A., van Rijn, R.D., Sprengler, P.A., Spoors, P.G., Shakespeare, W.C., Pietranico-Cole, S., Barbosa, J., Liu, J., Yao, W., Rohrer, S., and Smith, A.B., 3rd. 1998. Modulation of receptor and receptor subtype affinities using diastereomeric and enantiomeric monosaccharide scaffolds as a means to structural and biological diversity. A new route to ether synthesis. J. Med. Chem. 41:1382-1391. Hoppe-Seyler, F., Crnkovic-Mertens, I., Denk, C., Fitscher, B.A., Klevenz, B., Tomai, E., and Butz, K. 2001. Peptide aptamers: new tools to study protein interactions. J. Steroid. Biochem. Mol. Biol. 78:105-111. Horswill, A.R., Savinov, S.N., and Benkovic, S.J. 2004. A systematic method for identifying small-molecule modulators of proteinprotein interactions. Proc. Natl. Acad. Sci. U.S.A 101:15591-15596. Huang, J. and Schreiber, S.L. 1997. A yeast genetic system for selecting small molecule inhibitors of protein-protein interactions in nanodroplets. Proc. Natl. Acad. Sci. U.S.A. 94:13396-13401. Joung, J.K., Ramm, E.I., and Pabo, C.O. 2000. A bacterial two-hybrid selection system for studying protein-DNA and protein-protein interactions. Proc. Natl. Acad. Sci. U.S.A. 97:73827387. Khlebnikov, A., Datsenko, K.A., Skaug, T., Wanner, B.L., and Keasling, J.D. 2001.
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Homogeneous expression of the P(BAD) promoter in Escherichia coli by constitutive expression of the low-affinity high-capacity AraE transporter. Microbiology 147:3241-3247. Ladant, D. and Karimova, G. 2000. Genetic systems for analyzing protein-protein interactions in bacteria. Res. Microbiol. 151:711-720. Leanna, C.A. and Hannink, M. 1996. The reverse two-hybrid system: a genetic scheme for selection against specific protein/protein interactions. Nucleic Acids Res. 24:3341-3347. Marcotte, E.M., Pellegrini, M., Ng, H.L., Rice, D.W., Yeates, T.O., and Eisenberg, D. 1999. Detecting protein function and protein-protein interactions from genome sequences. Science 285:751-753. Miller, J. 1992. Procedures for working with lac. In A Short Course in Bacterial Genetics, pp. 71-80. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Naumann, T.A., Savinov, S.N., and Benkovic, S.J. 2005. Engineering an affinity tag for genetically encoded cyclic peptides. Biotechnol. Bioeng. 92:820-830. Norman, T.C., Smith, D.L., Sorger, P.K., Drees, B.L., O’Rourke, S.M., Hughes, T.R., Roberts, C.J. Friend, S.H., Fields, S., and Murray, A.W. 1999. Genetic selection of peptide inhibitors of biological pathways. Science 285:591-595.
the biosynthesis of backbone cyclic peptide libraries. Chem. Biol. 8:801-815. Seidman, C.E., Struhl, K., and Sheen, J. 2002. Introduction of plasmid DNA into cells. In Short Protocols in Molecular Biology, 5th ed. (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 1-29 to 1-31. John Wiley & Sons, New York. Serebriiskii, I.G., Mitina, O., Pugacheva, E.N., Benevolenskaya, E., Kotova, E., Toby, G.G., Khazak, V., Kaelin, W.G., Chernoff, J., and Golemis, E.A. 2002. Detection of peptides, proteins, and drugs that selectively interact with protein targets. Genome Res. 12:1785-1791. Tavassoli, A. and Benkovic, S.J. 2005. Genetically selected cyclic-peptide inhibitors of AICAR transformylase homodimerization. Angew. Chem. Int. Ed. Engl. 44:2760-2763. Toogood, P.L. 2002. Inhibition of protein-protein association by small molecules: approaches and progress. J. Med. Chem. 45:1543-1558. Vidal, M., Brachmann, R.K., Fattaey, A., Harlow, E., and Boeke, J.D. 1996. Reverse two-hybrid and one-hybrid systems to detect dissociation of protein-protein and DNA-protein interactions. Proc. Natl. Acad. Sci. U.S.A. 93:10315-10320. Vidal, M. and Endoh, H. 1999. Prospects for drug screening using the reverse two-hybrid system. Trends Biotechnol. 17:374-381.
Park, S.H. and Raines, R.T. 2000. Genetic selection for dissociative inhibitors of designated proteinprotein interactions. Nat. Biotechnol. 18:847851.
Walker, J.R., Roth, J.R., and Altman, E. 2001. An in vivo study of novel bioactive peptides that inhibit the growth of Escherichia coli. J. Pept. Res. 58:380-388.
Ramani, A.K., Bunescu, R.C., Mooney, R.J., and Marcotte, E.M. 2005. Consolidating the set of known human protein-protein interactions in preparation for large-scale mapping of the human interactome. Genome Biol. 6:R40.
Contributed by Alexander R. Horswill University of Iowa Iowa City, Iowa
Scott, C.P., Abel-Santos, E., Wall, M., Wahnon, D.C., and Benkovic, S.J. 1999. Production of cyclic peptides and proteins in vivo. Proc. Natl. Acad. Sci. U.S.A. 96:13638-13643.
Stephen J. Benkovic The Pennsylvania State University University Park, Pennsylvania
Scott, C.P., Abel-Santos, E., Jones, A.D., and Benkovic, S.J. 2001. Structural requirements for
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Determination of Protein Contacts by Chemical Cross-Linking With EDC and Mass Spectrometry Chemical cross-linking of proteins is an established method for obtaining rough structural information about protein complexes. By identifying the cross-linked residues of neighboring proteins, spatial alignments can be deduced and a rough three-dimensional model can be built. Similarly, in monomeric proteins, cross-links can reveal information about protein folding and domain orientation. In addition, analysis of the cross-linked products on SDS-PAGE gels can help partially distinguish intramolecular and intermolecular cross-links (see Anticipated Results).
UNIT 19.16
BASIC PROTOCOL
Mass spectral analysis of cross-linked proteins has emerged as a simple and efficient technique for the analysis of protein folding and protein-protein interactions. When a folded monomeric protein or a protein complex (multimeric protein) in a native conformation is chemically cross-linked, a covalent link between two amino acids is formed. The distance between these linked amino acids is defined by the three-dimensional folding of the protein and is “frozen” by the spacer length of the cross-linker. This distance is maintained during all subsequent purification and analysis stages. The more cross-links made and identified, the greater the number of conformational restraints that can be applied for low-resolution structural modeling. Currently many protein cross-linking reagents are commercially available and a large number of custom-synthesized reagents have been reported (Sinz, 2003; Trakselis et al., 2005). These include cross-linkers reactive to different amino acids and of various spacer lengths. The use of the cross-linking reagent 1-ethyl-3-[3-dimethylaminopropyl] carbodiimide hydrochloride (EDC) is described in this unit as an example. The general rules discussed below for the use of EDC (e.g., the use of buffers nonreactive to the cross-linker; cross-linker to protein ratio) apply for all cross-linkers. These and other guidelines are also found in the supplier’s instructions. EDC is known as a “zero-length” cross-linker and introduces a chemical bond between a negatively charged side chain carboxyl group (Glu or Asn) and a positively charged side chain amino group (Lys). The close proximity of the reacting groups allows a very precise pinpointing of the sites of protein interactions. The yield of cross-linked protein from this type of experiment varies significantly, depending on the nature and folding state of the protein, the accessibility of the reactive amino acids and the concentration of the cross-linker. It is recommended to start with a small amount of material and perform trial reactions with varied amounts of cross-linker. The conditions below represent a good starting point. NOTE: Throughout the protocol acetonitrile is used at different concentrations. It may extract additives and plasticizers from the plastic tubes which will result in contaminating peaks in the mass spectra. Before the experiment, half-fill all tubes with 70% acetonitrile in 0.1% trifluoroacetic acid. Close the tubes. Shake or vortex vigorously for about 1 min. Discard wash. Rinse with water and allow to air dry.
Materials Purified protein(s) to be cross-linked, of a known concentration (6 to 8 nmol minimum) 0.5 M sodium N-morpholinoethanesulfonate (MES buffer), pH 6.5 (see recipe) Identification of Protein Interactions Contributed by Lyuben N. Marekov Current Protocols in Protein Science (2007) 19.16.1-19.16.8 C 2007 by John Wiley & Sons, Inc. Copyright
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0.2 M [1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride] (EDC; Pierce) in 50 mM MES; prepare immediately before use 0.02 M sulfo-N-hydroxysuccinimide (Sulfo-NHS;Pierce) in 50 mM MES; prepare immediately before use. 0.5 M hydroxylamine hydrochloride 25 mM Tris·Cl, pH 8.0 (APPENDIX 2E) in acetonitrile:water (1:1) 100% acetonitrile Trypsin working solution (see recipe) 25 mM Tris·Cl, pH 8.0 in water 25 mM Tris·Cl, pH 8.0 in 20% (v/v) acetonitrile 80% H2 16 O/20% (v/v) acetonitrile 80% H2 18 O/20% (v/v) acetonitrile Modified trypsin, sequencing grade (Promega) 95% isotopically enriched H2 18 O (Sigma-Aldrich) 2.5% (v/v) trifluoroacetic acid in water 0.1% (v/v) trifluoroacetic acid in 50% (v/v) acetonitrile 0.1% (v/v) trifluoroacetic acid in water 0.1% formic acid in 50% (v/v) acetonitrile or methanol Dialysis membrane or micro dialysis cups of appropriate MWCO (Pierce) or gel filtration spin column (MacroSpin columns; http://nestgrp.com, no. SMM 025.25; also see UNIT 8.3) Scalpel Speedvac evaporator Clear 0.5- or 1.5-ml microcentrifuge tubes 30◦ C incubator Reversed-phase C18 ZipTip pipet tips (Millipore) or OMIX Tip C18 (Varian) Additional reagents and equipment for dialysis (APPENDIX 3B) or buffer exchange using a gel filtration column (UNIT 8.3), mass spectrometry (Chapter 16), SDS-PAGE gel electrophoresis (UNIT 10.1), and gel staining and destaining (UNIT 10.5) 1. Prepare the protein or proteins of interest in a solution of 50 mM MES buffer, pH 6.5, so that the concentration is ∼1 mg/ml. 2. Dialyze (APPENDIX 3B) extensively against 50 mM MES buffer, pH 6.5. Alternatively, perform a buffer exchange with a spin column. This step is crucial as trace amounts of carbonate, acetate, phosphate, or aminocontaining buffers from the protein preparation procedures will interfere with the crosslinking.
3. Add 10-fold molar excess of EDC over the carboxylic groups. To calculate the amount of carboxyl groups use the equation: (µg protein/molecular weight [kDa])×(number of glutamic and aspartic acid residues) = nmol protein carboxyls Example: 0.2 mg (200 µg) of a 40-kDa protein with 18 carboxylic groups contains: (200 [µg]/40 [kDa]) × (18) = 90 nmol carboxylic groups To calculate the amount of EDC required 90 (nmol carboxylic groups) × 10-fold molar excess = 900 nmol EDC 0.2 M EDC = 200 nmol/µl. Add 900/200 = 4.5 µl Protein Contacts by Chemical Cross-Linking & MS
4. Add 0.9 µl of 0.02 M sulfo-NHS to achieve a 50:1 EDC/sulfo-NHS ratio. 5. Incubate up to 4 hr at room temperature.
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6. At various time intervals (e.g., 15, 30, 60, 120 min) withdraw aliquots containing sufficient protein to be visualized on an electrophoresis gel. Immediately quench the reaction by adding hydroxylamine hydrochloride to a final concentration of 10 mM. Add the appropriate amount of (2× or 4×) reducing, denaturing SDS sample buffer. 7. Analyze the aliquots by denaturing SDS gel electrophoresis (UNIT 10.1), stain and destain the gels (UNIT 10.5). Depending on the results, repeat the experiment with modified conditions to optimize the reaction. 8. Perform a full-scale reaction. 9. Separate the products by 1-D SDS-PAGE (UNIT (UNIT 10.5).
10.1),
stain and destain the gels
Perform in-gel protein digestion Also see UNITS 11.3 and 16.4 for more information on enzymatic digestion of proteins for mass spectrometric analysis. 10. Using a clean scalpel cut out the stained band(s) of interest. 11. Estimate the volume of the excised bands using the equation below: Volume of band [mm3 ] = width × height × gel thickness. Sum the volume of all the bands. Multiply by 1.2 to determine the volume [µl] of trypsin solution needed for step 19.
12. Cut each gel slice into small pieces (<1 mm3 ) and place into a plastic tube. 13. To extract buffer and salts add 250 to 300 µl 25 mM Tris·Cl, pH 8.0 in acetonitrile:water (1:1) and vortex for 10 min. 14. Using gel loading pipet tip, remove the supernatant and discard. 15. Repeat steps 13 and 14 twice. 16. Add 250 to 300 µl 100% acetonitrile and vortex for 5 min. At this stage the gel pieces are dehydrated; they will shrink and turn opaque.
17. Using gel loading pipet tip, remove the supernatant and discard. 18. Evaporate the gel pieces to complete dryness in a Speedvac evaporator (∼10 min). 19. Add trypsin working solution to fully cover the gel pieces. The volume for each sample was determined in step 11.
20. Rehydrate the gel pieces for 10 min. If necessary add small amounts of (3 to 5 µl) trypsin working solution until the all the trypsin is absorbed. At this point there will be a 1- to 2-mm clear layer of excess trypsin solution above the gel pieces.
21. Aspirate unabsorbed trypsin working solution with a gel loading pipet tip. Carefully overlay with 25 mM Tris·Cl, pH 8.0 in water to cover the gel pieces by 1 to 2 mm. Removal of excess trypsin will reduce the amount of autolytic peptides. Overlaying with 25 mM Tris·Cl, pH 8.0 will prevent the gel pieces from drying during the next step.
22. Cap the tube. Incubate the sample at 37◦ C overnight. At this stage the samples can be kept frozen for up to a month at −70◦ C.
Extract peptides 23. To the gel pieces, add 50 µl 25 mM Tris·Cl, pH 8.0 in 20% acetonitrile, vortex 20 to 30 min, then centrifuge 30 sec at 700 × g, room temperature.
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24. Transfer supernatant to a clean tube. 25. Repeat steps 23 and 24. Combine supernatants. 26. Split the combined supernatants equally into two microcentrifuge tubes. Evaporate both to complete dryness in a Speedvac evaporator.
Perform differential isotopic labeling of peptides Also see UNIT 23.4 for more information on isotopic labeling. At this stage the trypsin will exchange the two 16 O atoms at the carboxyl end of the peptides generated in the previous stage for two 18 O atoms. 27. Reconstitute the first sample to its original volume (step 26) with nonisotopically labeled 80% H2 O/20% acetonitrile and vortex for a few minutes. Acetonitrile helps dissolve hydrophobic peptides that may have adsorbed to the surface of the plastic tube and also enhances the enzymatic activity of trypsin.
28. Reconstitute a vial (20 µg) of sequencing grade modified trypsin (Promega) with 20 µl water (H2 16 O). 29. Add 1 µl (1 µg) trypsin working solution to the sample. The remaining trypsin can be stored up to 2 months at −20◦ C in the original vial. Store in aliquots if more than one freeze-thaw cycle is expected.
30. Process the second sample identically (steps 27 to 29) except use 80% H2 18 O/20% acetonitrile and trypsin dissolved in 95% H2 18 O water. 31. Incubate both samples for 4 hr at 30◦ C. 32. Stop the isotopic exchange by adding 2.5% trifluoroacetic acid in regular Milli-Q water. The pH should be <4.0.
33. Combine the contents of both vials.
Desalt peptides and exchange solvent for MS analysis 34. Wet C18 ZipTip pipet tip as follows. Aspirate 10 µl of 50% acetonitrile in 0.1% trifluoroacetic acid and discard solvent. Repeat. 35. Equilibrate tip: Aspirate 10 µl of 0.1% trifluoroacetic acid in water and discard solvent. Repeat. 36. Load tip: Aspirate up to 10 µl of sample into the tip. Slowly dispense into the same tube. Repeat 5 to 7 times. 37. Rinse tip: Aspirate 10 µl of 0.1% trifluoroacetic acid in water and discard solvent. Repeat 3 to 5 times. 38. Elute peptides: a. For MALDI-TOF analysis: Aspirate 5 to 8 µl 0.1% trifluoroacetic acid in 50% acetonitrile. Discharge into a clean tube. Sample can be stored at −70◦ C for more than 6 months.
b. For LC/MS analysis: Elute with 5 to 8 µl of 0.1% formic acid in 50% acetonitrile or methanol. Discharge into a clean tube. Protein Contacts by Chemical Cross-Linking & MS
Sample can be stored at −70◦ C for more than 6 months.
39. Analyze the sample by MS. (For additional details, see UNIT 16.3).
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
MES buffer, 0.5 M Dissolve 19.5 g MES-hydrate (Sigma) in ∼180 ml Milli-Q water. Adjust pH to 6.5 with 10 N NaOH (∼ 7.0 ml). Bring volume to 200 ml with Milli-Q water. Use this as a 10× stock solution to prepare 50 mM working solutions.
Trypsin stock solution Reconstitute a vial (20 µg) of sequencing-grade modified trypsin (Promega) with 20 µl of 1 mM hydrochloric acid. The reconstituted enzyme is stable up to 2 weeks at 0◦ to 4◦ C.
Trypsin working solution Immediately before use add 1 µl of trypsin stock solution to 50 µl of 25 mM Tris·Cl, pH 8.0 in water.
COMMENTARY Background Information Ion-pairing interactions between acidic and basic amino acid residues play an important role in stabilization of the three-dimensional structure of proteins. The close proximity between the negatively and positively charged groups makes them ideal candidates for introducing a cross-link. Carbodiimides catalyze the formation of amide bonds between carboxylic acids and amines (DeTar and Silverstein, 1966) by activating the carboxyl group (I) to form an O-acylurea (III) (Fig. 19.16.1A). This intermediate can react with an amine (II) to form the amide (IV). However, the intermediate is short-lived in aqueous solutions and quickly hydrolyzes to a urea derivative and releases the free carboxyl group. The introduction of small amounts of water-soluble N-hydroxysuccinimide (NHS) to the reaction (Fig. 19.16.1B; Staros et al., 1986) enhances the coupling yields by transforming the Oacylurea (III) to a more stable active ester intermediate (V). Enzyme-catalyzed double 18 O labeling by trypsin, Lys-C, and Glu-C provides highly specific, complete, and stable isotopic labeling of the peptide products from a protein (Schnolzer et al., 1996). Due to the incorporation of two 18 O atoms in each C-terminus, normal peptides shift 4 mass units and cross-linked peptides, containing two C-termini, shift 8 mass units and can be easily identified in mass spectra (Back et al., 2002).
The digestion and labeling processes can be separated (Yao et al., 2003). Proteins can be cross-linked, solubilized, separated on a gel, and digested–all in water–and stored frozen. When the peptides are finally isolated and dried they are easily soluble in minimal amounts of water or 18 O water (expensive). Trypsin can incorporate/exchange 18 O atoms in predigested peptides at this stage (Yao et al., 2003). The cross-linked peptides can then be sequenced. During MS/MS sequencing the peptide backbone is fragmented producing a complex mixture of amino-(a-, b-, and c-type) and carboxy-(x-, y- and z-type) terminal fragment ions. The fragmentation is most common at the peptide bonds giving rise to the relatively intensive b- and y-type ions. The C-terminal y-type ions are easily identified by the mass spectrometer as 8 mass units doublet peaks and a “ladder sequence” can be extracted. An alternative method to identify crosslinked amino acids/peptides is to use a commercially available program GPMAW (General Protein/Mass Analysis for Windows; http://www.gpmaw.com). The user defines one or two protein sequences and a cross-linker and the software generates a database of all theoretical cross-links. Then the MS-identified crosslink masses can be matched against this database and the sequences of the participating peptides identified. The demo version of GPMAW is fully functional and will run 20 times.
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Figure 19.16.1 The mechanism of condensation (cross-linking) reaction between a protein side chain carboxyl (I) and amine (II) group. (A) Reaction with EDC alone. (B) NHS forms a more stable active ester (V) and increases the yield of the isopeptide (cross-link) bond (IV).
Protein Contacts by Chemical Cross-Linking & MS
There are occasions when proteins do not contain closely positioned ion pairs to allow the formation of cross-links. Pierce (www.piercenet.com) offers a variety of crosslinking reagents. They are reactive towards different amino acid side chains and their “spacer arms” can span between residues separated by ◦ a range of distances up to ∼20 A. (A technical handbook “Cross-Linking Reagents” can be ordered online). However, when the distance between two cross-linked amino acids increases, so does the ambiguity about their spatial orientation in the protein. In this case the more cross-links identified the greater the number of spatial constraints that can be applied when modeling the three-dimensional structure. Recently, an alternative method for isotopic labeling has been introduced by Pierce in which an equimolar mixture of identical lysine-to-lysine cross-link reagents, one of them with 4 hydrogen atoms and the other with
4 deuterium atoms in the linker, is used. The cross-linked peptides are again easily identified as doublets 4 mass units apart.
Critical Parameters and Troubleshooting Residual amounts of acetate, phosphate, carbonate, or amino groups from the initial protein purification steps will reduce the yield by consuming the cross-linker. Make sure that the protein is extensively dialyzed, free of contaminating buffers and salts. If ionic strength adjustment is desired use ultra-pure NaCl or KCl. If the protein is not highly soluble under the initial conditions, vary the pH between 5.7 and 6.9 (Sehgal and Vijay 1994). Lower pH favors the stability of the active ester (III; Fig. 19.16.1); higher pH favors the formation of the cross-link (IV), so pH 6.5 is a good average and a starting point. If starting at low pH, after a certain amount of time the pH may be carefully raised to
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pH 7.0 to 7.5 by stepwise addition of microliter amounts of 0.5 M HEPES, pH 8.2, vortexing and checking the pH on narrow-ranged indicator paper. The use of ammonium bicarbonate in the in-gel digestion buffer is the generally accepted procedure for mass spectroscopy (UNITS 11.3 & 16.4). During the subsequent freeze drying step most of it will evaporate and pH adjustment with buffer in (expensive) H2 18 O for the isotopic exchange step will be necessary. Use of Tris·Cl buffer avoids this problem. Depending on the nature of the protein other proteases may be used for digestion. Apart from trypsin, three other members of the serine family of proteases (e.g., Lys-C, Glu-C, and chymotrypsin) have been shown to incorporate 18 O at the C-terminus (Yao et al., 2003). In such cases, the same protease should be used in the labeling step. Some specific features of these proteases limit their application. Lys-C, cleaving at the carboxy-terminal side of lysine, is highly specific and can be used under identical conditions as trypsin. As it generates relatively large peptides (hence larger cross-linked peptides), its use is limited only to relatively basic peptides, where trypsin would produce numerous very short peptides. Glu-C, cleaving at the carboxy-terminal side of glutamic acid at nearly identical conditions (50 mM Tris·Cl, pH 8.3) is rather specific and can be used. However, published data about its use for mass spectroscopy are rather scarce. One reason for this is the fact that while trypsin
and Lys-C generated peptides that have (at least) one basic residue and ionize well in the mass-spectrometer, not all peptides generated by Glu-C do so and will not be detected. Still, Glu-C can be used by more experienced MS users. Chymotrypsin, however, is not as highly specific and should be avoided.
Anticipated Results Following the separation of the cross-linked proteins on a SDS-PAGE gel, two types of protein bands can be observed: (1) Monomer bands with (often) progressively decreasing molecular weight relative to the nonmodified protein. These bands are due to intramolecular cross-links which prevent the protein from completely unfolding in the electrophoresis SDS sample buffer. Excising these bands and analyzing them further will identify closely located amino acids and will provide information about the folding of the protein. (2) Higher molecular weight bands corresponding to dimers, trimers, and higher multimers. The cross-links deduced from analysis of these bands usually will carry information about the contact points on the surfaces of the interacting protein molecules, but also may originate from intramolecular cross-links. Excluding the cross-links determined from the monomer bands may facilitate the identification of the intermolecular links. Still, the discrimination between the two types of cross-link cannot always be straightforward in case of homodimeric (homo-oligomeric) proteins.
Figure 19.16.2 Portion of a MALDI-TOF spectrum displaying the isotopic distribution of a crosslinked H2 16 O/H2 18 O-double-labeled peptide.
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This protocol should produce a set of proteolytic peptides, all of which, other than the peptide containing the original C-terminus, are labeled >95% with two 18 O atoms. The 18 Olabeled peptides can then be mixed with the unlabeled ones and analyzed. In the mass spectrometer the masses of the uncross-linked peptides will be seen as doublets 4 mass units apart. The cross linked peptides will produce doublets 8 mass units apart (Fig. 19.16.2).
Time Considerations The most time-consuming part of the protocol is the extensive dialysis (2 or 3 changes of buffer), which usually takes up to 24 hr. The ingel digestion is usually carried out overnight. Extraction of peptides, 18 O-labeling and peptide purification can easily be accomplished in a day.
Sehgal, D. and Vijay, I.K. 1994. A method for the high-efficiency of water-soluble carbodiimidemediated amidation. Anal. Biochem. 218:87-91. Sinz, A. 2003. Chemical cross-linking and mass spectrometry for mapping three-dimensional structures of proteins and protein complexes. J. Mass Spectrom. 38:1225-37. Staros, J.V., Wright, R.W., and Swingle, D.M. 1986. Enhancement by N-hydroxysulfosuccinimide of water-soluble carbodiimide-mediated coupling reactions. Anal. Biochem. 156:220-222. Trakselis, M.A., Alley, S.C., and Ishmael, F.T. 2005. Identification and mapping of protein-protein interactions by a combination of cross-linking, cleavage, and proteomics. Bioconjugate Chem. 16:741-751. Yao, X., Afonso, C., and Fenselau, C. 2003. Dissection of proteolytic 18 O labeling: endoprotease catalyzed 16 O-to-18 O exchange of truncated peptide substrates. J. Proteome Res. 2:147-152.
Key References Acknowledgement This research was supported by the Intramural Research Program of the National Institute of Arthritis and Musculskeletal and Skin Diseases of the National Institutes of Health.
Literature Cited Back, J.W., Notenboom, V., de Koning, L.J., Muijsers, A.O., Sixma, T.K., de Koster, C.G., and de Jong, L. 2002. Identification of crosslinked peptides for protein interaction studies using mass spectrometry and 18 O labeling. Anal. Chem. 74:4417-4422. DeTar, D.F. and Silverstein, R. 1966. Reactions of carbodiimides. II. The reactions of dicyclohexylcarbodiimide with carboxylic acids in the presence of amines and phenols. J. Am. Chem. Soc. 88:1013-1019. Schnolzer, M., Jedrzejewski, P., and Lehmann, W.D. 1996. Protease-catalyzed incorporation of 18 O into peptide fragments and its application for protein sequencing by electrospray and matrix assisted laser desorption/ionization mass spectrometry. Electrophoresis 17:945-953.
Sinz, 2003. See above. An extensive review of cross-linkers, types of crosslinks and methods of MS analysis. Trakselis et al., 2005. See above. Recent advances and current experimental techniques for identifying protein contacts and networks by mass spectrometry.
Internet Resources http://www.gpmaw.com GPMAW (General Protein/Mass Analysis for Windows). Software with many useful features for handling and analyzing protein and peptide sequences. http://www.piercenet.com The company offers a wide choice of cross-linking reagents with varying spacer lengths and reactive towards different amino acid side chains.
Contributed by Lyuben N. Marekov National Institutes of Health Bethesda, Maryland
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CHAPTER 20 Quantitation of Protein Interactions INTRODUCTION
T
he highest level of protein structure is quaternary structure, formed by the noncovalent association of protein subunits. Usually the associating protein subunits have a folded, or tertiary, structure, although there are cases where unfolded proteins with only secondary structure directly associate to form folded multisubunit complexes (e.g., trimeric glucagon). Protein complexes can range from simple symmetrical homodimers formed of identical subunits to highly complex multisubunit structures such as viral capsids. Protein-protein interactions are important, indeed vital, to many biological events and processes. Analysis of interactions between proteins and detailed analysis of molecular interfaces—generally done in vitro using isolated proteins—are the bases for understanding many biological processes. A number of biochemical and biophysical techniques can be applied to study specific interactions involving proteins. This information, together with high-resolution structural information obtained by X-ray crystallography and NMR (Chapter 17), can provide an understanding of the molecular basis of these interactions. Methodologies for studying the identification and quantitation of protein interactions have become so important that the editors of this volume have decided to devote two chapters to these techniques. In Chapter 19, the emphasis is on the detection of specific protein interactions; in this chapter the material is more concerned with quantitative aspects— e.g., the measurement of binding strengths and rates of binding. Quantitation of binding is usually performed using biophysical techniques such as gel filtration, equilibrium dialysis, large-zone analytical size-exclusion chromatography (SEC), and titration microcalorimetry, to name a few; these techniques are reviewed in UNIT 20.1. Some of these methods rely on sophisticated equipment such as the analytical ultracentrifuge. It is recognized that many laboratories will not own such instruments, so the first units covering each method will be extended overviews designed both to supplement the information in the general chapter overview and to provide investigators with enough background to both follow the published literature and, if necessary, intelligently seek collaboration. These overviews will be complemented by actual experimental protocols to be published in future supplements.
In this vein, then, comes UNIT 20.2, the first extended overview. This unit discusses the measurement of protein interactions by optical biosensors (including the popular Biacore instrumentation). It includes a Strategic Planning section on how to proceed with an experiment—depending on whether the binding will be steady-state, kinetic, or competitive—and information on sample preparation, analysis of results, and troubleshooting of suboptimal experimental results. The second overview, UNIT 20.3, discusses the classical biochemical method of sedimentation equilibrium using an analytical ultracentrifuge. Presented are: background information; basic theory; strategies to use for planning experiments, and the analysis of sedimentation equilibrium data. Isothermal titration microcalorimetry (ITC) is a method for characterizing protein-protein and protein-ligand interactions by measuring the intrinsic heat (enthalpy) change of the reactions. UNIT 20.4 includes a basic protocol for determining observed binding parameters: Contributed by Paul T. Wingfield Current Protocols in Protein Science (2003) 20.0.1-20.0.2 Copyright © 2003 by John Wiley & Sons, Inc.
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protein (ligand) binding to a given protein, determination of binding affinity (Ka), and binding stoichiometry. A second protocol describes the determination of biochemical well-defined molecular binding parameters. This is experimentally more rigorous but can lead to an understanding of the binding molecular mechanism. Protein associations can also be measured by methods that require less sophisticated equipment than described in the previous units. One such method is large zone-analytical gel filtration chromatography originally pioneered by Ackers and co-workers. In UNIT 20.5 a variant of the original method is described, where high-performance liquid chromatography (HPLC) is used instead of low-pressure size exclusion chromatography (SEC). This modification results in great savings of both time and materials. Several light-scattering techniques have been described in detail in UNIT 7.8 and UNIT 8.3 deals with SEC. In UNIT 20.6, static light scattering is combined with SEC to allow the direct determination of the molecular weight of fractionated proteins. Protocols describe the application of this two-dimensional methodology for the determination of the mass of protein-protein and protein-carbohydrate complexes. The second basic type of experiment performed with the analytical ultracentrifuge is sedimentation velocity and this is the subject of UNIT 20.7 (the first type—sedimentation equilibrium—is discussed in UNIT 20.3). In sedimentation velocity, the concentration distribution of protein in an analytical centrifuge cell occurs as a function of both time and distance from the axis of rotation. Compared to equilibrium analysis, relatively high speeds are used which deplete solute near the cell meniscus, forming a boundary between buffer and protein solution. Analysis of the rate of boundary movement can yield information about molar masses of the sedimenting proteins as well as stoichiometries and equilibrium constants for their interactions. In future updates, experimental protocols will be provided for both modes of analytical centrifugation. Paul T. Wingfield
Introduction
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Overview of the Quantitation of Protein Interactions The biological function of many (and perhaps most) proteins involves reversible interactions with other proteins, nucleic acids, or other nonprotein ligands. Such interactions play many different roles in a wide range of cellular processes. A few examples are: (1) storing or transporting key metabolites (e.g., O2 storage by myoglobin); (2) forming and maintaining the quaternary structure of multisubunit enzymes; (3) specific binding and recognition events (antigen-antibody, hormone-receptor, transcription factor–promoter); and (4) self-assembly of large structures (e.g., microtubules or chromatin). Thus the quantitative characterization of such interactions represents an important part of understanding the function of such proteins and their role in these cellular events. The two fundamental properties that characterize any binding interaction are the stoichiometry (how many moles of each reactant combine to form the product) and the binding strength. The binding strength or “affinity” of the interaction is usually quantitatively expressed through either the equilibrium association constant, Ka, or its inverse, the dissociation constant, Kd, which for a reaction nA + mB ↔ AnBm are defined by:
Ka = Kd =
A n Bm A
n
B
m
A
n
B
m
A n Bm
The choice of using Ka or Kd is largely a matter of personal preference. The concentration terms in Equation 20.1.1 may be expressed in any concentration units, but molar units are generally preferred. The binding strength can also be characterized by the change in Gibbs free energy for the reaction, ∆G, which is related to the equilibrium constant through the relation ∆G = RT1n(Kd) = −RT1n(Ka), where R is the gas constant and T is the absolute temperature. It should always be remembered that such energy changes are not absolute, but are always relative to some reference state, which conventionally is taken as a 1 M solution of each reactant (and hence the preference for equilibrium constants in molar units).
Contributed by John S. Philo Current Protocols in Protein Science (1999) 20.1.1-20.1.13 Copyright © 1999 by John Wiley & Sons, Inc.
UNIT 20.1
The various experimental methods differ considerably in their ability to determine the stoichiometry and/or Kd as well as in the range of values they can measure. Table 20.1.1 lists the techniques that will be covered in this unit and summarizes the type of information each gives and the approximate range of Kd values it can measure. The good news is that there are often several of these methods that can be appropriately applied to a given system. Moreover, the continued emergence of new and improved instruments, software, and protocols broadens the choices and reduces the time, effort, and/or amount of material required for obtaining the desired information.
SOME COMMENTS ON TECHNIQUES Most of the methods in this unit detect binding interactions through a change in mass as binding occurs. In addition to mass spectrometry itself, this group includes sedimentation equilibrium (analytical and micropreparative), surface plasmon resonance (SPR), size-exclusion chromatography (SEC) with on-line light scattering, and chemical cross-linking. Techniques such as large-zone analytical SEC, sedimentation velocity, and some of the electrophoresis methods, while not strictly mass-based since the changes in hydrodynamic properties or mobility are related to changes in molecular shape as well as mass, can nonetheless reasonably be grouped with the true mass techniques. Changes in spectroscopic properties such as fluorescence polarization (Heyduk et al., 1996) may also be related to mass changes and hence certain spectroscopic approaches also belong in this group. As might be expected, methods that are members of this group are usually capable of determining stoichiometry, but some of these methods are not necessarily able to measure the equilibrium constant. For example, mass spectrometry, cross-linking, and light scattering/SEC are generally excellent at determining stoichiometry, but the amount of the highermass complex that they detect may not accurately reflect its proportion at equilibrium. It is also important to remember that some methods, such as sedimentation velocity and light scattering/SEC, involve dynamic physical separa-
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Micropreparative analytical ultracentrifugation (sedimentation equilibrium)
Change in solution mass Stoichiometry
SEC with light scattering Equilibrium dialysis and gel permeation, ultrafiltration, gel chromatography (Hummel-Dreyer) Large-zone analytical SEC Stoichiometry (Kd ∼10−4 to 10−13 M)
Change in hydrodynamic size
Change in solution mass Stoichiometry (Kd ∼10−3 to 10−13 M)
Stoichiometry (Kd ∼10−1 to 10−13 M)
Partitioning of free and bound ligand
Stoichiometry
Change in molecular mass
Chemical cross-linking
Stoichiometry
Binding properties measured (and range of Kd)
Change in molecular mass
Basis for detection
Mass spectrometry
Techniquea
30 min + readout
5 µld
100 µld
1000 µld
Harding et al., 1992; Wen et al., 1996 Andreu, 1985; Kido et al., 1985; Sophianopoulos and Sophianopoulos, 1985; Sebille et al., 1990 Ackers, 1973; Valdes and Ackers, 1979; Nenortas and Beckett, 1994
Loster and Josic, 1997; Kunkel et al., 1981
Loo, 1997; Roepstorff, 1997; Farmer and Caprioli, 1998
References
12 to 24 hr + Darawshe et al., 1993, readout (4 to 6 1995 samples per rotor)
30 to 60 min + readout
15 min to 24 hr + readout
30 to 60 min
15 min
0.001 µg; <1 µl
100 µg (20 kDa); 10 µg (200 kDa); 5 µl 10 µld
Typical analysis timec
Minimum sample mass or volume neededb
Table 20.1.1 Summary of Characteristics for Some Important Methods for Characterizing Protein Interactions
Overview of the Quantitation of Protein Interactions
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Wide range of potential on-line or off-line detection methods; may be labor-intensive depending on degree of automation Requires only preparative centrifuges; fewer data points and more labor-intensive than on-line instruments, but much greater specificity and sensitivity through off-line detection
May not detect weaker interactions; very helpful prelude to centrifugation methods, especially for glycoproteins Useful and fairly simple approach to determining quaternary structure and stoichiometry Gives true mass, independent of molecular shape “Low tech” and often labor-intensive, but still very useful and adaptable
Comments
Current Protocols in Protein Science
0.2 to 2 hr
3 hr
12 to 24 hr per rotor speed (9 to 21 samples) 2 to 5 hr (3 to 7 samples)
15 to 30 min (1 analyte, up to 4 surfaces)
—e
100 µg; 2000 µl
10 µg; 100 µl
10 µg; 400 µl
0.01 µg; 100 µl
Change in solution mass Stoichiometry (Kd ∼10−3 to 10−9 M)
Change in sedimentation coefficient
Change in mass bound to surface
Sedimentation equilibrium (in analytical ultracentrifuge) Sedimentation velocity (in analytical ultracentrifuge)
Surface plasmon resonance
Stoichiometry; kinetics (Kd ∼10−3-10−13 M)
Stoichiometry; conformational changes (Kd ∼10−3 to 10−9 M)
Myszka, 1997; Schuck, 1997; Morton and Myszka, 1998
Rivas and Minton, 1993; Laue, 1995; Schuster and Toedt, 1996 Schuster and Toedt, 1996; Lobert et al., 1997; Stafford, 1997
Bundle and Sigurskjold, 1994; Fisher and Singh, 1995; Doyle, 1997
Oberfelder and Lee, 1985; Heyduk et al., 1996; van Holde et al., 1998
Universal signal; works well for small molecules; useful additional thermodynamic information Powerful method when there is a significant change in mass; much more accessible with modern hardware and software Newer software makes data analysis fast and user-friendly; generally not as good as sedimentation equilibrium for Kd Broad range of Kd; high throughput; fairly insensitive to contaminants; potential problems from surface coupling
Applicability depends on nature of protein and/or ligand; chromophore label may be required
eRequired mass and volume depend on detection method.
dRequired mass depends on detection method; for radiolabeling, ELISA, or fluorescence detection the mass required may be extremely small.
cThis is the approximate time needed to carry out the experiment. For methods in which more than one sample may be run simultaneously, the number of samples is noted in parentheses. For methods in which the readout of concentration is performed separately (e.g., by ELISA or scintillation counting) the time for that step is not included.
have this volume of solution at a concentration high enough to saturate the interaction. These figures will vary by an order of magnitude or more depending on the equipment and exact protocol employed.
bApproximate minimum sample mass (detection limit) and minimum sample volume. For weaker interactions the sample requirement is often dictated more by the volume requirement, since it is necessary to
instrumentation.
aTechniques that primarily give information about stoichiometry are listed first, followed by those with good quantitation of binding affinity, with the latter arranged roughly in order of cost and complexity of the
Titration microcalorimetry
Change in spectroscopic Molar ratio property stoichiometry (possibly); conformational changes (Kd ∼10−5 to 10−11 M) Heat release or uptake Molar ratio; ∆H; ∆cp (Kd ∼10−3 to 10−9 M)
Spectroscopic methods
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tion of components and are not true equilibrium methods. With these methods, the results may depend on the rates of association-dissociation as well as the equilibrium constant. For relatively weak interactions with relatively fast kinetics, the band or boundary that is seen is a so-called “reaction boundary” and does not necessarily correspond to any single species or any particular stoichiometric complex. This complication does not apply to sedimentation equilibrium, even though there is some separation of components, because association-dissociation equilibrium is maintained at every position in the centrifuge cell. Large-zone analytical SEC also can be regarded as a true equilibrium technique because the concentration and composition of the plateau region matches what was present before the sample was loaded onto the column. One common drawback of these methods based on mass is that they may be insensitive to the binding of peptides or other ligands having a mass that is small compared to that of the protein. Under certain circumstances it may therefore be desirable to deliberately increase the mass of such ligands. For example, a peptide might be fused with a larger polypeptide or a small ligand modified by adding a defined polyethylene glycol polymer. The caveat here is, of course, that such modifications must not interfere with binding, so this approach usually requires knowledge that some region of the ligand is not important for binding.
Surface Plasmon Resonance
Overview of the Quantitation of Protein Interactions
SPR-based sensors have several unique characteristics relative to the other mass-based methods, which warrant some further discussion. First, SPR is unique among these methods in that the kinetics of the association and dissociation events can be measured explicitly, and this kinetic information can provide insights that may be difficult to obtain from equilibrium information alone. SPR actually provides two fairly different approaches to characterizing the binding affinity. In one approach, the equilibrium association constant is determined from the kinetic data by first determining the association and dissociation rates and then calculating their ratio. The alternative approach involves exposing the reactant solution to the measurement surface for a long time to allow the binding to reach equilibrium. The magnitude of this binding signal is then analyzed as a function of reactant concentration to obtain the equilibrium constant, in a fashion similar to other equilibrium methods.
Another significant difference in SPR is that one of the reactants is coupled to a surface. In principle, this should not matter, provided that this coupling does not produce steric hindrance or involve chemically modifying residues important for the interaction being studied. Random coupling methods, however, have the potential for producing a range of orientations and binding properties, so coupling via a specific site and/or in a specific orientation is preferred for detailed quantitative studies. Further, surface binding can be fundamentally different than solution binding if there is the potential for more than one molecule on the surface to simultaneously interact with a single analyte molecule in solution. A failure to properly consider and treat multivalent interactions on the surface can, and has, led to errors of many orders of magnitude in binding constants obtained by SPR (Kalinin et al., 1995; Ladbury et al., 1995). Thus independent knowledge of the reaction stoichiometry is quite helpful for the design and interpretation of SPR experiments. With current procedures (Schuck, 1997; Morton and Myszka, 1998; UNIT 20.2) such problems arising from multivalent interactions, as well a number of other potential artifacts in SPR, can usually be avoided. SPR is generally an excellent tool for studying protein-ligand interactions and interactions between two different proteins, but the required surface coupling limits its utility for characterizing protein self-associations. Lastly, it should not go unmentioned that one of the great virtues of SPR methods is that the amount of material required is usually quite small, particularly for the component bound to the measurement surface.
Analytical Ultracentrifugation (Sedimentation Equilibrium and Sedimentation Velocity) Once relegated to a few dedicated specialists, with improved hardware and automation this technique is again becoming widely used and more accessible to nonspecialists, and the quantity of sample required is also reduced. Among the strengths of analytical ultracentrifugation are that the governing principles are simple and well understood, and the theory and practice has been developed over many years. Sedimentation equilibrium has many virtues (and a long history of success) for detailed studies of complex interactions, among which are: (1) it is a true solution equilibrium method; (2) it can cover a broad range of concentrations (∼5 µg/ml to >100 mg/ml for
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proteins); (3) there are few restrictions on the choice of solvents; and (4) it can be applied over a fairly wide temperature range (0° to 40°C for the Beckman XL-I). Its primary drawback is that reaching equilibrium typically requires 12 to 24 hr for a 3-mm solution column, so the throughput is fairly low (but 9 to 21 samples can be run simultaneously), and some proteins are not sufficiently stable over these long time periods. The use of even shorter solution columns (<1 mm) with interference optics reduces equilibration time to a few hours or less, but results in fewer data points and less overall accuracy. This short-column approach is excellent for determining stoichiometry, for survey experiments, and for studying changes in weight-average mass as a function of ligand concentration or mixing ratios of different proteins. It should not be overlooked that sedimentation equilibrium experiments can be performed using preparative centrifuges, and this approach has advantages over the dedicated analytical instruments (Rivas and Minton, 1993). The collection of fractions at the end of the experiment allows any off-line method to be used to follow the concentration distribution, greatly extending the range of concentrations and specificity of detection methods. This also permits further separation of individual components by electrophoresis or chromatography prior to detection. Sedimentation velocity experiments require only 2 to 5 hr, and thus can be employed for labile proteins, and are now often done at protein concentrations of 10 to 100 µg/ml instead of the 2 to 10 mg/ml common in the past. Stafford’s dc/dt method of data analysis (Stafford, 1994) has been a boon to the field, and the stoichiometry of tight complexes can often be determined simply by observing the appearance of new peaks as reactants are mixed in various proportions (Stafford, 1997). Other recent developments in data analysis (Philo, 1994; Stafford, 1997; Schuck, 1998) now allow the simultaneous determination of diffusion coefficients along with the sedimentation coefficients, and thus calculation of the true mass, for nonassociating monomers as well as tightly associated oligomers or complexes, provided there is little dissociation during the experiment. In general, though, individual species or complexes are not resolved, and the data are analyzed to give an overall weight-average sedimentation coefficient that changes systematically as binding or self-association occurs. This latter approach is particularly useful for
studying ligand binding when that binding is linked to protein self-association, as exemplified by recent work on drug binding to tubulin by Lobert and Correia (Lobert et al., 1998). The fitting of weight-average sedimentation data to obtain Kd can be complicated by the fact that the sedimentation coefficients depend on shape and conformation as well as mass, and thus it is generally not possible to accurately predict the sedimentation coefficients of oligomers or protein-protein complexes, and these must be treated as additional adjustable fitting parameters. On the other hand, the sensitivity to conformation can allow studying the binding of even very small ligands when that binding is linked to a conformational change. The latest development in using sedimentation velocity for studying interactions is use of finite-element numerical methods to directly simulate and fit a binding model (Schuck, 1998; Stafford, 1999). This approach can potentially obtain information about kinetics as well as Kd.
Titration Microcalorimetry In titration microcalorimetry, interactions are detected through the heat that is released or absorbed as a binding event occurs. The fact that this heat of reaction signal is so universal, and does not require a significant change of mass or the presence of any chromophores, is one of the great strengths of this method. It is important to note that this method does not measure the true binding stoichiometry but rather measures only the molar ratio at which the reactants combine—i.e., it cannot distinguish a 1:1 from a 2:2 stoichiometry. This method can be applied to self-associating systems by detecting dissociation as the protein is diluted into buffer, but the limited concentration range, and lack of information about true stoichiometry, limit its utility in this area. Titration calorimetry is often the method of choice for binding of low-molecular-weight ligands, particularly when the affinity is fairly low. Moreover, it is unique in directly giving information about the binding enthalpy, ∆H, in addition to measuring equilibrium constants and molar ratios. Another unique aspect of this technique is that the proton binding or release associated with a protein-protein or protein-ligand interaction can be quantitated simply by changing buffer systems (Doyle et al., 1995; Baker and Murphy, 1996). On the other hand, phenomena other than the binding event of interest can sometimes generate significant amounts of heat, so titration calorimetry also has a unique set of arti-
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facts that must be avoided. Among the potential problems are heat generated by small differences in pH or salt concentration between the titrant and the solution being titrated, and aggregation and precipitation of proteins induced by the stirring of the calorimeter cell. The heat due to solvent differences can of course be eliminated by thorough dialysis, but when studying binding of low-molecular-weight compounds, dialysis is often not possible. Limited solubility of the reactants can also create difficulties and force the use of large injection volumes, which generally compromises the signal/noise. Overall, though, this technique can be very useful, can give highly precise data, and the instrumentation and ease of use continues to improve.
Methods Based on Separation or Partitioning
Overview of the Quantitation of Protein Interactions
The glamour and popularity of the some of the “high-tech,” instrumentation-intensive methods should not overshadow a number of relatively simple and inexpensive approaches for studying protein interactions. For proteinligand interactions, equilibrium dialysis or ultrafiltration often work well, and are still primary methods for studying drug binding to serum proteins (Sophianopoulos and Sophianopoulos, 1985; Sebille, 1990; Sebille et al., 1990). Gel filtration media can be used in either batch mode (Kido et al., 1985), for smallzone chromatographic analysis (HummelDreyer analysis and its variants; Hummel and Dreyer, 1962; Andreu, 1985), or for large-zone (frontal) chromatography (Valdes and Ackers, 1979; Nenortas and Beckett, 1994). Virtually any method with good precision can be used to monitor the concentration of the protein or ligand following separation (or on-line monitoring may be used for chromatography), allowing great flexibility and an enormous concentration range. A common problem with these approaches is that the ligand or protein may bind to the membranes or resins. Furthermore, since transport is involved, one must be cautious about whether the measurements reflect equilibrium rather than kinetic properties. The material requirements may also be high unless some effort is devoted to keeping volumes small, and these approaches can be fairly labor intensive. Additional separation-based methods for protein-ligand interactions that have been used successfully are steady-state and counterion electrophoresis and isoelectric focusing (Oberfelder and Lee, 1985) and affin-
ity-chromatography approaches (Winzor and De, 1989).
THE IMPORTANCE OF DATA ANALYSIS METHODS Another common theme among these techniques is the central role of numerical methods for analysis and interpretation of the data, and this aspect has proven daunting for some researchers. Fortunately, today’s desktop computers allow very sophisticated analyses to be done quickly and routinely, and the software has become increasingly user-friendly. Equally important, the data now are often acquired directly in digital form, and on-line computerized instrument control permits unattended operations and more complex data-acquisition protocols. To see the tremendous impact the sum of all these phenomenon can have, one need look no further than at analytical ultracentrifugation. With new computerized instrumentation, formerly onerous data acquisition and analysis chores now take only seconds, and new numerical methods for analysis have added new capabilities and approaches. The net result is that a field that had all but disappeared has been rejuvenated. In the area of data analysis, one frequently encounters the need to formulate specific models for use in analyzing the data. For example, protein self-association studies by large-zone SEC or sedimentation equilibrium are typically fitted to a specific assembly pathway such as monomer ↔ dimer ↔ tetramer. In such situations, it is always important to critically analyze whether the right model has been chosen, and to try to get independent information to support that choice of model. In this regard, information about the reaction stoichiometry (or at least the limiting assembly states) is often especially valuable. Another important phenomenon in data analysis is the more widespread use of “global fitting” techniques. In global fitting, multiple experiments taken under different conditions (and possibly by different techniques) are analyzed simultaneously, using a common set of parameters to describe the molecular interactions. Such global analysis provides a much more robust analysis, and often allows more information to be extracted from a given set of experiments. This approach is fairly commonly used for certain techniques (e.g., analytical ultracentrifugation and SPR data), but is still seldom used for data from more than one type of instrument.
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SENSITIVITY TO CONTAMINANTS AND PROTEIN QUALITY In general, the acquisition of accurate quantitative data on protein interactions will require highly purified, fully active protein samples, but a few of these techniques can be quite insensitive to the presence of contaminants or other proteins present as carriers. This is an area where the SPR sensors particularly excel, and titration calorimetry is also fairly insensitive to contaminants or carrier proteins. Unfortunately, it is not uncommon for even highly purified protein samples to contain some inactive or partially active material, and further degradation may occur during the course of the measurements. This will usually limit the accuracy and reproducibility of the quantitative results, and possibly may even lead to false interpretations of the data. For example, proteins normally found as tightly associated oligomers often contain a small fraction of partially-denatured monomers that do not self-associate, or at least do so very weakly. Such species are often termed “incompetent monomers” in the analytical ultracentrifugation community. Suppose one is trying to quantitate the reversible dissociation of this oligomer at low concentrations in order to establish whether the oligomer exists at physiologically relevant concentrations. The presence of any incompetent monomer can result in a large increase in the apparent Kd for the dissociation when little reversible dissociation occurs over the experimentally accessible concentration range. Thus, extrapolation of such data to a low physiological concentration could lead to a false conclusion about the physiologically relevant quaternary structure. Similarly, titration calorimetry is sometimes employed to study interactions with dissociation constants in the nanomolar range under conditions where the reactant concentrations are two to three orders of magnitude higher than the Kd. Under such conditions, it may be very difficult to distinguish a sample in which the majority of the protein binds extremely tightly (outside the range of the instrument) but a small portion of the protein binds more weakly, from one where 100% of the protein has a Kd that is close to the measurement limit of the instrument. Such potential artifacts from inactive or partially active species can, of course, be detected through inconsistencies in the apparent Kd values for samples studied at different concentrations. That is precisely why it is often important
to incorporate data from multiple experiments into the data analysis, and to cover as wide a portion of the binding isotherm as possible. Overall, it is always advantageous to have highly pure, well-characterized, and stable protein samples. Before embarking on a detailed analysis of a complex set of protein interactions, it may be wise to devote some further effort to purification and to finding solvent conditions that promote stabilization. For proteins with poor stability, it will also be prudent to choose methods that are more rapid, and perhaps to work at lower temperatures.
THE ADVENT OF RECOMBINANT PROTEINS ENHANCES OUR CAPABILITIES FOR STUDYING PROTEIN INTERACTIONS Biotechnology has had a profound influence on many areas of science and medicine, including the study of protein interactions. Many of the proteins being studied today are produced by recombinant technology, and would otherwise be unavailable in sufficient quantity (or totally unknown if they have been found through genomic or proteomic techniques). The quality of recombinant proteins (purity and homogeneity) can also often be superior to those from natural sources, in part because starting purification with larger amounts of protein allows more extensive purification protocols and more rigorous exclusion of side fractions. Aside from issues of quality and quantity, a major virtue of recombinant proteins is the ability to engineer them in ways helpful for interaction studies. This is particularly useful for adding new, specific chemical groups such as cysteines for purposes of specifically immobilizing (e.g., SPR) or tagging the protein (e.g., fluorophores or isotopes). Such engineering may also involve adding new sequence motifs such as epitopes for commercially available antibodies or even entire functional domains such as green fluorescent protein. Protein engineering can also be useful in enhancing certain physical properties of the protein. Among these are improving stability or solubility, removal of sites leading to heterogeneity (N-terminal processing, glycosylation), or even removing flexible loops. While some of these aspects may be more of interest in obtaining proteins better suited for highresolution structural determination, lowering the chemical and structural heterogeneity may also be desirable before undertaking extensive interaction analysis of complex systems.
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Last, but certainly not least, interaction studies done with site-directed mutants and/or proteins with deletions of individual functional domains allow a much more detailed interpretation of the interaction data and strongly enhance the analysis of structure/function relations.
THE VIRTUES OF USING MULTIPLE TECHNIQUES
Overview of the Quantitation of Protein Interactions
Seldom can any one method give complete and fully reliable information about protein interactions. Each method has certain strengths and weaknesses, and may depend on certain models and assumptions. Therefore it is highly desirable, and sometimes necessary, to use multiple techniques in studying any protein interaction. Fortunately many of the techniques covered in this unit are highly complementary to one another, and this is definitely a field where the whole is greater than the sum of its parts. It is always important to characterize the state of association of the protein(s) being studied. Such studies of self-association may be either the desired endpoint of an interaction analysis, or only preliminary to studying a protein-ligand interaction or the interactions between different proteins. Mass spectrometry provides, at the very least, an excellent beginning and foundation for other methods of studying self-association by verifying the covalent structure and determining an accurate monomer mass, and may also provide information about the presence of higher oligomers or binding of metal ions or other ligands (Anderegg et al., 1997). Light scattering/SEC is also very useful (and very quick) for determining whether the predominant form is a monomer, dimer, or other oligomer. While this is not usually a good method for directly measuring the equilibrium constants for self-associating systems, it can be ver y h elp fu l in id en tify ing limiting stoichiometries, and may identify other assembly intermediates or the presence of incompetent monomer. Light scattering/SEC therefore can be an excellent prelude or adjunct to sedimentation equilibrium or large-zone analytical SEC (often the best methods for fully characterizing a self-associating system) by providing independent information about appropriate assembly models. In principle the combination of on-line scattering with large-zone SEC should make an excellent tool for measuring binding affinities of mixed- or self-associations, but this has not yet been extensively exploited, and the sensitivity of the scattering detectors may not
be sufficient for studying stronger interactions (Kd < 10−7 to 10−8 M). A wide variety of methods can be applied to interactions between different proteins or protein-DNA interactions (so-called “mixed” or “heterogeneous” associations), and many are complementary rather than redundant. Once again a knowledge of stoichiometry can be very helpful in designing experiments and selecting proper models for data analysis, and mass spectrometry, light scattering/SEC, and chemical cross-linking can be very helpful in this regard. In many cases, several techniques may be appropriate for determining the Kd, and if so it is certainly helpful to use more than one, particularly for complex associations or when the information is critical. One field where there has been a particularly nice synergy between methods such as sedimentation equilibrium, SPR, light scattering/SEC, and titration calorimetry has been in studies of interactions between protein hormones and their receptors (Ward et al., 1994; Zhang et al., 1997; Philo and Hensley, 1998). By confining receptors to the surface of SPR sensors it has also been possible to more closely mimic the situation when the receptor is embedded in the cell membrane, and even multisubunit receptor complexes have been assembled on the sensor surface (Kalinin et al., 1995; Myszka et al., 1996). Comparison of such data with those from solution methods such as sedimentation equilibrium or titration calorimetry may allow a more thorough understanding of the importance of having the hormone interact simultaneously with multiple receptor subunits, and determination of the energetic consequences of having the receptors preoriented in a fashion enabling such interactions.
SOME THOUGHTS ON CHOOSING METHODS As mentioned earlier, the fact that most of the methods are based on changes in mass often limits their application to studying binding of low-molecular-weight compounds, so for such materials, titration calorimetry or a “low-tech” approach like equilibrium dialysis may be particularly advantageous. For protein-protein or protein-DNA interaction studies, the choice of method(s) for determining the Kd will often be dictated by the magnitude of that Kd. Both titration calorimetry and sedimentation equilibrium lack the sensitivity to measure Kd values below ∼10−8 to 10−9 M. Stronger interactions with Kd values down to ∼10−12 M can be studied by SPR or fluorescence techniques. By
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coupling them to highly sensitive radiometric, fluorometric, or ELISA assays, techniques such as equilibrium dialysis, ultrafiltration, micropreparative analytical ultracentrifugation, or large-zone analytical SEC can also study Kd values in the picomolar range or lower. At the other extreme, weak interactions with Kd values up to ∼10−3 M can be measured by analytical ultracentrifugation, titration calorimetry, SPR, equilibrium dialysis, or ultrafiltration. With very weak interactions, reactant solubility may limit the application of titration calorimetry, while nonspecific binding to the surfaces can be problematic for SPR, dialysis, and ultrafiltration. In other cases the choice of technique to apply to obtain the Kd of a mixed or self-association may be dictated by the amount of material that is available. For the less sensitive techniques such as sedimentation equilibrium and titration calorimetry, where this is most likely to be an issue, as a rule of thumb the minimum amount of protein needed to run one sample with good signal/noise ratio is ∼10 µg for sedimentation equilibrium (with a 3-mm column), and ∼100 µg for titration calorimetry. However, the amount of material needed is often dictated not by the inherent detection limits of a particular instrument, but rather by the necessity to run at least one sample at a high enough concentration to promote essentially full association (i.e., 10 times the Kd or preferably higher). Thus, the material requirement is often proportional to the product of the Kd and the minimum volume needed for this high-concentration sample. For the more sensitive methods, this volume requirement may actually be dictated more by sample-handling considerations than by how much material actually enters the instrument. For sedimentation equilibrium, the minimum volume is ∼100 µl, for sedimentation velocity ∼400 µl, and for titration calorimetry ∼2 ml. In SPR the material requirements for the reactant that will be coupled to the sensor surface is quite small; material requirements are usually only a potential concern for the reactant in solution. For flow-based SPR sensors the minimum volume for the solution reactant depends on flow rate and injection length, which in turn vary with the kinetics of the process and whether the instrument is operated in a kinetic or equilibrium-binding mode, but this volume is typically ∼100 µl. In large-zone SEC, the volume required depends on the column volume but is often several milliliters (or more).
OBTAINING ADDITIONAL THERMODYNAMIC INFORMATION Additional insight into protein interactions can be obtained by extracting solution thermodynamic quantities from the experimental data. As mentioned previously, titration calorimetry measures the binding enthalpy, ∆H, directly (and this quantity can be measured easily even when the Kd is too low to be measured). When the Kd is also known, the entropy change can then be calculated from the relation ∆G = RT1n(Kd) = ∆H − T∆S. The magnitudes of ∆H and ∆S may provide some insight into the nature of the interaction (e.g., whether it is predominantly hydrophobic or electrostatic). However these net quantities are sums of contributions from many effects, and are often dominated by interactions with the solvent, so structural interpretation is risky at best. A more promising approach to structural interpretation of the thermodynamic data comes from examining the change in heat capacity, ∆cp. This can be obtained by measuring the calorimetric ∆H at several temperatures and taking the slope of a plot of ∆H versus T, through the relation ∆cp = δ(∆H)/δT. This ∆cp can then be interpreted in structural terms, as either a change in buried surface area when the complex is formed (Murphy and Freire, 1992) or as the number of amino acid residues that go from a disordered to an ordered state as a consequence of the interaction (Spolar and Record, 1994). The apparent, or van’t Hoff, enthalpy change, ∆HvH, can also be obtained by measuring the Kd as a function of temperature (using any method) if it is assumed that ∆H and ∆S are constant with temperature (at least over some narrow range). This is done by plotting ln(Kd) versus 1/T (a van’t Hoff plot), taking the slope of a line through the data, and using the relation ∆HvH = Rδ(1nKd)/δ(1/T). The calorimetric and van’t Hoff enthalpies may differ significantly. The calorimetric ∆H includes all heats associated with the overall biochemical reaction, whereas ∆HvH includes only those reactions that directly influence the Kd, and thus may not include heats associated with solvation, ionization, or conformational changes if those processes have no net effect on the binding free energy (Bundle and Sigurskjold, 1994). It is also possible to take advantage of the temperature dependence of equilibrium constants to extend the range of affinities that can be covered by any given method. Since most reactions are exothermic (negative ∆H), they
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become weaker as the temperature is increased. Thus for studying very strong affinities it may be helpful to determine the Kd at high temperature and then correct this value to lower temperature using measured values for ∆H (and preferably ∆cp also). This approach has been used, for example, to study high-affinity antigen-antibody binding (Schwarz et al., 1995).
THERMODYNAMIC LINKAGE RELATIONSHIPS The concept of thermodynamic linkage has provided a powerful tool for probing and understanding protein interactions, and also serves as a basis for extending the range of affinities that can be probed. Consider first a protein, P, that can bind a ligand, L, and that also self-associates to form dimer. The ligand binding and association reactions form the thermodynamic cycle, illustrated in Figure 20.1.1. Because the total change in free energy must be independent of the pathway, if the dimer binds ligand with a different affinity than the monomer, it must also be true that the liganded form dimerizes differently than the unliganded form, such that ∆GL − ∆GLd = ∆GdL − ∆Gd (or equivalently KL/KLd = KdL/Kd). How does this exercise help us? Suppose the dimer binds ligand more strongly than monomer, so strongly that we cannot measure the affinity. This linkage relationship says that if we can determine the (lower) affinity of the monomer, we need only measure the monomerdimer dissociation equilibrium of the unliganded and liganded forms, and we may then calculate the ligand affinity of the dimer. This
P←
KL
K dL
Kd P2 ←
Overview of the Quantitation of Protein Interactions
←PL
K Ld
←(PL)2
Figure 20.1.1 Thermodynamic cycle for ligand binding and association reactions. KL and KLd are the association constants for ligand binding for the monomer and dimer, respectively, and Kd and KdL are the monomer-dimer association constants for the unliganded and liganded form of the protein, with corresponding free energies ∆GL, ∆GLd, ∆Gd, and ∆GdL.
very simple but powerful idea has been exploited with tremendous success, for example, by Gary Ackers and his co-workers in detailed studies of oxygen binding and subunit assembly for hemoglobin (Turner et al., 1981; Ackers, 1998), in probing protein-DNA interactions (Koblan and Ackers, 1991), and in many other systems. These same linkage concepts can also be applied when the protein’s affinity for a ligand varies with pH, ions such as calcium, or effectors such as nucleotide phosphates, rather than through self-assembly. For example, since titration calorimetry can fairly easily quantitate proton uptake and release, a reaction whose affinity is too high to be studied at neutral pH can be studied at a pH where the binding is weaker, and then corrected to neutral pH conditions using the linkage relations and the difference in proton binding (Doyle et al., 1995; Baker and Murphy, 1996).
LITERATURE CITED Ackers, G.K. 1973. Studies of protein ligand binding by gel permeation techniques. Methods Enzymol. 27:441-455. Ackers, G.K. 1998. Deciphering the molecular code of hemoglobin allostery. Adv. Protein Chem. 51:185-253. Anderegg, R.J., Wagner, D.S., Blackburn, R.K., Opiteck, G.J., and Jorgenson, J.W. 1997. A multidimensional approach to protein characterization. J. Prot. Chem. 16:523-526. Andreu, J.M. 1985. Measurement of protein-ligand interactions by gel chromatography. Methods Enzymol. 117:346-354. Baker, B.M. and Murphy, K.P. 1996. Evaluation of linked protonation effects in protein binding reactions using isothermal titration calorimetry. Biophys. J. 71:2049-2055. Bundle, D.R. and Sigurskjold, B.W. 1994. Determination of accurate thermodynamics of binding by titration microcalorimetry. Methods Enzymol. 247:288-305. Darawshe, S., Rivas, G., and Minton, A.P. 1993. Rapid and accurate microfractionation of the contents of small centrifuge tubes: Application in the measurement of molecular weight of proteins via sedimentation equilibrium. Anal. Biochem. 209:130-135. Darawshe, S., Merezhinskaya, N., and Minton, A.P. 1995. PhosphorImager enhancement of sedimentation equilibrium–quantitative polyacrylamide gel electrophoresis: A highly sensitive technique for quantitation of equilibrium gradients of individual components in mixtures. Anal. Biochem. 229:8-14. Doyle, M.L. 1997. Characterization of binding interactions by isothermal titration calorimetry. Curr. Opin. Biotechnol. 8:31-35.
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Doyle, M.L., Louie, G., Dal Monte, P.R., and Sokoloski, T.D. 1995. Tight binding affinities determined from thermodynamic linkage to protons by titration calorimetry. Methods Enzymol. 259:183-194. Farmer, T.B. and Caprioli, R.M. 1998. Determination of protein-protein interactions by matrix-assisted laser desorption/ionization mass spectrometry. J. Mass Spectrom. 33:697-704. Fisher, H.F. and Singh, N. 1995. Calorimetric methods for interpreting protein-ligand interactions. Methods Enzymol. 259:194-221. Harding, S.E., Rowe, A.J., and Horton, J.C. 1992. Analytical Ultracentrifugation in Biochemistry and Polymer Science. Royal Society of Chemistry, Cambridge, U.K. Heyduk, T., Ma, Y., Tang, H., and Ebright, R.H. 1996. Fluorescence anisotropy: Rapid, quantitative assay for protein-DNA and protein-protein interaction. Methods Enzymol. 274:492-503. Hummel, J.P. and Dreyer, W.J. 1962. Measurement of protein-binding phenomena by gel filtration. Biochim. Biophys. Acta 63:530-532. Kalinin, N.L., Ward, L.D., and Winzor, D.J. 1995. Effects of solute multivalence on the evaluation of binding constants by biosensor technology: Studies with concanavalin A and interleukin-6 as partitioning proteins. Anal. Biochem. 228:238244. Kido, H., Vita, A., and Horecker, B.L. 1985. Ligand binding to proteins by equilibrium gel penetration. Methods Enzymol. 117:342-346. Koblan, K.S. and Ackers, G.K. 1991. Energetics of subunit dimerization in bacteriophage lambda cI repressor: Linkage to protons, temperature, and KCl. Biochemistry 30:7817-7821. Kunkel, G.R., Mehrabian, M., and Martinson, H.G. 1981. Contact-site cross-linking agents. Mol.Cell. Biochem. 34:3-13. Ladbury, J.E., Lemmon, M.A., Zhou, M., Green, J., Botfield, M.C., and Schlessinger, J. 1995. Measurement of the binding of tyrosyl phosphopeptides to SH2 domains: A reappraisal. Proc. Natl. Acad. Sci. U.S.A. 92:3199-3203. Laue, T.M. 1995. Sedimentation equilibrium as thermo dy namic to ol. Methods Enzymol. 259:427-452. Lobert, S., Boyd, C.A., and Correia, J.J. 1997. Divalent cation and ionic strength effects on vinca alkaloid–induced tubulin self-association. Biophys. J. 72:416-427. Lobert, S., Ingram, J.W., Hill, B.T., and Correia, J.J. 1998. A comparison of thermodynamic parameters for vinorelbine and vinflunine-induced tubulin self-association by sedimentation velocity. Mol. Pharmacol. 53:908-915. Loo, J.A. 1997. Studying noncovalent protein complexes by electrospray ionization mass spectrometry. Mass Spectrom. Rev. 16:1-23. Loster, K. and Josic, D. 1997. Analysis of protein aggregates by combination of cross-linking re-
actions and chromatographic separations. J. Chromatogr. B. Biomed. Sci. Appl. 699:439-461. Morton, T.A. and Myszka, D.G. 1998. Kinetic analysis of macromolecular interactions using surface plasmon resonance biosensors. Methods Enzymol. 295:268-294. Murphy, K.P. and Freire, E. 1992. Thermodynamics of structural stability and cooperative folding behavior in proteins. Adv. Protein Chem. 43:313361. Myszka, D.G. 1997. Kinetic analysis of macromolecular interactions using surface plasmon resonance biosensors. Curr. Opin. Biotechnol. 8:50-57. Myszka, D.G., Arulanantham, P.R., Sana, T., Wu, Z.N., Morton, T.A., and Ciardelli, T.L. 1996. Kinetic analysis of ligand binding to interleukin2 receptor complexes created on an optical biosensor surface. Protein Sci. 5:2468-2478. Nenortas, E. and Beckett, D. 1994. Reduced-scale large-zone analytical gel filtration chromatography for measurement of protein association equilibria. Anal. Biochem. 222:366-373. Oberfelder, R.W. and Lee, J.C. 1985. Measurement of ligand-protein interaction by electrophoretic and spectroscopic techniques. Methods Enzymol. 117:381-399. Philo, J.S. 1994. Measuring sedimentation, diffusion, and molecular weights of small molecules by direct fitting of sedimentation velocity concentration profiles. In Modern analytical ultracentrifugation (T.M. Schuster and T.M. Laue, eds.) pp. 156-170. Birkhauser, Boston. Philo, J.S. and Hensley, P. 1998. Integrating analytical ultracentrifugation with other experimental approaches: A case study on erythropoietin interactions with its receptor. Chemtracts 11:969979. Rivas, G. and Minton, A.P. 1993. New developments in the study of biomolecular associations via sedimentation equilibrium. Trends. Biochem. Sci. 18:284-287. Roepstorff, P. 1997. Mass spectrometry in protein studies from genome to function. Curr. Opin. Biotechnol. 8:6-13. Schuck, P. 1997. Reliable determination of binding affinity and kinetics using surface plasmon resonance biosensors. Curr. Opin. Biotechnol. 8:498502. Schuck, P. 1998. Sedimentation analysis of noninteracting and self-associating solutes using numerical solutions to the Lamm equation. Biophys. J. 75:1503-1512. Schuster, T.M. and Toedt, J.M. 1996. New revolutions in the evolution of analytical ultracentrifugation. Curr. Opin. Structur. Biol. 6:650-658. Schwarz, F.P., Tello, D., Goldbaum, F.A., Mariuzza, R.A., and Poljak, R.J. 1995. Thermodynamics of antigen-antibody binding using specific antilysozyme antibodies. Eur. J. Biochem. 228:388394.
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Sebille, B. 1990. Methods of drug protein binding determinations. Fund. Clin. Pharmacol. 4 Suppl 2:151s-161s.
More advanced texts; coverage of binding equilibria and thermodynamics, spectroscopy, and centrifugation methods.
Sebille, B., Zini, R., Madjar, C.V., Thuaud, N., and Tillement, J.P. 1990. Separation procedures used to reveal and follow drug-protein binding. J. Chromatogr. 531:51-77.
Freifelder, D. 1982. Physical Biochemistry: Applications to Biochemistry and Molecular Biology. W.H. Freeman, New York.
Sophianopoulos, A.J. and Sophianopoulos, J.A. 1985. Ultrafiltration in ligand-binding studies. Methods Enzymol. 117:354-370. Spolar, R.S. and Record, M.T., Jr. 1994. Coupling of local folding to site-specific binding of proteins to DNA [see comments]. Science 263:777784. Stafford, W.F., III. 1994. Boundary analysis in sedimentation velocity experiments. Method Enzymol. 240:478-501.
Johnson, M.L. and Frasier, S.G. 1985. Nonlinear least-squares analysis. Methods Enzymol. 117:301-342. Good overview of the fitting of experimental data (a key part of all protein interaction studies), with some examples for ligand binding. van Holde et al., 1998. See above.
Stafford, W.F., III. 1997. Sedimentation velocity spins a new weave for an old fabric. Curr. Opin. Biotechnol. 8:14-24.
More advanced texts; coverage of binding equilibria and thermodynamics, spectroscopy, and centrifugation methods.
Stafford, W.F., III. 1999. Analysis of reversibly interacting macromolecular systems by time derivative sedimentation velocity. Methods Enzymol. (In Press)
Winsor, D.J. and Sawyer, W.H. 1995. Quantitative characterization of ligand binding. John Wiley & Sons, New York.
Turner, B.W., Pettigrew, D.W., and Ackers, G.K. 1981. Measurement and analysis of ligandlinked subunit dissociation equilibria in human hemoglobins. Methods Enzymol. 76:596-628. Valdes, R.J. and Ackers, G.K. 1979. Study of protein subunit association equilibria by elution gel chromatography. Methods Enzymol. 61:125412. van Holde, K.E., Johnson, W.C., Jr., and Ho, P.S. 1998. Principles of Physical Biochemistry. Prentice-Hall, Upper Saddle River, N.J.. Ward, L.D., Howlett, G.J., Discolo, G., Yasukawa, K., Hammacher, A., Moritz, R.L., and Simpson, R.J. 1994. High affinity interleukin-6 receptor is a hexameric complex consisting of two molecules each of interleukin-6, interleukin- 6 receptor, and gp-130. J. Biol. Chem. 269:2328623289. Wen, J., Arakawa, T., and Philo, J.S. 1996. Size-exclusion chromatography with on-line light-scattering, absorbance, and refractive index detectors for studying proteins and their interactions. Anal. Biochem. 240:155-166. Winzor, D.J. and De, J.J. 1989. Biospecific interactions: Their quantitative characterization and use for solute purification. J. Chromatogr. 492:377430. Zhang, J.G., Owczarek, C.M., Ward, L.D., Howlett, G.J., Fabri, L.J., Roberts, B.A., and Nicola, N.A. 1997. Evidence for the formation of a heterotrimeric complex of leukaemia inhibitory factor with its receptor subunits in solution. Biochem. J. 325:693-700.
KEY REFERENCES Overview of the Quantitation of Protein Interactions
Good introductory text; strong on spectroscopy and centrifugation methods
Cantor, C.R. and Schimmel, P.R. 1980. Biophysical chemistry. Part II: Techniques for the study of biological structure and function. W.H. Freeman, San Francisco.
An excellent but heavily mathematical text covering theory, data analysis, and interpretation. Methods based on phase separation, spectral changes, affinity and gel chromatography, and competitive binding are discussed in some detail, with some coverage of sedimentation, calorimetry, and other techniques. An entire chapter is devoted to DNA-ligand interactions (also see APPENDIX 5A).
INTERNET RESOURCES [email protected] (subscriptions) [email protected] (to post questions) Reversible Associations in Structural and Molecular Biology (RASMB) e-mail list server. This e-mail group list is for discussions and questions regarding the study of reversible interactions. The dominant focus is analytical ultracentrifugation (and nearly all the experts participate), but tremendous expertise in many other relevant areas is represented as well. http://www.bbri.org/RASMB/rasmb.html Reversible Associations in Structural and Molecular Biology (RASMB) Web site. Distributes public domain software for analytical ultracentrifugation. http://www.cauma.uthscsa.edu Distributes public domain software for analytical ultracentrifugation. Also maintains an archive of the RASMB discussions and tutorials on sedimentation methodology. http://www.hci.utah.edu/groups/interaction/index. htm This site by the Protein Interaction Facility at the Huntsman Cancer Institute nicely describes the basics of SPR, analytical ultracentrifugation, and titration calorimetry, and is the distribution point for the CLAMP package of public-domain SPR analysis software.
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http://www.beckmancoulter.com/beckman/biorsr ch/prodinfo/xla/xlaprod.asp Analytical ultracentrifugation products from Beckman Coulter. Useful application notes and bibliographies.
http://www.panvera.com/ls/fpabout.html Panvera Web site (fluorescence polarization). Useful application notes and bibliographies. http://www.wyatt.com
http://www.calscorp.com
Wyatt Technologies Web site (light scattering). Useful application notes and bibliographies.
Calorimetry Sciences Web site. Useful calorimetry application notes and bibliographies.
http://www.biacore.com
http://www.microcalorimetry.com
BIAcore Web site (SPR). Useful application notes and bibliographies.
Microcal Web site. Useful calorimetry application notes and bibliographies.
Contributed by John S. Philo Alliance Protein Laboratories Thousand Oaks, California
Quantitation of Protein Interactions
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Measuring Protein Interactions by Optical Biosensors
UNIT 20.2
Optical evanescent wave biosensors, such as surface plasmon resonance, resonant mirror, or waveguide biosensors, have been increasingly used during the last several years for the characterization of protein-protein interactions (Schuck, 1997a; Margulies et al., 1993, 1996; van der Merwe and Barclay, 1996; Malmborg and Borrebaeck, 1995; Garland, 1996). This includes both the determination of equilibrium binding constants, which reveal the magnitude of the binding energy, as well as the determination of the bimolecular rate constants describing the kinetics of the interaction. Studying protein interactions at a surface, as opposed to solution methods, has at least two fundamental advantages. First, the magnitude of the signal involved is independent of the affinity of the interaction over a wide range. Because one binding partner is confined to the surface (in the following referred to as ligand), the number of free binding sites that are under observation in an experiment is determined predominantly by the surface immobilization procedure (requiring usually only very small amounts of material). Therefore, the sensitivity of the method is limited essentially by the availability of the soluble binding partner (the analyte) in a well-defined state, at concentrations in the order of the inverse affinity constant. This allows studies of interactions with affinities spanning a range of at least 105 to 1010 M–1, and makes this method very attractive, in particular, for high-affinity interactions. Second, the separation of the surface-immobilized sites from the mobile binding partners in solution, if combined with an appropriate fluidic handling system, is virtually ideally suited for the real-time observation of the kinetics of the binding processes. Chemical on-rate constants of up to 105/Msec to 106/Msec and off-rate constants of between 10–5/sec and 10–1/sec are generally accessible. Information on the chemical on-rate constants, and on the lifetime of the complexes formed, can be of great value, for example, in the study of interactions involved in cellular signaling processes. However, these fundamental advantages are gained at the expense of additional experimental difficulties intrinsic to surface binding studies. These include the need for immobilization of one reactant at a relatively high local surface density (and the execution of binding studies under these conditions), the problems of nonspecific surface binding, possible multiple attachments to the surface through multivalent interactions, the problem of mass transport in kinetic experiments, and finally the possibility of surface potentials influencing the thermodynamics of the interaction (Leckband, 1997; Leckband et al., 1995). Although the different commercial instruments for optical biosensing are based on slightly different physical principles, they all exploit surface-confined electromagnetic fields for the precise and real-time measurement of the refractive index of the medium in the immediate vicinity of the sensor surface (Garland, 1996; Knoll, 1998; Lukosz, 1991). When ligands are immobilized to the sensor surface, reversibly interacting analytes will accumulate at the sensor surface when brought into the solution above. By virtue of the difference of the refractive index increments of proteins and most aqueous buffer solutions, an increase of the total refractive index at the surface will then be reported (Fig. 20.2.1). Removal of the analyte from the solution causes dissociation of the reversibly bound molecules from the surface sites, which is accompanied by a decrease in the refractive index at the sensor surface, generating a decrease in the signal. Since the changes in signal are, to a good approximation, proportional to the changes in the mass of surface-bound analyte, the steady-state signals can be analyzed in terms of the law of Contributed by Peter Schuck, Lisa F. Boyd, and Peter S. Andersen Current Protocols in Protein Science (1999) 20.2.1-20.2.22 Copyright © 1999 by John Wiley & Sons, Inc.
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mass action and Langmuir isotherms. The time course of the signal can be interpreted in the context of chemical binding kinetics. This basic principle, and consequently the derived strategies for studying protein-protein interactions, as well as most of the experimental procedures and potential complications, are the same in the different commercial instruments. Among the manufacturers are Biacore (http://www.biacore.com), Affinity Sensors (http://www.affinity-sensors.com), Intersens Instruments (Amersfoort, Netherlands), BioTul (http://www.biotul.com), and Artificial Sensing In-
A
analyte flow
cA
n surf
B
buffer flow c A= 0
C
(B) c A=0
n surf
(A) c A>0
regeneration
n surf
Time
Measuring Protein Interactions by Optical Biosensors
Figure 20.2.1 Schematic presentation of a typical optical biosensor experiment. Light is coupled into a structure that allows generation of surface-confined electromagnetic waves (e.g., surface plasmons in a gold film, or modes of a planar waveguide), which are sensitive to the refractive index of the solution in the range of the evanescent field, nsurf. Typical penetration depths of the sensitive volume into the solution are in the order of 100 nm. Ligands are attached to the sensor surface, as indicated by half-circles. (A) When analytes (full circles) are introduced into the solution above the surface, reversible interactions with the ligand lead to association events at a rate governed by koncAcL,free (solid arrows), and dissociation events at a rate ccomplexkoff (dashed arrows). In the association phase, association events outnumber dissociation events until a steady state is reached. (B) When the surface is washed with running buffer in the absence of analyte, only dissociation events take place. (C) Signal obtained from probing the refractive index nsurf during the sequential application of the configurations depicted in A and B, given in arbitrary units. Following the association phase (A) and the dissociation phase (B), usually a regeneration procedure is applied for removing all remaining analyte from the surface before a new experimental cycle takes place.
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struments (ASI AG, Zürich, Switzerland). The purpose of this unit is to provide a brief introduction to these techniques for the quantitative characterization of biomolecular interactions. At the time of the introduction of optical biosensors in biomedical research, these instruments were thought to offer a particularly simple technique for the rapid quantitative measurement of equilibrium binding constants as well as binding kinetics. The reader is cautioned that the apparent simplicity and rapidity of the method is deceptive, and that the ease of using some of the commercial data analysis software packages can be highly misleading. The reliability of many of the initially used approaches in this field could not be verified in subsequent critical assessments of the methods. However, the technique has evolved significantly during the last few years. This is mainly due to the refinement and increasing variety of immobilization strategies, the identification of sources of artifacts, the use of control experiments, and the maturation of the analytical techniques (most notably global modeling). Emphasis is given in this unit to introducing the reader to robust approaches that will result in reliable results. Although the technique is frequently presented as being label-free, one of the interacting proteins has to be attached in an active form to the sensor surface. Most frequently, this is accomplished through covalent coupling, which can introduce all the problems well-known in chromophoric labeling and related covalent protein modifications. For all these reasons, the use of this technique requires careful planning and the execution of a number of different experiments, with time and effort more or less comparable to any of the other techniques available for the study of protein-protein interactions. Optical biosensors are very versatile for the study of protein interactions. However, as described below (see Strategic Planning), the choice of the experimental strategy is usually dictated by the proteins under investigation. Only general guidelines can be given for this stage of the experiment. Detailed procedures for some of the most commonly employed immobilization procedures are described, followed by an introduction to the different strategies for the binding studies, and a brief description of some of the most commonly encountered problems. Next, the need for control experiments is emphasized, and the execution of a variety of such control experiments is described. Finally, some ideas are given for simple troubleshooting. Obviously, this introductory unit cannot be even remotely complete, or cover many of the more advanced techniques such as assembly of protein complexes (Andersen et al., 1999; Schuster et al., 1993). For many of these, the reader will be referred to the specialized literature cited throughout this unit. STRATEGIC PLANNING The first step in planning a biosensor experiment for the measurement of the interaction of two proteins requires the choice of the binding partner to be immobilized and the type of binding experiment that will be employed (steady-state, kinetic, or competition experiment, see Binding Experiments and Data Analysis). This involves the consideration of several factors related to size, purity, and chemical properties of the protein, as well as the stoichiometry of the interaction. In general, this will require the protein samples to be well characterized with respect to these properties. Size, Concentration, and Volume Optical biosensors detect changes in the refractive index at the sensor surface due to analyte binding. This signal is superimposed by an offset from the bulk refractive index differences of the buffers used). Because the refractive index increment of proteins is very similar in units of weight concentration, the signal obtained from saturation of a given
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number of surface sites will increase with increasing molar mass of the soluble protein (or other analyte). In some of the commercial instruments, the binding of an analyte with a molar mass of only 1000 Da can be detected without difficulty, given the commonly achievable surface densities of sites. There is virtually no upper limit for the size of the analyte. Size consideration is important before setting up the experiment if the two binding partners to be studied are of significantly different size. The choice of a larger species as the analyte is generally preferable because this provides a larger signal. However, the immobilization of a very small peptide is usually difficult to control, and in practice frequently leads to unsuitably high density of surface sites, which can introduce problems of steric hindrance and mass transport limitation (see discussion of Mass Transport Limitation). In the absence of any other considerations, it seems advantageous to immobilize the larger binding partner, in order to diminish the probability of steric hindrance problems brought about by binding to surface sites in close vicinity. This will also help to minimize the probability of the immobilization affecting the conformation of the binding epitope. On the other hand, peptide ligands often are far more resistant to the denaturing effects of repeated cycles of binding and regeneration, and can be expected to provide a more stable binding surface than most proteins. If one of the proteins under study is available only in small quantities, it is advantageous to immobilize this species, because during the course of a study usually only a few functionalized surfaces are needed, each requiring usually only a few micrograms of material. The analytes are used in larger amounts; in one complete set of kinetic experiments volumes of the order of 100 µl at concentrations of ten times the KD are typically required. In competition experiments, similar quantities of the immobilized species are additionally required. The active concentration of the analyte should be well known, because errors in this value will directly translate into proportional errors of both the binding constant KD and the chemical on-rate constant kon, resulting from analysis of the experiment (see Binding Experiments and Data Analysis). Purity The sample purity requirements in biosensor experiments can be relatively low. In principle, the criterion for sufficient purity is the absence of cross-reaction between the two samples, aside from the interaction of the proteins under study—i.e., that no binding occurs between any contaminating species in the ligand sample and in the analyte sample. In practice, this means that at least either one of the samples should be highly purified. However, very strict requirements have to be met to ensure the absence of contaminating multimeric aggregates of the mobile species (see discussion of Analyte Aggregates, below, and Davis et al., 1998). Oligomeric Structure, Stoichiometry, and Multivalent Binding
Measuring Protein Interactions by Optical Biosensors
It is important to avoid multiple attachment of a single analyte molecule to two or more surface sites. If this occurs, the lifetime of the surface-bound complex is greatly enhanced. This avidity effect depends strongly on the local density of surface sites (Muller et al., 1998), and the binding data obtained under these conditions are governed to a large extent by the specific properties of the functionalized surface, much more than the surface binding properties of the analyte. (It cannot be assumed that the avidity effect of multivalent binding to the sensor surface is similar to possible avidity effects in binding to cell surface receptors.) Therefore, if the proteins under study are subject to multimeric binding, the multivalent species must be chosen for immobilization. For example, in antibody-antigen interactions, the antibody should be immobilized and the antigen should be the mobile analyte (Fig. 20.2.2). For unknown stoichiometries, appropriate control experiments should be performed that can identify the presence of multivalent species.
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A koff
k (app)off <
kon
k (2)on>>kon
B k (2)on=kon
koff kon
k (2)off=koff
kon
koff
Figure 20.2.2 (A) The binding of a bivalent analyte can take place through the interaction with a single ligand molecule (left), or with two ligand molecules (right), depending on the local ligand density. If an unoccupied immobilized ligand molecule is within an accessible distance to an already singly bound analyte molecule, then the on-rate constant of the second interaction is strongly enhanced because of the entropic colocalization constraints. The overall dissociation rate constant of the double attached analyte is substantially reduceed (usually by orders of magnitude), because it requires the virtually simultaneous dissociation of both interactions, which is much more improbable than a single dissociation event. The overall binding kinetics obtained from multivalent analytes is highly complex. (B) If the configuration is reversed, with the bivalent binding partner immobilized to the sensor surface, then both valencies act independently (left), and (in the absence of cooperativity of the sites) lead to binding kinetics that are identical to that of the monovalent interactions (right).
For the same reason, the presence of even very small amounts of reversibly formed oligomers (self-association) or irreversibly formed oligomers can significantly interfere with the analysis of surface binding equilibria or kinetics. Because of the difficulties in detecting relatively weak or rapidly reversible oligomerization, solution techniques like analytical ultracentrifugation can be preferable to chromatographic or electrophoretic techniques. Reactive Properties: Susceptibility to Immobilization, Nonspecific Binding, and Regeneration The key to successful biosensor experiments is the immobilization of one protein species in an active, and, if possible, uniformly oriented state to the sensor surface. The ease of immobilization can be the major consideration for the choice of the immobilized species. Examples are the presence of biotin, histidine, or other tags on the molecule that allow specific capture, or the presence of a single accessible cysteine suitable for employing specific cross-linking chemistry. If a protein is produced by recombinant methods, appropriate tags can frequently be introduced. Analogously, the protein that exhibits less nonspecific surface binding is the preferable choice for an analyte. Many sensor surfaces are slightly hydrophobic, and it is highly recommended to measure the extent of binding of both proteins to the nonfunctionalized surface at the maximal concentrations to be used in the binding experiments (e.g., at
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concentrations equal to 10-fold the expected dissociation equilibrium constant, KD). Significant degrees of nonspecific binding can make the analysis of the surface binding kinetics very difficult. Most analytical strategies require a series of multiple association/dissociation cycles to be executed with the same surface. Therefore, conditions need to be established that allow removal of all surface-bound analyte, without permanently altering the conformation and activity of the immobilized ligand (but tolerating possible destruction of the analyte). Naturally, the choice of the regeneration procedure depends very much on the proteins. Frequently, helpful hints can be obtained from the procedure used to purify the ligand— e.g., conditions employed during ion-exchange or affinity chromatography. Examples of common regeneration procedures include exposure to low-pH glycine or HCl buffer for 1 or 2 min. For some sensitive ligands that interact weakly with their respective analytes, an extended washout with the binding buffer is the gentlest and most reliable regeneration procedure. After selection of a regeneration procedure, control experiments for the integrity of the surface and for the reproducibility of the observed signal should be performed. These should consist of several identical association/dissociation/regeneration cycles. Insufficient regeneration can be identified by residual signal increase after each cycle and/or a negative drift of the baseline. The loss of binding sites through too-harsh regeneration conditions is indicated by a maximal signal during the association phase, which decreases with each cycle. Regeneration is not required for the equilibriumtitration techniques (Schuck et al., 1998; Myszka et al., 1998). IMMOBILIZATION PROTOCOLS The goal of immobilization is the stable coupling of the ligand to the sensor surface in an active form. In addition to the obvious functional test of analyte binding during the course of the interaction study, probing the conformation with monoclonal antibodies against ligand epitopes can be highly useful. If possible, it is desirable to reverse the role of ligand and analyte to rule out artifacts of immobilization on the interaction. It is important to note that the immobilization methods differ in their requirement for the exposure of the ligand to relatively harsh conditions, such as pH values below the pI in the commonly employed amine coupling. A related consideration in the choice of the immobilization strategy is the orientation of the surface-immobilized ligand. Random coupling techniques, such as amine coupling, may create different subpopulations of differently reactive ligand molecules at the surface, while specific orientation techniques (e.g., through specific cysteines or through specifically attached biotin moieties) create uniform surfaces. An elegant method for allowing uniform surface attachment is the introduction of specific tags (such as cysteine or histidine) into recombinantly produced or synthetic ligands.
Measuring Protein Interactions by Optical Biosensors
Below are basic descriptions of three of the most commonly used immobilization techniques, which can be applied to the most commonly employed carboxymethylated dextran sensor surfaces. In any of these methods it is very important to control the number of surface sites, since several surface-related artifacts are invoked if the surface density of the proteins under study is too high. The density of immobilized protein can be measured as the difference of the signal before and after immobilization, which can be transformed into the maximal analyte binding capacity by multiplication with the molar mass ratio of the analyte and ligand. For kinetic experiments, generally a relatively low density of immobilized protein should be obtained—i.e., a binding capacity of 50- to 200-fold of the instrumental noise [∼50 to 200 units (RU) on a Biacore system]. For competition and for certain control experiments (see Competition Analysis), a larger binding capacity can be desirable. In practice, for initial orientation on the behavior of a particular system of interacting proteins, it is best to prepare several different surfaces of
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different ligand densities and to compare the analysis of each. This will be helpful for diagnosing the regimes where mass transport limitations may be present (see Mass Transport Limitations), or where high surface coverage may lead to steric hindrance of analyte binding to neighboring surface sites (leading to biphasic surface binding kinetics). Instruments that allow the on-line observation of a reference surface can be optimally utilized if the reference surface is treated as similarly as possible to the functionalized surface. This is important predominantly because of possible nonspecific interactions of the analyte with the sensor surface, and the possible changes in surface properties due to immobilization. For amine coupling, this can include the application of the activation/deactivation cycle to the reference surface, but without cross-linking of any protein, or with cross-linking of a nonfunctional control protein (for example an unrelated antibody of the same isotype). For the same reason, if using biotin-streptavidin coupling, it is advantageous to immobilize streptavidin to both surfaces. A variety of additional immobilization techniques are available in addition those described below, such as hydrazide coupling (O’Shannessy et al., 1992), Ni2+ chelate coupling (Gershon and Khilko, 1995; O’Shannessy et al., 1995; Sigal et al., 1996), coupling of hydrophobic groups (Stein and Gerisch, 1996), the use of self-assembled monolayers (Sigal et al., 1996) or supported lipid or hybrid bilayers (Plant et al., 1995; Ramsden et al., 1996), and immobilization to aminosilane-derivatized surfaces (Edwards et al., 1995; Buckle et al., 1993). For details on chemical modifications of the proteins, see Hermanson (1996). Additionally, capturing strategies can be used for reversible attachment of the ligand. More specific information on commercially available sensor surfaces, and step-by-step protocols for the corresponding immobilization techniques, can be found in the commercial instrument handbooks, such as the Biacore applications handbook or the IAsys applications notes. Amine Coupling Amine coupling is most frequently carried out using a sensor surface that is modified with carboxymethylated dextran. It relies on the electrostatic preconcentration of the ligand to the sensor surface, and therefore requires that the proteins be positively charged. The coupling buffer should be 1 to 2 pH units below the pI of the protein, of low ionic strength (to avoid charge screening), and not contain any primary amines. Effective pre-concentration can be tested before the activation of the surface. It is frequently necessary to test immobilization with several different buffers of slightly different pH; an adjustment by only one half pH unit can be crucial for a successful immobilization. In a typical protocol, the carboxyl groups of the sensor surface are activated by exposure for 5 to 7 min with 50 mM N-hydroxysuccinimide (NHS) and 200 mM N-ethyl-N′-(dimethylaminopropyl)carbodiimide (EDC). Next, the ligand is injected in 10 mM sodium acetate pH 4.5 buffer. To control the amount of cross-linked ligand, it is usually better to adjust the reaction time as compared to adjustment of the ligand concentration. Repeated injection/wash cycles can be used to differentiate the covalently attached ligand from electrostatically surface-bound ligand. Finally, the surface is deactivated with 1 M ethanolamine⋅HCl, pH 8.5, for 7 min. Biacore and IAsys provide kits and prepared buffers for this procedure. The NHS and EDC should be kept frozen separately, in aliquots, at −20°C until use. Avidin- or Streptavidin-Biotin Coupling Covalent immobilization of avidin or streptavidin allows the capture of biotinylated proteins. This method avoids the need to expose the protein to a pH lower than the pI, and therefore can better preserve activity. Procedures for biotinylating the analyte are found
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in Hermanson (1996). The affinity of biotin-avidin interaction is very high and leads to a very stable surface. It should be noted that streptavidin-coated surfaces tend to be more sticky and lead to an increased level of nonspecific binding of the analyte to the surface. Therefore, good blank control surfaces are important. The amine coupling protocol described above and a carboxymethylated dextran surface can be used for this procedure. The surface is activated with NHS/EDC (see discussion of Amine Coupling), the streptavidin (∼100 µg/ml in buffer of pH 4.5 to 5.0) or avidin (∼50 µg/ml in pH 4.5 buffer) is immobilized, and the surface is deactivated with ethanolamine. Note that, if commercially prepared streptavidin surfaces are used, this activation step can be omitted. The biotinylated protein is then injected; usually concentrations of 10 to 100 µg/ml and low flow rates are sufficient for efficient capture. Typical times for exposure of the surface are 10 to 30 min. Thiol-Disulfide Exchange Coupling This immobilization technique is frequently used for acidic proteins, for small peptides, and for cases when immobilization in a specific orientation is important. It also serves as a highly efficient coupling procedure that can be used if amine coupling or avidin-biotin coupling is unsuccessful. In this method, the immobilization is accomplished either by the introduction of an active disulfide group onto the sensor surface, which can react with thiols on the protein to be immobilized, or, vice versa, by the modification of the sensor surface with a thiol group, which then can react with active disulfides on the protein. The choice between these alternative procedures is dependent on the protein, i.e., the presence of accessible thiol groups, or the potential for introducing activated disulfide groups. The latter procedure may be accomplished, e.g., using 2-(2-pyridinyldithio)ethaneamine (PDEA) to derivatize carboxyl groups or succinimidyloxycarbonyl-α-methyl-α-(2pyridyldithio)toluene (SMPT) for derivatizing amino groups. It should be noted that, in general, thiol-disulfide exchange coupling cannot be used for studies in the presence of reducing agents. However, the use of covalent thioester coupling—e.g., with succinimidyl 4-(N-maleimidomethyl) cyclohexane-1-carboxylate (SMCC), sulfo-SMCC, or N-(γmaleimidobutyryloxy)succinimide ester (GMBS)—produces a surface that is stable against reducing agents, and allows one to specifically chose the site of coupling. In addition, the latter technique provides a small spacer between the protein and the surface, which may allow better accessibility (Khilko et al., 1995). BINDING EXPERIMENTS AND DATA ANALYSIS Optical biosensors provide data on the time course of analyte binding, and allow for a number of different experimental and analytical strategies for interaction analysis. Most rely on a repeated cycle of association, dissociation, and surface regeneration (Fig. 20.2.1). Among the most important analysis strategies are the kinetic analysis, which takes advantage of the real-time observation to obtain the kinetic rate constants of the reaction; the steady state analysis for measuring the affinity of the reaction; and the competition analysis for measuring the affinity of the interaction in solution. The choice may be constrained by the affinity and kinetics of the proteins under study. If possible, multiple approaches should be taken and the results should be compared for their consistency. Their optimal application requires approximate knowledge of the order of magnitude of the affinity of the interaction. This may make it necessary to perform these studies in several steps. Measuring Protein Interactions by Optical Biosensors
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100 90
CA =10 K D
80 CA = 4 K D
Signal (%)
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60 50
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40 30
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20 CA = 0.25 K D
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0 0
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Figure 20.2.3 Schematic presentation of an ideal 1:1 pseudo first-order reaction. Shown is a superposition of the binding progress curves that should be obtained in a series of experiments at different analyte concentrations. The data are given in percent of the signal at maximal binding, but the units of the signal do not affect the results of the data analysis. The arrows on the curves at the higher concentrations indicate that steady-state binding is attained, whereas, for the lower curves, longer association times would be required as part of a steady-state analysis.
Kinetic Analysis Experimental The experiment consists of repeated injections of the analyte at different concentrations (Fig. 20.2.3). It should be conducted at relatively low surface density of sites, to avoid kinetic artifacts arising from mass transport limitation. An analyte concentration range as wide as possible should be covered (at least from 1/10 of the KD to 10 times the KD), using two-fold or three-fold dilutions. The time allowed for the surface binding phase (injection time) should be long enough for the observation of significant curvature in the binding progress, since it is only the curvature that contains the information about the reaction rate constants. It is highly desirable that steady-state binding be attained, at least for the highest concentrations used. The observation time for the dissociation phase should at least be similar to the injection time. Data analysis If the analyte-ligand interaction follows a pseudo first-order reaction, L + A ⇔ LA then the measured binding progress (proportional to [LA]) is described by: dR = kon cA ( Rmax − R) − koff R dt Equation 20.2.1
where cA denotes the analyte concentration, R denotes the signal (in arbitrary units), and Rmax denotes the signal at maximum saturation of the surface sites. The goal is the calculation of the chemical on-rate and off-rate constants, kon and koff, which relate to the equilibrium constant as:
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k KD = off kon Equation 20.2.2
While it is possible, in principle, to transform the data as dR/dt versus R in the association phase and as log(R) versus time in the dissociation phase, where kobs = (koncA + koff) and koff, respectively, would represent the slopes of the obtained straight lines, this procedure is very unreliable. Instead, it is state-of-the-art procedure to directly fit Equation 20.2.1, or its integrated form: Rassoc (t ) =
Rmax [1 − exp(kon c A + koff )t ] koff 1+ kon c A Equation 20.2.3
Rdissoc (t ) = Rassoc (t d ) exp[ − koff (t − t d )] Equation 20.2.4
directly to the data (with td denoting the end of the association and the beginning of the dissociation phase, where cA = 0; O’Shannessy et al., 1993). This should be done without truncation of the data beyond possible regions of artifacts from buffer changes or injections. Global analysis should be employed, which allows one to analyze all experimentally observed curves at all analyte concentrations simultaneously. This results in unambiguous best-parameter estimates for the binding constants, and provides residuals that should be inspected for comparison of the model and the data. Software needed for global analysis can be obtained from the manufacturer or as shareware. It should be noted that exclusion of specific data regions that may be poorly described by the model will introduce a bias into the analysis. If the global analysis cannot be performed, and separate analyses of the individual binding curves of each experiment according to Equation 20.2.3 and Equation 20.2.4 are applied, then tests for consistency of the results are crucial (Schuck and Minton, 1996a). Important tests are: (1) the estimated extrapolated values of Rassoc (t → ∞) at different concentrations should be consistent with a Langmuir isotherm for the estimated KD (e.g., approaching half-saturation at cA = KD; see Figure 20.2.2); and (2) the extrapolated values of (kassoccA + koff) from the association phases should be consistent with the value of koff as estimated in the dissociation phase. An analysis of the accuracy of the kinetic rate constants can be found in (Ober and Ward, 1999a).
Measuring Protein Interactions by Optical Biosensors
Possible problems The data may be only poorly described by Equation 20.2.3 and Equation 20.2.4 (leading to a poor fit and systematically distributed residuals) and instead show multiphasic behavior. Although it is in principle possible to apply global modeling with multiple binding-sites models, this is not recommended, because it is very difficult and can easily lead to very large errors (up to several orders of magnitude). Such modeling is usually not very robust, and many different models may fit the same data equally well (Glaser and Hausdorf, 1996). The reader is also advised to be extremely cautious with some of the implemented models in commercial or shared software, because they may not be valid descriptions of real surface binding processes (for example, mass transport models or models for bivalent analytes). Frequently, the deviations of the data from the single
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exponential binding progress predicted in Equations 20.2.3 and Equation 20.2.4 are signatures of experimental artifacts, such as heterogeneity of the ligand (subpopulations of ligand with different binding properties through nonspecific immobilization), steric hindrance of binding to neighboring sites, or the presence of mass transport limitations or multivalent aggregates of the analyte (see discussion of Common Experimental Obstacles). Steady-State Experiments and Binding Isotherm Analysis This approach can yield both equilibrium constants and, if appended by a dissociation analysis, kinetic constants. Its main advantage with respect to the previously described technique (see discussion of Kinetic Analysis) is that it is much more robust and reliable because it is less affected by problems of potential multiphasic binding. Experimental If in the course of the association experiments a steady-state signal is observed, which is generally possible for interactions with relatively high koff, then a binding isotherm can be constructed (Figs. 20.2.3 and 20.2.4). In an experimental procedure similar to the kinetic analysis, a concentration range as wide as possible should be covered, with two-fold concentration steps starting from both far below (≤1/10) KD up to almost complete saturation of the surface sites, far above the KD (≥ 10 times the KD). The injection times required to attain steady state at the lower concentrations are longer (see Equation 20.2.3 and Equation 20.2.4), and exponential extrapolation may be used, in particular in the Biacore system, since the association times may be limited in the by the limited volume in the injection loop of the microfluidics. Exponential extrapolation is not advised, however, for concentrations above KD. For interactions that do not yield steady-state binding within a tolerable time interval for the association phase, or where the ligand does not withstand a regeneration procedure, equilibrium titration represents an experimental variant (Fig. 20.2.4). In this approach, the analyte concentration is increased in two-fold steps, allowing for attainment of steady state in each step, avoiding regeneration between the steps. This can be achieved by addition of a small aliquot of the analyte into a fixed volume of solution above the sensor surface (in cuvette-based sensors; Hall and Winzor, 1997), into a loop of recycling sample (slightly modified Biacore X; Schuck et al., 1998), or into the running buffer reservoir of a standard Biacore (Myszka et al., 1998). The fixed-volume techniques require far less sample volume. After the highest analyte concentration has been applied, the dissociation phase can be observed and analyzed in terms of Equation 20.2.4. This can give an estimate of koff, which can be used in conjunction with Equation 20.2.2 to calculate kon as koff/KD. Data analysis Although the data analysis could in principle be performed by Scatchard analysis, this is not recommended for statistical reasons. A better method is a plot of Rsteady-state versus log(cA), in which the KD can be easily estimated as the inflection point at half saturation of the typical sigmoid isotherm (Fig. 20.2.4). The functional form for modeling is: Rsteady −state (cA ) =
Rmax 1 + K D / cA
Equation 20.2.5
Even if the data cannot be fitted well with this expression, the 50% saturation will still represent a good estimate of the order of magnitude of an average KD. It is important,
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however, in such an approach, to have enough data points at high analyte concentrations for a reliable estimate of Rmax and the 50% saturation level, respectively. Then the robustness will be greater than that of the kinetic analysis of Equations 20.2.3 and Equation 20.2.4. Competition Analysis Competition experiments can be performed in different variations, but conceptually all use a calibration of the sensor signal as a function of free analyte concentration, followed
A
100 4KD
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0 0.01KD
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Figure 20.2.4 (A) Time course of an equilibrium titration experiment. Arrows indicate the timepoints of the step-wise increase in the analyte concentration. (B) Steady-state binding analysis according to Equation 20.2.5. In a plot of the steady-state signal versus the base-10 logarithm of the analyte concentration, the binding isotherm exhibits a typical sigmoid shape. The inflection point at 50% saturation determines KD, and the width of the curve is characterized by ∼10% saturation at 1/10 KD and ∼90% saturation at 10 KD (dotted lines). Visible inspection of the data in this representation already allows a robust parameter estimation.
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by experiments with mixtures of analyte and soluble ligand in different molar ratios. The basic presumption is that the soluble ligand competes with the immobilized ligand for the free analyte, which leads to a reduction of the surface binding. This gives information on the solution affinity of ligand and analyte, while the freedom from the need to explicitly model the surface binding drastically diminishes the influence of potential surface artifacts on the results (Nieba et al., 1996; Schuck et al., 1998). This analysis can be extended to any other competitor that completely inhibits ligand binding of the analyte when bound to the analyte. Experimental For low-affinity systems with high chemical off-rate constants, in general, steady-state conditions can be achieved. Accordingly, the binding isotherm can be taken as a “calibration” of the sensor signal as a function of the free analyte concentration. Experiments to derive the surface binding constant can be conducted as described above (see Steady-State Experiments and Binding Isotherm Analysis). In a second variation, for high-affinity systems, mass-transport limited conditions are established by using a large immobilization density. Under these conditions, the initial binding rate constant will be directly proportional to the free analyte concentration (dR/dt = ktcA). The initial binding rate constant should be measured in equidistant concentration intervals for analyte concentrations up to a few times (two-fold) the KD. After either one of these sets of calibration experiments, a set of experiments is performed using a constant concentration of analyte, cA,tot (which should be taken in the range of KD), premixed and equilibrated with different concentrations of the soluble form of the ligand. The ligand concentrations, cL, should be taken in two-fold steps, spanning the range from far below to far above the KD. Choice of the fixed analyte concentration is important, since too high a value for cA,tot would show competition only in the range of stoichiometric binding. For systems that do not withstand regeneration, a recycling competitive titration procedure has been developed (Schuck et al., 1998). Data analysis For the high-affinity variant of the experiment with a linear initial rate of binding, dR/dt = ktcA, a plot of dR/dt versus cA can be used to graphically calculate the amount of free analyte, cA,free, during the competition experiment, or, cA,free is calculated according to the analytical inversion cA,free(cL) = dR/dt(cL)/kt. Similarly, in place of the linear function, an empirical calibration function for dR/dt(cA) could be used. The obtained concentration of free analyte cA,free in experiments at different soluble ligand concentrations cL can be plotted as cA,free versus log(cL), which gives a competition isotherm with the characteristic sigmoid shape. It can be fitted with Equation 20.2.6, the solution of the quadratic equation of the mass action law combined with mass balance: cA,free = cA,tot −
FH
1 sol sol 2 cA,tot + cL + KD − (cA,tot + cL + KD ) − 4cA,tot cL 2
IK
Equation 20.2.6
which gives the solution binding constant of analyte and ligand, KDsol. If the low-affinity variant of the experiment was used, the steady-state analysis of the sensor response in the absence of soluble ligand can be performed according to Equation 20.2.5, giving Rmax and KDsurface. This can be used in a direct analysis of the competition steady-state response using: Quantitation of Protein Interactions
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A
analyte flow cA
depletion zone
c
B
buffer flow
cA=0
retention zone
cA>0
C
competitor in flow
cA=0
Signal (%)
D
100 competitor c L
80 60
rebinding
40 20
no rebinding
0 0
100
200
300
400
500
Time (sec)
Measuring Protein Interactions by Optical Biosensors
Figure 20.2.5 Mass transport effects on the surface binding process. (A) In the association phase, insufficient transport does not fully replenish the free analyte in a zone near the sensor surface (depletion zone). The inability to maintain the concentration cA in the depletion zone leads to a limited surface binding rate. (B) The corresponding effect of mass transport limitation in the dissociation phase is that the surface is insufficiently washed and dissociated analyte is insufficiently removed. This generates a zone near the surface in which free analyte (after dissociation from the ligand) can rebind to empty ligand sites before diffusing into the bulk flow. This zone is indicated as retention zone. (C) Introduction of an excess of the soluble form of the ligand as a competitor into the buffer of the dissociation phase helps prevent rebinding to the surface. Soluble ligand can bind to dissociated analyte before rebinding takes place, and the soluble complex can diffuse into the buffer flow. This can allow the measurement of koff free of mass transport effects. (D) Time course of dissociation during a mass transport induced rebinding situation (as depicted in panel B), and after introduction of a soluble competitor into the washing buffer (arrow). In the presence of the competitor, the signal reflects the chemical off-rate constant koff, while in the rebinding situation, the apparent rate governing the signal is reduced.
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R( cL ) =
LM N
surf / cA,tot − 1 + KD
FH
Rmax
1 sol sol 2 cA,tot + cL + K D − ( cA,tot + cL + K D ) − 4cA,tot cL 2
IK OP Q
Equation 20.2.7
Alternatively, both titrations can be analyzed simultaneously in a global fit to both isotherms, assuming the binding constants to the surface sites and in solution to be identical. COMMON EXPERIMENTAL OBSTACLES Two of the most important experimental problems are mass transport limitations and the effect of aggregates on the binding kinetics. The first difficulty is found most frequently (but not exclusively) in systems of medium to high affinity with high kon, whereas the second is observed predominantly in systems with lower affinity, where higher analyte concentrations are required. Other obstacles can be steric hindrance effects of binding to adjacent surface sites (O’Shannessy and Winzor, 1996), or the contribution of second-order kinetics to the binding process in cuvette systems (Edwards et al., 1998). Most of these processes scale with the immobilization level of the ligand. For this reason, a low ligand density is generally advantageous in minimizing potential problems. Special considerations for working at very low ligand densities are described in Ober and Ward (1999a,b). The use of control experiments (e.g., comparison of the KD from solution competition experiments with the KD obtained from surface binding kinetic measurements) is very useful in verifying the absence of artifacts. The role of mass transport and analyte aggregates will be explained in detail below. Mass Transport Limitation If the supply of analyte to the sensor surface is the limiting factor in the surface binding rate, the observed binding kinetics are mass transport limited. This will depend directly on both the density of the immobilized sites, and on the chemical on-rate constant of the interaction. In the association phase, this will produce a zone of depleted analyte concentration in the vicinity of the surface, while in the dissociation phase, this will lead to a zone of nonvanishing free analyte concentration, which is subject to rebinding-retention effect (Silhavy et al., 1975; Fig. 20.2.5). The quantitative influence of this process on the observed surface binding kinetics is governed by a highly complex reaction/diffusion/convection process (Yarmush et al., 1996; Schuck, 1996). Although the processes in the association and dissociation phase are conceptually closely related, it cannot be assumed that the ratio of the “apparent” rate constants still reflects a good estimate of the equilibrium constant. Also, it is strongly recommended that the nonspecialist reader refrain from modeling using mass-transport-corrected kinetic models. Instead, the primary goal is to identify mass transport limitations, and to establish experimental conditions avoiding transport effects. The following is a list of diagnostic indicators, in the order of reliability, for mass transport limited binding. 1. Dependence on surface capacity. This is a very good indicator of mass transport limitation, since the transport limit always directly scales with the density of active surface sites. For this reason, variations of the immobilization density of the ligand are recommended as routine tests demonstrating the absence of transport limitation and other surface-related artifacts. This test can fail in the presence of significant non-specific analyte binding to the surface.
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B F2
F3
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10 Concentration(µM)
100
Figure 20.2.6 Exemplary data of the interaction of TCR with superantigen (for details of this interaction see Andersen et al., 1999). (A) Elution profile of the TCR from size-exclusion chromatography, indicating the fractions used for kinetic analysis of the interaction with immobilized superantigen, as shown in (B). It should be noted that fraction 1 (bold solid line), fraction 2 (dashed line), and fraction 3 (solid line) all exhibit multiphasic binding, with different relative magnitudes of the slower component. It should also be noted that despite the lower concentration of fraction 1, which results in the lowest response in the association phase, the signal in the dissociation phase is highest. This slower kinetic component in the binding progress curve reflects different amounts of aggregates bound to the sensor surface. The aggregates have a slower dissociation because of their multivalent attachment. For comparison, the same interaction is shown in the reverse orientation, with immobilized TCR and soluble superantigen (dotted line). (C) Sequence of association-dissociation curves at different superantigen concentrations, binding to immobilized TCR. In this orientation, the association and the dissociation is much faster, virtually monophasic, and the binding is completely reversible, which provides further evidence that the slow kinetic components introduced in the different fractions of the soluble TCR sample in Panel B are artifacts. For quantitative analysis, the signal measured at a nonfunctionalized surface (dotted line) is subtracted in order to remove the bulk refractive index contribution of the analyte. (D) A plot of the net steady-state binding signal versus superantigen concentration allows the measurement of the affinity of the interaction free from artifacts due to TCR aggregation that would be introduced in the configuration of Panel B.
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Figure 20.2.7 Effects of the ligand density on the binding kinetics for interactions with relative low affinity (A) and medium affinity (B). Each panel shows the interaction of immobilized superantigen with single-chain TCR, observed at high ligand surface density (1700 RU, solid lines) and low ligand surface density (700 RU, dashed lines). For easier comparison, the signal obtained at the lower density surface was scaled proportionally. In panel A, a slow phase of binding in the association phase and a residual binding (possible slow dissociation of multivalently bound aggregates) is introduced at high ligand density, under otherwise identical conditions. For the interaction in panel B, the chemical off-rate constant is smaller than for the low-affinity interaction shown above. Nevertheless, from comparison of the binding curves at different ligand surface densities (under otherwise identical conditions) it is obvious that an increased ligand density has significant effects on the surface binding kinetics. This observation could be due to aggregation or to mass transport limitations, both of which are more likely at higher ligand densities.
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2. Increased dissociation rate when a soluble form of the ligand is injected (Fig. 20.2.5D). This always strongly indicates rebinding (mass transport limitation) if detected. Since the ligand by itself should not interact with the surface, its effect is the binding to analyte near the sensor surface, preventing it from rebinding and allowing diffusion and washout from the surface (Fig. 20.2.5C). Unfortunately, this effect may not be present for large ligands or ligands/analytes with high nonspecific binding if they exhibit limited diffusivity. 3. Double exponential dissociation phase. This will be seen only after dissociation from more than 50% occupation of all available surface sites. 4. Weak dependence on the flow rate (stirring speed in cuvette-based systems). Since the transport parameters only change with the cube root of the flow rate, i.e., generating only a factor of 2 when changing the flow rate by a factor of 10, this flow rate dependency can be difficult to detect. This is particularly true if the flow rate is only varied by a factor of 2, and the reaction is only partially transport influenced. Then, only a ∼10% change in the apparent koff would be expected in cases where the true koff is 100% larger than the apparent koff. 5. Linear association phase. This effect may not always be present, in particular at substantial transport limitation (Schuck, 1996; Schuck, 1997b; Schuck and Minton, 1996b). Generally, tests 1 and 2 are the most reliable. They also lead to experimental techniques that can be utilized to reduce or eliminate mass transport artifacts. The most effective way for reducing mass transport influence is lowering the surface density of the immobilized ligand. Higher flow rates give only comparatively very small improvements, but are connected with strongly increased sample volume requirements. If the surface density of the ligand cannot be reduced further without leading to an insufficient signal-to-noise ratio, then switching from kinetic experiments to steady-state or competition steady-state experiments is the best solution. This will give information on the equilibrium constant. The kinetic rate constants can then be estimated best from a saturation experiment (approaching complete saturation of all surface sites), followed by a dissociation phase during which soluble ligand is coinjected. The soluble ligand will minimize rebinding and allow the estimation of the chemical off-rate constant, from which the chemical on-rate constant can be determined via Equation 20.2.3. Analyte Aggregates Oligomeric aggregates of analyte can be troublesome in biosensor experiments in two different ways. First, if trace amounts of higher oligomers are present in the analyte sample, this will lead, in the association phase, to a slow accumulation at the sensor surface, which can be visible as a slower second phase of binding. As depicted in Figure 20.2.2, these multimeric analyte aggregates can have multiple interactions with immobilized ligand molecules, and therefore they will dissociate much more slowly than the monomeric analyte. Consequently, they will appear in the dissociation phase as a submoiety with a very low off-rate constant (Davis et al., 1998). The troublesome trace amounts of oligomers can be eliminated by careful chromatographic purification, or their influence can be minimized by exchanging the role of analyte and ligand (Davis et al., 1998; Andersen et al., 1999). This is illustrated in the example of Figure 20.2.6, which demonstrates the importance of both the sample preparation and the choice of ligand and analyte. Measuring Protein Interactions by Optical Biosensors
The second potentially problematic form of aggregation is a surface-induced multimerization of the analyte. Because the local macromolecular concentrations at the sensor surface are very high (e.g., in the order of 10mg/ml at a signal of 1000 RU in Biacore
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instruments), local crowding effects combined with non-specific interactions of the analyte can promote oligomer formation at the sensor surface (Fig. 20.2.7A; Minton, 1995, 1998). As with the influence of preformed aggregates, this process will lead to biphasic association and dissociation profiles, with the slower phase resulting from oligomer accumulation and dissociation, respectively. This process will be favored by higher surface concentrations—i.e., higher density of immobilized ligand, and by higher ligand affinity (Fig. 20.2.7). As with mass transport limitations, they can be detected by Table 20.2.1
Troubleshooting Guide for Measuring Protein Interactions by Optical Biosensors
Problem
Solution
No electrostatic preconcentration achieved; poor immobilization
Desalt protein (e.g., using spin column or microdialysis); decrease pH of buffer used for immobilization (should be below pI of protein) Analyte may be too hydrophobic, or there may be electrostatic interaction with surface. Increase salt or detergent concentration in running buffer.a If this does not result from high analyte concentrations, dialyze the analyte against running buffer, or use a spin column for buffer exchange Use increasingly harsher conditions for regeneration; test procedures used in affinity chromatography; check for strong nonspecific binding of the analyte; check for possible incomplete blocking of activated surface sites after immobilization Check for presence of free biotin in sample. Biotin on the analyte may not be accessible by surface-immobilized streptavidin; in this case try biotinylated linker Check for mass-transport artifacts; check for possible traces of aggregates and for formation of aggregates at the surface; change immobilization method (avoid random coupling, avoid large surface densities). If this is not successful, go to steady-state analysis methods. Use longer injections by increasing injection volume and/or decreasing flow rate. If sample volume is limiting, try an equilibrium titration Decrease immobilization density.b If immobilization density cannot be lowered further, go to steady-state analysis, combined with establishment of lower limit of kdis from dissociation after saturation. Potential signature of mass transport; lower the immobilization density Signature of mass transport limitation; lower the immobilization density
Nonspecific binding is high
High signal from buffer; refractive index changes
Analyte does not come off after regeneration
No binding of biotinylated sample to streptavidin surface
Kinetics does not follow 1:1 binding
No steady-state binding is reached in a flow system Mass transport limitation
Increasing slope in the association phase Increasing signal in the dissociation phase
aBe aware of effects of nonspecific binding on the binding kinetics, which substantially decreases the diffusivity of the
analyte across the sensor surface, potentially leading to mass transport artifacts that cannot be detected through change of the ligand density. bIncreasing the flow rate affects transport much less, but consumes much more sample.
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variation of the surface density of the ligand. They also can be reduced by lowering the surface density of the immobilized ligand, by size-exclusion chromatography immediately prior to the experiment, or by exchanging the role of ligand and analyte. Troubleshooting As with other complex biophysical techniques and all investigations involving biological samples, it is impossible to give guidelines which are even nearly complete or generally helpful for troubleshooting. Nevertheless, Table 20.2.1 presents a small list of possible solutions to potential problems, given in the hope that some readers may find them helpful. SUMMARY Binding studies with optical biosensors can be very powerful and versatile. Among the most important virtues are their high sensitivity and utility for a broad range of affinities, real-time detection allowing studies of binding kinetics, and relatively low requirements of sample volume. We have outlined some general strategies and described some of the most commonly used techniques. The work with protein interactions at a surface can introduce additional experimental difficulties as compared to solution methods. As a general rule in experiments with optical biosensors, it is highly recommended that analyses be performed in different ways so that the consistency of the results can be tested. Biosensors can be an excellent tool in the study of protein interactions, and become particularly powerful if combined with other methods. ACKNOWLEDGMENT The authors are grateful for helpful comments by D. Margulies in the preparation of the manuscript. They acknowledge D. Kranz and K. Karjalainen for providing the samples used for some of the experiments shown. Peter Andersen acknowledges grant support by the Danish Natural Sciences Research Council. LITERATURE CITED Andersen, P.S., Lavoie, P.M., Sekaly, R.P., Churchill, H., Kranz, D.M., Schlievert, P.M., Karjalainen, K., and Mariuzza, R.A. 1999. Role of the T cell receptor alpha chain in stabilizing TCR-superantigen-MHC class II complexes. Immunity 10:473-483. Buckle, P.E., Davies, R.J., Kinning, T., Yeung, D., Edwards, P.R., Pollard-Knight, D., and Lowe, C.R. 1993. The resonant mirror: A novel optical sensor for direct sensing of biomolecular interactions. Part II: Applications. Biosens. Bioelectron. 8:355-363. Davis, S.J., Ikemizu, S., Wild, M.K., and van der Merwe, P.A. 1998. CD2 and the nature of protein interactions mediating cell-cell recognition. Immunol. Rev. 163:217-236. Edwards, P.R., Gill, A., Pollard-Knight, D.V., Hoare, M., Buckle, P.E., Lowe, P.A., and Leatherbarrow, R.J. 1995. Kinetics of protein-protein interactions at the surface of an optical biosensor. Anal. Biochem. 231:210-217. Measuring Protein Interactions by Optical Biosensors
Edwards, P.R., Maule, C.H., Leatherbarrow, R.J., and Winzor, D.J. 1998. Second-order kinetic analysis of IAsys biosensor data: Its use and applicability. Anal. Biochem. 263:1-12.
Garland, P.B. 1996. Optical evanescent wave methods for the study of biomolecular interactions. Q. Rev. Biophys. 29:91-117. Gershon, P.D. and Khilko, S. 1995. Stable chelating linkage for reversible immobilization of oligohistidine tagged proteins in the Biacore surface plasmon resonance detector. J. Immunol. Methods. 183:65-76. Glaser, R.W. and Hausdorf, G. 1996. Binding kinetics of an antibody against HIV p24 core protein measured with real-time biomolecular interaction analysis suggest a slow conformational change in antigen p24. J. Immunol. Methods. 189:1-14. Hall, D.R. and Winzor, D.J. 1997. Use of a resonant mirror biosensor to characterize the interaction of carboxypeptidase A with an elicited monoclonal antibody. Anal. Biochem. 244:152-160. Hermanson, G.T. 1996. Bioconjugate techniques. Academic Press, San Diego. Khilko, S.N., Jelonek, M.T., Corr, M., Boyd, L.F., Bothwell, A.L.M., and Margulies, D.H. 1995. Measuring interactions of MHC class I molecules using surface plasmon resonance. J. Immunol. Methods. 183:77-94.
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Knoll, W. 1998. Interfaces and thin films as seen by bound electromagnetic waves. Annu. Rev. Phys. Chem. 49:569-638. Leckband, D.E. 1997. The influence of protein and interfacial structure on the self-assembly of oriented protein arrays. Adv. Biophys. 34:173-190. Leckband, D.E., Kuhl, T., Wang, H.K., Herron, J., Muller, W., and Ringsdorf, H. 1995. 4-4-20 antifluorescyl IgG Fab′ recognition of membrane bound hapten: Direct evidence for the role of protein and interfacial structure. Biochemistry 36:11467-11478. Lukosz, W. 1991. Principles and sensitivities of integrated optical and surface plasmon sensors for direct affinity sensing and immunosensing. Biosens. Bioelectronics. 6:215-225. Malmborg, A.C. and Borrebaeck, C.A. 1995. Biacore as a tool in antibody engineering. J. Immunol. Methods. 183:7-13. Margulies, D.H., Corr, M., Boyd, L.F., and Khilko, S.N. 1993. MHC Class I/peptide interactions: Binding specificity and kinetics. J. Mol. Recognit. 6:59-69. Margulies, D.H., Plaksin, D., Khilko, S.N., and Jelonek, M.T. 1996. Studying interactions involving the T-cell antigen receptor by surface plasmon resonance. Curr. Opin. Immunol. 8:262-270. Minton, A.P. 1995. Confinement as a determinant of macromolecular structure and reactivity. 2. Effects of weakly attractive interactions between confined macrosolutes and confining structures. Biophys. J. 68:1311-1322. Minton, A.P. 1998. Molecular crowding: Analysis of effects of high concentrations of inert cosolutes on biochemical equilibria and rates in terms of volume exclusion. Methods Enzymol. 295:127149. Muller, K.M., Arndt, K.M., and Plückthun, A. 1998. Model and simulation of multivalent binding to fixed ligands. Anal. Biochem. 261:149-158. Myszka, D.G., Jonsen, M.D., and Graves, B.J. 1998. Equilibrium analysis of high affinity interactions using Biacore. Anal. Biochem. 265:326-30. Nieba, L., Krebber, A., and Plückthun, A. 1996. Competition Biacore for measuring true affinities: Large differences from values determined from binding kinetics. Anal. Biochem. 234:155165. Ober, R.J., and Ward, E.S. 1999a. The influence of signal noise on the accuracy of kinetic constants measured by surface plasmon resonance experiments. Anal. Biochem. In press. Ober, R.J. and Ward, E.S. 1999b. The choice of reference cell in the analysis of kinetic data using Biacore. Anal. Biochem. 271:70-80. O’Shannessy, D.J. and Winzor, D.J. 1996. Interpretation of deviations from pseudo-first-order kinetic behavior in the characterization of ligand binding by biosensor technology. Anal. Biochem. 236:275-283.
O’Shannessy, D.J., Brigham-Burke, M., and Peck, K. 1992. Immobilization chemistries suitable for use in the Biacore surface plasmon resonance detector. Anal. Biochem. 205:132-136. O’Shannessy, D.J., Brigham-Burke, M., Soneson, K.K., Hensley, P., and Brooks, I. 1993. Determination of rate and equilibrium binding constants for macromolecular interactions using surface plasmon resonance: Use of nonlinear least squares analysis methods. Anal. Biochem. 212:457-468. O’Shannessy, D.J., O’Donnell, K.C., Martin, J., and Brigham-Burke, M. 1995. Detection and quantitation of hexa-histidine-tagged recombinant proteins on western blots and by a surface plasmon resonance biosensor technique. Anal Biochem. 229:119-24. Plant, A.L., Brigham-Burke, M., Petrella, E.C., and O’Shannessy, D.J. 1995. Phospholipid/alkanethiol bilayers for cell-surface receptor studies by surface plasmon resonance. Anal. Biochem. 226:342-348. Ramsden, J.J., Bachmanova, G.I., and Archakov, A.I. 1996. Immobilization of proteins to lipid bilayers. Biosens. Bioelectronics 11:523-528. Schuck, P. 1996. Kinetics of ligand binding to receptor immobilized in a polymer matrix, as detected with an evanescent wave biosensor. I. A computer simulation of the influence of mass transport. Biophys. J. 70:1230-1249. Schuck, P. 1997a. Use of surface plasmon resonance to probe the equilibrium and dynamic aspects of interactions between biological macromolecules. Annu. Rev. Biophys. Biomol. Struct. 26:541-566. Schuck, P. 1997b. Reliable determination of binding affinity and kinetics using surface plasmon resonance biosensors. Curr. Opin. Biotechnol. 8:498502. Schuck, P., and Minton, A.P. 1996a. Minimal requirements for internal consistency of the analysis of surface plasmon resonance biosensor data. Trends Biochem. Sci. 252:458-460. Schuck, P. and Minton, A.P. 1996b. Analysis of mass transport limited binding kinetics in evanescent wave biosensors. Anal. Biochem. 240:262-272. Schuck, P., Millar, D.B., and Kortt, A.A. 1998. Determination of binding constants by equilibrium titration with circulating sample in a surface plasmon resonance biosensor. Anal. Biochem. 265:79-91. Schuster, S.C., Swanson, R.V., Alex, L.A., Bourret, R.B., and Simon, M.I. 1993. Assembly and function of a quaternary signal transduction complex monitored by surface plasmon resonance. Nature 365:343-347. Sigal, G.B., Bamdad, C., Barberis, A., Strominger, J., and Whitesides, G.M. 1996. A self-assembled monolayer for the binding and study of histidine-tagged proteins by surface plasmon resonance. Anal. Chem. 68:490-497.
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Silhavy, T.J., Szmelcman, S., Boos, W., and Schwartz, M. 1975. On the significance of the retention of ligand by protein. Proc. Natl. Acad. Sci. U.S.A. 72:2120-2124.
O’Shannessy et al., 1992. See above.
Stein, T., and Gerisch, G. 1996. Oriented binding of a lipid-anchored cell adhesion protein onto a biosensor surface using hydrophobic immobilization and photoactive crosslinking. Anal. Biochem. 237:252-259.
General review of the method and its application.
van der Merwe, P.A. and Barclay, A.N. 1996. Analysis of cell-adhesion molecule interactions using surface plasmon resonance. Curr. Opin. Immunol. 8:257-261.
Web site for Biacore; extensive list of published biosensor applications
Yarmush, M.L., Patankar, D.B., and Yarmush, D.M. 1996. An analysis of transport resistances in the operation of Biacore; Implications for kinetic studies of biospecific interactions. Mol. Immunol. 33:1203-1214.
Key References Davis et al, 1998. See above. Contains a detailed description of analyte aggregation effects on the measured surface binding. Nieba et al., 1996. See above.
Collection of immobilization techniques. Schuck, 1997b. See above.
Internet Resources http://www.biacore.com
http://www.affinity-sensors.com Web site for Affinity Sensorsl; extensive list of published biosensor applications.
Contributed by Peter Schuck and Lisa F. Boyd National Institutes of Health Bethesda, Maryland Peter S. Andersen University of Maryland Rockville, Maryland
Demonstration how competition approaches can be used to circumvent kinetic artifacts.
Measuring Protein Interactions by Optical Biosensors
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Current Protocols in Protein Science
Analytical Centrifugation: Equilibrium Approach Equilibrium centrifugation, also called equilibrium sedimentation and equilibrium analytical ultracentrifugation, is a classical method of biochemistry for providing firstprinciple thermodynamic information about the molar mass, association energy, association stoichiometry, and thermodynamic nonideality of molecules in solution. Because it relies on the principal property of mass and the fundamental laws of gravitation, equilibrium sedimentation can be used to analyze the solution behavior of proteins of any type in a wide range of solvents and over a wide range of protein concentrations. For many questions about protein solution behavior, there is no satisfactory substitute method of analysis. This unit presents (1) background information about equilibrium sedimentation, (2) a review of the basic theory of equilibrium sedimentation, (3) a strategy to use when planning equilibrium sedimentation experiments, and (4) an overview of the analysis of sedimentation equilibrium data. The early literature covering the invariant foundations of sedimentation is still relevant (see references in Laue and Stafford, 1999), and beginners should review these early papers. This unit will stress what questions may be answered when performing sedimentation equilibrium analysis using the absorbance and interference optical detectors on the BeckmanCoulter XLI ultracentrifuge. In so doing, it will neglect techniques that use preparative centrifuges and postsedimentation analysis (Rivas and Minton, 1993), as well as density gradient sedimentation (Rickwood, 1984). Furthermore, space limitations prevent complete descriptions of the methods available for data analysis; therefore, references are made to the pertinent literature. The analytical ultracentrifuge is similar to a high-speed preparative centrifuge in that a spinning rotor provides a gravitational field large enough to sediment molecules (Ralston, 1993). The analytical ultracentrifuge is distinguished from the preparative centrifuge by specialized rotors, sample holders, and optical systems that permit the observation of samples during sedimentation (Fig. 20.3.1). Using present technology, a maximum of 32 samples may be analyzed in a single experiment (Ralston, 1993; Laue, 1995).
Contributed by Tom Laue Current Protocols in Protein Science (1999) 20.3.1-20.3.13 Copyright © 1999 by John Wiley & Sons, Inc.
UNIT 20.3
The defining characteristic of sedimentation equilibrium is the time-invariant concentration gradient that develops as the flux of sedimenting molecules is exactly balanced by the flux of diffusing molecules at each point in the sample holder, referred to as a cell. No distinct boundaries are observed; instead, a smooth concentration gradient is observed (Fig. 20.3.2). The accurate determination of this concentration gradient is the fundamental measurement needed for the analysis of a sedimentation equilibrium experiment. To make this measurement, the cells are passed through the optical paths of detectors by the spinning rotor (Fig. 20.3.1). Presently, there are two optical detectors for the XLI, one that uses absorbance and one that uses refractive index to measure the concentration at closely spaced radial intervals. A review of these two optical systems is available (Laue, 1996). Analysis of the concentration profiles provided by the optical systems provides rigorous insight into the solution behavior of sedimenting molecules. Interpretation of these data requires knowledge of the rotor speed, temperature, elapsed time, and buoyancy (Laue and Stafford, 1999).
THEORY Sedimentation equilibrium is a powerful method for describing the solution thermodynamic behavior of proteins. The most common thermodynamic behaviors exhibited by proteins include (1) single ideal component, (2) ideal, reversible self-association, (3) heterogeneous mixture of components, (4) ideal, reversible association between dissimilar proteins, and (5) nonideality. The theory behind these behaviors may be found in Laue and Stafford (1999) and references therein. These behaviors are not necessarily mutually exclusive, and it is often found that a protein exhibits both self-association and nonideality, or self-association and heterogeneity. Experimentally, then, it is important to be able to diagnose which of these five behaviors are important to a protein’s solution behavior. The goal of this section is to provide a brief mathematical description of the equilibrium concentration profiles for the five common behaviors. Based on these descriptions, a set of diagnostic tests will be presented.
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F
H A
G
B F
D
C
E
D
E
Figure 20.3.1 A schematic representation of an analytical ultracentrifuge. The rotor (C) has holes through it to hold sample containers commonly called cells (D). Each cell (see inset) consists of a centerpiece (G) with open-sided chambers called channels (H) to hold the liquid samples. There are two channels per sample, one containing the protein in its solvent and an adjacent one (not shown) containing only solvent to serve as an optical reference. The centerpiece, in turn, is sealed between windows (F) to permit the passage of light through the channels, thus allowing the cell contents to be viewed. Centerpieces are made out of a variety of tough inert materials. Depending on the type of experiment that will be performed, centerpieces can hold one, three, or four samples each. Rotors can hold either four or eight cells. As the rotor spins, each cell passes through the optical paths of two different detectors. The absorbance detector uses a pulsed xenon lamp (A) to provide a burst of light when a cell is aligned in the beam. The absorbance detector (E) uses a narrow slit and photomultiplier tube to determine the light intensity after the beam has passed through the sample. The slit is moved radially by a motor so that the absorbance profile, called an absorbance scan, can be determined. The second detector determines the refractive index difference between the sample and reference channels at each radial position. The light source (B) uses a laser diode to produce two radially directed narrow stripes of light, one that passes through the sample channel and one that passes through the reference channel. These two stripes of light are brought together to produce an interference image (Fig. 20.3.2) in which the difference in the refractive index at each radial position is displayed as the vertical displacement of a set of fringes.
Single Ideal Component Dilute solutions of a highly purified protein at physiological salt concentrations often behave like an ideal system with the protein as the only sedimenting component. For a single thermodynamically ideal component, the basic equation (Eqn. 20.3.1) describing the concentration as a function of radial position, c(r), is a simple exponential: 2
c( r ) = c0 e
σ
r − r0 2 2
= c0 e σξ
Equation 20.3.1 Analytical Centrifugation: Equilibrium Approach
where c0 is the concentration at a reference radius (r0, in cm), and σ is the reduced molecular weight:
σ=
Mbω 2 RT
Equation 20.3.2
In Equation 20.3.2, Mb is the buoyant mass (discussed below), ω is the centimeter-gramsecond (cgs) rotor speed (rpm π/30, in radians/sec), R is the cgs gas constant (3.14 × 107 erg/mole/°K), and T is the absolute temperature (°K). Notice that σ is directly proportional to the buoyant mass and the square of the rotor speed. Because of these dependencies, equilibrium sedimentation can be used to measure molecular weights ranging from ∼1000 to 20 × 106 Da. Experimentally, a single ideal component yields the same value of Mb regardless of the sample concentration or rotor speed.
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meniscus
solution
base
increasing c sedimentation diffusion increasing r increasing g
Figure 20.3.2 A Rayleigh interference refractive optical system image of one channel of a six-channel centerpiece (Fig. 20.3.4). This optical system provides an image in which the concentration at each radial position is represented by the vertical displacement of a set of equally spaced horizontal fringes. The center of rotation is to the left and the edge of the rotor is to the right. An air/liquid meniscus forms at the top of the channel, one in the reference sector and one in the sample sector. As slightly different volumes of liquid are used to fill these two sectors, the menisci do not appear at exactly the same radius. If the two menisci do become superimposed over the course of an experiment, there is a leak between the sample and reference sectors, indicating that the centerpiece is scratched and needs polishing. The sample is a protein at sedimentation equilibrium, where the flux of molecules towards the cell bottom due to sedimentation is exactly balanced by the flux of molecules towards the meniscus due to diffusion. The balance of fluxes yields an exponential concentration distribution. The base of the cell is formed by FC-43, a clear fluorocarbon liquid with a refractive index very close to that of water.
Ideal Reversible Self-Association The equation describing reversible self-association in an otherwise ideal solution is obtained by expanding Equation 20.3.1 in terms of the species:
c( r ) = ∑ c0 i e c01e
σi ξ
σ1ξ
=
+ c02 e
σ 2ξ
+ c03 e
σ 3ξ
+...
c0 n
Kal ⇔n =
c01
n
Equation 20.3.4
where [c0n] is the oligomer concentration and [c01] is the monomer concentration. Substituting Equation 20.3.4 into Equation 20.3.3: i
Equation 20.3.3
where i is the index for the components, c01 and σ1 refer to reference concentration and reduced buoyant molecular weight of the monomer, c02 and σ2 refer to these quantities for the dimer, and so forth. For a reversible mass-action association, all σ’s are related so that σ2 = 2σ1, σ3 = 3σ1, and so on. The monomer reference concentration, c01, is linked to the other reference concentrations, c0n, through the equilibrium constant:
c( r ) = ∑ ci ( r ) = ∑ c0 i K al ⇔ i e = c01e 3
σ1ξ
2
+ c01 K al ⇔ 2 e
c01 K al ⇔ 3 e
3σ1ξ
iσ i ξ
2 σ1ξ
+
+...
Equation 20.3.5
It is important to note in Equation 20.3.5 that the abundance of all of the associated species depends only on the reference concentration of the monomeric protein, c01. As c01 increases, the abundance of dimers, trimers, and so on
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increases relative to the concentration of the monomer, whereas dilution decreases their relative abundance. One way to describe the changes in the relative abundance of species is to use an average molecular weight (Van Holde, 1985). The weight-averaged molecular weight, Mw, is defined as:
Mw ≡
∑ ci Mi ∑ ci
Equation 20.3.6
There are procedures that determine Mw at each point in an equilibrium sedimentation distribution, and graphs of Mw as a function of either concentration or rotor speed are useful diagnostics (Yphantis, 1964; Roark and Yphantis, 1969). For a reversible ideal self-association, Mw increases with increasing protein concentration and decreases as a solution is made more dilute. Most importantly, Mw depends only on the concentration of the monomer and is independent of rotor speed.
Heterogeneous Mixtures Formation of irreversible aggregates is a common behavior of proteins. These aggregates can be of any size, resulting in a mixture of dimers, trimers, and so on whose concentration is not a reflection of the monomer concentration. The equation describing the sedimentation equilibrium of such a heterogeneous mixture is obtained by expanding Equation 20.3.1 in terms of distinct components:
Ideal Reversible Heterogeneous Association There are two common situations that lead to heterogeneous associations. The first is when monomers of an otherwise seemingly homogeneous solution self-associate, but with a range of equilibrium constants (Yphantis et al., 1978; Senear and Teller, 1981). In fact, careful scrutiny of nearly all self-associations seems to reveal some heterogeneity. Experimental treatment of these systems is outside the scope of this chapter, but the reader should be aware that this can be observed. The second situation occurs when unlike molecules within a mixture associate. Some of the most interesting interacting biochemical systems involve reversible associations between nonidentical components. More complicated expressions must be derived that explicitly take into account the concentrations of each component and the various species:
c( r ) = ∑ ci ( r ) = c01e
σ1ξ
m ′ ⇔m e Kal + ∑ c01
mσ1ξ
m
(self − association of component 1) + c02 e
σ2ξ
n ′′ ⇔n e Kal + ∑ c02
nσ 2 ξ
n
(self − association of component 2) j k ′′′⇔ j + k e Kal + ∑ c01c02
( jσ1 + kσ 2 ) ξ
j ,k
( heteroassociation ) Equation 20.3.8
c( r ) = ∑ c 0 i e
σi ξ
σ ξ c01e 1
=
+ c02 e
σ 2ξ
+ c03e
σ 3ξ
+...
Equation 20.3.7
Analytical Centrifugation: Equilibrium Approach
Each term on the right-hand side of this equation describes the separate sedimentation of one size of aggregate. However, unlike a reversible association (Equation 20.3.5), there is no relationship linking the different reference concentrations. Experimentally, a heterogeneous mixture leads to values of Mw that are either independent of protein concentration or, for reasons outside this discussion, decrease with increasing protein concentration. Furthermore, as higher molecular weight components are pelleted from the solution, Mw decreases, often dramatically, with increasing rotor speed.
where m, n, j, and k are stoichiometric coefficients describing the formation of species from a hetergeneous mixture of components. Although extracting thermodynamic quantities for extremely complicated systems is still beyond the capabilities of sedimentation equilibrium, two-component associations are amenable to study. The first step in studying such a system is to characterize the behavior of the individual components. Should one of the components reversibly self-associate, then the analysis becomes more difficult, though not intractable. Often, the analysis of a heterogeneous association is made easier by discriminating between components, either by spectral enhancement (Laue et al., 1993) or by spectral deconvolution (Schuck, 1994). Characterizing heterogeneous associations is an experimental challenge that requires the analysis of a large amount of data acquired over
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Current Protocols in Protein Science
a wide range of concentrations and at different concentration ratios of the components. The analysis of such data is outside the scope of this unit, but has been described elsewhere (Luckow et al., 1989). Diagnostically, a heterogeneous association exhibits increasing Mw with increasing protein concentration, though the extent of the increase in Mw will depend on the ratio of the concentrations of the components initially in the cell, as well as on the stoichiometry of assembly of the complex between the components. Also, there is likely to be some decrease in Mw with increasing rotor speed, especially if the two components differ significantly in Mb.
Nonideality Although a reversible association is a form of nonideality, in the context of sedimentation equilibrium the term nonideality is usually reserved for the repulsive interactions between protein molecules. These interactions tend to increase the apparent concentration, or chemical activity, ai, of a component, i, so that ai = γici, with the nonideality coefficient γi > 1. It is ai that is balanced by the gravitational potential, but ci that is measured by the optical systems. The effect of nonideality, therefore, is to suppress the observed concentration gradient (i.e., ci = ai/γi, with γi > 1). The two contributions to nonideality are the volume occupied by the protein molecules (called the excluded volume effect) and the charge-charge repulsion between like-charged
Table 20.3.1
proteins. Excluded volume contributions depend on the shape and flexibility of a molecule and are entropic. Charge-repulsion contributions are enthalpic and increase as the pair-wise product of the charges on the proteins, but decrease inversely with supporting electrolyte concentration. The two contributions to nonideality are additive, with charge-charge repulsion usually being the major contributor. Should the effects not be too severe, the effects of nonideality can be included in Equation 20.3.1 or Equation 20.3.5 by approximating the activity as a virial expansion of the concentration (Yphantis, 1964; Laue, 1995). Experimentally, nonideality is observed as a decrease in Mw with increasing concentration. For globular proteins having a moderate net charge (less than ∼10), nonideality will be minimal at concentrations <2 mg/ml in physiological solvents. For more highly charged proteins, charge-charge nonideality may be minimized by using solvents with higher salt concentrations. Highly asymmetric proteins, proteoglycans, and denatured proteins can exhibit significant excluded volume nonideality, which cannot be reduced by increased salt concentrations. Such nonideality must be taken into account in any subsequent analyses.
STRATEGY The five behaviors described above give rise to different rotor speed and protein concentration dependencies for Mw that may be used diagnostically (Table 20.3.1). This section will
Diagnostic Changes in Mw
Thermodynamic behavior
With increased rotor speed
With increased concentration
Ideal Ideal reversible self-association
None None
None Increase
Heterogeneous mixtures Ideal, reversible heterogeneous associations
Decrease
None
Slight decrease
Increase
Nonideality
None
Decrease
Comments
A slight decrease in Mw will be observed at rotor speeds high enough to deplete the cell concentration by effectively pelleting some of the sample There may be a slight decrease in Mw with increasing concentration The slight decrease in Mw with rotor speed is due to increased “unmixing” of the components at higher rotor speeds Salt-dependent concentration dependence indicates charge nonideality
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focus on establishing an experimental protocol for diagnosing and characterizing these different thermodynamic behaviors. Some experimental parameters are interdependent (e.g., protein concentration and optical detection method), and the correct choice of other experimental parameters will depend on the protein’s properties (e.g., rotor speed and monomer molecular weight). Preliminary sample handling is outlined in Figure 20.3.3 and is discussed in greater detail below.
Protein Concentration and Dilution It is necessary for measurements to cover a 100-fold range of protein concentrations in order to characterize the thermodynamic solution behavior of the protein. To characterize a reversible association, this protein concentration range must span a range sufficiently broad to include low concentrations where there is very little association as well as high concentrations where the associated species is the dominant form (Laue, 1995). Although covering this broad concentration range may seem to be a formidable challenge, a 100-fold concentration range may be covered in one experiment by examining three or four dilutions of a stock protein solution over an appropriate range of rotor speeds. The range of concentrations that is experimentally accessible is determined by the capabilities of the detectors and, for the absorbance detector, the characteristics of the protein. For
the refractive optical system, the lower concentration limit is ∼100 µg/ml, irrespective of the protein’s composition. For the absorbance optics, the lower concentration limit is determined partly by the sensitivity of the detector and partly by the protein’s extinction coefficient. For a “typical” protein, this corresponds to a lower usable concentration of ∼100 µg/ml at 280 nm, and ∼20 µg/ml at 230 nm. The general method for setting the protein concentration for the absorbance system is to measure the absorbance using a benchtop spectrophotometer. Keep in mind that the optical pathlength for the centrifuge centerpiece is 1.2 cm, rather than the usual 1.0 cm for a spectrophotometer cuvette. Thus, the absorbance readings for the centrifuge will be 20% higher than from the benchtop spectrophotometer. The upper concentration limit for a protein is often set by its solubility or availability. However, the optical systems also have upper limits of detection. For the refractive optics, the upper limit is not set by the concentration per se, but rather by the concentration gradient that develops during sedimentation (Yphantis, 1964; Laue, 1996). As a general rule, concentrations up to ∼100 mg/ml may be used with the refractive optics. For the absorbance optics, the upper concentration limit is determined by the sample absorbance, which should not exceed 1.5 OD at 280 nm, or 1.2 OD at 230 nm (Laue, 1996). Higher-concentration protein samples may be examined either by choosing
Sample preliminaries if sample has not been gel filtered during purification,do so before analysis
General sample handling estimate concentration and volume bring sample to dialysis equilibrium with buffer choose optical system and centerpiece material sedimentation equilibrium
Short column quick survey heteroassociations titrations
Analytical Centrifugation: Equilibrium Approach
sedimentation velocity
Short column detailed analysis low molecular weight heterogeneity
Figure 20.3.3 Flow chart of the protocol used to prepare a protein solution for analysis.
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r
A B C D
A B C cell type
two-channel double sector
six-channel high speed
eight-channel short column
absorbance compatible
yes
yes
no
dilution ratio not applicable from inner to outer radius
A 1:1 B 1:3 C 1:9
A 1:1 B 1:2 C 1:4 D 1:8
sample volume (µL)
50-110
15
50-450
Figure 20.3.4 Descriptions of centerpieces commonly used for sedimentation equilibrium analysis under operating conditions usually employed.
a wavelength where the absorbance readings fall in the range of 0.1 to 1.0 OD, or by using centerpieces with shorter optical pathlengths. The latter method is preferred, because merely changing wavelengths may lead to optical artifacts (Laue, 1996). At sedimentation equilibrium, the widest concentration span that can be measured from the meniscus to the base of a single channel covers a ∼10-fold range, with the bulk of the higher concentration data clustered in a few data points at the base. Thus, in order to have a sufficient number of data points covering the full 100-fold concentration range, it is necessary to make measurements on a dilution series of the initial sample. The dilution series should cover as wide a range of concentrations as possible, starting with the undiluted sample. All dilutions must be made using the dialysate (see Sample Handling, Solvent Considerations, and Initial Handling). The volume ratio for the dilutions depends on the type of centerpiece used in the experiment (Fig. 20.3.4; see Centerpiece and Window Type). If following these guidelines causes a sample to become too dilute for detection, a lower dilution ratio may be used. For multichannel centerpieces, the highest concentration sample is placed in the channel
closest to the center of the rotor, and the most dilute sample is placed in the channel closest to the edge of the rotor. By doing this, the dilute sample will be concentrated by the higher gravitational field at the edge of the rotor, thus providing a more useable signal from this sample, and steep concentration gradients are minimized by having the high concentration sample experience the lower gravitational field.
Optical Detection Method The advantages and disadvantages of the two optical systems may help in selecting which to use for a particular experiment. Table 20.3.2 presents the rationale for selecting which optical detector to use. If a sample does not lend itself to a clear choice based on these criteria, absorbance detection is easier to use. It is sometimes advantageous to use both optical systems. For example, the interference optical system provides a signal that is directly proportional to the weight concentration of the protein. Therefore, by dividing the absorbance reading by the interference reading, the extinction coefficient may be determined (Gray et al., 1995). Likewise, the ratio of the absorbance to the interference signal should remain constant and proportional to the extinction coefficient across a boundary. A systematic deviation in
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Table 20.3.2
Choosing an Optical System
Use absorbance if
Use refractive optics if
Use both to
Selectivity is required
Buffer absorbs in UV
Added sensitivity is required (e.g., using 230 nm) Solvent contains nondialyzable components
High precision is required
Determine the extinction coefficient Test for sample purity
Protein concentration is high
Extend the concentration range
Protein has low extinction coefficient Protein’s extinction coefficient varies with association state Short solution columns are used
this ratio indicates the presence of impurities (Laue, 1996). Finally, to study the widest concentration range, it is useful to combine data acquired using both optical systems. To do this, it is necessary to have appropriate conversion factors to adjust the concentrations obtained from the detectors (e.g., A280 or fringes) to a common concentration scale (e.g., mg/ml or molar). Procedures for doing this are available (Laue, 1996).
Centerpiece and Window Type
Analytical Centrifugation: Equilibrium Approach
Samples are contained in a centerpiece (Fig. 20.3.1). There are three types of centerpieces used for sedimentation equilibrium analysis (Fig. 20.3.4). The type used depends on the sort of question being addressed. For a simple analysis, the two-sector centerpiece may be used and has a number of advantages. (1) It is widely available, because it is also used for sedimentation velocity analysis. (2) It can accommodate a wide range of filling volumes. (3) It allows correction of data for optical imperfections using a water blank (Yphantis, 1964). (4) It permits both the absorbance and interference detectors to be used. (5) It is available in a variety of materials, thus providing compatibility with a wide range of solvents. To choose the filling volume, a compromise must be reached between a larger volume, which provides more data for analysis, and a smaller volume, which reduces the amount of sample needed and shortens the time required to reach equilibrium (Yphantis, 1960, 1964). A good compromise is a 110-µl sample, which results in a ∼3-mm-long column, ∼100 absorbance readings from meniscus to base,
and ∼10 hr to reach equilibrium (below). The greatest disadvantage of this sort of centerpiece is that it allows data to be collected for only one dilution per cell. The six-channel centerpiece offers the same advantages as the two-channel centerpiece, except that the six-channel centerpiece cannot be used for sedimentation velocity analysis and it holds a maximum of 110 µl sample per channel. These limitations are compensated by the increased number of dilutions that can be examined in a single cell. By using multiple cells, it is possible to cover a very wide concentration range in a single experiment. A version of this centerpiece is available that permits data correction for optical imperfections using water blanks acquired without disassembly of the windows, thus providing a 10-fold improvement in the precision of the data from the refractive optical system. The combination of allowing such data correction and providing for the analysis of a large number of samples makes this centerpiece the best choice for high-precision work. The eight-channel, or short-column, centerpiece is specifically designed for the rapid analysis of a large number of samples (Yphantis, 1960; Laue, 1992). Only 15 µl of sample is required, the time needed to reach equilibrium can be <1 hr, and up to 32 samples may be analyzed in a single experiment. This centerpiece is the best choice for conducting a rapid survey of behaviors under different solvent conditions (e.g., varying pH, ionic strength) or for working with scarce or unstable proteins. Although it is possible to analyze protein associations using short columns, fewer data points may be acquired over the shorter distance be-
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tween the meniscus and base, thus reducing the precision of the analysis. Furthermore, the geometry of the short-column centerpieces makes it difficult to acquire accurate data with absorbance optics. Also, because filling the centerpiece requires the removal of the top window, it is not possible to obtain an accurate water blank for data correction (Yphantis, 1960; Laue, 1992), again compromising the precision of the data acquired using these centerpieces. Cell windows are made from either fused silica or sapphire. Fused silica windows are far less expensive than sapphire windows, and are necessary for absorbance detection at wavelengths <260 nm. For the proper functioning of the refractive optical system, the increased hardness of sapphire windows is absolutely required, and they are acceptable for absorbance detection down to 260 nm. Keep in mind that sapphire windows are considerably more expensive to replace than fused silica windows, so they should be used only if needed for refractive detection.
Sample Handling, Solvent Considerations, and Initial Characterization Sample purity The steps for handling a sample are outlined in Figure 20.3.3. Only pure samples may be characterized using sedimentation equilibrium. Consequently, it is most important to eliminate aggregates and proteolytic fragments from the sample by gel filtration. Gel filtration should be done just prior to sedimentation analysis, as many proteins are prone to aggregation or proteolysis while stored. Solvent There are few limitations on the nature of the solvent used for sedimentation equilibrium. The solvent should include a pH buffer and sufficient salt to minimize the effects of charge on sedimentation (Laue and Stafford, 1999). For most proteins, an ionic strength of 30 to 50 mM is usually sufficient to suppress charge nonideality at protein concentrations <1 mg/ml. For highly charged proteins (e.g., histones or proteoglycans) or for high protein concentrations, the ionic strength may need to be higher and the data analysis must account for thermodynamic nonideality (see Methods of Analysis). If there are any unusual solvent components (e.g., high concentrations of dimethyl sulfoxide), it is important to check with
the manufacturer for its compatibility with the cell centerpiece material. If the absorbance optical system will be used, attention must be paid to the UV absorbance properties of the solvent components. Of particular concern at 280 nm is oxidized dithiothreitol or 2-mercaptoethanol. At shorter wavelengths (e.g., 230 nm), the list of problem components includes K+, Tris (or any nitrogencontaining component), materials with C=O bonds (e.g., EDTA), and others. In general, if absorbance optics are to be used, it is a good idea to measure the absorbance of the solvent against distilled water at the same wavelengths that will be used in the experiment. This absorbance should be <1.0 OD to minimize noise in the data (Laue, 1996). Solvent buoyancy The buoyancy of the solvent should be considered, especially if detergents or denaturants are used. The mass of a protein in solution (i.e., its effective mass) is its anhydrous mass minus the mass of the solvent it displaces. If M is the anhydrous protein molar mass (expressed in g/mol), the mass of the displaced solvent is Mvρ, where v is the protein’s partial specific volume (in ml/g) and ρ is the solvent density (in g/ml). It is the net or buoyant mass, Mb = M(1−vρ), that always appears in sedimentation equations. To a first approximation, v is the reciprocal of the density of the protein. Most often v is calculated from the composition of the protein (Laue et al., 1992; http://www.bbri. org/RASMB/rasmb.html; see Internet Resources). Any error in the buoyancy term propagates as a three-fold error in M. It is relatively simple to measure or calculate ρ to within a few tenths of a percent error, and v typically can be estimated to within 1%. This means that for a pure protein exhibiting ideal solution behavior, M can be determined to within 3%. Adjustment of the buoyancy factor is useful when examining proteins in detergent-containing solutions (Reynolds and McCaslin, 1985). In particular, the solvent density is adjusted so that the detergent is neutrally buoyant (i.e., it neither sinks nor floats). By making the detergent neutrally buoyant, it does not contribute to the protein’s buoyant mass. Consequently, all sedimentation is due to the protein, thus allowing determination of M without concern about the amount of detergent bound to the protein. It is even possible to characterize protein-protein interactions in detergent-containing solutions using this technique (Fleming et al., 1997).
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Buoyancy is also important when studying proteins in denaturing solvents, such as 8 M urea or 6 M guanidinium chloride, and methods have been developed to analyze sedimentation data in these solutions (Arakawa and Timasheff, 1985; Laue et al., 1992; http://www. bbri.org/RASMB/rasmb.html; see Internet Resources). Dialysis The sample should be brought to equilibrium by dialysis with the solvent to thermodynamically define the free concentrations (i.e., chemical activities) of the solvent components as being equal to that in the dialysate (Cassasa and Eisenberg, 1964). Therefore, the dialysate should be used for making any dilutions of the protein solutions. Dialysis also matches the optical properties of the solution and solvent, and the reference channels (Fig. 20.3.1) must be filled with dialysate so that accurate protein concentrations are measured by the absorbance optical system. For the refractive optical system, use of the dialysate for dilutions and as the optical reference is essential (Laue, 1996). Dialysis equilibrium may be achieved either by exhaustive dialysis (48 hr with stirring against three changes of solvent at a 100:1 v/v ratio of solvent to sample), by gel filtration, or by centrifugal gel filtration (Christoperson et al., 1979). There are certain solvent components, most notably detergents, that do not reach dialysis equilibrium using any of the methods listed. Under these circumstances, the experimenter must be aware that the free concentration of the nondialyzable component is not well defined, and that only the absorbance optics will give useful data (Laue, 1996).
Analytical Centrifugation: Equilibrium Approach
Protein concentration and sample volume Once the sample is prepared, a rough estimate should be made of the initial protein concentration and sample volume. This will help determine how many experiments can be conducted. For short-column centerpieces, a minimum of ∼50 µl stock solution will be necessary. For six-channel centerpieces, ∼250 µl will be needed. For the absorbance system, it is best if the initial protein concentration is ≥0.6 OD at whatever wavelength is to be used. For the refractive optical system, it is best if the initial protein concentration is ≥0.8 mg/ml. If the initial concentration is significantly less than these values, then the sample should be concentrated using a filter-type concentration method (e.g., a centrifugal concentrator, a vacuum dialysis
concentrator, or a passive dialysis concentrator). It should be noted that it is possible to retrieve samples after an experiment (typically 80% 90% of the initial amount), thus extending the number of experiments that can be performed on a single sample. Sedimentation velocity If there is sufficient material, sedimentation velocity analyses should be performed prior to conducting sedimentation equilibrium experiments. From such an analysis it is usually possible to determine whether or not the protein is associated in solution and whether the association is reversible (Laue and Stafford, 1999). For cases where there is no association, the monomer molecular weight can be determined. Likewise, for cases where complex formation is not reversible, it is often possible to determine the molecular weights and stoichiometries of the associated species. For many proteins, this level of characterization is sufficient, and the lengthier protocols of sedimentation equilibrium are not needed. However, sedimentation equilibrium will be needed to characterize reversible associations or thermodynamic nonideality, and will be necessary if hydrodynamic nonideality interferes with the interpretation of the sedimentation velocity experiments (Laue and Stafford, 1999).
Rotor Speed The shape of the concentration gradient (Fig. 20.3.2) largely depends on the mass of the protein and the rotor speed. For proper data analysis, a range of rotor speeds must be used that starts low enough that any large species may be examined without being packed at the cell bottom, and ends high enough that the smallest protein species will generate a gradient with significant curvature (Yphantis, 1964; Johnson et al., 1981). Usually, data are gathered at three or four rotor speeds, with the highest rotor speed being two to three times that of the lowest rotor speed. For a typical protein, a good starting speed can be calculated from rpm = 4.53 × 106/M1/2 if there is an estimate for the molecular weight of the protein. If the molecular weight of the protein is unknown, the lowest rotor speed will have to be determined empirically. To select the lowest rotor speed empirically, start centrifuging the samples at 3000 rpm to see whether a pronounced sigmoidal concentration gradient develops after 15 to 30 min. If no such concentration gradient develops, the rotor speed should be doubled to 6000 rpm, and the same test performed. This procedure should
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be continued until significant concentration redistribution is observed. Once this lower rotor speed is established, three or four higher rotor speeds should be chosen so that the highest rotor speed is two to three times higher than the lowest rotor speed. It is imperative that an experimental procedure start with the lowest rotor speed and always proceed from lower to higher speeds. If it is found that the initial rotor speed is too high, then it is best to stop the run, stir up the cell contents by gently rocking the rotor, and start the experiment over at a lower rotor speed. Simply decreasing the rotor speed may result in unreasonably large times required to reach equilibrium.
the slow aggregation of a protein, equilibrium may never be established. (4) Thermodynamic nonideality will shorten the time needed to achieve equilibrium. An empirical demonstration of equilibrium is often made by looking for a zero concentration difference when two concentration profiles, taken an hour or so apart, are subtracted (Yphantis, 1964). A better procedure is to monitor the minimized average differences in concentration profiles over time, as is done by MATCH (David Yphantis and Jeff Lary, available from ftp://rasmb.bbri.org). The results from MATCH also indicate the intrinsic noise level of the data. This is useful information when analyzing data using a nonlinear least squares approach.
Temperature The experimental temperature may be kept between 0° and 40°C. To prevent the possibility of a leak, the cell should be assembled (or retightened) at the same temperature as the experiment. In cases where the protein undergoes an association that is strongly temperature-dependent, it is important to have the samples, cells, and rotor preequilibrated at the experimental temperature so that sedimentation equilibrium is reached in a reasonable period of time.
Time to Reach Equilibrium Whereas hydrodynamics affects the length of time needed to reach equilibrium, only thermodynamics affects the equilibrium concentration gradient. The exact length of time needed to establish sedimentation equilibrium is a complicated function of the experimental parameters, and cannot be accurately predicted except in the simplest cases (Yphantis, 1960, 1964). Though attainment of equilibrium must be determined empirically, the relative time needed follows a few simple rules. (1) The equilibration time increases as the square of the distance from the meniscus to the base of a solution column (Fig. 20.3.2). Thus, the use of shorter solution columns will speed up equilibration considerably, although with loss of precision (Yphantis, 1960). (2) The time needed to reach equilibrium increases proportionally with the viscosity of the sample. For aqueous solutions, this leads to a temperature dependence such that a sample will take ∼1.5 times as long to reach equilibrium at 4°C as at 20°C, and will take ∼0.6 times as long at 40°C as at 20°C. (3) Kinetic processes, such as mass-action associations, usually increase the time needed to reach equilibrium. For some processes, such as
DATA ANALYSIS AND INTERPRETATION At the present time there is no comprehensive manual for the analysis of sedimentation equilibrium data. However, there are reviews that reference papers discussing the analysis of particular kinds of solution behavior (McRorie and Voelker, 1993; Laue, 1995; Laue and Stafford, 1999). Presented here are some guidelines for analyzing sedimentation equilibrium data, and a brief description of the most common analysis methods (also see UNIT 20.1). Depending on the question a researcher is trying to answer, the most difficult part of a sedimentation equilibrium experiment may be the accurate analysis of the data. If the question being addressed is largely qualitative (i.e., determining whether a protein is a multimer in solution) or diagnostic (i.e., determining the concentration-dependent behavior of a protein), then the analysis is usually straightforward. However, nature sometimes endows proteins with complicated solution behavior (association, nonideality) that may require a more detailed quantitative analysis (i.e., determining the stoichiometry and association energy of a complex). If solution behavior must be understood in greater detail, the analysis becomes more difficult. The difficulty stems from three sources. First, the raw data from an equilibrium sedimentation experiment consists of an exponential concentration curve, one of nature’s most boring functions (Fig. 20.3.2). Even if the curve is distorted somewhat by nonideality or self-association, such curves have no features (e.g., peaks or boundaries) to provide an experimenter with an intuitive feeling for what is going on in the solution. Second, fewer and
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fewer biochemists and molecular biologists are trained in solution thermodynamics. Thus, while the use of computer programs makes it possible to extract thermodynamic parameters from sedimentation equilibrium data, researchers are often uncertain about whether a parameter is significant or even reasonable. Third, solution thermodynamic data may lead to ambiguous interpretations (Laue, 1995). For example, it can be difficult to determine a unique stoichiometry to describe a reversible association. Such ambiguity is disconcerting to a researcher testing a nonambiguous hypothesis. This ambiguity is not a shortcoming of sedimentation equilibrium, nor of the researcher, but is an inherent trait of nature: thermodynamics and nature do not rely on models, but molecular biology does. It is unsettling to researchers to learn that sedimentation equilibrium is providing an accurate view of the ambiguities inherent in nature. There is no simple remedy for these problems except experience. For a novice in this area, it is best to contact an expert for guidance through the early stages of experimental design and data analysis. Expert advice is available from the Center to Advance Molecular Interaction Science (CAMIS, [email protected]) and from the ultracentrifuge users group ([email protected]). Exp er ien ce is gained rapidly, and with it, confidence.
Methods of Analysis
Analytical Centrifugation: Equilibrium Approach
Sedimentation equilibrium data can be analyzed by methods that fit into one of two broad categories, graphical analysis or nonlinear least squares fitting (Ralston, 1993; Laue, 1995, and references therein). Each of these methods has its advantages and disadvantages. The graphical methods, as exemplified by molecular weight moment analysis (Roark and Yphantis, 1969) and the Omega (or Psi) analysis (Haschemeyer and Bowers, 1970; Morris and Ralston, 1985), provide direct model-independent insight into solution thermodynamics. These graphs provide diagnostic information that can be useful guides for subsequent nonlinear least squares analyses. However, all graphical methods require manipulation of the data, which distorts the error space without increasing the information content. Hence, it is the author’s opinion that the extraction of thermodynamic parameters should be done by direct fitting of the data using nonlinear least squares analysis (Johnson et al., 1981). Nonlinear least squares analysis of the primary data—i.e., c(r) as a function of r—is the
most popular method for extracting thermodynamic parameters from sedimentation equilibrium experiments, with NONLIN the most widely used program for this purpose (Johnson et al., 1981). What makes NONLIN particularly powerful is that it permits the analysis of several data sets simultaneously, with some of the fitting parameters considered global to all data sets, while others are considered local to each data set. Data acquired at multiple rotor speeds, at different concentrations, and using both the absorbance and interference detectors may be fit simultaneously. By analyzing large quantities of data acquired over a wide range of concentrations, it is possible to obtain monomer molecular weights, association constants, stoichiometries, and virial coefficients for moderately complex systems. Versions of NONLIN for various computer operating systems are available free from the Reversible Association in Structural and Molecular Biology (RASMB) site, as are directions for its use (ftp://rasmb.bbri.org).
Data Interpretation A complete description of the interpretation of sedimentation data is beyond the scope of this unit. However, a detailed review exists (Laue et al., 1992) and an excellent computer program is available for assisting in the interpretation of sedimentation data (http://www. bbri.org/RASMB/rasmb.html; see Internet Resources). It is recommended that these be consulted.
LITERATURE CITED Arakawa, T. and Timasheff, S.N. 1985. Calculation of the partial specific volume of proteins in concentrated salt and amino acid solutions. Methods Enzymol. 117:60-65. Casassa, E.F. and Eisenberg, H. 1964. Thermodynamic analysis of multicomponent solutions. Adv. Protein Chem. 19:287-395. Christopherson, R.I., Jones, M.E., and Finch, L.R. 1979. A simple centrifuge column for desalting protein solutions. Anal. Biochem. 100:184-187. Fleming, K.G., Ackerman, A.L., and Engelman, D.M. 1997. The effect of point mutations on the free energy of transmembrane alpha-helix dimerization. J. Mol. Biol. 272:266-275. Gray, R.A., Stern, A., Bewley, T., and Shire, S.J. 1995. Rapid Determination of Spectrophotometric Absorptivity by Analytical Ultracentrifugation. Application Data Sheet A-1815A. Beckman Coulter Instruments, Fullerton, Calif. Haschemeyer, R.H. and Bowers, W.F. 1970. Exponential analysis of concentration or concentration difference data for discrete molecular
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weight distributions in sedimentation equilibrium. Biochemistry 9:435-445. Johnson, M.L., Correia, J.J., Yphantis, D.A., and Halvorson, H.R. 1981. Analysis of data from the analytical ultracentrifuge by nonlinear leastsquares techniques. Biophys. J. 36:575-588. Laue, T.M. 1992. Short Column Sedimentation Equilibrium Analysis for Rapid Characterization of Macromolecules in Solution. Technical Information DS835. Beckman Instruments, Palo Alto, Calif. Laue, T.M. 1995. Sedimentation equilibrium as a thermo dy namic to ol. Methods Enzymol. 259:427-452.
Rivas, G.A. and Minton, A.P. 1993. New developments in the study of biomolecular associations via sedimentation equilibrium. Trends Biochem. Sci. 18:284-287. Roark, D.E. and Yphantis, D.A. 1969. Studies of self-associating systems by equilibrium ultracentrifugation. Ann. N.Y. Acad. Sci. 164:245278. Schuck, P. 1994. Simultaneous radial and wavelength analysis with the Optima XL-A analytical ultracentrifuge. Progr. Colloid Polym. Sci. 94:113. Senear, D.F. and Teller, D.C. 1981. Thermodynamics of concanavalin A dimer-tetramer self-association: Sedimentation equilibrium studies. Biochemistry 20:3076-3083.
Laue, T.M. 1996. Solution Interaction Analysis: Choosing Which Optical System of the Optima XL-I Analytical Ultracentrifuge to Use. Application data sheet A-1821A. Beckman Coulter Instruments, Fullerton, Calif.
Van Holde, K.E. 1985. Sedimentation. In Physical Biochemistry, pp. 110-136. Prentice-Hall, Englewood Cliffs, N.J.
Laue, T.M. and Stafford, W.F. III. 1999. Modern applications of analytical ultracentrifugation. Annu. Rev. Biophys. Biomol. Struct. 28:75-100.
Yphantis, D.A. 1960. Rapid determination of molecular weights of peptides and proteins. Ann. N.Y. Acad. Sci. 88:586-601.
Laue, T.M., Shah, B.D., Ridgeway, T.M., and Pelletier, S.L. 1992. Computer-aided interpretation of analytical sedimentation data for proteins. In Analytical Ultracentrifugation in Biochemistry and Polymer Science (S.E. Harding, A.J. Rowe, and J.C. Horton, eds.) pp. 90-125. Royal Society of Chemistry, Cambridge.
Yphantis, D.A. 1964. Equilibrium ultracentrifugation of dilute solutions. Biochemistry 3:297- 317.
Laue, T.M., Senear, D.F., Eaton, S.F., and Ross, J.B.A. 1993. 5-Hydroxytryptophan as a new intrinsic probe for investigating protein-DNA interactions by analytical ultracentrifugation. Study of the effect of DNA on self-assembly of the bacteriophage cI repressor. Biochemistry 32:2469-2472. Luckow, E.A., Lyons, D.A., Ridgeway, T.M., Esmon, C.T., and Laue, T.M. 1989. Analysis of the interaction of coagulation factor V heavy chain with prothrombin and prethrombin-1 by analytical ultracentrifugation. Role of activated protein C in regulating this interaction. Biochemistry 28:2348-3454. McRorie, D.K. and Voelker, P.J. 1993. Self-associating Systems in the Analytical Ultracentrifuge. Beckman Coulter Instruments, Fullerton, Calif. Morris, M. and Ralston, G.B. 1985. Determination of the parameters of self-association by direct fitting of the omega function. Biophys. Chem. 23:49-61. Ralston, G.B. 1993. Introduction to Analytical Ultracentrifugation. Beckman Coulter Instruments, Fullerton, Calif. Reynolds, J.A. and McCaslin, D.R. 1985. Determination of protein molecular weight in complexes with detergent without knowledge of binding. Methods Enzymol. 117:41-53.
Yphantis, D.A., Correia, J.J., Johnson, M.L., and Wu, G. 1978. Detection of heterogeneity in selfassociating systems. In Physical Aspects of Protein Interactions (N. Catsimpoolas, ed.) pp. 275303. Elsevier/North-Holland, Amsterdam.
INTERNET RESOURCES ftp://rasmb.bbri.org Site for MATCH program (David Yphantis and Jeff Lary), used for monitoring minimized average differences in concentration profiles over time. [email protected] Contact for Center to Advance Molecular Interaction Science (CAMIS), for assistance with data analysis. [email protected] Contact for the ultracentrifuge users group at Boston Biomedical Research Institute, for assistance with data analysis. http://www.bbri.org/RASMB/rasmb.html Site for Sednterp (J. Philo, D.B. Hayes, and T.M. Laue), a program for the interpretation of sedimentation data for proteins. The program uses databases so that it is readily customized and extended.
Tom Laue University of New Hampshire Durham, New Hampshire
Rickwood, D. 1984. Centrifugation: A Practical Approach, 2nd ed. IRL Press, Washington, D.C. Quantitation of Protein Interactions
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Titration Microcalorimetry
UNIT 20.4
Isothermal titration calorimetry (ITC) is perhaps the most rigorous commercially available method for characterizing protein-ligand interactions. Interactions are detected by ITC from the intrinsic heat (binding enthalpy) change of the reaction. The method is applicable to a multitude of biomolecular interactions, including those that involve proteins, nucleic acids, carbohydrates, lipids, small organic molecules, metals, ions, and protons. ITC should be used when an accurate, quantitative measure of the ligand-binding properties of a protein is desired. A major strength of ITC over other methods (e.g., surface plasmon resonance) is that the technique is applicable to native, unmodified proteins in solution. This is important for proteins that lose or change their functional behavior when chemically modified or attached to a surface. ITC is also useful for evaluating qualitative questions such whether a proposed binding interaction occurs to begin with, or for quantitatively measuring the concentration of functionally active protein. Finally, if executed with proper control experiments, ITC can be a rich source of thermodynamic information about the molecular binding mechanism. STRATEGIC PLANNING There are two general approaches for undertaking an ITC study: determination of observed binding parameters and determination of biochemical binding parameters. Both approaches yield quantitative information about binding, but they differ in the extent to which they provide rigorous, biochemically well defined binding parameters. In the former approach the binding parameters tell us qualitatively about the binding interaction, but are not rigorous enough to be assigned unambiguously to a specific biochemical reaction. In the later approach, the goal is to rigorously determine physically well-defined parameters for a specific biochemical binding reaction. In this unit we separate the two analytical approaches into Basic Protocols 1 and 2, respectively. The differences in physical meanings between parameters obtained from the two protocols are listed in Table 20.4.1. Determination of observed binding parameters by Basic Protocol 1 is usually straightforward and can be done in a day. In these studies, one is interested in answering questions such as the following. Does a ligand bind a given protein? What is the affinity for an assumed simple, well-behaved bimolecular interaction? What fraction of a protein sample is functionally competent for binding? What is the likely stoichiometry for the interaction? These questions cover a wide range of practical issues and can be addressed readily by ITC according to Basic Protocol 1. Determination of biochemically well-defined molecular binding parameters by Basic Protocol 2, on the other hand, requires a different mindset and a significantly greater expenditure of reagents and time (which is true to some degree for all quantitative protein interaction studies, not just ITC). In these studies, the goal is to obtain a rigorous molecular-level characterization of the binding mechanism. For example, one may wish to measure the true molecular binding thermodynamics for an interaction (e.g., the binding Gibbs free energy and enthalpy changes) in order to understand what is happening at the molecular level in terms of hydration or protonation effects coupled to binding. The control experiments and corrections needed to resolve this information are described in Basic Protocol 2. Basic Protocol 2 is also more reliant than Basic Protocol 1 on the support protocols indicated in this unit (i.e., accurate determination of reactant concentrations, Contributed by Michael L. Doyle Current Protocols in Protein Science (1999) 20.4.1-20.4.24 Copyright © 1999 by John Wiley & Sons, Inc.
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thermal stability measurements, and self-association measurements). Once the biochemical binding thermodynamics have been determined for an interaction using Basic Protocol 2, one would naturally seek to use the thermodynamic information to learn about the binding molecular mechanism. The Commentary section of this unit mentions briefly some current progress along these lines. However, it should be recognized that molecu-
Table 20.4.1 Comparison of the Physical Meaning of Binding Parameters Determined from Basic Protocols 1 (Observed Values) Versus Those Obtained from Basic Protocol 2 (Biochemical Values)
Parametera
Comments
Kdobs
Measured by a single ITC experiment. May or may not equal true molecular affinity. May include contributions from complicating side reactions such as ligand-induced dimerization and unfolding.
Kd°′
Pertains to a specific molecular interaction under defined conditions of pH, temperature, ionic strength, etc. Reflects all aspects of binding, including conformational changes, changes in hydrogen bonds, van der Waals interactions, ionic bonds, hydration, protonation, ion binding, etc. Aggregation and unfolding side reactions confirmed not to be present
∆Hobs
Measured by a single ITC experiment. Usually includes artifactual heats due to buffer ionization. May or may not equal/approximate true molecular binding ∆H, depending on relative contribution from buffer ionization heats.
∆H°′
Pertains to a specific molecular interaction under defined conditions, as described for biochemical Kd. Corrected for buffer ionization heat artifacts, and possible coupled unfolding or aggregation side reactions.
∆Cpobs
Measured by ITC experiments done at several temperatures in a single buffer. May or may not equal/approximate true molecular binding ∆Cp, depending on temperature-dependence of buffer ionization heats
∆Cp°′
Pertains to a specific molecular binding interaction, as described for biochemical Kd. Corrected for buffer ionization heat artifacts over the temperature range studied. Since determination of ∆Cp requires measurements at various temperatures, ∆Cp may include changes in the nature of the molecular interaction with temperature, such as changes in extent of coupled protonation, or partial thermal unfolding.
Molar binding ratio (observed stoichiometry)
Measured by a single ITC experiment
Molecular stoichiometry
Equal to molar binding ratio from an ITC experiment only if both reactants are 100% active and their concentrations are accurately determined. Otherwise can be measured directly for protein-protein interactions by static light scattering or analytical ultracentrifugation
aObserved parameters are represented by the superscript “obs”; biochemical parameters are represented by the degree
Titration Microcalorimetry
and prime signs.
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lar-level interpretation of binding thermodynamics is a challenging and subtle subject, and an ongoing area of research. Proper interpretation of the data will in many cases require a more comprehensive understanding of the biothermodynamics field than is given in this unit. DETERMINING A MOLAR RATIO, OBSERVED AFFINITY (Kdobs), AND OBSERVED BINDING ENTHALPY CHANGE (∆Hobs) BY ITC
BASIC PROTOCOL 1
This protocol describes how to carry out an ITC experiment. The results can be used to: (1) test whether a binding interaction occurs, (2) measure the observed affinity of an interaction, (3) measure molar binding ratio of an interaction, and (4) measure the observed binding enthalpy change. An example data set from an ITC experiment is shown in Figure 20.4.1. Note that the observed binding enthalpy change ∆Hobs) that is measured by this protocol is not corrected for possible heats of buffer ionization and thus may not be equal to the molecular enthalpy change for the reaction being investigated (see Basic Protocol 2 and Table 20.4.1). Also note that the measured molar binding ratio may not be equal to the true molecular binding stoichiometry. The molar binding ratio is an empirical value equal to the ratio of the concentration of one reactant that reacts with a given concentration of another. It is related to the true molecular stoichiometry only to the extent that both reactant concentrations were previously measured accurately, and both reactants are 100% active for binding. Finally, the observed affinity (Kdobs) measured by this protocol may or may not pertain rigorously to a well-defined biochemical equilibrium binding constant, since no attempt is made to test whether the reactants undergo possible side reactions such as changes in aggregation state or thermal unfolding. Testing for these types of side reactions is described in Basic Protocol 2. Materials Proteins and ligands of interest Appropriate pH buffer Isothermal titration calorimeter (MicroCal, Calorimetry Sciences, Thermometric AB) 2.5-ml Hamilton gas-tight syringe with blunt-end needle long enough to reach volume of sample cell UV-visible spectrophotometer Micro combination pH electrode Degassing apparatus Nonlinear least-squares regression software (generally provided by the instrument manufacturer) Prepare the ITC 1. If necessary, calibrate the ITC instrument. Titration microcalorimetry data are measured with isothermal titration calorimeters (ITC) that have been designed specifically for the high-sensitivity requirements of characterizing biomolecular interactions. ITC instruments are calibrated by the manufacturers when delivered. However, occasionally the user should test the accuracy of the instrument. This can be done with some instruments according to specific instructions provided by the manufacturer. Alternatively one may wish to calibrate using an independent reaction with known binding thermodynamics. A suitable reaction system for this is the interaction of barium with 18-crown ether (Liu and Sturtevant, 1995). Both of these reactants are available in stable and high-purity form, and the binding affinity, stoichiometry, and enthalpy change have been documented in the literature. Quantitation of Protein Interactions
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Estimate reactant quantities needed The amounts of the reactants required for ITC depend on the affinity and, to a lesser extent, the binding enthalpy change. These parameters are unique to each reaction, and hence reagent needs for ITC can vary considerably. These parameters may not be known in advance at the start of an ITC study. It may be necessary to make assumptions about these parameters, carry out initial titrations, and optimize experimental design as needed.
A
Time (min) 0
20
40
60
80
100
120
µcal/sec
0
–50
–100
B kcal/mol of barium injected
0
–2
–4
–6
–8 0
0.5
1.0
1.5
2.0
2.5
Molar ratio (barium per crown ether)
Titration Microcalorimetry
Figure 20.4.1 Isothermal titration calorimetry (ITC) data for titration of 5.7 mM 18-crown-6 ether with 5-µL injections of 103 mM barium. (A) Raw power (µcal/sec) versus time tracing. At each injection an exothermic “spike” is seen. The area under each spike is proportional to the heat of binding of barium to crown ether. The first injection is only a preliminary injection for experimental set up purposes and is ignored during data analysis. (B) Amount of heat measured at each injection normalized to the number of moles of barium injected (kcal/mol) versus molar ratio of cumulative barium added per crown ether in the cell. Nonlinear least squares analysis of the data yields a molar ratio of 1.01 ± 0.01 barium ions per crown ether, an observed binding ∆H = −8.17 ± 0.01 kcal/mol barium, and an observed equilibrium dissociation constant Kd = 290 ± 2 µM.
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2. Estimate the expected equilibrium dissociation constant (Kd) for the binding interaction from existing information if possible. If not, make assumptions about the affinity (e.g., as in step 4b). The dissociation equilibrium constant, Kd, for a protein-ligand interaction is defined for a biochemical equilibrium M⋅L = M + L (where M is the protein macromolecule and L is the ligand) as: Kd = [M][L]/[M⋅L]. The affinity of an interaction can also be reported as an association equilibrium constant (e.g., Ka). Ka has units of inverse molarity. Kd has units of molarity, and is equal to the reciprocal of the association binding equilibrium constant (Kd = 1/Ka).
3. Choose which reactant goes into the cell (the “cell reactant”) and which goes into the syringe (the “syringe reactant”). Generally, the more expensive reactant would be placed inside the microcalorimeter reaction cell and the other reactant would be loaded into the injection syringe. The reasons for this are, first, the syringe has a dead volume and hence requires an excess of reactant to fill it, and second, that titrations are always carried out using an excess of the syringe reactant. The solubilities of the two reactants may also be an important factor. Since the syringe reactant must be prepared to ∼20-fold higher concentration, the relative solubilities of the reactants may govern which can go into the syringe. For proteins that have multiple ligand binding sites, it may be desirable to have the ligand in the syringe. In favorable cases, this may aid in deconvoluting the heterogeneous binding thermodynamics.
4a. If the expected Kd can be estimated: Calculate the concentration of cell reactant to use from the expected Kd. The concentration of the cell reactant should ideally be 10- to 50-fold higher than the Kd of the interaction (see Fig. 20.4.2), or at least 5 µM for very tight (Kd <100 nM) binding interactions. This calculation assumes the cell reactant has a single binding site for the syringe reactant. For multiple binding site cases, the relevant units of concentration are based on the number of equivalents of binding sites. For example, one would use 2.5 µM of a protein molecule that has 2 binding sites per molecule (5 µM binding sites total).
4b. If the Kd cannot be estimated beforehand: Assume the Kd is 1 µM or tighter and use a cell reactant concentration of 5 µM. The reagent needs for all interactions tighter than 1 µM are similar, and represent, approximately, the minimal amount needed for an ITC experiment. In general one would like to measure at least several titration heats of a few microcalories or more of binding heat signal. Any less than this could be too small to reliably measure with current instrumentation. Alternatively, if the Kd is suspected to be weaker than 1 µM, then a proportionately higher cell reactant concentration will be needed. For weak binding interactions, one may need to use large quantities of reactants in order to drive the binding reaction to a measurable extent. IMPORTANT NOTE: In order to detect a binding interaction, the concentrations of reactants need to be studied at concentrations higher than the expected Kd. Figure 20.4.1 shows simulated curves of the relation between reactant concentrations, affinity of the interaction, and the expected curvature and quality of the data. As pointed out previously (Wiseman et al., 1989), one should aim to have the concentration of the cell reactant between 4-fold and 1000-fold higher than the Kd of the binding reaction. Concentrations less than this may not have sufficient curvature to establish whether or not a binding interaction exists between the ligand and protein. On the other hand, concentrations above 1000-fold Kd should allow high-precision determination of the molar binding ratio and observed enthalpy change, but will not have enough information in the transition region to allow the Kd to be resolved.
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Fraction of injectant bound
1.0
∞
1000
30
200
0.8 5
0.6
0.4 1 0.2 0.2 0.0 0.0
0.5
1.0
1.5 Molar ratio
2.0
2.5
3.0
Figure 20.4.2 Resolvability of binding parameters from ITC data depends on the concentration of the cell reactant relative to the affinity of the interaction. Shown, as fraction injectant bound versus molar ratio of syringe reactant added per cell reactant, are simulated ITC curves at various “C parameter” values, (following the analysis of Wiseman et al. (1989). The C parameter is the ratio of concentration of reactant in the cell divided by the dissociation equilibrium constant, Kd. When the concentration of cell reactant is too low (C < 1), the data shape is very shallow and does not allow for determination of any of the interaction constants (Kd, molar ratio, or enthalpy change). When the cell reactant concentration is too high (C > 1000), the binding molar ratio and observed enthalpy change are well determined, but there is insufficient information in the transition region to resolve the affinity constant, Kd. Optimal C values to use for experiments are ∼10 to 100.
5. Calculate the concentration of syringe reactant as 20-fold higher than that of the cell reactant. 6. Calculate the total amount of cell and syringe reactants needed, based on the concentrations and volumes to be used. The volumes needed depend somewhat on the make and model of the ITC instrument being used. As ballpark figures, one may typically need 2.5 ml of the cell reactant solution and 1 ml of syringe reactant. This volume of cell reactant is enough for only one titration, whereas the volume of the syringe reactant will usually be enough for a few titrations.
Select experimental conditions 7. Choose a pH buffer that has a low heat of ionization in order to minimize artifactual heats of buffer ionization. Avoid using pH buffers that have large proton ionization heats (e.g., Tris). Any pH mismatches between syringe and cell reactant solutions will cause large heats of mixing during the titration, when the pH of the titrant solution gets titrated upon injection into the cell. Phosphate is a good buffer to use and has a small proton ionization enthalpy. Refer to Table 20.4.2 and references therein for a listing of buffers and their ionization heats.
Titration Microcalorimetry
Aside from minimizing mixing heats (steps 6 and 7), solution conditions with ITC are flexible. ITC can be done over a range of pH conditions, in the presence of glycerol, sucrose, reducing agents, or cosolvents such as 1% to 10% DMSO. Solvents that evaporate readily might create evaporative heat noise. Other chemicals that would undergo reactions during the experiment would also cause noise. Check with the instrument manufacturer for possible conditions (such as strong acids or bases) that might damage the calorimeter cell.
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Table 20.4.2 Ionization pKa, Enthalpy Change, and Heat Capacity Change Values for Some Buffers Commonly Used in ITC Studiesa
pKa(25)
∆Hion(25) (kcal/mol)b
∆Cpion (kcal/mol/°C)c
Cacodylate
6.3
−0.6d
NA
Citrate (pK3) Phosphate (pK2)
6.4 7.2
−0.8 0.9
−0.060 −0.040
PIPES MES
6.8 6.1
2.7e 3.7d
+0.005e NA
MOPS HEPES
7.2 7.5
5.3d 5.7f
NA NA
ACES Tricine
6.8 8.1
7.5d 7.7b
NA NA
Imidazole
7.1 8.1
8.7 11.4
+0.005 −0.015
Buffer
Tris
aValues refer to 25°C and are from Christensen et al. (1976) unless otherwise
indicated. These parameters depend on ionic strength and other solution conditions. ∆Cpion is assumed independent of temperature. Refer to Christensen et al. (1976) for a detailed compilation of ionization ∆Hion and ∆Cpion values. b∆Hion can be calculated as a function of temperature from Kirchhoff’s Law:
∆Hion(T) = ∆Hion(25) + ∆Cpion[T − 25]. cNA, not available. dValues at 30°C from Murphy et al. (1993). eValues from Baker and Murphy (1997). fValues calculated from dpK /dT reported by Stoll and Blanchard (1990). a
Match chemical compositions of reactant solutions 8. Prepare the reactant solutions according to the experimental needs outlined in steps 9 and 10. It is essential that the syringe and cell reactant solutions be precisely matched chemically in terms of pH and solvent components. Differences in salt or even small differences in solvent compositions will cause substantial background heats of mixing and may mask the binding heats for the reaction of interest. Dialysis is a preferred method for matching the chemical compositions of reactants to a set of buffer conditions. Microspin columns are also useful in this regard. If necessary, manual adjustment of concentrations and pH values can be done.
9a. For reactants that have masses large enough to be dialyzed: Dialyze reactant against buffer containing the desired solution conditions of the experiment. All solvent components, as well as the pH, should be matched to the dialysis buffer.
9b. For low-molecular-weight components that cannot be dialyzed: Dissolve samples directly into the reaction buffer. Record weight (solids) or volume (liquids) of sample dissolved and final volume of solution in order to allow the reactant concentration to be accurately calculated. Measure and compare the pH values of the dissolved sample and the dialyzed sample. pH should match to within 0.1 pH units. If not, adjust pH with small amounts of acid or base, being careful not to denature any protein reagents. It is preferable to adjust the pH of nonprotein reactants to match those of the dialyzed protein sample. A micro combination pH electrode is often needed for expensive reactants in small volumes.
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If dissolution of one of the reactants (such as a small organic molecule) requires addition of a nonaqueous solvent such as DMSO, record the final concentration of solvent used and add a matching amount to the other reactant.
Measure reactant concentrations 10a. For protein reactants: Determine concentrations by measuring absorbance at 205, 210, and/or 280 nm, and calculating the concentration from molar extinction coefficients. The accuracy in determining the concentration of the syringe reactant is critical to the accuracy of the binding thermodynamic parameters obtained by ITC. Molar extinction coefficients for proteins can usually be calculated to within 5% to 10% accuracy if the amino acid composition is known (UNIT 3.2). It is preferable that the UV-visible spectrum be measured from ∼240 to 400 nm. This will provide a quality check on the material. The spectral baseline should be nearly flat from 320 to 400 nm. Significant sloping baselines indicate potential light-scattering effects and may be diagnostic of the protein sample being aggregated (and possibly inactive). If the A280 measurement is in question, absorbance at 205 and/or 210 nm provides an independent measure of total amino acid concentration with a potential accuracy near 2% (see UNITS 3.1 & 3.2; a general discussion about the potential problems associated with the above approaches is found in UNIT 3.2).
10b. For non-protein reactants: Establish concentration by gravimetric or volumetric analysis using the weights and volumes recorded in step 9b. Alternatively measure the absorbance of the sample and calculate its concentration from a molar extinction coefficient, if available. IMPORTANT NOTE: The accuracy of the affinity and binding enthalpy change are directly proportional to the accuracy (and functional activity) with which the concentration of the reactant in the syringe was determined. For determination of the affinity constant, this may not be a big concern (a factor of two error is usually not a concern); however, small inaccuracies in the binding enthalpy change can be very misleading if one is to interpret the binding thermodynamics in a detailed way. It is thus important to determine the syringe reactant concentration as accurately as possible to obtain an accurate enthalpy change. On the other hand, the accuracies of the affinity and enthalpy change do not depend on the accuracy of the concentration of the cell reactant. The accuracy of the molar ratio (i.e., as a measure of the true molecular binding stoichiometry) depends equally on the accuracy and functional activity of the concentrations of both syringe and cell reactants.
Degas reactant solutions 11. Once reactant concentrations have been established, degas the samples by gently stirring under a vacuum for 10 min. Fill calorimeter cell and syringe 12. Prerinse the calorimeter cell with buffer in order to cleanse the reaction cell. Using a 3-ml syringe with blunt end needle long enough to reach the bottom of the sample cell, inject buffer into the cell just until the buffer arrives at the opening of the reaction cell port. Details for filling the calorimeter cell and syringe are specific to instrument makes and models. These instructions are but one example.
13. Remove the buffer from the cell using the 3-ml syringe. It will not be possible to remove all the sample buffer, and thus the sample will become slightly diluted (e.g., by ∼2%) when added to the cell.
Titration Microcalorimetry
14. Load the sample into the cell using the 2.5-ml Hamilton syringe. Place the needle on the bottom of the cell and slowly inject the sample while moving the syringe up and down a few millimeters so as to dislodge any air bubbles that may otherwise become trapped. When the sample can be observed to be coming up out of the reaction cell port, begin removing the syringe while continuing to slowly inject sample, so that air
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bubbles are not allowed to go down into the cell. Remove excess solution from around the cell porthole but leave sample cell filled to the top of the porthole. 15. Load the syringe with its reactant. Draw the solution up into the calorimeter syringe using, for example, a plastic syringe connected to the top filling port by flexible tubing. Slowly draw the sample up until full. When full, push the syringe plunger down to just below the filling port. Design the number and volume of injections, and initiate titration 16. Using the known concentrations of cell and syringe reactants together with the known effective reaction volume of the calorimeter being used (refer to the manufacturer’s documentation to find out the effective reaction volume of the cell), calculate the volume of injections that will be needed to reach the equivalence point (equimolar concentrations of cell and syringe reactants in the effective cell volume, taking into consideration the number of binding sites each has) of the titration after five or six injections. 17. Program the titration schedule into the software. Input a 3- or 4-min interval between each injection to allow each point along the titration sufficient time to reach equilibrium. Carry the titration out to a molar ratio of at least 2 or 3. Generally, it is desirable to have at least 5 or 6 data points before the equivalence point of the titration is reached, in order to better resolve the binding parameters from the data, especially the binding enthalpy change. More data points are preferable if reactants are plentiful and the concentration of active protein is not known, or if the number of binding sites is not known. It is best to keep the volumes of all the injections the same, since the background mixing heats will be different for different injection volumes. Although this can be corrected for, it requires an extra control titration to be done. Allow for enough injections to be made to carry the titration out to a molar ratio of at least 2 or 3. For tight binding conditions (e.g., Fig. 20.4.2, C >30) a final ratio of 2 should be enough to enable all the fitting parameters to be resolved. For weaker binding conditions (C < 30) the titration may need to be carried out to higher molar ratios in order to better define the curvature and asymptotic mixing heats.
18. Initiate the titration according to the manufacturer’s instructions. An example ITC data set is shown in Figure 20.4.1. ITC instruments are computer controlled. Once the titration schedule has been input, the titration can be initiated. If a binding interaction is not detected, see Troubleshooting, below.
Empty and clean syringe and cell 19. After finishing the ITC titrations, clean and store the calorimeter cell as recommended by the instrument manufacturer. If multiple titrations are being undertaken on the same day, it is usually not necessary to clean the cell between experiments. A simple rinsing of the cell with 2 to 3 volumes of buffer should be adequate.
Carry out a blank titration to measure background mixing heats 20. Repeat precisely the same titration schedule as above, keeping the syringe reactant in the syringe, but using buffer only in the calorimeter cell (no cell reactant). Even in the absence of a binding event between syringe and cell reactants, there will always be measurable “background” heats at every injection. These heats may be small or large, and constant or variable depending on conditions of the titration. Hence, in order to determine the absolute amount of heat produced for a binding event at each titration step, the background heats must be subtracted.
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There are a few cases where this is important. (1) This is critically important under weak binding conditions (e.g., where C < 5). In such cases the asymptotic background heat of mixing is not well defined from the primary titration, since the reaction may not have gone to completion. (2) It is also important under conditions where the syringe reactant changes aggregation state when diluted into the cell. Since the extent of dissociation here depends on its final concentration in the cell, these dissociation heats will vary with each injection. (3) Finally, it is important in cases where the syringe and cell reactant solutions not adequately matched chemically. For such cases, the mixing heats will also vary with each injection as the differences in chemical mismatching will become smaller with each injection.
A 0
5
10
Time (min) 15
20
25
0.2
µcal/sec
0.0
–0.2
–0.4
B
kcal/mol D1D2 injected
0 –4 –8 –12 –16 –20 –24 0
1
2
3
4
Molar ratio (D1D2/mAb)
Titration Microcalorimetry
Figure 20.4.3 Example of ITC data where the affinity is too tight to allow resolution of Kdobs. Note that only one data point is present in the transition region. The data is for an MAb-antigen (CD4, D1D2 domains) interaction with a known Kd°′ = 0.3 nM under these conditions, and the C value (see Fig. 20.4.2) is 3000. (A) Raw data shown as µcal/sec versus time. (B) Integration of the raw data gives the amount of heat produced at each injection normalized to the number of moles D1D2 injected. Reproduced from Doyle and Hensley (1998) with permission from Academic Press.
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A blank titration is not always needed. Very often, with tight binding conditions (C > 10), the titration can be carried out well beyond the equivalence point until there is no cell reactant available for binding. From that point onward, the injection heats are equivalent to blank injection heats. In the end, the need for doing a blank injection experiment depends on the specifics of an experiment and instrument as weighed against the accuracy needed from the analysis.
Analyze data by least-squares regression 21. Analyze the binding data using the nonlinear least-squares regression software provided by the instrument manufacturer. If necessary, correct the data first by subtracting the blank titration data from the binding data. Software for analyzing ITC data is generally provided by the instrument manufacturer.
22a. For single-site binding cases: Assuming there is sufficient binding heat signal and concentrations of reactants are favorable with respect to the C parameter (Fig. 20.4.2), perform curve fitting for three parameters: the molar ratio, binding equilibrium constant (either as Kdobs or its reciprocal), and ∆Hobs. It may be necessary to input initial guesses for some of these parameters to aid the curve-fitting process. If the binding constant for the interaction is too tight to be reliably resolved during data analysis (see Fig. 20.4.3 for an example), see Troubleshooting, below, for ways to weaken the affinity experimentally. Best-fit binding constants that have C values greater than ∼1000 should be regarded as at the limit of the instrumental capabilities and possibly incorrect.
22b. To validate tight binding constants that are at the instrumental limits: Test for reproducibility, change experimental conditions (temperature, pH, salt) to make the affinity weak enough that it can reliably be measured, or use an independent method (e.g., surface plasmon resonance; UNIT 19.4) to validate the affinity. Complex data analysis models are difficult to solve by ITC (or most any other method for that matter) and may require very high-precision data measured over a range of conditions. The underlying challenge for multiple site cases is the need to resolve (or assume) Kdobs, and ∆Hobs for every stoichiometric species. Cases with more than two types of binding sites can be solved in favorable instances with a sequential binding model (e.g., MicroCal, ITC Data Analysis in Origin Tutorial Guide, version 5.0). An example where a four-site binding case has been solved with ITC data is given in Bruzzese and Connelly (1997). General treatment of ligand binding thermodynamic models for proteins is given by Wyman and Gill (1990).
DETERMINING BIOCHEMICAL BINDING THERMODYNAMICS: ∆G°′, ∆H°′ AND ∆Cp°′ BY ITC Protein-ligand interactions are highly sensitive to solvent components such as pH, ionic strength, temperature, specific ions, and water activity. Consequently, the binding thermodynamics for protein-ligand interactions also depend on all these variables, and definition of a pure chemical reaction becomes very complicated. In order to recognize this fact, the authors refer here to the molecular binding thermodynamics of a protein-ligand interaction occurring under specified solution conditions as the biochemical binding thermodynamics for those conditions. See Alberty (1994) for more on the definition of biochemical thermodynamics, and Wiesinger and Hinz (1986) who used the term apparent thermodynamics to account for the same complexities inherent to protein-ligand interactions. The prime superscript used with the thermodynamic parameters indicates that they refer to a biochemical reaction. The superscript symbol (°) is a more general convention in the thermodynamics field and indicates that the standard-state concentration units are in molarity. There is still quite a lot of variability in the nomenclature used to report ligand
BASIC PROTOCOL 2
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binding biothermodynamic data. Nevertheless, the precise physical meaning of biothermodynamic parameters can be understood if they are defined explicitly, including the solution conditions under which they were measured. Determination of biochemical binding thermodynamics requires considerably more effort and care than determination of observed thermodynamics using Basic Protocol 1. First, the experimentally observed binding enthalpy change from a single ITC experiment usually comprises a mixture of the biochemical binding enthalpy change of interest plus artifactual heats of ionization of the pH buffer. This is the case whenever a binding reaction is coupled to uptake or release of protons, and is a common occurrence for protein interactions. Secondly, determination of the biochemical binding thermodynamics for a specified equilibrium reaction implies that no other side reactions are present, such as coupled thermal unfolding of proteins or coupled changes in aggregation state of the reactants. These types of side reactions for proteins are also common and must be considered on a case-by-case basis. This protocol involves correcting the observed binding enthalpy change for buffer ionization heats and controlling for possible side reactions that might corrupt the observed binding thermodynamics. The goal is to measure the biochemical Gibbs free energy, enthalpy, and heat capacity changes that accompany a specific binding interaction. From these parameters, one may calculate the entropy change. The resulting thermodynamic profile of the binding interaction is the first step in utilizing binding thermodynamics data to characterize molecular aspects of the binding mechanism. The results should also allow the temperature dependence of the binding free energy, enthalpy, and entropy changes to be calculated over the temperature range of the study. For reagents and equipment, see Basic Protocol 1. Determine the binding Gibbs free energy change 1. Calculate the binding Gibbs free energy as in steps 2 and 3, below. The binding Gibbs free energy change for a bimolecular biochemical equilibrium reaction can be calculated directly from its biochemical equilibrium dissociation constant, Kd°′ as: ∆G°′= RT ln Kd°′, where R is the gas constant (1.987 calK−1mol−1) and T is temperature in degrees K. In determining Kd°′ one needs to know whether or not the binding interaction is coupled to side reactions such as ligand-induced changes in aggregation state or thermal unfolding. It is also necessary to know the aggregation state of the syringe reactant in order to interpret the data. The following example pertains to a simple ligand binding protein case where there is only a single binding site per protein. Protein molecules having multiple ligand binding sites are significantly more complex to solve by ITC alone and go beyond the realm of the present unit.
2. Measure the self-association status of the protein(s) in their bound and unbound forms at concentrations where the Kdobs will be measured by ITC (see Support Protocol 1). For many proteins, changes in aggregation state upon ligand binding may not occur, or may not occur sufficiently to alter the measured binding parameters. However, one should be suspicious of certain classes of proteins such as those that are multimeric and allosterically regulated. If changes in protein aggregation state are found to occur between bound and unbound forms, a more complex model may be needed for analyzing the data (Doyle et al., 1986). This would represent a very challenging problem to solve with ITC alone.
3. Measure the thermal stability of the protein(s) to define the maximum temperature range under which the binding thermodynamics can measured (see Support Protocol 2). Titration Microcalorimetry
4. Measure Kdobs at a temperature where the protein(s) are known to be in their native folded structures (see Basic Protocol 1).
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5. If step 1 indicates no changes in aggregation state upon binding ligand, then Kdobs is equated with with Kd°′ and the calculation ∆G°′ = RTlnKd°′ is made. Determine ∆H°′ at constant temperature by correcting for buffer ionization heats Many, or perhaps most, protein-ligand reactions proceed with a concomitant net uptake or release of protons. These changes in protonation of the protein or ligand are due to changes in chemical environment of proton ionizable groups, such as histidines at neutral pH, for example. When protons are taken up or released, the buffer releases or takes up protons to maintain constant pH. Since heats of buffer ionization are in the kilocalorie range, they frequently make a substantial artifactual contribution to the observed heat signal. Fortunately, the buffer ionization heats can be corrected for as described below. 6. Measure ∆Hobs in three or more buffers having widely differing proton ionization heats (see Basic Protocol 1). Ideally, the ionization enthalpy changes of the buffers should vary by at least a few kilocalories. Table 20.4.1 lists ionization enthalpy changes for some commonly used buffers. The observed binding enthalpy changes for each of the titrations must be reasonably well resolved. That is, the C parameter should be greater than ∼5 and blank injection experiments should be done if needed.
7. Plot ∆Hobs versus ionization enthalpy of the pH buffers (Fig. 20.4.4). Fit the data by least-squares regression to determine the slope and the intercept. The biochemical binding enthalpy change, ∆H°′, is given by the intercept. The slope of the plot gives the number of protons thermodynamically coupled to binding at the specific pH and other solution conditions of the titrations. The data should be linear. Significant deviation from linearity could mean that one or more of the buffers has a specific interaction with the protein being studied that alters the protein’s function. For example, phosphate is known to bind to some proteins.
Determine the binding heat capacity change, ∆Cp°′, using buffer-corrected binding enthalpy changes measured at various temperatures The binding heat capacity change is defined as the temperature dependence of the binding enthalpy change. It is a parameter that plays a key role in interpreting the thermodynamics of an interaction, and for correcting the thermodynamics for temperature differences. 8. Measure buffer-corrected binding enthalpy changes, ∆H°′, according to steps 5 and 6 of this protocol, at several temperatures covering a range of at least 10°C. The exact temperature range used will depend on the thermal stabilities of the reactants. In general, the larger the range the better resolved and more informative the data. Aim for a 20° or 30°C range if possible.
9. Plot ∆H°′ versus temperature (Fig. 20.4.5). The slope of the plot is equal to the binding heat capacity change, ∆Cp°′. For typical protein-ligand interactions the plot should be linear (e.g., see Fig. 20.4.5). Deviation from linearity could mean that the binding reaction is coupled to temperaturedependent processes that make a significant contribution to the binding enthalpy change. Possible sources include thermal unfolding of the protein (Thomson et al., 1994), temperature-induced changes in native protein conformation, coupled protonation reactions that vary substantially with temperature (Koslov and Lohman, 1999), and changes in aggregation state (Doyle et al., 1986). Any such temperature-dependent changes in reaction mechanism may influence interpretation of the measured ∆Cp°′. When making measurements over a range of temperature, the molar ratio values should be constant. Significant
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–16 citrate
∆H obs (kcal/mol CD4)
–18
–20 HEPES –22
–24
ACES
–2
0
2
6
4
8
Buffer ionization ∆H i on (kcal/mol proton)
Figure 20.4.4 Observed binding enthalpy change from ITC depends on the buffer proton-ionization enthalpy whenever binding is coupled to uptake or release of protons. Data are for a monoclonal antibody (MAb) protein–antigen (CD4) interaction (Doyle et al., 1995). The slope of the plot yields the number of protons coupled to the antibody-antigen binding reaction, and the intercept yields the biochemical binding enthalpy change (∆H°′) corrected for buffer ionization artifact heats.
–14 –16 Binding ∆H ˚′ (kcal/mol)
–18 –20 –22 –24 –26 –28 –30 –32 20
25
30
35
40
45
50
Temperature (˚ C)
Titration Microcalorimetry
Figure 20.4.5 Temperature dependence of the biochemical binding enthalpy change for a MAb protein-antigen interaction (binding of CD4 to anti-CD4 MAb CE9.1). The slope of the line yields a binding heat capacity change equal to −0.69 kcal/mol/°C. Different protein-protein or protein-ligand interactions will have different slope and intercept values, but all protein-ligand binding enthalpy changes should exhibit a temperature dependence. Reproduced from Doyle and Hensley (1998) with permission from Academic Press.
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changes in molar ratio would indicate thermal degradation of one of the reactants or a fundamental change in binding mechanism. When reporting biothermodynamic parameters, one must always define explicitly whether they are observed or biochemical. In either case, the solution conditions that pertain to the measured parameters must also be reported.
MEASUREMENT OF THERMAL STABILITY To rigorously interpret biothermodynamic data, it is necessary to understand some physical properties of the reactants. It is important to characterize the thermal stabilities of the protein reactants in order to define the temperature range over which they are thermally stable. Otherwise, any thermally induced unfolding reactions could greatly alter the bioenergetics of the reaction and lead to misinterpretation of the data.
SUPPORT PROTOCOL 1
Using solution conditions relevant to the ITC experiments, measure the thermal stability of the protein(s) with differential scanning calorimetry (DSC; refer to UNIT 7.9). Scan a temperature range covering the range to be used for ITC studies, plus at least an additional 10° to 20°C on each end to define the baseline. DSC (Fig. 20.4.6) is the method of choice since it can cover the widest temperature range, and provides a more direct evaluation of protein unfolding processes. However, it may require a milligram or more of protein, depending on the nature of its unfolding properties.
350
–10
250
Ellipticity
C p (kcal/mol/degree)
300
200 150
–20 10 20 30 40 50 60 70 80 Temperature
100 50 0 10
20
30
40
50 60 Temperature
70
80
90
100
Figure 20.4.6 DSC data for the temperature dependence of the heat capacity of MAb CE9.1. Measurements were done at a scan rate 1°C/minute. The temperature-dependence of the MAb heat capacity Cp does not reveal any thermally induced transitions over the experimental temperature range of the present ITC studies, suggesting the MAb does not undergo temperature-induced structure changes. Unfolding begins to occur at 60°C. Multiple transitions indicate multiple domains and are typical for MAb unfolding. Inset is thermal unfolding of D1D2 domains of CD4 as measured by circular dichroism. Measurements were done at a scan rate of 1 degree/minute. Plot shows ellipticity in millidegrees versus temperature. D1D2 is stable up to ∼50°C. The maximum temperature used in ITC studies (47°C) is indicated by the vertical dotted line. Reproduced from Doyle and Hensley (1998) with permission from Academic Press.
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If protein reagents are expensive or access to a DSC instrument is limited, measure the thermal stability of the protein(s) using circular dichroism (CD) as a spectral monitor of protein secondary structure (refer to UNIT 7.6 and Eftink, 1995). Temperature-regulated cuvettes will be necessary to do this: either as Peltier-controlled or as water-jacketed cuvettes with a circulating temperature-controlled water bath. CD spectroscopy (Fig. 20.4.6) requires <0.1 mg of protein, but is a less direct monitor of protein unfolding, due partly to possible interference from optical artifacts. Fluorescence is another possible spectral method for evaluating protein thermal stability, but the analysis is more complicated due to temperature-dependent baseline changes in fluorescence as described by Eftink (1995). SUPPORT PROTOCOL 2
DETERMINATION OF PROTEIN ASSEMBLY STATES Evaluation of the self-association status of reactants and products is also important for properly interpreting thermodynamic data. Measure the aggregation state of the protein(s) using the conditions in the ITC studies. Use protein concentrations that correspond to those used in the ITC experiments. Methods for quantitative analysis of protein aggregation states include analytical ultracentrifugation (UNIT 20.3), size-exclusion chromatography with on-line light scattering, and largezone analytical gel filtration (UNIT 20.5). COMMENTARY Background Information A major goal in the post-genome era will be quantitative molecular characterization of protein-ligand interactions (Doyle and Hensley, 1997). Isothermal titration calorimetry will undoubtedly be a key tool for high-resolution characterization of protein interactions by virtue of its high thermodynamic information content, high accuracy, and general applicability. Titration microcalorimetry detects binding directly from the heat absorbed or released during a binding reaction (i.e., the binding enthalpy change). Because all molecular interactions are accompanied by a heat change, ITC is in principle a universal method for characterizing chemical and biochemical binding interactions. Although titration calorimetry has been used to characterize chemical reactions for decades, it was not until the 1980s (McKinnon et al., 1984; Wiseman et al., 1989) that the sensitivity was sufficient for characterization of biomolecules that can be produced only in limited quantities. ITC has been applied to a wide range of interactions (see Doyle, 1997 and references therein) including antibody-antigen, peptide-protein, lipid-protein, protein-receptor (soluble and membrane-bound), carbohydrate-protein, nucleic acid–protein, protein-protein, and small molecule/drug–protein interactions.
Titration Microcalorimetry
Advantages and disadvantages of the procedure ITC is probably the most rigorous method for accurate characterization of binding affinity and functional chemistry of biological macromolecules. There are several advantages that ITC has over other methods, listed as follows. 1. ITC has the ability to characterize interactions of unmodified biomolecules in solution, which improves the likelihood of determining accurate binding parameters. Some binding methods—e.g., enzyme-linked immunoassay (ELISA), magnetic beads, or surface plasmon resonance (SPR)—require covalent attachment of one of the reactants to a surface, a bead, or an optical label. Such modifications run the risk of altering, or abolishing, the functional binding properties of a biomolecule. 2. Binding equilibrium reactions are characterized by ITC under conditions of equilibrium, and the attainment of equilibrium can be monitored in real time during the course of the titration. This contrasts with some methods (e.g., ELISA) that require physical separation of protein-ligand and unbound ligand species. The separation step will often perturb the position of equilibrium and hence give an inaccurate measure of the affinity. 3. Equilibrium constants are measured directly, in contrast to kinetic methods (e.g., SPR),
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which predict equilibrium constants from the ratio of association and dissociation rate constants. Determination of the rate constants in turn relies on computational deconvolution from association and dissociation processes that are often complex or inadequately understood. 4. The generality of a binding heat detection method makes it applicable, in theory, to any binding interaction. 5. ITC has the potential for producing highresolution thermodynamic information about the molecular mechanism of an interaction. However, obtaining reliable thermodynamic information that has specific molecular physical meaning about an interaction requires some additional control experiments to be conducted, both by ITC and other methods. Drawbacks of ITC include the following. 1. The method has a relatively large protein sample requirement of at least 5 to 10 nmol. For proteins that are difficult or expensive to produce, this can be cost-prohibitive. 2. The throughput of about a few titrations per day is low compared to less quantitative methods such as 96-well fluorescence-based techniques. However, higher-throughput instrumentation with autosampler capability should soon be available. 3. Titrations are carried out at reactant concentrations of micromolar and above. The weaker the affinity of the interaction, the higher the concentrations of reactants that must be used. This becomes a limiting factor for lowsolubility reactants or for expensive reactants, and is especially a problem for measuring affinities weaker than micromolar. 4. The range of binding constants that has been measured by ITC is approximately millimolar to nanomolar. The weaker affinities could only be measured for very soluble and inexpensive reactants. Affinities tighter than about nanomolar cannot generally be measured by current ITC instrumentation; however, binding enthalpy changes and molar ratios can be readily determined for very tight binding interactions. Interpretation of binding thermodynamics An attractive aspect of ITC is the information-rich data it yields about binding thermodynamics. Protein-ligand interactions involve changes in hydrogen bonding, van der Waals interactions, ionic bonds, hydration of both protein and ligand, protonation, and ion binding. Moreover, many proteins, especially regulatory proteins, undergo substantial changes in conformation upon ligand binding. All of these mo-
lecular events have specific thermodynamic consequences that sum up to net, observed changes in Gibbs free energy, enthalpy, entropy, and heat capacity. Current research is aimed at deconvoluting binding thermodynamics into the contributions from each of these constitutive processes. However, it is clear that the problem is not a simple one. The first step in correlating binding thermodynamics with molecular mechanism is to obtain high-quality and well-defined thermodynamic parameters. The objective of Basic Protocol 2 in this unit is to guide the experimentalist through the various control experiments needed to obtain this information. Using this information to understand the molecular details of a protein-ligand interaction is a separate and very challenging task. Phenomenology. That said, observed binding thermodynamics can be used straightforwardly as a stringent molecular readout of the overall functional chemistry of a binding interaction, despite the large number of the underlying molecular processes. For example, one may compare the complete binding thermodynamic profiles for glycosylated and deglycosylated forms of a protein. If glycosylation affects the functional chemistry of the protein-ligand interaction, it is more likely to show up in a complete thermodynamic profile than a measure of affinity alone. Cases have been reported where mutationally or chemically altered proteins have exhibited changes in binding ∆H and ∆S terms that were “hidden” in affinity measurements alone, due to entropy-enthalpy compensation. Thus, it was only through knowledge of the thermodynamic parameters that the modifications playing a structural role in binding were identified (Murphy et al., 1995; Reedstrom and Royer, 1995; Evans et al., 1996; Pearce et al., 1996). Molecular mechanism. As mentioned above, the binding thermodynamics for proteins are comprised of a large number of molecular processes, and partitioning the thermodynamics into each of these processes is a major goal of current biothermodynamics research. General ideas concerning interpretation of biothermodynamic data for protein-ligand interactions can be found in recent articles. Calorimetric studies have succeeded in using biothermodynamic data to characterize various aspects of protein-ligand binding mechanisms such as the roles of coupled ion binding (Guinto and Di Cera, 1996; Lohman et al., 1996; Koslov and Lohman, 1998), protonation (Baker and Murphy, 1996, 1997; Koslov and Lohman, 1999), hydration (Bhat et al.,
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1994; Connelly et al., 1994, Chung et al., 1998), the amount of surface area buried in proteinprotein interactions (Murphy and Freire, 1992; Gomez and Freire, 1995), and the extent of conformational change coupled to binding (Spolar and Record, 1994). A detailed correlation between biothermodynamics and molecular binding mechanism is challenging and should take into account all of these factors. A rigorous study would be multidimensional in nature, and would be likely to include measurements of the biochemical binding thermodynamics (corrected for buffer ionization heats) as a function of temperature, salt concentration, and osmotic pressure. Salt effects can play a dominant role in the thermodynamics of protein-nucleic acid interactions (Lohman et al., 1996; Koslov and Lohman, 1998). Given the challenges of interpreting biochemical thermodynamics with the myriad of processes coupled to protein-ligand interactions, it is recommended that the reader gain a more comprehensive understanding of the biothermodynamics field than is presented in this unit.
Critical Parameters
Titration Microcalorimetry
Determination of observed binding thermodynamics Successful implementation of titration microcalorimetry relies on several factors. First, the solutions of the reactants in the syringe and cell must be chemically matched so that the background heats of mixing are small relative to the heats of reactant binding. Balancing the pH values and total concentrations of nonaqueous additive solvents such as DMSO or glycerol are especially important. Second, the reactants must be studied at concentrations that are relevant to the affinity of the interaction. In order to detect binding, the concentration of reactant in the cell should be at least four-fold higher than the equilibrium dissociation constant of the interaction (Fig. 20.4.2). For weak binding interactions where binding is not detected (i.e., weak relative to the reactant concentration in the cell) the titration may need to be repeated at much higher concentrations of reactants in order to drive forward the binding process. On the other hand, for tight binding interactions (tight relative to the concentration of reactant in the cell), binding may be easily detected, but the affinity constant may not be resolvable due to “stoichiometric binding conditions” (see Troubleshooting). In these cases, the concentrations of reactants should be
decreased until the Kd can be resolved. In general, the minimum concentrations that can be studied are ∼1 µM in the cell and 20 µM in the syringe. With lower concentrations it is unlikely that there will be enough heat to characterize the interaction. Hence, it may not be possible in general to directly measure affinity constants by ITC for very tight binding interactions (e.g., Kd <1 nmol) due to the instrumental sensitivity limits. Finally, the accuracy of the binding physical parameters determined by ITC depends heavily on (1) the accuracy with which the reactant concentrations were determined, and (2) the fractional activity of the reactants. In particular, the accuracy of the affinity constant and binding enthalpy change are directly proportional to the accuracy of the syringe reactant. Inaccuracies of two-fold in the concentration of syringe reactant will translate into a two-fold inaccuracy in the measured affinity constant. For most studies this is of little concern. However, a two-fold inaccuracy in the binding enthalpy change will cause serious problems in interpretation of the binding thermodynamics. The accuracy of the cell reactant concentration and its fractional activity affect neither the affinity constant nor the binding enthalpy change. But they do affect in direct proportion the accuracy of the binding molar ratio (and so does the accuracy of the syringe reactant concentration and fractional activity). Determination of biochemical binding thermodynamics Progression from determination of observed binding parameters (see Basic Protocol 1) to determination of rigorous, molecular-level, biochemical thermodynamics (see Basic Protocol 2) requires first that the binding model be clearly defined, particularly with regard to the thermal stability behavior of the reactants and possible changes in aggregation state upon binding. Typical analysis of ITC data involves fitting the data to simple single-site or two-site binding models, which assumes that the proteins are not undergoing ligand-induced changes in self-association or extent of thermal unfolding. If, for example, ligand binding is coupled to changes in self-association, the measured apparent thermodynamic parameters will include contributions from self-association. Analysis of thermal stability of biological macromolecules is highly relevant to ITC studies because one of the most information-rich parameters, ∆Cp, is determined from measurements made over a range of temperature. One
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must take care to conduct ITC measurements only under conditions where the protein(s) are in their native folded state(s). The binding ∆Hobs measured from a single ITC experiment usually includes heats due to buffer ionization. This is true whenever binding is coupled to protonation, as is often the case for biological macromolecules. Thus, determination of the biochemical binding ∆H°′ (Table 20.4.1) for the molecular interaction of interest requires measurements to be performed separately at constant pH but in buffers with different ionization enthalpies. An additional benefit of this analysis is that it also yields the number of protons that are coupled to binding. Characterization of the coupled protonation reactions is useful for understanding the molecular details of the binding reaction (Garcia-Fuentes et al., 1995; Baker and Murphy, 1996, 1997).
near zero at a given temperature, it will be nonzero at a distal temperature. Note that some proteins are labile at high temperatures, and hence the thermal stability of the reactants may need to be evaluated before increasing temperature (see Support Protocol 1). 3. Amplify binding heat signal with a different pH buffer Carry out the titration in a pH buffer that has a proton ionization enthalpy change that is distinct from the one used in the initial titration. The rationale here is that many protein-ligand interactions are accompanied by either release or uptake of protons. In the presence of a buffer, these protons are in turn absorbed or released by the buffer, with an accompanying ionization heat (Doyle et al., 1995). Consequently, the observed binding enthalpy from ITC may depend on the pH buffer used, as portrayed in Figure 20.4.4.
Troubleshooting
What to do if binding is too tight to measure Kd The underlying difficulty in measuring very tight binding constants lies in the challenge of being able to measure the concentration of free (unbound) ligand during a titration curve. The equilibrium dissociation constant is defined as the ratio Kd = [P][L]/[PL], where [PL] is the concentration of protein-ligand complex, [P] is unbound protein, and [L] is the unbound ligand concentration ([L] = [Ltotal added] − [Lbound]). For tight binding in ITC experiments (or any other method) all of the added ligand becomes bound to protein, so that [L] is immeasurably small and the Kd is unresolvable. This condition is called “stoichiometric binding” and refers to the fact that within the margins of error all the ligand added to the cell becomes bound until the cell reactant binding sites are fully titrated. An example of ITC data where the affinity is too tight to resolve the Kd is given in Figure 20.4.5. Some of the solutions to these problems are discussed below. 1. Lower the reactant concentrations. If binding is very tight relative to the concentration of reactant in the cell (Fig. 20.4.5), the affinity constant will not be resolvable from the data. The first step in addressing this problem is to lower the concentration of the reactant, particularly the one in the cell. Unfortunately, the lower limit of reactant in the cell that can be adequately studied is typically near 1 µM. With any lower concentrations, the total heat signal becomes too small to reliably characterize the reaction. The absolute lower limit varies by several fold and depends on the magnitude of the binding enthalpy change.
A troubleshooting guide for ITC experiments can be found in Table 20.4.3. What to do if binding is not detected If binding is not detected, either: (a) the affinity is too weak relative to the concentrations being studied, or (b) the binding enthalpy change is small (or a combination of both). There are several optimizations to try at this point. Increasing reactant concentrations is the most likely strategy to reveal binding. If that does not work, or if materials are insoluble or in limited supply, there are a couple of other strategies (changing temperature or buffer) that may amplify binding heats. These later strategies only apply for cases where reactant concentration in the cell is at least several-fold higher than the binding Kd. These strategies are discussed below in order of mention. 1. Increase reactant concentrations 10-fold or more, if possible. By increasing concentrations, one will increase the extent of binding for weak binding cases and produce more heat signal. 2. Amplify the binding heat signal with a different temperature. Change the experimental temperature by at least 5° or 10°C in either direction. It is theoretically possible that a binding interaction is occurring, but that the binding heat is coincidentally close to zero. All proteinligand interactions are accompanied by a temperature dependence to their binding enthalpy change (See Fig. 20.4.5). The parameter that describes this temperature-dependence is defined as the binding heat capacity change (Doyle and Hensley, 1998). Thus, if the binding heat is
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Table 20.4.3
Troubleshooting Guide for Isothermal Titration Calorimetry
Problem
Possible Cause
Solution
Noisy baseline when stirring starts
Misaligned or bent syringe
Verify that syringe/stirring apparatus is centered and well seated. Inspect syringe needle for bending. Replace syringe if bent.
Sample contains air bubbles
Remove sample from cell and refill cell with sample while avoiding bubble introduction
Mixing heats do not titrate away
Buffer mismatch between syringe and cell solutions
Test pH of both solutions and match if needed
Binding not observed
Reactant concentrations too low
Increase reactant concentration in cell to at least four-fold higher than the expected Kd. Adjust syringe reactant concentration as needed.
Binding heat is small
Amplify binding heat by increasing or decreasing temperature by several degrees
Binding heat is small
Change buffer to one with a different ionization heat to amplify heat of coupled protonation (if present)
Equivalence point not yet reached
Carry titration to higher molar ratio
Observed binding constant is close to instrumental limit
Affinity is too tight to measure directly
Increase temperature (exothermic reactions) to weaken affinity. Change other solution conditions (salt, pH) to possibly weaken affinity.
Unexpected value for molar ratio/stoichiometry
One or both reactant concentrations incorrect
Determine concentrations more quantitatively
One or both reactants partially inactive
Repurify reactants Evaluate whether protein(s) are folded under reaction conditions
Binding heat has a nonlinear dependence with temperature
Reaction more complex than a simple A + B = AB binding equilibrium
Evaluate thermal stability and ligand-induced aggregation changes
Observed binding constant does not vary with temperature according to van’t Hoff model
Reaction more complex than a simple A + B = AB binding equilibrium
Evaluate thermal stability and ligand-induced aggregation changes
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4
K d˚′ nM
3
2
1
0 20
25
30
35
40
45
50
Temperature (˚C)
Figure 20.4.7 Temperature-dependence of the binding equilibrium dissociation constant for a MAb-antigen (CD4) interaction. Solid curve is calculated from the integrated van’t Hoff equation (Equation 20.4.1) using the affinity constants measured at 45° and 47°C in combination with measured values of the binding ∆H°′ and ∆Cp°′. Squares represent affinities measured directly by titration calorimetry at higher temperatures. Circles represent affinity measured by an independent method by Doyle et al. (1995). Triangles represent affinity measured by surface plasmon resonance. Reproduced from Doyle and Hensley (1998) with permission from Academic Press.
2. Use a higher temperature. Typically the affinity of an interaction will become weaker as temperature increases. This is true when the binding enthalpy change is exothermic, which is almost always the case for protein-ligand interactions at 25°C or higher. For endothermic reactions, the affinity can be weakened by lowering temperature. The extent to which the affinity becomes weaker with temperature depends on the magnitude of the binding enthalpy change. Generally speaking, by increasing the temperature by 10°C, one might expect to lower the affinity by several fold. Thus, the new value of the affinity might be weak enough to be measured directly by ITC. Whether or not the reactants are stable at the new temperature should be evaluated by Support protocol 1 or 2). 3. Correcting the affinity constant to a standard temperature. By changing the temperature of the experiment, one measures affinity under new conditions, and it may be desirable to correct the measured affinity to some reference temperature. This can be done by using the van’t Hoff model for the temperature dependence of an equilibrium constant. The model assumes a simple equilibrium reaction, and seems to at least approximate well the temperature depend-
ence of protein-ligand reactions. The appropriate form of the integrated van’t Hoff equation for protein-ligand interactions is (Brandts and Lin, 1990): K d ( T ) = K d ( T0 ) exp
LM ∆H(T ) FG 1 − 1 IJ − N R HT T K ∆C F T T IO − − 1J P ln G R H T T KQ 0
0
p
0
0
Equation 20.4.1
Here, Kd(T) is the equilibrium dissociation constant calculated at a temperature T, and Kd(T0) is the equilibrium dissociation constant measured at temperature T0. ∆H(T0) is the biochemical binding enthalpy change (corrected for buffer ionization heats) measured at T0. The ∆Cp term is the binding heat capacity change (also corrected for buffer ionization heats) and provides a first-order correction for the temperature dependence of ∆H and a second-order correction to the temperature dependence of the equilibrium constant. An example of using the van’t Hoff model to calculate the temperature-
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dependence of the affinity constant for a protein-ligand interaction is given in Figure 20.4.7. It is important to note that there are some caveats in using the van’t Hoff model. Although the model has enjoyed wide acceptance in the biothermodynamics field and appears to describe the temperature dependence of substantial number of protein-folding and protein-ligand reactions, close examination with model compound interactions suggest it may be an oversimplification for interactions in aqueous solution (Liu and Sturtevant, 1995). Thus, the larger the temperature extrapolation used with the van’t Hoff equation, the greater the potential for both random and systematic error. Based on empirical results in the literature, the van’t Hoff model would be expected to be valid over at least a 10° to 20°C temperature range for protein-ligand interactions. But this could vary between different protein-ligand interactions. A key assumption of the model is that a simple binding equilibrium exists over the temperature range (e.g., no thermal unfolding of proteins). Troubleshooting of the instrumentation The instrumentation can be tested by making injections of water into water. Since there is no chemical mismatch or heat of dilution for water injected into water, the data should appear as a series of small injection heats that are constant for each injection. This control experiment is useful when it is suspected that the injection syringe may be bent, or some other instrumental hardware problem may exist.
Anticipated Results Binding heats should be visually distinct from background mixing heats. A transition curve should be observed which has an equivalence point near a molar ratio of one ligand per binding site on the protein.
Time Considerations Sample preparation time may vary depending on the time required for dialysis (which can be done in 1 hr with current methods) and for establishing solubility limits in the case of poorly soluble reactants such as small organic molecules. Setting up and running the ITC experiment takes ∼2 hr.
Literature Cited Alberty, R.A. 1994. Biochemical thermodynamics. Biochim. Biophys. Acta 1207:1-11.
Titration Microcalorimetry
Baker, B.M. and Murphy K.P. 1996. Evaluation of linked protonation effects in protein binding re-
actions using isothermal titration calorimetry. Biophys. J. 71:2049-2055. Baker, B.M. and Murphy, K.P. 1997. Dissecting the energetics of protein-protein interactions: The binding of ovomucoid third domain to elastase. J. Mol. Biol. 268:557-569. Bhat, T.N., Bentley, G.A., Boulot, G., Greene, M.I., Tello, D., Dall’Acqua, W., Souchon, H., Schwarz, F.P., Mariuzza, R.A., and Poljak, R.J. 1994. Bound water molecules and conformational stabilization help mediate an antigen-antibody association. Proc. Natl. Acad. Sci. U.S.A. 91:1089-93. Brandts, J.F. and Lin, L.-N. 1990. Study of strong to ultratight protein interactions using differential scanning calorimetry. Biochemistry 29:69276940. Bruzzese, F.J. and Connelly, P.R. 1997. Allosteric properties of inosine monophosphate dehydrogenase revealed through the thermodynamics of binding inosine 5′-monophosphate and mycophenolic acid: Temperature dependent heat capacity of binding as a signature of ligand-coupled conformational equilibria. Biochemistry 36:10428-10438. Christensen, J.J., Hansen, L.D., and Izatt, R.M (eds.). 1976. Handbook of Proton Ionization Heats and Related Thermodynamic Quantities. John Wiley & Sons, New York. Chung, E., Henriques, D., Renzoni, D., Zvelebil, M., Bradshaw, J.M., Waksman, G., Robinson, C.V., and Ladbury, J.E. 1998. Mass spectrometric and thermodynamic studies reveal the role of water molecules in complexes formed between SH2 domains and tyrosyl phosphopeptides. Structure 6:1141-1151. Connelly, P.R., Aldape, R.A., Bruzzese, F.J., Chambers, S.P., Fitzgibbon, M.J., Fleming, M.A., Itoh, S., Livingston, D.J., Navia, M.A., Thomson, J.A., and Wilson, K.P. 1994. Enthalpy of hydrogen bond formation in a protein-ligand binding reaction. Proc. Natl. Acad. Sci. U.S.A. 91:19641968. Doyle, M.L. 1997. Characterization of binding interactions by isothermal titration calorimetry Curr. Opin. Biotechnol. 8:31-35. Doyle, M.L. and Hensley, P. 1997. Experimental dissection of protein-protein interactions in solution. Adv. Mol. Cell Biol. 22A:279-237. Doyle, M.L. and Hensley, P. 1998. Tight ligand binding affinities determined from thermodynamic linkage to temperature by titration calorimetry. Methods Enzymol. 295:88-99. Doyle, M.L., Gill, S.J., and Cusanovich, M.A. 1986. Ligand-controlled dissociation of Chromatium vinosum cytochrome c′. Biochemistry 25:25092516. Doyle, M.L., Louie, G., Dal Monte, P., and Sokoloski, T. 1995. Tight binding affinities determined from thermodynamic linkage to protons by titration calorimetry. Methods Enzymol. 259:183-194.
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Eftink, M.R. 1995. Use of multiple spectroscopic methods to monitor equilibrium unfolding of proteins. Methods Enzymol. 259:487-512.
Spolar, R.S. and Record, M.T., Jr. 1994. Coupling of local folding to site-specific binding of proteins to DNA. Science 263:777-784.
Evans, L.J.A., Cooper A., and Lakey, J.H. 1996. Direct measurement of the association of a protein with a family of membrane receptors. J. Mol. Biol. 255:559-563.
Stoll, V.S. and Blanchard, J.S. 1990. Buffers: Principle and practice. Methods Enzymol. 182:24-38.
Garcia-Fuentes, L., Reche, P., Lopez-Mayorga, O., Santi, D.V., Gonzalez-Pacanowska, D., and Baron, C. 1995. Thermodynamic analysis of the binding of 5-fluoro-2′-deoxyuridine 5′-monophosphate to thymidylate synthetase over a range of temperatures. Eur. J. Biochem. 232:641-645. Gomez, J. and Freire, E. 1995. Thermodynamic mapping of the inhibitor site of the aspartic protease endothiapepsin. J. Mol. Biol. 252:337-350. Guinto, E.R. and Di Cera, E. 1996. Large heat capacity change in a protein-monovalent cation interaction. Biochemistry 35:8800-8804. Koslov, A.G. and Lohman, T.M. 1998. Calorimetric studies of E. coli SSB protein-single stranded DNA interactions: Effects of monovalent salts on binding enthalpy J. Mol. Biol. 278:999-1014. Koslov, A.G. and Lohman, T.M. 1999. Adenine base unstacking dominates the observed enthalpy and heat capacity changes for the Escherichia coli SSB tetramer binding to single-stranded oligoadenylates. Biochemistry 38:7388-7397. Liu, Y. and Sturtevant, J.M. 1995. Significant discrepancies between van’t Hoff and calorimetric enthalpies. II. Protein Sci. 4:2559-2561. Lohman, T.M., Overman, L.B., Ferrari, M.E., and Kozlov, A.G. 1996. A highly salt-dependent enthalpy change for Escherichia coli SSB proteinnucleic acid binding due to ion-protein interactions. Biochemistry 35:5272-5279. McKinnon, I.R., Fall, L., Parody-Morreale, A., and Gill, S.J. 1984. A twin titration microcalorimeter for the study of biochemical reactions. Anal. Biochem. 139:134-139. Murphy, K.P. and Freire, E. 1992. Thermodynamics of structural stability and cooperative folding behavior in proteins. Adv. Protein Chem. 43:31361. Murphy, K.P., Xie, D., Garcia, K.C., Amzel, L.M., and Freire, E. 1993. Structural energetics of peptide recognition: Angiotensin II/antibody binding. Proteins Struct. Funct. Genet. 15:113-120. Murphy, K.P., Freire, E., and Paterson, Y. 1995. Configurational effects in antibody-antigen interactions studied by microcalorimetry. Proteins Struct. Funct. Genet. 21:83-90. Pearce, K.H., Ultsch, M.H., Kelley, R.F., de Vos, A.M., and Wells, J.A. 1996. Structural and mutational analysis of affinity-inert contact residues at the growth hormone-receptor interface. Biochemistry 35:10300-10307. Reedstrom, R.J. and Royer, C.A. 1995. Evidence for coupling of folding and function in trp repressor: Physical characterization of the superrepressor mutant AV77. J. Mol. Biol. 253:266-276.
Thomson, J., Ratnaparkhi, G.S., Varadarajan, R., Sturtevant, J.M., and Richards, F.M. 1994. Thermodynamic and structural consequences of changing a sulfur atom to a methylene group in the M13Nle mutation in ribonuclease-S. Biochemistry 33:8587-8593. Wiesinger, H. and Hinz, H.J. 1986. Thermodynamic Data for Biochemistry and Biothermodynamics. pp. 211-226. Springer-Verlag, Berlin. Wiseman, T., Williston, S., Brandts, J.F., and Lin, L-N. 1989. Rapid measurement of binding constants and heats of binding using a new titration calorimeter. Anal. Biochem. 179:131-137. Wyman, J. and Gill, S.J. 1990. Binding and Linkage: The Functional Chemistry of Biological Macromolecules. University Science Books, Mill Valley, Calif.
Key References Baker and Murphy, 1996. See above. A comprehensive treatise on the use of ITC to characterize protonation reactions that are coupled to ligand binding. Bhatnagar, R.S. and Gordon, J.I. 1995. Thermodynamic studies of myristoyl-CoA-protein nmyristoyltransferase using isothermal titration calorimetry. Methods Enzymol. 250:467-486. Reviews practical aspects of ITC, including experimental design for tight/weak binding interactions. Bundle, D.R. and Sigurskjold, B.W. 1994. Determination of accurate thermodynamics of binding by titration microcalorimetry. Methods Enzymol. 247:288-304. Reviews practical and theoretical aspects of using ITC to characterize protein-carbohydrate interactions. Chung et al., 1998. See above. A detailed structural-thermodynamic study describing the role of water in achieving specificity, as well as the thermodynamic consequences. Connelly et al., 1994. See above. A detailed structural-thermodynamic study describing the role of water in achieving specificity, as well as the thermodynamic consequences. Cooper, A. and Johnson, C.M. 1994. Isothermal titration microcalorimetry. Methods Mol. Biol. 22:125-150. Reviews the fundamentals of conducting ITC studies. Fisher, H.F. and Singh, N. 1995. Calorimetric methods for interpreting protein-ligand interactions. Methods Enzymol. 259:194-221.
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A comprehensive review of the mechanics and practical issues involved in conducting ITC experiments. Includes discussion on data analysis and deconvolution of linked protons.
Correlated binding thermodynamics to extent of conformational change observed by X-ray and NMR structural analysis.
Gomez and Freire, 1995. See above.
Describes ITC instrumentation and data analysis.
A comprehensive thermodynamic study that dissects the individual contributions of amino acids to a peptide-protein binding interaction. This paper points out the specific properties of protein-ligand interactions (e.g., coupled protonation events and conformational entropies for amino acid sidechains) that must be taken into account when using thermodynamic approaches to structure-based drug design.
Wiseman et al., 1989. See above.
Internet Resources http://www.biophysics.org/biophys/society/bioh ome.htm Biophysical Society home page. Refer to Biophysics On-Line and Chapter on Thermodynamics links at this site. http://www.microcalorimetry.com/
Koslov and Lohman, 1998. See above.
http://www.calorimetry.com
Reports the large effects that specific salts can have on the thermodynamics of protein-nucleic acid interactions. Indicates that an accurate understanding of structure-thermodynamic relationships in binding specificity will require characterization of salt effects.
Web site of MicroCal (manufacturer of high-sensitivity microcalorimeters); includes background and literature references about calorimetry.
Lin, L.-N., Li, J., Brandts, J.F., and Weiss, R.M. 1994. The serine receptor of bacterial chemotaxis exhibits half-site saturation for serine binding. Biochemistry 33:6564-6570. An example where ITC was done with membranebound receptors. Lohman et al., 1996. See above. Reports the large effects that specific salts can have on the thermodynamics of protein-nucleic acid interactions. Indicates that an accurate understanding of structure-thermodynamic relationships in binding specificity will require characterization of salt effects.
http://www.calscorp.com Web site of Calorimetry Sciences Corporation (manufacturer of high-sensitivity microcalorimeters); includes background and literature references about calorimetry. http://www.thermometric.com Web site of Thermometric AB, Sweden (manufacturer of high-sensitivity microcalorimeters); includes background and literature references about calorimetry.
Contributed by Michael L. Doyle SmithKline Beecham Pharmaceuticals King of Prussia, Pennsylvania
Spolar and Record, 1994. See above.
Titration Microcalorimetry
20.4.24 Supplement 18
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Reduced-Scale Large-Zone Analytical Gel-Filtration Chromatography for Measurement of Protein Association Equilibria
UNIT 20.5
Protein association reactions contribute to many biologically significant phenomena and, as evidenced by the other entries in this chapter, measurement of a protein assembly process can be accomplished using a variety of approaches. The advantage of large-zone analytical gel-filtration chromatography (AGFC), originally developed by Ackers and co-workers (Ackers, 1975; Valdes and Ackers, 1979), over many other techniques is that it provides a method for quantitative measurement of a protein assembly reaction that, because of its relatively “low-tech” instrumental requirements, is accessible to virtually all biochemistry laboratories. The large-zone AGFC experiment is performed using a column—packed by the investigator with hydrophilic, compressible, size-exclusion resin, such as Sephadex—and a peristaltic pump capable of providing stable flow rates. Study of a protein assembly reaction by large-zone AGFC requires determination of the weight-average partition coefficient or average size over a protein concentration range that defines the association equilibrium on a calibrated size-exclusion column. The concentration dependence of the protein size is then analyzed using the appropriate model for the assembly reaction to obtain the energetics and stoichiometry of the process. The adaptation of large-zone AGFC for use with modern semicompressible, size-exclusion resin and high-performance liquid chromatography (HPLC) equipment described in this unit provides significant advantages over the original method in savings of both time and material. These large-zone AGFC experiments are performed using a 2.3-ml column packed with size-exclusion resin, an HPLC pump, a Teflon injection valve, a UV/visible detector with digital data collection capability, and a computer for data analysis. The validity of large-zone AGFC adapted to the HPLC format has been rigorously demonstrated (Nenortas and Beckett, 1994). The ten-fold reduction in column volume realized, as compared to the classical low pressure chromatography format, results in a dramatic savings of the material and time required to measure an equilibrium association constant. The method is of general use in the measurement of protein association processes. REDUCED-SCALE LARGE-ZONE ANALYTICAL GEL-FILTRATION CHROMATOGRAPHY In performing reduced-scale large-zone AGFC, one first prepares a size-exclusion column and verifies its suitability for use in large-zone experiments by analyzing a number of calibration standards of known size. Once a column is determined to be suitable for highly precise and accurate measurements of the sizes of known monomeric globular proteins, the protein of interest may be studied as a function of concentration. If the protein undergoes reversible assembly in the concentration range examined, a dependence of elution volume on protein concentration will be observed. Conversion of elution volume to molecular size yields information about the concentration dependence of the average size of the protein (see Support Protocol 1). Finally, the concentration dependence of average size can be analyzed using the appropriate stoichiometric model to obtain the stoichiometry and energetics of the protein assembly process (see Support Protocol 2). Contributed by Dorothy Beckett Current Protocols in Protein Science (1999) 20.5.1-20.5.13 Copyright © 1999 by John Wiley & Sons, Inc.
BASIC PROTOCOL
Quantitation of Protein Interactions
20.5.1 Supplement 18
Materials Gel-filtration resin (e.g., Sephacryl S-200, Amersham Pharmacia Biotech) Column running buffer (e.g., 50 mM Tris⋅Cl, pH 7.5/200 mM NaCl, 20°C) Column calibration standards (e.g., bovine serum albumin, hen egg ovalbumin, bovine pancreas ribonuclease A, bovine pancreas chymotrypsinogen A, glycine, and Blue Dextran 2000; Amersham Pharmacia Biotech) Protein sample to be tested Jacketed analytical glass columns (0.66-cm i.d.; 10-cm long; Omnifit) with end fitting and top column connection (Thomson Instrument; both with frits supplied by manufacturer) 5⁄ - and 1⁄ -in.-i.d. Tygon tubing 16 16 Ring stands with clamps Peristaltic pump High-performance liquid chromatography (HPLC) system with: HPLC pump (microbore), capable of delivering buffer at a flow rate of 0.152 ± 0.002 ml/min over long time periods (see Critical Parameters) Teflon syringe-loading sample injector (Rheodyne model 7125; Thomson Instrument) UV/visible detector and attached digital integrator of the detector signal 1⁄ -in.-o.d. Teflon tubing 16 0.25-cm-i.d. Teflon tubing Column-end-fitting Teflon ferrules, drilled to accept 1⁄16-in.-o.d. Teflon tubing Polymeric one-piece nut and ferrules Zero-dead-volume polymeric unions 20-µl Hamilton syringes Refrigerated circulating water bath Preweighed 1.5-ml conical microcentrifuge tubes Prepare resin for packing 1. Prepare gel-filtration resin for pouring the column by exchanging it into column running buffer at least five times. First exchange the resin in at least a two-fold excess (v/v) of buffer over the settled resin. Allow the resin to settled and then remove the buffer by aspiration. Repeat this procedure four more times. A semicompressible resin that will allow resolution of proteins in the desired size or molecular weight range should be used. See Critical Parameters and Troubleshooting for a discussion of size-exclusion resins. The salt concentration in the buffer should not be too low (e.g., <100 mM), as interaction between the protein and resin may occur at low salt concentration. The upper limit of the salt concentration depends on the particular resin. Some resins will compress at salt concentrations above 400 mM. All buffers should be prepared using Milli-Q-purified water or the equivalent, and should be filtered through 0.22-ìm filters prior to use.
2. Add buffer to yield a slurry composition of ∼40% (by volume) settled resin. Degas the 40% slurry and extra column running buffer under vacuum while preparing the column assembly for pouring. The degassing should be performed in a sealed flask, using an aspirator for ∼15 min.
Large-Zone Analytical Gel-Filtration Chromatography
Prepare column for packing 3. Connect two empty Omnifit columns in series using a short piece of 5⁄16-in.-i.d. Tygon tubing (Fig. 20.5.1). Connect the end fitting of the gel-filtration (bottom) column, including the frit, to the outlet tubing (1⁄16-in.-i.d., 20-in.-long Tygon tubing). The extra (top) column, which is identical to the bottom column, serves as a packing reservoir.
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column 2 (packing reservoir)
5/16-in. Tygon tubing connector
column 1 (column to be packed)
outlet tubing
bottom frit and end-fitting
Figure 20.5.1 Schematic drawing of the setup used for pouring the reduced-scale column for large-zone analytical gel-filtration chromatography. The lower column is to be packed and the upper is the reservoir for resin.
4. Attach the column assembly vertically to a ring stand using two clamps. 5. Add degassed buffer to the column assembly and remove bubbles from the bottom column frit by attaching the outlet tubing to a peristaltic pump and pumping buffer out of the column. 6. Remove the column outlet tubing from the pump and allow the buffer level to freely drain until it is even with the top of the bottom column frit. 7. Raise the column outlet tubing to the level of the top of the packing reservoir (Fig. 20.5.1), fasten the tubing to a second ring stand, and leave it unclamped in order to provide zero hydrostatic head pressure while pouring the column. Pack the column 8. Add the slurry dropwise to the column assembly until the level inside the packing reservoir is even with the column outlet tubing. Allow the resin to settle at zero hydrostatic head pressure for 15 min. Quantitation of Protein Interactions
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9. Decrease the outlet tubing level by 3 cm every 15 min, in order to slowly increase the hydrostatic pressure, until the resin is completely settled. Place a beaker under the outlet tubing to collect waste. This gradual increase in the pressure ensures even packing of the resin. The final column bed should have a 0.66-in. diameter, ∼6.8-cm length, and 2.33-ml volume. Since the total length of the two columns is ∼20 cm, ∼1 hr and 45 min is required for the complete settling process.
Finish the column 10. Connect a top column connection to the HPLC pump flow and allow the buffer to flow until the bubbles are expelled from its frit. 11. Remove the packing reservoir and the connector tubing from the column assembly, withdraw settled resin from the top of the bottom column, and insert the top column connection containing the frit. Remove enough resin to fit the column frit into the column with no dead volume remaining between the resin bed and the frit. These final steps should be carried out quickly without allowing introduction of air to the column.
12. Seat a short length of 1⁄16-in.-o.d. Teflon tubing in each of the column-end-fitting Teflon ferrules at the two ends of the column. Connect the other end of the tubing from the top of the column to a Teflon syringe-loading sample injector. Connect the tubing from the bottom of the column to the UV/visible detector. Each of these connections should be made through a polymeric one-piece nut and ferrules and zero-dead-volume polymeric unions. In order to minimize dead volume, make sure that the tubing lengths used to connect the column to the injector and detector are as short as possible. It is advisable to prepare these tubing connections prior to pouring the column. In addition, arrange the detector and pump so that the former is on the bottom. This will also ensure minimum dead volume.
13. Connect the jacketed column to a refrigerated circulating water bath adjusted to the desired experimental temperature. A temperature in the range of room temperature (20° to 25°C) should be used in order to avoid formation of bubbles. The buffer reservoir should be kept in this bath when the column is being run.
Evaluate column quality 14. Evaluate column quality by chromatographing small zones of a few (3 to 4) calibration standards. Prepare each standard in column running buffer at a final concentration of 1 to 2 mg/ml. Load a small zone in the sample injector loop for 6 sec with a Hamilton syringe, resulting in a 15-µl injection. Only columns that provide symmetrical peak shapes for small zones are suitable (Fig. 20.5.2, see Critical Parameters).
15. Once it is established that the shapes of the small zones are appropriate, equilibrate the column overnight by pumping buffer through it at a flow rate of 0.152 ml/min.
Large-Zone Analytical Gel-Filtration Chromatography
Calibrate the column 16. Calibrate the equilibrated column by making duplicate injections of small zones of glycine, Blue Dextran 2000, and the protein calibration standards prepared as described in Step 14. Load each small zone from the injector loop for 6 sec, corresponding to a 15-µl Teflon loop injection.
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0.20
Absorbance (280 nm)
0.15
0.10
0.05
0.00 0.0
1.0
2.0
3.0
Volume (ml)
Figure 20.5.2 Schematic representation of a small zone of protein run on the reduced-scale gel-filtration column. The peak shape should be nearly symmetrical as indicated in this figure.
17. Measure the column flow rate directly in each chromatographic analysis by weighing the column effluent, collected for a known time period into a preweighed 1.5-ml conical tube. Perform this measurement several times per run so that the mean and standard deviation of the flow rate are known for each run. The precision in the flow rate, which is determined by the precision of the HPLC pump, is critical to the success of the experiment.
18. Analyze the data obtained from the standards as described below (see Support Protocol 1), before proceeding to run large zones on the column. A suitable column will, on analysis of the linear plot of Stokes radius versus erfc−1σ, yield a correlation coefficient, or R2, ≥0.95. Once a column is established to be suitable, it can be operated continuously for ≤3 weeks with no apparent degradation in performance.
Perform chromatography of large zones 19. Construct large-zone sample loops from 0.25-cm-i.d. Teflon tubing to provide a total loop volume of 2.7 ml and attach to the injector sample loop using polymeric unions. 20. Prepare large zones of sample protein in running buffer at a total volume of 3.5 ml. The zones should be prepared in the concentration range over which the protein assembly takes place. For a monomer-dimer equilibrium process, ∼12 zones should be prepared that span the concentration range from 100% monomer to 100% dimer. The solution at each concentration should be prepared and equilibrated (in the constant temperature bath for the column) ∼15 min before loading.
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Absorbance (415 nm)
0.15
B
0.10
0.05
A
0.00 0.0
2.0
1.0
Vc
3.0
4.0
5.0
Volume (ml)
Figure 20.5.3 Characteristic large zone of oxyhemoglobin Ao, plateau [Hb dimer] = 5.62 µM in 100 mM Tris⋅Cl, pH 7.40 ± 0.01/100 mM NaCl/1 mM Na2 EDTA at 21.5°C, run on Sephacryl S-200. The centroid position, or Vc, is defined as the position on the elution volume scale of the solid vertical line that yields equal areas for regions A and B.
21. Inject the protein into the sample loop and load onto the column for 13.0 min, corresponding to a loaded sample of 1.95 ml. Repeat for all zones. Protein samples should be loaded in sufficient volume to obtain a plateau at the concentration of the loaded sample (Fig. 20.5.3). In order to achieve this on the column described, ≥1.95 ml sample must be loaded. If a longer column is poured a larger volume must be loaded. It is a good idea to check that a plateau has been achieved by comparing the absorbance (at 280 nm) in the presumed plateau region to that measured for the sample prior to loading. Equilibrium constant determinations are performed by running large zones at twelve concentrations and can be completed in one day. The concentrations of the large zones should span from 50-fold less to 50-fold more than the equilibrium dissociation constant for the assembly reaction. The concentrations of the protein in these samples should be evenly spaced on a logarithmic scale over this range. This is to ensure that data over the entire range from monomer to n-mer are obtained.
22. Obtain chromatograms by measuring the absorbance at a single wavelength versus time. For most proteins this wavelength is 280 nm. The sampling of the absorbance should be set at 5 points per second. Large-Zone Analytical Gel-Filtration Chromatography
23. Analyze chromatographic data (see Support Protocol 1) and determine the energetics and stoichiometry of the protein assembly process (see Support Protocol 2).
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DATA ANALYSIS Determination of Elution Volumes for Small Zones and Centroid Volumes for Large Zones Convert peak retention times to elution and centroid volumes by multiplication with the measured flow rate, which was determined by weighing volumes of column effluent collected over a known time period.
SUPPORT PROTOCOL 1
Measure elution volumes, Ve, for small zones as the elution volume at the peak apex. Elution volumes for large zones are analyzed as the centroids (equivalent sharp boundaries) of the leading or trailing boundaries of the zone (Ackers, 1975; Valdes and Ackers, 1979) as described below (see Fig. 20.5.3). Centroid volumes, Vc, are determined by trapezoidal integration of the data obtained from the integrator using MS-DOS 5.0 QBasic, developed in this laboratory. The centroid-volume calculation program requires input of ASCII-formatted y-axis data, supplied without file header information. The AUTOASCX file-conversion program available with the PE Nelson model 1020 integrator converts the y-axis raw data to an ASCII format. Other integrators should have the same capability. The centroid calculation program inputs every fifth data point of the ASCII-formatted y-axis data and stores the values in an array. Since the data collection rate is set to five data points per second, the array pointer is equivalent to the time in seconds. Two versions of the centroid calculation program have been developed in the author’s laboratory and are available upon request. The first version of the program requires the investigator to examine the y-axis data in a spreadsheet program and determine the average y-axis values that define the baseline before the leading edge, the plateau region at the leading edge, the plateau region at the trailing edge, and the baseline after the trailing edge. This version also requires inputs of the starting and ending times of the leading and trailing edges, respectively, as well as estimates for the centroid times of the leading and trailing edges. The second version of the centroid calculation program requires no input of parameters by the investigator. It uses empirically determined routines to calculate the parameters detailed above. This program cannot be used for large-zone profiles with baseline anomalies. A data collection rate of five data points per second results in very large ASCII input files for viewing in spreadsheet programs or plotting with graphing programs. Code for QBasic programs used to reduce the number of data points in an ASCII textfile is also available from the author. Column Calibration and Determination of Partition Coefficients and Stokes Radii for Proteins Calculate partition coefficients, σ, of protein small zones from the elution volumes of the zones using the following equation: σ = (Ve − Vo )/ Vi Equation 20.5.1
where Vo is the void volume of the column and Vi is the included volume. Vo corresponds to the elution volume of a small zone of Blue Dextran, while Vi + Vo is the elution volume of a small zone of glycine or some other small molecule that is completely included in the resin. Calibrate the column using the measured partition coefficients of the protein standards and the known Stokes radii of the calibration standards and applying the following linear equation:
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40
Stokes radius (Å)
35
30
25
20
15 0.4
0.5
0.6
0.7
0.8
erfc –1σ
Figure 20.5.4 Example of a calibration curve obtained from analyzing small zones of protein standards on a reduced-scale analytical gel-filtration column packed with Sephacryl S-200 HR resin. The standards from largest to smallest are BSA, ovalbumin, chymotrypsin, and RNase A. The solid line represents the best fit of the data to Equation 20.5.2.
a = ao + bo erfc −1σ Equation 20.5.2
where a is the molecular or Stokes radius (Å), erfc−1σ is the inverse error function complement of the partition coefficient (the values for erfc−1σ may be obtained from published tables; U.S. Government Printing Office, 1954), and ao and bo (the intercept and slope, respectively) are the linear calibration constants for the column (Ackers, 1967).The column calibration constants ao and bo are determined by linear least squares analysis of calibration standard data using Equation 20.5.2. The advantage of this method of calibrating the size-exclusion column over other traditional methods is that, by taking into account the realistic distribution of sizes of pores in the resin, it provides a more accurate relationship between molecular size and elution volume as represented by the partition coefficient. An example of a calibration curve for the type of column described in this unit is shown in Figure 20.5.4. Weight-average partition coefficients for self-associating protein large zones, denoted as σw, are calculated using Equation 20.5.3. σ w = (Vc − Vo )/ Vi Equation 20.5.3 Large-Zone Analytical Gel-Filtration Chromatography
where Vc is the centroid or equivalent sharp boundary volume of the large zone. The partition coefficients can be used to estimate the apparent or average Stokes radius of an interacting protein at a given concentration using the equation of the line determined from
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linear least squares analysis of the calibration data (Equation 20.5.2). Note that if the shape of a protein deviates significantly from spherical it will exhibit aberrant mobility in size-exclusion chromatography, which is typically evidenced in a larger-than-expected Stokes radius. DETERMINATION OF THE ENERGETICS AND STOICHIOMETRY OF THE PROTEIN ASSEMBLY PROCESS
SUPPORT PROTOCOL 2
Once the concentration dependence of the average size or partition coefficient of the protein is known (see Support Protocol 1) the data can be analyzed to obtain information about the stoichiometry and energetics of assembly. A monomer-dimer assembly process is the simplest to analyze, and the theory and procedure for analysis of data using this model is described in detail below. Other models are possible and it is important that they be tested in cases for which the stoichiometry of assembly is unknown (Valdes and Ackers, 1979). Monomer-Dimer Stoichiometric Model For a reversibly self-associating protein monomer, M1, there are theoretically several assembly steps available, which are indicated in the expressions M1 + M1 = M2, M1 + M2 = M3, through Mn-1 + M1 = Mn. Assuming a discrete assembly process (monomer to n-mer), the weight-average partition coefficient or σw for a mixture of monomer and n-mer can be expressed in terms of the weight fraction of monomer, α, σ w = ασ1 + (1 − α )σ n Equation 20.5.4
where α = C1/CT, C1 and CT are the unassembled monomer and total monomer concentrations, respectively, and σ1 and σn are the monomer and n-mer partition coefficients, respectively. The equilibrium association constant, Keq, can be expressed in terms of α as Keq =
C − αCT 1− α = T = n n −1 n α CT [ M1 ] (αCT ) [M n ]
n
Equation 20.5.5
Rearrangement of Equation 20.5.5 in terms of α and CT yields K eq CTn −1α n + α − 1 = 0 Equation 20.5.6
For the case of a monomer-dimer equilibrium where n = 2, solution of Equation 20.5.6 using the quadratic equation gives α=
2 2 CT + 4 K eq CT −1 + K eq
2 K eq CT Equation 20.5.7
Rearranging equation 20.5.4 using σn = σ2 gives:
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σ w = α ( σ1 − σ 2 ) + σ 2 Equation 20.5.8
Combination of Equation 20.5.7 with Equation 20.5.8 yields an expression for σw in terms of Keq, CT, σ1, and σ2. Nonlinear least squares analysis of σw versus concentration data provides values for the equilibrium constant, Keq, and the partition coefficients for the monomeric and dimeric proteins, σ1 and σ2. The analysis can be performed using any of the many programs that are commercially available (e.g., Origin; Microcal). Alternatively the NONLIN program developed by Dr. Michael Johnson may be used (Johnson and Faunt, 1992). Fitting of Data for a Monomer-Dimer Equilibrium Association Measurement Oxyhemoglobin is an example of a protein that undergoes a simple dimer-tetramer equilibrium and, since the hemoglobin dimer does not further dissociate to monomer at the concentrations relevant to the dimer-tetramer equilibrium, the equations that apply to a simple monomer-dimer association reaction also apply to this assembly reaction. Results of analysis of the σw versus hemoglobin dimer concentration, CT, data using Equations 20.5.7 and 20.5.8 provide the partition coefficients for the dimer, σD, and tetramer, σT, and the equilibrium association constant, Keq. σw = (
2 2 CT + 4 K eq CT −1 + K eq
2 K eq CT
)(σ D − σ T ) + σ T
Equation 20.5.9
1.00
Fraction Hb dimer
0.75
0.50
0.25
0.00 –2
Large-Zone Analytical Gel-Filtration Chromatography
–1
0 log [Hb dimer]
1
2
Figure 20.5.5 Association curve for the oxyhemoglobin Ao (Hb) dimer-tetramer equilibrium. Experimental fraction Hb dimer values (dots) were obtained at twelve micromolar Hb dimer concentrations. The solid line is simulated using the best resolved parameters determined by nonlinear least squares analysis.
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The experimental weight-average partition coefficient at each concentration is calculated using Equation 20.5.3. It can be related to weight fraction dimer, fD, using the following equation: fD = ( σ w − σ T )/( σ D − σ T ) Equation 20.5.10
where σT and σD are the experimentally determined tetramer and dimer partition coefficients obtained from nonlinear least squares analysis of the chromatography data using Equation 20.5.9. An association curve for the oxyhemoglobin dimer-tetramer reaction simulated using the best-fit parameters obtained from analysis of the experimental data is shown in Figure 20.5.5 along with the data. The equilibrium constant and Gibbs free energy for the dimer-tetramer association, Keq and ∆G, determined for the data are 1.0 ± 0.2 µM and −8.1 ± 0.1 kcal/mol, respectively. These are in excellent agreement with values determined using the standard large-zone AGFC technique (Ip and Ackers, 1977). The quality of the fit, which is judged by the square root of the variance (2% standard error) and the random distribution of residuals, indicates that the data conform well to this model, as expected. If the data deviate systematically from the best-fit curve, this may be an indication that the assembly model used to analyze the data is not correct. Other plausible models should then be tested. COMMENTARY Background Information Large-zone analytical gel filtration chromatography (AGFC) was developed as a tool for quantitative studies of macromolecular association in the 1960s. The method is analogous to other transport techniques including sedimentation and electrophoresis. The major advantage of AGFC over some of the other methods discussed in this book—including sedimentation, isothermal titration calorimetry, and surface plasmon resonance—is the low cost associated with the equipment required for the experiment. The original method described by Ackers and co-workers (Ackers, 1975; Valdes and Ackers, 1979) requires equipment found in any standard biochemistry laboratory. The two major weaknesses associated with the original technique include its large material and time requirements. The method described in this unit represents a scaling down of the original method for modern HPLC equipment. The equipment requirements for this method are not extraordinary for a biochemistry laboratory and the reduced-scale method represents significant savings of both material and time over the original large-zone AGFC (Nenortas and Beckett, 1994). As discussed in the protocol, the method is conceptually very simple and entails determination of the average size of a protein over the concentration range relevant to its association
from one oligomeric state to another. For example, if a protein undergoes a monomer-dimer transition, it is expected that this transition will be accompanied by a change in the average size of the protein in the relevant concentration range. Provided that the kinetics of the equilibrium are rapid relative to the time scale of the chromatography experiment, upon increasing the protein concentration from that at which the protein is fully monomeric to a concentration at which it is fully dimeric, the average size of the protein will increase. The size at concentrations at which there is a mixture of monomers and dimers will reflect a weight average of the contributions of the two species to the total species population. The goal of the size-exclusion chromatography experiment is to measure the average size as expressed in the weight-average partition coefficient (σw) as a function of the total protein concentration. As indicated in Equation 20.5.9, the change in protein size with protein concentration will reflect the specific association reaction that is relevant to the assembly process. The data are analyzed using the appropriate assembly model to determine the equilibrium constant for the reaction and the partition coefficients for the species that contribute to the reaction. In this unit only the simple monomer-dimer reaction has been discussed. It is possible for more complex equilibria to occur, and in those cases the data can be
Quantitation of Protein Interactions
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analyzed using the appropriate mathematical expressions. The reason for using large zones as opposed to small zones in determination of protein association equilibria is related to the dilution of a small zone that occurs in the course of its migration through the column. Because of this dilution, the protein concentration at the elution volume of a small zone is considerably lower than the concentration originally loaded on the column. The large-zone experiment, in which the leading and trailing edges of the zone are fed by a plateau of protein at the initial loading concentration, overcomes the dilution problem. Consequently the centroid of the zone represents the elution volume of the protein sample at the concentration at which it was originally loaded onto the column. The review by Becker (1988) provides a discussion of methods related to large-zone AGFC for study of protein association. In this unit the only method described for detection of chromatographically analyzed samples is UV/visible absorption. The use of fluorescence detection should increase the sensitivity of the method. Scintillation counting of collected fractions of the eluted proteins that have been labeled to very high specific activity extends the concentration range of protein detection down to the 10-picomolar range. Accessibility to these low concentrations extends the range of equilibrium constants measurable by this method to the nanomolar scale (Beckett et al., 1991).
Critical Parameters and Troubleshooting
Large-Zone Analytical Gel-Filtration Chromatography
Choice of size-exclusion resin Requirements of the resin used in large-zone AGFC are: (1) lack of interaction with the protein analytes, (2) fast diffusion rates of the interacting species in and out of the resin pores relative to the rate of chromatography (Valdes and Ackers, 1979), and (3) stability to a range of pHs and ionic strengths. The column described in this unit is prepared using Sephacryl S-200 HR resin (Pharmacia). Toyopearl HW-50 S (Toso Haas) was found to be unsuitable due to interaction with protein analytes as evidenced by tailing of small-zone peaks. Other size-exclusion resins may be used, but it is imperative that they be checked for interaction with proteins using the criterion of peak shape symmetry. In general Sephadex and the Bio-Gel P resins are good choices; however, care must be taken to avoid compression of the former resin. For a review of the properties of
size-exclusion resins the reader is referred to Dubin (1992). HPLC equipment The pump. In this experiment, column buffer is delivered at a flow rate of 0.152 ± 0.002 ml/min by an HPLC pump. The precision in the flow rate is critical for the success of the experiment. This is because the certainty with which an elution volume, and therefore the partition coefficient, of any protein can be measured is directly related to the precision of the flow rate. The author has used a Shimadzu LC-10AD pump, but any system that can operate at this low flow rate for long time periods (e.g., 36 hr) will do. The flow rate is measured for each analysis by weighing the column effluent, collected in a preweighed conical tube, for a measured period of time. The injector. A standard HPLC injector using stainless-steel sample loops proved unsuitable for use with the proteins analyzed due to sample carryover. Therefore, a Rheodyne model 7125 Teflon syringe-loading sample injector should be used to introduce samples. Small zones are loaded from the Teflon injector loop. Large-zone sample loops are constructed from 0.25-cm-i.d. Teflon tubing to provide a volume of 2.7 ml, and are attached to the injector sample loop using polymeric unions. The detector and integrator. Eluted zones of protein are detected using a UV/visible spectrophotometric detector and recorded on an integrator. Although, many other systems should be suitable, the author used a Shimadzu detector and a Perkin-Elmer Nelson Unit, respectively. It is important that the data from the integrator can be easily obtained in ASCII format. The linearity of the detector should be evaluated by measuring the absorbance of a series of weight dilutions of a protein stock on both the detector and a double-beam spectrophotometer. Dilutions should be prepared to give peak absorbance values between 0.05 and 1.0 AU for each wavelength. Absorbance measurements on both instruments should agree to within ±0.02 AU. The column. Omnifit analytical glass columns are packed to obtain a column gel bed of 0.66-cm i.d. and ∼6.8 cm length for a total column volume of 2.33 ml. A longer column can be used if it is required to obtain sufficient resolution in a particular associating system. Trailing of small zones Small zones should exhibit the typical Gaussian-shaped chromatographic peaks
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shown in Figure 20.5.2. If interaction of the protein standards with the resin occurs, as evidenced by small-zone peak tailing on the trailing edge, either the resin is unsuitable or a higher salt concentration should be used in the running buffer in order to disrupt the interaction of the protein with the resin.
for a simple protein-association equilibrium (i.e., monomer-dimer) requires 1 day to run. If a more complex equilibrium is occurring, data over a very broad concentration range may be required. In these cases data collection may require 3 days. Data analysis can be performed in 1 day.
Large zones Large-zone elution profiles are characterized by a relatively sharp leading boundary with a more diffuse trailing boundary separated by a plateau of constant concentration identical to the applied sample concentration (Fig. 20.5.4). These plateau concentrations should be verified by direct comparison of the detector absorbance to spectrophotometer absorbance, or by measuring the absorbance of the collected plateau column effluent on a spectrophotometer. In developing the reduced-scale large-zone AGFC, it has been found that while small-zone elution volume (Ve) and large-zone leading edge centroid volume (Vc) are, for a monomeric protein, consistently in good agreement, agreement with the trailing-edge Vc was found to depend on sample-loop volume. The minimum sample-loop volume that results in good agreement between the elution and centroid volumes of small and large zones was 2.7 ml. The HPLC sample introduction format requires buffer flow to load the sample from the loop. Diffusion occurs where the large-zone sample in the loop contacts the buffer during the 13.0 min sample loading time. In order to obtain trailing-edge data in agreement with leading-edge data, the loading process must be terminated before a sample that has been affected by dilution is loaded. Use of a 2.7-ml sample-loop volume results in good agreement between small-zone Ve and large-zone leading-edge and trailingedge Vc data. Note, however, that the leading edge information is sufficient in analysis of a protein assembly reaction.
Literature Cited
Time Considerations
Column packing requires ∼2 hr and initial calibration 20 min. The complete column calibration curve can be completed in 1 day. The series of large zones, twelve in total, required to determine the stoichiometry and energetics
Ackers, G.K. 1967. New calibration procedure for gel filtration columns. J. Biol. Chem. 242:32373238. Ackers, G.K. 1975. Molecular sieve methods of analysis. In The Proteins, Vol. 1 (H. Neurath, R.L. Hill, and C. Roeder, eds.) pp. 1-94. Academic Press, New York. Becker, G.W. 1988. Frontal boundary analysis in size exclusion chromatography of self-associating proteins. In Aqueous Size Exclusion Chromatography (P.L. Dubin, ed.) pp. 375-397. Elsevier Science Publishing, New York. Beckett, D., Koblan, K.S., and Ackers, G.K. 1991. Quantitative study of protein association at picomolar concentrations: The λ phage cI repressor. Anal. Biochem. 196:69-75. Dubin, P.L. 1992. Problems in aqueous size exclusion chromatography. In Advances in Chromatography, Vol. 31 (J.C. Giddings, E. Grushka, and P.R. Brown, eds.) pp.119-151. Marcel Dekker, New York. Ip, S.H.C. and Ackers, G.K. 1977. Thermodynamic studies on subunit association constants for oxygenated and unliganded hemoglobins. J. Biol. Chem. 252:82-87. Johnson, M.L. and Faunt, L.M. 1992. Parameter estimation by least squares methods. Methods Enzymol. 210:1-36. Nenortas, E. and Beckett, D. 1994. Reduced-scale large-zone analytical gel filtration chromatography for measurement of protein association equilibria. Anal. Biochem. 222:366-373. U.S. Government Printing Office. 1954. Tables of the Error Function and Its Derivative. N.B.S. Applied Mathematics Series 41, U.S. Government Printing Office. Washington, D.C. Valdes, R. and Ackers, G.K. 1979. Study of protein subunit association equilibria by elution gel chromatography. Methods Enzymol. 61:125142.
Contributed by Dorothy Beckett University of Maryland College Park, Maryland
Quantitation of Protein Interactions
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Size-Exclusion Chromatography with On-Line Light Scattering
UNIT 20.6
Several light-scattering techniques have been described in detail in UNIT 7.8. This unit will focus on the static light-scattering technique used in conjunction with size-exclusion chromatography (SEC-LS). SEC and static light scattering are complimentary techniques to determine the molecular weight of proteins. Integrating the two techniques offers numerous advantages over using either one in isolation. While batch-mode light scattering determines overall average molecular weights for heterogeneous and associating or interacting protein samples, size fractionation by SEC helps determine molecular weights of individual species in a sample solution. In addition, although SEC is a simple and fast method for estimating the molecular weights of many proteins based on their elution position, there are several problems with this conventional SEC approach. One is that the elution position depends not only on the molecular weight of the protein, but also on its shape. A protein with extended structure elutes earlier than expected and gives a higher molecular weight. Another problem is that the elution position will change if the protein has any tendency to interact with the column matrix. In addition, when dealing with a complex protein, i.e., glyco-, lipo-, or conjugated protein, SEC may not be able to determine its polypeptide molecular weight, since the carbohydrates usually have a disproportionately large effect on its elution position. When SEC is used together with light scattering, the molecular weight from this measurement is independent of the elution position, and for glycoproteins the molecular weight only of the polypeptide component can be determined if the extinction coefficient of the polypeptide alone is used in the analysis. There are several excellent reviews on the topic of SEC with on-line light scattering (Takagi, 1985, 1990; Stuting et al., 1992; Wyatt, 1993). This unit describes the use of size-exclusion chromatography with on-line light scattering, UV absorbance, and refractive index detectors (SEC-LS/UV/RI) to determine: (a) the molecular weight of simple proteins containing no carbohydrates (see Basic Protocol 1 and Alternate Protocols 1 and 3), (b) the molecular weight of glycoproteins (see Basic Protocol 3), and (c), most importantly, the molecular weight and stoichiometry of protein-protein complexes or protein-carbohydrate complexes (see Basic Protocols 2, 4, and 5, Alternate Protocol 4, and the Support Protocol). Multiangle light scattering (see Alternate Protocol 2) is also discussed. USING REFRACTIVE INDEX AND LIGHT SCATTERING TO CALCULATE THE MOLECULAR WEIGHT AND DEGREE OF SELF-ASSOCIATION OF PROTEINS CONTAINING NO CARBOHYDRATES (TWO-DETECTOR METHOD)
BASIC PROTOCOL 1
This method uses two detectors—one for light scattering (LS) and one for refractive index (RI), or, occasionally, UV absorbance (UV). For a protein or a complex that does not contain carbohydrates, lipids, or conjugates such as polyethylene glycol (PEG), the refractive index of a protein, dn/dc, is constant (∼0.186 ml/g) and nearly independent of its amino acid composition, where n is the refractive index and c is the concentration in g/ml. Hence, we can determine molecular weight of a protein, M, from the ratio of the signals from the two detectors, (LS) and (RI): Quantitation of Protein Interactions Contributed by Tsutomu Arakawa and Jie Wen Current Protocols in Protein Science (2001) 20.6.1-20.6.21 Copyright © 2001 by John Wiley & Sons, Inc.
20.6.1 Supplement 25
M = K ′ ( LS) / ( RI ) Equation 20.6.1
where K′ = KRI/[KLS(dn/dc)] (see Background Information for more details). This is the so-called “two-detector method.” It is the method most commonly used, but it is only valid when dn/dc is known, which is generally not true for glycoproteins, protein conjugates, or their complexes. An important feature of the two-detector method using RI is that it requires no knowledge of extinction coefficient of the proteins for data analysis. More applications using this two-detector method to study nonglycosylated proteins can be found in Narhi et al. (1993), Philo et al. (1993), Gombotz et al. (1994), Jackson and O’Brien (1995), and Zhang et al. (1995). A typical on-line SEC-LS/UV/RI system is diagrammed in Fig. 20.6.1. When using only two detectors, the LS and RI detectors are placed in the order indicated. NOTE: A helium or on-line vacuum degasser system, such as Agilent (P) 1100 Series vacuum degasser, should also be used, placed between the reservoir and HPLC pump. These treatments help minimize spikes in LS signal arising from particles and air bubbles. Materials Protein solutions of unknown molecular weight, at ∼1 mg/ml Calibration standards (also see recipe): ∼1 mg/ml ribonuclease, ovalbumin, and BSA in column buffer Elution (column) buffer (see recipe and UNIT 8.3) Size-exclusion (gel-filtration) chromatography column (UNIT 8.3) HPLC apparatus (UNITS 8.3 & 8.7) including light-scattering (miniDawn, Wyatt Technology) and refractive index (Agilent 1047A) detectors Additional reagents and equipment for gel-filtration chromatography (UNIT 8.3) and HPLC (UNIT 8.7)
light scattering detector
absorbance detector
solvent
pump
injector size-exclusion column
Size-Exclusion Chromatography with On-Line Light Scattering
refractive index detector
Figure 20.6.1 A typical configuration for SEC-LS/UV/RI (size-exclusion chromatography with on-line light scattering, UV absorbance, and refractive index detection).
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Prepare samples and calibrate instrument 1. Microcentrifuge unknown samples 10 min at maximum speed to remove large aggregates and particles, which would clog the filter and the SEC column. 2. Run a sufficient volume of elution (column) buffer through the SEC column to establish a stable baseline in both LS and RI detectors. 3. Inject 50 to 100 µl of calibration standard proteins into the SEC column. It is possible to mix and run the standards together as long as they are well resolved by the chromatography.
4. Determine the peak signals (height or width) from LS and RI detectors. Plot (LS)/(RI) versus molecular weight and fit the data points to a line passing through the origin, to determine the instrument constant, K′ (see Fig. 20.6.2). This protein standard method calibrates all detectors simultaneously, rather than one detector at a time. For an alternate calibration method, see Alternate Protocol 3.
4
LS/RI
3
2
1
0 0
20,000
40,000
60,000
80,000
Molecular weight
Figure 20.6.2 A typical plot of LS/RI versus the molecular weights of protein standards (RNase, ovalbumin, and BSA). 100 µl of each protein standard was injected onto a SEC column separately, to obtain more accurate data (slight overlap may occur for some SEC columns). Typical protein concentrations were 2.0, 1.5, and 1.5 mg/ml for RNase, ovalbumin, and BSA, respectively. Reproduced from Wen et al. (1996a) with permission of Academic Press.
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LS and RI signals (arbitrary units)
peak 2
peak 1
LS
RI
15
20
25
35
30
Retention time (min)
Figure 20.6.3 Chromatogram of BSA that contains monomers, dimers, and other oligomers. Commercial BSA from Sigma is a mixture of monomers, dimers, and higher oligomers. The molecular weight of peak 1 calculated from the two-detector method is 132,000, which agrees well with twice the BSA sequence molecular weight of 66,269, indicating that peak 1 is a BSA dimer. 100 µl of 4 mg/ml BSA was injected onto a Superose 6 column (Pharmacia) with PBS as eluent at 0.5 ml/min flow rate. The solid line is the LS signal and the dashed line is the RI signal. Reproduced from Wen et al. (1996a) with permission of Academic Press.
Calculate inter-detector delay correction 5. Inject a homogeneous protein solution to obtain a narrow peak. Use one of the protein standards.
6. Calculate the inter-detector volume. Determine difference in elution time between LS and RI detectors 7. Shift all subsequent RI peaks by the inter-detector volume. This compensates for the signal delay caused by the detectors being connected in series.
Determine sample molecular weight 8. Run samples of unknown molecular weight at two to three different concentrations. Size-Exclusion Chromatography with On-Line Light Scattering
9. Determine the LS and RI signal peaks (height or width). Calculate the molecular weight using Equation 1, above (also see Fig. 20.6.3).
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A
native RNase
LS and RI signals (arbitrary units)
RI
LS
B
reduced RNase RI
LS
C
18
native RNase
reduced RNase RI
RI
LS
LS
20
22
24
26
28
Retention time (min)
Figure 20.6.4 Chromatograms of native and reduced carboxymethylated (RCM)–RNase. 100 µl of each protein was injected onto a Superdex 75 column (Pharmacia) with PBS as eluent at a flow rate of 0.5 ml/min. (A) 1 mg/ml of native RNase; (B) 1 mg/ml of RCM-RNase; (C) chromatograms in panels A and B are put onto the same scale for comparing the LS/RI ratios of native and reduced RNase. The lines are the same as defined in Figure 20.6.3. Reproduced from Wen et al. (1996a) with permission of Academic Press.
Determine degree of self-association 10. Normalize the LS signal for one elution peak, so that the LS/RI ratio becomes one for that peak. The ratios of LS/RI for other peaks correspond to the degree of self-association. This self-association calculation is independent of molecular weight and works equally well for glycosylated proteins and proteins conjugated with other adducts. A combination of light-scattering data and elution position of a nonglycosylated protein sometimes can provide information not only on molecular weight but also on conformational change of the protein. To illustrate this, both native and reduced-carboxymethylated ribonuclease (RCM-RNase) were subjected to SEC as shown in Fig. 20.6.4. Despite the fact that the elution positions for native (Fig. 20.6.4A) and RCM-RNase (Fig. 20.6.4B) are very different, the molecular weights calculated from the two-detector method are the same for both, because their (LS)/(RI) ratios are the same. This can be clearly seen in Fig. 20.6.4C, in which the chromatograms are normalized to the same scale, as in Figure 20.6.3. If the molecular weights were calculated based only on their elution positions, there would be a different result, i.e., 13,700 for the native RNase and 41,000 for the RCM-RNase. The smaller elution volume of the RCM-RNase is due to unfolding. As shown by this example,
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SEC with light-scattering detection may be useful in some instances to provide information regarding the conformation of a protein, if the molecular weights derived from light scattering and from the elution position are compared. This also illustrates the importance of using the static, not dynamic scattering. If the dynamic scattering detector were used, one would not be able to demonstrate the monomeric state of RCM-RNase. ALTERNATE PROTOCOL 1
COMBINATION OF LS AND UV DETECTORS TO CALCULATE THE MOLECULAR WEIGHT OF NONGLYCOSYLATED PROTEINS (TWO-DETECTOR METHOD) One can use UV absorbance detection (UV), instead of RI, to monitor elution and hence determine protein concentration in the peak. The UV signal is given by: UV = K UV cε
Equation 20.6.2
where KUV is an instrument calibration constant and ε is the extinction coefficient of the protein (the absorbance of 1 mg/ml of a protein at 1-cm pathlength). In this case, instead of Equation 1, the following equation is used (see Background Information for derivation): M=
K UV
dn dc
K LS
2
×
ε ( LS )
( UV )
Equation 20.6.3
and hence with the same assumption that dn/dc is constant, the calibration constant can be determined by running the same protein standards as described in Basic Protocol 1. The ε used in the authors’ analyses for ribonuclease, ovalbumin, and BSA are 0.706, 0.735, and 0.670, respectively (Takagi, 1985). Thus, M can be determined from the linear plot of [ε (LS)]/(UV) versus M, if ε is known. ε can be calculated from the amino acid composition according to Equation 7 (see Basic Protocol 4). Use of UV may be preferred when RI has a low signal/noise ratio, as in the case where detergents are present. It has also an advantage when protein concentration is low, since UV has a higher sensitivity. ALTERNATE PROTOCOL 2
SEC/LS FOR LARGE PROTEINS: DEBYE ANALYSIS With a Wyatt Dawn multi-angle light-scattering detector (Wyatt Technologies), it is possible to measure light scattering at multiple angles of proteins eluting from an SEC column. In this case light-scattering signals in other angles need to be normalized to the calibrated 90° detector by injecting protein standards with narrow peaks, such as ovalbumin and BSA. Using protein concentration determined by RI (or UV), an angular dependence of light scattering (or molecular weight), known as a Debye plot, can be constructed. The molecular weight can be obtained from the intercept and the molecular radius can be obtained from the slope (see UNIT 7.8 and Wyatt manual). As described before, such analysis is only applicable for large proteins or protein complexes (mol. wt. >5 × 107).
Size-Exclusion Chromatography with On-Line Light Scattering
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A
UV
LS, UV, and RI signals (arbitrary units)
LS RI
B
UV
LS RI
C
LS
UV
RI 16
18
20
22
24
26
Retention time (min)
Figure 20.6.5 Chromatograms of TNF, sTNFR, and the mixture of sTNFR and TNF. 100 µl of each protein was injected onto a Superose 12 column (Pharmacia) with PBS as eluent at a flow rate of 0.5 ml/min. (A) TNF control sample (no sTNFR); (B) sTNFR control sample (no TNF); (C) a mixture of sTNFR and TNF made at a molar ratio of about three sTNFR per TNF. The solid line is the LS signal, the dashed line is the RI signal, and the dotted line is the UV signal. Reproduced from Wen et al. (1996a) with permission of Academic Press.
ABSOLUTE MOLECULAR WEIGHT CALIBRATION METHOD The details of the absolute calibration method can be found in Wyatt et al. (1988) and Wyatt (1993). In this method, the light scattering is calibrated using a pure solvent such as toluene instead of protein standards. Brief procedures for the Wyatt Dawn instrument are given here. The LS detector (at 90°) is calibrated with Rayleigh scattering of a pure solvent such as toluene. The RI detector also needs to be calibrated by using solutions (e.g., NaCl) with known dn/dc and with known concentrations. If the UV detector is used for the calculation, the UV calibration constant also needs to be calculated or measured. Currently the analytical software provided by manufacturers usually handles only two detectors at one time, even though signals from three detectors are monitored. For nonglycosylated proteins, with a constant value of dn/dc, one can use the software to calculate molecular weights from LS using either RI or UV for protein concentration measurements. However, for glycoproteins, the molecular weight and degree of glycosylation cannot both be independently determined using only two detectors. In this case, the three-detector method mentioned allows easy determination of the polypeptide molecular
ALTERNATE PROTOCOL 3
Quantitation of Protein Interactions
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Supplement 25
weight without measuring the dn/dc of the whole molecule (including the carbohydrates). When using the three-detector method, instrument calibration using standard proteins is much easier than the absolute calibration. BASIC PROTOCOL 2
CALCULATING THE STOICHIOMETRY OF A PROTEIN-PROTEIN COMPLEX CONTAINING NO CARBOHYDRATES USING LS/RI This protocol describes the method for determining the stoichiometry of a protein complex. If the complex does not contain carbohydrates or other conjugates, the two-detector method can be directly used to get the total polypeptide molecular weight of the complex and derive its stoichiometry. Such complexes may be either covalent or noncovalent, but in the latter case the affinity of the reversibly interacting components must be sufficiently high to maintain the integrity of the complex during chromatography. It is also important that the elution buffer be identical or close to the buffer used to prepare protein samples, since otherwise, the elution buffer may affect the binding between proteins. One can follow Basic Protocol 1 except that two molecules are mixed at various ratios at room temperature, centrifuged, and injected. Shown here is an example of the use of SEC-LS/UV/RI to study the stoichiometry of the complex of two nonglycosylated proteins, human tumor necrosis factor-α trimer (TNF) and the extracellular domain of the TNF type I receptor (sTNFR; Wen et al., 1996c). First, the molecular weight of each component needs to be characterized. The chromatograms obtained using a Pharmacia Superose 12 are shown in panels A and B of Fig. 20.6.5. Since both are nonglycosylated, the molecular weight can be calculated using the two-detector method and the calibration curve in Fig. 20.6.2. Thus, the molecular weight of each protein alone is 52,000 for TNF and 17,000 for sTNFR. A mixture made at a molar ratio of ∼3:1 sTNFR:TNF was then injected under the same experimental conditions, giving the chromatogram shown in Fig. 20.6.5C. Complexes of sTNFR with TNF elute as a broad distribution from 17.5 to 22 min. The molecular weight calculated by the two-detector method for the peak at around 18.5 min is 107,000, indicating that the stoichiometry of this complex is 3:1 sTNFR:TNF.
ALTERNATE PROTOCOL 4
CALCULATING THE STOICHIOMETRY OF PROTEIN-PROTEIN INTERACTIONS FOR NONGLYCOSYLATED PROTEINS USING LS/UV Following Alternate Protocol 1, it is also possible to employ UV, instead of RI, to do the same analysis for sTNFR or TNF using Equation 3, since ε for both proteins is known. However, for a complex formed by these proteins, the analysis is not so simple, since the ε value of the complex is not known. In order to overcome this problem, a self-consistent method as described below must be used to calculate the ε of the complex (from Equation 7) by assuming the stoichiometry, and then the molecular weight of the complex (obtained from Equation 3). If the assumption is correct, the calculated molecular weight should agree with the theoretical molecular weight from the assumed stoichiometry. For a more detailed description of the calculation protocol, see Basic Protocol 4.
BASIC PROTOCOL 3
Size-Exclusion Chromatography with On-Line Light Scattering
CALCULATING THE MOLECULAR WEIGHT OF GLYCOPROTEINS OR PROTEIN CONJUGATES USING THREE DETECTORS When a protein or a protein complex contains carbohydrates (or any other non-amino acid components that have a different dn/dc from a polypeptide and have near-zero absorbance at the wavelength used), its dn/dc is no longer known or constant because a carbohydrate usually has a different dn/dc than a polypeptide, and the carbohydrate content is normally unknown (and varies significantly from molecule to molecule). In such cases, we cannot
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use the RI detector to determine the concentration, c, for Equation 1, nor can we use Equation 3. We have to combine signals from all three detectors and use Equation 4: M=
( LS )( UV ) K LS K UV ε ( RI )2 2 K RI
Equation 20.6.4
where M and ε are the molecular weight and extinction coefficient of the entire glycoprotein, including carbohydrate. This equation is the basis for the “three-detector method.” However, in most cases the ε is unknown for glycoproteins, protein conjugates, or protein complexes. For this reason, therefore, Equation 4 is not commonly used, nor is the molecular weight of whole glycoproteins determined. The determination of ε requires a cumbersome experiment such as dry weight measurement (UNIT 3.4). What may be obtained relatively easily is the polypeptide extinction coefficient, εp, which is of course identical to ε for nonglycosylated proteins such as BSA. The value of εp can be either obtained from experimental data (such as quantitative amino acid analysis) or estimated with reasonable accuracy from the amino acid composition as in Equation 5 (Pace et al., 1995):
(
)
ε p ( 280 ) M−1cm −1 = 1/ MS ( no. of Trp × 5500 + no. of Tyr × 1490 + no. of Cys × 125 ) Equation 20.6.5
where MS (sequence molecular weight) is the polypeptide molecular weight and no. of Trp, no. of Tyr, and no. of Cys are the numbers of tryptophan, tyrosine, and disulfides in the protein, respectively. It is evident from this equation that MS can be readily obtained from the amino acid composition of the protein if the sequence is known. This is true for recombinant proteins that are encoded by the cDNA of known sequence. It is obvious from this that what this protocol tries to determine is not the absolute numerical values of the molecular weights, but the molecular weights for different peaks in the chromatogram, which may indicate the degree of self-association, the stoichiometry of complexes from interactions with other molecules, and conformational changes. All of this information cannot be derived from the sequence alone. If we use εp and select a wavelength where the carbohydrate (or conjugates) does not absorb, it is possible to algebraically eliminate all the contributions from the carbohydrates as described in Wen et al. (1996a), and calculate the molecular weights of polypeptide moiety, Mp, in glycoproteins, using the following equation: Mp =
( LS )( UV ) K LS K UV ε p ( RI )2 2 K RI
Equation 20.6.6
As shown in Equation 6, all contributions of the carbohydrate are canceled algebraically. Consequently, as shown in the example below, no knowledge of carbohydrate content is required for analysis. The polypeptide molecular weight can be measured directly as long as the polypeptide extinction coefficient is known. Equation 6 is the actual basis of the “three-detector method” used in the authors’ study of Chinese hamster ovary (CHO) and E. coli-derived recombinant stem cell factor (SCF). The results are shown below.
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LS, UV, and RI signals (arbitrary units)
24
26
28
30
32
Retention time (min)
Figure 20.6.6 Chromatograms of E. coli SCF. CHO SCF contains >30% carbohydrate and thus the three-detector method was used. 100 µl of 3.8 mg/ml of E. coli SCF was injected into a Superdex 200 column (Pharmacia) with PBS as eluent at a flow rate of 0.5 ml/min. The solid line is the LS signal, the dashed line is the RI signal, and the dotted line is the UV signal. Reproduced from Wen et al. (1996a) with permission of Academic Press.
1. Connect three detectors in series in the order of LS, UV, and RI (note that the order of LS and UV is interchangeable). 2. Prepare samples at 2 to 3 different protein concentrations, microcentrifuge 10 min at maximum speed, and use the clear supernatant for injection. 3. Calibrate the detector by injecting the same standard proteins as used in Basic Protocol 1. The instrument calibration constant, KRI2/(KLSKUV), can again be conveniently obtained by plotting [(LS)(UV)]/[εp(RI)2] against Mp. The εp values used in our analyses for RNase, ovalbumin, and BSA are the same as described in Basic Protocol 1.
4. The chromatograms obtained are shown in Fig. 20.6.6. Calculate the εp value from the amino acid composition. Note that the analysis requires no knowledge of exact carbohydrate content.
Size-Exclusion Chromatography with On-Line Light Scattering
5. The polypeptide molecular weight of CHO SCF calculated from the calibration curve of [(LS)(UV)]/[εp(RI)2] versus Mp is 38,000, which agrees quite well with the sequence molecular weight, 37,051, of the CHO SCF dimer (see Fig. 20.6.6). Even
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if SCF were not glycosylated, the three-detector method could be used, although it would not be necessary, with the same results. More applications using this three-detector method to study glycoproteins can be found in Arakawa et al. (1992) and Lu et al. (1995). The total molecular weight of CHO SCF estimated by the conventional SEC method is 113,000, whereas the experimental value (including carbohydrates) is 53,100 from sedimentation equilibrium (Arakawa et al., 1991). This error for the conventional SEC method is a consequence of both the asymmetric shape of CHO SCF and the disproportionately large effect of carbohydrates on the hydrodynamic size. This example illustrates the dangers of using the elution position alone to estimate the molecular weight of a protein.
DETERMINING THE STOICHIOMETRY OF A PROTEIN-PROTEIN COMPLEX CONTAINING CARBOHYDRATES
BASIC PROTOCOL 4
If a protein complex contains carbohydrates, the three-detector method is required to calculate its polypeptide molecular weight and determine its stoichiometry. In this section, we will focus on how to determine the stoichiometry of such a complex (Wen et al., 1996a,b). In order to use the three-detector method, we must know the polypeptide extinction coefficient of the complex. Since two UV-absorbing proteins form complexes, the extinction coefficient of polypeptide in the complex is no longer identical to that in the free state. In most common circumstances, only the polypeptide extinction coefficients of each protein in a complex is known, and thus we need to calculate the polypeptide extinction coefficient of the complex as a whole. The polypeptide extinction coefficient of a complex, εp, with a known stoichiometry (AmBn) can be calculated using the following equation: ε p = ( mε A MA + nε B MB ) / ( mMA + nMB ) Equation 20.6.7
where εA, εB, MA, and MB are the polypeptide extinction coefficients and molecular weights of A or B. After obtaining εp, we can calculate the polypeptide molecular weight by Equation 6 (or Equation 3 for nonglycosylated proteins in two-detector method; see Alternate Protocol 1), and, hence, determine the stoichiometry. It is obvious that this is a circular argument. On the one hand, we want to use Equation 5 to calculate the polypeptide extinction coefficient of the complex and then use Equation 6 to determine the corresponding molecular weight and the stoichiometry; on the other hand, the polypeptide extinction coefficient of the complex cannot be calculated from Equation 5 until the stoichiometry of the complex is known. In order to solve this conundrum, a self-consistent method has been developed. 1. Assume various possibilities for the stoichiometry of the complex, i.e., set m = 1 and n = 1, m = 1 and n = 2, m = 2 and n = 1, etc. Calculate molecular weight, MP, of the complex from each component, MA and MB, as: M P = mM A + nM B
Equation 20.6.8
2. Calculate εp from the assumed m and n using Equation 5.
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LS, UV, and RI signals (arbitrary units)
A
UV
LS
RI
B
UV
LS
RI
18
20
22
24
26
28
30
Retention time (min)
Figure 20.6.7 Chromatograms of sTrkB and the mixture of sTrkB and BDNF. 100 µl of each protein was injected onto a Superdex 200 column (Pharmacia) with PBS as eluent at a flow rate of 0.5 ml/min. (A) sTrkB control sample (no BDNF); (B) a mixture of sTrkB and BDNF made at a molar ratio of about two sTrkB per BDNF. The lines are the same as defined in Figure 20.6.5. Reproduced from Wen et al. (1996a) with permission of Academic Press.
3. Calculate experimental molecular weight of polypeptide in the complex using Equation 6 and the εp obtained above. 4. Compare the experimental value (from step 3) with the theoretical molecular weight, obtained from step 1. Correct assumptions for m and n should give an identical molecular weight between the experimental and the theoretical and will indicate the stoichiometry for the complex.
5. Analyze the results according to the Support Protocol. SUPPORT PROTOCOL
SAMPLE ANALYSIS: RECEPTOR-LIGAND INTERACTIONS This protocol provides two examples of the analysis of receptor-ligand interactions. Interactions of Extracellular Domains
Size-Exclusion Chromatography with On-Line Light Scattering
Interactions of the extracellular domains of the TrkB and TrkC neurotrophin receptors (sTrkB and sTrkC) with brain-derived neurotrophic factor (BDNF) and neurotrophin-3 (NT-3) provide one example of the analysis described in Basic Protocol 4 (Philo et al., 1994). It is evident that both sTrkB and sTrkC are glycosylated, and hence the three-de-
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Table 20.6.1
Summary of sTrkB/BDNF and sTrkC/NT-3 Resultsa
Samples
Assumed stoichiometry
Experimental ε mol. wt. from [ml/(mg⋅cm)] LS ± 5%
BDNF dimer NT-3 dimer sTrkB sTrkC sTrkB + BDNF mixture
— — — — 1sTrkB:1BDNF dimer 2sTrkB:1BDNF dimer 1sTrkC:1NT-3 dimer 2sTrkC:1NT-3 dimer
1.6 2.17 1.15 1.19 1.32 1.26 1.56 1.42
sTrkC + NT-3 mixture
27000 28000 44000 45000 110000 115000 118000 130000
Theoretical mol. wt. Correct assumption? from sequence 27300 27500 44100 44700 71000 116000 72000 117000
— — — — No Yes No Yes
aThe extinction coefficients and molecular weights in this table reflect only the polypeptide components of the complexes. Reproduced from Wen et al. (1996a) with permission of Academic Press. Bold letters indicate a correct stoichiometry.
tector method is needed to characterize their interactions with nonglycosylated ligands. When BDNF is injected into a Superdex 200 column with PBS as eluent, it shows no elution peak because of its interaction with the column matrix. Obviously if one wants to determine the molecular weight of BDNF, other columns or solvents must be used. For example, high-ionic-strength buffer may elute BDNF. However, such buffers may interfere with the interaction of BDNF with sTrkB. In this case, PBS can be used, since the complex of BDNF with sTrkB elutes. sTrkB is a glycoprotein and elutes with no indication of interaction with the column (Fig. 20.6.7A). An indication of such an interaction is a skewing of the elution peak, but as long as the injected materials elute and are equilibrated with the elution buffer, one can always calculate the molecular weight. The molecular weight determined from the three-detector method for sTrkB is 44,000, in agreement with the sequence molecular weight (Table 20.6.1), indicating that sTrkB exists as a monomer in solution. The chromatogram of a sTrkB/BDNF mixture made at a molar ratio of two sTrkB per BDNF dimer is shown in Fig. 20.6.7B. In order to calculate the molecular weight and stoichiometry of the sTrkB/BDNF complex, it is necessary to calculate its polypeptide extinction coefficient as discussed above. However, the extinction coefficient cannot be calculated until the stoichiometry of the complex is known. Therefore, we assume that either one (m = 1) or two (m = 2) sTrkB bind to one BDNF dimer (n = 1) and calculate the corresponding experimental (step 3) and theoretical (step 1) molecular weights. The results are summarized in Table 20.6.1. Obviously, the experimental molecular weight agrees with the theoretical one under the assumption that two sTrkB bind to one BDNF dimer. Therefore, we conclude that the stoichiometry, 2sTrkB:1BDNF dimer, is the correct one for the sTrkB/BDNF complex. Antigen-Antibody Interactions This protocol shows determination of stoichiometry of antigen-antibody interaction. Here an example is given using sHer2, which is the extracellular domain of erbB2/Her2 tyrosine kinase receptor, with monoclonal antibodies prepared against it (Kita et al., 1996; Wen et al., 1996b). In this case, both molecules are glycosylated. 100-µl samples of sHer2 and MAb controls, as well as mixtures of sHer2 with the MAb, were injected into the SEC-LS/UV/RI system separately. The results are summarized in Table 20.6.2 and the chromatograms are shown in Fig. 20.6.8. The molecular weight of the MAb was calculated as 139,000 using an average extinction coefficient value for MAbs of 1.4 ml/(mg⋅cm). The molecular weight of sHer2 was calculated with the extinction coefficient
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Table 20.6.2
Summary of Light-Scattering Results for sHer2 and MAbsa
Samples
Assumed stoichiometry
Experimental ε mol. wt. from [ml/(mg⋅cm)] LS ± 5%
sHer2 MAb 35 MAb 52 MAb 58 MAb 42b sHer+MAb35 mixture
— — — — — 1sHer2:1MAb35 2:1 3:1 1:2 1:3
0.85 1.4 1.4 1.4 1.4 1.24 1.14 1.08 1.31 1.41
69000 139000 151000 142000 136000 237000 261000 275000 226000 208000
Theoretical mol. wt. Correct assumption? from sequence — — — — — 208000 277000 346000 347000 486000
— — — — — No Yes No No No
aAll extinction coefficients and molecular weights in this table reflect only the polypeptide components of the complexes. Reproduced from
Wen et al. (1996a) with permission of Academic Press. Bold letters indicate a correct stoichiometry.
A RI
LS and RI signals (arbitrary units)
LS
B RI
LS
C
RI LS
16
18
20
22
24
26
Retention time (min)
Size-Exclusion Chromatography with On-Line Light Scattering
Figure 20.6.8 Chromatograms of MAb35, sHer2, and the mixture of sHer2 and MAb35. 100 µl of each protein was injected onto a Superdex 200 column (Pharmacia) with PBS as eluent at a flow rate of 0.5 ml/min. (A) MAb35 control sample (no sHer2); (B) sHer2 control sample (no MAb35). (C) A mixture of MAb35 and sHer2 made at a molar ratio of about 2.7 sHer2 per MAb35. The lines are the same as defined in Figure 20.6.3. Reproduced from Wen et al. (1996a) with permission of Academic Press.
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B
UV absorbance (280 nm, arbitrary units)
C
F
G D
E D
C E
B
20
25
30
35
Retention time (min)
Figure 20.6.9 Chromatograms of bFGF and HMWH mixtures (UV absorbance traces only). All samples contained 117 µM bFGF, and the amount of HMWH varied. The [HMWH]/[bFGF] molar ratio was 0.011:1 for B, 0.022:1 for C, 0.056:1 for D, 0.11:1 for E, 0.22:1 for F, and 0.43:1 for G. Sample A (bFGF control) is not shown here. 100 µl of each mixture was injected onto a Superdex 200 column (Pharmacia) with PBS as eluent at 0.5 ml/min flow rate. Reproduced from Wen et al. (1996a) with permission of Academic Press.
estimated from the amino acid sequence, and the result agrees well with its sequence molecular weight, indicating that sHer2 exists as a monomer in solution. The mixtures made with excess sHer2 all showed a peak eluting earlier than the MAb or sHer2 control. This indicates that complexes were formed in all mixtures. Different possibilities were assumed for the stoichiometry of each complex, and the corresponding experimental and theoretical molecular weights were thus calculated (Table 20.6.2). The experimental molecular weight is most consistent with the theoretical one under the assumption that the MAb binds two sHer2 molecules. More applications using this self-consistent three-detector method to study protein interactions can be found in Horan et al. (1995, 1996) and Philo et al. (1996a,b).
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Table 20.6.3
Summary of the Molecular Weights of bFGF/HMWH Complexesa
Sample letter in Fig. 20.6.9
Peak at 19 to 29 min Mixture molar ratio Intensity(based [HMWH]/[bFGF] Average mol. wt on UV)
Free bFGF at ∼34 min (based on UV)
A
Control (no HMWH) 0.011:1 0.022:1 0.056:1 0.11:1 0.22:1 0.43:1
B C D E F G
No peak
0.0
14
101000 102000 86000 84000 77000 44000
0.46 1.0 2.6 4.6 7.0 7.2
11 11 8.1 2.2 0.0 0.0
aAll molecular weights in this table reflect only the polypeptide components of the complexes. Reproduced
from Wen et al. (1996a) with permission of Academic Press.
BASIC PROTOCOL 5
ANALYSIS OF PROTEIN-HEPARIN INTERACTIONS USING THREE DETECTORS One can apply the three-detector method to a measurement of binding of a protein to a UV non-absorbing polymer, such as heparin. Here an example is given for binding of basic fibroblast growth factor (bFGF) to high-molecular-weight heparin (HMWH, M ≈ 16,000; Arakawa et al., 1994). By measuring the total polypeptide molecular weight of the complexes, we can determine how many bFGF molecules bind to one heparin molecule, i.e., how many molecules of bFGF are incorporated into the heparin-protein complex. Since heparin absorbs little light at 280 nm, the polypeptide molecular weight in the protein-heparin complex can be calculated by using the polypeptide extinction coefficient, εp (see Equation 8). In other words, when proteins bind to UV non-absorbing polymers, the polypeptide extinction coefficient in the complex is identical to that of free proteins. bFGF and HMWH are mixed at different ratios and injected into the SEC column. The eluent is monitored as in Basic Protocol 3. The chromatograms of bFGF and HMWH mixtures are shown in Figure 20.6.9, and the polypeptide molecular weights of each peak summarized in Table 20.6.3. The results indicate that each peak contains an average of 6 to 7 bFGFs (again no knowledge of heparin molecular weight is required for the calculation). Besides providing the molecular weight as described above, another important feature of SEC-LS/RI/UV (i.e., when three detectors are in use) is that all conventional SEC methods may also be used at any time because the data of UV and RI chromatograms are always included in the raw data file along with the LS data. In this particular example, we can use the UV chromatogram to determine the amount of bFGF remaining unbound to heparin. More detailed analysis, and the stoichiometry of low-molecular-weight heparin with bFGF and other protein-carbohydrate interactions, can be found in Kato et al. (1992), Arakawa et al. (1995), and Wen et al. (1997). It is obvious that the above method can apply not only to heparin, but also to PEG or any other conjugates that have a different dn/dc from a polypeptide and have near-zero absorbance at 280 nm.
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Elution (column) buffer Phosphate-buffered saline (PBS; APPENDIX 2E) is one commonly used column buffer. It works with many SEC columns. When a protein sticks to a column, excess sodium chloride may be added to the elution buffer to increase its ionic strength. Elution (column) buffers should be filtered before being added to the reservoir, and use of on-line filters may further improve the LS signals.
Protein calibration standards The commonly used standard proteins for obtaining the calibration curve are bovine serum albumin (BSA; M = 66,270), ovalbumin (M = 42,750), and ribonuclease (M = 13,600). Of course, one may use more than three proteins to obtain a more reliable calibration curve (Takagi, 1985).
COMMENTARY Background Information Light scattering has been one of the choices for determination of absolute molecular weight of macromolecules, although its use has been limited, in part, due to cumbersome sample handling measures required to minimize contamination by large particles and dust. This problem was eliminated upon introduction of laser light sources, which made flow-mode measurements possible. In addition, the laser light sources made it possible to obtain light scattering signals at low protein concentrations and for small proteins. In a flow-mode operation in conjunction with SEC, light scattering now can be used as a routine method to determine molecular weight of proteins in a standard biochemistry laboratory. Basic light scattering theory The basic light-scattering equation is: K ∗c
= R (θ ) 1 16 π2 2 2 1 + 2 〈rg 〉 sin ( θ / 2 ) + + 2 A2 c M 3λ Equation 20.6.9 where R(θ) is the excess intensity of light scattered at angle θ (i.e., the intensity due to the solute). K* is an optical parameter equal to [4π2n2(dn/dc)2]/(λ40NA), where c is the solute concentration in mg/ml, λ0 is the wavelength of the light in vacuum, M is the weight-average molecular weight, is the mean square radius of gyration, A2 is the second virial coef-
ficient, n is the refractive index, dn/dc is the refractive index increment of the solute, and NA is Avogadro’s number. At the low concentrations usually encountered during column chromatography, the virial coefficient term is negligible. In addition, the term [16π2sin2(θ/2)]/(3λ2) will also generally be negligible for proteins or 1 complexes with ⁄2 < 15 nm (Dollinger et al., 1992), which is true for a folded polypeptide with M < 5 × 107 (this approximation makes the approaches described in this unit valid for both single or multi-angle on-line light-scattering instruments available commercially). Thus, there is no angular dependence of light scattering. Under these conditions, the above basic light-scattering equation can be simplified to (K*c)/R(θ) = 1/M. Substituting K* with [4π2n2(dn/dc)2]/(λ40NA), the intensity of the light-scattering signal (LS) is given by:
( LS) = KLScM ( dn / dc )2 Equation 20.6.10 where KLS is an instrument calibration constant. Similarly, the refractive index signal, (RI), may be expressed as:
( RI ) = KRI c ( dn / dc ) Equation 20.6.11 where KRI is again an instrument calibration constant. These are the basic equations used in light-scattering measurements. Combining Equation 10 and Equation 11 leads to the twodetector method (see Basic Protocol 1).
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While the LS signal here refers to that determined at one angle, for some detectors there may be signals available at multiple angles (e.g., Dawn from Wyatt Technology). Except for proteins of high molecular weight, there should not be any significant angular dependence. Therefore, it is our general practice to use only the data for scattering at 90° (even though data are available at other angles), both for simplicity and because the 90° data usually have the highest signal/noise ratio. However, for multi-angle detectors it is certainly possible to use all the available data for the analysis, and the additional data may improve the overall accuracy of the molecular weight determination in some situations. For large proteins, it is possible to determine the mean radius of gyration with multi-angle detectors according to Debye analysis (see Alternate Protocol 2 and UNIT 7.8).
Critical Parameters Instrument In on-line SEC-LS/UV/RI, SEC serves as a filter and separation tool to remove large particles from protein samples and to fractionate proteins that may be heterogeneous in size. However, it may also generate noises. A stable pump is very important for LS and RI detectors, since the LS detectors are very sensitive to the particles from the column generated by the unstable flow and RI detectors are very sensitive to the pressure fluctuation. The three detectors following the column are in the following order: LS, UV, and RI. Although the position of the LS and UV detectors may be exchanged, the RI detector is normally the last one in the order since it is very sensitive to the back-pressure. As seen in Equation 10, besides UV and RI detectors, any other detectors that are able to determine protein concentration may be used together with the LS detector to determine a protein molecular weight with known dn/dc.
Size-Exclusion Chromatography with On-Line Light Scattering
Column selection Column selection is very important for obtaining protein recovery and separation. 1. Some proteins, and particularly their aggregates, have a tendency to stick to certain types of column matrices. This may cause recovery problems and result in an unusual elution position of the protein peak, causing it to overlap with the salt peak. Therefore, it is essential to choose columns that have the least tendency to bind the proteins. For proteins, in
general, polymer-based columns such as Superose or Superdex have a weaker tendency to bind proteins nonspecifically than silica-based columns, although silica-based columns, such as TosoHaas TSKgel G2000SWXL and TSKgel G3000SWXL, are routinely used in protein analysis by the authors. 2. The column should be long enough to bring the sample to complete equilibrium with the elution buffer and provide adequate separation. A complete separation allows determination of molecular weight of each species rather than the average molecular weight of the sample. Since an injection peak often appears at the void volume and the elution of salt peaks affects both RI and LS signals, the proper selection of column should allow the protein peak to elute between the void volume and salt peaks. 3. Particulate shedding from the column should be minimized. For most cases, a column such as TosoHaas TSKgel G2000SWXL, TSKgel G3000SWXL (7.8 mm i.d. × 30 cm length), or Pharmacia Superose or Superdex (10 × 300 mm) is adequate. Elution Elution buffers also play a critical role in performance of SEC. For most proteins, an eluent of sufficient ionic strength is required to minimize nonspecific binding of proteins. A conventional PBS (∼10 mM phosphate, ∼0.15 M NaCl) or PBS with a slightly higher salt concentration may provide enough ionic strength. Excessively high salt concentrations should be avoided, since this will change the dn/dc of proteins. The corrected dn/dc for proteins at different NaCl concentration may be derived as described in Arakawa and Wen (2000). Sample preparation In general, protein samples do not need to be dialyzed against the elution buffer before the measurements. However, when the protein elution position is close to the salt elution position, dialysis is recommended since it will minimize interference by the salt. Protein samples may be prepared in different concentrations according to the purpose of the analysis (the concentrations may vary from 0.05 mg/ml to 50 mg/ml, or in an even bigger range). However, the total mass injected should be controlled by adjusting the injection volume. Injection of too much sample may saturate the column and the detectors, and if the sample amount injected is not sufficient, it may not
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provide quality signals for analysis. A typical sample amount for each injection is between 20 µg and 200 µg, and this range may vary according to column size, detector sensitivity, and experimental objectives. Injection Before each injection, the column normally needs to be equilibrated with elution buffer until stable baselines are obtained for all detectors. A new column may require an extensive washing with the elution buffer before a stable LS signal can be obtained, since a new column normally generates more particles. An injection peak may appear at void volume, particularly for the light scattering signal, due to the pressure change. The baselines should be examined before each injection, because the baseline may not come back to the original level immediately after each run. If an automatic injector is used to inject a series of samples, the equilibration time between injections should be long enough to ensure that good baselines have been established for all detectors before each injection. Calibration, calculation, and applications In order to calculate molecular weights of proteins from light scattering, the instrument calibration constant must be determined. There are two major calibration methods, i.e., the absolute molecular weight calibration method (Wyatt, 1993) and the protein standard calibration method. Since the absolute molecular weight calibration method can only be carried out using specific analytical software, this unit focuses on the protein standard calibration method. There are several reasons why the authors prefer the protein standard method to calibrate the instrument: (a) the calibration can be easily done, (b) all detectors are calibrated at the same time, and (c) the method does not require specific analytical software. The protein standard calibration method has been used for many years in their laboratory and the data from this method is very reliable. Either peak height or peak area can be used to obtain the values of LS and RI signals when the sample is homogeneous. When the sample is heterogeneous and the resolution between peaks is poor, the authors use the peak height method. Using peak height, one can calculate the molecular weight distribution, slice-byslice, through the whole chromatogram and obtain number-average, weight-average, and z-average molecular weights, and polydispersity (Wyatt et al., 1988; Wyatt, 1993).
Troubleshooting The following are a series of potential problems that may arise in SEC-LS, along with recommendations for solving them. Weak or no signals 1. Make sure that all detectors are turned on. 2. Sometimes a protein may stick to a column. One may use standard proteins, such as BSA (inject 50 to 100 µg and use PBS as elution buffer), to test the instrument. If signals are found as usual for the standard proteins, this may indicate that the sample protein may stick to the column or the amount injected is too low. Noise in RI 1. Make sure that the change from one buffer to another is complete. This can be done by sufficient purging of the pump (it may take 10 to 30 min, depending on the size of the tubing from the buffer reservoir to the pump). 2. Make sure that the pump pressure is stable. A stable pump is very important for RI detectors. 3. Make sure that the RI detector is not subject to any back-pressure. 4. Let the RI detector warm up for a long enough period of time. Noise in LS 1. Make sure the column is washed sufficiently. It may take hours or a day to wash a column. Whenever changing elution buffers from one to another, it may take additional hours to wash the column. For a new column, it may take days to wash the column before a good LS baseline can be obtained, because a new column normally generates a large amount of particles. It is recommended that a new column be washed for at least 24 hr before connecting it to the detectors; otherwise, a large number of particles may reach the detectors. 2. Make sure that the pressure of the pump is stable, because the unstable flow may generate particles from the column. Abnormal molecular weight 1. Make sure that the signals for each detector are aligned correctly and the numbers for inter-detector delays are input correctly. If the signals for each detector are not aligned, this may cause abnormal molecular weight. 2. Make sure to input the correct dn/dc value when using absolute molecular weight calibration method.
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3. Make sure that light scattering and RI signals are clean enough and their signal/noise ratios are high enough. 4. When using UV and εP calculated from the polypeptide sequence, the calculation assumes that the polypeptide sequence is intact and that any conjugations present do not result in a change of εP. If the εP of the protein is no longer identical to that from the sequence, then such assumption will lead to abnormal molecular weight. The intactness of the polypeptide can be checked by N- and C-terminal sequencing or mass spectrometry for simple proteins. 5. When using LS at more than one angle, extrapolation to zero angle may lead to abnormal molecular weight, if normalization is done incorrectly. This is particularly true when signal is weak, since noise at angles other than 90° becomes too large. Check the molecular weight at 90°. As described in the previous section, there should be no angular dependence of LS for most proteins. The molecular weight calculated from LS at 90° should be correct.
Anticipated Results
Literature Cited Arakawa, T. and Wen, J. 2000. Refractive index of proteins in aqueous NaCl. Anal. Biochem. 280:327-329. Arakawa, T., Yphantis, D.A., Lary, J.W., Narhi, L.O., Lu, H.S., Prestrelski, S.J., Clogston, C.L., Zsebo, K.M., Mendiaz, E.A., Wypych, J., and Langley, K.E. 1991. Glycosylated and unglycosylated recombinant-derived human stem cell factors are dimeric and have extensive regular secondary structure. J. Biol. Chem. 266:1894218948. Arakawa, T., Langley, K.E., Kameyama, K., and Takagi, T. 1992. Molecular weights of glycosylated and nonglycosylated forms of recombinant human stem cell factor determined by low-angle laser light scattering. Anal. Biochem. 203:53-57. Arakawa, T., Wen, J., and Philo, J.S. 1994. Stoichiometry of heparin binding to basic fibroblast growth factor. Arch. Biochem. Biophys. 308:267-273. Arakawa, T., Holst, P., Narhi, L.O., Philo, J.S., Wen, J., Prestrelski, S.J., Zhu, X., Rees, D.C., and Fox, G.M. 1995. The importance of Arg40 and 45 in the mitogenic activity and structural stability of basic fibroblast growth factor: Effects of acidic amino acid substitutions. J. Prot. Chem. 14:263274.
In most cases, a symmetrical chromatogram such as that shown in Figure 20.6.3 is observed. The molecular weight of proteins in such peaks is calculated according to the protocols described in this unit. The molecular weight of a protein thus determined is an integer of the sequence molecular weight of the protein or a numerical sum of two proteins forming a complex. A skewed peak signifies heterogeneity in the stoichiometry of the complex.
Dollinger, G., Cunico, B., Kunitani, M., Johnson, D., and Jones, R. 1992. Practical on-line determination of biopolymer molecular weights by high-performance liquid chromatography with classical light-scattering detection. J. Chromatogr. 592:215-228.
Time Considerations
Horan, T., Wen, J., Arakawa, T., Liu, N., Brankow, D., Hu, S., Ratzkin, B., and Philo, J.S. 1995. Binding of Neu differentiation factor with the extracellular domain of Her2 and Her3. J. Biol. Chem. 270:24604-24608.
The time for each run varies according to the size and cleanliness of the column and the sensitivity required for the experiments. In general, a new, dirty, or large-size column requires a longer time to wash. The higher the sensitivity required, the longer it may take. For a typical experiment using a clean TosoHaas TSKgel G2000SWXL or TSKgel G3000SWXL (7.8 mm i.d. × 30 cm length) column with 0.5 ml/min flow rate, it may take 1 to 2 hr to equilibrate the column, 15 min to do one run, and about 10 to 60 min to analyze the data if everything goes well.
Gombotz, W.R., Pankey, S.C., Phan, D., Drager, R., Donaldson, K., Antonsen, K.P., Hoffman, A.S., and Raff, H.V. 1994. The stabilization of a hum an Ig M mon oclon al an tibo dy with poly(vinylpyrrolidone). Pharm. Res. 11:624632.
Horan, T., Wen, J., Narhi, L., Parker, V., Arakawa, T., and Philo, J. 1996. Dimerization of the extracellular domain of granuloycte-colony stimulating factor receptor by ligand binding: A monovalent ligand induces 2:2 complexes. Biochemistry 35:4886-4896. Jackson, C. and O’Brien, J.P. 1995. Molecular weight distribution of Nephila clavipes dragline silk. Macromolecules 28:5975-5977. Kato, A., Kameyama, K., and Takagi, T. 1992. Molecular weight determination and compositional analysis of dextran-protein conjugates using low-angle laser light scattering technique combined with high-performance gel chromatography. Biochim. Biophys. Acta 1159:22-28.
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Kita, Y., Tseng, J., Horan, T., Wen, J., Philo, J., Chang, D., Ratzkin, B., Pacifici, R., Brankow, D., Hu, S., Luo, Y., Wen, D., Arakawa, T., and Nicolson, M. 1996. ErbB receptor activation, cell morphology changes, and apoptosis induced by anti-Her2 monoclonal antibodies. Biochem. Biophys. Res. Commun. 226:59-69. Lu, H.S., Chang, D., Philo, J.S., Zhang, K., Narhi, O.L., Liu, N., Zhang, M., Sun, J., Wen, J., Yanagihara, D., Karunagaran, D., Yarden, Y., and Ratzkin, B. 1995. Studies on the structure and function of glycosylated and nonglycosylated neu differentiation factors. Similarities and differences of the alpha and beta isoforms. J. Biol. Chem. 270:4784-4791. Narhi, L.O., Rosenfeld, R., Wen, J., Arakawa, T., Prestrelski, S.J., and Philo, J.S. 1993. Acid-induced unfolding of brain-derived neurotrophic factor results in the formation of a monomeric “A state.” Biochemistry 32:10819-10825. Pace, C.N., Vajdos, F., Fee, L., Grimsley, G., and Gray, T. 1995. How to measure and predict the molar absorption coefficient of a protein. Prot. Sci. 4:2411-2423. Philo, J.S., Rosenfeld, R., Arakawa, T., Wen, J., and Narhi, L.O. 1993. Comparison of solution properties of human and rat ciliary neurotrophic factor. Biochemistry 32:10812-10818. Philo, J., Talvenheimo, J., Wen, J., Rosenfeld, R., Welcher, A., and Arakawa, T. 1994. Interactions of neurotrophin-3 (NT-3), brain-derived neurotrophic factor (BDNF), and the NT3.BDNF heterodimer with the extracellular domains of the TrkB and TrkC receptors. J. Biol. Chem. 269:27840-27846. Philo, J.S., Aoki, K.H., Arakawa, T., Narhi, L.O., and Wen, J. 1996a. Dimerization of the extracellular domain of the erythropoietin (EPO) receptor by EPO: One high-affinity and one low-affinity interaction. Biochemistry 35:1681-1691. Philo, J.S., Wen, J., Wypych, J., Schwartz, M.G., Mendiaz, E.A., and Langley, K.E. 1996b. Human stem cell factor dimer forms a complex with two molecules of the extracellular domain of its receptor, Kit. J. Biol. Chem. 271:6895-6902. Stuting, H.H., Krull, I.S., Mhatre, R., Krzysko, S.C., and Barth, H.G. 1992. High performance liquid chromatography of biopolymers using on-line laser light scattering photometry. LC-GC 7:402415. Takagi, T. 1985. Determination of protein molecular weight by gel permeation chromatography equipped with low-angle laser light scattering photometer. In Progress in HPLC (Paryez et al., eds.) vol. 1, pp. 27-41. VNU Science Press, Utrecht, The Netherlands.
Takagi, T. 1990. Application of low-angle laser light-scattering detection in the field of biochemistry. J. Chromatogr. 506:409-416. Wen, J., Arakawa, T., and Philo, J.S. 1996a. Size-exclusion chromatography with on-line light-scattering, absorbance, and refractive index detectors for studying proteins and their interactions. Anal. Biochem. 240:155-166. Wen, J., Arakawa, T., Talvenheimo, J., Welcher, A., Horan, T., Kita, Y., Tseng, J., Nicolson, M., and Philo, J.S. 1996b. A light-scattering/size exclusion chromatography method for studying the stoichiometry of a protein-protein complex. In Techniques in Protein Chemistry VII (D.R. Marshak, ed.) pp. 23-31. Academic Press, San Diego. Wen, J., Philo, J.S., Arakawa, T., Rosendahl, M., and Narhi, L.O. 1996c. A light-scattering/size exclusion chromatography method for studying the stoichiometry of a protein-protein complex. In Techniques in Protein Chemistry VII (D.R. Marshak, ed.) pp. 23-31. Academic Press, San Diego. Wen, J., Arakawa, T., Wypych, J., Langley, K.E., Schwartz, M.G., and Philo, J.S. 1997. Chromatographic determination of extinction coefficients of non-glycosylated proteins using refractive index (RI) and UV absorbance detectors: Applications for studying protein interactions by size exclusion chromatography with light-scattering, UV and RI detectors. In Techniques in Protein Chemistry VIII (D.R. Marshak, ed.) pp. 113-119. Academic Press, San Diego. Wyatt, P.J. 1993. Light-scattering and the absolute characterization of macromolecules. Anal. Chim. Acta 272:1-40. Wyatt, P.J., Jackson, C., and Wyatt, G.K. 1988. Absolute GPC determinations of molecular weights and sizes from light scattering Am. Lab. 20:86-89. Zhang, M.Z., Wen, J., Arakawa, T., and Prestrelski, S.J. 1995. A new strategy for enhancing the stability of lyophilized protein: The effect of the reconstitution medium on keratinocyte growth factor. Pharm. Res. 12:1447-1452.
Contributed by Tsutomu Arakawa Alliance Protein Laboratories, Inc. Thousand Oaks, California Jie Wen Amgen Inc. Thousand Oaks, California
We thank Dr. Gay-May Wu and Dr. John Philo for their many useful suggestions. Quantitation of Protein Interactions
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Analytical Ultracentrifugation: Sedimentation Velocity Analysis The analytical ultracentrifuge is a high-speed centrifuge with an optical system allowing observation of material during the sedimentation process. Several types of optical system are available for observation of the concentration of macromolecules as a function of radius and time. There are two basic modes of operation for the ultracentrifuge: velocity mode and equilibrium mode. In the equilibrium mode, relatively low speeds are used such that an exponential gradient of macromolecule is allowed to build with sedimentation, proceeding until it is opposed by diffusion so that no further concentration changes occur. When the system is at equilibrium, the radial dependence of the concentration distribution is determined by the molar mass distribution. Appropriate analysis of the concentration gradient allows determination of molar mass, stoichiometry, and equilibrium constants for interacting and noninteracting systems. The equilibrium mode is discussed in UNIT 20.3. The current unit concerns the velocity mode of operation.
SEDIMENTATION VELOCITY THEORY In a sedimentation velocity experiment, the concentration distribution evolves as a function of both radius and time. Normally, relatively high speeds are used so that a boundary is formed between the solution of sedimenting macromolecule and the buffer in which it is dissolved. Analysis of the rate of boundary movement and evolution of its shape can yield information about the molar masses of species present, as well as stoichiometries and equilibrium constants for their interactions. It is convenient to classify systems into two main categories: noninteracting and interacting systems. Each of these can be either ideal or nonideal. In addition, interacting systems can be either rapidly reversible, so that chemical equilibrium is maintained during sedimentation, or kinetically limited so that chemical equilibrium is not maintained during sedimentation. For a good general discussion of sedimentation velocity analyses, the reader is referred to the monographs by Schachman (1959) and Stafford and Schuster (1995). More detailed theoretical treatments can be found in Williams et al. (1958) and Claesson and Moring-Claesson (1961).
Contributed by Walter Stafford Current Protocols in Protein Science (2003) 20.7.1-20.7.11 Copyright © 2003 by John Wiley & Sons, Inc.
UNIT 20.7
Monodisperse Ideal System The simplest type of system that one is likely to encounter is the monodisperse ideal system. For a monodisperse macromolecular solution, one can determine the molar mass and frictional coefficient in a sedimentation velocity experiment. In the classical sedimentation velocity experiment, one measures the rate of movement of the midpoint of the boundary to determine the sedimentation coefficient of the macromolecule. More sophisticated methods accessible with desktop and larger computers are now more commonly used. Transport by sedimentation and diffusion in centrifugal field is described by the continuity equation, also know as the Lamm Equation (Lamm, 1929): 1 ∂ ∂c = − [r ( J sed − J dif )] r ∂r ∂t r Equation 20.7.1
where c is the concentration, which is a function of radius and time, r is the radial distance from the center of rotation, and t is the time of sedimentation from the beginning of the run. The Lamm equation relates the time dependence of the concentration distribution to the fluxes due to sedimentation, Jsed, and diffusion, Ddif, respectively. The Lamm equation is essentially a statement of the conservation of mass. The flux due to sedimentation is just the concentration times the velocity, and is given by:
J sed = c
dr dt
= csω2 r
Equation 20.7.2
where c is the concentration, ω is the angular velocity of the rotor, and r is the radius. The sedimentation coefficient (s) is a primary physical property of the molecule and is defined as the velocity of the particle in a gravitational field divided by the field strength: d /d s ≡ r2 t ω r Equation 20.7.3 Quantitation of Protein Interactions
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The sedimentation coefficient depends both on the molecular mass and shape according to the following relationship: s=
M (1 − νρ) Nf
computed from crystallographic coordinate data. The diffusion coefficient, D, is related to the frictional coefficient through the Stokes-Einstein equation: D=
RT Nf
Equation 20.7.4
where f is the frictional coefficient related to the shape by the Stokes equation:
Equation 20.7.9
and is defined by Fick’s First Law of diffusion:
∂c ∂x t
f = 6 πη0 Rs
J dif = − D
Equation 20.7.5
where ηo is the viscosity of the solvent and Rs is the Stokes radius. The Stokes radius is the radius of a hydrodynamically equivalent sphere. The frictional ratio, which is a measure of the deviation of the macromolecule from sphericity, is the ratio of the observed Stokes radius to the radius of a sphere with the same hydrated volume as the macromolecule and is defined by:
f f0
=
R0
where Ro is the radius of sphere with the same volume as the macromolecule (including hydration) and is given by: f 0 = 6πη0 R0 Equation 20.7.7
where Ro is given by:
)
1/3
3 ν2 + δ1ν10 M R0 = 4πN Equation 20.7.8
Analytical Ultracentrifugation: Sedimentation Velocity Analysis
where Jdiff is the flux per unit cross-sectional area due to diffusion. After substituting, the continuity equation becomes:
∂c 1 ∂ ∂c 2 2 = rD − 2ω r cs ∂t r r ∂r ∂r t Equation 20.7.11
Rs
Equation 20.7.6
(
Equation 20.7.10
where ν2 is the partial specific volume of the macromolecule, δ1 is the hydration coefficient, ν01 is the specific volume of pure water, M is the molar mass of the macromolecule, and N is Avogadro’s number. The frictional ratio forms the basis of shape analysis of macromolecules and can be compared to values of f/fo for various models of macromolecular shape ranging from ellipsoids of revolution to detailed bead or shell models
A noninteracting system can be composed of one or more independently sedimenting species which give rise to overlapping, additive, essentially Gaussian boundaries whose spreading is determined solely by diffusion. Interacting systems, on the other hand, give rise to so-called reaction boundaries. Reaction boundaries, by their very nature, cannot be resolved into individual contributions from the several species present. For example, for a selfassociating system, the Law of Mass Action must be obeyed at each point in a reaction boundary such that the amount of each species is determined by the equilibrium constant and total concentration. The shape of the boundary is determined by coupling sedimentation transport with re-equilibration such that mass action is obeyed at each point in the boundary. Having made that distinction, a description of each type of system is provided below along with the ways in which they may be analyzed.
Polydisperse, Noninteracting Systems For a polydisperse, noninteracting system, it must be determined that the system is in fact noninteracting by demonstrating that sedimentation rate and boundary shape are independent of concentration. Once concentration independence has been established, the boundary may be analyzed by various methods capable
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of resolving independent boundaries. A noninteracting system will be composed of several superimposable independent boundaries that are essentially Gaussian in shape. The boundary for each species is transported according to its sedimentation coefficient and it spreads according to its diffusion coefficient. In the cylindrical coordinate system of the rotating reference frame of the centrifuge cell, the shape of the boundary is not exactly a Gaussian, but is sufficiently close that it can be treated as Gaussian. The variance of the Gaussian curve that represents the boundary is proportional to the product Dt.
Polydisperse, Interacting Systems For interacting systems, a different approach has to be applied to determine the parameters describing the system. Interacting systems can be either single-component or multicomponent. An example of a single-component interacting system is a self-associating system such as a monomer-dimer or monomer-dimertetramer system. It can be characterized by a single initial total concentration and the equilibrium constants for each reaction. For an incompressible system, the concentration of each macromolecular species is uniquely determined by the equilibrium constants, and the total concentration according to the Law of Mass Action. Therefore, for a given monomerdimer system, for example, the amount of monomer and dimer present at each point in the cell is uniquely determined by the concentration at that point. Because the system is in reversible chemical equilibrium at each point, no resolution into monomer and dimer boundaries can occur. This type of boundary is called a reaction boundary, and its shape and evolution with time is characteristic of the stoichiometry and equilibrium constants (Cann and Goad, 1968).
Nonideality Nonideality can be manifested either hydrodynamically, through concentration dependence of the frictional coefficient, or thermodynamically, through concentration dependence of the activity coefficient. The former affects both sedimentation and diffusion, while the latter affects diffusion through its effect on the chemical potential gradient which is the driving force for diffusion. There are two main types of hydrodynamic concentration dependence: one due to backflow created as the macromolecule displaces buffer as it sediments, and the other due to the so-called primary charge effect.
The primary charge effect is a result of the requirement of electroneutrality. At low ionic strength, sedimentation of the macromolecule is retarded by the drag of its more slowly sedimenting counter-ions. At higher ionic strengths, where there is an excess of counterions, the effect of counter-ion drag becomes negligible. The effect of counter-ions on the diffusion coefficient is to increase the rate of diffusion because the counter-ions are more mobile than the macromolecule and tend to pull it along. The nonideal effects result in a decrease in sedimentation coefficient with an increase in protein concentration, and are treated commonly by the following mathematical formula: s (c ) =
s (c = 0 ) (1 + ks c )
Equation 20.7.12
where s is the sedimentation coefficient (see below), c is the concentration, and ks is the concentration dependence of the sedimentation coefficient. The value ks depends on a number of factors including the ionic strength, shape, and charge of the molecule. Sometimes a linear form is used, but the inverse form is applicable over a wider range of concentration. For the diffusion coefficient, D(c), the expression is:
D(c) = D(c = 0)
(1 + 2 BM1c ) (1 + ks c )
Equation 20.7.13
where M1 is the molecular mass of the diffusing component and B is the colligative second virial coefficient (Tanford, 1961), which depends on both excluded volume and charge. When determining either molar mass or shape from frictional information, it is best to work at the lowest concentrations possible to avoid nonideality. For interacting systems, concentrations near the value of the dissociation constant must be used, and for many systems of interest, these concentrations will be sufficiently low that nonideality effects can be ignored. At moderate concentrations, nonideality can be treated using the above equations. At high concentrations, other theoretical treatments are required (Minton, 1998). Quantitation of Protein Interactions
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Weight Average Sedimentation Coefficient The weight average sedimentation coefficient is defined as the mass concentration weighted average of the sedimentation coefficients of the species comprising the system, and is defined by: N
sw =
∑ ci si i =1 N
∑ ci i =1
Equation 20.7.14
Since the values of ci are uniquely determined by the equilibrium constants and total concentration, sw is a quantity completely determined by the thermodynamics of the system. The value of sw is derived from the measurement of the equivalent boundary position obtained by integrating over the boundary (Schachman, 1959; Stafford and Schuster, 1995). The observed dependence of sw on plateau concentration can be modeled to determine the stoichiometry and equilibrium constants (Timasheff et al., 1976; Na and Timasheff, 1986a,b; Lobert et al., 1995). An advantage of using sw is that its determination is completely independent of boundary shape since it is determined solely by the composition of the solution at the plateau concentration. The weight average value of s can be determined from g(s*) analysis (Stafford, 1994a,b). The g(s*) function is an apparent sedimentation coefficient distribution. It is the gradient of the concentration profile with respect to s*, which is defined by:
s* =
r ln 2 ω t rm 1
Equation 20.7.15
where rm is the radius of the meniscus and other variables are defined as for Equations 20.7.1 and 20.7.2.
Least-Squares Fitting to Solutions to the Lamm Equation
Analytical Ultracentrifugation: Sedimentation Velocity Analysis
A disadvantage of using sw is that it does not make use of the information contained in the evolution of the shape of the boundary. One way to make use of the boundary shape is to curve fit the reaction boundary with numerical solutions to the Lamm equation solved for
various reaction schemes (Todd and Haschemeyer, 1981). Various programs are available for fitting using numerical solutions to the Lamm equation. For noninteracting and self-associating systems, one can use the software Ultrascan, running under Linux, (Demeler and Saber, 1998), or SEDFIT, running under Windows (Schuck, 1998). For noninteracting, self-associating and hetero-associating systems the software ABCD_Fitter, running under Linux, DOS, and MacOS, or SEDANAL running under Windows 95/98/2000 (Stafford, 1998; Stafford and Sherwood, 2003) may be used. Analysis software packages may be obtained from the R ASMB anonymous FTP site (ftp://rasmb.bbri.org/rasmb/spin/). With the previous considerations in mind, the following section discusses the practical aspects of sedimentation velocity analysis. Many of the choices involved in the practical application of the techniques will be made with regard to the theory. Also, most of the results of sedimentation analysis are interpreted in terms of the theory, so if the experiments are not designed to reflect the requirements of the theory, the data will typically not be meaningful.
EXPERIMENTAL DESIGN AND PROTOCOLS Rotors and Cells See cells.
UNIT 7.5
for a discussion of rotors and
Sample Preparation for Sedimentation Velocity Analytical Ultracentrifugation For initial centrifugation runs, samples should be in the concentration range of 0.5 to 1.0 mg/ml for use with the Rayleigh optical system (refractive index) or in the range of 0.5 to 1.0 absorbance units (AU) at the appropriate wavelength for the absorbance optics. Preferably, samples should be of the highest purity possible. Ideally, the samples should be subjected to gel filtration just prior to sedimentation to remove aggregates. Samples also should be at “osmotic” equilibrium with their respective buffers This can be achieved either by 24-hr dialysis or by gel filtration. Either ordinary column gel filtration by Sephadex, FPLC, or HPLC can be used, or spin columns can be used (e.g., Christopherson et al., 1979). Spin columns (e.g., Bio-Spin columns from Bio-Rad) are convenient because
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there is effectively no dilution during buffer exchange. For samples prepared by dialysis and transported to another location, it is best to deliver the samples while they are still in their dialysis bags, e.g., in ∼25 to 50 ml of dialysis buffer in a conical screw-top centrifuge tube. As much of the air as possible should be removed from the dialysis bags, to minimize the potential for surface denaturation during transport. When brought to a laboratory for analysis by analytical centrifugation, samples prepared by ordinary gel filtration or spin columns must be accompanied by ∼25 to 50 ml of the same column buffer that was used to equilibrate the column. This will be used to make dilutions and as an optical reference. Since the refractive index match between the sample buffer and the reference buffer must be exact, an aliquot of buffer of nominally the same composition will not serve as an acceptable reference. Exhaustive dialysis (or its equivalent, gel filtration) has thermodynamic consequences related to the definition of macromolecular components; in order to maintain the composition of the macromolecular component (i.e., ratio of protein to counter-ions) upon dilution, the dilutions must be carried out with dialysate (Casassa and Eisenberg, 1964). Therefore, dialysis is recommended even if absorbance optics are being used. Resolution in a sedimentation velocity experiment increases as the boundary is transported toward the cell bottom; therefore, the cell should be filled as fully as possible to create the longest sedimentation path possible. In a polydisperse noninteracting sample, the separation between components is roughly proportional to the first power of time, while the contribution to boundary spreading due to diffusion is roughly proportional to the square root of time. Therefore, even though the boundary is spreading during the time of sedimentation, the resolution will still increase as the square root of time, and therefore of distance. Filling the cells With the screw ring of the assembled cell facing the investigator, the reference solution should be loaded on the left and the sample on the right side. Load 430 ml on each side to match the menisci as closely as possible. This is very important with the interference optical system, since that system measures the refractive index contribution from the protein by subtracting two relatively large numbers—the refractive index of the buffer from the refractive
index of the buffer plus protein. The protein makes a relatively small contribution to the total refractive index. If the menisci are not matched, then the contributions from the buffer, which also redistributes to a significant extent during the experiment, will not cancel at corresponding radial positions on the sample and reference sides; the result will be a changing background refractive index gradient that will make a contribution to the overall sedimentation pattern. Needless to say, this variable background contribution will complicate the analysis. Similar but much smaller contributions can be expected from the absorbance optical system if the buffer absorbs appreciably at the wavelengths of observation. It should also be noted that with interference optics, proper cancellation of the buffer contribution at corresponding radial positions requires that the axis of the cylinder lens be properly aligned perpendicularly to the radius bisecting the centerpiece. This alignment can be checked and verified by the user via a simple procedure (see below); however, the actual adjustment must be made by the manufacturer’s service technician. The author of this unit generally uses capillary-type synthetic boundary centerpieces to match the menisci exactly. Usually, the reference side is initially filled to 440 ml and the sample side to 420 ml. The rotor is accelerated to 3,000 to 10,000 rpm to allow the small amount of buffer to flow onto the sample side; when the menisci are matched, the rotor is stopped, removed, and gently shaken to remix the solution so that the run starts with a uniform initial concentration distribution. The extra step to match the menisci is especially important when working at protein concentrations below ∼0.1 mg/ml with interference optics. Cell alignment in the rotor It is extremely important that the cells be aligned properly in the centrifugal field. Because sedimentation proceeds in a radial direction, outward from the center of the rotor, the sample and reference compartments of the cells are sector-shaped to prevent material from colliding with the cell walls. If a cell is slightly misaligned, material can collect against a cell wall and build up to a concentration that will lead to unstable, convective sedimentation. Each hole in the rotor has a small line inscribed at the center of both the centripetal and centrifugal sides of the cell holes. The cell casing also has corresponding lines inscribed in the bottom. These lines must be aligned as carefully as possible to avoid convection. Use of a
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magnifying glass is recommended. One can check the alignment with the interference optics while the rotor is spinning by stepping through the delay until the image just starts to darken. The fringes should darken uniformly across the cell as it is adjusted. If the fringes at the top or the bottom start to darken before the rest of the image, then the cell is misaligned. It is highly recommended that the run be stopped and the misaligned cell readjusted before proceeding. Convection will confound the analysis and is to be avoided. Convection can be caused by scratches on the cell walls. Rotor temperature equilibration It is important that the rotor be at uniform temperature before starting the run. The Beckman Coulter XL-A/I centrifuges have two temperature-sensing mechanisms. At 100 microns of pressure the instrument switches from one to the other and there is a jump of as much as 4°C in the reported temperature. Therefore, it is important to make sure the pressure is below 100 microns during equilibration of the rotor, so it equilibrates at the expected temperature. Since the rough vacuum pump may not be able to pull the vacuum below 100 microns, the diffusion pump must be turned off to attain the full vacuum. The diffusion pump comes on only while the machine is running; therefore, the machine must be run at 0 rpm (not 3000 rpm) during the equilibration process by entering 0 rpm for the speed and pressing “START”. It may take up to an hour to equilibrate the rotor, depending on the initial temperature difference. It should be noted that the temperature of the rotor will decrease by about 0.8°C upon acceleration from 0 to 60,000 rpm, as the rotor stretches (Waugh and Yphantis, 1952); there will also be an increase in the sample temperature due to compression of the solution that will result in a net temperature difference of about 1°C between the sample and the rotor. It takes some time for this difference to dissipate, and it can lead to sample convection. The default acceleration rate of the XL is 400 rpm/sec. One might consider decreasing this value somewhat if convective disturbances of the boundaries are suspected, to allow more time for re-equilibration during acceleration.
Optical Systems Analytical Ultracentrifugation: Sedimentation Velocity Analysis
The XL series of analytical ultracentrifuges comes equipped with either an absorbance optical system (XL-A) or both absorbance and interference optical systems (XL-I). The absorbance optical system gives a profile of optical
density as a function radius and time; the interference optical system gives a profile of refractive index as a function of radius and time. The absorbance optical system allows analysis at up to three specific wavelengths but is hampered by buffer components that may absorb at the wavelengths of interest. The interference optical system has an intrinsically higher signal-tonoise ratio (by ∼5×) than the absorbance optics, and is not affected by absorbing buffer components unless they absorb at the laser wavelength. Machine setup With either optical system, it is desirable to record data as frequently as possible. For most methods of analysis the greatest signal-to-noise ratio is achieved with the largest number of scans acquired over a given time period. Absorbance optics The absorbance optical system is composed of a dual-beam, multiwavelength spectrophotometer that scans an image of the cell to produce a plot of absorbance as a function of radius. Up to three wavelengths may scanned in a single experiment. It is advantageous to collect each scan in the shortest possible time consistent with the desired point density and signal-to-noise ratio. The scan time on the XL-A is dependent on rotor speed and the radial increment: shorter scan rates can be achieved at higher speeds. The scan cycle time is typically 1 to 2 min. For typical velocity runs, the radial increment should be set to 0.03 to 0.05 and average 4 flashes of the lamp. Interference optics The scan cycle time for the interference system is on the order of 5 to 10 sec. It is necessary when using time derivative analysis (DCDT), for example, to gather as much data as possible to take advantage of its signal averaging features to the increase signal-to-noise ratio of the g(s*) plots. Most other methods of analysis will perform better if the data are sampled as frequently as possible. Adjust the laser timing so that the laser dwell is centered over the cell sectors or slits if using interference window holders. In the case of wide window holders, set the laser pulse width to 0.4° and then set the laser delay so that there is an equal angle to the edge of the sectors. For example, for a given cell, find the smallest delay angle at which the fringes just appear, then find the largest angle at which they disappear. Set
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the delay halfway between these two values. During this process, note whether the fringes disappear and reappear uniformly across the cell as these limiting delay angles are approached. If not, then the cell is not oriented correctly in the rotor hole. If the cell is misaligned, it is likely that convection of the sample will occur. The run should be stopped and the cell reoriented in the rotor with careful attention to the exact orientation of the scribe lines on the cell casing with respect to those on the rotor hole. For interference slits, the laser pulse width can be set to 1.4°. It is good idea to use interference window holders. They act to mask off a constant piece of the cell window and make the interference fringe patterns insensitive to any jitter in the laser delay timing pulses. This is especially important for proper cell blank corrections when performing equilibrium runs (see UNIT 7.5). Cylinder lens alignment check To check the cylinder lens alignment, exactly the same solution must be present in each sector. To accomplish this, a specially modified centerpiece is required. A double-sector centerpiece must be irreversibly modified by removing the rib between the two sectors (e.g., using a carefully applied coping saw). Assemble the cell and fill it with 60 mg/ml bovine serum albumin (or similarly expendable protein). Accelerate the rotor to 50,000 rpm for about an hour until the boundary is about 1/3 the way to the bottom, then decelerate the rotor to about 14,000 rpm and allow the protein to diffuse so that gradients are present throughout most of the cell. Initially, the gradients will be so steep that the light is bent completely out of the optical system; black bands will appear on either side of the boundary position and at the base of the cell. The edges of these regions represent the regions of the steepest gradient that it is possible to observe with this optical configuration. Now, if the cylinder lens is properly aligned, the fringes will be straight and level up to where they disappear, at the point of maximum gradient. If there is any significant curvature of the fringes as they enter the region of steepest gradient, the optics are not properly aligned and a trained service technician will be required to align them. Centrifuge cells The centerpieces normally used for sedimentation velocity analysis are the ordinary double-sector type. They are usually fabricated out of charcoal- or aluminum-filled epon. These centerpieces are nominally rated for
40,000 rpm; however, if both sectors are filled to the same level there will negligible distortion even up to 60,000 rpm. It is important to make sure the cells will not leak, because a leak from one sector will result in collapse of the center rib and destruction of the centerpiece. Choice of cell window material depends on the optical system. When using the absorbance optics, quartz windows are sufficient. However, when using the interference optics, sapphire windows are required. Sapphire windows are much less subject to compression and distortion than quartz at high centrifugal fields. Sapphire windows may be used also with the absorbance optical system as long as they are optically clear at the wavelengths used. The centrifugation run When the rotor temperature has equilibrated, start the run by accelerating to the desired speed; start recording data immediately so that any larger material or aggregates can be seen. If the presence of a wide range of sizes is suspected in the sample, continue the run for a while at a relatively low speed to determine if very large particles are present, then accelerate to the selected full speed for the rest of the run. Set the time between scans to “0" so that the machine will take scans as rapidly as possible. The actual time between scans will be determined by the CPU speed of the computer used to operate the XLA/I. Hard disk storage is very cheap today, especially with the available datacompression algorithms, so there is no reason not to acquire the maximum amount of data. Failure to do so may preclude certain types of analysis. Correction of sedimentation and diffusion coefficients to standard conditions It is common practice to correct raw sedimentation and diffusion coefficients to standard conditions of water at 20°C for the purposes of comparison with other experiments. Both density and viscosity corrections must be made to the sedimentation coefficient (Equation 20.7.16) while only viscosity corrections are made to the diffusion coefficient (Equation 20.7.17).
ηt0 ηtb (1 − ν2.0 ρ0 ) 0 η0 20 ηt (1 − ν2,b ρb )
s20, w = st ,b
Equation 20.7.16 Quantitation of Protein Interactions
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Experimental Design
D20, w = Dt ,b
0 ηt 0 η20
b ηt 0 ηt
Equation 20.7.17
η0t
where is the viscosity of water at the temperature, t, of the run; η020 is the viscosity of water at 20°C; ηbt is the viscosity of the buffer at the temperature of the run; ρo is the density of water at 20°C and ρb is the density of the buffer at the temperature of the run; ν2.0, and ν2,b are the partial specific volume of the macromolecule in water at 20°C and in buffer at the temperature of the run, respectively. The buoyancy correction term, (1 − ν2ρ), is used here by convention to represent the density increment (∂ρ/∂c2)T,µ.
DATA ANALYSIS AND INTERPRETATION
Analytical Ultracentrifugation: Sedimentation Velocity Analysis
The primary data are supplied as text files containing a two-line header with information about the conditions of the run followed by columns of data (i.e., radius and concentration) and sometimes a third column of standard errors of the concentration data. These concentration profile data may be analyzed by various methods to extract the hydrodynamic and thermodynamic information that the investigator desires. There are several software packages available that either transform the data into suitable visual form or that fit the profiles directly by least-squares techniques (Table 20.7.1). Before applying any of these techniques, distinctions must be made between the different types of system likely to be encountered, in order to choose the appropriate type of analysis. In general, systems can be divided into either ideal or nonideal, interacting or non-interacting, monodisperse or polydisperse, and single-component or multicomponent. An example of a single-component, monodisperse, ideal system is a globular protein near its isoelectric point at an ionic strength of 0.1, at about 1 mg/ml and not undergoing self-association. An example of a single-component, polydisperse ideal system is a monomer-dimer-tetramer self-associating system in rapidly reversible equilibrium near its isoelectric point, at an ionic strength of 0.1 at ∼1 mg/ml. An example of a two-component, polydisperse, ideal system is a weakly binding antigen-antibody system composed of four species—free antibody, free antigen, singly ligated antigen, and doubly ligated antigen—at an ionic strength of 0.1 at ∼1 mg/ml.
Starting with a sample of unknown properties, a run should be performed with three or four cells covering a wide range of loading concentrations. With interference optics, a typical protocol is to start with either 1 mg/ml or 0.3 mg/ml as the highest concentration and make 3-fold serial dilutions spanning a 27-fold range. With absorption optics, use a starting absorbance of 1.0 absorbance units. This initial run will provide information about both the polydispersity and possible concentration dependence. If concentration dependence can be ruled out at this stage, the system can be analyzed as either a monodisperse system or a polydisperse system composed of independent species. If concentration dependence is observed, it will be a combination of either association or nonideality, or both. Association is manifested as a weight-average sedimentation coefficient that increases with concentration, while nonideality is manifested as a weight average sedimentation coefficient that decreases with concentration. A nonideal, monoor polydisperse system can be analyzed as the sum of nonideal independent components. An interacting system, which is composed of a reaction boundary, must be analyzed by taking mass action into account during sedimentation. The distinction made above between ideal, nonideal, interacting, and independent species will determine which method of analysis must be employed. Use of DCDT-g(s*) or leastsquares-g(s*) analysis would be appropriate for the initial characterization and as a way of visualizing the general behavior of the system.
Data Analysis There is a wide range of software available for analysis of sedimentation velocity data. The reader is referred to Table 20.7.1 for a list of the most commonly used software packages. The g(s*) methods are model-independent and provide estimates of the range of sedimentation coefficient, number of species, their relative amounts, and degree of concentration dependence. In the absence of concentration dependence, one could use any one of the curve-fitting methods that use either approximate or numerical solutions to the Lamm equation to estimate number of components as well as their sedimentation and diffusion coefficients, relative amounts, and molecular weights. In the case of negative concentration-dependent single species—non-ideal systems—hydrodynamic nonideality and second virial coefficients can be estimated (Solovyova et al., 2001). For the
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Table 20.7.1
Software Available for Analysis of Sedimentation Velocity Dataa
Package
Author(s)
Abilities
Beckman Coulter
Various
DCDT
Stafford (1992)
DCDT+
Philo (2000)
DCDT-wd
LAMM
National Analytical Ultracentrifugation Facility (http://vm.uconn.edu/~wwwbiotc/ uaf.html) Behlke and Ristau (2002)
General package for velocity and equilibrium sedimentation analysis Time derivative g(s*) analysis for MacOS, including multi-speed wide distribution analysis Time derivative g(s*) analysis for Windows; includes fitting routines for molecular weight determination Time derivative g(s*) analysis for Windows, including multi-speed wide distribution analysis
SEDFIT
Holladay (1979); Holladay (1980)
SVEDBERG
Philo (1997)
SEDFIT, SEDPHAT
Schuck (2000)
SEDANAL
Stafford and Sherwood (2003)
ABCD_Fitter
Stafford (1998); Rivas et al. (1999)
ISODES_Fitter
Stafford (1998); Rivas et al. (1999)
ULTRASCAN
http://www.ultrascan.uthscsa.edu/
van Holde–Weischet
van Holde and Weischet (1979)
SEDNTERP
http://www.jphilo.mailway.com/ download.htm
Direct least squares fitting to approximate solutions of the Lamm equation. Useful for low-molecular-weight proteins (M < 20 kDa) and peptides. Direct least-squares fitting to concentration data of approximate solutions of the Lamm equation Direct least-squares fitting to concentration and concentration time-difference data for approximate solutions of the Lamm equation for molecular weight of noninteracting systems. Useful for low-molecular-weight proteins (M < 20 kDa) and peptides. A powerful suite of procedures for the analysis of sedimentation velocity data. Includes Lamm equation fitting, least-squares-g(s*), partial diffusion deconvolution with c(s) analysis. Least-squares fitting of concentration time difference data to numerical solutions of the Lamm equation for single- or multi-component, polydisperse interacting and noninteracting systems including self- and heteroassociating systems. Least squares fitting to concentration and concentration time difference data for numerical solutions of the Lamm equation for two component hetero-associating systems. A+B=C; C+B=D Least-squares fitting to concentration and concentration time difference data for numerical solutions of the Lamm equation for a single-component indefinite self-associating system A powerful general-purpose package for the analysis of sedimentation velocity and equilibrium data Extrapolation method for the reduction of the effects of diffusion. Useful for revealing heterogeneity in paucidisperse non-interacting systems. A general-purpose tool for the interpretation of sedimentation velocity and sedimentation equilibrium experiments. Calculates partial specific volume, hydration and other parameters from the atomic composition.
aMost of these software packages can be downloaded from the RASMB ftp site (ftp://rasmb.bbri.org/rasmb/spin/).
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analysis of interacting systems, Ultrascan, SEDFIT, SEDANAL, and ISODES_Fitter can be used for simple self-associations. For more complex interactions, ABCD_Fitter and SEDANAL can be used for analysis of multi-component systems forming heterologous complexes. SEDANAL can also deal with self-association of either reactants or products. SEDFIT is also useful for global fitting to data from dynamic light scattering used in conjunction with sedimentation data.
LITERATURE CITED Behlke, J. and Ristau, O. 2002. A new approximate whole boundary solution of the Lamm differential equation for the analysis of sedimentation velocity experiments. Biophys.Chem. 5:59-68. Cann, J.R. and Goad, W.B. 1968. Theory of transport of interacting systems of biological macromolecules. Adv. Enzymol. Relat. Areas Mol. Biol. 30:139-177. Casassa, E.F. and Eisenberg, H. 1964. Thermodynamic analysis of multicomponent solutions. Adv. Prot. Chem. 19:287-395. Christopherson, R.I., Jones, M.E., and Finch, L.R. 1979. A simple centrifuge column for desalting protein solutions. Anal. Biochem. 100:184-187. Claesson, S. and Moring-Claesson, I. 1961. Determination of the size and shape of protein molecules: Ultracentrifugation. In Laboratory Manual of Analytical Methods of Protein Chemistry (Including Polypeptides). (P. Alexander, ed.) Pergamon Press, New York. Demeler, B. and Saber, H. 1998. Determination of molecular parameters by fitting sedimentation data to finite-element solutions of the Lamm equation.Biophys. J. 74:444-454. Holladay, L. 1979. An approximate solution to the Lamm equation. Biophys. Chem. 10:187-190. Holladay, L. 1980. Simultaneous rapid estimation of sedimentation coefficient and molecular weight. Biophys. Chem. 11:303-308. Lamm, O. 1929. Die Differentialgleichung der Ultrazentrifugierung. Arkiv. Math. Astron. Fysik 21B(no. 2):1-4. Lobert, S., Frankfurter,.A., and Correia, J.J. 1995. Binding of vinblastine to phosphocellulose-purified and alpha beta-class III tubulin: The role of nucleotides and beta-tubulin isotypes. Biochemistry 34:8050-8060. Minton, A.P. 1998. Molecular Crowding: Analysis of the effects of high concentrations of inert cosolutes on biochemical equilibria and rates in terms of volume exclusion. Methods Enzymo. 295:127-149.
Analytical Ultracentrifugation: Sedimentation Velocity Analysis
Na, G.C. and Timasheff, S.N. 1986a. Interaction of vinblastine with calf brain tubulin: Multiple equilibria. Biochemistry 25:6214-6222. Na, G.C. and Timasheff, S.N. 1986b. Interaction of vinblastine with calf brain tubulin: Effects of magnesium ions. Biochemistry 25:6222-6228.
Philo, J.S. 1997. An improved function for fitting sedimentation velocity data for low-molecularweight solutes. Biophys. J. 72:435-444. Philo, J.S. 2000. A method for directly fitting the time derivative of sedimentation velocity data and an alternative algorithm for calculating sedimentation coefficient distribution functions. Anal. Biochem. 279:151-163. Rivas, G., Fernandez, J.A., and Minton, A.P. 1999. Direct observation of the self-association of dilute proteins in the presence of inert macromolecules at high concentration via tracer sedimentation equilibrium: Theory, experiment, and bioBiochemistry logical significa nce. 38:9379-9388. Rivas, G., Stafford, W.F., and Minton, A.P. 1999. Characterization of heterologous protein-protein interaction via analytical ultracentrifugation. Methods 19:194-212. Schachman, H.K. 1959. Ultracentrifugation in Biochemistry. Academic Press, New York. Schuck, P. 1998. Sedimentation analysis of noninteracting and self-associating solutes using numerical solutions to the Lamm equation.Biophys. J. 75:1503-1512. Schuck, P. 2000. Size-distribution analysis of macromolecules by sedimentation velocity ultracentrifugation and lamm equation modeling. Biophys. J. 78:1606-1619. Solovyova, A., Schuck, P., Costenaro, L., and Ebel, C. 2001. Non-ideality by sedimentation velocity of halophilic malate dehydrogenase in complex solvents. Biophys. J. 81:1868-1880. Stafford, W.F. 1992. Boundary analysis in sedimentation transport experiments: A procedure for obtaining sedimentation coefficient distributions using the time derivative of the concentration profile. Anal. Biochem. 203:295-301. Stafford, W.F. 1994a Boundary analysis in sedimentation velocity experiments. In Methods in Enzymology, Numerical Computer Methods, 240, part B (M.L. Johnson and L. Brand, eds.), pp. 478-501. Academic Press, Orlando, Fla. Stafford, W.F. 1994b. Sedimentation boundary analysis of interacting systems: Use of the apparent sedimentation coefficient distribution function. In Modern Analytical Ultracentrifugation: Acquisition and Interpretation of Data for Biological and Synthetic Polymer Systems (T.M. Schuster and T.M. Laue, eds.), pp. 119-137. Birkhäuser Boston, Boston, Mass. Stafford, W.F. 1998. Time difference sedimentation velocity analysis of rapidly reversible interacting systems: Determination of equilibrium constants by non-linear curve fitting procedures. Biophys. J. 74:A301. Stafford, W.F. and Schuster, T.M. 1995. Hydrodynamic methods. In Introduction to Biophysical Methods for Protein and Nucleic Acid Research (J. A. Glasel and M. P. Deutscher, eds.), pp. 111-145. Academic Press, San Diego.
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Stafford, W.F. and Sherwood, P. 2003. Analysis of heterologous interacting systems by sedimentation velocity: Curve-fitting algorithms for estimation of sedimentation coefficients and equilibrium constants. J. Biophys. Chem. In press.
Williams, J.W., Van Holde, K.E., Baldwin, R.L., and Fujita, H. 1958. The theory of sedimentation analysis. Chem. Rev. 58:715-806.
Tanford, C. 1961. Physical Chemistry of Macromolecules. John Wiley & Sons, New York.
Williams et al., 1958. See above.
Timasheff, S.N., Frigon, R.P., and Lee, J.C. 1976. A solution physical-chemical examination of the self-association of tubulin. Fed. Proc. 35:18861891.
Claesson and Moring-Claesson, 1961. See above.
Todd, G.P. and Haschmeyer, R.H. 1981. General solution to the inverse problem of the differential equation of the ultracentrifuge. Proc. Natl. Acad. Sci. U.S.A. 78:6739-6743. Van Holde, K.E. and Weischet, W.O. 1978. Boundary analysis of sedimentation-velocity experiments with monodisperse and paucidisperse solutes. Biopolymers 17:1387-1403. Waugh, D.F. and Yphantis, D.A. 1952. Rotor temperature measurement and control in the ultracentrifuge. Rev. Sci. Inst. 23:609-614.
KEY REFERENCES Schachman, 1959. See above. This selection of three pieces of older literature contains a tremendous amount of information that is very germane to modern day sedimentation analysis.
Contributed by Walter Stafford Analytical Ultracentrifugation Research Laboratory Boston Biomedical Research Institute Watertown, Massachusetts and Massachusetts General Hospital Harvard Medical School Boston, Massachusetts
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Spectroscopic Methods for the Determination of Protein Interactions
UNIT 20.8
Among the spectroscopic methods available in a standard biochemistry laboratory (e.g., absorption spectroscopy, fluorescence spectroscopy, circular dichroism), fluorescence spectroscopy is often the most suitable for studying the formation of protein complexes. This unit focuses on the use of fluorescence spectroscopy for the quantification of proteinprotein interactions. The methods presented are restricted to steady-state fluorescence, since time-resolved fluorescence experiments (which are used to determine the lifetimes of various species) require relatively expensive equipment and rather complicated data analysis techniques. Although studies of protein-protein interactions have been conducted using absorption spectroscopy or circular dichroism to monitor conformational changes upon interaction, fluorescence measurements offer a much broader range of possibilities and can usually be performed with only small amounts of protein. The fluorescence of a protein is characterized by its excitation and emission spectra and corresponding emission maximum (λmax ), as well as by its quantum yield (i.e., the percentage of absorbed photons that result in emitted photons) and anisotropy (i.e., the extent to which fluorescence intensity differs when measured in different directions). These parameters depend on the local environment of the fluorophore and therefore can change upon interaction of the protein with another protein or ligand. As a result, these characteristics can often be used to measure the extent of complex formation involving a given protein (Fig. 20.8.1).
Figure 20.8.1 Changes in fluorescence intensity and emission maximum wavelength (λmax ) upon complex formation. Compound S (gray line) does not contain tryptophan and thus does not contribute any fluorescence signal between 330 and 360 nm. Addition of compound S to protein A (dashed line), which contains five tryptophan residues, leads to an approximately twofold increase in fluorescence intensity and a shift in the fluorescence maximum from 335 nm to 328 nm upon complex formation (black solid line). AU, arbitrary units. Contributed by Yvonne Groemping and Nadja Hellmann Current Protocols in Protein Science (2005) 20.8.1-20.8.27 C 2005 by John Wiley & Sons, Inc. Copyright
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The source of fluorescence can be an intrinsic fluorophore, such as tryptophan or tyrosine; a covalently or noncovalently attached fluorescent dye (i.e., a label); or a fluorescent binding partner (e.g., a cofactor, such as NADH, or a nucleotide) that binds specifically to the complex under investigation. This unit provides an overview of fluorescent signals suitable for monitoring protein interactions. The protocols presented describe the determination of the optimal wavelength for a titration experiment (Support Protocol 1), the preparation of necessary controls (Support Protocol 2), and the execution of the actual titration (Basic Protocol 1). In addition, a section on the fluorescent labeling of proteins (Basic Protocol 2) and an introduction to appropriate data analysis are included (Basic Protocol 3). An example of a fluorescence titration experiment and the data analysis that follows is also presented. Many of the topics addressed in this unit can also be applied to other spectroscopic methods.
STRATEGIC PLANNING In the quantification of protein-protein interactions via spectroscopic methods, the aims are to obtain a significant signal change upon binding and to determine conditions under which the system is sufficiently stable to allow the performance of a titration experiment to evaluate binding affinity and stoichiometry. As a guideline, for best results, the signal should change by at least 30% upon binding. If the signal is unstable and decreases with time due to photobleaching or protein aggregation, it is imperative that these decreases not be misinterpreted as being indicative of binding events. The first option to examine is the use of intrinsic tryptophan (or tyrosine) fluorescence as a monitor for the interaction. If the associated change in signal is sufficiently large, then this option is preferable to external labeling, which requires additional time and may influence the binding properties of the protein. However, if the protein does not contain a suitable intrinsic fluorophore, an extrinsic label must be attached at an appropriate position within the protein. In such cases, it is advantageous to have structural information about the protein in order to determine a suitable position for the label. Extrinsic labels usually have a much higher quantum yield and therefore allow the use of much lower protein concentrations. Nonetheless, the labeling procedure can cause unwanted modification of the protein of interest, thereby affecting its activity or stability. Hence, it is crucial to verify that the activity (e.g., enzymatic activity or ligand-binding behavior) of the labeled protein is similar to that of the wild-type version.
Quantification of Protein Interactions Using spectroscopic techniques, it is possible to quantify protein-protein interactions to obtain two key parameters: the equilibrium binding constant (K; dissociation constant Kd = 1/K) and the stoichiometry value (p) for the binding partners under investigation. In this unit, the following assumption is made about the protein-protein interaction under study: Protein A can bind p copies of compound/protein S with equal affinity. For analysis of more complex binding mechanisms, see APPENDIX 5A.
Spectroscopic Methods for the Study of Protein Interactions
The following complexes can be formed upon interaction between A and S: AS, AS2 , . . . ASp . Spectroscopic methods rely on differences between the specific spectroscopic signal associated with one of the reaction partners (A or S) and the signals associated with the expected complexes (AS, AS2 , . . . ASp ). We assume further that the fluorescence of the complex is proportional to the number of molecules of S bound (as would be the case if, for example, protein A was a symmetrical dimer). Thus, the complex AS2 gives rise to twice the signal yielded by AS, and so forth. In order to obtain well-defined binding curves, the aim should be to maximize the fluorescence amplitude (F), which is the difference between the fluorescence of the complex (Fas ) and the sum
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Figure 20.8.2 Binding curve as inferred from changes in the fluorescence maximum. A simulated binding curve constructed using saturation values (solid circles) is compared with a “binding curve” generated using the observed change in λmax (open circles). Full saturation is normalized to 1. The simulated spectra on which the determination of λmax is based are shown in Figure 20.8.9. The change in λmax due to complex formation is not linear and thus cannot be used to represent the true binding curve.
of the individual fluorescence signals of protein A and compound S (Fa and Fs ); thus, F = Fas − Fa − Fs . To determine binding parameters, the change in fluorescence that results from the interaction is monitored while the concentration of one of the reaction partners is varied. The signal employed could be fluorescence intensity, anisotropy, fluorescence resonance energy transfer (FRET), or change in fluorescence intensity as a function of accessibility to a quencher molecule for the complex as compared with the unbound protein. Each type of signal is described below.) It may not be suitable to simply measure the shift in the emission λmax (open circles), since this shift is not necessarily linearly correlated with the amount of complex formed (see Fig. 20.8.2) and thus does not represent the true binding curve (solid circles).
Intrinsic Fluorescence Tryptophan is the dominant amino acid contributor to the fluorescence spectrum of a protein. The quantum yield and emission maximum wavelength (λmax ) associated with intrinsic fluorescence are very sensitive to the polarity of the environment (Hutnik et al., 1991; Xia et al., 2003). If tryptophan fluorescence is used as a signal for monitoring an interaction, an excitation wavelength that does not induce significant tyrosine fluorescence (e.g., 295 nm) should be chosen (Fig. 20.8.3). Tyrosine fluorescence is often less sensitive to changes in environment and thus produces a constant background signal upon excitation. Monitoring of emission at a wavelength close to the Raman band (see UNIT 7.7) should be avoided, especially when working with low protein concentrations. A common problem in the study of protein-protein interactions is that although both proteins contain several tryptophan residues, the fluorescence of only a few residues is affected by binding (Fig. 20.8.4). If only one of the proteins being studied contains
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Figure 20.8.3 Fluorescence emission spectra of 10 µM tryptophan and 10 µM tyrosine at excitation wavelengths (λex ) of 280 and 295 nm. AU, arbitrary units.
Spectroscopic Methods for the Study of Protein Interactions
Figure 20.8.4 Typical fluorescence titration data for a protein-protein interaction. Protein S was added in stepwise fashion to a solution of protein A (0.9 µM). The relative fluorescence signal (F[S] /F[S]=0 ) is plotted against the concentration of protein S. Binding of protein S quenches the fluorescence of protein A, which results in a decrease in signal. However, the relative change in fluorescence is only 17%, due to the presence of tryptophan residues not affected by the interaction. AU, arbitrary units.
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Figure 20.8.5 Effect of the fluorescence of compound S on binding data. Simulated data for the addition of S to a solution originally containing 15 µM protein A (Kd = 5 µM, p = 1) are shown. Fluorescence is plotted against the concentration of compound S. In the simplest case, the fluorescence signal arises only from protein A (squares). If compound S also contributes to the fluorescence signal (solid circles), a nonhyperbolic total fluorescence curve is observed (open circles). In order to correct for the contribution of S, linear regression of the data obtained in the absence of protein A is performed, and the results of the regression are subtracted from the raw binding data. Abbreviations: AU, arbitrary units; C¯ s , total concentration of compound S.
tryptophan residues, then the signal change may nonetheless be strong enough to allow determination of a binding curve. However, if both proteins contain tryptophan side chains, titration of the second component may lead to a linearly changing background (see Basic Protocol 3 and Fig. 20.8.5). If so, this background must be subtracted correctly to obtain meaningful results. Recently, the biosynthetic incorporation of irregular or unnatural amino acids, such as azatryptophan, has been reported. (Proteins can contain multiple tryptophan residues, making it difficult to assign changes in tryptophan fluorescence to a particular site. The advantage of using unnatural amino acids is that their fluorescence differs from the intrinsic fluorescence of the components used.) However, this is not a standard procedure and may not be feasible in a standard protein laboratory (Becker et al., 2003; Li et al., 2003).
Extrinsic Fluorescence Extrinsic labels (fluorescent dyes) can be attached to reactive cysteine residues or amino groups (e.g., those found in lysine residues or at the N termini of proteins). Cysteine residues are preferred for such reactions, because they are generally not very abundant in proteins (as compared with lysine), and labeling can therefore be quite specific. In some cases, excess cysteine residues may have to be removed or specific cysteine residues
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introduced using site-directed mutagenesis. It is very important to determine whether these mutations affect the stability, folding, or activity of the protein in question. If cysteine residues have to be introduced for labeling, there are two possibilities for their positioning. In one scenario, the label can be positioned at the putative interaction interface. This would maximize the probability of environmental change upon complex formation, but at the same time, the dye could also influence binding affinity or even inhibit complex formation. Alternatively, the label could be attached at a position far from the interface. This would be recommended if a conformational change was expected at the labeled site upon complex formation, as such a change would lead to a difference in fluorescence intensity. Another reason for choosing a remote position would be the expectation of a significant change in the rotational diffusion coefficient (see Background Information, “Fluorescence anisotropy”) upon complex formation, as remote labeling would allow the change in anisotropy to be monitored in such a case. A third strategy involves labeling both reaction partners and monitoring energy transfer (FRET; see below and Background Information) as an indicator of complex formation. It is important to ensure that the labeling procedure yields a homogenous population of labeled protein molecules. If, for example, some protein A molecules contain two labels, the associated Fa and Fas values might differ from the corresponding values for the singly labeled species, leading to more complicated binding curves. Since most fluorescent dyes have absorption and emission spectra in the visible region (500 to 600 nm), an unwanted contribution to the signal from the unlabeled protein component (i.e., tryptophan or tyrosine fluorescence) is unlikely, unless prosthetic groups are present. These groups, although often not fluorescent, can lead to signal changes due to their absorption properties (inner filter effect; see UNIT 7.7). Molecular Probes (http://www.probes.com) offers a broad selection of amino- and cysteine-specific labels and also provides detailed information about the labeling procedure, as well as background information on the dyes offered. Other companies, including Amersham Biosciences (CyDye fluors, convenient for labeling antibodies, proteins, and oligonucleotides), Sigma-Aldrich (Fluka fluorescent probes), Pierce (fluorescein and rhodamine labeling kits, designed for antibody labeling, but applicable to all proteins), Roche Applied Science, and PerkinElmer, to name a few, also provide fluorescent probes and easy-to-use labeling kits.
Fluorescence Resonance Energy Transfer (FRET) If the fluorescence intensity of (labeled) protein A does not change significantly upon binding to compound S, it may be possible to monitor binding using resonance energy transfer instead. This method does not depend on environmental changes around the fluorophore, but rather on the proximity of a particular pair of fluorophores (or a fluorophore and a chromophore), each attached to one of the two molecules under investigation.
Spectroscopic Methods for the Study of Protein Interactions
The spectroscopic requirement for FRET is a significant overlap between the emission spectrum of one fluorophore (the donor) and the absorption spectrum of the other (the acceptor; see Background Information). The structural requirement is that the distance between the donor and acceptor be minimal during the interaction. The efficiency of FRET depends inversely on the sixth power of the distance between donor and acceptor, providing a sensitive molecular ruler. The distance at which the energy transfer efficiency is 50% is defined as R0 (the F¨orster radius), the value of which depends on the spectroscopic properties of the pair of fluorophores used (see Background Information). A change in the extent of FRET can be measured either as a decrease in donor fluorescence or as an increase in acceptor fluorescence.
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Intensity or Anisotropy? When complex formation leads to a significant shift in the spectrum of the fluorophore or to a large change in the quantum yield, the intensity at a specific emission wavelength (associated with a particular excitation wavelength) serves as a suitable monitor for the quantification of complex formation. The excitation and emission wavelengths should be chosen to maximize the change in signal (see Support Protocol 1). However, if the spectral properties do not change significantly upon complex formation, it may be possible to use anisotropy to monitor the interaction (Bai et al., 1997; LeTilly and Royer, 1993). In this case, the excitation and emission wavelengths should be chosen to minimize the change in signal intensity upon complex formation, because the results have to be corrected for any such changes (see Background Information). One disadvantage associated with the use of anisotropy is that the signal-to-noise ratio is usually rather small. Fluorescence Quenching Upon binding of a fluorescent protein to another protein or ligand, a decrease in fluorescence intensity due to direct quenching of the fluorophore by the bound protein/ligand can occur (Kelly and von Hippel, 1976). In a standard titration experiment, this quenching would be indistinguishable from a decrease caused by a change in the polarity of the environment. The accessibility of the binding site to external quenchers (e.g., acrylamide) or dyes [e.g., anilinonaphthalenesulfonate (ANS)] can be used as an alternative signal in cases in which fluorescence intensity and anisotropy do not change significantly upon complex formation (UNIT 7.7). Specifically, titration of protein A with compound S can be performed in the presence of such a quencher or dye. One example of this approach involves the binding of ε-ADP to DNA B helicase (Jezewska and Bujalowski, 1997). FLUORESCENCE TITRATION TO MONITOR COMPLEX FORMATION AND DETERMINE BINDING CONSTANTS
BASIC PROTOCOL 1
This protocol describes how to perform a titration experiment using any of the signals mentioned in Strategic Planning—namely, fluorescence intensity of a single label, fluorescence intensity due to FRET (also see Background Information), or fluorescence intensity as influenced by the accessibility of the fluorophore to a quencher molecule. In this third case, the starting solution consists of protein A in the presence of a suitable amount of quencher. The same protocol applies if a change in anisotropy is employed to monitor complex formation (in which case a polarization device has to be set accordingly; see UNIT 7.7) or if the fluorescence of a third ligand or cofactor that binds specifically to the complex is chosen as the signal to be measured (also see Background Information).
Choice of concentrations It is essential that the concentrations of the proteins in both solutions be known and that no inactive protein fractions be present. In general, with regard to the protein solution in the reaction cuvette, use of the lowest concentration possible is recommended. Doing so minimizes possible inner filter effects, the amount of material used, and possible problems due to reduced solubility. In addition, if the concentration of the protein in the cuvette is significantly lower than the Kd for binding and the stoichiometry of the interaction is already known, data analysis becomes much simpler (see Basic Protocol 3). However, if the concentration is too low, protein content may be lost through binding to the cuvette walls (see Support Protocol 2). If stoichiometry and affinity values are to be determined from a single titration experiment, the concentration of protein A should be 1 to 3 times the expected Kd .
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Assuming that no information is known about the affinity of the interacting molecules, start with the following: Use a concentration of ∼0.5 to 1 µM in a volume of 1 ml for protein A, and add small volumes (1 to 5 µl) of a stock solution (∼100 µM) of compound S until the signal reaches saturation. An example of such a titration and a description of the subsequent data analysis are given in Basic Protocol 4. For the determination of the equilibrium constant, it is essential to obtain sufficient data points in the region between 10% and 90% complex formation. It is advisable to reach nearly complete saturation so that any deviations from the simplest (and therefore anticipated) binding behavior can be detected and the final fluorescence amplitude can be accurately determined. Thus, the titration increments should initially be relatively small and become larger toward the end of the titration. When preparing the stock solution of component S, two criteria should be taken into consideration. The volume added should be minimized (to minimize dilution of protein component A), but it should be large enough to allow reproducible results. Reproducibility should be confirmed by titrating a corresponding volume of a simple fluorescent compound (e.g., tryptophan). It should also be ensured that the buffer solution has the same viscosity (e.g., the same concentration of glycerol or saccharose, if present) as the titrant. Another option for keeping the concentration of protein A constant is to prepare solution S with the inclusion of protein A at the same concentration as is present in the cuvette. (Since solution S is usually quite concentrated, the fraction of S that is bound to A in this solution can be neglected.) This is a useful (although not very widely used) approach. As an example, in a titration of 1 µM protein A with 5-µl aliquots of a 100 µM solution of compound S, 1 µM protein A would be added to the stock solution of compound S. Then, regardless of the volume of stock solution added to the cuvette, the concentration of protein A in the cuvette would remain constant at 1 µM.
Materials Filtered and degassed buffer solution for diluting protein samples (see UNIT 7.7) Protein “A” and compound “S” solutions, purified and of known concentration Calibrated fluorescence spectrometer (see UNIT 7.7) Cleaned cuvettes (see UNIT 7.7) Stirring devices NOTE: Always make sure that the solution has reached the required temperature before starting the measurement. Extra time must be allowed for temperature equilibration if the protein solution has been on ice before the experiment, as fluorescence is strongly temperature dependent. Samples in plastic cuvettes tend to require more time to reach their final temperature than do samples in glass or quartz cuvettes. NOTE: All of the experiments mentioned in this unit are equilibrium experiments. Thus, it is very important to ensure that samples are at equilibrium (i.e., that the fluorescence reaches a constant value) after every titration step. Depending on the speed of complex formation at the concentrations chosen, it may be necessary to wait for some time (i.e., several minutes) before recording data.
Prepare spectrometer 1. Switch on the fluorescence spectrometer and allow it to warm up (see UNIT 7.7). 2. Set excitation and emission wavelengths (Support Protocol 1). Spectroscopic Methods for the Study of Protein Interactions
If photobleaching occurs, the shutters should be closed during equilibration (see Support Protocol 2).
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Perform titration 3. Fill cuvette with buffer solution and wait for solution to equilibrate at the desired experimental temperature. Add protein A, stir the sample if possible (see Support Protocol 2), and monitor the fluorescence signal (see UNIT 7.7). In some fluorescence spectrometers, a magnetic stirring device is already mounted below the cuvette holder. Alternatively, samples can be stirred manually using a small plastic stirrer. If the protein is sensitive to stirring, mixing can be achieved by gently pipetting the solution up and down or by applying a lid to the cuvette and inverting once or twice.
4. Once the fluorescence signal has stabilized, add stepwise aliquots of the stock solution of compound S. Stir solution continuously if possible (see Support Protocol 2). Measure an emission spectrum or acquire a time trace at a single wavelength (preferably for at least ∼50 sec, so the stability of the signal can be monitored). If a spectrum is obtained after each addition, select a wavelength and read the fluorescence value at this wavelength each time; plot the resulting fluorescence values against the concentration of compound S. This technique permits the investigator to judge whether the titration contains a sufficient number of data points. Saturation can also be estimated from an overlay of the spectra obtained. If fluorescence is recorded at a single wavelength, the titration may be monitored continuously (time trace) to measure the signal change throughout the experiment. In this case, the display would resemble the final binding curve. This kind of experiment requires a fluorimeter, in which the shutters close automatically upon opening of the lid. If a continuous time trace is not obtained, a spectrum must be recorded after each titration step, and the fluorescence intensities recorded at the emission wavelength must be plotted against the concentration of compound S.
5. Plot the observed equilibrium fluorescence against the concentration of compound S. Check the plot visually for unusual behavior in the final part of the binding curve (at levels near saturation). If there are any indications of processes such as photobleaching or damage to the protein, perform the control experiments described in Support Protocol 2. If photobleaching and other artifacts can be ruled out, the presence of multiple distinctive slopes in the plot may be an indication of nonspecific binding.
6. Analyze data using an appropriate model (see Basic Protocol 3).
INTRODUCTION OF A FLUORESCENT LABEL INTO A PROTEIN AND DETERMINATION OF THE DEGREE OF LABELING
BASIC PROTOCOL 2
Commonly used fluorescent labels include fluorescein, dansyl, rhodamine, and NBD [N,N -dimethyl-N-(acetyl)-N -(7-nitrobenz-2-oxa-1,3-diazol-4-yl)]. Molecular Probes offers a selection of new chromophores, including the Alexa dyes (coumarin derivatives), BODIPY, and others, that have very high extinction coefficients and quantum yields. Compared with “older” labels such as fluorescein, these newer compounds are quite photostable and much less sensitive to pH (with activity over a pH range of 4 to 10), and they are available for a large range of excitation and emission wavelengths. A disadvantage, however, is that these novel fluorescent probes contain very complex and bulky aromatic groups, which could disturb the interaction of the labeled protein with its ligand or potentially lead to nonspecific binding due to hydrophobic interactions. It should also be borne in mind that some of these newer compounds are less sensitive to changes in their environment compared with more traditional labeling compounds (e.g., fluorescein). Quantitation of Protein Interactions
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All of the fluorescent labels mentioned can be attached to cysteine residues (as is typically done with maleimides or iodoacetamides) or amino groups (as is typically done with succinimidyl esters) on the protein of interest. One easy way to label a protein is to use a commercially available labeling kit. These kits usually contain a premeasured amount of dye, so that only a certain amount of pure protein has to be added to the dye vial. They also include the buffer solutions and columns needed to purify the modified protein (typically gravity-feed size-exclusion columns or prepacked desalting columns). The following protocol explains how to label a protein with a fluorescent dye without using such a kit. The protocol can also be modified according to the instructions of the label’s supplier (e.g., to incorporate optimal pH and buffer conditions and optimal temperature and time for the labeling reaction).
Materials Protein, purified and of known concentration (5 to 20 mg/ml), to be labeled Reaction buffer (as recommended by supplier of labeling compound) 10 mg/ml fluorescent dye in DMSO or DMF: Use succinimidyl esters (or sulfonyl chlorides) for the modification of amine groups, and use maleimides (or iodoacetamide) for the modification of thiol groups (in cysteine residues) Filtered and degassed buffer solution (see UNIT 7.7) Dialysis membranes or Slide-A-Lyzer dialysis kits (Pierce) PD-10 or NAP-5 columns (Amersham Biosciences) Cuvettes (see UNIT 7.7) UV-vis spectrometer Additional reagents and equipment for dialysis (APPENDIX 3B) or buffer exchange using a desalting column (UNIT 8.3) CAUTION: Dimethylsulfoxide (DMSO) and dimethylformamide (DMF) should be handled with caution, as they are known to facilitate the entry of organic molecules into tissues. It is strongly recommended that double gloves be used when handling these solvents. NOTE: The amount of protein used for labeling depends mainly on the availability of the protein. Typically, 5 to 10 mg is used. NOTE: Some fluorescent dyes are very photosensitive; thus, the labeling procedure should be allowed to take place under minimal light. Ideally, samples should always be kept in the dark or wrapped in aluminum foil and, if possible, kept on ice during and between experiments.
Label ε-amino or thiol groups on target protein 1. Dialyze the protein to be labeled (APPENDIX 3B) using the recommended reaction buffer (e.g., 0.1 M sodium bicarbonate, pH 8.0), or else change buffer using an NAP desalting column (UNIT 8.3). Primary amine buffer compounds, such as Tris, should be avoided, especially when using amine-reactive dyes, as amine-containing buffer molecules will compete with the amino groups on the protein. Spectroscopic Methods for the Study of Protein Interactions
The pH of the labeling reaction should be chosen with care. For amine-reactive dyes, a basic pH (8 to 9) should be used if the ε-amino groups of lysine residues are to be labeled, whereas a lower pH (7 to 7.5) should be chosen if the protein is to be specifically labeled at the N terminus. Furthermore, the pH of the buffer solution should not be too close to (i.e., should differ by > 0.5 pH units from) the isoelectric point (pI) of the target protein.
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If thiol groups (on cysteine residues) are to be labeled, they must first be reduced. This is usually achieved by adding reducing reagents such as DTT or mercaptoethanol. Unfortunately, these reducing agents tend to react with the dye and thus compete with the labeling reaction; therefore, they should be avoided if possible. A suitable alternative is TCEP [Tris-(2-carboxyethyl)phosphine], which is much more stable than DTT and does not contain thiol groups. Nevertheless, TCEP may also react with certain labeling reagents, such as maleimides; thus, it is useful to check the supplier’s protocol for reaction buffer recommendations prior to labeling. In many cases, the protein can be reduced with DTT or TCEP and then rapidly desalted (e.g., using a small PD10 column) immediately before labeling.
2. Add 100 µl dye solution (1 mg) to 1 ml protein solution in a small vessel (e.g., a 2to 5-ml plastic tube). Stir continuously for 1 hr using a magnetic stirrer or by gently rocking the tube. Depending on the amount of protein used, the incubation time may have to be adjusted. If the protein is not stable at room temperature, the incubation can be performed at 4◦ C and allowed to continue for a longer period of time.
3. Remove free dye by applying the reaction solution to an equilibrated PD-10 or NAP-5 column and eluting with the appropriate buffer. Collect eluate. Ideally, a buffer similar to the one used in fluorescence titration will be used for elution. Free dye molecules can be separated from the conjugated protein using these desalting columns. (Typically, 0.5- to 1-ml columns are used.) Note, however, that this procedure does not separate labeled protein from unlabeled protein. Depending on the specific protein, samples can usually be stored in a refrigerator for 1 to 2 days between experiments. Samples should be frozen in liquid nitrogen and kept at −20◦ C for medium-length storage (up to 2 weeks) or at −70◦ C for long-term storage. Light-sensitive samples should always be wrapped in aluminum foil.
Determine yield of labeled protein 4. Using a UV-vis absorption spectrophotometer (UNIT 7.2), measure the absorbance of the solution at 280 nm and at the wavelength at which the dye exhibits dominant absorbance (denoted as λ). The concentrations of labeled and unlabeled protein can be determined as long as the label and the unlabeled protein exhibit distinct absorption spectra. Typically, the protein will have an aromatic absorbance around 280 nm. The total absorbance should be between 0.1 and 1.0 to ensure accurate readings; dilute protein solution with storage or desalting buffer if necessary to achieve this condition.
5. Calculate the concentrations of dye and protein in the solution. Concentrations of dye and protein are calculated differently depending on the spectral dye dye characteristics of these compounds. In the following equations, ε280 and ελ are the extinction coefficients of the dye at 280 nm and λ (the wavelength at which the dye exhibits p p dominant absorbance), respectively, and ε280 and ελ are the extinction coefficients of the protein at 280 nm and λ, respectively. Case 1: No spectral overlap:
Equation 20.8.1
Equation 20.8.2
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Case 2: Dye absorbs at 280 nm, but protein does not absorb at λdye :
Equation 20.8.1
Equation 20.8.3
Case 3: Dye absorbs at 280 nm, and protein absorbs at λdye :
Equation 20.8.4
Equation 20.8.3
All calculations assume an optical path length of 1 cm. The ratio of dye concentration to protein concentration equals the yield of labeled protein. Example: The following absorbances were obtained from a protein labeling reaction mixture (path length, 1 cm): OD280 = 2.0 and OD400 = 2.9. The extinction coefficients for the protein were 0.039 µM−1 cm−1 (280 nm) and 0.005 µM−1 cm−1 (400 nm), and the corresponding extinction coefficients for the dye were 0.010 µM−1 cm−1 and 0.090 µM−1 cm−1 , respectively. Calculation according to Case 3 above yields the following:
Equation 20.8.5
Thus, the yield of labeled protein is 30 µM/44 µM = 68%.
Spectroscopic Methods for the Study of Protein Interactions
Typical yields for the labeling of proteins vary considerably, from 30% to 100%, depending on the protein and labeling procedure used. Hence, samples can still contain unconjugated protein after the labeling reaction is run. Although this results in a decrease in the total change in fluorescence intensity upon binding, this decrease should not affect the standard titration experiments described here.
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ANALYSIS OF TITRATION EXPERIMENTS: DETERMINATION OF BINDING AFFINITY AND STOICHIOMETRY
BASIC PROTOCOL 3
This protocol describes corrections that may be necessary after a binding curve has been obtained and explains how to determine the binding affinity (K) and stoichiometry value (p) using the data yielded by a titration experiment.
Correct data The raw data (denoted Fobs ) obtained in the titration experiment (Basic Protocol 1) may deviate from an ideal binding curve for several reasons, including the following: (1) the contribution of compound S to the fluorescence signal (denoted Fadd ); (2) the dilution of protein A upon titration with compound S; and (3) bleaching or stirring artifacts. (Support Protocol 1 describes how to test for such artifacts.) Dilution effects can be avoided as described in Basic Protocol 1 or corrected for as described in this protocol. If corrections for both dilution effects and the contribution of compound S to the total fluorescence signal have to be made, then the latter correction must be made first. If compound S contributes to the fluorescence signal: If the added compound S contributes to the fluorescence signal, a fraction of the observed fluorescence will change linearly with the concentration of S. An example is shown in Figure 20.8.5. There are two possible ways to account for this: Either subtract the contribution prior to data analysis or include the contribution of S in the analysis (see Analyze Data below for a description of the latter option). Correction of data requires that the contribution of compound S be measured accurately. If the contribution of compound S is included in the data analysis, the additional measurements described below are not necessary; this may be advantageous if the amount of compound S available is limited. The first step in the correction process is to perform a titration as in Basic Protocol 1, but with protein A absent from the sample buffer in the cuvette. The recorded fluorescence signal should change linearly with the total concentration of S (C¯ s ; see Fig. 20.8.5, black circles). Next, using the fluorescence data obtained, perform a linear regression to determine the fractional fluorescence intensity of compound S (Fs ) Fadd = Fs C¯ s Once Fs has been determined from this equation, a corrected fluorescence reading (Fcorr ) for each data point in the titration curve from Basic Protocol 1 can be obtained using the following equation: Fcorr = Fobs − Fs C¯ s In Figure 20.8.5, linear regression analysis of the fluorescence data obtained by titrating with compound S in the absence of protein A yields a slope of Fs = 2 (black circles). Using this slope, the contribution of compound S to the signal observed in Basic Protocol 1 can be calculated (second column, Table 20.8.1). This contribution (denoted Fadd ) is then subtracted from Fobs (Fig. 20.8.5, open circles) to yield the corrected binding curve (Fig. 20.8.5, open squares).
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Table 20.8.1 Corrections for the Contribution of Compound S to the Total Fluorescence Signala,b
C¯ s
Fadd = 2C¯ s
Fobs
Fcorr = Fobs − Fadd
0
0
0
0
3
6
224
218
6
12
419
407
9
18
599
581
17
33
925
892
26
51
1101
1050
35
69
1242
1173
43
87
1317
1230
52
105
1360
1255
61
123
1427
1304
70
141
1495
1354
83
165
1513
1348
98
195
1640
1445
113
225
1641
1416
128
255
1724
1469
a C¯ , total concentration of compound S; F , contribution of compound S to observed fluorescence s add
signal; Fobs , total observed fluorescence signal; Fcorr , corrected fluorescence signal. b All fluorescence values in arbitrary units.
If dilution effects need to be corrected for: As a rule, if the solution containing protein A is diluted by more than 5% during titration, the following correction should be performed:
Equation 20.8.6
Here, Fobs is the fluorescence observed prior to correction for dilution effects (and after correction for the contribution of compound S, if necessary), V0 is the total volume of protein A solution at the beginning of the experiment, and dV is the total volume of compound S solution added during titration. Examples of data prior to and after correction for dilution effects are shown in Figure 20.8.6.
Analyze data
Spectroscopic Methods for the Study of Protein Interactions
The data analyses presented in APPENDIX 5A are based on the transformation of data into a form in which the concentration of the complex is plotted against the free ligand concentration. This requires knowledge of the stoichiometry value (p, which can be determined via stoichiometric titration, for example; see Fig. A.5A.2) prior to data analysis. For the most simple binding mechanism (p identical binding sites), such a transformation yields a hyperbolic binding curve.
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Figure 20.8.6 Quadratic binding curve and dilution effects. Typical experimental data were simulated for a case in which analysis based on a quadratic function was warranted. Black circles represent raw data, and triangles represent the data after correction for dilution. Curves correspond to the fits as discussed in the text. AU, arbitrary units.
If the stoichiometry cannot be determined separately, it is still possible to obtain at least the binding constant (K) using the experimental data, and under specific conditions, p can be determined as well. Here, two cases must be distinguished from one another:
Hyperbolic binding curve At concentrations of protein A that are much smaller than Kd , the concentration of complex present after each addition of compound S is always much smaller than the total concentration of S (see example in Basic Protocol 4). If [A] < 0.1 Kd , then at halfsaturation, the total and free concentrations of compound S differ by <5%. Thus, the concentration of free compound S (Cs ) can safely be approximated as the total concentration of S (C¯ s ). In this case, no information about stoichiometry can be obtained, but the binding constant can be determined. The corresponding binding curve is hyperbolic and can be described by the following equation:
Equation 20.8.7
where Fa is the fluorescence of the free protein A, Fas is the fluorescence of the complex AS, C¯ a is the total concentration of protein A, and F = Fas − Fa . The data in these curves can either be linearized (see APPENDIX 5A) or, preferably, analyzed with nonlinear fitting routines to determine Kd and pFC¯ a .
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Quadratic (or square root) binding curve If the concentration of protein A is ≥0.2 Kd , the concentration of free compound S present at each titration step is reduced significantly upon complex formation. In this case, mass balance must be used to obtain the relevant function for the concentration of complex formed:
Equation 20.8.8
This yields a quadratic equation for Cas , which denotes the concentration of occupied binding sites on protein A: 2 − (C ¯ s + Kd + pC¯ a )Cas + pC¯ a C¯ s = 0 Cas
The value of Cas that satisfies this quadratic equation is a square root function involving C¯ s and C¯ a (the total concentrations of compound S and protein A, respectively):
Equation 20.8.9
where g = C¯ s + Kd + pC¯ a and h = 4pC¯ a C¯ s Thus, the total fluorescence signal observed is given by:
Equation 20.8.10
where F = Fas − Fa . The determination of p via this method, although less exact than direct measurement in a stoichiometric experiment, might sometimes be the only possibility (e.g., when the reactants have limited solubility). In order to obtain a reliable estimate of the stoichiometry value p, the concentration of protein A should be 1× to 3× Kd (estimated value).
Inclusion of signal arising from compound S If compound S, which is used to titrate the fluorescent protein A, also contributes to the fluorescence signal, this contribution has to be taken into account. This can be done either by performing a separate titration of compound S in the absence of protein A (see above) or by explicitly including the contribution of compound S in the data analysis. In the latter case, the observed fluorescence is given by: F = C¯ a Fa + FCas + C¯ s Fs where F = Fas − Fa − Fs . Then, the parameters in the equation (either quadratic or hyperbolic) describing Cas , as well as the slope Fs , are to be optimized with regard to fitting of the experimental data. Spectroscopic Methods for the Study of Protein Interactions
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Note that when a nonfluorescent compound S binds nonspecifically to protein A and alters the fluorescence of the latter, the binding curve that results can resemble the curve that arises when fluorescent compound S contributes directly to the observed signal. However, if compound S is not fluorescent, then a titration involving compound S in the absence of protein A will yield no significant fluorescence signal. For the analysis of data in the event of nonspecific binding, refer to APPENDIX 5A.
EXAMPLE: STANDARD PROTEIN-PROTEIN TITRATION WITH DATA ANALYSIS As an example, the results of a hypothetical titration of 1 µM protein A with a 100 µM stock solution of compound S are presented in Figure 20.8.6. A 1-ml volume of buffer solution containing 1 µM protein A, which carries the fluorophore, is dispensed into the fluorimeter cell, and the fluorescence signal is recorded at a constant wavelength. Aliquots of the compound S solution (Table 20.8.2) are subsequently added to the protein A solution in the cell, with the fluorescence signal being monitored at this same wavelength after each addition. The binding curve corresponding to this sample experiment is depicted in Figure 20.8.6. A first estimation of Kd can be obtained as follows: 50% of protein A is complexed with compound S at a total concentration of ∼1 µM compound S. At this point on the curve, the concentration of free S is about 0.5 µM (assuming that p = 1), and therefore, Kd is estimated to be approximately 0.5 µM as well. This estimate suggests that the binding data obtained should be analyzed using a quadratic model, since the concentration of free S differs from the total concentration of S. Note that this difference would be even larger if the stoichiometry value p was greater than 1. Analysis of the data shown in Figure 20.8.6 using a quadratic model (Eq. 20.8.10) yields a Kd of 0.51 µM and a stoichiometry value (p) of 0.96.
Table 20.8.2 Sample Data from a Typical Protein-Protein Fluorescence Titrationa,b
Point no.
dV (µl)
[S] (µM)
[A] (µM)
Fobs
Fcorr = Fobs × (1000 + dV)/1000
0
0
0.0
1.000
4000
4000
1
3
0.3
0.997
4215
4228
2
3
0.6
0.994
4383
4409
3
3
0.9
0.991
4521
4562
4
3
1.2
0.988
4616
4671
5
3
1.5
0.985
4688
4759
6
3
1.8
0.982
4739
4824
7
5
2.2
0.978
4791
4901
8
5
2.7
0.973
4819
4954
9
10
3.7
0.963
4837
5020
10
10
4.6
0.954
4827
5059
11
20
6.4
0.936
4776
5101
12
20
8.1
0.919
4711
5126
a dV, volume of 100 µM compound S solution added; F, fluorescence signal; F , total observed fluorescence obs
signal; Fcorr , corrected fluorescence signal. b All fluorescence values in arbitrary units.
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Figure 20.8.7 Hyperbolic binding curve and dilution effects. Typical experimental data were simulated for a case in which analysis based on a hyperbolic function was warranted ([A] < 0.1Kd ). Black circles represent raw data, and triangles represent the data after correction for dilution. AU, arbitrary units.
Figure 20.8.7 shows data from a simulated experiment in which 0.01 µM protein A is titrated with 100 µM compound S under otherwise identical conditions. Here, when the total concentration of compound S is ∼0.5 µM, about 50% of protein A is complexed with S. Thus, the concentration of free S at this point is 0.5 µM − (50% × 0.01 µM) = 0.495 µM, assuming that p = 1. In this case, the concentration of free S can safely be approximated as the total concentration of S, and the binding curve should be analyzed using the hyperbolic model (Eq. 20.8.7).
SUPPORT PROTOCOL 1
DETERMINATION OF OPTIMAL EMISSION AND EXCITATION WAVELENGTHS FOR FLUORESCENCE TITRATION EXPERIMENTS In order to obtain a suitable binding curve for the quantification of a protein-ligand interaction, a sufficiently large signal change upon complex formation is required. The approximate excitation and emission maxima for the protein of interest are usually known. (If an extrinsic label is used, this information should be available from the supplier of the label.) The strategy presented here applies to the use of changes in fluorescence intensity as a monitor for complex formation.
Materials Filtered and degassed buffer solution for diluting protein samples (see UNIT 7.7) Protein “A” and compound “S” solutions, purified and of known concentration Calibrated fluorescence spectrometer (see UNIT 7.7) Cleaned cuvettes (see UNIT 7.7) Spectroscopic Methods for the Study of Protein Interactions
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1. Switch on the fluorescence spectrometer and allow it to warm up (UNIT 7.7). 2. Set the excitation wavelength as suggested by the dye manufacturer, or use 280 or 295 nm if tryptophan fluorescence is to be measured. Fill cuvette with buffer solution, add protein A, stir the sample if possible, and obtain an emission spectrum. Make certain that the buffer solution has equilibrated to the required temperature before performing this step.
3. Identify the wavelength corresponding to maximum emission in the emission spectrum obtained in step 2. Set the emission wavelength of the fluorescence spectrometer to this value, and acquire an excitation spectrum for protein A alone. The excitation spectrum must be corrected for the lamp spectrum (as the lamp monochromator intensity varies as a function of wavelength). Lamp spectra for such corrections are generally obtained using rhodamine B. Usually, lamp spectrum corrections are made by the supplier of the instrument prior to shipping, and the supplier also typically provides rhodamine B as a calibration standard for the investigator.
4. Repeat steps 2 and 3 using a solution of protein A saturated with compound S. The concentration of compound S necessary for saturation must be determined experimentally.
5. Repeat steps 2 and 3 for a solution containing only compound S at the saturating concentration, and also for buffer alone. 6. Choose the pair of excitation and emission wavelengths that yields the maximum difference between fluorescence signal background (i.e., signal attributable to buffer plus signal attributable to protein A and compound S individually) and signal attributable to the complex AS.
DETERMINATION OF THE STABILITY AND PROPENSITY FOR PHOTOBLEACHING OF A FLUORESCENT PROTEIN
SUPPORT PROTOCOL 2
In a typical titration experiment, compound S is titrated into a solution of protein A. Thus, because the same protein A sample is used throughout the course of the experiment, it is important to make sure that extended exposure to light and/or mechanical stress due to stirring do not induce alterations in the protein or protein-ligand complex.
Materials Filtered and degassed buffer solution for diluting protein samples (see UNIT 7.7) Protein “A” and compound “S” solutions, purified and of known concentration Calibrated fluorescence spectrometer (see UNIT 7.7) Cleaned cuvettes (see UNIT 7.7) NOTE: In the protocol that follows, it is assumed that protein A contains the fluorescent label.
Prepare spectrometer 1. Switch on the fluorescence spectrometer and allow it to warm up (UNIT 7.7). 2. Set the excitation and emission wavelengths as determined using Support Protocol 1. Quantitation of Protein Interactions
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Perform control assessments of protein A 3. Check the stability of the protein A solution by filling the cuvette with this solution under the same conditions as in the titration experiment (see Basic Protocol 1, step 3) and observing the fluorescence signal continuously for a period of time corresponding to the duration of the titration experiment (at least 30 min). Make certain that the solution has equilibrated to the required temperature before making fluorescence readings. A decrease in the fluorescence signal over time could indicate photobleaching, loss of protein content due to attachment of protein molecules to the cuvette walls, or mechanical damage to the protein as a result of stirring.
4. Check for photobleaching by filling the cuvette with a fresh aliquot of protein A solution and observing the fluorescence signal at intermittent time points (by briefly opening and then closing the shutter on the fluorimeter) over a period of time corresponding to the duration of the titration experiment. In total, ∼10 to 20 fluorescence signal readings should be obtained in this step. If the fluorescence signal decreases to the same extent as in step 3, then photobleaching can be ruled out. Photobleaching can be reduced by decreasing the excitation bandwidth or changing the excitation wavelength to one at which the intensity of the lamp is lower. The absolute signal will be reduced by these procedures, but such reductions can be partially compensated for by increasing the emission slit width (particularly when using a monochromator).
5. Check for mechanical damage and loss of protein content due to protein–cuvette wall interactions by repeating step 3 using a new aliquot of protein A solution. If the fluorescence signal decreases over time, compare the rate of decrease with and without stirring the solution. Photobleaching can sometimes be distinguished from mechanical damage/loss of protein content by comparing the fluorescence decreases observed with and without stirring. Switching off the stirrer should make the decrease in fluorescence less severe in the case of mechanical damage/loss of protein content due to protein–cuvette wall interactions, whereas it should make the decrease in fluorescence more severe in the case of photobleaching.
6. Repeat steps 3 through 5 using different concentrations of protein A solution. Mechanical damage usually results in a concentration-independent change in fluorescence, whereas the fraction of protein content lost to protein–cuvette wall interactions generally decreases with increasing protein concentration. If it is found that protein binding to cuvette walls is causing appreciable signal instability, it might be necessary to change buffer conditions, coat the cuvette walls with detergent (e.g., 0.05% Tween 20), or use higher concentrations of protein A (so that the relative loss of protein content becomes smaller). Another option is to add 5% to 10% (v/v) glycerol to the protein dilution buffer (step 3) to stabilize the protein. If detergents are used, their fluorescence profiles should be checked beforehand.
Perform control assessments of the AS complex 7. Perform steps 3 to 6 using samples that contain the protein complex AS. The concentration of complex should ideally be close to half-saturation to produce maximal effects due to changes in fluorescence resulting from adsorption, bleaching or degradation of A, S or AS. Spectroscopic Methods for the Study of Protein Interactions
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COMMENTARY Background Information For a detailed introduction to tryptophan and tyrosine fluorescence spectroscopy, including quenching and fluorescence anisotropy, see UNIT 7.7. This section provides a discussion on the use of extrinsic probes. Changing the environment of a probe can affect the intensity of the fluorescence signal, the spectral shape, or both. For most labels used in protein interaction assays, a blue shift in the emission maximum occurs when the label is transferred from a
hydrophilic to a more hydrophobic environment. In general, the spectroscopic probe should be located close to the region where the interaction partner is expected to bind (see Fig. 20.8.8). In such cases, the spectral change observed upon addition of compound S could look like the one depicted in Figure 20.8.9. Note the appearance of an isosbestic point, where the same fluorescence intensity is seen in both spectra. When spectra are overlaid
Figure 20.8.8 Schematic representation of a protein-protein interaction. Protein S binds to protein A, which is fluorescently labeled. Upon binding, the spectral properties and intensities of the emitted fluorescence change.
Figure 20.8.9 Simulated fluorescence spectra as might be observed in a titration experiment (fluorescence in arbitrary units, AU). Note the isosbestic point, where both spectra show the same fluorescence intensity. The existence of an isosbestic point indicates that only two interconverting fluorescent species (protein and complex) are present.
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Figure 20.8.10 Spectral requirements for fluorescence resonance energy transfer (FRET). Schematic excitation (dotted) and emission (solid) spectra for a donor species (gray) and an acceptor species (black) are shown. The region of spectral overlap is shaded. AU, arbitrary units.
with one another, the appearance of an isosbestic point indicates that only two fluorescent species exist in the system being studied. If binding of the partner S induces a conformational change in protein A, a fluorescent position far from the binding site might also be affected, and the resulting change in signal could be used to monitor complex formation (Hiratsuka, 1998). Another option for avoiding labeling near the protein-ligand interface is to employ FRET (UNIT 19.5). Fluorescence resonance energy transfer (FRET) Fluorescence resonance energy transfer describes the nonradiative transfer of excitedstate energy from a donor (D) to an acceptor (A). This process occurs as the result of a dipole-dipole coupling between the donor and the acceptor. The efficiency of energy transfer (E) is given by:
Equation 20.8.11 Spectroscopic Methods for the Study of Protein Interactions
Here, R0 , the so-called F¨orster radius, is characteristic of the specific donor-acceptor pair, and r is the actual distance between the
acceptor fluorophore on protein A and the donor fluorophore on compound S. At r = R0 , an energy transfer efficiency of 50% is obtained. The value of R0 depends on the overlap of the emission spectrum of the donor with the excitation spectrum of the acceptor (see Fig. 20.8.10) and on the relative orientation (κ) of the transition dipole moments of the donor and the acceptor. The latter feature is often not known or not constant during the lifetime of the donor, and therefore, it is usually approximated by κ2 = 2/3, which corresponds to a situation in which the orientation is randomized by rotational diffusion prior to energy transfer. Fortunately, variations in the actual value of κ2 do not appear to result in major errors in the distances calculated using FRET data (Lakowitz, 1999). Furthermore, when energy transfer is simply used as a signal to quantify the amount of complex formed, knowledge of the exact value of R0 is not necessary. The pronounced dependence of the efficiency of FRET on the distance between the two interacting species (1/r6 ) makes this method especially suitable for use as a molecular ruler. For example, FRET can be used to monitor the association of two proteins based on changes in the distance between them, as shown in Figure 20.8.11.
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Figure 20.8.11 Monitoring of complex formation using FRET. A schematic representation of the use of FRET to monitor the formation of a complex between protein A and compound S is shown. (A) Assume that compound S is labeled with a donor molecule (grey) and that protein A is labeled with an acceptor molecule (black). (B) In a solution in which both S and A are present but do not interact, irradiation with a wavelength corresponding to the excitation maximum for the donor leads to fluorescence of the donor, but to much less (if any) fluorescence of the acceptor (depending on its excitation spectrum). (C) However, when a complex between S and A is formed, such that the distance between acceptor and donor becomes small enough to allow FRET, the fluorescence of the donor decreases and the fluorescence of the acceptor increases. AU, arbitrary units.
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Examples of the use of FRET in the quantification of protein complex formation (either with another protein or with a small ligand) can be found in Cornea and Conn (2002), Johnson et al. (1993), and Wagner et al. (1995). The advantage of this method is that the fluorophores do not have to be located at the interaction interface, thus minimizing the potential for fluorescent labels to interfere with complex formation. The labels just have to be close enough (typically ≤ R0 ) in the final complex to allow significant energy transfer. F¨orster distances (R0 ) for typical donor-acceptor pairs range from 20 to 50 Å. The obvious disadvantages of FRET are that both compounds have to be labeled and that controls for unwanted effects of the labeling procedure have to be performed. Fluorescence anisotropy A detailed introduction to fluorescence depolarization is given in UNIT 7.7. By irradiating the sample with polarized light, molecules whose transition dipole moments are oriented parallel to the plane of polarization are preferentially excited. If the axes of the fluorophores maintain their orientation during this process, the emitted fluorescent light will also be polarized. Relative to this situation, the extent of polarization will decrease if the fluorophores exhibit rotational diffusion during the lifetimes of their excited states. In analogy to the translational diffusion coefficient, the rotational diffusion coefficient is a measure of how rapidly the rotation of a particular protein molecule occurs. This rotation leads to a change in the orientation of the emission transition dipole moment relative to the orientation of the excitation dipole moment. For small fluorescent compounds, the rotational diffusion coefficient is large enough to produce almost complete depolarization. The rotational diffusion coefficient of a fluorescent label will be reduced by attachment to protein A, which increases the mass of the rotating species. Upon binding of compound S to labeled protein A, the rotational diffusion coefficient can be further reduced. In order to characterize the degree to which the emitted fluorescence is polarized, two quantities are commonly used: polarization (P) and anisotropy (r) (see UNIT 7.7). In the quantitation of protein complexes, the use of anisotropy is preferred, because it yields simpler equations. Anisotropy is defined as: Spectroscopic Methods for the Study of Protein Interactions
Equation 20.8.12
where Ipar is the intensity of emission parallel to the excitation polarization and Iperp is the intensity of emission perpendicular to the excitation polarization. Important with regard to the effect of rotational diffusion on depolarization is the ratio of the lifetime of the fluorophore (τ ) to the rotational diffusion time (θ). According to the Perrin equation, the anisotropy observed in the presence of rotational diffusion (r) is related to the anisotropy observed in the absence of rotational diffusion (r0 ) as follows:
Equation 20.8.13
where D is the rotational diffusion coefficient. The anisotropy value can be used to determine the fraction of complexes formed, since it is related linearly to the fractional fluorescence intensity: robs = Fa ra + Fas ras where Fa and Fas are the fractional fluorescence values for the labeled protein and the complex, respectively, and ra and ras represent the anisotropy of the labeled protein and the complex, respectively. Thus, when using anisotropy to quantify complex formation, it is desirable to work at an emission wavelength at which the fluorescence intensity remains unchanged upon complex formation. By doing so, the equation above simplifies to: robs = fa ra + fas ras = ra + rfas where fas represents the fraction of binding sites that are saturated, fa represents the fraction of protein A molecules that are unbound (fa = 1 − fas ), and r = ras − ra . An example of the use of changes in tryptophan fluorescence anisotropy to monitor protein-protein interactions can be found in Lukas et al. (1986). Indirect quantification using a labeled ligand When the fluorescence of the system of interest does not change upon interaction and labeling of the interacting compounds is not feasible or successful, another approach, which involves the fluorescence of a third compound (usually a cofactor or fluorescent nucleotide), may be appropriate. This fluorescent ligand (referred to as “L” here) must bind specifically to the complex under investigation and must not interfere with the interaction between protein A and compound S.
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When such a ligand is used, the following binding reactions must be considered: K
A+S AS KL
ASL AS + L where L contains a label whose spectroscopic properties change significantly upon binding to the complex AS. Furthermore, it is assumed that L is present in large excess relative to the complex ([L] > 20 KL ). As a result, [ASL] = KL [L][AS] ≈ KL [L]tot [AS] = KL [L]tot K[A][S] = Kobs [A][S]. By observing the production of ASL at various concentrations of S (with the concentration of A fixed), the apparent binding constant (Kobs ) can be determined. Then, if KL and [L]tot are known, the binding constant (Kd = 1/K) can be calculated as well. Discussion of a similar approach, in which the product yielded by enzymatic activity interacts with a fluorescent component, can be found in Solomon and Stansbie (1984).
Critical Parameters To avoid nonspecific interactions, the protein and ligand should be as pure as possible. Temperature For fluorescence spectroscopy it is particularly important to maintain the chosen temperature since the fluorescence signal is temperature dependent (even if the interaction is not). The protein and ligand of interest should be sufficiently stable under the conditions chosen for the experiment, so that measurement of a complete titration curve is possible. Make sure that all solutions are at the required temperature and are in equilibrium before making experimental measurements. This is especially important if some of the solutions are kept on ice. Buffers The buffer solutions used in titration experiments have to meet two criteria: (1) they should create a stable environment for the protein and ligand under investigation, and (2) they should not interfere with the spectroscopic technique. Absorption and fluorescence spectra of the buffer solution over the relevant spectral range should be obtained so that any interference (inner filter effect or background signal) can be accounted for.
In particular, the following buffer solutions may create problems: (1) Imidazole is not a suitable buffer for absorption or fluorescence spectroscopy involving tryptophan fluorescence since it absorbs at the same wavelength as the fluorophore in this case. (2) Oxidized dithiothreitol absorbs light over a similar range of wavelengths as does tryptophan and should therefore be removed by dialysis; otherwise, reduced DTT should be added immediately before starting the experiment. In general, when fluorescence signals are low, the buffer fluorescence should always be subtracted. Buffers kept in plastic containers tend to contain small amounts of fluorescing species leached from the container material, with these species usually emitting in the region of tryptophan fluorescence. For all solutions, great care should be taken to ensure that no light-scattering particles are present. Thus, buffer solutions should be filtered, and it is recommended that all aspects of these experiments be kept as dust-free as possible.
Troubleshooting Spectral/kinetic data show sharp “spikes” This problem is typically due to light scattering from large particles (e.g., dust, aggregated protein). If spikes are observed, clarify the solutions (UNIT 7.7). Fluorescence decreases over time One possibility is that the solutions are not thermally equilibrated. This can be remedied by incubating solutions for a longer time at the required temperature before use. It is also possible that one or both of the reagents involved in the interaction are binding to the walls of the cuvette. Try using higher concentrations of the interacting molecules, or add 5% to 10% (v/v) glycerol to stabilize the protein. A third possibility is that the protein of interest is aggregating. Check to see if the solution is turbid. Stirring of the protein may cause damage, so try different methods of mixing, such as pipetting up and down or using small plastic paddles. Finally, if photobleaching is suspected, measure the change in fluorescence of the protein A solution over time according to the method described in Support Protocol 2.
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Linear increase in signal throughout titration, with no saturation The protein may be aggregating or polymerizing and may therefore exhibit a linear increase in fluorescence. Perform a control titration without protein A; specifically, add compound S in a stepwise fashion to a sample of buffer solution that does not contain protein A. Another possibility is that the Kd is very high (e.g., 100× the concentrations being used). Perform the titration using higher concentrations of A and S. Possible mixing or stirring artifacts Perform a buffer titration (stepwise addition of buffer, instead of compound S, to protein A solution) to determine whether the titration results are real or are an effect of the mixing or stirring procedure.
Spectroscopic Methods for the Study of Protein Interactions
Problems with fluorescent labeling of a protein (Basic Protocol 2) If the measured ratio of dye to protein is less than 1:1, a possible reason is that the dye concentration used was too low. An increase in dye concentration may lead to a higher yield of the labeled species. However, if the protein to be labeled contains unreactive amino acid residues, increasing the dye concentration will not change this ratio. Note that, in some cases, the extinction coefficient of the dye changes upon interaction with the protein, which means that the amount of conjugated protein appears to be lower than it actually is; in such cases, even an increase in label concentration may not change the calculated ratio. The easiest way to test for a change in the extinction coefficient is to measure the absorption of the label-protein complex and the free dye under native and denaturing conditions (e.g., 6 M guanidine-HCl). If the absorbance of the free dye is not changed by the presence of guanidine-HCl, but the absorbance of the dye in the conjugated protein is changed, then the difference must be due to the interaction of the dye with the protein. In this case, the amount of dye attached to the protein must be calculated using data from the denatured state. Increasing the dye concentration can lead to multiply labeled (or nonspecifically labeled) protein fractions, particularly when lysine residues are chosen for the attachment of the label. This is undesirable, because it is difficult to separate these differently labeled fractions. Nonetheless, it may be possible to separate such fractions using HPLC (UNIT 8.7).
In cases of multiple labeling (particularly amino-terminal labeling, where several lysine residues may be targeted), it may be important to determine the positions of the labels. This can be achieved by digesting the protein with trypsin (or another protease; UNIT 11.1) and analyzing the fragments using mass spectrometry (Chapter 16). In general, it is more desirable to retain a certain fraction of unlabeled protein (which simply results in a smaller change in signal) than to risk having multiply labeled proteins. Differently labeled proteins may have different fluorescence signals, which significantly complicate data analysis.
Anticipated Results If the protein does not aggregate or bind to the cuvette walls and does show a significant change in fluorescence upon complex formation, a binding curve such as the one shown in Figure 20.8.2 can be expected (assuming 1:1 stoichiometry and a simple binding mechanism). The equilibrium dissociation constant, as well as the stoichiometry value associated with the interaction, may be obtainable from the experimental data (Basic Protocol 3). Two examples of typical titrations, along with data analysis, are provided in Basic Protocol 3.
Time Considerations Performing a fluorescence titration and the necessary control experiments typically requires ∼2 hr. However, the time needed for titration depends largely on the rate of the binding event. If the binding interaction is very slow, the experiment may take considerably longer. In all cases, the reproducibility of the experiment must be verified. If one of the proteins has to be labeled, the experiment will require a considerably longer period of time. The labeling of the protein and the execution of the necessary control experiments (e.g., measuring the effects of labeling on protein activity and folding) may take several days. If a cysteine residue has to be introduced or removed prior to labeling, 2 to 3 weeks may be required before the fluorescence experiment can be started.
Literature Cited Solomon, M. and Stansbie, D. 1984. A coupled fluorometric rate assay for pyruvate dehydrogenase in cultured human fibroblasts. Anal. Biochem. 141:337-343. Bai, Y., Perez, G.M., Beechem, J.M., and Weil, P.A. 1997. Structure-function analysis of TAF130: Identification and characterization of a
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high-affinity TATA-binding protein interaction domain in the N terminus of yeast TAF(II)130. Mol. Cell. Biol. 17:3081-3093. Becker, C.F., Hunter, C.L., Seidel, R., Kent, S.B., Goody, R.S., and Engelhard, M. 2003. Total chemical synthesis of a functional interacting protein pair: The protooncogene H-Ras and the Ras-binding domain of its effector c-Raf1. Proc. Natl. Acad. Sci. U.S.A. 100:5075-5080. Cornea, A. and Conn, P.M. 2002. Measurement of changes in fluorescence resonance energy transfer between gonadotropin-releasing hormone receptors in response to agonists. Methods 27:333339. Hiratsuka, T. 1998. Prodan fluorescence reflects differences in nucleotide-induced conformational states in the myosin head and allows continuous visualization of the ATPase reactions. Biochemistry 37:7167-7176. Hutnik, C.M., MacManus, J.P., Banville, D., and Szabo, A.G. 1991. Metal-induced changes in the fluorescence properties of tyrosine and tryptophan site-specific mutants of oncomodulin. Biochemistry 30:7652-7660. Jezewska, M.J. and Bujalowski, W. 1997. Quantitative analysis of ligand-macromolecule interactions using differential dynamic quenching of the ligand fluorescence to monitor the binding. Biophys. Chem. 64:253-269. Johnson, D.A., Leathers, V.L., Martinez, A.M., Walsh, D.A., and Fletcher, W.H. 1993. Fluorescence resonance energy transfer within a heterochromatic cAMP-dependent protein kinase holoenzyme under equilibrium conditions: New insights into the conformational changes that result in cAMP-dependent activation. Biochemistry 32:6402-6410. Kelly, R.C. and von Hippel, P.H. 1976. DNA “melting” proteins: III: Fluorescence “mapping” of the nucleic acid binding site of bacteriophage T4 gene 32-protein. J. Biol. Chem. 251:7229-7239. Lakowitz, J.R. 1999. Principles of Fluorescence Spectroscopy. Kluwer Academic/Plenum Publishers, New York. LeTilly, V. and Royer, C.A. 1993. Fluorescence anisotropy assays implicate protein-protein interactions in regulating trp repressor DNA binding. Biochemistry 32:7753-7758. Li, Q., Du, H.N., and Hu, H.Y. 2003. Study of protein-protein interactions by fluorescence of tryptophan analogs: Application to immunoglobulin G binding domain of streptococcal protein G. Biopolymers 72:116-122.
Wagner, A., Simon, I., Sprinzl, M., and Goody, R.S. 1995. Interaction of guanosine nucleotides and their analogs with elongation factor Tu from Thermus thermophilus. Biochemistry 34:1253512542. Xia, X., Lin, J.T., and Kinne, R.K. 2003. Binding of phlorizin to the isolated C-terminal extramembranous loop of the Na+/glucose cotransporter assessed by intrinsic tryptophan fluorescence. Biochemistry 42:6115-6120.
Key References Gore, M.G. 2000. Spectrophotometry and Spectrofluorimetry: A Practical Approach. Oxford University Press, Oxford. Describes the theory and practical applications of many spectroscopic techniques, including fluorescence and absorption spectroscopy and circular dichroism. Emphasis is placed on the practical aspects of these techniques, and protocols for test experiments as well as lists of background literature are provided. A very good introduction to spectrometry. Lakowitz, 1999. See above. Provides basic and in-depth information on fluorescence spectroscopy, including an overview of extrinsic fluorophores. Two chapters discuss timeresolved measurements (advanced). Creighton, T.E. 1993. Proteins: Structures and Molecular Properties. W.H. Freeman, New York. A basic introduction to the use of spectroscopic methods for detecting conformational changes.
Internet Resources http://www.probes.com Molecular Probes Web site. Includes an online version of the Handbook of Fluorescent Probes and Research Products, with up-to-date background and technical information. An e-mail newsletter is also available.
Contributed by Yvonne Groemping Department of Biomolecular Mechanisms Max Planck Institute for Medical Research Heidelberg, Germany Nadja Hellmann Institute for Molecular Biophysics University of Mainz Mainz, Germany
Lukas, T.J., Burgess, W.H., Prendergast, F.G., Lau, W., and Watterson, D.M. 1986. Calmodulin binding domains: Characterization of a phosphorylation and calmodulin binding site from myosin light chain kinase. Biochemistry 25:1458-1464.
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Application of Amide Proton Exchange Mass Spectrometry for the Study of Protein-Protein Interactions
UNIT 20.9
Amide proton exchange experiments have traditionally been performed to study protein folding and unfolding by measuring the solvent exchange rates of amide protons that participate in hydrogen bond formation (Englander et al., 1997). In studies of proteinprotein interfaces, the hydrogen bonds examined are usually those that exhibit decreased solvent accessibility arising from factors such as side chain interactions and decreased loop mobility, because hydrogen bonds rarely form across interfaces. Amide groups on the interior of a protein exchange protons with solvent more slowly than do amide groups on the surface, and it is almost always the more rapidly exchanging surface amides that require monitoring in protein-protein interface studies. Typically, amides on the surface of a protein exchange protons on a time scale ranging from seconds to minutes. The corresponding exchange rate is slower than that for amides in unstructured peptides (Bai et al., 1993), but relatively faster than that for amide groups residing within the core of a protein (Dharmasiri and Smith, 1996). Early amide proton exchange experiments relied on tritium labeling and separation by chromatography; however, these early methods have been supplanted in recent years by NMR and mass spectrometry, both of which measure deuterium incorporation (Rosa and Richards, 1979; Englander and Mayne, 1992; Zhang et al., 1996). NMR has the advantage of being able to detect single amide groups. Among the disadvantages are that the protein must be of a size that is tractable for NMR spectroscopy (typically <40 kDa), that it must be expressible, and that it must be isotopically labeled with 15 N so that resonance assignments for the amide protons can be made. In contrast, mass spectrometric measurements avoid the need for isotopic labeling of the protein and can be performed on proteins of any size (UNIT 17.6). The disadvantage of mass spectrometry is that, at present, single-amide resolution is not possible. Mass spectrometric measurements rely on proteolytic digestion of the protein and analysis of the resulting peptide fragments. Thus, the resolution is limited by the sizes of these peptide fragments, although it can be improved if many overlapping fragments are obtained. Because the methodology of measuring amide proton exchange by mass spectrometry is already covered in UNIT 17.6, this unit focuses on the particulars of experiments used to study protein-protein interactions.
STRATEGIC PLANNING The protein-protein binding equilibrium constant plays an important role in determining whether interface protection is observed. If the proteins in a complex are dissociating and reassociating rapidly, then the amide protons will have time to exchange with solvent protons while the proteins are dissociated, and interface protection will not be observed. To address this issue, some knowledge of the binding equilibrium constant is required. Titration calorimetry (UNIT 20.4) is an excellent way to obtain the binding constant, and information on the kinetics of association and dissociation is best obtained from optical biosensor measurements (UNIT 20.2). In the systems the authors have studied by off-exchange, there are amides at the proteinprotein interface that are completely protected from exchanging protons with solvent as long as the two proteins remain bound (Baerga-Ortiz et al., 2004; Mandell et al., 2001). Quantification of Protein Interactions Contributed by Jeffrey G. Mandell, Abel Baerga-Ortiz, Carrie H. Croy, Arnold M. Falick, and Elizabeth A. Komives
20.9.1
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Figure 20.9.1
Legend at right.
H-Exchange MS to Study Protein-Protein Interactions
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For these completely solvent-excluded amides, the observed 1 H/2 H exchange rate is a function of the protein-protein dissociation and association rates, as well as the intrinsic rate of hydrogen exchange (kex ): kd
k
ex [RD ·L] RD + L → RH
ka
where RH is the protonated receptor, RD is the deuterated receptor, L is the ligand, kex is the intrinsic proton exchange rate (in min−1 ) for amides in the uncomplexed receptor, kd is the rate of dissociation of the complex (in min−1 ), and ka is the rate of association of the proteins undergoing complexation (in M−1 ·min−1 ). 1 H/2 H exchange at a proteinprotein interface could occur because of solvent accessibility to the interface, or it could occur because the complex has dissociated. A theoretical analysis shows that under the conditions of the 1 H/2 H exchange experiment presented in this unit (protein concentration, 3 µM), if Kd (which equals kd /ka ) is <2 nM, a 1:1 ratio of ligand to receptor is sufficient to ensure that essentially 100% of the receptors are bound to ligand throughout the experiment. If Kd is between 10 and 100 nM, then higher ratios of ligand to receptor are required to study 1 H/2 H exchange kinetics at the interface in the bound complex, and the interplay between binding kinetics and exchange kinetics becomes important (Mandell et al., 2001). In these cases, full knowledge of the binding kinetics is required prior to quantitative interpretation of the data. For protein-protein interactions that are in rapid equilibrium, the amides excluded at the interface should not show pH-dependent exchange rates (Mandell et al., 2001). This unit contains two protocols for studying protein-protein interfaces by monitoring amide proton exchange using matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) mass spectrometry (UNIT 16.2, UNIT 16.3, and UNIT 16.4). The simplest protocol, known as the “on-exchange” experiment (see Basic Protocol 1), is used to determine the location of the protein-protein interface. In this experiment, a comparison is made between surface amide deuteration in the uncomplexed proteins and surface amide deuteration in the protein-protein complex (Fig. 20.9.1A). The part of each protein that shows less deuteration in the complex than in the uncomplexed state is the interacting surface, or interface. On-exchange data, in combination with computational docking, have been used by Anand et al. (2003) to build a model of the protein kinase A holoenzyme complex. The “off-exchange” experiment (see Basic Protocol 2) yields interface protection data that pertain more to conformational changes (Fig. 20.9.1B). In this experiment, by first monitoring on-exchange (i.e., replacement of hydrogen atoms with deuterium atoms) in the separate proteins, one can assess which regions are highly solvent accessible prior to complexation; these regions are likely to be on the surface of the protein. If any of these highly accessible regions are subsequently found to be protected when the deuterated proteins are allowed to complex and off-exchange (i.e., replacement of deuterium atoms with hydrogen atoms) is measured, it is safe to assume that they are at the protein-protein interface, as it is unlikely that a highly accessible surface region will fold into the protein core so as to become completely inaccessible upon complex formation. Figure 20.9.1 (at left) Flow chart diagram of the two types of amide proton exchange experiments used to study protein-protein interfaces. (A) In the on-exchange experiment, the protein-protein complex shows a region in which less deuterium is incorporated when compared with control samples in which each protein is deuterated separately. (B) In the off-exchange experiment, each protein is allowed to incorporate deuterium separately, the deuterated proteins are allowed to complex, and then deuterium atoms are off-exchanged with hydrogen atoms by dilution in H2 O; residues located at the protein-protein interface are characterized by the retention of deuterons in the protein-protein complex as compared with a lack of retention in control experiments in which only one of the two proteins is present.
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The “off-exchange” experiment also allows discrimination between partially inaccessible and completely inaccessible regions of the interface. The ability of solvent H2 O to penetrate a protein-protein interface, as well as the idea that H2 O molecules are released from some portion of a protein’s surface when that protein interacts with another one, is a topic of current research interest in biophysics (Baerga-Ortiz et al., 2004; Mandell et al., 2001). BASIC PROTOCOL 1
IDENTIFICATION OF THE PROTEIN-PROTEIN INTERFACE VIA THE ON-EXCHANGE METHOD The simplest protocol used to study protein-protein interfaces by monitoring amide proton exchange using MALDI-TOF mass spectrometry is the “on-exchange” experiment, which is used to find the location of the protein-protein interface. In this experiment, a comparison is made between surface amide deuteration in the uncomplexed proteins and surface amide deuteration in the protein-protein complex. The part of each protein that shows less deuteration in the complex than in the uncomplexed state is the interacting surface, or interface. Thus, the experiment involves separately deuterating the individual proteins and the protein complex by incubation in D2 O buffer. After quenching of the exchange reaction and proteolytic digestion of each sample, mass spectrometry is used to reveal regions in the protein complex that were protected from exchange (Fig. 20.9.1A). In order to interpret the results of the “on-exchange” experiment, it is necessary to assume that protection occurs because of interface formation, and not because of conformational changes. If the structures of the interacting proteins are known, it will be readily apparent as to whether this assumption is valid. Prior to carrying out an amide proton exchange experiment, it is necessary to determine the pepsin digestion conditions that result in the greatest yield of peptide fragments from the protein under study, and it is also necessary to identify the peptides that are obtained from pepsin digestion. Determination of optimal digestion conditions and identification of peptide fragments only need to be performed once for a particular protein and should be performed under conditions identical to those that will be used in the actual amide proton exchange experiment. Matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) mass spectrometry (UNIT 16.2, UNIT 16.3, and UNIT 16.4) is used in this protocol. The methodology for measuring amide proton exchange by MALDI-TOF mass spectrometry is relatively simple, and no modification of commercially available instruments is required (Mandell et al., 1988a,b). Mass spectrometric measurement of amide proton exchange was pioneered by David Smith in the early 1990s, using electrospray ionization (Zhang and Smith, 1993). Amide proton exchange results identical to those obtained using Basic Protocol 1 can be achieved using an electrospray mass spectrometer. Furthermore, high-resolution sequencing information similar to that obtained using a MALDI-TOF mass spectrometer can also be obtained using a tandem quadrupole time-of-flight mass spectrometer.
Materials
H-Exchange MS to Study Protein-Protein Interactions
10× low-salt buffer (e.g., 25 mM Tris·Cl, 25 mM NaCl, 50 mM Na/KHPO4 ) Purified receptor and ligand proteins 0.1% and 2.0% (v/v) trifluoroacetic acid (TFA; highest purity possible) 2 to 3 mg/ml immobilized pepsin in 6% (w/v) cross-linked beaded agarose (Pierce Biotechnology) Matrix solution for MALDI-TOF mass spectrometry (see recipe) Peptide mass standard mixture (e.g., Sequazyme Calibration Mixture 2; Applied Biosystems) Deuterium oxide (D2 O; 99.996% purity; Cambridge Isotope Laboratories)
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pH meter with InLab 423 electrode (Mettler Instruments) Cold box, 4◦ C (Isotemp Chromatography Refrigerator; Fisher Scientific) Microcentrifuge, 4◦ C (in cold box) 0.5-ml thin-walled microcentrifuge tubes (USA/Scientific) MALDI target plates, 4◦ C (in cold box; chilled overnight in a plastic case to prevent absorption of atmospheric H2 O) Vacuum desiccator (with liquid nitrogen trap) located adjacent to MALDI-TOF mass spectrometer MALDI-TOF mass spectrometer (UNIT 16.2, UNIT 16.3, and UNIT 16.4) C18 analytical column Additional reagents and equipment for reversed-phase high-performance liquid chromatography (HPLC; UNIT 8.7) and post-source decay sequencing (UNIT 16.1), carboxypeptidase Y C-terminal sequencing (UNIT 11.8), or electrospray ionization tandem mass spectrometric (ESI MS/MS) sequencing (UNIT 16.10 and UNIT 16.11) NOTE: The amide proton exchange reaction is highly sensitive to temperature; exchange is rapid at room temperature but slows dramatically when the reaction sample is chilled to 4◦ C. Thus, all sample manipulations subsequent to the timed incubation of the receptor, ligand, or receptor-ligand complex in D2 O must be performed at 4◦ C. The simplest experimental setup that makes this possible involves having a cold box in the same room as the mass spectrometer. Optimize pepsin digestion conditions 1. In 10× low-salt buffer, prepare a 100 pmol/µl sample of one of the two proteins under study. Transfer a 1.2-µl aliquot of this sample to each of several 0.5-ml microcentrifuge tubes, and dilute each aliquot with 12 µl H2 O to yield a sample that is ∼1× with respect to the buffer solution. To screen digestion conditions, four to ten separate samples will be required.
2. Dilute each protein sample (protein in ∼1× buffer, from step 1) 10-fold by adding 120 µl of 0.1% TFA, and measure the pH of the resulting solution. For each sample, use 2% TFA to adjust the pH to match the original pH of the added 0.1% TFA solution (pH 2.0 to 2.5), making note of how much TFA was required, and then chill the sample to 4◦ C in the cold box. When preparing on-exchange samples in subsequent steps, the volume of 2% TFA needed to achieve the desired (optimum) pH will be known, making it unnecessary to monitor the pH of each experimental sample while adding 2% TFA.
3. Immediately before use, transfer immobilized pepsin (25 µl of pepsin beads, in the form of 50 µl of the 1:1 pepsin bead slurry supplied by the manufacturer) to each of several 1.5-ml microcentrifuge tubes. Wash the immobilized pepsin twice, each time by adding 1 ml chilled 0.1% TFA, pH 2.0 to 2.5, per tube; vortexing; microcentrifuging 2 min at 7000 × g, 4◦ C; and then removing all supernatants. Pepsin is used to cleave proteins into fragments of 8 to 20 amino acids (Zhang and Smith, 1993). Pepsin beads can be washed and reused ∼5 times, but they begin to lose their activity 4 hr after acidification. The pH of the pepsin digestion is controlled by the pH of the 0.1% TFA used to wash the pepsin beads in this step. The washing pH for each tube in this step should match the pH of a sample from step 2.
4. Add each protein sample (step 2) to a separate tube of immobilized pepsin (washed at the pH of that sample) and incubate at 4◦ C, using each sample to test a different combination of incubation time and pH.
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The completeness and specificity of pepsin cleavage depend on incubation time and pH; therefore, it is necessary to vary these two parameters to find the conditions that lead to optimal pepsin digestion product yields. The optimal incubation time typically ranges from 1 to 10 min, and the optimal pH typically ranges from 2.0 to 2.5.
5. Microcentrifuge the digested protein samples 1 min at 14,000 × g, 4◦ C. For each sample, transfer 20-µl aliquots of the supernatant to separate 0.5-ml thin-walled microcentrifuge tubes and immediately freeze in liquid nitrogen. Store aliquots up to 2 weeks at −80◦ C. In experimental runs, on-exchange samples should be quenched (i.e., treated with TFA), digested, and frozen in <15 min total, since the median time for amide deuteron back exchange under quench conditions is ∼15 min.
6. In the following way, obtain a MALDI-TOF mass spectrum of at least one 20-µl aliquot of digest mixture obtained under each distinct set of digestion conditions. a. Quickly defrost the frozen digestion sample by warming the sample tube by hand, with periodic flicking of the tube to help break up/liquefy the sample. Defrosting should be completed within 30 sec of removing the sample from frozen storage.
b. Once the sample has been defrosted, transfer the tube to a prechilled microcentrifuge tube rack sitting in the cold box. c. Transfer 5 µl of sample to a clean, prechilled microcentrifuge tube, add an equal volume of prechilled matrix solution, and mix. d. Spot 1 µl of the resulting mixture onto a prechilled MALDI target plate, and then immediately transfer the target plate to a vacuum desiccator and apply moderate vacuum. If many samples are to be analyzed, it will be necessary to have at least two additional target plates (i.e., aside from the one in use) being chilled in the cold box at any given time, so that new samples can be prepared without having to wait for a target plate to cool to the appropriate temperature. A SpeedVac evaporator can be used in place of a vacuum desiccator to dry the target plate. The strength of the vacuum should be such that it takes ∼1 min for the spot to dry. Slow drying of the spot under moderate vacuum or by cycling the vacuum on and off results in higher-quality sample analysis, presumably because it leads to improved crystal growth.
e. Once the plate has dried, transfer it as quickly as possible (i.e., within 10 sec) to the mass spectrometer and acquire a MALDI-TOF spectrum in accordance with the manufacturer’s instructions. 7. Based on the MALDI-TOF data obtained in step 6, identify the pepsin digestion conditions that result in the highest product yield. It is important that the conditions used in the actual amide proton exchange experiment be identical to those used in the optimized pepsin digestion so that proteolysis products can be accurately identified.
H-Exchange MS to Study Protein-Protein Interactions
Identify pepsin proteolysis products by mass spectrometry 8. Thaw another aliquot of the digest mixture produced under the optimized proteolysis conditions, combine the aliquot with a peptide mass standard mixture, and analyze by MALDI-TOF mass spectrometry (step 6; add mass standard to digest mixture before combining a 5-µl aliquot with matrix solution). Calibrate the mass spectrum using the masses of the standards. A typical mass standard mixture will require 1:300 dilution before being added to the pepsin digest mixture for MALDI-TOF MS analysis.
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Figure 20.9.2 (A) Example of a matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) mass spectrum of pepsin digest products. The inset shows that the high resolution of the time-of-flight mass analyzer allows baseline resolution of the mass envelope for each peptide. Each peptide’s mass envelope results from the natural abundance of 13 C within that peptide, such that the peak representing the fraction of peptide molecules containing one 13 C atom appears one mass unit to the right of the monoisotopic peak, the peak representing the fraction containing two 13 C atoms appears two mass units to the right of the monoisotopic peak, and so on. (B) Sequence coverage map for the protein whose peptide digest mass spectrum is shown in panel A. Each of the lines under the sequence represents a peptide fragment whose mass matches that of a peak observed in the mass spectrum.
The MALDI-TOF mass spectrum acquired in this step contains all pepsin digest products that will be observed for this protein in subsequent deuterium exchange experiments (Fig. 20.9.2A). If a typical, well-tuned MALDI-TOF instrument is used, the masses of the peptides should be accurate to within 20 ppm (tolerance of 0.02 for a peptide of mass 1000), provided that internal standards are used.
9. From the calibrated spectrum, determine the MH+ masses of all observed pepsin digest products. Using any of a number of search engines available on the Internet, search the entire sequence of the protein of interest and identify all potential peptide fragments whose masses match (to within 20 ppm) an MH+ mass that appears in the calibrated spectrum. For some of the observed MH+ masses, there will be only one peptide fragment that falls within the 20-ppm tolerance window, and so a peak assignment can be made unambiguously. Once at least two peptide fragments are identified unambiguously in this manner, they should be used as internal calibration standards in all subsequent mass spectra.
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10. In the following way, identify all peptide fragments in the digestion mixture that have not yet been identified unambiguously. a. Thaw another five 20-µl aliquots of the digest mixture produced under the optimized proteolysis conditions and pool them. Separate the peptides in the pooled mixture using a C-18 analytical column and reversed-phase HPLC (UNIT 8.7), and collect each peak fraction. Each peptide fragment that is responsible for a peak in the mass spectrum will be found in one of the HPLC-separated fractions and can be readily sequenced from that purified fraction.
b. Sequence each previously unidentified peptide obtained in substep a by mass spectrometric methods. HPLC separation of ∼1 nmol of protein will provide sufficient peptide for postsource decay sequencing (UNIT 16.1) or carboxypeptidase Y C-terminal sequencing (UNIT 11.8), both of which are MALDI-based methods, or for ESI MS/MS sequencing (UNIT 16.10 and UNIT 16.11) using an ion trap or tandem quadrupole-time-of-flight mass spectrometer. It is important to note that while pepsin cleavage is not predictable, it is completely reproducible. Therefore, the sequencing of peptide fragments resulting from the digestion of a particular protein need only be carried out once prior to a set of amide proton exchange experiments.
11. Map all identified peptide fragments back onto the sequence of the protein to determine sequence coverage (Fig. 20.9.2B). If there are regions of the protein that are not covered by the peptide fragments found in the mass spectrum, it may be necessary to change the pepsin digestion conditions, perhaps by including up to 1 M urea as a denaturant (if the protein is particularly stable) or by using a different protein concentration. Buffers can also play a role, with low salt concentrations (<10 mM) generally being preferable for all types of mass spectrometry; thus, reducing the salt content in the initial protein solution often improves sequence coverage.
12. Repeat steps 1 to 11 using the other of the two proteins being studied. 13. Prepare a 100 pmol/µl sample of the receptor protein in 10× buffer, and then prepare a separate 100 pmol/µl sample of the ligand protein in 10× buffer. In each of several 0.5-ml microcentrifuge tubes, mix 0.6 µl of the receptor protein solution and 0.6 µl of the ligand protein solution and then dilute with 12 µl H2 O. Using the resulting samples, repeat steps 2 to 11 to identify the optimal digestion conditions for the receptor-ligand complex.
Perform amide proton exchange experiments to determine time course of deuteration for each protein 14. In the following way, prepare several microcentrifuge tubes (one for each onexchange time point—0, 1, 2, 5, 10, and 20 min, typically) containing the quench solution for one of the two proteins being studied. a. To each tube, add 120 µl of chilled 0.1% TFA (4◦ C) at the optimal digestion pH for the protein in question. b. To each tube, add the appropriate volume of 2% TFA for maintaining the optimal digestion pH after the addition of a sample of the protein (as determined in step 2). c. Chill the tubes at 4◦ C in the cold box. H-Exchange MS to Study Protein-Protein Interactions
15. For each tube from step 14, prepare a tube of immobilized pepsin beads as described in step 3, making sure to wash at the optimal digestion pH.
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16. In 10× buffer, prepare a 100 pmol/µl sample of one of the two proteins under study. Transfer a 1.2-µl aliquot of this sample to each of several 0.5-ml microcentrifuge tubes, and dilute each aliquot with 12 µl D2 O. Incubate each of the resulting samples for a different length of time (ranging from 0 to 30 min, except for one sample, for which incubation should last 24 hr) at the temperature of interest (typically 25◦ to 37◦ C). The sample that is incubated for 24 hr is subsequently used to correct experimental results for the effects of back exchange (Support Protocol 1, step 5).
17. At the end of each sample’s incubation period, quench on-exchange by adding the contents of one tube of quench buffer to the sample, transfer the resulting mixture to a tube of immobilized pepsin beads, and incubate for the optimal length of time (as identified in step 7) at 4◦ C. When digestion is complete, centrifuge, aliquot, freeze, and store the digest mixture as described in step 5. 18. Repeat steps 14 to 17 for the other of the two proteins being studied.
Perform amide proton exchange experiment to identify protein-protein interface 19. Prepare several microcentrifuge tubes (one for each on-exchange time point) containing the quench solution for the receptor-ligand complex (see step 14). 20. For each tube from step 19, prepare a tube of immobilized pepsin beads as described in step 3, making sure to wash at the optimal digestion pH for the receptor-ligand complex. 21. Prepare a 100 pmol/µl sample of the receptor protein in 10× buffer, and then prepare a separate 100 pmol/µl sample of the ligand protein in 10× buffer. In each of several 0.5-ml microcentrifuge tubes, mix 0.6 µl of the receptor protein solution and 0.6 µl of the ligand protein solution and then dilute with 12 µl D2 O. Incubate each of the resulting samples for a different length of time (ranging from 0 to 30 min, except for one sample, for which incubation should last 24 hr) at the temperature of interest. Alternatively, a 100 pmol/µl sample of the protein complex in 10× buffer can be prepared in advance and used in place of the separate receptor and ligand solutions that are mixed together in this step. The sample that is incubated for 24 hr is subsequently used to correct experimental results for the effects of back exchange (Support Protocol 1, step 5).
22. At the end of each sample’s incubation period, quench on-exchange by adding the contents of one tube of quench buffer to the sample, transfer the resulting mixture to a tube of immobilized pepsin beads, and incubate for the optimal length of time (as identified in step 7) at 4◦ C. When digestion is complete, centrifuge, aliquot, freeze, and store the digest mixture as described in step 5.
Obtain mass spectra of amide proton exchange samples 23. Obtain mass spectra of the digested amide proton exchange samples (steps 17, 18, and 22) as described in step 6, substeps a to e. Analyze all mass spectrometric data as described in Support Protocol 1. Amide proton exchange samples for the receptor, ligand, and receptor-ligand complex should all be analyzed on the same day using the same batch of matrix solution, as doing so produces more consistent results. In addition, only a single block of mass spectrometer time will need to be reserved if all spectra are obtained on the same day. For each protein, the interface region is identified as the area that shows decreased solvent accessibility in the complex as compared with the uncomplexed state (Fig. 20.9.3). Quantification of Protein Interactions
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Figure 20.9.3 In the on-exchange experiment, the interacting proteins, both separately and in the complexed state, are incubated in deuterated buffer for varying lengths of time prior to quenching, pepsin digestion, and mass spectrometric analysis. (A) Example of a peptide mass envelope resulting from pepsin digestion of (i) nondeuterated CheB protein; (ii) a deuterated protein complex involving CheB; and (iii) CheB alone after being subjected to the same deuteration period as the complex in (ii). The mass envelope broadens somewhat upon deuteration, and there is less deuteration of the peptide in the protein-protein complex than in the protein alone. (B) Graph of number of deuterons incorporated into the peptide shown in panel A versus deuteration time. Circles, uncomplexed CheB; squares, complexed CheB.
SUPPORT PROTOCOL 1
ANALYSIS OF EXPERIMENTAL DATA FROM THE ON-EXCHANGE EXPERIMENT Materials MALDI-TOF mass spectra of pepsin digest products from all receptor, ligand, and receptor-ligand complex on-exchange samples (Basic Protocol 1) MALDI-TOF mass spectra of pepsin digest products from nondeuterated receptor, ligand, and receptor-ligand complex samples (Basic Protocol 1) Software for displaying multiple mass spectra in stacked layout (provided by manufacturer of the mass spectrometer) CAPP software package (Mandell et al., 1988b) KaleidaGraph 3.0 (Synergy Software) or other nonlinear fitting software 1. Display the mass spectra from Basic Protocol 1, step 23, using software that allows the stacking of plots, which will facilitate accurate assignment of monoisotopic peaks (Fig. 20.9.3). Even in the mass spectra of nondeuterated proteins, a “mass envelope” will be observed for each peptide fragment. Within each mass envelope, the lowest-mass peak (termed the MH+ peak) corresponds to the version of the peptide fragment in which all carbons are 12 C atoms, and the next highest–mass peak corresponds to a version of the peptide fragment in which one 13 C atom is present. It is possible to calculate the expected relative heights of the peaks in a mass envelope based on the natural abundance of 13C and on the number of carbon atoms in the peptide fragment.
H-Exchange MS to Study Protein-Protein Interactions
Because the mass of 2 H is exactly one mass unit greater than that of 1 H, the presence of a 2 H atom in a peptide fragment will shift the entire mass envelope for that fragment by one mass unit. Usually, 2 H incorporation also leads to broadening of the mass envelope, although this depends on the distribution (across individual molecules) of the number of 2 H atoms incorporated.
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2. Stack the mass spectrum of each deuterated sample on top of the mass spectrum of the corresponding nondeuterated sample. Use the known masses of the peptide fragments in the nondeuterated sample (see Basic Protocol 1, step 9) to calibrate the mass spectra of the corresponding deuterated samples. For some deuterated peptide fragments, a monoisotopic peak will not be present in the mass envelope, and so higher-mass peaks within the same envelope will need to be used. If necessary, these peaks can be identified in the uncalibrated (other than instrument calibration) stacked plots, and since they differ from their corresponding MH+ peaks by an exact integer number of mass units, they can be used for internal calibration.
3. Calculate the average mass of each peptide fragment in each deuterated sample and each nondeuterated sample by determining the centroid of the fragment’s mass envelope using CAPP software (Mandell et al., 1998b). Determine the raw amount of deuterium present in each peptide fragment at each on-exchange time point by calculating the difference between that fragment’s average mass in each D2 O labeling sample and its average mass in the nondeuterated sample in which it appears. CAPP software is available free of charge from the author ([email protected]). The difference between the average mass of a peptide fragment in a deuterated sample and the mass of that same fragment in the nondeuterated sample equals the raw amount of deuterium that has been incorporated. However, when using MALDI-TOF 1 H/2 H exchange data, two corrections (steps 4 and 5) are required to convert the raw result into the actual number of deuterons that have exchanged positions with protons.
4. In the following way, correct each raw result (step 3) for deuterium content at the rapidly exchanging sites in each peptide due to the presence of a residual amount of deuterium in the MALDI-TOF sample. a. Based on the extent to which D2 O was diluted during labeling, pepsin digestion, and subsequent MALDI-TOF sample preparation, calculate the percentage of deuterium present in the MALDI-TOF sample spot. For example, if the experiment involved mixing 100% D2 O with a concentrated protein solution in a 90:10 ratio (for protein labeling), diluting the resulting mixture by a factor of 10 with TFA (for pepsin digestion), and then diluting the digest mixture by a factor of 2 with matrix solution (for MALDI-TOF sample preparation), then the residual deuterium content in the sample spot would be 90% × 1/10 × 1/2 = 4.5% (0.045).
b. Calculate the total number of rapidly exchanging proton positions in the peptide fragment of interest (3 at the N-terminus and 1 at the C-terminus, plus 1 for every Asp, Glu, Cys, His, Ser, Thr, Tyr, or Trp residue; 2 for every Asn, Gln, or Lys residue; and 4 for every Arg residue). c. Multiply the proportion of residual deuterium present in the MALDI-TOF sample spot (substep a) by the number of rapidly exchanging proton positions in the peptide fragment of interest (substep b), and subtract the resulting quantity from the raw result obtained in step 3. 5. In the following way, correct all non–24-hr incubation results from step 4 for back exchange to determine the true amount of deuterium incorporation during the labeling period. a. For each 24-hr labeling sample, using the results from step 4, determine what percentage of potential amide proton exchange sites in each peptide fragment are actually deuterated. In each sample, identify the peptide fragment containing the highest percentage of deuterated sites.
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The percentage of nondeuterated sites in the most deuterated peptide fragment is taken to be the percentage of sites at which back exchange occurred in each corresponding sample during analysis. For example, if the most deuterated peptide fragment in the 24-hr labeling sample shows deuteration at 71% of all possible amide proton exchange sites, then the overall back exchange frequency is calculated to be 100% − 71% = 29%. On-exchange will have run its complete course in some, but not all, regions of a globular protein by 24 hr.
b. Obtain fully corrected results by dividing each non–24-hr incubation result from step 4 by the percentage of deuterated sites in the most deuterated peptide fragment from the corresponding 24-hr labeling sample. For example, if a peptide fragment obtained from a receptor labeling sample was calculated (up to this point) to contain an average of 5 deuterons per molecule, and the maximum percentage of deuterated sites in any peptide fragment obtained from the 24-hr receptor labeling sample was 71%, then the actual amount of deuterium incorporation into the fragment of interest would be 5/0.71 = 7 deuterons per molecule. Since all peptide fragments are analyzed simultaneously in the MALDI-TOF experiment, the same back exchange correction term is used for all fragments obtained from the same type of sample (receptor, ligand, or receptor-ligand complex).
6. Using the corrected results obtained in step 5, construct a plot of deuterium incorporation versus labeling time for each peptide fragment yielded by each type of sample. Use KaleidaGraph 3.0 (Synergy Software) or any other appropriate nonlinear fitting program to fit the data in each plot to the following two-exponential model, which accounts for positions at which exchange is very rapid (i.e., amides that are fully solvent accessible) and positions at which exchange occurs more slowly (i.e., amides with reduced solvent accessibility in the folded protein):
where D is the total number of deuterons that have undergone on-exchange by time t; Nfast is the number of deuterons exchanging at the faster rate, kfast ; and Nslow is the number of deuterons exchanging at the slower rate, kslow . All instances of rapid exchange will have occurred by the first time point, so kfast can be fixed to improve fitting accuracy. Other floating parameters (Nfast , Nslow , and kslow ) are completely insensitive to changes in the value of kfast over a range from 10 min−1 to 100 min−1 , and so kfast can be set to 30 min−1 , the median amide proton exchange rate for rapidly exchanging amide protons in the P13 domain of E. coli mannose permease (Gemmecker et al., 1993). Depending on the range of incubation times sampled and the number of data points, a single exponential fit will sometimes be sufficient to describe the data obtained. In such a case, both the number of deuterons undergoing on-exchange and the rate of on-exchange should be fitted parameters. The plots obtained from analysis of the mass envelope in Figure 20.9.3A are shown in Figure 20.9.3B. In this example, the peptide fragment being analyzed corresponds to an interface region. It is clear from the raw data that this region is protected in the complex (Fig. 20.9.3A), and this protection is confirmed at various on-exchange time points out to 5 min (Fig. 20.9.3B). H-Exchange MS to Study Protein-Protein Interactions
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DETERMINATION OF THE SOLVENT ACCESSIBILITY OF A PROTEIN-PROTEIN INTERFACE VIA THE OFF-EXCHANGE METHOD
BASIC PROTOCOL 2
The rate of off-exchange of deuterons from a protein-protein interface can provide additional information about the solvent accessibility of the interface (Mandell et al., 2001). The protocol described here includes measurement of the on-exchange of deuterium into each of the proteins involved in an interaction. After on-exchange is measured, a suitable set of on-exchange time points are chosen, and the protein samples corresponding to those time points are then combined to allow complex formation, after which the complex is diluted in H2 O for the off-exchange portion of the experiment (Fig. 20.9.1B). With both the on-exchange and off-exchange properties of the interface available, the effects of conformational changes and the effects of interface protection can more easily be distinguished.
Additional Materials (also see Basic Protocol 1) H2 O buffer: 10 mM Tris·Cl (APPENDIX 2E), pH 6.6 to 7.9, as appropriate 1. Determine optimal digestion conditions for the separate proteins of interest and the complex and obtain on-exchange data for each of the separate proteins (but not the complex) by performing Basic Protocol 1, omitting steps 19 to 22 (Fig. 20.9.1). On-exchange data provide information about the solvent accessibility of various surface regions on the proteins of interest. In addition, 24-hr on-exchange data are used to obtain back exchange correction factors for use in off-exchange data analysis.
2. For each off-exchange sample that is to be generated, prepare a tube of immobilized pepsin beads as described in Basic Protocol 1, step 3, making sure to wash at the optimal digestion pH. 3. Prepare several on-exchange samples of the receptor protein and the ligand protein as described in Basic Protocol 1, step 16, and incubate these samples at 25◦ C for a sufficient length of time to deuterate the surface of the protein (i.e., at least 10 min). 4. At the end of the on-exchange incubation period, combine a 6-µl aliquot of each receptor sample with a 6-µl aliquot of a ligand sample in the same 0.5-ml microcentrifuge tube, mix, and incubate 2 min at 25◦ C to allow complex formation. 5. Initiate off-exchange by adding 120 µl H2 O buffer to each sample tube, and then incubate at 25◦ C for times ranging from 1 to 30 min. A typical off-exchange experiment involves testing of five to eight different time points.
6. At the end of each off-exchange sample’s incubation period, quench the sample by plunging the tube into ice water and rapidly adding a predetermined amount of 2% trifluoroacetic acid (∼6 µl) to bring the pH of the sample to the optimal digestion pH. The appropriate amount of 2% TFA to add is determined ahead of time, as in Basic Protocol 1, step 2, but with H2 O buffer taking the place of 0.1% TFA in the sample containing the protein complex.
7. Using the pepsin beads prepared in step 2, perform protein digestion (under the optimal conditions identified in step 1) followed by mass spectrometric analysis of each off-exchange sample (see Basic Protocol 1, steps 4 to 6). 8. Perform control experiments in which each protein is analyzed separately for offexchange by repeating steps 2 to 7, using D2 O in place of the other protein in step 4.
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Figure 20.9.4 Sample off-exchange data. A mass envelope for a single peptide fragment at a single off-exchange time point is shown; however, data from several different time points should be collected to check for consistency in the data. (A) Nondeuterated peptide fragment from the uncomplexed protein. (B) Peptide fragment from the uncomplexed protein after on-exchange for 10 min. (C) Peptide fragment from the uncomplexed protein after on-exchange for 10 min followed by off-exchange for 10 min. Some residual deuteration is seen, due to the fact that some D2 O remains in the H2 O buffer used for off-exchange. (D) Peptide fragment from the complexed protein after on-exchange (performed on each component of the complex separately) for 10 min followed by off-exchange (performed on the bound complex) for 10 min. In the bound complex, the peptide retains deuterium throughout off-exchange, whereas in the uncomplexed state, it does not. This indicates that the peptide is probably part of the protein-protein interface region.
Comparison of receptor-ligand complex data with control data will reveal interfaces as sites at which deuterium (from the on-exchange portion of the experiment) is retained in the complex but not in the uncomplexed protein (Fig. 20.9.4).
9. Analyze off-exchange data as described in Support Protocol 2. ALTERNATE PROTOCOL
DETERMINATION OF THE SOLVENT ACCESSIBILITY OF A PROTEIN-PROTEIN INTERFACE VIA THE OFF-EXCHANGE METHOD USING AN AGAROSE-BOUND INTERACTION PARTNER When both proteins involved in the interaction of interest are large (>40 kDa each), it may be necessary to remove one of them from the mixture prior to mass spectrometric analysis so that good sequence coverage from the observed pepsin fragments can be obtained. In the example presented here, removal of an antibody prior to analysis of the epitope-containing protein is required for successful epitope mapping (Baerga-Ortiz et al., 2002). In such a case, the protein to be removed is covalently linked to agarose beads and eventually separated from its interaction partner by microcentrifugation.
H-Exchange MS to Study Protein-Protein Interactions
Additional Materials (also see Basic Protocol 1) Antibody of interest Protein G-agarose beads (Sigma)
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20 mM dimethylpimelimidate (Pierce Chemical) Epitope-containing protein of interest 1:1 mixture of 1-propanol/0.1% (v/v) trifluoroacetic acid (TFA; pH 2.25), chilled to 4◦ C Additional reagents and equipment for buffer exchange (UNIT 4.4) Perform on-exchange experiment for epitope-containing protein 1. Determine optimal digestion conditions and obtain on-exchange data for the epitopecontaining protein by performing Basic Protocol 1, omitting steps 12 to 13 and steps 18 to 22. Prepare antibody beads 2. Using the guidelines provided by the manufacturer, bind the antibody of interest to Protein G-agarose beads and perform cross-linking using 20 mM dimethylpimelimidate. Wash the beads, perform buffer exchange (UNIT 4.4), and then store the beads in 30-µl aliquots according to the manufacturer’s instructions. Immediately before use, exchange the antibody beads into deuterated buffer (10× protein buffer added to D2 O to a final buffer concentration of 1×). See Basic Protocol 1 for indications regarding appropriate 10× protein buffers.
3. Perform control experiments to ascertain nonspecific binding of the epitopecontaining protein to the Protein G-agarose beads and to determine the binding capacity of the antibody beads. These control experiments entail performing the procedure that follows (steps 4 to 10) using a number of different samples. To test for nonspecific binding, samples containing different amounts of Protein G beads (not linked to the antibody of interest) should be used, and to test for binding capacity, samples containing different amounts of antibodycross-linked beads should be used. The amount of beads that yields a reproducible mass spectrometer signal while minimizing nonspecific binding should be used in all subsequent experiments.
Perform off-exchange experiment 4. For each sample that is to be generated, prepare a tube of immobilized pepsin beads as described in Basic Protocol 1, step 3, making sure to wash at the optimal digestion pH. 5. In several separate 0.5-ml microcentrifuge tubes, add 1 µl of 100 pmol/µl epitopecontaining protein to 9 µl deuterated buffer and incubate 10 min at 25◦ C to allow deuteration of surface amides. Beginning at the same time, resuspend 30-µl aliquots of antibody beads in 270 µl deuterated buffer and incubate 10 min at 25◦ C. 6. Microcentrifuge the antibody bead suspensions 1 min at 14,000 × g, 25◦ C. Remove and discard 270 µl of supernatant from each, and add a 10-µl deuterated epitopecontaining protein sample to the remaining 30-µl aliquot of antibody beads. 7. Initiate off-exchange by diluting each sample with an additional 360 µl of H2 O. Immediately centrifuge each diluted sample 1 min at 14,000 × g, 25◦ C, and then discard 360 µl of supernatant. Resuspend each of the remaining 40-µl aliquots in 360 µl H2 O, and then incubate each for a different length of time (0 to 30 min) before centrifuging once more for 1 min at 14,000 × g, 25◦ C, and discarding as much supernatant as possible (leaving a volume of ∼30 µl, most of which should be antibody beads, in each tube). If the final centrifugation is performed immediately after resuspension of the antibody beads in H2 O, then the time from initiation of off-exchange to completion of the final centrifugation should be ∼2 min. During these 2 min, off-exchange of deuterons from the
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complex is occurring. Longer off-exchange time points are acquired by allowing additional time to pass between resuspension of the beads and centrifugation of the resulting sample.
8. For each sample, quench the reaction by adding 30 µl of a chilled 1:1 mixture of 1-propanol and 0.1% TFA (pH 2.25). Centrifuge each quenched sample 1 min at 14,000 × g, 4◦ C. The 1:1 1-propanol/0.1% TFA solution not only stops off-exchange but also elutes the binding protein from the antibody, which is covalently attached to the beads.
9. Add each supernatant from step 8 (30 µl) to a tube of pepsin beads prepared in step 4 and incubate under the optimal digestion conditions for the epitope-containing protein. Process all digestion samples as described in Basic Protocol 1, step 5. 10. Obtain mass spectra of all digestion samples as described in Basic Protocol 1, step 6, substeps a to e.
Perform control experiments and analyze data 11. Perform the control experiment (no beads) in the following way. a. In each of several 0.5-ml microcentrifuge tubes, add 1 µl of 100 pmol/µl binding protein to 9 µl deuterated buffer and incubate 10 min at 25◦ C. b. Initiate off-exchange by diluting each sample with an additional 90 µl of H2 O, and incubate each sample at 25◦ C for a length of time corresponding to one of the experimental off-exchange time points. When incubation is complete, transfer 10 µl of the sample to a tube containing 30 µl of a chilled 1:1 mixture of 1-propanol and 0.1% TFA (pH 2.25) to quench the reaction. c. Prepare an immobilized pepsin bead slurry. Add 100 µl of pepsin slurry to each quenched sample and incubate under the optimal digestion conditions for the epitope-containing protein. Process all digestion samples as described in Basic Protocol 1, step 5. d. Obtain mass spectra of all digestion samples as described in Basic Protocol 1, step 6, substeps a to e. In control samples, D2 O is diluted 10-fold in H2 O during off-exchange, while in experimental samples, D2 O is diluted by a factor of ∼100. Despite this difference, retention of deuterium in the complex as compared with the uncomplexed protein should be observed. Differences in D2 O dilution are corrected for during data analysis.
12. Analyze off-exchange data as described in Support Protocol 2. SUPPORT PROTOCOL 2
H-Exchange MS to Study Protein-Protein Interactions
ANALYSIS OF OFF-EXCHANGE DATA An analogous procedure to the one used to analyze on-exchange data (see Support Protocol 1), including corrections for back exchange and residual deuteration, should be used for off-exchange data. The kinetics of off-exchange typically fit best to either a biexponential or a triexponential model. The rate constants in the triexponential model correspond to amides that are not protected from solvent in the complex and remain susceptible to rapid exchange (rate constant set at 30 min−1 ; see Support Protocol 1), amides that are partially protected and thus undergo proton exchange at an intermediate rate (typically 0.1 to 0.5 min−1 ), and amides that are completely protected and thus undergo proton exchange at a slow rate (typically <0.04 min−1 ), respectively. The biexponential model is used when the triexponential model fails to lead to any improvement in the fitting of the experimental data. In such cases, typically, there are no sites at which exchange occurs at an intermediate rate. The triexponential equation is as follows:
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where D is the total number of deuterons that have undergone off-exchange by time t; Nfast is the number of deuterons exchanging at the fast rate, kfast ; Ninter is the number of deuterons exchanging at the intermediate rate, kintermed ; and Nslow is the number of deuterons exchanging at the slow rate, kslow .
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Matrix solution for MALDI-TOF 5 mg/ml α-cyano-4-hydroxycinnamic acid (Sigma; recrystallized once from ethanol) in 1:1:1 (v/v/v) acetonitrile/ethanol/0.1% (v/v) trifluoroacetic acid Adjust pH to 2.2 with 2% (v/v) trifluoroacetic acid Chill on ice for at least 2 hr prior to use Prepare fresh before each use COMMENTARY Background Information Identification of pepsin digest products It is important to determine the identity of the pepsin digest products in a reliable manner. Several bioinformatics tools and mass spectrometry data analysis software packages provide the option of entering peptide fragment masses along with a protein sequence and purport to identify pepsin digest peptides based on this information. These algorithms are only correct some 85% of the time, as they are based on the most common, but not the only, pepsin cleavage sites. Therefore, in order to determine peptide identities reliably, the approach described in Basic Protocol 1 must be used. Requirement for structural information It is important to note that the results of these types of experiments are extremely difficult to interpret in the absence of a structure of at least one of the proteins present in the complex. If a structure is available, then one can “map” the results onto the structure and observe where the changes in proton exchange behavior are occurring. If the segments found to be protected from solvent in the complex all coalesce to a single surface of the protein, then this surface is very likely to be the proteinprotein interface. If the segments found to be protected from solvent are on different parts of the molecule, with regions in-between that do not show protection, it is likely that a conformational change is occurring upon proteinprotein complex formation.
Critical Parameters and Troubleshooting Amide proton exchange rates are strongly dependent on pH, and therefore, control of
the pH throughout the exchange experiments is critical for obtaining reproducible, highquality data. Measurements of pH should be made on nondeuterated mock solutions to avoid electrode isotope effects. Even when deuteration is to be performed on 10-µl protein samples, prior to the first experiment, five 10-µl mock samples should be pooled so that the pH of the mock solution (and, thus, the pH of the actual deuteration solution) can be determined exactly. Measurements of solutions of >50 µl can be accurately made using an InLab 423 pH electrode (Mettler Instruments). In the authors’ experience, however, probes that purport to measure pH in smaller volumes of solution are very inaccurate. Deuterated buffer solutions should be prepared from 1 M stock buffers (in H2 O) in exactly the same manner as the mock solutions.
Anticipated Results Finding the solvent-inaccessible “core” of the interface The first step in finding the solventinaccessible “core” of the interface is to ensure that the complex remains 100% bound throughout the experiment. In one case, the authors studied the binding of two proteins for which the Kd was 120 nM. This interaction showed increasing amounts of deuterium retention at interface peptides as the ratio of one protein to the other was increased from 2.6:1 to 7:1, corresponding to an increase from 98.77% bound complex to 99.78% bound complex (Fig. 20.9.5). The change in the amount of bound complex was highly significant, because at the 2.6:1 ratio, the interacting proteins were apart for 1.5 sec during the 2 min offexchange period, compared with 0.3 seconds
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Figure 20.9.5 Plot showing the relationship between ligand concentration and percentage of receptor molecules bound to the ligand in a typical amide proton exchange experiment. To ensure that 100% of receptor molecules are bound, ligand-to-receptor ratios > 1:1 are required when Kd ≥ 10 nM. Theoretical curves for Kd = 1 nM (circles), Kd = 10 nM (squares), and Kd = 50 nM (triangles) are shown. Actual experimental data from an interaction for which Kd = 120 nM most closely resembled the Kd = 50 nM curve.
H-Exchange MS to Study Protein-Protein Interactions
when a 7:1 ratio was used, and this difference was enough to decrease the observed amount of deuteration. Thus, to ensure that the observed 1 H/2 H exchange rate was influenced by amides at the interface of the protein complex and not by complex dissociation, a ratio of 7:1 was used for the quantitative measurement of the exchange rate. The possibility of nonspecific binding arises with high ratios of ligand to receptor, and so comparisons should be made using lower ligand-to-receptor ratios as well; the same surface regions should show slowed exchange in the complex at ratios of 7:1 and 2.6:1, consistent with a lack of nonspecific binding. Once conditions are found in which the complex remains 100% bound during the experiment, it is sometimes possible to observe amide deuterons that do not off-exchange with solvent H2 O protons. Such peptides become deuterated quickly during on-exchange but then retain their deuterons throughout the offexchange period (Mandell et al., 2001). An important check on this result involves changing the pH, because the off-exchange reaction (at physiological pH) is a base-catalyzed reaction of OH− with an amide deuteron. Thus, by changing the pH from 7.4 to 8.4, the concen-
tration of OH− is increased 10-fold, and therefore, the rate of this base-catalyzed reaction is also increased 10-fold. If exchange occurs more rapidly at the higher pH, then the amides in question must be at least partially accessible to solvent. On the other hand, if these amides only undergo exchange when the complex dissociates, then pH will have no effect on the exchange rate, since this rate will be limited by the rate of dissociation. Examples of the kinetic plots obtained from a solvent-inaccessible region and a partially solvent-accessible region of the thrombin-thrombomodulin complex are shown in Figure 20.9.6. Differentiating conformational changes from interface protection It is often difficult to differentiate between changes in solvent accessibility that arise from conformational changes and those that arise from interface protection. Sometimes when proteins interact, a region of the protein becomes more accessible to solvent in the complex, and this can readily be interpreted as resulting from a conformational change. An example of this type of result is seen in the regulatory subunit of protein kinase A (Anand et al., 2002).
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Figure 20.9.6 Kinetic plots of amide deuteration after data analysis to correct for back exchange and residual side-chain deuteration. (A) On-exchange data for a solvent-accessible region of an uncomplexed protein at two different pH values, 6.5 (circles) and 7.9 (squares). (B) Off-exchange data for the same solvent-accessible region in the uncomplexed protein (squares) and in the bound complex (triangles). The data obtained from the uncomplexed protein show that off-exchange occurs as rapidly as on-exchange did, and the difference between the two curves indicates that amides are protected from off-exchange in the complex. Identical data were obtained at both pH 6.5 and pH 7.9. (C) On-exchange data for a partially solvent-accessible region of the uncomplexed protein at two different pH values, 6.5 (circles) and 7.9 (squares). (D) Off-exchange data for the same partially accessible region indicate that at pH 6.5, some deuterium is retained in the bound complex (triangles), but only for a short time. No deuterium retention was observed at pH 7.9 (data not shown).
For regions that show “protection” from amide proton exchange in the bound complex, it is important to map the observed peptide fragments back onto the structure of the protein. This will reveal whether the region corresponding to the peptides in question is on the protein’s surface. In the example of thrombomodulin, one of the peptide fragments found to be fully protected in the complex was completely exposed to solvent and susceptible to rapid on-exchange in the control experiment involving the uncomplexed protein. Such data
are the most rigorous indicator of the location of the interface that can be obtained from amide proton exchange experiments, and regions that exhibit such behavior can unambiguously be assigned to the interface. If a peptide fragment exhibiting protection in the complex runs from an interior region of the uncomplexed protein to a surface region, then assignment of the fragment to the interface can sometimes be made by interpreting the time courses of on-exchange and offexchange in the uncomplexed protein at two
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different pH values (for example, 7.4 and 8.4). If the off-exchange kinetics are insensitive to pH, then it is likely that the peptide is located at the interface. However, if protection is only observed when the amides are deuterated at the higher of the two pH values, and not at the lower pH, then it is likely that the amides being deuterated at the higher pH are those located on the interior. This would indicate the possibility that a conformational change is occurring in the complex, since the amides in question are not on the surface of the protein. Finally, if a region incorporates deuterium slowly at both pH levels, any observed protection in the complex should be interpreted with caution.
Time Considerations Because amide proton exchange experiments are time sensitive, it is important to set up all materials (e.g., chilled solutions, tubes) ahead of time so that once deuteration is begun, samples can be processed rapidly and consistently. It is best to do one experiment at a time, as this leads to better temporal reproducibility. The number of separate experiments required and the range of deuteration times explored will depend on the protein-protein interaction of interest, but five to six time points (range, 1 to 20 min) typically are sufficient for sampling the changes that occur on a protein’s surface. The time required for a typical experiment varies greatly depending on the instrumentation available and the proteins being studied. In particular, there are large variations in the length of time required to accurately determine the identities of the pepsin digest products, which can range from a few days to weeks. The rest of the protocol, including planning (1 day), pilot studies (1 day), execution of the actual experiment (2 to 3 days), and data analysis (2 to 3 days), typically takes 1 to 2 weeks.
Literature Cited Anand, G.S., Hughes, C.A., Jones, J.M., Taylor, S.S., and Komives, E.A. 2002. Amide H/2 H exchange reveals communication between the cAMP- and catalytic subunit-binding sites in the regulatory subunit of protein kinase A. J. Mol. Biol. 323:377-386.
H-Exchange MS to Study Protein-Protein Interactions
Anand, G.S., Law, D., Mandell, J.G., Snead, A.N., Tsigelny, I., Taylor, S.S., Ten Eyck, L., and Komives, E.A. 2003. Identification of the protein kinase A regulatory RIα-catalytic subunit interface by amide H/2 H exchange and protein docking. Proc. Natl. Acad. Sci. U.S.A. 100:1326413269.
Baerga-Ortiz, A., Bergqvist, S.P., Mandell, J.G., and Komives, E.A. 2004. Two different proteins that compete for binding to thrombin have opposite kinetic and thermodynamic profiles. Protein Sci. 13:166-176. Baerga-Ortiz, A., Hughes, C.A., Mandell, J.G., and Komives, E.A. 2002. Epitope mapping of a monoclonal antibody against human thrombin by H/D-exchange mass spectrometry reveals selection of a diverse sequence in a highly conserved protein. Protein Sci. 11:1300-1308. Bai, Y., Milne, J.S., Mayne, L., and Englander, S.W. 1993. Primary structure effects on peptide group hydrogen exchange. Proteins 17:75-86. Dharmasiri, K. and Smith, D.L. 1996. Mass spectrometric determination of isotopic exchange rates of amide hydrogens located on the surfaces of proteins. Anal. Chem. 68:2340-2344. Englander, S. and Mayne, L. 1992. Protein folding studied using hydrogen-exchange labeling and two-dimensional NMR. Annu. Rev. Biophys. Biomol. Struct. 21:243-265. Englander, S., Mayne, L., Bai, Y., and Sosnick, T. 1997. Hydrogen exchange: The modern legacy of Linderstrøm-Lang. Protein Sci. 6:1101-1109. Gemmecker, G., Jahnke, W., and Kessler, H. 1993. Measurement of fast proton exchange rates in isotopically labeled compounds. J. Am. Chem. Soc. 115:11620-11621. Mandell, J.G., Baerga-Ortiz, A., Akashi, S., Takio, K., and Komives, E.A. 2001. Solvent accessibility of the thrombin-thrombomodulin interface. J. Mol. Biol. 306:575-589. Mandell, J.G., Falick, A.M., and Komives, E.A. 1998a. Identification of protein-protein interfaces by decreased amide proton solvent accessibility. Proc. Natl. Acad. Sci. U.S.A. 95:1470514710. Mandell, J.G., Falick, A.M., and Komives, E.A. 1998b. Measurement of amide hydrogen exchange by MALDI-TOF mass spectrometry. Anal. Chem. 70:3987-3995. Rosa, J.J. and Richards, F.M. 1979. An experimental procedure for increasing the structural resolution of chemical hydrogen-exchange measurements on proteins: Application to ribonuclease S peptide. J. Mol. Biol. 133:399-416. Zhang, Z., Post, C.B., and Smith, D.L. 1996. Amide hydrogen exchange determined by mass spectrometry: Application to rabbit muscle aldolase. Biochemistry 35:779-791. Zhang, Z. and Smith, D.L. 1993. Determination of amide hydrogen exchange by mass spectrometry: A new tool for protein structure elucidation. Protein Sci. 2:522-531.
Contributed by Jeffrey G. Mandell, Abel Baerga-Ortiz, Carrie H. Croy, Arnold M. Falick, and Elizabeth A. Komives University of California, San Diego La Jolla, California
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Circular Dichroism to Study Protein Interactions
UNIT 20.10
Proteins play a wide variety of functional roles in biological systems, including catalysis, transport, regulation, defense, and maintenance of cell architecture. All of these functions depend on specific interactions between proteins and other molecules. These interactions must be characterized in structural and energetic terms to understand the functions of proteins in a cellular context. An initial goal might be merely to demonstrate that a particular protein interacts with a particular ligand and perhaps explore the specificity of this interaction by surveying a range of other potential ligands. If an interaction is demonstrated, the high-resolution structural methods of X-ray crystallography or nuclear magnetic resonance (NMR) could be used to provide atomic-level structural information about the interaction. However, a second, and perhaps more important, goal is to explore the energetics of the interaction, in both thermodynamic and kinetic terms, and for this, a range of other methods can be employed. There are two principal types of these latter methods. The first type includes those that involve measurement of a change in some parameter of the protein (or its interacting partner or both) on complex formation. This parameter is often a spectroscopic signal such as absorption, fluorescence (UNIT 20.8), or circular dichroism (CD), but could also be a change in the stability of the protein towards thermal or chemical denaturation (Greenfield, 2004a) or in the enthalpy of the system, which could be detected by isothermal titration calorimetry (Cooper and Johnson, 1994). The second type of method is based on exploiting the change in mass, which occurs on complex formation, e.g., using equilibrium dialysis or ultracentrifugation to separate the complex from one or more of the isolated components (van Holde et al., 2006), or surface plasmon resonance, which detects the change in the intensity of reflection of polarized light of the immobilized complex (McDonnell, 2001). This unit focuses on the application of CD to study the interactions of proteins. A comprehensive discussion of the assessment of CD, its advantages and disadvantages over other spectroscopic techniques, and factors to consider in determining the suitability of CD for studying the protein interaction in question is given in Strategic Planning. Protocols follow, which describe how to perform a CD titration to monitor binding (see Basic Protocol 1) and the subsequent analysis of CD data to determine binding constants (see Basic Protocol 2). A series of support protocols provide important ancillary methods and considerations: analysis of far-UV CD data to determine secondary structure content (see Support Protocol 1); assessment of stability of the protein during titration (see Support Protocol 2); choice of buffer and solvent system (see Support Protocol 3); determination of protein concentration (see Support Protocol 4); calibration of CD instrument (see Support Protocol 5); and choice of instrument parameters (see Support Protocol 6). Finally, the Commentary section includes some background information on CD as a spectroscopic technique and the units used.
STRATEGIC PLANNING As with other spectroscopic techniques, to apply CD to study the binding of a protein (P) to another species (ligand, L), there must be a change in signal to distinguish the complex (PL) from the isolated components P and L. This is discussed in the following section. Contributed by Sharon M. Kelly and Nicholas C. Price Current Protocols in Protein Science (2006) 20.10.1-20.10.18 C 2006 by John Wiley & Sons, Inc. Copyright
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Assessment of CD in the Study of Protein-Ligand Interactions In the case of two proteins interacting, it is generally the case that substantial changes in secondary structural components do not occur upon complex formation. Since the isolated proteins are each likely to have significant CD signals in the far-UV region, this can make any changes in the CD signals in this region that occur upon complex formation relatively small and thus difficult to measure accurately. However, it is possible to use CD to measure a change in stability of the protein upon complex formation as a result of thermal or chemical denaturation, even when there is little or no change in its spectrum. The changes in stability can be analyzed to yield the free energy (and hence equilibrium constant) for the interaction. This approach has been extensively used to characterize a number of interactions between different muscle proteins (Greenfield, 2004a). However, when two proteins interact, there are more likely to be significant changes in the tertiary structures of one or both components upon complex formation; this should lead to significant changes in the near-UV CD spectrum. For example, the association of insulin monomers to form dimers and then hexamers is accompanied by changes in the environment of surface Tyr side chains, so this process can be readily monitored by near-UV CD, although there is no significant change in the far-UV CD spectrum (Wood et al., 1975). This type of approach is aided if only one of the proteins involved has aromatic amino acid side chains or a particular non-protein cofactor, since these will provide convenient signals for monitoring the association. The latter case occurs in the reductase and P450 domains of the flavocytochrome BM3 from Bacillus megaterium, where the covalent linking of the two domains in the intact protein leads to changes in the environment of the heme cofactor in the P450 domain, which can be detected by CD (Munro et al., 1994). There is no detectable change in the secondary structure of the individual domains when they are fused in the intact protein, implying that the structural changes are confined to the local environment of the heme cofactor. CD is widely applicable to study protein–small-molecule ligand interactions and interpretation of data is easier when absorption bands of the protein and the ligand do not overlap. It is possible to monitor the changes in the protein (secondary and/or tertiary structure) or the ligand upon complex formation. Typically, a small-molecule ligand (such as a drug) will possess either no or a very small CD signal when free in solution. However, upon binding to a defined site on a protein, it will acquire chirality and hence display a measurable CD signal, providing a convenient method to monitor complex formation. It is important to note that CD is much more sensitive than absorption spectroscopy in this regard, since there is usually only a small difference in the absorption spectrum of the ligand upon binding (Munro et al., 1996).
CD to Study Protein Interactions
From the intensity and appearance of the CD signals, it may be possible to draw conclusions about the changes in conformation of the protein or environment of the components upon complex formation. For example, the addition of Ca2+ ions to troponin-C leads to a marked increase in its helical content, and the change in ellipticity at 222 nm (an increase of ∼65%) provides a convenient means of monitoring the formation of the complex (Smith et al., 1994). Addition of MoO4 2− ions to the ModE binding protein leads to a marked change in the CD spectrum in the Trp region of the spectrum (see Fig. 20.10.1A) and the changes in ellipticity at 285 nm can be used to monitor the association; there is little change in the far-UV CD spectrum of ModE upon binding MoO4 2− (see Fig. 20.10.1B; Anderson et al., 1997). Site-directed mutagenesis studies have shown that the changes in the near-UV CD spectrum arise principally from changes in the environment of the Trp 186 residue in each subunit of the dimeric protein (Boxer et al., 2004). It should be noted that complex formation can lead to a marked change in spectroscopic signal even when the estimated contents of helix and sheet are essentially unchanged, as in the case of Ca2+ binding to the adenylyl cyclase toxin (CyaA) of Bordetella pertussis,
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Figure 20.10.1 CD spectra of the molybdate-sensing protein ModE from E. coli in the absence (solid line) and presence (dashed line) of 1 mM molybdate. Panels A and B represent the near-UV and far-UV CD spectra, respectively. The data are adapted from Anderson et al. (1997) and Boxer et al. (2004).
where it appears that the spectral changes (Rose et al., 1995) in the far-UV CD spectrum (Fig. 20.10.2) may reflect alterations in the orientation of helices with respect to each other rather than the actual content of helices (Cheung et al., unpub. observ.). In this case, the binding of Ca2+ could be monitored by changes in the ellipticity at 220 nm or 200 nm; there is a proportionally greater change in the latter signal (150% compared with 30%). One particular class of protein-ligand interactions for which CD is well suited is proteinnucleic acid interactions, since at equivalent molar ratios, nucleic acids tend to make only a small contribution to signals in the far-UV region, where the peptide bonds of the protein are dominant. In addition, proteins make only a small contribution in the near-UV region where the major signals are from the nucleic acids. Thus, it is possible to assess changes in secondary structure of the protein component as well as changes
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Figure 20.10.2 Far-UV CD spectra of the adenylyl cyclase toxin of Bordetella pertussis (CyaA) in the absence (solid line) and presence (dashed line) of 1 mM calcium chloride. The data are adapted from Cheung et al. (unpub. observ.).
in the nucleic acid component (such as the content of duplex) upon complex formation. Examples include the binding of DNA to the transcription factor NF-κB (Matthews et al., 1995) and the binding of DNA to DNA methyltransferase systems (Baldwin et al., 1994; Hornby et al., 1994).
Advantages and disadvantages of CD compared with other spectroscopic techniques The main advantages of CD for studying protein-ligand interactions are that the technique can be used to study a wide range of interactions since changes in signal from either the protein or the ligand (or both) are likely to occur upon complex formation. In many cases, it is possible to interpret changes in the CD signals in structural terms, e.g., in terms of the secondary structure of the protein (far-UV) or the integrity of the ligand-binding site. In addition, the interactions can be studied under a wide range of experimental conditions such as protein concentration, pH, temperature, ionic strength, etc. In more specialized applications, it is possible to measure the rates of complex formation using time-resolved CD, with a dead time generally in the range 1 to 5 msec for stopped-flow CD instruments. Suppliers of stopped-flow CD instruments include BioLogic and Applied Photophysics. The principal disadvantages of CD are the relatively large amounts of material required, particularly for near-UV and visible CD studies (typically, ∼1 mg protein per titration), and the fact that complex formation between two protein species might give rise to relatively small changes in the far-UV CD. Although it may be possible to measure changes in stability upon complex formation and hence derive the appropriate free energy values, this is a much more demanding task than a simple spectroscopic titration. In addition, because the CD signals from biological systems are relatively small, experiments must be carefully designed to ensure that reliable data are obtained.
Establishing the suitability of CD for studying protein interactions CD to Study Protein Interactions
Before undertaking detailed studies of a protein-ligand interaction, it is important to perform a number of preliminary experiments.
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First, the CD spectra of the isolated components (P and L) and the spectrum of the protein in the presence of ligand (the PL complex) should be recorded. These spectra should be corrected for any contribution of solvent by running suitable blank spectra. From the corrected spectra (that of PL compared with the sum of those of P and L), it is possible to decide whether a suitable spectral change has occurred and what wavelength region is best for monitoring the interaction. Ideally, the change in signal at the chosen wavelength should be at least 25%. It is important to assess the extent to which the ligand might contribute to the CD signal at this wavelength. Some conclusions might be drawn about structural changes that occur upon complex formation. If no significant change in the spectrum occurs after mixing the components, it does not necessarily mean that an interaction has not occurred. In such a case, it might be possible to measure a change in stability towards denaturation, as was done for the interaction between fragments of the muscle proteins E-tropomodulin and α-tropomyosin (Greenfield, 2004a). Second, the range of protein and ligand concentrations over which complex formation occurs should be assessed by one or more preliminary titration experiments so that a more detailed study of the interaction can be planned. CD spectra can be acquired over a wide range of protein concentrations, depending on cell pathlength and spectral ranges, giving an extra degree of flexibility in designing experiments. Third, the stability of the protein and the ligand under the conditions of the binding experiment should be established (UNIT 20.8; and Support Protocol 2).
CD TITRATION TO MONITOR BINDING This protocol describes how a titration experiment should be performed in terms of the component solutions (protein, ligand, etc.) and the correct choice of experimental parameters to obtain reliable data. Further information is given in Support Protocols 1 through 6.
BASIC PROTOCOL 1
Solutions Required The protein studied should be of suitable purity (>95% homogenous as determined by SDS-PAGE) and dissolved in a suitable buffer system to yield a clear solution, to avoid problems due to light scattering (UNIT 7.6; Kelly et al., 2005). The buffer should be appropriate for the pH range being studied (i.e., used at a pH within one unit of the relevant pKa ), should not have an unacceptably high absorbance (i.e., the absorbance should be ≤1.0) over the range of wavelengths studied; this is especially important in the far-UV region (see Support Protocol 3). Suitable buffers would include 10 mM sodium phosphate, borate, or Tris·Cl. The concentration of the protein solution should be determined by a reliable method (see Support Protocol 4). In addition, it is important that the conditions maintain the protein in a native state, although this can be difficult to prove unless there is a validated assay for its biological activity (see Support Protocol 2). The ligand studied should ideally be dissolved in the same buffer as the protein. However, some organic ligands may not be sufficiently soluble to allow preparation of a concentrated stock solution, so it may be necessary to dissolve them initially in a water-miscible solvent such as ethanol, acetonitrile, dimethylformamide, or dimethylsulfoxide, before dilution into the aqueous buffer system. The ligand concentration should be accurately determined (usually by spectroscopy). The effects of any residual organic solvents added during the titration on the protein and any interference caused to the CD spectrum by high absorbance must be assessed in separate experiments. Quantitation of Protein Interactions
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Instrument There are three main suppliers of CD spectrometers (spectropolarimeters), namely AVIV Associates, Jasco, and Applied Photophysics. Of these, the Applied Photophysics instruments are particularly suitable for fast kinetic studies using CD. These are described in UNIT 7.6. A supply of high purity nitrogen gas is required to displace oxygen to prevent damage to the optics by the build-up of ozone and to reduce absorbance by oxygen at wavelengths <200 nm. Details of suitable nitrogen sources are given in UNIT 7.6 and Kelly et al. (2005). After switching the instrument on, the spectropolarimeter and lamp should be allowed to warm up for 30 min to achieve stability. Calibration of the wavelength and signal amplitude of the spectropolarimeter should be checked regularly. Details of the procedures are given in UNIT 7.6 and Kelly et al. (2005), and summarized in Support Protocol 5. The correct choice of instrument parameters (bandwidth, time constant/acquisition time, scan speed, number of accumulations) have been described by UNIT 7.6 and Kelly et al. (2005), and are summarized in Support Protocol 6.
Cells The cells used for CD should be made of fused quartz and designated as “strain-free for polarimetry” to avoid optical artifacts. The major suppliers of cells for CD are Hellma and Starna. Different types of cells are available, namely cylindrical, rectangular, and demountable, which will provide a range of pathlengths and geometries (UNIT 7.6). For far-UV, it is often necessary to use cells of short pathlength (<1 mm), so as to prevent the absorbance becoming excessive. Demountable cells (which consist essentially of two quartz plates, one of which is machined out to the required depth) can provide very short pathlengths down to as little as 10 µm, but can be difficult to handle and thermostat efficiently. Although only a very small volume of sample (<0.1 ml) is required, it is impossible to recover any substantial fraction of this after recording the CD spectrum. Cylindrical cells (with two holes to allow filling and emptying with a microsyringe) provide short pathlengths (down to 0.1 mm) without requiring a large sample volume and it is possible to recover most of the sample. However, it is difficult to mix solutions efficiently in a cylindrical cell and there can be problems with cleaning such cells especially when sticky protein deposits have formed. Rectangular cells generally provide longer pathlengths and require larger sample volumes, but are more suitable for mixing solutions during titrations. For accurate work, it is important to check the actual pathlengths of cells; this can be done by interferometry or by measurements of the absorbance of a standard solution of potassium chromate (see Kelly et al., 2005). The extinction coefficient at 372 nm of potassium chromate in 0.05 M KOH is 4830 M−1 cm−1 ; thus, a 0.01 M (1.942 g/liter) solution in a cell of pathlength 0.1 mm has an absorbance of 0.483 (Haupt, 1952). The cleaning and general care of cells is described in detail in (2005).
CD to Study Protein Interactions
UNIT 7.6
and Kelly et al.
Choice of Concentration of Protein The choice of the concentration of protein will depend on a number of factors, including how much protein is available and strength/avidity of the protein-ligand interaction. When a suitable protein concentration has been chosen, the appropriate pathlength can
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Table 20.10.1 Absorbance Values for Proteins in the UVa
Wavelength (nm)
Absorbance (1 mg/ml solution in a 1-cm pathlength cell)
190
65
193
68
200
45
205
32
210
21
215
15
220
11
230
5
280
≈1b
310
<0.05c
320
0c
a The absorbance values shown are to be regarded as typical for proteins, rather than exact values. b This value depends on the Trp and Tyr content of the protein; values for A 280 are found to range
from 0 to 4. c Assuming that no cofactor (e.g., FAD, pyridoxal-5 -phosphate) absorbing in this range is present
and that scattering due to aggregates is not significant.
be determined using the criterion that the maximum absorbance should not exceed ∼1.0. The absorbance values for a typical protein are shown in Table 20.10.1; at wavelengths <230 nm, the absorption is predominantly due to peptide bonds, which occur at an essentially constant frequency in most proteins (Kelly et al., 2005). Thus, for a 1 mg/ml solution of protein, a cell pathlength of ≤0.01 cm (0.1 mm) is required to bring the A190 into the required range. If the protein concentration is 0.1 mg/ml, the pathlength should be <1 mm. In the near-UV and visible regions, the protein concentration would be typically 0.5 to 1.0 mg/ml in a cell of pathlength 5 or 10 mm. The strength of the interaction may well play a part in deciding a suitable protein concentration for the titration (see Basic Protocol 2).
Collection of Data Collecting the data for the titration of protein and ligand usually involves one of three types of experimental approach. In each case, it is important to allow suitable time for equilibration at each concentration of added ligand (usually at least 1 min), before the signal is recorded. For many purposes it is sufficient to record the CD signal at the chosen wavelength (this is best done by averaging the signal over a period of 1 min), although in some cases, it may be important to record a complete spectrum after each addition. The latter approach would allow a more complete description of the structural changes in the protein, which occur as a consequence of ligand binding, but would significantly add to the time required for data collection and may be difficult if the protein is not stable over long periods (see Support Protocol 2). If the ligand makes a contribution to the signal at the chosen wavelength, it is necessary to correct for this in the manner described in UNIT 20.8. (1) Small aliquots (typically ≤1% of the total volume) of a stock solution of ligand are added to a solution of protein and the signal recorded after each addition. During this process, the concentration of the protein should not change significantly (the overall
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change should be <10%) and the observed signal can be corrected for this prior to analysis. It may be necessary to use two or more stock solutions of different concentration to achieve suitable coverage of the saturation curve. For example, if the dissociation constant of the PL complex is 100 µM, ligand concentrations of 12.5, 25, 50, 75, 100, 150, 200, 300, 400, and 500 µM would cover the range from 11% to 83% saturation. These could be achieved using ligand stock solutions of 2.5 mM (12.5 to 100 µM), 5 mM (150 and 200 µM), and 10 mM (300, 400, and 500 µM). In general, the ligand concentrations used in a titration should range from ∼0.2 times the Kd to 5 times the Kd . The titration requires use of a cell that will allow suitable mixing (e.g., with a small magnetic stirrer). Suitable motor-driven syringes are commercially available, e.g., the BioLogic device can be readily interfaced with a Jasco spectropolarimeter. (2) A series of solutions are prepared with the protein concentration constant and the concentration of ligand varied so as to span the titration range, as described above. While this approach avoids the dilution of protein, which occurs as above, it is much less economical in terms of the amount of protein required and suffers from the disadvantage that there are random errors caused by multiple pipettings in preparing the solutions. (3) Two solutions, one containing protein alone (experiment starting point), and the other containing the same concentration of protein effectively saturated with ligand (experiment ending point), are mixed in a systematic fashion. Thus, initially the CD signal or CD spectrum of the protein alone is recorded, then a defined volume (typically 20%) of this solution is withdrawn and replaced with the same volume of the ligand-saturated solution and the CD signal or spectrum subsequently recorded. A further defined volume of this solution is withdrawn and replaced by the end-point solution before recording the new signal or spectrum. By increasing the proportion of the ligand-saturated (end-point) solution in a suitable manner (with no change in the protein concentration), the saturation curve can be obtained. Programmable mixing devices, which allow the user to specify the required points on the saturation curve, may be supplied by spectropolarimeter manufacturers, including Jasco and AVIV. These devices also allow the data to be analyzed to determine the binding constants. The choice of method will depend on several factors including the availability of equipment, the amount of material available, and any constraints imposed by the sensitivity of CD technique and the appropriate concentration of protein required to obtain suitable data for analysis of the binding curve (see Basic Protocol 2). BASIC PROTOCOL 2
ANALYSIS OF CD DATA TO DETERMINE BINDING CONSTANTS General Considerations Many of the key principles of analyzing titration data are analogous to those described for the analysis of fluorescence data (UNIT 20.8).
CD to Study Protein Interactions
As with all spectroscopic techniques, it is necessary to assume that the change in signal observed is proportional to the degree of saturation of binding sites on the protein. While this is likely to be true in most cases, there could be situations involving multiple binding sites that interact cooperatively in which the change in signal does not accurately reflect the saturation of sites, especially where the spectroscopic signal reflects a widespread conformational change in the protein, rather than a more local effect. In such cases, it is necessary to use a technique for measuring binding based on physical separation, such as equilibrium dialysis (van Holde et al., 2006; APPENDIX 5A) with a signal such as radioactivity to measure ligand concentrations.
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To analyze the binding data (e.g., using non-linear curve fitting or a Scatchard plot) to obtain reliable values for both the dissociation constant (Kd ) and the number of binding sites (n), it is necessary to obtain data points across an appropriate proportion of the saturation curve, ideally between 15% and 85% saturation. CD is useful for examining interactions within the range of dissociation constants from 1 µM to 1 mM (i.e., association constants in the range 106 to 103 M−1 ). This is a convenient range, especially for most enzyme-substrate or enzyme-inhibitor interactions, and for many interactions involved in transport or regulatory roles of proteins. For weak interactions (i.e., Kd >1 mM) it can be difficult to observe changes in the signal of the protein without using very high ligand concentrations, which may lead to problems with high absorbance. Although it may be possible to obtain a reasonable estimate of the Kd , it is very difficult to establish the number of binding sites under these conditions. For strong interactions (i.e., Kd <1 µM), it is difficult to obtain a suitable balance between free and bound protein without using low (<1 µM) protein concentrations; these conditions may produce signals below the limit of detection by the CD technique. In these situations, it is usually possible to obtain the number of binding sites (n) by performing titrations at high protein concentrations (i.e., well above the Kd ), but it is difficult to measure the Kd accurately.
Analysis using different binding models Consider the titration of a protein P, with ligand L (L could also be a second protein). The initial concentration of P is designated [P0 ]; at any point in the titration, the total concentration of L added is [Lt ] and the change in CD signal is θ. For the interaction, the dissociation constant is defined as Kd (i.e., Kd = ([P] [L])/[PL]), where [P] and [L] are the concentrations of free protein and ligand, respectively. The maximum change in CD signal, i.e., when P is saturated with ligand, is θmax .
Weak binding Weak binding (where essentially all L added can be considered as being free) would apply when [P0 ] in molar terms is <0.2 Kd . For example, if a 30-kDa protein were present at a concentration of 1 mg/ml (33 µM), weak binding refers to a Kd >165 µM. If the protein were present at 0.2 mg/ml, weak binding would refer to a Kd >33 µM. As ligand is added to protein, relatively little of the ligand is bound and, hence, it is possible to assume that the free ligand concentration equals the total ligand concentration. Hyperbolic binding occurs when a single molecule of ligand binds to a protein (or for a protein with multiple binding sites for a ligand, when the sites act independently of each other). Equation 20.10.1 (which is formally analogous to the Michaelis-Menten equation for enzyme kinetics or the Langmuir isotherm for binding) holds in such situations:
Equation 20.10.1
The data obtained, i.e., values of θ as a function of Lt , can be analyzed by nonlinear curve-fitting procedures (e.g., programs in SigmaPlot or Microcal Origin software) to yield values of K and θmax , and to give standard errors of these estimates. The result of nonlinear curve-fitting is illustrated in Figure 20.10.3A. Panel A illustrates nonlinear fitting to Equation 20.10.1.
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A
16 14 12
∆θ (mdeg)
10 8 6 4 2 0 0
B
20
40
60
100
80
120
140
160
180
[Lt] (µM) 0.50 0.45 0.40
∆θ/[Lt] (mdeg/µM)
0.35 0.30 0.25 0.20 0.15 0.10 0.05 0.00 0
2
4
6
8
10
12
14
16
∆θ (mdeg)
Figure 20.10.3 Analysis of binding data. Panel A illustrates nonlinear fitting to Equation 20.10.1. The resultant values are θmax = 15.97 mdeg (standard error 0.03 mdeg), and Kd = 34.9 µM (standard error 0.2 µM). Panel B shows a Scatchard plot according to Equation 20.10.2. In this plot, the slope = −1/Kd ; hence, Kd can be calculated to be 35 µM. The intercept on the y-axis = θmax ; hence, θmax = 16 mdeg. Whereas the nonlinear fitting is useful for indicating the errors in the estimates of the binding parameters, the Scatchard plot can indicate whether there are systematic deviations from the hyperbolic binding model. In this example, the ligand concentration is assumed to be much larger than that of the protein, so that [L]free can be set equal to [Lt ] in the plots.
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Alternatively, the data can be analyzed by a linear transformation of the hyperbolic equation, such as the Scatchard plot:
Equation 20.10.2
A plot of θ/[Lt ] against θ gives a straight line with a slope = −1/Kd and an x-axis intercept of θmax (Fig. 20.10.3B). The nonlinear analysis approach is very widely used to analyze binding data, because of the ready availability of appropriate programs. However, it is still valuable to employ a linear transformation method such as the Scatchard plot to verify that the data conform to the hyperbolic binding equation (i.e., that the deviations of the experimental data from the theoretical model are random and show no systematic trend). Cooperative binding occurs where the protein has several binding sites for L and the binding of ligand at one site influences the binding at other sites. In these cases, the Hill equation (Equation 20.10.3), where h is the Hill coefficient, is often used to fit the data:
Equation 20.10.3
The data (θ as a function of [Lt ]) are best analyzed by nonlinear curve-fitting procedures (programs in SigmaPlot or Microcal Origin) to give estimates of the parameters K, h, and θmax . Alternatively, if θmax can be estimated, it is possible to analyze the data graphically in terms of a Hill plot (a plot of log (θ/(θmax − θ)) versus log [Lt ]; Fig. 20.10.4), in which the slope = h, and the point of intersection with the x-axis = (log K)/h. In this case, K is not a true dissociation constant but is related to the concentration
Figure 20.10.4 Analysis of specimen binding data using the Hill plot according to Equation 20.10.3. The slope of the plot is the Hill coefficient; this can be calculated to be 2.5, indicating a significant degree of cooperativity in ligand binding. From the point of intersection of line with the x-axis, the value of K can be calculated to be 5500 µM (Equation 20.10.4).
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of L at which 50% of the binding sites are saturated ([Lt ]0.5 ), by Equation 20.10.4:
Equation 20.10.4
Constructing a Hill plot is a valuable approach to confirm the applicability of the Hill equation (Eq. 20.10.3) to the analysis of the binding data. The value of h provides a measure of the degree of cooperativity in the system. When h = 1, the binding is hyperbolic in nature, i.e., the sites behave independently of each other. When h > 1, the binding is said to display positive cooperativity, i.e., binding at the first site strengthens the subsequent binding of ligand to other sites. If h < 1, the system is said to show negative cooperativity. The Hill coefficient can take values up to n, the number of binding sites; if h = n, there is extreme cooperativity and essentially there is an equilibrium between P and PLn only.
Tight binding Tight binding (where a significant fraction of added ligand is bound to protein) would apply when [P0 ] is ≥0.2 Kd for the interaction. Thus, in the case of a 30-kDa protein at 1 mg/ml (33 µM), strong binding would refer to a Kd of ≤165 µM; at 0.2 mg/ml, strong binding would refer to a Kd of ≤33 µM. As ligand is added to the protein, a significant fraction of the ligand is bound so the assumption can no longer be made that all the added ligand can be regarded as free ligand. It is necessary to solve a quadratic equation (Eqn. 20.10.5) describing the change in CD signal (θ) at a given added concentration of ligand ([Lt ]) to determine the dissociation constant (Kd ) and the maximum change in CD signal (θmax ) for the interaction.
Equation 20.10.5
This equation is formally analogous to those described in UNIT 20.8 for the analysis of fluorescence data and by Greenfield (2004a) for the analysis of CD data. (It should be noted that the equation (Equation 9) given by Greenfield is written in terms of the association constant for the interaction rather than the dissociation constant as in the present case; the dissociation constant is merely the reciprocal of the association constant). The data can be analyzed in terms of Equation 20.10.5 by fitting programs available within Sigma Plot or Microcal Origin, to give estimates of θmax and Kd (UNIT 20.8). It is necessary to enter initial trial values of these parameters so the program can determine the best-fit values with the standard errors of the estimates. Errors in the estimates of the parameters Kd and n can be taken as an indication of the quality of the experimental data, and the pattern of residuals (i.e., the pattern of differences between the calculated and experimental degrees of saturation as a function of ligand concentration) can be taken as a measure of the appropriateness of the binding model used. Ideally, the error in Kd should be <10% of its actual value and the residuals should be distributed in a random fashion about the zero line. CD to Study Protein Interactions
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ANALYSIS OF FAR-UV CD DATA TO DETERMINE SECONDARY STRUCTURE CONTENT
SUPPORT PROTOCOL 1
A number of procedures have been described to use the data from far-UV CD experiments to determine estimates of the secondary structure composition of proteins. Most of these methods employ basis datasets consisting of the CD spectra of proteins of various fold classes whose structures have been solved to high resolution by X-ray crystallography. Details of these methods are given in review articles by Sreerama and Woody (2004) and Greenfield (2004b). An online server DICHROWEB (see Internet Resources) has been developed, hosted by Birkbeck College, University of London, which allows CD data to be entered in a variety of formats and analyzed by the most useful algorithms with a choice of databases (Lobley et al., 2002; Whitmore and Wallace, 2004). The following points should be kept in mind when judging the reliability of an analysis. (1) The goodness-of-fit parameter normalized root mean square deviation (NRMSD) is the single most useful measure of how well the theoretical CD spectrum calculated from the derived secondary structure composition matches the experimental data over the entire wavelength range. The NRMSD can take values from 0 (perfect fit) to 1 (no fit whatsoever). A value of NRMSD <0.1, and ideally <0.05, should be the goal. A value >0.25 probably indicates an error in the analysis procedure. (2) The comparison between the calculated and experimental spectra should be inspected for signs of systematic differences. This might happen if a membrane protein was being compared with a reference set of soluble proteins since there could be small wavelength shifts in the absorption maxima, reflecting the changes in polarity between an aqueous solvent and a lipid bilayer. (3) The results of using different algorithms should be compared. Secondary structure analysis can be considered reliable if very similar estimates are obtained after employing a number of different procedures. Further details can be found in Whitmore and Wallace (2004), UNIT 7.6, and Kelly et al. (2005).
ASSESSMENT OF STABILITY OF PROTEIN DURING TITRATION It is important to check that the protein is stable over the time course of the experiment. If spectra are recorded at intermediate points in the titration, this can be up to 2 to 3 hr, whereas if the CD signal is monitored at one wavelength only, the titration is likely to be complete within 30 min. It is important to check that the protein is stable under the conditions of pH, buffer, temperature, and other parameters used when it is being irradiated by the high-intensity Xe-arc source in conventional spectropolarimeters. The easiest way to establish that the protein is stable is to perform a blank titration in which buffer containing no ligand is added to the protein solution under the same conditions, and then to check for (1) structural stability by comparing the CD spectra in the farUV and near-UV regions at the beginning and end of the experiment, (2) formation of aggregates by recording absorption spectra over the range 240 nm to 400 nm to detect light scattering, and (3) functional stability by performing suitable assays (e.g., enzyme activity, binding capacity) to confirm that biological activity has not been damaged. When very high intensity synchrotron radiation CD beam lines are used, it appears that some damage to proteins can occur; this is more likely to be due to local heating of the sample leading to denaturation, rather than to photochemical damage resulting from the formation of reactive free-radical species (Wien et al., 2005).
SUPPORT PROTOCOL 2
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SUPPORT PROTOCOL 3
CHOICE OF BUFFER AND SOLVENT SYSTEM In choosing a buffer and solvent system for the binding experiment, several factors should be borne in mind. These include: (1) Is a certain buffer system required for the stability of the protein? (2) Does the buffer system contain components that might interfere with the CD measurements, e.g., by showing a high absorbance at the relevant wavelengths? (3) Does the buffer potentially interfere with the titration, e.g., by interacting with the protein or the ligand? An example of interference by buffer system components would be the use of phosphate buffer in a titration in which Ca2+ ions were added to a protein, since Ca2+ ions form a strong, insoluble complex with phosphate. Suitable buffers for far-UV CD experiments (down to 190 nm or 180 nm) include phosphate, Tris, and borate. Chloride ions absorb strongly below ∼195 nm, and if it is necessary to use a reasonably high ionic strength to maintain protein stability, fluoride or sulfate salts, which do not absorb significantly, should be added. Another ligand that can cause problems is imidazole, which is often used in the purification of His-tagged proteins; this absorbs strongly <225 nm and should be removed by extensive dialysis. In any case, a baseline spectrum should be recorded with the buffer to check for any potential complications. A more detailed discussion is given in Schmid (1989), UNIT 7.6, and Kelly et al. (2005).
SUPPORT PROTOCOL 4
DETERMINATION OF PROTEIN CONCENTRATION Accurate determination of the protein concentration is important not only to allow reliable analysis of the CD spectrum in terms of the secondary structure composition, but also to allow accurate estimates of the stoichiometry of ligand binding. Several methods are available but the most widely used (especially for expressed recombinant proteins) is based on measurements of the A280 using a value for the extinction coefficient calculated from the amino acid composition (Gill and von Hippel, 1989; Pace et al., 1995; UNIT 3.1), using Equations 20.10.6 and 20.10.7. The ProtParam tool within the ExPASy Web site (see Internet Resources) will calculate the extinction coefficient of a user-entered protein sequence according to Equation 20.10.7.
Equation 20.10.6
Equation 20.10.7
Where nW , nY , and nC are the numbers of tryptophan, tyrosine, and cysteine residues, respectively, in a polypeptide chain of molecular mass M (Da). For Equation 20.10.6, Gill and von Hippel (1989) gave the coefficient for nC as 120; the value of 60 is correct, since two cysteine side chains are required to form disulfide bonds. Both equations assume that all cysteines form disulfide bonds; if this is known not to be the case, the terms in nC are neglected. The equation of Gill and von Hippel (1989) refers to denatured protein, that of Pace et al. (1995) and UNIT 3.1 to native protein. CD to Study Protein Interactions
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This method is reliable provided certain conditions are met: (1) there is no contribution from light scattering; (2) there is no other chromophore (e.g., a flavin or heme cofactor) in the protein; and (3) there is no other absorbing contaminant (e.g., nucleic acids or oligonucleotides). Current Protocols in Protein Science
Assessment of whether these conditions are fulfilled can be readily made if the absorption spectrum is recorded over a suitable range (e.g., 400 nm to 240 nm). More detailed discussions of the methods available are given in Kelly et al. (2005) and UNIT 3.1.
CALIBRATION OF CD INSTRUMENT A number of compounds have been recommended for magnitude calibration of spectropolarimeters. The most commonly used compound is 1S-(+)-10-camphorsulfonic acid (CSA) because it has convenient CD peaks in the near- and far-UV regions (290.5 nm and 192.5 nm, respectively). Standard solutions of the free acid form of CSA (molecular mass 232.3 Da) are typically prepared at 0.6 mg/ml, which should have an absorbance of 0.0894 at 285 nm in a 1-cm pathlength cell. In a 1-cm pathlength cell, this solution should have an ellipticity of +202 mdeg at 290.5 nm and −420 mdeg at −192.5 nm. The ratio of the absolute signals at 192.5 nm and 290.5 nm is a good test of the performance of the instrument; the ratio should be in the range of 2.05 to 2.10. Because the free acid form of CSA is hygroscopic, the ammonium salt (molecular mass = 249.3) is sometimes preferred. A 0.6 mg/ml solution of this salt has an absorbance of 0.0833 at 285 nm in a 1-cm pathlength cell. In a 1-cm pathlength cell, this solution should have ellipticities of +188 mdeg and −391 mdeg at 290.5 nm and 192.5 nm, respectively.
SUPPORT PROTOCOL 5
The instrument manual should be consulted to ascertain how the spectropolarimeter should be adjusted if necessary. Further details of the various standards used for calibration are given in UNIT 7.6 and Kelly et al. (2005).
CHOICE OF INSTRUMENT PARAMETERS As the spectropolarimeter is used to detect very small differences between the absorbance of the right and left circularly polarized components of plane polarized light, it is not surprising that careful attention must be paid to various parameters to obtain the best results. There are four such parameters that can be adjusted, namely: bandwidth, time constant (or response time or dwell time), scan rate, and number of scans. The purpose of this section is to provide guidance as to how these can be chosen correctly, although it is not possible to give definitive recommendations that will apply under all circumstances. Further details are given in UNIT 7.6 and Kelly et al. (2005).
SUPPORT PROTOCOL 6
The bandwidth is a measure of the precision with which a monochromator will transmit light of a specified wavelength. Larger bandwidths will allow more light to fall on the sample and hence reach the photomultiplier detector, but will compromise the ability to resolve spectral features. For routine CD studies, the bandwidth should be set at 1 nm, though smaller values may be required to resolve fine structure in the near-UV spectrum of proteins. The time constant is a measure of the time over which the CD data are averaged. If a CD spectrum is being recorded, it is important that the scan rate and the time constant are chosen so as not to distort the spectrum. For example, if the scan rate is high for the chosen time constant, the slow response of the instrument can distort the spectrum from its true shape. A useful guideline for the choice of parameters is provided by Equation 20.10.8, where W = width at half-height of the spectral feature under investigation:
Equation 20.10.8
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Thus, if the bandwidth is 1 nm, combinations of time constant and scan rate of 2 sec and 20 nm/min or 0.25 sec and 200 nm/min could be chosen. Adjustments would have to be made if the bandwidth were decreased in order to resolve fine spectral detail. Increasing the number of scans will improve the signal-to-noise ratio; this ratio is proportional to the square root of the number of scans. A choice of the number of scans to be used will depend on various factors including the stability of the sample, the time available for the experiment and the stability of the instrument.
COMMENTARY Background Information CD as a spectroscopic technique CD (see UNIT 7.6) is based on the differential absorption of the left and right circularly polarized components of plane-polarized light by compounds that display chirality (optical activity). The background and practical aspects of the study of proteins by CD have been reviewed recently in UNIT 7.6 and Kelly et al. (2005). Other recent review articles, which deal with specific aspects of the use of CD to study protein interactions, include those by Drake (2001) and Greenfield (2004a). The former article emphasizes the interactions of proteins with small ligands such as drug molecules, the latter focuses on proteinprotein interactions. Proteins give rise to CD signals in various spectral regions corresponding to the types of chromophores present. In the region of peptide bond absorption (far-UV, 240 nm to 180 nm), the signals largely reflect the different types of regular secondary structure (α-helix, β-sheet, turn, etc.) present and it is possible to analyze the CD spectrum to obtain reliable estimates of the contents of these structural features (Greenfield, 2004b; UNIT 7.6; Whitmore and Wallace, 2004; see Support Protocol 1). In the region of aromatic amino acid absorption (near-UV, 320 nm to 260 nm), information on the tertiary structure of the protein can be obtained, since the magnitude and appearance of the spectra reflect the environment of these amino acid side chains (UNIT 7.6; Kelly et al., 2005). Finally, in those proteins that contain non-protein cofactors such as flavin or heme groups, the CD spectrum in the region where the cofactor absorbs provides a measure of the structural integrity of its binding site in the protein. CD to Study Protein Interactions
A note on units CD spectropolarimeters measure the difference in absorbance between the L (left) and
R (right) circularly polarized components (A = AL − AR ), but will generally display this in terms of the ellipticity (θ) in millidegrees (see UNIT 7.6 and Fig. 20.10.3 for a more detailed discussion). It should be noted that θ = tan−1 (b/a), where b and a are the minor and major axes of the ellipse, which would result if the two components were to be recombined after passage through the sample. There is a simple numerical relationship between A and θ (in millidegrees), namely θ = 33,000 A (or more precisely, θ = 32,980 A). It is important to remember that in most, if not all, biological studies, the observed CD signals are very small, with differences in absorbance (A = AL − AR ) of the order of 3 × 10−4 (corresponding to an ellipticity of ∼10 mdeg). As described in Basic Protocol 1, experiments must be carefully designed to ensure that reliable results are obtained using the CD technique. CD data are routinely normalized by scaling to molar concentrations of either the whole molecule or the repeating unit of the polymer. For the far-UV CD, the repeating unit is the peptide bond and data are expressed in terms of the mean residue weight (MRW), which is defined by MRW = M/(N – 1), where M is the molecular mass of the polypeptide chain in daltons and N is the number of amino acids in the chain (the number of amide bonds = N – 1). The mean residue ellipticity at a wavelength λ([θ]MRW,λ ) is given by Equation 20.10.9, where θλ is the observed ellipticity (in millidegrees) at wavelength λ, d is the pathlength (cm), and c is the concentration (mg/ml):
Equation 20.10.9
If the molar concentration (m) of a solute is known, the molar ellipticity at a wavelength
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λ ([θ]molar, λ ) is given by Equation 20.10.10, where θλ and d have the same meaning as in Equation 20.10.9.
Greenfield, N.J. 2004a. Circular dichroism analysis for protein-protein interactions. Methods Mol. Biol. 261:55-78. Greenfield, N.J. 2004b. Analysis of circular dichroism data. Methods Enzymol. 383:282-317.
Equation 20.10.10
The units of [θ]MRW,λ and [θ]molar,λ are deg cm2 dmol−1 . For data in absorption units, it is usual to calculate the molar differential extinction coefficient, ε. If the observed difference in absorbance at a certain wavelength of a solution of molar concentration m in a cell of pathlength d (cm) is A, then ε is given by Equation 20.10.11.
Equation 20.10.11
The units of molar differential extinction coefficient are M−1 cm−1 . There is a simple numerical relationship between [θ]MRW and ε, namely [θ]MRW = 3298 × ε. There is some debate about the correct choice of molar units for near-UV and visible CD as to whether it should be the MRW (repeating unit) or the whole polypeptide chain. In any work, it is essential to specify the actual molar unit used.
Literature Cited Anderson, L.A., Palmer, T., Price, N.C., Bornemann, S., Boxer, D.H., and Pau, R.N. 1997. Characterisation of the molybdenumresponsive ModE regulatory protein and its binding to the promoter region of the modABCD (molybdenum transport) operon of Escherichia coli. Eur. J. Biochem. 246:119-126. Baldwin, G.S., Kelly, S.M., Price, N.C., Wilson, G.W., Connolly, B.A., and Hornby, D.P. 1994. Ligand-induced conformational states of a deoxy-cytidine specific DNA methyltransferase. J. Mol. Biol. 235:545-553. Boxer, D.H., Zhang, H., Gourley, D.G., Hunter, W.N., Kelly, S.M., and Price, N.C. 2004. Sensing of remote oxyanion binding at the DNA binding domain of the molybdate-dependent transcription regulator, ModE. Org. Biomol. Chem. 2:2829-2837. Cooper, A. and Johnson, C.M. 1994. Isothermal titration microcalorimetry. Methods Mol. Biol. 22:137-150.
Haupt, G.W. 1952. An alkaline solution of potassium chromate as a transmittancy standard in the ultraviolet. J. Res. Natl. Bur. Stand. 48:414423. Hornby, D.P., Whitmarsh, A., Pinabarsi, H., Kelly, S.M., Price, N.C., Shore, P.D., Baldwin, G.S., and Waltho, J. 1994. The DNA recognition subunit of a DNA methyltransferase is predominantly a molten globule in the absence of DNA. FEBS Lett. 355:57-60. Kelly, S.M., Jess, T.J., and Price, N.C. 2005. How to study proteins by circular dichroism. Biochim. Biophys. Acta 1751:119-139. Lobley, A., Whitmore, L., and Wallace, B.A. 2002. DICHROWEB: An interactive Website for the analysis of protein secondary structure from circular dichroism spectra. Bioinformatics 18:211212. Matthews, J.R., Nicolson, J., Jaffray, E., Kelly, S.M., Price, N.C., and Hay, R.T. 1995. Conformational changes induced by DNA binding of NF-κB. Nucl. Acids Res. 23:3393-3402. McDonnell, J.M. 2001. Surface plasmon resonance: Towards an understanding of the mechanisms of biological molecular recognition. Curr. Opin. Chem. Biol. 5:572-577. Munro, A.W., Lindsay, J.G., Coggins, J.R., Kelly, S.M., and Price, N.C. 1994. Structural and enzymological analysis of the interaction of isolated domains of cytochrome P-450 BM3. FEBS Lett. 343:70-74. Munro, A.W., Lindsay, J.G., Coggins, J.R., Kelly, S.M., and Price, N.C. 1996. Analysis of the structural stability of the multidomain enzyme flavocytochrome P-450 BM3. Biochim. Biophys. Acta 1296:127-137. Pace, C.N., Vajdos, F., Fee, L., Grimsley, G., and Gray, T. 1995. How to measure and predict the molar absorption coefficient of a protein. Prot. Sci. 4:2411-2423. Rose, T., Sebo, P., Bellalou, J., and Ladant, D. 1995. Interaction of calcium with Bordetella pertussis adenylate cyclase toxin. J. Biol. Chem. 270:26370-26376. Schmid, F.X. 1989. Spectral methods of characterizing protein conformation and conformational changes in protein structure. In Protein Structure: A Practical Approach, (T.E. Creighton, ed.) pp. 252-285. Oxford University Press, Oxford.
Drake, A.F. 2001. Circular dichroism. In ProteinLigand Interactions: Structure and Spectroscopy (S.E. Harding and B.Z. Chowdhry, eds.) pp. 123167. Oxford University Press, New York.
Smith, L., Greenfield, N.J., and HitchcockDeGregori, S.E. 1994. The effects of deletion of the amino-terminal helix on troponin C function and stability. J. Biol. Chem. 269:98579863.
Gill, S.C. and von Hippel, P.H. 1989. Calculation of protein extinction coefficients from amino acid sequence data. Anal. Biochem. 182:319-326.
Sreerama, N. and Woody, R.W. 2004. Computation and analysis of protein circular dichroism spectra. Methods Enzymol. 383:318-351.
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van Holde, K.E., Johnson, W.C., and Ho, P.S. 2006. Principles of Physical Biochemistry, 2nd ed. Chapter 14. Pearson Education, Upper Saddle River, N.J. Whitmore, L. and Wallace, B.A. 2004. DICHROWEB, an online server from protein secondary structure analyses from circular dichroism spectroscopic data. Nucl. Acids Res. 32:W668-W673. Wien, F., Miles, A.J., Lees, J.G., Vronning Hoffmann, S., and Wallace, B.A., 2005. VUV irradiation effects on proteins in high-flux synchrotron radiation circular dichroism spectroscopy. J. Synchrotron Radiat. 12:517-523. Wood, S.P., Blundell, T.L., Wollmer, A., Lazarus, N.R., and Nevill, R.W.J. 1975. Relation of conformation and association of insulin to receptor binding. X-ray and circular-dichroism studies on bovine and hystricomorph insulins. Eur. J. Biochem. 55:531-542.
Internet Resources http://www.gla.ac.uk/ibls/pcf/circular dichroism. htm The home page of the Scottish Circular Dichroism Facility contains an explanation of the theory and practice of CD and has links to other important CD sites. http://www.cryst.bbk.ac.uk/cdweb/html/home.html The DICHROWEB server for online analysis of farUV CD spectra (Lobley et al., 2002; Whitmore and Wallace, 2004). http://us.expasy.org/tools/protparam.html The ProtParam tool within the ExPASy Website calculates the extinction coefficient for a native protein according to the equation proposed by Pace et al. (1995) (Equation 20.10.6). Also see UNIT 3.1.
Contributed by Sharon M. Kelly and Nicholas C. Price University of Glasgow Glasgow, United Kingdom
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Quantitative Determination of Protein Stability and Ligand Binding by Pulse Proteolysis
UNIT 20.11
Determining protein stability is an essential procedure for studying protein structures and energetics. Determination of the effects of mutations on protein stability is frequently needed to assess the role of particular amino acid residues in protein structure and folding (Lim and Sauer, 1989; Matouschek et al., 1989). Direct assessment of mutational effects on stability can be critical in interpreting experimental observations, particularly in structure/function studies. Destabilizing mutations frequently cause loss-of-function phenotypes, and determining the stability of the mutant protein gives the most direct evidence for this occurrence. Because ligand binding causes changes in the global stability of a protein, stability determination can also be used to study ligand-binding affinity or to screen ligand-protein interactions. The pulse proteolysis method allows for quantitative determination of protein stability by exploiting the fact that folded conformations are more resistant to proteolysis than unfolded conformations (Park and Marqusee, 2005; Fig. 20.11.1). In this method, protein solutions are prepared in varying concentrations of urea. After the protein solution reaches conformational equilibrium, proteolysis is performed with an excess of protease for a time window short enough to only digest the unfolded proteins in the sample. The amount of undigested protein in each reaction is an indicator of the fraction of folded protein, and can be quantified using SDS-PAGE. By plotting the amount of remaining (folded) protein versus urea concentration, the midpoint of the urea-induced unfolding transition (Cm ) is determined (see Background Information). This Cm can then be used to determine the stability of the protein in water. Unlike conventional methods for protein stability determination, pulse proteolysis does not require the intense use of specialized biophysical instrumentation, such as circular dichroism (CD) or fluorescence. Because pulse proteolysis uses SDS-PAGE to detect the fraction of folded protein, the amount of protein required is quite minimal compared to standard spectroscopic methods. Moreover, impure protein samples and even proteins in crude cell lysate can be studied by pulse proteolysis. Thus, this method is a useful
Figure 20.11.1 Principle of pulse proteolysis. Because of the difference in proteolytic susceptibility, only unfolded protein is digested completely by pulse proteolysis. Quantitation of Protein Interactions Contributed by Chiwook Park and Susan Marqusee Current Protocols in Protein Science (2006) 20.11.1-20.11.14 C 2006 by John Wiley & Sons, Inc. Copyright
20.11.1 Supplement 46
alternative when expensive instrumentation is not available or quick preliminary results are desired before carrying out more thorough biophysical characterizations. Pulse proteolysis is also useful for determining ligand-binding affinity, especially when the ligand is not compatible with spectroscopic methods. Effectively, pulse proteolysis takes a snapshot of a protein’s conformational ensemble at equilibrium. For the method to work, the folded proteins must not be digested or unfold within the time period of the pulse (usually about a minute) around the Cm . If a protein unfolds significantly during the pulse of proteolysis, both folded as well as unfolded proteins in the equilibrium mixture will appear digested. Also, in proteins with an extremely flexible region, the protein may undergo significant proteolysis during the pulse without global unfolding. In both of these cases, the fraction of folded protein is underestimated by pulse proteolysis. A simple method for verifying that proteolysis of the folded protein population is not affecting the results is by directly monitoring the proteolysis kinetics of the protein at or near the Cm . In this unit the Basic Protocol describes the use of pulse proteolysis to determine global protein stability. The Support Protocol provides a procedure to check the degree of apparent digestion of folded protein by pulse proteolysis. The possible systematic error in Cm determination from apparent proteolysis of folded protein is discussed in Background Information.
STRATEGIC PLANNING Pulse proteolysis measures the fraction of folded protein at each urea concentration using proteolysis followed by electrophoresis. The key parameter that can be determined with pulse proteolysis is Cm , the denaturant concentration where the fraction of folded protein is 0.5 (see Fig. 20.11.2). Because of the cooperativity observed in the equilibrium unfolding of proteins, the fraction folded changes most notably in the region around the Cm , the transition zone (Fig. 20.11.2). Data points in this region are most critical for determining Cm accurately. The more data points in the transition zone, the more reliable the determined Cm . The number of samples attainable in a single pulse proteolysis experiment, however, is limited, because the complete sample set needs to be analyzed on a single SDS-polyacrylamide gel to obtain a quantitative comparison. The practical number of data points for one pulse proteolysis experiment is therefore about eight to twelve.
Protein Stability & Ligand Binding by Pulse Proteolysis
Figure 20.11.2 Experimental design of the distribution of urea concentrations. (A) Equilibrium unfolding of a protein with an even distribution of urea concentration for an initial survey. (B) Equilibrium unfolding of a protein with more data points in the transition zone when an approximate Cm is known.
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Current Protocols in Protein Science
When nothing is known a priori about the stability of a protein, two trials of pulse proteolysis are recommended. The first is a preliminary survey to obtain a rough estimate of the Cm , and the second trial is an experiment tuned to the transition zone in order to determine Cm accurately. For the first trial, evenly distributed urea concentrations are recommended to cover a wide range of denaturant concentration (Fig. 20.11.2A). For the second trial, the distribution of urea concentrations is selected to have more data points in the transition zone by narrowing concentration intervals near the Cm estimated from the first trial (Fig. 20.11.2B). The Basic Protocol includes suggestions for the distribution of urea concentrations for both experiments. Thermolysin is the protease of choice for pulse proteolysis. It has broad substrate specificity, favoring hydrophobic residues, and is active even in 8 M urea. The broad optimal pH range (pH 5 to 8) of thermolysin allows its use in various pH conditions. Thermolysin requires Ca2+ for its structural integrity; therefore, buffers containing phosphate are not recommended because of formation of insoluble calcium phosphate. For proper quantification of the protein of interest on a gel, the protein should not co-migrate with thermolysin (mol. wt. 33 kDa). In addition, quenching of thermolysin activity with EDTA results in minor fragments produced by autolytic cleavage of the enzyme (migrating at approximately 8 kDa, 24 kDa, and 25 kDa) that may interfere with quantification. This autolytic cleavage can be prevented by modifying the quenching method (see Troubleshooting).
DETERMINATION OF GLOBAL PROTEIN STABILITY BY PULSE PROTEOLYSIS
BASIC PROTOCOL
This protocol describes the basic procedure for determining protein stability by pulse proteolysis (Fig. 20.11.3). Each set of pulse proteolysis data requires less than 1 mg of protein. When the protein is purified to homogeneity, the experiment can be performed with much less protein (<100 µg). For a determination of ligand-binding affinity, two sets of pulse proteolysis experiments need to be carried out, one without ligand and the other with ligand. The change in protein stability is then used to determine Kd , the dissociation equilibrium constant for the protein-ligand complex (see Background Information).
Materials 5 to 10 mg/ml protein solution of known concentration in an appropriate buffer 8.0 M urea 1.0 M CaCl2 50 mM EDTA (pH 8.0) 10 mg/ml thermolysin (see recipe) Sample buffer (e.g., UNIT 10.1) Sypro Red staining solution (Molecular Probes) Typhoon Scanner (GE Healthcare) Image analysis software (e.g., ImageJ, a freeware available for download at http://rsbweb.nih.gov/ij; or ImageQuant, GE Healthcare) Additional reagents and equipment for performing SDS-PAGE (UNIT 10.1) Establish folding-unfolding equilibrium in urea 1. Using 8.0 M urea and 1.0 M CaCl2 prepare two 1.0 ml samples of 0.5 mg/ml protein solutions in a desired buffer containing 10 mM CaCl2 , one sample without urea (solution A) and one sample containing 6.0 M urea (solution B). IMPORTANT NOTE: Because Ca2+ is required for thermolysin activity, pulse proteolysis cannot be performed in buffers containing phosphate (due to calcium phosphate precipitation). Current Protocols in Protein Science
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20.11.3 Supplement 46
Figure 20.11.3
Schematic diagram of procedures in the Basic Protocol.
IMPORTANT NOTE: Dithiothreitol (DTT) may inhibit thermolysin. If a reducing agent is needed, use only a minimal concentration (<0.5 mM) of DTT or try Tris(2carboxyethyl)phosphine hydrochloride (TCEP) instead. Be careful to make the two solutions exactly the same except for the concentration of urea. If a urea concentration higher than 6.0 M is desired, use 9.0 M urea stock solution or dry urea powder. The actual urea concentration in solution B can be determined accurately by measuring refractive indices of solutions with a refractometer (Pace, 1986) and using the following equation, where n is the difference in refractive indices of solution A and solution B: [urea] = 117.66n + 29.753 n2 + 185.56n3 .
Protein Stability & Ligand Binding by Pulse Proteolysis
Lower concentrations of protein can be used as long as they are amenable to quantitative detection by SDS-PAGE. When using Sypro Red dye, the protein concentration can be as low 30 µg/ml.
2. Mix solution A and solution B according to Table 20.11.1 to make 100-µl reactions with different urea concentrations.
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Table 20.11.1 Preparation of Protein Solutions in Different Urea Concentrations for an Initial Survey
Urea concentration (M)
Solution A (0 M urea) (µl)
Solution B (6.0 M urea) (µl)
0.0
100
0
0.6
90
10
1.2
80
20
1.8
70
30
2.4
60
40
3.0
50
50
3.6
40
60
4.2
30
70
4.8
20
80
5.2
10
90
6.0
0
100
Table 20.11.2 Suggested Preparation of Protein Solutions in Different Urea Concentrations for a Protein with an Expected Cm of ∼3 M
Urea concentration (M)
Solution A (0 M urea) (µl)
Solution B (6.0 M urea) (µl)
0.0
100
0
0.9
85
15
1.8
70
30
2.4
60
40
2.7
55
45
3.0
50
50
3.3
45
55
3.6
40
60
4.2
30
70
5.1
15
85
6.0
0
100
When an approximate Cm is known, more samples with urea concentrations near the Cm can be prepared as suggested in Table 20.11.2, which allows one to acquire more data points in the transition zone (Fig. 20.11.2B). Samples from a single set of pulse proteolysis experiments need to be separated by SDSPAGE on the same gel for the most accurate comparison. The number of reactions is therefore limited by the number of wells in the gel. Gels with twelve to fifteen wells are recommended for most applications.
3. Incubate samples overnight at 25◦ C. The reactions need to be incubated long enough to reach equilibrium. Longer incubations might be required for proteins with slow relaxation kinetics. However, because urea can covalently modify primary amines in proteins, extended incubations are not recommended. Quantitation of Protein Interactions
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Pulse proteolyze proteins in urea 4. Prepare 5-µl aliquots of 50 mM EDTA (pH 8.0) in microcentrifuge tubes (one for each reaction set up above). EDTA is used to quench the proteolysis reaction by thermolysin. The tubes with dispensed EDTA need to be capped to prevent evaporation, which can result in a change in volume and cause systematic errors in analysis.
5. Add 2 µl of thermolysin stock solution (10 mg/ml) to a reaction and vortex briefly. Addition of thermolysin stock solution increases the concentration of NaCl in the reaction by 50 mM. However, because pulse proteolysis is only suitable for systems that do not reestablish a new equilibrium within a minute, the stability determined by pulse proteolysis is that of the buffer conditions before the addition of protease.
6. After 1 min, remove 15 µl of the reaction and mix it with the 5 µl of 50 mM EDTA (pH 8.0) predispensed in a microcentrifuge tube. The pulse proteolysis results are, in principle, robust to the variation in the pulse length; reasonable errors in pulse length (<15 sec) are tolerable.
7. Repeat 5 and 6 for the rest of the reaction samples.
Determine amount of intact protein by electrophoresis 8. Add 20 µl sample buffer to each quenched reaction and load 10 µl of each sample onto an SDS polyacrylamide gel. Perform SDS-PAGE (see UNIT 10.1). Accurate pipetting is critical for quantitative determination of band intensities.
9. Stain the gel with Sypro Red staining solution according to the manufacturer’s instructions. Coomassie staining (UNIT 10.5) is an acceptable alternative to this fluorescent dye. However, Sypro Red has a broader linear dynamic range and better sensitivity.
10. Quantify the band intensity of the intact protein in each lane after scanning the stained gel with a laser scanner. For gels stained with Coomassie, a densitometer or a generic photo scanner can be used. However, linearity of the detection method should be confirmed with samples of known concentrations. For quantification of the band intensities, image-analysis software is required. ImageJ, a freeware available for download at http://rsbweb.nih.gov/ij/, can be an alternative to expensive commercial programs.
Determine Cm by curve-fitting 11. Plot band intensities (I) versus urea concentration. 12. Determine the Cm by fitting the band intensities to Equation 20.11.1:
Equation 20.11.1
where I0 , R, and T are the intensity of the folded protein at time zero, the gas constant, and temperature in degrees Kelvin, respectively. Protein Stability & Ligand Binding by Pulse Proteolysis
IMPORTANT NOTE: This curve-fitting is to determine the Cm only. The Gunf o and mvalue determined by this curve-fitting are not reliable due to an insufficient number of data points in the transition zone. The standard errors of Cm values in triplicate experiments are usually <0.1 M.
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Calculate Gunf o 13. Estimate the m-value of the protein with Equation 20.11.2.
Equation 20.11.2
m-values, which represent the dependence of the unfolding transition on denaturant concentration, are known to correlate linearly with the size of proteins. The coefficient of −0.013 in the case of urea denaturation was determined by a statistical analysis of proteins with known m-values (Myers et al., 1995). In pulse proteolysis, this m-value estimated by statistical data is more reliable than the m-value determined in step 12.
14. Calculate protein stability, Gunf ◦ , the free energy of global unfolding, using Equation 20.11.3. Representative values are shown in Table 20.11.3; see Anticipated Results for more details.
Equation 20.11.3
VERIFICATION OF RESISTANCE OF FOLDED PROTEINS TO THE PULSE OF PROTEOLYSIS
SUPPORT PROTOCOL
This Support Protocol describes the procedure to verify that a target protein is suitable for a pulse proteolysis experiment (Fig. 20.11.4). If a significant population of folded protein unfolds and is digested during the pulse, the fraction folded, as determined by pulse proteolysis, will be underestimated. To assure that digestion of the folded protein during the one-min pulse is negligible, a pilot experiment should be performed in which the protein is first digested at or near the Cm , and the rate of proteolysis measured. This procedure can also be performed with the same samples prepared for the Basic Protocol, as long as the approximate Cm is known. Simply choose the sample with a urea concentration close to the Cm . For the rest of samples, pulse proteolysis is performed as described in Basic Protocol. For the chosen sample, the proteolysis reaction is incubated further, and samples needed to measure proteolysis kinetics other than the 1 min are prepared as described below.
Materials 5 to 10 mg/ml protein solution of known concentration in a desired buffer 8.0 M urea 1.0 M CaCl2 10 mg/ml thermolysin (see recipe) 50 mM EDTA (pH 8.0) Sample buffer (UNIT 10.1) Sypro Red staining solution (Molecular Probes) Typhoon Scanner (GE Healthcare) Image analysis software (e.g., ImageJ, a freeware available for download at http://rsbweb.nih.gov/ij; or ImageQuant, GE Healthcare) Additional reagents and equipment for performing SDS-PAGE (UNIT 10.1) 1. Using 8.0 M urea and 1.0 M CaCl2 prepare 200 µl of 0.5 mg/ml protein solutions in a desired buffer containing urea with the concentration near the Cm . From this 200-µl reaction, 6 to 10 15-µl samples are taken at different time points. A 100-µl reaction is sufficient for six samples.
2. Incubate overnight at 25◦ C. Current Protocols in Protein Science
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Figure 20.11.4
Schematic diagram of procedures in the Support Protocol.
3. For each time point, prepare 5-µl aliquots of 50 mM EDTA (pH 8.0) in microcentrifuge tubes. 4. Add 4 µl of 10 mg/ml thermolysin stock solution to the reaction and vortex briefly. 5. At each designated time, take 15 µl of the reaction and mix it with 5 µl of 50 mM EDTA (pH 8.0) to quench the thermolysin. Suggested time points are 1 min, 2 min, 3 min, 4 min, 6 min, 8 min, and 10 min.
6. Add 20 µl sample buffer, perform SDS-PAGE with the quenched samples, and analyze band intensities by staining with Spyro Red and laser scanning as described in the Basic Protocol. Protein Stability & Ligand Binding by Pulse Proteolysis
20.11.8 Supplement 46
If the change in band intensity over time is negligible, the protein is suitable for pulse proteolysis. If a decrease in band intensity is observed, it is necessary to assess how significantly this affects the determination of Cm as in the following step. Current Protocols in Protein Science
7. Optional: Plot the intensity change versus time and fit the curve to a first-order rate equation:
Equation 20.11.4
where I is the intensity of the intact protein band on the SDS PAGE gel, I0 is the intensity of the folded protein at time zero, k is the kinetic constant for proteolysis of folded protein, and t is the length of incubation. I0 corresponds to the real amount of folded protein at this urea concentration, and the difference between I0 and I at 1 min is the experimental error due to the susceptibility of the population of folded proteins to pulse proteolysis. Even if some of the folded population is digested in a minute, pulse proteolysis might be still valid (see Background Information).
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Thermolysin stock solution (10 mg/ml) 10 mg thermolysin (lyophilized, Sigma) 500 µl thermolysin-dilution solution (see recipe) After dissolving lyophilized thermolysin in the thermolysin-dilution solution, centrifuge the mixture for 10 min at maximum speed, 25◦ C in a benchtop centrifuge to remove any insoluble material. Determine protein concentration by measuring A280 in 20 mM sodium phosphate buffer (pH 6.5) containing 6.0 M guanidinium chloride with ε = 2.3 cm−1 (mg/ml)−1 at 280 nm (Gill and von Hippel, 1989). Adjust the concentration to 10 mg/ml by adding an appropriate amount of thermolysin-dilution solution. Divide into 100-µl aliquots and store up to 6 months at −20◦ C. This stock solution can be thawed and frozen freely without any noticeable decrease in activity. Thermolysin-dilution solution (Inouye et al., 1998) 50 ml 5.0 M NaCl 1.0 ml 1.0 M CaCl2 H2 O to 100 ml Store indefinitely at room temperature unless evaporation is an issue. COMMENTARY Background Information Equilibrium unfolding of proteins by denaturant Protein stability is expressed as the Gibbs free energy of global unfolding (Gunf o ), the difference in stability between the unfolded (U) and folded (native; N) state. The equilibrium for global unfolding (assuming a twostate reaction) is shown in Equation 20.11.5:
Equation 20.11.5
where R is the gas constant and T is temperature in Kelvin. For stable proteins, Kunf is too small to detect directly under native conditions. Therefore, protein stability is usually determined by monitoring the denaturation of a protein using a chemical denaturant such as urea (Fig. 20.11.5). Protein stability decreases as the denaturant concentration is increased. At higher denaturant concentration Gunf o becomes close to 0, and both folded and unfolded proteins coexist in observable amounts; this range of denaturants where both folded and unfolded proteins are significantly populated is called the transition zone. In the transition zone, Kunf is determined directly from the
Quantitation of Protein Interactions
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Figure 20.11.5 Equilibrium unfolding of proteins by a chemical denaturant. Fraction of folded protein at each denaturant concentration is determined experimentally and then used to calculate Gunf (H2 O) with the linear extrapolation method (LEM).
observed fraction of folded proteins. Empirically, protein stability (Gunf ◦ ) is found to show a linear relationship with denaturant concentration (D), as shown in Equation 20.11.6 and Figure 20.11.5 (Greene and Pace, 1974):
Equation 20.11.6
where Gunf ◦ (H2 O) is the global stability of the protein in water and m, typically −0.5 to −5.0 kcal/(mol·M), is the dependence of the global stability on denaturant concentration. Therefore, Gunf ◦ (H2 O) is determined from the Gunf ◦ values measured in the transition zone by the linear extrapolation method (LEM).
Protein Stability & Ligand Binding by Pulse Proteolysis
Principle of pulse proteolysis Spectroscopic methods, such as CD and fluorescence, that can differentiate folded and unfolded proteins are frequently used to determine protein stability. The fraction of folded protein in an equilibrium mixture is calculated by using a linear combination of the folded and unfolded signal. In pulse proteolysis, the difference in proteolytic susceptibility between folded and unfolded proteins is used to determine this fraction of folded proteins in the transition zone. For enzymatic cleavage of an amide bond, the substrate peptide chain needs to be bound in an extended conformation to the active site of a proteolytic enzyme (Tyndall et al., 2005). When a protein is tightly folded, its peptide backbone is rarely in this extended conformation or not usually exposed to solvent. Proteins, therefore, need to be unfolded, at least locally if not globally, to be digested by a pro-
tease (Linderstrøm-Lang, 1938; Imoto et al., 1986; Park and Marqusee, 2004). This requirement for conformational change prior to proteolysis confers folded proteins thermodynamic and/or kinetic protection against proteolysis. This protection results in a difference in proteolytic susceptibility between folded and unfolded proteins. In pulse proteolysis, the length of digestion is chosen to be long enough to digest the unfolded proteins completely, but still short enough not to allow any digestion of the population of folded molecules (Fig. 20.11.1). As a result, only folded proteins remain intact after the pulse of proteolysis in the mixture of folded and unfolded proteins. By quantifying the amount of remaining proteins, one can determine how much protein was in the folded conformation at equilibrium. To discriminate the folded and unfolded fractions as accurately as possible, an excess of protease is used for pulse proteolysis, since unfolded protein will be digested faster in a higher concentration of protease. If the proteolysis rate for the folded population is limited by the rate for a conformational change to a susceptible conformation, the proteolysis rate for that fraction of proteins will be independent of protease concentration. Therefore, the more protease used in pulse proteolysis, the greater the difference in proteolysis rates between the folded and unfolded proteins. The protocols described here use 0.2 mg/ml thermolysin for pulse proteolysis. In the absence of urea, 0.2 mg/ml thermolysin digests unfolded proteins within a second. Although the activity of the enzyme is lower in higher concentration of urea, this concentration of thermolysin can digest unfolded proteins completely within a minute even in 6.0 M urea.
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Current Protocols in Protein Science
Fraction of folded protein
f (C ′) N
m
0.5
∆C m
C m′
Cm
Denaturant (M)
Figure 20.11.6 Errors in Cm determination from proteolysis of the folded protein populations. Only the transition zones in equilibrium unfolding curves are shown here for clarity. The apparent fraction of folded protein (solid line) is less than the actual fraction of folded protein (dashed line), which leads to a systematic error in Cm determination (Cm ).
Assessment of the error from digestion of folded protein When the population of folded protein molecules is affected by proteolysis during the pulse, the fraction of folded proteins will be underestimated. How much error does this cause in the resulting Cm ? The systematic error in Cm (Cm ) can be estimated when both the m-value and the kinetic constant for proteolysis of the folded fraction at the apparent Cm are known. Equation 20.11.7 shows the fraction of folded protein ( fN ) as a function of denaturant concentration (D), which is a modified form of Equation 20.11.1.
The relationship between fN and fN at Cm is expressed by setting t = 1 min in Equation 20.11.4:
Equation 20.11.9
where k is the rate constant for proteolysis of folded protein population at Cm . Because the apparent fraction of folded protein is 1/2 after 1 min of pulse at the apparent Cm , the half-life for proteolysis of folded population under this condition cannot be less than 1 min, which means that k is always smaller than ln2 (∼0.693 min−1 ). Combining Equation 20.11.8 and Equation 20.11.9 yields Equation 20.11.10.
Equation 20.11.7
When folded protein population is digested during the 1-min pulse proteolysis, the apparent fraction of folded protein ( fN ) and the apparent Cm (Cm ) is different from the actual values (Fig. 20.11.6). When D = Cm , fN is 1/2 by definition. The actual fN at Cm is calculated with Equation 20.11.8.
Equation 20.11.10
By rearranging this equation, Cm is expressed as a function of m and k in Equation 20.11.11.
Equation 20.11.11
Equation 20.11.8
In Figure 20.11.7, Cm is plotted as a function of k with different m-values at 25◦ C. The graph shows that this systematic error is not
Quantitation of Protein Interactions
20.11.11 Current Protocols in Protein Science
Supplement 46
Figure 20.11.7 Systematic errors in Cm as a function of the kinetic constant for proteolysis of the folded protein population (k). Cm is calculated by using Equation 20.11.11 with m-values of −2 kcal/(mol·M) (solid line), −3 kcal/(mol·M) (long-dashed line), and −4 kcal/(mol·M) (shortdashed line).
so significant when k is less than 0.2 min−1 . However, the error in Cm determination is increased significantly when k approaches ln2 (∼0.693 min−1 ). When k = 0.1 min−1 and m = −2 kcal/(mol·M), Cm is estimated as 0.062 M. Considering that the standard error in Cm in triplicate pulse proteolysis experiments is <0.1 M, the error occurring from the proteolysis of folded protein population is insignificant. Also, the error is inversely proportional to the m-values, which means that proteolysis of folded protein populations by pulse proteolysis causes smaller errors with larger proteins. When k is known, the apparent Cm can be also corrected by using Equation 20.11.11. Ligand binding and global stability The conformational stability of a protein is increased when it is associated with (i.e., bound to) a ligand whose binding requires the folded conformation of the protein. Equation 20.11.12 shows this relationship in a simplest form:
Equation 20.11.12 Protein Stability & Ligand Binding by Pulse Proteolysis
20.11.12 Supplement 46
where L is ligand, Kd is the dissociation equilibrium constant for the ligand-protein complex, and Kunf is the unfolding equilibrium constant for the protein in the absence of ligand (apo-form of the protein). With this rela-
tionship, the apparent unfolding equilibrium constant, Kunf, app is shown as:
Equation 20.11.13
Therefore, the apparent thermodynamic stability of the protein (Gunf,app ◦ ) can be expressed as:
Equation 20.11.14
Equation 20.11.14 indicates that the dissociation equilibrium constant of the complex (Kd ) can be determined by measuring the effect of ligand binding on the conformational stability of a protein. For this application, protein stability is determined without its ligand and then with its ligand. The stability determined without a ligand corresponds to Gunf ◦ , and the stability determined with a ligand corresponds to Gunf,app ◦ . Kd can then be determined using Equation 20.11.14, where Gunf ◦ , Gunf,app ◦ , and [L] is known. For a reliable determination of Kd , the difference between Gunf ◦ and Gunf,app ◦ should be much greater than the experimental errors. Therefore, the ligand concentration needs to be several orders of magnitude larger than the Kd . For example, when 3 kcal/mol of energy difference is desirable, the ligand concentration Current Protocols in Protein Science
should be 100-fold higher than the Kd . Because Kd is unknown in most practical applications, the highest possible ligand concentration will ensure a reliable determination of Kd by maximizing the difference, except in the cases when the ligand stabilizes the protein so much that urea cannot unfold the protein. When dealing with a high concentration of ligand, one should be aware of the chance that the ligand might cause nonspecific stabilization (or destabilization) by means other than complex formation. Determination of ligandbinding affinity by monitoring stability is more suitable for proteins of smaller sizes. Because m-values are linearly proportional to the size of proteins, smaller proteins manifest greater differences in Cm than larger proteins when their stabilities are increased by similar amounts. This greater change in Cm helps in the achievement of more reliable determination of Kd as described above.
Critical Parameters Quantitative analysis of SDS-PAGE gels Because pulse proteolysis relies on the quantification of intact protein by SDS-PAGE, accurate loading onto the gels and reliable determination of band intensities are critical. One should take a great care to ensure reproducible pipetting. For reliable quantification of protein from gels, the amount of sample used in SDSPAGE must be within the linear range of the employed staining and scanning method. The linearity of each detection method can be confirmed easily by running an SDS PAGE gel with serially diluted standard protein samples and measuring band intensities after staining and scanning. Pulse proteolysis can be performed over a broad range of protein concentration as long as the amounts of protein loaded on the gels are within the linear range of detection. Applicability of a broad range of protein concentrations is a useful feature for studying thermodynamic stability of multimeric proteins. Determination of urea concentration The experimental error in determining urea concentration propagates directly into the error in Cm determination. Because of the hygroscopic nature of urea, the concentration of a urea solution cannot be reliably determined from the weight of urea powder, and use of a refractometer is highly recommended. The difference in refractive indices between a urea solution and a reference buffer solution (n) allows an accurate determination of urea con-
centration, i.e., [urea] = 117.66n + 29.753 n2 + 185.56n3 (Pace, 1986).
Troubleshooting
When the urea concentration is >7 M, unfolded protein may not be digested completely with a 1-min pulse because thermolysin activity is diminished in 7 M urea. (Park and Marqusee 2004; 2005). If this impedes a reliable determination of Cm , increasing the pulse length or the amount of thermolysin may solve this problem. The resistance of folded protein to proteolysis under this new condition needs to be verified by the procedure described in the Support Protocol. Another possible reason for a persistent intact protein band at higher urea concentrations is the autolysis of thermolysin. Thermolysin produces autolysis fragments when the reaction is quenched by EDTA, though the degree of autolysis is somewhat dependent on reaction conditions (Dahlquist et al., 1976). If one of the autolysis fragments happens to co-migrate with the protein of interest, it will appear as though the protein is resistant to thermolysin at high concentration of urea. If this impedes proper quantification, one can eliminate autolysis of thermolysin by quenching the reaction with phosphoramidon before adding EDTA. To inhibit thermolysin with phosphoramidon, 1 µl of 200 µM phosphoramidon is mixed with 14 µl of thermolysin reaction. Then, 5 µl of 50 mM EDTA is mixed with the quenched reaction to prevent thermolysin from being reactivated by dissociation of phosphoramidon during boiling.
Anticipated Results Pulse proteolysis allows determination of the fraction of folded protein molecules in a urea solution. If an experiment is designed properly, this fraction can be used to calculate both the Cm and Gunf ◦ of a protein. By determining Gunf ◦ in the presence and absence of a ligand, the Kd of the protein–ligand complex can also be obtained. Table 20.11.3 shows values for Gunf ◦ determined by the pulse proteolysis method (described here) and by CD (for comparison) for RNase H* (recombinant Escherichia coli ribonuclease HI) and two variants, as well as for maltose binding protein (Park and Marqusee, 2005).
Time Considerations Setting up reactions covering a range of urea concentrations typically requires ∼1 hr.
Quantitation of Protein Interactions
20.11.13 Current Protocols in Protein Science
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Table 20.11.3 Representative Gunf ◦ Determined by Pulse Proteolysis and Circular Dichroism at pH 8.0
Protein
Cm (M)
Gunf ◦ (kcal/mol) by proteolysisa
Gunf ◦ (kcal/mol) by CDb
5.14
10.3
11.0 ± 0.3
I53A RNase H
*
3.89
7.8
8.9 ± 0.3
I53D RNase H
*
1.97
3.9
3.7 ± 0.1
Maltose binding protein
2.78
13.3
14.6 ± 0.7
Maltose binding protein + 100 µM maltose
3.59
17.2
–
E.coli RNase H*
a G
◦
determined by proteolysis was calculated by multiplying the Cm from pulse proteolysis and estimated m-values. b G ◦ was determined by circular dichroism experiments with 0.030 mg/ml protein in the same buffer as used unf for proteolysis. unf
For establishment of folding-unfolding equilibrium in urea, overnight incubation is recommended for most applications. If the conformational relaxation of the protein is slow, a longer incubation time may be required to establish equilibrium. The pulse proteolysis step usually requires ∼1 hr. Electrophoresis, staining, and quantification typically take 2 to 3 hr. When experimental settings are optimized, four to six sets of experiments can be finished in 2 days. If Coomassie staining is used instead of fluorescent dyes, the procedure requires several hours more for proper staining and destaining.
Literature Cited Dahlquist, F.W., Long, J.W., and Bigbee, W.L. 1976. Role of Calcium in the thermal stability of thermolysin. Biochemistry-US 15:1103-1111. Gill, S.C. and von Hippel, P.H. 1989. Calculation of protein extinction coefficients from amino acid sequence data. Anal. Biochem. 182:319-326. Greene, R.F. and Pace, C.N. 1974. Urea and guanidine-hydrochloride denaturation of ribonuclease, lysozyme, α-chymotrypsin, and βlactoglobulin. J. Biol. Chem. 249:5388-5393.
Lim, W.A. and Sauer, R.T. 1989. Alternative packing arrangements in the hydrophobic core of lambda repressor. Nature 339:31-36. Linderstrøm-Lang, K. 1938. Peptide bonds in globular proteins. Nature 142:996. Matouschek, A., Kellis, J.T., Jr., Serrano, L., and Fersht, A.R. 1989. Mapping the transition state and pathway of protein folding by protein engineering. Nature 340:122-126. Myers, J.K., Pace, C.N., and Scholtz, J.M. 1995. Denaturant m values and heat capacity changes: Relation to changes in accessible surface areas of protein unfolding. Protein Sci. 4:2138-2148. Pace, C.N. 1986. Determination and analysis of urea and guanidine hydrochloride denaturation curves. Methods Enzymol. 131:266-280. Park, C. and Marqusee, S. 2004. Probing the high energy states in proteins by proteolysis. J. Mol. Biol. 343:1467-1476. Park, C., and Marqusee, S. 2005. Pulse proteolysis: A facile method for quantitative determination of protein stability and ligand binding. Nature Methods 2:207-212. Tyndall, J.D., Nall, T., and Fairlie, D.P. 2005. Proteases universally recognize beta strands in their active sites. Chem. Rev. 105:973-999.
Imoto, T., Yamada, H., and Ueda, T. 1986. Unfolding rates of globular proteins determined by kinetics of proteolysis. J. Mol. Biol. 190:647649.
Contributed by Chiwook Park Purdue University West Lafayette, Indiana
Inouye, K., Kuzuya, K., and Tonomura, B. 1998. Sodium chloride enhances markedly the thermal stability of thermolysin as well as its catalytic activity. Biochim. Biophys. Acta 1388:209-214.
Susan Marqusee University of California Berkeley, California
Protein Stability & Ligand Binding by Pulse Proteolysis
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Current Protocols in Protein Science
CHAPTER 21 Peptidases INTRODUCTION
P
eptidases, the family of enzymes that are designed to cleave peptide bonds within protein chains, play extremely important roles in biology. Also known as proteases or proteolytic enzymes, the ability to digest proteins, which led to their discovery in the first place, is only one among many essential functions carried out by peptidases. For example, the cascades that lead to the formation of a blood clot (coagulation) and its eventual breakdown (fibrinolysis) are catalyzed by a series of peptidases operating sequentially in the circulation. Recent discoveries of cellular events that result in programmed cell death, or apoptosis, have revealed the essential role of a specialized type of peptidase, the caspases. Other advances have pinpointed proteolytic processing steps in developmental events in embryonic tissues in plants and animals. It is now understood that limited proteolysis controls the expression of the activity of prohormone molecules, as well as the release of receptors from cell surfaces. The processes listed above are in no way comprehensive, and are only given as examples of essential roles played by peptidases. In a more pragmatic sense, peptidases have played vital roles in the development of the field of biochemistry and cellular biology. The discovery of the selective cleavage at only a few amino acids by some readily purified digestive enzymes provided the critical reagents for protein sequencing work. Fragmentation of a large protein into smaller pieces remains an essential step in the process of primary protein sequence determination. Even in today’s world of genomics, protein sequence analysis is important. Isolation of a protein toxin, for example, can be followed by N-terminal sequence analysis, and, following fragmentation and separation, the sequencing of the C-terminal fragment. This information would permit the design of 5′- and 3′-oligonucleotide probes for the cloning of the gene for the toxin. New advances in mass spectroscopic analysis have led to the use of trypsin digests of complex mixtures followed by mass analysis of fragments to identify proteins expressed in specific cells and tissues. Chapter 16 presents methods for mass spectrometry while Chapter 22 provides overviews and protocols on proteome analysis. Another area where peptidases are used as reagents is in the study of the exposed segments of another protein. Peptidases do not, as a rule, attack the tightly folded core of a protein target, but are able to bind to and cleave any “floppy bits” on the surface that may unwind to present themselves as cleavage points. Thus, a map of the points within a sequence where peptidase cleavage occurs provides one piece of information on the three-dimensional arrangement of that protein. Peptidases play important roles in many infectious diseases, and, as such, have become valid targets for drug discovery. To accomplish this, the properties of any peptidase from an infectious organism must be studied thoroughly and compared to similar enzymes of the human or animal host. This information can then permit design and development of selective drugs targeted against the harmful species. This chapter will present overview units to give readers a background in the essential characteristics of several classes of peptidases. In future units, methods for the purification and assay of important peptidases will be presented. Due to the large number of proteolytic enzymes already known and the estimate that roughly 2% of all genes code for peptidases (e.g., ca. 400-1000 for the human genome), it is important to place newly discovered enzymes into Peptidases
Contributed by Ben M. Dunn Current Protocols in Protein Science (2002) 21.0.1-21.0.2 Copyright © 2002 by John Wiley & Sons, Inc.
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families. Thus, the properties of the new enzyme may be directly compared to those of closely related family members, which is simpler than having to make the comparison to all peptidases. Alan Barrett has provided a rational method for this analysis, as described in UNIT 21.1. When a new peptidase is discovered, the sequence can be readily compared to all known peptidases, and patterns of catalytic amino acids can be recognized that will mark the new sequence as to family type. When an unusual peptidase is discovered, the classification method developed by Barrett will highlight the similarities and differences with known enzymes, and will allow a thorough analysis of possible antecedents. Peptidases have been divided for some time into four major classes, based on the catalytic residues. The threonine class represented by the proteosome has recently joined serine, cysteine-, aspartic, and metallopeptidases. Dieter Brömme presents a description of the cysteine peptidase class (UNIT 21.2). The family relationships, mechanism of action, and structure of the aspartic peptidases are presented in UNIT 21.3. Metallopeptidases form a large (and growing) family of enzymes that play significant roles in cellular processes (UNIT 21.4). In particular, metallopeptidases have become an extremely hot topic of research due to their involvement in a wide range of disease processes. The discovery that poly-ubiquitination marks proteins for degradation by the 26S proteosome has revolutionized the field of intracellular proteolysis. The purification of the 26S and 20S proteosome particles from S. cerevisiae are detailed in UNIT 21.5, while purification of the core 20S proteosome catalytic complex from bovine pituitaries is described in UNIT 21.6. The field of proteolysis is inextricably joined with the field of peptidase inhibitors. Prevention of unwanted proteolysis is a fundamental biological process. For the Serine Peptidase family, a large number of proteins have been discovered to serve as endogenous inhibitors. Purifications of several Serpins(Serine Peptidase Inhibitors) are described in UNIT 21.7. Apoptosis, or programmed cell death, has been dissected by the discoveries of several integrated pathways that lead to activation of intracellular enzymes. The Caspases are described in an overview unit (UNIT 21.8). With the discovery of new peptidases accelerating in recent years, new methods for analysis of the activity of these enzymes are needed. UNIT 21.9 describes the construction and use of Green Fluorescent Protein-labeled protein substrates for sequence-specific peptidases. One of the largest classes of peptidases is the Serine family. Historically, these enzymes were among the most useful in early biochemical studies as trypsin (Lys or Arg) and chymotrypsin (large hydrophobic) provide complementary cleavage specificities for protein fragmentation studies. Liz Hedstrom presents an overview of this important class in UNIT 21.10. Production by recombinant means, conversion from pro-enzyme forms to mature enzymes, and purification by affinity methods are important procedures for peptidases, especially for generating mutants for structure-function analyses. Marina Parry (UNIT 21.11) describes specific methods for the production of members of the serine peptidase family. Many proteolytic enzymes function in an intracellular environment. Methods for the analysis of the cellular content of peptidases are discussed by Kevin Wang (UNIT 21.12). General methods for the study of secreted peptidases or those obtained following cell lysis are also detailed. Methods for expression, purification, and functional analysis of the important intracellular caspase family of cysteine peptidases (UNIT 21.13) and aspartic peptidases (UNIT 21.14) are provided. Two important general methods for analysis of the precursor form of proteolytic enzymes (zymography; UNIT 21.15) and the sensitive measurement of peptidase activity using fluorogenic substrates (UNIT 21.16) provide a solid basis for characterization of all types of these enzymes. Introduction
Ben M. Dunn
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Proteases BIOLOGICAL IMPORTANCE OF PROTEASES Proteases are essential to the existence of all kinds of living organisms, and many of the reasons are fairly obvious. For example, living organisms are composed largely of proteins, and commonly need to obtain the amino acid units from which they are synthesized by breaking down preexisting protein molecules. It also follows that the processes of growth and remodeling of cells, as well as the tissues of multicellular organisms, require breakdown of old protein molecules in concert with the synthesis of new ones. In addition to the gross hydrolysis of proteins to amino acids, there are myriad forms of limited proteolysis that are no less vital. For instance, many newly synthesized protein molecules require proteolytic processing to convert them to their biologically active forms. Viruses are commonly dependent upon proteases to segment their polyproteins. In both prokaryotic and eukaryotic cells, secreted proteins require proteolytic removal of the signal peptide in the secretory pathway. Many proteases are themselves synthesized as proenzymes that are catalytically inactive, and are subsequently activated proteolytically at a biologically appropriate time and place. This proteolytic “switching” of the activity of proteins represents an important mechanism of biological regulation. Unlike regulation through reversible binding to receptors, this process is irreversible, and being catalytic it requires only trace amounts of the effector molecule and can therefore provide a large amplification factor. This is elegantly demonstrated by the “cascade” of sequential activation of proenzymes in the blood coagulation process (Lorand and Mann, 1993). Proteolysis can terminate the activity of a protein just as easily as it can initiate it. The way in which the caspases mediate apoptosis by surgical cleavage of enzymes that are vital to the life of the cell is a dramatic example of this (Cohen, 1997); however, there are many others. For instance, the proteolytic destruction of proteins and polypeptides with signaling functions serves to keep the biological signals local in time and space. Life and death interactions between organisms can be mediated by proteolysis. Because animals are made of protein, they can be in-
UNIT 21.1 vaded by means of proteolysis, but proteolysis contributes to defensive systems too, as in the recognition of the peptide fragments of foreign proteins that triggers an immune response. Consistent with the biological importance of proteases, it is found that ∼2% of all genes encode these enzymes (Rawlings and Barrett, 1999); therefore, over 1000 are expected to be found in the human genome. About 300 human proteases are known at the time of this writing (see Internet Resources), but more are rapidly being discovered.
THE TERMINOLOGY OF PROTEASES The great interest in proteases as subjects of research, taken together with their large number, makes it essential that scientists have unambiguous terminology to use when communicating information about them. The term protease was introduced in the late nineteenth century to describe an enzyme that breaks down a protein. In the 1920s and 1930s, a need arose to distinguish two major groups of proteases. The enzymes of one group cut the polypeptide chain internally to produce large fragments; these were termed proteinases (Grassmann and Dyckerhoff, 1928) or endopeptidases. The enzymes of the second group acted near an end of the polypeptide chain to liberate products of one or a few amino acid residues; these were termed exopeptidases (Bergmann and Ross, 1936). A key passage from the paper of Bergmann and Ross reads: “It is therefore possible to classify the two types of peptidase as exopeptidases, which are restricted to terminal peptide linkages, and endopeptidases, which are not thus restricted. The exopeptidases, such as aminopeptidase and carboxypeptidase, should have a very restricted action on the gigantic protein molecules, because these have so few terminal groups. The endopeptidases, on the other hand, should be more suited for digesting proteins, and it is to be expected that the proteinases will fall largely in this group...”. One crucial point in the above quotation is the use of peptidase as a general term for proteolytic enzymes. Previously, peptidase had tended to be used for enzymes acting preferentially on oligopeptides (Grassmann and Dyckerhoff, 1928), but Bergmann’s usage has much to be said for it. Although “protease” remains the term most commonly used to refer Peptidases
Contributed by Alan J. Barrett Current Protocols in Protein Science (2000) 21.1.1-21.1.12 Copyright © 2000 by John Wiley & Sons, Inc.
21.1.1 Supplement 21
NH2
P3
P2
P1
P1′
P2′
S3
S2
S1
S1′
S2′
COOH
Figure 21.1.1 Scheme for the terminology of specificity subsites of proteases. The diagram shows the specificity subsites and the complementary features of the substrate (adapted from Berger and Schechter, 1970) and the recommendations of NC-IUBMB (see Internet Resources)). This scheme, for the active site of an endopeptidase, shows how substrate-binding sites are considered to be located on either side of the catalytic group, in the active site cleft. The subsites are numbered S1, S2, and so on, away from the catalytic site towards the N-terminus, and S1′, S2′, etc. towards the C-terminus. The residues of the substrate (and products) that fit are given equivalent identifiers with P in place of S. The subsites are usually thought of as binding amino acid side chains but there can also be important interactions with the polypeptide backbone. In exopeptidases the cleft is likely to be “blind” on one side, not extending beyond S1 or S1′ for an aminopeptidase or a carboxypeptidase, respectively.
to proteolytic enzymes in general, the more logical term is peptidase, which can be thought of as a short form of “peptide bond hydrolase”. The “peptidase” root has been used to create all of the terms for the subtypes of proteolytic activities. Already, the terms endopeptidase, exopeptidase, aminopeptidase, and carboxypeptidase have been encountered, and these and other “peptidase” terms now play an indispensable role in communication about proteolytic enzymes. For this reason, peptidase is the word that the Nomenclature Committee of the International Union of Biochemistry and Molecular Biology (NC-IUBMB) recommends as the general term for all proteolytic enzymes (NC-IUBMB, 1992; see Internet Resources). It is possible to use the old familiar terms protease and proteinase as synonyms of peptidase and endopeptidase, respectively, but the simple logic of peptidase and its derivatives is particularly appreciated by students and newcomers to the field of proteolytic enzymes, and the author’s personal inclination is to encourage its increasing use.
Specificity-Subsite Terminology Crystallographic structures show that the active site of a protease is commonly located in a groove on the surface of the molecule between adjacent structural domains, and the substrate specificity is dictated by the properties of binding sites arranged along the groove on one or both sides of the catalytic machinery that is
responsible for hydrolysis of the peptide bond. Accordingly, the specificity of a peptidase is described by use of a conceptual model in which each specificity subsite is able to accommodate the side chain of a single amino acid residue. The sites are numbered from the catalytic site, S1, S2...Sn towards the N-terminus of the substrate, and S1′, S2′...Sn′ towards the C-terminus. The amino acids they accommodate are numbered P1, P2...Pn, and P1′, P2′...Pn′, respectively, as shown in Figure 21.1.1.
THREE WAYS IN WHICH THE MANY PROTEASES MAY USEFULLY BE SUBDIVIDED Any field of science needs a system of classification for the objects with which it deals, especially when they are numerous. The Linnaean system of classification and nomenclature for living organisms brought order out of chaos in biology two centuries ago, and in the present day, proteins are in need of something similar. It seems likely that no one system will be applied to proteins as a whole; rather, what will happen is that separate systems will be established for particular subgroups, which will make use of characteristics that happen to work well for that particular group of proteins. For the classification of proteases there are three aspects that have proved useful in this way.
Proteases
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The Type of Proteolytic Reaction Catalyzed Enzymes are proteins that catalyze chemical reactions, and it is therefore entirely logical to classify them in terms of the reactions they catalyze. The primary division of proteases on enzymological grounds is into exopeptidases and endopeptidases as has been explained (see The Terminology of Proteases). An exopeptidase acts only near one end of a polypeptide chain. Those acting at a free N-terminus liberate a single amino acid residue, a dipeptide, or a tripeptide (i.e., aminopeptidases, dipeptidylpeptidases, and tripeptidyl-peptidases respectively). Those acting at a free C-terminus liberate a single residue (carboxypeptidases) or a dipeptide (peptidyl-dipeptidases). Other exopeptidases are specific for hydrolysis of dipeptides (dipeptidases) or remove terminal residues that are substituted, cyclized, or linked by isopeptide bonds (i.e., peptide linkages other than those of α-carboxyl to α-amino groups; omega peptidases). Endopeptidases are able to cleave internal bonds in polypeptide chains. No satisfactory general classification of these by specificity is possible, so they are divided on the basis of catalytic mechanism (see The Type of Catalytic Mechanism). Most endopeptidases will act on a polypeptide chain of any length, so that proteins are Table 21.1.1
amongst the substrates, but a subset of endopeptidases are restricted to acting on substrates smaller than proteins; these are termed “oligopeptidases” (Rawlings et al., 1991). The approach of classifying peptidases by reaction catalyzed (as far as possible), and then by catalytic type, is followed in the comprehensive recommendations on enzyme nomenclature of the Nomenclature Committee of the International Union of Biochemistry and Molecular Biology (NC-IUBMB). This committee is the successor to the Enzyme Commission, and the index numbers that it assigns to enzymes are still described as “EC” numbers. The latest printing of the EC list is in Enzyme Nomenclature (NCIUBMB, 1992), but the current, updated version can be found on the World Wide Web at http://www.chem.qmw.ac.uk/iubmb/enzyme/ EC34/ (see Internet Resources). In the EC system, all enzymes are divided between six classes, amongst which the hydrolases form class 4 and the proteases (recommended to be called peptidases) form subclass 3.4. The 14 sub-subclasses of this subclass are shown in Table 21.1.1.
The Type of Catalytic Mechanism It was first pointed out by Hartley (1960) that proteolytic enzymes can be grouped according to the chemical nature of the catalytic
The EC System of Classification of Peptidasesa
Sub-subclass Peptidase type 3.4.11 3.4.13 3.4.14 3.4.15 3.4.16 3.4.17 3.4.18 3.4.19 3.4.21 3.4.22 3.4.23 3.4.24 3.4.25 3.4.99
Aminopeptidases Dipeptidases Dipeptidyl-peptidases Peptidyl-dipeptidases Serine-type carboxypeptidases Metallocarboxypeptidases Cysteine-type carboxypeptidases Omega peptidases Serine endopeptidases Cysteine endopeptidases Aspartic endopeptidases Metalloendopeptidases Threonine endopeptidases Endopeptidases of unknown type Total
Number of entries 20 11 8 3 4 19 1 11 77 28 34 70 1 0 287
aThe data apply to the Web edition of Enzyme Nomenclature following the
incorporation of Supplements 1 to 6 (http://www.chem.qmw.ac.uk/iubmb/enzyme/ EC34; see Internet Resources).
Peptidases
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Table 21.1.2
Catalytic Types of Proteasesa
Type
Examples
Typical inhibitors
Aspartic Cysteine Metallo
Pepsin, cathepsin E Papain, cathepsin K Thermolysin, vertebrate collagenase, carboxypeptidase A Trypsin, prolyl oligopeptidase Proteasome gpr endopeptidase, prepilin type IV peptidase
Pepstatin Iodoacetate 1,10-phenanthroline
Serine Threonine Unknown
Diisopropyl fluorophosphate (Lactacystin for some) None of the above
aFive types of proteases are recognized on the basis of the chemistry of the catalytic site, and some are still of unknown
catalytic type. More information about the example enzymes listed here can be obtained from the Web pages of the EC list and the MEROPS database (see Internet Resources).
site, and this has proved valuable. The modern system of classification of proteases in this way now provides for the five groups of aspartic-, cysteine-, metallo-, serine-, and threoninetypes of proteases (Table 21.1.2). The threonine type was recently added to the four recognized by Hartley (Seemüller et al., 1995). Historically, the catalytic type of the active site has generally been identified by use of type-specific inhibitors. Now that the deduced amino acid sequence is commonly available at an early stage in the study of a protease, it is usually possible to recognize that the enzyme is homologous to proteases for which the catalytic type is already known, and to conclude that it will be the same. If not, site-directed mutagenesis of potential catalytic residues may be instructive. In the serine-, cysteine- and threonine-type proteases the –OH or –SH group on the side chain of the amino acid at the heart of the catalytic site acts as a nucleophile in catalysis, forming an acyl enzyme intermediate. Sometimes this ester or thiol ester can break down in a transamination reaction rather than hydrolysis. In this unit, all of the serine-, cysteine-, and threonine-types of proteases are referred to as “protein-nucleophile” enzymes. In contrast, the aspartic and metallopeptidases seem to activate a bound water molecule to serve as the nucleophile, and it is seldom possible to detect any covalent intermediate in catalysis. Although much remains to be learned about the catalytic mechanisms of the aspartic- and metallopeptidases, the authors find it useful to describe them as “water-nucleophile” enzymes. There are a significant number of proteases for which the catalytic mechanism remains to be determined.
Structural Relationships As has been shown, the classification of proteases by the type of reaction catalyzed and by the catalytic mechanism, as implemented in the EC system, places each of the peptidases in one of just 14 sub-subclasses, with only 6 of the sub-subclasses available for the very numerous endopeptidases (comprising 210 of the total of 287 entries). This inevitably lumps together peptidases that are very different, and very importantly, it takes no account of structural groupings of peptidases that would reflect evolutionary relationships. Rawlings and Barrett (1993) described a new form of classification that was designed to group the peptidases in a way that would reflect essential structural features and evolutionary relationships, taking advantage of the wealth of amino acid sequence data that was becoming available. This has come to be called the “MEROPS” system (for reasons that are no longer important). In the MEROPS system, proteases are classified by use of structural features that are believed to reflect evolutionary relationships. Similarities in amino acid sequence are used to group closely related peptidases in families. Groups of families that show evidence of having had a common origin are then grouped together in clans. Each family of proteases is formed around a founding member or type example (such as pepsin in family A1) in the following manner. First, the part of the molecule that is responsible for proteolytic activity is identified, and is termed the “peptidase unit” (http://www. merops.co.uk; see Internet Resources). The family is then built up by adding proteases that show statistically significant similarity in amino acid sequence to the peptidase unit of
Proteases
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the type example, or of another existing member of the family. The comparisons are made with strict parameters such that there is no realistic possibility of two peptidases being placed in the same family when they are not truly homologous. The use of the peptidase units for comparisons eliminates many problems that could arise from the fact that proteases are often mosaic and contain additional nonpeptidase domains that are shared with other groups of proteins. Some families contain two or more distinct groups of proteases that differ greatly in primary structure. When a dendrogram for a family shows branches separated by deep divergences (150 protein alignment matrix, or PAM, units; UNIT 2.1) then the branches are recognized as subfamilies. Each family is assigned a simple identifier that starts with a letter denoting the catalytic type of the peptidases it contains (i.e., A, C, M, S, T, or U for aspartic, cysteine, metallo, serine, threonine, or unknown type; see Internet Resources). The family identifier is completed with a number that is assigned sequentially as the families are recognized in the MEROPS classification. Subfamily names are derived from the family name with the addition of a letter also assigned sequentially, e.g., M10A, M10B, and M10C for the subdivisions of family M10. Although proteases in different families have no significant similarity in amino acid sequence, there may still be evidence that they share a common origin. This can come from similar three-dimensional structures or conserved sequence motifs around the catalytic amino acids. The families of peptidases showing these distant relationships are grouped together in a “clan”. The identifier of a clan in the MEROPS system starts with a letter based on the catalytic type or types of the families it contains, and is completed by a second letter of the alphabet assigned sequentially as the clan is recognized. A few clans contain families of more than one of the types C, S, and T, and the letter P is used in forming the identifier of these. Thus, clan PA contains several families of serine peptidases but also includes families of cysteine peptidases from RNA viruses. It is probable that a catalytic serine has mutated to cysteine with retention of catalytic activity in the evolution of this clan. The families and clans that are recognized in the MEROPS system at the time of writing are summarized in Table 21.1.3 in the Appendix, but the reader should see the Web version of the MEROPS database
for current information (http://www.merops. co.uk; see Internet Resources). A MEROPS identifier for each protease is constructed from the family identifier, padded to three characters when necessary, followed by a decimal point and a three-digit number. An example would be S01.151 for trypsin in family S1.
THE MEROPS DATABASE REFLECTS THE GROWTH OF KNOWLEDGE ABOUT PROTEASES The rate of discovery of new proteases is so rapid that no printed publication can ever be up to date. In an attempt to overcome this problem, the MEROPS database has been established on the World Wide Web (http://www.merops. co.uk; Rawlings and Barrett, 1999, 2000). There are three major sets of pages in the database, representing the three levels of classification: peptidases (in which each peptidase has its own “PepCard”), families (with a “FamCard” for each), and clans (“ClanCards”). Each set of cards pages has a name index, and in addition, the peptidases are indexed by source organism and MEROPS identifier. In addition to its thousands of links to other sources of information about proteases, the MEROPS database contains a comprehensive list of the alternative names of the enzymes, images of the molecules of most of the proteases for which three-dimensional structures have been published, sequence alignments and dendrograms for many families and reference lists. The proteases that are brought together into one family or clan on the basis of their structural similarities and homology in the MEROPS system tend to have common characteristics that distinguish them from the proteases in other families and clans. For this reason, the assignment, which can be made when the deduced amino acid sequence is all the information one has about a putative protease, can lead to predictions about its proteolytic activity and sometimes biological functions.
Literature Cited Berger, A. and Schechter, I. 1970. Mapping the active site of papain with the aid of peptide substrates and inhibitors. Philos.Trans. R. Soc. Lond. Ser. B Biol. Sci. 257:249-264. Bergmann, M. and Ross, W.F. 1936. On proteolytic enzymes. X. The enzymes of papain and their activation. J. Biol. Chem. 114:717-726. Cohen, G.M. 1997. Caspases: The executioners of apoptosis. Biochem. J. 326:1-16. Peptidases
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Grassmann, W. and Dyckerhoff, H. 1928. Über die Proteinase und die Polypeptidase der Hefe. 13. Abhandlung über Pflanzenproteasen in der von R. Willstätter und Mitarbeitern begonnen Untersuchungsreihe. Hoppe-Seylers Z. Physiol. Chem. 179:41-78.
Internet Resources
Hartley, B.S. 1960. Proteolytic enzymes. Annu. Rev. Biochem. 29:45-72.
http://www.chem.qmw.ac.uk/iubmb/enzyme
Lorand, L. and Mann, K.G. (eds.) 1993. Mammalian Blood Coagulation Factors and Inhibitors, Part A. Methods Enzymol. 222:1-195.
http://merops.co.uk/ Web site of the MEROPS database that provides links to much information on proteases that is available on the Internet.
Web site for the latest update of the nomenclature for enzymes of the Nomenclature Committee of the International Union of Biochemistry and Molecular Biology.
NC-IUBMB (Nomenclature Committee of the International Union of Biochemistry and Molecular Biology). 1992. Enzyme Nomenclature 1992. Recommendations of the Nomenclature Committee of the International Union of Biochemistry and Molecular Biology on the Nomenclature and Classification of Enzymes. Academic Press, Orlando, Fla.
http://www.chem.qmw.ac.uk/iubmb/enzyme/EC34
Rawlings, N.D. and Barrett, A.J. 1993. Evolutionary families of peptidases. Biochem. J. 290:205-218.
Contributed by Alan J. Barrett Babraham Institute Babraham, Cambridge, United Kingdom
Rawlings, N.D. and Barrett, A.J. 1999. MEROPS: The peptidase database. Nucl. Acids Res. 27:325331. Rawlings, N.D. and Barrett, A.J. 2000. MEROPS: The peptidase database. Nucl. Acids Res. 28:323325. Rawlings, N.D., Polgár, L., and Barrett, A.J. 1991. A new family of serine-type peptidases related to prolyl oligopeptidase. Biochem. J. 279:907908. Seemüller, E., Lupas, A., Stock, D., Löwe, J., Huber, R., and Baumeister, W. 1995. Proteasome from Thermoplasma acidophilum: A threonine protease. Science 268:579-582.
Key References Barrett, A.J., Rawlings, N.D., and Woessner, J.F. (eds.). 1998. Handbook of Proteolytic Enzymes. Academic Press, London. This recent volume is the most comprehensive single publication on proteolytic enzymes that has so far been produced.
Web site for the latest update of the nomenclature for peptidases of the Nomenclature Committee of the International Union of Biochemistry and Molecular Biology.
APPENDIX: SUMMARY OF FAMILIES AND CLANS RECOGNIZED IN THE MEROPS SYSTEM The identifier of a clan in the MEROPS system starts with a letter based on the catalytic type or types of the families it contains, and is completed by a second letter of the alphabet assigned sequentially as the clan is recognized (see Structural Relationships for more information). The families and clans that are recognized in the MEROPS system at the time of writing are summarized in Table 21.1.3, but the reader should see the Web version of the MEROPS d atabase f or current information (http://www.merops.co.uk; see Internet Resources). IMPORTANT NOTE: The MEROPS identifier listed in the following table is that of the protease example only.
Proteases
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Table 21.1.3
Family identifier
Overview of the MEROPS Classification of Proteases
Subfamily Protease name (source organism) designation
Protein-nucleophile proteases Catalytic type: cysteine Clan CA C1 A Papain (Carica papaya) B Bleomycin hydrolase (Saccharomyces cerevisiae) C2 Calpain 2 (Homo sapiens) C6 Helper component proteinase (potato virus Y) C7 p29 proteinase (Cryphonectria hypovirus) C8 p48 proteinase (Cryphonectria hypovirus) C9 nsP2 proteinase (Sindbis virus) C10 Streptopain (Streptococcus pyogenes) C12 Ubiquitin C-terminal hydrolase UCH-L1 (Homo sapiens) C16 Papain-like endopeptidase 1 (murine hepatitis virus) C21 Endopeptidase (turnip yellow mosaic virus) C23 Endopeptidase (apple stem pitting virus) C27 Endopeptidase (rubella virus) C28 Leader proteinase (foot-and-mouth disease virus) C29 Papain-like endopeptidase 2 (murine hepatitis virus) C31 PCPα endopeptidase (lactate-dehydrogenase-elevating virus) C32 PCPβ endopeptidase (Lelystad virus) C33 Nsp2 cysteine endopeptidase (equine arteritis virus) C34 Putative papain-like endopeptidase (apple chlorotic leaf spot virus) C35 Putative papain-like endopeptidase (apple stem grooving virus) C36 Papain-like endopeptidase (beet necrotic yellow vein virus) C39 Processing peptidase PedD (Pediococcus acidilactici) C41 Cysteine proteinase (hepatitis E virus) C42 Papain-like endopeptidase (sugar beet yellows virus) C47 Staphopain (Staphylococcus aureus) Clan CD C11 Clostripain (Clostridium histolyticum) C13 Legumain (Canavalia ensiformis) C14 Caspase-1 (Rattus norvegicus) C25 Gingipain R (Porphyromonas gingivalis) Clan CE C5 Adenain (human adenovirus type 2) Clan CF C15 Pyroglutamyl-peptidase I (Bacillus amyloliquefaciens) Clan CG C17 ER-60 proteinase (Homo sapiens)
Protease MEROPS identifier
C01.001 C01.085 C02.003 C06.001 C07.001 C08.001 C09.001 C10.001 C12.001 C16.001 C21.001 C23.001 C27.001 C28.001 C29.001 C31.001 C32.001 C33.001 C34.001 C35.001 C36.001 C39.002 C41.001 C42.001 C47.001 C11.001 C13.001 C14.001 C25.001 C05.001 C15.001 C17.001 continued
21.1.7 Current Protocols in Protein Science
Supplement 21
Table 21.1.3
Family identifier
Overview of the MEROPS Classification of Proteases, continued
Subfamily designation
Protease MEROPS identifier
Protease name (source organism)
Clan CH C46 Hedgehog protein (Drosophila melanogaster) Families not assigned to clans C22 Cysteine proteinase (Trichomonas vaginalis) γ-glutamyl hydrolase (Rattus norvegicus) C26 C40 Dipeptidyl-peptidase VI (Bacillus sphaericus) Catalytic type: serine Clan SB S8 A B C Clan SC S9 A B C S10 S15 S28 S33 S37 Clan SE S11 S12 S13 Clan SF S24 S26 S41
A B A B
C46.001 C22.001 C26.001 C40.001
Subtilisin (Bacillus licheniformis) Kexin (Saccharomyces cerevisiae) Tripeptidyl-peptidase II (Homo sapiens)
S08.001 S08.070 S08.090
Prolyl oligopeptidase (Sus scrofa) Dipeptidyl-peptidase IV (Homo sapiens) Acylaminoacyl-peptidase (Homo sapiens) Carboxypeptidase C (Saccharomyces cerevisiae) X-Pro dipeptidyl-peptidase (Lactococcus lactis) Lysosomal Pro-X carboxypeptidase (Homo sapiens) Prolyl aminopeptidase (Neisseria gonorrhoeae) PS-10 peptidase (Streptomyces lividans)
S09.001 S09.003 S09.004 S10.001 S15.001 S28.001 S33.001 S37.001
D-Ala-D-Ala carboxypeptidase A (Bacillus
S11.001
stearothermophilus) D-Ala-D-Ala carboxypeptidase B (Streptomyces sp.) D-Ala-D-Ala peptidase C (Escherichia coli)
S12.001 S13.001
Pepressor LexA (Escherichia coli) Signal peptidase I (Escherichia coli) Signalase (Saccharomyces cerevisiae) Tsp protease (Escherichia coli) Tricorn core protease (Sulfolobus solfataricus)
S24.001 S26.001 S26.010 S41.001 S41.005
Clan SH S21 Assemblin (human cytomegalovirus) Clan SK S14 Endopeptidase Clp (Escherichia coli) S49 Protease IV (Escherichia coli) Families not assigned to clans S16 Endopeptidase La (Escherichia coli) S18 Omptin (Escherichia coli) S19 Chymotrypsin-like protease (Coccidioides immitis) S38 Chymotrypsin-like protease PrtB (Treponema denticola)
S21.002 S14.001 S49.001 S16.001 S18.001 S19.001 S38.001 continued
Proteases
21.1.8 Supplement 21
Current Protocols in Protein Science
Table 21.1.3
Family identifier
Overview of the MEROPS Classification of Proteases, continued
Subfamily designation
Mixed catalytic type Clan PA C3 A B C D E C4 C24 C30 C37 C38 S1
A B C D E
S3 S6 S7 S29 S30 S31 S32 S35 S43 Clan PB C44 C45 S45 T1 T2 T3 T4
A B
Protease name (source organism)
Protease MEROPS identifier
Picornain 3C (poliovirus type 1) Picornain 2A (poliovirus type 1) Picornain 3C (foot and mouth disease virus) 24 kDa proteinase (cowpea mosaic virus) Picornain 3C (hepatitis A virus) NIa proteinase (plum pox potyvirus) 3C endopeptidase (rabbit hemorrhagic disease virus) Picornain 3C-like polyprotein endopeptidase (murine hepatitis virus) Processing peptidase (Southampton virus) Putative processing peptidase (parsnip yellow fleck virus) Chymotrypsin (Bos taurus) Glutamyl endopeptidase (Staphylococcus aureus) Protease Do (Escherichia coli) Lysyl endopeptidase (Achromobacter lyticus) Streptogrisin A (Streptomyces griseus) Togavirin (Sindbis virus) IgA1-specific serine-type prolyl endopeptidase (Neisseria gonorrhoeae) Flavivirin (yellow fever virus) Hepacivirin (hepatitis C virus) P1 proteinase (plum pox potyvirus) NS3 polyprotein peptidase (cattle viral diarrhea virus) Serine endopeptidase (equine arteritis virus) 36 kDa protease (apple stem grooving virus) Porin D (Pseudomonas aeruginosa)
C03.001 C03.020 C03.008 C03.003 C03.005 C04.001 C24.001 C30.001
Glutamine PRPP amidotransferase precursor (Homo sapiens) Acyl-coenzyme A:6-aminopenicillanic acid acyl-transferase precursor (Penicillium chrysogenum) Penicillin amidohydrolase precursor (Escherichia coli) Archaean proteasome (Thermoplasma acidophilum) HslVU protease component HslV (Escherichia coli) Glycosylasparaginase precursor (Homo sapiens) γ-glutamyltransferase (Escherichia coli) Aminopeptidase DmpA (Ochrobactrum anthropi)
C44.001
C37.001 C38.001 S01.152 S01.269 S01.273 S01.280 S01.261 S03.001 S06.001 S07.001 S29.001 S30.001 S31.001 S32.001 S35.001 S43.001
C45.001 S45.001 T01.002 T01.006 T02.001 T03.001 T04.001 continued
Peptidases
21.1.9 Current Protocols in Protein Science
Supplement 21
Table 21.1.3
Family identifier
Overview of the MEROPS Classification of Proteases, continued
Subfamily designation
Protease MEROPS identifier
Protease name (source organism)
Water-nucleophile proteases Catalytic type: aspartic/carboxyl Clan AA A1 Pepsin A (Homo sapiens) A2 HIV1 retropepsin (human immunodeficiency virus type 1) A3 A Endopeptidase (cauliflower mosaic virus) B Bacilliform virus putative protease (rice tungro bacilliform virus) A9 Polyprotein peptidase (human spumaretrovirus) A11 A Copia transposon (Drosophila melanogaster) B SIRE-1 peptidase (Glycine max) A12 Retrotransposon bs1 endopeptidase (Zea mays) A13 Retrotransposon peptidase (Drosophila buzzatii) A16 Tas retrotransposon peptidase (Ascaris lumbricoides) A17 Pao retrotransposon peptidase (Bombyx mori) A18 Putative proteinase of Skippy retrotransposon (Fusarium oxysporum) Clan AB A6 Nodavirus endopeptidase (flock house virus) A21 Tetravirus endopeptidase (Nudaurelia capensis omega virus) Families not assigned to clans A4 Aspergillopepsin II (Aspergillus niger) A5 Thermopsin (Sulfolobus acidocaldarius) A7 Pseudomonapepsin (Pseudomonas sp. 101) A8 Signal peptidase II (Escherichia coli) A22 Presenilin 1 (Homo sapiens) Catalytic type: metallo Clan MA M1 Membrane alanyl aminopeptidase (Homo sapiens) M2 Germinal peptidyl-dipeptidase A (Homo sapiens) M4 Thermolysin (Bacillus stearothermophilus) M5 Mycolysin (Streptomyces cacaoi) M6 Immune inhibitor A (Bacillus thuringiensis) M7 Snapalysin (Streptomyces lividans) M8 Leishmanolysin (Leishmania major) M9 A Microbial collagenase (Vibrio alginolyticus) B Microbial collagenase (Clostridium perfringens) M10 A Collagenase 1 (Homo sapiens) B Serralysin (Serratia marcescens) C Fragilysin (Bacteroides fragilis) M11 Gametolysin (Chlamydomonas reinhardtii)
A01.001 A02.001 A03.001 A03.002 A09.001 A11.001 A11.051 A12.001 A13.001 A16.001 A17.001 A18.001
A06.001 A21.001
A04.002 A05.001 A07.001 A08.001 A22.001
M01.001 M02.004 M04.001 M05.001 M06.001 M07.001 M08.001 M09.001 M09.002 M10.001 M10.051 M10.020 M11.001 continued
Proteases
21.1.10 Supplement 21
Current Protocols in Protein Science
Table 21.1.3
Overview of the MEROPS Classification of Proteases, continued
Family identifier
Subfamily designation
M12
A B C
Protease name (source organism)
Astacin (Astacus fluviatilis) Adamalysin (Crotalus adamanteus) ADAM 10 (Bos taurus) M13 Neprilysin (Homo sapiens) M30 Hyicolysin (Staphylococcus hyicus) M36 Fungalysin (Aspergillus fumigatus) M48 Ste24p endopeptidase (Saccharomyces cerevisiae) Other HEXXH metallopeptidase families M3 A Thimet oligopeptidase (Rattus norvegicus) B Oligopeptidase F (Lactococcus lactis) M26 IgA1-specific metalloendopeptidase (Streptococcus sanguis) M27 Tentoxilysin (Clostridium tetani) M32 Carboxypeptidase Taq (Thermus aquaticus) M34 Anthrax lethal factor (Bacillus anthracis) M35 Penicillolysin (Penicillium citrinum) M41 FtsH endopeptidase (Escherichia coli) M43 Cytophagalysin (Cytophaga sp.) M50 S2P protease (Homo sapiens) M51 Sporulation factor SpoIVFB (Bacillus subtilis) Clan MC M14 A Carboxypeptidase A (Homo sapiens) B Carboxypeptidase E (Bos taurus) γ-D-glutamyl-(L)-meso-diaminopimelate peptidase I C (Bacillus sphaericus) Clan MD M15 A Zinc D-Ala-D-Ala carboxypeptidase (Streptomyces albus) B VanY D-Ala-D-Ala carboxypeptidase (Enterococcus faecium) C Endolysin (bacteriophage A118) M45 VanX D-Ala-D-Ala dipeptidase (Enterococcus faecium) Clan ME M16 A Pitrilysin (Escherichia coli) B Mitochondrial processing peptidase (Saccharomyces cerevisiae) M44 Metalloendopeptidase (Vaccinia virus) Clan MF M17 Leucyl aminopeptidase (Bos taurus) Clan MG M24 A Methionyl aminopeptidase I (Escherichia coli) B X-Pro aminopeptidase (Escherichia coli)
Protease MEROPS identifier M12.001 M12.141 M12.210 M13.001 M30.001 M36.001 M48.001 M03.001 M03.007 M26.001 M27.001 M32.001 M34.001 M35.001 M41.001 M43.001 M50.001 M51.001 M14.001 M14.005 M14.008
M15.001 M15.010 M15.020 M45.001 M16.001 M16.003 M44.001 M17.001 M24.001 M24.004 continued
Peptidases
21.1.11 Current Protocols in Protein Science
Supplement 22
Table 21.1.3
Family identifier Clan MH M18 M20 M25 M28
M40 M42 Clan MJ M38 Clan MK M22
Overview of the MEROPS Classification of Proteases, continued
Subfamily designation
A B A B C
Protease MEROPS identifier
Protease name (source organism)
Aminopeptidase I (Saccharomyces cerevisiae) Glutamate carboxypeptidase (Pseudomonas sp.) Gly-X carboxypeptidase (Saccharomyces cerevisiae) X-His dipeptidase (Escherichia coli) Leucyl aminopeptidase (Vibrio proteolyticus) Aminopeptidase (Streptomyces griseus) IAP aminopeptidase (Escherichia coli) Carboxypeptidase Ss1 (Sulfolobus solfataricus) Glutamyl aminopeptidase (Lactococcus lactis)
M18.001 M20.001 M20.002 M25.001 M28.002 M28.003 M28.005 M40.001 M42.001
β-aspartyl dipeptidase (Escherichia coli)
M38.001
O-sialoglycoprotein endopeptidase (Pasteurella haemolytica)
M22.001
Clan ML M52 HYBD endopeptidase (Escherichia coli) Families not assigned to clans M19 Membrane dipeptidase (Homo sapiens) β-lytic metalloendopeptidase (Achromobacter lyticus) M23 M29 Aminopeptidase T (Thermus aquaticus) M37 Lysostaphin (Staphylococcus simulans) M49 Dipeptidyl-peptidase III (metallo-form) (Rattus norvegicus) Proteases of unknown catalytic type U3 gpr endopeptidase (Bacillus megaterium) U4 Sporulation factor SpoIIGA (Bacillus subtilis) U6 Murein endopeptidase (Escherichia coli) U9 Prohead proteinase (bacteriophage T4) U12 Prepilin type IV peptidase (Escherichia coli) α-aspartyl dipeptidase (Escherichia coli) U28 U29 Cardiovirus endopeptidase 2A (encephalomyocarditis virus) U32 Proteinase C (Porphyromonas gingivalis) U34 Dipeptidase A (Lactobacillus helveticus) U39 NS2-3 protease (hepatitis C virus) U40 Protein P5 murein endopeptidase (bacteriophage P-6) U43 Endopeptidase (infectious pancreatic necrosis virus) U44 Npro endopeptidase (hog cholera virus) U46 PfpI endopeptidase (Pyrococcus furiosus) U48 CAAX prenyl protease 2 (Saccharomyces cerevisiae)
M52.001 M19.001 M23.001 M29.001 M37.001 M49.001
U03.001 U04.001 U06.001 U09.001 U12.001 U28.001 U29.001 U32.001 U34.001 U39.001 U40.001 U43.001 U44.001 U46.001 U48.001
aThe MEROPS identifier is that of the specific example listed.
21.1.12 Supplement 22
Current Protocols in Protein Science
Papain-like Cysteine Proteases The name “cysteine proteinase” refers to the protease’s nucleophilic cysteine residue, which forms a covalent bond with the carbonyl group of the scissile peptide bond in substrates. By far the most numerous family among cysteine proteases is formed by papain-like proteases that have been classified as the C1 family (UNIT 21.1; Rawlings and Barrett, 1994). Papain-like cysteine proteases are found in viruses, plants, primitive parasites, invertebrates, and vertebrates. Most of the members of the C1-family are endopeptidases, but some of them act as exopeptidases or have mixed types of activity. Mammalian papain-like cysteine proteases are also known as cathepsins, a name originally introduced by Willstädter and Bamann in 1929 describing acidic intracellular proteases. Due to recent advances in deciphering the human genome, the number of human cathepsins has increased from four to eleven members. In addition, two novel papain-like cysteine proteases, mouse cathepsin P and rat cathepsin Q, have been described, for which human orthologs are presently unknown. The name cathepsin is derived from the Greek word “καθεψειν” which means “to digest.” It should be noted that the term cathepsin is also used for members of the aspartate and serine protease classes. This section is focused on mammalian C1-family cysteine proteases, designated as cathepsins in the following unit.
SEQUENCE HOMOLOGIES AND EVOLUTIONARY RELATEDNESS OF CATHEPSINS Cathepsins share a high sequence homology with papain, a plant protease isolated from papaya fruits. Presently, eleven human members have been cloned and partially or fully characterized. Some of the cathepsins have been cloned simultaneously in different laboratories, resulting in multiple designations. Table 21.2.1 summarizes gene assignment numbers, various names given to these proteases, length of the protein domains in amino acids, and chromosomal localization of the human genes. The nomenclature of cysteine-dependent cathepsins is presently under review. All cathepsins have in common that their genes encode a protein containing a 15- to 21-amino acid–long signal sequence, a propeptide of varying lengths between 41 and 251 amino acids, and a mature domain of 214 to 260 amino
UNIT 21.2
acids. The highest conservation is observed in the mature domain of the polypeptide that represents the catalytically active protease. Table 21.2.2 shows sequence identities and similarities between the cathepsins and papain. All cathepsins have in common a conserved active site that is formed by cysteine, histidine, and asparagine residues. The cysteine residue is always followed by a tryptophan residue; the histidine residue by a small and neutral amino acid and branched hydrophobic residues such as valine, leucine, or isoleucine; and the asparagine residue is embedded into the AsnSer-Trp motif. The second residue in the mature domain next to the processing site of the cathepsin zymogens is always a proline which may protect the proteases against N-terminal truncation by lysosomal aminopeptidase activity. In addition, various other residues and motifs are conserved among the cathepsins. Figure 21.2.1 depicts the sequence alignment of the mature cathepsins of the eleven presently known human C1 family members, and two novel sequences recently identified in mouse and rat, and highlights residues of identity and similarity. Using a phylogenetic analysis program, cathepsins can be grouped into a cathepsin L-like group (cathepsins L, V, S, K, and H), a cathepsin F-like group (cathepsins F and W), and a cathepsin B-like group which comprises, besides cathepsin B, other cathepsins that are not well assigned (such as cathepsins Z and C; Fig. 21.2.2).
CATALYTIC MECHANISM OF CATHEPSINS The catalytic site in the cysteine proteases of the C1 family is formed by a cysteine residue (Cys-25), a histidine residue (His-159), and an asparagine residue (Asn-175; papain numbering). The geometry of the catalytic site in cathepsins is similar to that in serine proteases where the cysteine residue is replaced by a serine, and the asparagine by an aspartate residue. Cys-25 and His-159 form an ion pair which is stabilized by Asn-175. In contrast to serine proteases, the ion pair is already present in the ground state of the enzyme and does not depend on the interaction with a substrate. Therefore, cysteine proteases can be regarded as “a priori” activated enzymes (Polgar and Halasz, 1982). The thiolate cysteine leads a nucleophilic attack on the carbonyl carbon of Peptidases
Contributed by Dieter Brömme Current Protocols in Protein Science (2000) 21.2.1-21.2.14 Copyright © 2000 by John Wiley & Sons, Inc.
21.2.1 Supplement 21
Papain-like Cysteine Proteases
21.2.2
Supplement 21
Current Protocols in Protein Science
Cathepsin B (human) Cathepsin L (human) Cathepsin H (human) Cathepsin I (rabbit) Cathepsin S (human) Cathepsin C (human) DPPI (human) Cathepsin J (rat) Cathepsin O (human) Cathepsin K (human) Cathepsin O (human) Cathepsin O2(human) Cathepsin X (human) OC2 (rabbit) Cathepsin W (human) Lymphopain (human) Cathepsin V (human) Cathepsin U (human) Cathepsin L2 (human) Cathepsin F (human) Cathepsin Z (human) Cathepsin X (human) Cathepsin P (human) Cathepsin Y (rat) Cathepsin P (mouse) Cathepsin J (mouse) Cathepsin Q (rat) 86 99
108 96
251 41
95 104
21 15
19 17
19 20
18 20
P43234 NM_000396 U13665 S79895 U20280 P43236 AAC32181 P56202 BAA25909 AAC23598 O60911 AAD41790 NP_001327 AAC61477 AAC63141 BAA82844 AAD41898 AAF13142 AAF01247
98 209
16 21
A42482 P53634
62 96 98
Prodomain
17 17 17
Signal peptide
219
221
214 242
221
249
214 215
217 233
260 220 220
Mature domain
Length (amino acids)
P07858 NP_001903 P09668
Gene accession number
Mammalian Cathepsins
Protease and synonyms
Table 21.2.1
342
334
484 303
334
376
321 329
331 463
339 333 335
Prepro-enzyme
11.q13.1-3 20q13
9q21
11q13.1-3
4q31-32 1q21
1q21 11q14.1-3
8q22-23 9q21-22 15q24-25
Chromosomal localization
Table 21.2.2 Protein Sequence Identities and Similarities (%) of Mature Human Cathepsins and the Plant Protease Papain (protein sequence similarities are given in parenthesis)
Cathepsin
L
L
100
V S K H
V
S
K
H
80 55 58 45 (88) (73) (74) (63) 100 56 57 46 (73) (72) (61) 100 58 44 (71) (60) 100 47 (61) 100
F W O B Z C Papain the scissile bond of the bound substrate, which resolves via a tetrahedral intermediate into a covalently bound acyl enzyme under release of the first product (acylation). This step is followed by hydrolysis of the acyl enzyme with water, forming a second tetrahedral intermediate that finally splits into free enzyme and second product (deacylation). The formation and cleavage of tetrahedral intermediates are accompanied by appropriate transition states. The sequence of the reaction is depicted in Figure 21.2.3A. The catalytic site is located in the center of the substrate binding region, which is responsible for the substrate specificity of individual enzymes. Originally, Schechter and Berger (1967) defined seven binding sites (designated as subsites) for amino acids in substrates, with four sites binding amino acid residues N-terminal with respect to the scissile bond (S1, S2, etc.) and three sites in the C-terminal direction (S1′, S2′, etc.). The substrate residues are designated as P1, P2, etc., and P1′, P2′, etc., residues (Fig. 21.2.3B). In all cathepsins where a three-dimensional structure is known, the S2
F
W
O
B
Z
C
Papain
41 (57) 42 (57) 36 (54) 41 (56) 40 (54) 100
31 (49) 33 (49) 31 (49) 32 (50) 32 (49) 37 (53) 100
37 (52) 37 (51) 37 (53) 36 (50) 35 (52) 34 (51) 26 (41) 100
29 (43) 29 (43) 31 (47) 27 (44) 29 (47) 24 (42) 25 (41) 26 (39) 100
32 (48) 32 (49) 32 (47) 30 (47) 30 (42) 28 (42) 23 (36) 31 (44) 28 (41) 100
36 (50) 35 (50) 33 (51) 32 (46) 39 (54) 32 (49) 28 (42) 29 (45) 34 (48) 33 (45) 100
40 (54) 43 (56) 43 (57) 44 (57) 41 (55) 41 (51) 31 (46) 31 (47) 29 (41) 26 (40) 33 (48) 100
subsite forms a well-defined binding pocket which defines the so-called primary specificity of the enzyme. The other subsites are less structurally defined and form, at best, shallow groves on the surface of the protein. With the exception of cathepsin C which forms a tetramer (Dolenc et al., 1995), all other cathepsins are monomeric.
SUBCELLULAR AND TISSUE LOCALIZATION OF CATHEPSINS Cathepsins are known as acidic lysosomal proteases. All cathepsins are expressed as preproenzymes. The preregion which represents a typical signal sequence of ∼14 to 21 amino acids is responsible for the importation of the proteases into the endoplasmic reticulum (see Table 21.2.1). Moreover, all cathepsins possess putative N-linked mannose-6-phosphate glycosylation sites that are used to target proteases into lysosomes. The only nonlysosomal human papain-like cysteine protease is cathepsin W, which appears to be retained to the endoplasmic reticulum (Wex et al., 1998). Peptidases
21.2.3 Current Protocols in Protein Science
Supplement 21
hCTSLmat hCTSVmat hCTSSmat hCTSKmat hCTSHmat hCTSFmat hCTSWmat hCTSOmat hCTSBmat hCTSZmat hCTSCmat mCTSPmat rCTSQmat
A L L A Y A V L L L L L L
P P P P P P P P P P P P P
R K D D P P F L A K T D K
S S S S S E S R S S S Y F
V V V V V W C F F W W K V
D D D D D D D D D D D D D
W W W Y W W W W A W W W W
R R R R R R R R R R R R R
10 E K K K E K K K K K S K K V D K E Q N V N V E E N E
G G G G G G A Q W D H G G
N G P G G -
Q V I -
C N N -
P -
Y Y C Y F A A V T Y F Y Y
V V V V V V I V I A V V V
T T T T S T S T K S S T T
20 P V P V E V P V P V K V P I Q V E I I T P V P V R V
K K K K K K K R R R R R R
N N Y N N D D N D N N N K
Q Q Q Q Q Q Q Q Q Q Q Q Q
G K G G G G K Q G H A G G
Q Q S Q A M N M S I S K G
P -
Q -
Y -
30 C G C G C G C G C G C G C N C G C G C G C G C G C S
S S A S S S C G S S S S S
C C C C C C C C C C C C C
W W W W W W W W W W Y W W
A A A A T A A A A A S A A
F F F F F F M F F H F F F
S S S S S S A S G A A A P
A A A S T V A V A S S A V
T T V V T T A V V T M A T
40 G A G A G A G A G A G N G N G A E A S A G M G A G A
L L L L L V I V I M L I I
E E E E E E E E S A E E E
G G A G S G T S D D A G G
Q Q Q Q A Q L A R R R Q Q
M M L L I W W Y I I I M M
F F K K A F R A C N R F F
R R L K I L I I I I I W K
K K K K A N S K H K L K K
50 T T T T T Q F G T N R K T N T T -
hCTSLmat hCTSVmat hCTSSmat hCTSKmat hCTSHmat hCTSFmat hCTSWmat hCTSOmat hCTSBmat hCTSZmat hCTSCmat mCTSPmat rCTSQmat
G -
A A N -
G G G G G G W K H W S G G
R K K K K T D P V P Q N K
L L L L M L F L S S T L L
I V V L L L V E V T P T I
S S T N S S D D E L I P P
L L L L L L V L V L L L L
S S S S A S S S S S S S S
60 E E A P E E V V A V P V V
Q Q Q Q Q Q H Q E Q Q Q Q
N N N N Q E E Q D N E N N
L L L L L L L V L V V L L
V V V V V L L I L I V L I
D D D D D D D D T D S D D
C C C C C C C C C C C C C
S S S V A D G S C G S S S
T S G -
70 G P R P E K E N Q D - - - - S - - K T K P
Q G Q G Y G D G F N K M R C Y N M C - N Q Y V G Q G
N N N N D G N G A A N N
E Q K Y K D Y D G Q K R
G G G G A G G G S G G G
C C C C C C C C C C C C C
N G N G N G G G Q G M G H G N G N G E G E G Q S LW
80 G L G F G F G Y G L G L G F G S G Y G N G F G T G N
M M M M P P V T P D P A T
D A T T S S W L A L Y H Y
Y R T N Q N D N E S L Q N
A A A A A A A A A V I A A
F F F F F Y F L W W A F F
Q Q Q Q E S I N N D G E Q
Y Y Y Y Y A T W F Y K Y Y
V V I V I I V L W A Y V V
90 Q D K E I D Q K L Y K N L N N K T R H Q A Q L K L H
N N N N N L N M K H D N N
G -
L -
V -
S -
G G K R K G S Q G G F K G
G G G G G G G V G I G G G
K L P -
100 L D L D I D I D I M L E L A L V Y E D E L V L E L E
hCTSLmat hCTSVmat hCTSSmat hCTSKmat hCTSHmat hCTSFmat hCTSWmat hCTSOmat hCTSBmat hCTSZmat hCTSCmat mCTSPmat rCTSQmat
S S S S G T S K S T E A A
E E D E E E E D H C E E E
E E A D D D K S V N A A A
G -
S S S A T D D E C C T T
Y Y Y Y Y Y Y Y R F Y Y
P P P P P S P P P N P P P
Y Y Y Y Y Y F F Y Y Y Y Y
110 E A V A K A V G Q G Q G Q G K A S I Q A T G E G E R
T V M Q K H K Q P K T K K
E D D E D M V N P D D D E
R C Q -
A E E -
H C -
H D -
V K -
N F -
120 - - - - - - - - G S N Q - - - -
R C -
E E Q E G Q H G P G S G G
S I K S Y S R L P T P P V
C C C C C C C C C C C C C
K K Q M K N H H T N K R R
Y Y Y Y F F P Y G E M Y Y
N R D N Q S K F E F K R N
P P S P P A K S G K E S P
130 K Y E N K Y T G G K E K Y Q G S D T E C D C E N K N
S S R K A A K H P H F A S
V V A A I K V S K A R S S
G C I -
A A A A G V A F S R Y A A
N N T K F Y W S K N Y N K
D D C C V I I I I Y S I I
T T S R K N Q K C T S T T
G G K G D D D G E L E D G
140 F V F T Y T Y R V A S V F I Y S P G WR Y H Y V F V
D V E E N E M A Y V Y N V
I V L I I L L Y S G V L L
P A P P T S Q D P D G P P
K P Y E I Q N F T Y G P E
G G G Y S Y G F -
D K S Y -
Q L G -
D S -
150 - - - - - - - - K H - - - - -
hCTSLmat hCTSVmat hCTSSmat hCTSKmat hCTSHmat hCTSFmat hCTSWmat hCTSOmat hCTSBmat hCTSZmat hCTSCmat mCTSPmat rCTSQmat
Y -
G -
Y -
N -
S -
Y -
S -
V -
160 - - - - - - - - S N - G C - - -
Q K R N D N N Q S G N N S
E E E E E E E E E R E E E
K K D K E Q H D K E A L D
A A V A A K R E D K L Y V
LM LM L K L K M V L A I A M A I M MM M K LW LM
K K E R E A Q K A A L V D
A A A A A W Y A E E E A A
170 V A V A V A V A V A L A L A L L I Y I Y L V V A V A
T T N R L K T T K A H S T
V V K V Y R Y F N N H I K
G G G G N G G G G G G G G
P P P P P P P P P P P P P
I I V V V I I L V I M V I
S S S S S S T V E S A S A
V V V V F V V V G C V A T
A A G A A A T I A G A A G
180 I D M D V D I D F E I N I N V D F S I M F E I D V H
A A A A V A M A V A V A V
G G R S T F K V Y T Y S I
H H H L Q G P S S E D H S
E S P T D M L W D R D D S
S S S S Q Q S S
F F F F F F L F L F F F
L Q F Q M Y Y L A L R R
F F L F M R R Q L N H F F
190 Y K Y K Y R Y S Y R H G K G D Y Y K Y T Y K Y N Y Q
E S S K T I V L S G K G K
G G G G G S I G G G G G G
I I V V I R K G V I I I V
Y Y Y Y Y P A I Y Y Y Y Y
F F Y Y S L T I Q A H Y H
E E E D S R P Q H E H E E
P P P E T P T H V Y T P P
D D S S S L T H T Q G N K
200 C S C S C T C N C H C S C D C S G E D T L R C S C S
hCTSLmat hCTSVmat hCTSSmat hCTSKmat hCTSHmat hCTSFmat hCTSWmat hCTSOmat hCTSBmat hCTSZmat hCTSCmat mCTSPmat rCTSQmat
S S S K P P S M T D S S
T P -
P F -
N -
P -
F -
E K Q D D W Q G E Y Y
D N N N K L L E M Y L F -
210 M D L D V N L N V N I D V D A N G G I N T N V N V N
H H H H H H H H H H H H H
G G G A A A S A A V A A A
V V V V V V V V I V V V V
L L L L L L L L R S L L L
V V V A A L L I I V L V V
V V V V V V V T L A V V V
G G G G G G G G G G G G G
Y Y Y Y Y Y F F W W Y Y Y
220 G F G F G D G I G E G N G S D K G V G I G T G S G F
E E L Q K R V T E S D E E
S G K S G G
T A S D N
E N E V E
S S E K T
G -
I -
W -
230 - - - - - - A E - - - - - - -
T -
V -
S -
S -
Q -
S -
Q -
P -
240 - - - - - - Q P - - - - - - -
P A -
D N N K N S H G N D S D D
N N G G G D P S G G G G G
N S K N I V T T T T M N N
K K E K P P P P P E D N N
Y Y Y H Y F Y Y Y Y Y Y Y
W W W W W W W W W W W W W
L L L I I A I I L I I L L
250 V K V K V K I K V K I K L K V R V A V R V K I K I K
hCTSLmat hCTSVmat hCTSSmat hCTSKmat hCTSHmat hCTSFmat hCTSWmat hCTSOmat hCTSBmat hCTSZmat hCTSCmat mCTSPmat rCTSQmat
N N N N N N N N N N N N N
S S S S S S S S S S S S S
W W W W W W W W W W W W W
G G G G G G G G N G G G G
E P H E P T A S T E T E K
E E N N Q D Q S D P G E R
W W F W W W W W W W W W W
G G G G G G G G G G G G G
260 M G S N E E N K M N E K E K V D D N E R E N M N L R
G G G G G G G G G G G G G
Y Y Y Y Y Y Y Y F W Y Y Y
V V I I F Y F A F L F M M
K K R L L Y R H K R R Q K
M I M M I L L V I I I I I
A A A A E H H K L V R A A
K K R R R R R M R T R K K
D D N N G G G G G S G D D
270 R R K N K G K N K S S S Q T Y T H N R N
K -
N N N N N G N N D D D N N
H H H A M A T V H G E H H
C C C C C C C C C K C C C
G G G G G G G G G G A G A
I I I I L V I I I A I I I
A A A A A N T A E R E A A
S T S N A T K D S Y S S S
280 A A A A F P L A C A M A F P S V E V N L I A L A L A
S S S S S S L S V A V S Q
Y Y Y F Y S T S A I A Y Y
P P P P P A A I G E A P P
T N E K I V R F I E T N T
V V I M P V V V P H P I V
290
300
L V D Q K P D M K P R V S C P P R T D Q Y W E K I C T F G D P I V I P K L F
21.2.4 Supplement 21
Current Protocols in Protein Science
cathepsin V cathepsin L cathepsin P 97
cathepsin Q cathepsin S
93 cathepsin K
cathepsin H cathepsin F 93 cathepsin W
98
cathepsin C 55
88
cathepsin B
60
62 55
cathepsin Z cathepsin O
Figure 21.2.2 Phylogenetic dendrogram of mammalian cathepsins based on their amino acid sequences of the mature proteases. Support for the internal branches of the tree topology is shown in percent.
The tissue distribution of cathepsins varies from ubiquitous to highly tissue-specific. The originally described cathepsins B, L, and H are characterized by a widespread expression throughout most tissues, suggesting housekeeping functions. A similar ubiquitous tissue distribution has been observed for cathepsins O, F, Z, and C (Velasco et al., 1994; Rao et al., 1997; Santamaria et al., 1998; Wang et al., 1998). In contrast, the expression of cathepsins S, V, K, and W is restricted to well-defined tissues. Cathepsin S was the first papain-like protease identified to show a specific tissue distribution. It is predominantly expressed in lymphatic tissues and immune-related cell types such as those found in lymph nodes, spleen, and antigen-presenting cells (Kirschke
et al., 1986; Chapman et al., 1997). The closely related cathepsin K is predominantly expressed in bone-degrading osteoclasts as well as related giant multinucleated cells, and constitutes a critical protease in bone resorption (Brömme and Okamoto, 1995; Drake et al., 1996). However, at lower expression levels, cathepsin K is also expressed in various epithelial cells found in the bronchia, bile duct, urethra, and thyroid gland (Bühling et al., 1999; Haeckel et al., 1999). Cathepsin V, which shares an 80% sequence identity with the ubiquitously expressed cathepsin L, is specifically expressed in the thymus, testis, and retina (Adachi et al., 1998; Brömme et al., 1999). The most celltype-specific C1 cysteine protease is cathepsin W. Cathepsin W is exclusively expressed in
Figure 12.2.1 (at left) Multiple amino acid alignment of mammalian mature cathepsins. The amino acid sequences have been extracted from GenBank (see accession numbers in Table 21.2.1). Multiple sequence alignment was performed using ClustalW of MacVector (Oxford Molecular Group, PLC). Identical amino acid residues are darkly shaded, similar amino acids are lightly shaded, and unrelated residues have a white background. Peptidases
21.2.5 Current Protocols in Protein Science
Supplement 21
A
tetrahedral intermediate _ free enzyme forms ion-pair
O R
R C
C substrate
Cys25-S _
N H
O N H
Cys25-S
R′
+H N
R′
+H N
His159
His159
NH
NH
R C
H2N
O
R′
Cys25-S
H2O
N His159
NH acyl-enzyme
B
substrate H2N
P4
P3
S4
S3
P2
P1
P1′
P2′
P3′
S1
S1′
S2′
S3′
COOH
S2 cathepsin
Figure 21.2.3 (A) Mechanism of substrate hydrolysis by papain-like cysteine proteases. The thiol group of the free enzyme attacks the carbonyl carbon of a substrate’s peptide bond and forms an unstable tetrahedral intermediate which disintegrates into an acyl enzyme under release of the C-terminal portion of the substrate. Subsequently, the acyl enzyme hydrolyzes into the free enzyme and the N-terminal portion of the substrate. (B) Substrate binding region of cathepsins. Protease subsites are designated as S1, S2, S3, S4, S1′, S2′, and S3′ and the appropriate amino acid positions in the substrate are designated as P1, P2, P3, P4, P1′, P2′, and P3′ respectively. The scissile bond in substrates is between P1 and P1′.
Papain-like Cysteine Proteases
21.2.6 Supplement 21
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Table 21.2.3
Activity, Tissue Distribution, and Potential Function of Mammalian Cathepsins
Protease
Activity
Tissue distribution
Primary function
Cathepsin B Cathepsin L Cathepsin H Cathepsin O Cathepsin F Cathepsin C
Endo- and exopeptidase Endopeptidase Endo- and exopeptidase Endopeptidase Endopeptidase Exopeptidase
Ubiquitous Ubiquitous Ubiquitous Ubiquitous Ubiquitous Ubiquitous
Cathepsin Z
Exopeptidase
Ubiquitous
Cathepsin S
Endopeptidase
Cathepsin K Endopeptidase Cathepsin V Endopeptidase Cathepsin W Not known at this time
Specific (antigen presenting cells) Specific (osteoclasts) Specific (thymus, testis Specific (NK cells)
Housekeeping Housekeeping Housekeeping Housekeeping Housekeeping Housekeeping and granzyme processing Housekeeping and processing? Antigen presentation
Cathepsin P Cathepsin Q
Specific (placenta) Specific (placenta)
Not known at this time Not known at this time
cytotoxic lymphocytes and natural killer cells (Linnevers et al., 1997b; Brown et al., 1998; Wex et al., 1998). Table 21.2.3 summarizes the expression pattern and the potential function of human C1-family cathepsins. For mouse cathepsin P and rat cathepsin Q, human orthologs are presently not known (Sol-Church et al., 1999, 2000).
BIOLOGICAL AND INTRINSIC SUBSTRATE SPECIFICITY B iolo gical substrate specificity of cathepsins is determined by: (1) the intrinsic substrate specificity of the individual protease based on structural properties; (2) the selective expression of individual cathepsins in specialized cells; and (3) the accessibility of specific substrates to the protease. All three features are equally important for the understanding of the substrate specificity of cysteine-dependent cathepsins. Cells with general metabolic functions are likely to express ubiquitously distributed housekeeping proteases such as cathepsins L, B, and H, but probably also F, C, and Z. Cathepsins L and F are very potent endopeptidases with no known in vivo preference for physiologically relevant substrates. Their second-order rate constants towards small synthetic peptide substrates are the highest meas-
Bone resorption Antigen presentation? Cytotoxic function in NK cells? Not known at this time Not known at this time
ured amongst known cathepsins (Wang et al., 1998). Cathepsins B, H, C, and Z are primarily exopeptidases, although all of them also show endopeptidase activities to a various degree. Cathepsins B and Z are carboxypeptidases, with cathepsin B being preferentially a dipeptidyl carboxypeptidase (Aronson and Barrett, 1978; Nagler et al., 1999). In contrast, cathepsins H and C are aminopeptidases, with cathepsin C acting as a dipeptidyl aminopeptidase (Lindley, 1972; Takahashi et al., 1988). Combined endopeptidase and exopeptidase activities are best suited for the terminal protein breakdown within lysosomes. However, the presence of ubiquitously expressed cathepsins in specialized cells may indicate a highly specific function for a rather nonspecific protease. For example, it has been well demonstrated that cathepsin C is responsible for processing of cytotoxic lymphocyte resident granzyme precursors whose activation requires the removal of an N-terminal activation peptide (Pham and Ley, 1999). Thus, the specific function of cathepsin C in cell-mediated immunity is based on the cathepsin’s general ability to cleave off N-terminal dipeptides and the collective presence of cathepsin C and the zymogens of granzyme substrates in highly specialized granula of cytotoxic lymphocytes. Cathepsin K is an example where biological substrate specificity Peptidases
21.2.7 Current Protocols in Protein Science
Supplement 21
Table 21.2.4
Selected Substrates of Mammalian Cathepsinsa
Protease
Protein substrates
Cathepsin B
Collagens (telopeptide release), fibrinogen, fibronectin, laminin, osteocalcin, osteonectin, glucagon, fructose-1,6-bisphosphate aldolase, proinsulin, thyroglobulin, proteoglycans Invariant chain, collagens (telopeptide release), elastin, fibronectin, laminin, osteonectin, osteocalcin, proteoglycans, glucagon, insulin, fructose-1,6bisphosphate aldolase Invariant chain, elastin, proteoglycans, gelatin Type I collagen, type II collagen, osteonectin, osteopontin, proteoglycans, elastin, gelatin Invariant chain Invariant chain Progranzymes, angiotensin, secretin, glucagon Albumin, prorenin, α1-antitrypsin, fibrinogen, angiotensin, bradykinin
Cathepsin L
Cathepsin S Cathepsin K Cathepsin V Cathepsin F Cathepsin C Cathepsin H
aThe interested reader is referred to Kirschke et al. (1995) for more detail.
Papain-like Cysteine Proteases
Table 21.2.5
S2 Constituting Amino Acids of Human Cathepsins L, V, S, K, F, and B
Cathepsin L
Cathepsin V
Cathepsin S
Cathepsin K
Cathepsin F
Cathepsin B
Leu69 Met70 Ala135 Met161 Ala214
Phe69 Met70 Ala135 Leu161 Ala214
Phe70 Met71 Gly137 Val162 Phe211
Tyr67 Met68 Ala134 Leu160 Leu209
Leu67 Pro68 Ala133 Ile152 Met207
Tyr75 Pro76 Ala173 Gly Glu245
is defined by the enzyme’s unique ability to degrade specific substrates such as type I and II collagens (Garnero et al., 1998, Kafienah et al., 1998), its predominant expression in specialized collagen-degrading osteoclasts (Brömme and Okamoto, 1995; Drake et al., 1996), and the abundant presence of collagen substrates in bone and cartilage. In contrast, the specific processing of regulatory proteins such as the invariant chain of MHC class II complexes depends primarily on the selective expression of individual cathepsins in antigenpresenting cells, and not on a unique specificity of the proteases towards the invariant chain. Most cathepsins with endopeptidase activity are able to selectively degrade the invariant chain (D. Brömme, unpub. observ.), but only cathepsin S plays a biologically relevant role in MHC class II antigen presentation in bone marrow–derived APCs, due to its selective expression in these cells (Riese et al., 1996; Shi et al., 1999). In contrast, in thymic epithelial cells, cathepsin L appears to be responsible for invariant chain processing (Nakagawa et al.,
1998). Table 21.2.4 lists selected protein substrates hydrolyzed by cathepsins. The intrinsic specificity of cathepsins is determined by their three-dimensional structure, which is responsible for the geometry and accessibility of the substrate binding region. The best characterized subsite in cathepsins is S2, which forms a deep pocket accommodating the P2 substrate amino acid residue. Table 21.2.5 shows the critical amino acid residues forming the S2 subsite in cathepsins L, V, S, K, F, and B, and Figure 21.2.4 displays the S2-P2 specificity of these proteases to a substrate series with varying P2 residues. All cathepsins listed in Table 21.2.5 except cathepsins K and S prefer an aromatic phenylalanine over a leucine residue in their S2 subsite. A replacement of Gly137 in cathepsin S by an alanine residue, which is present in other papain-like proteases, leads to a reversal of the preference of cathepsin S for leucine over the phenylalanine residue (Brömme et al., 1994). However, the presence of an alanine in this position cannot solely explain the preferred acceptance of aromatic
21.2.8 Supplement 21
Current Protocols in Protein Science
1.2
1.0
Relative activity
0.8
0.6
0.4
0.2
0.0
F L V R F L V R F L V R F L V R F L V R F L V R CTSL CTSV CTSS CTSK CTSF CTSB
Figure 21.2.4 Substrate P2 amino acid preferences for cathepsins L, V, K, S, F, and B. kcat/Km values for the cathepsin-mediated hydrolysis of Z-X-Arg-MCA (X = Phe, Leu, Val, Arg). Values were normalized to the best substrate = 1.
residues in the P2 position since cathepsin K has also an alanine in this position and clearly prefers leucine over phenylalanine. Three-dimensional structural analysis revealed that Leu-209 in cathepsin K would prevent optimal binding of an aromatic residue (McGrath et al., 1997). Leu-209 is located at the bottom of the S2 subsite and is the ortholog residue to Glu245 in cathepsin B. The negative charge of Glu-245 is responsible for the binding of basic residues such as arginine or lysine in the S2 subsite of cathepsin B, a residue excluded in S2 for all other cathepsins (Hasnain et al., 1993). The exopeptidase activity of cathepsins B and H is explained by additional protein loops in their respective structures. Residues 184 to 197 in cathepsin B forms an occluding loop which blocks an endopeptidase-efficient binding of the enzyme to the substrate. Two histidine residues residing in the middle of the occluding loop interact with the carboxyl group of the
C-terminus of substrates, and thus favor the dipeptidyl carboxypeptidase activity (Musil et al., 1991). In cathepsin H, an octapeptide derived from the proregion protrudes backwards into the S2 subsites, and thus only allows the occupation of the S1′ and S1 subsites, explaining the enzyme’s aminopeptidase activity (Guncar et al., 1998).
CATHEPSINS AND DISEASES The ability of cathepsins to efficiently degrade extracellular matrix (ECM) proteins suggests their involvement in degenerative diseases and cancer. Depending on the cell types involved in ECM degradation, ubiquitously expressed proteases such as cathepsins L and B, but also selectively expressed proteases such as cathepsins K and S, may contribute to tissue destruction. Numerous reports in the literature describe circumstantial evidence that cathepsins B and L contribute to tumor progresPeptidases
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Papain-like Cysteine Proteases
sion and metastasis. This evidence is based on observations describing an increased secretion of procathepsins B and L by various tumor cell lines, peripheral accumulation of cathepsin B in tumor cells at the site of invasion, inhibition of matrix invasion by general and cathepsin B–specific inhibitors, and is corroborated by specific antibodies against cathepsin L in tumor mouse models (Rozhin et al., 1994; Weber et al., 1994; De Stefanis et al., 1997; Mai et al., 2000). Although there is ample evidence that links cathepsins B and L to tumor progression, reported results frequently show poor correlation of cathepsin expression levels to indicators of tumor invasiveness, including shortened survival, recurrence, and tumor stage. However, it has been clearly shown that cathepsin B is found at higher levels in a number of human cancers (i.e., colorectal, lung, and bladder), suggesting that it may be useful as a diagnostic and prognostic marker (Koblinski et al., 2000). Recent progress in the understanding of proteolytic mechanisms of bone remodeling have clearly implicated cathepsin K as a critical cysteine protease in normal and pathological bone resorption. Bone resorption is a two step process of mineral solubilization and protein matrix degradation by osteoclasts. As described above, cathepsin K is the major proteolytic component of osteoclasts and has a potent and unique collagenolytic activity. Its specific inhibition leads to a reduction of bone resorption in osteoclast pit assays and to a decrease of surrogate markers of bone erosion in osteoporosis models (Thompson et al., 1997; Xia et al., 1999). However, the most convincing fact confirming the critical role of cathepsin K in bone remodeling was the finding that the autosomal recessive bone sclerosing disorder, pycnodysostosis (OMIM 265800), resulted from the deficient activity of cathepsin K (Gelb et al., 1996). On the molecular level, deficiency in cathepsin K activity leads to an accumulation of undigested collagen fibrils in osteoclastic lysosomes and to an increase in demineralized areas underneath osteoclasts as seen in pycnodysostosis (Everts et al., 1985). Besides their role in ECM degradation, cathepsins have been implicated in various aspects of MHC class II-related antigen processing and presentation. Before launching a T cell response to an antigen, the latter must be processed and presented by antigen-presenting cells (APC). The processing of the antigen is probably mediated by aspartic as well as cysteine proteases. Using specific cathepsin B inhibi-
tors, it has been convincingly shown that cathepsin B is essential for the processing of selected antigens in macrophages (Katunuma et al., 1994). Before the processed antigen can be presented, the MHC class II-complex must be freed from the invariant chain which blocks the groove of the antigen binding site. The removal of the invariant chain is catalyzed by cathepsin S in bone-marrow derived APCs (Shi et al., 1999), and probably by cathepsin L in thymic epithelial cells (Nakagawa et al., 1998). The finding that cathepsins S and L are essential for antigen presentation raises the possibility to selectively modulate immune responses by inhibiting these cathepsins. Recent studies have demonstrated that the abrogation of cathepsin S activity significantly reduces the inflammatory reaction in collagen-induced arthritis (Nakagawa et al., 1999). Thus, a targeted inhibition of cathepsin S might be beneficial for the treatment of arthritis and asthma, or useful in transplantation. Cathepsin C deficiency has been recently identified as the cause of the Papillon-Lefèvre syndrome or keratosis palmoplantaris with periodontopathia (OMIM 245000). Cathepsin Cdeficient patients loose their permanent teeth at age fourteen and suffer from keratosis of the palms and soles (Toomes et al., 1999). It is thought that cathepsin C deficiency leads to an impairment of the immunological properties of the tissue surrounding teeth and the processing of keratins. It is known that the failure to activate granzymes in cytotoxic lymphocytes and natural killer cells compromises the ability of these cells in the immune response against infections (Pham and Ley, 1999). To date, only cathepsins K and C deficiencies have been implicated in inherited diseases. Cathepsin knockout mice have been described for cathepsins L (Nakagawa et al., 1998), B (Deussing et al., 1998), K (Saftig et al., 1998; Gowen et al., 1999), S (Nakagawa et al., 1999; Shi et al., 1999), and C (Pham and Ley, 1999). Cathepsin null mice have been very useful tools to elucidate the functional role of cathepsins in health and disease. Cathepsins have been implicated in the following human degenerative and inflammatory diseases: osteoporosis, pycnodysostosis, osteoarthritis, rheumatoid arthritis, inflammatory bowel disease, periodontitis, muscular dystrophy, chronic obstructive pulmonary disease (COPD), acute respiratory distress syndrome (ARDS), Alzheimer’s disease, progressive myoclonus epilepsy, as well as tumor in-
21.2.10 Supplement 21
Current Protocols in Protein Science
vasion and metastasis. There is also evidence for the involvement of cathepsins in the following infectious diseases: malaria, trypanosomiasis, and schistosomiasis. In addition, parasite-expressed papain-like cysteine proteases have been demonstrated to be crucial for the viability and/or the invasion process of the pathogens. It has been shown that specific inhibitors targeted against those cysteine proteases have an anti-parasitic activity and might represent a new approach to fight against parasite-mediated diseases (McKerrow, 1999; McKerrow et al., 1999).
RECOMBINANT EXPRESSION, PURIFICATION, AND ASSAYS OF CATHEPSINS Native cathepsins are relatively easy to isolate from lysosomal preparations. Using standard gradient centrifugation methods for the isolation of lysosomes, cathepsins can be the purified to homogeneity in milligram quantities (Kirschke et al., 1977). This approach is effective for ubiquitously expressed cathepsins which are present in easily accessible organs such as the liver, kidney, or placenta. It should be noted that the majority of cathepsin activity in tissues is inhibited by endogenous cystatins when samples undergo total homogenization that includes the rupture of lysosomal membranes. In contrast, cathepsins expressed in highly specialized cells, such as cathepsin K or W in osteoclasts or natural killer cells, are difficult or impossible to isolate from natural sources, and they must be produced as recombinant enzymes. Recombinant expression of cathepsins is the most effective way to obtain sufficient amounts of protease material for crystallization, assay development, protein chemical characterization, and high-throughput screening of inhibitors. Presently, various expression systems have been established. These include the baculovirus insect cell (Brömme and McGrath, 1996), yeast (Brömme et al., 1993, Linnevers et al., 1997), E. coli (Kopitar et al., 1996; D’Alessio et al., 1999), and mammalian cell systems (Ren et al., 1996). Although the yields of active proteases obtained by these different methods are difficult to predict and vary strongly for individual enzymes, the baculovirus and the Pichia pastoris systems (UNITS 5.4-5.5 and UNIT 5.7) seem to be most effective. The Pichia pastoris expression system seems to be advantageous in two respects: (1) the low cost in terms of culture conditions, and (2) the fact that the expression host does
not express endogenous papain-like cysteine proteases which constitute a serious contamination problem in mammalian cell and baculovirus systems. A summary of recombinant protease expression techniques is given in Brömme and Schmidt (1999). Purification of active cathepsins is best performed using various column chromatography steps, which must be optimized for individual proteases. Cathepsins have been purified based on (1) their net charge, i.e., ion-exchange chromatography (Wang et al., 1998); (2) their surface properties, i.e., hydrophobic interaction chromatography (Brömme et al., 1996, 1999); or (3) by high-affinity ligands, i.e., immobilized reversible inhibitors (Rich et al., 1986). Thiopropyl Sepharose 6B chromatography is an elegant approach to purify various cathepsins (Brocklehurst et al., 1987; Brömme et al., 1993). The method is based on a disulfide formation with the free active site thiol of the enzyme at acidic pH, thus utilizing the specific ion pair present in cathepsins. Other free cysteine residues would only bind the thiolSepharose at alkaline pH. However, an efficient purification requires a washing of the columns at neutral to alkaline pH, which may inactivate some of the highly neutral pH-sensitive cysteine proteases such as cathepsin L. The activity of cathepsins is best monitored by synthetic fluorogenic substrates such as methyl coumarylamides (MCA). All known cathepsins with endopeptidase activity can be easily followed by their Z-Phe-Arg-MCA cleaving activity (e.g., cathepsins L, B, K, S, V, O, and F). Cathepsins with aminopeptidase activity are measured with substrates such as Arg-MCA (cathepsin H) or Ala-Ala-MCA (cathepsin C). Highly specific synthetic substrates for individual cathepsins are rare. ZArg-Arg-MCA is selective for cathepsin B, and Z-Gly-Pro-Arg-MCA and Z-Val-Val-ArgMCA are preferentially hydrolyzed by cathepsin K and cathepsin S, respectively (Brömme et al., 1989, 1993; Knight, 1980; Aibe et al., 1996). It should be noted that both substrates are also efficient for other cathepsins and cannot be used to unambiguously differentiate these cathepsins from related activities. A differentiation of a mix of cathepsin activities always requires various combinations of substrate and inhibitor assays. Naphthylamide substrates are useful for the localization of cathepsin activities in cryo-tissue sections or in cell cultures (van Noorden et al., 1987, Xia et al., 1999). Peptidases
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LITERATURE CITED
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Aibe, K., Yazawa, H., Abe, K., Teramura, K., Kumegawa, M., Kawashima, H., and Honda, K., 1996. Substrate specificity of recombinant osteoclast-specific cathepsin K from rabbits. Biol. Pharm. Bull. 19:1026-1031.
Bühling, F., Gerber, A., Häckel, C., Krüger, S., Köhnlein, T., Brömme, D., Reinhold, D., Ansorge, F., and Welte, T. 1999. Expression of cathepsin K in lung epithelial cells. Am. J. Respir. Crit. Care Med. 20:612-619.
Aronson, N.N.J. and Barrett, A.J. 1978. The specificity of cathepsin B. Hydrolysis of glucagon at the C-terminus by a peptidyldipeptidase mechanism. Biochem. J. 171:759-765.
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Brocklehurst, K., Kowlessur, D., O’Driscoll, M., Patel, G., Quenby, S., Salih, E., and Templeton W, Thomas, E.W., and Willenbrock, F. 1987. Substrate-derived two-protonic-state electrophiles as sensitive kinetic specificity probes for cysteine proteinases. Activation of 2-pyridyl disulphides by hydrogen-bonding. Biochem. J. 244:173-81.
D’Alessio, K.J., McQueney, M.S., Brun, K.A., Orsini, M.J., and Debouck, C.M. 1999. Expression in Escherichia coli, refolding, and purification of human procathepsin K, an osteoclast-specific protease. Protein Expr. Purif. 15:213-220.
Brömme, D. and McGrath, M.E., 1996. High level expression and crystallization of recombinant human cathepsin S. Protein Sci. 5:787-791.
De Stefanis, D., Demoz, M., Dragonetti, A., Houri, J.J., Ogier-Denis, E., Codogno, P., Baccino, F.M., and Isidoro, C. 1997. Differentiation-induced changes in the content, secretion, and subcellular distribution of lysosomal cathepsins in the human colon cancer HT-29 cell line. Cell Tissue Res. 289:109-117.
Brömme, D. and Okamoto, K. 1995. Human cathepsin O2, a novel cysteine protease highly expressed in osteoclastomas and ovary. Molecular cloning, sequencing and tissue distribution. Biol. Chem. Hoppe-Seyler 376:379-384.
Deussing, J.R.W., Saftig, P, Peters, C., Ploegh, H.L., and Villadangos, J.A. 1998. Cathepsins B and D are dispensable for major histocompatibility complex class II-mediated antigen presentation. Proc. Natl. Acad. Sci. U.S.A. 95:4516-4521.
Brömme, D. and Schmidt, B.F. 1999. Functional expression of recombinant proteases. In Proteolytic Enzymes—Tools and Targets (E.E. Sterchi and W. Stöcker, eds.) Springer-Verlag, New York.
Dolenc, I., Turk, B., Pungercic, G., Ritonja, A., and Turk, V. 1995. Oligomeric structure and substrate induced inhibition of human cathepsin C. J. Biol. Chem. 270:21626-21631.
Brömme, D., Steinert, A., Friebe, S., Fittkau, S., Wiederanders, B., and Kirschke, H. 1989. The specificity of bovine spleen cathepsin S. A comparison with liver cathepsins L and B. Biochem. J. 264:475-481. Brömme, D., Bonneau, P.R., Lachance, P., Wiederanders, B., Kirschke, H., Peters, C., Thomas, D.Y., Storer, A.C., and Vernet, T. 1993. Functional expression of human cathepsin S in Saccharomyces cerevisiae. Purification and characterization of the recombinant enzyme. J. Biol. Chem. 268:4832-4838. Brömme, D., Bonneau, P.R., Lachance, P., and Storer, A.C. 1994. Engineering the S2 subsite specificity of human cathepsin S to a cathepsin L-and cathepsin B-like specificity. J. Biol. Chem. 269:30238-30242. Brömme, D., Okamoto, K., Wang, B.B., and Biroc, S. 1996. Human cathepsin O2, a matrix proteindegrading cysteine protease expressed in osteoclasts. Functional expression of human cathepsin O 2 in Spodoptera frugiperda and characterization of the enzyme. J. Biol. Chem. 271:2126-2132. Papain-like Cysteine Proteases
tial, enzymatic characterization, and chromosomal localization. Biochemistry 38:2377-2385.
Adachi, W., Kawamoto, S., Ohno, I., Nishida, K., Kinoshita, S., Matsubara, K., and Okubo, K., 1998. Isolation and characterization of human cathepsin V: A major proteinase in corneal epithelium. Investig. Ophthalmol. Vis. Sci. 39: 17891796.
Brömme, D., Li, Z., Barnes, M., and Mehler, E. 1999. Human cathepsin V functional expression, tissue distribution, electrostatic surface poten-
Drake, F.H., Dodds, R.A., James, I.E., Connor, J.R., Debouck, C., Richardson, S., Lee-Rykaczewski, E., Coleman, L., Rieman, D., Barthlow, R., Hastings, G., and Gowen, M. 1996. Cathepsin K, but not cathepsins B, L, or S, is abundantly expressed in human osteoclasts. J. Biol. Chem. 271:1251112516. Everts, V., Aronson, D.C., and Beertsen, W. 1985. Phagocytosis of bone collagen by osteoclasts in two cases of pycnodysostosis. Calcif. Tissue Int. 37:25-31. Garnero, P., Borel, O., Byrjalsen, I., Ferreras, M., Drake, F.H., McQueney, M.S., Foged, N.T., Delmas, P.D., and Delaisse, J.M. 1998. The collagenolytic activity of cathepsin K is unique among mammalian proteinases. J. Biol. Chem. 273:32347-32352. Gelb, B.D., Shi, G.P., Chapman, H.A., and Desnick, R.J. 1996. Pycnodysostosis, a lysosomal disease caused by cathepsin K deficiency. Science 273:1236-1238. Gowen, M., Lazner, F., Dodds, R., Kapadia, R., Feild, J., Tavaria, M., Bertoncello, I., Drake, F., Zavarselk, S., Tellis, I., Hertzog, P., Debouck, C., and Kola, I.. 1999. Cathepsin K knockout mice develop osteopetrosis due to a deficit in matrix
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degradation but not demineralization. J. Bone Miner. Res. 14:1654-63.
preliminary crystallographic studies of an inhibitor complex. Protein Sci. 6:919-921.
Guncar, G., Podobnik, M., Pungercar, J., Strukelj, B., Turk, V., and Turk, D. 1998. Crystal structure of porcine cathepsin H determined at 2.1 A resolution: Location of the mini-chain C-terminal carboxyl group defines cathepsin H aminopeptidase function. Structure 6:51-61.
Mai, J., Waisman, D.M., and Sloane, B.F. 2000. Cell surface complex of cathepsin B/annexin II tetramer in malignant progression. Biochim. Biophys. Acta 1477:215-230.
Haeckel, C., Krueger, S., Buehling, F., Brömme, D., Franke, K., Schuetze, A., Roese, I., and Roessner, A. 1999. Expression of cathepsin K in the human embryo and fetus. Dev. Dyn. 216:89-95. Hasnain, S., Hirama, T., Huber, C.P., Mason, P., and Mort, J.S. 1993. Characterization of cathepsin B specificity by site-directed mutagenesis. Importance of Glu245 in the S2-P2 specificity for arginine and its role in transition state stabilization. J. Biol. Chem. 268:235-240. Kafienah, W., Brömme, D., Buttle, D.J., Croucher, L.J., and Hollander, A.P. 1998. Human cathepsin K cleaves native type I and II collagens at the N-terminal end of the triple helix. Biochem. J. 331:727-732. Katunuma, N., Matsunaga, Y., and Saibara, T. 1994. Mechanism and regulation of antigen processing by cathepsin B. Adv. Enzyme Regul. 34:145-158. Kirschke, H., Langner, J., Wiederanders, B., Ansorge, S., and Bohley, P. 1977. Cathepsin L. A new proteinase from rat-liver lysosomes. Eur. J. Biochem. 74:293-301. Kirschke, H., Schmidt, I., and Wiederanders, B. 1986. Cathepsin S. The cysteine proteinase from bovine lymphoid tissue is distinct from cathepsin L (EC 3.4.22.15). Biochem. J. 240:455-459. Kirschke, H, Barrett, A.J., and Rawlings, N.D. 1995. Proteinases 1: Lysosomal cysteine proteinases. In Protein Profiles 2 (14):1588-1643, Academic Press. Knight, C.G. 1980. Human cathepsin B. Application of the substrate N-benzyloxycarbonyl-L-arginyl-L-arginine2-naphthylamide to a study of the inhibition by leupeptin. Biochem. J. 189(3):447-453. Koblinski, J.E., Ahram, M., and Sloane, B.F. 2000. Unraveling the role of proteases in cancer. Clin. Chim. Acta 291:113-135. Kopitar, G., Dolinar, M., Strukelj, B., Pungercar, J., and Turk, V. 1996. Folding and activation of human procathepsin S from inclusion bodies produced in Escherichia coli. Eur. J. Biochem 236:558-562. Lindley, H. 1972. The specificity of dipeptidyl aminopeptidase I (cathepsin C) and its use in peptide sequence studies. Biochem. J. 126:683-685. Linnevers, C., Smeekens, S.P., and Brömme, D. 1997a. Human cathepsin W, a putative cysteine protease predominantly expressed in CD8+ Tlymphocytes. FEBS Lett. 405:253-259. Linnevers, C.J., McGrath, M.E., Armstrong, R., Mistry, F.R., Barnes, M., Klaus, J.L., Palmer, J.T., Katz, B.A., and Brömme, D. 1997b. Expression of human cathepsin K in Pichia pastoris and
McGrath, M.E., Klaus, J.L., Barnes, M.G., and Brömme, D. 1997. Crystal structure of human cathepsin K complexed with a potent inhibitor. Nature Struct. Biol. 4:105-109. McKerrow, J.H. 1999. Development of cysteine protease inhibitors as chemotherapy for parasitic diseases: Insights on safety, target validation, and mechanism of action. Int. J. Parasitol. 29:833937. McKerrow, J.H., Engel, J.C., and Caffrey, C.R. 1999. Cysteine protease inhibitors as chemotherapy for parasitic infections. Bioorg. Med. Chem. 7:639-644. Musil, D., Zucic, D., Turk, D., Engh, R.A., Mayr, I., Huber, R., Popovic, T., Turk, V., Towatari, T., Katunuma, N., and Bode, W. 1991. The refined 2.15 Å X-ray crystal structure of human liver cathepsin B: The structural basis for its specificity. EMBO J. 10:2321-2330. Nagler, D.K., Zhang, R., Tam, W., Sulea, T., Purisima, E.O., and Menard, R. 1999. Human cathepsin X: A cysteine protease with unique car boxypeptidase ac tivity. Biochemistry 38:12648-12654. Nakagawa, T., Roth, W., Wong, P., Nelson, A., Farr, A., Deussing, J., Villadangos, J.A., Ploegh, H., Peters, C., and Rudensky, A.Y. 1998. Cathepsin L: Critical role in Ii degradation and CD4 T cell selection in the thymus. Science 280:450-453. Nakagawa, T.Y., Brissette, W.H., Lira, P.D., Griffiths, R.J., Petrushova, N., Stock, J., McNeish, J.D., Eastman, S.E., Howard, E.D., Clarke, S.R., Rosloniec, E.F., Elliott, E.A., and Rudensky, A.Y. 1999. Impaired invariant chain degradation and antigen presentation and diminished collagen-induced arthritis in cathepsin S null mice. Immunity 10:207-217. Pham, C.T. and Ley, T.J. 1999. Dipeptidyl peptidase I is required for the processing and activation of granzymes A and B in vivo. Proc. Natl. Acad. Sci. U.S.A. 96:8627-8632. Polgar, L. and Halasz, P. 1982. Current problems in mechanistic studies of serine and cysteine proteinases. Biochem. J. 207:1-10. Rao, N.V., Rao, G.V., and Hoidal, J.R. 1997. Human dipeptidyl-peptidase I. Gene characterization, localization, and expression. J. Biol. Chem. 272:10260-10265. Rawlings, N.D. and Barrett, A.J. 1994. Families of cysteine peptidases. Methods Enzymol. 244:46186. Ren, W.P., Fridman, R., Zabrecky, J.R., Morris, L.D., Day, N.A., and Sloane, B.F. 1996. Expression of functional recombinant human procathepsin B in mammalian cells. Biochem. J. 319:793-800. Peptidases
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Rich, D.H., Brown, M.A., and Barrett, A.J. 1986. Purification of cathepsin B by a new form of affinity chromatography. Biochem. J. 235:731734. Riese, R.J., Wolf, P., Brömme, D., Natkin, L.R., Villadangos, J.A., Ploegh, H.L., and Chapman, H.A. 1996. Essential role for cathepsin S in MHC class II-associated invariant chain processing and peptide loading. Immunity 4:357-366. Rozhin, J., Sameni, M., Ziegler, G., and Sloane, B.F. 1994. Pericellular pH affects distribution and secretion of cathepsin B in malignant cells. Cancer Res. 54:6517-6525. Saftig, P., Wehmeyer, O., Hunziker, E., Jones, S., Boyde, A., Rommerskirch, W., and von Figura, K. 1998. Impaired osteoclastic bone resorption leads to osteopetrosis in cathepsin K-deficient mice. Proc. Natl. Acad. Sci. U.S.A. 95:1345313458.
the active site. Proc. Natl. Acad. Sci. U.S.A. 94:14249-14254. Toomes, C., James, J., Wood, A.J., Wu, C.L., McCormick, D., Lench, N., Hewitt, C., Moynihan, L., Roberts, E., Woods, C.G., Markham, A., Wong, M., Widmer, R., Ghaffar, K.A., Pemberton, M., Hussein, I.R., Temtamy, S.A., Davies, R., Read, A.P., Sloan, P., Dixon, M.J., and Thakker, N.S. 1999. Loss-of-function mutations in the cathepsin C gene result in periodontal disease and palmoplantar keratosis. Nature Genet. 23:421-424. van Noorden, C.J.F., Vogels, I.M., Everts, V., and Beertsen, W. 1987. Localization of cathepsin B activity in fibroblasts and chondrocytes by continuous monitoring of the formation of a final fluorescent reaction product using 5-nitrosalicylaldehyde. Histochem. J. 19:483-487.
Santamaria, I., Velasco, G., Pendas, A.M., Fueyo, A., and Lopez-Otin, C. 1998. Cathepsin Z, a novel human cysteine proteinase with a short propeptide domain and a unique chromosomal location. J. Biol. Chem. 273:16816-16823.
Velasco, G., Ferrando, A.A., Puente, X.S., Sanchez, L.M., and Lopez-Otin, C. 1994. Human cathepsin O. Molecular cloning from a breast carcinoma, production of the active enzyme in Escherichia coli, and expression analysis in human tissues. J. Biol. Chem. 269:27136-27142.
Schechter, I. and Berger, A. 1967. On the size of the active site in proteases. I. Papain. Biochem. Biophys. Res. Commun. 27:157-62.
Wang, B., Shi, G.P., Yao, P.M., Li, Z., Chapman, H.A., and Brömme, D. 1998. Human cathepsin F. J. Biol. Chem. 273:32000-32008.
Shi, G.P., Villadangos, J.A., Dranoff, G., Small, C., Gu, L., Haley, K.J., Riese, R., Ploegh, H.L., and Chapman, H.A. 1999. Cathepsin S required for normal MHC class II peptide loading and germinal center development. Immunity 10:197-206.
Weber, E., Gunther, D., Laube, F., Wiederanders, B., and Kirschke, H. 1994. Hybridoma cells producing antibodies to cathepsin L have greatly reduced potential for tumour growth. Cancer Res. Clin. Oncol. 120:564-567.
Sol-Church, K., Frenck, J., Troeber, D., and Mason, R.W. 1999. Cathepsin P, a novel protease in mouse placenta. Biochem. J. 343:207-209.
Wex, T., Levy, B., Smeekens, S.P., Ansorge, S., Desnick, R.J., and Brömme, D. 1998. Genomic structure, chromosomal localization, and expression of human cathepsin W. Biochem. Biophys. Res. Commun. 248:255-261.
Sol-Church, K., Frenck, J., and Mason, R.W. 2000. Cathepsin Q, a novel lysosomal cysteine protease highly expressed in placenta. Biochem. Biophys. Res. Commun. 267:791-795. Takahashi, T., Dehdarani, A.H., and Tang, J. 1988. Porcine spleen cathepsin H hydrolyzes oligopeptides solely by aminopeptidase activity. J. Biol. Chem. 263:10952-10957. Thompson, S.K., Halbert, S.M., Bossard, M.J., Tomaszek, T.A., Levy, M.A., Zhao, B., Smith, W.W., Abdel-Meguid, S.S., Janson, C.A., D’Alessio, K.J., McQueney, M.S., Amegadzie, B.Y., Hanning, C.R., DesJarlais, R.L., Briand, J., Sarkar, S.K., Huddleston, M.J., Ijames, C.F., Carr, S.A., Garnes, K.T., Shu, A., Heys, J.R., Brad beer, J., Zembryski, D., and LeeRykaczewski, L. 1997. Design of potent and selective human cathepsin K inhibitors that span
Willstädter, R. and Bamann, E. 1929. Über die Proteasen der Magenschleimhaut. Hoppe-Seylers Z. Physiol. Chem. 180:127-143. Xia, L., Kilb, J., Wex, H., Lipyansky, A., Breuil, V., Stein, L., Palmer, J.T., Dempster, D.W., and Brömme, D. 1999. Localization of rat cathepsin K in osteoclasts and resorption pits: Inhibition of bone resorption cathepsin K-activity by peptidyl vinyl sulfones. Biol. Chem. 380:679-687.
Contributed by Dieter Brömme Mount Sinai School of Medicine New York, New York
Papain-like Cysteine Proteases
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Overview of Pepsin-like Aspartic Peptidases The aspartic peptidase family of enzymes (also known as aspartic proteinases; A1 in the MEROPS database (Rawlings and Barrett, 2000); http://www.merops.co.uk/) has been identified by the occurrence of the sequence Asp-Thr-Gly (Davies, 1990; Barrett et al., 1998). This sequence is critical—the aspartic acid in this sequence provides one-half of the catalytic machinery of these enzymes; the ThrGly sequence following the Asp is essential for the formation of a unique catalytic conformation that also defines an enzyme of this class. In single-chain enzymes, two separate AspThr-Gly sequences separated by 170 to 190 amino acids are required to form the complete catalytic site. In the homodimeric enzymes of the retroviral aspartic proteinase subfamily (A2 in the MEROPS database, http://www.merops.co.uk/), each monomeric unit has one Asp-Thr-Gly sequence. In either type of aspartic peptidase, the folding of the chains brings the two aspartates together to create the catalytic apparatus. In evolutionary terms, the retroviral enzymes are derived from an ancestor that diverged to give,
UNIT 21.3
through gene duplication, single chain enzymes with two Asp-Thr-Gly sequences. Although the Asp-Thr-Gly sequence is the main feature of this class of enzymes, an aspartic peptidase has other elements that define its unique structure and fold. The single chain enzymes can also be described as two-domain proteins (Orengo et al., 1999), with mainly beta structure [CATH lexicon: 2 170 10 10], as detailed in Figure 21.3.1. Both occurrences of the Asp-Thr-Gly sequence are always preceded by two hydrophobic amino acids. The fungal members of the family (e.g., rhizopepsin and aspergillopepsin) diverge slightly in that they have an aspartate residue two positions before the first catalytic Asp (Barrett et al., 1998). In porcine pepsin, 43 residues after the first Asp, a conserved tyrosine residue appears in the pepsin-like family A1. In the family A2 of retroviral peptidases, this Tyr is missing. The tyrosine residue plays an important role in defining the active site pockets that bind substrate side-chain residues, separating the S1 binding pocket from the S2′ pocket.
Figure 21.3.1 Schematic diagram of a typical aspartic proteinase. The backbone is represented as a shaded ribbon, with the N-terminal domain on the left and the C-terminal domain on the right. Note the high degree of beta structure (dark gray ribbons) and relatively low occurrence of alpha helices (black spirals). The catalytic aspartic acids are represented as space-filling spheres in the middle of the diagram. Peptidases Contributed by Ben M. Dunn Current Protocols in Protein Science (2001) 21.3.1-21.3.6 Copyright © 2001 by John Wiley & Sons, Inc.
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In addition, a conserved glycine residue in the sequence Leu-Gly-Ile occurs 47 residues after the tyrosine. The glycine residue is conserved for structural reasons; it passes through the wide loop containing the Asp-Thr-Gly sequence to form the “Psi-loop.” The Leu and Ile residues fill up space above and below the Gly so that the Psi-loop is “locked” in place. The Psi-loop motif is repeated in the second half of the protein. In porcine pepsin, 87 residues following the second Asp-Thr-Gly is LeuGly-Asp. Again, the Gly fits nicely in the strand running through the wide loop, and the Leu and Asp side chains provide the steric bulk to lock the strand in place. These features imply that the folding of the protein must occur in a way that allows the strand to be near the Asp-ThrGly loop before the final folding. The strand could not insert through the loop due to the size of the residues before and after the glycine residue. Figures 21.3.2 and 21.3.3 detail these features.
SEQUENCE HOMOLOGIES AND EVOLUTIONARY RELATEDNESS The MEROPS database currently lists 200 members of family A1 of clan AA. The clan designation AA refers to the use of water as a nucleophile, with the water molecule bound by two catalytic Asp residues. All members of clan AA are from eukaryotic organisms or viruses or virus-like elements. Family A1 refers to the members that have a statistically significant sequence relationship to porcine pepsin, the first member identified. The reader is referred to the MEROPS database for additional information (http://www.merops.co.uk/).
CATALYTIC MECHANISM It is now generally accepted that catalysis by members of the pepsin family involves general acid–general base catalysis of the attack of a water molecule upon the peptide bond of a bound substrate. The aspartate residue in the second domain (Asp 215 in pepsin numbering) serves as the general base to abstract a proton from a water molecule that can be seen in the crystal structures of aspartic proteinases without bound inhibitors. The water molecule si-
Overview of Pepsin-like Aspartic Peptidases
multaneously attacks the carbonyl carbon while the first aspartate residue (Asp 32) provides electrophilic assistance to the carbonyl oxygen. The resulting tetrahedral intermediate (Fraser et al., 1992; Veerapandian et al., 1992) can break down if the nitrogen of the peptide bond acquires a proton from solvent, or through inversion of the nitrogen so that Asp 215 can give back the proton it took from the attacking water molecule. In either case, the amine group would then be a suitable leaving group, resulting in the formation of the amine and carboxyl products of the reaction. The final step, which could be rate limiting in some cases, would be the departure of the products from the active site (Fig. 21.3.4).
SUBCELLULAR AND TISSUE LOCALIZATION OF ASPARTIC PROTEINASES The biological niche occupied by aspartic peptidases is extremely broad, although their presence is generally restricted to acidic environments within cells and tissues. The parent member, porcine pepsin, and other gastric enzymes are synthesized in the chief and mucous neck cells of the fundic mucosa. The enzyme is synthesized as a pro-proenzyme with the signal sequence being removed in the usual fashion in the ER. The proenzyme is packaged into granules and stored inside the cells. Under the control of several signals, the granules fuse with the cell surface membrane and release their contents into the stomach. The drop in pH experienced is dramatic, leading to protonation of groups within the active site and self-processing to the mature enzyme. In contrast, cathepsin D and E are intracellular enzymes (Saku et al., 1991). The former is also made as a proenzyme and targeted to the lysosomes of cells by the mannose-6-phosphate pathway. There, it is activated to the mature form by the action of unknown cysteine proteinases within the organelle. The vacuoles of many yeast species also contain enzymes of this class. Cathepsin E, on the other hand, while also remaining inside the cell, does not reside in the lysosomes, but rather exists within the vesicular network of the cells.
~30-35AA~Hb-Hb-D-T-G-S-S~45-50-AA~Y-X-S/T~45-50AA~ L-G-I~90-95AA~Hb-Hb-D-T/S-G-T-S~85-90AA~L-G-D~ 20-30AA Figure 21.3.2 Sequence template for an aspartic proteinase. Single letter codes used for amino acids; Hb = hydrophobic amino acid.
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A
B
C
Figure 21.3.3 (A) Psi loop structure, showing how the Leu-Gly-Ile sequence fits through the wide loop of the catalytic Asp residue, with Leu below the wide loop, and Ile above it. (B) Psi loops of porcine pepsin. (C) Hydrogen bonding (dashed lines) within and between the two Asp-Thr-Gly active site motifs of porcine pepsin. The Thr of one motif hydrogen bonds to the Asp of the other motif and vice versa.
Peptidases
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1
2
2.575 A 2.580 A
D215
D32
Figure 21.3.4 Structure of hypothetical tetrahedral intermediate in catalysis by aspartic proteinases. Oxygen atom 1 is derived from the water molecule that attacks the carbonyl group of the substrate, and atom 2 is the original carbonyl oxygen. Asp 215 (D215) assists the attack by taking a proton from the water molecule, and ASP 32 (D32) adds a proton to the carbonyl oxygen.
Overview of Pepsin-like Aspartic Peptidases
The malarial parasite, Plasmodium falciparum, utilizes hemoglobin degradation as a food source during the erythrocytic phase of its life cycle (Francis et al., 1997). At least two enzymes, plasmepsins I and II, are known to be found in the food vacuole of the parasite and may be involved in the process of degradation of host cell hemoglobin. Many members of the fungi family are also secreted enzymes, playing a role in supplying nutrients through degradation of proteins encountered in the environment around the cells. Aspartic proteinases of plants diverge slightly from the sequence template given above in that they include a 105 amino acid insert (“plant-specific insert”) that most likely plays a targeting role within the plant cell (Kervinen et al., 1999). These enzymes can be either freely soluble or membrane bound. The newly discovered “mamepsins” of the human, which have been implicated in the pathology of Alzheimer’s Disease (Sinha et al., 1999), contain C-terminal extensions that can provide type-I membrane anchorage. As the genome sequencing efforts continue, more examples of membrane-bound aspartic peptidases will almost certainly be discovered.
BIOLOGICAL AND INTRINSIC SUBSTRATE SPECIFICITY Aspartic proteinases exhibit a range of substrate specificity, from the extremely broad to exquisitely specific. Enzymes that are charged with a general role in degradation of nutrient proteins, such as the gastric pepsins and the secreted fungal enzymes, tend to have powerful, broad specificity. The acidic milieu aids in the unfolding of substrate proteins, so that the enzymes can readily attack the exposed peptide bonds. On the other extreme, human renin is very selective, cleaving only one bond within the sequence of human angiotensinogen (Green et al., 1990). In addition, animal renins are unable to cleave human angiotensinogen. It may be that the structure of renin has evolved to recognize the cleavage sequence within a partly unfolded substrate protein. While the overall structure of renin is very similar to that of other members of the family, the active site is more restricted, leading to the suggestion that there is less adaptability in the fit between substrate and enzyme.
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ASPARTIC PEPTIDASES AND DISEASE Aspartic proteinases may play many roles in human disease, although in many cases the speculations outnumber the proofs. For many years, gastric pepsin was suspected of contributing to the formation of stomach ulcers. In cases of cancer of the esophagus, pepsinogen secretion is stimulated and the excessive amounts of pepsin that result could play a role in extensive degradation of the esophagus and upper stomach (Varis et al., 1991). Human cathepsin D has long been suspected of involvement in breast cancer, again due to its abundant secretion in necrotic tissues (Rochefort et al., 2000). However, it is unclear whether this is a cause or a consequence of the lesions. Furthermore, the activity in an extracellular environment that is not acidic could be limiting. The enzyme is over-expressed in some tumor cell lines. Renin plays a role in regulation of blood pressure in mammals, and has long been the target of drug discovery for control of hypertension. In fact, the rapid advances in the discovery of effective drugs for HIV infection, with HIV protease as the target, were largely based on prior experience in studies of renin. The mamepsins, recently reported membrane-bound enzymes, have been implicated in the aberrant processing of beta amyloid protein to yield the fragments that precipitate and contribute to plaque formation that leads to Alzheimer’s Disease. Microorganisms that cause human disease are numerous and many employ aspartic proteinases to gain entry into cells and tissues. Fungi such as Candida albicans produce secreted enzymes of this class that have been shown to be virulence factors for disease. The chestnut blight fungus, Cryphonectria parasitica, produces endothiapepsin, whose structure was one of the first members of the family to be solved by X-ray crystallography (Blundell et al., 1990). The fungus has a devastating effect upon the American chestnut tree. As mentioned earlier, aspartic peptidases in the malaria parasite, Plasmodium falciparum, may be involved in the supply of nutrients derived from hemoglobin degradation. Cysteine proteinases are also involved in this process, and the recent discovery of several additional genes has complicated the scenario.
RECOMBINANT EXPRESSION, PURIFICATION, AND ASSAYS OF ASPARTIC PEPTIDASES In addition to purification of naturally occurring enzymes from extracellular secretions or tissues, aspartic proteinases can be produced by recombinant means. Expression in bacteria, plant cells, yeast, and mammalian systems has been reported for various members of the family. In general, expression of the proenzyme form is necessary in order to permit proper folding of the polypeptide chain. Several early attempts to produce the mature form of the enzyme, lacking the 40- to 50-residue prosegment, were unsuccessful, whereas the proenzymes can be produced in inclusion bodies in bacterial expression systems and refolded. The yields are highly variable, depending on the sequence, and there is a need for improvements in the expression of many enzymes of this family. Assays of enzymes in this class vary from the classical Anson hemoglobin digestion assay, where the absorbance of solubilized peptide fragments is quantified, to high throughput fluorescence assays utilizing synthetic peptides. General and specific assays for peptidases will be described in other units in this chapter.
Literature Cited Barrett, A.J., Rawlings, N.D., and Woessner, J.F. 1998. The Handbook of Proteolytic Enzymes, pp. 799-986. Academic Press, San Diego. Blundell, T.L., Jenkins, J.A., Sewell, B.T., Pearl, L.H., Cooper, J.B., Tickle, I.J., Veerapandian, B., and Wood, S.P. 1990. X-ray analysis of aspartic proteinases: The 3-dimensional structure at 2.1 A resolution of endothiapepsin. J. Mol. Biol. 211:919-941. Davies, D.R. 1990. The structure and function of the aspartic proteinases. Annu. Rev. Biophys. Chem. 19:189-215. Francis, S.E., Sullivan, D.J., and Goldberg, D.E. 1997. Hemoglobin metabolism in the malaria parasite Plasmodium falciparum. Annu. Rev. Microbiol. 51:97-123. Fraser, M.E., Strynadka, N.C.J., Bartlett, P.A., Hanson, J.E., and James, M.N.G. 1992. Crystallographic analysis of transition-state minics bound to penicillopepsin-phosphorus-containing peptide analogs. Biochemistry 31:52015214. Green, D.W., Aykent, S., Gierse, J.K., and Zupec, M.E. 1990. Substrate specificity of recombinant human renal renin: Effect of histidine in the P2 su bsite on pH dependence. Biochemistry 29:3126-3133. Peptidases
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Kervinen, J., Tobin, G.J., Costa, J., Waugh, D.S., Wlodawer, A., and Zdanov, A. 1999. Crystal structure of plant aspartic proteinase prophytepsin: Inactivation and vacuolar targeting. EMBO J. 18:3947-3955.
Varis, K., Kekki, M., Harkonen, M., Sipponen, P., and Samloff, I.M. 1991. Serum pepsinogen-1 and serum gastrin in the screening of atrophic pangastritis with high risk of gastric cancer. Scand. J. Gastroenterol. 26:117-113.
Orengo, C.A., Michie, A.D., Jones, S., Jones, D.T., Swindells, M.B., and Thornton, J.M. 1999. CATH: A hierarchic classification of protein domain structures. Structure 5:1093-1108.
Veerapandian, B., Cooper, J.B., Sali, A., Blundell, T.L., Rosati, R.L., Dominy, B.W., Damon, D.B., and Hoover, D.J. 1992. Direct observation by x-ray analysis of the tetrahedral intermediate of aspartic proteinases. Protein Sci. 1:322-328.
Rawlings, N.D. and Barrett, A.J. 2000. MEROPS: The peptidase database. Nucl. Acids Res. 28:323325. Rochefort, H., Garcia, M., Glondu, M., Laurent, V., Liaudet, E., Rey, J.M., and Roger, P. 2000. Cathepsin D in breast cancer: Mechanisms and clinical applications, a 1999 overview. Clin. Chim. Acta 291:157-170. Saku, T., Sakai, H., Shibata, Y., Kato, Y., and Yamamoto, K. 1991. An immunocytochemical study on distinct intracellular localization of cathepsin D and cathepsin D in human gastric cells and various rat cells. J. Biochem. 110:956964. Sinha, S., Anderson, J.P., Barbour, R., Basi, G.S., Caccavello, R., Davis, D., Doan, M., Dovey, H.F., Frigon, N., Hong, J., Jacobson-Croak, K., Jewett, N., Keim, P., Knops, J., Lieberburg, I., Power, M., Tan, H., Tatsuno, G., Tung, J., Schenk, D., Seubert, P., Suomensaari, S.M., Wang, S.W., Walker, D., Zhao, J., McConlogue, L., and John, V. 1999. Purification and cloning of amyloid precursor protein beta-secretase from human brain. Nature 402:537-540.
Key Reference Barrett et al., 1998. See above. The definitive compilation of facts for the peptidase world. Authoritative but concise reviews by experts on each enzyme, all presented in a common format, make this a must-have for any laboratory involved in proteolytic enzymes.
Internet Resource http://www.merops.co.uk/ MEROPS website.
Contributed by Ben M. Dunn University of Florida College of Medicine Gainesville, Florida
Overview of Pepsin-like Aspartic Peptidases
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Metalloproteases Metalloproteases (metallopeptidases) are composed of a diverse group of endopeptidases and exopeptidases. They participate in many biological processes such as embryonic development, morphogenesis, processing of peptide hormones, release of cytokines and growth factors, cell-cell fusion, cell adhesion and migration, intestinal absorption of nutrients, viral polyprotein processing, bacterial cell wall biosynthesis, and metabolism of antibiotics (examples are listed in Table 21.4.1). Because metalloproteases play key roles in many normal biological processes, their aberrant activities have been implicated in diseases such as arthritis, cancer, cardiovascular diseases, nephritis, disorders in the central nervous system, fibrosis, and infection, as well as others. Neurotoxins of tetanus and botulism, and anthrax toxin lethal factor are also metalloproteases (Table 21.4.1). Metallopeptidases contain a divalent metal ion at the active site. The catalytic metal ion is usually coordinated by three amino acid sidechain ligands. A water molecule that is essential for hydrolysis of the peptide bond also coordinates with the metal ion as a fourth ligand in the active form of metallopeptidase. In most cases the metal ion is zinc, but in some cases it is cobalt, manganese, or nickel. The known metal ligands in metallopeptidases are His, Glu, Asp, or Lys. The occurrence of these ligands in the catalytic sites is His>>Glu>Asp>Lys (Hooper, 1996). An unpaired Cys is found in the propeptides of matrix metalloproteinases (MMPs). This Cys coordinates the zinc ion of the active site as a fourth ligand, which keeps the zymogens of MMPs inactive. Upon activation the propeptide is removed and a water molecule replaces the cysteine to form an active MMP (Woessner and Nagase, 2000). Binding of the substrate to the catalytic site displaces the water molecule from the zinc ion, and in its place the carbonyl group of the substrate interacts with the zinc. Unlike serine and cysteine proteases, metalloproteases do not form covalent intermediates during peptide bond hydrolysis, a characteristic shared with aspartic proteases (UNIT 21.3), and the hydrolysis of a peptide bond by the metallopeptidase is mediated by the nucleophilic attack of a water molecule on the carbonyl of the scissile bond.
UNIT 21.4 Most metallopeptidases require only one metal ion for catalysis, but some enzymes possess two metal ions that act cocatalytically. All cobalt and manganese metalloproteases possess two metal ions in the active site. A diagnostic feature of metallopeptidases is that chelating agents such as 1,10-phenanthroline or EDTA inhibit enzymic activity. Dithiothreitol and cysteine are often inhibitory as they interact with the catalytic metal ion.
CLASSIFICATION The cu rr ent MEROPS database (http://www.merops.co.uk/merops/merops.htm) classifies metallopeptidases into forty-six families (see Table 21.1.3). The families are grouped into ten clans based on metal ion binding motifs and similarities in 3-D structure (Table 21.4.2). Only representative examples of metallopeptidases are discussed here. For more detailed properties of each enzyme, please consult the Handbook of Proteolytic Enzymes (Barrett et al., 1998).
Most Metalloproteases Including Glu-Zincins and Metzincins Have the HEXXH Motif The majority of metallopeptidases have the catalytic metal ion binding motif, HEXXH, where X is any amino acid. The enzymes whose third metal ligand is His, Glu, or Asp are grouped as Clan MA (Rawlings and Barrett, 2000) and are all zinc metalloproteases. Clan MA is the largest clan of the metalloproteinases, consisting of 25 families. Some of these families are composed of members with the zinc binding motif HEXXH(Xn)E, where n is at least 14 residues, and the underlined residues are zinc ligands. These proteases are also called “glu-zincins,” as they possess a glutamic acid as a third zinc ligand (Hooper, 1994). Typical enzymes of the HEXXH(Xn)E zinc binding motif are: thermolysins and the related bacterial metallopeptidases (family M4), animal peptidyl-dipeptidase A (angiotensin I-converting enzyme; family M2), and endothelinconverting enzyme (family M13). Others include aminopeptidase B, lysyl aminopeptidase, membrane analanyl aminopeptidase (myeloid leukemia marker CD13; family M1), mycolysin (family M5), Clostridium collagenase (family M9), hyicolysin (family M30), fungaPeptidases
Contributed by Hideaki Nagase Current Protocols in Protein Science (2001) 21.4.1-21.4.13 Copyright © 2001 by John Wiley & Sons, Inc.
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Biological Functions of Metallopeptidases
Biological processes Antibiotic resistance β-Lactamase activity Vancomycin resistance
Enzyme [family] Membrane dipeptidase [M19] VanX D-Ala-D-Ala-dipeptidase (vancomycin-resistant Enterococcus faecolis host) [M45]
Bacteriolytic activity Resolution of peptidoglycan N-acetylmuramoyl-L-alanine amidase (not peptidase) [M15] Resolution of peptidoglycan β-lytic metalloendopeptidase (Lysobacter enzymogenes) [M23], staphylolysin [M23], (Staphylococcus aureas) lysostaphin [M37] Biosynthesis Biosynthesis of bacterial Zinc D-Ala-D-Ala carboxypeptidase [M15] cell walls γ-D-glutamyl-meso-(L)-diaminopimelate Sporulation (Bacillus) peptidase [M14] Development Cell-cell fusion ADAMs [M12] Cell migration MMPs [M12] Morphogenesis ADAMs [M12], bone morphogenetic protein 1 (BMP-1) [M12], MMPs [M10], tolkin (Drosophila) [M12], tolloid [M12] Release of cytokines and ADAMs [M12], MMPs [M10] growth factors Effects on immune systems Disruption of insect Immune inhibitor A [M6] immune system IgA, IgG degradation IgA specific metalloendopeptidase [M26], mirabilysin [M10] Infection Diarrhea Fragilysin (Bacteriodes fragilis) [M10] Opportunistic bacterial Coccolysin (Enterococcus faecalis) [M4], infection pseudolysin (Pseudomonas aeruginosa) [M4], aeruginolysin (Pseudomonae aeruginosa) [M10] Protozoan infection Leishmanolysin (Leishmania) [M8] Viral polyprotein processing Hepatitis C virus endopeptidase 2 [M44], vaccinia virus proteinase [M44] Virulence factor activation Hemagglutinin/protease 9 (Vibrio cholerae) of toxins of Vibrio cholerae [M4] Nutrition Intestinal absorption of folic Glutamate carboxypeptidase II [M28] acid Peptide and protein processing ADAM 17 (TACE) [M12] Alzheimer’s precursor protein processing (α-secretase activity) Cell signaling/release of a S2P protease [M50], SpoIVFB (Bacillus transcription factor subtilis) [M51], ADAM-10 [M12] continued Metalloproteases
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Table 21.4.1
Biological Functions of Metallopeptidases, continued
Biological processes
Enzyme [family]
Inactivation of leukotriene D4 to leukotriene E4 Mitochondrial/chloroplast protein processing and trafficking
Membrane dipeptidase [M19]
Chloroplast stromal processing peptide [M16], mitochondrial processing peptidase [M16], mitochondrial intermediate peptidase [M3] Peptide hormone and Carboxypeptide E (removal of C-terminal neurotransmitter processing basic residues) [M14], metallocarboxypeptide D [M14] Processing of peptide Endothelin-converting enzyme [M13], hormone peptidyl dipeptidase A (angiotensin I-converting enzyme) [M2] Processing of precursor Chloroplast stromal processing peptidase proteins [M16], mitochondrial processing peptidase [M16] Procollagen processing and Procollagen N-endopeptidase fibril formation (ADAM-TS2) [M12] Removal of N-terminal Methionyl aminopeptidase I and II [M24] methionine Shedding of cell surface ADAM 10, ADAM 17 (TACE) [M12] molecule Yeast mating α-factor Ste 24 protease [M48] maturation Reproduction Blastocyst implantation MMPs [M10] Embryo hatching Choriolysins (teleost fish, the medaka) [M12], envelysin (sea urchin MMP) [M10] Fertilization ADAM1 [M12] Ovulation MMPs [M10] Gametolysin (Chlamydomonas reinhardtii) Removal of cell walls [M11] during green algae gamete mating Tissue remodeling Bone remodeling ADAMs [M12], MMPs [M10] Extracellular matrix MMPs [M10] turnover/breakdown Wound/fracture healing MMPs [M10] Toxins Anthrax Anthrax toxin lethal factor (Bacillus anthracis) [M34] Cytoxicity and hemolytic Legionella metalloendopeptidase [M4] Hemorrhagic toxin Snake venom metalloproteinase reprolysins (e.g., atrolysin A, ruberlysin, mutalysins, janarhagin) [M12] Tetanus and botulism Tentoxilysin (Clostridium tetani), bontoxilysin (Clostridium botulinum) [M27]
Peptidases
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Table 21.4.2
Family
Clans and Families of Metallopeptidasesa
Subfamily
Example (organism)
Clan MA: Water nucleophile; water bound by single zinc ion ligated to two His (within the motif HEXXH) and Glu, His or Asp M1 Membrane alanyl aminopeptidase (Homo sapiens) M2 Peptidyl-dipeptidase A (angiotensin I converting enzyme; Homo sapiens) M4 Thermolysin (Bacillus stearothermophilus) M5 Mycolysin (Streptomyces cacoi) M6 Immune inhibitor A (Bacillus thuringiensis) M7 Snapalysin (Streptomyces lividans) M8 Leishmanolysin (Leishmania major) Vibrio collagenase (Vibrio alginolyticus) M9 A Clostridium collagenase (Clostridium B perfringens) M10 A Collagenase 1 (Homo sapiens) B Serralysin (Serratia marcescens) C Fragilysin (Bacteroides fragilis) M11 Gametolysin (Chlamydomonas reinhardii) M12 A Astacin (Astacus fluviatilis) B Adamalysin (Crotalus adamanteus) C ADAM 17 (Homo sapiens) M13 Neprilysin (Homo sapiens) M30 Hyicolysin (Straphylococcus hyicus) M36 Fungalysin (Aspergillus fumigatus) M48 Ste24 endopeptidase (Saccharomyces cerevisiae) Clan MX: Other metallopeptidases with HEXXH motif M3 A Thimet oligopeptidase (Rattus norvegicus) B Oligopeptidase F (Lactococcus lactis) M26 IgA1-specific metalloendopeptidase (Streptococcus sanguis) M27 Tentoxyilysin (Clostridium tetani) M32 Carboxypeptidase Taq (Thermus aquaticus) M34 Anthrax lethal factor (Bacillus anthracis) M35 Penicillolysin (Penicillum citrinum) M41 FtsH endopeptidase (Escherichia coli) M43 Cytophagolysin (Cytophaga sp.) Clan MC: Water nucleophile; water bound by single zinc ion ligated to His, Glu (within the motif HXXE) and His M14 A Carboxypeptidase A (Homo sapiens) B Carboxypeptidase E (Bos taurus) γ-D-glutamyl-(L)-meso-diaminopimelate C peptidase I (Bacillus sphaericus) Clan MD: Water nucleophile; water bound by single zinc ion ligated to His, Asp and His M15 A Zinc D-Ala-D-Ala carboxypeptidase (Streptomyces albus) B Van Y D-Ala-D-Ala carboxypeptidase (Enterococcus faecium) C Endolysin (bacteriophage A118)
Metalloproteases
continued
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Table 21.4.2
Family
Clans and Families of Metallopeptidasesa, continued
Subfamily
Example (organism)
M45
Van X D-Ala-D-Ala dipeptidase (Enterococcus faecium) Clan ME: Water nucleophile; water bound by single zinc ion ligated to two His (within the motif HXXEH) and Glu M16 A Pitrilysin (Escherichia coli) B Mitochondrial processing peptidase (Saccharomyces cerevisiae) M44 Metalloendopeptidase (Vaccinia virus) Clan MF: Water nucleophile; water bound by two zinc ions ligated by Lys, Asp, Asp, Asp, Glu M17 Leucyl aminopeptidase (Bos taurus) Clan MG: Water nucleophile; water bound by two cobalt or manganese ions ligated by Asp, Asp, His, Glu, Glu M24 A Methionine aminopeptidase I (Escherichia coli) B X-proaminopeptidase (Escherichia coli) Clan MH: Water nucleophile; water bound by two zinc ions ligated by His (or Asp), Asp, Glu, Asp (or Glu), His M18 Aminopeptidase I (Saccharomyces cerevisiae) M20 A Glutamate carboxypeptidase (Pseudomonas sp.) B Gly-X carboxypeptidase (Saccharomyces cerevisiae) M25 X-His dipeptidase (Escherichia coli) M28 A Leucyl aminopeptidase (Vibrio proteolyticus) B Aminopeptidase (Streptomyces griseus) C IAP aminopeptidase (Escherichia coli) M40 Sulfolobus carboxypeptidase (Sulfolobus solfataricus) M42 Glutamyl aminopeptidase (Lactococcus lactis) Clan MJ: Water nucleophile; water bound by two nickel ions ligated by His, His, Lys, His, His, Asp or a single zinc ion ligated by two His (within the motif HXH) and a third, as yet unidentified, residue β-aspartyl dipeptidase (Escherichia coli) M38 Clan MK: Water nucleophile; water bound by single zinc ion probably ligated to two His (within the motif HXEXH) and an as yet unidentified third ligand; probable actin-like fold M22 O-sialoglycoprotein endopeptidase (Pasteurella haemolytica) Clan ML: Water nucleophile; water bound by single nickel ion ligated by Glu, Asp, His M52 HybD endopeptidase (Escherichia coli) Clan MM: Water nucleophile; water bound by single zinc ion ligated to two His (within the motif HEXXH) and Asp; membrane proteins with intramembrane active site M50 S2P protease (Homo sapiens) M51 Sporulation factor SpoIVFB (Bacillus subtilis) continued
Peptidases
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Table 21.4.2
Family
Clans and Families of Metallopeptidasesa, continued
Subfamily
Example (organism)
Metallopeptidase families not yet assigned to clan M19 Membrane dipeptidase (Homo sapiens) β-lytic metalloendopeptidase (Achromobacter M23 lyticus) M29 Aminopeptidase T (Thermus aquaticus) M37 Lysostaphin (Staphylococcus simulans) M44 Dipeptidyl-peptidase III (metallo-form) (Rattus norvegicus) aAdopted from Rawlings and Barrett (2000).
Metalloproteases
lysin (family M36), and Ste 24p protease (family M48). Metalloproteases with the HEXXHXXGXXH motif are called “metzincins.” This term was coined by Bode et al. (1993) for the matrix metalloproteinases (MMPs) also called “matrixins” (subfamily A of family M10), the bacterial serralysins (subfamily B of family M10), astacins (subfamily A of family M12), and reprolysins (subfamily B of family M12), based on the crystal structures of representative members of those subfamilies. The zinc binding ligands of metzin cin s ar e th ree histidines of the HEXXHXXGXXH motif. In addition, they all possess a conserved Met-containing 1,4-turn in a similar conformation, called a “Met-turn,” which forms a hydrophobic base for the catalytic zinc. Pregnancy-associated plasma protein A, which has been characterized as a protease that cleaves insulin-like growth factor binding protein 4, is considered to be a new family belonging to metzincins (Lawrence et al., 1999). Because subfamilies of the matrixins (MMPs), the astacins, and the reprolysins consist of a large number of related metalloproteases, these subfamilies are commonly referred to as the matrixin (or MMP) family, the astacin family, and the reprolysin family, respectively. The matrixins have been characterized as key proteases that degrade extracellular matrix components such as collagens, proteoglycans, and glycoproteins, and they consist of more than thirty members of plant, insect, nematode, and vertebrate origin (Woessner and Nagase, 2000). The astacins include astacin, meprin, and procollagen C-endopeptidase. The reprolysins include snake venom metalloproteinases (e.g., adamalysins, atrolysins) and ADAMs
(i.e., a disintegrin and metalloprotease domain). ADAMs are type 1 transmembrane proteins, but only about half of the members have proteolytic activity, while the rest have an incomplete zinc binding motif and/or lack the glutamic acid essential for catalysis (Black and White, 1998). The members, often referred to as the ADAM family in the literature, include metalloproteases that shed cell surface molecules (e.g., ADAM-9, ADAM-10, tumor necrosis factor converting enzyme/ADAM-17) and soluble forms of the enzymes, the ADAMTS metalloproteases (i.e., ADAM with thrombospondin domain). ADAM-TSs lack the transmembrane and a cytoplasmic domain, but instead have one or more thrombospondin type I motifs. These include procollagen N-endopeptidase (ADAM-TS3), aggrecanase 1 (ADAMTS4), and aggrecanase 2 (ADAM-TS5). Not all ADAM metalloproteinases have been characterized for their proteolytic activities. Oth er enzymes that contain the HEXXHXXGXXH zinc binding motif are leishmanolysin (family M8), a membrane bound metalloprotease expressed at the surface of the protozoan parasite Leishmania, and gametolysin, which digests cell walls during green algae gamete mating (family M11). Clan MA also contains the enzymes with the zinc binding motif of HEXXHXXGXXD. This includes immune inhibitor A (Bacillus thuringiensis; family M6) and synapalysin (Streptomyces neutral proteinase A; family M7). Immune inhibitor A degrades antibacterial proteins, cecropins, and attacins from several species of insect, inhibiting the ability of the insect immune system to kill invading bacteria. There are ten metallopeptidase families with the HEXXH motif that are not classified as Clan MA, because of structural dissimilarities. They
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include thimetoligopeptidase (family M3), Ig A-specific metalloproteinase (Streptococcus sanguis; family M26), tentoxilysin (Clostridium tetani; family M27), carboxypeptidase Taq (Thermus aquaticus; family M32), anthrax lethal factor (Bacillus anthracis; family M34), FtsH endopeptidase (family M41), and others (see Table 21.1.2). FtsH protease is found in bacteria, mitochondria, and chloroplasts. Bacterial FtsH is an ATP-dependent metalloprotease that degrades the σ32 factor during cell division. S2P protease (family 50) and sporulation factor SpoIVFB (family 51), which also have the HEXXH motif, are assigned to Clan MM (see below).
Clans MC and MD Include Carboxypeptidases The catalytic zinc of enzymes in Clan MC is tetrahedrally coordinated by two histidines, one glutamic acid, and a water molecule. One of the histidines and the glutamic acid are in the HXXE motif and the third ligand (His) is located 103 to 143 residues C-terminal to this motif. Clan MC is composed of only family M14, which includes three subfamilies: carboxypeptidases A, B, T, and U, and mast cell carboxypeptidase A in subfamily A; carboxypeptidases D, H, and M, metallocarboxypeptidase and other unassigned carboxypeptidases in subfamily B; and γ-D-glutamyl-meso-(L)-diaminopimelate peptidase I from Bacillus sphaevicus in subfamily C. Metallocarboxypeptidase D has three carboxypeptidase-like units, followed by a transmembrane domain and a cytosolic tail. Only the first two carboxypeptidase units are active. Carboxypeptidase M is membrane bound via glycosyl-phosphatidylinositol (GPI); however, most members of family M14 are not membrane bound. γ-D-glutamyl-meso-diaminopimelate peptidase I is involved in turnover and lysis of bacterial cell wall, which removes the C-terminal and meso-diaminopimelic acid (msA2pm) or the C-terminal dipeptide ms-A2pmD-Ala by its unique ability to cleave the γ-D-glutamyl linkage of L-Ala-γ-D-Glu∼(L)msA2pm or L-Ala-γ-D-Glu∼(L)msrA2pm(L)-D-Ala, respectively. Clan MD is composed of only one family, M15. The family contains zinc D-Ala-D-Ala carboxypeptidase from Streptomyces, which is involved in the biosynthesis of bacterial cell wall peptidoglycans. The 3-D structure of the enzyme indicates that His154, Asp161, and His197 are the zinc ligands. The enzyme consists of an N-terminal domain homologous to
a C-terminal domain of N-acetyl muramoylAla amidase (E.C. 3.5.1.28) from Bacillus and a C-terminal domain with the zinc ligands and the active site. The polypeptide fold of zinc D-Ala-D-Ala carboxypeptidase is similar to that of the N-terminal domain of the sonic hedgehog protein from mouse, expressed in the notochord, and responsible for the local and longrange induction of ventral cell types within the neural tube and somites. The N-terminal domain of the sonic hedgehog protein has a zinc ion coordination site (His, Asp, and His), and it is postulated that this protein may have a peptidase activity.
Metalloproteases with the HXXEH Motif: Clan ME Several metalloproteases have the zinc ligand motif HXXEH, in which the glutamic acid is thought to participate in catalysis. This is in contrast to the HEXXH of Clan MA. They are grouped as clan ME, and families M16 and M44 belong to this clan. Typical members of M16 are pitrilysin (bacterial), insulysin, nardilysin, and mitochondrial processing peptidase. The third zinc ligand of pitrilysin is Glu169 located 77 residues C-terminal from the HXXEH motif. Family M44 includes vaccinia virus polyprotein processing endopeptidase and hepatitis C virus endopeptidase 2.
Metallopeptidases with Two Metal Ions for Catalysis: Clans MF, MG, and MH Some metallopeptidases have two metal ions at their active site. They are found in Clans MF, MG, and MH. All known metallopeptidases with two catalytic metal ions are exopeptidases. Leucyl aminopeptidases from eukaryotes and prokaryotes belong to family M17, and this is the only family in Clan MF; the enzymes consist of the N-terminal and the C-terminal domains. The 3-D structure of cattle leucyl aminopeptidase indicates it is a homohexamer and the active sites are located in the interior of the complex. Each enzyme subunit has two zinc ions in the C-terminal domain, and zinc ions are pentahedrally coordinated with Asp255, Asp 332, and Glu 334 for one zinc, and Asp255, Lys250, Asp273, and Glu334 for the other (Sträter and Lipscomb, 1998). Clan MG also consists of one family (family M24). Representative members are methionyl aminopeptidases and X-Pro aminopeptidases. Methionine aminopeptidases are involved in the processing of the initiator methionine from cytoplasmic proteins. Methionyl aminopepti-
Peptidases
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dase I (type I enzyme) is found in prokaryotes and eukaryotes, including humans, but not in archaebacteria, whereas methionyl aminopeptidase II (type II enzyme) is found in archaebacteria and eukaryotes (Bradshaw et al., 1998). E. coli type I enzyme has a two-domain structure, each possessing a similar fold. This is considered to have arisen from gene duplication and it is speculated that the ancestral enzyme was active as a homodimer. Two cobalt ions are coordinated by two aspartic acids, two glutamic acids, and one histidine. Bacterial X-Pro aminopeptidase is a manganese-dependent enzyme and eukaryotic X-Pro aminopeptidase is a zinc-dependent enzyme. The eukaryotic enzymes are homodimers, and the bacterial enzyme forms a homodimer or a homotetramer depending on the concentration. Conserved metal-binding ligands characteristic of methionyl aminopeptidases are present. Six families comprise Clan MH (M18, M20, M25, M28, M40, and M42). They contain aminopeptidases and carboxypeptidases. Examples are: aminopeptidase I (family M18); glutamate carboxypeptidase (family M20); bacterial X-His dipeptidases (family M25); mammalian membrane-bound glutamate carboxypeptidase II (family M28); Streptomyces aminopeptidase (family M28); Vibrio leucyl proteolyticus aminopeptidase (family M28); carboxypeptidase from the thermoacidophilic archaean Sulfolobus solfataricus (family M40); and Bacillus aminopeptidase I (family M42). Aminopeptidase I, Pseudomonas glutamate carboxypeptidase, Streptomyces aminopeptidase, and Vibrio aminopeptidase have two zinc ions at the active site. The zinc ligands in Vibrio aminopeptidase of family M28 are Asp117, Glu152, and His256 for one zinc ion, and His97, Asp117, and Asp179 for the other. Mammalian glutamate carboxypeptidase II converts folylpoly-γ-glutamate to pteroylglutamate (folate), the process required for intestinal uptake. It also degrades the neuropeptide Ac-Asp-Glu in the brain. It is predicted that the enzyme has two catalytic zinc ions.
Clans MJ and MK
Metalloproteases
Clan MJ includes only family M38, and β-aspartyl dipeptidase (E. coli) belongs to this family. Asp and Asn can undergo spontaneous cyclization reaction, which forms the peptide bond linked through its side chain β-carboxyl group rather than α-carboxyl in normal peptide. This isoaspartyl β-carboxyl bond is generally resistant to proteolysis, but it is cleaved by
β-aspartyl dipeptidase, releasing an L-isoaspartyl residue from the N-terminus of a dipeptide. The enzyme is homologous to bacterial dihydroorotase, which generates N-carbamoyl-Laspartate from (S)-dihydroorate, and other amidohydrolases. Metal ligands for these enzymes are considered to be one aspartic acid and four histidines with the DXHXH motif and possible two other histidines (Kim and Kim, 1998). Only family M22 comprises Clan MK, and contains O-sialoglycoprotein endopeptidases (Pasteurella haemlytica). The enzyme cleaves only proteins that are heavily O-sialoglycosylated such as glycophorin from erythrocytes and leukocyte surface antigen CD34, CD43, CD44, and CD45. The metal ion binding site is thought to be two histidines in the HXEXXH motif. The third ligand is not identified.
A Nickel Metalloprotease HybD Endopeptidase: Clan ML HybD endopeptidase (E. coli) belongs to family M52 of Clan ML. The enzyme cleaves the precursor of the large hydrogenase subunit after a histidine or arginine residue in the C-terminal consensus motif DPCXXCXXH(R). The two cysteines of this motif and two cysteines in the N-terminal motif RXCXXC are ligands to the nickel or both nickel and iron of the hydrogenase. The metal ion of HybD endopeptidase is coordinated with Glu16, Asp62, His93, and a water molecule (Fritsche et al., 1999). The metal ion that coordinates this site is postulated to be the nickel ion that binds to the C-terminal nickel binding motif of the hydrogenase precursor. Interestingly, the processing of the hydrogenase precursor by the enzyme is not inhibited by phenylmethlysulfonyl fluoride, EDTA, or EGTA (Rossman et al., 1995).
Membrane Metalloproteases with an Intramembranous Active Site: Clan MM Two metalloproteases are thought to have their active site within the membrane and the cleavage site of the substrate is also located within the potential transmembrane segment (Lewis and Thomas, 1999; Yu and Kroos, 2000). A first example is human S2P protease, which releases the membrane-bound transcription factors—i.e., the so-called sterol-regulatory element-binding proteins (SREBPs)— from rough endoplasmic reticulum by cleaving a peptide bond located within a transmembrane domain. S2P is a polytopic membrane protein containing a putative zinc-binding motif HEXXH; the active site is thought to be located
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within the membrane of rough endoplasmic reticulum (Rawson et al., 1997). Additional family members are found in Drosophila melanogaster, Caenorhabditis elegans, Schistosoma mansoni, and Sulfolobus solfataricus. These enzymes are classified as family M50 of Clan MM. Another metalloprotease whose active site is located within the membrane is sporulation factor SpoIVFB (Bacillus subtilis), which processes pro-σK factor to σK, and is involved in transcriptional regulation of genes associated with morphogenesis. SpoIVFB also contains the HEXXH motif. The enzyme is classified as a member of family M5 of Clan MM.
Families That Are Not Assigned to a Clan As mentioned above, eight metallopeptidase families that have the HEXXH motif are not assigned to clans. In addition, there are five families that do not possess the HEXXH motif in the protein sequence and are therefore not assigned to clans (see UNIT 21.1). They include GPI-anchored membrane dipeptidase (family 19), β-lytic metalloendopeptidase (Achromobacter lyticus; family M23), aminopeptidase T (Thermus aquaticus; family M29), lysostaphin (Staphylococcus simulans; family M37), and dipeptidyl-peptidase III (metallo-form from Rattus norvegicus; family M49). The major physiological substrate of membrane dipeptidase is thought to be leukotriene D4 (peptidyl leukotriene), a potent broncho- and vasoconstrictor, which is converted to inactive leukotriene E4. β-Lytic metallopeptidase has an ability to lyse other bacteria and some soil nematodes, and lysostaphin cleaves staphylococci peptidoglycans and especially lyses Staphylococcus aureus.
CATALYTIC MECHANISMS The 3-D structures of many metallopeptidases have been determined and has suggested the mechanisms of peptide bond hydrolysis by metallopeptidases. The two most extensively studied enzymes are thermolysin (Matthews, 1988) and carboxypeptidase A (Christianson and Lipscomb, 1989). The mechanics of peptide hydrolysis by thermolysin is shown in Figure 21.4.1. In thermolysin, two histidines of the HEXXH motif (residues 142 to 146), Glu166 located 20 residues C-terminal from the motif, and a water molecule are the four ligands for the catalytic zinc ion. Upon binding of the peptide substrate to the active site of the enzyme, the water molecule is displaced from
the zinc atom and the carbonyl oxygen of the substrate interacts with the zinc. The nucleophilic attack on the peptide bond is mediated by a water molecule. This reaction is assisted by the carbonyl group of Glu143 in the HEXXH motif, which serves as a general base to draw a proton from the water molecule and assists the nucleophilic attack of the water molecule on the carbonyl carbon of the scissile bond. The carbonyl oxygen of the tetrahedral intermediate of the substrate is then stabilized by protons donated from Tyr157 and His231. The Glu143 donates a proton to the nitrogen of the leaving group. During this reaction no covalent intermediates are formed between the substrate and the enzyme. A similar catalytic mechanism has been proposed for MMPs and astacins; however, MMPs lack residues equivalent to Tyr157 and His231. In the case of astacin (family M12) and serralysin (family M10), a tyrosine residue (Tyr149 and Tyr216, respectively) occupies the same position as His231 of thermolysin. This tyrosine is a fourth protein ligand of the catalytic zinc ion in astacin and serralysin. When a transition analog inhibitor is bound to astacin, Tyr149 moves away from the zinc ion and forms a hydrogen bond with the inhibitor. A similar catalytic mechanism also applies to carboxypeptidase A. In this case, however, Arg145 interacts with the C-terminal carboxyl group of the substrate or a peptide inhibitor analog, and this interaction is stabilized by protons donated by Asn144 and Tyr248. A hydrogen bond is formed between Arg127 and an oxygen from a putative tetrahedral intermediate (Christianson and Lipscomb, 1989).
MOSAIC STRUCTURES OF METALLOPEPTIDASES AND SUBSTRATE SPECIFICITY A number of metallopeptidases are mosaic proteins containing domains without catalytic activities. These noncatalytic domains have been shown to be essential for some enzymes to express particular proteolytic activities and biological functions. In many cases, however, the functions of noncatalytic domains are yet to be investigated. Some examples are discussed below. Collagenases (MMPs-1, -8, -13, and -18) in the matrixins consist of an N-terminal protease domain and a C-terminal hemopexin-like domain, which are linked by a proline-rich hinge region. While the catalytic domain alone hydrolyzes synthetic peptides or noncollagenous proteins, without the hemopexin-like domain
Peptidases
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Glu143 Phe114 Ala113
O (–)
O
O
Glu143 Phe114 Ala113
O
Asn112
O
O
Asn112
H H
H
O
O R1
CH
H N
C
CH
CH
NH R1′ R1′ + HN
OH Tyr157
O
Zn
Tyr157
O (–)
Asn112
R1
NH
R1′
HN Zn
Asn112 NH2
O
H3 CH C(–) +N CH
+
2+
O
O
O
NH2
HH C N CH
OH
His231
Glu143 Phe114 Ala113
+
O(–)
HN 2+
O
CH
NH
R1′ +
Tyr157
O
H R1
NH2
H N CH
OH
His231
O
C O(–)
Glu143 Phe114 Ala113
O (–)
O
O R1
O 2+ + Zn Zn2
H
NH2
His231
O
R1′
OH Tyr157
+
Zn2
NH H+N
His231
Figure 21.4.1 Mechanism of peptide hydrolysis proposed for thermolysin. The substrate is shown in bold.
Metalloproteases
these enzymes are unable to cleave triple helical collagens. Hemopexin-like domains are found in most other matrixins (e.g., stromelysins, gelatinases, metalloelastase, membrane-type MMPs), but they do not appear to participate in determining their substrate specificities as examined so far; however, inactive pro-forms of MMP-2 (gelatinase A) and MMP-9 (gelatinase B) bind to TIMP-2 and TIMP-1, respectively (Woessner and Nagase, 2000). These interactions are through the C-terminal hemopexin domain of the proenzyme and the C-terminal domain of the inhibitor. The N-terminal inhibitory domain of the TIMP is therefore unblocked, and the complex functions as an inhibitor. Gelatinases A (MMP-2) and B
(MMP-9) possess three repeats of a fibronectin type II-like domain inserted into the protease domain. They are essential for efficient gelatinolytic and elastolytic activities of these metalloproteases (for review see Woessner and Nagase, 2000). ADAM metalloproteases are membranebound enzymes. The extracellular domains of ADAMs consist of domains similar to those found in snake venom reprolysins containing a disintegrin-like domain and a cysteine-rich domain. The disintegrin-like domains of snake venom reprolysins bind to the platelet integrin GPIIb/IIa (αIIb/β3) and prevent platelet aggregation. The disintegrin domain of ADAM2 (fertilin β) binds to integrin α6β1, and that of
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ADAM15 to integrins α6β1, αvβ3, and α5β1 (for review see Schlöndorff and Blobel, 1999). Some ADAMs shed extracellular domains of transmembrane proteins as demonstrated for cytokines, growth factors, receptors, adhesion molecules, and other cell surface proteins. It is postulated that interaction of noncatalytic domains with cell surface molecules is important for shedding of cell surface molecules, but it is yet to be elucidated how these domains affect the hydrolysis of protein substrates (Black and White, 1998). The members of the astacin subfamily M12A, except for astacin, are also mosaic proteins. Procollagen C-endopeptidase has three CUB (i.e., complement proteins (C1r/C1s)Uegf-BMP1) domains and an epidermal growth factor (EGF)-like domain. Meprins have a MAM (i.e., meprin, A-5 protein, receptor protein tyrosine phosphatase µ) and a MATH (i.e., meprin and TRAF homology) domain, a domain called the AM (i.e., after Math), an EGF-like domain, a transmembrane domain, and a cytoplasmic domain. The functions of these domains in terms of the protease action are not clearly understood. Tetanus and botulism neurotoxins are produced as a single polypeptide of 150 kDa consisting of three domains. Proteolysis of an exposed loop generates the heavy chain (100 kDa) with Hc and Hn domains, and the light chain (50 kDa) of the metalloprotease, either tentoxilysin or bontoxilysin (family M27). The Hc domain is responsible for binding to the presynaptic terminal of the neuromuscular junction and the Hn domain is involved in translocation of the metalloprotease domain into the cytoplasm. Thus, nonproteolytic domains assist to locate the metalloproteinase domain at the site where they cleave vesicle-associated membrane proteins. The yeast and human type I methionyl aminopeptidases have a 125- and 134-amino acid N-terminal extension, respectively. These Nterminal regions possess putative zinc finger consensus sequences of the type C2C2 and C2H2. The function of these domains is not known, but it is suggested that they may be involved in ribosomal binding and a cotranslational processing of the N-terminal methionyl residue. Human type II methionyl aminopeptidase also has an N-terminal extension, but there is no significant sequence similarities between type I and type II in this region. The type II extension does not have any recognizable motif. Bacterial equivalents lack the N-terminal extension.
There are sufficient examples that demonstrate the importance of noncatalytic domains of metallopeptidases for the expression of specific biological functions of protease specificities; however, further studies are clearly necessary to investigate possible roles of noncatalytic domains in determining protease specificity. Identification of such functions may allow us to propose novel means to intervene in specific proteolytic activities.
METALLOPROTEASE INHIBITORS Protease inhibitors are essential partners for the investigation of proteolytic enzymes. They are useful not only for characterization of proteases but also for inhibition of unwanted proteolysis in an experimental system. Inhibitors may also have therapeutic value. Chelating agents such as EDTA and 1,10-phenanthroline are commonly used to block metallopeptidase activities in in vitro experiments. Synthetic inhibitors have been designed to inhibit specific metalloproteases. They commonly contain a chelating moiety, such as a carboxyl, a thiol, a phosphorous, or a hydroxamic acid group. The chelating group is attached to a series of other groups that fit the specificity pocket of a particular metallopeptidase. Because unbalanced activities of metalloproteases are involved in a variety of diseases, they are considered good therapeutic targets. Captopril, a thiol compound, is a potent inhibitor of peptidyl dipeptidase A (angiotensin I converting enzyme), and is used to treat hypertension. A recent advancement includes the design of a compound with a meta-substituted benzofused macrocyclic lactam that inhibits both peptidyl dipeptidase A and neprilysin with IC50 values of 4 nM and 8 nM, respectively (Ksander et al., 1997). Such dual inhibitors may be potent antihypertensives due to an increase of the circulating levels of arterial natriuretic peptide resulting from the inhibition of neprilysin and also due to a decrease of angiotensin II resulting from the inhibition of peptidyl dipeptidase A. Many synthetic inhibitors have been designed for MMPs, enzymes that participate in breakdown of extracellular matrix compounds (Clendeninn and Appelt, 2000; Woessner and Nagase, 2000). Some inhibitors selective towards particular MMPs have been generated, but they were tested with only a limited number of MMPs. Furthermore, many MMP inhibitors appear to inhibit metalloproteases other than MMPs such as ADAMs. Because many of these enzymes are likely to be biologically important, a longterm use of metalloprotease inhibitors with the
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Table 21.4.3
Natural Protein Inhibitors of Metallopeptidases
Inhibitor
Target enzyme
Metalloprotease inhibitor from Habu snake serum Oprin
Habu venom metalloproteases
Potato carboxypeptidase inhibitor Serralysin inhibitor Streptomyces metalloproteinase inhibitor TIMP-1 TIMP-2 TIMP-3 TIMP-4
Metalloproteases
Adamolysin, Atrolysin A and B, Adamolysin Carboxypeptidase B Serralysin Thermolysin (Pseudomonas aeruginosa) metalloproteases, Thermolysin MMPs (poor for MMP-14) MMPs ADAM10, ADAM17, ADAM-TS4, ADAM-TS5, MMPs MMPs
aim of blocking MMP activity may require careful scrutiny. While a large number of synthetic metalloprotease inhibitors have been designed, only a small number of protein inhibitors of metalloproteases have been found (Table 21.4.3). These inhibitors are often specific to a particular family or subfamily of metallopeptidases. For example, tissue inhibitors of metalloproteinases (TIMPs) inhibit MMPs, but not metalloproteases belonging to other families. Currently four isoforms of TIMPs (TIMPs 1 to 4) have been described; however, recent studies showed that TIMP-3, but not other TIMPs, can inhibit ADAM17 (Amour et al., 1998). Other examples found in vertebrates are oprin from opossum serum that inhibits snake venom metalloprotease atrolysin A and B (Catanese and Kress, 1992) and a 323-amino acid glycoprotein from the Habu snake Trimerasurus flavoviridis that inhibits metalloproteases in Habu venom (Yamakawa and Omori-Satoh, 1992). The inhibitor from Habu contains two copies of a cystatin-like domain, but it does not inhibit cysteine proteases. A 12-kDa inhibitor from Streptomyces nigrescens TK-23 inhibits thermolysin and Pseudomonas aeruginosa metalloproteases (Seeram et al., 1997). Microbial protein inhibitors of 10 kDa for serralysins are also found in Erwinia chrysanthemi (Letoffe et al., 1989), Serratia marcescens (Kim et al., 1995), and P. aeruginosa (Duong et al., 1992). A carboxypeptidase inhibitor is found in potatoes (Ryan et al., 1974). Because many metallopeptidases play important roles in biological and pathological processes, these enzyme activities are likely to
be regulated by the endogenous inhibitors. In this regard it is surprising to see only a very limited number of natural inhibitors have been found so far. Future efforts to search for naturally occurring protein inhibitors of metalloproteases are necessary for our understanding of the balance of metalloprotease activities. The gene coding for such inhibitors may be useful for tissue-targeted delivery for therapeutic purposes.
LITERATURE CITED Amour, A., Slocombe, P.M., Webster, A., Butler, M., Knight, S.G., Smith, B.J., Stephens, P.E., Shelley, C., Hutton, M., Knäuper, V., Docherty, A.J., and Murphy, G. 1998. TNF-α converting enzyme (TACE) is inhibited by TIMP-3. FEBS Lett. 435:39-44. Barrett, A.J., Rawlings, N.D., and Woessner, J.F. (eds.) 1998. Handbook of Proteolytic Enzymes. Academic Press, London. Black, R.A. and White, J.M. 1998. ADAMs: Focus on the protease domain. Curr. Opin. Cell Biol. 10:654-659. Bode, W., Gomis-Rüth, F.-X., and Stockler, W. 1993. Astacins, serralysins, snake venom and matrix metalloproteinases exhibit identical zincbinding environments (HEXXHXXGXXH and Met-turn) and topologies and should be grouped into a common family, the “metzincins.” FEBS Lett. 331:134-140. Bradshaw, R.A., Arfin, S.M., and Walker, K.W. 1998. Methionyl aminopeptidase type II. In Handbook of Proteolytic Enzymes (A.J. Barrett, N.D. Rawlings, and J.F. Woessner, eds.) pp. 1399-1403. Academic Press, London. Catanese, J.J. and Kress, L.F. 1992. Isolation from opossum serum of a metalloproteinases inhibitor homologous to human α 1B-glycoprotein. Biochemistry 31:410-418.
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Christianson, D.W. and Lipscomb, W.N. 1989. Carboxypeptidase A. Acc. Chem. Res. 22:62-69. Clendeninn, N.J. and Appelt, K. 2000. Matrix Metalloproteinase Inhibitors in Cancer Therapy. Humana Press, Totowa, N.J. Duong, F., Lazdunski, A., Cami, B., and Murgier, M. 1992. Sequence of a cluster of genes controlling synthesis and secretion of alkaline protease in Pseudomonas aeruginosa: Relationship to other secretory pathways? Gene 121:47-54. Fritsche, E., Paschos, A., Beisel, A.-G., Bock, A., and Huber, R. 1999. Crystal structure of the hydrogenase maturating endopeptidase HYBD from Escherichia coli. J. Mol. Biol. 288:989998. Hooper, H.M. 1996. Zinc Metalloproteases in Health and Disease. Taytor and Francis, London. Hooper, N.M. 1994. Families of zinc metalloproteinases. FEBS Lett. 354:1-6. Kim, G. and Kim, H.-S. 1998. Identification of the structural similarity in the functionally related amidohydrolases acting on the cyclic amide ring. Biochem. J. 330:295-302. Kim, K.S., Kim, T.U., Kim, I.J., Byun, S.M., and Shin, Y.C. 1995. Characterization of a metalloprotease inhibitor protein (SmaPI) of Serratia marcescens. Appl. Environ. Microbiol. 61:30353041. Ksander, G.M., de Jesus, R., Yuan, A., Ghai, R.D., McMartin, C., and Bohacek, R. 1997. Meta-substituted benzofused macrocyclic lactams as zinc metalloprotease inhibitors. J. Med. Chem. 40:506-514. Lawrence, J.B., Oxvig, C., Overgaard, M.T., Sottrup-Jensen, L., Gleich, G.J., Hays, L.G., Yates, J.R. III, and Conover, C.A. 1999. The insulinlike growth factor (IGF)-dependent IGF binding protein-4 protease secreted by human fibroblasts is pregnancy-associated plasma protein-A. Proc. Natl. Acad. Sci. U.S.A. 96:3149-3153. Letoffe, S., Delepelaire, P., and Wandersman, C. 1989. Characterization of a protein inhibitor of extracellular proteases produced by Erwinia chrysanthemi. Mol. Microbiol. 3:79-86. Lewis, A.P. and Thomas, P.J. 1999. A novel clan of zinc metallopeptidases with possible intramembrane cleavage properties. Protein Sci. 8:439442. Matthews, B.W. 1988. Structural basis of the actions of thermolysin and related zinc peptidases. Acc. Chem. Res. 21:333-340.
Rawlings, N.D. and Barrett, A.J. 2000 MEROPS: The peptidase database. Nucl. Acids Res. 28:323325. Rawson, R.B., Zelenski, N.G., Nijhawan, D., Ye, J., Sakai, J., Hasan, M.T., Chang, T.Y., Brown, M.S., and Goldstein, J.L. 1997. Complementation cloning of S2P, a gene encoding a putative metalloprotease required for intramembrane cleavage of SREBPs. Mol. Cell 1:47-57. Rossman, R., Maier, T., Lottspeich, F., and Böck, A. 1995. Characterization of a protease from Escherichia coli involved in hydrogenase maturation. Eur. J. Biochem. 227:545-550. Ryan, C.A., Hass, G.M., and Kuhn, R.W. 1974. Purification and properties of a carboxypeptidase inhibitor from potatoes. J. Biol. Chem. 249:5495-5499. Schlöndorff, J. and Blobel, C.P. 1999. Metalloprotease-disintegrins: Modular proteins capable of promoting cell-cell interactions and triggering signals by protein-ectodomain shedding. J. Cell Sci. 112:3603-3617. Seeram, S.S., Hiraga, K., Saji, A., Tashiro, M., and Oda, K. 1997. Identification of reactive site of a proteinaceous metalloproteinase inhibitor from Streptomyces nigrescens TK-23 J. Biochem. 121:1088-1095. Sträter, N. and Lipscomb, W.N. 1998. Leucyl aminopeptidase (animal and plant). In Handbook of Proteolytic Enzymes (A.J. Barrett, N.D. Rawlings, and J.F. Woessner, eds.) pp. 1384-1389. Academic Press, London. Woessner, J.F. and Nagase, H. 2000. Matrix Metalloproteinases and TIMPs. Oxford University Press, Oxford, UK. Yamakawa, Y. and Omori-Satoh, T. 1992. Primary structure of the antihemorrhagic factor in serum of the Japanese Habu: A snake venom metalloproteinase inhibitor with a double-headed cystatin domain. J. Biochem. 112:583-589. Yu, Y.T. and Kroos, L. 2000. Evidence that SpoIVFB is a novel type of membrane metalloprotease governing intercompartmental communication during Bacillus subtilis sporulation. J. Bacteriol. 182:3305-3309.
Contributed by Hideaki Nagase Imperial College School of Medicine London, United Kingdom
Peptidases
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Purification and Characterization of Proteasomes from Saccharomyces cerevisiae
UNIT 21.5
The proteasome plays a central role in eukaryotic cells, since it is responsible for the degradation of specific proteins involved in a large range of cellular processes. Analysis of proteasome mechanisms of action, or in vitro reconstitution, or dissection of the complex biological pathways in which it partakes, requires having a reliable source of pure active proteasome. Although the biologically relevant form of the proteasome is usually considered to be the 26S proteasome, this unit describes different methods for purification and study of both 26S and 20S proteasomes from Saccharomyces cerevisiae cells. The 20S proteasome, which serves as the proteolytic core of the 26S proteasome, is relatively easy to isolate due to its resistance to harsh chemical and physical treatments. Satisfactory purification of Saccharomyces cerevisiae 26S proteasome has proven more difficult to achieve, the major problem being to avoid its dissociation into its subcomponents (i.e., the 20S proteasome and the 19S regulatory particle); however, over the past years, several procedures have been published that allow efficient purification of 26S proteasomes for in vitro studies. Most of them rely on classical chromatographic steps (i.e., mainly anion-exchange and gel filtration), although affinity chromatography using specific tags on proteasome subunits has also been used more recently. In the present unit, protocols allowing rapid purification of pure 26S (see Basic Protocol 1 and Alternate Protocol) and 20S (see Basic Protocol 2) proteasomes from yeast cells are described, and Support Protocols 1 to 5 detail the various assays that can be used to assess proteasome activity. It is recommended that the vacuolar proteases–deficient strain pep4 be used; however, these protocols have been successfully used on a variety of strains, including wild-types. It is worth emphasizing that these procedures can also be used to purify (to a reasonable level) proteasomes from mammalian cells. PURIFICATION OF YEAST 26S PROTEASOME BY ANION-EXCHANGE CHROMATOGRAPHY AND GEL FILTRATION
BASIC PROTOCOL 1
This protocol is based on published procedures (Rubin et al., 1996; Glickman et al., 1998a). It should lead to consistent results provided that certain considerations are respected (see Critical Parameters). Materials Saccharomyces cerevisiae (e.g., pep4) cells YPD (APPENDIX 4L) or other appropriate medium 26S lysis buffer (see recipe) 1 M Tris base DEAE Affi-Gel Blue resin (Bio-Rad) 26S chromatography buffer (see recipe) with and without 50, 100, 150, and 500 mM NaCl High-resolution “Q” anion-exchange resin (e.g., Pharmacia Mono-Q or Resource Q; Bio-Rad Bio-scale Q); also see UNIT 8.2 Gel-filtration resin with high-molecular-weight resolution (e.g., Sephacryl S-400HR or Superose 6HR; Amersham/Pharmacia); also see UNIT 8.3 French press or equivalent Cheesecloth Appropriately sized chromatography columns (see Critical Parameters) FPLC-type chromatography system Contributed by Michael Glickman and Olivier Coux Current Protocols in Protein Science (2001) 21.5.1- 21.5.17 Copyright © 2001 by John Wiley & Sons, Inc.
Peptidases
21.5.1 Supplement 24
Additional reagents and equipment for testing peptidase activity (see Support Protocol 1) and performing ion-exchange (UNIT 8.2) and gel-filtration chromatography (UNIT 8.3) Prepare lysate 1. Grow Saccharomyces cerevisiae cells to an OD of >1 in YPD or other appropriate medium. Pellet cells by brief centrifugation (e.g., 10 min at 2000 to 3000 × g). 2. Resuspend cell pellet in 2 vols 26S lysis buffer. 3. Lyse cells using a French press or other device appropriate for lysing large quantities of yeast cells. Yeast cells are efficiently lysed using a French press or Mill device. A bead-beater is not recommended for proteasome purification and sonication is inefficient for lysis of yeast cells.
4. Titrate cell extract to pH 7.4 using 1 M Tris base. 5. Clarify extract by centrifugation for 20 min at 30,000 × g, 4°C. Pass supernatant through two layers of cheesecloth. Perform DEAE Affi-Gel Blue chromatography 6. Load extract onto a DEAE Affi-Gel Blue column equilibrated with 26S chromatography buffer without NaCl. The size of the columns used will vary based on the amount of proteasome to be purified. The rule of thumb is 1 ml DEAE Affi-Gel Blue resin for each milliliter pelleted yeast cells. This first fractionation of the yeast extract can be done using a low-pressure purification system or even a peristaltic pump.
7. Rinse with one column volume of 26S chromatography buffer without NaCl using an FPLC-like chromatography system, followed by three column volumes 26S chromatography buffer supplemented with 50 mM NaCl. 8. Elute with three column volumes of 26S chromatography buffer supplemented with 150 mM NaCl, collecting 8 to 10 fractions. 9. Identify fractions containing proteasomes using the peptidase activity assay (see Support Protocol 1). Perform high-resolution “Q” anion-exchange chromatography 10. Load proteasome-containing fractions onto a high-resolution “Q” anion-exchange column (also see UNIT 8.2) equilibrated in 26S chromatography buffer supplemented with 100 mM NaCl. 11. Rinse the column with 26S chromatography buffer until the OD returns to baseline. 12. Elute bound proteins with 26S chromatography buffer, using a linear gradient from 100 mM NaCl to 500 mM NaCl over 10 column volumes. Collect 15 to 20 fractions. Proteasomes elute at a salt concentration of around 300 to 330 mM NaCl. The size of the anion-exchange and gel-filtration columns needs to be calculated based on total protein yield at each step and on the protein-binding capacity of these resins (see manufacturer’s instructions).
13. Identify proteasome-containing fractions by testing for peptidase activity (see Support Protocol 1). Purification of Yeast Proteasome
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14. Concentrate fractions that contain peptidase activity to a volume suitable for loading onto a gel-filtration column. A quick and efficient method for concentration is a spin filter with a molecular weight cutoff of 30,000 (e.g., Centricon 30; Amicon). Concentration should also include changing the buffer to 26S chromatography buffer in order to remove excess NaCl after the anion exchange step.
Perform gel-filtration chromatography 15. Load concentrated fractions onto a gel-filtration column (also see UNIT 8.3) with high-molecular-weight resolution, equilibrated with 26S chromatography buffer supplemented with 100 mM NaCl. 16. Elute the proteasome isocratically in 26S chromatography buffer supplemented with 100 mM NaCl. A rather wide peak of peptidase activity will elute immediately after the dead volume, corresponding to the multiple forms of the 26S proteasome followed by 20S proteasome.
17. Pool the first 30% to 50% of peak fractions. Concentrate if necessary. This corresponds to the higher-molecular-weight forms of the 26S proteasome (see Fig. 21.5.1). The remainder of the peak is enriched in 20S and is also contaminated with additional proteins; however, it can be further purified if necessary. A more accurate assessment of purity and distribution of 26S and 20S forms can be achieved by native PAGE (see Support Protocol 5), or by testing column fractions for peptidase activity both in the absence and in the presence of 0.02% (w/v) SDS (see Support Protocol 1).
26S
20S
26Sa 26Sb
20S
Figure 21.5.1 Visualization of proteasome using in-gel peptidase overlay assay. Purified 26S and 20S proteasomes (left lane and right lane, respectively) were resolved by nondenaturing PAGE as described (see Support Protocol 5). The gel was incubated in 0.1 mM LLVY-AMC and photographed under UV light. Fluorescent bands indicate the presence of free AMC, which is released by the proteases in the gel. Note that the 26S proteasome appears in two forms: the slower migrating symmetric form in which two 19S regulatory caps are attached to a single 20S core (26Sa), and a slightly faster migrating asymmetric form, composed of one 19S and one 20S subcomplex (26Sb). Free 20S migrates faster than either of the 26S proteasomes.
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ALTERNATE PROTOCOL
PURIFICATION OF YEAST 26S PROTEASOME BY AFFINITY CHROMATOGRAPHY Large-scale affinity purification of 26S proteasome by epitope-tagging a single proteasomal subunit is problematic due to the high molecular weight of the proteasome (2.5 MDa) and the large number of subunits (≥64). Nevertheless, for specific purposes, affinity purification can be rapid and sufficient. For example, nickel-NTA affinity purification can be used to rapidly enrich the extract with active proteasomes from strains expressing His6 epitope–tagged proteasomes. Likewise, immunopurification of proteasomes with epitope tags can yield highly purified proteasomes (Enenkel et al., 1998; Verma et al., 2000); however, certain tags require elution conditions that inactivate the complex. A number of yeast strains expressing single 20S or 19S subunits tagged with the His6 epitope are available (Rubin et al., 1996; Glickman et al., 1998b) and nickel-NTA affinity purification can be used to rapidly obtain partially purified 26S proteasomes from these strains. Purification can follow standard published protocols for nickel-NTA affinity purification directly from clarified yeast cell extracts (see Basic Protocol 1, steps 1 to 5); however, it should be noted that after loading the column with cell extract, nonspecifically bound proteins should be washed off using 26S chromatography buffer (see recipe) supplemented with 15 mM imidazole and 100 mM NaCl. Proteasomes are then eluted with 26S chromatography buffer supplemented with 100 mM imidazole and 100 mM NaCl.
BASIC PROTOCOL 2
PURIFICATION OF YEAST 20S PROTEASOME BY CONVENTIONAL CHROMATOGRAPHY In order to purify 20S instead of 26S proteasomes, several modifications (as compared with 26S proteasome purification; see Basic Protocol 1) are introduced in this procedure. Most importantly, Mg2+ and ATP are omitted in the lysis buffer and during the first steps, in order to promote dissociation of the 26S proteasome. Addition of NaN3 in the lysis buffer helps to block ATP regeneration in the extract. Materials Saccharomyces cerevisiae (e.g., pep4) cells YPD (APPENDIX 4L) or other appropriate medium 20S lysis buffer (see recipe) 1 M Tris base High-capacity anion-exchange resin (e.g., Q-Sepharose or Q-Fast Flow; Amersham); also see UNIT 8.2 25 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E), with 100 mM, 200 mM, and 1 M NaCl 1 M MgCl2 (APPENDIX 2E) Hydroxyapatite column (Bio-Rad); also see UNIT 8.6 40 and 400 mM potassium phosphate buffer pH 7.2 (APPENDIX 2E) High molecular weight–resolving gel (e.g., Sephacryl S300HR or Superose 6HR; Amersham); also see UNIT 8.3 French press or equivalent Cheesecloth Appropriately sized chromatography columns (see Critical Parameters) Additional reagents and equipment for ion-exchange (UNIT 8.2), gel-filtration (UNIT 8.3), and hydroxylapatite chromatography (UNIT 8.6), and testing peptidase activity (see Support Protocol 1) or performing the native gel procedure (see Support Protocol 5)
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Prepare lysate 1. Grow Saccharomyces cerevisiae to an OD of >1 in YPD or other appropriate medium. Pellet by brief centrifugation (e.g., 10 min at 2000 to 3000 × g). 2. Resuspend cell pellet in 2 vols 20S lysis buffer. 3. Lyse cells using a French press or other lysing device appropriate for lysing large quantities of yeast cells. Yeast cells are efficiently lysed using a French press or Mill device. A bead-beater is not recommended for proteasome purification and sonication is inefficient for lysis of yeast cells.
4. Titrate cell extract to pH 7.4 using 1 M Tris base. 5. Clarify extract by centrifugation for 20 min at 30,000 × g, 4°C. Pass supernatant through two layers of cheesecloth. Perform anion-exchange chromatography 6. Load extract onto a high-capacity anion-exchange column (e.g., Q-Sepharose; also see UNIT 8.2), equilibrated with 25 mM Tris⋅Cl, pH 7.4 with 200 mM NaCl. 7. Elute with 25 mM Tris⋅Cl using a linear gradient from 200 to 500 mM NaCl over 10-column volumes. 8. Identify and pool fractions containing 20S proteasome by testing for peptidase activity (see Support Protocol 1). Perform hydroxyapatite chromatography 9. Add 1 M MgCl2 to the sample to a final concentration of 10 mM (to enhance binding to the hydroxyapatite column), and load onto a hydroxyapatite column (also see UNIT 8.6) equilibrated with 40 mM potassium phosphate, pH 7.2. 10. Elute with linear gradient from 40 to 400 mM potassium phosphate over 20 column volumes. Collect 20 fractions. 20S particles elute at around 200 mM phosphate.
11. Pool fractions, concentrate, and exchange the buffer to 25 mM Tris⋅Cl, pH 7.4, containing 100 mM NaCl. A quick and efficient method for concentration is a spin filter with a molecular weight cutoff of 30,000 (e.g., Centricon 30; Amicon).
Perform gel-filtration chromatography 12. Load sample onto a high-molecular-weight resolving column. 13. Elute isocratically in 25 mM Tris⋅Cl, pH 7.4, supplemented with 100 mM NaCl. 14. Identify 20S containing fractions by screening for peptidase activity (see Support Protocol 1) or using the native gel procedure (see Support Protocol 5). PEPTIDASE ACTIVITY ASSAY FOR 26S AND 20S PROTEASOME During purification of proteasomes the authors recommend, whenever possible, that the activity be tested by using the fluorogenic peptide N-succinyl-Leu-Leu-Val-Tyr 7-amido4-methylcoumarin (N-suc-LLVY-AMC; Bachem or Sigma). This assay is fast, sensitive, easy to install, and rather specific for the 26S proteasome (i.e., the 20S proteasome can also hydrolyze this peptide, but its specific activity is much lower). Testing hydrolysis of this substrate in the presence or absence of low concentrations of SDS allows discrimination between the two forms of the proteasome (see below). The proteasome cleaves this peptide after the Tyr residue, liberating the AMC moiety that then fluoresces blue.
SUPPORT PROTOCOL 1
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Materials Protein fractions to be analyzed (see Basic Protocols 1 and 2 and Alternate Protocol) 10 mM N-suc-LLVY-AMC (Bachem or Sigma) in DMSO Activity buffer (see recipe) 1% (w/v) SDS 1-ml quartz or appropriate type of plastic cuvette Fluorometer with 380-nm excitation and 440-nm emission wavelength 1. Incubate protein fractions to be analyzed in the presence of 100 µM (final) N-sucLLVY-AMC in activity buffer (dilute the peptide from a 10 mM stock) with a reaction volume of 50 to 100 µl at 30°C. 2. Stop the reaction after 15 min by adding 1 ml 1% (w/v) SDS. 3. Measure fluorescence for each sample and a blank in a 1-ml quartz or appropriate type of plastic cuvette, with an excitation wavelength of 380 nm, and an emission wavelength of 440 nm. The blank (i.e., no protein) should have very little fluorescence in this assay. The same assay can also be performed in the presence of very low concentrations (i.e., ≤0.02%) of SDS; however, the optimal concentration has to be determined in each case. In the presence of 0.02% (w/v) SDS, peptidase activity of the 20S proteasome is activated several-fold, while the 26S proteasome is either unaffected or slightly inhibited; however, the proteasome should not be incubated with the SDS prior to addition of the peptide, since both 20 and 26S proteasomes can be rapidly inactivated by the SDS. This procedure allows easy discrimination between the two forms of the proteasome, and can be used to determine their respective elution profiles after chromatography. SUPPORT PROTOCOL 2
PREPARATION OF POLY-UBIQUITINATED LYSOZYME Although the peptidase assay described above is convenient and fast, and should be used in the course of the purification, peptides are not substrates of proteasomes in cells. Thus, this assay does not measure the physiologic activity of the complex. The ultimate activity test of the 26S proteasome is to measure the degradation of polyubiquitinated proteins that are the actual specific substrates of this complex in vivo but are not hydrolyzed by 20S proteasomes. Unfortunately, ubiquitinated proteins are hard to produce in large amounts. With time, it will become easier to perform in vitro ubiquitination of various proteins using recombinant ubiquitination enzymes as more and more ubiquitination pathways will be dissected; however, since at present those possibilities are limited, a method is presented here to prepare a polyubiquitinated lysozyme substrate, which can be performed in any laboratory equipped for handling 125I-protein samples. This procedure involves the purification of ubiquitination enzymes using a ubiquitin (Ub) affinity column. Chemical Radioactive Labeling of Lysozyme This procedure is derived from a published protocol (Parker, 1990).
Purification of Yeast Proteasome
Materials 10 mg/ml chicken egg white lysozyme 0.5 M potassium phosphate buffer, pH 7.6 (APPENDIX 2E) 100 mCi/ml Na[125I] (ICN or Amersham-Pharmacia Biotech) 3.6 mg/ml chloramine T in 0.5 M potassium phosphate buffer, pH 7.6 (APPENDIX 2E): prepare immediately before use 10 mg/ml sodium metabisulfite in 0.5 M potassium phosphate buffer, pH 7.6 (APPENDIX 2E): prepare immediately before use
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PD-10 column (Pharmacia) 20 mM Tris⋅Cl, pH 7.5/20 mM NaCl/1 M DTT (APPENDIX 2E) Gamma counter CAUTION: Iodine is extremely volatile, and all necessary precautions for handling radioactive vapor should be taken. Carefully follow the appropriate laboratory guidelines (APPENDIX 2B). 1. Combine the following in a 1.5-ml microcentrifuge tube: 100 µl 10 mg/ml lysozyme (1 mg) 180 µl 0.5 M potassium phosphate buffer 20 µl 100 mCi/ml Na[125I] (2 mCi) 100 µl 3.6 mg/ml chloramine T (360 µg) 2. Mix for 30 sec at room temperature. 3. Stop the reaction by adding 100 µl (1 mg) freshly prepared sodium metabisulfite in 0.5 M potassium phosphate buffer, pH 7.6. 4. Apply the reaction to a PD-10 column equilibrated with 20 mM Tris⋅Cl, pH 7.5/20 mM NaCl/1 mM DTT. 5. Collect ∼20 0.5-ml fractions. Count 1 µl of each fraction in a gamma counter and pool the fractions corresponding to the first peak of radioactivity. Purification of Ubiquitination Enzymes This procedure is derived from previous published work (Tamura et al., 1991). It allows the sequential elution of two heterogeneous fractions from a Ub-affinity column. The first fraction (DTT eluate) contains the Ub-activating enzyme E1 and the Ub-carrier proteins (E2s) present in reticulocyte fraction II. The second fraction (pH 9/KCl eluate) contains several proteins, including the so-called “N-end rule” Ub-protein ligase (E3α) necessary for lysozyme ubiquitination. Materials Ub-Sepharose column (Ciechanover et al., 1982) Ub-Sepharose loading buffer (see recipe) >25 mg protein/ml fraction II (FII; Hershko et al., 1983) supplemented to 5 mM MgCl2 and 2 mM ATP Ub-Sepharose wash buffer (see recipe) E1/E2 elution buffer (see recipe) E3 elution buffer (see recipe) 1 M Tris⋅Cl, pH 7.2 (APPENDIX 2E) 50 mM Tris⋅Cl, pH 7.2 (APPENDIX 2E)/0.02% NaN3 Ubiquitination enzymes storage buffer Ultrafiltration device (e.g., 10,000-MWCO Centricon; Amicon) NOTE: All steps are carried out at room temperature unless otherwise indicated. Use ∼1 ml Ub-Sepharose for every 100 mg FII. 1. Equilibrate a Ub-Sepharose column with 4 column volumes Ub-Sepharose loading buffer. The Ub-Sepharose column (10 to 20 mg Ub/ml resin) is prepared as described in Ciechanover et al. (1982). Use activated-CH Sepharose (Pharmacia), according to the manufacturers instructions.
2. Load FII supplemented to 5 mM MgCl2 and 2 mM ATP onto the column at a low flow rate. Collect the flow through and reload it or, alternatively, combine the Ub-Sepharose resin and FII with agitation at room temperature.
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FII is DEAE-bound proteins usually prepared from rabbit reticulocytes as in Hershko et al. (1983). The supplemented Mg2+ and ATP are necessary for activation of Ub by E1.
3. Wash the column with 4 column volumes Ub-Sepharose loading buffer. 4. Wash the column with 4 column volumes Ub-Sepharose wash buffer. 5. Add 4 column volumes E1/E2 elution buffer and collect the eluate on ice. The eluate (i.e., the DTT eluate) should contain E1 and E2.
6. Add 6 column volumes E3 elution buffer. Collect the fractions on ice in a tube containing 1⁄10 the elution volume 1 M Tris⋅Cl, pH 7.2. The fraction (i.e., the pH 9/KCl eluate) should contain E3. The 1 M Tris⋅Cl, pH 7.2, is added to neutralize the eluate.
7. Wash the column with 50 mM Tris⋅Cl, pH 7.2/0.02% NaN3. The column can be stored at 4°C for future use for months.
8. Concentrate the different eluates using an ultrafiltration device. 9. Change the buffer to ubiquitination enzymes storage buffer by successive dilution and concentration in the same device, or by dialysis (APPENDIX 3B). Store the enzymes at −80°C for weeks. Synthesis of Polyubiquitinated Lysozyme (Ub-Lysozyme) Materials >106 cpm/µg [125I]lysozyme E1 and E2 (i.e., DTT eluate) E3 (i.e., KCl/pH 9 eluate) 1 M Tris⋅Cl, pH 9.0 (APPENDIX 2E) 5 to 10 mg/ml ubiquitin (His6-tagged Ub recommended; see below) 0.1 M ATP 1 M MgCl2 (APPENDIX 2E) 1 M DTT (APPENDIX 2E) PD-10 column (Pharmacia) Ni-NTA column buffer (see recipe) with and without 10 mg/ml lysozyme Ni-NTA resin (Qiagen) 1 M stock imidazole, pH 8 Ub-lysozyme storage buffer (see recipe) Appropriately sized chromatography column (see Critical Parameters) Gamma counter Additional equipment and reagents for electrophoresis, autoradiography (UNIT 10.11), and dialysis (APPENDIX 3B) Set up the reaction 1. Mix the following and incubate 1 hr at 37°C:
Purification of Yeast Proteasome
50 µg 106 cpm/µg [125I]lysozyme ≥300 µg E1 and E2 (i.e., DTT eluate) ≥300 µg E3 (i.e., pH 9/KCl) 100 µl 1 M Tris⋅Cl, pH 9.0 (100 mM final) 1 to 1.5 mg/ml ubiquitin (if possible containing His6-Ub) 50 µl 0.1 M ATP (final 5 mM) 10 µl 1 M MgCl2 (final 10 mM) 1 µl 1 M DTT (final 1 mM) Add H2O to 1.0 ml
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In practice, use the DTT and pH 9/KCl eluates from the Ub-affinity column. Use as much material as possible, since these fractions correspond to a mixture of proteins. If possible, use a mixture of normal ubiquitin (Sigma) and recombinant His6-tagged ubiquitin (Beers and Callis, 1993). This enables purifying the Ub-lysozyme from unconjugated lysozyme using a nickel column. A mixture of 1⁄3 His6-Ub, 2⁄3 normal Ub is often used, since there are multiple Ub moieties per molecule Ub-lysozyme. After the reaction, keep an aliquot for analysis by SDS-PAGE and autoradiography. Although the reaction mixture will contain unconjugated lysozyme, it can be used “as-is” for degradation assays by the 26S proteasome, since the free lysozyme is not a substrate for this enzyme; however, if His6-Ub has been used for the conjugation reaction, it is possible to separate the free and the conjugated lysozyme. It is important to keep in mind that, after purification, the recovery of ubiquitinated lysozyme is low, due to the poor solubility of these molecules.
Purify Ub-Lysozyme by nickel-column chromatography 2. Equilibrate a PD-10 column with Ni-NTA column buffer supplemented to 1 mg/ml lysozyme. The lysozyme in the buffer is used to saturate the column for nonspecific binding.
3. Load the ubiquitination mixture on the column and collect approximately ten 1-ml fractions. Count 1 µl of each fraction with a gamma counter. Keep the one or two fractions containing the peak of radioactivity. 4. Load the ubiquitination mixture (i.e., the pooled fractions from the PD-10 column) onto a column of ∼0.5 ml Ni-NTA equilibrated with Ni-NTA column buffer. 5. Wash the column with several column volumes of Ni-NTA column buffer. 6. Elute the bound proteins using 2.5 column volumes Ni-NTA column buffer, containing increasing concentrations (i.e., 100, 200, 300, 400, and 500 mM) of imidazole, pH 8, diluted from a 1 M stock. The Ub-lysozyme should be eluted at 200 to 300 mM imidazole.
7. Count each fraction and analyze the Ub-lysozyme by electrophoresis and autoradiography (UNIT 10.11). 8. Dialyze (APPENDIX 3B) the fractions containing polyubiquitinated lysozyme against Ub-lysozyme storage buffer. Store frozen in aliquots at −80°C. The shelf life is dictated by the specific activity of the radiolabeled enzyme (i.e., how long it can be detected in a given application).
PREPARATION OF RADIOLABELED CASEIN This is a quick and easy method to make large amounts of a stable radiolabeled protein substrate that can be used to test proteasome activity. Casein hydrolysis by the proteasome can be tested together with Ub-lysozyme degradation in order to compare the ability of the proteasome to proteolyze both ubiquitinated and nonubiquitinated proteins. In addition, this is an important substrate to test the proteolytic activity of the 20S proteasome, since this complex cannot proteolyze ubiquitinated proteins.
SUPPORT PROTOCOL 3
Materials 15 mg/ml β-casein in H2O 200 µl 1 M potassium phosphate buffer, pH 8 (APPENDIX 2E) 250 µCi [14C]formaldehyde in ampule (ICN) 200 µl 0.2 M sodium cyanoborohydride (NaCNBH3) Small desalting column (e.g., PD-10 column; Pharmacia) Additional equipment and reagents for dialysis (APPENDIX 3B)
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1. Mix 1 ml 15 mg/ml β-casein and 200 µl 1 M potassium phosphate buffer, pH 8, and add directly into the [14C]formaldehyde ampule. 2. Immediately add 200 µl 0.2 M sodium cyanoborohydride (NaCNBH3). 3. Leave on ice 3 hr. 4. Remove unreacted [14C]formaldehyde and exchange buffer to desired buffer using a small desalting column or dialysis (APPENDIX 3B). SUPPORT PROTOCOL 4
DEGRADATION OF PROTEIN SUBSTRATES BY THE 26S PROTEASOME This protocol allows the rapid measurement of 26S proteasome activity against radioactive protein substrates, whether ubiquitinated or not (see Support Protocols 2 and 3). 20S proteasomes can be tested in this manner too, although in this case Mg2+ and ATP can be omitted from the reaction. After degradation, proteins are precipitated by TCA (trichloroacetic acid), which leaves the peptides generated by the proteasome soluble. After centrifugation, radioactivity in the supernatant will be proportional to the proteolytic activity of the enzyme tested. Additional Materials (also see Support Protocol 1) Enzyme (see Basic Protocols 1 and 2 and Alternate Protocol) Radioactive substrate (see Support Protocols 2 and 3) Activity buffer (see recipe) Carrier protein (e.g., BSA) 10% or 20% TCA Appropriate instrument for measuring radioactivity 1. Incubate the enzyme with the radioactive substrate in activity buffer for 30 min at 30°C. 2. After the reaction is completed, add 20 µg carrier protein (e.g., BSA), and 10% or 20% TCA to a final concentration of 5%. Keep on ice 15 min. 3. Microcentrifuge the precipitated undigested proteins 15 min at maximum speed. Recover the supernatant and count the radioactivity with the appropriate apparatus. A blank assay, with buffer instead of the enzyme, must be performed for each experiment to measure the background hydrolysis of the substrate. If possible, a measurement should be performed at time 0 and each assay should be done in duplicate or triplicate.
SUPPORT PROTOCOL 5
Purification of Yeast Proteasome
NATIVE GEL ELECTROPHORESIS/IN-GEL PEPTIDASE ASSAY OF THE PROTEASOME This procedure allows the researcher to rapidly resolve the different forms of the proteasome and to analyze their activity by in-gel hydrolysis of the peptide Suc-LLVYAMC. The method is relatively fast and presents two major advantages. First, it is very sensitive (26S proteasome present in a few microliters of cell extract can be visualized). Second, it is much better than any other method in resolving the different forms of the proteasome. Materials 1.5-mm-thick 4% polyacrylamide gel (see recipe) 1× native gel electrophoresis buffer, pH 8.1 to 8.4 (see recipe) 5× native gel loading buffer: 50% glycerol to which a tiny speck of xylene cyanol is added Proteasome sample (see Basic Protocols 1 and 2 and Alternate Protocol) Native gel assay buffer (see recipe)
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Electrophoresis apparatus UV lamp Camera and appropriate filters Additional equipment and reagents for casting and running native acrylamide gels (UNIT 10.3), and staining gels with Coomassie blue (UNIT 10.5) NOTE: All equipment and reagents must be prechilled to 4°C. 1. Polymerize a 1.5 mm thick 4% polyacrylamide gel. 2. Install the gel into an electrophoresis chamber. Use 1× native gel electrophoresis buffer for running buffer. Place at 4°C. 3. Dilute 5× native gel loading buffer (1:5 dilution) in the proteasome sample. Load an amount of the buffered sample that is appropriate to the size of the well onto the gel. 4. Run the gel at 80 to 100 V (∼20 mA) at 4°C, until the xylene cyanol dye runs out of the bottom of the gel (i.e., 1.5 to 2 hr). 5. Remove the gel carefully from the glass plates and incubate in 15 ml native gel assay buffer for 15 min at 30°C. The liquid must totally cover the gel to allow even diffusion of the substrate.
6. Analyze the gel using a UV lamp. Bands corresponding to the proteasome fluoresce blue (440 nm). Fluorescent bands can be photographed with any camera (fit appropriate filter if necessary). See the example in Figure 21.5.1.
7. Optional: Stain the gel with Coomassie blue (UNIT 10.5). The major difficulty in this procedure is to handle the gel after electrophoresis without breaking it. Two rules must be observed. First, be careful with the choice of gloves (some latex gloves stick strongly to the gel). Second, manipulate the gel as a whole, for example by rolling it prior to transfer from the gel plate to the incubation box, rather than “tugging” at its corners.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
20S lysis buffer 25 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) 1 mM EDTA (APPENDIX 2E) 1 mM NaN3 Store up to several weeks at 4°C. 26S lysis buffer Supplement 26S chromatography buffer (see recipe) to 4 mM ATP (final) from a 0.25 M stock. As this reagent contains ATP, prepare fresh. 26S chromatography buffer 10% (v/v) glycerol 25 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) 10 mM MgCl2 (APPENDIX 2E) 1 mM ATP 1 mM DTT (APPENDIX 2E) As this reagent contains ATP and DTT, prepare fresh Peptidases
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Activity buffer 20 mM Tris⋅Cl, pH 7.5, 30°C (APPENDIX 2E) 5 mM MgCl2 (APPENDIX 2E) 1 mM ATP 1 mM DTT (APPENDIX 2E) As this reagent contains ATP and DTT, prepare fresh E1/E2 elution buffer 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 20 mM DTT (APPENDIX 2E) 100 mM KCl (APPENDIX 2E) 0.1 mM EDTA (APPENDIX 2E) As this reagent contains DTT, prepare fresh E3 elution buffer 50 mM Tris⋅Cl, pH 9.0 (APPENDIX 2E) 1 M KCl (APPENDIX 2E) 2 mM DTT (APPENDIX 2E) As this reagent contains DTT, prepare fresh Native gel assay buffer 20 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) 5 mM MgCl2 (APPENDIX 2E) 1 mM ATP 0.1 mM Suc-LLVY-AMC (Bachem or Sigma) As this reagent contains ATP, prepare fresh Native gel electrophoresis buffer, pH 8.1 to 8.4, 5×, 1× 5× native electrophoresis buffer, pH 8.1 to 8.4 0.45 M Tris base 0.45 M boric acid 10 mM MgCl2 (APPENDIX 2E) Store up to several weeks at 4°C. 1× native gel electrophoresis buffer, pH 8.1 to 8.4 Dilute 5× native electrophoresis buffer with water and add 1 M DTT (APPENDIX 2E) and 0.25 M ATP to 1 mM (each). As this reagent contains ATP and DTT, prepare fresh. Ni-NTA column buffer 50 mM sodium phosphate buffer, pH 7.8 (APPENDIX 2E) 300 mM NaCl (APPENDIX 2E) 15 mM imidazole Store up to several weeks at 4°C. Polyacrylamide gel, 4%, 1.5-mm-thick 3 ml 5× native gel electrophoresis buffer (see recipe) 1.5 ml 40% 37.5:1 acrylamide:bisacrylamide 60 µl 0.25 M ATP (1 mM final) 15 µl 1 M DTT (1 mM final; APPENDIX 2E) The gel is polymerized by adding 120 µl 10% ammonium persulfate (APS) and 12 µl N,N,N′,N′-tetramethylethylenediamine (TEMED). See UNIT 10.3 for details on assembling native (nondenaturing) gels. The above recipe is for a minigel (15 ml). No stacking gel is necessary. Purification of Yeast Proteasome
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Ubiquitination enzymes storage buffer 20 mM Tris⋅Cl, pH 7.8 1 mM DTT 10% (v/v) glycerol As this reagent contains DTT, prepare fresh Ub-lysozyme storage buffer 20 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 5 mM NaCl (APPENDIX 2E) 20% glycerol Store up to several weeks at 4°C. Ub-sepharose loading buffer 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 2 mM ATP 5 mM MgCl2 (APPENDIX 2E) 0.2 mM DTT (APPENDIX 2E) As this reagent contains ATP and DTT, prepare fresh Ub-sepharose wash buffer 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 100 mM KCl (APPENDIX 2E) 0.1 mM EDTA (APPENDIX 2E) Store up to several weeks at 4°C. COMMENTARY Background Information The ubiquitin (Ub)–proteasome system is responsible for most of the intracellular proteolysis in growing cells, especially in the degradation of short-lived proteins (Rock et al., 1994; Hershko and Ciechanover, 1998). In this system, the proteolytic core is a 700-kDa threonine protease called the 20S proteasome. This enzyme is a hollow cylinder made of 28 (14 different) subunits assembled in four heptameric rings, and possesses at least six active sites that are facing into an internal proteolytic chamber (Seemuller et al., 1995; Groll et al., 1997). The 20S proteasome associates with the 19S regulatory complex (also called the regulatory particle) to form a 2500 kDa complex called the 26S proteasome that is responsible for degradation of polyubiquitinated proteins (Coux et al., 1996; Glickman et al., 1998b; Voges et al., 1999). Two forms of the 26S proteasome can be distinguished in crude cell extracts, corresponding to molecules with one 20S proteasome associated to one or two 19S complexes (see Figure 21.5.1); it is unclear at present whether both forms coexist in the cell. Association of the 19S complex to the 20S proteasome is a critical event, since it brings with it multiple activities essential for 26S proteasome function, including a subunit able to
bind polyubiquitin chains, an isopeptidase that catalyzes the release of free Ub, and several ATPases whose likely functions are to activate the 20S proteasome and to facilitate the unfolding of substrates or their translocation into the catalytic chamber (Rubin and Finley, 1995; Coux et al., 1996; Braun et al., 1999). It is thus likely that 26S proteasome assembly from its subcomplexes is a regulated process, as indicated by results obtained upon activation of oocytes of various species (Kawahara and Yokosawa, 1994). Although the mechanisms regulating this assembly are not understood, initial experiments showed that in ATP-depleted rabbit reticulocyte extracts, degradation of polyubiquitinated proteins could be reconstituted from three complexes: CF-1, -2, and -3, upon addition of ATP (Ganoth et al., 1988). Since it was later shown that CF-3 is the 20S proteasome (Eytan et al., 1989), it is believed that CF-1 and CF-2 together correspond to what is now termed the 19S complex, whose association to the 20S proteasome is also ATP-dependent (Chu-Ping et al., 1994). First identified in rabbit reticulocytes, the 26S proteasome has been purified from a variety of eukaryotic cells. It is a highly conserved molecule, both at the levels of structure and subunit composition. It must be emphasized, Peptidases
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Table 21.5.1
Troubleshooting Guide to Yeast Proteasome Purification
Problem
Possible cause
Solution
Low proteasome purification yield
pH drops after lysis
Check and recheck pH after lysis. Titrate with 1 M Tris base to pH 7.4. Work fast. Elute from Affi-Gel Blue column within 3 hr of lysis. Remake ATP and Mg2+ concentrations in the buffers. Enrich lysis buffer with additional ATP or Mg ions2+. Immediately exchange the buffer of proteasome- containing samples after anion-exchange step to a low-salt buffer (e.g., 26S chromatography buffer). Concentrate samples after each step Work fast. Run gel-filtration column overnight (day one). Use S400 instead of Superose 6, or vice versa, in gel filtration step Collect smaller fraction from gel-filtration step. Pool only homogeneous fractions as tested by SDS-PAGE. Add an additional purification step, such as affinity purification if applicable, or hydroxyapatite chromatography
Procedure takes too long Proteasome is purified mainly as 20S Mg⋅ATP concentrations are too low form, while very little 26S is present Exposure to high NaCl concentrations for too long
Samples are too dilute Entire procedure exceeds two days Contaminants in final preparation
Gel filtration step is inefficient Gel filtration fractions are too large
An additional purification step is necessary
Purification of Yeast Proteasome
however, that the purified complex is probably only the core structure of the enzyme that exists in vivo. A number of proteins that are not found in highly purified 26S proteasome, but which are nevertheless important for its proper functioning, are probably associated with it in cruder fractions (Glickman et al., 1998b; Papa et al., 1999). Purification of the 26S proteasome was achieved first in cells from higher eukaryotes (Peters et al., 1991). Most of the procedures relied on differential ultracentrifugation as an early step to separate the proteasome from the bulk of smaller intracellular proteins; however, when the same methods were applied to yeast extracts, they yielded essentially the 20S proteasome. A possibility was that during the long centrifugation the 26S proteasome could have been damaged by other proteins of the extract (e.g., the vacuolar proteases liberated upon breaking of the cells). Indeed, when the centrifugation step was omitted and instead a rapid ion-exchange chromatography step was performed, satisfactory purification of yeast proteasome was obtained (Rubin et al., 1996).
Critical Parameters As mentioned above, the major difficulty in purifying the 26S proteasome is to avoid its dissociation into its subcomplexes: the 20S proteasome and the 19S complex. Towards this goal, several general principles must be respected. It is important to work fast, since the level of dissociation of the 26S proteasome will increase with time. This first recommendation is especially true during the initial stages of the purification, until the complex is separated from the bulk of intracellular proteins, since other proteins in crude extracts seem able to damage the complex. As a rule of thumb, it seems fair to say that the faster the first ion exchange step is performed, the better the yield of the experiment. It is also recommended that all buffers be supplemented with MgCl2 and ATP, since Mg⋅ATP is necessary for 26S proteasome assembly and activity. In fact, when purified, the yeast complex reacts well to the absence of ATP at 4°C, indicating that Mg⋅ATP is not required for its stability at this temperature; however, this observation does not seem to apply to crude
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26S
110 KDa
19S
30 KDa
20S
Figure 21.5.2 SDS-PAGE migration-pattern of 26S subunits. Purified 26S proteasomes were analyzed by SDS-PAGE. The gel was stained with Coomassie blue. The 17 different 26S subunits range between 110 kDa and 30 kDa, while the 14 20S subunits migrate in between the 25 and 35 kDa range.
fractions and it should be emphasized that it also does not apply when the temperature is raised to physiologic values. At 30°C, the yeast 26S proteasome will dissociate rapidly in the absence of ATP. High salt concentrations are detrimental to 26S proteasome stability. This complex should never be subjected to NaCl concentrations above 400 mM, and should not be stored for prolonged periods of time in salt concentrations above 250 mM. It is important to keep the protein concentration as high as possible during the entire procedure to limit the dissociation of the complex. This recommendation implies that it is important to carefully determine the appropriate size of the columns to avoid unnecessary dilution of the sample. This is especially true for the gel-filtration step, during which dilution is inevitable. Also, as with many fragile multisubunit complexes, it is recommended that glycerol be added in all buffers, as it is known to stabilize the proteasome. Abstain from multiple freeze/thawing of the samples.
Store purified proteasomes as concentrated samples (≥0.5 mg/ml) in the presence of ≥10% glycerol, avoid multiple freeze/thaw cycles, and maintain pH >7. Also, limit exposure to high salt conditions.
Troubleshooting Purification of the proteasome is relatively straightforward, as long as the critical parameters mentioned above are respected. Problems can occur with 26S proteasome purification if the pH is not well controlled, if the complex is exposed for too long to high salt concentration, or if the protein concentration is too low for too long. A list of commonly encountered problems is presented in Table 21.5.1. Finally, it has been observed that certain mutations in the proteasome result in higher fragility of the complex, especially when exposed to elevated salt concentrations during anion exchange (see for example Glickman et al., 1998a). If the possibility exists to use a tagged version of the proteasome, try to substitute the ion-exchange steps with affinity chromatography.
Peptidases
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Table 21.5.2
Standard Purification Experiment
Purification step Clarified cell extract 50 ml DEAE Affi-Gel Blue 25 ml Resource Q 100 ml S400
mg (total)
Peptidase activity (arbitrary units)
Specific activity
Fold purification
100 67 28a 15b
0.0267 0.223 3.11 9.8
1 8 116 367
3750 300 9 1.53
aThe big drop in total peptidase activity during the anion-exchange procedure is probably due to loss of intact proteasome
holoenzyme upon exposure to high-salt conditions (see Critical Parameters). 20S particles have a lower specific activity than intact 26S proteasome holoenzyme, thus disintegration of proteasomes into 20S and 19S subcomplexes will result in a loss of total activity. Proteasome eluting from the anion exchange column should not be maintained in the high salt concentration for long periods of time; the buffer exchange step should be performed immediately. bAdditional peptidase activity is present in fractions that contain mainly 20S particles.
Anticipated Results
Literature Cited
The protocol for purifying 26S proteasome (see Basic Protocol 1) is reliable, and should lead to the purification of an easily identifiable multisubunit complex with the following earmarks when analyzed by SDS-PAGE (see Figure 21.5.2): a characteristic doublet at ∼100 kDa, a group of proteins in the 50 to 60 kDa range, and a group of proteins in the range of 20 to 35 kDa. The latter correspond to the 20S proteasome subunits. The yield of the procedure is variable, due to the lack of stability of the 26S proteasome. As an example, the purification results of a standard experiment are given in Table 21.5.2. The starting sample was 60 ml cells pelleted from 6 liters wild-type yeast grown in YPD.
Beers, E.P. and Callis, J. 1993. Utility of polyhistidine-tagged ubiquitin in the purification of ubiquitin-protein conjugates and as an affinity ligand for the purification of ubiquitin-specific hydrolases. J. Biol. Chem. 268:21645-21649.
Time Considerations As it is important to work fast (see Critical Parameters), delay must be avoided between steps. All columns must be ready and equilibrated before lysing the cells. The whole procedure (see Basic Protocol 1) should be performed in less than 2 days for best results. Running the extract on a DEAE Affi-Gel Blue column should be finished 3 hr after cell harvesting. Running the sample on a Mono-Q column or equivalent can be performed in 2 to 3 hr. The best strategy is to run the lengthy gel-filtration column during the night of the first day; however, as the concentration and buffer exchange preceding this stage is often time-consuming, another possibility is to leave the sample in the concentrator after buffer exchange overnight, and load the gel-filtration column the following day.
Braun, B.C., Glickman, M., Kraft, R., Dahlmann, B., Kloetzel, P.M., Finley, D., and Schmidt, M. 1999. The base of the proteasome regulatory particle exhibits chaperone-like activity. Nat. Cell Biol. 1:221-226. Chu-Ping, M., Vu, J.H., Proske, R.J., Slaughter, C.A., and Demartino, G.N. 1994. Identification, purification, and characterization of a high molecular weight, ATP-dependent activator (PA700) of the 20-S proteasome. J. Biol. Chem. 269:3539-3547. Ciechanover, A., Elias, S., Heller, H., and Hershko, A. 1982. “Covalent affinity” purification of ubiquitin-activating enzyme. J. Biol. Chem. 257:2537-2542. Coux, O., Tanaka, K., and Goldberg, A.L. 1996. Structure and functions of the 20S and 26S proteasomes. Annu. Rev. Biochem. 65:801-847. Enenkel, C., Lehmann, A., and Kloetzel, P.M. 1998. Subcellular distribution of proteasomes implicates a major location of protein degradation in the nuclear envelope-ER network in yeast. EMBO J. 17:6144-6154. Eytan, E., Ganoth, D., Armon, T., and Hershko, A. 1989. ATP-dependent incorporation of 20S protease into the 26S complex that degrades proteins conjugated to ubiquitin. Proc. Natl. Acad. Sci. U.S.A. 86:7751-7755. Ganoth, D., Leshinsky, E., Eytan, E., and Hershko, A. 1988. A multicomponent system that degrades proteins conjugated to ubiquitin. Resolution of factors and evidence for ATP-dependent complex formation. J. Biol. Chem. 263:1241212419.
Purification of Yeast Proteasome
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Glickman, M.H., Rubin, D.M., Coux, O., Wefes, I., Pfeifer, G., Cjeka, Z., Baumeister, W., Fried, V.A., and Finley, D. 1998a. A subcomplex of the proteasome regulatory particle required for ubiquitin-conjugate degradation and related to the COP9-signalosome and eIF3. Cell 94:615623. Glickman, M.H., Rubin, D.M., Fried, V.A., and Finley, D. 1998b. The regulatory particle of the Saccharomyces cerevisiae proteasome. Mol. Cell. Biol. 18:3149-3162. Groll, M., Ditzel, L., Lowe, J., Stock, D., Bochtler, M., Bartunik, H.D., and Huber, R. 1997. Structure of 20S proteasome from yeast at 2.4 Å resolution. Nature 386:463-471. Hershko, A. and Ciechanover, A. 1998. The ubiquitin system. Annu. Rev. Biochem. 67:425479. Hershko, A., Heller, H., Elias, S., and Ciechanover, A. 1983. Components of ubiquitin-protein ligase system. Resolution, affinity purification, and role in protein breakdown. J. Biol. Chem. 258:8206-8214. Kawahara, H. and Yokosawa, H. 1994. Intracellular calcium mobilization regulates the activity of 26 S proteasome during the metaphase-anaphase transition in the ascidian meiotic cell cycle. Dev. Biol. 166:623-633. Papa, F.R., Amerik, A.Y., and Hochstrasser, M. 1999. Interaction of the Doa4 deubiquitinating enzyme with the yeast 26S proteasome. Mol. Biol. Cell 10:741-756. Parker, C.W. 1990. Radiolabeling of proteins. Methods Enzymol. 182:721-737. Peters, J.M., Harris, J.R., and Kleinschmidt, J.A. 1991. Ultrastructure of the approximately 26S complex containing the approximately 20S cylinder particle (multicatalytic proteinase/proteasome). Eur. J. Cell Biol. 56:422-432.
Rock, K.L., Gramm, C., Rothstein, L., Clark, K., Stein, R., Dick, L., Hwang, D., and Goldberg, A.L. 1994. Inhibitors of the proteasome block the degradation of most cell proteins and the generation of peptides presented on MHC class 1 molecules. Cell 78:761-771. Rubin, D.M. and Finley, D. 1995. Proteolysis. The proteasome: A protein-degrading organelle. Curr. Biol. 5:854-858. Rubin, D.M., Coux, O., Wefes, I., Hengartner, C., Young, R.A., Goldberg, A.L., and Finley, D. 1996. Identification of the Gal4 suppressor Sug1 as a subunit of the yeast 26S proteasome. Nature 379:655-657. Seemuller, E., Lupas, A., Stock, D., Lowe, J., Huber, R., and Baumeister, W. 1995. Proteasome from thermoplasma acidophilum—a threonine protease. Science 268:579-582. Tamura, T., Tanaka, K., Tanahashi, N., and Ichihara, A. 1991. Improved method for preparation of ubiquitin-ligated lysozyme as substrate of ATPdependent proteolysis. FEBS Lett. 292:154-158. Verma, R., Chen, S., Feldman, R., Schieltz, D., Yates, J., Dohmen, J., and Deshaires, R.J. 2000. Proteasomal proteomics: Identification of nucleotide-sensitive proteasome interacting proteins by mass spectrometric analysis of affinitypurified proteasomes. Mol. Biol. Cell 11:34253439. Voges, D., Zwickl, P., and Baumeister, W. 1999. The 26S Proteasome: A molecular machine designed for controlled proteolysis. Annu. Rev. Biochem. 68:1015-1068.
Contributed by Michael Glickman The Technion Haifa, Israel Olivier Coux CRBM-CNRS Montpellier, France
Peptidases
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Purification of the Eukaryotic 20S Proteasome
UNIT 21.6
The 20S proteasome is the catalytic core of the major extralysosomal proteolytic system of the cell. Combination of the 20S proteasome with a complex of regulatory proteins forms the 26S proteasome, which in turn is responsible for the recognition and degradation of ubiquitin-protein conjugates. The constitutive form of the 20S proteasome can be conveniently purified as a stable and homogeneous preparation from bovine pituitaries. The Basic Protocol consists of an initial ammonium sulfate fractionation step followed by sequential anion-exchange, gel-filtration, and phenyl-Sepharose chromatography. An optional second gel filtration step is added at the end if necessary. A Support Protocol details an enzyme assay used in evaluating proteasomal activity. For the purposes of this discussion, “chymotrypsin-like activity” refers to the catalytic component that preferentially cleaves peptide bonds after hydrophobic amino acids. “Trypsin-like activity” refers to the catalytic component that cleaves peptide bonds after basic amino acids. Finally, “peptidylglutamyl peptide bond–hydrolyzing activity” refers to the catalytic component that cleaves peptide bonds after acidic amino acids. PURIFICATION OF THE 20S PROTEASOME FROM BOVINE PITUITARIES The bovine pituitary proteasome is quite stable and was historically the first preparation purified (Wilk and Orlowski, 1980). Many other procedures have been used to prepare the proteasome, including rapid affinity purification employing a monoclonal antibody coupled to Sepharose CL-4B (Hendil and Uerkvitz, 1991).
BASIC PROTOCOL
This protocol describes a relatively straightforward means for obtaining essentially homogeneous 20S proteasome. The purification procedure relies on anion-exchange, gel-filtration, and hydrophobic-interaction chromatography steps. Since the proteasome is a relatively abundant molecule, its purification is not particularly difficult. After each step of the purification protocol, the total volume should be measured and a small aliquot saved for assessment of enzyme activity and determination of protein concentration. One should construct a table of purification similar to Table 21.6.1. After each chromatographic step, the elution of total protein and enzyme should be graphed. One must use one’s best judgment in deciding which tubes to pool. It is usually preferable to sacrifice some yield in favor of higher purity. Results after each step should be compared to the table to confirm that there are no gross discrepancies in either yield or purity.
Table 21.6.1
Purification Scheme for Bovine Pituitaries, 200 g
Activity
Step
Vol. (ml)
Protein (mg/ml)
U/ml
U/mg
Total U
Yield (%)
Homogenate Supernatant (NH4)2SO4 DEAE-Toyopearl AcA22 Phenyl-Sepharose Second AcA22
885 1085 46 44 32 3a 10
30 12.5 30 0.76 0.52 1.6 0.38
0.29 0.27 5.9 2.81 2.81 14.1 3.84
0.01 0.021 0.20 3.70 5.4 8.83 10.0
257 293 271 123 90 42.3 38.4
100 114 105 48 35 16.5 14.9
aAfter concentration and dialysis.
Peptidases Contributed by Sherwin Wilk and Wei-Er Chen Current Protocols in Protein Science (2001) 21.6.1-21.6.9 Copyright © 2001 by John Wiley & Sons, Inc.
21.6.1 Supplement 25
NOTE: All chromatographic steps are conducted in a cold room at 4°C. Materials Frozen bovine pituitaries (Pel-Freez) 10 mM Tris-EDTA buffer, pH 7.5 (see recipe) Ammonium sulfate, ultrapure or enzyme grade Toyopearl DEAE-650M anion-exchange resin (Rohm and Haas) Eluting buffer: 300 mM Tris-EDTA buffer, pH 7.5 (see recipe) Nitrogen source Ultrogel AcA22 gel-filtration resin (BioSepra) AcA22 buffer: 10 mM Tris-EDTA buffer, pH 7.5 (see recipe), containing 0.1 M NaCl Phenyl-Sepharose 6 Fast Flow (high sub; Amersham Pharmacia Biotech) Starting buffer: 10 mM Tris-EDTA buffer, pH 7.5 (see recipe), containing 1 M ammonium sulfate Blender Refrigerated centrifuge Glass rod Dialysis tubing (16 mm diameter, 3500 MWCO; soak in distilled water and bring to boil, then decant and repeat two more times) 2.5-cm inner diameter glass chromatography column (25 cm length) for ion-exchange chromatography Peristaltic pump Amicon 50-ml ultrafiltration cell and PM 10 ultrafiltration membranes 5 × 50–cm glass chromatography column for gel-filtration chromatography 1-cm diameter glass chromatography column (25 cm length) for phenyl-Sepharose chromatography Additional reagents and equipment for dialysis (APPENDIX 3B), assaying proteasome enzymatic activity (see Support Protocol), and SDS-PAGE (UNIT 10.1) NOTE: Since it takes about 2 days to equilibrate the Ultrogel AcA22 gel filtration column (see step 23), it should be prepared before starting the purification protocol. Also be aware that the phenyl-Sepharose column (step 26) must be equilibrated overnight; therefore the experimental timetable must be arranged accordingly. Prepare the bovine pituitary homogenate 1. Weigh out ∼200 g frozen bovine pituitaries. 2. Remove excess blood by suspending the pituitaries in 500 ml cold 10 mM Tris-EDTA buffer, pH 7.5, and decanting the liquid. Repeat the wash. 3. Working in the cold room, transfer about half of the pituitaries to a blender and add 300 ml of cold 10 mM Tris-EDTA buffer, pH 7.5. 4. Homogenize at low speed for 30 sec, then let stand 15 min. Homogenize again at high speed for 1 min. Let stand for 15 min, then homogenize two more times at high speed for 1 min at 15-min intervals. Repeat with the second half of the pituitary suspension and combine the homogenates. Measure the total volume, total protein, and activity. Record values on purification table (Table 20.6.1). Save a 1-ml aliquot.
Purification of the Eukaryotic 20S Proteasome
Bovine pituitaries are difficult to homogenize. A prolonged period of homogenization will heat the sample and produce excessive foaming. Therefore, homogenization is conducted in several “pulses,” allowing 15-min intervals after each pulse for the sample to cool. The object is to produce a smooth and cold homogenate while minimizing foaming.
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Prepare the supernatant 5. Centrifuge the homogenate 30 min at 15,000 × g, 4°C. 6. Decant and save the supernatant. Transfer the pellets to the blender, add 200 ml cold 10 mM Tris-EDTA buffer, pH 7.5, and homogenize at high speed for 30 sec. 7. Centrifuge 30 min at 15,000 × g, 4°C. Combine the two supernatant fractions and measure the total volume, protein content, and activity. Record in table. Fractionate with 40% to 60% ammonium sulfate 8. Place a flask surrounded with ice and containing a magnetic stirring bar on a magnetic stirring plate. Decant the supernatant into the flask and adjust the speed of the magnetic stirrer to achieve efficient stirring without foaming. 9. Gradually add solid ammonium sulfate to 40% saturation (243 g/liter) to the cooled and stirred solution over a 30-min period. Continue stirring for an additional 30 min. The ammonium sulfate should be labeled “ultrapure” or “enzyme grade.” Lesser-quality material may contain trace amounts of heavy metals that could inactivate the enzyme. The ammonium sulfate added should be granular in consistency. Break up any lumps of ammonium sulfate before addition. It is necessary to avoid a local high concentration of ammonium sulfate, which could result in premature precipitation of the proteasome after the first addition (in this step) or in precipitation of unwanted proteins after the second addition (in step 10).
10. Centrifuge 30 min at 20,000 × g, 4°C. Remove the supernatant and measure its volume. Add ammonium sulfate to 60% saturation (132 g/liter) over a 30-min period. The volume of the solution may expand after the first addition of ammonium sulfate, and therefore it is necessary to measure the volume before the second addition.
11. Stir an additional 30 min and centrifuge 30 min at 20,000 × g, 4°C. 12. Discard the supernatant and dissolve the precipitate in a minimal amount (∼40 ml) of Tris-EDTA buffer, pH 7.5, with the aid of a glass rod. 13. Dialyze overnight against 7 liters cold 10 mM Tris-EDTA buffer, pH 7.5, using a 3500 MWCO dialysis membrane (APPENDIX 3B). Centrifuge the dialyzed enzyme for 10 min at 15,000 × g, 4°C, and discard the precipitate. Measure total volume, protein content, and activity. Record in table. Purify by anion-exchange chromatography 14. Suspend 75 ml Toyopearl DEAE-650M in 500 ml 10 mM Tris-EDTA buffer, pH 7.5, and measure the pH. Allow the ion-exchange resin to settle and decant the buffer. 15. Wash the resin three times, each time with 500 ml of 10 mM Tris-EDTA buffer, pH 7.5. Repeat if necessary until the pH of the suspension reaches 7.5. 16. Pour the exchanger into a 2.5-cm column (25 cm length) and allow the buffer to drain through the column without letting the column run dry. 17. Connect a peristaltic pump to the column with polyethylene tubing and apply the dialyzed sample at a rate of 1 ml/min. Allow the clear dialyzed enzyme solution to pass through the column, but do not let the column run dry. 18. Prepare a buffer head by carefully adding several milliliters of 10 mM Tris-EDTA buffer, pH 7.5, to the top of the column and connect the column to a buffer reservoir containing 10 mM Tris-EDTA buffer, pH 7.5. Wash the column at a flow rate of 1 ml/min until the absorbance of the effluent at 280 nm reaches 0.1. Peptidases
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19. Establish a linear gradient between 250 ml of Tris-EDTA buffer, pH 7.5 and 250 ml of eluting buffer, and elute the enzyme at a flow rate of 1 ml/min, collecting 6 ml per tube. 20. Measure the absorbance at 280 nm of every other tube against a buffer blank and assay every other tube for enzymatic activity (see Support Protocol). The enzyme begins eluting at about 0.2 M buffer.
21. Combine enzymatically active fractions and concentrate to ∼8 ml in an ultrafiltration cell under nitrogen pressure through a PM 10 membrane. Measure total volume, protein content, and activity. Record in table. Other anion exchangers such as DE-52 (Whatman) and DEAE-Sephacel (Pharmacia) can be used in place of the Toyopearl DEAE-650M. Although the Toyopearl exchanger gives better resolution, flow rates are slower. The Toyopearl manufacturer recommends packing the column under gentle pressure; however, the authors have obtained excellent results with simple gravitational packing.
Perform first Ultrogel AcA22 gel-filtration chromatography 22. Prepare a 1:1 (v/v) slurry of Ultrogel AcA22 in AcA22 buffer sufficient to pack a 5 cm × 50 cm column. Following the instructions of the manufacturer or any manual on gel-filtration chromatography, carefully pack the column. 23. Equilibrate the column with AcA22 buffer by washing with 2 liters of that buffer at a flow rate of 30 ml/hr. 24. Load the sample onto the column and elute with AcA22 buffer at the same flow rate. 25. Collect 8-ml fractions. Measure the absorbance at 280 nm of every other tube against a buffer blank and assay every other fraction for enzymatic activity (see Support Protocol). Combine enzymatically active fractions. Measure total volume of pooled active fraction, protein content, and activity. Record in table. Good separations on a gel-filtration column require care in packing. Since it takes about 2 days to equilibrate the column, it should be prepared before starting the purification protocol. The enzyme is obtained in the same buffer to be used in the next chromatography step. Therefore, there is no need for buffer exchange.
Perform phenyl-Sepharose chromatography 26. Suspend 10 ml phenyl-Sepharose 6 Fast Flow (high sub) in starting buffer and pour the suspension into a 1-cm diameter column. Wash the column overnight with the starting buffer at a flow rate of 10 ml/hr. 27. Add ammonium sulfate to the enzyme solution to a final concentration of 1 M. Load the enzyme onto the column and wash with starting buffer until the absorbance at 280 nm reaches baseline. 28. Establish a linear gradient between 125 ml of starting buffer and 125 ml 10 mM Tris-EDTA buffer, pH 7.5 (without ammonium sulfate). Elute at a flow rate of 20 ml/hr, collecting 3-ml fractions. Assay every other fraction for enzymatic activity (see Support Protocol). Examine enzymatically active fractions by SDS-PAGE (UNIT 10.1).
Purification of the Eukaryotic 20S Proteasome
In addition to other contaminating proteins, phenyl-Sepharose removes Hsp90, which copurifies with the proteasome and is quite abundant (Wagner and Margolis, 1995). The preparation at this stage is more than 95% pure. A 40-kDa contaminant elutes somewhat earlier than the proteasome. SDS-PAGE will reveal whether the final gel-filtration step is necessary, and, if so, will also indicate which samples to pool.
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29. Optional: Perform second AcA22 gel-filtration chromatography (steps 23 to 25). 30. Concentrate the enzymatically active samples as in step 21. Measure total volume, protein content, and activity. Record in table. Examine enzymatically active fractions by SDS-PAGE (UNIT 10.1). Dialyze against 10 mM Tris-EDTA buffer, pH 7.5, and store in aliquots at −80°C. ENZYME ASSAY TO ANALYZE PROTEASOME-CATALYZED CLEAVAGE The assay is based on the proteasome-catalyzed cleavage of a chromogenic group (p-nitroaniline) or a fluorogenic group (7-amino-4-methyl coumarin) from an appropriate substrate. In the method described below, liberated p-nitroaniline is converted to a red-violet colored product by a standard diazotization procedure.
SUPPORT PROTOCOL
NOTE: All incubations are conducted in a shaking incubator at 37°C. Materials 0.05 M Tris⋅Cl, pH 7.5 (APPENDIX 2E) Substrate: 10 mM N-benzyloxycarbonyl-Gly-Gly-Leu-p-nitroanilide (Z-GGL-pNA; Sigma) in DMSO Enzyme (e.g., fraction from chromatographic purification of 20S proteasome; see Basic Protocol) 10% (w/v) trichloroacetic acid (TCA) 0.1% (w/v) NaNO2 0.5% (w/v) ammonium sulfamate 0.05% N-(1-naphthyl)ethylenediamine⋅2HCl (NNEDA; see recipe) 10 mM p-nitroaniline stock in methanol (prepare fresh) Shaking incubator Tabletop centrifuge CAUTION: Wear gloves when handling trichloroacetic acid. Store the NaNO2 and NNEDA solutions in dark bottles. Store the stock substrate solutions in the freezer. 1. Mix 190 µl of 0.05 M Tris⋅Cl, pH 7.5, with 10 µl of 10 mM Z-GGL-pNA. Prepare a parallel reagent blank by mixing 10 µl of 10 mM Z-GGL-pNA substrate and 240 µl of 0.05 M Tris⋅Cl, pH 7.5. Either Z-GGL-pNA or 10 mM succinyl-Leu-Leu-Val-Tyr-7-amido-4-methylcoumarin (sucLLVY-AMC) in DMSO can be used to monitor the purification. The former substrate is preferred because it is more cost-effective.
2. Start the reaction by adding 50 µl of the enzyme solution to the first (assay) tube. Incubate both the assay tube and the reagent blank for 30 min at 37°C in a shaking incubator. 3. Stop the reaction by adding 0.25 ml of 10% TCA to each of the tubes. Centrifuge at 5000 rpm for 5 min in a tabletop centrifuge to remove precipitated protein. Centrifugation is no longer necessary when sampling after the first ion-exchange chromatographic step (see Basic Protocol, step 21).
4. From each tube, transfer 0.25 ml clear supernatant to a fresh test tube, add 0.25 ml of 0.1% NaNO2, mix, and wait 3 min. 5. Add 0.25 ml of 0.5% ammonium sulfamate, mix, and wait 2 min. Add 0.5 ml of 0.05% NNEDA and mix vigorously. Peptidases
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6. Read the absorbance of the chromogen against the blank in a spectrophotometer set at 540 nm. 7. Construct a standard curve by carrying 10 to 100 nmol p-nitroaniline through steps 4 to 6. Calculate the number of nmol product formed from this curve. If the amount of p-nitroaniline released exceeds the linear portion of the standard curve, rerun the incubation using less sample, adjusting the total volume with 0.05 M Tris⋅Cl, pH 7.5, to 250 µl. The spectrophotometric method relies on the formation of a red-violet colored chromogen formed from diazotized and NNEDA-coupled pNA. This reaction requires an acidic pH. The advantage of the spectrophotometric procedure is its simplicity. The assay can be readily scaled down to a reaction volume of 100 ìl (remember to appropriately scale down the standard curve). Proteasome activity is also commonly measured by a fluorometric procedure. In this case, the fluorogenic substrate suc-LLVY-AMC is used. The advantages of the fluorometric method are its greater sensitivity and its ability to be used in a continuous assay. For continuous assay of AMC production, the reaction mixture is monitored in a fluorometer at an excitation wavelength of 370 nm and an emission wavelength of 460 nm. For an end-point fluorometric assay, the reaction mixture can be stopped with 0.3 ml 1% SDS plus 1 ml 0.1 M sodium borate buffer, pH 9.1.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
N-(1-naphthyl)ethylenediamine⋅2 HCl (NNEDA), 0.05% in 95% ethanol Dissolve 0.5 g NNEDA (Sigma) in 50 ml H2O and dilute to 1 liter with 100% ethanol. Store in a dark bottle at room temperature. Tris-EDTA buffer (pH 7.5), 300 mM and 10 mM 300 mM: Prepare a solution of 300 mM Trizma free base (Sigma) in water. Titrate to pH 7.5 using powdered EDTA free acid. 10 mM: Dilute 300 mM Tris-EDTA buffer 1:30 in water. Use of sodium salts of EDTA is not recommended for preparing this solution, as the presence of extraneous cations may affect enzyme activity.
COMMENTARY Background Information
Purification of the Eukaryotic 20S Proteasome
The eukaryotic proteasome is a high-molecular-weight proteinase (700,000 Da) composed of four rings each containing 7 subunits. The outer rings are composed of subunits designated as α-type, and the inner rings are composed of subunits designated as β-type. The molecule is a symmetrical dimer, each half containing 7 distinct α-type and 7 distinct βtype subunits (Baumeister et al., 1998). SDSPAGE analysis of the 20S proteasome yields a series of closely spaced bands from 22 to 32 kDa (Fig. 21.6.1). The proteasome was initially described as a “multicatalytic protease complex” (Orlowski and Wilk, 1981). It contains three well-defined catalytic activities which cleave peptide bonds
after acidic, basic, and hydrophobic amino acids. The active site nucleophile is an N-terminal threonine, which is present on three distinct β-type subunits (Seemuller et al., 1995). The active sites are sequestered within an internal cavity of the molecule (Groll et al., 1997) thereby protecting the cell from uncontrolled proteolysis by this abundant proteinase. The catalytically active constitutive β-type subunits can be replaced by three γ-interferon up-regulated subunits to form an “immunoproteasome.” The immunoproteasome is believed to play a role in the generation of antigens by the MHC class I system (Tanaka et al., 1997). Proteasomes from various tissues are mixtures of constitutive and immunoproteasomes. The bovine pituitary preparation is almost exclu-
21.6.6 Supplement 25
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1
2
3
4
5
6
7
8
94 67 43
30
20
14
Figure 21.6.1 SDS-PAGE analysis of the fractions obtained in the proteasome purification protocol described in Table 21.6.1 (12.5% gel). Lanes 1 and 8, molecular mass markers in kDa; lane 2, supernatant; lane 3, after (NH4)2SO4 fractionation; lane 4, after DEAE Toyopearl chromatography; lane 5, after the first AcA22 chromatography step; lane 6, after phenyl-Sepharose chromatography; lane 7, after the second AcA22 chromatography step.
sively constitutive proteasome, whereas the preparation from bovine spleen is primarily immunoproteasome (Eleuteri et al., 1997). Since the catalytic properties of constitutive and immunoproteasome differ, studies on the bovine pituitary preparation can yield less ambiguous results than preparations from other tissues. The three catalytic activities of the proteasome have been designated as chymotrypsinlike (cleavage after a hydrophobic amino acid), trypsin-like (cleavage after a basic amino acid), and peptidylglutamylpeptide bond–hydrolyzing (PGPH; cleavage after an acidic amino acid; Wilk and Orlowski, 1983). Thus far, the major proteasome inhibitors described target the chymotrypsinlike activity. The subunits responsible for each catalytic activity have been identified (Orlowski and Wilk, 2000). Whereas mutation of the trypsinlike and PGPH active site threonines has little effect on cell growth or the degradation of ubiquitin-protein conjugates, the opposite is true when the N-terminal threonine of the subunit responsible for the chymotrypsinlike activity is mutated (Arendt and Hochstrasser, 1997). Purification of the yeast proteasome is described in UNIT 21.5.
Critical Parameters and Troubleshooting In any enzyme purification protocol, it is best to proceed efficiently from one step to the next to minimize losses due to instability or to proteolysis. Therefore, prior to the start of the procedure, all buffers should be prepared, the ion-exchangers equilibrated at the proper pH, and the gel-filtration column properly packed and washed. The initial purification procedure described in 1983 contained two sequential DEAESephacel chromatographic steps followed by a single Ultrogel filtration step (Wilk and Orlowski, 1983). This procedure was modified by Orlowski and Michaud to include a second Ultrogel chromatographic separation which served to remove trace contaminants occasionally seen in the earlier procedure. More recently, Orlowski determined that much greater resolution could be obtained when Toyopearl DEAE-650M was used in place of DEAESephacel (M. Orlowski, pers. comm.), and the authors have confirmed this. Some batches of bovine pituitaries contain relatively large amounts of a copurifying major protein migrating as 92 kDa on SDS-PAGE gels. Although this protein, which the authors of this unit identified as Hsp90 by western blotting, can be
Peptidases
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almost totally removed at the last gel filtration step, minor contamination still remains. It is of interest that Wagner and Margolis (1995) have reported that Hsp90 is a major component of the proteasome preparation from bovine lenses obtained from 1-month-old animals but is not found in preparations from 2-year-old animals. We therefore suspect that occasional lots of bovine pituitaries may contain organs obtained from young animals. Hsp90 binds very tightly to phenyl-Sepharose. We therefore modified the procedure to include a phenyl-Sepharose step, which effectively removes Hsp90 from the proteasome preparation. At this stage, the preparation is near homogeneity and should be acceptable for most studies. If trace contaminants are visible after SDS-PAGE, a final gelfiltration step will remove these. SDS-PAGE analysis after the various steps of the purification procedure is shown in Figure 21.6.1. The bovine pituitary proteasome is reversibly inhibited by low concentrations of Na+ and K+. This was the basis of our original nomenclature of the enzyme as “cation-sensitive neutral endopeptidase” (Wilk and Orlowski, 1980). Other proteasome preparations are similarly inhibited (Seol et al., 1989). A Tris-EDTA gradient, rather than an NaCl gradient, is used to elute the enzyme from the anion exchange column. At the end of the procedure, the preparation is dialyzed against 10 mM Tris-EDTA buffer, pH 7.5 to remove NaCl.
Anticipated Results Table 20.6.1 summarizes results from two representative preparations and should serve as a guide. It can be seen that ∼4 mg of purified proteasome can be obtained from 200 g of bovine pituitaries.
Time Considerations Day 1 Carry the purification protocol through the ammonium sulfate fractionation step. Dialyze the resuspended 40% to 60% ammonium sulfate precipitate overnight. Day 2 Load the sample onto the anion-exchange column, wash the column during the day, and establish the gradient for overnight elution.
Purification of the Eukaryotic 20S Proteasome
Day 3 Assay the samples from the anion-exchange column. Pool and concentrate the active frac-
tions and apply to the gel-filtration column for overnight chromatography. Day 4 Assay the samples from the gel-filtration column. Pool and concentrate the active fractions, add ammonium sulfate, apply to the phenyl-Sepharose column, and wash overnight with the starting buffer. Day 5 Elute the enzyme, assay the fractions and examine each of the active fractions by SDSPAGE. If necessary, the samples may be concentrated and chromatographed a second time by gel filtration.
Literature Cited Arendt, C.S. and Hochstrasser, M. 1997. Identification of the yeast 20S proteasome catalytic centers and subunit interactions required for active-site formation. Proc. Natl. Acad. Sci. U.S.A. 94:7156-7161. Baumeister, W., Walz, J., Zuhl, F., and Seemuller, E. 1998. The proteasome: Paradigm of a self-compartmentalizing protease. Cell 92:367-380. Eleuteri, A.M., Kohanski, R.A., Cardozo, C., and Orlowski, M. 1997. Bovine spleen multicatalytic proteinase complex (proteasome). Replacement of X, Y, and Z subunits by LMP7, LMP2, and MECL1 and changes in properties and specificity. J. Biol. Chem. 272:11824-11831. Groll, M., Ditzel, L., Lowe, J., Stock, D., Bochtler, B., Bartunik, H., and Huber, R. 1997. Structure of 20S proteasome from the yeast at 2.4 Å resolution. Nature 386:463-471. Hendil, K.B. and Uerkvitz, W. 1991. The human multicatalytic proteinase: Affinity purification using a monoclonal antibody. J. Biochem. Biophys. Methods 22:159-165. Orlowski, M. and Wilk, S. 1981. A multicatalytical protease complex that forms enkephalin and enkephalin containing peptides. Biochem. Biophys. Res. Comm. 101:814-822. Orlowski, M. and Wilk, S. 2000. Catalytic activities of the 20S proteasome, a multicatalytic proteinase complex. Arch. Biochem. Biophys. 383:116. Seemuller, E., Lupas, A., Stock, D., Lowe, J., Huber, R., and Baumeister, W. 1995. Proteasome from Thermoplasma acidophilum: A threonine protease. Science 268:579-582. Seol, J.H., Park. S.C., Ha, D.B., Chung, C.H., Tanaka, K., and Ichihara, A. 1989. Na+, K+-specific inhibition of protein and peptide hydrolyses by proteasomes from human hepatoma tissues. FEBS Lett. 247:197-200. Tanaka, K., Tanahashi, N., Tsurumi, C., Yokota, K.-Y., and Shimbara, N. 1997. Proteasomes and antigen processing. Adv. Immunol. 64:1-38.
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Wagner, B.J. and Margolis, J.W. 1995. Age-dependent association of isolated bovine lens multicatalytic proteinase complex (proteasome) with heat shock protein 90, an endogenous inhibitor. Arch. Biochem. Biophys. 323:455-462. Wilk, S. and Orlowski, M. 1980. Cation-sensitive neutral endopeptidase: Isolation and purification of the bovine pituitary enzyme. J. Neurochem. 35:1172-1182. Wilk, S. and Orlowski, M. 1983. Evidence that pituitary cation-sensitive neutral endopeptidase is a multicatalytic protease complex. J. Neurochem. 40:842-849.
Contributed by Sherwin Wilk and Wei-Er Chen Mount Sinai School of Medicine New York, New York
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Serpins (Serine Protease Inhibitors)
UNIT 21.7
Serpins are a class of proteins involved in the regulation of serine and other types of proteases (Huber and Carrell, 1989; Potempa et al., 1994; Church et al., 1997). To date, DNA and protein sequencing have identified >400 members of the serpin superfamily (Whisstock et al., 1999). Interestingly, not all serpins are of an inhibitory nature (RemoldO’Donnell, 1993). Noninhibitory serpins include ovalbumin, angiotensinogen, and pigment epithelium-derived factor. In addition to being found in humans, serpins have been found in other mammals, insects, plants, and viruses. In humans, the majority of serpins regulate the functions of proteases involved in the body’s response to injury. This includes roles in coagulation, fibrinolysis, inflammation, wound healing, and tissue repair (Huber and Carrell, 1989; Potempa et al., 1994; Church et al., 1997). Serpins have been implicated in various animal and human pathologies by the loss of a functional serpin gene through deletion or mutation, which results in a deficiency or a defect in functional protein. Examples of serpin-related pathologies include venous thromboembolic disease (antithrombin), breast cancer metastasis (maspin), emphysema (α1-protease inhibitor), and hereditary angioedema (C1 inhibitor) (Huber and Carrell, 1989; Potempa et al., 1994; Church et al., 1997). In this unit, the purification (see Basic Protocol 1) and assay (see Support Protocol) of antithrombin (AT; historically called antithrombin III) are first described. Then, protocols to determine the second-order rate constant of AT inhibition of thrombin in the absence and presence of heparin (see Basic Protocols 2 and 3) are presented. For a partial list of other serpins and their purification methods refer to Table 21.7.1. Antithrombin is a glycosaminoglycan-binding serpin that is found in human plasma at a concentration of 3 to 5 µM. The target proteases of AT include the following blood-coagulation proteases: thrombin, Factors Xa, IXa, XIa, and XIIa. CAUTION: Human blood products are a biohazard and must be handled and disposed of according to institutional guidelines. See APPENDIX 2A for general precautions. PURIFICATION OF HUMAN ANTITHROMBIN BY COLUMN CHROMATOGRAPHY
BASIC PROTOCOL 1
This purification protocol describes the isolation of human antithrombin (AT) from human plasma (Griffith et al., 1985b). After an initial polyethylene glycol (PEG) precipitation step to clear-out excess fats/lipids from the plasma, the protocol continues with barium citrate adsorption of citrated human plasma to remove vitamin K–dependent blood coagulation factors (e.g., Factors IX, X, and prothrombin). Partial purification is achieved by performing another PEG fractionation of the plasma. Finally, the PEG-fractionated plasma is subjected to heparin-Sepharose column chromatography. By utilizing the heparin affinity of AT and specific ionic strength buffers, AT can be isolated in a pure form. The final purity and enzymatic activity of the isolated AT can be assessed through SDS-PAGE and activity assays. Materials 12 to 14 U frozen citrated human plasma (available from the American Red Cross) Soybean trypsin inhibitor PEG 8000 0.5 M (v/v) benzamidine solution (store up to 1 week at 4°C) 1 M barium chloride (BaCl2; store indefinitely at 4°C) TCE buffer (see recipe)
Peptidases Contributed by Susannah J. Bauman, Herbert C. Whinna, and Frank C. Church Current Protocols in Protein Science (2001) 21.7.1-21.7.14 Copyright © 2001 by John Wiley & Sons, Inc.
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Table 21.7.1
Purification Methods for Selected Serpins
Serpin
Source
α1-Antichymotrypsin Human plasma α1-Antitrypsin
Human plasma
Antithrombin III
Human plasma
C1 inhibitor
Human plasma
CrmA
Heparin Cofactor II
Plasminogen activator inhibitor-1 Plasminogen activator inhibitor-2 Protease nexin 1
Protein C inhibitor
SERP-1
Matrices used
Reference
Blue Sepharose CL-6B; calf thymus DNA-Sepharose Glutathione Sepharose; Q Sepharose Heparin Sepharose
Chang and Lomas (1998)
DEAE cellulose; Bio-Gel A-5m; concanavalin A Sepharose Ni-NTA agarose; Recombinant DEAE Sepharose; histidine tagged protein in E. coli or Glutathione Sepharose; or recombinant GST DEAE Sephacel in line with glutathione fusion protein in Human 143 cells Sepharose Human plasma Heparin Sepharose; Q Sepharose; Sephacryl S-300; or dextran sulfate; Q-Sepharose; heparin Sepharose; Sephadex G-150 Sephacryl S-200; Recombinant protein in E. coli hydroxylapatite; heparin agarose Human placenta DEAE Sepharose; hydroxylapatite; phenyl Sepharose Human fibroblasts Dextran sulfate Sepharose; PN-1 mAb-Sepharose; or dextran sulfate Sepharose; DEAE Sepharose in line with dextran sulfate Sepharose Human plasma DEAE Sepharose; dextran sulfate-agarose; Ultrogel AcA-44; DEAE-Sephacel MonoQ; Recombinant protein in BGMK Superdex 75 cells
Lomas et al. (1993) Griffith et al. (1985a) Harpel and Cooper (1975)
Quan et al. (1995) or Ray et al. (1992)
Griffith et al. (1985a) or Tollefsen et al. (1982)
Lawrence et al. (1990) Wun and Reich (1987) VanNostrand et al. (1988) or Farrell et al. (1986)
Suzuki et al. (1983) or as modified in Pratt et al. (1989) Nash et al. (1998)
Serpins (Serine Protease Inhibitors)
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Heparin-Sepharose in 5.0-cm × 20-cm column (heparin-Sepharose can be purchased from Amersham Pharmacia Biotech or prepared using unbleached heparin and CnBr activation of Sepharose; see Griffith et al., 1985b) TCE buffer (see recipe) containing 2 M NaCl 32°C water bath 4-liter beakers, clean Magnetic stir plate and stir bars 1-liter centrifuge bottles Centrifuge, 4°C Fraction collector Additional reagents and equipment for measuring protein concentration by spectrophotometry (UNIT 3.1), SDS-PAGE (UNIT 10.1), and activity assay (see Support Protocol) Thaw plasma and perform initial PEG fractionation 1. Thaw 12 to 14 U (∼3 liters) frozen citrated human plasma in 32°C water bath and pour thawed plasma into clean 4-liter beaker. Remove plasma from bags immediately after thawing to maintain temperature <10°C.
2. To plasma, add 10 mg/liter soybean trypsin inhibitor, 33.3 g/liter PEG 8000, and 2 ml/liter of 0.5 M benzamidine solution. 3. Stir plasma solution for 30 min at 4°C. Transfer plasma solution to 1-liter centrifuge bottles. 4. Centrifuge solution 30 min at 4000 × g, 4°C, to remove insoluble material. 5. Decant supernatants into a clean 4-liter beaker and discard pellet (precipitate). Perform barium citrate adsorption 6. To supernatant, slowly add (over ∼1-2 min) 10% of the original plasma volume of 1 M BaCl2 and 2 ml/liter of 0.5 M benzamidine solution. Add both slowly and with adequate stirring to ensure quick dispersal throughout the plasma.
7. Stir solution for 30 min at 4°C. Transfer the solution to centrifuge bottles. 8. Centrifuge solution 30 min at 4000 × g, 4°C, to remove insoluble material. 9. Decant supernatants into a clean 4-liter beaker. At this point, it is possible to store the supernatant covered with plastic wrap overnight at 4°C. Also, the precipitate at this step may be further processed for isolation of vitamin K-dependent coagulation factors (see Griffith et al., 1985b).
Perform PEG fractionation 10. To supernatant, slowly add 100 g/liter PEG 8000. Stir solution 30 min at room temperature. Add both slowly and with adequate stirring to avoid clumping of PEG 8000. If this solution had been stored overnight at 4°C, it should be warmed to room temperature before adding PEG 8000.
11. Transfer solution to 1-liter centrifuge bottles and centrifuge solution 30 min at 4000 × g, 4°C, to remove insoluble material. Peptidases
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12. Carefully decant supernatant so as not to disrupt pellet. Discard supernatant and transfer pellets into a clean 4-liter beaker. 13. Resuspend pellets in 3 liters TCE buffer containing 100 g/liter PEG 8000, pH 7.4. Stir solution 60 min at room temperature to resuspend pellets. Check occasionally so that stir bar does not become stuck in the pellet.
14. Centrifuge solution 30 min at 4000 × g, 4°C, to remove insoluble material. 15. Decant supernatant into clean 4-liter beaker and discard precipitate. 16. To supernatant, add 150 g/liter PEG 8000. Stir solution for 60 min at room temperature. 17. Centrifuge solution 30 min at 4000 × g, 4°C, to obtain precipitate. Decant supernatant and discard. Purify protein by heparin-Sepharose column chromatography 18. Resuspend pellets in 3 liters TCE buffer, pH 7.4. 19. Load resuspended solution onto heparin-Sepharose column pre-equilibrated in 2 liters TCE buffer, pH 7.4. Wash column consecutively with 2 liters each TCE buffer, pH 7.4, with 0.25 M NaCl (to remove heparin cofactor II), 0.4 M NaCl (to remove fibronectin), and 0.75 M NaCl (to remove any residual trace proteins).
20. Elute AT with TCE buffer, pH 7.4, containing 2.0 M NaCl, and collect 11-ml fractions. 21. Run 3 liters TCE buffer, pH 7.4, over column to re-equilibrate. Assess isolated product 22. Measure the A280 of each fraction (see UNIT 3.1) to determine the position of the peak. 23. Run activity assay on fractions to determine specific activity (see Support Protocol). 24. Run 10% SDS-PAGE (UNIT 10.1) on activity-containing fractions to determine purity. 25. Pool appropriate fractions (see Support Protocol). Measure the A280, run activity assay, and run 10% SDS-PAGE on pooled material. As explained in the Support Protocol, the fractions to be pooled are those that contain the highest degree of thrombin-inhibition activity relative to the protein (A280) values. The pooled fractions can be dialyzed against HNPN buffer (see Support Protocol, step 9) to establish the correct buffer composition for subsequent assays.
26. Dispense into desired aliquots, label, and freeze pooled material at −20°C for later use. Aliquots can be stored for 6 to 12 months at −20°C. The usual recovery is 60% to 100% of total plasma AT. A DEAE-Sepharose (2.5-cm × 10-cm) column equilibrated in HNPN containing 0.25 M NaCl, pH 7.4, can be used to remove trace contaminating heparin that may occasionally leach off heparin-Sepharose. Under these conditions the AT does not stick to this matrix and continues through the matrix in the void volume.
Serpins (Serine Protease Inhibitors)
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Current Protocols in Protein Science
AN ASSAY FOR ANTITHROMBIN IN ELUTED FRACTIONS In this assay, fractions collected in Basic Protocol 1 are tested for antithrombin activity. This procedure uses the ability of AT to inhibit thrombin in the presence of the glycosaminoglycan heparin at a rapid rate. Those fractions that do contain AT will inhibit thrombin in the presence of heparin during the allotted incubation time. Following the incubation of AT with heparin and thrombin, a thrombin substrate (Chromozyme TH) with polybrene is added to the reaction to quantitate the remaining thrombin activity. Buffer containing heparin alone with thrombin is an important control.
SUPPORT PROTOCOL
Materials AT fractions (see Basic Protocol 1) HNPN buffer (see recipe) Thrombin/heparin solution (see recipe) Chromozyme TH/polybrene solution (see recipe) Microtiter plates coated with BSA (see recipe) Kinetic plate reader (Molecular Devices) 1. Add 2 µl of individual AT prep fractions and 48 µl of HNPN buffer into the wells of a microtiter plate coated with BSA. 2. Prepare a control sample of 50 µl HNPN buffer alone. These wells will be the thrombin alone control. They will assess 100% thrombin activity.
3. At t = 0 sec, add 50 µl of thrombin/heparin solution to each well. 4. At t = 30 sec, add 50 µl Chromozyme TH/polybrene solution to each well. 5. Place plate in kinetic plate reader and monitor absorbance change for 3 min at 405 nm. A kinetic experiment of this type is described in Lottenberg et al. (1983).
6. Compare fractions to the control of thrombin alone. 7. Express samples as percentages of thrombin control. 8. Pool samples that contain the highest degree of inhibition activity relative to protein A280 values (i.e., lowest thrombin activity remaining). 9. Dialyze the pooled samples two times against 4 liters of HNPN buffer (see Basic Protocol 2), and determine protein concentration using a specific absorption coefficient value of 0.624 ml/mg(cm) at 280 nm and an Mr of 56,600 (Church et al., 1987). AN ASSAY OF ANTITHROMBIN INHIBITION IN THE ABSENCE OF GLYCOSAMINOGLYCAN—“PROGRESSIVE ANTITHROMBIN ACTIVITY” This assay allows one to determine a second-order rate constant for AT inhibition of thrombin in the absence of glycosaminoglycan under pseudo–first order conditions. Thrombin is incubated with the AT sample for a specified time. The residual thrombin activity in the reaction is measured by the ability of the remaining thrombin to hydrolyze the substrate, Chromozyme TH. The substrate hydrolysis is subsequently quenched by the addition of a 50% acetic acid solution. By comparing the amount of substrate hydrolysis (via color development at A405) in the presence and absence of a known amount of serpin, one can calculate a second-order rate constant. NOTE: A key to the success of this assay is the accurate timing of incubations and measurements of solutions within the reaction. This includes inhibitor samples as well as thrombin-only controls.
BASIC PROTOCOL 2
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Materials HNPN/polybrene solution (see recipe) Chromozyme TH/polybrene solution (see recipe) Antithrombin (AT) solutionn (see recipe), purified and characterized (see Basic Protocol 1 and Support Protocol) HNPN buffer (see recipe for buffer) Thrombin solution (see recipe) 50% (v/v) glacial acetic acid Microtiter plates coated with BSA (see recipe) Kinetic plate reader (Molecular Devices) 1. To four wells (to be averaged) of a microtiter plate coated with BSA, add 15 µl HNPN/polybrene solution (0.1 mg/ml final) and 10 µl AT solution (100 to 200 nM final). In addition, add to two wells (to be averaged) 15 µl HNPN/polybrene solution and 10 µl HNPN solution to create a thrombin-only control (no inhibitor). 2. At t = 0 sec, add 25 µl thrombin solution to wells (1 nM final) to begin the incubation of AT and thrombin. 3. At various times from t = 10 to 90 min, add 50 µl of Chromozyme TH/polybrene solution to wells to begin the association of remaining thrombin with substrate. For a given amount of AT, one can use an approximate k2 of 1-2 × 105 M−1 min−1 to estimate a t1⁄2 for the inhibition reaction, and choose time points that bracket above and below this t1⁄2 estimate. The incubation time of thrombin with inhibitor for this assay is calculated by deriving the time necessary to obtain ∼50% inhibition of the protease with the amount of serpin being used. This is done with the equation t1⁄2 = −ln(0.5)/k2[I], where k2 is the expected second order rate constant of thrombin inhibition by AT (listed above) and [I] is the concentration of AT being used.
4. After a suitable incubation time, add 50 µl of 50% glacial acetic acid solution to wells when A405 approaches 0.1 to 0.15. The association time (reaction time of Chromozyme TH with thrombin/inhibitor solution) for this assay is determined by monitoring the A405 of the reaction. The A405 should fall on the linear portion of the hydrolysis curve of Chromozyme TH and not exceed 10% substrate hydrolysis to avoid product inhibition. This is achieved by doing several “thrombin-only” control reactions (described in step 1 above), following the change in A405, and determining the point at which the hydrolysis becomes nonlinear. Alternatively, the plate could be read continuously (“kinetically”) without adding acid, as in Support Protocol, step 5.
5. Read the A405 on a microtiter plate reader. 6. Calculate the second-order rate constant using the equation: k2 = −ln(a)/([I]t) where a is residual protease activity measured as A405 of reaction with serpin divided by A405 of reaction without serpin; [I] is the initial concentration of serpin; and t is the protease/serpin association time.
Serpins (Serine Protease Inhibitors)
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Current Protocols in Protein Science
Rate of thrombin inhibition (k2 × 108 M–1 min–1)
10
8
6
4
2
0.01
0.1
1
10
100
1000
Heparin (µg/ml)
Figure 21.7.1 The effect of heparin concentration on thrombin inhibition by purified antithrombin. The rate of thrombin inhibition by AT was measured as a function of heparin concentration, as described here, and plotted as k2 (× 108 M−1 min−1).
AN ASSAY OF ANTITHROMBIN INHIBITION IN THE PRESENCE OF GLYCOSAMINOGLYCAN—“HEPARIN COFACTOR ACTIVITY”
BASIC PROTOCOL 3
This assay measures a second-order rate constant of AT inhibition of thrombin in the presence of heparin under pseudo–first order conditions. Thrombin is incubated with the AT sample and various amounts of heparin for a specified time. The residual thrombin activity in the reaction is measured by the ability of thrombin to hydrolyze a substrate, Chromozyme TH. The substrate hydrolysis is subsequently quenched by the addition of a 50% acetic acid solution. By comparing the amount of substrate hydrolysis (via color development at A405) in the presence and absence of a known amount of serpin, one can calculate a second-order rate constant. This assay also allows one to create a “heparin template curve” by plotting the second-order rate constants versus the amount of heparin (see Figure 21.7.1). Heparin template curves permit the comparison of serpin samples in their ability to inhibit thrombin maximally and to qualitatively assess their affinities for heparin. NOTE: The key to the success of this assay is the accurate timing of incubations and measurements of solutions within the reaction. This includes inhibitor samples as well as thrombin-only controls. Materials Heparin solutions (see recipe) Antithrombin (AT) solution (see recipe), purified and characterized (see Basic Protocol 1 and Support Protocol) HNPN buffer (see recipe for buffer) Thrombin solution (see recipe) Chromozyme TH/polybrene solution (see recipe) 50% (v/v) glacial acetic acid Microtiter plates coated with BSA (see recipe) Table-top centrifuge suitable for microtiter plates Kinetic plate reader (Molecular Devices) Peptidases
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1. To four wells (to be averaged) of a microtiter plate coated with BSA, add 15 µl heparin solution and 10 µl AT solution (5 to 10 nM final). To two wells (to be averaged), add 15 µl of heparin solution and 10 µl of HNPN buffer to create a thrombin-only control (no inhibitor). For a heparin template curve, create wells with various amounts of heparin (e.g., a range containing 0.01, 0.05, 0.1, 0.2, 0.5, 1, 2, 5, 10, 20, 50, 100, 200, 500, and 1000 ìg/ml as final concentrations in the reaction).
2. At t = 0 sec, add 25 µl of thrombin solution to wells (0.5 to 1.0 nM final) to begin the incubation of AT, thrombin, and heparin. The association time of thrombin with inhibitor for this assay is “calculated” by deriving the time necessary to obtain ∼50% inhibition of the protease with the amount of serpin being used at each individual heparin concentration (that is, using different association times for each different heparin concentration). This is done with the equation t1⁄2 = −ln(0.5)/k2[I], where k2 is the expected second-order rate constant of thrombin inhibition by AT at the given heparin concentration and [I] is the concentration of AT being used. Alternatively, one could estimate an approximate t1⁄2 at the optimal heparin concentration and use the same association time throughout the assay at the various heparin concentrations.
3. At a given t (as determined above, usually 15 to 45 sec), add 50 µl Chromozyme TH/polybrene solution to wells to begin the association of remaining thrombin with substrate. The association time (reaction time of Chromozyme TH with thrombin/inhibitor solution) for this assay is determined by monitoring the A405 of the reaction. Using the conditions described (0.5 to 1.0 nM thrombin and 150 ìM Chromozyme TH; final concentrations), the A405 should not exceed 0.15, which would exceed the linear portion of the hydrolysis of the Chromozyme TH under the conditions of the assay (see Basic Protocol 2, step 4, second annotation).
4. After a suitable incubation time, add 50 µl of 50% acetic acid solution to wells when A405 approaches 0.1 to 0.15. 5. Before reading A405, centrifuge each reaction in a table-top centrifuge 10 min at 2000 × g to remove precipitated heparin. Transfer 75 µl of assay solution from each well into a new microtiter plate. The heparin/polybrene complex precipitates under acidic conditions; expect higher heparin concentrations (≥50 µg/ml).
6. Read plate at A405 on a reader. 7. Calculate the second-order rate constant using the equation: k2 = −ln(a)/([I]t) where a is the residual protease activity measured at A405 of reaction with serpin divided by A405 of reaction without serpin; [I] is the concentration of serpin; and t is the protease/serpin association time. 8. Create heparin “template curves” by plotting the second-order rate constant versus heparin concentration with the heparin concentration being displayed on a log scale (see Fig. 21.7.1).
Serpins (Serine Protease Inhibitors)
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Antithrombin solution Prepare 500 to 1000 nM human antithrombin (for final concentration in the assay wells of 10 to 200 nM) in HNPN buffer (see recipe). Make only enough of this solution for the assays to be done. This is determined by the number of samples to be tested.
Chromozyme TH/polybrene solution Make up 300 µM Chromozyme TH (150 µM final; tosyl-Gly-Pro-Arg-p-nitroanilide, Boehringer Mannheim) and 2 to 4 mg/ml polybrene (1 to 2 mg/ml final; note that in the presence of heparin the final polybrene concentration must be equal to or in excess of the heparin concentration; assays in the absence of heparin can be done with 0.1 mg/ml polybrene) in HNPN buffer (see recipe). Make only enough of this solution for the assays to be done. This is determined by the number of fractions to be tested. Store up to a few days at 4°C; the stock Chromozyme TH (50 mM) is prepared in DMSO and is stable in the dark for months at room temperature. The stock polybrene solution (200 mg/ml) is made in distilled water and is stable indefinitely at 4°C (and is used throughout these assays including the “progressive” antithrombin activity).
Heparin solutions Make up heparin solutions in a range of 0 to 3.33 mg/ml heparin (0 to 1 mg/ml final) in HNPN buffer (see recipe). Make only enough of this solution for the assays to be done. This is determined by the number of sample to be tested. Store up to 1 week at 4°C.
HNPN buffer 20 mM HEPES, pH 7.4 150 mM NaCl 0.1% PEG 8000 0.02% (w/v) NaN3 Store up to 1 year at room temperature HNPN/polybrene solution HNPN buffer (see recipe) containing 0.667 mg/µl polybrene Make only enough of this solution for the assays to be done. This is determined by the number of samples to be tested. Prepare fresh. Polybrene (hexadimethrine bromide) can be obtained from Sigma-Aldrich.
Microtiter plates coated with BSA Add 200 µl of a 2 mg/ml solution of BSA in HNPN buffer (see recipe) to each well of a 96-well U-bottom microtiter plate. Let plates coat a minimum of overnight at 4°C. Remove all of BSA/HNPN solution before using plate for assay. Store up to several weeks at 4°C. Chicken egg-white ovalbumin can be used in place of BSA to avoid possible contamination by bovine serum proteins.
Peptidases
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TCE buffer 0.05 M Tris⋅Cl, pH 7.4 (APPENDIX 2E) 0.02 M citric acid 5.0 mM EDTA 0.02% (w/v) sodium azide (NaN3) Store up to several months at 4°C Thrombin/heparin solution HNPN buffer (see recipe) containing: 1 to 2 nM human α-thrombin (0.5 to 1.0 nM final) 2.0 mg/ml BSA (optional) 1.0 µg/ml heparin (0.5 µg/ml final) Make only enough of this solution for the assays to be done. This is determined by the number of fractions to be tested. Prepare fresh. Human α-thrombin can be obtained from either Enzyme Research Laboratories or Haematologic Technologies, or it can be prepared and purified from human plasma as described by Church and Whinna (1986). Chicken egg-white ovalbumin can be used in place of BSA to avoid possible contamination by bovine serum proteins.
Thrombin solution HNPN buffer (see recipe) containing: 1 to 2 nM human α-thrombin (0.5 to 1 nM final) 2 mg/ml BSA (optional) Make only enough of this solution for the assays to be done. This is determined by the number of samples to be tested. Prepare fresh. Human α-thrombin can be obtained from either Enzyme Research Laboratories or Haematologic Technologies, or it can be prepared and purified from human plasma as described by Church and Whinna (1986). Chicken egg-white ovalbumin can be used in place of BSA to avoid possible contamination by bovine serum proteins.
COMMENTARY Background Information
Serpins (Serine Protease Inhibitors)
Serpins are a superfamily of proteins that are typically ≥400 amino acids in length. The prototypical serpin is α1-proteinase inhibitor, also known as α1-antitrypsin (Loebermann et al., 1984; Huber and Carrell, 1989). Serpins have a highly organized tertiary structure consisting of three β-pleated sheets, referred to as A-C, and nine α-helices referred to as A-I (Loebermann et al., 1984; Huber and Carrell, 1989). Serpins differ in one portion of their structure, the carboxyl portion of the protein called the reactive site loop (Carrell and Travis, 1985). This is the region of a serpin that docks to the active site of a target protease. The reactive site loop amino acids are labeled from amino to carboxyl terminus as P15-P14-P13…P3-P2P1-P1′-P2′-P3′-P4′-P5′ (Schechter and Berger, 1967). In this method of labeling, the target protease recognizes the P1-P1′ bond, also called the reactive site, as the ideal substrate.
Inhibitory serpins function to inhibit their target proteases by presenting themselves to the protease as a pseudosubstrate (Huber and Carrell, 1989; Potempa et al., 1994; Gettins and Olson, 1996; Church et al., 1997). The protease, upon trying to cleave the reactive site bond (P1-P1′), is trapped in a stable 1:1 complex with the inhibitor. The reactive site loop of a serpin is flexible and can adopt a variety of conformations unlike standard-mechanism inhibitors that retain a rigid, “canonical” conformation (Hopkins et al., 1995; Stein and Carrell, 1995). In the different conformations, the amino terminal portion of the reactive site loop is inserted to various degrees into the A β-sheet. Insertion is not necessary for canonical conformation of the reactive site loop. The residues P10-P15 of the serpin are termed the hinge region (Hopkins et al., 1993). The substitution of residues with larger side chains impedes loop insertion and decreases the rate of stable serpin/protease
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Table 21.7.2
Selected Serpins and Their Target Proteases
Serpin
Putative target proteases
α1-Antichymotrypsin α1-Antitrypsin Antithrombin III C1 Inhibitor CrmA
Cathepsin G Neutrophil elastase Thrombin, factor Xa C1s, kallikrein Interleukin-1β-converting enzyme, granzyme B Thrombin Plasminogen activators (t- and u-) Plasminogen activators (t- and u-) Thrombin, plasmin, urokinase, trypsin Activated protein C, thrombin, thrombin-thrombomodulin complex, acrosin, urokinase Plasminogen activators (t- and u-), plasmin, thrombin, factor Xa
Heparin cofactor II Plasminogen activator inhibitor-1 Plasminogen activator inhibitor-2 Protease nexin 1 Protein C inhibitor (plasminogen activator inhibitor-3) SERP-1
complex formation. Crystal structures of serpins have been shown to cleave reactive sites− fully inserted, uncleave reactive sites−partially inserted, and uncleave reactive sites−fully accessible (Loebermann et al., 1984; Huber and Carrell, 1989; Stein et al., 1990; Mottonen et al., 1992; Skinner et al., 1998). The position of the reactive site loop in a final serpin/protease complex is controversial, and in this debate between full- or partial-loop insertion, there is evidence of both sides to support their claims (Church et al., 1997). A number of serpins are able to bind glycosaminoglycans (Rosenberg and Damus, 1973; Tollefsen and Blank, 1981; Suzuki et al., 1983; Skinner et al., 1998). Some serpins, such as AT, are believed to bind proteoglycans located on endothelial cells, thus providing a large pool of available inhibitor (de Agostini et al., 1990; Cooper et al., 1996). The serpins AT, heparin cofactor II, protein C inhibitor, plasminogen activator inhibitor-1, and protease nexin1 have enhanced activities in the presence of glycosaminoglycans (Gettins and Olson, 1996; Church et al., 1997). The glycosaminoglycans act as catalysts by binding to specific regions (typically helices) within the serpins such as the A- and D-helices for AT, the D-helix for heparin cofactor II, the A+- and H-helices for protein C inhibitor, and the D-helix region for plasminogen activator inhibitor-1 and protease nexin-1 (Gettins and Olson, 1996; Church et al., 1997). By utilizing these unique structural properties of serpins, one can purify the specific
serpins using heparin-Sepharose. Additionally, one can assess their broad diversity of target proteases using purified enzymes and specific chromogenic substrates (see Table 21.7.2).
Critical Parameters The greatest source of lost protein in purification occurs, most likely, in the early stages of selective PEG precipitation; however, the majority of AT is recovered by heparin-Sepharose chromatography. Loss of activity is always a concern with protein purification, and AT solutions typically remain active for a few weeks at 4°C and for years at −20°C or −80°C. The speed of purification of AT is important, thus, working consecutive days to finish the purification procedure is recommended. Heparin-Sepharose—heparin leaching is always an issue, especially if one has prepared one’s own matrix using cyanogen bromide activation. The newer matrices from Amersham Pharmacia Biotech are excellent, notably the Hi-Trap Heparin-Sepharose, in terms of binding affinity and lack of leaching-off of the heparin. For smaller scale preparations of AT (about ≤100 ml of plasma), plasma could be directly applied to a Hi-Trap Heparin-Sepharose matrix and with appropriate wash steps and subsequent purification procedures, AT can be obtained in a relatively pure form (O’Reilly et al., 1999). Peptidases
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Troubleshooting
DEAE-Sepharose (2.5-cm × 10-cm) can be used to remove leached heparin from the first column as described above. α- versus β- forms of AT may be important to consider, in that a minor portion (∼5% to 10%) of AT termed β-AT is lacking a carbohydrate attached at Asn 135 and binds unusually well to heparin-Sepharose, eluting at a significantly higher ionic strength that the majority of AT. Timing in assays is a critical parameter to assess, in that the “rate” at which AT inhibits thrombin in the presence of heparin is remarkably fast and careful timing and assay technique is crucial for reliable inhibition rate constant measurements.
Anticipated Results The described purification of AT will allow isolation of 60% to 100% of the AT from the citrated human plasma. The isolated product should be pure when visualized on SDS-PAGE. The assay of AT activity in the presence and absence of heparin will yield second-order rate constants. Wild-type protein should have a second-order rate constant of ∼1-2 × 105 M−1 min−1 in the absence of heparin and ∼5-10 × 108 M−1 min−1 in the presence of optimal heparin concentration (1 to 5 µg/ml). A heparin “template curve” can be created by plotting the secondorder rate constant versus the heparin concentration in log scale as shown in Figure 21.7.1. The point at which the curve peaks is considered the optimal heparin concentration for inhibition. This value can be used qualitatively to compare the heparin affinity of a given sample of AT. The lower the value of optimal heparin concentration, the greater the affinity of AT for heparin.
Time Considerations
Serpins (Serine Protease Inhibitors)
The purification of AT can easily be done in the span of 2 to 3 days. There is a potential stopping point at step 9, at which point the supernatant can be stored overnight at 4°C. The purification can also be stopped after fractions are collected from the heparin-Sepharose column. Fractions can be stored overnight at 4°C for activity assays the following day. The activity assays performed in the presence or absence of heparin will take a matter of hours depending on the incubation and association times used. The time required for color development of the thrombin substrate must also be considered.
Literature Cited
Carrell, R.W. and Travis, J. 1985. α1-Antitrypsin and the serpins: Variations and countervariations. Trends Biochem. Sci. 10:20-24. Chang, W.-S.W. and Lomas, D.A. 1998. Latent α1antichymotrypsin: A molecular explanation for the inactivation of α1-antichymotrypsin in chronic bronchitis and emphysema. J. Biol. Chem. 273:3695-3701. Church, F.C. and Whinna, H.C. 1986. Rapid sulfopropyl-disk chromatographic purification of bovine and human thrombin. Anal. Biochem. 157:77-83. Church, F.C., Meade, J.B., and Pratt, C.W. 1987. Stucture-function relationships in heparin cofactor II: Spectral analysis of aromatic residues and absence of a role for sulfhydryl groups in thromb in in hibition. Arch. Biochem. Biophys. 259:331-340. Church, F.C., Cunningham, D.D., Ginsburg, D., Hoffman, M., Stone, S.R., and Tollefsen, D.M. 1997. Chemistry and biology of serpins. Adv. Exp. Med. Biol. 425:358. Cooper, S.T., Neese, L.L., DiCuccio, M.N., Liles, D.K., Hoffman, M., and Church, F.C. 1996. Vascular localization of the heparin-binding serpins antithrombin, heparin cofactor II, and protein C inhibitor. Clin. Appl. Thrombosis/Hemostasis 2:185-191. de Agostini, A.I., Watkins, S.C., Slayter, H.S., Youssoufian, H., and Rosenberg, R.D. 1990. Localization of anticoagulantly active heparin sulfate proteoglycans in vascular endothelium: Antithrombin binding on cultured endothelial cells and perfused rat aorta. J. Cell Biol. 111:12931304. Farrell, D.H., Van, N.W.E., and Cunningham, D.D. 1986. A simple two-step purification of protease nexin. Biochem. J. 237:907-912. Gettins, P. and Olson, S.T. 1996. Serpins: Structure, Function and Biology, 1st edition. R.G. Landes, Austin, Tex. Griffith, M.J., Noyes, C.M., and Church, F.C. 1985a. Reactive site peptide structural similarity between heparin cofactor II and antithrombin III. J. Biol. Chem. 260:2218-2225. Griffith, M.J., Noyes, C.M., Tyndall, J.A., and Church, F.C. 1985b. Structural evidence for leucine at the reactive site of heparin cofactor II. Biochemistry 24:6777-6782. Harpel, P.C. and Cooper, N.R. 1975. Studies on human plasma C1 inactivator-enzyme interactions. J. Clin. Invest. 55:593-604. Hopkins, P.C.R., Carrell, R.W., and Stone, S.R. 1993. Effects of mutations in the hinge region of serpins. Biochemistry 32:7650-7657. Hopkins, P.C.R., Crowther, D.C., Carrell, R.W., and Stone, S.R. 1995. Development of a novel recombinant serpin with potential antithrombotic properties. J. Biol. Chem. 270:11866-11871.
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Huber, R. and Carrell, R.W. 1989. Implications of the three-dimensional structure of α1-antitrypsin for structure and function of serpins. Biochemistry 28:8951-8966. Lawrence, D.A., Strandberg, L., Ericson, J., and Ny, T. 1990. Structure-function studies of the SERPIN plasminogen activator inhibitor type 1. J. Biol. Chem. 265:20293-20301. Loebermann, H., Tukuok, R., Deisenhofer, J., and Huber, R. 1984. Human α1-proteinase inhibitor crystal structure analysis of two crystal modifications, molecular model and preliminary analysis of the implications for function. J. Mol. Biol. 107:531-556. Lomas, D.A., Evans, D.L.I., Stone, S.R., Chang, W.-S.W., and Carrell, R.W. 1993. Effect of the Z mutation on the physical and inhibitory properties of α1-antitrypsin. Biochemistry 32:500-508. Lottenberg, R., Hall, J.A., Blinder, M., Binder, E.P., and Jackson, C.M. 1983. The action of thrombin on peptide p-nitroanilide substrates. Substrate selectivity and examination of hydrolysis under different reaction conditions. Biochim. Biophys. Acta. 742:539-557. Mottonen, J., Strand, A., Symersky, J., Sweet, R.M., Danley, D.E., Geoghegan, K.F., Gerard, R.D., and Goldsmith, E.J. 1992. Structural basis of latency in plasminogen activator inhibitor-1. Nature 355:270-273. Nash, P., Whitty, A., Handwerker, J., Macen, J., and McFadden, G. 1998. Inhibitory specificity of the anti-inflammatory myxoma virus serpin, SERP1. J. Biol. Chem. 273:20982-20991. O’Reilly, M.S., Pirie-Shepherd, S., Lane, W.S., and Folkman, J. 1999. Antiangiogenic activity of the cleaved conformation of the serpin antithrombin. Science 285:1926-1928. Potempa, J., Korzus, E., and Travis, J.T. 1994. The serpin superfamily of proteinase inhibitors: Structure, function, and regulation. J. Biol. Chem. 269:15957-15960. Pratt, C.W., Macik, B.G., and Church, F.C. 1989. Protein C inhibitor: Purification and proteinase reactivity. Thromb. Res. 53:595-602. Quan, L.T., Caputo, A., Bleakley, R.C., Pickup, D.J., and Salvesen, G.S. 1995. Granzyme B is inhibited by the cowpox virus serpin cytokine response modifier A. J. Biol. Chem. 270:1037710379. Ray, C.A., Black, R.A., Kronheim, S.R., Greenstreet, T.A., Sleath, P.R., Salvesen, G.S., and Pickup, D.J. 1992. Viral inhibition of inflammation: Cowpox virus encodes an inhibitor of the interleukin-1β converting enzyme. Cell 69:597604.
Schechter, I. and Berger, A. 1967. On the size of the active site in proteases I. Papain. Biochem. Biophys. Res. Commun. 27:157-162. Skinner, R., Chang, W.S., Jin, L., Pei, X., Huntington, J.A., Abrahams, J.P., Carrell, R.W., and Lomas, D.A. 1998. Implications for function and therapy of a 2.9 A structure of binary-complexed antithrombin. J. Mol. Biol. 283:9-14. Stein, P.E. and Carrell, R.W. 1995. What do dysfunctional serpins tell us about molecular mobility and disease? Struct. Biol. 2:96-113. Stein, P.E., Leslie, A.G.W., Finch, J.T., Turnell, W.G., Mclaughlin, P.J., and Carrell, R.W. 1990. Crystal structure of ovalbumin as a model for the reactive centre of serpins. Nature 347:99-102. Suzuki, K., Nishioka, J., and Hashimoto, S. 1983. Protein C inhibitor: Purification from human plasma and characterization. J. Biol. Chem. 258:163-168. Tollefsen, D.M. and Blank, M.K. 1981. Detection of a new heparin-dependent inhibitor of thrombin in human plasma. J. Clin. Invest. 68:589-596. Tollefsen, D.M., Majerus, D.W., and Blank, M.K. 1982. Heparin cofactor II: Purification and properties of a heparin-dependent inhibitor of thrombin in human plasma. J. Biol. Chem. 257:21622169. VanNostrand, W.E., Wagner, S.L., and Cunningham, D.D. 1988. Purification of a form of protease nexin 1 that binds heparin with a low affinity. Biochemistry 27:2172-2177. Whisstock, J.C., Irving, J.A., Bottomley, S.P., Pike, R.N., and Lesk, A.M. 1999. Serpins in the Caenorhabditis elegans g e no me. Proteins 36:31-41. Wun, T.-C. and Reich, E. 1987. An inhibitor of plasminogen activation from human placenta: Purification and characterization. J. Biol. Chem. 262:3646-3653.
Key References Church et al., 1997. Proceedings of the International Symposium on the chemistry and biology of serpins held April 13 to 16, 1996, in Chapel Hill, North Carolina. Provides overviews of numerous serpins. Gettins and Olson, 1996. Provides basic descriptions of action of various serpins. Griffith et al., 1985a. Describes the purification and subsequent assay of human antithrombin III and heparin cofactor II.
Remold-O’Donnell, E. 1993. The ovalbumin family of serpin proteins. FEBS Lett. 315:105-108. Rosenberg, R.D. and Damus, P.S. 1973. The purification and mechanism of action of human antithrombin-heparin cofactor. J. Biol. Chem. 248:6490-6505. Peptidases
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Silverman, G.A., Bird, P.I., Carrell, R.W., Church, F.C., Coughlin, P.B., Gettins, P.G., Irving, J.A., Lomas, D.A., Luke, C.J., Moyer, R.W., Pemberton, P.A., Remold-O’Donnell, E., Salvesen, G.S., Travis, J., and Whisstock, J.C. 2001. The serpins are an expanding superfamily of structurally similar but functionally diverse proteins. Evolution, mechanism of inhibition, novel functions, and a revised nomenclature. J. Biol. Chem. 276:33293-33296.
Contributed by Susannah J. Bauman, Herbert C. Whinna, and Frank C. Church The University of North Carolina at Chapel Hill Chapel Hill, North Carolina
Serpins (Serine Protease Inhibitors)
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Caspases The name caspase stands for cysteine aspartyl-specific protease (alternatively cysteine aspartyl-specific protease): i.e., cysteine proteases with a strict specificity for an aspartate residue in the P1 position (N-terminal to the scissile bond) of the substrate. The catalytic site configuration and structural fold classify them as members of clan CD, which also contains bacterial gingipains (Eichinger et al., 1999; also see Table 21.1.3). Caspases are found in the intracellular environment of all multicellular organisms so far analyzed, including the nematode Caenorhabditis elegans, Drosophila melanogaster, and mammals. The first member of this family to be described was caspase-1 (Black et al., 1989; Kostura et al., 1989) which was cloned and sequenced in 1992 (Cerretti et al., 1992; Thornberry et al., 1992). It was shown to be responsible for the production of interleukin 1β (IL-1β) from its precursor in monocytes and was originally named ICE (interleukin 1β-converting enzyme). Yuan et al. (1993) reported the sequence of a gene (CED3) essential for the propagation of apoptosis in C. elegans, and demonstrated its homology with ICE. This discovery predicted, for the first time in the field of apoptosis research, a functional activity—proteolysis—associated with an apoptotic gene, galvanizing the field and laying the ground for the rapid characterization and analysis of novel caspases. Since the discovery of ICE and Ced-3, at least 10 other caspases have been identified in humans (Fig. 21.8.1), each of them expected to play particular roles in living cells. It is clear from all the biological and biochemical studies on caspases that they are essential for one of two important biological processes: inflammatory signaling and related functions (the cytokine activator branch) and cell death (the other branch). The latter branch contains caspases that initiate the apoptotic process (initiator branch) and proteases that are responsible for the execution phase of apoptosis (execution branch). An exception to this classification is caspase-14, which does not participate in apoptosis or in cytokine activation but is rather expressed in differentiated keratinocytes; its activation is associated with terminal epidermal differentiation (Lippens et al., 2000).
UNIT 21.8 STRUCTURE, SEQUENCE HOMOLOGY AND EVOLUTIONARY RELATEDNESS OF CASPASES All caspases are synthesized as inactive precursors (zymogens) and show a similar domain organization (Fig. 21.8.2A). The ∼30 kDa catalytic domains are well conserved and contain all of the enzymatic machinery. Two other types of well-recognized domains can be found at the amino-terminal region of several caspases: CARD (caspase activation and recruitment domain; Hofmann et al., 1997) and DED (death effector domain; Chinnaiyan et al., 1995). Both domains are distantly related, and are implicated in homophilic interactions with other proteins that allow the respective caspase zymogens to be recruited to protein complexes to promote their activation. Activation of caspases containing CARD and DED domains normally leads to cleavage of such domains and release of the active catalytic portion, with the exception of caspase-9. Caspase-3, -6, and -7 have a short N-peptide (23-28 residues) that is removed during activation. Its role is not yet known but there have been suggestions that removal of the N-peptide of caspase-7 by caspase-3 is necessary for its activation by caspase-8 (Yang et al., 1998) and that the Npeptide of caspase-3 may act as an apoptotic silencer in vivo (Meergans et al., 2000). Importantly, cleavage of the N-terminal domains is not required for activation; rather, internal cleavage within the catalytic domain is usually the activating event (Fig. 21.8.2B). Cleavage within the catalytic domain results in the formation of two chains that participate in forming the catalytic and substrate-binding sites (Walker et al., 1994; Wilson et al., 1994). Furthermore, analysis of 3-D structures reveals that active caspases are generally found as dimers of catalytic domains (Fig. 21.8.3A) linked primarily through hydrophobic interactions. The crystallographic structures of caspase-1 (Walker et al., 1994; Wilson et al., 1994), caspase-3 (Rotonda et al., 1996; Mittl et al., 1997; Riedl et al., 2001), caspase-7 (Wei et al., 2000; Chai et al., 2001; Huang et al., 2001), caspase-8 (Blanchard et al., 1999; Watt et al., 1999), and caspase-9 (Renatus et al., 2001) have been solved and they all show the same basic fold (Fig. 21.8.3A). The subunits of each catalytic domain are αβα sandwiches, folded Peptidases
Contributed by Jean-Bernard Denault and Guy S. Salvesen Current Protocols in Protein Science (2001) 21.8.1-21.8.16 Copyright © 2001 by John Wiley & Sons, Inc.
21.8.1 Supplement 26
–100
0
Casp-1
100
200
300
400
amino acids
404
CARD CARD
377
Casp-5
CARD
418
Casp-13
CARD
377
Ced-3
416
CARD DED DED
DED
479
DED
479 N
277
Casp-6
N
293
Casp-7
336
N
Casp-14
executioners
Casp-3
initiators
Casp-10
435
CARD
Casp-9
Casp-8
508
CARD
Casp-2
cytokine activators
Casp-4
242 large small subunits
Figure 21.8.1 Caspase family members. Schematic representation of human caspases along with Ced-3 from C. elegans. The three groups (i.e., cytokine activators, initiators, and executioners) are based on demonstrated or presumed physiological functions. CARD domain (light gray), DED domain (hatched), and the small N-peptide of executioner caspases are also depicted in the diagram. The scale is set in accordance to caspase-1 numbering and the dotted line indicates the starting point of the large subunit (dark gray; amino acid 161 in caspase-1). Domains were delineated using the conserved domain database (CDD) and the Pfam algorithm.
Caspases
into a compact cylinder with six-stranded βsheets in the center surrounded by five helices positioned on opposing sides of the plane formed by the β-sheets. Despite the presence of extra strands in some structures, the overall arrangement is kept intact. The catalytic dimer contains monomers arranged by two-fold symmetry, with the two active sites found at opposite ends of the dimer. The cleaved C-terminus of the large subunit and the N-terminus of the small subunit are close to each other. This configuration leads to a proposal for the structure of the zymogen in which the two subunits of the active caspase molecule arise from different subunits of each zymogen by a domainswapping model (Walker et al., 1994; Wilson et al., 1994). This model implies that the zymogen conformation is extremely close to the active conformation. An even more appealing
possibility is that each subunit of the active molecule comes from the pro-enzyme and that cleavage of the inter-domain linker causes a rearrangement of subunit orientation (see Fig. 21.8.2B). The latter model has the advantage of explaining why pro-caspases are inactive, whereas the first model suggests that the catalytic site is already in an active conformation. In the absence of a 3-D structure of a caspase zymogen, biochemical data are equivocal regarding the mechanism and zymogen conformation (Gu et al., 1995). In a close view of the catalytic site of caspase-3 (Fig. 21.8.3B), the enzyme is depicted with Ac-Asp-Glu-Val-Asp-CHO bound to the active cysteine in a tetrahedral conformation. The catalytic dyad His237/Cys285 is found in the large subunit (amino acid numbering is according to caspase-1) and the S1 subsite is
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A His237 N-segment
Cys285
Arg341 small
large
Asp297
Asp
B
proteolysis
Figure 21.8.2 (A) Domain organization of caspases. All caspases possess an N-terminal segment, frequently and confusingly called a pro-domain, with a length that varies from 3 kDa (caspase-3 and -6) to 24 kDa (caspase-8 and -10). In most proteases a pro-domain signifies an activation peptide, but in caspases removal of the N-peptide does not result in activation, although its presence is required for the recruitment of several members of the family to their activation complexes. The N-peptide is followed by the large subunit (17 to 21 kDa) and a small subunit (10 to 13 kDa) which constitute the catalytic domain of the enzyme. A small linker segment (12 to 39 amino acids) separates the large and small subunits. This segment contains the cleavage sites whose proteolysis usually activates the enzyme. Residues important for catalysis (Cys285 and His237), maturation (Asp), activation (Asp297), and S1 substrate binding pocket (Arg341) are indicated. (B) Caspase activation hypothesis. The 3-D structure of the active form of several caspases has been determined, but the conformation of the zymogen is open to conjecture. The large and small subunits are represented as dark or light gray shapes joined by the linker segment (dotted line) that could adopt one of two configurations, each generating a different conformation of the zymogen. The upper model shows the caspase in the almost final conformation, needing only proteolysis to achieve the active state of the dimer, whereas the lower model implies a conformational change to form the active site (star) and, thus, to obtain the active dimer. The main difference is that, in the upper model, the subunits are swapped within the putative caspase zymogen dimer. Catalytic sites are shown as stars (a hatched star represents inactive site).
formed by side chains from both large and small subunits (Arg179, Gln283, and Arg341). Subsites S2-S4 are predominantly set by the small subunit. It is thus the variation in amino acid sequence of the small subunit that is responsible for the variable extended substrate specificity of caspases. The main chain of the substrate forms an antiparallel β-strand with the main chain of enzyme residues 339-341. Interestingly, this mode of substrate alignment is very
different from that achieved by other cysteine protease families, but is shared by the chymotrypsin family of serine proteases. Multiple sequence alignment of caspases (Fig. 21.8.4) and similarity determinations (Table 21.8.1) reveal the following features. It is clear that the N-terminal portions of caspases show no obvious homology, accounting for the various domains found in that region and the varying lengths of the amino-terminal segment.
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Figure 21.8.3 Structure of caspase-3. (A) Ribbon model of caspase-3 dimer. The large (light) and small (dark) subunit of each monomer are shown. Six sheets from each monomer are shown in a head-to-tail configuration. Sheets and strands are numbered and N- and C-terminal ends of each subunit are indicated. The active cysteine (*) is presented as a ball-and-stick model to show the catalytic sites located on the opposite side of the dimer molecule, both facing toward the reader. Derived from PDB file 1CP3 using WebLab Viewer. (B) Surface model of a close view of caspase-3 active site with Ac-Asp-Glu-Val-Asp-CHO inhibitor (P1-P4 right to left) bound to the active Cys285. His237 of the catalytic dyad, Gly238 of the oxyanionic hole, and Arg179 (partially hidden by His237), Gln283, and Arg341 that form the primary substrate pocket are shown as darker surfaces. A dotted line delineates the large and small subunits boundary and shows that binding subsites are predominantly set by the small subunit of caspase. Generated with PDB file 1PAU as above. Caspases
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On the contrary, the catalytic core of caspases is well conserved, especially around the catalytic dyad His238/Cys285. Many other residues are conserved in all caspases, including key residues for the formation of the S1 pocket and the oxyanion hole. The alignment suggests that secondary structure is conserved among all caspases, which is borne out by the existing caspase 3-D structures (caspase-1, -3, -7, -8, and -9). Cytokine activator caspases (caspase1, -4, -5, and -13) show greater similarity than any other groups. It is also noteworthy that lower sequence identity is generally found between this group and all other caspases (less than 20% in overall identity and lower that 25% in large/small subunit comparison). This low similarity implies a greater evolutionary distance, possibly accounting for the development of a nonapoptotic role of this branch of the caspases. The phylogenic tree, in which cytokine activator caspases segregate well from other caspases (Fig. 21.8.5), also illustrates this fact. Other distinct similarity groupings include the apical caspase-8 and -10, which play a similar role in receptor-stimulated caspase activation, and caspase-3 and -7, which play a combined role in the execution phase of apoptosis. Interestingly, database mining leads to the possibility that caspase-3 and -7 may have diverged from a common ancestor at a stage between bony fishes and frogs.
CATALYTIC MECHANISM AND SUBSTRATE SPECIFICITY Catalysis Caspases have a stringent specificity for an aspartate residue in the S1 substrate pocket. The S1 pocket is formed by cooperation of at least three residues (Arg179, Gln283, and Arg341) that form hydrogen bonds with the carboxylate group of the P1 aspartate. This tightly controlled environment prevents other amino acids such as glutamate from binding in the S1 site; indeed, the selectivity (kcat/Km) for Asp over Glu reaches as high as 20,000-fold for caspase3. As with other cysteine and serine proteases, the role of the catalytic site is to force the formation of the tetrahedral intermediate, and although enzyme-mechanism investigations on caspases are surprisingly lacking, the following mechanism can be proposed based on available 3-D structures (Fig. 21.8.6). Once the substrate is bound, the NH backbone of Gly238 and Cys285 forming the oxyanionic hole donates a hydrogen bond to the carbonyl oxygen, thus polariz-
ing the carbonyl group of the scissile bond. The nucleophilic Cys285 can now attack the electrophilic carbon. During or before this, the proton of the thiol group may be passed to the neighboring His237. This proton now can act as a catalytic acid by protonating the amine group of the leaving P1′ residue. During deacylation, His237 may polarize the water molecule needed to complete the hydrolysis and formation of the second tetrahedral intermediate. In many proteases, the side chain of a third residue participates in the catalysis by orienting the general base (histidine in this case). In caspases, it is the backbone of residue 177 (a proline in five caspases) that has been proposed to play that role. This hypothesis is hard to test, since interaction with the backbone cannot be assessed by mutagenesis. Caspases display maximal activity at neutral pH (6.8 to 7.2), under reducing conditions, and at the ionic strength of the cytosol (Stennicke and Salvesen, 1997; Garcia-Calvo et al., 1999). At neutral pH, the active cysteine would be polarized by the catalytic histidine. Interestingly, caspase structures argue against prepolarization of the catalytic thiol (Cys285), because the distance to His237 is too large for an ionic bond. This distance, 5.3 to 6.0 Å depending on the caspase, is much larger than the one found in unrelated cysteine proteases such as papain (3.75 Å). Moreover, the scissile bond of the substrate lies between the catalytic Cys and His side chains, which is very unusual since other protease families have their catalytic nucleophile and general base residue on the same side of the scissile bond. This suggests that His237 does not participate directly in the generation of the catalytic Cys nucleophile, but rather is important for the protonation of the α-amine of the leaving group and the generation of a nucleophilic water molecule required for deacylation. These observations suggest that caspases utilize a catalytic dyad, and not a triad. Significantly, a similar distance (5.8 Å) is found in the distant caspase homolog Arggingipain.
Substrate Specificity Although caspases share a common P1 preference for Asp, they generally also share a slight preference for His in S2, Glu in S3, and small uncharged residues in P1′ (Thornberry et al., 1997; Stennicke et al., 2000). The main specificity determinant that distinguishes the functional caspase groups from one another is the nature of the S4 pocket. The cytokine activators prefer aromatic residues (Trp or Tyr) in S4, the
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A Casp-1 Casp-4 Casp-5 Casp-13 Casp-2 Casp-14 ced3 Casp-8 Casp-10 Casp-9 Casp-3 Casp-7 Casp-6
Casp-1 Casp-4 Casp-5 Casp-13 Casp-2 Casp-14 ced3 Casp-8 Casp-10 Casp-9 Casp-3 Casp-7 Casp-6
Casp-1 Casp-4 Casp-5 Casp-13 Casp-2 Casp-14 ced3 Casp-8 Casp-10 Casp-9 Casp-3 Casp-7 Casp-6
Caspases
120 130 140 150 160 | | | | | ------------NPAMPTSSGSEGNVKLCSLEEAQRIWKQKSAEIYPIMDKSSRTRLALI ------------------------ALKLCPHEEFLRLCKERAEEIYPIKERNNRTRLALI QKITSVKPLLQIEAGPPESAESTNILKLCPREEFLRLCKKNHDEIYPIKKREDRRRLALI -----------------ESVGSAATLKLCPHEEFLKLCKERAGEIYPIKERKDRTRLALI ---------------TVEHSLDNKDGPVCLQVKPCTPEFYQTHFQLAYRLQSRPRGLALV ------------------------------------MSNPRSLEEEKYDMSGARLALILC ----------------------------------APTISRVFDEKTMYRNFSSPRGMCLI --------------------------------------SESQTLDKVYQMKSKPRGYCLI ----------------------------------VKTFLEALPRAAVYRMNRNHRGLCVI -------------------------IGSGGFGDVGALESLRGNADLAYILSMEPCGHCLI ---------------------------------------SGISLDNSYKMDYPEMGLCII -----------AKPDRSSFVPSLFSKKKKNVTMRSIKTTRDRVPTYQYNMNFEKLGKCII ----------------------------------AFYKREMFDPAEKYKMDHRRRGIALI <- hA -> <- s1 170 180 190 200 210 | | | | | ICNEEFDSIP----------RRTGAEVDITGMTMLLQNLGYSVDVKKNLTASDMTTELEA ICNTEFDHLP----------PRNGADFDITGMKELLEGLDYSVDVEENLTARDMESALRA ICNTKFDHLP----------ARNGAHYDIVGMKRLLQGLGYTVVDEKNLTARDMESVLRA ICNTEFDHMP----------PRNGAALDILGMKQLLEGLGYTVEVEEKLTARDMESVLWK LSNVHFTGE-------KELEFRSGGDVDHSTLVTLFKLLGYDVHVLCDQTAQEMQEKLQN VTK-----------------AREGSEEDLDALEHMFRQLRFESTMKRDPTAEQFQEELEK INNEHFEQMP----------TRNGTKADKDNLTNLFRCMGYTVICKDNLTGRGMLLTIRD INNHNFAKAREKVPKLHSIRDRNGTHLDAGALTTTFEELHFEIKPHDDCTVEQIYEILKI VNNHSFT----------SLKDRQGTHKDAEILSHVFQWLGFTVHIHNNVTKVEMEMVLQK INNVNFCRE-------SGLRTRTGSNIDCEKLRRRFSSLHFMVEVKGDLTAKKMVLALLE INNKNFHKS-------TGMTSRSGTDVDAANLRETFRNLKYEVRNKNDLTREEIVELMRD INNKNFDKV-------TGMGVRNGTDKDAEALFKCFRSLGFDVIVYNDCSCAKMQDLLKK FNHERFFWH-------LTLPERRRTCADRDNLTRRFSDLGFEVKCFNDLKAEELLLKIHE -> <- s2 -> <- s3 -> <s3a> <s3b><- hD ->
Figure 21.8.4 Multiple amino acids alignment of human caspases and C. elegans Ced-3. The caspase-1 numbering system is used. Identical amino acid residues are black-shadowed, conserved substitutions are gray-shadowed, and semi-conserved residues are grayed. Known (caspase-1, -8, -10, -9, -3, -7, -6) and predicted (other caspases) processing sites are underlined. Multiple alignment was performed using CLUSTALW and manual alignment was used to reflect structural features. For that particular purpose, caspase-4, -5 and -13 were compared to caspase-1, caspase-7 and -6 were compared to caspase-3, and caspase-10 was compared to caspase-8 and/or -9; caspase-2, Ced-3, and caspase-14 were aligned using CLUSTALW. (continued on next page)
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B
Casp-1 Casp-4 Casp-5 Casp-13 Casp-2 Casp-14 ced3 Casp-8 Casp-10 Casp-9 Casp-3 Casp-7 Casp-6
Casp-1 Casp-4 Casp-5 Casp-13 Casp-2 Casp-14 ced3 Casp-8 Casp-10 Casp-9 Casp-3 Casp-7 Casp-6
270 280 290 300 310 | | | | | NCPSLKDKPKVIIIQACRGDSPGVVW--FKDSVGVSGNLSLPTTEEFEDD---------NCLSLKDKPKVIIVQACRGANRGELW--VRDSPASLEVASSQSSENLEED---------NCLSLKDKPKVIIVQACRGEKHGELW--VRDSPASLAVISSQSSENLEAD---------NCLSLKDKPKVIIVQACRGANRGELW--VSDSPPALADSFSQSSENLEED---------NCPSLQNKPKMFFIQACRGDETDRGVD-QQDGKNHAGSPGCEESD--------------NCQALRAKPKVYIIQACRGEQRDPGETVGGDEIVMVIKD--------------------NAPRLANKPKIVFVQACRGERRDNGFP-VLDSVDGVPAFLRRGWDNRD--GPLFNFLGCV KCPSLAGKPKVFFIQACQGDNYQKGIPVETDSEEQPYLEMD------------------QCPRLAEKPKLFFIQACQGEEIQPSVSIEADALNPEQAPTSLQD---------------SCPSLGGKPKLFFIQACGGEQKDHGFEVASTSPEDESPGSNPEPD------ATPFQEGLR RCRSLTGKPKLFIIQACRGTELDCGI--ETDSGVDD-----------------------RCKTLLEKPKLFFIQACRGTELDDGI--QADSGPINDTD--------------------KCHSLVGKPKIFIIQACRGNQHDVPVI-PLDVVDNQTEKLDTNITEVD-----------<- s4 -> <- s5 -> <s6 320 330 340 350 360 | | | | | ------AIKKAHIEKDFIAFCSSTPDNVSWRHPTMGSVFIGRLIEHMQEYACS-CDVEEI ------AVYKTHVEKDFIAFCSSTPHNVSWRDSTMGSIFITQLITCFQKYSWC-CHLEEV ------SVCKIHEEKDFIAFCSSTPHNVSWRDRTRGSIFITELITCFQKYSCC-CHLMEI ------AVYKTHVEKDFIAFCSSTPHNVSWRDIKKGSLFITRLITCFQKYAWC-CHLEEV AGKEKLPKMRLPTRSDMICGYACLKGTAAMRNTKRGSWYIEALAQVFSERACD-MHVADM ------SPQTIPTYTDALHVYSTVEGYIAYRHDQKGSCFIQTLVDVFTKRKG---HILEL RPQVQQVWRKKPSQADILIAYATTAQYVSWRNSARGSWFIQAVCEVFSTHAKD-MDVVEL --LSSPQTRYIPDEADFLLGMATVNNCVSYRNPAEGTWYIQSLCQSLRERCPRGDDILTI ---------SIPAEADFLLGLATVPGYVSFRHVEEGSWYIQSLCNHLKKLVPRHEDILSI TFDQLDAISSLPTPSDIFVSYSTFPGFVSWRDPKSGSWYVETLDDIFEQWAHS-EDLQSL ----DMACHKIPVEADFLYAYSTAPGYYSWRNSKDGSWFIQSLCAMLKQYADK-LEFMHI ----ANPRYKIPVEADFLFAYSTVPGYYSWRSPGRGSWFVQALCSILEEHGKD-LEIMQI ----AASVYTLPAGADFLMCYSVAEGYYSHRETVNGSWYIQDLCEMLGKYGSS-LEFTEL -> <-s7> <-
370 380 390 400 | | | | Casp-1 FRKVRFSFEQPD----------GRAQMPTTERVTLTRCFYLFPGH Casp-4 FRKVQQSFETPR----------AKAQMPTIERLSMTRYFYLFPGN Casp-5 FRKVQKSFEVPQ----------AKAQMPTIERATLTRDFYLFPGN Casp-13 FRKVQQSFEKPN----------VKAQMPTVERLSMTRYFYLFPGN Casp-2 LVKVNALIKDREGYAPGTEFHRC-KEMSEYC-STLCRHLYLFPGHPPT Casp-14 LTEVTRRMAEAELVQEGK----ARKTNPEIQ-STLRKRLYLQ ced3 LTEVNKKVACGFQTSQGSNIL---KQMPEMT-SRLLKKFYFWPEARNSAV Casp-8 LTEVNYEVSNKDDKKNMG------KQMPQPT-FTLRKKLVFPSD Casp-10 LTAVNDDVSRRVDKQGT------KKQMPQPA-FTLRKKLVFPVPLDALSI Casp-9 LLRVANAVSVKGIY----------KQMPGCF-NFLRKKLFFKTS Casp-3 LTRVNRKVATEFESFSFDATFHAKKQIPCIV-SMLTKELYFYH Casp-7 LTRVNDRVARHFESQSDDPHFHEKKQIPCVV-SMLTKELYFSQ Casp-6 LTLVNRKVSQRRVDFCKDPSAIGKKQVPCFA-SMLTKKLHFFPKSN hF -> < s8 >
Figure 21.8.4 (continued from previous page) Structural determinants [helices (h) and strands (s) number] are shown below the sequence alignment. Amino acids prior to caspase-1 Gln147 residues and residues from the inter-domain Asp297-Asp326 were not aligned. Sequences used for the alignment can be retrieved from GenBank with the following accession numbers: caspase-1, P29466; caspase-4, XP_006262; caspase-5, AAA75172; caspase-13, NM_003714; caspase-2, P42575; caspase-14, AAD16173; Ced-3, A49429; caspase-8, AAC50602; caspase-10, AAC50644; caspase-9, AAC50640; caspase-6, AAC50168; caspase-3, AAA74929; caspase-7, AAC51152. Peptidases
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60:67
62:64 — 76 77 26 21 19 14 19 22 23 18 16
—
51 45 49 23 22 22 16 15 22 24 16 20
81:83 — 68 22 22 21 13 18 17 23 18 19
Casp-5
Casp-4
Casp-1 60:64 82:87 76:71 — 22 22 22 15 18 20 24 17 16
Casp-13 33:31 32:24 32:29 29:26 — 26 26 17 21 27 27 23 24
Casp-2 30:11 27:12 29:16 27:13 28:30 — 26 27 25 22 29 27 26
Casp-14
Percent Identity Between Caspases and Caspase Subunitsa
33:30 31:21 34:26 32:22 37:32 23:35 — 22 16 26 29 22 30
Ced-3 19:10 19:10 21:22 35:15 28:30 33:37 — 37 26 35 29 24
20:18
Casp-8 21:24 25:22 22:25 35:12 22:35 31:31 41:62 — 24 32 31 27
22:21
25:24 25:23 25:23 23:25 36:20 27:15 24:35 35:32 33:33 — 32 25 27
Casp-10 Casp-9
27:27 25:26 25:26 25:24 34:26 29:35 29:40 41:35 29:39 39:35 — 49 37
Casp-3
23:18 25:18 24:18 26:20 32:26 25:35 29:40 41:36 36:40 36:36 49:65 — 33
Casp-7
20:25 20:21 27:16 20:17 31:23 26:32 31:34 34:15 31:36 31:32 37:52 35:45 —
Casp-6
aSequences used are the same as for Figure 21.8.3 and data were generated by pairwise alignment using CLUSTALW. The amino acid identity (%) of the large subunit (Gln147-Asp297 caspase-1 numbering) and the small subunit (Asp326-end) are presented in boldface and normal text respectively (separated by a colon). The lower section of the table reflects overall identity between pro-caspases (entire sequences were used).
Casp-1 Casp-4 Casp-5 Casp-13 Casp-2 Casp-14 Ced-3 Casp-8 Casp-10 Casp-9 Casp-3 Casp-7 Casp-6
Table 21.8.1
Caspases
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Caspase-1 Caspase-5
cytokine activators
Caspase-4 Caspase-13 Ced3 Caspase-14 Caspase-2 Caspase-9 Caspase-8
initiators
Caspase-10 Caspase-3 Caspase-7
executioners
Caspase-6
0.1
Figure 21.8.5 Phylogenic tree of caspases. Phylogenic tree was obtained by comparing sequences from all caspases starting at Gln147 (caspase-1 numbering). The length of each branch indicates distance from node. Grouping of caspases into three functional groups (Fig. 21.8.1) is also reflected by the phylogenic tree, with the exception of caspase-2, which is closer to the cytokine activator group than the proposed initiator group, although the biological function of caspase-2 is not yet well known. C. elegans Ced-3 seems more closely related to cytokine activators group than the other groups based on sequences.
apical caspase-8 and -9 prefer hydrophobic residues such as Leu and Ile, and the executioner caspases, caspase-3 and -7, prefer Asp at this position. These preferences account in part for caspase specificity on natural substrates. For example, IL-1β is processed at a Tyr-ValHis-Asp site by caspase-1 (Cameron et al., 1985), caspase-8 and -9 process the zymogens of executioner caspase-3 and -7 at Ile-Glu-ThrAsp and Ile-Gln-Ala-Asp, respectively, and caspase-3 and caspase-7 themselves process the death substrates, such as poly(ADP-ribose) polymerase, which is involved in DNA repair (Lazebnik et al., 1994), DFF45/ICAD, which regulates DNA fragmentation (Liu et al., 1997), and the 70-kDa protein component of the U1 small nuclear ribonucleoprotein, which is involved in mRNA splicing (Casciola-Rosen et
al., 1994) at Asp-Xaa-Xaa-Asp sites. Some complexity in understanding the role of presumptive apoptotic caspases is revealed by the preference of caspase-2 for Asp-Xaa-Xaa-Asp (executioner-type) and caspase-6 for Val-XaaXaa-Asp (initiator-type) motifs (Thornberry et al., 1997). These groupings are in contrast to the expected position of caspase-2 as an apical caspase (Duan and Dixit, 1997), and caspase-6 as an executioner (Orth et al., 1996; Faleiro et al., 1997). Using 2-D gel electrophoresis, it has been shown that a limited set of proteins are cleaved during apoptosis (∼100; Brockstedt et al., 1998; Otto et al., 1998). About fifty of them have been identified so far, but only a few have been directly demonstrated to be implicated in apoptosis. The vast majority may be either cleaved
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oxyanionic hole formed by backbone Cys285 amino acid Gly238 amides
1
tetrahedral intermediate
O
O
2
3 R Cys285
C
S
R N H
Cys285
R′
S
+ H N
C N H
R′
+H N His237
His237
NH
NH
free enzyme
R
COOH
5
O
R Cys285
C
H2N
4
R′
S O
H
H N + His237 NH
Figure 21.8.6 Catalytic mechanism of caspases. The role of the catalytic site of a protease is to force the formation of the tetrahedral intermediate. Once bound to the S1-S4 pocket, the scissile bond is close to the active Cys285, which is probably prepolarized by the neighboring His237 at neutral pH. The oxyanionic hole, formed by Gly238 and Cys285 backbones, causes the carbonyl group of the peptide bond to be electrophilic, and thus susceptible to an attack by the free thiol group of the active Cys285 (step 1). This step leads to the formation of the first tetrahedral intermediate (step 2). This step is reversible since no bond is yet broken. The other possible path leads to the formation of the acyl-enzyme moiety (step 3) with loss of the C-terminal part of the substrate (P′ residues). Finally, a water molecule polarized by His237 can attack the acyl-enzyme carbon (step 4) leading to the second tetrahedral intermediate (black arrows) and deacylation of the enzyme (step 5, dotted arrows) to the free enzyme state ready for a new round of catalysis (grayed portion). “R” represents the N-terminal portion of the substrate whereas “R′” is the C-terminal segment.
Caspases
as part of the general breakdown of the cell or as a “post-mortem” proteolysis. Most of the time this is attributed to caspase-3. The reason for this is two-fold. First, caspase-3 is the most abundant caspase in cells (around 200 nM). Secondly, it has a kcat/KM for an Asp-Glu-ValAsp containing internally quenched fluoro-
genic substrate which is at least six times higher than its closest relative, caspase-7 (Stennicke et al., 2000), which is present at a 5-fold lower concentration (Stennicke et al., 1999). This constitutes the main obstacle to the in vivo study of caspase functions, since the activity of
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other caspases is frequently overpowered by caspase-3 activity.
ACTIVATION AND INHIBITION OF CASPASES Activation A linker segment containing conserved aspartate residues separates the large and small subunits of the caspase catalytic domain. Though not obligatory (Zhou and Salvesen, 1997), processing at these residues within the linker is the general process of caspase activation. A significant exception to this rule is found with caspase-9, which is not activated by proteolysis, but rather by recruitment to its cofactor Apaf-1 (Rodriguez and Lazebnik, 1999; Stennicke et al., 1999). However, all other known caspases seem to require proteolysis in the linker segment for activation, and two mechanisms have been proposed to explain this process: auto-activation and trans-activation. Upon ligation of a death receptor (Fas or TNF-R1, extrinsic pathway of apoptosis activation), the apical caspase-8, by virtue of its DEDs, is recruited in an oligomeric assembly with adapter proteins (FADD for instance) to the cytosolic face of the cell membrane, inducing the formation of a death-inducing signaling complex (DISC; Muzio et al., 1996). This clustering results in pro-caspase activation via a mechanism known as the induced proximity model (Salvesen and Dixit, 1999), which is related to the loose constraint placed on the caspase-8 zymogen (Muzio et al., 1998). Pro-caspase-8 displays limited enzymatic activity (1%) of the mature, fully processed protease, and it is said to have a zymogenicity of 100. This degree of activity is much higher than other protease zymogens. The induced proximity model states that this activity is sufficient to induce auto-activation when the local concentration of zymogen is very high, as it would be the case at the cytosolic face of the DISC, for example. Once activated, caspase-8 can process and activate executioner caspase-3 and -7. It is interesting to note that in the induced proximity hypothesis, caspases downstream of initiator caspases should have high zymogenicities (constrained zymogens), since they will be activated by trans-activation. In fact, pro-caspase3 and -7 have zymogenicities of >10,000 and are extremely well constrained (Stennicke and Salvesen, 2000b). Though trans-activation of executioner caspase zymogens by apical caspases is now a well-established way to activate apoptosis, this process was originally iden-
tified by examining the function of the granzyme B, a serine protease that has specificity for a P1 aspartate (Odake et al., 1991). Granzyme B is responsible for cytotoxic lymphocyte-induced apoptosis of infected cells, and operates primarily by activating the executioner caspase zymogens (Darmon et al., 1996; Martin et al., 1996; Quan et al., 1996). This is a beautiful example of convergent evolution where a completely unrelated fold has adapted to perform an enzymatic activity shared by apical caspases.
Inhibition Nonspecific caspase inhibitors include cysteine-modifying reagents (Cerretti et al., 1992; Thornberry et al., 1992), and some metal cations such as zinc (Stennicke and Salvesen, 1997). More specific inhibitors have been designed based on short peptides adapting to the specificity pockets listed earlier. These inhibitors generally have a reactive group such as chloro- or fluoro-methylketone or acyloxymethylketone replacing the P1′ residue. Tetrapeptide aldehydes are also commonly employed, though they tend to be slow-reacting. Free side chains are required to react with caspase specificity sites, but for in vivo or in cellulo usage, compounds with protected aspartate/glutamate residues (O-methylation, for example) should be employed. Unfortunately, none of the current commercial inhibitors are specific enough to enable dissection of individual caspase pathways in vivo (GarciaCalvo et al., 1998). Natural caspase inhibitors comprise members of three unrelated families, cowpox virus CrmA, baculovirus p35, and the IAPs (inhibitors of apoptosis proteins). Of these, only the IAPs are endogenous to animals, since the other two are virally encoded. To date seven different IAPs have been identified in humans, and with the genome sequencing project completed this is likely to represent the total number. IAPs are a family of endogenous proteins, several of which possess inhibitory activity towards caspases. They inhibit caspases in a 1:1 stochiometric ratio by forming a tight complex with the protease. One of them, XIAP (Xlinked IAP), is formed of 3 consecutive BIR domains, followed by a RING domain implicated in ubiquitin-regulated degradation. Recent experiments have demonstrated that the BIR2 domain inhibits caspase-3 and -7 with a Ki of 2 to 5 nM (Riedl et al., 2001), whereas the BIR3 domain inhibits caspase-9 with Ki of 50 nM (Sun et al., 2000). The crystal structure of
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caspase-7 with the BIR2 domain of XIAP has revealed that the inhibitor binds in reverse orientation with respect to the one adopted by a substrate and blocks access to the substrate binding cleft (Chai et al., 2001; Huang et al., 2001; Riedl et al., 2001). Induction of apoptosis would then be a process in which caspase activity is raised over a certain threshold set by IAPs. Although higher organisms have some need to be able to suppress undesired apoptosis, this is even more important in the case of viruses, which invade the host to propagate. Because viruses induce an immune response, infected cells become targets for immune-induced apoptosis by cytotoxic lymphocytes. For example, cowpox virus CrmA, a member of the serpin family, rapidly and efficiently inhibits apical caspase-8 and the pro-inflammatory caspase-1 with inhibition rates in the 106 to 107 M−1 sec−1 range (Zhou et al., 1997). The p35 protein from baculovirus is a caspase-specific and potent inhibitor of the executioner caspases, caspase-1 and apical caspase-8, -10, and -12 with inhibition rates in the range of 105 M−1 sec−1 (Zhou et al., 1998). Both of these caspase inhibitors are essential for viral infectivity, since their deletion renders the respective viruses nonvirulent, emphasizing the importance of caspase pathways in antiviral surveillance.
EXPRESSION, PURIFICATION AND CHARACTERIZATION OF CASPASES
Caspases
Expression of multiple caspases with overlapping substrate specificity renders purification from tissue and cells a tedious process. Furthermore, the high expression level of caspase-3 and its high enzymatic activity would contaminate any other purification procedure. The establishment of prokaryotic expression systems is an effective and rapid method to express and purify caspases. Since caspase expression is generally toxic to E. coli, inducible repressed expression is generally required. Caspase cDNAs may be cloned into a vector in fusion to a C-terminal poly-histidine tag, with expression driven by the T7 promoter (pET expression system from Novagen). E. coli strains such as BL21(DE3), expressing the T7 RNA polymerase under the IPTG (isopropylβ-D-thiogalactopyranoside)–inducible pTac promoter, are generally used (Stennicke and Salvesen, 1999). Expression conditions vary depending on whether one wants to obtain the zymogen or the active form of the enzyme, but
some general rules apply. Bacteria are generally grown from a diluted overnight culture in 2× YT medium with appropriate antibiotics at 37°C and induced with IPTG while expression is carried out at 30°C. Expression time varies for each caspase and depends on which form is to be purified. For example, induction time of 2 hr may give the zymogen form, but longer expression (4 to 6 hr) results in fully active enzyme with very little or no pro-enzyme left. Even shorter periods of induction (30 min) are sometimes necessary to yield the zymogen form with intact N-terminal peptide, as is the case for caspase-3 and -7. Purification is carried out using zinc immobilized on a chelating resin. Cells are harvested and lysed by sonication in a Tris buffer containing sodium chloride. Cleared lysate is loaded on a column, washed, and eluted with an imidazole gradient. This simple protocol results in >95% pure enzyme, and, generally, >1 mg of protein is obtained from a liter of medium. An alternative expression procedure for some caspases utilizes refolding of individually expressed large and small catalytic subunits, followed by affinity purification on caspasespecific tetrapeptide aldehyde chromatography (Garcia-Calvo et al., 1999). Two types of substrate are routinely used to assay caspases. Chromogenic peptides with a para-nitroanilide moiety are frequently used to assay caspases when a high concentration of enzyme is used—e.g., for enzyme titration and activation analysis (Thornberry, 1993; Stennicke and Salvesen, 2000a). More sensitive assays are conducted with the same peptides coupled to the fluorogenic 7-amido-4-trifluoromethyl coumarin (Afc) or 7-amido-4methyl coumarin (Amc) group (Thornberry, 1993; Stennicke and Salvesen, 2000a). In both cases, the reporter group is attached to the C-terminal of a short tetrapeptide that reflects the caspase’s specificity. Although none of the substrates are selective enough to allow quantitation in complex mixtures, purified caspases may be quantitated as follows. Ac-Asp-GluVal-Asp-Afc is the substrate of choice for executioner caspase-3 and -7, Ac-Leu-Glu-HisAsp-Afc is used for caspase-9, Ac-Ile-Glu-ThrAsp -Af c f or casp ase-8 and -10, and Ac-Tyr-Val-Ala-Asp-Afc for caspase-1, -4, and -5 (Thornberry et al., 1997). Since the minimal requirement for caspases is the P1 Asp, DEVD substrates can be substituted for others, and enzyme concentration can be adjusted to obtain adequate readout of the assay (Stennicke and Salvesen, 2000a).
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CASPASES AND DISEASE STATE Mammals contain two biologically distinct subfamilies of caspases, one of which participates in the processing of pro-inflammatory cytokines whereas the other is required to initiate and execute the apoptotic response during programmed cell death (Fig. 21.8.1). Since the normal healthy adult human produces close to 1011 to 1012 cells per day, the body can be considered to be in a rapid state of proliferation. This proliferation is balanced by programmed cell death to maintain a constant cell number. Terminally differentiated cells are relatively resistant to apoptosis; however, proliferating cells need to be more sensitive to apoptosis to prevent overgrowth. Too much apoptosis results in degenerative disease of post-mitotic cells, and too little results in oncogenesis. Inappropriate increases in cell death have been reported in AIDS, neurodegenerative disorders, and ischemic injury, whereas decreases in cell death contribute to cancer, autoimmune diseases, and restenosis. Deregulation of cell proliferation and resistance to apoptosis are hallmarks of all cancerous cells. To acquire both phenotypes (and the underlying genotypes), a cell needs to undergo a set of genetic alterations and deregulation events. One class of growth-deregulating mutations affects signaling proteins, such as Ras, which is necessary to survival pathways. Another may involve the major late-G1 cell-cycle checkpoint controlled by pRB (Harbour and Dean, 2000) and its regulators (Sherr, 1996) and/or the over-expression of Myc (Baudino and Cleveland, 2001). Since Myc deregulation is a strong inducer of apoptosis via the intrinsic pathway (mitochondrion), a concomitant increase in cell death resistance is necessary. For example, up-regulation or over-expression of one of the two anti-apoptotic proteins Bcl-2 or Bcl-XL is able to block Myc-induced apoptosis (Fanidi et al., 1992). Many stress signals (DNA damage, telomere erosion, cell-cycle deregulation, hypoxia, loss of support, or chemotherapy) encountered during tumor progression activate p53 and result in p53-dependent apoptosis, identifying this protein as a major obstacle to tumor cells (Evan and Vousden, 2001). It is thus not surprising that more than 50% of all tumor cell lines have altered p53 gene expression or activity (Hollstein et al., 1991, 1996). Resistance to apoptosis generally involves many changes, the vast majority of which are found upstream of caspases (p53 mutation for instance). For example, mutation or deletion of the caspase-9 activator Apaf-1 is
a relatively frequent event in malignant melanoma (Soengas et al., 2001). Gene ablation experiments in mice have established that ablation of any of the major caspase genes (caspase-3, -8, and -9) is lethal, suggesting that inherited mutation of these genes is an unlikely probability (Evan and Vousden, 2001) and such mutations must be acquired by tumor cells. Little is known about mutation of caspases leading to disease. The only example to date is a nonsense mutation of caspase-10 gene that is associated with the nonmalignant auto-immune lymphoproliferative syndrome/ALPS type II (Van Der Werff Ten Bosch et al., 2001). The deletion of exon 3 of the caspase-3 gene in the MCF7 breast carcinoma cell line (Janicke et al., 1998) renders these cells more resistant to a variety of apoptotic stimuli, concomitant with the central role of this caspase in the execution phase of the cell death program. The next few years will undoubtedly reveal many as yet unknown pathological implications of these proteases, and will certainly identify these proteins as major therapeutic targets.
LITERATURE CITED Baudino, T.A. and Cleveland, J.L. 2001. The Max network gone mad. Mol. Cell. Biol. 21:691-702. Black, R.A., Kronheim, S.R., Merriam, J.E., March, C.J., and Hopp, T.P. 1989. A pre-aspartate-specific protease from human leukocytes that cleaves pro-interleukin-1b. J. Biol. Chem. 264:5323-5326. Blanchard, H., Kodandapani, L., Mittl, P.R.E., Di Marco, S., Krebs, J.F., Wu, J.C., Tomaselli, K.J., and Grütter, M.G. 1999. The three-dimensional structure of caspase-8: An initiator enzyme in apoptosis. Structure 27:1125-1133. Brockstedt, E., Rickers, A., Kostka, S., Laubersheimer, A., Dorken, B., Wittmann-Liebold, B., Bommert, K., and Otto, A. 1998. Identification of apoptosis-associated proteins in a human Burkitt lymphoma cell line: Cleavage of heterogeneous nuclear ribonucleoprotein A1 by caspase 3. J. Biol. Chem. 273:28057-28064. Cameron, P., Limjuco, G., Rodkey, J., Bennett, C., and Schmidt, J.A. 1985. Amino acid sequence analysis of human interleukin 1 (IL-1): Evidence for biochemically distinct forms of IL-1. J. Exp. Med. 162:790-801. Casciola-Rosen, L.A., Miller, D.K., Anhalt, G.J., and Rosen, A. 1994. Specific cleavage of the 70-kDa protein component of the U1 small nuclear ribonucleoprotein is a characteristic biochemical feature of apoptotic cell death. J. Biol. Chem. 269:30757-30760.
Peptidases
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Cerretti, D.P., Kozlosky, C.J., Mosley, B., Nelson, N., Van Ness, K., Greenstreet, T.A., March, C.J., Kronheim, S.R., Druck, T., Cannizzaro, L.A., Huebner, K., and Black, R.A. 1992. Molecular cloning of the interleukin-1b converting enzyme. Science 256:97-100. Chai, J., Shiozaki, E., Srinivasula, S.M., Wu, Q., Dataa, P., Alnemri, E.S., and Yigong Shi, Y. 2001. Structural basis of caspase-7 inhibition by XIAP. Cell 104:769-780. Chinnaiyan, A.M., O’Rourke, K., Tewari, M., and Dixit, V.M. 1995. FADD, a novel death domain– containing protein, interacts with the death domain of Fas and initiates apoptosis. Cell 81:505512. Darmon, A.J., Ley, T.J., Nicholson, D.W., and Bleackley, R.C. 1996. Cleavage of CPP32 by granzyme B represents a critical role for granzyme B in the induction of target cell DNA fragmentation. J. Biol. Chem. 271:21709-21712. Duan, H. and Dixit, V.M. 1997. RAIDD is a new “death” adaptor molecule. Nature 385:86-89. Eichinger, A., Beisel, H.G., Jacob, U., Huber, R., Medrano, F.J., Banbula, A., Potempa, J., Travis, J., and Bode, W. 1999. Crystal structure of gingipain R: An Arg-specific bacterial cysteine proteinase with a caspase-like fold. EMBO J. 18:5453-5462. Evan, G.I. and Vousden, K.H. 2001. Proliferation, cell cycle and apoptosis in cancer. Nature 411:342-348. Faleiro, L., Kobayashi, R., Fearnhead, H., and Lazebnik, Y. 1997. Multiple species of CPP32 and Mch2 are the major active caspases present in apoptotic cells. EMBO J. 16:2271-2281. Fanidi, A., Harrington, E.A., and Evan, G.I. 1992. Cooperative interaction between c-myc and bcl2 proto-oncogenes. Nature 359:554-556. Garcia-Calvo, M., Peterson, E.P., Leiting, B., Ruel, R., Nicholson, D.W., and Thornberry, N.A. 1998. Inhibition of human caspases by peptidebased and macromolecular inhibitors. J. Biol. Chem. 273:32608-32613. Garcia-Calvo, M., Peterson, E.P., Rasper, D.M., Vaillancourt, J.P., Zamboni, R., Nicholson, D.W., and Thornberry, N.A. 1999. Purification and catalytic properties of human caspase family members. Cell Death Differ. 6:362-369. Gu, Y., Wu, J.W., Faucheu, C., Lalanne, J.L., Diu, A., Livingston, D.J., and Su, M.S.S. 1995. Interleukin-1-beta converting enzyme requires oligomerization for activity of processed forms in vivo. EMBO J. 14:1923-1931. Harbour, J.W. and Dean, D.C. 2000. The Rb/E2F pathway: Expanding roles and emerging paradigms. Genes Dev. 14:2393-2409. Hofmann, K., Bucher, P., and Tschopp, J. 1997. The CARD domain: A new apoptotic signalling motif. Trends Biochem. Sci. 22:155-156.
Caspases
Hollstein, M., Sidransky, D., Vogelstein, B., and Harris, C.C. 1991. p53 mutations in human cancers. Science 253:49-53.
Hollstein, M., Shomer, B., Greenblatt, M., Soussi, T., Hovig, E., Montesano, R., and Harris, C.C. 1996. Somatic point mutations in the p53 gene of human tumors and cell lines: Updated compilation. Nucl. Acids Res. 24:141-146. Huang, Y., Park, Y.C., Rich, R.L., Segal, D., Myszka, D.G., and Wu, H. 2001. Structural basis of caspase inhibition by XIAP: Differential roles of the linker versus the BIR domain. Cell 104:781790. Janicke, R.U., Sprengart, M.L., Wati, M.R., and Porter, A.G. 1998. Caspase-3 is required for DNA fragmentation and morphological changes. J. Biol. Chem. 273:9357-9360. Kostura, M.J., Tocci, M.J., Limjuco, G., Chin, J., Cameron, P., Hillman, A.G., Chartrain, N.A., and Schmidt, J.A. 1989. Identification of a monocyte specific pre-interleukin 1b convertase activity. Proc. Natl. Acad. Sci. U.S.A. 86:5227-5231. Lazebnik, Y.A., Kaufmann, S.H., Desnoyers, S., Poirier, G.G., and Earnshaw, W.C. 1994. Cleavage of poly(ADP-ribose) polymerase by a proteinase with properties like ICE. Nature 371:346347. Lippens, S., Kockx, M., Knaapen, M., Mortier, L., Polakowska, R., Verheyen, A., Garmyn, M., Zwijsen, A., Formstecher, P., Huylebroeck, D., Vandenabeele, P., and Declercq, W. 2000. Epidermal differentiation does not involve the pro-apoptotic executioner caspases, but is associated with caspase-14 induction and processing. Cell Death Differ. 7:1218-1224. Liu, X., Zou, H., Slaughter, C., and Wang, X. 1997. DFF, a heterodimeric protein that functions downstream of caspase-3 to trigger DNA fragmentation during apoptosis. Cell 89:175-184. Martin, S.J., Amarante-Mendez, G.P., Shi, L., Chuang, T.-S., Casiano, C.A., O’Brien, G.A., Fitzgerald, P., Tan, E.M., Bokoch, G.M., Greenberg, A.H., and Green, D.R. 1996. The cytotoxic cell protease granzyme B initiates apoptosis in a cell-free system by proteolytic processing and activation of the ICE,Ced3 family protease, CPP32, via a novel two-step mechanism. EMBO J. 15:2407-2416. Meergans, T., Hildebrandt, A.K., Horak, D., Haenisch, C., and Wendel, A. 2000. The short prodomain influences caspase-3 activation in HeLa cells. Biochem. J. 349:135-140. Mittl, P.R., Di Marco, S., Krebs, J.F., Bai, X., Karanewsky, D.S., Priestle, J.P., Tomaselli, K.J., and Grutter, M.G. 1997. Structure of recombinant human CPP32 in complex with the tetrapeptide acetyl-Asp-Val-Ala-Asp fluoromethyl ketone. J. Biol. Chem. 272:6539-6547. Muzio, M., Chinnaiyan, A.M., Kischkel, F.C., O’Rourke, K., Shevchenko, A., Ni, J., Scaffidi, C., Bretz, J.D., Zhang, M., Gentz, R., Mann, M., Krammer, P.H., Peter, M.E., and Dixit, V.M. 1996. FLICE, a novel FADD-homologous ICE/CED-3-like protease, is recruited to the CD95 (Fas/APO-1) death-inducing signaling complex. Cell 85:817-827.
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Muzio, M., Stockwell, B.R., Stennicke, H.R., Salvesen, G.S., and Dixit, V.M. 1998. An induced proximity model for caspase-8 activation. J. Biol. Chem. 273:2926-2930. Odake, S., Kam, C.M., Narasimhan, L., Poe, M., Blake, J.T., Krahenbuhl, O., Tschopp, J., and Powers, J.C. 1991. Human and murine cytotoxic T lymphocyte serine proteases: Subsite mapping with peptide thioester substrates and inhibition of enzyme activity and cytolysis by isocoumarins. Biochemistry 30:2217-2227. Orth, K., Chinnaiyan, A.M., Garg, M., Froelich, C.J., and Dixit, V.M. 1996. The CED-3/ICE-like protease Mch2 is activated during apoptosis and cleaves the death substrate lamin A. J. Biol. Chem. 271:16443-16446. Otto, A., Muller, E.C., Brockstedt, E., Schumann, M., Rickers, A., Bommert, K., and WittmannLiebold, B. 1998. High performance two dimensional gel electrophoresis and nanoelectrospray mass spectrometry as powerful tool to study apoptosis-associated processes in a Burkitt lymphoma cell line. J. Protein Chem. 17:564-565. Quan, L.T., Tewari, M., O’Rourke, K., Dixit, V., Snipas, S.J., Poirier, G.G., Ray, C., Pickup, D.J., and Salvesen, G. 1996. Proteolytic activation of the cell death protease Yama/CPP32 by granzyme B. Proc. Natl. Acad. Sci. U.S.A. 93:19721976. Renatus, M., Stennicke, H.R., Scott, F.L., Liddington, R.C., and Salvesen, G.S. 2001. A novel self-priming mechanism drives the activation of caspase 9. Submitted for publication. Riedl, S.J., Renatus, M., Schwarzenbacher, R., Zhou, Q., Sun, S., Fesik, S.W., Liddington, R.C., and Salvesen, G.S. 2001. Structural basis for the inhibition of caspase-3 by XIAP. Cell 104:791800. Rodriguez, J. and Lazebnik, Y. 1999. Caspase-9 and APAF-1 form an active holoenzyme. Genes Dev. 13:3179-3184. Rotonda, J., Nicholson, D.W., Fazil, K.M., Gallant, M., Gareau, Y., Labelle, M., Peterson, E.P., Rasper, D.M., Tuel, R., Vaillancourt, J.P., Thornberry, N.A., and Becher, J.W. 1996. The threedimensional structure of apopain/CPP32, a key mediator of apoptosis. Nature Struct. Biol. 3:619-625. Salvesen, G.S. and Dixit, V.M. 1999. Caspase activation: The induced-proximity model. Proc. Natl. Acad. Sci. U.S.A. 96:10964-10967. Sherr, C.J. 1996. Cancer cell cycles. Science 274:1672-1677. Soengas, M.S., Capodieci, P., Polsky, D., Mora, J., Esteller, M., Opitz-Araya, X., McCombie, R., Herman, J.G., Gerald, W.L., Lazebnik, Y.A., Cordon-Cardo, C., and Lowe, S.W. 2001. Inactivation of the apoptosis effector Apaf-1 in malignant melanoma. Nature 409:207-211. Stennicke, H.R. and Salvesen, G.S. 1997. Biochemical characteristics of caspases-3, -6, -7, and -8. J. Biol. Chem. 272:25719-25723.
Stennicke, H.R. and Salvesen, G.S. 1999. Caspases: Preparation and characterization. Methods 17:313-319. Stennicke, H.R. and Salvesen, G.S. 2000a. Caspase assays. Methods Enzymol. 322:91-100. Stennicke, H.R. and Salvesen, G.S. 2000b. Caspases: Controlling intracellular signals by protease zymogen activation. Biochim. Biophys. Acta. 1477:299-306. Stennicke, H.R., Deveraux, Q.L., Humke, E.W., Reed, J.C., Dixit, V.M., and Salvesen, G.S. 1999. Caspase-9 can be activated without proteolytic processing. J. Biol. Chem. 274:8359-8362. Stennicke, H.R., Renatus, M., Meldal, M., and Salvesen, G.S. 2000. Internally quenched fluorescent peptide substrates disclose the subsite preferences of human caspases 1, 3, 6, 7 and 8. Biochem. J. 350:563-568. Sun, C., Cai, M., Meadows, R.P., Xu, N., Gunasekera, A.H., Herrmann, J., Wu, J.C., and Fesik, S.W. 2000. NMR structure and mutagenesis of the third bir domain of the inhibitor of apoptosis protein XIAP. J. Biol. Chem. 275:33777-33781. Thornberry, N.A. 1993. Interleukin-1b converting enzyme. Methods Enzymol. 244:615-632. Thornberry, N.A., Bull, H.G., Calaycay, J.R., Chapman, K.T., Howard, A.D., Kostura, M.J., Miller, D.K., Molineaux, S.M., Weidner, J.R., Aunins, J., Elliston, K.O., Ayala, J.M., Casano, F.J., Chin, J., Ding, G.J.F., Egger, L.A., Gaffney, E.P., Limjuco, G., Palyha, O.C., Raju, S.M., Rolando, A.M., Salley, J.P., Yamin, T.T., and Tocci, M.J. 1992. A novel heterodimeric cysteine protease is required for interleukin-1beta processing in monocytes. Nature 356:768-774. Thornberry, N.A., Rano, T.A., Peterson, E.P., Rasper, D.M., Timkey, T., Garcia-Calvo, M., Houtzager, V.M., Nordstrom, P.A., Roy, S., Vaillancourt, J.P., Chapman, K.T., and Nicholson, D.W. 1997. A combinatorial approach defines specificities of members of the caspase family and granzyme B. J. Biol. Chem. 272:1790717911. Van Der Werff Ten Bosch, J., Otten, J., and Thielemans, K. 2001. Autoimmune lymphoproliferative syndrome type III: An indefinite disorder. Leuk. Lymphoma 41:55-65. Walker, N.P.C., Talanian, R.V., Brady, K.D., Dang, L.C., Bump, N.J., Ferenz, C.R., Franklin, S., Ghayur, T., Hackett, M.C., Hammill, L.D., Herzog, L., Hugunin, M., Houy, W., Mankovich, J.A., McGuiness, L., Orlewicz, E., Paskind, M., Pratt, C.A., Reis, P., Summani, A., Terranova, M., Welch, J.P., Xiong, L., and Möller, A. 1994. Crystal structure of the cysteine protease interleukin-1beta-converting enzyme: A (p20/p10)2 homodimer. Cell 78:343-352. Watt, W., Koeplinger, K.A., Mildner, A.M., Heinrikson, R.L., Tomasselli, G., and Watenpaugh, K.D. 1999. The atomic resolution structure of human caspase-8, a key activator of apoptosis. Structure 27:1135-1143. Peptidases
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Wei, Y., Fox, T., Chambers, S.P., Sintchak, J., Coll, J.T., Golec, J.M., Swenson, L., Wilson, K.P., and Charifson, P.S. 2000. The structures of caspases1, -3, -7 and -8 reveal the basis for substrate and inhibitor selectivity. Chem. Biol. 7:423-432. Wilson, K.P., Black, J.A., Thomson, J.A., Kim, E.E., Griffith, J.P., Navia, M.A., Murcko, M.A., Chambers, S.P., Aldape, R.A., Raybuck, S.A., and Livingston, D.J. 1994. Structure and mechanism of interleukin-1 beta converting enzyme. Nature 370:270-275. Yang, X., Stennicke, H.R., Wang, B., Green, D.R., Janicke, R.U., Srinivasan, A., Seth, P., Salvesen, G.S., and Froelich, C.J. 1998. Granzyme B mimics apical caspases. Description of a unified pathway for trans-activation of executioner caspase-3 and -7. J. Biol. Chem. 273:34278-34283. Yuan, J., Shaham, S., Ledoux, S., Ellis, H.M., and Horvitz, H.M. 1993. The C. elegans cell death gene ced-3 encodes a protein similar to mammalian interleukin-1b-converting enzyme. Cell 75:641-652.
Zhou, Q. and Salvesen, G.S. 1997. Activation of pro-caspase-7 by serine proteases includes a non-canonical specificity. Biochem. J. 324:361364. Zhou, Q., Snipas, S., Orth, K., Dixit, V.M., and Salvesen, G.S. 1997. Target protease specificity of the viral serpin CrmA: Analysis of five caspases. J. Biol. Chem. 273:7797-7800. Zhou, Q., Krebs, J.F., Snipas, S.J., Price, A., Alnemri, E.S., Tomaselli, K.J., and Salvesen, G.S. 1998. Interaction of the baculovirus anti-apoptotic protein p35 with caspases: Specificity, kinetics, and characterization of the caspase/p35 complex. Biochemistry 37:10757-10765.
Contributed by Jean-Bernard Denault and Guy S. Salvesen The Burnham Institute La Jolla, California
Caspases
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Use of GFP as a Reporter for the Analysis of Sequence-Specific Proteases
UNIT 21.9
Sequence-specific proteases are endopeptidases that cleave adjacent to a specific amino acid sequence motif. Substrates for sequence-specific proteases are polypeptides containing a scissile site flanked by structural domains with specific recognition sequences. The necessity for a specific amino acid sequence within a larger polypeptide precludes the use of simple peptide substrates such as those routinely employed to assay catabolic peptidases. Sequence-specific peptidases are routinely assayed by synthesizing a radioactive substrate in vitro and using SDS-PAGE (UNIT 10.1) to follow the appearance of the cleavage product. The time-consuming nature of this assay is an impediment to the biochemical characterization of these important enzymes. The ability to engineer recombinant fusion proteins presents a facile means to generate virtually any polypeptide substrate in biochemical amounts. By including a convenient N-terminal affinity handle and an easily assayed reporter protein C-terminal to the scissile bond, sequence-specific protease activity is easily measured as the amount of reporter protein remaining in the supernatant after removal of uncleaved substrate by affinity chromatography. As described in this unit, an N-terminal 6×His tag provides a convenient purification handle for small-scale batch separation of cleaved and uncleaved substrate (see Support Protocol). Green fluorescent protein (GFP), a naturally fluorescent protease-resistant protein, is a sensitive reporter protein for the spectrofluorometric assay of cleaved substrate (see Basic Protocol). Together, the 6×His tag and GFP can be used to design substrates suitable for the rapid, sensitive, high-throughput assay of virtually any sequence-specific protease, allowing detailed studies of these fascinating enzymes. An alternate assay that monitors fluorescent resonance energy transfer to assay site-specific proteases is also discussed (see Alternate Protocol). STRATEGIC PLANNING Any commercially available expression system can be used to produce the His-tagged GFP-fusion substrate. One convenient vector is the Novagen pET28 series, which contains a T7 RNA polymerase binding site to control expression, a region encoding a 6×His tag, followed by a multiple cloning site for insertion of the substrate-GFP fusion protein. PCR provides a convenient method for cloning the substrate coding region by incorporating an NdeI site at the 5′ end and a BamHI site at the 3′ end allowing the GFP coding region to be cloned in-frame as a BamHI/NotI fragment obtained from any of the Clontech EGFP vectors. The substrate-GFP fusion is transformed into a suitable E. coli host (XL-1 Blue; JM 109, and others). Transformed colonies are screened for plasmid, and used as a source of engineered plasmids for transformation into the E. coli host (BL-21 DE3) for expression of the GFP-fusion substrate under control of the inducible T7 RNA polymerase present in the host strain. Isopropyl-β-D-galactopyranoside induction produces high levels of soluble EGFP in 2 to 3 hr. High level induction of soluble substrateGFP fusions is not always obtained under these conditions. Aeration, growth, temperature, and induction level influence the amount of soluble substrate-GFP fusion protein obtained. Optimal induction conditions must be determined empirically for each substrateGFP fusion.
Peptidases Contributed by Gautam Sarath and Steven D. Schwartzbach Current Protocols in Protein Science (2001) 21.9.1-21.9.10 Copyright © 2001 by John Wiley & Sons, Inc.
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BASIC PROTOCOL
FLUORESCENT ASSAY OF SITE-SPECIFIC PROTEASE This procedure describes a rapid sensitive fluorescent assay for sequence-specific proteases. Saturating amounts of a His-tagged-substrate-GFP fusion protein are incubated with a sequence-specific protease for varying time periods. At the end of the reaction, the mixture is incubated with nickel-charged metal-chelate resin. His-tagged undigested substrate-GFP fusion protein and His-tagged substrate are bound to the resin and separated from the proteolytically released GFP by centrifugation. Proteolytic activity is determined by measuring GFP remaining in the supernatant using a spectrofluorometer with an excitation of 484 nm and an emission of 509 nm. Materials Purified stock substrate-GFP fusion protein (see Support Protocol) 5× or 10× stock buffers for protease assay Stock solutions of additives for optimal enzyme activity Extract containing sequence-specific protease Milli-Q water or equivalent 5× binding buffer (see recipe), ice cold Washed metal-chelate affinity resin, e.g., 1:1 (w/v) aqueous slurry of Talon beads (Clontech) Stock solutions of sequence-specific protease inhibitor (optional) 1.7-ml microcentrifuge tubes Rocker-shaker, 4°C Spectrofluorometer/fluorescence plate reader Assemble assay tubes and run reaction 1. Pipet 500 µl containing 25,000 fluorescence units (λex = 484 nm; λem = 509 nm) of substrate-GFP fusion protein (see Support Protocol) into a 1.7-ml microcentrifuge tube placed on ice. Add 300 µl of 5× buffer solution, additives, extract containing sequence-specific protease, and enough Milli-Q water to get a final volume of 1500 µl. Assemble duplicate tubes for each reaction and appropriate controls. The source of sequence-specific protease and the expected relative activity will determine the conditions for isolation and concentrations required in the assay. The substrate-GFP fusion protein should be at a concentration of 30,000 to 60,000 fluorescence units/ml and the amount added adjusted accordingly. Saturating substrate levels for a fixed amount of enzyme must be determined empirically. Each assay should include a tube without enzyme that is used to determine the residual substrate remaining in the supernatant after batch metal-chelate chromatography. A tube containing boiled enzyme should also be included to ensure that substrate release is enzymatic.
2. Cap tubes, mix gently, and incubate at the appropriate temperature. IMPORTANT NOTE: Some substrate-GFP fusion proteins precipitate if shaken too vigorously.
Stop the reaction and separate released GFP from substrate-GFP fusion protein 3. At three evenly spaced time points, withdraw 450 µl from the reaction and mix on ice in a 1.7-ml microcentrifuge tube containing 200 µl of ice-cold 5× binding buffer, 200 µl of a 1:1 aqueous slurry of Talon resin, and 150 µl water. Incubate 30 min on a gently rocking shaker at 4°C. Use of GFP as a Reporter for the Analysis of Sequence-Specific Proteases
Dilution of the reaction mixture into ice-cold reagents and incubation at 4°C rapidly stops the reaction. Some substrate-GFP fusion proteins are heat stable and the reaction can be stopped by denaturing the protease by incubating 5 min at 50° to 70°C. Reactions can also be rapidly terminated by addition of an enzyme inhibitor. Shaking to obtain adequate
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mixing of the resin with the assay solution is critical to achieve reproducible removal of undigested and cleaved substrate. Preliminary experiments should be performed to determine the minimum amount of metal-chelate resin needed to remove the maximum fraction of added substrate.
4. Microcentrifuge tubes 30 sec at maximum speed, 4°C, to sediment resin containing uncleaved substrate-GFP fusion protein. Carefully pipet 800 µl of the supernatant into a new labeled 1.7-ml microcentrifuge tube. Tubes can be stored on ice for a few hours or, with some constructs, overnight prior to fluorometry. Note that some substrate-GFP fusions can precipitate upon storage so it is best to quantitate the samples as soon as possible.
Determine released GFP by spectrofluorometry 5. Measure the fluorescence of the assay reactions in a spectrofluorometer using an excitation of 484 nm and an emission of 509 nm. Subtract the fluorescence of the control lacking enzyme from each time point to determine GFP released by proteolysis. It is important to predetermine the linearity of the reaction over time. Once the time scales for linear release of GFP from the fusion substrate in the presence of a known quantity of enzyme or crude extract is known, appropriate time points can be chosen for rate determinations.
SUBSTRATE-GFP FUSION PROTEIN PURIFICATION USING METAL-CHELATE CHROMATOGRAPHY
SUPPORT PROTOCOL
This procedure describes a method for the isolation and purification of recombinant His-tagged substrate-GFP fusion proteins. The example presented here is for a substrateGFP fusion protein, SSU-10-GFP, used to assay the Euglena polyprotein processing peptidase. A cDNA encoding a protein composed of the small subunit of ribulose-bisphosphate carboxylase (SSU) linked to GFP by the decapeptide recognition sequence for the polyprotein-processing protease was cloned into the pET28 vector. The encoded protein is composed of an N-terminal His-tagged thrombin cleavage site SSU-decapeptide-GFP with the His-tagged thrombin cleavage site contributed by the pET28 vector. A crude protein extract is produced by five freeze-thaw cycles followed by centrifugation. The His-tagged substrate-GFP fusion protein is purified by metal-chelate chromatography. This removes substrate-fusion protein that does not bind to the metal-chelate resin and E. coli proteases that will interfere with the enzyme assay. Materials E. coli transformed with expression vector encoding His-tagged substrate-GFP fusion protein LB medium (UNIT 5.2) 5× binding buffer (see recipe) 1:1 (w/v) aqueous slurry of Talon resin Elution buffer (see recipe) Enzyme assay buffer 26°C rotary shaker for growing bacterial cultures 15-ml culture tubes 2-liter flasks Spectrofluorometer 250-ml centrifuge bottles Refrigerated centrifuge 50-ml conical centrifuge tubes
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Sonicator with microtip 26-ml polycarbonate bottles for ultracentrifuge Ultracentrifuge with a 70 Ti rotor (Beckman) or equivalent 1-ml gravity flow chromatography columns Additional reagents and equipment for dialysis (APPENDIX 3B) Harvest bacterial cells expressing substrate-GFP fusion protein 1. Grow a 5-ml E. coli culture in LB medium in a 15-ml culture tube overnight at 26°C. In the morning, inoculate 750 ml LB medium in a 2-liter flask with the 5-ml overnight culture. Grow overnight at 26°C shaking at 175 rpm. This growth procedure has been used to produce a number of different soluble substrateGFP fusion proteins from pET28 plasmids transformed into E. coli BL21(DE3). Production of substrate begins well after stationary phase has been achieved at a culture density A600 of 1.8 to 2.2. Maximum induction is usually attained after 24 hr. Conditions of aeration, time, and temperature must be evaluated for each construct.
2. Measure substrate-GFP fusion protein levels in a spectrofluorometer by placing an aliquot of the culture directly into the spectrofluorometer. Blank the spectrofluorometer against an aliquot of the culture supernatant prepared by microcentrifuging 1 min at maximum speed. 3. When the culture fluorescence is between 200 and 300 fluorescence units/ml, transfer culture into 250-ml centrifuge bottles and harvest the culture by centrifuging 5 min at 4000 × g, 4°C. A large portion of SSU-GFP fusion protein is often in fluorescent inclusion bodies that can not be solubilized for use in protease assays. The typical yield of purified substrate is 20,000 fluorescent units per 750 ml culture. The yield for each construct must be determined empirically.
Isolate soluble recombinant substrate-GFP fusion protein 4. Resuspend the cell pellet in 15 ml of 5× binding buffer/750 ml of culture, and place in a 50-ml plastic conical centrifuge tube. Freeze suspension at −70°C. Cells can be stored for 3 to 4 months at −70°C. It is best to accumulate a large quantity of cells and process the amount needed just prior to an assay. It is most efficient to process cell suspensions in batches containing cells from a 750 ml culture. Some substrate-GFP fusions precipitate over time after purification, so it is best to use the purified substrate as quickly as possible.
5. Thaw frozen cells in a 30°C water bath and repeat freeze-thaw cycle five times. 6. Sonicate cells using five 15-sec bursts at the maximum power setting for a microtip. Loss of viscosity indicates sonication is complete.
7. Clarify lysate by centrifuging 15 min at 27,000 × g, 4°C. The supernatant contains all of the soluble substrate-GFP fusion protein. A large amount of fluorescent protein is recovered in the pellet. This represents protein present in inclusion bodies and unbroken cells. Additional cycles of freeze thawing and sonication do not release significant amounts of soluble substrate-GFP fusion protein.
8. Transfer supernatant to a 26-ml polycarbonate bottle and ultracentrifuge supernatant 1 hr at 100,000 × g (70 Ti rotor), 4°C. Use of GFP as a Reporter for the Analysis of Sequence-Specific Proteases
The brown translucent pellet contains negligible amounts of substrate-GFP fusion protein. This step removes material that reduces the flow rate of the metal-chelate column. Omission of this step does not decrease the yield of purified substrate-GFP fusion protein but it
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substantially increases the time needed for column chromatography. The lysate can be stored overnight at 4°C.
Purify recombinant substrate-GFP fusion protein 9. Pour 2 ml of a 1:1 aqueous slurry of Talon resin into a small 1-ml gravity flow chromatography column. All chromatography steps are performed at 4°C.
10. Equilibrate column with 5 vol of 5× binding buffer. Load substrate on column. 11. Wash column with 5× binding buffer until eluant is clear. A significant amount of substrate-GFP fusion protein is not retained by the column reflecting the fact that significant amounts of fusion protein in the crude bacterial extract is in a conformation that can not bind the metal-chelate resin.
12. Elute bound substrate from column with elution buffer. The bright green color of GFP is clearly visible under fluorescent lights allowing column fractions to be collected by hand. The most intense fractions are pooled.
13. Dialyze substrate-GFP fusion protein against enzyme assay buffer and store at 4°C. The substrate should be at a concentration of 40,000 to 60,000 fluorescence units/ml. The substrate can be concentrated with a centrifugal membrane concentrator. For some substrate fusion proteins, storage at 4°C can result in a loss of the ability to bind the metal-chelate resin. Stability of metal-chelate binding ability must be determined empirically for each substrate.
FLUORESCENT RESONANCE ENERGY TRANSFER ASSAY OF SITE-SPECIFIC PROTEASES
ALTERNATE PROTOCOL
An alternative GFP based fluorescent assay for site-specific proteases is based on fluorescent resonance energy transfer between two GFP spectral variants linked by the protease target sequence (Mitra et al., 1996; UNIT 19.5). Cleavage of the substrate separates the two GFPs so fluorescence resonance energy transfer (FRET) does not occur and the amount of cleaved substrate is determined by the decreased fluoresence of the longer wavelength GFP. FRET is a complex function of the distance between the chromophores and the angle formed by their dipole moments (Stryer, 1978). The requirement for the substrate to fold so that the GFPs flanking the protease cleavage site are properly positioned for FRET to occur complicates substrate design limiting the utility of this assay. A second limitation of this assay is that fluorescence spectra must be measured for each sample because protease activity is determined from the ratio of fluorescence of the individual GFPs. Calculation of activity is more time consuming than the direct fluorescence measurements needed to determine the amount of GFP remaining in the supernatant after batch metal-chelate chromatography. Although the FRET based assay is a sensitive alternative for determining the activity of site-specific proteases, measuring the direct release of GFP from a substrate-GFP fusion protein is a simpler method that is more suitable for the rapid analysis of multiple samples and multiple substrates.
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Binding buffer, 5× 0.25 M NaH2PO4, pH 8.0 0.5 M NaCl (APPENDIX 2E) 0.05 M Tris⋅Cl, pH 8.0 (APPENDIX 2E) Prepare a stock solution of 0.5 M NaHPO4, pH 8.0 by adding 0.5 M NaH2PO4 to adjust a solution of 0.5 M Na2HPO4 to pH 8.0. Mix 50 ml of 0.5 M NaHPO4, pH 8.0, 5 ml of 1 M Tris⋅Cl, pH 8.0, 10 ml of 5 M NaCl and 35 ml water. Store up to 6 months at 4°C. Elution buffer 60 mM imidazole 30 mM NaCl 1.2 mM Tris⋅Cl, pH 7.9 Store up to 6 months at 4°C COMMENTARY Background Information
Use of GFP as a Reporter for the Analysis of Sequence-Specific Proteases
The Basic Protocol describes a simple rapid assay for structure and sequence-specific proteases. The defining feature of structure and sequence-specific proteases is that they recognize a structural element or amino acid sequence within a polypeptide producing two cleavage products of a defined size. Measurements of protease activity are based upon either the disappearance of the uncleaved polypeptide or the appearance of the cleavage product. This assay entails separating the cleavage products by size, usually by SDS gel electrophoresis (UNIT 10.1). Radiolabeled substrates produced by in vitro translation can be used allowing quantitative analysis of substrate and product levels present in the gel (Enomoto et al., 1997). Alternatively, western blotting can be used to detect the reaction products (UNIT 10.10). The requirement to run each reaction on an SDS gel and the additional time required to develop a radioactive or western blot signal makes this an extremely cumbersome assay suitable for only a limited number of samples. End-point chromogenic assays are among the most commonly used enzyme assays. An enzymatic reaction produces a compound that either absorbs light or fluoresces at a wavelength differing from that of the substrate. Measurements of absorbance or fluorescence are simple, highly sensitive and rapid making these large throughput assays. The assay described in this protocol presented diagrammatically in Figure 21.9.1 represents an assay for site-specific proteases with all the advantages
of a chromogenic endpoint assay. The basis of this assay is the ability of the autofluorescent protein GFP to be fused to the C-terminus of most proteins without altering the functional properties of the fusion partner (Tsien, 1998). This essentially enables any site-specific protease substrate to be converted into a chromogenic substrate that, upon proteolytic cleavage, produces a fluorescent and nonfluorescent product. Rather than using a size difference to separate the uncleaved substrate from the cleaved reaction products, the chromogenic substrate-GFP fusion protein contains an N-terminal His tag allowing batch metal-chelate chromatography to be used to separate unreacted substrate from the fluorescent GFP cleavage product. The tedious process of running an SDS gel is replaced by a 0.5-hr incubation with metal-chelate beads and microcentrifugation for 30 sec. Fluorometry of the supernatant replaces the time consuming quantitation of product by western blotting, or radioactivity measurements. The assay was developed using purified thrombin as a site-specific protease (Pehrson et al., 1999). The major requirements for an enzyme assay, easy sensitive measurement of released product, linearity of initial rates of substrate catalysis with respect to enzyme and substrate concentration, and ability to assay enzyme at saturating substrate concentrations were achieved. The fluorometric assay has been successfully used to study the chloroplast-localized polyprotein-processing protease of Euglena. This processing protease has been quite diffi-
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sequence-specific protease cleavage site polyhistidine linker measure fluorescence at start of assay GFP (t = 0) to obtain total available substrate GFP - fusion protein substrate add assay buffer + enzyme + stabilizers – wait suitable time (1 to 30 min) 1. add inhibitors/or otherwise quench enzymatic reaction 2. add His-bound resin – incubate with shaking for 30 min
centrifuge pellet undigested substrate + nonfluorescent product discard
supernatant released GFP measure fluorescence
Figure 21.9.1 Flow diagram of fluorescent sequence-specific protease assay.
cult to characterize using standard methods for the analyses of sequence-specific proteases (Enomoto et al., 1997). A number of Euglena chloroplast proteins are synthesized as polyprotein precursors (Houlne and Schantz, 1988; Chan et al., 1990; Muchhal and Schwartzbach, 1992). A decapeptide with two conserved and four invariant positions (bold), AMFAXXGXKX, links mature units within all known Euglena polyprotein precursors (Schiff et al., 1991). The polyprotein-processing protease is a chloroplast enzyme(s) that cleaves at both the N and C terminus of the decapeptide (Enomoto et al., 1997). The small subunit of ribulose-bis-phosphate-carboxylase (SSU) is a Euglena chloroplast protein derived from a polyprotein precursor (Chan et al., 1990). The pET 28 expression system was used to produce a His-tagged recombinant substrateGFP fusion protein, SSU-10-GFP composed of an N-terminal His-tagged thrombin cleavage site-SSU-decapeptide-GFP with the Histagged thrombin cleavage site contributed by the pET28 vector. When a crude membranefree chloroplast lysate (Enomoto et al., 1997) was incubated with the recombinant polyprotein substrate SSU-10-GFP, the rate of GFP release was linear over an 18-min period (Fig.
21.9.2) demonstrating the utility of this assay protocol.
Critical Parameters and Troubleshooting Induction This assay requires large quantities of soluble recombinant His-tagged substrate-GFP fusion protein. The pET 28 vector was chosen because it provides a simple method for cloning a gene encoding a His-tagged substrate-GFP fusion protein with gene expression controlled by a tightly regulated inducible promoter. The Clontech EGFP gene was chosen as the fusion partner because it has a high fluorescent yield and most importantly, high levels of soluble protein expression can be obtained in E. coli grown at 37°C. As a rule, the authors find substrate-GFP fusions are less soluble than GFP alone and high yields of soluble substrateGFP fusion proteins are not obtained in cells grown at 37°C. Temperature, extent of expression induction, and aeration affect the yield of soluble fluorescent substrate GFP-fusions. Growth conditions must be empirically determined for each substrate-GFP fusion. The growth conditions presented in this protocol have been successfully used to produce large Peptidases
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Fluorescence
800 – 700 –
SSU-10-GFP
600 – 500 – 400 – SSU-GFP
300 – 200 – 100 – –
–
–
–
–
0– 0
5
10
15
20
Time (min)
Figure 21.9.2 Rate of GFP release from the decapeptide containing recombinant polyprotein, SSU-10-GFP, and the recombinant polyprotein, SSU-GFP, lacking the decapeptide. Substrates were incubated with chloroplast extract and at the indicated times, uncleaved GFP containing substrate was bound to metal-chelate resin. The resin was recovered by centrifugation and unbound GFP remaining in the supernatant was quantitated by spectrofluorometry.
amounts of a number of substrate-GFP fusions. The low rate of leaky expression when cells enter stationary phase is much more compatible with soluble protein accumulation than is the high expression rates produced by induction with IPTG. In cases where it has been impossible to obtain soluble fluorescent substrateGFP fusions, truncations of the substrate portion of the fusion often increase solubility witho ut alterin g reco gn ition b y the sequence-specific peptidase.
Use of GFP as a Reporter for the Analysis of Sequence-Specific Proteases
Sensitivity The simplicity of the assay derives from the use of metal-chelate chromatography to separate uncleaved fluorescent substrate from the fluorescent GFP released by proteolysis. The sensitivity of the assay depends on the amount of uncleaved substrate remaining in the supernatant after metal-chelate chromatography. The larger the amount of uncleaved substrate remaining in the supernatant, the greater the amount of product that must be released to produce a significant increase in supernatant fluorescence. For many substrate-GFP fusion proteins, as much as 50% of the fluorescent protein in a bacterial lysate does not bind the metal-chelate resin and remains in the supernatant. Metal-chelate chromatography of the crude bacterial lysate purifies the binding competent fraction of the substrate-GFP fusion protein. Less than 2% of the purified substrateGFP fusion protein remains in the supernatant
at the end of an assay after batch chromatography with a saturating amount of metal-chelate resin. Purification of the substrate-GFP fusion protein has the additional advantage that proteolytic inhibitors and proteases present in the crude bacterial lysate are also removed. The fraction of the purified substrate-GFP fusion protein that does not bind resin increases during storage at 4°C. The rate of binding loss must be determined for each substrate as there are large differences in substrate stability. It is prudent to always include a boiled enzyme and minus enzyme control to monitor each series of reactions for the fraction of residual uncleaved substrate-GFP fusion protein remaining in the supernatant. Cleavage specificity The major disadvantage of the fluorescent assay is that the size of the cleavage product is unknown. Cleavage at any site C-terminal to the the His-tag releases GFP contributing to the measured reaction rate. Assays utilizing SDSgel electrophoresis are not plagued by this problem as reaction rates are determined by measuring the rate of appearance of a protein having a specific size (Enomoto et al., 1997). Cleavage at a site other than the target site would produce a protein product having a different size and the amount of this protein would not contribute to the overall reaction rate. This problem can be detected by comparing reaction rates using a substrate-GFP fusion protein con-
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SSU-10-GFP 3 min 15 min
SSU-GFP 3 min 15 min
cut SSU-GFP
GFP
Figure 21.9.3 Western blot of GFP released from the decapeptide containing recombinant polyprotein, SSU-10-GFP, and the recombinant polyprotein, SSU-GFP, lacking the decapeptide. Substrates were incubated with chloroplast extract and at the indicated times, uncleaved GFP containing substrate was bound to metal-chelate resin. The resin was recovered by centrifugation and unbound proteins remaining in the supernatant were separated by SDS-polyacrylamide gel electrophoresis. Proteins were blotted to nitrocellulose and GFP containing proteins were detected using anti-GFP as the primary antibody and horseradish peroxidase as the secondary antibody.
taining the putative sequence-specific cleavage site and a second substrate-GFP fusion protein that lacks the protease cleavage site but is otherwise identical to the first substrate. The utility of this approach is demonstrated using the Euglena polyprotein-processing protease as an example. Equal aliquots of a Euglena chloroplast lysate were incubated with the substrate fusion protein SSU-10-GFP and the substrate fusion protein SSU-GFP. The only amino acid sequence difference between the two substrates is the presence of the polyprotein-processing protease-recognition sequence, MAAMTGEKD, in SSU-10-GFP. There was a low but significant rate of GFP release from SSU-GFP (Fig. 21.9.2). The rate of GFP release from SSU-10-GFP, the decapeptide containing substrate, was, however, >3-fold greater than the rate of release from SSU-GFP, the substrate lacking the decapeptide. The rate of GFP release from SSU-10-GFP and SSU-GFP was linear over an 18-min period. Immunoblots of the proteins remaining in the supernatant after metal-chelate batch chromatography of chloroplast extracts incubated with SSU-10-GFP identified two GFP-containing proteins as reaction products (Fig. 21.9.3). The largest protein, cut SSU-GFP, is similar in size to the uncleaved recombinant polyprotein substrate suggesting cleavage within the N-terminal region. The smaller proteolytic products, GFP, are the size expected for decapeptide-GFP and free GFP released by cleavage at the N- or C-terminus of the decapeptide. The amount of all detected proteins increased between 3 and
15 min of incubation with enzyme extract indicative of release through proteolytic activity. The larger product, cut SSU-GFP, is the only product seen when SSU-GFP was used as substrate indicating that it is formed by cleavage at a site other than the polyprotein processing peptidase cleavage site. Taken together, these results show that a substrate-GFP fusion protein lacking the protease target site provides a control for determining reaction specificity.
Anticipated Results This assay provides a simple, rapid, accurate, sensitive method for measuring levels of site-specific proteases over at least one order of magnitude. Sufficient substrate can be produced to allow for determination of kinetic parameters and inhibitor cross-sections. Substrate-GFP fusion proteins with site directed mutations within the protease recognition sequence are easily produced allowing identification of the critical sequence determinants for substrate recognition. The simplicity and rapidity of the assay makes it ideally suited for the assay of the large number of samples generated during enzyme purification.
Time Considerations Production and purification of the substrateGFP fusion protein requires ∼3 days. The initial bacterial innoculum is grown overnight and used to start a large-scale culture that is grown for 24 hr. Cell harvesting and resuspension in sonication buffer requires ∼30 min and the cell suspension can be frozen at −70°C for sub-
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sequent processing. Preparation of the highspeed supernatant containing the substrateGFP fusion protein requires ∼3 hr including 1.25 hr of centrifugation. Purification of the substrate by metal-chelate chromatography requires ∼4 hr. It is convenient to start preparing the high-speed supernatant in the afternoon and to run the metal-chelate column overnight. The protease assay itself requires ∼2 hr for 20 to 40 samples.
Pehrson, J.C., Weatherman, A., Markwell, J., Sarath, G., and Schwartzbach, S.D. 1999. The use of GFP for the facile analysis of sequence-specific proteases. BioTechniques 27:28-32.
Literature Cited
Tsien, R.Y. 1998. The green fluorescent protein. Annu. Rev. Biochem. 67:509-544.
Chan, R.L., Keller, M., Canaday, J., Weil, J.H., and Imbault, P. 1990. Eight small subunits of Euglena ribulose 1-5 bisphosphate carboxylase/oxygenase are translated from a large mRNA as a polyprotein. EMBO J. 9:333-338. Enomoto, T., Sulli, C., and Schwartzbach, S.D. 1997. A soluble chloroplast protease processes the Euglena polyprotein precursor to the light harvesting chlorophyll a/b binding protein of photosystem II. Plant Cell Physiol. 38:743-746. Houlne, G. and Schantz, R. 1988. Characterization of cDNA sequences for LHCI apoproteins in Euglena gracilis: The mRNA encodes a large precursor containing several consecutive divergent polypeptides. Mol. Gen. Genet. 213:479486. Mitra, R.D., Silva, C.M., and Youvan, D.C. 1996. Fluorescence resonance energy transfer between blue-emitting and re-shifted excitation derivatives of the green fluorescent protein. Gene 173:13-17. Muchhal, U. and Schwartzbach, S.D. 1992. Characterization of a Euglena gene encoding a polyprotein precursor to the light harvesting chlorophyll a/b binding protein of photosystem II. Plant Mol. Biol. 18:287-299.
Schiff, J.A., Schwartzbach, S.D., Osafune, T., and Hase, E. 1991. Photocontrol and processing of LHCPII apoprotein in Euglena: Possible role of golgi and other cytoplasmic sites. J. Photochem. Photobiol. Biol. 11:219-236. Stryer, L. 1978. Fluorescence energy transfer as a spectroscopic ruler. Annu. Rev. Biochem. 47:819-846.
Key References Enomoto et al., 1997. See above. Demonstrates the difficulties inherent in using radiolabeled substrates produced by in vitro translation and SDS gel electrophoresis to assay site-specific protease activity. Tsien, 1998. See above. A comprehensive review of the biochemical properties of GFP and its uses in biological research.
Contributed by Gautam Sarath University of Nebraska Lincoln, Nebraska Steven D. Schwartzbach University of Memphis Memphis, Tennessee
This work was supported by National Science Foundation grants MCB 9630817 and MCB 00800345 (S.D.S.) and the Center for Biotechnology, University of Nebraska-Lincoln.
Use of GFP as a Reporter for the Analysis of Sequence-Specific Proteases
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An Overview of Serine Proteases FAMILIES OF SERINE PROTEASES The term “serine protease” denotes a mechanistic class of protease reactions that utilize a catalytic Ser residue. These proteases are involved in a wide variety of physiological processes, including digestion, development, blood coagulation, fibrinolysis, immune response, prohormone processing, signal transduction, and complement fixation. Over 700 serine proteases have been identified, making these enzymes among the most abundant in nature. This unit will focus on “classic” serine proteases that contain the catalytic triad Ser-His-Asp (Figure 21.10.1). This catalytic triad is found in at least three structurally unrelated contexts, indicating that it has evolved at least three independent times. The prototypes of these three structural classes are trypsin, subtilisin, and serine carboxypeptidase II (reviews of the structures of these enzymes can be found in Remington, 1993; Perona and Craik, 1995). Using the MEROPS nomenclature outlined in UNIT 21.1, these enzymes represent clans PA(S) [formerly clan SA; the new nomenclature links these proteases to the structurally-related clan of cysteine proteases PA(C)], SB and SC, respectively (Rawlings and Barrett, 1994). Six other clans of serine proteases have been identified that utilize different chemical strategies to activate Ser. Clans SH and SN utilize alternate triads involving Ser/His/His and Ser/His/Glu, respectively. Clans SE, SF, SL, and SM utilize “catalytic dyads” involving Ser/Lys or Ser/His. The mammalian digestive proteases trypsin, chymotrypsin, and elastase are among the best characterized of all enzymes and the trypsin clan is the largest among all protease groups (for a good review of the catalytic properties of trypsin, see Kossiakoff, 1987). Members have been found in bacteria, archae, fungi, plants,
UNIT 21.10
animals, and viruses—every phylum except archezoa and protozoa. Many physiological processes involve cascades of sequential activations of trypsin-like proteases, including development, blood coagulation, fibrinolysis, and complement fixation. The catalytic-triad residues are His57, Asp102, and Ser195 (chymotrypsinogen numbering system). The active site spans a crevice between two β-barrel domains. Trypsin-like proteases are usually monomeric, although dimeric and tetrameric examples also exist. Many of these proteases are modular. For example, tissue plasminogen activator, one of the key proteases of fibrinolysis, contains finger, growth factor, and two kringle domains in addition to the protease domain. These proteases are often glycosylated; removal of the glycosyl groups usually does not affect activity. Clan SB includes subtilisin, a bacterial protease commonly used as an additive in laundry detergents, as well as proteases involved in prohormone processing such as KEX2 protease, furin, and PC2. Members of this clan are found in organisms representing every phylum except archezoa. These proteases have a single domain comprised of seven β sheets surrounded by nine α helices. The catalytic triad is found in the order Asp-His-Ser. Clan SC is also widely distributed, with members found in every phylum except archezoa and viruses. Examples include carboxypeptidase C, prolyl aminopeptidase, and prohormone processing enzymes such as dipeptidyl peptidase II, angiotensinase C and the yeast KEX1 gene product. These proteases have a structure known as the α/β hydrolase fold that contains an 11-stranded β sheet structure surrounded by 15 α helices. This fold is also found in esterases and other hydrolytic
O O–
HN
Asp
N His
H O Ser
Figure 21.10.1 The catalytic triad. Peptidases Contributed by Lizbeth Hedstrom Current Protocols in Protein Science (2001) 21.10.1-21.10.8 Copyright © 2001 by John Wiley & Sons, Inc.
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folding and secretion of the mature protease (Silen et al., 1989). The prodomain binds in the active site and acts as a template for the folding of the protease domain. The prodomain is removed and degraded by α-lytic protease. This mechanism is also widespread; examples are found in subtilisin-like enzymes as well as other protease classes (Baker et al., 1993).
enzymes. The catalytic triad is found in the order Ser-Asp-His.
SERINE PROTEASE PRECURSORS Virtually all proteases are translated as inactive precursors. In serine proteases, pro regions can consist of a few residues or separate domains of several hundred residues. Small pro regions prevent the formation of an active protease conformation as exemplified by the conversion of trypsinogen to trypsin. Although most of the structure of trypsinogen is identical to trypsin, the S1 site is deformed (Huber and Bode, 1978). Enterokinase activates trypsinogen by removing eight residues from the N-terminus. The new N-terminus, Ile16, forms a salt bridge with Asp194, triggering a conformational change that orders the active site. Similar conformational rearrangements are found in the conversion of chymotrypsinogen to chymotrypsin and in the protease cascades that drive blood coagulation, complement fixation and fibrinolysis. Large pro regions operate by a different mechanism as exemplified by the 166 residue prodomain of the bacterial enzyme α-lytic protease. This enzyme is also a member of the trypsin clan. The prodomain is required for
THE MECHANISM OF THE SERINE PROTEASE REACTION The reaction involves the attack of the active site Ser on the scissile peptide bond much like cysteine proteases utilize the active site Cys residue (UNIT 21.2). The Ser is activated by a general base as shown in Figure 21.10.2. In the catalytic triad enzymes, His acts as a general base, supported by a hydrogen bond to the Asp. The oxyanion of the resulting tetrahedral intermediate is stabilized by an “oxyanion hole” formed by mainchain amides in the case of clan PA(S) or an Asn residue in clan SB. The general base transfers the proton to the amine leaving group. The tetrahedral intermediate collapses with expulsion of the product amine, resulting in an acylenzyme intermediate. This acylenzyme is attacked by water, again with the assistance of the general base, to form a second
O R
C
NH
R*
NH2-R*
O–
enz
O
R
O H
O
NH
R*
R O
O
H
H
enz
enz BH+
B
B
enz
enz
enz
O O enz
R
O–
OH
H R
O H O
enz
B
enz BH+ enz
An Overview of Serine Proteases
Figure 21.10.2 The mechanism of the serine protease reaction.
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E+
O R–C–X
Km =
kcat =
k1 k–1
O
E• R–C–X
k–1k3 k1(k2+k3)
=
O
k2
E–O–C–R
k3 HOH
O E + R–C–O–
X–
Kdk3 (k2+k3)
k2k3 (k2+k3)
Figure 21.10.3 The kinetic mechanism of the serine protease reaction.
tetrahedral intermediate. Collapse of this intermediate, with the protonation of the Ser by the general base, releases the product acid and returns the enzyme to its original state. The tetrahedral intermediates are unstable; their presence is mainly inferred from model reactions. In contrast, the presence of an acylenzyme intermediate is well established. Consequently, the kinetics of the reaction can be described in three steps: (1) binding (k1, k-1), (2) acylation (k2), and (3) deacylation (k3) (Figure 21.10.3). Unfortunately, the idea that acylation is rate-limiting for amide substrates, demonstrated for poor substrates of chymotrypsin, has become ingrained in textbooks. Deacylation is actually rate-limiting for good substrates (Hedstrom et al., 1992; Rockwell and Fuller, 2001). Note that since an intermediate is present, Km can no longer be described as a binding constant; low values of Km usually reflect large values of k2, the acylation rate constant, rather than high substrate affinity.
SPECIFICITY OF SERINE PROTEASES Serine proteases can have endo- or exopeptidase activity; substrate specificity can be broad or extraordinarily narrow as required for their physiological function. Digestive proteases generally have few requirements for substrate. For example, chymotrypsin requires only a large hydrophobic residue in the P1 position (e.g., Trp, Phe, Tyr, Leu; nomenclature of Schechter and Berger, as described in Figures 21.1.1 and 21.2.3). Similarly, trypsin prefers positively charged P1 residues (Arg and Lys) and elastase prefers small alkyl side chains
(Ala and Val). In the trypsin family, the specificity of the S1 site can often be predicted by the residue at position 189 (Perona and Craik, 1995). This residue is usually an Asp for proteases that recognize Arg/Lys while proteases that prefer hydrophobic substrates usually contain Ser at this position. Note that in the literature, proteases are often described as chymotrypsin-like, trypsin-like, or elastase-like to denote similar substrate preferences rather than structural or even mechanistic similarity. For other serine proteases, substrate recognition can extend beyond the S1 site. Such proteases can be extraordinarily specific. Enterokinase recognizes Asp-Asp-Asp-Asp-Lys in the P5-P1 positions. Factor D will only hydrolyze its natural protein substrate, the factor C3b⋅B⋅Mg2+ complex, and fails to hydrolyze peptides comprised of sequences spanning the cleavage site (Taylor et al., 1999). All serine proteases, regardless of clan, bind peptides in an extended β sheet–like conformation (Perona and Craik, 1995). This conformation is also found in the inhibitory loops of many naturally occurring protease inhibitors (see below). To the first approximation, the specificity of the subsites is additive and not context-dependent. This is an important observation for the determination of substrate specificity because the optimal interactions at each subsite can be determined in separate experiments and combined to create an optimal substrate. Various combinatorial strategies have been developed to rapidly determine specificity (see below). Interestingly, the optimal substrate determined with these methods does not necPeptidases
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O R
O
HOH R
X
+ XH + H +
O–
X=
NH
NO2
p-nitroaniline (pNA) absorbance wavelength 410 nm,
O
NO2
p-nitrophenol absorbance wavelength 400 nm, ε = 5400 M–1 cm–1 at pH 6.5
ε = 8480 M –1 cm –1
thiobenzylester (SBzl) reaction monitored with thiol detection reagents –1 DTNB, absorbance wavelength 412 nm, ε = 13,600 M cm–1 –1 DTP, absorbance wavelength 324 nm, ε = 19,800 M cm –1
S
CH3 7-amino-4-methyl coumarin (AMC) excitation wavelength 380 nm, emission wavelength 460 nm
NH
O
O
CF3
NH
O
O
7-amino-4-trifluoromethyl coumarin (AFC) excitation wavelength 400 nm, emission wavelength 505 nm absorbance wavelength 380 nm, ε = 12,600 M –1cm –1
Figure 21.10.4 Commonly used leaving groups in serine protease assays.
An Overview of Serine Proteases
essarily correspond to the natural substrate (Ding et al., 1995). Specificity can also be determined by interactions outside the S-P sites and modulated by protein factors and other allosteric effectors. Thrombin is an excellent example of this phenomenon (Stubbs and Bode, 1995). The thrombin catalyzed conversion of fibrinogen to fibrin is the key reaction in blood coagulation. Thrombin has a positively charged exosite that mediates the binding of fibrinogen. In addition, thrombomodulin converts thrombin into an anticoagulation factor. Thrombomodulin binds to thrombin via the exosite, blocking fibrinogen binding; thrombomodulin also changes the specificity of the S sites so that the thrombinthrombomodulin complex preferentially
cleaves and activates protein C, which in turn cleaves the coagulation factors Va and VIIIa, limiting the clotting reaction (Esmon, 2000). Many examples of such allosteric activation and modulation of serine protease activity exist.
ASSAYS OF SERINE PROTEASES Most serine proteases will also hydrolyze esters and amides where the leaving group is replaced with a chromogenic or fluorogenic group. Such compounds ignore the specificity of the S′ sites. Nevertheless, these assays are generally preferred because they are simple, rapid, sensitive, and continuous. Libraries of such compounds allow the rapid determination of S-site specificity (Rano et al., 1997; Harris et al., 2000). A wide variety of such substrates
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O E+ R
C X
Ks
O E• R
C X
O
k2
E
O
C R
k3 HOH
O E+ R
C O–
X– k4
O E+
R
C NHR′
R′NH2
Figure 21.10.5 Acyl transfer reactions for determining S′ specificity.
are commercially available. Some of the more common leaving groups are shown in Figure 21.10.4. S1′ specificity is more difficult to determine. The acylenzyme intermediate can react with amine nucleophiles to form peptide bonds as shown in Figure 21.10.5. This reaction is the reverse of peptide hydrolysis and therefore provides analogous specificity data (Schellenberger et al., 1994). This reaction can be performed with mixtures of peptide nucleophiles, allowing the rapid determination of S′ specificity (Schellenberger et al., 1994). Phage display can also be used to determine both S and S′ specificity (Matthews and Wells, 1993; Smith and Shi, 1995).
INHIBITORS This section briefly introduces some common commercially available inhibitors/inactivators of serine proteases. Many other classes of inhibitors have been developed and several appear to have clinical promise (Leung et al., 2000).
inhibitors are TPCK and TLCK, which inactivate chymotrypsin and trypsin, respectively. Other examples include Phe-Pro-Arg-CMK, a potent inactivator of thrombin and Z-Gly-LeuPhe-CMK, a cathepsin G inactivator. A wide variety of CMK inhibitors are commercially available. These compounds will also inhibit Cys proteases, and are, therefore, not diagnostic for protease mechanism.
Active Site Titrants Compounds that form stable acylenzymes inactivate serine proteases. If these compounds release a chromophore or fluorophore, they can be used to readily measure the concentration of active sites. Examples of such reagents include p-nitrophenyl p′-guanidinobenzoate (pNGB) and 4-methylumbelliferyl p-guanidinobenzoate (MUGB) for trypsin. Note that PNGB can also be used for chymotrypsin; presumably this reagent binds backwards with the guanidino group in the S1′ site and the p-nitrophenyl group in the S1 site.
Transition State Analogs General Inhibitors of Serine Proteases Many broad spectrum inactivators of serine proteases are commercially available (Figure 21.10.6). Diisopropylfluorophosphate (DFP) was among the first inactivators identified for serine proteases. DFP irreversibly modifies the active site Ser. However, DFP is a potent neurotoxin, and its use has declined as less toxic alternatives such as PMSF, AEBSF, and 3,4-dichloroisocoumarin have become available. Sensitivity to these compounds is diagnostic for a serine protease. Chloromethylketones (CMK) are among the most useful of the many classes of serine protease inhibitors. These compounds form irreversible adducts, crosslinking the Ser and His residues. CMK inactivators recapitulate the substrate specificity; the best inactivators match the S/P interactions of an optimal substrate. Perhaps the most well known CMK
Aldehyde and ketones form tetrahedral adducts with the active site Ser. These adducts are transition state analogs, mimicking the tetrahedral intermediates formed during the normal reactions (compare Fig. 21.10.2 and 21.10.6). These inhibitors are reversible; potency depends on the correct S-P interactions. Leupeptin (Ac-Leu-Leu-Arg-CHO) is the best known member of this class of inhibitors. Note that this class of inhibitors is also effective against cysteine proteases where a similar tetrahedral transition state analog is formed.
Small Molecule Reversible Inhibitors A number of reversible competitive inhibitors of serine proteases have also found wide use, especially for proteases with trypsin-like specificity. The two most prevalent are benzamidine and ε-aminocaprioc acid. These compounds compete with substrates for the S1 site.
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O
O
E-OH
O P F O
DFP forms an irreversible adduct with the active site Ser
O P O E O
O
O
E-OH
R S F
PMSF and AEBSF form irreversible adducts with the active site Ser
R S O E
O
O
PMSF, R =
AEBSF, R =
H2NCH2CH2
O
O
O
E-OH
O E
Cl
Cl
Cl
O X E
3,4-dichloroisocoumarin
E-OH
O
R
Cl
O– R O E N
CMK TPCK, R = Tosyl-Phe TLCK, R = Tosyl-Lys
O
R
+ N H
O–
E-OH H
3,4-dichloroisocoumarin forms an adduct with the active site Ser and can also crosslink to another enzyme residue
O E
R H
CMK inhibitors crosslink the active site Ser and His
E
aldehydes form tetrahedral adducts with the active site Ser that mimic the transition state of the hydrolysis reaction
Figure 21.10.6 Inhibitors/inactivators of serine proteases.
Protein Inhibitors of Serine Proteases
An Overview of Serine Proteases
Nature has created a vast array of inhibitors to combat and regulate serine proteases (Laskowski and Kato, 1980). Several families of reversible protein inhibitors exist. These inhibitors exactly complement their target protease, forming extremely high affinity complexes. For example, aprotinin (also known as bovine pancreatic trypsin inhibitor or BPTI) to trypsin (with Kd = 0.1 pM). Aprotinin is also a potent inhibitor of other trypsin-like enzymes. The
protease-inhibitor interaction is mediated by a loop that binds in the active site cleft. This loop is constrained in the canonical substrate conformation; interactions mimic substrate-protease binding. These inhibitors can also interact with other regions of the protease. Serpins are irreversible inhibitors of serine proteases (Lawrence, 1997; UNIT 21.7). The term serpin was originally derived from SERine Protease INhibitor, but now designates a class of protein structures; not all serpins inhibit serine
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proteases. These inhibitors form stable acylenzyme adducts with their target proteases. Examples include antithrombin III, α2-macroglobulin, antitrypsin, and antichymotrypsin. These inhibitors can be very specific, inactivating their target enzyme but leaving proteases with similar specificity unimpaired.
EXPRESSION OF SERINE PROTEASES Serine proteases have been successfully expressed in bacterial, yeast, insect, and mammalian expression systems (for examples see Madison et al., 1989; Hedstrom et al., 1991; Castellino et al., 1993; Huang et al., 2001). Unfortunately, recombinant expression remains an exercise in trial and error. Serine proteases that are naturally secreted are best expressed as secreted recombinant proteins. Many proteases have been successfully refolded from inclusion bodies. In general, proteases are best expressed as zymogens. The N-terminus of proteases with small pro regions can be substituted to create more convenient cleavage sites. While large pro regions can not be substituted, they can be expressed in trans (Silen and Agard, 1989). In addition, since the protease must degrade its prodomain, changing substrate specificity can also require changing the cleavage site between the pro region and the protease domain. As always, care must be taken to ensure that heterologous expression correctly recapitulates postranslational modifications such as glycosylation and gamma carboxylation.
LITERATURE CITED Baker, D., Shiau, A.K., and Agard, D.A. 1993. The role of pro regions in protein folding. Curr. Opin. Cell Biol. 5:966-970. Castellino, F.J., Davidson, D.J., Roasen, E., and McLinden, J. 1993. Expression of human plasminogen cDNA in lepidopteran insect cells and analysis of asparagine-linked glycosylation patterns of recombinant plasminogens. Methods Enzymol. 222:168-185. Ding, L., Coombs, G.S., Strandberg, L., Navre, M., Corey, D.R., and Madison, E.L. 1995. Origins of the specificity of tissue-type plasminogen activator. Proc. Natl. Acad. Sci. U.S.A. 92:7627-7631. Esmon, C.T. 2000. Regulation of blood coagulation. Biochim. Biophys. Acta. 1477:349-360. Harris, J.L., Backes, B.J., Leonetti, F., Mahrus, S., Ellman, J.A., and Craik, C.S. 2000. Rapid and general profiling of protease specificity by using combinatorial fluorogenic substrate libraries. Proc. Natl. Acad. Sci. U.S.A. 97:7754-7759.
Hedstrom, L., Graf, L., Stewart, C., Rutter, W.J., and Phillips, M.A. 1991. Modulation of enzyme specificity by site-directed mutagenesis. Methods Enzymol. 22:671-687. Hedstrom, L., Szilagyi, L., and Rutter, W.J. 1992. Converting trypsin to chymotrypsin: The role of surface loops. Science 255:1249-1253. Huang, X., Knoell, C.T., Frey, G., Hazegh-Azam, M., Tashjian, A.H.J., Hedstrom, L., Abeles, R.H., and Timasheff, S.N. 2001. Modulation of recombinant human prostate-specific antigen: Activation by hofmeister salts and inhibition by azapeptides. Appendix: Thermodynamic interpretation of the activation by concentrated salts. Biochemistry 40:11734-11741. Huber, H. and Bode, W. 1978. Structural basis for the activation and action of trypsin. Acct. Chem. Res. 11:114-122. Kossiakoff, A.A. 1987. Catalytic properties of trypsin. In Biological Macromolecules and Assemblies (F. A. Jurnak and A. McPherson, eds.) pp. 369-412. John Wiley & Sons, New York. Laskowski, M. and Kato, I. 1980. Protein inhibitors of proteinases. Annu. Rev. Biochem. 49:593-626. Lawrence, D.A. 1997. The serpin-proteinase complex revealed. Nat. Struct. Biol. 4:339-340. Leung, D., Abbenante, G., and Fairlie, D.P. 2000. Protease inhibitors: Current status and future prospects. J. Med. Chem. 43:305-341. Madison, E.L., Goldsmith, E.J., Gerard, R.D., Gething, M.J., and Sambrook, J.F. 1989. Serpin-resistant mutants of human tissue-type plasminogen activator. Nature 339:721-724. Matthews, D.J. and Wells, J.A. 1993. Substrate phage: Selection of protease substrates by monovalent phage display. Science 260:1113-1117. Perona, J.J. and Craik, C.S. 1995. Structural basis of substrate specificity in the serine proteases. Prot. Sci. 4:337-360. Rano, T.A., Timkey, T., Peterson, E.P., Rotonda, J., Nicholson, D.W., Becker, J.W., Chapman, K.T., and Thornberry, N.A. 1997. A combinatorial approach for determining protease specificities: Application to interleukin-1beta converting enzyme (ICE). Chem. Biol. 4:149-155. Rawlings, N.D. and Barrett, A.J. 1994. Families of serine peptidases. Methods Enzymol. 244:19-61. Remington, S.J. 1993. Serine carboxypeptidases: A new and versatile family of enzymes. Curr. Opin. Biotechnol. 4:62-68. Rockwell, N.C. and Fuller, R.S. 2001. Direct measurement of acylenzyme hydrolysis demonstrates rate-limiting deacylation in cleavage of physiological sequences by the processing protease Kex2. Biochemistry 40:3657-3665. Schellenberger, V., Turck, C.W., and Rutter, W.J. 1994. Role of the S′ subsites in serine protease catalysis. Active site mapping of rat chymotrypsin, rat trypsin, alpha-lytic protease and cercarial protease from Schistosoma mansoni. Biochemistry 33:4251-4257. Peptidases
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Silen, J.L. and Agard, D.A. 1989. The alpha-lytic protease pro-region does not require a physical linkage to activate the protease domain in vivo. Nature 341:462-464. Silen, J.L., Frank, D., Fujishige, A., Bone, R., and Agard, D.A. 1989. Analysis of prepro-alphalytic protease expression in Escherichia coli reveals that the pro region is required for activity. J. Bacteriol. 171:1320-1325. Smith, M.M. and Shi, L. 1995. Rapid identification of highly active and selective substrates for stromelysin and matrilysin using bacteriophage peptide display libraries. J. Biol. Chem. 270:6440-6449. Stubbs, M.T. and Bode, W. 1995. The clot thickens: Clues provided by thrombin structure. Trends Biochem. Sci. 20:23-28.
INTERNET RESOURCES http://www.biochem.wustl.edu/∼protease/ index.html Rose and Di Cera serine protease home page. http://merops.iapc.bbsrc.ac.uk/ MEROPS data base. http://www.expasy.ch/cgi-bin/lists?peptidas.txt ExPASy website.
Contributed by Lizbeth Hedstrom Brandeis University Waltham, Massachusetts
Taylor, F.R., Bixler, S.A., Budman, J.I., Wen, D., Karpusas, M., Ryan, S.T., Jaworski, G.J., SafariFard, A., Pollard, S., and Whitty, A.A. 1999. Induced fit activation mechanism of the exceptionally specific serine protease, complement factor D. Biochemistry 38:2849-2859.
An Overview of Serine Proteases
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Over-Expression and Purification of Active Serine Proteases and Their Variants from Escherichia coli Inclusion Bodies
UNIT 21.11
Heterologous expression of recombinant proteins in E. coli often results in the formation of insoluble and inactive protein aggregates commonly referred to as inclusion bodies. Inclusion bodies are normally formed in the cytoplasm. Alternatively, if a secretion vector is used, they can also form in the periplasmic space. To obtain the native, correctly folded, hence, active form of the protein from such aggregates, four steps are usually performed. (1) The bacterial cells are lysed and the aggregates, heavily contaminated with E. coli cell wall and outer membrane components, are collected by centrifugation. (2) The contaminants are largely removed by selective extraction with detergents, salts, and low concentrations of either urea or guanidinium⋅HCl to produce so-called washed pellets. (3) The aggregates are solubilized or extracted with strong protein denaturants such as guanidinium⋅HCl or urea and an oxido/shuffling system based on oxidized (GSSG) and reduced glutathione (GSH). Extraction with a denaturant simultaneously dissociates protein-protein interactions and unfolds the protein. Redox buffers facilitate the later oxidation and refolding of the protein through thiol/disulfide exchange reactions. Normally, GSSG/GSH ratios of 5:1 to 10:1 are used, which are similar to those found in vivo in the endoplasmic reticulum. As a result, the extracted protein consists ideally of unfolded monomers with sulfhydryl groups, if present, in the reduced state. If the purity of the inclusion body preparation is poor, some purification prior to folding is recommended since folding a pure or partially purified protein leads to higher yields of folded protein. (4) To obtain the native protein, it must be folded into the correct tertiary and quartenary structure to acquire its functional and biological properties. Thus, protein folding and disulfide bond formation (“oxidation”) must both occur in a concomitant manner. Co-solvents are included in the refolding mixture to maintain solubility during folding and to avoid or minimize aggregation. These basic steps result in a significant purification of the recombinant protein. However, if necessary, the folded protein is further purified. Gel-filtration chromatography is a useful final step for removing aggregates or misfolded protein. This unit describes the over-expression and purification of active serine proteases and their variants from Escherichia coli inclusion bodies. The challenge is to purify, solubilize, and fold the recombinant protein into its native and biologically active conformation in sufficiently high amounts (i.e., milligram range) using simple purification schemes. The strategy includes the folding and purification of a stable zymogen precursor protein, and its later activation with the appropriate convertase to the less stable but active protease. It should be emphasized that although most of the protocols described in this chapter are applied to a specific example, namely the purification of microplasmin to homogeneity, they are fairly representative of the methods and approaches used in general for laboratory-scale preparation of other recombinant serine proteases. The critical steps and how this template protocol can be adapted for the purification of other serine proteases are described (see Critical Parameters and Troubleshooting). The individual characteristics of a given protein (e.g., solubility, isoelectric point, pH stability, and the number of sulfhydryl residues) will dictate the conditions required for any particular folding process or purification step. Method optimization is usually determined empirically.
Peptidases Contributed by Marina A. A. Parry Current Protocols in Protein Science (2002) 21.11.1-21.11.19 Copyright © 2002 by John Wiley & Sons, Inc.
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Preparation of a cell pellet, cell lysis, and purification of inclusion bodies from E. coli cultures are presented (see Basic Protocol 1). Basic Protocol 1 also describes a method to resolubilize the protein in guanidinium⋅HCl, to refold and convert it into its native conformation. Basic Protocol 2 details the activation of the zymogen microplasminogen to the catalytically active microplasmin and provides an affinity chromatography method to purify microplasmin to homogeneity. Support Protocols 1 and 2 describe assays to characterize the purified recombinant microplasmin. A test to follow the presence of activity in the samples (see Support Protocol 1), together with an active-site titration protocol to determine the number of active-sites per mole of total protein (see Support Protocol 2) are provided. NOTE: Some of the protocols described in this unit may be covered by patent rights. In particular, nonacademic or commercial users should consult the patent literature before using these protocols. Research directed to commercial use is considered to be an infringement of relevant patent rights. Any other type of research does not infringe on existing patent rights. BASIC PROTOCOL 1
OVER-EXPRESSION AND PURIFICATION OF MICROPLASMINOGEN AND ITS VARIANTS FROM ESCHERICHIA COLI INCLUSION BODIES The high-level expression of microplasminogen results in the formation of inclusion bodies, which consist of insoluble protein aggregates. To obtain highly purified and functional microplasminogen, the E. coli cells are first lysed with a French press and the inclusion bodies are purified from soluble and particulate bacterial cell contaminants using a selective extraction procedure (UNIT 6.3). Subsequent washing and centrifugation steps in the presence of high salt and Triton X-100 remove contaminating bacterial proteins. The inclusion bodies are then solubilized in guanidinium⋅HCl and a mixture of reduced and oxidized glutathione to produce fully unfolded and reduced protein. In subsequent steps, the protein is simultaneously folded and oxidized by the removal of protein denaturant in the presence of co-solvents, which facilitate appropriate disulfide bond formation and minimize the formation of “sticky” folding intermediates (see UNITS 6.4 & 6.5). This protocol yields highly enriched and native microplasminogen. Materials 2× TYkan/amp medium (see recipe), 37°C Glycerol stock of cells harboring the construct of interest 500 mM (250×) isopropyl-β-D-thiogalactopyranosidase (IPTG; see recipe) Lysis buffer (see recipe) Lysozyme stock (see recipe) 1 M MgCl2 stock (APPENDIX 2E) 0.2 mg/ml DNase I stock (see recipe) Wash buffer 1 (see recipe) Wash buffer 2 (see recipe) Denaturing buffer (see recipe), freshly made 1 M HCl Dialysis buffer 1 (see recipe), 4°C Refolding buffer (see recipe) Dialysis buffer 2 (see recipe), 4°C
Expression and Purification of Serine Proteases
15-ml culture tubes 1, 2-, and 3-liter baffled culture flasks French press (Spectronic Instruments) or sonicator (Sonics & Materials)
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Tissue homogenizer (e.g., Potter-Elvejhem, Braun Biotech) End-over-end rotator pH-indicator paper 0 to 14 or 4 to 7 (Merck) Dialysis tubing, ∼3.2-cm diameter (Spectrum Laboratories) 0.4-µm nitrocellulose membrane cut to fit vacuum filtration apparatus (Schleicher & Schuell) or 0.7-µm glass microfiber filters (Whatman), optional Tangential flow ultrafiltration equipment with membrane (Pall Filtron) Additional reagents and equipment for absorbance measurements (OD600 and OD280; UNIT 3.1), centrifugation (UNIT 4.1), dialysis (APPENDIX 3B), chromatography (Chapter 8), and SDS-PAGE (UNIT 10.1) Inoculate overnight cultures, induce, and prepare cell pellet 1. For a 6-liter culture, inoculate two tubes of 5 ml 2× TYkan/amp medium with 100 µl of a glycerol stock of the corresponding bacteria in 15-ml culture tubes and grow for 3 to 5 hr at 150 rpm in a 37°C incubator shaker. NOTE: Glycerol stocks should be made from an uninduced culture grown to OD600 0.5 with 10% glycerol and stored at −80°C. Typically, the cultures are grown in 15-ml sterile falcon tubes, where the lid is not completely closed, to facilitate aeration.
2. Transfer both cultures into a 1-liter baffled culture flask with 300 ml of 2× TYkan/amp and incubate overnight at 37°C with shaking. Typically, a baffled shaker flask is filled to 1⁄3 of its total volume to avoid spilling and contamination of the shaking incubator.
3. Distribute the overnight culture into six 2- to 3-liter baffled culture shaker flasks, each containing 1 liter of 2× TYkan/amp and continue shaking until absorbance at 600 nm is 0.5 to 0.8 (∼90 min). 4. Add to each culture flask 4 ml of 500 mM IPTG to 2 mM final concentration. Grow for an additional 4 to 6 hr in a 37°C incubator with shaking. Addition of IPTG to 2 mM final concentration induces the expression of the desired protein.
5. Harvest the cell pellet by centrifuging 20 min at 6000 × g, 4°C. Discard the supernatant. The cell pellet can be stored in polyethylene bags as a flattened paste for several months at −80°C.
Lyse bacterial cells and extract inclusion bodies 6. Resuspend the pellet in 75 ml lysis buffer. Add lysozyme to 0.25 mg/ml (1:10 dilution from stock) and incubate for 30 min to 1 hr on ice. 7. Disrupt cells in a French press by ∼1000 psig (three cycles; UNIT 6.2). Freezing and thawing is not a recommended cell lysis method because of its relatively low efficiency. However, cell lysis can be also accomplished by sonication on ice. The extent of lysis can be assessed by examining the morphology of the cells under a light microscope. An aggregated appearance indicates that the cells are adequately disrupted, whereas the presence of numerous singular rod shaped cells indicates that further sonication is required.
8. Add 150 µl of 1 M MgCl2 (2 mM final concentration) and 3.75 ml of 0.2 mg/ml DNase I (10 µg/ml final concentration). Incubate 30 min at 25°C to digest the DNA. Alternatively, 750 ìg DNase I can be added to the lysis mixture as a powder.
Peptidases
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9. Centrifuge the lysate for 30 min at 20,000 × g, 4°C. Discard the soluble fraction and continue with the insoluble pellet. The pellet can be stored for several months at −20°C at this stage.
Purify inclusion bodies to remove contaminating E. coli proteins 10. Resuspend the pellet with the help of a tissue homogenizer in 38 ml of wash buffer 1. Incubate for an additional 30 min at room temperature with end-over-end rotation. 11. Recover the inclusion bodies by centrifuging for 30 min at 20,000 × g, 4°C. Discard the supernatant. 12. Resuspend the pellet in 38 ml wash buffer 2 with the help of a tissue homogenizer for 5 min. Rotate the suspension end-over-end for 15 min. 13. Centrifuge for 30 min at 20,000 × g, 4°C. Discard the supernatant. 14. Repeat steps 12 and 13. 15. Weigh out the purified inclusion bodies to estimate the yield and compare performances of different preparations. The pellet can be stored for several months at −20°C at this stage.
Denature inclusion bodies in denaturing buffer 16. Solubilize the inclusion bodies with the help of a tissue homogenizer in 24 ml of denaturing buffer and rotate end-over-end overnight. This step should be carried out carefully since excess oxygen could oxidize the denatured protein and impair correct refolding.
17. Using pH-indicator paper, adjust the pH to 5.0 with 1 M HCl and centrifuge for 30 min at 20,000 × g, 4°C. Repeat step 16. 18. Dialyze the solubilized protein in dialysis tubing (∼3.2-cm diameter) for a 24-hr period against 100 vol of dialysis buffer 1 at 4°C, with three buffer changes every ∼8 hr. Fill dialysis bags only 3⁄4 of the way to allow for any volume increase during dialysis.
19. Remove eventual precipitates by centrifuging 30 min at 20,000 × g, 4°C. 20. Read absorbance at 280 nm and run a sample on an SDS-PAGE gel (UNIT 10.1) to estimate the total amount and the purity of the protein in the preparation. Because guanidium⋅HCl forms a precipitate with SDS, it is necessary to remove the former before carrying out SDS-PAGE. For a detailed protocol, see Preparation of Samples Containing Guanidine Hydrochloride for SDS-PAGE (UNIT 6.3, Support Protocol).
Refold and concentrate protein 21. Determine the total volume of the dialyzed material and divide it into three equal aliquots. The total volume of the dialyzed material is typically ∼50 ml for a 6-liter culture preparation. Thus, the three aliquots are typically ∼17 ml each.
22. Initiate refolding by diluting one of the three aliquots in 100 vol of refolding buffer. Incubate 8 to 24 hr at room temperature. Keep the other two aliquots at 4°C until use. Expression and Purification of Serine Proteases
Typically, the total volume of refolding buffer is ∼1.7 liters. To achieve a rapid dilution and to assist proper refolding, the dilution is done drop-wise while carefully rotating the flask.
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23. Add the second aliquot to the refolding mixture and incubate an additional 8 to 24 hr at room temperature. 24. Add the third aliquot to the refolding mixture and incubate an additional 8 to 24 hr at room temperature. 25. Centrifuge the refolding mixture for 90 min at 6000 × g at room temperature. Flaky precipitates that are difficult to remove by centrifugation can be filtered through a 0.4-ìm nitrocellulose membrane or a 0.7-ìm glass microfiber filter.
26. Concentrate the solution in a tangential flow ultrafiltration concentrator until the absorbance at 280 nm is ∼1.0 (∼1 mg/ml). 27. Transfer the concentrate to a dialysis membrane and dialyze the concentrated sample for a 24-hr period at 4°C against 50 vol of dialysis buffer 2, with three buffer changes every ∼8 hr. Fill dialysis membranes only 3⁄4 of the way to allow for any volume increase during dialysis.
28. Remove precipitated proteins by centrifugation for 30 min at 20,000 × g, 4°C. Determine the OD 280 nm against the dialysis buffer to estimate the protein concentration and the yield of the preparation. The preparation can be stored up to six months at −80°C.
29. If the preparation shows only one band by SDS-PAGE, do not perform any further purification (Figure 21.11.1). In this case, no further purification steps were required since the aim of the preparation was to obtain the active serine protease and not the inactive zymogen. Typical criteria to decide whether to embark on further purification steps are, e.g., the presence of more than one band by SDS-PAGE (UNIT 10.1; Coomassie Blue or silver staining) and the presence of aggregates as detected by monodispersity measurements or native PAGE gels. The latter techniques allow one to follow the refolding efficiency.
A
B
Figure 21.11.1 (A) Refolded microplasminogen purified from inclusion bodies. The preparation showed only one band by SDS-PAGE, and thus, no further purification steps were required. Complete bacterial cell lysis, as well as careful washing of the insoluble inclusion bodies, were critical steps in this preparation. (B) Affinity purified microplasmin. The preparation showed only one band on a silver stained SDS-PAGE gel.
Peptidases
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BASIC PROTOCOL 2
ACTIVATION OF THE ZYMOGEN MICROPLASMINOGEN AND PURIFICATION OF THE CATALYTICALLY ACTIVE MICROPLASMIN The refolded microplasminogen protein must be proteolytically activated to purify it by affinity chromatography. The microplasminogen activator urokinase (uPA), also a serine protease, cleaves microplasminogen specifically at the Arg15-Val16 bond, leaving the cleaved propeptide attached to the catalytically active domain by disulfide bridges. Concomitantly to the activation cleavage, a conformational change takes place, leading to an active-site exposure in microplasmin (Parry et al., 2000; Wang et al., 2000). For convenience, the activating convertase (in this case urokinase) is covalently coupled to CNBr-activated Sepharose 4B through free amino groups. In this way, the urokinaseSepharose beads can be used repeatedly and can be easily removed by centrifugation. Cleavage of the zymogen propeptide and conversion of microplasminogen to microplasmin can be monitored by reduced SDS-PAGE at defined time intervals and by a suitable chromogenic assay (see Support Protocols 1 and 2; Figure 21.11.2). The final affinity purification step makes use of the newly exposed active-site and separates microplasmin from all impurities including nonactivated microplasminogen. The positively charged serine protease inhibitor benzamidine specifically binds to the negatively charged carboxylate group of Asp194 of microplasmin. Elution with acetic acid, pH 3, leads to protonation of Asp194 and subsequent dissociation of the microplasmin–benzamidine complex. This protocol yields highly active and pure microplasmin. Materials Urokinase plasminogen activator (uPA; Sigma) CNBr-activated Sepharose 4B (Amersham Pharmacia) Recombinant microplasminogen protein (see Basic Protocol 1) 5 M NaCl (APPENDIX 2E) 87% glycerol Urokinase-Sepharose storage buffer (see recipe), optional Benzamidine-Sepharose column (Sigma) Wash buffer for benzamidine-Sepharose column (see recipe) Elution buffer 1 or 2 for benzamidine-Sepharose column (see recipe) Regeneration buffer for benzamidine-Sepharose column (see recipe) End-over-end rotator Additional reagents and equipment for absorbance measurements (UNIT 3.1), centrifugation (UNIT 4.1), dialysis (APPENDIX 3B), chromatography (Chapter 8), and SDS-PAGE (UNIT 10.1) Activate recombinant protein 1. Immobilize urokinase plasminogen activator (uPA) on CNBr-activated Sepharose 4B (UNIT 9.3). The uPA-Sepharose beads can be used repeatedly and are easily removed by centrifugation (see below).
2. Prepare the recombinant microplasminogen protein for activation. Add to a ∼1 mg/ml recombinant microplasminogen protein solution 1/10 vol of 5 M NaCl and 87% glycerol to 25% final concentration. Expression and Purification of Serine Proteases
Typically, 0.21 ml of 5 M NaCl and 8.4 ml of 87% glycerol are added to 21 ml microplasminogen at ∼1 mg/ml. The addition of 25% glycerol prevents microplasmin autoproteolysis upon activation.
21.11.6 Supplement 27
Current Protocols in Protein Science
c TYTX
ef XTg
TYTW
TYTV
TYTU
TYTT T
UT
VT Z[\]^_`ab
d
T
TYg
V
X
WT
h
XT
i
UV
jklmnop[qk^p[\]^_`ab
Figure 21.11.2 Time course of microplasminogen activation. (A) The activity increases in a hyperbolic fashion towards a limiting value. (B) On a reduced SDS-PAGE gel, only one band is present at t = 0, a lower molecular weight band increasingly appears with incubation time. These results correlate with the disappearance of microplasminogen and the appearance of catalytically active microplasmin domain. Under reducing conditions, the disulfide-linked propeptide of microplasminogen (∼2 kDa) dissociates from the catalytically active microplasmin.
3. Activate 1 volume of this activation mixture by adding 1⁄10 vol of uPA-Sepharose beads. Incubate overnight at room temperature with gentle rotation to resuspend the beads. Pilot-scale experiments are usually necessary to optimize the conditions for removing the propeptide using urokinase. The typical parameters to vary are enzyme/substrate ratios and incubation time and temperature.
4. Monitor the removal of the zymogen propeptide by a suitable chromogenic assay (see Support Protocols 1 and 2) and by reduced SDS-PAGE at defined time intervals (Figure 21.11.2). 5. Separate the uPA-Sepharose beads by centrifugation for 15 min at 400 × g, 4°C, to stop the activation reaction. After use, perform a centrigufation wash on the uPA-Sepharose with urokinas-Sepharose storage buffer. The uPA-Sepharose beads can be stored for several months as a 50% slurry at 4°C in urokinase-Sepharose storage buffer.
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Purify on a benzamidine-Sepharose column 6. Apply the supernatant containing the activated protease to a benzamidine-Sepharose column equilibrated with benzamidine-Sepharose column wash buffer. Because of the high glycerol content of the activation mixture, the flow rate should not exceed 0.2 ml/min. Alternatively, the affinity chromatography can be carried out by binding the protein to the resin in a batch procedure and by washing and eluting the benzamidineSepharose on a sintered glass filter.
7. Wash the benzamidine-Sepharose column with two column volumes of benzamidineSepharose column wash buffer. 8. Elute with one column volume of benzamidine-Sepharose column elution buffer 1. Collect the eluate until baseline is reached by reading absorbance at 280 nm. Alternatively, the column can be eluted with benzamidine-Sepharose column elution buffer 2. Typical flow rates are ∼1 ml/min. Microplasmin is very unstable and autodigests within hours to minutes at a neutral pH. Thus, microplasmin should be inactivated prior to storage. Elution buffers 1 and 2 provide two alternatives to inactive microplamin while eluting it from the affinity column. Elution buffer 1 protonates the active site of microplasmin (pH 3), and this renders the protease inactive. Elution buffer 2 contains a serine protease inhibitor (benzamidine) that reversibly blocks microplamin’s active site. The choice of buffer depends on the later use of the preparation.
9. Regenerate the column in benzamidine-Sepharose column regeneration buffer until the pH of the flow-through equals that of the running buffer. The total amount of purified protein and its concentration can be measured by SDS-PAGE (UNIT 10.1), OD280 (UNIT 3.1), or Bradford assay (UNIT 3.4). SUPPORT PROTOCOL 1
ASSAY TO DETERMINE THE ACTIVITY OF THE RECOMBINANT MICROPLASMIN This protocol describes an assay to follow the presence of microplasmin activity. The catalytic activity of microplasmin is assayed in a chromogenic assay using a colorless substrate (H-D-valyl-leucyl-L-lysine-p-nitroaniline), which is hydrolyzed by microplasmin to release yellow p-nitroaniline (Castellino and Powell, 1981; Wu et al., 1987). The increase in absorbance at 405 nm is proportional to the amount of active microplasmin in the assay. The measurements should take place in the “linear range” where ∼10% of the total substrate concentration is being hydrolyzed. Materials S-2251 chromogenic substrate (Chromogenix) Assay buffer (see recipe) 1.5-ml plastic cuvettes (Amersham Pharmacia) Additional reagents and equipment for absorbance measurements (UNIT 3.1) 1. Prepare a stock solution of 3 to 4 mM S-2251 chromogenic substrate by dissolving it in Milli-Q-purified water. Read absorbance at 316 nm and determine the substrate concentration using the extinction coefficient, ε = 1.27 × 104 mol−1 cm−1. 2. Prepare the assay buffer. A larger stock can be prepared in advance since the assay buffer can be frozen in 10-ml aliquots for several months at −20°C until use.
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3. Prepare a substrate-buffer mix by diluting the S-2251 chromogenic substrate stock solution to a final concentration of 200 µM in assay buffer. 4. Dispense 800-µl aliquots of the substrate-buffer mix into 1.5-ml plastic cuvettes. Place them in a spectrophotometer prepared for readings at 405 nm and set absorbance to baseline (autozero). 5. Start the reaction by adding 1 to 5 µl of samples containing microplasmin and stir. The samples should be prepared and measured in duplicates or triplicates.
6. Immediately follow the increase in absorbance at 405 nm over time (5 to 10 min). The slope of the graph (dA/min) divided by the volume of sample added to the assay (µl) is a relative measurement of the microplasmin activity in the solution. The absorbance should increase in a linear fashion over time (“initial rate,” “linear range”), e.g., the substrate turnover should not exceed 10% of its total concentration. In this particular case, a range up to ∼0.6 OD units at 405 nm assures working in the linear range.
ASSAY TO DETERMINE THE ACTIVE-SITE CONCENTRATION The actual active-site concentration of microplasmin is determined in the presence of increasing amounts of bovine pancreatic trypsin inhibitor (BPTI) in the chromogenic assay described above. By extrapolation, the inhibitor concentration necessary to achieve a 100% microplasmin inhibition is calculated. This inhibitor concentration corresponds to the concentration of active-sites of microplasmin on a 1:1 molar ratio.
SUPPORT PROTOCOL 2
Materials Substrate-buffer mix (see Support Protocol 1, steps 1 to 3) 0.5 µM protease inhibitor (bovine pancreatic trypsin inhibitor, BPTI) 0.1 M acetic acid, pH 3.0 1.5-ml plastic cuvettes Additional reagents and equipment for amino acid analysis (UNIT 11.9), determination of protein concentration (UNIT 3.1), Lowry or Bradford methods (UNIT 3.4) 1. Prepare a substrate-buffer mix as described in Support Protocol 1, steps 1 to 3. 2. Prepare a 0.5 µM solution of protease inhibitor, BPTI, in water. Determine its accurate concentration preferably by amino acid analysis (UNIT 11.9) in triplicate. 3. Measure the protein (enzyme) concentration by OD280 (UNIT 3.1) or by the Lowry or Bradford methods (UNIT 3.4). Using this information, dilute the microplasmin solution in 0.1 M acetic acid, pH 3.0, to a final concentration of 5.5 µM. Microplasmin is inactive in acetic acid, pH 3.0, and remains stable under these experimental conditions until use. Microplasmin is stable for several weeks at an acidic pH.
4. Prepare a range of enzyme-inhibitor complexes as shown in Table 21.11.1. Namely, keep a constant enzyme concentration and increase BPTI concentrations with the addition of assay buffer to a final volume of 100 µl. Keep the tubes on ice. It is preferable to prepare samples in duplicate or triplicate to increase the accuracy of the measurement.
5. Pipet 800-µl aliquots of buffer-substrate mix into 1.5-ml plastic cuvettes, place them in a spectrophotometer prepared for readings at 405 nm and set absorbance to baseline (autozero).
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Table 21.11.1
Pipetting Scheme for the Active-Site Titration of Microplasmin
Tube number
Microplasmin volume (µl)
Microplasmin concentration (pmol)
BPTI volume (µl)
BPTI Assay buffer concentration (µl) (pmol)
Molar ratio (BPTI/microplasmin)
1 2 3 4 5 6
10 10 10 10 10 10
55 55 55 55 55 55
76 48 36 24 12 0
38 24 18 12 6 0
0.69 0.44 0.33 0.22 0.11 0
A
14 42 54 66 78 90
Temperature N/A
0.900
5
6
4 3
Absorbance
2
1 –0.05 0.00
Time (sec)
300
B 0.22 0.20 0.18
Absorbance/time
0.16 0.14 0.12 0.10 0.08 0.06 0.04 0.02 0.00 0.0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
1.0
1.1
BPTI:microplasmin
Expression and Purification of Serine Proteases
21.11.10 Supplement 27
Figure 21.11.3 Active-site titration with increasing BPTI concentrations. (A) Initial rates decrease with increasing inhibitor concentrations. Measurements are carried out in the linear range. Tubes no. 1 through 6 indicated from Table 21.11.1. (B) Active-site titration of microplasmin. The slopes (∆ Absorbance/time) obtained in A are plotted against the BPTI/microplasmin molar ratio. The specific activity of the purified protein was calculated as the percentage of active sites per moles of total protein and was typically shown to be >95% active. Current Protocols in Protein Science
6. Add the 100 µl enzyme-inhibitor complexes prepared above, mix, and immediately follow the increase in activity for 5 to 10 min. Calculate the slopes of the graph in dA/min. 7. Plot the slopes (initial rates versus the inhibitor concentrations or the inhibitor/enzyme molar ratios; Table 21.11.1, Figure 21.11.3). Draw a straight line through the linear part of the plot and extrapolate on the x-axis, the inhibitor concentration necessary to achieve 100% inhibition. This inhibitor concentration corresponds to the active microplasmin concentration on a 1:1 molar basis (Figure 21.11.3). Only data points that decrease linearly with the increasing inhibitor concentration should be used for data evaluation. Data points that asymptotically approach the x-axis should be left out, since they are affected by the dissociation constant of the complex.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Assay buffer 0.1 M HEPES, pH 7.4 0.1 M NaCl (APPENDIX 2E for 5 M) 1 mM EDTA (APPENDIX 2E for 0.5 M) 1 mg/ml PEG 4000 Store up to 6 months, −20°C Denaturing buffer 6 M guanidinium⋅HCl 100 mM Tris⋅Cl, pH 8.2 (APPENDIX 2E for 1 M) 20 mM EDTA (APPENDIX 2E for 0.5 M) 150 mM oxidized glutathione 15 mM reduced glutathione Prepare on day of use. Dialysis buffer 1 6 M guanidinium⋅HCl, pH 5.0 20 mM EDTA (APPENDIX 2E for 0.5 M) Prepare just before use. Dialysis buffer 2 10 mM Tris⋅Cl, pH 7.2 (APPENDIX 2E for 1 M) 100 mM NaCl (APPENDIX 2E for 5 M) Store up to several weeks, 4°C DNase I stock, 20× 0.2 mg/ml DNase I from bovine pancreas (Roche Molecular) in 50 mM Tris⋅Cl, pH 7.2 (APPENDIX 2E for 1 M) Filter sterilize using a 0.22-µm filter Dispense into 2-ml aliquots, store up to 6 months, −20°C Elution buffer 1 for benzamidine-Sepharose column 0.1 M acetic acid, pH 3.0 Store up to several weeks at room temperature Elution buffer 2 for benzamidine-Sepharose column 5 mM Tris⋅Cl, pH 7.0 (APPENDIX 2E for 1 M) 100 mM NaCl (APPENDIX 2E for 5 M) 50 mM benzamidine Store up to several weeks, room temperature Peptidases
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IPTG stock, 250× 6 g isopropyl-β-D-thiogalactopyranosidase (IPTG) with water to 50 ml (final 500 mM) Filter sterilize using a 0.22-µm filter, dispense into 2-ml aliquots, and store up to 6 months, −20°C Lysis buffer 50 mM Tris⋅Cl, pH 7.2 (APPENDIX 2E) Lysozyme stock, 10× 2.5 mg/ml chicken eggwhite lysozyme (Sigma) in 50 mM Tris⋅Cl, pH 7.2 (APPENDIX 2E) Dispense into 2-ml aliquots and store up to 6 months, −20°C Refolding buffer 50 mM Tris⋅Cl, pH 8.5 (APPENDIX 2E for 1 M) 0.5 M arginine 1 mM EDTA (APPENDIX 2E for 0.5 M) 50 mM CaCl2 0.5 mM cysteine 100 mM NaCl (APPENDIX 2E for 5 M) 1% PEG 4000 Prepare on day of use Regeneration buffer for benzamidine-Sepharose column 100 mM Tris⋅Cl, pH 8.5 (APPENDIX 2E for 1 M) 500 mM NaCl (APPENDIX 2E for 5 M) Store up to several weeks, 4°C TY medium, 2× 16 g tryptone (final 2% w/v) 10 g yeast extract (final 1% w/v) 5 g NaCl (final 0.5% w/v) H2O to 1 liter Autoclave immediately after preparation. Store for several weeks, tightly closed, at 4°C. TYkan/amp medium, 2× 1 liter 2× TY medium (see recipe) 100 mg ampicillin (final 100 µg/ml) 50 mg kanamycin (final 50 µg/ml) Store up to several weeks, tightly closed, at 4°C Urokinase-Sepharose storage buffer 100 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E for 1 M) 500 mM NaCl (APPENDIX 2E for 5 M) Store up to 6 months, 4°C
Expression and Purification of Serine Proteases
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Wash buffer 1 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E for 1 M) 60 mM EDTA (APPENDIX 2E for 0.5 M) 1.5 M NaCl (APPENDIX 2E for 5 M) 6% Triton X-100 Store up to several weeks at room temperature Wash buffer 2 50 mM Tris⋅Cl, pH 7.2 (APPENDIX 2E for 1 M) 60 mM EDTA (APPENDIX 2E for 0.5 M) Store up to several weeks at room temperature Wash buffer for benzamidine-Sepharose column 5 mM Tris⋅Cl, pH 7.0 (APPENDIX 2E for 1 M) 100 mM NaCl (APPENDIX 2E for 5 M) Store up to several weeks, 4°C COMMENTARY Background Information In the process of fibrinolysis, as well as in coagulation, anticoagulation, and the complement system, serine proteases play crucial roles (Bachman, 1994). They work in amplification cascades where an active serine protease activates another proproteinase (Davie et al., 1991). These cascades offer a number of regulatory steps, where cofactors play crucial roles in enhancing the substrate presentation to the enzyme and modulating their specificity towards substrates and inhibitors, making a “specificity-switch” possible (Parry et al., 2000). More recently, serine proteases have been reported to play roles in tissue remodeling, asthma, and also viral and bacterial infections (Chapman, 1997; Lottenberg, 1997). The overall structure of serine proteases shows that they are folded into two six-stranded β-barrels that are thought to have evolved by gene duplication. The active-site cleft and the catalytic residues are located at the junction of both barrels. Their surface is made up of various turn structures and loops, and two short α-helices (Bode and Schwager, 1975). The surface loops around the active-site cleft provide the amino acid side chains that contribute to the formation of the S3-S′3 subsites, and thus, determine the protease specificity (Hedstrom et al., 1992). Plasmin is a key serine protease in the fibrinolytic system. Its activation from the zymogen plasminogen results in the dissolution of blood clots, and also promotes pericellular proteolysis, cell migration, and tissue remodeling (Bachman, 1994; Chapman, 1997). Interestingly, pathogenic cells recruit plasmin proteolytic activity to their cell surface to facilitate
cell invasion and migration through tissue layers (Murakami et al., 2001). The surface-bound plasmin is resistant to inhibition. Plasminogen is a modular protein that contains a pre-activation peptide (PAP) followed by five kringle domains and a catalytic C-terminal serine proteinase domain (Sottrup-Jensen et al., 1975). The kringle domains of plasmin(ogen) contain lysine binding sites that mediate its association to fibrin and cellular surfaces. In this way, the catalytic activity of plasmin is directed and localized. The physiological plasminogen activators, tissue plasminogen activator (tPA) and urokinase plasminogen activator (uPA), activate plasminogen by limited proteolysis at the Arg15(561)-Val16(562) bond (nomenclature based on the topological equivalence to chymotrypsinogen, with the plasminogen numbering given in parenthesis; Madison and Sambrook, 1993; Ke et al., 1997). The site of activation cleavage to yield plasmin divides the plasminogen molecule into the N-terminal socalled A-chain and the C-terminal proteolytically active B-chain. Microplasmin and microplasminogen are truncated forms of plasmin and plasminogen consisting of a short 20-residue A-chain connected by two disulfide bridges to the catalytically active 229-residue B-chain (Parry et al., 1998; Wang et al., 1998). In this unit, the protocols for the over-expression and purification of active serine proteases and their variants from Escherichia coli inclusion bodies are described. The strategy includes the folding and purification of a stable zymogen precursor protein, and its later activation to the less stable but active protease with the appropriate convertase. The methods are
Peptidases
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robust and reproducible, and enable the purification of a protease to homogeneity in high yields. Using protocols based on the one described here, at least eight different serine proteases and also mutants thereof, were refolded and purified successfully (Kopetzki et al., 1993; Hopfner et al., 1997; Parry et al., 1998; Wilharm et al., 1999). In Basic Protocol 1, the over-expression and purification of microplasminogen and their variants from Escherichia coli inclusion bodies is described. Basic protocol 2 describes the activation of the zymogen microplasminogen and the purification of the catalytically active microplasmin. Support Protocols 1 and 2 provide methods for following the protease activity and determining its active site concentration. Critical Parameters and Troubleshooting points out the critical steps and explains how this template protocol can be optimized for the purification of other serine proteases.
Critical Parameters and Troubleshooting Bacterial culture conditions and inclusion body formation Inclusion bodies can be easily observed by phase-contrast microscopy at high magnification (1000×) as dense bodies usually located at the polar extremities of the cell (Figure 21.11.4; Seckler and Jaenicke, 1992; Oberg et al., 1994;). The formation of inclusion bodies can occasionally protect proteins against proteolysis and can also allow accumulation of proteins normally toxic to the cell. It also simplifies the purification of the protein albeit in a denatured/aggregated state. The protein must then be refolded into a native-like conformation. The formation of inclusion bodies is not an intrinsic property of a particular protein and can sometimes be forced. Successful approaches include: changing the host strain (e.g., BL21
A
B
Expression and Purification of Serine Proteases
Figure 21.11.4 Escherichia coli culture observed by phase-contrast microscopy. (A) Bacteria that grew without IPTG addition. (B) Bacteria that grew with the addition of IPTG to induce protein expression. Inclusion bodies can be easily seen as dense bodies located at the polar extremities of the cells.
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Current Protocols in Protein Science
(DE3), B834 (DE3), UT5600), adding glucose (2%) to the medium to repress a leaky promotor, optimizing the IPTG concentration necessary for induction, changing the temperature of induction (usually increasing it), changing the E. coli promotor to induce a stronger protein expression (T7 promotor), controlling the growth conditions by using a different medium (Luria Bertani broth versus the richer 2× TY medium), optimizing the time course of protein expression (3 to 4 hr, 4 to 6 hr, or even overnight), and combinations thereof. Moreover, the recombinant protein may be located in both the insoluble and soluble fractions of the cell (mixed-phase expression). In these cases, the choice of the purification strategy must take into account the expected yield and the purpose of preparation. Bacterial cell lysis Once it has been established that the recombinant protein is insoluble (UNIT 6.1), extra care should be taken to ensure complete cell lysis (UNITS 6.2 & 6.3). If unbroken cells are present in the insoluble fraction, they will leak contaminants when the pellets are extracted with strong protein denaturants.
18
Absorbance/time
15
Treatment of the cells with 0.25 mg/ml lysozyme prior to homogenization will improve the efficiency of cell lysis (UNIT 6.5). Homogenization by freezing and thawing is not recommended. However, lysis by a French press or sonication on ice usually gives good results. Proper operation of the French press and sonicator is described in UNIT 6.2. The extent of lysis can be assessed by examining the morphology of the cells under a light microscope. Addition of MgCl2 to a final concentration of 2 mM and DNase I to a final concentration of 10 µg/ml reduces the viscosity of the lysate, and facilitates the downstream handling of the sample (Hopfner et al., 1997; Parry et al., 1998; Wilharm et al., 1999). The choice of cell lysis buffer is not critical, however, the buffer pH should be above 6.2 as many soluble E. coli proteins precipitate at slightly acidic pH values. Inclusion body purification Other major sources of contamination are outer-membrane and cell-wall material, most of which can be pre-extracted with detergents such as Triton X-100 (0.5% to 5%), with high salt (1.5 M NaCl), with urea (1 to 4 M), or
18
18
A
15
B
15
12
12
12
9
9
9
6
6
6
3
3
3
0
0
7
Tris Tris/arg Pi Tris/PEG Buffer
8.5 PH
9
10
18
18 15
8
D
15
RT
0
15 12
9
9
9
6
6
6
3
3
3
4 C
0
0.5 0.7 0.9 Arginine (mM)
F
0
0
0.1 0 NaCl (mM)
0.3
18
E
12
12
C
0
10 20 30 CaCl 2 (mM)
40
0.5 1 1.5 2 Cysteine (mM)
Figure 21.11.5 Effect of different co-solvents in the refolding efficiency of microplasminogen. (A) Buffer composition; Tris alone (1M), with addition of arginine (Arg) or PEG4000, phosphate buffer (Pi). (B) Buffer pH. (C) Arginine concentration. (D) Salt and temperature. (E) CaCl2 concentration. (F) Cysteine concentration.
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21.11.15 Current Protocols in Protein Science
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combinations thereof (UNIT 6.3; Kopetzki et al., 1993; Hopfner et al., 1997; Parry et al., 1998; Wilharm et al., 1999). After pre-extraction, the aggregates should be washed (two times) with buffer alone to remove excess detergent and salt. When attempting to develop a reproducible folding protocol, varying amounts of detergent carry-over may influence the outcome. To obtain a complete dissolution of the pellets and to improve the effectiveness of the extraction and washing steps, some form of mechanical dispersion is often required. Homogenization at room temperature with a Potter-Elvejhem-style tissue homogenizer as described is often adequate and can greatly improve the purity of the preparation. Although insoluble proteins are less susceptible to proteolysis than native proteins, inclusion of EDTA (e.g., 60 mM) and possibly other protease inhibitors could prove useful. Denaturation in guanidinium⋅HCl The washed pellets containing the insoluble inclusion body protein are extracted with guanidinium⋅HCl (UNIT 6.3). To obtain complete dissolution of the pellets, the use of a PotterElvejhem-style tissue homogenizer as described above is often adequate; however, sonication should be used if the pellets are especially recalcitrant (UNIT 6.5). Heating the solution will also aid protein solubilization (10 to 15 min at 50° to 60°C). Excess heating without adequate cooling, whether direct or from the sonication, should be avoided. Derivatization of the solubilized protein is accomplished by inclusion of an oxido/shuffling system based on oxidized (GSSG) and reduced glutathione (GSH). Normally, GSSGGSH ratios of 5:1 to 10:1 are used (Kopetzki et al., 1997; Hopfner et al., 1997; Parry et al., 1998; Wilharm et al., 1999). Redox buffers facilitate the later oxidation and refolding of the protein through thiol/disulfide exchange reactions. To reduce the rate of GSH loss due to air oxidation, EDTA should be included in the buffer.
Expression and Purification of Serine Proteases
Refolding Aggregation during the folding process can be minimized by using solvent additives collectively called “co-solvents.” Presumably, the co-solvents minimize intermolecular associations between the hydrophobic and, thus, sticky folding intermediates. For theoretical and practical discussions of folding or aggregation, see UNITS 6.4 & 6.5.
Successfully used co-solvents include arginine (0.5 to 5 M), polyethylene glycol, nonionic detergents and lipids, cationic detergents, salts (NaCl, CaCl2) and nondenaturing concentrations of urea or guanidinium⋅HCl (1 to 1.5 M). Additives such as ammonium sulfate, glycerol, sucrose, ligands and enzyme substrates, inhibitors, or cofactors have also been used to enhance protein folding (Kopetzki et al., 1997; Hopfner et al., 1997; Parry et al., 1998; Wilharm et al., 1999). The optimal concentration and ratios of reagents are established empirically (Figure 21.11.5). Refolding efficiencies can be followed by activity tests and/or by detecting the presence of aggregates in the sample. Techniques such as native PAGE (UNIT 10.3) and dynamic light scattering (UNIT 7.8) prove helpful in optimizing the composition of the refolding buffer. Tangential flow concentration Refolding is typically achieved by rapid step-wise dilution to increase the yields. Depending on the scale of the preparation, this results in large volumes containing dilute refolded protein. Moreover, the presence of cosolvents often slows down the concentration procedure. In these cases, the use of tangential flow concentrators dramatically speeds up the ultrafiltration (UNIT 4.4; commercially available from Filtron). Purification of the refolded protein Careful cell lysis and inclusion body purification often proves rewarding and results in preparations of very good purity as assessed by SDS-PAGE (Figure 21.11.1). However, the presence of aggregates should be monitored by native PAGE (UNIT 10.3) or dynamic light scattering (UNIT 7.8) as discussed above (see Refolding). Further purification steps might be needed, e.g., gel filtration chromatography (UNITS 6.3 & 8.3). Depending on the protein to be purified, anion exchange chromatography (UNIT 8.2; S-Sepharose or Q-Sepharose) or affinity chromatography (e.g., heparin-agarose; UNIT 9.3) give good results. Activation of the recombinant zymogen The appropriate choice of the activating enzyme or “convertase” should rely on literature searches, availability, and price (Kopetzki et al., 1997; Hopfner et al., 1997; Parry et al., 1998; Wilharm et al., 1999). Immobilization of the convertase on Sepharose beads is advantageous, since these beads can be re-used and are
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media preparation & inoculation of overnight cultures
induction & preparation cell pellet
cell lysis washes to remove contaminating E.coli proteins denaturation in guanidinium•HCl
dialysis
ultracentrifugation to remove insoluble contaminants refolding 1
refolding 2
refolding 3
concentration
zymogen activation & active site titration
Figure 21.11.6 Overview of the steps required for over-expression, purification, and actication of a serine protease from inclusion bodies. Each block in the flow chart represents the steps taht can be done in one day.
easily separated from the preparation by centrifugation at 400 × g. First of all, pilot-scale tests need to be done to find the optimal activation conditions. Upon addition of the convertase, the activity of the preparation can be followed by a suitable chromogenic/fluorogenic assay at defined time intervals. Removal of the propeptide can also be monitored by reduced SDS-PAGE (UNIT 10.1) since there is usually a detectable mass difference between the zymogen and the active protein (Figure 21.11.2). The digestion should be as complete as possible at the specific cleavage site, but over-digestion may result in nonspecific cleavage at other sites and should be avoided. It is also important to avoid denaturation of the convertase by vigorous stirring or overheating, since partially denatured proteases often exhibit some loss of specificity and cleave at unpredictable sites The activated serine protease can then be purified by affinity chromatography over an inhibitor-Sepharose column (e.g., benzamidine-Sepharose or BPTI-Agarose; UNIT 9.3). In this way, nonactivated zymogen, released propeptide, and other contaminants can be
separated by making use of the newly exposed active-site. An active-site titration with a tight binding inhibitor of accurately known concentration should be done to finally characterize the quality of the preparation.
Anticipated Results The steps involved in the over-expression and purification of microplasmin from inclusion bodies are described in Figure 21.11.6. The refolded microplasminogen purified from inclusion bodies showed only one band by SDSPAGE (UNIT 10.1), and thus, no further purification steps were required. Complete cell lysis and careful washing (UNIT 6.3) of the insoluble inclusion bodies were crucial steps in this preparation (Figure 21.11.1). Upon incubation with urokinase-Sepharose quantitative microplasminogen activation was achieved, as assessed by SDS-PAGE (UNIT 10.1) and activity tests (Figure 21.11.2; see Support Protocol 1). Active-site titration of the purified microplasmin shows a very good specific activity of >95% (Figure 21.11.3; see Support Protocol 2). Using protocols based on the one described here, at least eight different serine proteases and
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also mutants thereof, were refolded successfully (Kopetzki et al., 1993; Hopfner et al., 1997; Parry et al., 1998; Wilharm et al., 1999). Typically, ∼0.3 g of purified inclusion bodies culture were obtained per liter. The final yields were nevertheless very variable: ten-fold differences were observed ranging from 2 mg to 22 mg of refolded material per liter culture. Time considerations The steps required for over-expression, purification, and activation of a serine protease from inclusion bodies can be done in 9 days. Basic Protocol 1 for over-expression and purification of microplasminogen and their variants from Escherichia coli inclusion bodies takes 8 days, from the glycerol stock to the concentrated and purified protein. The activation of the zymogen to the catalytically active microplasmin and its affinity purification takes ∼2 days. However, as stated in the protocol, the preparation can be interrupted at different stages, and some of the steps involve little manual handling (such as the dialysis and refolding steps). At the end of day 2, the cell paste can be frozen, as well as the cell lysate or the purified inclusion bodies at the end of day 3. When starting a new project, additional time should be set aside to optimize the culture and over-expression conditions and especially the refolding conditions, as discussed in the Troubleshooting section (Figures 21.11.4 and 21.11.5).
Literature Cited Bachman, F. 1994. Fibrinolysis. In Haemostasis and Thrombosis. (A.L. Bloom, C.D. Forbes, D.P. Thomas, and E.G.D. Tuddenham, eds.), pp. 549625. Churchill Livingstone, London. Bode, W. and Schwager, P. 1975. The refined crystal structure of bovine beta-trypsin at 1.8 A resolution. II. Crystallographic refinement, calcium binding site, benzamidine binding site and active site at pH 7.0. J. Mol. Biol. 98:693-717. Bode, W., Engh, R., Hopfner, K.-P., Huber R., and Kopetzki, E. July 1999. Chimeric serine proteases. EP-00927764. Castellino, F.J. and Powell, J.R. 1981. Human plasminogen. Methods Enzymol. 80:365-378. Chapman, H.A. 1997. Plasminogen activators, integrins, and the coordinated regulation of cell adhesion and migration. Curr. Opin. Cell Biol. 9:714-724.
Expression and Purification of Serine Proteases
Davie, E.W., Fujikawa, K., and Kisiel, W. 1991. The coagulation cascade: Initiation, maintenance, and regulation. Biochemistry 30:10363-10370.
Hedstrom, L., Szilagyi, L., and Rutter, W.J. 1992. Converting trypsin to chymotrypsin: The role of surface loops. Science 255:1249-1253. Hopfner, K.P., Brandstetter, H., Karcher, A., Kopetzki, E., Huber, R., Engh, R.A., and Bode, W. 1997. Converting blood coagulation factor IXa into factor Xa: Dramatic increase in amidolytic activity identifies important active site determinants. Embo. J. 16:6626-6635. Jenne, D., Wilharm, E., Huber, R., Bode, W., and Parry, M. Procedure to prepare active serine proteases and their inactive variants. International patent filed. Ke, S.H., Coombs, G.S., Tachias, K., Navre, M., Corey, D.R., and Madison, E.L. 1997. Distinguishing the specificities of closely related proteases. Role of P3 in substrate and inhibitor discrimination between tissue-type plasminogen activator and urokinase. J. Biol. Chem. 272:16603-16609. Kopetzki, E. and Hopfner, K.-P. December 1997. Recombinant blood-coagulation proteases. WO09747737. Kopetzki, E., Rudolph, R., and Grossmann, A. 1993. Recombinant core streptavidin. U.S, patent US5,489,528 and European patent EP612,325. Lottenberg, R. 1997. A novel approach to explore the role of plasminogen in bacterial pathogenesis. Trends Microbiol. 5:466-468. Madison, E.L. and Sambrook, J.E. 1993. Probing structure-function relationships of tissue-type plasminogen activator by oligonucleotide-mediated site-specific mutagenesis. Methods Enzymol. 223:249-271. Murakami, M., Towatari, T., Ohuchi, M., Shiota, M., Akao, M., Okumura, Y., Parry, M.A., and Kido, H. 2001. Mini-plasmin found in the epithelial cells of bronchioles triggers infection by broadspectrum influenza A viruses and Sendai virus. Eur. J. Biochem. 268:2847-2855. Oberg, K., Chrunyk, B.A., Wetzel, R., and Fink, A.L. 1994. Nativelike secondary structure in interleukin-1 beta inclusion bodies by attenuated total reflectance FTIR. Biochemistry 33:26282634. Parry, M.A., Fernandez-Catalan, C., Bergner, A., Huber, R., Hopfner, K.P., Schlott, B., Guhrs, K.H., and Bode, W. 1998. The ternary microplasmin-staphylokinase-microplasmin complex is a proteinase-cofactor-substrate complex in action. Nat. Struct. Biol. 5:917-923. Parry, M.A., Zhang, X.C., and Bode, I. 2000. Molecular mechanisms of plasminogen activation: Bacterial cofactors provide clues. Trends Biochem. Sci. 25:53-59. Seckler, R. and Jaenicke, R. 1992. Protein folding and protein refolding. FASEB J. 6:2545-2552. Sottrup-Jensen, L., Zajdel, M., Claeys, H., Petersen, T.E., and Magnusson, S. 1975. Amino-acid sequence of activation cleavage site in plasminogen: Homology with “pro” part of prothrombin. Proc. Natl. Acad. Sci. U.S.A. 72:2577-2581.
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Wang, X., Lin, X., Loy, J.A., Tang, J., and Zhang, X.C. 1998. Crystal structure of the catalytic domain of human plasmin complexed with streptokinase. Science 281:1662-1665. Wang, X., Terzyan, S., Tang, J., Loy, J.A., Lin, X., and Zhang, X.C. 2000. Human plasminogen catalytic domain undergoes an unusual conformational change upon activation. J. Mol. Biol. 295:903-914. Wilharm, E., Parry, M.A., Friebel, R., Tschesche, H., Matschiner, G., Sommerhoff, C.P., and Jenne, D.E. 1999. Generation of catalytically active granzyme K from Escherichia coli inclusion bodies and identification of efficient granzyme K inhibitors in human plasma. J. Biol. Chem. 274:27331-27337. Wu, H.L., Shi, G.Y., and Bender, M.L. 1987. Preparation and purification of microplasmin. Proc. Natl. Acad. Sci. U.S.A. 84:8292-8295.
Contributed by Marina A. A. Parry Actelion Pharmaceuticals Ltd. Allschwil, Switzerland
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Assaying Proteases in Cellular Environments
UNIT 21.12
To better study protease activation, regulation, and inhibition physiologically or pharmacologically, it is often advantageous to use cell-based systems. The introduction of the cellular environment poses additional challenges for the assay of the protease of interest. In this unit, various methods for monitoring protease activity in cell-based systems will be discussed— including procedures for intracellular proteases (see Basic Protocols 1 to 3) as well as for those that might be secreted into cell-conditioned medium (see Basic Protocols 4 and 5). The possibility of using an endogenous protein substrate (see Basic Protocol 2) versus an introduced peptide substrate (see Basic Protocol 1) to monitor intracellular protease activity will be discussed. The scope of the unit will be restricted to mammalian cell system. It is also stressed that specific considerations or requirements for a particular protease or peptidase are often needed in order to optimize the assay. Readers are advised to refer to other units that cover the protease of interest (UNITS 21.1-21.4; UNIT 21.8; UNITS 21.10 & 21.11). MONITORING INTRACELLULAR PROTEASE ACTIVITY IN SITU WITH CELL-PERMEABLE PEPTIDE SUBSTRATE
BASIC PROTOCOL 1
If the protease of interest is located intracellularly, ideally one would like to observe and measure the protease activity without perturbing the cellular environment (e.g., by lysing the plasma membrane). For certain proteases, it is possible to introduce a small fluorogenic peptide substrate to which the cell membrane is permeable. Thus, once the peptide is introduced into the cells, the protease can be activated by the addition of stimulus, and the protease activity can be tracked either continuously or as end-point measurement, using a fluorometer. The use of colorimetrically detectable substrates (e.g., peptide-p-nitroanilide) usually does not yield a signal strong enough for detection, and is therefore not generally recommended. An example of the use of the above mentioned procedure is the study of the calcium-activated protease (calpain). A cell-permeable peptide substrate such as succinyl-Leu-LeuVal-Tyr-7-amino-4-chloromethylcoumarin (SLLLVY-AMC) can be introduced, and the protease is then activated by a calcium-channel opener, maitotoxin or calcium ionophore (e.g., A23187). The fluorescence derived from the release of AMC is monitored in a 12-well plate format (Wang et al., 1996). A similar peptide (t-butoxycarbonyl-Leu-MetAMC) which is conjugated with glutathione intracellularly (as cell trap) has also been described (Rosser et al., 1993). Similarly, rhodamine 110–linked cell-permeable peptide substrates or the fluorogenic peptide substrate PhiPhiLux (OncoImmunin, Inc.) for caspases can be used in this manner (Hug et al., 1999; Komoriya et al., 2000). Materials Cell line or primary cell culture with demonstrated expression of calpain I or II, e.g., human neuroblastoma SH-SY5Y cells (Wang et al., 1996) DMEM medium, serum-free (Sigma), containing 0.8 mM CaCl2 40 mM stock solution of succinyl-Leu-Leu-Val-Tyr-7-amido-4-methylcoumarin (SLLVY-AMC; Sigma) in DMSO Calcium channel opener: 1 µM maitotoxin (MTX) or 10 mM calcium ionophore A 23187 (Calbiochem) in DMSO Cell culture medium, serum-free, containing 5 mM EDTA Calpain inhibitor I (Ac-Leu-Leu-nLeu-CHO; Calbiochem) 12-well plates (tissue–culture treated; Corning) Fluorescence plate reader (e.g., Millipore Cytofluor 2300) Additional reagents and equipment for cell culture (APPENDIX 3C) Peptidases Contributed by Kevin K.W. Wang Current Protocols in Protein Science (2002) 21.12.1-21.12.14 Copyright © 2002 by John Wiley & Sons, Inc.
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600
Fluorescence units
500
400
300
200
100
0 Basal
+10 µM calpain inhibitor I
MTX
+10 µM calpain inhibitor I
Figure 21.12.1 Protease activity monitored by cell-permeable fluorogenic peptide substrate. SLLVY-hydrolysis by calpain is measured in SY5Y cells without calcium channel opener maitotoxin (basal; open bar) or with maitotoxin (MTX, treatment for 60 min; dark bar). Each of these conditions were duplicated in the presence of 10 µM calpain inhibitor I represented by the shorter bar to the right of each of the above bars.
1. Grow human neuroblastoma SH-SY5Y cells to cell density of 1 × 106 cells/well (80% confluence) in 12-well plates (see APPENDIX 3C for basic cell culture techniques). 2. Wash the culture wells three times with serum-free medium. Reintroduce 0.5 ml of serum-free medium containing 0.8 mM CaCl2 into each of the wells. 3. To each well, add 0.5 ml of serum-free medium containing a quantity of succinylLeu-Leu-Val-Tyr-7-amido-4-methylcoumarin (SLLVY-AMC; add as 40 mM stock in DMSO) sufficient to achieve a final concentration of 80 µM (Rosser et al., 1993). 4. Add calcium channel opener MTX (as 1 µM stock) to a final concentration of 0.1 to 1 nM, or calcium ionophore A23187 (as 10 mM stock) to a final concentration of 10 µM. Incubate the plates at room temperature or at 37°C. As negative control, use medium with 5 mM EDTA added. Duplicate each condition in the presence of 10 µM calpain inhibitor I. Results of an experiment set up in this manner are shown in Figure 21.12.1
5. Measure fluorescence (excitation 380 nm and emission 420 nm with slit width set at 15 nm and 20 nm, respectively) every 15 to 30 min, up to 120 min, with a Millipore Cytofluor 2300 fluorescence plate reader.
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MONITORING INTRACELLULAR PROTEASE ACTIVITY IN SITU BY AUTOLYTIC ACTIVATION AND ENDOGENOUS SUBSTRATE PROTEIN DEGRADATION
BASIC PROTOCOL 2
A powerful means to monitor intracellular protease activity is by monitoring the integrity and/or level of the protease, as well as its endogenous protein substrates, instead of introducing an artificial substrate. The general technique calls for standard cellular protein extraction, SDS-PAGE (UNIT 10.1), immunoblotting (UNIT 10.10), and then the detection with a specific antibody (Fig. 21.12.2). Most proteases exist as zymogens that are proteolytically or autolytically activated—e.g., cathepsin B, L, D, calpains, caspases and matrix metalloproteases (MMPs). Similarly distinct endogenous protein substrates have been identified for a number of intracellular proteases such as calpain (α-spectrin), caspases (poly(ADP)ribose polymerase or PARP), and proteasome (e.g., iκB-α). With calpain and caspase, the substrates are generally cleaved into fragments with smaller molecular weight, readily distinguishable from the intact proteins. The proteasome generally degrades proteins into very small peptides. Thus, the disappearance or intensity reduction of the intact protein band, rather than protein fragments, is expected. Also, inhibition of proteasome (e.g., lactacystin) would result in an accumulation of the high-molecularweight ubiquitin-conjugates of the protein of interest, since ubiquitinization is the biochemical step preceding proteasome-mediated degradation (Figueiredo-Pereira et al., 1994). Materials Cell type expressing protease of interest Serum-free DMEM medium (Sigma), containing 0.8 mM CaCl2 Calcium channel opener: maitotoxin (MTX) or calcium ionophore (Calbiochem) Lysis buffer I (see recipe) SDS sample buffer (see recipe) 4% to 20% polyacrylamide gradient Tris-glycine mini-gel (Novex) Rainbow-colored molecular weight markers (RPN756: Amersham)
A
B intact protein substrate
zymogen activated protease
protein fragment
Figure 21.12.2 Protease proteolytic/autolytic activation and endogenous substrate protein fragmentation visualized after SDS-PAGE and immunoblotting. (A) An antibody to the protease of interest detects protease zymogen in resting cells (left lane), but detects the partially processed and activated form of the protease when cells are stimulated or challenged (right lane). (B) Similarly, an antibody against an endogenous substrate of the protease of interest detects the intact protein in resting cells (left lane) and its major immunoreactive fragment of lower molecular weight in challenged cells (right lane).
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Running buffer (see recipe) TBST (see recipe) 5% (w/v) nonfat milk in TBST (see recipe for TBST) Primary antibody: Monoclonal anti-calpain I antibody diluted 1:500 in TBST or monoclonal anti-α-spectrin antibody diluted 1:3000 in TBST (both antibodies available from Chemicon; see recipe for TBST) Secondary antibody: biotinylated anti-mouse IgG diluted 1:1000 in TBST (see recipe for TBST) Alkaline phosphatase–conjugated avidin diluted 1:3000 in TBST (see recipe for TBST) 12-well plates (tissue culture–treated; Corning) PVDF membrane (Novex), presoaked with 100% methanol Semidry electrotransfer unit (e.g., Bio-Rad) Additional reagents and equipment for protein assay (UNIT 3.4), SDS-PAGE (UNIT 10.1), electroblotting onto PVDF membranes (UNIT 10.7), and immunoblot detection (UNIT 10.10) Prepare cells 1. Select a cell type that expresses the protease of interest (e.g., human SH-SY5Y cells, which express calpain). Grow cells to cell density of 1 × 106 cells/well (80% confluence) in 12-well plates (see APPENDIX 3C for basic cell culture techniques). The presence of an endogenous substrate protein for the protease of interest (e.g., alpha II-spectrin) should also been confirmed before hand.
2. Wash the cultures three times with serum-free medium. Reintroduce 0.5 ml of serum-free medium into each well. 3. Add calcium channel opener MTX to a final concentration of 0.1 to 1 nM, or calcium ionophore A23187 to a final concentration of 10 µM. Incubate the plates at room temperature or at 37°C for 60 to 90 min. Total volume in wells at the end of step 4 should be 1 ml.
4. Add 100 µl of lysis buffer I to each well to arrest any further protease activities. 5. Microcentrifuge the contents of each well for 5 min at maximum speed, 4°C, and collect the supernatant (cell lysate). Samples can be used immediately or stored at −45°C for up to 3 weeks.
Determine extent of autolysis or endogenous substrate fragmentation by immunoblotting 6. Perform protein assay (UNIT 3.4) on sample. Add 5 µl SDS sample buffer to ∼10 to 20 µl of sample (10 to 20 µg protein based on protein assay; use water to make total volume, 40 µl). 7. Load protease samples into the wells of a 4% to 20% polyacrylamide gradient Tris-glycine mini-gel. Run rainbow-colored molecular weight markers alongside for subsequent molecular-weight estimation. 8. Run electrophoresis in running buffer at 125 V for 2.2 hr at room temperature.
Assaying Proteases in Cellular Environments
9. Transfer proteins onto a PVDF membrane with Tris-glycine buffer system (using a semidry electrotransfer unit; Bio-Rad) at 20 mA for 1.5 to 2 hr (UNIT 10.7). Wash membranes four times with TBST.
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10. Perform immunoblotting as follows: a. Block blots with 5% (w/v) nonfat milk in TBST for 30 min. Wash membranes four times with TBST. b. Probe with monoclonal anti-calpain I diluted 1:500 in TBST or monoclonal anti-α-spectrin antibody diluted 1:3000 in TBST for 2 hr or overnight. Wash membranes four times with TBST. c. Probe with biotinylated anti-mouse IgG secondary antibody diluted 1:1000 in TBST for 2 hr. Wash membranes four times with TBST. d. Probe with alkaline phosphatase–conjugated avidin diluted 1:3000 in TBST for 30 min. Wash membranes four times with TBST. 11. Develop the blot with nitro blue tetrazolium and 5-bromo-4-chloro-3-indolyl-phosphate (UNIT 10.10). The inert zymogen in resting cells (80 kDa) and autolyzed active form of calpain (78 kDa) in challenged cells can be visualized on the blot probed with anti-calpain I antibody (see Fig. 21.12.2; Wang et al., 1996). Similarly, for proteolysis of endogenous substrate a II-spectrin, the blot probed with anti-a-spectrin would detect the intact protein (280 kDa) in resting cells, and its major immunoreactive fragment of lower molecular weight (150 kDa) in challenged cells, respectively (see Fig. 21.12.2; Wang et al., 1996). Densitometric analysis of immunoblots was performed using a color scanner (Umax UC630) and the NIH program Image 1.5.
MONITORING PROTEASE ACTIVITY LEVELS AS REFLECTED IN CELL LYSATE
BASIC PROTOCOL 3
Many proteases exist as inactive zymogen or proenzyme and need to be proteolytically or autolytically activated (e.g., cathepsin B, calpains, and caspases). Thus, by extracting cell lysate, it is possible to detect the level of “activated” form of the protease of interest. Caspase-3 is an excellent example. Its proenzyme form, 32 kDa, is active, but is only activated by a two-step process: (1) processing at the interface region between the p20 and p12 subunits and (2) autolytic truncation at the N-terminal. The end product is a fully activated p17-p12 heterodimeric enzyme capable of hydrolyzing synthetic substrate such as Acetyl-DEVD-AMC (Nath et al., 1996). Thus, by using a nonionic detergent (Triton X-100)based cell lysis and total cellular protein extraction method as described in this protocol, the activation of caspase-3 can be readily monitored under apoptotic conditions. Again, specific inhibitors of caspase-3 should be used to validate the specificity of the assay. Cautions should be exercised regarding the specificity of the synthetic protease substrate, since the total protein extract is likely to contain various other proteases that might hydrolyze the same substrates. It is also worth noting that if an endogenous inhibitor for the protease of interest is present in the cell lysate, one will be measuring the net activity of the protease over its endogenous inhibitor. That is, the protease activity could be partially or completely masked by the inhibitor. Examples of endogenous protein inhibitors are serpins (UNIT 21.7) for serine proteases (UNITS 21.10 & 21.11), cystatins (stefins) for cysteine protease (UNIT 21.2), and TIMPs (tissue inhibitors of metalloproteases) for metalloproteases (UNIT 21.4). Inhibitors of apoptosis proteins (IAPs) are specific inhibitors of caspases (UNIT 21.8), and calpastatin is specific for calpain I and II. Materials Human neuroblastoma SH-SY5Y cells growing in 12-well plates Serum-free DMEM medium (Sigma) containing 0.8 mM CaCl2 Staurosporine (Calbiochem) TBS-EDTA (see recipe), room temperature Lysis buffer II (see recipe), 4°C Glycerol
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Substrate solution for caspase-3 (see recipe) Caspase inhibitor: carbobenzoxy-Asp-CH2OC(=O)-2,6-dichlorobenzene (Z-D-DCB; Bachem) 12-well plates (tissue culture–treated; Corning) Fluorescence plate reader (e.g., Millipore Cytofluor 2300) 1. Wash adherent human neuroblastoma SH-SY5Y cells (in 12-well plates) three times with serum-free DMEM. Challenge cells by exposure to 0.5 µM staurosporine also in serum-free DMEM medium for 0, 1, 3, 6, and 24 hr. 2. Wash cells three times with serum-free DMEM, then two times with 1 ml of room temperature TBS-EDTA. Suspension cells, may be washed three times with serum-free medium, each time by resuspending in 1 ml medium and centrifuging at 5,000 × g, 4°C, and finally resuspending in 1 ml TBS-EDTA and centrifuging again to recover cell pellet in a 1.5-ml microcentrifuge tube.
3. Lyse cells directly in the bottoms of the wells (or cell pellets in microcentrifuge tubes if working with suspension cells) in in lysis buffer II for 60 to 90 min at 4°C. 4. Clear lysates by centrifuging 5 min at 5,000 × g, 4°C, and retaining the supernatant. Add glycerol to 50% (v/v) and store at –70°C. 5. To assay caspase-3 activity, add 25 µl of cell lysate to 225 µl substrate solution for caspase-3 (see recipe), in the absence or presence of a caspase inhibitor such as 10 µM carbobenzoxy-Asp-CH2OC(=O)-2,6-dichlorobenzene (Z-D-DCB), or other protease inhibitor, depending on the enzyme being assayed (optional). 6. Measure fluorescence (excitation 380 nm ± 15 nm and emission 460 nm ± 15 nm) every 15 to 30 min, up to 60 min, with a Millipore Cytofluor 2300 fluorescence plate-reader or equivalent. BASIC PROTOCOL 4
MONITORING ACTIVITY OF SECRETED PROTEASE IN SITU Mast cells are characterized by their abundant content of tryptase and chymase, which are also secreted in fully active forms. Both of these are serine proteases (Algermissen et al., 1999; UNITS 21.10 & 21.11). Since they are sensitive to the serine protease inhibitor diisopropylfluorophosphate (DFP), one possible way of detecting their activity or level of expression is by incubating cell lysate or whole cells with radioactive DFP. Cell samples may be processed for gel or immunoblotting and analyzed by autoradiography. Another approach is by the application of selective and cell-penetrating inhibitors such as selective chymase inhibitors—e.g., Z-Ile-Glu-Pro-Phe-CO(2)Me or Ala-Ala-Phe-chloromethylketone (AAPCK)—or tryptase inhibitors—e.g., APC 366 ([N-(1-hydroxy-2-naphthoyl)-Larginyl-L-prolinamide hydrochloride]) and BABIM ([bis(5-amidino- 2benzimidazolyl)methane); also see Table 21.12.1. These agents, when added to mast cell culture, prevent mast cell IgE-dependent activation and degranulation (Rice et al., 1998; He et al., 1999). Since both of these proteases are released as fully active enzymes, it is possible to collect cell-conditioned medium and monitor by hydrolytic activities of (Boc)-Phe-Ser-Arg-p-nitroanilide by tryptase or succinyl-Ala-Ala-Pro-Phe-p-nitroanilide by chymase.
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Table 21.12.1
Selective Protease Inhibitors Compatible with Cell Culture Environment
Protease
Selective inhibitors
Calpain I and II Caspases Cathepsin Ba Cathepsin D Cathepsin Kb Cathepsin L Chymase MMPs
MDL28170; PD150606, calpain inhibitor I Cbz-Asp-CH2-DCB; Ac-Asp-Glu-Val-Asp-CHO CA-074 Pepstatin A Cbz-Leu-Leu-CH2OMe Boc-Leu-Gly-aziridine-2, 3-dicarboxylic acid-(OEt)2 Cbz-Ile-Glu-Pro-Phe-CO(2)Me; AAPCK EDTA; TAPI-1; HS-CH2-(R)-CH(CH2CH(CH3)2)-CO-Phe-Ala-NH2 lactacystin; MG132; PSI APC-366; BABIM OM-99-2
Proteasome Tryptase APP β-secretasec aButtle et al. (1992) bThompson et al. (1999) cGhosh et al. (2001)
Materials Human mast cells or other cells that express chymase/tryptase, growing in culture Serum-free RPMI medium Immunoglobulin for activation and degranulation of mast cells, e.g., IgE 1 M HEPES, pH 7.4 Substrate solution for tryptase/chymase (see recipe) Serine protease inhibitors (UNIT 21.7): Diisopropyl fluorophosphate (for both serine proteases; Sigma) Tosyllysine chloromethyl ketone (TLCK, for tryptase; Sigma) Tosylphenylalanine chloromethyl ketone (TPCK, for chymase; Sigma) 12-well plates (tissue culture–treated; Corning) 96-well microtiter plates UV-visible spectrophotometer with microtiter plate reader (e.g., Spectramax; Molecular Devices) Prepare cell-conditioned medium 1. Plate 0.5–1 × 106 human mast cells or other cells that express chymase/tryptase in 12-well culture plates, using 1 ml serum-free RPMI medium per well. 2. Immunologically activate and degranulate mast cells by adding immunoglobulin, e.g., 10 to 100 µg/ml IgE. Allow activation to proceed for 2 hr at 37°C. 3. After 2 hr incubation, collect 500 µl cell-conditioned medium containing the released chymase and tryptase. Microcentrifuge 10 min at 10,000 × g, 4°C, to remove cell debris. Retain the supernatant. 4. Add 50 µl of 1 M HEPES, pH 7.4, to achieve a final concentration of 100 mM, and store at 4°C or quick-freeze on dry ice and store at −45°C until use. Assay protease activity 5. Add 20 to 100 µl of the conditioned medium to each well of a 96-well plate containing substrate solution for tryptase or chymase, for a final volume of 250 µl. Reserve some of these wells as negative controls and add serine protease inhibitors to these—2 mM diisopropyl fluorophosphate (for both serine proteases), 100 mM tosyllysine chlo-
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romethyl ketone (TLCK, for tryptase), 100 mM tosylphenylalanine chloromethyl ketone (TPCK, for chymase), or the more selected tryptase/chymase inhibitors outlined above. 6. Monitor release of chromophore nitroanaline at 405 nm with a UV-visible microtiter plate reader spectrophotometer. BASIC PROTOCOL 5
Assaying Proteases in Cellular Environments
USING ZYMOGRAPHY TO MEASURE ACTIVITY OF CELL-SECRETED MMPs Matrix metalloproteases (MMPs; e.g., gelatinases and stromelysins; UNIT 21.4) are zinc-requiring proteases that are secreted into intercellular space. They are designated as MMP-1 through MMP-12; examples include MMP-2 (gelatinase A), MMP-9 (gelatinase B), and MMP-3 (stromelysin-1). Their major targets are extracellular matrix (ECM) proteins, such as collagen. Some of them are inducible when the cells are stimulated. MMPs also have a reputation of being capable of refolding back into active enzyme following removal of a denaturing detergent such as SDS. A zymography technique which exploits this refolding behavior is commonly used to assay these enzymes. Preferred substrates for the MMP such as collagen, gelatin, or casein are copolymerized into an acrylamide gel matrix and samples of cell-conditioned medium are subjected to denaturing SDS-PAGE. The MMP bands in the gel are then reactivated by removal of SDS, which is replaced with Triton X-100. The activated but immobilized MMPs thus hydrolyze the protein substrate in the local environment. Upon completion of the digestion, the substrate gel is stained with Coomassie blue. Over a uniformly stained gel, the MMPs should show up as clearing bands (Wang, 1999). Gelatinase A (70 kDa, active form 62 kDa) and B (92 kDa, active form 82 kDa) can be simultaneously detected due to their different sizes. This zymogram method has been used successfully to quantify level of various MMPs, as well as to monitor the intracellular conversion of MMP zymogens to their activated forms. MMP inhibitors such as HONHCOCH2CH(CH2CH(CH3)2)-CO-Nal-Ala-NHCH2CH2NH2 (TAPI-1) or HS-CH2-(R)-CH(CH2CH(CH3)2)-CO-Phe-Ala-NH2 (Calbiochem) could be used to selectively inhibit MMP-mediated biochemical processes in cell environment (Crowe et al., 1995; Table 21.12.1). A modification of this method has been used to detect the level of endogenous MMP inhibitors called tissue inhibitors of MMPs (TIMP 1 to 3). In this case, both the substrate protein (e.g., gelatin) and the MMP are incorporated into the polymerized gel. The cell lysate or cell-conditioned medium containing the TIMP is be loaded onto the gel lanes. After electrophoresis, the zymogram is treated as before. However, on final analysis, this zymogram will have a generally clear background because the substrate is digested with the embedded MMP, and the TIMP bands should appear as Coomassie-stained blue bands, since the MMP is inhibited locally, leaving the gelatin undegraded (Oliver et al., 1999). Materials Human fibroblasts growing in cell culture DMEM medium (Sigma), serum free Phorbol ester: e.g., phorbol-12-myristate-13-acetate to induce MMP-2 and MMP-9 100 mM EGTA SDS sample buffer (see recipe) without 2-mercaptoethanol Precast gelatin PAGE gel (Novex) Running buffer (see recipe) SDS-PAGE molecular-weight standards (broad-range, Bio-Rad) Reactivation buffer: 2.5% (v/v) Triton X-100/100 mM glycine, pH 8.3 Proteolysis buffer: 100 mM glycine, pH 8.3 Fixing/destaining solution: 5:4:1 methanol:water:glacial acetic acid 0.25% (w/v) Coomassie blue R-250 in fixing/destaining solution 2.5% (v/v) acetic acid
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Additional reagents and equipment for SDS-PAGE (UNIT 10.1) Prepare the gel and samples 1. Challenge human fibroblasts (in 1 ml of serum-free DMEM medium) with phorbol ester (100 nM phorbol-12-myristate-13-acetate) for 24 hr to induce MMP-2 and MMP-9. 2. Add 25 µl of 100 mM EGTA stock solution to the cell-conditioned medium and collect 35-µl medium samples for MMP assay. 3. Add 5 µl of SDS sample buffer (total volume 40 µl) and keep at room temperature or stored at −20°C. Do not use reducing agent such as 2-mercaptoethanol or dithiothreitol.
Perform electrophoresis 4. Pre-run the precast gelatin PAGE gel in running buffer for 15 min at 125 V, room temperature. 5. Load the protease samples into the wells of the gel. Load SDS-PAGE molecular weight markers alongside for subsequent molecular weight estimation. 6. Run electrophoresis in running buffer at 125 V for 2.2 hr at room temperature. Reactivate the enzyme and allow proteolysis to occur 7. Remove gel from electrophoresis apparatus and incubate in two changes of reactivation buffer at 20° to 25°C over a period of 1 hr. 8. Incubate gel in three changes of proteolysis buffer at 20° to 25°C over a period of 3 hr. Continue incubating the gels in the same buffer for 6 to 24 hr at ambient temperature. Fix and stain the gel 9. At the end of the proteolysis reaction, incubate the gels in two changes of water over a period of 1 hr, then fix in fixing/staining solution for 30 min. 10. Stain gel with 0.25% (w/v) Coomassie blue R-250 in fixing/destaining solution for 30 min, then destain with several changes of fixing/destaining solution over a period of 2 to 5 hr. 11. Store the gel in 2.5% (v/v) acetic acid. The gels can be observed, imaged, or quantified densitometrically.
REAGENTS AND SOLUTIONS Use deionized or distilled water in all recipes and protocol steps. For common stock Solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Lysis buffer I (for Basic Protocol 1) 50 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) 100 mM NaCl 1% (v/v) Triton X-100 2.5 mM EGTA 2.5 mM EDTA 1× protease inhibitor cocktail (Complete Mini; Boehringer Mannheim) Store up to 3 months at 4°C Peptidases
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Lysis buffer II (for Basic Protocol 3) 20 mM Tris⋅Cl, pH 7.4 at 4°C (APPENDIX 2E) 150 mM NaCl 1 mM DTT 5 mM EDTA 5 mM EGTA 1% (v/v) Triton X-100 Store up to 3 months at 4°C Running buffer 0.2% (w/v) SDS 25 mM Tris base 192 mM glycine, pH 8.3 Store up to 1 month at room temperature SDS sample buffer 10% (w/v) SDS 20% (v/v) glycerol 0.004% (w/v) bromphenol blue 2 mM 2-mercaptoethanol (omit for MMP assay) 200 mM Tris⋅Cl, pH 6.8 (APPENDIX 2E) Store up to 1 year at −10°C Substrate solution for caspase-3 100 µM peptide substrate Ac-DEVD-AMC (Bachem Bioscience) 100 mM HEPES, pH 7.4 10% (v/v) glycerol 1 mM EDTA 10 mM DTT Prepare fresh Substrate solution for tryptase/chymase Prepare 100 mM HEPES containing 200 µM (Boc)-Phe-Ser-Arg-p-nitroanilide (for tryptase; available from Sigma; see Takai et al., 2000) or succinyl-Ala-Ala-Pro-Phep-nitroanilide (for chymase; available from Peptides International; see Caughey et al., 1988). Make fresh before use. TBS-EDTA 20 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) 155 mM NaCl 1 mM EDTA Store up to 3 months at 4°C TBST 230 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) 150 mM NaCl 0.2% Tween 20 Store up to 1 month at room temperature COMMENTARY Background Information Assaying Proteases in Cellular Environments
The presence of the cellular environment introduces a large number of proteins and enzymes other than proteases, as well as other biochemicals, into the protease assay system.
Some of these could inhibit the activity of the protease of interest (resulting in false negatives) or, alternatively, contribute to the hydrolysis of the protease substrate employed (resulting in false positives). In this unit, several different
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methods for monitoring protease activity in cell-based systems have been described. For cysteine proteases (UNIT 21.10), the use of fluorogenic cell-permeable peptide substrates seems to work well (see Basic Protocol 1). Alternatively, synthetic peptide substrates in cell lysate work well for both cysteine proteases and aspartic proteases (see Basic Protocol 3; UNITS 21.2 & 21.3). MMP (UNIT 21.4) and other metalloprotease activity can easily be monitored using synthetic peptide substrate (see Basic Protocol 4) or zymography (see Basic Protocol 5) with cell-conditioned medium, since these proteases are usually secreted. However, cell lysate can also be alternatively used. For serine proteases that are secreted, again, an assay using synthetic peptide substrates with cell lysate is the method of choice. If all else fails, usually one can fall back on monitoring protease autolysis or endogenous substrate proteolysis by immunoblotting (see Basic Protocol 2; UNIT 10.10). This method has been proven to work for a large number of protease family members including calpain, caspases, proteasome, cathepsins, and amyloid precursor protein (APP) β- and γ-secretases.
Critical Parameters and Troubleshooting Selecting a cell line The choice of cell type is obviously very important. If no protease activity is detected using the protocols outlined in this unit, one of the most likely possibilities is that a cell type was used that does not provide enough of the protease of interest. Thus, to select a cell line, the primary criterion is the presence of a reasonably high level of the protease of interest. Other considerations that would favor a particular cell line are the lack of other proteases that could hydrolyze the same substrate, as well as low levels of endogenous inhibitor(s) of the protease of interest—such as calpastatin for calpain, serpin for chymase and other serine proteases, inhibitor of apoptosis proteins (IAPs) for caspases, and tissue inhibitor of matrix metalloprotease (TIMPs) for matrix metalloproteases. The choice of peptide substrate is also very important. The primary goal is to select a substrate that (1) is selectively hydrolyzed by the protease of interest and (2) is cell-membrane permeable. Often, the starting point would be those peptide substrates that are used in an in vitro assay for the protease.
The use of selective protease inhibitors The use of selective protease inhibitors is a very powerful technique for validating the specificity of a cell-based protease system. However, the most common limitations on this technique are the lack of permeability and selectivity of the protease inhibitor. Sometimes, by applying higher concentrations of a peptide inhibitor with low cell permeability, one can get a sufficient amount of the inhibitor into the cell. Selectivity is a more important issue. For example, several calpain-inhibiting peptide aldehydes (such as calpain inhibitor I and II) also cross-inhibit threonine protease proteasome. Yet, a calcium-binding-site directed calpain inhibitor, PD150606 (Calbiochem), does not inhibit proteasome (Yuen and Wang, 1998; Table 21.12.1). Conversely, a threonine-modifying lactacystin is highly specific for proteasome. Other proteasome inhibitors, including CbzLeu-Leu-Leu-CHO (MG 132), and Cbz-IleGlu(OtBu)-Ala-Leu-H (PSI), are also more selective for proteasome than for calpain. These compounds are also available from Calbiochem; see Table 21.12.1. Thus, the use of more than one inhibitor is strongly advised. Iodinated calpain inhibitors have been used successfully in labeling activated calpain in activated platelets (Anagli et al., 1993). Similarly, two biotinlabeled caspase inhibitors—Ac-YVK(Biotin)D-amk (Yamin et al., 1996) and CbzVK( Bio tin) D-f mk (Ar mstr ong et al., 1996)—have been utilized successfully to label caspase-3 in apoptotic cell lysate. Among the cysteine protease cathepsins K, L and B, there are now documented inhibitors that show subclass selectivity (Yamamoto et al., 1997; Marquis et al., 1999; Schirmeister, 1999; Table 21.12.1). The aspartic protease inhibitor pepstatin A has been used successfully in inhibiting cathepsin D activity in intact cells (Deiss et al., 1996). Keeping the protease alive It should be noted that the proteases are sensitive to changes in pH, temperature, and other environmental factors. It is advisable that the cells be freshly prepared and used to measure protease activity as soon as possible. Constant medium temperature (37°C) and pH (7.2 to 7.4) should be maintained. For assessing protease activity in cell lysate, a nonionic detergent (e.g., Triton X-100 or CHAPS) should be used so as not to denature the protease. Precautions should also be taken to preserve the lysates at cold temperatures (4°C to −20°C) throughout the procedures. The addition of
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MCA formation (arbitrary units)
300
DEVD-AMC
200
100
YVAD-AMC
0 0
5
10
15
20
Time (hr)
Figure 21.12.3 Protease activity measurement in cell lysate. SH-SY5Y cells are challenged with staurosporine. Caspase-3 assay is run with cell lysate by measuring the hydrolysis of fluorometric substrates for either caspase-1 (Ac-YVAD-MCA, circles) or caspase-3 (Ac-DEVD-MCA, squares). Assayed cell lysates are collected from SH-SY5Y cells at 0, 1, 3, 6, and 16 hr following staurosporine challenge.
glycerol (40% to 50% v/v) also helps stabilize protease activity in cell lysate.
Anticipated Results Basic Protocol 1 In the experiment represented by Figure 21.12.1, a cell-permeable calpain substrate, succinyl-Leu-Leu-Val-Tyr-7-amido-4-methyl coumarin (SLLVY-AMC, 80 µM), was preincubated with neuroblastoma SH-SY5Y cells; calpain activation is then triggered with the addition of maitotoxin (MTX). The increase in fluorescence due to liberated AMC is readily observed. This signal is inhibited by the addition of either calpain inhibitor I (see Fig. 21.12.1) or EDTA (not shown in figure).
Assaying Proteases in Cellular Environments
Basic Protocol 2 In the experiment represented by Figure 21.12.2, SH-SY5Y cells are first subjected to MTX or A23187 treatment. Cell lysates were analyzed by Western blots. Antibody to calpain 1 detected the pro-form of the protease (zymogen) in resting cells, while the partially processed and activated form of calpain is detected in cells that are stimulated with MTX or A23187. Similarly, an antibody against an en-
dogenous substrate αII-spectrin would detect the intact αII-spectrin (280 kDa) in resting cells and detects its major immunoreactive fragment of lower molecular weight MTX- or A23187challenged cells (150 kDa). Basic Protocol 3 In the experiment represented by Figure 21.12.3, SH-SY5Y cells following staurosporine challenge are lysed in a Triton X-100–containing buffer. Caspase-3 activity is monitored in the cell lysate with fluorometric substrates for either caspase-1 (Ac-YVAD-AMC) or caspase-3 (Ac-DEVD-AMC). In this case, only the pro-apoptosis caspase-3 activity is increased. Basic Protocol 4 Mast cells will secrete chymase and tryptase following treatment with 100 µg/ml IgE. Thus, cell-conditioned medium is collected and assayed with either 200 µM of benzoyl-L-LysGly-Arg-p-nitroanilide for tryptase or succinyl-L-Val-Pro-Phe-p-nitroanilide for chymase (Caughey et al., 1988). The expected results are similar to those obtained in the caspase-3 assay in Basic Protocol 3.
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Time (hr) 0
6
12
18
24
MMP-9
MMP-2
Figure 21.12.4 Using zymography to monitor cell-secreted MMP activity. Cell-conditioned medium (25 µl) from human fibroblasts challenged with PMA for 0 to 24 hr is collected and subjected to gelatin zymography. The lower band is MMP-2 (70 kDa) while the upper band is MMP-9 (92 kDa), as resolved by SDS-PAGE before zymographic development.
Basic Protocol 5 In the experiment represented by Figure 21.12.4, cell-conditioned medium (25 µl) from human fibroblasts, challenged for 0 to 24 hr with phorbol ester (phorbol-12-myristate-13acetate; 100 nM), was collected and subjected to gelatin zymography. MMP-2 (70 kDa) and MMP-9 (92 kDa) were first resolved by SDSpolyacrylamide-3 gel electrophoresis (PAGE). The MMP bands in the gel were then renatured and allowed to hydrolyze the embedded gelatin. When the gels were subsequently fixed and stained with Coomassie blue, the MMP bands would appear as clearing bands over a uniform blue-stained gel background (Fig. 21.12.4). The intensity of the clearing bands correlates with the amount of secreted MMPs.
Time Considerations Basic Protocol 1 Cell preparation requires 6 to 16 hr and the protease assay requires 2 to 3 hr. Basic Protocol 2 Cell lysate preparation requires 6 to 24 hr and immunoblot analysis rquires 6 to 16 hr.
Basic Protocol 3 Cell lysate preparation requires 6 to 24 hr and the protease assay requires 2 to 3 hr. Basic Protocol 4 Preparation of cell-conditioned medium rquires 6 to 24 hr and the protease assay requires 2 to 3 hr. Basic Protocol 5 Cell preparation requires 6 to 24 hr. and the zymography assay requires 8 hr.
Literature Cited Algermissen, B., Laubscher, J.C., Bauer, F., and Henz, B.M. 1999. Purification of mast cell proteases from murine skin. Exp. Dermatol. 8:413418. Anagli, J., Hagmann, J., and Shaw, E. 1993. Affinity labelling of the Ca(2+)-activated neutral proteinase (calpain) in intact human platelets. Biochem. J. 289:93-99. Armstrong, R.C., Aja, T.J., Hoang, K.D., Gaur, S., Bai, X., Alnemri, E.S., Litwack, G., Karanewsky, D.S., Fritz, L.C., and Tomaselli, K.J. 1996. Activation of the CED3/ICE-related protease CPP32 in cerebellar granule neurons undergoing apoptosis but not necrosis. J. Neurosci. 17:553-562. Buttle, D.J., Murata, M., Knight, C.G., and Barret, A.J. 1992. CA074 methyl ester: A proinhibitor
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for intracellular cathepsin B. Arch. Biochem. Biophys. 299:377-380.
lies in neuronal apoptosis. Biochem. J. 319:683690.
Caughey, G.H., Viro, N.F., Lazarus, S.C., and Nadel, J.A. 1988. Purification and characterization of dog mastocytoma chymase: Identification of an octapeptide conserved in chymotryptic leukocyte proteinases. Biochim. Biophys. Acta. 952:142-149.
Oliver, G.W., Stetler-Stevenson, W.G., and Kleiner, D.E. 1999. Zymography, Casein zymography and reverse zymography: Activity assays for proteases and their inhibitors. In Proteolytic Enzymes: Tools and Targets (E.E. Sterchi and W. Stocker, eds.) Unit 5, pp. 63-76. Springer-Verlag, Heidelberg, Germany.
Crowe, P.D., Walter, B.N., Mohler, K.M., OttenEvans, C., Black, R.A., and Ware, C.F. 1995. A metalloprotease inhibitor blocks shedding of the 80-kD TNF receptor and TNF processing in T lymphocytes. J. Exp. Med. 181:205-210.
Assaying Proteases in Cellular Environments
Rice, K.D., Tanaka, R.D., Katz, B.A., Numerof, R.P., and Moore, W.R. 1998. Inhibitors of tryptase for the treatment of mast cell-mediated diseases. Curr. Pharm. Des. 4:381-396.
Deiss, L.P., Galinka, H., Berissi, H., Cohen, O., and Kimchi, A. 1996. Cathepsin D protease mediates programmed cell death induced by interferongamma, Fas/APO-1 and TNF-alpha. EMBO J. 15:3861-3870.
Rosser, B.G., Powers, S.P., and Gores, G.J. 1993. Calpain activity increases in hepatocytes following addition of ATP: Demonstration by a novel fluorescent approach. J. Biol. Chem. 268:2359323600.
Figueiredo-Pereira, M.E., Berg, K.A., and Wilk, S. 1994. A new inhibitor of the chymotrypsin-like activity of the multicatalytic proteinase complex (20S proteasome) induces accumulation of ubiquitin-protein conjugates in a neuronal cell. J. Neurochem. 63:1578-1581.
Schirmeister, T. 1999. New peptidic cysteine protease inhibitors derived from the electrophilic alpha-amino acid aziridine-2,3-dicarboxylic acid. J. Med. Chem. 42:560-572.
Fukami, H., Okunishi, H., and Miyazaki, M. 1998. Chymase: Its pathophysiological roles and inhibitors. Curr. Pharm. Des. 4:439-453.
Takai, S., Sakaguchi, M., Jin, D., and Miyazaki, M. 2000. New method for simultaneous measurements of mast cell proteases in human vascular tissue. Clin. Exp. Pharmacol. Physiol. 27:700704.
Ghosh, A.K., Bilcer, G., Harwood, C., Kawahama, R., Shin, D., Hussain, K.A., Hong, L., Loy, J.A., Nguyen, C., Koelsch, G., Ermolieff, J., and Tang, J. 2001. Structure-based design: Potent inhibitors of human brain memapsin 2 (beta-secretase). J. Med. Chem. 44:2865-2868.
Thompson, S.K., Halbert, S.M., DesJarlais, R.L., Tomaszek, T.A., Levy, M.A., Tew, D.G., Ijames, C.F., and Veber, D.F. 1999. Structure-based design of non-peptide, carbohydrazide-based cathepsin K inhibitors. Bioorg. Med. Chem. 7:599-605.
He, S., Gaca, M.D., McEuen, A.R., and Walls, A.F. 1999. Inhibitors of chymase as mast cell-stabilizing agents: Contribution of chymase in the activation of human mast cells. J. Pharmacol. Exp. Ther. 291:517-523.
Wang, K.K.W. 1999. Detection of proteolytic enzymes using protein substrates. In Proteolytic Enzymes: Tools and Targets (E.E. Sterchi and W. Stocker, eds.) Unit 4, pp. 49-62. Springer-Verlag, Heidelberg, Germany.
Hug, H., Los, M., Hirt, W., and Debatin, K.M. 1999. Rhodamine 110-linked amino acids and peptides as substrates to measure caspase activity upon apoptosis induction in intact cells. Biochemistry 38:13906-13911.
Wang, K.K.W., Hajimohammadreza, I., Raser, K.J., and Nath, R. 1996. Maitotoxin induces calpain activation in SH-SY5Y neuroblastoma cells and cerebrocortical cultures. Arch. Biochem. Biophys. 331:208-214.
Komoriya, A., Packard, B.Z., Brown, M.J., Wu, M.L., and Henkart, P.A. 2000. Assessment of caspase activities in intact apoptotic thymocytes using cell-permeable fluorogenic caspase substrates. J. Exp. Med. 191:1819-1828.
Yamamoto, A., Hara, T., Tomoo, K., Ishida, T., Fujii, T., Hata, Y., Murata, M., and Kitamura, K. 1997. Binding mode of CA074, a specific irreversible inhibitor, to bovine cathepsin B as determined by X-ray crystal analysis of the complex. J. Biochem. 121:974-977.
Marquis, R.W., Ru, Y., Yamashita, D.S., Oh, H.J., Yen, J., Thompson, S.K., Carr, T.J., Levy, M.A., Tomaszek, T.A., Ijames, C.F., Smith, W.W., Zhao, B., Janson, C.A., Abdel-Meguid, S.S., D’Alessio, K.J., McQueney, M.S., and Veber, D.F. 1999. Potent dipeptidylketone inhibitors of the cysteine protease cathepsin K. Bioorg. Med. Chem. 7:581-588.
Yuen, P.-W. and Wang, K.K.W. 1998. Calpain inhibitors: Novel neuroprotectants and potential anticataractic agents. Drug Future 23:741-749.
Nath, R., Raser, K.J., Stafford, D., Hajimohammadreza, I., Posner, A., Allen, H., Talanian, R.V., Yuen, P.-W., Gilbertsen, R.B., and Wang, K.K.W. 1996. Nonerythroid alpha-spectrin breakdown by calpain and ICE-like protease(s) in apoptotic cells: Contributory roles of both protease fami-
Contributed by Kevin K.W. Wang Pfizer Global Research and Development Ann Arbor, Michigan
Yamin, T.T., Ayala, J.M., and Miller, D.K. 1996. Activation of the native 45-kDa precursor form of interleukin-1-converting enzyme. J. Biol. Chem. 271:13273-13282.
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Expression, Purification, and Characterization of Caspases
UNIT 21.13
Caspases are a family of cysteine protease necessary for apoptosis and the inflammatory response. There are 14 different caspases in mammals but only 11 are present in humans. They have a strict specificity for aspartic acid residues at the P1 position within their substrate. Other important determinants are the P2 to P4 residues, which confer a rather loose selectivity (Thornberry et al., 1997) and the P1(residue, which can greatly influence the efficiency of cleavage of a given sequence (Stennicke et al., 2000). Because of these considerations, no diagnostic synthetic substrate can be assigned to any caspase, but only a subset of preferred substrates. Nevertheless it is possible to delineate the role of individual caspases by the identification of their intracellular protein substrates. Inflammatory caspases (-1, -4, and -5) can process cytokines, such as IL-1β and IL-18 (Kuida et al., 1995; Li et al., 1995). Apoptotic caspases can be further subdivided into initiator (caspase-2, -8, -9, and -10) and executioner caspases (-3, -6, and -7). Whereas the main substrates of initiator caspases are executioner caspases, the latter ones process the many cellular proteins to produce apoptosis. On the molecular level, caspase activation requires rearrangement of loops to position the catalytic machinery and substrate binding sites. This is achieved by dimerization (caspase-8, -9, -10, and probably caspase-2; Renatus et al., 2001; unpub. observ.) or by cleavage of the linker region bounding the large and small subunits of executioner caspases, since those enzymes are already dimers as latent zymogens. Cleavage of initiator caspases is most likely a sign that they have been activated than a direct mark of their activation. Further information on the mechanism of caspase activation can be found in UNIT 21.8. The signaling cascade leading to caspase activation could originate from the outside the cell (extrinsic pathway) or from the inside (intrinsic pathway). In both cases, initiator caspases will transmit the signal to the executioners. Levels of complexity are provided by amplification loops, endogenous inhibitors, and many cell-specific molecules. Expression of multiple caspases (also see UNIT 21.8) with overlapping substrate specificities renders purification of these enzymes from the animal tissue and cells that naturally contain them a tedious process. However, prokaryotic expression systems allow for quick and effective expression and purification of caspases. Because caspases are toxic when expressed in E. coli, a tightly regulated expression system, such as the pET expression system from Novagen, is often needed. Briefly, the cDNA of interest is subcloned in-frame with an N/C-terminal polyhistidine sequence downstream of the T7 bacteriophage promoter. The plasmid is transformed into an E. coli strain such as BL21(DE3) that expresses the T7 RNA polymerase under an IPTG (isopropyl-β-D-thiogalactopyranoside)-inducible promoter. This expression system allows better repression of the transgene than direct expression system under a pTac promoter. The expressed protein is then purified on a Ni2+-resin. For caspase-1,-4, and -5, an alternative expression and purification procedure has been developed because these caspases, expressed as described in the present unit, tend to be insoluble. The alternative procedure utilizes refolding of individually expressed large and small catalytic subunits, followed by affinity purification using tetrapeptide aldehyde chromatography (Garcia-Calvo et al., 1999). This unit describes a procedure for expression and purification of caspase-2, -3, -6, -7, -8, -9, and -10 (see Basic Protocol 1). Expression conditions vary depending on whether one wants to obtain the zymogen or the active form of the enzyme, but some general rules apply to all and are discussed here.
Peptidases
Contributed by Jean-Bernard Denault and Guy S. Salvesen Current Protocols in Protein Science (2002) 21.13.1-21.13.15
21.13.1
Copyright © 2002 by John Wiley & Sons, Inc.
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STRATEGIC PLANNING The design of the DNA construct in the expression vector is of paramount importance for correct expression and depends on which caspase and which form of the caspase is desired. For example, full-length executioner caspases (-3, -6, and -7) should be expressed as C-terminal His-tag fusion proteins because the N-terminal peptide is proteolytically removed during expression. Expression of caspase-2, -8 -9, -10 is greatly augmented if the N-terminal domain is removed. This is because the death effector domains (DEDs) and caspase recruitment domain (CARD) of initiator caspases probably oligomerize, resulting in insoluble protein, or an increase of toxicity of such proteins in E. coli. For most cases, it is appropriate to use a truncated version, ∆DEDs-caspase-8∆ (Stennicke and Salvesen, 1999) or CARD-caspase-9∆ (Stennicke et al., 1999), with an N-terminal His-tag, although C-terminus His-tagged ∆CARD-caspase-9 could work. Vectors such as pET-23b(+) for C-terminal His-tag is routinely used with the desired cDNA subcloned into the NdeI and XhoI restriction sites, whereas N-terminal His-tag constructs are made by subcloning an NdeI-BamHI cDNA into pET-15b. In the former case, the NdeI site is used because it encodes an initiating methionine close to the ribosome binding site. BASIC PROTOCOL 1
EXPRESSION OF CASPASES IN E. coli This procedure describes the expression of recombinant caspase using the pET expression system from Novagen in the E. coli strain BL21(DE3) from Stratagene. Expressed protein is purified on a Ni2+-chelating resin. A BL21(DE3) clone carrying the plasmid of interest is expanded to the desired culture volume in a rich broth, and expression of the recombinant protein is induced by the addition of IPTG to the growing culture during log phase. After the necessary expression period, bacteria are harvested and cell lysate is prepared by sonication. The purification protocol described here is not different in principle from many other protocols dealing with histidine affinity tag (UNIT 9.4), but has been optimized for caspases. NOTE: The key to success with this procedure is the selection of appropriate expression conditions for a given clone. Because some caspases are toxic to E. coli, plasmids are unstable despite the relatively tight regulation of that expression system even if further repression is imposed using pLysS (see Commentary). It is thus strongly suggested that a freshly transformed colony be used. It is important that the plasmid be maintained in a strain that does not contain the DE3 gene (T7 RNA polymerase) to avoid selection of deleterious mutations of the plasmid. Materials Freshly transformed (UNIT 10.1) BL21(DE3) cells (Stratagene) carrying the appropriate construct, on selective plates 2× TY medium (APPENDIX 4A) containing 100 µg/ml ampicillin (added from 50 mg/ml ampicillin stock) 1 M 1-isopropyl-β-D-thiogalactopyranoside (IPTG) stock solution Chelating Sepharose Fast Flow Resin (Amersham Pharmacia Biotech) Resuspension buffer (see recipe) 0.1 M NiSO4, filter-sterilized Washing buffer (see recipe) Elution buffer (see recipe)
Expression, Purification, and Characterization of Caspases
15-ml culture tubes Orbital shaker incubator (with speed up to 275 rpm preferred) Bacterial culture flasks (flask volume depends on the final culture volume) 1-liter baffled culture flasks
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Spectrophotometer cuvettes with 1-cm light path 0.5-liter centrifuge bottles or tangential flow concentrator system Refrigerated centrifuges with Sorvall SLA-3000 and SM-34 or equivalent rotors 50-ml polypropylene disposable screw-cap tubes Sonicator with large probe Disposable filter units (e.g., Millipore Stericup 0.45-µm HV Durapore membrane or equivalent) Chromatography column: e.g., Econo-Pac 0.7 × 5.0 cm (Bio-Rad) with funnel and flow adaptor, or equivalent setup Gradient former (e.g., Life Technologies Model 150) Peristaltic pump (able to deliver a flow rate of 1.0 to 1.5 ml/min) Fraction collector NOTE: All reagents and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly. Prepare culture and induce caspase expression 1. In a 15-ml culture tube, inoculate 2 ml of 2× TY medium containing 100 µg/ml ampicillin with a small to medium colony of freshly transformed BL21(DE3) cells. Incubate ∼8 hr at 37°C with vigorous shaking (225 rpm) in an orbital shaker incubator. This is the primary culture.
2. In a bacterial culture flask, dilute the primary culture 100-fold into fresh 2× TY medium containing 100 µg/ml ampicillin and incubate as in step 1 for ∼16 hr (overnight). Allow slightly more than 20 ml for each liter of final culture, to compensate for any evaporation. This is the secondary culture. If baffled flasks are used, a culture to flask volume ratio of 1⁄2 or less is recommended; otherwise a ratio of 1⁄5 or less is preferable for other types of flasks.
Table 21.13.1
Expression Parameters for Various Caspases and Intermediate Formsa
Caspases
Forms
Casp-3
Active Zymogen C285A Active C285A Active Full-length zymogen Zymogen N peptide C285A Active Active Active C285A Active
Casp-6 Casp-6 Casp-7
∆DEDs-Casp-8d Casp-9d ∆CARD-Casp-9d Casp-10d
IPTG (mM)
Expression time (hr)
0.2 0.5 0.5 0.02-0.05 0.5 0.2 0.2 0.2 0.5 0.2 0.4 0.4 0.4 0.02-0.05
4-6 0.5b 4-6 16c 4-6 16c 0.5 1.5-2 4-6 4-6 3-5 3-5 4-6 3-5
aThe table only provides suggested starting points; actual parameters should be optimized. bActivation of caspase-3 is extremely effective so it is not possible to obtain the zymogen form with the N-peptide
still attached unless mutations are made at processing sites. cOvernight expression is necessary to obtain full conversion of the zymogen to the active enzyme state. dThe high zymogenicity of initiator caspases does not allow expression of these zymogen forms.
Peptidases
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3. Set up as many 1-liter baffled culture flasks as needed (each flask should contain no more than 0.5 liter of medium) by diluting the secondary culture 50-fold into 2× TY medium containing 100 µg/ml ampicillin and incubate as in step 1 until the optical density at 600 nm (OD600) is between 0.5 and 0.7 (∼2 to 4 hr). Use sterile medium as a spectrophotometer blank. The culture growth may exhibit an initial lag, but the OD600 will increase rapidly once the cells resume growing (doubling time of ∼30 min). The time needed to reach the required OD600 is not necessarily an indication of yield or toxicity. Total growth time may vary significantly between experiments, but should not exceed 7 hr. The baffled flasks increase aeration, which favors growth and protein expression.
4. Induce expression by adding to each flask a sufficient quantity of 1 M IPTG stock solution to achieve the appropriate final concentration (Table 21.13.1). Incubate at 30°C with vigorous shaking (250 to 275 rpm) for the appropriate period of time (indicated in Table 21.13.1). Expression at 30°C with vigorous shaking is preferred because it slows growth and protein expression, allowing more time for protein folding. The time of expression may vary from 30 min to 16 hr, depending on the protein form needed (see Table 21.13.1).
5. Once the expression period is over, transfer the culture to 0.5-liter centrifuge bottles and centrifuge cells 10 min at 3900 × g (5000 rpm in a Sorvall SLA-3000 rotor or equivalent), 4°C. If more than 3 liters are to be harvested, cleared medium can be removed from the centrifuged bottles and more of the culture can be added on top of the cell pellet for recentrifugation as above. Alternatively, a tangential flow concentrator apparatus (UNIT 4.4) can be used and cells washed with resuspension buffer.
6. Resuspend cell pellet in 10 to 20 ml of 4°C resuspension buffer per liter of original culture volume. Purify immediately or store at −20°C (for <1 week) or −80°C (for >1 week up to 6 months) in 50-ml polypropylene disposable screw cap tubes or equivalent (maximum of 45 ml per tube). It is preferable to resuspend cells before freezing because thick cell pellets are more difficult to resuspend. Furthermore, thawing in resuspension buffer will assist cell lysis.
Prepare cell lysate 7. If cells were frozen, thaw them in warm water (<37°C). Place the suspension (contained in a beaker) on an ice bath, and, using an ultrasonic homogenizer, sonicate 2 min at full power with 50% duty cycle (on for 0.5 sec, then off for 0.5 sec). The time needed for proper lysis depends on the volume used and the amount of cells. A 2-min sonication is generally sufficient for 50 ml of cell suspension. At first, viscosity will increase due to release of genomic DNA, but will diminish as DNA is sheared by sonication. The cells are usually adequately lysed when the suspension turns a slightly darker color and becomes clearer. To minimize proteolysis in the sample, it is essential to keep the cells on ice throughout the sonication procedure. Avoid excessive sonication, as this can lead to copurification of E. coli host proteins. Avoid frothing during sonication, which can denature the fusion protein. CAUTION: Wear sound-protection earmuffs to protect ears from ultrasonic noise. Because sonication will generate some aerosol, use the sonicator in a microbiological hood if possible.
8. Transfer the lysate to centrifuge tubes and centrifuge 30 min at 18,000 × g (12,000 rpm in a Sorvall SM-34 rotor or equivalent), 4°C. Expression, Purification, and Characterization of Caspases
It is highly recommended to check for caspase activity prior to purification. At this point, the purification column can be set up (steps 10 to 14 below).
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9. Filter the supernatant through a 0.45-µm disposable filter unit. Wash the filter unit with 5 ml of resuspension buffer to increase recovery and place solution on ice. This step is essential to remove any debris from the clarified lysate that did not precipitate during the centrifugation step.
Prepare column 10. Pour 1 to 5 ml of Chelating Sepharose Fast Flow resin into an empty chromatography column (1.0 cm or less in diameter; e.g., Econo-Pac 0.7 × 5.0-cm) and let liquid (ethanol) drain. Allow 1 ml of resin for 2 to 5 mg of recombinant protein. The amount of resin should not be <1 ml (flow rate will be too fast) or >5 ml (too slow). Using too much resin will increase nonspecific protein binding and decrease purity of the recombinant protein solution. Because the desired recombinant protein has the highest affinity for the resin, keeping the amount of resin low will increase the purity. Qiagen Ni-NTA superflow and HIS-Select HC Affinity gel from Sigma have also been used with success, but check the protein capacity of each resin.
11. Wash column with 5 bed volumes of Milli-Q water to remove ethanol, and let drain. If using resin already containing Ni2+, skip step 12.
12. Add 2 ml of 0.1 M NiSO4 solution per ml of drained resin. Let the nickel solution drain through. 13. Wash column with 5 bed volumes of Milli-Q water to remove excess NiSO4. The column should be light blue at this point.
14. Equilibrate column with 5 bed volumes of resuspension buffer and transfer the column to a 4°C location, e.g., a cold room. The column is now ready for use.
Purify protein 15. With a funnel tightly attached to the top of the column, pour the filtrate from step 9 onto the equilibrated column and let the lysate drain through the resin by gravity flow. A 3-cm long tube can be attached to the column outlet to increase flow rate if necessary.
16. Wash column with 10 bed volumes of 4°C washing buffer (at least 40 ml) and let drain. This step can be done overnight.
17. Apply 10 ml of 4°C resuspension buffer to the column. Do not let column drain completely. This will reduce the salt concentration from the washing step. The following steps can be carried out at room temperature.
18. With liquid still on top of the resin bed, attach the flow adaptor to the column inlet and connect the column outlet to the fraction collector. Set up a gradient former upstream of a peristaltic pump and connect the pump to the flow adaptor. 19. Elute protein with a 25-ml linear gradient of 0 to 200 mM imidazole using resuspension buffer (no imidazole) and elution buffer (high imidazole). Set flow rate to 1.0 to 1.5 ml/min and collect fractions of 1.0 to 1.5 ml. Keep eluted fractions on ice. A total gradient volume of 25 ml is sufficient to provide good separation of nonspecific proteins and recombinant protein.
20. Optional: Apply an additional 5 ml of elution buffer directly to the column to elute any remaining material.
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Analyze fractions 21. Measure the absorbance of each fraction at 280 nm (without diluting the sample) using a 1-cm-path length quartz cuvette. Store fractions at 4°C until analysis (sooner is better). Protein concentration can be more accurately estimated using the Edelhoch relationship (Edelhoch, 1967), which uses the observed absorbance of Trp, Tyr, and Cys content of the protein as a basis for calculating the extinction coefficient.
22. Analyze a 10- to 20-µl aliquot of each fraction by SDS-PAGE using procedures suitable for small polypeptides (UNIT 10.1). An enzymatic assay (see Basic Protocol 2) can also be performed on fractions of interest, but should not replace the SDS-PAGE analysis. All fractions should be analyzed for each caspase being purified, but, as experience is gained, it may only be necessary to analyze a few fractions.
23. Pool fractions containing the protein of interest, dispense into small aliquots in microcentrifuge tubes, and store at −80°C (for 4 months to 2 years depending on caspases). BASIC PROTOCOL 2
CASPASE ENZYMATIC ASSAY This assay is based on the enzymatic hydrolysis of peptidic substrates and can be used to determine caspase activity in purified fractions, as well as to test for proper enzyme expression in cell lysates prior to purification. The amount of sample needed varies greatly depending on whether it is crude extract or a purified fraction, and on whether it is a highly active caspase (caspase 3), or a caspase with low intrinsic activity (caspase 9), as well as on the level of expression and volume of culture used. Optimal sample quantity should thus be determined empirically for each experiment. This protocol is designed for either a chromogenic assay using p-nitroaniline (pNA) or a fluorogenic assay using 7-amino-4trifluoromethylcoumarin (AFC) or 7-amino-4-methylcoumarin (AMC). Chromogenic assays are ∼100-fold less sensitive, but do not require a fluorometer. If such an instrument is available, AFC is preferred over AMC because it can also be quantitated colorimetrically at 380 nm if desired, and has less interference from cell extracts. N-terminal blocking groups on substrate could be either acetyl- (Ac-) or benzoxycarbonyl (Z-). The preferred peptidic substrates suitable for caspases are listed in Table 21.13.2. It is worth mentioning that the substrates listed in Table 21.13.2 are only preferred substrates and not specific substrates. The same remarks apply for the corresponding caspase inhibitors. There are no specific synthetic caspase substrates or inhibitors; there are just degrees of reactivity, some of them not even very high. Materials 2× caspase buffer (see recipe) Caspase-containing samples of interest Chromogenic or fluorogenic substrate (see recipe and Table 21.13.2) 96-well flat-bottom plate (opaque for fluorescent assays and clear for chromogenic assays) Plate reader (colorimetric or fluorometric) ideally with regulated incubator NOTE: The method below assumes a plate-reading 96-well format instrument. If one is not available, then volumes can be increased proportionally for assays in 1-ml cuvettes.
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1. Add 50 µl of 2× caspase buffer to as many wells of 96-well flat-bottom plate(s) as there are samples to be tested. Also add 50 µl of the buffer to two extra wells, one to measure the effect of the buffer alone (blank) and one that will serve to determine the background signal of the substrate. The final volume per well (in a 96-well plate) will be 100 ìl.
2. Adjust the volume of the caspase-containing sample to 30 µl with Milli-Q water (or equivalent) and add the 30 µl of sample to the appropriate buffer-containing wells. Mix well and incubate 15 min at 37°C to allow activation. Add 30 µl of Milli-Q water (or equivalent) to the background well and 50 µl of Milli-Q water (or equivalent) to the blank well. The incubation allows warming of the sample and reduces the cysteine residues. Some caspase preparations contain high levels of enzymatic activity and it may be necessary to dilute samples by as much as 1000-fold before analysis. This will become clear when undiluted samples rapidly deplete the substrate.
3. Prepare a 5× solution of the appropriate substrate in Milli-Q water and warm to 37°C. The full final concentration of substrate in the assay should be 200 ìM for pNA substrates and 100 ìM for AFC/AMC substrates.
4. With a repeating pipet, add 20 µl of the 5× substrate to each well (with the exception of the blank) and mix thoroughly. For pNA substrates, read the absorbance of samples at 405 to 410 nm. For fluorogenic substrates, use the following parameters: AFC, λEX = 405 nm, λEM = 500 nm; AMC, λEX = 380 nm, λEM = 460 nm. Make all readings at 37°C. A common mistake is not being aware of substrate depletion in the assay, which can lead to significant miscalculation of enzyme activity. It is essential to collect data continuously over 30 to 60 min instead of using end-point analysis because rapid substrate hydrolysis could lead to substrate depletion. The continuous monitoring method also allows more accurate determination of the rate of hydrolysis in the linear portion of the signal-versustime data.
5. Transform the enzymatic activity in the assay in relative fluorescence units (rfu)/min or absorbance units/min to rate of hydrolysis (µM/min) using a standard curve of pNA/AFC/AMC (see Reagents and Solutions, discussion under caspase substrates recipe). Use a straight portion of the signal-versus time curve of each sample to determine the rate of hydrolysis (this avoids substrate depletion effects, which lead to errors in rate measurements). Use the rate of a sample lacking enzyme to correct the obtained value for background hydrolysis of the substrate in the assay conditions. If purification fractions (or pure enzyme sample) are analyzed, the rate of hydrolysis is a direct measurement of the caspase activity in those samples. It could be used to obtain the activity profile of a purification scheme (plot as a histogram of rate for each fraction). If the sample is a cell extract (i.e., an enzyme mixture), the rate represents the sum of all contributing components to the hydrolysis of the given substrate.
TITRATION OF RECOMBINANT CASPASE This protocol allows determination of the active enzyme concentration in a homogenous preparation of caspase. It cannot be used to determine the concentration of a particular caspase from mixtures such as cell extracts because active-site titrants are not specific. It is a two-step assay. First the enzyme is incubated with the titrant in concentrations ranging from 0 to 4 times the estimated caspase concentration, in a minimal volume in assay buffer, for 30 min at 37°C. After enzyme-titrant equilibrium is reached, the assay is diluted by
SUPPORT PROTOCOL
Peptidases
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Table 21.13.2
Preferred Peptidic Substrates Suitable for Caspasesa,b
Caspase
Preferred substrates
Caspase-1 Caspase-2 Caspase-3 Caspase-4 Caspase-5 Caspase-6
Tyr-Val-Ala-Asp Val-Asp-Val-Ala-Aspc Asp-Glu-Val-Asp Tyr-Val-Ala-Asp Tyr-Val-Ala-Asp Val-Glu-Ile-Aspd Asp-Glu-Val-Asp Asp-Glu-Val-Asp Ile-Glu-Thr-Aspd Asp-Glu-Val-Asp Val-Glu-Ile-Asp Leu-Glu-His-Aspd Ile-Glu-Thr-Asp Ile-Glu-Thr-Asp
Caspase-7 Caspase-8
Caspase-9 Caspase-10
aTalanian et al. (1997); Thornberry et al. (1997) Stennicke et al. (2000). Further details can be
found in Stennicke and Salvesen (1999). It is important to keep in mind that the preferred substrate cannot be used to distinguish between caspases in a complex mixture such as animal cell lysate, because they are not specific substrates. bN-terminal blocking groups on substrate could be either acetyl- (Ac-) or benzoxycarbonyl (Z-). cNot yet well established. dFirst substrate is preferred but others are suitable.
the addition of an excess of substrate (100 to 200 µM) and the remaining activity is determined as in a simple enzymatic assay. The titrant of choice is the irreversible caspase inhibitor benzoxycarbonyl-Val-Ala-Asp-fluoromethylketone (Z-VAD-FMK) because it is a reasonably effective, commercially available, and broad-range inhibitor. The concentration of active enzyme is determined by plotting the relative hydrolysis rate against the concentration of Z-VAD-FMK. The x-axis intercept of the tangent to the slope at low titrant concentration gives the active caspase concentration in the assay. The best titration (accurate enzyme concentration measurement) is achieved when all the titrant has reacted with the enzyme. This is mostly obtained at high enzyme concentration; pNA-conjugated substrates are thus better than fluorogenic substrates for titration because, by being less sensitive, they allow the use of more enzyme. Further details can be found in Stennicke and Salvesen (1999). Additional Materials (also see Basic Protocol 2) 10 mM Z-VAD-FMK (see recipe) 250 µM pNA-conjugated caspase substrate (see recipe for caspase substrates) in 1× caspase buffer (see recipe for 2× buffer) 1. Estimate the concentration of caspase in the sample either using the Edelhoch relationship (Edelhoch, 1967; also see Basic Protocol 1, step 21 annotation), or, more simply, by assuming 80% purity and that 1 absorbance unit at 280 nm is equivalent to 1 mg/ml of caspase. Transfer enough sample to a microcentrifuge tube so that the estimated caspase concentration is 200 nM in 1× caspase buffer. Prepare slightly more volume than needed, to account for spillage; typically 200 µl will be needed per 16 wells of a 96-well plate (see step 4). Incubate for 15 min at 37°C to allow activation in the caspase buffer. Expression, Purification, and Characterization of Caspases
2. Using a 10 mM stock solution of Z-VAD-FMK, prepare a 2/3 serial dilution of this inhibitor in 1× caspase buffer to cover a 1000-fold concentration range with 2 µM (1 µM final during inhibition) as the highest concentration.
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A 2/3 serial dilution over 16 wells will titrate an enzyme over a concentration range of 1 ìM to 2 nM.
3. Pipet 10 µl of each serial dilution into the corresponding wells of a 96-well microtiter plate. Also include a sample lacking inhibitor to serve as the uninhibited control sample. 4. Transfer 10 µl of diluted enzyme (from step 1) to each well, mix thoroughly, and incubate for 30 min at 37°C (for caspase-6, incubate 10 to 15 min). 5. With a repeating pipet, rapidly add 80 µl of 250 µM pNA-conjugated substrate diluted in 1× caspase buffer, mix thoroughly, and collect data over a 30-min period (see Basic Protocol 2, step 4). Because the enzyme concentration will be high in the uninhibited sample, substrate depletion may occur within 5 min, especially with caspase-3. It is thus critical to read samples as quickly as possible. One approach is to add substrate starting with wells containing the highest Z-VAD-FMK concentration and perform one titration at a time. Alternatively, the enzyme inhibitor mixtures can be diluted 10-fold before addition of the reporter substrate.
Absorbance at 280 nm
A
B
2.5
kDa
2.0
200 97 66
1.5
45 1.0
31
0.5
21 14 6
minus Npep zymogen large (p20) small (p13/12)
2.5 1
2
3
4
5
6
7
8
9 10 11 12 13 14 15 16 17
5
9
10 11
12
13
14 15
16
Fraction no.
C
D 2.0
0.15
3.4 5.1 7.7 11.6
0.10
17.3 26 0.05
titer is 40.4 nM
2.0
y = -44.6x + 1.8 r 2= 0.99
1.5
1.5 rate (µM/mln)
Absorbance at 405 nm
0 nM
1.0 1.0 0.5 0.0
39
0.5
0
10
20
30
40
50
58.5 132 0.00
0.0 0
5
10
15 Time (min)
20
25
30
0
200
400
600
800
1000
Z-VAD-FMK (nM)
Figure 21.13.1 Purification of caspase-7. Caspase-7 was purified from a 1-liter culture induced with 0.2 mM IPTG for 8 hr. (A) The absorbance at 280 nm of each column fraction and (B) the SDS-PAGE analysis of some fractions are presented. Arrows point to caspase-7-related proteins. (C) Raw data of a typical titration with Z-VAD-FMK of purified caspase-7 and Ac-DEVD-pNA as a substrate. The different rates of substrate hydrolysis between 2 and 15 min (bracket) are linear for all time points and are plotted against the inhibitor concentration to obtain the titration curve (D). The bracket in panel D indicates the values used to obtain the active caspase-7 concentration (see inset) by linear regression through the filled data points. Inclusion of two extra data points (open circles) still results in a high regression coefficient (r2 = 0.98) but the calculated enzyme concentration shifts from 40.4 to 49.6 µM, resulting in overestimation of the active enzyme concentration.
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6. Plot the remaining activity (e.g., measured activity) in relative fluorescence units (rfu) or absorbance units against the Z-VAD-FMK molar concentration of the inhibition reaction (step 4). Extrapolate the tangent of the curve at low titrant concentration and read the x-axis intercept to determine the concentration of active enzyme in the assay (see Fig. 21.13.1D). Multiply this value by the dilution factor used to prepare the enzyme sample (step 1). The value obtained is the active enzyme concentration in the initial, undiluted sample.
7. Optional: To obtain a more accurate active-enzyme concentration, repeat the titration using 16 concentrations of Z-VAD-FMK ranging from 0.1- to 2-fold the initially determined enzyme concentration. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Caspase buffer, 2× 20 mM piperazine-N,N′-bis(2-hydroxypropanesulfonic acid) (PIPES), pH 7.2 200 mM NaCl 20% sucrose 0.2% (w/v) 3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate (CHAPS) 2 mM EDTA Filter sterilize Store up to 1 year at room temperature On day of use, add reducing agent from freshly prepared stock solution: dithiothreitol (DTT) to 20 mM or 2-mercaptoethanol (2-ME) to 40 mM. Discard buffer at end of the day. Caspase substrates Preferred peptidic substrates suitable for the different caspases are listed in Table 21.13.2. Prepare p-nitroaniline (pNA)-conjugated substrates (Bachem Bioscience) as 20 mM stock solutions in dimethylsulfoxide (DMSO) and prepare AFC/AMC-conjugated substrates (ICN Biochemicals) as 10 mM stock solutions in DMSO. Substrate solutions are stable for at least 1 year at −20°C. IMPORTANT NOTE: The purity of commercially available substrates varies among providers and from lot to lot within the same manufacturer. A simple way to get accurate and reproducible values using commercial substrates is to titrate them against their respective hydrolyzed chromophore (pNA) or fluorophore (AFC/AMC) as follows. 1. A solution of pNA/AFC (in water)/AMC (in ethanol) is used as a reference. Determine the concentration using the molar extinction coefficient (ε) of each compound: pNA ε410 nm = 8,800 M−1cm−1; AFC, ε380 nm = 12,600 M−1cm−1; AMC, ε354 nm = 17,800 M−1cm−1. 2. Dissolve a known amount of substrate, estimated as the amount indicated on the product label, in DMSO to obtain twice the final concentration (e.g., 20 mM instead of 10 mM final for AFC/AMC substrate). Combine a small portion with excess enzyme to obtain complete hydrolysis of the substrate. 3. Serially dilute the pNA/AFC/AMC stock solution in assay buffer and compare the results to the hydrolyzed substrate to determine the actual concentration of “hydrolyzable” substrate in the original stock. 4. Dilute the substrate stock solution to 10/20 mM concentration. Expression, Purification, and Characterization of Caspases
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Elution buffer 50 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 100 mM NaCl 200 mM imidazole Filter sterilize (do not autoclave) Store up to 1 year at room temperature The buffer can be made as a 5× stock solution and diluted in Milli-Q water at time of assay. IMPORTANT NOTE: The purity of the imidazole is very important. It must have a low absorbance at 280 nm. Imidazole meeting ACS specifications is not suitable and will produce significant background absorbance. The reagent needs to have 0.005% β-NADH equivalent as a fluorescence blank (e.g., Sigma cat. no. I-0250).
Resuspension buffer 50 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 100 mM NaCl Autoclave or filter sterilize Store up to 1 year at room temperature The buffer can be made as a 10× stock solution and diluted in Milli-Q water at time of assay.
Washing buffer 50 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 500 mM NaCl Autoclave or filter-sterilize Store up to 1 year at room temperature The buffer can be made as a 10× stock solution and diluted in Milli-Q water at time of assay.
Z-VAD-FMK, 10 mM Prepare 10 mM benzoxycarbonyl-Val-Ala-Asp-fluoromethylketone (Z-VADFMK; ICN Biochemicals) in DMSO. The solution is stable for at least 3 months at −20°C, 3 days at 4°C, and three freeze-thaw cycles without losing activity. It is important to use the unmethylated derivative (ICN Biochemicals) since the methylated form will not react stoichiometrically with caspases. The absolute accuracy of the titration is dependent on the purity of the titrant and the accuracy with which it is weighed. If available, use an analytical microbalance. Assume that the compound is 95% pure, unless otherwise stated on the manufacturer’s data sheet.
COMMENTARY Background Information Purification by immobilized metal ion affinity chromatography (IMAC) was first reported by Porath et al. (1975). Since then, the simplicity and efficacy of this purification system, along with the availability of many expression vectors encoding a polyhistidine tag and relatively inexpensive affinity resin, have greatly contributed to its popularity. This unit offers a simple and quick procedure to obtain milligram amounts of enzymatically pure recombinant caspase. Basic Protocol 1 can be used to express wild-type caspase-2, -3, -6, -7, -8, -9, and -10, as well as numerous mutants of each of them along with other noncaspase proteases and proteins. The procedure
is reproducible as long as satisfactory protein expression is obtained. The pET expression system uses a promoter (T7 bacteriophage promoter) that is not recognized by the endogenous E. coli transcription machinery, thereby reducing leakage of expression. It is the T7 RNA polymerase that is under the regulation of an IPTG-inducible promoter. This system is less likely to leak than one with a simple IPTG-inducible promoter. It is thus a method of choice for caspases whose activity is toxic to E. coli. The expression phase gives highest average yields when induced at 30°C. In most basic expression protocols, the expression step lasts for the time needed for the culture to reach latency and maximal protein yield. In the case of caspases, expression periods can be
Peptidases
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Table 21.13.3
Troubleshooting Guide for Caspase Expression and Purification
Problem
Possible cause
Solution
No expression
Colony picked is too old Bad clone Construct problem
Poor yield
Protein too toxic
Retransform the plasmid Try another clone Try in vitro translation or immunoblot with His-tag antibody Try another clone Optimize IPTG concentration and/or induce expression at OD600 = 0.9-1.1 Use the pLysS plasmid (Stratagene) Express at 22°C and/or use lower IPTG concentration Use less resin
Protein has poor solubility Purified protein not clean Poor recovery form the resin Flow rate too slow
Caspase detected in flow-through
Protein precipitates in the fraction
Caspase not fully processed Caspase-6 activity lost during titration Caspase concentration is overestimated
Caspase concentration is underestimated
Expression, Purification, and Characterization of Caspases
Too much resin Protein still on the column
Add more elution buffer directly to the column DNA not sufficiently sheared Increase sonication time Lysate too concentrated Attach a 3-cm tube to the column to increase flow rate pH of resuspension buffer too low Check pH of buffer and/or lysate with pH paper Not enough resin Apply flow-through to another column Unknown Reload the flow-through on the same column pH too close to isoelectric point Lower the pH of (pI) or protein is too hydrophobic resuspension/washing/elution buffer to 7.2-7.4 Dilute protein as it elutes from the column or increase total gradient volume Increase expression time to allow Insufficient expression or not conversion of zymogen enough time allowed for processing Half-life of caspase-6 is 30 min in Cut incubation time with assay buffer Z-VAD-FMK to 10 min Incomplete titrant-enzyme reaction Use highest possible enzyme concentration Increase time with Z-VAD-FMK Too much zymogen left (zymogen Use a preparation with as little may react with the titrant but will zymogen as possible not hydrolyze substrate) Still some zymogen left Increase expression time to allow conversion of zymogen
varied to obtain particular forms of caspase. For example 30 min of expression yields singlechain pro-caspase-7 with the N-peptide still attached; 1.5 to 2 hr of expression yields single-chain pro-caspase-7 lacking the N-peptide; and 16 hr of expression yields the fully mature enzyme (see Table 21.13.1). Activation of all
caspases in E. coli is autocatalytic. This is because the caspase concentration reached inside the bacteria is high enough (estimated to reach as high as 1 mM) so the low level of enzymatic activity of the zymogen (termed zymogenicity) is sufficient to provoke autoactivation (see UNIT 21.8). For that reason, despite the
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fact that the maximum yield is usually obtained after few hours (e.g., 8 hr), incubating the culture for a longer period of time (overnight) may help conversion of the zymogen to the active form. Sonication is described here as a simple means to generate a bacterial lysate. This method has the advantage of being quick and the additional advantage of being able to shear DNA, so that the clarified lysate is not viscous. However, several alternative procedures could be used to prepare E. coli lysates. In principle, any nondenaturing lysis method—e.g., lysozyme (UNIT 6.5) or French press (UNIT 6.2)— could be substituted for the one described here, with caveats. Lysis protocols that use enzymes to d isru pt the cell wall, such as lysozyme/DNAse/RNAse, are suitable as long as care is taken to choose enzyme supplies that are protease-free. Furthermore, any protocols must be compatible with the chelating resin procedure, should not use strong reducing agents, metallic cations, or chelators, and should have a pH in the range of 7.2 to 8.0. Chemicals that have the property to order water molecules—e.g., cosmotropic compounds such as polyethylene glycols (PEGs) and various ammonium salts—should be avoided since they could modify some intrinsic properties of caspases (J.B. Denault and G.S. Salvesen, unpub. observ.). Finally, protease inhibitors are not necessary for any of the wild-type caspases and probably most of their mutants since it is generally the expressed caspase that constitutes the likely threat to itself. Numerous researchers and manufacturers have described IMAC purification from E. coli lysates. Batch purification of polyhistidinetagged proteins is an option, but it generally results in a less pure protein preparation than a column method. This is mainly because many proteins have affinity for the Ni2+-resin such as the E. coli slyD rotamase (GenBank accession no., P30856) and are not further separated during the elution from the resin. Selective elution from the resin is best achieved using an imidazole gradient to elute a column. Although a pH gradient would theoretically yield similar results, there is danger of protein precipitation if the pH is too close to the isoelectric point (pI) of the recombinant protein.
Critical Parameters Because caspases are toxic to E. coli, plasmids are unstable in some expression strains such as BL21(DE3). For this reason, it is recommended that plasmids be manipulated and
stored in nonexpressing strains, and that only freshly transformed clones be used. It is not unusual to try two or three clones before finding one that gives good expression, especially since cell lysates can easily be assayed for caspase activity before purification. To reduce leakage of protein expression, co-maintenance of the pLysS plasmid (Stratagene) along with the caspase expression vector can be used. This was found to assist expression of large amount of caspase-3 in several instances. pLysS is maintained in a standard E. coli stain such as DH5α or TG1 in the presence of 25 µg/ml of chloramphenicol. A strain of BL21(DE3) carrying the pLysS plasmid can also be established, but should not be relied on to propagate the plasmid. In general, low amounts of recombinant protein in the cell lysate will allow more nonspecific proteins to bind to the unoccupied resin, resulting in poor purification (e.g., compare Fig. 21.13.1B and Fig. 21.13.2E). The best purifications are achieved when the amount of recombinant protein in the lysate matches as closely as possible the capacity of the amount of resin used. It is thus suggested that not too much resin be used. It is noteworthy that any affinity purification scheme should employ a buffer system compatible with the reagents used in the IMAC procedure (e.g., without metallic cations, chelating agents, or strong reducing chemicals). See UNIT 9.4 for a more thorough discussion of factors affecting metal chelate affinity chromatography. Because the purification procedure relies on the affinity of a polyhistidine sequence toward a chelated metallic ion, pH of the bacterial lysate is critical. At pH 8.0, histidine residues are fully charged. Nevertheless, the pH can be modified if the recombinant protein is likely to precipitate at pH 8.0. For example, fully activated caspase-7 has a pI of 7.8, so purification is performed at pH 7.2 (but not lower).
Troubleshooting A list of potential problems, causes, and solutions for expression, purification, and characterization of caspases is given in Table 21.13.3.
Anticipated Results A complete expression example is shown in Figure 21.13.1 and several examples of purification analysis are shown in Figure 21.13.2. Expression of active caspases should produce a large and a small subunit (Figure 21.13.2A, B and D), but in many cases multiple large
Peptidases
21.13.13 Current Protocols in Protein Science
Supplement 30
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Figure 21.13.2 Purification profiles of various caspase proteins. Caspases were purified using parameters outlined in Table 21.13.1. Proteins were separated by gradient SDS-PAGE using the 2-amino-2-methyl-1,3-propanediol buffer system (Bury, 1981). Arrows indicate both large and small subunits with the approximate protein mass in parentheses. In some cases more than one large (∆CARD-Casp-9) or small (Casp-7) subunit are obtained. The asterisks (Casp-6, ∆DEDs-Casp-8, and Casp-9) indicate caspase zymogen or a processing intermediate that has not fully converted to large and small subunits.
Expression, Purification, and Characterization of Caspases
(Figure 21.13.2F) or small (Figure 21.13.2E) subunits may be observed. Most likely it is the caspase activity itself that generates these extra bands by proteolysis of the various processing sites found in the linker region of caspases (see UNIT 21.8). Furthermore, it should be kept in mind that biosynthesis of an active caspase molecule occurs through the proteolysis of a zymogen and all processing intermediates could be found in a given preparation (asterisks in Figure 21.13.2B, C, and D). For example, caspase-6 does not remove it N-peptide efficiently, so it is often observed that two large subunits are present, one of them still containing the N-peptide (Figure 21.13.2B, asterisk). Yield may vary greatly from one caspase to another and for the various forms being expressed, but it is the rule that 1 mg/liter of recombinant protein can typically be obtained without significant problems.
Time Considerations Titration takes about 1 hr. Setting up the cultures requires very little time (15 min). The entire purification procedure can be completed in less than 8 hr, depending on the volume being processed. Overall, the entire protocol can be accomplished in 3 days including the time needed for growing cultures.
Literature Cited Bury, A. 1981. Analysis of protein and peptide mixtures: Evaluation of three sodium dodecyl sulphate-polyacrylamide gel electrophoresis buffer systems. J. Chromatog. 213:491-500. Edelhoch, H. 1967. Spectroscopic determination of tryptophan and tyrosine in proteins. Biochemistry 6:1948–1954. Garcia-Calvo, M., Peterson, E.P., Rasper, D.M., Vaillancourt, J.P., Zamboni, R., Nicholson, D. W., and Thornberry, N. A. 1999. Purification and catalytic properties of human caspase family members. Cell Death Diff. 6:362-369. Kuida, K., Lippke, J.A., Ku, G., Harding, M.W., Livingston, D.J., Su, M.S.S., and Flavell, R.A.
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1995. Altered cytokine export and apoptosis in mice deficient in interleukin-1-β converting enzyme. Science 267:2000-2003.
rescent peptide substrates disclose the subsite preferences of human caspases 1, 3, 6, 7 and 8. Biochem. J. 350:563-568.
Li, P., Allen, H., Bannerjee, S., Franklin, S., Herzog, L., Johnston, C., McDowell, J., Paskind, M., Rodman, L., Salfeld, J., Towne, E., Tracey, D., Wardwell, S., Wei, F.-Y., Wong, W., Kamen, R., and Seshardi, T. 1995. Mice deficient in IL-1βconverting enzyme are defective in production of mature IL-1β and resistant to endotoxic shock. Cell 80:401-411.
Talanian, R.V., Quinlan, C., Trautz, S., Hackett, M.C., Mankovich, J.A., Banach, D., Ghayur, T., Brady, K.D., and Wong, W.W. 1997. Substrate specificities of caspase family proteases. J. Biol. Chem. 272:9677-9682.
Porath, J., Carlsson, J., Olsson, I., and Belfrage, G. 1975. Metal chelate affinity chromatography, a new approach to protein fractionation. Nature 258:598-599. Renatus, M., Stennicke, H.R., Scott, F.L., Liddington, R.C., and Salvesen, G.S. 2001. Dimer formation drives the activation of the cell death protease caspase 9. Proc. Natl. Acad. Sci. U.S.A. 98:14250-14255. Stennicke, H.R. and Salvesen, G.S. 1999. Caspases: Preparation and characterization. Methods 17:313-319.
Thornberry, N.A., Rano, T.A., Peterson, E.P., Rasper, D.M., Timkey, T., Garcia-Calvo, M., Houtzager, V.M., Nordstrom, P.A., Roy, S., Vaillancourt, J.P., Chapman, K.T., and Nicholson, D.W. 1997. A combinatorial approach defines specificities of members of the caspase family and granzyme B: Functional relationships established for key mediators of apoptosis. J. Biol. Chem. 272:17907-17911.
Contributed by Jean-Bernard Denault and Guy S. Salvesen The Burnham Institute La Jolla, California
Stennicke, H.R., Renatus, M., Meldal, M., and Salvesen, G.S. 2000. Internally quenched fluo-
Peptidases
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Expression, Purification, and Characterization of Aspartic Endopeptidases: Plasmodium Plasmepsins and “Short” Recombinant Human Pseudocathepsin
UNIT 21.14
Aspartic endopeptidases are a class of proteolytic enzymes found in mammals, plants, fungi, and viruses. Obtaining sufficient quantities of aspartic peptidases for kinetic and structural studies has long been a significant obstacle. Although these peptidases can often be purified from their natural hosts, recombinant methods provide an expedient way to obtain large amounts of these enzymes. This unit describes methods for the expression and purification of recombinant aspartic endopeptidases, specifically malarial plasmepsins and a nonglycosylated, single-chain procathepsin D termed “short” pseudocathepsin D (Beyer and Dunn, 1996). The cDNA of the desired peptidase is cloned in-frame into a pET3a expression system (Conner and Udey, 1990), and is subsequently transformed into an E. coli strain, such as BL21(DE3) or BL21(DE3)pLysS cells, for high-level expression of the recombinant protein. The cell lines containing λDE3 lysogen express T7 RNA polymerase under the control of an isopropyl-β-D-thiogalactopyranoside (IPTG)–inducible promoter, which permits controlled expression of the target peptidase. At high cell density in rich medium, the recombinant protein accumulates in intracytoplasmic inclusion bodies and can be isolated from the insoluble fraction of lysed cells, solubilized, and refolded (Conner and Udey, 1990). The refolded zymogens are purified to homogeneity utilizing anion-exchange chromatography in conjunction with gel filtration or pepstatinyl-agarose affinity chromatography to remove improperly/incompletely folded forms of the protein. The protocols in this unit address the expression, refolding and purification of the known malarial aspartic peptidase zymogens of P. vivax, P. ovale, and P. malariae, as well as plasmepsins 2 and 4 from P. falciparum (Westling et al., 1997, 1999) and human “short” pseudocathepsin D. The Basic Protocol describes the expression of these proteins in E. coli; Support Protocols 1, 2, 3, and 4 describe assays for the enzymes in the various fractions of purification. STRATEGIC PLANNING The construction and expression of recombinant proteins has become a routine process, and there are currently numerous different expression systems in which recombinant proteins can be produced. The use of plasmids to express soluble proteins with histidine tags, or fusion proteins, can be combined with affinity chromatography to yield substantial amounts of a homogeneous protein of interest. Alternatively, recombinant proteins can be expressed as inclusion bodies in vectors such as pET3a and subsequently refolded into an active protein. The latter method has proven to be a viable way to express aspartic endopeptidases, including the plasmepsins and cathepsin D. Inclusion bodies must be purified from the expression host, denatured under reducing conditions, and refolded into a conformationally active enzyme. The refolding of denatured protein is an inefficient process; therefore it is suggested that detailed experiments be conducted to optimize several critical parameters that affect the refolding yield. The solubilization and refolding conditions are a function of the physical and chemical properties of the particular protein. Thus, pilot experiments should be performed to optimize the protein and urea concentration, temperature, ionic strength, pH, thiol-shuffling reagent ratios, and other parameters (see UNITS 6.4 & 6.5) for different proteins and variants of the same protein. Also, it is vital Peptidases Contributed by Bret B. Beyer, Nathan E. Goldfarb, and Ben M. Dunn Current Protocols in Protein Science (2003) 21.14.1-21.14.24 Copyright © 2003 by John Wiley & Sons, Inc.
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to perform refolding time-course experiments (UNIT 6.5) to determine the optimal time for refolding. There is a balance between refolding and stability that must be evaluated to ensure maximal purification yield. Once the optimal refolding conditions are determined, the target protein must be purified with chromatography to remove any contaminating proteins or conformational variants of the target protein. Since the pET3a expression vector does not incorporate a polyhistidine tag or fusion protein, the purification methods are limited. Anion-exchange, gel-filtration, and pepstatinyl-agarose affinity chromatography are sufficient to purify the aspartic peptidase zymogens described in this unit. If purification of the mature enzyme is desired, the optimal point at which to renature/activate the enzyme must be determined. As with refolding, it is suggested that activation conditions be scrutinized when studying different variants. Finally, synthesis of pepstatinyl-agarose is strongly recommended over purchasing the reagent. The purchased material tends to be less efficient in binding cathepsin D, as a result of weaker coupling of the pepstatin to the amino-hexyl agarose. Synthesized material made according to the method of Huang et al. (1979) should provide a resin capable of binding 4 mg cathepsin D per ml resuspended resin. BASIC PROTOCOL
EXPRESSION OF ASPARTIC ENDOPEPTIDASES IN E. COLI This procedure describes the expression and purification of recombinant aspartic endopeptidases using the pET expression system (Novagen) in the BL21(DE3) strains of E. coli (Stratagene). After an induction period with IPTG, bacteria are harvested and cell lysate is prepared with a French pressure cell. The inclusion bodies are isolated from lysed cells by a series of centrifugations, solubilized under denaturing conditions, and refolded by dialysis. Post-dialysate containing mostly the protein of interest is then subjected to column chromatography to yield a homogeneous enzyme sample. The cells used for inoculating the expression flasks should be from a cell stock of a single positive colony stored at −80°C in 15% glycerol. The use of a cell stock is strongly recommended, as this reduces the need to use freshly transformed colonies (UNIT 5.2).
Expression, Purification, and Characterization of Aspartic Endopeptidases
Materials E. coli BL21(DE3) cells (Stratagene), growing on LB plates (APPENDIX 4A), containing plasmepsin gene in pET3a or pTCPSD2 construct of cathepsin D (see APPENDIX 4D for introduction of plasmid DNA into E. coli) LB medium (APPENDIX 4A) Glycerol (spectrophotometric grade), sterile M9 medium (for cathepsin D; APPENDIX 4A) 50 mg/ml ampicillin stock 1 M isopropyl-β-D-thiogalactopyranoside stock (IPTG; UNIT 5.2) 6× SDS sample buffer (UNIT 10.1) 95% ethanol Resuspension buffers A, B, C, and D (see recipes; for plasmepsins) or A2 and B2 (see recipes; for cathepsin D); prechilled DNase I (Sigma) 27% (w/v) sucrose (density, 1.1 g/ml) TE buffer, pH 8.0 (APPENDIX 2E) Solubilization buffer A (for plasmepsins; see recipe) or solubilization buffer A2 (for cathepsin D; see recipe) Dialysis buffer A (for plasmepsins): 20 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) IEX starting buffer (for plasmepsins): 20 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E), filter-sterilized IEX elution buffer (for plasmepsins): 20 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E)/1 M NaCl, filter-sterilized
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Superdex 75 prep grade gel filtration resin (optional, for plasmepsins; Amersham Pharmacia Biotech) Rapid dilution buffer (for cathepsin D): 10 mM Tris⋅Cl, pH 8.7 (APPENDIX 2E; filter through Whatman no. 4 filter paper to remove particulates that may act as seeds for protein aggregation) Oxidized glutathione (for cathepsin D; Sigma) 95% formic acid (for cathepsin D; Sigma) Pepstatinyl-agarose affinity resin (for cathepsin D; synthesized by the method of Huang et al., 1979) Pepstatinyl-agarose equilibration buffer (for cathepsin D; see recipe), 25°C Pepstatinyl-agarose washing buffer (for cathepsin D; see recipe), 25°C Pepstatinyl-agarose elution buffer (for cathepsin D; see recipe), 4°C DEAE equilibration buffer (for cathepsin D; see recipe), 4°C DEAE buffer A (for cathepsin D): 10 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) DEAE buffer B (for cathepsin D): 10 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E)/80 mM NaCl, pH 8.0 15-ml capped culture tubes Orbital shaker incubator (preferably with speed up to 275 rpm) 250-ml baffled bacterial culture flask (or larger flask depending on final culture volume) 2.8- or 4-liter Erlenmeyer flasks 1-ml syringe with 27-G needle 0.5-liter centrifuge bottles Beckman refrigerated centrifuge with JA-20 rotor (or equivalent) 50-ml disposable plastic conical screw-cap centrifuge tubes 30-ml Corex centrifuge tubes with rubber jackets Dialysis tubing (for plasmepsins), SpectraPor, MWCO 12,000 to 14,000 (smaller MWCO can be used if necessary): prepare as described in manufacturer’s instructions and APPENDIX 3B FPLC system (Amersham Pharmacia Biotech) including: 5-ml HiTrap Q Sepharose HP anion-exchange column Computer-controlled linear gradient mixer Peristaltic pump capable of 1.0 to 1.5 ml/min flow rate Fraction collector UV spectrophotometer and 1-cm path-length quartz cuvettes 1.5 × 100–cm FPLC column with flow adapter (for plasmepsins, if optional gel-filtration step is to be performed; available from Bio-Rad) C 16/20 chromatography column (for cathepsin D; Amersham Pharmacia Biotech) Büchner funnel (for cathepsin D) Whatman no. 4 filter paper (for cathepsin D) Dialysis tubing (for cathepsin D), MWCO 6000 to 8000: prepare as described in manufacturer’s instructions and APPENDIX 3B Additional reagents and equipment for transformation (APPENDIX 4D) and growth of E. coli (APPENDIX 4B), isolating plasmid DNA (APPENDIX 4C), DNA sequencing (Ausubel et al., 2003, Chapter 7), monitoring bacterial growth by spectrophotometry (UNIT 5.3), SDS-PAGE (UNITS 10.1-10.4), lysis of bacteria using a French pressure cell (UNIT 6.2), dialysis (APPENDIX 3B), measuring protein concentration by spectrophotometry (UNIT 3.4), calculating protein concentration (UNIT 7.2), activity assays for plasmepsins (Support Protocols 1 and 2), fluorogenic assay for cathpsin D (Support Protocol 3), silver staining of protein gels (UNIT 10.5), and titration of recombinant cathepsin D (Support Protocol 4) NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly.
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Set up culture and express protein This protocol is for a 1-liter expression. 1. Identify and store positive clones as follows. a. Pick several individual colonies of the expression cell line harboring the plasmid construct of interest from a newly transformed LB plate (see APPENDIX 4B & 4D). b. Grow separate overnight cultures in 5 ml of liquid LB medium (APPENDIX 4A & 4B) in 15-ml capped culture tubes at 37°C for 12 to 16 hr with vigorous shaking (225 rpm). c. The next day, remove 850 µl of each overnight culture and transfer to separate sterile microcentrifuge tubes each containing 150 µl of sterile glycerol. Vortex and store at −80°C. d. Verify positive clones by isolating plasmid (APPENDIX 4C) and sequencing the DNA insert (see Chapter 7 in Ausubel et al., 2003). Use positive clones as a cell stock for the inoculation of expression medium. 2. Using aseptic technique, pipet 50 ml of LB medium (for plasmepsins) or 40 ml of M9 medium (for cathepsin D) into a 250-ml baffled flask and add 50 mg/ml ampicillin stock to a final concentration of 50 µg/ml. Inoculate medium with a loop of bacteria from a positive cell stock (APPENDIX 4B) and incubate at 37°C for 12 to 16 hr with vigorous shaking (225 rpm). 3. Prepare a 2.8- or 4-liter flask containing 1 liter of LB medium with 50 µg/ml ampicillin (added from 50 mg/ml stock). Inoculate flask with 40 ml of the overnight bacterial culture from step 2, thereby resulting in a 4% inoculation. Incubate at 37°C with vigorous shaking until the optical density at 600 nm (OD600) is between 0.5 and 0.7 absorbance units (AU; UNIT 5.3), using sterile LB medium as a blank. If the enzyme is toxic to the cells, which is manifested as failure of the cells to grow, it may be necessary to inoculate at a lower level, i.e., 1% or 2%.
4. Transfer a 1-ml aliquot of the culture to a microcentrifuge tube (for later SDS-PAGE analysis; UNITS 10.1-10.4) and record the OD600 for this sample (determined in step 3). Add 1 M IPTG stock to final concentration of 1 mM for all plasmepsins, or to a final concentration of 0.5 mM for cathepsin D, to each of the flasks prepared in step 3, to induce expression of the target protein. Incubate at 37°C with vigorous shaking (250 to 275 rpm). 5. At 1, 2, and 3 hr after addition of IPTG, record the OD600 of each sample and aliquot 1 ml of the culture into a microcentrifuge tube. Immediately microcentrifuge the 1-ml aliquots for 2 min at maximum speed. Aspirate supernatant and resuspend the pellet in a solution of 100 µl of TE buffer, pH 8.0, plus 20 µl of SDS sample buffer. 6. Lyse the cells by passing through a 1-ml syringe equipped with a 27-G needle. Between samples, wash the syringe with 95% ethanol, then deionized, distilled water. Store the samples at −20°C until ready to perform SDS-PAGE analysis. 7. Boil each sample, then perform SDS-PAGE (UNIT 10.1) and observe the extent of protein expression (Fig. 21.14.1), normalizing the intensity of the bands for the protein of interest with respect to the OD600 of the aliquot taken (see steps 3 and 4). Expression, Purification, and Characterization of Aspartic Endopeptidases
If the protein is poorly expressed after 3 hr, then the concentration of IPTG used for protein induction should be investigated and optimized. The addition of IPTG marks the beginning of protein expression induction (T=0) which should be complete by ∼3 hr post-induction (T=3); however, the length of the induction period is is an additional parameter that should be varied to obtain optimal expression levels.
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8. Once the expression period is complete (according to the assessment made in step 7), transfer LB cultures (step 5) into preweighed 0.5-liter centrifuge bottles and harvest cells by centrifuging 10 min at 7700 ×g (8,000 rpm in a Beckman JA-20 rotor), 4°C. Remove supernatant, invert centrifuge bottles on paper towels to remove the bulk of the liquid, and determine weight of cell pellet. If the expression volume is >1 liter for a single clone, decant the cleared supernatant from the centrifugation bottles, add the remaining cell culture on top of the pellet, and continue harvesting the cells. This will reduce the total number of pellet-containing bottles that must be resuspended in the next step.
9. Resuspend the pellet either in chilled resuspension buffer A (for plasmepsins) or chilled resuspension buffer A2 (for cathepsin D) at 4.2 ml of buffer per gram of cell pellet. Add 80 Kunitz units of DNase I for every milliliter of resuspension buffer added to each cell pellet. Purify inclusion bodies immediately (step 10) or store resuspended pellets at –20°C (for <1 week) or –80°C (for >1 week) in 50-ml disposable screw-cap conical tubes or equivalents. Alternately, cell pellets can be frozen at –20°C after completing step 8 and resuspended as described in this step immediately prior to the inclusion body purification. The thick cell pellets resulting from centrifugation are difficult to resuspend. Repeated pipetting and/or vigorous vortexing will be required to obtain the desired suspension.
Isolate inclusion bodies 10. If cells were previously resuspended and frozen in resuspension buffer, thaw them in warm water (<37°C) for 5 min. If cell pellets were frozen directly after centrifugation, resuspend as described in step 9. From this point on, it is important to keep cell lysate on ice (or preferably to perform the whole procedure in a cold room at 4°C) to prevent proteolysis of the target protein.
11. Using a French pressure cell (UNIT 6.2), lyse cells twice using the settings recommended by the manufacturer and collect the lysate in a 250-ml flask. For an SLM-Aminco press with a 20-K manual-fill cell, the maximum pressure is 1000 psi. The pressure induced by this apparatus disintegrates the cell walls of the E. coli, and the cell contents are thereby released into the resuspension buffer. The total number of runs through the press required to achieve cell lysis depends on the total volume of the resuspended cells. Typically, a total of two passes through the pressure cell is sufficient to completely destroy the bacterial cell walls. The cells are usually adequately lysed at the point when the suspension turns a slightly darker color and becomes clearer. To minimize proteolysis in the sample, it is essential to keep the cells on ice throughout the procedure. CAUTION: The French pressure cell utilizes high pressure produced by a hydraulic press, which, if misused, can result in serious bodily injury. Follow all safety procedures outlined in the manufacturer’s instructions and do not exceed the recommended pressures and settings. See UNIT 6.2 for additional detail.
12. Incubate the cell lysates for 15 min at 37°C to allow the DNase I to degrade any genomic DNA. Place the samples on ice after this incubation. 13. Gently layer up to 18 ml of cell lysate over a 10-ml 27% sucrose cushion (density, 1.1 g/ml) in a 30-ml Corex tube. Centrifuge 30 min at 12,000 × g, 4°C, in a swinging-bucket rotor to isolate inclusion bodies that sediment through the sucrose solution (Taylor et al., 1986). Save 100 µl of the resulting supernatant for SDS-PAGE and label it as supernatant 1 (“Sup1”). Remove remaining supernatant and save until the SDS-PAGE in step 17 is to be performed. If the majority of the target protein is found in the inclusion body pellet, then the supernatants can be discarded in subsequent inclusion body purifications. Peptidases
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14. Resuspend each pellet obtained from the previous centrifugation in 5 ml of resuspension buffer B (for plasmepsins) or 5 ml of resuspension buffer B2 (for cathepsin D). Gently layer suspension over a second 10-ml 27% sucrose cushion. Centrifuge again as described in step 13. Retain 100 µl of the resulting supernatant for SDS-PAGE (“Sup2”). Remove remaining supernatant. For cathepsin D, use preweighed 30-ml Corex tubes for this centrifugation. The inclusion body pellet is difficult to resuspend. It is helpful to spread the pellet over the inside of the glass centrifuge tube while washing it down the side. To ensure that adequate resuspension is achieved, the material should be slowly dispensed through a 50-ml serological pipet down the side of the glass tube while inspecting for clumps of inclusion body. Repeat resuspension until no clumps are visible.
15a. For plasmepsins: Resuspend the pellet from step 14 in 15 ml of resuspension buffer C and centrifuge at 12,000 × g for 15 min in a swinging bucket rotor at 4°C. Save 100 µl of the resulting supernatant for SDS-PAGE (“Sup3”). Remove remaining supernatant. Resuspend the pellet in 40 ml of resuspension buffer D and centrifuge in preweighed 50-ml plastic centrifuge tubes for 15 min at 12,000 × g, 4°C. Save 100 µl of the resulting supernatant for SDS-PAGE (“Sup4”); remove the remaining supernatant. 15b. For cathepsin D: Proceed directly to step 16 after aspirating the supernatant produced in step 14. 16. Allow the preweighed tubes to sit inverted on a paper towel for ∼15 min to remove excess buffer from the tubes. Weigh each tube and subtract the tare weight to obtain the total weight of the inclusion bodies. Resuspend the inclusion bodies in TE buffer, pH 8.0, at a final concentration of 50 mg/ml. Divide purified inclusion bodies into 1-ml aliquots in microcentrifuge tubes and store at −80°C. 1 ml is a convenient aliquot volume for solubilization that circumvents unnecessary freeze/thaw cycles.
17. Place 10 µl of the resuspended inclusion bodies in a new 1.5-ml microcentrifuge tube. Add 20 µl SDS sample buffer and 90 µl TE buffer, pH 8.0, to this tube, label as “IBs” (for inclusion bodies), and store at 4°C until ready to perform SDS-PAGE. Add 20 µl of SDS sample buffer to each of the tubes from steps 13 to 15 (Sup1 through Sup4) and store at 4°C. Analyze these samples using SDS-PAGE (UNIT 10.1) to observe the efficiency of inclusion body purification (Fig. 21.14.1). Solubilize inclusion bodies 18. Remove the 50 mg/ml aliquots of inclusion bodies (step 16) from the –80°C freezer and thaw on ice. 19. Slowly add the thawed inclusion bodies dropwise to a sufficient volume of solubilization buffer A (for plasmepsins) or solubilization buffer A2 (for cathepsin D) for a final concentration of 1 mg/ml for plasmepsins or 2 mg/ml for cathepsin D. 20. Allow inclusion bodies to dissolve with gentle stirring for 45 min at room temperature. For the plasmepsins, if solubilization buffer appears to contain undissolved inclusion bodies after stirring, centrifuge the insoluble material at 13,500 × g for 30 min at room temperature. Use the clarified supernatant in the next step. Expression, Purification, and Characterization of Aspartic Endopeptidases
The solubilization conditions for the plasmepsins and cathepsin D have been optimized at room temperature (25°); this parameter may vary for other proteins.
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A
mol. wt. kDa marker T=0 207
T=3
S1
S2
S3
S4
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118 81.0
52.5 rPM2 36.2 29.9
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B kDa
mol. wt. marker
T=0
T=1
T=2
T=3
Sup
IB
81.0
52.5 rHuCatD 36.2
29.9
20.7
Figure 21.14.1 SDS-PAGE analysis of the expression and inclusion body purification of aspartic endopeptidases. (A) A 12% Tris-glycine SDS-PAGE of recombinant plasmepsin 2 zymogen at different stages of expression and purification in E. coli. The gel shows plasmepsin 2 (42 kDa) at time zero of IPTG induction (T=0), 3 hr post–IPTG induction (T=3), supernatants from each centrifugation of the inclusion body purification (S1 to S4), purified inclusion bodies (IB), and post-dialysate (PD). Note the overexpression of plasmepsin 2 at 3 hr post-induction and the relatively pure post-dialysate. Molecular weights of the markers are identified, as well as the bands corresponding to plasmepsin 2. (B) SDS-PAGE analysis depicting overexpression of rHuCatD in E. coli upon addition of IPTG. The far left lane corresponds to molecular weight markers. rHuCatD migrates at 42 kDa in comparison to the markers. Shown in the T=0 lane are endogenous E. coli proteins before induction. Lanes T=1, T=2, and T=3 illustrate the overexpression of rHuCatD. rHuCatD does not get exported to the supernatant, as shown in the lane labeled Sup. Lane IB shows the purity of the inclusion body preparation.
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Refold inclusion bodies For plasmepsin: 21a. Place solubilized protein into prepared dialysis tubing. 22a. Dialyze solubilized protein against 4 liters (for 250 mg of IBs dissolved in 250 ml) of dialysis buffer A (20 mM Tris⋅Cl, pH 8.0) at room temperature for 4 hr with stirring (see APPENDIX 3B for general dialysis protocols). The total volume of dialysis buffer used in the five dialyses performed in steps 22a to 24a should be 80× the volume of solubilization buffer used to dissolve the inclusion bodies in step 19. Thus, a total of 20 liters is used for the dialysis of 250 mg protein dissolved in 250 ml. The volume of dialysis buffer must therefore be scaled according to the amount of inclusion bodies to be refolded.
23a. Place the dialysis tubing into 4 liters of fresh dialysis buffer A and continue the dialysis for an additional 6 hr at 4°C. 24a. Repeat this process three more times for an additional 6 hr (minimum) to 18 hr (maximum) of dialysis at 4°C. If any insoluble protein is present after the dialysis is complete, centrifuge the sample for 30 min at 40,000 × g, 4°C. Save 10 ml of the solution from the final dialysis and label it PD (post-dialysate) for measurement of kinetics (see Support Protocols 1 and 2) and protein gel analysis (Fig. 21.14.1).
25a. Keep post-dialysate at 4°C until chromatography is to be performed. If chromatography cannot be performed immediately after the dialysis, then add spectrophotometric grade glycerol to a final concentration of 20% (v/v) and store at –20°C for up to 1 week.
26a. Set up the FPLC system according to the manufacturer’s instructions. Equilibrate HiTrap Q Sepharose HP anion-exchange column with 5 column volumes of IEX starting buffer (20 mM Tris⋅Cl, pH 8.0, filtered) at a flow rate of up to 5 ml/min. After column equilibration, verify that the pH and ionic strength of the flowthrough are correct. All subsequent column chromatography steps are performed at 4°C.
27a. Load post-dialysate onto column at a flow rate no greater than 2.0 ml/min. Using a higher flow rate may result in decreased binding of the protein to the ion-exchange column, thereby resulting in a loss of protein into the flowthrough. Do not exceed the binding capacity of the column (∼55 mg of protein for the 5-ml HiTrap Q column). There is no set volume of post-dialysate that one should load onto the column since the concentration varies for each dialysis. Instead, obtain the concentration of the protein in the post-dialysate by measuring A280 (UNIT 3.4) and load a volume that results in 50 mg of protein. The concentration of the protein in the sample can be estimated using the Equations 7.2.6 and 7.2.7, given in UNIT 7.2.
28a. Elute protein with a linear gradient of 0 to 1 M NaCl using a computer-controlled gradient mixer that mixes the starting buffer (20 mM Tris⋅Cl/0 M NaCl) and the elution buffer (20 mM Tris⋅Cl/1 M NaCl) over a fixed amount of time (typically 60 min). Set flow rate to 2.0 ml/min and collect 2.0-ml fractions. Keep eluted fractions at 4°C.
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29a. If the FPLC is equipped with a UV monitor, identify the fractions containing protein by examining the absorbance trace. If not, then measure the absorbance of each fraction spectrophotometrically (UNIT 3.4) at 280 nm (without diluting the sample) using a 1-cm path-length quartz cuvette. The concentration of protein in the sample can be estimated using Equations 7.2.6 and 7.2.7 in UNIT 7.2. Keep fractions at 4°C.
30a. Analyze a 10- to 20-µl aliquot of each post–ion exchange fraction by SDS-PAGE (UNIT 10.1). An enzymatic chromogenic assay (Support Protocol 1) can also be performed on fractions of interest but should not replace the SDS-PAGE analysis. For initial purification runs of plasmepsins, all fractions should be analyzed by SDS-PAGE and activity assays (see Support Protocol 1). However, once the elution and activity profile is determined for several purifications, only a few selected fractions are required to be tested for activity.
31a. Pool fractions containing the protein of interest and analyze by SDS-PAGE (UNIT 10.1). If the SDS-PAGE analysis of the purification reveals the plasmepsin protein is not homogeneous, then further purify the pooled fractions by gel filtration (step 32a). Otherwise, pool the desired fractions and quantify the concentration of active enzyme (see Support Protocol 2). Dispense the enzyme solution into small aliquots in microcentrifuge tubes and store at 4°C for up to 3 months or at –80°C in 10% (v/v) spectrophotometric-grade glycerol for up to 2 years. 32a. Optional (if pooled fractions in step 31a are not homogeneous): Pack a 1.5 × 100 cm chromatography column, fitted with a flow adapter, with Superdex 75 prep-grade gel-filtration resin and equilibrate in IEX starting buffer (20 mM Tris⋅Cl, pH 8.0) on the FPLC. Load pooled post–anion exchange fractions (step 31a) onto the column. Separate proteins using a 1 ml/min flow rate of the starting buffer and collect 1-ml fractions. Analyze fractions by SDS-PAGE (UNIT 10.1) and enzymatic chromogenic assay (Support Protocol 1). Pool the desired fractions and quantify the concentration of active enzyme (see Support Protocol 2). Concentration of the anion-exchange fractions using a 10,000 MWCO centrifugation concentrator and/or multiple gel filtration column runs may be necessary to purify the entire post–anion exchange sample.
For cathepsin D 21b. Add solubilized inclusion bodies (from step 20) dropwise into rapid dilution buffer (10 mM Tris⋅Cl, pH 8.7, filtered to removed particulates) while stirring the buffer at a moderate rate to achieve a 100-fold dilution with a final inclusion body concentration of 20 µg/ml. From this point on, the rapid dilution buffer is only stirred when adding oxidized glutathione and during pH titrations. Stirring should be avoided where possible, to avoid aggregation of misfolded protein. When performing a scale-up, it is recommended that a separatory funnel or similar device be used to add inclusion bodies dropwise.
22b. Add oxidized glutathione to a final concentration of 0.5 mM. Stir briefly until dissolved. 23b. Let stand at 25°C for the predetermined optimal refolding time. Pilot experiments should be performed to determine the optimal refolding time for every enzyme and variant.
24b. Activate refolded protein by titrating to pH 3.7 with 95% formic acid for the predetermined optimal activation time. Peptidases
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25b. Dispense an appropriate amount of pepstatinyl-agarose affinity resin into a C 16/20 chromatography column. Equilibrate resin with ∼2 to 5 times the resin bed volume of pepstatinyl-agarose equilibration buffer at a flow rate of at least 1 ml/min (gravity-flow). If the flow rate is slow, the resin may need to be changed. Additionally, the pre-filters on the intake lines often become clogged, as well as the filter on the resin bed support. Change these accessories if necessary. If the column is not to be used immediately, store the resin in 0.02% sodium azide at 4°C to prevent microbial degradation of the resin. Always wear gloves when working with sodium azide.
26b. Clean all inlet and outlet tubing to column by running ethanol and then water through the lines. 27b. Filter the refolded/activated material gently by weak vacuum, so as to avoid frothing (which can promote precipitation of protein), using a Büchner funnel and Whatman no. 4 filter paper, to remove large aggregates of protein and particulates that will clog the affinity resin and result in slow flow rates. 28b. Apply material to column at a flow rate of at least 1 ml/min at 25°C. For large-scale purifications of 4 liters of refolding solution, a flow rate of 2 to 5 ml/min is typically used, with an overnight loading. It is important to monitor the flow-through for the presence of the desired protein. If protein is present in the flow-through, a slower flow rate may be desired. Likewise, it is important not to exceed the capacity of the resin. Approximately 1 ml of resin slurry is sufficient to bind 80 mg of refolded/activated cathepsin D.
29b. Wash the resin with 2 to 5 times the resin bed volume with pepstatinyl-agarose equilibration buffer. Do not overwash the resin. Washing with too much buffer may result in washing off some bound protein.
30b. Wash the resin with 2 to 5 times the resin bed volume with pepstatinyl-agarose washing buffer. The purpose of this wash step is to lower the buffering strength in order to facilitate a sharp pH step elution.
31b. Elute the bound protein in a single step with 4°C pepstatinyl-agarose elution buffer. The elution buffer is the only buffer chilled to 4°C. Cold elution buffer appears to help with protein elution. It is important to elute by adding just enough buffer to cover the resin while maintaining approximately a 1-ml buffer head. This avoids dilution of protein in elution buffer. Store fractions on ice.
32b. Measure activity of eluted fractions using a fluorogenic assay (Support Protocol 3). Continue to collect fractions until no activity is detected. Pool fractions containing desired activity and add spectrophotometric-grade glycerol to a final concentration of 10% (v/v). Freeze material at –20°C. Elution of protein must be evaluated by measuring activity. The detergent present in the pepstatinyl-agarose column buffers interferes with protein concentration determination at 280 nm, so spectrophotometric methods must be avoided.
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33b. Desalt the pooled pepstatinyl-agarose column fractions by placing 35 to 40 ml of sample in a 6000 to 8000 MWCO dialysis tubing and dialyzing (APPENDIX 3B) against 1 liter of DEAE equilibration buffer, 4°C, for several hours, then against 4 liters DEAE equilibration buffer, 4°C, overnight. Complete removal of salt is paramount to the success of the subsequent ion-exchange chromatography. Two dialyses against very large volumes of buffer with constant stirring are sufficient for complete dialysis. One pitfall in dialysis is not leaving sufficient room in the dialysis tubing for the expansion of the bag that will be caused by diffusion. This will result in incomplete dialysis. It is important to leave room for a 20- to 30-ml increase in volume.
34b. Set up the FPLC system according to the manufacturer’s instructions. Equilibrate HiTrap Q Sepharose HP anion-exchange column with 5 column volumes of DEAE buffer A at a flow rate of 1 ml/min. After column equilibration, verify that the pH and ionic strength of the flow-through is correct. All subsequent column chromatography steps are performed at 4°C.
35b. Apply dialyzed sample to column at a flow rate of 1 ml/min at 4°C and collect flow-through for SDS-PAGE and activity analyses. 36b. Wash column with 5 column volumes DEAE buffer A at a flow rate of 1 ml/min, saving wash for SDS-PAGE and activity analyses. 37b. Elute protein at a flow rate of 1 ml/min at 4°C with a linear gradient of 0 to 80 mM NaCl using a computer-controlled gradient mixer that mixes DEAE buffers A and B, for a total of 55 column volumes of gradient. 38b. Monitor eluted fractions for cathepsin D activity as described in Support Protocol 3. Pool fractions containing the enzyme, unless higher concentrations of enzyme are required for subsequent analyses, in which case do not pool. 39b. Analyze purity of material by SDS-PAGE (UNIT 10.1) followed by silver staining (UNIT 10.5). Quantify the concentration of active enzyme (see Support Protocol 4). 40b. Add spectrophotometric-grade glycerol to the pooled enzyme sample to a final concentration of 10% (v/v), divide into 1-ml aliquots, and store at –20°C for up to 2 months. “Short” rHuCatD is stable in solution at 4°C, pH 8.0 for at least a week. This may be a valuable option to avoid freeze/thaw or the addition of cryoprotectant. Otherwise, it is recommended to freeze the samples in 10% glycerol.
ENZYMATIC PLASMEPSIN CHROMOGENIC ASSAY The assay listed below describes the use of a chromogenic octapeptide substrate to test the activity of the post-dialysate and the post–anion exchange fractions obtained from column chromatography (see Basic Protocol). The assay measures the rate of cleavage of the substrate on the amino terminal side of a p-nitophenylalanine chromogenic reporter group, which is translated into a decrease in absorbance over 284 to 324 nm as measured by a diode array spectrophotometer (Dunn et al., 1994; Westling et al., 1997). The substrate sequence used in these assays is Lys-Pro-Ile-Glu-Phe-Nph-Arg-Leu, such that Nph is a p-nitophenylalanine, and is a useful sequence for all the plasmepsins purified to date (Westling et al., 1997, 1999). The use of chromogenic and fluorogenic substrates with aspartic endopeptidases is described in numerous publications (Dunn et al., 1994; Westling, 1997; Yasuda et al., 1999). The amount of enzyme sample needed varies greatly depending on whether it is from post-dialysate or a purified fraction. However, all plasmepsins have relatively the same intrinsic activity, such that the volumes used in the
SUPPORT PROTOCOL 1
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assay should not need to be varied more than 10-fold. This remains true only if similar concentrations of inclusion bodies were denatured/refolded and if comparable amounts of the enzyme were purified from the ion exchange chromatography procedure. Materials Chromogenic plasmepsin substrate stock (see recipe) 5× plasmepsin assay buffer: 0.5 M sodium formate, pH 4.5, filter-sterilized Plasmepsin-containing fraction to be analyzed (see Basic Protocol) Capless 1.5-ml microcentrifuge tubes 1-cm path-length quartz cuvettes Diode array spectrophotometer with a seven-cuvette multi-cell transport thermostatted to 37°C with a circulating water bath 1. Set up assay as follows. The final volume in the 1-cm quartz cuvette at step 3 will be 250 ìl.
a. Using a primary stock solution of substrate, typically with a concentration of 6 to 8 mM as determined by amino acid analysis, make a secondary stock solution of 1250 µM substrate. When prepared as described (see Reagents and Solutions), the concentration of the substrate stock solution is typically 6 to 8 ìM. However, amino acid analysis must always be performed to obtain an exact concentration for the stock, so that secondary dilutions are of known concentration.
b. For each fraction to be tested for activity, aliquot 20 µl of the substrate secondary stock solution into a capless 1.5-ml microcentrifuge tube. This volume of substrate will yield a final substrate concentration in the enzymatic reaction of 100 ìM.
c. To a second series of corresponding microcentrifuge tubes, add 50 µl of 5× plasmepsin assay buffer, 175 µl of sterile-filtered water, and 5 µl of the post-ion exchange fraction to be analyzed for plasmepsin (or other volume, adjusting the water volume to compensate). The final DMSO concentration used to dissolve the peptide substrate (see Reagents and Solutions) should be <0.1% in the activity assay, as a higher concentration could adversely affect enzyme activity. The volume of the enzyme to be added will vary depending upon the concentration of the plasmepsin in the sample. If more or less enzyme is required to observe the cleavage of the substrate, then the volume of the water must be adjusted accordingly to keep the final reaction volume fixed at 250 ìl. Additionally, a master mix of the water and buffer can be made to reduce the manifestation of pipetting error in the kinetic reaction. This is not critical when testing the activity of fractions, but becomes important when determining kinetic parameters with purified enzyme.
2. Incubate both the substrate-containing and the enzyme/buffer/water-containing capless microcentrifuge tubes at 37°C for 3 min. 3. After the incubation period:
Expression, Purification, and Characterization of Aspartic Endopeptidases
a. Add the contents of the tube containing the water, buffer, and enzyme into the corresponding tube containing the substrate. Vortex briefly b. Transfer to a 1-cm path-length quartz cuvette and begin acquiring data. Monitor the average decrease in absorbance on the spectrophotometer from 284 to 324 nm minus the average absorbance at 400 to 420 nm. Allow the reaction proceed for a minimum of 200 sec.
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The resulting data will be a change in absorbance with respect to time, which is used to calculate the initial velocity of the rates of cleavage. The rates of cleavage observed in this assay are proportional to the amount of active enzyme in the sample. The absorbance maximum for a Phe-Nph bond is 278 to 280 nm. As this bond is cleaved, the maximum will shift to 270 to 272 nm. The average decrease in absorbance is measured from 284 to 324 nm over time to obtain a rate for the cleavage reaction. The average decrease in the range of 284 to 324 nm has a significantly larger value than the decrease from 278 to 280 nm, or the increase from 270 to 272 nm; therefore the acquisition of data of the former, 40-nm range results in a more sensitive assay, and also reduces the propagation of errors that could arise from measuring only a single wavelength. It is important to note that not all spectrophotometers have this capability, so a scan must be performed before and after peptide hydrolysis to establish the optimal wavelength for subsequent assays. Subtracting the absorbance from 400 to 420 nm is known as “subtracting the drift.” For a Phe-Nph-containing peptide, the reading from 400 to 420 nm should be close to zero whether the peptide is intact or completely hydrolyzed, but the lamp in the spectrophotometer can gradually change its zero reading for various reasons; the subtraction of the reading from 400 to 420 nm is done to account for these fluctuations. The activity of the enzyme present in each post–ion exchange fraction is determined by the initial velocities of cleavage obtained for these reactions, which is recorded as ∆A/sec. Since the concentration of substrate does not vary for these reactions (step 1b), the different rates correspond to the varying concentration of active enzyme present in each of the post-dialysate or post-IEX fractions.
c. Fit a line tangent to the initial linear portion of the hydrolysis reaction curve so that the calculated slope yields the initial velocity for the reaction (Dunn et al., 1994). Pool purified fractions that show evidence of cleavage and store up to 3 months at 4°C; alternatively add spectrophotometric-grade glycerol to a final concentration of 10% (v/v) and store up to 2 years at –80°C. Selection of samples is a qualitative process; fractions showing activity are pooled, whereas fractions without activity are discarded. The pooled fractions are used in subsequent spectroscopic assays for active-site titrations, KM, and kcat determinations for various peptide substrates, or Ki analyses with aspartic peptidase inhibitors.
TITRATION OF RECOMBINANT PLASMEPSINS This assay allows determination of the total active enzyme concentration in a given sample fraction. The concentration of active enzyme is determined using a highly specific aspartic endopeptidase inhibitor, pepstatin A. The rates of cleavage in the presence of various inhibitor concentrations are fitted to the competitive tight binding inhibition equation for reversible inhibitors [ v = (v0/(2 × E)) × (E – I – Kap + sqrt((E – I – Kap)2 + 4 × E × Kap)) ] in the Enzyme Kinetics Module (Version 1.0) of SigmaPlot 2000 (Version 6.10). In these assays, it is important that the enzyme concentration be much greater than the inhibitor concentration in the assay; if the ratio of free enzyme to the Ki of the inhibitor is ≥100, then almost all of the inhibitor present in the reaction is bound to the enzyme. Under these conditions, the binding is so tight that the enzyme is titrated by the inhibitor (Morrison 1969; Bieth 1995).
SUPPORT PROTOCOL 2
Materials Pepstatin A stock (see recipe) Enzyme Kinetics Module (Version 1.0) of SigmaPlot 2000 (Version 6.10; Sigma) Additional reagents and equipment for enzymatic plasmepsin chromogenic assay (Support Protocol 1)
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1. Prior to the active-site titration described below, perform the plasmepsin chromogenic assay using varying amounts of the fraction to be analyzed (Support Protocol 1) to determine the volume of enzyme to use for the remainder of the titration protocol. This step is necessary since the enzyme can vary in its activity depending on the efficiency of refolding and purification of each preparation. Essentially, the enzyme concentration to be used in the titration is determined by performing the enzymatic plasmepsin assay described in Support Protocol 1 with varying amounts of post-column material (see step 1 of Support Protocol 1). Use a volume of enzyme that yields a readily measurable rate of cleavage, since subsequent reactions will have reduced rates of cleavage due to the presence of pepstatin.
Determine initial velocities of cleavage The first assay (steps 2 to 7) is to determine the initial velocities of cleavage for six different substrate concentrations in the absence of inhibitor with a fixed amount of enzyme (typically post-anion-exchange pooled fractions, determined by screen of various amounts of enzyme) to determine a Michealis constant for this enzyme and substrate. The final assay volume for all reactions in this protocol is 250 µl. 2. Prepare a six dilutions of the plasmepsin substrate such that the concentration of these secondary stocks range from 250 to 1500 µM. When 20 ìl of these secondary stocks are used in a 250 ìl enzymatic reaction, the final concentration of the substrate is 20 to 120 ìM in the assay (Dunn et al., 1994).
3. Into six microcentrifuge tubes, aliquot 20 µl of each secondary stock of the plasmepsin substrate so that each tube has a different concentration (final concentrations of 20 to 120 µM, in 20 µM increments, usually suffice). 4. In a 1.5-ml capped microcentrifuge tube, prepare a reaction cocktail containing the required amounts of water, 5× plasmepsin assay buffer, and purified enzyme (see Support Protocol 1). In order to compensate for pipetting errors, the volume of the reaction cocktail prepared should be greater than the minimum required volume.
5. After mixing the reaction cocktail, pipet 230 µl of it into a second series of six microcentrifuge tubes. Incubate the six tubes containing substrate and the six tubes containing the aliquots of the reaction cocktail at 37°C for 3 min to allow for the conversion of plasmepsin zymogen into mature enzyme. The conversion of zymogen to mature enzyme may vary for different plasmepsins, as well as for different aspartic peptidases. The time and pH required for conversion should be evaluated in preliminary studies using SDS-PAGE analysis (UNIT 10.1).
6. After the incubation period, add the contents of the tube containing the reaction mixture aliquot into the tube containing the substrate. Vortex briefly and transfer to 1-cm quartz cuvettes in the multi-cell transport of the spectrophotometer. 7. Begin scanning the decrease in absorbance on the spectrophotometer from 284 to 324 nm minus the absorbance at 400 to 420 nm. Let the reaction proceed for a minimum of 200 sec. See Support Protocol 1 for details. The resulting data will be a change in absorbance with respect to time, which is used to calculate the initial velocity of the rates of cleavage. The initial velocities for each reaction are plotted versus the corresponding concentration of substrate and fit using the MichaelisMenten equation: Expression, Purification, and Characterization of Aspartic Endopeptidases
vi = (Vmax × [S])/(KM + [S]) The values of Vmax and apparent KM are calculated in the Single Substrate Study option of the Enzyme Kinetics Module of SigmaPlot by entering the substrate concentrations
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analyzed and rates obtained from the spectrophotometer. The plots of vi versus [S] are drawn by selecting “Michaelis-Menten” in the “Equations” and “Graphs” windows within the “Module” window.
Peform enzymatic assays in the presence of inhibitor 8. Set up a second series of reactions as described in the steps above, but include the inhibitor pepstatin in the reaction cocktail (step 4) such that the final concentration is sufficient to give a 25% to 30% decrease in the rates of cleavage observed in the absence of pepstatin, as determined by pilot experiments. Repeat the scan using the inhibitor-containing reaction mix. The absorbance decrease resulting from this run will be plotted to give an apparent increase in Km, which results from the presence of pepstatin. The volume of water in this reaction cocktail must be decreased to compensate for the presence of the inhibitor solution. However, the volume added to the six capless microcentrifuge tubes will remain 230 ìl, so the addition of the substrate will keep the final reaction volume fixed at 250 ìl.
9. Repeat the assay as described in step 8, but increase the inhibitor concentration such that a 50% to 60% decrease in the initial velocities of cleavage (as determined by pilot experiments) is observed with respect to the sample without inhibitor. The initial velocities of this reaction will result in an increased apparent Km that is higher than both of the previous assays.
10. Fit the initial velocities for each substrate/inhibitor concentration to the tight-binding inhibitor equation in the Enzyme Kinetics Module of SigmaPlot. Using the Tight Binding Inhibition option, enter the initial velocity rates for the assay in the absence of inhibitor and the corresponding substrate concentrations (obtained from step 7). Next, sequentially enter the values for the two data sets that contained pepstatin. To complete the analysis, select Michaelis-Menten and Lineweaver-Burk under “Graphs,” which will plot the data sets as vi versus [S] or 1/v versus 1/[S], respectively. Also, select “Competitive Tight” under “Equations” for the program to calculate the values of Vmax and apparent KM (based on the samples without inhibitor) as well as Ki and Etotal using the rates from the samples with inhibitor present. If the enzyme concentration used was sufficiently high, then the equation will yield an accurate concentration of the total active enzyme in the sample with <20% error. If the amount of enzyme in the assay was too low, then the Enzyme Kinetics Module of SigmaPlot will properly fit the data to the tight binding inhibitor equation, and a Ki with a low percent error for the pepstatin inhibitor will result. If this occurs, increase the amount of enzyme used in the assay.
ENZYMATIC CATHEPSIN D FLUOROGENIC ASSAY This protocol describes the use of a fluorescent substrate to detect activity of the refolded/activated cathepsin D in the eluted fractions from Basic Protocol 1. This increase in sensitivity over chromogenic substrates is ideal for situations where protein concentration is low. The substrate contains a highly fluorescent (7-methoxycoumarin-4-yl) acetyl moiety and a quenching 2,4-dinitrophenyl group. Upon cleavage of the Phe-Phe bond, the fluorescence at an excitation wavelength of 328 nm and emission wavelength of 393 increases due to diminished quenching, which in turn results from the separation of the fluorescent and quenching groups (Yasuda et al., 1999).
SUPPORT PROTOCOL 3
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Materials Cathepsin D assay buffer: 1 M sodium formate, pH 3.7 Cathepsin D–containing fraction to be analyzed (see Basic Protocol 1) 100 µM fluorogenic cathepsin D substrate stock (see recipe) 96-well round-bottom microtiter plates Fluorescence multi-well plate reader with regulated incubator Multichannel pipettor 1. Set up assay as follows. The final assay volume in the well is 200 ìl.
a. For each assay to be performed, pipet 40 µl cathepsin D assay buffer, 120 µl water, and 20 µl of eluted fraction into one well. b. For each assay to be performed, pipet 20 µl of 100 µM substrate stock into a separate well. The final concentration of substrate in the assay should be 10 ìM. The final DMSO concentration is <0.1%.
c. Insert plate into reader and incubate for 3 min at 37°C. The fluorescent substrate should be protected from light at all times. Exposure to light can lead to photobleaching of the fluorophore.
2. Eject plate and transfer the buffer/water/enzyme mixtures to the wells containing substrate. Immediately start assay, which includes a 3-sec mixing step, by monitoring an increase in fluorescence intensity produced by substrate cleavage at an emission wavelength of 393 nm with excitation at 328 nm. Ensure that proper filters are used, and collect data for at least 3 min. It may be necessary to increase the volume of eluted fraction in order to observe substrate hydrolysis. This is particularly the case with variants that are less efficient in refolding and activation.
3. Quantitate total enzymatic activity by plotting the initial rates of substrate cleavage as a function of the eluted fraction number, to obtain a peak activity profile. To maximize the sample concentration, assay individual post-column enzyme fractions for active enzyme concentration by active site titration as in Support Protocol 4. Alternatively, the active peak fractions can be pooled together and the final volume recorded. One should be aware that pooling the samples will result in sample dilution, which may be a grave pitfall if the protein concentration is low. In order to determine the total activity of the pooled material, assay an aliquot for activity and calculate the total activity as follows: (measured rate in AFU/sec ÷ volume of material used in activity assay) × total volume of pooled active peak material Note that this is a total activity measurement, not a determination of active enzyme concentration. If an active enzyme concentration is desired, one should proceed with an active site titration as described in Support Protocol 4. SUPPORT PROTOCOL 4
Expression, Purification, and Characterization of Aspartic Endopeptidases
TITRATION OF RECOMBINANT CATHEPSIN D This protocol permits the determination of active enzyme concentration of eluted fractions using a fluorogenic substrate. The amount of active enzyme in each assay is determined by competitive inhibition with the potent inhibitor pepstatin (Isovaleryl-Val-Val-Sta-AlaSta) (Sigma). The resulting curve was fitted with the Dixon equation on the Enzyme Kinetics module of SigmaPlot.
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Materials Pepstatin A stock (see recipe) Enzyme Kinetics Module (Version 1.0) of SigmaPlot 2000 (Version 6.10; Sigma) Additional reagents and equipment for enzymatic cathepsin D fluorogenic assay (Support Protocol 1) 1. Prior to the active-site titration described in the subsequent steps, perform the cathepsin D fluorogenic assay (Support Protocol 3) using varying amounts of the enzyme-containing fraction and 10 µM substrate. This assay permits determination of the necessary amount of enzyme required for the active site titration. It is important to use an enzyme concentration high enough to give reliable rates when incubated with a range of inhibitor concentrations. This is empirically determined and will vary with the success of purification.
2. In a row of wells, make inhibitor dilutions in water in a total volume of 50 µl covering a range of concentrations with at least 5 nM increments. Include a no-inhibitor control for each repeat of the assay. The final assay volume in the well is 200 ìl (see Support Protocol 3). Pipetting accuracy is critical for the success of an active-site titration.
3. Make a cocktail containing the appropriate amount (see Support Protocol 3) of enzyme-containing fraction, cathepsin D assay buffer, and water. It is important to make more cocktail than will be needed to allow for pipetting errors. For example, for 8 wells, the cocktail is scaled up 12.5 times.
4. Add 180 µl of the enzyme, buffer, and water cocktail to each well containing an inhibitor dilution, changing tips each time to avoid carryover of inhibitor. Note that the volume of the cocktail plus inhibitor is 230 ìl. This extra volume will prevent pipetting errors by allowing enough volume to transfer accurately.
5. Add 20 µl of 10 µM cathepsin D substrate to a separate row of 8 wells. 6. Insert plate into reader and incubate for 3 min at 37°C. 7. Set a multichannel pipettor to pick up 200 µl and dispense 180 µl. 8. Carefully pick up 200 µl of the cocktail and dispense 180 µl in rows containing substrate. Inspect pipet tips after picking up the cocktail/inhibitor mix to be sure that all of the tips are picking up an even quantity of liquid. If uneven pipetting is suspected, eject entire sample back into wells and verify that the pipet tips are securely attached to the device.
9. Inset plate and start assay. Record data for at least 5 min. 10. Plot remaining activity as a function of inhibitor concentration and fit with the Dixon equation for tight binding reversible competitive inhibition using the SigmaPlot software. Once conditions are established, this method provides reproducible determinations with 10% to 20% error. If errors are high, check all dilutions and either repeat or add inhibitor concentrations. A common mistake is using to little enzyme that will result in nontitrating conditions. In this case, fitting the data will yield Ki values but not enzyme concentration.
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Cathepsin D substrate stock solution, fluorogenic, 100 ìM Synthesize fluorogenic substrates [e.g., MOCAc-Gly-Lys-Pro-Ile-Leu-Phe-PheArg-Leu-Lys(Dnp)-Arg] using an Applied Biosystems automatic peptide synthesizer (ABI model 430A or 432A). Dissolve in 20% (v/v) DMSO. Centrifuge substrate solution on a 0.45 µm Costar cellulose acetate centrifuge tube filter (Corning) for 5 min at 20,000 × g to remove particulates. Quantify by amino acid analysis on a Beckman System 6300 High Performance Amino Acid Analyzer (also see UNIT 3.2). Store peptides up to 1 month at 4°C (Dunn et al., 1994). Fluorescent substrates should always be protected from light.
Pepstatin stock solution, 0.6 mM Dissolve 10 mg pepstatin A (Isovaleryl-Val-Val-Sta-Ala-Sta; e.g., Sigma) in 1 ml of 90:10 ethanol:acetic acid. Centrifuge the pepstatin A solution on a 0.45 µm Costar cellulose acetate centrifuge tube filter (Corning) for 5 min at 20,000 × g to remove particulates. Quantify by amino acid analysis on a Beckman System 6300 High Performance Amino Acid Analyzer (also see UNIT 3.2). Store inhibitor stock up to 2 months at 4°C, if storing longer, the stock solution should be quantified again by amino acid analysis. Pepstatinyl-agarose equilibration buffer 0.1 M sodium formate, pH 3.7 0.4 M NaCl 0.05% Brij-35 Store up to 6 weeks at room temperature Pepstatinyl-agarose washing buffer 0.01 M sodium formate, pH 3.7 0.4 M NaCl 0.05% Brij-35 Store up to 6 weeks at room temperature Pepstatinyl-agarose elution buffer 20 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 0.4 M NaCl 0.5% Brij 35 Store up to 6 weeks at 4°C Use prechilled at 4°C
Expression, Purification, and Characterization of Aspartic Endopeptidases
Plasmepsin substrate stock solution, chromogenic, 6 to 8 mM Synthesize p-nitrophenylalanine-containing peptide substrates for plasmepsin (e.g., Lys-Pro-Ile-Glu-Phe-Nph-Arg-Leu) using an Applied Biosystems automatic peptide synthesizer (ABI model 430A or 432A). Weigh out 10 mg of the chromogenic octapeptide substrate into a microcentrifuge tube. Dissolve the peptide in 200 µl DMSO, vortex well, and then add sufficient 10% (v/v) formic acid to bring the volume to 1 ml. Centrifuge substrate solution on a 0.45 µm Costar cellulose acetate centrifuge tube filter (Corning) for 5 min at 20,000 × g to remove particulates. Quantify by amino acid analysis on a Beckman System 6300 High Performance Amino Acid Analyzer (also see UNIT 3.2). Store peptides up to 1 month at 4°C (Dunn et al., 1994).
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Resuspension buffer A 10 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 20 mM MgCl2 5 mM CaCl Filter-sterilize Store up to 6 weeks at 4°C Resuspension buffer A2 50 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) 150 mM NaCl 1 mM MgCl2 Store up to 6 weeks at 4°C Resuspension buffer B 10 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 1 mM EDTA 2 mM 2-mercaptoethanol (2-ME) 100 mM NaCl Filter-sterilize Store up to 6 weeks at room temperature Resuspension buffer B2 50 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) 150 mM NaCl 1 mM MgCl2 1% (v/v) Triton X-100 Store up to 6 weeks at 4°C Resuspension buffer C 50 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 5 mM EDTA 5 mM 2-mercaptoethanol (2-ME) 0.5% Triton X-100 Filter-sterilize Store up to 6 weeks at room temperature Resuspension buffer D 50 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 5 mM EDTA 5 mM 2-mercaptoethanol (2-ME) Filter-sterilize Store up to 6 weeks at room temperature Solubilization buffer A 8 M urea, deionized and filtered (see recipe) 50 mM 3-(cyclohexylamino)-1-propanesulfonic acid (CAPS), pH 10.5 5 mM EDTA 200 mM 2-mercaptoethanol Prepare fresh The urea must be deionized and filtered before the other reagents are added (see recipe for urea, below).
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Solubilization buffer A2 8 M urea, deionized and filtered (see recipe) 50 mM 3-(cyclohexylamino)-1-propanesulfonic acid (CAPS), pH 10.7 50 mM 2-mercaptoethanol Prepare fresh The urea must be deionized and filtered before the other reagents are added (see recipe for urea, below). The solution must be prepared fresh prior to each use, because urea can break down.
Urea, 8 M, deionized and filtered Dissolve urea in distilled, deionized water, pH 6.0 to 8.0, in a volume equal to 40% of the final volume of the solubilization buffer A or A2 (see recipes) to be prepared. For example, if 250 ml of 8 M urea is required, add the urea to 100 ml of water. Use gentle stirring and moderate heat, between 25° and 30°C to assist in dissolving the urea at this high molar concentration. After urea is solubilized, add a mixed-bed ion-exchange resin (e.g., Dowex or Amberlite) and stir at 25°C for 30 min. The mixed-bed resin is used to remove any breakdown products of urea, such as cyanate, that can adversely react with the protein of interest. Use a Büchner funnel with filter suction to remove the resin, then add the remaining reagents necessary to prepare solubilization buffer A or A2 (see recipes), and add water to achieve the desired final volume. The solution must be prepared fresh prior to each use, because urea can break down.
COMMENTARY Background Information
Expression, Purification, and Characterization of Aspartic Endopeptidases
Recombinant techniques have become an important tool in the quest to obtain large quantities of enzyme, especially in cases where native sources have been deemed insufficient. The pET vector expression system represents one method to exploit the translational machinery of an appropriate host bacterium to synthesize a target protein. The stringently controlled T7 bacteriophage promoter prevents undesired leaky expression in the absence of IPTG, especially if the host harbors the plasmid for T7 lysozyme (pLysS or pLysE cell lines). The expression of recombinant aspartic peptidases is complete in a relatively short time (3 hr) at 37°C as compared to the expression of soluble proteins, which often require lower temperatures and longer induction periods. The refolding and column chromatography techniques utilized for the plasmepsins have proven efficient since their inception, even with the newly identified enzymes from the P. falciparum genome project (for example, plasmepsin 4). Certain peptidases, such as cathepsin D, are more difficult to refold and purify by these methods, and require different approaches in order to obtain a homogenous enzyme. Pepstatinyl-agarose has been utilized to purify cathepsin D for a number of years; however the zymogenic form coelutes with the mature enzym e. An additional ion-exchange
chromatography is performed to isolate the mature enzyme from the zymogen and improper folding intermediates. This step takes advantage of the difference in pI between the zymogen and the mature enzyme and is successful in producing a homogenous preparation of mature enzyme. Finally, all aspartic peptidase share a common trait in that they have acidic pH optima. Although these values can vary slightly, malarial aspartic endopeptidases and cathepsin D typically have pH optima at 4.5 and 3.7, respectively. Although an acidic pH in this range is a good starting point, any newly isolated or refolded enzyme should be tested in numerous pH buffers to elucidate its preferred enzymatic conditions.
Critical Parameters and Troubleshooting Of all the processes involved in the protocols described in this unit, two are more problematic than the others: choice of expression vector and host expression cell line, and determination of inclusion-body refolding conditions. There are several concerns that one must address when discussing protein expression. The selection of the expression vector is of utmost importance, especially if one wishes to express a soluble and/or fusion protein. In the case of the aspartic endopeptidases described in this unit, the inclusion body form of the protein is preferred, so a
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sion of the target gene, perhaps preventing the possible toxicity of the peptidases to the host cell. Lastly, the properties of the expression cell line should be examined when choosing an expression system. There are numerous cell lines commercially available, including but not limited to the strains BL21(DE3)pLysS (Novagen), BL21-STAR-(DE3)pLysS (Invitrogen), and BL21-CodonPlus-(DE3)-RIL (Stratagene). In all of these cells, the protein of
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vector/host combination that produces large quantities of protein is desired. pET3a has proven to be a viable plasmid, but other vectors, such as pET15b, can be used if a histidine tag is desired (Gulnik et al., 2002). Also, the toxicity of the target protein to the bacterial host is a major concern, but this pitfall has not been observed with the plasmepsins or cathepsin D. During the logarithmic growth phase of the bacteria, the pLysS strain reduces leaky expres-
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207 118 81 52.5 rPM2 36.2 29.9
Figure 21.14.2 Elution profile and SDS-PAGE analysis of plasmepsin 2 from ion-exchange chromatography. (A) Plasmepsin 2 post-dialysate was loaded onto a HiTrap Q Sepharose HP anion-exchange column and purified using a linear NaCl gradient from 0 to 1 M (straight diagonal line) in 20 mM Tris⋅Cl, pH 8.0, at 4°C over a period of 60 min. “% B” refers to the percentage of the IEX elution buffer (containing 1 M NaCl) in the gradient; the salt-free IEX starting buffer would be “A.” Flow rate was 2 ml/min and 2-ml fractions were collected. Recombinant plasmepsin 2 zymogen began to elute at approximately 220 mM NaCl as shown by the solid curve with the peak. Chromogenic assays performed at pH 4.5 with the post-ion exchange fractions revealed the active plasmepsin to be in fractions 17-19 (dashed curve with peak). Additionally, gel-filtration chromatography verified that fractions 16-19 contained properly folded plasmepsin, whereas the remaining fractions contained unfolded or misfolded enzyme (Fig. 21.14.4). (B) Plasmepsin 2 was separated on a 12% Tris-glycine acrylamide gel. Recombinant plasmepsin 2 in zymogen form migrates at approximately 42,000 Da. The gel reveals the majority of plasmepsin 2 is found in fractions 17-19, which corresponds to the fractions that contained the most activity as shown in panel A.
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fewer opportunities to form improper folding intermediates. Another method is to start with a high-pH buffer and slowly dialyze it to a lower pH. For cathepsin D, the solubilization and refolding of inclusion bodies is the most sensitive step of this protocol and all parameters must be strictly controlled. Additionally, refolding and activation times are typically determined by evaluating the emergence of proteolytic activity. Since refolding is done at very dilute concentrations of protein, concentration of the sample is required. The temperature of refolding, ionic strength of the buffers, amount of reducing and/or oxidizing agents, and the length of each buffer change should also be investigated (see UNITS 6.4 & 6.5 for suggested conditions), as these parameters affect all of the previously mentioned pitfalls. Unfortunately, there is no straightforward method by which to screen these conditions; one must simply test them systematically by trial and error. Lastly, the quality of the pepstatinyl-agarose used in the affinity purification of cathepsin D is extremely important. Commercially available pepstatinyl-agarose manufactured utilizing a cyanogen bromide–activated resin appears less stable than NHS-DCC activated resin synthesized from the method of Huang et. al. (1979).
interest was overexpressed quite well. However, it was noted that the total amount of expression was superior with the BL21-STAR(DE3)pLysS cell line (which provided a 2- to 3-fold greater inclusion body yield). These cells encode a truncated RNaseE enzyme (rne131) that lacks the ability to degrade mRNA, resulting in an increase in mRNA stability (Grunberg-Manago, 1999; Lopez et al., 1999). This property of the BL21-STAR-(DE3)pLysS, in conjunction with the tight control as a result of the pLysS plasmid that produces T7 lysozyme, is a logical explanation for the higher amounts of inclusion bodies produced by these cells. The conditions under which a protein undergoes proper refolding are difficult to discern for many constructs. However, the malarial endopeptidases described in this unit do not require extreme conditions to yield milligram amounts of active enzyme. These proteins easily refold by simply lowering the concentration of denaturants in the buffered system. If proteins precipitate during this process, then alternate methods must be investigated to prevent this occurrence. One possible remedy includes decreasing the urea concentration with each buffer change, thereby allowing the protein to adopt its proper conformation slowly and with
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zymogenic rHuCatD mature rHuCatD
30 20 14
Expression, Purification, and Characterization of Aspartic Endopeptidases
Figure 21.14.3 SDS-PAGE analysis illustrating purification of “short” pseudo rHuCatD from the refolding mixture. The far left lane corresponds to molecular weight markers. Lane 1 represents full length rHuCatD from the refolding mixture. Lane 2 shows the pepstatinyl-agarose copurification of the zymogen with the mature enzyme (the faint proenzyme band in this gel does not reproduce well). Lane 3 illustrates the ion-exchange purified “short” pseudo rHuCatD migrating at about 42 kDa. The single band indicates purity to near homogeneity as judged by silver staining.
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volume of 16 liters is typically employed for each refolding preparation. For both enzymes, these amounts typically yield 0.2 to 2 mg of pure enzyme. For any new construct, be it a novel enzyme or a mutant of an existing endopeptidase, one should analyze the gamut of refolding parameters discussed above (see Critical Parameters). Lastly, the purification of plasmepsins and cathepsin D using anion-exchange or pepstatinyl-agarose affinity chromatography, respectively, is typically sufficient to give homogenous enzyme (Figs. 21.14.2 and 21.14.3). This is primarily due to the efficiency of the inclusion body purification protocol, which removes other bacterial proteins. If a construct is not homogenous after these steps, a gel-filtration column (Superdex 75 or Sephacryl 100 resin) can be used to separate any contaminating proteins or misfolded forms (Fig. 21.14.4).
Anticipated Results SDS-PAGE gels for the different expression and purification fractions of plasmepsin 2 and “short” recombinant human cathepsin D are shown in Figures 21.14.1A and 21.14.1B, respectively. After 3 hr post-IPTG induction, the expression levels of the target peptidase are maximal. As observed with plasmepsin 2 and cathepsin D, the purification from a 1-liter expression culture typically yields about 500 to 600 mg of wet inclusion body pellet. The total inclusion body yield will vary depending upon which peptidase is being expressed; therefore, choose the expression host that yields the largest wet weight of inclusion bodies per liter of medium for each enzyme. The solubilization and refolding conditions remain constant for each of the plasmepsins, but this is likely to be due to their high primary sequence and structural similarities. For the plasmepsins, solubilized protein is denatured at 1 mg/ml in 250 ml of buffer for each refolding assay. However, refolding is an inefficient process for cathepsin D, so at a concentration of 20 µg/ml, a refolding
Time Considerations The protocols described in this section for the plasmepsins can be completed in fewer than
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Figure 21.14.4 Elution profile of Superdex 75 gel filtration chromatography assay using post–anion exchange fractions of plasmepsin 2. Plasmepsin 2 post–anion exchange fractions (post-IEX) from Figure 21.14.2 were loaded onto a Superdex 75 gel filtration column (GF75) and separated at 4°C in 20 mM Tris⋅Cl, pH 8.0. Flow rate was 2 ml/min and 1-ml fractions were collected. Bio-Rad Gel Filtration Chromatography Standards are shown and are labeled according to the molecular weight of each peak. Fractions 16 to 19 obtained from post-IEX, termed peak 1 from IEX, were pooled and separated on the GF75 column. The elution profile of this peak corresponds to the molecular weight standard of 44,000 Da, suggesting the plasmepsin 2 is in its properly folded tertiary form. Fractions 20 to 24 of IEX, termed peak 2 from IEX, were pooled and concentrated 10-fold prior to application onto the GF75 column. The elution profile of these samples suggests these post-IEX fractions contain unfolded and/or misfolded protein, as well as properly folded enzyme.
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5 days. It should take ∼1 day to express and harvest the cells, 1 day for inclusion body purification/solubilization, 2 days for the five buffer changes of dialysis, and 1 day for protein purification and enzymatic analysis. For the ambitious, this process can be shortened if the expression and inclusion body purification/solubilization are performed on the same day. For native human pseudocathepsin D material, 3 days of refolding plus 2 days of activation are required. The pepstatinyl-agarose column is an overnight procedure, as well as the dialysis step. The ion-exchange step can be done in about 4 to 5 hr depending on the sample volume. Taken together, the entire cathepsin D protocol can be completed in 8 days.
Gulnik, S.V., Afonina, E.I., Gustchina, E., Yu, B., Silva, A.M., Kim, Y., and Erickson, J.W. 2002. Utility of (His)6 tag for purification and refolding of proplasmepsin-2 and mutants with altered activation properties. Protein Expr. Purif. 24:412-419.
Literature Cited
Taylor, G., Hoare, M., Gray, D., and Marston, F. 1986. Biotechnology 4:553-557.
Ausubel, F.M., Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G., Smith, J.A, and Struhl, K. (eds.). 2003. Current Protocols in Molecular Biology. John Wiley & Sons, New York. Beyer, B.M. and Dunn, B.M. 1996. Self-activation of recombinant human lysosomal procathepsin D at a newly engineered cleavage junction, “short” pseudocathepsin D. J. Biol. Chem. 271:15590-15596. Bieth, J.G. 1995. Theoretical and practical aspects of proteinase inhibition kinetics. Methods Enzymol. 248:59-84. Conner, G.E. and Udey, J.A. 1990. Expression and refolding of recombinant human fibroblast procathepsin D. DNA Cell Biol. 9:1-9. Dunn, B.M., Scarborough, P.E., Davenport, R., and Swietnicki W. 1994. Analysis of proteinase specificity by studies of peptide substrates: The use of UV and fluorescence spectroscopy to quantitate rates of enzymatic cleavage. Methods Mol. Biol. 36:225-243. Grunberg-Manago, M. 1999. Messenger RNA stability and its role in control of gene expression in bacteria and phages. Annu. Rev. Genet. 33:193-227.
Huang, J.S., Huang, S.S., and Tang, J. 1979. Cathepsin D isozymes from porcine spleens: Large scale purification and polypeptide chain arrangements. J. Biol. Chem. 254:11405-11417. Lopez, P.J., Marchand, I., Joyce, S.A., and Dreyfus, M. 1999. The C-terminal half of RNase E, which organizes the Escherichia coli degradosome, participates in mRNA degradation but not rRNA processing in vivo. Mol. Microbiol. 33:188-199. Morrison, J.F. 1969. Kinetics of the reversible inhibition of enzyme-catalysed reactions by tightbinding inhibitors. Biochim. Biophys. Acta. 185:269-286.
Westling, J., Yowell, C.A., Majer, P., Erickson, J.W., Dame, J.B., and Dunn, B.M. 1997. Plasmodium falciparum, P. vivax, and P. malariae: A comparison of the active site properties of plasmepsins cloned and expressed from three different species of the malaria parasite. Exp. Parasitol. 87:185-193. Westling, J., Cipullo, P., Hung, S.H., Saft, H., Dame, J.B., and Dunn, B.M.. 1999. Active site specificity of plasmepsin II. Protein Sci. 8:2001-2009. Yasuda, Y., Kageyama, T., Akamine, A., Shibata, M., Kominami, E., Uchiyama, Y., and Yamamoto, K. 1999. Characterization of new fluorogenic substrates for the rapid and sensitive assay of cathepsin E and cathepsin D. J. Biochem. (Tokyo) 125:1137-1143.
Contributed by Bret B. Beyer, Nathan E. Goldfarb, and Ben M. Dunn University of Florida Gainesville, Florida
Expression, Purification, and Characterization of Aspartic Endopeptidases
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Zymography of Metalloproteinases
UNIT 21.15
Zymography is an electrophoretic technique enabling visualization of the number and approximate size of proteinases in a sample on the basis of their hydrolysis of a gel-embedded protein substrate such as proteoglycan (Barrett, 1966), gelatin (Itoh et al., 1998), and casein (Leber and Bakuil, 1997; Zeng et al., 2002). The technique is particularly useful for analyzing the proteinase composition of complex biological samples, because visualization depends directly on proteolytic activity (Feitosa et al., 1998; Jain et al., 2001). Zymography is also widely used to study various aspects of matrix metalloproteinase (MMP) function (Kjeldsen et al., 1993; Itoh et al., 1998). Gelatin zymography, a widely used technique in the study of MMPs, is described in this unit. In principle, it is also applicable for use with many proteinases if the enzyme retains activity after electrophoresis and renaturation. This unit presents a representative zymography assay using matrix metalloproteinase (MMP) and a gelatin or casein substrate (see Basic Protocol), as well as protocols for other substrates (see Alternate Protocol 1), real-time zymography (i.e., using a fluorescent protein substrate; see Alternate Protocol 2), and reverse zymography for the study of proteinase inhibitors (see Alternate Protocol 3). These procedures employ the 2-amino-2-methyl-1,3-propanediol (ammediol) buffer system introduced by Bury (1981), but many other SDS-PAGE systems (e.g., Laemmli; UNIT 10.1) have been successfully used by others (Hummel et al., 1996; Hawkes et al., 2001). NOTE: For an overview of matrix metalloproteinases and tissue inhibitors of metalloproteinases (TIMPs), see UNIT 21.4. ZYMOGRAPHY OF MATRIX METALLOPROTEINASES USING A GELATIN OR CASEIN SUBSTRATE
BASIC PROTOCOL
To provide a uniformly dispersed substrate, the peptide (e.g., gelatin) is included in the standard polyacrylamide gel solution and the gel is then poured and polymerized as usual. Since the substrate is entrapped in the pores of the polyacrylamide gel, it does not migrate out of the gel during electrophoresis. Proteinase samples are denatured with sodium dodecyl sulfate (SDS), but not boiled or reduced, and then electrophoresed. The separated proteinases are renatured within the gel by replacing the SDS with a nonionic detergent such as Triton X-100. The gel is then incubated in a buffer suitable for enzymatic activity, allowing the renatured proteinases to digest the gel-bound substrate protein in a zone around their electrophoresed position. These zones of digestion are visualized using Coomassie brilliant blue R250, with the areas of digestion appearing clear against a blue-stained background of undigested protein. Materials Acrylamide/bisacrylamide solution (see recipe) Separating gel buffer (see recipe) 10× casein or gelatin substrate solution (see recipes) Sucrose solution (see recipe) 10% (w/v) ammonium persulfate (see recipe) TEMED n-Butanol, H2O saturated Stacking gel buffer (see recipe) MMP sample 1 mM APMA (see recipe) Peptidases Contributed by Linda Troeberg and Hideaki Nagase Current Protocols in Protein Science (2003) 21.15.1-21.15.12 Copyright © 2003 by John Wiley & Sons, Inc.
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2× sample loading buffer (see recipe) Proteinase standards (e.g., recombinant MMP-1, -2, or -3) 1× anode and cathode reservoir buffers (see recipes) Sample known to degrade gelatin or casein (e.g., recombinant culture supernatents) Molecular mass standards (e.g., BioRad low-range SDS-PAGE standards) Enzyme renaturing buffer (see recipe) Developing buffer, 37°C (see recipe) Staining solution (see recipe) Destaining solution (see recipe) Electrophoresis apparatus (e.g., ∼9-cm × 6-cm × 0.75-mm minigel system) and comb Gel-loading pipet tips or Hamilton syringe Constant voltage power supply Sealed plastic container Additional equipment and reagents for preparing and running acrylamide gels (UNIT 10.1) 1. Choose an appropriate percentage separating gel and prepare according to Table 21.15.1. Mix the solution well and pour between electrophoresis plates to a height ∼1.5 to 2 cm from the top (UNIT 10.1). Overlay with water-saturated n-butanol and allow the gel to polymerize ∼30 min. The choice of acrylamide percentage depends on the molecular weight of the proteinase to be visualized. Proteins with higher molecular masses resolve better on lower percentage gels, while those with lower molecular masses resolve better on higher percentage gels. For analysis of MMPs, 10% gels are usually appropriate, as they resolve proteins between 20 and 100 kDa. Alternatively, Novex casein and gelatin zymograms are commercially available from Invitrogen (Kleiner and Stetler-Stevenson, 1994). Also, different substrates can be used (see Alternate Protocol 1).
2. Prepare a stacking gel by mixing the following: 142 µl acrylamide/bisacrylamide 285 µl stacking gel buffer 285 µl sucrose solution 428 µl H2O 14 µl 10% (w/v) ammonium persulfate 3 µl TEMED. Decant the n-butanol from the separating gel, gently rinse with water (e.g., from a squirt bottle) to remove any remaining alcohol, and immediately pour the stacking gel solution on top of the separating gel. Insert the well comb, taking care not to trap bubbles under the comb. Allow the stacking gel to polymerize 15 to 20 min at room temperature. The stacking gel polymerizes much faster than the separating gel (∼5 to 10 min), so work as quickly as possible after adding TEMED. A stacking gel with the same percentage acrylamide is used irrespective of the acrylamide concentration in the separating gel. There should be at least 1-cm stacking gel below the comb to ensure proper stacking of proteins, which is a prerequisite for good resolution in the separating gel. Refer to UNIT 10.1 for more information. Zymography of Metalloproteinases
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Table 21.15.1
Recipes for Zymogram Separating Gelsa,b
Stock solution Acrylamide/bisacrylamide solution Separating gel buffer 10× casein or gelatin substrate solution Sucrose solution Water 10% (w/v) ammonium persulfate TEMED
Final acrylamide concentration in gel 6%
7.5%
10%
12%
0.8 ml 1.0 ml 0.4 ml 0.86 ml 0.94 ml 14 µl 1.5 µl
1.0 ml 1.0 ml 0.4 ml 0.86 ml 0.74 ml 14 µl 1.5 µl
1.32 ml 1.0 ml 0.4 ml 0.86 ml 0.42 ml 14 µl 1.5 µl
1.61 ml 1.0 ml 0.4 ml 0.86 ml 0.13 ml 14 µl 1.5 µl
aAbbreviations: TEMED N,N,N′,N′-tetramethylethylenediamine. bRecipes given are sufficient for one ∼9-cm × 6-cm × 0.75–mm minigel.
3. Optional: Activate proenzyme (zymogen) samples by incubating 1 to 4 hr at 37°C in the presence of 1 mM APMA diluted from a 20 mM stock. Mix 5 to 20 µl activated sample with an equal volume of 2× sample loading buffer. Most MMP proenzymes can be activated using these conditions. See Nagase (1997) for a discussion of the current hypothesis of APMA activation. Note that this loading buffer contains no reducing agents or EDTA, which would inactivate matrix metalloproteinases (MMPs). Do not boil the sample, as this will irreversibly inactivate MMPs.
4. Optional: If the identity of the proteinase is known, prepare a standard curve of proteinase standards (i.e., at least five known concentrations) and coanalyze (see Figure 21.15.1). 5. Assemble the gel in the electrophoresis apparatus according to the manufacturer’s instructions and UNIT 10.1. Pour 1× anode and cathode reservoir buffers into the lower and upper chambers, respectively. 6. Load samples, proteinase standards (if proteinase is known), a sample known to degrade gelatin or casein (e.g., recombinant culture supernatants; positive control), unconditioned medium (negative control), and molecular mass standards into appropriate wells using gel-loading pipet tips or a Hamilton syringe. Be especially careful to avoid spillage between lanes for samples that contain a high concentration of proteinase activity or leave empty lanes between such samples. Thoroughly wash the Hamilton syringe between samples.
7. Electrophorese using a constant voltage power supply at 150 V (nonlimiting current) until the bromophenol blue reaches the bottom of the gel (UNIT 10.1). A 10% acrylamide gel should take ∼45 min to complete.
8. After electrophoresis, disassemble the gel apparatus and place the gels in a sealed plastic or glass container filled with ∼20 ml enzyme renaturing buffer. Wash 15 min at room temperature with gentle agitation on an orbital shaker. Repeat this procedure four times using fresh buffer each time, for a total washing time of 1 hr. This process exchanges SDS within the gel with the nonionic detergent Triton X-100, facilitating renaturation of enzyme samples. Incubate one gel per container so that all areas of each gel are thoroughly washed. It is not necessary to remove the stacking gel. Peptidases
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9. Transfer gel to ∼30 ml developing buffer and incubate 2 hr to overnight at 37°C. An appropriate incubation time should be empirically determined depending on the required level of sensitivity. As little as 5 pg MMP-2 can be detected using an overnight incubation. If incubating overnight, ensure that the container is airtight to prevent evaporation. Real-time zymography can also be performed (see Alternate Protocol 2).
10. Decant the developing buffer and add staining solution. Stain 20 to 30 min at room temperature and then destain thoroughly (>2 hr) using multiple changes of destaining solution until clear bands of MMP activity are visible against the dark blue background. The gel can be stored in destaining solution up to a few months at room temperature. For long-term storage, gels can be dried using a commercial gel drier. Refer to UNIT 10.5 for more information on Coomassie brilliant blue R250 staining and drying gels.
A B 1 2 3 4 5 6 7 8 9 10
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MMP-2 (pg)
Zymography of Metalloproteinases
Figure 21.15.1 Recombinant proMMP-2 was activated by 1 hr incubation with 1 mM APMA at 37°C and its active concentration confirmed by titration against TIMP-2 (Troeberg et al., 2002). Various concentrations of recombinant MMP-2 were analyzed on a 7.5% gelatin zymogram (upper panel) incubated at 37°C for either 4 hr or 16 hr, A and B, respectively. Loads were as follows: lane 1, 2 pg; lane 2, 5 pg; lane 3, 10 pg; lane 4, 25 pg; lane 5, 50 pg; lane 6, 100 pg; lane 7, 200 pg; lane 8, 300 pg; lane 9, 400 pg; lane 10, 500 pg; lane 11, 600 pg; and lane 12, 1 ng. Empty lanes were left between the highest concentrations of MMP-2 to ensure accurate quantitation. Pixel volume was quantified by arbitrary units (given by the software) and plotted against the amount of MMP-2 (lower panel), with the 4 hr and 16 hr incubations represented with closed and open circles, respectively. The dashed lines show that band intensity is linearly related to MMP-2 concentration, but the linear range of enzyme concentration varies with the incubation time (i.e., a longer incubation results in more substrate being cleaved and subsequently the signal becomes saturated at lower concentrations of enzyme relative to a shorter incubation). In this example, the relationship is linear up to ∼1.3 × 106 intensity units. Empty lanes have left between the highest concentrations of MMP-2 to ensure accurate quantitation.
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11. If proteinase standards were included, construct a calibration curve using the known concentrations of the enzyme under study and the area of digestion produced. Compare this with the area of digestion of a test sample to quantify results. The linear range for such analysis varies with the enzyme analyzed (and its ability to digest the protein substrate used) and with the incubation time. As little as 1 to 10 pg MMP-2 (Fig. 21.15.1; Kleiner and Stetler-Stevenson, 1994) or 32 pg MMP-9 (Leber and Balkwill, 1997) can be detected on gelatin zymograms. The area of digestion can be quantified using a densitometer and associated software. Specific analysis techniques will vary with the software available. The gel in Figure 21.15.1 was scanned using a Bio Rad GS-710 Calibrated Imaging Densitometer and analyzed using Phoretix 1D software (Non-Linear Dynamics). The image should be inverted to give black bands on a white background. Place equally sized rectangular areas around each band, choosing an area just large enough to encompass the largest band. Background can be subtracted either locally or globally. Suitable background subtraction techniques are best determined empirically depending on the gel (intensity of bands, darkness of background) and the software available. Pixel density within the areas is then quantified and plotted against the amount of MMP-2. Band intensity is linearly related to MMP-2 concentration over a range that varies with the incubation time. In Figure 21.15.1, the relationship is linear up to ∼1.3 × 106 units. Empty lanes have been left between the highest concentrations of MMP-2 to ensure accurate quantitation.
ZYMOGRAPHY OF MATRIX METALLOPROTEINASES USING SUBSTRATE PROTEINS OTHER THAN GELATIN
ALTERNATE PROTOCOL 1
Gelatin is widely used as a substrate for zymography because it is readily available, economical, and serves as a substrate for several MMPs (see Basic Protocol). However, any protein that is digested by the proteinase of interest and that remains soluble in the buffer system used for electrophoresis can be used as an alternative to gelatin. For example, proteins such as collagen (0.2 to 1 mg/ml; Gogly et al., 1998), fibronectin (2 mg/ml; Feitosa et al., 1998), and fibrinogen (2 mg/ml; Feitosa et al., 1998) have been used (all available from Sigma). When working with a novel enzyme, it may be beneficial to try several possible substrates to determine which yields the best results. Although able to detect low picogram quantities of enzymes (Gogly et al., 1998), collagen zymograms are technically difficult and not widely used. Collagen has a tendency to precipitate in buffers commonly used for electrophoresis and considerable care must be taken to prevent denaturation of the collagen to gelatin in order to prevent detection of gelatinase in addition to the collagenase activity. For example, the temperature of the gel must not exceed 25°C at any time, including during electrophoresis. REAL-TIME ZYMOGRAPHY With Coomassie staining of zymograms (see Basic Protocol), it is necessary to estimate a suitable development time before irreversibly fixing and staining proteins. To eliminate this inherent uncertainty, the technique of real-time zymography has been developed, using a fluorescent protein as the substrate. Digestion is visualized directly in the developing buffer using a UV transilluminator with areas of digestion visible as dark bands against a fluorescent background. Fluorescent substrates include 0.5 mg/ml FITCgelatin (heat-denatured FITC-collagen) and 0.5 mg/ml FITC-casein (Hattori et al., 2002), which have been used for analyzing MMP-2, -3, and -9. FITC-gelatin zymograms allow for detection of as little as 16 pg MMP-2 and 3 pg MMP-9 after incubation for 24 hr. In fact, FITC-casein is five times more sensitive for MMP-3 than conventional casein zymography.
ALTERNATE PROTOCOL 2
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In addition to the substrates mentioned above, a short peptide sequence known to be hydrolyzable by the proteinase of interest can be tagged with the fluorescent 4-methylcoumaryl-7-amide (MCA) moiety. For example, trypsin and chymotrypsin activity have been detected using Boc-Gln-Ala-Arg-MCA and Suc-Ala-Ala-Pro-Phe-MCA respectively (Yasothornsrikul and Hook, 2000). In this case, activity is visible as zones of fluorescence against a dark background. ALTERNATE PROTOCOL 3
REVERSE ZYMOGRAPHY FOR STUDY OF PROTEINASE INHIBITORS A variation of zymography (see Basic Protocol) can be used to analyze inhibitors of matrix metalloproteinases such as tissue inhibitors of metalloproteinases (TIMPs). In this version, reverse zymography, a proteinase (commonly MMP-2) is embedded in the gel in addition to the protein substrate (e.g., gelatin). The TIMPs are then electrophoresed, and during the development step, the renatured proteinase digests the background substrate throughout the gel, except at positions in the gel where inhibitors are present. Upon staining, the digested background appears pale blue, while areas of inhibition are visible as reddish-purple zones. It is always necessary to run a companion SDS-PAGE gel (without gelatin or proteinase) to check that these zones represent true areas of inhibition, and not just Coomassie-stained proteins; inhibitors will be visible on the reverse zymogram, but absent on the companion SDS-PAGE gel. As little as 1 pg TIMP-2 and 40 pg TIMP-1 can be detected using MMP-2 and gelatin (Oliver et al., 1997). This technique has been used to investigate the presence of TIMPs in biological samples (Noha et al., 2000), the role of TIMP-2 in MMP-2 activation (Itoh et al., 1998), and various aspects of TIMP function (Yeow et al., 2002). FITC-labeled proteins (see Alternate Protocol 2) can also be used for reverse zymography. In this case, visualization is directly dependent on inhibition of proteolytic activity and thus permits easier discrimination between inhibitors and contaminating protein bands than gelatin-based techniques (Hattori et al., 2002). REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Acrylamide/bisacrylamide solution 30% acrylamide (Severn Biotech) 0.8% bisacrylamide (Severn Biotech) Store up to 6 months at 4°C. This reagent is commercially available as 37.5:1 acrylamide/bisacrylamide.
Ammonium persulfate, 10% (w/v) Dissolve 0.5 g ammonium persulfate in a final volume of 5 ml water. Store up to 1 month at −20°C. Anode reservoir buffer, 4×, 1× Dissolve 26.28 g of 2-amino-2-methyl-1,3-propanediol⋅HCl (ammediol; 84 mM) in ∼900 ml water and adjust to pH 8.23 with HCl. Adjust volume to 1000 ml with water. Store up to 1 month at 4°C. Dilute to 1× with water as required. The anode reservoir is the lower chamber and is indicated by red markings.
Zymography of Metalloproteinases
APMA, 20 mM Dissolve 7 mg (20 mM) 4-aminophenylmercuric acetate (APMA; ICN Biochemicals) in 1 ml of 80 mM NaOH with vortexing. Store up to 1 month protected from light at −20°C.
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Casein substrate solution, 10× Dissolve 800 mg casein (BDH) in 100 ml of 0.1 N NaOH. Heat the solution to 37°C with occasional vortexing until no undissolved material is visible. Store up to several months at −20°C. Cathode reservoir buffer, 4×, 1× Dissolve 17 g ammediol (41 mM), 12 g glycine (40 mM ), and 4 g SDS (0.4%) in ∼900 ml water. Adjust volume to 1000 ml with water. Check that the pH is ∼9.39 and adjust with NaOH or HCl as necessary. Store up to 1 month at 4°C. Dilute to 1× with water as required. The cathode reservoir is the upper chamber and is usually indicated by black markings.
Destaining solution Combine 1.5 liters methanol, 3.5 liters water, and 50 ml formic acid. Store up to several months at room temperature. Developing buffer Dissolve 6.055 g Tris base (50 mM), 11.69 g NaCl (200 mM), 0.7 mg ZnCl2, 0.74 g CaCl2⋅2 H2O (5 mM), and 0.2 g NaN3 (0.02%) in ∼900 ml water. Adjust to pH 7.5 with HCl and adjust volume to 1000 ml with water. Store up to 1 month at 4°C. Enzyme renaturing buffer Add 25 ml Triton X-100 to developing buffer (see recipe). Store up to 1 month at 4°C. Final concentrations are 200 mM NaCl, 5 mM CaCl2, 5 ìM ZnCl2, 2.5% (v/v) Triton X-100, 0.02% (w/v) NaN3, and 50 mM Tris⋅Cl, pH 7.5.
Gelatin substrate solution, 10× Add 800 mg porcine-skin gelatin (Sigma) to 100 ml water in a glass container. To ensure a uniformly stained background, dissolve the protein substrate fully before incorporation into the gel. Heat in a microwave oven until the solution just boils. Do not allow to boil over. Swirl thoroughly to ensure that all of the protein is in solution. Store up to one week at 4°C or several months at −20°C. Sample loading buffer, 2× Dissolve 0.2 g SDS (2%) and 0.01 g bromophenol blue (0.1%) in 5 ml stacking gel buffer (see recipe) and add 5 ml glycerol (40%). Store up to 1 month at −20°C. Separating gel buffer Dissolve 4.63 g ammediol⋅HCl (110 mM) and 0.02 g sodium azide (NaN3; 0.02%) in ∼90 ml water. Adjust pH to 8.96 with HCl and adjust volume to 100 ml with water. Store up to 1 month at 4°C. Stacking gel buffer Dissolve 3.51 g ammediol⋅HCl (84 mM ) and 0.02 g NaN3 (0.02%) in ∼90 ml water. Adjust pH to 8.37 with HCl and adjust volume to 100 ml with water. Store up to 1 month at 4°C. Staining solution Dissolve 2.5 g Coomassie brilliant blue R-250 (0.125%) in a mixture of 1.25 liters methanol, 0.5 liters acetic acid, and 0.75 liters water. Store up to several months at room temperature.
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Sucrose solution Dissolve 50 g sucrose and 0.02 g NaN3 in ∼50 ml water. Add 30 µl toluene and adjust volume to 100 ml with water. Store up to 1 month at 4°C. COMMENTARY Critical Parameters and Troubleshooting See UNIT 10.1 for a thorough discussion of important technical considerations as well as problems that can occur with preparing and running SDS polyacrylamide gels. The following is a brief list of problems that are specific to zymograms. 1. Background too pale. Inadequate incorporation of the substrate protein into the gel can cause problems. Remake the gelatin solution, taking care that the correct amount of protein has been added. Gelatin solutions should not be stored at 4°C for more than ∼1 week, to prevent contamination with bacteria, which may secrete proteolytic enzymes. 2. Background streaky or blotchy. To achieve a uniformly stained background, it is crucial that the substrate protein be completely dissolved prior to casting the gel. Hold the protein solution up to the light and swirl to check for any undissolved material. Gelatin solutions should be heated briefly in the microwave or incubated at 37°C with occasional vortexing to ensure that all protein is dissolved. 3. No zones of digestion are evident. As a positive control, incorporate appropriate quantities of enzymes that are known to digest the substrate—e.g., MMP-2 (100 pg for an overnight incubation) or MMP-9 (100 pg for an overnight incubation) for gelatin zymograms, and MMP-1 (500 pg for an overnight incubation) for casein zymograms. If these positive controls are visible, the test sample may be devoid of the enzymes that digest the particular protein substrate or the proteinase concentration may be lower than expected; either concentrate the solution by nondenaturing means or increase the incubation time. If these positive controls are not visible, check that the sample loading buffer does not contain any reagents that would inactivate the proteinases—e.g., for MMPs, this buffer should not contain EDTA. Also check that the samples have not been boiled, as this will irreversibly inactive the proteinases, and that the renaturing solution contains the correct amount of Triton X-100, which is required to renature the proteinases after electrophoresis. Gels must be washed at Zymography of Metalloproteinases
least four times for 15 min each to achieve adequate renaturation. 4. Smeared zones of digestion. Smearing is a sign that too much substrate protein has been digested. Decrease the incubation time or load less enzyme.
Anticipated Results Figure 21.15.2 shows zymograms of various recombinant MMPs using gelatin (panel A) or casein (panel B) substrate stained with Coomassie brilliant blue. The substrate background stains dark blue, while the MMPs show up as lightly stained zones of digestion, having degraded the protein substrate around the position to which they have electrophoresed. On gelatin zymograms, as little as 10 pg MMP-2 generates a visible zone of digestion (Fig. 21.15.2A, lane 3), while 500 pg MMP-1 and -3 are required to generate a clear zone of digestion. This indicates that gelatin is a better substrate for MMP-2 and -9 than MMP-1 and -3. Similar amounts of MMP-1 and -3 are required if casein is used as the substrate protein (Fig. 21.15.2B), but since MMP-2 does not hydrolyze casein effectively, casein zymograms are more useful for analyzing MMP-1 and -3, while excluding MMP-2 activity. The molecular mass of proteinases can be estimated from their position relative to molecular mass markers; however, samples are not boiled or reduced prior to electrophoresis, so their migration may not accurately reflect their molecular mass. Additionally, inclusion of a protein substrate in the gel retards the migration of proteins through the gel in a nonlinear manner (Hummel et al., 1996). Accurate assessment of molecular mass is thus not possible on a zymogram. Samples of known MMPs can be run alongside test samples for more accurate estimation of molecular mass, but alignment of a band of interest with a known MMP cannot be taken as proof of identity. Further characterization, such as immunoblotting (UNIT 10.10) or removal of the proteinase from the test sample by immunoprecipitation (UNIT 9.8), is required for unequivocal identification. MMP proenzymes are inactive in solution, but visible on zymograms. As shown in Figure 21.15.2A, zymography permits visualization of
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activation should not be confused with in vitro activation by proteinases or APMA, which is carried out prior to electrophoresis. For example, 72-kDa proMMP-2 (Fig. 21.15.2A, band a) is converted to 68-kDa active MMP-2 (lane 3) by treatment with APMA prior to electrophoresis. Furthermore, MMPs are often present in biological samples in inactive complexes with their endogenous inhibitors, TIMPs. These complexes dissociate upon treatment with SDS and, as MMP and TIMP electrophorese to different positions and remain separated during development of the zymogram, TIMPcomplexed MMPs appear as active enzymes on zymograms (see also Fig. 21.15.4; Itoh et al., 1998).
proenzyme forms of MMPs—e.g., proMMP-2 in lane 2 (band a). MMPs are maintained in their proenzyme form by coordination of the catalytic zinc atom in the MMP active site by a cysteine residue in the propeptide region. They are activated by proteolytic processing of the propeptide, which generates an active lowermolecular-mass form (Nagase, 1997). Zymography, however, permits visualization of these normally inactive proenzymes. Treatment of MMPs with SDS perturbs their tertiary structure, disrupting the interaction of the propeptide with the catalytic zinc atom. Upon renaturation of the enzymes after electrophoresis, the proenzymes are autocatalytically activated and the protein substrate digested in a region corresponding to the molecular mass of the zymogen. This characteristic of zymography can be useful and has permitted study of proMMP-2 activation by zymography (Itoh et al., 1998). Care must be taken not to infer in vivo activity from a zymogram band that actually corresponds with a proenzyme inactive in solution. Also, this zymogram-dependent
High-molecular-mass MMP complexes Gels of biological samples often contain bands of proteolytic activity at masses in excess of any known MMPs, which are often actually complexes of MMP-9. MMP-9 (92 kDa) forms a disulfide-bonded homodimer of ∼200 kDa
B
A
250 kDa 150 100 75 50 37 25
1
2
3
4
5
M 6
7
8
9 10
1
2
3
4
5
M
6
7
8
9 10
Figure 21.15.2 Various recombinant MMPs and conditioned culture media from different cell lines were analyzed on 7.5% acrylamide gelatin (A) or casein (B) zymograms, and developed overnight at 37°C. Lane 1 shows 500 ng of 43-kDa recombinant MMP-1. Lane 2 shows 100 pg of 72-kDa proMMP-2 (band a) and the 68-kDa active form of MMP-2 (band b) purified from the culture medium of human cervical fibroblasts. Lane 3 contains 10 pg of 68-kDa APMA-activated MMP-2. Lane 4 shows 250 ng of 18.5-kDa recombinant MMP-3 catalytic domain. Lanes 5 to 10 show culture media collected from MMP-expressing cells. In lane 5, 2 µl of HT1080 (human fibrosarcoma) culture supernatant treated with the phorbol ester 12-O-tetradecanoylphorbol 13-acetate (TPA) produce zones of digestion likely to correspond to the 92-kDa proMMP-9 (band c), 83-kDa active form of MMP-9 (band d), 72-kDa proMMP-2, and 68-kDa MMP-2. Lanes 6 to 8 show 2-µl loads of three fibroblast cell lines that produce mainly proMMP-2 and MMP-2 (lane 6, human embryonic lung fibroblasts; lane 7, human fibrosarcoma; lane 8, human uterine cervical fibroblasts). Lane 9 shows a 2-µl load of cultured human rheumatoid synovial fibroblasts, which produce 92-kDa proMMP-9 and 83-kDa MMP-9, in addition to proMMP-2/MMP-2, whereas only proMMP-2 is visible in the 2 µl culture medium from cultured human chondrocytes shown in lane 10. Protein molecular mass markers are shown in lane M. Fewer bands of digestion are evident on the casein zymogram (B), since MMP-2 and -9 digest this substrate poorly; however, MMP-1 and -3 are clearly visible on both casein and gelatin zymograms.
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none
EDTA
E-64
pepA
AEBSF 250 kDa 150 100 75 50 37 25
M rE M
rE
H
H
rE
M rE
M rE H
Figure 21.15.3 A sample containing 100 pg recombinant proMMP-2/MMP-2 (rE) and 2 µl conditioned medium from TPA-treated HT1080 (H) were run on a 7.5% gelatin zymogram as in Figure 21.15.2. The effects of various proteinase inhibitors on activity were investigated. Compared with zymograms incubated in the absence of inhibitors (none), addition of 20 µM E-64, 10 µM pepstatin A (pepA), or 1 mM AEBSF to the renaturing and developing buffers had no effect on enzyme activity. Addition of 20 mM EDTA, however, abolished all zones of gelatin digestion, confirming that the enzymes are metalloproteinases. Lane M contains protein molecular-mass markers.
250 kDa 150
α2M/MMP-2 complexes
100 75 proMMP-2 active MMP-2
50
MMP-2 catalytic domain
37 25 1
Zymography of Metalloproteinases
2
3
4
M
5
6
Figure 21.15.4 MMP samples were analyzed on a 7.5% acrylamide gelatin zymogram, incubated in developing buffer overnight at 37°C. Lane 1 shows 100 pg MMP-2 activated with 1 mM APMA, which results in a mixture of the 69-kDa full-length active form and the truncated 45-kDa catalytic domain. Preincubation of this sample with an equimolar amount of TIMP-2 for 1 hr at 37°C has no effect on the observed pattern of digestion because the enzyme/inhibitor complex dissociates in sample loading buffer (lane 2). Addition of a 50-fold molar excess of α2M to MMP-2 for 30 min at 37°C reduces the activity of both the 68- and 45-kDa proteolytically active species (lane 3). α2M differs from TIMP-2 in that the α2M/MMP-2 complex is stable in sample loading buffer. Preincubation of MMP-2 with TIMP-2 protects it from association with α2M (lane 4), since the MMP-2/TIMP-2 complex is not proteolytically active and hence unable to bind to α2M. Lane 5 shows a mixture of 72-kDa proMMP-2, 68-kDa MMP-2, and 45-kDA MMP-2 catalytic domain, preincubated with TIMP-2. Addition of α2M to this sample has no effect on the zones of hydrolysis observed for proMMP-2 as the zymogen is proteolytically inactive (lane 6). The MMP-2/TIMP-2 complex is similarly inactive and does not interact with α2M (lane 6). The truncated catalytic domain, however, interacts poorly at TIMP-2 and becomes associated with α2M (Itoh et al., 1998), shifting its zone of digestion from 45 kDa. A zone of lysis can often be observed associated with α2M. Lane M shows protein molecular mass markers.
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(Kjeldsen et al., 1993). Additionally, MMP-9 forms SDS-stable disulfide-bonded complexes with a 25-kDa protein called neutrophil gelatinase-associated protein (NGAL), producing a complex of 125 kDa (Kjeldsen et al., 1993).
It is crucial to add the inhibitors at the stated concentrations during all renaturing steps as well as in the development step to ensure reliable results.
Time Considerations Specific inhibitors to confirm class of proteinase To confirm that the observed proteinase is a metalloproteinase, specific proteinase inhibitors can be added to the renaturing and developing buffers. As shown in Figure 21.15.3, metalloproteinases (MMP-2 and -9 in this example) are inhibited by addition of 20 mM EDTA to the renaturing and developing buffers, but unaffected by addition of L-trans-epoxysuccinyl-leucylamido(4-guanidino)butane (E64), which inhibits cysteine proteinases at 20 µM; phenylmethylsulfonyl fluoride (PMSF) or 4-(2-aminoethyl)-benzenesulfonyl fluoride (AEBSF), which inhibit serine proteinases at 1 mM; or pepstatin A, which inhibits aspartic proteinases at 10 µM. α2-Macroglobulin (α2M) is a large plasma proteinase inhibitor that only binds to active proteinases and is therefore useful for identifying whether matrix metalloproteinases (MMPs) extracted from tissues or found in conditioned media are proteolytically active (Itoh et al., 1995). Most active endoproteinases from all four classes of proteinases (UNIT 21.1) readily cleave the bait region of α2M, which triggers a conformational change in the inhibitor and traps the proteinase within the molecule, hence rendering the proteinase inactive against large protein substrates (Barrett, 1981). Most α2M/proteinase complexes are SDS-stable so addition of α2M reduces activity at the normal zymogram position of the enzyme. While α2M complexes are inactive against protein substrates in solution, addition of SDS to the enzyme solution reduces the band at the enzyme’s molecular weight as the enzyme runs together with the α2M on the gel near the top of the zymogram where α2M runs. Figure 21.15.4 shows the effects of α2M on migration of active MMP-2 and inactive MMP-2/TIMP-2 complexes. Treatment of MMP-2 with α2M reduces the enzyme’s zone of digestion at 68 kDa (compare lane 1 with lane 3). Preincubation of MMP-2 with TIMP-2 generates a proteolytically inactive complex and hence protects the enzyme from interaction with α2M (compare lanes 2 and 4). Addition of α2M has no effect on the zone of digestion for either proMMP-2 or proMMP-2/TIMP-2, as both are proteolytically inactive (comparing lanes 5 and 6).
Preparation and polymerization of separating and stacking gels requires 1 to 2 hr. Sample preparation takes 10 to 15 min. Gels take ∼1 hr to load and run. Washing of gels in Triton X-100 (renaturation) takes 1 hr, and the development step varies between 2 and 20 hr, depending on the sensitivity required. Staining takes ∼30 min and destaining takes ∼2 hr.
Literature Cited Barrett, A.J. 1966. Chondromucoprotein-degrading enzymes. Nature 211:1188-1190. Barrett, A.J. 1981. Alpha 2-macroglobulin. Methods Enzymol. 80C:737-754. Bury, A.F. 1981. Analysis of protein and peptide mixtures. Evaluation of three sodium dodecyl sulfate-polyacrylamide gel electrophoresis buffer systems. J. Chromatogr. 213:491-500. Feitosa, L., Gremski, W., Veiga, S.S., Elias, M.C., Graner E., Mangili, O.C., and Brentani, R.R. 1998. Detection and characterization of metalloproteinases with gelatinolytic, fibronectinolytic and fibrinogenolytic activities in brown spider (Loxosceles intermedia) ven om. Toxicon 36:1039-1051. Gogly, B., Groult, N., Hornebeck, W., Godeau, G., and Pellat, B. 1998. Collagen zymography as a sensitive and specific technique for the determination of subpicogram levels of interstitial collagenase. Anal. Biochem. 255:211-216. Hattori, S., Fujisaki, H., Kiriyama, T., Yokoyama, T., and Irie, S. 2002. Real-time zymography and reverse zymography: A method for detecting activities of matrix metallopeptidases and their inhibitors using FITC-labeled collagen and casein as substrates. Anal. Biochem. 301:27-34. Hawkes, S.P., Hongxia, L., and Taniguchi, G.T. 2001. Zymography and reverse zymography for detecting MMPs and TIMPs. In Matrix Metallopeptidase Protocols (I.M. Clark, ed.) pp. 399410. Humana, Totowa, N.J. Hummel, K.M., Penheiter, A.R., Gathman, A.C., and Lilly, W.W. 1996. Anomalous estimation of protease molecular weights using gelatin-containing SDS-PAGE. Anal. Biochem. 233:140-142. Itoh, Y., Binner, S., and Nagase, H. 1995. Steps involved in the activation of pro-matrix metallopeptidase 2 (progelatinase A) and tissue inhibitor of metallopeptidases (TIMP)-2 by 4-aminophenylmercuric acetate. Biochem. J. 308:645-651.
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Itoh, Y., Ito, A., Iwata, K., Tanzawa, K., Mori, Y., and Nagase, H. 1998. Plasma membrane-bound tissue inhibitor of metallopeptidases (TIMP)-2 specifically inhibits matrix metallopeptidase 2 (gelatinase A) activated on the cell surface. J. Biol. Chem. 273:24360-24367.
Oliver, G.W., Leferson, J.D., Stetler-Stevenson, W.G., and Kleiner, D.E. 1997. Quantitative reverse zymography: Analysis of picogram amounts of metallopeptidase inhibitors using gelatinase A and B reverse zymograms. Anal. Biochem. 244:161-166.
Jain, A., Nanchahal, J., Troeberg, L., Green, P., and Brennan, F. 2001. Production of cytokines, vascular endothelial growth factor, matrix metallopeptidases, and tissue inhibitor of metallopeptidases 1 by tenosynovium demonstrates its potential for tendon destruction in rheumatoid arthritis. Arthritis Rheum. 44:1754-1760.
Troeberg, L., Tanaka, M., Wait, R., Shi, Y.E., Brew, K., and Nagase, H. 2002. E. coli expression of TIMP-4 and comparative kinetic studies with TIMP-1 and TIMP-2: Insights into the interactions of TIMPs and matrix metallopeptidase 2 (gelatinase A). Biochem. 41:15025-15035.
Kjeldsen, L., Johnsen, A.H., Sengeløv, H., and Borregaard, N. 1993. Isolation and primary structure of NGAL, a novel protein associated with human neutrophil gelatinase. J. Biol. Chem. 268:1042510432. Kleiner, D.E. and Stetler-Stevenson, W.G. 1994. Quantitative zymography: Detection of picogram quantities of gelatinases. Anal. Biochem. 218:329-329. Leber, T.M. and Balkwill, F.R. 1997. Zymography: A single step staining method for quantitation of proteolytic activity on substrate gels. Anal. Biochem. 249:24-28. Nagase, H. 1997. Activation mechanisms of matrix metallopeptidases. Biol. Chem. 378:151-160. Noha, M., Yoshida, D., Watanabe, K., and Teramoto, A. 2000. Suppression of cell invasion on human malignant glioma cell lines by a novel matrixmetallopeptidase inhibitor SI-27: In Vitro study. J. Neurooncol. 48:217-223.
Yasothornsrikul, S. and Hook, V.Y. 2000. Detection of proteolytic activity by fluorescent zymogram in-gel assays. Biotechniques 28:1166-1168. Yeow, K.M., Kishnani, N.S., Hutton, M., Hawkes, S.P., Murphy, G., and Edwards, D.R. 2002. Sorby’s fundus dystrophy tissue inhibitor of metallopeptidases-3 (TIMP-3) mutants have unimpaired matrix inhibitory activities, but affect cell adhesion to the extracellular matrix. Matrix Biol. 21:75-88. Zeng, Z.S., Shu, W.P., Cohen, A.M., and Guillem, J.G. 2002. Matrix metallopeptidase-7 expression in colorectal cancer liver metastases: Evidence for involvement of MMP-7 activation in human cancer metastases. Clin. Cancer Res. 8:144-148.
Contributed by Linda Troeberg and Hideaki Nagase Imperial College London London, United Kingdom
This work is supported by a Wellcome Trust grant no. 057508. The authors thank Abhilash Jain, Nidhi Sofat, and Yoshifumi Itoh for providing conditioned medium from cultured human wrist joint synovium, human chondrocytes, and TPA-treated HT1080, respectively.
Zymography of Metalloproteinases
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Monitoring Metalloproteinase Activity Using Synthetic Fluorogenic Substrates
UNIT 21.16
Fluorogenic synthetic substrates are commonly used to monitor the activity of proteinases in vitro. This unit describes an assay for matrix metalloproteinases (MMPs) using the substrate MCA-Pro-Leu-Gly∼Leu-Dpa(Dnp)-Ala-Arg-NH2, where MCA is the fluorophore (7-methoxycoumarin-4-yl)acetate, Dnp is the quencher dinitrophenyl, and the tilde (∼) represents the peptide-bond target of hydrolysis. This substrate was first described by Knight et al. (1992) for the assay of matrix metalloproteinase (MMP)-1, -2, and -3, and is now widely used as a general MMP substrate (Fig. 21.16.1). Protocols are given for both stopped-time (see Basic Protocol 1) and continuous (see Basic Protocol 2) assays. The former is suitable for assaying MMP activity in a large number of samples, while the latter is commonly used when establishing an assay protocol or investigating kinetic aspects of enzyme behavior. Other fluorogenic peptide and protein substrates are also discussed, along with nonfluorogenic alternatives. MMP concentration can be estimated from absorbance at 280 nm, but for accurate determination, a method for titrating the amount of proteinase is presented (see Support Protocol). For a more thorough discussion of MMPs and TIMPs, see UNIT 21.4. NOTE: Perform all assays at least in duplicate.
MCA
Dnp
Pro-Leu-Gly~Leu-Dpa-Ala-Arg-NH2
MMP cleavage
MCA Dnp Pro-Leu-Gly Leu-Dpa-Ala-Arg-NH2
Figure 21.16.1 Schematic diagram of MMP hydrolysis of MCA-Pro-Leu-Gly∼Leu-Dpa(Dnp)-AlaArg-NH2. This MMP substrate contains a methoxycoumarin (MCA) fluorophore and a dinitrophenyl (Dnp) quencher located on opposite sides of the susceptible peptide bond, indicated by a tilde (∼). The excitation peak of the quencher overlaps with the emission peak of the fluorophore, allowing the quencher to absorb the energy from the fluorophore in a distance-dependent manner. This prevents fluorescence of the intact substrate by a process of fluorescence resonance energy transfer (FRET). Upon hydrolysis of the susceptible peptide bond, the quencher and the fluorophore become physically separated and the fluorescence of the MCA group can be detected at 393 nm upon excitation at 325 nm.
Peptides
Contributed by Linda Troeberg and Hideaki Nagase
21.16.1
Current Protocols in Protein Science (2003) 21.16.1-21.16.9 Copyright © 2003 by John Wiley & Sons, Inc.
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BASIC PROTOCOL 1
STOPPED-TIME ASSAYS FOR MMPs USING A FLUOROGENIC SUBSTRATE Stopped-time assays can be performed in either a microwell-plate or in microcentrifuge tubes, depending on the type of fluorometer available and the sensitivity required. Use a 96-well plate if a microwell-plate fluorometer (e.g., Molecular Devices SPECTRAmax) is available or choose the microcentrifuge tube assay for a cuvette-based fluorometer (e.g., Perkin Elmer LS-50B). Cuvette-based fluorometers tend to be more sensitive, but each sample must be read individually, increasing the total assay time. Microwell-plate fluorometers such as the SPECTRAmax are better suited for routine assay of larger numbers of samples. The choice of enzyme concentration will depend on the identity of the MMP, as some hydrolyze the substrate faster than others. For example, MMP-2 hydrolyzes it rapidly so a final enzyme concentration of 0.1 to 1 nM is appropriate. For MMP-3, which hydrolyzes the substrate more slowly, a final concentration of 1 to 10 nM is more suitable. If the concentration of the MMP within the sample is unknown, various dilutions covering a 10- to 50-fold range in concentration should be tested. Materials 2–2000 nM MMP solution TNC assay buffer (see recipe) at 37°C 0.5 M EDTA 3 µM MCA-Pro-Leu-Gly∼Leu-Dpa(Dnp)-Ala-Arg-NH2 (see recipe) 10× stop solution (see recipe) 96-well microplate Spectrofluorometer—e.g., LS-50B luminescence spectrometer (Perkin Elmer) or SPECTRAmax Gemini XS spectrofluorometer (Molecular Devices) For microplate format 1a. In an appropriate number of wells in a 96-well microplate, mix 10 µl of 2 to 2000 nM MMP solution with 90 µl TNC assay buffer. As negative controls, include one set of wells with 10 µl TNC assay buffer instead of enzyme, and another with 10 µl MMP solution, 2 µl of 0.5 M EDTA (10 mM final), and 88 µl TNC assay buffer. Equilibrate 10 min at 37°C. If necessary, the volume of enzyme added can be altered up to a maximum of 100 ìl. An optional positive control can be included (e.g., 1 to 10 nM recombinant MMP). See Critical Parameters for information on how to evaluate MMP inhibitors.
2a. Add 100 µl of 3 µM MCA-Pro-Leu-Gly∼Leu-Dpa(Dnp)-Ala-Arg-NH2 (1.5 µM final). Substrate hydrolysis begins as soon as the substrate is added, so add the substrate as quickly as possible. A multichannel pipettor can be used.
3a. Incubate 10 min to 1 hr at 37°C. Add 20 µl of 10× stop solution to each well in the order that the substrate was added as quickly as possible to ensure that all wells have been incubated with substrate for the same amount of time. Stop solution contains EDTA which stops the reaction by chelating the Zn2+ required for MMP activity.
Monitoring Metalloproteinase Activity With Fluorogenic Substrates
Increasing the time of incubation will increase the amount of product generated and hence the fluorescence signal. When establishing an assay for a particular enzyme, it is useful to prepare multiple wells for a particular enzyme concentration and stop the reaction after different times (e.g., 5, 10, 20, 30, and 60 min) to establish an appropriate assay time for further experiments. See Commentary concerning establishing the linear range of the assay.
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4a. Read the fluorescence using an excitation wavelength of 325 nm and an emission wavelength of 393 nm according to the manufacturer’s instructions. For microcentrifuge tube format 1b. To each microcentrifuge tube, mix 50 µl of 2 to 2000 nM MMP solution with 450 µl TNC assay buffer. As negative controls, include one set of wells with 50 µl TNC assay buffer instead of enzyme, and another with 50 µl MMP solution, 10 µl of 0.5 M EDTA, and 440 µl TNC assay buffer. Equilibrate 10 min at 37°C. If necessary, the volume of enzyme added can be increased to a maximum of 500 ìl in an appropriate buffer. Positive controls can be included as described in step 1a. See Critical Parameters for information on how to evaluate MMP inhibitors.
2b. Add 500 µl of 3 µM MCA-Pro-Leu-Gly∼Leu-Dpa(Dnp)-Ala-Arg-NH2 (1.5 µM final). 3b. Incubate 10 min to 1 hr at 37°C. Add 100 µl of 10× stop solution to each tube to each well as quickly as possible and in the order that the substrate was added to ensure that all wells have been incubated with substrate for the same amount of time. 4b. Sequentially aspirate the assay solutions into an appropriate cuvette and read the fluorescence using an excitation wavelength of 325 nm and an emission wavelength of 393 nm according to the manufacturer’s instructions. CONTINUOUS ASSAY FOR MMPs USING A FLUOROGENIC SUBSTRATE Continuous assays permit monitoring of enzyme-catalyzed reactions in more detail than their stopped-time counterparts and are usually performed when investigating the enzyme mechanism or rate at which proteinases interact with inhibitors (Salvesen and Nagase, 2001). Such assays are conducted directly in a fluorometer, with the increase in substrate hydrolysis monitored semicontinuously over the assay period rather than once at the end. Some machines (e.g., the LS-50B; Perkin Elmer) allow for detailed monitoring of a single cuvette, with readings taken several times a second. Other fluorometers (e.g., SPECTRAmax; Molecular Devices) allow for less detailed monitoring of several samples in a microwell-plate format.
BASIC PROTOCOL 2
Materials 3 µM MCA-Pro-Leu-Gly∼Leu-Dpa(Dnp)-Ala-Arg-NH2 (see recipe) TNC assay buffer (see recipe) 2–2000 nM MMP solution 10× stop solution (see recipe) Spectrofluorometer with 37°C circulating water bath—e.g., LS-50B luminescence spectrometer (Perkin Elmer) or SPECTRAmax Gemini XS spectrofluorometer (Molecular Devices) NOTE: Volumes given are for the cuvette method. For microplate assays, scale the volumes down 5-fold. 1. In appropriate cuvette, mix 500 µl of 3 µM MCA-Pro-Leu-Gly∼Leu-Dpa(Dnp)-AlaArg-NH2 with 450 µl TNC assay buffer. Place the cuvette in a spectrofluorometer with circulating water bath set at 37°C. Equilibrate 10 min. If it is necessary to vary the volume of enzyme added, the volume of TNC assay buffer should be varied accordingly to give a total volume of 500 ìl.
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2. Add 50 µl of 2 to 2000 nM MMP solution, and mix well. Close the fluorometer lid as quickly as possible and monitor the increase in product fluorescence for as long as required using an excitation wavelength of 325 nm and an emission wavelength of 393 nm according to manufacturer’s instructions. Substrate hydrolysis begins as soon as the enzyme is added. The increase in product fluorescence is monitored from the time the lid is closed. The length of time for which fluorescence should be monitored depends on the purpose of the assay. To determine the amount of enzyme present in a sample, it is sufficient to monitor the reaction for 2 to 5 min, allowing determination of the initial rate of enzyme activity; however, if the purpose of the assay is to monitor association with an inhibitor, then activity should be monitored until equilibrium is reached, which can take in excess of 1 hr.
3. Discard the reaction product, wash the cuvette well with water, and repeat the process (steps 1 to 3) for as many reactions as desired. The cuvette should periodically be cleaned more thoroughly by soaking in 1% (w/v) sodium hypochlorite for ∼1 hr. A dilute detergent solution can also be used. Take care to completely wash out this cleansing solution before resuming measurements. SUPPORT PROTOCOL
QUANTITATION OF PEPTIDASE ACTIVITY BY TITRATION AGAINST TIMPS MMP concentrations can be estimated from their absorbance at 280 nm (e.g., molar extinction coefficients are 64,390 M−1cm−1 for human MMP-1, 122,870 M−1cm−1 for MMP-2, and 67,640 M−1cm−1 for MMP-3. Values for other MMPs can be determined using the sequence analysis tools on SwissProt (http://ca.expasy.org). However, some purified MMPs (e.g., MMP-2) are prone to autocatalytic degradation, and their active concentration cannot be accurately determined from their absorbance at 280 nm. In this case, the active concentration of an MMP preparation can be determined by titrating it against a tissue inhibitor of metalloproteases (TIMP) solution of known concentration. TIMPs are very stable molecules whose concentrations can be accurately calculated from their A280. Human TIMP-1 and TIMP-2 have molar extinction coefficients of 24,750 and 31,720 M−1cm−1, respectively. TIMPs are commercially available from Chemicon, R & D Systems, and Oncogene Research Products. To titrate the enzyme, incubate ∼20 nM MMP (estimated from its A280) 1 hr with a range (5 to 30 nM) of known TIMP concentrations at 37°C in a total volume of 100 µl. This will allow the formation of stable, inactive TIMP/MMP complexes. Assay the active MMP against 100 µl of 3 µM MCAPro-Leu-Gly∼Leu-Dpa(Dnp)-Ala-Arg-NH2 as described above (see Basic Protocol 1). Stopped-time assays are recommended for titrations, since multiple samples must be assayed. Calculate the concentration of MMP based on the minimum concentration of TIMP required for complete inhibition and the fact that TIMP/MMP complexes have 1:1 stoichiometry (example given in Troeberg et al., 2002). REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Monitoring Metalloproteinase Activity With Fluorogenic Substrates
MCA-Pro-Leu-Gly∼Leu-Dpa(Dnp)-Ala-Arg-NH2, 3 ìM Prepare a 10 mM solution by dissolving 11 mg MCA-Pro-Leu-Gly∼Leu-Dpa(Dnp)Ala-Arg-NH2 (Bachem or Peptides International) in 1 ml dimethyl sulfoxide (DMSO). Stir at room temperature until the substrate is fully dissolved. Store protected from light (i.e., in a brown bottle or wrapped in aluminum foil) up to 12 months at −20°C. It is convenient to prepare a 100 µM solution by diluting 10 µl of the 10 mM solution into 990 µl TNC assay buffer (see recipe). Store as described
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Table 21.16.1
Fluorogenic Peptide Substrates Commonly Used to Assay MMPsa
Substrateb,c
MMPs assayed
MCA-Pro-Leu-Gly∼Leu-Dpa(Dnp)- Ala-Arg-NH2
-1, -2, -3, -7 Standard, multipurpose substrate -2, -3 Multipurpose
MCA-Pro-Lys-Pro-Gln-Gln-Phe-Phe-Gly∼Leu-Lys-(Dnp)-Gl MCA-Pro-Leu-Ala∼Cys(Mob)-Tro-Ala-Arg-Dpa(Dnp)NH2
MCA-Arg-Pro-Lys-Pro-Tyr-Ala∼Nva-Trp-Met-Lys(Dnp)-NH MCA-Arg-Pro-Lys-Pro-Val-Glu∼Nva-Trp-Arg-Lys(Dnp)-NH2 DABCYL-γ-Abu-Pro-Gln-Gly∼Leu-Glu(EDANS)-Ala Lys-NH2
-3, -14
-3, -2, -9
Use
Useful for MMP-3 and MMP-14 Multi-purpose
Reference Knight et al. (1992) Nagase et al. (1994) Muncha et al. (1998)
Nagase et al. (1994) -3 Preferential Nagase et al. MMP-3 substrate (1994) -1, -2, -3, -9 Complex Beekman et al. biological samples (1996)
aAbbreviations: ABU, L-α-aminobutyryl(s)-2-amino-butanoyl; DABCYL, 4-(4-dimethylaminophenylazo)benzoyl; Dnp, dinitrophenol; Dpa, L-α,
β-diaminopropionic acid; EDANS, 5-[(2-aminoethyl)amino]napththalene-1-sulfonic acid; Mob, 4-methoxybenzyl; Nva, norvaline. bThese fluorogenic peptide substrates, sold by Bachem or Peptides International, can be used to assay MMP activity in vitro. The included references
describe their hydrolysis by the listed MMPs, but it should be remembered that these substrates may also be hydrolyzed by other enzymes not listed below. cThe tilde (∼) represents the target of hydrolysis.
for the 10 mM solution. Immediately before use, prepare a 3 µM working solution by diluting 150 µl of the 100 µM solution to 5 ml with TNC assay buffer. This is discarded after use. Dnp is the quencher moiety dinitrophenyl, Dpa is 2,3-diaminopropionic acid MCA is the fluorophore (7-methoxycoumarin-4-yl)acetate. See Critical Parameters and Table 21.16.1 for information on other proteinase substrates.
Stop solution, 10× Add 37.2 g disodium EDTA dihydrate (100 nM) and 0.2 g NaN3 (3 mM) to ∼900 ml water while stirring. Adjust pH to 8.0 with HCl and continue stirring until all of the salts dissolve. Adjust volume to 1000 ml with water. Store up to 3 months at room temperature. TNC assay buffer Dissolve 6.055 g Tris base (50 mM), 8.766 g NaCl (0.15 M), 1.47 g CaCl2⋅2 H2O (10 mM) and 0.2 g NaN3 (0.02%) in ∼900 ml of water. Adjust pH to 7.5 at room temperature with HCl and volume to 1000 ml with water. Store up to 1 month at room temperature. COMMENTARY Background Information Fluorogenic groups Numerous substrates are available to monitor the activity of proteinases in vitro. Some of these substrates are proteins, but most are small synthetic peptides similar to the peptide se-
quences recognized in natural protein substrates. Most in vitro substrates are labeled in some way so that their hydrolysis generates an easily detectable signal. MMP substrates most commonly utilize fluorogenic groups such as 7-methoxycoumarin, which are highly sensitive and easy to use.
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Fluorogenic groups (or fluorophores) are excited at one wavelength, known as the excitation wavelength, and emit this energy at a higher wavelength. In addition to a fluorophore, MMP substrates commonly also contain a quencher substituent group located on the opposite side of the susceptible peptide bond. The quencher, as its name suggests, prevents fluorescence emission from the fluorophore in the intact substrate by a process known as fluorescence resonance energy transfer (FRET). This occurs when the excitation wavelength of the quencher is within the range of the emission wavelengths of the fluorophore, enabling the quencher to absorb the emitted light and hence prevent fluorescence when the two are in close proximity (e.g., in the uncleaved substrate). Upon peptide hydrolysis, the quencher and fluorophore are separated, allowing the fluorophore to generate an unquenched signal. The substrate described in this protocol is MCA-ProLeu-Gly∼Leu-Dpa(Dnp)-Ala-Arg-NH2, where MCA is the fluorophore and Dnp the quencher (Fig. 21.16.1). Michaelis-Menten kinetics To obtain meaningful data from enzyme activity assays, it is necessary to have at least a working understanding of the Michaelis-Menten equation, which describes the hyperbolic relationship between the substrate concentration ([S]) and the initial rate of substrate hydrolysis (v) for a fixed enzyme concentration. This is explained in detail in the legend to Figure 21.16.2. The true Km value of each MMP for MCA-Pro-Leu-Gly∼Leu-Dpa(Dnp)-AlaArg-NH2 is not known, because absorptive quenching effects prevent use of this substrate at sufficiently high concentrations to allow accurate Km calculation (Knight et al., 1992). Nevertheless, at [S] <10 µM, the initial rate of substrate hydrolysis by MMP-1, -2, and -3 is proportional to the substrate concentration, indicating that Km is greater than this value. In the protocol described above, MCA-Pro-LeuGly∼Leu-Dpa(Dnp)-Ala-Arg-NH2 is used at a concentration (1.5 µM final) below its Km for these MMPs, which hence function at a rate below Vmax.
Critical Parameters Monitoring Metalloproteinase Activity With Fluorogenic Substrates
Other 7-methoxycoumarin peptide substrates The rate at which a fluorogenic peptide is hydrolyzed by a particular MMP depends on the substrate preference of the enzyme. For
instance, MCA-Pro-Leu-Gly∼Leu-Dpa(Dnp)Ala-Arg-NH2 is readily hydrolyzed by MMP-2 (kcat/Km = 629,000 M−1⋅sec−1) and MMP-7 (kcat/Km = 169,000 M−1⋅sec−1) and less readily by MMP-1 (kcat/Km = 14,800 M−1⋅sec−1) and MMP-3 (kcat/Km = 23,000 M−1⋅sec−1; Knight et al., 1992). For monitoring the presence of MMPs in a sample (e.g., during purification), any substrate that gives a sufficiently detectable signal can be used, while for kinetic studies, it is often desirable to use as sensitive a substrate as possible (i.e., high kcat/Km). Table 21.16.1 shows some of the commercially available substrates suitable for assaying MMPs. Fluorogenically labeled proteinase substrates Various fluorogenic peptide substrates that mimic native triple-helical collagen have been described (Lauer-Fields et al., 2001). These are not specific for collagenases, but may be useful for studying the triple-helicase activity of the collagenases MMP-1, -2, -8, -13, and -14. Fluorescein-labeled gelatin, commercially available as DQ gelatin from Molecular Probes, is used for in vitro and cell culture assay of MMP activity (D’Angelo et al., 2001). Nonfluorogenic labels Protein substrates can be easily radiolabeled in vitro by acetylation using [3H]acetic anhydride (Cawston and Barrett, 1979) or by reduction and carboxymethylation using [3H]iodoacetic acid (Nagase, 1995). Protocols for the synthesis of 14C-radiolabeled peptide substrates have also been described (Brownell et al., 1994). Biotinylation of substrate proteins provides a simple, nonradioactive alternative (Ratnikov et al., 2000). Biotinylated-gelatin and -collagen assay kits are sold by Chemicon. Hydroxamates and other inhibitors Various synthetic inhibitors have been developed with the goal controlling MMP activity in a variety of pathological conditions, ranging from cancer to arthritis. These can be used in vitro to confirm the presence of a metalloproteinase in a sample. The hydroxymate (e.g., 10 µM GM6001, Elastin Products) is preincubated with 0.1 to 10 nM MMP for 1 hr at 37°C to allow equilbrium to be reached. The synthetic substrate is then added as described (see Basic Protocol 1). Comparing treated samples with untreated controls will indicate whether the compound inhibits MMP activity. The inhibitory properties of the compound can be studied in more detail (e.g., Ki determined) by analyz-
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ing the rate with which equilibrium between peptidase and inhibitor is reached (discussed in detail in Salvesen and Nagase, 2001).
Troubleshooting Substrate background is too high Always perform a negative substrate control, including just the substrate and the assay buffer. This gives a measure of the inherent substrate fluorescence or background. If this value is high, the substrate may already be hydrolyzed. Make a fresh working solution and, if necessary, a fresh stock solution. Assay buffers can become contaminated with bacteria, which secrete enzymes that commonly hydrolyze peptide substrates. As another negative control, add stop solution to a mixture of assay buffer and enzyme before addition of the substrate. This control will reveal the presence of fluorogenic material in the enzyme solution. Fluorescence does not increase when enzyme is added There are several possible reasons for such a result. First, the enzyme concentration may
be too low. Add a higher concentration or greater volume of enzyme, or increase the incubation time. The enzymes may be present in an inactive zymogen form. Most MMP zymogens can be activated by addition of organomercurials such as 4-aminophenylmercuric acetate (APMA). Test whether addition of 1 mM APMA for 1 to 4 hr at 37°C increases the activity. Alternatively, they may be complexed with inhibitors such as TIMPs, in which case it will not to possible to observe their activity using this method. Zymography may provide a suitable alternative (UNIT 21.15). The assay buffer may not be suitable for enzyme activity. For MMPs, ∼10 mM Ca2+ must be present in the assay buffer, which must have a pH 7.0 to 7.8. The sample may have been treated in a manner that has inactivated the proteinases. Enzymes may be visible by immunoblot (UNIT 10.10), but will not be active against synthetic substrates in the presence of reducing agents or if they have been boiled.
ν Vmax
Vmax 2
ν =
Vmax × [S] Km + [S]
[S] Km
Figure 21.16.2 Schematic diagram of the Michaelis-Menten equation. The equation (inset) describes the hyperbolic relationship between substrate concentration ([S]) and the initial rate of substrate hydrolysis (v) for a fixed enzyme concentration. At low substrate concentration, v increases linearly with [S]. Any increase in [S] causes a corresponding increase in the rate of enzyme activity. With increasing [S], however, the enzyme eventually becomes saturated with substrate and operates at its maximum velocity (Vmax). Vmax is directly proportional to kcat, the turnover number, which represents the maximum number of substrate molecules hydrolyzed by one molecule of enzyme per second. Km, the Michaelis constant, is defined as the substrate concentration that gives v = 0.5 Vmax and gives an indication of the affinity of the enzyme for the substrate. If the enzyme concentration is doubled, Vmax doubles, but Km remains constant.
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Anticipated Results Figure 21.16.3 shows typical results for MMP-2 hydrolysis of MCA-Pro-LeuGly∼Leu-Dpa(Dnp)-Ala-Arg-NH2. Fo r 0.1 and 0.2 nM MMP-2, the rate of substrate hydrolysis is linear throughout the 30-min assay period. Additionally, the rate of substrate hydrolysis (v) by 0.2 nM MMP-2 is double that of the 0.1 nM reaction, indicating that at these enzyme concentrations, v is proportional to the enzyme concentration ([E]). For 0.4 nM MMP-2, however, v is linear (and twice v for 0.2 nM
MMP-2) for only the first 10 to 15 min, after which it decreases as the substrate concentration becomes limiting. Figure 21.16.3B shows results for the stopped-time assays. Note that if several stopped-time assays are conducted for each enzyme concentration (as shown), then the same conclusions can be drawn as from Figure 21.16.3A. If only one stopped-time assay is conducted at 30 min, however, it will not be clear that v is nonlinear for 0.4 nM MMP-2. In this case, if a test sample were analyzed, an incorrect [E] would thus be inferred from the
Fluorescence (arbitary units)
A
Fluorescence (arbitary units)
B
5
10
15
20
25
30
Time (min)
Monitoring Metalloproteinase Activity With Fluorogenic Substrates
Figure 21.16.3 Typical results for continuous and stopped-time assays of 0.1 (closed circle), 0.2 (open circles), or 0.4 (closed square) nM MMP-2 hydrolysis of 15 µM MCA-Pro-Leu-Gly∼LeuDpa(Dnp)-Ala-Arg-NH2. (A) Continuous assays of 1-ml reactions (readings taken every 0.1 sec) performed in an LS-50B luminescence spectrometer (Perkin Elmer). Fluorescence values for the substrate alone (substrate blank) have been subtracted to generate the data shown. For 0.1 and 0.2 nM MMP-2, the rate of substrate hydrolysis (v) is linear throughout the assay period, indicating that v is proportional to the enzyme concentration ([E]) throughout. For 0.4 nM MMP-2, however, v is linear only for the first 15 min of the assay and then decreases, indicating that the substrate becomes limiting and that v is no longer proportional to [E]. (B) Stopped-time assays of 200-µl reactions stopped at 5, 10, 15, 20, 25 or 30 min, measured at 25°C in microplates, and read in a SPECTRAmax spectrofluorometer. In contrast with the continuous assay (A), where fluorescence data is collected from each assay every 0.1 sec, here data is collected only once from each assay, after its termination. Data points represent average fluorescence of triplicate measurements.
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fluorescence read at 30 min. It is thus crucial to establish the linear range for each enzyme assay—i.e., the range of enzyme concentrations and assay times for which v is directly proportional to [E]. This is normally dictated by the percentage hydrolysis of the substrate: the assay remains linear provided that no more than 5% to 10% of the total amount of substrate is hydrolyzed during an assay. The assay shown in Figure 21.16.3 is linear up to ∼10% substrate hydrolysis. While MCA-Pro-Leu-Gly∼Leu-Dpa(Dnp)Ala-Arg-NH2 is efficiently hydrolyzed by many MMPs, it is not a specific substrate and can also be hydrolyzed by other metalloproteinases as well as proteinases of other classes. Thus, it is not possible to infer that a test sample showing activity against this substrate contains an MMP. For example, the bacterial metalloproteinase thermolysin readily cleaves this substrate. To ascertain that an MMP is present, it is necessary to add specific MMP inhibitors such as naturally occurring TIMPs (see Table 21.4.3 in UNIT 21.4).
Time Considerations Stopped-time assays take about 30 min to prepare and substrate hydrolysis is typically monitored for 10 min to 1 hr. Longer incubations can be performed if increased sensitivity is required. All of the samples in a microplate can be read within 2 min, but if using a cuvettebased fluorometer, each sample can take up to 1 min to read. Continuous assays generally take longer to perform, with each assay ranging from 10 min to several hours depending on the purpose of the assay.
Literature Cited Beekman, B., Drijfhout, J.W., Bloemhoff, W., Ronday, H.K., Tak, P.P., and TeKoppele, J.M. 1996. Convenient fluorometric assay for matrix metallopeptidase activity and its application in biological media. FEBS Lett. 390:221-225. Brownell, J., Earley, W., Kunec, E., Morgan, B.A., Olyslager, B., Wahl, R.C., and Houck, D.R. 1994. Comparison of native matrix metallopeptidases and their recombinant catalytic domains using a novel radiometric assay. Arch. Biochem. Biophys. 314:120-125.
Cawston, T.E. and Barrett, A.J. 1979. A rapid and reproducible assay for collagenase using [114 C]acetylated collagen. Anal. Biochem. 99:340345. D’Angelo, M., Billings, P.C., Pacifici, M., Leboy, P.S., and Kirsch, T. 2001. Authentic matrix vesicles contain active metallopeptidases (MMP). J. Biol. Chem. 276:11347-11353. Knight, C.G., Willenbrock, F., and Murphy, G. 1992. A novel coumarin-labelled peptide for sensitive continuous assays of the matrix metallopeptidases. FEBS Lett. 296:263-266. Lauer-Fields, J.L., Broder, T., Sritharan, T., Chung, L., Nagase, H., and Fields, G.B. 2001. Kinetic analysis of matrix metallopeptidase activity using fluorogenic triple-helical substrates. Biochemistry 40:5795-5803. Muncha, A., Cuniasse, P., Kannan, R., Beau, F., Yiotakis, A., Basset, P., and Dive, V. 1998. Membrane type-1 matrix metallopeptidase and stromelysin-3 cleave more efficiently synthetic substrates containing unusual amino acids in their P1 positions. J. Biol. Chem. 273:27632768. Nagase, H. 1995. Human stromelysins 1 and 2. Methods Enzymol. 248:449-470. Nagase, H., Fields, C.G., and Fields, G.B. 1994. Design and characterization of a fluorogenic substrate selectivity hydrolyzed by stromelysin 1 (matrix metallopeptidase-3). J. Biol Chem. 269:20952-20957. Ratnikov, B., Deryugina, E., Leng, J., Marchenko, G., Dembrow, D., and Strongin, A. 2000. Determination of matrix metallopeptidase activity using biotinylated gelatin. Anal. Biochem. 286:149-155. Salvesen, G. and Nagase, H. 2001. Inhibition of proteolytic enzymes. In Proteolytic Enzymes: A Practical Approach, Second edition (R.J. Benyon and J.S. Bond, eds.) pp 105-130. Oxford University Press, Oxford. Troeberg, L., Tanaka, M., Wait, R., Shi, Y.E., Brew, K., and Nagase, H. 2002. E. coli expression of TIMP-4 and comparative kinetic studies with TIMP-1 and TIMP-2: Insights into the interactions of TIMPs and matrix metallopeptidase 2 (gelatinase A). Biochemistry 41:15025-15035.
Contributed by Linda Troeberg and Hideaki Nagase Imperial College London London, United Kingdom
This work is supported by Wellcome Trust grant no. 057508.
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Applications for Chemical Probes of Proteolytic Activity
UNIT 21.17
The tools used to study peptidase biology have been evolving at a rapid pace as interest in this field continues to grow. At the heart of virtually all peptidase research have been small molecule inhibitors. Furthermore, the discovery that specific peptidase inhibitors can lead to beneficial effects in certain disease indications ignited vigorous work by chemists to design small molecules that bind tightly within the active site of a peptidase, thus preventing access to substrates. As a result of this work, covalent and noncovalent inhibitors have been designed for virtually every major class of peptidase. Many of these reagents have been developed with the goal of creating new therapeutic drugs, while still others are used as “benchmark” compounds to assess the contribution of a general peptidase family to a given biological process. Regardless of the common uses for these compounds, there has recently been a push to re-engineer old inhibitors for new applications. The application of fluorescent, radioactive, and biotin tags to general inhibitor scaffolds has led to the creation of new families of activity-based probes (ABPs) that can be used to covalently label peptidases in complex biological samples. These reagents are proving to be useful for applications ranging from simple screening for recombinant peptidase expression efficiency to imaging specific peptidases in vivo during tumor progression. This unit outlines the process of probe selection, probe tagging using reporters, labeling of recombinant targets and crude protein extracts, labeling of peptidase targets in cell culture systems, and finally affinity purification and inhibitor screening using ABPs. The unit begins by outlining the process of selecting the appropriate reagent for the peptidase or peptidase family of interest (see Strategic Planning). Probes have been designed to label many different classes of peptidases ranging from caspases to the proteasome. Table 21.17.1 presents a list of commonly used probes as well as sources and references for use of these probes. It is highly recommended that readers refer to this table for probe selection and background reading regarding probe use and target peptidase biology. Sources of probes can vary, and some are commercially available while others need to be obtained through collaborations/gifts from laboratories that routinely synthesize and use the probes. In some cases probes can only be purchased in a form that requires in-house labeling with a tag such as radioactive iodine (125I). For this reason, simple protocols are included that can be used to radioiodinate a probe (see Basic Protocol 1 and Alternate Protocol 1). While this type of labeling strategy requires some preparation, in many cases these 125I-labeled probes provide the cleanest and most easily interpretable results. However, probes that can be obtained with fluorescent and biotin tags already attached provide for the least amount of setup requirements, are easy to work with, and often provide equally high-quality results. In some cases, probes can be obtained commercially that require attachment of biotin or fluorescent tags. Biotin and fluorophore labeling of ABPs are described in Alternate Protocol 2. Each of the probe-tagging strategies requires different detection methods, so protocols outlining detection methods for each label type are included (see Basic Protocol 2 and Alternate Protocols 3, 4, and 5). In many cases, a probe is unique in terms of its overall reactivity with specific targets within a given set of experimental conditions. Therefore, a section is included to help the investigator deal with some of the common parameters that may need to be optimized to provide desired results (see Basic Protocol 3). These parameters will depend on the kinetics of probe labeling and may require optimization of conditions such as protein and Peptidases Contributed by Matthew Bogyo, Amos Baruch, Douglas A. Jeffery, Doron Greenbaum, Anna Borodovsky, Huib Ovaa, and Benedikt Kessler
21.17.1
Current Protocols in Protein Science (2004) 21.17.1-21.17.35 Copyright ©2004 by John Wiley & Sons, Inc.
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Table 21.17.1
List of Commonly Used Peptidase Probes and Their Sources
Peptidase family
Specific targets
Available probes
Source and/or reference
Cysteine peptidases Papain family Biotin tags Cathepsins/calpains DCG-04 Cathepsin B NS-196 Fluorescent tags Cathespins/calpains BODIPY-DCG-04 Radiolabels Cathespins/calpains Cbz-Try-Ala-N2 JPM-565 JPM-OEt Cathepsin B LHVS-PhOH MB-074 Caspasesa Biotin tags General caspase Biotin-X-VAD(OMe)-fmk Caspase 1,4 Z-VK-X-(biotin)-D(OMe) -fmk Biotin-YVAD-cmk Biotin-YVAD-faom Caspase 3,6,7,10 Ac-YVK(biotin)-D-aom Biotin-DEVD-CHO Biotin-X-D(OMe)E (OMe)VD(OMe)-fmk Fluorescent tags General caspase SR-VAD-fmk FAM-VAD-fmk Caspase 1
FAM-YVAD-fmk
Caspase 2
FAM-VDVAD-fmk
Caspase 3
SR-DEVD-fmk FAM-DEVD-fmk
Caspase 6
FAM-VEID-fmk
Caspase 8
FAM-LETD-fmk
Caspase 9
FAM-LEHD-fmk
Caspase 10
FAM-AEVD-fmk
Caspase 13
FAM-LEED-fmk
Greenbaum et al. (2000) Schaschke et al. (2000) Greenbaum et al. (2002a) Enzyme System Products Shi et al. (1992) Bogyo et al. (2000) Bogyo et al. (2000) Bogyo et al. (2000)
Calbiochem Calbiochem Calbiochem Calbiochem Calbiochem Calbiochem Calbiochem
Immunochemistry Technologies, LLC Immunochemistry Technologies, LLC Immunochemistry Technologies, LLC Immunochemistry Technologies, LLC Immunochemistry Technologies, LLC Immunochemistry Technologies, LLC Immunochemistry Technologies, LLC Immunochemistry Technologies, LLC Immunochemistry Technologies, LLC Immunochemistry Technologies, LLC Immunochemistry Technologies, LLC continued
Applications for Chemical Probes of Proteolytic Activity
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Table 21.17.1
List of Commonly Used Peptidase Probes and Their Sources, continued
Peptidase family
Specific targets
Available probes
Source and/or reference
HA tags HA-Ub-VS HA-Ub-Cl HA-Ub-Br2 HA-Ub-Br3 HA-Ub-VME HA-Ub-BsPh
Borodovsky et al. (2002) Borodovsky et al. (2002) Borodovsky et al. (2002) Borodovsky et al. (2002) Borodovsky et al. (2002) Borodovsky et al. (2002)
Deubiquintinating enzymes
Arg-gingipain
Serine peptidases All hydrolyases
Biotin tags BiRK Radiolabels 3H-DFP
Biotin tags FP-Biotin FP-Peg-Biotin Chymotrypsin-like Biotin-AAF-cmk All hydrolyases
All hydrolyases
Threonine peptidases
Fluorescent tags FP-Peg-TRM FP-fluorescein FP-Peg-fluorescein
Mikolajcyk et al. (2003) PerkinElmer Life and Analytical Sciences Liu et al. (1999) Kidd et al. (2001) Enzyme Systems Products Patricelli et al. (2001) Liu et al. (1999) Patricelli et al. (2001)
The proteasome Radiolabels NP-LLL-VS
Cabiochem, Bogyo et al. (1997)
Biotin tags Epoxomicin biotin Meng et al. (1999) AdaLys(Bio)AhX3L3-VS Calbiochem, Kessler et al. (2001) aWhile there are commercially available inhibitors and probes for specific caspases, in reality the specificity of these
reagents is far from absolute. Care should be taken when interpreting results using “specific” caspase inhibitors.
probe concentration, temperature of the labeling reaction, pH, and buffer ingredients. Changing these conditions can have a significant effect on the labeling performance of some probes. Therefore, this protocol is a good starting point for working with any new probe. The authors have dedicated a set of protocols to the use of the ABPs on live cell systems (see Basic Protocol 4 and Alternate Protocols 6 and 7). While the in situ labeling method described in these protocols provides for the most “native” environment for reaction of probes with target peptidases, it does not work for all probes, as many are non–cell permeable. In some cases, it is desirable to use a non–cell permeable reagent to assay the activity of peptidases on the cell surface or in the culture medium (see Alternate Protocol 7). Peptidases
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Regardless of whether the probe has been used in cellular/tissue extract or on intact cells, the paramount concern for most peptidase biologists is the ability to identify specific targets that have been modified by a probe. Protein identification can be accomplished in any of a number of ways. The most common methods include isolation using a specific antibody or direct affinity purification of probe-bound peptidases followed by identification by mass spectrometry. The steps required to perform each of these identification methods are outlined (see Basic Protocol 5). The final basic protocol of this unit describes the use of ABPs in conjunction with small molecule inhibitors to determine potency and selectivity in intact cells and cell extracts (see Basic Protocol 6). This final application of the probe allows the simultaneous screening of compounds against several peptidase targets by making use of a simple competition assay in whole-cell homogenates. The assay requires initial optimization of inhibition kinetics and some knowledge of the mechanism of action of the compounds being assayed. It is likely to prove to be one of the most powerful applications of ABPs to peptidase biology. STRATEGIC PLANNING Choosing a Probe Activity-based probes come in many shapes and sizes and can be obtained from a number of different sources (Table 21.17.1). Over the past decade there has been a marked increase in the number of commercially available peptidase inhibitors. With this increase in the number of available compounds has come an increase in the number of general covalent inhibitors that contain biotin or fluorescent tags, allowing for their immediate use as ABPs. Still others are sold in a form that is amenable to labeling with tags such as radioactive iodine. Even with the ever-expanding list of commercially available reagents, some probes remain available only through academic laboratories that have authored publications on their use. Table 21.17.2 lists some of the major peptidase families that have been targeted with ABPs as well as relevant probes and their availability. The ABPs created to date have focused mainly on the serine/threonine and cysteine peptidase families, as these enzymes utilize a covalent acyl-enzyme intermediate. While some examples of covalent inhibitors of metallopeptidases and aspartyl peptidases do exist, none have been developed into a tagged form for labeling in complex protein samples. Table 21.17.2 Commercially Available Irreversible Inhibitors of Peptidases for Use as Specificity Controls
Peptidase family
Specific targets
Cysteine peptidases
Papain family Cathepsins/calpains Cathepsin B Caspases General and specific caspase inhibitors
Serine peptidases Applications for Chemical Probes of Proteolytic Activity
All hydrolyases
Available probes
Source
E-64 E-64d (cell permeable) CA-074 CA-074 Me (cell permeable)
Sigma Sigma Calbiochem Calbiochem
Wide variety similar to the tagged probes listed in Table 21.17.1; see Calbiochem catalog DFP PMSF AEBSF
Calbiochem
Sigma Sigma Sigma
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One of the most critical parameters to consider when selecting a probe is the type of tagging method used. For many of the peptidase families that have been targeted with ABPs, multiple probe families have been developed. Some of the issues to consider when choosing a tagging method are listed in Table 21.17.3. There are pros and cons to each method, and many people find that initial work with biotin-tagged reagents makes the most sense, as it has the greatest versatility and requires the least amount of special equipment. However, if sensitivity and background labeling are problematic, other methods can prove to be more optimal. The fluorescent reagents, while reducing the time required to analyze labeling patterns, require the use of a flatbed fluorescence scanner to create gel images and quantify labeling (see Alternate Protocol 3 for details). Several companies make these instruments, and they are becoming more commonly available at companies and universities. Radioactive probes often provide very clean labeling profiles with high sensitivity, but the use of radioactive materials makes handling and clean-up more difficult. Table 21.17.3 Considerations for Choosing a Tagging Method
Tagging method
Pros
Cons
Biotin tag
Does not require special equipment for analysis
Requires a two-step gel and blot protocol
Can be used to purify targets
Prevents labeling of peptidases inside an intact cell
Can be used for labeling of cell-surface peptidases on intact cells without contamination by intracellular peptidases
Poor solubility Presence of endogenously biotinylated proteins can obscure labeled peptidases Linear quantitation of labeled proteins is difficult Requires a good flatbed scanner for analysis of gels Fluorophores are hydrophobic and can have poor solubility
Fluorescent tag
HA-tag
Radioactive tag
Rapid, real-time scanning and quantitation of labeled peptidases in gel without processing or blotting Can be used to label intracellular peptidases in live cells Improved sensitivity over biotin
Can only be used to directly isolate targets if good antibody to fluorophore is available Starting materials can be expensive Excellent signal-to-noise ratio due Probes with this tag are not cell to availability of high-affinity permeable antibodies (e.g., 12CA5) Can be used to isolate and identify target proteins by mass spectrometry Requires work with radioactive Good signal-to-noise properties materials make analysis of data unambiguous Simple gel processing without blotting
Cannot be used to directly isolate targets
Improved sensitivity over biotin
Peptidases
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A second issue to consider if one has the option of multiple probes is the type of functional group used for covalent modification. As indicated by the long list of probes in Table 21.17.1, there are many different types of reactive “warheads” used in ABPs. In some cases, there are few choices, because the warhead directs the reactivity of a probe towards a specific peptidase family. For example, serine peptidase ABPs mainly utilize a fluorophosphonate warhead (Liu et al., 1999). There are some examples of chloromethyl ketone–based probes that covalently react with serine peptidases. However, these functional groups react with cysteine peptidases as well, and also show broad reactivity in crude cell extracts (Bogyo et al., unpub. observ.). For this reason, chloromethyl ketones are likely to be useful as probes only when working with recombinant peptidases where the sample complexity is limited. For the cysteine peptidase family, there are many probe options for each of the major target families. Extensive work on probes of papain family peptidases has created a host of reagents that can be used as general labels (see references in Table 21.17.1). Many of these reagents show different levels of selectivity towards members of this family, and the reader is advised to consult the literature to determine the best probe for a desired application. Probes for the caspase family have been used extensively over the past decade and currently there are many probes available commercially (see references in Table 21.17.1). However, it should be noted that while some of the probes are listed as specific for a single caspase, often the level of specificity is far from absolute. Furthermore, some of the probes work well in some assays and not in others, so it may be necessary to try multiple probes to find an optimal reagent. Probes based on large proteins with an epitope tag for detection at one end and a covalent warhead at the other have recently been designed to target deubiquitinating enzymes (DUBs; Borodovsky et al., 2002). These peptidases make contacts with multiple sites on the ubiquitin protein and therefore are not subject to labeling by small molecule probes. A clever strategy has been established to build protein-based probes of the enzyme class containing a range of reactive groups (see references in Table 21.17.1). These reagents require significant effort to prepare, but are very powerful tools for studying this family of peptidases (Borodovsky et al., 2001; Vugmeyster et al., 2002). Finally, several probe families have been designed to target the multi-catalytic proteasome (see references in Table 21.17.1). The first of these probes (NP-LLL-VS) was designed to be radioiodinated, and is cell permeable. Recent work to extend the peptide backbone has resulted in vinyl sulfone–based probes that are equally reactive towards each of the three primary active sites and contain a biotin tag (AdaLys(Bio)AhX3L3-VS). BASIC PROTOCOL 1
Applications for Chemical Probes of Proteolytic Activity
LABELING/TAGGING OF SMALL MOLECULE ACTIVITY-BASED REAGENTS USING THE IODOGEN METHOD This protocol describes the incorporation of a radioactive tag into small molecule activity-based reagents. This procedure can be used to convert a standard suicide inhibitor into an activity-based probe. The protocol outlines the incorporation of the radioactive isotope 125I into a small molecule probe; 125I labeling of a protein probe is described in Alternate Protocol 1. Labeling of probes with 125I using this protocol can be accomplished for any small molecule that contains a phenol group and for any protein that contains a tyrosine residue (UNIT 3.3). The following procedure discusses coupling of 125I using the Iodogen method. Other labeling approaches are feasible as well, although the authors have found this method to be the simplest and most effective.
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It should also be noted that this iodination protocol results in addition of multiple iodine atoms to a single tyrosine residue (mono- and di-iodo products are the most common). This is not a problem in most cases, unless the addition of a second iodine results in steric hindrance for binding to the target. Also, the addition of one or two iodines to the phenol ring leads to a change in the pK of the phenolic proton. In many cases, heterogeneous labeling can be resolved in an isoelectric focusing gel (i.e., multiple spots for a single peptidase in a 2-D gel due to mixtures of mono- and di-iodinated probes). For the most part, these issues should not affect simple 1-D gel analysis. If this is a problem, it is possible to purify the mono- and di-iodinated products individually while also removing noniodinated inhibitors using a standard C18 HPLC system. However this is an inconvenient procedure and should only be used if necessary. Materials Compound to be labeled 50% (v/v) ethanol Iodogen, Iodogen-coated beads, or Iodogen-coated microcentrifuge tubes (all available from Pierce) Chloroform 100 mCi/ml Na[125I] (3.7 GBq/mmol; Amersham Biosciences) 50 mM sodium phosphate, pH 7.4 (APPENDIX 2E) Acetonitrile C18 SepPak light gel cartridges (Waters) Column stand 10- and 30-ml Luer-Lok disposable syringes Lead pig γ counter and tubes CAUTION: This procedure involves handling a γ-emitting isotope and must be done with extreme care and in an assigned location with proper radiation safety equipment. Also see APPENDIX 2B. Be sure to use tips with filters in them to minimize contamination of the pipettor with volatile iodine. It is difficult to avoid this contamination, so it is desirable to have a dedicated set of pipettors for work with volatile radioactive iodine. Purification of the labeled probe also removes all volatile free iodine, so pipet tips without filters can be used with purified labeled probe. 1. Dilute compound to be labeled to 5 mM in 50% ethanol, for a final volume of 15 µl. This can be achieved by diluting a 50 mM DMSO stock solution of the compound to be labeled 1:10 with 50% ethanol. The solubility of the compound should be tested before performing the iodination. If the compound is soluble in 50% ethanol at 5 mM, this is advantageous, because the amount of DMSO present in the labeling reaction should be minimized. For efficient iodination, DMSO should be kept <15%.
2. Prepare a 1 mg/ml solution of Iodogen in chloroform and add 100 µl to a 1.5-ml microcentrifuge tube. Evaporate to dryness using a Speedvac evaporator or other rotary evaporation device. Alternatively, place a single Iodogen-coated bead in the bottom of a 1.5-ml microcentrifuge tube. Beads work as well as Iodogen-coated tubes, but are much more expensive in the long term. Also, if cost is not a issue, Pierce sells microcentrifuge tubes that have been precoated with Iodogen.
3. Add 35 µl of 50 mM sodium phosphate, pH 7.4 to the Iodogen-coated tube (or tube containing the Iodogen bead) followed by 10 µl (1 mCi) of Na[125I] and 12.5 µl of the 5 mM stock of compound to be labeled (from step 1). Carefully flick tube to mix contents and incubate for 15 min at room temperature. Peptidases
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It is possible to perform several iodinations simultaneously by starting reactions in a staggered format with 10 to 15 min between samples.
4. Place a SepPak C18 cartridge vertically in a column stand. Prepare column by wetting with 5 ml acetonitrile (dispensed using a full 10-ml disposable syringe) followed by equilibration with 5 ml of 5 mM sodium phosphate buffer pH 7.4. (dispensed using a full 30-ml disposable syringe; leave the syringe with the 25 ml of remaining phosphate buffer attached to the SepPak column to prevent drying of the column). Save the syringes with the remaining liquid inside for use in washing and elution. 5. At the end of the incubation in step 3, add 100 µl of 50 mM sodium phosphate buffer, pH 7.4, to the reaction mixture, remove the 30-ml buffer syringe from the column (see step 4), replace it with a 10-ml syringe that has its plunger removed, and load the reaction mixture into this 10-ml syringe (the reaction mixture will only fill the very tip of the syringe). 6. Gently push reaction mixture into the column using the plunger, being careful not to push air into the column. Remove the plunger and fill syringe with 10 ml of sodium phosphate buffer, pH 7.4. Replace plunger and rinse column with this buffer. Discard empty syringe in radioactive waste container. Wash column with the remainder of the 50 mM sodium phosphate buffer, pH 7.4, in the 30-ml syringe from step 4. 7. Elute iodinated compound with the 5 ml of acetonitrile remaining in the 10-ml syringe from step 4. Collect three 1-ml fractions into microcentrifuge tubes and store in a lead pig. Pipet 2 µl of each fraction and drop the tip with sample inside into a tube for counting using a γ counter. Most of the iodinated material should elute in the first two fractions. Iodination efficiency will typically be 30% to 50%, and one should expect to obtain anywhere from 5 × 105 to 2 × 106 cpm from 2 ìl of the peak fraction. The authors usually use the peak fraction (almost always the first fraction) and discard the others.
8. Use radiolabeled compound as probe. Once a labeled probe is obtained using this method, it can be added directly to biological samples without purification or concentration. The authors typically use 2 ìl of the peak fraction in 100 ìl of biological sample, although this may need to be optimized for the particular probe and sample. For discussion on how to optimize these conditions, see Basic Protocol 3. ALTERNATE PROTOCOL 1
Applications for Chemical Probes of Proteolytic Activity
125
I LABELING OF SMALL AMOUNTS OF A PROTEIN-BASED PROBE
This protocol has been used for the preparation of 10 to 40 µg of radiolabeled ubiquitinbased probes (Borodovsky et al., 2001). Materials Sephadex G-25 coarse resin (Amersham Biosciences) 0.2 mg/ml bovine serum albumin (BSA) in 50 mM sodium phosphate buffer, pH 7.5 (see APPENDIX 2E for buffer), 4°C Protein to be labeled 50 mM sodium phosphate buffer, pH 7.5 (APPENDIX 2E) 1 M Na2HPO4 1 M NaH2PO4 Iodogen-coated microcentrifuge tubes (see Basic Protocol 1, step 2) 100 mCi/ml Na[125I] (3.7 GBq/mmol; Amersham Biosciences) 0.4 mg/ml tyrosine 0.1 M NaI (unlabeled)
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Tabletop centrifuge with swinging-bucket rotor (e.g., IEC Clinical) 1-ml syringes Glass wool 10-ml round-bottom Falcon tubes CAUTION: This procedure involves handling a γ-emitting isotope and must be executed with extreme care and in a designated location with proper radiation safety equipment. Also see APPENDIX 2B. 1. Pre-swell 2 g Sephadex G-25 resin in 10 ml 0.2 mg/ml BSA and 50 mM sodium phosphate buffer, pH 7.5, overnight at 4°C or for 3 hr at room temperature. 2. Wash the resin 2 to 3 times with fresh 50 mM sodium phosphate buffer, pH 7.5, each time by centrifuging 1 min at 14,000 × g , 4°C, discarding the solution after each wash. Maintain the resin at 4°C. 3. Use a small amount of glass wool to plug a 1-ml syringe and fill with the prepared G-25 resin suspension to the 1-ml mark, using a Pasteur pipet. 4. Before use, place each column in a round-bottom 10-ml Falcon tube and centrifuge 2 min at 2000 rpm in a tabletop centrifuge with swinging bucket rotor. 5. Remove the lid of a 1.5-ml microcentrifuge tube and place the tube at the bottom of a new 10-ml round-bottomed Falcon tube as a collection vial. Place each column over one of these collection vials in a round-bottom Falcon tube prior to loading the sample. 6. Dilute the protein to be labeled with 50 mM sodium phosphate buffer, pH 7.5, to a final concentration of 10 µM, and check that the pH remains at 7 to 8 (adjust with 1 M Na2HPO4 or NaH2PO4 solution if necessary). Add the solution to a Iodogen-coated microcentrifuge tube. 7. Add 10 µl (1 mCi) of Na[125I], vortex vigorously, and incubate sample on ice for 30 min. 8. Quench the reaction by adding a 0.25 vol of 0.4 mg/ml tyrosine. Incubate sample 1 min at room temperature. 9. Add 8 µl of 0.1 M NaI to the sample. Add 50 mg/ml lysozyme stock to a final concentration of 1 mg/ml lysosyme, as a carrier protein. If more than 100 ìg of protein was used for the iodination reaction, it is not necessary to add lysozyme as a carrier protein.
10. Load 94 µl of the protein radiolabeling reaction from step 9 per G-25 column. Centrifuge each column 2 min at 2000 rpm in tabletop centrifuge with swingingbucket rotor. 11. Discard columns and vials and count 2 µl of the sample (recovered in the microcentrifuge tubes under the columns) in a γ counter. About 100 ìl of sample should be recovered from each column. Iodination efficiency will typically be 30% to 50%. and one should expect 5 × 105 to 2× 106 cpm from 2 ìl of the collected fraction. The radiolabeled protein probes can be divided into aliquots and can be stored up to 1 month at −80°C.
Peptidases
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ALTERNATE PROTOCOL 2
TAGGING SMALL MOLECULES WITH BIOTIN OR FLUOROPHORE This protocol describes labeling of small-molecule probes with a tag such as biotin or a fluorophore. This is carried out using deprotonated primary amine groups on a probe, and utilizes tags containing a succinimidyl ester (e.g., N-hydroxysuccinimide; NHS), which are available from several commercial sources. Coupling can be accomplished under either aqueous or organic conditions depending on the molecule to be labeled. In most cases, when labeling small molecules, organic solvents are preferred. It should be noted that this procedure is not significantly different from any NHS-ester/primary amine coupling protocol supplied by the manufacturer of the reagents. However, caution should be taken with prolonged incubation times, as in some cases this can lead to decomposition of the electrophilic probe. The method described here utilizes biotin (UNIT 3.6) and BODIPY fluorophore (Molecular Probes) tags, however, any NHS ester-containing tag could be used. Coupling times and solvents may need to be optimized based on type of NHS ester tag used. Materials Amine-containing reagent DMSO Diisopropylethylamine (DIEA) NHS ester-containing tag (e.g., biotin or BODIPY fluorophore) C18 column (UNIT 8.7) 50% (v/v) acetonitrile in H2O Liquid nitrogen 50-ml polypropylene tubes Wide-gauge needle Additional reagents and equipment for HPLC (UNIT 8.7) 1. In a 1.5-ml microcentrifuge tube, prepare a 10 µM stock solution of amine-containing reagent in 100 µl of DMSO. Add 2 equivalents of DIEA and mix well. 2. Prepare a 9 µM solution of NHS ester-containing tag in 100 µl of DMSO. 3. Add predissolved tag solution to the amine-containing reagent mixture. Flick tube to mix its contents and incubate 2 hr at room temperature with mild agitation. Using the amine-containing reagent in a slight excess will increase coupling efficiency and make subsequent HPLC purification easier. The reaction can be monitored by analytical HPLC (see step 4). In many cases the reaction does not proceed to completion and extended incubation times do not improve yield; therefore, reactions should be monitored early and stopped after 2 hr. At the end of the incubation, the reaction mixture can be analyzed and purified by HPLC, or, if this step cannot be accomplished on the same day, residual active esters must be quenched by adding ethanolamine (adjusted to pH 7.5 with HCl) to a final concentration of 50 ìM for an additional 30 min. The reaction mixture can then be kept frozen at −20°C until the purification step is performed.
4. Dilute 2 µl of reaction mixture into 10 µl of 50% acetonitrile and analyze by reversed-phase HPLC using a C18 column (UNIT 8.7).
Applications for Chemical Probes of Proteolytic Activity
In most cases, the tag to be attached is more hydrophobic than the reagent to be tagged. Thus, the free amine-containing reagent will elute before the tagged reagent, and unbound succinimidyl ester-containing tag will elute later than the tagged reagent. Obtaining elution profiles of the starting material beforehand is useful when analyzing the reaction HPLC profile.
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5. Purify reaction mixture by reversed-phase HPLC and pool probe-containing elution fractions in a 50-ml polypropylene tube. The analytical HPLC should provide a map of the general reaction profile that can be used to determine which peak is the desired product.
6. Puncture several holes in the cap of the tube containing the pooled fractions using a wide-gauge needle. Freeze pooled fractions in liquid nitrogen and lyophilize to obtain a dry powder. Redissolve the lyophilized power in 500 µl to 1 ml of 50% acetonitrile and transfer to a weighed microcentrifuge tube. Dry down on lyophilizer or Speedvac evaporator, then obtain weight. DETECTION OF LABELED PROBES The following protocols present a variety of methods for detecting probe-modified proteins. This set of protocols can be used for any proteome, whether it is a mixture of recombinant proteins, a lysate, or intact cells. However, labeling protocols should be optimized for the proteome and the type of probe being used. A systematic way to optimize the labeling protocol is described in Basic Protocol 3. Four procedures are described here: (1) detection of biotinylated probes via immunoblotting (see Alternate Protocol 4); (2) detection of hemagglutinin antigen (HA)–tagged protein probes via immunoblotting (see Alternate Protocol 5); (3) detection of radioactive probes on autoradiograms (see Basic Protocol 3); and (4) detection of fluorescent-tagged probes via fluorescent scanning (see Alternate Protocol 3). All of these procedures require prior separation of the probe-labeled protein sample by SDS-PAGE (UNIT 10.1). While detection of radioactive and fluorescent probes can be performed directly in the polyacrylamide gel, detection of biotinylated or HA-tagged probe requires a blotting step and is therefore more time-consuming. Detection of Radiolabeled Probes by Autoradiography
BASIC PROTOCOL 2
Materials Sample: probe-modified protein mixture of interest Fixation solution (see recipe) 50% (v/v) methanol Whatman 3MM filter paper PhosphorImager (Amersham Biosciences; optional) Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and autoradiography (UNIT 10.11) 1. Separate samples by SDS-PAGE (UNIT 10.1) using a gel with a polyacrylamide concentration that will yield the best separation of the proteins of interest. The desired percentage of acrylamide in the separating gel depends on the molecular size of the protein being separated. Generally, use 5% gels for SDS-denatured proteins of 60 to 200 kDa, 10% gels for SDS-denatured proteins of 16 to 70 kDa, and 15% gels for SDS-denatured proteins of 12 to 45 kDa (Table 10.1.1). Note that the free probe migrates at the dye front of the gel-resolved samples. The bromphenol blue migration line is usually a good indicator of the location of the free probe on the gel. Running samples so that the free probe migrates out of the gel will produce a better signal-to-noise ratio in the subsequent detection step. Care should be taken when discarding the gel running buffer because it contains a substantial amount of radioactive material. Peptidases
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2. Carefully place gel in a dish containing fixation solution. Fix for 1 hr at room temperature with gentle shaking. Rinse gel with 50% methanol for 10 min, then with distilled water for 10 min. Place fixed gel on a 3MM filter paper and dry using a gel dryer. Gels can be stained prior to drying with various protein staining reagents such as Coomassie blue or silver stain (UNIT 10.5).
3. Expose dried gel to film (autoradiography; UNIT 10.11) or to a PhosphorImager screen. Typical exposure times for freshly iodinated probes will range from 1 hr to overnight. In some cases longer exposures will be necessary; however, if exposure times are too long it may be necessary to optimize the labeling protocol (see Basic Protocol 3). ALTERNATE PROTOCOL 3
Detecting Fluorescently Labeled Probes Additional Materials (also see Basic Protocol 2) High-intensity flat-bed fluorescent scanner (e.g., Typhoon; Molecular Dynamics) 1. Follow the same electrophoresis procedure as described for use with radioiodinated probes (see Basic Protocol 2, step 1). 2. When electrophoretic separation is complete place the gel in double-distilled water for several minutes. Some fluorescent scanners allow analysis of gels while still in the electrophoresis plates. This requires that the gel plates have low fluorescence emission and absorption characteristics, so most plastic plates will not work for this application. The benefit of that method is that it is possible to monitor resolution of labeled proteins during a run. After scanning, the gel can be returned to the gel chamber and proteins can be further separated if the desired resolution has not yet been achieved.
3. Scan gel using a fluorescence scanner. There are currently several commercially available fluorescence scanners. The use of fluorescent probes requires a high-intensity flat-bed laser scanner. It is not possible to use simple UV light boxes and CCD cameras, as the intensity of signal is not strong enough and quantification is not reliable. The authors use the Molecular Dynamics Typhoon scanner, which is available with several different excitation lasers and can be equipped with various filters for emission detection. These scanners are also useful for detection of blots using fluorescent secondary reagents, and also can be used to scan radioactive gels using a phosphor screen. Alternatively, scanners are available from Hitachi and Bio-Rad that contain lasers capable of sensitive detection of fluorescent probes.
4. If the background signal is high or proteins of interest are co-migrating with free probe, incubate the gel in fixation solution for 2 hr and perform several washes with 50% methanol to reduce the background signal. ALTERNATE PROTOCOL 4
Detecting Biotinylated Probes Materials Sample: biotin-modified protein mixture of interest Immunoblot transfer buffer (see recipe) Immunoblot blocking solution (see recipe) Vectastain Elite Kit (Vector Laboratories) PBST: PBS (APPENDIX 2E) containing 0.1% (v/v) Tween 20 Chemiluminescent HRP substrate (e.g., ECL, Pierce)
Applications for Chemical Probes of Proteolytic Activity
Additional reagents and equipment for SDS-PAGE (UNIT 10.1) and protein transfer to PVDF or nitrocellulose membranes (UNIT 10.7)
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1. Separate labeled sample using SDS-PAGE (UNIT 10.1) and transfer the protein to a PVDF or nitrocellulose membrane (UNIT 10.7), using immunoblot transfer buffer (see Reagents and Solutions) in place of the transfer buffer in UNIT 10.7. For most applications nitrocellulose works well and is less expensive than PVDF.
2. Block membrane by incubating with immunoblot blocking solution for 1 hr at room temperature with gentle agitation. It is possible to use a milk-based blocking solution; however, since milk contains biotin, the membrane must be washed thoroughly with PBST before addition of avidin-HRP conjugates.
3. During the incubation in step 2, mix 2 drops of A and B Vectastain components (from Vectastain Elite Kit) in 10 ml of PBST and incubate 30 min according to the manufacturer’s protocol. The VectaStain reagent is made up of two components, avidin and biotinylated HRP. Mixture of the two reagents results in conjugates that contain an open biotin binding site and two to three HRP molecules bound via the remaining biotin-binding sites. This multimeric complex produces superior signal in immunoblots. There are various other commercially available streptavidin-HRP reagents that can be used in this protocol. However, these reagents are made up of monomeric HRP-avidin conjugates that provide poorer signal-to-noise ratios.
3. Wash membrane three times, each time for 5 min with PBST. 4. Incubate membrane with the prepared Vectastain reagent from step 2 for 1 hr at room temperature. 5. Wash membrane at least three times, each time for 15 min, with PBST. 6. Incubate membrane with appropriate volume of chemiluminescent HRP substrate using manufacturer instructions and expose to film. Detecting HA-Tagged Protein-Based Probes Using Anti-HA Immunoblot The protein-based probes described here are not cell-permeable and can be used only to target active deubiquitinating enzymes (DUBs) that are either pure or present in cell extracts. For this purpose, cells are usually lysed using methodologies that do not involve detergents (glass beads, several cycles of freeze and thaw, or French press) in order to minimize interference with enzyme activity. More recent studies reveal that most DUBs retain their activity in detergent lysis protocols employing 0.1% NP-40. The amounts of probe and extracts used for labeling experiments need to be optimized depending on the nature of the biological sample. Usually, extracts prepared from cell lines contain a larger number of active DUBs as compared to extracts derived from tissue samples. The incubation times are also sample-dependent, but optimal labeling is typically observed after one hour at 37°C. HA-tagged probes can also be used to isolate and identify target peptidases by immunoprecipitation followed by analysis using mass spectrometry-based methods (see Basic Protocol 5 and Borodovsky, 2002).
ALTERNATE PROTOCOL 5
In order to have an optimal separation of labeled DUBs using ubiquitin-based probes, 10% or gradient SDS-PAGE gels should be used (UNIT 10.1). Precast gradient minigels provide excellent resolution, but larger gels such as those provided by Bio-Rad or other vendors can significantly increase resolution. In this protocol, a Bio-Rad wet transfer apparatus is used. Blotting protocols are identical to standard immunoblotting protocols described in UNITS 10.7, 10.8, and 10.10. Peptidases
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BASIC PROTOCOL 3
OPTIMIZING IN VITRO LABELING OF PEPTIDASES BY ACTIVITY-BASED PROBES This protocol describes the conditions that need to be optimized to obtain robust and reproducible labeling of peptidases by ABPs in vitro. A basic understanding of enzyme kinetics and protein biochemistry is assumed. This protocol is designed to help researchers achieve the maximum amount of specific peptidase labeling while minimizing nonspecific background labeling for the purpose of visualizing and identifying active peptidases in a solution. As such, this protocol can be used to assess the total amount of active peptidases in a solution but not to assess their specific activities. See Basic Protocol 6 for details on how to assess specific peptidase activity using ABPs. The conditions that need to be established are shown in Table 21.17.4. Since the ABPs described in this unit all behave as suicide substrates, it is natural to think of the reaction between the ABP and the enzyme of interest as an enzyme-substrate interaction. The parameters in Table 21.17.4 are basic conditions that enzymologists test when trying to optimize the activity of an enzyme with a specific substrate. A reasonable range for the different parameters is given as a starting point for the optimization. At this point it is assumed that an ABP and source of enzyme (purified or crude) has been chosen and all that remains is to mix the two and analyze the results on an SDS-PAGE gel (see Basic Protocol 2). The basic protocol outlined below starts with the optimization of probe concentration and time of labeling. Once that is completed, different temperatures and buffer conditions can be tested to improve specific labeling. Materials Solution of peptidase (purified or crude) 10 mM stock of ABP in 100% DMSO Specificity control (see Critical Parameters) Tris hydrochloride and Tris base EDTA Triton X-100 1 M dithiothreitol (DTT) 10 mM stock of specific inhibitor in 100% DMSO 4× SDS-PAGE sample buffer (UNIT 10.1) Silanized microcentrifuge tubes (National Scientific; APPENDIX 3E) 80°C heating block or water bath 96-well polystyrene plates Additional reagents and equipment for SDS-PAGE analysis of tagged ABPs (see Basic Protocol 2 or Alternate Protocol 3, 4, or 5) 1. Thaw the enzyme solution on ice and the stock of ABP and specificity control at room temperature or 37°C. The enzyme and ABP are more stable if stored at −80°C and −20°C respectively. 100% DMSO stocks will not thaw on ice. It is very important to have the correct specificity control in place before doing any optimization. The probes described in this unit contain active electrophiles and are capable of reacting nonspecifically with proteins that contain strong nucleophiles. See Critical Parameters for more discussion.
Applications for Chemical Probes of Proteolytic Activity
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Table 21.17.4
Optimization Parameters
Parameter
Range
Rationale
ABP concentration
Biotinylated probe: 1-50 µM Fluorescent probe: 0.01-5 µM Iodinated probe: 1–5 × 106 total cpm
The concentration of ABP used will depend primarily on the tag that is being used. Iodinated tags can be used in very small quantities because of the sensitivity of detection. Fluorescent probes can be used at low nanomolar concentrations because of their sensitivity and should not be used at high concentrations due to high background. Biotinylated probes generally give less background but are harder to detect, so they should be used at higher concentrations. For purified enzyme the Changing the concentration of enzyme most important issue is level of detection since affects the amount of nonspecific binding should enzyme bound by the ABP. There also needs to not be an issue. For crude mixtures, it is more be enough enzyme important to keep the present to be able to detect it without having concentration of non-target enzymes/proteins low, so as to use so much as to to minimize background. create nonspecific interactions between the ABP and the enzyme or other proteins. Increasing the amount of Labeling time is most important to optimize with time can increase crude mixtures. Labeling for background in some too long can increase cases. With highly specific probes, labeling nonspecific labeling and lead to protein degradation increases with time without large increase in by other peptidases in the background labeling. In mixture. all cases, labeling will saturate at some prolonged incubation time depending on the kinetics of labeling. Most enzymes have evolved Temperature has a to work best at 37°C, but dramatic effect on enzyme activity and will labeling can occur quite robustly at room therefore affect the amount of labeling by the temperature. High temperatures can cause ABP protein denaturation.
Enzyme/protein Purified enzyme: concentration 0.01-1 µM Crude mixture: 0.1-5 mg/ml
Time
10 min to 3 hr
Temperature
20°C to 37°C
Comments
Changing the concentration of the ABP changes the amount of ABP specifically bound to enzyme and nonspecifically bound to enzyme and other proteins. Concentration can have a dramatic effect on signal-to-noise ratios.
continued Peptidases
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Table 21.17.4
Optimization Parameters, continued
Parameter
Range
Rationale
pH
Lysosomal cysteine peptidases: pH 4-6 Other peptidases: pH 6.5-8.5
Ionic strength
0-150 mM NaCl or KCl
Reducing agent 0-5 mM DTT or 2-mercaptoethanol
Detergent
Specificity controls
0.01-1.0% (v/v) nonionic detergent (NP-40 or Triton X-100) or zwitterionic detergent (CHAPS) 10-100 fold excess of an irreversible active-site inhibitor of the enzyme, or preheating of enzyme to 80°C
Comments
Other than the lysosomal cysteine peptidases that prefer pH 4-6, most enzymes work best around a physiological pH of 7.4. Choose a buffering system that works well in the range needed. It is not recommended to go Ionic strength can sometimes effect enzyme above physiological concentrations of salt, i.e., activity and will >150 mM therefore affect the amount of labeling by the ABP Some enzymes are Cysteine peptidases are adversely affected by particularly susceptible to oxidation of cysteine side loss of activity by oxidation chains Some cysteine peptidases of Detergents can sometimes effect enzyme the cathepsin family are sensitive to nonionic activity and will detergents therefore affect the amount of labeling by the ABP The best control is a potent, It is critical to know specific, irreversible which covalent modifications are due to active-site inhibitor such as those listed in Table modification of the active site of the enzyme 21.17.2. In the absence of target and which are due this, preheating should be used. to nonspecific nucleophiles present in the reaction mixture pH can have a dramatic effect on enzyme activity and will therefore affect the amount of labeling by the ABP
2. Prepare 1 ml of reaction buffer at room temperature; a typical reaction buffer for use with probes in vitro is as follows: 50 mM Tris⋅Cl, pH 7.5 1 mM EDTA, pH 8.0 0.01% Triton X-100 2.5 mM DTT. If necessary, this reaction buffer can be stored at 4°C up to 6 months.
3. Dilute enzyme in reaction buffer to 1 mg/ml for a crude lysate or 0.1 µM for a purified enzyme. Prepare 500 µl of diluted enzyme.
Applications for Chemical Probes of Proteolytic Activity
4. Aliquot 250 µl of diluted enzyme into each of two silanized microcentrifuge tubes. Add 1.25 µl of specific inhibitor to one tube (50 µM final concentration) and 1.25 µl of 100% DMSO to the other. Allow reaction to proceed for 30 min at room temperature. Alternatively, heat one aliquot of enzyme to 80°C for 10 min, then microcentrifuge 5 min at maximum speed, room temperature, and pipet the supernatant into a new tube.
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This step is used to ensure the specificity of any labeling observed with the ABP. An irreversible active site inhibitor of the target enzyme will prevent the subsequent modification of the enzyme by the ABP. Heating to 80°C will denature the enzyme, leading to loss of activity and the ability to specifically interact with the ABP.
5. Prepare 5-fold serial dilutions of the ABP in 100% DMSO (final concentrations of 500, 100, 20, and 4 µM). 10 ìl of each dilution is sufficient.
6. Divide each of the two enzyme solutions from step 4 (pretreated with inhibitor/heat and untreated) into four 60-µl aliquots. Add 0.6 µl of each of the ABP dilutions prepared in step 5 to a corresponding aliquot of each of the two enzyme solutions (to yield ABP final concentrations of 5, 1, 0.2, and 0.04 µM). 7. At 10, 30, 90, and 180 min, remove 14 µl from each tube and add it to a corresponding well of a 96-well plate containing 5 µl of 4× SDS-PAGE sample buffer and 1 µl of 1M DTT. 8. After the last time point, heat plate to 80°C for 5 min and load all of the samples on an SDS-PAGE gel (see Basic Protocol 2 or Alternate Protocol 3, 4, or 5). Analysis of the results will indicate which conditions should be used for further labeling experiments. Altering the other parameters in Table 21.17.4 can significantly increase the amount of labeling obtained. It is straightforward to incorporate some of the other buffer conditions or changes in temperature into the protocol described above. See Troubleshooting for more details on data analysis.
IN SITU LABELING OF LYSOSOMAL CYSTEINE PEPTIDASES WITH ACTIVITY-BASED PROBES Basic Protocol 4 and Alternate Protocols 6 and 7 discuss the use of ABPs for detecting enzyme activity within intact cells. In many ways this is simpler than the in vitro method (Basic Protocol 3), since there is no need to optimize the labeling buffer system. The main consideration for these experiments is probe permeability. The probe should be compatible with the cellular spatial distribution of the peptidase(s) of interest. For example, when labeling lysosomal cathepsins, it is necessary to use a membrane-permeable probe. However, if the enzyme active site is located in the extracellular compartment, as with many serine hydrolases, a non-cell-permeable probe can be used. Another consideration is the time of labeling. In many cases, it is advantageous to perform rapid labeling to monitor enzymatic activity in a specific cellular time window or when conducting competition experiments (see Basic Protocol 6) using reversible inhibitors. In such cases, the kinetic properties of the probe, in particularly KD and kinact, should be compatible with the overall experimental design. The presence or absence of serum during labeling should also be taken into consideration. Firstly, some probes will bind proteins in the serum such as albumin through hydrophobic interactions, thereby preventing their access to the cells. Secondly, the commonly used fetal bovine serum contains serpins that can irreversibly inactivate cell surface serine peptidases. Finally, it is difficult to analyze activity of secreted peptidases in the presence of serum. In some cases, if the presence of serum is required, it is possible to use biochemically defined serum-free medium supplement that can be obtained from commercial vendors. This protocol focuses on in situ labeling of lysosomal cysteine peptidases. NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. NOTE: All reagents and equipment coming into contact with live cells must be sterile, and aseptic technique should be used accordingly. Peptidases
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BASIC PROTOCOL 4
Labeling of Lysosomal Cysteine Peptidases in Intact Cells Using 125I-Labeled Activity-Based Probes Materials Cells Serum-containing medium appropriate for cells Radiolabeled activity-based probe (e.g., [125I ]JPM-OEt, a cell-permeable probe) Inhibitor: e.g., 10 mM E-64d (Calbiochem) in 100% DMSO Phosphate-buffered saline (PBS; APPENDIX 2E) Cell lysis buffer (see recipe) 12-well tissue culture plate Rubber policeman Additional reagents and equipment for SDS-PAGE (UNIT 10.1), detemination of protein concentration (UNIT 3.4), and detection of radiolabeled probes (see Basic Protocol 2) 1. Plate sufficient cells in wells of a 12-well dish in 1 ml of serum-containing medium a day prior to the scheduled labeling experiments to achieve 70% to 80% confluency at time of experiment. 2. When cells have reached 70% to 80% confluency, add relevant inhibitor to control wells and incubate 1 hr. Add probe(s) to a final concentration of 106 cpm/ml medium and continue to incubate for an additional 1 hr. Design of a typical in situ labeling experiment in a 12-well dish is described in Table 21.17.5.
3. Transfer cells to an ice-bed and discard medium in a proper safety container. Wash cells with 2 ml of cold PBS per well. 4. Lyse cells by adding 200 µl of cell lysis buffer directly to wells, and dislodge cells using a rubber policeman. Transfer lysed cells to a microcentrifuge tube and microcentrifuge 10 min at maximum speed, 4°C. 5. Determine protein concentration (UNIT 3.4) of supernatant. Analyze labeled proteins by SDS-PAGE (UNIT 10.1) and continue with detection as specified in Basic Protocol 2. The proposed lysis buffer for this protocol is also compatible with 2-D gel separation (UNIT 10.4). Following lysis, the protein solution can be diluted in an appropriate isoelectric focusing sample buffer and resolved in the first dimension using an ampholyte pH gradient or commercially available IPG strips. ALTERNATE PROTOCOL 6
Labeling of Lysosomal Cysteine Peptidases in Intact Cells Using Fluorophore-Labeled Activity-Based Probe This protocol is almost identical to the one using a radiolabeled ABP (see Basic Protocol 4). The main difference is that the labeling is accomplished in the absence of serum since the probe used here binds to serum proteins. Additional Materials (also see Basic Protocol 4) Serum-free medium appropriate for cells Fluorescently labeled activity based probe (e.g., BODIPY-DCG-04, a cell-permeable probe)
Applications for Chemical Probes of Proteolytic Activity
Additional reagents and equipment for detection of fluorescently labeled probes (see Alternate Protocol 3)
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Table 21.17.5 Example of Experimental Design for in situ Labeling
No. of cells
Medium Inhibitor volume volume
2 × 105
1 ml
Pretreatment 2 × 105 with inhibitor
1 ml
Experiment Probe labeling
None: use 2 µl of pure DMSO as mock treatment 2 µl of 500× inhibitor/DMSO stock
Probe volume
Preincubation time/temp. with inhibitor
Incubation time/temp. with probe
2 µl of 500× probe 1 hr at 37°C stock solution
1 hr at 37°C
2 µl of 500× probe 1 hr at 37°C stock
1 hr at 37°C
1. Plate sufficient cells in wells of a 12-well dish in 1 ml of serum-containing medium a day prior to the scheduled labeling experiments to achieve 70% to 80% confluency at time of experiment. 2. When cells have reached 70% to 80% confluency, wash cells twice, each time with 2 ml of serum-free medium per well, and add 1 ml of serum-free medium per well. 3. Add relevant inhibitor to control wells and incubate 1 hr. Add probe to a final concentration of 1 µM to cell medium and continue to incubate for an additional 2 hr. 4. Transfer cells to an ice-bed and discard medium in a proper safety container. Wash cells with 2 ml of cold PBS per well. 5. Lyse cells by adding 200 µl of cell lysis buffer directly to wells, and dislodge cells using a rubber policeman. Transfer lysed cells to a microcentrifuge tube and microcentrifuge 10 min at maximum speed, 4°C. 6. Determine protein concentration (UNIT 3.4). Analyze labeled proteins by SDS-PAGE (UNIT 10.1) and continue with detection as specified in Alternate Protocol 3. Labeling of Lysosomal Cysteine Peptidases in Intact Cells Using Biotinylated Activity-Based Probes
ALTERNATE PROTOCOL 7
This protocol is designed to label extracellular peptidases since the biotinylated versions of most ABPs are non-cell-permeable. The labeling can be done in the presence or absence of serum. Conditioned medium from labeled cells can also be analyzed for detection of secreted hydrolases. In this case, a serum-free medium should be used. Medium can be directly analyzed on an SDS-PAGE gel or concentrated first using common protein extraction methods such as ammonium sulfate precipitation or acetone precipitation (both described in UNIT 4.5). Additional Materials (also see Basic Protocol 4) Biotinylated activity-based probe (e.g., biotin-DCG-04, a non-cell-permeable probe) Additional reagents and equipment for detection of biotinylated probes (see Alternate Protocol 4) 1. Plate sufficient cells in wells of a 12-well dish in 1 ml of serum-containing medium a day prior to the scheduled labeling experiments to achieve 70% to 80% confluency at time of experiment. Peptidases
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2. When cells have reached 70% to 80% confluency, wash cells twice with 2 ml of serum-free medium per well and add 1 ml of serum-free medium per well. 3. Add relevant inhibitor to control wells and incubate 1 hr. Add probe to a final concentration of 1 µM to cell medium and continue to incubate for an additional 2 hr. 4. Transfer cells to an ice-bed and discard medium in a proper safety container. Wash cells with 2 ml of cold PBS per well. 5. Lyse cells by adding 200 µl of cell lysis buffer directly to wells and dislodge cells using a rubber policeman. Transfer lysed cells to a microcentrifuge tube and microcentrifuge 10 min at maximum speed, 4°C. 6. Determine protein concentration (UNIT 3.4). Analyze labeled proteins by SDS-PAGE (UNIT 10.1) and continue detection as specified in Alternate Protocol 4. BASIC PROTOCOL 5
STRATEGIES FOR IDENTIFICATION OF ACTIVE PEPTIDASE BY IMMUNOPRECIPITATION To identify a labeled peptidase of interest, specific antibodies against known peptidases can be used in an immunoprecipitation (IP) experiment (see Harlow and Lane, 1988, for an excellent reference on antibodies; also see UNIT 9.8). This method works well when using an ABP that targets a specific family of peptidases, for example the papain family cysteine peptidase probe DCG-04 (Greenbaum et al., 2000). The identity of an unknown cysteine peptidase can be established using a panel of commercially available antibodies to the known lysosomal cathepsin peptidases. This method does not work for ABPs that target larger families of enzymes, e.g., the serine hydrolase probes containing fluorophosphonate warheads, because some of the labeled enzymes may not yet be characterized (Liu et al., 1999). A useful application of immunoprecipitation is to isolate an enzyme of interest from a mixture of proteins that are labeled by the ABP. If there are a large number of specifically or nonspecifically labeled proteins in the mixture, the labeled enzyme of interest could be obscured after SDS-PAGE analysis. Immunoprecipitation of the desired target can be used to purify it from the other labeled proteins. There are many sources of antibodies that can be used, including polyclonal rabbit serum, purified polyclonal antibodies, monoclonal ascites fluid or tissue culture fluid, and purified monoclonal antibodies. Researchers are encouraged to consult Harlow and Lane (1988) for more details on the immunoprecipitation protocols available. Two protocols are provided here for immunoprecipitation under native or denaturing conditions. Some antibodies work better under one or the other condition, although the native immunoprecipitation protocol is the more common. Strategies for identification of active enzymes by purification and mass spectrometry The most precise way to identify a labeled enzyme of interest is to first purify it and then identify it by mass spectrometry (for review see Rappsilber and Mann, 2002). In this experiment, proteins are purified by affinity chromatography (Chapter 9) or immunoprecipitation (UNIT 9.8) and separated by SDS-PAGE (UNIT 10.1). Proteins in the gel are stained with silver or Coomassie blue (UNIT 10.5), excised from the gel, and proteolyzed, most commonly with trypsin (UNIT 11.3). These tryptic peptide fragments are then identified by mass spectrometry (MS) or coupled HPLC/mass spectrometry (LC/MS; Chapter 16). This method can provide unambiguous identification of the labeled enzyme.
Applications for Chemical Probes of Proteolytic Activity
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The labeled enzymes are purified most easily using biotinylated ABPs and streptavidinagarose affinity chromatography (Wilchek and Bayer, 1990), but they can also be purified by immunoprecipitation with antibodies directed against the fluorescent tags often used in ABPs (available from the manufacturer). The strepavidin-based enzyme purification is a straightforward biochemical procedure that is primarily limited by the amount of starting material available and the number of nonspecific proteins that copurify with the labeled enzyme of interest. In general, this purification and MS identification method requires 2 to 4 orders of magnitude more starting material than either 2-D gel analysis or immunoprecipitation. The purity of the labeled enzymes obtained can be quite high using streptavidin-agarose affinity chromatography, but endogenously biotinylated proteins and avidin itself can sometimes interfere with visualization by SDS-PAGE and identification of labeled enzymes by MS. In an immunoprecipitation experiment, the presence of large amounts of IgG can contaminate the samples used for subsequent identification by MS. Researchers are encouraged to read Harlow and Lane (1988) for strategies to decrease the amount of antibody copurifying with the enzyme of interest. Few laboratories have the combined skills, money, and equipment required to identify purified enzymes using MS, although this is changing as mass spectrometers become less expensive and more user friendly. Some laboratories will collaborate to help identify proteins by MS; another option is simply to pay someone to do the identification. Many large universities are establishing proteomics core facilities that should be able to provide this service as well. All of these MS options are expensive in different ways, and identification by MS should only be carried out when it is absolutely essential to identify a labeled enzyme. No matter which option is chosen, it is imperative to speak with an expert in protein identification by MS, as it requires specialized equipment, protocols, and expertise to be successful. It is critical that the final preparation of the enzyme to be identified be compatible with subsequent MS analysis methods. Affinity purification of biotinylated enzymes by immunoprecipitation is described in Basic Protocol 6. This purification can be performed under native or denaturing conditions and both options are detailed. A list of key references is provided at the end of this unit for the subsequent staining, digestion, and MS analysis of tryptic peptides. Materials Biotinylated enzyme Specific antibody and control antibody IP wash buffer (see recipe) Protein A or protein G coupled to agarose beads (Amersham Biosciences) 10% (w/v) SDS (APPENDIX 2E) RIPA dilution buffer (see recipe) RIPA buffer (see recipe) End-over-end rotator, 4°C 80°C water bath Additional reagents and equipment for labeling enzymes (see Basic Protocol 3) and SDS-PAGE (UNIT 10.1; also see Basic Protocol 2) To immunoprecipitate labeled enzymes under native conditions 1a. Perform a labeling reaction as outlined in Basic Protocol 3. The amount of starting material will depend on the intensity of the signal from the label on the enzyme of interest. Generally speaking, robust labeling can be achieved with <100 ìg of crude lysate no matter what detection method is used. Peptidases
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2a. Quench the labeling reaction (see Basic Protocol 3) and add an equal volume of ice-cold IP wash buffer. Cool the tube on ice for 10 min. The reaction can be quenched by adding an excess of an irreversible active site inhibitor.
3a. Add the antibody as undiluted serum to the tube on ice. The amount of antibody to add will depend on the source of the antibody and the concentration of the antibody in that solution. It may be necessary to perform a pilot experiment titrating the amount of antibody to determine how much is needed for robust and specific immunoprecipitation. It is also critical to include the correct negative control antibody.
4a. Allow the antibody-enzyme complex to form on ice for 60 min. Microcentrifuge the solution 5 min at maximum speed, room temperature, to pellet any debris. Transfer supernatant to a new tube. Incubating for longer than 60 min will not significantly increase the amount of complex formed.
5a. Add 10 to 25 µl bead volume of protein A–or protein G–agarose beads prewashed in cold IP wash buffer according to the manufacturer’s instructions. Put the tube on an end-over-end rotator at 4°C and allow the protein A– or G–antibody complex to form for 60 min. The amount of protein A or G agarose beads added will depend on the amount of antibody added. Again, incubating for longer than 60 min will not significantly increase the amount of complex formed.
6a. Microcentrifuge sample 10 sec at 1000 × g, 4°C to pellet the beads, and discard supernatant. 7a. Wash pellet three times, each time with 1 ml of IP wash buffer at 4°C, using the centrifugation conditions in step 6a. 8a. Remove all the liquid using a pipet equipped with a gel-loading pipet tip inserted all the way into the beads at the bottom of the tube. Any beads that stick to the tip can be rubbed off on the inside of the tube.
9a. Add 20 to 40 µl of 1× SDS-PAGE sample buffer, heat tube to 80°C for 5 min, and load sample on SDS-PAGE gel (UNIT 10.1; see Basic Protocol 2 for conditions). To immunoprecipitate labeled enzymes under denaturing conditions 1b. Perform a labeling reaction as outlined in Basic Protocol 3. 2b. To quench the reaction, add 1/10 volume of 10% SDS and heat the tube at 80°C for 10 min. 3b. Microcentrifuge briefly to bring liquid to bottom of tube and add 9 vol of RIPA dilution buffer. 4b. Cool the tube on ice for 10 min. 5b. Add antibodies and protein A or G beads and perform incubations exactly as described in the native IP protocol (steps 3a to 5a), except prewash the beads in RIPA buffer before adding them to the tube. 6b. After forming the protein complexes, pellet the beads to the bottom of the tube as described for the native IP protocol (step 6a).
Applications for Chemical Probes of Proteolytic Activity
7b. Wash the beads three times, each time with 1 ml of RIPA buffer at 4°C (see step 6a for centrifugation conditions). 8b. Remove all liquid and resuspend the beads in SDS-PAGE sample buffer as described for the native IP protocol (steps 8a to 9a).
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AFFINITY PURIFICATION OF BIOTINYLATED ENZYMES This protocol requires a lot of starting material and a well-established and well-controlled labeling reaction that has been previously optimized (see Basic Protocol 3). The amount of starting material required will depend on the abundance of the target in the crude mixture. Once biotinylated enzymes have been identified on a small scale, the reaction can be scaled up very easily.
ALTERNATE PROTOCOL 8
Materials Biotinylated enzyme of interest PBS (APPENDIX 2E) Streptavidin-agarose beads (Pierce) PBS (APPENDIX 2E)/0.1% (w/v) SDS PD-10 desalting columns (Amersham Biosciences) 15-ml conical tubes (Falcon) 80°C water bath End-over-end rotator Tabletop centrifuge Additional reagents and equipment for labeling of peptidases (see Basic Protocol 3), desalting (UNIT 8.3), and SDS-PAGE (UNIT 10.1 and Basic Protocol 2) 1. Perform a large labeling reaction, starting with 10 to 50 mg of crude lysate (see Basic Protocol 3) in a 2.5- or 5-ml reaction. Note that the concentration of lysate used here may be higher than that used on a small scale, perhaps as high as 10 mg/ml total protein. This is important so the reaction can be desalted more easily (see step 2).
2. Stop the labeling reaction by desalting on a PD-10 column equilibrated with PBS, according to the manufacturer’s instructions (also see UNIT 8.3). Elute into a 15-ml conical tube. 3.5 ml of desalted material will be obtained for every 2.5 ml of reaction volume.
3. For denaturing conditions, add 140 µl of 10% SDS for every 3.5 ml of desalted labeling reaction (0.4% final concentration of SDS). Heat reaction at 80°C for 10 min. Denaturing the proteins before streptavidin-agarose affinity chromatography can sometimes increase the accessibility of the biotin tag to the streptavidin, especially if there is steric hindrance from the tertiary structure of the folded native enzyme. Denaturation and the presence of SDS do not significantly affect the ability of streptavidin and biotin to interact.
4. Dilute the desalted reaction with 3 vol PBS (10.5 ml for every 3.5 ml desalted reaction). For the denatured reaction, this serves to decrease the concentration of SDS to 0.1%.
5. Add 20 µl of streptavidin-agarose beads prewashed in PBS. The amount of beads added will depend on the amount of biotinylated protein present in the reaction. The capacity of the streptavidin-agarose beads is quite high, ∼3 mg biotinylated protein/ml of beads, and it is important to minimize the amount of beads and thereby minimize the amount of avidin monomer contaminating the purified enzyme after elution (see step 10). 20 ìl of beads will ideally bind 60 ìg of biotinylated enzymes, which is much less enzyme than is expected to be present in even 50 mg of total crude lysate.
6. Incubate tubes 60 min at room temperature with rotation on an end-over-end rotator. Peptidases
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7. Spin the beads at 1000 × g in a tabletop centrifuge, then remove ∼13.5 ml of supernatant, leaving ∼0.5 ml on top of the beads. Transfer the liquid and the beads to a microcentrifuge tube. 8. Wash the beads five times, each time with 1 ml of PBS (nondenaturing) or PBS/0.1% SDS (denaturing), using the centrifugation conditions in step 7. 9. Remove all the liquid using a pipet equipped with a gel-loading tip inserted all the way into the beads. Any beads that stick to the tip can be rubbed off on the inside of the tube.
10. Add 20 to 40 µl of 1× SDS-PAGE sample buffer, heat tube to 80°C for 5 min, and load sample on SDS-PAGE gel (UNIT 10.1; see Basic Protocol 2 in the current unit for conditions). It is a recommended to load as much material as possible in a single lane on a gel to maximize the signal from the staining procedure. BASIC PROTOCOL 6
ACTIVITY-BASED PROBE COMPETITION ASSAYS TO CHARACTERIZE SMALL MOLECULE PEPTIDASE INHIBITORS This protocol describes the use of ABPs to identify and characterize reversible and irreversible inhibitors of peptidases. Recently, several examples of the use of ABPs for identifying new specific peptidase inhibitors have been described (Greenbaum et al., 2002a,b). The compounds that resulted from the assay were used to assign a physiological function to specific peptidase target. These studies serve as excellent references and provide an example of what to expect when performing a competition-based screen. This application is useful when there is no source of purified enzyme or when an appropriate substrate is not available for a more classical enzymatic assay. Moreover, multiple peptidases can be assayed simultaneously. It is particularly useful in tissue culture cells or even in live animals, where it is otherwise extremely difficult or impossible to monitor enzyme-inhibitor interactions. This protocol is based on the competition between the ABP and an active-site-directed inhibitor for the target enzyme. When the inhibitor is bound, it prevents the ABP from binding and irreversibly modifying the enzyme. The resulting labeling yields a readout of the relative specific activity (the amount of label incorporated per unit time per unit of enzyme) of the untreated enzyme and the enzyme in the presence of an inhibitor. An IC50 value (the concentration of inhibitor required to effect a 50% decrease in specific activity) can be calculated that often closely matches the Ki as measured by more standard enzymatic assays. Finally, ABPs can be used not only to monitor potency of an inhibitor (the IC50 value) but also the specificity of an inhibitor for one enzyme family member over another. If a crude mixture is used that contains several related enzymes that are labeled with a single ABP, a single reaction with an inhibitor can show whether that inhibitor has specificity for one family member over another (see Fig. 21.17.1).
Applications for Chemical Probes of Proteolytic Activity
To perform this experiment, it is necessary to optimize the labeling reaction between the ABP and the enzymes in the target proteome. The reaction can be carried out in vitro (see Basic Protocol 3), in situ (see Basic Protocol 4), or even in vivo (not covered in this unit). However, the single most important parameter that needs to be established to have success with this application is to determine when to stop the reaction and analyze the results. ABPs are irreversible inhibitors of their enzyme targets. This means that the concentrations of enzyme and ABP (which can be considered a substrate of the enzyme) in the reaction are decreasing over time (see Fig. 21.17.2). In Henri-Michaelis-Menten kinetics, it is most straightforward to measure the specific activity of an enzyme when the amount of enzyme and substrate do not change over the time the reaction is being monitored (Fersht, 1985). The fact that both are decreasing in the reaction between enzyme and ABP
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poses a problem for monitoring specific activity. This can be overcome by stopping the reaction at a point where the amount of label incorporated into the untreated enzyme is still increasing linearly with time (see Figure 21.17.2). Under these conditions, it can be assumed that the concentrations of enzyme and ABP have not decreased enough to significantly affect the specific activity of the enzyme. Quantification of labeling is most easily done with a fluorescent or radioactive tag (see Basic Protocol 2). It is possible to quantify the amount of labeling by a biotinylated tag using immunoblot analysis (see Basic Protocol 2) but is more time consuming, has a more limited dynamic range, and requires a densitometer capable of reading the signal from exposed films. This protocol assumes that there is a suitable ABP with a fluorescent or radioactive tag.
B
A O
O N H
H N
O
OH
inhibitor
N
O
125
probe: I-JPM-565 competitor: 0 0.1 1
O
10
50 100
M 40 kDa 33 kDa Cat B
lysate/cell
Cat S 20 kDa
probe
O H N
general cysteine protease inhibitor - E-64
OH O
N H
O O 125
probe: I-JPM-565 increasing conc.
competitor:
0
0.1
1
10
50
100
M
40 kDa 33 kDa Cat B Cat S 20 kDa
indirect competition Cat B specific protease inhibitor - MB-074
Figure 21.17.1 Competition analysis to generate inhibitor potency and selectivity profiles using an activity-based probe (ABP) as a readout. (A) Schematic representation of the general competition protocol. Samples are preincubated with a range of concentrations of an inhibitor of interest followed by labeling with a small-molecule ABP carrying a tag (star). Samples are analyzed by SDS-PAGE followed by detection of probe-labeled peptidases. As the inhibitor binds to a target enzyme in the sample, it blocks labeling by the ABP. This is visualized by loss of enzyme labeling as the concentration of inhibitor is increased. Inhibitor selectivity and potency can be determined by quantification of labeling for each target enzyme throughout the range of inhibitor concentrations. (B) Examples of competition profiles obtained for total cellular extracts treated with a general papain family peptidase inhibitor (top panel) and a cathepsin B specific inhibitor (bottom panel) followed by labeling with the general probe of papain family peptidases [125I]JPM-565 (Table 21.17.1 contains references for this probe). Details of the competition protocol can be found in Basic Protocol 6. Peptidases
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Amount of enzyme
unlabeled enzyme
line through early time points
labeled enzyme
Time
Figure 21.17.2 Hypothetical curves showing the increase in labeled enzyme (dashed line) and decrease in free enzyme over time. The shaded area shows the time of the reaction during which the labeling is still linear. The time point at which to stop the reaction should be chosen from within the linear portion of the labeling reaction.
Materials Source of target enzyme(s) (in solution or in live tissue culture cells) Activity-based probe with a fluorescent or radioactive tag (see Basic Protocol 1 or Alternate Protocol 1 or 2) Small molecule inhibitors to be tested: as 0.1 mM stocks in DMSO Dimethylsulfoxide (DMSO) Graphing program (e.g., Microsoft Excel) 96-well plate Multichannel pipettor capable of dispensing 1 to 20 µl Additional reagents and equipment for peptidase labeling in vitro (see Basic Protocol 2) or in situ (see Basic Protocol 4), SDS-PAGE (UNIT 10.1 and Basic Protocol 2), and autoradiography (UNIT 10.11) Optimize time of reaction 1. Set up a labeling reaction according to Basic Protocol 3, but do not add the fluorescent or radioactive ABP. This can be an in vitro or in situ labeling experiment. The size of the reaction should be sufficient to take aliquots for six time points from an in vitro reaction, or should consist of five identical wells of tissue culture cells as an in situ experiment. At this point, all the parameters described in Basic Protocol 3 and Basic Protocol 4 should be optimized.
2. Start the reaction by adding the ABP and at times of 10, 20, 30, 60, 90, and 180 min for in vitro experiments or 30, 60, 120, 180, 240 min for in situ experiments, remove an aliquot, and stop the reaction as described in Basic Protocol 3 or Basic Protocol 4. Applications for Chemical Probes of Proteolytic Activity
3. Analyze the reactions by SDS-PAGE (UNIT 10.1 and Basic Protocol 2) and fluorescence scanning, autoradiography (UNIT 10.11 and Basic Protocol 2), or phosphor imaging.
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4. Using a graphing program, plot the amount of label incorporated on the y axis and time on the x axis (see Fig. 21.17.2). 5. Draw lines through different combinations of the early time points to determine the time period during which the amount of labeling increases linearly with time (R2 > 0.95 using linear regression). 6. Select a reaction termination time point from within this linear portion of the graph. Because small variations in reaction conditions can affect the incorporation of label in the target enzyme, it is important to be in the heart of the linear portion of the labeling reaction, not near the point where it becomes nonlinear.
Screen a library of potential inhibitors 7. In a 96-well plate, set up reactions to label the enzyme of interest, but do not add the ABP. Again, this can be done either in vitro or in situ.
8. Add 1/100 volume of DMSO alone or the inhibitors to the wells (final concentration of 1 µM of each inhibitor). Incubate reaction 30 min. For a standard reversible inhibitor at room temperature or above, 30 min is more than enough time for the binding reaction to reach equilibrium.
9. Using a multichannel pipettor, add the ABP to the wells and allow the reaction to proceed until the time determined in step 6. 10. Stop the reactions and analyze the results by SDS-PAGE (UNIT 10.1 and Basic Protocol 2) and fluorescence scanning, autoradiography (UNIT 10.11 and Basic Protocol 2), or phosphor imaging. Potent inhibitors will cause at least a 50% decrease in enzyme labeling at 1 ìM and those that do can be tested further to determine IC50 values (see below).
Measure the IC50 value of an inhibitor for the enzyme of interest 11. Set up a labeling reaction but do not add the ABP. Set up enough reactions for an untreated control and four reactions for each inhibitor to be tested. 12. Add 1/100 volume of DMSO or a dilution series of the inhibitor to be tested in 100% DMSO. A good starting range for an inhibitor dilution series is 10, 1, 0.1, and 0.01 ìM. Incubate the reaction for 30 min.
13. Add the ABP and allow the reaction to proceed until the time determined in step 6. 14. Analyze the results by SDS-PAGE (UNIT 10.1 and Basic Protocol 2) and fluorescence scanning, autoradiography (UNIT 10.11 and Basic Protocol 2), or phosphorimaging. 15. To determine the IC50 value, plot the data with the label incorporated on the y axis and the inhibitor concentration on the x axis (see Fig. 21.17.3). Use a logarithmic scale on the x axis. In the ideal experiment this will create a straight line. The IC50 value of the compound can be calculated by determining the equation describing the line and extrapolating from that equation the concentration of compound required to obtain a 50% reduction in enzyme labeling compared to the untreated control. The IC50 value can be more accurately determined by repeating the experiment with concentrations of inhibitor clustered more closely around the IC50 (5 to 10 concentration points within an order of magnitude on either side of the measured IC50). Peptidases
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Linear regression extrapolated from linear portion of graph
50% of maximal activity 1,200
1,000
$
y = – 270x + 1715
&
Activity units
"
800
!
6
7
"
#
$
%
600 400 200 0 0
1
2
3 4 Log[inhibitor]
5
8
9
Log[IC50] (Calculated from linear trendline equation]
Figure 21.17.3 Determining the IC50 value of an inhibitor by competition labeling using an ABP. Sample of a typical graph generated from competition labeling of a target peptidase in the presence of increasing concentrations of an inhibitor. As the inhibitor binds in the active site of the peptidase, it blocks labeling by the ABP. By plotting the intensity of peptidase labeling (activity units) versus log[inhibitor] it is possible to generate a curve as shown here. A linear regression can be extrapolated from the linear portion of curve and used to determine the IC50 value of the inhibitor.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Cell lysis buffer 50 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) 1% (w/v) CHAPS Peptidase inhibitor cocktail (Sigma) 50 µM E-64 (Calbiochem) Store up to 3 months at 4°C Fixation solution 50% (v/v) methanol 12% (v/v) acetic acid 38% (v/v) distilled deionized H2O Store up to 6 months at room temperature Immunoblot blocking solution PBS (APPENDIX 2E) containing: 0.1% (v/v) Tween 20 2% (w/v) casein 0.01% (w/v) sodium azide Prepare fresh for use Applications for Chemical Probes of Proteolytic Activity
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Immunoblot transfer buffer For 1 liter of 10× stock solution 58 g Tris base 29 g glycine Store stock solution up to 1 year at room temperature For 1× blotting transfer buffer, combine the following in H2O: 10% (v/v) methanol 10% (v/v) 10× stock solution (see above) Prepare fresh IP wash buffer 50 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 150 mM NaCl 0.5% (v/v) NP-40 1 mM EDTA, pH 8.0 (APPENDIX 2E) Store at 4°C for up to 6 months RIPA buffer 1× PBS (APPENDIX 2E) 1% (v/v) Triton X-100 1% (w/v) sodium deoxycholate 0.1% (w/v) SDS 2 mM EDTA Store at room temperature for up to 6 months RIPA dilution buffer 1.1× PBS (APPENDIX 2E) 1.1% (v/v) Triton X-100 1.1% (w/v) sodium deoxycholate 2 mM EDTA (APPENDIX 2E) Store at room temperature for up to 6 months COMMENTARY Background Information
Critical Parameters
Recent advances in the use of small molecule and protein affinity tags have greatly enhanced biochemical and cell biological studies of peptidases. Activity-based probes (ABPs) have been developed that target several major classes of peptidases, including the lysosomal cysteine peptidases, caspases, DUBs, serine peptidases, and the proteasome. The authors have compiled a complete list of the currently available reagents for use as ABPs (Table 21.17.1) and readers are strongly encouraged to obtain the cited references before embarking on any experiments using a probe. While the protocols presented in this unit provide all the necessary experimental details, the reader is referred to the individual published articles for further information regarding the background of specific target peptidase families.
The process of generating an ABP from unlabeled starting material requires optimization of the labeling reaction. In the case of radioactive iodine, the labeling protocol is somewhat flexible, as labeling efficiency is typically low and it is difficult to remove unlabeled compound after completion of the labeling reaction. The authors have found that the general procedures outlined in Basic Protocol 1 and Alternate Protocol 1 work well for virtually all small molecule and protein probes. The only potential problem with this method arises in the case where an electrophile or probe scaffold is sensitive to the radical-mediated labeling conditions. If this is a concern, a simple cold iodination should be performed, and the products analyzed by mass spectrometry and/or NMR. In the case where biotin or other activated esters are used to tag a free amine-containing probe, the most critical parameters to optimize are solvent composition and reaction
Peptidases
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Applications for Chemical Probes of Proteolytic Activity
time. Many amines will not react to completion, and prolonging the reaction times will not improve yields. The authors recommend careful monitoring of the reaction using analytical HPLC. Once a probe has been selected and prepared, the critical parameters for their use vary based on the method of detection used to analyze peptidase labeling. When using radioactive probes, the main consideration is the length of time the gel is incubated in wash solution. In many cases, the free probe may remain in the gel and can lead to an extremely wide band of signal that can mask primary bands of interest. Further washing of the gel or switching to a different percentage gel can fix this problem. When using biotinylated ABPs, detection is based on a relatively standard blotting method. For this protocol, the reagents used for detection of the biotin molecule are critical. Many of the commonly used streptavadin-HRP conjugates produce high background signals in affinity blots. The authors have found that Vectastain products are effective and have very low background due to the coupling method used to generate the strepavidin-HRP species. Another critical parameter to consider when blotting concerns the wash conditions used prior to addition of the HRP reagents. In general, this protocol can be optimized in a manner similar to optimization of a standard immunoblot (UNIT 10.7). When using fluorescent reagents, the critical parameters are to use the proper scanner, equipped with an optimal laser, and also to use an optimal probe concentration and labeling time to minimize the background signal. Since there are many parameters in a typical labeling reaction that can be altered, it is recommended to start by altering enzyme concentration and probe concentration, since these are likely to have the most dramatic impact on labeling. In most cases, buffer conditions for an enzyme or enzyme family can be obtained from the literature. If not, the typical buffer recipes given in this unit are a good place to start. It is very important to have the correct specificity control in place before undertaking any optimization. The probes described in this unit contain active electrophiles and are capable of reacting nonspecifically with proteins that contain strong nucleophiles. Irreversible activesite inhibitors of many families of enzymes are available (see Table 21.17.2) and should be used in every experiment. A pre-heat control (achieved by heating the enzyme-containing sample to 70°C for 10 min) also works well to distinguish between specific labeling of pepti-
dases from nonspecific labeling of irrelevant proteins. For affinity purification, the two most important issues are having enough purified protein on the gel and the purity of the preparation. Both parameters will have a direct effect on the ability to identify the protein by trypsinolysis and mass spectrometry. Identification of a protein should be relatively easy with >1 pmol protein (40 ng of a 40-kDa protein) on the gel; it is feasible with 0.1 to 1.0 pmol protein, and extremely difficult with <0.1 pmol of protein on the gel. The presence of contaminating proteins will have a detrimental effect on digestion and identification, especially if the unknown protein is present in low abundance and the contaminant is present in high abundance. The two most common sources of contaminants in streptavidin-affinity chromatography are monomeric and dimeric avidin (14 and 28 kDa respectively) and abundant endogenously biotinylated proteins. Avidin monomer comes from the streptavidin-agarose beads used for affinity purification, so it is important not to use too much during the purification. Endogenously biotinylated proteins can be eliminated by incubating the lysate with streptavidin-agarose beads prior to labeling with the ABP. For the trypsinolysis and mass spectrometry analysis, the most important issue is to know who is going to do the mass spectrometry and in what form they need the sample to be prepared. The identification of tryptic peptides in a mass spectrometer is technically challenging and is very sensitive to contaminants such as detergent and salts. Unless one has significant experience in this area, it is absolutely necessary to consult MS collaborators before embarking on the purification and analysis of the results (also see Chapter 16). As mentioned in the introduction, the most important parameter that needs to be optimized when performing competition experiments is the time of peptidase labeling. In order to measure a relative specific activity of labeling in the presence or absence of an active-site inhibitor, the ABP must be labeling the enzyme linearly with respect to time. The labeling reaction and detection of labeling must be optimized and quantifiable. Once the time of the reaction has been established, it is best to then use a known inhibitor of the enzyme to verify that the competition is proceeding as expected.
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are also endogenously biotinylated proteins that will be detected on streptavidin-HRP immunoblots. These nonspecific interactions can be ruled out as bona fide enzyme targets by incorporating the specificity controls mentioned above. The amount of label incorporated into the nonspecific proteins can be significantly altered by using less probe, a lower concentration of protein, and shorter incubation times. See Figure 21.17.4 for an example of a crude protein extract labeled with probes tagged with biotin, 125I, and fluorescent tags. In an immunoprecipitation, it is expected that the antibody will bind with high specificity to the target(s) of interest and not to other proteins. Antibody raised against a specific protein or peptide from a specific protein
Troubleshooting Table 21.17.6 has been included to assist in troubleshooting some of the more critical protocols. The table is divided according to the various Basic Protocols.
Anticipated Results Most enzymes will react strongly and specifically with well-designed ABPs. With a purified enzyme, it is expected that there will only be one major band labeled on the SDS gel, but this will depend on the purity of the preparation. In a crude lysate, there could be dozens of enzymes labeled, depending on the probe and the lysate. There will likely be nonspecific labeling of proteins that contain a strong nucleophile somewhere on their surface. There
B
A
probe:
biotin
radioactive
blot
autoradiography
O H N O
OH probe
N H
O
#1-
O
#2#3#4-
lysate/cell
detection method: red
fluorescent green blue
yellow
#1SDS-PAGE and autoradiography
#2#3#4-
direct labeling
detection method:
fluorescent scanner
Figure 21.17.4 Direct labeling of peptidases with small molecule probes. (A) Schematic representation of a direct labeling experiment. Complex protein mixtures from cells are incubated with a small molecule ABP carrying a tag (star). After covalent modification of peptidases, samples are analyzed by SDS-PAGE and tag is detected via blotting, autoradiography, or fluorescence scanning. (B) Examples of general labeling profiles of papain family cysteine peptidases in total rat liver homogenates. Profiles were obtained for the same sample labeled with a set of identical ABPs that differ only in the type of tag used. Examples include results for a radioactive tag, a biotin tag, and four different colored fluorescent tags. The method used for detection of the labeled bands is listed below. For details of labeling and detection method see Basic Protocols 2 and 3.
Peptidases
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Table 21.17.6
Troubleshooting Guide for Using Chemical Probes to Detect Proteolytic Activity
Problem
Possible cause
Solution
Probe synthesis and detection (Basic Protocols 1 and 2) Poor efficiency of iodination
Too much DMSO
Poor efficiency of coupling to fluorophore or biotin
Hydrolysis of probe
Low signal-to-noise ratio following detection protocols
High background
Low signal
In situ labeling problems
Too few bands
Too many bands
Start from a more concentrated DMSO stock solution Increase the ratio of tag to probe and shorten incubation time (<2 hr) Load less protein on gel Wash membrane for longer (for biotin labeling) Incubate with fixation solution (for radioactive and fluorescent labeling) Load more protein on gel Incubate membrane with higher concentration of HRP reagent (for biotin labeling) Label in the absence of serum in the growth medium Increase labeling duration Check cell intactness Reduce probe concentration
Labeling optimization (Basic Protocol 3) No labeled enzymes on gel ABP not working
High background labeling
Specificity control does not work
Applications for Chemical Probes of Proteolytic Activity
Use a positive control enzyme that is known to be labeled by the ABP Perform immunoblot to No enzymes in confirm the presence of target mixture or enzymes enzymes. Be extremely careful are inactivated while handling protein mixtures Wrong detection Double check which tag is method used present on the ABP Concentration of ABP Increase concentration of ABP or enzyme too low or enzyme Buffer conditions Optimize labeling conditions incorrect as described Concentration of ABP Reduce concentration of ABP or enzyme too high or enzyme Problem with See troubleshooting tips for detection method Basic Protocol 2 Using the wrong Check Table 21.17.2 for the correct specificity control for small-molecule the enzyme family being specificity control targeted Enzyme not denatured Try heating the enzyme upon heating solution for longer or at higher temperatures. Add SDS to 1% to completely inactivate the enzyme before heating. continued
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Table 21.17.6 continued
Troubleshooting Guide for Using Chemical Probes to Detect Proteolytic Activity,
Problem
Possible cause
Solution
Too few bands
Label in the absence of serum in the growth medium Increase labeling duration Check cell intactness Reduce probe concentration
In situ labeling (Basic Protocol 4) In situ labeling problems
Too many bands Isolation of labeled proteins (Basic Protocol 5) No labeled enzyme in the IP
Labeling did not work Reoptimize labeling as in Basic Protocol 3 Antibodies do not Try to IP a recombinant form work in the IP of the labeled enzyme High background in the IP Too much antibody Reduce the concentration of was used antibody in the reaction Antibody not specific Affinity purify the antibody against the target enzyme (see Harlow and Lane No proteins visible on gel after Enzymes not labeled Check labeling by avidin-HRP affinity purification immunoblot (see Basic Protocol 2) Add more streptavidin-agarose Not enough streptavidin-agarose added Protein staining did Run a lane of a known amount not work of protein standard to determine sensitivity of staining procedure Check that sample buffer is Proteins were not correct eluted from streptavidin-agarose No peptides detected in mass In-gel digestion did Perform in-gel digestion using spectrometer not work a known amount of a standard protein (e.g. Not enough protein Scale up purification procedure purified and run on gel Competition with inhibitors (Basic Protocol 6) No labeling of enzyme
Inhibitors do not prevent labeling of enzyme
Labeling conditions Reoptimize labeling as in Basic not well established Protocol 3 Detection not working See troubleshooting tips for Basic Protocol 2 Not enough inhibitor Add a higher concentration of added inhibitors Inhibitors are not Try different inhibitors. Use a potent known potent inhibitor as a positive control. Incubate inhibitors with Inhibitors have not enzyme for longer time period reached equilibrium before adding ABP binding conditions with enzyme continued Peptidases
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Table 21.17.6 continued
Applications for Chemical Probes of Proteolytic Activity
Troubleshooting Guide for Using Chemical Probes to Detect Proteolytic Activity,
Problem
Possible cause
Solution
Enzyme labeling is not linear with respect to time
Too much ABP in the reaction Too little enzyme in the reaction Problems with detection method
Decrease concentration of ABP
should immunoprecipitate only that protein, and the result should be a single labeled band on the gel. These profiles will look much cleaner than those of direct loads (Fig. 21.17.4) but in most cases the signal will be weaker due to loss of sample during the precipitation. For the affinity purification, it is expected that every enzyme that appeared on the avidinHRP immunoblot (see Basic Protocol 2) will also be present after purification and staining of the gel with Coomassie blue or silver stain. There may be more bands present in the purification, which are either contaminants or proteins associated with the labeled enzymes. This distinction can be made by performing a mock purification from a lysate that was pretreated with a control inhibitor prior to labeling with the ABP. The contaminating proteins should be present in both the biotinylated and nonbiotinylated purification, whereas associated proteins should be present only in the biotinylated preparation. Associated proteins will not be present if the denaturing purification procedure is used. Competition studies with ABPs and unlabeled inhibitors typically result in profiles in which distinct labeled peptidase bands disappear upon addition of increased concentrations of an inhibitor (Fig. 21.17.1). Using this method, it should be possible to measure an IC50 value for an inhibitor of a peptidase in crude sample that closely matches the Ki determined for that same peptidase using standard enzymatic assays. The IC50 value measured in tissue culture cells could be considerably higher, because the inhibitor needs to get into the cells and find the target enzyme before being pumped out of the cell or metabolized. It is not uncommon to obtain IC50 values that are an order of magnitude higher in tissue culture cells in comparison with those determined in
Increase the concentration of enzyme Make sure that detection is linear in the [enzyme] vs time range being used. This can be done by serially diluting labeled enzyme and making sure that the detected signal is linear with dilution.
solution. Some inhibitors will be ineffective simply because they are not cell permeable.
Time Considerations Iodination of small molecule or protein probes take about 1 hr. Coupling of a fluorophore or a biotin tag to an ABP requires a 3-hr incubation, followed by HPLC purification, which can take an additional hour. Subsequently, the HPLC elution fractions that contain the tagged probe need to be lyophilized for 16 to 24 hr. In vitro labeling experiments are usually fast and easy to set up. Labeling the sample with the ABP takes 1 to 2 hr. Preincubation with inhibitors prior to labeling with ABPs normally takes about the same amount of time. In situ labeling can be done in 2 to 12 hr depending on the type of probe and the tag being used. Typically, for labeling cysteine peptidases within cells using a fluorescent probe, a 1- to 3-hr incubation is sufficient. Lysis of cells following labeling usually takes 30 min to 1 hr. Detecting the signal in the both in vitro and in situ labeled samples takes different amounts of time depending on the type of tag used. For radioactive probes, detection requires 1 to 2 days. For biotinylated probes, detection takes 4 to 5 hr. For fluorescent probes, detection is fast, and only requires separation by SDSPAGE and fluorescence scanning, which altogether take 1 to 2 hr. Purification and identification of labeled peptidases takes 2 days, in which biochemical purification is performed on the first day; identification by mass spectroscopy could be completed in the next day. Typically, identification of proteins by LC/MS takes 1 hr per sample.
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Literature Cited Bogyo, M., McMaster, J.S., Gaczynska, M., Tortorella, D., Goldberg, A.L., and Ploegh, H. 1997. Covalent modification of the active site threonine of proteasomal beta subunits and the Escherichia coli homolog HslV by a new class of inhibitors. Proc. Natl. Acad. Sci. U.S.A. 94:6629-6634. Bogyo, M., Verhelst, S., Bellingard-Dubouchaud, V., Toba, S., and Greenbaum, D. 2000. Selective targeting of lysosomal cysteine peptidases with radiolabeled electrophilic substrate analogs. Chem. Biol. 7:27-38. Borodovsky, A., Kessler, B.M., Casagrande, R., Overkleeft, H.S., Wilkinson, K.D., and Ploegh, H.L. 2001. A novel active site-directed probe specific for deubiquitylating enzymes reveals proteasome association of USP14. EMBO J. 20:5187-5196. Borodovsky, A., Ovaa, H., Kolli, N., Gan-Erdene, T., Wilkinson, K.D., Ploegh, H.L., and Kessler, B.M., 2002. Chemistry-based functional proteomics reveals novel members of the deubiquitinating enzyme family. Chem. Biol. 9:1149-1159. Fersht, A. 1985. Enzyme Structure and Mechanism, 2nd ed. W.H. Freeman, New York. Greenbaum, D.C., Medzihradszky, K.F., Burlingame, A., and Bogyo, M. 2000. Epoxide electrophiles as activity-dependent cysteine peptidase profiling and discovery tools. Chem. Biol. 7:569-581. Greenbaum, D.C., Baruch, A., Hayrapetian, L., Darula, Z., Burlingame, A., Medzihradszky, K.F., and Bogyo, M. 2002a. Chemical approaches for functionally probing the proteome. Mol. Cell. Proteomics 1:60-68. Greenbaum, D.C., Baruch, A., Grainger, M., Bozdech, Z., Medzihradszky, K.F., Engel, J., DeRisi, J., Holder, A.A., and Bogyo, M. 2002b. A role for the peptidase falcipain 1 in host cell invasion by the human malaria parasite. Science 298:2002-2006. Harlow, E. and Lane, D. 1988. Antibodies: A Laboratory Manual, 1st ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. Kessler, B.M., Tortorella, D., Altun, M., Kisselev, A.F., Fiebiger, E., Hekking, B.G., Ploegh, H.L., and Overkleeft, H.S. 2001. Extended peptidebased inhibitors efficiently target the proteasome and reveal overlapping specificities of the catalytic beta-subunits. Chem. Biol. 8:913-929. Kidd, D., Liu, Y., and Cravatt, B.F. 2001. Profiling serine hydrolase activities in complex proteomes. Biochemistry 40:4005-4015. Krupa, J.C. and Mort, J.S. 2000. Optimization of detergents for the assay of cathepsins B, L, S, and K. Anal. Biochem. 283:99-103. Liu, Y., Patricelli, M.P., and Cravatt, B.F. 1999. Activity-based protein profiling: The serine hydrolases. Proc. Natl. Acad. Sci. U.S.A. 96:14694-14699. Meng, L., Mohan, R., Kwok, B.H., Elofsson, M., Sin, N., and Crews, C.M. 1999. Epoxomicin, a potent and selective proteasome inhibitor, exhibits in vivo antiinflammatory activity. Proc. Natl. Acad. Sci. U.S.A. 96:10403-10408.
Mikolajczyk, J., Boatright, K.M., Stennicke, H.R., Nazif, T., Potempa, J., Bogyo, M., and Salvesen, G.S. 2003. Sequential autolytic processing activates the zymogen of Arg-gingipain. J. Biol. Chem. 278:10458-10464. Patricelli, M.P., Giang, D.K., Stamp, L.M., and Burbaum, J.J. 2001. Direct visualization of serine hydrolase activities in complex proteomes using fluorescent active site-directed probes. Proteomics 1:1067-1071. Rappsilber, J. and Mann, M. 2002: What does it mean to identify a protein in proteomics? Trends Biochem. Sci. 27:74-78. Schaschke, N., Assfalg-Machleidt, I., Lassleben, T., Sommerhoff, C.P., Moroder, L., and Machleidt, W. 2000. Epoxysuccinyl peptide-derived affinity labels for cathepsin B. FEBS Lett. 482:91-96. Shi, G.P., Munger, J.S., Meara, J.P., Rich, D.H., and Chapman, H.A. 1992. Molecular cloning and expression of human alveolar macrophage cathepsin S, an elastinolytic cysteine peptidase. J. Biol. Chem. 267:7258-7262. Vugmeyster, Y., Borodovsky, A., Maurice, M.M., Maehr, R., Furman, M.H., and Ploegh, H.L. 2002. The ubiquitin-proteasome pathway in thymocyte apoptosis: Caspase-dependent processing of the deubiquitinating enzyme USP7 (HAUSP). Mol. Immunol. 39:431-441. Wilchek, M. and Bayer, E.A. 1990. Introduction to avidin-biotin technology. Methods Enzymol. 184:5-13.
Key References Havlis, J., Thomas, H., Sebela, M., and Shevchenko, A. 2003. Fast-response proteomics by accelerated in-gel digestion of proteins. Anal. Chem. 75:1300-1306. In-gel digestion of protein bands. Shevchenko, A., Wilm, M., Vorm, O., and Mann, M. 1996. Mass spectrometric sequencing of proteins silver-stained polyacrylamide gels. Anal. Chem. 68:850-858. Silver staining and MS sequencing. Rappsilber, J. and Mann, M., 2002. See above. MS sequencing.
Contributed by Matthew Bogyo Stanford University Stanford, California Amos Baruch and Douglas A. Jeffery Celera Genomics South San Francisco, California Doron Greenbaum University of California San Francisco, California Anna Borodovsky, Huib Ovaa, and Benedikt Kessler Harvard Medical School Boston, Massachusetts
Peptidases
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CHAPTER 22 Gel-Based Proteome Analysis INTRODUCTION
P
roteomics is a new discipline that originated in the mid-1990s and has grown rapidly in a very short time. As with many new fields, there is some disagreement concerning what proteomics really encompasses, and some feel that proteomics is simply a new name for the protein biochemistry studies that many have been pursuing for years. However, these broad definitions do not accurately reflect the paradigm shift driven by several key enabling technological advances in the 1990s that led to the birth of proteomics as a unique approach to biology, namely availability of complete genome sequences and high sensitivity mass spectrometry methods to identify proteins. Perhaps the best current definition of proteomics is “any large-scale protein-based systematic analysis of the proteome or defined sub-proteome from a cell, tissue, or entire organism.” Proteomics is the logical next step after genome sequencing, and should enhance our understanding of cellular and disease processes, as well as our ability to develop new therapies. However, proteomics is more complicated and more challenging than sequencing genomes because genomes are finite and usually static over the lifetime of a cell while proteomes are constantly changing. In addition, the protein repertoire of higher eukaryotes is far greater than the number of open reading frames in the genome because individual genes often produce many different protein products that have important functional differences. These include alternatively spliced proteins and heterogeneous post-translational modifications. This chapter will describe gel-based methods and related technologies for analyzing proteomes. A future chapter will address non-gel based methods of separating and analyzing proteomes including multi-dimensional chromatographic separations coupled directly to mass spectrometers. Antibody and other protein arrays will be described as well. Subsequent chapters will address targeted proteomes and proteome bioinformatics.
The current chapter begins with a general overview of proteomics (UNIT 22.1) focusing on the three most common types of proteome studies: 1) quantitative protein profile comparisons, 2) analysis of protein-protein interactions, and 3) compositional analysis of simple proteomes or subproteomes, such as organelles or large protein complexes. The complexity of different types of proteomes, the merits of targeted versus global proteome studies, and the advantages of alternative separation and analysis technologies are discussed. UNIT 22.2 describes protein profile comparisons using differential fluorescent labeling of two different samples that are then mixed and separated using a single two-dimensional gel. This method eliminates the gel-to-gel variation that frequently occurs with 2-D gels, and thereby improves comparisons of protein profiles and reduces the number of gels that need to be run to quantitatively compare protein profiles from two or more samples. Sample preparation is a major challenge for most proteome studies regardless of whether gel- or non-gel-based separation and analysis methods are used. Proper experimental design and reproducibility of sample preparation must be carefully considered to avoid wasting substantial effort studying sample processing artifacts or experimental variations unrelated to the biological or pathological question that is being studied. When analyzing Contributed by David Speicher Current Protocols in Protein Science (2003) 22.0.1-22.0.2 Copyright © 2003 by John Wiley & Sons, Inc.
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tissue samples, as opposed to cell lines or single cell organisms, tissue heterogeneity raises two potential problems. First, the presence of multiple cell types increases the total number of proteins present, which complicates separation and analysis. More importantly, when comparing two or more specimens, variations in the numbers of different types of cells can obscure changes in protein profiles with a specific cell type of interest, such as cancer cells. A recently developed approach for decreasing the heterogeneity of tissue specimens is laser capture microdissection (UNIT 22.3). However, the total amount of protein that can be isolated in a realistic timeframe is currently rather small, and requires very high-sensitivity subsequent analysis methods. Three such high-sensitivity methods also described in this unit are: 2-D PAGE coupled with sensitive detection methods, immunoblotting after 1-D or 2-D electrophoresis, and surface enhanced laser desorption ionization mass spectrometry (SELDI MS). UNIT 22.4 describes methods for preparing samples for quantitative comparisons using 2-dimensional gels. These protocols address dispersion and solubilization of commonly used types of samples including cultured cells, solid tissues, biological fluids, plant tissues and cells, and isolated organelles. Over the past several years, increasing emphasis has been placed on the importance of sample prefractionation prior to 2-D gel analyses. This is because the number of protein spots that can be typically separated on a full sized 2-D gel (~18 × 18 cm) is usually less than 2000, while the number of unique protein components in the simplest proteomes of higher eukaryotes is thought to be at least 20,000 to 50,000. Effective strategies for detecting larger portions of complex proteomes and particularly low abundance proteins are needed. At present, some of the most promising approaches involve isolation of organelles or systematic fractionation of a proteome into a modest number of fractions or pools for subsequent gel-based or non-gel protein profiling analyses. UNIT 22.5 describes one very powerful electrophoretic prefractionation method that can be used to isolate either intact organelles or pools of proteins based upon their pIs. Future units will address other prefractionation strategies to simplify complex proteome samples, image analysis of gels, internet resources for 2-D gel databases, and identification of proteins in gels. David Speicher
Introduction
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Overview of Proteome Analysis WHAT IS PROTEOME ANALYSIS? The term “proteome” was first introduced in the mid-1990s by Wilkins and Williams to indicate the entire “PROTEin” complement expressed by a “genOME” of a cell, tissue, or entire organism (Wilkins et al., 1996). The proteome of a cell is therefore that cell’s specific protein complement in a defined physiological context at a specific point in time. Proteome analysis or “proteomics” is a relatively new discipline that has been growing rapidly since its inception. As with many new fields there is some confusion concerning what proteomics really encompasses. For example, some investigators regard proteomics as a new name for an older discipline, namely protein biochemistry. A related viewpoint is to define proteomics as any study of proteins. While there is some merit to these perspectives, they do not adequately reflect the paradigm shift driven by several key enabling technological advances (see below) that occurred in the 1990s and led to the birth of proteomics as a unique approach to biology. Specifically, instead of focusing on single proteins or simple macromolecular complexes as in most previous protein biochemical studies, proteomics takes a broader, more comprehensive, aggressive, and systematic approach to investigation of biological systems. Proteomics substantially overlaps but does not entirely encompass “functional genomics,” i.e., the functional analysis of genes. On the other hand, “structural genomics,” the recent, focused large-scale effort to determine threedimensional structures of many proteins in a high-throughput mode, should be regarded as a separate complementary discipline. Similarly, most large-scale nucleic acid–based studies, including mutagenesis, gene knockouts, or nucleotide microarrays, are not considered part of proteomics. However, this distinction is particularly fuzzy for nucleotide microarrays, which measure mRNA levels. Although it is now well established that these measurements do not accurately reflect expression levels for all proteins, they do accurately detect quantitative changes of some proteins and therefore overlap with proteomic analysis of quantitative changes. Some researchers feel that the inception of proteomics can be traced back to the mid-1970s, when 2-D gel electrophoresis (2DE) was first used to separate proteins and
quantitatively compare changes related to biological questions of interest. This method remains one of the major tools for the quantitative comparison of protein levels among two or more samples (protein profiling). However, the early 2-D gel studies were extremely limited in their analytical scope, since efficient technologies to identify and further characterize the separated proteins did not exist at that time. The best current definition of proteomics is “any large-scale protein-based systematic analysis of the proteome or defined sub-proteome from a cell, tissue, or entire organism.” Although many technologies contribute to current proteome analysis studies, two key enabling advances occurred in the 1990s that made large-scale, systematic studies of proteins feasible. First, large-scale genome sequencing efforts began producing databases containing complete or nearly complete genomes for a number of prokaryotes, followed by important model eukaryotic organisms including S. cerevisiae and C. elegans. Subsequently, the human genome has been essentially completed and the mouse sequence is nearing completion. The importance of complete genome sequences for enabling proteome studies is reflected by considering earlier 2-D gel studies that were similar to many current proteome comparisons. When quantitative differences were observed in these early 2-D comparisons, it was feasible, but not easy, to obtain some protein sequence information for the most abundant proteins in the 1980s using Edman sequencing. However, at that time only a relatively small portion of protein sequences for any organism were present in sequence databases and therefore most experimental sequence data did not match known proteins. Therefore, further interpretation of the importance of protein changes or pursuit of further studies of these proteins could not, typically, be readily accomplished. Second, mass spectrometry advances in the 1990s enabled rapid, high sensitivity identification of proteins if the corresponding protein sequences were in available databases. Hence, both developments were essential for enabling effective proteomic studies of any species where the complete genome was sequenced. Proteomics represents the next logical step after genome sequencing and promises to enhance our understanding of cellular and disease processes, as well as our ability to develop new
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therapies. However, analysis of proteomes is much more complicated and challenging than sequencing genomes for three reasons. First, in higher eukaryotes, a single gene often produces many different forms of the protein including alternative spliceoforms and diverse post-translational modifications of the protein such as N-terminal processing, glycosylation, phosphorylation, ubiquitination, and sulfation, and these different forms of the proteins usually have important functional differences. Second, genomes are largely static during the lifetime of a cell or organism, but proteomes are highly dynamic and can change substantially, often over very short time scales, due to external stimuli and during development, growth, and aging. One frequently cited illustration of the dramatic impact of proteome changes in a single organism is the comparison between a caterpillar and its development into a butterfly— i.e., a single genome can result in two different proteomes with strikingly different phenotypes. Although sequencing a genome is a finite problem, the constant dynamics of proteomes means that an essentially infinite number of proteomes can be produced during the lifetime of a cell or organism. Third, the technologies available for analyzing proteomes are currently not as robust as nucleic acid based methods and further improvements are needed to optimally analyze the most complex proteomes.
PROTEOME COMPLEXITY, EXPERIMENTAL DESIGN, AND SAMPLE PREPARATION
Overview of Proteome Analysis
As the above discussion suggests, the challenges in proteome analysis increase in higher organisms. Most prokaryotes have simple genomes ranging from less than 1,000 open reading frames (ORFs) to about 4,500 ORFs, and there is minimal post-translational processing of proteins in these systems. Therefore, the maximum number of unique proteins that can be produced is close to the number of ORFs, and these proteomes are relatively simple, limiting the maximum potential complexity of these proteomes. Yeast, the simplest eukaryote, is more complex, with over 6,000 ORFs and moderate amounts of post-translational processing. In contrast, human proteomes are extremely complex with between 35,000 and 50,000 ORFs (not all ORFs have been identified at present), alternative splicing of many mRNAs, and extensive, variable post-translational modifications of many proteins. It is too early in the history of proteome analysis of human cells and tissues to accurately estimate
the total number of structurally and functionally distinct protein components that can be produced from a single genome, but a reasonable estimate is probably on the order of 1 million or more distinct components. The average number of unique protein components that occur at any given time in a single human cell type is similarly not well defined, but estimates of 20,000 to 50,000 or more seem likely for most cells. Wide differences in protein abundance levels (dynamic range) in most cells and organisms further complicate protein profile comparisons because the dynamic range of current separation and detection methods is more limited than the range of protein abundances in most proteomes. The range of protein abundances in even simple prokaryotes will often exceed the dynamic range of protein-profiling methods. Protein copy numbers in a single cell will frequently range from about 10,000,000 copies for major structural proteins such as actin to <100 copies for low-abundance regulatory proteins and some enzymes, for a dynamic range of >105. Biological fluids usually have even wider dynamic ranges than cells and tissues. Human plasma or serum has a few major proteins that comprise more than 90% of the total protein in the sample. Albumin, the most abundant protein, is present at about 40 mg/ml while cytokines and other low-abundance proteins are present at a few pg/ml for a dynamic range of about 1010. The first critical factor in a successful proteome analysis experiment is selection of an experimental design that minimizes protein changes unrelated to the biological question (i.e., protein profile “noise”) that is being addressed. This requires selecting samples for comparison where the only factor that should affect protein levels is the desired experimental parameter, and sample processing conditions must be used that do not introduce artifactual changes in protein profiles. In this context it is important to recognize that most experimental manipulations, including those reflecting a biological question or those caused by poorly controlled experimental parameters, will typically result in quantitative changes of many proteins rather than just a few proteins. Extraneous changes introduced due to faulty experimental design can result in much wasted effort spent in unnecessary protein identification and characterization. But most importantly, it is often difficult or impossible to easily distinguish between observed changes caused by the biologi-
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cal question being tested versus changes caused by experimental design problems. One example of an experiment that would probably have extensive extraneous protein profile noise would be evaluation of protein changes related to tumor progression by comparing different cancer cell lines derived from different-stage tumors. Because most cancer cells have impaired DNA repair and apoptosis control, cell lines are usually aneuploid (having an abnormal number of chromosomes) and have extensive random chromosomal translocations that result in very different basic protein profiles in which many changes may not be related to stage of tumor development. Also, when working with cell lines, it is essential that growth conditions and external signals, such as extent of confluency, pH, nutrient depletion, and pH changes be maintained as consistently as possible to avoid introducing unnecessary changes into protein profiles. Sample harvesting and processing are other key steps, and potential variable artifacts that can occur at these stages include differing extents of solubilization, proteolysis, oxidation, and dephosphorylation. For many studies involving higher organisms, direct analysis of tissue specimens rather than cells in culture frequently has the advantage of more closely reflecting events in the intact organism. However, the presence of multiple cell types in a specimen further increases the number of protein components present in a sample, and if the proportions of cell types change between compared samples, many observed protein changes may be due to the differing proportions of cell types rather than to changes within the targeted cell type. To circumvent or at least minimize this problem, laser capture microdissection, fluorescence activated cell sorting (FACS), and magnetic bead capture are some of the major methods that have been developed to isolate more homogenous cell populations from tissues.
GLOBAL AND TARGETED PROTEOMICS The aim of global proteomics studies is to examine all of the proteins in a cell or tissue simultaneously. For example, a typical global analysis might involve the quantitative comparison of the protein profiles in normal cells and diseased cells, or comparison of a cell line before and after exposure to a drug. The goal is to identify specific proteins connected with the disorder or drug treatment. Such investigations may result in the discovery of new drug targets
and diagnostic markers, provide mechanistic insights, or define critical signaling pathways. For example, protein expression profiling has resulted in the annotation of some of the proteins that are differentially regulated in normal human luminal and myoepithelial breast cells (Page et al., 1999). Such studies are also contributing to the compilation of databases for the study of bladder cancer (Celis et al., 1999) and the human prostate (Nelson et al., 2000), while a number of potential cancer biomarkers have been identified by similar means (Srinivas et al., 2001). Although the value of successfully elucidating cellular and pathological processes on a proteome-wide level is indisputable, the scale of the undertaking is not trivial and becomes more challenging as the complexity of the organism increases, as discussed above. Thus, the ability to effectively analyze proteins on a global scale is subject to the capacities of proteomic technologies for the resolution of a large number of proteins with distinct properties, as well as their detection over a wide range of concentrations. There are multiple alternative methods for enhancing the comprehensiveness of protein profile analyses of complex proteins, including prefractionation of the proteome by various methods, followed by analysis of all fractions. Another strategy for addressing the greater complexity of most proteomes relative to currently available separation and analysis methods is to use targeted proteomics. This approach focuses on a well defined subset of the proteome that can be reproducibly isolated. Reduced complexity is an inherent advantage of this approach. The closer match between protein complexity of many subproteomes and available analytical tools suggests that it may be more efficient and feasible to combine the results of multiple sub-proteomic investigations to define complete proteomes than to employ a direct global proteome analysis approach. Examples of sub-proteomes that are being studied include large macromolecular complexes and cellular machines, specific classes of proteins, and organelles (see Table 22.1.1). Although the same analytical technologies may usually be employed for global and targeted proteomic studies, the latter studies require specific initial strategies to isolate the appropriate sub-proteome components. Frequently, multiple alternative methods are available for isolating particular subsets of the proteome. For example, phosphoproteins can be isolated using antibodies against phosphoamino acids
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Table 22.1.1
Examples of Targeted Proteome Studies
Target
Example
Potential isolation strategies
Macromolecular complexes and cellular machines
Ribosomes, nuclear pore complex, spliceosome complex, p24 complex, transcription factor complexes Proteins of EGFR signal transduction pathway, PDGFβ-receptor signal transduction pathway, all phosphotyrosine-labeled proteins Serine proteases, cysteine proteases, protease subclasses Cell surface glycoproteins, shifts in glycosylation patterns in cancer Secreted into serum-free medium, serum/plasma, other biological fluids Golgi, lysosome, mitochondria, chloroplast, nucleus, plasma membrane
Subcellular fractionation, immunoaffinity columns, transfection of fusion protein with affinity tag Anti-phosphoamino acid antibodies, chemical modification with biotinylated tag
Phosphoproteins
Enzymes Glycoproteins Secreted proteins
Organelle
(Pandey et al., 2000) or by direct chemical modification of the phosphoamino acids using reactive groups coupled to affinity tags (Oda et al., 2001; Molloy and Andrews, 2001; Steen and Mann, 2002). Targeted proteomics is not only appealing in terms of reduced sample complexity, but it also concentrates on unraveling more focused biological questions, such as the components and interactions in cellular machines or key cellular processes and pathways. Protein phosphorylation, for example, is a central feature of signal-transduction pathways and mediates protein-protein interactions associated with cell cycle control. Furthermore, it is primarily protein interactions and multi-protein complexes, rather than proteins in isolation, that mediate biological processes. Global and targeted proteomics therefore define the two major levels of scope for proteome analysis studies. Although there are multiple alternative criteria that can be used to further classify different types of proteome studies, one of the simplest classifications defines the three most common types as: (1) quantitative protein profile comparisons, (2) protein-protein interaction studies, and (3) protein composition analysis. Each type of study can have a global or targeted scope. Overview of Proteome Analysis
Covalent enzyme active site reagents with affinity tag Lectin affinity columns, chemical modification with biotinylated tag Fluid collection
Subcellular fractionation, affinity tag to surface receptors
QUANTITATIVE PROTEIN PROFILING Quantitative protein profile comparisons that ask either targeted or global questions are the most common type of proteomic studies. Profile comparisons between two or more experimental conditions such as normal versus diseased cells or tissues, time courses after drug treatment, or responses to environmental stimuli or stresses can provide highly informative insights into biological processes and diseases.
Two Dimensional Gel Electrophoresis (2-DE) Most of the protein-profiling data currently available have been obtained using 2-DE. The basic method has been used for more than 25 years, and although it has a number of substantial limitations when applied to global protein profiling of complex proteomes (see discussion below; also see Table 22.1.2), it remains the most powerful integrated separation method for proteins. This superior resolution arises from the highly efficient coupling of the two individual separation strategies that constitute 2-DE. Proteins are first separated according to their individual isoelectric points by isoelectric focusing (IEF). The pH gradients necessary for IEF were historically established using soluble carrier ampholytes. Immobilized pH gradients (IPGs) were subsequently developed where the buffering components (immobilines) for the
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Table 22.1.2
Method
Comparison of Quantitative Protein Profiling Methods
Advantages
Disadvantages
Electrophoresis Difficult to automate, some variability Poor separation of large, very small, very acidic, very basic, and hydrophobic proteins Time consuming, technically demanding Resolves only the 1000 to 2000 most abundant proteins in sample Limited protein loads; cannot readily detect low-abundance proteins All of above except variability 2-D DIGE All of above Eliminates gel-to-gel variation, reduces Difficult to excise spots due to shift of labeled relative to unlabeled number of gels required Broader linear dynamic range compared protein Variations in fluorescent labeling to Coomassie or silver stains between samples may cause spot shifts Stable isotope labeling 2-DE
High-resolution, well established method Resolves many post-translationally modified forms of proteins Resolves most alternative spliceoforms Resolves most highly homologous proteins
ICAT
Can be automated Can scale up total protein analyzed to detect lower-abundance proteins Only cysteine-containing peptides are analyzed to reduce complexity Detects large, hydrophobic, very acidic, and very basic proteins Can combine with gel methods to detect ratios of incompletely resolved proteins
Incorporation All ICAT advantages except no of 18O enrichment to reduce complexity Simple method, fewer steps than ICAT All peptides except C-terminal are labeled
Metabolic All of ICAT advantages except no stable isotope enrichment to reduce complexity labeling All proteins and peptides are labeled Small changes in abundance can be accurately determined
Isotopic N-terminal labeling
Similar advantages to 18O method Simplifies interpretation of MS/MS
Limited to comparing two samples at a time Insensitive to post-translational modifications Proteins without cysteine are missed Cannot resolve most spliceoforms or closely homologous proteins Typically does not profile more proteins than 2-D gels (<1500) but different bias–some low-abundance proteins Limited to comparing two samples at a time No affinity method to simplify mixture Only 4-Da separation between peptide pairs Trypsin digestion variations and peptide losses contribute to apparent differences Typically profiles fewer proteins than 2-D gels Only feasible to label simple organisms Limited to comparing two samples at a time No affinity method to simplify mixture Typically profiles less proteins than 2-D gels fewer Similar disadvantages to 18O method continued
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Table 22.1.2
Method
Comparison of Quantitative Protein Profiling Methods, continued
Advantages
Disadvantages
Arrays High-throughput Antibody/ antigen arrays Extensive multiplexing with minimal sample feasible High sensitivity Readily automated
Overview of Proteome Analysis
pH gradient were covalently incorporated into the gel matrix. This resulted in increased reproducibility of 2-D gels and provided greater control over pH gradients. Following IEF, the IPG strip with the focused proteins is placed on the top of an SDS slab gel and proteins are separated in the orthogonal direction based upon size of the polypeptide chain. Proteins are then detected using conventional gel stains, typically colloidal Coomassie, Sypro Ruby, or a silver stain. Digital images are obtained using a scanner or CCD camera, and the intensity of the spots arising from the detection step should reflect the amount of a specific protein present in the sample. For example, a typical quantitative experiment might involve the separation of proteins from an experimental sample on one set of replicate 2-D gels and proteins from a control sample on a parallel set of 2-D gels (Fig. 22.1.1). Specialized 2-D gel image analysis software matches the protein patterns by correcting for gel-to-gel variations and compares matched spot intensities. Proteins of interest, that is, those showing substantial changes in amount, are then excised from the gel and subjected to in-gel proteolytic digestion. The resulting peptides are then analyzed by mass spectrometry (MS) in order to identify the proteins from which they were derived using appropriate database search algorithms. One advantage of 2-DE is that in most cases even a single change in a side-chain charge can be detected in the IEF dimension. In addition, changes in total protein mass that are larger than 5% to 10% can usually be readily detected. Hence 2-DE can detect many but not all changes in post-translational modifications, shifts between protein isoforms that result from alternative mRNA splicing, and proteolytic processing events. Another advantage of this
Nascent technology Preparation of reagents with appropriate specificities and affinities very time consuming Limited by range of available receptor molecules Not discovery-based, i.e., must know that target may be present High background signals (non-specific interactions) possible when biological fluids are analyzed
method is that 2-DE comparisons are not limited to simultaneous comparisons of only two or three samples. This is particularly an advantage for time-course studies or studies of many different related samples. However, 2-DE has a number of disadvantages as well (Table 22.1.2), and these limitations have encouraged development of alternative non-gel-based protein profiling methods. The most serious limitations with 2-DE are: (1) the relatively narrow dynamic range of detection, which is usually 2 to 3 orders of magnitude; (2) the limited number of proteins that can typically be separated and visualized using protein stains, usually about 1,000 to 1,500; (3) the fact that only the most abundant proteins in the sample are visualized; (4) the fact that multiple classes of proteins are poorly recovered or entirely lost; and (5) the observation that, although IPG gels have improved reproducibility compared with older soluble ampholyte-based systems, substantial gel-to-gel variation still exists. Protein classes that are largely incompatible with 2-DE include: any proteins insoluble in the extraction buffer used, proteins larger than ∼100 kDa, proteins/peptides smaller than about 6 to 10 kDa, very hydrophobic proteins, and very acidic and very basic proteins. The dynamic range of proteins displayed on 2-D gels is limited by a combination of the maximum feasible total protein load and the sensitivity of the stain used. In addition, in some cases, low-abundance proteins are obscured by high-abundance proteins. Some of the most promising strategies for expanding resolving power of 2-D gels employ various prefractionation methods to increase the number of proteins separated and/or to detect less abundant proteins. Approaches include sequential extractions with increasingly stronger solubilization solutions (Molloy et al.,
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A
B sample A (experimental)
sample B (reference)
sample A (experimental)
sample B (reference)
2-D electrophoresis
2-D electrophoresis
label with cyanine dye A
label with cyanine dye B
image analysis
image analysis
mix samples
separation on single 2-D gel match spots and compare intensities
image analysis (compare signal intensities of fluors) excise proteins of interest excise proteins of interest in-gel digestion
protein identification by MS
in-gel digestion
protein identification by MS
Figure 22.1.1 Quantitative protein profiling by 2-D gel electrophoresis. (A) Conventional 2-D gel electrophoresis (2-DE). Samples to be compared could be extracts from cells, tissues, bodily fluids or single-cell organisms. In most protein profiling studies, two or more samples are compared, i.e, the experimental (diseased/altered) and control (reference) conditions. Extracts are separated on one or more of various broad- and narrow-pH-range 2-D gels, and after separation and staining, computer-assisted image analysis is used to match spots and compare spot intensities to detect significant quantitative changes. Proteins of interest are excised, digested, and identified by mass spectrometry. (B) 2-D differential gel electrophoresis (2-D DIGE). Individual samples are initially labeled with distinct cyanine dyes before being combined and separated on a single 2-D gel. Signal intensities of the different fluors are compared during image analysis. Proteins of interest are excised, digested, and identified by mass spectrometry.
1998) and subcellular fractionation (Huber et al., 1996; Cordwell et al., 2000). Various chromatographic methods have also been used. However, all these methods result in substantial cross-contamination between fractions, and variable cross-contamination of many components between adjacent fractions severely complicates quantitative comparisons of protein profiles. A high-resolution method that interfaces well with 2-DE is microscale solution isoelectrofocusing (µsol-IEF), which has been applied to E. coli extracts, serum proteins, and human cancer cell extracts (Zuo and Speicher, 2000, 2002; Zuo et al., 2001). The µsol-IEF device incorporates a preparative electrophoresis method with key advantages over other preparative electrophoresis methods. It is simpler, can cover the full pH range, has smaller sample
volumes that do not require subsequent concentration, and, most importantly, it yields highresolution separations between fractions.
Two-Dimensional Differential Gel Electrophoresis (2-D DIGE) This is a variation of the 2-DE technique that has several advantages over the conventional method (Table 22.1.2). The 2-D DIGE method (Unlu et al., 1997) involves separately labeling two or three different protein extracts with distinct fluorophores, which are cyanine dyes (Amersham Biosciences), by covalently modifying lysine residues in the proteins (see UNIT 22.2). These dyes have positive charges to replace the loss of the positive charge on the lysine side chain, and the masses of the dyes are similar to each other. Samples are reacted under conditions that label only a few lysines
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per molecule. As long as the extent of reaction is similar between the samples to be compared, the mass shift will be uniform and the pI should be essentially the same as the unreacted protein. The fluorophores have distinguishable spectra, and therefore the individual fluorescently labeled protein samples are combined and separated by conventional 2-DE on a single gel (Fig. 22.1.1). The proteins from each sample can then be visualized separately by imaging, using appropriate excitation and emission wavelengths for the different dyes. The differences between the quantities of the individual proteins from each sample can then be determined using specialized 2-D image-analysis software designed for this system. The detection of differences between two protein samples is greatly simplified by 2-D DIGE in comparison with conventional 2-DE because both protein samples to be compared are separated on a single gel, which eliminates gel-to-gel variation. In addition to the greatly improved spot matching resulting from use of a single gel, the number of parallel and replicate gels required to obtain reliable results is greatly reduced. Furthermore, fluorescence imparts the ability to detect proteins over a much broader linear dynamic range of concentrations than the colorimetric gel stains (Patton, 2000). One potential limitation of this method is that excision of spots of interest for identification by MS is not as straightforward as for conventional 2-D gels. Coomassie or silverstained gels can be readily excised manually or using a spot cutting robot because protein spots are visible. However, excision of fluorescent spots is more difficult. One solution is to use an excision robot that can “see” fluorescent spots. Another potential solution is to produce a hard-copy image that exactly matches the gel size and place it below the gel to guide excision. However, excision remains unreliable with these methods because, with minimal labeling conditions, only a few percent of a specific protein is labeled and this minor fluorescent population is generally shifted to a slightly higher mass position due to the mass of the covalently bound dye. Therefore, the position of the bulk amount of unlabeled protein may be estimated as being shifted about one spot diameter down (lower mass), but this strategy can lead to excision of a protein other than the one of interest (Patton, 2000). Another potential resolution to this problem is to match 2-D DIGE gel patterns with corresponding preparative colloidal Coomassie 2-D gel patterns for spot excision from the colloidal Coomassie gel
(Tonge et al., 2001). However, in this study about 40% of the spots of interest could not be reliably identified by this method due to the lower sensitivity of the colloidal Coomassie stain.
MS-based Protein Profiling Methods Using Stable Isotope-Based Methods MS analysis of tryptic peptides has emerged as the predominant method for identifying proteins for most proteome studies, including the gel-based methods described above. Furthermore, the limitations of 2-D gels, the high sensitivity and high throughput of current mass spectrometers, and the potential for automating sample processing prior to MS has led to development of multiple non-gel-based protein profiling methods as alternatives to 2-DE. Depending upon the complexity of the initial protein samples to be analyzed, most methods use one, two, or three dimensions of chromatographic separations followed by MS analysis. Some methods separate and analyze the intact proteins, or more commonly tryptic peptides of the entire protein mixture are produced, separated, and analyzed by MS. However, MS is not intrinsically quantitative and this limitation is most commonly circumvented by introducing a stable isotope (e.g., 15N, 13C, 2H) into one of two samples to be compared (De Leenheer and Thienpont, 1992). The two protein samples are then combined, proteolyzed, separated, and analyzed by MS. The labeling of one sample generates internal standards for the individual peptides, since each protein has a heavy isotope and natural abundance isotope form. Furthermore, owing to the generally identical physicochemical characteristics shared by the two isotopic forms of the same peptide, relative abundance is usually not altered by the separation step. The corresponding peptides from the two samples will differ in mass by the number of heavy atoms incorporated in the stable isotope– labeling experiment, and comparison of peak heights for each peptide pair should reflect the relative abundances of the protein in the two different samples. Isotope-coded affinity tag (ICAT) One of the best known and most versatile methods of stable isotope labeling for protein profiling studies is the ICAT method. Two forms of a cysteine-specific labeling reagent with a biotin affinity tag, known respectively as heavy and light ICAT, are used in this procedure (Gygi et al., 1999). Heavy ICAT is distinguished by the presence of eight deute-
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A
XX
XX X = deuterium (heavy ICAT) X = hydrogen (light ICAT)
XX biotin
XX linker
thiol reactive group
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reference
react sample with heavy ICAT
react sample with light ICAT
mix protein samples
protease digestion
avidin affinity column
step elution from ion exchange column
Intensity
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8 Da
1 2 3 4 MS scan of multiple peptides for isotope ratios
MS/MS scan for protein identification
Figure 22.1.2 Isotope-coded affinity tag (ICAT) method. (A) The ICAT reagent consists of a biotin affinity tag that binds to avidin, a linker region incorporating deuterium (heavy ICAT reagent) or hydrogen (light ICAT reagent) atoms, as well as a thiol-specific reactive group that can covalently attach to a cysteine residue. (B) Proteins in the experimental and reference samples are derivatized with the different isotopic forms of the ICAT reagent. The two samples are then combined and subjected to proteolysis and modified cysteine-containing peptides are enriched by avidin affinity chromatography. An optional step that can greatly expand the number of detected proteins is to apply the peptide mixture from the avidin column to an ion-exchange column that is eluted with multiple steps of increasing ionic strength. Fractions are subsequently applied to a microcapillary reversed-phase (µLC) column interfaced directly with a mass spectrometer that performs scans in MS and MS/MS modes. In the MS spectrum, the ratio of the peak areas for each peptide pair (gray bars represent the heavy ICAT peptides) indicates the relative abundance of the corresponding protein in the initial samples (left lower panel). The MS/MS spectra (right lower panel) are used to determine protein identities by database searches.
rium atoms in the reagent’s linker region (Fig. 22.1.2A), while in light ICAT this portion of the reagent contains hydrogen atoms instead. This creates a mass difference of 8 Da between the two ICAT forms. To compare two protein
samples (Fig. 22.1.2B), each sample is labeled with a different form of the ICAT reagent under conditions where all cysteines are derivatized (end-point labeling), and the samples are combined and subjected to proteolysis. ICAT-la-
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beled peptides are recovered using avidin affinity chromatography, taking advantage of the biotin tag. This greatly simplifies the peptide mixture because cysteine is a relatively low-frequency amino acid in proteins and therefore only a few percent of the peptides in the digest are tagged. The enriched peptides are subsequently applied to a microcapillary reversedphase HPLC (µLC) column, from which light and heavy ICAT-labeled forms of the same peptide, that is, peptide pairs, coelute. The µLC column is usually interfaced directly with a mass spectrometer through an electrospray interface and a combination of MS (intact peptide masses) and MS/MS (fragmentation of specific peptides to produce daughter ion spectra) is automatically performed. The ratio of the peak areas for each peptide pair is indicative of the relative abundance of the corresponding protein in the initial samples. The MS/MS spectra are used to search databases using appropriate algorithms to correlate the observed spectra with hypothetical MS/MS spectra from sequences in the database to identify the protein. Simple protein mixtures, such as from targeted proteome analyses, might be analyzed using a single µLC reversed-phase gradient. Alternatively, an increased number of proteins can be identified by adding an ion-exchange separation between the avidin affinity step and the µLC column (Fig. 22.1.2B). The ICAT method is an attractive technique for quantitative protein profiling because it can be used to label virtually any type of sample, while metabolic isotopic labeling (see below) has much more limited utility. The ICAT method is an interesting alternative to 2-DE, and in contrast to 2-DE, the amount of protein that can be processed is theoretically unlimited. This is an important strength of the approach since increasing the amount of starting material improves analysis of lower-abundance proteins. It is also possible to enrich such proteins by prefractionation prior to derivatization with ICAT. As noted above, the tagging and isolation of only cysteine residues greatly reduces the peptide mixture complexity, but this can also be regarded as a disadvantage because proteins that do not contain cysteine are not detected by this method, although reagents with different reactivities could be synthesized (Gygi et al., 2000). Another potential problem lies in the avidin affinity step during which nonspecific interactions between the purification matrix and the peptides may occur. This could potentially compromise the recovery of peptides originating from low-copy proteins (Graves
and Haystead, 2002). Furthermore, the large ICAT reagent (∼500 Da) remains attached to the peptides throughout the procedure. This can cause difficulties with respect to database searching algorithms used to identify proteins using the MS/MS spectra, particularly in the case of peptides that comprise less than 7 residues (Gygi et al., 2000). Finally, although the ICAT method is quite promising, it and other multidimensional chromatography methods currently require many hours or days of mass spectrometer instrument time and computer time, and in only a few cases have more proteins been quantitatively profiled than the 1,000 to 1,500 proteins that can be separated in a single high-resolution 2-D gel. Incorporation of 18O A second stable isotope–labeling method for proteome studies uses 18O-enriched water during the protease digestion of protein samples (Yao et al, 2001). When a protein sample is digested with trypsin in 18O-enriched water, the hydrolysis of the peptide bond adds two 18O molecules to the C-terminal amino acid. The second sample in a comparison experiment is digested in unenriched water, and the trypsin digests are then mixed before separation and MS analysis is performed as described for the method above. In this case, the corresponding peptides differ by 4 Da, and therefore the separation of isotopic envelopes is much less than with the ICAT reagent. This is a very simple procedure that involves fewer steps than the ICAT method. However, in addition to the reduced mass separation of the paired peptides, H2[18O] is expensive, and variations in degree of digestion of the two samples could contribute to apparent quantitative differences. Metabolic stable isotope labeling A third stable isotope labeling method is to metabolically label specimens by growing simple organisms in medium enriched with a stable isotope. For example, in one study, one pool of yeast cells was grown using medium containing natural isotopes (i.e., 14N) while the second pool of yeast cells, which were unable to express the cyclin CLN2, were grown using medium enriched in 15N (Oda et al., 1999). Following a suitable period of growth, the two samples were combined prior to extraction of proteins of interest. The proteins were separated by RP-HPLC and subsequently by 1-D SDS-PAGE. Peptides of selected proteins were generated by in-gel digestion and analyzed by MS to determine their relative abundance and
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identity. Changes in abundance of more than 20% could be ascertained using this approach. Furthermore, it was possible to establish the relative abundance and identity of each of two different proteins present in the same band of the SDS-PAGE gel. Although this specific study used targeted proteomics and 1-D SDS gels to separate proteins of interest, global protein profiles could have been obtained by using multidimensional chromatography methods followed by MS analysis. An alternative strategy for metabolic stable isotope labeling uses rare isotope–depleted instead of stable isotope–enriched media. For example, in one study a batch of E. coli cells was grown on normal medium while a second batch of cells was grown on medium depleted of rare isotopes (Pasa-Tolic et al., 1999). The depleted medium was also supplemented with CdCl2 in order to study the E. coli cadmium stress response. After metabolic labeling, the two batches of cells were combined, lysed, and dialyzed. The expression ratios for 200 proteins were determined using capillary IEF combined with Fourier transform ion cyclotron resonance mass spectrometry (CIEF-FTICR). A key strength of FTICR MS instruments is that they generate data of greater accuracy and resolution than other mass spectrometers. One of the main drawbacks associated with metabolic isotope labeling is that it is not widely applicable, because only cells from simple organisms are generally amenable to this technique. Another disadvantage is that even for simple organisms, stable isotope–enriched media are relatively expensive. Isotopic N-terminal labeling A fourth differential isotope labeling method uses positively charged succinimide esters to label the N-terminals of tryptic peptides produced from protein mixtures. In a recent study, this labeling method was combined with 2-DE to increase reliability of quantitative comparisons (Munchbach et al., 2000). Two protein samples were obtained from E. coli cells supplied with either unlimited or restricted levels of a carbon source, the samples were separated individually by 2-DE, and spots of interest were excised. Following succinylation to modify lysine side chains and protease digestion, peptides originating from each of the two samples were differentially labeled by Nterminal nicotinylation with either 1-([H4]nicotinoyloxy)succinimide ester (H4-Nic-NHS) or 1-([D4]nicotinoyloxy)succinimide ester (D4-Nic-NHS). The two peptide samples were
then combined and analyzed by MS and MS/MS to determine the relative abundances of the proteins and their identities. This strategy was shown to afford certain advantages over approaches that solely rely on Coomassiestained 2-D gels for relative quantification. With this method, the determination of isotopic ratios is not limited by stain saturation and it is possible to compare spots that are too weakly stained for quantification by 2-D analysis software. Furthermore, individual ratios for two distinct proteins that migrate to the same spot on the 2-D gel could be ascertained. In contrast to ICAT, this procedure does not preclude the analysis of cysteine-free proteins. Although 2DE formed part of the protocol described by Munchbach and colleagues, the labeling method could conceivably be applied to complex peptide mixtures produced by solution digestion with proteases, and the labeled peptides would then be separated by multidimensional chromatography followed by MS analysis, which is similar to the strategies described above for the other stable isotope labeling methods.
Antibody/Antigen Arrays A protein array consists of numerous specific receptor or capture proteins (usually antibodies) that are immobilized or printed on a surface such as a glass slide in a regular pattern (also see Protein-Protein Interaction Studies, below). Proteins in a sample are captured by the receptors, and this association can be detected by a number of alternative methods. While conceptually similar to nucleotide microarrays, protein arrays present more complex challenges including: (1) maintaining the printed proteins in native, functional states; (2) development of hundreds or thousands of capture reagents with binding affinities in an appropriate range similar to other capture reagent affinities used in the same array; (3) avoiding nonspecific reactivity to the capture reagent and nonspecific interactions with specifically associating targets; and (4) development of quantitative detection methods with an adequate linear range. Of particular importance is the degree of multiplexing that the array allows, that is, the number of different molecules that can be assayed simultaneously. An important factor that may limit extent of multiplexing is the ability to obtain capture reagents with similar affinities, or, alternatively, to develop assays that accommodate a broader range of affinities while maintaining an adequate linear range of detection.
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Many array-based methods for protein quantification are variations of the ELISA (enzyme-linked immunosorbent assay) technique. One of the most common approaches is illustrated by a study utilizing a simple array of seven different antibodies to cytokines spotted onto the bottoms of wells in polystyrene plates (Moody et al., 2001). Cytokines captured by these bound ligands were then analyzed using biotinylated detection antibodies that bound to different epitopes than the capture antibodies. A streptavidin–horseradish peroxidase (SAHRP) conjugate and a chemiluminescent substrate were used, and signals were detected using a CCD camera. This assay was used to measure the changes in levels of cytokines secreted into growth medium by peripheral blood mononuclear cells (PBMCs) treated with lipopolysaccharide (LPS), and by THP-1 cells treated with LPS alone or in combination with the antiinflammatory drug dexamethasone. An advantage of chemiluminescence is its high sensitivity, which allows the measurement of cytokines at concentrations as low as 1 pg/ml (Moody et al., 2001). Another protein array method immobilizes antibodies or antigens on appropriate surfaces and uses differentially fluorescently-labeled samples to directly detect protein binding to the immobilized reagents. In one representative study, either antibodies (to detect antigens) or antigens (to detect antibodies) were immobilized on a microscope slide, and proteins in individual samples to be compared were labeled with distinct cyanine dyes (Haab et al., 2001), analogous to the detection strategies used for some nucleotide arrays or for 2-D DIGE. The fluorescently-labeled samples were then mixed and applied to the appropriate array. The different intensities of the distinct fluorescent signals at each spot on the array reflected the relative abundance of the protein captured at that position from the different samples. In this study, antigen arrays (i.e., where antigens acted as capture reagents) were associated with greater accuracy than antibody arrays. It was proposed that this was probably due to differences in labeling efficiency and stability of the different types of fluorescently tagged molecules; i.e., since the overall structures of antibodies are similar, they can be similarly labeled at lysine residues in their respective Fc regions. In contrast, many antigens may not have easily accessible groups for derivatization with the fluorescent dyes that neither inactivate the protein nor interfere with antibody binding, and the greater stability of antibodies in solution,
relative to many antigens, might also enhance performance of antigen arrays. Results from certain antigen-antibody pairs demonstrated that arrays could be sufficiently sensitive to detect certain proteins at physiologically relevant levels (Haab et al., 2001). However, when serum was added to mixtures of purified antigens, antibody arrays exhibited reduced accuracy, apparently due to higher background, which led to the recommendation that sample protein concentrations be limited to <1 mg/ml for optimal performance. Although this concentration limit would currently preclude facile detection of lower-abundance proteins in bodily fluids such as serum, alternative strategies may prove to be beneficial, such as prefractionation of samples and enhanced blocking of nonspecific adsorption (Haab et al., 2001). Although acceptable sample-labeling methods must be developed, one potential advantage of this method, compared with the dual-antibody method described above, is that only one antibody with appropriate affinity is needed per antigen. An alternative method of enhancing fluorescent signals in antibody assays (Schweitzer et al., 2000) is to use the rolling circle amplification (RCA) method (Lizardi et al., 1998). Proteins captured by immobilized antibodies are subsequently coupled to antibody-DNA conjugates. The latter each consist of the 5′ end of an oligonucleotide primer attached to an immunoglobulin. A circular DNA molecule is added that hybridizes to the primer. In the presence of DNA polymerase and nucleotides, rolling circle amplification takes place, generating a long single DNA molecule consisting of tandemly linked copies of the circle. This molecule remains bound to the antibody and can be detected by hybridization of multiple fluorescently-labeled complementary oligonucleotide probes or by supplying fluorescently-labeled nucleotides for incorporation into the amplified DNA. Using such an “immunoRCA” sandwich assay an improvement of up to 2 to 3 orders of magnitude in sensitivity compared with conventional ELISAs was obtained, and the relative abundances of 51 cytokines secreted into growth medium by Langerhans cells were analyzed (Schweitzer et al., 2002). Biosensor antibody-antigen arrays differ most markedly from the arrays described above in terms of the detection mode. In the case of biosensors, the layer to which receptor molecules are linked undergoes a physicochemical change in response to analyte capture. This change is converted to an electrical signal, al-
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lowing the amount of analyte bound to be determined. An example of this type of array is the piezoelectric (Pz) crystal device, also referred to as the quartz crystal microbalance (QCM), consisting of receptor molecules coupled to an oscillating metal-coated crystal surface (Lin et al., 2000). The oscillation frequency is determined by both the mass of the crystal and by the mass of proteins coupled to its surface. The capture of an analyte increases the mass of bound proteins, which in turn decreases the crystal’s oscillation frequency. The change in mass is thus transduced via the change in oscillation frequency into an electrical signal, allowing the quantity of bound analyte to be ascertained. A QCM-based method has been used, for example, to measure serum allergen-specific IgE levels, although the results were not always consistent with those obtained using a commercially available conventional IgE immunoassay test kit (Su et al., 2000a). In general, the main problems associated with the QCM approach relate to non-specific binding and low sensitivity for small molecules (small mass changes). However, modifications of the basic QCM method are being developed to address these issues (Su et al., 2000b). Biosensor arrays that exploit other physico-chemical changes are reviewed elsewhere (Jenkins and Pennington, 2001). Antibody/antigen array technology represents one of the most promising recent developments in protein profiling methods. Although most studies are at the proof-of-principle stage and further improvements are needed, antibody (or antibody mimic) arrays are likely to ultimately offer the highest-sensitivity, highest-throughput methods for quantitatively comparing large numbers of proteins in complex protein extracts. A major obstacle to widespread adoption of the antibody array approach for a comprehensive range of proteomic studies is its dependence on analyte-specific receptor molecules that must be individually prepared and tested for appropriate reactivity and affinity properties. It is particularly time-consuming and expensive to prepare each immunological reagent by immunizing animals and then performing affinity purification of resulting antibodies. One alternative to this approach is to use in vitro selection methods to obtain antibodies or antibody mimics with the desired specificity and affinity (Holt et al., 2000; Lohse and Wright 2001).
PROTEIN-PROTEIN INTERACTION STUDIES The second major type of proteome analysis is systematic analysis of protein-protein interactions. These studies often complement data obtained from quantitative protein profile comparisons and can overlap targeted quantitative protein profile comparisons that examine changes in large protein complexes and molecular machines. As with quantitative protein profiling, protein-protein interactions can be pursued either with a global scope—e.g., identifying all binary protein-protein interactions in yeast—or with a targeted scope—e.g., identifying all proteins that interact with protein X, followed by all proteins that interact with protein X’s binding partners. Most cellular processes are mediated by multi-protein complexes or pathways comprising interacting proteins. Hence, systematic identification of many protein-protein interactions is expected to provide new insights into cellular pathways and networks. Similarly, identification of which proteins of known function associate with a protein of unknown function can provide insight into the latter’s role and the underlying cellular mechanisms involved. For these reasons, protein-protein interaction maps for a number of different organisms are being constructed (Auerbach, 2002). For example, over 1,200 interactions between Helicobacter pylori proteins have been identified, connecting 46.6% of the organism’s genome (Rain et al., 2001).
Immunoaffinity and Biochemical Isolation of Protein Complexes The most direct method of identifying interacting proteins is to isolate a macromolecular complex, followed by identification of all components using MS methods. A number of alternative conventional protein biochemical methods can be used to isolate macromolecular complexes, including subcellular fractionation for very large complexes and cellular machines such as ribosomes, the proteosome, and the nuclear pore complex. For many macromolecular complexes, one of the simplest and most common isolation methods is immunoaffinity purification, if an appropriate antibody to one of the interacting proteins is available. Typically, the antibody is covalently coupled to a chromatographic resin to minimize interference from the antibody during later MS identification of components. The complex is then extracted in an appropriate buffer, bound to the immobilized antibody, washed to remove nonspecifically bound proteins, and eluted from the
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column. Proteins in the complex can be identified by running 1-D gels followed by identification of stained bands by MS methods after in-gel digestion. Alternatively, the proteins in the solution eluted from the affinity column can be identified directly by protease digestion of the protein mixture in solution followed by microcapillary reversed-phase chromatography interfaced with a tandem mass spectrometer (µLC-MS/MS). The reversed-phase separation can be used by itself or in combination with prior batchwise ion-exchange column elution to produce a multidimensional chromatography method capable of separating >100 components in large macromolecular complexes and cellular machines such as the ribosome (McCormack et al., 1997; Link et al., 1999). The chromatography and MS methods are similar to those described above for non-gel-based quantitative protein profiling methods, except that no stable isotopes are needed here because this type of study typically does not involve quantitative comparisons.
Protein Complex Isolation Using Engineered Affinity Tags
Overview of Proteome Analysis
An alternative to immunoaffinity isolation of macromolecular complexes is to engineer one or preferably two affinity tags into a protein of interest, followed by transfection of the fusion protein into an appropriate host cell. Two recent large-scale studies in S. cerevisiae demonstrate the utility of systematic global analyses of protein-protein interactions by isolating complexes (Gavin et al., 2002; Ho et al., 2002). In these studies, DNA encoding tagged (bait) proteins was introduced into yeast cells. Complexes that formed between bait fusion proteins and other cellular proteins were isolated using the affinity tag, and constituent proteins in the complexes were identified using MS. This approach allowed Gavin and colleagues to characterize 232 multi-protein complexes and to propose the functions of 231 proteins whose individual roles had not been previously ascertained (Gavin et al., 2002). The extent of novel information and insights into cellular processes generated by these studies is remarkable; although a significant proportion of false positives were encountered and some previously characterized interactions were not identified (Gavin et al., 2002; Ho et al., 2002). However, this simply demonstrates the need for independent validation of most observations from large scale proteomics studies. Actually, one of the biggest challenges with characterization of either immunoaffinity or
affinity-tag isolated protein complexes is to distinguish moderate-affinity physiological interactions from nonspecific interactions and other false positives. Frequently, the specificity of high-affinity interactions can be tested by increasing the stringency and/or number of wash steps prior to elution of the complex. However, the use of more extensive washes or more stringent wash buffers is likely to dissociate moderate-affinity interactions. Multiple alternative criteria and controls may need to be used to help determine which proteins are “sticky” and will be observed in isolates of multiple different complexes, and which are moderate-affinity specific interactions. Usually the best method to ultimately distinguish specific and nonspecific interactions is to perform reciprocal binding or reciprocal immunoprecipitation experiments to demonstrate that all putative interacting partners actually interact.
Protein-Protein Interaction Arrays Protein arrays may also be used to define interactions between proteins. Affinity-tagged proteins can be purified following expression from cloned ORFs and screened in an array format. In one study, 5,800 glutathione S-transferase (GST)-ORF constructs were cloned, expressed and affinity purified prior to being printed onto slides, creating a yeast proteome microarray that comprised 80% of yeast proteins (Zhu et al., 2001). The arrayed fusions were then probed with biotinylated calmodulin and interactions were detected using cyaninelabeled streptavidin. This led to the identification of many previously known as well as unknown calmodulin-interacting proteins. The proteome array was also probed with biotinylated lipids to identify phospholipid-interacting proteins, demonstrating that the method may be used to ascertain function by determining interactions between proteins and other types of biomolecules. Proteome arrays or arrays comprising particular classes of proteins may also be screened for specific biochemical activities (Martzen et al., 1999; Zhu 2000). Other approaches for identifying protein-protein interactions include phage display screens (Hufton et al., 1999; Zozulya et al., 1999).
Yeast Two-Hybrid Screens A common, simple, gene-based assay for protein-protein interactions is the yeast two-hybrid system (Fields and Song, 1989). In this case, a “bait” is constructed consisting of a protein or protein domain of interest fused to the DNA-binding domain of a transcription
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factor, while a potential interacting partner or “prey” is fused to the activation domain of the transcription factor. Interactions between the two test proteins, following the mating of bait- and prey-expressing clones are indicated by the activation of a reporter gene, since bait-prey complex formation brings the DNA-binding domain and activation domain into close proximity. Two forms of the yeast two-hybrid system are currently employed for proteomic studies (Auerbach et al., 2002). The array or matrix approach involves mating each yeast clone expressing one bait protein with individual arrayed colonies that each express different prey. One of the benefits associated with this approach is that false positives may be quickly identified, leading to the assurance that the system is working correctly. If, for example, it is observed that a specific prey protein interacts with every bait tested, the result is probably indicative of a false positive. One of the disadvantages of the approach arises from the incompatibility between yeast and certain full-length proteins from other organisms. Such proteins, for example, may be degraded, may be toxic to the yeast cells, may not fold properly, or may lack post-translational modifications that are essential for function. The second large-scale yeast two-hybrid approach, the library screening method, entails the production of a library comprising cells expressing different prey. The library is divided into pools, each of which is mated with individual yeast clones expressing one of the bait proteins. A key strength of this approach is that it enables the analysis of full-length proteins as well as those expressed from random ORF fragments. Expression from the latter may generate protein moieties that are compatible with yeast cells, even if this is not the case for the full-length proteins from which they are derived. In this system, DNA encoding positive prey has to be isolated from positive clones and sequenced to identify the interacting proteins, in contrast to the array approach, where prey can be identified according to their position in the array. The library screening method is more expensive and time consuming than the arraybased technique because it entails analysis of a greater number of clones. In general, yeast two-hybrid studies are not suitable for hydrophobic transmembrane proteins or transcription factors. Adaptations of the yeast two hybrid system and other genetic screening methods have been devised to address these shortcomings (Auerbach, 2002). Also of concern is the substantial number of
false positives that are often observed, and the fact that the library screening method is likely to produce more false positives than the array approach (Auerbach et al., 2002).
PROTEIN COMPOSITION ANALYSIS The third major type of proteome analyses involves the detection and identification of as many protein components as possible in a simple to moderately complex proteome or subproteome. The distinction relative to quantitative protein profiling is that in protein composition studies, comparisons of two or more samples and quantitation of protein levels are not involved. Instead, the goal is to attempt to define the total protein complement of a proteome such as that of prokaryotes or yeast, or all the proteins present in a cellular fraction or compartment such as mitochondria. There is some overlap with analysis of protein-protein interactions because compositional analysis of all proteins in a ribosome or large macromolecular complex will contribute to our knowledge of interacting complexes, although a simple compositional analysis of large complexes will not define the specific proteins that directly interact within the ribosome or complex. The preferred analysis methods for composition proteomics are various related multidimensional chromatographic methods coupled to tandem mass spectrometry, an approach that has been referred to as multidimensional protein identification technology or “MudPIT” (Washburn et al., 2001; see Fig. 22.1.3). The process recently has been automated and a detection dynamic range of about 104 has been demonstrated (Wolters et al., 2001). It should be emphasized that the non-gel-based separation and protein detection techniques that are suitable for compositional analysis are also suitable for quantitative protein profiling. The major difference is that stable isotopes are not required for compositional studies because quantitation is not needed. Conversely, the MudPIT method, which was initially used for qualitative descriptions of compositions of proteomes, can be readily made into a quantitative method by using stable isotope labeling (Washburn et al., 2002). Typically, 2-D gels are not the best choice for composition studies because quantitative comparisons of many different samples, which are the best applications for 2-D gels, are not needed. Most importantly, if 2-D gels are used for compositional studies, all detectable spots would need to be digested and
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protein sample
denature and alkylate
solution digestion with immobilized trypsin
step elution linear elution (ionic strength) (ethanol)
SCX
RP
µLC MS/MS
Figure 22.1.3 Schematic representation of multi-dimensional protein identification technology (MudPIT). Proteins are denatured, alkylated, and digested prior to being loaded onto a biphasic microcapillary column containing strong cation exchange (SCX) and reverse phase (RP) matrices. Ionic strength is increased to elute a fraction of peptides from the SCX phase onto the RP matrix. Peptides are recovered from the latter using a linear ethanol gradient and subsequently analyzed by µLC MS/MS in order to identify them. The process is iterative, ionic strength being increased (step elution) to displace the next fraction of peptides from the SCX matrix to the RP portion of the column.
identified by MS methods after completing the 2-D gels. In contrast, non-gel-based methods include identification of all resolved proteins as an integral part of the analysis.
SUPPORTING TECHNOLOGIES
Overview of Proteome Analysis
To provide an introduction to the large and rapidly changing field of proteomics, this overview has focused on the scope of proteome analyses, the three major types of studies that are most commonly pursued in proteome analyses, and some of the major separation and MS-based identification techniques. However, it should be stressed that there are many other critical areas of proteomics that could be examined in similar depth. The subjects discussed above, as well as others, will be topics of future detailed units in this chapter. One area of rapid growth is automation, which will positively impact most proteome methods by improving throughput and reproducibility. Some for-profit proteomics ventures have already invested heavily in extensive proprietary automation. In addition, automation provided by commercial vendors is a major growth area. Although very little progress has been made in commercial products for automating gel electrophoresis, substantial progress has been made in most other areas of proteomics, including robotic spot cutters to excise spots and bands from gels, robots for 96-well plate processing including antibody arrays, protein-protein interaction screens. and in-gel digests, as well as automation for conducting MudPIT (Wolters et al., 2001) and similar chromatographic separations. Another major growth area encompasses bioinformatics including software, databases, and computer hardware. It is quite clear that
proteomics, like genomics, involves generation of very large datasets, which must be organized, stored, and made accessible in logical ways (Harry et al. 2000). One pressing need is to develop databases that will handle large datasets with diverse types of data. In addition, while further progress in all areas of proteomics-related computational informatics would be advantageous, there is a particular need for improved databases and data-mining tools to improve the efficiency of follow up in silico investigations and further experimental design after initial protein identifications are made using MS data.
CONCLUSION AND FUTURE PERSPECTIVES An interesting perspective on the sequencing of the human genome is that its completion is not an end to the project, but instead, is the beginning of the real task of understanding what the genome means. In other words, how are different proteomes regulated and how do these different proteomes produce observed phenotypes? In the short time since the conceptualization of the proteome, the information that has emerged as a result of proteomic studies, particularly targeted protein profiling, compositional studies, and global interaction screens, has already been quite impressive. One can expect that proteomics will continue to provide major insights into diseases and biological mechanisms, including identification of genes implicated in complex diseases such as cancer that will lead to new diagnostic and therapeutic targets. Finally an interesting question is “which proteome technology(ies) will be used 5 years from now?” Proteomics has both rejuvenated
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older technologies, especially 2-DE and the yeast two-hybrid system, and stimulated rapid progress in newer fields, particularly mass spectrometry, software, automation, and nanotechnology. Recent progress in antibody arrays has been particularly intriguing. However, there are a nearly infinite number of proteomes for each genome and each human gene can produce on average about 20 different protein components. In addition, unlike nucleic acids, proteins exhibit a wide range of properties and behavior. Therefore, it seems unlikely that any single method or technology platform will be able to best address most proteome analysis problems within the near future. Instead it is more likely that all of the major current analytical technologies including 1-D and 2-D gels, multidimensional chromatography coupled with MS instruments, and antibody arrays will continue to play important and often complementary roles in proteome analysis studies for the near future.
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Yates, III, J.R. 1999. Direct analysis of protein complexes using mass spectrometry. Nature Biotechnol. 17:676-82. Lizardi, P.M., Huang, X., Zhu, Z., Bray-Ward, P., Thomas, D.C., and Ward, D.C. 1998. Mutation detection and single-molecule counting using isothermal rolling-circle amplification. Nature Genet. 19:225-232. Lohse, P.A. and Wright, M.C. 2001. In vitro protein display in drug discovery. Curr. Opin. Drug Discov. Devel. 4:198-204. Martzen, M.R., McCraith, S.M., Spinelli, S.L., Torres, F.M., Fields, S., Grayhack, E.J., and Phizicky, E.M. 1999. A biochemical genomics approach for identifying genes by the activity of their products. Science 286:1153-1155. McCormack, A.L., Schieltz, D.M., Goode, B., Yang, S., Barnes, G., Drubin, D., and Yates, III, J.R. 1997. Direct analysis and identification of proteins in mixtures by LC/MS/MS and database searching at the low-femtomole level. Anal. Chem. 69:767-776. Molloy, M.P. and Andrews, P.C. 2001. Phosphopeptide derivatization signatures to identify serine and threonine phosphorylated peptides by mass spectrometry. Anal. Chem. 73:5387-5394. Molloy, M.P., Herbert, B.R., Walsh, B.J., Tyler, M.I., Traini, M., Sanchez, J.C., Hochstrasser, D.F., Williams, K.L., and Gooley, A.A. 1998. Extraction of membrane proteins by differential solubilization for separation using two-dimensional gel electrophoresis. Electrophoresis 19:837-844. Moody, M.D., Van Arsdell, S.W., Murphy, K.P., Orencole, S.F., and Burns, C. 2001. Array-based ELISAs for high-throughput analysis of human cytokines. Biotechniques. 31:186-194. Munchbach, M., Quadroni, M., Miotto, G., and James, P. 2000. Quantitation and facilitated de novo sequencing of proteins by isotopic N-terminal labeling of peptides with a fragmentationdirecting moiety. Anal. Chem. 72:4047-4057. Nelson, P.S., Clegg, N., Eroglu, B., Hawkins, V., Bumgarner, R., Smith, T., and Hood, L. 2000. The prostate expression database (PEDB): Status and enhancements in 2000. Nucleic Acids Res. 28: 212-213. Oda, Y., Huang, K., Cross, F.R., Cowburn, D., and Chait, B.T. 1999. Accurate quantitation of protein expression and site-specific phosphorylation. Proc. Natl. Acad. Sci. U.S.A. 96: 6591-6596. Oda, Y., Nagasu, T., and Chait, B.T. 2001. Enrichment analysis of phosphorylated proteins as a tool for probing the phosphoproteome. Nature Biotechnol. 19:379-382.
Overview of Proteome Analysis
Analysis of receptor signaling pathways by mass spectrometry: Identification of vav-2 as a substrate of the epidermal and platelet-derived growth factor receptors. Proc. Natl. Acad. Sci. U.S.A. 97:179-184. Pasa-Tolic, L., Jensen, P. K., Anderson, G.A., Lipton, M.S., Peden, K.K., Martinovic, S., Tolic, N., Bruce, J.E., and Smith, R.D. 1999. Highthroughput proteome-wide precision measurements of protein expression using mass spectrometry. J. Am. Chem. Soc. 121:7949-7950. Patton W.F. 2000. A thousand points of light: The application of fluorescence detection technologies to two-dimensional gel electrophoresis and proteomics. Electrophoresis 21:1123-1144. Rain, J.C., Selig, L., De Reuse, H., Battaglia, V., Reverdy, C., Simon, S., Lenzen, G., Petel, F., Wojcik, J., Schachter, V., Chemama, Y., Labigne, A., and Legrain, P. 2001. The protein-protein interaction map of Helicobacter pylori. Nature 409: 211-215. Schweitzer, B., Wiltshire, S., Lambert, J., O’Malley, S., Kukanskis, K., Zhu, Z., Kingsmore, S.F., Lizardi, P.M., and Ward, D.C. 2000. Inaugural article: Immunoassays with rolling circle DNA amplification: A versatile platform for ultrasensitive antigen detection. Proc. Natl. Acad. Sci. U.S.A. 97:113-119. Schweitzer, B., Roberts, S., Grimwade, B., Shao, W., Wang, M., Fu, Q., Shu, Q., Laroche, I., Zhou, Z., Tchernev, V.T., Christiansen, J., Velleca, M., and Kingsmore, S.F. 2002. Multiplexed protein profiling on microarrays by rolling-circle amplification. Nat. Biotechnol. 20:359-65. Srinivas, P.R., Srivastava, S., Hanash, S., and Wright, G.L. Jr. 2001. Proteomics in early detection of cancer. Clin Chem. 47:1901-11. Steen, H. and Mann, M. 2002. A new derivatization strategy for the analysis of phosphopeptides by precursor ion scanning in positive ion mode. J. Am. Soc. Mass Spectrom. 13:996-1003. Su, X., Chew, F.T., and Li, S.F. 2000a. Piezoelectric quartz crystal based label-free analysis for allergy disease. Biosens. Bioelectron. 15:629-639. Su, X., Chew, F.T., and Li, S.F. 2000b. Design and application of piezoelectric quartz crystal-based immunoassay. Anal. Sciences 16:107-114. Tonge, R., Shaw, J., Middleton, B., Rowlinson, R., Rayner, S., Young, J., Pognan, F., Hawkins, E., Currie, I., and Davison, M. 2001. Validation and development of fluorescence two-dimensional differential gel electrophoresis proteomics technology. Proteomics 1:377-396. Unlu, M., Morgan, M.E., and Minden, J.S. 1997. Difference gel electrophoresis: A single gel method for detecting changes in protein extracts. Electrophoresis 18:2071-2077.
Page, M.J., Amess, B., Townsend, R.R., Parekh, R., Herath, A., Brusten, L., Zvelebil, M.J., Stein, R.C., Waterfield, M.D., Davies, S.C., and O’Hare, M.J. 1999. Proteomic definition of normal human luminal and myoepithelial breast cells purified from reduction mammoplasties. Proc. Natl. Acad. Sci. USA. 96:12589-12594.
Washburn, M.P., Wolters, D., and Yates, III, J.R. 2001. Large-scale analysis of the yeast proteome by multidimensional protein identification technology. Nature Biotechnol. 19:242-247.
Pandey, A., Podtelejnikov, A.V., Blagoev, B., Bustelo, X.R., Mann, M., and Lodish, H.F. 2000.
Washburn, M.P., Ulaszek, R., Deciu, C., Schieltz, D.M., and Yates, III, J.R. 2002. Analysis of quan-
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Contributed by Nadeem Ali-Khan, Xun Zuo, and David W. Speicher The Wistar Institute Philadelphia, Pennsylvania
Gel-Based Proteome Analysis
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Protein Profiling Using Two-Dimensional Difference Gel Electrophoresis (2-D DIGE)
UNIT 22.2
Two-dimensional polyacrylamide gel electrophoresis (2-D PAGE) has been widely used over the past three decades to resolve and investigate the abundance of several thousand proteins in a single sample. The technique has enabled identification of the major proteins in a tissue or subcellular fraction by mass spectrometric methods. In addition, 2-D PAGE has been used to compare relative abundances of proteins in related samples, such as mutant and wild-type forms or proteins from altered environments. To date, the majority of comparative protein profiling studies have produced qualitative data, which have enabled the investigator to determine whether expression of a particular protein is increased or decreased. However, this information provides no measure of the extent of this expression change so this approach is unsuitable for clustered data analysis that ultimately may provide an insight into functionality. Quantitative proteomics allows co-expression patterns to be studied, and proteins showing similar expression trends suggest membership of the same functional groups. However, there are still surprisingly few examples of quantitative analysis of changes in protein abundance. Quantitation has been hampered by a series of factors. Silver staining, being more sensitive than Coomassie staining methods, has been widely used for high-sensitivity protein visualization on 2-D PAGE, but it is unsuitable for quantitative analysis, as it has a limited dynamic range, and the most sensitive of silver staining methods are also incompatible with protein identification methods based on mass spectrometry. More recently, the Sypro family of post-electrophoretic fluorescent stains has offered a better dynamic range and sensitivity than silver staining, but is expensive to use. Another problem is the irreproducibility of 2-D gels; no two gels run identically, and corresponding spots between the two gels have to be matched prior to quantification. These aforementioned factors all add variability into the system, thus making it unsuitable for accurate quantitation. Difference gel electrophoresis (DIGE), first described some time ago (Ünlü et al., 1997), has the potential to overcome these issues. This technique relies on pre-electrophoretic labeling of samples with one of three spectrally distinct fluorescent dyes: cyanine-2 (Cy2), cyanine-3 (Cy3), or cyanine-5 (Cy5). The samples are all electrophoresed in one gel and viewed individually by scanning the gel at different wavelengths, thus circumventing problems with spot matching between gels. Image analysis programs can then be used to generate volume ratios for each spot, which essentially describe the intensity of a particular spot in each test sample, and thus enable expression differences to be identified and quantified. This unit describes the DIGE procedure in terms of sample preparation from various types of cells (see Support Protocol), labeling of proteins (see Basic Protocol) and points to consider in the downstream processing of fluorescently labeled samples. For applications of the DIGE technique in the literature see Tonge et al. (2001), Kernec et al. (2001), Gharbi et al. (2002), and Zhou et al. (2002). LABELING OF PROTEIN WITH CYANINE DYES (CyDye DIGE FLUORS) FOR 2-D PAGE
BASIC PROTOCOL
This procedure describes the preparation of protein samples for labeling with N-hydroxysuccinimide (NHS) ester–modified cyanine fluors, determination of the optimal protein concentration and protein/fluor ratio to be used for efficient labeling, and a stepwise guide to labeling. After labeling, the samples are applied to 2-D gels that are then scanned and the resulting images are analyzed. Figure 22.2.1 shows the procedure for labeling two samples in which protein profiles are to be compared. Gel-Based Proteome Analysis Contributed by Kathryn S. Lilley Current Protocols in Protein Science (2002) 22.2.1-22.2.14 Copyright © 2002 by John Wiley & Sons, Inc.
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protein sample 1
protein sample 2
N+ (CH2)4
N+ (CH2)2
N+ (CH2)4
CO2H
CH3
CO2R
label with Cy3 in dark 30 min at 4°C
N CH3
label with Cy5 in dark 30 min at 4°C
quench unreacted fluor by adding 1 mM lysine in dark 10 min at 4°C and mix samples
2-D gel electrophoresis
Figure 22.2.1 Scheme for labeling two samples whose protein profiles are to be compared by 2-D DIGE technique.
CyDye DIGE fluors can be purchased commercially from Amersham Biosciences. They are formulated as NHS esters that react with primary amines. For efficient labeling to take place, an optimum ratio of fluor to protein must be maintained. Typically, 50 µg of protein should be labeled with 200 to 400 pmol of fluor. In cases where larger quantities of labeled proteins are required, it is advisable to maintain this ratio to achieve optimal labeling. Before embarking on large-scale experiments, always check the compatibility of a new sample preparation protocol (e.g., see Support Protocol) with DIGE fluors by labeling a small amount of prepared sample and run on a 1-D mini gel and scan for fluorescence. Compare the amount of fluorescence incorporated with that of a control sample. Materials Protein sample (see Support Protocol) Protein determination kit (e.g., BioRad DC protein assay, BioRad Laboratories; or PlusOne 2-D Quant kit, Amersham Biosciences) Protein concentration kit (e.g., PlusOne 2-D Clean-Up kit, Amersham Biosciences; or PerfectFOCUS, Genotech), optional 0.1 M ammonium acetate in methanol, cold 80% acetone Lysis buffers (see recipe) 400 pmol/µl CyDye DIGE fluors (see recipe) 10 mM L-lysine IPG sample buffer (see recipe) Ampholine sample buffer (see recipe) Protein Profiling Using Two-Dimensional Difference Gel Electrophoresis
0.5-ml microcentrifuge tubes Low-fluorescence glass gel plates Gel fluorescence scanner (e.g., ProXPRESS, PE Biosystems; or Typhoon 9200/9400, Amersham Biosciences)
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Image analysis software: DeCyder software (Amersham Biosciences) is the only analysis software specifically designed for use with DIGE and capable of using one of the CyDye-labeled images as an internal standard, simplifying gel-to-gel analysis. Other software packages can also be used such as Phoretix and Progenesis (Nonlinear Dynamics), MELANIE (Geneva Bioinformatics S.A.) AlphaMatch 2D (Alpha Innotech) PDQuest (BioRad Laboratories), Z3 and Z4000 (Compugen). Additional reagents and equipment for 2-D PAGE (UNIT 10.4) Quantitate protein in sample 1. Determine the protein concentration of all samples in lysis buffer before labeling, e.g., using a kit such as BioRad DC protein assay or PlusOne 2-D Quant kit. There are numerous kits available for determining protein concentration, but it is imperative that the kit employed is appropriate for samples containing detergents.
2. Adjust protein solution to a concentration of 5 to 10 mg/ml using an appropriate lysis buffer. If necessary, concentrate protein samples using either one of several commercial kits such as PlusOne 2-D Clean-Up kit or PerfectFOCUS, or by following steps 3 to 9. If using a kit, proceed to step 10. This concentration allows for an efficient labeling reaction in a small volume.
Concentrate protein by precipitation and resuspension 3. Record the sample volume and add 5 volumes of cold 0.1 M ammonium acetate in methanol and leave for 12 hr or overnight at –20°C. Use this sample volume as the reference volume for subsequent wash steps (steps 5 and 7).
4. Centrifuge for 10 min at 1400 × g, 4°C and remove the supernatant. A pellet of protein should be visible at this stage.
5. Wash the pellet in 5 vol of 80% 0.1 M ammonium acetate in methanol/20% water. 6. Centrifuge for 10 min at 1400 × g, 4°C and remove the supernatant. 7. Wash the pellet with ∼1 vol of 80% acetone, agitate briefly, and centrifuge as described in step 6. 8. Dry pellet for 15 min by leaving the sample tube open in a laminar flow cabinet. 9. Dissolve the pellet in a smaller volume of the appropriate lysis buffer (see Table 22.2.2) and re-measure the protein concentration (step 1). Label proteins 10. Add 50 µg of protein in ≤20 µl to a 0.5-ml microcentrifuge tube. 11. Add 1 µl (400 pmol) of freshly diluted working fluor solution and mix thoroughly by vortexing. The recommended ratio of fluor/protein is 400 pmol:50 ìg. If the ratio is too low, labeling will be inefficient; if it is too high, there is a possibility that multiple labeling events will take place resulting in smearing of spots on 2-D PAGE gels. If a larger amount of protein is to be labeled, add a correspondingly larger amount of dye to maintain the fluor/protein ratio.
12. Centrifuge 30 sec at 12,000 × g, 4°C, to ensure the reagents are at the bottom of the tube and leave on ice for 30 min in the dark. Gel-Based Proteome Analysis
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13. Add 1 µl of 10 mM L-lysine (in MilliQ or equivalent quality water) to quench the reaction. Vortex and centrifuge 30 sec at 12,000 × g, 4°C and leave on ice in the dark for an additional 10 min. The labeled proteins are now stable for 3 months if stored at −70°C. If larger volumes of fluor are used, be sure to add the same volume of 10 mM L-lysine solution as working fluor solution.
14. Pool differentially labeled samples that are to be run on the same gel. Prepare labeled samples for 2-D PAGE 15. Before carrying out isoelectric focusing of labeled proteins, either by immobilized pH gradient (IPG) strips (UNIT 10.4) or ampholine tube gels (UNIT 10.4), dilute the sample with an equal volume of IPG sample buffer or ampholine sample buffer (whichever is appropriate) and incubate on ice for 10 min. If a lysis buffer was used where CHAPS was substituted with another detergent, and 7 M urea/2 M thiourea was substituted for 8 M urea, these alterations can be maintained in the sample buffer. If a lysis buffer was used that contained 2% SDS, the sample must be diluted such that the final percentage of SDS in the sample applied to the IPG strip is <0.2%. Greater amounts of SDS severely compromise the isoelectric focusing of proteins.
16. Perform 2-D PAGE (see UNIT 10.4) using low-fluorescence glass plates. For 7-cm IPG strips used in conjunction with mini gels, use 5 to 20 ìg of sample per gel; for 13-cm IPG strips, use 50 to 200 ìg of sample per gel; and for 24-cm IPG strips, use 100 to 500 ìg of sample per gel. See Critical Parameters for a discussion of special precautions that should be taken when running 2-D PAGE gels.
Capture and analyze gel images 17. Scan gels using a system that is compatible with CyDye DIGE fluors such as the ProXPRESS or the Typhoon 9200/9400 series. Acquire images with 100-µm pixel resolution for accurate determination of image information. The excitation and emission wavelengths for all three fluors are given in Table 22.2.1. Image acquisition can be achieved using a variety of scanners or digital imagers most of which are based on photomultiplier tubes or charged coupled devices (CCDs). See Critical Parameters for a discussion of important factors to consider when scanning 2-D DIGE gels.
18. Analyze scanned images using, e.g., DeCyder differential analysis software. More detail on image analysis is provided in the Experimental Design section of the Commentary. Typically, comparative analysis of images from a gel results in spots, and relative intensities or normalized spot volumes are expressed as ratios. DeCyder software (Amersham Biosciences) is the only analysis software specifically designed for use with DIGE and capable of using one of the CyDye-labeled images as an internal standard, simplifying gel-to-gel analysis.
Table 22.2.1
CyDye DIGE Fluor Excitation and Emission Wavelengths
Excitation maxima λ Protein Profiling Using Two-Dimensional Difference Gel Electrophoresis
Cy2 Cy3 Cy5
480 540 620
Emission maxima λ 530 590 680
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PREPARATION OF PROTEIN FOR Cy LABELING REACTIONS Protein samples prepared from a variety of different sources can be rendered suitable for labeling with CyDye DIGE fluors. This protocol describes protein extraction from cultured cells, plant cells, bacterial and yeast cells, Drosophila, serum or culture media, and tissue samples. After extraction, all samples should be stored in aliquots at –70°C until required for labeling.
SUPPORT PROTOCOL
Materials Cultured cells, plant cell extracts, bacterial or yeast cells, Drosophila, serum or culture media (secreted proteins), or tissue samples Wash solution: 10 mM Tris⋅Cl, pH 8.0 and 5 mM magnesium acetate Lysis buffer (see recipe; see Table 22.2.2) Phenol saturated with TE (Sigma-Aldrich) TE saturated with phenol (see recipe) Cell wash buffer: 75 mM PBS (APPENDIX 2E) 0.9% saline Additional reagents and equipment for protein precipitation (see Basic Protocol) NOTE: Typical cell lysing conditions involve resuspending a cell pellet in an appropriate lysis buffer and incubating for 30 to 60 min on ice. A typical cell washing procedure involves resuspending a cell pellet in wash solution at ≥2× the volume of the cell pellet, gently vortexing, and then re-centrifuging. For cultured cells 1a. Wash pelleted cells with an appropriate solution to remove any reagents such as primary amines from the growth medium that would interfere with labeling. This wash solution should be chosen such that it does not lyse the cells, e.g., an isotonic buffer such as PBS. Wash solutions should typically not contain primary amines, although Tris solutions at a concentration of ≤10 mM are permissible.
2a. Centrifuge cells 10 min at 12,000 × g, 4°C, and lyse with an appropriate lysis buffer (see recipes in Table 22.2.2). For plant cell extracts For preparations from total plant cell extracts, it is necessary to remove plant metabolites such as polyphosphates and carbohydrates prior to labeling. A successful method of achieving this involves precipitation of proteins at a phenol/aqueous interface. 1b. To a solution or suspension of protein extract, add an equal volume of phenol saturated with TE. 2b. Vortex ten times in 10-sec bursts between incubation on ice for a total of 10 min. 3b. Centrifuge for 5 min at 1400 × g, 4°C. 4b. Remove and discard the top aqueous layer. There should be a visible layer of protein at the interface of the phenol and aqueous layer; be careful not to disturb this when removing the top layer.
5b. Add two volumes of TE saturated with phenol. 6b. Repeat steps 2b and 3b. 7b. Remove and discard aqueous phase 8b. Proceed to the protein precipitation procedure described in Basic Protocol, steps 3 to 9.
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9b. After precipitation, resuspend pellets in an appropriate lysis buffer. An appropriate lysis buffer is one that will most effectively solubilize the proteins of interest, e.g., a lysis buffer containing a detergent such as ABS14 may be more effective in solubilizing membrane-associated proteins.
For bacterial and yeast cells 1c. Wash pelleted cells in the cell wash buffer (centrifuge 10 min at 12,000 × g, 4°C). Repeat wash and pellet cells two times. 2c. Resuspend cells in an appropriate lysis buffer. It is advisable to add a protease inhibitor cocktail (see recipe for lysis buffer for protease inhibitor component compatibility) to the lysis buffer.
3c. Sonicate the cells on ice three times in 10-sec bursts, ensuring that the temperature of the sample does not rise due to the presence of urea in the lysis buffer. Heating will result in carbamylation of primary amines within the proteins, leading to changes in pI.
4c. For best results, precipitate proteins as described in Basic Protocol, steps 3 to 9. Resuspend pellets in appropriate lysis buffer (DNase can be used at typical concentrations if required). For Drosophila 1d. Homogenize whole flies directly into an appropriate lysis buffer. For best results, precipitate proteins as described in Basic Protocol, steps 3 to 9, and resuspend pellets in appropriate lysis buffer. Generally, 100 ìg of protein can be extracted from an individual Drosophila fly. An efficient method of extracting protein from Drosophila is to place frozen flies in a 1.5- or 2.0-ml microcentrifuge tube and add a volume of lysis buffer equal to the volume of flies used. The flies can then be ground using a plastic micropestle. An appropriate lysis buffer is one that will most effectively solubilize the proteins of interest, e.g., a lysis buffer containing a detergent such as ABS14 may be more effective in solubilizing membrane-associated proteins.
For serum or culture media (secreted proteins) 1e. Precipitate proteins as described in Basic Protocol, steps 3 to 9, and resuspend pellets in an appropriate lysis buffer. Proteins added to culture medium, such as fetal calf serum proteins, will also precipitate and will be present on subsequent gels.
For tissue samples 1f. Rinse in 0.9% saline and homogenize in an appropriate lysis buffer. In all cases, check the pH of the samples in lysis buffer before labeling. It is imperative that the final pH of this solution is between 8.0 and 8.8 for efficient labeling to occur.
Protein Profiling Using Two-Dimensional Difference Gel Electrophoresis
Sufficient tissue should be used to produce the required amount of protein for the planned experiment. This will depend on the experimental plan and the type of tissue used. Generally, 0.2 g of tissue in 5 ml lysis buffer will give a concentration of 5 to 10 mg/ml. The best homogenization method to use will also depend on the type of tissue and the size of the sample necessary to provide the required amount.
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PROTEIN IDENTIFICATION FROM DIGE GELS BY MASS SPECTROMETRIC TECHNIQUES
ALTERNATE PROTOCOL
DIGE labeling of proteins is compatible with in-gel digestion of proteins to peptides by proteases, and with subsequent identification by mass spectrometric techniques such as peptide mass fingerprinting. It is necessary to post-stain the gel with colloidal Coomassie or mass spectrometry-compatible silver stain to visualize the proteins for spot excision. With DIGE minimal fluor labeling, the majority of the protein on a gel is unlabeled and as such is not visualized on the DIGE images. The fluor molecules add ∼500 Da to the labeled proteins, causing the labeled and unlabeled proteins to migrate to different positions within the SDS-PAGE gel–this is most significant with low molecular weight species. Therefore, to pick sufficient protein for in-gel trypsin digestion and identification by mass spectrometry, it is necessary to use a total protein stain to visualize and identify the unlabeled proteins, unless an automated system with appropriate fluorescence detection is used to excise the spots of interest for a DIGE gel, or one that can import spot coordinates from a scanned image. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Ampholine sample buffer 9.8 M urea 2% carrier ampholytes (Amersham Biosciences) 4% NP40 20 mg/ml DTT Aliquots without DTT and carrier ampholytes may be stored up to 12 months at −20°C. Add DTT and carrier ampholytes immediately before use. CyDye DIGE fluors Stock solution: CyDye DIGE fluors (Cy2, Cy3, Cy5) are purchased as a powder from Amersham Biosciences. For long-term storage, dissolve the powder in dimethyl formamide (DMF) to a final stock concentration of 1 nmol/µl. This solution is stable for 2 months at –70°C. Store the stock fluor solution in small (2-µl) aliquots to avoid freeze-thaw cycles. Working solution (400 pmol/µl): Dilute the fluor stock solution to 400 pmol/µl for the labeling protocol. Once diluted to a working concentration, the fluors are stable for 2 weeks at −20°C. Use high-quality DMF with a water content of <0.005%. DMF will degrade over time to form amine compounds that will reduce the efficiency of labeling. It is therefore recommended to replace DMF with a fresh bottle at least every 3 months. CAUTION: DMF is harmful upon inhalation or contact with skin. Handle with a double layer of gloves in a chemical fume hood. Follow all precautions listed with this chemical.
IPG sample buffer 20 mg/ml DTT 2% IPG buffers or carrier ampholytes (Amersham Biosciences) 4% CHAPS 8 M urea Prepare fresh
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Lysis buffers Table 22.2.2 describes several useful lysis buffers that are appropriate for preparation of DIGE samples. The following are recommended detergent components: a. ASB14 (amidosulphobetaine-14 ) [182750], Calbiochem b. CHAPS (3-[(3-chloamidopropyl)dimethylammonio]-1-propane-sulfonate; Sigma) c. SB3-10 (N-decyl-N,N-dimethyl-3-ammonio-1-propanesulfonate; Sigma) at concentrations <1% d. NP-40 (ethylphenyl-polyethylene glycol; USB) at concentrations >1% However, the following components should be excluded from all lysis buffers used in DIGE labeling procedures: a. Any compound containing primary amines b. Reducing agents: >2 mg/ml DTT or >1 mM TCEP; 2-mercaptoethanol at any concentration c. Buffers: >5 mM HEPES, CHES, PIPES; or ampholine or IPG buffers at any concentration d. Detergents: >1% TritonX-100, SDS, or NP-40 For more details about the effect of detergents on protein solubilization, see Santoni et al. (2000).
e. Protease inhibitors: Any preparation containing AEBSF or >10 mm EDTA Use other standard protease inhibitors at recommended concentrations.
Table 22.2.2
Recommended Lysis Buffers for 2-D DIGE Sample Preparation
Lysis buffer
Reagent
Concentration
CHAPS Urea Tris⋅Cl, pH 8.0-9.0 Magnesium acetate
4% (w/v) 8M 10-30 mM 5 mM
CHAPS Urea Thiourea Tris/HCl pH 8.0-9.0 Magnesium acetate
4% (w/v) 7M 2M 10-30 mM 5 mM
ASB14b Urea Thiourea Tris⋅Cl pH 8.0-9.0 Magnesium acetate
2% (w/v) 7M 2M 10-30 mM 5 mM
SDS Tris⋅Cl pH 8.0-9.0 Magnesium acetate
2% (w/v) 10-30 mM 5 mM
Standarda
Thiourea-containinga
ASB14-containinga
SDS-containingc
aDo not heat solutions containing urea and thiourea in order to aid solubilization.
Protein Profiling Using Two-Dimensional Difference Gel Electrophoresis
bASB14 can be substituted with NP40 or SB3-10, providing the final concentration of NP40 before
labeling is <1%. Use 2% SB3-10. cSDS-containing solutions must be diluted to a final concentration of ≤0.2% before successful
isoelectric focusing can take place.
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TE saturated with phenol Make a 100 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) and 1 mM EDTA solution. Add phenol saturated with TE solution (Sigma-Aldrich) dropwise and vortex after the addition of each drop. Repeat the dropwise additions until a lower partitioned phenol layer is visible. CAUTION: Phenol is toxic on contact with skin and if swallowed. Handle with a double layer of gloves in a chemical fume hood. Follow all precautions listed with this chemical.
COMMENTARY Background Information This unit describes the methods required to profile proteins using 2-D difference gel electrophoresis (DIGE). This method was first described in 1997 by Jon Minden and co-workers (Ünlü et al., 1997) and involves the pre-electrophoretic labeling of samples with one of three spectrally distinct fluors, cyanine-2 (Cy2), cyanine-3 (Cy3), or cyanine-5 (Cy5). More than one set of samples, each labeled with a different Cy dye, is run on one gel and viewed individually by scanning the gel at different wavelengths, thus circumventing problems with spot matching between gels. Image analysis programs are then used to generate volume ratios for each spot. Volume ratios essentially describe the intensity of a particular spot in each test sample, and thus enable expression differences to be identified and quantified. This methodology has since been commercialized by Amersham Biosciences and all three CyDye DIGE fluors are available. CyDye DIGE fluors have a reactive N-hydroxysuccinimide ester group that forms a covalent bond with the ε-amino group of lysine side chains (see Fig 22.2.1 for CyDye DIGE fluor structures). Labeling reactions are designed such that the stoichiometry of protein to fluor results in only 1% to 2% of the total number of lysine residues being labeled. The fluors also carry a net charge of +1 so the pI of the protein is maintained with labeling. The three fluors have essentially equivalent molecular weights; each labeling event adding ∼500 Da to the mass of the protein The employment of DIGE labeling for comparing the relative abundance of proteins between two or several samples has many benefits: Reproducibility Most protein profiling studies to date have required the test samples to be compared on separate 2-D gels, with subsequent visualization of the spots with silver or Coomassie staining. Inter-gel variation has been problematic and spot matching using warping algorithms
that are built into various gel analysis software packages have been employed with varying success. The CyDye DIGE fluors are matched for both mass and charge, therefore, a protein labeled with any of these fluors will migrate to the same position on the 2-D gel, thus eliminating intra-gel variation. Sensitivity Labeling with CyDye DIGE fluors is the most sensitive method of protein detection currently available with detection of 125 pg of a single protein and a linear response in protein concentration over at least five orders of magnitude. In comparison, the limits of detection with silver stain is ∼1 ng of protein with a dynamic range of no more than two orders of magnitude (Tonge et al., 2001; Lilley et al., 2002). Multiplexing The specific spectral properties of the three CyDye DIGE fluors enable multiplexing of up to three separate protein mixtures on the same 2-D gel. Internal standards The use of DIGE labeling enables internal standardization to be achieved along with matching of protein patterns across sets of gels. A pool of all the samples within the experiment can be labeled with one of the three fluors. This pooled sample can then be run in every gel in an experimental set and, as it contains every protein from every sample, it further negates the problem of inter-gel variation. This approach allows accurate and statistically relevant quantitation of differences between large numbers of samples. The minimal labeling of proteins with CyDye DIGE fluors is compatible with the downstream processing commonly used to identify proteins within spots on 2-D gels showing changes in abundance between sets of samples. Identification is achieved using a variety of mass spectrometric methods, all of which rely
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on the production of peptides by proteolysis within excised gel pieces. The protease most frequently employed is trypsin, which cleaves the peptide bond on the C-terminal side of lysine and arginine residues. Minimal labeling results in only 1% to 2% of lysine residues being modified and therefore does not significantly reduce the number of available tryptic sites. It is unlikely that a peptide modified with a CyDye DIGE fluor would be extracted from a gel piece, hence interference by the fluors in peptide mass fingerprinting and de novo sequencing mass spectrometric techniques is minimal.
Protein Profiling Using Two-Dimensional Difference Gel Electrophoresis
Experimental design The design of a protein profiling analysis experiment using DIGE is crucial to the degree of statistical significance that can be assigned to the resulting data. Traditional 2-D electrophoresis suffers from two major sources of variation: (1) experimental and (2) biological variation. With DIGE, many sources of experimental variation can be controlled by the inclusion of an internal standard on each gel. Experimental variation. Variation introduced by experimental procedures can occur at numerous positions within the DIGE procedure. First, as with traditional 2-D PAGE, sample preparation must be carefully controlled and maximally replicated between sets of samples. Ideally, multiple preparations of samples are required to ensure that changes in protein profiles are a function of the biological system and not a function of variation in sample preparations. There are several points within this process where particular care should be taken; one is where preparation protocols require the addition of protein solutions such as BSA to maintain protein concentration during extraction procedures. Such additions should be avoided if possible. The extent of proteolysis between samples during preparation may also be a variable. Variations in the extent of proteolysis should be assessed by differentially labeling replicates, running them on the same gel, and comparing the spot profiles. Alternative protease inhibitors should be considered during sample preparation if consistent profiles are not observed. Second, variation can be introduced during labeling. Care should be taken to ensure that approximately equal amounts of protein are added to each labeling reaction. To some extent, as with traditional 2-D PAGE, different protein loads are normalized at the analysis stage. It is crucial to load similar amounts of protein on the gel for accurate quantitation of lower abun-
dance species; the use of greatly varying amounts of protein will result in a particular protein being absent from one image as the amount of fluorescence associated with it will fall below the threshold of detection. Variations in the electrophoretic properties of proteins labeled with specific CyDye DIGE fluors is occasionally observed, mostly as a result of a shift in the apparent pI of the protein. To check for this, it is advisable to run the sample labeled with all the fluors to be used on the same gel; consistent profiles should be observed. Third, gel-to-gel variation will occur due to differences in electrophoretic conditions, different IEF strips, or ampholine tubes, different second dimensions, particularly in situations where pre-poured gels are not purchased from commercial sources, and operator-to-operator variation. Gel-to-gel variation can be controlled to some extent by the use of internal standards as described later in this section. Lastly, variation will be introduced during analysis due to poor detection and background subtraction. Observed differences in the effective potencies of the fluors can occur depending on the preparation of the fluor stocks and buffer components used in sample preparation. This can contribute towards different levels of background fluorescence on the gel images. There are more noticeable differences in background levels between images within one gel due to the different fluorescent characteristics of acrylamide at the various wavelengths used for excitation of Cy2, Cy3, and Cy5. The different background levels are accounted for in the downstream analysis. The use of the pooled internal standard within each gel negates these minor differences associated with labeling with different fluors, as well as reducing gel-to-gel variation associated with traditional 2-D PAGE. In conclusion, it is necessary to assess the experimental variation in the DIGE process for any new set of samples. This is achieved by running sets of gels where the same sample is labeled with all fluors to be used, loaded onto one gel, and full analysis carried out. This will also give an indication of the inherent error of the system and suggest the threshold of significance or a fold change above which indicates true expressional changes. Biological variation. Biological variation between individual organisms and cultures also leads to variation. This variation should be assessed carefully to ensure that only biologically significant changes in protein profiles are taken into consideration. The only mechanism by which biological variation can be monitored
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is by preparing samples from biological replicates of each group and comparing profiles. This is best achieved by using an internal standard as described below. Another approach would be to simply pool a large number of biological replicates to form an average sample, but this is not always advisable. Unless the extent of possible variation between individuals is assessed, there is no way of knowing whether the ’average’ sample is skewed by an outlier. Accuracy of data. To achieve accurate and meaningful results, consideration must be given to methods employed to normalize data for both biological and experimental variation. Most analysis packages allow for normalization between different fluorescent images of the same gel. This type of normalization accounts for differences in the amount of material used in individual labeling reactions or differences in the potencies of fluors used. Normalization tends to either use a global theme or uses individual spot or spot intensities. Global normalization assumes that the sum of the pixel intensities for each image should be the same and applies a correction factor to achieve this. This approach can be lead to errors in analysis especially in cases where a very abundant protein species or numerous proteins are present in much greater quantities in one sample, leading to a significant increase in the sum of the pixel values for the associated image. Global normalization in these situations will lead to an underestimation of spot volumes in the case of the image containing such a protein and hence will yield skewed data. Normalization using a single spot (or group of selected spots) assumes that a spot is identified, which is present in all images to be compared whose expression does not vary within the system being tested (sometimes referred to as a housekeeping protein). The spot volume of this spot should be equivalent across all images. A correction factor is calculated to achieve equivalence for this spot and is then applied to all other detected spots. This approach is the least desirable method of normalization and is only valid in situations where the identity of a spot is known and where other experimental data shows that expression of this “reference” protein does not change under the experimental conditions used. Comigration of another protein whose expression is affected by experimental conditions with this reference protein would also result in corruption of the normalization. A variation of this method would be to include a constant amount of a standard protein into labeling reactions, employing a protein that migrates to a discrete
position on the 2-D gel. The volume of the spot formed by this protein should be consistent between gels and a correction factor can be applied if this is not the case. Finding a suitable protein is not trivial and this type of normalization does not account for situations where there is a very abundant protein or where numerous proteins are present in much greater quantities in one sample, leading to relative skewing of the amount of protein added to a labeling reaction. A more robust method that addresses experimental and biological variation as well as normalization between labeling reactions and gel images involves the use of the Cy2 labeling reaction to form an internal standard. The internal standard is created by combining aliquots from all biological samples in the experiment. This internal standard is then run on every gel in an experiment and spot volumes from the Cy3 and Cy5 images are expressed as a ratio of the corresponding spot in the Cy2 image. This approach has a number of advantages: (1) Improved accuracy of spot volume quantitation as all spots are present in the internal standard, hence the effect of gross changes in expression between samples is minimized. (2) Ability to link numerous gels in a large experimental design. The Cy2 image should be identical in each gel, thus allowing meaningful statistical analyses. (3) Discrimination of gel-to-gel variation from biological variation and inconsistencies in sample preparation.
Critical Parameters Sample preparation Always check new sample preparation protocols before committing precious sample to a 2-D DIGE experiment. This is easily achieved by labeling a small amount of prepared sample with just one of the CyDye DIGE fluors and running it on a 1-D minigel alongside a control sample known to be labeled with these fluors. Scan the gel and compare fluorescence intensities between the new preparation and the control sample. It is advisable to check the pH of the samples in lysis buffer before labeling, as efficient labeling will only take place between pH 8.0 and 8.8. It is imperative to use lysis buffers that do not contain compounds with primary amine groups, as these will react with the fluors and reduce the effective concentration of the fluors. It is also important to choose lysis buffers that do not contain significant concentrations of salts or ionic detergents as these will interfere with isoelectric focusing.
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Labeling The recommended ratio of fluor to protein is 400 pmol:50 µg. A sub-optimal ratio will result in inefficient labeling, while the use of higher ratios may result in multiple and variable labeling events and subsequent problems during resolution of labeled protein into discrete spots in the second dimension of 2-D PAGE. (Each fluor addition will lead to a gain in effective mass 500 Da.) Larger amounts of proteins can be labeled, but in these situations a corresponding increase of fluor must be included to maintain the fluor/protein ratio. Where larger volumes of fluors are used, the same volume of L-lysine solution as working fluor solution must be added for effective quenching of the labeling reaction. Any reactive fluor present after quenching can potentially react with buffer components added in the later stages of the DIGE process and lead to spurious fluorescence on the final 2-D gel. 2-D PAGE considerations Photobleaching of the fluors must be prevented by running both dimensions in the dark or covering gel rigs to exclude light. Alu-
A
minium foil is not recommended because it can cause problems with instruments. To prevent unacceptable levels of background fluorescence in subsequent imaging systems that scan through gel plates, use lowfluorescence glass plates. Prior to preparing gels, wash glass plates thoroughly, with a final Milli-Q water wash followed by drying with lint-free tissues. Filter the acrylamide solution to remove dust particles that can lead to scanning problems and fibers from clothing that may introduce unwanted keratins into the system. At all times, wear powder-free latex or nitrile gloves and work in an environment that is as dust-free as possible. Scanning 2-D DIGE gels Save data as recorded without data compression as this can affect the accuracy of recording and the amount of retrievable information upon transport into analysis packages. For accurate quantitation, do not exceed the maximum pixel intensity of the instrument. At the maximum pixel intensity, saturation of the detector system is reached resulting in inaccu-
C
B
no difference presence/absence up/down-regulation
Protein Profiling Using Two-Dimensional Difference Gel Electrophoresis
Figure 22.2.2 Typical gel images from a 2-D DIGE experiment. The images are from a 2-D analytical gel (pH 4 to 7), which was loaded with 50 µg of a total protein extract from a wild-type strain of Erwinia carotovorea labeled with Cy3 (A) and 50 µg of total protein extract from a mutant strain of Erwinia carotovorea labeled with Cy5 (B). The images were acquired using a 2920-2D Master Imager (Amersham Biosciences). The images were exported as 16-bit TIFF images for analysis. To produce this figure, 8-bit TIFF versions of the images were imported into Adobe Photoshop version 5 and false colored, green for Cy3, red for Cy5 and the two images overlaid (C). Key: yellow, no difference; green/red, presence/absence; orange, up/down-regulation. This black and white facsimile of the figure is intended only as a placeholder; for full-color version of figure go to http://www.currentprotocols.com/colorfigures.
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rate volume measurements, and in the case of CCD based systems, risk of bleedover of signal from one pixel to its neighboring pixels. The linear dynamic range of DIGE labeling is five orders of magnitude; the dynamic range of the imaging system used may be significantly less than this and will have great impact on the quality of data recorded. In extreme cases, two different image intensity settings may be employed, resulting, in the case of the higher setting, in some spots with pixel intensities at saturation. Specks of dust within the gel and on the surface of gel plates may give rise to pixel intensities at the instrument maximum. It is advisable to assess which pixel is giving the highest reading on an image. Spot morphology will indicate a “real” spot or dust. Save individual images as 16-bit TIFF (tagged image file format) images for import into most commercially available 2-D gel image analysis packages. This is the most commonly accepted format for files to be exchanged between different software applications and platforms.
Troubleshooting This section deals with failure to produce fluorescently labeled proteins on 2-D gels, which can be attributed to inadequate sample preparation or labeling protocols and not failures caused by problems with the 2-D electrophoresis process. A troubleshooting guide to 2-D PAGE can be found in UNIT 10.4 Lysis The most common problem associated with sample preparation is low yield of protein from cell lysis. When preparing samples from bacterial or yeast culture, check that the cell densities are optimal for the extraction. It may be necessary to increase sonication times to improve the amount of protein extracted from such samples. In the case of samples from other sources, changing to a lysis buffer containing a different detergent, such as ASB14, and altering the incubation time may improve the yield of extracted proteins. Where there is insufficient protein in the sample, use the protein precipitation methods described in this unit and resuspend in a smaller volume of lysis buffer to achieve the optimal protein concentration for labeling. Labeling with CyDye DIGE fluors There are several potential problems that can occur when labeling proteins with CyDye DIGE fluors that will result in weak fluorescent signals on the 2-D gel image.
Certain reagents interfering with correct determination of protein concentration. When determining the concentration of a protein solution to be used for labeling, use a kit that is compatible with solutions containing thiourea and the various detergents that are used in the lysis buffer and were used to prepare the sample. Failure to use an appropriate kit may result in overestimation of the concentration of protein in the sample. Incorrect pH of sample for labeling reaction. Check the pH of the sample immediately before labeling. If the pH is <8.0, increase it by adding an additional volume of the lysis buffer, which contains 30 mM Tris⋅Cl, pH 9 to 10. Buffer components are incompatible. Check whether the sample contained any thiol reagents or primary amines at concentrations in excess of those stipulated in the sample preparation section of this unit. It may be worth reprecipitating the sample and resuspending in a compatible sample buffer. Fluor is degraded. The fluor solution may be degraded due to hydrolysis of the NHS-ester if DMF containing a higher-than-recommended water content is used in its preparation. DMF will degrade over time to produce amine compounds with which the CyDye DIGE fluors will react, thus, reducing the effective concentration of fluor in labeling reactions. Always store fluor solutions in the dark to minimize photodegradation that will reduce the efficiency of dye labeling. Irreproducible fluorescent spot patterns Variations in spot patterns can result from several factors. Proteolysis during sample preparation will lead to the appearance of truncated species on 2-D gels, hence be sure to include an appropriate set of protease inhibitors into the lysis buffer used if proteolysis is evident. Variable yields from the protein extraction procedure will lead to inconsistent spot patterns. Optimize the extraction process using alternative lysis buffers to improve reproducibility of extraction. Addition of proteins such as BSA and DNase may lead to irreproducible patterns. Wherever possible, avoid the use of these proteins during sample preparation and wear gloves at all times to prevent contamination with keratins. Finally, dust particles on the gel plates or within the gel matrix can also lead to spurious spots. Filter all gel solutions before polymerization and use lint-free tissues to clean all glass plate surfaces. Gel-Based Proteome Analysis
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Anticipated Results The end point of the 2-D DIGE process results in the acquisition of two or more fluorescent images from a single 2-D gel (Figure 22.2.2). Each image represents the proteins present in samples labeled with a specific fluor. Analysis of the images will generate volume ratios for each spot, which essentially describe the intensity of a particular spot in each test sample, and thus enable expression differences to be identified and quantified. Gel images and their associated spot volumes should be reproducible when the process is duplicated.
Time Considerations The process of labeling samples with CyDy DIGE fluors takes ~1 hr. The 2-D electrophoresis process requires a variable length of time depending on the system used and the size of the final 2-D gel. Similarly, image capture and analysis will depend on the scanner and software employed.
Literature Cited Gharbi, S., Gaffney, P., Yang, A., Zvelebil, M.J., Cramer, R., Waterfield, M.D., and Timms, J.F. 2001. Evaluation of two-dimensional differential gel electrophoresis for proteomic expression analysis of a model breast cancer cell system. Mol. Cell. Prot. 1:91-98. Kernec, F., Ünlü, M., Labeikovsky, W., Minden, J.S., and Koretsky, A.P. 2001. Changes in mitochondrial proteome from mouse hearts deficient in creatine kinase. Physiol. Genom. 6:117-128.
Lilley, K.S., Razzaq, A., and Dupree, P. 2002. Twodimensional gel electrophoresis for the definition of protein function. Curr. Opin. Chem. Biol. 6:46-50. Santoni, V., Kieffer, S., Desclaux, D., Masson, F., and Rabilloud, T. 2000. Membrane proteomics: Use of additive main effects with multiplicative interaction model to classify plasma membrane proteins according to their solubility and electrophoretic properties. Electrophoresis 21:33293344. Tonge, R., Shaw, J., Middleton, B., Rowlinson, R., Rayner, S., Young, J., Pognan, F., Hawkins, E., Currie, I., and Davison, M. 2001. Validation and development of fluorescence two dimensional differential gel electrophoresis proteomics technology. Proteomics 1:377-396. Ünlü, M., Morgan, M.E., and Minden, J.S. 1997. Difference gel electrophoresis: A single gel method for detecting changes in protein extracts. Electrophoresis 18:2071-2077. Zhou, G., Hongmei, L., DeCamp, D., Chen, S., Shu, H., Gong, Y., Flaig, M., Gillespie, J.W., Hu, N., Taylor, P.R., Emmert-Buck, M.R., Liotta, L.A., Petracoin, E.F., and Zhao, Y. 2002. 2D Differential in-gel electrophoresis for the identification of esophageal scans cell cancer-specific protein markers. Mol. Cell. Prot. 1:117-124.
Internet resources http://www4.amershambiosciences.com An Amersham Biosciences Ettan DIGE user manual can be downloaded as a pdf file.
Contributed by Kathryn S. Lilley University of Cambridge Cambridge, United Kingdom
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Laser Capture Microdissection for Proteome Analysis
UNIT 22.3
Tissues vary in their heterogeneity but often are composed of many different cell types. For example, the main types of cell within the kidney include proximal tubule epithelial cells, distal tubule epithelial cells, collecting duct epithelial cells, endothelial cells, mesangial cells, fibroblasts, and muscle cells. In pathological circumstances, the situation may be even more complex, with diseased cells adjacent to normal cells and the presence of necrotic areas and additional infiltrating cells, e.g., lymphocytes and monocytes. Attempting to understand the biology or pathology of particular cells against such a heterogeneous background is therefore a challenge. Laser capture microdissection (LCM) is a recently introduced approach that allows the enrichment of specific cell types for subsequent analysis. To isolate specific cell types, frozen tissue sections are first stained, most commonly using modified rapid hematoxylin and eosin staining protocols that ensure protein integrity and compatability with subsequent analytical techniques. Alternative stains such as methyl green or toluidine blue are also suitable. All of these techniques are described in the LCM protocol below (see Basic Protocol 1). If visualization of the specific cells is not adequate using histological stains, then it is possible to use modified immunolabeling methods such as silver-enhanced gold labeling to highlight the cells of interest prior to LCM (see Alternate Protocol 1). The processing of the tissue samples post-surgery is crucial (see Support Protocol 1), as only frozen material and not archival paraffin-embedded material is suitable for subsequent protein analysis. Following staining, cells of interest are then captured by LCM (see Basic Protocol 1). Essentially this is carried out by visualizing the cells of interest through a Perspex cap coated with a thermolabile film. A laser beam is fired at the cells, melting the film and causing it to fuse with the underlying cells so that they are retained on the cap when it is lifted from the section (Fig. 22.3.1). The cells are then solubilized from the cap using an appropriate buffer. As the cell lysates obtained are usually contained in only a few microliters and may be of relatively low concentration, sensitive protein assay methods have to be used to quantify the protein extracted (see Support Protocol 2). A technique which has received a lot of attention as a possible subsequent analysis method is two-dimensional polyacrylamide electrophoresis (2-D PAGE), where proteins are separated in the first dimension by isoelectric focusing using immobilized pH gradient (IPG) strips and in the second dimension by SDS-PAGE (see Basic Protocol 2 and UNIT 10.4). This combination of separation modalities allows a much higher resolution of the proteins that are present than would be achieved with either technique alone, and also provides information about two independent features of the proteins—i.e., isoelectric point and molecular weight. If particular proteins are the target of the investigation, an alternative approach is to use immunoblotting (western blotting), which can be done with as little as a few hundred cells (see Basic Protocol 3 and UNIT 10.10), provided that the necessary antibodies are available. Surface enhanced laser desorption ionization mass spectrometry (SELDI-MS) is an emerging technology for protein analysis. Proteins selectively immobilized on the surface of ProteinChips (Ciphergen Biosystems) on the basis of conventional chromatographic chemistries are profiled using a SELDI mass spectrometer with a time-of-flight detector (see Basic Protocol 4). The high sensitivity of this technology allows the use of small amounts of sample such as those generated from LCM. A further level of selection is Contributed by Rachel A. Craven and Rosamonde E. Banks Current Protocols in Protein Science (2003) 22.3.1-22.3.27 Copyright © 2003 by John Wiley & Sons, Inc.
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laser
extraction buffer
cap
section slide
Figure 22.3.1 Illustration of the basic principles of laser capture microdissection as described in the text.
generated by the use of different extraction buffers for the sample. Differing buffers can also be selected to increase or decrease the stringency of selection, and different sample matrix preparations can be used to optimize ionization in the different mass regions of the spectra generated (see Critical Parameters and Troubleshooting). This unit also includes details of the alternative laser-based systems which are now available for microdissection, as well as examples of results generated using the approaches described in the unit and with other approaches such as protein microarrays and immunoassays (see Commentary). The Commentary also includes a discussion of the limitations of the LCM approach, as well as a likely time frame for the procedure and a troubleshooting guide. NOTE: High-purity water (Milli-Q or equivalent) should be used throughout for all solutions. Chemicals should be at minimum Analar grade or equivalent, unless otherwise indicated. BASIC PROTOCOL 1
Laser Capture Microdissection for Proteome Analysis
LASER CAPTURE MICRODISSECTION (LCM) Prior to LCM, tissue sections must be stained to allow microscopic selection of areas of interest. The LCM procedure below includes a modification of standard histochemical staining protocols, with very brief incubation times and the incorporation of protease inhibitors into staining baths, that has been successful for maintaining sample integrity for subsequent protein analysis by a number of analytical techniques. Hematoxylin and eosin is perhaps the most commonly used histochemical stain for visualizing tissue morphology, but other stains such as methyl green and toluidine blue have also been used successfully. However, tissue-section processing must be carefully controlled to minimize in vitro artifacts and interference with downstream protein analysis, and to maximize protein recovery. Work-up experiments should be carried out to assess recovery by analyzing protein extracted from undissected stained tissue sections and comparing it to that obtained from unprocessed tissue. In the subsequent steps, the use of LCM for the isolation of particular cell types from stained tissue sections is described. The Arcturus PixCell II LCM system consists of an inverted microscope mounted with a near-infrared laser that can be focused on a selected area of tissue, and a joystick that controls the position of the stage, linked to a workstation which allows image archiving as well as real time visualization of the section being dissected. LCM relies on the use of a cap with a thermolabile film that is placed in contact
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with the tissue section. Localized firing of the laser causes the film to melt, fusing it with the underlying tissue. Captured tissue is then selectively removed when the cap is lifted. NOTE: It is recommended that images of the tissue be captured before and after dissection and stored as a record. Materials OCT-embedded tissue blocks (see Support Protocol 1) 70% and 100% (v/v) ethanol Mayer’s hematoxylin (see recipe) Scott’s water (see recipe) 1% (w/v) eosin (BDH Chemicals) Complete Mini Protease Inhibitor Tablets (Roche Molecular Biologicals) Xylene Solubilization buffer compatible with protein analysis to be performed Cryostat (Leica CM1850 or equivalent) Glass slides, cleaned by dipping in ethanol Arcturus PixCell II laser capture microdissection system CapSure LCM caps or CapSure HS caps (Arcturus) Humidified chamber (prepared by placing wet tissues inside the base of a plastic pipet-tip box) 0.5-ml Eppendorf Safe-Lock tubes Prepare stained tissue sections 1. Cut 8-µm sections from OCT-embedded fresh-frozen tissue onto ethanol-dipped glass slides and place on dry ice. The cryostat blade should be thoroughly cleaned with ethanol between samples to avoid contamination. Sections should ideally be used the same day, although they may be stored at −80°C for subsequent use. The effect of storage should be evaluated, as problems may be encountered with loss of antigen reactivity and protein degradation.
2. Prepare Coplin jars containing Mayer’s hematoxylin, Scott’s water, deionized water, and 1% eosin. Add one Complete Mini Protease Inhibitor Cocktail tablet each to the Coplin jars containing the hematoxylin and the eosin stain, to prevent protein degradation. 3. Defrost slides and fix sections in 70% ethanol for 1 min. Determine the optimum length of time of defrosting empirically; in practice, 4 min presents no problems for subsequent protein analysis and ensures adequate staining and retention of the section on the slide. However, the use of considerably shorter times (seconds) has also been advocated.
4. Stain with hematoxylin and eosin by immersing the slides as follows: 30 sec in Mayer’s hematoxylin Rinse in Milli-Q water Rinse again in Milli-Q water 10 sec in Scott’s water Rinse in Milli-Q water 20 sec in 1% eosin Rinse in Milli-Q water Rinse again in Milli-Q water.
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Staining times should be minimized to prevent possible interference with subsequent protein analysis techniques, while still allowing adequate visualization of the tissue. Other histochemical stains can be used in place of hematoxylin/eosin such as 1% (w/v) aqueous methyl green (stain for 1 min) or 0.025% (w/v) toluidine blue O in 50 mM sodium phosphate buffer, pH 5.5 (see APPENDIX 2E for buffer; stain for 5 sec). For all staining protocols, keep incubations to a minimum to reduce artifacts and interference with subsequent analysis.
5. Dehydrate sections by immersing slides as follows: 30 sec in 70% ethanol 1 min in 100% ethanol 5 min in xylene 5 min (again) in xylene. Use fresh ethanol and xylene solutions to ensure adequate dehydration of sections for optimal microdissection.
6. Allow sections to dry and proceed immediately to LCM. Protein remains intact for at least 2 hr following staining. However, dissect sections immediately to avoid absorption of moisture, which can affect LCM.
Carry out laser capture microdissection 7. Place the slide on the stage of the LCM apparatus and hold in position using the vacuum. 8. Place the CapSure LCM cap over the tissue section using the micromanipulator arm. 9. Locate an area for dissection. A visualizer can be temporarily inserted into the cap holder to improve the morphology of the tissue section while areas of tissue are being selected for dissection. The morphology of cryosections, which is inferior to that of paraffin-embedded material, can still be problematic, and it can be useful to stain and mount a parallel section for examination.
10. Select the laser diameter required for dissection. There are three laser sizes that can be employed for dissection—7.5, 15, and 30 ìm. Within each setting, laser size can be more finely adjusted by altering the power and duration of the laser pulse. Start with the default settings and initially fire at an area of the cap that is not covering tissue to check that the laser is focused and to gain an approximate idea of the diameter of the laser at those settings. Adjust as necessary prior to dissecting tissue and during dissection if necessary.
11. Fire the laser. Lift the cap at intervals to “capture” the tissue and check the dissection. If areas of tissue are to be dissected, the repetitive firing mode can be used, with the position of the laser being precisely adjusted using the joystick.
12. Following LCM, examine the cap surface to confirm the efficiency of capture. Captured tissue should be selectively removed when the cap is lifted. Two problems can occur: lifting of surrounding nonspecific tissue, which may be removed using the sticky edge of a “Post-It” note, and inefficient lifting of captured material. For further discussion of these problems, see Critical Parameters and Troubleshooting.
Laser Capture Microdissection for Proteome Analysis
Solubilize captured protein for subsequent analysis 13a. If very small volumes (≤10 ìl) of extraction buffer are to be used: Invert the cap and pipet buffer on to the cap surface, then incubate the cap in a humidified chamber or invert an Eppendorf Safe-Lock tube over the cap to prevent evaporation. Extract for 15 to 30 min.
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13b. With larger extraction volumes: insert the cap into an Eppendorf Safe-Lock tube containing buffer and invert the tube, taking care to mix thoroughly to ensure the tissue is completely coated with buffer. Extract for 15 to 30 min. Alternatively, strip the film off the cap and insert into buffer. Details of the specific extraction buffers and conditions used for different downstream analysis are provided in Support Protocol 2 and Basic Protocols 2 to 4; also see Critical Parameters and Troubleshooting.
14. Vortex and microcentrifuge to collect the extract. Check the cap surface to ensure tissue solubilization. IMMUNOLABELING OF TISSUE SECTIONS FOR LCM Immunolabeling provides an alternative to histochemical stains (see Basic Protocol 1) in the capture of cells of interest by LCM. Standard enzymatic-based detection systems may be suitable, but it would be necessary to confirm that these do not result in extensive protein modifications. If an LCM with a fluorescent attachment is being employed, use of fluorescent detection is an obvious choice. In other cases silver enhancement of gold labeling can be used prior to protein analysis, although yield can be affected and care must be taken to optimize the protocol to minimize selective protein loss and protein profile alterations. The choice of fixative is critical. Acetone may be ideal for some antibodies, but protein loss from the sections is far greater during processing. Ethanol results in better protein recovery but may not be suitable for some antibodies.
ALTERNATE PROTOCOL 1
Additional Materials (also see Basic Protocol 1) TBS (see recipe) Complete Mini Protease Inhibitor Tablets (Roche Molecular Biologicals) Acetone Primary antibody against protein of interest Gold-labeled secondary antibody (British BioCell International) Silver-enhancing kit (British BioCell International) Replace steps 1 to 5 of Basic Protocol 1 with the following: 1. Cut 8-µm sections from OCT embedded fresh frozen tissue onto ethanol-dipped glass slides and place on dry ice. The cryostat blade should be thoroughly cleaned with ethanol between samples to avoid contamination. Sections should ideally be used the same day, although they may be stored at −80°C for subsequent use. The effect of storage should be evaluated, as problems may be encountered with loss of antigen reactivity and protein degradation.
2. Defrost slides and fix sections in 70% ethanol or 100% acetone for 1 min. 3. Add one Complete Mini Protease Inhibitor tablet per 10 ml of TBS (for 1× final concentration of protease inhibitor cocktail). Dilute the primary antibody (against the protein of interest) to the appropriate concentration in the protease inhibitor–containing TBS. Incubate slide with primary antibody in TBS for 5 min. 4. Dilute the gold-labeled secondary antibody to the appropriate concentration in protease inhibitor–containing TBS. 5. Briefly wash the slide with TBS, then incubate with gold-labeled secondary antibody in TBS for 5 min.
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6. Briefly wash in water and treat with the silver-enhancing kit according to the manufacturer’s instructions. 7. Briefly wash in water, then counterstain by immersing slide as follows: 30 sec in Mayer’s hematoxylin Rinse in Milli-Q water Rinse again in Milli-Q water 10 sec in Scott’s tap water Rinse in Milli-Q water. 8. Dehydrate sections by immersing slides as follows: 30 sec in 70% ethanol 1 min in 100% ethanol 5 min in xylene 5 min (again) in xylene. 9. Allow sections to dry and proceed immediately to LCM (see Basic Protocol 1, steps 7 to 14). SUPPORT PROTOCOL 1
COLLECTION AND STORAGE OF TISSUE FOR ANALYSIS BY LCM The most important resource for approaches utilizing LCM is a good-quality tissue bank. This protocol describes the collection and processing of tissue samples following surgical resection, with the specific aim of preserving DNA, RNA, and protein integrity for downstream molecular analysis. An important factor in tissue banking is selection of representative areas of tissue, for example, macroscopically non-necrotic areas of tumor and distal normal tissue, for which it is essential to follow the guidance of a pathologist. For human samples (subject to the approval of relevant ethical bodies) where diagnostic use has to be prioritized, the cooperation of surgeons and pathologists is required. A critical factor for successful tissue sample banking is rapid processing to minimize any degradation or other changes in the protein composition. The authors routinely complete the whole process below within 30 min of the removal of tissue from the patient, with no apparent protein degradation. However, if circumstances allow processing to be completed more rapidly, this is obviously advantageous. NOTE: This protocol has been used successfully with kidney, bladder, cervix, and ovary samples, and is likely to be suitable for use with other tissues. However, this should be confirmed at an early stage in tissue banking and modified if necessary. Materials Tissue of interest freshly obtained from patient Transport medium (see recipe), ice-cold RPMI 1640 medium (Life Technologies) Complete Mini Protease Inhibitor Tablets (Roche Molecular Biologicals) PBS (see recipe), ice-cold Isotonic (250 mM) sucrose (8.6 g sucrose, H2O to 100 ml; prepare immediately before use), ice-cold OCT embedding medium (BDH) Liquid nitrogen
Laser Capture Microdissection for Proteome Analysis
Sterile disposable scalpels Cryostat chucks, precooled on dry ice
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1. Excise blocks of tissue from the sample using a sterile scalpel and place in ice-cold transport medium. 2. Cut into the required number of blocks of appropriate size for storage. The size is dependent on the tissue and subsequent use, but generally blocks of 3 to 5 mm in each dimension are suitable; this size allows rapid freezing and is adequate for subsequent sectioning.
3. Prepare ice-cold PBS in plastic petri dishes or other suitable containers on ice. Wash the tissue blocks briefly in the PBS, then in ice-cold isotonic sucrose. The sucrose wash is carried out to minimize salt carryover, which can interfere if whole tissue is used for electrophoresis.
4. Touch the tissue block against a piece of paper to remove excess liquid. Place a drop of OCT on the pre-cooled chuck, and once it has started to freeze, place the tissue on the OCT. Cover with a drop of OCT, taking care not to form air bubbles. Once completely frozen, remove the embedded tissue from the chuck. Material embedded in this way appears to be stable for several years when stored at −80°C or in liquid nitrogen.
QUANTIFICATION OF PROTEIN IN MATERIAL CAPTURED BY LCM USING SYPRO RUBY
SUPPORT PROTOCOL 2
The amount of protein captured by LCM is not usually sufficient for standard protein assays. If laser sizes are carefully controlled, normalization can be based on relative numbers of laser shots or cells captured, although obviously these may only be regarded as estimates. Alternative possibilities are immunoblotting for housekeeping proteins or comparison of a small aliquot by one-dimensional PAGE and silver staining against titrated standardized tissue lysates. With samples captured for SELDI-MS (see Basic Protocol 4) into 10 mM Tris⋅Cl, pH 8.0/1% (w/v) NP-40, a membrane-bound assay based on the fluorescent dye SYPRO Ruby (Molecular Probes) can be used. This assay is sensitive in the nanogram range and can tolerate some additional buffer components, such as 8 M urea. Materials 10 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 10 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E)/ 0.1% (w/v) NP-40 (prepare fresh; add NP-40 from 10% w/v stock solution) Bovine serum albumin (BSA) Samples from LCM (see Basic Protocol 1) in 10 mM Tris⋅Cl, pH 8.0/1% NP-40 SYPRO Ruby protein blot stain (Molecular Probes) MultiScreen-HA sterile 96-well plate (mixed cellulose ester membrane; Millipore) Fluorescence plate reader (BMG FLUOstar or equivalent reader with excitation filter at either 280 or 450 nm and emission filter at 618 nm) 1. Dilute samples (see Basic Protocol 1) 10-fold in 10 mM Tris⋅Cl, pH 8.0. 2. Prepare solutions of BSA (0 to 40 ng/ml) in 10 mM Tris⋅Cl, pH 8.0/0.1% NP-40 to construct a standard curve. 3. Pipet 5 µl of each standard (step 2) and diluted sample into a well of a sterile 96-well plate and incubate for 10 min at room temperature. Note the sterile plates must be used, as the nonsterile equivalents do not work with this assay, although the reasons for this are unknown.
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4. Wash wells four times with water, each time for 1 min. This is most easily achieved by alternately pipetting 200 ìl of water into each well and flicking to empty.
5. Dilute SYPRO Ruby 1:24 in water. Add 50 µl of the diluted stain to each well and incubate plate for 15 min in the dark with shaking. 6. Wash wells again as in step 4. 7. Read signals using a fluorescent plate reader (excitation filter at either 280 or 450 nm, emission filter at 618 nm). The standard curve will typically start to plateau at the higher protein concentrations. Samples may need to be diluted further to ensure that measurements fall within the linear portion of the standard curve. BASIC PROTOCOL 2
TWO-DIMENSIONAL GEL ELECTROPHORESIS OF MATERIAL CAPTURED BY LCM 2-D PAGE is the central separation tool in proteomic studies. This technique separates proteins on the basis of two independent characteristics: pI in the first dimension (isoelectric focusing) and molecular weight in the second dimension (SDS-PAGE). This technique allows up to 2000 proteins to be resolved in a single standard large-format gel. A description of the principles of 2-D PAGE and detailed experimental procedures can be found in UNIT 10.4. Here, a brief protocol for each of the steps involved in analysis of samples by 2-D PAGE is given, namely using immobilized pH gradient (IPG) strips with the IPGPhor apparatus (Amersham Pharmacia Biotech) for the first dimension, reducing SDS-PAGE for the second dimension, and silver staining for subsequent protein detection. CAUTION: See UNIT 10.4 for safety considerations regarding chemicals and electrophoretic apparatus used in this protocol. Materials Cells of interest captured by LCM (see Basic Protocol 1) 2-D PAGE lysis buffer (see recipe) Re-swell buffer (see recipe) DryStrip cover fluid (Amersham Pharmacia Biotech) Molecular-weight markers (Novex Mark12 unstained standards, or equivalent) 1% LMP agarose (see recipe) Equilibration buffer (see recipe) containing 10 mg/ml dithiothreitol (DTT) and a trace of bromphenol blue Equilibration buffer (see recipe) containing 40 mg/ml iodoacetamide and a trace of bromphenol blue SDS-PAGE running buffer (see recipe) 50% (v/v) methanol/10% (v/v) acetic acid OWL silver stain (Autogen Bioclear) 0.5-ml Eppendorf Safe-Lock tubes 18-cm pH 3-10NL IPG strips (Amersham Pharmacia Biotech) IPGphor and 18-cm strip holders (Amersham Pharmacia Biotech) Equilibration tray ISO-DALT tank, casting unit, and glass plates (Amersham Pharmacia Biotech) Gel scanner (Molecular Dynamics Personal Densitometer or equivalent)
Laser Capture Microdissection for Proteome Analysis
Additional reagents and equipment for 2D-PAGE (UNIT 10.4)
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Solubilize protein from captured material 1. Capture cells of interest (see Basic Protocol 1 through step 12). 2. Insert the cap into an Eppendorf Safe-Lock tube containing 50 µl 2-D PAGE lysis buffer. 3. Invert the tube and mix well to ensure the cap surface is completely coated in buffer. 4. Leave for ∼30 min at room temperature, then vortex. 5. Microcentrifuge briefly to collect the extract. To allow sufficiently concentrated samples to be prepared for 2-D PAGE, captured material from several successive caps can be solubilized into the same aliquot of 2-D PAGE lysis buffer. Lysates are stable for several hours at room temperature, but should be frozen if not analyzed the same day. Samples should not be freeze-thawed, but several aliquots collected over different days may be pooled for analysis.
Run the first dimension 6. Dilute the protein extract with re-swell buffer to give a final volume of 450 µl. Generally <100 ìl of the total volume is comprised of 2-D lysis buffer, since greater amounts can interfere with focusing, although adequate results have been obtained with up to 200 ìl. The optimal protein load for 18-cm 3-10NL IPG strips using this protocol is 30 ìg. However, with samples procured by LCM, significantly less material is generally obtained, with a concomitant decrease in the number of spots seen, but adequate profiles can be produced.
7. Pipet the sample into a strip holder, running evenly between the electrodes. 8. Thaw the IPG strip, remove the backing cover, and place it gel-side-down over the sample. It is important that the gel side of the strip not be handled and that the sample is spread evenly along the strip with no air bubbles trapped.
9. Overlay with DryStrip cover fluid and apply the strip holder lid. 10. Rehydrate and focus using the IPGphor program below with a surface temperature of 20°C, maximum power of 5 W, current limit of 0.05 mA per strip, and a total focusing time of 65 kV-h. 30 V 200 V 500 V 1000 V 1000 to 8000 V 8000 V
13 hr 1 hr 1 hr 1 hr 1 hr to end
Following isoelectric focusing, run the second dimension immediately or store strips between plastic sheets in an X-ray film cassette at –80°C. Run the second dimension For global protein profiling, the percentage of the resolving gel should be chosen to maximize the information in the protein profile obtained. Although this is sample-dependent, for analysis of whole tissue extracts a 10% T resolving gel is generally suitable or alternatively a gradient gel can be used. For details on how to pour gels using the DALT casting system, see UNIT 10.4 and the ISO-DALT user manual. Store resolving gels at 4°C for up to 2 weeks.
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11. Pour a 1-cm 4% T stacking gel on top of the resolving gel, allow it to set, then wash the top of the gel with SDS-PAGE running buffer. This is an optional step which few groups use, but the authors find that it results in increased visualization of high-molecular-weight proteins.
12. Dilute an appropriate volume of molecular-weight markers into 1% LMP agarose, pipet 20-µl drops onto Parafilm, and allow to set. Novex Mark12 unstained standards diluted 1 in 10 with agarose provide a suitable load for subsequent silver staining.
13. Equilibrate the strip (from step 10) for 15 min, gel-side-up, with 5 ml of equilibration buffer containing 10 mg/ml DTT and a trace of bromphenol blue (∼1 drop of 0.25% w/v per 10 ml). It is convenient to carry out strip equilibrations in an equilibration tray on a shaking platform.
14. Remove and re-equilibrate the strip for 10 min with 5 ml equilibration buffer containing 40 mg/ml iodoacetamide and a trace of bromphenol blue (∼1 drop of 0.25% w/v per 10 ml). 15. Decant the buffer from the gel from step 11 and insert a molecular-weight standard on top of the stacking gel, 1 cm in from one end.
3
pI
10 200
116 97
66 55
36.5
31
21.5
Laser Capture Microdissection for Proteome Analysis
Figure 22.3.2 An example of 2-D PAGE analysis of laser capture microdissected material. 30,000 15-µm laser shots of cervical epithelium were procured by laser capture microdissection (LCM). Protein extracts were prepared and separated by 2-D PAGE. 930 protein species were resolved (Craven and Banks, 2002; reproduced with permission of Academic Press).
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16. Remove the strip from equilibration buffer, rinse with SDS-PAGE running buffer, and place on top of the stacking gel with the cathode end next to the markers. Ensure that the strip lies flat with no air bubbles trapped between the strip and the gel surface.
17. Overlay with 1% LMP agarose and allow to set. 18. Electrophorese overnight at 12.5°C in SDS-PAGE running buffer at 18 mA/gel. Fix and stain the gel 19. Fix each gel overnight in 500 ml of 50% methanol/10% acetic acid. Alternatively gels can be fixed for 2 hr in 250-ml volumes of fixative with two changes.
20. Stain using OWL silver stain according to the manufacturer’s instructions, using 300-ml volumes for reagent steps and 500 ml for water washes. Alternative stains such as Coomassie Blue and SYPRO Ruby, or silver staining methods, can also be used.
21. Scan the gel using a Molecular Dynamics Personal densitometer or equivalent, and analyze (by eye or using gel-analysis software). Figure 22.3.2 shows an example 2-D PAGE analysis of laser capture microdissected material.
ANALYSIS OF MATERIAL CAPTURED BY LCM USING IMMUNOBLOTTING
BASIC PROTOCOL 3
Immunoblotting (western blotting) is a highly sensitive method for analysis of particular molecules of interest. Similarly, groups of molecules such as phospho- or glycoproteins can be analyzed with phosphoamino acid–specific antibodies or lectins, respectively. Proteins separated by 1-D or 2-D electrophoresis are transferred to a membrane for subsequent probing. For routine blotting procedures, Towbin’s transfer buffer (see Reagents and Solutions) and nitrocellulose membrane can be employed. For some proteins modifications such as inclusion of SDS or changes in the methanol concentration of the transfer buffer, or the use of PVDF membrane, can improve results. A full consideration of such issues is presented in UNIT 10.10. In a routine immunoblotting protocol, the membrane is blocked to prevent nonspecific binding, then probed with primary antibody. It is then probed with a secondary antibody directly conjugated to an enzyme—usually horseradish peroxidase (HRP)—and detection is achieved using chemiluminescence. For increased sensitivity, a biotinylated secondary antibody can be used, followed by streptavidin-HRP, in which case the additional step gives further amplification. One problem with such a strategy is detection of endogenous biotin-containing enzymes, which are abundant in many tissues including kidney and liver. An alternative high-sensitivity approach is the Envision system developed by Dako, which uses an HRP-labeled polymer conjugated to secondary antibodies. Developed for use in immunohistochemistry, this system gives good results in immunoblotting procedures. Envision reagents are currently available for use with rabbit and mouse primary antibodies. NOTE: The system described below is based on minigels, which are ideal for rapid use with the small amounts of material from LCM, but the protocols apply equally to large-format or 2-D gels. Gel-Based Proteome Analysis
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Materials Cells of interest captured by LCM (see Basic Protocol 1) SDS-PAGE lysis buffer (see recipe) SDS-PAGE running buffer (see recipe) Molecular weight markers (Invitrogen MultiMark multicolored standards, or equivalent) Towbin’s transfer buffer (see recipe) TBS-T (see recipe) containing 1%, 5%, and 10% (w/v) nonfat dry milk Primary antibody against protein of interest TBS-T (see recipe) Envision kit (Dako; available for use with rabbit or mouse primary antibodies) Super Signal (PerBio) 0.5-ml Eppendorf Safe-Lock tubes Minigel system (BioRad mini-PROTEAN II and Mini Trans-Blot cell or equivalent) Whatman filter paper Hybond C Super nitrocellulose membrane (Amersham Pharmacia Biotech) Additional reagents and equipment for one-dimensional SDS-PAGE (UNIT 10.1) and immunoblotting (UNIT 10.10) Solubilize protein from captured material 1. Insert the cap containing the captured cells (see Basic Protocol 1) into a 0.5-ml Eppendorf Safe-Lock tube containing 25 µl SDS-PAGE lysis buffer. 2. Invert the tube and mix well to ensure the cap surface is completely coated in buffer. 3. Leave for ∼30 min at room temperature. 4. Microcentrifuge briefly to collect the extract. Run SDS-PAGE gel and transfer protein to membrane 5. Separate the proteins by 1-D SDS-PAGE (UNIT 10.1) using a mini-gel system and a resolving gel percentage suitable for the molecular weight of the protein being analyzed. Also include molecular weight standards. Electrophorese at 125 V in SDS-PAGE running buffer. 6. Equilibrate gel in Towbin’s transfer buffer for 30 min. 7. Cut six Whatman filter papers and nitrocellulose membranes to the size of the gel and presoak in Towbin’s transfer buffer. Also see UNIT 10.10 for general principles of immunoblotting.
8. Prepare the gel-transfer sandwich and assemble the blotting module, including the Bio-Ice cooling unit, according to the manufacturer’s instructions. 9. Transfer with stirring at 100 V (160 to 200 mA) for 1 hr at room temperature. Carry out immunoblotting The following steps should be carried out at room temperature with gentle agitation on an orbital shaker. 10. Block for 2 hr in TBS-T containing 10% (w/v) nonfat dry milk. Laser Capture Microdissection for Proteome Analysis
For certain applications such as probing with phospho-specific antibodies or lectins, milk proteins are unsuitable blocking agents. Some milk preparations may also be unsuitable
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1
2
3 major vault protein
b-Actin
Figure 22.3.3 Immunoblot analysis of laser captured renal cell carcinoma tissue. 1250 (lane 1), 500 (lane 2), and 250 (lane 3) 7.5 µm laser shots of renal cell carcinoma were analyzed by immunoblotting with antibodies to major vault protein (upper panel) and β-actin (lower panel) (Craven and Banks, 2002; reproduced with permission from Academic Press).
for streptavidin-based detection systems due to their biotin content. Alternatives include bovine serum albumin (BSA) or human serum albumin (HSA).
11. Dilute primary antibody to appropriate concentration in TBS-T containing 1% (w/v) non-fat dry milk and incubate blot at room temperature for 1 to 2 hr. It may be necessary to incubate some antibodies overnight at 4°C.
12. Wash with TBS-T four times, each time for 5 min. 13. Incubate for 1 hr in Envision diluted 1:100 to 1:1000 in TBS-T containing 5% (w/v) nonfat dry milk. Alternatively, incubate in biotinylated secondary antibody diluted in TBS-T containing 1% (w/v) nonfat dry milk, then wash four times in TBS-T, each time for 5 min, and incubate 1 hr in streptavidin-HRP diluted in TBS-T containing 1% (w/v) nonfat dry milk.
14. Wash with TBS-T four times, each time for 5 min. 15. Develop with Super Signal according to the manufacturer’s instructions. Figure 22.3.3 shows an immunoblot of laser-captured renal cell carcinoma tissue.
SELDI MASS SPECTROMETRY This protocol describes the use of SELDI mass spectrometry (Ciphergen Biosystems) to analyze LCM-procured samples (see Fig. 22.3.4). Following LCM, protein is solubilized with one of several possible extraction buffers, quantified, and applied to the surface of a ProteinChip. ProteinChips consist of metal strips containing defined “sample spots” (usually 8-, 16- or 24-spot) with specific surface coatings including those designed to immobilize hydrophobic molecules, anions, cations, or metal-binding proteins. Additionally, ProteinChips with preactivated surfaces are available, which can be used to immobilize antibodies and immunoprecipitate specific proteins from a complex mixture. After sample application and addition of sample matrix (an energy-absorbing molecule that mediates ionization of the sample), the ProteinChips are analyzed using the SELDI-MS apparatus. The principles of MALDI mass spectrometry are described in detail in UNITS 16.3 & 16.4. Essentially, a laser beam is fired at the sample, causing ionization of the analytes. As the molecules become positively charged and desorb, changing state from solid phase to gaseous phase, they “fly” from the positive surface of the ProteinChip on application of a voltage differential and are detected by a detector, in this case a “time-of-flight” (TOF) detector which determines the time taken for the molecules to leave the ProteinChip
BASIC PROTOCOL 4
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laser
2. Nonspecifically bound proteins and components such as salts and detergents are washed away.
ionization
Intensity
1. Sample is applied directly to the ProteinChip spot surface. In appropriate binding buffer, proteins bind via specific and nonspecific interactions.
Mass/charge 3. EAM is added.
4. Protein mass profile is determined by SELDI-TOF mass spectrometry.
5. Data are collected and analyzed.
Figure 22.3.4 Illustration of the basic principles of SELDI sample analysis as described in the text.
surface and reach the detector. Because the flight times of molecules are proportional to their mass, a profile of molecules ionized is generated with each peak resolved on the basis of mass/charge ratio. The high sensitivity of this technology (femtomole to attomole range) allows the use of small amounts of sample such as that generated from LCM. The method is ideally suited for analyzing molecules <20 kDa in size, thus complementing analyses by techniques such as 2-D PAGE. The identification of proteins present in specific peaks represents a challenge because, although tentative identifications can be made on the basis of mass, this has to be confirmed independently, e.g., using antibodies to deplete the sample and then monitoring the changes in peak profile. Alternatively mass spectrometric sequencing can be used, generally following on-chip or larger-scale purification strategies. The use of SELDI for analyzing LCM-captured material is relatively new, and hence protocols are continuously evolving. Factors such as solubilization, binding, and wash buffers, as well as the ProteinChip type and sample matrix, influence the results obtained. Sample processing can also be important. These factors need to be determined empirically (see Commentary). The SELDI running protocols and data analysis methods are complex and beyond the scope of this protocol. They are described in the large user manuals supplied with the equipment, and the equipment should only be run with the technical assistance of experienced personnel. Particular issues that should be considered are the reproducibility of sample and ProteinChip preparation and calibration and normalization of the SELDIMS data. Figure 22.3.5 shows an example of SELDI analysis of cervical epithelium and stromal fractions from laser capture microdissected material.
Laser Capture Microdissection for Proteome Analysis
CAUTION: Extreme care needs to be exercised when using several of the reagents, particularly trifluoroacetic acid (also see APPENDIX 2A).
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10 kDa
12 kDa
14 kDa
16 kDa
stroma
epithelium
Figure 22.3.5 An example of SELDI analysis of cervical epithelium and stromal fractions from laser capture microdissected material. 2500 15-µm laser shots of cervical epithelium were microdissected and analyzed by SELDI using an NP2 ProteinChip. The profile of the stromal tissue is also shown. Differences in spectral profiles are evident, with a much more complex profile, containing more peaks, in the epithelial sample (Craven and Banks, 2002; reproduced with permission from Academic Press).
Materials Cells of interest captured by LCM (see Basic Protocol 1) 10 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E)/ 0.1% (w/v) NP-40 (prepare fresh; add NP-40 from 10% w/v stock solution) 20% (v/v) acetonitrile/0.2% (v/v) trifluoroacetic acid (see recipe) Sinapinic acid matrix solution, saturated (see recipe) 50% acetonitrile (see recipe) 10% (v/v) acetonitrile/0.1% v/v trifluoroacetic acid (see recipe for 20% acetonitrile/0.2% TFA; dilute 1:1 with H2O) 10 mM HCl 100 mM ammonium acetate pH 6.5 (see recipe) 50 mM nickel sulfate (see recipe) 2× PBS/NaCl (see recipe) SlickSeal low-protein-binding tubes (National Scientific Supply) NP2, H4, SAX2, WCX2, and/or IMAC3 ProteinChips (Ciphergen) SELDI mass spectrometer (Ciphergen) Humidified chamber (prepared by placing wet tissues inside the base of a plastic pipet-tip box) Solubilize of proteins from captured material 1. Invert the cap containing the captured cells (see Basic Protocol 1) and add 5 µl of 10 mM Tris⋅Cl, pH 8.0/1% NP-40 directly to the cap surface to cover the captured material. 2. Incubate for 15 min at 4°C. 3. Using a fine tip, remove the solubilized protein, pipetting the solution up and down several times to ensure maximal recovery. 4. Transfer the solubilized material to a SlickSeal tube. Proceed with minimum delay to SELDI analysis, keeping the sample at 4°C, or store sample at –80°C. Gel-Based Proteome Analysis
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Prepare the ProteinChip Separate sets of alternative steps for each type of ProteinChip are provided below. The amount of protein loaded per ProteinChip should be standardized, preferably by total protein quantification; loading by cell number is possible although this procedure is less accurate and may influence the profiles generated. The exact amount to be loaded should be determined on a case-by-case basis, but peaks can be seen from analysis of as few as 50 cells; however, far more complex profiles are generated with higher protein loads. Larger volumes of solutions can be loaded using the SELDI bioprocessor, details of which can be found in the SELDI user manuals. NOTE: Care should be taken when applying solutions to the ProteinChip surface to avoid touching the ProteinChip surface with the pipet tip. In addition, powder-free gloves should be worn, as spurious peaks can be generated by inadvertent powder contamination. The analysis should be done as soon as possible due to the deterioration of matrix solution and the ProteinChip should be stored in the dark until analyzed. For the NP2 ProteinChip (normal phase) 5a. Mix the solubilized LCM sample (from step 4) with an equal volume of 20% acetonitrile/0.2% TFA and apply 5 µl to the NP2 ProteinChip surface. 6a. Allow to dry. 7a. Wash each sample spot with three separate water washes and allow to dry. For each wash, rapidly pipet 5 ìl of water up and down ∼10 times using a micropipettor with a fine tip.
8a. Add 0.35 µl of saturated sinapinic acid matrix solution to each spot and allow to dry. 9a. Repeat step 8a. 10a. Analyze the ProteinChip by SELDI according to the manufacturer’s instructions for the SELDI mass spectrometer. For the H4 protein chip (hydrophobic) 5b. Pretreat each sample spot of the H4 ProteinChip by adding 5 µl of 50% v/v acetonitrile. 6b. Mix the solubilized LCM sample (from step 4) with an equal volume of 20% acetonitrile/0.2% TFA. 7b. Remove the acetonitrile from each sample spot and apply 5 µl of sample to the H4 ProteinChip surface, taking care not to allow the sample spots to dry. 8b. Incubate the ProteinChip in a humidified chamber for 20 min. 9b. Remove the ProteinChip from the humidified chamber and wash each sample spot four times with 10% acetonitrile/0.1% TFA. The first two washes should be performed by rapidly pipetting 5 ìl of 10% acetonitrile/0.1% TFA up and down ∼10 times for each wash using a micropipettor with a fine tip. The second two washes should be performed by leaving the wash solution on the sample spot for 5 min.
10b. Remove the final wash and allow to dry. 11b. Add 0.35 µl of saturated sinapinic acid matrix solution to each sample spot and allow to dry. Laser Capture Microdissection for Proteome Analysis
12b. Repeat step 11b.
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13b. Analyze the ProteinChip by SELDI according to the manufacturer’s instructions for the SELDI mass spectrometer. For the SAX2 ProteinChip (strong anion exchange) 5c. Wash the SAX2 ProteinChip sample spots by adding 5 µl of 10 mM Tris⋅Cl, pH 8.0/1% NP-40 to each sample spot. After 5 min, remove the solution and replace with a further 5 µl of 10 mM Tris⋅Cl, pH 8.0/1% NP-40 and leave for a further 5 min. 6c. Mix the solubilized LCM-captured sample (from step 4) with an equal volume of 10 mM Tris⋅Cl, pH 8.0/1% NP-40. Remove the wash solution from the sample spots and apply 5 µl of the diluted sample to each sample spot. 7c. Incubate in a humidified chamber at room temperature for 30 min. 8c. Remove the ProteinChip and perform five separate washes with 10 mM Tris⋅Cl, pH 8.0/1% NP-40, followed by two separate water washes, on each sample spot. For each wash, rapidly pipet a 5-ìl volume up and down ∼10 times using a micropipettor with a fine tip.
9c. Remove the water and add 0.35 µl of saturated sinapinic acid matrix solution to each sample spot while it is still moist. Allow to dry. 10c. Repeat step 9c. 11c. Analyze the ProteinChip by SELDI according to the manufacturer’s instructions for the SELDI mass spectrometer. For the WCX2 ProteinChip (weak cation exchange) 5d. Prepare the WCX2 ProteinChip surface by adding 5 µl of 10 mM HCl to each sample spot surface and then incubating in a humidified chamber for 10 min at room temperature. 6d. Remove the HCl and wash each sample spot with water for 3 separate washes. For each wash, rapidly pipet 5 ìl of water up and down ∼10 times using a micropipettor with a fine tip
7d. Remove the water from each sample spot and replace with 5 µl of 100 mM ammonium acetate pH 6.5. Allow to stand for 5 min. 8d. Mix equal volumes of solubilized LCM captured material (from step 4) and 100 mM ammonium acetate, pH 6.5. 9d. Remove the ammonium acetate solution from the sample spots and apply 5 µl of sample to the sample spot surface. 10d. Incubate for 30 min in a humidified chamber. 11d. Using the same technique as in step 6d, perform three separate washes of each sample spot with 100 mM ammonium acetate, pH 6.5, followed by two washes with water. 12d. Add 0.35 µl of saturated sinapinic acid matrix solution to each sample spot while it is still moist, and allow to dry. 13d. Repeat step 12d. 14d. Analyze the ProteinChip by SELDI according to the manufacturer’s instructions for the SELDI mass spectrometer. Gel-Based Proteome Analysis
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For the IMAC3 ProteinChip (immobilized metal affinity) 5e. Add 5 µl of 50 mM nickel sulfate to the sample spots and incubate for 5 min at room temperature. Remove the solution and repeat the process. 6e. Remove the nickel sulfate and perform two washes with 5 µl of 1× PBS/NaCl using the same technique as in as in step 5e. 7e. Mix solubilized LCM-captured material (from step 4) with 2× PBS/NaCl in equal volumes and apply 5 µl to each sample spot surface. 8e. Incubate the ProteinChip in a humidified chamber for 1 hr. 9e. Remove the sample solution from each sample spot and wash five times with 1× PBS/NaCl, then twice with water. For each wash, rapidly pipet a 5-ìl volume up and down ∼10 times using a micropipettor with a fine tip.
10e. Add 0.35 µl of saturated sinapinic acid matrix solution to each sample spot while it is still moist, and allow to dry. 11e. Repeat step 10e. 12e. Analyze the ProteinChip by SELDI according to the manufacturer’s instructions for the SELDI mass spectrometer. REAGENTS AND SOLUTIONS Use Milli-Q purified water or equivalent for the preparation of all solutions. All chemicals should be of at least Analar grade or equivalent unless otherwise indicated. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Acetonitrile, 50% (v/v) Prepare by mixing equal amounts of HPLC-grade acetonitrile with water. Make fresh solutions each day to avoid potential concentration change due to evaporation. Take care to use solvent-resistant containers such as polypropylene microcentrifuge tubes. Acetonitrile (20% v/v)/trifluoroacetic acid (0.2% v/v) 200 µl HPLC-grade acetonitrile 798 µl H2O 2 µl trifluoroacetic acid (TFA; concentrated, sequencing grade) This solution should be made on the day of use. The stock TFA should be colorless; if it is brown or yellow it should be discarded using methods determined after consultation with the local Safety Officer. Extreme care should be taken when using TFA, as is it is an extremely hazardous chemical and the concentrated acid should only be used in a fume hood and dispensed wearing suitable protective clothing. Consult the Material Safety Data Sheet for handling details and disposal.
Agarose, 1% (w/v) 1 g low-melting point (LMP) agarose 1× SDS-PAGE running buffer (see recipe) to 100 ml Add a few drops of 0.25% (w/v) bromphenol blue Mix, then dissolve with heating in a microwave oven Make fresh on day of use Laser Capture Microdissection for Proteome Analysis
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Ammonium acetate, 100 mM, pH 6.5 0.77 g ammonium acetate Dissolve in H2O Adjust pH to 6.5 with acetic acid H2O to 100 ml Store up to 3 months at 4°C Equilibration buffer 180 g urea 150 ml glycerol 10 g SDS 50 ml 0.5 M Tris⋅Cl, pH 6.8 (APPENDIX 2E) H2O to 500 ml Store at room temperature for up to 2 weeks. Final concentrations are 6 M urea, 30% (v/v) glycerol, 2% (w/v) SDS, 0.05 M Tris⋅Cl, pH 6.8.
Mayer’s hematoxylin Prepare solution 1 by mixing: 3 g hematoxylin 20 ml 100% ethanol Prepare solution 2 by dissolving the following in 850 ml of H2O, in the order indicated: 0.3 g sodium iodate 1 g citric acid 50 g chloral hydrate 50 g aluminium potassium sulfate Mix solutions 1 and 2. Add 120 ml of glycerol and mix again. Store in dark bottles up to several months at room temperature; filter on the day of use. Nickel sulfate, 50 mM 1.31 g NiSO4⋅6H2O 100 ml H2O Store up to 1 month at 4°C 2-D PAGE lysis buffer 21.9 g urea 7.9 g thiourea H2O to 50 ml Add 0.5g Amberlite MB-1 ion-exchange resin and stir at room temperature for 10 min. Filter through Whatman no. 1 filter paper. To 48 ml of the above add: 2 g CHAPS 0.5 g DTT 1 ml 40% stock solution of Pharmalyte pH 3 to 10 (Amersham Pharmacia Biotech) 500 ml Pefabloc (Roche Molecular Biologicals; 100 mg/ml stock) Store in aliquots at −80°C; do not refreeze thawed aliquots Final concentrations are 7 M urea, 2 M thiourea, 4% (w/v) CHAPS, 1% (w/v) DTT, 0.8% (v/v) Pharmalyte pH 3 to 10, and 1 mg/ml Pefabloc.
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PBS, pH 7.3 1 tablet of Dulbecco’s A PBS (Oxoid) H2O to 100 ml Make fresh PBS/NaCl, 2× 2.92 g NaCl 1 tablet of Dulbecco’s A PBS (Oxoid) 100 µl 10% (w/v) NP-40 (Roche) H2O to 50 ml Store up to 1 month at 4°C Re-swell buffer 21.9 g urea 7.9 g thiourea H2O to 50 ml Add 0.5g Amberlite MB-1 ion-exchange resin and stir at room temperature for 10 min. Filter through Whatman no. 1 filter paper. To 48 ml of the above, add: 2 g CHAPS 230 mg DTT 250 µl 40% stock solution of Pharmalyte pH 3 to 10 (Amersham Pharmacia Biotech) Add 3 to 5 drops 0.25% (w/v) bromphenol blue per 50 ml buffer Store up to 3 months at −20°C Final concentrations are 7 M urea, 2 M thiourea, 4% (w/v) CHAPS, 0.46% (w/v) DTT, 0.2% (v/v) Pharmalyte pH 3to 10, and a trace of bromphenol blue.
Scott’s water 100 ml of 10% (w/v) magnesium sulfate 50 ml of 7% (w/v) sodium bicarbonate H2O to 1 liter Store up to 6 months at room temperature SDS-PAGE lysis buffer 2.0 ml 0.5 M Tris⋅Cl, pH 6.8 (see recipe) 1.6 ml glycerol 3.2 ml of 10% (w/v) SDS 0.8 ml 2-mercaptoethanol 0.4 ml 0.1% (w/v) bromphenol blue This gives a 2× solution. Store up to 3 months at 4°C
SDS-PAGE running buffer, 10× 30.3 g Tris base 144.0 g glycine 10.0 g SDS H2O to 1 liter Store at room temperature To make working solution, dilute 1 in 10 with H2O Final concentrations for the working solution are 25 mM Tris, 192 mM glycine, and 1% (w/v) SDS. When using with the Iso-DALT tank. it is more convenient to make 1× running buffer up in the tank according to the manufacturer’s instructions. Laser Capture Microdissection for Proteome Analysis
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Sinapinic acid matrix solution (saturated) Combine the following, in the order indicated: 5 mg sinapinic acid matrix (Ciphergen) 125 µl HPLC-grade acetonitrile 125 µl 1% (v/v) trifluoroacetic acid (TFA) Vortex thoroughly Allow to stand 5 min Microcentrifuge 2 min at maximum speed Store protected from light Prepare fresh each day TBS, pH 7.6, 10× 24.2 g Tris base 80 g NaCl Dissolve in H2O Adjust pH to 7.6 with HCl H2O to 1 liter Store at room temperature To make working solution, dilute 1 in 10 with H2O (final concentrations of the working solution are 20 mM Tris⋅Cl and 140 mM NaCl). TBS-T 100 ml 10× TBS (see recipe) 1 ml Tween 20 899 ml H2O Final concentrations are 20 mM Tris⋅Cl, 140 mM NaCl, and 0.1% (v/v) Tween 20.
Towbin’s transfer buffer 3.03 g Tris base 14.4 g glycine 200 ml methanol H2O to 1 liter Final concentrations are 25 mM Tris, 192 mM glycine, and 20% (v/v) methanol.
Transport medium Dissolve one Complete Mini Protease Inhibitor Tablet in 20 ml RPMI 1640 medium. Prepare fresh immediately before use and keep on ice. COMMENTARY Background Information Cell enrichment or purification strategies The diversity and complexity of the normal cellular makeup of tissues or organs, which reflects their varying biological functions, can be altered not only during pathological states but also as a result of normal physiological changes. For example, profound cellular changes can be seen in the ovary and endometrium during the menstrual cycle. In disease, there may not only be alterations in the cells that are normally present—e.g., the development of malignant clones—but other cell types may also be present and environmental
influences on the affected tissue may change. In cancer, for example, increased angiogenesis, the presence of infiltrating cells, and areas of hypoxia are often seen. It is therefore a challenge to unravel the changes occurring in specific cellular compartments with the use of whole tissue generating a significant amount of background “noise” against which results must be interpreted. The manual approaches used to address this problem include gross dissection of tissue samples with a scalpel, manual microdissection, and selective ultraviolet ablation of unwanted areas of tissue. However, limitations of accuracy with these methods and the relatively high
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risk of contamination have made other approaches desirable. Alternative strategies for purifying specific cell types, for example, using enzymatic digestion of tissues followed by manual removal of epithelial layers, or employing immunoaffinity methods, have been used with some success. However, the former techniques may be influenced by the generation of in vitro artifacts and the latter limited by the availability of suitable antibodies. Consequently, there has been much interest in the potential improvement offered by the use of innovative laser-assisted microdissection systems to selectively procure specific cell types for subsequent proteomic analysis.
Laser Capture Microdissection for Proteome Analysis
Laser-assisted microdissection systems There are now several commercially available laser-based microdissection systems that have been developed specifically for enriching specific cell types from tissue sections. The methodology incorporated into this unit has very much focused on the use of the first developed system, i.e., the Pixcell LCM (Arcturus). This system was developed in collaboration between scientists at the National Institutes of Health in the U.S. and the bioengineering company Arcturus (Emmert-Buck et al., 1996). This system is the one used in the authors’ laboratory and has been the most widely used to date. However many aspects of the protocols and the experimental considerations discussed therein apply equally to the other systems, which differ only in the “mechanics” of the dissection. The principle underlying the cell capture of the PixCell LCM system is shown in Figure 22.3.1, and is based on an inverted microscope with a near-infrared laser mounted above it, a manipulator arm, and a computerized imagestorage system. Fixed and stained sections are placed on the microscope platform, and a Perspex cap with a 6-mm-diameter transparent thermo-labile ethylene vinyl acetate film on its lower surface is then placed on the section, with the film in direct contact with the tissue. The laser beam (variable diameter 7.5 to 30 µm) is fired through the cap at selected cells, and the heat generated causes localized melting of the film and fusion with the targeted cells below. Heat damage to the tissue is minimized by using relatively short laser pulses (up to 5 msecs). When the cap is then lifted from the section, the dissected cells, fused with the cap, are also removed and can be solubilized using an appropriate extraction buffer. Recently a semi-automated version of the PixCell has been developed.
The alternative laser-assisted microdissection approaches are noncontact and nonheating and are based on ablating tissue around the cells of interest. The PALM system (PALM Mikrolaser Technologie; http://www.palm-mikrolaser.com/) uses “laser microbeam microdissection” (LMM) with “laser pressure catapulting” (LPC; Schutze and Lahr, 1998). An ultraviolet laser microbeam cuts around the area of interest, with a membrane backing often used to support the section and prevent contamination. After microdissection, a brief pulse of the laser is used to catapult the dissected tissue into a collecting tube containing the appropriate extraction buffer. The most recently developed commercially available system, the Leica AS LMD (Leica), employs a similar microdissection strategy, but the slide is inverted, allowing freed dissected material to be collected in a tube below the section, largely under the force of gravity (Kolble, 2000). Both these systems are semiautomated, with computerdriven dissection of areas delineated on a computer screen. The relative advantages and disadvantages of each system have been reviewed (Cornea and Mungenast, 2002) with particular regard to resolution, ease of use, sample preparation protocols, and sample visualization. Compatability of LCM and protein analysis These laser-assisted microdissection systems have been used extensively in studies utilizing nucleic acids, where amplification by PCR potentially allows analysis down to the single-cell level. Such amplification is not possible with protein-based approaches, and hence far more dissection is generally required. Fewer studies have used this approach so far; however, there are an increasing number of papers in the literature illustrating the potential of such sample selection in proteomics-based approaches. The initial report describing protein extraction and global profiling of laser captured cells generated from frozen tissue sections after brief fixation and hematoxylin and eosin staining (Banks et al., 1999) demonstrated the feasibility of using LCM with 2-D PAGE. Subsequently, alternative tissue processing regimens involving staining with methyl green or toluidine blue, as well as immunolabeling techniques, have been shown to be compatible with LCM and 2-D PAGE. However, staining protocols should be optimized for specific tissue types, as the degree of staining can adversely affect the focusing of the first-dimension gels and the recovery of protein (Craven et al., 2002). This is also the case for subsequent
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analysis of stained material by SELDI-MS, where effects of staining on some peaks have been seen (R.E. Banks, unpub. observ.). Several additional factors to be considered are very tissue-dependent and can only be determined empirically. Tens of thousands of laser shots/cells with dissection times of several hours or even days per sample may be needed in order to generate enough material for subsequent analysis by 2-D PAGE, although much less sample is needed for approaches such as immunoblotting and SELDI. The degree of purity of the microdissected cells will vary with the tissue and cell type being obtained, and the benefit that LCM affords in terms of enrichment in the protein profile relative to the starting material should also be considered (Craven et al., 2002). Applications of LCM in proteomics Several successful LCM/2-D PAGE studies have now been carried out. As an early example, the comparative analysis of approximately 50,000 normal and malignant esophageal cells dissected from two patient samples demonstrated 98% similarity between the normal and malignant protein profiles of almost 700 resolved proteins. Ten proteins were present uniquely in tumor and seven in normal epithelium. The identification of annexin I as one of the proteins lost in tumors (Emmert-Buck et al., 2000) was subsequently validated using immunoblotting to examine further microdissected patient samples, confirming complete loss or dramatic reduction of annexin 1 in both esophageal and prostate cancers (Paweletz et al., 2000a). This change was also shown to occur in premalignant lesions. Such studies elegantly illustrate the use of 2-D PAGE, with its requirement for large amounts of microdissected tissue, as the initial step to determine differentially expressed proteins, followed by validation on a larger sample set using immunoblotting, with the necessity for much less material. With the more sensitive chemiluminescent detection systems now available and sensitivity levels in the picogram range for many proteins, depending on antibody quality, several studies have shown that protein peaks can be profiled using as few as 200 to 2500 microdissected cells, depending on the protein abundance. The combination of microdissection and immunoblotting complements immunohistochemistry, confirming the cellular localization but additionally providing information on the form of protein in terms of its molecular size. Although not covered specifi-
cally in the protocols here, LCM has also been used to obtain cells for subsequent immunoassay of specific analytes, in this case the tumor marker prostate specific antigen (PSA) in normal and malignant prostate epithelium (Simone et al., 2000). The range of values in cancer cells was up to 7-fold higher than in normal cells, although there was overlap between the sets of samples, and PSA concentrations from LCMcaptured tissue broadly reflected the immunohistochemical expression of this marker. The first study to use SELDI-MS to analyze microdissected material was part of a larger study examining the specific forms of several prostate cancer markers in tissue samples and biological fluids. Several proteins were found to be differentially expressed between microdissected normal and malignant cells when analyzed on an anion-exchange ProteinChip, with a peak detected only in the prostate carcinoma samples being found to have the same molecular weight as purified prostate-specific membrane antigen (PSMA; Wright et al., 1999). These lysates were generated from only 2000 to 3000 microdissected cells. Other preliminary studies using SELDI profiling of microdissected tissue lysates illustrate the potential for this approach in comparing normal and diseased tissues, although no large-scale studies have yet been carried out, and the full selective capabilities of the different ProteinChip surfaces have not yet been exploited. However, even without the benefit of a chipsurface-based selection, a study using a single mastectomy sample demonstrated that direct analysis of 1250 cells following addition of sinapinic acid to the cap film, immobilized on a standard stainless steel MALDI target, could generate distinct reproducible spectra containing 20 to 50 peaks from four cell types (Palmer-Toy et al., 2000). Issues regarding which extraction buffers are optimal for subsequent mass spectrometric analysis (MALDI or SELDI) have yet to be addressed widely. Similarly, the amounts of material needed per sample may vary with ProteinChip type, extraction buffer, and tissue, but reports indicate that reproducible protein “fingerprints” can be generated using as few as 25 to 100 cells per sample (Paweletz et al., 2000b; Von Eggeling et al., 2000). However, far more protein peaks can be seen with increasing loads, and generally 500 to 2000 cells dissected using either the PixCell LCM or PALM systems have been used to demonstrate discriminatory profiles with maximal information (Paweletz et al., 2000b; Palmer-Toy et al., 2000; Westphal et al., 2002).
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It is extremely likely that laser-assisted microdissection technologies will form part of the armamentarium in the development of protein microarrays—an approach being explored increasingly in various formats to allow the simultaneous high-throughput profiling of many proteins in large numbers of samples, with only relatively small amounts of sample being required. Using a reversed-phase array approach, solubilized microdissected prostate tissue has been used to generate several microarrays (up to 1000 tissue lysate spots/slide) using lysates resulting from only 500 to 3,000 laser shots (typically representing 2,500 to 15,000 cells). By probing the arrays with a range of antibodies, the transition from normal epithelium through prostate intraepithelial neoplasia (PIN) to invasive cancer was mapped with decreased levels of cleaved poly(ADP-ribose) polymerase (PARP) and cleaved caspase-7, increased phosphorylation of Akt, and decreased phosphorylation of ERK being interpretated as representing activation of pro-survival events during tumor progression (Paweletz et al., 2001). Antibody arrays have also been used to analyze microdissected samples (2,500 to 3,500 cells) of squamous cell carcinoma of the oral cavity, where stromal and epithelial compartments have been analyzed through distinct stages of disease progression (Knezevic et al., 2001). Multiple protein differences were found in both diseased epithelial cells and surrounding stromal cells, with many of the identified proteins being involved in signal transduction pathways, illustrating the potential importance of communications between different tissue elements in cancer.
Critical Parameters and Troubleshooting
Laser Capture Microdissection for Proteome Analysis
Only aspects of the tissue processing and preparation, the laser capture microdissection processes per se, and tissue solubilization are discussed in this section. Issues relating to gel electrophoresis are discussed in UNITS 10.1 & 10.4 and those related to immunoblotting in UNIT 10.10. Critical paramters for immunohistochemical staining are discussed in Hofman (2002). Maintaining protein integrity and maximizing protein recovery are critical aspects of an LCM-based approach. With carefully controlled sample banking and storage, rapid staining protocols, inclusion of protease inhibitors in the stains, and the use of extraction buffers optimized to prevent protein degradation, proteins can be successfully extracted for downstream
analysis. If protein degradation appears to be a problem, examination of non-processed tissue sections should be performed to eliminate the possibility that tissue banking per se is the source of the problem. If the banked tissue is of good quality, then the subsequent processing steps should be examined. Protein degradation can be encountered if sections are stored for prolonged periods either at −80°C or on dry ice prior to staining or following staining and prior to LCM. However, tissue section processing (fixation and staining) is perhaps the stage at which most problems can occur with respect to total and selective protein recovery, and it is essential to determine the extent and variability of in vitro generated artifacts. Ethanol has been used successfully as a fixative in a number of studies, but acetone fixation, which gives improved results with many antibodies in immunohistochemistry, has been associated with impaired protein recoveries. In general, staining should be kept to a minimum to reduce problems with downstream analysis. For example, overstaining with eosin or toluidine blue interferes with isoelectric focusing. Incomplete tissue removal following dissection is one of the most frequently encountered problems. If sections are of varying thickness or uneven, the interface between the film and section may not be adequate to allow capture of cells. This may also occur following repositioning of the cap if large numbers of cells have been captured onto the film surface. Inefficient lifting is also caused by moisture in the tissue sections, which can be overcome by ensuring that fresh ethanol and xylene are used for tissue section dehydration and by processing sections immediately prior to dissection and controlling room temperature and humidity. The laser settings (power and duration) can also be altered to improve tissue transfer to the cap and changing the length of time slides are defrosted prior to dissection may improve dissection efficiency. The choice of slides is also critical. SuperFrost Plus charged slides or poly-L-lysine-coated slides may aid adhesion of tissue sections during staining procedures, but the adhesive forces may be too great to allow good capture and lift of dissected material. Plain glass microscope slides usually result in sufficient immobilization of sections to prevent loss during processing, but generally allow capture of dissected material more readily. Section thickness can also be reduced, although this naturally impacts on protein recovery. At the other end of the spectrum are problems associated with nonspecific tissue re-
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moval, resulting in sample contamination. PrepStrip Tissue Preparation Strips can be used to remove loosely bound material from the section surface prior to dissection, or the adhesive edge of a Post-It note can be used to remove nonspecific tissue from the cap surface. Alternatively CapSure HS caps (Arcturus) can be used for non-contact LCM. The laser diameter should also be carefully controlled to reduce capture of unwanted tissue areas. SELDI extraction buffers, ProteinChip loading buffers, and sample matrix The Tris⋅Cl/NP-40 buffer described is fully compatible with SELDI analysis but is not optimal for total protein solubilization. Alternative buffers have been described for solubilizing microdissected samples and incorporate a range of chaotropes (such as urea), detergents (0.1% to 1% w/v CHAPS; 0.5% to 1% w/v Triton X-100; 1% w/v MEGA-10; 1% w/v octyl-β-glucopyranoside; and 0.1% w/v SDS) and reducing agents (5 mM β-mercaptoethanol). Protease inhibitors or chelating agents such as 1 mM EGTA have also been added. An additional possibility is the use of 2-D PAGE buffer (see Basic Protocol 2; also see Reagents and Solutions). Components of buffers must be compatible with subsequent analysis; for example, the use of EDTA and DTT should be avoided with IMAC ProteinChips. Similarly, particular detergents such as SDS, or high salt concentrations, can interfere with mass spectrometry, so thorough washing steps are needed. It is recommended that appropriate initial experiments be carried out using nondissected tissue sections to optimize conditions for any particular application. The binding buffers for each ProteinChip can be modified to alter the stringency of the binding conditions. With the NP2 ProteinChip, the use of increasing salt (50 to 500 mM), detergent (0.01% to 0.1% v/v), or organic solvent (0% to 50% v/v), or decreasing the pH, will increase the selectivity. For the H4 ProteinChip, the inclusion of 50 to 100 mM NaCl will increase the hydrophobic interactions, whereas detergents present in the sample will have the converse effect and may need to be diluted out to maximize binding. The SAX2 and WCX2 ProteinChips bind anions and cations respectively, and selectivity can be altered by varying the pH of the binding buffers used. Proteins with a low isoelectric point (pI) bind to the SAX2 ProteinChip, but as buffer pH is lowered below the pI of the protein, the protein will bear a net positive charge and will bind
more weakly. Conversely, the WCX2 ProteinChip binds proteins with a high pI, and as buffer pH is increased above the pI of the protein, the protein will bear a net negative charge and bind more weakly. For the IMAC ProteinChip, proteins bind to the chelated metal ion through histidine, tryptophan, cysteine, and phosphorylated amino acids. Alternative metal ions that can be used to prime the ProteinChip include copper and gallium, which may alter the profile of bound proteins. Hydrophobic and ionic interactions can be minimized by increasing the NaCl concentration in the binding buffer to up to 1 M and including 0.1% Triton X-100. It is also possible to carry out “dry-down” sample application, whereby the surface chemistries of the particular ProteinChip surfaces are not exploited to yield specific profiles, but, rather, the sample is applied as described for the H4 ProteinChip and is then allowed to dry completely and is briefly washed, whereupon the matrix is added and the samples are analyzed by SELDI (see Basic Protocol 4, steps 5b to 13b). This essentially profiles all readily ionizable molecules present that remain after fairly nonstringent application. Care should be taken with this approach if using extraction buffers with high concentrations of salts or detergents, or reagents such as urea that may interfere with subsequent analysis. The energy-absorbing molecule (EAM) described here for the sample matrix is sinapinic acid, which is most suited for a wider range of molecular masses, particularly those >5 kDa. If the region <5 kDa is of particular interest, an alternative EAM is CHCA (α-cyano-4-hydroxycinnamic acid).
Anticipated Results The specific results will be very dependent on the particular tissue/cells being dissected, but, in general, using the tissue processing and staining protocols described here, protein integrity should be preserved and 2-D gels produced with well resolved and focused spots, provided that enough sample material is dissected. For a large-format 18-cm 2-D gel (containing ampholytes of pH 3 to 10), this would ideally be in the region of 30 µg total protein, generating 1200 to 2000 spots, although in practice this is rarely obtained using LCM due to the dissection time involved. The protein loads actually achieved tend to be considerably less, and result in <1000 protein spots. As a guide, an 8-µm thick frozen section of kidney approximately 2 to 3 mm square would yield 6 to 8 µg total protein. Approximate estimates of
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likely protein yields can be determined by processing and extracting whole tissue sections and estimating the percentage occupied by the cells of interest. The sensitivity of immunoblotting for detecting specific proteins in laser-captured material is dependent on the abundance of the specific proteins being detected and the quality of the antibody. As examples, the authors have been able to detect actin and major vault protein in <250 7.5-µm shots (very approximately equivalent to cell number) with no evidence of protein degradation. The complexity of the results obtained with material analyzed by SELDI is generally dependent on the material, solubilization process, and ProteinChip type selected. Differences between epithelial compartments and stromal elements of tissues can be seen very clearly at submicrogram levels of material per sample spot—indeed few molecules can be seen to ionize in stromal samples, presumably due to the preponderance of larger-molecular-weight proteins. Far more complex profiles are obtained with epithelial cells, and profiles can be generated using <100 cells, although more information is likely to result from additional material as described previously.
Time Considerations The processing of sections for analysis takes <20 min. Dissection can take anything from 10 min to several days, with the products of several dissections being pooled to ensure sufficient material for the downstream application. Enough material for immunoblotting or SELDI may be obtained in a few minutes for each sample, whereas tens of thousands of shots may be necessary, over several hours or days, to obtain sufficient material for 2-D PAGE. This is partly determined by the nature of the tissue in terms of ease of dissection and abundance of cell type of interest. A consideration of the time taken for 2-D PAGE has already been provided in UNIT 10.4, where large-format gels may take 3 days to complete. 1-D immunoblots, however, can be accomplished in a single day. Preparation of SELDI ProteinChips using dissected material generally takes between 15 min and 2 hr, depending on the ProteinChip type.
Literature Cited Laser Capture Microdissection for Proteome Analysis
Preliminary findings. Electrophoresis 20:689700. Cornea, A. and Mungensat, A. 2002. Comparison of current equipment. Methods Enzymol. 356:3-12. Craven, R.A. and Banks, R.E. 2002. Use of laser capture microdissection to selectively obtain distinct populations of cells for proteomic analysis. Methods Enzymol. 356:33-49. Craven, R.A., Totty, N., Harnden, P., Selby, P.J., and Banks, R.E. 2002. Laser capture microdissection and two-dimensional polyacrylamide gel electrophoresis: Evaluation of tissue preparation and sample limitations. Am. J. Pathol. 160:815-822. Emmert-Buck, M.R., Bonner, R.F., Smith, P.D., Chuaqui, R.F., Zhuang, Z., Goldstein, S.R., Weiss, R.A., and Liotta, L.A. 1996. Laser capture microdissection. Science 274:998-1001. Emmert-Buck, M.R., Gillespie, J.W., Paweletz, C.P., Ornstein, D.K., Basrur, V., Appella, E., Wang, Q.H., Huang, J., Hu, N., Taylor, P., and Petricoin, E.F. 2000. An approach to proteomic analysis of human tumors. Mol. Carcino. 27:158-165. Hofman, F. 2002. Immunohistochemistry. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober), 21.4.1-21.4.23. John Wiley & Sons, New York. Knezevic, V., Leethanakul, C., Bichsel, V.E., Worth, J.M., Prabhu, V.V., Gutkind, J.S., Liotta, L.A., Munson, P.J., Petricoin, E.F., and Krizman, D.B. 2001. Proteomic profiling of the cancer microenvironment by antibody arrays. Proteomics 1:1271-1278. Kolble, K. 2000. The LEICA microdissection system: Design and applications. J. Mol. Med. 78:B24-B25. Palmer-Toy, D.E., Sarracino, D.A., Sgroi, D., LeVangie, R., and Leopold, P.E. 2000. Direct acquisition of matrix-assisted laser desorption/ionization time-of-flight mass spectra from laser capture microdissected tissues. Clin. Chem. 46:1513-1516. Paweletz, C.P., Ornstein, D.K., Roth, M.J., Bichsel, V.E., Gillespie, J.W., Calvert, V.S., Vocke, C.D., Hewitt, S.M., Duray, P.H., Herring, J., Wang, Q.H., Hu, N., Linehan, W.M., Taylor, P.R., Liotta, L.A., Emmert-Buck, M.R., and Petricoin, E.F., III. 2000a. Loss of annexin 1 correlates with early onset of tumorigenesis in esophageal and prostate carcinoma. Cancer Res. 60:6293-6297. Paweletz, C.P., Gillespie, J.W., Ornstein, D.K., Simone, N.L., Brown, M.R., Cole, K.A., Wang, Q.-H., Huang, J., Hu, N., Yip, T.-T., Rich, W.E., Kohn, E.C., Linehan, W.M., Weber, T., Taylor, P., Emmert-Buck, M.R., Liotta, L.A., and Petricoin, E.F. 2000b. Rapid protein display profiling of cancer progression directly from human tissue using a protein biochip. Drug Dev. R. 49:34-42.
Banks, R.E., Dunn, M.J., Forbes, M.A., Stanley, A., Pappin, D., Naven, T., Gough, M., Harnden, P., and Selby, P.J. 1999. The potential use of laser capture microdissection to selectively obtain distinct populations of cells for proteomic analysis:
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Paweletz, C.P., Charboneau, L., Bichsel, V.E., Simone, N.L., Chen, T., Gillespie, J.W., EmmertBuck, M.R., Roth, M.J., Petricoin, III E.F., and Liotta, L.A. 2001. Reverse phase protein microarrays which capture disease progression show activation of pro-survival pathways at the cancer invasion front. Oncogene 20:1981-1989. Schutze, K. and Lahr, G. 1998. Identification of expressed genes by laser-mediated manipulation of single cells. Nature Biotechnol. 16:737-742. Simone N.L., Remaley, A.T., Charboneau, L., Petricoin, E.F., Glickman, J.W., Emmert-Buck, M.R., Fleisher, T.A., and Liotta, L.A. 2000. Sensitive immunoassay of tissue cell proteins procured by laser capture microdissection. Am. J. Pathol. 156:445-452. von Eggeling, F., Davies, H., Lomas, L., Fiedler, W., Junker, K., Claussen, U., and Ernst, G. 2000. Tissue-specific microdissection coupled with ProteinChip array technologies: Applications in cancer research. Biotechniques 29:1066-1070. Westphal, G., Burgemeister, R., Friedemann, G., Wellmann, A., Wernert, N., Wollscheid, V., Becker, B., Vogt, T., Knuchel, R., Stolz, W., and Schutze, K. 2002. Noncontact laser catapulting: A basic procedure for functional genomics and proteomics. Methods Enzymol. 356:80-97. Wright, Jr. G.L., Cazares, L.H., Leung, S.-M., Nasim, S., Adam, B.L., Yip, T.-T., Schellhammer, P.F., Gong, L., and Vlahou, A. 1999. ProteinChip surface enhanced laser desorption/ionization (SELDI) mass spectrometry: A novel protein biochip technology for detection of prostate cancer biomarkers in complex protein mixtures. Prostate Cancer Prostatic Dis. 2:264-276.
Key References Conn, P.M. (ed.). 2002. Laser Capture Microscopy and Microdissection. In Methods in Enzymology, Vol. 356. Academic Press, San Diego, Ca. A whole volume devoted to all aspects of this technology, from a description of the different systems through to protocols and applications for laserbased cell selection for subsequent analysis of either proteins or nucleic acids.
Internet Resources http://www.arctur.com/ The homepage for Arcturus. This Web site and the sites below are the homepages of the companies supplying the different laser-based microdissection systems and the SELDI-MS. Their listing does not constitute an endorsement of their products by the authors but indicates where further technical information can be obtained. Some of the sites provide limited protocols, links to other relevant sites, and also listings of relevant publications. http://www.palm.spacenet.de/ The homepage for PALM. http://www.leica-microsystems.com/ The homepage for Leica. http://www.ciphergen.com/ The homepage for Ciphergen.
Contributed by Rachel A. Craven and Rosamonde E. Banks St. James’s University Hospital Leeds, United Kingdom
Gel-Based Proteome Analysis
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Preparing Protein Extracts for Quantitative Two-Dimensional Gel Comparison
UNIT 22.4
This unit provides methods for protein extraction and removal of interfering compounds for subsequent analysis of the proteins by two-dimensional (2-D) gel electrophoresis (UNIT 10.4). An important caveat is that these methods are intended only for two-dimensional separations in which the first dimension is isoelectric focusing (IEF). Thus, it is essential that the sample preparation process address the chemical constraints inherent in the IEF process and be reproducible between extractions to allow quantitative analysis of the resulting 2-D gels, the latter being crucial for quantitative proteomics analysis using 2-D electrophoresis (e.g., UNITS 10.4 & 22.2). The procedures described in this unit address the first consideration by solubilizing the proteins in a solution similar to that used during IEF (i.e., a strongly denaturing solution of low ionic strength). The extraction solution usually contains high concentrations of urea and sometimes ancillary chaotropes such as thiourea, as well as nonionic and zwitterionic detergents, a disulfide bond reducer, and other reagents such as carrier ampholytes (used here as a buffer). While simple dilution of pure protein samples in such solution is convenient as a sample preparation process, the situation is much more complicated with complex samples where the concentration of many interfering compounds (namely anything other than proteins) must not exceed a critical threshold. This threshold will depend upon many parameters (e.g., amount of sample loaded, geometry of the 2-D electrophoresis setup) and is therefore very difficult to predict. Some cellular components (e.g., DNA) strongly interfere with isoelectric focusing and must always be removed unless present in very low amounts (e.g., in mitochondrial preparations). Other components (e.g., salts, lipids, neutral polysaccharides) interfere much less severely and need to be removed only when highly abundant in the biological material of interest. Consequently, the optimized extraction process is strongly sample-dependent and cannot be described for every type. Nevertheless, the protocols in this unit serve as general guidelines and provide good starting points for sample preparation optimization. Methods are listed according to type of biological material, starting with animal cells, nuclei, and bacteria (see Basic Protocol 1), animal tissues (see Alternate Protocol 1), plasma or serum (see Alternate Protocol 2), and other biological fluids (see Alternate Protocol 3). These are followed by extraction techniques for plant tissues and cells (see Basic Protocol 2), and finally subcellular organelles other than nuclei (see Basic Protocol 3). NOTE: Refer to protocols concerning electrophoresis on immobilized pH gradient (IPG) gels and second dimension electrophoresis of IPG gels for more information on 2-D gel electrophoresis (UNIT 10.4). PROTEIN EXTRACTION FROM CULTURED ANIMAL CELLS, CELL NUCLEI, AND BACTERIA
BASIC PROTOCOL 1
This protocol is broadly applicable for extracting protein from a wide variety of biological materials. It is designed to remove nucleic acids very efficiently and is therefore suitable for preparation of nuclear proteins. It is also fairly simple and does not involve a protein precipitation step. It is not suited for the extraction of very basic proteins, however, and thus cannot accommodate materials that are very rich in hydrophobic compounds. For these reasons, this protocol is not suitable for plant cells or tissues, which are instead addressed in a separate method (see Basic Protocol 2). Gel-based Proteome Analysis Contributed by Mireille Chevallet, Christophe Tastet, Sylvie Luche, and Thierry Rabilloud Current Protocols in Protein Science (2003) 22.4.1-22.4.9
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Materials Animal or bacterial cells, or cell nuclei (UNIT 4.2) Phosphate-buffered saline (PBS; APPENDIX 2E) Isotonic buffer (see recipe) 1.25× lysis buffer (see recipe) 40% carrier ampholytes (Amersham Pharmacia Biotech), pI range 3 to 10 Thick-walled polyallomer ultracentrifuge tube Thick laboratory plastic film Ultracentrifuge with fixed-angle rotor Additional reagents and equipment for fractionation of cell nuclei (UNIT 4.2) and measuring protein concentration (UNIT 3.4) 1a. For animal or bacterial cell suspensions: Centrifuge cells for 5 min at 1000 × g, 4 °C. 1b. For adherent animal cells: Scrape into an appropriate tube and centrifuge 5 min at 1000 × g, 4°C. 1c. For cell nuclei: Prepare a nuclei pellet (UNIT 4.2) and use directly in the subsequent steps. 2. Wash the resulting pellet twice in 10 vol phosphate buffered saline (PBS), then once in 10 vol isotonic buffer. Estimate pellet volume and resuspend in the same volume of isotonic buffer. Transfer to a thick-walled polyallomer ultracentrifuge tube and measure the volume. The polyallomer tube is needed to tolerate the high pH of lysis buffer (close to 10). Cellulose ester and polycarbonate tubes are not suitable for ultracentrifugation at this high pH.
3. Add 4 vol 1.25× lysis buffer. Put a small piece of thick laboratory plastic film (e.g., Parafilm) on top of the tube and mix by inversion. The solution should become very viscous at this stage and almost impossible to pipet. It is for this reason that lysis is performed directly in the ultracentrifuge tube.
4. Incubate 30 min to 1 hr at room temperature. 5. Ultracentrifuge in a fixed-angle rotor 1 hr at 200,000 × g, room temperature. A tight, colorless pellet of nucleic acids should be present at the bottom of the tubes. Take care not to disturb it with the pipet. The solution should be less viscous at this point.
6. Collect the supernatant and estimate the protein concentration by Bradford assay (UNIT 3.4). 7. Add 1/100 vol of 40% carrier ampholytes, pI range 3 to 10, and mix. Store frozen up to 1 year at –80°C. ALTERNATE PROTOCOL 1
Preparing Protein Extracts for Quantitative Two-Dimensional Gel Comparison
PROTEIN EXTRACTION FROM ANIMAL TISSUES This protocol is similar to the one presented for animal cells (see Basic Protocol 1), but homogenization of the tissue is required to accomplish total extraction of the proteins. Additional Materials (also see Basic Protocol 1) Animal tissue, frozen Tissue homogenizer adapted to the hardness of the tissue 1. Mince fresh animal tissue into small pieces (~20 cubic millimeters) with scissors and estimate the tissue volume.
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2. Add four volumes of 1.25× lysis buffer and homogenize at room temperature. A Potter Elvejhem homogenizer is sufficient for soft tissues (e.g., liver), but a helix homogenizer may be required for more fibrous tissue (e.g., pituitary gland).
3. Clarify the lysate by centrifuging 30 min at 15,000 × g, room temperature. 4. Decant supernatant to a polyallomer tube and ultracentrifuge in a fixed-angle rotor 1 hr at 200,000 × g, room temperature. Store up to 1 year at −80°C. PROTEIN EXTRACTION FROM PLASMA OR SERUM Plasma and serum are liquids very rich in proteins and contain a relatively low amount of salts, nucleic acids, and lipids. A beneficial effect of an initial extraction by hot SDS has been reported, however (Hochstrasser et al., 1988).
ALTERNATE PROTOCOL 2
Additional Materials (also see Basic Protocol 1) Serum or plasma SDS buffer (see recipe) 1.25× denaturing buffer (see recipe) 20° and 100°C (i.e., boiling) water bath 1. Mix one volume serum or plasma with 1.5 vol SDS buffer. Heat 3 min at 100°C in a boiling water bath. 2. Cool to 20°C in a water bath and add 4 vol of 1.25× denaturing buffer. Use immediately or prepare 0.1- to 0.2-ml aliquots and store up to 1 year at −80°C. When calculating the volume of concentrated denaturing buffer, compare with the total plasma volume plus the added SDS buffer volume.
PROTEIN EXTRACTION FROM OTHER BIOLOGICAL FLUIDS BY ABSORPTION-DESORPTION
ALTERNATE PROTOCOL 3
Unlike plasma and serum, most other biological fluids (e.g., urine, sweat, cerebrospinal fluid, conditioned culture solution) have a relatively low protein content and a rather high salt concentration. They should therefore be desalted and the protein concentrated. This can be achieved by concentration-dialysis methods, but these methods usually induce significant protein losses. The adsorption-desorption method described below is more reliable. Additional Materials (also see Basic Protocol 1) Biological fluid (other than plasma or serum) 10% diatomaceous earth suspension (see recipe) 1.25× denaturing buffer (see recipe) 1. Estimate the protein concentration of the biological fluid sample using a suitable protein assay (UNIT 3.4). 2. Mix the sample with 1/4 vol of 10% diatomaceous earth suspension. Agitate 1 hr at room temperature using a rotary shaker. 3. Centrifuge 5 min at 15,000 × g, 4°C, and remove supernatant. 4. Add 4 pellet volumes 1.25× denaturing buffer. Agitate 30 min at room temperature. 5. Pellet the diatomaceous earth by centrifuging 10 min at 15,000 × g, room temperature. Recover the supernatant and store up to 1 year at −80°C.
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BASIC PROTOCOL 2
PROTEIN EXTRACTION FROM PLANT TISSUES AND CELLS In contrast to animal samples (see Basic Protocol 1 and Alternate Protocols 1 to 3), plant samples are generally very rich in protease activities and in interfering materials, such as cell wall polysaccharides, polyphenolic compounds, pigments, and other solvent-soluble materials. Materials Plant tissue or cell pellet Acid denaturing solution (see recipe) 0.05% (w/v) DTT in acetone 1× denaturing buffer (see recipe) Mortar and pestle Ultracentrifuge with fixed-angle rotor Additional reagents and equipment for measuring protein concentration (UNIT 3.4) 1. Grind the plant tissue or cell pellet under liquid nitrogen using a motor and pestle. Let the nitrogen evaporate and mix the powder with ∼10 vol acid denaturing solution. 2. Precipitate the proteins by incubating 45 min at −20°C and then centrifuging the resulting suspension in a fixed-angle rotor 15 min at 35,000 × g, 4°C. 3. Decant the supernatant, resuspend the pellet in 0.05% (w/v) DTT in acetone, and extract by incubating 1 hr at −20°C. 4. Centrifuge the suspension 15 min at 35,000 × g, 4°C. 5. Discard the supernatant and dry the pellet under vacuum. Resolubilize the pellet in 1× denaturing buffer. Incubate 1 hr at room temperature. This step is critical for correct protein recovery. Incubating 30 min in a water sonicator can also be helpful.
6. Ultracentrifuge the suspension 30 min at 200,000 × g, 20°C. Recover the supernatant and measure the protein concentration (UNIT 3.4). Store up to 1 year at −80°C. Carrier ampholytes interfere slightly with the protein assay. The blank and standard samples must contain the same volume of denaturing buffer as the sample to be tested. If the protein concentration of the sample is out of the linear range of the assay, dilutions must be made in denaturing buffer and the same volume of sample used throughout. BASIC PROTOCOL 3
PROTEIN EXTRACTION FROM SUBCELLULAR ORGANELLES OTHER THAN NUCLEI Subcellular organelles usually contain very low amounts of salts and nucleic acids, but can be very rich in membranes and thus in lipids and membrane proteins. In this case, incorporation of specific detergents in the protein extraction process can be very helpful to maximize the yield of the protein extraction. This combination of reducing agents and detergents prevents protein determination by most protein assays, however. Thus, the protein determination must be carried out on the organelle preparation prior to extraction.
Preparing Protein Extracts for Quantitative Two-Dimensional Gel Comparison
Materials Isolated organelles (e.g., UNIT 4.2) Isotonic buffer (see recipe) 20% (w/v) detergent solution in 50% (v/v) ethanol, 30°C 1.25× chaotrope solution, 30°C (see recipe) Ultracentrifuge with fixed-angle rotor Additional reagents and equipment for determining protein concentration (UNIT 3.4)
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1. At the end of their preparation, wash isolated organelles with isotonic buffer and resuspend in a small volume (e.g., 1 pellet vol) of isotonic buffer. 2. Determine the protein concentration of the solution using a suitable protein assay (UNIT 3.4). Aliquot the organelle suspension and store frozen at −80°C. The protein concentration should be ≥20 mg/ml.
3. Determine the suspension volume needed for the 2-D gel electrophoresis experiment. Prepare the final denaturing solution by adding 1 vol (i.e., equivalent to the volume of organelle suspension) 20% (w/v) detergent solution in 50% ethanol to 4 vol 1.25× chaotrope solution, both warmed to 30°C. Immediately add the organelle suspension to this denaturing buffer. This preparation may seem cumbersome, but it is the only way to make sure that the sample is always in contact with chaotropes and detergent while allowing relatively unstable but powerful chaotrope-detergent pairs to be used. Warming the detergent and chaotrope solutions usually prevents detergent precipitation, which is generally very temperature-sensitive. The amount and identity of detergent to use must be empirically determined; however, the usual concentration is 2% (final). A silver-stained analytical gel requires 100 pg of protein; however, up to 1 mg of protein can be loaded if required.
4. Incubate (i.e., extract) 1 hr at room temperature. Ultracentrifuge in a fixed-angle rotor 30 min at 200,000 × g, 20°C. Use the supernatant immediately. Some detergents are poorly compatible with urea and thiourea. The resulting mix is therefore very difficult to resolubilize if frozen. This is why storage of these solutions is not recommended. It is much more convenient to aliquot and freeze the initial organelle suspension.
REAGENTS AND SOLUTIONS Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Acid denaturing solution Prepare a 10% (w/v) solution of trichloracetic acid in acetone. Store up to 3 months at 4°C. On the day of use, add 0.05% (w/v) DTT. Trichloroacetic acid is very hygroscopic. Ideally, it should be dried by rotary evaporation under vacuum prior to use. Otherwise, a new bottle should be opened and the acetone added directly to the bottle without prior weighing.
Chaotrope solution, 1.25× In the bottom of an appropriate tube, place sufficient solids to prepare a solution of 8.75 M urea and 2.5 M thiourea. Add 40% (w/v) carrier ampholytes (Amersham Pharmacia Biotech), pI range 3 to 10, and 1 M DTT to achieve final concentrations of 0.5% (w/v) and 50 mM, respectively. Add water to volume, bearing in mind that 1 g urea occupies a volume of 0.75 and 1 g thiourea occupies 1 ml. (As an example, for a 10 ml solution, add 5.26 g urea, 1.9 g thiourea, 0.125 ml of 40% (w/v) carrier ampholytes, 0.5 ml of 1 M DTT, and 3.53 ml water.) Completely dissolve the solids, but do not warm above 37°C to avoid urea decomposition. Prepare 1-ml aliquots and store frozen up to 1 year at –20°C. Two freeze-thaw cycles are permissible per aliquot. The solution can be prepared very conveniently in a graduated, disposable, polystyrene centrifuge tube. The use of a water sonicating bath is very helpful to achieve complete dissolution of the solids. Thawing can be aided by bath sonication or by incubation in a warm (30°C) water bath. Gel-based Proteome Analysis
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Denaturing buffer, 1×, 1.25× For the 1.25× solution, prepare 1.25× chaotrope solution (see recipe), except after the addition of thiourea, add 3-[(3-cholamidopropyl)dimethylamino]-1-propanesulfonate (CHAPS) to 5% (w/v). Prepare and store as described for the chaotrope solution. When adding water, bear in mind that 1 g CHAPS occupies 1 ml. If necessary, dilute to 1× with water. As an alternative, the 1× solution can be prepared directly from solids and stock solutions as described for chaotrope solution using the following final concentrations: 7 M urea, 2 M thiourea, 4% (w/v) CHAPS, 40 mM DTT, and 0.4% (w/v) carrier ampholytes, pI range 3 to 10.
Diatomaceous earth suspension Suspend 1 g sulfuric acid–washed diatomaceous earth (Sigma) in 10 ml water, shaking thoroughly. Centrifuge 10 min at 2000 × g, 4°C. Remove the water by pipetting. Repeat this washing procedure twice. Resuspend in 10 ml water and store up to 3 months at 4°C. Isotonic buffer 0.25 M sucrose 1 mM EDTA 10 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) Store up to 1 month at 4°C Lysis buffer, 1.25× 8.75 M urea 2.5 M thiourea 5% (w/v) CHAPS 25 mM spermine base, fresh 50 mM dithiothreitol Prepare and store as described for 1.25× chaotrope solution (see recipe) Generally, 1 M stock solutions are prepared for spermine base (do not use spermine tetrahydrochloride) and dithiothreitol. Spermine base dissolves easily in water, but both the dry and dissolved forms readily absorb carbon dioxide from the air. Therefore, the stock solution should be prepared directly in a newly opened bottle without weighing. The weight provided by the supplier is considered as correct. Aliquot the stock solution (1 ml) and store frozen up to 1 year at –20°C.
SDS buffer 10% (w/v) SDS 2% (w/v) DTT Store in 0.1-ml aliquots up to 1 year at –20°C COMMENTARY Background Information
Preparing Protein Extracts for Quantitative Two-Dimensional Gel Comparison
Methods of protein preparation for two-dimensional electrophoresis must take into account the constraints required for isoelectric focusing, the first dimension of typical two-dimensional electrophoresis. These constraints include the requirement for low-ionic-strength conditions and the exclusion of solubilizers that may bind proteins and subsequently modify their pI. Hence, salts and ionic detergents cannot be used freely in such approaches. In addition, all molecular species in the sample that may interfere with the isoelectric focusing
process must be removed during this extraction step, while preserving the solubility of protein. Such interfering compounds include salts, nucleic acids, and lipids. Finally, proteins should not be exposed to conditions that may induce chemical modification or degradation, either from the enzymes present in the sample or from the chemical conditions used for sample preparation. Given so many constraints, a complete and perfect protein solubilization method for twodimensional electrophoresis is an almost unreachable Holy Grail. For example, separating
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proteins from nucleic acids and removing the latter without carrying over proteins usually requires high salt concentrations and/or ionic detergents or chaotropes; conditions incompatible with two-dimensional electrophoresis. As another example, removal of lipids is usually achieved by extraction into organic solvents, but this precipitates proteins. In addition, it appears that some proteins are soluble in these organic solvents (e.g., see Molloy et al., 1999) and that others, once precipitated, will never redissolve, especially when low-ionic-strength conditions must be used (e.g., Luche et al., 2003). Since proteomics experiments generally imply comparative analysis of a sample series, important emphasis must be placed on the reproducibility of sample preparation protocols. Reproducibility can be compromised by poor absolute protein extraction in addition to the fact that significant sample variation is often encountered in proteomics studies. Thus, protocols used for sample preparation for two-dimensional electrophoresis are always imperfect compromises. Due to the constraints on ionic chemicals, the final protein extraction solution, which must be closely related to the isoelectric focusing gel solution, usually contains high concentrations of nonionic chaotropes (e.g., urea and its derivatives, but not guanidinium salts) and nonionic or zwitterionic detergents. In most cases, however, the sample preparation protocol is not simple dilution of the biological material of interest into the chaotrope-detergent solution. Owing to the severity of the artifacts that can be induced by nonproteinaceous compounds or by the lytic activities present in the sample, the protocol must be designed to minimize these artifacts. In addition, because interfering chemicals tend to vary much more between samples than the proteins of interest themselves, the specifics of the extraction protocol are heavily sample dependent. Consequently, most two-dimensional gel electrophoresis sample preparation protocols include steps that are specifically designed to remove or inactivate some interfering chemicals. For example, salt removal by protein adsorption (see Alternate Protocol 3; Maresh and Dunbar, 1988) or removal of cell wall polysaccharides and hydrophobic compounds by the TCA/acetone procedure (see Basic Protocol 2; Damerval et al., 1986). Additional benefits of TCA are its ability to degrade RNA, liberate even very basic proteins (e.g., ribosomal pro-
teins; Görg et al., 1998), and inhibit proteases almost instantly by denaturation. When the above constraints are met, then some variegation can be applied to maximize the protein yield. This variegation usually deals with the nature and amounts of chaotropes (Rabilloud et al., 1997) and detergents used (Chevallet et al., 1998). In some cases, low amounts of SDS have been shown to increase protein extraction, as described for serum (Hochstrasser et al., 1988). It must be kept in mind, however, that SDS is useful only at the initial solubilization stage and must be displaced from proteins at the IEF stage by the molar excess of the chaotrope-nonionic detergent mixture. Thus, the presence of SDS in the sample limits the amount that can be loaded on a 2-D gel.
Critical Parameters and Troubleshooting A guide for efficient sample preparation is difficult to put forward because of the extreme dependence of the process on the sample itself. Most often, the quality of the sample can be judged only at the very end of the process (i.e., when the spots are analyzed for their protein content by mass spectrometry). At this stage, if the spots do not match with the identified proteins in terms of pI and especially molecular weight, the first suspicion should be alteration by contaminating proteases (see below and Harder et al., 1999). The only anomalies which can be easily detected at the two-dimensionalpattern stage are those induced by excess salt or by nucleic acids. Excess salt can be detected as soon as the end of the first dimension in immobilized pH gradients by the presence of physical bulges at the end(s) of the IPG strip. An empty zone will be present at the corresponding location on the 2-D gel. Lysis speed Rapid cell or organelle lysis is extremely important for successful sample preparation. Slow lysis usually allows significant lytic activities by proteases and increases the time for precipitation phenomena. The conditions for lysis and precipitation have important consequences in protocol design. It is always better to lyse a well-dispersed sample (i.e., a very fine powder or, even better, a concentrated and homogeneous suspension or solution) by dilution into a concentrated lysis solution than to apply a lysis solution to a pellet. In the latter case, the gradient of penetration of the lysis solution into the pellet usually results in the formation of a
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the pellet, further slowing penetration of the lysis solution. The result is almost always an irreproducible and poor extraction, with extensive proteolytic degradation. If cells must be frozen prior to use, they must be frozen as a live suspension in solution containing cryoprotectants—e.g., 10% (v/v) DMSO. They are then thawed and treated as live cells. Even so, some cell lysis always occurs at this stage, and the use of fresh cell suspension is always recommended.
transient zone in which intermediate denaturing conditions prevail. This is the worst possible situation. For example, with regard to protease activity, intermediate denaturing conditions will cause many proteins to be at least partially denatured and thus expose proteasesensitive sites, but the proteases themselves will generally not be denatured since they are generally fairly denaturation-resistant (Colas des Francs et al., 1985). In addition, intermediate denaturing conditions usually disrupt subunit interactions, but do not denature the actual subunits (Elbaum et al., 1974), thereby providing the opportunity for these proteins to engage in artifactual interactions with other cellular components. If, for example, some nonnuclear proteins interact with DNA, they will be removed along with the DNA and thus will be lost in the final extract. The above-mentioned phenomena are the rationale for which the protocols described in this unit so often use sample dilution as the lysis step. Proceeding with this trend, frozen cell pellets probably represent the worst samples. During the thawing process, cell lysis begins in the pellet itself and DNA unwinding occurs. This in turn induces significant viscosity within
A
Preparing Protein Extracts for Quantitative Two-Dimensional Gel Comparison
Proteases Another important and related problem is linked to the activity of proteases in the sample. Because these proteins are very efficient and very difficult to inhibit, especially when starting with a complex sample containing a wide variety of proteases, it is crucial to minimize the amount of time the sample spends in conditions in which proteases are active. This in turn implies that treatment of the sample with other enzymes (e.g., nucleases to remove DNA and RNA or glycosidases to simplify the protein pattern) must be used with great caution. Nucleic acid removal is best achieved under strongly denaturing conditions.
B
Figure 22.4.1 Two-dimensional gels of HeLa (A) and E. coli (B) total cell extracts prepared according to the procedure for cultured animal and bacterial cells (see Basic Protocol 1). In the first dimension, isoelectric focusing was performed using 0.1 mg protein on a pH 4 to 8 gradient and focused for 60,000 V hr. In the second dimension, 10% polyacrylamide gel electrophoresis was performed for 5 hr hr at 200 V, followed by detection with silver staining. Note the very low number of streaks, the round shape of most spots, and the high spot density in the two gels, indicative of a correct sample preparation.
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Macromolecular interference Interference by nucleic acids can sometimes be detected directly at the lysis stage by poor homogeneity of the sample and high viscosity not eliminated by ultracentrifugation. In cases with lower amounts of nucleic acids, the interference is only detected in the 2-D gel pattern by intense vertical streaks (DNA) or a grid-like pattern (RNA) as described in Rabilloud et al. (1986). Interference by other molecules (e.g., polysaccharides, lipids) is generally detected by a much lower than expected number of spots compared to sample load. Generally speaking, a 0.1-mg load of a complex extract on a wide pH range (e.g., 4 to 7 or wider) coupled with silver staining should produce more than 1000 protein spots. Much lower numbers are indicative of interference by other cellular compounds. This phenomenon can be investigated by mixing the suspect sample with a well-behaved sample—e.g., an E. coli or a HeLa total cell extract prepared as described (see Basic Protocol 1)—and comparing the resulting pattern with the one obtained with this reference sample alone. A significant reduction in spot number is then indicative of interference. Extensive streaking on the gel is difficult to interpret, as it can arise from problems in the isoelectric focusing, SDS electrophoresis, or sample preparation steps. Here again, the use of reference sample can be useful to trace the problems.
Anticipated Results The extreme dependency on the biological source makes specific result prediction nearly impossible. Instead, examples of gels obtained with such reference samples are presented (Fig. 22.4.1). They show minimal streaking, good spot shapes, and a high density of spots.
Time Considerations The extraction typically takes 3 to 4 hr, except for subcellular organelles other than nuclei (see Basic Protocol 3), which takes 30 min.
Literature Cited Chevallet, M., Santoni, V., Poinas, A., Rouquié, D., Fuchs, A., Kieffer, S., Rossignol, M., Lunardi, J., Garin, J., and Rabilloud, T. 1998. New zwitterionic detergents improve the analysis of membrane proteins by two-dimensional electrophoresis. Electrophoresis 19:1901-1909.
Colas des Francs, C., Thiellement, H., and De Vienne, D. 1985. Analysis of leaf proteins by twodimensional electrophoresis. Plant Physiol. 78:178-182. Damerval, C., De Vienne, D., Zivy, M., and Thiellement, H. 1986. Technical improvements in twodimensional electrophoresis increase the level of genetic variation detected in wheat-seedling proteins. Electrophoresis 7:52-54. Elbaum, D., Pandolfelli, E.R., and Herskovits, T.T. 1974. Denaturation of human and Glycera dibranchiata hemoglobins by the urea and amide classes of denaturants. Biochemistry 13:12781284. Görg, A., Boguth, G., Obermaier, C., and Weiss, W. 1998. Two-dimensional electrophoresis of proteins in an immobilized pH 4-12 gradient. Electrophoresis 19:1516-1519. Harder, A, Wildgruber, R., Nawrocki, A., Fey, S.J., Larsen, P.M., and Görg, A.1999. Comparison of yeast cell protein solubilization procedures for two-dimensional electrophoresis. Electrophoresis 20:826-829. Hochstrasser, D.F., Harrington, M.G., Hochstrasser, A.C., Miller, and M.J, Merril, C.R. 1988. Methods for increasing the resolution of two-dimensional protein electrophoresis. Anal. Biochem. 173:424-435. Maresh, G.A. and Dunbar, B.S. 1988. A simple, inexpensive method for the salt-free concentration of proteins for analysis by two-dimensional gel electrophoresis. Electrophoresis 9:54-57. Molloy, M.P., Herbert, B.R., Williams, K.L., and Gooley, A.A. 1999. Extraction of Escherichia coli proteins with organic solvents prior to twodimensional electrophoresis. Electrophoresis 20:701-704. Rabilloud, T., Hubert, M., and Tarroux, P. 1986. Procedures for two-dimensional electrophoretic analysis of nuclear proteins. J. Chromatogr. 351:77-89. Rabilloud, T., Adessi, C., Giraudel, A., and Lunardi, J. 1997. Improvement of the solubilization of proteins in two-dimensional electrophoresis with immobilized pH gradients. Electrophoresis 18:307-316.
Contributed by Mireille Chevallet, Christophe Tastet, Sylvie Luche, and Thierry Rabilloud Départment Résponse et Dynamique Cellulaire/BioEnergetique Cellulaire et Pathologique, Commissariat à l’Energie Atomique Grenoble, France
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Isolation of Organelles and Prefractionation of Protein Extracts Using Free-Flow Electrophoresis
UNIT 22.5
Free-flow electrophoresis (FFE) is a highly versatile technique for the separation of a wide variety of charged or chargeable analytes such as low-molecular weight organic compounds, peptides, proteins, protein complexes, membranes, organelles, and whole cells. This separation is achieved utilizing a variety of different separation principles including isoelectric focusing (IEF), zone electrophoresis (ZE), and isotachophoresis (ITP). FFE distinguishes itself from other fractionation technologies like chromatography or gel-based electrophoresis in several ways. The most significant is the matrix-free fractionation principle. The analyte is injected into a thin, laminar film of aqueous separation buffer and deflected by an electric field perpendicular to the flow direction. The absence of any kind of matrix is directly linked to high sample recoveries of >90%, fast fractionation times of <20 min, and a high sample throughput of up to 50 mg/hr. The rapid-throughput advantage is further strengthened by the continuous application of samples into the continuous laminar separation film. This allows the preparative processing of sample volumes of up to 100 ml during a single 24-hr run. One of the major obstacles in the analysis of proteomes is the extreme complexity of any given cell or body fluid. Reduction of this complexity is a prerequisite for systematic and comprehensive protein analyses and can be achieved at two different stages: on the protein level by fractionation of crude protein mixtures, e.g., whole-cell lysates, or on a subcellular level by isolation and purification of distinct subcellular entities (organelles). This unit focuses on the most important FFE applications in a proteomics context, i.e., the use of native as well as denaturing IEF-FFE for the prepurification of complex protein mixtures for subsequent detailed analysis (see Basic Protocol 1 and Alternate Protocol) and the use of ZE-FFE as a fine purification method for prepurified organelles (see Basic Protocol 2). NATIVE FFE FRACTIONATION OF CRUDE PROTEIN MIXTURES USING THE ISOELECTRIC FOCUSING MODE
BASIC PROTOCOL 1
The preparation and execution of a native isoelectric focusing FFE experiment for the fractionation of crude protein mixtures is presented in this protocol. The protocol uses a ProTeam FFE instrument from Tecan (Fig. 22.5.1)—the only commercially available free-flow electrophoresis device. The separation media are based on Prolyte reagents, also supplied by Tecan. In contrast to classical ampholytes, Prolytes are exactly defined mixtures of low-molecular-weight (<300 Da) organic acids, bases, and zwitterions. This allows for highly reproducible runs as well as easy removal of the Prolytes from the analytes, in combination with a pH gradient that is linear from at least 3.0 to 11.5. Glycerol is added to the separation media to increase the viscosity, whereas hydroxypropylmethylcellulose (HPMC) is added to suppress electroendosmosis. Materials 1:1 (w/w) glycerol/isopropanol Isopropanol (2-propanol) Kerosene, low-odor ProTeam FFE SPADNS (sulfanilic acid azochromotrop; Tecan) Anode circuit solution (100 mM H2SO4; see recipe) Contributed by Peter J. A. Weber, Gerhard Weber, and Christoph Eckerskorn Current Protocols in Protein Science (2003) 22.5.1-22.5.21 Copyright © 2003 by John Wiley & Sons, Inc.
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Figure 22.5.1 ProTeam FFE instrument.
Cathode circuit solution (100 mM NaOH; see recipe) Anode stabilization medium for Basic Protocol 1 (see recipe) Separation medium 1 for Basic Protocol 1 (see recipe) Separation medium 2 for Basic Protocol 1 (see recipe) Separation medium 3 for Basic Protocol 1 (see recipe) Cathode stabilization medium for Basic Protocol 1 (see recipe) Counterflow medium for Basic Protocol 1 (see recipe) ProTeam FFE pI marker (Tecan) Concentrated protein solution for analysis ProTeam FFE system (Tecan) equipped with: Recirculating water cooler and 6-mm i.d. tubing (supplied with instrument, but other water cooler of power ≥350 W may be used) 7 × 0.64–mm i.d. (orange-white) media tubes 1 × 1.42–mm i.d. (yellow-yellow) counterflow tube 1 × 0.51–mm i.d. (orange-yellow) sample tube 0.4-mm spacer Anode and cathode membranes 0.65-mm filter paper strips Lint-free paper towels Balance capable of weighing ∼10 mg to 2 kg 96-well microtiter plates 96-well deep-well plates 96-well absorbance reader (optional; for quality controls) equipped with two filters (λabs= 500 to 530 nm and λabs= 400 to 420 nm) Isolation of Organelles and Proteome Prefractionation
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Disassemble and clean the instrument 1. Switch on the cooler and set the temperature to 10°C. The separation chamber has to be in thermal equilibrium before closing, to avoid stressrelated material damage.
2. Move the separation chamber to the vertical position. 3. Attach the fractionation plate to the front part of the separation chamber via the magnetic holder. 4. Open the clamps from the outside to the inside in a pairwise fashion. 5. Open the separation chamber by carefully pulling the front part. 6. Put electrode membranes into a 1:1 mixture of glycerol and isopropanol. In case of using previously used membranes, be careful not to switch the membrane that covered the anode channel with the membrane that covered the cathode channel.
7. Put the paper strips in high-purity water. In case of using previously used paper strips, be careful not to switch the paper strip that covered the anode channel with the paper strip that covered the cathode channel.
8. Make sure that the sample tube is connected to the sample inlet S13 (see Fig. 22.5.2) and that the two other sample inlets (S11 and S12) are closed.
counterflow inlet
I8 counterflow reservoir
A1 B1 C1
G12H12
fractionation tubes
electrode seals
external separation chamber seal
internal separation chamber seal
anode (red plug)
cathode (black plug)
sample inlets
media inlets
S11 S12 S13
I1 I2 I3 I4 I5 I6 I7
Gel-Based Proteome Analysis Figure 22.5.2 Schematic of the separation section of the ProTeam FFE instrument. Current Protocols in Protein Science
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The inlets suggested here generally give the best results, but in some cases it may be necessary to apply the samples via another sample inlet. For example, basic proteins should be applied at an acidic pH, i.e., via inlet S11.
9. Clean the interior walls of the separation chamber between the electrode ducts with clean, lint-free paper towels and the following solvents, in the order indicated: High-purity water Isopropanol Kerosene Isopropanol (again) High-purity water (again). Use a separate paper towel for each cleaning operation. The cleaning process should be repeated twice in the case of a new installation and after repairs or service. On no account should silicone, oil, grease, or adhesive reach the separation-chamber interior, as this could affect separation performance. Never use acetone under any circumstances. Prolonged treatment with kerosene and isopropanol might damage seals or other parts of the separation chamber. Use only powder-free PVC or nitrile gloves. Do not use powdered or silicone-treated gloves under any circumstances.
Assemble the instrument 10. Wet the 0.4-mm spacer with high-purity water and place it on the front plate of the separation chamber. Ensure even distance to the electrode seals and make sure that the separation media inlets do not get covered. 11. Place the anode membrane on the anode and the cathode membrane on cathode, starting from the top to the bottom, with the smooth side of the membrane facing the electrode seal. The membrane must not protrude over the electrode seal.
12. Congruently place the anode paper strip on the anode membrane and the cathode paper strip on the cathode membrane, proceeding from the top to the bottom. 13. Reduce the force of all separation-chamber clamps by turning counter-clockwise. 14. Briskly close the separation chamber front part with both hands on the second connecting bar from the top, ensuring that the separation chamber front plate is parallel with the back plate. 15. Close the middle pair of clamps simultaneously, then close adjacent clamps in a pairwise fashion. 16. Increase the clamping force in a pairwise fashion, starting from the center, by opening the pair of clamps, turning the clamps clockwise, and closing the clamps. The water drops underneath the spacer should get displaced—i.e., the spacer should appear clear afterwards.
17. Visually inspect to see whether the membranes and filter papers have moved. Reopen the chamber and repeat the positioning process if required. 18. Place the fractionation plate on the fraction collector housing. Fill the instrument 19. Tilt the separation chamber 45°. 20. Turn the bubble trap in the filling position. Isolation of Organelles and Proteome Prefractionation
21. Place all media and counterflow tubes in a bottle containing high-purity water. Never use separation media directly, because in that case it will be impossible to remove all the bubbles from the separation chamber.
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22. Place the sample tube in an empty microcentrifuge tube without tightening the screw. 23. Open the three-way stopcock on the counterflow tube in all directions and open the luer-lock closure on the cock. 24. Introduce the upper counterflow tube outlet into the fractionation tray and introduce the three-way stopcock into the bottle grid. 25. Start media pump (direction “IN”), then close the tube cassettes and continue pumping with a delivery rate of 50 rpm until the separation chamber is halfway filled. 26. Reverse the pumping direction and empty the separation chamber until the inlet areas of the spacer are reached and all air bubbles are gone. To accomplish this, a reduction of the pump speed to 20 rpm is advisable.
27. Reverse the pumping direction of the media pump once more and fill the chamber, without any air bubbles, with a delivery rate of 50 rpm to a level just underneath the fractionation tubes. 28. Proceed at maximum speed until the counterflow reservoir is nearly full. 29. To fill the last part of the counterflow reservoir without bubbles, reduce the speed to 20 rpm. 30. After the counterflow reservoir has been filled completely, fill the counterflow tube, including the bubble trap, again at maximum speed. The bubble trap is located close to the counterflow inlet and serves to prevent the transport of bubbles into the separation chamber.
31. Reduce the flow rate to 20 rpm and connect the luer-lock closure of the three-way stopcock by holding both openings up. Light tapping prevents air bubbles from remaining in the stopcock or luer adapter.
32. Close the remaining opening of the three-way stopcock. The fractionation tubes will start dripping.
33. Tilt the chamber to the horizontal position. 34. Tap fractionation plate on the fractionation collector housing and check to make sure that all 96 fractionation tubes deliver evenly. If a fractionation tube fails to deliver, connect a syringe to the corresponding fractionation tube and suck until liquid is coming out of the tube. Whenever restarting the media pump (direction “IN”) use the highest flow rate for some seconds until all fractionation tubes are dropping.
35. Dry the outside of the separation chamber and control for any kind of leakage. If a leak is detected, the chamber must be emptied, opened, and closed again.
36. Start the sample pump with 4 rpm (direction “IN”). Tighten the adjusting screw until the tube starts to deliver, then tighten it an additional quarter turn. 37. Change the pumping direction to “OUT” and run until no air is left in the tube, then stop the sample pump. Quality control I: check the flow profile 38. Dilute 500 µl of the red dye ProTeam FFE SPADNS with 50 ml of high-purity water. 39. Switch off the media pump. Place the media tubes of inlets I2, I4, and I6 (see Fig. 22.5.2) in the dye solution and leave the remaining media tubes of inlet I1, I3, I5, and I7, as well as the counterflow tube of inlet I8, in high-purity water. Never put the dye solution in inlets I1 or I7, because this would contaminate the electrodes.
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40. Run the media pump at 40 rpm (direction “IN”) and collect a 96-well plate after 6 min. Red and colorless stripes that flow in parallel, with identical width and sharp boundaries, should appear in the separation chamber. This can be evaluated and documented by collecting a 96-well plate and measuring the OD at 500 to 530 nm with an appropriate reader. If this is not the case, the following reasons may apply: (1) uneven adjustment of the clamps; (2) surface contamination of the separation chamber; (3) incorrectly installed spacer, seals, paper strips, or membranes; (4) partially clogged or leaking media tubes; (5) opened sample pump adjusting screw; (6) defective tubes or tubes with different diameters. In all cases try to change or correct the corresponding parameter.
41. Switch off media pump. Return the media tubes of inlets I2, I4, and I6 (see Fig. 22.5.2) to high-purity water. 42. Run media pump at 40 rpm (direction “IN”) and rinse for 6 min. Calibrate pumps 43. To calibrate the sample pump, fill a microcentrifuge tube with ∼1.5 ml of high-purity water and weigh accurately to a milligram. The sample pump must be calibrated after each adjustment of the sample-pump screw.
44. Immerse the sample tube in the microcentrifuge tube containing the water. On the touch screen of the ProTeam FFE apparatus, call up the dialog for “Calibrate sample pump” and follow the instructions. Again weigh the calibration vessel, then save the calibration. 45. To calibrate the media pump, fill an appropriate bottle with approximately 200 ml of high-purity water and weigh accurately to 10 mg. The sample pump should be calibrated every week, or after changing the sample tubes.
46. Immerse the media tubes (inlets I1 to I7 without counterflow tube I8; see Fig. 22.5.2) in the bottle of high-purity water. Call up the dialogue “Calibrate media pump” and follow the instructions. Again weigh the calibration vessel (bottle), then save the calibration. Fill the separation chamber with separation media 47. Make sure that the media pump is switched off. Immerse liquid circuit tubes in the appropriate media: for anode circuit (+ve), immerse in anode circuit solution (100 mM H2SO4); for the cathode circuit (–ve), immerse in cathode circuit solution (100 mM NaOH). 48. Close the safety cover of the electrode circuit bottles and the media pump. Turn on the electrode pump. Check whether the electrode pump delivers properly (flowing air bubbles in the electrode ducts). 49. Immerse media tubes in the corresponding bottles containing separation media and counterflow tube in the corresponding bottle containing counterflow medium. Arrange the media tubes by bottle as follows: Immerse tube for inlet 1 in anode stabilization medium for Basic Protocol 1 Immerse tube for inlet 2 in separation medium 1 for Basic Protocol 1 Immerse tubes for inlets 3 to 5 in separation medium 2 for Basic Protocol 1 Immerse tube for inlet 6 in separation medium 3 for Basic Protocol 1 Immerse tube for inlet 7 in cathode stabilization medium for Basic Protocol 1 Immerse tube for inlet 8 in counterflow medium for Basic Protocol 1. Isolation of Organelles and Proteome Prefractionation
50. Turn on the media pump at 20 rpm for 3 sec (direction “OUT”). Tap the media bottles against the bottle grid, switch on the media pump with 20 rpm (direction “IN”), rinse the chamber with media for 15 min.
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Begin operation of the instrument 51. Set the flow rate to ∼57 ml/hr, the voltage to 1500 V, and the current to 50 mA. Switch on the high voltage. The high voltage cannot be switched on if (a) the safety circuit is open, (b) the electrode pump is switched off, (c) the media pump is switched off, or (d) a connected cooler is not turned on.
52. Wait ∼15 min until the current reached a stable minimum of ∼22 ± 3 mA. Quality control II: check the performance 53. As a control of the performance of the instrument, dilute the pI markers 1:10 with separation medium 3. The pI markers represent a mixture of one red and six yellow dyes having different pIs. The mixture is provided together with the Prolyte solutions.
54. Apply the pI markers at sample inlet S13 (see Fig. 22.5.2) at a sample flow rate of 0.5 ml/hr. As soon as colored drops are visible at the fractionation, wait an additional 10 min. For evaluation, collect the separated pI marker fractions in a 96-well plate by placing it on the drawer and moving it under the fractionation tubing outlets. The 96-well plate can be evaluated by measuring the OD of each well at 400 to 420 and 500 to 530 nm with an appropriate reader. Cross-contaminations during the collection are avoided by tapping the fractionation plate on the fractionation collector housing just before introducing and removing the drawer. The width of the individual pI markers should be no more than 3 to 4 wells, otherwise the separation media have not been prepared properly or the separation chamber has not been assembled properly.
Prepare, apply, and fractionate the sample 55. Dilute concentrated sample solution at least 1:5 (for a final concentration of ∼1 to 5 mg/ml; this can be increased for subsequent experiments if no precipitation occurs) with separation medium 3 for Basic Protocol 1. The chemical and physical properties of the sample and the separation media should be similar. This applies particularly to the density, the conductivity, and the viscosity. For this reason, it is recommended that a concentrated sample solution be prepared and diluted at least 1:5 with separation medium. The total salt concentration of the sample should be no more than 25 to 50 mM. If the salt content of the sample is too high, it should either be desalted or diluted with separation media. Most samples are colorless, which means that it is difficult to detect the position of the sample during the run. For this reason, ProTeam FFE SPADNS can be added to the sample to a concentration of 1%. Turbid protein samples are an indication that protein precipitation has already occurred or that insoluble cell components are still present. Turbid sample solutions must be clarified by filtration or centrifugation before use. It is possible to add nonionic detergents such as CHAPS, octylglucoside, or Triton X-114 (to concentration of 0.1% to 1%) and reducing agents like DTT (up to 50 mM) or uncharged phosphines to the sample.
56. To avoid carryover when applying a new sample, make sure that the separation chamber and the sample tube get rinsed for 30 min (media pump direction “IN”, ∼57 ml/hr; sample pump direction “OUT”, 2 ml/hr). 57. Stop the sample pump to apply the sample. Snip the end of the sample tube. Snipping the end of the sample tube creates a small bubble in the tube which serves as a sharp border between media and sample.
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58. Immerse the sample tube into the sample vial. Turn on the sample pump (direction “IN”) with a high flow rate until the bubble reaches the sample inlet. Apply the sample with a flow rate of ∼1 ml/hr. 59. As soon as the red dye reaches the fractionation manifold, start collecting the sample fractions in a 96-well microtiter plate for ∼5 to 10 min by placing it on the drawer and moving it under the fractionation tubing outlets. Cross-contaminations during the collection are avoided by tapping the fractionation plate on the fractionation collector housing just before introducing and removing the drawer. These early fractions should be collected separately, because the fractionation pattern might not have reached its equilibrium state. When using only a small amount of sample, this separate pre-sample collection should be omitted, but this might result in a lower resolution.
60. Switch to a 96-well deep-well plate and collect the sample fractions until the red dye leaves the separation chamber. Regularly check the sample volume and the media volume.
pH = 4.1
pH = 5.5
fraction 39
fraction 40
fraction 41
Isolation of Organelles and Proteome Prefractionation
Figure 22.5.3 Ultra-zoom 2-D gels of three consecutive fractions of crude rat liver homogenate separated by IEF-FFE.
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IMPORTANT NOTE: Under no circumstances must bubbles be allowed to enter the separation chamber, because this would necessitate the interruption of any given run, as well as the emptying and refilling of the separation chamber.
61. Regularly check for precipitation of the sample in the separation chamber. Precipitation will appear as white lines in the separation chamber. Precipitation can be tolerated as long as no immobile “islands” are formed. If these are observed, a reduction of the sample delivery rate or of the concentration of the sample is required.
62. Switch to an extra 96-well microtiter plate to collect the remaining sample fractions until the red dye is collected completely. As with the early fractions (see annotation to step 59), these late fractions should be collected separately because the fractionation pattern may no longer be in its equilibrium state. When using only a small amount of sample, this separate post-sample collection should be omitted, but this might result in a lower resolution A representative result of Basic Protocol 1 is illustrated in Figure 22.5.3.
Perform active rinsing to clean the apparatus 63. Switch off high voltage. Stop the media pump and the electrode pump. Make sure that the separation chamber is positioned horizontally. 64. Place the counterflow tube as well as the media tubes in a bottle containing at least 1 liter of high-purity water. Place the sample tube in an empty microcentrifuge tube. 65. Operate the media pump for 10 min with 15 rpm (direction “IN”). Operate the media pump for additional 30 min with 50 rpm (direction “IN”). At the same time rinse the sample tube with 4 rpm (direction “OUT”). 66. Rinse the electrodes at the same time (always wear gloves when handling electrode media): a. Remove the two “longer” electrode tubes signed with “+” or “−” from the electrode media. b. Operate the electrode pump until the electrode ducts are almost free of electrode media, then switch off the electrode pump. c. Remove the bottles with electrode media. Place all four electrode tubes in a vessel containing ∼500 ml of high-purity water. d. Operate electrode pump for ∼10 min. e. Remove the two “longer” electrode tubes signed with “+” or “−” from the high-purity water. f. Operate electrode pump until electrode ducts are almost free of water, then switch off electrode pump and remove vessel with water. Perform passive rinsing to clean the apparatus 67. Exchange 96-well plate drawer for rinse tray with approximately 2 liters of high-purity water. Operate media pump at 20 rpm (direction “IN”). Dunk fractionation plate into the rinse tray, then stop media pump. 68. Place counterflow and media tubes on bottle grid and remove vessel with water. Turn bubble trap in the draining-position. Tilt the chamber 45° upwards. 69. Open every pair of clamps and relax clamping force for three complete revolutions (turn counter-clockwise) and close it again. Open upper and lower pairs of clamps completely.
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70. Remove the media pump safety cover and release the tube cassettes by pushing on the lower right side. Release the adjusting screw of the sample pump and place the sample tube on the bottle grid. Switch off the FFE and the cooler. If the FFE is to be used within the next 24 hr, it should remain in this state. If it is not to be used within the next 24 hr, open the chamber on the next day. Rinse the spacer and filter paper with high-purity water and store them dry. Store the membranes in 1:1 glycerol:isopropanol. ALTERNATE PROTOCOL
DENATURING FFE FRACTIONATION OF CRUDE PROTEIN MIXTURES USING THE ISOELECTRIC FOCUSING MODE This protocol is typically used to work under denaturing conditions and to increase the solubility of the proteins. It differs from Basic Protocol 1 in that a new set of separation media containing 8 M urea are used, and in certain corresponding adjustments regarding protocol parameters like voltage, current, and flow rate. Additional Materials (also see Basic Protocol 1) Anode stabilization medium for Alternate Protocol (see recipe) Separation medium 1 for Alternate Protocol (see recipe) Separation medium 2 for Alternate Protocol (see recipe) Separation medium 3 for Alternate Protocol (see recipe) Cathode stabilization medium for Alternate Protocol (see recipe) Counterflow medium for Alternate Protocol (see recipe) Carry out Basic Protocol 1 with the following variations at the indicated steps: 8a. Make sure that the sample tube is connected with the sample inlet S12 (instead of S13; see Fig. 22.5.2). 49a. Immerse media tubes and counterflow tubes as follows: Immerse tube for inlet 1 in anode stabilization medium for Alternate Protocol Immerse tube for inlet 2 in separation medium 1 for Alternate Protocol Immerse tubes for inlets 3 and 4 in separation medium 2 for Alternate Protocol Immerse tube for inlet 5 in separation medium 3 for Alternate Protocol Immerse tubes for inlets 6 an 7 in cathode stabilization medium for Alternate Protocol Immerse tube for inlet 8 in counterflow medium for Alternate Protocol. 51a. Set the voltage to 1200 V (and the current to 50 mA as in Basic Protocol 1). 52a. Wait ∼15 min until the current has reached a stable minimum of ∼17 ± 2 mA. 53a. Dilute the pI markers 1:10 with separation medium 2 instead of separation medium 3. 54a. Apply the pI markers at sample inlet S12 instead of S13. 55a. Dilute concentrated sample solution at least 1:5 in separation medium 2 for Alternate Protocol.
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FFE PURIFICATION OF ORGANELLES USING THE ZONE ELECTROPHORESIS MODE
BASIC PROTOCOL 2
The preparation and execution of a zone electrophoresis FFE experiment for the fractionation of prepurified organelles is presented in this protocol. Like the isoelectric focusing procedure (see Basic Protocol 1), the protocol uses a ProTeam FFE instrument (Fig. 22.5.1). Sucrose is added to the separation media as an osmotic expander whereas EDTA is added to prevent the aggregation of organelles. Materials 1:1 (w/w) glycerol/isopropanol Isopropanol (2-propanol) Kerosene low odor ProTeam FFE SPADNS (sulfanilic acid azochromotrop; Tecan) Anode/cathode circuit solution for Basic Protocol 2 (see recipe) Anode/cathode stabilization medium for Basic Protocol 2 (see recipe) Separation medium for Basic Protocol 2 (see recipe) Anode/cathode stabilization medium for Basic Protocol 2 (see recipe) Counterflow medium for Basic Protocol 2 (see recipe) Organelle suspension for analysis ProTeam FFE system (Tecan) equipped with: Recirculating water cooler and 6-mm i.d. tubing (supplied with instrument, but other water cooler of power ≥350 W may be used) 7 × 0.64–mm i.d. (orange-white) media tubes 1 ×1.42–mm i.d. (yellow-yellow) counterflow tube 1 ×0.25–mm i.d. (orange-blue) sample tube 0.5-mm spacer Anode and cathode membranes 0.8-mm filter paper strips Lint-free paper towels Balance capable of weighing ∼10 mg to 2 kg within 1 mg of accuracy 96-well microtiter plates 96-well deep-well plates 96-well absorbance reader (optional) equ ipped with an appropriate filter for light scattering experiments (λabs ∼420 nm) Additional reagents and equipment for purification of organelles (UNIT 4.2) 1. Disassemble and clean the FFE instrument as in the isoelectric focusing procedure (see Basic Protocol 1, steps 1 to 9), except set the cooler to 5°C. Assemble the instrument 2. Make sure that the sample tube (inlet S13; see Fig. 22.5.2) protrudes 3 to 4 mm into the separation chamber. Orient this protruding end in such a way that it gets squashed between the front and the back plate of the separation chamber, parallel to the direction of the media flow after closing the separation chamber. 3. Wet the 0.5-mm zone electrophoresis spacer with high-purity water and place it on the front plate of the separation chamber. Ensure even distance to the electrode seals and take care that the separation media inlets do not get covered. 4. Continue with the assembly of the instrument as described in Basic Protocol 1, steps 11 to 18, except use 0.8-mm filter paper strips in step 12 of that protocol. Do not use the 0.65-mm filter paper strips that are used with the 0.4-mm spacer, because they are thinner than the ones used for the 0.5 mm spacer.
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5. Fill the instrument (see Basic Protocol 1, steps 19 to 37). 6. Perform quality control I by checking the flow profile (see Basic Protocol 1, steps 38 to 42). 7. Calibrate the pumps (see Basic Protocol 1, steps 43 to 46). 8. Fill the separation chamber with separation media as described in Basic Protocol 1, steps 47 to 50, except: a. In step 47 of Basic Protocol 1, immerse the liquid circuit tubes for both the anode circuit (+ve) and the cathode circuit (–ve) in electrode anode/cathode circuit solution for Basic Protocol 2 (acetic acid/triethanolamine/EDTA). b. In step 49 of Basic Protocol 1, immerse the media tubes as follows: Immerse tube for inlet 1 in anode/cathode stabilization medium for Basic Protocol 2 Immerse tubes for inlets 2 to 6 in separation medium for Basic Protocol 2 Immerse tube for inlet 7 in anode/cathode stabilization medium for Basic Protocol 2 Immerse tube for inlet 8 in counterflow medium for Basic Protocol 2. Begin operation of the instrument 9. Set the flow rate to ∼280 ml/hr, the voltage to 750 V, the current to 150 mA and the power to 200 W. To set the power you have to leave the “Manual Control” menu and switch to the “Options” menu.
10. Switch on the high voltage. The current will reach a value of approximately 120 to 135 mA. The high voltage cannot be switched on if (a) the safety circuit is open, (b) the electrode pump is switched off, (c) the media pump is switched off, or (d) a connected cooler is not turned on.
Prepare, apply, and fractionate the sample 11. Pre-purify the organelles by a classical technology like density-gradient centrifugation (UNIT 4.2). 12. Dilute concentrated suspension of organelles at least 1:5 with separation medium for Basic Protocol 2 or centrifuge the organelles and resuspend with separation medium. The chemical and physical properties of the sample and the separation medium must be similar. This applies particularly to the density, the conductivity and the viscosity. As a starting point, the final protein concentration of the organelles should be around 1 to 2 mg/ml.
13. Check whether the organelles sediment or not by watching if they settle to the bottom of the vessel within 10 to 15 min. If they sediment, turn the fractionation chamber to the vertical position; otherwise keep it in the horizontal position. Organelles that settle to the bottom would experience an additional force (friction) which would lead to band broadening. This can be avoided by using the vertical mode even though this mode is much more sensitive to density differences of the separation media. The samples are colorless, but since most organelles scatter light, they can be visualized by means of a flashlight. Isolation of Organelles and Proteome Prefractionation
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14. To avoid carryover when applying a new sample, make sure that the separation chamber and the sample tube get rinsed for 30 min (media pump direction “IN”, ∼280 ml/hr; sample pump direction “OUT”, 2 ml/hr). 15. Stop the sample pump to apply the sample. 16. Immerse the sample tube into the sample vial. Turn on the sample pump (direction “IN”) with a high flow rate until the sample reaches the sample inlet. Apply the sample with a flow rate of ∼ 1 ml/hr. If the organelle band is too “fat” reduce the flow rate, if it is hardly visible increase the flow rate.
17. Follow the progress of separation with a flashlight. If the major organelle band migrates too rapidly towards the anode, increase flow rate or reduce voltage. If the band migrates too slowly, either lower the flow rate or increase the voltage. 18. As soon as the organelles reach the fractionation manifold, start collecting the sample fractions in a 96-well microtiter plate for ∼5 min by placing it on the drawer and moving it under the fractionation tubing outlets. Cross-contaminations during the collection are avoided by tapping the fractionation plate on the fractionation collector housing just before introducing and removing the drawer. These early fractions should be collected separately, because the fractionation pattern might not have reached its equilibrium state. When using only a small amount of sample, this separate pre-sample collection should be omitted, but this might result in a lower resolution.
0.055
0.050
OD (595 nm)
0.045
0.040
0.035
0.030
0.025 1
6 11 16 21 26 31 36 41 46 51 56 61 66 71 76 81 86 91 96 Fraction number
Figure 22.5.4 Light-scattering profile of a ZE-FFE run of yeast mitochondria prepurified by classical centrifugation techniques. The double peak around fraction 30 corresponds to two distinct populations of highly pure mitochondria. The respective fractions can be pooled, concentrated by centrifugation, and analyzed by electron microscopy or 2-D gel electrophoresis.
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19. Switch to a 96-well deep-well plate and collect the sample fractions until the organelles are reaching the end of the separation chamber. Regularly check the sample volume and the media volume. IMPORTANT NOTE: Under no circumstances must bubbles be allowed to enter the separation chamber, because this would necessitate the interruption of any given run, as well as emptying and refilling of the separation chamber.
20. Switch to an extra 96-well microtiter plate to collect the remaining sample fractions until the organelles are completely collected. As with the early fractions (see annotation to step 18), these late fractions should be collected separately because the fractionation pattern may no longer be in its equilibrium state. When using only a small amount of sample, this separate post-sample collection should be omitted, but this might result in a lower resolution A representative result of Basic Protocol 2 is illustrated in Figure 22.5.4.
21. End the operation by rinsing the apparatus as in Basic Protocol 1, steps 63 to 70, but prolong the operation of the media pump at 50 rpm from 30 min to ≥1 hr (see Basic Protocol 1, step 65), because the sucrose used in the media in this protocol offers perfect conditions for growth of contaminating microorganisms if not completely removed. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
General instructions for preparing FFE media 1. All media must be freshly prepared every day using the stock solutions mentioned in the recipes. 2. It is imperative the different components of each medium be measured by weighing, since the results are more accurate, and the high viscosity of glycerol and HPMC prevents accurate volumetric measurements. A balance capable of weighing 10 mg to 2 kg is needed. 3. It is recommended the pH and conductivity of the solutions be checked, if specified. The instruments recommended for this are: (a) a pH meter with micro pH electrode (diameter = 3 to 5 mm); and (b) conductivity meter (∼10 to 200 mS). 4. Heating of urea leads to its decomposition. The temperature of the media should not exceed the working temperature by more than 15°C. 5. Inhomogenous media prevent good results. Anode/cathode circuit solution (+ve and -ve) for Basic Protocol 2 100 mM acetic acid 100 mM triethanolamine 10 mM EDTA, pH 7.4 There is no actual consumption of this solution over the course of the experiment. 400 g is the minimum amount that is needed to run workday experiments. Use larger amounts for long-term (i.e., >12-hr) experiments.
Anode/cathode stabilization medium for Basic Protocol 2 200 mM sucrose 100 mM acetic acid 100 mM triethanolamine Isolation of Organelles and Proteome Prefractionation
10 mM EDTA, pH 7.4 The actual consumption of this solution over the course of an experiment is ∼80 g/hr.
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Anode circuit (+ve) solution (100 mM H2SO4) for Basic Protocol 1 and Alternate Protocol Prepare 400 g of 100 mM H2SO4 by weighing 40 g of 1 M H2SO4 and adding 360 g of high-purity water. Stir until thoroughly mixed. There is no actual consumption of this solution over the course of the experiment. 400 g is the minimum amount that is needed to run workday experiments. Use larger amounts for long-term (i.e., >12-hr) experiments.
Anode stabilization medium for Alternate Protocol Prepare 68.9 g (∼60 ml) of this solution by combining the following in the order indicated: 6 g 1 M H2SO4 9 g high-purity water 25 g HPMC/glycerol/H2O stock for Alternate Protocol (see recipe) 28.9 g urea Dissolve by stirring at lukewarm temperature The actual composition of this solution over the course of an experiment is ∼27g/hr. The final composition is 14.5% (w/w) glycerol; 0.12% (w/w) HPMC; 100 mM H2SO4; and 8 M urea.
Anode stabilization medium for Basic Protocol 1 Prepare 70 g of this solution by combining the following in the order indicated: 70 g 1 M H2SO4 10.5 g high-purity H20 52.5 g HPMC/glycerol/H2O stock for Basic Protocol 1 (see recipe) Mix thorougly by stirring The actual consumption of this medium over the course of an experiment is ∼9 g/hr. The final composition is: 25% (w/w) glycerol; 0.2% (w/w) HPMC; and 100 mM H2SO4.
Cathode circuit (−ve) solution (100 mM NaOH) for Basic Protocol 1 and Alternate Protocol Prepare 400 g of 100 mM NaOH by weighing 40 g of 1 M NaOH and adding 360 g of high-purity water. Stir until thoroughly mixed. There is no actual consumption of this solution over the course of the experiment. 400 g is the minimum amount that is needed to run workday experiments. Use larger amounts for long-term (i.e., >12-hr) experiments.
Cathode stabilization medium for Alternate Protocol Prepare 137.8 g (∼120 ml) of this solution by combining the following in the order indictated: 12 g 1 M NaOH 18 g high-purity water 50 g HPMC/glycerol/H2O stock for Alternate Protocol (see recipe) 57.8 g urea Dissolve by stirring at lukewarm temperature The actual consumption of this solution over the course of an experiment is ~20 g/hr. The final composition is: 14.5% (w/w) glycerol; 0.12% (w/w) HPMC; 100 mM NaOH; and 8 M urea.
Cathode stabilization medium for Basic Protocol 1 Prepare 70 g of this solution by combining the following in the order indicated: 7 g 1 M NaOH 10.5 g high-purity water 52.5 g HPMC/glycerol/H2O stock for Basic Protocol 1 (see recipe) Mix thoroughly by stirring The actual consumption of this solution of the course of an experiment is ∼9 g/hr. The final composition is 25% (w/w) glycerol; 0.2% HPMC; and 100 mM NaOH. Gel-Based Proteome Analysis
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Counterflow medium for Alternate Protocol Prepare 330.5 g (∼288 ml) of this solution by combining the following in the order indicated: 120 g HPMC/glycerol/H2O stock for Alternate Protocol (see recipe) 72 g high-purity water 138.5 g urea Dissolve by stirring at lukewarm temperature The actual consumption of this solution over the course of an experiment is ∼46 g/hr. The final composition is: 14.5% (w/w) glycerol; 0.12% (w/w) HPMC; and 8 M urea.
Counterflow medium for Basic Protocol 1 Prepare 330 g of this solution by combining the following in the order indicated: 247.5 g HPMC/glycerol/H2O stock for Basic Protocol 1 (see recipe) 82.5 g high-purity water Mix thoroughly by stirring The actual consumption of this solution over the course of an experiment is ∼41 g/hr. The final composition is: 25% (w/w) glycerol/0.2% (w/w) HPMC.
Counterflow medium for Basic Protocol 2 Prepare 280 mM sucrose. The actual consumption of this solution over the course of an experiment is ∼470 g/hr.
HPMC/glycerol/H2O stock for Alternate Protocol 40% (w/w) ProTeam FFE HPMC 0.8% (Tecan) 40% (w/w) ProTeam FFE glycerol (Tecan) 20% (w/w) high-purity H2O HPMC/glycerol/H2O stock for Basic Protocol 1 33% (w/w) ProTeam FFE HPMC 0.8% (Tecan) 33% (w/w) ProTeam FFE glycerol (Tecan) 33% (w/w) high-purity H2O Separation medium 1 for Alternate Protocol Prepare 68.9 g (∼60 ml) of this solution by combining the following in the order indicated: 8 g Prolyte 1 (Tecan) 7 g high-purity water 25 g HPMC/glycerol/H2O stock for Alternate Protocol (see recipe) 28.9 g urea Dissolve by stirring at lukewarm temperature Check that pH is 4.83 ± 0.10 Check that conductivity is 207 ± 15 µS/cm The actual consumption of this solution over the course of an experiment is ~10 g/hr. The final composition is 14.5% (w/w) glycerol; 0.12% (w/w) HPMC; 11.6% (w/w) Prolyte 1; and 8 M urea.
Separation medium 2 for Alternate Protocol Prepare 137.8 g (∼120 ml) of this solution by combining the following in the order indicated: 26.7 g Prolyte 2 (Tecan) 3.3 g high-purity water 50 g HPMC/glycerol/H2O stock for Alternate Protocol (see recipe) 57.8 g urea Dissolve by stirring at lukewarm temperature Check that pH = 7.54 ± 0.10 Check that conductivity is 347 ± 20 µS/cm Isolation of Organelles and Proteome Prefractionation
The actual consumption of this solution over the course of an experiment is ∼20 g/hr. The final composition is 14.5% (w/w) glycerol; 0.12% (w/w) HPMC; 19.4% (w/w) Prolyte 2; and 8 M urea.
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Separation medium 3 for Alternate Protocol Prepare 68.9 g (∼60 ml) of this solution by combining the following in the order indicated: 10 g Prolyte 3 5 g high-purity water 25 g HPMC/glycerol/H2O stock for Alternate Protocol (see recipe) 28.9 g urea Dissolve by stirring at lukewarm temperature Check that pH is 10.26 ± 0.17 Check that conductivity is 235 ± 20 µS/cm The actual consumption of this solution over the course of an experiment is ∼10 g/hr. The final composition is 14.5% (w/w) glycerol; 0.12% (w/w) HPMC; 14.5% (w/w) Prolyte 3; and 8 M urea. Separation medium 1 for Basic Protocol 1 Prepare 70 g of this solution by combining the following in the order indicated: 10 g Prolyte 1 (Tecan) 7.5 g high-purity H2O 52.5 g HPMC/glycerol/H2O stock for Basic Protocol 1 (see recipe) Mix thoroughly by stirring Check that pH is 4.00 ± 0.10 Check that conductivity is 295 ± 25 µS/cm The actual consumption of this solution over the course of an experiment is ∼9 g/hr. The final composition is: 25% (w/w) glycerol; 0.2% (w/w) HPMC; and 14.3% (w/w) Prolyte 1.
Separation medium 2 for Basic Protocol 1 Prepare 210 g of this solution by combining the following in the order indicated: 42 g Prolyte 2 (Tecan) 10.5 g high-purity water 157.5 g HPMC/glycerol/H2O stock for Basic Protocol 1 (see recipe) Mix thoroughly by stirring Check that pH is 6.98 ± 0.10 Check that conductivity is 495 ± 15 µS/cm The actual consumption of this solution over the course of an experiment is ∼27 g/hr. The final composition is 25% (w/w) glycerol; 0.2% (w/w) HPMC; and 20% (w/w) Prolyte 2.
Separation medium 3 for Basic Protocol 1 Prepare 70 g of this solution by combining the following in the order indicated: 10 g Prolyte 3 (Tecan) 7.5 g high-purity water 52.5 g HPMC/glycerol/H2O stock for Basic Protocol 1 (see recipe) Mix thoroughly by stirring Check that pH is 9.85 ± 0.10 Check that conductivity is 302 ± 12 µS/cm The actual consumption of this solution over the course of an experiment is ∼9 g/hr. The final composition is 25% (w/v) glycerol; 0.2% (w/w) HPMC; and 14.3% (w/w) Prolyte 3.
Separation medium for Basic Protocol 2 280 mM sucrose 10 mM acetic acid 10 mM triethanolamine 1 mM EDTA, pH 7.4 The actual consumption of this solution over the course of an experiment is ∼200 g/hr.
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COMMENTARY Background Information
Isolation of Organelles and Proteome Prefractionation
The principles of free-flow electrophoresis were first described by Barrollier et al. (1958) and Hannig (1961). During the following years, the technique was further improved and commercialized by several companies, including Bender & Hobein (Munich, Germany). It was successfully used for zone electrophoresis separations of all kinds of charged compounds ranging from low-molecular-weight organic compounds, peptides, proteins, and organelles, to whole cells. This is documented by several hundreds of publications from the 1960s to the 1980s. Nevertheless, stagnation of the technology, reduced interest in fields like organelle separations, the rise of new fields like molecular biology, and improvements in other techniques like RP-HPLC for peptides and small proteins and flow cytometry for cells gradually ousted FFE and its manufacturers from the market. Only one small German company (Dr. Weber GmbH, Ismaning, Germany; now part of Tecan) managed to survive, with their OCTOPUS instrument (now called ProTeam FFE, after being completely redesigned). They addressed the technological and experimental problems with a variety of innovative concepts (i.e., development of a reproducible IEF process with high resolution and improvement of the robustness of the FFE process by means of stabilization and counterflow media) and combined it with the inherent advantages of FFE. These advantages include the following. a. FFE is carried out in the absence of any matrix, which leads to high sample recovery, fast fractionation, and high sample throughput. The fact that no solid matrix is involved also allows combination of FFE with all kinds of downstream analysis techniques like LC-MS and 2-D electrophoresis b. FFE is based on a continuous principle, which allows virtually unlimited preparative fractionations. c. FFE is a gentle procedure which allows the fractionation of viable cells and active enzymes. With the advent of the proteomics era, FFE is experiencing a second spring, because it offers a portfolio of methods for the rising fractionation and prefractionation needs of proteins (Baldermann et al., 1998; Bernardo et al., 2000; Hoffmann et al., 2001; Lasch et al., 1998; Maida et al., 2000; Song et al., 1998) and organelles (Amigorena et al., 1994; Bard et al.,
2002; Kushimoto et al., 2001; Mohr and Voelkl, 2002; Pierre et al., 1997; Thery et al., 2001). Alternative methods for the (pre-)fractionation of proteins include liquid-based electrophoresis techniques like Rotofor (Bio-Rad), Isoprime (Amersham Biosciences), and Gradiflow (Gradipore, Frenchs Forest, Australia). These methods prove to be useful for certain purification procedures, but all of them depend on some kind of compartmentalization with a membrane/grid/matrix. The passage of a given protein from one compartment to another, based on a pH gradient, for example, inevitably leads to heavy losses of material, because the gradient within the membrane/grid/matrix will be very steep, i.e., most of the proteins of a protein mixture will focus directly in the membrane/grid/matrix. Other methods for the fractionation of organelles are predominantly based on centrifugation (see, e.g., UNIT 4.2). FFE is not intended to replace these methods, but rather uses the resulting fractions to further increase their purity.
Critical Parameters and Troubleshooting Potential mechanical and/or electrical problems that might be encountered during instrument use are addressed in the troubleshooting section of the manufacturer’s manual. Therefore, this discussion will focus on the most common problems encountered with the sample, the separation media, and the handling of the instrument. Separation chamber assembly The proper assembly of the separation chamber is probably the most crucial point. If this is not done well, there is no way of getting significant results from any FFE experiment, and a whole day and precious sample might be wasted. For this reason there are two qualitycontrol experiments included in the protocols. The first is done with no voltage applied and serves to control the laminar flow profile in the separation chamber and the integrity of the tubing. The second is done under “real” fractionation conditions and serves to control the actual performance of the separation chamber. The test mimics the fractionation of the sample by the fractionation of a mixture of seven dyes of different pI, using the same separation media, voltage, temperature, flow rate, etc., that will be used for the actual separation. Even
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though the execution of these control experiments takes some time, it is strongly recommended that they not be skipped. Air bubbles in the separation chamber Entry of air bubbles into the separation chamber must be avoided by all means. Bubbles will massively disturb the laminar-flow profile and usually destroy the fractionation pattern. In the case of IEF-FFE experiments (Basic Protocol 1 and Alternate Protocol), air bubbles in the first third of the separation chamber might be tolerable, because the focusing principle might make up for the distortion. Due to the lack of a focusing force during ZE-FFE experiments (Basic Protocol 2), any kind of bubble in the separation chamber—no matter where it is—causes irreparable distortions. In some cases it might be possible to get rid of air bubbles by reversing the direction of the flow, but in most cases the only recourse is to stop the run, flush the separation chamber with water, empty the separation chamber, open it, and restart the whole process by filling the chamber with water. It is better to prevent the problem by regularly checking the liquid level of the separation medium and the sample. Reagent purity It is important to use only high-purity water as well as chemicals of the highest grade available. This is particularly valid for detergents or other additives that are used for sample preparation or modification of separation media. Media viscosity and density All media should have the same viscosity and density, because differences will inevitably lead to unpredictable phase boundary effects and/or turbulences. This is especially valid for sample solutions and for zone electrophoresis experiments, due to the absence of a focusing effect that could compensate for the distortions. For this reason, all separation solutions should be prepared very carefully. To improve accuracy it is strongly recommended to exclusively add all compounds by weight and not by volume. For example glycerol and HPMC solutions are very viscous, which prevents accurate volumetric measurements. Sample preparation The chemical and physical properties of the sample and the separation media should be similar. To match the density, viscosity, and conductivity of the sample solution and the separation media it is recommended that a con-
centrated sample solution be prepared and that it be diluted at least 1:5 with separation medium. If the sample proteins are not stable at the pH of the separation media, or if the sample concentration would be too low after dilution with the separation medium, the components of the separation media can be added directly to the sample. However, this has to be done very accurately to yield the same viscosity and density as for the separation media. In the case of organelles (see Basic Protocol 2), these can be concentrated by centrifuging and resuspending with the separation medium. Detailed information regarding sample preparation is included in the protocols. The total salt concentration of the sample should be no more than 25 to 50 mM. If the salt content of the sample is too high, it should either be desalted or diluted with separation media. Turbid protein samples are an indication that protein precipitation has already occurred or that insoluble cell components are still present. Turbid sample solutions must be clarified by filtration or centrifugation before use. Protein precipitation Two factors increase the tendency of proteins to precipitate during isoelectric focusing experiments. One is the low solubility of proteins at their isoelectric point, and the other is the high local concentration of focused proteins. Very often, these factors lead to the precipitation of one or a few proteins having the highest concentration and/or the lowest solubility at their pI. This phenomenon can be easily detected by sharp white lines occurring in the separation chamber. As long as these lines stay sharp and keep moving, there is nothing to worry about and one can keep on fractionating the sample. Measures must be taken only when “flakes” (which might clog the fractionation outlets) or stationary “islands” (i.e., stationary broader regions within the sharp white lines which look like a snake that has swallowed its prey, and which might cause flow turbulences and local overheating) are starting to form. In such cases, either the sample flow must be reduced, the sample must be diluted, or nonionic or zwitterionic detergents, e.g., CHAPS, Triton X-100, n-octyl-β-D-glucopyranoside, must be added at concentrations below or around their CMC values (also see APPENDIX 1B).
Anticipated Results With an isoelectric focusing FFE experiment (Basic Protocol 1 and Alternate Protocol) a complex protein mixture should get fractionated in such a way that the majority of individ-
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ual proteins can be detected in a maximum of three consecutive wells out of the 96. A large number of proteins should be further limited to one or two wells (see Fig. 22.5.3). On the other hand, a few highly abundant proteins or proteins that tend to interact with surfaces or with other proteins might have a broader distribution. As a general observation, the resolution in native media is slightly better than in urea-containing media. The resolution of an organelle population after zone electrophoresis FFE should be in a range of 5 to 15 wells (see Fig. 22.5.4). The substantially better resolution of IEF experiments is based on the focusing effect of the pH gradient, which is absent in a zone electrophoresis experiment. The recovery of the samples is highly dependent on the amount of sample. In principle, the absence of any matrix generally allows much higher recoveries as compared to other methods, but the (semi-)preparative dimensions of the instrument cause some losses due to sample adsorption to inner surfaces. When dealing with milliliter and milligram amounts of sample and run times of >2 hr, one can expect to have recoveries >90%, if no precipitation occurs. With microliter, microgram, and smaller amounts of sample, the yield is inescapably limited to less than 50%. The dilution of protein samples after FFE equals ∼3 when dealing with a concentrated sample, based on the following realistic assumptions: (1) the sample is injected into the separation chamber with a flow rate of 1 ml/hr; and (2) the total flow rate of the separation media is ∼60 ml/hr. The flow rate of the counterflow adds about an additional two-thirds of the flow rate of the separation media, based on the comparison of the inner diameter of the counterflow tube and that of the separation media tubes. This means that the total flow that reaches the 96-well plate is ∼100 ml/hr, which is ∼1 ml/well/hr. As mentioned previously, most of the proteins get focused to up to three fractions, which is 3 ml/hr. Coming back to a sample flow rate of 1 ml/hr, this amounts to a 3-fold dilution. It is worth noting that dilute samples could even be concentrated by FFE by applying them via a separation media tube (∼9 ml/hr), instead of the sample tube. The dilution of organelle samples after FFE is not very important, because organelles can easily be concentrated by centrifugation. Isolation of Organelles and Proteome Prefractionation
Time Considerations Time requirements for all steps involved in cleaning, assembling, filling, calibrating, and checking the performance of the instrument, including preparation of the sample and the separation media, add up to ∼2 to 3 hr for experienced users. The actual time needed for the application and fractionation of a sample is highly dependent on the sample amount, concentration, and solubility. The expression “run time” does not make sense in this context, because FFE is a continuous technique, in contrast to classical gel-based electrophoresis or chromatography. The application of 1 ml of sample per hr containing 1 mg of protein serves as a good starting point. For soluble cytosolic protein mixtures, this could easily be increased to 2 to 3 ml of sample per hr containing 10 to 20 mg of protein. Total application time is virtually unlimited based on the continuous fractionation principle. Washing time between the application of different samples is about 30 min, and active rinsing time at the end of operation is ∼45 min. Passive rinsing is done overnight without any hands-on time.
Literature Cited Amigorena, S., Drake, J.R., Webster, P., and Mellman, I. 1994. Transient accumulation of new class II MHC molecules in a novel endocytic co mpartment in B ly mph ocytes. Nature 369:113-119. Baldermann, C., Lupas, A., Lubieniecki, J., and Engelhardt, H. 1998. The regulated outer membrane protein Omp21 from Comamonas acidovorans is identified as a member of a new family of eight-stranded beta-sheet proteins by its sequence properties. J. Bacteriol. 180:3741-3749. Bard, F., Patel, U., Levy, J.B., Horne, W.C., and Baron, R. 2002. Molecular complexes that contain both c-Cbl and c-Src associate with Golgi membranes. Eur. J. Cell. Biol. 81:26-35. Barrollier, J., Watzke, E., and Gibian, H. 1958. Einfache Apparatur für die trägerfreie präparative Durchlauf-Elektrophorese. Z. Naturforschung 13b:754-755. Bernardo, K., Krut, O., Wiegmann, K., Kreder, D., Micheli, M., Schafer, R., Sickman, A., Schmidt, W.E., Schroder, J.M., Meyer, H.E., Sandhoff, K., and Kroenke, M. 2000. Purification and characterization of a magnesium-dependent neutral sphingomyelinase from bovine brain. J. Biol. Chem. 275:7641-7647. Hannig, K. 1961. Die trägerfreie kontinuierliche Elektrophorese und ihre Anwendung. Z. Anal. Chem. 181:244-254. Hoffmann, P., Ji, H., Moritz, R.L., Connolly, L.M., Frecklington, D.F., Layton, M.J., Eddes, J.S., and Simpson, R.J. 2001. Continuous free-flow electrophoresis separation of cytosolic proteins
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from the human colon carcinoma cell line LIM 1215: A non-two-dimensional gel electrophoresis-based proteome analysis strategy. Proteomics 1:807-818. Kushimoto, T., Basrur, V., Valencia, J., Matsunaga, J., Vieira, W.D., Ferrans, V.J., Muller, J., Appella, E., and Hearing, V.J. 2001. A model for melanosome biogenesis based on the purification and analysis of early melanosomes. Proc. Natl. Acad. Sci. U.S.A. 98:10698-10703. Lasch, J., Moschner, S., Sann, H., Zellmer, S., and Koelsch, R. 1998. Aminopeptidase P−A cell surface antigen of endothelial and lymphoid cells: Catalytical and immuno-histotopical evidences. Biol. Chem. 379:705-709. Maida, R., Krieger, J., Gebauer, T., Lange, U., and Ziegelberger, G. 2000. Three pheromone-binding proteins in olfactory sensilla of the two silkmoth species Antheraea polyphemus and Antheraea pernyi. Eur. J. Biochem. 267:2899-2908. Mohr, H., and Voelkl, A. 2002. Isolation of peroxisomal subpopulations from mouse liver by immune free-flow electrophoresis. Electrophoresis 23:2130-2137. Pierre, P., Turley, S.J., Gatti, E., Hull, M., Meltzer, J., Mirza, A., Inaba, K., Steinman, R.M., and Mellman, I. 1997. Developmental regulation of
MHC class II transport in mouse dendritic cells. Nature 388:787-792. Song, J.-F., Liu, T., Shen, X., Wu, G.-D., and Xia, Q.-C. 1998. Application of free-flow electrophoresis to the purification of trichosanthin from a crude product of acetone fractional precipitation. Electrophoresis 19:1097-1103. Thery, C., Boussac, M., Veron, P., Ricciardi-Castagnoli, P., Raposo, G., Garin, J., and Amigorena, S. 2001. Proteomic analysis of dendritic cell-derived exosomes: A secreted subcellular compartment distinct from apoptotic vesicles. J. Immunol. 166:7309-7318.
Key References Krivankova, L. and Bocek, P. 1998. Continuous free-flow electrophoresis. Electrophoresis 19:1064-1074. Excellent review of the principles of the method, the possible techniques and a variety of applications.
Contributed by Peter J. A. Weber, Gerhard Weber, and Christoph Eckerskorn Tecan Munich GmbH Kirchheim, Germany
Gel-Based Proteome Analysis
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Protein Profiling by Microscale Solution Isoelectrofocusing (MicroSol-IEF)
UNIT 22.6
The great complexity and broad dynamic range of most eukaryotic proteomes, together with limitations of the existing 2-D gel approach and of alternative MS/MS methods, suggest that sample prefractionation is essential for more comprehensive coverage and reliable detection of low-abundance proteins in complex proteomes. An efficient and reproducible new method for this purpose is microscale solution isoelectrofocusing (MicroSol-IEF), which seamlessly interfaces with narrow-pH-range 2-D gels, enhancing data obtained from protein profiling studies, including quantitative proteome comparisons (Zuo and Speicher, 2000, 2002, Zuo et al., 2002). Complex proteomes, such as mammalian cell extracts and biological fluids, are separated into well resolved fractions on the basis of isoelectric point (pI) by means of the MicroSolIEF device, now commercially available from Invitrogen as the ZOOM IEF Fractionator. This consists of tandem small-volume (650- to 700-µl) chambers separated by thin, porous, acrylamide gel membranes that contain Immobilines (Amersham Biosciences) at specific pHs. After proteins are isoelectrically focused to equilibrium, each chamber contains the proteins with pIs between the pHs defined by the two boundary membranes, resulting in concentrated fractions that can be further analyzed using 2-D gels and other separation methods. This unit focuses on the MicroSol-IEF method for prefractionation of complex proteomes using the ZOOM IEF Fractionator with commercially available pH partition membranes (see Basic Protocol 1). Since the fractionator was designed with a flexible configuration, it is possible to create alternative setups for this device to separate samples into different numbers of fractions with various pH ranges and volumes (see Alternate Protocol 1). As only a few pH partition membranes are produced commercially, a detailed method is provided for preparing custom pH membranes to create alternative MicroSol-IEF configurations (see Support Protocol 1). In addition, methods are provided for evaluating the effectiveness of the prefractionation using 1-D and 2-D gel electrophoresis (see Support Protocols 2 and 3). Approaches for quantitative protein profiling that incorporate MicroSol-IEF are also discussed (see Strategic Planning).
STRATEGIC PLANNING In its simplest form, the MicroSol-IEF approach serves as a crucial prefractionation step for the comprehensive analysis of a single complex proteome. However, this approach can be integrated into broader experimental schemes that seek to provide more meaningful proteomic data, i.e., quantitative protein profiling. In this context, the versatility of MicroSol-IEF becomes clear when one considers how it can be interfaced with other proteomic approaches, both non-gel-based and gel-based, and dependent on or independent of different protein labeling species (Fig. 22.6.1). A label-free approach involves the use of narrow, overlapping 2-D gels for comparing <100-kDa proteins from two samples (see Basic Protocol 2) and large-pore 1-D gels for large (>100-kDa) proteins (see Support Protocol 4) following prefractionation. Alternatively, proteins may be labeled with cyanine dyes before prefractionation as part of an approach that combines MicroSol-IEF with two-dimensional differential gel electrophoresis (2-D DIGE; see Support Protocol 5 and UNIT 22.2). Prefractionation may also be integrated with stable isotope labeling and multi-dimensional LC/LC-MS/MS procedures (see Alternate Protocol 2). Non-Gel-Based Proteome Analysis Contributed by Xun Zuo, KiBoem Lee, Nadeem Ali-Khan, and David W. Speicher Current Protocols in Protein Science (2005) 22.6.1-22.6.29 C 2005 by John Wiley & Sons, Inc. Copyright
22.6.1 Supplement 39
Microscale Solution Isoelectrofocusing (MicroSol - IEF)
Figure 22.6.1
Legend at right.
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PREFRACTIONATION OF PROTEOMES INTO FIVE FRACTIONS USING THE ZOOM IEF FRACTIONATOR
BASIC PROTOCOL 1
This protocol pertains to the ZOOM IEF configuration in which six commercially available pH membranes (Invitrogen) are used to create five separation chambers.
Materials 1× Microsol-IEF anode buffer (see recipe) 1× Microsol-IEF cathode buffer (see recipe) Protein sample (e.g., cell extract) MicroSol-IEF sample buffer (see recipe) ZOOM IEF Fractionator (Invitrogen; Fig. 22.6.2) pH partition membranes (ZOOM pH Disks; Invitrogen): six different pH membranes (pH 3.0, 4.6, 5.4, 6.2, 7.0, and 10.0) 0.22-µm Ultrafree-MC micro-filter (Millipore) Power supply capable of operating at low currents (0.1 to 1 mA) and at least 1000 V 24-well tissue culture plates (Becton Dickinson Labware) Assemble the ZOOM IEF Fractionator 1. Organize the constituent parts of the device (Fig. 22.6.2). 2. Place O-rings on the separation chambers and insert a plug into the port of each chamber (see Fig. 22.6.3). 3. Insert the anode end sealer into the chamber assembly tube, followed by a spacer. 4. Insert a chamber (with its port plug and O-ring) into the assembly tube. The O-ring should be facing down. The use of a spacer, rather than a membrane, between the electrode chamber and this first potential separation chamber means that, in this configuration, the first chamber will actually contain anode electrode buffer (Fig. 22.6.4A).
5. Using forceps to hold the membrane’s edge (Fig. 22.6.5A), place a pH 3.0 partition membrane into the inserted separation chamber. 6. Insert the next separation chamber, making sure that the port plug is in place and the O-ring is facing down. Push the chamber into the chamber assembly tube until it is flush with the top of the tube (Fig. 22.6.5B). 7. Repeat steps 4 to 6 for the pH 4.6, 5.4, 6.2, 7.0, and 10.0 membranes and separation chambers (Fig. 22.6.5A and B). Invitrogen now produces pH 9.1 and 12.0 membranes, in addition to the six membranes referred to here.
8. Insert a spacer followed by the cathode end sealer (Fig. 22.6.5C).
Figure 22.6.1 (at Left) Strategic planning for interfacing MicroSol-IEF prefractionation with downstream protein analyses for comprehensive protein profiling and quantitative comparisons. As shown in the top portion of the figure, nonlabeled protein samples should be fractionated individually with a ZOOM IEF Fractionator and the fractions subsequently analyzed on overlapping narrow pH range 2-D gels for smaller proteins (<100 kDa) and on large pore 1-D gels for the large proteins (>100 kDa). Alternatively, either 2-D DIGE (A) or ICAT (B) reagents can initially label protein samples that are combined for a single MicroSol-IEF prefractionation. As depicted in panel A, for further analysis of the fluorescently labeled fractions, the same 1-D and 2-D gels as described for the unlabeled proteins can be used. To interface with conventional LC-MS/MS methods (see panel B), the MicroSol-IEF fractions are run into a 1-D mini-gel for a very short distance (∼5 mm) to remove interfering chemicals from the samples. After in-gel digestion of the entire gel sample from each fraction, the peptides are fractionated and purified by a SCX column followed by analysis with LC-MS/MS methods.
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Figure 22.6.2 The ZOOM IEF Fractionator. (A) A completely assembled device and lid. (B) The major internal components of the device needed for prefractionation using seven pH ranges: top row, eight O-rings that will be placed on the grooves of the chambers; middle row (from left to right), anode end sealer, seven oval chambers (with plugs on and grooved sides facing up), cathode end sealer, and cathode end screw cap; bottom row, eight membrane partitions (ZOOM pH disks), which are acrylamide/Immobiline gels cast on oval polyethylene disks that will be placed between the chambers.
9. Fasten the cathode end screw cap onto the assembly tube (Fig. 22.6.5D). Do not overtighten, as excess pressure distorts the Teflon chambers and causes air leaks and subsequent sample loss from leaking chambers due to electro-osmosis.
10. Insert the assembly tube containing the anode electrode and all the separation chambers into the cathode electrode chamber. 11. Fill the anode and cathode electrode chambers with 1× IEF anode and cathode buffer, respectively (∼19 ml per chamber). Typically, 8 M urea and 2 M thiourea are included in the electrode buffers to minimize electro-osmosis during IEF (see Reagents and Solutions).
Prepare and load a protein sample into the ZOOM IEF Fractionator 12. Prepare a protein sample (e.g., cell extract) in MicroSol-IEF sample buffer. Microscale Solution Isoelectrofocusing (MicroSol - IEF)
Depending upon the number of separation chambers and the amount and volume of the sample to be used, the sample prepared in sample buffer can be further diluted with additional sample buffer to a final volume equal to the total volumes of the chambers
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Figure 22.6.3 Preparation of a sample chamber for the ZOOM IEF Fractionator. (A) The unassembled unit parts; (B) insertion of a plug on the sample chamber through the fill port provided; (C) placement of an O-ring on the chamber’s groove.
that will be loaded with sample. See Commentary for information about final IPG buffer concentrations.
13. Immediately prior to loading, filter the sample through a 0.22-µm Ultrafree-MC microfilter unit by microcentrifuging 10 min at 10,000 × g, room temperature, to remove any particulate material. 14. Remove the fill plugs from the chambers and load the sample into the desired separation chamber(s) at ∼700 µl per chamber. Load sample buffer into the remaining chambers. Note that since there are spacers between the electrodes and terminal separation chambers, the latter contain electrode buffers, not sample or sample buffer. The protein sample can be loaded into one or more of the five innermost separation chambers for prefractionation. Figure 22.6.6 illustrates the loading of a sample into one central chamber.
15. Insert the plugs into the ports of all the chambers. It is important to leave a small gap of air in each chamber after loading the sample or buffer and to tightly seal the chamber ports with the plugs to minimize electro-osmosis during IEF, because excessive electro-osmosis will deplete most of the liquid from the affected chamber and result in poor protein separation.
Run ZOOM IEF to fractionate sample proteins 16. Focus proteins by gradually increasing the applied voltage from ∼50 V, without allowing the current to exceed 1 mA, until a maximum of 1000 V is attained. Prefractionation of a highly complex proteome such as mammalian cell extract typically requires a power supply capable of operating at a voltage of at least 1000 V and at currents as low as 0.1 mA. If possible, set maximum current and power limits at 1 mA and 1 W, respectively. Depending upon the programming capacity of the power supply, voltage can be increased manually in a stepwise manner or automatically as the current falls.
Non-Gel-Based Proteome Analysis
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Figure 22.6.4 (A) A schematic of the basic ZOOM IEF Fractionator unit containing five pH-range separation chambers. The device consists of small Teflon chambers (total volume ∼750 µl; preferred sample loading volume 650 µl) separated by six commercial pH partition membranes (pH values as indicated). In this configuration for five fractions, the two terminal chambers are coupled to the electrode chambers with spacers equal in thickness to the partition membranes but with a central hole equal to the bore of the chambers. Hence, these terminal chambers and the adjacent electrode chambers are contiguous and will be loaded with anode and cathode IEF electrode buffers respectively, instead of sample. (B) A schematic illustration of a ZOOM IEF Fractionator unit containing the maximum of seven pH-range separation chambers. This configuration for the seven fractions is similar to that for the five fractions (panel A), except that there are no spacers and eight custom-made pH partition membranes (pH as indicated) are placed between the chambers and terminal chambers with electrodes.
17. After the current drops to a stable value (e.g., <0.5 mA), continue focusing at 1000 V for a further 30 min, then switch the power supply off. MicroSol-IEF conditions, including the total time required for prefractionation of a protein mixture, are dependent on the sample type, quantity of protein, sample ionic strength and number of chambers initially loaded with sample. Some simple samples, such as prokaryotic cell extracts, may require less time and lower voltages for adequate focusing, as previously reported (Zuo and Speicher, 2000). The manufacturer also recommends a maximum voltage of 600 V. However, many complex samples, such as mammalian cell or tissue extracts and biological fluids, may be incompletely focused unless higher voltages, e.g., 1000 V, are used. It should be noted that excessive heat and electrical force may damage the pH partition membranes. In the authors’ laboratory, voltage ramping conditions for the separation of human breast cancer cell extract were: 50 V for 1 hr, 100 V for 1.5 hr, 200 V for 30 min, 350 V for 30 min, 600 V for 20 min, and finally 1000 V until the lowest possible current was reached (0.2 to 0.3 mA) and maintained for 30 min (a total of 1 to 1.5 hr at 1000 V). The total run time for prefractionation of this sample was thus about 5 hr. Microscale Solution Isoelectrofocusing (MicroSol - IEF)
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Figure 22.6.5 Assembly of chambers and partition membranes for the ZOOM IEF Fractionator. (A) Placement of a pH partition membrane into a separation chamber using forceps. (B) After placing a partition membrane, the next separation chamber is carefully inserted with the O-ring side down. (C) After all membranes and separation chambers are inserted, the cathode end sealer is added. (D) After inserting all components, the cathode end screw cap is screwed onto the top of the illustrated assembly to compress the O-rings, forming seals between the tandem Teflon chambers and end units.
Figure 22.6.6 Loading of a protein sample into the ZOOM IEF Fractionator. This figure shows the loading of an entire protein sample into a single central chamber.
Collect fractions and extract proteins from partition membranes 18. Withdraw fractions through the port of each chamber using a pipet and transfer to 1.5-ml microcentrifuge tubes. 19. Rinse the inside walls of the individual chambers and surfaces of the partition membranes by adding a small volume (∼100 µl) of MicroSol-IEF sample buffer to each chamber and gently agitating the device briefly. Combine each rinse with the appropriate fraction.
Non-Gel-Based Proteome Analysis
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20. Remove the pH partition membranes and transfer them to individual wells of a 24well tissue culture plate, each containing 200 µl MicroSol-IEF sample buffer. Shake for 30 min at room temperature. Remove sample buffer to a clean tube and repeat membrane protein extraction with fresh sample buffer. Combine the two extractions from each membrane in a single 1.5-ml microcentrifuge tube. 21. Analyze fractions and membrane extracts by 1-D (UNIT 10.1) or 2-D (UNIT 10.4) electrophoresis. Divide any samples and membrane extracts that will not be used immediately into aliquots and store up to 1 year or more at −80◦ C. ALTERNATE PROTOCOL 1
SEPARATING PROTEOMES INTO SEVEN FRACTIONS USING CUSTOM-MADE PARTITION MEMBRANES As an alternative to the typical separation of a proteome into five fractions (see Basic Protocol 1), the ZOOM IEF Fractionator can also be configured to perform a sevenfraction separation (Fig. 22.6.4B). Four very narrow pH fractions, each separated by 0.5 pH units, encompass a range of pH 4.4 to 6.4 in the center of the apparatus. Most eukaryotic proteomes have a high proportion of proteins with pI values in this range. The procedures for preparing and loading the sample into the ZOOM IEF device, running the fractionator, collecting the fractions, and extracting the proteins from the partition membranes, are as described in Basic Protocol 1. The configuration should be selected based on the particular prefractionation goal. For example, mammalian serum contains albumin accounts for more than 50% of its total protein content and severely restricts the amount of sample that can be applied to a 2-D gel. Therefore, it is useful to sequester albumin in a single narrow pH range fraction (e.g., pH 5.4 to 5.8) to enhance the detection of low-abundance proteins in other fractions (Zuo et al., 2001, Zuo and Speicher, 2002). Custom-made membranes of the type used in Basic Protocol 1 can be prepared at desired pH values (Support Protocol 1) according to the requirements of the experiment. Furthermore, the actual number of fractions as well as the pH ranges and volumes of the fractions can easily be varied. This is achieved by altering the placement of spacers and membranes, as shown in Table 22.6.1. For example, the volume of a fraction can be equal to the optimal load volume (∼650 µl) of the chamber housing it, or two or more chambers can be connected by placing spacers instead of membranes between them to Table 22.6.1 Alternative Configurations for ZOOM IEF Fractionator Units
pH membrane/spacer placementa
Separation mode 1
2
3
4
5
6
7
8
5 fractions, uniform volumes
S
3.0
4.6
5.4
6.2
7.0
10.0
S
5 fractions, 2 double volumes
3.0
4.6
S
5.4
6.2
S
7.0
10.0
4 fractions, uniform minimal volumes
S
S
3.0
5.4
6.2
7.0
10.0
S
3 fractions, variable volumes
3.0
S
5.4
S
6.2
S
S
10.0
7 fractions, uniform volumes
3.0
4.4
4.9
5.4
5.9
6.4
8.1
10.0
5 fractions, uniform minimal volumes
S
3.0
4.9
5.4
6.4
8.1
10.0
S
4 fractions, variable volumes
3.0
S
S
5.0
6.3
S
7.7
10.0
Commercially available membranes
Custom-made membranes
Microscale Solution Isoelectrofocusing (MicroSol - IEF)
a Either a pH partition membrane or a spacer (S) is placed between the chambers at the indicated positions in the unit.
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increase the volume of one or more selected fractions. Where feasible, experiments should be designed so that fraction volumes and protein concentrations are directly compatible with downstream analysis steps without the need to concentrate fractions. For example, a 650-µl fraction volume, plus a 150-µl rinse of the chamber, yields a total volume of 800 µl. This entire fraction can be analyzed on duplicate 24-cm IPG strips using in-gel rehydration as the loading method (UNIT 10.4).
Additional Materials (also see Basic Protocol 1) Eight custom-made pH partition membranes (see Support Protocol 1) For this protocol, follow steps 1 to 21 of Basic Protocol 1, making the following modifications. Insert a pH 3.0 partition membrane instead of a spacer between the anode end sealer and the first separation chamber. Insert the other custom-made membranes in order (see Fig. 22.6.4B), ensuring that the pH 10.0 membrane is between the cathode end sealer and the last separation chamber, instead of a spacer.
PREPARATION OF AN ACRYLAMIDE/IMMOBILINE PARTITION MEMBRANE AT A SPECIFIC pH
SUPPORT PROTOCOL 1
The pH partition membranes for the ZOOM IEF Fractionator are acrylamide/Immobiline gels cast on strong (polyethylene) support disks. The partition membranes can be made with different acrylamide gel concentrations, but in general these should remain as low as possible for large pore sizes while maintaining sufficient strength, typically 4%T/10%C (where T represents total acrylamide and C represents the cross-linker), so that very large proteins can be efficiently focused. Currently, six different Immobilines (pK values 3.6, 4.6, 6.2, 7.0, 8.5, and 9.3) are available as 0.2 M stock solutions from Amersham Biosciences. Various Immobilines are mixed to produce a buffering capacity at any desired pH between 3.0 and 10.0. Typically, the amount and types of Immobilines used for desired pH values are calculated using a computer program, “Doctor pH,” originally developed by Giaffreda et al. (1993) and commercially available from Amersham Biosciences. Alternatively, one can use pH recipes from formula tables that have been calculated using this software (Pharmacia Biotech, 1997). Generally, there are two steps in preparing partition membranes. First, Immobilines of different pK values are mixed for the desired pH values. Second, acrylamide/Immobiline gels are cast on porous polyethylene disks (∼0.67 mm thickness) that provide mechanical strength. The 4%T/10%C acrylamide gels containing Immobilines at specific pH values are cast on thin support disks to facilitate the migration of proteins, including very large ones (≥500 kDa), through the membranes. The general method for preparing partition membranes at desired pH values is described below. Water used for all reagents should be a high-quality research grade such as Milli-Q or equivalent.
Materials 0.2 M Immobiline stock solutions of desired pH ranges (Amersham Biosciences) 1 M Tris base 1 M acetic acid 30%T/10%C acrylamide/bisacrylamide stock (see recipe) 87% (v/v) glycerol (Amersham Biosciences) 40% ammonium persulfate (Bio-Rad), prepare immediately before use TEMED (Bio-Rad) 12% (v/v) glycerol Partition membrane storage solution (see recipe)
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Table 22.6.2 Preparation of Immobilinea Mixtures at Specific pH Values
pH 3.0
pH 4.4
pH 4.9
pH 5.4
pH 5.9
pH 6.4
pH 8.1
pH 10.0
pK 3.6
1548 µl
615 µl
479 µl
331 µl
280 µl
228 µl
—
—
pK 4.6
60 µl
236 µl
321 µl
348 µl
438 µl
529 µl
206 µl
60 µl
pK 6.2
60 µl
472 µl
363 µl
247 µl
204 µl
161 µl
760 µl
—
pK 7.0
—
48 µl
89 µl
107 µl
144 µl
181 µl
299 µl
1707 µl
pK 8.5
—
—
—
—
—
—
218 µl
128 µl
pK 9.3
96 µl
138 µl
258 µl
314 µl
424 µl
532 µl
17 µl
356 µl
Water
5.74 ml
5.88 ml
5.91 ml
6.11 ml
5.98 ml
5.87 ml
5.92 ml
5.25 ml
Total volume
∼7.5 ml
∼7.5 ml
∼7.5 ml
∼7.5 ml
∼7.5 ml
∼7.5 ml
∼7.5 ml
∼7.5 ml
a Immobilines (each at 0.2 M; Amersham Biosciences) at different pK values are combined, with the final Immobiline concentration in each gel membrane being ∼12 mM. When the seven-fraction separations are conducted, custom partition membranes are used, which are cast using the above Immobiline mixtures at the indicated pH values. Alternatively, the commercially available pH 3.0 and 10.0 partition membranes can be used to complete the pH series required (see Table 22.6.1).
15-ml conical centrifuge tubes Doctor pH software (Amersham Biosciences) Gel support films (Bio-Rad Laboratories) ∼ 10.4 × 12.6–cm lid from box of 1-ml pipet tips (e.g., Fisher) Repel-Silane (Amersham Biosciences) 1.5-mm slab gel spacers (Bio-Rad) Precut oval disks of hydrophilic porous polyethylene, 0.67 mm thick (Invitrogen) Sharp scalpel or razor blade 24-well tissue culture plates (Becton Dickinson Labware) Platform shaker Sealing tape (Fisher) Prepare an Immobiline mixture at a specific pH 1. Mix the appropriate amount of each 0.2 M stock Immobiline and water in a 15-ml centrifuge tube. As described above, the amount of Immoblines required for pHs ranging from pH 3.0 to 10.0 can be calculated using the “Doctor pH” software (Giaffreda et al., 1993). Table 22.6.2 lists recipes that have been modified from precalculated amounts of Immobilines (Pharmacia Biotech, 1997) for the eight pH partition membranes commonly used in prefractionation of mammalian cell extracts (see Fig. 22.6.4B). The resulting pH ranges of the fractions match the commercially available large IPG strips described in Basic Protocol 2.
2. Measure the pH of the mixture immediately using a pH meter with an accuracy of at least 0.01 pH unit. 3. If the observed pH deviates from that required, carefully adjust to the desired pH by adding a small amount of the most acidic (pK 3.6) or most basic (pK 9.3) stock Immobiline. 4. Titrate the Immobiline mixture to pH ∼6.5 using either 1 M Tris base or 1 M acetic acid. Microscale Solution Isoelectrofocusing (MicroSol - IEF)
Adjusting the pH of the Immobiline mixture to ∼6.5 is important for efficient polymerization of the acrylamide/Immobiline gel. Tris and acetic acid are not covalently incorporated into the gel matrix and will be washed out after polymerization.
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Figure 22.6.7 Placement of materials for casting custom pH partition membranes. The plastic cover from a 1.0-ml pipettip box is used as a convenient container. Typical placement of 24 oval porous polyethylene disks is shown—i.e., the discs are distributed (without touching each other) on top of a silanized gel support film with four 1.5-mm gel spacers around the edges. As discussed in the text, it is important to pipet the gel solution directly onto the disks to remove air trapped in the disc pores before adding extra solution to the remaining area.
Cast acrylamide/Immobiline gel on support disks 5. Trim two gel support films to ∼10.4 × 12.6–cm, so they can fit inside the lid of a 1-ml pipet tip box. 6. Coat the two support films with Repel-Silane according to the manufacturer’s instructions. 7. Insert one silanized film into the pipet-tip box lid and place 1.5-mm gel spacers at the edges of the film (see Fig. 22.6.7). 8. Place 24 oval porous polyethylene disks on the film, ensuring they do not touch each other (Fig. 22.6.7). Blank disks are precoated with a surfactant; do not rinse or wash prior to use. The porous polyethylene is also available as large sheets from POREX. Non-Gel-Based Proteome Analysis
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Table 22.6.3 Acrylamide/Immobiline Gel Working Solution
Component
Volume
Final concentration
Immobiline mixture (at specific pH)a
7.50 ml
∼12 mM
Acrylamide/bis stock (30%T/10%C)b
3.33 ml
4%T/10%C
Glycerol (87% v/v)
3.45 ml
12% (v/v)
Milli-Q H2 O
10.70 ml
—
Ammonium persulfate (40% w/v)
20 µl
0.08% (w/v)
TEMED
10 µl
0.04% (v/v)
Total volume
∼25 ml
—
a See Support Protocol 1, step 4. b See recipe in Reagents and Solutions.
9. Prepare acrylamide/Immobiline gel working solution by combining the Immobiline mixture of desired pH (step 4) with 30%T/10%C acrylamide/bis stock, 87% glycerol, Milli-Q water, ammonium persulfate, and TEMED, according to Table 22.6.3. The 25-ml gel working solution is sufficient for casting 24 disk membranes at a specific pH.
10. Immediately degas the working solution for ∼1 min. 11. Immediately pipet the working solution directly onto the disks until the solution is completely absorbed inside the polyethylene pores and all air is displaced. Continue adding gel solution to uniformly cover the remaining areas of the film until the liquid is as high as the 1.5-mm spacers at the edges of the film. 12. Immediately cover the solution and disks with the second silanized support film. Leave the gel to polymerize at room temperature (∼2 hr). 13. After polymerization is complete, remove the disks and trim excess gel from both sides of each disk using a sharp scalpel or razor blade. Gently trim excess gel from the edges of each disk also. 14. Immediately transfer each disk into an individual well of a 24-well tissue culture plate. 15. Wash disks three times, each time by adding 2 ml of 12% glycerol to each of the disk-containing wells, shaking the plate on a platform shaker at room temperature, then aspirating the glycerol solution. Sufficient washing is necessary to remove the acetic acid or Tris and polymerization byproducts.
16. After the final wash, replace the 12% glycerol solution with 1 ml of the partition membrane storage solution in each well and tightly seal the plate with the sealing tape. Partition membranes can be used immediately or stored at 4◦ C for up to at least 6 weeks without losing effectiveness.
Microscale Solution Isoelectrofocusing (MicroSol - IEF)
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EVALUATION OF THE EFFECTIVENESS OF ZOOM IEF PREFRACTIONATION USING 1-D MINIGELS
SUPPORT PROTOCOL 2
A simple initial check of fractionation results involves running proportional amounts of all fractions and partition membrane extracts on 1-D 10% SDS minigels. Proteins resolved on 1-D gels can be visualized (UNIT 10.5) by using either Coomassie blue stain or highersensitivity stains such as Sypro Ruby or silver stains (e.g., SiverQuest Silver Staining Kit, Invitrogen), by means of which overall protein recoveries and distribution among fractions and membrane partitions can be observed. In addition, estimates of total protein in each lane can be obtained using 1-D gel image analysis software such as Discovery Series Quantity One (version 4.2.0, Bio-Rad). These 1-D gels are particularly useful for preliminary assessments regarding the reproducibility of replicate prefractionation runs. Representative results from prefractionation of a human breast cancer cell extract into seven pH ranges using the ZOOM IEF Fractionator with the eight custom-made partition membranes (see Fig. 22.6.4B for schematic) are shown in Figure 22.6.8. Silver stain was used in this case to maximize detection sensitivity, but a disadvantage of this stain is its narrow dynamic range. Thus, many bands from the fractions are outside the linear range of the stain. As a result, minor bands in the membrane extracts appear somewhat overemphasized. Using 1-D gels for evaluating the effectiveness of MicroSolIEF prefractionation is a high-throughput approach, but the gels provide only limited data. For example, they cannot be used to verify the pH boundaries of the fractions. 2-D gels (see Support Protocol 3) allow a much fuller evaluation of the prefractionation.
Figure 22.6.8 An initial evaluation of MicroSol-IEF fractions using a 1-D mini SDS gel. After 3.0 mg of a MCF-7/6 human breast cancer cell extract were fractionated into seven fractions (see Fig. 22.6.4B for schematic), small proportional aliquots of the fractions and the extracts from partition membranes were separated on a 10% SDS Bis-Tris NuPAGE minigel (Invitrogen), and the gel was silver stained. Lane definitions: F1 to F7, fractions 1 to 7; M1 to M8, extractions from the partition membranes with pH 3.0, 4.4, 4.9, 5.4, 5.9, 8.1, and 10, respectively. Protein loads were amounts derived from ∼8 µg of original cell extract.
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Materials Protein fractions from ZOOM IEF fractionator (see Basic Protocol 1, steps 18 and 19) Protein extracts from pH partition membranes (see Basic Protcol 1, step 20) Unfractionated protein sample (e.g., cell extract) that was the starting material for the fractionation in Basic Protocol 1 SDS sample buffer (UNIT 10.1) NuPAGE 10% Bis-Tris gels (Invitrogen) Molecular weight markers (Invitrogen) 20× NuPAGE MES SDS Running Buffer (Invitrogen) Novex Mini-Cell electrophoresis device (Invitrogen) Gel-image analysis software (e.g., Discovery Series Quantity One, version 4.2.0) Additional reagents and equipment for 1-D SDS-PAGE (UNIT 10.1) and staining of gels (UNIT 10.5) 1. In separate tubes, mix small volumes of the fractions, the partition membrane extracts, and the initial unfractionated sample with SDS sample buffer to achieve a final concentration of 1× SDS sample buffer. 2. Load the samples on a NuPAGE 10% Bis-Tris Gel. Also load molecular weight markers. The amounts of each fraction or extract loaded into each lane should be proportional to one another for reliable comparisons, i.e., the same percentage of the total volume of each sample should be loaded so that staining intensities directly reflect protein recoveries. For example, if the final volumes of all fractions are 800 ml and all membrane extracts are 400 µl, one might load 10 µl of each fraction and 5 µl of each membrane extract.
3. Run the gel (also see Running buffer.
UNIT 10.1)
using a Novex Mini-Cell and NuPAGE MES SDS
4. Stain the gel (UNIT 10.5) to visualize proteins. The gel can be most easily stained with Coomassie blue. However, Sypro Ruby or silver stain is recommended if higher detection sensitivity is required.
5. Analyze the gel image using computer software (e.g., Discovery Series Quantity One, version 4.2.0). See Anticipated Results for a description of what should be observed. SUPPORT PROTOCOL 3
Microscale Solution Isoelectrofocusing (MicroSol - IEF)
EVALUATION OF THE EFFECTIVENESS OF MicroSol-IEF PREFRACTIONATION USING 2-D GELS Separation of small aliquots of each fraction and membrane extracts on appropriate 3.0 pH unit–wide 2-D gels is an efficient and reliable way to determine whether proteins in a fraction are within the intended pH range and whether sharp protein distribution boundaries are observed near the expected pH. For this purpose either 7-cm or 11-cm IPG gels can be used to form part of a relatively high-throughput 2-D electrophoresis procedure. The 11-cm IPG gels are strongly preferred because they have ∼1.6× greater separation distance than 7-cm IPG strips. Furthermore, medium-size SDS gels (Criterion gel format, Bio-Rad), to which the 11-cm IPG gels are applied, are not significantly more difficult or time-consuming to run compared with multiple mini-gels for the 7-cm IPG strips. Representative 2-D gel results for analysis of the three central fractions from the same MicroSol-IEF experiment evaluated by 1-D gels in Figure 22.6.8 are shown in Figure 22.6.9. These 2-D maps confirmed that MicroSol-IEF effectively separated the proteins with pI values in the range of pH 4.9 to 6.4 into three well-defined fractions, since clear boundaries near the expected pH values were obtained and only minimal amounts of proteins were observed outside the expected pH ranges of fractions. It is important to
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Figure 22.6.9 Further evaluation of the effectiveness of the MicroSol-IEF fractionation using small mid-pH range 2-D gels. Samples and conditions were as shown in Fig. 22.6.8. The samples from three center pH fractions were focused on 11-cm IPG strips, followed by separation using Tris-Tricine SDS gels (Bio-Rad Criterion gel format, ∼11 × 9 cm). The amount of sample applied to IPG strips was derived from ∼40 µg of original cell extract, and the pH values of the IPG strips used is indicated above the images. The expected fraction pH boundaries are indicated by vertical dashed lines. Gels were stained using Sypro Ruby.
check these middle fractions since the majority of proteins in eukaryotic proteomes have pI values within the pH range they encompass.
Materials Protein fractions from ZOOM IEF fractionator (see Basic Protocol 1, steps 18 and 19) IPG buffers (Amersham Biosciences): pH ranges of the buffers are chosen in accordance with the IPG strips Reducing buffer (see recipe) Alkylating buffer (see recipe) IPG strips: mid-pH-range (3.0 pH units wide where possible), either 7 cm (Invitrogen) or 11 cm (from Bio-Rad or Amersham Biosciences) Second-dimension gels (also see UNIT 10.4): 10% SDS-polyacrylamide gels, either mini-size (7 × 7 cm) pre-cast NuPAGE gels (Invitrogen) or medium-size (11 × 9 cm, Bio-Rad Criterion gel format) Gel-image analysis software (e.g., Melanie 2-D Gel Analysis, version 4.0) Additional reagents and equipment for 2-D electrophoresis using immobilized pH gradient gel strips (UNIT 10.4, Basic Protocol 2) and staining of gels (UNIT 10.5) 1. Dilute aliquots from the fractions and partition membrane extracts to be analyzed with MicroSol-IEF sample buffer and add the appropriate IPG buffers (carrier ampholytes), depending on the IPG strips being used. The final volume of the sample to be loaded onto an IPG strip depends on the length of the strip. For the loading of 11-cm strips by in-gel rehydration, ∼240-µl samples should be prepared. Final IPG buffer concentrations must be determined empirically for the fractionated sample in question. For the purposes of human breast cancer cell extracts, ∼0.2% IPG buffer (final) produces the best results for the evaluation of prefractionation. See UNIT 10.4 for further information about 2-D electrophoresis.
2. Load the samples on to the IPG strips by in-gel rehydration (UNIT 10.4). The pH range of the strip should be wider than that of the fraction being analyzed. For example, pH 3 to 6, 4 to 7, and 6 to 9 IPG strips are suitable for checking the fractions obtained using a configuration like that illustrated in Figure 22.6.4A and B, except the most basic fractions, which should be checked on a pH 6 to 11 IPG strip
3. Run IPG-IEF (first dimension; see UNIT 10.4).
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4. Equilibrate the IPG strips in reducing buffer for 10 min at room temperature, then equilibrate in alkylating buffer for 10 min at room temperature. Transfer them to the second-dimension gels (also see UNIT 10.4). 5. Stain the gels using appropriate staining reagents (e.g., Coomassie blue, Sypro Ruby, or silver stain). 6. Analyze the gel images using appropriate software (e.g., Melanie 2-D Gel Analysis, version 4.0). See Anticipated Results for a description of what should be observed. BASIC PROTOCOL 2
DETECTION OF FRACTIONATED PROTEINS USING HIGH-RESOLUTION, NARROW-pH-RANGE 2-D GELS Prefractionation of proteomes using the ZOOM IEF Fractionator seamlessly interfaces with the use of narrow-pH-range IPG gels, since the acrylamide/Immobiline partitions for ZOOM IEF are made with the same pH precision that is inherent in IPG technology (Righetti, 1983), and the same buffer with denaturing and reducing properties is used for both MicroSol-IEF and IPG-IEF systems. Furthermore, separating a complex protein sample into fractions with pH ranges that match IPG strips is especially advantageous for the isoelectric focusing of the proteins on the gel. This is because the initial MicroSol-IEF procedure effectively eliminates the interfering proteins with pI values outside the range of the IPG strip and minimizes “nonideal” behavior (e.g., precipitation, aggregation, or protein-protein interactions) during IPG-IEF. The reduction in aberrant isoelectric focusing allows more reliable quantitative comparisons of proteomes. To maximize spot separation and resolution of fractionated proteins on narrow-pH-range 2-D gels, separation distances over the pH range of each fraction should be maximized. Ideally, a series of overlapping IPG strips, each with a slightly wider pH range (± 0.1 pH unit) than the corresponding MicroSol-IEF fraction, should be used to maximize separation distances. Matching 1.0 pH unit–wide IPG strips for the three center fractions (each 0.8 pH units wide) obtained using the six commercially available partition membranes (see Fig. 22.6.4A and Table 22.6.1) are available for minigels, i.e., 7-cm IPG strips (Invitrogen). However, the separation distance obtained on these small gels is similar to that obtained by running an unfractionated sample on a single larger pH 4 to 7 IPG strip (18 or 24 cm), thus largely negating the advantages of using fractionated samples and narrow-pH-range 2-D gels. Although longer custom IPG strips could be prepared in individual laboratories, it is time-consuming and quite labor-intensive to make them. More importantly, using the custom-made IPG gels will reduce the reproducibility of 2-D separation between different laboratories.
Microscale Solution Isoelectrofocusing (MicroSol - IEF)
A practical strategy for interfacing the seven fractions obtained from the MicroSol-IEF by using custom pH partition membranes (see Fig. 22.6.4B) with large commercially available IPG strips, to maximize both total separation distance and throughput, is shown in Table 22.6.4. The pH ranges of these seven fractions have been selected in part so that the MicroSol-IEF fractions can be separated on readily available 24-cm narrow-pH-range gels. After IEF, regions of the IPG strip that are more than 0.1 pH unit outside the pH range of the MicroSol-IEF fraction are cut off to reduce the IPG strip size to 18 cm (Fig. 22.6.10). The second-dimension gels can then be run on 18-cm wide gels. This strategy produces a total effective IEF separation distance of about 86 cm over the range of pH 3 to 10 (see Table 22.6.4) with improved spot separation and resolution compared to that obtained using conventional 18-cm IPG strips and with no disadvantages when compared to running 24-cm IPG strips and larger second-dimension gels (see Fig. 22.6.10). Since the time required and difficulty of running second-dimension gels increase as the gel size increases, this down-sizing of the second-dimension gel improves throughput without
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Table 22.6.4 Strategy for Interfacing MicroSol-IEF Fractions with Large Commercially Available IPG Strips for Optimal Separation of Proteins
Fractions
F1
F2
F3
F4
F5
F6
F7
MicroSol fractions
3.0-4.4
4.4-4.9
4.9-5.4
5.4-5.9
5.9-6.4
6.4-8.1
8.1-10
Commercial 24-cm IPGs
3.0-6.0
4.0-5.0
4.5-5.5
5.0-6.0
5.5-6.7
6.0-9.0
7.0-10
pH range (trimmed to 18 cm)
3.0-5.25
4.25-5.0
4.75-5.5
5.25-6.0
5.7-6.6
6.2-8.45
7.75-10
Effective IEF separation (cm per pH unit)
8
24
24
24
20
8
8
Effective separation distance
Total ∼86 cm for the 7 fractions
sacrificing resolution. In addition, smaller second-dimension gels provide cost savings in terms of gel reagents and staining. When this strategy is used with commercial 24-cm IPG strips to analyze prefractionated samples (see Table 22.6.4), protein loading capacity and separation quality are much improved compared with what is obtained by separating unfractionated samples on either a single broad pH range 2-D gel or a series of narrow pH range 2-D gels. In addition, partition membrane extracts from a MicroSol-IEF run can be combined with appropriate adjacent soluble fractions for 2-D gel separation to minimize the sample losses from membrane trapping.
Materials Protein fractions from ZOOM IEF fractionator (see Basic Protocol 1, steps 18 and 19) MicroSol-IEF sample buffer (see recipe) IPG buffers (Amersham Biosciences): pH ranges chosen in accordance with the IPG strips used Reducing buffer (see recipe) Alkylating buffer (see recipe) Commercial 24-cm IPG strips: pH 3 to 6 and pH 7 and 10 strips (Bio-Rad) or pH 4.0 to 5.0, 4.5 to 5.5, 5.0 to 6.0, 5.5 to 6.7, and 6 to 9 strips (Amersham Biosciences) Second-dimension gels (also see UNIT 10.4): large-size (18 × 19 cm) 10% SDS-polyacrylamide gels, custom-made using the Bio-Rad Protean II format Gel-image analysis software (e.g., Melanie 2-D Gel Analysis, version 4.0) Additional reagents and equipment for 2-D electrophoresis using immobilized pH gradient gel strips (UNIT 10.4) and staining of gels (UNIT 10.5) 1. Dilute aliquots from the fractions to be analyzed with MicroSol-IEF sample buffer (components listed in Basic Protocol 1) and add the appropriate IPG buffers (carrier ampholytes), depending on the IPG strips being used. For the loading of 24-cm strips by in-gel rehydration, prepare samples with a final volume of ∼470 µl. Final IPG buffer concentrations must be determined empirically for each sample. For human breast cancer cell extracts, a mixture of narrow- and broad-range IPG buffers at relatively high concentrations (∼0.5% to 0.8% final) produces the best results for detection of fractionated proteins. See UNIT 10.4 for further information about 2-DE.
2. Load samples on to the IPG strips by in-gel rehydration (UNIT 10.4). 3. Run IPG-IEF (first dimension; see UNIT 10.4).
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Figure 22.6.10 Effect of IPG gel length and second-dimension gel size on resolution and sensitivity when analyzing 0.5 pH unit MicroSol-IEF fractions. The fractionated proteins (pH 4.7 to 5.2) were obtained from prefractionation of a human breast cancer MCF-7/6 cell extract and separated on commercial 24- or 18-cm IPG strips (pH 4.5 to 5.5). After IEF, different-sized second-dimension gels were evaluated. IPG strips were either used in directly matching second dimension gels (A and C) or the excess IPG gel regions from a 24-cm IPG strip (where the pH is outside the fraction’s pH range) were trimmed so that the IPG gel (∼0.7 pH units) could be applied to an 18-cm-wide second-dimension gel (B). All seconddimension separations were performed on 10% Tris-Tricine SDS gels, and proteins were visualized using silver staining. Protein loads were adjusted for differences in 2-D gel volumes and resulting spot sizes so that similar staining intensities were obtained. Fractionated sample, derived from the following amounts of original cell extract, was used (left to right): 150 µg, 100 µg, and 100 µg. As shown in gel images, less sample was loaded on the trimmed 18-cm IPG gel (B) than on the 24-cm IPG gel (A), with similar sensitivity, but the trimmed 18-cm IPG gel did not have negative effects on detection and resolution. However, compared with the same proteins that were directly loaded on the commercial 18-cm IPG strip (C), the trimmed 18-cm IPG gel (B) had a higher number of spots detected and higher resolution.
4. After IEF, trim (either from one end or both ends) the 24-cm strips to 18 cm (see Table 22.6.4). 5. Equilibrate the IPG strips in reducing buffer for 10 min at room temperature, then equilibrate in alkylating buffer for 10 min at room temperature. Transfer them to the second-dimension gels (also see UNIT 10.4). Microscale Solution Isoelectrofocusing (MicroSol - IEF)
6. Stain the gels using appropriate staining reagents (e.g., Coomassie blue, Sypro Ruby, or silver stain). 7. Analyze the gel images using appropriate software (e.g., Melanie 2-D Gel Analysis, version 4.0).
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Figure 22.6.11 Comparison of large proteins in closely related human breast cancer cells with low and high metastatic potential using MicroSol-IEF and large pore 1-D SDS gels. The >100 kDa regions from a silver-stained 3% to 8% Tris-Acetate NuPAGE minigel shows the comparison of MCF-7/AZ cells (low metastatic potential, indicated by –) with MCF-7/6 cells (high metastatic potential, indicated by +). Lane definitions: ext, cell extracts without prefractionation (4 µg); F1 to F7, MicroSol-IEF fractions 1 to 7, respectively (see Fig. 22.6.4B for schematic), derived from ∼15 µg of the original cell extracts. Protein bands were quantitatively analyzed and compared using the Bio-Rad Discovery Series Quantity One (version 4.2.0) software. Triangles highlight proteins with greater than two-fold increase in density compared with the alternative cell line.
DETECTION OF LARGE FRACTIONATED PROTEINS USING HIGH RESOLUTION, LARGE-PORE 1-D GELS
SUPPORT PROTOCOL 4
Large proteins (>100 kDa) present insurmountable problems for 2-D gels because they typically focus poorly and inconsistently as a result of poor solubility near their pI values. In addition, their large size restricts their diffusion into IPG gels, and they may be recovered from IPG gels with poor efficiency. Hence, large-pore 1-D SDS gels are recommended for routine detection of large proteins (>100 kDa) after a MicroSol-IEF prefractionation, to complement data obtained from the 2-D gel analysis described in Basic Protocol 2. Typically, 1-D SDS gels with a gradient acrylamide concentration capable of resolving proteins ≥500 kDa are suitable for detecting large proteins (>100 kDa) in the fractions. One major advantage is that the complexity of individual bands is substantially reduced by prefractionation of the sample, which facilitates quantitative comparisons of the large proteins (100 to 500 kDa) in complex proteomes (Zuo et al., 2002). A comparison of prefractionated large proteins (>100 kDa) from closely related human breast cancer cell lines with low and high metastatic potentials is shown in Figure 22.6.11. Few differences were observed between the two cell lines when the unfractionated cell extracts were compared, due to the great sample complexity which led to the presence of multiple proteins in individual bands. In contrast, many quantitative differences were readily detected in the fractionated samples.
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Materials Protein fractions from ZOOM IEF fractionator (see Basic Protocol 1, steps 18 and 19) Unfractionated protein sample (e.g., cell extract) that was the starting material for the fractionation in Basic Protocol 1 SDS sample buffer (UNIT 10.1) NuPAGE 3% to 8% Tris-Acetate gels (Invitrogen). 20× Novex Tris-Acetate SDS Running Buffer (Invitrogen) Novex Mini-Cell electrophoresis device (Invitrogen) Gel-image analysis software (e.g., Discovery Series Quantity One, version 4.2.0) Additional reagents and equipment for1-D SDS-PAGE (UNIT 10.1) and staining of gels (UNIT 10.5) 1. In separate tubes, mix small volumes of the fractions and the initial unfractionated sample with SDS sample buffer to achieve a final concentration of 1× SDS sample buffer (see Basic Protocol 1 in UNIT 10.1). Proportional amounts of protein from each fraction and unfractionated sample should be loaded on the gel for reliable comparisons, i.e., the same percentage of the total sample volume.
2. Load the protein samples on to NuPAGE 3% to 8% Tris-Acetate Gels. Also load molecular weight markers. 3. Run the gel (also see UNIT 10.1) using a Novex Mini-Cell and Novex Tris-Acetate SDS Running Buffer. 4. Stain the gels using Coomassie blue, Sypro Ruby, or silver stains (UNIT 10.5). 5. Analyze the gel image using appropriate software (e.g., Discovery Series Quantity One, version 4.2.0). SUPPORT PROTOCOL 5
COMBINATION OF ZOOM IEF WITH TWO-DIMENSIONAL DIFFERENTIAL GEL ELECTROPHORESIS (2-D DIGE) FOR QUANTITATIVE COMPARISON OF PROTEINS Two-dimensional differential gel electrophoresis (2-D DIGE) is a relatively new approach to quantitative proteome analysis using 2-D gels. This method involves separately labeling two or three different protein extracts with distinct fluorophores (cyanine dyes) by covalently modifying lysine residues in the proteins (Unlu et al., 1997). The ability to combine two or three labeled samples on a single 2-D gel eliminates problems associated with gel-to-gel variation.
Microscale Solution Isoelectrofocusing (MicroSol - IEF)
As described above, MicroSol-IEF prefractionation can greatly enhance detection capacities of 1-D and 2-D gels. In fact, the MicroSol-IEF method, with its smaller sample volume requirements, is particularly well suited for use with 2-D DIGE. In addition, all the approaches using subsequent slightly overlapping narrow-range 2-D gels and largepore 1-D (Support Protocol 4) gel methods described in the above protocols are readily compatible with 2-D DIGE (see Fig. 22.6.1A). When 2-D DIGE is used, any subtle variations in MicroSol-IEF and on the 1-D and 2-D gels are no longer a problem, because the two or three labeled samples are used together in a mixture at each separation step. Combination of MicroSol-IEF with 2-D DIGE also significantly reduces the gel-related workload, since the number of narrow-pH-range 2-D gels and large-pore 1-D gels that are required for analyses of the fractions will be greatly reduced. However, the major disadvantages are those inherent to 2-D DIGE, i.e., the method is often less sensitive
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than silver staining, and it can be difficult to match observed fluorescent changes with the corresponding unlabeled proteins for identification by mass spectrometry.
Materials Protein samples Additional materials and equipment for labeling proteins as for 2-D DIGE (UNIT 22.2, Basic Protocol), protein fractionation using the ZOOM IEF Fractionator (see Basic Protocol 1 or Alternate Protocol 1 in this unit), and detection of fractionated proteins using high-resolution, narrow-pH-range 2-D gels (see Basic Protocol 2 in this unit) or high-resolution, large-pore 1-D gels (see Support Protocol 4 in this unit) 1. Label two protein samples individually with different fluorescent dyes as for 2-D DIGE (e.g., Cy3 and Cy5; see UNIT 22.2, Basic Protocol). 2. Combine the two labeled samples and prefractionate the mixture using the ZOOM IEF Fractionator (see Basic Protocol 1 or Alternate Protocol 1). 3. Run the fractionated samples on overlapping narrow pH range 2-D gels for small (<100 kDa) proteins (see Basic Protocol 2) or large-pore 1-D gels for large (>100 kDa) proteins (see Supporting Protocol 4). 4. Capture and analyze gel images (see UNIT 22.2, Basic Protocol).
ANALYSIS OF FRACTIONATED PROTEINS USING LC-MS/MS METHODS Mass spectrometry (MS) analysis of tryptic peptides is an important non-gel approach that usually uses one or two liquid chromatographic separations prior to MS and MS/MS analysis (LC-MS or LC/LC-MS/MS) to reduce sample complexity. Using isotope-coded affinity tags (ICAT) to label proteins prior to MS analysis makes quantitative comparison of two samples possible (Gygi et al., 1999; 2002).
ALTERNATE PROTOCOL 2
Although ICAT and other MS-based methods alone can detect lower abundance proteins than 2-D gels, they are not capable of reliably detecting more than about 1000 to 2000 protein species, and throughput is similar to that obtained with 2-D gels. Fortunately, these non-gel-based methods can benefit from prior prefractionation of complex protein samples. A strategy combining ICAT, MicroSol-IEF, and LC-MS/MS methods to improve protein profiling and quantitative comparison is illustrated in Figure 22.6.1B. Reducing sample complexity by MicroSol-IEF prefractionation should allow detection and reliable quantitation of more proteins than the typical LC-MS/MS methods when they are employed without sample prefractionation.
Materials Protein samples ICAT reagents (Applied Biosystems) 10% SDS polyacrylamide gel (UNIT 10.1) Additional reagents and equipment for ICAT (UNIT 23.2, Basic Protocol 1), protein fractionation using the ZOOM IEF Fractionator (see Basic Protocol 1 in this unit), SDS-PAGE (UNIT 10.1), and LC-MS/MS (UNIT 23.1) 1. Label two protein samples individually with heavy and light forms of ICAT reagents, respectively (see UNIT 23.2, Basic Protocol 1). 2. Combine the isotope-labeled samples and perform MicroSol-IEF to fractionate proteins into five fractions (see Basic Protocol 1).
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3. Load an aliquot from each fraction into a separate lane of a 10% SDS-polyacrylamide gel and electrophorese for a short distance (∼5 mm). Running the fractionated samples into a short SDS gel is a convenient method to remove the reagents in the samples that will otherwise interfere with trypsin digestion and MS/MS analysis.
4. Digest the proteins in the gel (one digest per fraction) with trypsin (see UNIT 11.3). 5. Fractionate the digested sample (peptide mixture) using a strong-cation exchange (SCX) capillary column and analyze using LC-MS/MS (see Basic Protocol 1 in UNIT 23.1).
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Acrylamide/bis stock, 30%T/10%C Place 27.0 g acrylamide and 3.0 g bisacrylamide (“bis”) in a 150-ml beaker. Add ∼70 ml Milli-Q water. Mix with magnetic stir bar at room temperature until completely dissolved (∼20 to 30 min). Add extra Milli-Q water to a final volume of 100 ml. Filter the solution through Whatman filter paper or a 0.22-µm filter membrane to remove undissolved particles. Store at 4◦ C and use within 1 month. CAUTION: Acrylamide is highly toxic; see APPENDIX 2A for safety guidelines.
Alkylating buffer 50 mM Tris·Cl, pH 8.0 (APPENDIX 2E) 6 M urea 2% (w/v) SDS 30% (v/v) glycerol 2.5% (w/v) iodoacetamide MicroSol-IEF anode buffer, 1× Prepare 20 ml on day of use by adding the following to a 25-ml beaker: 2 ml 10× IEF anode buffer (Bio-Rad) 8 ml Milli-Q H2 O 9.6 g urea 3.04 g thiourea Mix with magnetic stir bar at room temperature until completely dissolved (∼30 to 40 min). Adjust volume to 20 ml using Milli-Q water. Check pH and adjust to 2.9 to 2.95 by adding 85% phosphoric acid (∼100 µl). Final concentrations are ∼7 mM phosphoric acid, 8 M urea, and 2 M thiourea. The initial pH will be higher than 3.0 (∼3.65 to 3.82) due to the presence of urea and thiourea.
Microscale Solution Isoelectrofocusing (MicroSol - IEF)
MicroSol-IEF cathode buffer, 1× Prepare 20 ml on day of use by adding the following to a 25-ml beaker: 2 ml 10× IEF cathode buffer (Bio-Rad) 8 ml Milli-Q water 9.6 g urea 3.04 thiourea 40 mg DTT Mix with magnetic stir bar at room temperature until completely dissolved (∼30 to 40 min). Adjust volume to 20 ml using Milli-Q water. Measure pH (should be 10.10 to 10.40; do not adjust). Final concentrations are 20 mM lysine, 20 mM arginine, 8 M urea, 2 M thiourea, and 0.2% DTT).
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MicroSol-IEF sample buffer 8 M urea 2 M thiourea 4% (w/v) CHAPS 1% (w/v) DTT 0.2% to 0.5% (v/v) IPG buffer (pH 3 to 10; Amersham Biosciences) Store in 1-ml aliquots at −20◦ C and use within 3 months.
Partition membrane storage solution, pH 6.8: 10 mM sodium phosphate buffer, pH 6.8 (APPENDIX 2E) 12% (v/v) glycerol 0.4% (w/v) acrylamide 2 mM sodium azide (NaN3 ) Store at 4◦ C and use within 1 month. CAUTION: Acrylamide is highly toxic; see APPENDIX 2A for safety guidelines.
Reducing buffer 50 mM Tris·Cl, pH 8.0 (APPENDIX 2E) 6 M urea 2% (w/v) SDS 30% (v/v) glycerol 1% (w/v) DTT COMMENTARY Background Information Currently existing methods for quantitatively comparing protein profiles, including two-dimensional polyacrylamide gel electrophoresis (2-D PAGE) and non-gel methods such as isotope coded affinity tag methods (ICAT) and other LC/LC-MS/MS approaches, have limited resolution and modest dynamic ranges for detection (UNIT 22.1; Gygi et al., 1999; Quadroni and James 1999; Ranish et al., 2003). As a result, these methods can only detect small portions of complex proteomes such as eukaryotic cell extracts. One strategy to increase the proportion of complex proteomes that can be quantitatively compared by existing protein profiling methods is to fractionate these samples into a number of well resolved pools prior to downstream analysis using 2-D gels or non-gel-based methods (Righetti et al., 2001; Zuo et al., 2004). Preparative solution-phase IEF separation was reported as one of the most promising approaches for fractionating complex proteomes to enhance protein profiling (Weber and Bocek, 1998; Wall et al., 2000; Hoffmann, et al., 2001; Righetti et al., 2003). In general, solution IEF prefractionation approaches use one of two separation principles, these being: (1) “free solution IEF” where soluble carrier ampholytes are used to create and maintain
pH gradients across a single focusing chamber (e.g., the Rotofor marketed by Bio-Rad); and (2) “multicompartment solution IEF,” where pH-selective partitions with protein-scale pores divide a series of tandem chambers containing the sample (e.g., the IsoPrime marketed by Amersham Biosciences). However, all the older preparative solution IEF devices have at least one or more major drawbacks when applied to sample prefractionation for comprehensive proteome analysis, these being: (1) they have relatively large separation chambers that require loading a high initial sample volume; (2) they produce large-volume, dilute fractions that must be concentrated prior to downstream analysis methods; (3) they exhibit poor resolution resulting in fractions with substantially overlapping proteins; (4) they use carrier amphoytes for generating the pH gradient, which usually results in a pH range devoid of very acidic and very basic pH values; and (5) they involve complicated procedures and require expensive instrumentation. The Micro-Scale Solution IEF (MicroSolIEF) method developed in the authors’ laboratory is convenient, relatively simple, economical, and directly compatible with sample sizes commonly used for proteome experiments (Zuo and Speicher, 2000). It utilizes the same “multi-compartment solution IEF”
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Microscale Solution Isoelectrofocusing (MicroSol - IEF)
principle originally described by Righetti et al. (1989, 1990) and described above. However, unlike other preparative solution IEF methods and devices, it is capable of efficiently separating microgram to milligram quantities of complex protein mixtures (Zuo and Speicher, 2000, Zuo et al., 2002, 2004), making it highly compatible with 2-D gels for further analyses. Complex proteomes are fractionated based upon protein isoelectric points (pI) by utilizing protein-permeable acrylamide partitions that contain covalently-bound Immobilines at different pHs between small tandem separation chambers. After isoelectric focusing, each separation chamber will contain proteins having pI values between the pH values defined by the two boundary membranes. Prefractionation of complex proteomes such as mammalian cell extracts (Zuo et al., 2002) and serum (Zuo et al., 2001; Zuo and Speicher, 2002) produced well resolved pH pools that could be directly applied to IPG-based 2-D gels or 1-D SDS gels without concentration. The ZOOM IEF Fractionator (Invitrogen) is a commercial device for performing MicroSol-IEF prefractionation that is more user-friendly than earlier homemade devices. The current version of the ZOOM IEF device can be set up in a flexible manner to produce various numbers of fractions, pH ranges, and fraction volumes. Many gel-based and non-gel-based methods interface with the MicroSol-IEF fractions for further analyses, but slightly overlapping narrow-pH-range 2-D gels (for <100 kDa) and large-pore 1-D gels (for >100 kDa) are particularly useful for quantitative protein profiling (Zuo et al., 2004). Compared with direct analysis of unfractionated samples on a series of narrow-pH-range 2-D gels, prefractionation of complex proteomes using MicroSol-IEF has many advantages (Zuo and Speicher, 2000, 2002, Zuo et al., 2002), including: (1) higher resolution on narrow-pH-range gels with fewer artifacts; (2) the ability to use higher sample loads, which increases dynamic range and detection of lower-abundance proteins; (3) the ability to detect far more total proteins; and (4) conservation of proteome samples. In addition, MicroSol-IEF prefractionation is compatible with most downstream non-gel protein profiling methods including LC-MS/MS and LC/LC-MS/MS (Speicher et al., 2004). Recently, additional partition membranes have become available from Invitrogen, including pH 12.0 membranes. This now extends fractionation over the pH range of 3.0 to 12.0.
Critical Parameters Protein sample and sample buffer As with 2-D gels, the protein samples for MicroSol-IEF should be initially prepared and/or significantly diluted using the MicroSol-IEF solubilization buffer (sample buffer), which has denaturing and reducing effects, prior to prefractionation. Typically, critical components in the MicroSol-IEF sample buffer are denaturants, reducing agents, and nonionic detergents, which are similarly required for running IPG-based 2-D gels (Rabilloud, 1999). Actually, use of the same sample buffer for both MicroSol-IEF and IPG-IEF separations is especially advantageous for interfacing the two systems when the fractionated samples are subjected to further analysis on 2-D gels, because the samples can be directly applied to the first-dimension IPG gels without any sample treatment. In addition, the reagents in the buffer that promote sample solubility and effective focusing in one IEF system should produce similar beneficial effects in the other. Specifically, the combination of urea and thiourea is superior to the use of urea alone for solublizing proteins (Rabilloud et al., 1997; Rabilloud, 1998). Similarly, DTT is the most commonly used reducing agent for IEF, despite its mild negative charge, which might cause it to migrate out of high-pH separation chambers during MicroSol-IEF. Addition of a small amount of DTT (e.g., ∼0.2%) to the MicroSol-IEF cathode buffer could minimize this depletion, although it does not appear to be a major concern if the prefractionation is completed in a few hours. Although it has been suggested that tributyl phosphine (TBP) is superior to DTT because it has no electrical charge and is a more powerful reducing agent (Herbert et al., 1998), the authors of this unit found that TBP is highly unstable and rapidly degraded during both MicroSol-IEF and IPG-IEF. Therefore, the authors strongly prefer not to use TBP in any IEF system. The presence of soluble ampholytes spanning the full pH range (e.g., IPG buffer pH 3 to 10, Amersham Biosciences; or pH 3 to 12, Invitrogen) is necessary for optimal separation of proteins in both IEF systems. However, the actual minimum amount of ampholytes needed for optimal MicroSol-IEF appears to vary somewhat with type of sample, protein load, and pH ranges of the fractions. The authors suggest that 0.2% to 0.5% IPG buffer be included in the MicroSol-IEF sample buffer as a starting point. Although isoelectric focusing of proteins using a ZOOM IEF Fractionator results in
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a desalting effect, high ionic strength should be avoided in initial protein samples that are subjected to the MicroSol-IEF prefractionation because of several negative consequences. First, high salt concentrations in the ZOOM IEF device will produce high conductivity and, since current is typically limited to 1 mA, voltages will remain low for extended periods of time at higher conductivities, thus greatly increasing total IEF time. Second, high ionic strength within separation chambers can result in high electroosmosis, reducing the effectiveness of focusing. In general, the final concentration of total salts in the sample should be <10 mM when introduced into a ZOOM IEF Fractionator. Ionic detergents such as SDS should be avoided or kept to a minimal final concentration (<0.1%). Ideally, protein samples such as cell extracts should be prepared in low-ionic-strength buffer and be further diluted with the MicroSol-IEF sample buffer prior to sample loading. In addition, the concentration of carrier ampholytes will affect the initial conductivity, total focusing time, and the minimum current that is reached at the final 1000 V focusing step. The negative effects of high ampholyte concentration may be counterbalanced by the beneficial effects that higher ampholyte concentrations may bring in terms of protein solubility at high protein loads. In general, the final concentration of ampholytes in the sample buffer should be <0.5%, but higher concentrations may be necessary for very complex samples. pH partition membranes It is critical to prepare acrylamide/ Immobiline membranes with reproducible pH values and buffering capacities to ensure reproducible separations. Although theoretically only two Immobilines are required to define any pH, in practice, more than two Immobilines are needed to ensure more uniform buffering capacity (Giaffreda et al., 1993). Also, it is suggested that a total Immobiline concentration between 10 and 20 mM be used in the acrylamide gel, because the authors have found that higher concentrations (>20 mM) usually cause gels to swell during washing and use. In addition, where large amounts of Immobiline are present in the acrylamide gel (e.g., >40 mM final concentration), the efficiency of Immobiline incorporation begins to decrease. Such inconsistent incorporation of different Immobilines into a covalent acrylamide matrix will cause pH errors (Giaffreda et al., 1993). In some experiments, the authors have observed minor deviations for the actual pH
ranges of MicroSol-IEF fractions compared with the expected pH ranges when the proteins were separated on very narrow-pH-range (∼1.0- to 1.2-unit) IPG gels (Amersham Biosciences). Most MicroSol-IEF partition membrane pH values showed fairly consistent shifts of ∼0.1 to 0.2 pH units below the expected pHs on the IPG gels. Possible reasons for this minor pH deviation are as follows. First, the MicroSol-IEF partition membrane pH values are determined in the absence of denaturing agents, while the IPG strip pH values are defined in the presence of a high amount of urea and thiourea. Second, different Immobilines may be incorporated into the acrylamide gel matrix with varying efficiencies, which can skew the actual pH. When pH deviations are observed between MicroSol-IEF pH partition membranes and IPG gels, it is impossible to determine which pH is more accurate, since the pH values of these gels cannot be measured directly even with an advanced surface electrode (Giaffreda et al., 1993). However, in practice, the minor pH differences observed are not a major concern, since the MicroSol-IEF partition membrane pHs appear to be highly reproducible between batches when the same procedure is followed, and the reproducibility of MicroSol-IEF prefractionation is more important than absolute accuracy of pH values for individual fractions. Although various concentrations of acrylamide gels for covalently binding Immobilines can be used for the pH partition membranes, gels should ideally be prepared using low total acrylamide (%T) and high crosslinker (%C) concentrations to create large pore sizes that can facilitate rapid transfer of all proteins at high protein loads. On the other hand, the gels should be strong enough to remain inside the pores of the polyethylene disks during electrophoresis. The authors of this unit suggest 4%T/10%C acrylamide/bis gels. In early experiments, as with the IsoPrime device (Amersham Biosciences), the authors used glass-fiber filters (∼1.5-mm thickness) as supports for the partition membranes (Zuo and Speicher, 2000; Zuo et al., 2001), but subsequently found that hydrophilic polyethylene (∼0.67 mm thickness) was superior to the glass fibers (Zuo and Speicher, 2002; Zuo et al., 2002) as they are stronger and thinner. This also minimizes partition gel volumes to facilitate migration of proteins. The large pores of acrylamide gels cast on the thin support disks should allow effective separation of large proteins and minimize their precipitation on the membrane surfaces.
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Microscale Solution Isoelectrofocusing (MicroSol - IEF)
Sample loading methods Since proteins are separated based on their pI values using the ZOOM IEF Fractionator, there is substantial flexibility in how the protein sample can be loaded into the device for prefractionation. For example, in the case of the seven-fraction device illustrated in Fig. 22.6.4B, the initial sample can be loaded into any single chamber, several chambers, or all of the chambers. The advantages of different loading methods are sample dependent. For example, it may be advantageous to load an entire serum sample into the chamber whose pH range encompasses the pI of albumin, the most abundant protein constituting >50% of total serum protein, because this loading method could facilitate migration of other proteins through partition membranes and reduce the potential for protein precipitation in the partitions. However, it would be a disadvantage to load an entire cell extract into a single chamber, because this loading increases the local initial protein concentration compared with more distributed loading strategies, and this higher protein concentration could lead to protein precipitation and aggregation early in the focusing step. Also, if the sample is too dilute to fit into one or several separation chambers, but would fit into all separation chambers, then it is usually preferable to load the sample into all chambers rather than risk sample losses during a concentration step. Some proteins show poor focusing in terminal chambers when the sample is loaded into all separation chambers. This may be because ampholytes migrate more rapidly than proteins, and the pH values in the terminal chambers may reach pH extremes before proteins that are unstable at extreme pHs have sufficient time to migrate out of these terminal chambers, which may result in precipitation and aggregation of the proteins. Hence, a general recommendation is to load cell or tissue extracts into central chambers and avoid loading proteins into the most acidic and most basic chamber if the initial sample volume allows. The maximum sample loading capacity appears to depend on the type of sample, loading position, and total number of separation chambers. In general, higher loads can be used for clean samples that contain primarily proteins, compared with extracts that contain large amounts of lipids and nucleic acids. In the authors’ experience, for complex samples such as mammalian cell or tissue extracts, up to 3 mg of protein is suitable for a ZOOM IEF device with seven separation chambers.
Higher loads are possible for less complex proteomes such as prokaryotic cell extracts, cytoplasmic fractions, and biological fluids. However, overloaded samples will result in protein deposition on partition-membrane surfaces and incomplete separation. Sample loading capacity may also be limited if the pI of an abundant protein exactly matches the pH of any partition membrane. These proteins may block the pores owing to their deposition on the membrane surfaces or may overwhelm the buffering capacity of the Immobilines in the membrane. The load capacity discussed above matches well with the maximum feasible loads for subsequent narrow pH range 2-D gels (Zuo et al., 2001, 2002; Zuo and Speicher, 2002).
Troubleshooting Problems associated with MicroSol-IEF that might occur and suggestions on how to resolve them are listed in Table 22.6.5.
Anticipated Results MicroSol-IEF prefractionation After a protein sample is separated by a ZOOM IEF Fractionator, the fractions should be recovered in similar volumes to those loaded. A small deviation is acceptable (e.g., if 650 µl of sample were initially loaded into each chamber, it is normal to recover ∼550 to 680 µl). However, if <50% of original volumes is recovered in any fraction after MicroSolIEF, the experiment should be discarded, since this indicates that one or more chambers were not adequately sealed or that excessive electroosmosis occurred, which usually results in poor separation. Checking MicroSol-IEF fractions on 1-D gels After electrophoresing all fractions and extracts from the partition membranes, the following should be observed on the stained gels (see Fig. 22.6.8). 1. All the proteins observed in the initial unfractionated sample should be present in and distributed throughout the fractions. 2. Central pH fractions should contain more proteins than the terminal pH fractions and overall protein band patterns should be different for different fractions. 3. The majority of proteins should be in fractions, though some can be found in the extracts from partition membranes due largely to the proteins with pI values equal to the pH values of partitions.
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Table 22.6.5 Troubleshooting Preparation of Partition Membranes and Performing MicroSol-IEF Prefractionation
Problem
Causes
Solution
Difficulty in polymerizaton of acrylamide/Immobiline gels; gels become swollen during washing process
Excesive Immobiline concentrations in the gels
Make final concentration of Immobilines in gel at <20 mM
Gel solution pH is not close to neutral prior to polymerization
Adjust the gel solution pH to ∼6.5 prior to gel polymerization
Power supply has a low current shut-off feature
Use a power supply capable of operating at low current (<0.1 mA) and at high voltage (≥1000 V) with no low current shut-off feature
Air bubbles formed in chambers
Avoid trapping any air bubbles in the chambers when loading sample or buffers. Break up bubbles with a gel-loading tip if some form.
Very high electro-osmosis occurred during MicroSol IEF
Ensure urea and thiourea concentrations in the electrode buffers are the same as in the sample buffer
No current for isoelectric focusing
Uneven volumes recovered from chambers (too high and/or too low volumes after prefractionation)
Ensure total salt concentration is <10 mM and final concentration of IPG buffer is <0.5% in the sample prior to loading for prefractionation After loading a suitable volume (∼650 to 675 µl), ensure that a small gap of air remains in each chamber and tightly seal the chamber ports with the plugs Do not overtighten end cap, as excess pressure distorts the Teflon chambers and keeps chamber plugs from sealing properly. Check O-rings on chamber plugs for possible damage Fractions are not well defined pH pools after prefractionation
Liquid leaking between chambers
Use partition membranes with final concentration of acrylamide at not less than 4%T, cast on polyethylene disks
Incorrect pH or pH changed in the partition membranes used
Ensure assembly of the pH partition membranes in the right order and use them before expiration dates
High salt in sample or protein overloaded
Prepare initial protein sample in low ionic strength buffer; load total protein amount within the device’s capacity
IEF terminated before components focused to equilibrium
Do not terminate IEF until final voltage (e.g., 1000 V) is achieved and the lowest stable current (e.g., <0.5 mA) remains unchanged for at least 30 min
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Checking MicroSol-IEF fractions on 2-D gels The following should be observed on the stained 2-D gels (see Fig. 22.6.9). 1. Clear pH boundaries found at, or close to, the expected pH values on the 2-D gels. 2. The majority of protein spots detected within the pH boundaries of the fraction and only minimal spots overlapping between fractions. 3. More spots detected in the central pH fractions than the terminal pH fractions.
Time Considerations Preparation of custom pH partition membranes requires ∼2 to 3 days for eight specific pHs (24 membranes for each pH). MicroSolIEF prefractionation can be performed in 1 day. Evaluation of MicroSol-IEF effectiveness using 1-D mini SDS gels takes 1 to 2 days for 5 to 7 fractions, while evaluation of MicroSol-IEF effectiveness using small or medium size 2-D gels requires 3 to 5 days for 5 to 7 fractions.
Rabilloud, T. 1999. Solubilization of proteins in 2-D electrophoresis: An outline. Methods Mol. Biol. 112:9-19. Rabilloud, T., Adessi, C., Giraudel, A., and Lunardi, J. 1997. Improvement of the solubilization of proteins in two-dimensional electrophoresis with immobilized pH gradients. Electrophoresis 18:307-316. Ranish, J.A., Yi, E.C., Leslie, D.M., Purvine, S.O., Goodlett, D.R., Eng, J., and Aebersold, R. 2003. The study of macromolecular complexes by quantitative proteomics. Nat. Genet. 33:349355. Righetti, P.G. 1983. Isoelectric focusing: Theory, Methodology and Application. Elsevier, Amsterdam. Righetti, P.G., Wenisch, E., and Faupel, M. 1989. Preparative protein purification in a multi-compartment electrolyser with immobiline membranes. J. Chromatogr. 475:293-309.
Literature Cited
Righetti, P.G., Wenisch, E., Jungbauer, A., Katinger, H., and Faupel, M. 1990. Preparative purification of human monoclonal antibody isoforms in a multi-compartment electrolyser with immobiline membranes. J. Chromatogr. 500:681696.
Giaffreda, E., Tonani, C., and Righetti, P.G. 1993. A pH gradient simulator for electrophoretic techniques in a Windows environment. J. Chromatogr. 630:313-327.
Righetti, P.G., Castagna, A., and Herbert, B. 2001. Prefractionation techniques in proteome analysis: A new approach identifies more lowabundance proteins. Anal. Chem. 73:320-326.
Gygi, S.P., Rist, B., Gerber, S.A., Turecek, F., Gelb, M.H., and Aebersold, R. 1999. Quantitative analysis of complex protein mixtures using isotope-coded affinity tags. Nat. Biotechnol. 17:994-999.
Righetti, P.G., Castagna, A.C., Herbert, B.H., Reymond, F.R., and Rossier, J.S. 2003. Prefractionation techniques in proteome analysis. Proteomics 3:1397-1407.
Gygi, S.P., Rist, B., Griffin, T.J., Eng, J., and Aebersold, R. 2002. Proteome analysis of low-abundance proteins using multidimensional chromatography and isotope-coded affinity tags. J. Proteome Res. 1:47-54. Herbert, B.R., Molloy, M.P., Walsh, B.J., Gooley, A.A., Bryson, W.G., and Williams, K.L. 1998. Improved protein solubility in 2-D electrophoresis using tributyl phosphine. Electrophoresis 19:845-851. Hoffmann, P., Hong, J., Moritz, R.L., Connolly, L.M., Frecklington, D.F., Layton, M.J., Eddes, S., and Simpson, R.J. 2001. Continuous free-flow electrophoresis separation of cytosolic proteins from the human colon carcinoma cell line LIM 1215: A non two-dimensional gel elecrophoresis–based proteome analysis strategy. Proteomics 1:807-818. Pharmacia Biotech. 1997. Isoelectric membrane formulas for IsoPrime Purification of proteins. IsoPrime Protocol Guide #1. Pharmacia Biotech, Uppsala, Sweden. Microscale Solution Isoelectrofocusing (MicroSol - IEF)
dimensional electrophoresis. Electrophoresis 19:758-760.
Quadroni, M. and James, P. 1999. Proteomics and automation. Electrophoresis 20:664-677. Rabilloud, T. 1998. Use of thiourea to increase the solubility of membrane proteins in two-
Speicher, D.W., Lee, K., Tang, H.Y., Echan, L., AliKhan, N., Zuo, X., and Hembach, P. 2004. Current challenges in proteomics: Mining low abundance proteins and expanding protein profiling capacities. In Proceedings of the 16th International Mass Spectrometry Conference (A.E. Ashcroft, A.G. Brenton, and J.J. Monaghan, eds). 16:37-57. Unlu, M., Morgan, M.E., and Minden, J.S. 1997. Difference gel electrophoresis: A single gel method for detecting changes in protein extracts. Electrophoresis 18:2071-2077. Wall, D.B., Kachman, M.T., Gong, S., Hinderer, R., Parus, S., Misek, D.E., Hanash, S.M., and Lubman, D.M. 2000. Isoelectric focusing nonporous RP HPLC: A two-dimensional liquidphase separation method for mapping of cellular proteins with identification using MALDI-TOF mass spectrometry. Anal. Chem. 72:1099-1111. Weber, G. and Bocek, P. 1998. Recent developments in preparative free flow isoelectric focusing. Electrophoresis 19:1649-1653. Zuo, X. and Speicher, D.W. 2000. A method for global analysis of complex proteomes using sample prefractionation by solution isofocusing prior to two-dimensional electrophoresis. Anal. Biochem. 284:266-278.
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Zuo, X. and Speicher, D.W. 2002. Comprehensive analysis of complex proteomes using microscale solution isoelectrofocusing prior to narrow pH range two-dimensional electrophoresis. Proteomics 2:58-68. Zuo, X., Echan, L., Hembach, P., Tang, H.Y., Speicher, K.D., Santoli, D., and Speicher, D.W. 2001. Towards global analysis of mammalian proteomes using sample prefractionation prior to narrow pH range two-dimensional gels and using one-dimensional gels for insoluble and large proteins. Electrophoresis 22:1603-1615. Zuo, X., Hembach, P., Echan, L., and Speicher, D.W. 2002. Enhanced analysis of human breast cancer proteomes using micro-scale solution isoelectrofocusing combined with high resolution 1-D and 2-D gels. J. Chromatogr. B 782:253-265. Zuo, X., Lee, K., and Speicher, D.W. 2004. Electrophoretic prefractionation of proteomes for comprehensive analysis of proteomes. In Proteome Analysis: Interpreting the Genome (D.W. Speicher, ed.), pp. 93-118. Elsevier Science Inc., New York.
Key References Zuo and Speicher, 2000. See above. The first publication for development of proteome prefractionation using microscale solution IEF. Zuo and Speicher, 2002. See above. Demonstrated the enhanced protein profiling of mouse serum by combining narrow pH range 2-D gels and prefractionation by microscale solution IEF. Zuo et al., 2002. See above. The first publication to demonstrate the combination of microscale solution IEF and downstream high resolution 1-D and 2-D gels for improvement of breast cancer proteome analyses.
Contributed by Xun Zuo, KiBoem Lee, Nadeem Ali-Khan, and David W. Speicher The Wistar Institute Philadelphia, Pennsylvania
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Chapter 23 Non-Gel-Based Proteome Analysis INTRODUCTION
P
roteomics is a new discipline that originated in the mid-1990s and has grown extremely rapidly in a very short time. This, in turn, has driven improvements in diverse technologies. Most proteome studies fall into one of three types: (1) compositional analysis where one attempts to identify all proteins present in a proteome or sub-proteome; (2) protein profile comparisons where one attempts to quantitatively compare changes in the levels of as many proteins as possible, and (3) analysis of protein-protein interactions. The first two types of studies often utilize the same separation and analysis methods and differ only in the need for quantitation in the protein profiling studies. The oldest and most commonly used methods for separating complex protein mixtures for proteomics utilize two-dimensional gel electrophoresis and related gel methods (Chapter 22). Twodimensional gels remain as the gold standard for protein profiling, to which all alternative methods should be compared. However, there is substantial interest in recently developed and emerging protein profiling and compositional analysis methods that do not rely on 2-D dimensional gels. This chapter describes these non-2-D gel based separation and analysis methods. Many of these methods can be more readily automated than gel methods and may develop into higher-throughput methods.
describes the most commonly used non-gel proteomics method which relies on multi dimensional chromatography of peptides prior to mass spectrometry analysis to separate and analyze proteomes. Such multidimensional methods can be used to quantitatively compare two samples if one sample is tagged with a stable isotope. UNIT 23.2 describes one popular method for both isotopically tagging samples for comparisons and for simplifying the peptide mixtures, namely the isotope-coded affinity tag (ICAT) method. UNIT 23.4 provides one of a number of alternative methods for differentially tagging samples with stable isotopes using H2[18O] during or after trypsin digestion. An alternative strategy to fragmenting complex protein samples with trypsin prior to separating the resulting very complex peptide mixtures (bottom-up methods) is to first separate the proteins prior to mass spectrometry analysis and protein identification (top-down methods) using multi-dimensional liquid chromatography (UNIT 23.3). Future units will describe additional top-down and bottom-up methods. UNIT 23.1
David Speicher
Non-Gel-Based Proteome Analysis Contributed by David Speicher Current Protocols in Protein Science (2003) 23.0.1-23.0.2 Copyright © 2003 by John Wiley & Sons, Inc.
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Analysis of Protein Composition Using Multidimensional Chromatography and Mass Spectrometry
UNIT 23.1
This unit describes coupling multidimensional microcapillary liquid chromatography (LC) of peptides with tandem mass spectrometry (MS/MS) for identifying proteins in complex biological samples. Computational comparison of acquired tandem spectra with genomic sequence databases and data processing produces a list of proteins in the original sample (UNITS 16.6 & 16.10). Originally published under the term direct analysis of large protein complexes (DALPC; Link et al., 1999), it is typically now published as multidimensional protein identification technology (MudPIT; Washburn et al., 2001). Compared to single-dimension chromatography LC-MS/MS (UNIT 16.9), multidimensional LCMS/MS chromatography has dramatically increased resolution and loading capacity for identifying large numbers of proteins with a wide dynamic range of abundances. While numerous examples of multidimensional separations have been described (Moore et al., 1996; Opiteck et al., 1997), the combination of strong-cation exchange (SCX) with reversed-phase (RP) chromatography has proven to be most successful (Link et al., 1999; Washburn et al., 2001). The protein identification approaches in this unit are in contrast to methodologies that use gel electrophoresis to fractionate complex protein mixtures followed by in-gel digestion of the proteins and mass spectrometric identification (UNITS 16.4, 16.8, & 16.9). Starting with either a purified protein complex, a subcellular protein fraction, or a whole cell lysates, the complex mixture of proteins in solution is first denatured and reduced (UNIT 11.1). Second, the mixture of proteins is cleaved either enzymatically (i.e., digestion) or chemically to generate a mixture of peptide fragments (see UNITS 11.1 & 11.4). The complex peptide mixture is fractionated and analyzed by multidimensional liquid chromatography coupled with electrospray ionization (ESI) and automated MS/MS (see below). The acquired tandem mass spectra are computationally compared to protein sequences in either protein or translated nucleic acid databases (UNITS 16.6 & 16.10). The list of peptides significantly matching sequences in the databases are used to generate a list of proteins in the original protein complex, subcellular compartment, or whole cell lysate. The first basic protocol (see Basic Protocol 1) describes an offline approach for coupling multidimensional liquid chromatography with electrospray ionization (ESI) and tandem mass spectrometry (MS/MS). It involves off-line SCX fractionation of peptides followed by RP-LC-MS/MS of each fraction. The second basic protocol (see Basic Protocol 2) describes a novel inline multidimensional fractionation using a single, biphasic SCX-RP microcapillary column directly coupled to an ESI mass spectrometer. Both approaches have demonstrated the ability to identify >1000 different proteins from biological samples (Washburn et al., 2001; Gygi et al., 2002). While different configurations and instruments for performing multidimensional chromatography coupled to mass spectrometry can be considered, this unit focuses on high sensitivity approaches for analyzing limited amounts of starting material (Link et al., 1999; Washburn et al., 2001; Sanders et al., 2002). Support protocols describe construction of both fritted (see Support Protocol 1) and fritless (see Support Protocol 2) capillary columns, as well as a method for packing them (see Support Protocol 3). CAUTION: Always wear safety glasses when working with a pneumatic loading device. Non-Gel-Based Proteome Analysis Contributed by Andrew J. Link and Jennifer L. Jennings, and Michael P. Washburn Current Protocols in Protein Science (2003) 23.1.1-23.1.25 Copyright © 2003 by John Wiley & Sons, Inc.
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STRATEGIC PLANNING When deciding to use multidimensional chromatography coupled with mass spectrometry, several decisions need to be made. If the technique will only be used for a small number of samples, off-line SCX fractionation of the sample coupled with manual loading and analysis of each fraction by LC-MS/MS is an option. If repeated analysis of complex samples is planned, computer-controlled automation of the processes will be essential. Automation greatly improves the efficiency of the process since samples can be processed reproducibly and unattended. Autosampling of off-line SCX peptide fractions for RP-LCMS/MS analysis or computer-controlled MudPIT should be considered if multidimensional chromatography will be used routinely. Regardless of which approach is used, the process generates thousands of tandem mass spectra that need to be analyzed using MS/MS search algorithms such as SEQUEST (Thermo Finnigan), Mascot (Matrix Science), or MS-Tag (University of California, San Francisco). Search times against a large protein database and processing the result into a list of proteins in the original sample will quickly become an issue. Off-line processing of the data using either multiprocessor computers or distributive computing should be considered. BASIC PROTOCOL 1
OFF-LINE SCX FRACTIONATION COUPLED TO RP-LC-ESI-MS/MS This protocol describes the procedure for performing multidimensional separations in two separate steps. A complex peptide mixture is loaded onto a strong-cation exchange (SCX) capillary column and discrete fractions of peptides are then displaced from the column using a salt gradient. Each fraction is loaded onto a RP capillary column and analyzed using RP-LC-ESI-MS/MS. The acquired tandem mass spectra are searched against a sequence database using a MS/MS search algorithm to identify the proteins in the original sample. Off-line SCX fractionation of peptides can also be performed using standard HPLC methods (UNIT 8.7). Because of the relatively high flow rates and composition of the SCX mobile phases, the SCX fractions are typically lyophilized and the peptides resolubilized in 0.5% acetic acid prior to RP-LC-ESI-MS/MS. To assure peptides bind to the SCX column, digested protein samples are desalted prior to SCX fractionation using an off-line RP trapping cartridge. Several vendors offer trapping systems for desalting peptide mixtures prior to SCX fractionation. Materials 2% and 95% acetonitrile/0.1% TFA 70% formic acid/30% isopropanol Enzymatically digested or chemically cleaved protein sample 0.5% acetic acid SCX buffers A, B, C, and D (see recipes) Angiotensin solution (see recipe) High-pressure helium Trapping system: 0.5-, 20-, or 200-µg reversed-phase (RP) trapping cartridges (Microm BioResources) RP trapping cartridge holder with syringe port and capillary fitting (Microm BioResources) 10- and 50-µl syringes (Hamilton)
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SCX apparatus: 50- and 75-µm-i.d. × 365-µm-o.d. fused-silica capillary (FSC; Polymicro Technology) Quaternary HPLC pump (Agilent Technologies) PEEK micro tee (Upchurch) PEEK microtight adapter (Upchurch) PEEK 380-µm-i.d. microtight sleeves (Upchurch) Capillary SCX column (see Support Protocol 3) PEEK microtight true zero dead volume (ZDV) union (Upchurch) Syringe port (Upchurch) 10-µl sample loop (Upchurch) Quartz capillary cleaving tool (Scientific Instrument Services) Reversed-phase (RP) apparatus: Binary HPLC pump (Agilent Technologies) PEEK micro tee (Upchurch) PEEK 380-µm-i.d. microtight sleeves (Upchurch) Capillary RP column (see Support Protocol 3) 50- and 75-µm-i.d. × 365-µm-o.d. fused-silica capillary (FSC; Polymicro Technology) 0.025-in.-o.d. gold wire (Scientific Instrument Services) Pneumatic loading vessel (James Hill Instruments) 1⁄ -in. to 0.4-mm Vespel ferrules (Chromatography Research Supplies) 16 20-µm-i.d. × 365-µm-o.d. uncoated emitter tip with 10-µm opening (New Objective) 5-µl graduated glass capillary (Sigma) Microspray ionization source with x-y-z manipulator ESI mass spectrometer Additional equipment and reagents for analyzing MS/MS data (UNIT 16.10) NOTE: Routine use of a trapping cartridge is recommended for desalting peptides and filtering samples. Prepare RP trapping cartridge 1. Place the RP trapping cartridge into the trap holder. Connect the syringe port and the capillary fitting to the trap holder. Use the appropriate size RP cartridge for the sample quantity.
2. Using a 50-µl syringe, inject 50 µl of 95% acetonitrile/0.1% TFA through the trap cartridge. This highly organic solvent removes any nonpolar contaminants.
3. Inject 50 µl of 70% formic acid/30% isopropanol through the trap. This extremely stringent solvent washes away any contaminants and removes any previous sample peptides.
4. Inject 50 µl of 2% acetonitrile/0.1% TFA through the trap. This step equilibrates the trap to ensure all the organic solvent is removed to allow sample peptide trapping. Additional injections of 2% acetonitrile/0.1% TFA can be performed prior to sample loading to ensure the trap is equilibrated.
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Prepare peptide sample 5. Pellet solid material that may plug the trapping cartridge by centrifuging the peptide sample 1 min at 14,000 × g, 4°C. 6. Aspirate the sample into a clean 50-µl syringe. Desalt peptides 7. Slowly inject the sample through the equilibrated trap. During the injection of the sample, the flowthrough fraction can be collected in a microcentrifuge tube to recover peptides that fail to bind to the RP cartridge.
8. Inject 50 µl of 2% acetonitrile/0.1% TFA through the trap. Additional washes with 2% acetonitrile/0.1% TFA may be necessary if the original peptide sample contains excessive amounts of salts, chaotropes, or detergents.
9. Inject 50 µl of 95% acetonitrile/0.1% TFA through the trap and collect the eluate into a labeled microcentrifuge tube. This solution is highly organic and elutes the peptides from the trap. This is the peptide fraction.
Remove solvent and resuspend peptides 10. Freeze the desalted peptide sample on dry ice and then lyophilize to a volume of 1 to 2 µl using a Speedvac evaporator. Excessive lyophilization can lead to loss of low abundance peptides.
11. Resuspend the peptides in 0.5% acetic acid to a final volume of 10 µl. Clean and store trapping cartridge 12. Clean and equilibrate the microtrap with two 50-µl washes of 95% acetonitrile/0.1% TFA, then two 50-µl washes of 70% formic acid/30% isopropanol, and finally two 50-µl washes of 2% acetonitrile/0.1% TFA. 13. Disassemble the trap holder and plug the ends of the trap to prevent the RP cartridge from drying out. The RP cartridge can be removed from the holder and stored indefinitely in 0.5% acetic acid at room temperature.
Assemble capillary SCX-HPLC apparatus When working with limited amounts of material, SCX fractionation is performed in capillary columns to minimize sample losses. To reduce the HPLC flow rate to the capillary column, the SCX column is connected to a splitting tee (see Fig. 23.1.1). The SCX column is conditioned with angiotensin prior to loading sample peptides to block nonspecific, irreversible binding sites in the column. 14. Assemble the off-line SCX apparatus as follows (Fig. 23.1.1):
Protein Analysis by Multidimensional Chromatography & Mass Spec
a. Connect the typical 1⁄16-in. transfer line from the HPLC pump (intakes connected to SCX buffers A, B, C, and D) to a 75-µm-i.d. × 365-µm-o.d. × 10-cm long FSC transfer line using a PEEK microtight adapter and a PEEK 380-µm-i.d. microtight sleeve. b. Connect this 75-µm-i.d. × 365-µm-o.d. FSC transfer line from the PEEK microtight adapter to a PEEK micro tee using a PEEK 380-µm-i.d. microtight sleeve. c. Connect the capillary SCX column on the opposite arm using a PEEK microtight ZDV union, a 10-cm piece of 75 × 365–µm FSC, and PEEK microtight sleeves.
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d. Prepare a restrictor line by connecting a 30-cm piece of 50-µm-i.d. × 365-µm-o.d. FSC through the third arm of the tee using a PEEK microtight sleeve. The micro tee splits the flow between the SCX capillary column and the restrictor line. The HPLC pump is set at a constant flow rate and the length of the restrictor line is adjusted to deliver the desired flow rate to the HPLC column. This splitter assembly allows the HPLC pump to be operated at flow rates that accurately deliver and adequately mix precise gradients. Typically, the pump is set at 300 ìl/min and the restrictor line is shortened to give a 300-to-1 flow split for an effective 1ìl/min flow rate to the HPLC column. The flow through the restrictor line is collected in a waste bottle and discarded.
Adjust flow rate and equilibrate SCX column 15. Set the HPLC pump to 100% SCX buffer A with a flow rate of 300 µl/min. Hold the HPLC pump flow rate constant throughout the offline SCX fractionation. The composition of the flow changes in a series of increasing salt steps during the offline SCX fractionations.
16. Trim the 50 × 365–µm restrictor line using the quartz capillary cleaving tool until a flow rate of 1 µl/min through the SCX column is reached as measured for 1 min using a 5-µl graduated glass capillary pipet. Measurement of the flow-rate and adjustment of the split line may have to be repeated several times until the target flow rate is reached.
17. Equilibrate the column 10 min with 100% SCX buffer A (i.e., solvent A).
PEEK microtight PEEK micro tee FSC PEEK microtight ZDV union ZDV adaptor (75-µm i.d. × 365-µm o.d.)
flow direction
FSC (75-µm i.d. × 365-µm o.d.)
PEEK sleeve (380-µm i.d. × 0.025-in. o.d.)
1/16-in. transfer line from HPLC pump
SCX column
50-µm i.d. × 365-µm o.d. FSC restrictor line
waste
Figure 23.1.1 Off-line SCX assembly. The device couples a quaternary HPLC pump to a capillary SCX column for fractionating complex peptide mixtures. The micro tee splits the flow between the SCX capillary column and the restrictor line. The assembly allows the HPLC pump to be operated at flow rates that accurately deliver and adequately mix precise gradients. The length of the restrictor line is adjusted to deliver the desired flow rate to the HPLC column. Typically, the pump is set at 300 µl/min and the restrictor line is cut to give a 300-to-1 flow split for an effective 1 µl/min flow rate to the HPLC column. The ZDV union enables a sample loop to be easily installed between the micro tee and the SCX column (see Fig. 23.1.2). Abbreviations: FSC, fused-silica capillary; HPLC, high-performance liquid chromatography; PEEK, polyetheretherketone; SCX, strong cation exchange; ZDV, zero dead volume.
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A
syringe port
sample loop
PEEK ZDV union
FSC (75 × 365)
B flow direction
flow splitter
PEEK sleeve
PEEK microtight FSC ZDV adaptor 75-µm i.d × 365-µm o.d.
sample loop
PEEK microtight ZDV adaptor 50-µm i.d. × 365-µm o.d. FSC restrictor line
FSC (75 × 365)
PEEK sleeve
PEEK microtight PEEK microtight ZDV adaptor ZDV union SCX column
transfer line assembly from HPLC pump (see Figure 23.1.1 for details)
Figure 23.1.2 Loading assembly for off-line SCX capillary column. The assembly allows a peptide mixture to be quickly loaded onto the SCX capillary column without a pneumatic loading device (see Fig. 23.1.4). In (A), a peptide mixture in a volume equal to the size of the sample loop is loaded into the loop. In (B) the filled sample loop is installed inline between the flow splitter and the SCX column. The flow from the HPLC pump loads the sample from the sample loop onto the SCX column. Abbreviations: FSC, fused-silica capillary; HPLC, high-performance liquid chromatography; PEEK, polyethe- retherketone; SCX, strong cation exchange; ZDV, zero dead volume.
Prepare sample loading loop 18. Connect a syringe port to a 10-µl sample loop using a PEEK ZDV union (Fig. 23.1.2A). Connect the opposite end of the loop to a piece of 75 × 365–µm FSC using a PEEK Microtight ZDV adapter and a PEEK sleeve. HPLC injection valves are avoided to eliminate carryover from previous samples. The sensitivity of mass spectrometry makes sample carryover a major concern. Although peptide samples can be loaded onto the SCX column using a pneumatic loading vessel, the use of a sample loop and the HPLC pump allows higher pressure to be used to quickly and reliably load peptide samples onto the capillary SCX HPLC columns.
19. Use a 10-µl syringe to clean and equilibrate the 10-µl sample loop with two 10-µl washes of SCX buffer B, then two 10-µl washes of SCX buffer C, and finally two 10-µl washes of SCX buffer A.
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Load angiotensin 20. Inject 10 µl angiotensin solution into the loop. 21. Remove the syringe port and connect the angiotensin-filled sample loop inline to the SCX column as follows: a. Set the HPLC pump flow to 0. b. Disconnect the sample loop from the syringe port and PEEK ZDV union, leaving a free sample loop end (Fig. 23.1.2A). Attach a PEEK microtight adapter to the free sample-loop end. c. Remove the ZDV union joining the SCX column and the FSC connected to the micro tee (Fig. 23.1.1) d. Join the newly added adapter connected to the sample loop to the free end of the FSC connected to the micro tee (Fig. 23.1.2B). e. Connect the free ZDV adapter attached to the sample loop directly to the ZDV union joining the SCX column. f. Set the HPLC flow rate to 300 µl/min at 100% SCX buffer A. The SCX column is disconnected from the tee, and the sample loop is connected inline between the tee and the column using two PEEK Microtight adapters (see Fig. 23.1.2B).
22. Run 100% SCX buffer A for 15 min. This allows angiotensin to transfer from the loop to the SCX column.
23. Remove the sample loop as follows: a. Set the HPLC flow rate to 0. b. Disconnect the microtight adapter from FSC that attaches the sample loop to the micro tee (splitter tee) (Fig. 23.1.2B). c. Disconnect the microtight union from the FSC that attaches the sample loop to the SCX column (Fig. 23.1.2B). d. Connect the SCX column with the attached ZDV union to the micro tee (Fig. 23.1.1). e. Set the HPLC flow rate to 300 µl/min at 100% SCX buffer A. Removing the sample loop reduces the dead volume in the assembly during the SCX fractionation.
Block nonspecific binding sites 24. Run 100% SCX buffer B for 15 min across the SCX column. The flow of angiotensin through the column blocks nonspecific binding sites.
25. Equilibrate the SCX column 15 min with 100% SCX buffer A. 26. Clean and equilibrate the sample loop with two 10-µl washes of SCX buffer B, then two 10-µl washes of SCX Buffer C, and finally two 10-µl washes of SCX buffer A. Load sample 27. Inject 10 µl of the desalted peptides (step 11) into the sample loop as described in step 20. Connect the loop to SCX column as described in step 21. 28. Run 100% SCX buffer A for 15 min and collect the flow-through in a fresh tube. The flow-through fraction is later frozen on dry ice and lyophilized using a Speedvac evaporator. This fraction is resuspended in 6 ìl of 0.5% acetic acid and analyzed by RP-LS-MS/MS to identify peptides that fail to bind to the SCX column.
29. Disconnect the SCX column, remove the sample loop, and reconnect the column to the tee as described in step 23.
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Fractionate peptides 30. Perform a series of 6-min step gradients of 10%, 20%, 30%, 40%, 60%, 80%, and 100% SCX buffer B versus A. Collect the 6-min (6-µl) fractions in labeled tubes. For this first dimension fractionation, the step gradients can be programmed on the quaternary HPLC pump. The 6-ìl fractions can be collected in either 0.5-ml microcentrifuge tubes or autosampler tubes. The number of fractionation steps can be increased or decreased depending upon the complexity of the sample.
31. Collect separate 6-min fractions of 100% SCX buffer C and then 100% SCX buffer D in labeled tubes. Desalt these fractions as described in steps 1 to 10. Resuspend in 6 µl of 0.5% acetic acid. Analyze by RP-LC-MS/MS to identify peptides tightly associated with the SCX column. These two additional high-salt washes elute peptides that are tightly bound to the SCX column.
emitter tip (20-µm i.d. × 365 µm o.d., 10-µm opening)
RP column fused silica transfer line from HPLC pump (75-µm i.d. × 365-µm o.d.)
PEEK microtight ZDV union x -y-z micromanipulator stage
PEEK micro tee
capillary opening into the mass spectrometer
fused silica capillary (75-µm i.d. × 365-µm o.d.)
gold wire (0.025-in. o.d.)
PEEK sleeve
to ESI voltage source
PEEK ESI-restrictor micro tee
50-µm-i.d. × 365-µm-o.d. FSC restrictor line
waste
Protein Analysis by Multidimensional Chromatography & Mass Spec
Figure 23.1.3 RP-LC-ESI-MS assembly. The schematic shows a system for performing microcapillary RP-LC-ESI-MS. The micro tee splits the flow from the HPLC pump between the RP capillary column and the ESI-restrictor micro tee. The ESI-restrictor tee provides a liquid-metal interface to apply a voltage for a stable electrospray at the RP column’s emitter tip during the RP gradients. The length of the FSC restrictor line is adjusted to achieve the desired flow rate to the RP column. The x-y-z manipulator is used to precisely position the RP column emitter tip in front of the opening into the mass spectrometer. To load peptide samples, the RP column is disconnected from the micro tee and inserted into the pneumatic device, shown in Fig. 23.1.4, containing the peptide sample. Abbreviations: ESI, electrospray ionization; FSC, fused silica capillary; HPLC, high-performance liquid chromatography; PEEK, polyetheretherketone; RP, reversed phase; ZDV, zero dead volume.
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Turn off SCX apparatus 32. Equilibrate the SCX column 10 min with 100% SCX buffer A and turn off the pump. Assemble RP-HPLC and mass spec apparatus The following steps describe the RP-LC-ESI-MS assembly and process used to analyze the SCX fractions (see Fig. 23.1.3). Prior to analysis, the RP column is conditioned with angiotensin to block nonspecific binding sites to minimize sample losses and to check the performance of the chromatography and mass spectrometry systems. 33. Prepare the RP-apparatus as follows (see Fig. 23.1.3): a. Connect the transfer line from the binary HPLC pump to the center arm of a PEEK micro tee using a PEEK 380-µm-i.d. microtight sleeve. b. Connect the capillary RP column to one arm of the tee using a PEEK sleeve. c. Connect a PEEK ESI-restrictor micro tee to the opposite arm using a 75 × 365–µm FSC and PEEK sleeves. d. Connect a 0.025-in.-o.d. gold wire through the center arm of the ESI-restrictor tee and attach to the ESI voltage source. e. Connect a 30-cm piece of 50 × 365–µm FSC restrictor line through the third arm of the ESI-restrictor tee using a PEEK sleeve. The ESI-restrictor tee is the site where the voltage is applied to produce ESI. A voltage of ∼2.2 kV is applied to the gold wire during ESI. Any gas bubbles generated by electrolysis from the ESI voltage are directed to the restrictor line and waste.
Adjust flow rate and equilibrate RP column 34. Set the HPLC pump to 100% solvent A (0.5% acetic acid) with a flow rate of 200 µl/min. 35. Measure the flow rate through the column for 1 min using a 5-µl graduated glass capillary pipet. Trim the 50 × 365–µm restrictor line using the capillary cleaving tool until a flow rate of 0.2 to 0.5 µl/min through the RP column is obtained. Measurement of the flow rate and adjustment of the split line may have to be repeated several times until the target flow rate is achieved.
36. Run a blank RP-HPLC gradient (see step 50 for example) of acetonitrile to pack the RP resin and condition the column. A 15-min gradient from 0% to 60% acetonitrile is typically used to condition the column. The column is re-equilibrated with 100% solvent A (0.5% acetic acid) for 10 min before loading a peptide sample.
Load angiotensin using a pneumatic vessel 37. Place a tube containing 5 µl angiotensin solution (0.5 pmol) into a pneumatic loading vessel and tightly attach the lid of the vessel (Fig. 23.1.6). 38. Disconnect the column from the PEEK assembly and insert the open end of the column into the top of the loading vessel until the capillary reaches the bottom of the sample tube. Pull the column up slightly off the bottom of the sample tube. Leaving a small gap between the capillary and the bottom of the tube reduces the probability of any solid particulates at the bottom of the tube clogging the capillary column.
39. Secure the column to the bomb by tightening the compression nut on the Swagelok fitting at the top of the loading vessel. Tightening the Swagelok fitting’s nut compresses the Vespel ferrule around the FSC (Fig. 23.1.4). Non-Gel-Based Proteome Analysis
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fused silica capilary (365-µm o.d.)
bolts securing lid to base
regulator
1/16-in. × 0.4-µm i.d. vespel ferrule in a 1/16-in. swagelok fitting to 1/8-in. male NPT
3-way valve O-ring
high pressure helium tank
tube with sample or column packing material
vent 1/8-in. NPT to 1/8-in. swagelok fitting
Teflon tube holder
stir plate
Figure 23.1.4 Stainless steel pneumatic loading vessel. The device is used to slurry-pack capillary HPLC columns with RP or SCX material. It is also used to load peptides samples onto the microcapillary columns. When loading peptides onto the HPLC columns, the stir plate is not used. When the three-way valve is turned, ∼500 to 1000 psi helium forces the sample into the capillary column. After the injection, the pressure is released. The pneumatic vessel is fabricated from ∼1⁄2-in. stainless steel. The top is made from the same thickness steel and is attached by bolts to the base. An O-ring seals the top and base. Abbreviations: NPT, National Pipe Thread.
40. Apply pressure to the loading vessel by first setting the regulator on the helium gas cylinder to 500 to 1000 psi, then opening the three-way valve. Load angiotensin onto the RP column 41. Measure the volume loaded using a 5-µl calibrated glass pipet to collect the displaced volume from the end of the column. 42. Release the pressure in the loading vessel using the three-way valve once the sample is loaded. 43. Reinstall the column in the union and run an HPLC gradient. The RP column can be plumbed for performing microspray mass spectrometry on the angiotensin as it elutes from the column (see steps 44 to 52). Monitoring the retention time and signal intensity of the angiotensin peptide can be used to check the performance of the chromatography and mass spectrometry system. For the HPLC gradient, a 60-min gradient from 0% to 40% acetonitrile followed by a 10-min gradient from 40% to 60% acetonitrile can be used. This is identical to the HPLC gradient used for RP-LC-ESI-MS/MS analysis of the SCX fractions in step 50 of this protocol.
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Prepare RP column for LC-ESI-MS-MS of SCX fractions 44. Use a quartz capillary cutter to trim the ragged end of the emitter tips. Connect the emitter tip to the fritted end of the RP column using a PEEK microtight ZDV union and PEEK 380-µm-i.d. microtight sleeves. 45. Mount the emitter tip assembly to the x-y-z manipulator of the microspray ionization source at the entrance of the mass spectrometer. The RP column and emitter tip assembly is mounted to an x-y-z manipulator at the entrance of the mass spectrometer. The manipulator allows for fine adjustment of the column position with respect to the mass spectrometer. The emitter tip is extremely fragile. Avoid letting the tip strike a solid surface.
46. Start the HPLC flowing at 200 µl/min solvent A (0.5% acetic acid) and recheck the RP column flow rate (step 35). Separate SCX fraction by RP chromatography 47. Load a SCX fraction onto the RP column using the loading vessel as described in steps 37 to 43. For DALPC, a FAMOS plate autosampler (Dionex) can be used to automatically load SCX fractions onto the RP column. This avoids the tedious process of manually loading SCX fractions and allows fractions to be analyzed unattended. When using the autosampler, the PEEK flow splitting tee is placed in-line between the HPLC pump and the autosampler (see Fig. 23.1.5). The HPLC flow from the autosampler is connected to one arm of the PEEK ESI tee. The RP column is attached to the opposite arm of the ESI tee. A method for rapidly loading fractions using an autosampler and a novel vented RP column assembly has been described (Licklider et al., 2002). The MudPIT approach described in this unit (see Basic Protocol 2) employs a biphasic SCX/RP and computer control to automate most of the process.
48. Set the HPLC for 100% solvent A (0.5% acetic acid) with a flow rate of 200 µl/min. 49. Carefully position the emitter tip at the entrance of the mass spectrometer. Use a camera monitor to assist in positioning the emitter tip at the optimal position. The tip of the emitter is centered 1 to 5 mm from the orifice of the capillary opening into the mass spectrometer.
50. Program the HPLC using the following as an example: 20 min isocratic at 0% acetonitrile (100% of 0.5% acetic acid) 60 min gradient from 0% to 40% acetonitrile 10 min gradient from 40% to 60% acetonitrile. There are many gradient variations and RP buffers that can be used.
Analyze fractions by ESI-MS 51. Carefully wick away any large drops from the emitter tip. Start the mass spectrometer and the HPLC gradient and collect MS/MS data on peptides as they elute from the RP column. A voltage of ∼2.2 kV is applied to the gold wire during ESI. Typical data acquisition settings consist of a continual cycle beginning with one MS scan with a m/z scan range of 300 to 2000, which records all the m/z values of ions present at that moment in the gradient, followed by three rounds of MS/MS. The MS/MS scans fragment the three most abundant ions recorded in the first MS scan. Dynamic exclusion is activated to improve protein identification capacity by avoiding the repeated fragmentation of abundant ions. Non-Gel-Based Proteome Analysis
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emitter tip (20-µm i.d. × 365-µm o.d.,10-µm opening) RP column
gold wire (0.025-in. o.d.) to ESI voltage source
FSC (75-µm i.d. × 365-µm o.d.)
PEEK sleeve
PEEK microtight ZDV union x -y -z micromanipulator stage
PEEK ESI micro tee
capillary opening into the mass spectrometer
FSC fused silica transfer (75-µm i.d. × 365-µm o.d.) line from HPLC pump (75-µm i.d. × 365-µm o.d.)
autosampler 50-µm-i.d. × 365-µm-o.d. FSC restrictor line waste
Figure 23.1.5 RP-LC-ESI-MS autosampler assembly. The schematic shows a system for the automated RP-LC-ESI-MS analysis of SCX peptide fractions collected offline. The system allows SCX fractions to be automatically loaded onto the RP capillary column and analyzed. The legend of Figure 23.1.3 describes the functions of the different features shown in the figure. Abbreviations: ESI, electrospray ionization; FSC, fused-silica capillary; HPLC, high-performance liquid chromatography; PEEK, polyetheretherketone; RP, reversed phase; ZDV, zero dead volume.
52. When the ESI-LC-MS/MS run is complete, turn off the ESI voltage. 53. Analyze each of the remaining SCX fractions (steps 28, 30, and 31) in the same way. Be sure to equilibrate the RP column for 10 min with 0.5% acetic acid before loading the next SCX fraction. Process ESI-MS/MS data 54. Upon completion of the run, process the acquired MS/MS data essentially as described in UNIT 16.10. BASIC PROTOCOL 2
Protein Analysis by Multidimensional Chromatography & Mass Spec
MULTIDIMENSIONAL PROTEIN IDENTIFICATION TECHNOLOGY (MudPIT) In multidimensional protein identification technology (MudPIT), a digested protein complex is loaded directly onto a biphasic SCX-RP capillary column that directly elutes into an ESI mass spectrometer. The peptides are sequentially eluted from the capillary column and fragmented in the mass spectrometer. In this protocol, MudPIT is carried out using a Thermo Finnigan LCQ ion trap mass spectrometer and an Agilent quaternary HPLC pump. The biphasic SCX-RP MudPIT column is interfaced directly to the mass spectrometer and HPLC pump using a PEEK micro cross assembly (see Fig. 23.1.6). The HPLC pump is directly controlled by LCQ Xcalibur software. Within the Instrument Setup of Xcalibur, the gradient of each individual step is programmed. A typical analysis consists of a fully automated 14-step chromatography run on highly complex mixtures, but any number of steps can be applied to the column.
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There are several emerging choices of systems for DALPC/MudPIT-type analyses. Systems are becoming available for purchase from a variety of instrument manufacturers including the ProteomeX from Thermo Finnigan and a Q-TOF-CapLC system from Micromass/Waters. For researchers who wish to pursue this route, it is essential that the 2-D-HPLC and mass spectrometry systems are compatible with each other. For example, in this protocol, the chromatography is controlled by the mass spectrometer’s software. Materials MudPIT buffers A, B, C, and D (see recipes) Enzymatically digested or chemically cleaved protein sample Quaternary HPLC pump (Agilent Technologies) and Xcalibur software (Thermo Finnigan) PEEK micro-cross (Upchurch) PEEK 380-µm-i.d. microtight sleeves (Upchurch) Biphasic MudPIT column (see Support Protocol 2) 50-µm-i.d. × 365-µm-o.d. fused-silica capillary (Polymicro Technology) 0.025-in.-o.d. gold wire (Scientific Instrument Services) ESI voltage source 5-µl graduated glass capillary pipet Quartz capillary cleaving tool (Scientific Instrument Services)
to ESI voltage source
MudPIT column (not drawn to scale) (100-µm i.d. × 365-µm o.d.) gold wire (0.025-in. o.d.)
fused silica transfer line from quaternary HPLC pump
SCX
RP capillary opening into the mass spectrometer
PEEK sleeve PEEK micro-cross 50-µm-i.d. × 365-µm-o.d. FSC restrictor line
waste
Figure 23.1.6 MudPIT assembly. The schematic shows the biphasic microcapillary column and electrospray interface system for performing multidimensional protein identification technology (MudPIT). The length of the FSC restrictor line is adjusted to achieve the desired flow rate to the MudPIT column. To load samples, the column is disconnected from the micro-cross and a peptide sample is loaded using the pneumatic loading vessel (see Fig. 23.1.4). The HPLC and mass spectrometer are controlled simultaneously by means of the user interface of the mass spectrometer. A precolumn application of ∼1.8 kV at the liquid-metal interface produces a stable electrospray during the RP gradients. The MudPIT assembly is mounted to an x-y-z manipulator stage at the capillary opening of the mass spectrometer. The stage supports the microcross along with connections and allows for fine adjustment of the MudPIT column emitter tip with respect to the entrance to the mass spectrometer. Abbreviations: FSC, fused-silica capillary; HPLC, high-performance liquid chromatography; MudPIT, multidimensional protein identification technology; PEEK, polyetheretherketone.
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Pneumatic loading vessel (James Hill Instruments) 1⁄ -in. to 0.4-mm Vespel ferrules (Chromatography Research Supplies) 16 LCQ mass spectrometer (ThermoFinnigan) Microspray ionization source with x-y-z manipulator Additional equipment and reagents for loading from a pneumatic vessel (see Basic Protocol 1, steps 38 to 42) and analyzing MS/MS data (UNIT 16.10) 1. Connect the transfer line from the quaternary HPLC pump to one arm of the PEEK micro-cross using a PEEK 380-µm-i.d. microtight sleeve (Fig. 23.1.6). 2. Connect the biphasic MudPIT capillary column to the opposite arm of the cross using a PEEK sleeve. 3. Connect a 30-cm piece of 50-µm-i.d. × 365-µm-o.d. FSC restrictor line to the micro-cross using a PEEK sleeve. 4. Connect a 0.025-in.-o.d. gold wire through the opposite arm of the micro-cross and attach to the ESI voltage source. A voltage of ∼1.8 kV is applied to the gold wire during ESI.
5. Set the HPLC pump to 100% MudPIT buffer A with a flow rate of 150 µl/min. Measure the flow rate through the column for 1 min using a 5-µl graduated glass capillary pipet. Adjust the length of the 50 × 365–µm restrictor line using a quartz capillary cleaving tool to achieve an effective flow rate of 0.15 to 0.25 µl/min through the MudPIT column. Measurement of the flow-rate and adjustment of the split line may have to be repeated several times until the target flow-rate is obtained. To improve the accuracy of the flow rate calibration, one can increase the collection time over which the flow rate is measured.
6. Pellet any insoluble material from the sample by centrifuging 10 min at 14,000 × g, 4°C. If a pellet is visible, transfer the supernatant to a fresh tube. Even minute amounts of solid will plug the capillary column.
7. Disconnect the MudPIT column from the micro-cross and load the peptide mixture onto the biphasic column using a pneumatic loading vessel as described (see Basic Protocol 1, steps 38 to 42 and Fig. 23.1.6). 8. Reconnect the MudPIT column to the micro-cross. Position the emitter trip of the column within 5 mm of the orifice of the mass spectrometer’s capillary opening. Set the HPLC pump flow rate to 150 µl/min and 100% MudPIT buffer A, and recheck the flow rate to the column as described in step 5. Adjust the length of the restrictor line if necessary. The micro-cross assembly is mounted to an x-y-z manipulator at the entrance of the mass spectrometer. The manipulator allows for fine adjustment of the column position with respect to the mass spectrometer.
9. Program the Instrument Methods in the Xcalibur software.
Protein Analysis by Multidimensional Chromatography & Mass Spec
For the LCQ, the HPLC gradients and data-dependent acquisition of tandem spectra is programmed through the Xcalibur software. Typical data acquisition settings consist of a continual cycle beginning with one MS scan, with a m/z range of 400 to 1400, which records all the m/z values of ions present at that moment in the gradient, followed by three rounds of MS/MS. The MS/MS scans fragment the three most abundant ions recorded in the first MS scan. Dynamic exclusion is activated to improve protein identification capacity by avoiding the repeated fragmentation of abundant ions.
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10. Program the following sequential 14 gradients steps into the Gradient Program portion of the LCQ Xcalibur: Step 1
Steps 2–11
Step 12
Step 13
Step 14
0-min hold at 100% MudPIT buffer A (starting composition) 70-min gradient from 0% to 80% MudPIT buffer B 10-min hold at 80% MudPIT buffer B 5-min hold at 100% MudPIT buffer A 2-min salt wash at x% MudPIT buffer C (begin with 10%) 3-min hold at 100% MudPIT buffer A 10-min gradient from 0% to 10% MudPIT buffer B 90-min gradient from 10% to 45% MudPIT buffer B 5-min hold at 100% MudPIT buffer A 20-min salt wash at 100% MudPIT buffer C 3-min hold at 100% MudPIT buffer A 10-min gradient from 0% to 10% MudPIT buffer B 90-min gradient from 10% to 45% MudPIT buffer B 5-min hold at 100% MudPIT buffer A 20-min salt wash at 100% MudPIT buffer D 3-min hold at 100% MudPIT buffer A 10-min gradient from 0% to 10% MudPIT buffer B 90-min gradient from 10% to 45% MudPIT buffer B 5-min hold at 100% MudPIT buffer A 20-min salt wash at 100% MudPIT buffer D 3-min hold at 100% MudPIT buffer A 10-min gradient from 0% to 10% MudPIT buffer B 130-min gradient from 10% to 100% MudPIT buffer B
The iterative gradient steps 2 to 11 use the following x% MudPIT buffer C in the salt washes: 10%, 20%, 30%, 40%, 50%, 60%, 70%, 80%, 90%, and 100%. For all gradient steps, the specified buffer is mixed with MudPIT buffer A. When two concentrations are given for a solution (e.g., 10% to 45% MudPIT buffer B), it should be assumed that a linear ramp should be programmed over the time given.
11. Start the mass spectrometer and HPLC pumps. For the LCQ, the Sequence Setup window records the file name, the directory in which to record the mass spectrometry data, and the instrument method to use during the run (i.e., Basic Protocol steps 9 and 10). Once everything is programmed and the MudPIT column is positioned optimally in front of the entrance to the mass spectrometer, the run is started.
12. Upon completion of the run, process the acquired MS/MS data as described in UNIT 16.10. CONSTRUCTING FRITTED CAPILLARY HPLC COLUMNS Two procedures are given in this unit for making either a fritted or fritless (see Support Protocol 2) microcapillary column from fused-silica capillary (FSC). For the successful ESI LC-MS/MS analysis of low femtomole (nanogram) amounts of peptides, microcapillary HPLC is required. The columns are later packed (see Support Protocol 3) with chromatography media for fractionating peptides. This protocol describes construction of a fritted capillary column containing a porous ceramic frit at one end of the FSC (Cortes et al., 1987). A pneumatic loading device or “packing bomb” is used to wash the inside of the microcapillary column and condition the frit (see Fig. 23.1.4). These columns will be used for off-line SCX fractionation and RP-LC-ESI-MS. For LC-ESI-MS, an emitter needle is later connected to the fritted column using a zero dead volume (ZDV) union.
SUPPORT PROTOCOL 1
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Materials Methanol High-pressure helium Potassium silicate (KASIL #1; The PQ Corporation) Formamide 1 M ammonium nitrate 1 M HCl Acetonitrile 75-µm-i.d. × 365-µm-o.d. fused-silica capillary (FSC; Polymicro Technology) Quartz capillary tubing cleaving tool (Chromatography Research Supplies) Pneumatic loading vessel (James Hill Instruments) 1⁄ -in. to 0.4-mm Vespel ferrules (Chromatography Research Supplies) 16 Stereomicroscope 100°C heat block 1. Cut an ∼30-cm piece of 75-µm-i.d. × 365-µm-o.d. FSC with a capillary tubing cleaving tool. A longer or shorter length of capillary tubing may be desired depending on the column application.
2. Place a 1.5-ml microcentrifuge tube filled with methanol into a pneumatic loading device. Secure the lid by tightening the bolts that attach the lid to the base. The lid has a Swagelok fitting containing a vespel ferrule.
3. Feed the FSC down through the ferrule until the end reaches the bottom of the tube. Tighten the ferrule to secure the FSC (Fig. 23.1.4). Tighten the compression nut on the Swagelok fitting to compress the Vespel ferrule around the FSC (Fig. 23.1.4).
4. Apply pressure to the loading vessel by first setting the regulator on the high-pressure helium cylinder to 500 psi, and then opening the three-way valve. In this way, wash the inside of the capillary with 100% methanol for 3 to 5 sec. 5. Release the pressure to the vessel using the three-way valve. 6. Dry the inside of the column by repeating steps 4 and 5 without methanol in the loading vessel. This will blow helium through the column.
7. In order, place 170 µl potassium silicate (KASIL #1) and 30 µl formamide into a 0.5-ml microcentrifuge tube. Vortex 15 sec. Centrifuge for 1 min at 2000 × g, room temperature. Make sure to add the KASIL to the tube first. A small lucent precipitate should appear at the bottom of the tube.
8. Quickly dip one end of the capillary into the middle of the potassium silicate mixture. Avoid the upper and lower layers of the potassium silicate solution.
9. View the length of the frit using a stereomicroscope. The optimal frit is 2 to 3 mm in length. Longer frits create excessive back pressure.
10. Wipe off the outside of the capillary with a lint-free tissue (e.g., Kimwipe). Protein Analysis by Multidimensional Chromatography & Mass Spec
11. Bake the fritted capillary tip at 100°C in a heat block for a minimum of 3 hr. The fritted end of the column can be placed under a heating block set at 100°C. An oven set at 100°C can also be used to bake the frits.
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12. Following baking, check the frit length under the stereomicroscope. The frit should be ∼2 to 3 mm. Excessively long frits can be trimmed using a capillary tubing cleaving tool.
13. Using the pneumatic loading vessel, wash the frit for 30 sec with the following solutions as described (step 4): 1 M ammonium nitrate 1 M HCl Water Acetonitrile. Use a Teflon tube holder in the pneumatic loading device when washing with acetonitrile. Acetonitrile will dissolve most plastics, especially Plexiglas. A correctly constructed frit will form a steady stream of small droplets when pressurized during each wash step. If no droplets form, trim the frit with a capillary tubing cleavage tool. If a steady spray of liquid is observed when washing, discard the column. A defective frit can be cleaved from the FSC and the procedure repeated starting at step 2. If necessary, the end of the frit can be polished using 400-grit sandpaper to square the end. The porous ceramic fritted columns can be stored at room temperature without any special precautions until ready to pack with chromatography media (see Support Protocol 3).
CONSTRUCTING FRITLESS CAPILLARY HPLC COLUMNS This protocol describes construction of a fritless column containing a ∼3-µm orifice at the end of the fused-silica capillary using a laser micropipet puller. The restriction prevents the packing material from passing through the column. For ESI-LC-MS/MS, the integrated column and emitter needle are connected to an electrospray interface. An alternative approach for constructing an integrated microcapillary HPLC column and electrospray needle is described in UNIT 16.9.
SUPPORT PROTOCOL 2
Materials Methanol 100-µm-i.d. × 365-µm-o.d. fused-silica capillary (FSC; Polymicro Technology) Alcohol lamp P-2000 laser micropipet puller (Sutter Instruments) Quartz capillary tubing cleaving tool (Chromatography Research Supplies) 1. Cut a 50-cm piece of 100-µm-i.d. × 365-µm-o.d. fused-silica capillary (FSC). 2. Make a 1-in. window in the center of FSC by holding it over the flame from an alcohol lamp until the polyimide coating has been charred. Remove the charred material by carefully wiping the capillary with a lint-free tissue (e.g., Kimwipe) soaked in methanol. Once the polyimide coating is removed, the exposed quartz glass is extremely fragile.
3. Place the window portion of the capillary into a P-2000 laser micropipet puller. 4. Position the exposed capillary in the center of the mirrored chamber of the puller. 5. Align the fused silica in the grooves on each side of the mirror and secure it in place using the supplied small vises.
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6. Pull two columns from the FSC using the following, typical, four-step parameter setup for pulling ∼3-µm tips from a 100-µm-i.d. × 365-µm-o.d. as FSC a guide (all other values set to zero): Heat = 320 Heat = 310 Heat = 300 Heat = 290
Velocity = 40 Velocity = 30 Velocity = 25 Velocity = 20
Delay = 200 Delay = 200 Delay = 200 Delay = 200.
Fritless columns can be stored at room temperature until ready to pack with chromatography media (see Support Protocol 3). Avoid letting the tip strike a hard surface since the exposed quartz is extremely fragile. SUPPORT PROTOCOL 3
PACKING CAPILLARY HPLC COLUMNS This protocol describes packing of fritted or fritless FSC columns. For performing off-line SCX fractionation of peptide mixtures followed by RP-ESI-LC-MS/MS (see Basic Protocol 1), separate SCX and RP columns should be packed. For inline multidimensional separations (MudPIT; see Basic Protocol 2), a single biphasic SCX-RP column is constructed. A pneumatic loading vessel is used to pack the capillary HPLC columns (see Fig. 23.1.4). If the researcher prefers not to construct individual columns or purchase a laser puller, packed 1-D and 2-D fused-silica microcapillary columns, named PicoFrit and Biphasic PicoFrit, may be purchased from New Objective and other vendors. Materials Isopropanol Reverse-phase (RP) packing material (Perceptive Poros R-10 or Agilent Technologies 5-µm XDB-C18) Strong cation exchange (SCX) packing material (Whatman 5-µm Partisphere SCX) High-pressure helium gas 0.5% acetic acid Microstir bar 1.8-ml glass vial (Chromatography Research Supplies) Sonicator Pneumatic loading vessel (James Hill Instruments) 1⁄ -in. to 0.4-mm Vespel ferrules (Chromatography Research Supplies) 16 Empty fritted or fritless column (see Support Protocol 1 or 2) Stereomicroscope Prepare slurry 1. Place a microstir bar into a 1.8-ml glass vial. Add 0.6 ml isopropanol and 8 mg packing material to the glass vial. A thin or thick slurry will determine how quickly the column will pack. Separate vials of SCX and RP packing material are prepared. Methanol can be substituted for isopropanol when preparing the packing slurry.
2. Vortex to resuspend the packing material and sonicate 5 min to prevent aggregation of particles. Protein Analysis by Multidimensional Chromatography & Mass Spec
Load packing material in pneumatic loading vessel 3. Transfer the slurry to a pneumatic loading vessel and place the loading vessel on a stir plate. Turn on the stir plate to keep the packing material suspended. Secure the lid to the loading vessel by tightening the bolts that attach the lid to the base.
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4. Measuring from the frit end, place a mark on the empty fritted or fritless column showing the desired packing height. For off-line SCX, a fritted 75 × 365–ìm column is typically packed to a height of 6 cm with SCX material. For RP-LC-MS, a fritted 75 × 365–ìm column is packed to a height of 12 cm with Poros C18 material. For a biphasic MudPIT column, a 100 × 365–ìm fritless column is first packed with 10 cm of Agilent XDB-C18 5-ìm RP material followed by 4 cm of 5-ìm SCX material.
5. Feed the FSC down through the vespel ferrule in the Swagelok fitting on the lid of a pneumatic loading vessel until the end reaches the bottom of the tube. Pull the column so that the capillary rests just off the bottom of the glass vial and stir bar and tighten the ferrule to secure the FSC. Pack column 6. Apply pressure to the loading vessel by first setting the regulator on the high-pressure helium gas cylinder to 500 to 1000 psi, then opening the three-way valve. 7. Place the fritted end of a column under the stereomicroscope to observe the packing. A steady stream of packing material should be seen flowing into the capillary. For fritless columns, the tip may need to be opened slightly when packing the column. To do this, touch the tip to a hard surface.
8. When the column has been packed to the mark, slowly turn off the pressure to the vessel at the three-way valve. For the individual SCX and RP capillary columns, proceed to step 12. Slowly releasing the pressure prevents the packing material from unpacking.
Prepare biphasic column 9. For the biphasic column, replace the RP slurry in the loading vessel with a Partisphere SCX packing material slurry. 10. Reinstall the RP-packed column into the pneumatic loading vessel (step 5). 11. Pressurize the loading vessel to 1000 psi and continue packing the biphasic column with 4 cm of SCX material as described (steps 6 to 8). Wash and store column 12. Replace the tube containing the slurry with a 1.5-ml microcentrifuge tube filled with 0.5% acetic acid. 13. Wash the column for 10 min using the loading vessel. Store the column in 0.5% acetic acid until ready to use. The column must be completely submerged in 0.5% acetic acid to prevent the packing resin from drying out. Columns can be stored indefinitely at room temperature in 0.5% acetic acid.
14. Before using for any biological samples, run a blank HPLC gradient across the column to condition it. Conditioning the column is important to firmly pack the resin. Multiple blank HPLC gradients may be required to obtain a reproducible baseline. If high sensitivity applications are planned, load 0.5 pmol of angiotensin peptide onto the new column and run an HPLC gradient. Angiotensin binds to nonspecific binding sites in the column and minimizes nonspecific, irreversible binding of sample peptides. Non-Gel-Based Proteome Analysis
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Angiotensin solution 0.1 pmol/µl human angiotensin I 0.5% acetic acid Prepare fresh each time MudPIT buffer A 5% acetonitrile 0.02% heptafluorobutyric acid (HFBA) Store up to 1 month at room temperature MudPIT buffer B 80% acetonitrile 0.02% HFBA Store up to 1 month at room temperature MudPIT buffer C 250 mM ammonium acetate 5% acetonitrile 0.02% HFBA Store up to 1 month at room temperature MudPIT buffer D 500 mM ammonium acetate 5% acetonitrile 0.02% HFBA Store up to 1 month at room temperature SCX buffer A 0.5% acetic acid 2% acetonitrile Store up to 1 month at room temperature SCX buffer B 250 mM ammonium acetate 0.5% acetic acid 2% acetonitrile Store up to 1 month at room temperature SCX buffer C 0.5 M KCl 0.5% acetic acid 2% acetonitrile Store up to 1 month at room temperature SCX buffer D 1 M NaCl 0.5% acetic acid 2% acetonitrile Store up to 1 month at room temperature Protein Analysis by Multidimensional Chromatography & Mass Spec
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COMMENTARY Background Information Chromatography combinations Multidimensional peptide separation typically exploits two or more independent physical properties to fractionate a peptide mixture into individual components. Separation methods that rely on independent physical properties are considered orthogonal. Physical properties of peptides typically exploited include size, charge, hydrophobicity, and biological interaction or affinity. Different types of chromatography may be used as long as they are largely independent and the components resolved in one dimension remain resolved in the second. Components that are not separated in the first separation step are typically resolved in the second dimension. Various combinations of multidimensional peptide separation approaches coupled with mass spectrometry have been published, including size-exclusion/reversed-phase chromatography (Opiteck et al., 1997), affinity/reversed-phase chromatography (Geng et al., 2000; Spahr et al., 2000), immobilized metal-affinity/reversed phase chromatography (Ficarro et al., 2002), anionexchange/reversed phase chromatography (Holland et al., 1995), cation-exchange/reversed phase chromatography (Link et al., 1999; Davis et al., 2001; Wolters et al., 2001; Gygi et al. 2002), and various chromatography separations with capillary electrophoresis (Moore et al., 1996; Tong et al., 1999; Cao et al., 2000). For identifying proteins, the combination of strong cation exchange (SCX) and reversed-phase chromatography, coupled with mass spectrometry, has proven to be the most successful. Peak capacity Peak capacity is the number of individual components that can be resolved by a separation method. Multidimensional separations are used to dramatically increase the peak capacity when any single parameter does not provide the required resolution and capacity. Giddings provides a mathematical model showing that, if the multidimensional separations are orthogonal, then the total peak capacity is the product of the individual peak capacities of each separation (Giddings, 1987). For the MudPIT approach described in this unit, the peak capacity has been estimated to be >20,000 (Wolters et al., 2001).
Strong cation exchange chromatography In this unit, SCX is used to resolve peptides in the first dimension. At pH <3, negative charges at the carboxyl groups and the C terminus are neutralized by complete protonation. This leaves the arginine, lysine, and histidine residues, plus the N terminus to contribute to a net positive charge of the peptide. The fully protonated peptides are then fractionated by SCX chromatography. Although the major property governing peptide retention during ion-exchange chromatography involves ionic interactions, most ion-exchange resins also exert some hydrophobic influence. This is commonly referred to as a mixed-mode effect. The mixed-mode effect partially explains why peptides with the same number of positively charged residues can often be resolved by ionexchange chromatography. Counter ions: To elute peptides from SCX columns, a counterion is typically used. While potassium chloride (KCl) is routinely used to elute peptides from SCX beds, this salt is incompatible with ESI-MS and must be removed before starting the RP gradient and ESIMS/MS. Unfortunately, high concentrations of KCl are notoriously difficult to remove completely by washing. Instead, ammonium acetate or other volatile salts can be used to elute peptides from the SCX column. These volatile salts are more compatible with ESI and the elution results are similar to KCl. Ion-pairing reagents: To improve RP chromatographic performance, ion-pairing reagents are required. While the ion-pairing reagent TFA is routinely used for RP-HPLC of peptides with UV detection, TFA suppresses ionization during ESI. Although producing broader chromatographic peaks, weak acids, such as acetic and formic acid, are used as ion-pairing reagents to minimize ion suppression. More recently, heptafluorobutyric acid (HFBA) has been substituted for these weak acids. HFBA showed an improved combination of the peak sharpness and minimal ion suppression (Wolters et al., 2001). Load capacity: Load capacity is defined as the maximum amount of material that can be loaded in a separation while maintaining chromatographic resolution. Multidimensional separations are used to increase the load capacity when analyzing low-abundance or trace components in a peptide mixture. Large amounts of peptides that would typically overwhelm the resolution and capacity of a single
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RP column can be loaded and fractionated by SCX. For the capillary columns described in this unit, up to 400 µg of complex peptides have been loaded and fractionated using SCX chromatography (Washburn et al., 2001). For offline SCX systems, milligram amounts of peptides can be fractionated and later analyzed by RP-LC-MS/MS (Gygi et al., 2002). The increase in loading capacity and peak capacity coupled with high sensitivity mass spectrometry enables low-abundance proteins to be identified in complex protein samples (Washburn et al., 2001; Sanders et al., 2002). Electrospray ionization-mass spectrometry The sensitivity of ESI-MS is inversely proportional to the flow rate. The use of low flow rates (<0.5 µl/min) in capillary RP-HPLC increases the sensitivity of detection by orders of magnitude compared to standard RP-HPLC columns using flow rates of ≥50 µl/min. For successful multidimensional analysis of low femtomole (nanogram) amounts of peptides, capillary RP-HPLC coupled to ESI-mass spectrometry is required. DALPC or MudPIT avoids several of the drawbacks inherent when gel electrophoresis and mass spectrometry are coupled for the separation and identification of proteins in complex mixtures. These limitations include limited fractionation ranges, protein insolubility, and limited recovery of material. Multidimensional chromatography combined with sophisticated data-dependent mass spectrometry data acquisition methods (e.g., dynamic exclusion) have increased the dynamic range of DALPC and MudPIT for detecting low-abundance proteins in complex mixtures. As mass spectrometry has the sensitivity to identify proteins in quantities undetectable by silver staining, direct analysis of protein complexes enables these rare proteins to be reproducibly identified.
Protein Analysis by Multidimensional Chromatography & Mass Spec
HPLC columns This unit describes two approaches for constructing capillary HPLC columns. The porous ceramic fritted columns are fabricated from potassium silicate solution. The porosity of these frits is estimated to be between 3000 and 5000 Å (Cortes et al., 1987). These frits have proven to be simple and reproducible to make. In addition, they are exceedingly rugged and minimize band spreading. The fritted columns tend to be less prone to clogging compared to the fritless columns. Attaching an emitter tip enables these columns to be used for high sensitivity ESI-LC-MS. For low-flow systems,
emitter tips with <10-µm-i.d. openings can be attached to produce a stable electrospray. The emitter tip is required for a stable electrospray using the low flow rates in this unit. If the emitter tip clogs during sample loading or MS analysis, the removable tip can simply be replaced without having to make a new column and loss of precious samples. The fritless columns use a narrowly constricted orifice as a frit. The tip diameter must be small enough to restrict the packing material from passing through. This simple one-piece design minimizes dead volume and peak broadening. Because of the small orifice of the tip, buffers and samples must be free of insoluble particles that can clog the tip. When packing the column, it may be necessary to lightly touch the tip to a hard surface to open the end slightly. Because of the one-piece design, when the tip of the fritless column becomes clogged, a new column typically must be used. To reproducibly make the uniform shaped tip of the fritless columns, a laser micropipet puller is essential. Since the reversed-phase gradients expose the SCX phase of the MudPIT column to high concentrations of organic solvents, the mixedmode effects in the SCX phase are minimized. SCX/RP-ESI-MS/MS Off-line desalting and SCX fractionation combined with automated RP-ESI-MS/MS is a robust and reproducible combination for analyzing a variety of samples. Dilute peptide mixtures can be concentrated and desalted using the RP trapping cartridges with a minimal loss of material. The trapping cartridges are less prone to clogging compared to the capillary columns and serve to filter particulates. Using the loading loop procedure described in this unit, the off-line SCX capillary columns can be robustly and rapidly loaded with a minimum of sample loss. In addition, this approach can be used to directly analyze peptides present in detergents that would be incompatible with a RP-LC-ESI-MS. For example, proteins in 0.2% NP-40 detergent-containing buffer have been successfully identified (Link, unpub. observ.). The nonionic detergents can be effectively washed away during SCX fractionation. Comprehensive systems A multidimensional system is considered comprehensive when the entire eluant from the first dimension separation is transferred to the second dimension. The MudPIT protocol is a comprehensive multidimensional fractionation. Using the biphasic SCX-RP column, all
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the peptides eluted from a SCX fraction are transferred to the RP section of the column. The comprehensive nature of MudPIT makes it a highly sensitive approach for identifying proteins in complex mixtures.
Critical Parameters When making the fritted capillary columns, the length of the frit should be minimized for optimal performance. Excessively long frits produce high flow resistance. If the back-pressure is excessive, the frit can be trimmed using a capillary tubing cleaving tool to increase the flow rate. For the fritless columns, all the buffers and samples should be filtered or centrifuged to remove insoluble particles that can clog the tip. Complete desalting of peptides mixtures is essential for binding of peptides to the SCX column. The buffer components typically used for trypsin digestion of proteins can severely reduce the binding of peptides to the SCX resin. Routine use of a trapping cartridge is recommended for desalting peptides and filtering the samples. The SCX-RP multidimensional fractionation and identification system tends to be biased against low molecular weight (<10 kDa) and acidic proteins. Typically, low molecular weight proteins generate fewer tryptic peptides in the m/z ratio range of the mass spectrometer compared to higher molecular weight proteins. Acidic proteins have fewer lysine and arginine residues and consequently generate fewer tryptic peptides. Coupling multidimensional peptide fractionation with tandem mass spectrometry can routinely generate tens of thousands of tandem spectra. To identify the proteins or post-translationally modified peptides, each spectra typically needs to be searched against a translated nucleotide sequence or protein database that can contain millions of characters. The time required for the routine processing of this much data on a single processor is prohibitive. Offline processing of data using multiple processors is typically necessary to correlate mass spectral data with the sequences in the sequence databases. In addition, the resulting collection of significantly correlating peptide sequences needs to be assembled into a list of likely proteins or modifications in the original sample.
Troubleshooting Successfully working with low flow rates and capillary columns requires practice and
finesse. The PEEK connections joining the fused-silica capillaries should be only finger tight. Excessive tightening of the connections will crush the PEEK sleeves and fused-silica capillary tubing. The FSC should be able to slide through the PEEK sleeve. If the FSC becomes tightly bound in the PEEK sleeve, replace the sleeve or the connection if necessary. The ends of the FSC should be square with no jagged edges, especially for the emitter tips. Jagged edges tend to break off and clog the emitter tips or fritless columns. A quartz capillary tubing cutting tool should be used to produce a clean, square cut at the ends of the FSC. With the assemblies properly connected and the HPLC pump on, a small steadily increasing droplet should form at the end of the capillary or emitter tip. If no flow is detected, check the PEEK connections. For the fritted RP columns, remove the emitter tip and check the flow through the column. Replacing the emitter tip can usually resolve the problem. If the fritted SCX/RP or biphasic MudPIT column is not flowing and all the connections have been checked, it is likely the column is clogged and needs to be replaced. When working with capillary columns, solid particles will plug the columns. It is essential that all samples be centrifuged prior to loading onto the capillary columns. UNIT 16.9 describes additional precautions and troubleshooting tips for working with RP capillary columns and ESI mass spectrometry. With the ESI voltage on, the droplet at the end of the fritted RP and MudPIT column should disappear. If the droplet disappears, ions are being formed. Failure to detect ions is typically caused by the emitter tip not being aligned properly with the capillary opening to the mass spectrometer. A closed circuit television with magnifying optics is typically used to monitor the alignment of the emitter tip with the capillary opening of the mass spectrometer. Continued failure to detect ions can be caused by improper ESI tune setting or problems with the mass analyzer. If the droplet on the emitter tip fails to disappear when the ESI is on, the emitter tip may be broken. The integrity of the tip can be checked using a stereomicroscope. For the fritted RP columns, a new tip can be used to replace the broken or clogged tip. The electrical connections should be checked if the tip and flow rate appear normal. The front surfaces of the mass spectrometer capillary opening should be cleaned to assure
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the potential difference between the emitter tip and capillary is maintained. For high sensitivity analysis of samples, the mass spectrometer’s capillary opening and ionization optics should be clean and operating parameters optimized. The mass spectrometer’s ion optics should be tuned and calibrated according to the manufacturer’s recommendations. Routine cleaning and tuning of the ionization optics is necessary to maintain high sensitivity. Before analyzing experimental samples, the performance of the chromatography and mass spectrometry systems is typically tested using a peptide standard (e.g., angiotensin) to assure conditions are optimal and all systems are functioning properly.
Anticipated Results The two approaches described in this unit should yield large numbers of high quality tandem mass spectra from complex biological protein samples. Using capillary columns with their low flow rates allows the two approaches to identify proteins in the low femtomole range. Published studies have used these protocols to identify >100 proteins from purified protein complexes and >1000 proteins starting from whole-cell lysates (Washburn et al., 2001; Sanders et al., 2002).
Time Considerations Because of the number of SCX fractions and long RP gradient times to resolve complex peptide mixtures, DALPC and MudPIT experiments are relatively long processes. When performing off-line SCX fractionations, expect 3 to 4 hr of time to desalt and SCX fractionate the peptide mixture. Using the 90-min gradients in the first method (see Basic Protocol 1), the automated RP-LC-ESI-MS/MS analysis of 12 SCX fractions requires ∼20 hr. The 15-step MudPIT protocol requires ∼30 hr to complete. For both protocols, the run times can be shortened or lengthened, depending on the number of SCX fractionation steps and the length of the RP gradients.
Literature Cited Cao, P. and Stults, J.T. 2000. Mapping the phosphorylation sites of proteins using on-line immobilized metal affinity chromatography/capillary electrophoresis/electrospray ionization multiple stage tandem mass spectrometry. Rapid Commun. Mass. Spectrom. 14:1600-1606.
Cortes, H.J., Pfeiffer, C.D., Richter, B.E., and Stevens, T.C. 1987. Porous ceramic bed supports for fused silica packed capillary columns used in liquid chromatography. J. High Res. Chrom. 10:446-448. Davis, M.T., Beierle, J., Bures, E.T., McGinley, M.D., Mort, J., Robinson, J.H., Spahr, C.S., Yu, W., Luethy, R., and Patterson, S.D. 2001. Automated LC-LC-MS-MS platform using binary ion-exchange and gradient reversed-phase chromatography for improved proteomic analyses. J. Chromatogr. B. Biomed. Appl. 752:281-291. Ficarro, S.B., McCleland, M.L., Stukenberg, P.T., Burke, D.J., Ross, M.M., Shabanowitz, J., Hunt, D.F., and White, F.M. 2002. Phosphoproteome analysis by mass spectrometry and its application to Saccharomyces cerevisiae. Nat. Biotechnol. 20:301-305. Geng, M., Ji, J., and Regnier, F.E. 2000. Signaturepeptide approach to detecting proteins in complex mixtures. J. Chromatogr. A 870:295-313. Giddings, J.C. 1987. Concepts and comparisons in multidimensional separation. J. High Res. Chrom. 10:319-323. Gygi, S.P., Rist, B., Griffin, T.J., Eng, J., and Aebersold, R. 2002. Proteome analysis of low-abundance proteins using multidimensional chromatography and isotope-coded affinity tags. J. Proteome Res. 1:47-54. Holland, L.A. and Jorgenson, J.W. 1995. Separation of nanoliter samples of biological amines by a comprehensive two-dimensional microcolumn liquid chromatography system. Anal. Chem. 67:3275-3283. Licklider, L.J., Thoreen, C.C., Peng, J., and Gygi, S.P. 2002. Automation of nanoscale microcapillary liquid chromatography-tandem mass spectrometry with a vented column. Anal. Chem. 74:3076-3083. Link, A.J., Eng, J., Schieltz, D.M., Carmack, E., Mize, G.J., Morris, D.R., Garvik, B.M., and Yates, J.R. III 1999. Direct analysis of protein complexes using mass spectrometry. Nat. Biotechnol. 17:676-682. Moore, A.W. Jr., Larmann, J.P. Jr., Lemmo, A.V., and Jorgenson, J.W. 1996. Two-dimensional liquid chromatography-capillary electrophoresis techniques for analysis of proteins and peptides. Methods Enzymol. 270:401-419. Opiteck, G.J. and Jorgenson, J.W. 1997. Two-dimensional SEC/RPLC coupled to mass spectrometry for the analysis of peptides. Anal. Chem. 69:2283-2291. Sanders, S.L., Jennings, J., Canutescu, A., Link, A.J., and Weil, P.A. 2002. Proteomics of the eukaryotic transcription machinery: Identification of proteins associated with components of yeast TFIID by multidimensional mass spectrometry. Mol. Cell. Biol. 22:4723-4738.
Protein Analysis by Multidimensional Chromatography & Mass Spec
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Spahr, C.S., Susin, S.A., Bures, E.J., Robinson, J.H., Davis, M.T., McGinley, M.D., Kroemer, G., and Patterson, S.D. 2000. Simplification of complex peptide mixtures for proteomic analysis: Reversible biotinylation of cysteinyl peptides. Electrophoresis 21:1635-1650. Tong, W., Link, A., Eng, J.K., and Yates, J.R. III 1999. Identification of proteins in complexes by solid-phase microextraction/multistep elution/capillary electrophoresis/tandem mass spectrometry. Anal. Chem. 71:2270-2278. Washburn, M.P., Wolters, D., and Yates, J.R. III. 2001. Large-scale analysis of the yeast proteome by multidimensional protein identification technology. Nat. Biotechnol. 19:242-247. Wolters, D.A., Washburn, M.P., and Yates, J.R. III 2001. An automated multidimensional protein identification technology for shotgun proteomics. Anal. Chem. 73:5683-5690.
Key References Kinter, M. and Sherman, N.E. 2000. Protein Sequencing and Identification Using Tandem Mass Spectrometry. Wiley-Interscience, New York.
This is an excellent, easy-to-read tutorial on mass spectrometry for identifying proteins and posttranslational modifications. Many of the fundamental applications in this unit are described in detail from the view of the bench scientist. Link, et. al., 1999. See above. This publication demonstrates the original biphasic MudPIT approach and the use of multidimensional technology to directly analyze protein complexes. Wolters, et. al., 2001. See above. This technical publication optimizes and automates MudPIT for analysis of complex peptide mixtures.
Contributed by Andrew J. Link and Jennifer L. Jennings Vanderbilt University School of Medicine Nashville, Tennessee Michael P. Washburn Torrey Mesa Research Institute San Diego, California
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Quantitative Protein Profile Comparisons Using the Isotope-Coded Affinity Tag Method
UNIT 23.2
The quantitative analysis of protein profiles using the isotope-coded affinity tag (ICAT) methodology enables pairwise comparison of protein expression in biological samples, such as cell and tissue extracts and biological fluids. The measurement of relative changes in protein expression is achieved by selective in vitro alkylation of proteins at the amino acid cysteine with either an isotopically heavy reagent containing 13C isotopes (i.e., the 13 C-ICAT) or an isotopically normal reagent with no 13C isotopes (i.e., the 12C-ICAT), which is 9 Da lighter (Basic Protocol 1). As illustrated in Figure 23.2.1, the two labeled protein mixtures are mixed and digested with trypsin, creating a mixture of peptides, most of which are not labeled with the ICAT reagent. To reduce complexity at the point of microcapillary high-performance liquid chromatography (HPLC) separation and tandem mass spectrometry, the peptide mixture is fractionated by strong cation-exchange (SCX) chromatography (Basic Protocol 2). The complexity of each SCX fraction is further reduced by approximately an order of magnitude by selective isolation of ICAT-labeled peptides on a monomeric avidin column (the ICAT label contains a biotin group, but the actual structure is proprietary), allowing unlabeled peptides to pass through (Basic Protocol 3). After a washing step, the ICAT-labeled peptides are released from the column by a low–trifluoroacetic acid (TFA) concentration wash that disrupts the avidin-biotin interaction. Finally, the biotin tag is removed from the peptides by a high–TFA concentration incubation (Basic Protocol 4). The retentate from each avidin affinity column, which contains predominantly ICATlabeled peptides, is then separated by reversed-phase microcapillary HPLC (RP-µLC), a procedure that is not described here. This separation may be conducted on-line with direct feed to an electrospray ionization tandem mass spectrometer or directly onto a MALDI plate for subsequent analysis. For example, ICAT-labeled peptide tandem mass spectra can be matched to peptide amino acid sequence using the SEQUEST algorithm and relative quantitation can be performed using XPRESS software (see Internet Resources). Applied Biosystems offers all reagents and accessories necessary to perform an ICAT experiment (i.e., Cleavable ICAT Reagent Kit for Protein Labeling, Monoplex Version; see Internet Resources). LABELING PROTEINS WITH ACID-CLEAVABLE ISOTOPE-CODED AFFINITY TAG REAGENT
BASIC PROTOCOL 1
This procedure describes solubilization of proteins in isotope-coded affinity tag (ICAT) labeling buffer. If proteins are in a solution that is incompatible with labeling buffer ingredients, such as >0.5% (w/v) SDS or other detergents (see Critical Parameters), then the samples must first be cleaned up by trichloroacetic acid (TCA) precipitation (UNIT 3.4) or other compatible methods. The resulting pellet is resolubilized in the appropriate labeling buffer. It is sometimes necessary to first attempt solubilization of the pellet in a small volume of labeling buffer that contains a relatively high (1% to 2% w/v) concentration of SDS. Once the pellet is solubilized, the solution can be diluted to reduce the SDS concentration. The following protocol is based on labeling 500 µg protein with each of the two ICAT reagents. If an amount other than 500 µg is used, the volume of the labeling buffer and the stoichiometric ratio of ICAT reagent can be adjusted proportionally. Non-Gel-Based Proteome Analysis Contributed by Eugene C. Yi and David R. Goodlett Current Protocols in Protein Science (2003) 23.2.1-23.2.11 Copyright © 2003 by John Wiley & Sons, Inc.
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denature protein and reduce free sulfhydryls 12C-ICAT
13C-ICAT
(light)
(heavy)
label with light or heavy ICAT reagent
combine
tryptic digestion
cation-exchange factionation 50c
50c
50c
50c
50c
50c
50c
avidin affinity purification acid treatment to remove biotin tag
multidimensional purification
Figure 23.2.1 Flow chart of the quantitative analysis of protein profiles with cleavable isotopecoded affinity tag (ICAT) reagent.
Materials Protein pellets, dried, from control and test samples Labeling buffer: 0.05% (w/v) SDS/50 mM Tris⋅Cl, pH 8.3/5 mM EDTA, pH 8.0/6 M urea; prepare fresh from stock solutions (APPENDIX 2E) Bovine IgG 250 mM Tris(2-carboxyethyl) phosphine (TCEP) 50 mM Tris⋅Cl, pH 8.3 (APPENDIX 2E) Cleavable ICAT reagents (heavy and light; Applied Biosystems) 1 M dithiothreitol (DTT; APPENDIX 2E) Sequencing-grade trypsin (Promega) 6 N acetic acid Tube rocker (e.g., Baxter Scientific Products)
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Additional reagents and equipment for determining protein concentration (UNIT 3.4), SDS–polyacrylamide gel electrophoresis (SDS-PAGE; UNIT 10.1), and silver staining (UNIT 10.5)
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1. Solubilize dried protein pellets from control and test samples in two separate microcentrifuge tubes using ≤100 µl each freshly prepared labeling buffer and agitate. If the pellet does not fully solubilize upon agitation, the samples should be sonicated in a water bath sonicator until the pellet dissolves. The water bath should be kept cool (<37°C) while sonicating. To maintain an effective concentration of ICAT reagent, it is highly recommended that the sample volume be kept at ∼100 ìl.
2. Use a small aliquot of solubilized sample to separately determine the amount of protein in the control and test samples using bovine IgG as a standard protein (UNIT 3.4). The authors routinely use the Bio-Rad Protein Assay kit.
3. Transfer a volume of each sample containing 500 µg protein to a fresh tube and add labeling buffer to 90 µl. Save an aliquot (∼1 µg) from each sample to monitor the labeling efficiency by SDS-PAGE (step 9). 4. Add 2 µl of 250 mM TCEP (5 mM final) to the 500-µg protein samples. Check the pH of both solutions and adjust to 8.3 with 50 mM Tris⋅Cl, if needed. TCEP is water soluble and less toxic than tributylphosphine (TBP). It is therefore more user friendly than TBP. More importantly, TCEP adducts do not interfere with selection of ICAT-labeled peptides for collision-induced dissociation (CID) because of their high relative hydrophilicity. It is, however, acidic, so the sample solution requires careful buffering to retain a pH of ∼8.3. TCEP can drastically change the labeling buffer pH and can directly react with the ICAT reagent if present in excessively high concentrations. To counter these potential problems, TCEP should be used at a relatively low concentration (1 to 5 mM), and the pH of the solution should be checked and adjusted as needed.
5. Add ∼5-fold molar excess of cleavable ICAT reagent over free sulfhydryls to each vial, using the light reagent for one sample and the heavy reagent for the other, to achieve a final concentration of 3.0 mM in 100 µl sample solution. Mix well and gently shake suspensions using a tube rocker for 90 min at room temperature in the dark. Molar amount of cysteines in samples can be estimated as follows: a) Assume the average molecular weight of the proteins in a sample is 50 kDa. b) Determine the molar amount of proteins using the protein concentration (i.e., 500 ìg) and the average molecular weight of proteins (i.e., 50 kDa): 0.5 × 10−3 g/50,000 g/mol = 10 × 10−9 mol protein c) Assume an average number of cysteines per protein (e.g., six cysteines per protein) and calculate the molar amount of cysteines: 60 × 10−9 mol cysteine/100 ìl = 0.6 mM d) For a 5-fold molar excess (or 3 mM) of ICAT reagent, use two 100-ìg tubes of ICAT reagent (each with 175 nmol ICAT reagent) for 3.5 mM final. This slightly higher ICAT concentration is acceptable, as long as sufficient DTT is added (step 6).
6. Quench the reactions by adding DTT from a 1 M stock solution to yield a 10-fold molar excess of DTT with respect to the ICAT reagent. Mix and incubate 5 min at room temperature. Save an aliquot (∼1 µg) of each sample to monitor labeling efficiency. For example, add DTT to 35 mM final concentration if the ICAT reagent concentration is 3.5 mM Failure to add a sufficient concentration of DTT could cause unwanted labeling after the two ICAT-labeled samples are combined. The ICAT-labeled protein mixtures can be stored at −80°C for up to a few months.
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7. Combine the labeled control and test samples and dilute the combined sample with 50 mM Tris⋅Cl, pH 8.3, until urea (from labeling buffer) is at a final concentration of 1 M. 8. Add 1:100 (w/w) sequencing-grade trypsin/protein and incubate overnight at 37°C. Quench reaction by adding 6 N acetic acid to pH 3.0. Save an aliquot (∼1 µg) of each sample to monitor completeness of tryptic digestion. 9. Analyze ICAT labeling efficiency and tryptic digestion by SDS-PAGE of the saved aliquots from steps 3, 6, and 8 followed by silver staining (Fig. 23.2.2). The trypsinized ICAT-labeled peptide mixtures can be stored at −80°C for a few months; for an optional fractionation by strong cation-exchange chromatography (see Basic Protocol 2), they should be used immediately . With the described labeling conditions, ICAT labeling of cysteinyl residues is complete within 1 hr. It is important, however, to evaluate the labeling efficiency. Analysis with a 12.5% (w/v) SDS–polyacrylamide gel is suitable for evaluating the labeling efficiency and the trypsinization of the protein mixtures. As shown in Figure 23.2.2, slower electrophoretic mobility of ICAT-labeled proteins that contain several cysteines (lanes 3 and 4) can be observed. Proteins that are lower in abundance or that have few cysteines may not, however, show an appreciable shift on a gel. After trypsinization, the digested ICAT-labeled protein mixtures can be detected at the bottom of the gel (lane 5). BASIC PROTOCOL 2
PURIFICATION AND FRACTIONATION OF PEPTIDES LABELED WITH ISOTOPE-CODED AFFINITY TAGS BY STRONG CATION-EXCHANGE CHROMATOGRAPHY This procedure describes the cation-exchange purification and fractionation of trypsinized peptide mixtures. For less complex samples or where limited protein is available (e.g., ∼100 µg total protein), use of the cation-exchange cartridge found in the isotope-coded affinity tag (ICAT) Reagent Kit (Applied Biosystems) to prepare a single fraction for avidin affinity purification should be considered. For highly complex samples (i.e., 0.5 to 1 mg total protein), however, fractionation by strong cation-exchange (SCX) chromatography into 25 to 50 fractions is recommended. There is a trade-off between the number of SCX fractions collected, the amount of starting protein, and the resultant quality of peptide tandem mass spectra; too little protein (e.g., 100 µg) fractionated into too many fractions (e.g., 50) will yield poor results from mass spectrometry (MS) analysis.
1
Quantitative Protein Profile Comparisions Using ICAT
2
3
4
5
Figure 23.2.2 SDS-PAGE analysis (12.5%, w/v) of nonlabeled (lanes 1 and 2), isotope-coded affinity tag (ICAT)-labeled (lanes 3 and 4), and trypsinized (lane 5) protein mixtures. Proteins and peptides were detected by silver staining.
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Materials ICAT-labeled protein sample (see Basic Protocol 1) 3 N phosphoric acid Cation-exchange buffer B: 10 mM potassium phosphate/600 mM KCl/25% (v/v) acetonitrile, pH 3.0; store at room temperature Cation-exchange buffer A: 10 mM potassium phosphate/25% (v/v) acetonitrile, pH 3.0; store at room temperature 2.1 × 200–mm PolySULFOETHYL A column (PolyLC) High-performance liquid chromatography (HPLC) system with binary pump and fraction collector 1-ml glass vials with polyethylene snap caps (e.g., Waters Chromatography) 2-ml microcentrifuge tubes (e.g., Micro Tube; Sarstedt) 1. Deactivate and equilibrate a 2.1 × 200–mm PolySULFOETHYL A strong cation-exchange (SCX) column as recommended by the manufacturer and set up in an HPLC system. It is also good practice to run standard peptide mixtures to assess the column performance prior to running samples.
2. Acidify an ICAT-labeled sample with 3 N phosphoric acid to pH 3.0. Remove any precipitates by centrifugation for 10 min at 1400 × g, 37°C and load the clarified sample onto the SCX column. Acidification may precipitate salts and trypsin.
3. Elute ICAT-labeled peptides from the SCX column using a linear gradient of 0 to 100% cation-exchange buffer B in cation-exchange buffer A over 60 min and collect fractions in 1-ml glass vials placed in 2-ml microcentrifuge tubes at a suitable interval (e.g., 1 min). The length of the gradient and the sample collection size can be optimized to suit the sample size and complexity.
AFFINITY PURIFICATION WITH AVIDIN AFFINITY CARTRIDGES OF PEPTIDES LABELED WITH ISOTOPE-CODED AFFINITY TAGS In the isotope-coded affinity tag (ICAT) procedure, strong cation-exchange (SCX) fractionation is performed prior to avidin affinity purification to remove trypsin, which will digest avidin.
BASIC PROTOCOL 3
Materials Loading buffer: 2× PBS (APPENDIX 2E) Cation-exchange fractions (see Basic Protocol 2) Wash buffer 1: 1× PBS (APPENDIX 2E) Wash buffer 2: 50 mM ammonium bicarbonate, pH 8.3/20% (v/v) methanol Elution buffer: 0.4% (v/v) trifluoroacetic acid (TFA)/30% (v/v) acetonitrile Avidin cartridges and cartridge holders (Applied Biosystems) Ring stand for mounting cartridge 1000-µl gas-tight syringe with stainless steel needle (e.g., Hamilton) 1-ml glass vials with polyethylene snap caps (e.g., Waters Chromatography) NOTE: All buffers should be stored at 4°C for up to 3 months.
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1. Clean and activate an avidin cartridge as recommended by the manufacturer and place it in a cartridge holder supported by a ring stand. With appropriate care, the avidin cartridge can be reused up to 50 times. The cartridge can also be stored in PBS with 0.1% (w/v) sodium azide at 4°C for up to 3 months.
2. Use a 1000-µl gas-tight syringe with stainless steel needle to transfer 500 µl loading buffer to each cation-exchange fraction. Save an aliquot (∼1 µg) of neutralized sample for monitoring the avidin purification process. The loading buffer neutralizes the fractions to pH 7.2. The same syringe can be used for all buffers, but it must be washed thoroughly with water between the different solutions. The protein amount in each cation-exchange fraction can be estimated based on the number of fractions collected (see Basic Protocol 2) and the initial amount of protein in the sample.
3. Slowly load (∼1 drop/sec) a neutralized fraction (typically ∼1 ml) onto the avidin cartridge and collect and save the flowthrough in a 1-ml glass vial. 4. Add an additional 500 µl loading buffer and collect flowthrough in the same vial. Store the combined flowthrough at −80oC to monitor the avidin purification efficiency. Optionally, this flowthrough can be used to corroborate protein identifications based on tandem mass spectrometry (MS/MS) spectra of the ICAT-labeled peptides or to locate peptides with post-translational modifications not present on the ICAT-labeled peptides.
5. Inject 1 ml wash buffer 1. Discard the output. 6. Inject 1 ml wash buffer 2. Discard the output. 7. Fill the gas-tight syringe with 800 µl elution buffer and slowly inject (∼1 drop/sec) the first 50 µl of buffer. Discard the output. 8. Elute the labeled peptides (∼1 drop/sec) with the remaining 750 µl elution buffer and collect in a clean glass vial. 9. Clean and activate cartridge and repeat steps 3 to 8 for the remaining cation-exchange fractions. The manufacturer states that the avidin cartridge is good for purification of ≤50 cationexchange fractions. The avidin column efficiency can also be monitored by SDS-PAGE. The fractions can be stored up to 3 months at −80°C. BASIC PROTOCOL 4
CLEAVAGE OF THE BIOTIN AFFINITY TAG FROM PEPTIDES LABELED WITH ISOTOPE-CODED AFFINITY TAGS The cleavage procedure described here follows the protocol provided by Applied Biosystems as part of the isotope-coded affinity tag (ICAT) Reagent Kit. Materials Affinity-purified ICAT-labeled peptide fractions in glass vials (see Basic Protocol 3) Cleaving reagents A and B (Applied Biosystems) Appropriate solvent for microcapillary liquid chromatography–tandem mass spectrometry (µLC-MS/MS) analysis
Quantitative Protein Profile Comparisions Using ICAT
Speedvac evaporator (Savant) 1000-µl glass syringe with stainless steel needle (e.g., Hamilton) 1-ml glass vials with polyethylene snap caps (e.g., Waters Chromatography)
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1. Dry each affinity-purified ICAT-labeled peptide fraction completely in a Speedvac evaporator. In general, drying samples that are basic together with those that are acidic in the same evaporator should be avoided because salts will form in the sample and inside the Speedvac.
2. Prepare the cleaving reagent by using a 1000-µl glass syringe with stainless steel needle to combine 95:5 (v/v) cleaving reagent A/cleaving reagent B in a 1-ml glass vial. Vortex to mix. Be sure to use a glass vial and a glass syringe with stainless steel needle to transfer the cleaving reagent because the reagent contains a strong acid.
3. Transfer 100 µl cleaving reagent to each dried fraction in the glass vials. Cap vials and incubate 2 hr at 37°C. 4. Dry fractions completely in the Speedvac. 5. Add the appropriate solvent to the dried pellet. Vortex, centrifuge, carefully transfer to an appropriate tube, and store at −80°C until ready for µLC-MS/MS or up to 3 months. The sample is usually dissolved in LC/MS loading buffer, such as 0.1% (v/v) formic acid or 0.4% (v/v) acetic acid, in a 250-ìl microcentrifuge tube.
COMMENTARY Background Information The isotope-coded affinity tag (ICAT) method has enabled the rapid and comprehensive comparative characterization of complex protein samples (Gygi et al., 1999; Han et al., 2001; Baliga et al., 2002; Guina et al., 2003; Ranish et al., 2003). Compared with the traditional two-dimensional gel electrophoresis (2DE)-based approach to proteomics, the ICAT method allows direct analysis of hydrophobic membrane proteins. It also serves as a readily available screening tool for generating new hypotheses for testing by allowing simultaneous western blot of many proteins. Additionally, the ICAT method is easily amenable to high-throughput data generation and can dramatically increase the detection sensitivity of proteins both qualitatively and quantitatively (Goodlett and Aebersold, 2001; Flory et al., 2002; Goodlett and Yi, 2002). The following section is intended to orient the new user and further explain the method. The protocol described herein is specific to the second-generation cysteine-specific ICAT reagent. It consists of a biotin affinity tag; an acid-cleavable linker that allows removal of the biotin tag prior to mass spectrometry (MS) analysis; a mass tag linker, which has nine 13C molecules that replace nine 12C molecules in the heavy reagent (whereas no 12C molecules are replaced in the light reagent); and an io-
doacetamide-reactive group that specifically reacts with cysteinyl thiols. Stable isotopes are incorporated into proteins by selective alkylation of cysteines with the ICAT reagents under denaturing and reducing conditions. Peptides from the proteins in one sample that were labeled with the light ICAT reagent serve as the internal standards to normalize the data of the other group of proteins labeled with the heavy ICAT reagent. After the two ICAT-labeled protein mixtures are combined and the proteins are digested to peptides with trypsin (Basic Protocol 1), the protocols in this unit employ the standard ICAT process: pairwise comparison together with peptide separation by strong cation exchange (SCX), avidin affinity chromatography (Basic Protocol 3), removal of the biotin tag (Basic Protocol 4), and on-line reversed-phase microcapillary column chromatography (RP-µLC), which is not covered here. An optional fractionation technique can be performed to enrich low-abundance proteins or reduce the complexity of the mixture (Basic Protocol 2). Once the protein samples are combined, any protein losses that are due to processing the sample will be shared, and thus no bias will be introduced into the final ICAT ratio. In the standard ICAT process, all peptides are typically fractionated by SCX chromatography into 25 to 50 1-min fractions (i.e., that contain peptides of OD214
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Quantitative Protein Profile Comparisions Using ICAT
≥0.2) from an hour-long linear salt gradient, and each fraction then undergoes avidin affinity chromatography. Less-complex samples, such as samples with little protein (<100 µg) or samples from immunoprecipitation should not, however, be over-fractionated. These samples do require a simple one-step cleanup by SCX chromatography to remove excess trypsin, which will digest avidin. The peptide complexity of each fraction is further reduced by approximately an order of magnitude by selectively isolating ICAT-labeled peptides using a monomeric avidin cartridge. Note that peptides from the avidin affinity column flowthrough can be saved to later corroborate identifications made from single ICAT-labeled peptides. This sequential fractionation dramatically simplifies the complex peptide mixture and significantly enhances the detection and identification of low-abundance proteins in conjunction with automated, datadependent tandem mass spectrometry (MS/MS). Each fraction from the avidin affinity column separation is subjected to RP-µLC and is identified by automated data-dependent MS/MS using a strategy that involves a single MS survey scan or full scan followed by four MS/MS scans. Acquiring more than about five MS/MS scans prior to conducting the next survey scan, however, will lead to increased error in the measured ICAT ratio. The recent introduction of a second-generation ICAT reagent provides two specific advantages over the original reagent. These are: (1) the use of 13C instead 2H as the heavy replacement element in the ICAT reagent, which eliminates the small mobility shift during HPLC separation of ICAT-labeled peptides and allows for more accurate quantification, and (2) the use of an acid-cleavable linker to remove the bulky biotin group, which results in an improvement in the quality of peptide, MS/MS data and thus should result in more protein identifications. Subsequent quantitative data analysis can be performed using software such as XPRESS (see Internet Resources). XPRESS automates the process of quantitatively analyzing the elution profile of all identified peptides and comparing them against their ICAT partners. Other software packages currently under development at the Institute for Systems Biology will allow for a higher level of automated ICAT ratio analysis. Note also that most MS instrument manufacturers plan to have some sort of quantitative software available, but the quality and timing of release of these products is uncertain.
Critical Parameters Prior to ICAT labeling, it is necessary to remove salts and detergents that may interfere with the labeling reaction. Specifically, it is important to remove DTT because the sulfhydryl groups of the reagent may couple to the ICAT reagent. Tributylphosphine (TBP) or any nucelophilic chemicals that could compete with the labeling reaction should also be removed. In addition, to maintain an optimal SDS concentration of 0.5% (w/v) for the labeling reaction, it is convenient to remove SDS and reconstitute the proteins in a solution containing the desired SDS concentration. It is also necessary to accurately estimate the amount of protein in each sample to be labeled. Labeling solutions should be prepared fresh for each experiment from stock solutions. Note that urea will react with Tris buffer and thus may hinder subsequent labeling steps. Relatively high percentages of SDS (e.g., 1% to 2% [w/v] SDS) and agitation are often required to properly solubilize the protein pellets. One can then dilute the mixture with buffer to the optimal 0.5% (w/v) SDS concentration. To ensure complete ICAT labeling of the proteins and thus accurate relative abundances, it is imperative to estimate the number of free sulfhydryls in each sample prior to the labeling reaction. The number of free sulfhydryls can be estimated from an average molecular weight and an average number of cysteines per protein as described in Basic Protocol 1. Additionally, to ensure that the final ICAT ratios reflect the relative abundances of proteins in the original mixtures, it is important that the proteins be completely denatured. A combination of 6 M urea and 0.5% (w/v) SDS works well for this purpose. Tris(2-carboxyethyl) phosphine (TCEP), for several reasons not given here, is the best agent for reducing cysteine disulfides in this process. The excess ICAT reagent needed for high labeling efficiency can, however, lead to side reactions, such as the formation of TCEP-ICAT adducts (Smolka et al., 2001). To minimize this problem, use no more than the recommended amount of TCEP. Note also that TCEP can drastically change the pH of the labeling buffer; it is advisable to check the pH of the labeling mixture after adding TCEP. TBP is avoided because the product of the condensation reaction between the ICAT reagent and TBP elutes with ICAT-labeled peptides during RP-µLC. This reduces the number of actual ICAT-labeled peptide pairs that will be selected for collisioninduced dissociation (CID) because the ion
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selection routines cannot distinguish between peptide and nonpeptide contaminants. The ICAT reaction volume for 500 µg of protein sample should not exceed 100 µl to maintain effective concentration of the ICAT reagent for maximum labeling efficiency. Prior to tryptic digestion of the labeled proteins, the reaction mixture should be diluted 10-fold to 0.6 M urea and 0.05% (w/v) SDS to enable efficient trypsin activity prior to digestion. To avoid the introduction of plasticizers into the sample, a glass vial (and glass syringe equipped with a stainless steel needle) should be used whenever handling a strong acid, such as trifluoroacetic acid (TFA), during the sample preparation process. Glass vials are recommended for collecting fractions during the cation-exchange, avidin purification, and acidcleaving processes. Prior to cation-exchange fractionation, column performance should be checked with standard peptide mixtures. The length of the gradient for the ion exchange step and the sample collection size can be optimized according to the amount and complexity of the sample. The cation-exchange column and avidin car-
tridge should be properly cleaned and stored as recommended by the manufacturer.
Troubleshooting The process of comparative sample analysis can be divided into several critical steps for the purpose of monitoring reaction progress and sample recovery. First, ICAT labeling efficiency and tryptic digestion efficiency should be checked by SDS-PAGE using aliquots saved from each relevant step. Depending on the number of cysteines in a protein, a shift in gel mobility between labeled and unlabeled proteins should be observed if the proteins are properly labeled (Fig. 23.2.2). It is difficult, however, to visualize such shifts for those proteins that are low in abundance or that contain few cysteines. Second, the amount of peptide available for liquid chromatography–tandem mass spectrometry (LC-MS/MS) analysis can be estimated from the OD214 values produced during cation-exchange high-performance liquid chromatography (HPLC) separation (e.g., OD214 ≥ 0.5 from 500 µg total protein sample, with a 2.1 × 200–mm PolySULFOETHYL A column). If the OD214 is too low (e.g., ≤0.1), it is possible that the starting amount of protein
0.5
OD214
0.4 0.3 0.2 0.1 0.0
10
20
30
40
50
60
70
Fraction number
Figure 23.2.3 Fractionation by strong cation-exchange chromatography. Trypsinized peptide mixtures from 500 µg of total isotope-coded affinity tag (ICAT)-labeled protein sample were injected onto a 2.1 × 200–mm PolySULFOETHYL A column. The ICAT-labeled peptides were eluted using a linear gradient of 600 mM KCl from 0 to 100% B in 60 min, and fractions were collected in glass vials (placed in 2-ml microcentrifuge tubes) at 2-min intervals. The fraction number corresponds to the time (in minutes) of fraction collection.
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Light 1419.936 +1 AREA:9.79E+07
100 V 757.4
90 80
Relative abundance
70 60
1040 1050 1060 Heavy 1428.000 +1 AREA:1.16E+08
1070
1040 1050
1070
1080
1090
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# * 933.7 988.4 992.9
50 40
*
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*779.7
10
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* 1412.0
924.7 739.6
30
0 400
3.03E+07
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1047.9
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1200
1400
1600
1800
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Figure 23.2.4 Mass spectrometry (MS) scan of isotope-coded affinity tag (ICAT)-labeled peptides acquired by a Thermoquest LCQ ion trap mass spectrometer. Single- (v), double- (*), and triple- (#) charged ICAT peptide pairs show mass differences of 9.0, 4.5, and 3.3 Da, respectively, between heavy and light ICAT-labeled peptides. The inset figure is a MS scan trace of the indicated ICAT peptide pair obtained from the XPRESS quantification software program.
Quantitative Protein Profile Comparisions Using ICAT
was measured incorrectly or that most of the sample was lost during preparation. Finally, if ion signals are not in the satisfactory range of the MS, peptides might not have bound to the avidin affinity column. This can happen as a result of a failure of the avidin column or of an improper pH in the loading and elution buffers. The authors recommend saving all avidin column flowthrough (loading and washing buffer) to ensure that the sample can be recovered.
fractions of interest are processed. The relative abundance of a given ICAT-labeled peptide pair (i.e., the heavy to light ICAT ratio) can, in theory, vary over a few orders of magnitude. When analyzing whole-cell lysates, however, it is more common to observe ICAT ratios that vary from 1 to 100 with <20% error; generally, any ratio above two can be considered to be of interest, although this will depend on the biological system that is being studied.
Anticipated Results
Time Considerations
A typical result from ICAT analysis of 500 µg of total protein from a yeast lysate will generate 250 to 300 protein identifications per SCX-avidin fraction analyzed (i.e., this assumes injection of ∼30% of a fraction equivalent to OD214 ∼0.2 and base peak ion intensity between 5.0 × 108 and 1 × 109; see Figs. 23.2.3 and 23.2.4). Thus, the total number of protein identifications from the experiment will be close to 2000 or higher once the remaining
Typically, ICAT sample preparation requires 3 days. This includes the ICAT labeling, tryptic digestion, sample cleanup by cation-exchange cartridge, and avidin affinity chromatography purification. The optional cation-exchange fractionation followed by the avidin purification may take an additional day. Use of an automated multidimensional chromatography system (e.g., VISION; Applied Biosystems), which can carry out the SCX and avidin
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affinity column separations to generate 50 fractions in 24 hr, will save time. Details of the authors’ MS setup for analysis of all the fractions using an ion trap MS are found elsewhere (Baliga et al., 2002; Shiio et al., 2002; Yi et al., 2002; Guina et al., 2003; Ranish et al., 2003), but from start to finish, 3 to 5 days of continuous MS time is required to analyze all 50 fractions, depending on the type and length of HPLC gradient used.
Literature Cited Baliga, N.S., Pan, M., Goo, Y.A., Yi, E.C., Goodlett, D.R., Dimitrov, K., Shannon, P., Aebersold, R., Ng, W.V., and Hood, L. 2002. Coordinate regulation of energy transduction modules in Halobacterium sp. analyzed by a global systems approach. Proc. Natl. Acad. Sci. U.S.A. 99:1491314918. Flory, M.R., Griffin, T.J., Martin, D., and Aebersold, R. 2002. Advances in quantitative proteomics using stable isotope tags. Trends Biotech. 12:S23-S29. Goodlett, D.R. and Aebersold, R. 2001. Mass spectrometry in proteomics. Chem. Rev. 101:269295. Goodlett, D.R. and Yi, E.C. 2002. Proteomics without polyacrylamide: Qualitative and quantitative uses of tandem mass spectrometry in proteome analysis. Funct. Integr. Genomics. 2:138-153. Guina, T., Purvine, S.O., Yi, E.C., Eng, J., Goodlett, D.R., Aebersold, R., and Miller, S.I. 2003. Quantitative proteomic analysis indicates increased synthesis of a quinolone by Pseudomonas aeruginosa isolates from cystic fibrosis airways. Proc. Natl. Acad. Sci. U.S.A. 100:2771-2776. Gygi, S.P., Rist, B., Gerber, S.A., Turecek, F., Gelb, M., and Aebersold, R. 1999. Quantitative analysis of complex protein mixtures using isotope coded affinity tags. Nat. Biotechnol. 17:994-999.
Han, D.K., Eng, J., Zhou, H., and Aebersold, R. 2001. Quantitative profiling of differentiationinduced microsomal proteins using isotopecoded affinity tags and mass spectrometery. Nat. Biotechnol. 19:946-951. Ranish, J.A., Yi, E.C., Leslie, D.M., Purvine, S.O., Goodlett, D.R., Eng, J., and Aebersold, R. 2003. The study of macromolecular complexes by quantitative proteomics. Nat. Genet. 33:349355. Shiio, Y., Donohoe, S., Yi, E.C., Goodlett, D.R., Aebersold, R., and Eisenman, R.N. 2002. Quantitative proteomic analysis of Myc oncoprotein function. EMBO J. 21:5088-5096. Smolka, M.B., Zhou, H., Purkayastha, S., and Aebersold, R. 2001. Optimization of the isotopecoded affinity tag-labeling procedure for quantitative proteome analysis. Anal. Biochem. 297:25-31. Yi, E.C., Mrelli, M., Lee, H., Purvine, S.O., Aebersold, R., Aitchison, J.D., and Goodlett, D.R. 2002. Approaching complete peroxisome characterization by gas-phase fractionation. Electrophoresis 23:3205-3216.
Internet Resources http://www.proteomecenter.org/software.php Includes details of proteomics software tools from the Institute for Systems Biology, including the XPRESS software. http://www.appliedbiosystems.com/products/ productdetail.cfm?ID=153 Provides further information on Applied Biosystem’s ICAT reagents.
Contributed by Eugene C. Yi and David R. Goodlett Institute for Systems Biology Seattle, Washington
Non-Gel-Based Proteome Analysis
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Proteomic Analysis Using 2-D Liquid Separations of Intact Proteins From Whole-Cell Lysates
UNIT 23.3
This unit describes procedures for 2-D liquid separations of proteins from whole-cell lysates. Protocols for protein isoelectric point (pI) fractionation in the first dimension include the use of liquid isoelectric focusing (IEF; see Basic Protocol 1) and chromatofocusing (see Basic Protocol 2). The liquid IEF provides a pI-based fractionation using a batch-phase electrophoretic method, while chromatofocusing uses a column-based chromatographic method to generate the pH gradient. With either method, a second-dimension fractionation is provided in the liquid phase using nonporous silica-based reversed-phase HPLC (NPS-RP-HPLC; see Basic Protocol 3) to generate a 2-D liquid map of the protein content of the cell. Procedures for HPLC separation of the pI fractions of intact proteins are also described. The eluate of the 2-D liquid fractionation is directly coupled to a mass spectrometer for on-line detection of the intact molecular weights (MW) of proteins. As a result, a multidimensional map of protein expression is obtained that characterizes cellular proteins by pI, hydrophobicity, and intact molecular weight. Such expression maps are useful for differential proteomic comparison between different cell samples. PROTEIN FRACTIONATION BY ISOELECTRIC FOCUSING USING THE ROTOFOR CELL
BASIC PROTOCOL 1
The first dimension of the 2-D liquid separations presented in this unit is based upon isoelectric point (pI) fractionation carried out in a manner analogous to 2-D gel electrophoresis (UNIT 10.4). In this protocol, pI fractionation is achieved by liquid phase isoelectric focusing (IEF). There are several ways to accomplish liquid-phase IEF (see UNIT 10.2 for general principles of IEF), including the use of carrier ampholytes, isoelectric membranes, and continuous flow electrophoresis (UNIT 22.5). In the present discussion, procedures are described for carrier ampholyte–based liquid IEF separations, which are analogous to separations using carrier ampholyte gels (Scopes, 1994). Liquid IEF has the advantages of high sample capacity, ability to fractionate over a broad pH range, and ability to separate cytosolic proteins in addition to membrane-bound proteins and proteins of high molecular weight. The liquid-phase IEF separation is performed using a commercial device called the Rotofor (Bio-Rad). The Rotofor separates proteins in solution based on pI, which is the pH value at which the net charge of a protein is zero. The amino acid sequence and the environment determine the charge on the protein. A protein is positively charged when the pH of the environment is lower than its pI and negatively charged when the pH of the environment is higher than its pI. Upon application of electric potential, positively charged proteins move toward the anode and negatively charged proteins move toward the cathode until they are neutral, i.e., the pI is approached (See Fig. 23.3.1). Thus, the proteins are focused at their respective pIs. If the proteins attempt to diffuse out of their focused zones, they will ionize and the electric field will push them back to their pI. This protocol describes a typical Rotofor experiment including materials and step-by-step separation procedures. The procedures are similar for the mini-Rotofor and standard Rotofor. With the following protocol, a wide range of protein samples can be fractionated into 20 pI fractions over a range of pH 3 to 12 using the standard commercial apparatus. In addition, these experiments have been devised to be compatible with a second dimension of liquid separation and further analysis by mass spectrometry.
Non-Gel-Based Proteome Analysis
Contributed by Kan Zhu, Fang Yan, Kimberly A. O’Neil, Rick Hamler, Linda Lin, Timothy J. Barder, and David M. Lubman
23.3.1
Current Protocols in Protein Science (2003) 23.3.1-23.3.28 Copyright © 2003 by John Wiley & Sons, Inc.
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pH 3
10 NH+3
anode +
NH+3
NH+3
NH2
NH2
NH2 COOH
NH2
NH2
NH2
COOH
NH2
cathode – COOH
COOH COOH COOH COOH COOH COOH COOH COO– COO– positively charged negatively charged neutral electric field
Figure 23.3.1 Electrophoretic focusing in the Rotofor. Denaturing conditions are utilized for isoelectric focusing, so proteins will not be present in their native folded states as depicted here.
To obtain optimum performance with the Rotofor, it is necessary to quantify proteins from cell lysates before the Rotofor fractionation. Each collected pI fraction should also be analyzed quantitatively for protein to calculate overall recovery. The Bradford assay is a convenient method of protein quantitation, considering its compatibility with many detergents, chaotropes, and reducing agents. However, some reducing reagents and detergents may interfere with the assay. Check the compatible reagent lists that accompany the assay kits before using. The sample concentration must be within the linear range of the standard curve, from 1.2 to 10 µg/ml (for Bradford microassay). A dilution from 40 to 400 is usually performed for cell lysates and fractionated samples. In order to eliminate the possible interference from the cell lysis buffer or running buffer, the same volume of corresponding buffer is mixed with dye, standard, and water and analyzed as a blank. Materials Protein sample: see e.g., Support Protocol 1; concentration determined, e.g., by Bradford assay (UNIT 3.4) or Bio-Rad RC-DC Protein Assay Kit Cathode electrolyte: 0.1 M NaOH Anode electrolyte: 0.1 M H3PO4 Rotofor running buffer (see recipe) Bio-Lyte 3/10 ampholyte (Bio-Rad) Rotofor system (Bio-Rad) Circulating water cooling system Power supply with constant power capability 50-ml conical centrifuge tube 50-ml syringe with 19-G, 1.5-in. needle Harvesting tubes: 12 × 75-mm culture tubes Cold trap Vacuum pump pH meter Additional reagents and equipment for protein assay (UNIT 3.4) 2-D Liquid Separations of Whole-Cell Lysates
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Prepare and pre-run the Rotofor system 1. Equilibrate anion-exchange membrane in 0.1 M NaOH and cation-exchange membrane in 0.1 M H3PO4 overnight. Ion-exchange membranes isolate the electrolytes from the focusing chamber while allowing current to flow. Anion- and cation-exchange membranes are not interchangeable. Once they are wet, they should be kept in cathode and anode electrolyte solution, respectively.
2. Assemble the Rotofor system according to the manual. The standard Rotofor and mini-Rotofor have different sets of membrane cores and focusing chambers. The standard Rotofor holds 60 ml and can handle milligram to grams of total protein, while the mini-Rotofor can hold 18 ml and is more appropriate for micrograms to milligrams of total protein.
3. Connect circulation water and power supply to the Rotofor system. The cooling system is important for heat dissipation. The two ports on the cooling finger are interchangeable.
4. Seal the harvesting port of the Rotofor with sealing tape. Fill the Rotofor chamber with distilled water, then seal the loading port with tape. Apply 5 W for 5 min, then remove the tape and discard the water. This is a necessary step to remove residual ionic contaminants from the membrane core and the ion-exchange membranes.
Load sample and begin run 5. Seal the harvesting port with tape. Before loading, mix the protein sample with the Rotofor running buffer in a 50-ml conical centrifuge tube for a total volume of 18 ml (for Mini-Rotofor), making sure that the final concentration of ampholytes matches that of the protein, per Table 23.3.1. Load the sample/running buffer mixture into the Rotofor chamber using a 50-ml syringe with a 19-G, 1.5-in. needle, taking care that no air bubbles are trapped. Seal the loading port with tape. During filling, air bubbles can be trapped in the system, especially with the mini-Rotofor. This will cause fluctuations in voltage and discontinuity in the electric field. Bubbles must be removed before the separation starts (see Troubleshooting). Gloves should be worn when handling the tape; otherwise skin keratins may contaminate samples.
6. Apply 12 W (mini-Rotofor) or 15 W (standard Rotofor) of constant power to the chamber (current decreases with time while voltage increases), maintaining the temperature at ∼15°C by setting the coolant temperature to 4°C. The separation process takes about 4 hr and is complete when the voltage remains stable for ∼30 min. When potential is applied to the sample solution, charged proteins and carrier ampholytes move toward their pI values, so that the initial current is high. With the increase of time, carrier ampholytes and some of the proteins reach their pIs and are neutralized, so that the number of charged species decreases and the current decreases with time. Do not wait too long after the voltage is stable; otherwise, the established pH gradient will collapse. Table 23.3.1
Ampholyte Concentration Corresponding to Sample Concentrationa
Protein concentration
Ampholytes (v/v)
>2 mg/ml 1 mg/ml 0.5 mg/ml 0.25 mg/ml
2% 1.5% 1% 0.5%
aThis information was taken from the Bio-Rad Rotofor instruction manual.
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Harvest fractionated sample 7. Place 20 harvesting tubes into the rack inside the harvest box. Connect the outlet of the harvest box with the cold trap that is connected to a vacuum pump. Start vacuum pump. The cold trap is used to protect the vacuum pump and is not supplied with the Rotofor system. A typical glass cold trap contains one inlet, which is connected to the outlet of the harvesting box, and one outlet, which is connected to the vacuum pump.
8. Turn the power supply off and disconnect it. Remove the tape from the loading port and punch the 20 pins through the tape on the harvesting ports. Focused fractions are aspirated from the focusing chamber by the vacuum pump. To prevent uneven harvesting, make sure needles are pushed uniformly through the sealing tape into the chamber. These procedures need to be performed quickly and without shaking the apparatus, so that diffusion of proteins after removal of the electric field is minimized.
9. Measure pH of the 20 collected fractions with a pH meter (Fig. 23.3.2). Determine protein concentration of each fraction (UNIT 3.4 or Bio-Rad RC-DC Protein Assay kit). The fractions are now ready for a second-dimension separation, e.g., see Basic Protocol 3. If the samples are not used right after fractionation, they should be stored frozen at –80°C. It is best to analyze them within 6 months.
10. Disassemble and clean the Rotofor system. Clean the ion-exchange membranes with distilled water, detergent (e.g., ∼10% Alconox), or 0.1 M NaOH to remove residual proteins and then place them back into fresh equilibration solution (see step 1). The membranes are reusable if kept wet. The membrane core should also be thoroughly washed with distilled water and is reusable. ALTERNATE PROTOCOL 1
AMPHOLYTE-FREE PREPARATIVE ISOELECTRIC FOCUSING OVER NARROW pH RANGES IN THE ROTOFOR CELL Additional Materials (also see Basic Protocol 1) Rotolyte buffer-pair mixtures (Bio-Rad)
14 12
pH
10 8 6 4 2 0 1
2-D Liquid Separations of Whole-Cell Lysates
3
5
7
9 11 13 Fraction number
15
17
19
Figure 23.3.2 A typical pH gradient achieved in a Rotofor experiment.
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In Basic Protocol 1, the pH gradient is maintained by thousands of ampholytes. In this ampholyte-free method, only one of 13 buffer pairs is used at a time. When voltage is applied, the buffer pair generates a pH gradient between the individual pKa values of the buffers. The simple mixture of the buffer pair cannot provide as wide a linear range as the ampholyte-based method; the typical pH range used in this method is within 1 to 2 pH units. This is a useful method for concentrating a limited number of proteins over a small pH range, but is not suitable for profiling proteins in a complex biosystem. The method eliminates possible interferences from ampholytes, thereby extending its applicability to protein samples incompatible with ampholytes. The thirteen buffer pairs used in this method are listed in the Bio-Rad Rotofor manual. This series of buffer pairs cover the pH range from pH 2.9 to 11.0. Prior to the experiment, the pI of the target proteins should be known. For optimum resolution, ratios of the selected buffer pairs can be adjusted so that the pI of the protein to be purified falls near the middle of the established pH gradient. The experimental procedure here is essentially the same as that for the ampholyte-based method (see Basic Protocol 1). However, the running buffer and electrolytes used in the anodic and cathodic cells are different. The running buffer is usually composed of 50% (v/v) water/sample and 50% (v/v) Rotolyte buffer-pair mixtures. A high concentration of buffer mixtures is important for preventing pH distortion when charged proteins migrate. Anode and cathode electrolytes are categorized in the Bio-Rad Rotolyte instruction manual. Appropriate electrolytes should be used to minimize the pH distortion at the acidic and basic ends. ISOELECTRIC FOCUSING USING THE ROTOFOR WITH ION-EXCHANGE RESIN AS ELECTROLYTE
ALTERNATE PROTOCOL 2
Additional Materials (also see Basic Protocol 1) AG50W-X8 cation exchanger (hydrogen form; Bio-Rad) AG1-X8 anion exchange resin (hydroxide form; Bio-Rad) One of the limitations of the Rotofor is the pH gradient distortion at the acidic and basic ends. During electrophoresis, ampholytes and proteins migrate and focus at their pI points in the chamber; however, in the anode and cathode cells, gas generated during electrolysis pushes acid and base through the ion-exchange membranes and into the focusing chamber. Using ion-exchange resins in both electrodes where free hydronium and hydroxide ions are in equilibrium with ions attached to the resin can alleviate this problem. The result is that a limited number of free ions are pushed through the membrane and the pH gradient distortion is minimized. In this alternative procedure, the focusing experiment is conducted in the same manner as described in Basic Protocol 1; however, instead of loading 0.1 M phosphoric acid in the anode and 0.1 M sodium hydroxide in the cathode, AG50W-X8 cation exchanger (hydrogen form) and AG1-X8 anion exchange resin (hydroxide form) are applied to the anode and cathode, respectively. Prior to the experiment, the resin should be washed with deionized water three times, after which it should be resuspended in deionized water for a volume of ∼20 ml, drawn up into a 50-ml syringe without a needle, and injected into the electode cell until the cell is ∼80% full. When using resins in the electrode chambers, Rotofor running times can be greatly increased as compared to the standard operating conditions.
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BASIC PROTOCOL 2
PROTEIN FRACTIONATION BY CHROMATOFOCUSING Chromatofocusing (CF; also see UNIT 8.5) has been used as an alternative method for performing a pI separation in the first dimension of the 2-D liquid separation scheme (Sluyterman, 1978; Sluyterman, 1981a,b; Liu and Anderson, 1997; Strong and Frey, 1997; Fountoulakis et al., 1998; Hutchens, 1998; Chong et al., 2001). CF is a column-based chromatographic method that uses a weak anion exchange column to generate a descending linear pH gradient. In contrast to conventional ion exchange chromatography, the exchanger is titrated using ampholytes (Polybuffer, Amersham Pharmacia Biotech) rather than salts as the titration media. The result is a pH separation where fractionation can be performed down to <0.1 pH units. The CF method has been marketed for many years by Amersham Pharmacia Biotech and is used mainly as a preparative method to purify a limited number of proteins for further chromatographic analysis (Pharmacia Biotech, 1985). However, with recent proprietary advances in ion-exchange column technology and polyampholyte buffers, this method has demonstrated the capability of fractionating large numbers of proteins from mammalian cell lysates as the first step in a 2-D liquid chromatography setup. This improved 2-D column technology is available through Eprogen, Inc., under the name ProteoSep 2-D Protein Mapping Kit. In the CF process, pH is monitored using an on-line pH probe mounted in a flow cell; consequently, each pH range can be automatically collected using a fraction collector. The use of CF allows efficient interfacing to the second-dimension NPS-RP-HPLC (see Basic Protocol 3) separation without additional sample manipulation. The use of on-line pH monitoring also provides a basis for reproducibly collecting pH fractions from different samples. A second dimension such as NPS-RP-HPLC can then be used to analyze each pI fraction. The result is a 2-D UV map of protein expression, with high reproducibility. Furthermore, this technique provides excellent fractionation and relatively pure proteins in the liquid phase that can be collected for further characterization. The method below has a protein loading capacity comparable to 2-D gels, i.e., between 100 µg and 5 mg. Most importantly, the use of CF as the first dimension of separation allows the entire 2-D liquid separation process to be readily automated. Materials Protein sample: see, e.g., Support Protocol 1; concentration determined, e.g., by Bradford assay UNIT 3.4 or Bio-Rad RC-DC Protein Assay kit CF start buffer (see recipe for pH ranges 8.5 to 4 or 7 to 4) CF elution buffer (see recipe for pH ranges 8.5 to 4 or 7 to 4) 1 M NaCl Desalting column: PD-10 (Sephadex G-25; Amersham Pharmacia Biotech) NPS-RP-HPLC column (recommended): 33 × 4.6–mm Micra Platinum column (Eprogen) HPLC/FPLC system running at ambient temperature and at a flow rate of 0.2 ml/min, with fraction collector, UV detector (280 nm), pH electrode, flow cell, and flow meter Chromatofocusing (CF) column: HPCF 1-D column (Eprogen) Additional reagents and equipment for desalting and buffer exchange by column chromatography (UNIT 8.3)
2-D Liquid Separations of Whole-Cell Lysates
NOTE: The CF buffers (see Reagents and Solutions for recipes) bracket the pH range over which the separation is to be performed. CF Buffer solutions are also commercially available through Eprogen as part of the 2-D Proteosep kit. It is preferable to use the premixed buffers to ensure reproducible results.
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Exchange samples into CF start buffer 1. Equilibrate the PD-10 column with ∼25 ml of CF start buffer (also see UNIT 8.3). 2. Load 2.5 ml of sample onto PD-10 column (if the sample volume is <2.5 ml, bring to volume with CF start buffer). Discard the eluate. 3. Add 3.5 ml of CF start buffer to the PD-10 column after the sample has entered the column bed. Collect the eluate until all 3.5 ml of the buffer has entered the bed. Test sample preparation on RP-HPLC column (recommended) 4. Inject 50 µl of exchanged sample onto a NPS-RP-HPLC column to check the detailed reversed-phase elution profile of the sample prior to pI fractionation. This second-dimension RP-HPLC analysis is highly recommended prior to running the first-dimension CF analysis. If any sample preparation problems have occurred, they will be readily identified after analysis of the chromatogram from this step. This diagnostic measure may also help the investigator extend the lifetime of the column, since the distribution of the hydrophobicities of proteins can be obtained in the prerun. More hydrophobic proteins will elute from the column later, and if there are more hydrophobic proteins in the sample, extensive cleaning is required for the CF and reversed-phase columns to extend their lifetime.
Relative intensity at 214 mn (mV)
Depending on the complexity and amount of proteins expected to be in the sample, a very “spiky”– or “porcupine”–looking chromatogram (Fig. 23.3.3) should be obtained at this point. One should not proceed to the CF fractionation step if one does not see a protein profile that makes sense for the sample being analyzed.
200 190 180 170 160 150 140 130 120 110 100 90 80 70 60 50 40 30 20 10 0 5
7
9
11 13 15 Retention time (min)
17
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Figure 23.3.3 Chromatogram of hepatocyte cell culture sample exchanged with start buffer (optional pre-run on second-dimension NPS-RP column, without previous CF; see Basic Protocol 2, step 4).
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Perform first-dimension analysis by chromatofocusing 5. Before injection, flush the CF column with 10 column volumes (CV; for the HPCF 1-D column, 1 CV = 0.87 ml) of 100% distilled water, filtered with a 0.45-µm filter. 6. Equilibrate the CF column with 100% CF start buffer until the pH of the effluent reaches that of the start buffer and the baseline is stable (∼30 CV). 7. Inject 1.0 to 3.0 ml of sample onto the CF column. Loading 1 to 5 mg of total protein is recommended to allow for UV detection of lowerabundance proteins. With injections of 10 to 100 ìg of total protein, it is possible to obtain a separation of highly expressed proteins, but proteins expressed in low abundance will not be readily observed using UV detection. The number of proteins observed is related to the total protein load, the concentration of the individual protein, and the UV absorption detection of the protein. When studying samples that contain a dominant protein (e.g., serum or urine), loading on the high end of the recommended total protein amount (1 to 5 mg) may cause overload of the UV signal in the pI fraction where the dominant protein elutes.
8. Wash CF column with 100% CF start buffer at a flow rate of 0.2 ml/min while monitoring the baseline absorbance. Collect this wash as a single fraction since all proteins with pIs >8.5 will elute from the column during this wash step. After the UV absorbance value returns to baseline, stop the flow of start buffer. The wash step will take ∼10 to 30 min.
9. Before beginning the pH gradient, flush the tubing and pump head with 100% elution buffer (volume depending on system configuration) to reduce dead volume. Start the gradient by running 100% elution buffer at 0.2 ml/min. Allow 65 to 70 min to perform the full pH gradient; at ∼60 min the pH of the eluate should be 4.0 ± 0.1. Collect fractions at chosen pI intervals, generally 0.1 to 0.3 pH units, using fraction collector. Monitor and record the pH at the start and end of each fraction collected. Software for automatic monitoring of the pH and reproducible fraction collection has been developed by Dr. Steven J. Parus ([email protected]) of the University of Michigan, Department of Chemistry.
80
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injection of 3.0 ml of 3.5 ml Fao cell lysate exchanged with SB, 0.2 ml/min
8.6
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Retnetion time (min)
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Figure 23.3.4 UV profile of a typical chromatofocusing experiment.
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10. After completing pH run, wash HPCF column with 10 CV of 1.0 M NaCl. Collect the UV peak(s) as a fraction for analysis with the second-dimension column (see e.g., Fig. 23.3.4). 11. Transfer fractions from the fraction collector that are to be analyzed in the second dimension to appropriate vials for use with the HPLC autosampler. Freeze fractions at –20°C or below if the second dimension analysis is not to be performed within 8 to 10 hr. Basic Protocol 3 provides an example of a second-dimension separation.
12. Wash the CF column with 10 CV of deionized water. The HPCF 1-D column can be stored in water at room temperature up to 1 to 2 days, but should be stored in 100% isopropanol for long periods of time (>2 days) at room temperature.
CELL LYSIS FOR PROTEIN FRACTIONATION BY CHROMATOFOCUSING OR ISOELECTRIC FOCUSING USING THE ROTOFOR
SUPPORT PROTOCOL 1
Cell lysis is a process that extracts proteins from cells. In order to keep extracted proteins intact in solution, the pH of the lysis buffer is controlled close to physiological pH using Tris buffer, and proteases are inhibited by protease inhibitors. Furthermore, various nonionic chaotropes, detergents, and reducing reagents can be used to enhance the solubility of proteins. In both the electrophoretic-based separation (liquid-phase IEF, Rotofor) and the ion-exchange-based separation (chromatofocusing), the ions will cause the focusing and protein binding, respectively, to deteriorate so maintaining low ionic strength is a key issue. Materials Cells for analysis 50 mM Tris⋅Cl, pH 7.8 to 8.2 (APPENDIX 2E; any pH between these values can be used) containing 1 mM PMSF (add from 0.5 M stock in DMSO or isopropanol; see note below) General lysis buffer (see recipe) Membrane lysis buffers A and B (see recipes) 0.1 M sodium carbonate (ice cold) 50 mM Tris⋅Cl, pH 7.3 (APPENDIX 2E) containing 1 mM PMSF (add from 0.5 M stock in DMSO or isopropanol; see note below) Ultrasonicator with microtip probe (e.g., Branson; for bacterial lysis) Bath sonicator (for membrane lysis) Centrifuge (refrigerated centrifuge capable of 20,000 × g for general lysis procedure; ultracentrifuge capable of 150,000 × g for membrane lysis procedure) Dry ice/acetone bath NOTE: PMSF has a half-life in aqueous solution of ∼1 hr. Therefore, a stock solution of 0.5 M PMSF should be made in DMSO or isopropanol and should be added to the aqueous lysis solutions (50 mM Tris⋅Cl and membrane lysis buffer A) to a final concentration of 0.5 mM (i.e., half the original concentration of 1 mM) every hour. The DMSO or isopropanol stock can be stored at 4°C for up to 1 month. CAUTION: Be aware that PMSF is highly toxic and solutions containing this compound must be handled with caution at all times. Non-Gel-Based Proteome Analysis
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For bacterial cells 1a. Suspend the 10-mg bacterial cell pellet in 0.4 ml of 50 mM Tris⋅Cl, pH 7.8 to 8.2. Vortex vigorously, then sonicate using an ultrasonicator with a microtip in an ice bath. Repeat this procedure twice, each time for 30 sec, to break the cell membranes. 2a. Mix the suspension into 1.6 ml of general lysis buffer. The final cell lysate consists of 6 M urea, 2 M thiourea, 10% glycerol, 50 mM Tris, 2% OGL, 5 mM TCEP, and 1 mM protease inhibitor.
3a. Centrifuge solution at >20,000 × g for 60 min. 4a. Remove the supernatant from the cell debris. Store at −20°C until use. This sonication method works well for most Gram-negative bacteria such as E. coli. It may not work well for Gram-positive bacteria and other specialized bacteria with strong cell membranes. In these cases, it is recommended to use a much more rigorous method to break the cell membrane, such as the French press (UNITS 6.1 & 6.7).
For human cell lines 1b. Mix ∼0.4 ml of a single-cell suspension of cell line sample containing 1 × 108 cells into 1.6 ml of general lysis buffer and vortex vigorously. If cells are in pellet form, suspend the cell pellet in 0.4 ml of 50 mM Tris⋅Cl, pH 7.8 to 8.2, and vortex, then add 1.6 ml of general lysis buffer and vortex vigorously. The final cell lysate consists of: 6 M urea, 2 M thiourea, 10% glycerol, 50 mM Tris, 2% OGL, 5 mM TCEP, 1 mM of protease inhibitor.
2b. Centrifuge the lysate 60 min at >20,000 × g, 4°C. 3b. Remove the supernatant from the cell debris. Store at −20°C until use. For membrane protein lysis 1c. Suspend cell pellet equivalent to 6 × 108 cells in 3 ml of membrane lysis buffer A. Vortex vigorously. 2c. Lyse cells by placing tubes in an dry ice/acetone bath to quickly freeze cells, then allowing them to thaw at room temperature. Vortex thoroughly and freeze again. Repeat this cycle a total of five times. Sonicate in a bath sonicator on ice for 30 min, then vortex again. 3c. Repeat step 2c two additional times for a total of 15 freeze/thaws and three sonications. 4c. Centrifuge 20 min at 2800 × g, 4°C, to remove unlysed cells. Remove supernatant and set aside on ice. 5c. Resuspend the pellet from step 4c in 1 ml of membrane lysis buffer A, repeat steps 2c to 4c, and pool the resulting supernatant with that set aside in step 4c. 6c. Add 10 volumes of 0.1 M sodium carbonate to the pooled supernatant. Agitate/stir on ice for 1 hr. 7c. Collect membranes by centrifuging 1 hr at 150,000 × g, 4°C. Remove and discard supernatant.
2-D Liquid Separations of Whole-Cell Lysates
8c. Resuspend pellet by adding 1 ml of 50 mM Tris⋅Cl, pH 7.3, and vortexing. Collect membranes by centrifuging for 1 hr at 150,000 × g, 4°C. Remove and discard supernatant. Repeat this wash step two more times for a total of three washes of the membrane pellet. This serves to remove soluble protein contaminants in the membrane preparation.
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9c. Solubilize the membrane pellet in 1.2 ml of membrane lysis buffer B and vortex vigorously until pellet dissolves. Use more buffer if necessary for solubilization. Store at –20°C until use. This membrane extraction procedure isolates all cellular membranes including the plasma membrane as well as organelle membranes such as mitochondria, Golgi, and vesicles.
NONPOROUS REVERSED-PHASE CHROMATOGRAPHY INTERFACED TO ESI-MS Reversed-phase chromatography (RP-HPLC; also see UNITS 8.7 & 11.6) is a widely used separation method with powerful resolving capability, reproducibility, and recovery. Separation in RP-HPLC is based on hydrophobicity; thus, it serves as an orthogonal separation method to the first-dimension pI separation. In RP-HPLC, after a protein sample has adsorbed to the column material, a gradient of increasingly hydrophobic mobile phase causes a protein molecule to elute from the column when its affinity for the mobile phase exceeds its affinity for the stationary phase. When the organic concentration in the mobile phase reaches a particular value, the protein desorbs from the stationary phase and enters the mobile phase to elute from the column. Hydrophobic proteins interact more strongly with the bonded stationary phase (conventionally alkyl groups such as C4, C8, and C18) than hydrophilic proteins and, consequently, elute from the column later during the separation. Under optimal conditions, protein peak retention times vary only approximately 0.1 min (Wall, 2000, 2001; Kachman, 2002) over a total separation time of 30 min for complex protein sample mixtures. High reproducibility of retention times using nonporous silica (NPS) columns permits collection of protein peaks for further analysis, including peptide mass fingerprinting using MALDI-MS (Chong et al., 1999; Wall et al., 1999; Chong et al., 2000; UNIT 16.5). In addition, this technique permits comparison of chromatographic profiles for differential proteomic studies.
BASIC PROTOCOL 3
Strategic Planning for Interfacing NPS RP-HPLC to ESI-MS Electrospray ionization Electrospray ionization (ESI) is an ideal interface between RP-HPLC and mass spectrometry (MS). The HPLC eluate can be directly introduced on-line to the ESI source. Eluate solution is sprayed through a capillary with high voltage applied, producing multiply charged droplets. The charges are dispersed on the droplet surface until the charge-charge repulsion exceeds the surface tension, forcing the droplet to split into smaller droplets. Heated desolvation gas in the ESI source aids in decreasing droplet size by vaporizing solvent molecules, causing greater charge repulsion and droplet instability. This process continues until all solvent has evaporated and the sample molecules become multiply charged gas-phase ions. ESI is a soft ionization method where proteins are detected as intact, multiply charged molecular ions with little or no fragmentation. Since proteins possess multiple sites capable of accepting protons, the mass spectrum appears as an umbrella of multiply charged ions. The intact molecular weight (MW) of the protein is obtained by deconvoluting the multiply charged peaks in the ESI-MS spectrum (see Fig. 23.3.5). Desalting Fractions from the pH dimension contain reagents other than proteins. Chaotropes, detergents, and ampholyte molecules can both interfere with and suppress sample ionization and detection. They suppress ionization of sample molecules by competing for charges. In addition, the large number of ionized reagent species may saturate the mass spectrometer detector and decrease the signal-to-noise ratio of detection (Cech and Enke, 2001). Detergents and metal ions can form adducts with proteins, leading to the detection of artifactual protein modifications. Therefore, desalting before ESI-MS is necessary to
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obtain high-sensitivity spectra without adducts or other artifacts. RP-HPLC provides an excellent means of sample desalting, since proteins can be concentrated onto the head of the column and remain on the column during a water wash. Multiple injections of deionized water are used to wash off various unwanted reagents. The Rotofor fractions require more water injections for washing than chromatofocusing fractions, since there are more contaminants that need to be eliminated. Interfacing pH-separated fractions to NPS-RP-HPLC After pH fractionation, proteins remain in the liquid phase. Sample concentration and desalting are achieved on the RP column. Consequently, no further sample treatment is necessary for interfacing to the pH fractions to liquid chromatography. Considering the differences in sample capacity for the standard Rotofor, mini-Rotofor, and chromatofocusing, a protein assay (UNIT 3.4) is usually performed to quantify each fraction so that the optimal amount of protein is introduced onto RP column. All pH-separated fractions should be analyzed individually to maximize the separation capacity for both dimensions. The greater the number of fractions obtained in the
A 100
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39500 40000
44155.0 42369.0 42732.0 43274.0 43765.0
40500 41000 41500 42000 42500 Mass (Da)
43000 43500 44000
Figure 23.3.5 NPS-RP-HPLC/ESI-TOF mass spectrum of β-actin and γ-actin. (A) Combined ESI-TOF spectra of β-actin and γ-actin. (B) MaxEnt deconvoluted β-actin and γ-actin.
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first-phase separation, the better the resolution achieved in the pH dimension and in the subsequent HPLC separation; however, the greater the number of fractions obtained, the greater the amount of work required in the second dimension. In the case of the Rotofor, up to 20 fractions are obtained in the first dimension. However, in CF, if one uses a 0.1–pH unit fractionation, then up to 45 fractions may be obtained over the entire range, each of which needs to be interfaced to RP-HPLC. The running conditions in the pH dimension are different from those in the RP-HPLC. In the pH dimension, proteins are focused to their pI values, where protein solubility is at a minimum, making chaotropes and detergents necessary to prevent protein precipitation. In RP-HPLC, the mobile phase pH is very acidic because of the addition of TFA and FA. Chaotropes and detergents are not necessary and are actually detrimental to the ESI process. The change in pH and medium may cause some protein precipitation. In order to prolong the lifetime of the RP column, it is necessary to mix the sample with mobile phase A (no organic modifier) to minimize precipitation and centrifuge the mixture at 14,000 rpm for 10 min to eliminate particulates. Column selection In order to select an appropriate column for RP-HPLC, factors such as particle size, column diameter, pore size, and coating should be considered (Scopes, 1994). Decreasing particle size helps intraparticulate mass transfer and increases column efficiency and resolution. The major disadvantage is the increased back-pressure that accompanies the use of smaller particles at the same flow rate. However, at an acceptable working pressure, a smaller particle size provides improved separation. This protocol employs 1.5-µm ODS(C18) nonporous silica (NPS) particles (Eprogen, Inc.). NPS columns provide a faster high-resolution separation with better protein recovery than comparable porous columns (Gooding and Regnier, 1990; Hanson et al., 1996; Banks and Gulcicek, 1997; Barder et al., 1997). Slow and restricted diffusion of protein molecules in conventional porous packing materials limits the speed of protein separation while maintaining satisfactory resolution and detection sensitivity. Moreover, recovery is low due to protein adsorption within pores in the stationary phase preventing protein elution. Considering the follow-up analysis with MS, high speed and efficiency are essential for several reasons. First, a high-speed separation results in less MS data to be processed and a subsequent gain in sample throughput. Also, ESI-MS is a concentration-sensitive technique; therefore, high efficiency in the RP-HPLC separation implies better sensitivity in MS separation (Niessen and Tinke, 1995). Nonporous media eliminate issues of macromolecular pore adsorption and stagnant mobile phase in the intraparticle void volume associated with porous stationary phases. As a result, higher column efficiency is obtained with much improved resolution (Chen and Horvath, 1995; Hanson et al., 1996; Lee, 1997; Yu and El Rassi, 1997; Yang and Lee, 1999). A disadvantage is decreased surface area, hence sample capacity, which is compensated for by using very small (1.5-µm) silica particles. Decreasing the length of the column alleviates the problem of high back-pressure associated with small particles. In addition, at elevated temperatures, NPS packing materials are generally more stable than their porous counterparts. Overall, the high efficiency of NPS 1.5-µm packing allows for reproducible, high resolution, high-speed protein separations. In RP-HPLC, the surface of the stationary phase is hydrophobic. Rigid supports such as silica are covalently bonded with alkyl groups, typically C4, C8, and C18. The interaction between protein and the immobilized alkyl groups increases with longer alkyl chain. Although stationary phases with shorter alkyl chains have demonstrated better recovery, short-chain, polar, bonded-phase silica experiences poor stability in reversed-phase operation at the low pH and elevated temperature required for efficient protein separations Non-Gel-Based Proteome Analysis
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(Kastner, 2000). Consequently, C18 (ODS) stationary phase is still the choice for interfacing RP-HPLC to ESI-MS. Small-diameter columns are more easily interfaced to ESI-MS because of their lower optimum flow rate; however, the sample capacity of narrow-bore columns is low. For regular columns, the optimum flow rate is usually too high (∼1 ml/min) to be efficiently ionized by ESI. Flow splitting after the column is necessary for effectively interfacing such columns (≥4.6 mm in diameter) with ESI-MS (Niessen and Tinke, 1995). The NPS 3-mm diameter columns (33 or 53 mm in length) are optimal in these studies. The flow rates used are generally 0.2 to 0.4 ml/min through the column and 0.1 to 0.2 ml/min after the split. There are three kinds of NPS columns: ODS-I, ODS-II, and ODS-IIIE, available through Eprogen. NPS ODS-I is polymeric bonded phase with low surface silanol activity and no silanol end capping. The stability over a broad pH range makes the ODS-I phase suitable for analyzing fractions in the range of pH 2 to 10. NPS ODS-II phase is monomeric in nature; it provides a more hydrophilic surface and higher selectivity than the ODS-I phase for the protein separation (see volume 3 of the Eprogen manual for Micra HPLC columns; http://www.eprogen.com/hplc/catalog/cataloglowrez.pdf). It is the premium choice for the separation of proteins after CF fractionation due to its excellent peak shape, higher resolution, and peak capacity. However, the ODS-II stationary phase is not as stable as ODS-I within the basic pH range. The mobile phase pH for ODS-II is from pH 2 to 9; ODS-I can go up to pH 10, but for storage, it is better to keep these columns between pH 5 and 9. ODS-IIIE has monomeric binding like ODS-II except it is end-capped, which provides faster separation for small molecules and high stability for basic species. In order to select the right column for a separation, particle size, pressure, recovery, stability, sample capacity, and efficiency need to be balanced. The authors typically use NPS 1.5 µm-particle size packed columns. In the case of UV detection, generally, 4.6 × 33–mm columns have been used. When interfacing to ESI-MS, the 3 × 33–mm NPS columns have been used. In cases where large amounts of sample need to be collected for further analysis, an 8 × 33–mm column has been used with a flow rate of 1 ml/min. Temperature control Protein separations require the use of elevated column temperatures to achieve good separation efficiency and reproducibility. At elevated temperatures, the mobile phase viscosity is reduced and mass transfer in solution is enhanced. High pressure due to the use of small-dimension packing materials is alleviated, allowing faster separations (Chen and Horvath, 1995). Moreover, at higher temperatures, the sorption kinetics of the sample are accelerated; as a result, plate height decreases and column efficiency increases. Temperature can also affect protein structure and the reversed-phase surface. This could alter sample interaction with the reversed-phase surface and, as a result, resolution may be improved or degraded. Hence, stable temperature control is necessary for reproducible separation. In RP-HPLC, the use of an acidic hydro-organic mobile phase generally denatures proteins, so that using elevated temperatures does not further complicate the separation. One must be careful about using a temperature that is too high, where proteins may be labile. Furthermore, the stationary phase may not be stable at elevated temperatures. For RP-HPLC separations using nonporous silica, a column temperature of 60° to 65°C is generally most effective. At this temperature, rapid, high-resolution separations are obtained on columns packed with 1.5 µm NPS ODS without problems in column or protein stability. 2-D Liquid Separations of Whole-Cell Lysates
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Mobile phase In order to choose the organic component of the mobile phase, solute solubility, ionization efficiency, and miscibility of the eluent components should be considered. The eluting strength of the commonly used organic eluents follows the order: methanol
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Table 23.3.2
A Sample Gradient for NPS-RP-HPLC Separation
Organic (%B)
Duration (min)
5-15 15-25 25-31 31-41 41-47 47-67 67-100
1 2 3 10 3 4 1
Materials Sample fractions from pH separation (see Basic Protocol 1 or 2), with the protein concentration of each fraction determined (UNIT 3.4) 100% acetonitrile NPS-RP-HPLC Solvent A: dissolve 1 ml trifluoroacetic acid (TFA) and 3 ml formic acid (FA) in 996 ml ultrapure water NPS RP-HPLC Solvent B: Dissolve 1 ml TFA and 3 ml FA in 996 ml HPLC-grade acetonitrile HPLC system including: HPLC Pump RP column (NPS ODS-II, 3 × 33.0 cm; Eprogen) Column heater Splitter Syringe Ferrules PEEK tubing Clean and equilibrate the system 1. Purge the HPLC system with deionized water to remove salt. Inorganic salts may precipitate in the organic solvent in the tubing and the column.
2. Equilibrate column by first washing with 100% acetonitrile for 180 min followed by several blank gradients (from 100% water to 100% acetonitrile). Running a “blank gradient” means, specifically, that at a flow rate of 1 ml/min, one should apply the same gradient that will be used in the sample separation and make sure that the baseline is stable. Typically, one blank gradient is enough, but sometimes two to three blank gradients may be necessary.
Prepare and load sample 3. Dilute samples with starting mobile phase (NPS RP-HPLC solvent A) and centrifuge 10 min at 25,000 × g, room temperature, to remove any precipitated protein and other particulates. Proper sample preparation is important to extend the lifetime of the column.
4. Remove the supernatant from the centrifuged sample and inject it onto the column. Wash the column with multiple injections of deionized water until a water peak is observed. 2-D Liquid Separations of Whole-Cell Lysates
The wash step is recommended to eliminate detergents and salts which can interfere with MS analysis. When monitored at 214 nm with a UV detector, water absorption is lower than the mobile phase absorption, so injection of water will result in a negative peak.
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550
Relative intensity at 214 nm (mV)
500 450 400 350 300 250 200 150 100 4
8
12 16 20 Retention time (min)
24
28
Figure 23.3.6 The NPS-RP-HPLC chromatogram of chromatofocusing fraction pH 5.27 to 5.47 of colon cancer HCT116 cell line (UV 214 nm).
Prepare and optimize ESI-MS 5. Adjust the splitter to make sure the flow to the ESI-MS is optimized. Optimal results can be obtained when a 150 to 200 ìl/min flow is introduced to the mass spectrometer. The rule of thumb is to make sure that the eluate vaporizes completely. An easy means to check this (for the Micromass LCT mass spectrometer) is to ensure that there is no liquid accumulating on the baffle which is positioned opposite to the electrospray capillary.
Run NPS-RP-HPLC 6. Start gradient and data collection (Fig. 23.3.6). 7. To regenerate the column, rinse with 5 column volumes of deionized water, run 100% NPS RP-HPLC solvent B through the column for 30 min, then perform a sawtooth gradient for 20 cycles. Column regeneration and cleaning is important for prolonging column lifetime and reproducible separation results. Some proteins tend to show memory effects, so a sawtooth gradient is recommended. Memory effects refer to phenomena whereby, because proteins elute at a certain acetonitrile percentage, they will not elute off the column until the same acetonitrile percentage is reached again. A sawtooth gradient is a fast gradient from 100% water to 100% acetonitrile, followed by a gradient from 100% acetonitrile to 100% water, within the same time. The process is repeated several times.
8. Analyze data (see Support Protocol 2).
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fractions (I, I + 1, ...,J)
LC-MS for fraction I
LC-MS for fraction (I + 1) ...
LC-MS for fraction J
deconvolute fraction I
deconvolute fraction (I + 1) ...
deconvolute fraction J
combine fraction I
combine fraction (I + 1)
...
combine fraction J
Display fractions I...J according to their pI with Proteo Vue
Figure 23.3.7 Flow chart of 2-D map data process.
SUPPORT PROTOCOL 2
INTEGRATING pI, MW, AND ABUNDANCE INFORMATION INTO 2-D MAP As shown in Figure 23.3.7, fractions obtained from pH-based separations are subjected to LC-MS. In order to construct a liquid-phase 2-D map, fractions within the target pH range (fractions I to J in Fig. 23.3.7) are each analyzed by LC-MS (see Basic Protocol 3). One such liquid-phase 2-D map is shown in Figure 23.3.8. After fractionation of the whole-cell lysate of MCF10A, which is a normal breast epithelial cell line, using the Rotofor cell, each of the 20 fractions are analyzed by LC-MS. The MW value and peak area information are obtained from each deconvolution and combined for each fraction. The combined data are displayed in parallel according to their pH. A band on the 2-D map represents each protein, and its abundance is proportional to the optical density of the band. Bands with the same MW value in adjacent fractions with very close pI values may indicate an overlapping of a highly abundant protein; however, they are different proteins if their pI values are different. Furthermore, corresponding pH fractions from different samples are compared by placing resulting lane maps side by side (see Fig. 23.3.9). In the LC-MS experiment, a mass spectrum is acquired every second, so a typical 30-min run generates a tremendous amount of information. In order to convert raw MS data (as shown in Fig. 23.3.5A) into protein MW (as in Fig. 23.3.5B), a large number of deconvolutions need to be performed to obtain quantitative data. Although deconvolution software programs such as MaxEnt (Micromass, available from Waters) extract MW information for complex protein mixtures, the deconvolution is a time-consuming process that currently limits the throughput.
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100974
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81620
MW(Da)
71943
62266
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42912
33235
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13881
4204 2.71 3.45 4.48 5.25 5.53 5.73 5.81 5.94 6.16 6.58 6.87 7.02 7.14 7.27 7.44 7.69 8.07 8.43 9.34 10.55
Fraction pH
Figure 23.3.8 A complete two-dimensional mass map depicting MW versus pI of the MCF10A breast epithelial cell line. The x axis shows measured pH values of each of 20 Rotofor fractions and the y axis shows MW values from deconvoluted ESI-TOF-MS spectra.
Materials Personal computer (speed >200 MHz) MassLynx and MaxEnt (Micromass) MSCombine (S. Parus, University of Michigan) ProteoVue (Eprogen, Inc.) Data processing for one LC-MS run 1. Combine mass spectra containing the same multiply charged peaks into one combined mass spectrum using MassLynx (Micromass). One mass spectrum is acquired every second. For a protein eluting for 10 sec, 10 mass spectra should be combined into one spectrum. Non-Gel-Based Proteome Analysis
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MW(Da)
ES2 Fraction 7
MDAH Fraction 7
OSE Fraction 7
Figure 23.3.9 A two-dimensional mass map of MW versus pI comparing ES2, MDAH2774, and OSE ovarian cell lines for Rotofor fraction no. 7.
2. Still using MassLynx select the m/z range in the combined mass spectrum for deconvolution by zooming to the selected m/z range. In the mass spectrum, peaks below 800 are usually considered to be noise. It will take more time to deconvolute a raw spectrum with wider m/z range. Moreover, artifact peaks may appear in the deconvoluted spectrum.
3. Deconvolute the selected segment of spectrum with MaxEnt. The deconvoluted spectrum is displayed as peak intensity versus MW (Da).
4. Activate the “spectrum center function” in MassLynx, check the “areas” in the pop-up window for spectrum center, and perform center for the deconvoluted mass spectrum. 5. Zoom in to the protein peak and copy the MW value and peak area information into the MS Windows clipboard by using the “copy spectrum list” function in MassLynx. Paste into Notepad and save as a text file. 6. Repeat steps 1 to 5 through the LC-MS TIC (total ion count) chromatogram manually or using software if available. 7. Combine all text files into one combined text file with MSCombine. 8. Display the combined text file with ProteoVue. This is one lane in a 2-D map. In order to obtain a 2-D map covering the entire target pH range, steps 1 to 8 should be repeated for each fraction in the range. 2-D Liquid Separations of Whole-Cell Lysates
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
CF elution buffer, pH 7 to 4 Place the following in a 250-ml beaker: 90.09 g urea 25 ml Polybuffer 74 (Amersham Pharmacia Biotech) Ultrapure H2O to ∼210 ml Stir at room temperature until completely dissolved. Degas under vacuum by sonication or by sparging with helium. Adjust pH to 4.0 by adding saturated iminodiacetic acid (the pH measurement should be performed at the same temperature as that of CF experiments). Add 500 mg of n-octyl-β-D-glucopyranoside (OGL) and stir until dissolved. Transfer solution to 250-ml volumetric flask, rinse the beaker twice with water, and transfer to the flask. Add water to 250-ml mark. Mix well and store at 2° to 4°C in the dark. All solvents should be filtered through a 0.22-ìm nylon filter. Final concentrations are: 10% (v/v) Polybuffer, 6 M urea, and 0.2% (w/v) OGL.
CF elution buffer, pH 8.5 to 4 Place the following in a 250-ml beaker: 90.09 g urea 17.5 ml Polybuffer 74 (Amersham Pharmacia Biotech) 7.5 ml Polybuffer 96 (Amersham Pharmacia Biotech) Ultrapure H2O to ∼210 ml Stir at room temperature until completely dissolved. Degas under vacuum by sonication or by sparging with helium. Adjust pH to 4.0 by adding saturated iminodiacetic acid. Add 500 mg of n-octyl-β-D-glucopyranoside (OGL) and stir until dissolved. Transfer the solution to 250-ml volumetric flask, rinse the beaker twice with water, and transfer to the flask. Add water to 250-ml mark. Mix well and store at 2° to 4°C in the dark. All solvents should be filtered through a 0.22-ìm nylon filter. Final concentrations are: 10% (v/v) Polybuffer, 6 M urea, and 0.2% (w/v) OGL.
CF start buffer, pH 7 to 4 Place the following in a 500-ml beaker: 180.18 g urea 2.615 g Bis-Tris Ultrapure H2O to ∼440 ml Stir at room temperature until completely dissolved. Degas under vacuum by sonication or by sparging with helium. Adjust pH to 7.1 by adding saturated iminodiacetic acid. Add 1 g n-octyl-β-D-glucopyranoside (OGL) and stir until dissolved. Transfer the solution to 500-ml volumetric flask, rinse the beaker twice with water, and transfer to the flask. Adjust with water to 500-ml mark. Mix well and store the solution at 2° to 4°C in the dark. All solvents should be filtered through a 0.22-ìm nylon filter. Final concentrations are: 6 M urea, 25 mM Bis-Tris,and 0.2% (w/v) OGL.
CF start buffer, pH 8.5 to 4 Place the following in a 500-ml beaker: 180.18 g urea 1.865 g triethanolamine (or 3.529 g Bis-Tris propane) Ultrapure H2O to ∼440 ml.
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Stir at room temperature until completely dissolved. Degas under vacuum, by sonication, or by sparging with helium. Adjust pH to 8.5 by adding saturated iminodiacetic acid. Add OGL and stir until dissolved. Transfer the solution to 500-ml volumetric flask, rinse the beaker twice with water, and transfer to the flask. Add water to 500-ml mark. Mix well and store 2° to 4°C in the dark. All solvents should be filtered through a 0.22-ìm nylon filter. Final concentrations are: 6 M urea, 25 mM Bis-Tris propane, and 0.2% (w/v) OGL.
General lysis buffer Place the following in a 100-ml beaker: 22.52 g urea 9.52 g thiourea 0.304 g Tris hydrochloride 1.25 g n-octyl-β-D-glucopyranoside (OGL) 0.0896 g TCEP 1.868 ml of 0.5 M phenylmethylsulfonyl fluoride (PMSF) stock (prepared in DMSO or isopropanol) 20 ml 12.5% (v/v) glycerol Stir at room temperature until completely dissolved. Transfer all of the solution to a 50-ml volumetric flask. Add 12.5% (v/v) glycerol to the 50-ml mark. Dispense the above buffer solution into separate 2.0-ml vials, then store them at −20°C. Final concentrations are: 7.5 M urea, 2.5 M thiourea, 12.5% (v/v) glycerol, 50 mM Tris, 2.5% (w/v) n-octyl-β-D-glucopyranoside (OGL), 6.25 mM Tris-(carboxyethyl)phosphine hydrochloride (TCEP), and 1.25 mM of PMSF. Protease inhibitor cocktail from Sigma may also be used with mammalian cells; add 100 ìl of the cocktail solution per 100 ml of lysis buffer, or to 1.25 mM if the protease inhibitor cocktail molarity is known.
Membrane lysis buffer A 50 mM Tris⋅Cl, pH 7.3 (APPENDIX 2E) 250 mM sucrose 2 mM EDTA 2 mM phenylmethylsulfonyl fluoride (PMSF; add from 0.5 M stock in DMSO or isopropanol) PMSF has a half-life in aqueous solution of ∼1 hr. Therefore, a stock solution of 0.5 M PMSF should be made in DMSO or isopropanol and should be added to the aqueous lysis solutions (50 mM Tris⋅Cl and membrane lysis buffer A) to a final concentration of 0.5 mM every hour.
Membrane lysis buffer B 40 mM Tris⋅Cl. pH 7.3 (APPENDIX 2E) 7 M urea 2 M thiourea 4% (w/v) n-octyl-β-D-glucopyranoside (OGL) 2% (w/v) CHAPS 2% (w/v) N-decyl-N,N-dimethyl-3-ammonio-1-propane-sulfonate (SB3-10; Eprogen) 10 mM TCEP (add from 0.5 M stock solution; see recipe) Store up to 1 month at 4°C
2-D Liquid Separations of Whole-Cell Lysates
Rotofor running buffer A sample recipe is given below for a running buffer containing 1% ampholytes; the actual final ampholyte concentration that will be loaded in the Rotofor cell should match the protein concentration as detailed in Table 23.3.1. The volume of water in the recipe should be scaled up or down accordingly when both the volume of cell
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lysate and the volume of ampholyte are changed to match the total amount of proteins in the sample. For 50 ml running buffer: Place 18.018 g urea into 23.4 ml distilled water (or other volume scaled according to amount of ampholyte added), stir until it dissolves; then add 7.612 g thiourea, 0.25 g n-octyl-β-D-glucopyranoside (OGL), 5 ml glycerol, 0.5 ml (or other volume to achieve appropriate ampholyte concentration; see Table 23.3.1) Bio-Lyte 3/10 ampholyte (Bio-Rad), and 200 µl 0.5 M tris(2-carboxyethyl)phosphine (TCEP) stock solution (see recipe). Store up to 1 month at 4°C. Since 1 g urea accounts for a volume of 0.75 ml and 1 g thiourea a volume of 1 ml, this contribution to the final volume should be considered when preparing the buffer solution. Therefore, one should begin with no more than 23.4 ml of distilled water, otherwise the final volume will exceed 50 ml, resulting in incorrect final concentrations of running buffer constituents. The dissolution of urea is an endothermic process. Heating the solution will speed the process. Thiourea itself has very low solubility; however, its solubility increases in urea solution. Because urea decomposes at high temperatures, keep the temperature below 50°C. OGL is a nonionic detergent, which helps solubilize membrane proteins and phospholipids. n-octyl β-D-galactopyranoside, an isoform of octyl-β-D-glucopyranoside (OGL), is also a good nonionic detergent, but it is much more expensive than the glucopyranoside isoform. Final concentrations are: 6 M urea, 2 M thiourea, 0.5% (w/v) OGL, 1% (v/v) ampholytes, 10% (v/v) glycerol, and 2 mM TCEP. Gloves are required when handling chemicals. TCEP, thiourea, and OGL are harmful for ingestion, inhalation, and skin contact.
TCEP stock solution, 0.5 M Dissolve 1.433 g tris(2-carboxyethyl)phosphine (TCEP) in 10 ml distilled water. TCEP should be handled in the hood with gloves. Inhalation, ingestion, or skin contact is harmful. Store at 4°C when not in use.
COMMENTARY Background Information Isoelectric focusing is a powerful technique in which proteins are focused to narrow bands under the influence of an electric field. A stable and uniform pH gradient, convection control, and pH measurement are three key factors for the success of any IEF method. In the Rotofor, a stable pH gradient is established by carrier ampholytes, which are composed of synthetic polyamino-carboxylic acids or polyamino-sulfonic acids with molecular weights of ∼300 to 900 (Osterman, 1984). Under the influence of an electric field, these amphoteric molecules migrate to their respective pI values and a pH gradient with increasing pH from anode to cathode is created. The high buffering capacity and conductivity of ampholytes prevent the distortion of the pH gradient when charged proteins are present; consequently, a stable and uniform pH gradient is obtained in the liquid phase. During the operation of the Rotofor, rotating the focusing chamber around the focusing axis
counteracts convection. Increasing the viscosity of the running buffer can further control convection. In gel-based IEF, convection is prevented by the use of a solid gel support. Convection is more efficiently eliminated because liquid molecules are restricted from moving. However, sample recovery from gels is more difficult, considering that focused proteins must be removed from the gel support for further analysis. Using the Rotofor, the focused proteins are still in solution phase; therefore, it is easy to extract proteins from the device without remixing. In the authors’ experience, the Mini-Rotofor chamber can fractionate >50 mg of proteins, and the standard Rotofor can fractionate gram-level samples. Compared to the maximum loadability of a gel, i.e., ∼5 mg, the Rotofor is at least 10 times higher. In addition, scale-up can be easily achieved by going from the mini-Rotofor to the preparative model, in which the volume of running solution is increased. These advantages of the Rotofor and the degree of fractionation achieved in the pH
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dimension make it an excellent method for prefractionation of complex protein mixtures. Since Bio-Rad released the Rotofor in the 1980s, it has been successfully used in isolating proteins from serum, cell cultures, tissues, and cerebrospinal fluid (Davidsson et al., 1999; Puchades et al., 1999; Wall et al., 2000; Davidsson et al., 2001; Gustafsson et al., 2001; Wall et al., 2001; Kachman et al., 2002). When coupled with other orthogonal techniques such as reversed-phase liquid chromatography and mass spectrometry, it can be used as an improved alternative to 2-D gel electrophoresis in profiling protein content in complex mixtures (Wall et al., 2000; Wall et al., 2001; Kachman et al., 2002; Wall et al., 2002; Wang et al., 2002). In the proteomic era, it will play an important role in sample preparation with its ability to enrich low-abundance proteins from complex biological mixtures.
Critical Parameters The most critical aspects of running the Rotofor include protein solubilization during operation, salt concentration, and temperature control. Once these parameters are optimized, excellent fractionation in the pH dimension can be achieved. The Rotofor has two different focusing chambers, standard and mini. The difference between them is sample volume and, consequently, the amount of sample that can be loaded. The standard Rotofor chamber holds 60 ml of solution containing milligrams to grams of total protein, while the mini-Rotofor can only hold 18 ml of sample with micrograms to milligrams of total protein. As a result, each pI fraction obtained from the standard Rotofor is ∼3 ml, while each pI fraction is 800 µl for the mini-Rotofor. In order to obtain the desired concentration of the focused protein, a protein assay (see UNIT 3.4) is necessary to determine the
total protein loaded and the volume into which it should be dissolved. In the Rotofor device, proteins are focused according to their pIs; however, the solubility of a protein reaches its minimum at the pI point. In order to prevent proteins from precipitating out of solution, it is necessary to break disulfide bonds as well as to disrupt protein-protein intramolecular interactions. Charged compounds would be a perfect choice to disrupt intramolecular interactions because they provide a hydrophobic microenvironment around protein molecules and electrostatic repulsion between molecules. Unfortunately, such charged compounds are not compatible with IEF, with NPSRP-HPLC separations, or with subsequent mass spectrometric analyses. Thus, nonionic chaotrope mixtures (e.g., urea and thiourea) and nonionic and/or zwitterionic detergents (e.g., octylglucoside) should be added to efficiently solubilize proteins (Rabilloud, 2000). The recommended chaotropes and detergents are listed in Table 23.3.3. The presence of protein disulfide bonds may also lead to aggregation and decrease in protein solubility. Disulfide bonds are disrupted by thiol compounds such as mercaptoethanol and dithiothreitol (DTT), and also by the nonthiol reducing compound tris(2-carboxyethyl)phosphine (TCEP). The latter compound is favored because it is an effective reagent with the added advantages of being less toxic and odorless. The loading of concentrated sample into the Rotofor requires a high ampholyte concentration to maintain a linear pH over the pH range and to keep proteins soluble at their pIs. Sample quantification is necessary to determine the optimal ampholyte concentration for the separation (see Table 23.3.1). In some cases, the solubility of the protein of interest may rely on ionic strength; in these instances, the ampholyte concentration can be increased.
Table 23.3.3 Recommended Solubilizing Agents for the Rotofor Systema
Nonionic detergents 0.1%-3.0% (w/v) digitonin 0.1%-3.0% (w/v) octylglucoside 0.1%-3.0% (v/v) Triton X-114 2-D Liquid Separations of Whole-Cell Lysates
Zwitterionic detergents 0.1%-3.0% (w/v) CHAPS 0.1%-3.0% (w/v) CHAPS
Reducing agents
Chaotropic agents
5-20 mM DTE
1.0-8.0 M urea
5-20 mM DTE
2 M thiourea
1-5 mM 2-ME
0.1%-2.0% (w/v) glycine 0.1%-2.0% (w/v) proline
2 mM TCEP aFrom the Bio-Rad Rotofor Instruction Manual.
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During the Rotofor separation process, the proteins at the focused zone tend to diffuse away because of the concentration gradient. This diffusion decreases the focusing effect and results in decreased resolution. If one considers that the diffusion rate is proportional to the temperature in solution, maintaining low temperature, hence low diffusion, is essential for improved resolution. In addition, many proteins, especially enzymes, are labile to temperature. Efficiently dissipating the heat generated during IEF is important to maintain the activity of proteins. Typically, in a Rotofor experiment, a water-cooling system (e.g., a closed-cycle chiller system) is connected with the sample chamber through a cooling finger. The temperature of the chiller should be set at the lowest temperature that prevents proteins from precipitating out of the running buffer This cooling improves both the resolution and the activity. In addition, the use of glycerol stabilizes the hydration shell around proteins, thus effectively solubilizing them. Furthermore, glycerol increases the viscosity of the sample solution, which results in less diffusion and improved resolution. The resolving power of IEF is proportional to the square root of the voltage gradient. When separating a sample with a high salt content, the high conductance of running solution will decrease the effective voltage across the sample and narrow the linear portion of the pH gradient. In this case, dialysis should be performed to desalt the sample before the Rotofor separation. The ionic strength should be controlled within 10 mM.
Troubleshooting Rotofor Maintaining proteins in solution phase is essential for operation of the Rotofor. Typical solubilizing agents include chaotropes, detergents, and reducing agents. Chaotropes disrupt protein-protein interactions; detergents provide a hydrophobic environment for proteins; reducing agents such as DTT, TCEP, and mercaptoethanol disrupt disulfide bonds. Protein focusing in the Rotofor is an electrophoretic process where high salt concentrations in the running solution affect focusing and generate heat. Therefore, all reagents added should be nonionic. The recommended solubilizing reagents for the Rotofor system are shown in Table 23.3.3. Air bubbles trapped in the Rotofor system can cause voltage fluctuations and an unstable
pH gradient. In the worst case, air bubbles may break the electrical connection. After the Rotofor cell is filled with sample solution and ampholytes, the whole assembly should be turned 180° and tapped on both electrolyte chambers to remove possible air bubbles trapped between the membrane and the sample. Air bubbles tend to be a more significant problem in the case of the mini-Rotofor than in the preparative device. In a successful Rotofor run, the pH gradient should be a linear curve in the middle fractions, as shown in Figure 23.3.2. If the ampholyte in the running buffer is insufficient, i.e., not enough buffer capacity, sample proteins can distort the pH gradient. This problem can be easily resolved by adding more ampholyte. Air bubbles and other contaminants in the system can be another source that results in distortion of the pH gradient. In addition, uneven sample harvesting of the Rotofor fractions after the separation will have the same result. Phosphoric acid (0.1 M) and sodium hydroxide (0.1 M) are recommended for the anode and cathode electrolyte, respectively. The diffusion of protons and hydroxide ions through the ion exchange membrane is inevitable in the focusing run, so that the first several channels maybe very acidic (<3) and the last channels very basic (>10). The result is that the pH gradient is distorted at both ends. One can improve the linearity of the pH gradient by using a solid-phase matrix, such as anion and cation exchange resins, at both ends. In a normal Rotofor run, the running current increases at the beginning then decreases with time. If the anode and cathode are inadvertently reversed, the current will increase with time and no current decrease is observed. Strong overlapping of proteins across pI lanes occurs if the Rotofor is overloaded or the sample becomes aggregated. A protein assay and an effective solubilizing solution are strongly recommended before the Rotofor run. Overlapping cannot be completely overcome without membrane separators between adjacent fractions. In order to minimize overlapping across lanes, prefractionated samples can be pooled and loaded onto the Rotofor cell for a second run. Additional ampholyte is not necessary since the prefractionated samples already contain ampholytes covering the pH range. Other separation techniques can also be applied to purify target proteins since the Rotofor provides samples that are sufficiently fractionated. Non-Gel-Based Proteome Analysis
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NPS RP-HPLC interfaced to ESI-MS Band broadening, decreased retention time, tailing, and elevated pressure are usually observed for “dirty” columns. These may result from irreversible adsorption of proteins and accumulated impurities from samples and solvents. Column cleaning after each separation is required for reproducible separation and prolonged column lifetime. Flushing the column in the reverse direction with 100% organic solvent such as acetonitrile or isopropanol is an effective means of removing hydrophobic substances and strongly retained molecules on top of the column bed. At least 10 CV of organic solvent should be applied, followed by running a blank gradient. The cleaning cycle should be repeated until no peaks are observed on the blank run. If the column pressure is irreversibly increased, sonicating the column frits in organic solvent may help. Microbiological growth can occur if the system and column are not handled correctly. When not in use, both system and column should be flushed and stored in 100% organic solvent.
8.5 including the subsequent salt gradient is usually greater than 80%. NPS RP-HPLC interfaced to ESI-MS A successful NPS-RP-HPLC separation of a first-phase IEF or CF fraction can typically separate at least 100 protein peaks in a 30-min run. The proteins that elute are generally purified proteins free of salt; however, the proteins are denatured because of the denaturing RPHPLC conditions. Since MS has the capability to resolve coeluting proteins and RP-HPLC concentrates proteins into narrow bands, the combination of LC and MS provides high resolution and sensitivity. The method has excellent reproducibility for detection of specific proteins. ESI-MS detects the intact MW of protein with high accuracy. The recovery of total protein is generally higher in such liquid-based separation techniques as compared to 2-D gels. The recovery of most proteins under 100 kDa from the NPS columns varies from 65% to 95% (Wall et al., 1999).
Time Considerations Anticipated Results Rotofor A successful Rotofor separation concentrates proteins into a fraction with a pH that is the same as the pI of the protein. The pH distribution is linear and shallow in the middle pH range, as shown in Figure 23.3.2, but is distorted at the two ends because of the diffusion of hydronium and hydroxide ions from the anode and cathode to the chamber. The fractionation is reasonably clean in the mid-pH range if the separation is not overloaded. Overlapping of proteins over a pH range is common in complex samples and especially for highly abundant proteins. Additionally, fraction overlapping occurs at the high pH due to cathodic drift. Often refractionation with the Rotofor or with other separation techniques (such as NPSRP-HPLC) is performed to purify the proteins for further analysis.
2-D Liquid Separations of Whole-Cell Lysates
Chromatofocusing A successful chromatofocusing run fractionates proteins according to their pIs. Proteins with higher pIs elute off the column earlier in the gradient. If run properly, a reproducible linear pH gradient for a given start and elution pH can be observed (See Fig. 23.3.2). One can obtain excellent fractionation to <0.1 pH unit for complex mixtures of proteins. The total protein recovery over a range from pH 4.0 to
Rotofor It takes ∼45 min to set up the Rotofor. A Rotofor run takes 4 to 5 hr for optimum separation of proteins across the pH range.Cleaning of the Rotofor device takes ∼1 hr. Chromatofocusing The time required for a CF experiment varies with the length of column, flow rate, complexity of the sample, and volume of sample. A higher flow rate makes the pH gradient steeper, resulting in a shorter run with lower pH resolution. At 0.2 ml/min elution using an HPCF 1D column, a typical separation takes ∼1 hr to complete. NPS RP-HPLC interfaced to ESI-MS A single NPS-RP-HPLC separation generally takes from 10 to 30 min. When coupled to the ESI-MS, a run time of ∼30 min is generally used. Cleaning the column after the separation may require 1 hr or longer depending on the complexity of sample and the lifetime of the column. For a relatively new column and simple protein mixtures, it takes ∼1 hr to clean the column; however, for an old column and complex samples, the authors recommend that the column be cleaned for 3 hr.
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Literature Cited Banks, J.F. and Gulcicek, E.E. 1997. Rapid peptide mapping by reversed-phase liquid chromatography on nonporous silica with on-line electrospray time of flight mass spectrometry. Anal. Chem. 69:3973-3978. Barder, T.J., Wohlman, P.J., Thrall, C., and DuBois, P.D. 1997. Fast chromatography and nonporous silica. LC-GC 15:918-926. Cech, N.B. and Enke, C.G. 2001. Practical implications of some recent studies in electrospray ionization fundamentals. Mass Spectrom. Rev. 20:362-387. Chen, H. and Horvath, C. 1995. High-speed highperformance liquid chromatography of peptides and proteins. J. Chromatogr. A 705:3-20. Chong, B.E., Lubman, D.M., Rosenspire, A., and Miller, F.R. 1999. Protein profiles and identification of HPLC isolated proteins of cancer cell lines using MALDI-TOF MS. Rapid Commun. Mass Spectrom. 13:1808-1812. Chong, B.E., Kim, J., Lubman, D.M., Tiedje, J.M., and Kathariou, S. 2000. Use of non-porous reversed-phase high-performance liquid chromatography for protein profiling and isolation of proteins induced by temperature variations for Siberian permafrost bacteria with identification by matrix-assisted laser desorption ionization time-of-flight mass spectrometry and capillary electrophoresis-electrospray ionization mass spectrometry. J. Chromatogr. B 748:167-177. Chong, B.E., Yan, F., Lubman, D.M., and Miller, F.R. 2001. Chromatofocusing nonporous reversed-phase high-performance liquid chromatography/electrospray ionization time-of-flight mass spectrometry of proteins form human breast cancer whole cell lysates: A novel two-dimensional liquid chromatography/mass spectrometry method. Rapid Commun. Mass Spectrom. 15:291-296. Davidsson, P., Westman, A., Puchades, M., Nilsson, C.L., and Blennow, K. 1999. Characterization of protein from human cerebrospinal fluid by a combination of preparative two-dimensional liquid-phase electrophoresis and matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Anal. Chem. 71:642-647. Davidsson, P., Paulson, L., Hesse, C., Blennow, K., and Nilsson, C.L. 2001. Proteome studies of human cerebrospinal fluid and brain tissue using a preparative two-dimensional electrophoresis approach prior to mass spectrometry. Proteomics 1:444-452. Fountoulakis, M., Langen, H., Gray, C., and Takacs, B. 1998. Enrichment and purification of proteins of Haemophilus influenzae by chromatofocusing. J. Chromatogr. A 806:279-291. Gooding, K.M. and Regnier, F.M. 1990. HPLC of Biological Macromolecules: Methods and Applications. Marcel Dekker, New York.
Gustafsson, E., Thoren, K., Larsson, T., Davidsson, P., Karlsson, K., and Nilsson, C.L. 2001. Identification of proteins from Escherichia coli using two-dimensional semi-preparative electrophoresis and mass spectrometry. Rapid Commun. Mass Spectrom. 15:428-432. Hanson, M., Unger, K.K., Mant, C.T., and Hodges, R.S. 1996. Optimization strategies in ultrafast reversed-phase chromatography of proteins. Trans. Anal. Chem. 15:102-110. Hutchens, T.W. 1998. Chromatofocusing. In Protein Purification: Principles, High-Resolution Methods, and Application, 2nd ed. (J.C. Janson, L. Ryden, eds.) pp. 149-174. John Wiley & Sons, New York. Kachman, M.T., Wang, H., Schwartz, D.R., Cho, K.R., and Lubman, D.M. 2002. A 2-D liquid separations/mass mapping method for interlysate comparison of ovarian cancers. Anal. Chem. 74:1779-1791. Kastner, M. 2000. Protein Liquid Chromatography. Elsevier, New York. Lee, W.C. 1997. Protein separation using non-porous sorbents. J. Chromatogr. B 699:29-45. Liu, Y.S. and Anderson, D.J. 1997. Gradient chromatofocusing high-performance liquid chromatography. 1. Practical aspects. J. Chromatogr. A 762:207-217. Mehlis, B. and Kertscher, U. 1997. Liquid chromatography mass spectrometry of peptides of biological samples. Anal. Chim. Acta. 352:71-83. Niessen, W.M.A. and Tinke, A.P. 1995. Liquid-chromatography mass-spectrometry: General principles and instrumentation. J. Chromatogr. A 703:37-57. Nilsson, C.L., Puchades, M., Westman, A., Blennow, K., and Davidsson, P. 1999. Identification of proteins in human pleural exudate using twodimensional preparative liquid-phase electrophoresis and matrix-assisted laser desorption/ionization mass spectrometry. Electrophoresis 20:860-865. Nilsson, C.L., Larsson, T., Gustafsson, E., Karlsson, K., and Davidsson, P. 2000. Identification of protein vaccine candidates from Helicobacter pylori using a preparative two-dimensional electrophoretic procedure and mass spectrometry. Anal. Chem. 72:2148-2153. Osterman, L.A. 1984. Methods of Protein and Nucleic Acid Research. Springer-Verlag, New York. Phamacia Biotech. 1985. FPLC Ion-Exchange and Chromatofocusing: Principles and Methods. Pharmacia Biotech AB, Uppsala, Sweden. Puchades, M., Westman, A., Blennow, K., and Davidsson, P. 1999. Analysis of intact proteins from cerebrospinal fluid by matrix-assisted laser desorption/ionization mass spectrometry after two-dimensional liquid-phase electrophoresis. Rapid Commun. Mass Spectrom. 13:2450-2455. Rabilloud, T. 2000. Proteome Research: Two-Dimensional Gel Electrophoresis and Identification Methods. Springer-Verlag, New York.
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Scopes, R.K. 1994. Protein Purification: Principles and Practice, 3rd ed. Springer-Verlag, New York. Sluyterman, L.A.A.E. and Elgersma, O. 1978. Chromatofocusing: Isoelectric-focusing on ionexchange columns. 1. General principles. J. Chromatogr. 150:17-30. Sluyterman, L.A.A.E. and Wijdenes, J. 1978. Chromatofocusing: Isoelectric-focusing on ion-exchange columns. 2. Experimental verification. J. Chromatogr. 150:31-44. Sluyterman, L.A.A.E. and Wijdenes, J. 1981a. Chromatofocusing. 3. The properties of a DEAE-agarose anion-exchanger and its suitability for protein separations. J. Chromatogr. 206:429-440. Sluyterman, L.A.A.E. and Wijdenes, J. 1981b. Chromatofocusing. 4. Properties of an agarose polyethyleneimine ion-exchanger and its suitability for protein separations. J. Chromatogr. 206:441-447. Strong, J.C. and Frey, D.D. 1997. Experimental and numerical studies of the chromatofocusing of dilute proteins using retained pH gradients formed on a strong- base anion-exchange column. J. Chromatogr. A 769:129-143. Wall, D.B., Lubman, D.M., and Flynn, S.J. 1999. Rapid profiling of induced proteins in bacteria using MALDI-TOF mass spectrometric detection of nonporous RP HPLC-separated whole cell lysates. Anal. Chem. 71:3894-3900. Wall, D.B., Kachman, M.T., Gong, S., Hinderer, R., Parus, S.J., Misek, D.E., Hanash, S.M., and Lubman, D.M. 2000. Isoelectric focusing nonporous RP HPLC: A two-dimensional liquid-phase separation method for mapping of cellular proteins with identification using MALDI-TOF mass spectrometry. Anal. Chem. 72:1099-1111.
Wall, D.B., Kachman, M.T., Gong, S., Parus, S.J., Long, M.W., and Lubman, D.M. 2001. Isoelectric focusing nonporous silica reversed-phase high-performance liquid chromatography/electrospray ionization time-of-flight mass spectrometry: a three-dimensional liquid-phase protein separation method as applied to the human erythroleukemia cell-line. Rapid Commun. Mass Spectrom. 15:1649-1661. Wall, D.B., Parus, S.J., and Lubman, D.M. 2002. Three-dimensional protein map according to pI, hydrophobicity and molecular mass. J. Chromatogr. B 774:53-58. Wang, H., Kachman, M.T., Schwartz, D.R., Cho, K.R., and Lubman, D.M. 2002. A protein molecular weight map of ES2 clear cell ovarian carcinoma cells using a two-dimensional liquid separations/mass mapping technique. Electrophoresis 23:3168-3181. Yang, Y.J. and Lee, M.L. 1999. Theoretical optimization of packed capillary column liquid chromatography using nonporous particles. J. Microcol. Sep. 11:131-140. Yu, J. and El Rassi, Z. 1997. High performance liquid chromatography of small and large molecules with nonporous silica-based stationary phases. J. Liq. Chrom. Rel. Technol. 20:183-201.
Contributed by Kan Zhu, Fang Yan, Kimberly A. O’Neil, Rick Hamler, and David M. Lubman The University of Michigan Ann Arbor, Michigan Linda Lin and Timothy J. Barder Eprogen, Inc. Darien, Illinois
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Quantitative Protein Analysis Using Proteolytic [18O]Water Labeling As the sequencing of many species’ genomes is completed, the study of the protein complement to the genome, the proteome, has emerged as a dynamic field of research. A common approach to characterizing the various states in which a protein exists is comparative proteomics. In this method, the relative amounts of proteins present in two or more samples are compared. An enzyme-catalyzed method for 18O labeling can be used to determine the relative amounts of the proteins present and is described in detail in this unit. Pools of proteins are enzymatically digested in parallel in [16O]- (i.e., standard deionized) and [18O]water. In the latter pool, two atoms of 18O are incorporated into the carboxyl terminus of each new peptide. Comparative proteomic studies are performed by mixing the pool of unlabeled peptide (generated in [16O]water) and the pool of isotopelabeled peptide (from [18O]water) and analyzing the peptide pairs by mass spectrometry. Relative quantification is derived from ratios of the isotope pairs. Tandem mass spectrometry experiments provide sequence information from the peptides, which is used to identify the proteins.
UNIT 23.4
BASIC PROTOCOL
This protocol contains optional, but recommended, steps to prepare a protein sample for proteolytic digestion and 18O labeling. This step will reduce and alkylate cysteine residues, as well as denature the protein. Alkylating the protein sample will allow more efficient digestion and avoid mixed disulfide bonds between digested peptides. Figure 24.3.1 gives an outline of the procedure. Note that both pools (i.e., labeled and unlabeled) must first be digested by trypsin before comparative proteomic investigation. Materials Protein samples 50 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E; prepare fresh) Urea (Sigma) 100 mM DTT (Sigma) 0.5 M iodoacetamide (Sigma) 10 mM Tris⋅Cl, pH 7.4 (APPENDIX 2E) Modified trypsin, sequencing grade (Promega) Immobilized trypsin on Poros (Applied Biosystems) 80% isotopically enriched H218O (stock is >95% 18O; Isotech)/20% acetonitrile 80% deionized H216O/20% acetonitrile Benchtop rotator BioSpin 6 Column (BioRad) IsoPro 3.0 software (http://members.aol.com/msmssoft) Additional reagents and equipment for mass spectrometry (Chapter 16) 1. Prepare the protein or proteins of interest in a freshly made solution of 50 mM Tris⋅Cl, pH 8, so that the concentration is ∼1 µg/µl. For the purpose of this protocol, the concentration is 100 ìg protein in 100 ìl buffer.
2. Add 0.024 g urea. The urea concentration is now 4 M.
3. Reduce cysteine residues by adding 10 µl of 100 mM DTT and rotating 1 hr at room temperature. The DTT concentration is ∼10 mM. Contributed by Kristy J. Reynolds and Catherine Fenselau Current Protocols in Protein Science (2003) 23.4.1-23.4.8 Copyright © 2003 by John Wiley & Sons, Inc.
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protein pool 1 (reduced, alkylated and desalted in 50 mM Tris pH 8)
protein pool 2 (reduced, alkylated and desalted in 50 mM Tris pH 8)
digest with trypsin overnight 37°C
digest with trypsin overnight 37°C
add immobilized trypsin
add immobilized trypsin
dry completely in vacuum centrifuge
dry completely in vacuum centrifuge
resuspend in H2 16O
resuspend in H2 18O
unlabeled peptide pool
18O 2
labeled peptide pool
rotate for several hours
rotate for several hours
Relative intensity
mix samples and analyze by mass spectrometry
m/z unlabeled peptide 18O2 labeled peptide
Figure 23.4.1 Schematic representation of the enzyme-catalyzed 18O-labeling strategy for comparative proteomics with digestion and labeling steps decoupled.
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4. Alkylate the reduced cysteine residues by adding 30 µl of 0.5 M iodoacetamide and rotate 1 hr in the dark at room temperature. The iodoacetamide concentration is approximately ten times the DTT concentration.
5. Desalt the reduced and alkylated protein sample using a BioSpin 6 column. Recover the sample in 10 mM Tris, pH 7.4 and pool the recovered samples. These columns are provided by the manufacturer in buffer solution. Directions are also provided about how to remove the buffer, as well as how to load, wash, and recover the protein sample. The manufacturer reports 90% to 95% recovery. One spin column can desalt 75 ìl, so two columns used in parallel are necessary. This procedure also removes excess iodoacetamide.
Perform initial protein digest 6. Add 20 µl of 50 mM Tris⋅Cl, pH 8, directly to one vial of 20 µg sequencing-grade modified trypsin The final trypsin concentration is 1 ìg/ìl.
7. Add 1 µl (i.e., 1 µg) dissolved trypsin per 50 to 100 µg protein in the sample This is 0.5 to 1 ìl when 100 ìg protein has been dissolved in 100 ìl (step 1). The remaining trypsin can be stored up to 6 months at −80°C in conveniently sized aliquots to avoid repetitive freeze-thaw cycles.
8. Check the pH to ensure that it is between 7 and 8. The pH can be adjusted by adding additional Tris⋅Cl buffer.
9. Incubate overnight at 37°C to allow complete digestion. The following section outlines the steps for incorporating two atoms of 18O into each peptide C-terminus (Fig. 23.4.1). Only one of the peptide pools in the comparative proteomic experiment is labeled with 18O; however, the procedure is applied to both pools in order to ensure experimental homology.
18O 2
16O 2
Relative intensity
100
I0
I4
75
100
M0
75 16O18O
50
I2
50
25
25
0
0
M2
M4 Mass/charge
Figure 23.4.2 Electrospray mass spectrum of a pair of doubly charged peptides, unlabeled (I0) and labeled (I4), and the theoretical molecular ion profile of the unlabeled peptide. (A) Observed isotopic distribution of 18O and 16O mixed samples. (B) Theoretical isotopic distribution without 18O labeling.
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Proteolytic 18O[water] labeling 10. Clean the immobilized trypsin by washing with 5 vol. water. Centrifuge in a benchtop centrifuge 1 min at 8000 rpm and discard the supernatant. Repeat two more times. Immobilized trypsin must be cleaned prior to use.
11. Retrieve the digested protein sample from the 37°C water bath (step 9). Add an amount of immobilized trypsin to the peptide sample equal to ∼20% of the sample volume. This is ∼20 ìl if 100 ìg protein was dissolved in 100 ìl buffer (step 1).
12. Dry the peptide and immobilized trypsin mixture completely in a vacuum evaporator (∼1 hr at room temperature). Confirm by visual inspection. Adding the trypsin beads before drying the sample allows for more homogeneous mixing.
13. Resuspend the pool to be labeled in a mixture of 80% isotopically enriched [18O]water/20% acetonitrile to a final volume of 100 µl. Resuspend the unlabeled peptide counterpart in a mixture of 80% [16O]water/20% acetonitrile]. The organic solvent enhances trypsin efficiency. Store remaining H218O protected from dilution by ambient moisture up to 2 years at −20°C.
14. Rotate ∼5 hr at room temperature. Recover the peptides in the supernatant following a 2 min microcentrifugation at 8000 rpm. Additional or less time may be necessary depending on the particular peptides in the sample.
15. Mix the labeled and unlabeled peptide pools and analyze by mass spectrometry (Fig 23.4.2; also see Chapter 16). Samples can be analyzed by nanospray, electrospray, LC-MS, or MALDI. The method is compatible with most commercial mass spectrometers as long as they are able to resolve 4 Da. The mixing ratio can be varied from 1:1 and taken into account in the calculations.
Interpretation and protein quantitation 16. Calculate values for the theoretical peak areas for the monoisotope peak of a peptide with known composition (M0), the peak 2 Da higher (M2), and the peak 4 Da higher (M4) for each peptide using IsoPro 3.0 software. The elemental composition and charge state are entered into the software program and the program calculates the theoretical isotopic distribution. The distribution of 13C isotopes in the unlabeled and labeled peptides is assumed to be identical.
17. Calculate the ratios of [18O/16O] peptides using the following equation. I4 − Ratio 1 =
M4 M I0 − 2 M0 M0
M2 1 M2 I0 + I2 I0 I2 − M0 2 M0 I0
where I0, I2, and I4 are the observed peak areas for the monoisotopic peak for the peptides without 18O label, the peak 2 Da higher, and the peak 4 Da higher (double 18 O labeled peak), respectively. Quantitative Protein Analysis Using Proteolytic [18O]Water Labeling
This equation takes into account not only the ratio of labeled and unlabeled peak areas, but also contributions from incomplete labeling and corrects for the theoretical isotopic distribution of a peptide.
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unlabeled protein O
O– protease
H218O
18OH 2
18O– + H N 2
NH
NH R′
single 18O-labeled peptide O
O
R′
R′
serine-enzyme double 18O labeled peptide 18O–
18O–
OH
16 18O + H2 O
R′
R′
O
18OH 2
protease
O
18O
H218O R′
serine-enzyme
Figure 23.4.3 Schematic representation of the mechanism of double single step.
18
O incorporation with digestion and labeling in a
If the peptide sequence is not known, an estimate of the 18O/16O ratio can be made by taking the ratio of the peak areas of I4 and I0.
COMMENTARY Background Information Enzyme-catalyzed 18O labeling enables relative protein quantitation in high throughput proteomic strategies. Historically, comparative proteomic studies have been carried out using two-dimensional gel electrophoresis (UNITS 10.4 & 22.1). Currently, there is a push to find alternatives to two-dimensional gel electrophoresis because of its limited dynamic range, loss of hydrophobic proteins, and tedious procedures. Enzyme-catalyzed 18O labeling provides highly specific isotopic labeling of all but one of the peptide products from each protein. Labeling has been demonstrated to be complete and stable (Schnolzer et al., 1996; Yao et al., 2001; Reynolds et al., 2002). The stability of the label makes the method compatible with multiple separation techniques that can add to the diversity of the experiment (e.g., affinity chromatography to isolate cysteine-containing peptides). The double 18O label can be introduced by various enzymes including trypsin, Lys-C, chymotrypsin, and Glu-C, as long as the same proteolytic enzyme is used to generate the peptides. The mechanism of incorporation is shown schematically in Figure 23.4.3. The enzyme covalently bonds the target residue and
the intermediate then reacts with [18O]- or [16O]water. Double 18O labeling is achieved because the enzyme rebinds the C-terminal residue of the peptide product. In order to achieve complete double 18O labeling, the sample must be digested in highly enriched (>95%) [18O]water. Comparative proteomic studies are carried out by incubating one pool of proteins or peptides with an enzyme in [16O]water (unlabeled) and the other pool with the same enzyme in [18O] (double 18O labeled). The two pools are then mixed and analyzed by mass spectrometry. The labeled and unlabeled peptides co-elute chromatographically and appear as isotopic doublets in the mass spectrometer (Fig. 23.4.2). Therefore, the relative abundance of each peptide can be directly quantitated based on peak areas. The double 18O label is located at the peptide C-terminus. This label aids peptide sequencing in tandem mass spectrometry experiments because the y-ions, or C-terminicontaining ions, carry the double label and can be readily identified in the mass spectrum. It then becomes trivial to walk down the sequence by following the y-ion doublets. The parent protein is then identified by querying the se-
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glycosylated peptide unlabeled 100 triple 18O labeled
Relative intensity
75
non-glycosylated peptide double 18O unlabeled labeled
50
25
0 468
470
1239 Mass/charge
1242
1245
1248
Figure 23.4.4 Electrospray mass spectrum of two peptides, which were deglycosylated and digested in [18O]water in parallel with the [16O] counterparts. A triply charged peptide that did not contain a glycosylation site, and a singly charged peptide that had been glycosylated are detected with the respective 4 and 6 Da mass increments. (Reynolds et al., 2002) Reprinted with permission.
Quantitative Protein Analysis Using Proteolytic [18O]Water Labeling
quence information in a protein database (e.g., Mascot, Matrix Science). Previous studies on 18O proteolytic labeling performed digestion and labeling in one step (Yao et al., 2001; Reynolds et al., 2002). The current method, detailed in this unit, decouples the digestion and labeling steps (Yao et al., 2003). The decoupling adds to the versatility of the method because each step can be optimized individually. For example, if a high urea concentration is necessary to maintain a protein in solution, the sample could be digested by Lys-C and then the more soluble peptides could be labeled by trypsin. Most importantly, the twostep procedure eliminates the need to redissolve dried proteins in [18O]water before the label is incorporated. This method has also been applied to determ ine sites o f N- linked glycosylation (Reynolds, et al., 2002). When N-linked sugars are enzymatically cleaved from a protein by glycopeptidase [18O]water, a single 18O atom is incorporated. Following further digestion in [18O]water by trypsin, Glu-C, or other equivalent enzymes, the peptides that contained Nglycosylation sites have three 18O labels and are readily recognized by mass spectrometry, as
shown in Figure 23.4.4. The site can usually be located by mass changes in the sequence ions from a tandem mass spectrometry experiment. In summary, proteolytic 18O labeling is a flexible, global, and robust strategy for comparative proteomics. Currently, this method is being employed in the authors’ laboratory to compare the proteomes of drug-susceptible and -resistant breast cancer cell lines.
Critical Parameters In this unit, proteolytic 18O labeling is described using trypsin to catalyze isotope incorporation. (Other proteases are known to catalyze double 18O incorporation, including Lys-C and Glu-C.) Since these enzymes are necessary to digest the protein and catalyze isotope incorporation into the peptide products, the sample conditions must be compatible with the enzyme—i.e., proper pH and salt concentrations should be used to keep the protease folded and functional. The volumes and concentrations described in this unit may be scaled up or down to suit the experiment.
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18O 2
16O 2
100
Relative intensity
75
50
25
0 570
572
574
576
578
580
Mass charge DoxR/susceptible:1.2
Figure 23.4.5 Electrospray mass spectrum of doubly charged N-terminally acetylated peptide ASGVAVSDGVIK from human cofilin isolated from MCF-7 human breast cancer cell lines. The unlabeled peptide was isolated from drug-susceptible cells, while the double 18O labeled peptide was isolated from doxorubicin-resistant cells. The ratio of cofilin in the two cell lines was calculated as resistant/susceptible: 1.2 ± 0.2.
Troubleshooting
Time Considerations
If complete double 18O incorporation is not observed, additional incubation time or addition of a second aliquot of enzyme may be necessary. Also, check the pH to ensure that the enzyme is functional.
The labeling step has been evaluated using trypsin with a mixture of peptides terminating in lysine or arginine (Yao et al., 2003). The time required for 95% incorporation of two 18O atoms varied from 5 to 90 min.
Anticipated Results
Literature Cited
This protocol should produce a set of proteolytic peptides, all of which, other than the peptide containing the original C-terminus, are labeled >95% with two 18O atoms. The 18O2 labeled peptides can then be mixed with the unlabeled pool and relative quantitative information derived by mass spectrometry. The procedure has been successfully applied to study adenovirus strains using MALDI-TOF mass spectrometry (Yao et al., 2001). The method has also been used to quantitate changes in phosphorylation (Bonefant et al., 2003). Figure 23.4.5 shows an electrospray mass spectrum of the human protein cofilin isolated from a human breast cancer cell line. The 18O2 labeled peptide was isolated from drug-susceptible cells, while the unlabeled peptide counterpart was isolated from doxorubicin-resistant cells. The peptides were observed in a 1:1 ratio, so no change in regulation was observed.
Bonenfant, D., Schmelzle, T., Jacinto, E., Crespo, J.L., Mini, T., Hall, M.N., and Jeno, P. 2003. Quantitation of changes in protein phosphorylation: A simple method based on stable isotope labeling and mass spectrometry. Proc. Natl. Acad. Sci. U.S.A. 100:880-885. Reynolds, K.J., Yao, X., and Fenselau, C. 2002. Proteolytic 18O Labeling for Comparative Proteomics: Evaluation of Endoprotease Glu-C as the Catalytic Agent. J. Proteome Res. 1:27-33. Schnolzer, M., Jedrzejewski, P., and Lehmann, W.D. 1996. Protease-catalyzed incorporation of 18O into peptide fragments and its application for protein sequencing by electrospray and matrixassisted laser desorption/ionization mass spectrometry. Electrophoresis. 17:945-953. Yao, X., Freas, A., Ramirez, J., Demirev, P.A., and Fenselau, C. 2001. Proteolytic 18O labeling for comparative proteomics: Model studies with two serotypes of adenovirus. Anal. Chem. 73:28362842. Non-Gel-Based Proteome Analysis
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Yao, X., Afonso, C., and Fenselau, C. 2003. Dissection of proteolytic 18O labeling: Endoproteasecatalyzed 16O-to-18O exchange of truncated peptide substrates. J. Proteome Res. 2:147-152.
Contributed by Kristy J. Reynolds and Catherine Fenselau University of Maryland College Park, Maryland
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Protein Microarrays
UNIT 23.5
Recent technological advances have resulted in the miniaturization of ligand-binding assays, and highly sensitive systems have been developed that allow the quantification of an analyte from a minute amount of sample. The ability to miniaturize a large number of assays gives rise to the possibility of designing systems capable of determining thousands of analytes in parallel within a single experiment. Such miniaturized and parallelized analytical systems were initially employed to study gene expression, and the terms DNA chip and DNA microarray were used to describe the technology. While these initial experiments showed the exceptional potential of microarrays, their use is not limited to generating nucleic acid expression patterns. In recent years, it has been demonstrated that parallel analysis of proteins is feasible, and systems using proteins as immobilized capture molecules have been developed. Although the problems encountered when working with proteins are different, the underlying principle of a solid-phase binding assay is the same for both DNA and protein microarrays. Protein microarrays can be powerful tools that are capable of simultaneously determining a large number of parameters from complex samples. These samples are incubated on arrays of numerous microspots, each of which contains a single, well-defined immobilized capture molecule. The most commonly used format is the antibody microarray, which is used to identify and quantitate target proteins of interest. Novel applications include protein affinity assays in a highly parallel mode and protein interaction studies. The different types of binding molecules immobilized in microspots include proteins, peptides, and low-molecular weight compounds. The ability to identify post-translational modifications shows that protein array-based methods go beyond simple expression analysis, and systems that are capable of determining enzymatic activity on arrays of immobilized substrates allow detailed protein characterization. These examples show that microarrays composed of proteins have developed into analytical tools, and such microarrays have recently begun to play a major role in the analysis of proteomes. The design and generation of specific capture microarrays depends on knowledge of the target analytes and on the availability of corresponding highly specific capture agents. With DNA or RNA, it is possible to predict and to synthesize in vitro a nearly infinite number of high-affinity and high-specificity capture molecules—namely, the complementary strands of the nucleic acids of interest. For proteins, such specific capture reagents are often not available, and the generation of specific binding molecules is time consuming and expensive. Today, the most applicable molecules are antibodies, and while several interesting alternatives are under development, most protein microarrays rely on the use of antibodies as immobilized capture reagents. As a result, the number of binding events that can be monitored in one experiment is dependent on the number of capture molecules used, and is low compared to that possible with DNA chips. Miniaturized and multiplex immunoassays can be regarded as excellent solutions for applications in which several parameters of a single sample must be analyzed in parallel. For large-scale proteomics research, it can further be assumed that high-content protein microarrays will accelerate the identification of new biomarkers. In contrast to such highcontent multiplex analyses, diagnostic assays usually focus on a small, clearly defined set of analytes. For example, in immune or allergy diagnostics, it is necessary to screen patient sera for the presence or absence of a relatively small number of different types of autoantibodies or antibodies directed against parasitic and viral antigens or allergens. In particular, in the field of autoimmune diagnostics, protein microarrays offer an excellent option when patient sera have to be screened for the presence or absence of different types Contributed by Stefan Kramer, Thomas O. Joos, and Markus F. Templin Current Protocols in Protein Science (2005) 23.5.1-23.5.20 C 2005 by John Wiley & Sons, Inc. Copyright
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of autoantibodies. Autoantigens used as diagnostic markers for autoimmune diseases (e.g., systemic rheumatic diseases) can be immobilized in a microarray, and different types of autoantibodies can be accurately quantified from <1 ml of patient serum. Technically, such diagnostic applications closely resemble the procedures that are required for setting up an array-based proteome analysis, since the principle (namely, the observation of a binding reaction at a site containing an immobilized capture molecule) is the same. Microarray-based assay systems are used to carry out solid-phase binding assays and share many characteristics with well-known macrosystems, like 96-well ELISA experiments. Actually, protein assays in ELISA format are good candidates for miniaturization and are likely to work in an array system. While the difference in size is obvious, the miniaturization of the assay does not result in lower detection limits. It is one of the remarkable properties of microarray-based systems that miniaturization can actually lead to an increase in sensitivity. The “microspot concept” developed by Ekins et al. (1989) shows that a ligand-binding assay achieves its highest sensitivity when capture molecules are immobilized to a solid phase only within a very small area. The diameter of a microspot, the single assay point found in an array, is typically <200 µm, and hundreds of spots can therefore be immobilized in an area of one square centimeter.
Generation of protein microarrays The production of arrays of microspots requires expensive arraying robots, and the readout of the chips is performed using sensitive chip readers that possess a high spatial resolution. Due to the high level of interest in DNA microarrays, dedicated equipment has been developed over the last 10 years, and the systems now available show a high degree of maturity. The impact of the results generated with DNA chips has led to the establishment of DNA microarray core facilities at a growing number of universities and companies. Since most of the equipment required to perform a protein microarray experiment is present in these service units, it has become possible for many investigators to consider microarrays as an experimental approach to solving scientific problems. Support Protocol 1 describes how protein microarrays can be generated using the equipment found in laboratories working with DNA microarrays. Different technologies are used to generate microspots on a solid support. Most common are contact printing devices that use steel needles to transfer a small volume of a solution (<0.5 nl) to an activated support material. A second commonly employed arraying system uses non-contact printers that are similar to the ink-jet printers found in an office; in such systems, small droplets of liquid are shot toward the array support. The use of protein solutions is possible in both types of systems, and, in most cases, no significant problems are encountered, despite the fact that the machines were constructed for arraying nucleic acids. The manuals accompanying the equipment should be consulted, as standard printing protocols developed for DNA are likely to work with proteins as well. Nevertheless, some of the additives frequently used to stabilize proteins in solution can produce problems during the arraying process. Detergents can lead to foam formation, which results in missing spots, and solutions with a high viscosity (e.g., solutions with a high protein or glycerol concentration) can also present problems, especially for non-contact printing devices.
Substrates
Protein Microarrays
An important issue in the design of protein microarrays is the choice of substrate on which the microspots are to be printed. Most of the commercially available supports used for the immobilization of capture molecules are made of glass, but plastic and polymer membranes can also be used. The immobilization of proteins occurs through noncovalent protein-surface interactions with hydrophobic (e.g., nitrocellulose, polystyrene) or
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positively charged (e.g., polylysine, aminosilane) surfaces. Covalent attachment using a variety of chemically activated surfaces (e.g., aldehydes, epoxies, active esters) is also possible, as is attachment via specific bimolecular interactions (e.g., streptavidin-biotin, polyhistidine-nickel chelation) in the generation of protein microarrays.
Array stabilization measures A very important issue during microarray production is the post-print processing of the generated arrays. In contrast to DNA arrays, the immobilized proteins are susceptible to denaturation, which leads to the loss of function of the capturing reagent. Effective procedures for stabilizing proteins immobilized on a solid support have been developed for coated microwell plates used in ELISA-type assay systems. The strategies for stabilizing protein microarrays are similar, and arrays that can be stored over a period of two to three months can easily be produced. Stabilization procedures are usually combined with a blocking process, which is important for improving the performance of the microarray. By blocking the unsaturated binding sites on the substrate surface, the background signal of the assay can be reduced significantly, and an optimized blocking protocol can lead to a gain in sensitivity of one order of magnitude.
Applications of protein microarrays The possibility of detecting small amounts of a number of different analytes from a few microliters of sample in one experiment makes the use of microarrays attractive for analytical purposes. This unit focuses on the use of microarrays consisting of defined capture molecules, namely antibodies and purified antigens. The degree of multiplex (i.e., the number of different analytes determined simultaneously in one experiment) for the described setup is usually limited to several dozen different binding reactions, due to the low number of specific capture molecules available. This unit deals with two types of assays that, while technically very similar, address different types of questions. Support Protocol 1 describes the immobilization of a number of defined proteins in an array format to allow screening for the presence of an expected type of binding molecule. In addition, the arrangement of a parallelized autoimmune diagnostic assay in which known autoantigens are immobilized is shown. Basic Protocol 1 presents an array-based approach to serum diagnostics that allows the parallel detection of a number of specific immunoglobulins from the immunoglobulin pool found in serum. Detection is performed using a single detection reagent—namely, an antibody specific for the bound autoantibodies. Such an assay presents an example of a simple multiplex system that contains a variety of arrayed proteins and detects binding to the capture proteins using one detection reagent. Comparable systems are used to detect infection or allergy (Fall et al., 2003; Harwanegg et al., 2003; Bacarese-Hamilton et al., 2004). A variation of the setup has also proven to be useful for characterizing monoclonal antibodies with respect to specificity and cross-reactivity (Lueking et al., 1999). The Alternate Protocol discusses the use of parallelized quantitative sandwich immunoassays. In such assays, a collection of specific antibodies is immobilized in the array and, after sample application, the corresponding set of detection antibodies is added to quantify the amount of analyte present in the sample. This type of setup has been termed micro-ELISA, due to the similarity of the system to the common sandwich ELISA. The major effort in setting up such a system is in the selection of antibodies. While a given assay, considered individually, may work in the miniaturized format, the combination of such assays very often leads to a significant loss in sensitivity, due to the cross-reactivities of the antibodies employed.
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Other major applications of protein microarrays (namely, parallelized screening for protein-protein interactions and the arraying of complex protein mixtures for biomarker identification) are discussed briefly in the Commentary. BASIC PROTOCOL
IMMUNOASSAY USING PROTEIN MICROARRAYS: DETECTION OF AUTOANTIBODIES Protein microarrays are versatile tools that can be adapted to a variety of applications. This protocol, which gives an example of the use of microarrays consisting of defined capture molecules, provides an array-based approach to serum diagnostics and is used to detect autoantibodies from human serum in an immunological assay. More than 25 different autoantigens have been used to generate microarrays using this protocol (Joos et al., 2000; Robinson et al., 2002). An example of an array layout consisting of 20 different autoantigens is given in Figure 23.5.1, and the antigens chosen can easily be adapted to the requirements of the assay. The assay procedure closely resembles the immunochemical staining procedures established in many immunological and histological laboratories. Actually, many of the protocols used in these laboratories can be adapted to protein microarrays. The generated readout is a fluorescence signal that is quantified using a dedicated microarray scanner. Therefore, the type of equipment found in laboratories that work with DNA microarrays must be used.
Figure 23.5.1 Schematic picture of a protein microarray. The drawing shows a standard arraying substrate (7.5 cm × 2.5 cm) that has been used to create a protein microarray consisting of 12 identical subarrays. Each of the subarrays in this example consists of 144 microspots measuring 100 to 200 µm in diameter. Protein Microarrays
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Materials PBS (APPENDIX 2E) Wash buffer: 0.1% Tween 20 (v/v) in PBS (store up to one month at 4◦ C) Serum samples to be analyzed Dilution buffer (see recipe) Detection antibody (biotinylated goat anti–human IgG antibody; Jackson Dianova) 5 µg/ml Cy5-labeled streptavidin (Jackson Dianova, Prozyme) in dilution buffer ProPlate multiarray slide module (Grace Bio-Labs, distributed by Invitrogen) Protein microarrays consisting of various autoantigens (see Support Protocol) Plate shaker Compressed (oil-free) air or nitrogen source Microarray scanner (see Support Protocol) NOTE: All steps are performed at room temperature. The use of a plate shaker set at 150 rpm is recommended throughout the entire procedure.
Perform immunoassay 1. Attach the ProPlate slide module to a microarray-containing slide according to the manufacturer’s recommendations. Each module has a gasket that forms 16 wells around 16 separate arrays (each consisting of up to 300 spots) such that a maximum of 16 assays can be performed in parallel on one slide. This gasket significantly simplifies the handling of slides during the assay. The distances between the wells are compatible with the microtiter plate format; therefore, standard pipetting equipment, such as multichannel pipettors, can be used to process the protein microarrays. ProPlate trays that hold up to four slide modules are available. These trays have the footprint of a microtiter plate and are compatible with robotic fluid-handling devices.
2. Rehydrate the arrays by pipetting 100 µl PBS into each subarray and allowing to sit for 30 min. Briefly rinse the subarrays three times by pipetting with wash buffer and then add 100 µl wash buffer to each subarray. Incubate 5 min at room temperature, mixing several times by gently shaking slides at ∼150 rpm on a plate shaker. A subarray framed with a disposable gasket requires 25 to 100 µl for incubation; during rehydration and washing, the use of 100 µl is recommended. Always use fresh wash buffer for each wash step. When processing multiple subarrays, it is most convenient to use a multichannel pipettor. After the rehydration step, the arrays must not be allowed to dry, as drying will lead to a significant increase in background signal.
3. Dilute the serum samples 1:200 in dilution buffer. Remove the wash buffer from each subarray by pipetting, and then add 25 to 40 µl of diluted serum. Depending on the expected amount of autoantibody in the sample, the dilution can be varied from 1:50 to 1:10,000. The ratio 1:200 has been chosen because it very often allows clear identification of low-positive samples (i.e., samples containing autoantibodies that, upon binding, yield low fluorescence signals). Although a 1:200 serum dilution may result in a low signal intensity, a signal that is clearly above background is expected for a positive sample at this dilution.
4. Incubate the slides, while shielded from light, for 30 to 60 min at room temperature. Prolonged incubation and elevated incubation temperatures (e.g., 37◦ C) increase detection signals by allowing the formation of more antibody-antigen complexes. Care should be taken to avoid sample evaporation. To this end, the use of a lid or a sealing film is recommended for longer incubation times.
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5. Use a pipet to remove the sample solution from each subarray, taking care to avoid contamination of the neighboring subarrays. Rinse each subarray five times with 100 µl wash buffer, then apply 100 µl wash buffer to each subarray, and incubate for 5 min. The wash procedure is important for maximizing the quality of the results obtained; poor washing leads to a significant increase in background and therefore limits the dynamic range of the assay.
6. Dilute the biotinylated anti–human IgG antibody to a final concentration of 2.5 µg/ml in dilution buffer. Remove the wash buffer from each framed: Subarray by pipetting, and then overlay each subarray with 40 to 70 µl of the diluted detection antibody and incubate 45 min at room temperature. Secondary antibodies are generally used at concentrations between 0.5 and 5 µg/ml; 1 µg/ml is usually a good starting concentration.
7. Remove the antibody solution from each of the subarrays and wash as in step 5. 8. Remove wash buffer and incubate arrays with 40 to 100 µl 5 µg/ml Cy5-labeled streptavidin solution for 20 min. Cy5 has been chosen as a fluorophore label because it is widely used for microarray applications. On planar arrays, it often results in a lower background compared with the analog dye Cy3. Other dyes (especially the Alexa dyes from Molecular Probes, and R-phycoerythrin) can also produce satisfactory, if not superior, results.
9. Perform a final wash step as described in step 5, remove slides from slide modules, and rinse slides twice (10 sec per rinse), first with tap water and then with distilled water. 10. Dry the slides in a stream of oil-free air or nitrogen.
Detect and evaluate fluorescence signals 11. Analyze the dry slides with a microarray scanner using the channel that corresponds to the fluorophore used. Consult the manual accompanying the microarray scanner for proper settings with regard to parameters such as wavelength, filters, laser power, and photomultiplier gain. Care must be taken to avoid saturation in the generated array image. Even in the most intense spots, the signal must not exceed half the maximum value. For the 16-bit TIFF file that is normally generated, values >30,000 units must be avoided. Data analysis requires software that is capable of identifying microspots and quantifying signal intensity. Such software is usually part of the scanner software package and can be found at DNA microarray facilities. ALTERNATE PROTOCOL 1
Protein Microarrays
PROTEIN-PROTEIN INTERACTION STUDIES ON PROTEIN MICROARRAYS Conventional biochemical methods such as immunoprecipitation coupled with immunoblotting are of limited use when screening for novel protein-protein interactions, since such methods allow the analysis of only a small number of possible interactions at one time. As a result, genetic screens (e.g., the yeast two-hybrid system) have had an enormous impact, making it possible to generate huge data sets and integrate these data into interaction maps of high complexity (see Chapters 19 and 20). A second method that allows screening for protein-protein interactions on a large scale involves a two-step affinity precipitation procedure coupled to mass spectrometry for exploring the composition of protein complexes (Rigaut et al., 1999). Protein microarray technology represents a complementary method for studying protein interactions in vitro. Microarray-based
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studies of protein-protein interactions require purified proteins, which are usually obtained as tagged fusion proteins from cloned genes. By arraying sets of recombinant proteins in the microarray format, dozens to thousands of potential capture molecules are immobilized such that binding to a defined reaction partner, a protein in solution, can be detected. If sets of purified proteins are available, the generation of arrays can be performed as described in the Support Protocol. Alternatively, prefabricated protein microarrays can be purchased and used for the detection of protein-protein interactions. Commercially available arrays include the Yeast Protoarray PPI Microarray (Invitrogen), which consists of 5800 expressed reading frames from yeast, and the ProtoArray Human Protein Microarray (Invitrogen), made from 1800 expressed human cDNAs. To probe the interaction arrays, a purified protein is added, and binding to the interaction partners must subsequently be detected. The method described in Basic Protocol 1 uses an antibody against the binding partner from the fluid phase, and if antibodies raised against the protein of interest are available, then the procedure outlined in that protocol remains applicable. Very often, such antibodies are not available, and alternative methods therefore have to be employed. The use of recombinantly expressed proteins that carry a detectable tag is the most convenient method for detecting binding events. Here, an antibody against the tag is used as the detecting reagent, and again, Basic Protocol 1 can be used to generate a fluorescent signal on the microarray. A second set of recombinant fusion proteins that corresponds to the set immobilized on the array can be especially useful, provided that this second set contains a tag different from the one found in the immobilized set. This strategy allows one to probe each of the proteins against all members of the set without changing the detection system. More importantly, it also allows the generation of a second type of array that consists of the same proteins, and this array can be used to confirm positive interaction results in a reciprocal experiment. As an alternative, the proteins used as probes can be tagged directly. In particular, biotinylation procedures (UNIT 3.6) have proven to be reliable and convenient.
PARALLELIZED SANDWICH IMMUNOASSAYS ON PROTEIN MICROARRAYS
ALTERNATE PROTOCOL 2
The miniaturization and parallelization of sandwich immunoassays is possible, and systems have been built that are capable of quantifying more than 20 different analytes on protein microarrays with high sensitivity. Some of the principal problems that have to be solved when setting up such systems are known from the development of the standard sandwich immunoassay. The identification of an appropriate pair of antibodies for a specific and sensitive assay can be time consuming. When such a pair is available, implementation of the assay typically becomes a routine process. This is also true for the miniaturization of a sandwich immunoassay. With the availability of a matched antibody pair, the microspot assay can usually be set up without significant problems. Parallelization presents a different problem, since every capture antibody in the array requires its own specific detection antibody. Therefore, even for low-complexity arrays (i.e., those with <50 different capture molecules), cross-reactivity between antibodies and between different analytes becomes a major issue. Combination of assays very often leads to a significant loss in sensitivity, and the inclusion of assays for certain analytes can turn out to be very difficult. Nevertheless, it often is possible to combine 10 or more analytes on a microarray and still obtain sensitive measurements from a 10-µl sample. In these types of assays, the major advantage typically is the small amount of sample required, rather than the high degree of multiplexing that is achievable using DNA microarrays. The procedure for performing a sandwich assay is essentially identical to the Basic Protocol. The only difference is the number of detection antibodies required for the assay.
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SUPPORT PROTOCOL
GENERATION OF PROTEIN MICROARRAYS The production of microarrays on chip surfaces is done using dedicated microspot arrayers. The most commonly found arraying robots are contact printers equipped with needles and ink jet printers that deposit pico-to-nanoliter droplets on the chip surface. Both types of printers are suitable for generating protein microarrays. The manuals accompanying the available arrayer should be consulted, and a standard printing protocol developed for DNA should be chosen for the printing of proteins. An important issue when planning an experiment is the design of the array. It is desirable to spot two to four replicates of each immobilized protein and to include different control spots that allow one to monitor the performance of the assay (e.g., a dilution series of the detection antibody for verification of the detection system). A useful feature in any array is the presence of marker spots. These spots should be easily detectable, and they usually are made using dye-coupled proteins (fluorophore-labeled antibodies or BSA). The marker spots, which are typically positioned in the corners of the array, allow determination of the orientation of the array. For most arrays, spots are printed with 375 or 500 µm centerto-center spacing in both the X and Y directions. This spacing allows the generation of several protein microarrays on a 2.5 × 7.5–cm arraying substrate. A proposed layout for a microarray consisting of 12 subarrays is shown in Figure 23.5.1. In this layout, four replicates of the immobilized proteins are placed adjacent to each other, and marker spots are located in the corners indicate the orientation of the array. In protein microarrays, a spacing of 4 to 6 mm between adjacent subarrays has proven to be useful, since this allows the separation of the arrays by a mechanical barrier and the processing of 12 individual samples on subarrays each consisting of >100 spots. During the planning of the assay, a substrate for the protein microarray must be chosen. A wide variety of substrates are commercially available (Table 23.5.1), and while they differ in terms of their composition and surface chemistry, all can produce excellent results. Most of the supports used for the immobilization of capture molecules are made of glass, but plastic slides and polymer membranes are also available. The immobilization of proteins is often done using noncovalent protein-surface interactions. Covalent attachment using a variety of chemically activated surfaces (e.g., aldehydes, epoxies, active esters) is also common, and hydrogels in which the proteins of interest are embedded in a water-containing matrix have been developed as well. The quality of the results obtained is dependent on a variety of factors. The surface structure has important effects on background noise and is one of the major factors that determines the signal-to-noise ratio. Background noise is caused by surface irregularities, which lead to light scattering, nonspecific adsorption, and differences in the immobilization efficiency of the printed capture molecules. Certain surfaces (e.g., membrane-coated slides) can show a high degree of autofluorescence and are therefore only of limited use when fluorescence-based readout systems are used. In addition, the binding capacity of the surface determines the maximal signal that can be generated. Another important issue in the immobilization of proteins is the functionality of the capture molecules. Denaturation effects caused by binding to the surface can lead to a loss of signal intensity. These effects do not present a major problem in the immobilization of stable molecules (e.g., antibodies), but they must be taken into account for certain proteins, and it also is advisable to test different types of substrates for such effects. In this protocol, the substrate of choice is aldehyde-activated glass slides. The same substrate was chosen for the Basic Protocol, since these slides have been proven to yield consistent results in the described experimental setup. Depending on the proteins to be immobilized, other surfaces may yield better results.
Protein Microarrays
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Table 23.5.1 List of Commercially Available Microarray Slides That Have Been Tested with the Described Experimental Setup
Surface chemistry
Binding mode/ reactive groups
Product
Manufacturer/distributor
Aldehyde groups
Covalent/amine groups
CSS silylated slides
CEL Associates TeleChem Internationala
Aminosilane
Adsorption, ionostatic interaction
GAPS II
Corning
Poly-L-lysine
Adsorption, ionostatic interaction
Poly-Prep slides
Sigma-Aldrich
Modified polyacrylamide gel
Adsorption, interactions with the 3-D matrix
Hydrogel-coated slides
Packard Biochip Technologiesb
Epoxy
Covalent/amino, thiol, or hydroxyl groups
AdvaSlide AD100
Advalytixc Nunc A/S
Nitrocellulose polymer
Adsorption
FAST slides
Schleicher & Schuell
Amine-reactive cross-linked polymer
Covalent/amine groups
CodeLink activated slides
Amersham Bioscience
Activated heat-stable polymer
Hydrophobic
MaxiSorp
Nunc A/S
a http://www.arrayit.com. b http://www.perkinelmer.com. c http://www.advalytix.de.
Materials Printing buffer: 0.01% (w/v) bromophenol blue in PBS (store up to one month at 4◦ C) 1.5 mg/ml purified protein or capture antibody solutions Stained marker protein (e.g., antibody labeled with Cy5, Jackson Immunoresearch; Alexa 647-labeled BSA, Molecular Probes) Blocking buffer (see recipe) Dedicated microarraying robot: The arrayers listed below have been tested with the described protocol and yield satisfactory results; other arraying robots are likely to produce comparable results. GMS 417 Arrayer (Affymetrix) BioChip Arrayer (Perkin-Elmer) Nanoplotter (GeSiM) MicroGrid (Genomic Solutions) Aldehyde-activated glass slides (CSS; CEL Associates; distributed by Telechem) Microarray scanner (optional): The scanners listed below have been used for the analysis of protein microarrays and yield satisfactory results; other scanners are likely to produce comparable results: GMS 418 Microarray Scanner (Affymetrix) ScanArray Express (Perkin-Elmer) Axon Genepix Microarray Scanner (Molecular Devices) Humidified chamber (e.g., pipet tip box containing a damp Kimwipe) Glass slide staining rack
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Print protein microarrays 1. For each capture protein, add one volume of printing buffer to two volumes of the capture protein solution to yield 1 mg/ml protein in the printing solution. The amount of printing solution to prepare is dependent on a variety of factors. Usually, it is not the number of arrays to be produced that determines the required volume, but rather the arrayer employed for the printing process and/or the plate used as a reservoir for the printing solution. Consult the arrayer manual for recommended volumes. A volume of 25 µl in the commonly used 384-well plate is sufficient to produce several thousand microspots. If possible, the concentrations of the proteins to be immobilized are kept high (>0.5 mg/ml). If diluted protein solutions are used, the maximal signal intensity decreases significantly, and the sensitivity of the assay drops correspondingly. Different buffer systems can be used for the printing process; alternate buffers include HEPES and 2-(N-morpholino)ethanesulfonic acid. Amine-containing buffers (e.g., Tris, glycine) present a problem for the covalent immobilization of proteins on amine-reactive slides. Glycerol (up to 25% v/v) can be included in the printing buffer to allow long-term storage (up to 3 months at −20◦ C) of source plates containing proteins that are sensitive to being frozen. In systems that use a noncovalent immobilization strategy, detergents often cause problems by blocking the array surface. In contrast, some arraying substrates that use covalent binding are able to tolerate detergents (e.g., 0.15% CHAPS, 0.05% Tween 20) in the printing buffer. The presence of bromophenol blue in the printing solution results in a microspot that can be visualized due to its color. Pictures taken with a standard microarray scanner (set to 532 or 635 nm) directly after printing can provide quality control for the generated microarray. During the blocking and assay procedures, the dye is washed away and therefore does not interfere with the results of the experiment.
2. Dilute the labeled marker protein 1:1000 in printing buffer. Position marker spots in the corners of each subarray (see array layout in Fig. 23.5.2).
Protein Microarrays
Figure 23.5.2 Example of an array layout for detecting autoimmune diseases. The array is composed of 20 different autoantigens (thyroid peroxidase, myeloperoxidase, Sm, SmB histone, ssDNA, dsDNA, U1 snRNP A, U1 snRNP C, snRNP 68 kDa, Ro52, Ro60, Scl-70, CENP-B, Jo-1, PM-Scl 100, β2-glycoprotein 1, La, Topo 1, PDH, and PARP; for details on the diagnostic relevance of these well characterized autoimmune proteins, see Rose and Mackay, 1998). The different capture molecules were arrayed in quadruplicate, and the following controls were included: BSA-Cy5, a general control placed at the corners of the array to establish the orientation of the system; human IgG, a positive control for the detection of bound human IgG; and MBP (E. coli maltose binding protein), a negative control.
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3. Load the arrayer with the activated glass slides. Program the arrayer and print the desired number of arrays using a standard printing protocol for the chosen arraying robot. If possible, the arraying process should be performed under controlled environmental conditions; room temperature (20◦ C) and 65% to 70% humidity have proven to be optimal. Nevertheless, good results can be obtained for most proteins (e.g., antibodies) without such controls.
Process arrays after printing 4. Remove glass slides containing the printed arrays from the arraying robot. 5. Optional: Scan the generated arrays in a microarray scanner for quality control. Quality control of the generated arrays is highly recommended during establishment of the method. The researcher should be aware of the integrity of the spotted array and the morphology of the generated spots. (The presence of bromophenol blue dye facilitates detection of irregularities at an early phase of the experiment.) Quality control scans also allow determination of the exact locations of all spots in the array and can therefore be useful for designing a grid in the image analysis program. Solutions to some common quality control problems are listed in Table 23.5.2.
6. Incubate the slides overnight at room temperature in a light-protected humidified chamber (e.g., a pipet tip box containing a damp Kimwipe). This incubation allows covalent linkages to be made between the aldehyde groups on the slide surface and the free amino groups of the spotted proteins via the formation of a Schiff base. Efficient immobilization requires an extended post-printing incubation (a minimum of 8 to 16 hr). When using a particularly fragile, thermally sensitive, or impure protein, the incubation can be carried out for a longer time at a lower temperature (e.g., overnight at 4◦ C).
7. Block the slides by placing them in a slide dish with ∼5 ml blocking buffer per slide and incubating for 30 min at room temperature. Remove excess buffer and let the slides dry in an upright position using a glass slide staining rack. Excess buffer can easily be removed by rapping the edge of the slide on a paper towel resting on the laboratory bench. Blocking has been shown to improve the lower detection limit in most assays by reducing nonspecific adsorption. For the majority of capture molecules, no significant loss of probe activity will be caused by the described blocking procedure. The protein film that forms on the slides after blocking significantly improves the stability of the capture molecules in the array.
8. Store slides in the dark at room temperature for up to 3 months.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Blocking buffer PBS (APPENDIX 2E) containing: 1.5% (w/v) BSA 5% (w/v) lowfat dried milk Store 50-ml aliquots up to 6 months at –20◦ C. Non-Gel-Based Proteome Analysis
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Table 23.5.2 Troubleshooting Guide for Protein Microarrays
Problem
Possible cause
Solution
Missing spots and/or small satellite spots
Poorly cleaned arrayer pins or tips
Clean print pins (contact arrayer) or tips (non-contact arrayer) thoroughly; for details, see manufacturer’s recommendations
High viscosity of sample High amount of detergent
Check for foam formation Dilute or dialyze printing solution (UNIT 4.4) Test different arrayer system if available
Poorly cleaned pins or tips
Thoroughly clean print pins (contact arrayer) or tips (non-contact arrayer)
Detergent in sample that is not compatible with surface
Dilute or dialyze printing solution (UNIT 4.4) Test alternative arraying substrate
Poorly visible spots, no marker spots present
General immobilization problem
Check humidity and temperature during immobilization Test different batch of slides when using chemically labile surfaces (i.e., active ester–activated surfaces) Check printing buffer Test different type of surface
Poorly visible spots, marker spots present
Low capture-molecule concentration
Concentrate printing solution
Interfering substance in printing solution
For covalent immobilization strategies (e.g., aldehyde, epoxy, active ester), check for the presence of amine-containing compounds (Tris, glycine, and azides interfere with the immobilization); remove interfering substances by dialysis (UNIT 4.4), ultrafiltration (UNIT 4.4), or gel filtration chromatography (UNIT 8.3).
Denatured capture proteins
Test alternative arraying substrate Concentrate printing solution (UNIT 4.4) Test alternate buffer compositions with and without stabilizing proteins such as BSA and with and without detergents/stabilizing agents such as trehalose or glycerol
Arrayer problem: tips move away from spot too quickly
Consult arrayer manual Reduce movement speed of tips
Blocking problem
Add a greater volume of blocking solution Add blocking solution more quickly
Irregular spot morphology Diffuse or very large spots
Spots appear smeared or as comets
continued
Protein Microarrays
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Table 23.5.2 Troubleshooting Guide for Protein Microarrays, continued
Problem
Possible cause
Solution
No blocking step performed or blocking was unsuccessful
Perform blocking step or extend blocking period
Drying of the array during the assay procedure
Place slides in a humidified environment to reduce evaporation; do not let slide dry Work with a higher volume of washing buffer
Insufficient washing; washing steps are critical for lowering the background and are frequently performed inadequately
Wash thoroughly for the time and number of washes recommended in the protocol Use fresh changes of wash buffer for each wash step Increase volume and/or number of wash steps Extend washing time
Unsuitable combination of arraying substrate and detection system. 632 nm laser (Cy5 channel) often gives lower background. Use of membranes as arraying substrates can lead to high background when fluorescence is used as a readout.
Use a different combination of substrate and detection system
Cross-reactivity of capture spot with detection antibody; this is often the major problem encountered during experiments involving a higher degree of multiplex
Check cross-reactivity of antibodies in assay blank Try different antibodies
Low amount of functional capture molecule in spot
Test alternate buffer compositions with and without stabilizing proteins such as BSA and with and without detergents/stabilizing agents such as trehalose or glycerol See entries above regarding irregular spot morphology
Low analyte concentration
Extend analyte incubation time and/or increase incubation temperature (e.g., incubate overnight at 4◦ C, increase incubation period at room temperature from 2 hr to 4 hr, incubate 2 hr at 37◦ C instead of 2 hr at room temperature)
Low affinity of binding reaction
Shorten washing procedure Look for alternative capture molecule
Poorly matched sandwich antibody pair
Use a different sandwich pair
Problems with the detection system
Assess the performance of the detection antibody and/or the fluorophore conjugate Include control spots that allow monitoring of the performance of the detection system
Background High or uneven slide background
High signal from spot lacking analyte
Signal Low assay signal
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Dilution buffer PBS (APPENDIX 2E) containing: 1.5% (w/v) BSA 2.5% (w/v) lowfat dried milk 0.1% (v/v) Tween 20 Store 3-ml aliquots up to 6 months at –20◦ C COMMENTARY Background Information
Protein Microarrays
The principles of miniaturized ligandbinding assays were first described more than a decade ago. In their ‘ambient analyte theory,’ Roger Ekins and coworkers (Ekins et al., 1989) explained why a miniaturized assay can be more sensitive than a conventional macroscale ligand-binding assay. They also introduced the concept of the microspot, in which capture molecules are immobilized to the solid phase within a very small area. Although the amount of capture molecules present in such a system is low, a high density of molecules can be obtained in the microspot. Due to the limited number of capture molecules, only a small number of analyte molecules can be captured during an assay procedure, and, thus, the concentration of free analyte in a sample is not changed significantly by the binding reaction. This issue, which can arise even for analytes present at a low concentration and for high-affinity binding reactions, is operative under what have been termed ambient analyte conditions. The fact that no visible changes in concentration occur means that the binding reaction takes place at the highest possible analyte concentration, and, therefore, a high sensitivity can be achieved. Because the capture molecule– analyte complex is concentrated within the small area of a microspot, microassays result in a much higher signal compared with macrospot assays. Therefore, the highest signal intensities per unit area and optimal signalto-noise ratios can be achieved under ambient analyte conditions in a miniaturized system. In such systems, femtomolar concentrations of analytes such as thyroid-stimulating hormone (TSH) and hepatitis B surface antigen (HbsAG) have been detected (Finckh et al., 1998). Miniaturization also allows parallelization, which, in combination with the sensitivities that are achievable, shows that microarray technology has enormous potential for diagnostic applications. The huge interest that microarray-based assays have evoked comes from work using DNA
chips. The possibility of determining thousands of binding events using highly parallelized DNA chips perfectly suited the needs of genome analysis, and consequently, these chips became a key technology in a new area of research, the field of genomics. Today, DNA microarray analysis is a robust and reliable method that allows a global view of the transcriptome of a living cell or organism to be obtained through a single experiment (The Chipping Forecast, 1999; Kapranov et al., 2002; The Chipping Forecast II, 2002; Cawley et al., 2004). Technologies established for DNA chip applications have been adapted to cater to the needs of protein microarray– based research. There are hundreds of companies and technology providers on the market offering products such as microarray surfaces, arrayers, and detection systems for the fast-growing DNA and protein microarray community (http://www.biochipnet.com). Microarrays can be seen as tools that have reached a certain level of maturity, and the use of protein arrays will become more and more important during the coming years. Nevertheless, it has to be stated that technologies and applications are still under development and that many of the published applications have not yet become routinely used methods. Protein microarrays for diagnostic applications Miniaturized and multiplexed immunoassays offer an attractive solution for applications in which several parameters of a single sample have to be analyzed in parallel. This consideration is important in autoimmune diagnostics, in which patient sera are screened for the presence or absence of a number of different types of autoantibodies (Joos et al., 2000; Robinson et al., 2002). Autoantigens used as diagnostic markers for autoimmune conditions such as systemic rheumatic diseases have been immobilized in a microarray, and different types of autoantibodies have been accurately determined from less than 1 µl of patient serum. This reflects the enormous
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potential of protein microarrays employed to study humoral responses against a large number of antigens. Microarrays have also been successfully used to detect the presence of specific IgG and IgM antibodies directed against parasitic and viral antigens such as those from Toxoplasma gondii, cytomegalovirus, herpes simplex virus, papillomavirus, and rubella virus (Mezzasoma et al., 2002; Opalka et al., 2003; Bacarese-Hamilton et al., 2004). The analytical sensitivity of these assays was similar to the sensitivity obtained using standard ELISA technology (Mezzasoma et al., 2002). Subsequent critical steps will involve the validation of array-based assay systems and the clarification of diagnostic needs. Allergy diagnostics might find microarrays to be quite a suitable format, as a sufficiently large number of parameters must be determined in this field. Microarrays represent a very interesting technological alternative in allergy diagnostics (Wiltshire et al., 2000; Bacarese-Hamilton et al., 2002), as the identification of diseasecausing allergenic molecules is a costly and time-consuming procedure. Miniaturized and multiplexed ligand-binding assays can circumvent most of the current limitations associated with this procedure. Planar microarrays (i.e., microarrays that have been created on planar surfaces, such as activated glass slides) are very well suited to multiplex immunoassays. Accurate quantification and control of assay performance and reproducibility can be achieved by including several positive and negative controls and/or internal calibration standards. Thus, multiplexed miniaturized sandwich immunoassays have the potential to become robust and reliable diagnostic tools. Besides planar microarrays, bead arrays present a very interesting alternative, especially when the number of parameters to be determined is low. Bead-based assay systems employ up to 100 different populations of color- or size-coded microspheres as solid supports for the capture molecules. A flow cytometer, which is able to identify the different beads by their color or size, quantifies the amount of captured target on each bead by measuring fluorescence found on the surface of the bead after an immunoassay (Templin et al., 2004). Sandwich immunoassays have also been adapted to the microarray format (Huang, 2001; Schweitzer et al., 2002; Tam et al., 2002). Examples include highly specific and sensitive protein microarrays, and systems capable of quantifying more then 20 different
cytokines from conditioned media and patient sera are possible; however, the parallelization of sandwich assays shows limitations, and only a few examples of systems more complex than this are described. Cross-reactivity of certain detection antibodies with immobilized capture antibodies or nonspecific analytes is likely to make the setup of highly multiplexed sandwich immunoassays difficult. Protein microarrays in proteomic research Aside from more focused diagnostic protein microarray approaches, enormous efforts are being undertaken to transfer the DNA microarray approach to the world of proteins so that a large number of proteins of interest can be quantitated. The potential for array-based proteomic approaches has recently been discussed in several reviews (Cahill, 2000; Emili and Cagney, 2000; Templin et al., 2003). Protein microarray technology can be used to simultaneously determine levels of a large number of target proteins. Specifically, cell extracts that are quantitatively labeled with fluorescent markers are incubated in an array consisting of specific antibodies, after which bound proteins can be detected using conventional DNA microarray instrumentation. Sreekumar et al. (2001) used a comparative approach to study changes in protein expression levels in colon carcinoma cells after an ionizing radiation treatment. Lysates from treated and untreated cells were labeled with different fluorescent dyes, mixed, and allowed to bind to a microarray containing 146 distinct antibodies directed against proteins involved in apoptosis, stress response, and the cell cycle. This comparative analysis revealed the up-regulation of eleven distinct proteins and the down-regulation of a single protein in treated cells. Until then, only five of the eleven up-regulated proteins had been known to be involved in the response to ionizing radiation treatment. Likewise, Knecevic et al. (2001) were able to show cancer-specific alterations in the expression of proteins using a microarray containing 368 antibodies. Cancerous oral epithelium, normal cells, and cells surrounding these tissues were microdissected, lysed, biotinylated, and incubated in the arrays, and bound proteins were detected using biotin-based signal amplification. Eleven distinct proteins with altered expression levels in correlation with tumor progression were identified. These approaches reflect the power of antibody microarrays in determining changes in protein expression in
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a single experiment. Results obtained with antibody microarrays must be verified and confirmed using standard methods, like immunohistochemical staining and immunoblotting, since one must be aware that some antibodies exhibit strong cross-reactivity. In addition, proteins are often assembled in multiprotein complexes. Thus, a strong signal on a microspot can result not only from the presence of a large number of target molecules but also from the capture of a huge labeled complex. Recently, microarrays consisting of several hundred monoclonal antibodies have become commercially available (BD Biosciences Clontech, http://www.bdbiosciences.com; Sigma-Aldrich, http://www.sigma-aldrich. com; Hypromatrix, http://www.hypromatrix. com). Protein microarrays also offer the opportunity to study protein-protein interactions by immobilizing purified recombinant proteins. Interaction assays may use other purified proteins, enriched protein fractions, or complete cell lysates. An interesting example of a global protein interaction study involved a yeast proteome chip generated from recombinant protein probes corresponding to all 5800 known open reading frames in yeast (Zhu et al., 2001). The yeast proteome was then probed with biotinylated calmodulin to test for protein-protein interactions. Many of the known calmodulin-interacting proteins, such as the calmodulin-dependent kinases and the calcineurins, were identified as binding partners of calmodulin. This experiment demonstrated the ability of microarrays to screen for protein-protein interactions at a proteomewide level. At the same time, 33 new potential binding partners of calmodulin were identified, and convincing data were presented indicating that these interactions were real. This study clearly demonstrated the advantages of an approach using a proteome chip, and the chip has since been made commercially available (Yeast Protoarray PPI Microarray; Invitrogen). Using this technology, an entire proteome can be prepared and directly screened in vitro for a wide variety of activities, including protein-drug and protein-lipid interactions, which might not be accessible by other approaches. The generation of arrays containing tens of thousands of proteins for global proteome analysis in humans and other eukaryotes is feasible using similar procedures. In addition, Invitrogen has developed and made commercially available a microarray consisting of 1800 recombinant human proteins.
There is substantial interest in using protein microarrays for the identification of disease and tumor markers. As mentioned above, possible approaches include the use of antibody arrays for comparative analysis of protein expression in normal tissue and diseased and cancerous tissues to identify aberrantly expressed proteins. The molecules identified by such assays might represent new markers (Hanash, 2003). An alternative approach is the use of reversed-phase protein microarrays. In contrast to previously described systems, this type of system involves a complex mixture of proteins immobilized on a microspot. Lysates are prepared from cultured cells or microdissected tissues (e.g., tissues procured by laser capture microdissection) and arranged in a microarray format in miniature dilution series. The microarrays are then screened with specific antibodies for the presence of defined target proteins. Characteristic features of reversed-phase microarrays include high linearity and excellent sensitivity. In addition, the sample proteins need not be labeled (Paweletz et al., 2001). Reversed-phase arrays can use denatured lysates so that the retrieval of antigens does not pose problems. Nondenatured lysates can also be used, even when directly derived from laser capture microdissection–procured tissue cells, to identify the target protein of interest as well as to elucidate protein–protein, protein–DNA, and/or protein-RNA interactions. The samples arrayed in a dilution series provide an internal standard. Direct quantitative assessments can also be made using reversed-phase microarrays, because measurements lie within the linear dynamic range of the antibody-analyte interaction at any given point on the dilution curve (Petricoin et al., 2002). This approach is similar to proteome analysis using surface-enhanced laser desorption and ionization (SELDI) technology, which relies on mass spectrometry as the readout system; detailed information on SELDI can be found in UNIT 22.3. SELDI is a tool that is useful in rapid screening for differences in total protein content and in the detection of small proteins and peptides. However, its sensitivity is much lower in comparison with fluorescently-labeled capture targets on antibody microarrays. These examples show that protein microarrays are powerful tools in the study of complex processes, but it must be reemphasized that protein microarray analysis is a technology that is still under development.
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Critical Parameters and Troubleshooting The printing of microarrays from proteins in solution requires dedicated microarrayers. The arraying robots available have been constructed to print nucleic acids, and in many cases, no information on the printing of proteins is available. Nonetheless, standard printing protocols developed for DNA are often applicable and should be followed for the first set of experiments. Problems can occur during the generation of microarrays, and the most commonly encountered difficulties involve incompletely printed arrays with missing spots. Incomplete printing is often due to protein solutions that contain large amounts of detergents or glycerol. If such a problem occurs, the protein solution should be diluted in suitable printing buffer or, if necessary, dialyzed against a suitable printing buffer. During the planning of the experiment, a substrate for the generation of protein microarrays must be chosen. Table 23.5.1 lists commercially available substrates that can be used for the generation of protein arrays. Several of these surfaces may have to be tested before optimal results are obtained for a specific assay. However, the choice of substrates can be narrowed by considering the following questions. Are the proteins to be arrayed particularly labile? Does the buffer contain substances that are interfering with the immobilization chemistry? Are the surfaces compatible with the chosen readout system? The functionality of the arrayed proteins as capture molecules is of prime interest when choosing a substrate. Denaturation effects that are caused by the binding of the protein can occur and can lead to a decrease in signal intensity. This is particularly important when proteinprotein interaction assays involving many different types of proteins are performed. The generation of capture arrays consisting of immunoglobulins is not critically dependent on the choice of substrate, and all of the substrates listed in Table 23.5.1 have been used with good results for this type of protein array. The composition of the buffer that contains the protein is a second factor that determines the choice of substrate. The presence of detergents, which can block substrate surfaces, limits the use of substrates that rely on noncovalent immobilization; in addition, aminecontaining buffers (e.g., Tris, glycine) interfere with covalent attachment via amino groups. Highly sensitive assays that use fluorescence as the readout system benefit from an optically
flat surface and from low autofluorescence by the arraying-substrate. The signal intensity that can be obtained in a binding assay is dependent on the density of the molecules in the microspot and also on the functionality of the immobilized proteins. Three-dimensional surfaces (e.g., branched polymers, gel-like structures, nitrocellulose membranes) facilitate the achievement of a high density of molecules in the microspot. The functionality of the arrayed proteins as capture molecules is mainly determined by denaturation effects that are caused by binding to the substrate surface. A simple way of reducing such effects involves the use of highly concentrated protein solutions (>1 mg/ml). This helps yield the high capture-molecule density that is required for a sensitive assay, and at the same time, the high concentration seems to stabilize the proteins in the microspot. The problem of denaturation of the capture molecules is seen in direct connection with the stability of the generated array. Without stabilization, a loss of capture activity is seen within days. Stabilization procedures that have been developed for 96-well plates work well for protein microarrays, and the procedure described in the Support Protocol has been adapted from the 96-well plate format. The described treatment allows storage of the generated arrays for a period of two to three months. Stabilization procedures can be combined with blocking of the surface, which is important for improving the performance of the microarray assay. During blocking, unsaturated binding sites on the substrate surface become occupied, and the background signal of the assay can be reduced significantly as a result. An optimized blocking protocol can lead to a gain in sensitivity by one order of magnitude. Table 23.5.2 describes problems that may be encountered when constructing and using protein microarrays, along with explanations of possible causes and recommendations for overcoming or avoiding these problems.
Anticipated Results The use of protein microarrays allows the measurement of several dozen parameters in one reaction. For systems like the described autoimmune assay (Basic Protocol), the degree of multiplex is limited only by the number of antigen preparations available. The sensitivity of such systems is high, and therefore, the amount of serum required to perform an assay can be <1 µl. The quality (i.e., selectivity and specificity) of the assay is comparable to a 96
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microwell–based assay system when using the same antigen preparations; usually, miniaturization does not change the performance of an assay, and problems observed in the macroassay are also found in the miniaturized array format. The signal-to-noise ratio of a properly designed system is well above 100, and therefore, the dilution of the sample is not a critical factor. The described autoimmune assay works with serum dilutions of 1:50 to 1:1000. The proposed 200-fold dilution has proven to be useful for the set of antigens arrayed, and the identification and classification of positives can be achieved within this one dilution. Figure 23.5.2 shows an example of an autoimmune array that has been used routinely to detect anti-nuclear autoantibodies in human serum (Joos et al., 2000). The setup of a multiplex sandwich assay is often more complicated. Although systems that are capable of measuring more than 30 different cytokines in a multiplexed assay have been described (Schweitzer et al., 2002; Wang et al., 2002), this degree of multiplex is still the exception, and novel technological developments in this area are required to make highly parallel systems possible. Nevertheless, it is possible to run parallelized immunoassays with sensitivity comparable to that of standard microwell ELISA systems, and the described setup has been used to detect different cytokines in the low pg/ml range; commercial suppliers of such arrays are Novagen, Perbio, and Schleicher & Schuell. At the moment, the major advantage of parallelized and miniaturized sandwich immunoassays is the small amount of sample required per analyte, and not the high degree of multiplexing that is seen in DNA microarray experiments.
before starting the arraying. With these initial experiments, the generation of arrays can be straightforward. An array layout must be created (see example in Fig. 23.5.2), and the arraying robot must be programmed to generate the arrays. The spotting of a dozen arrays, each composed of 12 subarrays, will take a few hours. The exact time is dependent on the number of spots and arrays to be printed. It is advantageous to start the arraying process in the morning. It is also possible to array overnight if no problems with the reliability of the arraying robot have been observed. Blocking and stabilization of the arrays must be done on the following day to minimize problems associated with the stability of the arrayed proteins. Again, time requirements are dependent on the number of arrays generated, but blocking usually requires less than 3 hr. The blocked arrays are stable when stored under appropriate conditions, and assays can be performed over the course of several weeks using the same batch of microarrays. The actual assay starts with a dehydration step, followed by different incubation and washing steps, and the time required is similar to that for other immunoassay procedures (usually 3 to 5 hr). After drying, the arrays are stable, and scanning can be performed immediately or, if the arrays are stored properly in the dark, up to several days later. The scanning process is fast, requiring only a few minutes, and depending on the scanning software, it allows an initial evaluation of the data. A more detailed evaluation requires dedicated microarray evaluation software and is dependent on the amount of data generated. If such an evaluation is performed, more time may be required for quantification and visualization of the results.
Literature Cited Time Considerations
Protein Microarrays
While the actual time required to run a microarray-based immunoassay as described is only 1 to 4 hr, the experimenter should be aware that the setup of a protein microarray–based analytical system requires thorough planning. The planning starts with obtaining access to a microarraying robot and a microarray scanner. This dedicated equipment is now present in microarray core facilities at many companies and universities. The proteins chosen to fill the array should be tested in a standard immunoassay format to obtain an indication of their quality; when considering the use of arrays for miniaturized sandwich immunoassays, it is advisable that a pair of antibodies for each analyte be tested
Bacarese-Hamilton, T., Ardizzoni, A., Gray, J., and Crisanti, A. 2004. Protein arrays for serodiagnosis of disease. Methods Mol. Biol. 278:271-284. Bacarese-Hamilton, T., Mezzasoma, L., Ingham, C., Ardizzoni, A., Rossi, R., Bistoni, F., and Crisanti, A. 2002. Detection of allergen-specific IgE on microarrays by use of signal amplification techniques. Clin. Chem. 48:1367-1370. Cahill, D.J. 2000. Protein arrays: A high-throughput solution for proteomics research? In Proteomics: A Trends Guide (W. Blackstock, ed.) pp. 47-51. Elsevier Science Publishing, New York. Cawley, S., Bekiranov, S., Ng, H.H., Kapranov, P., Sekinger, E.A., Kampa, D., Piccolboni, A., Sementchenko, V., Cheng, J., Williams, A.J., Wheeler, R., Wong, B., Drenkow, J., Yamanaka, M., Patel, S., Brubaker, S., Tammana, H., Helt, G., Struhl, K., and Gingeras, T.R. 2004. Unbiased mapping of transcription factor
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binding sites along human chromosomes 21 and 22 points to widespread regulation of noncoding RNAs. Cell 116:499-509. Ekins, R.P. 1989. Multi-analyte immunoassay. J. Pharm. Biomed. Anal. 7:155-168. Emili, A.Q. and Cagney, G. 2000. Large-scale functional analysis using peptide or protein arrays. Nat. Biotechnol. 18:393-397. Fall, B.I., Eberlein-Konig, B., Behrendt, H., Niessner, R., Ring, J., and Weller, M.G. 2003. Microarrays for the screening of allergenspecific IgE in human serum. Anal. Chem. 75:556-562. Finckh, P., Berger, H., Karl, J., Eichenlaub, U., Weindel, K., Hornauer, H., Lenz, H., Sluka, P., Weinreich, G.E., Chu, F., and Ekins, R. 1998. Microspot—an ultrasensitive microarray-based ligand assay system: A practical application of ambient analyte assay theory. Proc. U.K. Natl. Ext. Qual. Assess. Serv. Meeting 3:155-165. Hanash, S. 2003. Disease proteomics. Nature 422:226-232. Harwanegg, C., Laffer, S., Hiller, R., Mueller, M.W., Kraft, D., Spitzauer, S., and Valenta, R. 2003. Microarrayed recombinant allergens for diagnosis of allergy. Clin. Exp. Allergy 33:7-13. Huang, R.P. 2001. Simultaneous detection of multiple proteins with an array-based enzyme-linked immunosorbent assay (ELISA) and enhanced chemiluminescence (ECL). Clin. Chem. Lab. Med. 39:209-214. Joos, T.O., Schrenk, M., Hopfl, P., Kroger, K., Chowdhury, U., Stoll, D., Schorner, D., Durr, M., Herick, K., Rupp, S., Sohn, K., and Hammerle, H. 2000. A microarray enzymelinked immunosorbent assay for autoimmune diagnostics. Electrophoresis 21:2641-2650. Kapranov, P., Cawley, S.E., Drenkow, J., Bekiranov, S., Strausberg, R.L., Fodor, S.P., and Gingeras, T.R. 2002. Large-scale transcriptional activity in chromosomes 21 and 22. Science 296:916-919. Knezevic, V., Leethanakul, C., Bichsel, V.E., Worth, J.M., Prabhu, V.V., Gutkind, J.S., Liotta, L.A., Munson, P.J., Petricoin, E.F. 3rd, and Krizman, D.B. 2001. Proteomic profiling of the cancer microenvironment by antibody arrays. Proteomics 1:1271-1278. Lueking, A., Horn, M., Eickhoff, H., Bussow, K., Lehrach, H., and Walter, G. 1999. Protein microarrays for gene expression and antibody screening. Anal. Biochem. 270:103-111. Mezzasoma, L., Bacarese-Hamilton, T., Di Cristina, M., Rossi, R., Bistoni, F., and Crisanti, A. 2002. Antigen microarrays for serodiagnosis of infectious diseases. Clin. Chem. 48:121-130. Opalka, D., Lachman, C.E., MacMullen, S.A., Jansen, K.U., Smith, J.F., Chirmule, N., and Esser, M.T. 2003. Simultaneous quantitation of antibodies to neutralizing epitopes on virus-like particles for human papillomavirus types 6, 11, 16, and 18 by a multiplexed luminex assay. Clin. Diagn. Lab. Immunol. 10:108-115.
Paweletz, C.P., Charboneau, L., Bichsel, V.E., Simone, N.L., Chen, T., Gillespie, J.W., Emmert-Buck, M.R., Roth, M.J., Petricoin, I.E., and Liotta, L.A. 2001. Reverse phase protein microarrays which capture disease progression show activation of pro-survival pathways at the cancer invasion front. Oncogene 20:19811989. Petricoin, E.F., Zoon, K.C., Kohn, E.C., Barrett, J.C., and Liotta, L.A. 2002. Clinical proteomics: Translating benchside promise into bedside reality. Nat. Rev. Drug. Discov. 1:683-695. Rigaut, G., Shevchenko, A., Ruiz, B., Wilm, M., Mann, M., and Seraphin, B. 1999. A generic protein purification method for protein complex characterization and proteome exploration. Nat. Biotechnol. 17:1030-1032. Robinson, W.H., DiGennaro, C., Hueber, W., Haab, B.B., Kamachi, M., Dean, E.J., Fournel, S., Fong, D., Genovese, M.C., de Vegvar, H.E., Skriner, K., Hirschberg, D.L., Morris, R.I., Muller, S., Pruijn, G.J., van Venrooij, W.J., Smolen, J.S., Brown, P.O., Steinman, L., and Utz, P.J. 2002. Autoantigen microarrays for multiplex characterization of autoantibody responses. Nat. Med. 8:295-301. Rose, N.R. and Mackay, I.R. 1998. The Autoimmune Diseases. Academic Press, San Diego. Schweitzer, B., Roberts, S., Grimwade, B., Shao, W., Wang, M., Fu, Q., Shu, Q., Laroche, I., Zhou, Z., Tchernev, V.T., Christiansen, J., Velleca, M., and Kingsmore, S.F. 2002. Multiplexed protein profiling on microarrays by rolling-circle amplification. Nat. Biotechnol. 20:359-365. Sreekumar, A., Nyati, M.K., Varambally, S., Barrette, T.R., Ghosh, D., Lawrence, T.S., and Chinnaiyan, A.M. 2001. Profiling of cancer cells using protein microarrays: Discovery of novel radiation-regulated proteins. Cancer Res. 61:7585-7593. Tam, S.W., Wiese, R., Lee, S., Gilmore, J., and Kumble, K.D. 2002. Simultaneous analysis of eight human Th1/Th2 cytokines using microarrays. J. Immunol. Methods 261:157-165. Templin, M.F., Stoll, D., Bachmann, J., and Joos, T.O. 2004. Protein microarrays and multiplexed sandwich immunoassays: What beats the beads? Comb. Chem. High Throughput Screen. 7:223229. Templin, M.F., Stoll, D., Schwenk, J.M., Potz, O., Kramer, S., and Joos, T.O. 2003. Protein microarrays: Promising tools for proteomic research. Proteomics 3:2155-2166. The Chipping Forecast. 1999. Nat. Genet. 21 Suppl:1-60. The Chipping Forecast II. 2002. Nat. Genet. 32 Suppl:461-552. Wang, C.C., Huang, R.P., Sommer, M., Lisoukov, H., Huang, R., Lin, Y., Miller, T., and Burke, J. 2002. Array-based multiplexed screening and quantitation of human cytokines and chemokines. J. Proteome Res. 1:337-343.
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Wiltshire, S., O’Malley, S., Lambert, J., Kukanskis, K., Edgar, D., Kingsmore, S.F., and Schweitzer, B. 2000. Detection of multiple allergen-specific IgEs on microarrays by immunoassay with rolling circle amplification. Clin. Chem. 46:1990-1993. Zhu, H., Bilgin, M., Bangham, R., Hall, D., Casamayor, A., Bertone, P., Lan, N., Jansen, R., Bidlingmaier, S., Houfek, T., Mitchell, T., Miller, P., Dean, R.A., Gerstein, M., and Snyder, M. 2001. Global analysis of protein activities using proteome chips. Science 293:2101-2105.
Contributed by Stefan Kramer, Thomas O. Joos, and Markus F. Templin NMI—Natural and Medical Science Institute at the University of T¨ubingen T¨ubingen, Germany
Protein Microarrays
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CHAPTER 24 Targeted Proteomics INTRODUCTION roteomics is a relatively new discipline that has grown rapidly, both in terms of expectations and capabilities, during the past decade. Chapters 22 (Gel-Based Proteome Analysis) and 23 (Non-Gel-Based Proteome Analysis) focus primarily on proteinprofiling techniques and associated analysis strategies. The techniques described in those chapters can be applied to different types of samples having a wide range of complexity. Comprehensive (global) proteome analysis involves attempts to identify and, in some cases, quantitatively compare as many proteins as possible in the sample of interest. However, comprehensive analysis of complex proteomes, such as those from cells or tissues of higher eukaryotic organisms, is complicated by the fact that the complexity of the proteome being analyzed usually greatly exceeds the analytical capacity of common current protein-profiling methods such as 2-D gels or LC/LC-MS/MS analysis.
P
A useful strategy that frequently effectively addresses the high complexity of higher eukaryotic proteomes is to focus on a specific subproteome (targeted proteomics). The key factor in pursuing targeted proteomics is to define a subset of a complete proteome that can be reproducibly isolated. The closer match between the protein complexity of most subproteomes and available analytical methods makes targeted proteomics a very attractive approach to proteomics-based exploration of biological processes and diseases. In terms of cellular complexity, subproteomes can range from simple macromolecular complexes to cellular machines to organelles. Other strategies for targeting subproteomes are to isolate specific classes of proteins, such as phosphoproteins or tyrosine-phosphorylated proteins, glycoproteins, or oxidatively modified proteins, to name only a few. Frequently, several alternative methods are available for isolating specific subproteomes. As noted above, one of the most critical considerations is the reproducibility of the subproteome isolation method. Another critical consideration is the specificity of the isolation method used; since no method is completely specific, an important issue to consider is how one will distinguish contaminants from proteins that belong to the targeted subproteome. Another advantage of targeted proteomics, in addition to matching analytical methods, is that more specific biological questions can be addressed, compared with global proteomic analyses. This usually facilitates interpretation of the biological significance of study results. The first unit in this chapter (UNIT 24.1) addresses analysis of low-abundance proteins in plasma as an example of analysis of biological fluids. The most common goal of biological fluid proteome analysis is discovery of disease-related biomarkers. In order to detect low-abundance proteins that are potential biomarkers, effective analysis of plasma or serum requires depletion of high-abundance proteins. Alternative methods for detecting potential biomarkers are to isolate the low-molecular-weight subproteome or to focus on specific subsets of proteins, such as glycoproteins, proteases, or oxidatively modified proteins. The second unit in this chapter (UNIT 24.2) provides an overview of organelle proteomics, including several protocols for isolating nuclei from different sources. It also crossreferences units in other chapters that describe conventional and electrophoretic organelle isolation methods that can be used for proteomics studies. Contributed by David W. Speicher Current Protocols in Protein Science (2006) 24.0.1-24.0.2 C 2006 by John Wiley & Sons, Inc. Copyright
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Future units in this chapter will explore isolation of classes of protein such as phosphoproteins, glycoproteins, proteases, and cell-surface proteins. Coverage of subproteomes related to cellular complexity will include macromolecular complexes, cellular machines, and isolation of entire organelles. David W. Speicher
Introduction
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Human Serum and Plasma Proteomics
UNIT 24.1
The human circulatory system provides a unique source of proteins that can potentially be used to evaluate both normal and disease conditions. Serum and plasma are easily accessible fluids and, in theory, contain protein signatures from every cell within the body. However, systematic analysis of serum and plasma proteomes is extremely challenging due to the huge dynamic range of protein concentrations, the large number of proteins present, and extensive physiological variation. Albumin, the most common protein in serum, comprises more than 50% of serum protein content and is present at mg/ml concentrations, while most potential disease biomarkers are thought to be present at ng/ml levels or less. Approximately 99% of the protein content of serum is comprised of 22 major proteins (Tirumalai et al., 2003), which, with their various isoforms, dominate and obscure less abundant proteins. Current serum and plasma proteome analysis strategies typically seek to overcome the wide dynamic range problem by employing protein-depletion methods, as well as by more completely fractionating the proteome using more than two dimensions of protein/peptide separations. This unit focuses on several different methods of human serum/plasma fractionation and analysis. One of the most powerful current protein-profiling strategies utilizes major protein depletion coupled to other protein-separation techniques such as MicroSol IEF fractionation and SDS-PAGE analysis followed by LC-MS/MS (see Basic Protocol 1). More common basic approaches involve the depletion of abundant proteins followed by analysis either via 2-D gels (see Alternate Protocol 1) or using offline Multidimensional Protein Identification Technology or MudPIT (see Alternate Protocol 2). Additional protein-profiling strategies are also feasible, as discussed in Strategic Planning.
STRATEGIC PLANNING Sample Types and Sample Collection The most basic choice involved in proteomic analysis of human blood proteins is whether serum or plasma will be analyzed. When studying human diseases, an initial consideration should be given to potential archives of samples that can be utilized. Sample archives have the advantage of providing a large number of samples for study, coupled with the disadvantage of potential variability. The characteristics of a serum or plasma archive must be fully examined before proceeding with its use. Details such as duration and temperature of storage and volume and type of samples are more difficult to control when dealing with a third-party archive, and records appropriate to the proposed experimental procedures may not have been adequately kept. Blood-sample archives are traditionally dominated by serum rather than plasma samples. If an adequately annotated and maintained archive is not available, it may be a worthwhile endeavor to collect new samples. Rai et al. (2005), based upon a comparison showing a large number of peptides in serum that are not present in plasma, recommend the use of platelet-depleted plasma. This ex vivo generation of peptides may be the result of platelets releasing peptides, or of the effects of blood clotting (Misek et al., 2005; Tammen et al., 2005). Peptides generated in this manner may interfere with some studies. The collection of samples should be a consistent, well defined procedure, in order to maintain the integrity of the experiment and to avoid introduction of bias related to sample collection and handling. Numerous factors must be considered in the collection and handling of specimens. Collection should be performed in a consistent manner, using Targeted Proteomics Contributed by Ian Musselman and David W. Speicher Current Protocols in Protein Science (2005) 24.1.1-24.1.18 C 2005 by John Wiley & Sons, Inc. Copyright
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the same type of sample-collection device each time. For plasma analysis, a coagulation inhibitor must be used; typical options include citrate, EDTA, and heparin. Citrate and EDTA inhibit coagulation by chelating divalent cations, which subsequently inhibits enzymes involved in blood clotting (Tammen et al., 2005). Heparin, which functions through the activation of antithrombin III, displays slightly different elution characteristics in LC-MS analysis, potentially due to chromatographic interference (Tammen et al., 2005). The anticoagulating factor, as well as the manner in which it is used, should be kept consistent throughout the collection of all samples to assure a low level of variance.
Sample Handling Recent studies show that, with serum, even long-term exposure to room temperature only causes moderate levels of proteolytic degradation (Rai et al., 2005). This is in great contrast to cell lysates, for which short periods of exposure usually result in drastic proteolytic degradation. However, despite this relative level of stability, Rai and colleagues found that even at −20◦ C substantial proteolysis and modification of some proteins still occurs. For this reason, samples should be stored only for minimal and well defined time ranges at either room temperature or 4◦ C prior to quick freezing aliquots for storage at −80◦ C or lower. The number of freeze/thaw cycles per sample should also be kept to a minimum, preferably a maximum of one or at most two. Storage in liquid nitrogen is preferable, but probably not generally practical, and storage at −80◦ C appears to work nearly as well. Rai et al. (2005) found that the addition of protease inhibitors (PI) afforded a nearly immediate and detectable positive benefit by stabilizing the sample. Thus, the presence of PI at the earliest stage possible, such as during phlebotomy, could result in a substantial reduction of protein degradation. However, use of PIs that are either proteins themselves or reagents that modify amino acid side chains, such as 4-(2-aminoethyl)benzenesulfonyl fluoride (AEBSF), should be avoided, as they may react with other proteins in addition to the intended target proteases (Rai et al., 2005).
Human Serum and Plasma Proteomics
Dynamic Range and Complexity of Plasma and Serum The dynamic range of protein concentrations in serum and plasma presents a unique challenge in the field of proteomics. Serum and plasma, with typical protein concentrations of around 60 to 80 mg/ml, are dominated by a small number of high-abundance proteins. The range of protein concentrations in serum or plasma span at least 1012 orders of magnitude in concentration (Tirumalai et al., 2003). The six most abundant proteins (albumin, transferrin, haptoglobin, α1-antitrypsin, IgG, and IgA) account for about 85% of the proteome. These proteins also have a substantial number of isoforms, which can be seen as multiple gel spots in Figure 24.1.1 (Echan et al., 2005). Albumin, with a typical concentration of 35 to 50 mg/ml, accounts for >50% of the total serum protein content (Tirumalai et al., 2003). Some protein concentration ranges and their relative abundance levels are displayed in Figure 24.1.2. Serum and plasma are thought to contain a very complex set of proteins shed from most cell types in the body; however, most shed proteins will be present at far lower concentrations than the dominant proteins. This has led to the creation of high-performance protein depletion columns, one example being the Agilent Multiple Affinity Removal System (MARS) column. Depletion of the top six proteins substantially increases the volume of serum that can be analyzed by most methods (Fig. 24.1.1). However, when 2-D gels are used to analyze depleted serum or plasma, most of the “low-abundance” spots revealed by these higher serum loads are minor forms of high- or moderate-abundance proteins (Echan et al., 2005).
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Figure 24.1.1 2-D gels of human serum before and after depletion using the Multiple Affinity Removal System. Samples were separated on 18-cm pH 3-10 L IPG strips and silver stained. (A) Gel from 100 µg of nondepleted human serum. The high-abundance proteins mask the lower-abundance proteins. (B) Proteins after depleting the top six proteins from 1 mg of depleted plasma. The larger sample load shows additional spots that were not visible due to the presence of highabundance proteins in A. However, many of the new forms visible in B are minor forms of high- and moderate-abundance proteins. (C) The bound fraction highlights the locations of the major proteins that were depleted. The large numbers of major isoforms for each of these proteins are indicated by the ellipses. Most unlabeled spots are proteolytic fragments of the major proteins (Echan et al., 2005).
Although protein-depletion methods allow improved detection of low-abundance proteins, depletion decreases throughput of human serum/plasma analysis, and antibody affinity resins are quite expensive. Currently, there are two predominant types of major protein-depletion affinity resins including dyes and protein A or G for albumin and immunoglobulins, respectively, or immunoaffinity columns. Immunoaffinity resins such as the MARS HPLC columns are expensive, and also require complex instrumentation, but have been found to be most successful in regard to depletion (Echan et al., 2005). Dye-based affinity resins are more economical but are less effective and less specific than
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Figure 24.1.2 Protein concentration ranges in human serum and plasma (from Specialty Laboratories; see Internet Resources).
immunoaffinity resins. Another consideration is the potential for low-abundance proteins to be bound to higher-abundance proteins or to bind nonspecifically to the affinity resin, resulting in their depletion. Echan et al. (2005) found that when the HPLC MARS column is used, all major protein spots depleted were isoforms of the targeted high-abundance proteins. Nonetheless, there is always the possibility that a low-abundance protein of interest might be recovered in the bound fraction. Hence, it is generally a good idea to analyze both the bound and unbound fractions, regardless of the depletion method used.
Human Serum and Plasma Proteomics
The exact level of complexity of human serum/plasma is not well defined, but serum and plasma probably represent the most complex proteomes in the body, potentially containing nearly all of the proteins present in the body. This extensive complexity is reflected by the fact that recent studies have identified between 1000 and 3000 proteins in serum and plasma (Anderson et al., 2004; Chan et al., 2004; Shen et al., 2004; Omenn et al., 2005), yet even in these largest available protein datasets, a large portion of the identified proteins were in fact unique to each study (Omenn et al., 2005). While there was extensive overlap of higher-abundance proteins between data sets, as expected, the minimal overlap of identifications of lower-abundance proteins in different analyses indicates that small portions of a very large pool of low-abundance proteins are being somewhat randomly sampled.
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Table 24.1.1 Comparisons of Common Human Serum and Plasma Proteome Analysis Methodsa
Method
Advantages
Disadvantages
No. of proteins typically found in serum
2-D electrophoresis
High resolution
Difficult to automate
400-700 spots
Well established method
Some variability between gels
Resolves many post-translationally modified proteins Resolves spliceoforms
Poor separation/recovery of large, very small, very acidic, very basic, and hydrophobic proteins
Resolves highly homologous proteins
Technically demanding
Relative and absolute quantitation
Limited protein loads Limited dynamic range: about 102 to 104 depending on type of stain
2-D DIGE
Same advantages as 2-D electrophoresis Same as 2-D electrophoresis except variability between gels Eliminates gel-to-gel variation Broader linear dynamic range compared to silver or colloidal Coomassie staining
500-700 spots
May be difficult to excise correct spots due to shift caused by labeling
Relative quantitation Shotgun proteomics (MudPIT)
Capable of detecting a few low-abundance proteins
Low to moderate throughput
Improved dynamic range compared to 2-D gels
Technically demanding
Increased sensitivity
Expensive equipment required
Can be automated
Does not detect most isoforms or PTMs
Relative quantitation and limited multiplexing with stable isotope labels such as iTRAQ
May not distinguish between closely related sequences
Multidimensional Capable of detecting the largest number Low throughput methods (>2-D) of proteins including many Technically demanding low-abundance proteins Automation and increased throughput are feasible
Expensive equipment required
Wide dynamic range, up to 109
Limited detection of different isoforms and related sequences
Increased sensitivity Relative quantitation and limited multiplexing with stable isotope labels such as iTRAQ Pattern High throughput recognition Relatively straightforward method methods (SELDI) Readily automated
Questionable reproducibility
100-350
>1000
<100
Potential interference from variable proteolysis of major serum proteins Questionable sensitivity continued
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Table 24.1.1 Comparisons of Common Human Serum and Plasma Proteome Analysis Methodsa , Continued
Method
Advantages
Disadvantages
No. of proteins typically found in serum
Antibody arrays
High throughput
Emerging technology
N/A
Extensive multiplexing with minimal sample consumption
Very time-consuming reagent preparation
High sensitivity
Limited by available receptor molecules
Readily automated
Not discovery based
Absolute or relative quantitation depending upon detection method
Potentially high background signals
a Some data have been extracted from UNIT 22.1.
Sample Number Versus Analysis Throughput The minimum number of samples that need to be analyzed in a given study is a critical consideration in serum/plasma proteome analysis experimental design. Currently the only high-throughput proteomic approaches are antibody arrays (UNIT 23.5) and SELDIMS (UNIT 22.3). Antibody arrays are limited by the number of appropriate antibodies available, and SELDI has questionable sensitivity and reproducibility (Diamandis, 2004) and preferentially interrogates the low-molecular-weight proteome. In serum and plasma, the low-molecular-weight proteome is dominated by proteolytic fragments of highand medium-abundance proteins. Other proteomic methods have substantially lower throughput. Powerful multidimensional approaches (see Basic Protocol 1), in particular, may yield the most extensive protein profiles. However, it may take several weeks to prepare and analyze each sample by such methods, allowing only a small number of samples to be analyzed. Another consideration is that samples run at different times must be stringently monitored or compared to a uniform reference to control for parameters that may bias comparisons between samples and controls or temporally separated analyses. Protein-Profiling Techniques Table 24.1.1 lists some major protein-profiling techniques that are applicable to serum and plasma proteome analyses. The technique chosen should take into account the experimental goals. For example, 2-D gel-based approaches are particularly well suited for analyzing protein isoforms and post-translational modifications, but lack the ability to detect low-abundance proteins. BASIC PROTOCOL 1
Human Serum and Plasma Proteomics
MULTIDIMENSIONAL PROTEIN AND PEPTIDE SEPARATION This protocol presents a multidimensional protein and peptide fractionation strategy capable of identifying a large number of serum or plasma proteins over at least nine orders of magnitude of concentration, including a substantial number of proteins at the ng/ml level. It consists of three sequential protein-fractionation steps: major protein depletion via reversed-phase high-precison liquid chromatography (RP-HPLC), MicroSol isoelectric focusing (IEF) fractionation, and SDS-PAGE, followed by liquid chromatography– tandem mass spectrometry (LC-MS/MS). The scheme was developed primarily by integrating and optimizing orthogonal separation techniques (Tang et al., 2005). This protocol involves the use of HPLC (see UNIT 8.7). In addition, this protocol may be made quantitative by using either ICAT (UNIT 23.2) or iTRAQ stable isotope labeling methods.
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Materials Human serum or plasma sample 1× Agilent MARS Buffer A 1× Agilent MARS Buffer B Human serum or plasma sample Isopropanol 1 M NaOH 10 mM sodium phosphate buffer, pH 7.0 (APPENDIX 2E) 1 M Tris·Cl, pH 8.3 (APPENDIX 2E) Urea Dithiothreitol (DTT) Dimethylacrylamide Thiourea 3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate (CHAPS) IPG buffer (carrier amphoytes, pH 3 to 10 L; Amersham Biosciences) 200 mM ammonium bicarbonate in 50% (v/v) acetonitrile 0.02 µg/µl modified trypsin (Promega) in 40 mM ammonium bicarbonate/10% acetonitrile, pH 8.1 Argon or nitrogen source 40 mM ammonium bicarbonate in 10% (v/v) acetonitrile, pH 8.1 Concentrated acetic acid Solvent A: 0.58% (v/v) acetic acid in Milli-Q or HPLC-grade H2 O Solvent B: 0.58% acetic acid in acetonitrile HPLC system (UNIT 8.7) 1× Agilent MARS column, 4.6 × 50–mm or 4.6 × 100–mm 0.22-µm spin filters Microcentrifuge 5-kDa MWCO spin separators (Amicon Ultra 4; Millipore) Gel-cutting device (e.g., MEF-1.5 Gel Cutter; The Gel Company, http://www.gelcompany.com/) Digestion tubes: microcentrifuge tubes precleaned by rinsing several times in 0.1% trifluoroacetic acid/50% acetonitrile ◦ 37 C shaking water bath 37◦ C thermostatically controlled oven Reversed-phase 75 µm i.d. × 15 cm length PicoFrit column (New Objective, Inc.), packed with MAGIC C18 resin, 5 µm particle size (Michrom BioResources) Linear ion-trap mass spectrometer with microscale autosampler and injection loop (also see Chapter 16) MS/MS data analysis software (e.g., Sequest; http://fields.scripps.edu/sequest/). Additional reagents and equipment for HPLC (UNIT 8.7), SDS-PAGE (UNIT 10.1), protein assay (UNIT 3.4), Zoom IEF fractionation (UNIT 22.6), running Zoom IEF fractions on 1-D minigels (UNIT 22.6), staining with colloidal Coomassie blue (UNIT 10.5), and RP-HPLC (UNIT 8.7), and mass spectrometry (Chapter 16) Connect the MARS column 1. Before connecting the column, purge the lines of the HPLC system (UNIT 8.7) with 1× Agilent MARS Buffer A and MARS Buffer B at a 1.0 ml/min flow rate for 10 min. MARS (Multiple Affinity Removal System) is a registered trademark of Agilent.
2. Attach the 1× Agilent MARS column as described in UNIT 8.7. Equilibrate the column with Buffer A at a flow rate of 1 ml/min for 4 min at room temperature. Program the instrument based on Table 24.1.2. Maximum pressure should be 120 bars. 1× Agilent MARS Buffers A and B should be the only mobile phases. Current Protocols in Protein Science
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Table 24.1.2 HPLC Programs for Several MARS Column Configurations
4.6 × 50–mm column Stage
Time (min)
%B
4.6 × 100–mm column
Flow rate
Time (min)
%B
4.6 × 50–mm + 100 mm in series
Flow rate
Time (min)
%B
Flow rate
1
0.00
0.00
0.25
0.00
0.00
0.50
0.00
0.00
0.25
2
9.00
0.00
0.25
10.00
0.00
0.50
27.00
0.00
0.25
3
9.01
100.00
1.00
10.01
100.00
1.00
27.01
100.00
1.00
4
12.50
100.00
1.00
17.00
100.00
1.00
37.50
100.00
1.00
5
12.60
0.00
1.00
17.01
0.00
1.00
37.80
0.00
1.00
20.00
0.00
1.00
28.00
0.00
1.00
60.00
0.00
1.00
Deplete serum or plasma sample 3. Dilute the human serum or plasma sample 5× with MARS Buffer A (1 part serum to 4 parts buffer A). The amount of sample used should be based on the capacity of the column(s) available and the sample size desired. The 4.6 × 50–mm column has a capacity of about 15 to 20 µl of human serum, while the 4.6 × 100–mm column has a capacity of 30 to 40 µl of human serum. If the volume of serum or plasma to be depleted exceeds column capacity, prepare one large sample and perform multiple depletions.
4. Filter diluted sample using a 0.22-µm spin filter by microcentrifuging for 1 min at 16,000 × g. This will help to remove particles that would clog or contaminate the column.
5. Prior to injecting the first sample of the day, and at the end of the day, run a blank (no injection), then reequilibrate the column using buffer A. 6. Inject the diluted, filtered serum. The injection volume should be based on the column’s capacity. A single 4.6 × 50–mm column can be used with 75 to 100 µl of diluted serum or plasma, while a single 4.6 × 100–mm column will deplete 150 to 200 µl of diluted sample. Running multiple columns in tandem provides a combinatorial increase in sample based on volume (i.e., a 4.6 × 50–mm and a 4.6 × 100–mm column in series can accommodate 225 to 300 µl of diluted serum or plasma in a single run).
7. Collect 0.5-ml flowthrough fractions as indicated in Figure 24.1.3. The high-abundance proteins targeted for depletion will bind to the column. Fractions should be analyzed as soon as possible; however they can be stored at 4◦ C overnight or at −20◦ C for at least a month if needed.
8. Elute bound proteins from the column using MARS Buffer B at a flow rate of 1 ml/min, for a time based on the capacity of the column (Table 24.1.2). Collect 0.5-ml fractions. Collection of eluted fractions is strongly advised. 1-D SDS-PAGE (UNIT 10.1) analysis of these fractions is useful to evaluate column efficiency and capacity as well as nonspecific binding of nontargeted proteins to the bound targeted proteins or column resin.
9. Regenerate the column by running MARS Buffer A according to Stage 5 in Table 24.1.2. Human Serum and Plasma Proteomics
10. Repeat steps 1 to 9 as needed to produce sufficient depleted sample. 11. Disconnect and store the column at 4◦ C in MARS Buffer A.
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Figure 24.1.3 MARS HPLC depletion of six abundant proteins. (A) Chromatogram from a typical MARS HPLC depletion of human serum. (B) Typical 1-D SDS-PAGE gel of initial sample and resulting fractions.
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12. Flush HPLC with water for 30 min at 1 ml/min, then with isopropanol for 30 min at 1 ml/min. The washing steps flush the high-salt MARS buffers from the HPLC system with water.
13. Wet two or more 5-kDa MWCO spin separators by centrifuging 1 ml of water through them. Pool all eluted fractions and concentrate to 1 ml using a spin separator according to the manufacturer’s instructions. Adjust to pH 7.0 using 1 M NaOH. 14. Pool all flowthrough fractions and concentrate to 200 µl using a spin separator. 15. Desalt flowthrough concentrate by adding 1.8 ml of 10 mM sodium phosphate buffer, pH 7.0, and concentrating to 200 µl again using a spin separator. Repeat at least four times to reduce ionic strength and replace the MARS Buffer A. The concentrated flowthrough fraction should represent ∼15% of the initial protein amount applied to the MARS column. The bound fraction should represent the remaining 85%. The protein content of these fractions should be determined using a standard protein assay (UNIT 3.4) to determine sample recovery
Alkylate and perform microscale solution IEF fractionation 16. Add sufficient 1 M Tris·Cl, pH 8.3, and dry urea to the concentrated flow-through fraction to yield final concentrations of 100 mM Tris·Cl and 8 M urea at a pH of 8.3 in a 1.0-ml final volume. 17. Reduce proteins by adding DTT to a final concentration of 20 mM. Incubate for 30 min with shaking at room temperature. 18. Alkylate proteins by adding dimethylacrylamide to a final concentration of 50 mM. Incubate for 30 min with shaking at room temperature. 19. Terminate reaction by adding an additional 60 mM DTT and incubating for 15 min at 37◦ C. 20. Precipitate salts and reagents by adding 9 vol of −20◦ C acetone and incubating 1 hr at room temperature. Microcentrifuge 20 min at maximum speed, room temperature, then remove (and discard) supernatant and air dry pellet. 21. Add the following reagents to the pellet achieve the indicated final concentrations in a final volume of 1.95 ml:
Urea to 8 M Thiourea to 2 M CHAPS to 4% (w/v) DTT to 1% (w/v) Carrier ampholytes (IPG buffer), pH 3-10 L to 0.2% (v/v). It is best to create a master mix of all of the above components except the carrier ampholytes, and to add the ampholytes immediately before proceeding to fractionation. The sample is thus reconstitituted in 1× MicroSol IEF sample buffer.
22. Assemble the Zoom IEF fractionator (UNIT 22.6). The number and pH of membranes used in the Zoom IEF fractionator should be determined based on experimental needs. A seven-chamber approach will increase the level of fractionation of the sample; however, the membranes will have to be custom made (UNIT 22.6). This substantially increases the total protocol time. For this reason, using a five-chamber approach, such as one with commercial membranes having pH values of 3.0, 4.6, 5.4, 6.2, 7.0, and 12.0, is more expedient. Human Serum and Plasma Proteomics
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23. Load 650 µl of sample into the three separation chambers bounded by the pH 4.6 and 7.0 membranes (see UNIT 22.6) of the Zoom IEF fractionator. Run the Zoom IEF fractionator as in Basic Protocol 1 of UNIT 22.6. Current Protocols in Protein Science
Perform 1-D SDS-PAGE and trypsin digestion 24. Further fractionate samples by running 1-D minigels (UNIT 22.6); however, in this case run each fraction on a separate gel at maximum load. Stain using colloidal Coomassie blue (UNIT 10.5). Electrophoresing the fractions on separate gels helps to prevent cross-contamination between samples. Loading the maximum amount helps to increase the total sample digested and analyzed using LC-MS/MS. Optionally fractions with low concentrations, such as the most acidic and most basic fractions, can be concentrated by acetone precipitation.
25. Electrophorese the samples until desired separation is achieved: i.e., 4 or 6 cm. 26. Slice a lane of each gel into uniform 1-mm slices using the gel-cutting device. Place slices into precleaned digestion tubes. Generally two adjacent slices should be combined in each digestion tube. This may be adjusted, depending on the resolution desired.
27. Destain gel slices twice, each time by shaking for 45 min with 200 µl of 200 mM ammonium bicarbonate/50% acetonitrile at 37◦ C. Discard the destaining solution at the end of each treatment. 28. Dry gel pieces for about 30 min using a Speedvac evaporator. 29. Rehydrate gel bands by adding 20 µl of 0.02 µg/µl trypsin in 40 mM ammonium bicarbonate/10% acetonitrile, pH 8.1. Blanket with argon or nitrogen and incubate 16 to 18 hr at 37◦ C in a thermostatically controlled oven. Heating blocks or water baths can result in condensation of the small volume of liquid at the top of the tube, which may result in inefficient protein cleavage.
30. Transfer supernatant (“extract 1”) to a precleaned microcentrifuge tube. 31. Add an additional 20 µl of 40 mM ammonium bicarbonate/10% acetonitrile, pH 8.1, to extracted gel slices and incubate 45 min at 37◦ C. 32. Remove supernatant, combine with first extract, and add 4 µl concentrated acetic acid to reduce pH to ∼3. Store up to several months (or longer) at −80◦ C or continue immediately with LC-MS/MS analysis.
Perform LC-MS/MS with a linear ion trap and autosampler 33. Connect a 75 µm i.d. × 15 cm PicoFrit column, packed with MAGIC C18 resin, 5 µm particle size, to the mass spectrometer. Also, equip a microvolume autosampler with a 10-µl injection loop. Chapter 16 describes basic mass spectrometry techniques.
34. Set the flow rate to 200 nl/min and equilibrate the column with 1% Solvent B. 35. Inject 1 to 10 µl of each trypsin digest over a 10-min period with 1% Solvent B. Program the HPLC to execute the following gradient:
1% to 28% Solvent B over 50 min 28% to 50% Solvent B over 14 min 50% to 80% Solvent B over 5 min Return to 1% Solvent B over 1 min. The total run time is 75 min. The injection volume can be adjusted based on intensity of protein staining in individual digests to prevent overloading the column. To help minimize carryover between samples, a blank (no injection) should be run between samples. A reasonable blank cycle time is 35 min, giving a total time per sample of 120 min.
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36. Set the mass spectrometer to scan m/z from 375 to 1600, followed by data-dependent acquisition of MS/MS data of the dominant peaks. 37. Analyze the resulting data using MS/MS analysis software (e.g., Sequest). ALTERNATE PROTOCOL 1
MAJOR PROTEIN DEPLETION FOLLOWED BY 2-D GEL ANALYSIS One relatively simple proteome-analysis strategy for human serum or plasma involves the use of two-dimensional gel electrophoresis, preferably after depleting as many proteins as possible. This protocol describes depletion of six abundant proteins in human serum/plasma samples followed by analysis with 2-D PAGE. Alternatively, 2-D DIGE (UNIT 22.2) could be used.
Additional Materials (also see Basic Protocol 1) 1.1× IEF sample buffer (see recipe) Equilibration buffer I: add 463 mg, dithiothreitol (DTT) to 100 ml equilibration buffer base solution (see recipe) immediately before use Equilibration buffer II: add 4.37 g iodoacetamide to 100 ml equilibration buffer base solution (see recipe) immediately before use 24-cm pH 3-10 L IPG strips (Amersham Biosciences) 25 × 20 cm, 1 mm thick 10% Tris·glycine polyacrylamide gels (UNIT 10.4) Gel-image analysis software (e.g., Melanie 4; GeneBio) Additional reagents and equipment for 2-D electrophoresis using immobilized pH gradient gel strips (UNIT 10.4, Basic Protocol 2) and staining of gels (UNIT 10.5). 1. Deplete, desalt, and concentrate the desired volume of human serum or plasma samples as described in Basic Protocol 1 steps 1 to 15. 2. Dilute 40 µl of the concentrated serum or plasma from Basic Protocol 1, step 15, to 400 µl using 1.1× IEF sample buffer. Following depletion, the flowthrough protein from as much as 2 mg of original serum or plasma sample may be used for 2-D gel analysis. The amount of protein depleted should be dictated by the amount needed for 2-D gel analysis. Bound and flowthrough fractions from the depletion may be run in the same manner; however, substantially less bound fraction should be loaded, i.e., on the order of 100 µg, when using high-sensitivity MS-compatible silver stains such as Invitrogen’s Silver Quest.
3. Warm 24-cm pH 3-10 L IPG strips to room temperature. Alternatively 18-cm IPG strips can be used, although these gels will produce slightly lower resolution than 24-cm gels. IPG gels shorter than 18 cm are not recommended for such complex samples.
4. Load samples on to the IPG strip(s) by in-gel rehydration (UNIT Protocol 2). 5. Run IPG-IEF (first dimension; gradient at 20◦ C:
UNIT 10.4,
10.4,
Basic
Basic Protocol 2) using the following
1 hr rehydration 12 hr rehydration with 50-V step and hold Linear gradient to 250 V for 15 min Linear gradient to 10,000 V for 3 hr Hold at 10,000 V for 60,000 V-hr (overnight) Decrease to 500 V and hold. Human Serum and Plasma Proteomics
6. Equilibrate the IPG strip(s) in equilibration buffer I for 10 min at room temperature, then discard the buffer.
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7. Equilibrate the IPG strip(s) in equilibration buffer II for 10 min at room temperature. 8. Transfer IPG strip(s) to 25 × 20 cm, 1-mm thick second-dimension 10% Tris·glycine polyacrylamide gels. Run at 100-mA constant current with external cooling (UNIT 10.4, Basic Protocol 4). 9. Stain the gel using desired staining reagents such as Coomassie blue or silver stain (UNIT 10.5). Analyze gel images using gel-image analysis software (e.g., Melanie 4 from GeneBio).
MAJOR PROTEIN DEPLETION FOLLOWED BY OFFLINE MULTIDIMENSIONAL PROTEIN IDENTIFICATION TECHNOLOGY (MudPIT)
ALTERNATE PROTOCOL 2
Multidimensional Protein Identification Technology (MudPIT) on LC/LC-MS/MS employs the use of two chromatography steps to fractionate digested peptides (Link et al., 1999). Typically this involves the use of a strong cation exchange (SCX) column, followed by reversed-phase (RP) chromatography. This can be performed in either an online tandem manner or as two separate steps using offline SCX. The online approach requires a considerably more sophisticated HPLC apparatus with four pumps, and has more limited sample capacity. Offline SCX approaches can use either a basic dual-pump HPLC or other devices such as spin columns for the SCX fractionation. This protocol deals with analyzing major protein depleted serum via the two-step offline procedure (see UNIT 23.1).
Additional Materials (also see Basic Protocol 1) 50 mM ammonium bicarbonate in 10% (v/v) acetonitrile, pH 8.1 Modified trypsin (Promega) Additional reagents and equipment for SCX and RP-LC-MS/MS (UNIT 23.1, Basic Protocol 1) 1. Deplete, desalt, and concentrate the desired volume of human serum or plasma samples as described in Basic Protocol 1 steps 1 to 15. Reduce and alkylate sample as described in Basic Protocol 1, steps 16 to 20. 2. Dilute sample by adding 3 vol of 50 mM ammonium bicarbonate/10% acetonitrile, pH 8.1, to reduce urea concentration to 2 M. A low level of urea is needed to keep the proteins soluble and at least partially denatured. A high level of urea will inactivate the trypsin in the subsequent step.
3. Add 2 µg modified trypsin per 100 µg of protein and digest overnight at 37◦ C. 4. Perform SCX fractionation, then perform LC-MS/MS on SCX fractions using a linear ion trap mass spectrometer and autosampler (UNIT 23.1, Basic Protocol 1).
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see suppliers index.
Equilibration buffer base solution 0.05 M Tris·Cl, pH 8.3 (APPENDIX 2E) 6 M urea 30% (v/v) glycerol 2% sodium dodecyl sulfate (SDS; APPENDIX 2E) Milli-Q H2 O to 200 ml final volume Prepare immediately before use, do not store
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IEF sample buffer, 1.1× 9 M urea 2 M thiourea 4% (w/v) 3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate (CHAPS) 0.1 M dithiothreitol (DTT) Divide solution with above components into 1-ml aliquots and store up to several months at –20◦ C Just before use, add carrier ampholytes, pH 3-10L (IPG buffer; Amersham Biosciences) to 1.6% (v/v) final concentration COMMENTARY Background Information
Human Serum and Plasma Proteomics
Human plasma and serum proteomes may contain protein markers for every biological and disease process currently occurring in the body. Blood collection is minimally invasive, which makes blood an extremely attractive biological fluid for detecting disease biomarkers. However, human blood analysis is extremely challenging due to the dynamic range and complexity of the blood proteome. Anderson and Anderson (2002) divide the plasma proteome into eight different design and function groups as detailed below, which helps illustrate the complexity of the problem. 1. Proteins secreted by solid tissues with specific primary functions in the blood: These are the high-abundance classical plasma proteins, secreted to remain and function in the plasma. Albumin is the most obvious example. 2. Immunoglobulins: Millions of different unique antibodies are routinely circulating in the plasma of each adult. 3. “Long distance” receptor ligands: These comprise peptide and protein hormones such as insulin. 4. “Local” receptor ligands: These comprise cytokines and other local mediators of cellular response. 5. Temporary passengers: These are defined as proteins traversing the plasma from their place of secretion to their place of primary function. 6. Tissue leakage products: These are proteins that normally function in specific cell types, but are released into the blood as a result of cell death or damage. 7. Aberrent secretions: These are proteins that should not be secreted normally, but happen to be released by diseased tissues. 8. Foreign proteins: These include proteins from infectious organisms and parasites. These classes of proteins all contribute to the complexity of human plasma. Groups 6, 7,
and 8 are the ones thought most likely to contain biomarkers, and are thus the most interesting classes for the types of studies described in this unit. Other groups, such as 1 and 5, may also play a role in some diseases and other biological processes, and, therefore, could merit investigation, as well (Anderson et al., 2002). The dynamic range of human plasma and serum presents another major hurdle. Most current analytical techniques use a maximum of two separation dimensions, e.g., 2-D gels or MudPIT, and have a limited ability to examine a broad dynamic range. 2-D gels are capable of detecting proteins over concentration ranges of a maximum of about 3 to 4 orders of magnitude. In human serum and plasma, the 22 most abundant proteins represent 99% of the protein content (Tirumalai et al., 2003) and the concentrations range from tens of milligrams per ml to pg/ml or less. The very wide range of concentrations in serum and plasma makes more than two fractionation steps essential for detection of low-abundance proteins. However, even these techniques provide only a sampling of the total pool of proteins present in the blood (Omenn et al., 2005). Sample collection and handling There are many points in the collection and handling of samples at which substantial variation and error can occur. In order to minimize this, protocols should be established and followed rigidly. All sample collection and handling conditions should be carefully monitored and recorded. Serum and plasma are relatively resistant to degradation at room temperature compared to cell lysates. However, a detectable level of ex vivo proteolysis and other modifications still occur (Rai et al., 2005). For this reason, length of time prior to freezing should be kept minimal and consistent. Also, the number of freeze/thaw cycles should be minimized and samples should be stored at −80◦ C, or preferably in liquid nitrogen.
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In a further effort to avoid the degradation of samples, protease inhibitors (PIs) can be employed during the course of experimentation, including during blood collection. Processing requires a minimum of 10 min for plasma and 30 min for serum after the blood is drawn. During this time, biological processes are occurring, including proteolysis (Marshall et al., 2003). To explore this issue further, the addition of PIs at the time of blood draw was analyzed using SELDI in a recent study. There was some evidence that samples in which the PIs were added at the time of blood draw decreased proteolysis (Rai et al., 2005). However, the effects of PIs are not entirely clear. On 2-D gels, there is no direct evidence that PIs offer any protection against a decrease in molecular weight, and some PIs that react covalently with targeted proteins may cause certain human plasma protein isoforms to shift to a higher pI than normal. This is apparently due to covalent modifications of nontargeted plasma proteins caused by the PI (Schuchard et al., 2005). For this reason, care should be taken when working with PIs, and those that cause covalent modifications should be avoided (Rai et al., 2005). One other factor to consider is whether to use serum or plasma. Currently, most archives of clinical specimens are composed of serum. However, due to the blood-clotting process, which involves activation of proteases, there is reason to believe that conditions such as temperature and processing time at collection will substantially affect sample variability. There is also the possibility that certain proteins will bind to the clotting components in an uncontrolled manner, adding further variation to the sample. Serum also has a tendency to show a number of intense peptide signals produced by the blood clotting proteolysis, which may interfere with certain types of analysis such as SELDI. These factors must be taken into consideration when deciding on what type of sample and analysis method to use (Rai et al., 2005). Serum and plasma proteomics analysis techniques There are currently many different methods of protein identification employed in proteomics. Part of the success of these techniques comes from the ability to mix and match basic methods to increase the capacity for analyzing extremely complex mixtures that cannot effectively be analyzed by one method alone. The field of serum and plasma proteomics continues to evolve rapidly in an attempt to overcome
the twin issues of dynamic range and sample complexity. Most of the current state-of-theart protein-profiling strategies employ a series of integrated methods that use more than two dimensions of separation. This is illustrated in Figure 24.1.4, which shows potential ways of combining protein separation, peptide separation, and MS/MS analysis. The methods described in this unit could be termed protein identification discovery methods. Discovery-based methods are those that make no prior assumptions about what proteins are present. One type of discovery-based method, which employs pattern matching, is represented by the SELDI-MS technology and related approaches. Some studies using this technology have claimed exceptionally good results (Petricoin et al., 2002; Villanueva et al., 2004); however, there is considerable debate about the merits and reliability of this approach (Check et al., 2004, Diamandis et al., 2004). SELDI-MS is one of the few high-throughput protein-profiling analysis methods, and requires less technical skill than most other methods. There are, in particular, questions regarding the reproducibility of the technique, which provides good sensitivity and resolution only in the low-mass range. Recently, patternmatching approaches involving peptides rather than proteins have been employed in combination with a MudPIT strategy. In this approach, the pattern created by the m/z value versus the retention time on the LC column is compared across samples. However, since this technique employs the use of an LC-MS/MS device, each spot in the pattern can be easily identified as a specific peptide. In contrast, it is usually quite difficult to identify the proteins or peptides responsible for features of interest in SELDI-MS patterns. The only other high-throughput proteomics analysis tool currently available utilizes antibody arrays. Similar to multiplexed ELISAs, these assays trap and quantitate specific proteins using antibodies or antibody mimics. Antibody arrays have the advantage of being extremely high throughput, but are limited to existing antibodies. Because not all antibodies can function in such assays, the number of potential proteins that can be profiled is severely limited. Mass spectrometry is not inherently quantitative, which presents some difficulty when comparing different human serum or plasma samples. To provide reliable quantification with most MS methods, stable isotope labeling strategies are usually used. Since metabolic labeling is not feasible for human subjects,
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Figure 24.1.4 Alternative protein and peptide separation methods and MS or MS/MS analysis methods. This flow diagram illustrates the most common protein separation, peptide separation and mass spectrometry methods that can be integrated into multimethod protein-profiling strategies for plasma or serum analysis.
Human Serum and Plasma Proteomics
post-isolation stable isotope labeling strategies such as ICAT, iTRAQ, or labeling during trypsin digestion with H2 [18 O] must be used. However, any variations in protein yield prior to the labeling step produces experimental noise. Alternative quantitative techniques applicable to serum and plasma analysis include: 2-D gels, 2-D DIGE, and antibody arrays. Until recently, two-dimensional gel electrophoresis has dominated the proteomics field. It is one of the most established technologies, and has been in use for almost 30 years. 2-D gels consist of an initial isoelectric focusing (IEF) phase followed by an SDS gel. This separation, based on both pI and size, is capable of detecting many different isoforms of proteins. It also provides absolute quantitation and allows ready comparison of quantitative differences between samples. However, there are substantial disadvantages as well. 2-D gels have a limited dynamic range, and in the case of serum or plasma, the abundant proteins, with their many isoforms, cover most of the gel and mask lowerabundance proteins. Depletion helps address
this problem to some extent by allowing larger loads (Fig. 24.1.1A and B), but even at higher loads, most proteins observed are in the µg/ml range or higher. The technique is also time consuming and difficult, requiring both skill and experience, and reproducibility remains suboptimal.Two-dimensional differential gel electrophoresis (2-D DIGE; UNIT 22.2) is a variation of the 2-D gel technique that uses cyanine dyes to label samples prior to electrophoresis. Each sample is labeled with a different dye (Cy2, Cy3, and Cy5), which allows a direct quantitative comparison of up to three samples, or more commonly two samples and a reference standard, on the same gel. This largely alleviates the reproducibility problems of standard 2-D gels. One potential limitation is that the dyes cause a slight offset from the actual location of the bulk of the protein because only a small percentage of each protein is actually labeled with the dye. This could lead to errors in excising and correctly identifying spots of interest. Nonetheless, the ability to compare three different samples at once without complications that result from gel-to-gel variability makes this an extremely useful technique.
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Multidimensional analysis is any strategy that combines several different basic methods to increase resolution and dynamic range of detection. These strategies tend to have better sensitivity, with increased detection of low-abundance proteins. In contrast, basic, conventional protein-profiling methods typically utilize, at most, two dimensions of separation, e.g., 2-D gels, 2-D DIGE, and MudPIT (LC/LC-MS/MS). MudPIT can detect a greater number of lower-abundance proteins than 2-D methods. However, even with this method, only a very limited number of lowabundance proteins are usually detected, because it only has two dimensions of separation after the complex mixture is digested to tryptic peptides. However, most multidimensional methods are fairly low-throughput and usually require state-of-the-art HPLC instrumentation and high-performance ESI mass spectrometers that are not necessarily available in all labs. The 4-D strategy discussed in Basic Protocol 1 is a good example of a multidimensional analysis with a wide dynamic range, which can also be made quantitative through the use of ICAT, iTRAQ, or other stable isotope labeling strategies.
Critical Parameters and Troubleshooting One of the most important parameters involved in the analysis of human serum/plasma is an understanding of the time and cost required. While combining multiple fractionation strategies can result in powerful multidimensional analysis strategies, throughput can take a dramatic hit very quickly. Four critical parameters for selecting the best analysis strategy are: careful evaluation of the minimum number of samples that must be analyzed to accomplish a research task, how many proteins must be detected, what concentration ranges must be detected, and what types of changes are important. For example, if detection of isoforms or post-translational modifications is important, the best strategy is likely to incorporate 2-D DIGE. Another critical parameter is whether quantitative comparisons are to be performed or if the goal is simply to detect as many peptides as possible. Major protein depletion of human serum or plasma as described here is HPLC-based, so much of the potential troubleshooting is discussed in UNIT 8.7. Other issues that must be considered are specific to the column itself; these include potential overloading of the column and optimization of specific samples. Both of these issues can be addressed by care-
fully analyzing the amount of sample introduced onto the column and evaluating the unbound and bound fractions on 1-D and 2-D gels. Another consideration is that these antibody columns contain covalently limited functional proteins. Therefore, care must be exercised to avoid exposure of the columns to proteolysis or conditions that could irreversibly denature or otherwise modify the antibodies. Troubleshooting for MicroSol IEF is discussed in UNIT 22.6, and troubleshooting of 2-D gels and 2-D DIGE are discussed in UNITS 10.4 & 22.2, respectively. Troubleshooting of MudPIT is discussed in UNIT 23.1, and troubleshooting for RP-HPLC and LC-ESI MS/MS is discussed in UNIT 16.9.
Anticipated Results The protocols detailed in this unit provide several different options for proteomic analysis of human serum or plasma using three or four separation dimensions. Each of these methods has been successfully employed for identification of blood proteins. Basic Protocol 1 allows identification of the largest number of proteins, detecting more than 1000 proteins with high confidence. These identifications will range from major proteins (mg/ml) to detection of a substantial number of proteins at the low ng/ml level and a few proteins at the pg/ml level. Alternate Protocol 1 should facilitate the analysis of major serum proteins in the mg/ml and µg/ml range, including most isoforms of these proteins. Alternate Protocol 2 should identify on the order of 100 to 350 proteins with high confidence.
Time Considerations Multidimensional proteomic analysis of human serum or plasma is for the most part a low-throughput process. The depletion portion of Basic Protocol 1 will take ∼1 day, including setting up the HPLC, running up to four replicate runs, and flushing the column and HPLC system for storage. The 1-D gels take ∼1 day, and in-gel digestion of gel slices takes about 1.5 days. The entire Basic Protocol 1 will take a total of about 2 to 3 weeks, depending on the total number of gel slices analyzed and the length of the HPLC gradients utilized. Alternate Protocol 1 will take about 1 day per sample for major protein depletion and 3 to 5 days to run up to twelve 2-D gels. Alternate Protocol 2 will take about 1 day per sample for major protein depletion, 1 day for the tryptic digestion, 1 day to perform the SCX column separation, and 2 to 4 days to perform the LCMS/MS analysis, depending on the number of
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SCX fractions analyzed and the length of the HPLC gradient.
Literature Cited Anderson, N.L. and Anderson, N.G. 2002. The human plasma proteome: History, character, and diagnostic prospects. Mol. Cell. Proteomics 1:845-867. Anderson, N.L., Polanski, M., Pieper, R., Gatlin, T., Tirumalai, R.S., Conrads, T.P., Veenstra, T.D., Adkins, J.N., Pounds, J.G., Fagan, R., and Lobley, A. 2004. The human plasma proteome: A nonredundant list developed by combination of four separate sources. Mol. Cell. Proteomics 3:311-326. Chan, K.C., Lucas, D.A., Hise, D., Schaefer, C.F., Xiao, Z., Janini, G.M., Buetow, K.H., Issaq, H.J., Veenstra, T.D., and Conrads, T.P. 2004. Analysis of the human serum proteome. Clinical Proteomics 1:101-226. Check, E. 2004. Proteomics and cancer: Running before we can walk? Nature 429:496-497. Diamandis, E.P. 2004. Mass spectrometry as a diagnostic and a cancer biomarker discovery tool: Opportunities and potential limitations. Mol. Cell. Proteomics 3:367-378. Echan, L.A., Tang, H.Y., Ali-Khan, N., Lee, K., and Speicher, D.W. 2005. Depletion of multiple high abundance proteins improves protein profiling capacities of human serum and plasma. Proteomics 5:3292-3303. Link, A.J., Eng, J., Schieltz, D.M., Carmack, E., Mize, G.J., Morris, D.R., Garvik, B.M., and Yates, J.R. 3rd. 1999. Direct analysis of protein complexes using mass spectrometry. Nat. Biotechnol. 17:676-682. Marshall, J., Kupchak, P., Zhu, W., Yantha, J., Vrees, T., Furesz, S., Jacks, K., Smith, C., Kireeva, I., Zhang, R., Takahashi, M., Stanton, E., and Jackowski, G. 2003. Processing of serum proteins underlies the mass spectral fingerprinting of myocardial infarction. J. Proteome. Res. 2:361-372. Misek, D.E., Kuick, R., Wang, H., Galchev, V., Deng, B., Zhao, R., Tra, J., Pisano, M.R., Amunugama, R., Allen, D., Walker, A.K., Strahler, J.R., Andrews, P., Omenn, G.S., and Hanash, S.M. 2005. A wide range of protein isoforms in serum and plasma uncovered by a quantitative intact protein analysis system. Proteomics 5:3343-3352.
Human Serum and Plasma Proteomics
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Omenn, G.S., States, D.J., Adamski, M., Blackwell, T.W., Menon, R., Hermjakob, H., Apweiler, R., Haab, B.B., Simpson, R.J., Eddes, J.S., Kapp, E.A., Moritz, R.L., Chan, D.W., Rai, A.J., Admon, A., Aebersold, R., Eng, J., Hancock, W.S., Hefta, S.A., Meyer, H., Paik, Y.K., Yoo, J.S., Ping, P., Pounds, J., Adkins, J., Qian, X., Wang, R., Wasinger, V., Wu, C.Y., Zhao, X., Zeng, R., Archakov, A., Tsugita, A., Beer, I., Pandey, A., Pisano, M., Andrews, P., Tammen, H., Speicher, D.W., and Hanash, S.M. 2005. Overview of the HUPO Proteome Project: Results from the pilot phase with 35 collaborating laboratories and multiple analytical groups, generating a core dataset of 3020 proteins and a publically available database. Proteomics 5:3226-3245.
Petricoin, E.F., Ardekani, A.M., Hitt, B.A., Levine, P.J., Fusaro, V.A., Steinberg, S.M., Mills, G.B., Simone, C., Fishman, D.A., Kohn, E.C., and Liotta, L.A. 2002. Use of proteomic patterns in serum to identify ovarian cancer. Lancet 359:572-577. Rai, A.J., Gelfand, C.A., Haywood, B.C., Warunek, D.J., Yi, J., Schuchard, M.D., Mehigh, R.J., Cockrill, S.L., Scott, G.B., Tammen, H., SchulzKnappe, P., Speicher, D.W., Vitzthum, F., Haab, B.B., Siest, G., and Chan, D.W. 2005. HUPO plasma proteome project specimen collection and handling: Towards the standardization of parameters for plasma proteome samples. Proteomics. 5:3262-3277. Schuchard, M.D., Mehigh, R.J., Cockrill, S.L., Lipscomb, G.T., Stephan, J.D., Wildsmith, J., Valdes-Camin, R., Kappel, W.K., Rai, A.J., and Scott, G.B. 2005. Artifactual isoform profile modification following treatment of human plasma or serum with protease inhibitor, monitored by 2-dimensional electrophoresis and mass spectrometry. Biotechniques. 39:239-247. Shen, Y., Jacobs, J.M., Camp, D.G. 2nd, Fang, R., Moore, R.J., Smith, R.D., Xiao, W., Davis, R.W., and Tompkins, R.G. 2004. Ultra-high-efficiency strong cation exchange LC/RPLC/MS/MS for high dynamic range characterization of the human plasma proteome. Anal. Chem. 76:11341144. Tammen, H., Schulte, I., Hess, R., Menzel, C., Kellmann, M., Mohring, T., and Schulz-Knappe, P. 2005. Peptidomic analysis of human blood specimens: Comparison between plasma specimens and serum by differential peptide display. Proteomics 5:3414-3422. Tang, H.Y., Ali-Khan, N., Echan, L.A., Levenkova, N., Rux, J.J., and Speicher, D.W. 2005. A novel four-dimensional strategy combing protein and peptide separation methods enables detection of low-abundance proteins in human plasma and serum proteomes. Proteomics. In press. Tirumalai, R.S., Chan, K.C., Prieto, D.A., Issaq, H.J., Conrads, T.P., and Veenstra, T.D. 2003. Characterization of the low molecular weight human serum proteome. Mol Cell. Proteomics. 2:1096-1103. Villanueva, J., Philip, J., Entenberg, D., Chaparro, C.A., Tanwar, M.K., Holland, E.C., and Tempst, P. 2004. Serum peptide profiling by magnetic particle-assisted, automated sample processing and MALDI-TOF mass spectrometry. Anal. Chem. 76:1560-1570.
Internet Resources http://www.specialtylabs.com Specialty Laboratories publications, Santa Monica, Ca. August 10, 2005. This web site provided the information shown in Figure 24.1.2. Searching the site for specific proteins will provide information on abundance ranges.
Contributed by Ian Musselman and David W. Speicher The Wistar Institute Philadelphia, Pennsylvania Current Protocols in Protein Science
Organelle Proteomics
UNIT 24.2
With advances in proteomics studies, it becomes increasingly obvious that currently available proteomics technologies are insufficient to handle the enormous complexity of biological samples. There are two major approaches for improvement. The first is to increase the resolving power of the proteomics technologies, i.e., increase the number of different proteins that can be separated by the analytical techniques used (e.g., chromatographic or electrophoretic separations). While this is, in some sense, the ideal way, it is also the most difficult and the one for which the results are most difficult to anticipate. The second approach is to reduce the complexity of the sample to be analyzed, so it more closely fits the current performance capacities of the proteomics toolbox. For eukaryotic cells, one method of sample simplification is to fractionate the cells into various component structures such as organelles (i.e., membrane-bound subcellular structures delineated by lipid membranes), or even smaller functional cellular components, which are increasingly referred to as subcellular machines (e.g., nuclear pores and nucleoli). The advantage is that each isolated component is far less complex than the complete cell in terms of the population of proteins present. Thus, a current trend in proteomics is to restrict analysis to selected subcellular components instead of using whole-cell lysates. In addition to providing a better analysis depth (see Critical Parameters) by reducing the complexity of the sample to better accommodate the performance limits of the methods, this approach also provides additional biologically valuable information such as subcellular localization of proteins. Another advantage is that targeted proteomics allows the investigator to ask more focused questions than are possible using global protein profile comparisons of whole cells or tissues. Organelle-targeted proteomics typically follows a series of steps that lead to the preparation of the subcellular structure(s) of interest with sufficient purity and reproducibility. Specific protocols for subcellular fractionation of tissue culture cells (UNIT 4.3) and isolation of organelles using free-flow electrophoresis (UNIT 22.5) have been described As the cited units deal with the isolation of membrane-derived organelles, representative protocols are provided here for isolation of nuclei from cultured cells and tissues. The steps following sample preparation in organelle-targeted proteomics concern the proteomic analysis per se. This subject is addressed in Chapter 22 (especially UNITS 22.1, 22.2, & 22.4), with specific aspects also described in Chapters 10 (Electrophoresis) and 16 (Mass Spectrometry), and is consequently not discussed here. However, a brief Background Information section is provided to compare specific aspects of organelle proteomics to the more general aspects discussed in the units mentioned above.
STRATEGIC PLANNING For targeted proteomics studies, strategic planning will depend on whether a single or several components isolated from the biological source are to be studied. For a single component (e.g., nuclei), specific isolation procedures can be used that may result in damage to, or gross contamination of, other structures, which are then discarded in this type of analysis. Contamination or damage to other components cannot be tolerated when several components are to be analyzed. However, single-component-oriented procedures usually result in higher purity of the target component. For example, nuclei are resistant to nonionic detergent because of the lamina, which is not the case for most other organelles (e.g., Golgi, mitochondria, endoplasmic reticulum). Thus, isolation of nuclei in the presence of low concentrations of nonionic detergents (see Basic Protocol 2) Targeted Proteomics Contributed by Pierre Lescuyer, Mireille Chevallet, Sylvie Luche, and Thierry Rabilloud Current Protocols in Protein Science (2006) 24.2.1-24.2.9 C 2006 by John Wiley & Sons, Inc. Copyright
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destroys other organelles that would otherwise contaminate the preparation of nuclei (e.g., mitochondria). Conversely, simultaneous isolation of multiple organelles from the same source prevents the use of such methods, as membrane integrity must be preserved. In this case, the cells must be lysed mechanically or osmotically. Osmotic lysis is more gentle, but can be used only for cultured cells. Mechanical lysis, which induces more organelle breaking, is used for tissues or for cells that are resistant to osmotic lysis (e.g., plant cells, yeast). Differential centrifugation strategies using both the density of the organelles and their sedimentation velocity are then used to separate the various types of organelles (typically nuclei, mitochondria, and microsomes/lysosomes). However, the resulting fractions are never pure, and contamination of microsomes by broken mitochondria is very common. The presence of lysosomal proteins in mitochondrial proteomic maps is also very common. Starting from these enriched fractions, further purification can be afforded by immunopurification (UNITS 9.5 & 9.8) or free-flow electrophoresis (UNIT 22.5). While these methods usually result in pure organelles, their yield is often low and a significant portion of the starting material is lost. BASIC PROTOCOL 1
WORKFLOW FOR ORGANELLE PROTEOMICS ANALYSIS This protocol illustrates the basic workflow for organelle proteomics analysis. Detailed protocols showing examples of organelle preparation and centered on nuclei isolation are provided in Basic Protocol 2 and Alternate Protocols 1 and 2. 1. Isolate the component(s) of interest from the biological source. For all organelles except nuclei, follow the protocols described in UNITS 4.3 & 22.5. For the direct isolation of nuclei, follow Basic Protocol 2 or Alternate Protocol 1 in this unit. For the simultaneous isolation of nuclei and other organelles, follow Alternate Protocol 2. The basic point of departure for the isolation of subcellular components is first to lyse the cell under conditions that do not damage the components of interest. This is why nuclei, which are limited by a lamina, can be isolated with nonionic detergents (e.g., using Alternate Protocols 1 and 2), while membrane-limited organelles cannot. This means that the use of detergents in nuclei isolation will dramatically lower the extent of contamination by organelles such as lysosomes or mitochondria. Once the cells have been opened, the components of interest are generally purified by differential centrifugation (UNIT 4.2), exploiting differences in the sedimentation velocity and/or buoyant density of the various subcellular components. It must be stressed, however, that there is often a significant overlap in these parameters for various subcellular components. As a result, significant cross-contamination is often encountered—for example, between mitochondria and lysosomes. It is also often difficult to resolve the total microsomes into the plasma membrane, endoplasmic reticulum (ER), and Golgi components. For such delicate final purification, an interesting and useful method is free-flow electrophoresis (UNIT 22.5). However, this technique requires a special apparatus. Furthermore, increased purity is usually accompanied by lowered yields. Finally, many subcellular components are linked to the cytoskeleton. As this latter component is very strongly assembled, contamination of subcellular structures with cytoskeleton is commonplace and very difficult to overcome.
2. Extract the proteins as described in UNIT 22.4 3. Analyze the proteins by the appropriate proteomics methods, e.g., those described in UNITS 22.1 & 22.2. Organelle Proteomics
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The choice of method will depend on several, often mutually exclusive, considerations. As an example, membrane proteins are poorly analyzed by techniques based on twodimensional electrophoresis because of solubility problems in the isoelectric focusing dimension. However, alternative techniques that perform adequately with membrane proteins, such as those based on SDS electrophoresis followed by LC/MS-MS, are not quantitative and offer poor performance in the analysis of post-translational modifications because of the low number of peptides analyzed per protein. The probability of detecting a modified peptide is therefore significantly reduced. The final choice of the experimental proteomics approach will then depend on the aspect that is judged to be most important in the particular study.
DIRECT ISOLATION OF NUCLEI FROM CULTURED CELLS This protocol describes the rapid and direct isolation of nuclei from cultured cells. It is based on the use of nonionic detergents such as Triton X-100 to dissolve the lipid bilayers present in the cells, including the plasma membrane and all the inner membrane–bound inner organelles (e.g., Golgi, ER, mitochondria), which are thus irreversibly lost. It also dissolves the ER-derived cisternae forming the outer nuclear membrane, usually resulting in pure nuclei.
BASIC PROTOCOL 2
Materials Cells growing in tissue culture Cell washing solution (see recipe), ice cold Cell lysis solution (see recipe), ice cold Nuclei wash solution (see recipe), ice cold Nuclei storage solution (see recipe), ice cold Liquid nitrogen Refrigerated centrifuge 1. Harvest cells and collect by centrifuging 5 min at 1000 × g, 4◦ C. Remove the supernatant. Wash twice, each time by adding 10 vol of ice-cold cell washing solution, resuspending by pipetting, centrifuging again as before, and removing the supernatant. Estimate the volume of the packed cell pellet. This protocol works for cells without a cell wall, i.e., mainly mammalian cells. It also works for cultured insect cells. Because nuclei often represent ∼10% of the cell protein mass, 10 to 100 million cells are needed to be able to carry out a successful protemics analysis.
2. Resuspend the cell pellet by vortexing in 10 vol of ice-cold cell lysis solution. Incubate on ice for 20 min, vortexing for 20 sec every 5 min. 3. Collect the nuclei by centrifuging 5 min at 1000 × g, 4◦ C. Remove the supernatant. Resuspend pellet by vortexing in the same volume of ice-cold cell lysis solution that was used for the first lysis. Incubate on ice for 5 min, then centrifuge again as before. Discard the supernatant. 4. Resuspend pellet in the same volume of ice-cold nuclei wash solution that was used for the first lysis. Incubate on ice for 5 min, then centrifuge as described in step 3. Decant and discard supernatant. Repeat this rinse one additional time. 5. After the last centrifugation, estimate the nuclei pellet volume and resuspend thoroughly by vigorous pipetting and vortexing in a volume of storage solution equal to the estimated pellet volume. Divide into 0.1-ml aliquots and store at −80◦ C (stable for at least 1 year), or preferably in liquid nitrogen. See Support Protocol to assess purity.
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ALTERNATE PROTOCOL 1
DIRECT ISOLATION OF NUCLEI FROM CELLS WITH WALLS OR TISSUES USING MECHANICAL LYSIS Basic Protocol 2 employs an osmotic/detergent lysis which is efficient only for cultured cells devoid of a cell wall. Cells with a wall or tissues require a more efficient, vigorous procedure, namely mechanical lysis.
Additional Materials (also see Basic Protocol 2) Tissue sample or cells with walls Dounce glass-glass homogenizer or blade-type homogenenizer (e.g., Waring blender) for samples resistant to lysis 1. Cut the tissue of interest into small pieces with scissors and wash the pieces with cell washing solution. Estimate the tissue volume once settled. Homogenize the tissue pieces in 10 pellet volumes of cell lysis solution using a Dounce glass-glass homogenizer (ten strokes are usually sufficient). For tissues (e.g., plant) that are very resistant to lysis, blade-type homogenizers (e.g., Waring blender) are generally used. While causing much more efficient cell breakage, they also lead to nuclei breakage, thereby decreasing the overall yield. As a rule, the gentlest homogenizer leading to efficient cell disruption should be used. This is why Dounce homogenizers are used for most soft tissues.
2. Collect and wash the nuclei as described in Basic Protocol 2. ALTERNATE PROTOCOL 2
SIMULTANEOUS PURIFICATION OF NUCLEI AND OTHER ORGANELLES FOR PROTEOMICS STUDIES Basic Protocols 1 and 2 use direct disruption of the cells in a buffer containing Triton X-100. As this results in irreversible loss of most organelles other than the nuclei, these protocols cannot be used when other organelles are also to be collected. In this case, cell-disruption protocols respecting the integrity of the organelles are used (see UNIT 4.3). These protocols generally provide a nuclei-rich pellet at the end of the first low-speed centrifugation. This pellet contains nuclei, unbroken cells, and contaminating organelles (e.g., mitochondria or membrane sheets). Further purification of this crude nuclei preparation is therefore advisable and is described in this protocol.
Additional Materials (also see Basic Protocol 2) Starting material: crude nuclei pellet provided by a detergent-free cell lysis procedure (UNIT 4.3) Cheesecloth 1. Filter the nuclei pellet resulting from a detergent-free cell lysis procedure (UNIT 4.3) through cheesecloth to collect unlysed pieces of tissue. Centrifuge 5 min at 1000 × g, 4◦ C, to collect nuclei contaminated with unbroken cells and other large organelles (e.g., mitochondria). 2. Continue as described in steps 2 to 5 of Basic Protocol 2. Resuspend the initial nuclei pellet in 10 volumes using a Dounce glass-glass homogenizer. Five strokes are usually sufficient. 3. Collect and wash the nuclei as described in Basic Protocol 2.
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EVALUATION OF NUCLEI PURITY BY MICROSCOPIC EXAMINATION Because of the use of a detergent, the nuclei prepared according to Basic Protocol 2 or Alternate Protocols 1 or 2 are generally devoid of contamination by other organelles. The specific enzyme test described in UNIT 4.3 does not provide an accurate estimate of purity. The most likely contaminants in these nuclei preparations are cytoskeletal proteins and ribosomes. Although these are difficult to quantify, a good estimate of nuclei purity can be provided by microscopic examination using the following protocol.
SUPPORT PROTOCOL
Materials Microscopy stain solution (see recipe) Nuclei suspension (see Basic Protocol 2 or Alternate Protocols 1 or 2) Microscope slide and coverslip Light microscope with 200× magnification 1. Add one drop of microscopy stain solution to one drop of nuclei suspension on a microscope slide. Cover with a coverslip and leave at room temperature for 10 min before examination on a microscope. Magnification should be at least 200×, and phase contrast provides additional information. The staining solution contains two cationic stains: pyronin Y and methyl green. Methyl green has a strong affinity for DNA, while pyronin Y stains RNA, and mainly rRNA, which is the most abundant in cells Nuclei appear light green. One or two pink-colored nucleoli should be visible inside the nuclei. Contaminating cytoplasm results in pink (i.e., ribosome-containing) material outside the nuclei. The cytoskeleton is visible under phase contrast as refringent filaments.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Cell lysis solution 10 mM HEPES, pH 7.5 (adjust pH with NaOH) 1 mM dithiothreitol (DTT) 1 mM spermidine trihydrochloride 0.25 mM spermine tetrahydrochloride 0.5 mM EDTA (APPENDIX 2E) 0.1% (w/v) Triton X-100 Store solution with above components up to 1 week at 4◦ C Just before use, add 30 µM of 3,4 dichloroisocoumarin (Fluka; add from 30 mM stock prepared in dimethylformamide, stored at −20◦ C) Cell washing solution 10 mM sodium phosphate buffer, pH 7.5 (APPENDIX 2E) 0.125 M NaCl Store up to 3 months at 4◦ C Microscopy stain solution Dissolve 150 mg of methyl green and 250 mg of pyronin Y in 2.5 ml of 95% ethanol. Add 50 mM sodium acetate buffer, pH 4.8 (APPENDIX 2E) to 100 ml. Add 40 g sucrose and dissolve by mixing. Store up to 3 at 4◦ C in tightly stoppered flasks (to avoid evaporation of the acetic acid). Targeted Proteomics
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Nuclei storage solution 10 mM HEPES, pH 7.5 (adjust pH with NaOH) 1 mM dithiothreitol (DTT) 25% (v/v) glycerol 5 mM MgCl2 Store up to 3 months at 4◦ C Nuclei wash solution 10 mM HEPES, pH 7.5 (adjust pH with NaOH) 1 mM dithiothreitol (DTT) 0.2 M sucrose 2 mM MgCl2 Store up to 1 week at 4◦ C COMMENTARY Background Information
Organelle Proteomics
The objective of targeted proteomics studies is to analyze a defined subset of cellular proteins in order to adapt (reduce) the complexity of the sample to the performance limits of the analysis tools. Although the trend toward such an approach began some time ago (Wilkins et al., 1998; Gygi et al., 2000), more recently it has become increasingly apparent that the proteomics toolbox, i.e., the combination of analytical techniques used for proteomics (chromatography, electrophoresis, and mass spectrometry), cannot accommodate the complexity of real biological samples. This holds true for all the proteome analysis methods described to date, whether based on 2-D gels (Gygi et al., 2000) or not (Wu et al., 2003). As a result of the above mentioned limitations, the idea of analyzing only defined cellular subsets of proteins became popular a few years ago (Rabilloud et al., 1998; Jung et al., 2000). These subsets can be obtained by biochemical fractionation of wholecell or tissue extracts (Fountoulakis et al., 1997; Zuo et al., 2000), but the performance (especially the resolution) of classical biochemical fractionation techniques is somewhat limited. Another approach, namely organelle proteomics, was developed to take advantage of the existing compartmentalization of eukaryotic cells. In addition to providing samples of reduced complexity for proteomics analysis, organelle proteomics provides cell biology–relevant information, e.g., detecting the presence of “new” or “unexpected” proteins in a given organelle compartment. This information can, in turn, facilitate important cell biological observations (Gagnon et al., 2002). The relevance of organelle proteomics depends on the purity and reproducibility of
the organelle preparation. These parameters function at different levels to modulate the quality of the final proteomics analysis. For example, organelle preparations are often contaminated by proteins originating from different parts of the cell. This contamination alters the downstream proteomics analysis at two levels. First of all, contamination introduces misleading information about the subcellular location of proteins. While this is not a major problem for well known proteins that can be easily reassigned to their known location, this can be a significant problem for poorly described proteins that can be erroneously assigned to an organelle just because they contaminate the preparation of the organelle of interest. Secondly, the extent of contamination may vary from one preparation to another, thereby varying the proportion of proteins really belonging to the organelle of interest. This will, in turn, complicate the comparative analysis of organelles under two circumstances of interest.
Critical Parameters The purity of the organelle preparation plays a major role in the accuracy of the proteomics analysis. Samples of low purity will contain contaminating proteins that will also be present in the organelle map. These contaminants can lead to misinterpretation of the results, especially when unexpected proteins are present in the map. Another important consequence of organelle purity is the depth of the resulting proteomics analysis, i.e., the ability to detect proteins of low abundance. One example of this phenomenon is provided by the analysis of mitochondria (Zischka et al., 2003). Even for organelle proteomics, the limiting factor is the capacity of the proteomics toolbox.
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Consequently, when organelle samples of low purity are analyzed, contaminating proteins occupy a substantial fraction of the analysis space (2-D gels or peptide MS peaks) and prevent minor, bona fide organelle proteins from being detected and analyzed. This problem gradually disappears with increasing organelle purity, allowing the analysis of minor proteins (Zischka et al., 2003). A second critical point is the reproducibility of the organelle preparation. While poor reproducibility is a minor problem for cartographic (mapping) proteomics purposes (i.e., to determine the protein composition of a given sample), it becomes a major problem when comparative, physiology-oriented studies are carried out, as is common in clinical proteomics. Problems with reproducibility can be difficult to overcome, since the changes induced by the pathology studied or upon the physiological process of interest can interfere with the organelle preparation. As an example, any physiological alteration resulting in a major change in the cytoskeleton, a common major source of contamination in organelle preparations, will alter the purity of the final organelle preparation. This will be reflected by changes in the proteomics map when comparing the different physiological situations of interest. However, these changes are artifactual and, although linked to a change in the cell physiology, are not really related to the organelle of interest. Such a situation can lead to misinterpretation of the results, e.g., erroneously assigning an organelle a role in a pathology. The robustness of the protocol used for organelle preparation is therefore quite critical in such comparative studies, and rather difficult to establish, as a protocol performing well with control samples can fail insidiously with other samples used in the study. Another major caveat in targeted proteomics is sample consumption, as exemplified by proteomics of nuclear proteins. Nuclei represent 10% of the total cell protein mass, and the isolation procedures have a yield close to 50%. This means, in turn, that 5% of the initial sample is used at the end and 95% of the protein content is lost. While such losses can be accommodated in some experimental models, they are usually often not tolerable when the sample supply is limited, e.g., in clinical proteomics. Another important issue for targeted proteomics is the choice of technology, which is necessarily a function of the aims of the study. 2-D gel-based technologies are the best choice when quantitative changes and/or changes in
post-translational modifications are important in the design of the study. As mentioned above, this is seldom the case in targeted proteomics studies. Moreover, these technologies usually involve a high rate of sample consumption and a strong bias against some classes of proteins, such as membrane proteins. Conversely, peptide separation–based approaches (e.g., MudPIT; UNIT 23.1; Washburn et al., 2001) do not have the same drawbacks as 2-D gels and are therefore highly complementary to 2-D gels. Their advantages in targeted proteomics are linked to their low sample consumption (Pflieger et al., 2002) and to the fact that they do not have the same biases against classes of proteins. Membrane proteins present in the sample are therefore more easily analyzed (Pflieger et al., 2002). These positive features are shared both by methods relying only on peptide separation (Washburn et al., 2001) and by methods using a first step of protein separation by SDS electrophoresis and then a peptide-separation step (e.g., Rout et al., 2000; Bell et al., 2001; Pflieger et al., 2002).
Anticipated Results One type of information that can be gained from targeted proteomics studies is identification of the components present in a sample of interest. Recent examples can be found in organelle proteomics studies of mitochondria (Taylor et al., 2002; Lescuyer et al., 2003), Golgi (Bell et al., 2001), and the peroxisome (Yi et al., 2002). In terms of nonorganelle subcellular components or cellular machines, examples include the nuclear pore (Rout et al., 2000) and the nucleolus (Andersen et al., 2002; Scherl et al., 2002). These studies highlight the fact that the proteomics aspect is not necessarily the most difficult part of the analysis. Validation of the data, i.e., the confirmation of the assignment made to the organelle via the proteomics approach, is much more difficult and time consuming (Rout et al., 2000). In some instances, the targeted proteomics approach can identify unexpected but verified localizations of proteins. Such discoveries can lead to more cell physiology–oriented studies, as exemplified in Gagnon et al. (2002). The second-stage type of targeted proteomics lies in studies of protein dynamics, in which changes in protein levels within an organelle proteome that occur during various physiological conditions are investigated. Examples can be found of investigations using this approach even before the word proteomics existed (Howe and Solter, 1979; Rabilloud
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et al., 1991). These approaches are, however, somewhat difficult to use because of sampleconsumption issues. As a matter of fact, concentrating the proteomics approach on a single cell compartment wastes most of the cellular proteins. This, in turn, limits this type of approach to biological materials that can be obtained repeatedly in gram quantities. In conclusion, although the choice of the most appropriate proteomics approach for targeted proteomics is important, it must not mask the fact that the real bottleneck in the field is clearly the reproducible preparation of pure subcellular structures.
Time Considerations
Lescuyer, P., Strub, J.M., Luche, S., Diemer, H., Martinez, P., Van Dorsselaer, A., Lunardi, J., and Rabilloud, T. 2003. Progress in the definition of a reference human mitochondrial proteome. Proteomics 3:157-167. Pflieger, D., Le Caer, J.P., Lemaire, C., Bernard, B.A., Dujardin, G., and Rossier, J. 2002. Systematic identification of mitochondrial proteins by LC-MS/MS. Anal. Chem. 74:2400-2406. Rabilloud, T., Pennetier, J.L., Hibner, U., Vincens, P., Tarroux, P., and Rougeon, F. 1991. Stage transitions in B-lymphocyte differentiation correlate with limited variations in nuclear proteins. Proc. Natl. Acad. Sci. U.S.A. 88:1830-1834.
This section is devoted only to the organelle preparation steps, as the time considerations for each type of proteomic analysis are adequately described elsewhere (e.g., UNITS 22.1, 22.4, 23.1, & 23.3). Simple nuclei isolation, as described in Basic Protocol 2, yields pure nuclei in a time frame of 2 hr starting from the initial cell pellet.
Rabilloud, T., Kieffer, S., Procaccio, V., Louwagie, M., Courchesne, P.L, Patterson, S.D., Martinez, P., Garin, J., and Lunardi, J. 1998. Twodimensional electrophoresis of human placental mitochondria and protein identification by mass spectrometry: Toward a human mitochondrial proteome. Electrophoresis 19:1006-1014.
Literature Cited
Scherl, A., Coute, Y., Deon, C., Calle, A., Kindbeiter, K., Sanchez, J.C., Greco, A., Hochstrasser, D., and Diaz, J.J. 2002. Functional proteomic analysis of human nucleolus. Mol. Biol. Cell 13:4100-4109.
Andersen, J.S., Lyon, C.E., Fox, A.H., Leung, A.K., Lam, Y.W., Steen, H., Mann, M., and Lamond, A.I. 2002. Directed proteomic analysis of the human nucleolus. Curr. Biol. 12:1-11. Bell, A.W., Ward, M.A., Blackstock, W.P., Freeman, H.N., Choudhary, J.S., Lewis, A.P., Chotai, D., Fazel, A., Gushue, J.N., Paiement, J., Palcy, S., Chevet, E., Lafreniere-Roula, M., Solari, R., Thomas, D.Y., Rowley, A., and Bergeron, J.J. 2001. Proteomics characterization of abundant Golgi membrane proteins. J. Biol. Chem. 276:5152-5165. Fountoulakis, M., Langen, H., Evers, S., Gray, C., and Takacs, B. 1997. Two-dimensional map of Haemophilus influenzae following protein enrichment by heparin chromatography. Electrophoresis 18:1193-1202. Gagnon, E., Duclos, S., Rondeau, C., Chevet, E., Cameron, P.H., Steele-Mortimer, O., Paiement, J., Bergeron, J.J., and Desjardins, M. 2002. Endoplasmic reticulum–mediated phagocytosis is a mechanism of entry into macrophages. Cell 110:119-131. Gygi, S.P., Corthals, G.L., Zhang, Y., Rochon, Y., and Aebersold, R. 2000. Evaluation of two-dimensional gel electrophoresis-based proteome analysis technology. Proc. Natl. Acad. Sci. U.S.A. 97:9390-9395.
Organelle Proteomics
Jung, E., Heller, M., Sanchez, J.C., and Hochstrasser, D.F. 2000. Proteomics meets cell biology: The establishment of subcellular proteomes. Electrophoresis 21:3369-3377.
Howe, C.C. and Solter, D. 1979. Cytoplasmic and nuclear protein synthesis in preimplantation mouse embryos. J. Embryol. Exp. Morphol. 52:209-225.
Rout, M.P., Aitchison, J.D., Suprapto, A., Hjertaas, K., Zhao, Y., and Chait, B.T. 2000. The yeast nuclear pore complex: Composition, architecture, and transport mechanism. J. Cell Biol. 148:635-651.
Taylor, S.W., Warnock, D.E., Glenn, G.M., Zhang, B., Fahy, E., Gaucher, S.P., Capaldi, R.A., Gibson, B.W., and Ghosh, S.S. 2002. An alternative strategy to determine the mitochondrial proteome using sucrose gradient fractionation and 1D PAGE on highly purified human heart mitochondria. J. Proteome Res. 1:451-458. Wilkins, M.R., Gasteiger, E., Sanchez, J.C., Bairoch, A., and Hochstrasser, D.F. 1998. Twodimensional gel electrophoresis for proteome projects: The effects of protein hydrophobicity and copy number. Electrophoresis 19:15011505. Washburn, M.P., Wolters, D., and Yates, J.R. 3rd. 2001. Large-scale analysis of the yeast proteome by multidimensional protein identification technology. Nat. Biotechnol. 19:242-247. Wu, C.C., MacCoss, M.J., Howell, K.E., and Yates, J.R. 2003. A method for the comprehensive proteomic analysis of membrane proteins. Nat. Biotechnol. 21:532-538. Yi, E.C., Marelli, M., Lee, H., Purvine, S.O., Aebersold, R., Aitchison, J.D., and Goodlett, D.R. 2002. Approaching complete peroxisome characterization by gas-phase fractionation. Electrophoresis 23:3205-3216. Zischka, H., Weber, G., Weber, P.J., Posch, A., Braun, R.J., Buhringer, D., Schneider, U.,
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Nissum, M., Meitinger, T., Ueffing, M., and Eckerskorn, C. 2003. Improved proteome analysis of Saccharomyces cerevisiae mitochondria by freeflow electrophoresis. Proteomics 3:906-916. Zuo, X. and Speicher, D.W. 2000. A method for global analysis of complex proteomes using sample prefractionation by solution isoelectrofocusing prior to two-dimensional electrophoresis. Anal. Biochem. 284: 266-278.
Contributed by Pierre Lescuyer, Mireille Chevallet, Sylvie Luche, and Thierry Rabilloud CEA Grenoble Grenoble, France
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Glycoproteomics Using Chemical Immobilization
UNIT 24.3
Quantitative analysis of glycoproteins is a powerful approach to profiling extracellular proteins, including transmembrane proteins, cell surface proteins, and secreted proteins. These methods are used to identify disease-associated proteins and cell-surface antigens from body fluids or tissues for diagnostic and therapeutic purposes. The methods described in this unit allow the analysis of transmembrane proteins, cell surface proteins, and secreted proteins from cells, cell culture medium, body fluids, and tissues. This unit describes methods for solid-phase extraction of glycopeptides and subsequent identification of glycopeptides as well as their glycosylation sites. The Basic Protocol is based on the digestion of glycoproteins to peptides, conjugation of glycopeptides to a solid support using hydrazide chemistry, and specific release of formerly glycosylated peptides using enzymatic or chemical methods (see Fig. 24.3.1A). The recovered peptides are then identified by tandem mass spectrometry. An Alternate Protocol is also described, and it is based on the coupling of intact glycoproteins first to a solid support using hydrazide chemistry, removal of nonglycosylated peptides with protease cleavage, and specific release of formerly glycosylated peptides using enzymatic or chemical methods (see Fig. 24.3.1B). The difference between the two approaches lies in the order in which the glycopeptides are captured to the solid support. In the first approach, proteins from a biological protein mixture are first digested to peptides. Glycopeptides are then oxidized and captured on a hydrazide resin. Nonglycosylated peptides are washed away while the covalently linked glycosylated peptides remain on the solid support. In the second approach, the glycoproteins from a biological protein mixture are first captured to hydrazide resin. After the nonglycosylated proteins are washed away, the immobilized glycoproteins are proteolyzed on the solid support. Nonglycosylated peptides are then washed away while the glycosylated peptides remain on the solid support. In both approaches the formerly N-linked glycosylated peptides are finally released from the solid phase using peptideN-glycosidase F (PNGaseF). Considerations in deciding which of the two approaches might be most appropriate are discussed in the introduction to the Alternate Protocol. A chemical method (e.g., β-elimination) may be used for the release of the formerly O-linked glycosylated peptides (e.g., Greis et al., 1996). Support protocols describe preparation of protein samples from cultured cells and their medium (Support Protocols 1 and 2), body fluids (Support Protocol 3), and tissues (Support Protocol 4) prior to glycopeptide analyses. Support Protocol 5 gives methods for determining the efficiency of tryptic digestion or coupling using SDS-PAGE and silver staining. Support Protocol 6 describes the use of a reverse-phase column for desalting peptides, an essential step in the coupling reactions, and for purifying resin-released O-linked glycosylated peptides. NOTE: Use Milli-Q-purified water or equivalent in all solutions and protocol steps unless otherwise indicated.
SOLID-PHASE EXTRACTION OF GLYCOPEPTIDES USING HYDRAZIDE RESIN The Basic Protocol describes one of two variant protocols for the isolation and identification of N-linked glycopeptides in complex biological samples. In this approach, proteins from a biological protein mixture are first digested to peptides. Glycopeptides Contributed by Hui Zhang Current Protocols in Protein Science (2007) 24.3.1-24.3.14 C 2007 by John Wiley & Sons, Inc. Copyright
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Figure 24.3.1 Schematic diagram of quantitative analysis of glycopeptides using (A) the Basic Protocol for glycopeptide capture or (B) the Alternate Protocol for glycoprotein capture.
are then oxidized and covalently captured on a hydrazide resin. Nonglycosylated peptides are washed away while the covalently linked glycosylated peptides remain on the solid support. (see Fig. 24.3.1A). At the glycopeptide-resin coupling step, both N-linked and O-linked glycopeptides are conjugated (Zhang et al., 2003). After removing nonglycosylated peptides, the glycopeptides are then released from the solid support. The formerly N-linked glycosylated peptides are released from the solid phase using peptide-N-glycosidase F (PNGaseF). PNGaseF treatment also results in conversion of the glycosylated asparagine to aspartic acid, thus generating a one unit mass shift at the formerly glycosylated asparagine residues that is detectable in a highresolution mass spectrometer and is diagnostic for the glycosylation site. Therefore, in a single analysis, the method identifies N-linked glycosylated peptides, the site(s) of Nlinked glycosylation, and the relative quantity of the identified glycopeptides. There are no equivalent glycosidases that release O-linked oligosaccharides in the way that PNGaseF specifically releases N-linked oligosaccharides from the solid support. To successfully release O-linked oligosaccharides, a chemical method, such as β-elimination, may be used for the release of the formerly O-linked glycosylated peptides (e.g., see Greis et al., 1996).
Materials
Glycoproteomics Using Chemical Immobilization
1 mg protein sample (Support Protocols 1 to 4) Lysis buffer: 50 mM KHPO4 , pH 8 (APPENDIX 2E)/0.1% RapiGest (Waters): prepare fresh Trifluoroethanol (TFE) 50 mM tris (2-carboxyethyl) phosphine (TCEP; Pierce): store up to 3 months at −20◦ C 250 mM iodoacetamide: prepare fresh 50 mM and 100 mM ammonium bicarbonate (NH4 HCO3 ) buffer, pH 7.8: prepare fresh 0.5 mg trypsin/ml 50 mM acetic acid: translation grade trypsin (Promega) supplied with 50 mM acetic acid Coupling buffer: 100 mM sodium acetate/150 mM NaCl, pH 5.5; store up to 3 months at room temperature 50 mM sodium periodate: prepare fresh and hold in the dark
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Hydrazide resin (Bio-Rad) 80% acetonitrile/0.1% trifluoroacetic acid (TFA) 1.5 M NaCl Peptide-N-glycosidase F (PNGase F; NEB) 100 mM NaOH 0.3 M and 0.4% (v/v) acetic acid: store up to 3 months at room temperature HPLC solvent A: 0.4% acetic acid/0.005% heptafluorobutyric acid (HFBA); store up to 3 months at room temperature HPLC solvent B: acetonitrile, 0.4% acetic acid/0.005% HFBA; store up to 3 months at room temperature 60◦ C water bath Sonicator (e.g., Sonicator 3000; Misonix) Test tube rocker 1-ml glass vial with polyethylene snap cap (e.g., Waters), three for each sample SpeedVac concentrator Four-port union (Upchurch Scientific) constructed entirely of polyetheretherketone (PEEK; Yi et al., 2003) connecting to: Microcapillary HPLC column (10 cm × 75 µm i.d.) packed with MAGIC C18 resin (Michrom BioResources) or equivalent Peptide trap cartridge packed with MAGIC C18 resin (Michrom BioResources) High-voltage ESI emitter Waste line FAMOS autosampler (Dionex) or equivalent HP1100 solvent delivery system (Hewlett-Packard) or equivalent LCQ ion trap mass spectrometer (Thermo Finnigan) or equivalent SEQUEST software (Eng et al., 1994) Protein database appropriate to the samples of interest PeptideProphet (Proteome Software, http://www.proteomesoftware.com) Additional reagents and equipment for performing SDS-PAGE (Support Protocol 5 and UNIT 10.1) and desalting and purifying peptides (Support Protocol 6) Digest glycoproteins 1. Dissolve 1 mg protein in 100 µl lysis buffer in a 1.5-ml microcentrifuge tube. 2. To denature the proteins add 100 µl TFE to sample, close the tube, and incubate 1 hr at 60◦ C. 3. To reduce the proteins, sonicate 2 min at setting 5 in an ice-water bath, add 20 µl of 50 mM TCEP, and incubate 30 min at 60◦ C. 4. To alkylate the proteins, add 20 µl of 250 mM iodoacetamide and incubate 30 min at room temperature. 5. Dilute the sample by adding 1 ml of 50 mM ammonium bicarbonate buffer, pH 7.8 to reduce the TFE concentration below 10%. 6. Remove 5 µg of undigested proteins for analysis by SDS-PAGE (Support Protocol 5). 7. Add sequencing grade trypsin in a 1:50 (w/w) ratio of trypsin:protein to the sample and incubate 4 hr at 37◦ C, with gentle shaking on a tube rocker. 8. Remove 5 µg of digested peptides and analyze by SDS-PAGE (Support Protocol 5) along with the sample retained in step 6 to determine completeness of tryptic digestion prior to coupling of glycopeptides to the hydrazide resin. 9. Desalt digested peptides using a reverse-phase column (Support Protocol 6), and dry the peptides in a 1-ml glass vial using a SpeedVac.
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Couple glycopeptides to hydrazine resin 10. Dissolve peptides in 80 µl coupling buffer, and add 20 µl of 50 mM freshly prepared sodium periodate to oxidize the glycopeptides. 11. Mix well and gently shake the suspension using a tube rocker 60 min at 4◦ C in the dark. 12. While the peptides are oxidizing, prepare 50 µl per sample of 50% (v/v) hydrazide resin slurry in water. a. Wash the hydrazide resin with 10 column volumes of water. b. Centrifuge the resin 5 min at 3,000 × g, 20◦ C and discard the supernatant. c. Repeat the wash one more time and remove the water. 13. Desalt the oxidized peptides from step 10 using a reverse-phase column (see Support Protocol 6) and elute the peptides with 1 ml of 80% acetonitrile/0.1% TFA. Do not dry the peptides. 14. Add the desalted peptides (in the 80% acetonitrile/0.1% TFA) directly to the prepared hydrazide resin. Mix well and gently shake the suspension using a tube rocker for at least 4 hr at room temperature. 15. Remove the uncoupled peptides by washing the resin three times with 700 µl of each of the following solutions:
1.5 M NaCl H2 O 100 mM ammonium bicarbonate buffer. Remove the supernatant by centrifuging 5 min at 3,000 × g, 20◦ C after each wash step. 16. Resuspend the resin in 100 µl freshly prepared 100 mM ammonium bicarbonate buffer.
Release N-linked glycopeptides from resin 17. Release N-linked glycopeptides from the resin by adding 3 µl PNGase F and incubating overnight at 37◦ C with mixing. 18. Centrifuge the resin 5 min at 1000 × g, 20◦ C and transfer the supernatant to a glass vial. 19. Wash the hydrazide resin twice by adding 200 µl of 100 mM ammonium bicarbonate buffer and centrifuging 5 min at 3,000 × g, 20◦ C. Combine the washes with the supernatant in the glass vial.
Optional: For analysis of O-linked glycopeptides 20. Release O-linked glycopeptides from the resin by incubating hydrazide resin in 100 µl of 100 mM NaOH 4 hr at 45◦ C. 21. Terminate the reaction by adding 100 µl of 0.3 M acetic acid on ice (Greis et al., 1996). 22. Purify the released glycopeptides using a reverse-phase column (see Support Protocol 6). Glycoproteomics Using Chemical Immobilization
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23. Dry in a 1-ml glass vial using a SpeedVac. 24. Resuspend peptides in 10 µl of 0.4% acetic acid for mass spectrometry analysis (2 µl for each analysis). Current Protocols in Protein Science
Figure 24.3.2 Set-up of a four-port union. Port 1: a µ-LC pulled tip reverse-phase column; port 2: a waste line; port 3: a C18 peptide trap cartridge; port 4: an ESI (electrospray ionization) line.
Separate and analyze peptides by tandem mass spectrometry 25. Separate the peptides by online reverse-phase HPLC using HPLC solvents A and B. Connect the microcapillary C18 column, the C18 peptide trap cartridge, µ-ESIemitter/µ-LC pulled tip column combination, high voltage ESI line, and waste line to separate ports of a four-port union . See Figure 24.3.2 for a schematic illustration of setup and UNIT 8.7 for details on the chromatography system.
26. Equilibrate the trap cartridge and microcapillary column 15 min with solvent B. 27. Inject the formerly glycosylated peptides into the C18 peptide trap cartridge using a FAMOS autosampler. The peptides are then passed through a 10 cm × 75 µm i.d. microcapillary C18 column.
28. Apply a linear gradient from 5% to 32% solvent B (acetonitrile) over 100 min at a flow rate of ∼300 nl/min for reverse-phase liquid chromatography using an HP1100 solvent delivery system. The eluting peptides are directly ionized by electrospray ionization and detected by the mass spectrometer. Specific peptide ions are automatically selected and fragmented by collision-induced dissociation (CID) to generate a tandem mass spectrum (MS/MS spectrum or CID mass spectrum).
29. Operate the mass spectrometer in a data-dependent MS/MS mode. In this mode a full-scan mass spectrum is followed by one or more tandem mass spectra, and the precursor ion is selected “on the fly” from the previous scan. An m/z ratio for an ion that has been selected for fragmentation is placed in a list and dynamically excluded for 3 min from further fragmentation. See UNITS 16.8 & 16.10 for detailed instructions.
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30. Query acquired MS/MS spectra against an appropriate protein database using SEQUEST software. Set the database search parameters to the following modifications: carboxyaminomethylated cysteines as static modifications, oxidized methionines and a conversion of Asn to Asp as dynamic modifications. The search parameters indicate that the release of the peptides is PNGase F-catalyzed and occurs at the original site of carbohydrate attachment to the peptide/protein (i.e., the N-glycosite) as dynamic modifications. No other constraints are included for database searches.
31. Statistically analyze database search results using PeptideProphet to effectively compute a probability for the likelihood of each identification being correct (on a scale of 0 to 1) in a data-dependent fashion (Keller et al., 2002). A PeptideProphet probability score of ≥0.9 is used as a filter to remove low probability peptide identifications. ALTERNATE PROTOCOL
SOLID-PHASE EXTRACTION OF GLYCOPROTEINS USING HYDRAZIDE RESIN To analyze glycoproteins rather than glycosylated peptides only, capture can be performed at the protein rather than the peptide level. After covalent capture of glycoproteins onto the hydrazide resin, a protease digestion step releases nonglycopeptides, which are then optionally collected for analysis via mass spectrometry. This approach will increase the number of unique peptides identified from each protein and therefore increase the confidence of protein identifications. However, for proteins with multiple glycosylation sites, solid-phase capture of glycoproteins may not immobilize all the glycosylation sites to the solid support because of steric effects. In this case, the nonimmobilized glycosylation sites from glycoproteins are released from the solid support after protease digestion, which may decrease the efficiency and sensitivity of the analysis.
Additional Materials (also see Basic Protocol) 80% acetonitrile 100% methanol Size-exclusion column, two for each sample: SEC Macro spin column, P-6 (>6 kDa) gel filtration (The Nest Group) 1. Equilibrate two size-exclusion columns for each sample with 200 µl coupling buffer for 15 min and remove buffer by centrifuging 5 min at 2,500 × g, 20◦ C. Repeat this procedure two more times. For buffer exchange of proteins, a size-exclusion column is used instead of the Sep-Pak column used for peptides in Support Protocol 6.
Prepare proteins and resin 2. Add 1 mg protein sample in 80 µl total volume to one of the equilibrated sizeexclusion columns to exchange proteins to coupling buffer. 3. Collect the protein samples into a new collection tube by centrifuging 5 min at 2500 × g, 20◦ C. 4. To oxidize glycoproteins add 20 µl of 50 mM sodium periodate, mix well and incubate 60 min at 4◦ C in the dark, with gentle shaking using a tube rocker. 5. While the proteins are incubating, prepare 50 µl per sample of 50% (v/v) hydrazide resin slurry in water. Glycoproteomics Using Chemical Immobilization
a. Wash hydrazide resin with 10 column volumes of water. Centrifuge the resin 5 min at 3000 × g, 20◦ C and discard the water.
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b. Equilibrate hydrazide resin with 10 column volumes of coupling buffer. Centrifuge the resin 5 min at 3000 × g, 20◦ C and discard the coupling buffer. c. Resuspend the hydrazide resin with 25 µl of coupling buffer to make a 50% slurry. 6. Add the oxidized protein sample from step 3 to the second prepared size-exclusion column to remove excess sodium periodate by exchanging proteins to coupling buffer. 7. Collect the protein samples into a new collection tube by centrifuging 5 min at 2,500 × g. Retain 5 µg of the protein before adding the hydrazide resin to analyze by SDS-PAGE along with the uncoupled proteins from step 9.
Couple proteins to resin 8. Add hydrazide resin equilibrated in coupling buffer to the sample. Mix well and incubate 4 hr at room temperature, with gentle shaking using a tube rocker. 9. Centrifuge the resin 5 min at 1000 × g, 20◦ C and save the supernatant. Retain a volume of the supernatant equivalent to the volume of protein solution retained in step 7 to analyze by SDS-PAGE. Proteins remaining in the supernatant can be analyzed by SDS-PAGE and compared with the precoupling protein profile of the sample retained in step 7 (see Support Protocol 5 for details).
10. Wash the protein-coupled resin three times by adding 1 ml lysis buffer. Centrifuge 5 min at 3,000 × g, 20◦ C and remove lysis buffer after each wash. 11. Resuspend the protein-coupled resin with 50 µl of lysis buffer.
Remove unglycosylated protein 12. To denature the proteins add 100 µl of TFE to sample and incubate at 60◦ C for 1 hr. 13. To reduce the proteins add 20 µl of 50 mM TCEP and incubate 30 min at 60◦ C. 14. To alkylate the proteins add 20 µl of 250 mM iodoacetamide and incubate 30 min at room temperature. 15. Dilute the sample by adding 1 ml of 50 mM ammonium bicarbonate buffer, pH 7.8 to reduce the TFE concentration below 10%. 16. Add trypsin solution to sample and incubate 4 hr at 37◦ C, with gentle shaking. 17. Centrifuge the hydrazide resin 5 min at 1000 × g, 20◦ C. Save the supernatant containing nonglycosylated peptides. This fraction of peptides can be purified using reverse-phase columns (see Support Protocol 6) and dried in 1 ml glass vial with a SpeedVac for analysis by mass spectrometry for identification of additional peptides other than glycopeptides from glycoproteins (follow Basic Protocol from step 24).
18. Wash the glycopeptides on the hydrazide resin three times each with 700 µl of the following solutions:
1.5 M NaCl 80% acetonitrile 100% methanol H2 O 100 mM ammonium bicarbonate buffer. Remove supernatant by centrifuging 5 min at 3,000 × g, 20◦ C after each wash step. 19. Continue with steps 16 to 31 in the Basic Protocol to release and analyze the glycopeptides. Current Protocols in Protein Science
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SUPPORT PROTOCOL 1
EXTRACTION OF GLYCOPROTEINS FROM CULTURED CELLS Proteins from cells are extracted prior to the capture of glycopeptides (Basic Protocol) or glycoproteins (Alternate Protocol). NOTE: All solutions and equipment coming into contact with cells must be sterile, and proper aseptic technique should be used accordingly.
Materials Cells of interest, in culture Cell culture medium containing serum and growth factors (e.g., DMEM with 10% FBS; see APPENDIX 3C) Cell culture medium without serum Lysis buffer: 50 mM KHPO4 , pH 8 (APPENDIX 2E)/ 0.1% RapiGest (Waters); prepare fresh Humidified, 5% CO2 , 37◦ C cell incubator Sonicator (e.g., Sonicator 3000; Misonix) Additional reagents and equipment for determining cell number (APPENDIX 3C) 1. Culture cells in medium supplemented with serum or growth factors. Grow cells to confluence in a humidified incubator with 5% CO2 at 37◦ C. 2a. To wash adherent cells: Discard the original medium, add 20 ml serum-free medium, swirl, and discard the medium. Repeat four more times. 2b. To wash cells in suspension: Centrifuge the suspension 5 min at 1000 × g, 20◦ C and discard the medium. Resuspend the cells in 20 ml serum-free medium, centrifuge, and discard medium. Repeat four more times. 3. Harvest adherent cells by scraping and add enough lysis buffer to result in 108 cells/ml, or add lysis buffer to the pellet of washed cells (from the suspension culture) to result in 108 cells/ml. 4. Sonicate cells 30 sec at setting 5 in an ice-water bath. Repeat sonication three times. 5. Centrifuge sonicated cell lysates 10 min at 10,000 × g, 4◦ C. 6. Remove supernatant and use for glycopeptide (Basic Protocol) or glycoprotein (Alternate Protocol) capture. Store samples up to 3 months at −80◦ C before using for capture protocols. SUPPORT PROTOCOL 2
COLLECTION AND CONCENTRATION OF SECRETED GLYCOPROTEINS FROM CELL CULTURE MEDIUM Proteins secreted from cultured cells are extracted from serum-free cell culture medium to avoid contamination by serum proteins that normally supplement culture medium. Secreted proteins are generally too dilute and must be concentrated from cell culture medium prior to the capture of glycoproteins or glycopeptides. NOTE: All solutions and equipment coming into contact with cells must be sterile, and proper aseptic technique should be used accordingly.
Materials
Glycoproteomics Using Chemical Immobilization
Cells of interest, in culture Cell culture medium containing serum and growth factors (e.g., DMEM with 10% FBS; see APPENDIX 3C) Cell culture medium without serum Lysis buffer: 50 mM KHPO4 , pH 8 (APPENDIX 2E)/ 0.1% RapiGest (Waters); prepare fresh
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Humidified, 5% CO2 , 37◦ C cell incubator Centrifugal filter devices (e.g., CentriprepYM10; Millipore) 1. Culture cells in medium supplemented with serum or growth factors. Grow cells to confluence in a humidified incubator with 5% CO2 at 37◦ C. 2. Wash cells five times with serum-free medium as in Support Protocol 1, step 2a or 2b. 3. Incubate cells in a serum-free medium for the required time. The incubation time required for protein secretion into the medium varies from ∼1 hr to overnight, depending on the cell type. If an experimental treatment of cells is needed prior to collection of secreted proteins, the treatment can be done at the time of switching to a serum-free medium.
4. Collect cell culture medium by centrifuging 10 min at 1,500 × g, 20◦ C to remove any cells or cell debris. 5. Concentrate proteins in the supernatant to 10 mg/ml using centrifugal filter devices according to the manufacturer’s directions. 6. Store samples up to 3 months at −80◦ C before using the Basic Protocol or Alternate Protocol to capture glycopeptides or glycoproteins, respectively.
COLLECTION AND CONCENTRATION OF GLYCOPROTEINS FROM PLASMA AND OTHER BODY FLUIDS
SUPPORT PROTOCOL 3
Proteins from body fluids are present in varying concentrations; therefore, adjustments of proteins to certain concentrations (∼10 mg/ml) are usually performed prior to performing the Basic or Alternate Protocol. In general, proteins from plasma are diluted with PBS to 10 mg/ml. Proteins from most other body fluids (e.g., cerebrospinal fluid, urine, breast milk, saliva, lung lavage fluid, and pancreatic juice) need to be concentrated to 10 mg/ml using centrifugal filter devices (e.g., Centriprep YM10; Millipore). Protein samples can also be desalted after concentration by diluting the concentrated proteins with coupling buffer, and repeating the concentration procedure using the centrifugal filter devices three times.
EXTRACTION OF GLYCOPROTEINS FROM SOLID TISSUES Proteins from solid tissues are first extracted mechanically and subsequently digested to peptides prior to the capture of glycopeptides using the Basic Protocol.
SUPPORT PROTOCOL 4
Materials 100 mg frozen tissue Lysis buffer Razor blade Sonicator (e.g., Sonicator 3000; Misonix) 1. Slice frozen tissue into 1 to 3 mm3 thick pieces and incubate 2 to 3 min in 1 ml lysis buffer with periodic vortexing. 2. Sonicate the sample 1 min at setting 5 in an ice-water bath five times, waiting 1 min each time between sonication to prevent overheating. 3. Clear sonicated cell lysates by centrifuging 10 min at 10,000 × g, 4◦ C. 4. Remove supernatant and use for capture protocols. Store samples up to 3 months at −80◦ C until used for capture protocols.
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SUPPORT PROTOCOL 5
DETERMINATION OF EFFICIENCY OF GLYCOPROTEIN TRYPTIC DIGESTION OR GLYCOPEPTIDE/GLYCOPROTEIN-HYDRAZIDE RESIN COUPLING USING SDS-PAGE AND SILVER STAINING The tryptic digestion of proteins or the coupling of glycoproteins to hydrazide resin is monitored by SDS-PAGE to check the two critical steps of the solid-phase extraction of glycopeptides procedure of the Basic Protocol. Aliquots of proteins/peptides are removed from the sample both before and after tryptic digestion or before and after coupling to the hydrazide resin. The expected result for efficient tryptic digestion is that protein bands at high molecular weight essentially disappear upon trypsin digestion, while the expected result for efficient coupling of glycoproteins to hydrazide resin is that some protein bands disappear after coupling.
Materials 5 µg protein/peptide samples (Basic Protocol, steps 6 and 8; Alternate Protocol, steps 7 and 9) 3× sample loading buffer (Cell Signaling Technology) 50% methanol/10% acetic acid solution Silver stain kit (Bio-Rad) including: Silver stain sensitizing solution Staining solution Developing solution Stopping solution Gel dryer Flatbed scanner Additional reagents and equipment for SDS-PAGE (UNIT 10.1) Prepare proteins/peptides and perform SDS-PAGE 1. Mix protein/peptide samples with sufficient 3× sample loading buffer to yield a 1× concentration of sample loading buffer. 2. Denature proteins by heating the samples 5 min at 95◦ C. Cool on ice. 3. Microcentrifuge samples 5 min at maximum speed. 4. Load proteins onto an SDS-polyacrylamide gel and electrophorese 1 hr at 200 V, following the specific instructions provided with the electrophoresis system (also see UNIT 10.1).
Fix and stain gel 5. After electrophoresis, fix the gel by agitating 30 min in a 50% methanol, 10% acetic acid solution. 6. Wash the gel in five changes of deionized water, 20 min each wash. 7. Incubate the gel 30 min in silver stain sensitizing solution. 8. Following incubation, wash the gel in three changes of deionized water, 30 sec each wash. 9. Incubate the gel 20 min in staining solution. 10. After incubation, wash the gel in three changes of deionized water, 60 sec each wash. 11. Incubate the gel up to 10 min in developing solution. Glycoproteomics Using Chemical Immobilization
Proteins are visualized as dark brown bands, and longer incubation times will increase the background.
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12. Stop the development by incubating at least 10 min with stopping solution. 13. Wash the gel in deionized water. 14. Dry the gel on a gel drier and scan/photograph the gel. The expected result for efficient tryptic digestion is that protein bands at high molecular weight essentially disappear upon trypsin digestion, i.e., most silver stained bands >5 kDa present before digestion disappear after the digestion. The expected result for efficient coupling of glycoproteins to hydrazide resin is that some protein bands disappear after coupling, i.e., some silver stained bands are present before and absent after the coupling.
DESALTING AND PURIFYING PEPTIDES USING A REVERSE-PHASE COLUMN
SUPPORT PROTOCOL 6
Peptides can be purified using a reverse-phase column prior to mass spectrometry analysis to remove any contamination that might interfere. The purified peptides can also be resuspended in a different buffer for the purposes of desalting or buffer exchange for different reaction conditions.
Materials 80% acetonitrile/0.1% trifluoroacetic acid (TFA) 0.1% and 1% TFA Peptide samples (Basic Protocol, steps 7, 12, and 21; Alternate Protocol, step 17) 1-cc reverse-phase columns (C18, Sep-Pak; Waters), four columns for each sample 1. Clean reverse-phase column by adding 1 ml of 80% acetonitrile/0.1% TFA and drain the solution by gravity. Repeat this procedure one more time. 2. Equilibrate the reverse-phase column with 1 ml of 0.1% TFA. Drain the solution by gravity. Repeat this procedure one more time. 3. Adjust the pH of peptide sample to 3 using 1% TFA. 4. Load peptide solution onto the equilibrated reverse-phase column. 5. Wash the reverse-phase column with 1 ml of 0.1% TFA and drain the solution by gravity. Repeat this procedure one more time. 6. Elute the peptides into a new collection tube with 1 ml of 80% acetonitrile/0.1% TFA.
COMMENTARY Background Information Protein glycosylation plays a significant role in protein sorting and secretion into extracellular environments (Roth, 2002). In addition, the aberrant glycosylation of proteins is a fundamental characteristic of many diseases, including cancer and diabetes. However, the most significant benefits of selective isolation and proteomic analysis of glycoproteins originate from the fact that the number of glycopeptides in the human proteome is relatively modest. Typically, carbohydrates are linked to the side chains of serine or threonine residues (O-linked glycosylation) or to asparagine residues (N-linked glycosylation).
N-linked glycosylation sites generally fall into the N-X-S/T sequence motif in which X denotes any amino acid except proline (Bause, 1983). Because glycosylation only occurs on extracellular domains of membrane proteins, cell surface proteins, and secreted proteins, the total number of tryptic peptides generated from human proteins containing glycosylation sites is modest. Yet, this limited set of peptides represents the majority of membrane, cell surface, and secreted proteins because most of the proteins with domains in the extracellular environment are glycosylated (Zhang et al., 2006b). This results in a very limited number of peptides for each mass detected by mass
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spectrometry, indicating that a defined glycopeptide from the proteome can be assigned to each peptide mass observed if the glycopeptides can be specifically isolated and analyzed. Therefore, it is apparent that a method that can specifically and efficiently isolate glycopeptides would be of significant interest for proteomic analysis of extracellular proteins, including secreted proteins from body fluids. This methodology will significantly increase the throughput and sensitivity for proteomic analysis of extracellular proteins for biomarker discovery or identification of immunotherapy targets. However, after many years of efforts in the field, the Protein Information ResourcesProtein Sequence Database (http://pir. georgetown.edu/pirwww/search/textpsd.shtml) contained only 172 experimentally confirmed human glycoproteins (Kaji et al., 2003) before two new methods for the identification of N-linked glycopeptides in complex biological samples were reported recently (Kaji et al., 2003; Zhang et al., 2003). The difference between the two approaches lies in the mechanisms by which the glycopeptides are captured to solid support. In the first method, glycoproteins or glycopeptides are captured to solid support by chemical immobilization as described in this unit (Zhang et al., 2003). Periodate oxidation converts the cis-diol groups of carbohydrates to aldehydes, which then form covalent hydrazone bonds with hydrazide groups immobilized on the solid support. The second approach is based on a lectin column–mediated affinity capture (Kaji et al., 2003). After immobilization of glycopeptides on a solid support using either method, peptideN-glycosidase is then used to release the Nlinked glycopeptides followed by analysis of mass spectrometry to identify the N-linked glycosylation sites and quantify the relative abundance of glycopeptides. For glycopeptide capture using chemical immobilization, peptides containing either N-linked or O-linked oligosaccharides are covalently conjugated to a solid support. Nonglycosylated peptides can be removed by extensive washing before the release of glycopeptides; therefore, the glycopeptides can be specifically enriched (over 90% enrichment; Zhang et al., 2005). The specific release of different types of glycopeptides from solid support can be achieved by varying glycosidases or cleavage reagents. For glycopeptide capture using lectin affinity columns, glycopeptides containing certain
glycan structures can be selectively enriched by employing different lectin columns. The specificity of glycopeptide capture using a lectin column may not be as high as can be achieved with the chemical immobilization method. This is due to the less efficient removal of nonglycopeptides by the mild washing conditions used with the lectin affinity column. However, different glycans can be selectively analyzed using a lectin column specific to a particular glycan structure. Glycoproteomic analysis using chemical immobilization of glycopeptides has been successfully applied to the analysis of cell surface proteins and for serum proteome profiling (Zhang et al., 2003; Desiere et al., 2005; Deutsch et al., 2005; Li et al., 2005; Pan et al., 2005; Zhang et al., 2005; Zhang et al., 2006a). Interestingly, albumin, the most abundant serum protein, does not appear to contain any N-linked glycosylation sites and is therefore transparent to the method, allowing for more efficient identification and quantification of the lower abundance glycoproteins (Zhang et al., 2003; Zhang et al., 2006a,b; Zhang et al., 2005).
Critical Parameters Glycopeptide capture using chemical immobilization was first introduced in 2003 (Zhang et al., 2003), and it consists of four steps: (1) Glycoprotein oxidation: periodate oxidation converts the cis-diol groups of carbohydrates to aldehydes. (2) Coupling: the aldehydes react with hydrazide groups immobilized on a solid support to form covalent hydrazone bonds. The nonglycosylated proteins are removed by washing. (3) Proteolysis: the immobilized glycoproteins are proteolyzed, and nonglycosylated peptides of glycoproteins are removed by washing. (4) Release: formerly N-linked glycopeptides are released from the solid support by PNGase F treatment. For the oxidation step, there are two parameters to consider. First, glycoproteins or glycopeptides should be devoid of interfering compounds (e.g., compounds containing amino or cis-diol groups). The interfering compounds are generally removed from proteins using desalting columns and from peptides using reverse-phase columns. Second, different glycans can be oxidized at different temperatures and concentrations of periodate; 10 mM sodium periodate and 4◦ C is used to oxidize most oligosaccharides.
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For the coupling step, the ratio of protein to hydrazide resin is critical for optimal specificity and efficiency of glycopeptide capture. A high ratio of protein to hydrazide resin results in incomplete capture of all glycopeptides and therefore low yield. A low ratio of protein to hydrazide resin results in nonspecific binding of nonglycopeptides to the hydrazide resin, or nonspecific binding of glycopeptides released by PNGase F treatment; this will result in low specificity and low recovery of the glycopeptides. The amount of hydrazide resin used in the protocols described in this unit is based on capture of glycopeptides from plasma using commercially available hydrazide resin. Different protein samples will have different levels of glycosylated proteins. For a new type of resin or a protein sample type, determination of the optimal amount of hydrazide resin for coupling is strongly recommended. For the proteolysis step, it is necessary to completely proteolyze proteins to peptides. For this purpose, denaturing proteins effectively facilitates the protease digestion. Removing nonglycosylated peptides from the solid support by intensive washing is a critical step in increasing specificity. For the release step, enzymes or chemicals should be pure and active to effectively release the glycopeptides and avoid the nonspecific release of peptides.
Troubleshooting No glycopeptides identified 1. Incomplete protease digestion: check the saved fraction after digestion and compare it with the protein sample before digestion (see Support Protocol 5). 2. Insufficient coupling: this may be due to interfering compounds (such as compounds containing amino or cis-diol groups) present in the protein sample during the oxidation or coupling step. Low specificity 1. Incomplete protease digestion: check the saved fraction after digestion and compare it with the protein sample before digestion (see Support Protocol 5). 2. Insufficient washing: make sure all the hydrazide resin is completely suspended for each washing step. Low yield 1. Incomplete protease digestion: check the saved fraction after digestion and compare it with the protein sample before digestion (see Support Protocol 5).
2. Insufficient coupling: this may be due to interfering compounds (such as compounds containing amino or cis-diol groups) present in protein samples during oxidation or coupling step. 3. Inefficient releasing of glycopeptides from hydrazide resin: check the activity and purity of enzymes and chemicals. 4. Ratio of protein to hydrazide resin too high or too low: test different ratios.
Anticipated Results
For N-linked glycopeptides, ∼1% of the original protein mass is recovered as formerly N-linked glycosylated peptides. Analysis of 1 µg of recovered peptides using a 3-D ion trap such as an LCQ mass spectrometer generally identifies over 100 unique N-linked glycopeptides representing at least 50 unique glycoproteins in a single analysis. The number of unique N-linked glycopeptides and glycoproteins increases when the samples are analyzed using recently advanced mass spectrometers such as a linear ion trap (LTQ) or LTQ FT. For the identified peptides, over 90% of peptides contain the consensus N-linked glycosylation motif (N-X-T/S motif, X can be any amino acid except P; Zhang et al., 2005).
Time Considerations Both the Basic Protocol and Alternate Protocol require three incubations of 4 hr to overnight for proteolysis, coupling, and releasing. However, less than 3 hr is required between the incubations for measuring samples, mixing, centrifuging, and transferring.
Literature Cited Bause, E. 1983. Structural requirements of Nglycosylation of proteins. Studies with proline peptides as conformational probes. Biochem. J. 209:331-336. Desiere, F., Deutsch, E.W., Nesvizhskii, A.I., Mallick, P., King, N.L., Eng, J.K., Aderem, A., Boyle, R., Brunner, E., Donohoe, S., Fausto, N., Hafen, E., Hood, L., Katze, M.G., Kennedy, K.A., Kregenow, F., Lee, H., Lin, B., Martin, D., Ranish, J.A., Rawlings, D.J., Samelson, L.E., Shiio, Y., Watts, J.D., Wollscheid, B., Wright, M.E., Yan, W., Yang, L., Yi, E.C., Zhang, H., and Aebersold, R. 2005. Integration with the human genome of peptide sequences obtained by high-throughput mass spectrometry. Genome Biol. 6:R9 Deutsch, E.W., Eng, J.K., Zhang, H., King, N.L., Nesvizhskii, A.I., Lin, B., Lee, H., Yi, E.C., Ossola, R., and Aebersold, R. 2005. Human Plasma Peptide Atlas. Proteomics 5:34973500.
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Eng, J., McCormack, A.L., and Yates, J.R. 3rd. 1994. An approach to correlate tandem mass spectral data of peptides with amino acid sequences in a protein database. J. Am. Soc. Mass Spectrom. 5:976-989.
Zhang, H., Li, X.J., Martin, D.B., and Aebersold, R. 2003. Identification and quantification of Nlinked glycoproteins using hydrazide chemistry, stable isotope labeling and mass spectrometry. Nat. Biotechnol. 21:660-666.
Greis, K.D., Hayes, B.K., Comer, F.I., Kirk, M., Barnes, S., Lowary, T.L., and Hart, G.W. 1996. Selective detection and site-analysis of O-GlcNAc-modified glycopeptides by betaelimination and tandem electrospray mass spectrometry. Anal. Biochem. 234:38-49.
Zhang, H., Yi, E.C., Li, X.J., Mallick, P., KellySpratt, K.S., Masselon, C.D., Camp, D.G. 2nd, Smith, R.D., Kemp, C.J., and Aebersold, R. 2005. High throughput quantitative analysis of serum proteins using glycopeptide capture and liquid chromatography mass spectrometry. Mol. Cell Proteomics 4:144-155.
Kaji, H., Saito, H., Yamauchi, Y., Shinkawa, T., Taoka, M., Hirabayashi, J., Kasai, K., Takahashi, N., and Isobe, T. 2003. Lectin affinity capture, isotope-coded tagging and mass spectrometry to identify N-linked glycoproteins. Nat. Biotechnol. 21:667-672. Keller, A., Nesvizhskii, A.I., Kolker, E., and Aebersold, R. 2002. Empirical statistical model to estimate the accuracy of peptide identifications made by MS/MS and database search. Anal. Chem. 74:5383-5392. Li, X.J., Yi, E.C., Kemp, C.J., Zhang, H., and Aebersold, R. 2005. A software suite for the generation and comparison of peptide arrays from sets of data collected by liquid chromatography-mass spectrometry. Mol. Cell Proteomics 4:1328-1340. Pan, S., Zhang, H., Rush, J., Eng, J., Zhang, N., Patterson, D., Comb, M.J., and Aebersold, R. 2005. High throughput proteome screening for biomarker detection. Mol. Cell Proteomics 4:182-190. Roth, J. 2002. Protein N-glycosylation along the secretory pathway: Relationship to organelle topography and function, protein quality control, and cell interactions. Chem. Rev. 102:285-303. Yi, E.C., Lee, H., Aebersold, R., and Goodlett, D.R. 2003. A microcapillary trap cartridgemicrocapillary high-performance liquid chromatography electrospray ionization emitter device capable of peptide tandem mass spectrometry at the attomole level on an ion trap mass spectrometer with automated routine operation. Rapid Commun. Mass Spectrom. 17:20932098.
Zhang, H., Liu, A.Y., Loriaux, P., Wollscheid, B., Zhou, Y., Watts, J., and Aebersold, R. 2006a. Mass spectrometric detection of tissue-derived proteins in blood. Mol. Cell. Proteomics 6:6471. Zhang, H., Loriaux, P., Eng, J., Campbell, D., Keller, A., Moss, P., Bonneau, R., Zhang, N., Zhou, Y., Wollscheid, B., Cooke, K., Yi, E.C., Lee, H., Peskind, E.R., Zhang, J., Smith, R.D., and Aebersold, R. 2006b. UniPep, a database for human N-linked glycosites: A resource for biomarker discovery. Genome Biol. 7:R73.
Key References Zhang et al., 2003. See above Describes a method to identify and quantify Nlinked glycoproteins. It also applies the method to enrich cell surface proteins and efficiently profile serum proteins by removing albumin from serum samples. Kaji et al., 2003. See above Describes mass spectrometry analysis of protein glycosylations using combined techniques of lectin affinity purification and isotope-coded tagging.
Internet Resources http://www.unipep.org UniPep, a database for human N-linked glycosites: a resource for biomarker discovery.
Contributed by Hui Zhang Johns Hopkins University Baltimore, Maryland
Glycoproteomics Using Chemical Immobilization
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CHAPTER 25 Proteome Bioinformatics INTRODUCTION omputational challenges associated with proteomic studies have recently emerged as some of the most critical and limiting factors in this rapidly evolving discipline. The increased importance of computational analyses has resulted in large part from recent improvements in mass spectrometers, which have become dramatically faster and more sensitive. Most proteomic studies rely upon LC-MS/MS analysis of complex peptide mixtures as the last step in the overall proteome analysis, followed by correlation of the resulting MS/MS spectra with protein sequence databases using algorithms such as Sequest (UNIT 16.10). As a result of the increased speed and sensitivity of mass spectrometers, dramatically expanded datasets can be obtained from a single LC-MS/MS run. Furthermore, more extensive fractionations of complex samples are being performed prior to LC-MS/MS in efforts to detect and analyze larger numbers of proteins in complex proteomes and subproteomes (see Chapter 24). These improvements have resulted in major bottlenecks in analyzing, displaying and interpreting proteomic data.
C
When sequence databases and MS/MS datasets were small, data analysis was relatively manageable and it was feasible to manually review all spectra for peptides and proteins that were identified to attempt to validate the accuracy of the computer generated matches. As MS/MS datasets have expanded and proteomes have been fractionated into many subsets that result in many LC-MS/MS analyses for a single proteome, the need for improved computational tools for most stages of the proteome analysis process has become critical. The complexity of many current datasets has rapidly outstripped the feasibility of manually validating data, yet it is vital that false positive identification levels be well defined and maintained at low levels. Otherwise, the proteomics literature and results databases will be rapidly polluted with incorrect and misleading information. In addition to the challenges of distinguishing false positive identifications from true positive identifications and managing large datasets, there are a number of important downstream challenges. One issue is that the most comprehensive sequence databases are incompletely annotated and contain redundant entries for single proteins. It can be quite challenging to eliminate redundant protein identifications from large datasets and simultaneously distinguish which splice forms and closely related members of protein families are actually present in complex samples. Another problem is that not all codingregion single nucleotide polymorphisms (SNPs) are represented in sequence databases. A SNP that is present in the sample analyzed but absent from the database used to interpret the MS/MS data can lead to an incorrect identification for that peptide. Another important challenge is linking biologically relevant observations with knowledge contained in the scientific literature. A major hurdle is the diversity and inconsistency of protein names. A given protein may appear in the scientific literature under a variety of different names, some of which are not necessarily represented in the descriptors in sequence databases and cannot be readily mined using existing databases. Hence, there are challenges at every stage of the proteomics field that are being addressed by improved computational tools, improvements to databases, and development Proteome Bioinformatics Contributed by David W. Speicher Current Protocols in Protein Science (2005) 25.0.1-25.0.2 C 2005 by John Wiley & Sons, Inc. Copyright
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of standards for representing and reporting proteomics data. UNIT 25.1 addresses analysis of protein-protein interactions (the interactome) as well as a number of questions and issues raised above. Future units will address proteomics data standards, tools for analyzing large datasets, strategies for distinguishing true positive from false positive protein identifications, comparisons of alternative search algorithms, and related issues. David W. Speicher
Introduction
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Bioinformatics Analysis for Interactive Proteomics The advent of genome sequencing projects has made it possible to analyze protein-protein interactions on the genome scale. The complete genomes of more than 100 bacteria, as well as several eukaryotes, have been sequenced. The complete genome sequence provides information about all the proteins in the genome, i.e., the proteome. Such information makes possible, for the first time, the characterization of all the protein-protein interactions in an organism, which is referred to as the protein interaction map or "interactome" (Legrain et al., 2001). Several high-throughput technologies have been developed to characterize the protein-protein interaction map, in which life reveals itself not as a mere collection of proteins, but rather as a sophisticated network. In other words, we can see not only the trees but also the forest. Since proteinprotein interaction data can provide information about the biological context in which an individual protein plays its cellular role, the protein-interaction map provides a unique approach to addressing challenges in this postgenome era, especially for understanding roles of many newly discovered genes, the functions of which have not been characterized. For example, only one-third of all 6200 predicted yeast genes were functionally characterized when the complete sequence of the yeast genome became available. At present, 3800 yeast genes have been characterized by genetic or biochemical techniques, and an additional 600 genes have been characterized based on homology with known functions in other organisms. This leaves about 1800 genes with unknown functions (Kumar and Snyder, 2001). Another challenge in the post-genome era is to understand how the proteins encoded in the genome interact with each other. The protein-interaction map can provide essential information in addressing this challenge, especially in helping to understand cellular biological machineries and biological pathways. High-throughput protein-protein interaction data are generated from technology driven experiments, which provide rich information with an ever increasing volume of data. Understanding the biological significance of raw outputs from experiments is becoming the bottleneck in the analysis of high-throughput
UNIT 25.1
protein-protein interaction data. This problem calls upon a number of aspects of bioinformatics: First, databases are needed to store, document, and disseminate increasing amounts of protein-protein interaction data, and visualization tools are needed to display and navigate the interaction network. Second, because heterogeneity in data quality and a high noise level are inherent in the high-throughput nature of the experimental techniques, computational and statistical methods are required for prediction of protein-protein interactions and to handle data quality control with reliability assessments and validation. Finally, new computational tools are in demand for biological discovery and hypotheses validation, based on computational analyses of high-throughput protein-protein interaction data, in conjunction with other biological data. Computational approaches for exploiting protein-protein interaction data will not only provide tools for biologists, but will also result in important scientific insights into cellular mechanisms. This unit addresses technical details related to these three issues.
Experimental Methods for Protein-Protein Interaction Identification There are many experimental methods for identification of protein-protein interactions and characterization of their biological importance. Traditionally, protein-protein interactions have been studied on an individual basis using low-throughput technologies, e.g., immunoprecipitations (Phizicky and Field, 1995) and the glutathione S-transferase (GST) pulldown method (Martzen et al., 1999.) In the high-throughput proteomics approaches, several techniques have been used (Table 25.1.1; Pelletier and Sidhu, 2001). To effectively analyze protein-protein interaction data, it is important to know the source of the data together with the strengths and limitations of the associated experimental techniques. Different experimental methods can identify different types of protein-protein interactions. Some technologies (e.g., yeast twohybrid, protein chip, and phage display) can detect binary interactions, while others (e.g., immunoprecipitation, GST pull-down, and Proteome Bioinformatics
Contributed by Yu Chen and Dong Xu Current Protocols in Protein Science (2005) 25.1.1-25.1.14 C 2005 by John Wiley & Sons, Inc. Copyright
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Table 25.1.1 Current Major Technologies for Studying Protein-Protein Interactions
Method
Experimental condition
Interaction
Throughput
Noise level
Immunoprecipitations (UNIT 9.8)
In vitro
Complex
Low
Low
Pull-down
In vivo
Complex
Low
Low
Yeast two-hybrid (UNIT 19.2)
In vivo
Binary
High
High
Mass spectrometry (Chapter 16)
In vitro
Complex
High
High
Protein chip (UNIT 23.5)
In vitro
Binary
High
High
Phage display (Mullaney and Pallavicini, 2001)
In vitro
Binary
High
High
Figure 25.1.1 (A) Visualization of 8286 protein-protein interactions from the BIND database generated with the Pajek visualization tool (http://vlado.fmf.uni-lj.si/pub/networks/pajek/). (B) Cytoscape (http://www.cytoscape.org/) graphic output for 359 protein-protein interactions in yeast detected by two-hybrid assays.
Bioinformatics Analysis for Interactive Proteomics
mass spectrometry) can identify protein complexes. In a protein interaction graph (e.g., Fig. 25.1.1), a binary interaction can be represented as an edge (line) with the two interacting proteins as nodes (dots). A protein complex can be regarded as a connected graph. However, the topology of the interactions, i.e., which pairs of proteins within a complex physically interact with each other, is unknown. Therefore, exact binary information cannot be obtained about a protein complex. True proteinprotein interactions within a complex may not be detectable in a binary interaction study because of multibody effects (i.e., interaction of two proteins that requires the presence of other proteins to help form a stable complex). One major issue with high-throughput experimental technologies is the generation
of false negatives and false positives. Proteins interact with one another over a wide range of affinities and time scales. Such interactions often occur at the limits of experimental detection, and interactions that are physiologically unimportant can result in experimental noise. Each technique has its own strengths and weaknesses in detecting certain types of interactions, and different techniques result in different noise levels. One should take into account technique biases and limitations when performing computational analyses of the data. For example, most current techniques are not effective in characterizing protein interactions involving integral membrane proteins because these techniques cannot provide the membrane environment, without which membrane proteins unfold. To overcome
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this shortcoming, genetic screening systems have been developed for assaying membrane protein interactions, e.g., the Ras recruitment system (Broder et al., 1998), the G protein– based screening system (Ehrhard et al., 2000), and the split-ubiquitin system (Johnsson and Varshavsky, 1994). Genome-wide protein-protein interaction studies have been carried out in many organisms including bacteriophage T7 (Bartel et al., 1996), Hepatitis C virus (HCV; Flajolet et al., 2000), Helicobacter pylori (Rain et al., 2001), Caenorhabditis elegans (Walhout et al., 2000), Saccharomyces cerevisiae (Uetz et al., 2000; Ito et al., 2001; Gavin et al., 2002; Ho et al., 2002), and mouse (Suzuki et al., 2001). The examples described in this unit focus on S. cerevisiae, which is not only a good model organism for eukaryotes, but also contains the most protein-protein interaction data generated thus far. The following descriptions of experimental technologies used in yeast studies demonstrate the types and volumes of data that have been generated.
Yeast two-hybrid The yeast two-hybrid system (UNIT 19.2), first described by Field and Song (1989), is the most widely used method for detecting protein-protein interactions in highthroughput mode. The yeast two-hybrid technique involves two fusions: a bait protein fused to the DNA-binding domain of a transcription factor, and potential interacting partners (prey) fused to the activation domain of the same transcription factor. An interaction between the bait and prey brings the two components of the transcription factor into spatial association, resulting in the formation of a functional transcription factor, which induces the expression of a specific reporter gene, thereby allowing such interactions to be detected. It should be noted that this approach forces the proteinprotein interaction between the bait and prey to occur in nuclei, and some errors in measuring protein interactions may result from this restriction. In recent years, variations of the two-hybrid system have been developed (Drees, 1999), and the methods have also been extended to other organisms others, including bacteria and viruses (Uetz and Hughes, 2000). Many protein-protein interaction data have been generated using the two-hybrid system. In a proteome-wide study in yeast by Uetz et al. (2000), two experiments were performed, one with a low-throughput protein array and one with a high-throughput array. In the lowthroughput array, 192 bait proteins were tested
against a set of 6000 prey proteins, and a total of 281 binary interactions were identified. The high-throughput approach used the complete set of 6000 yeast proteins as baits against the complete set of 6000 prey proteins. The second approach identified 692 interacting protein pairs involving 817 unique proteins as either bait or prey proteins. An independent, large-scale project by Ito et al. (2001) was also conducted for the entire yeast proteome. This study detected 3278 proteins involved in 4589 protein-protein interactions.
Mass spectrometry A typical approach to identifying proteins in a protein complex is separation of the various proteins of an extract by gel electrophoresis followed by mass spectrometric analysis of the protein derived from the gel spots (see UNIT 16.1). Polypeptides can then be identified by searching protein databases for the experimentally determined molecular weights. High throughput is achieved by using automated matrix-assisted laser desorption/ionization (MALDI; see UNIT 16.2), providing a list of the masses of the peptide fragments. Recently, Gavin et al. (2002) have taken a new approach, called tandem affinity purification (TAP), to screening protein-protein interactions on the proteome scale. This method is particularly effective for identifying protein complexes that contain three or more components. First, the authors modified the genetic sequences of hundreds of proteins to yield bait proteins with specific amino acid tags. They then expressed these modified proteins in yeast cells, where they formed physiological complexes with other proteins. By using the tag, each bait protein was pulled out of the cell lysate, usually retrieving the entire complex along with it. The extracted proteins were identified using MALDI mass spectrometry. Note that the tags may perturb some protein interactions and yield erroneous data. By using TAP, Gavin et al. (2002) identified 1440 distinct proteins within 232 multiprotein complexes in yeast after processing 1739 genes as baits. Of these complexes, 91% contained at least one protein of unknown function. Ho et al. (2002) used a similar approach, which they termed high-throughput mass spectrometric protein complex identification (HMS-PCI). This group constructed an initial set of 725 tagged proteins, from which they identified 3617 associated proteins, covering about 25% of the yeast proteome.
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Protein chip Another approach to generating a proteinprotein interaction map is protein-chip technology (Zhu et al., 2000; also see UNIT 23.5). In this approach, proteins are expressed, purified, and screened in a high-throughput system in which a large number of proteins are attached to a planar substrate (chip) as discrete spots at known locations; the proteins maintain their folded conformations and their ability to interact specifically with other proteins. A solution containing labeled protein(s) to be tested is incubated with the chip, and the chip is then washed. Specific interactions between proteins on the chip and proteins from the solution are indicated by the position of the label. In addition to the rapid simultaneous measurement of large numbers of samples, protein chip technology has substantial advantages over conventional methods, especially the high signal-to-noise ratio, the small amount of sample needed, and high sensitivity. On the other hand, attachment of proteins to chips can disrupt some protein interactions.
Phage display Phage display is another method for studying protein-protein interactions (Mullaney and Pallavicini, 2001). It is based on the ability of bacteriophage to express engineered proteins on their surface coat. Diverse libraries such as peptides, antibodies, and protein domains corresponding to gene fragments can be displayed on the virus coat by way of a foreign DNA sequence artificially inserted into the phage genome. By immobilizing a protein on an affinity support, phages with display proteins binding to the immobilized protein can be selected from the library. The phages selected on the basis of their interaction with the immobilized protein can be enriched, and the proteins on the phages that interact with the immobilized proteins can be identified.
Protein-Protein Interaction Databases and Visualization Tools
Bioinformatics Analysis for Interactive Proteomics
Data management is critical when using high-throughput protein-protein interaction systems. Several Internet databases have been established to collect, retrieve, and describe protein-protein interactions (see Table 25.1.2), and protein interactions retrieved from the literature were also included in the databases (Ono et al., 2001). These databases typically have user-friendly interfaces, and most also provide good searching capability. One can search the interactions of a particu-
lar protein by querying its open reading frame (ORF) name or its gene name. A functional annotation, when available, is usually given for a protein participating in an interaction, and the experimental source and reference for the interaction are also provided in most databases. Such information can help evaluate errors and validate protein-protein interactions in experimental data. In addition, protein-protein interaction data in a centralized database provide a starting point (input) for computer programs that analyze protein-protein interaction on the proteome scale. Data visualization helps users process, interpret and act upon data. With more and more protein-protein interaction data collected into databases, a long list of interacting proteins or a table of protein pairs falls short of capturing and understanding the complex relationships underlying protein interaction data. The information presented in the form of images is more intelligible since it transforms the original data into a geometric representation. A visualization tool usually includes data manipulation functions to retrieve data, remove unwanted data, and reorganize data. Visualization tools can further help researchers validate interactions from different experimental sources, make sense of interaction paths, and construct hypotheses for biological pathways. A protein interaction network is represented as a graph with proteins as nodes and interactions as edges (Fig. 25.1.1), facilitating location of proteins by positions on a map. Once a protein has been identified in the graph layout, its immediate and indirect neighbors can be displayed and the relationships between this protein and any other protein can be revealed. To make best use of protein interaction networks, the integrated systems of database, graphics, and computational tools allow the simultaneous analysis of various types of biological data relating to proteins and interactions. For example, PIMRider provides an integrated platform for the exploration of protein interaction maps and other genomic/proteomic information (Schachter, 2002). Most protein-protein interaction databases provide visualization tools (see Table 25.1.2). InterViewer (http://wilab.inha.ac.kr/protein/) is a free tool to query and visualize protein-protein interaction data, which can then be further refined or navigated to explore protein interactions. A Web applet (http://www.charite.de/bioinformatics) can be implemented to browse protein-protein interactions. It enables the display of interaction relationships based on neighboring distance and
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Xenarios et al. (2002)
Database of Interacting Proteins
Biomolecular Interaction Bader et al. Network Database (2001)
Munich Information Center for Protein Sequences
Molecular Interaction Database
Biomolecular Relations in Information Transmission and Expression
Pathcalling Yeast Interaction Database
Hybrigenics
The General Repository for Interaction Datasets
DIP
BIND
MIPS
MINT
BRITE
PathCalling
PIMRider
GRID
a Number of binary interactions and number of protein complexes.
—
—
—
Duan et al. (2002)
Zanzoni et al. (2002)
Mewes et al. (2000)
Reference
Database name
Acronym
Table 25.1.2 Online Protein-Protein Interaction Databases
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http://biodata.mshri.on.ca/grid/
http://pim.hybrigenics.com/
http://curatools.curagen.com/pathcalling portal/
http://www.genome.ad.jp/brite/
http://cbm.bio.uniroma2.it/mint/
http://mips.gsf.de/proj/yeast/CYGD/db/
http://bind.ca/
http://dip.doe-mbi.ucla.edu
Web site
14,318
1400
4549
6971
4586
11,200
16644
18,494
1050
1306
Yes
Yes
Yes
No
Yes
No
Yes
Yes
Database sizea Visual Binary Complex tool?
Yes
Yes
Yes
Yes
Yes
Yes
Yes
Yes
Yes
No
No
Yes
Yes
Yes
Yes
No
Academic Commercial free free access? access?
biological functions. By choosing two such parameters, users can find interactions of interest. Cytoscape (http://www.cytoscape.org/) is another bioinformatics software platform for visualizing molecular interaction networks. It can integrate protein interactions with gene expression profiles and other biological data. The inputs include various molecular interactions such as protein-protein and protein-DNA interaction pairs. As a key feature, Cytoscape provides an algorithm that identifies active sub-network/pathway modules in a graph. Currently, Cytoscape runs under Java on both Linux and Windows platforms. Pajek (http://vlado.fmf.unilj.si/pub/networks/pajek/) is a program for Windows used to analyze large networks. It is free to noncommercial users. In addition to network visualization, Pajek provides other feature functions such as partition, permutation, clustering, and hierarchical decomposition. Nine widely used protein-protein interaction databases are described below, together with their visualization tools (if available). Table 25.1.2 summarizes information about the databases including, acronym, name, literature reference, Web site, size as of June 2003 (number of binary interactions and number of protein complexes), availability of a visualization tool, and permission for free academic or commercial use.
DIP DIP provides an integrated tool for browsing and extracting information about proteinprotein interactions, which are either generated from various high-throughput experiments or collected from literature searches. The vast majority of data are derived from yeast, Helicobacter pylori and humans. DIP allows the visual representation and navigation of protein-protein interaction networks. The reproducibility of a given interaction can be assessed visually by the thickness of the line between two proteins. A related tool, LiveDIP, also integrates the protein-protein interactions network with large-scale gene expression data.
BIND
Bioinformatics Analysis for Interactive Proteomics
The BIND database stores various interactions between molecular compounds including protein-protein, protein-RNA, protein-DNA, and protein-ligand interactions. Descriptions of interactions include subcellular localizations of the proteins involved and experimental conditions used to observe the interactions. This database also contains information about
molecular complexes and pathways. BIND can be visually navigated using a Java applet. BIND also provides a framework for users to build their own protein-protein interaction databases.
MIPS The MIPS Comprehensive Yeast Genome Database (CYGD) provides the proteinprotein interaction data together with sequence and function information for all the genes in the budding yeast Saccharomyces cerevisiae. The protein-protein interaction data, as well as other compiled yeast data including functional classification category, subcellular localization category, and EC number category, are available to download as text files.
MINT The MINT database stores data, extracted from the scientific literature, about functional interactions between proteins and includes information about the effects of enzymatic modifications of one of the partners. The interaction data can be extracted and visualized graphically. Presently MINT contains 4586 interactions, 782 of which are genetic interactions.
BRITE BRITE is a database of binary proteinprotein interactions retrieved from the literature, as well as two-hybrid data for S. cerevisiae and H. pylori proteins.
PathCalling The PathCalling database (http://curatools. curagen. com / pathcalling portal / ) contains yeast protein-protein interaction data from high-throughput yeast two-hybrid experiments. Visualization tools are also available for depicting the protein interaction network. Figure 25.1.2 shows the graph for interaction mapping around the protein Nup100p. Each node represents one protein and each edge marks an interaction. All the immediate neighbors and next immediate neighbors in the protein interaction map for Nup100p are displayed. By clicking a node, one can navigate its neighbors and obtain detailed descriptions of the gene products, including amino acid sequences and protein functions.
PIMRider PIMRider contains protein-protein interaction data for Helicobacter pylori, which has been studied using genome-wide two-hybrid
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Figure 25.1.2 The protein interaction map in yeast around Nup100p from PathCalling (http://curatools.curagen.com/pathcalling portal/). A gene is represented as a node and a proteinprotein interaction is indicated as an edge.
assays (Kanehisa et al., 2002). A graph of 1400 protein-protein interactions can be viewed and assigned a PIM Biological Score (PBS score; PBS is a registered trademark of Hybrigenics) that quantifies the confidence level of each interaction, allowing filtering of interactions based on their confidence levels. The PBS score takes into account the characteristics of the libraries screened and the target organisms as well as the results of the screens. The PBS score ranges from 0 (the best) to 1 (the worst). In PIMRider, tools have also been developed to identify the specific protein domain involved in a given interaction and to infer the interaction pathways between two proteins.
GRID GRID is a database of genetic and physical interactions. At present it includes 14,138 unique protein-protein interactions, including the data from MIPS and BIND. Osprey Network Visualization System (http:// biodata.mshri.on.ca/osprey/index.html) is integrated into the database to allow users to visualize searched results. Users can also upload their own data sets and visualize the interaction maps.
Quality of Protein-Protein Interaction Data To effectively use high-throughput proteinprotein interaction data for biological inference, it is essential to evaluate the coverage and reliability of data. The systematic proteinprotein interaction assay methods described above clearly show that a single screening rarely captures all proteins capable of interacting with a given bait protein. In the
yeast two-hybrid array, even multiple screenings with different two-hybrid variants usually produce only partial overlaps. Several different complementary methods are necessary to identify as many legitimate interactions as possible. Two examples illustrate the complementarity of the two-hybrid and mass-spectrometry methods. The first example involves interactions with the RNA polymerase I–specific transcription initiation factor Rrn10p (YBL025W). Using mass spectrometry, Ho et al. (2002) demonstrated that there was no detectable protein complex involving Rrn10p. Yeast two-hybrid assays (Uetz et al., 2000) detected interaction between Rrn10p and Rrn9p, which was also validated by other experiments. It was found from genome-wide analysis of mRNA abundance that the expression level of Rrn10p is much lower than the average value (Holstege et al., 1998); thus, it is likely that the protein complex may not be formed due to the low abundance of Rrn10p. It is also possible that this pair of proteins have only a transient interaction rather than forming a stable complex. On the other hand, yeast twohybrid assays are independent of protein concentration and are capable of detecting transient and weak interactions. A converse example involves Cmd1p, a small, ubiquitous Ca2+ -binding protein that regulates a wide variety of proteins and processes in all eukaryotes (Cyert, 2001). Yeast two-hybrid analysis cannot detect any interactions with this protein, while mass spectrometry using different bait proteins detects several complementary protein complexes (see Fig. 25.1.3). In response to a Ca2+ signal, Cmd1p binds Ca2+ and consequently undergoes a conformational change that allows it to bind and
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Figure 25.1.3 Three Cmd1p related protein complexes identified by mass spectrometry. The annotated Cmd1p interacting partners (i.e., proteins with known functions), shown in dark color, are derived from experimental results reviewed in Cyert (2001).
Bioinformatics Analysis for Interactive Proteomics
activate a host of target proteins. The absence of native physiological conditions (i.e., insufficient Ca2+ concentration), may prevent the yeast two-hybrid assay from detecting the interaction. Confidence in the authenticity of a reported interaction is increased by observations of the same interaction using different methods. For example, if an interaction is detected by two distinct experiments, the joint observations enhance the confidence level for this particular interaction. Large-scale twohybrid screening can identify some classes of systematic false positive results using multiple, independent screening experiments under standardized conditions. For example, some interactions related to particular proteins tend to appear repeatedly with unrelated baits, indicating false positive data. Automated literature searching is another resource for validating protein-protein interactions generated by high-throughput techniques. The more often two protein names appear in the same paper, the better the possibility that they may interact with each other. While such information may not be entirely reliable for validating protein-protein interactions, it can at least provide clues for evaluating the authenticity of an interaction. Computational approaches, as discussed in the next section, can also be
used to assess the reliability of observed highthroughput protein-protein interactions. The internal relationships between proteinprotein interactions and other biological data such as subcellular localization data, protein function and structure, and gene expression profiles provide a basis for cross-validating the data and offering more information than a single source of data can achieve (Chen and Xu, 2003). For example, two proteins having a true interaction often have related functions, similar cellular role, correlated gene expression pattern, and the same subcellular localization.
In Silico Prediction of Protein-Protein Interactions In addition to systematic analyses of protein interactions using high-throughput experiments, a number of computational methods have been developed for predicting proteinprotein interactions based on protein/DNA sequence information using computer-based modeling (in silico). Predicted protein-protein interactions are less reliable than results generated from high-throughput experiments. However, they expand the scope of experimental data and are useful in assessing experimentally detected protein-protein interactions. These in silico methods include three major computational areas: gene
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fusion, conserved genetic neighborhood, and co-occurrence of genes in genomes.
Gene fusion is based on the observation that the homologs of a pair of interacting proteins in one organism may be fused into a single protein chain in another organism (Marcotte et al., 1999). For example, two separate proteins A and B in organism X are naturally expressed as a single fusion protein in another organism Y. A and B as protein domains in organism Y generally interact with each other, and this implies that A and B as separate proteins in organism X probably interact too.
from 89 species. STRING contains a unique scoring framework based on different types of interactions and associations, integrated into a single confidence score. The graphical representation of the network of inferred, weighted protein interactions provides a highlevel view of protein interaction and functional linkage, facilitating the analysis of modularity in biological processes. The user inputs a gene/protein identifier or, alternatively, the raw amino acid sequence of a protein. The result will be a list of predicted interacting proteins ranked by their confidence scores, together with graphics for navigation of the functional categories of the protein interactions.
Conservation of gene neighborhood
Predictome
Proteins with a conserved genetic neighborhood in bacteria, i.e., a group of genes arranged in tandem, which often appear in a similar arrangement in other related genomes, tend to interact with each other to form complexes (Overbeek et al., 1999). Operons represent one such conserved gene context. Identification of operons or other “conserved gene contexts” can provide clues about sets of proteins that may form complexes. This can be done through discovering the sequential arrangement of genes in a microbial genomic sequence and conserved gene context across multiple related microbial genomes.
Predictome (http://predictome.bu.edu/) stores predicted interactions among the proteins from 44 genomes based on the three computational methods discussed above (see In Silico Prediction of Protein-Protein Interactions). In addition, published experimental data of protein-protein interactions from yeast two-hybrid, coimmunoprecipitation, and other methods are included in the database. Users input gene/protein identifiers and species names. The output is a graphic view of the interactions together with functional annotations for related proteins.
Co-occurrence of genes in genome
Biological Inference Through Protein Interaction Data
Gene fusion
During evolution, functionally linked proteins tend to be either conserved together or completely eliminated in a new species. Such information can be represented by a phylogenetic profile that records the presence or absence of a protein in every known genome in a phylogenetic tree (Pellegrini et al., 1999). It has been shown that proteins having similar profiles tend to have physical interactions and to be functionally linked.
Databases of Predicted Protein-Protein Interactions Several databases of predicted proteinprotein interactions including visualization tools are available for online searches. Two of them, STRING and Predictome, are described below.
STRING The database STRING (http://www.bork. embl-heidelberg.de/STRING/) is a resource for the exploration and analysis of protein interactions and functional associations. STRING currently contains 261,033 gene sequences
Protein-protein interaction networks contain information about individual proteins on multiple levels, including their partners, functions, interactive complexes, and biological pathways in which they are involved. Thus, protein-protein interaction data are useful for assigning functions to uncharacterized gene products and for constructing biological pathways, especially signal transduction pathways. However, biological inference from proteinprotein interaction data is not trivial, given the complexity and errors in protein interaction maps. Manual analyses subsequent to computational predictions are often necessary.
Protein function prediction A protein often performs its function by interacting with other proteins having the same or related cellular functions. Discovering that a protein of unknown function interacts with one or more proteins of known function provides a valuable clue to the role of the novel gene product through “guilt by association.” For example, if protein X (uncharacterized)
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Figure 25.1.4 Output generated by GENFAS (http://digbio.missouri.edu/genefas/) to predict the function of the hypothetical gene YER079W. “Index” indicates the gene ontology (GO) hierarchical level. “Reliability Score” is used to rank the predicted functions. “Probability” gives the likelihood of prediction accuracy. “GO identifier” is the entry ID for GO.
Bioinformatics Analysis for Interactive Proteomics
is found to interact with protein Y and protein Z, and both Y and Z are components of the DNA transcription processing machinery, then it is likely that protein X is also involved in this process, perhaps as part of the complex containing Y and Z. Based on such an approach, high-throughput proteinprotein interaction data provide a good coverage for many novel proteins, the functions of which cannot be assigned by sequence comparison. Schwikowski et al. (2000) collected 2709 published protein-protein interaction data in yeast and clustered them based on cellular role and subcellular localization annotated in the Yeast Proteome Database (YPD; http://www.proteome.com/). They compiled a list of about 370 proteins with unknown functions that interact with at least one protein with a known function. Among them, 29 proteins have two or more interacting partners with a common function. Prediction of function through proteinprotein interaction data is nontrivial since the data are noisy with many false positives and false negatives. Moreover, protein-protein interaction information alone may be insufficient for predictive purposes. Hence, it is important to integrate different sources of data, such as protein interaction, genomic sequence,
and gene expression data, to provide better predictions of protein functions. GENFAS (http://digbio.missouri.edu/genefas/) was developed to annotate unknown proteins’ functions by using all available genome-scale data in yeast S. cerevisiae, e.g., protein-protein interaction data, genetic interaction data, and microarray data. Given a gene/protein identifier or a protein sequence, GENFAS predicts the protein function based on combined probability calculated from various high-throughput data in the format shown in Figure 25.1.4. The possible functions of the protein according to the gene ontology (GO) annotation indices are listed according to reliability scores.
Biological pathway construction Protein-protein interaction networks not only allow assignment of cellular functions to novel proteins, but also provide the resources for constructing biological pathways. Physical protein-protein interactions often provide the backbones for determining biological pathways. For example, in a signaltransduction pathway, protein-protein interactions generally play the main role in the signal-transduction cascade. Therefore, highthroughput protein-protein interaction data, in conjunction with other high-throughput
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Figure 25.1.5 Relationship between biological pathway and protein-protein interaction data. (A) The MAPK signaling pathway for filamentation (i.e., the connections between genes) taken from KEGG. (B) A MAPK signal transduction pathway constructed using protein interaction data.
data and protein function annotations, can help construct in silico biological pathways, at least partially. Figure 25.1.5 illustrates a validation of this approach. As a test, the author chose the well-known MAPK signal transduction pathway of filamentation from the KEGG databases at GenomeNet (http://www. genome.ad.jp/kegg/; Kanehisa et al., 2002; see Fig. 25.1.5A). This pathway can be rebuilt using protein-protein interaction data, as shown in Figure 25.1.5B. A Web server (http://digbio.missouri.edu/ genepath) has been developed to systematically and automatically derive partial biological pathways in the yeast Saccharomyces cerevisiae using high-throughput biological data, including yeast two-hybrid data, protein complexes identified from mass spectroscopy, genetic interactions, and microarray gene expression data (Chen et al., 2003a). The inputs of the server are two termini of a pathway, i.e., the upstream starting protein (e.g., a signal sensor YBL062W) and the downstream terminal protein (e.g., a transcription factor YNL119W). The output is the predicted protein-interaction chain between the two terminal proteins. Figure 25.1.6 is a snapshot of a predicted pathway generated by the server when using YBL062W
and YNL119W as the two terminal proteins. Protein annotation information such as function classification and subcellular localization is also given. This server can help piece together much more complicated pathways in yeast (Chen et al., 2003b).
Summary As the major focus in biological research is moving from genome to proteome, analyses of protein-protein interactions will play an increasingly important role. High-throughput protein-protein interaction data provide valuable global information, although current data deliver limited coverage of all possible protein interactions and contain false positives. With improved high-throughput experimental techniques, more high-quality protein-protein interaction data will be generated. Moreover, well developed databases and integrated computational tools will enable biologists to easily navigate protein-protein interaction networks and make new biological discoveries. It may not be long before computational analysis of protein-protein interaction data becomes a routine procedure in many biological laboratories, just as sequence homology research is today.
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Figure 25.1.6 Output generated by the Web server http://digbio.missouri.edu/genepath showing a predicted biological pathway using products of YBL062W and YNL119W as the two terminal proteins. “GO MF ID” indicates GO molecular function identification codes. The numbers in the Function row indicate GO hierarchical levels.
Acknowledgments This work was funded by the U.S. Department of Energy Genomes to Life program under the project “Center for Molecular and Cellular Systems” (http://www.ornl.gov/ GenomestoLife), under Contract DE-AC0500OR22725, managed by UT-Battelle, LLC.
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Ho, Y., Gruhler, A., Heilbut, A., Bader, G.D., Moore, L., Adams, S.L., Millar, A., Taylor, P., Bennett, K., Boutilier, K., Yang, L., Wolting, C., Donaldson, I., Schandorff, S., Shewnarane, J., Vo, M., Taggart, J., Goudreault, M., Muskat, B., Alfarano, C., Dewar, D., Lin, Z., Michalickova, K., Willems, A.R., Sassi, H., Nielsen, P.A., Rasmussen, K.J, Andersen, J.R., Johansen, L.E., Hansen, L.H., Jespersen, H., Podtelejnikov, A., Nielsen, E., Crawford, J., Poulsen, V., Sorensen, B.D., Matthiesen, J., Hendrickson, R.C., Gleeson, F., Pawson, T., Moran, M.F, Durocher, D., Mann, M., Hogue, C.W., Figeys, D., and Tyers, M. 2002. Systematic identification of protein complexes in Saccharomyces cerevisiae by mass spectrometry. Nature 415:180-183.
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Holstege, F.C., Jennings, E.G., Wyrick, J.J., Lee, T.I., Hengartner, C.J., Green, M.R., Golub, T.R., Lander, E.S., and Young, R.A. 1998. Chromosomal landscape of nucleosome-dependent gene expression and silencing in yeast. Cell 95:717728. Ito, T., Chiba, T., Ozawa, R., Yoshida, M., Hattori, M., and Sakaki, Y. 2001. A comprehensive two-hybrid analysis to explore the yeast protein interactome. Proc. Natl. Acad. Sci. U.S.A. 98:4569-4574. Johnsson, N. and Varshavsky, A. 1994. Split ubiquitin as a sensor of protein interactions in vivo. Proc. Natl. Acad. Sci. U.S.A. 91:1034010344. Kanehisa, M., Goto, S., Kawashima, S., and Nakaya, A. 2002. The KEGG databases at GenomeNet. Nucl. Acids Res. 30:42-46.
Mullaney, B.P. and Pallavicini, M.G. 2001. Proteinprotein interactions in hematology and phage display. Exp. Hematol. 29:1136-1146. Ono, T., Hishigaki, H., Tanigami, A., and Takagi, T. 2001. Automated extraction of information on protein-protein interactions from the biological literature. Bioinformatics 17:155-161.
Pellegrini, M., Marcotte, E.M., Thompson, M.J., Eisenberg, D., and Yeates, T.O. 1999. Assigning protein functions by comparative genome analysis: Protein phylogenetic profiles. Proc. Natl. Acad. Sci. U.S.A. 96:4285-4288. Pelletier, J. and Sidhu, S. 2001. Mapping proteinprotein interactions with combinatorial biology methods. Curr. Opin. Biotechnol. 12:340347. Phizicky, E.M. and Field, S. 1995. Protein-protein interactions: Methods for detection and analysis. Microbiol. Rev. 59:94-123. Rain, J.C., Selig, L., De Reuse, H., Battaglia, V., Reverdy, C., Simon, S., Lenzen, G., Petel, F., Wojcik, J., Schachter, V., Chemama, Y., Labigne, A., and Legrain, P. 2001. The proteinprotein interaction map of Helicobacter pylori. Nature 409:211-215. Schachter, V. 2002. Protein-interaction networks: From experiments to analysis. Drug Discov. Today 7:S48-S54. Schwikowski, B., Uetz, P., and Fields, S. 2000. A network of protein-protein interactions in yeast. Nat. Biotechnol. 18:1257-1261. Suzuki, H., Fukunishi, Y., Kagawa, I., Saito, R., Oda, H., Endo, T., Kondo, S., Bono, H., Okazaki, Y., and Hayashizaki, Y. 2001. Protein-protein interaction panel using mouse full-length cDNAs. Genome Res. 11:1758-1765.
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Uetz, P. and Hughes, R.E. 2000. Systematic and large-scale two-hybrid screens. Curr. Opin. Microbiol. 3:303-308. Uetz, P., Giot, L., Cagney, G., Mansfield, T.A., Judson, R.S., Knight, J.R., Lockshon, D., Narayan, V., Srinivasan, M., Pochart, P., Qureshi-Emili, A., Li, Y., Godwin, B., Conover, D., Kalbfleisch, T., Vijayadamodar, G., Yang, M., Johnston, M., Fields, S., and Rothberg, J.M. 2000. A comprehensive analysis of protein-protein interactions in Saccharomyces cerevisiae. Nature 403:623627. Walhout, A.J., Soredella, R., Lu, X., Hartley, J.L., Temple, G.F., Brasch, M.A., Thierry-Mieg, N., and Vidal, M. 2000. Protein interaction mapping in C. elegans using proteins involved in vulval development. Science 287:116-122. Xenarios, I., Salwinski, L., Duan, X.J., Higney, P., Kim, S., and Eisenberg, D. 2002. DIP, the Database of Interacting Proteins: A research tool for studying cellular networks of protein interactions. Nucl. Acids Res. 30:303-305.
Zanzoni, A., Montechi-Palazzi, L., Quondam, M., Ausiello, G., Helmer-Citterich, M., and Cesareni, G. 2002. MINT: A Molecular INTeraction database. FEBS Lett. 513:135-140. Zhu, H., Klemic, J.F., Chang, S., Bertone, P., Casamayor, A., Klemic, K.G., Smith, D., Gerstein, M., Reed, M.A., and Snyder, M. 2000. Analysis of yeast protein kinases using protein chips. Nat. Genet. 26:283-289.
Contributed by Yu Chen Monsanto Company St. Louis, Missouri Dong Xu University of Missouri—Columbia Columbia, Missouri
Bioinformatics Analysis for Interactive Proteomics
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CHAPTER 26 Protein Engineering INTRODUCTION he growth of protein science over the past 50 years has culminated in the ability to design and produce proteins by recombinant technology. What was a dream in the 1960s is now a reality. This chapter will describe methodologies for accomplishing both parts of protein engineering: the design and the production aspects. Many new technologies will be added as the chapter develops over time.
T
The chapter begins with a unit from Pfeffer and colleagues on the use of a cysteine scanning mutagenesis to identify surface interaction regions of a protein (UNIT 26.1). The methodology for introduction of cysteine residues described in this unit is also useful in other applications. Arnold and colleagues detail the reactions needed to construct libraries of chimeras of distantly related or nonrelated genes through the combination of gene fragments using specific overhangs (UNIT 26.2). Detailed protocols for the steps necessary for incorporation of isotopically enriched amino acids into proteins for multi-dimensional NMR studies are described by Long and Edison (UNIT 26.3). Additional units planned for future updates to this chapter include: fusions to promote folding, including high-throughput screens for solubility, addition of extensions to either terminus to target proteins or to test function of ends, addition of inserts within a sequence for targeting or to introduce a functional site, methods for humanizing proteins and antibodies, methods for creating domains and multimers, descriptions of protein design principles, and methods for engineering solubility and stability into proteins for functional reasons or to create products that could be used in humans or animals. In addition, units will be included on site-directed mutagenesis for: (a) studies of protein function; (b) introducing or deleting glycosylation sites; (c) introducing metal-binding sites; (d) introducing cysteines for creation of disulfide linkages within proteins; (e) creating soluble mutants for crystallography; and (f) adding cysteine residues for cross-linking to other proteins or to DNA. Ben M. Dunn
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Misincorporation Proton-Alkyl Exchange (MPAX): Engineering Cysteine Probes into Proteins
UNIT 26.1
A method to obtain a map of protein topology is described in this unit. When combined with a high-throughput screen for detecting mutations that perturb ligand binding, this method provides a very useful tool to identify amino acid residues that contribute to protein-protein interfaces. This approach makes use of misincorporation proton-alkyl exchange (MPAX) to determine the topology of individual amino acid residues within a folded protein. Cysteine residues are incorporated at random and at a low frequency when the protein is expressed in E. coli cells harboring specific mutant cysteine tRNAs. These “misincorporator tRNAs” introduce cysteine in place of a given amino acid. The resulting misincorporated peptides contain one cysteine residue on average per protein molecule, in place of a specific amino acid. The chemical reactivity of the cysteine to an alkylating reagent is used in combination with 2-nitro-5-thiocyanobenzoic acid–driven chemical cleavage of the protein backbone to measure the relative burial of that residue in the folded protein structure. A surface map is thereby generated, which reveals the identity of surface-localized hydrophobic residues that are likely to contribute to hydrophobic protein-protein interfaces. Mutant protein pools generated with distinct misincorporator tRNAs can also be screened simultaneously in a single affinity chromatography step to reveal differences in ligand binding affinity. In this unit, the use of MPAX technology to misincorporate cysteine residues (see Basic Protocol 1) and chemical cleavage of the protein backbone at the misincorporation site (see Basic Protocol 2) are described. Protocols to determine the solvent accessibility of misincorporated cysteines in the context of native versus unfolded proteins are then presented (see Basic Protocols 3 and 4). Basic Protocol 5 describes how to analyze MPAX data to enable prediction of binding interfaces. In addition, the combination of MPAX with affinity chromatography to provide a high-throughput method to select for affinity-altered mutant proteins in vitro (see Basic Protocol 6) is described. Note that this protocol is independent of Basic Protocols 3 through 5. These methods require high-resolution separation of small peptides by gel electrophoresis (see Support Protocol 1). Finally, this unit describes the assembly of a synthetic gene that can be used to achieve maximal cysteine misincorporation (see Support Protocol 2 and 3).
STRATEGIC PLANNING Naturally occurring cysteines To simplify the biochemical readout of cysteine misincorporation, cysteine codons in the gene of interest may be replaced, for instance with alanine codons, by conventional site-directed mutagenesis (see Critical Parameters for further discussion). This is helpful because naturally occurring cysteine residues that have been modified by 2-nitro 5-thiocyanobenzoic acid are cleaved with 84% efficiency (Silverman and Harbury, 2002a). This results in a major cleavage product that may occlude the signal from any misincorporated cysteine that is nearby in the protein sequence (see Table 1 in Silverman and Harbury, 2002b). In addition, signal from misincorporated cysteines that are distal to both the protein kinase A (PKA) tag and the naturally occurring cysteine only arise Protein Engineering Contributed by Alondra Schweizer Burguete, Pehr B. Harbury, and Suzanne R. Pfeffer Current Protocols in Protein Science (2005) 26.1.1-26.1.23 C 2005 by John Wiley & Sons, Inc. Copyright
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from peptides that remain uncleaved at the naturally occurring cysteine (and still contain a PKA tag).
Protein kinase A labeling motif Peptides containing misincorporated cysteines are detected by end-radiolabeling. For this purpose, a protein kinase A recognition sequence is introduced at one end of the polypeptide. Thus, the gene of interest should be prepared to include this motif (encoding amino acids Nter -GRRASIY-Cter ). Gene assembly Cysteine misincorporation is enabled by co-expression of a gene of interest together with specific cysteine misincorporator tRNAs in E. coli. Misincorporator tRNAs recognize a subset of naturally occurring triplets that encode a given amino acid. Maximal misincorporation is possible when a synthetic gene is utilized, in which the codons are selected for optimal recognition by the tRNAs employed. Support Protocol 2 can be followed to assemble a gene containing exclusively amino acid triplet codons that match the specific misincorporator tRNAs employed (see Table 26.1.2). Support Protocol 2 is not essential but will provide maximal cysteine misincorporation. BASIC PROTOCOL 1
INTRODUCTION OF CYSTEINE RESIDUES BY TRANSLATIONAL MISINCORPORATION This protocol describes the replacement of a specific amino acid within a polypeptide with cysteine, using translational misincorporation in E. coli. The process requires a mutant cysteine tRNA bearing an anticodon triplet that matches the amino acid to be replaced (see Table 26.1.2). In the example shown in Figure 26.1.1, the cysteine tRNA anticodon has been mutated to that of valine (GAC). Simultaneous expression of this mutant tRNA and a gene of interest results in the introduction of cysteine at a random valine position in the growing polypeptide chain. This technique makes use of vectors (pMPAX) encoding specific mutant E. coli cysteine transfer RNAs and cysteinyl tRNA synthetase (Silverman and Harbury, 2002a). This protocol describes how to co-express a gene of interest together with a pMPAX vector to yield an ensemble of peptides containing cysteine misincorporated at different positions. The protein can then be purified according to individual protocols.
Materials pMPAX vectors (Harbury laboratory, Stanford University; [email protected]) Kanamycin-resistance plasmid (pET28a, Novagen; pET28a pH6 PKA and pET28a PKA pH6, Harbury laboratory, Stanford University) BL21(DE3) E. coli cells (Novagen) LB plates (APPENDIX 4A) containing 30 µg/ml kanamycin and 100 µg/ml carbenicillin LB medium (EMD Biosciences) containing 30 µg/ml kanamycin and 100 µg/ml carbenicillin 1× PBS (see recipe) M63 medium containing 30 µg/ml kanamycin and 100 µg/ml carbenicillin (see recipe) 1 M isopropyl-β-D-thiogalactopyranoside (IPTG) MPAX: Engineering Cysteine Probes into Proteins
Incubator and shaker (37◦ C and 30◦ C) Culture flasks, sterile Additional reagents and equipment for electroporation (UNIT 5.10)
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Figure 26.1.1 Translational misincorporation of cysteine residues at valine positions by use of a mutant cysteine transfer RNA.
1. Mix ∼1 µg each of pMPAX vector and a kanamycin-resistance plasmid containing the gene of interest under the control of a T7 promoter. Note that the gene of interest is fused to a sequence encoding a protein kinase A recognition motif. Use sterile technique through step 7.
2. Electroporate (UNIT 5.10) the DNA mixture from step 1 into BL21(DE3) E. coli cells. Electroporation is the method of choice for transforming E. coli because this method yields a high number of double transformants. It is recommended to transform E. coli cells with the gene of interest in the absence of pMPAX and grow on kanamycin plates and medium as a negative control.
3. Spread 50 µl of transformed cells onto LB plates containing 30 µg/ml kanamycin and 100 µg/ml carbenicillin. Incubate overnight at 37◦ C. 4. Transfer a single colony into 50 ml LB containing 30 µg/ml kanamycin and 100 µg/ml carbenicillin at the end of the day. Shake overnight at 37◦ C. 5. Wash cells with 1× PBS, centrifuge 15 min at 5000 × g, 4◦ C, decant supernatant, and resuspend in 50 ml of 1× PBS.
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6. Inoculate cells 1:50 into M63 medium containing 30 µg/ml kanamycin and 100 µg/ml carbenicillin. Shake at 37◦ C until cells reach an OD600 of ∼0.5 (4 to 6 hr). An initial expression volume of 100 to 500 ml is recommended. This can subsequently be scaled up depending on the amounts of protein required.
7. Add IPTG to a final concentration of 1 mM. Shake for 3 hr at 30◦ C or according to a protein-specific optimized protocol. 8. Centrifuge cells 15 min at 5000 × g, 4◦ C. Discard supernatant. Lyse cells and purify the protein of interest by an appropriate protocol (see UNITS 6.1, 6.2, 6.3, 6.6, 6.7 & 8.1). Non-lysed bacterial pellets can be frozen in liquid nitrogen and stored for up to 6 months at −80◦ C. BASIC PROTOCOL 2
CHEMICAL CLEAVAGE AT CYSTEINE MPAX takes advantage of the fact that cysteine residues within a polypeptide chain can effect polypeptide cleavage upon cyanylation with 2-nitro 5-thiocyanobenzoic acid (NTCB) followed by incubation in ammonia (Fig. 26.1.2; Jacobson et al., 1973; Wu and Watson, 1998; Silverman and Harbury, 2002a; Burguete et al., 2004). In this protocol, proteins containing misincorporated cysteine residues (see Basic Protocol 1) are first unfolded with guanidine·HCl to make the sulfhydryls accessible and then incubated with NTCB. Cyanylated proteins are then precipitated with trichloroacetic acid (Arnold and Ulbrich-Hofmann, 1999). The protein backbone is cleaved upon incubation at increased pH in the presence of ammonia. Cleavage products are resolved in a Tris·Tricine gel and can subsequently be quantified on a phosphorimager. Cysteine misincorporation occurs at a very low frequency (<1 cysteine per polypeptide, on average). Therefore, this analysis yields a distinct and characteristic cleavage pattern where each band on the gel corresponds to a specific amino acid position in the protein sequence. It is useful to include a background control sample containing a protein that was expressed in the absence of misincorporator tRNA and run it parallel to the sample. Additional control reactions could be performed by
MPAX: Engineering Cysteine Probes into Proteins
Figure 26.1.2 Cyanyaltion of a cysteine sulfhydryl and subsequent base-mediated site-specific cleavage of the protein backbone.
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deleting a reagent, e.g., in absence of CNTB or ammonia, to verify a specific cleavage pattern.
Materials Cysteine-misincorporated, purified protein (see Basic Protocol 1) 10× PKA buffer (see recipe) 100 mM dithiothreitol (DTT) 1% Triton X-100 10 mg/ml bovine serum albumin 5 U/µl protein kinase A suspended in 1× protein kinase A buffer (see recipe) 10 µCi/µl adenosine triphosphate-γ-33 P (3000 Ci/mmol) 10 µM adenosine triphosphate (cold) 2 M Bicine, pH 8.6 8 M guanidine·HCl 1 M 2-nitro-5-thiocyanobenzoic acid (NTCB) in dry dioxane 0.05% (w/v) deoxycholic acid sodium salt 50% (v/v) trichloroacetic acid Acetone 8 M urea 0.1 M ammonium hydroxide (NH4 OH) 3× Tricine loading dye (see recipe) Tris·Tricine gel (see Support Protocol 1) 1.5-ml microcentrifuge tubes Vortex Speed-vacuum centrifuge 1. Combine 10 ng to 5 µg of cysteine-misincorporated, purified protein in a 1.5-ml microcentrifuge tube with the following reagents:
5 µl 10× PKA Buffer 3 µl 100 mM DTT 2 µl 1% Triton X-100 2 µl 10 mg/ml BSA 0.2 µl 5 U/µl protein kinase A suspended in 1× protein kinase A buffer H2 O to 44.5 µl Then, add: 0.5 µl 10 µCi/µl ATP-γ-33 P (3000 Ci/mmol) 5 µl 10 µM cold ATP Incubate 15 min at 37◦ C. The addition of 0.04% Triton X-100 is optional and does not interfere with subsequent analysis. Also, the use of ATP-γ -33 P is critical because it yields a sharp protein band signal. The total reaction volume is 50 µl.
2. Add an equal volume of 200 mM Bicine in 8 M guanidine·HCl, pH 8.6, and mix. Add 50 mM NTCB (5 µl) from a fresh 1 M stock in dry dioxane and mix. Incubate 5 min at room temperature. An “equal volume” in this case would be 50 µl. If the labeling reaction volume is altered, then the “equal volume” added in step 2 should be changed accordingly. Prepare the guanidine·HCl/Bicine mix just prior to addition and check the pH.
3. Add 1 ml of 0.05% sodium deoxycholic acid and mix well. Add 100 µl of 50% trichloroacetic acid, mix by inversion, and centrifuge 1 min at 14,000 × g, room temperature.
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4. Wash pellets two times with 600 µl acetone and vigorous vortexing at room temperature. Dry in a speed-vacuum centrifuge. Heating of the centrifuge is not necessary as the acetone will evaporate within minutes. It is important to remove all NTCB in this step. Residual NTCB will cause nonspecific cleavage of the protein backbone when the pH is increased with ammonium hydroxide.
5. Resuspend pellets in 10 µl of 8 M urea, 0.1 M NH4 OH. Incubate 1 hr at room temperature to cleave polypeptides. Residual TCA can reduce the pH in this step. Alternatively, the pellets can be resuspended in 10 µl of 8 M urea, 0.8 M NH4 OH. Incubate 1 hr at room temperature and open tubes to allow the NH4 OH to evaporate. Wait ∼20 min until the pH < 10. The incubation time can be used to prepare a Tris·Tricine gel (see Support Protocol 1).
6. Add 5 µl of 3× Tricine loading dye. Analyze 5 µl of resultant solution on a Tris·Tricine gel (see Support Protocol 1). BASIC PROTOCOL 3
NATIVE CYSTEINE ALKYLATION Cysteines introduced by misincorporator tRNAs (see Basic Protocol 1) can be used as structural probes to determine the degree of burial at specific positions in the native protein (Silverman and Harbury, 2002a; Burguete et al., 2004). The chemical reactivity of a cysteine sulfhydryl group with the alkylating agent iodoacetamide provides a measure of residue burial in the native protein. Solvent-exposed sulfhydryls that are alkylated resist modification by NTCB and are not subject to chemical cleavage; buried thiols are protected from alkylation by protein structure and can be revealed by subsequent denaturation and NTCB-driven cleavage (see Basic Protocol 2). This protocol describes how to monitor the kinetics of iodoacetamide alkylation at all cysteine misincorporation positions in conjunction with NTCB cleavage. A reaction in the absence of iodoacetamide provides a “zero time point” and alkylation in the presence of guanidine·HCl yields maximal alkylation or the “infinite time point.” Thus, both extremes of the kinetic reaction are represented.
Materials Labeled protein (see Basic Protocol 2, steps 1 and 2) 2 M Bicine, pH 8.6 Quench solution (see recipe) 100 mM iodoacetamide 8 M guanidine·HCl Tris·Tricine gel (see Support Protocol 1) Prepare labeled protein 1. Label 1 to 10 µg of protein according to Basic Protocol 2, steps 1 and 2, in a total volume of 150 µl. Prepare zero-time-point sample. 2. Combine 5 µl of 2 M Bicine, pH 8.6, and 75 µl water and add 20 µl radiolabeled sample. 3. Add 100 µl quench solution and store on ice. For the zero time point, native protein is incubated in the absence of iodoacetamide. Upon denaturation, all cysteines become cyanylated by NTCB and yield cleavage bands that correspond to every misincorporation position in the protein. MPAX: Engineering Cysteine Probes into Proteins
Quenched samples may be stored up to 1 week at −20◦ C (not longer due to decay of radiolabel) if not analyzed by gel electrophoresis directly after completion of the time course experiment.
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Prepare infinite-time-point sample 4. Combine 10 µl of 2 M Bicine, pH 8.6, 10 µl of 100 mM iodoacetamide, 30 µl of 8 M guanidine·HCl, and 30 µl water. Add 20 µl radiolabeled sample and incubate for at least 50 min at room temperature. It is important that the pH of the sample is 8.6. Bicine is included because the addition of guanidine·HCl might lower the pH in the sample and cause inefficient alkylation. In the so-called, infinite-time-point sample, the protein is unfolded with guanidine·HCl so that all the cysteine sulfhydryls are in contact with solvent and become alkylated. Thus, no sites are available for NTCB modification; this results in the lack of chemical cleavage and the absence of cleaved peptides on the gel.
Determine time course of protein alkylation 5. Combine 25 µl of 2 M Bicine, pH 8.6, 50 µl of 100 mM iodoacetamide, and 325 µl water. Add 100 µl radiolabeled sample. Mix and incubate at room temperature. 6. Remove 100-µl aliquots at intervals (e.g., after 1, 5, 20, and 50 min) and add 100 µl quench solution. Store quenched samples on ice. 7. At the final time point, also add 100 µl quench solution to the infinite-time-point sample. 8. Denature, cleave, and analyze all samples according to Basic Protocol 2, steps 2 to 6.
Figure 26.1.3 Representative graph and data fit for a time-course alkylation of eight cysteine substitutions of valine (V), leucine (L), or isoleucine (I) residues in TIP47. (Reprinted with permission from Burguete et al., 2004.)
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9. Determine the rate of alkylation at all misincorporated amino acid positions by fitting the data to an exponential decay function. Divide the sec−1 rate by 10−2 (10 mM iodoacetamide) to determine a rate constant in M−1 sec−1 units. Alkylation will lead to the disappearance of a given band. Thus, bands obtained in the time course experiment will appear weaker with increasing time of alkylation; the rate of their disappearance reflects their relative reactivity to iodoacetamide in the native protein (see Fig. 26.1.3). BASIC PROTOCOL 4
INTRINSIC ALKYLATION RATE The protection factor for a given cysteine residue in the folded protein is a measure of its relative burial and thus, accessibility to chemical modification. It can be determined by comparing the rates at which that residue is alkylated in both the native and the unfolded states (Silverman and Harbury, 2002a; Burguete et al., 2004). This terminology is adapted from the nuclear magnetic resonance hydrogen/deuterium exchange literature. This protocol describes how to determine the alkylation rate for fully exposed, misincorporated cysteines (see Basic Protocol 1) in proteins that are denatured in 2.4 M guanidine·HCl. The iodoacetamide concentration is kept low (0.8 mM) to achieve a logistically feasible time frame for the experiment. For the infinite time point, the iodoacetamide concentration is increased to 10 mM. Keep in mind that a ten-fold increase or decrease in iodoacetamide concentration results in a ten-fold faster or slower alkylation rate, respectively. The alkylation rate is also dependent on the pH and will be ten times slower for every pH unit below 9. It is therefore extremely important to perform the reactions at the recommended pH 8.6. Note that the observed rate reflects pseudo-first order kinetics with units of sec−1 and needs to be converted to a rate constant with the units M−1 sec−1 for comparison with alkylation rate constants of native protein. It is convenient to determine the intrinsic alkylation rate in a sample containing only a few misincorporated cysteines, at both buried and exposed positions. Similar rates of alkylation at buried and exposed sites confirm that the protein is indeed fully denatured. As a control, the rates of alkylation at the same positions should be determined in the native protein.
Materials Labeled protein 2 M Bicine, pH 8.6 8 M guanidine·HCl Quench solution (see recipe) 100 and 8 mM iodoacetamide Tris·Tricine gel (see Support Protocol 1) Label protein 1. Label 1 to 10 µg of protein according to Basic Protocol 2, steps 1 and 2, in a total volume of 150 µl. Prepare zero-time-point sample 2. Combine 10 µl of 2 M Bicine, pH 8.6, 30 µl of 8 M guanidine·HCl, and 40 µl water. Add 20 µl radiolabeled sample. MPAX: Engineering Cysteine Probes into Proteins
3. Add 100 ml quench solution and store on ice.
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Prepare infinite-time-point sample 4. Combine 10 µl of 2 M Bicine, pH 8.6, 10 µl of 100 mM iodoacetamide, 30 µl of 8 M guanidine·HCl, and 30 µl water. Add 20 µl radiolabeled sample and incubate at room temperature. Alkylate over time 5. Combine 40 µl of 2 M Bicine, pH 8.6, 40 µl of 8 mM iodoacetamide, 120 µl of 8 M guanidine·HCl, and 120 µl water. Add 80 µl radiolabeled sample. Mix and incubate at room temperature. It may be necessary to adjust the iodoacetamide concentration to fit the measurable range for the particular system studied. If cysteines remain unalkylated at the last time point, try increasing the iodoacetamide concentration slightly. If all cysteines become alkylated within the first time point, try lowering the iodoacetamide concentration. Keep in mind that a ten-fold increase or decrease in iodoacetamide concentration results in a ten-fold faster or slower alkylation rate, respectively. The alkylation rate is also dependent on the pH and will be ten times slower for every pH unit below 9. Make sure to perform the reactions at the recommended pH.
6. Remove 100-µl aliquots after 10, 30, and 60 sec and add 100 µl quench solution. Store quenched samples on ice. Prepare tubes containing 100 µl quench solution each before the experiment and keep on ice. This facilitates the addition of aliquots at the desired time point.
7. At the final time point, add 100 µl quench solution to the infinite-time-point sample. 8. Denature, cleave, and analyze all samples according to Basic Protocol 2, steps 2 to 6. 9. Determine the intrinsic rate of alkylation by averaging the decay rates of all detected cysteine probes. For example: This example illustrates the determination of the intrinsic alkylation rate constant as reported in Burguete et al. (2004). The pseudo-first order rate constants (the rate constants at a fixed iodoacetamide concentration) for cysteines at five isoleucine positions in a denatured sample were determined at 0.8 mM iodoacetamide. Residue Pseudo-first order rate constant k (sec−1 ) I333 0.018 I348 0.020 I376 0.030 I386 0.032 I425 0.038 The first order rate constant can then be calculated (I333 in this example) as: 0.018 sec−1 /(0.8 × 10−3 M) = 22.5 M−1 sec−1 The rate constants for the other residue positions in the sample and their average are then determined as follows: Residue I333 I348 I376 I386 I425
First order rate constant (M−1 sec−1 ) 22.5 25.0 37.5 40.0 47.5
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And therefore, the intrinsic rate constant (average) is as follows: (22.5 + 25.0 + 37.5 + 40.0 + 47.5)/5 = 34.5 M−1 sec−1 10. Determine a protection factor according to the following formula: Protection factor = kintrinsic /kobserved Example: Calculate the protection factor for I333: 34.5 M−1 sec−1 /1.19 M−1 sec−1 = 29 Residue I333 I348 I376 I386 I425
First order rate constant in native protein (M−1 sec−1 ) 1.19 0.18 0.25 1.85 16.67
Protection factor 29 192 138 19 2
Protection factors range from 1 (for a cysteine sulfhydryl that is fully exposed to solvent) to 104 or greater (for a cysteine sulfhydryl that is fully buried within the protein structure). BASIC PROTOCOL 5
PREDICTION OF BINDING INTERFACES When analyzing MPAX data, it is useful to plot the protection factor against residue position in the amino acid sequence (see Fig. 26.1.4). This type of graph provides useful information about protein topology and allows ready detection of buried and or solventexposed regions in the polypeptide (Burguete et al., 2004). Protection factors obtained by nuclear magnetic resonance (UNIT 17.5) hydrogen exchange measurements have been used in an analogous way in a number of studies to characterize protein topology (Raschke and Marqusee, 1998). Aromatic and hydrophobic residues are enriched at binding interfaces (Lo Conte et al., 1999). MPAX provides a rapid means to identify amino acid residues that have the potential to contribute to hydrophobic protein-protein interactions. The cores of proteins of known structures have a defined amino acid composition (Janin, 1979). Thus, different amino acids have different probabilities of being buried in a folded protein. Measured protection factors for different residue positions can be directly compared to the burial propensity of the amino acid that was replaced. This type of plot readily identifies candidate residues that are more solvent-exposed than would be expected from their occurrence in protein cores of known structures. Such residues have a high burial propensity but a low protection factor (Burguete et al., 2004).
BASIC PROTOCOL 6
SELECTION OF MUTANTS WITH ALTERED LIGAND OR PARTNER BINDING
MPAX: Engineering Cysteine Probes into Proteins
In this protocol, an ensemble preparation of proteins containing different, randomly misincorporated cysteine residues (see Basic Protocol 1) are screened as a pool for alterations in their ability to bind ligand (Burguete et al., 2004). The mutant proteins are allowed to bind to a ligand (or partner protein), which is then captured on a solid support and unbound material is washed from the column. Bound proteins are eluted and subjected to NTCB-mediated chemical cleavage (see Basic Protocol 2) and separation by gel electrophoresis (see Support Protocol 1). The eluted protein containing a mixture of individual mutants is then directly compared to the total starting protein preparation.
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Figure 26.1.4 Protection factor map of 48 cysteine substitutions in TIP47 replacing valine, leucine, isoleucine, phenylalanine, and glutamine residues. (Reprinted with permission from Burguete et al., 2004.)
Individual cysteine mutants with increased or decreased ligand binding, as monitored by their enrichment or loss from the ligand-bound fraction, are readily detected in this comparison. This protocol requires that the ligand be coupled to a solid support using a different means than that used to purify mutant proteins. For example, the misincorporated protein could bear a 6-histidine tag for purification purposes while the binding partner could be immobilized using an antibody epitope tag. It is important to verify the biochemical properties of selected mutant proteins. For this purpose, individual mutants can be prepared by conventional site-directed mutagenesis of the wild type gene. The resulting protein preparation, in which every molecule carries the same mutant residue, can then be analyzed for binding to known ligands or partners by established protocols to confirm the MPAX findings. In addition, the folded nature and overall secondary structure can be confirmed by circular dichroism or other methods.
Materials Radiolabeled protein (see Basic Protocol 2) Ligand or partner protein Solid support Binding buffer 4 M guanidine·HCl 100 mM sodium Bicine, pH 8.6 1 M 2-nitro-5-thiocyanobenzoic acid (NTCB) in dry dioxane Tris·Tricine gel (see Support Protocol 1) Scintillation counter
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1. Radiolabel protein according to Basic Protocol 2, steps 1 and 2. 2. Bind to ligand or partner protein according to established protocol. 3. Bind ligand/protein complex to a solid support according to individual protocol. Glutathione S-transferase (GST) is a commonly used tag for immobilization and/or purification of proteins. The tagged protein is supported on glutathione agarose beads according to the manufacturer’s instructions. A GST tag has been used successfully in this approach (Burguete et al., 2004). In some cases, it may be preferable to invert the order of steps 2 and 3, depending upon the specific application.
4. Wash in binding buffer to remove unbound protein. 5. Elute in 4 M guanidine·HCl and 100 mM sodium Bicine, pH 8.6. The protein can alternatively be eluted according to other protocols and chemical cleavage performed according to Basic Protocol 2, steps 2 to 6.
6. Add 50 mM NTCB from a fresh 1 M stock in dry dioxane. Incubate 5 min at room temperature. 7. Perform Basic Protocol 2, steps 3 to 6. 8. Determine radioactivity in eluted protein and total protein in a β-scintillation counter. Load equal counts of eluted and total protein on a Tris·Tricine gel (see Support Protocol 1). 9. Quantify band intensities corresponding to different amino acid residue positions and determine signal ratios internally within a single sample. It is useful to identify a reference band that does not change in intensity between samples (assuming equal loading of different samples in different lanes). The relative intensity of other bands in comparison with the reference band in a given lane is then determined as a percentage of the reference band intensity.
10. Compare relative band intensities from eluted and total protein samples. A mutant that binds better to ligand will be overrepresented in the bound fraction and the ratio of eluted/total mutant protein will be >1. Alternatively, if a cysteine mutant displays decreased ligand affinity, the corresponding band will be underrepresented in the bound fraction and the ratio of eluted/total mutant protein will be <1. Specific cysteine mutants can then be prepared individually by conventional site-directed mutagenesis of the wild type gene for further characterization. SUPPORT PROTOCOL 1
TRIS·TRICINE GEL TO RESOLVE SMALL PEPTIDES FROM MPAX EXPERIMENTS This protocol makes use of Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis with a discontinuous buffer system to give high-resolution separation of small peptides (Schagger and von Jagow, 1987; also see Alternate Protocol 1 of UNIT 10.1). It is highly recommended to use this protocol to analyze samples from Basic Protocols 2, 3, 4, and 6. This gel consists of four different layers, which include resolving, intercalating, stacking, and comb gels that are essential for the resolution of peptides that only differ in a few amino acids in size.
MPAX: Engineering Cysteine Probes into Proteins
CAUTION: Note that the voltages and currents used during electrophoresis are dangerous and potentially lethal. It is extremely important to read UNIT 10.1 to understand how to run a gel and to follow the “Safety Considerations” outlined under “Electricity and Electrophoresis.”
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Materials 1.5% (w/v) agarose solution (see recipe) 40% (v/v) 19:1 acrylamide 3× gel buffer (see recipe) 3× comb buffer (see recipe) Glycerol 1% (w/v) Coomassie brilliant blue G250 10% (w/v) ammonium persulfate TEMED Butanol 1× cathode buffer (see recipe) 10× anode buffer (see recipe) Glass plates (20 × 20–cm and 18 × 20–cm) Spacers (0.5-mm) Clamps Whatman filter paper Comb (0.5-mm; 19 wells ∼0.5-cm wide) Gel-running apparatus Capillary pipet tips Power supply Plastic wrap Heated vacuum gel dryer Phosphor image plate and casette Phosphorimager Prepare Tris·Tricine gel 1. Pre-assemble two scrupulously clean glass plates with 0.5-mm spacers on the side and bottom edges. Hold in place with clamps. 2. Heat 1.5% agarose solution in a microwave without lid for 1 min or until boiling. Cool for 1 min. 3. Use a glass pipet to seal the spacers by applying warm 1.5% agarose along the outside edges. The warm agarose will enter the space by capillary force.
4. Combine resolving gel components (see Table 26.1.1) and pour quickly into the preassembled gel sandwich until 3.5 cm from top edge. Carefully add a layer of neat butanol (∼ 2 mm) and leave for 10 min or until fully polymerized. Add ammonium persulfate and TEMED just prior to pouring. Butanol provides a barrier to oxygen, which will inhibit polymerization. Do not use propanol or ethanol as these tend to mix with the gel solution.
5. Rinse with water to remove the butanol. Hold gel upside down and insert a small piece of filter paper between the glass plates parallel to one of the spacers to absorb remaining water. Be careful not to disturb the gel surface. 6. Repeat steps 4 and 5 for intercalating and stacking layers. Make these layers each ∼1 cm high. 7. Combine comb-gel reagents and fill the rest of the chamber. 8. Quickly insert the comb as close to the stacking layer as possible without disturbing it. Avoid trapping air bubbles. Protein Engineering
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Table 26.1.1 Comb, Stacking, Intercalating, and Resolving Gel Components
Order of layers
4 (top)
3
2
1 (bottom)
Reagent 40% 19:1 acrylamide 3× gel buffer 3× comb buffer Glycerol dH2 O 1% Coomassie brilliant blue G250 Add just prior to pouring: 10% ammonium persulfate TEMED
Comb 1.0 ml – 1.7 ml – 2.3 ml 15 µl
Stacking 0.5 ml 1.2 ml – – 3.3 ml –
Intercalating 1.3 ml 1.7 ml – 0.5 ml 1.5 ml –
Resolving 8.0 ml 6.0 ml – 1.8 ml 2.2 ml –
50 µl 7 µl
50 µl 7 µl
50 µl 7 µl
100 µl 20 µl
9. Allow the comb gel to polymerize (∼10 to 20 min) at room temperature and then remove bottom spacer. Keep side clamps attached while removing bottom spacer to inhibit accidental displacement of the plates.
Run and dry gel 10. Place gel in running apparatus and fill the cathode (upper) chamber with 1× cathode buffer. Fill the anode (lower) chamber with 1× anode buffer. 11. Remove comb and carefully rinse wells with a pipet. 12. Load 5 to 10 µl of sample at the bottom of each well with a capillary pipet tip. Loading larger volumes will result in band thickening and suboptimal resolution. Loading samples in every other well will increase resolution.
13. Run gel at 145 V, 145 mA, and 100 W for 16 hr or until dye front reaches bottom of gel. 14. Transfer the gel to a piece of Whatman filter paper and cover with plastic wrap. Dry the gel on a heated vacuum gel dryer for 2.5 hr at 75◦ C. Cool to room temperature before breaking the vacuum. Position the gel sandwich on a flat surface with the smaller glass plate facing up. Insert a metal spatula 0.5 cm into the comb gel and gently split the two glass plates apart. Carefully remove the small plate (the gel usually sticks to the bottom, large plate). Cover gel with a 30 × 30–cm piece of Whatman filter paper. Do not apply pressure on the filter paper as the gel will attach to the glass plate. Turn the sandwich upside down. Carefully remove the large glass plate and apply a 30 × 30–cm piece of plastic wrap onto the gel. Avoid trapping air bubbles. Cut off excess paper at edges and dry the gel in a preheated dryer to avoid cracking the gel.
15. Expose the dried gel on an image plate overnight to 2 days and quantify bands on a phosphorimager. SUPPORT PROTOCOL 2
MPAX: Engineering Cysteine Probes into Proteins
GENE ASSEMBLY This protocol describes a procedure to assemble a synthetic gene (Stemmer et al., 1995) containing codons complementary to those of specific misincorporator tRNAs (see Basic Protocol 1 and Table 26.1.2). Misincorporation seems to follow normal wobble base pairing rules. As an example, the Val(GAC) tRNA misincorporates cysteine at the same frequency at GUC or GUU codons, but not at GUG or GUA codons (Silverman and Harbury, 2002a). Hence, if a wild-type gene is co-expressed with the Val(GAC) tRNA, cysteine will be misincorporated at fewer than all possible valine positions.
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Table 26.1.2 Suggested Codon Usage for Misincorporation Proton-Alkyl Exchange
Misincorporator tRNA anticodon
RNA codon
Codon usage for DNA gene assembly
Tyr
GUA
UAC
TAC
Ile
GAU
AUC
ATC
Val
GAC
GUC
GTC
Leu
GAG
CUC
CTC
His
GUG
CAC
CAC
Phe
GAA
UUC
TTC
Trp
CCA
UGG
TGG
Ser
GCU
AGC
AGC
Asp
GUC
GAC
GAC
Ala
AGC
GCU
GCT
Arg
ACG
CGU
CGT
Thr
AGU
ACU
ACT
Asn
GUU
AAC
AAC
Gln
UUG
CAA
CAA
Glu
UUC
GAA
GAA
Gly
ACC
GGU
GGT
Lys
UUU
AAA
AAA
Pro
AGG
CCU
CCT
Amino acid
Maximal misincorporation can be achieved by expressing a gene carrying exclusively codons that match those of the co-expressed misincorporator tRNAs. This protocol describes the design and assembly of such a gene from oligonucleotides using the polymerase chain reaction (PCR). Note that synthesis and purification of oligos (steps 2 through 16) can be omitted by ordering full-length gel-purified oligos from a synthesis facility. The assembled gene is cloned into either of two vectors, pET28a pH6 PKA or pET28a PKA pH6 (Harbury laboratory, Stanford University), using the restriction sites EcoRI and HindIII or BamHI and HindIII, respectively. The restriction enzymes used in this protocol are intended for cloning into the pET28a pH6 PKA vector. Reading APPENDIX 4E of this manual and UNITS 2.1A, 2.11, & 2.12 in Current Protocols in Molecular Biology (CPMB) for directions on purification of DNA and synthesis of oligonucleotides is recommended. In addition, APPENDIX 4I and APPENDIX 4J of this manual and UNITS 3.1, 3.16, & 3.17 in CPMB can be consulted for instructions on how to digest DNA with restriction endonucleases, and to subclone and amplify DNA. CAUTION: Voltages and currents used during electrophoresis are dangerous and potentially lethal. Before undertaking this protocol, read UNIT 10.1 to understand how to run a gel and follow the “Safety Considerations” outlined under “Electricity and Electrophoresis.”
Materials Sense and antisense oligonucleotides (5-nmol scale) Ammonia Formamide loading dye (see recipe)
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Urea-acrylamide gel (see Support Protocol 3) 1× TBE (see recipe) Ethidium bromide solution in TBE (see recipe) Sep-Pak C18 cartridge (Vac 1 cc, 100 mg; Waters) 50% (v/v) acetonitrile 2 M and 100 mM triethylammonium acetate EcoRI and HindIII restriction enzymes pET28a pH6 PKA plasmid modified to provide an N-terminal histidine tag and a C-terminal protein kinase A recognition site 2-ml screw-cap tubes 50◦ C incubator Speed vacuum centrifuge Power supply Metal spatula Low-intensity UV source Scalpel Metal spatula Rocker Glass fiber paper (Whatman) Additional reagents and equipment for chloroform extraction and ethanol precipitation (APPENDIX 4E) Design and deprotect oligonucleotides 1. Design a gene using a single codon for each amino acid according to Table 26.1.2 and replace any cysteine codons with GCT (alanine). Avoid any EcoRI and HindIII restriction sites as these will interfere with later cloning procedures. The amino acid codons in Table 26.1.2 match the anticodons in the mutant cysteine misincorporator tRNAs used for co-expression. It is recommended that cysteine codons be removed, as they may complicate the biochemical readout of cysteine misincorporation (see Strategic Planning and Critical Parameters).
2. Synthesize sense and antisense oligonucleotides on a 5-nmol scale), 40 bp long, and with a 20-bp overlap to cover the entire gene. 3. Resuspend each oligonucleotide in 40 µl water and transfer 2 µl to a 2-ml screw-cap tube. 4. To deprotect the oligonucleotides, add 0.5 ml ammonia, cap tightly, and incubate overnight at 50◦ C. 5. Cool to 22◦ C and evaporate the ammonia in a speed vacuum centrifuge.
Isolate full-length oligonucleotides 6. Resuspend oligonucleotides in 7 µl formamide loading dye. 7. Pre-run a urea-acrylamide gel for 45 min at 600 V in 1× TBE. 8. Load oligonucleotides in separate lanes and run gel at 600 V for ∼1 hr or until the loading dye runs out from the bottom of the gel. 9. Turn off the power supply and carefully lift the front glass plate with a metal spatula. 10. Carefully transfer the gel to a tray containing ethidium bromide in 1× TBE and incubate 15 min on a rocker to stain DNA. MPAX: Engineering Cysteine Probes into Proteins
11. Visualize DNA with a low-intensity UV source and excise and pool the top, fulllength oligonucleotides with a clean scalpel.
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Avoid exposing unprotected skin and eyes to intense UV sources. If the UV light is aimed upwards, wear a UV protective face shield when standing near the source.
12. Homogenize gel pieces in 2 ml water with a metal spatula, place in a screw-cap tube and rotate overnight at 37◦ C. 13. Filter through Whatman glass fiber paper and collect liquid.
Desalt oligonucleotides 14. Wash a Sep-Pak C18 cartridge with 2 ml of 100% acetonitrile and then with 2 ml of 2 M triethylammonium acetate. 15. Load sample slowly and reapply flow-through onto the column several times. 16. Wash with 2 ml of 100 mM triethylammonium acetate, 1 ml water, and elute with 1 ml of 50% acetonitrile.
Assemble gene 17. Use the desalted primer mix to assemble the gene in a PCR reaction (APPENDIX 4J). Set up the thermal cycler using the following parameters: 36 cycles:
30 sec 40 sec 60 sec
94◦ C 42◦ C 72◦ C.
When analyzed by agarose gel electrophoresis (APPENDIX 4F) the product should appear as a smear ranging from small to large sized DNA.
18. Reamplify the PCR product in a second reaction generated by 5 and 3 end primers containing restriction sites for EcoRI and HindIII, respectively, using the following parameters: 36 cycles:
30 sec 40 sec 60 sec
94◦ C 52◦ C 72◦ C.
Agarose gel electrophoresis analysis should reveal a blurry band around the predicted size of the gene.
19. Chloroform extract and ethanol precipitate the second PCR product (APPENDIX 4E). 20. Cleave concatamers with restriction endonucleases EcoRI and HindIII (APPENDIX 4I) to generate a single gene assembly product. Agarose gel electrophoresis analysis should reveal a single sharp band of the predicted size.
21. Clone the assembled gene into a pET28a pH6 PKA plasmid, modified to provide an N-terminal histidine tag and a C-terminal protein kinase A recognition site (Silverman and Harbury, 2002a), generating pH6 GENE PKA.
UREA-ACRYLAMIDE GEL This protocol provides a method to purify oligonucleotides by use of a urea-acrylamide gel.
SUPPORT PROTOCOL 3
Materials Urea-acrylamide gel (see recipe) TEMED Glass plates, spacers, and combs (see Support Protocol 1)
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1. Assemble glass plates according to Support Protocol 1, steps 1 to 3. 2. Mix and pour urea-acrylamide gel quickly into the preassembled gel-sandwich. 3. Quickly insert a 0.5-mm comb with ∼ 0.5-cm wide wells. 4. Allow the gel to polymerize (∼10 to 20 min) at room temperature and then remove bottom spacer.
REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Agarose solution, 15% (w/v) 1.5 g agarose Add H2 O to 100 ml Heat to dissolve Store ≤1 week at room temperature Anode buffer, 10× 242 g Tris base (2 M final) Add H2 O to <1 liter Adjust pH to 8.8 with HCl Correct volume to 1 liter with H2 O Store ≤3 months at room temperature Cathode buffer, 1× 12 g Tris base (100 mM final) 18 g Tricine (100 mM final) 1 g SDS (1 g/liter final) Add H2 O to 1 liter Store ≤3 months at room temperature Comb buffer, 3× 36 g Tris base (0.3 M final) 54 g Tricine (0.3 M final) 3 g SDS (3 g/liter final) Add H2 O to 1 liter Store ≤3 months at room temperature Ethidium bromide solution in TBE 4 µl ethidium bromide (10 mg/ml in water) 100 ml 1× TBE (see recipe) Prepare fresh for each experiment CAUTION: Ethidium bromide is a powerful mutagen and moderately toxic. Gloves should be worn when working with solutions that contain this dye.
MPAX: Engineering Cysteine Probes into Proteins
Formamide loading dye 0.2 ml 0.5 M EDTA (10 mM final) 0.1% (w/v) bromphenol blue 0.1% (w/v) xylene cyanol FF 10 ml deionized formamide Store ≤4 months at room temperature
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Gel buffer, 3× 363 g Tris base (3 M final) 3 g SDS (3 g/liter final) Add H2 O to <1 liter Adjust pH to 8.6 with HCl Correct volume to 1 liter with H2 O Store ≤ 3 months at room temperature M63 medium containing 30 µg/ml kanamycin and 100 µg/ml carbenicillin 14 g (NH4 )2 SO4 (100 mM final) 2 g glucose (11 mM final) 100 µl 0.5% (w/v) thiamine HCl in H2 O (15 mM final) 1 ml 1 M MgSO4 (1 mM final) 1 mg FeSO4 (0.1% w/v final) 50 ml 1 M KH2 PO4 pH 7.5 (50 mM final) 1 ml 30 mg/ml kanamycin (30 µg/ml final) 1 ml 100 mg/ml carbenicillin (100 µg/ml final) Add H2 O to 1 liter; filter sterilize Store ≤2 days at room temperature Phosphate buffered saline (PBS), 10× 160 g NaCl (1.37 M final) 4 g KCl (26.8 mM final) 4 g KH2 PO4 (14.7 mM final) 43.4 g Na2 HPO4 ·7H2 O (80.9 mM final) Add H2 O to 2 liters Adjust pH to 7.5 Store ≤3 months at room temperature if sterile Protein kinase A buffer, 10× 500 µl 1 M Tris·Cl, pH 7.5 (0.5 M final) 100 µl 1 M MgCl2 (0.1 M final) 400 µl H2 O Store ≤12 months at −20◦ C Quench solution 25 mM β-mercaptoethanol 0.1 mg/ml BSA Prepare fresh for each experiment TBE, 10× 215.7 g Tris base (0.89 M final) 110 g Boric acid (0.89 M final) 14.9 g Na2 EDTA·2H2 O (0.02 M final) Add H2 O to just <2 liters Adjust pH to 8.2 to 8.5 Correct volume to 2 liters with H2 O Store ≤3 months at room temperature
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Tricine loading dye, 3× 250 µl 4 M Tris·Cl, pH 6.5 (1 M final) 250 µl glycerol (25% v/v final) 480 µl 10% (w/v) SDS (4.8% w/v final) 20 µl 1% (w/v) Coomassie brilliant blue G250 (0.02% w/v final) Store ≤12 months at −20◦ C Urea-acrylamide gel 12.6 g urea (7 M final) 3 ml 10× TBE (see recipe) 11.25 ml acrylamide, 19:1 (40% w/v) Add H2 O to 30 ml Then add: 150 µl 10% (w/v) ammonium persulfate in H2 O (0.05% w/v final) 15 µl TEMED (0.05% v/v final) Prepare fresh for each experiment COMMENTARY Background Information
MPAX: Engineering Cysteine Probes into Proteins
Methods for the identification of protein interaction domains often rely on truncated or mutated recombinant proteins. A disadvantage of this approach is that binding interfaces represent three-dimensional surfaces and cannot be fully identified using protein truncations or point mutations. Knowledge of the threedimensional structure of the proteins under study is helpful, but in the absence of a cocrystal of two binding partners, a significant amount of work is still required to define binding interfaces. Previous methods for mapping protein structure through the measurement of amino acid side chain solvent accessibility were based on acylation of lysine residues, oxidation of methionine residues, and alkylation of cysteine residues (Doonan and Fahmy, 1975; de Arruda et al., 1992; Hanai and Wang, 1994; Doering and Matsudaira, 1996; Tu and Wang, 1999). However, data collection was limited to a few naturally occurring probes. Recently, a technique named misincorporation protonalkyl exchange (MPAX) was developed, which makes use of translational misincorporation to introduce multiple structural probes into a protein structure (Silverman and Harbury, 2002a). This method was named analogous to nuclear magnetic resonance hydrogen/deuterium exchange and makes use of the reactivity of cysteine sulfhydryls to an alkylating reagent as a reporter of protein structure. The approach described here makes use of MPAX to rapidly identify surface hydrophobic residues that contribute to protein-protein interaction interfaces. Small quantities (micrograms) of protein are required to perform all
of the protocols described in this unit. This, in combination with the short time required for each assay, makes the method a highly attractive complement to high-resolution structural techniques such as X-ray crystallography and nuclear magnetic resonance. The technique is not limited by protein size or solubility. It has also proven very useful for proteins that are difficult to crystallize (Burguete et al., 2004). A central advantage lies in the rapid generation of a complete set of single amino acid mutant proteins that can be analyzed simultaneously for alterations in ligand binding or partner protein interaction. Mutation to a cysteine residue does not generally disrupt normal protein topology because cysteine is small and amphiphilic and can easily be accommodated on the surface or within the core of folded proteins. The relative burial of cysteine residues in folded proteins correlates well with the expected burial of the particular amino acids that were replaced (Silverman and Harbury, 2002a,b; Burguete et al., 2004). A further advantage of this technique, compared with crystallography, is that it detects the dynamics of a molecule in solution rather than a single, immobile, structural form. Unstructured regions of the protein and/or flexible loops are fully “visible” with this technology (Burguete et al., 2004). Moreover, residues that seem buried in a crystal structure may unexpectedly exchange with solvent due to protein dynamics in solution. An alternative use of MPAX for detection of protein interfaces has been described (Silverman and Harbury, 2002a). Amino acids in binding interfaces were detected through direct protection from solvent by ligand binding.
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Such residues displayed a slowed alkylation rate in the presence of ligand as compared to reactions carried out in its absence. It should be possible to apply this method to the detection of conformational changes, in the presence or absence of cofactors and/or ligands. In addition, Basic Protocols 2 and 3 can be performed in the presence of any number of additional proteins that do not contain a PKA recognition site. The methods described here are thus also applicable to large protein complexes, partially folded proteins, and membrane proteins. With regard to the specific chemistry utilized herein, solvent-accessible cysteine thiols are reactive to alkylation by iodoacetamide. The alkylating reagent is quenched with a 2.5fold molar excess of β-mercaptoethanol together with BSA. Non-alkylated cysteines that are buried in the protein can be detected by unfolding in guanidine·HCl and subsequent reaction with 2-nitro-5-thiocyanobenzoic acid (NTCB). A four-fold molar excess of NTCB over β-mercaptoethanol is used to cyanylate cysteine sulfhydryls that have not been selectively alkylated. Subsequent exposure to pH 8 to 9 results in cleavage at the amino acid that has been modified by NTCB (see Fig. 26.1.2; Jacobson et al., 1973).
Critical Parameters While the general MPAX strategy of replacing hydrophobic residues with cysteine residues does not appear to significantly disturb protein stability, the effect of replacing conserved cysteine residues with alanine may be large in some cases (Silverman and Harbury, 2002b). It is, therefore, recommended to avoid replacing conserved cysteine codons when preparing the gene of interest (see Strategic Planning). A protein of interest must be expressed in minimal medium to achieve efficient misincorporation. Furthermore, it is critical to use ATP-γ-33 P to obtain sharp protein bands upon phosphorimaging. The accuracy of the iodoacetamide concentration is critical to obtain reproducible results and to compare rate constants determined at different concentrations (native versus intrinsic rates). The alkylation rate is directly proportional to the iodoacetamide concentration (a ten-fold increase in concentration results in a ten-fold faster alkylation rate and vice versa). Because the cysteine alkylation chemistry is pH-sensitive (it requires the deprotonated form of the sulfhydryl, pKa ∼9), it is critical to perform the assay at the indicated pH. In addition, it is important to exclude any
reagents containing free sulfhydryls in significant concentrations, as they will quench the alkylating reagent.
Troubleshooting Nonspecific background backbone cleavage If excess NTCB has not been completely removed before raising the pH of the protein solution with ammonia, the NTCB will cleave the protein backbone nonspecifically. It is important to remove excess NTCB fully during the TCA precipitation, if necessary, with additional acetone washes. Vortexing for prolonged periods of time can help dissolve the TCA pellet. Autophosphorylation of PKA and subsequent nonspecific cleavage can cause background bands in the absence of a PKA substrate. Include a control sample with protein expressed in the absence of misincorporator tRNA to verify that no spurious bands are generated. Always include BSA in labeling reactions. Full length PKA and BSA do not enter the resolving gel and will not interfere with the analysis. Bands appear at the infinite alkylation time point Addition of guanidine·HCl in the infinite time point can lower the pH of the solution. The Bicine concentration is therefore doubled in samples that are alkylated in the presence of guanidine·HCl (see Basic Protocols 3 and 4). It is important to check the solution pH after guanidine·HCl has been added. Chemical cleavage is not efficient TCA that remains in the pellet after the acetone washes in Basic Protocol 2, step 5, might reduce the pH in the cleavage reaction. If this is a problem, the pellets can be resuspended in 10 µl of 8 M urea, 0.8 M ammonia in step 5. After cleavage, the ammonia is evaporated in open tubes before the samples are loaded onto the Tricine gel. High ammonia concentrations in samples will interfere with gel analysis. Band intensities cannot be fitted to exponential decay functions This is a typical result of loading unequal protein quantities from samples corresponding to different time points. Analyze equal amounts of radioactivity for different samples on the Tris·Tricine gel. The signals from individual bands can also be normalized for total lane intensity to facilitate comparison.
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Bands appear blurry It is critical to label the protein with 33 P (and not 32 P) to obtain the sharpest possible band signals from peptides that are resolved in the Tris·Tricine gel. In addition, this problem can be addressed by carefully loading a small volume (∼5 µl) of sample into the bottom of the well of the Tris·Tricine comb gel. Avoid getting sample on the side of the wells. This can be done using a capillary pipet tip. Dry the gel immediately after peptide separation. Make sure that gel interfaces are smooth and that air bubbles are not trapped in the gel. Since any movement of the gel on the image plate results in a distortion of the signal, it is recommended to tape the dried gel onto the image plate.
Anticipated Results Cysteine misincorporation and chemical cleavage at cysteine are highly reproducible techniques. The variation of measured alkylation rates between individual experiments is very small. MPAX in combination with affinity chromatography has previously been used to identify residues in binding interfaces and to detect surface hydrophobic residues that are likely candidates for protein-protein interaction (Burguete et al., 2004). Presuming that the protein of interest can be purified in microgram quantities, these methods are very likely to reveal amino acids that contribute to proteinprotein interaction.
Time Considerations
MPAX: Engineering Cysteine Probes into Proteins
Preparation of LB plates and medium prior to starting Basic Protocol 1 should take no longer than 1 to 2 hr. Bacterial expression of protein together with misincorporator tRNAs in minimal medium requires approximately the same time as expression without misincorporator tRNAs in rich medium. This includes transformation of E. coli and plating onto agar on day 1, which can be done in 1.5 hr. Inoculation of LB medium by individual bacterial colonies on day 2 should take no longer than 20 min. On day 3, cells are inoculated into minimal medium (20 min) and grown for 4 to 6 hr. Induction with IPTG takes 10 min and the expression time varies according to individual protocols and can be 1 hr to overnight. The duration of purification on day 4, again, varies according to individual protocols. As an example, a normal histidine-tagged protein can be purified in 4 to 6 hr and dialysis can be set up to proceed overnight. Thus, the total handson time for preparation of protein containing
misincorporated cysteine should be no longer than 11 hr within a total time frame of 4 days. Chemical cleavage at cysteine positions is typically performed within 3 hr, with the addition of 1.5 hr for the native cysteine alkylation protocol or the intrinsic alkylation rate protocol (see Basic Protocols 2 to 4). Tris·Tricine gels are prepared in 1 to 2 hr depending on the time required for polymerization (see Support Protocol 1). Gel set-up, counting of samples, and loading requires ∼2 hr and gels are run overnight. The total drying time for gels is ∼3.5 hr on the subsequent day. Thus, MPAX assays require 1 full day of hands-on time with the addition of a total of 20 min on day 2 for gel drying. The exposure time for radioactive gels on image plates can range from a few hours to 2 days, depending on signal intensity. Data analysis including gel scanning, band quantification, data fitting using software, and determination of protection factors can take 1 to 3 days, depending on the skill and experience of the investigator. The protocol describing how to select residues that contribute to binding interfaces (see Basic Protocol 6) can be performed in 1 day, assuming that purified proteins are available and with the addition of gel-processing time as described above. Budget ∼2 weeks for the gene assembly process (see Support Protocols 2 and 3), excluding the time required for oligonucleotide synthesis. Additional time will be required for correction of any point mutations detected in the assembled gene, sequencing, and cloning. Finally, plan 1 to 3 weeks for designing a gene that will be useful for the planned experiments considering the points raised under Strategic Planning.
Literature Cited Arnold, U. and Ulbrich-Hofmann, R. 1999. Quantitative protein precipitation from guanidine hydrochloride-containing solutions by sodium deoxycholate/trichloroacetic acid. Anal. Biochem. 271:197-199. Burguete, A.S., Harbury, P.B., and Pfeffer, S.R. 2004. In vitro selection and prediction of TIP47 protein-interaction interfaces. Nat. Methods 1:55-60. de Arruda, M.V., Bazari, H., Wallek, M., and Matsudaira, P. 1992. An actin footprint on villin. Single site substitutions in a cluster of basic residues inhibit the actin severing but not capping activity of villin. J. Biol. Chem. 267:13079-13085. Doering, D.S. and Matsudaira, P. 1996. Cysteine scanning mutagenesis at 40 of 76 positions in villin headpiece maps the F-actin binding site
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Doonan, S. and Fahmy, H.M. 1975. Specific enzymic cleavage of polypeptides at cysteine residues. Eur. J. Biochem. 56:421-426.
Tu, B.P. and Wang, J.C. 1999. Protein footprinting at cysteines: Probing ATP-modulated contacts in cysteine-substitution mutants of yeast DNA topoisomerase II. Proc. Natl. Acad. Sci. U.S.A. 96:4862-4867.
Hanai, R. and Wang, J.C. 1994. Protein footprinting by the combined use of reversible and irreversible lysine modifications. Proc. Natl. Acad. Sci. U.S.A. 91:11904-11908.
Wu, J. and Watson, J.T. 1998. Optimization of the cleavage reaction for cyanylated cysteinyl proteins for efficient and simplified mass mapping. Anal. Biochem. 258:268-276.
Janin, J. 1979. Surface and inside volumes in globular proteins. Nature 277:491-492.
Key References
Jacobson, G.R., Schaffer, M.H., Stark, G.R., and Vanaman, T.C. 1973. Specific chemical cleavage in high yield at the amino peptide bonds of cysteine and cystine residues. J. Biol. Chem. 248:6583-6591.
This is the first “blind” study where protein structure is mapped using the MPAX protocol and subsequently compared to crystallographic data. A method to select for residues that contribute to protein-interaction interfaces is developed and MPAX technology is also used to enable prediction of residues that lie in protein-protein interfaces. Residues that are highly likely to contribute to partner protein binding are selected by screening a population of different cysteine misincorporation mutants in a single affinity chromatography step.
and structural features of the domain. Biochemistry 35:12677-12685.
Lo Conte, L., Chothia, C., and Janin, J. 1999. The atomic structure of protein-protein recognition sites. J. Mol. Biol. 285:2177-2198. Raschke, T.M. and Marqusee, S. 1998. Hydrogen exchange studies of protein structure. Curr. Opin. Biotechnol. 9:80-86. Schagger, H. and von Jagow, G. 1987. Tricinesodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa. Anal. Biochem. 166:368-379. Silverman, J.A. and Harbury, P.B. 2002a. Rapid mapping of protein structure, interactions, and ligand binding by misincorporation proton-alkyl exchange. J. Biol. Chem. 277:30968-30975. Silverman, J.A. and Harbury, P.B. 2002b. The equilibrium unfolding pathway of a (beta/alpha)8 barrel. J. Mol. Biol. 324:1031-1040. Stemmer, W.P., Crameri, A., Ha, K.D., Brennan, T.M., and Heyneker, H.L. 1995. Single-step assembly of a gene and entire plasmid from large numbers of oligodeoxyribonucleotides. Gene 164:49-53.
Burguete et al., 2004. See above.
Silverman and Harbury, 2002a. See above. MPAX technology to footprint protein structure at a single amino acid resolution is developed and described here for the first time. The substrate-binding site for triosephosphate isomerase (β/α)8 barrel is accurately mapped and the stability of the barrel is determined. In addition, a mass spectrometry method to measure the alkylation rate at misincorporated cysteines is described.
Contributed by Alondra Schweizer Burguete, Pehr B. Harbury, and Suzanne R. Pfeffer Stanford School of Medicine Stanford, California
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Combinatorial Recombination of Gene Fragments to Construct a Library of Chimeras
UNIT 26.2
This unit contains protocols that allow creation of libraries of chimeras through combinatorial assembly of gene fragments (Fig. 26.2.1). The experimenter specifies where recombination is allowed to occur, giving the method several advantages over annealingbased and truncation-based methods for chimeragenesis. Among these advantages are that many genes of any identity can be recombined with up to eight recombination sites, and no frame-shifted chimeras are produced. Additionally, unlike total gene synthesis using oligonucleotides, many different chimeras are made simultaneously, it is easy to create full-length genes with few or no point mutations, and fewer oligonucleotides are required for the procedure, reducing the cost of constructing chimeras. Furthermore, it is easy to modify the protocol to incorporate desirable insertions, deletions, or rearrangements in the genes being recombined. Combinatorial recombination is achieved by assembling blocks of sequence with specific overhangs, or sticky ends. A “block” is a portion of sequence that has a particular set of overhangs and position in the gene; it may be from any parent. A “fragment” is a block from a specific parent gene (Fig. 26.2.1). The overhangs permit even unrelated fragments to be ligated in the correct order while allowing independent assortment of fragments with identical overhangs. Genes of any identity can be recombined so long as they share 3 to 5 base pairs of identity at the desired recombination site. The Basic Protocol focuses on the mechanistic details necessary for creating a combinatorial gene library. The Basic Protocol limits the size of the blocks and the number of recombination sites. It is also intended to incorporate fragments from each parent gene at all blocks. Alternate protocols have been provided to circumvent these limitations. Choosing the recombination sites effectively, however, is a key part of a successful experiment and is discussed in the Commentary.
STRATEGIC PLANNING Block Size Choosing the number and locations of recombination sites is an essential component of the experimental design, and a more detailed discussion of how to choose recombination sites is provided in the Commentary. However, a few details resulting from these choices affect the construction methodology and are discussed here. First, the smallest block should be larger than ∼40 bp for the standard procedure to work correctly. Blocks smaller than this are lost during the purification steps that remove the tag sequences (∼30 bp). However, Alternate Protocol 1 provides options for using blocks smaller than 40 bp. Number of Recombination Sites The complexity of the procedure increases significantly with the number of recombination sites. This protocol is based on ligation reactions, which are inherently inefficient. The authors have found it difficult to ligate more than four DNA pieces together in a single reaction and achieve sufficient yield. Therefore, in order to have more than four independent blocks, two separate ligation steps are performed with a cloning step between them. Choosing four or eight blocks takes the best advantage of these two ligation steps. Choosing a different number of blocks results in fewer than four blocks in one
Contributed by Michelle M. Meyer, Kaori Hiraga, and Frances H. Arnold Current Protocols in Protein Science (2006) 26.2.1-26.2.17 C 2006 by John Wiley & Sons, Inc. Copyright
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Figure 26.2.1 By combinatorially recombining three genes with eight independently assorting blocks 38 = 6561 possible chimeras can be formed in a single experiment. Each positional block of sequence is numbered 1 to 8, and at each block there are three possible fragments labeled 1 to 3. The designation 1.3 refers to the first block with the fragment from parent three.
or more of the mini-libraries. Using four or fewer recombination sites allows a significantly shorter procedure, where the final construction phase can be omitted. Alternate Protocol 2 details a protocol that incorporates more than eight blocks for using more recombination sites.
Restriction Sites The restriction sites used for cloning during construction and for the final cloning of the library should be different. This can be achieved by using a different plasmid for the construction steps, or by using a different set of restriction sites for construction. Using different sites prevents cross-ligation during the final ligation phase of construction. It also allows the mini-libraries to be inversely cloned into plasmids that have leaky or constitutive expression without fear that low-level expression of the mini-libraries will affect the fragment proportions in the final library. Several restriction endonucleases are used as examples in the Basic Protocol but can be substituted with any restriction enzyme of the researcher’s choice. BASIC PROTOCOL
Combinatorial Recombination of Gene Fragments
CONSTRUCTION OF A COMBINATORIAL GENE LIBRARY This protocol is split into six phases—two design phases and four construction phases. The first design phase determines the recombination sites and the gene fragments between them. Additionally, the “tag” sequences used to generate specific overhangs at the recombination sites are designed. PCR primers to insert the tags into the genes are designed during the second design phase. The construction phases are summarized in Figure 26.2.2. The first construction phase introduces tag sequences into the genes that will be recombined. The blocks are also divided into groups of four or fewer to facilitate ligation steps during construction phase 3. The product of the first construction phase is a set of plasmids; each group of four blocks has a plasmid for each gene recombined. For example, three genes with eight blocks (two groups) would have a set of 3 × 2 = 6 plasmids (Fig. 26.2.2). These plasmids should be sequenced to ensure that no mutations have been introduced during the construction process.
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Figure 26.2.2 Overview of the construction process broken down into the four construction phases. This figure demonstrates the recombination of three parental genes, broken into eight blocks (seven recombination sites). First, tag sequences that will allow specific overhangs to be generated are inserted into the genes using PCR. Next, the tag-inserted genes are cut with a type IIB restriction enzyme to expose the DNA fragments with desired overhangs, and the tag sequences are removed. The DNA fragments are then ligated together to form two mini-libraries, which are cloned individually. Finally, the two mini-libraries are ligated to form full-length genes. Sequences cloned and transformed into E. coli are shown with the plasmid backbone.
The second construction phase generates sequence blocks that ligate independently through cleavage of the tag-inserted genes with a type IIB restriction enzyme that leaves specified overhangs. The third construction phase ligates these blocks together to generate mini-libraries that contain 12 of the full-length gene. These mini-libraries are cloned separately and provide an opportunity to check for proper construction. The final construction phase ligates the mini-libraries together to generate full-length genes and a full-size library.
Materials Custom-synthesized oligonucleotides for PCR primers Gene sequences for proteins to be recombined High-fidelity DNA polymerase for PCR, e.g., Pfu DNA gel extraction kit (Zymogen Zymoclean Gel DNA Recovery Kit or Qiagen Qiaquick Gel Extraction Kit) Current Protocols in Protein Science
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PstI restriction endonuclease SalI restriction endonuclease DNA purification kit (Zymogen DNA Clean and Concentrator or Qiagen Qiaquick PCR Purification Kit) Plasmid for construction (e.g., pBC KS+; Stratagene) Calf alkaline phosphatase (Tabor, 1987a) BsaXI restriction endonuclease T4 DNA ligase (Tabor, 1987b) NdeI restriction endonuclease 150 × 15 mm LB agar plates (APPENDIX 4A) with antibiotic for transformation SapI restriction endonuclease BamHI restriction endonuclease HindIII restriction endonuclease Plasmid for final library cloning and expression (e.g., pET28) 16◦ C water bath Additional reagents and equipment for the polymerase chain reaction (APPENDIX 4J), agarose gel electrophoresis (APPENDIX 4F), restriction enzyme digestion (APPENDIX 4I), introduction of plasmid DNA into cells (APPENDIX 4D), preparation of plasmid DNA (APPENDIX 4C), DNA sequencing (Ausubel et al., 2006, Chapter 7), quantitation of nucleic acids with absorption spectroscopy (APPENDIX 4K) NOTE: All restriction enzymes mentioned in this protocol are examples, and may be replaced with other restriction enzymes compatible with the design of the experiment. SapI and BsaXI cleave outside their recognition sequences and should only be replaced with enzymes that have similar properties.
Design phase 1: Choose recombination sites and design tag sequences 1. Choose the type IIB restriction site to be used during construction to generate 5 overhangs. Type IIB restriction enzymes cut asymmetrically outside of their recognition sites, allowing any desired overhang to be generated. The authors have used both BsaXI (leaves 3-bp 5 overhang) and BaeI (leaves 5-bp 5 overhang) for construction and have found that BsaXI works better. BsaXI is used in this example. Because BsaX I leaves 3-bp overhangs, the recombination sites require 3 bp of identity in the parental sequences.
2. Choose recombination sites using methods discussed in Background Information. The recombination sites chosen using these methods may not be experimentally feasible and may have to be adjusted. A recombination site should have identical base pairs in all the parental genes to form the overhangs, as shown for 3 bp in Figure 26.2.3A. The number of base pairs required varies depending on the Type IIB enzyme chosen.
3. Make a list of the recombination sites with the overhangs (Fig 26.2.3B). The DNA between recombination sites is referred to generally as a block. A fragment is a block from a specific gene.
4. Starting at the N-terminus, group the blocks so that four or fewer sequential blocks are in each group. These groups will make up the mini-libraries that are each treated separately until the final construction phase (Fig 26.2.3B).
5. Separately list the recombination site overhangs internal to each mini-library. Combinatorial Recombination of Gene Fragments
There will be one recombination site that is not in either group. This site will be used to join the two mini-libraries in the final construction phase (Fig 26.2.3B).
6. Ensure that the overhangs will not cross-ligate within each list of recombination sites.
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Figure 26.2.3 (A) A recombination site must have at least 3 bp which are, or can be made, identical for all the genes to be recombined. For the recombination site shown, the last base pair of the codon can be changed to accommodate the recombination site, without changing the amino acid sequence of the proteins. (B) The recombination sites are listed with the unique overhangs and then grouped for the mini-libraries. Within each group, these overhangs must be different. However, the same overhang can be repeated in different groups (e.g., recombination sites II and V have the same overhang, but they are in different mini-libraries). (C) The template for the tag sequence contains the BsaXI (bold) and NdeI (italics) restriction sites. (D) A tag for each recombination site is formed by first adding the overhang bases, as listed in panel B. (E) The remaining bases in the tag are filled in using bases that will not cross-anneal during PCR.
They must be unique and nonpalindromic. For example CTA and TAG will cross-ligate, and AATT will cross-ligate with itself.
7. Design template tag sequences. For each mini-library a set of tag sequences will be inserted into the gene at the recombination sites. Some portions of these tag sequences are identical for all recombination sites. To design the tags, first make a template tag as shown in Figure 26.2.3C. The BsaXI recognition site is shown in bold and a second restriction site (NdeI) site is shown in italics. S, X, Y, and N represent bases in the tag that can be any base in this template, but will be filled in as the tags are designed.
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Figure 26.2.4 (A) An overview of the PCR protocol necessary to form the tag-inserted sequences. (B) Primers to insert the tag sequences into each gene are used to amplify the fragments before and after the restriction site, and to form part of the tag. They overlap in order to allow PCR-based assembly of the fulllength construct with the tags inserted. (C) There are joining primers at the ends of the mini-libraries. These primers contain SapI (bold) restriction sites to generate the overhangs and secondary sites (italics) for cloning. Lowercase letters are bases added to the 5 end of the primer to ensure efficient restriction enzyme cleavage. (D) The N- and C-terminal primers each contain two restriction sites, one for cloning during construction (italics) and a second for cloning into the final expression plasmid (bold).
This template will be different for different type IIB restriction enzymes, depending on their recognition and cleavage sites. The second restriction enzyme will be used to remove unwanted tags during the procedure and should be robust and ideally cut in the same buffer as the type IIB enzyme. For demonstration purposes here, BsaX I is used as the IIB restriction site and NdeI as the secondary site. Neither enzyme should cut within any of the genes. Combinatorial Recombination of Gene Fragments
8. For each recombination site, except the final joining site (see step 5), copy the template tag and replace the S portions with the overhang bases as shown (Fig. 26.2.3D).
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9. Next, replace the X, Y, and N portions of each tag with bases that are ∼50% GC and that will not cross-anneal during PCR or generate a restriction site used during the construction process (Fig. 26.2.3E).
Design phase 2: Design primers 10. For each gene, design two primers to insert each tag. One primer amplifies the DNA fragment before the tag, and the second amplifies the DNA fragment after the tag (Fig. 26.2.4). The 5 portions of the primers match the tag sequence and overlap (Tm ≥ ∼50◦ C) to allow PCR-based construction of the gene with tags inserted (Fig. 26.2.4B). The 3 portion of the primer is unique for each gene.
11. For each gene, design primers for the N- and C-termini. These need to contain two restriction sites, each at the 5 end for cloning into plasmids. The restriction sites for the final expression plasmid should be closest to the gene DNA, and the restriction sites for the construction plasmid should be placed at the 5 end of the primer (Fig. 26.2.4D). SalI and PstI are used here as example restriction sites for the N- and C-termini during construction and BamHI and HinIII restriction sites for the final cloning. The reason for using two different plasmids is discussed in Critical Parameters. However, two sets of restriction sites are required for the procedure.
12. Design primers for junction of mini-libraries. See considerations below for cloning final library. The last primers are those that allow the mini-libraries to be cloned independently during the first three stages of construction and then to be joined during the final stage. This also involves two restriction sites in each primer. Each primer must contain a restriction site that allows cleavage outside of the recognition site to expose the desired 5 overhang (Fig. 26.2.4C). Choose this restriction enzyme and design the primer to ensure correct cleavage to release overhangs. For this step, it is better to use a type II restriction site with a smaller number of base-pairs between the recognition and cleavage sites than BsaXI. SapI generally produces good results and is used here as an example. It is important to note that many of these enzymes are asymmetric cutters, and the site must be designed in the correct orientation.
13. Design primers for junction of mini-libraries. See considerations below for cloning during construction. In addition, restriction sites for cloning during construction are required at the 5 ends of each joining primer (Fig. 26.2.3C). These restriction sites should be the same as those used for cloning the N- and C-termini into the construction plasmid. The primer that forms the C-terminal portion of the first mini-library (amplifying block D) has a PstI site, and the primer that forms the N-terminal portion of the second mini-library (amplifying block E) has a Sal I site, so that both mini-libraries can be cloned into the same plasmid. At this point one should have a list of primers that includes a forward and reverse primer for each fragment.
14. Examine primers and starting gene sequences to ensure that none of the restriction sites used during construction are present except where desired.
Construction phase 1: Generate tag-inserted plasmids 15. Use PCR (APPENDIX 4J) to amplify each fragment with the appropriate primers designed above. Use the full-length gene as template. A high-fidelity polymerase, such as Pfu, is recommended for all PCR.
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16. Use preparatory agarose gel electrophoresis (APPENDIX 4F) to separate the individual PCR products, excise the band of the correct size, and purify it from the agarose using a gel extraction kit. 17. For each gene, connect the PCR products of adjacent fragments using PCR. Use the two fragments amplified above as template and the outer 5 and 3 primers from the reactions carried out previously (Fig. 26.2.4A) as primers. 18. Repeat steps 16 and 17 on the PCR products until the full-length (four or fewer) group of fragments is generated. 19. Digest the product with PstI and SalI (APPENDIX 4I) and remove the enzymes using a DNA purification kit, or heat-kill the enzymes according to the manufacturer’s instructions. Ligate the product into the construction plasmid cut with the same enzymes using T4 ligase according to the ligase manufacturer’s instructions. Transform the product into E. coli (APPENDIX 4D). 20. Pick several colonies, isolate the DNA, and perform an analytical digest with SalI and PstI to verify full-length tag-inserted sequences (APPENDIX 4C). Sequence the DNA (Ausubel et al., 2006, Chapter 7) to confirm that no mutations were introduced during the PCR. There should be a separate plasmid for each gene and each mini-library (i.e., for three genes and two mini-libraries, six plasmids are needed).
Construction phase 2: Generate library pieces This construction phase is performed separately with each mini-library (Fig. 26.2.2). 21. Prepare a large quantity (20 to 100 µg scale or larger; APPENDIX 4C) of each tag-inserted plasmid. Instead of using plasmid DNA, the tag-inserted gene can be PCR amplified using Pfu polymerase. This may result in a few extra mutations in the final library, but can save some time as steps 23 to 25 are no longer necessary.
22. Quantify the concentration of DNA using absorption spectroscopy (APPENDIX 4K) and mix together 3 pmol of each tag-inserted plasmid. 3 pmol is roughly 1 µg/kb of plasmid (i.e., 3 pmol is 9 µg of a 3-kb plasmid). This amount does not need to be exact, but is a guide to the appropriate scale necessary for the procedure. It is important that the DNA be mixed in the molar ratio desired in the library. Even if the tag-inserted gene is PCR amplified (see step 21), the resulting DNA must be quantified and mixed appropriately.
23. Digest with PstI and SalI restriction endonucleases (APPENDIX 4I) to remove the insert from the construction plasmid. This step is unnecessary if the tag-inserted genes were PCR amplified in step 21.
24. Dephosphorylate with calf alkaline phosphatase according to the manufacturer’s instructions (also see Tabor, 1987a). This will ensure that no ligation occurs at the termini of the insert, which would interfere with the rest of the reaction during construction phase 3. This step is unnecessary if the tag-inserted genes were PCR amplified in step 21.
Combinatorial Recombination of Gene Fragments
25. Use preparatory agarose gel electrophoresis (APPENDIX 4F) to isolate the dephosphorylated inserts and remove the plasmid backbone. Purify the inserts using a gel extraction kit. This removes the unnecessary plasmid DNA from subsequent reactions where it may hinder the process.
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This step is unnecessary if the tag-inserted genes were PCR amplified in step 21.
26. Cut the insert with BsaXI (or other type IIB restriction site, as discussed during the design phase) according to the manufacturer’s instructions to expose the 5 overhangs on the blocks. 27. Remove the released tags and BsaXI using a DNA purification kit according to the manufacturer’s instructions. It is important that the tags be removed from the reaction. Not all DNA purification kits remove small pieces of DNA. Blocks smaller than the cutoff for the DNA purification kit are also removed during this step. Running an analytical agarose electrophoresis gel to verify that smaller fragments are not lost is often worthwhile here. If using small blocks, see Alternate Protocol 1.
Construction phase 3: Generate multiple mini-libraries This construction phase is done separately for each mini-library (Fig. 26.2.2). 28. Ligate the fragments together using T4 ligase (also see Tabor, 1987b) for 4 hr at 16◦ C. Heat-kill the ligase after the reaction according the ligase manufacturer’s instructions. The ideal concentration of 5 ends in the reaction is 0 .15 µM. Assuming a 70% yield on the above steps starting with N × 3 pmol of each plasmid (where N is the number of genes with which the procedure started), N × 50 µl is ∼0.15 µM. There should be N × 6 Weiss units of ligase added to the reaction, proportional to the amount of DNA. One Weiss unit of T4 DNA ligase converts 1 nmol of 32 P from pyrophosphate into Norit-adsorbable material in 20 min at 37◦ C (Weiss et al., 1968). One Weiss unit equals ∼67 cohesive-end units.
29. Digest the ligated insert with BsaXI and NdeI according to the manufacturer’s instructions (also see APPENDIX 4I). Heat-kill the restriction enzymes or remove them using a DNA purification kit. Make sure to use the appropriate buffer conditions. Purification of the DNA may be required after the ligation and prior to the restriction digest.
30. Amplify the ligated product by PCR (APPENDIX 4J) using the appropriate joining primers and N- or C-termini primers. For each mini-library, use every possible primer pair, resulting in four reactions for two genes, nine for three genes, etc. Start with small amounts of template; it is important that all the PCR reactions work on the same batch of ligated template.
31. Purify the PCR products using preparatory agarose gel electrophoresis and a gel extraction kit. Make sure the pieces are appropriately sized. Bands slightly larger than expected may indicate the tags were not completely removed and bands smaller than expected may indicate that one or more of the fragments was not correctly incorporated. It may be helpful to use PCR products from unaltered parent genes as controls to ensure the band sizes are correct.
32. Quantify the gel-purified product using absorption spectroscopy (APPENDIX 4K) and mix the reactions in the molar ratio desired in the final library. This step is only necessary if the PCR reactions did not have roughly equivalent yields or if there are significant variations between samples during the purification of the PCR products. However, better quantification will only decrease undesirable biases in the final library. Protein Engineering
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33. Digest the mixed reaction with SalI and PstI, then remove the enzymes using a DNA purification kit (or heat-kill the enzymes). Using T4 ligase (Tabor, 1987b), ligate the insert into the construction plasmid cut with the same enzymes. 34. Transform the product into E. coli (APPENDIX 4D) and plate the bacteria on 150 × 15–mm plates. The number of colonies containing a full- length insert should be significantly (i.e., 2 orders of magnitude) greater than the maximum possible number of combinations in the mini-library (e.g., 81 for 3 genes and 4 blocks). This is to ensure that all combinations are represented in the final library. The authors recommend the use of 150 × 15–mm plates for this transformation to make step 36 easier. It is possible to estimate the complexity of the library by plating a small aliquot of the transformation onto a separate plate in order to count the colonies.
35. Pick 10 to 20 colonies for each mini-library, isolate the DNA, and perform an analytical digest with SalI and PstI to determine insert incorporation rate and sequence to confirm correct construction.
Construction phase 4: Construct full-length genes 36. Pool the colonies from each mini-library transformation and isolate the DNA (APPENDIX 4C). If there is insufficient cell mass for an effective DNA preparation, then grow the cells for a few hours to increase the cell mass. Quantitate the DNA using absorption spectroscopy (APPENDIX 4K). 37. Digest 3 pmol of the N-terminal mini-library with SalI to linearize the plasmid (APPENDIX 4I). Digest 3 pmol of the C-terminal mini-library with PstI to linearize the plasmid. This should be the restriction enzyme that is not next to SapI in the mini-library.
38. Dephosphorylate the overhangs with calf alkaline phosphatase (Tabor, 1987a). This prevents any cross-ligation during step 42.
39. Purify the linearized plasmid with a DNA purification kit. 40. Digest the plasmid with SapI (APPENDIX 4I) to release the library inserts. 41. Purify the inserts using preparative agarose gel electrophoresis (APPENDIX 4F) and a gel extraction kit. 42. Ligate the inserts together for 4 hr at 16◦ C using T4 ligase (Tabor, 1987b). Heat-kill the ligase after the reaction. The ideal concentration of 5 ends in the reaction is 0 .15 µM. Assuming a 70% yield in the above steps, that is ∼40 µl. Approximately 6 Weiss units of ligase should be sufficient for this reaction (see step 28 for information about Weiss units). It may be beneficial to purify the correct-size insert using preparative agarose gel electrophoresis and a gel extraction kit, but this is optional.
43. Digest the ligated product with BamHI and HindIII according to the manufacturer’s instructions (also see APPENDIX 4I). Remove the enzymes using a DNA purification kit or heat-kill the enzymes. 44. Cut the final expression plasmid (e.g., pET28) with the same enzymes, then ligate the cut inserts into the cut plasmid using T4 ligase (Tabor, 1987b). Combinatorial Recombination of Gene Fragments
45. Transform into E. coli (APPENDIX 4D) to generate the full library.
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LIBRARY CONSTRUCTION TO INCORPORATE SMALL (<40 bp) BLOCKS This is a modification of the Basic Protocol to add small blocks (<40 bp) into the library. Blocks of this size cannot be used in the Basic Protocol because they are lost during the purification steps. Incorporating many blocks in this manner is not recommended because they cannot be sequenced to ensure integrity before they are incorporated into the library.
ALTERNATE PROTOCOL 1
Additional Materials (also see Basic Protocol) T4 polynucleotide kinase (Tabor, 1987a) Kinase buffer with 1 mM ATP Thermal cycler capable of 1◦ C/sec ramp 1. Perform steps 1 to 27 of the Basic Protocol, performing steps 2 to 4 below in parallel with the Basic Protocol steps, then proceed to step 5 of this protocol. 2. For each gene, design two oligonucleotides that, when annealed, form the fragment with the appropriate 5 overhangs. It is very important that these oligonucleotides be short (<50 bp) and of the highest purity available (PAGE purified). The small fraction of oligonucleotides containing single-basepair deletions will result in frame shifts in the library, and all mutations that occur in the oligonucleotides will be transferred into the library.
3. Phosphorylate each oligonucleotide using T4 polynucleotide kinase (see Tabor, 1987a, or purchase phosphorylated from the manufacturer). The recommended concentration of oligonucleotide in the reaction is ∼3 µM, but follow the manufacturer’s instructions. Most kinase buffers do not contain ATP, which therefore must be supplemented to 1 mM.
4. Anneal oligonucleotide pairs by combining equimolar amounts, heating the mixture to 100◦ C and cooling at 1◦ C/sec to 16◦ C. Alternatively, cool slowly to room temperature by placing tubes on bench after heating.
5. Add 6 pmol (2 µl) of each annealed oligonucleotide pair to the ligation reaction in step 28 of the Basic Protocol (construction phase 3). Add twice as much of the phosphorylated oligonucleotides to the reaction as compared to the cut pieces that would otherwise be used in Basic Protocol 1, because the phosphorylation reaction is inefficient and many oligonucleotides may not be phosphorylated.
LIBRARY CONSTRUCTION TO RECOMBINE MORE THAN 8 BLOCKS This is a modification of the site-directed recombination Basic Protocol to allow the recombination of more than eight blocks. The authors do not recommend increasing the number of blocks in the mini-libraries to more than four. However, additional minilibraries can be constructed and added during the final ligation step in construction phase 4. This protocol outlines a few key modifications of the design of additional mini-libraries and the final construction phase.
ALTERNATE PROTOCOL 2
1. During the design phase, make sure that none of the overhangs generated for the final ligation cross-ligate (similar to ensuring that no overhangs within a mini-library cross-ligate, as in step 6 of the Basic Protocol). 2. During the final construction phase, use the following procedure for mini-libraries not at the N- and C-termini instead of steps 37 to 41. For the N- and C-terminal mini-libraries use steps 37 to 41 from the Basic Protocol.
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a. Digest 3 pmol of the mini-library with SapI according to the manufacturer’s instructions (also see APPENDIX 4I). b. Purify the released insert using preparative gel electrophoresis (APPENDIX 4F) and a gel extraction kit. 3. Following step 42 (construction phase 4), perform the optional preparative agarose gel purification of the correct-size ligation product. This step is not optional if there are more than two mini-libraries.
4. If there is not sufficient ligation product for cloning, PCR-amplify the ligation product essentially as in steps 30 to 32 (construction phase 3) of the Basic Protocol. Use all possible pairs of N- and C-terminal primers. Continue the ligation protocol starting at step 43 (construction phase 4). ALTERNATE PROTOCOL 3
LIBRARY CONSTRUCTION USING A SUBSET OF GENE FRAGMENTS It may be desirable to allow only certain genes to occur at some positions in the combinatorial library. This protocol can be easily modified to remove one or more of the gene fragments from the synthesis reaction. However, this must be planned accordingly during the design phase. The essential component of the method is creation of sequence blocks with specific overhangs that allow correct ligation. To remove a block, simply leave it out and create a tag sequence appropriate for the flanking blocks. To do this, design a tag specific to the gene with the two different overhangs required by the flanking blocks rather than the same overhang on both ends. It is important to make sure that all fragments are present in desired molar proportions and that each block has the appropriate overhangs when cut with the type IIB restriction enzyme.
COMMENTARY Background Information
Combinatorial Recombination of Gene Fragments
Site-directed recombination was developed as an alternative to PCR and truncationbased methods of recombining distantly related genes (Hiraga and Arnold, 2003). PCRbased methods are usually limited to genes with more than 70% sequence identity (Joern et al., 2002) and truncation-based methods lead to a large number of nonviable variants due to insertions, deletions, and frame-shift issues (Ostermeier et al., 1999a,b; Lutz et al., 2001; Sieber et al., 2001). Additionally, with these methods there is little or no control over recombination site number or location within the product genes. Site-directed recombination trades the blind approach used in such methods for a more directed tactic that can take advantage of additional information to choose recombination sites that result in a high proportion of folded chimeras. Additionally, the resulting populations have well defined sequences that allow a much more detailed analysis of the collection of proteins generated in the experiment (Meyer et al., 2003; Drummond et al., 2005). Alternate Protocol 3 for recombining only specific gene fragments can also be used to build libraries that contain
a subset of the possible parental genes at any given position (Saraf et al., 2005). In order to generate a population with a large percentage of folded and potentially functional chimeras by recombining distantly related proteins, it is necessary to choose recombination sites effectively. The recombination sites chosen dictate the chimeras that will be constructed. Recombination sites between related proteins have been chosen, with varying degrees of success, to correspond with regions of high sequence identity (Burson and Khosla, 2000) as well as with boundaries of exons (Back and Chappell, 1996), secondary structure elements (Jermutus et al., 2001; Koenderink et al., 2001), and clear structural domains (Nicot et al., 2002; Roman et al., 2003). As the sequence identity between the genes to be recombined decreases, the integration of structural information becomes more and more critical to obtaining a highly folded population. In order to meet this need, several computational methods have been developed that rate chimeras based on sequence and structure information (Voigt et al., 2002; Saraf et al., 2004; Hernandez and LeMaster, 2005). Comparison
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with existing data on chimeric enzymes has shown that they are at least somewhat effective at predicting which chimeras are more likely to retain function. However, most of these methods have not been thoroughly tested. The authors of this unit have developed a simple computational model for rating chimeras that takes into account both the protein structure and the sequence of the proteins being recombined to predict which chimeras are more likely to retain the parental fold. The metric used, SCHEMA disruption (E), is calculated by first identifying all amino acid pairs that are contacting in the three-dimensional structure (heavy atoms separated by less than ◦ 4.5 A). A chimera’s disruption is then determined by counting the number of contacting pairs where the identities of the amino acids have changed (“broken contacts”). A contact is not broken if the new amino acid pairing is found in any of the parental proteins. Using this method, a chimera can be assessed in silico for likelihood of folding before it is constructed in the laboratory (Silberg et al., 2004). The authors have shown that this metric is better at determining which chimeras will function than simply counting the number of mutations in the chimeras (Meyer et al., 2003). The authors have applied SCHEMA disruption to library design (choosing the best recombination sites) by developing an algorithm which minimizes the average disruption for all chimeras in a library at many levels of mutation. This algorithm, RASPP (Recombination as a Shortest Path Problem), uses changes in minimum block length to identify libraries with the least disruption at a range of levels of average mutation (Endelman et al., 2004). The output of RASPP is a curve that contains the libraries with the lowest disruption at various levels of average mutation. Disruption can be transformed into a percentage of chimeras folded, if desired. However, the relationship between disruption and percentage folded must be calibrated using previously collected data (Meyer et al., 2003; Otey et al., 2004). At the Web site listed under Internet Resources, the authors provide MATLAB code to perform RASPP and disruption calculations based on a protein structure and an amino acid sequence alignment of the parental proteins.
Critical Parameters The most critical parameter in building a site-directed recombination library is primer design. It is very easy to make a mistake during the primer design that will detrimentally
affect the construction procedure. Be sure to check all of the DNA created during the procedure for all of the restriction sites used to ensure that there are no extra sites. Be aware that sites may be created during the construction process due to the insertion and deletion of the tag sequences. The best way to ensure that there are no extra sites is to build, in silico, all of the intermediate DNA fragments from the designed tags, gene sequences, and plasmid sequences, and check them individually, stepping through each phase of the procedure with the DNA sequences. Most of the Basic Protocol involves standard molecular biology techniques strung together. However, it is essential that every step work with high efficiency. The PCR reactions, especially those involving the joining and N- or C-termini primers, should be optimized, and the primers for each gene should be specific. The ligations and restriction digests may also be optimized to ensure a higher success rate. There are several places where plasmids are transformed into E. coli. These can serve as places to stop and make sure the procedure is proceeding as planned by sequencing the DNA. It is also essential that the transformed mini-libraries have many more colonies than the expected complexity of the mini-library (at least 1 to 2 orders of magnitude).
Troubleshooting Table 26.2.1 lists some of the more common problems that may be encountered using the protocols described in this unit, along with explanations of possible causes of the problems and suggested approaches for overcoming these barriers.
Anticipated Results The library produced by this method is likely to be biased, with some fragments occurring disproportionately in comparison to the desired molar ratio. This can be minimized through careful quantitation of the plasmids during steps 22 and 32 of the Basic Protocol and through proper mixing. However, it is unlikely that the library will be completely unbiased. The bias can be determined by sequencing a set of unselected chimeras. Sequencing of a large number of chimeras is easily achieved in high-throughput format by DNA hybridization (Meinhold et al., 2003). For a two-parent library, the average percentage of each fragment is 50% ± 11%. For two separate three parent libraries the standard deviations are higher, with the average percentage of each fragment 33% ± 13 and
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Table 26.2.1 Troubleshooting Guide for Creating a Combinatorial Gene Library
Problem
Probable Cause
Solution
Construction phase 1 (steps 15-20) Individual blocks amplify but Not enough overlap between the assembly PCRs are not the primers in the tag regions working
Lower annealing temperature. Design new primers with more overlap.
Tags did not insert correctly
Raise annealing temperature. Design new primers with more distinct regions in the tags.
Tag overlaps cross-annealed during PCR
Construction phase 3 (steps 28-35) PCR after ligation yields bands that are too small
One of the blocks is missing because it is too small (<40 bp)
Use Alternate Protocol 1 to add in the piece right before the ligation
PCR after ligation yields bands that are too large
The digests with BsaXI and NdeI to remove tags after ligation are not working correctly
Optimize these digests using plasmid DNA
DNA purification is not removing cut tags correctly
Try DNA purification kit from a different manufacturer
Dephosphorylation in step 24 was not complete
Optimize dephosphorylation
Combination of issues listed above. Also check to make sure that BsaXI is cutting to release the fragments
Run analytical agarose gel prior to the ligation to ensure that pieces are cut
One (or more) PCR reaction Not enough template present after ligation gives no product
Increase amount of template. Check the proportionality of the genes added to the mix.
PCR after ligation yields multiple bands
None of the PCR reactions work after ligation
PCR reactions do not all produce similar yields
Combinatorial Recombination of Gene Fragments
PCR conditions are not optimized
Optimize PCR reactions on plasmids with small amounts of template
Template is not full length
Make sure there are no BsaXI or NdeI sites within the gene
There is no template
Run an analytical gel to check concentration of PCR template. There should be something visible in the amount added to the PCR, although it may be a smear.
Template does not contain equal proportions of the different parental fragments
Check the proportionality of the genes added to the mix. Make sure all genes cut with BsaXI individually. Mix the PCR reactions so that an equal quantity of DNA is used from each reaction.
Pieces not in correct order in Check overhangs for mini-libraries/extra pieces palindromic sequences, or found in libraries sequences that could possibly cross-anneal
Ligate at a higher temperature (20◦ -25◦ C)
continued
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Table 26.2.1 Troubleshooting Guide for Creating a Combinatorial Gene Library, continued
Problem
Probable Cause
Solution
Tags still present in mini-libraries
BsaXI and NdeI are not removing tags correctly
Optimize the restriction digest using plasmid DNA
DNA purification not Try DNA purification kit removing tags prior to ligation from a different manufacturer Block missing in mini-library Block is too small and getting lost in DNA clean-up used to remove tags Block from one gene is missing in mini-library
Use Alternate Protocol 1
There is a restriction site in the Check to make sure that the piece block is not cleaved by one of the enzymes used One of the tags is not cleaving Check the tag sequences at completely both ends for the BsaXI site
Construction phase 4 (steps 36-45) Mini-libraries not assembling Cross-ligation of overhangs in correct order
Check overhangs for palindromes
Incomplete dephosphorylation Optimize dephosphorylation Mini-libraries not assembling, or missing a component
Inefficient SapI digestion
Not enough insert to ligate into final expression plasmid
Optimize the SapI digest using plasmid Cut more DNA. PCR amplify the full-length product prior to cutting with HindIII/BamHI to generate more product.
Alternate Protocol 1: Adding small blocks Blocks do not incorporate in mini-library
Phosphorylation inefficient
Optimize phosphorylation reaction. ATP may be missing from buffer
Oligonucleotides designed incorrectly
Make sure that the oligonucleotides, when annealed, leave the necessary overhangs
33% ± 19%. While most positions (>60%) typically show even percentages of each fragment, there are always some positions where the distribution of fragments is such that one fragment is either not present in significant quantity (<10%) or is dominant (>70%). However, usually the mutation frequency is very low (0.007%), and most tags are correctly removed (in 40 randomly chosen chimeras, a single uncleaved tag sequence being identified).
Time Considerations The amount of time the design phase will take varies depending on the complexity of the library and how the recombination sites are chosen. Construction phase 1 will take 1 to 2 Current Protocols in Protein Science
days, and several days of DNA sequencing will then be required to ensure that the taginserted plasmids are correct. Construction phases 2 and 3 can be completed in a single, very long day, but it is easier to perform them over 2 or even 3 days, allowing the PCR amplification to proceed overnight. As with most molecular biology reactions, these can be stored overnight at −20◦ C after almost any step in the procedure. Following construction phase 3, it is also recommended that correct sequences be confirmed, which may take a few days. The final construction phase 4 can be done in a single day. An ideal construction with no problems will take ∼2 to 3 weeks to complete, depending on the turnaround time for DNA sequencing. However, due
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to the number of steps that may require optimization, it is more likely to take even an experienced molecular biologist 4 to 8 weeks to complete the first time. This time is not all dedicated to the experiment, but also includes growing time for the E. coli necessary to produce DNA.
Literature Cited Ausubel, F.M., Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G., Smith, J.A, and Struhl, K. (eds.). 2006. Current Protocols in Molecular Biology. John Wiley & Sons, Hoboken, N.J. Back, K. and Chappell, J. 1996. Identifying functional domains within terpene cyclases using a domain-swapping strategy. Proc. Natl. Acad. Sci. U.S.A. 93:6841-6845. Burson, K. K. and Khosla, C. 2000. Dissecting the chain length specificity in bacterial aromatic polyketide synthases using chimeric genes. Tetrahedron 56:9401-9408. Drummond, D.A., Silberg, J.J., Meyer, M.M., Wilke, C.O., and Arnold, F.H. 2005. On the conservative nature of intragenic recombination. Proc. Natl. Acad. Sci. U.S.A. 102:5280-5385. Endelman, J.B., Silberg, J.J., Wang, Z.-G., and Arnold, F.H. 2004. Site-directed protein recombination as a shortest-path problem. Protein Eng. Des. Sel. 17:589-594. Hernandez, G. and LeMaster, D.M. 2005. Hybrid native partitioning of interactions among nonconserved residues in chimeric proteins. Proteins: Structure, Function and Bioinformatics 60:723-731. Hiraga, K. and Arnold, F.H. 2003. General method for sequence-independent site-directed chimeragenesis. J. Mol. Biol. 330:287-296. Jermutus, L., Terrier, M., Pasamontes, L., van Loon, A.P.G.M., and Lehmann, M. 2001. Structurebased chimeric enzymes as an alternative to directed enzyme evolution: Phytase as a test case. J. Biotechnol. 85. Joern, J.M., Meinhold, P., and Arnold, F.H. 2002. Analysis of shuffled gene libraries. J. Mol. Biol. 316:643-656. Koenderink, J.B., Swarts, H.G.P., Stronks, C., Hermsen, H.P.H., Willems, P.H.G.M., and de Pont, J.J.H.H.M., 2001. Chimeras of X+, K+ATPases. J. Biol. Chem. 276:111705-11711. Lutz, S., Ostermeier, M., Moore, G.L., Maranas, C.D., and Benkovic, S.J. 2001. Creating multiple-crossover DNA libraries independent of sequence identity. Proc. Natl. Acad. Sci. U.S.A. 98:11248-11253.
Combinatorial Recombination of Gene Fragments
Meinhold, P.M., Joern, J.M., and Silberg, J.J. 2003. Analysis of shuffled libraries by oligonucleotide probe hybridization. In Methods in Molecular Biology Vol. 231: Directed Evolution Library Creation (F. H. Arnold and G. Georgiou, eds.) pp. 177-188. Humana Press, Totowa, N.J.
Meyer, M.M., Silberg, J.J., Voigt, C.A., Endelman, J.B., Mayo, S.L., Wang, Z.-G., and Arnold, F.H. 2003. Library analysis of SCHEMA-guided recombination. Protein Sci. 12:1686-1693. Nicot, C., Relat, J., Woldegiorgis, Haro, D., and Marrerro, P.F. 2002. Pig liver carnitine palmitoyltransferase. J. Biol. Chem. 277. Ostermeier, M., Nixon, A.E., and Benkovic, S.L. 1999a. Incremental truncation as a strategy in the engineering of novel biocatalysts. Bioorg. Med. Chem. 7:2139-2144. Ostermeier, M., Shim, J.H., and Benkovic, S.J. 1999b. A combinatorial approach to hyrbid enzymes independent of DNA homology. Nat. Biotechnol. 17:1205-1209. Otey, C.R., Silberg, J.J., Endelman, J.B., Bandara, G., and Arnold, F.H. 2004. Functional evolution and structural conservation in chimeric cytochromes P450: Calibrating a structure-guided approach. Chem. Biol. 11:309-318. Roman, L.J., McLain, J., and Masters, B.S.S. 2003. Chimeric enzymes of cytochrome P450 oxidoreductase and neuronal nitric-oxide synthase reductase domain reveal structural and functional differences. J. Biol. Chem. 278:25700-26707. Saraf, M.C., Horswill, A.R., Benkovic, S.J., and Maranas, C.D. 2004. FamClash: A method for ranking the activity of engineered enzymes. Proc. Natl. Acad. Sci. U.S.A. 101:4142-4147. Saraf, M.C., Gupta, A., and Maranas, C.D. 2005. Design of combinatorial protein libraries of optimal size. Proteins: Structure, Function and Bioinformatics 60:769-777. Sieber, V., Martinez, C.A., and Arnold, F.H. 2001. Libraries of hybrid proteins from distantly related sequences. Nat. Biotechnol. 19:456-460. Silberg, J.J., Endelman, J.B., and Arnold, F.H. 2004. SCHEMA-guided protein recombination. In Methods in Enzymology Vol. 228 Protein Engineering (D.E. Robinson and J.P. Noel, eds.) pp. 35-41. Elsevier Academic Press, Amsterdam. Tabor, S. 1987a. Phosphatases and kinases. In Current Protocols in Molecular Biology (F.M. Ausubel, Brent, R., R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 3.10.1-3.10.5. John Wiley & Sons, Hoboken, N.J. Tabor, S. 1987b. DNA ligases. In Current Protocols in Molecular Biology (F.M. Ausubel, Brent, R., R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 3.14.1-3.14.4. John Wiley & Sons, Hoboken, N.J. Voigt, C.A., Martinez, C., Wang, Z-G., Mayo, S.L., and Arnold, F.H. 2002 Protein building blocks preserved by recombination. Nat. Struct. Biol. 9:552-558. Weiss, B., Jacquemin-Sablon, A., Live, T.R., Faree, G., and Richardson, C.C. 1968. Enzymatic breakage and joining of deoxyribonucleic acid. J. Biol. Chem. 243:4543-4555.
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Internet Resources http://www.che.caltech.edu/groups/fha/ Web site that supplies RASPP code for determining optimal recombination points.
Contributed by Michelle M. Meyer, Kaori Hiraga, and Frances H. Arnold California Institute of Technology Pasadena, California (Kaori Hiraga’s current affiliation is: New York State Department of Health Albany, New York)
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Incorporation of Isotopically Enriched Amino Acids
UNIT 26.3
Isotopic labeling of proteins represents an important tool for the application of nuclear magnetic resonance (NMR), X-ray or neutron diffraction crystallography, and mass spectrometry (MS) methods to investigate the structure and dynamics of proteins alone or with interacting ligands. Uniform isotope labeling of recombinant proteins is especially useful in NMR methodology. Isotopic labeling strategies are determined by protein type and size, production method, and NMR spectral overlap, as well as the methods used for determining the phase of crystallographic diffraction data and the type of information being sought. The great majority of the uniformly isotope-labeled proteins reported in the literature thus far have been expressed in E. coli (Goto and Kay, 2000). However, complex proteins often cannot be expressed in E. coli in an active, correctly folded, and post-translationally modified form, and thus a eukaryotic host cell, e.g., a baculovirus expression system (UNITS 5.4 & 5.5), is used for expression. Smaller proteins (<100 amino acids) can also be synthesized and isotopically labeled via solid phase peptide synthesis. Proteins which are not expressed well in cells or are toxic can be produced using cell-free expression systems, such as wheat germ (UNIT 5.18). Finally, proteins isolated from native sources can be chemically modified to introduce isotopic labels.
STRATEGIC PLANNING Isotopic Labeling of Proteins Expressed Recombinantly Using E. coli Uniform labeling This unit covers a variety of methods for incorporating isotopic labels into proteins expressed recombinantly using E. coli (see Basic Protocol 1 and Alternate Protocols 1 to 6). Under aerobic conditions, bacteria can utilize glucose, pyruvate, acetate, succinate and glycerol as sole carbon sources, and ammonium salts (chloride, nitrate and sulfate being the most common) as sole nitrogen sources to synthesize 20 amino acids (Lian and Middleton, 2001; Fig. 26.3.1). Uniform labeling with 15 N is relatively simple and costeffective using defined media containing [15 N]-ammonium chloride or sometimes [15 N]ammonium sulfate as the sole source of nitrogen. The uniform 15 N labeling of proteins is essential for the implementation of 1 H/15 N-HSQC (heteronuclear single quantum coherence), a fundamental technique in protein NMR. Triple resonance experiments (1 H/13 C/15 N) allow for sequential assignment of uniformly 13 C/15 N-labeled recombinant proteins. The uniform 13 C-labeling of proteins can be achieved equally effectively by using either [13 C6 ]-glucose or sodium [1,2-13 C] acetate as the sole source of carbon. By supplying specifically 13 C-labeled glucose or glycerol as the sole source of carbon, certain carbon sites can be 13 C-labeled to a varying degree while other sites remain completely unlabeled. The labeling scheme of isotopic enrichment at alternating carbon sites is achieved using [1-13 C]-glucose or [2-13 C]-glycerol as the carbon source. The opposite labeling pattern is obtained by growth in a [1,3-13 C2 ]-glycerol enriched medium (LeMaster and Kushlan, 1996). The labeling scheme of alternating carbon atoms is desirable for NMR applications to overcome the scalar and dipolar coupling interactions introduced between directly bonded 13 C nuclei. Uniform labeling of selected amino acids Selective labeling of certain amino acids can also be used to generate information about protein folding and conformation. Hydrophobic residues such as Ile, Leu, and Val are usually situated in the protein core and are particularly useful for determining global folds. Protein Engineering Contributed by Mini Samuel-Landtiser, Cherian Zachariah, Chris R. Williams, Arthur S. Edison, and Joanna R. Long
26.3.1
Current Protocols in Protein Science (2007) 26.3.1-26.3.49 C 2007 by John Wiley & Sons, Inc. Copyright
Supplement 47
glucose
glycerol
glucose-6-phosphate His PENTOSEPHOSPHATE PATHWAY glyceraldehyde-3-phosphate
Phe Tyr Trp
chorismate
glycerate-3-phosphate
dihydroxyacetone phosphate C1 METABOLISM Gly Ser
phosphoenolpyruvate
Cys
pyruvate
Ala ANAPLEROTIC PATHWAY
hydroxyethyl thiamine pyrophosphate α - ketoisovalerate
Val
acetate
oxaloacetate GLYOXYLATE isocitrate aspartate CYCLE semialdehyde α -ketoglutarate malate
Lys
CITRIC ACID CYCLE
homoserine
succinate
Thr
Leu
acetyl CoA
Asp
Asn
GLYCOLYTIC PATHWAY
Glu Gln Pro Arg
Met
α - ketobutyrate
Ile
Figure 26.3.1 2001).
hydroxyethyl thiamine pyrophosphate
Amino acid synthesis pathways in E. coli (adapted from Liam and Middleton,
The methyl groups of Ala, Val, Leu, and Ile are derived from the methyl group of pyruvate because pyruvate is the direct biosynthetic precursor of these residues. These metabolic pathways are exploited in the generation of recombinant proteins in which the methyl groups of these residues are labeled differently from the rest of the protein molecule.
Incorporation of Isotopically Enriched Amino Acids
NMR sensitivity and resolution can be greatly improved in larger proteins with deuterium labeling. The use of deuterium (2 H) incorporation into proteins has been used for several years as a tool for simplifying proton spectra and improving proton relaxation properties. When proteins are expressed in bacteria grown in 2 H2 O medium containing protonated [3-13 C]-pyruvate, this results in a labeling pattern in which only the methyl groups of the aliphatic residues (Ala, Val, Leu and Ile) are 13 C labeled and are partially protonated (13 C2 H2 H; Lee et al., 1997; Ishima et al., 1999). Other side chain carbon and most hydrogen atoms are 12 C and 2 H, respectively. The overexpression of proteins in bacteria grown in 2 H2 O medium containing protonated [13 C3 ]-pyruvate as the sole carbon source results in uniform 13 C labeling and complete deuteration at the Cα and nearly complete deuteration at the Cβ positions of all amino acids (see Fig. 26.3.2 for Cα and Cβ positions; see Fig. A.1A.1 for structural information about all the amino acids). However, the methyl
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Figure 26.3.2 Cα and Cβ positions in an amino acid. See APPENDIX 1A for structural information about the 20 natural amino acids.
groups of the aliphatic residues Ala, Val, Leu, and Ile are 13 C labeled and remain highly protonated (Rosen et al., 1996). The use of [1-13 C]-glucose and [2-13 C]-glycerol as the sole carbon source results in relatively extensive labeling of the predominantly hydrophilic amino acids derived from the citric acid cycle. In the study of membrane proteins, where the membrane-embedded segments of these proteins consist of primarily hydrophobic amino acids, it would be advantageous to filter out the 13 C signals of the these hydrophilic amino acids in order to simplify the NMR spectra. The TEASE (ten-amino-acid selective and extensive; see Alternate Protocol 4) 13 C labeling protocol selectively and extensively labels ten predominantly hydrophobic amino acids that include Gly, Cys, Trp, Phe, Tyr, Ala, Val, Leu, Ser and His, which are produced from glycolysis and the pentose phosphate pathways (Hong and Jakes, 1999). At the same time, the biosynthesis of the amino acids from the citric acid cycle which include Asp, Asn, Met, Thr, Lys, Ile, Glu, Gln, Pro, and Arg can be blocked by supplementing the expression media with the unlabeled forms of these amino acids.
Selective labeling For some applications it is desirable to selectively label the protein or peptide at only one or a few amino acid residues. This selective labeling approach is used in NMR studies to filter out unwanted signals from overlapping resonances and thereby simplifies the NMR spectra so that unequivocal resonance assignments can be made. In MS the selective labeling of proteins is used to assist with the identification and quantification of peptide fragments of proteins. The incorporation of isotopic labels into a protein within a desired amino acid type, either uniformly or at specific atomic sites, while leaving the other amino acids unlabeled can be achieved by supplementing media with isotopically enriched amino acids. While bacteria are able to synthesize all their amino acids beginning from a sole nitrogen and carbon source, they will utilize a preformed amino acid, when available, in preference to synthesizing that amino acid. 19 F
spin labeling has been used for NMR studies of biological systems for over three decades. The use of 19 F NMR in the study of protein structure and function is a proven technique since the high-sensitivity fluorine nucleus is readily incorporated into amino acids without compromising the structure or function of the protein (Gerig, 2001). Dipolar interactions can lead to 19 F-1 H nuclear Overhauser effects (NOEs), which can provide information about internuclear distances. Several of the fluorinated analogs of the aromatic amino acids, e.g., 2-, 3- and 4-fluorophenylalanine, 6- and 5-fluorotryptophan and 3-fluorotyrosine (Fig. 26.3.3), can be obtained commercially. There are only a few examples of naturally occurring fluorine-containing compounds in biological systems. Synthetic methods are available for the preparation of fluorinated derivatives of most of the common amino acids. By far the most widely used method for incorporating fluorinated amino acids into the primary sequence of proteins is biosynthesis of the protein by bacteria and yeast (e.g., Kranz et al., 1996).
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Figure 26.3.3 Commonly used fluorinated amino acids: (A) 2-fluorophenylalanine, (B) 3-fluorophenylalanine, (C) 4-fluorophenylalanine, (D) 6-fluorotryptophan, (E) 5-fluorotryptophan and (F) 3-fluorotyrosine.
Isotopic Labeling of Proteins Expressed in Eukaryotic Host Cells Numerous proteins require eukaryotic host cells for expression. At present the baculovirus expression system (UNITS 5.4 & 5.5) is one of the most frequently used eukaryotic expression systems, although only a few reports of uniform isotope labeling in proteins using this system have been published. One of the major advantages of the eukaryotic expression systems is the ability to produce functional heterologous proteins such as enzymes, antibodies and receptors. Basic Protocol 2 presents an adaptation of the Gateway Technology (also see UNIT 5.17) to transfer the gene of interest into a Gateway vector, followed by recombination with the baculovirus genome, and expression of either unlabeled or labeled heterologous protein in insect cells using a commercially available labeling medium. Although recombinant protein expression in yeast is a viable alternative, this topic is beyond the scope of this unit (see UNITS 5.6 to 5.8).
Incorporation of Isotopically Enriched Amino Acids
Isotopic Labeling of Proteins Expressed in Cell-Free Systems Cell-free systems which couple transcription-translation reactions in vitro offer numerous advantages over the most widely used E. coli-based protein expression system. About onethird of bacterial proteins and a higher fraction of eukaryotic proteins cannot be expressed in E. coli (Vinarov et al., 2004). Cell-free methods for protein expression utilizing both prokaryotic and eukaryotic sources for cell extracts have been used to express proteins that undergo rapid proteolysis in cells (Goff and Goldberg, 1987), membrane proteins (Klammt et al., 2004), proteins that are toxic (Henrich et al., 1982; Chrunyk et al., 1993) and those that form inclusion bodies in cells (Chrunyk et al., 1993). Basic Protocol 3 presents a method for isotopic enrichment in a cell-free environment based on E. coli extracts. Cell-free protein synthesis is used to produce labeled recombinant proteins for NMR structural studies due to its advantages of fast expression, small reaction volumes, and efficient utilization of labeled precursors. Hence, cell-free systems can prove to be economical when coupled with the apparent lack of toxicity of the expressed protein (Ozawa et al., 2006).
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Isotopic Labeling of Peptides Made via Solid Phase Peptide Synthesis Basic Protocol 4 describes the preparation and incorporation of isotopically enriched amino acids using Fmoc-based solid phase peptide synthesis (SPPS; Chan and White 2000). This approach is expedient for the incorporation of isotopes at very specific points in small proteins although it is less cost effective than recombinant approaches. Support Protocol 3 describes preparation of an Fmoc-protected amino acid and Support Protocol 4 provides a ninhydrin-based method for monitoring the amino acid coupling reaction. Isotopic Labeling by Chemical Modification of Natural Proteins For proteins that are isolated rather than synthesized, chemical modification is possible to introduce isotopes at chemically reactive side-chains. Additionally, chemical exchange can be used to introduce 2 H labels at positions that have exchangeable protons, such as the amides in the protein backbone. Chemical exchange can be used as a very sensitive measure of protein folding rates and mechanisms. These techniques are described in Basic Protocol 5 and Alternate Protocol 7. The techniques described in this unit can also be used to isotopically enrich individual protein domains. The segmental labeling of proteins, in which fragments of the sequence with arbitrary isotopic composition are recombined, represents a significant step toward the ability to control the sequential, regiospecific, and stereospecific isotopic composition of virtually every atom in the protein. Any approach to segmental labeling involves the production of fragments with the appropriate isotopic labeling and the condensation of the fragments to produce the intact target protein. The most effective approach for chemical fragment condensation of synthetic unprotected peptides is native chemical
Figure 26.3.4 Schematic illustrating native chemical ligation (NCL) using thioesters (adapted from Cowburn and Muir, 2001).
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ligation (NCL) in which the chemoselective reaction between a C-terminal thioester and an N-terminal cysteine residue of the other fragment results in a native peptide as shown in Figure 26.3.4. Although this approach has been applied to obtain peptides, small proteins or protein domains (up to ∼150 residues), incorporating isotopic labels in larger proteins is difficult due to the limited size of the ligated fragments.
Isotopic Labeling of Proteins by Self-Catalytic Protein Splicing (Inteins) The ligation of two protein fragments under very mild conditions has been shown to be possible through a self-catalytic protein splicing process involving naturally occurring domains called inteins. In the course of intein excision from a polypeptide chain, the flanking regions (called exteins) are joined together by a native peptide bond. The majority of the residues involved in this reaction reside within the intein and only an
Incorporation of Isotopically Enriched Amino Acids
Figure 26.3.5 Schematic outline of segmental isotope labeling of proteins using trans-splicing strategy (adapted from Cowburn and Muir, 2001).
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N-terminal cysteine, threonine or serine residue is required on the C-terminal extein for ligation to occur. There are currently two strategies for segmental labeling which use inteins: trans-splicing and expressed protein ligation (EPL) (Ralf et al., 2004). In the trans-splicing strategy (Fig. 26.3.5), the intein is split into two fragments and expressed as a fusion to either the N-terminal or C-terminal fragment of the protein target. The two fusion proteins are combined in a denatured form and refolded to generate an active intein that catalyzes the splicing reaction to ligate the target polypeptides. For expressed protein ligation, the production of a C-terminal thioester relies on mutations of the intein stopping the reaction sequence of Figure 26.3.5 prior to the rearrangement, in the side reaction of step 2. The use of intein technologies for the incorporation of isotopically labeled amino acids is a specialized technique requiring suitable plasmids and vectors, specific to the protein of interest to be labeled. For detailed information to assist in developing a protocol specific to the protein of interest, see David et al., 2004 and Cowburn and Muir, 2001.
ISOTOPIC LABELING OF PROTEINS PRODUCED RECOMBINANTLY IN E. col i Almost any DNA sequence under control of the T7 lac promoter can be transcribed by phage T7 RNA polymerase in E. coli expression systems (Studier and Moffatt, 1986; Studier et al., 1990). In pET expression systems, T7 RNA polymerase is under control of the lacUV5 promoter in DE3 lysogen host strains such as BL21 (DE3). The target DNA sequence is subcloned into a pET vector downstream of the T7lac promoter, which can only be transcribed by T7 RNA polymerase in the host strain. Under noninducing conditions, little if any T7 RNA polymerase is present in the cell, and consequentially little or no target protein is produced. Upon induction either by addition of isopropyl1-β-D-thiogalactopyranoside (IPTG; see Basic Protocol 1) or changes in sugar levels (see Alternate Protocol 1), T7 RNA polymerase is produced, repression of the T7lac promoter is removed, and the target gene is transcribed. The specificity and high activity of T7 RNA polymerase guarantees that the cell’s protein synthesis factory will almost exclusively generate target protein from the target transcript, reaching levels comparable to that of total cell protein. To isotopically enrich recombinantly produced proteins, a fully defined medium is used which contains isotopically enriched sources of carbon, nitrogen, deuterium, or specific amino acids. The basic medium consists of sources of phosphate, nitrogen, sulfate, magnesium and trace metals and a primary carbon and energy source such as glucose or glycerol. This medium can be supplemented with either isotopically labeled amino acids or specific metabolic precursors to enrich only specific amino acids or types of amino acids (see Alternate Protocols 2, 3, 4, 5, and 6). The media are prepared fresh from autoclaved or filter-sterilized concentrated stock solutions as needed in proper growth vessels. Initial cultures grown to saturation in a noninducing medium are used for stable storage of the plasmid-containing cells, and also for inoculation of scaled-up volumes of enriched and labeled media for protein expression. For optimum growth to high density and production of large quantities of recombinant protein, it is important to obtain the highest aeration of the growth media possible by the use of baffled flasks and/or culture volumes 25% (or less) of the growth flask volume with rapid shaking (i.e., ∼300 rpm). Growth incubation temperature and carbon source will affect the time it takes to reach saturation of the cultures, and low temperature incubation (i.e., 18◦ C to 20◦ C) can be used to improve target protein solubility. Protein Engineering
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Pilot experiments using small cultures (i.e., 1.5 ml in 17 × 100–mm polypropylene tubes) should be done prior to scaling up to confirm proper growth and induction characteristics. Initial analysis includes: (1) titering of the culture on agar plates containing selective antibiotic or no antibiotic to determine the percent of total cells capable of forming colonies on selective antibiotic, and therefore still harboring viable target plasmid, (2) recording the absorbance at 600 nm in a spectrophotometer to determine when the culture reaches saturation, (3) measuring the pH to ensure that the cultures have not become too acidic for continued growth (i.e., pH 5.5 or lower) and (4) polyacrylamide gel electrophoresis of total cell proteins in sodium dodecyl sulfate (SDS-PAGE; UNIT 10.1) to directly observe appearance of over-expressed target protein as a function of induction time. See UNIT 5.3 for more details on optimizing growth and induction conditions for E. coli. NOTE: For an overview and details of general recombinant protein expression including expression strategies, secretion of proteins, and use of fusion tags, see Chapter 5. BASIC PROTOCOL 1
UNIFORM ISOTOPIC LABELING OF PROTEINS THROUGH EXPRESSION IN E. col i USING IPTG INDUCTION This protocol describes the heterologous expression of the protein of interest in transformed E. coli cells containing an expression vector which is regulated by the T7 lac promoter. The bacteria are grown in minimal medium containing isotopically enriched nutrients which cause the amino acids synthesized by the bacteria to contain isotopically 13 C-enriched carbon atoms and/or 15 N-enriched nitrogen atoms. This method relies on induction of the plasmid by the addition of isopropyl-1-thio-β-D-galactopyranoside (IPTG) to produce the protein of interest. An alternate protocol for protein production via auto-induction by carefully balancing the levels of glycerol and lactose is also described (see Alternate Protocol 1). This protocol describes the preparation of 1 liter of transfected E. coli cells in two 4-liter flasks in a shaker incubator at 37◦ C, monitoring cell growth by determining the optical density at 600 nm (OD600 ), and subsequent induction of protein production. Cells should not be grown for extended periods of time after induction because this leads to cell lysis and proteolysis by the released proteases. This protocol can be scaled up for use in a fermentor (UNIT 5.3).
Materials Lysogenic expression host, e.g., E. coli BL21 (DE3) transformed with a pET or similar vector containing a T7 lac promoter (EMD Biosciences) and the gene for the protein of interest (see Chapter 5) LB agar plate containing the appropriate antibiotic (APPENDIX 4A) Overnight medium (see recipe) Antibiotic solutions (see recipes) appropriate for expression host and vector Glucose-based minimal medium, isotopically enriched (see recipe) 0.5 M isopropyl-1-thio-β-D-galactopyranoside (IPTG; Research Products International): sterilize by passing through a 0.22-µm filter and store in 5-ml and 500-µl aliquots up to 6 months at −20◦ C 250-ml and 4-liter culture flasks, sterile and preferably baffled 37◦ C orbital shaker incubator 500-ml centrifuge tubes High speed refrigerated centrifuge with appropriate rotors (e.g., Beckman J2-HS, with JA 10 and JA 20 rotors) Incorporation of Isotopically Enriched Amino Acids
Additional reagents and equipment for performing SDS-PAGE (UNIT 10.1) and purifying recombinant proteins (see Chapter 6)
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1. Grow the expression host containing the gene for the protein of interest on an LB plate containing the appropriate antibiotic overnight at 37◦ C. 2. Pick a single colony and inoculate a 250-ml flask containing 50 ml of overnight medium and 0.05 to 0.1 ml of the appropriate antibiotic solution. Incubate the flask overnight at 37◦ C, with shaking at 250 to 300 rpm. 3. The following morning, determine the OD600 against water as a blank to verify that the culture has reached stationary phase with an OD600 >1.5. Dilute the sample 10-fold to ensure a reading between 0.1 and 1. This preparation can be used to create both 8% glycerol freezer stocks, which can be stored indefinitely at −80◦ C, as well as working stocks, which can be stored up to 2 weeks at 4◦ C. Pilot experiments using 1 to 4 ml of cultures can be performed to optimize growth temperature, IPTG concentration, and protein expression times before beginning largescale production with isotopically enriched medium.
4. Use 5 ml of the overnight culture to inoculate each of the two 4-liter baffled flasks containing 500 ml isotopically enriched glucose-based minimal medium equilibrated to 37◦ C and 0.5 to 1 ml of the appropriate antibiotic solution. Glucose-based minimal medium is prepared as indicated in Reagents and Solutions. The labeled sources of 13 C and 15 N can be used individually or in combination and are added depending on the desired nature of the labeling.
5. Incubate the flask 2 to 4 hr at 37◦ C, with shaking at 250 to 300 rpm until the bacterial culture reaches an OD600 of 0.6 to 0.8. 6. Remove an initial 1-ml sample from each flask and save for SDS-PAGE analysis as a control for uninduced cells, and either immediately treat as described in step 10 or freeze the sample until ready to perform step 10. 7. Add 0.5 ml of 0.5 M IPTG to each flask for a final concentration of 0.5 mM IPTG to induce protein expression. IPTG concentration can be vector and protein dependent. IPTG levels may be optimized in the 0.1 to 1 mM range using pilot experiments after step 3.
8. Incubate the flask 2 to 4 hr at 37◦ C with shaking at 250 to 300 rpm. Remove 1 ml from each culture and save for SDS-PAGE analysis. Check the final OD600 . The final OD600 is checked to make sure that protein expression did not lead to large amounts of cell death. An OD600 of 1 to 2 is expected. The sample will need to be diluted before taking a reading.
9. Harvest the cells by centrifuging the culture in 500-ml centrifuge tubes 20 min at 10,000 × g, 4◦ C. Decant the supernatant and store the cell pellet up to 6 months at −80◦ C. 10. Analyze the 1-ml samples taken before and after induction using SDS-PAGE (UNIT 10.1) to check for protein expression. If a distinct protein band in the induced sample is not observed, more pilot experiments need to be done.
11. Lyse the cells and purify the induced recombinant protein using the appropriate protocols (see Chapter 6).
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ALTERNATE PROTOCOL 1
UNIFORM ISOTOPIC LABELING OF PROTEINS THROUGH EXPRESSION IN E. coli USING AUTO-INDUCTION The mechanism of auto-induction arises from the way bacteria regulate the use of carbon and energy sources. In the presence of glucose, a primary energy source, uptake of lactose by the cells is prevented, and unintended induction is blocked. For cells continuing to grow to high density, glucose is depleted, and lactose uptake via lactose permease occurs. Therefore, in auto-inducing media, induction by lactose is controlled until such a time that cells reach a moderate to high density. The auto-induction stage will therefore occur when cell growth is near saturation resulting in a maximum amount of target protein production. Fully defined auto-inducing media have been formulated based on the original work of Studier (Studier, 2005), and can be used for 15 N and 13 C isotopic labeling of high levels of target protein produced in high density cultures (Tyler et al., 2005). The following protocol describes formulation of 1 liter of fully defined auto-induction medium supplemented with uniformly 15 N- or 13 C- ([U-15 N]- or [U-13 C]-) labeled compounds. Use of this medium induces protein production via auto-induction by carefully balancing the levels of glycerol and lactose in the minimal medium (Studier, 2005). Uniform labeling of target protein with 15 N can be accomplished by including [U-15 N]labeled (NH4 )2 SO4 or NH4 Cl in the medium. Uniform isotopic labeling with 13 C is slightly more complicated due to the use of several carbon sources in auto-inducing media. [U-13 C]-Glucose would be an economical and primary candidate as a 13 C source, however, it prevents auto-induction. [U-13 C]-labeled glycerol is the next obvious choice because it is used by the cells as efficiently as glucose but does not prevent auto-induction. Lactose, the inducing agent, will also contribute some carbon for incorporation into the target protein when glucose becomes depleted. Therefore, it is prudent to optimize the transfer of 13 C to target protein from [U-13 C]-glycerol while minimizing carbon transfer from lactose. The recipes and growth methods given below should provide good starting points for auto-induction of uniformly-labeled target proteins with 15 N, 13 C, or both 15 N and 13 C isotopes for NMR. This protocol describes the preparation of 1 liter of transfected E. coli cells in two 4-liter flasks in a shaker incubator at 37◦ C, monitoring cell growth by determining the optical density at 600 nm (OD600 ), and subsequent induction of protein production. Cells should not be grown for extended periods of time after induction because this leads to cell lysis and proteolysis by the released proteases. This protocol can be scaled up for use in a fermentor (UNIT 5.3).
Additional Materials (also see Basic Protocol 1) Auto-induction minimal medium, isotopically enriched (see recipe) 1. Grow the expression host containing the gene for the protein of interest on an LB plate containing the appropriate antibiotic overnight at 37◦ C. 2. Pick a single colony and inoculate a 250-ml flask containing 50 ml of overnight medium and 0.05 to 0.1 ml of the appropriate antibiotic solution. Incubate the flask overnight at 37◦ C, with shaking at 250 to 300 rpm. 3. The following morning, determine the OD600 against water as a blank to verify that the culture has reached stationary phase with an OD600 >1.5. Dilute the sample 10-fold to ensure a reading between 0.1 and 1. Incorporation of Isotopically Enriched Amino Acids
This preparation can be used to create both 8% glycerol freezer stocks, which can be stored indefinitely at −80◦ C, as well as working stocks, which can be stored up to 2 weeks at 4◦ C.
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Pilot experiments using 1 to 4 ml of cultures can be performed to optimize growth temperature and protein expression times before beginning large-scale production with isotopically enriched medium.
4. Use ∼5 ml of this initial culture to inoculate 500 ml isotopically enriched autoinduction minimal medium equilibrated to the desired growth temperature (usually 37◦ C) in each of two 4-liter flasks. The labeled sources of 13 C and 15 N can be used individually or in combination and are added depending on the desired nature of the labeling (see recipe in Reagents and Solutions section).
5. Incubate 6 to 8 hr with shaking at 250 to 300 rpm until they are visibly cloudy. 6. Remove and freeze 1 ml for SDS-PAGE analysis (UNIT 10.1) as a control when verifying overexpression of protein as a function of induction time. 7. Grow 500-ml cultures to high density (OD600 >1.5) with shaking at 300 rpm at the desired temperature. Note that the growth temperature will affect the time it takes to reach saturation of the cultures, and low-temperature incubation (i.e., 18◦ to 25◦ C) will result in saturation times ≥24 hr. Following the OD600 of these cultures will help to determine saturation behavior under the given conditions. Remove 1 ml from each culture and save for SDS-PAGE analysis.
8. Harvest the cells by centrifugation for 20 min at 10,000 × g at 4◦ C. Cell pellets should be stored up to 6 months at −80◦ C.
9. Analyze the 1-ml samples taken after induction using SDS-PAGE (UNIT 10.1) to check for protein expression. If a distinct protein band in the induced sample is not observed, more pilot experiments need to be done.
10. Lyse the cells and purify protein using appropriate protocols for the induced protein (see Chapter 6).
UNIFORM ISOTOPIC LABELING OF SPECIFIC CARBONS IN PROTEINS EXPRESSED IN E. coli This protocol describes the heterologous expression of the protein of interest in the presence of a minimal medium to selectively label particular carbons in the recombinant protein. The pattern of carbon labeling is determined by which carbons in the carbon source are isotopically enriched. Choices include: (a) 2-13 C-glycerol or 1,3-13 C2 glycerol to label alternating carbons in the protein, (b) 1-13 C- sodium acetate or 2-13 Csodium acetate to label alternating carbons, (c) 3-13 C-pyruvate to label the methyl groups on Ala, Ile, Leu, and Val residues, or (d) the use of [3,3 -13 C2 ]-α-ketoisovalerate and [3-13 C]-α-ketobutyrate with 12 C glucose to label only the Leu, Val, and Ile-δ1 methyl groups (Hajduk et al., 2000). This protocol relies on induction of protein production by the addition of isopropyl-1-thio-β-D-galactopyranoside (IPTG) to produce the protein of interest. An alternative method for protein production via auto-induction involves carefully balancing the levels of glycerol and lactose and is described in Alternate Protocol 1. This protocol describes the preparation of 1 liter of transfected E. coli cells in two 4-liter flasks in a shaker incubator at 37◦ C, monitoring cell growth by determining the optical density at 600 nm (OD600 ), and subsequent induction of protein production. Cells should not be grown for extended periods of time after induction because this leads to cell lysis and proteolysis by the released proteases. This protocol can be scaled up for use in a fermentor (UNIT 5.3).
ALTERNATE PROTOCOL 2
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Additional Materials (also see Basic Protocol 1) Specific carbon source-based minimal medium, isotopically enriched (see recipe) 500× α-ketoisovalerate and α-ketobutyrate (see recipe), labeled 1. Grow the expression host containing the gene for the protein of interest on an LB plate containing the appropriate antibiotic overnight at 37◦ C. 2. Pick a single colony and inoculate a 250-ml flask containing 50 ml of overnight medium and 0.05 to 0.1 ml of the appropriate antibiotic solution. Incubate the flask overnight at 37◦ C, with shaking at 250 to 300 rpm. 3. The following morning, determine the OD600 against water as a blank to verify that the culture has reached stationary phase with an OD600 >1.5. Dilute the sample 10-fold to ensure a reading between 0.1 and 1. This preparation can be used to create both 8% glycerol freezer stocks, which can be stored indefinitely at −80◦ C, as well as working stocks, which can be stored up to 2 weeks at 4◦ C.
4. Centrifuge 10 ml of the overnight culture for 5 min at 10,000 × g. Discard the supernatant. 5. Resuspend the cell pellet in 10 ml of isotopically enriched specific carbon sourcebased medium. Specific carbon source-based minimal media are prepared as indicated in Reagents and Solutions. Alternate labeling can either be accomplished with 2-13 C-glycerol, 1,3-13 C2 glycerol, 1-13 C sodium acetate or 2-13 C sodium acetate. Cell growth and protein induction is faster in the glycerol medium due to a lag phase when switching to acetate as the carbon source. To selectively label Ala, Ile, Leu, and Val methyl groups, 3-13 C pyruvate is used, or for more specific methyl group labeling (Val, Leu, and Ile-δ 1 ) cells are grown on 12 C glucose, and [dimethyl-13 C2 ]-α-ketoisovalerate, and [methyl-13 C]-α-ketobutyrate are added just prior to induction. The incorporation of 15 N can also be accomplished by using isotopically enriched 15 NH4 Cl in the minimal medium.
6. Use 5 ml of the resuspended pellet to inoculate each of the two 4-liter baffled flasks containing 500 ml isotopically enriched specific carbon source-based minimal medium equilibrated to 37◦ C and 0.5 to 1 ml of the appropriate antibiotic solution. 7. Incubate the flask at 37◦ C with shaking at 250 to 300 rpm until the bacterial culture reaches an OD600 of 0.6 to 0.8. This should take 2 to 4 hr at 37◦ C for glycerol or glucose, longer for other carbon sources.
8. Optional: If using [3,3 -13 C2 ]-α-ketoisovalerate and [3-13 C]-α-ketobutyrate in combination with 12 C glucose, at this point add 1 ml of the 500× stock solutions of the labeled α-ketoisovalerate and α-ketobutyrate to each flask and incubate for an additional 30 min. 9. Remove an initial 1-ml sample from each flask and save for SDS-PAGE analysis as a control for uninduced cells and either immediately treat as described in step 14 or freeze the sample until ready to perform step 14. 10. Add 0.1 to 1.0 ml of 0.5 M IPTG to each flask for a final concentration of 0.1 to 1.0 mM IPTG to induce protein expression. Incorporation of Isotopically Enriched Amino Acids
11. Incubate the flask at 37◦ C in an incubator shaker at 250 to 300 rpm for 2 to 4 hr after induction. 12. Remove 1 ml from each culture and save for SDS-PAGE analysis. Check final OD600 .
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13. Harvest the cells by centrifuging the culture in 500-ml centrifuge tubes for 20 min at 10,000 × g, 4◦ C. Decant the supernatant and store the cell pellet at −80◦ C. 14. Analyze the 1-ml samples taken before and after induction using SDS-PAGE (UNIT 10.1) to check for protein expression. If a distinct protein band in the induced sample is not observed, more pilot experiments need to be done.
15. Lyse the cells and purify protein using appropriate protocols for the induced protein (Chapter 6).
UNIFORM DEUTERATION WITH CONCOMITANT 13 C AND 15 N INCORPORATION IN PROTEINS EXPRESSED IN E. coli
ALTERNATE PROTOCOL 3
This protocol describes the heterologous expression of the protein of interest in the presence of a minimal medium made with 2 H2 O to deuterate the protein and either (1) uniformly 13 C, 15 N and 2 H label all the amino acids present in the recombinant protein, (2) uniformly 13 C and 15 N label the protein and achieve close to complete deuteration at the amide and Cα carbons and >80% deuteration at the Cβ carbons, (3) 13 C-label and partially protonate (13 C2 H2 H) the methyl groups of aliphatic residues Ala, Val, Leu and Ile, or (4) uniformly 13 C-label the protein, with only the Val, Leu, and Ile-δ1 methyl groups remaining highly protonated (Gardner et al., 1997; Goto et al., 1999). The choice among these four separate labeling schemes is dependent on the size of the protein and the intended NMR experiments. Initial overnight stocks are first conditioned to the carbon source that will be utilized and then grown in deuterated media. This protocol describes the preparation of 1 liter of transfected E. coli cells in two 4-liter flasks in a shaker incubator at 37◦ C, monitoring cell growth by determining the optical density at 600 nm (OD600 ), and subsequent induction of protein production. Cells should not be grown for extended periods of time after induction because this leads to cell lysis and proteolysis by the released proteases. This protocol can be scaled up for use in a fermentor (UNIT 5.3).
Additional Materials (also see Basic Protocol 1) Carbon source conditioning medium (see recipe) Deuterated minimal medium (see recipe) 500× [3,3 -2 H2 ]-13 C-α-ketobutyrate and [3-2 H]-13 C-α-ketoisovalerate stock solutions (see recipe), if required 250-ml centrifuge tubes, optional 30-ml centrifuge tubes 1. Grow the expression host containing the gene for the protein of interest on an LB plate containing the appropriate antibiotic overnight at 37◦ C. 2. Pick a single colony and inoculate a 250-ml flask containing 50 ml of overnight medium equilibrated to 37◦ C and 0.05 to 0.1 ml of the appropriate antibiotic solution. Incubate the flask overnight at 37◦ C, with shaking at 250 to 300 rpm. 3. The following morning, determine the OD600 against water as a blank to verify that the culture has reached stationary phase with an OD600 >1.5. Dilute the sample 10-fold to ensure a reading between 0.1 and 1. This preparation can be used to create both 8% glycerol freezer stocks, which can be stored indefinitely at −80◦ C, as well as working stocks, which can be stored up to 2 weeks at 4◦ C.
4. Centrifuge 10 ml of the overnight culture for 5 min at 10,000 × g. Discard the supernatant. Protein Engineering
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5. Resuspend the cell pellet in 10 ml of carbon source conditioning medium and use 5 ml to inoculate each of two 250-ml baffled flasks containing 50 ml carbon source conditioning medium and 0.05 to 0.1 ml of the appropriate antibiotic solution. Carbon source conditioning medium is prepared as indicated in Reagents and Solutions. The carbon source selected depends on the ultimate choice of medium for isotopic labeling. The cells should only be switched serially to a specific carbon source and then to a deuterated medium to ensure optimal cell growth and protein expression.
6. Incubate the flask at 37◦ C in an incubator shaker at 200 to 250 rpm until the bacterial culture reaches an OD600 of 0.6. This should take 2 to 4 hr at 37◦ C if using glucose or glycerol, longer if using other carbon sources.
7. Centrifuge each culture for 5 min at 10,000 × g and discard the supernatant. 8. Resuspend each cell pellet in 500 ml of deuterated minimal medium in a 4-liter flask along with 0.5 to 1 ml of the appropriate antibiotic solution. The deuterated minimal medium is prepared as indicated in Reagents and Solutions. The carbon source depends on the type of 13 C labeling to be accomplished. For full 2 H and 13 C labeling, either U-13 C, 2 H-D-glucose or U-13 C, 2 H sodium acetate is used. To achieve full 13 C labeling and almost complete deuteration of the amides and Cα carbons and >80% deuteration of the Cβ carbons, 13 C3 -pyruvate can be used (the methyl groups in L, V, I, and A will be protonated). To 13 C label only the methyl groups of A, V, L, and I with partial protonation of the methyl groups, 3-13 C-pyruvate is used. For more selective protonation, U-13 C,2 H-D-glucose is used initially with addition of [3,3 -2 H2 ]-13 C-α-ketobutyrate and [3-2 H]-13 C-α-ketoisovalerate just prior to induction. The incorporation of 15 N can also be accomplished by using isotopically enriched 15 NH4 Cl in the minimal medium.
9. Incubate the flask 2 to 4 hr at 37◦ C in an incubator shaker at 200 to 250 rpm until the bacterial culture reaches an OD600 of 0.6. 10. Optional: If using [3,3 -2 H2 ]-13 C-α-ketobutyrate and [3-2 H]-13 C-α-ketoisovalerate in combination with U-13 C,2 H glucose, add 1 ml of the 500× stock solution of the labeled α-ketoisovalerate and α-ketobutyrate to each flask and incubate for an additional 30 min. 11. Remove an initial 1-ml sample from each flask and save for SDS-PAGE analysis as a control for uninduced cells and either immediately treat as described in step 15 or freeze the sample until ready to perform step 15. 12. Add 0.1 to 1.0 ml of 0.5 M IPTG to each flask for a final concentration of 0.1 to 1.0 mM IPTG to induce protein expression. 13. Incubate the flask 2 to 4 hr at 37◦ C, with shaking at 250 to 300 rpm. Remove 1 ml from each culture and save for SDS-PAGE analysis. Check final OD600 . 14. Harvest the cells by centrifuging the culture in 250-ml or 500-ml centrifuge tubes for 20 min at 10,000 × g, 4◦ C. Decant the supernatant and store the cell pellet at −80◦ C. 15. Analyze the 1-ml samples taken before and after induction using SDS-PAGE (UNIT 10.1) to check for protein expression. If a distinct protein band in the induced sample is not observed, more pilot experiments need to be done. Incorporation of Isotopically Enriched Amino Acids
16. Lyse the cells and purify protein using the appropriate protocols for the induced protein (Chapter 6).
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SELECTIVE ISOTOPIC LABELING OF PROTEINS THROUGH EXPRESSION IN BACTERIA USING THE “TEASE” LABELING SCHEME
ALTERNATE PROTOCOL 4
This protocol describes the heterologous expression of the protein of interest in the presence of a minimal medium to isotopically label the carbon atoms and nitrogen atoms of ten predominantly hydrophobic amino acids (Gly, Cys, Trp, Phe, Tyr, Ala, Val, Leu, Ser and His) present in the recombinant protein. This protocol describes the preparation of 1 liter of transfected E. coli cells in two 4-liter flasks in a shaker incubator at 37◦ C, monitoring cell growth by determining the optical density at 600 nm (OD600 ), and subsequent induction of protein production. Cells should not be grown for extended periods of time after induction because this leads to cell lysis and proteolysis by the released proteases. This protocol can be scaled up for use in a fermentor (UNIT 5.3).
Additional Materials (also see Basic Protocol 1) TEASE minimal medium (see recipe) 250-ml (optional) centrifuge tubes 1. Grow the expression host containing the gene for the protein of interest on an LB plate containing the appropriate antibiotic overnight at 37◦ C. 2. Pick a single colony and inoculate a 250-ml flask containing 50 ml of overnight medium equilibrated to 37◦ C and 0.05 to 0.1 ml of the appropriate antibiotic solution. Incubate the flask overnight at 37◦ C, with shaking at 250 to 300 rpm. 3. The following morning, determine the OD600 against water as a blank to verify that the culture has reached stationary phase with an OD600 >1.5. Dilute the sample 10-fold to ensure a reading between 0.1 and 1. This preparation can be used to create both 8% glycerol freezer stocks, which can be stored indefinitely at −80◦ C, as well as working stocks, which can be stored up to 2 weeks at 4◦ C.
4. Centrifuge 10 ml of the overnight culture for 5 min at 10,000 × g. Discard the supernatant. 5. Resuspend the cell pellet in 10 ml of TEASE minimal medium. 6. Use 5 ml of the resuspended pellet to inoculate each of the two 4-liter baffled flasks containing 500 ml 37◦ C TEASE minimal medium and 0.5 to 1 ml of the appropriate antibiotic solution. TEASE minimal medium is prepared using isotopically 13 C-labeled glycerol as indicated in Reagents and Solutions. The incorporation of 15 N can also be accomplished by using isotopically enriched 15 NH4 Cl in the salts solution for the TEASE minimal medium.
7. Incubate the flask 2 to 4 hr at 37◦ C in an incubator shaker at 250 to 300 rpm until the bacterial culture reaches an OD600 of 0.6 to 0.8. 8. Remove an initial 1-ml sample from each flask and save for SDS-PAGE analysis as a control for uninduced cells and either immediately treat as described in step 13 or freeze the sample until ready to perform step 13. 9. Add 0.1 to 1.0 ml of 0.5 M IPTG to each flask for a final concentration of 0.1 to 1.0 mM IPTG to induce protein expression. 10. Incubate the flasks at 37◦ C in an incubator shaker at 250 to 300 rpm for 2 to 4 hr after induction. 11. Remove 1 ml from each culture and save for SDS-PAGE analysis (UNIT 10.1). Check final OD600 .
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12. Harvest the cells by centrifuging the culture in 250- or 500-ml centrifuge tubes for 20 min at 10,000 × g, 4◦ C. Decant the supernatant and store the cell pellet up to 6 months at −80◦ C. 13. Analyze the 1-ml samples taken before and after induction using SDS-PAGE (UNIT 10.1) to check for protein expression. If a distinct protein band in the induced sample is not observed, more pilot experiments need to be done.
14. Lyse the cells and purify protein using appropriate protocols for the induced protein (Chapter 6). ALTERNATE PROTOCOL 5
PREPARATION OF SELECTIVELY LABELED PROTEINS IN E. col i BY INCORPORATION OF SPECIFICALLY LABELED AMINO ACIDS This protocol describes the heterologous expression of the protein of interest in the presence of selectively labeled amino acids added to the minimal medium just prior to induction. This protocol describes the preparation of 1 liter of transfected E. coli cells in two 4-liter flasks in a shaker incubator at 37◦ C, monitoring cell growth by determining the optical density at 600 nm (OD600 ), and subsequent induction of protein production. Cells should not be grown for extended periods of time after induction because this leads to cell lysis and proteolysis by the released proteases. This protocol can be scaled up for use in a fermentor (UNIT 5.3).
Additional Materials (also see Basic Protocol 1) 50× amino acid stock solution (see recipe) 30-ml and 250-ml (optional) ml centrifuge tubes Grow bacterial cells and express labeled protein 1. Grow the expression host containing the gene for the protein of interest on an LB plate containing the appropriate antibiotic overnight at 37◦ C. 2. Pick a single colony and inoculate a 250-ml flask containing 50 ml of overnight medium equilibrated to 37◦ C and 0.05 to 0.1 ml of the appropriate antibiotic solution. Incubate the flask overnight at 37◦ C, with shaking at 250 to 300 rpm. 3. The following morning, determine the OD600 against water as a blank to verify that the culture has reached stationary phase with an OD600 >1.5. Dilute the sample 10-fold to ensure a reading between 0.1 and 1. This preparation can be used to create both 8% glycerol freezer stocks, which can be stored indefinitely at −80◦ C, as well as working stocks, which can be stored up to 2 weeks at 4◦ C.
4. Centrifuge 10 ml of the overnight culture for 5 min at 10,000 × g. Discard the supernatant. Resuspend the cell pellet in 10 ml of glucose-based minimal medium and use 5 ml to inoculate each of the two 4-liter baffled flasks containing 500 ml glucose-based minimal medium and 0.5 to 1 ml of the appropriate antibiotic solution equilibrated to 37◦ C. The growth medium is not isotopically enriched.
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5. Incubate the flask 2 to 4 hr at 37◦ C in an incubator shaker at 200 to 250 rpm until the bacterial culture reaches an OD600 of 0.6 to 0.8. At this point add 10 ml of 50×stock solution (with labeled amino acid) to each flask and incubate for 30 min. 6. Remove an initial 1-ml sample from each flask and save for SDS-PAGE analysis as a control for uninduced cells and either immediately treat as described in step 10 or freeze the sample until ready to perform step 10. Current Protocols in Protein Science
7. Add 0.1 to 1.0 ml of 0.5 M IPTG to each flask for a final concentration of 0.1 to 1.0 mM IPTG to induce protein expression. 8. Incubate the flasks at 37◦ C in an incubator shaker at 250 to 300 rpm for 2 to 4 hr after induction. Remove 1 ml from each culture and save for SDS-PAGE analysis (UNIT 10.1). Check final OD600 . 9. Harvest the cells by centrifuging the culture in 250- or 500-ml centrifuge tubes for 20 min at 10,000 × g, 4◦ C. Decant the supernatant and store the cell pellet up to 6 months at −80◦ C. 10. Analyze the 1-ml samples taken before and after induction using SDS-PAGE (UNIT 10.1) to check for protein expression. If a distinct protein band in the induced sample is not observed, more pilot experiments need to be done.
11. Lyse the cells and purify protein using appropriate protocols for the induced protein (Chapter 6).
PREPARATION OF 19 F-LABELED PROTEINS IN E. col i USING 19 F-LABELED AROMATIC AMINO ACIDS
ALTERNATE PROTOCOL 6
Currently, there are several synthetic methods for production of fluorine-containing analogs of amino acids, and therefore peptides and proteins. The incorporation of these fluorinated materials into biopolymeric materials has been accomplished through various strategies including chemical synthesis and biosynthesis by organisms. This protocol describes the biosynthetic incorporation of 19 F-labeled aromatic amino acids (fluorotyrosine, fluorotryptophan or fluorophenylalanine) in a protein of interest. This protocol describes the preparation of 1 liter of transfected E. coli cells in two 4-liter flasks in a shaker incubator at 37◦ C, monitoring cell growth by determining the optical density at 600 nm (OD600 ), and subsequent induction of protein production. Cells should not be grown for extended periods of time after induction because this leads to cell lysis and proteolysis by the released proteases. This protocol can be scaled up for use in a fermentor (UNIT 5.3).
Additional Materials (also see Basic Protocol 1) Fluorine-labeling medium (see recipe) Fluorinated amino acid stock: 50× fluorophenylalanine or 50× fluorotryptophan or 50× fluorotyrosine (see recipes) 2-liter baffled flasks, sterile 30-ml and 250-ml (optional) centrifuge tubes 1. Grow the expression host containing the gene for the protein of interest on an LB plate containing the appropriate antibiotic overnight at 37◦ C. 2. Pick a single colony and inoculate a 250-ml flask containing 50 ml of overnight medium equilibrated to 37◦ C and 0.05 to 0.1 ml of the appropriate antibiotic solution. Incubate the flask overnight at 37◦ C, with shaking at 250 to 300 rpm. 3. The following morning, determine the OD600 against water as a blank to verify that the culture has reached stationary phase with an OD600 >1.5. Dilute the sample 10-fold to ensure a reading between 0.1 and 1. This preparation can be used to create both 8% glycerol freezer stocks, which can be stored indefinitely at −80◦ C, as well as working stocks, which can be stored up to 2 weeks at 4◦ C.
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4. Centrifuge 10 ml of the overnight culture for 5 min at 10,000 × g. Discard the supernatant. 5. Resuspend the cell pellet in 10 ml of fluorine-labeling medium and use 5 ml to inoculate each of the two 4-liter baffled flasks containing 500 ml fluorine-labeling medium and 0.5 to 1 ml of the appropriate antibiotic solution. The growth medium is not isotopically enriched and contains the aromatic amino acids which will not be fluorinated.
6. Incubate the flask 2 to 4 hr at 37◦ C in an incubator shaker at 200 to 250 rpm until the bacterial culture reaches an OD600 of 0.6 to 0.8. At this point, add 10 ml of a 50× stock solution of the fluorinated amino acid to each flask and incubate for 30 min. 7. Remove an initial 1-ml sample from each flask and save for SDS-PAGE analysis as a control for uninduced cells and either immediately treat as described in step 13 or freeze the sample until ready to perform step 13. 8. Add 0.1 to 1.0 ml of 0.5 M IPTG to each flask for a final concentration of 0.1 to 1.0 mM IPTG to induce protein expression. 9. Incubate the flasks at 37◦ C in an incubator shaker at 250 to 300 rpm for 2 to 4 hr after induction. 10. Remove 1 ml from each culture and save for SDS-PAGE analysis (UNIT 10.1). Check final OD600 . 11. Harvest the cells by centrifuging the culture in 250- or 500-ml centrifuge tubes for 20 min at 10,000 × g, 4◦ C. 12. Decant the supernatant and store the cell pellet up to 6 months at −80◦ C. 13. Analyze the 1-ml samples taken before and after induction using SDS-PAGE (UNIT 10.1) to check for protein expression. If a distinct protein band in the induced sample is not observed, more pilot experiments need to be done.
14. Lyse the cells and purify protein using appropriate protocols for the induced protein (Chapter 6). BASIC PROTOCOL 2
Incorporation of Isotopically Enriched Amino Acids
EXPRESSION OF UNIFORMLY 15 N OR 13 C/15 N LABELED RECOMBINANT PROTEINS USING THE GATEWAY TECHNOLOGY AND THE BACULODIRECT BACULOVIRUS EXPRESSION SYSTEMS For several reasons protein production methods using the baculovirus-based expression systems have become popular for producing heterologous proteins in insect cells. The baculovirus expression systems allow for easy production of functional heterologous proteins because the insect cells possess the necessary cellular components to facilitate proper folding and/or post-translational modifications of eukaryotic proteins. An outline of the principles involved in the Gateway Technology (Invitrogen) cloning method for protein expression is described in UNIT 5.17. For an overview of the baculovirus expression system and recombinant protein expression in insect cells refer to UNITS 5.4 & 5.5. There are only a limited number of examples of labeled amino acid incorporation into proteins expressed in baculovirus-infected Sf 9 cells (DeLange et al., 1998; Creemers et al., 1999; Br¨uggert et al., 2003; Strauss et al., 2003). This unit describes the procedure for cloning the gene of interest, using the Gateway Technology, into the pENTR Directional TOPO vector (see Invitrogen Manual 25-0434, Version E, 2004); the Gateway LR recombination reaction for transferring the gene of interest into the baculovirus genome and transfection of insect cells (Sf 9; see Invitrogen Manual
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25-0650, Version F); and the method for expressing unlabeled or uniformly labeled 15 N or 13 C/15 N recombinant proteins (Strauss et al., 2005) in insect cells (Spodoptera frugiperda) using the commercially available BioExpress 2000 insect cell medium (Cambridge Isotopes Laboratories).
Materials DNA template containing the gene of interest Appropriate primers to produce the PCR product Thermostable, proofreading polymerase, e.g., AccuPrime Taq high fidelity DNA polymerase kit (Invitrogen): includes 10× PCR buffer pENTR/D-TOPO Cloning Kit (Invitrogen): includes One Shot Top10 chemically competent E. coli, salt solution, and SOC medium LB agar plates containing 50 µg/ml kanamycin (Research Products International or see APPENDIX 4A) LB medium (APPENDIX 4A) BaculoDirect linear DNA, 300 ng/vial (Invitrogen) TE buffer, pH 8.0 (APPENDIX 2E) LR Clonase II enzyme mix (Invitrogen) 2 mg/µl proteinase K (Invitrogen) Sf 9, Sf 21, or High Five insect cells (Invitrogen) Complete growth medium Sf-900 II SFM (Invitrogen) Grace’s Insect Medium, unsupplemented (Invitrogen) Cellfectin Reagent (Invitrogen) Complete growth medium Sf-900 II SFM with antibiotics (Invitrogen) Ganciclovir (Invitrogen) BioExpress-2000 insect cell medium: unlabeled (Cambridge Isotopes Laboratories) or BioExpress-2000 insect cell medium: U-15 N, 98% (Cambridge Isotopes Laboratories) or BioExpress-2000 insect cell medium: U-13 C 92-98%, U-15 N, 92-95% (Cambridge Isotopes Laboratories) or BioExpress-2000 insect cell medium: 13 C/15 N for L-methionine, L-leucine, L-isoleucine, and L-tyrosine (Cambridge Isotopes Laboratories) 25◦ C, 27◦ C, 37◦ C, and 42◦ C water baths 37◦ C orbital shaker incubator 27◦ C and 37◦ C incubators 6-well tissue culture plates Microscope Plastic bag and paper towels 50-ml centrifuge tubes, sterile 500-ml Erlenmeyer flasks Additional reagents and equipment for PCR (APPENDIX 4J), agarose gel electrophoresis (APPENDIX 4F), restriction analysis (APPENDIX 4I; optional), DNA plasmid miniprep (APPENDIX 4C), DNA quantitation (APPENDIX 4K), determination of plaque forming units (UNIT 5.5), and cell quantitation using a hemacytometer (APPENDIX 3C) Generate blunt-end PCR product 1. Generate a blunt-end PCR product of the gene of interest using appropriate primers and any suitable thermostable proofreading DNA polymerase (see APPENDIX 4J for PCR methodology). For a detailed explanation of the different TOPO vectors and PCR primers, see the manufacturer’s instruction manual 25-0434, Version E.
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2. Purify and verify the PCR-amplified product by agarose gel electrophoresis (APPENDIX 4F) and DNA sequencing. Typically, the DNA is sent to a sequencing facility.
3. Perform the 6-µl TOPO cloning reaction using the purified PCR product and the appropriate TOPO vector kit. Combine
0.5 µl to 4 µl fresh PCR product (step 1) 1 µl salt solution supplied with cloning kit Sterile H2 O to a final volume of 5 µl 1 µl TOPO vector. 4. Mix the reaction gently, incubate 5 min at room temperature (22◦ C to 23◦ C), and then place the reaction mix on ice. A 0.5:1 to 2.0:1 molar ratio of PCR product to TOPO vector must be used to obtain high TOPO cloning efficiency. For PCR products >1 kb or multiple PCR products, the reaction time may be increased up to 30 min to obtain more colonies. The TOPO cloning reaction can be stored overnight at −20◦ C. For a detailed explanation of TOPO TA cloning see Invitrogen’s TOPO TA Cloning instruction manual (25-0184), currently version U.
Transform the PCR product into E. coli 5. Add 2 µl TOPO cloning reaction (from step 3) into a vial of One Shot chemically competent E. coli cells. Mix gently. Do not pipet the mixture to mix. 6. Incubate the vial of cells on ice 5 to 30 min. 7. Heat shock the cells 30 sec at 42◦ C. 8. Transfer the cells to ice. 9. Add 250 µl SOC medium at room temperature. 10. Cap the tube and incubate the cells horizontally 1 hr at 37◦ C, with shaking at 200 rpm. 11. Plate 50 to 200 µl of the transformed cells on a warm LB/kanamycin plate and incubate overnight at 37◦ C. 12. Grow individual colonies in 5 ml LB and isolate (using DNA miniprep methods; APPENDIX 4C) and quantitate (APPENDIX 4K) plasmid DNA. Analyze by PCR (APPENDIX 4J) or restriction analysis (APPENDIX 4I), and verify the sequence of the construct to confirm the orientation.
Perform recombination reaction 13a. For each reaction mix: Combine the following components in 1.5-ml microcentrifuge tubes at room temperature. 1-2 µl DNA from step 12 (50 to 150 ng/µl in TE buffer, pH 8.0) 5 µl BaculoDirect linear DNA (150 ng/µl) TE Buffer, pH 8.0 to a final volume of 8 µl. 13b. For the positive control: Combine the following components in 1.5-ml microcentrifuge tubes at room temperature. Incorporation of Isotopically Enriched Amino Acids
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1 µl pENTR/CAT control (100 ng/µl) 5 µl BaculoDirect linear DNA (150 ng/µl) 2 µl TE Buffer, pH 8.0 The control DNA comes with the cloning kit. Current Protocols in Protein Science
14. Thaw the LR Clonase II enzyme mix on ice, and vortex briefly. 15. To each sample and the positive control, add 2 µl LR Clonase II enzyme mix. Mix well by tapping the tube gently. Do not pipet the mixture or vortex to mix 16. Incubate the reactions 1 hr at 25◦ C. 17. Add 1 µl of proteinase K solution to the reaction and incubate 10 min at 37◦ C. This is the LR recombination reaction.
Prepare Sf 9 or Sf 21 cells 18. Seed 8 × 105 Sf 9 cells in log phase of growth in 2 ml complete growth medium in a 6-well plate. Distribute the cells evenly in all the wells by gently tapping the plate. 19. Incubate the cells 1 hr at 27◦ C to allow the cells to adhere to the plate. Observe the cells under a microscope to verify that the cells are attached.
Transfect Sf 9 or Sf 21 cells 20. Prepare the following mixtures in sterile microcentrifuge tubes: a. Transfection mixture A 5 µl LR recombination reaction (from step 17) 100 µl Grace’s insect medium (unsupplemented) b. Transfection mixture B 6 µl Cellfectin reagent 100 µl Grace’s insect medium (unsupplemented). The amounts of cells, Cellfectin reagent, and viral DNA are optimized for one well in a 6-well tissue culture plate. The entire volume (∼1000µl) from steps 20 to 22 is for 1 well.
21. Combine transfection mixture A and transfection mixture B. Mix gently by inversion, and incubate the mixture 15 to 45 min at room temperature. 22. Add 800 µl of Grace’s insect medium (unsupplemented) to the transfection mix. Mix gently by inversion. 23. Remove the medium by pipetting from the cells in the wells of the tissue culture plates. Wash the cells with fresh Grace’s insect medium (unsupplemented). Remove the wash medium. 24. Add the entire amount of the transfection mix dropwise to the cells. The drops must be distributed evenly over the cells.
25. Incubate the cells 5 hr at 27◦ C. 26. Remove the transfection mixture by pipetting and add 2 ml of complete growth medium with antibiotics and 100 mM ganciclovir to each well. 27. Place the plates in a plastic bag with moist paper towels to prevent loss of moisture, and incubate 96 hr at 27◦ C. The transfected cells should show signs of viral infection (cell lysis, disruption, and clearing in the monolayer) in ∼96 hr.
28. Collect the medium from each of the wells and centrifuge 5 min at 4000 × g. Check the supernatant containing the recombinant baculovirus stock for plaque forming units (pfus; see UNIT 5.5) and store at 4◦ C, protected from light. This is the P1 viral stock. Generally the P1 viral stock is of low titer (1 × 10 5 to 1 × 10 6 plaque forming units (pfu)/ml. Hence, a high titer P2 stock (5 × 10 7 to 1 × 10 8 pfu/ml) must be obtained to use for protein expression.
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29. Seed 8 × 105 Sf 9 (or Sf 21) cells in 2 ml complete growth medium with 100 µM ganciclovir in a 6-well plate. Distribute the cells evenly in all the wells by gently tapping the plate. 30. Incubate the cells for 1 hr at 27◦ C to allow the cells to adhere to the plate. Observe the cells under a microscope to verify that they are attached. 31. Add 5 µl of P1 viral stock (from step 28) into each well. Place the plates in a plastic bag with moist paper towels to prevent loss of moisture, and incubate 72 hr at 27◦ C. 32. Collect the medium from each of the wells and centrifuge for 5 min at 4000 × g. Check the supernatant containing the recombinant baculovirus stock for plaque forming units (pfus; see UNIT 5.5) and store at 4◦ C, protected from light. This is the P2 viral stock.
Express labeled recombinant protein 33. Preculture Sf 9, Sf 21, or High Five cells in 100 ml complete growth medium Sf-900 II in a 500-ml Erlenmeyer flask for ∼3 days at 27◦ C, with shaking at 90 rpm to a final cell density of ∼4–5 × 106 (determined using a hemacytometer; see APPENDIX 3C). 34. Inoculate five 500-ml Erlenmeyer flasks, each containing 100 ml Sf-900 II medium, with 10 ml of cell culture from step 33. Incubate cells ∼3 days at 27◦ C, with shaking at 90 rpm, or until the cell density reaches 4 × 106 cells/ml (determined using a hemacytometer; see APPENDIX 3C). 35. Remove 380 ml of the medium with cells and centrifuge in eight sterile 50-ml centrifuge tubes 10 min at 400 × g, room temperature. 36. Remove the supernatant (by pipetting) as thoroughly as possible. Resuspend and combine the cells in 150 ml BioExpress unlabeled or labeled medium, depending upon the desired label. 37. Transfer five 30-ml aliquots of the resuspended cells into each of five 500-ml Erlenmeyer flasks containing 70 ml of BioExpress unlabeled or labeled medium. To express uniform 15 N-labeled protein use BioExpress-2000 (insect cell) U-15 -98% medium, for uniformly labeled 13 C/15 N protein, use BioExpress-2000 (insect cell) U15 N/U-13 C 92% to 95% labeled medium. BioExpress-2000 (insect cell) 13 C/15 N-labeled medium (for L-methionine, L-leucine, L-isoleucine, and L-tyrosine) can also be used to selectively label methionine, leucine, isoleucine, and tyrosine in the expressed protein.
38. Add 1.5 ml P2 recombinant baculovirus suspension with a titer value of ∼2 × 108 pfu/ml (see step 32) to each of the flasks. This results in a final MOI of 2.
39. Incubate the cells for 3 days at 27◦ C and 90 rpm. 40. Harvest the cells by centrifuging the cells at 400 × g at room temperature. Store the cell pellets up to 6 months at −80◦ C until used to purify the protein of interest.
ISOTOPIC LABELING OF PROTEINS PRODUCED IN CELL-FREE EXTRACTS
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Cell-free protein synthesis couples transcription and translation reactions in vitro, and is an effective means for producing proteins that are toxic to host cells or are quickly degraded by intracellular proteases (Nirenberg and Matthaei, 1961). Additionally, proteins that aggregate in inclusion bodies during in vivo production can sometimes be produced in a soluble form via cell-free synthesis. Finally, because little amino acid metabolism occurs in cell-free extracts, this method provides a means for incorporating specifically labeled amino acids into proteins with minimal scrambling of the isotope. Current Protocols in Protein Science
Basic Protocol 3 describes the cell-free reaction for producing isotopically enriched proteins. Support Protocols 1 and 2 describe the production of the S30 E. coli extract and T7 RNA polymerase necessary for cell-free production, respectively.
CELL-FREE PROTEIN EXPRESSION USING S30 EXTRACT AND T7 RNA POLYMERASE
BASIC PROTOCOL 3
Initial expression trials should be performed using a small-scale expression reaction. Once the protein expression and purification methodologies have been optimized, a large-scale expression system (using the optimized parameters from the small-scale reaction) with isotopically enriched amino acids can be used to make the isotopically labeled protein (see Torizawa et al., 2004).
Materials DEPC-treated H2 O: 500 µl DEPC per 100 ml H2 O 1.4 M CH3 COONH4 (ammonium acetate) 0.5 M (CH3 COO)2 Mg (magnesium acetate) Amino acid mixture (see recipe) 0.645 M creatine phosphate LM mixture (see recipe) 1 mg/ml high copy DNA template plasmid with gene for protein of interest and T7 promoter in DEPC-treated H2 O 11 mg/ml T7 RNA polymerase (Support Protocol 2 or Fermentas Life Sciences) RNase inhibitor (Fermentas Life Sciences) 10 mg/ml creatine kinase (Roche) S30 extract (Support Protocol 1) Small-scale dialysis solution (see recipe) Large-scale dialysis solution (see recipe) 5-mm or 15-mm diameter Spectra/Por CE 50,000 MWCO dialysis tubing (Spectrum Laboratories) 10-mm or 20-mm (i.d.) test tubes 37◦ C shaking water bath Additional reagents and equipment for performing SDS-PAGE (UNIT 10.1) and purifying proteins (Chapter 6) 1. Recommended: Rinse all labware with DEPC-treated water and autoclave just prior to use.
Set up a small-scale reaction 2a. Set up a small (500 µl) expression reaction by combining the following: 9.8 µl 1.4 M CH3 COONH4 (see annotation) 15 µl 0.5 M (CH3 COO)2 Mg 20 µl amino acid mixture 40 µl 0.645 M creatine phosphate 125 µl LM mixture 10 µl 1 mg/ml high copy DNA template plasmid 4.5 µl T7 RNA polymerase 1.25 µl RNase inhibitor 12.5 µl 10 mg/ml creatine kinase 150 µl S30 extract DEPC-treated H2 O (see recipe) to 500 µl. Use [15 N]- NH4 OOCCH3 if expressing 15 N labeled protein.
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3a. Place 0.5 ml of the reaction solution in the 5-mm dialysis tubing and place in a 10-mm test tube containing 2 ml of the small-scale dialysis solution. 4a. Place the test tube at an angle in a shaking water bath and incubate 7 hr at 37◦ C, with shaking at 150 rpm. 5a. Determine protein expression levels of a sample withdrawn from the dialysis chamber by separating the proteins using SDS-PAGE (UNIT 10.1) and comparing the brightness of the protein bands to the brightness of a standard marker.
Set up a large-scale reaction 2b. Set up a large (6 ml) expression reaction by combining the following: 117.6 µl 1.4 M CH3 COONH4 (see annotation below) 180 µl 0.5 M (CH3 COO)2 Mg 240 µl amino acid mixture 480 µl 0.645 M creatine phosphate 1500 µl LM mixture 120 µl 1 mg/ml high copy DNA template plasmid 54 µl T7 RNA polymerase 15 µl RNase inhibitor 150 µl 10 mg/ml creatine kinase 1800 µl S30 extract DEPC-treated H2 O to a final volume of 6 ml. Use [15 N]-CH3 COONH4 if expressing 15 N labeled protein.
3b. Place 6 ml reaction solution in the 15-mm dialysis tubing and place in a 20-mm test tube containing 24 ml large-scale dialysis solution. 4b. Incubate the test tube at an angle in a shaking water bath 7 hr at 37◦ C, with shaking at 150 rpm. The amino acid mixture is substituted by (a) 15 N-labeled algal hydrolysate (4 mg/ml) to express [U-15 N] labeled protein; (b) 2 H, 13 C, 15 N-labeled algal hydrolysate (4 mg/ml) to express [U-2 H, 13 C,15 N] labeled protein; and (c) 13 C, 15 N-labeled amino acids (1.7 mg/ml) to express[U-13 C, 15 N]-labeled protein.
5b. Determine protein expression levels by SDS-PAGE (UNIT 10.1) of a sample withdrawn from the dialysis tubing. 6b. Combine both the reaction and dialysis solutions, concentrate, and purify the protein of interest using appropriate protocols (see Chapter 6). SUPPORT PROTOCOL 1
Incorporation of Isotopically Enriched Amino Acids
PREPARATION OF S30 EXTRACT FROM E. coli S30 extract refers to the supernatant from the centrifugation of the E. coli cell lysate at 30,000 × g that contains the required cellular components for translation of mRNA. This protocol describes the growth of 10 liters of E. coli cells, followed by extraction and purification to yield an extract capable of high-yield protein synthesis with little endogenous, natural abundance amino acids. A purified extract maximizes the incorporation of isotopically enriched amino acids during cell-free expression. This protocol takes 2 to 3 days and yields enough extract for ∼15 large scale reactions or >100 small scale cell-free reactions (Torizawa et al., 2004).
Materials E. coli cells lacking RNase activity, e.g., BL21 star (DE3) or A19 (met B, RNA), Invitrogen LB agar plate containing the appropriate antibiotic (APPENDIX 4A)
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LB medium (APPENDIX 4A) S30 buffer (see recipe), chilled to 4◦ C 1 M dithiothreitol (DTT) or solid DTT Activation solution (see recipe) Polyethylene glycol 8000 (PEG 8000; Sigma-Aldrich) 500-ml flasks 4-liter baffled culture flasks, sterile 37◦ C orbital shaker incubator 250- and 500-ml centrifuge tubes High-speed refrigerated centrifuge with appropriate rotors (e.g., Beckman J2-HS, with JA 10 and JA 20 rotors) French Press 20.4 mm Spectra/Por dialysis tubing, 6-8,000 MWCO (Spectrum Laboratories) 2 Hi-Prep 26/10 columns (GE Healthcare) 1. Recommended: Rinse all labware with DEPC-treated water (see recipe) and autoclave just prior to use.
Grow and lyse bacterial cells 2. Grow an E. coli strain lacking RNase activity on an LB plate containing the appropriate antibiotic (depending on the cell line) overnight at 37◦ C. 3. Pick a single colony and inoculate a 500-ml flask containing 100 ml of LB medium equilibrated to 37◦ C. Incubate the flask overnight at 37◦ C, with shaking at 250 to 300 rpm. 4. The following morning, determine the OD600 against water as a blank to verify that the culture has reached stationary phase with an OD600 >1.5. Dilute the sample 10-fold to ensure a reading between 0.1 and 1.0. 5. Add 100 ml of the overnight culture to 10 liters LB medium equilibrated to 37◦ C, dispense among ten 4-liter flasks, and incubate 2 to 4 hr at 37◦ C, with shaking, until the cultures reach an OD600 of 0.7.
Harvest and lyse cells 6. Harvest the cells by centrifuging the culture in 500-ml centrifuge tubes 20 min at 10,000 × g, 4◦ C. 7. Decant the supernatant and resuspend all the cell pellets in a total volume of 200 ml chilled S30 buffer. 8. Centrifuge the suspension in 250-ml centrifuge tubes 20 min at 10,000 × g, 4◦ C. 9. Decant the supernatant and resuspend the cell pellet in 200 ml of fresh, chilled S30 buffer. 10. Repeat steps 8 and 9 two more times for a total of four washes in S30 buffer. The cell pellet can be stored at −80◦ C overnight after the third wash and the fourth wash can be done the next morning before proceeding to step 11.
11. Suspend the cells in 23 ml of chilled S30 buffer. Lyse the cells using a French press with a pressure of 20,000 psi.
Purify S30 extract 12. Add 23 µl of 1 M DTT solution (or 3.55 mg of DTT) to the cell lysate to achieve a final concentration of 1 mM. Centrifuge the lysate 30 min at 30,000 × g, 4◦ C. 13. Collect the top 24 ml of the supernatant and centrifuge 30 min at 30,000 × g,
4◦ C.
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14. Carefully collect the top 18 ml of the supernatant and mix with 5.4 ml of activation solution. Incubate the mixture 80 min at 37◦ C, with shaking at 80 rpm. 15. Using 6,000-8,000 molecular weight cutoff (MWCO) dialysis tubing, dialyze the 23.4 ml of extract against 2 liters of prechilled S30 buffer 45 min at 4◦ C. 16. Change the buffer and dialyze 45 min at 4◦ C two more times for a total of three dialyses. 17. Equilibrate two HiPrep 26/10 columns for gel filtration with 3 to 5 column volumes of S30 buffer (200 ml/column). Load the extract onto the columns and collect the first 35 ml of the eluant. The columns can be run either in parallel or in series.
18. Concentrate the eluant by placing it in 6,000 to 8,000 MWCO dialysis tubing and dialyzing against a mixture of 367 ml S30 buffer and 333 ml PEG-8000 1 hr at 4◦ C. 19. Dialyze the concentrated extract in 2 liters of S30 buffer 1 hr at 4◦ C. 20. Dispense the extract into 2-ml aliquots in microcentrifuge tubes and store up to 6 months at −80◦ C. SUPPORT PROTOCOL 2
PREPARATION OF T7 RNA POLYMERASE In vitro translation systems use the efficient bacteriophage T7 RNA polymerase to form mRNA by transcription for subsequent translation. This protocol describes the overexpression of recombinant T7 RNA polymerase from a 1-liter culture of E. coli transformed with pAR1219 and induced with IPTG. The resulting protein is purified via ion exchange chromatography, dialyzed into buffer containing 50% glycerol, and stored until needed for cell-free expression. For an overview and details about recombinant protein expression, including expression strategies, secretion of proteins, fusion tags and proteins, refer to Chapter 5.
Materials Lysogenic expression host (E. coli, such as BL21 (DE3) transformed with pAR1219 (Davanloo et al., 1984) LB agar plate containing the appropriate antibiotic (APPENDIX 4A) LB medium (APPENDIX 4A) 500× ampicillin (see recipe for antibiotic solutions) 1 M Isopropyl-1-thio-β-D-galactopyranoside (IPTG; Research Products International) Lysis buffer for T7 RNA polymerase (see recipe), chilled to 4◦ C Sodium deoxycholate (Sigma-Aldrich) Tris buffer for T7 RNA polymerase, pH 8.1 (see recipe), chilled to 4◦ C 2 M ammonium sulfate: prepare fresh and sterilize by passing through a 0.22-µm filter 10% (w/v) polyethyleneimine (Sigma): prepare fresh and sterilize by passing through a 0.22-µm filter Ammonium sulfate, saturated (see recipe) Dialysis buffer (see recipe) 5 M NaCl 0.2 M PMSF (Sigma-Aldrich Co.) in isopropanol Glycerol Incorporation of Isotopically Enriched Amino Acids
250-ml and 4-liter baffled culture flasks, sterile 37◦ C orbital shaker incubator 500-ml centrifuge tubes
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High speed refrigerated centrifuge with appropriate rotors (Beckman J2-HS, with JA 10 and JA-20 rotors) Sonicator with 3/6-mm Vibracell probe tip (Sonics & Materials) 20.4-mm dialysis tubing, 6000 to 8000 MWCO 20-ml HiTrap SP FF (GE Healthcare) or Macro-prep high S (Bio-Rad) column Additional reagents and equipment for performing SDS-PAGE (UNIT 10.1) and spectrophotometrically determining protein concentration (UNIT 3.1) Grow bacterial cells 1. Grow the expression host bacterial strain transformed with pAR1219 on an LB plate containing the appropriate antibiotic overnight at 37◦ C. 2. Pick a single colony and inoculate a 250-ml flask containing 50 ml of LB medium and 0.05 to 0.1 ml of the appropriate antibiotic solution, e.g., ampicillin if E. coli strain BL21(DE3) is used. Incubate the flask overnight at 37◦ C, with shaking at 250 to 300 rpm. 3. The following morning, determine the OD600 against water as a blank to verify that the culture has reached stationary phase with an OD600 >1.5. Dilute the sample 10-fold to ensure a reading between 0.1 and 1. 4. Use 5 ml of the overnight culture to inoculate each of two 4-liter baffled flasks each containing 500 ml of LB medium and 1 ml of 500× ampicillin equilibrated to 37◦ C. 5. Incubate the flasks 2 to 4 hr at 37◦ C, with shaking at 250 to 300 rpm until the bacterial culture reaches an OD600 of 0.5. 6. Remove an initial 1-ml sample from each flask and save for SDS-PAGE analysis (UNIT 10.1) as a control for uninduced cells and either immediately treat as described in step 11 or freeze the sample until ready to perform step 11.
Express recombinant T7 RNA polymerase 7. Add 0.5 ml of 1 M IPTG to each flask for a final concentration of 0.5 mM IPTG to induce protein expression. 8. Incubate the flask at 37◦ C in an incubator shaker at 250 to 300 rpm for 8 hr after induction. Remove 1 ml from each culture and save for SDS-PAGE analysis (UNIT 10.1). Check final OD600 . 9. Harvest the cells by centrifuging the culture in 500-ml centrifuge tubes 20 min at 10,000 × g, 4◦ C. 10. Decant the supernatant and resuspend the cell pellets in a total volume of 25 ml chilled lysis buffer and stir for 15 min. 11. Analyze the 1-ml samples taken before and after induction using SDS-PAGE (UNIT 10.1) to check for protein expression.
Fractionate cell lysate by ammonium sulfate precipitation 12. Add 200 mg of sodium deoxycholate to the lysed solution from step 10 for a final concentration of 0.8% (w/v). Stir the solution for 15 min. 13. Sonicate the suspension on ice with a probe-tip sonicator using 10 short bursts (10 sec) with 1-min pauses between bursts. 14. Add 15 ml of chilled 2 M ammonium sulfate to the lysate. Bring the total volume to 150 ml with chilled Tris buffer. 15. Slowly add 15 ml of chilled 10% (w/v) polyethyleneimine (adjusted to pH 7.0 with HCl) and stir 20 min at 4◦ C.
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16. Centrifuge 15 min at 39,000 × g, 4◦ C and collect the supernatant. 17. Add 135 ml of saturated ammonium sulfate to the supernatant and stir for 15 min at 4◦ C. 18. Centrifuge for 15 min at 12,000 × g, 4◦ C. Decant the supernatant, resuspend the pellet in 45 ml of chilled dialysis buffer and add 900 µl of 5 M NaCl and 22 µl of 0.2 M PMSF.
Purify recombinant T7 RNA polymerase 19. Transfer the suspension to 6,000-8,000 MWCO dialysis tubing. Dialyze the suspension in 2 liters of dialysis buffer to which 40 ml of 5 M NaCl and 1 ml of 0.2 M PMSF has been added, 3 hr at 4◦ C. 20. Place sample into fresh buffer, keeping the salt and PMSF concentrations constant and dialyze an additional 3 hr at 4◦ C. Repeat the dialysis a third time. 21. Remove the suspension from the dialysis tubing and add 50 ml chilled dialysis buffer and 50 µl of 0.2 M PMSF (no additional NaCl is added). 22. Equilibrate a HiTrap SP FF or Macro-prep high S column with 100 ml of chilled dialysis buffer to which 1 ml of 5 M NaCl and 100 µl of 0.2 M PMSF has been added. 23. Add the protein suspension to the column and wash with 100 ml of chilled dialysis buffer to which 1 ml of 5 M NaCl has been added. 24. Elute the T7 RNA polymerase with 100 ml of chilled dialysis buffer to which 4 ml of 5 M NaCl has been added. Collect the fractions exhibiting absorbance at 280 nm. 25. Place the fractions in 6,000 to 8,000 MWCO dialysis tubing and dialyze twice against 2 liters of dialysis buffer to which 40 ml of 5 M NaCl has been added, for 6 hr at 4◦ C each time. 26. Dialyze a third time against 1 liter dialysis buffer to which 20 ml of 5 M NaCl and 1 liter of glycerol have been added, 6 hr at 4◦ C. 27. Remove the protein solution from the dialysis tubing and centrifuge for 20 min at 39,000 × g, 4◦ C. Save the supernatant. 28. Determine the protein concentration (UNIT 3.1). Concentrate the protein if necessary to ∼10 mg/ml and store up to 6 months at −20◦ C and longer at −80◦ C. An absorbance of 0.7 at OD280 corresponds to 1 mg/ml. An Amicon Centricon (Millipore) may be used if protein concentration is required. BASIC PROTOCOL 4
Incorporation of Isotopically Enriched Amino Acids
INCORPORATION OF AN ISOTOPICALLY ENRICHED AMINO ACID IN A PEPTIDE VIA SOLID PHASE PEPTIDE SYNTHESIS UTILIZING 9-FLUORENYLMETHYLOXYCARBONYL (Fmoc) AMINO ACIDS Solid phase peptide synthesis (SPPS) is based on the sequential addition of α-amino and side-chain protected amino acid residues to an insoluble polymeric support. The growing peptide chain is subjected to treatment with mild base to remove the baselabile 9-fluorenylmethyloxycarbonyl (Fmoc) group, which is used for α-amino protection followed by addition of the next amino acid. Final cleavage of the completed peptide from the resin and removal of the side-chain protecting groups is done with trifluoroacetic acid (TFA). This protocol describes the solid phase synthesis of a peptide using the appropriate isotopically labeled Fmoc-amino acids of interest. See UNIT 18.2 for additional information and applications.
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Materials Solid phase resin with carboxy terminal amino acid attached (EMD Biosciences) Dichloromethane (DCM; Fisher Scientific) N-methylpyrrolidone (NMP; Sigma-Aldrich) 25% (v/v) piperidine (Sigma-Aldrich) in NMP Isotopically labeled and unlabeled Fmoc-protected amino acids (Support Protocol 3 or Cambridge Isotope Laboratories) N-hydroxybenzotriazole (HOBt; EMD Biosciences) Benzotriazole-1-yl-oxy-tris-pyrrolidino-phosphonium hexafluorophosphate (PyBOP; EMD Biosciences) Diisopropylethylamine (DIPEA; Aldrich) Acetic anhydride solution (see recipe) King’s reagent (see recipe) or TFA/TIS/H2 O solution: 9 ml trifluoroacetic acid (TFA; Acros)/2.5 ml triisopropylsilane (TIS; Sigma-Aldrich)/2.5 ml H2 O solution Diethyl ether, chilled Nitrogen gas (optional) peptide reaction vessel (Sigma) 100-ml round-bottom flask Rotary evaporator equipped with a base trap filled with NaOH pellets Additional reagents and equipment for performing the ninhydrin test (Support Protocol 4) Swell and wash resin 1. Place the solid phase resin (0.25 mM, 1 equivalent) in a peptide reaction vessel. The resin is supplied as a solid, quantified in mmol of active group/gram of resin support. Use this number to calculate the grams of resin needed for 0.25 mmol of active sites.
2. Add 15 ml of dichloromethane (DCM) to cover the solid phase resin. 3. Gently agitate the solid phase resin mixture for 15 min. 4. Remove the DCM by application of a vacuum. 5. Wash the resin with 15 ml N-methylpyrrolidone (NMP) four times.
Deprotect resin 6. Add 20 ml of 25% (v/v) piperidine in NMP to cover the resin. 7. Gently agitate the solid phase resin mixture for 30 min. 8. Drain the reagents. 9. Wash the resin with 15 ml of NMP four times.
Couple Fmoc-amino acid with solid phase peptidyl resin 10. Weigh out 4 equivalents (Eq) of the isotopically labeled Fmoc-amino acid (see Support Protocol 3), 4 Eq (153 mg) of HOBt and 4 Eq (520 mg) of PyBOP. 11. Add 15 ml of NMP to effect complete dissolution of the reagents. 12. Add 8 Eq (348 µl, 258 mg) of DIPEA to the solution and mix thoroughly. 13. Add the Fmoc-amino acid/HOBt/PyBOP/DIPEA solution to the N-deprotected solid phase resin. Protein Engineering
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14. Gently agitate the resin in this solution for 2 hr. 15. Drain the reagents. 16. Wash the resin with 15 ml of NMP four times. 17. Transfer a few beads of resin to a small test tube and perform the Ninhydrin test (see Support Protocol 4) to determine the amount of free amino groups. If the resin gives a positive color test, repeat the coupling reaction with fresh reagents. Serine, asparagine, and aspartic acid do NOT produce the expected dark blue/purple color typical of free primary amino groups. Therefore, a double coupling step should be employed for the addition of these amino acid residues.
18. Wash the resin with 15 ml of NMP four times.
Cap unreacted terminal amino groups 19. Add 15 ml acetic anhydride solution to the peptidyl resin and agitate gently for 20 min. Use the standard Ninhydrin test (Support Protocol 4) to monitor the reaction. DO NOT cap if hydroxyl groups on serines or threonines are unprotected; skip steps 20 to 22 and proceed to repeat step 6 if adding more residues or step 23 if peptide is complete.
20. Remove the reaction solution by filtration. 21. Wash the product with 15 ml of (1:1, v/v) NMP/DCM four times. The solid phase peptidyl resin is now ready for Fmoc deprotection. Repeat steps 6 through 21 for each additional amino acid residue to be added.
Cleave completed peptide and remove side chain protecting groups 22. In a 100-ml round bottom flask combine resin and 10 ml/g resin of King’s reagent or TFA/ TIS/H2 O solution and stir for 1.5 to 2 hr. CAUTION: TFA is a very strong acid with a low boiling point. Wear eye protection, use gloves, and handle only in a fume hood. The choice of cleavage solution is peptide sequence dependent.
23. Using a 50-ml sintered-glass funnel, filter the TFA/TIS/H2 O solution and rinse the round-bottom flask and funnel with additional TFA/TIS/H2 O solution. Collect and combine all the TFA-containing fractions. 24. Concentrate the TFA solution to 1 to 2 ml by rotary evaporation, using a base trap. The base trap is a glass vessel containing NaOH pellets to neutralize any acid before it reaches the vacuum pump.
25. Using a disposable glass pipet, drip the residual TFA solution slowly into 50 ml of chilled diethyl ether to precipitate the peptide. 26. Centrifuge diethyl ether solution 5 min at 4000 × g, 4◦ C to collect the precipitated peptide. CAUTION: Avoid open flames near the centrifuge.
27. Decant the diethyl ether layer, add 45 ml of diethyl ether, and repeat centrifugation. Repeat this step three more times for a total of four washes. Incorporation of Isotopically Enriched Amino Acids
28. Dry the crude peptide under a low nitrogen stream or vacuum and store up to 6 months at 4◦ C until ready for chromatography. Keep the peptide dry.
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PREPARATION OF A 9-FLUORENYLMETHYLOXYCARBONYL (Fmoc)-PROTECTED AMINO ACID
SUPPORT PROTOCOL 3
Although the isotopically enriched analogs of most amino acids with the 9fluorenylmethoxycarbonyl (Fmoc) protecting group are commercially available, it is cost effective and in some cases necessary to obtain the isotopically labeled amino acid and perform the Fmoc-protection oneself. This protocol describes the protection of appropriately isotopically labeled amino acids with the 9-fluorenylmethoxycarbonyl (Fmoc) protecting group. This protocol is only appropriate for amino acids which do not have reactive side chains (Gly, Ala, Val, Ile, Phe, Pro, Trp, Leu, and Met) or side chains with limited reactivity (Ser, Thr, Tyr).
Materials Isotopically labeled amino acid (Cambridge Isotope Laboratories) 10% (w/v) Na2 CO3 solution, warm Fmoc-O-succinimide (EMD Biosciences) 1,4-dioxane Toluene/acetic acid (10:1 ratio) Diethyl ether HCl, concentrated and 1 N Ethyl acetate MgSO4 , anhydrous 50-ml round-bottom flask Aluminum-backed thin-layer chromatography (TLC) plates (Whatman) TLC tank UV light source 250-ml separatory funnel Rotary evaporator 1. Dissolve 2.25 mmol of the isotopically labeled amino acid in 10 ml of warm (50◦ C) 10% (w/v) Na2 CO3 . Gentle heating of the solution will assist in dissolution.
2. Dissolve 850 mg (2.5 mmol) of Fmoc-O-succinimide in a minimal volume (∼5 ml) of dioxane in a 50-ml round bottom flask. CAUTION: Dioxane is toxic, and the liquid and vapor are flammable. Use gloves, eye protection, and good ventilation.
3. Add the Na2 CO3 solution dropwise to the 50-ml flask containing the dioxane while stirring, and continue stirring 2 to 24 hr at room temperature. Monitor the course of the reaction by thin-layer chromatography, using 10:1 toluene:acetic acid as the eluting solvent system. a. Using a capillary pipet, spot ∼1 µl of the reaction mixture near the bottom of a TLC plate and allow the spot to dry. As controls, spot 1 µl of each of the starting amino acids and the Fmoc-O-succinimide. b. Gently (without splashing) place the plate into a TLC tank containing 10:1 toluene:acetic acid so the spot is near the surface of solution but not submersed in it. c. Cover the tank and allow the solvent front to migrate at least 75% of the height of the plate. d. Remove plate and allow it to air-dry. Protein Engineering
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e. View reaction products using a handheld UV light to look for the disappearance of the spot corresponding to the starting amino acid in the reaction mix. 4. If the reaction is not complete at 24 hr, add an additional 300 mg (0.88 mmol) of Fmoc-O-succinimide dissolved in ∼2 ml of dioxane and stir for another 24 hr. 5. Dilute the reaction mixture by adding ∼30 ml distilled water. 6. Transfer the reaction mixture to a 250-ml separatory funnel and wash three times with 20 ml of diethyl ether each time. Use the aqueous layer to proceed with purification of the product, but save the ether layers until successfully completing the protocol. 7. Add sufficient concentrated HCl, with stirring and cooling, to the aqueous layer to acidify it to pH 2. 8. Extract the aqueous layer three times with ethyl acetate, 50 ml each time. Use the ethyl acetate layer to proceed with purification of the product, but save the aqueous layers until successfully completing the protocol. 9. Use the separatory funnel to wash the combined ethyl acetate extracts twice with 100 ml of 1 N HCl, then once with 100 ml of distilled water. 10. Add anhydrous MgSO4 to the ethyl acetate solution and allow to stand for at least 30 min to remove residual water. 11. Filter the ethyl acetate solution and remove ethyl acetate via rotary evaporation. Crystals of the Fmoc-protected amino acid should form and there should be no ethyl acetate smell. Purity can be assessed by TLC as in step 4, using commercially available Fmoc-amino acids as controls. SUPPORT PROTOCOL 4
NINHYDRIN TEST TO MONITOR AMINO ACID COUPLING REACTION The ninhydrin test is based upon reaction of ninhydrin with free primary amines to form a violet colored compound, known as Ruhemann’s purple. The formation of Ruhemann’s purple indicates the free amine in the remaining resin sample taken after the coupling reaction (i.e., the unreacted amino group on the growing, resin-attached peptide).
Materials Sample (e.g., amino acid coupling reaction from Basic Protocol 4) Solution 1: 5% (w/v) ninhydrin (Fisher) in ethanol Solution 2: 1 mM KCN in pyridine (see recipe) Solution 3: 80% (w/v) phenol in ethanol 15-ml glass test tube 100◦ C hotplate 1. Transfer a few resin beads to a 15-ml glass test tube and add 2 drops each of solutions 1, 2, and 3. CAUTION: Potassium cyanide is extremely toxic. Follow all precautions recommended in the material safety data sheet (MSDS).
2. Mix well and heat to 100◦ C for 3 min. The appearance of a dark blue/purple color indicates the presence of free amine. Incorporation of Isotopically Enriched Amino Acids
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ISOTOPIC LABELING OF AMINO ACIDS IN A PROTEIN VIA CHEMICAL EXCHANGE USING DEUTERATED WATER
BASIC PROTOCOL 5
Chemical exchange of the protons in a protein is accomplished by dissolving the protein in deuterated water for short periods of time at an acidic pH. The exchangeable NH and OH protons are replaced by deuterons from the solvent. The conditions described below are optimized for a labeling time of ∼30 min for an unfolded protein. The kinetics of the proton/deuteron exchange can be altered by changing the pH of the solution (Rohl et al., 1992; Englander and Poulsen, 1969). For stable, folded proteins, the exchange of interior amide protons may take from hours up to days. For proteins which fold reversibly, the addition of guanidinium chloride will increase the rate of proton/deuteron exchange (Kotik et al., 1995). The protein can then be refolded via dialysis in a deuterated buffer solution. This protocol describes deuteration of amino acids in a polypeptide via solvent exchange.
Materials Protein or peptide of interest 1 M hydrochloric acid Deuterated water (Cambridge Isotope Laboratories) containing 150 mM NaCl and 5 mM sodium phosphate, 5◦ C 100-ml round-bottom flask Magnetic stir bar and stirrer pH meter Lyophilizer 1. Dissolve the protein in distilled water to a concentration of 1 to 2 mM in a roundbottom flask equipped with a magnetic stir bar. 2. Adjust the pH of the protein solution to 3.0 by adding 1 M HCl dropwise while stirring. 3. Lyophilize the protein sample to dryness and store at −20◦ C. 4. Dissolve the protein sample to a concentration of 1 to 2 mM in cold deuterated water (2 H2 O) containing 150 mM NaCl and 5 mM sodium phosphate and allow to stand 30 min at 5◦ C.
ISOTOPIC LABELING OF AMINO ACIDS IN A PROTEIN VIA CHEMICAL EXCHANGE USING DEUTERATED GUANIDINIUM CHLORIDE
ALTERNATE PROTOCOL 7
Treatment of a protein with deuterated guanidinium hydrochloride causes the denaturation of the protein and allows chemical exchange of all interchangeable NH and OH protons, including those that were inaccessible in the native conformation of the protein. Refolding of the protein to the native state can be achieved by dialysis of the protein in deuterated water. Following lyophilization, incubation of the native peptide in distilled water affords protein in which only the interior OH and NH protons are substituted with deuterons. The pH of the final buffer should be chosen based on the exchange rates needed for the NMR experiment.
Materials Protein or peptide of interest 1 M HCl 6 M deuterated guanidinium chloride (Cambridge Isotope Laboratories) in deuterated water (Cambridge Isotope Laboratories) 150 mM NaCl/5 mM sodium phosphate solution Deuterated water (Cambridge Isotope Laboratories)
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10-ml round-bottom flask Magnetic stir bar and stirrer pH meter Lyophilizer Dialysis tubing (protein-dependent MWCO) 1. Dissolve the protein sample in distilled water to a concentration of 1 to 2 mM in a round-bottom flask equipped with a magnetic stir bar. 2. Adjust the pH of the protein solution to 3.0 by adding 1 M HCl dropwise while stirring. 3. Lyophilize the peptide sample to dryness and store at −20◦ C. 4. Dissolve the protein sample to a concentration of 1 to 2 mM in 6 M deuterated guanidinium chloride in deuterated water and allow to stand for 30 min at room temperature. 5. Dialyze the peptide solution into deuterated water containing 150 mM NaCl and 5 mM sodium phosphate for 24 hr at 4◦ C. The dialysate reservoir should be replaced at least once during this time period. The sample may now be used fully deuterated or the exchangeable deuterons may be replaced by proceeding with steps 6 & 7.
6. Lyophilize the peptide sample to dryness. 7. Replace the lost water with an equal volume of distilled water containing 150 mM NaCl and 5 mM sodium phosphate and allow to stand for 30 min at room temperature. BASIC PROTOCOL 6
SELECTIVE ISOTOPIC LABELING OF PROTEINS BY CHEMICAL MODIFICATION: REDUCTIVE ALKYLATION OF PRIMARY AMINES This protocol describes the selective isotope labeling of proteins by covalently modifying the lysine residues and N-terminus with C2 H3 or 13 CH3 as a means of introducing isotopic labels into the protein with minimal disruption to protein function and structure. This reaction, in which 2 H- or 13 C-labeled formaldehyde is reacted with the primary amine and subsequently reduced to the tertiary amine by sodium cyanoborohydride (NaCNBH3 ) is very specific because the reductive methylation occurs only at the NH2 terminus and the side chain groups of lysine residues (Jentoft and Dearborn, 1983; Spooner and Watts, 1991).
Materials 2 mg/ml solution of protein of interest in 50 mM phosphate or HEPES buffer, pH 7.0 0.1 M sodium cyanoborodeuteride solution (NaCNB2 H3 ; Cambridge Isotope Laboratories) or 0.1 M sodium cyanoborohydride solution (NaCNBH3 ; Aldrich) 0.1 M 2 H-labeled formaldehyde solution (see recipe) or 0.1 M 13 C-labeled formaldehyde solution (see recipe) 0.1 M NaCl 5-ml test tube Dialysis tubing with MWCO appropriate for the size of the protein of interest Incorporation of Isotopically Enriched Amino Acids
1. Add 1 ml of ∼2 mg/ml protein in 50 mM phosphate or HEPES buffer, pH 7.0, to a 5-ml test tube.
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2a. For 2 H-labeling: Add 0.1 ml of 0.1 M sodium cyanoborodeuteride to the test tube and mix well. 2b. For 13 C-labeling: Add 0.1 ml of 0.1 M sodium cyanoborohydride to the test tube and mix well. CAUTION: Sodium cyanoborohydride and sodium cyanoborodeuteride are highly toxic. Wear protective eyewear, gloves, and clothing, and work only in a fume hood. The NaCNBH3 (or NaCNB2 H3 ) solution should be prepared on the day of use. The amount of NaCNBH3 (or NaCNB2 H3 ) added should be adjusted if the protein to be labeled has an abnormally high or low lysine content, to give an approximately 10-fold molar excess over the free amino groups.
3a. For 2 H-labeling: Add 0.1 ml of 0.1 M 2 H-labeled formaldehyde to the test tube and mix well. 3b. For 13 C-labeling: Add 0.1 ml of 13 C-labeled formaldehyde to the test tube and mix well. 4. Incubate the mixture 2 hr at room temperature or overnight at 4◦ C. 5. Dialyze (see APPENDIX 3B) the reaction mixture extensively against several changes of 0.1 M NaCl or any appropriate buffer.
SELECTIVE ISOTOPIC LABELING OF PROTEIN SULFHYDRYL GROUPS BY COVALENT CHEMICAL MODIFICATION
BASIC PROTOCOL 7
This protocol describes the introduction of isotopic labels into a protein via covalent modification of cysteine sulfhydryl groups with a chemical reagent. One example of such a reagent is 3-bromo-1,1,1-trifluoroacetone (BTFA), which is effective for introducing an 19 F labeled moiety onto cysteine residues (Brauer and Sykes, 1986). Iodoacetamide can be used to add an 15 N-enriched or 13 C-enriched moiety.
Materials Protein sample Denaturing or nondenaturing reducing buffer (see recipes) 3-Bromo-1,1,1-trifluoroacetone (BTFA) or iodoacetamide with either 13 C or 15 N enrichment (Cambridge Isotope Laboratories) Dialysis tubing (if denaturing) with MWCO appropriate for the size of the protein of interest Dialysis buffer (see recipe) 250-ml round-bottom flask Magnetic stir bar and stirrer 1. Dissolve protein at a concentration of 50 µM in denaturing or nondenaturing reducing buffer to a final volume of 1 ml. Stir at 37◦ C for 1 hr. 2. Add either 0.19 mg of BTFA (if introducing 19 F) or 0.29 mg of iodoacetamide (with either 13 C or 15 N enrichment) and stir for an additional hour. The level of isotopic enrichment is dependent on the accessibility of the cysteine residues in the protein. It will be higher in a denaturing buffer, but refolding of the protein can be problematic.
3. If using denaturing buffer, dialyze (see appropriate buffer.
APPENDIX 3B)
the reaction solution into an
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REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX. Store sterile media components at room temperature, unless otherwise indicated, for periods of time appropriate to their chemical and microbiological stability.
Acetic anhydride solution 190 µl acetic anhydride (Sigma-Aldrich) 175 µl diisoproylethylamine (DIPEA; Sigma-Aldrich) N-methylpyrrolidone (NMP) to a final volume of 15 ml Prepare fresh just prior to use Activation solution Combine using sterilized stock solutions just prior to use. 286 mg Tris-acetate 100 µl 0.5 M Mg(CH3 COO)2 (magnesium acetate) 24 µl 1 M DTT 1.08 ml 1 M Tris buffer, pH 8.0 (APPENDIX 2E) 8.6 µl 2.5 mM amino acid mixture (see recipe) 93 mg phospho(enol)pyruvic acid monopotassium salt (Sigma) 3.75 ml 100 mM ATP 36 units pyruvate kinase (USB) H2 O to a final volume of 5.4 ml Amino acid mixture (2.5 mM) 223 mg alanine 435 mg arginine 330 mg asparagine 333 mg aspartic acid 303 mg cysteine 365 mg glutamine 368 mg glutamic acid 188 mg glycine 388 mg histidine 328 mg isoleucine 328 mg leucine 365 mg lysine 373 mg methionine 413 mg phenylalanine 288 mg proline 263 mg serine 298 mg threonine 508 mg tryptophan 453 mg tyrosine 293 mg valine H2 O to a final volume of 100 ml Sterilize by passing through a 0.22-µm filter, divide into 1-ml aliquots, and store up to 6 months at −20◦ C. Incorporation of Isotopically Enriched Amino Acids
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Amino acids solution, 50× (lacking cysteine, tyrosine and methionine) 1 g each of: monosodium glutamate asparagine monohydrate arginine monohydrochloride lysine monohydrochloride histidine monohydrochloride monohydrate 1 g each of free acids of: aspartic acid alanine praline glycine threonine serine glutamate valine leucine isoleucine phenylalanine tryptophan H2 O to a final volume of 100 ml If necessary, adjust the pH to ∼7 with 1 M NaOH. Sterilize by passing through a 0.22-µm filter, divide into 10-ml aliquots, and store up to 6 months at −20◦ C.
Amino acid stock solution, 50× 1 g of selectively isotopically enriched amino acid (Cambridge Isotope Laboratories) H2 O to a final volume of 100 ml If necessary, adjust the pH to ∼7 with 1 M NaOH. Sterilize by passing through a 0.22-µm filter, divide into 10-ml aliquots, and store up to 6 months at −20◦ C. The isotopically enriched amino acid is generally >99% labeled. It may be the salt of the amino acid.
Ammonium sulfate, saturated 62 g ammonium sulfate H2 O to a final volume of 150 ml Adjust pH to 7.0 with 1 M Tris buffer, pH 7.0 (APPENDIX 2E) Store up to 6 months at room temperature Antibiotic solutions, 500× 500× ampicillin solution Dissolve 0.25 g of ampicillin (Research Products International) in 10 ml water. 500× chloramphenicol solution Dissolve 0.125 g of chloramphenicol (Research Products International) in 10 ml deionized water. 500× kanamycin solution Dissolve 0.5 g of kanamycin (Research Products International) in 10 ml deionized water. Sterilize by passing through a 0.22-µm filter, divide into 5-ml and 500-µl aliquots, and store up to 6 months at −20◦ C.
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Auto-induction carbon source, 50× 37.5 g glycerol 2.5 g D-glucose 0.5 g α-lactose monohydrate H2 O to a final volume of 100 ml Sterilize by passing through a 0.22-µm filter and store up to 6 months at room temperature Use 99% [13 C3 ]-glycerol if full 13 C labeling of the protein is intended.
Auto-induction minimal medium Combine the following sterilized stock solutions just prior to use: 50 ml 20× salts solution (cooled to <50◦ C; see recipe) 20 ml 50× auto-induction carbon source (see recipe) 2 ml 1 M MgSO4 0.2 ml 1000× metals (see recipe) 2 ml 500× antibiotic solution (plasmid-dependent; see recipe) H2 O to a final volume of 1 liter To prepare 13 C labeled minimal medium: use 99% [13 C3 ]-glycerol (Cambridge Isotopes Laboratories) in the auto-induction carbon source recipe. To prepare 15 N labeled minimal medium: substitute 99% 15 NH4 Cl (Cambridge Isotopes Laboratories) for unlabeled NH4 Cl, as noted in the salts solution recipe. To prepare 15 N, 13 C labeled minimal medium: substitute 99% 15 NH4 Cl (Cambridge Isotopes Laboratories) for unlabeled NH4 Cl and use 99% [13 C3 ]-glycerol (Cambridge Isotopes Laboratories) is as the sole carbon source. The labeled sources of 13 C and 15 N can be used individually or in combination and are added depending on the desired nature of the labeling.
Auxotroph amino acids, 50× If incorporating fluorotryptophan 1 g each of tyrosine and phenylalanine H2 O to a final volume of 100 ml If incorporating fluorophenylalanine 1 g each of tyrosine and tryptophan H2 O to a final volume of 100 ml If incorporating fluorotyrosine 1 g each of tryptophan and phenylalanine H2 O to 100 ml If necessary, adjust the pH to ∼7 with 1 M NaOH. Sterilize by passing through a 0.22-µm filter, divide into 10-ml aliquots, and store up to 6 months at −20◦ C. 13
C-labeled formaldehyde solution, 0.1 M 32 mg 2 H- labeled paraformaldehyde (Cambridge Isotope Laboratories) Bring to a final volume of 10 ml with water in a test tube. Seal the test tube and hydrolyze overnight at 100◦ to 110◦ C. Prepare just prior to use.
Incorporation of Isotopically Enriched Amino Acids
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Carbon source conditioning medium Combine, using sterilized stock solutions, just prior to use.
50 ml 20× salts solution (cooled to <50◦ C; see recipe; no labeling required) 20 ml specific carbon source (see below) 2 ml 1 M MgSO4 0.2 ml 1000× trace metals solution (see recipe) 10 ml 100× vitamin solution (see recipe) 2 ml 500× antibiotic solution (plasmid dependent; see recipe) H2 O to a final volume of 1 liter For uniform isotopic labeling with isotopically enriched glucose: use a 20% (w/v) solution of unlabeled glucose in the conditioning medium. For uniform isotopic labeling with isotopically enriched sodium acetate: use a 15% (w/v) solution of unlabeled sodium acetate in the conditioning medium. For selective isotopic labeling with isotopically enriched pyruvate: use a 15% (w/v) solution of unlabeled pyruvate in the conditioning medium. For selective isotopic labeling with isotopically enriched glucose, α-ketoisovalerate and α-ketobutyrate: use a 20% (w/v) solution of unlabeled glucose in the conditioning medium.
Denaturing reducing buffer 5 ml 1 M Tris buffer, pH 7 (APPENDIX 2E) 57 g guanidinium·HCl 28.7 mg tris(carboxyethyl) phosphine (TCEP) Adjust final volume to 100 ml with H2 O Prepare fresh prior to use Deuterated minimal medium Combine the following, using sterilized stock solutions, just prior to use.
50 ml 20× salts (cooled to <50◦ C; see recipe) 20 ml specific carbon source (see below) 2 ml 1 M MgSO4 0.2 ml 1000× metals (see recipe) 2 ml 500× antibiotic (plasmid-dependent; see recipe) 2 H2 O (Cambridge Isotope Laboratories) to a final volume of 1 liter To uniformly label carbon and hydrogen: use a 20% (w/v) solution of 99% [U13 C,2 H]-glucose or a 15% (w/v) solution of 99% [U-13 C,2 H]-sodium pyruvate (Cambridge Isotopes Laboratories) as the sole carbon source. Use the same carbon source as in the conditioning medium, except isotopically enriched. To ensure full deuteration use salts solution and carbon source solution prepared with 2 H2 O. To fully 13 C label and almost completely deuterate the amides and Cα carbons and deuterate the Cβ carbons to >80%: use a 15% (w/v) solution of [U-13 C]-pyruvate as the sole carbon source (the methyl groups in the Leu, Val, le, and Ala residues will be protonated). To protonate only the Leu, Val, and Ile δ 1 methyl groups: use 20% (w/v) of [U13 C,2 H]-glucose and, in the protocol, add a solution of [3-2 H]-α-ketoisovalerate and [3,3 -2 H2 ]-α-ketobutyrate just prior to induction. To prepare 15 N, 13 C-labeled minimal medium: substitute 99% 15 NH4 Cl (Cambridge Isotopes Laboratories) for unlabeled NH4 Cl in the salts solution. Current Protocols in Protein Science
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Dialysis buffer Combine, using sterilized stock solutions, just prior to use.
230 ml 1 M sodium phosphate buffer, pH 7.7 (see APPENDIX 2E) 3.36 g EDTA 1.77 g DTT 575 ml glycerol H2 O to a final volume of 11.5 liters Store up to 3 days at room temperature FeCl3 (0.1 M) in diluted HCl Combine the following in the indicated order. 99 ml H2 O 1 ml concentrated HCl 1.622 g FeCl3 Sterilize by passing through a 0.22-µm filter and store up to 6 months at room temperature.
Fluorine-labeling medium Combine, using sterilized stock solutions, just prior to use. 50 ml 20× salts solution (cooled to <50◦ C; see recipe; no labeling required) 20 ml 20% (w/v) glucose 2 ml 1 M MgSO4 0.2 ml 1000× trace metals solution (see recipe) 10 ml 100× vitamin solution 2 ml 500× antibiotic (plasmid dependent; see recipe) 20 ml 50× auxotroph amino acids (see recipe) H2 O to a final volume of 1 liter Fluorinated amino acid stocks, 50× 50× fluorophenylalanine 1 g fluorophenylalanine H2 O to a final volume of 100 ml 50× fluorotryptophan 1 g fluorotryptophan H2 O to a final volume of 100 ml 50× fluorotyrosine 1 g fluorotyrosine H2 O to a final volume of 100 ml If necessary, adjust the pH to ∼7 with 1 M NaOH. Sterilize by passing through a 0.22-µm filter, divide into 10-ml aliquots, and store up to 6 months at −20◦ C.
Glucose-based minimal medium
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Combine, using sterilized stock solutions, just prior to use. 50 ml 20× salts solution (cooled to <50◦ C; see recipe) 20 ml 20% (w/v) glucose 2 ml 1 M MgSO4 0.2 ml 1000× trace metals solution (see recipe) 2 ml 500× antibiotic (plasmid dependent; see recipe) H2 O to a final volume of 1 liter continued
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To prepare 13 C labeled minimal medium: use 99% [13 C6 ]-glucose (Cambridge Isotopes Laboratories) as the sole carbon source. To prepare 15 N labeled minimal medium: substitute 99% 15 NH4 Cl (Cambridge Isotopes Laboratories) for unlabeled NH4 Cl in the salts solution. To prepare 15 N, 13 C labeled minimal medium: substitute 99% 15 NH4 Cl (Cambridge Isotopes Laboratories) is substituted for unlabeled NH4 Cl and use 99% [13 C6 ]glucose, (Cambridge Isotopes Laboratories) as the sole carbon source. 2
H-labeled formaldehyde solution, 0.1 M Add 32 mg 2 H-labeled paraformaldehyde (Cambridge Isotope Laboratories) to 10 ml water in a test tube. Seal the test tube and hydrolyze overnight at 100◦ C to 110◦ C. Prepare just prior to use.
α-Ketoisovalerate and α-ketobutyrate, 500×
500 mg α-ketoisovalerate (Cambridge Isotope Laboratories) 250 mg α-ketobutyrate (Cambridge Isotope Laboratories) H2 O to a final volume of 10 ml Sterilize by passing through a 0.22-µm filter, divide into 5-ml and 500-µl aliquots, and store up to 6 months at −20◦ C. 13
C-labeling of these components should be consistent with the desired protein labeling. See notes for the medium being prepared with this solution.
King’s reagent 8.25 ml trifluoroacetic acid (TFA; Sigma-Aldrich) 500 µl phenol (Sigma-Aldrich) 500 µl thioanisol (Sigma-Aldrich) 250 µl ethanedithiol (Sigma-Aldrich) 500 µl H2 O The phenol, thioanisol, and ethanedithiol may be premixed in a ratio of 2:2:1 and stored up to 6 months in a bottle in the fume hood. CAUTION: This mixture is very hazardous and noxious; prepare and store in a fume hood.
Large-scale dialysis solution 470.4 µl of 1.4 M CH3 COONH4 (ammonium acetate) 720 µl of 0.5 M Mg(CH3 COO)2 (magnesium acetate) 960 µl amino acid mixture (see recipe) 1920 µl of 0.645 M creatine phosphate DEPC-treated H2 O to 24 ml Prepare fresh just prior to use Use [15 N]-CH3 COONH4 if expressing 15 N-labeled protein.
LM mixture 22 ml 2 M HEPES·KOH, pH 7.5 33.4 ml 6 M KOOCCH3 (potassium acetate; include only for large-scale protein expression) 210 mg DTT 530 mg ATP continued
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338 mg CTP 335 mg GTP 310 mg UTP 172 mg cAMP 28 mg folinic acid 140 mg tRNA (Roche) 64 ml of 50% (w/v) PEG-8000 (include only for large-scale protein expression) DEPC-treated H2 O to a final volume of 200 ml Dispense in 2-ml aliquots and store up to 6 months at −20◦ C Lysis buffer for T7 RNA polymerase Combine, using sterilize stock solutions, just prior to use. 1.25 ml 1 M Tris buffer, pH 8.1 (APPENDIX 2E) 29 mg NaCl 15 mg EDTA 4 mg dithiothreitol (DTT) 7.5 mg lysozyme ∼1 µl 0.2 M PMSF 100 µl 1 mg/ml solution of bacitracin (Calbiochem) 60 µl 1 mg/ml solution of benzamidine (Sigma) H2 O to 25 ml Nondenaturing reducing buffer 5 ml 1 M MOPS buffer, pH 7.0 373 mg KCl 28.7 mg tris(carboxyethyl) phosphine (TCEP) 20 mg MgCl2 H2 O to a final volume of 100 ml Store up to 3 days at room temperature Overnight medium Combine, using sterilized stock solutions, just prior to use. 2.5 ml 20× salts solution (cooled to <50◦ C) 1.25 ml 20% glucose 0.1 ml 1 M MgSO4 10 µl 1000× metals 0.5 ml 100× vitamins 1 ml 50× amino acids solution (lacking cysteine, tyrosine, and methionine), see recipe 0.1 ml 500× antibiotic (plasmid-dependent) H2 O to a final volume of 50 ml S30 Buffer
Incorporation of Isotopically Enriched Amino Acids
Combine, using sterilized stock solutions, just prior to use. 50 ml 1 M Tris buffer, pH 8.2 (APPENDIX 2E) 5.83 g Mg(CH3 COO)2 (magnesium acetate) 29.43 g KOOCCH3 (potassium acetate) 771 mg DTT 2.5 ml 2-mercaptoethanol H2 O to a final volume of 5 liters
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Salts solution, 20× 268.1 g Na2 HPO4 ·7 H2 O 136.1 g KH2 PO4 53.5 g unlabeled or [U-15 N]NH4 Cl 14.2 g Na2 SO4 H2 O to 1 liter Divide the salt solution into 200-ml aliquots and sterilize by autoclaving 15 min at 15 psi (1.05 kg/cm2 ) on liquid cycle. Store up to 6 months at room temperature. Use unlabeled NH4 Cl if 15 N labeling is not required.
Small-scale dialysis solution 39.2 µl 1.4 M CH3 COONH4 (ammonium acetate; see recipe) 60 µl 0.5 M Mg(CH3 COO)2 (magnesium acetate; see recipe) 80 µl amino acid mixture (see recipe) 160 µl 0.645 M creatine phosphate 500 µl LM mixture (see recipe) DEPC-treated H2 O to 2 ml Use [15 N]-CH3 COONH4 if expressing 15 N-labeled protein.
Solution 2 65 mg KCN (Sigma-Aldrich; 1 mM final concentration) Dissolve and bring final volume to 20 ml with H2 O Take 400 µl of this solution and dilute to 20 ml with pyridine CAUTION: Potassium cyanide is extremely toxic. Follow all precautions recommended in the MSDS.
Specific carbon source-based minimal medium Combine the following sterilized stock solutions: 50 ml 20× salts solution (cooled to <50◦ C; see recipe) 20 ml specific carbon source (see below) 2 ml 1 M MgSO4 0.2 ml 1000× trace metals solution (see recipe) 10 ml 100× vitamin solution (see recipe) 2 ml 500× antibiotic solution (plasmid-dependent; see recipe) H2 O to a final volume of 1 liter To label alternate carbon atoms: a 20% (w/v) solution of 99% [1-13 C]-glucose or 99% [2-13 C]-glycerol (Cambridge Isotopes Laboratories) is used as the sole carbon source. Alternatively, to label alternate carbon atoms: use a 15% (w/v) solution of 1-13 C sodium acetate or 2-13 C sodium acetate To label carbons in the opposite pattern: use a 20% (w/v) solution of 99% [1,313 C ]-glycerol (Cambridge Isotopes Laboratories) as the carbon source. 2 To label 50% of carbons at all positions: use a 50/50 mix of 15% (w/v) solutions of 1-13 C and 2-13 C sodium acetate. To label only the methyl groups on Ala, Ile, Leu, and Val residues: use a 15% w/v solution of 3-13 C-pyruvate. continued
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To 13 C label only the L, V, and I-δ 1 methyl groups: use 20% (w/v) of unlabeled glucose and in the protocol add a solution of [dimethyl-13 C2 ]-α-ketoisovalerate and [methyl-13 C]-α-ketobutyrate just prior to induction. To prepare 15 N, 13 C labeled minimal medium: substitute 99% 15 NH4 Cl (Cambridge Isotopes Laboratories) for unlabeled NH4 Cl.
TEASE amino acids, 50× 1 g each of: monosodium glutamate asparagine monohydrate arginine monohydrochloride lysine monohydrochloride 1 g each free acids of: aspartic acid proline threonine serine glutamate isoleucine H2 O to 100 ml If necessary, adjust the pH to ∼7 with 1 M NaOH. Sterilize by passing through a 0.22-µm filter, dispense into 20-ml aliquots, and store up to 6 months at −20◦ C.
TEASE minimal medium Combine the following, using sterilized stock solutions, just prior to use.
50 ml 20× salts (cooled to <50◦ C; see recipe) 20 ml 20% (w/v) 2-13 C glycerol 2 ml 1 M MgSO4 0.2 ml 1000× metals (see recipe) 2 ml 500× antibiotic (plasmid-dependent; see recipe) 20 ml 50× TEASE amino acids (see recipe) H2 O to a final volume of 1 liter Trace metal solution, 1000× 50 ml 0.1 M FeCl3 in ∼60 mM HCl (see recipe) 2 ml 1 M CaCl2 2 ml 0.5 M MnCl2 2 m1 0.5 M ZnSO4 0.5 ml 0.4 M CoCl2 0.5 ml 0.4 M CuCl2 0.5 ml 0.4 M NiCl2 0.5 ml 0.4 M Na2 MoO4 0.5 ml 0.4 M Na2 SeO3 0.5 ml 0.4 M H3 BO3 Sterilize by passing through a 0.22-µm filter and store up to 6 months at room temperature. Tris buffer for T7 RNA polymerase Combine, using sterilized stock solutions, just prior to use. Incorporation of Isotopically Enriched Amino Acids
11.25 ml 1 M Tris buffer, pH 8.1 (APPENDIX 2E) 175 mg NaCl 88 mg EDTA continued
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23 mg DTT H2 O to a final volume of 150 ml Vitamin stock solution, 100× (Venters et al., 1995) To 100 ml of H2 O, add: 10 mg biotin 10 mg choline-Cl 10 mg folic acid 10 mg niacinamide 10 mg D-pantothenate 10 mg pyridoxal 1 mg riboflavin Sterilize by passing through a 0.22-µm filter, divide into 1- to 5-ml aliquots, and store up to 6 months at −20◦ C.
COMMENTARY Background Information Isotope enrichment increases sensitivity, can improve resolution, and permits the selection of specific interactions in the application of NMR, neutron diffraction, and mass spectrometry to biomolecules and thus is used extensively when determining protein structure or measuring dynamics. The isotopic labeling strategy selected is determined by the source of the protein, the physical technique which will be used, the molecular size of the protein, and the particular goals of the experiment. The focus of this unit is on isotope enrichment for NMR applications since this field has the greatest variety of labeling strategies and protocols. However, many of the protocols are equally well-suited to specific labeling strategies for mass spectrometry and neutron diffraction applications, with some minor adaptations. In solution NMR spectroscopy, general assignments of resonances and structural measurements can be made on small peptides and proteins (<50 amino acids) without isotopic labeling using currently available NMR techniques. However, for obtaining dynamic information or for studying proteins of ∼50 to 80 residues, universal isotopic labeling of the nitrogen atoms with 15 N is recommended. This allows measurement of sequential connectivities along the protein backbone, as well as measurement of other inter-residue interactions. For intermediate-sized proteins (∼80 to 160 amino acids), universal isotopic labeling of two nuclei (13 C, 15 N) is warranted. For large proteins (>160 amino acids), isotopic labeling of three nuclei (13 C, 15 N, 2 H) is often required to minimize the deleterious effects of 1 H-1 H dipolar couplings. For proteins requiring uni-
versal isotope incorporation, recombinant protein production is typically necessary due to the prohibitive expense of isotope incorporation via chemical synthesis or via nutrient enrichment in higher organisms. When possible, protein production should be carried out utilizing a prokaryotic organism such as E. coli, since this route is the most cost effective and allows the most flexibility. However, if protein production with a prokaryotic system is not possible, or a protein with post-translational modifications is desired, the use of a eukaryotic expression system, such as the baculovirus system, may be warranted. Additionally, it is possible to have tissue culture media custommade with isotopically enriched reagents, but due to the prohibitive cost and specialized application of this approach, a protocol is not presented for it in this unit. Finally, in recent years, the development of cell-free systems has led to another avenue for isotope labeling which can, with extensive practice, be a very efficient means of producing isotopically enriched proteins. In solid state NMR, specific atoms are often isotopically enriched. This can be accomplished either via synthetic methods, such as solid phase peptide synthesis and chemical modification, or by recombinant methodology using selectively labeled reagents. Given the selective nature of the labeling, thus far most published work has relied on samples which were either made synthetically or produced recombinantly in E. coli. However, with the advent of more sophisticated, less expensive, commercially available cellfree systems, this may become the means of choice for producing isotopically enriched samples.
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The strategies used for labeling proteins via recombinant protein production parallel those used for the production of their unlabeled analogues. Thus, this unit contains protocols that are similar to those previously published. The major modifications are either to the carbon source, other specifically-labeled reagents, or slight changes at key steps in the protein production process. These changes are made to introduce the specific isotope labels of interest in the final protein. From a practical standpoint, isotopic labeling of proteins using cell culture depends on high expression levels of the proteins under the culture conditions used, a high percentage of incorporation of isotopic label into the protein, limited cross-labeling of amino acid residues if specific residues are being labeled, and a minimization of the costs involved. Recombinant protein expression from E. coli is covered in UNITS 5.1 – 5.3. The best method to enhance the yield of expressed labeled proteins in bacteria is to use labeled algal lysates as substrates. But the costs involved make this method one of the most expensive methods currently available. Thus, the protocols outlined in this unit rely primarily on the use of minimal media along with isotopically enriched reagents. Protein production using the baculovirus expression system is described in detail in UNITS 5.4 & 5.5. Most formulations for insect culture that are currently available do not have a well defined composition to account for sources of H, C, and N in the recombinant protein product. However, there are a few commercially available products that are specially formulated for labeling proteins in insect cell culture. Due to the specialized nature of this approach, producing isotopically labeled proteins in insect cells is relatively more expensive. Thus, for proteins that cannot be synthesized in a prokaryotic system, the cell-free approach to protein production is advantageous in that it allows for the robust production of eukaryotic proteins (see UNIT 5.18 for a detailed overview). For proteins which cannot be expressed recombinantly, the isolated and purified protein can be chemically modified with isotopicallylabeled reagents in order to perform spectroscopic analyses. For smaller peptides, isotopes can be incorporated in the course of chemically synthesizing the peptide. This unit contains protocols for both chemically synthesizing peptides and chemically modifying proteins with isotopically labeled reagents. These protocols are similar to those outlined in other units with the exception that the isotopically enriched reagent is always the limiting reagent
in order to minimize cost. An overview of the field of synthetic peptides and proteins is given in UNIT 18.1, along with additional information regarding common side reactions, synthesizing modified residues and the selection of common coupling reagents. A very detailed discussion about the selection of the solid support resin is presented in UNIT 18.7. UNITS 15.1 and UNIT 15.2 provide overviews of the modification of cysteine residues and amino groups, respectively. These units contain the analogous protocols for chemically modifying proteins in the absence of isotopic labels.
Critical Parameters and Troubleshooting For most of the protocols in this unit, the quantities of reagents have been adjusted such that the reagent that is the source of the isotope label has been set as the limiting reagent for cost effectiveness. In all cases it is recommended that the protocol first be performed without labeled reagents and optimized with regard to the protein of interest. Optimization of cell growth and expression times can significantly improve the overall yield and should be done prior to scaling up and using isotopically enriched reagents. Most recombinant proteins are currently expressed in E. coli. However, to reduce the cost of producing isotopically labeled proteins, various strategies have been tried, such as the addition of supplements to minimal medium and the implementation of fermentors. Since one of the most important factors in protein production is the density of viable cells in the medium, initial growth of E. coli cells can be carried out in an unlabeled, enriched medium such as LB medium. The cells can then be exchanged into an isotopically defined minimal medium prior to induction of protein synthesis. This is because fully labeled cells are not an absolute requirement for the incorporation of higher levels of isotopes in proteins. If minimal medium is used to grow the cells, exchanging the medium used for fresh medium prior to induction also may play a role in protein production by the removal of toxic byproducts of regular metabolism, thus enhancing levels of protein expression. The critical parameters and yield of expressed labeled proteins in insect cells are described in UNIT 5.5. One of the major drawbacks of protein expression in insect cells is the time frame involved in expressing proteins. It would typically require 9 to 12 days to complete a round of expression. As in the case of
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protein expression in bacterial cells, switching the insect cells from a normal medium such as SF900 to the labeled medium such as BioExpress-2000 may alter the expression of proteins, so growth and expression times will need to be verified prior to culturing and expressing large quantities of cells.
Anticipated Results Many of the protocols described in this unit are very similar to protocols for producing or chemically synthesizing unlabeled proteins and peptides. Before using a protocol to make a labeled sample, it should be tested and optimized for producing or synthesizing the unlabeled analogues. Once this has been done, utilizing these protocols for label incorporation is fairly straightforward. When using E. coli, the yield of labeled recombinant protein varies depending on the growth conditions, carbon source, and type of protein expressed. The yield of certain soluble proteins using shaken cultures and IPTG induction could be as high as 40 to 80 mg/liter of minimal medium, whereas others may be much lower. The use of labeled auto-induction media has been shown to yield amounts comparable to those in enriched media using IPTG induction. For the synthesis and chemical modification protocols, the yields are similar to those expected for solid phase peptide synthesis and peptide chemical modification described in previous units, and the degree of isotope incorporation throughout is expected to be high (≥90%).
Time Considerations Time considerations for recombinant protein expression in E. coli are reviewed in UNITS 5.2 & 5.3. In addition to the time required to test the various parameters such as cell type, plasmid stability and growth conditions required for normal protein expression, 2 to 3 days should be allocated to test the expression of the proteins in the analogous unlabeled media. The growth and expression of labeled proteins may vary depending on the source of labels from the substrates used. One of the major drawbacks of protein expression in insect cells (i.e., the baculovirus expression system) is the time frame involved for expressing proteins. Approximately 3 days is required to generate the PCR product, perform the TOPO cloning reaction, transform the cells, and isolate and analyze the purified plasmids. It generally takes up to 7 days to generate
a high titer of P2 viral stock. This is followed by up to 5 days for protein expression. Isotope incorporation through solid phase peptide synthesis can take up to 3 weeks to make a 24-amino acid peptide, if amino acid coupling is done at 3 cycles/day (which can fit into a conventional working day), and includes time allotted for side chain deprotection, wetting and drying of the resin. The purification step (RP-HPLC or other liquid chromatography) can take several days, depending on the quantity of the peptide. Processing of the purified peptide (concentration and lyophilization) can take 1 to 2 days (depending on the volume). The preparation of Fmoc-protected amino acids can take up to 2 days, including recrystallization of the crude product. The incorporation of isotope labels by chemical exchange requires several hours for the lyophilization step (this depends on the volume of the frozen peptide solution), while the exchange step typically requires only 30 min for an unfolded protein. The dialysis step (required for refolding of an unfolded protein) usually requires 6 to 24 hr. The chemical modification step to incorporate isotope labels requires 2 hr and the dialysis step typically requires an additional 6 to 24 hr.
Literature Cited Brauer, M. and Sykes, B.D. 1986. 19 F nuclear magnetic resonance studies of selectively fluorinated derivatives of G- and F-actin. Biochemistry 25:2187-2191. Br¨uggert, M., Rehm, T., Shankar. S., Georgescu, J., and Holak, T.A. 2003. A novel medium for expression of proteins selectively labeled with 15 N-amino acids in Spodoptera frugiperda (Sf9) insect cells. J. Biomol. N.M.R. 25:335-348. Chan, W.C. and White, P.D. 2000. FMOC Solid Phase Peptide Synthesis: A Practical Approach, Oxford University Press, Oxford. Chrunyk, B.A., Evans, J., Lillquist, J., Young, P., and Wetzel, R. 1993. Inclusion-body formation and protein stability in sequence variants of interleukin-β. J. Biol. Chem. 268:18053-18061. Cowburn, D. and Muir, T.W. 2001. Segmental isotopic labeling using expressed protein ligation. Method Enzymol. 339:41-54. Creemers, A.F.L., Klaassen, C.H.W., BoveeGeurts, P.H.M., Kelle, R., Kragl, U., Raap, J., de Grip, W.J., Lugtenburg, J., and de Groot, H.J.M. 1999. Solid state 15 N NMR evidence for a complex Schiff base counterion in the visual G-protein receptor rhodopsin. Biochemistry 38:7195-7199. Davanloo, P., Rosenberg, A.H., Dunn, J.J., and Studier, F.W. 1984. Cloning and expression of the gene for bacteriophage T7 RNA polymerase. Proc Natl Acad Sci U.S.A. 81:2035-2039.
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David, R., Richter, P.O., and Beck-Sickinger, A.G. 2004. Expressed protein ligation, methods and applications. Eur J. Biochem. 271:663-677. DeLange, F., Klaassen, C.H.W., Wallace-Williams, S.E., Bovee-Geurts, P.H.M., Liu, X-M., de Grip, W.J., and Rothschild, K.J. 1998. Tyrosine structural changes detected during the photoactivation of rhodopsin. J. Biol. Chem. 273:2373523739. Englander, S.W. and Poulsen, A. 1969. Hydrogentritium exchange of the random chain polypeptide. Biopolymers 7:379-393. Gardner, K.H. and Kay, L.E. 1997. Production and incorporation of 15 N, 13 C, 2 H (1 H-δ1 methyl) isoleucine into proteins for multidimensional NMR studies. J. Am. Chem. Soc. 119:75997600. Gerig, J.T. 2001. Fluorine NMR. Biophysics Textbook Online. 1-35. Goff, S.A. and Goldberg, A.L. 1987. An increased content of protease LA, the Lon gene product, increases protein degradation and blocks growth in Escherichia coli. J. Biol. Chem. 262:45084515. Goto, N.K. and Kay, L.E. 2000. New developments in isotope labeling strategies for protein solution NMR spectroscopy. Curr. Opin. Struc. Biol. 10:585-592. Goto, N.K., Gardner, K.H., Mueller, G.A., Willis, R.C., and Kay, L.E. 1999. A robust and costeffective method for the production of Val, Leu, Ile (δ1) methyl-protonated 15 N-, 13 C-, 2 Hlabeled proteins. J. Biomol. N.M.R. 13:369374. Hajduk, P.J., Augeri, D.J., Mack, J., Mendoza, R., Yang, J., Betz, S.F. and Fesik, S.W. 2000. NMR-based screening of proteins containing 13 C-labeled methyl groups. J. Am. Chem. Soc. 122:7898-7904. Henrich, B., Lubitz, W. and Plapp, R. 1982. Lysis of Escherichia coli by induction of cloned φ X174 genes. Mol. Gen. Genet. 185:493-497. Hong, M. and Jakes, K. 1999. Selective and extensive 13 C labeling of a membrane protein for solid-state NMR investigations. J. Biomol. N.M.R. 14:71-74. Ishima, R., Louis, J.M., and Torchia, D.A. 1999. Transverse 13 C relaxation of CHD2 methyl isotopmers to detect slow conformational changes of protein side chains. J. Am. Chem. Soc. 121:11589-11590. Jentoft, N. and Dearborn, D.P.G., 1983. Protein labeling by reductive alkylation. Methods in Enzymolgy 19:570-579. Klammt, C., Lohr, F., Schafer, B., Haase, W. Dotsch, V., Ruterjans, H., Glaubitz, C., and Bernhard, F. 2004. High level cell free expression and specific labeling of integral membrane proteins. Eur. J. Biochem. 271:568-580. Incorporation of Isotopically Enriched Amino Acids
Kotik, M., Radford, S.E., and Dobson, C.M. 1995. Comparison of the refolding of hen lysozyme from dimethyl sulfoxide and guanidinium chloride. Biochemistry 34:1714-1724.
Kranz, James K., Lu, J. and Hall, K.B. 1996. Contribution of the tyrosines to the structure and Function of the human U1A N-terminal RNA binding domain. Protein Sci. 5:1567-1583. Lee, A.L., Urbauer, J.L. and Wand, J.A. 1997. Improved labeling strategy for 13 C relaxation measurements of methyl groups in proteins. J. Biomol. N.M.R. 9:437-440. LeMaster, D.M. and Kushlan, D.M. 1996. Dynamical Mapping of E. coli Thioredoxin via 13 C NMR Relaxation Analysis. J. Am. Chem. Soc. 118:9255-9264. Lian, L-Y. and Middleton, D.A. 2001. Labelling approaches for protein structural studies for solution-state and solid-state NMR. Prog. Nucl. Mag. Res. Spec. 39:171-190. Nirenberg, M.W. and Matthaei, J.H. 1961. The dependence of cell-free protein synthesis in E. coli upon naturally occurring or synthetic polyribonucleotides. Proc. Natl. Acad. Sci. U.S.A. 47:1588-1602. Ozawa, K., Wu, P.S., Dixon, N.E., and Otting, G. 2006. 15 N-labeled proteins by cell-free protein synthesis. Strategies for high-throughput NMR studies of proteins and protein-ligand complexes. FEBS J. 273:4154-4159. Ralf, D., Richter, M.P.O., and Beck-Sickinger, A.G. 2004. Expressed protein ligation. Eur. J. Biochem. 271:663-677. Rohl, C.A., Scholtz, J.M., York, E.J., Stewart, J.M. and Baldwin, R.L. 1992. Kinetics of amide proton exchange in helical peptides of varying chain lengths. Interpretation by the Lifson-Roig Equation. Biochemistry 31:1263-1269. Rosen, M.K., Gardner, K.H., Willis, R.C., Parris, W.E., Pawson, T., and Kay, L.E. 1996. Selective methyl group protonation of perdeuterated proteins. J. Mol. Biol. 263: 627-636. Sambrook, J., Fritsch, E.F., and Maniatis, T. 1989. Molecular Cloning: A Laboratory Manual, 2nd ed. Cold Spring Harbor Press, Cold Spring Harbor, New York. Spooner, P.J. and Watts, A., 1991. Reversible unfolding of cytochrome c upon interaction with cardiolipin bilayers. 1.Evidence from deuterium NMR measurements. Biochemistry 30:38713879. Strauss, A., Bitsch, F., Fendrich, G., Cutting, B., Fendrich, G., Graff, P., Liebetanz, J., Zurini, M. and Jahnke, W. 2003. Amino-acid-type selective isotope labeling of protein expressed in Baculovirus-infected cells useful for NMR studies. J. Biomol. N.M.R. 26:347-372. Strauss, A., Bitsch, F., Fendrich, G., Graff, P., Knecht, R., Meyhack, B. and Jahnke, W. 2005. Efficient uniform isotope labeling of ab1 kinase expressed in Baculovirus-infected cells. J. Biomol. N.M.R. 31:343-349. Studier, F.W. 2005. Protein production by autoinduction in high density shaking cultures. Protein Expr. Purif. 41:207-234. Studier, F.W. and Moffatt, B.A. 1986. Use of bacteriophage T7 RNA polymerase to direct selective
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high-level expression of cloned genes. J. Mol. Biol. 189:113-130. Studier, F.W., Rosenberg, A.H., Dunn, J.J., and Dubendorff, J.W. 1990. Use of T7 RNA polymerase to direct expression of cloned genes. Methods Enzymol. 185:60-89. Torizawa, T., Shimizu, M., Taoka, M., Miyano, H., and Kainosho, M. 2004. Efficient production of isotopically labeled proteins by cell-free synthesis: A practical protocol. J. Biomol. N.M.R. 30:311-325. Tyler, R.C., Sreenath, H.K., Singh, S., Aceti, D.J., Bingman, C.A., Markley, J.L., and Fox, B.G. 2005. Auto-induction medium for the production of [U-15 N]- and [U-13 C, U-15 N]-labeled proteins for NMR screening and structure determination. Protein Expr. Purif. 40:268-278.
Fields, G.B. and Noble, R.L. 1990. Solid phase peptide synthesis utilizing 9-fluorenylmethoxycarbonyl amino acids, Int. J. Peptide Protein Res. 35:161-214. An overview of solid phase peptide synthesis methodology. pENTR Directional TOPO Cloning Kit Instruction Manual, Version G, 2006. Invitrogen, Carlsbad, Calif. Comprehensive instruction manual for the pENTR Directional TOPO Cloning Kits for directional cloning of a blunt-end PCR product into a vector for entry into Invitrogen’s Gateway system. Sambrook et al., 1989. See above. A resource for general molecular biology cloning and expression techniques.
Venters, R.A., Huang, C-C., Farmer II, B.T., Trolard, R., Spicer, L.D., and Fierke, C.A. 1995. High-level 2 H/13 C/15 N labeling of proteins for NMR studies. J. Biomol. N.M.R. 5:339-344.
Strauss et al., 2005. See above. Describes the use of commercially available BioExpress media to express both unlabeled and isotopically labeled recombinant proteins using insect cells.
Vinarov, D.A., Lytle, B.L., Peterson, F.C., Tyler, E.M., Volkman, B.F., and Markley, J.L. 2004. Nat. Methods 1:1-5.
Studier, 2005. See above. This provides a detailed account of auto induction of recombinant proteins in E. coli.
Key References BaculoDirect Baculovirus Expression Systems Instruction Manual, Version F, 2004. Invitrogen, Carlsbad, Calif. This manual describes the steps involved in forming a recombinant baculovirus with the gene of interest, which can subsequently be used to transfect insect cells for protein expression, using the BaculoDirect Baculovirus Expression System. Chan, W.C. and White, P.D. 2000. Fmoc Solid Phase Peptide Synthesis: A Practical Approach. Oxford University Press, Oxford. Covers the essential procedures for the production of linear peptides as well as new methodological advances in the field.
Torizawa et al., 2004. See above. This provides an excellent account of the protocol for the expression of labeled proteins using cell-free systems which includes modifications such as continuous diffusion of by-products and replenishments of precursors and energy regeneration.
Contributed by Mini Samuel-Landtiser, Cherian Zachariah, Chris R. Williams, Arthur S. Edison, and Joanna R. Long University of Florida Gainesville, Florida
Englander, S.W. and Poulsen, A. 1969. Hydrogentritium exchange of the random chain polypeptide. Biopolymers 7:379-393. Describes the kinetics of hydrogen exchange in proteins as a functin of pH.
Protein Engineering
26.3.49 Current Protocols in Protein Science
Supplement 47
USEFUL DATA
APPENDIX 1
Characteristics of Amino Acids
APPENDIX 1A
The physical properties of the amino acids determine the structure and function of the proteins in which they are found. Some useful details and relevant physical characteristics of the amino acids can be found in Table A.1A.1. Information about modified amino acids is provided in Table A.1A.2. The chemical structures of the amino acids, and an explanation of the role these structures play in enzymes, can be found in Figure A.1A.1. The three-dimensional structure of proteins is largely determined by the packing of their hydrophobic cores; the properties of amino acids that govern this packing are their relative hydrophobicities, which are discussed in UNIT 2.2, and their shapes and volumes, which can be assessed by referring to the space-filling models shown in Figure A.1A.2. Post-translational modifications will change the mass of a protein or peptide; values for some common mass changes are listed in Table A.1A.3. The changes in mass resulting from modification of proteins with different carbohydrates are shown in Table A.1A.4.
LITERATURE CITED Matthew, J.B., Friend, S.H., Botelho, L.H., Lehman, L.D., Hanania, G.I.H., and Gurd, F.R.N. 1979. Biochem. Biophys. Res. Commun. 81:416-421.
Useful Data Current Protocols in Protein Science (1995) A.1A.1-A.1A.8 Copyright © 2000 by John Wiley & Sons, Inc.
A.1A.1 Supplement 4
Table A.1A.1
Compositions and Masses of the Twenty Commonly Occurring Amino Acid Residuesa,b
Monoisotopic mass
Average mass
pKa of ionizing side chainc
Occurrence in proteins (%)
C3H5NO C6H12N4O
71.03711 156.10111
71.0788 156.1876
— ~11.5–12.5 (12)
8.3 5.7
Asparagine (Asn, N) Aspartic acid (Asp, D)
C4H6N2O2 C4H5NO3
114.04293 115.02694
114.1039 115.0886
— 3.9–4.5 (4)
4.4 5.3
Cysteine (Cys, C) Glutamic acid (Glu, E)
C3H5NOS C5H7NO3
103.00919 129.04259
103.1448 129.1155
8.3–9.5 (9) 4.3–4.5 (4.5)
1.7 6.2
Glutamine (Gln, Q) Glycine (Gly, G)
C5H8N2O2 C2H3NO
128.05858 57.02146
128.1308 57.0520
— —
4.0 7.2
Histidine (His, H) Isoleucine (Ile, I)
C6H7N3O C6H11NO
137.05891 113.08406
137.1412 113.1595
6.0–7.0 (6.3) —
2.2 5.2
Leucine (Leu, L) Lysine (Lys, K)
C6H11NO C6H12N2O
113.08406 128.09496
113.1595 128.1742
— 10.4–11.1 (10.4)
9.0 5.7
Methionine (Met, M) Phenylalanine (Phe, F)
C5H9NOS C9H9NO
131.04049 147.06841
131.1986 147.1766
— —
2.4 3.9
Proline (Pro, P) Serine (Ser, S)
C5H7NO C3H5NO2
97.05276 87.03203
97.1167 87.0782
— —
5.1 6.9
Threonine (Thr, T) Tryptophan (Trp, W)
C4H7NO2 C11H10N2O
101.04768 186.07931
101.1051 186.2133
— —
5.8 1.3
Tyrosine (Tyr, Y) Valine (Val, V)
C9H9NO2 C5H9NO
163.06333 99.06841
163.1760 99.1326
9.7–10.1 (10.0) —
3.2 6.6
Name
Composition
Alanine (Ala, A) Arginine (Arg, R)
aFor corresponding structures, see Figure A.1A.1 and Figure A.1A.2. bThe
molecular mass of a normally terminated and unmodified peptide or protein may be calculated by summing the masses of the appropriate amino acid residues and adding the masses of H and OH for the N and C termini, respectively. In cases where cysteines are linked to form disulfide bridges, the mass of two hydrogen atoms should be subtracted for each disulfide bridge in the molecule. Specifically, monoisotopic masses were calculated using the atomic masses of the most abundant isotope of the elements: C = 12.0000000, H = 1.0078250, N = 14.0030740, O = 15.9949146, and S = 31.9720718. Average masses were calculated using the atomic weights of the elements: C = 12.011, H = 1.00794, N = 14.00674, O = 15.9994, and S = 32.066.
cThese values are included for anyone wishing to make approximate isoelectric point determinations based on protein composition. Values for the terminal
residues depend on the identity of the residue: α-amino, pKa 6.8–8.2 (8.0); α-carboxyl, pKa 3.2–4.3 (3.6). Values in parentheses are based on those given by Matthew et al. (1978) and provide a good starting point for determinations.
Characteristics of Amino Acids
A.1A.2 Supplement 4
Current Protocols in Protein Science
Table A.1A.2
Compositions and Masses of Unusual and Modified Amino Acid Residuesa
Name
Composition
Monoisotopic mass
Carboxymethylcysteine (Cme) Carboxyamidomethyl cysteine (Cam) Pyridylethylcysteine (PECys) Acrylamide-modified cysteine (AACys) Cysteic acid [Cys(O3H)] Dehydroalanine (Dha) 4-Carboxyglutamic acid (Gla) Homoserine (Hse) Hydroxylysine (Hyl) Hydroxyproline (Hyp) Isovaline (Iva) Norleucine (Nlc) Ornithine (Orn) Pyroglutamic acid (Pyr) Sarcosine (Sar)
C5H7NO3S C5H8N2O2S C10H12N2OS C6H10N2O2S C3H5NO4S C3H3NO C4H5NO C4H7NO2 C6H12N2O2 C5H7NO2 C5H9NO C6H11NO C5H10N2O C5H5NO2 C3H5NO
161.01466 160.030650 208.06703 174.04631 150.99393 69.02146 173.03242 101.04768 144.08988 113.04768 99.06841 113.08406 114.07931 111.03203 71.03711
Average mass 161.1815 160.195940 208.23239 174.22363 151.1430 69.0630 173.1253 101.1051 144.1736 113.1161 99.1326 113.1595 114.1473 111.1002 71.0788
aMonoisotopic
masses were calculated using the atomic masses of the most abundant isotope of the elements: C = 12.0000000, H = 1.0078250, N = 14.0030740, O = 15.9949146, and S = 31.9720718. Average masses were calculated using the atomic weights of the elements: C = 12.011, H = 1.00794, N = 14.00674, O = 15.9994, and S = 32.066.
Useful Data
A.1A.3 Current Protocols in Protein Science
Supplement 4
A Amino acids with dissociable protons 10.5 OH
acidic
4.1 COOH
3.9 COOH H +
H3N
COO
8.4 SH
13.7 OH
H +
–
H3N
aspartate
H
COO
–
glutamate
+
H3N
H
COO
+
–
H3N
cysteine
H
COO
+
–
H3N
tyrosine
COO
–
serine
12.5
6.0 H + N
NH 2 + NH2 NH
10.5 NH3 +
N H H +
H3N
H
COO
+
–
COO
H3N
histidine
H +
–
COO
H3N
–
arginine
lysine
basic
Other amino acids with polar side chains O O
NH2
NH2
H +
H3N
COO
asparagine
+
H3N
COO
glutamine
H
H3C
H –
OH
–
+
H3N
H COO
–
threonine
Characteristics of Amino Acids
A.1A.4 Supplement 4
Current Protocols in Protein Science
B Nonpolar amino acids
H +
H3N
H
H COO
– N COO H + H
–
glycine
+
H3C
H
H3N
COO
proline
–
alanine
H
CH3
N
S CH3 H +
H3N
COO
–
+
H3N
tryptophan
CH3
COO
–
+
H3C –
leucine Figure A.1A.1 Line drawings of the amino acids. The amino acids are roughly divided into three groups: amino acids with dissociable protons (A), other amino acids with polar side chains (A), and nonpolar amino acids (B). These groupings are designed to facilitate an understanding of enzymology and the thermodynamics of protein folding. In this representation, hydrogens are omitted except in showing ionization or stereochemistry. In the case of arginine the delocalized positive charge is indicated by dashed double bonds. At stereocenters, bold lines indicate a group is coming out of the page toward the viewer, while hashed lines indicate that the group goes into the page away from the viewer. Amino acids with dissociable protons are generally intimately involved in the chemistry of enzymes. Acidic and basic groups can form salt bridges to substrates or to each other. They can also act as proton donors/acceptors in mechanisms that rely on acid/base catalysis. The polar side chains of some of these amino acids (notably cysteine, serine, and histidine) can act as nucleophiles. The pKa values for the free amino acids are shown, but these values
+
H3N
H3N
COO
–
methionine
CH3 H H
CH3
H3N
COO
H
valine
H +
H
H3C
COO
isoleucine
H –
+
H3N
COO
–
phenylalanine
can markedly change when these groups are buried in proteins. The pKas of the α-amino groups range from 8.7 to 10.7, while the pKas of the α-carboxylates range from 1.8 to 2.4 (see Table A.1A.1). Amino acids with polar side chains (A) can form hydrogen bonds to substrates or to each other. Cysteine, serine, and tyrosine could also be included in this group, as the ionized forms of these amino acids do not generally perform structural roles in proteins. In general, these amino acids (and the amino acids with dissociable protons) will be found on the surfaces of proteins. Cysteine is an exception, since it is slightly hydrophobic and can often be buried as a disulfide bond. The nonpolar amino acids (B) are often found in the interiors of proteins or in hydrophobic substrate-binding pockets. They interact with one another like jigsaw pieces, forming tight-fitting associations that have a density similar to that of an amino acid crystal. Proline is buried less frequently than might be expected because of its predominance in turns, which are often found on the periphery of a protein.
Useful Data
A.1A.5 Current Protocols in Protein Science
Supplement 4
glycine
aspartate
methionine
alanine
valine
glutamate
serine
cysteine
asparagine
leucine
histidine
threonine
proline
isoleucine
lysine
glutamine
phenylalanine
C
N H
arginine
tyrosine
tryptophan
S
O
Figure A.1A.2 Space-filling representations of the amino acids. The amino acids are arranged in order of size. The conformations shown maximize the two-dimensional area but are not necessarily the most stable geometries.
Characteristics of Amino Acids
A.1A.6 Supplement 4
Current Protocols in Protein Science
Table A.1A.3 Mass Changes Due to Some Post-Translational Modifications of Peptides and Proteinsa
Modificationb Common modifications Pyroglutamic acid formation from Gln Disulfide bond (cystine) formation C-terminal amide formation from Gly Deamidation of Asn and Gln Methylation Hydroxylation Oxidation of Met Proteolysis of a single peptide bond Formylation Acetylation Carboxylation of Asp and Glu Phosphorylation Sulfation Cysteinylation Glycosylation with pentoses (Ara, Rib, Xyl) Glycosylation with deoxyhexoses (Fuc, Rha) Glycosylation with hexosamines (GalN, GlcN) Glycosylation with hexoses (Fru, Gal, Glc, Man) Modification with lipoic acid (amide bond to lysine) Glycosylation with N-acetylhexosamines (GalNAc, GlcNAc) Farnesylation Myristoylation Biotinylation (amide bond to lysine) Modification with pyridoxal phosphate (Schiff base to lysine) Palmitoylation Stearoylation Geranylgeranlylation Glycosylation with N-acetylneuraminic acid (sialic acid, NeuAc, NANA, SA) Glutathionylation Glycosylation with N-glycolylneuraminic acid (NeuGe) 5′-Adenosylation Modification with 4′-Phosphopantetheine ADP-ribosylation (from NAD) Adventitious modifications Acrylamide Glutathione 2-Mercaptoethanol
Monoisotopic mass change
Average mass change
−17.0265 −2.0157 −0.9840 −0.9840 14.0157 15.9949 15.9949 18.0106 27.9949 42.0106 43.9898 79.9663 79.9568 119.0041 132.0423 146.0579 161.0688 162.0528 188.0330 203.0794 204.1878 210.1984 226.0776 231.0297 238.2297 266.2610 272.2504 291.0954
−17.0306 −2.0159 −0.9847 −0.9847 14.0269 15.9994 15.9994 18.0153 28.0104 42.0373 44.0098 79.9799 80.0642 119.1442 132.1161 146.1430 161.1577 162.1424 188.3147 203.1950 204.3556 210.3598 226.2994 231.1449 238.4136 266.4674 272.4741 291.2579
305.0682 307.0903 329.0525 339.0780 541.0611
305.3117 307.2573 329.2091 339.3294 541.3052
71.0371 304.0712 75.9983
71.0788 304.3038 76.1192
aTo obtain the molecular mass of a modified peptide or protein, the appropriate mass changes should be algebraically added
to the molecular mass calculated for the unmodified molecule. more extensive list of modifications is available from the Delta mass site at http://www.mbcf.dfci.harvard.edu/docs/DeltaMass.html.
bA
Useful Data
A.1A.7 Current Protocols in Protein Science
Supplement 27
Table A.1A.4
Mass Increments for Common Carbohydrate Structures
In-chain massa Sugarsb
Monoisotopic
Chemical average
1+
2+
3+
1+
2+
3+
Deoxyhexose Hexose
146.058 162.053
73.0 81.0
48.7 54.0
146.144 162.143
73.0 81.1
48.7 54.0
Hexosamine N-acetylhexosamine
161.069 203.080
80.5 101.6
53.7 67.7
161.158 203.196
80.6 101.6
53.7 67.7
N-acetylneuraminic acid N-glycolylneuraminic acid
291.095 307.090
145.6 153.6
97.0 102.4
291.259 307.257
145.6 153.6
97.1 102.4
Hex-HexNAc NeuAc-Hex-HexNAc
365.132 656.228
182.6 328.1
121.7 218.7
365.339 656.598
182.7 328.3
121.8 218.9
(Hex)3-HexNAc-HexNAc (Hex)3-HexNAc-(dHex)HexNAc
892.317 1038.375
446.2 519.2
297.4 346.1
892.821 1038.965
446.4 519.5
297.6 346.3
Biantennary (1 dHex; 0 NeuAc) Triantennary (1 dHex; 0 NeuAc)
1768.640 2133.772
884.3 1066.9
589.5 711.3
1769.643 2134.982
884.8 1067.5
589.9 711.7
Tetraantennary (1 dHex; 0 NeuAc)
2498.904
1249.5
833.0
2500.321
1250.2
833.4
aEnd-group mass (+18.011 for monoisotopic, +18.015 for chemical average) + in-chain mass = molecular mass of carbohydrate. Determined M of r peptide (attachment site = Asn) − in-chain mass of carbohydrate. bAbbreviations:
dHex, deoxyhexose; Hex, hexose; HexN, hexosamine; HexNAc, N-acetylhexosamine; NeuAc, N-acetylneuraminic acid.
Characteristics of Amino Acids
A.1A.8 Supplement 27
Current Protocols in Protein Science
Commonly Used Detergents
APPENDIX 1B
Detergents are polar lipids that are soluble in water. The presence of both a hydrophobic and hydrophilic portion makes these compounds very useful for lysis of lipid membranes, solubilization of antigens, and washing of immune complexes. TYPES OF DETERGENTS A large variety of detergents are available (Helenius et al., 1979). For biochemical studies, they are usually categorized according to the type of hydrophilic group they contain— anionic, cationic, amphoteric, or nonionic. Tables A.1B.1 and A.1B.2 list commonly used members of each type. In general, nonionic and amphoteric detergents are less denaturing for proteins than ionic detergents. Sodium cholate and sodium deoxycholate are the least denaturing of the commonly used ionic detergents. Two properties of detergents are important in their consideration for biological studies: the critical micelle concentration (CMC) and the micelle molecular weight (Table A.1B.1). The CMC is the concentration at which monomers of detergent molecules combine to form micelles; each detergent micelle has a characteristic micelle molecular weight. Detergents with a high micelle molecular weight, such as nonionic detergents, are difficult to remove from samples by dialysis. The CMC and the micelle molecular weight will vary depending on the buffer, salt concentration, pH, and temperature. In general, adding salt will lower the CMC and raise the micelle size. Table A.1B.1
Physical Properties of Commonly Used Detergentsa,b
Detergent
mp (°C)
Molecular weight (Da) Monomer
Micelle
CMC % (w/v)
M
Anionic SDS Cholate Deoxycholate
206 201 175
288 430 432
18,000 4,300 4,200
0.23 0.60 0.21
8.0 × 10−3 1.4 × 10−2 5.0 × 10−3
Cationic C16TAB
230
365
62,000
0.04
1.0 × 10−3
Amphoteric LysoPC CHAPS Zwittergent 3-14
— 157 —
495 615 364
92,000 6,150 30,000
0.0004 0.49 0.011
7.0 × 10−6 1.4 × 10−3 3.0 × 10−4
Nonionic Octyl glucoside Digitonin C12E8 Lubrol PX Triton X-100 Nonidet P-40 Tween 80
105 235 — — — — —
292 1,229 542 582 650 603 1,310
8,000 70,000 65,000 64,000 90,000 90,000 76,000
0.73 — 0.005 0.006 0.021 0.017 0.002
2.3 × 10−2 — 8.7 × 10−5 1.0 × 10−4 3.0 × 10−4 3.0 × 10−4 1.2 × 10−5
a Reprinted with permission from IRL Press (see Jones et al., 1987). bAbbreviations: C TAB, hexadecyl trimethylammonium bromide; CMC, critical micelle concentration; LysoPC, 16
lysophosphatidylcholine; mp, melting point; SDS, sodium dodecyl sulfate.
Useful Data Contributed by John E. Coligan Current Protocols in Protein Science (1998) A.1B.1-A.1B.3 Copyright © 1998 by John Wiley & Sons, Inc.
A.1B.1 Supplement 11
Table A.1B.2
Chemical Properties of Commonly Used Detergentsa,b
Ionic detergents
Nonionic detergents
SDS CHO DOC C16 LYS CHA ZWI
OGL DIG C12 LUB TNX NP-40 T80
Property Strongly denaturingc Dialyzable Ion exchangeabled Complexes ions Strong A280 Assay interference Cold precipitates High cost Availability Toxicity Ease of purification Radiolabeled Defined composition Auto-oxidation
+ + + + − − + − + − + + + −
− + + + − − − − + − + + + −
− + + + − − + − + − + + + −
+ + + − − − + − + − + − + −
+/− − − − − − − + + − +/− + + −
− + − − − − − + + − + − + −
+/− +/− − − − − − + +/− − + − + −
− + − − − − − + + − − + + −
− − − − − − − + + − + −
− − − − − − − − − − − − − − − +/− +/− +/− +/− +/− − − + + − − +/− +/− +/− +/− − − − − − + − − − − +/− + + + + − − − − − − − − − − + + + + + − − − − − + + + + +
aAdapted from IRL Press (see Jones et al., 1987). bAbbreviations: C , C E ; C , hexadecyl trimethylammonium bromide; CHA, CHAPS; CHO, cholate; DIG, digitonin; 12 12 8 16
DOC, deoxycholate; LUB, lubrol PX; LYS, lysophosphatidylcholine; NP-40, Nonidet P-40; OGL, octyl glucoside; SDS, sodium dodecyl sulfate; T80, Tween 80; TNX, Triton X-100; ZWI, Zwittergent 3-14. c Denaturing refers to disruption of secondary and tertiary protein structure. dIonic detergents are unsuitable for ion-exchange chromatography (UNIT 8.2).
CHOICE OF DETERGENTS Ionic detergents are very good solubilizing agents, but they tend to denature proteins by destroying native three-dimensional structures. This denaturing ability is useful for SDS-PAGE (UNIT 10.1), but is detrimental where native structure is important, as when functional activities must be retained (antibody activity is usually retained in <0.1% SDS). Nonionic and mildly ionic detergents are less denaturing and can often be used to solubilize membrane proteins while retaining protein-protein interactions. The following detergent properties are detrimental in certain procedures: 1. Phenol-containing detergents (e.g., Triton X-100 and NP-40) have a high absorbance at 280 nm and hence interfere with protein monitoring during chromatography (most ionic detergents do not absorb at 280 nm; Brij- and Lubrol-series detergents are nonionic detergents that do not have substantial absorbance at 280 nm). Phenol-containing detergents also induce precipitation in the Folin protein assay (but they can be used with the Bradford protein assay; UNIT 3.4). Finally, they are readily iodinated and so should not be present during radioiodination. 2. Many detergents have a very high micelle molecular weight (Table A.1B.1), which makes their use in gel filtration impossible since protein sizes are insignificant relative to the micelle size. In addition, such detergents cannot be readily removed by dialysis. 3. Sodium cholate and sodium deoxycholate are insoluble below pH 7.5 or above an ionic strength of 0.1%. SDS will often crystallize below 20°C. Detergents
4. Ionic detergents interfere with nondenaturing electrophoresis (UNIT 10.3) and isoelectric focusing (UNIT 10.2).
A.1B.2 Supplement 11
Current Protocols in Protein Science
Detergents can be removed or exchanged for other detergents by a variety of procedures (Hjelmeland, 1979; Furth et al., 1984; Harlow and Lane, 1988). Ionic and amphoteric detergents can usually be removed by dialysis UNIT 4.4. Pierce makes Extracti-Gel D for removing a variety of detergents from protein solutions (Pierce Immunotechnology Catalog and Handbook on Protein Modification). LITERATURE CITED Furth, A.J., Bolton, H., Potter, J., and Priddle, J.D. 1984. Separating detergents from proteins. Methods Enzymol. 104:318-328. Harlow, E. and Lane, D. 1988. Detergents. In Antibodies: A Laboratory Manual, pp. 687-689. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. Helenius, A., McCaslin, D.R., Fries, E., and Tanford, C. 1979. Properties of detergents. Methods Enzymol. 56:734-749. Hjelmand, L.M. 1979. Removal of detergents from membrane proteins. Methods Enzymol. 182:277-282. Jones, O.T., Earnest, J.P. and McNamee, M.G. 1987. Solubilization and reconstitution of membrane proteins. In Biological Membranes: A Practical Approach (J. Findlay and W.H. Evans, eds.), pp. 142-143. IRL Press, Oxford.
KEY REFERENCES Harlow and Lane, 1988. See above. Provides properties of commonly used detergents and means of removing them from proteins. Hjelmeland, L.M. and Chrambach, A. 1984. Solubilization of functional membrane proteins. Methods Enzymol. 182:305-318. Describes properties of detergents and how to use them to solubilize proteins. Johnstone, A. and Thorpe, R. 1982. Isolation and fractionation of lymphocytes. In Immunochemistry in Practice, pp. 94-101. Blackwell Scientific, Oxford. Provides details in use of detergents to solubilize cells and membranes. Neugebaur, J.M. 1990. Detergents: An overview. Methods Enzymol. 182:239-282. Provides details on detergent properties and how to choose one for a particular application.
Contributed by John E. Coligan National Institute of Allergy and Infectious Diseases Bethesda, Maryland
Useful Data
A.1B.3 Current Protocols in Protein Science
Supplement 11
Conversion Factors and Half-Life Information for Radioactivity
APPENDIX 1C
Discussion of radioactivity may use different terms for measurements of radioactivity and dose. Conversion factors for these terms are presented in Table A.1C.1. In addition, when investigators are working with a particular radiolabeled compound, it is necessary to calculate the amount of remaining radioactivity based on the amount of time that has elapsed from the reference date and the half life of the radioisotope (see Fig. A.1C.1). Decay factors for the commonly used shorter half life radioisotopes are presented in Table A.1C.2. For information about working with radioisotopes, see Guide for Safe Handling of Radioisotopes (APPENDIX 2B).
Table A.1C.1
Conversion Factors for Radioactivity
Measurement of Radioactivity The SI unit for measurement of radioactivity is the Becquerel: 1 Becquerel (Bq) = 1 disintegration per second The more commonly encountered unit is the Curie (Ci): 10 1 Ci = 3.7 × 10 Bq 12 = 2.22 × 10 disintegrations per minute (dpm) 7 9 1 millicurie (mCi) = 3.7 × 10 Bq = 2.22 × 10 dpm 4 6 1 microcurie (µCi) = 3.7 × 10 Bq = 2.22 × 10 dpm Conversion factors: 3 4 1 day = 1.44 × 10 min = 8.64 × 10 sec 5 7 1 year = 5.26 × 10 min = 3.16 × 10 sec counts per minute (cpm) = dpm × (counting efficiency) Measurement of Dose The SI unit for energy absorbed from radiation is the Gray (Gy): 1 Gy = 1 joule/kg Older units of absorbed energy are the rad and Roentgen (R): −2 1 rad = 100 ergs/g = 10 Gy 1 R = 0.877 r in air = 0.93 − 0.98 r in water and tissue The SI unit for radiation dosage is the Sievert (Sv), which takes into account the empirically determined relative biological effectiveness (RBE) of a given form of radiation: dosage [Sv] = RBE × dosage [Gy] RBE =
(biological effect of a dose of standard radiation [Gy]) (biological effect of a dose of other radiation [Gy])
RBE = 1 for commonly encountered radionuclides The older unit for dosage is the rem (Roentgen-equivalent-man): 1 rem = 0.01 Sv
Useful Data Current Protocols in Protein Science (1998) A.1C.1-A.1C.2 Copyright © 1998 by John Wiley & Sons, Inc.
A.1C.1 Supplement 11
Table A.1C.2 Decay factors for calculating the amount of radioactivity present at a given time after a reference date. For example, a vial containing 1.85 MBq (50 µCi) of an 35S-labeled compound on the reference date will have the following activity 33 days later: 1.85 × 0.770 = 1.42 MBq; 50 × 0.770 = 38.5 µCi.
Days 0 1.000 0.794 0.630 0.500 0.397 0.315 0.250 0.198 0.157 0.125 0.099 0.079 0.063
Days
131I
0 3 6 9 12 15 18 21 24 27 30 33 36
2 0.977 0.776 0.616 0.489 0.388 0.308 0.244 0.194 0.154 0.122 0.097 0.077 0.061
4 0.955 0.758 0.602 0.477 0.379 0.301 0.239 0.189 0.150 0.119 0.095 0.075 0.060
6 0.933 0.741 0.588 0.467 0.370 0.294 0.233 0.185 0.147 0.117 0.093 0.073 0.058
8 0.912 0.724 0.574 0.456 0.362 0.287 0.228 0.181 0.144 0.114 0.090 0.072 0.057
10 0.891 0.707 0.561 0.445 0.354 0.281 0.223 0.177 0.140 0.111 0.088 0.070 0.056
12 0.871 0.691 0.548 0.435 0.345 0.274 0.218 0.173 0.137 0.109 0.086 0.069 0.054
14 0.851 0.675 0.536 0.425 0.338 0.268 0.213 0.169 0.134 0.106 0.084 0.067 0.053
16 0.831 0.660 0.524 0.416 0.330 0.262 0.208 0.165 0.131 0.104 0.082 0.065 0.052
18 0.812 0.645 0.512 0.406 0.322 0.256 0.203 0.161 0.128 0.102 0.081 0.064 0.051
6 0.979 0.756 0.583 0.450 0.348 0.269 0.207 0.160 0.124 0.095 0.074 0.057 0.044
12 0.958 0.740 0.571 0.441 0.340 0.263 0.203 0.157 0.121 0.093 0.072 0.056 0.043
18 0.937 0.724 0.559 0.431 0.333 0.257 0.199 0.153 0.118 0.091 0.071 0.054 0.042
24 0.917 0.708 0.547 0.422 0.326 0.252 0.194 0.150 0.116 0.089 0.069 0.053 0.041
35S
30 0.898 0.693 0.533 0.413 0.319 0.246 0.190 0.147 0.113 0.088 0.068 0.052 0.040
Days
33P
Hours 0 1.000 0.824 0.679 0.559 0.460 0.379 0.312 0.257 0.212 0.175 0.144 0.119 0.098 0.080
0 4 8 12 16 20 24 28 32 36 40 44 48 52
Half-life: 8.04 days Hours 0 1.000 0.772 0.596 0.460 0.355 0.274 0.212 0.164 0.126 0.098 0.075 0.058 0.045
Half-life: 14.3 days
36 0.879 0.678 0.524 0.405 0.312 0.241 0.186 0.144 0.111 0.086 0.066 0.051 0.039
42 0.860 0.664 0.513 0.396 0.306 0.236 0.182 0.141 0.109 0.084 0.065 0.050 0.039
48 0.842 0.650 0.502 0.387 0.299 0.231 0.178 0.138 0.106 0.082 0.064 0.049 0.038
54 0.824 0.636 0.491 0.379 0.293 0.226 0.175 0.135 0.104 0.080 0.063 0.048 0.037
60 0.806 0.622 0.481 0.371 0.286 0.221 0.171 0.132 0.102 0.079 0.061 0.047 0.036
66 0.789 0.609 0.470 0.363 0.280 0.216 0.167 0.129 0.100 0.077 0.059 0.046 0.035
Weeks
0 20 40 60 80 100 120 140 160 180 200 220 240
32P
Half-life: 60.0 days
Days
Days
125I
0 1 2 3 4 5 6 7 8 9 10 11 12
12 0.976 0.804 0.662 0.546 0.449 0.370 0.305 0.251 0.207 0.170 0.140 0.116 0.095 0.078
24 0.953 0.785 0.646 0.533 0.439 0.361 0.298 0.245 0.202 0.166 0.137 0.113 0.093 0.077
36 0.930 0.766 0.631 0.520 0.428 0.353 0.291 0.239 0.197 0.162 0.134 0.110 0.091 0.075
48 0.908 0.748 0.616 0.507 0.418 0.344 0.284 0.234 0.192 0.159 0.131 0.108 0.089 0.073
60 0.886 0.730 0.601 0.495 0.408 0.336 0.277 0.228 0.188 0.155 0.127 0.105 0.086 0.071
72 0.865 0.712 0.587 0.483 0.398 0.328 0.270 0.223 0.183 0.151 0.124 0.102 0.084 0.070
84 0.844 0.695 0.573 0.472 0.389 0.320 0.264 0.217 0.179 0.147 0.121 0.100 0.082 0.068
Half-life: 87.4 days Days 0 1.000 0.946 0.895 0.847 0.801 0.758 0.717 0.678 0.641 0.607 0.574 0.543 0.514
1 0.992 0.939 0.888 0.840 0.795 0.752 0.711 0.673 0.636 0.602 0.569 0.539 0.510
2 0.984 0.931 0.881 0.833 0.788 0.746 0.705 0.667 0.631 0.597 0.565 0.534 0.506
3 0.976 0.924 0.874 0.827 0.782 0.740 0.700 0.662 0.626 0.592 0.560 0.530 0.502
4 0.969 0.916 0.867 0.820 0.776 0.734 0.694 0.657 0.621 0.588 0.556 0.526 0.498
5 0.961 0.909 0.860 0.814 0.770 0.728 0.689 0.652 0.616 0.583 0.552 0.522 0.494
6 0.954 0.902 0.853 0.807 0.764 0.722 0.683 0.646 0.612 0.579 0.547 0.518 0.490
Half-life: 25.4 days Days 0 1.000 0.761 0.579 0.441 0.336 0.256 0.195 0.148 0.113 0.086 0.065 0.050 0.038
0 10 20 30 40 50 60 70 80 90 100 110 120
1 0.973 0.741 0.564 0.429 0.327 0.249 0.189 0.144 0.110 0.084 0.064 0.048 0.037
2 0.947 0.721 0.549 0.418 0.318 0.242 0.184 0.140 0.107 0.081 0.062 0.047 0.036
3 0.921 0.701 0.534 0.406 0.309 0.236 0.179 0.136 0.104 0.079 0.060 0.046 0.035
4 0.897 0.683 0.520 0.395 0.301 0.229 0.174 0.133 0.101 0.077 0.059 0.045 0.034
5 0.872 0.664 0.506 0.385 0.293 0.223 0.170 0.129 0.098 0.075 0.057 0.043 0.033
6 0.849 0.646 0.492 0.374 0.285 0.217 0.165 0.126 0.096 0.073 0.055 0.042 0.032
7 0.826 0.629 0.479 0.364 0.277 0.211 0.161 0.122 0.093 0.071 0.054 0.041 0.031
8 0.804 0.612 0.466 0.355 0.270 0.205 0.156 0.119 0.091 0.069 0.053 0.040 0.030
9 0.782 0.595 0.453 0.345 0.263 0.200 0.152 0.116 0.088 0.067 0.051 0.039 0.030
Number of half-lives elapsed 6
5
2
1
3
3
4
4
5
6
7
8 9 10
1
2
15
20
25
30
40
50
60 70 80 90 100
Percent of radioactivity remaining
Figure A.1C.1 Correlation of loss of radioactivity with elapsing half-lives of an isotope.
Radioactivity
A.1C.2 Supplement 11
Current Protocols in Protein Science
Common Conversion Factors
APPENDIX 1D
Table A.1D.1 lists some of the more common conversion factors for units of measure used throughout Current Protocols manuals, while Table A.1D.2 gives prefixes indicating powers of ten for SI units. Table A.1D.3 gives conversions between temperatures on the Celsius (Centigrade) and Fahrenheit scales. Celsius temperatures are converted to Fahrenheit temperatures by multiplying the Celsius figure by 9, dividing by 5, and adding 32, or by multiplying the Celsius figure by 1.8 and adding 32. Fahrenheit is converted to Celsius by subtracting 32 from the Fahrenheit figure, multiplying by 5, and dividing by 9. In Table A.1D.3, the center figure represents the temperature one has read on one of the scales; the figure to the left is the conversion of that figure into Celsius if read in Fahrenheit, while that to the right represents the conversion to Fahrenheit if read in Celsius: e.g., the temperature 88 Fahrenheit converts to 31.1°C, while the temperature 88 Celsius converts to 190.4°F.
Table A.1D.1
Unit of Measurement Conversion Chart
To convert:
Into:
Use the multiplier:
amperes per square centimeter (amp/cm2)
amperes per square inch (amp/in.2) amperes per square meter (amp/m2)
6.452 104
amperes per square inch (amp/in.2)
amperes per square centimeter (amp/cm2) amperes per square meter (amp/m2)
0.1550 1.55 × 103
ampere-hours (amp-hr)
coulombs (C) faradays
3.6 × 103 3.731 × 10−2
atmospheres (atm)
bar millimeters of mercury (mmHg) or torr tons per square foot (tons/ft2)
1.01325 760 1.058
bar
atmospheres (atm) dynes per square centimeter (dyn/cm2) kilograms per square meter (kg/m2) pounds per square foot (lb/ft2) pounds per square inch (lb/in.2 or psi)
0.9869 106 1.020 × 104 2,089 14.50
British thermal units (Btu)
ergs gram-calories (g-cal) horsepower-hours (hp-hr) joules (J) kilogram-calories (kg-cal) kilogram-meters (kg-m) kilowatt-hours (kW-hr)
1.0550 × 1010 252.0 3.931 × 104 1,054.8 0.2520 107.5 2.928 × 10−4
British thermal unit per minute (Btu/min)
foot-pounds per second (ft-lb/sec) horsepower (hp) watts (W)
12.96 2.356 × 10−2 17.57
bushels
cubic feet (ft3) cubic inches (in.3) cubic meters (m3) liters quarts, dry
1.2445 2,150.4 3.524 × 10−2 35.24 32.0 continued
Current Protocols in Protein Science (1999) A.1D.1-A.1D.8 Copyright © 1999 by John Wiley & Sons, Inc.
A.1D.1 Supplement 17
Table A.1D.1
Unit of Measurement Conversion Chart, continued
To convert:
Into:
Use the multiplier:
degrees Celsius or Centigrade (°C)
degrees Fahrenheit (°F) Kelvin (K)
(°C × 9⁄5) + 32 (°C) + 273.15
degree Fahrenheit (°F)
degrees Celsius (°C) Kelvin (K)
5⁄
centimeters (cm)
feet (ft) inches (in.) kilometers (km) meters (m) miles millimeters (mm) mils yards
3.281 × 10−2 0.3937 10−5 10−2 6.214 × 10−6 10.0 393.7 1.094 × 10−2
centimeters per second (cm/sec)
feet per minute (ft/min) feet per second (ft/sec) kilometers per hour (km/hr) meters per minute (m/min) miles per hour (miles/hr) miles per minute (miles/min)
1.1969 3.281 × 10−2 3.6 × 10−2 0.6 2.237 × 10−2 3.728 × 10−4
coulombs (C)
faradays
1.036 × 10−5
coulombs per square centimeter (C/cm2)
coulombs per square inch (C/in.2) coulombs per square meter (C/m2)
64.52 104
coulombs per square inch (C/in.2)
coulombs per square centimeter (C/cm2) coulombs per square meter (C/m2)
0.1550 1.55 × × 103
cubic centimeters (cm3)
cubic feet (ft3) cubic inches (in.3)
3.531 × 10−5 6.102 × 10−2
cubic meters (m3) cubic yards gallons, U.S. liquid liters pints, U.S. liquid quarts, U.S. liquid
10−6 1.308 × 10−6 2.642 × 10−4 10−3 2.113 ×10−3 1.057 × 10−3
days
hours (hr) minutes (min) seconds (sec)
24.0 1.44 × 103 8.64 × 104
degrees (of angle; °)
minutes (min) quadrants, of angle radians (rad) seconds (sec)
60.0 1.111 × 10−2 1.745 × 10−2 3.6 × 104
drams
grams (g) grains ounces, avoirdupois (oz)
1.7718 27.3437 6.25 × 10−2
dynes (dyn)
joules per centimeter (J/cm) joules per meter (J/m) or newtons (N) kilograms (kg) pounds (lb)
10−7 10−5 1.020 × 10−6 2.248 × 10−6
9 × (°F − 32) [5⁄9 × (°F − 32)] + 273.15
continued
A.1D.2 Supplement 17
Current Protocols in Protein Science
Table A.1D.1
Unit of Measurement Conversion Chart, continued
To convert:
Into:
Use the multiplier:
faradays
ampere-hours (amp-hr) coulombs (C)
26.80 9.649 × 10−4
foot-pounds per minute (ft-lb/min)
British thermal units per minute (Btu/min) foot-pounds per second (ft-lb/sec) horsepower (hp) kilogram-calories per minute (kg-cal/min) kilowatts (kW)
1.286 × 10−3 1.667 × 10−2 3.030 × 10−5 3.24 × 10−4 2.260 × 10−5
grams (g)
decigrams (dg) dekagrams (Dg) dynes (dyn) grains hectograms (hg) kilograms (kg) micrograms (µg) milligrams (mg) ounces, avoirdupois (oz) ounces, troy pounds (lb)
10 0.1 980.7 15.43 10−2 10−3 106 103 3.527 × 10−2 3.215 × 10−2 2.205 × 10−3
horsepower (hp)
horsepower, metric
1.014
inches (in.)
centimeters (cm) feet (ft) meters (m) miles millimeters (mm) yards
2.540 8.333 × 10−2 2.540 × 10−2 1.578 × 10−5 25.40 2.778 × 10−2
inches of mercury (in. Hg)
atmospheres (atm) kilogram per square centimeter (kg/cm2) kilograms per square meter (kg/m2) pounds per square foot (lb/ft2) pounds per square inch (lb/in.2 or psi)
3.342 × 10−2 3.453 × 10−2 345.3 70.73 0.4912
joules (J)
British thermal units (Btu) ergs foot-pounds (ft-lb) kilogram-calories (kg-cal) kilogram-meters (kg-m) newton-meter (N-m) watt-hours (W-hr)
9.480 × 10−4 107 0.7376 2.389 × 10−4 0.1020 1 2.778 × 10−4
Kelvin (K)
degrees Celsius (°C) degrees Fahrenheit (°F)
K − 273.13 [(K − 273.13) × 9⁄5] + 32
kilolines
maxwells (Mx)
103
kilometers (km)
centimeters (cm) feet (ft) inches (in.) meters (m) miles yards
105 3,281 3.937 × 104 103 0.6214 1,094 continued
A.1D.3 Current Protocols in Protein Science
Supplement 22
Table A.1D.1
Unit of Measurement Conversion Chart, continued
To convert:
Into:
Use the multiplier:
kilowatts (kW)
British thermal units per minute (Btu/min) foot-pounds per minute (ft-lb/min) horsepower (hp) kilogram-calories per minute (kg-cal/min)
56.92 4.426 × 104 1.341 14.34
liters
bushels, U.S. dry cubic centimeters (cm3) cubic feet (ft3) cubic inches (in.3) cubic meters (m3) cubic yards gallons, U.S. liquid gallons, imperial kiloliter (kl) pints, U.S. liquid quarts, U.S. liquid
2.838 × 10−2 103 3.531 × 10−2 61.02 10−3 1.308 × 10−3 0.2642 0.21997 10−3 2.113 1.057
maxwells (Mx)
webers (W)
10−8
micrograms (µg)
grams (g)
10−6
microliters (µl)
liters
10−6
milligrams (mg)
grams (g)
10−3
milligrams per liter (mg/liter)
parts per million (ppm)
1.0
millihenries (mH)
henries (H)
10−3
milliliters (ml)
liters
10−3
millimeters (mm)
centimeters (cm) feet (ft) inches (in.) kilometers (km) meters (m) miles
0.1 3.281 × 10−3 3.937 × 10−2 10−6 10−3 6.214 × 10−7
millimeters of mercury (mmHg) or torr
atmospheres (atm) kilograms per square meter (kg/m2) pounds per square foot (lb/ft2) pounds per square inch (lb/in.2 or psi)
1.316 × 10−3 136.0 27.85 0.1934
nepers (Np)
decibels (dB)
8.686
newtons (N)
dynes (dyn) kilograms force (kg) pounds, force (lb)
105 0.10197162 4.6246 × 10−2
ohms (Ω)
megaohms (MΩ) microhms (µΩ)
106 10−6
ounces, avoirdupois
drams grains grams (g) pounds (lb) ounces, troy tons, metric
16.0 437.5 28.349527 6.25 × 10−2 0.9115 2.835 × 10−5 continued
A.1D.4 Supplement 22
Current Protocols in Protein Science
Table A.1D.1
Unit of Measurement Conversion Chart, continued
To convert:
Into:
Use the multiplier:
ounces, fluid
cubic inches (in3) liters
1.805 2.957 × 10−2
ounces, troy
grains grams (g) ounces, avoirdupois (oz) pounds, troy
480.0 31.103481 1.09714 8.333 × 10−2
pascal (P)
newton per square meter (N/m2)
1
newtons (N)
21.6237
atmospheres (atm) inches of mercury (in. Hg) kilograms per square meter (kg/m2) pounds per square inch (lb/in2 or psi)
4.725 × 10−4 1.414 × 10−2 4.882 6.944 × 10−3
pounds per square inch (lb/in.2 or psi)
atmospheres (atm) inches of mercury (in. Hg) kilograms per square meter (kg/m2) pounds per square foot (lb/ft2) bar
6.804 × 10−2 2.036 703.1 144.0 6.8966 × 10−2
quadrants, of angle
degrees (°) minutes (min) radians (rad) seconds (sec)
90.0 5.4 × 103 1.571 3.24 × 105
quarts, dry
cubic inches (in.3)
pounds, force (lb) pounds per square foot
(lb/ft2)
67.20 (cm3)
quarts, liquid
cubic centimeters cubic feet (ft3) cubic inches (in.3) cubic meters (m3) cubic yards gallons liters
radians (rad)
degrees (°) minutes (min) quadrants seconds (sec)
57.30 3,438 0.6366 2.063 × 105
watts (W)
British thermal units per hour (Btu/hr) British thermal units per min (Btu/min) ergs per second (ergs/sec)
3.413 5.688 × 10−2 107
webers (Wb)
maxwells (M) kilolines
108 105
946.4 3.342 × 10−2 57.75 9.464 × 10−4 1.238 × 10−3 0.25 0.9463
torr see millimeter of mercury
Useful Data
A.1D.5 Current Protocols in Protein Science
Supplement 22
Table A.1D.2 Power of Ten Prefixes for SI Units
Prefix
Factor
Abbreviation
atto femto pico nano micro milli centi deci deca hecto kilo myria mega giga tera peta exa
10−18 10−15 10−12 10−9 10−6 10−3 10−2 10−1 101 102 103 104 106 109 1012 1015 1018
a f p n µ m c d da h k my M G T P E
Common Conversion Factors
A.1D.6 Supplement 22
Current Protocols in Protein Science
Table A.1D.3 Celsius/Fahrenheit Temperature Conversion Chart
Degrees Celsius (°C)
Temperature
Degrees Fahrenheit (°F)
−17.8 −17.2 −16.7 −16.1 −15.6 −15.0 −14.4 −13.9 −13.3 −12.8 −12.2 −11.7 −11.1 −10.6 −10.0 −9.4 −8.9 −8.3 −7.8 −7.2 −6.7 −6.1 −5.6 −5.0 −4.4 −3.9 −3.3 −2.8 −2.2 −1.7 −1.1 −0.6 0.0 0.6 1.1 1.7 2.2 2.8 3.3 3.9 4.4 5.0 5.6 6.1 6.7 7.2 7.8 8.3 8.9 9.4 10.0 10.6
0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51
32.0 33.8 35.6 37.4 39.2 41.0 42.8 44.6 46.4 48.2 50.0 51.8 53.6 55.4 57.2 59.0 60.8 62.6 64.4 66.2 68.0 69.8 71.6 73.4 75.2 77.0 78.8 80.6 82.4 84.2 86.0 87.8 89.6 91.4 93.2 95.0 96.8 98.6 100.4 102.2 104.0 105.8 107.6 109.4 111.2 113.0 114.8 116.6 118.4 120.2 122.0 123.8 continued Useful Data
A.1D.7 Current Protocols in Protein Science
Supplement 17
Table A.1D.3 Celsius/Fahrenheit Temperature Conversion Chart, continued
Degrees Celsius (°C)
Temperature
Degrees Fahrenheit (°F)
11.1 11.7 12.2 12.8 13.3 13.9 14.4 15.0 15.6 16.1 16.7 17.2 17.8 18.3 18.9 19.4 20.0 20.6 21.1 21.7 22.2 22.8 23.3 23.9 24.4 25.0 25.6 26.1 26.7 27.2 27.8 28.3 28.9 29.4 30.0 30.6 31.1 31.7 32.2 32.8 33.3 33.9 34.4 35.0 35.6 36.1 36.7 37.2 37.8
52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86 87 88 89 90 91 92 93 94 95 96 97 98 99 100
125.6 127.4 129.2 131.0 132.8 134.6 136.4 138.2 140.0 141.8 143.6 145.4 147.2 149.0 150.8 152.6 154.4 156.2 158.0 159.8 161.6 163.4 165.2 167.0 168.8 170.6 172.4 174.2 176.0 177.8 179.6 181.4 183.2 185.0 186.8 188.6 190.4 192.2 194.0 195.8 197.6 199.4 201.2 203.0 204.8 206.8 208.4 210.2 212.0
Common Conversion Factors
A.1D.8 Supplement 17
Current Protocols in Protein Science
LABORATORY GUIDELINES, EQUIPMENT, AND STOCK SOLUTIONS
APPENDIX 2
Laboratory Safety
APPENDIX 2A
Persons carrying out the protocols in this manual may encounter the following hazardous or potentially hazardous materials (1) pathogenic and infectious biological agents and (2) toxic chemicals and carcinogenic, mutagenic, or teratogenic reagents (Table A.2A.1). Most governments regulate the use of these materials; it is essential that they be used in strict accordance with local and national regulations. Cautionary notes are included in many instances throughout the manual, and some specific guidelines are provided below (and in references therein). However, we emphasize that users must proceed with the prudence and precautions associated with good laboratory practice, under the supervision of personnel responsible for implementing laboratory safety programs at their institutions. Guidelines for the safe use of radioisotopes are presented in APPENDIX 2B. SAFE USE OF BIOHAZARDS AND INFECTIOUS BIOLOGICAL AGENTS Precautions described in this section should be applied to the routine handling of viable pathogenic microorganisms, as well as all human-derived materials, because they may harbor dangerous pathogens such as human immunodeficiency virus (HIV), hepatitis B virus (HBV), cytomegalovirus (CMV), Epstein-Barr virus (EBV), and a host of bacterial pathogens. In addition to the guidelines provided herein, experimenters can find a wealth of information about handling infectious agents in the government publications Biosafety in Microbiological and Biomedical Laboratories and Working Safely with HIV in the Research Laboratory (see Literature Cited). Routine Precautions When Working with Biohazards The following practices are recommended for all laboratories handling potentially dangerous microorganisms, whether pathogenic or not: 1. Decontaminate all work surfaces after each working day using an appropriate disinfectant. Decontaminate all spills of viable material. See discussion under Disinfectants for Biohazards. 2. Decontaminate all liquid or solid wastes that have come in contact with viable material. 3. Do not pipet by mouth. 4. Do not allow eating, drinking, smoking, or application of cosmetics in the work area. Do not store food in refrigerators that contain laboratory supplies. 5. Wash hands with disinfectant soap or detergent after handling viable materials and before leaving the lab. Do not handle telephones, doorknobs, or other common utensils without disinfecting hands. 6. When handling viable materials, minimize creation of aerosols. 7. Wear lab coats (preferably disposable) when in work area, but do not wear them away from the work area. 8. Wear disposable latex gloves when handling viable materials. These should be disposed of as biohazardous waste. Change gloves if they are directly contaminated.
Laboratory Guidelines, Equipment, and Stock Solutions
Contributed by George Lunn and Gretchen Lawler
A.2A.1
Current Protocols in Protein Science (2002) A.2A.1-A.2A.35 Copyright © 2002 by John Wiley & Sons, Inc.
Supplement 28
Table A.2A.1
Commonly Used Hazardous Chemicalsa
Chemical
Hazards
Acetic acid, glacial Acetonitrile
Corrosive, flammable liquid Flammable liquid, teratogenic, toxic Carcinogenic, mutagenic Carcinogenic, toxic
Acridine orange Acrylamide
Alcian blue 8GX Alizarin red S (monohydrate) p-Amidinophenylmethanesulfonyl fluoride (APMSF) 7-Aminoactinomycin D (7-AAD) 4-(2-Aminoethyl)benzenesulfonyl fluoride (AEBSF) Ammonium hydroxide, concentrated Azure A Azure B Benzidine (BDB) Bisacrylamide Boron dipyrromethane derivatives (BODIPY dyes) Brilliant blue R 5-Bromodeoxyuridine (BrdU) Cetylpyridinium chloride (CPC) Cetyltrimethylammonium bromide (CTAB) Chloroform Chlorotrimethylsilane Chromic/sulfuric acid cleaning solution
Remarksb
See Basic Protocol 2 Use dust mask; polyacrylamide gels contain residual acrylamide monomer and should be handled with gloves; acrylamide may polymerize with violence on melting at 86°C See Basic Protocol 2
Enzyme inhibitor
See Basic Protocol 11
Carcinogenic Mutagenic, enzyme inhibitor
See Basic Protocol 11
Corrosive, lachrymatory, toxic Mutagenic Mutagenic Carcinogenic, toxic Toxic Toxic Carcinogenic, mutagenic Mutagenic, teratogenic, photosensitizing Toxic Corrosive, teratogenic, toxic Carcinogenic, teratogenic, toxic Carcinogenic, corrosive, flammable liquid, toxic Carcinogenic, corrosive, oxidizer, toxic
Chromomycin A3 (CA3) Congo red Coomassie brilliant blue G Crystal violet Cresyl violet acetate Cyanides (e.g., KCN, NaCN)
Teratogenic, toxic Mutagenic, teratogenic Mutagenic
Cyanines (e.g., Cy3, Cy5) Cyanogen bromide (CNBr)
Toxic Toxic
Mutagenic Toxic
See Basic Protocol 2 See Basic Protocol 2 See Basic Protocol 1
See Basic Protocol 2
Reacts violently with water; see Basic Protocol 3 Replace with suitable commercially available cleanser See Basic Protocol 2 See Basic Protocol 2 See Basic Protocol 2 See Basic Protocol 2 Contact with acid will liberate HCN gas; see Basic Protocol 4 See Basic Protocol 4 continued
A.2A.2 Supplement 28
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Table A.2A.1
Commonly Used Hazardous Chemicalsa, continued
Chemical
Hazards
2′-Deoxycoformycin (dCF, pentostatin) 4′,6-Diamidino-2-phenylindole (DAPI) Diaminobenzidine (DAB) 1,4-Diazabicyclo[2,2,2]-octane (DABCO)
Teratogenic, toxic Mutagenic Carcinogenic Toxic
Dichloroacetic acid (DCA) Dichloromethane (methylene chloride)
Carcinogenic, corrosive, toxic Carcinogenic, mutagenic, teratogenic, toxic Corrosive, flammable liquid, toxic Carcinogenic, toxic Carcinogenic, teratogenic, toxic Highly toxic, cholinesterase inhibitor, neurotoxin Corrosive, flammable liquid, toxic Carcinogenic, toxic Flammable liquid, toxic
Diethylamine (DEA) Diethylpyrocarbonate (DEPC) Diethyl sulfate Diisopropyl fluorophosphate (DFP) Dimethyldichlorosilane Dimethyl sulfate (DMS) Dimethyl sulfoxide (DMSO) Diphenylamine (DPA) 2,5-Diphenyloxazole (PPO) Dithiothreitol (DTT) Eosin B Erythrosin B Ether
Ethidium bromide (EB) Ethyl methanesulfonate (EMS) Fluorescein and derivatives 5-Fluoro-2′-deoxyuridine (FUdR) Fluoroorotic acid (FOA) Formaldehyde
Remarksb
See Basic Protocol 1 Forms an explosive complex with hydrogen peroxide
See Basic Protocol 5 See Basic Protocol 11 See Basic Protocol 3 See Basic Protocol 5 Enhances absorption through skin
Teratogenic, toxic Toxic Toxic Carcinogenic, mutagenic Flammable liquid, toxic
Formamide Formic acid
Mutagenic, toxic Carcinogenic, toxic Carcinogenic, toxic Teratogenic, toxic Toxic Carcinogenic, flammable liquid, teratogenic, toxic Teratogenic, toxic Corrosive, toxic
Glutaraldehyde Guanidinium thiocyanate Hoechst 33258 dye Hydrochloric acid, concentrated
Corrosive, teratogenic, toxic Toxic Mutagenic, toxic Corrosive, teratogenic, toxic
See Basic Protocol 2 See Basic Protocol 2 May form explosive peroxides on standing; do not dry with NaOH or KOH See Basic Protocol 2 or 6 See Basic Protocol 5
May explode when heated >180°C in a sealed tube
continued
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Table A.2A.1
Commonly Used Hazardous Chemicalsa, continued
Chemical
Hazards
Remarksb
Hydrogen peroxide (30%)
Carcinogenic, corrosive, mutagenic, oxidizer
Hydroxylamine
Corrosive, flammable, mutagenic, toxic Carcinogenic Corrosive, toxic Carcinogenic, mutagenic, toxic Carcinogenic, mutagenic Carcinogenic, toxic Stench, toxic Teratogenic, toxic Teratogenic, toxic Carcinogenic, mutagenic, teratogenic, toxic Mutagenic, toxic Carcinogenic, toxic Teratogenic, toxic Mutagenic
Avoid bringing into contact with organic materials, which may form explosive peroxides; may decompose violently in contact with metals, salts, or oxidizable materials; see Basic Protocol 7 Explodes in air at >70°C
3-β-Indoleacrylic acid (IAA) Iodine Iodoacetamide Janus green B Lead compounds 2-Mercaptoethanol (2-ME) Mercury compounds Methionine sulfoximine (MSX) Methotrexate (amethopterin) Methylene blue Methyl methanesulfonate (MMS) Mycophenolic acid (MPA) Neutral red Nigrosin, water soluble Nitric acid, concentrated Nitroblue tetrazolium (NBT) Orcein, synthetic Oxonols Paraformaldehyde Phenol
See Basic Protocol 8 See Basic Protocol 2
See Basic Protocol 9
See Basic Protocol 2 See Basic Protocol 5 See Basic Protocol 2 See Basic Protocol 2
Corrosive, oxidizer, teratogenic, toxic Toxic See Basic Protocol 2
Phenylmethylsulfonyl fluoride (PMSF) Phorbol 12-myristate 13-acetate (PMA) Phycoerythrins (PE) Piperidine Potassium hydroxide, concentrated
Toxic Toxic Carcinogenic, corrosive, teratogenic, toxic Enzyme inhibitor Carcinogenic, toxic Toxic Flammable liquid, teratogenic, toxic Corrosive, toxic
Propane sultone Propidium iodide (PI) Pyridine Rhodamine and derivatives Rose Bengal Safranine O
Carcinogenic, toxic Mutagenic Flammable liquid, toxic Toxic Carcinogenic, teratogenic Mutagenic
Readily absorbed through the skin See Basic Protocol 11
Produces a highly exothermic reaction when solid is added to water See Basic Protocol 5 See Basic Protocol 2 or 6
See Basic Protocol 2 See Basic Protocol 2 continued
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Table A.2A.1
Commonly Used Hazardous Chemicalsa, continued
Chemical
Hazards
Remarksb
Sodium azide
Carcinogenic, toxic
Adding acid liberates explosive volatile, toxic hydrazoic acid; can form explosive heavy metal azides, e.g., with plumbing fixtures—do not discharge down drain; see Basic Protocol 10
Sodium deoxycholate (Na-DOC) Sodium dodecyl sulfate (sodium lauryl sulfate, SDS) Sodium hydroxide, concentrated
Carcinogenic, teratogenic, toxic Sensitizing, toxic
Sodium nitrite Sulfuric acid, concentrated
SYTO dyes Tetramethylammonium chloride (TMAC) N,N,N′,N′-Tetramethyl-ethylenediamine (TEMED) Texas Red (sulforhodamine 101, acid chloride) Toluene Toluidine blue O Nα-p-Tosyl-L-lysine chloromethyl ketone (TLCK) N-p-Tosyl-L-phenylalanine chloromethyl ketone (TPCK) Trichloroacetic acid (TCA) Triethanolamine acetate (TEA) Trifluoroacetic acid (TFA) Trimethyl phosphate (TMP) Trypan blue Xylenes
Corrosive, toxic
A highly exothermic reaction ensues when the solid is added to water
Carcinogenic Corrosive, oxidizer, teratogenic, toxic Reaction with water is very exothermic; always add concentrated sulfuric acid to water, never water to acid Toxic Toxic Corrosive, flammable liquid, toxic Toxic Flammable liquid, teratogenic, toxic Mutagenic, toxic Toxic, enzyme inhibitor
See Basic Protocol 2 See Basic Protocol 11
Toxic, mutagenic, enzyme inhibitor
See Basic Protocol 11
Carcinogenic, corrosive, teratogenic, toxic Carcinogenic, toxic Corrosive, toxic Carcinogenic, mutagenic, teratogenic May explode on distillation Carcinogenic, mutagenic, teratogenic See Basic Protocol 2 Flammable liquid, teratogenic, toxic
aFor extensive information on the hazards of these and other chemicals as well as cautionary details, see Bretherick (1986), O’Neil (2001), Furr (2000),
Lewis (1999), Lunn and Sansone (1994a), and Bretherick et al. (1999). bCAUTION: These chemicals should be handled only in a chemical fume hood by knowledgeable workers equipped with eye protection, lab coat, and gloves. The laboratory should be equipped with a safety shower and eye wash. Additional protective equipment may be required.
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9. Control pest populations. Windows in the lab that can be opened must be equipped with screens to exclude insects. 10. Use furniture that is easy to clean—i.e., with smooth, waterproof surfaces and as few seams as possible. 11. Keep biohazard waste in covered containers free from leaks. Use orange bags or red bags as required by institutional procedure (see discussion under Disposal of Biohazards). Autoclave and dump hazardous waste without undue delay. Disinfectants for Biohazards Major laboratory suppliers sell disinfectants based on quaternary ammonium compounds that are acceptable for routine biohazard decontamination (see SUPPLIERS APPENDIX). These include Roccal (Baxter), Vesphene II (Fisher), and industrial disinfectants such as concentrated Lysol. 10% chlorine bleach may also be used for decontamination. An antimicrobial liquid soap (e.g., Vionex; Fisher) should be provided in a dispenser near the sink so that no one need handle the outside of the container to use it. Disposal of Biohazards Most institutions have defined procedures for disposal of biohazardous waste, but the following are common to all of these systems: 1. All contaminated material should be placed in autoclavable bags, which should be contained in a plastic trash pail or wire frame. If large numbers of disposable pipets or other pointed instruments are being used, it may be necessary to double-bag the material. Autoclavable biohazard bags are sold by all major laboratory supply houses. In some institutions it is necessary to color code the biohazard waste (e.g., orange bags for less dangerous waste and red ones for suspected HIV-containing material). All of these bags are marked with the universal biohazard symbol. 2. At time of disposal, the bags are loosely closed (not completely sealed) with temperature-sensitive autoclave tape (also widely available from supply houses), placed in an autoclavable basin, and sterilized at 121°C. When the tape indicates that sterilization temperature has been achieved, it is then possible to dispose of the waste by ordinary means. 3. At many institutions, contaminated hypodermic needles, scalpels, broken glass, and other sharp objects must be disposed of separately. These must be placed in appropriate “sharps” containers (e.g., Baxter), which may be autoclaved when full. SAFE USE OF HAZARDOUS CHEMICALS It is not possible in the space available to list all the precautions required for handling hazardous chemicals. Many texts have been written about laboratory safety (see Literature Cited and Key References). Obviously, all national and local laws should be obeyed, as well as all institutional regulations. Controlled substances are regulated by the Drug Enforcement Administration (http://www.doj.gov/dea). By law, Material Safety Data Sheets (MSDSs) must be readily available. All laboratories should have a Chemical Hygiene Plan (29CFR Part 1910.1450); institutional safety officers should be consulted as to its implementation. Help is (or should be) available from your institutional Safety Office; use it.
Laboratory Safety
Chemicals must be stored properly for safety. Certain chemicals cannot be easily or safely mixed with and should not be stored near certain other chemicals, because their reaction is violently exothermic or yields a toxic product. Some examples of incompatibility are
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Table A.2A.2
Examples of Chemical Incompatibility
Chemical
Incompatible with
Acetic acid
Aldehydes, bases, carbonates, chromic acid, ethylene glycol, hydroxides, hydroxyl compounds, metals, nitric acid, oxidizers, perchloric acid, peroxides, phosphates, permanganates, xylene Acids, amines, concentrated nitric and sulfuric acid mixtures, oxidizers, plastics Copper, halogens, mercury, oxidizers, potassium, silver Acids, aldehydes, carbon dioxide, carbon tetrachloride or other chlorinated hydrocarbons, halogens, ketones, plastics, sulfur, water Acids, aldehydes, amides, calcium hypochlorite, hydrofluoric acid, halogens, heavy metals, mercury, oxidizers, plastics, sulfur Acids, alkalis, chlorates, chloride salts, flammable and combustible materials, metals, organic materials, phosphorus, reducing agents, sulfur, urea Acids, aluminum, dibenzoyl peroxide, oxidizers, plastics Any reducing agent Acids, heavy metals, oxidizers Acetaldehyde, alcohols, alkalis, amines, ammonia, combustible materials, ethylene, fluorine, hydrogen, ketones (e.g., acetone, carbonyls), metals, petroleum gases, sodium carbide, sulfur Acids, ethanol, fluorine, organic materials, water Alkali metals, calcium hypochlorite, halogens, oxidizers Sodium Acids, ammonium salts, finely divided organic or combustible materials, powdered metals, sulfur Acetylene or other hydrocarbons, alcohols, ammonia, benzene, butadiene, butane, combustible materials, ethylene, flammable compounds (e.g., hydrazine), hydrogen, hydrogen peroxide, iodine, metals, methane, nitrogen, oxygen, propane (or other petroleum gases), sodium carbide, sodium hydroxide Ammonia, hydrogen, hydrogen sulfide, mercury, methane, organic materials, phosphine, phosphorus, potassium hydroxide, sulfur Acetic acid, acetone, alcohols, alkalis, ammonia, bases, benzene, camphor, flammable liquids, glycerin (glycerol), hydrocarbons, metals, naphthalene, organic materials, phosphorus, plastics Acetylene, calcium, hydrocarbons, hydrogen peroxide, oxidizers Acids (organic or inorganic) Acids, alkaloids, aluminum, iodine, oxidizers, strong bases Ammonium nitrate, chromic acid, halogens, hydrogen peroxide, nitric acid, oxidizing agents in general, oxygen, sodium peroxide All other chemicals
Acetone Acetylene Alkali metals, alkaline earth metals Ammonia (anhydrous)
Ammonium nitrate
Aniline Arsenical materials Azides Bromine
Calcium oxide Carbon (activated) Carbon tetrachloride Chlorates Chlorine
Chlorine dioxide
Chromic acid, chromic oxide
Copper Cumene hydroperoxide Cyanides Flammable liquids
Fluorine
continued
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Table A.2A.2
Examples of Chemical Incompatibility, continued
Chemical
Incompatible with
Hydrocarbons (liquid or gas) Hydrocyanic acid Hydrofluoric acid
See flammable liquids
Hydrogen peroxide Hydrogen sulfide Hydroperoxide Hypochlorites Iodine Mercury Nitric acid Nitrites Nitroparaffins Oxalic acid Oxygen
Perchloric acid Peroxides, organic Phosphorus (white) Potassium chlorate
Potassium perchlorate Potassium permanganate Selenides and tellurides Silver Sodium Sodium nitrate Sodium peroxide
Sulfides Sulfuric acid
Alkali, nitric acid Ammonia, metals, organic materials, plastics, silica (glass, including fiberglass), sodium All organics, most metals or their salts, nitric acid, phosphorus, sodium, sulfuric acid Acetylaldehyde, fuming nitric acid, metals, oxidizers, sodium, strong bases Reducing agents Acids, activated carbon Acetylaldehyde, acetylene, ammonia, hydrogen, metals, sodium Acetylene, aluminum, amines, ammonia, calcium, fulminic acid, lithium, oxidizers, sodium Acids, nitrites, metals, most organics, plastics, sodium, sulfur, sulfuric acid Acids Amines, inorganic bases Mercury, oxidizers, silver, sodium chlorite All flammable and combustible materials, ammonia, carbon monoxide, grease, metals, oil, phosphorus, polymers All organics, bismuth and alloys, dehydrating agents, grease, hydrogen halides, iodides, paper, wood Acids (organic or mineral), avoid friction, store cold Air, alkalis, oxygen, reducing agents Acids, ammonia, combustible materials, fluorine, hydrocarbons, metals, organic materials, reducing agents, sugars Alcohols, combustible materials, fluorine, hydrazine, metals, organic matter, reducing agents, sulfuric acid Benzaldehyde, ethylene glycol, glycerin, sulfuric acid Reducing agents Acetylene, ammonium compounds, fulminic acid, oxalic acid, ozonides, peroxyformic acid, tartaric acid Acids, carbon dioxide, carbon tetrachloride, hydrazine, metals, oxidizers, water Acetic anhydride, acids, metals, organic matter, peroxyformic acid, reducing agents Acetic anhydride, benzaldehyde, benzene, carbon disulfide, ethyl acetate, ethyl or methyl alcohol, ethylene glycol, furfural, glacial acetic acid, glycerin, hydrogen sulfide, metals, methyl acetate, oxidizers, peroxyformic acid, phosphorus, reducing agents, sugars, water Acids Alcohols, bases, chlorates, perchlorates, permanganates of potassium, lithium, sodium, magnesium, calcium
Laboratory Safety
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listed in Table A.2A.2. When in doubt, always consult a current MSDS for information on reactivity, handling, and storage. Chemicals should be separated into general hazard classes and stored appropriately. For example, flammable chemicals such as alcohols, ketones, aliphatic and aromatic hydrocarbons, and other materials labeled flammable should be stored in approved flammable storage cabinets, with those also requiring refrigeration being kept in explosion-proof refrigerators. Strong oxidizers must be segregated. Strong acids (e.g., sulfuric, hydrochloric, nitric, perchloric, and hydrofluoric) should be stored in a separate cabinet well removed from strong bases and from flammable organics. An exception is glacial acetic acid, which is both corrosive and flammable, and which must be stored with the flammables. Facilities should be appropriate for working with hazardous chemicals. In particular, hazardous chemicals should be handled only in chemical fume hoods, not in laminar flow cabinets. The functioning of the fume hoods should be checked periodically. Laboratories should also be equipped with safety showers and eye-wash facilities. Again, this equipment should be tested periodically to ensure that it functions correctly. Other safety equipment may be required depending on the nature of the materials being handled. In addition, researchers should be trained in the proper procedures for handling hazardous chemicals as well as other laboratory operations—e.g., handling of compressed gases, use of cryogenic liquids, operation of high-voltage power supplies, and operation of lasers of all types. Before starting work, know the physical and chemical hazards of the reagents used. Wear appropriate protective clothing and have a plan for dealing with spills or accidents; coming up with a good plan on the spur of the moment is very difficult. For example, have the appropriate decontaminating or neutralizing agents prepared and close at hand. Small spills can probably be cleaned up by the researcher. In the case of larger spills, the area should be evacuated and help should be sought from those experienced in and equipped for dealing with spills—e.g., the institutional Safety Office. Protective equipment should include, at a minimum, eye protection, a lab coat, and gloves. In certain circumstances other items of protective equipment may be necessary (e.g., a face shield). Different types of gloves exhibit different resistance properties (Table A.2A.3). No gloves resist all chemicals, and no gloves resist any chemicals indefinitely. Disposable gloves labeled “exam” or “examination” are primarily for protection from biological materials (e.g., viruses, bacteria, feces, blood). They are not designed for and usually have not been tested for resistance to chemicals. Disposable gloves generally offer extremely marginal protection from chemical hazards in most cases and should be removed immediately upon contamination before the chemical can pass through. If possible, design handling procedures to eliminate or reduce potential for contamination. Never assume that disposable gloves will offer the same protection or even have the same properties as nondisposables. Select gloves carefully and always look for some evidence that they will protect against the materials being used. Inspect all gloves before every use for possible holes, tears, or weak areas. Never reuse disposable gloves. Clean reusable gloves after each use and dry carefully inside and out. Observe all common-sense precautions—e.g., do not pipet by mouth, keep unauthorized persons away from hazardous chemicals, do not eat or drink in the lab, wear proper clothing in the lab (sandals, open-toed shoes, and shorts are not appropriate). Order hazardous chemicals only in quantities that are likely to be used in a reasonable time. Buying large quantities at a lower unit cost is no bargain if someone (perhaps you) has to pay to dispose of surplus quantities. Substitute alcohol-filled thermometers for mercury-filled thermometers, which are a hazardous chemical spill waiting to happen.
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Table A.2A.3
Chemical Resistance of Commonly Used Glovesa,b
Chemical *Acetaldehyde Acetic acid *Acetone Ammonium hydroxide *Amyl acetate Aniline *Benzaldehyde *Benzene Butyl acetate Butyl alcohol Carbon disulfide *Carbon tetrachloride *Chlorobenzene *Chloroform Chloronaphthalene Chromic acid (50%) Cyclohexanol *Dibutyl phthalate Diisobutyl ketone Dimethylformamide Dioctyl phthalate Epoxy resins, dry *Ethyl acetate Ethyl alcohol *Ethyl ether *Ethylene dichloride Ethylene glycol Formaldehyde Formic acid Freon 11, 12, 21, 22 *Furfural Glycerin Hexane Hydrochloric acid Hydrofluoric acid (48%) Hydrogen peroxide (30%) Ketones Lactic acid (85%) Linseed oil Methyl alcohol Methylamine Methyl bromide *Methyl ethyl ketone *Methyl isobutylketone Methyl methacrylate Monoethanolamine
Neoprene gloves
Latex gloves
Butyl gloves
Nitrile gloves
VG VG G VG F G F P G VG F F F G F F G G P F G VG G VG VG F VG VG VG G G VG F VG VG G G VG VG VG F G G F G VG
G VG VG VG P F F P F VG F P P P P P F P F F P VG F VG G P VG VG VG P G VG P G G G VG VG P VG F F G F G G
VG VG VG VG F F G P F VG F P F P F F G G G G F VG G VG VG F VG VG VG F G VG P G G G VG VG F VG G G VG VG VG VG
G VG P VG P P G F P VG F G P E F F VG G P G VG VG F VG G P VG VG VG G G VG G G G G P VG VG VG G F P P F VG continued
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Table A.2A.3
Chemical Resistance of Commonly Used Glovesa,b, continued
Chemical Morpholine Naphthalene Naphthas, aliphatic Naphthas, aromatic *Nitric acid Nitric acid, red and white fuming Nitropropane (95.5%) Oleic acid Oxalic acid Palmitic acid Perchloric acid (60%) Perchloroethylene Phenol Phosphoric acid Potassium hydroxide Propyl acetate i-Propyl alcohol n-Propyl alcohol Sodium hydroxide Styrene (100%) Sulfuric acid Tetrahydrofuran *Toluene Toluene diisocyanate *Trichloroethylene Triethanolamine Tung oil Turpentine *Xylene
Neoprene gloves
Latex gloves
Butyl gloves
Nitrile gloves
VG G VG G G P
VG F F P F P
VG F F P F P
G G VG G F P
F VG VG VG VG F VG VG VG G VG VG VG P G P F F F VG VG G P
P F VG VG F P F G VG F VG VG VG P G F P G F G P F P
F G VG VG G P G VG VG G VG VG VG P G F P G P G F F P
F VG VG VG G G F VG VG F VG VG VG F G F F F G VG VG VG F
aPerformance varies with glove thickness and duration of contact. An asterisk indicates limited use. Abbreviations: VG,
very good; G, good; F, fair; P, poor (do not use). bAdapted from the July 8, 1998, version of the DOE OSH Technical Reference Chapter 5 (APPENDIX C at
http://tis.eh.doe.gov/docs/osh_tr/ch5c.html). For more information also see Forsberg and Keith (1999).
Although any number of chemicals commonly used in laboratories are toxic if used improperly, the toxic properties of a number of reagents require special mention. Chemicals that exhibit carcinogenic, corrosive, flammable, lachrymatory, mutagenic, oxidizing, teratogenic, toxic, or other hazardous properties are listed in Table A.2A.1. Chemicals listed as carcinogenic range from those accepted by expert review groups as causing cancer in humans to those for which only minimal evidence of carcinogenicity exists. No effort has been made to differentiate the carcinogenic potential of the compounds in Table A.2A.1. Oxidizers may react violently with oxidizable material (e.g., hydrocarbons, wood, and cellulose). Before using any of these chemicals, thoroughly investigate all its characteristics. Material Safety Data Sheets are readily available; they list some hazards but vary widely in quality. A number of texts describing hazardous properties are listed at the end of this Appendix (see Literature Cited). In particular, Sax’s Dangerous Properties of Industrial Materials, 10th ed. (Lewis, 1999) and the Handbook of Reactive Chemical Hazards, 6th ed. (Bretherick et al., 1999) give comprehensive listings of known hazardous
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properties; however, these texts list only the known properties. Many chemicals, especially fluorochromes, have been tested only partially or not at all. Prudence dictates that, unless there is good reason for believing otherwise, all chemicals should be regarded as volatile, highly toxic, flammable human carcinogens and should be handled with great care. Waste should be segregated according to institutional requirements, for example, into solid, aqueous, nonchlorinated organic, and chlorinated organic material, and should always be disposed of in accordance with all applicable federal, state, and local regulations. Extensive information and cautionary details along with techniques for the disposal of chemicals in laboratories have been published (Bretherick, 1986; Lunn and Sansone, 1994a; O’Neil, 2001; Furr, 2000). Some commonly used disposal procedures are outlined in Basic Protocols 1 to 11. Incorporation of these procedures into laboratory protocols can help to minimize waste disposal problems. Alternate Protocols 1 to 7 describe decontamination methods for some of the chemicals. Support Protocols 1 to 9 describe analytical techniques that are used to verify that reagents have been decontaminated; with modification, these assays may also be used to determine the concentration of a particular chemical. DISPOSAL METHODS A number of procedures for the disposal of hazardous chemicals are available; protocols for the disposal and decontamination of some hazardous chemicals commonly encountered in molecular biology laboratories are listed in Table A.2A.4. These procedures are necessarily brief; for full details consult the original references or a collection of these procedures (see Lunn and Sansone, 1994a). CAUTION: These disposal methods should be carried out only in a chemical fume hood by workers equipped with eye protection, a lab coat, and gloves. Additional protective equipment may be necessary. BASIC PROTOCOL 1
DISPOSAL OF BENZIDINE AND DIAMINOBENZIDINE Benzidine and diaminobenzidine can be degraded by oxidation with potassium permanganate (Castegnaro et al., 1985; Lunn and Sansone, 1991a). This protocol presents a method for decontamination of benzidine and diaminobenzidine in bulk. This method can also be adapted to the decontamination of benzidine and diaminobenzidine spills (see Alternate Protocol 1). These compounds can also be removed from solution using horseradish peroxidase in the presence of hydrogen peroxide (see Alternate Protocol 2). Destruction and decontamination are >99%. Support Protocol 1 is used to test for the presence of benzidine and diaminobenzidine. Materials Benzidine or diaminobenzidine tetrahydrochloride dihydrate 0.1 M HCl (for benzidine) 0.2 M potassium permanganate: prepare immediately before use 2 M sulfuric acid Sodium metabisulfite 10 M potassium hydroxide (KOH) Additional reagents and equipment for testing for the presence of aromatic amines (see Support Protocol 1)
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Table A.2A.4
Protocols for Disposal of Some Hazardous Chemicals
Protocol
Disposal method for
Basic Protocol 1 Alternate Protocol 1 Alternate Protocol 2 Support Protocol 1
Benzidine and diaminobenzidine Spills of benzidine and diaminobenzidine Aqueous solutions of benzidine and diaminobenzidine Analysis for benzidine and diaminobenzidine
Basic Protocol 2 Alternate Protocol 3 Support Protocol 2
Biological stains Large volumes of dilute biological stains Analysis for biological stains
Basic Protocol 3
Silanes
Basic Protocol 4 Support Protocol 3
Cyanide and cyanogen bromide Analysis for cyanide
Basic Protocol 5
Dimethyl sulfate, diethyl sulfate, methyl methanesulfonate, ethyl methanesulfonate, diepoxybutane, 1,3-propane sultone Analysis for dimethyl sulfate, diethyl sulfate, methyl methanesulfonate, ethyl methanesulfonate, diepoxybutane, 1,3-propane sultone
Support Protocol 4 Basic Protocol 6 Alternate Protocol 4 Alternate Protocol 5 Alternate Protocol 6 Support Protocol 5
Ethidium bromide and propidium iodide Equipment contaminated with ethidium bromide Ethidium bromide in isopropanol containing cesium chloride Ethidium bromide in alcohols Analysis for ethidium bromide and propidium iodide
Basic Protocol 7
Hydrogen peroxide
Basic Protocol 8
Iodine
Basic Protocol 9 Alternate Protocol 7 Support Protocol 6
Mercury compounds Waste water containing mercury Analysis for mercury
Basic Protocol 10 Support Protocol 7 Support Protocol 8
Sodium azide Analysis for sodium azide Analysis for nitrite
Basic Protocol 11 Support Protocol 9
Enzyme inhibitors Analysis for enzyme inhibitors
1. For each 9 mg benzidine, add 10 ml of 0.1 M HCl or for each 9 mg diaminobenzidine tetrahydrochloride dihydrate, add 10 ml water. Stir the solution until the aromatic amine has completely dissolved. 2. For each 10 ml of solution, add 5 ml freshly prepared 0.2 M potassium permanganate and 5 ml of 2 M sulfuric acid. Allow the mixture to stand for ≥10 hr. 3. Add sodium metabisulfite until the solution is decolorized. 4. Add 10 M KOH to make the solution strongly basic, pH >12. CAUTION: This reaction is exothermic.
5. Dilute with 5 vol water and pass through filter paper to remove manganese compounds. 6. Test the filtrate for the presence of aromatic amines (i.e., benzidine or diaminobenzidine; see Support Protocol 1).
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7. Neutralize the filtrate with acid and discard. ALTERNATE PROTOCOL 1
DECONTAMINATION OF SPILLS INVOLVING BENZIDINE AND DIAMINOBENZIDINE Additional Materials (also see Basic Protocol 1) Glacial acetic acid 1:1 (v/v) 0.2 M potassium permanganate/2 M sulfuric acid: prepare immediately before use Absorbent material (e.g., paper towels, Kimwipes) High-efficiency particulate air (HEPA) vacuum (Fisher) Additional reagents and equipment for testing for the presence of aromatic amines (see Support Protocol 1) CAUTION: This procedure may damage painted surfaces and Formica. 1. Remove as much of the spill as possible using absorbent material and high-efficiency particulate air (HEPA) vacuuming. 2. Wet the surface with glacial acetic acid until all the amines are dissolved, then add an excess of freshly prepared 1:1 (v/v) 0.2 M potassium permanganate/2 M sulfuric acid to the spill area. Allow the mixture to stand ≥10 hr. 3. Ventilate the area and decolorize with sodium metabisulfite. 4. Mop up the liquid with paper towels. Squeeze the solution out of the towels and collect in a suitable container. Discard towels as hazardous solid waste. 5. Add 10 M KOH to make the solution strongly basic, pH ≥12. CAUTION: This reaction is exothermic.
6. Dilute with 5 vol water and filter through filter paper to remove manganese compounds. 7. Test the filtrate for the presence of aromatic amines (i.e., benzidine or diaminobenzidine; see Support Protocol 1). 8. Neutralize the filtrate with acid and discard it. 9. Verify complete decontamination by wiping the surface with a paper towel moistened with water and squeezing the liquid out of the towel. Test the liquid for the presence of benzidine or diaminobenzidine (see Support Protocol 1). Repeat steps 1 to 9 as necessary. ALTERNATE PROTOCOL 2
Laboratory Safety
DECONTAMINATION OF AQUEOUS SOLUTIONS OF BENZIDINE AND DIAMINOBENZIDINE The enzyme horseradish peroxidase catalyzes the oxidation of the amine to a radical which diffuses into solution and polymerizes. The polymers are insoluble and fall out of solution. Additional Materials (also see Basic Protocol 1) Aqueous solution of benzidine or diaminobenzidine 1 N HCl or NaOH 3% (v/v) hydrogen peroxide Horseradish peroxidase (see recipe) 1:1 (v/v) 0.2 M potassium permanganate/2 M sulfuric acid 5% (w/v) ascorbic acid Porous glass filter or Sorvall GLC-1 centrifuge or equivalent
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Additional reagents and equipment for testing for the presence of aromatic amines (see Support Protocol 1) 1. Adjust the pH of the aqueous benzidine or diaminobenzidine solution to 5 to 7 with 1 N HCl or NaOH as required and dilute so the concentration of aromatic amines is ≤100 mg/liter. 2. For each liter of solution, add 3 ml of 3% hydrogen peroxide and 300 U horseradish peroxidase. Let the mixture stand 3 hr. 3. Remove the precipitate by filtering the solution through a porous glass filter or by centrifuging 5 min at room temperature in a benchtop centrifuge to pellet the precipitate. The precipitate is mutagenic and should be treated as hazardous waste.
4. Immerse the porous glass filter in 1:1 (v/v) 0.2 M potassium permanganate/2 M sulfuric acid. Clean the filter in a conventional fashion and discard potassium permanganate/sulfuric acid solution as described for benzidine and diaminobenzidine (see Basic Protocol 1). 5. For each liter of filtrate, add 100 ml of 5% ascorbic acid. 6. Test the filtrate for the presence of aromatic amines (see Support Protocol 1). 7. Discard the decontaminated filtrate. ANALYTICAL PROCEDURES TO DETECT BENZIDINE AND DIAMINOBENZIDINE Reversed-phase HPLC (Snyder et al., 1997) is used to test for the presence of aromatic amines. The limit of detection is 1 µg/ml for benzidine and 0.25 µg/ml for diaminobenzidine.
SUPPORT PROTOCOL 1
Materials Decontaminated aromatic amine solution 10:30:20 (v/v/v) acetonitrile/methanol/1.5 mM potassium phosphate buffer (1.5 mM K2HPO4/1.5 mM KH2PO4) (benzidine) or 75:25 (v/v) methanol/1.5 mM potassium phosphate buffer (diaminobenzidine) 250-mm × 4.6-mm-i.d. Microsorb C-8 reversed-phase HPLC column (Varian) or equivalent Additional reagents and equipment for reversed-phase liquid chromatography (Snyder et al., 1997) Analyze the decontaminated aromatic amine solution by reversed-phase HPLC using a 250-mm × 4.6-mm-i.d. Microsorb C-8 column or equivalent. To detect benzidine, elute with 10:30:20 (v/v/v) acetonitrile/methanol/1.5 mM potassium phosphate buffer at a flow rate of 1.5 ml/min and UV detection at 285 nm. To detect diaminobenzidine, elute with 75:25 (v/v) methanol/1.5 mM potassium phosphate buffer at a flow rate of 1 ml/min and UV detection at 300 nm.
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BASIC PROTOCOL 2
DISPOSAL OF BIOLOGICAL STAINS Biological stains (Table A.2A.5), as well as ethidium bromide and propidium iodide, can be removed from solution using the polymeric resin Amberlite XAD-16. The decontaminated solution may be disposed of as nonhazardous aqueous waste and the resin as hazardous solid waste. The volume of contaminated resin generated is much smaller than the original volume of the solution of biological stain, so the waste disposal problem is greatly reduced. The final concentration of any remaining stain should be less than the limit of detection (see Support Protocol 2 and Table A.2A.5). In each case decontamination should be >99%. This protocol describes a method for batch decontamination in which the resin is stirred in the solution to be decontaminated and removed by filtration at the end of the reaction time. Large volumes of biological stain can be decontaminated using a column (see Alternate Protocol 3). For full details refer to the original literature (Lunn and Sansone, 1991b) or a compilation (Lunn and Sansone, 1994a). Materials Amberlite XAD-16 resin (Supelco) 100 µg/ml biological stain in water Additional reagents and equipment for testing for the presence of biological stain (see Support Protocol 2) For batch decontamination of 20 ml stain 1a. Add 1 g Amberlite XAD-16 to 20 ml of 100 µg/ml biological stain in water. Table A.2A.5
Decontamination of Biological Stains
Compound Acridine orange Alcian blue 8GX Alizarin red S Azure A Azure B Brilliant blue R Congo red Coomassie brilliant blue G Cresyl violet acetate Crystal violet Eosin B Erythrosin B Ethidium bromide Janus green B Methylene blue Neutral red Nigrosin Orcein Propidium iodide Rose Bengal Safranine O Toluidine blue O Trypan blue
Time required for complete decontamination
Volume of solution (ml) decontaminated per gram resin
18 hr 10 min 18 hr 10 min 10 min 2 hr 2 hr 2 hr 2 hr 30 min 30 min 18 hr 4 hr 30 min 30 min 10 min 2 hr 2 hr 2 hr 3 hr 1 hr 30 min 2 hr
20 500 5 80 80 80 40 80 40 200 40 10 20 80 80 500 80 200 20 20 20 80 40
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For aqueous solutions having stain concentrations other than 100 ìg/ml, use proportionately greater or lesser amounts of resin to achieve complete decontamination. For solutions of erythrosin B, use 2 g Amberlite XAD-16 for 20 ml stain.
2a. Stir the mixture for at least the time indicated in Table A.2A.5. For batch decontamination of larger volumes of stain 1b. Add 1 g Amberlite XAD-16 to the volume of a 100 µg/ml biological stain in water indicated in Table A.2A.5. 2b. Stir the mixture for at least 18 hr. 3. Remove the resin by filtration through filter paper. 4. Test the filtrate for the presence of the biological stain (see Support Protocol 2). 5. Discard the resin as hazardous solid waste and the decontaminated filtrate as liquid waste. CONTINUOUS-FLOW DECONTAMINATION OF AQUEOUS SOLUTIONS OF BIOLOGICAL STAINS
ALTERNATE PROTOCOL 3
For treating large volumes of dilute aqueous solutions of biological stains (Table A.2A.5), it is possible to put the resin in a column and run the contaminated solution through the column in a continuous-flow system (Lunn et al., 1994). Limited grinding of the Amberlite XAD-16 resin increases its efficiency. Additional Materials (also see Basic Protocol 2) 25 µg/ml biological stain in water Methanol (optional) 300-mm × 11-mm-i.d. glass chromatography column fitted with threaded adapters and flow-regulating valves at top and bottom nut and insert connectors, and insertion tool (Ace Glass) or 300-mm × 15-mm-i.d. glass chromatography column (Spectrum 124010, Fisher) Glass wool 1.5-mm-i.d. × 0.3-mm-wall Teflon tubing Waring blender (optional) Rubber stopper fitted over a pencil QG 20 lab pump (Fluid Metering) Additional reagents and equipment for testing for the presence of biological stain (see Support Protocol 2) Using a slurry of Amberlite XAD-16 1a. Prepare a 300-mm × 11-mm-i.d. glass chromatography column. To prevent clogging of the column outlet, place a small plug of glass wool at the bottom of the chromatography column. Connect 1.5-mm-i.d. × 0.3-mm wall Teflon tubing to the adapters using nut and insert connectors. Attach the tubing using an insertion tool. 2a. Mix 10 g Amberlite XAD-16 and 25 ml water in a beaker and stir 5 min to wet the resin. Using a finely ground Amberlite XAD-16 slurry 1b. Prepare a 300-mm × 15-mm-i.d. glass chromatography column. To prevent clogging of the column outlet, place a small plug of glass wool at the bottom of the chromatography column.
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Table A.2A.6 Breakthrough Volumes for Continuous-Flow Decontamination of Biological Stains
Breakthrough volume (ml) Compound Acridine orange Alizarin red S Azure A Azure B Cresyl violet acetate Crystal violet Ethidium bromide Janus green B Methylene blue Neutral red Safranine O Toluidine blue O
Limit of detection
1 ppm
5 ppm
465 120 615 630 706 1020 260 170 420 >2480 365 353
>990 150 810 882 >1396 >1630 312 650 645 >2480 438 494
>990 240 >975 >1209 >1396 >1630 416 >870 1050 >2480 584 606
2b. Grind 20 g Amberlite XAD-16 with 200 ml water for exactly 10 sec in a Waring blender. 3. Pour the resin slurry into the column through a funnel. As the resin settles, tap the column with a rubber stopper fitted over a pencil to encourage even packing. Attach a QG 20 lab pump. 4. Pump the 25-µg/ml biological stain solution through the column at 2 ml/min. Alternatively, gravity flow coupled with periodic adjustment of the flow-regulating valve can be used.
5. Check the effluent from the column for the presence of biological stain (see Support Protocol 2). Stop the pump when stain is detected. Table A.2A.6 lists breakthrough volumes at different detection levels for a number of biological stains.
6. Discard the decontaminated effluent and the contaminated resin appropriately. 7. Many biological stains can be washed off the resin with methanol so the resin can be reused. Discard the methanol solution of stain as hazardous organic liquid waste. SUPPORT PROTOCOL 2
ANALYTICAL PROCEDURES TO DETECT BIOLOGICAL STAIN Depending on the biological stain, the filtrate or eluate from the decontamination procedure can be analyzed using either UV absorption (A) or fluorescence detection (F). Materials Filtrate or eluate from biological stain decontamination (see Basic Protocol 2 or Alternate Protocol 3) pH 5 buffer (see recipe) 1 M KOH solution 20 µg/ml calf thymus DNA in TBE electrophoresis buffer, pH 8.1 (APPENDIX 2E)
Laboratory Safety
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Table A.2A.7
Methods for Detecting Biological Stainsa
Compound
Reagentb
Acridine orange Alcian blue 8GX Alizarin red S Azure A Azure B Brilliant blue R Congo red Coomassie brilliant blue G Cresyl violet acetate Crystal violet Eosin B Erythrosin B Ethidium bromide Janus green B Methylene blue Neutral red Nigrosin Orcein Propidium iodide Rose Bengal Safranine O Toluidine blue O Trypan blue
DNA solution 1 M KOH
pH 5 buffer
pH 5 buffer 1 M KOH DNA solution
Procedure
Wavelength(s)(nm)
Limit of detection (ppm)
F A A A A A A A
ex 492, em 528 615 556 633 648 585 497 610
0.0032 0.9 0.46 0.15 0.13 1.0 0.25 1.7
F A A F F A A A A A F F F A A
ex 588, em 618 588 514 ex 488, em 556 ex 540, em 590 660 661 540 570 579 ex 350, em 600 ex 520, em 576 ex 460, em 582 626 607
0.021 0.1 0.21 0.025 0.05 0.6 0.13 0.6 0.8 1.15 0.1 0.04 0.03 0.2 0.22
aAbbreviations: A, absorbance; em, emission; ex, excitation; F, fluorescence bSee Support Protocol 2
Test the filtrate or eluate from the biological stain decontamination procedure using the appropriate method as indicated in Table A.2A.7. Traces of acid or base on the resin may induce color changes in the stain. Accordingly, with cresyl violet acetate or neutral red, mix aliquots of the filtrate with 1 vol pH 5 buffer before analyzing. With alizarin red S and orcein, mix aliquots of the filtrate with 1 vol of 1 M KOH solution before analyzing. Increase the fluorescence of solutions of acridine orange, ethidium bromide, and propidium iodide by mixing an aliquot of the filtrate with an equal volume of 20 ìg/ml calf thymus DNA in TBE electrophoresis buffer, pH 8.1. Let the solution stand 15 min before measuring the fluorescence.
DISPOSAL OF CHLOROTRIMETHYLSILANE AND DICHLORODIMETHYLSILANE Silane-containing compounds are hydrolyzed to hydrochloric acid and polymeric siliconcontaining material (Patnode and Wilcock, 1946).
BASIC PROTOCOL 3
1. Hydrolyze silane-containing compounds by cautiously adding 5 ml silane to 100 ml vigorously stirred water in a flask. Allow the resulting suspension to settle. 2. Remove any insoluble material by filtration and discard it with the solid or liquid hazardous waste. 3. Neutralize the aqueous layer with base and discard it.
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BASIC PROTOCOL 4
DISPOSAL OF CYANIDES AND CYANOGEN BROMIDE Inorganic cyanides (e.g., NaCN) and cyanogen bromide (CNBr) are oxidized by sodium hypochlorite (NaOCl; e.g., Clorox) in basic solution to the much less toxic cyanate ion (Lunn and Sansone, 1985a). Destruction is >99.7%. Materials Cyanide (e.g., NaCN) or cyanogen bromide (CNBr) 1 M NaOH 5.25% (v/v) sodium hypochlorite (NaOCl; i.e., standard household bleach) Additional reagents and equipment for testing for the presence of cyanide (see Support Protocol 3) 1. Dissolve cyanide in water to give a concentration ≤25 mg/ml or dissolve CNBr in water to give a concentration ≤60 mg/ml. If necessary, dilute aqueous solutions so the concentration of NaCN or CNBr does not exceed the limit.
2. Mix 1 vol NaCN or CNBr solution with 1 vol 1 M NaOH and 2 vol fresh 5.25% NaOCl. Stir the mixture 3 hr. IMPORTANT NOTE: With age bleach may become ineffective; use of fresh bleach is strongly recommended.
3. Test the reaction mixture for the presence of cyanide (see Support Protocol 3). 4. Neutralize the reaction mixture and discard it. SUPPORT PROTOCOL 3
ANALYTICAL PROCEDURE TO DETECT CYANIDE This protocol is used to detect cyanide or cyanogen bromide at ≥3 µg/ml. Materials Cyanide or cyanogen bromide decontamination reaction mixture (see Basic Protocol 4) Phosphate buffer (see recipe) 10 mg/ml sodium ascorbate in water: prepare fresh daily 100 mg/liter NaCN in water: prepare fresh weekly 10 mg/ml chloramine-T in water: prepare fresh daily Cyanide detection reagent (see recipe) Sorvall GLC-1 centrifuge or equivalent 1. If necessary to remove suspended solids, centrifuge two 1-ml aliquots of the cyanide or cyanogen bromide decontamination reaction mixture 5 min in a benchtop centrifuge, room temperature. Add each supernatant to 4 ml phosphate buffer in separate tubes. 2. If an orange or yellow color appears, add 10 mg/ml freshly prepared sodium ascorbate dropwise until the mixture is colorless, but do not add more than 2 ml. 3. Add 200 µl of 100 mg/liter NaCN to one reaction mixture (control solution). 4. Add 1 ml freshly prepared 10 mg/ml chloramine-T to each tube. Shake the tubes and let them stand 1 to 2 min. 5. Add 1 ml cyanide detection reagent, shake, and let stand 5 min.
Laboratory Safety
A blue color indicates the presence of cyanide. If destruction has been complete and the analytical procedure has been carried out correctly, the treated reaction mixture should be colorless and the control solution, which contains NaCN, should be blue.
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6. Centrifuge tubes 5 min, room temperature, if necessary to remove suspended solids. Measure the absorbance at 605 nm with appropriate standards and blanks. DISPOSAL OF DIMETHYL SULFATE, DIETHYL SULFATE, METHYL METHANESULFONATE, ETHYL METHANESULFONATE, DIEPOXYBUTANE, AND 1,3-PROPANE SULTONE
BASIC PROTOCOL 5
Dimethyl sulfate is hydrolyzed by base to methanol and methyl hydrogen sulfate (Lunn and Sansone, 1985b). Subsequent hydrolysis of methyl hydrogen sulfate to methanol and sulfuric acid is slow. Methyl hydrogen sulfate is nonmutagenic and a very poor alkylating agent. The other compounds can be hydrolyzed with base in a similar fashion (Lunn and Sansone, 1990a). Destruction is >99%. A method to verify that decontamination is complete is also provided (see Support Protocol 4). NOTE: The reaction times given in the protocol should give good results; however, reaction time may be affected by such factors as the size and shape of the flask and the rate of stirring. The presence of two phases indicates that the reaction is not complete, and stirring should be continued until the reaction mixture is homogeneous. Materials Dimethyl sulfate, diethyl sulfate, methyl methanesulfonate, ethyl methanesulfonate, diepoxybutane, or 1,3-propane sultone 5 M NaOH Additional reagents and equipment for testing for the presence of dimethyl sulfate, diethyl sulfate, methyl methanesulfonate, ethyl methanesulfonate, diepoxybutane, or 1,3-propane sultone (see Support Protocol 4) For bulk quantities of dimethyl sulfate 1a. Add 100 ml dimethyl sulfate to 1 liter of 5 M NaOH. Stir the reaction mixture. 2a. Fifteen minutes after all the dimethyl sulfate has gone into solution, neutralize the reaction mixture with acid. For bulk quantities of diethyl sulfate 1b. Add 100 ml diethyl sulfate to 1 liter of 5 M NaOH. Stir the reaction mixture 24 hr. 2b. Neutralize the reaction mixture with acid. For bulk quantities of methyl methanesulfonate, ethyl methanesulfonate, diepoxybutane, and 1,3-propane sultone 1c. Add 1 ml methyl methanesulfonate, ethyl methanesulfonate, or diepoxybutane, or 1 g of 1,3-propane sultone to 10 ml of 5 M NaOH. Stir the reaction mixture 1 hr for 1,3-propane sultone, 2 hr for methyl methanesulfonate, 22 hr for diepoxybutane, or 24 hr for ethyl methanesulfonate. 2c. Neutralize the reaction mixture with acid. 3. Test the reaction mixture for the presence of the original compound (see Support Protocol 4). 4. Discard the decontaminated reaction mix.
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SUPPORT PROTOCOL 4
ANALYTICAL PROCEDURE TO DETECT THE PRESENCE OF DIMETHYL SULFATE, DIETHYL SULFATE, METHYL METHANESULFONATE, ETHYL METHANESULFONATE, DIEPOXYBUTANE, AND 1,3-PROPANE SULTONE This protocol is used to verify decontamination of solutions containing dimethyl sulfate, diethyl sulfate, methyl methanesulfonate, ethyl methanesulfonate, diepoxybutane, or 1,3-propane sultone. The detection limit for this assay is 40 µg/ml for dimethyl sulfate, 108 µg/ml for diethyl sulfate, 84 µg/ml for methyl methanesulfonate, 1.1 µg/ml for ethyl methanesulfonate, 360 µg/ml for diepoxybutane, and 264 µg/ml for 1,3-propane sultone. Materials Reaction mixture containing dimethyl sulfate, diethyl sulfate, methyl methanesulfonate, ethyl methanesulfonate, diepoxybutane, or 1,3-propane sultone 98:2 (v/v) 2-methoxyethanol/acetic acid 5% (w/v) 4-(4-nitrobenzyl)pyridine in 2-methoxyethanol Piperidine 2-Methoxyethanol 1. Dilute an aliquot of the reaction mixture with 4 vol water. 2. Add 100 µl diluted reaction mixture to 1 ml of 98:2 (v/v) 2-methoxyethanol/acetic acid. Swirl to mix. 3. Add 1 ml of 5% (w/v) 4-(4-nitrobenzyl)pyridine in 2-methoxyethanol. Heat 10 min at 100°C, then cool 5 min in ice. 4. Add 0.5 ml piperidine and 2 ml of 2-methoxyethanol. 5. Measure the absorbance of the violet reaction mixture at 560 nm against an appropriate blank. The absorbance of a decontaminated solution should be 0.000.
BASIC PROTOCOL 6
DISPOSAL OF ETHIDIUM BROMIDE AND PROPIDIUM IODIDE Ethidium bromide and propidium iodide in water and buffer solutions may be degraded by reaction with sodium nitrite and hypophosphorous acid in aqueous solution (Lunn and Sansone, 1987); destruction is >99.87%. This reaction may also be used to decontaminate equipment contaminated with ethidium bromide (see Alternate Protocol 4; Lunn and Sansone, 1989) and to degrade ethidium bromide in organic solvents (see Alternate Protocol 5 and Alternate Protocol 6; Lunn and Sansone, 1990b). Ethidium bromide and propidium iodide may also be removed from solution by adsorption onto Amberlite XAD-16 resin (see Basic Protocol 2). Materials Ethidium bromide– or propidium iodide–containing solution in water, buffer, or 1 g/ml cesium chloride 5% (v/v) hypophosphorous acid: prepare fresh daily by diluting commercial 50% reagent 1/10 0.5 M sodium nitrite: prepare fresh daily Sodium bicarbonate Additional reagents and equipment for testing for the presence of ethidium bromide or propidium iodide (see Support Protocol 5)
Laboratory Safety
1. If necessary, dilute the ethidium bromide– or propidium iodide–containing solution so the concentration of ethidium bromide or propidium iodide is ≤0.5 mg/ml.
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2. For each 100 ml solution, add 20 ml of 5% hypophosphorous acid solution and 12 ml of 0.5 M sodium nitrite. Stir briefly and let stand 20 hr. 3. Neutralize the reaction mixture by adding sodium bicarbonate until the evolution of gas ceases. 4. Test the reaction mixture for the presence of ethidium bromide or propidium iodide (see Support Protocol 5). 5. Discard the decontaminated reaction mixture. DECONTAMINATION OF EQUIPMENT CONTAMINATED WITH ETHIDIUM BROMIDE
ALTERNATE PROTOCOL 4
Glass, stainless steel, Formica, floor tile, and the filters of transilluminators have been successfully decontaminated using this protocol. No change in the optical properties of the transilluminator filter could be detected, even after a number of decontamination cycles. Materials Equipment contaminated with ethidium bromide Decontamination solution (see recipe) Sodium bicarbonate Additional reagents and equipment for testing for the presence of ethidium bromide (see Support Protocol 5) 1. Wash the equipment contaminated with ethidium bromide once with a paper towel soaked in decontamination solution. The pH of the decontamination solution is 1.8. If this would be too corrosive for the surface to be decontaminated, wash with a paper towel soaked in water instead.
2. Wash the surface five times with paper towels soaked in water using a fresh towel each time. 3. Soak all the towels 1 hr in decontamination solution. 4. Neutralize the decontamination solution by adding sodium bicarbonate until the evolution of gas ceases. 5. Test the decontamination solution for the presence of ethidium bromide (see Support Protocol 5). 6. Discard the decontamination solution and the paper towels as nonhazardous liquid and solid wastes. DECONTAMINATION OF ETHIDIUM BROMIDE IN ISOPROPANOL SATURATED WITH CESIUM CHLORIDE
ALTERNATE PROTOCOL 5
Materials Ethidium bromide in isopropanol saturated with cesium chloride Decontamination solution (see recipe) Sodium bicarbonate Additional reagents and equipment for testing for the presence of ethidium bromide (see Support Protocol 5)
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1. If necessary, dilute the ethidium bromide in isopropanol saturated with cesium chloride so the concentration of ethidium bromide is ≤1 mg/ml. 2. For each volume of ethidium bromide solution, add 4 vol decontamination solution. Stir the reaction mixture 20 hr. 3. Neutralize the reaction mixture by adding sodium bicarbonate until the evolution of gas ceases. 4. Test the reaction mixture for the presence of ethidium bromide (see Support Protocol 5). 5. Discard the decontaminated solution. ALTERNATE PROTOCOL 6
DECONTAMINATION OF ETHIDIUM BROMIDE IN ISOAMYL ALCOHOL AND 1-BUTANOL Materials Ethidium bromide in isoamyl alcohol or 1-butanol Decontamination solution (see recipe) Activated charcoal Sodium bicarbonate Separatory funnel Additional reagents and equipment for testing for the presence of ethidium bromide 1. If necessary, dilute the ethidium bromide in isoamyl alcohol or 1-butanol so the concentration is ≤1 mg/ml final. 2. For each volume of ethidium bromide solution, add 4 vol decontamination solution. Stir the two-phase reaction mixture rapidly for 72 hr. 3. For each 100 ml of reaction mixture, add 2 g activated charcoal. Stir another 30 min. 4. Filter the reaction mixture. 5. Neutralize the filtrate by adding sodium bicarbonate until the evolution of gas ceases. Separate the layers using a separatory funnel. More alcohol may tend to separate from the aqueous layer on standing.
6. Test the alcohol and aqueous layers for the presence of ethidium bromide. 7. Discard the alcohol and aqueous layers appropriately. Discard the activated charcoal as solid waste. The aqueous layer contains 4.6% 1-butanol or 2.3% isoamyl alcohol. SUPPORT PROTOCOL 5
ANALYTICAL PROCEDURE TO DETECT ETHIDIUM BROMIDE OR PROPIDIUM IODIDE This protocol is used to verify that solutions no longer contain ethidium bromide or propidium iodide. The limits of detection are 0.05 parts per million (ppm) for ethidium bromide and 0.1 ppm for propidium iodide. Materials Reaction mixture containing ethidium bromide or propidium iodide TBE buffer, pH 8.1 (APPENDIX 2E) 20 µg/ml calf thymus DNA in TBE buffer, pH 8.1 1. Mix 100 µl reaction mixture containing ethidium bromide or propidium iodide with 900 µl TBE buffer, pH 8.1.
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2. Add 1 ml of 20 µg/ml calf thymus DNA in TBE, pH 8.1. Prepare a blank solution (100 µl water + 900 µl TBE + 1 ml of 20 µg/ml calf thymus DNA) and control solutions containing known quantities of ethidium bromide or propidium iodide. Let the mixtures stand 15 min. 3. To detect ethidium bromide, measure the fluorescence with an excitation wavelength of 540 nm and an emission wavelength of 590 nm. To detect propidium iodide, measure the fluorescence with an excitation wavelength of 350 nm and an emission wavelength of 600 nm. If a spectrophotofluorometer is not available, fluorescence of ethidium bromide can be qualitatively determined using a hand-held UV lamp on the long-wavelength setting (Lunn and Sansone, 1991c).
DISPOSAL OF HYDROGEN PEROXIDE Hydrogen peroxide can be reduced with sodium metabisulfite (Lunn and Sansone, 1994b).
BASIC PROTOCOL 7
Materials 30% hydrogen peroxide 10% (w/v) sodium metabisulfite 10% (w/v) potassium iodide 1 M HCl 1% (w/v) starch indicator solution 1. Add 5 ml of 30% hydrogen peroxide to 100 ml of 10% sodium metabisulfite. Stir the mixture at room temperature until the temperature starts to drop, indicating that the reaction is over. 2. Test for the presence of hydrogen peroxide by adding a few drops of the reaction mixture to an equal volume of 10% potassium iodide. Add a few drops of 1 M HCl to acidify the reaction mixture, then add a drop of 1% starch indicator solution. A deep blue color indicates the presence of excess oxidant. If necessary, add more 10% sodium metabisulfite until the starch test is negative.
3. Discard the decontaminated mixture. REDUCTION OF IODINE Iodine is reduced with sodium metabisulfite to iodide (Lunn and Sansone, 1994b).
BASIC PROTOCOL 8
Materials Iodine 10% (w/v) sodium metabisulfite 1 M HCl 1% (w/v) starch indicator solution 1. Add 5 g iodine to 100 ml of 10% sodium metabisulfite. Stir the mixture until the iodine has completely dissolved. 2. Acidify a few drops of the reaction mixture by adding a few drops of 1 M HCl. Add 1 drop of 1% starch indicator solution. A deep blue color indicates the presence of iodine. If reduction is not complete, add more sodium metabisulfite solution.
3. Dispose of the decontaminated solution.
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BASIC PROTOCOL 9
DISPOSAL OF MERCURY COMPOUNDS Solutions of mercuric acetate can be decontaminated using Dowex 50X8-100, a strongly acidic gel-type ion-exchange resin with a sulfonic acid functionality. Solutions of mercuric chloride can be decontaminated using Amberlite IRA-400(Cl), a strongly basic gel-type ion-exchange resin with a quaternary ammonium functionality. The final concentration of mercury is <3.8 ppm (Lunn and Sansone, 1994a). On a small scale it is most convenient to stir the resin in the solution to be decontaminated, but on a larger scale, or for routine use, it may be more convenient to pass the solution through a column packed with the resin. Although the volume of waste that must be disposed of is greatly reduced using this technique, a small amount of waste (i.e., the resin contaminated with mercury) remains and must be discarded appropriately. Resin can be regenerated by washing with acid, but the concentrated metal-containing solution generated by this must also be disposed of appropriately. Mercury may also be removed from laboratory waste water using a column of iron powder (see Alternate Protocol 7). Support Protocol 6 is used to detect the presence of mercury. Materials Solution containing ≤1600 ppm mercuric acetate or ≤1350 ppm mercuric chloride Dowex 50X8-100 ion-exchange resin or Amberlite IRA-400(Cl) ion-exchange resin Additional reagents and equipment to test for the presence of mercury (see Support Protocol 6) 1a. For mercuric acetate: For each 200 ml of solution containing ≤1600 ppm mercuric acetate, add 1 g Dowex 50X8-100 ion-exchange resin. Stir the mixture 1 hr, then filter through filter paper. 1b. For mercuric chloride: For each 200 ml of solution containing ≤1350 ppm mercuric chloride, add 1 g Amberlite IRA-400(Cl) ion-exchange resin. Stir the mixture 6 hr, then filter through filter paper. The speed and efficiency of decontamination will depend on factors such as the size and shape of the flask and the rate of stirring.
3. Test the filtrate for the presence of mercury (see Support Protocol 6). 4. Discard the decontaminated filtrate and the mercury-containing resin appropriately. ALTERNATE PROTOCOL 7
DECONTAMINATION OF WASTE WATER CONTAINING MERCURY Laboratory waste water that contains mercury is decontaminated by passing it through a column of iron powder. The mercury forms mercury amalgam and stays on the column. Some metallic mercury remains in solution but this can be removed by aeration. The final concentration of mercury is <5 ppb (Shirakashi et al., 1986). Materials Iron powder, 60 mesh Waste water containing ≤2.5 ppm mercury 6-mm-i.d. column 1. Pack a 6-mm-i.d. column with 1 g of 60-mesh iron powder. Use a fresh column for each treatment.
Laboratory Safety
2. Pass ≤2 liters of water containing ≤2.5 ppm of mercury through the column at a flow rate of 250 ml/hr.
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Solutions containing a higher concentration of mercury may also be treated, but this will result in a higher final concentration of mercury (e.g., treating a 100-ppm solution in this fashion yielded 33 ppb final). Some iron ends up in solution and can be removed by adjusting the pH to 8. The resulting precipitated Fe(OH)3 can then be removed by filtration.
3. Aerate the resulting effluent to remove traces of metallic mercury and continue aeration 30 min after the last of the effluent has emerged from the column. Vent the metallic mercury removed from the solution by aeration into the fume hood or capture it in a mercury trap. The effluent contains <5 ppb mercury. The presence of iodide or polypeptone may necessitate several treatments to reduce the mercury to an acceptable level.
ANALYTICAL PROCEDURE TO DETECT MERCURY Atomic absorption spectroscopy with detection at 253.7 nm, a slit width of 0.7 nm, and a limit of detection of 3.8 ppm can be used to determine the concentration of mercury in solution for experiments involving ion-exchange resins. A Hiranuma mercury meter model HG-1 can be used for experiments involving iron powder.
SUPPORT PROTOCOL 6
DISPOSAL OF SODIUM AZIDE Sodium azide can be oxidized by ceric ammonium nitrate (Manufacturing Chemists Association, 1973) to nitrogen (Mason, 1967) or by nitrous acid (National Research Council, 1983) to nitrous oxide (Mason, 1967); destruction is >99.996%. Sodium azide in buffer solution may also be degraded by the addition of sodium nitrite (Lunn and Sansone, 1994a). The reaction proceeds much more readily at low pH, but if sufficient sodium nitrite is added, it will proceed to completion even at high pH. At low pH, it may be possible to completely degrade the azide present in the buffer with less than the amount of sodium nitrite indicated. However, the reaction mixture must be carefully checked to ensure that no azide remains (see Support Protocol 7). At high pH it is possible for unreacted azide to remain in the presence of excess nitrite. Residual nitrite can be detected using Support Protocol 8.
BASIC PROTOCOL 10
CAUTION: Some toxic nitrogen dioxide may be produced as a by-product of these reactions, so they should always be carried out in a chemical fume hood. Materials Sodium azide or solution containing sodium azide Ceric ammonium nitrate 10% (w/v) potassium iodide 1 M HCl 1% (w/v) starch indicator solution Sodium nitrite 4 M sulfuric acid Additional reagents and equipment to test for the presence of sodium azide (see Support Protocol 7) or nitrite (see Support Protocol 8) Decontamination using ceric ammonium nitrate 1a. For each gram of sodium azide, add 9 g ceric ammonium nitrate to 30 ml of water, and stir until it has dissolved. 2a. Dissolve each gram of sodium azide in 5 ml water. Add this solution to the ceric ammonium nitrate solution at the rate of 1 ml each min. Stir 1 hr. If the reaction is carried out on a larger scale, an ice bath may be required for cooling.
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3a. Check that the reaction is still oxidizing by adding a few drops of the reaction mixture to an equal volume of 10% potassium iodide. Acidify the mixture with 1 drop 1 M HCl and add 1 drop 1% starch indicator solution. The deep blue color of the starch-iodine complex indicates that excess oxidant is present. If excess oxidant is not present, add more ceric ammonium nitrate.
4a. Test for the presence of sodium azide (see Support Protocol 8). 5a. Discard the decontaminated reaction mixture. Decontamination using sodium nitrite 1b. For each 5 g sodium azide, dissolve 7.5 g sodium nitrite in 30 ml water. 2b. Dissolve each 5 g sodium azide in 100 ml water. Add the sodium nitrite solution with stirring. Slowly add 4 M sulfuric acid until the reaction mixture is acidic to litmus. Stir 1 hr. CAUTION: It is important to add the sodium nitrite, then the sulfuric acid. Adding these reagents in reverse order will generate explosive, volatile, toxic hydrazoic acid. If the reaction is carried out on a large scale, an ice bath may be required for cooling.
3b. Check that there is excess nitrous acid in the reaction. Add a few drops of the reaction mixture to an equal volume of 10% potassium iodide. Acidify the mixture with 1 drop 1 M HCl. Add 1 drop starch indicator solution. The deep blue color of the starch-iodine complex indicates that excess nitrous acid is present. If excess nitrous acid is not present, add more sodium nitrite.
4b. If excess nitrous acid is present, test for the presence of sodium azide (see Support Protocol 7). 5b. Discard the decontaminated reaction mixture. Decontamination of sodium azide in buffer 1c. If necessary, dilute the buffer solution with water so the concentration of sodium azide is ≤1 mg/ml. 2c. For each 50 ml buffer solution add 5 g sodium nitrite. Stir the reaction 18 hr. 3c. Test for the presence of sodium azide (see Support Protocol 7). 4c. Discard the decontaminated reaction solution. SUPPORT PROTOCOL 7
Laboratory Safety
ANALYTICAL PROCEDURES TO DETECT SODIUM AZIDE Sodium azide is analyzed by reacting azide ion with 3,5-dinitrobenzoyl chloride to form 3,5-dinitrobenzoyl azide, which can be detected by reversed-phase HPLC. The limit of detection of this assay is 0.2 µg/ml sodium azide. This protocol works only in the absence of nitrite; verify that all of the nitrite has been destroyed by sulfamic acid by using the method detailed later in this Appendix (see Support Protocol 8). Materials Reaction mixture from sodium azide treated with ceric ammonium nitrate or sodium nitrite 1 M KOH Acetonitrile Sodium azide indicator solution (see recipe) 0.2 M HCl
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20% (w/v) sulfamic acid 3,5-dinitrobenzoyl chloride 50:50 (v/v) acetonitrile/water Sorvall GLC-1 centrifuge or equivalent 25-cm × 4.6-mm-i.d. Microsorb C-8 reversed-phase HPLC column (Varian) or equivalent Additional reagents and equipment for reversed-phase liquid chromatography (Snyder et al., 1997) To analyze for azide in the presence of ceric salts 1a. To a 10-ml aliquot of the reaction mixture from sodium azide treated with ceric ammonium nitrate add 40 ml water. Add 5 ml of this diluted solution to 3 ml of 1 M KOH and mix by shaking. If <3 ml of 1 M KOH is used, precipitation of ceric salts will not be complete.
2a. Centrifuge the mixture 5 min, room temperature. 3a. Remove 2 ml supernatant and add to 1 ml acetonitrile. Add 1 drop sodium azide indicator solution, add 0.2 M HCl dropwise until the mixture turns yellow, then add 1 drop more. To analyze for azide in the presence of nitrite 1b. To 5 ml of the reaction mixture from sodium azide treated with sodium nitrite add ≥1 ml sulfamic acid to remove excess nitrite. Let stand ≥3 min. More sulfamic acid solution may be required for strongly basic reaction mixtures or those containing high concentrations of nitrite. Complete removal of nitrite can be checked by using a modified Griess reagent (see Support Protocol 8). At high pH the reaction between azide and nitrite is quite slow, so the presence of excess nitrite does not mean that all the azide has been degraded.
2b. Add 1 drop sodium azide indicator solution, then basify the mixture by adding 1 M KOH until it turns purple (typically, 3 to 10 ml are required). 3b. Add 2 ml acetonitrile. Add 0.2 M HCl dropwise until the mixture turns yellow, then add 1 drop more. If >1 ml sulfamic acid is used, add 4 ml acetonitrile.
4. Prepare a 10% (w/v) solution of 3,5-dinitrobenzoyl chloride in acetonitrile. 5. Add 50 µl of 10% dinitrobenzoyl chloride/acetonitrile to the reaction mix (step 3a or 3b). Shake the mixture and let it stand ≥3 min. Longer standing times have no effect on the HPLC analysis. However, it is crucial to use freshly prepared 3,5-dinitrobenzoyl chloride solution within minutes of its preparation. It is generally most convenient to prepare all the analytical samples with the fresh solution at the beginning of the day and analyze them over the course of the day.
6. Analyze 20 µl of each reaction mixture by reversed-phase HPLC (Snyder et al., 1997) using a mobile phase of 50:50 (v/v) acetonitrile/water with a flow rate of 1 ml/min and UV detection at 254 nm. The peak for 3,5-dinitrobenzoyl azide elutes at ∼9 min.
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SUPPORT PROTOCOL 8
ANALYTICAL PROCEDURE TO DETECT NITRITE This protocol uses a modified Griess reagent to test for the presence of nitrite. The limit of detection of this assay is 0.06 µg/ml nitrite. A similar procedure uses N-(1-naphthyl)ethylenediamine (Cunniff, 1995). Materials α-Naphthylamine 15% (v/v) aqueous acetic acid Sulfanilic acid solution (see recipe) Reaction mixture treated to remove excess nitrite (see Support Protocol 7, step 1b) 1. Prepare the modified Griess reagent by boiling 0.1 g α-naphthylamine in 20 ml water until it dissolves. While the solution is still hot, pour it into 150 ml of 15% aqueous acetic acid. Add 150 ml sulfanilic acid solution. This reagent should be stored at room temperature in a brown bottle. CAUTION: α-Naphthylamine is a carcinogen.
2. Add 3 ml of the reaction mixture treated to remove excess nitrite to 1 ml modified Griess reagent. Let stand 6 min at room temperature. 3. Measure the absorbance at 520 nm against a suitable blank. BASIC PROTOCOL 11
DISPOSAL OF ENZYME INHIBITORS The enzyme inhibitors p-amidinophenylmethanesulfonyl fluoride (APMSF), 4-(2-aminoethyl)benzenesulfonyl fluoride (AEBSF), phenylmethylsulfonyl fluoride (PMSF; Lunn and Sansone, 1994c), diisopropyl fluorophosphate (DFP; Lunn and Sansone, 1994d), Nα-p-tosyl-L-lysine chloromethyl ketone (TLCK), and N-p-tosyl-L-phenylalanine chloromethyl ketone (TPCK; Lunn and Sansone, 1994c) may be degraded by reaction with 1 M NaOH. Destruction is >99.8% (except TPCK >98.3%). The exact reaction conditions depend on the solvent (see Table A.2A.8). The solutions that were decontaminated are representative of those described in the literature. Materials Solutions of APMSF, AEBSF, PMSF, DFP, TLCK, or TPCK in buffer, DMSO, isopropanol, or water 1 M NaOH Glacial acetic acid Additional reagents and equipment for testing for the presence of the enzyme inhibitors (see Support Protocol 9) 1. If necessary, dilute the solutions with the same solvent so that the concentrations given in Table A.2A.8 are not exceeded. Bulk quantities of AEBSF, PMSF, and TPCK may be dissolved in isopropanol and bulk quantities of APMSF and TLCK may be dissolved in water at the concentrations shown in Table A.2A.8. Bulk quantities of DFP (a liquid) may be mixed directly with 1 M NaOH, taking care to make sure that all the DFP has mixed thoroughly, in the ratio shown in Table A.2A.8 (e.g., 40 ìl DFP with 1 ml of 1 M NaOH).
2. Add 1 M NaOH so that the ratio of solution to 1 M NaOH is that listed in Table A.2A.8. Laboratory Safety
3. Shake to ensure complete mixing, check that the solution is strongly basic (pH ≥12), and allow to stand for the time given in Table A.2A.8.
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Table A.2A.8
Conditions for the Destruction of Enzyme Inhibitors
Compound
Concentration
AEBSF AEBSF AEBSF APMSF APMSF APMSF APMSF DFP DFP DFP DFP PMSF PMSF PMSF TLCK TLCK TLCK TPCK TPCK TPCK
1 mM 20 mM 20 mM 2.5 mM 25 mM 25 mM 100 mM 10 mM 200 mM pure 10 mM 10 mM 100 mM 100 mM 1 mM 5 mM 5 mM 1 mM 1 mM 1 mM
Solvent Buffer(pH 5.0-8.0) DMSO Isopropanol Buffer(pH 5.0-8.0) DMSO 50:50 isopropanol:pH 3 buffer Water Buffer (pH 6.4-7.2) DMF — Water Buffer (pH 5.0-8.0) DMSO Isopropanol Buffer (pH 5.0-8.0) DMSO Water Buffer (pH 5.0-8.0) DMSO Isopropanol
Solution: 1 M NaOH
Time
1:0.1 1:10 1:10 1:0.1 1:5 1:5 1:5 1:0.2 1:2 1:25 1:0.2 1:0.1 1:5 1:5 1:0.1 1:5 1:0.1 1:0.1 1:0.1 1:0.1
1 hr 24 hr 24 hr 1 hr 24 hr 24 hr 24 hr 18 hr 18 hr 1 hr 18 hr 1 hr 24 hr 24 hr 18 hr 18 hr 18 hr 18 hr 18 hr 18 hr
4. Neutralize the reaction mixture with acetic acid and test for the presence of residual enzyme inhibitor (see Support Protocol 9). 5. Discard the decontaminated reaction mixture. ANALYTICAL PROCEDURES TO DETECT ENZYME INHIBITORS DFP can be determined using a complex procedure involving the inhibition of chymotrypsin activity. For more information, refer to Lunn and Sansone (1994d). A gas chromatographic method has also been described by Degenhardt-Langelaan and Kientz (1996). AEBSF, APMSF, PMSF, TLCK, and TPCK may be determined by reversed-phase HPLC (Snyder et al., 1997). The chromatographic conditions and limits of detection are shown in Table A.2A.9 (Lunn and Sansone, 1994c).
SUPPORT PROTOCOL 9
Materials Decontaminated enzyme inhibitor solutions Acetonitrile (HPLC grade) Water (HPLC grade) 0.1% (v/v) trifluoroacetic acid in water 10 mM phosphate buffer, pH 7 250-mm × 4.6 mm-i.d. Microsorb C-8 reversed-phase HPLC column (Varian) or equivalent Additional reagents and equipment for reversed-phase liquid chromatography (Snyder et al., 1997)
Laboratory Guidelines, Equipment, and Stock Solutions
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Table A.2A.9 HPLC Conditions for Enzyme Inhibitors
Compound
Mobile phase
AEBSF
40:60 (v/v) acetonitrile:0.1% trifluoroacetic acid 40:60 (v/v) acetonitrile:0.1% trifluoroacetic acid 50:50 (v/v) acetonitrile:water 40:60 (v/v) acetonitrile:0.1% trifluoroacetic acid 48:52 (v/v) acetonitrile:10 mM pH 7 phosphate buffer
APMSF
PMSF TLCK
TPCK
Detector
Retention time
Limit of detection
UV 225 nm
9.5 min
0.1 µg/ml
UV 232 nm
7.7 min
0.5 µg/ml
UV 220 nm
8 min
0.9 µg/ml
UV 228 nm
9.5 min
0.37 µg/ml
UV 228 nm
10.5 min
2 µg/ml
Analyze the decontaminated enzyme inhibitor solutions by reversed-phase HPLC using a 250-mm × 4.6-mm-i.d. Microsorb C-8 reversed-phase column, or equivalent, using the conditions shown in Table A.2A.9. In each case, the injection volume was 20 µl, the separation occurred at ambient temperature, and the flow rate was 1 ml/min. Check the analytical procedures by spiking an aliquot of the acidified reaction mixture with a small quantity of a dilute solution of the compound of interest. REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Cyanide detection reagent Stir 3.0 g barbituric acid in 10 ml water. Add 15 ml of 4-methylpyridine and 3 ml concentrated HCl while continuing to stir. Cool and dilute to 50 ml with water. Store at room temperature. CAUTION: This reaction is exothermic.
Decontamination solution Dissolve 4.2 g sodium nitrite (0.2 M final) and 20 ml hypophosphorous acid (3.3% w/v final) in 300 ml water. Prepare fresh. Horseradish peroxidase Dissolve hydrogen-peroxide oxidoreductase (EC 1.11.1.7 [Type II]; specific activity 150 to 200 purpurogallin U/mg, Sigma) in 1 g/liter sodium acetate to give 30 U/ml. Prepare fresh daily. For small-scale reactions, a more dilute solution can be used to avoid working with inconveniently small volumes.
pH 5 buffer 2.04 g potassium hydrogen phthalate (0.05 M final) 38 ml 0.1 M potassium hydroxide (15 mM) H2O to 200 ml Store at room temperature Laboratory Safety
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Phosphate buffer 13.6 g monobasic potassium phosphate (KH2PO4; 0.1 M final) 0.28 g dibasic sodium phosphate (Na2HPO4; 2 mM final) 3.0 g potassium bromide (KBr; 25 mM final) 1 liter H2O Store at room temperature Potassium bromide is necessary to make the assay for cyanide work correctly.
Sodium azide indicator solution 0.1 g bromocresol purple (0.4% final) 18.5 ml 0.01 M potassium hydroxide (KOH; 7.4 mM final) H2O to 25 ml Store at room temperature Sulfanilic acid solution Dissolve 0.5 g sulfanilic acid in 150 ml of 15% (v/v) aqueous acetic acid. Use immediately. LITERATURE CITED Bretherick, L. (ed.) 1986. Hazards in the Chemical Laboratory, 4th ed. Royal Society of Chemistry, London. Bretherick, L., Urben, P.G., and Pitt, M.J. 1999. Bretherick’s Handbook of Reactive Chemical Hazards, 6th ed. Butterworth-Heinemann, London. Castegnaro, M., Barek, J., Dennis, J., Ellen, G., Klibanov, M., Lafontaine, M., Mitchum, R., van Roosmalen, P., Sansone, E.B., Sternson, L.A., and Vahl, M. (eds.) 1985. Laboratory Decontamination and Destruction of Carcinogens in Laboratory Wastes: Some Aromatic Amines and 4-Nitrobiphenyl. IARC Scientific Publications No. 64. International Agency for Research on Cancer, Lyon, France. Cunniff, P. (ed.) 1995. Official Methods of Analysis of the Association of Official Analytical Chemists, 16th ed., Ch. 4, p. 14. Association of Official Analytical Chemists, Arlington, Va. Degenhardt-Langelaan, C.E.A.M. and Kientz, C.E. 1996. Capillary gas chromatographic analysis of nerve agents using large volume injections. J. Chromatogr. A.723:210-214. Forsberg, K. and Keith, L.H. 1999. Chemical Protective Clothing Performance Index Book, 2nd ed. John Wiley & Sons, New York. Furr, A.K. (ed.) 2000. CRC Handbook of Laboratory Safety, 5th ed. CRC Press, Boca Raton, Fla. Lewis, R.J. Sr. 1999. Sax’s Dangerous Properties of Industrial Materials, 10th ed. John Wiley & Sons, New York. Lunn, G. and Sansone, E.B. 1985a. Destruction of cyanogen bromide and inorganic cyanides. Anal. Biochem. 147:245-250. Lunn, G. and Sansone, E.B. 1985b. Validation of techniques for the destruction of dimethyl sulfate. Am. Ind. Hyg. Assoc. J. 46:111-114. Lunn, G. and Sansone, E.B. 1987. Ethidium bromide: Destruction and decontamination of solutions. Anal. Biochem. 162:453-458. Lunn, G. and Sansone, E.B. 1989. Decontamination of ethidium bromide spills. Appl. Ind. Hyg. 4:234-237. Lunn, G. and Sansone, E.B. 1990a. Validated methods for degrading hazardous chemicals: Some alkylating agents and other compounds. J. Chem. Educ. 67:A249-A251. Lunn, G. and Sansone, E.B. 1990b. Degradation of ethidium bromide in alcohols. BioTechniques 8:372-373. Lunn, G. and Sansone, E.B. 1991a. The safe disposal of diaminobenzidine. Appl. Occup. Environ. Hyg. 6:49-53. Lunn, G. and Sansone, E.B. 1991b. Decontamination of aqueous solutions of biological stains. Biotech. Histochem. 66:307-315. Lunn, G. and Sansone, E.B. 1991c. Decontamination of ethidium bromide spills-author’s response. Appl. Occup. Environ. Hyg. 6:644-645. Lunn, G. and Sansone, E.B. 1994a. Destruction of Hazardous Chemicals in the Laboratory, 2nd ed. John Wiley & Sons, New York. Lunn, G. and Sansone, E.B. 1994b. Safe disposal of highly reactive chemicals. J. Chem. Educ. 71:972-976. Lunn, G. and Sansone, E.B. 1994c. Degradation and disposal of some enzyme inhibitors. Scientific note. Appl. Biochem. Biotechnol. 48:57-59.
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Lunn, G. and Sansone, E.B. 1994d. Safe disposal of diisopropyl fluorophosphate (DFP). Appl. Biochem. Biotechnol. 49:165-171. Lunn, G., Klausmeyer, P.K., and Sansone, E.B. 1994. Removal of biological stains from aqueous solution using a flow-through decontamination procedure. Biotech. Histochem. 69:45-54. Manufacturing Chemists Association. 1973. Laboratory Waste Disposal Manual. p. 136. Manufacturing Chemists Association, Washington, D.C. Mason, K.G. 1967. Hydrogen azide. In Mellor’s Comprehensive Treatise on Inorganic and Theoretical Chemistry, Vol. VIII (Suppl. II) pp. l-15. John Wiley & Sons, New York. National Research Council. 1983. Prudent Practices for Disposal of Chemicals from Laboratories, p. 88. National Academy Press, Washington, D.C. O’Neil, M.J. (ed.) 2001. The Merck Index, 13th ed. Merck & Co., Whitehouse Station, N.J. Patnode, W. and Wilcock, D.F. 1946. Methylpolysiloxanes. J. Am. Chem. Soc. 68:358-363. Shirakashi, T., Nakayama, K., Kakii, K., and Kuriyama, M. 1986. Removal of mercury from laboratory waste water with iron powder. Chem. Abstr. 105:213690y. Snyder, L.R., Kirkland, J.J., and Glajch, J.L. 1997. Practical HPLC Method Development, 2nd ed. John Wiley & Sons, New York.
KEY REFERENCES The following are good general references for laboratory safety. American Chemical Society, Committee on Chemical Safety. 1995. Safety in Academic Chemistry Laboratories, 6th ed. American Chemical Society, Washington, D.C. Castegnaro, M. and Sansone, E.B. 1986. Chemical Carcinogens. Springer-Verlag, New York. DiBerardinis, L.J., First, M.W., Gatwood, G.T., and Seth, A.K. 2001. Guidelines for Laboratory Design, Health and Safety Considerations, 3rd ed. John Wiley & Sons, New York. Fleming, D.D., Richardson, J.H., Tulis, J.J., and Vesley, D. 1995. Laboratory Safety, Principles and Practices, 2nd ed. American Society for Microbiology, Washington, D.C. Freeman, N.T. and Whitehead, J. 1982. Introduction to Safety in the Chemical Laboratory. Academic Press, San Diego. Fuscaldo, A.A., Erlick, B.J., and Hindman, B. (eds.) 1980. Laboratory Safety, Theory and Practice. Academic Press, San Diego. Lees, R. and Smith, A.F. (eds.) 1984. Design, Construction, and Refurbishment of Laboratories. Ellis Horwood, Chichester, United Kingdom. Montesano, R., Bartsch, H., Boyland, E., Della Porta, G., Fishbein, L., Griesemer, R.A., Swan, A.B., and Tomatis, L. (eds.) 1979. Handling Chemical Carcinogens in the Laboratory, Problems of Safety. IARC Scientific Publications No. 33. International Agency for Research on Cancer, Lyon, France. National Research Council. 1995. Prudent Practices in the Laboratory: Handling and Disposal of Chemicals. National Academy Press, Washington, D.C. Occupational Health and Safety. 1993. National Safety Council, Chicago. Pal, S.B. (ed.) 1991. Handbook of Laboratory Health and Safety Measures, 2nd ed. Kluwer Academic Publishers, Hingham, Mass. Rosenlund, S.J. 1987. The Chemical Laboratory: Its Design and Operation: A Practical Guide for Planners of Industrial, Medical, or Educational Facilities. Noyes Publishers, Park Ridge, N.J. Young, J.A. (ed.) 1991. Improving Safety in the Chemical Laboratory: A Practical Guide, 2nd ed. John Wiley & Sons, New York.
INTERNET RESOURCES http://www.ilpi.com/msds/index.html Where to find MSDSs on the internet. Contains links to general sites, government and nonprofit sites, chemical manufacturers and suppliers, pesticides, and miscellaneous sites. http://www.OSHA.gov OSHA web site. http://www.osha-slc.gov/OshStd_data/1910_1450.html Text of OSHA Standard 29 CFR 1910.1450: Occupational Exposure to Hazardous Chemicals in Laboratories. Laboratory Safety
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http://www.osha-slc.gov/OshStd_data/1910_1000_TABLE_Z-1.html Tables of permissible exposure limits (PELs) for air contaminants. http://www.osha-slc.gov/OshStd_data/1910_1000_TABLE_Z-2.html Tables of PELs for toxic and hazardous substances. http://hazard.com/msds/index.php Main site for Vermont SIRI. One of the best general sites to start a search. Browse manufacturers alphabetically (for sheets not in the SIRI collection) or do a keyword search in the SIRI MSDS database. Lots of additional safety links and information. http://siri.uvm.edu/msds Alternate site for Vermont SIRI. http://tis.eh.doe.gov/docs/osh_tr/ch5.html DOE OSH technical reference chapter on personal protective equipment.
Contributed by George Lunn Baltimore, Maryland Gretchen Lawler (chemical resistance of gloves) Purdue University West Lafayette, Indiana
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Safe Use of Radioisotopes The use of radioisotopes to label specific molecules in a defined way has greatly furthered the discovery and dissection of biochemical pathways. The development of methods to inexpensively synthesize such tagged biological compounds on an industrial scale has enabled them to be used routinely in laboratory protocols, including many detailed in this manual. Although most of these protocols involve the use of only microcurie (µCi) amounts of radioactivity, some (particularly those describing the metabolic labeling of proteins within cells; see UNIT 3.4) require amounts on the order of tens of millicuries (mCi). In all cases where radioisotopes are used, depending on the quantity and nature of the isotope, certain precautions must be taken to ensure the safety of the scientist. This appendix outlines a few such considerations relevant to the isotopes most frequently used in biological research. In designing safe protocols for the use of radioactivity, the importance of common sense, based on an understanding of the general principles of isotopic decay and the importance of continuous monitoring with a hand-held radioactivity monitor (e.g., Geiger counter), cannot be overemphasized. In addition, it is also critical to take into account the rules, regulations, and limitations imposed by each specific institution. These are usually not optional considerations: an institution’s license to use radioactivity normally depends on strict adherence to such rules. Many of the protocols described have evolved (and are evolving still) over the years and through the millicuries in the Department of Molecular Biology and Virology at the Salk Institute. The authors are indebted to those who have trained them in the safe use of radioactivity, in particular to the members of the Salk Institute Radiation Safety Department. Most of the designs for the shields and other safety equipment shown in Figures A.2B.1, A.2.B2, and A.2B.3 were created at the Salk Institute in collaboration with Dave Clarkin and Mario Tengco. Safety equipment of similar design is available from several commercial vendors, including CBS Scientific and Research Products International (see SUPPLIERS APPENDIX).
BACKGROUND INFORMATION The Decay Process As anyone who has taken a basic chemistry
course will remember, each element is characterized by its atomic number, defined as the number of orbital electrons or the number of protons in the nucleus of that atom. Isotopes of a given element exist because some atoms of each element, while by definition having the same number of protons, have a different number of neutrons and therefore a different nuclear weight. It should be noted that the number of electrons outside the nucleus remains the same for all isotopes of a given element, and so all isotopes of a given element are equivalent with respect to their chemical reactivity. Radioactive decay occurs when subatomic particles are released from the nucleus of an atom of a heavy isotope. This often results in the conversion of an atom of one isotope to an isotope of a different element, because the original isotope’s atomic number changes after decay. The subatomic particles released from naturally occurring radioisotopes are of three basic types: α and β particles and γ rays. An α particle is essentially the nucleus of a helium atom, or two protons plus two neutrons. It is a relatively large, heavy particle that moves slowly and usually only across short distances before it encounters some other atom with which it interacts. These particles are released from isotopes with large nuclei (atomic number >82; e.g., plutonium or uranium); such isotopes are not commonly used in biological research. In contrast to α particles, β particles are light, high-speed charged particles. Negatively charged β particles are essentially electrons of nuclear origin that are released when a neutron is converted to a proton. Release of a β particle thus changes the atomic number and elemental status of the isotope. γ radiation has both particle and wave properties; its wavelength falls within the range of X-ray wavelengths. The distinction between γ rays and X rays was made when primitive X-ray machines produced X rays with a wavelength longer than those of the γ rays produced naturally by radioisotopes. Modern X-ray machines produce a much broader spectrum of wavelengths, including γ radiation; currently this sort of X-ray radiation is termed γ when it is of nuclear origin. Unlike β-particle release, the release of γ radiation by itself produces an isotopic change rather than an elemental one; however, the resultant nuclei are unstable and often decay further, releasing β particles. The energy of all α particles and γ rays
Contributed by Jill Meisenhelder and Kentaro Semba Current Protocols in Protein Science (1995) A.2B.1-A.2B.12 Copyright © 2000 by John Wiley & Sons, Inc.
APPENDIX 2B
Laboratory Guidelines, Equipment, and Stock Solutions
A.2B.1 Supplement 1
Table A.2B.1
Nuclide
Physical Characteristics of Commonly Used Radionuclidesa
Half-life
Emission
Energy, max (MeV)
Range of emission, max 0.42 cm (air) 21.8 cm (air) 610 cm (air) 0.8 cm (water) 0.76 cm (Plexiglas) 49 cm 24.4 cm (air) 0.2 mm (lead) 165 cm (air) 2.4 cm (lead)
H C 32 b P
12.43 years 5370 years 14.3 days
β β β
0.0186 0.156 1.71
33 b
25.4 days 87.4 days 60 days 8.04 days
β β γ β γ
0.249 0.167 0.27–0.035 0.606 0.364
3
14
P S 125 c I 131 c I 35
Approx. specific activity at 100% enrichment (Ci/mg)
Atom resulting Target organ from decay
9.6 4.4 mCi/mg 285
3 2 14 7 33 16
156 43 14.2 123
33 16 35 17 125 52 130 54
He N S
S Cl Te Xe
Whole body Bone, fat Bone
Bone Testes Thyroid Thyroid
aTable
compiled based on information in Lederer et al. (1967) and Shleien (1987). shielding is Plexiglas; half-value layer measurement is 1 cm. cRecommended shielding is lead; half-value layer measurement is 0.02 mm. bRecommended
Safe Use of Radioisotopes
(measured in electron volts) is fixed, because they are of specific composition or wavelength. The energy of β particles, however, varies depending on the atom they originate from (and with concomitant release of neutrinos or antineutrinos that serve to balance the conservation of energy aspect of the decay equation). Thus there are (relatively) high-energy β particles released during the decay of 32P and low-energy β particles released when tritium (3H) decays. Isotopic decay usually involves a chain or sequence of events rather than a single loss of a particle, because the resultant, equally unstable atoms try to achieve equilibrium. During this course of decay, secondary forms of radiation can be generated that may also pose a hazard to workers. For example, when highenergy β particles released during the decay of 32P encounter the nuclei of atoms with a large atomic number, a strong interaction occurs. The β particle loses some energy in the form of a photon. Such photons are called Bremsstrahlung radiation; they are detectable using a monitor suitable for the detection of γ or X rays. Following their release, α, β, and γ emissions (as well as secondary forms of radiation) travel varying average distances at varying average speeds, depending on their energy and the density of the material through which they are moving. The distance they actually travel before encountering either the electrons or nucleus of another atom is termed their degree of penetrance. This value is expressed as an average for each type of particle. The energy of the
particles released (and therefore their potential penetrance) thus dictates what type of shielding, if any, is necessary for protection against the radiation generated by the decay of a given isotope. α, β, and γ emissions all have the potential, upon encountering an atom, to knock out its electrons, thereby creating ions. Thus, these three types of emissions are called ionizing radiation. The formation of such ions may result in the perturbation of biological processes: therein lies the danger associated with radioactivity! Table A.2B.1 provides a summary of the factors discussed above for several isotopes commonly used in biochemical research.
Measuring Radioactivity and Individual Exposure to It The radioactivity of a given substance is measured in terms of its ionizing activity. A Curie by definition is the amount of radioactive material that will produce 3.4 × 1010 disintegration (ions) per second. This, not coincidentally, happens to be the number of disintegrations that occur during the decay of one gram of radium and its decay products. Exposure to such radiation is measured as the amount of energy absorbed by the recipient, which, of course, is directly related to the potential damage such radiation may cause. One rad is the dose of radiation that will cause 100 ergs of energy to be absorbed per gram of irradiated material. Another unit commonly used to measure radiation doses is the rem; this is related to the rad but takes into account a “quality
A.2B.2 Supplement 1
Current Protocols in Protein Science
factor” based on the type of ionizing radiation being received. For β particles and γ or X rays this factor is 1; therefore, rems β equal rads β. In contrast, the quality factor associated with α particles is 20, so an exposure of one rad due to α particles would be recorded as 20 rem. The amount or dose of radiation received by materials (cells, scientists, etc.) near the source depends not only on the specific type and energy (penetrance) of the radiation being produced, but also on the subject’s distance from the source, the existence of any intervening layers of attenuating material (shielding), and the length of time spent in the vicinity of the radiation source. To best measure such doses, every person working with or around radioactivity should wear an appropriate type of radiation detection badge (in addition to carrying a portable radiation monitor that can give an immediate, approximate reading.) This is normally a requirement (not an option) for compliance with an institution’s license to use radioisotopes. Such badges are usually furnished by the safety department and collected at regular intervals for reading by a contracted company. At most institutions, the old-style film badges have been replaced with the more accurate TLDs (thermoluminescent dosimeters). These take advantage of chemicals such as calcium or lithium fluorides which, following exposure to ionizing radiation, will luminesce at temperatures below their normal thermal luminescence threshold. Different types of badges are sensitive to different types of radiation: always be sure to wear one that is appropriate for detecting exposure to the isotope being used! In most places pregnant women are asked to wear a more sensitive (and more expensive) dosimeter to better monitor their (and the developing fetus’) exposure. Most often workers will be asked to wear a radiation detection badge on the labcoat lapel in order to measure whole-body radiation. When working with 32P or 125I it is also advisable to wear a ring badge to measure exposure to the unshielded extremities (fingers). The limit set for “acceptable” exposure to whole-body radiation is several-fold less than the limit set for extremities. Nevertheless, we have found that the exposure recorded on ring badges is often significant with respect to the limit for extremities set by our institution. What is known about the effects on humans of exposure to low levels of radiation (i.e., levels which would be received when briefly handling mCi or µCi amounts of radioactivity)? Not much, for the obvious reason that direct
studies have not been undertaken. Accordingly, guidelines for exposure levels are set using extrapolations—either by extrapolating down from population statistics obtained following accidents or disasters (the Chernobyl meltdown, atomic bombings) or by extrapolating up from numbers obtained from animal experiments. Each form of extrapolation is subject to caveats, and given that predictions based on such extrapolations cannot be perfect, most health and safety personnel aim for radiation exposure levels said to be ALARA or “as low as reasonably achievable.” Further discussion of exposure limits and the statistics on which they are based may be found in B. Shleien’s health physics text (Shleien, 1987). Limiting exposure to radiation can be accomplished by adjusting several parameters of the exposure: the duration of exposure, distance from the source, and the density of the material (air, water, shielding) between the individual and the source. Time is of the essence When designing any experiment using radioactivity, every effort should be made to limit the time spent directly handling the vials or tubes containing the radioactive material. Speed should be encouraged in all manipulations, though not to the point of recklessness! Have everything needed for the experiment ready at hand before the radioactivity is introduced into the work area. Distance helps to determine dose When possible, experiments involving radioactivity should be performed in an area separate from the rest of the lab. Many institutions require that such work be performed in a designated “hot lab”; however, if many people in the laboratory routinely use radioisotopes, it is less than feasible to move them all into what is usually a smaller space. No matter where an individual is working, it is his or her responsibility to monitor the work area and ensure his or her own safety and the safety of those working nearby by using adequate shielding. Obviously, when handling the radioactive samples, it is necessary to work rapidly behind any required shielding. To protect bystanders, remember that the intensity of radiation from a source (moving through air) falls off in proportion to the square of the distance. Thus, if standing 1 foot away from a source for 5 min would result in an exposure of 45 units, standing 3 feet away for the same amount of time would result in an exposure (1/3)2 of 45 units,
Laboratory Guidelines, Equipment, and Stock Solutions
A.2B.3 Current Protocols in Protein Science
or 5 units. This factor is also relevant when considering the storage of large amounts of radioactivity, particularly 125I or 32P, as no amount of shielding can completely eliminate radiation.
Safe Use of Radioisotopes
Shielding is the key to safety As mentioned above, the energy of the particle(s) released during the decay of an isotope determines what, if any, type of shielding is appropriate. β particles released during the decay of 14C and 35S possess roughly ten times the energy of those released when 3H decays. All three β particles are of relatively low energy, do not travel very far in air, and cannot penetrate solid surfaces. No barriers are necessary for shielding against this type of β radiation. The major health threat from these isotopes occurs through their accidental ingestion, inhalation, or injection. β particles released during the decay of 32P have 10-fold higher energy than those released from 14C and pose a significant threat to workers. (One reported hazard is the potential for induction of cataracts in the unshielded eye.) The fact that these high-energy beta particles can potentially generate significant amounts of Bremsstrahlung radiation is the reason that low-density materials are used as the primary layer of shielding for 32P β radiation. Water, glass, and plastic are suitable low-density materials (as opposed to lead). Obviously water is unsuitable as a shielding layer for work on the bench, although it does a reasonable job when samples are incubating in a water bath. Shields made from a thickness of glass sufficient to stop these particles would be extremely heavy and cumbersome (as well as dangerous if dropped). Fortunately, plastic or acrylic materials—variously called Plexiglas, Perspex, or Lucite—are available for shielding against 32P β radiation. Shields as well as storage boxes constructed of various thicknesses of Plexiglas are necessary equipment in laboratories where 32P is used. When mCi amounts of 32P are used at one time it is necessary to also block the Bremsstrahlung radiation by adding a layer of high-density material (such as 4 to 6 mm lead) to the outside of the Plexiglas shield (covering the side farthest from the radioactive source). γ rays released during the decay of 125I have much higher penetrance than the β particles from 32P decay; this radiation must be stopped by very-high-density material, such as lead. Lead foil of varying thicknesses (2 to 6 mm) can be purchased in rolls and can be cut and molded to cover any container, or taped to a
Plexiglas shield (used in this instance for support). Obviously this latter arrangement has the disadvantage that it is impossible to see what one is doing through the shield. For routine shielding of manipulations involving 125I, it is useful to purchase a lead-impregnated Plexiglas shield that is transparent, albeit inevitably very heavy (as well as relatively expensive). Although it seems logical that the use of more radioactivity necessitates the use of thicker layers of shielding, it is also true that no shielding material is capable of completely stopping all radiation. When deciding how thick is “thick enough,” consult the half-value layer measurement for each type of shielding material. This number gives the thickness of a given material necessary to stop half the radiation from a source; Table A.2B.2 lists half-value layer measurements for the β particles released from isotopes commonly used in the biology laboratory. In general, 1 to 2 cm of Plexiglas and/or 0.02 mm of lead are sufficient to shield the amounts of radioactivity used in our experiments.
GENERAL PRECAUTIONS Before going on to a discussion of specific precautions to be taken with individual isotopes, a short list of general precautions to be taken with all isotopes seems pertinent: 1. Know the rules. Be sure that each individual is authorized to use each particular isotope and uses it in an authorized work area. 2. Don the appropriate apparel. Whenever working at the lab bench, it is good safety practice to wear a labcoat for protection. Disposable paper/synthetic coats of various styles are commercially available: at $4 each these may be conveniently thrown out if contaminated with radioactivity during an experiment, rather than held for decay as might be preferable with cloth coats costing about $30 each. As an alternative, disposable sleeves can be purchased and worn over those of the usual cloth coat. Other necessary accessories include radiation safety badges, gloves, and protective eyewear. It’s handy to wear two pairs of gloves at once when using radioactivity; when the outer pair becomes contaminated, it is possible to strip it off and continue working without interruption. 3. Protect the work area as well as the workers. Lab benches and the bases of any shields used should be covered with some sort of disposable, preferably absorbent, paper sheet. Underpads or diapers (the kind normally
A.2B.4 Current Protocols in Protein Science
Table A.2B.2
Shielding Radioactive Emissiona
β emitters Energy Mass (mg)/cm2 to reduce (MeV) intensity by 50% 0.1 1.0 2.0 5.0
Thickness (mm) to reduce intensity by 50% Water Glass Lead Plexiglas
1.3 48 130 400
0.013 0.48 1.3 4.0
0.005 0.192 0.52 1.6
0.0011 0.042 0.115 0.35
0.0125 0.38 1.1 4.2
γ emitters Energy (MeV)
Thickness of material (cm) to attenuate a broad beam of γ-rays by a factor of 10 Water Aluminum Iron Lead
0.5 1.0 2.0 3.0 aFrom Dawson et
54.6 70.0 76.0 89.0
20.3 24.4 32.0 37.0
6.1 8.2 11.0 12.0
1.8 3.8 5.9 6.4
al. (1986). Reprinted with permission.
used in hospitals) are convenient for this purpose. 4. Use appropriate designated equipment. It is very convenient, where use justifies the expense, to have a few adjustable pipettors dedicated for use with each particular isotope. Likewise, it is good practice to use only certain labeled centrifuges and microcentrifuge rotors for radioactive samples so that all the lab’s rotors do not become contaminated. Although such equipment should be cleaned after each use, complete decontamination is often not possible. A few pipettors or a single microcentrifuge can easily be stored (and used) behind appropriate shielding. Actually, contamination of the insides and tip ends of pipettors can be greatly reduced by using tips supplied with internal aerosol barriers; these have recently become very popular for setting up PCR reactions and are available to fit a variety of pipettors. To prevent contamination of the outside of the pipettor’s barrel, simply wrap the hand-grip in Parafilm, which can be discarded later. Several manufacturers sell disposable paper inserts for their microcentrifuges that protect the wall of the rotor chamber from contamination that might spin off the outside of sample tubes. Trying to fashion homemade liners of this sort is not recommended, as we have had disastrous experiences using laminated adhesive paper that “unstuck” during microcentrifuge spins. These liners would get caught by the rotor,
shattering sample tubes and creating an even bigger mess! 5. Know where to dispose of radioactive waste, both liquid and solid. Most institutions require that radioactive waste be segregated by isotope. This is done not only so that appropriate shielding can be placed around waste containers, but so that some waste can be allowed to decay prior to disposal through normal trash channels. With a decreasing number of radioactive waste disposal facilities able or willing to accept radioactive waste for burial (and a concomitant increase in dumping charges from those that still do) this practice of on-site decay can save an institution thousands of dollars a year in disposal charges. 6. Label your label! It is only common courtesy (as well as common sense) to alert coworkers to the existence of anything and everything radioactive that is left where they may come in contact with it! A simple piece of tape affixed to the sample box—with the investigator’s name, the amount and type of isotope, and the date written on it—should do the trick. Yellow hazard tape printed with the international symbol for radioactivity is commercially available in a variety of widths. 7. Monitor radioactivity early and often. Portable radiation detection monitors are essential equipment for every laboratory using radioactivity. No matter how much or how little radioactivity is being used, the investigator
Laboratory Guidelines, Equipment, and Stock Solutions
A.2B.5 Current Protocols in Protein Science
should keep a hand-held monitor nearby—and it should be on! Turn it on before touching any radioactivity to avoid contaminating the monitor’s switch. Use a monitor with the appropriate detection capacity (β for 35S and 32P; γ for 125I) before, during, and after all procedures. The more frequently fingers and relevant equipment are monitored, the more quickly a spill or glove contamination will be detected. Such timely detection will keep both the potential mess and the cleanup time to a minimum. Because low-energy β emitters such as 3H cannot be detected using such monitors, wipe tests of the bench and equipment used are necessary to ensure that contamination of the work area did not occur.
SPECIFIC PRECAUTIONS The following sections describe precautions to be taken with individual isotopes in specific forms. Although the sections dealing with 35Sor 32P-labeling of proteins in intact cells are presented in terms of mammalian cells, most of the instructions are also pertinent (with minimal and obvious modifications) to the labeling of proteins in other cells (e.g., bacterial, insect).
Working with 35S
Safe Use of Radioisotopes
Using 35S to label cellular proteins and proteins translated in vitro As discussed above, the β radiation generated during 35S decay is not strong enough to make barrier forms of shielding necessary. The risk associated with 35S comes primarily through its ingestion and subsequent concentration in various target organs, particularly the testes. Although willful ingestion of 35S seems unlikely, accidental or unknowing ingestion may be more common. As reported several years ago (Meisenhelder and Hunter, 1988), 35S-labeled methionine and cysteine, which are routinely used to label proteins in intact cells and by in vitro translation, break down chemically to generate a volatile radioactive component. The breakdown occurs independent of cellular metabolism. Thus the radioactive component is generated to the same extent in stock vials as in cell culture dishes. The process seems to be promoted by freezing and thawing 35S-labeled materials. The exact identity of this component is not known, although it is probably SO2 or CH3SH. What is known is that it dissolves readily in water and is absorbed by activated charcoal or copper. The amount of this volatile radioactive component released, despite stabilizers added by
the manufacturers, is about 1/8000 of the total radioactivity present. The amount of this radioactivity that a scientist is likely to inhale while using these compounds is presumably even smaller. Nevertheless, such a component can potentially contaminate a wide area because of its volatility, and also tends to concentrate in target organs. Thus, it is advisable to thaw vials of 35S-labeled amino acids in a controlled area such as a hood equipped with a charcoal filter. This charcoal filter will become quite contaminated and should be changed every few months. If such an area is not available, the stock vial should be thawed using a needle attached to a charcoal-packed syringe to vent and trap the volatile compound. Anyone who has ever added 35S-labeled amino acids to dishes of cells for even short periods knows that the incubator(s) used for such labelings quickly become highly contaminated with 35S. Such contamination is not limited to the dish itself, nor to the shelf on which the dish was placed. Rather, the radioactive component’s solubility in water allows it to circulate throughout the moist atmosphere of the incubator and contaminate all the inside surfaces of the incubator. For this reason, in laboratories where such metabolic labelings are routine, it is highly convenient to designate one incubator to be used solely for working with 35S-labeled samples. Such an incubator can be fitted with a large honeycomb-style filter, the size of an incubator shelf, made of pressed, activated charcoal. These filters are available from local air-quality-control companies. Such a filter will quickly become quite contaminated with radioactivity and should therefore be monitored and changed as necessary (usually about every three months if the incubator is used several times a week). The water used to humidify the incubator will also become quite “hot” (contaminated with radioactivity); keeping the water in a shallow glass pan on the bottom of the incubator makes it easy to change it after every use, thus preventing contamination from accumulating. Even with the charcoal filter and water as absorbents, the shelves, fan, and inner glass door of the incubator will become contaminated, as will the tray on which the cells are carried and incubated. Routine wipe tests and cleaning when necessary will help to minimize potential spread of this contamination. If such labelings are done infrequently or there is no “spare” incubator, dishes of cells can be placed in a box during incubation. This box should be made of plastic, which is generally
A.2B.6 Current Protocols in Protein Science
more easily decontaminated than metal. Along with the dishes of cells, a small sachet made of activated charcoal wrapped loosely in tissue (Kimwipes work well) should be placed in the box. If the box is sealed, it will obviously need to be gassed with the correct mixture of CO2; otherwise small holes can be incorporated into the box design to allow equilibration with the incubator’s atmosphere. In either case, the incubator used for the labeling should be carefully monitored for radioactivity after each experiment.
Working with 32P ìCi amounts of 32P The amount of 32P-labeled nucleotide used to label nucleic acid probes for northern or Southern blotting is typically under 250 µCi, and the amount of [γ-32P]ATP used for in vitro phosphorylation of proteins does not usually exceed 50 µCi for a single kinase reaction (or several hundred µCi per experiment). However, handling even these small amounts, given the time spent on such experiments, can result in an unacceptable level of exposure if proper shielding is not employed. With no intervening shielding, the dose rate 1 cm away from 1 mCi 32P is 200,000 mrads/hr; the local dose rate to basal cells resulting from a skin contamination of 1 µCi/cm2 is 9200 mrads/hr (Shleien, 1987). Such a skin contamination could be easily attained though careless pipetting and the resultant creation of an aerosol of radioactive microdroplets, because the concentration of a typical stock solution of labeled nucleotide may be 10 µCi/µl. For proper protection during this sort of experiment, besides the usual personal attire (glasses, coat, safety badges, and gloves) it is necessary to use some form of Plexiglas screen between the body and the samples (see Fig. A.2B.1, panel A). Check the level of radiation coming through the outside of the shield with a portable monitor to be sure the thickness of the Plexiglas is adequate. Hands can be shielded from some exposure by placing the sample tubes in a solid Plexiglas rack, which is also useful for transporting samples from the bench to a centrifuge or water bath (see Fig. A.2B.1, panel B). Experiments of these types often include an incubation step performed at a specific temperature, usually in a water bath. Although the water surrounding the tubes or hybridization bags will effectively stop β radiation, shielding should be added over the top of the tubes (where
there is no water)—e.g., a simple flat piece of Plexiglas. If the frequency of usage justifies the expense, an entire lid for the water bath can be constructed from Plexiglas. When hybridization reactions are performed in bags, care should be taken to monitor (and shield) the apparatus used to heat-seal the bags. The waste generated during the experiments should also be shielded. It is convenient to have a temporary waste container right on the bench. Discard pipette tips and other solid waste into a beaker lined with a plastic bag and placed behind the shield. This bag can then be emptied into the appropriate shielded laboratory waste container when the experiment is done. Liquid waste can be pipetted into a disposable tube set in a stable rack behind the shield (see Fig. A.2B.1, panel C). When radiolabeled probes or proteins must be gel-purified, it may be necessary to shield the gel apparatus during electrophoresis if the samples are particularly hot. Be advised that the electrophoresis buffer is likely to become very radioactive if the unincorporated label is allowed to run off the bottom of the gel; check with a radiation safety officer for instructions on how to dispose of such buffer. It is also prudent to check the gel plates with a portable detection monitor after the electrophoresis is completed, because they sometimes become contaminated as well. mCi amounts of 32P In order to study protein phosphorylation in intact mammalian cells, cells in tissue culture dishes are incubated in phosphate-free medium with 32P-labeled orthophosphate for a period of several hours or overnight to label the proteins. The amount of 32P used in such labelings can be substantial. Cells are normally incubated in 1 to 2 mCi of 32P/ml labeling medium; for each 6-cm dish of cells, 2.5 to 5 mCi 32P may be used. When this figure is multiplied by the number of dishes necessary per sample, and the number of different samples in each experiment, it is clear that the amount of 32P used in one experiment can easily reach 25 mCi or more. Because so much radioactivity is used in the initial labeling phase of such experiments, it is necessary for a researcher to take extra precautions in order to adequately shield him or herself and coworkers. When adding label to dishes of cells, it is important to work as rapidly as possible. An important contribution to the speed of these manipulations is to have everything that will be needed at hand before even introducing the
Laboratory Guidelines, Equipment, and Stock Solutions
A.2B.7 Current Protocols in Protein Science
A
12 in. (300 mm)
12-14 in. (300-350 mm)
18-24 in. (450-600 mm)
14 in. (350 mm)
12-15 in. (300-375 mm) L shield
T shield
B
3 in. (75 mm)
12 in. (300 mm) rack for microcentrifuge tubes
C 3.5 in. (90.5 mm)
tube holder for liquid waste
Figure A.2B.1 Plexiglas shielding for 32P. (A) Two portable shields (L and T design) made of 0.5-in. Plexiglas. Either can be used to directly shield the scientist from the radioactivity he or she is using. Turned on its side, the L-shaped shield can be used to construct two sides of a cage around a temporary work area, providing shielding for other workers directly across or to the sides of the person working with 32P. (B) Tube rack for samples in microcentrifuge tubes. (C) Tube holder for liquid waste collection.
Safe Use of Radioisotopes
label into the work area. Prepare the work area, arranging shielding and covering the bench with diapers, in advance. Set out all necessary items, including any pipettors and tips needed, a portable detection monitor, extra gloves, and a cell house (see Fig. A.2B.2, panel A). Work involving this much radioactivity should be done behind a Plexiglas shield at least 3⁄ in. (2 cm) thick; the addition of a layer of 4 lead to the outside lower section of this shield to stop Bremsstrahlung radiation is also needed. If one shield can be dedicated to this purpose at a specific location, a sheet of lead several centimeters thick can be permanently screwed to the Plexiglas (as shown in Fig. A.2B.2, panel B); however, this lead makes the shield extremely heavy and therefore less than
portable. If space constraints do not permit the existence of such a permanent labeling station, a layer or two of lead foil can be taped temporarily to the outside of the Plexiglas shield. Again, each worker should take care to shield not only him or herself, but bystanders on all sides. Handling of label should be done away from the central laboratory if possible to take maximum advantage of distance as an additional form of shielding. It is also advisable not to perform such experiments in a tissue culture room or any other room that is designed for a purpose vital to the whole laboratory. An accident involving this much 32P would seriously inconvenience future work in the area, if not make it altogether uninhabitable! If care is taken to minimize the amount of time the dish
A.2B.8 Current Protocols in Protein Science
A
door 2 in. (5 cm)
3.5 in. (9 cm) 7.5 in. (19 cm)
cell house
B
Plexiglas 0.75 -1 in. (20-25 mm) thick
lead plate 0.25 in. (4-6 mm) thick
Plexiglas 0.5 in. (12.5 mm) thick
leaded shield
C i.d. = 5.75 in. (140 mm) 1.25 in. (45 mm)
i.d. = 2.5 in. (65mm)
5.5 in. (135 mm) square block
box made of 0.5 in. (12.5 mm) thick Plexiglas rack and storage box
Figure A.2B.2 (A) Box for cell incubation (a “cell house”). (B) Stationary leaded shield. (C) Sample storage rack and box made of 0.5-in. Plexiglas. Abbreviations: i.d., interior dimension.
of cells is open when adding the label, use of a controlled air hood to prevent fungal or bacterial contamination of the cells should not be necessary. Once the label has been added to the dishes of cells, they will also need to be shielded for transport to and from the incubator and other work areas. Plexiglas boxes that are open at one end (for insertion of the dishes) and have a handle on top (for safe carrying) make ideal “cell houses” (see Figure A.2B.2, panel A). A Plexiglas door that slides into grooves at the open end is important to prevent dishes from sliding out if the box is tilted at all during transport. If this door is only two-thirds the height of the house wall, the open slot thus created will allow equilibration of the CO2 level
within the house with that in the incubator. Obviously, this slot will also allow a substantial stream of radiation to pass out of the cell house, so the house should be carried and placed in the incubator with its door facing away from the worker (and others)! Following incubation with label and any treatments or other experimental manipulations, the cells are usually lysed in some type of detergent buffer. It is probably during this lysis procedure that a worker’s hands will receive their greatest exposure to radiation, because it is necessary to directly handle the dishes over a period of several minutes. It is therefore very important to streamline this procedure and use any shielding whenever possible. If the cell lysates must be made at 4°C, as
Laboratory Guidelines, Equipment, and Stock Solutions
A.2B.9 Current Protocols in Protein Science
i.d. = 5 in. (125 mm)
i.d. = 6.75 in. (175 mm)
i.d. = 10.75 in. (275 mm) box for solid waste
Figure A.2B.3 Box for solid waste collection made of 0.5-in. Plexiglas. Abbreviation: i.d., interior dimension.
Safe Use of Radioisotopes
required by most protocols, working on a bench in a cold room is preferable to placing the dishes on a slippery bed of ice. In either case, make the lysate using the same sort of shielding (with lead if necessary) that was used when initially adding the label. Pipet the labeling medium and any solution used to rinse unincorporated radioactivity from the cells into a small tube held in a solid Plexiglas holder (shown in Fig. A.2B.1, panel C). The contents of this tube can later be poured into the appropriate liquid waste receptacle. If possible, it is a good practice to keep this high-specific-activity 32P liquid waste separate from the lower-activity waste generated in other procedures so that it can be removed from the laboratory as soon as possible following the experiment. If it is necessary to store it in the laboratory for any time, the shielding for the waste container should also include a layer of lead. The solid waste generated in the lysis part of these experiments (pipet tips, disposable pipets, cell scrapers, and dishes) is very hot and should be placed immediately into some sort of shielded container to avoid further exposure of the hands. A Plexiglas box similar in design to that in Figure A.2B.3 is convenient; placed to the side of the shield and lined with a plastic bag, it will safely hold all radioactive waste during the experiment and is light enough to be easily carried to the main laboratory waste container where the plastic bag (and its contents) can be dumped after the experiment is done. If the lid of the box protrudes an inch or so over the front wall, it can be lifted using the back of a hand, thus decreasing the possibility of contaminating it with hot gloves.
When scraping the cell lysates from the dishes, it is good practice to add them to microcentrifuge tubes that are shielded in a solid Plexiglas rack; this will help to further reduce the exposure to which the hands are subjected. At this point, the lysates are usually centrifuged at high speed (>10,000 × g) to clear them of unsolubilized cell material. Use screw-cap tubes for this clarification step, as these will contain the labeled lysate more securely than flip-top tubes, which may open during centrifugation. No matter what type of tube is used, the rotor of the centrifuge often becomes contaminated, most probably due to tiny drops of lysate (aerosol) initially present on the rim of the tubes that are spun off during centrifugation. Monitor the rotor and wipe it out after each use. The amount of 32P taken up by cells during the incubation period varies considerably, depending on the growth state of the culture as well as on the cell type and its sensitivity to radiation. This makes it difficult to predict the percentage of the radioactivity initially added to the cells that is incorporated into the cell lysate; however, this figure probably does not exceed 10%. Thus, the amount of radioactivity being handled decreases dramatically after lysis, making effective shielding much simpler. However, at least ten times more radioactivity than is usual in other sorts of experiments is still involved! It is easy to determine if the shielding is adequate—just use both β and γ portable monitors to measure the radiation coming through it. Again, be sure to check that people working nearby (including those across the bench) are also adequately shielded. It is
A.2B.10 Current Protocols in Protein Science
sometimes necessary to construct a sort of cage of Plexiglas shields around the ice bucket that contains the lysates. At the end of the day or the experiment, it may be necessary to store radioactive samples; in some experiments, it may be desirable to save the cell lysates. These very hot samples are best stored in tubes placed in solid Plexiglas racks that can then be put into Plexiglas boxes (see Fig. A.2B.2, panel C). Such boxes may be of similar construction to the cell houses described above; however, they should have a door that completely covers the opening. Be sure to check the γ radiation coming through these layers and add lead outside if necessary.
Working with 33P Using 33P-labeled nucleotides to label nucleic acid probes or proteins Several of the major companies that manufacture radiolabeled biological molecules have recently introduced nucleotides labeled with 33P (both α- and γ-labeled forms). 33P offers a clear advantage over 32P with respect to ease of handling, because the energy of the β particles it releases lies between that of 35S and 32P and thus its use does not require as many layers of Plexiglas and lead shielding as for 32P. In fact, the β radiation emitted can barely penetrate through gloves and the surface layer of skin, so the hazard associated with exposure to even millicurie amounts of 33P is thought to be insignificant (as reported in the DuPont NEN product brochure). Gel bands visualized on autoradiographs of 33P-labeled compounds are sharper than bands labeled with 32P because the lower-energy β radiation does not have the scatter associated with that from 32P. The halflife of 33P is also longer (25 days compared to 14 days for 32P). Despite its higher cost, these features have led many researchers to choose 33P-labeled nucleotides for use in experiments such as band/gel shift assays where discrimination of closely-spaced gel bands is important. The best way to determine what degree of shielding is needed when using 33P is to monitor the source using a portable β monitor and add layers of Plexiglas as necessary.
Working with 125I Using 125I to detect immune complexes (Western blots) 125I that is covalently attached to a molecule such as staphylococcal protein A is not volatile and therefore is much less hazardous than the
unbound or free form. Most institutions do not insist that work with bound 125I be performed in a hood, but shielding of the γ radiation is still necessary. Lead is a good high-density material for stopping these γ rays; its drawbacks are its weight and opacity. Commercially available shields for 125I are made of lead-impregnated Plexiglas—though heavy, these are at least seethrough. Alternatively, a piece of lead foil may be taped to a structural support, although this arrangement does not provide shielding for the head as a worker peers over the lead! Incubations of the membrane or blot with the [125I] protein A solution and subsequent washes are usually done on a shaker. For shielding during these steps, a piece of lead foil may simply be wrapped around the container. Solutions of 125I can be conveniently stored for repeated use in a rack placed in a lead box. Using 125I to label proteins or peptides in vitro Any experiments that call for the use of free, unbound 125I should be done behind a shield in a hood that contains a charcoal filter to absorb the volatile iodine. Most institutions require that such experiments be done in a special hot lab to which access is limited. Ingested or inhaled iodine is concentrated in the thyroid; a portable γ monitor should therefore be used to scan the neck and throat before beginning and after completing each experiment. Similar scans should routinely be performed on all members of any laboratory in which unbound iodine is used.
DEALING WITH ACCIDENTS Despite the best intentions and utmost caution, accidents happen! Accidents involving spills of radioactivity are particularly insidious because they can be virtually undetectable yet pose a significant threat to laboratory workers. For this reason it is best to foster a community spirit in any laboratory where radioisotopes are routinely used—a sense of cooperativity that extends from shielding each other properly to helping each other clean up when such accidents occur. The specific measures to be taken following an accident involving radioactivity naturally depend on the type and amount of the isotope involved, the chemical or biological hazards of the material it is associated with, and the physical parameters of the spill (i.e., where and onto what the isotope was “misplaced”). However, following any accident there are several immediate steps that should be taken:
Laboratory Guidelines, Equipment, and Stock Solutions
A.2B.11 Current Protocols in Protein Science
Supplement 1
1. Alert coworkers as well as Radiation Safety personnel to the fact that there has been an accident. This will give them the opportunity to shield themselves if necessary—and to help clean up as well! 2. Restrict access to and away from the site of the accident to ensure that any uncontained radioactive material is not spread around the laboratory. When leaving the site be sure to monitor the bottoms of the shoes as well as the rest of the body. 3. Take care of all contaminated personnel first, evacuating others if necessary. If anyone’s skin is contaminated, first use a portable monitor to identify specific areas of contamination. Then wipe these areas with a damp tissue to remove as much surface radioactivity as possible. Try to scrub only small areas at a time to keep the contamination localized. If the contamination is not easily removed with paper tissues, try a sponge or an abrasive pad, but be careful not to break the skin! Sometimes soaking is required: do this only after all easily removed contamination is gone and keep the soaked area to a minimum. Contaminated strands of hair can be washed (or perhaps a new hairstyle may be in order). 4. When attempting to clean any contaminated equipment, floors, benches, etc., begin by soaking up any visible radioactive liquid with
an absorbent material. Use a small amount of soap and water to clean the contaminated area, keeping the area wiped each time to a minimum to avoid smearing the contamination over an even greater surface. Many surfaces prove resistant to even Herculean cleaning efforts; in these instances the best that can be done is to remove all contamination possible and then shield whatever is left until the radioactivity decays sufficiently for safety.
LITERATURE CITED Dawson, R.M.C., Elliot, D.C., Elliott, W.H., and Jones, K.M. (eds.) 1986. Data for Biochemical Research. Alden Press, London. Lederer, C.M., Hollander, J.M., and Perlman, I. (eds.) 1967. Table of Radioisotopes, 6th edition. John Wiley & Sons, New York. Meisenhelder, J. and Hunter J. 1988. Radioactive protein-labeling techniques. Nature 335:120. Shleien, B. (ed.) 1987. Radiation Safety for Users of Radioisotopes in Research and Academic Institutions. Nuclear Lectern Associates, Olney, Maryland.
Contributed by Jill Meisenhelder and Kentaro Semba The Salk Institute La Jolla, California
Safe Use of Radioisotopes
A.2B.12 Supplement 1
Current Protocols in Protein Science
Centrifuges and Rotors
APPENDIX 2C
Centrifugation runs described in this book usually specify a relative centrifugal force (RCF; measured in × g), corresponding to a speed (in rpm) for a particular centrifuge and rotor model. As available equipment will vary from laboratory to laboratory, the investigator must be able to adapt these specifications to other centrifuges and rotors. The relationship between RCF and speed (rpm) is determined by the following equation: RCF =1.12r (rpm ⁄ 1000)2
where r is the rotating radius between the particle being centrifuged and the axis of rotation. In most cases, an accurate conversion from speed to relative centrifugal force (or vice versa) can be obtained using the maximum value of r—or rmax—equal to the distance between the axis of rotation and the bottom of the centrifuge tube as it sits in the well or bucket of the rotor. Table A.2C.1 provides rmax values for commonly used rotors manufactured by Du Pont (Sorvall), Beckman, Fisher, and IEC. There are situations (e.g., where an adapter is used to fit a smaller tube into a larger rotor well) where rmax will not accurately represent the effective rotating radius. In such cases, the manual for the rotor should be consulted to obtain the appropriate value of r. As an alternative to use of the above equation, the nomograms in Figures A.2C.1 and A.2C.2 make it possible to determine the RCF where speed and rmax are known, or the speed where RCF and rmax are known. This is done by aligning a ruler across the two known values and reading the unknown value at the point where the ruler crosses the remaining column. Figure A.2C.1 should be used for centrifuge runs <21,000 rpm, while Figure A.2C.2 should be used for faster spins. NOTE: In this manual, for spins involving microcentrifuges built to the Eppendorf standard, a shortened style of reference including only the speed (in rpm) is used. All of these instruments have approximately the same rotating radius; hence the same speed will yield the same RCF value from machine to machine. Microcentrifuge spins may also be described as at “top speed” or “maximum speed,” meaning 12,000 to 14,000 rpm, which is the maximum speed for all Eppendorf-type microcentrifuges. CAUTION: Do not exceed maximum rotor speed! For Beckman ultracentrifuges, the maximum speed for each rotor is denoted by its name, e.g., the maximum speed of the Beckman VTi 80 rotor is 80,000 rpm. This speed refers only to centrifugation of solutions below a particular allowed density, which differs among rotors (see user manual). For centrifugation of high-density solutions, rotor maximum speed can be determined as: 1 reduced rpm = rpmmax(A/B) ⁄2, where A = allowed density and B = density of solution. A = 1.7 g/ml for several vertical rotors (including VTi 80 and VTi 50), and 1.2 g/ml for several swinging-bucket rotors (including SW 55 Ti, 28, 28.1, 40 Ti, and 50.1). For gradients using heavy salts such as CsCl, particularly at low temperatures, maximum rpm should be reduced to prevent precipitation (see user manual).
Laboratory Guidelines, Equipment, and Stock Solutions Current Protocols in Protein Science (1995) A.2C.1-A.2C.5 Copyright © 2000 by John Wiley & Sons, Inc.
A.2C.1 Supplement 1
180
21,000 20,000
90,000
50,000 150
15,000
10,000 10,000 100 90
5000
80 5000 70 1000 60 500 50 200
2000
40 100
50 40 30
1000
30 20
25 Radial Distance (mm)
15.7
750
Relative centrifugal field (xg)
Rotor speed (rpm)
Figure A.2C.1 Nomogram for conversion of relative centrifugal force to rotor speed in low-speed centrifuge runs. For faster centrifugations, use Figure A.2C.2. A more precise conversion can be obtained using the equation at the beginning of this appendix. See Table A.2C.1 for rotating radii of commonly used rotors.
Centrifuges and Rotors
A.2C.2 Supplement 1
Current Protocols in Protein Science
80,000 75,000 70,000 65,000 60,000
200
1,000,000 900,000 800,000 700,000 600,000 500,000
180
400,000
160
300,000
140 120
50,000
40,000
200,000
100
70
100,000 90,000 80,000 70,000 60,000
60
50,000
90 80
30,000
40,000 50
30,000
40
20,000 20,000
30
20
10,000 9000 8000 7000 6000 5000 4000 3000 2000
10 1000 Radial Distance (mm)
Relative centrifugal field (xg)
10,000 Rotor speed (rpm)
Figure A.2C.2 Nomogram for conversion of relative centrifugal force to rotor speed in high-speed centrifuge runs. For slower centrifugations, use Figure A.2C.1. A more precise conversion can be obtained using the equation at the beginning of this appendix. See Table A.2C.1 for rotating radii of commonly used rotors.
A.2C.3 Current Protocols in Protein Science
Supplement 1
Table A.2C.1
Rotor modela
Maximum Rotating Radii for Common Rotors, Grouped by Centrifuge Model
rmax (mm)
For Sorvall centrifuge models GLC-1, GLC-2, GLC-2B, GLC-3, GLC-4, RT-6000B, T-6000, T-6000B A/S400 140 H-1000B 186 HL-4 with 50-ml bucket 180 HL-4 with 100-ml bucket 204 HL-4 with Omni-Carrier 163 M and A-384 (inner row) 91 M and A-384 (outer row) 121 SP/X and A-500 (inner row) 82 SP/X and A-500 (outer row) 123 For Sorvall centrifuge models RC-3, RC-3B, RC-3C H-2000B H-4000 and HG-4L H-6000A HL-8 with Omni-Carrier HL-8 with 50-ml bucket HL-8 with 100-ml bucket HL-2 and HL-2B LA/S400
261 230 260 221 238 247 166 140
For Sorvall centrifuge models RC-2, RC-2B, RC-5, RC-5B, RC-5C GSA GS-3 HB-4 HS-4 with 250-ml bucket SA-600 SE-12 SH-80 SM-24 (inner row) SM-24 (outer row) SS-34 SV-80 SV-288 TZ-28
145 151 147 172 129 93 101 91 110 107 101 90 95
For Sorvall ultracentrifuges T-865 T-865.1 T-875 T-880 T-1270 TFT-80.2 TFT-80.4 For Beckman GP series centrifuges GA-10 GA-24 GA-24 with adapter for 10-ml tubes Centrifuges and Rotors
91 87.1 87.1 84.7 82 65.5 60.1 123 123 108
Rotor modela
rmax (mm)
GH-3.7 (buckets) GH-3.7 (microplate carrier) GH-3.8 (buckets) GH-3.8 (microplate carrier)
204 168 204 168
For Beckman TJ-6 series centrifuges TA-10 TA-24 TA-24 with adapter for 10-ml tubes TH-4 (stainless-steel buckets) TH-4 (100-ml tube holders) TH-4 (microplate carrier) For Beckman AccuSpin AA-10 AA-24 AA-24 with adapter for 10-ml tubes AH-4
123 108 123 186 201 165 123 108 123 163
For Beckman J6 series centrifuges JR-3.2 JS-2.9 JS-3.0 JS-4.0 JS-4.2 JS-4.2SM JS-5.2 Microplate carrier (6-bucket rotors) Microplate carrier (4-bucket rotors)
206 265 254 226 254 248 226 214 192
For Beckman J2-21 series centrifuges JA-10 158 JA-14 137 JA-17 123 JA-18 132 112 JA-18.1 (25° angle) 116 JA-18.1 (45° angle) JA-20 108 JA-20.1 115 JA-21 102 JCF-Z 89 JCF-Z with small pellet core 81 JE-6B 125 JS-7.5 165 JS-13 142 JS-13.1 140 JV-20 93
continued
A.2C.4 Supplement 1
Current Protocols in Protein Science
Table A.2C.1 continued
Maximum Rotating Radii for Common Rotors, Grouped by Centrifuge Model
Rotor modela For Beckman series L7 and L8 ultracentrifuges SW 25.1 SW 28 SW 28.1 SW 30 SW 30.1 SW 40 Ti SW 41 Ti SW 50.1 SW 55 Ti SW 60 Ti SW 65 Ti Type 15 Type 19 Type 21 Type 25 Type 30 Type 30.2 Type 35 Type 40 Type 40.3 Type 42.1 Type 42.2 Ti Type 45 Ti Type 50 Type 50 Ti Type 50.2 Ti Type 50.3 Ti Type 50.4 Ti (inner row) Type 50.4 Ti (outer row) Type 55.2 Ti Type 60 Ti Type 65 Type 70 Ti Type 70.1 Ti Type 75 Ti
rmax (mm)
129.2 161.0 171.3 123.0 123.0 158.8 153.1 107.3 108.5 120.3 89.0 142.1 133.4 121.5 100.4 104.8 94.2 104.0 80.8 79.5 98.6 104 103 70.1 80.8 107.9 79.5 96.4 111.4 100.3 89.9 77.7 91.9 82.0 79.7
Rotor modela
rmax (mm)
Type 80 Ti VAC 50 VC 53 VTi 50 VTi 65 VTi 65.2 VTi 80
84.0 86.4 78.8 86.6 85.4 87.9 71.1
For Beckman Airfuge ultracentrifuge A-95 A-100/18 A-100/30 A-110 ACR-90 (2.4-ml liner) ACR-90 (3.5-ml liner) Batch rotor EM-90
17.6 14.6 16.5 14.7 11.8 13.4 14.6 13.0
For Beckman TL-100 series ultracentrifuges TLA-100 38.9 TLA 100.1 38.9 TLA-100.2 38.9 TLA-100.3 48.3 TLA-45 55.1 TLS-55 76.4 TLV-100 35.7 Miscellaneous centrifuges and rotorsb Clay Adams Dynac —c Fisher Centrific 113 Fisher Marathon 21K with 160 4-place rotor 155 IEC Clinical centrifuge with 4-place swinging-bucket rotor IEC general-purpose —c centrifuge models HN, HN-SII, and Centra-4
aSorvall centrifuges and rotors are a product of Du Pont Company Medical Products, Beckman centrifuges are a product
of Beckman Instruments, IEC centrifuges are a product of International Equipment Co., Clay Adams Dynac centrifuges are a product of Becton Dickinson Labware, and Fisher centrifuges are a product of Fisher Scientific. For ordering information see SUPPLIERSAPPENDIX. bThese instruments are often loosely referred to as “clinical,” “tabletop,” or “low-speed” centrifuges. cThese instruments accept a wide range of trunnion-ring rotors with variable rotating radii, as well as fixed-angle and swinging-bucket rotors that in turn accept a variety of adapters making it possible to spin different numbers tubes of various sizes. For instance, the commonly used IEC 958 trunnion-ring rotor may be adjusted to radii ranging from 137 to 181 mm, depending on the trunnion-ring chosen. It is therefore necessary to consult the manual for the specific system being used to obtain an accurate speed to RCF conversion.
Laboratory Guidelines, Equipment, and Stock Solutions
A.2C.5 Current Protocols in Protein Science
Supplement 1
Standard Laboratory Equipment
APPENDIX 2D
Special equipment is also itemized in the materials list of each protocol. We have not attempted to list all items required for each procedure, but rather have noted those items that might not be readily available in the laboratory or that require special preparation. Listed below are standard pieces of equipment in the modern biochemistry laboratory, i.e., items used extensively in this manual and thus not included in the individual materials lists. See SUPPLIERS APPENDIX for contact information for commercial vendors of laboratory equipment. Autoclave Balances, analytical and preparative. Bench protectors, plastic-backed (including “blue pads.”) Centrifuges low-speed (6,000 rpm) and high-speed (20,000 rpm) refrigerated centrifuges and an ultracentrifuge (20,000 to 80,000 rpm) are required for many procedures. At least one microcentrifuge that holds standard 0.5and 1.5-ml microcentrifuge tubes is essential. It is also useful to have a tabletop swingingbucket centrifuge with adapters for spinning 96-well microtiter plates. NOTE: Centrifuge speeds are provided as × g or as rpm (with example rotor models) throughout the manual. Readers should consult the nomogram in APPENDIX 2C to convert these speeds to their own rotor models.
Heating blocks thermostat-controlled metal heating blocks that hold test tubes and/or microcentrifuge tubes are very convenient for carrying out enzymatic reactions. Heat-sealable plastic bags and apparatus Hoods, chemical and microbiological. Ice maker Incubator (37°C) for growing bacteria. We recommend an incubator large enough to hold a “tissue culture” roller drum that can be used to grow 5-ml cultures in standard 18 × 150–mm test tubes. A convenient and durable tube roller is made by New Brunswick Scientific.
Computer (PC or Macintosh) and printer
Incubator/shaker(s) an enclosed shaker (such as the New Brunswick Controlled Environment Incubator Shaker) that can spin 4-liter flasks is essential for growing 1-liter E. coli cultures. A rotary shaking water bath (New Brunswick R76) is useful for growing smaller cultures in flasks.
Containers assortment of plastic and glass dishes for gel and membrane washes.
Light box for viewing gels and autoradiograms.
Darkroom and developing tanks or X- Omat automatic X-ray film developer (Kodak).
Liquid nitrogen
Cold room (4°C) or cold box.
Dry ice Filtration apparatus for collecting acid precipitates on nitrocellulose filters or membrane. Flasks, glass (e.g., Erlenmeyer, beveled shaker). Fraction collector Freezers for −20° and −70°C incubation and storage. Geiger counter Gel dryer Gel electrophoresis equipment at least one full-size horizontal apparatus and one horizontal minigel apparatus, one vertical full-size and minigel apparatus for polyacrylamide protein gels, and specialized equipment for two-dimensional protein gels as required. Current Protocols in Protein Science (1995) A.2D.1-A.2D.2 Copyright © 2000 by John Wiley & Sons, Inc.
Lyophilizers Magnetic stirrers (with heater is useful) and stir–bars. Microcentrifuge, Eppendorf-type, maximum speed 12,000 to 14,000 rpm. Microcentrifuge tubes, 1.5-ml and 0.5-ml. Microwave oven to melt agar and agarose. Mortar and pestle Paper cutter large size, for 46 × 57–cm Whatman paper sheets. Paper towels Parafilm Pasteur pipets and bulbs pH meter pH paper
Laboratory Guidelines, Equipment, and Stock Solutions
A.2D.1 Supplement 1
Pipettors that use disposable tips and dispense 1 to 1000 µl. It is best to have a set for each full-time researcher, and sets dedicated for radioactive experiments.
Scalpels and blades
Plastic wrap (e.g., Saran Wrap).
Seal-A-Meal bag sealer or equivalent.
Polaroid camera and UV transilluminator for taking photographs of stained gels.
Shakers, orbital and platform, room temperature or 37°C.
Policemen, rubber or plastic
Spectrophotometer UV and visible.
Power supplies 300-volt power supplies are sufficient for polyacrylamide gels; a 2000- to 3000-volt power supply is needed for some applications.
Speedvac evaporator (Savant). Tissue culture equipment CO2 incubator, phase contrast microscope, liquid nitrogen storage container, and laminar flow hood.
Racks, to hold test tubes and microcentrifuge tubes.
UV light sources, long- and short-wave, stationary or hand-held.
Radiation shield (Lucite or Plexiglas).
Vortex mixers
Radioactive waste containers, for liquid and solid waste.
Water baths at least two with 80°C capacity.
Scintillation counter Scissors
Refrigerator, 4°C.
Water purification equipment such as MilliQ system (Millipore) or its equivalent.
Safety glasses
X-ray film cassettes and intensifying screens
Standard Laboratory Equipment
A.2D.2 Current Protocols in Protein Science
Commonly Used Reagents
APPENDIX 2E
BUFFERS AND STOCK SOLUTIONS This collection describes the preparation of buffers and reagents used in the manipulation of proteins (see Table A.2E.1). When preparing solutions, use water purified using a Milli-Q apparatus (Millipore) or equivalent and reagents of the highest grade available. Sterilization—by filtration through a 0.22-µm filter—is recommended for most applications. CAUTION: Handle strong acids and bases carefully. Ammonium acetate, 10 M Dissolve 385.4 g ammonium acetate in 150 ml H2O Add H2O to 500 ml Dithiothreitol (DTT), 1 M Dissolve 15.45 g DTT in 100 ml H2O Store at −20°C EDTA (ethylenediamine tetraacetic acid), 0.5 M (pH 8.0) Dissolve 186.1 g Na2EDTA⋅2H2O in 700 ml H2O Adjust pH to 8.0 with 10 M NaOH (∼50 ml) Add H2O to 1 liter Begin titrating before the sample is completely dissolved. EDTA, even in the disodium salt form, is difficult to dissolve at this concentration unless the pH is increased to between 7 and 8.
Ethidium bromide, 10 mg/ml Dissolve 0.2 g ethidium bromide in 20 ml H2O Mix well and store at 4°C in dark CAUTION: Ethidium bromide is a mutagen and must be handled carefully.
Table A.2E.1
Molarities and Specific Gravities of Concentrated Acids and Bases
Acid/base Acids Acetic acid (glacial) Formic acid Hydrochloric acid Nitric acid Perchloric acid Phosphoric acid Sulfuric acid Bases Ammonium hydroxide Potassium hydroxide Potassium hydroxide Sodium hydroxide
Molecular weight
% by weight
Molarity (approx.)
1 M solution (ml/liter)
Specific gravity
60.05
99.6
17.4
57.5
1.05
46.03
98.00 98.07
90 98 36 70 60 72 85 98
23.6 25.9 11.6 15.7 9.2 12.2 14.7 18.3
42.4 38.5 85.9 63.7 108.8 82.1 67.8 54.5
1.205 1.22 1.18 1.42 1.54 1.70 1.70 1.835
35.0
28
14.8
67.6
0.90
56.11 56.11 40.0
45 50 50
11.6 13.4 19.1
82.2 74.6 52.4
1.447 1.51 1.53
36.46 63.01 100.46
Current Protocols in Protein Science (1998) A.2E.1-A.2E.5 Copyright © 1998 by John Wiley & Sons, Inc.
Laboratory Guidelines, Equipment, and Stock Solutions
A.2E.1 Supplement 12
HCl, 1 M Mix in the following order: 914.1 ml H2O 85.9 ml concentrated HCl KCl, 1 M 74.6 g KCl H2O to 1 liter MgCl2 , 1 M 20.3 g MgCl2⋅6H2O H2O to 100 ml MgSO4 , 1 M 24.6 g MgSO4⋅7H2O H2O to 100 ml NaCl, 5 M 292 g NaCl H2O to 1 liter NaOH, 10 M Mix in the following order: 476 ml H2O 524 ml 50% (v/v) NaOH Alternatively: Dissolve 400 g NaOH in 450 ml H2O Add H2O to 1 liter Phenol, buffered 8-hydroxyquinoline Liquefied phenol, redistilled 50 mM Tris base (unadjusted pH ∼10.5) 50 mM Tris⋅Cl, pH 8.0 (see recipe) TE buffer, pH 8.0 (see recipe) Add 0.5 g of 8-hydroxyquinoline to a 2-liter glass beaker containing a stir bar. Gently pour in 500 ml of liquefied phenol or melted crystals of redistilled phenol (melted in a water bath at 65°C). The phenol will turn yellow due to the 8-hydroxyquinoline, which is added as an antioxidant. Add 500 ml of 50 mM Tris base. Cover the beaker with aluminum foil and stir 10 min at low speed with magnetic stirrer at room temperature. Let phases separate at room temperature. Gently decant the top (aqueous) phase into a suitable waste receptacle. Remove what cannot be decanted with a 25-ml glass pipet and a suction bulb. Add 500 ml of 50 mM Tris⋅Cl, pH 8.0. Repeat equilibration with 500 ml of 50 mM Tris⋅Cl, pH 8.0, twice. The pH of the phenol phase can be checked with indicator paper and should be 8.0. If it is not, repeat equilibration until this pH is obtained. Add 250 ml of 50 mM Tris⋅Cl, pH 8.0, or TE buffer, pH 8.0, and store at 4°C in brown glass bottles or clear glass bottles wrapped in aluminum foil. Phenol prepared with 8-hydroxyquinoline as an antioxidant can be stored ≤2 months at 4°C. Phenol must be redistilled before use, because oxidation products of phenol can damage and introduce breaks into nucleic acid chains. Redistilled phenol is commercially available. Regardless of the source, the phenol must be buffered before use.
Commonly Used Reagents
CAUTION: Phenol can cause severe burns to skin and damage clothing. Gloves, safety glasses, and a lab coat should be worn whenever working with phenol, and all manipulations should be carried out in a fume hood. A glass receptacle should be available exclusively for disposing of used phenol and chloroform.
A.2E.2 Supplement 12
Current Protocols in Protein Science
Table A.2E.2 Preparation of 0.1 M Sodium and Potassium Acetate Buffersa
Desired pH
Solution A (ml)
Solution B (ml)
3.6 3.8 4.0 4.2 4.4 4.6 4.8 5.0 5.2 5.4 5.6
46.3 44.0 41.0 36.8 30.5 25.5 20.0 14.8 10.5 8.8 4.8
3.7 6.0 9.0 13.2 19.5 24.5 30.0 35.2 39.5 41.2 45.2
aAdapted
by permission from CRC, 1975.
Table A.2E.3 Preparation of 0.1 M Sodium and Potassium Phosphate Buffersa
Desired pH 5.7 5.8 5.9 6.0 6.1 6.2 6.3 6.4 6.5 6.6 6.7 6.8 aAdapted
Solution A (ml) 93.5 92.0 90.0 87.7 85.0 81.5 77.5 73.5 68.5 62.5 56.5 51.0
Solution B (ml)
Desired pH
6.5 8.0 10.0 12.3 15.0 18.5 22.5 26.5 31.5 37.5 43.5 49.0
6.9 7.0 7.1 7.2 7.3 7.4 7.5 7.6 7.7 7.8 7.9 8.0
Solution A (ml) 45.0 39.0 33.0 28.0 23.0 19.0 16.0 13.0 10.5 8.5 7.0 5.3
Solution B (ml) 55.0 61.0 67.0 72.0 77.0 81.0 84.0 87.0 90.5 91.5 93.0 94.7
by permission from CRC, 1975.
Phosphate-buffered saline (PBS) 10× × stock solution, 1 liter: 80 g NaCl 2 g KCl 11.5 g Na2HPO4⋅7H2O 2 g KH2PO4
Working solution, pH ∼7.3: 137 mM NaCl 2.7 mM KCl 4.3 mM Na2HPO4⋅7H2O 1.4 mM KH2PO4
Potassium acetate buffer, 0.1 M Solution A: 11.55 ml glacial acetic acid/liter (0.2 M) Solution B: 19.6 g potassium acetate (KC2H3O2)/liter (0.2 M) Referring to Table A.2E.2 for desired pH, mix the indicated volumes of solutions A and B, then dilute with H2O to 100 ml. This may be made as a 5- or 10-fold concentrate by scaling up the amount of potassium acetate in the same volume. Acetate buffers show concentration-dependent pH changes, so check concentrate pH by diluting an aliquot to the final concentration. To prepare buffers with pH intermediate between the points listed in Table A.2E.2, prepare closest higher pH, then titrate with solution A.
Laboratory Guidelines, Equipment, and Stock Solutions
A.2E.3 Current Protocols in Protein Science
Supplement 12
Potassium phosphate buffer, 0.1 M Solution A: 27.2 g KH2PO4 per liter (0.2 M) Solution B: 45.6 g K2HPO4⋅3H2O per liter (0.2 M) Referring to Table A.2E.3 for desired pH, mix the indicated volumes of solutions A and B, then dilute with H2O to 200 ml. This may be made as a 5- or 10-fold concentrate by scaling up the amount of potassium phosphate in the same volume. Phosphate buffers show concentration-dependent pH changes, so check concentrate pH by diluting an aliquot to the final concentration.
SDS, 20% Dissolve 20 g SDS (sodium dodecyl sulfate or sodium lauryl sulfate) in H2O to 100 ml total with stirring (it may be necessary to heat the solution slightly to fully dissolve the powder). Filter sterilize using a 0.45-µm filter. Sodium acetate, 3 M Dissolve 408 g sodium acetate⋅3H2O in 800 ml H2O Add H2O to 1 liter Adjust pH to 4.8 or 5.2 (as desired) with 3 M acetic acid Sodium acetate buffer, 0.1 M Solution A: 11.55 ml glacial acetic acid/liter (0.2 M) Solution B: 27.2 g sodium acetate (NaC2H3O2⋅3H2O)/liter (0.2 M) Referring to Table A.2E.2 for desired pH, mix the indicated volumes of solutions A and B, then dilute with H2O to 100 ml. (See Potassium acetate buffer recipe for further details.) Sodium phosphate buffer, 0.1 M Solution A: 27.6 g NaH2PO4⋅H2O per liter (0.2 M) Solution B: 53.65 g Na2HPO4⋅7H2O per liter (0.2 M) Referring to Table A.2E.3 for desired pH, mix the indicated volumes of solutions A and B, then dilute with H2O to 200 ml. (See Potassium phosphate buffer recipe for further details.) SSC (sodium chloride/sodium citrate), 20× 3 M NaCl (175 g/liter) 0.3 M Na3citrate⋅2H2O (88 g/liter) Adjust pH to 7.0 with 1 M HCl TAE (Tris/acetate/EDTA) electrophoresis buffer Working solution, pH ∼8.5: 50× stock solution: 40 mM Tris⋅acetate 242 g Tris base 2 mM Na2EDTA⋅2H2O 57.1 ml glacial acetic acid 37.2 g Na2EDTA⋅2H2O H2O to 1 liter TBE (Tris/borate/EDTA) electrophoresis buffer 10× stock solution, 1 liter: 108 g Tris base (890 mM) 55 g boric acid (890 mM) 40 ml 0.5 M EDTA, pH 8.0 (see recipe; 20 mM) TE buffer 10 mM Tris⋅Cl, pH 7.4, 7.5, or 8.0 (or other pH; see recipe) 1 mM EDTA, pH 8.0 Commonly Used Reagents
A.2E.4 Supplement 12
Current Protocols in Protein Science
Tris⋅Cl [tris(hydroxymethyl)aminomethane], 1 M Dissolve 121 g Tris base in 800 ml H2O Adjust to desired pH with concentrated HCl Mix and add H2O to 1 liter NOTE: The pH of Tris buffers changes significantly with temperature, decreasing approximately 0.028 pH units per 1°C. Tris-buffered solutions should be adjusted to the desired pH at the temperature at which they will be used. Because the pKa of Tris is 8.08, Tris should not be used as a buffer below pH ∼7.2 or above pH ∼9.0. LITERATURE CITED Chemical Rubber Company. 1975. CRC Handbook of Biochemistry and Molecular Biology, Physical and Chemical Data, 3rd ed., Vol. 1. CRC Press, Boca Raton, Fla.
Laboratory Guidelines, Equipment, and Stock Solutions
A.2E.5 Current Protocols in Protein Science
Supplement 12
COMMONLY USED TECHNIQUES
APPENDIX 3
Use of Protein Folding Reagents
APPENDIX 3A
DENATURANTS Chemical agents that denature proteins are used to extract proteins from inclusion bodies as well as to study the conformation of purified proteins (Pace, 1986). The most common denaturants are guanidine hydrochloride (guanidine⋅HCl) and urea. These reagents and methods for their purification are described in this section.
Guanidine Hydrochloride High-purity guanidine⋅HCl (also referred to as guanidinium chloride) can be purchased from several vendors as a crystalline solid or concentrated solution (usually 8 M) in water. Because dissolution of the salt is an endothermic process, when preparing concentrated solutions from the solid salt it is convenient to heat the mixture (in a glass container) in a domestic microwave oven. When the best grades of commercial guanidine⋅HCl (>99% pure) are used, a 6 M solution will be clear and colorless. Concentrated solutions made from practical-grade guanidine⋅HCl (usually 80% to 90% cheaper) may appear slightly hazy and be colored light brown. For large-scale preparative work, where the purchase of sufficient high-purity reagent would be prohibitive, solutions of practical-grade guanidine⋅HCl can be cleaned up adequately by filtration with 0.45- to 0.5-µm filters followed by decolorization with activated charcoal. Other applications, such as folding experiments (especially those involving spectroscopic monitoring), require higherquality reagent; in such cases it is necessary to recrystallize the salt or purchase the ultrapure grade. Methods for purifying guanidine⋅HCl by recrystallization have been described by Nozaki (1972). Table A.3A.1 Solutions
Because guanidine⋅HCl is hydroscopic, for critical studies, such as conformational analysis, accurate measurements of molarity should be made using a refractometer (e.g., the Abbe3L refractometer from Milton Roy; see SUPPLIERS APPENDIX). Using Table A.3A.1 as a guideline for quantities, guanidine⋅HCl and water (or buffer) can be weighed in a container to give the approximate molarities indicated. A table of the refractive indexes of guanidine⋅HCl solutions from 0.057 M to 8.51 M (in increments of ∼0.06 M) at 25°C is provided by Nozaki (1972). There have been no reports, at least to the author’s knowledge, of specific chemical modification of proteins resulting from exposure to guanidine⋅HCl or the contaminants found in the commercially purchased forms. However, the common contaminant ammeline, which is characterized by strong UV absorbance, may inhibit certain enzymes (Nozaki and Tanford, 1967; Nozaki, 1972, and references therein). Guanidine⋅HCl solutions are stable at room temperature for several days (for longer periods, store at 4°C). Solutions are stable over normal working pH ranges—i.e., pH 2.0 to 10.5. At alkaline pH (>11), biguanidine may form (Nozaki and Tanford, 1967).
Urea High-purity (>99%) urea can be purchased from many different vendors and is only approximately twice as expensive as practicalgrade material. Urea solutions slowly decompose to form ammonium cyanate, which reacts readily with various protein functional groups. An 8 M solution of urea incubated at room temperature >pH 7.0 will contain ∼4 mM cyanate after 7 days and ∼20 mM at equilibrium.
Properties of Urea and Guanidine⋅HCl
Property Molecular weight Solubility (at 25°C)a Melting point (°C) A260 (6 M in water)
Urea 60.06 10.49 M 133-137 <0.06
Guanidine⋅HCl 95.53 8.54 M 186.5-188.5 <0.03
aSolubilities at 5°C: urea, ∼8 M; guanidine⋅HCl, ≥8 M.
Contributed by Paul T. Wingfield Current Protocols in Protein Science (1995) A.3A.1-A.3A.4 Copyright © 2000 by John Wiley & Sons, Inc.
Commonly Used Techniques
A.3A.1 CPPS
Table A.3A.2
Preparation of Urea and Guanidine⋅HCl Solutions
Urea Molarity 1M 2M 3M 4M 5M 6M 7M 8M 9M 10 M
Quantitya 0.63 1.32 2.09 2.93 3.88 4.95 6.16 7.54 9.15 11.02
Vol (ml)b 10.52 11.04 11.61 12.24 12.96 13.76 14.68 15.73 16.95 18.38
Guanidine⋅HCl Concentration (g/liter)c 60.06 120.12 180.18 240.24 300.30 360.36 420.42 480.48 540.54 600.60
Quantitya
Vol (ml)b
1.03 2.23 3.65 5.36 7.45 10.09 13.51 18.14 — —
10.82 11.71 12.76 14.05 15.63 17.63 20.23 23.77 — —
Concentration (g/liter)c 95.53 191.06 286.59 382.12 477.65 573.18 668.71 764.24 — —
aGiven weight of reagent added to 10 g water to generate a solution of the indicated molarity. bThe final volume resulting from the addition of indicated amount of reagent to 10 g water was determined using the
specific gravity of the solution calculated as described by Kawahara and Tanford (1966). Volume estimates can also be made by assuming 1 g of urea or guanidine⋅HCl increases the volume by 0.763 ml (i.e., the partial specific volume = 0.763). The density of water at 25°C is 0.9971 (10 g = 10.029 ml). cWeight of reagent made up with water or buffer to a final volume of 1 liter.
Protein Folding Reagents
Formation of cyanate occurs faster at higher temperatures (e.g., equilibrium is reached in 30 min at 100°C), and it may therefore be helpful to keep the solution cold. Cyanate reacts readily with the amino, sulfhydryl, imidazole, tyrosyl, and carboxyl groups of proteins. Only the reaction with amino groups results in stable products at alkaline pH (Means and Feeney, 1971). The carbamylation of α-amino (N-terminal) and ε-amino groups (lysine side chains) results in the formation of a neutral product, homocitrulline; hence there is a loss of one positive charge for each residue modified. The standard method for removing cyanate is deionization using a mixed-bed ion-exchange resin (e.g., AG 501-X8, Bio-Rad; see SUPPLIERS APPENDIX). The batch method (adding resin to the solution, incubating, then filtering) is recommended for volumes <1 liter, whereas the column method (using resin packed in a chromatography column) is best for larger volumes (see Bio-Rad Bulletin 1825 and UNIT 8.2). Cyanate removal can be monitored by determining the solution conductivity; there are also simple specific assays available (Means and Feeney, 1972). For a detailed purification method, involving both recrystallization and deionization, see Prakash et al. (1981). Cyanate can also be removed by acidification to pH 2 with HCl, followed by a 1-hr incubation during which the cyanate breaks down to carbon diox-
ide and ammonium ions, and neutralization with Tris base. The solution can then be used as is. For most applications it is recommended that solutions be made with ultrapure urea and used within 24 hr. A scavenger should be included to react with slowly forming cyanate; buffers containing primary amines, such as Tris and glycine, are suitable for this purpose. It should be noted that urea may increase the measured pH of aqueous solutions (see Bull et al., 1964). As with guanidine⋅HCl, refractometry can be used to accurately determine the concentration of solutions, although solutions prepared by weight (Table A.3A.2) are normally accurate enough for most applications (Pace, 1986).
Effectiveness According to Pace (1986) and references therein, guanidine⋅HCl is generally 1.5 to 2.5 times more effective per mole as a protein denaturant than urea.
Other Reagents Guanidine thiocyanate is another denaturant of interest that is often used to elute proteins from immunoaffinity columns. Although guanidine thiocyanate is rarely used in preparative folding of recombinant proteins or in conformational analysis of proteins in general, it is
A.3A.2 Current Protocols in Protein Science
Table A.3A.3
Reduced and Oxidized Sulfhydryl Reagents
Absorbance (nm) (liter/mol/cm)e
SHf pKa
E′o (mV)g
192-195
260 (5.8) 280 (2.3)
8.63 (α) 9.45 (β)
−240
612.63
178
250 (342) 260 (271) 280 (111) 300 (30)
2-MEb HEDc
78.13 154.25
— 25-27
DTT
154.25
42-44
>260 (≈ 0, if pure) 245 λmax (380) 260 (310) 280 (200) >270 (≈ 0)
Oxidized DTTd
152.20
132
Reagent
MWa
MP (°C)
GSH
307.33
GSSG
283 λmax (273) 310 (110)
—
9.5 —
8.3 9.5
−330
—
aAbbreviations: DTT, dithiothreitol; GSH, reduced glutathione (γ-L-glutamyl-L-cysteinylglycine, anionic thiol);
GSSG, oxidized glutathione; HED, 2-hydroxyethyl disulfide; MP, melting point; MW, molecular weight. bPure liquid is 14.3 M (density = 1.114 g/ml) with a boiling point of 157° − 158°C. cLiquid is 8.2 M (density = 1.261 g /ml). dOxidized DTT (dihydroxy-1,2-dithiane) can be prepared from DTT by oxidation with ferricyanide (Cleland,
1964). Commercial material can be purified according to Creighton (1984). eThe values in parenthesis are the molar absorbencies at the wavelengths indicated. fIn amino thiols, the pK of the amine and thiol groups overlap; relevant pK ’s are indicated as α and β. a a gThe standard redox potential (at pH 7, 25°C); refers to the reduction potential for thiol-disulfide pairs: 2RSH = R-S-S-R + 2H++ + 2e−. Note that the lower the E′o, the stronger the reductant—e.g., DTT is a better reductant
than GSH.
2.5 to 3.5 times more effective than guanidine⋅HCl, and is available (albeit expensive) in high purity. Other chaotropic reagents, mainly inorganic salts, are briefly reviewed and referenced in the Commentary section of UNIT 6.3. As there are few (if any) reports in the literature regarding the use of these reagents in solubilizing inclusion bodies, detailed discussion of their properties is not included. Future updates of this section will, however, address any important new reagents.
SULFHYDRYL REAGENTS AND OXIDO-SHUFFLING SYSTEMS Sulfhydryl reagents (reducing agents) are used to prevent the oxidation of protein thiol groups. When proteins are extracted from inclusion bodies it is standard practice to include a reducing agent to prevent random oxidation of cysteine residues. Denatured proteins are subsequently folded under conditions that fa-
vor native disulfide formation. This process, referred to as oxidative folding, is carried out using “oxido-shuffling” (or oxidative regeneration) systems consisting of low-molecularweight thiol/disulfide pairs (UNITS 6.4 & 6.5). The thiols and disulfides commonly used to reduce and oxidize recombinant proteins are shown in Table A.3A.3 and discussed in this section. Further details on these and other reducing agents are reviewed by Jocelyn (1987).
Dithiothreitol Dithiothreitol (DTT), also known as Cleland’s reagent, is the preferred reductant for protein reduction. DTT and its isomer dithioerythritol (DTE)—which can also be used as a protein reductant—have similar properties (and prices), except that at lower pH the former is more effective. This is because the pKa values for DTE are higher (9.0 and 9.9) than those of DTT (8.3 and 9.5); hence, a higher pH is required to ionize DTE to its functional form.
Commonly Used Techniques
A.3A.3 Current Protocols in Protein Science
The reducing capacity of both reagents increases greatly over the pH range 7 to 9.5. On the other hand, the reagents are quenched by acidification to
2-Mercaptoethanol 2-Mercaptoethanol (2-ME) is also commonly used as a protein reducing agent. 2-ME is a volatile liquid that should be stored at 4°C and dispensed under a fume hood. A 100-mM stock solution of 2-ME can be made by diluting 6.99 ml of the reagent with solvent to 1 liter. 2-ME is not as potent a reductant as DTT, as it has a higher pKa (Table A.3A.3), and is used in the concentration range of 5 to 100 mM. Air oxidation of 2-ME is best monitored by measuring the increase in absorbance at 260 nm (Table A.3A.3). Freshly opened bottles contain 0.1% to 6.7% of the oxidation product 2-hydroxyethyl disulfide (HED; Wetlaufer et al., 1987).
Glutathione Glutathione (GSH) and its oxidized form GSSG are present in most cells at concentrations of 1 to 10 mM. In E. coli, the ratio of GSH/GSSG is 50:1 to 200:1 (Hwang et al., 1992), and the system is responsible for maintaining cytoplasmic proteins in a reduced state. GSH is not normally used as a reductant in analytical protein chemistry—its main utility is as a component of oxidative redox buffers (see discussion of Oxido-Shuffling Systems). However, GSH can be used to maintain unfolded protein in inclusion-body extracts in the reduced state (analogous to its role in vivo). Stock solutions of 0.1 M GSH can be prepared in water or the reagent can be directly added to buffers. In either case the pH should be adjusted with base to the required pH.
Oxido-Shuffling Systems Protein Folding Reagents
DTT (for reviews, see Creighton, 1984, and Wetlaufler, 1984), which are shown in Table A.3A.3. All six reagents are commercially available (the scarcest, HED, can be obtained from Aldrich; see SUPPLIERS APPENDIX) and are normally used as purchased, without further purification.
LITERATURE CITED Bio-Rad Bulletin 1825. Sample preparation: A guide to methods and applications. Bio-Rad Laboratories, Hercules, Calif. Bull, H.R., Breese, K., Ferguson, G.L., and Swenson, G.L. 1964. The pH of urea solutions. Arch. Biochem. Biophys. 104:297-304. Cleland, W.W. 1964. Dithiothreitol, a new protective reagent for SH groups. Biochemistry 3:480-482. Creighton, T.E. 1984. Disulfide bond formation in proteins. Methods Enzymol. 107:305-329. Hwang, C., Sinskey, A.J., and Lodish, H.F. 1992. Oxidized redox state of glutathione in the endoplasmic reticulum. Science 257:1496-1502. Jocelyn, P.C. 1987. Chemical reduction of disulfides. Methods Enzymol. 143:246-264. Kawahara, K. and Tanford, C. 1966. Viscosity and density of aqueous solutions of urea and guanidine hydrochloride. J. Biol. Chem. 241:3228-3232. Means, G.M. and Feeney, R.E. 1971. Chemical Modification of Proteins. Holden-Day, San Francisco. Nozaki, Y. 1972. The preparation of guanidine hydrochoride. Methods Enzymol. 26:43-50. Nozaki, Y. and Tanford, C. 1967. Acid-base titrations in concentrated guanidine hydrochloride. Dissociation constants of the guanidinium ion and some amino acids. J. Amer. Chem. Soc. 89:736742. Pace, C.N. 1986. Determination and analysis of urea and guanidine hydrochloride denaturation curves. Methods Enzymol. 131:266-280. Prakash, V., Loucheux, C., Scheufele, S., Gorbunoff, M.J., and Timasheff, S.N. 1981. Interactions of proteins with solvent components in 8 M urea. Arch. Biochem. Biophys. 210:455-464. Wetlaufer, D.B. 1984. Nonenzymatic formation and isomerization of protein disulfides. Methods Enzymol. 107:301-304. Wetlaufer, D.B., Branca, P.A., and Chen, G.X. 1987. The oxidative folding of proteins by disulfide plus thiol does not correlate with redox potential. Protein Eng. 1: 141-146.
Contributed by Paul T. Wingfield National Institutes of Health Bethesda, Maryland
The commonly used oxido-shuffling systems used in folding recombinant proteins are GSH/GSSG, 2-ME/HED, and DTT/oxidized
A.3A.4 Current Protocols in Protein Science
Dialysis
APPENDIX 3B
Conventional dialysis separates small molecules from large molecules by allowing diffusion of only the small molecules through selectively permeable membranes. Dialysis is usually used to change the salt (small-molecule) composition of a macromolecule-containing solution. The solution to be dialyzed is placed in a sealed dialysis membrane and immersed in a selected buffer; small solute molecules then equilibrate between the sample and the dialysate. Concomitant with the movement of small solutes across the membrane, however, is the movement of solvent in the opposite direction. This can result in some sample dilution (usually <50%). This section describes dialysis of large- and small-volume samples using cellulose membranes with pore sizes designed to exclude molecules above a selected molecular weight (Basic and Alternate Protocols). A Support Protocol describes preparation of membranes for dialysis. LARGE-VOLUME DIALYSIS This protocol describes the use of membranes, prepared using the support protocol, for dialysis of samples in large, easily handled volumes, typically 0.1 to 500 ml.
BASIC PROTOCOL
Materials Dialysis membrane (see Support Protocol) Clamps (Spectrapor Closures, Spectrum, or equivalent) Macromolecule-containing sample to be dialyzed Appropriate dialysis buffer 1. Remove dialysis membrane from ethanol storage solution and rinse with distilled water. Secure clamp to one end of the membrane or knot one end with double-knots. Always use gloves to handle the dialysis membrane because the membrane is susceptible to cellulolytic microorganisms. See support protocol for a discussion of commercially available membranes and preparation and storage procedures. Clamps are less likely to leak than knots, but either type of closure should be carefully tested as described in steps 2 and 3.
2. Fill membrane with water or buffer, hold the unclamped end closed, and squeeze membrane. A fine spray of liquid indicates a pinhole in the membrane; discard and try a new membrane. Pinholes are rare but cause traumatic sample loss. Hard squeezing will cause some liquid to seep from the bag; this is normal.
3. Replace the water or buffer in dialysis membrane with the macromolecule-containing sample and clamp the open end. Again, squeeze to check the integrity of the membrane and clamps. If dialyzing a concentrated or high-salt sample, leave some space in the clamped membrane; there will be a net flow of water into the sample, and if sufficient pressure builds up the membrane can burst.
4. Immerse dialysis membrane in a beaker or flask containing a large volume (relative to the sample) of the desired buffer. Dialyze at least 3 hr at the desired temperature with gentle stirring of the buffer. Dialysis rates are dependent on membrane pore size, sample viscosity, and ratio of membrane surface to sample volume. Temperature has little effect on dialysis rate, but low Contributed by Louis Zumstein Current Protocols in Protein Science (1995) A.3B.1-A.3B.4 Copyright © 2000 by John Wiley & Sons, Inc.
Commonly Used Techniques
A.3B.1 CPPS
temperatures (i.e., 4°C) are usually chosen to enhance macromolecule stability. Common salts will equilibrate across a 15,000-MWCO membrane (see Support Protocol) in ∼3 hr with stirring. Some compounds, especially detergents above their critical micelle concentration (CMC), dialyze very slowly.
5. Change dialysis buffer as necessary. Usually two to three dialysis buffer changes are sufficient. For example, when 100 mM Tris⋅Cl is removed from a protein for sequence analysis or other amino-reactive chemistry, two equilibriums against a 1000-fold volume excess of buffer will decrease the Tris concentration 106-fold, to 100 nM; three dialyses will decrease the concentration to 100 pM.
6. Remove dialysis membrane from the buffer. Hold the membrane vertically and remove excess buffer trapped in end of membrane outside upper clamp. Release upper clamp and remove the sample with a Pasteur pipet. ALTERNATE PROTOCOL
SMALL-VOLUME DIALYSIS For solution volumes less than ∼100 µl, the use of dialysis membrane as described above can result in unacceptable sample loss. The method described below can easily dialyze volumes as small as 10 µl. The sample is held in a 0.5-ml microcentrifuge tube, with dialysis membrane covering the open end of the tube. Additional Materials 0.5-ml microcentrifuge tube Cork borer 1. Cut a hole in the cap of a 0.5-ml microcentrifuge tube using a cork borer. Be sure that there are no rough edges on the inside of the cap (which will be in contact with the dialysis membrane). The success of this protocol is dependent on a tight-fitting cap, so make sure the cap is well-seated. On the other hand, an excessively tight-fitting cap can rip the membrane when it is inserted. Sarstedt tubes work well. Alternatively, some researchers use a microcentrifuge tube without a cap. The sample is placed in the microcentrifuge tube, a dialysis membrane is placed over the top of the tube, and the membrane is secured with a rubber band. With this method, however, there is a risk that a small sample—e.g., 10 to 20 µl—may be lost around the edge of the tube.
2. Place sample in the microcentrifuge tube, cover tube opening with dialysis membrane, and snap on the tube cap in which a hole has been cut. This places a fair amount of stress on the dialysis membrane, so use a fairly thick membrane and make sure there are no rough edges on the cap.
3. To get the sample in contact with the dialysis membrane, gently centrifuge the microcentrifuge tube in an inverted position in a tabletop centrifuge. 4. Keep the tube inverted, immerse it in dialysis buffer, and anchor it so that it will remain inverted and immersed. This is easily done by inserting tubes in a styrofoam or foam sheet (e.g., Fisherbrand foam microcentrifuge tube rack) and placing the inverted tubes on the surface of the dialysis buffer.
5. Use a bent Pasteur pipet to blow out any air bubbles trapped under the cap to allow dialysis buffer to contact the membrane. Dialysis
A.3B.2 Current Protocols in Protein Science
This step is usually not necessary if the membrane/rubber band method is used (see step 1 annotation).
6. Stir the dialysis buffer and dialyze at an appropriate temperature for at least 3 hr (see Basic Protocol, step 4). 7. To recover the sample, remove microcentrifuge tube from the buffer and centrifuge briefly right-side-up. Commercially available alternatives to this method include use of individual hollow-fiber filters from Spectrum (with sample capacities of 1 to 140 µl), and many different microdialysis machines, both single-sample and multisample (Spectrum, Cole-Parmer, Hoefer Pharmacia, and others).
SELECTION AND PREPARATION OF DIALYSIS MEMBRANE Dialysis membranes are available in a number of thicknesses and pore sizes. Thicker membranes are tougher, but restrict solute flow and reach equilibrium more slowly. Pore size is defined by “molecular weight cutoff” (MWCO)—i.e., the size of the smallest particle that cannot penetrate the membrane. The MWCO designation should be used only as a very rough guide; if accurate MWCO information is required, it should be determined empirically (see Craig, 1967, for a discussion of parameters affecting MWCO). Knowledge of the precise MWCO is usually not required, however; it is necessary only to use a membrane with a pore size that is much smaller than the macromolecule of interest. For most plasmid and protein dialyses, a MWCO of 12,000 to 14,000 is appropriate, whereas a MWCO of 3500, 2000, or even 1000 is appropriate for peptides.
SUPPORT PROTOCOL
Most dialysis membranes are made of derivatives of cellulose. They come in a wide variety of MWCO values, ranging from 500 to 500,000, and also vary in cleanliness, sterility, and cost. Spectrum has the most impressive inventory. The least expensive membranes come dry on rolls; these contain glycerol to keep them flexible as well as residual sulfides and traces of heavy metals from their manufacture. If glycerol, sulfur compounds, or small amounts of heavy metals are problematic, cleaner membranes should be purchased or membranes should be prepared as described below. Protein dialysis should only be done with clean membranes. Additional Materials 10 mM sodium bicarbonate 10 mM Na2EDTA, pH 8.0 20% to 50% (v/v) ethanol 1. Remove membrane from the roll and cut into usable lengths (usually 8 to 12 in.). Always use gloves to handle dialysis membrane, as it is susceptible to a number of cellulolytic microorganisms. Membrane is available as sheets or preformed tubing.
2. Wet membrane and boil it for several minutes in a large excess of 10 mM sodium bicarbonate. 3. Boil several minutes in 10 mM Na2EDTA. Repeat. Boiling speeds up the treatment process but is not necessary. A 30-min soak with some agitation can substitute for the boiling step.
4. Wash several times in distilled water. 5. Store at 4°C in 20% to 50% ethanol to prevent growth of cellulolytic microorganisms. Alternatively, bacteriostatic agents (e.g., sodium azide or sodium cacodylate) may be used for storage; however, ethanol is preferred for ease and convenience.
Commonly Used Techniques
A.3B.3 Current Protocols in Protein Science
LITERATURE CITED Craig, L.C. 1967. Techniques for the study of peptides and proteins by dialysis and diffusion. Methods Enzymol. 11:870-905. McPhie, P. 1971. Dialysis. Methods Enzymol. 22:23-32. Describe the theory and practice of dialysis and diffusion.
Contributed by Louis Zumstein Introgen Therapeutics, Inc. Houston, Texas
Dialysis
A.3B.4 Current Protocols in Protein Science
Techniques for Mammalian Cell Tissue Culture
APPENDIX 3C
Tissue culture technology has found wide application in biological research. Monolayer cell cultures are utilized in cytogenetic, biochemical, and molecular laboratories for research as well as diagnostic studies. In most cases, cells or tissues must be grown in culture for days or weeks to obtain sufficient numbers of cells for analysis. Maintenance of cells in long-term culture requires strict adherence to aseptic technique to avoid contamination and potential loss of valuable cell lines. The first section of this appendix discusses basic principles of sterile technique. An important factor influencing the growth of cells in culture is the choice of tissue culture medium. Many different recipes for tissue culture media are available and each laboratory must determine which medium best suits their needs. Individual laboratories may elect to use commercially prepared medium or prepare their own. Commercially available medium can be obtained as a sterile and ready-to-use liquid, in a concentrated liquid form, or in a powdered form. Besides providing nutrients for growing cells, medium is generally supplemented with antibiotics, fungicides, or both to inhibit contamination. The second section of this appendix discusses medium preparation. As cells grown in monolayer reach confluency, they must be subcultured or passaged. Failure to subculture confluent cells results in reduced mitotic index and eventually in cell death. The first step in subculturing is to detach cells from the surface of the primary culture vessel by trypsinization or mechanical means. The resultant cell suspension is then subdivided, or reseeded, into fresh cultures. Secondary cultures are checked for growth, fed periodically, and may be subsequently subcultured to produce tertiary cultures, etc. The time between passaging cells varies with the cell line and depends on the growth rate. The Basic Protocol describes subculturing of a monolayer culture grown in petri dishes or flasks. Support Protocols 1 to 4 describe freezing of monolayer cells, thawing and recovery of cells, counting cells using a hemacytometer, and preparing cells for transport. The Alternate Protocol describes freezing suspension cells. STERILE TECHNIQUE It is essential that sterile technique be maintained when working with cell cultures. Aseptic technique involves a number of precautions to protect both the cultured cells and the laboratory worker from infection. The laboratory worker must realize that cells handled in the lab are potentially infectious and should be handled with caution. Protective apparel such as gloves, lab coats or aprons, and eyewear should be worn when appropriate (Knutsen, 1991). Care should be taken when handling sharp objects such as needles, scissors, scalpel blades, and glass that could puncture the skin. Sterile disposable plastic supplies may be used to avoid the risk of broken or splintered glass (Rooney and Czepulkowski, 1992). Frequently, specimens received in the laboratory are not sterile, and cultures prepared from these specimens may become contaminated with bacteria, fungus, or yeast. The presence of microorganisms can inhibit growth, kill cell cultures, or lead to inconsistencies in test results. The contaminants deplete nutrients in the medium and may produce substances that are toxic to cells. Antibiotics (penicillin, streptomycin, kanamycin, or gentamycin) and fungicides (amphotericin B or mycostatin) may be added to tissue Contributed by Mary C. Phelan Current Protocols in Protein Science (1997) A.3C.1-A.3C.14 Copyright © 1997 by John Wiley & Sons, Inc.
Commonly Used Techniques
A.3C.1 Supplement 7
culture medium to combat potential contaminants (see Table A.3C.1). An antibiotic/antimycotic solution or lyophilized powder that contains penicillin, streptomycin, and amphotericin B is available from Sigma. The solution can be used to wash specimens prior to culture and can be added to medium used for tissue culture. Similar preparations are available from other suppliers. All materials that come into direct contact with cultures must be sterile. Sterile disposable dishes, flasks, pipets, etc., can be obtained directly from manufacturers. Reusable glassware must be washed, rinsed thoroughly, then sterilized by autoclaving or by dry heat before reusing. With dry heat, glassware should be heated 90 min to 2 hr at 160°C to ensure sterility. Materials that may be damaged by very high temperatures can be autoclaved 20 min at 120°C and 15 psi. All media, reagents, and other solutions that come into contact with the cultures must also be sterile; medium may be obtained as a sterile liquid from the manufacturer, autoclaved if not heat-sensitive, or filter sterilized. Supplements can be added to media prior to filtration, or they can be added aseptically after filtration. Filters with 0.20- to 0.22-µm pore size should be used to remove small gram-negative bacteria from culture media and solutions. Contamination can occur at any step in handling cultured cells. Care should be taken when pipetting media or other solutions for tissue culture. The necks of bottles and flasks, as well as the tips of the pipets, should be flamed before the pipet is introduced into the bottle. If the pipet tip comes into contact with the benchtop or any other nonsterile surface, it should be discarded and a fresh pipet obtained. Forceps and scissors used in tissue culture can be rapidly sterilized by dipping in 70% alcohol and flaming. Although tissue culture work can be done on an open bench if aseptic methods are strictly enforced, many labs prefer to perform tissue culture work in a room or low-traffic area reserved specifically for that purpose. At the very least, biological safety cabinets are recommended to protect the cultures as well as the laboratory worker. In a laminar flow hood, the flow of air protects the work area from dust and contamination and acts as a barrier between the work surface and the worker. Many different styles of safety hoods are available and the laboratory should consider the types of samples being processed and the types of potential pathogenic exposure in making their selection. Manufacturer recommendations should be followed regarding routine maintenance checks on air flow and filters. For day-to-day use, the cabinet should be turned on for at least 5 min prior to beginning work. All work surfaces both inside and outside of the hood should be kept clean and disinfected daily and after each use.
Techniques for Mammalian Cell Tissue Culture
Some safety cabinets are equipped with ultraviolet (UV) lights for decontamination of work surfaces. However, use of UV lamps is no longer recommended, as it is generally ineffective (Knutsen, 1991). UV lamps may produce a false sense of security as they maintain a visible blue glow long after their germicidal effectiveness is lost. Effectiveness diminishes over time as the glass tube gradually loses its ability to transmit short UV wavelengths, and it may also be reduced by dust on the glass tube, distance from the work surface, temperature, and air movement. Even when the UV output is adequate, the rays must directly strike a microorganism in order to kill it; bacteria or mold spores hidden below the surface of a material or outside the direct path of the rays will not be destroyed. Another rule of thumb is that anything that can be seen cannot be killed by UV. UV lamps will only destroy microorganisms such as bacteria, virus, and mold spores; they will not destroy insects or other large organisms (Westinghouse Electric Company, 1976). The current recommendation is that work surfaces be wiped down with ethanol instead of relying on UV lamps, although some labs use the lamps in addition to ethanol wipes to decontaminate work areas. A special metering device is available to measure the output
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of UV lamps, and the lamps should be replaced when they fall below the minimum requirements for protection (Westinghouse Electric Company, 1976). Cultures should be checked routinely for contamination. Indicators in the tissue culture medium change color when contamination is present: for example, medium that contains phenol red changes to yellow because of increased acidity. Cloudiness and turbidity are also observed in contaminated cultures. Once contamination is confirmed with a microscope, infected cultures are generally discarded. Keeping contaminated cultures increases the risk of contaminating other cultures. Sometimes a contaminated cell line can be salvaged by treating it with various combinations of antibiotics and antimycotics in an attempt to eradicate the infection (e.g., see Fitch et al., 1991). However, such treatment may adversely affect cell growth and is often unsuccessful in ridding cultures of contamination. PREPARING CULTURE MEDIUM Choice of tissue culture medium comes from experience. An individual laboratory must select the medium that best suits the type of cells being cultured. Chemically defined media are available in liquid or powdered form from a number of suppliers. Sterile, ready-to-use medium has the advantage of being convenient, although it is more costly than other forms. Powdered medium must be reconstituted with tissue culture grade water according to manufacturer’s directions. Distilled or deionized water is not of sufficiently high quality for medium preparation; double- or triple-distilled water or commercially available tissue culture water should be used. The medium should be filter-sterilized and transferred to sterile bottles. Prepared medium can generally be stored ≤1 month in a 4°C refrigerator. Laboratories using large volumes of medium may choose to prepare their own medium from standard recipes. This is the most economical approach, but it is time-consuming. Basic media such as Eagle’s minimal essential medium (MEM), Dulbecco’s modified Eagle medium (DMEM), Glasgow modified Eagle medium (GMEM), and RPMI 1640 and Ham F10 nutrient mixtures (e.g., Life Technologies) are composed of amino acids, glucose, salts, vitamins, and other nutrients. A basic medium is supplemented by addition of L-glutamine, antibiotics (typically penicillin and streptomycin sulfate), and usually serum to formulate a “complete medium.” Where serum is added, the amount is indicated as a percentage of fetal bovine serum (FBS) or other serum. Some media are also supplemented with antimycotics, nonessential amino acids, various growth factors, and/or drugs that provide selective growth conditions. Supplements should be added to medium prior to sterilization or filtration, or added aseptically just before use. The optimum pH for most mammalian cell cultures is 7.2 to 7.4. Adjust pH of the medium as necessary after all supplements are added. Buffers such as bicarbonate and HEPES are routinely used in tissue culture medium to prevent fluctuations in pH that might adversely affect cell growth. HEPES is especially useful in solutions used for procedures that do not take place in a controlled CO2 environment. Most cultured cells will tolerate a wide range of osmotic pressure and an osmolarity between 260 and 320 mOsm/kg is acceptable for most cells. The osmolarity of human plasma is ∼290 mOsm/kg, and this is probably the optimum for human cells in culture as well (Freshney, 1993). Fetal bovine serum (FBS; sometimes known as fetal calf serum, FCS) is the most frequently used serum supplement. Calf serum, horse serum, and human serum are also used; some cell lines are maintained in serum-free medium (Freshney, 1993). Complete
Commonly Used Techniques
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Table A.3C.1 Working Concentrations of Antibiotics and Fungicides for Mammalian Cell Culture
Additive Penicillin Streptomycin sulfate Kanamycin Gentamycin Mycostatin Amphotericin B
Final concentration 50–100 U/ml 50–100 µg/ml 100 µg/ml 50 µg/ml 20 µg/ml 0.25 µg/ml
medium is supplemented with 5% to 30% (v/v) serum, depending on the requirements of the particular cell type being cultured. Serum that has been heat-inactivated (30 min to 1 hr at 56°C) is generally preferred, because heat treatment inactivates complement and is thought to reduce the number of contaminants. Serum is obtained frozen, then is thawed, divided into smaller portions, and refrozen until needed. There is considerable lot-to-lot variation in FBS. Most suppliers will provide a sample of a specific lot and reserve a supply of that lot while the serum is tested for its suitability. The suitability of a serum lot depends upon the use. Frequently the ability of serum to promote cell growth equivalent to a laboratory standard is used to evaluate a serum lot. Once an acceptable lot is identified, enough of that lot should be purchased to meet the culture needs of the laboratory for an extended period of time. Commercially prepared media containing L-glutamine are available, but many laboratories choose to obtain medium without L-glutamine, and then add it to a final concentration of 2 mM just before use. L-glutamine is an unstable amino acid that, upon storage, converts to a form cells cannot use. Breakdown of L-glutamine is temperature- and pH-dependent. At 4°C, 80% of the L-glutamine remains after 3 weeks, but at near incubator temperature (35°C) only half remains after 9 days (Barch et al., 1991). To prevent degradation, 100× L-glutamine should be stored frozen in aliquots until needed. As well as practicing good aseptic technique, most laboratories add antimicrobial agents to medium to further reduce the risk of contamination. A combination of penicillin and streptomycin is the most commonly used antibiotic additive; kanamycin and gentamycin are used alone. Mycostatin and amphotericin B are the most commonly used fungicides (Rooney and Czepulkowski, 1992). Table A.3C.1 lists the final concentrations for the most commonly used antibiotics and antimycotics. Combining antibiotics in tissue culture media can be tricky, as some antibiotics are not compatible, and one may inhibit the action of another. Furthermore, combined antibiotics may be cytotoxic at lower concentrations than is true for the individual antibiotics. In addition, prolonged use of antibiotics may cause cell lines to develop antibiotic resistance. For this reason, some laboratories add antibiotics and/or fungicides to medium when initially establishing a culture but eliminate them from medium used in later subcultures.
Techniques for Mammalian Cell Tissue Culture
All tissue culture medium, whether commercially prepared or prepared within the laboratory, should be tested for sterility prior to use. A small aliquot from each lot of medium is incubated 48 hr at 37°C and monitored for evidence of contamination such as turbidity (infected medium will be cloudy) and color change (if phenol red is the indicator, infected medium will turn yellow). Any contaminated medium should be discarded.
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TRYPSINIZING AND SUBCULTURING CELLS FROM A MONOLAYER 2
A primary culture is grown to confluency in a 60-mm petri dish or 25-cm tissue culture flask containing 5 ml tissue culture medium. Cells are dispersed by trypsin treatment and then reseeded into secondary cultures. The process of removing cells from the primary culture and transferring them to secondary cultures constitutes a passage or subculture.
BASIC PROTOCOL
Materials Primary cultures of cells HBSS without Ca2+ and Mg2+ (see recipe), 37°C 0.25% (w/v) trypsin/0.2% EDTA solution (see recipe), 37°C Complete medium with serum: e.g., DMEM supplemented with 10% to 15% (v/v) fetal bovine serum (complete DMEM-10; see recipe), 37°C Sterile Pasteur pipets 37°C warming tray or incubator Tissue culture plasticware or glassware including pipets and 25-cm2 flasks or 60-mm petri dishes, sterile NOTE: All incubations are performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. 1. Remove all medium from primary culture with a sterile Pasteur pipet. Wash adhering cell monolayer once or twice with a small volume of 37°C HBSS without Ca2+ and Mg2+ to remove any residual FBS, which may inhibit the action of trypsin. Use a buffered salt solution that is Ca2+- and Mg2+-free to wash cells. Ca2+ and Mg2+ in the salt solution can cause cells to stick together. If this is the first medium change, rather than discarding medium that is removed from primary culture, put it into a fresh dish or flask. The medium contains unattached cells that may attach and grow, thereby providing a backup culture.
2. Add enough 37°C trypsin/EDTA solution to culture to cover adhering cell layer. 3. Place plate on a 37°C warming tray 1 to 2 min. Tap bottom of plate on the countertop to dislodge cells. Check culture with an inverted microscope to be sure that cells are rounded up and detached from the surface. If cells are not sufficiently detached, return plate to warming tray for an additional minute or two.
4. Add 2 ml of 37°C complete medium. Draw cell suspension into a Pasteur pipet and rinse cell layer two or three times to dissociate cells and to dislodge any remaining adherent cells. As soon as cells are detached, add serum or medium containing serum to inhibit further trypsin activity that might damage cells. If cultures are to be split 1/3 or 1/4 rather than 1/2, add sufficient medium such that 1 ml of cell suspension can be transferred into each fresh culture vessel.
5. Add an equal volume of cell suspension to fresh dishes or flasks that have been appropriately labeled. Alternatively, cells can be counted using a hemacytometer (see Support Protocol 3) or Coulter counter and diluted to the desired density so a specific number of cells can be added to each culture vessel. A final concentration of ∼5 × 104 cells/ml is appropriate for most subcultures. For primary cultures and early subcultures, 60-mm petri dishes or 25-cm2 flasks are generally used; larger petri dishes or flasks (e.g., 150-mm dishes or 75-cm2 flasks) may be used for later subcultures. Cultures should be labeled with patient name, lab number, date of subculture, and passage number.
Commonly Used Techniques
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6. Add 4 ml fresh medium to each new culture. Incubate in a humidified 37°C, 5% CO2 incubator. If using 75-cm2 culture flasks, add 9 ml medium per flask. Some labs now use incubators with 5% CO2 and 4% O2. The low oxygen concentration is thought to simulate the in vivo environment of cells and to enhance cell growth.
7. If necessary, feed subconfluent cultures after 3 or 4 days by removing old medium and adding fresh 37°C medium. 8. Passage secondary culture when it becomes confluent by repeating steps 1 to 7, and continue to passage as necessary. SUPPORT PROTOCOL 1
FREEZING HUMAN CELLS GROWN IN MONOLAYER CULTURES It is sometimes desirable to store cell lines for future study. To preserve cells, avoid senescence, reduce the risk of contamination, and minimize effects of genetic drift, cell lines may be frozen for long-term storage. Without the use of a cryoprotective agent freezing would be lethal to the cells in most cases. Generally, a cryoprotective agent such as dimethylsulfoxide (DMSO) is used in conjunction with complete medium for preserving cells at −70°C or lower. DMSO acts to reduce the freezing point and allows a slower cooling rate. Gradual freezing reduces the risk of ice crystal formation and cell damage. Materials Log-phase monolayer culture of cells in petri dish Complete medium Freezing medium: complete medium (e.g., DMEM or RPMI; see recipes) supplemented with 10% to 20% (v/v) FBS and 5% to 10% (v/v) DMSO, 4°C Benchtop clinical centrifuge (e.g., Fisher Centrific or Clay Adams Dynac) with 45°C fixed-angle or swinging-bucket rotor 1. Trypsinize log-phase monolayer culture of cells from plate (see Basic Protocol, steps 1 to 4). It is best to use cells in log-phase growth for cryopreservation.
2. Transfer cell suspension to a sterile centrifuge tube and add 2 ml complete medium with serum. Centrifuge 5 min at 300 to 350 × g (∼1500 rpm in Fisher Centrific rotor), room temperature. Cells from three or more dishes from the same subculture of the same patient can be combined in one tube.
3. Remove supernatant and add 1 ml of 4°C freezing medium. Resuspend pellet. 4. Add 4 ml of 4°C freezing medium, mix cells thoroughly, and place on wet ice. 5. Count cells using a hemacytometer (see Support Protocol 3). Dilute with more freezing medium as necessary to get a final cell concentration of 106 or 107 cells/ml. To freeze cells from a nearly confluent 25-cm2 flask, resuspend in roughly 3 ml freezing medium.
6. Pipet 1-ml aliquots of cell suspension into labeled 2-ml cryovials. Tighten caps on vials.
Techniques for Mammalian Cell Tissue Culture
7. Place vials 1 hr to overnight in a −70°C freezer, then transfer to liquid nitrogen storage freezer. Alternatively, freeze cells in a freezing chamber in the neck of a Dewar flask according to manufacturer’s instructions. Some laboratories place vials directly into the liquid nitrogen
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freezer, omitting the gradual temperature drop. Although this is contrary to the general recommendation to gradually reduce the temperature, laboratories that routinely use a direct-freezing technique report no loss of cell viability on recovery. Keep accurate records of the identity and location of cells stored in liquid nitrogen freezers. Cells may be stored for many years and proper information is imperative for locating a particular line for future use.
FREEZING CELLS GROWN IN SUSPENSION CULTURE Freezing cells from suspension culture is similar in principle to freezing cells from monolayer. The major difference is that suspension cultures need not be trypsinized.
ALTERNATE PROTOCOL
1. Transfer cell suspension to a centrifuge tube and centrifuge 10 min at 300 to 350 × g (∼1500 rpm in Fisher Centrific centrifuge), room temperature. 2. Remove supernatant and resuspend pellet in 4°C freezing medium at a density of 106 to 107 cells/ml. Some laboratories freeze lymphoblastoid lines at the higher cell density because they plan to recover them in a larger volume of medium and because there may be a greater loss of cell viability upon recovery as compared to other types of cells (e.g., fibroblasts).
3. Transfer 1-ml aliquots of cell suspension into labeled cryovials and freeze as for monolayer cultures. THAWING AND RECOVERING HUMAN CELLS When cryopreserved cells are needed for study, they should be thawed rapidly and plated at high density to optimize recovery.
SUPPORT PROTOCOL 2
CAUTION: Protective clothing, particularly insulated gloves and goggles, should be worn when removing frozen vials or ampules from the liquid nitrogen freezer. The room containing the liquid nitrogen freezer should be well-ventilated. Care should be taken not to spill liquid nitrogen on the skin. Materials Cryopreserved cells stored in liquid nitrogen freezer 70% (v/v) ethanol Complete medium (e.g., DMEM or RPMI; see recipes) containing 20% FBS (see recipe), 37°C Tissue culture dish or flask NOTE: All incubations are performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. 1. Remove vial from liquid nitrogen freezer and immediately place it into a 37°C water bath. Agitate vial continuously until medium is thawed. The medium usually thaws in <60 sec. Cells should be thawed as quickly as possible to prevent formation of ice crystals which can cause cell lysis. Try to avoid getting water around the cap of the vial.
2. Wipe top of vial with 70% ethanol before opening. Some labs prefer to submerge the vial in 70% ethanol and air dry before opening.
3. Transfer thawed cell suspension into a sterile centrifuge tube containing 2 ml warm complete medium containing 20% FBS. Centrifuge 10 min at 150 to 200 × g (∼1000 rpm in Fisher Centrific), room temperature. Discard supernatant. Cells are washed with fresh medium to remove residual DMSO.
Commonly Used Techniques
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4. Gently resuspend cell pellet in small amount (∼1 ml) of complete medium/20% FBS and transfer to properly labeled culture dish or flask containing the appropriate amount of medium. Cultures are reestablished at a higher cell density than that used for original cultures because there is some cell death associated with freezing. Generally, 1 ml cell suspension is reseeded in 5 to 20 ml medium.
5. Check cultures after ∼24 hr to ensure that cells have attached to the plate. 6. Change medium after 5 to 7 days or when pH indicator (e.g., phenol red) in medium changes color. Keep cultures in medium with 20% FBS until cell line is reestablished. If recovery rate is extremely low, only a subpopulation of the original culture may be growing; be extra careful of this when working with cell lines known to be mosaic. SUPPORT PROTOCOL 3
DETERMINING CELL NUMBER AND VIABILITY WITH A HEMACYTOMETER AND TRYPAN BLUE STAINING Determining the number of cells in culture is important in standardization of culture conditions and in performing accurate quantitation experiments. A hemacytometer is a thick glass slide with a central area designed as a counting chamber. The exact design of the hemacytometer may vary; the one described here is the Improved Neubauer from Baxter Scientific (Fig. A.3C.1). The central portion of the slide is the counting platform, which is bordered by a 1-mm groove. The central platform is divided into two counting chambers by a transverse groove. Each counting chamber consists of a silver footplate on which is etched a 3 × 3–mm grid. This grid is divided into nine secondary squares, each 1 × 1 mm. The four corner squares and the central square are used for determining the cell count. The corner squares are further divided into 16 tertiary squares and the central square into 25 tertiary squares to aid in cell counting. Accompanying the hemacytometer slide is a thick, even-surfaced coverslip. Ordinary coverslips may have uneven surfaces, which can introduce errors in cell counting; therefore, it is imperative that the coverslip provided with the hemacytometer be used in determining cell number. Cell suspension is applied to a defined area and counted so cell density can be calculated. Materials 70% (v/v) ethanol Cell suspension 0.4% (w/v) trypan blue or 0.4% (w/v) nigrosin, prepared in HBSS (see recipe) Hemacytometer with coverslip (Improved Neubauer, Baxter Scientific) Hand-held counter Prepare hemacytometer 1. Clean surface of hemacytometer slide and coverslip with 70% alcohol. Coverslip and slide should be clean, dry, and free from lint, fingerprints, and watermarks.
2. Wet edge of coverslip slightly with tap water and press over grooves on hemacytometer. The coverslip should rest evenly over the silver counting area.
Techniques for Mammalian Cell Tissue Culture
Prepare cell suspension 3. For cells grown in monolayer cultures, detach cells from surface of dish using trypsin (see Basic Protocol).
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4. Dilute cells as needed to obtain a uniform suspension. Disperse any clumps. When using the hemacytometer, a maximum cell count of 20 to 50 cells per 1-mm square is recommended.
Load hemacytometer 5. Use a sterile Pasteur pipet to transfer cell suspension to edge of hemacytometer counting chamber. Hold tip of pipet under the coverslip and dispense one drop of suspension. Suspension will be drawn under the coverslip by capillary action. Hemacytometer should be considered nonsterile. If cell suspension is to be used for cultures, do not reuse the pipet and do not return any excess cell suspension in the pipet to the original suspension.
coverslip
grid load cell suspension
1mm
2
1 3 4
1mm
5
Figure A.3C.1 Hemacytometer slide (Improved Neubauer) and coverslip. Coverslip is applied to slide and cell suspension is added to counting chamber using a Pasteur pipet. Each counting chamber has a 3 × 3–mm grid (enlarged). The four corner squares (1, 2, 4, and 5) and the central square (3) are counted on each side of the hemacytometer (numbers added).
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6. Fill second counting chamber. Count cells 7. Allow cells to settle for a few minutes before beginning to count. Blot off excess liquid. 8. View slide on microscope with 100× magnification. A 10× ocular with a 10× objective = 100× magnification. Position slide to view the large central area of the grid (section 3 in Fig. A.3C.1); this area is bordered by a set of three parallel lines. The central area of the grid should almost fill the microscope field. Subdivisions within the large central area are also bordered by three parallel lines and each subdivision is divided into sixteen smaller squares by single lines. Cells within this area should be evenly distributed without clumping. If cells are not evenly distributed, wash and reload hemacytometer.
9. Use a hand-held counter to count cells in each of the four corner and central squares (Fig. A.3C.1, squares numbered 1 to 5). Repeat counts for other counting chamber. Five squares (four corner and one center) are counted from each of the two counting chambers for a total of ten squares counted. Count cells touching the middle line of the triple line on the top and left of the squares. Do not count cells touching the middle line of the triple lines on the bottom or right side of the square.
Calculate cell number 10. Determine cells per milliliter by the following calculations: cells/ml = average count per square × dilution factor × 104 total cells = cells/ml × total original volume of cell suspension from which sample was taken. 104 is the volume correction factor for the hemacytometer: each square is 1 × 1 mm and the depth is 0.1 mm.
Stain cells with trypan blue to determine cell viability 11. Determine number of viable cells by adding 0.5 ml of 0.4% trypan blue, 0.3 ml HBSS, and 0.1 ml cell suspension to a small tube. Mix thoroughly and let stand 5 min before loading hemacytometer. Either 0.4% trypan blue or 0.4% nigrosin can be used to determine the viable cell number. Nonviable cells will take up the dye, whereas live cells will be impermeable to it.
12. Count total number of cells and total number of viable (unstained) cells. Calculate percent viable cells as follows: % viable cells =
number of unstained cells × 100 total number of cells
13. Decontaminate coverslip and hemacytometer by rinsing with 70% ethanol and then deionized water. Air dry and store for future use.
Techniques for Mammalian Cell Tissue Culture
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PREPARING CELLS FOR TRANSPORT 2
Both monolayer and suspension cultures can easily be shipped in 25-cm tissue culture flasks. Cells are grown to near confluency in a monolayer or to desired density in suspension. Medium is removed from monolayer cultures and the flask is filled with fresh medium. Fresh medium is added to suspension cultures to fill the flask. It is essential that the flasks be completely filled with medium to protect cells from drying if flasks are inverted during transport. The cap is tightened and taped securely in place. The flask is sealed in a leak-proof plastic bag or other leak-proof container designed to prevent spillage in the event that the flask should become damaged. The primary container is then placed in a secondary insulated container to protect it from extreme temperatures during transport. A biohazard label is affixed to the outside of the package. Generally, cultures are transported by same-day or overnight courier.
SUPPORT PROTOCOL 4
Cells can also be shipped frozen. The vial containing frozen cells is removed from the liquid nitrogen freezer and placed immediately on dry ice in an insulated container to prevent thawing during transport. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of cell buffers. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Complete DMEM Dulbecco’s modified Eagle medium, high-glucose formulation (e.g., Life Technologies), containing: 5%, 10%, or 20% (v/v) FBS, heat-inactivated (optional; see recipe) 1% (v/v) nonessential amino acids 2 mM L-glutamine (see recipe) 100 U/ml penicillin 100 µg/ml streptomycin sulfate Filter sterilize and store ≤1 month at 4°C Throughout this manual, the percentage of serum (usually fetal bovine serum, FBS) used in a protocol step is indicated by a numeral hyphenated to the base medium name. Thus, “complete DMEM-10” indicates that 10% FBS is used. Absence of a numeral indicates that no serum is used. DMEM containing 4500 mg/liter D-glucose can be obtained from Life Technologies. DMEM is also known as Dulbecco’s minimum essential medium.
Complete RPMI RPMI 1640 medium (e.g., Life Technologies) containing: 2%, 5%, 10%, 15%, or 20% FBS, heat-inactivated (optional; see recipe) 2 mM L-glutamine (see recipe) 100 U/ml penicillin 100 µg/ml streptomycin sulfate Filter sterilize and store ≤1 month at 4°C FBS (fetal bovine serum) Thaw purchased fetal bovine serum (shipped on dry ice and kept frozen until needed). Store 3 to 4 weeks at 4°C. If FBS is not to be used within this time, aseptically divide into smaller aliquots and refreeze until used. Store ≤1 year at −20°C. Repeated thawing and refreezing should be avoided as it may cause denaturation of the serum.
Commonly Used Techniques
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Inactivated FBS (FBS that has been treated with heat to inactivate complement protein and thus prevent an immunological reaction against cultured cells) is useful for a variety of purposes. It can be purchased commercially or made in the lab. To inactivate FBS, heat serum 30 min to 1 hr in a 56°C water bath. Alternatively, FBS may be inactivated through radiation treatment. L-glutamine,
0.2 M (100×) Thaw commercially prepared frozen L-glutamine or prepare an 0.2 M solution in water, aliquot aseptically into usable portions, then refreeze. For convenience, L-glutamine can be stored in 1-ml aliquots if 100-ml bottles of medium are used and in 5-ml aliquots if 500-ml bottles are used. Store ≤1 year at −20°C. Many laboratories supplement medium with 2 mM L-glutamine—1% (v/v) of 100× stock— just prior to use.
HBSS (Hanks balanced salt solution) 5.4 mM KCl 0.3 mM Na2HPO4 0.4 mM KH2HPO4 4.2 mM NaHCO3 1.3 mM CaCl2 0.5 mM MgCl2 0.6 mM MgSO4 137 mM NaCl 5.6 mM D-glucose 0.02% (w/v) phenol red (optional) Add H2O to 1 liter and adjust pH to 7.4 HBSS can be purchased from Biofluids or Whittaker.
To prepare HBSS without Ca2+ and Mg2+, omit the CaCl2, MgCl2, and MgSO4. Trypsin/EDTA solution Prepare in sterile HBSS (see recipe) or 0.9% (w/v) NaCl: 0.25% (w/v) trypsin 0.2% (w/v) EDTA Store ≤1 year (until needed) at −20°C Specific applications may require different concentrations of trypsin. Trypsin/EDTA solution is commercially available in various concentrations including 10×, 1×, and 0.25% (w/v). It is received frozen from the manufacturer and can be thawed and aseptically divided into smaller volumes. Preparing trypsin/EDTA from powdered stocks may reduce its cost; however, most laboratories prefer commercially prepared solutions for convenience. EDTA (disodium ethylenediamine tetraacetic acid) is added as a chelating agent to bind Ca2+ and Mg2+ ions that can interfere with the action of trypsin.
COMMENTARY Background Information
Techniques for Mammalian Cell Tissue Culture
At its inception in the early twentieth century, tissue culture was applied to the study of tissue fragments in culture. New growth in culture was limited to cells that migrated out from the initial tissue fragment. Tissue culture techniques evolved rapidly, and since the 1950s culture methods have allowed the growth and study of dispersed cells in culture (Freshney,
1993). Cells dispersed from the original tissue can be grown in monolayers and passaged repeatedly to give rise to a relatively stable cell line. Four distinct growth stages have been described for primary cells maintained in culture. First, cells adapt to the in vitro environment. Second, cells undergo an exponential growth phase lasting through ∼30 passages. Third, the
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growth rate of cells slows, leading to a progressively longer generation time. Finally, after 40 or 50 passages, cells begin to senesce and die (Lee, 1991). It may be desirable to study a particular cell line over several months or years, so monolayer cultures can be preserved to retain the integrity of the cell line. Aliquots of early passage cell suspensions are frozen, then thawed, and cultures reestablished as needed. Freezing monolayer cultures prevents changes due to genetic drift and avoids loss of cultures due to senescence or accidental contamination (Freshney, 1993). Cell lines are commercially available from a number of sources, including the American Type Culture Collection (ATCC) and the Human Genetic Mutant Cell Repository at the Coriell Cell Repository (CCR). These cell repositories are a valuable resource for researchers who do not have access to suitable patient populations.
Critical Parameters Use of aseptic technique is essential for successful tissue culture. Cell cultures can be contaminated at any time during handling, so precautions must be taken to minimize the chance of contamination. All supplies and reagents that come into contact with cultures must be sterile and all work surfaces should be kept clean and free from clutter. Cultures should be 75% to 100% confluent when selected for subculture. Growth in monolayer cultures will be adversely affected if cells are allowed to become overgrown. Passaging cells too early will result in a longer lag time before subcultures are established. Following dissociation of the monolayer by trypsinization, serum or medium containing serum should be added to the cell suspension to stop further action by trypsin that might be harmful to cells. When subculturing cells, add a sufficient number of cells to give a final concentration of ∼5 × 104 cells/ml in each new culture. Cells plated at too low a density may be inhibited or delayed in entry into growth stage. Cells plated at too high a density may reach confluence before the next scheduled subculturing; this could lead to cell loss and/or cessation of proliferation. The growth characteristics for different cell lines vary. A lower cell concentration (104 cells/ml) may be used to initiate subcultures of rapidly growing cells, and a higher cell concentration (105 cells/ml) may be used to initiate subcultures of more slowly growing cells. Adjusting the initial cell concentration
permits establishment of a regular, convenient schedule for subculturing—e.g., once or twice a week (Freshney, 1993). Cells in culture will undergo changes in growth, morphology, and genetic characteristics over time. Such changes can adversely affect reproducibility of laboratory results. Nontransformed cells will undergo senescence and eventual death if passaged indefinitely. The time of senescence will vary with cell line, but generally at between 40 and 50 population doublings fibroblast cell lines begin to senesce. Cryopreservation of cell lines will protect against these adverse changes and will avoid potential contamination. Cultures selected for cryopreservation should be in log-phase growth and free from contamination. Cells should be frozen at a concentration of 106 to 107 cells/ml. Cells should be frozen gradually and thawed rapidly to prevent formation of ice crystals that may cause cells to lyse. Cell lines can be thawed and recovered after long-term storage in liquid nitrogen. The top of the freezing vial should be cleaned with 70% alcohol before opening to prevent introduction of contaminants. To aid in recovery of cultures, thawed cells should be reseeded at a higher concentration than that used for initiating primary cultures. Careful records regarding identity and characteristics of frozen cells as well as their location in the freezer should be maintained to allow for easy retrieval. For accurate cell counting, the hemacytometer slide should be clean, dry, and free from lint, scratches, fingerprints, and watermarks. The coverslip supplied with the hemacytometer should always be used because it has an even surface and is specially designed for use with the counting chamber. Use of an ordinary coverslip may introduce errors in cell counting. If the cell suspension is too dense or the cells are clumped, inaccurate counts will be obtained. If the cell suspension is not evenly distributed over the counting chamber, the hemacytometer should be washed and reloaded.
Anticipated Results Confluent cell lines can be successfully subcultured in the vast majority of cases. The yield of cells derived from monolayer culture is directly dependent on the available surface area of the culture vessel (Freshney, 1993). Overly confluent cultures or senescent cells may be difficult to trypsinize, but increasing the time of trypsin exposure will help dissociate resistant cells. Cell lines can be propagated to get
Commonly Used Techniques
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sufficient cell populations for cytogenetic, biochemical, and molecular analyses. It is well accepted that anyone can successfully freeze cultured cells; it is thawing and recovering the cultures that presents the problem. Cultures that are healthy and free from contamination can be frozen and stored indefinitely. Cells stored in liquid nitrogen can be successfully thawed and recovered in over 95% of cases. Several aliquots of each cell line should be stored to increase the chance of recovery. Cells should be frozen gradually, with a temperature drop of ∼1°C per minute, but thawed rapidly. Gradual freezing and rapid thawing prevents formation of ice crystals that might cause cell lysis. Accurate cell counts can be obtained using the hemacytometer if cells are evenly dispersed in suspension and free from clumps. Determining the proportion of viable cells in a population will aid in standardization of experimental conditions.
Fitch, F.W., Gajewski, T.F., and Yokoyama, W.M. 1991. Diagnosis and treatment of mycoplasmacontaminated cell cultures. In Current Protocols in Immunologoy (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. A.3.9-A.3.12. John Wiley & Sons, New York.
Time Considerations
Key Reference
Establishment and maintenance of mammalian cell cultures require a regular routine for preparing media and feeding and passaging cells. Cultures should be inspected regularly for signs of contamination and to determine if the culture needs feeding or passaging.
Literature Cited Barch, M.J., Lawce, H.J., and Arsham, M.S. 1991. Peripheral Blood Culture. In The ACT Cytogenetics Laboratory Manual, 2nd ed. (M.J. Barch, ed.) pp. 17-30. Raven Press, New York.
Freshney, R.I. 1993. Culture of Animal Cells. A Manual of Basic Techniques, 3rd ed. Wiley-Liss, New York. Knutsen, T. 1991. In The ACT Cytogenetics Laboratory Manual, 2nd ed. (M.J. Barch, ed.) pp. 563-587. Raven Press, New York. Lee, E.C. 1991. Cytogenetic Analysis of Continuous Cell Lines. In The ACT Cytogenetics Laboratory Manual, 2nd ed. (M.J. Barch, ed.) pp. 107-148. Raven Press, New York. Rooney, D.E. and Czepulkowski, B.H. (eds.) 1992. Human Cytogenetics: A Practical Approach, Vol. I. Constitutional Analysis, 2nd ed. IRL Press, Washington, D.C. Westinghouse Electric Company. 1976. Westinghouse sterilamp germicidal ultraviolet tubes. Westinghouse Electric Corp., Bloomfield, NJ.
Lee, E.C. 1991. See above. Contains pertinent information on cell culture requirements including medium preparation and sterility. Also discusses trypsinization, freezing and thawing, and cell counting.
Contributed by Mary C. Phelan Greenwood Genetic Center Greenwood, South Carolina
Techniques for Mammalian Cell Tissue Culture
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Current Protocols in Protein Science
Importing Biological Materials Scientific cooperation is no longer confined to laboratories at the same institution, nor between laboratories within the boundaries of the United States. More than ever, scientists are forming collaborations with their peers outside the U.S., leading not only to the exchange of knowledge and ideas, but also to the exchange of biological reagents. Thus, investigators need be aware that an import permit may be required to obtain purified proteins, chemically synthesized materials, cell lines, tissues, and other biological materials from abroad. Before requesting any biological material from a laboratory outside the U.S., it is the responsibility of the “receiver” to ascertain whether an import permit is required, to apply for the permit if one is determined to be necessary, and to provide the “shipper” with an official copy of the permit to be included with the shipped material. There are two government agencies in place to issue permits for the importation of most biologicals. Materials derived from all animals are subject to regulations set forth by the United States Department of Agriculture (USDA) and must be cleared by USDA inspectors at the port of arrival before entry into the United States is authorized. In general, an import permit issued by the USDA is necessary for receipt of any animal-derived material or any biological material (including proteins) that has been in contact with materials of animal origin. The USDA does not have regulatory authority over the importation of human or nonhuman primate material unless it is produced via tissue culture, due to the use of bovine material (fetal bovine serum or calf serum) in tissue culture. The U.S. Public Health Service (UPHS) has jurisdiction over human and nonhuman primate materials. According to UPHS regulations, any etiologic agent, animal host, or vector of human disease may not be imported in the United States nor distributed after importation without being accompanied by a permit issued by the Director of the Centers for Disease Control. Failure to obtain the appropriate import permit will result in the shipment being held at the port of entry until a permit is applied for and issued (a 4- to 6-week process), or it may result in shipment confiscation and destruction by the quarantine officer present.
APPENDIX 3D
IMPORTING RESTRICTED BIOLOGICAL MATERIALS The following is a list of restricted biological materials requiring an import permit: proteins, monoclonal and polyclonal antibodies, antisera, immunoglobins, recombinant products, enzymes, hormones, immunoassay components or kits, animal tissues, blood, cells, cell lines, RNA/DNA extracts, plasmids, vectors, microorganisms—e.g., fungi, bacteria, viruses—and other products that are derived from or have been in contact with animal-derived materials. In addition, materials received from countries shown not to be free of foot-andmouth disease (see Title 9, code of Federal Regulations, Part 94.1) require an import permit. Note that the USDA will not permit the importation of cell cultures, monoclonal antibodies, ascites fluid, or bovine serum from countries where rinderpest and foot-and-mouth disease are present unless the imported materials are determined to be virus-free. Cell cultures being imported into the U.S. must have been grown only with fetal bovine serum of U.S., Canadian, New Zealand, or Australian origin. Material exposed to bovine products originating in the United Kingdom, Switzerland, Ireland, France, or Portugal may NOT be used, either directly or indirectly, in any animals due to the presence of bovine spongiform encephalopathy (BSE) in those countries, and the material is restricted to in vitro uses only.
CRITERIA FOR HANDLING IMPORTED RESTRICTED MATERIALS An import permit is only valid for transfer of the material(s) listed on the permit (a permit can cover more than one item), and for shipment between the two laboratories listed on the permit. Therefore, for each laboratory abroad from which biological materials are to be imported, a separate import permit is required. Once biological materials have been received from abroad, the import permit still imposes regulations on how the materials are handled by the receiver. Import permits do not usually authorize direct or indirect exposure of domestic animals—work is limited to in vitro uses only. In addition, permits are only valid for work conducted or directed by the permit holder at his or her facilities. Materials cannot be removed to another location, nor distributed to others, without USDA authorization. There Commonly Used Techniques
Contributed by Paula Wolf Current Protocols in Protein Science (1997) A.3D.1-A.3D.3 Copyright © 1997 by John Wiley & Sons, Inc.
A.3D.1 Supplement 7
is one exception: monoclonal antibodies produced by hybridoma cell lines may be commercially distributed for in vitro uses.
IMPORTING NONRESTRICTED BIOLOGICAL MATERIALS Any of the restricted materials that do not contain animal- or cell culture-derived products or additives such as albumin, serum, or gelatin, or were not exposed to any infectious agents of agricultural concern, do not need an official import permit. However, an accompanying declaration is required with each shipment to indicate that the material does not contain any animal- or cell culture-derived products or additives. This information should be supplied with each shipment in a clear and concise manner, and be available for the USDA Port of Entry Inspector’s review. A separate memo or letter should be included with the shipping documents, such as U.S. Customs declaration and invoice. Note, whether a particular biological material requires an import permit cannot be decided by the investigator; the investigator must contact the USDA for a decision well in advance of making arrangements for importing the material.
PACKAGING BIOLOGICAL MATERIALS FOR IMPORTATION
Importing Biological Materials
Imported biological materials are subject to packaging and shipping requirements of various federal and international regulations. Proper packaging is the primary consideration and of utmost importance in the safe transportation of hazardous materials. If the biological material being shipped is infectious or radioactive, this information has to be noted on the outside of the package. In addition, if dry ice is used for shipment, that must be clearly stated on the package. It is of course advantageous to the investigator that the shipped material be packaged in the most efficient way possible, as well as in a form which will ensure its stability. If proteins are to be shipped, it may be better to send the samples lyophilized or blotted on a membrane, rather than in liquid form. This will omit the need for dry ice or ice packs in the packaging, which will save on shipping costs as well as ensure the samples will be stable if receipt of the package meets with unforeseen delays. If sending materials that require dry ice or ice packs, such as antibodies or antisera, there must be enough dry ice to allow the package to maintain the appropriate temperature during shipping for approximately 5 days.
This requires utilizing much larger packaging boxes than are normally used to ship the same material within the United States. When shipping cell lines, sometimes it is better to send cells in culture, rather than frozen aliquots. To send cells in culture, the tissue culture flask containing the live cells must be filled to the very top with the appropriate medium, tightly capped, and the seal secured with parafilm. If the tissue culture flask is not completely filled with medium, the resulting air pockets will cause the flasks to explode in flight. Thus, the packaging of any biological material for shipping is an import consideration to ensure success of the scientific collaboration!
OBTAINING AN IMPORT PERMIT APPLICATION The Division of Veterinary Services of the USDA Animal and Plant Health Inspection Service (APHIS) administers regulatory programs to control the import/export of biological materials. Import permits can be obtained from either local USDA port offices or from APHIS directly: USDA, APHIS, VETERINARY SERVICES, National Center for Import-Export Products Program, 4700 River Road Unit 40, Riverdale, MD 20737-1231, Tel: (301) 7347830, Fax: (301) 734-8226. When submitting an import application, (1) obtain the import permit well in advance of the proposed shipping date; (2) inform the supplier of the USDA’s requirements, and do not allow shipment of materials until an import permit has been issued and the supplier has received a copy of the permit; (3) list all the potential U.S. ports of entry; (4) specify whether the material is for in vitro or in vivo use and whether it is for commercial distribution or for research in one’s own laboratory; (5) list any treatments the material has undergone, such as processing and purification steps involving pH, heat, chromatography, or other methods; and (6) indicate the source(s) of nutrient factors in culture media, cell line designation and history, whether enzymes were used, and what types of viruses (if any) are being studied in the laboratory of origin (all must be listed).
RESOURCES APHIS Web Site More information regarding APHIS programs and import/export can be obtained on their internet home page at http://www.aphis.usda.gov.
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Fax Service of Veterinary Services, National Center for Import-Export By calling (301) 734-4952, the investigator can access an automated document retrieval system and arrange to have an import application faxed directly, and choose from a menu of documents that describe general instructions
for completing the application, as well as information regarding the importation of biological materials from 18 distinct categories. Contributed by Paula Wolf Massachusetts Institute of Technology Cambridge, Massachusetts
Commonly Used Techniques
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Silanizing Glassware
APPENDIX 3E
Glassware is silanized (siliconized) to prevent adsorption of solute to the glass surface or to increase its hydrophobicity. This is particularly important when dealing with low concentrations of particularly “sticky” solutes such as single-stranded nucleic acids or proteins.
BASIC PROTOCOL
Materials Chlorotrimethylsilane or dichlorodimethylsilane Vacuum pump Desiccator, equipped with a valve 1. In a fume hood, place glassware or equipment to be silanized into desiccator along with a beaker containing 1 to 3 ml of chlorotrimethylsilane or dichlorodimethylsilane. CAUTION: Chlorotrimethylsilane and dichlorodimethylsilane vapors are toxic and highly flammable.
2. Connect desiccator to vacuum pump until silane starts to boil, and close connection to pump (maintaining vacuum in desiccator). Leave the desiccator evacuated and closed until liquid silane is gone (∼1 to 3 hr). During the incubation the silane will evaporate, be deposited on the surface of the glassware, and polymerize. Do not leave the desiccator attached to the vacuum pump. This will suck away the silane, minimizing deposition and damaging the pump.
3. Open desiccator in a fume hood, and leave open for several minutes to disperse silane vapors. 4. If desired, bake or autoclave the glassware or apparatus. Autoclaving or rinsing with water removes the reactive chlorosilane end of the dimethylsiloxane polymer generated by dichlorodimethylsilane.
COMMENTARY Untreated glass contains silicate and silanol groups that can act as ion-exchange and nucleophilic centers. To mask these groups and decrease the hydrophilicity of the surface, various reactive silanes are frequently used to coat the glass surface. The same or related chemistries can be used to introduce functional groups, including large molecules, onto the glass surface. On contact with a silanol (SiOH) group, chlorosilanes react to give HCl and a siloxane Si-O-Si linkage. For most applications “silanizing” or “siliconizing” a piece of glassware or equipment means introducing onto the glass surface either a short polymer of dimethylsiloxane Me Me Me Me | | | | glass−−O−−Si−−O−−Si−−O−−Si−− …O−−Si−−Cl | | | | Me Me Me Me
Contributed by Brian Seed Current Protocols in Protein Science (1998) A.3E.1-A.3E.2 Copyright © 1998 by John Wiley & Sons, Inc.
using dichlorodimethylsilane, or a trimethylsiloxane cap Me | glass−−O−−Si−−Me | Me
using chlorotrimethylsilane. Polydimethylsiloxane is silicone oil; the use of dichloromethylsilane essentially gives a light coating of oil to the glassware. The polymerization of dichlorodimethylsilane is mediated by residual water adsorbed to the glass surface; chlorosilanes also react with water and alcohols to give HCl and silanol or alkoxysilane, respectively. Thus, in the polymerization reaction, the residual water reacts with the chloride endgroup to give HCl and a silanol, which then reacts with more dichlorodimethylsilane and more water to give
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polymers. Chlorotrimethylsilane just gives a trimethyl cap to the silanol or silicate group. Items too large to fit in a desiccator can be silanized by briefly rinsing with or soaking in a solution of approximately 5% dichlorodimethylsilane in various volatile organic solvents such as chloroform or heptane. The organic solvent is removed by evaporation, depositing the dichlorodimethylsilane on the surface. This approach is particularly useful for
treating glass plates for denaturing polyacrylamide sequencing gels. CAUTION: If a flammable solvent is used, do not bake the glassware until the solvent is completely evaporated.
Contributed by Brian Seed Massachusetts General Hospital and Harvard Medical School Boston, Massachusetts
Silanizing Glassware
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Current Protocols in Protein Science
Protein Precipitation Using Ammonium Sulfate
APPENDIX 3F
BASIC THEORY The solubility of globular proteins increases upon the addition of salt (<0.15 M), an effect termed salting-in. At higher salt concentrations, protein solubility usually decreases, leading to precipitation; this effect is termed salting-out (Green and Hughes, 1955). Salts that reduce the solubility of proteins also tend to enhance the stability of the native conformation. In contrast, salting-in ions are usually denaturants. The mechanism of salting-out is based on preferential solvation due to exclusion of the cosolvent (salt) from the layer of water closely associated with the surface of the protein (hydration layer). The hydration layer, typically 0.3 to 0.4 g water per gram protein (Rupley et al., 1983), plays a critical role in maintaining solubility and the correctly folded native conformation. There are three main protein-water interactions: ion hydration between charged side chains (e.g., Asp, Glu, Lys), hydrogen bonding between polar groups and water (e.g., Ser, Thr, Tyr, and the main chain of all residues), and hydrophobic hydration (Val, Ile, Leu, Phe). In hydrophobic hydration, the configurational freedom of water molecules is reduced in the proximity of apolar residues. This ordering of water molecules results in a loss of entropy and is thus energetically unfavorable. When salt is added to the solution, the surface tension of the water increases, resulting in increased hydrophobic interaction between protein and water. The protein responds to this situation by decreasing its surface area in an attempt to minimize contact with the solvent—as manifested by folding (the folded conformation is more compact than the unfolded one) and then self-association leading to precipitation. Both folding and precipitation free up bound water, increasing the entropy of the system and making these processes energetically favorable. Timasheff and his colleagues provide a detailed discussion of these complex effects (e.g., Kita et al., 1994; Timasheff and Arakawa, 1997). It should be mentioned that the increase in surface tension of water by salt follows the well-known Hofmeister series, shown below (see Parsegian, 1995, and references therein). Hence, as an approximation, those salts that favor salting-out raise the surface tension of water the highest. As (NH4)2SO4 has much a higher solubility than any of the phosphate salts, it is the reagent of choice for salting-out. ← increasing precipitation (salting-out) ANIONS: PO34− > SO 24 − > CH 3COO − > Cl − > Br − > ClO 4− > SCN − CATIONS: NH 4+ > Rb + > K + > Na + > Li + > Mg 2 + > Ca 2 + > Ba 2 + increasing chaotropic effect (salting-in) → COMMON APPLICATIONS Concentration of Proteins Because precipitation is due to reduced solubility and not denaturation, pelleted protein can be readily resolubilized using standard buffers. After concentration, the protein is well suited for gel filtration (UNIT 8.3) whereby the buffer can be exchanged and the remaining ammonium sulfate removed. Alternately, the protein can be dissolved in a nonprecipitatContributed by Paul Wingfield Current Protocols in Protein Science (1998) A.3F.1-A.3F.8 Copyright © 1998 by John Wiley & Sons, Inc.
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ing concentration of (NH4)2SO4 (e.g., 1 M) and then applied to a hydrophobic interaction matrix (UNIT 8.4). Protein Purification Practical details of selective precipitation are presented in UNIT 4.5, and an example in the purification of interleukin 1β is given in UNIT 6.2. Salt precipitation has been widely used to fractionate membrane proteins (Schagger, 1994). Due to bound lipid and/or detergents, ammonium sulfate precipitates have lower density than protein-only precipitates. During centrifugation, these precipitates will often float to the top of tube rather than pelleting; the use of swing-out rotors is recommended. Crystallization is a traditional method of protein purification. Jakoby (1971) describes a general method that involves extracting (NH4)2SO4-precipitated protein with successively dilute (NH4)2SO4 solutions at low temperature. Folding and Stabilization of Protein Structure As mentioned above, (NH4)2SO4 and other neutral salts stabilize proteins by preferential solvation (Timasheff and Arakawa, 1997). Proteins are often stored in (NH4)2SO4, which inhibits bacterial growth and contaminating protease activities. Protein unfolded by denaturants such as urea can be pushed into native conformations by the addition of (NH4)2SO4 (Mitchinson and Pain, 1986). A practical application is the folding of recombinant proteins. For example, HIV-1 Rev expressed in E. coli was solubilized using urea, purified by ion-exchange chromatography in the presence of urea, then folded by the addition of 0.5 to 1.0 M (NH4)2SO4 (Wingfield et al., 1991). Basic Calculations Basic definitions Percentage (%) saturation concentration of (NH4)2SO4 in solution as % of maximum solubility at the given temperature. For example, at 0°C, a 100% saturated solution is 3.9 M. Specific volume (sp. vol.) volume occupied by 1 g of (NH4)2SO4 (ml/g) = inverse of density. At 0°C, if 706.8 g of (NH4)2SO4 is added to 1 L of water the volume = 1000 ml + volume occupied by the salt (706.8 × 0.5281 ml) = total volume of 1373.26 ml. The molarity = 3.9 M. Calculating quantities of (NH4)2SO4 to be added By weight. The following equation is used to calculate the weight of solid (NH4)2SO4 to be added to 1 liter of solution of initial concentration S1 to produce final saturation S2: G (S − S ) weight (g) = sat 2 1 1 − ( PS2 ) where: Gsat = grams of (NH4)2SO4 contained in 1 liter of saturated solution. For example, at 0°C, Gsat = 515.35 (see Table A.3F.1). S1 and S2 are fractions of complete saturation; for example, a 20% saturation is expressed as 0.2. Protein Precipitation Using Ammonium Sulfate
P = (sp. vol. × Gsat)/1000. For example, P = 0.2722 and 0.2945 at 0°C and 25°C, respectively.
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Table A.3F.1
Density and Molarity of Ammonium Sulfate Solutionsa,b
Temperature (°C) (NH4)2SO4 (g) added to 1 liter of water to give saturated solution (NH4)2SO4 (g) per liter saturated solution Molarity of saturated solution Density (g/ml) Specific volume in saturated solution (ml/g)
0
10
20
25
706.8
730.5
755.8
766.8
515.35 3.90 1.2428 0.5281
524.60 3.97 1.2436 0.5357
536.49 4.06 1.2447 0.5414
541.80 4.10 1.2450 0.5435
aMolecular bAdapted
weight of (NH4)2SO4 = 132.14. from Dawson et al. (1986).
By volume. The following equation is used to calculate volume of saturated (NH4)2SO4 solution to be added to 100 ml of solution to increase saturation from S1 to S2: volume (ml) =
100( S2 − S1 ) 1 − S2
For example, to raise 100 ml of 0.2 saturated solution to 0.70 saturation: 100 ml (0.2) + x ml (1.0) = (100 + x ml) 0.70 20 + x = 70 + 0.7x x = 166.66 ml Hence, 166.66 ml of saturated solution is added to 100 ml of 20% saturated solution to give 266.66 ml of 70% saturated solution. AMMONIUM SULFATE TABLES The tables shown are taken from Wood (1976). Table A.3F.2 gives the weight of (NH4)2SO4 to be added to a solution to obtain the desired concentration. Table A.3F.3 gives the volume of a 3.8 M solution to add to obtain a desired concentration. Tables A.3F.4 and A.3F.5 give the final volumes after the addition of the solid salt or a 3.8 M solution, respectively. The concentration of (NH4)2SO4 is expressed in molarity (corresponding % saturation is indicated in Table A.3F.2). The data is valid for solutions at 0°C, and the variation of specific volume with concentration is taken into account. For a table referring to solutions at 25°C, see Green and Hughes (1955). See following pages for tables.
Commonly Used Techniques
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Supplement 13
Initial molarity
0.00 0.20 0.40 0.60 0.80 1.00 1.20 1.40 1.60 1.80 2.00 2.20 2.40 2.60 2.80 3.00 3.20 3.40 3.60 3.80 3.90
Percent saturation
0.0 5.1 10.3 15.4 20.5 25.7 30.8 35.9 41.1 46.2 51.3 56.5 61.6 66.8 71.9 77.0 82.2 87.3 92.4 97.6 100.0
Final molarity
0.00 26.7 54.0 81.9 0.00 27.0 54.7 0.00 27.4 0.00
111 83.0 55.4 27.7 0.00
140 112 84.2 56.2 28.1 0.00
170 142 114 85.5 57.1 28.6 0.00
202 173 144 116 87.0 58.1 29.1 0.00
234 205 176 147 118 88.5 59.1 29.6 0.00
267 238 209 179 150 120 90.2 60.2 30.2 0.00
302 272 243 213 183 153 122 91.9 61.4 30.7 0.00
338 308 278 247 217 186 156 125 93.7 62.6 31.3 0.00
375 344 314 283 252 221 190 159 127 95.7 63.9 32.0 0.00
413 383 352 321 289 258 226 194 162 130 97.7 65.2 32.7 0.00
453 422 391 359 328 296 264 231 199 166 133 99.8 66.7 33.4 0.00
495 464 432 400 368 335 303 270 236 203 170 136 102 68.2 34.2 0.00
539 507 475 442 409 376 343 310 276 242 208 174 139 104 69.8 35.0 0.00
585 552 519 486 453 420 386 351 317 282 248 213 178 142 107 71.5 35.8 0.00
632 599 566 533 499 465 430 395 360 325 290 254 218 182 146 110 73.2 36.7 0.00
682 649 615 581 546 512 477 441 405 369 333 297 260 224 187 150 112 75.0 37.6 0.00
707 673 639 605 570 535 499 464 428 391 355 318 281 244 207 169 132 94.0 56.1 18.1 0.00
0.00 0.20 0.40 0.60 0.80 1.00 1.20 1.40 1.60 1.80 2.00 2.20 2.40 2.60 2.80 3.00 3.20 3.40 3.60 3.80 3.90
Grams of Ammonium Sulfate to Add to 1 Liter of Solution at 0°C
Table A.3F.2
Protein Precipitation Using Ammonium Sulfate
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0.00 0.20 0.40 0.60 0.80 1.00 1.20 1.40 1.60 1.80 2.00 2.20 2.40 2.60 2.80 3.00 3.20 3.40
Initial molarity
Final molarity
Milliliters of a 3.8 M Ammonium Sulfate Solution to Add to 1 Liter of Solution at 0°C
0.00 55.3 117 185 0.00 58.4 124 0.00 61.9 0.00
263 197 132 65.9 0.00
351 281 211 141 70.5 0.00
452 377 302 227 152 75.9 0.00
570 489 408 327 246 164 82.3 0.00
709 621 534 446 357 268 179 89.7 0.00
875 779 683 587 490 393 295 197 98.7 0.00
1077 972 866 760 652 545 437 328 219 110 0.00
1330 1213 1094 975 855 735 613 492 369 247 124 0.00
1655 1522 1387 1252 1115 978 840 702 562 423 282 141 0.00
2088 1933 1777 1620 1462 1303 1143 981 819 657 494 330 165 0.00
2693 2508 2322 2135 1946 1756 1565 1372 1179 984 789 593 396 198 0.00
3600 3371 3140 2907 2673 2437 2199 1959 1718 1475 1232 988 742 496 248 0.00
5111 4809 4503 4194 3884 3570 3255 2936 2616 2294 1971 1645 1319 991 662 332 0.00
8134 7684 7228 6768 6305 5837 5366 4891 4412 3931 3447 2961 2472 1981 1489 994 498 0.00
0.00 0.20 0.40 0.60 0.80 1.00 1.20 1.40 1.60 1.80 2.00 2.20 2.40 2.60 2.80 3.00 3.20 3.40
Table A.3F.3
Supplement 13
0.00 0.20 0.40 0.60 0.80 1.00 1.20 1.40 1.60 1.80 2.00 2.20 2.40 2.60 2.80 3.00 3.20 3.40 3.60 3.80 3.90
Initial molarity
Final molarity
Final Volume in Millimeters After Addition of Solid Ammonium Sulfate to 1 Liter of Solution at 0°C
1000 1010 1023 1033 1000 1011 1023 1000 1012 1000
1046 1035 1024 1012 1000
1060 1049 1037 1025 1013 1000
1074 1063 1052 1039 1027 1014 1000
1090 1079 1067 1054 1042 1028 1014 1000
1106 1095 1083 1070 1057 1044 1030 1015 1000
1123 1112 1100 1087 1074 1060 1046 1031 1016 1000
1142 1130 1118 1105 1091 1077 1063 1048 1032 1016 1000
1161 1149 1137 1124 1110 1096 1081 1065 1050 1033 1017 1000
1181 1169 1157 1143 1129 1115 1100 1084 1068 1052 1035 1018 1000
1203 1191 1178 1164 1150 1135 1120 1104 1088 1071 1054 1036 1018 1000
1225 1213 1200 1186 1172 1156 1141 1125 1108 1091 1073 1056 1037 1019 1000
1249 1237 1223 1209 1194 1179 1163 1147 1130 1112 1094 1076 1058 1039 1019 1000
1275 1262 1248 1233 1218 1203 1187 1170 1152 1135 1116 1098 1079 1060 1040 1020 1000
1301 1288 1274 1259 1244 1228 1211 1194 1176 1158 1140 1121 1101 1082 1062 1041 1021 1000
1329 1316 1301 1286 1271 1254 1237 1220 1202 1183 1164 1145 1125 1105 1085 1064 1043 1022 1000
1359 1345 1330 1315 1299 1282 1265 1247 1228 1209 1190 1170 1150 1129 1109 1087 1066 1044 1022 1000
1373 1359 1344 1329 1313 1296 1278 1260 1242 1222 1203 1183 1162 1142 1121 1099 1077 1055 1033 1011 1000
0.00 0.20 0.40 0.60 0.80 1.00 1.20 1.40 1.60 1.80 2.00 2.20 2.40 2.60 2.80 3.00 3.20 3.40 3.60 3.80 3.90
Table A.3F.4
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0.00 0.20 0.40 0.60 0.80 1.00 1.20 1.40 1.60 1.80 2.00 2.20 2.40 2.60 2.80 3.00 3.20 3.40
Initial molarity
Final molarity
Final Volume in Millimeters After Addition of 3.8 M Ammonium Sulfate Solution to 1 Liter of Solution at 0°C
1000 1051 1109 1174 1000 1055 1117 1000 1059 1000
1248 1187 1126 1063 1000
1333 1268 1202 1135 1068 1000
1432 1362 1291 1219 1147 1074 1000
1547 1471 1394 1317 1239 1160 1080 1000
1683 1601 1517 1433 1348 1262 1176 1088 1000
1847 1757 1665 1573 1479 1385 1290 1194 1097 1000
2047 1947 1846 1743 1640 1535 1430 1324 1216 1108 1000
2298 2186 2072 1957 1841 1723 1605 1486 1365 1244 1122 1000
2621 2493 2363 2232 2099 1966 1831 1694 1557 1419 1280 1141 1000
3051 2902 2751 2598 2444 2289 2131 1973 1813 1652 1491 1328 1164 1000
3654 3476 3294 3112 2927 2741 2553 2363 2171 1979 1785 1590 1394 1198 1000
4560 4337 4111 3883 3652 3420 3185 2948 2709 2469 2227 1984 1740 1494 1248 1000
6070 5773 5472 5168 4862 4552 4240 3924 3606 3287 2965 2642 2316 1989 1661 1331 1000
9091 8646 8196 7741 7282 6818 6350 5878 5402 4922 4441 3956 3469 2979 2488 1994 1498 1000
0.00 0.20 0.40 0.60 0.80 1.00 1.20 1.40 1.60 1.80 2.00 2.20 2.40 2.60 2.80 3.00 3.20 3.40
Table A.3F.5
LITERATURE CITED Dawson, R.M.C., Elliot, D.C., Elliot, W.H., and Jones, K.M. 1986. Data for Biochemical Research (3rd ed.), p. 537. Oxford Science Publications, Clarendon Press, Oxford. Green, A.A. and Hughes, W.L. 1955. Protein solubility on the basis of solubility in aqueous solutions of salts and organic solvents. Methods Enzymol. 1:67-90. Jakoby, W.B. 1971. Crystallization as a purification technique. Methods Enzymol. 22:248-252. Kita, Y., Arakawa, T., Lin, T.-Y., and Timasheff, S. 1994. Contribution of the surface free energy perturbations to protein-solvent interactions. Biochemistry 33:15178-15189. Mitchinson, C. and Pain, R.H. 1986. The effect of sulphate and urea on the stability and reversible unfolding of β-lactamase from Staphylococcus aureus. J. Mol. Biol. 184:331-342. Parsegian, V.A. 1995. Hopes for Hofmeister. Nature 378:335-336. Rupley, J.A., Gratton, E., and Careri, G. 1983. Water and globular proteins. Trend Biochem. Sci. 8:1822.
Timasheff, S.N. and Arakawa, T. 1997. Stabilization of protein structure by solvents. In Protein Structure: A Practical Approach. 2nd ed. (T.E. Creighton, ed.) pp. 349-364. IRL Press at Oxford University Press, Oxford. Wingfield, P.T., Stahl, S.J., Payton. M.A., Vankatesan, S., Misra, M., and Steven, A.C. 1991. HIV-1 Rev expressed in recombinant Escherichia coli: Purification, polymerization and conformational properties. Biochemistry 30:7527-7534. Wood, W.I. 1976. Tables for the preparation of ammonium sulfate solutions. Anal. Biochem. 73:250-257.
KEY REFERENCE Wood, 1976. See above. Tables A.3F.2-A.3F.5 are reprinted from this publication.
Contributed by Paul Wingfield National Institutes of Health Bethesda, Maryland
Schagger, H. 1994. Chromatographic techniques and basic operations in membrane protein purification. In A Practical Guide to Membrane Protein Purification (G. von Jagow and H. Schagger, eds.) pp. 23-57. Academic Press, San Diego.
Protein Precipitation Using Ammonium Sulfate
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Statistics: Detecting Differences Among Groups In this appendix, we will consider some of the statistical tests most commonly used (and misused) in biological research. The tests discussed here are those used for comparisons among groups (e.g., t test and ANOVA). A number of other important areas (e.g., linear regression, correlation, and goodness-of-fit testing) are not covered. The purpose of the Appendix is to enable you to determine rapidly the most appropriate way to analyze your data, and to point out some of the most common errors to avoid. Toward this end, we have included a flow chart to provide a quick guide to choosing the right statistical test (Fig. A.3G.1). Instead of including the voluminous statistical tables necessary to perform these tests, we assume you will have access to a spreadsheet software program such as Microsoft Excel or to a statistical software package to do the actual calculations involved and to supply critical values. We include calculations for some simpler tests to aid in the interpretation of test statistics. Clearly, this Appendix is a very superficial treatment of statistics! There are of course many statistics texts available; a few that the authors have found particularly useful are listed at the end of this Appendix (see Literature Cited and Key References).
BASIC STATISTICAL BACKGROUND Samples and Independence It is important to think about how you will collect your data and what statistical tests you will perform before you begin your experiment. Suppose you want to compare the immune response of normal mice with that of mice carrying a mutation of interest. You examine a sample of mice from each of these populations. For any statistical test to be valid, all subjects in a sample must come from the same population, and subjects must be selected independently of each other. For example, selecting the same mouse twice would violate the independence assumption; on the other hand, selecting mutant mice from different genetic backgrounds to form one sample would violate the assumption that all mice in a sample are coming from the same population. For some experiments, subjects may be measured twice (or more) by design, and this information is
incorporated into the statistical test. For example, mouse immune response could be measured before and after a treatment (see discussion of Paired t Test), or several measurements may be made from each mouse (see discussion of Multiple Comparison Testing). However, each mouse in the sample is still chosen independently of other mice.
Hypothesis Testing Statistical tests always test a null hypothesis, H0. The null hypothesis usually states that there is no difference between two (or more) populations. If the null hypothesis is true, any observed differences between samples from these populations arose by chance alone. The alternative hypothesis, Ha, states that differences observed in the samples reflect a real difference between the populations from which they were drawn. We usually hope to reject H0, in order to show something interesting. In our immune response experiment, for example, our H0 would be that mutant mice have the same response as normal mice. We hope to be able to reject H0, and show that mutant mice have a different immune response from normal mice. Although statistical tests can never tell us with absolute certainty whether our samples arose from two different populations, they can give us an idea about how probable our H0 is. What we would like to know is the probability that H0 is true. Unfortunately, statistical tests cannot tell us this probability. What they do tell us is a related probability—i.e., the probability that we would see at least as large a difference between samples as we do, given that H0 is true. This probability is the famous p value. For example, if p = 0.50, it means that there was a 50% chance of getting a difference between samples at least as large as we did, if the populations we sampled were really the same. This is a large probability, so we would accept the null hypothesis H0, and conclude that there is no difference between the two populations we sampled. The accepted standard is that we should reject H0 if α ≤ 0.05—i.e., there was only a 5% chance (or less) of getting at least as large a difference between samples as we did, if the populations sampled were really the same. The value of p below which we will reject the H0 is called the α value; α is usually chosen as 0.05. A difference between samples that
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results in a p value of ≤α is called statistically significant. Note that statistical tests are not black and white; if α = 0.05, there is a 5% chance that we will reject the null hypothesis when it is actually true! This is called a Type I error. You might think we could avoid this error by setting α to an even smaller value, say, 0.01 or 0.001. However, if we do this, we are likely to accept H0 when it is false! This is called a Type II error. Setting α = 0.05 is a standard that
no
data in samples independent?
is generally accepted as a good compromise between Type I and Type II errors. In practice, once you have chosen a statistical test, you calculate (or the computer calculates) the value of a statistic from your sample data using a formula. You then compare this value to a critical value in a table (or provided by the computer) to decide whether to accept or reject H0. The critical value (e.g., Fc for the F test, or tc for the t test; see discussion of The
no
paired data, or repeated measures?
yes
redesign experiment
yes
calculate descriptive statistics mean and standard deviation); construct histograms of data
no
samples approximately normal with similar variances?
may try data transformation, especially if data are proportions, ratios, or counts
yes yes
repeated measures?
transformed data approximately normal with similar variances
no
no yes
repeated measures?
yes
2 samples?
no
Friedman test
no yes
means same? yes
ANOVA
paired?
2 samples?
done
variances equal?
paired t test yes
no Mann-Whitney test
no
no
t test assuming t test assuming equal variances unequal variances or Mann-Whitney test
Statistics: Detecting Differences Among Groups
means same?
yes
yes Wilcoxon signed rank or sign test
parametric multiple comparisons parametric tests
Kruskal-Wallis test
paired?
no
yes
no
yes
done
no
nonparametric multiple comparisons nonparametric tests
Figure A.3G.1 Flow chart for comparisons among groups.
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t Test) is determined by the α value you have decided upon (usually, α = 0.05) and the degrees of freedom (df) of the data, which vary depending on the test being performed. Usually, you reject H0 if your statistic is greater than the critical value. Most tables (and statistical packages) give critical values with one-tailed and two-tailed levels of significance. You are doing a twotailed test when your Ha does not specify the direction of difference between populations. You perform a one-tailed test when your Ha predicts, based, for example, on a theory that you have, that populations will differ in a particular way. For example, if you think that there may be a difference between the immune response of mutant and normal mice, but you can’t predict whether the mutants will do better or worse, you should perform a two-tailed test. If you predict that the response of the mutant mice will be reduced (due to the loss of your gene of interest), you could perform a onetailed test. There is some controversy in the literature over the use of one-tailed tests. When in doubt, perform a two-tailed test; it is always more conservative (i.e., less likely to reject the null hypothesis).
CHOOSING A TEST— EXPLORATORY DATA ANALYSIS
What Are Parameters? In order to perform parametric tests, we need to calculate parameters—i.e., numbers that summarize the data in some descriptive way. The statistical parameters we want to calculate are the mean (average) and amount of variability of the population (variance and standard deviation). Your data are a sample from a large population. We estimate the population parameters by sampling a subset of the population. The larger the sample size (n), the closer your sample statistics are to the true population parameters. Any spreadsheet program or statistical package will allow you to determine the mean, variance, and standard deviation of your data; we show the formulas for these parameters below so that you can better interpret what such programs are telling you about the data. The_ following is the equation for the sample mean x: n
∑x
i
i =1
x=
n
Here, xi is the value of the ith data point, and n is the number of data points in the sample. The sample variance (s2) is expressed as follows: n
Parametric and Nonparametric Tests It can be difficult to sort out what test is appropriate to apply to your data when confronted with a bewildering array of possible tests and formulas. One important distinction you need to make is between parametric and nonparametric tests. Parametric tests (such as the t test) assume your data follow a statistical distribution with known mathematical properties (the normal distribution, for the parametric tests we will discuss). Often, parametric tests require that the variability of all the samples you are comparing be the same. Nonparametric tests make fewer assumptions about the distribution of your data. Nonparametric tests are thus less likely than parametric tests to commit a Type I error (rejecting H0 when it is true), but are usually less powerful (that is, more likely to commit a Type II error, and fail to detect a difference between groups). However, with data sets of reasonable size (say, 25 or more data points per group), nonparametric tests are often nearly as powerful as parametric ones. In order to determine which type of test is appropriate to your data, the first thing you need to do is examine the characteristics of that data.
s = 2
∑ (x
i
− x)
i =1
n −1
The sample standard deviation (s), is just the square root of the variance: s=
s2
You can think of the standard deviation as the average deviation from the mean; the magnitude of s tells you how much variability there is in the data. If your data are normally distributed, then ∼68% of the data should fall within 1 standard deviation of the mean and ∼95% of your data should fall within 2 standard deviations of the mean. The final parameter of interest is the standard error of the mean (s.e.m.; also called the standard error): s.e.m. =
s
n The standard error of the mean tells you how much the mean of a sample of size n taken from your population varies. That is, if you were to
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take a number of different samples of size n from your population, the mean of each sample would be slightly different. If you were to plot the means of a number of such samples, they would be distributed across a range of values, but there would be far less variability in those values than in the data of the original samples. While the standard deviation gives you a description of the variability of your data, the s.e.m. is the parameter used to compare two sample means and to determine whether or not they are statistically different. For this reason, the s.e.m. is what is normally plotted as the “error bars” around the mean on a graph showing samples that are being compared. Note that the s.e.m. does not tell you much about the variability of your original data; you can see from the equation for s.e.m. that even highly variable data can give a small s.e.m. if the sample size is large enough. Suppose we are interested in the ability of neurons to sprout neurites in response to growth factors. We want to compare a population of neurons exposed to nerve growth factor (NGF), a factor of known importance, with a population exposed to factor X. Our H0 would be that neurite outgrowth in response to NGF is the same as neurite outgrowth in response to factor X. We hope to be able to reject H0, and show
Table A.3G.1
Neurite Outgrowth Data
Neurite outgrowth (µm) NGF
Factor X
22.5 35.4 38.2 40.4 41.3 46.7 55.4 62.5 81.2 99.2
43.2 56.9 60.2 62.4 65.7 75.8 85.4 91.3 95.3 105.2
Table A.3G.2 Sample Parameters from Neurite Outgrowth Data
Statistics: Detecting Differences Among Groups
_ Mean (x) Variance (s2) Standard deviation (s) Standard error (s.e.m.)
NGF
Factor X
52.28 535.00 23.13 7.31
74.14 388.26 19.70 6.23
that neurons respond differently to factor X. Tables A.3G.1 and A.3G.2 show a (rather small) data set, with calculated sample parameters. If these data are normally distributed, then ∼68% of the data for NGF should fall within one standard deviation of the mean—i.e., between 29.15 and 75.41 (52.28 ± 23.13). In fact, 7⁄ or 70% of the data points fall within these 10 limits. About 95% of the data should fall within two standard deviations of the mean—between 6.02 and 98.54 (52.28 ± 2 × 23.13). In fact, 90% of the data fall within these limits. The distribution of factor X data is similar. This is reasonable agreement with the normal distribution, given the small sample sizes.
Determining Whether Data Are Normally Distributed with Equal Variances In order to determine whether to use a parametric or nonparametric test on your data, it is important to determine whether the data are normally distributed, with equal variances. An example of a statistical test for normality found in many computer packages is the KolmogoravSmirnoff test (Zar, 1984). We will discuss tests for homogeneity of variance later. However, you can often find out most of what you need to know about your data by simply plotting it and looking at it. For reasonably large data sets, the parametric tests that we will discuss are not much affected by small departures from normality (this property is called being “robust” to small departures from normality). As a rule of thumb, if you have ≥25 samples in each group that you are comparing, you can simply plot your data using a frequency histogram and examine it visually to see if it falls roughly into a bell-shaped curve. Our neurite outgrowth data is plotted in Figure A.3G.2. In this example, the data points are continuous numbers, so in order to make a histogram, we had to divide the data up into arbitrary “bins” (e.g., 0 to 20, 20 to 40, and so forth). The samples in our data set look approximately normally distributed, although the sample sizes are a bit small to be really sure. The variability in the two data sets also looks similar by eye. Looking back at our sample parameters, the calculated variance and standard deviation of the two data sets look reasonably similar. If you have a small data set, it may be difficult to determine visually whether your data are normally distributed, and statistical tests also do not have much power to make the determination. If you use a parametric test in this situation, and the data are really not drawn
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from a normal distribution, then you may reject your null hypothesis when it is actually true. If you use a nonparametric test, you may accept your null hypothesis when it is false. It is more conservative to use nonparametric tests in this situation; but you can see that the best solution (when possible) is to collect reasonably large data sets! If visual inspection suggests your data are not normally distributed, or if the variance differs widely among groups, it is sometimes useful to perform a transformation (see discussion of Data Transformations). The transformed data points may then form a normal distribution with similar variances among samples, and parametric statistical tests can be performed on the transformed data points. If neither the original nor the transformed data produce approximately normally distributed data with similar variances, then nonparametric statistical tests generally should be used.
Data Transformations The parametric tests that we will describe usually assume that the data are normally distributed, and that the variances are the same for all groups being compared (this latter property is known as “homoscedasticity”). There are a number of common situations—e.g., in which the data are expressed as proportions or counts of randomly occurring objects or events— where these assumptions are violated, but you
can still use a parametric test if you first perform a data transformation. Data expressed as proportions of 0 to 1.00 (i.e., 0% to 100%) are not normally distributed. However, you can make them nearly normally distributed, with reasonably homogeneous variances among groups, by performing the following transformation for each data point, and then using the p′ values for all of your calculations (e.g., mean, standard deviation, and test statistic):
p′ = arcsin
p
When the data are counts of randomly occurring objects (such as the number of butterflies per square foot of a field) or events (such as the number of cars passing by a location), the data will follow a Poisson rather than a normal distribution. Group variances will then be proportional to their means. In this case, the square root transformation, shown in the following equation, should be performed. As with proportions, statistical tests are then performed on the transformed values.
x′ =
x + 0.5
If variances among groups are very different and if the standard deviations are proportional to the means of your groups, the logarithmic transformation can be used:
5
Number of neurites
4
3
2
1
0 20
40
60
80
100
120
Neurite outgrowth (µ m)
Figure A.3G.2 Neurite outgrowth histogram. Black bars correspond to results for nerve growth factor (NGF) gray bars correspond to results for “factor X.”
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x ′ = log( x + 1) If your data are expressed as ratios, a logarithmic transformation is likely to make them more nearly normally distributed (for an example, see discussion of Ratio Paired t Tests). Other types of transformations are possible (Zar, 1984). If your data still don’t fit the assumptions of a parametric test after being transformed, then a nonparametric test should be used.
STATISTICAL TESTS FOR COMPARISONS BETWEEN TWO UNPAIRED GROUPS The t Test The t test is a parametric test that allows us to determine whether the means of two samples are statistically different. The nonparametric equivalent is the Mann-Whitney test (see discussion below; also called the Wilcoxon Rank Sum test). Assumptions of the t test 1. Only two samples are being compared. If more than two samples are compared, you must use a test specifically designed for multiple comparisons, such as ANOVA (see discussion under Multiple Comparison Testing). Using multiple t tests to compare among three or
more samples is invalid, and the computed p values will be wrong. This is one of the most common errors in the use of the t test. 2. Subjects in samples were chosen independently. If repeated measurements were made from individual subjects, ANOVA (or the nonparametric equivalent) should be used. 3. The two samples were not paired in any way (such as before-and-after measurements or matched cases and controls). If samples were paired, the paired t test (or the nonparametric Wilcoxon Signed Rank test; see discussion below) should be used. 4. The samples are approximately normally distributed. If this assumption is not met, either transform your data (see discussion of Data Transformations) or use the nonparametric Mann-Whitney test instead. The simplest form of the t test assumes that both samples have the same variance. A somewhat more complicated form is necessary if the sample variances are different. Therefore, in order to determine which form of the t test to use, we must first determine whether the sample variances are statistically different, by using the variance ratio test, or F test. Figure A.3G.3 shows a flow chart for the t test. Performing the F test The F test is summarized as follows:
perform F test (H0: sample variances are the same; s 1 2 = s 22 )
accept H 0 (variances are the same)
perform t test assuming equal variances (H0: sample means are the same; x 1 = x 2 )
Statistics: Detecting Differences Among Groups
reject H 0 (variances are different)
perform t test assuming unequal variances (H0 : sample means are the same; x 1 = x 2); if variances are wildly different, use a nonparametric test or try a data transformation
Figure A.3G.3 _ _ Flow chart for the t test. Here, we want to compare sample 1 and sample 2, with means x1 and x2, variances s12 and s22, and sample sizes n1 and n2. Whichever t test you use, accepting H0 for the t test says that the sample means are the same; rejecting H0 for the t test says that the sample means are different.
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H0 : s12 = s22 F=
s12 s22
or F =
s22 s12
, whichever is larger
(F will always be ≥ 1) reject H0 if F > Fc ; accept H0 if F < Fc In the above equations, F is referred to as the “F statistic,” s12 and s22 are the variances of sample 1 and sample 2, respectively, and Fc is the critical value. Statistical computer packages will report the F statistic, along with the appropriate critical value, Fc. Note that if the variances of the two samples are very different, F will be large, so it makes sense to reject H0 if F > Fc. If we reject H0, we would use the t test assuming unequal variances. If we accept H0, we would use the t test assuming equal variances. If the differences in variance are extreme, a data transformation may be indicated or the (nonparametric) MannWhitney test could be performed instead of the t test. Fc is chosen based on the significance level (usually α = 0.05), and the degrees of freedom (df) in the data. The value of df is equal to one less than the number of data points in each sample. In the case of the experiment in Table A.3G.1, which will be used as an example here, there are 10 data points in each sample, so df = 9 for each sample. Fc is usually written “Fnumerator df, denominator df, α.” For our example, Fc would be written F9,9,0.05(2). The 2 in parentheses denotes that this is a two-tailed critical value. If the computer package reports both a one-tailed and two-tailed critical value, use the two-tailed value. You would only use a onetailed value if you had reason to think before computing the variances that one variance would be larger than the other. When in doubt, use a two-tailed test. Performing the t test The t test with equal variances and sample sizes is summarized as follows:
H0 : x1 = x2 t=
x1 − x2 s12 + s22 n1
reject H0 if t > tc ; accept H0 if t < t c
In the above equations, t is referred to as the “t statistic” and tc is the critical value. Statistical packages will report t and tc. To choose tc from a table, the number of degrees of freedom is calculated as the sum of the number of data points in samples 1 and 2, minus 2 (df = n1 + n2 − 2). The critical value, tc, is often written tdf,0.05(2) for a two-tailed test. As with the F test, use a two-tailed test if both oneand two-tailed critical values are given. Examining the formula for the t statistic, you can see that t will be large if the difference between the sample means is large. However, as the variances of the samples get larger, the t statistic gets smaller. Thus, the equation makes intuitive sense; you will be able to reject the null hypothesis if the difference between means is large and the scatter in the data is small, so that there is little overlap between the two samples. There are several different formulas for the t test with unequal variances, which give slightly different results; they are not presented here (see Zar, 1984). The t test with unequal variances tests the same H0 and follows the same intuitive reasoning given above for the t test with equal variance. However, the calculation of df for this test is quite different than for the test with equal variance—they are calculated using a more complicated formula—a statistical package will calculate them, and print them for you. Example of the t test Let us assume that our neurite outgrowth data (see Table A.3G.1 and Table A.3G.2) fit all the assumptions for the t test, and let us perform the t test on this data set. H0 for the F test: variances of the two samples are equal. F = 535.00/388.26 = 1.378. Fc = F9,9,0.05(2) = 4.03. Thus, F < Fc, so we accept H0 and conclude that the variances of the two samples are not statistically different. We proceed to the t test assuming equal variances. H0 for the t test: means of the two samples are equal. We use a spreadsheet to calculate the t statistic. The computer reports: t = 2.275 tc = t18,0.05(2) = 2.10. Thus, t > tc, so we reject H0 and conclude that the means of the two samples are different, with statistical significance at the α = 0.05 level. The calculated p value for our data is p = 0.035.
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This means that the probability of observing by chance a difference in means at least as large as our data show is only 3.5%. Note that some spreadsheets print out negative values for the t statistic; in this case, simply use the absolute value.
(e.g., if two values tie for 8th largest, they would have been 8th and 9th largest, they each get a rank of 8.5). 2. Sum the ranks for each sample. Call the sums R1 and R2. 3. Calculate the U and U′ statistics according to the following equations:
The Mann-Whitney Test The Mann-Whitney test (also called the Wilcoxon Rank Sum test) is the nonparametric equivalent to the t test, and is used to determine if two samples are statistically different, with no assumptions about whether the data are normally distributed. This test is nearly as powerful as the t test. When you perform the Mann-Whitney test, you are comparing the order, or rank, of the data rather than its numerical value. To perform the Mann-Whitney test, you rank your entire data set (treating all data from both samples as a single list). For our neurite outgrowth data, the largest data point is 105.2; it has a rank of 1. The next largest is 99.2, with a rank of 2, and so on. You then sum the ranks for each individual data set. If your two samples have similar values, the ranks would tend to go back and forth between samples. For example, the highest-ranking value might be in sample 1, the next two highest in sample 2, the next in sample 1, and so on. If you then summed the ranks from each sample, the two sums would be about equal. On the other hand, if sample 1 had mostly larger numbers than sample 2, the ranks of sample 1 would all be small numbers, and the sum of the ranks of sample 1 would be much smaller than that of sample 2. The Mann-Whitney test tells us how different the rank sums must be to indicate a significant difference between the two samples. Assumptions of the Mann-Whitney test Assumptions 1 to 3 of the t test (see discussion of The t Test) apply also to the Mann-Whitney test. The Mann-Whitney test, however, makes no assumption about normality of data.
Statistics: Detecting Differences Among Groups
Performing the Mann-Whitney test The Mann-Whitney test is summarized as follows. H0: There is no significant difference between the two samples (of sizes n1 and n2). 1. Order all the data from both samples into a single list, from greatest to smallest value, keeping track of which sample the data came from. Tied values get the average of the two ranks they would have had if they had not tied
U = n1n2 +
n1 (n1 + 1) 2
− R1
U ′ = n1n2 − U 4. If either U or U′ is greater than the critical value, Uc (U0.05(2),n1,n2, in a table of U values; see, e.g., Zar, 1984), then reject H0. 5. With more than 20 measurements in one or both samples, the Z statistic is used, as calculated by the following equation. Only U or U′ needs to be calculated in this case, not both.
Z=
U − U − 0.5 s
where U= s=
n1n2 2
and
n1n2 (n1 + n2 + 1) 12
The Z statistic is approximately normally distributed, even though the original data may not be at all normally distributed. For α = 0.05, if Z > 1.96, H0 is rejected. Example of the Mann-Whitney test Let us say that we were a bit uncomfortable assuming that our neurite outgrowth data (see Table A.3G.1 and Table A.3G.2) are really normally distributed, and we decide to perform the Mann-Whitney test instead of the t test. These data are shown ranked for the Mann-Whitney test in Table A.3G.3. From the data in Table A.3G.3 we can make the following calculations, based on the above equations for the Mann-Whitney test: U = 100 + 55 − 134 = 21 U′ = 100 − 21 = 79 Uc = U0.05, 10, 10 = 77 Since U′ > Uc, H0 is rejected and it is concluded that the two samples are significantly different. Note that both the t and Mann-Whitney statistics were very close to the critical values for these data. The power of the Mann-Whitney
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Table A.3G.3 Neurite Outgrowth Data Ranked for the Mann-Whitney Test
Outgrowth
Rank (NGF)
NGF 22.5 35.4 38.2 40.4 41.3 46.7 55.4 62.5 81.2 99.2 Factor X 43.2 56.9 60.2 62.4 65.7 75.8 85.4 91.3 95.3 105.2 Rank sum
Rank (factor X)
20 19 18 17 16 14 13 9 6 2
— — — — — — — — — —
— — — — — — — — — — 134
15 12 11 10 8 7 5 4 3 1 76
test to reject H0 is almost as great as that of the t test.
COMPARING TWO PAIRED GROUPS: PAIRED t TEST, WILCOXON SIGNED RANK TEST, AND SIGN TEST When to Use a Paired Analysis A paired analysis is indicated under the following circumstances. 1. You measure a variable before and after some treatment is performed upon a subject. 2. You match subjects in pairs (e.g., for age, sex, and exposure) and then treat one of the subjects and not the other (or the other receives alternative treatment). 3. You compare relatives (e.g., siblings or parent/child). 4. You perform an experiment several times, each time with experimental and control preparations treated in parallel.
Paired t Test The paired t test is a parametric test used to compare two groups whose members are paired. When the assumptions of a paired t test are met, another approach is to perform a repeated-measures ANOVA (see discussion under Multiple Comparison Testing) with only two measures. However, most nonstatisticians find the paired t test easier to understand and perform. Assumptions of the paired t test 1. Pairs must be representative of a larger population. Each pair must be selected independently of other pairs. 2. Pairs must be made before data is collected. 3. The calculated differences between the two members of each pair must be approximately normally distributed. Performing the paired t test The paired t test is summarized as follows.
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Assume you have a set of n pairs: x11, x21, x12, x22,....through x1n, x2n. H0: There is no difference between members of pairs. _ (i.e., the mean difference between the pairs, d, is equal to 0). 1. For each pair, calculate the difference between the pairs (di = x1i − _ x2i). 2. Calculate the mean, d , and the s.e.m. of these differences. 3. Calculate the t statistic according to the following equation:
t=
d
s.e.m. 4. Determine tc from a table using df = n − 1. Note that each pair counts as 1 in computing the df; for the unpaired t test, each data point counted as 1. 5. If t > tc, reject H0. Example of the paired t test For our neurite outgrowth data, suppose that instead of looking at two different sets of neurons treated separately with NGF and factor X, we were looking at neurons treated sequentially, first with NGF and then with factor X. We were able to keep track of each individual neurite, and want to know if neurites continued to grow out in the presence of factor X after NGF was removed. We will analyze the same data as before, but now it is paired, as shown in Table A.3G.4 and Figure A.3G.4. H0: There is no change in neurite outgrowth when factor X is added to the medium _ d = 21.86 s.e.m. = 5.03 t = 21.86/5.03 = 4.35 tc = t9,0.05(2) = 2.26 t > tc; therefore reject H0. The p value for the paired t test is p = 0.0018 (the p value for the unpaired test on the same data was p = 0.035). Clearly, having data that are paired can give us much more power to reject H0. If we are uncertain whether our differences are normally distributed, we can use a nonparametric alternative (either the Wilcoxon Signed Rank test, or the Sign test; see discussion below).
Ratio Paired t Tests Statistics: Detecting Differences Among Groups
In many situations, such as measurement of enzyme activity or protein expression, data are presented graphically as percent of control, making statistical analysis difficult. The ratio paired t test allows a straightforward analysis
of such data. For example, you might want to look at densitometric data from a western blot of your favorite gene’s expression in a mutant animal, compared to the signal from a wild-type animal. For each experiment, the densitometry units can be adjusted arbitrarily, and may vary widely depending on how good your probe is and how long the exposure was. However, the ratio of the mutant (m) to wild-type (wt) readings is what is important. In this case, what you really want to know is whether the ratio of mutant to wild-type is significantly different from 1 (implying that gene X is expressed either more or less in the mutant than in the wild type). Ratios are not normally distributed, but logarithms of ratios are quite likely to be normally distributed. The logarithm of a ratio can be written as a difference:
m = log(m) − log(wt) wt
log
Thus, to analyze data of this type, simply take the logarithm of each data point and perform a paired t test on this transformed data (Table A.3G.5). If the transformed data do not appear to be approximately normal, the Wilcoxon Signed Rank test can be used instead. The null hypothesis for the transformed data is that the mean difference is 0, as before; this means that for the original (untransformed) data, the null hypothesis is that the ratio of mutant to wild-type expression is 1. Rejecting the null hypothesis says that the amount of expression in mutant and wild-type animals is significantly different. Let us perform the calculation as follows on the data in Table A.3G.5. H0: Ratio of mutant to wild-type expression = 1 (there is no difference between mutant and wild-type animals). Transform data, and perform paired t test of differences of logarithms of data as described for the paired t test. _ d = 0.1843 s.e.m. = 0.019755 t = 9.33 tc = t2,0.05(2) = 4.30 t > tc Thus, reject H0 and conclude that there is a significant difference in expression between mutant and wild-type animals. (the calculated p value was 0.01). Note that this is a very significant result, even with only three pairs of data.
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Table A.3G.4
Paired Neurite Outgrowth Data
Neurite outgrowth (µm) NGF
Factor X
22.5 35.4 38.2 40.4 41.3 46.7 55.4 62.5 81.2 99.2
43.2 56.9 65.7 62.4 75.8 85.4 95.3 60.2 105.2 91.3
Difference 20.7 21.5 27.5 22.0 34.5 38.7 39.9 −2.3 24 −7.9
6
Number of neurites
5 4 3 2 1 0 10
0
20
30
40
Change in outg rowth (µm)
Figure A.3G.4 Change in neurite outgrowth after addition of factor X, in cells treated first with NGF.
Table A.3G.5 Original and Transformed Densitometry Data from Western Blotting of Mutant Versus Wild-Type Gene Expressiona
Mutant
Wild-type
500
300
6000 750
4200 500
Ratio: mutant/ wild-type
log(mutant)
1.67 1.43 1.5
2.6990 3.7781 2.8751
log(wild-type) 2.4771 3.6232 2.6990
log(mutant) − log(wild-type) 0.2218 0.1549 0.1761
aValues expressed in arbitrary densitometry units.
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Table A.3G.6 Differences Computed from Neurite Outgrowth Data, and Ranked for Use in the Wilcoxon Signed Rank Test
Difference
Rank (−)
20.7 21.5 27.5 22.0 34.5 38.7 39.9 −2.3 24 −7.9 Rank Sum
— — — — — — — 1 — 2 3
Wilcoxon Signed Rank Test The Wilcoxon Signed Rank test is a nonparametric equivalent to the paired t test. It is used for data that may not be normally distributed. Assumptions of the Signed Rank test Assumptions 1 and 2 of the paired t test apply also to the Wilcoxon Signed Rank test. An additional assumption is made that the sampled population is symmetrical about the median. If the data do not fit this assumption very well, the Sign test (see discussion below) can be performed. Performing the Signed Rank test The Wilcoxon Signed Rank test is summarized as follows. H0: There are no differences between members of pairs. 1. For each pair, calculate the difference between the members of the pair. Keep track of the sign. 2. Rank the absolute value of all differences from low to high. 3. Sum the ranks of the negative differences (call the sum T−) and the ranks of the positive differences (call the sum T+). 4. Reject H0 if either T− or T+ is less than or equal to the critical value, Tc, as determined from a Wilcoxon Signed Rank table.
Statistics: Detecting Differences Among Groups
Example of the Signed Rank test If we were concerned about the normality of our neurite outgrowth differences (Table A.3G.1 and Table A.3G.2), we could perform the Wilcoxon Signed Rank test on these data
Rank (+) 3 4 7 5 8 9 10 — 6 — 52
rather than the paired t test we did earlier. Consider the data in Table A.3G.6. H0: There is no change in neurite outgrowth when factor X is added to the medium. T− = 3; T+ = 52 Tc = 8 T− < Tc; therefore reject H0.
The Sign Test The Sign test is another nonparametric paired test that comes in especially handy when you are sure of the direction of a difference between the members of your pair (positive or negative), but you may not know exactly how much difference there is. Assumptions of the Sign test The assumptions of the Sign test are the same as those of the Wilcoxon Signed Rank test, but no symmetry assumption is made. This is a less powerful test. Performing the Sign test The Sign test is summarized as follows. H0: There are no differences between members of pairs. 1. For each pair, record whether the first member is greater than the second (+) or less than the second (−). Ignore any pairs with equal members. C+ is the total number of (+) pairs; C− is the total number of (−) pairs. 2. Reject H0 if either C+ or C− is less than or equal to the critical value of C for the Sign test (Cc).
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Example of the Sign test H0: There is no change in neurite outgrowth when factor X is added to the medium. For our neurite outgrowth data (Table A.3G.6), C+ = 8 and C− = 2. For n = 10 and p = 0.05, the critical value, Cc is equal to 1. Therefore, H0 cannot be rejected.
MULTIPLE COMPARISONS Why Not Lots of t Tests? If we find that we want to compare multiple groups in an experiment, then the t test just will not do. Using pairwise comparisons ignores the experimental effects in our design in all but the two groups being compared. Even if it were appropriate to use multiple pairwise comparisons (and it is not), we would still run afoul of the problem of hypothesis rejection. If we used the typical α = 0.05, that would mean that for every 20 pairwise tests we ran, we would expect to get the wrong answer for one of them. This is not a desirable situation. There are a number of methods, however, that avoid these problems by allowing us to consider our entire design in our model for analysis. The most common in current literature is the analysis of variance (ANOVA). There are classical and nonparametric methods to analyze variance differences, and we will first consider the classical, or parametric case.
Assumptions of ANOVA As with any statistical test, there are several assumptions of the classical analysis of variance that need to be examined. Some of these are more critical than others, and we will try to make that clear. In this chapter, we are dealing only with one-way analysis of variance; additional assumptions hold for more complex ANOVA procedures. If in doubt about the ability of your own data to meet these criteria, use nonparametric alternatives, for which the assumptions are less stringent. Independent observations Observations must be independent even if you end up doing nonparametric analysis. Data points need to have been collected such that the magnitude of one observation does not affect another. Certain tests, such as the repeatedmeasures analysis of variance or Friedman’s nonparametric analysis of variance, however, allow for multiple measurements on a single subject. Care should be taken to avoid interdependence of data during your experimental design because no amount of adjustment or
transformation can rectify the problems inherent in nonindependent data. An example of nonindependent data would be taking three measurements of pH from a single cell culture dish and entering them as n = 3 in your ANOVA. In this case, n = 1, and you are just as well off taking a single measurement as you are averaging the three readings. Continuous variable This assumption is not an extremely strong one as long as you bear several things in mind. The data need to be numerical and ordinal. Continuous data, that is, data that can take on any possible numerical value, are the kind of data for which ANOVA was designed to be used. Deviations from this will affect the sensitivity of the test. It is permissible for data to be discrete (having only a certain set of possible values, like counting numbers) if the numerical values have a fairly wide range. For ANOVA, this would mean something on the order of a 1 to 10 scale or greater. The numbers applied to observations must have some tie to a “greater than or less than” scale; they cannot simply be a classification system of your data. If you have discrete data, you should pay closer attention to the adherence of your data to the additional assumptions detailed below, or just choose a nonparametric test from the outset. Homogeneity of variance This is probably the strongest assumption of ANOVA, since, as we will see later, the whole test hinges on the variability within each of our experimental groups being similar. As in the case of the t test (see discussion of The t Test), the variances for each group need to be tested for similarity. The big difference here, however, is that for the greater-than-two-groups case, there is no way to continue with the test if the variances of the groups are significantly different. Instead, we need to transform the data mathematically to make the variances more similar, or move on to a nonparametric equivalent. Checking for homogeneity of the variances should start with drawing a frequency histogram of each group and seeing how spread out the data are. It is also a good idea to calculate variances for each group and see if the numbers look close. In many cases, this is all the checking that needs be done. If you remain unsure, there is a simple test called the Fmax test, which is a simple matter of setting up the following ratio: Commonly Used Techniques
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largest group variance in the study smallest group variance in the study
= Fmax
This is equivalent to using an F ratio of two groups in the t test (see discussion of The t Test), but the tabled critical value to which it is compared is from a cumulative F distribution. Degrees of freedom are calculated as n − 1, where n is the number of observations in a single group. If n varies among groups, then the lowest value should be chosen, to be most conservative in the test. As in the case of the t test, if you fail to reject the null hypothesis, then the variances can be assumed to be roughly equivalent. Normality Although this is touted as an important assumption in most classical statistics, it is not as strong an assumption in practice for ANOVA as the assumption of homogeneous variances. Again, using a frequency histogram to give an idea of the shape of your distribution is often a good enough test of normality (see discussion of Determining Whether Data Are Normally Distributed with Equal Variances). If the data are distributed with a marked hump toward the average value and trailing off to either side, then ANOVA will probably work fine. Computer programs provide several different tests of normality that can also be used for this purpose.
Partitioning the Variance It may seem odd that we wish to analyze variance when, in the long run, our question is to find the differences in means among groups in our experiment. Analysis of variance, however, takes advantage of the underlying assumption that the variation in a measurable quantity is similar within all groups in a population. One or more of these groups, however, may have a very different mean relative to the other groups. If this were the case, the total variance of the whole population would increase relative to the amount seen in each group. It is this observation that gives us the null hypothesis for ANOVA, which is that the means of all the groups are the same: H0: µ1 = µ2 = µ3 = ... = µn
Statistics: Detecting Differences Among Groups
In ANOVA, we break down the total variance into two basic components: within-group and among-group variance. Within-group or error variance is the naturally occurring variance among individuals due to inherent genetic differences and the differential effects of environment that each individual experiences. This
contrasts with among-group or treatment variance, which is the variance observed among groups due to perturbations of the experiment. The treatment variance is the mathematical difference between the total variation observed in our experimental model and the natural variance measured across individuals (the error variance). Partitioning the variance into components allows us to establish the ratio of the two components of variance, defined as Fsas shown in the following equation: among-group (treatment variance) within-group (error) variance
= Fs
Testing the Null Hypothesis After partitioning the variance into its two components (see discussion of Partitioning the Variance), we can then compare our calculated Fs to the F distribution, which is tabled in numerous statistics texts. The tables are usually organized with the critical values listed for many combinations of the numerator (treatment) versus denominator (error) degrees of freedom. Treatment degrees of freedom are the number of groups minus 1, and the error degrees of freedom are the total number of individual observations in all groups minus the number of groups. While it is unlikely that you will ever have to look up tabled F values, since computers do it for us, it is important to understand the convention used for F ratios. The F value given for a particular tested proportion is written as: F(treatment df, error df, α) = critical value from table. So, the critical value of F for an hypothetical experiment with a rejection level (α) of 0.05, four treatment groups, and a total experimental population of 48 organisms would be written as: F(3,44,0.05) = 2.82. If α is chosen as 0.05, it is often omitted from the statistical reporting for brevity’s sake. This critical value is important in our analysis because, if our calculated F ratio gives a value higher then the critical value from the table, we reject our null hypothesis. For ANOVA, the null hypothesis is that all the means for all the groups are equal. Rejecting the null hypothesis tells us only one thing: that at least one of the groups differs from the others. It does not tell us how many differ or from how many of the other groups any one might differ. For this answer we need to proceed to a method of multiple-means comparisons.
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Table A.3G.7 Hypothetical Optical Density Data for Bacterial Cultures
Control 0.53 0.57 0.6 0.49 0.42 0.4 0.41
Treatment A
Treatment B
0.37 0.44 0.44 0.39 0.62 0.58 0.6
Treatment C
0.52 0.55 0.59 0.59 0.7 0.72 0.73
0.7 0.66 0.72 0.69 0.55 0.53 0.5
Table A.3G.8 Descriptive Summary Table for Hypothetical Optical Density Data Shown in Table A.3G.7
Groups Control Treatment A Treatment B Treatment C
Count
Sum
Mean
Variance
7 7 7 7
3.42 3.44 4.4 4.35
0.488571 0.491429 0.628571 0.621429
0.006581 0.011081 0.007448 0.008381
Table A.3G.9 Single-Factor ANOVA Table for Hypothetical Optical Density Data Shown in Tables A.3G.7 and Table A.3G.8a
Source of Variation
SS
df
MS
F
p value
Fc
Among groups Within groups Total
0.12778214 0.20094286 0.328725
3 24 27
0.042594 0.008373 —
5.09 — —
0.01 — —
3.01 — —
aAbbreviations: F , critical value of F statistic; SS, sum of squares; MS, mean square. c
The ANOVA Table Assuming a hypothetical experiment such as the one summarized in Table A.3G.7 and Table A.3G.8, the output from any ANOVA computer package is likely to produce an ANOVA table similar to Table A.3G.9, originally generated by Microsoft Excel. The data are hypothetical optical density readings for plasmid-transformed bacteria that have been transformed by three different experimental protocols, along with a control that has not been transformed. The first part of the data analysis is a simple descriptive summary table (Table A.3G.8) that gives the sample size, mean, and variance for each group. Based on this table, it is fairly clear that the variances within groups are similar, but that the means for some of the groups seem to be different from others. The ANOVA table (Table A.3G.9) then gives the variance compo-
nents and degrees of freedom accounted for among and within groups. The “SS” column indicates the sum of squares for each component, and the “MS” indicates mean square. Sum of squares is the summation of the square of the amount by which each observation differs from the total mean. The mean square is the same as the variance, and is calculated by dividing the SS by the appropriate degrees of freedom. The mean square within-groups value is often called the “mean square error,” and figures prominently into tests of differences among the means. The table also reports the calculated F ratio for the among versus within variance comparison. The computer then provides the exact p value associated with this ratio and sample size, and further indicates the critical F value that is necessary for rejection at the 0.05 level. Commonly Used Techniques
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Which Means Are Different? We have finally reached the point where we are ready to answer the question we asked, which was whether the average value of one or more of our groups differs from the others. While statisticians differ on this issue, it is the authors’ opinion that if you fail to reject the null hypothesis of your ANOVA you should not test for significant means differences unless you planned certain comparisons before you ran the experiment. That having been said, it is possible that you will find significant means differences even if you fail to reject the ANOVA null hypothesis. If that is the case, however, it is worth asking how robust your initial test was—did you have large-enough sample sizes, were your variances “equal enough,” and were the distributions of your data groups normal or at least all similarly shaped? If not, you would probably be better off reworking your analysis with a nonparametric equivalent. Assuming that you have rejected H0, how can you tell which means differ? Again, a first approximation is to look at the average values for each group and see which appear most different from the others. Next, you will want to proceed to a test that will provide you with a p value for those differences. For the special the case of testing a control versus treatments, the Dunnett test is appropriate. Other test names you might see are Bonferroni or Tukey; computer packages have many options and you should refer to the statistical documentation of your package to determine which option best suits your data. Since means comparisons are cumbersome to do by hand and are part of most statistical packages, we will not go into the specific calculations here. Make sure, however, that you understand how the computer is reporting the significance of your data to you. The computer will usually give you a p value. As you perform more mean-versus-mean comparisons, your acceptable α level will change, and the computer rarely keeps track of this, so it is important to know how to calculate these adjustments in α. One of these techniques is discussed below.
Statistics: Detecting Differences Among Groups
Planned or unplanned comparisons The use of certain techniques in your final means comparison will depend, in part, on whether you planned before the experiment among which groups you were going to look for differences (a priori tests) or whether you waited to peruse your data first (a posteriori). This turns out to be a theoretically very complex subject and one that is not worth delving into
here. A detailed treatment can be found in Sokal and Rohlf (1981). It is worth pointing out here a common mistake in choosing significance levels for multiple means comparisons. In general, planned comparisons allow the researcher a greater rejection level (bigger α or p value) than unplanned comparisons. The reasoning is simple: if you first look at your data, then choose your comparisons, you are introducing a bias to your analysis. To account for this, the level of rejection needs to be tightened. Even if you have planned your comparisons, however, it is easy to abuse the power of multiple means tests. For example, declaring a priori that you are going to conduct all possible pairwise comparisons in your experiment is tantamount to using multiple t tests (see discussion of Why Not Lots of t Tests?). If you have k groups in your analysis, the number of all possible pairwise comparisons would be k(k − 1)/2 Since each comparison is not independent of all others, there is a need to alter the significance level of the tests. In order to use the experiment-wise error (i.e., α = p = 0.05), the degrees of freedom for all the tests you perform should not exceed k − 1. So, if you have four groups (a,b,c,d), you might declare to test a versus b,c,d (1 df); a versus d (1 df); and b,c versus d (1 df). That would burn up 3 degrees of freedom at α = 0.05. Note that in the very common case of comparing each treatment to a control, we stay within this limit as well (assume a is the control and compare it pairwise with each of the three treatments b,c,d). If we were to go beyond this level of comparison, however, we would need to adjust the rejection level for all the tests we perform to account for a loss of independence. A simple and conservative technique that does this is the Bonferroni method (Sokal and Rohlf, 1987). To make an adjustment to the experiment-wise error term, α, calculate a test significance, α′, by dividing the experiment-wise error by the number of degrees of freedom used in your selected tests (dfT): α′ =
α dfT
Applying this technique to the example above, if we choose to compare the control a to each of the three treatment groups (3 df), and perform two additional internal treatment tests of b versus c,d (1 df) and c versus d (1 df), we have a total of 5 degrees of freedom, which exceeds our “allowable” number by 2. The
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Table A.3G.10 Turbidity Data (see Table A.3G.7) Scored on a Scale of 1 (least turbid) to 10 (most turbid)
Control
Treatment A
Treatment B
Treatment C
5 5 5 5 6 5 5
1 3 2 3 2 5 1
4 4 6 5 6 7 6
8 9 9 8 8 9 10
Table A.3G.11
Turbidity Data (see Table A.3G.10) Ranked from 1 to N
Control
Rank sums Rank averages
11.5 11.5 11.5 11.5 18.5 11.5 11.5 87.5 12.50
Treatment A 1.5 5.5 3.5 5.5 3.5 11.5 1.5 32.5 4.64
Bonferroni method would then require that we adjust α′ as follows:
α′ =
0.05
= 0.01 5 Therefore, for all the tests we perform, including all three of the control-versus-treatment tests, we have decreased our rejection level to p < 0.01, a level that could make a big difference in the significance of our results. This should point out the importance of good experimental design and of choosing only those tests that are important to the experimental goals.
The Nonparametric ANOVA Equivalent If the data you end up with do not meet the assumptions of ANOVA, you can still perform multiple comparisons with a nonparametric test. For a one-way ANOVA, the nonparametric equivalent most popularly used is the KruskalWallis one-way analysis of variance. The assumptions of the Kruskal-Wallis test are less stringent than those of the parametric ANOVA. The samples are drawn from a population with a continuous distribution and the sampling must be random. The samples must be inde-
Treatment B 7.5 7.5 18.5 11.5 18.5 21 18.5 103 14.71
Treatment C 23 26 26 23 23 26 28 175 25.00
pendent. Their values must be rankable, that is, on at least an ordinal scale. Many computer programs provide the option of calculating the Kruskal-Wallis statistic. It is simple to do by hand, and is worth outlining here. In the hypothetical example of the optical densities for transformed bacteria strains (see Table A.3G.7, Table A.3G.8, and Table A.3G.9), assume that the researcher, instead of using a spectrophotometer, simply used an ordinal scale of 1 to 10 to approximate the turbidity of the sample (see Table A.3G.10). (Okay, so it’s not very precise science, but, who knows, maybe the spec was broken that day and the samples needed to be measured!) In this case, there is an expectation that the scores would not be distributed in such a way as to meet the criteria for a parametric ANOVA. To proceed through the analysis, it is then necessary to find the average value for the ranks of the data in each of the groups. To do this, we will give the data across all groups a rank score. The smallest number will have the lowest rank, and so on until we have ranked all the data. Equivalent values are termed “ties” and are given the average rank of the tied observations (see Table A.3G.11).
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From these ranks, the test statistic (KW) is calculated according to the following formula:
KW =
12
k
∑n R N ( N − 1) j
j =1
2 j
− 3( N + 1)
In the above equation, N = total population size, nj = number of observations in a group (note that groups need not__be equal in size), k = number of groups, and Rj= rank average for each group. For small samples (usually k ≤ 3 with nj ≤ 5) there are special tabled values for the KW statistic. For larger samples, the resulting value is compared to a χ2 distribution with (k − 1) degrees of freedom. If the KW value is larger than the tabled valued at a chosen α, then H0 is rejected. As in the case of the parametric ANOVA, a significant result implies that at least one of the groups is different from the rest. A means comparison test needs to be done to show which group it is. For the data in Table A.3G.11, the calculation is as follows:
12 7(12.5)2 + 7(4.64)2 (28)(29) +7(14.71)2 + 7(25)2 − [3(28 + 1)]
KW =
= 18.45 The critical value for α = 0.05, with df = (k − 1) = 3, is 7.82. Since our KW test statistic is larger than that, our test is significant and we reject H0. A problem that we might have encountered with our data, however, is that we ended up with many tied ranks. There are mathematical corrections for ties that will yield a slightly higher KW statistic than the calculation without the correction. This means that if you reject H0 without the correction, you will also reject with the correction, so it is unnecessary if your results are significant. If, on the other hand, if your significance is marginal and you are concerned about the ties in your data, you may choose to perform a ties correction. Siegel and Castellan (1988) gives one such correction. For our example, the ties correction would yield KW = 19.0.
Nonparametric Means Comparison Statistics: Detecting Differences Among Groups
Having demonstrated a significant difference somewhere among the groups with the Kruskal-Wallis test (see discussion of The Nonparametric ANOVA Equivalent), it is then appropriate to proceed to find out where those
differences are. The technique that we will apply will allow us to perform only pairwise comparisons, but it is adjusted so that all possible pairwise comparisons can be made with the same level of rejection. Treatment versus treatment Multiple comparisons among all treatments, as in the case of ANOVA, require attention to the error inherent in the entire experiment relative to each pairwise test we choose to do. To account for this, we adjust our test statistic by the total possible pairwise comparisons in the experiment. Whether we plan to do only some of them or all of them is irrelevant to the adjustment; however, there is a special case, outlined below, if all we wish to do is compare a single control to several treatments. The many possible differences between pairs of observations in a data set will become approximately normal as the sample size increases, regardless of the underlying distribution of the data. This allows us to use the normal distribution probabilities to test for significant differences between groups. In effect, we can use the normal distribution to establish a confidence interval for the difference between two groups. If they fall beyond this value, we can say that their rank average values differ significantly. To be able to use the normal approximation, we first need to adjust the value we look up in a normal probability table (z) by the number of pairwise group comparisons possible in the experiment:
zα′ = z α
k ( k −1)
In this equation, k is the number of groups in the experiment. For the data in Table A.3G.11, if we chose an experiment-wise α = 0.05, this would yield zα′ = z0.05/(4)(3) = z0.004. The numerical value of zα′ can be looked up on a normal table or derived from a computer program. In our case, z0.004 = 2.65. Now that we have determined the correct normal approximation for our test, we apply it to determine the critical value for rejection at α = 0.05 (not α′ = 0.004) between any two groups in the population. If the mathematical difference between the two rank averages of the groups is greater than the critical value, then we reject at p < 0.05. The calculation of the critical value (CV) is dependent on sample sizes:
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Table A.3G.12 Pairwise Comparisons of Hypothetical Turbidity Ranked Data in Table A.3G.11
Significancea
Comparison
Calculation
Control vs. A Control vs. B Control vs. C A vs. B A vs. C B vs. C
|12.5 − 4.64| = 7.86 |12.5 − 14.71| = 2.21 |12.5 − 25| = 12.5 |4.64 − 14.71| = 10.07 |4.64 − 25| = 20.36 |14.71 − 25| = 10.29
NS NS * NS * NS
aNS, not significant; asterisk refers to differences significant at p < 0.05.
CV = zα′
N ( N + 1) 1 12
1 + nu nv
In the above equation, N is the total population size and nu and nv are the sample sizes of the two groups that you are comparing (they may be different sizes). For our example (see Table A.3G.11), since all group sizes are the same, the CV will be identical for any and all of the six possible pairwise comparisons. If sample sizes are different, you need to calculate a new CV for each comparison: CV = 2.65
28(29) 1
1 + 12 7 7
= 11.65 To determine which differences are significant, we first calculate the absolute value of the difference between all pairs of rank averages (Table A.3G.12). If the difference exceeds the critical value, then the difference can be determined to be significant at p < 0.05. From this analysis it is clear that while the entire model was quite significant the differences in the data set are due mostly to the large value of Treatment C. Control versus treatment The test above provides a powerful nonparametric tool for comparing multiple samples. A researcher often is interested, however, in a less broad model—one that looks only at the differences between a control and two or more treatments. Following the same procedure as above, it is possible to approximate these differences with a normal distribution. A different correction is needed, however, to account for fewer,
more defined comparisons. In this case the following equation applies:
zα′ = z α
2( k −1)
For our example, this equation would yield zα′ = z0.005/(2)(3) = z0.008 = 2.41. Because fewer comparisons are being done, this smaller z value will lead to a smaller critical value and provide greater probability of finding a difference between the control and a treatment. Furthermore, if we expected a priori that our treatments would yield a directional result (either smaller or greater, but not both) relative to the control, we could reduce this number even more by adjusting by α/(k − 1) rather than α/2(k − 1). Since we did not have this expectation in our data, we will use the two-tailed option and calculate as follows:
CV = zα′
N ( N + 1) 1 12
1 + nc nu
In the equation above, N is the total population size, nc is the sample size of the control, and nu is the sample size of the treatment being compared. Again, because all of our treatments have the same number of observations, we need calculate only one value for CV. CV = 2.41
28(29) 1
1 + 12 7 7
= 10.60 This value is then compared as above to the differences between the control and treatment rank averages. We would reject any difference greater than 10.60. Despite the tighter rejection
Commonly Used Techniques
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Table A.3G.13 Hypothetical Immunoprecipitation Data for Repeated-Measures ANOVA
IgG precipitate (mg/ml) Mouse ID A B C D E F G H
Day 1
Day 2
Day 3
Day 4
Day 5
0 0.7 0.09 0.01 0.6 0.2 0 0.2
0 2.5 1.8 0.01 5.5 1.1 0.05 0.5
0.02 3.2 4 0.6 6.8 2.4 0.9 0.7
0.7 4.5 7.1 1.8 9.6 2.5 3 2.5
1.3 3 2.2 0.1 8.5 1.5 2.7 6
Table A.3G.14 Repeated-Measures ANOVA Table for Hypothetical Immunoprecipitation Data in Table A.3G.13a
Source of variation
SS
df
Rows Columns Error Total
121.8885 68.29064 55.75068 245.9298
7 4 28 39
MS 17.41265 17.07266 1.991096 —
F
p value
8.745258 8.574505 — —
1.12E-05 0.00012 — —
Fc 2.359258 2.714074 — —
aAbbreviations: F , critical value of F statistic; SS, sum of squares; MS, mean square. c
interval (compared with 11.65 from the all comparisons model) the only treatment that differs significantly from control at p < 0.05 is Treatment C. Note that the previously significant comparison (A versus C) is not valid in this model.
Repeated Measures—A Special Case Sometimes we wish to collect more than one sample from an individual organism in our study. Given that we would expect there to be less variability in samples taken from one individual than in samples taken among several individuals, it would seem that we need to adjust our analysis of variance somewhat. And indeed we do. Instead of a simple one-way analysis of variance, we perform a repeatedmeasures analysis of variance. This technique allows us to statistically control for the variance attributable to the inherent differences in the test subjects relative to the variance in response to the treatments. Once you determine that a difference exists somewhere in the data set, you can test for means differences in the same way as described above for one-way ANOVA. Statistics: Detecting Differences Among Groups
ANOVA. For our parametric ANOVA example, let us consider the following case (see Table A.3G.13). You are producing an antibody in a strain of mice and want to know when antibody production reaches a peak. To collect data, you perform immunoprecipitation on serum collected for 5 days from eight study animals. As you can see, certain animals have higher values overall than others, some respond with IgG production very quickly, others lag a bit and so on. If we were to do a one-way ANOVA on these data, the variability among the animals (rather than across days) might compromise the significance of our test. To avoid this problem, the repeated-measures ANOVA calculates variances for the rows and columns separately. The final output for such an analysis would resemble Table A.3G.14, originally produced by Microsoft Excel. In this table there are significance values reported for the rows (mice) and columns (days). For our example, both factors are clearly significant. This implies that there is a difference in the precipitation of IgG over the 5 days of measurement and a difference in IgG expression in different mice.
Parametric The same assumptions hold true for the repeated measures ANOVA as for the one-way
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Table A.3G.15 Hypothetical Immunoprecipitation Data in Table A.3G.13 Ranked for Friedman’s Analysis of Variance
IgG precipitate (ranks)
Mouse ID A B C D E F G H Rank sums
Day 1
Day 2
1.5 1 1 1.5 1 1 1 1 9
1.5 2 2 1.5 2 2 2 2 15
Nonparametric If you have repeated measures that violate the assumptions of the parametric ANOVA, then there is a nonparametric equivalent for the repeated-measures ANOVA as well. As in the turbidity example above, a common case would be when a response was scored on an ordinal scale rather than measured on a continuous one. The nonparametric equivalent in this case is the Friedman’s analysis of variance. The data are handled in a fashion similar to the KruskalWallis test (see discussion of The Nonparametric ANOVA Equivalent), except that ranks are assigned only within an individual rather than across the entire data set. In this way, an individual that overall got higher responses than the others would not contribute unduly to the overall variance of the population. To illustrate, let us simply use the immunoprecipitation data in Table A.3G.13. We begin by ranking the values within each individual from 1 to 5 (see Table A.3G.15). We then sum the ranks for each column. Under the null hypothesis, the ranks should be distributed in random order for any individual, which would result in the column rank sums being approximately equal. Clearly the rank sums are not equal, but are the differences significant? We can determine this by using the Friedman test and comparing the resulting value to a χ2 distribution with k − 1 degrees of freedom for our selected α level of rejection. The calculation of the statistic (Fr) is similar to the calculation of the Kruskal-Wallis statistic:
Fr =
12
k
∑R Nk (k + 1) j =1
2 j
− 3N (k + 1)
Day 3
Day 4
Day 5
3 4 4 4 3 4 3 3 28
4 5 5 5 5 5 5 4 38
5 3 3 3 4 3 4 5 30
In the above equation, N is the number of rows (subjects), k is the number of columns (treatments), and Rj is equal to the rank sum of each column. For our example, this would yield the following:
12 (81 + 225 + 784 + 1444 + 900) (8)(5)(6)
Fr =
− (3)(8)(6) Fr = 27.7 The χ2 critical value for α = 0.05 with 4 degrees of freedom is 9.48. Since our Fr statistic is greater than that, we reject the null hypothesis and conclude that there are differences in IgG production over the course of 5 days (columns), irrespective of differences inherent in the test subjects (rows).
CONCLUSION While the methods for dealing with comparisons of data outlined above are just a small sampling of those available, we hope they add to the arsenal of tools available to biologists. By performing simple descriptive analyses of your data and following the guidelines for test selection outlined in this Appendix, you should be able to avoid many of the statistical pitfalls frequently encountered in groups comparisons.
LITERATURE CITED Siegel, S. and Castellan, N.J., Jr. 1988. Nonparametric Statistics for the Behavioral Sciences, 2nd ed. McGraw-Hill, New York. Sokal, R.R. and Rohlf, F.J. 1981. Biometry, 2nd ed. W.H. Freeman, New York. Sokal, R.R. and Rohlf, F.J. 1987. Introduction to Biostatistics, 2nd ed. W.H. Freeman, New York.
Commonly Used Techniques
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Zar, J.H. 1984. Biostatistical Analysis, 2nd ed. Prentice-Hall, Englewood Cliffs, New Jersey.
KEY REFERENCES Hampton, R.E. 1994. Introductory Biological Statistics. William. C. Brown Publishers, Dubuque, Iowa. Provides excellent outlines for many statistical tests and uses relevant, comprehensible biological examples.
Motulsky, H. 1995. Intuitive Biostatistics. Oxford University Press, New York. A nice, nonmathematical introduction to biostatistics, covering standard topics as well as many areas important to biology that are often omitted from introductory texts.
Contributed by Phil Robakiewicz and Elizabeth F. Ryder Worcester Polytechnic Institute Worcester, Massachusetts
Statistics: Detecting Differences Among Groups
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Analyzing Radioligand Binding Data
APPENDIX 3H
A radioligand is a radioactively labeled drug that can associate with a receptor, transporter, enzyme, or any protein of interest. The term ligand derives from the Latin word ligo, which means to bind or tie. Measuring the rate and extent of binding provides information on the number of binding sites, and their affinity and accessibility for various drugs. While physiological or biochemical measurements of tissue responses to drugs can prove the existence of receptors, only ligand binding studies (or possibly quantitative immunochemical studies) can determine the actual receptor concentration. Radioligand binding experiments are easy to perform, and provide useful data in many fields. For example, radioligand binding studies are used to: 1. Study receptor regulation, for example during development, in diseases, or in response to a drug treatment. 2. Discover new drugs by screening for compounds that compete with high affinity for radioligand binding to a particular receptor. 3. Investigate receptor localization in different organs or regions using autoradiography. 4. Categorize receptor subtypes. 5. Probe mechanisms of receptor signaling, via measurements of agonist binding and its regulation by ions, nucleotides, and other allosteric modulators. This unit reviews the theory of receptor binding and explains how to analyze experimental data. Since binding data are usually best analyzed using nonlinear regression, this unit also explains the principles of curve fitting with nonlinear regression. For more general information on analyses of receptor data, see books by Limbird (1996) and Kenakin (1993). BINDING THEORY The Law of Mass Action Binding of a ligand to a receptor is a complex process involving conformational changes and multiple noncovalent bonds. The details aren’t known in most cases. Despite this complexity, most analyses of radioligand binding experiments successfully use a simple model, called the law of mass action: → ligand + receptor ← ligand ⋅ receptor The model is based on these simple ideas: 1. Binding occurs when ligand and receptor collide (due to diffusion) with the correct orientation and sufficient energy. The rate of association (number of binding events per unit of time) equals [ligand] × [receptor] × kon, where kon is the association rate constant in units of M−1 min−1. 2. Once binding has occurred, the ligand and receptor remain bound together for a random amount of time. The rate of dissociation (number of dissociation events per unit time) equals [ligand ⋅ receptor] × koff, where koff is the dissociation rate constant expressed in units of min−1. 3. After dissociation, the ligand and receptor are the same as they were before binding. Commonly Used Techniques Contributed by Harvey Motulsky and Richard Neubig Current Protocols in Protein Science (2000) A.3H.1-A.3H.55 Copyright © 2000 by John Wiley & Sons, Inc.
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The equilibrium dissociation constant Kd Equilibrium is reached when the rate at which new ligand⋅receptor complexes are formed equals the rate at which they dissociate:
[ligand ] × [receptor ] × kon = [ligand ⋅ receptor ] × koff Rearrange to define the equilibrium dissociation constant Kd.
[ligand] × [receptor ] = koff [ligand ⋅ receptor ] kon
= Kd
The Kd, expressed in units of moles/liter or molar (M), is the concentration of ligand that occupies half of the receptors at equilibrium. To see this, set [ligand] equal to Kd in the equation above. In this case, [receptor] must equal ligand⋅receptor, which means that half the receptors are occupied by ligand. Affinity The term affinity is often used loosely. If the Kd is low (e.g., pM or nM), that means that only a low concentration of ligand is required to occupy the receptors, so the affinity is high. If the Kd is larger (e.g., µM or mM), a high concentration of ligand is required to occupy receptors, so the affinity is low. The term equilibrium association constant (Ka) is less commonly used, but is directly related to the affinity of a compound. The Ka is defined to be the reciprocal of the Kd, so it is expressed in units of liters/mole. A high Ka (e.g., > 108 M−1) would represent high affinity. Because the names sound familiar, it is easy to confuse the equilibrium dissociation constant (Kd, in molar units) with this dissociation rate constant (Koff, in min −1 units), and to confuse the equilibrium association constant (Ka, in liter/mole units) with the association rate constant (Kon, in M−1 min−1 units). To help avoid such confusion, equilibrium constants are written as capital “K” and the rate constants with a lower case “k.” A wide range of Kd values are seen with different ligands. Since the Kd equals the ratio koff/kon, compounds can have different Kd values for a receptor either because the association rate constants are different, the dissociation rate constants are different, or both. In fact, association rate constants are all pretty similar (usually 108 to 109 M−1 min−1, which is about two orders of magnitude slower than diffusion), while dissociation rate constants are quite variable (with half-times ranging from seconds to days). Fractional occupancy at equilibrium Fractional occupancy is defined as the fraction of all receptors that are bound to ligand. The law of mass action predicts the fractional receptor occupancy at equilibrium as a function of ligand concentration. fractional occupancy =
[ligand ⋅ receptor ] = [ligand ⋅ receptor ] [receptor ]total [receptor ] + [ligand ⋅ receptor ]
A bit of algebra creates a useful equation. Multiply both numerator and denominator by [ligand] and divide both by [ligand ⋅ receptor]. Then substitute the definition of Kd. fractional occupancy= Analyzing Radioligand Binding Data
[ligand ] [ligand ] + Kd
The approach to saturation as [ligand] increases is slower than one might imagine (see Fig. A.3H.1). Even using radioligand at a concentration equal to nine times its Kd will only lead to its binding to 90% of the receptors.
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0 1 4 9 99
0% 50% 80% 90% 99%
Percent occupancy at equilibrium
Fractional occupancy [ligand]/K d at equilibrium
100
50
0 0 1
2 3 4 5 6 7 8 9 10 [Radioligand]/K d
Figure A.3H.1 Occupancy at equilibrium. The fraction of receptors occupied by a ligand at equilibrium depends on the concentration of the ligand compared to its Kd.
Assumptions of the law of mass action Although termed a “law,” the law of mass action is simply a model. It is based on these assumptions: 1. All receptors are equally accessible to ligands. 2. All receptors are either free or bound to ligand. The model ignores any states of partial binding. 3. Neither ligand nor receptor are altered by binding. 4. Binding is reversible. If these assumptions are not met, there are two choices. One choice is to develop a more complicated model, which is beyond the scope of this unit. The other choice is to analyze the data in the usual way, but interpret the result as an empirical description of the system without attributing rigorous meanings to the Kd and rate constants. Nonspecific Binding In addition to binding to the receptors of physiological interest, radioligands also bind to other (nonreceptor) sites. Binding to the receptor of interest is termed specific binding. Binding to other sites is called nonspecific binding. Because of this operational definition, nonspecific binding can represent several phenomena: 1. The bulk of nonspecific binding represents some sort of interaction of the ligand with membranes. The molecular details are unclear, but nonspecific binding depends on the charge and hydrophobicity of a ligand—but not its exact structure. 2. Nonspecific binding can also result from binding to receptor transporters, or to enzymes not of interest to the investigator (e.g., binding of epinephrine to serotonin receptors). 3. In addition, nonspecific binding can represent binding to the filters used to separate bound from free ligand. In many systems, nonspecific binding is linear with radioligand concentration. This means that it is possible to account for nonspecific binding mathematically, without ever measuring nonspecific binding directly. To do this, measure only total binding experimentally, and fit the data to models that include both specific and nonspecific components (see More Complicated Situations, below). Commonly Used Techniques
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Most investigators, however, prefer to measure nonspecific binding experimentally. To measure nonspecific binding, first block almost all specific binding sites with an unlabeled drug. Under these conditions, the radioligand only binds nonspecifically. This raises two questions: which unlabeled drug should be used and at what concentration? The most obvious choice of drug to use is the same compound as the radioligand, but unlabeled. In many cases, this is necessary as no other drug is known to bind to the receptors. Most investigators avoid using the same compound as the hot and cold ligand for routine work because both the labeled and unlabeled forms of the drug will bind to the same specific and nonspecific sites. This means that the unlabeled drug will reduce binding purely by isotopic dilution. When possible, it is better to define nonspecific binding with a drug chemically distinct from the radioligand. The concentration of unlabeled drug should be high enough to block virtually all the specific radioligand binding, but not so much that it will cause more general physical changes to the membrane that might alter specific binding. If studying a well-characterized receptor, a useful rule of thumb is to use the unlabeled compound at a concentration equal to 100 times its Kd for the receptors, or 100 times the highest concentration of radioligand, whichever is higher. The same results should be obtained from defining nonspecific binding with a range of concentrations of several drugs. Ideally, nonspecific binding is only 10% to 20% of the total radioligand binding. If the nonspecific binding makes up more than half of the total binding, it will be hard to get quality data. If the system exhibits a great deal of nonspecific binding, use a different kind of filter, wash with a larger volume of buffer or a different temperature buffer, or use a different radioligand. Ligand Depletion The equations that describe the law of mass action include the variable [ligand], which is the free concentration of ligand. All the analyses presented later in this unit assume that a very small fraction of the ligand binds to receptors (or to nonspecific sites), so that the free concentration of ligand is approximately equal to the concentration added. In some experimental situations, the receptors are present in high concentration and have a high affinity for the ligand. A large fraction of the radioligand binds to receptors (or nonspecific sites), depleting the amount of ligand remaining free in solution. The discrepancy is not the same in all tubes or at all times. Many investigators use this rule of thumb: if <10% of the ligand binds, don’t worry about ligand depletion. If possible, design the experimental protocol to avoid situations where >10% of the ligand binds. This can be done by using less tissue in the assays; however, this will also decrease the number of counts. An alternative is to increase the volume of the assay without changing the amount of tissue. In this case, more radioligand will be needed. If radioligand depletion cannot be avoided, the depletion must be accounted for in the analyses. There are several approaches. 1. Measure the free concentration of ligand in every tube.
Analyzing Radioligand Binding Data
2. Calculate the free concentration in each tube by subtracting the number of cpm (counts per minute) of total binding from the cpm of added ligand. This method works only for saturation binding experiments, and cannot be extended to analysis of competition or kinetic experiments. One problem with this approach is that experimental error in determining specific binding also affects the calculated value of free ligand concentration. When fitting curves, both x and y would include experimental
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error, and the errors will be related. This violates the assumptions of nonlinear regression. Using simulated data, Swillens (1995) has shown that this can be a substantial problem. Another problem is that the free concentration of radioligand will not be the same in tubes used for determining total and nonspecific binding. Therefore specific binding cannot be calculated as the difference between the total binding and nonspecific binding. 3. Fit total binding as a function of added ligand using an equation that accounts both for nonspecific binding and for ligand depletion (Swillens, 1995). By analyzing simulated experiments, more reliable results are obtained than those obtained from calculating free ligand by subtraction. SATURATION BINDING EXPERIMENTS Saturation binding experiments determine receptor number and affinity by determining specific binding at various concentrations of the radioligand. Because this kind of experiment can be graphed as a Scatchard plot (more accurately attributed to Rosenthal, 1967), they are sometimes called “Scatchard experiments.” The analyses depend on the assumption that the incubation has reached equilibrium. This can take anywhere from a few minutes to many hours, depending on the ligand, receptor, temperature, and other experimental conditions. Since lower concentrations of radioligand take longer to equilibrate, use a low concentration of radioligand (perhaps 10% to 20% of the estimated Kd) when measuring how long it takes the incubation to reach equilibrium. Experimenters typically use 6 to 12 concentrations of radioligand. Theory of Saturation Binding Nonspecific binding Analysis of saturation binding curves requires accounting for nonspecific binding. Although it is possible to account for nonspecific binding mathematically by analyzing total binding (see More Complicated Situations, below), most investigators assess nonspecific binding experimentally by measuring radioligand binding in the presence of a concentration of an unlabeled compound that binds to essentially all the receptors. Since all the receptors are occupied by the unlabeled drug, the radioligand only binds nonspecifically. Once the nonspecific binding has been determined, subtract it from total binding to calculate specific binding. There are two ways to do this. 1. Experimentally measure nonspecific binding at each concentration of radioligand. Calculate specific binding as total binding minus nonspecific binding at each concentration. 2. An alternative approach relies on the observation that nonspecific binding is generally proportional to the concentration of radioligand (within the concentration range used in the experiment). This means that a graph of nonspecific binding as a function of radioligand binding is generally linear, as shown in Figure A.3H.2. Once nonspecific binding is observed to be linear with radioligand concentration in the system, linear regression can be used to find the best-fit line through the nonspecific binding data. Specific binding is calculated by subtracting the nonspecific binding predicted by that line from the total binding measured at each concentration of radioligand. The major advantage of this approach is that measurements of nonspecific binding at each concentration of radioligand are not needed. For example, one can measure total binding at eight concentrations of radioligand, and nonspecific binding at only four concentrations.
Commonly Used Techniques
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[3H] Meproadifen bound (µM)
[3H]Mesulergine bound (fmol/mg protein)
6000 4000 2000 0 0
5 10 Free [Mesulergine] (nM) total
0.4 0.3 0.2 0.1 0 0
1
2
3
Free [Meproadifen] (µM)
nonspecific
Figure A.3H.2 Examples of nonspecific binding. (A) [3H]Mesulergine binding to serotonin receptors has low nonspecific binding (<25% of total binding at the highest concentrations). (B) [3H]Meproadifen binding to the ion channel of nicotinic receptors has high nonspecific binding (>50%).
Binding
total
specific nonspecific
[Radioligand]
Figure A.3H.3 Total binding, specific binding, and nonspecific binding for a saturation binding experiment.
Panel A of Figure A.3H.2 shows data from a nearly ideal system, where nonspecific binding is less than 25% of total binding. Panel B shows a less ideal system where nonspecific binding is over 50% of total binding at high ligand concentrations. If nonspecific binding were much higher than this, it would be very difficult to get reliable results. Equations used to calculate binding Specific binding at equilibrium equals fractional occupancy times the total receptor number (Bmax), and depends on the concentration of radioligand ([L]): specific binding = fractional occupancy × Bmax =
Analyzing Radioligand Binding Data
Bmax × [L ] Kd + [L ]
This equation describes a rectangular hyperbola or a binding isotherm. [L] is the concentration of free radioligand, the value plotted on the x axis (see Fig. A.3H.3). Bmax is the total number of receptors and is expressed in the same units as the y values (i.e., cpm, sites/cell, or fmol/mg protein). Kd is the equilibrium dissociation constant (expressed in the same units as [L], usually nM). Figure A.3H.3 shows the total binding, specific binding, and nonspecific binding for a hypothetical experiment.
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A
B
Bound/free
Bound
B max
50%
Kd
Free
slope = −
Bound
1 Kd
B max
Figure A.3H.4 Displaying results as a Scatchard plot. (A) Specific binding as a function of free radioligand. (B) Transformation of Scatchard data to a plot.
Analysis of Saturation Binding Curves Using nonlinear regression to determine Bmax and Kd Follow these steps to analyze the data with nonlinear regression: 1. Calculate specific binding at each concentration of ligand (or, in rare cases, decide to analyze only total binding; see More Complicated Situations, below). 2. Convert the specific binding data from counts per minute to more useful units such as fmol/mg protein or sites per cell. 3. Define x as the radioligand concentrations in nM or pM. Define y as the specific binding in fmol/mg or sites per cell. 4. Fit the data to this equation. y = Bmax × x ( K d + x )
5. If the curve fitting program does not provide initial values (sometimes called estimated values) automatically, estimate Bmax as the largest value of y and estimate Kd as 0.2 times the largest value of x. Are the results reasonable? Before accepting the results of the curve fit, ask the questions listed in Table A.3H.1 to determine whether the results are reasonable. If the results are not reasonable, the experimental protocol may need revision. Also check that the data are being analyzed correctly. In addition, it’s possible that the system is more complex than the simple one-site binding model. To determine whether the system follows the assumptions of the simple model, consider the points in Table A.3H.2. Displaying results as a Scatchard plot Before nonlinear regression programs were widely available, scientists transformed data to make a linear graph and then analyzed the transformed data with linear regression. There are several ways to linearize binding data, but Scatchard plots (more accurately attributed to Rosenthal, 1967) are used most often. As shown in Figure A.3H.4, the x axis of the Scatchard plot represents specific binding (usually labeled “bound”) and the y axis is the ratio of specific binding to concentration of free radioligand (usually labeled “bound/free”). Bmax is the x intercept; Kd is the negative reciprocal of the slope.
Commonly Used Techniques
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Table A.3H.1
Evaluating the Results of Saturation Binding Curve Analysis
Question
Comment
Does the calculated curve go near the data points?
If the curve doesn’t go near the data, then something went wrong with the curve fit, and the “best-fit” values of Bmax and Kd should be ignored.
Were sufficient concentrations of radioligand used?
Ideally, the highest concentration should be at least 10 times the Kd. Calculate the ratio of the highest radioligand concentration used divided by the Kd reported by the program (both in nM or pM). The ratio should be greater than 10.
Is the Bmax reasonable?
Typical values for Bmax are 10 to 1000 fmol binding sites per milligram of membrane protein, 1000 sites per cell, or 1 receptor per square micron of membrane. If using cells transfected with receptor genes, then the Bmax may be 10 to 100 times larger than these values. Typical values for Kd of useful radioligands range between 10 pM and 100 nM. If the Kd is much lower than 10 pM, the dissociation rate is probably very slow and it will be difficult to achieve equilibrium. If the Kd is much higher than 100 nM, the dissociation rate will probably be fast, and may result in the loss of binding sites during separation of bound from free radioligand. Nonlinear regression programs report the uncertainty of the best-fit values for Bmax and Kd as standard errors and 95% confidence intervals. Divide the SE of the Bmax by the Bmax, and divide the SE of the Kd by the Kd. If either ratio is much larger than ∼20%, look further to determine why. Divide the nonspecific binding at the highest concentration of radioligand by the total binding at that concentration. Nonspecific binding should usually be less than 50% of the total binding.
Is the Kd reasonable?
Are the standard errors too large? Are the confidence intervals too wide?
Is the nonspecific binding too high?
Table A.3H.2
Evaluating the Assumptions of Saturation Binding Analysis
Assumption
Comment
Binding has reached equilibrium.
It takes longest for the lower concentrations to equilibrate, so test equilibration time with the lowest concentration of radioligand. See Theory: Comparing One- and Two-Site Models, below.
There is only one population of receptors.
Analyzing Radioligand Binding Data
Only a small fraction of the radioligand binds, therefore the free concentration is essentially identical to the concentration added.
Compare the cpm obtained for total binding to the amount of ligand. If the ratio is greater than 10% at any concentration, this assumption has been violated. Increase the volume of the reaction but use the same amount of tissue.
There is no cooperativity. Binding of a ligand to one binding site does not alter the affinity of another binding site.
See Cooperativity, below.
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B 40
2 Bound/free
Specific binding
A
20
0
1
0 0
50
100 [Ligand]
150
0
20
40
60
Bound
Figure A.3H.5 Why Scatchard plots (though useful for displaying data) should not be used for analyzing data. (A) Experimental data with best-fit curve determined by nonlinear regression. (B) Scatchard plot of the data. The solid line corresponds to the Bmax and Kd determined by nonlinear regression in panel A. The dashed line was determined by linear regression of transformed data in panel B. The results of linear regression of the Scatchard plot are not the most accurate values for Bmax (x intercept) or Kd (negative reciprocal of the slope).
When making a Scatchard plot, there are two ways to express the y axis. 1. One choice is to express both free ligand and specific binding in cpm so the ratio bound/free is a unitless fraction. The advantage of this choice is that you can interpret y values as the fraction of radioligand bound to receptors. If the highest y value is large (>0.10), then the free concentration of radioligand will be substantially less than the added concentration, and the standard analyses will yield inaccurate values for Bmax and Kd. In this situation, either revise the experimental protocol or use special analysis methods that deal with ligand, as discussed previously. The disadvantage of this choice of units is that the slope of the line cannot be interpreted without performing unit conversions. 2. An alternative is to express the y axis as the ratio of units used to display bound and free on the saturation binding graph (i.e., sites/cell/nM or fmol/mg/nM). While these values are hard to interpret, they simplify calculation of the Kd, which equals the negative reciprocal of the slope. The specific binding units cancel when calculating the slope. The negative reciprocal of the slope is expressed in units of concentration (nM) which equals the Kd. The problem with using Scatchard plots to analyze saturation binding experiments While Scatchard plots are very useful for visualizing data, they are not the most accurate way to analyze data. The problem is that the linear transformation distorts the experimental error. Linear regression assumes that the scatter of points around the line follows a Gaussian distribution and that the standard deviation is the same at every value of x. These assumptions are not true with the transformed data. A second problem is that the Scatchard transformation alters the relationship between x and y. The value of x (bound) is used to calculate y (bound/free), and this violates the assumptions of linear regression. Since these assumptions are violated, the Bmax and Kd values determined by linear regression of Scatchard-transformed data are likely to be further from the actual values than the Bmax and Kd determined by nonlinear regression. Nonlinear regression produces the most accurate results, whereas a Scatchard plot produces only approximate results. Figure A.3H.5 illustrates the problem of transforming data. The left panel shows data that follows a rectangular hyperbola (binding isotherm). The solid curve was determined by
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Total binding (cpm)
0.125 0.25 0.5 1.0 2.0 4.0
818 826 1856 1727 3452 3349 6681 6055 10077 9333 13715 13277
15,000
Nonspecific binding (cpm)
94
88
354
375
1573
1525
total Bound (cpm)
[Radioligand] (nM)
10,000
5,000 nonspecific 0 0
1
2
3
125 I-[Sar 1 ,IIe8 ]-ang
4 II (nM)
Figure A.3H.6 Sample saturation binding experiment. The ligand binding to angiotensin receptors in a membrane preparation was measured. Total and nonspecific binding are shown.
nonlinear regression. The right panel is a Scatchard plot of the same data. The solid line shows how that same curve would look after a Scatchard transformation. The dotted line shows the linear regression fit of the transformed data. The transformation amplified and distorted the scatter, and thus the linear regression fit does not yield the most accurate values for Bmax and Kd. In this example, the Bmax determined by the Scatchard plot is ∼25% too large and the Kd determined by the Scatchard plot is too high. The errors could just as easily have gone in the other direction. The experiment in Figure A.3H.5 was designed to determine the Bmax with little concern for the value of Kd. Therefore, it was appropriate to obtain only a few data points at the beginning of the curve and many in the plateau region. Note however how the Scatchard transformation gives undo weight to the data point collected at the lowest concentration of radioligand (the lower left point in panel A, the upper left point in panel B). This point dominates the linear regression calculations on the Scatchard graph. It has “pulled” the regression line to become shallower, resulting in an overestimate of the Bmax. Again, although it is inappropriate to analyze data by performing linear regression on a Scatchard plot, it is often helpful to display data as a Scatchard plot. Many people find it easier to visually interpret Scatchard plots than binding curves, especially when comparing results from different experimental treatments. Example of a Saturation Binding Experiment Raw data Figure A.3H.6 shows duplicate values for total binding of six concentrations of a radioligand to angiotensin receptors on membranes of cells transfected with an angiotensin gene (R. Neubig, unpub. observ.). The figure also shows nonspecific binding (assessed with 10 µM unlabeled angiotensin II) at three concentrations of radioligand.
Analyzing Radioligand Binding Data
Calculating specific binding Since nonspecific binding was only determined at three concentrations of radioligand, the standard method of subtracting each nonspecific value from the corresponding total value cannot be used. Instead, the fact (confirmed in other experiments) that nonspecific binding is proportional to radioligand concentration is relied upon, and the best-fit value of nonspecific binding is subtracted from each total binding value. This can be done in one step by choosing “remove baseline analysis” in GraphPad Prism software. Alternatively:
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Table A.3H.3
[Radioligand] (nM)
Calculating Specific Binding
Total binding (cpm) Duplicate 1 Duplicate 2
Computed nonspecific binding (cpm)
Calculated specific binding (cpm)
Specific binding (fmol/mg)
Duplicate 1 Duplicate 2
Duplicate 1 Duplicate 2
0.125
818
826
34
784
792
17.9
18.1
0.25 0.5
1856 3452
1727 3349
82 180
1774 3272
1645 3169
40.5 74.8
37.6 72.4
1.0 2.0
6681 10077
6055 9333
375 766
6306 9311
5680 8567
144.1 212.8
129.8 195.8
4.0
13715
13277
1547
12168
11730
278.1
268.1
1. Use linear regression. The best fit line through the nonspecific binding data is: nonspecific binding in cpm = −15.25 + 390.5 ([radioligand ] in nM ) 2. Use this equation to calculate nonspecific binding at each of the six radioligand concentrations. 3. Subtract that calculated value from the observed total binding to compute specific binding (Table A.3H.3). Converting units Convert from cpm to fmol/mg using the amount of protein in each tube (0.01 mg), the efficiency of the counting (90%), and the specific radioactivity of the ligand (2190 Ci/mmole). fmol mg =
cpm 2.22 × 10 dpm Ci × 0.90 cpm dpm × 2190 Ci mmol × 10 −12 mmol fmol × 0.01 mg 12
NOTE: 1. Receptors in membrane preparations are often expressed as fmol of receptor per milligram of membrane protein. One fmol is 10−15 moles. 2. Counting efficiency is the fraction of the radioactive disintegrations that are detected by the counter. This example uses a radioligand labeled with 125I, so the efficiency (90%) is very high. 3. The Curie (C:) is a unit of radioactivity and equals 2.22 × 1012 radioactive disintegrations per minute. 4. The value 2190 Ci/mmole is worth remembering. It is the specific activity of ligands iodinated with 125I, when every molecule is labeled with one iodine. 5. Simplifying the equation, simply divide the cpm by 43.756 (see Table A.3H.3). Fitting a curve to determine Bmax and Kd When fitting the example data to a curve, one must decide whether to enter the data as six points or twelve. Entering each replicate individually is better, as it provides more data to the curve fitting procedure. This should be avoided only when the replicates are not independent (i.e., when experimental error in one value is likely to affect the other value as well). In this case each replicate was determined in a separate tube poured over a separate filter, and all the data were obtained from one membrane preparation. Except
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Specific binding (fmol/mg)
300
200
100
0 0
1
2
3
4
5
[125I-[Sar1 , IIe8]-ang II](nM)
Figure A.3H.7 Specific binding with the best-fit curve determined by nonlinear regression. These data are the same as those shown in Figure A.3H.6 and Table A.3H.3.
for errors in preparing the radioligand dilutions, experimental errors will affect each value independently. Follow these steps to fit the data. 1. If the data-fitting program understands the concept of duplicates, then enter the data with radioligand concentration as six x values and the duplicate values of specific binding at each concentration. If the program does not understand how to deal with duplicates, enter each concentration value twice in the x column, to fill twelve rows. Enter the specific binding data as a column of twelve y values. 2. Choose nonlinear regression and choose or enter the one-site binding equation. Expressed in the format of most curve fitting systems, it is: y = ( Bmax × x ) ( K d + x )
3. If the chosen nonlinear regression program does not provide initial values automatically, estimate values for Bmax and Kd. For Bmax, enter a value a bit higher than the highest value in the data, perhaps 300 for this example. For Kd, estimate the concentration of radioligand that binds to half the sites, perhaps 0.5 nM. These estimated values do not have to be very accurate. 4. Start the curve fit, and note the results. The best-fit value of Bmax is 429. It is expressed in the same units as the y values entered (fmol/mg). The best-fit value of Kd is 2.27. It is expressed in the same units as the x values entered (nM). 5. Graph the specific binding with the best-fit curve as shown in Figure A.3H.7. Creating a Scatchard plot A Scatchard plot is a graph of specific binding vs. the ratio of specific binding to free radioligand. For specific binding, the two replicates are averaged (individual replicates could have been shown). For the example in Figure A.3H.8, bound/free is expressed as fmol/mg divided by nM. Analyzing Radioligand Binding Data
Figure A.3H.8 shows the Scatchard transformation of the specific binding data. Since it is not appropriate to determine the Kd and Bmax from linear regression of a Scatchard plot, derive the solid line on the graph from the best-fit values using nonlinear regression:
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18 39 73 136 203 272
143.5 155.5 146.5 136.3 101.7 68.0
200 Bound/free (fmol/mg/nM)
Specific binding Bound/free (fmol/mg protein) fmol/mg/nM
150 100 50 0 0
200
400
600
Bound (fmol/mg)
Figure A.3H.8 Scatchard transformation of the data from Figure A.3H.7. The solid line was created (as explained in the text) from the best-fit values of Bmax and Kd determined from nonlinear regression. This is the correct line to show on a Scatchard plot. The dashed line was determined by linear regression of the Scatchard transformed data. It is shown here for comparison only; it is not informative or helpful.
1. The x intercept of the Scatchard plot is Bmax, which equals 429 by nonlinear regression, so one end of the line is at x = 429, y = 0. 2. The slope of the line is the negative reciprocal of the Kd. Since the Kd is 2.27 nM, the slope must be −1/2.27, which equals −0.4405 nM−1. 3. The y intercept divided by the x intercept equals the negative slope. We know the slope and the x intercept, so can derive the y intercept. It equals −slope × x intercept = 0.4405 × 429 = 188.1. 4. Draw the line from x = 0, y = 188.1 to x = 429, y = 0, as in Figure A.3H.8. Figure A.3H.8 also shows the dotted line derived by linear regression of the Scatchard transformed data. This is shown only to emphasize the difference between it and the best-fit line derived from nonlinear regression. The linear regression line should not be used for data analysis and does not aid data presentation. Critiquing the experiment This example is not an ideal experiment. Consider these points: The highest concentration of radioligand used (4 nM) is not even twice the Kd (2.27 nM). Ideally the highest concentration of radioligand should be ten times the Kd. In addition, the specific binding of the first few points lies below the best fit curve. There are many possible explanations for this, including chance, but it may be because the system is not at equilibrium. The lowest concentrations take the longest to equilibrate, so it is possible that the first few concentrations had not equilibrated, resulting in an underestimate of specific binding at equilibrium. More Complicated Situations Measuring total binding only Rather than measure both total and nonspecific binding at each concentration of ligand, measure only total binding. Then fit the data to the equation below, which defines total binding as a function of ligand concentration. Total binding is the sum of specific binding and nonspecific binding. Fit the data to this equation to find best-fit values of Bmax, Kd,
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and NS (the slope of the graph of nonspecific binding as a function of radioligand concentration): total binding = specific binding + nonspecific binding =
Bmax × [L ] K d + [ L]
+ NS × [L ]
This approach assumes that nonspecific binding is proportional to [ligand]. This assumption is reasonable if the nonspecific binding is due to general binding to membranes, but may not be reasonable if some of the nonspecific binding represents binding to receptors or transporters other than the one being studied. To get useful results with this approach requires high-quality data and at least ten data points, including some well above the ligand Kd. Two classes of binding sites If the radioligand binds to two classes of binding sites, the specific binding data can be fit to this equation: specific binding =
Bmax1 × [L ] Kd1 + [L ]
+
Bmax2 × [L] Kd2 + [L]
This equation assumes that the radioligand binds to two independent noninteracting binding sites, and that the binding to each site follows the law of mass action. A comparison of the one-site and two-site fits will be addressed later in this unit (see Theory: Comparing One- and Two-Site Models, below). Meaningful results will be obtained from a two-site fit only if you have ten or more data points spaced over a wide range of radioligand concentrations. Binding should be measured at radioligand concentrations below the high-affinity Kd and above the low-affinity Kd. Homologous competitive binding curves Some investigators determine the Kd and Bmax of a ligand by holding the concentration of the radioligand constant and competing with various concentrations of the unlabeled ligand. This approach will be discussed below. COMPETITIVE BINDING EXPERIMENTS Theory of Competitive Binding Using competitive binding curves Competitive binding experiments measure the binding of a single concentration of labeled ligand in the presence of various concentrations (often twelve to sixteen) of unlabeled ligand. Competitive binding experiments are used to: 1. Validate an assay. Perform competitive binding experiments with a series of drugs whose potencies at the receptor of interest are known from functional experiments. Demonstrating that these drugs bind with the expected potencies, or at least the expected order of potency, helps prove that the radioligand has identified the correct receptor. This kind of experiment is crucial, because there is usually no point studying a binding site unless it has physiological significance.
Analyzing Radioligand Binding Data
2. Determine whether a drug binds to the receptor. Thousands of compounds can be screened to find drugs that bind to the receptor. This can be faster and easier than other screening methods.
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3. Investigate the interaction of low affinity drugs with receptors. Binding assays are only useful when the radioligand has a high affinity (Kd <100 nM or so). A radioligand with low affinity generally has a fast dissociation rate constant, and so won’t stay bound to the receptor while washing the filters. To study the binding of a low affinity drug, use it as an unlabeled competitor. 4. Determine receptor number and affinity by using the same compound as the labeled and unlabeled ligand (see Homologous Competitive Binding Curves, below). Performing the experiment Competitive binding experiments use a single concentration of radioligand and require incubation until equilibrium is reached. That raises two questions: how much radioligand should be used, and how long does it take to equilibrate? There is no clear answer to the first question. Higher concentrations of radioligand result in higher counts and thus lower counting error, but these experiments are more expensive and have higher nonspecific binding. Lower concentrations save money and reduce nonspecific binding but result in fewer counts from specific binding and thus more counting error. Many investigators choose a concentration approximately equal to the Kd of the radioligand for binding to the receptor, but this is not universal. In general, you should aim for a minimum of 1000 cpm from specific binding in the absence of competitor. Many investigators’ first thoughts are that binding will reach equilibration in the time it takes the radioligand to reach equilibrium in the absence of competitor. It turns out that this may not be long enough. Incubations should last four to five times the half-life for receptor dissociation as determined in a dissociation experiment. Equations for competitive binding Competitive binding curves are described by this equation: total radioligand binding = NS +
( total − NS)
log D − log( IC50 ) 1+10 [ ]
The x axis of Figure A.3H.9 shows varying concentrations of unlabeled drug on a log scale. The y axis can be expressed as cpm or converted to more useful units like fmol bound per milligram protein or number of binding sites per cell. Some investigators like to normalize the data from 100% (no competitor) to 0% (nonspecific binding at maximal concentrations of competitor). The top of the curve shows a plateau at the amount of radioligand bound in the absence of the competing unlabeled drug. This equals the parameter total in the equation. The bottom of the curve is a plateau equal to nonspecific binding; this is nonspecific (NS) in the equation. These values are expressed in the units of the y axis. The difference between the top and bottom plateaus is the specific binding. Note that this not the same as Bmax. When using a low concentration of radioligand (to save money and avoid nonspecific binding), only a fraction of receptors will be bound (even in the absence of competitor), so specific binding will be much lower than the Bmax. The concentration of unlabeled drug that results in radioligand binding halfway between the upper and lower plateaus is called the IC50 (inhibitory concentration 50%), also called the EC50 (effective concentration 50%). The IC50 is the concentration of unlabeled drug that blocks half the specific binding, and it is determined by three factors: 1. The Ki of the receptor for the competing drug. This is what is to be determined. It is the equilibrium dissociation constant for binding of the unlabeled drug—the concen-
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Total radioligand binding
Total
NS
Log(IC50) Log[unlabeled drug]
Figure A.3H.9 Schematic of a competitive binding experiment.
tration of the unlabeled drug that will bind to half the binding sites at equilibrium in the absence of radioligand or other competitors. The Ki is proportional to the IC50. If the Ki is low (i.e., the affinity is high), the IC50 will also be low. 2. The concentration of the radioligand. If a higher concentration of radioligand is used, it will take a larger concentration of unlabeled drug to compete for the binding. Therefore, increasing the concentration of radioligand will increase the IC50 without changing the Ki. 3. The affinity of the radioligand for the receptor (Kd). It takes more unlabeled drug to compete for a tightly bound radioligand (small Kd) than for a loosely bound radioligand (high Kd). Using a radioligand with a smaller Kd (higher affinity) will increase the IC50. Calculate the Ki from the IC50, using the equation of Cheng and Prusoff (1973). Ki =
IC50 [radioligand] 1+ Kd
Remember that Ki is a property of the receptor and unlabeled drug, while IC50 is a property of the experiment. By changing experimental conditions (changing the radioligand used or changing its concentration), the IC50 will change without affecting the Ki. This equation is based on the following assumptions: 1. Only a small fraction of either the labeled or unlabeled ligand has bound. This means that the free concentration is virtually the same as the added concentration. 2. The receptors are homogeneous and all have the same affinity for the ligands. 3. There is no cooperativity—binding to one binding site does not alter affinity at another site. 4. The experiment has reached equilibrium. 5. Binding is reversible and follows the law of mass action. Analyzing Radioligand Binding Data
6. The Kd of the radioligand is known from an experiment performed under similar conditions.
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Percent specific binding
100
90%
50 10%
81-fold
0 10 −9
10 −8
10 −7
10 −6
10 −5
10 −4 10 −3
[Unlabeled drug] (M)
Figure A.3H.10 Steepness of a competitive binding curve. This graph shows the results at equilibrium when radioligand and competitor bind to the same binding site. The curve will descend from 90% binding to 10% binding over an 81-fold increase in competitor concentration.
If the labeled and unlabeled ligand compete for a single binding site, the steepness of the competitive binding curve is determined by the law of mass action (see Fig. A.3H.10). The curve descends from 90% specific binding to 10% specific binding with an 81-fold increase in the concentration of the unlabeled drug. More simply, nearly the entire curve will cover two log units (100-fold change in concentration). Analyzing Competitive Binding Data Using nonlinear regression to determine the Ki Follow these steps to determine the Ki with nonlinear regression. 1. Enter the x values as the logarithm of the concentration of unlabeled compound, or enter the concentrations, and use the program to convert to logarithms. Since log(0) is undefined, the log scale cannot accommodate a concentration of zero. Instead enter a very low concentration. For example, if the lowest concentration of unlabeled compound is 10−10 M, then enter −12 for the zero concentration. 2. Enter the y values as cpm total binding. There is little advantage to converting to units such as fmol/mg or sites/cell. There is also little advantage to converting to percent specific binding. 3. Select the competitive binding equation (TOP is binding in the absence of competitor, BOTTOM is binding at maximal concentrations of competitor, logIC50 is the logarithm base 10 of the IC50): y = NS +
TOTAL − NS 1 + 10 x − log IC50
4. If the chosen nonlinear regression program doesn’t provide initial estimates automatically, enter these values. For NS, enter the smallest y value. For TOTAL, enter the largest y value. For log(IC50), enter the average of the smallest and largest x values. 5. If the data do not form clear plateaus at the top and bottom of the curve, consider fixing top or bottom to constant values. TOTAL can be fixed to the binding measured in the absence of competitor and NS to binding measured in the presence of a large concentration of a standard drug known to block radioligand binding to essentially all receptors.
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6. Start the curve fitting to determine TOTAL, NS, and log(IC50). 7. Calculate the IC50 as the antilog of log(IC50). 8. Calculate the Ki using this equation: Ki =
IC50 [radioligand] 1+ Kd
When to set total and NS constant In order to determine the best-fit value of IC50, the nonlinear regression program must be able to determine the 100% (total) and 0% (nonspecific) plateaus. If there is data over a wide range of concentrations of unlabeled drug, the curve will have clearly defined bottom and top plateaus and the program should have no trouble fitting all three values (both plateaus and the IC50). With some experiments, the competition data may not define a clear bottom plateau. If data are fit in the usual way, the program might stop with an error message, or it might find a nonsense value for the nonspecific plateau (it might even be negative). If the bottom plateau is incorrect, the IC50 will also be incorrect. To solve this problem, define the nonspecific binding from other data. All drugs that bind to the same receptor should compete for all specific radioligand binding and reach the same bottom plateau value. When running the curve fitting program, set the bottom plateau of the curve (NS) to a constant equal to binding in the presence of a standard drug known to block all specific binding. Similarly, if the curve doesn’t have a clear top plateau, set the total binding to be a constant equal to binding in the absence of any competitor. Interpreting the Results of Competitive Binding Curves Are the results reasonable? Table A.3H.4 presents some questions to consider when determining if the results are reasonable and logical. Do the data follow the assumptions of the analysis? Table A.3H.5 lists the assumptions.
Table A.3H.4
Analyzing Radioligand Binding Data
Evaluating the Results of Competitive Binding Curve Analyses
Question
Comment
Is the log(IC50) reasonable?
The IC50 should be near the middle of the curve, with at least several concentrations of unlabeled competitor on either side of it.
Are the standard errors too large? Are the confidence intervals too wide?
The SE of the log(IC50) should be <0.5 log unit (ideally much less).
Are the values of TOTAL and NS reasonable?
TOTAL should be near the binding observed in the absence of competitor. NS should be near the binding observed in the presence of a maximal concentration of competitor. If the best-fit value of NS is negative, consider fixing it to a constant value equal to nonspecific binding.
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Table A.3H.5
Evaluating the Assumptions of Competitive Binding Analyses
Assumption
Comment
Binding has reached equilibrium.
Competitive binding incubations take longer to incubate than saturation binding incubations. Incubate for 4 to 5 times the half life for radioligand dissociation.
There is only one population of receptors Only a small fraction of the radioligand binds, therefore the free concentration is essentially identical to the concentration added. There is no cooperativity. Binding of a ligand to one binding site does not alter the affinity of another binding site.
See Theory: Comparing One- and Two-Site Models. Compare the total binding in the absence of competitor in cpm, to the amount of ligand added in cpm. If the ratio is >10% at any concentration, then you’ve violated this assumption. See Cooperativity.
Why determine log(IC50) rather than IC50? The equation for a competitive binding curve (see Theory of Competitive Binding, Equations for competitive binding, above) looks a bit strange since it combines logarithms and antilogarithms (10 to the power). A bit of algebra simplifies it: y = nonspecific +
( total − nonspecific ) [drug] 1+ IC50
Fitting data to this equation results in the same best-fit curve and the same IC50. However, the confidence interval for the IC50 will be different. Which confidence interval is correct? With nonlinear regression, the standard errors of the variables are only approximately correct. Since the confidence intervals are calculated from the standard errors, they too are only approximately correct. The problem is that the real confidence interval may not be symmetrical around the best-fit value. It may extend further in one direction than the other. However, nonlinear regression programs always calculate symmetrical confidence intervals (unless you use advanced techniques). Therefore, when writing the equation for nonlinear regression, choose variables so the uncertainty is as symmetrical as possible. Because data are collected at concentrations of unlabeled drug equally spaced on a log axis, the uncertainty is symmetrical when the equation is written in terms of the log(IC50), but is not symmetrical when written in terms of IC50. Confidence intervals are more accurate when the equation is written in terms of the log(IC50). Figure A.3H.11 (R. Neubig, unpub. observ.) shows competition of unlabeled yohimbine for labeled UK14341 (an α2 adrenergic agonist). 1. Enter the data into a nonlinear regression program. Enter the logarithm of concentration of the unlabeled ligand in the x column, and the triplicate values of total binding in the x columns. If the selected program does not allow entry of triplicate values, enter each log of concentration three times. 2. Fit the data to a one-site competitive binding curve. If necessary, enter it in this format: NS y = NS + Total − ( x − log IC50 ) 1 + 10
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Binding (cpm) in triplicate
−12.0 −10.0 −9.5 −9.0 −8.5 −8.0 −7.5 −7.0 −6.5 −6.0 −5.5 −5.0
454 4380 4554 9 4803 4213 4604 4353 4278 4508 4192 4156 3972 4053 4420 4191 3453 3018 3024 2587 2946 2367 1295 1405 1402 888 886 880 603 591 612 555 580 559 555 521 545
[3H] UK 14304 bound (cpm)
Log [competitor](M)
5000 4000 3000 2000 1000 0 0
–10 –9 –8 –7 –6 –5 Log[yohimbine]
Figure A.3H.11 Example of a competitive binding experiment. Yohimbine competes for radioligand binding to α2 receptors on membranes.
3. If the nonlinear regression program does not provide initial values automatically, estimate the values of the variables. TOTAL is the top plateau of the curve, so estimate its value from the highest data values, perhaps 4500. NS is the bottom plateau, so estimate its value from the lowest data values, perhaps 500. Log(IC50) is the x value in the middle of the curve. From looking at the data, estimate its value as −7. None of these estimates has to be very accurate, and the nonlinear regression will probably work fine even if the estimates are fairly different than the values listed here. 4. Note the best-fit results: NS = 530.3, TOTAL = 4418, and log(IC50) = −A.3H32. 5. Convert the log(IC50) to the IC50 by taking the antilog. IC50 = 29.4 nM. 6. Convert the IC50 to Ki. To do this, the concentration of radioligand used (2.0 nM) and its Kd for the receptors (0.88 nM, determined in a separate saturation binding experiment not shown here) must be known. Ki =
IC50 29.4 nM = = 8.98 nM 2.0 nM radioligand ] [ 1+ 1+ 0.88 nM Kd
Homologous Competitive Binding Curves A competitive binding experiment is termed homologous when the same compound is used as the hot and cold ligand. The term heterologous is used when the hot and cold ligands differ. Homologous competitive binding experiments can be used to determine the affinity of a ligand for the receptor and the receptor number. In other words, the experiment has the same goals as a saturation binding curve. Because homologous competitive binding experiments use a single concentration of radioligand (which can be low), they consume less radioligand and thus are more practical when radioligands are expensive or difficult to synthesize. Analyzing Radioligand Binding Data
To analyze a homologous competitive binding curve, the following assumptions must be made:
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1. The receptor has identical affinity for the labeled and unlabeled ligand. If you choose an iodinated radioligand, you should also use an iodinated unlabeled compound (using nonradioactive iodine), because iodination often changes the binding properties of ligands. 2. There is no cooperativity. 3. There is no ligand depletion. The methods in this section assume that only a small fraction of ligand binds. In other words, the method assumes that free concentration equals the concentration added. 4. There is only one class of binding sites. It is difficult to detect a second class of binding sites unless the number of lower affinity sites vastly exceeds the number of higher affinity receptors (because the single low concentration of radioligand used in the experiment will bind to only a small fraction of low affinity receptors). Analyze a homologous competitive binding curve using the same equation used for a one-site heterologous competitive binding to determine the top and bottom plateaus and the IC50. The Cheng and Prusoff equation (see Theory of Competitive Binding, above) can be used to calculate the Ki from the IC50. In the case of a homologous competitive binding experiment, assume that the hot and cold ligand have identical affinities so that Kd and Ki are the same. An algebraic rearrangement yields: K d = K i = IC50 − [radioligand ]
Set the concentration of radioligand in the experimental design, and determine the IC50 from nonlinear regression. The difference between the two is the Kd of the ligand (assuming hot and cold ligands bind the same). The difference between the top and bottom plateaus of the curve represents the specific binding of radioligand at the concentration used. Depending on how much radioligand is used, this value may be close to the Bmax or far from it. To determine the Bmax, divide the specific binding by the fractional occupancy, calculated from the Kd and the concentration of radioligand. Bmax =
TOP − BOTTOM TOP − BOTTOM = fractional occupancy [radioligand ]
( Kd + [radioligand])
Example of homologous competitive binding Figure A.3H.12 shows data from a binding experiment using [3H]yohimbine to quantify α2 adrenergic receptors to compete with unlabeled yohimbine. Since there is no reason to think that the tritium label will alter yohimbine’s affinity for the receptor, the method from the previous section can be used to quantify receptor number and affinity. 1. Enter the data into a nonlinear regression program. Enter the logarithm of concentration as x and cpm bound as y. The first point represents a control with no yohimbine. Since the log of zero is undefined, this cannot be shown on a log scale. Instead enter this value as −12 (the exact value is a bit arbitrary). 2. Fit the data using nonlinear regression to a sigmoidal equation; or, for manual entry, enter it like this: y = BOTTOM +
( TOP − BOTTOM ) 1 + 10(
x − logIC50 )
Commonly Used Techniques
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[3H]yohimbine bound (cpm)
12,500
5 nM yohimbine
10,000
K d = 38 nM B max = 95, 200 cpm
7,500 5,000 2,500 0 −inf
−9
−8
−7
−6
−5
Log[yohimbine]
Figure A.3H.12 Example of homologous competitive binding experiment. The hot and cold ligands are identical.
3. Note the best-fit results: BOTTOM = 350.6; TOP= 11,510; log(IC50) = −7.370; and IC50 = 42.6 nM. 4. Compute the Ki as the difference between the IC50 and the amount of radioligand added to all tubes. Ki = 42.6 nM − 5.0 nM = 37.6 nM. 5. Calculate the fraction of the receptors occupied by the 5 nM [3H]yohimbine used in the experiment. Fractional occupancy = [ligand]/([ligand] + Kd) = 5/(5.0 + 37.6)= 11.73%. 6. Calculate the specific binding of 5 nM radioligand as the difference between the top and bottom plateaus. Specific binding = 11,510 − 350 = 11,160 cpm. 7. Divide the specific binding by the fractional occupancy to determine the total number of binding sites, Bmax. It equals 11,160/0.1173 = 95,140 cpm. 8. Finally convert to more useful units. In this example there were 6 × 104 cells per well, the specific activity of the [3H]yohimbine was 78 Ci/mmole, and the scintillation counting efficiency was 33%. Calculate receptors/cell using the equation: fmol mg =
95,140 cpm × 6.02 × 1020 receptors mmol
2.22 × 1012 dpm Ci × 0.33cpm dpm × 78 Ci mmol × 60, 000 cells = 1.67 × 10 7 receptors cell
The Slope Factor or Hill Slope Many competitive binding curves are shallower than predicted by the law of mass action for binding to a single site. The steepness of a binding curve can be quantified with a slope factor, often called a Hill slope. A one-site competitive binding curve that follows the law of mass action has a slope of −1.0. If the curve is more shallow, the slope factor will be a negative fraction (i.e., −0.85 or −0.60; see Fig. A.3H.13). The slope factor is negative because the curve goes downhill. To quantify the steepness of a competitive binding curve (or a dose-response curve), fit the data to this equation: Analyzing Radioligand Binding Data
total radioligand binding = NS +
( Total − NS)
1 + 10
(logIC50 −log[D]) × slope factor
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Percent specific binding
100
50
slope = −1.00 slope = −0.75 slope = −0.50
0 −9
−8
−7
−6
−5
−4
−3
Log[unlabeled competitor]
Figure A.3H.13 Examples of slope factors. The slope factor quantifies the steepness of the curve, and is determined by nonlinear regression of competitive binding data. It is not the same as the slope of the curves at the midpoints.
The slope factor is a number that describes the steepness of the curve. In most situations, there is no way to interpret the value in terms of chemistry or biology. If the slope factor differs significantly from −1.0, then the binding does not follow the law of mass action with a single site. Explanations for shallow binding curves include: 1. Heterogeneous receptors. The receptors do not all bind the unlabeled drug with the same affinity. This can be due to the presence of different receptor subtypes, or due to heterogeneity in receptor coupling to other molecules such as G proteins. In Fig. A.3H.12, the slope factor equals −0.78. 2. Negative cooperativity. Binding sites are clustered (perhaps several binding sites per molecule) and binding of the unlabeled ligand to one site causes the remaining site(s) to bind the unlabeled ligand with lower affinity. 3. Curve fitting problems. If the top and bottom plateaus are not correct, then the slope factor is not meaningful. Don’t try to interpret the slope factor unless the curve has clear top and bottom plateaus. KINETIC BINDING EXPERIMENTS Theory of Binding, Kinetic Dissociation experiments A dissociation binding experiment measures the “off rate” of radioligand dissociating from the receptor. Perform dissociation experiments to fully characterize the interaction of ligand and receptor and confirm that the law of mass action applies. They may also be used to help design the experimental protocol. If the dissociation is fast, filter and wash the samples quickly so that negligible dissociation occurs. It may also require lowering the temperature of the buffer used to wash the filters, or switching to a centrifugation or dialysis assay. If the dissociation is slow, then the samples can be filtered at a more leisurely pace, because the dissociation will be negligible during the wash. Commonly Used Techniques
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To perform a dissociation experiment, first allow ligand and receptor to bind, perhaps to equilibrium. At that point, block further binding of radioligand to receptor using one of these methods: 1. If the tissue is attached to a surface, remove the buffer containing radioligand and replace with fresh buffer without radioligand. 2. Spin the suspension and resuspend in fresh buffer. 3. Add a very high concentration of an unlabeled ligand (perhaps 100 times its Ki for that receptor). It will instantly bind to nearly all the unoccupied receptors and block binding of the radioligand. 4. Dilute the incubation by a large factor, perhaps a 20- to 100-fold dilution. This will reduce the concentration of radioligand by that factor. At such a low concentration, new binding of radioligand will be negligible. This method is only practical when using a fairly low radioligand concentration so its concentration after dilution is far below its Kd for binding. 5. After initiating dissociation, measure binding over time (typically 10 to 20 measurements) to determine how rapidly the ligand dissociates from the receptors. The law of mass action predicts that dissociation of radioligands from receptors follows this equation: total binding = NS + ( Total − NS) × e− koff t Total binding and nonspecific binding (NS) are expressed in cpm, fmol/mg protein, or sites/cell. Time (t) is usually expressed in minutes. The dissociation rate constant (koff) is expressed in units of inverse time, usually min−1. Since it is hard to think in those units, it helps to calculate the half-life for dissociation, which equals ln(2)/koff or 0.6931/koff. In one half-life, half the radioligand will have dissociated (see Fig. A.3H.14). In two half-lives, three quarters of the radioligand will have dissociated, etc. Typically the dissociation rate constant of useful radioligands is between 0.001 and 0.1 min−1. If the dissociation rate constant is any faster, it would be difficult to perform radioligand binding experiments as the radioligand would dissociate from the receptors while you wash the filters. If the dissociation rate constant is any slower, it would be hard to reach equilibrium.
Total binding
total
NS t 1/2
Analyzing Radioligand Binding Data
Time
Figure A.3H.14 Schematic of a dissociation kinetic experiment.
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Specific binding
max
Time
Figure A.3H.15 Schematic of an association kinetic experiment.
Association binding experiments Association binding experiments are used to determine the association rate constant. This value is useful to characterize the interaction of the ligand with the receptor. It also is important as it permits the determination of how long it takes to reach equilibrium in saturation and competition experiments. To perform an association experiment, add a single concentration of radioligand and measure specific binding at various times thereafter. Association of ligand to receptors (according to the law of mass action) follows this equation:
(
specific binding = Max × 1 − e− kob ×t
)
In Figure A.3H.15, note that the maximum binding (Max) is not the same as Bmax. The maximum (equilibrium) binding achieved in an association experiment depends on the concentration of radioligand. Low to moderate concentrations of radioligand will bind to only a small fraction of all the receptors no matter how long binding is allowed to proceed. Note that the equation used for fitting does not include the association rate constant, kon, but rather contains the observed rate constant, kob, which is expressed in units of inverse time (usually min−1). The kob is a measure of how quickly the incubation reaches equilibrium, and in the case of a simple bimolecular binding reaction is defined by this equation: kob = koff + kon × [radioligand ]
The equation defines kob as a function of three factors: 1. The association rate constant, kon. This is what is to be determined. If kon is larger (faster), kob will be larger as well. 2. The concentration of radioligand. When using more radioligand, the system will equilibrate faster and kob will be larger. 3. The dissociation rate constant, koff. It may be surprising to discover that the observed rate of association depends in part on the dissociation rate constant. This makes sense because an association experiment doesn’t directly measure how long it takes radioligand to bind, but rather measures how long it takes the binding to reach equilibrium. Equilibrium is reached when the rate of the forward binding reaction
Commonly Used Techniques
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equals the rate of the reverse dissociation reaction. If the radioligand dissociates quickly from the receptor, equilibrium will be reached faster, but there will be less binding at equilibrium. If the radioligand dissociates slowly, equilibrium will be reached more slowly and there will be more binding at equilibrium. To calculate the association rate constant, usually expressed in units of M−1 min−1, use the following equation. Typically ligands have association rate constants of ∼108 M−1 min−1. kon =
kob − koff [radioligand ]
Analyzing Association Binding Data Using nonlinear regression to determine koff 1. Enter the x data as time in minutes. 2. Enter the y data as total binding in cpm. 3. Choose the exponential dissociation equation. It may be expressed as: y = Span × exp ( −K × x ) + Plateau
4. If the chosen nonlinear regression program does not provide initial estimates of the variables, enter these values. Span is the specific binding at time zero and is estimated as the first y value minus the last y value. Plateau is the binding after a long time, and reflects nonspecific binding, which does not change with time. Estimate it as the last y value. K is the dissociation rate constant (koff). Estimate it by dividing 0.69 by an estimate of the half-time of dissociation. 5. Start the nonlinear regression procedure. 6. Calculate the half-life of dissociation from the rate constant. half-life = ln ( 2 ) koff = 0.693 K
Using nonlinear regression to determine kon 1. Enter the x data as time in minutes. 2. Enter the y data as specific binding in cpm. 3. Choose the exponential association equation. It may be expressed as: y = ymax ×{1 − exp ( −K × x )} 4. If the nonlinear regression program does not provide initial estimates of the variables, enter these values. ymax is the specific binding at equilibrium, which is estimated as the last y value. K is the observed rate of association (kob). Estimate it by dividing 0.69 by an estimate of the time it takes to achieve half-maximal binding. 5. Start the nonlinear regression procedure to determine k and ymax. Note that k is the observed rate constant (kob) in units of inverse time (min−1) if you entered x in minutes. This is not the same as the association rate constant. 6. Calculate the association rate constant from the observed rate constant and the dissociation rate constant. Use the following equation, where [radioligand] is expressed in molar and kob and koff are expressed in min−1. Analyzing Radioligand Binding Data
kon = ( kob − koff ) [radioligand ]
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ln(specific binding)
slope = −k off
Time
Figure A.3H.16 Schematic of a dissociation kinetic experiment shown on a log scale. The y axis plots the natural log of specific binding.
Displaying dissociation data on a log plot Figure A.3H.16 shows a plot of ln(Bt/B0) versus time. The graph of a dissociation experiment will be linear if the system follows the law of mass action with a single affinity state. Bt is the specific binding at time t; B0 is specific binding at time zero. The slope of this line will equal −koff. The log plot will only be linear when taking the logarithm of specific binding as a fraction of binding at time zero. Don’t use total binding. Use the natural logarithm, not the base ten log in order for the slope to equal −koff. If you use the base 10 log, then the slope will equal −2.303 times koff. Use the log plot only to display data, not to analyze data. A more accurate rate constant will be obtained by fitting the raw data using nonlinear regression. Interpreting the Results Are the results reasonable? Table A.3H.6 presents some questions that should be addressed when determining if the results are reasonable. Using Kinetic Data to Test the Law of Mass Action Standard binding experiments are usually fit to equations derived from the law of mass action. Kinetic experiments provide a more sensitive test than equilibrium experiments to determine whether the law of mass action actually applies for the system of interest. To test the law of mass action, ask these questions: Does the Kd calculated from kinetic data match the Kd calculated from saturation binding? According to the law of mass action, the ratio of koff to kon is the Kd of receptor binding: Kd =
koff kon
The units are consistent: koff is in units of min−1 and kon is in units of M−1min−1, so Kd is in units of M.
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If binding follows the law of mass action, the Kd calculated in this way should be the same as the Kd calculated from a saturation binding curve. Does kob increase linearly with the concentration of radioligand? The observed association rate constant, kob, is defined by this equation: kob = koff + kon × [radioligand ]
Therefore, association rate experiments performed at various concentrations of radioligand should look like Figure A.3H.17. As the concentration of radioligand is increased, the observed rate constant increases linearly. If the binding is more complex than a simple mass action model (such as a binding step followed by a conformational change) the plot of kob vs. [radioligand] may plateau at higher radioligand concentrations. The y intercept of the line equals koff. If the law of mass action applies to the system, the koff determined in this way should correspond to the koff determined from a dissociation experiment. Finally, this kind of experiment provides a more rigorous determination of kon than the value obtained with a single concentration of radioligand. Is specific binding 100% reversible, and is the dissociated ligand chemically intact? Nonspecific binding at “time zero” should equal total binding at the end (plateau) of the dissociation. In other words, all of the specific binding should dissociate after a sufficiently long period of time. Use chromatography to analyze the radioligand that dissociates to prove that it has not been altered.
Table A.3H.6
Analyzing Radioligand Binding Data
Evaluating the Results of Association Binding Analyses
Question
Comment
Was data collected over a long enough period of time?
Dissociation and association data should plateau, so the data obtained at the last few time points should be indistinguishable.
Is the value of kon reasonable?
The association rate constant, kon, depends largely on diffusion, so the value is similar for many ligands. Expect a result of ∼108 M−1 min−1.
Is the value of koff reasonable?
If the koff is >1 min−1, the ligand has a low affinity for the receptor. Most likely, dissociation will occur during separation of bound and free ligands. If koff is <0.001 min−1, attaining equilibrium will be difficult as the half-time of dissociation will be greater than 10 hr! Even if one waits that long, other reactions may occur that ruin the experiment.
Are the standard errors too large? Are the confidence intervals too wide?
Examine the SE and the confidence intervals to gauge the level of confidence to give the rate constants.
Does only a tiny fraction of radioligand bind to the receptors?
The standard analyses of association experiments assume that the concentration of free radioligand is constant during the experiment. This will be approximately true only if a tiny fraction of the added radioligand binds to the receptors. Compare the maximum total binding in cpm to the amount of added radioligand in cpm. If that ratio exceeds ∼10%, revise the experimental protocol.
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k ob (mi n −1)
slope = k on
k off [Radioligand] Figure A.3H.17 Schematic of observed association rate constants as a function of radioligand concentration. Higher concentrations of radioligand equilibrate more quickly. The slope of the line equals the association rate constant (kon); the y intercept is the dissociation rate constant (koff).
Is the dissociation rate the same when dissociation is initiated from different amounts or times of receptor occupation? If the ligand binds to a single site and obeys the law of mass action, the dissociation rate constant is independent of the amount of radioligand used or the time before initiating dissociation. Is there cooperativity? If the law of mass action applies, binding of a ligand to one binding site does not alter the affinity of another binding site. This also means that dissociation of a ligand from one site should not change the dissociation of ligand from other sites. To test this assumption, compare dissociation initiated by adding an unlabeled ligand with dissociation initiated by infinite dilution. The two rate constants should be identical (see Competitive Binding with Two Sites, below). Kinetics of Competitive Binding The standard methods of kinetic binding determine the kon and koff for a labeled ligand. Competitive binding can be used to determine the kon and koff of an unlabeled ligand. Add the two ligands at the same time, and measure radioligand binding over time. Use the following information to set up the equation (Motulsky and Mahan, 1984). Define the following variables. k1 association rate constant of radioligand (M−1 min−1) k2 dissociation rate constant of radioligand (min−1) k3 association rate constant of unlabeled ligand (M−1 min−1) k4 dissociation rate constant of unlabeled ligand (min−1) [radioligand] concentration of labeled drug (M) [unlabeled drug] concentration of unlabeled drug (M) S an arbitrary designation of an intermediate variable Bmax total number of binding sites (same units as specific binding, usually cpm) Commonly Used Techniques
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t time (min) K A = k1 × [radioligand ] + k2 K B = k3 × [unlabeled ligand ] + k4
+ 4 × k1 × k3 × [radioligand ] × [unlabeled ligand ]
S=
(KA − KB )
KF =
KA + KB + S 2
2
KS =
KA + KB − S 2
specific binding = Bmax × k1 × [radioligand ] k4 × ( KF − KS ) k4 − KF − KF ⋅t k4 − KS − KS ⋅t + ×e − ×e K F − KS KF KS K F × KS
Many data points are needed at early time points for this method to work. When fitting the data, set k1 and k2 to constant values determined from standard kinetic experiments. Set Bmax to a constant value determined in a saturation binding experiment. The concentrations of labeled and unlabeled compound are also constants, set by your experimental design. Fit the data to determine k3 and k4. TWO BINDING SITES Several receptor molecules frequently evolve for a single hormone or neurotransmitter. Also, many ligands bind to more than one receptor subtype. Saturation Binding Experiments with Two Sites When the radioligand binds to two classes of receptors, analyze the data by using this equation. specific binding = y =
Bmax1 × [L ] Kd1 + [L ]
+
Bmax2 × [L] Kd2 + [L]
Panel A of Figure A.3H.18 shows specific binding to two classes of receptors present in equal quantities, whose Kd values differ by a factor of ten. Panel B shows the transformation to a Scatchard plot. In both graphs the dotted and dashed lines show binding to the two individual receptors; the sum in each graph is represented by a solid curve. Note that the graph of specific binding is not obviously biphasic. It is very hard to see the presence of two binding affinities by just looking. The best way to detect the second site is to fit data to one- and two-site curves, and let the nonlinear regression program compare the two fits (see Theory: Comparing One- and Two-Site Models, below). The curvature of the Scatchard plot is not dramatic and can easily be obscured by experimental scatter. Note the location of the solid and dashed line in the Scatchard plot. The two components of a biphasic Scatchard are not the asymptotes of the curve. Competitive Binding with Two Sites Competitive binding experiments are often used in systems where the tissue contains two classes of binding sites (e.g., two subtypes of a receptor). Analysis of these data are straightforward if the following assumptions are met: Analyzing Radioligand Binding Data
1. There are two distinct classes of receptors. For example, a tissue could contain a mixture of β1 and β2 adrenergic receptors.
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2. The unlabeled ligand has distinct affinities for the two sites. 3. The labeled ligand has equal affinity for both sites. 4. Binding has reached equilibrium. 5. A small fraction of both labeled and unlabeled ligand bind. This means that the concentration of labeled and unlabeled ligand added is very close to the free concentration in all tubes. Based on these assumptions, binding follows the equation: F 1− F + y = NS + ( Total − NS) log[D]− log( IC50 A ) log[D]− log( IC50 B) 1 + 10 1 + 10 This equation has five variables: the total and nonspecific binding (the top and bottom binding plateaus), the fraction of binding to receptors of the first type of receptor (F), and the IC50 of the unlabeled ligand for each type of receptor. If the Kd and concentration of
200
Specific binding
A
100
0 0
25
50
75
100
[Radioligand] 75
Bound/free
B
50
25
0 0
50
100
150
200
Specific binding
Figure A.3H.18 Saturation binding to two classes of receptors. The two receptor types are present in equal quantities, but have Kd values that differ by a factor of ten. (A) Binding to the two individual receptor types are shown as dashed curves. The sum (observed experimentally) is shown as a solid curve. It is not obviously biphasic. (B) Scatchard transformation. The curvature of the overall Scatchard plot (solid) is subtle, and it would be easy to miss the curvature if the data were scattered. Note that the Scatchard plots for the individual receptors (dashed) are not asymptotes of the two-site Scatchard plot (solid).
Commonly Used Techniques
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the labeled ligand is known, the IC50 values can be converted to Ki values (see Analyzing Competitive Binding Data, above). Since there are two different kinds of receptors with different affinities, a biphasic competitive binding curve might be expected. In fact, a biphasic curve is seen only in unusual cases where the affinities are extremely different. More often, the two components are blurred together into a shallow curve. For example, Figure A.3H.19 shows competition for two equally abundant sites with a tenfold (one log unit) difference in IC50. Careful observation will reveal that the curve is shallow (it takes more than two log units to go from 90% to 10% competition), but two distinct components are not visible. Cooperativity In the standard mass action model, each binding site is independent. The standard mass action assumes that there is no cooperativity. Cooperativity occurs when binding of a ligand to one binding site affects binding to adjacent sites. Usually these binding sites are on the same molecule. If binding of one ligand increases the affinity of an adjacent site, this is positive cooperativity. If binding of one ligand decreases the affinity of an adjacent site, this is negative cooperativity. It is impossible to distinguish negative cooperativity from multiple independent binding sites (with different affinities) from data collected at equilibrium. Kinetic experiments are needed. To distinguish between multiple independent binding sites and negative cooperativity, compare the dissociation rate after initiating dissociation by infinite dilution with the dissociation rate when initiated by addition of a large concentration of unlabeled drug. If the radioligand is bound to multiple noninteracting binding sites, the dissociation will be identical in both experimental protocols as shown in panel A of Figure A.3H.20. Note that the y axis is shown using a log scale. If there were a single binding site, the dissociation data would be expected to appear linear on this graph. With two binding sites, the graph is curved, even on a log axis (assuming the radioligand is present at high enough concentration to bind appreciably to both sites).
Specific binding
Panel B shows ideal dissociation data when radioligand is bound to interacting binding sites with negative cooperativity. The data are different depending on how dissociation was initiated. If dissociation is initiated by infinite dilution, the dissociation rate will change over time. The dissociation of some radioligand will leave the remaining ligand bound more tightly. When dissociation is initiated by addition of cold drug, all the
IC50 site A
IC50 site B
Log[unlabeled competitor]
Analyzing Radioligand Binding Data
Figure A.3H.19 Two site competitive binding curve. The radioligand binds identically to two kinds of receptors, but these two receptors have a tenfold difference in affinity for the competitor. The curve is shallow, but not obviously biphasic.
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receptors are always occupied by ligand (some hot, some cold) and dissociation occurs at its maximal unchanging rate. Theory: Comparing One- and Two-Site Models Why not just compare sum of squares or R2? In a least squares analysis of data (either linear or nonlinear), the computer program will give an R2 value and the sum of the squared deviations from the theoretical fit in the experimental result. The smaller the sum-of-squares value and the higher the R2, the better the theory fits the data. However, a two-site model will almost always fit the data better than a one-site model. A three-site model fits even better, and a four-site model better yet! As more variables (sites) are added to the equation, more inflection points are added to the curve, so it gets closer to the points. The sum of squares gets smaller; R2 gets higher. Statistical calculations (such as the F test described below) should be used to see whether these changes are larger than expected by chance. Reality check Before performing statistical comparisons, however, look at whether the results make sense. Sometimes the two-site fit gives results that are clearly nonsense. Disregard a two-site fit when: (1) the two IC50 or Kd values are almost identical—the data are probably fit quite well by a single-site model; (2) one of the IC50 or Kd values is outside the range of data; (3) one of the sites has a very small fraction of the receptors—if there are too few sites, the IC50 or Kd cannot be determined reliably. (4) The best-fit values for the bottom and top plateaus are far from the range of y values observed in the experiment (applies to competitive binding curves only).
1.00
Bt /B0
A
0.10
0.01 1.00
Bt /B0
B
0.10
infinite dilution
0.01
excess unlabeled Time
Figure A.3H.20 Discriminating between binding to two (or more) binding sites (top) and negative cooperativity. With negative cooperativity, dissociation will be faster when initiated by adding excess unlabeled ligand than when initiated by infinite dilution.
Commonly Used Techniques
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If the two-site fit seems reasonable, test whether the difference between the one- and two-site fit is statistically significant. Using the F test to compare one- and two-site fits Even if the simpler one-site model is correct, the fit is expected to be worse (have the higher sum of squares) because it has fewer inflection points (more degrees of freedom). In fact, statisticians have proven that the relative increase in the sum of squares (SS) is expected to equal the relative increase in degrees of freedom (DF). In other words, if the one-site model is correct it would be expected that:
(SS1 − SS2 )
SS2 ≈ ( DF1 − DF2 ) DF2
If the more complicated two-site model is correct, then the relative increase in sum of squares (going from two-site to one-site) is expected to be greater than the relative increase in degrees of freedom:
(SS1 − SS2 )
SS2 > ( DF1 − DF2 ) DF2
Follow these steps to compare the two models: 1. Fit the data to the simpler (one-site) model and record the sum of squares (SS1) and degrees of freedom (DF1). 2. Fit the data to the more complicated (two-site) model and record the sum of squares (SS2) and degrees of freedom (DF2). 3. Look at whether the two-site model makes sense. If the best-fit values don’t make sense (or the values for the two sites are almost the same), then discard the two-site model and accept the one-site model. 4. Compare SS2 with SS1. If for some reason SS2 is larger than SS1, then the two-site fit is worse than the one-site fit and should be discarded. Accept the one-site fit. In most cases SS1 is larger, and further calculations will be needed. 5. Calculate the F ratio, which quantifies the relationship between the relative increase in sum of squares and the relative increase in degrees of freedom. F=
(SS1 − SS2 ) SS2 (DF1 − DF2 ) DF2
The equation for calculating F is usually presented in this equivalent form (see Table A.3H.7 for corresponding ANOVA table). F=
(SS1 − SS2 ) (DF1 − DF2 ) SS2 DF2
Table A.3H.7 Fitsa
Analyzing Radioligand Binding Data
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DFn = (DF1 − DF2 ), DFd = DF2
ANOVA Table for Comparison of One- and Two-Site
Mean square
Source of variation
Sum of squares
DF
Difference
SS1 − SS2
DF1 − DF2 SSI − SS2 DF1 - DF2
Model 2 (complicated)
SS2
DF2
Model 1 (simple)
SS1
DF1
SS2/DF2
aANOVA, analysis of variance.
Current Protocols in Protein Science
Binding (cpm)
3000
2000
1000
two sites one site
0 –10
–9
–8 –7 –6 Log[competitor]
–5
–4
Figure A.3H.21 The solid curve shows the fit to an equation describing competition for a single class of receptors. The dashed curve shows the fit to an equation describing competition for binding to two classes of receptors.
6. Use a table or program to determine the P value. When doing so, degrees of freedom should be entered for both the numerator (DFn) and denominator (DFd). The numerator has (DF1 − DF2) degrees of freedom. The denominator has DF2 degrees of freedom. If the one-site model is correct, an F ratio near one and a large P value are expected. If the two-site fit is correct, a large F ratio and a small P value would be seen. The P value can be small for two reasons. One possibility is that the two-site model is correct. The other possibility is that the one-site model is correct, but random scatter led the two-site model to fit better by chance. The P value tells how rarely this coincidence would occur. More precisely, the P value answers this question: If the one-site model is really correct, what is the chance that data would randomly fit the two-site model so much better? If the P value is smaller than a preset threshold (set to the arbitrary value of 0.05 by tradition), conclude that the two-site model is significantly better than the one-site model. Figure A.3H.21 compares a one-site and two-site competitive binding curve. The results are shown in Table A.3H.8. Table A.3H.8 Comparison of One-Site and Two-Site Competitive Binding Curve
Degrees of freedom Sum of squares
Two-site
One-site
% increase
10 129,800
12 248,100
20.00 91.14
In going from the two-site to the one-site model, two degrees of freedom are gained, because the one-site model has two fewer variables. Since the two-site model has 10 degrees of freedom (15 data points minus 5 variables), the degrees of freedom increased 20%. If the one-site model were correct, the sum of squares would be expected to increase ∼20% just by chance. In fact, the sum of squares increased 91%. The percent increase was 4.56 times higher than expected (91.1/20.0 = 4.56). This is the F ratio (F = 4.56), and it corresponds to a P value of 0.039. If the one-site model is correct, there is only a 3.9%
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Percent specific binding
100 +GTP
50
–GTP
0 Log[agonist]
Figure A.3H.22 Schematic of agonist competition for binding to a receptor linked to a G protein. In the absence of GTP (left) the curve is shallow (and in this extreme case, biphasic). In the presence of GTP (or an analog) the curve is shifted to the right and is steeper.
chance that randomly obtained data would fit the two-site model so much better. Since this is below the traditional threshold of 5%, conclude that the two-site model fits significantly better than the one-site model. AGONIST BINDING Receptors Linked to G Proteins The most studied example of agonist binding is the interaction of agonists with receptors that are linked to G proteins. This is studied by comparing the competition of agonists with radiolabeled antagonist binding in the presence and absence of GTP (or its analogs). These experiments are done in membrane preparations to wash away endogenous intracellular GTP. Without added GTP, the competitive binding curves tend to be shallow. When GTP or an analog is added, the competitive binding curve is of normal steepness. Figure A.3H.22 shows the results of an idealized experiment. The extended ternary complex model can partially account for these findings (and others). In this model, receptors can exist in two states, R and R*. The R* state has a high affinity for agonist and preferentially associates with G proteins to form an R*G complex. Although some receptors may exist in the R* state in the absence of agonist, the binding of agonist fosters the transition from R to R*, and thus promotes interaction of the receptor with G protein to form the ternary complex HR*G. The extended ternary complex model is shown in Figure A.3H.23.
Analyzing Radioligand Binding Data
The agonist binding curve is shallow (showing high and low affinity components) in the absence of GTP because some receptors interact with G proteins and others do not. The receptors that do interact with G proteins bind agonist with high affinity, while those that do not interact bind with low affinity. Not all receptors can bind to G proteins because either the receptors are heterogeneous, the G proteins are limiting, or the membrane compartmentation prevents some receptors from interacting with G proteins. If all the receptors could interact with G proteins, the expectation would be a high affinity, competitive binding curve in the absence of GTP. In the presence of GTP (or an analog), the HR*G complex is not stable, so the G protein dissociates into its αGTP and βγ subunits, and is uncoupled from the receptor. With GTP present, only a tiny fraction of receptors
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H+R+G
HR+G
H+R+G
HR+G
H+R+G
HR+G
H+R*+G
HR*+G
H+RG
HRG
H+RG
HRG
H+R*G
HR*G
Simple model
Ternary complex model
Extended ternary complex model
Figure A.3H.23 Models for agonist binding to receptors linked to G proteins. H, hormone or agonist; R, receptor; G, G protein.
are coupled to G at any given time, so the agonist competition curves are of low affinity and normal steepness as if only R was present and not RG. Although the extended ternary complex model is very useful conceptually, it is not very useful when analyzing data. There are simply too many variables. The simpler ternary complex model shown in Figure A.3H.23 has fewer variables, but still too many to reliably fit with nonlinear regression. For routine analyses, most investigators fit data to the much simpler two-state model shown in the figure. This model allows for receptors to exist in two affinity states (R and RG), but does not allow conversion between them. It is easy to fit data to this simpler model using a two-site competition curve model. Since the model is too simple, the high and low affinity dissociation constants derived from the model should be treated merely as empirical descriptions of the data and should not be thought of as true molecular equilibrium constants. Other Kinds of Receptors By definition, the binding of agonists to receptors makes something happen. So it is not surprising that agonist binding is often more complicated than the simple mass action model. For example, binding of agonists to nicotinic acetylcholine receptor causes a conformational change characterized by a high affinity binding of the agonist and desensitized receptors, and insulin binding to its receptor shows negative cooperativity due to dimerization of the receptors. ANALYZING DATA USING NONLINEAR REGRESSION Radioligand binding data are best analyzed using nonlinear regression to fit curves through the data. The Problem with Using Linear Regression on Transformed Data Before the age of microcomputers, scientists transformed their data to make a linear graph, and then analyzed the transformed data with linear regression. Examples include Lineweaver-Burke plots of enzyme kinetic data, Scatchard plots of binding data, and logarithmic plots of kinetic data. These methods are outdated, and should not be used to analyze data. The problem is that the linear transformation distorts the experimental error. Linear regression assumes that the scatter of points around the line follows a Gaussian distribution and that the standard
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deviation is the same at every value of x. These assumptions are usually not true with transformed data. A second problem is that some transformations alter the relationship between x and y. For example, in a Scatchard plot the value of x (bound) is used to calculate y (bound/free), and this violates the assumptions of linear regression. For an example of this, see Analysis of Saturation Binding Curves, The Problem with Using Scatchard Plots. Since the assumptions of linear regression are violated, the results of linear regression are incorrect. The values derived from the slope and intercept of the regression line are not the most accurate determinations of the receptor number, rate constants, or dissociation constants. Considering all the time and effort put into collecting data, the best possible analysis technique should be used, and nonlinear regression produces the most accurate results. Although linear regression is usually inappropriate for analyzing transformed data, it is often helpful for displaying transformed data because many people find it easier to visually interpret linear data. This makes sense because the human eye and brain evolved to detect edges (lines), not to detect rectangular hyperbolas or exponential decay curves. Comparison of Linear and Nonlinear Regression A line is described by a simple equation that calculates y from x, slope, and intercept. The purpose of linear regression is to find values for the slope and intercept that define the line that best fits the data. More precisely, it finds the line that minimizes the sum of the squares of the vertical distances of the points from the line. The goal of minimizing the sum of squares in linear regression can be achieved quite simply. A bit of algebra (shown in many statistics books) derives equations that define the best-fit slope and intercept. Put the data in, do a few calculations, and the answers come out. There is no chance for ambiguity. Nonlinear regression fits data to any equation that defines y as a function of x and one or more variables. Like linear regression, it finds the values of those variables that minimize the sum of the squares of the vertical distances of the points from the curve. With the exception of a few special cases (like linear regression), it is not possible to solve the equations directly to find the best-fit values of the variables. Instead nonlinear regression requires an iterative approach that requires use of a computer. To analyze data with nonlinear regression, the program will require an equation (model) that defines y as a function of x and one or more variables (i.e., Kd, Bmax, or rate constants). It will also require an estimate (or guess) for the best-fit value for each variable in the equation (some programs provide the initial estimates automatically). Every nonlinear regression program follows these steps: 1. Using the initial values provided, the program calculates a predicted value of y for each value of x. It then compares the actual y values with the predicted y values, and calculates the sum of the squares of the differences between observed and predicted y values.
Analyzing Radioligand Binding Data
2. The program then adjusts the variables to improve the fit and reduce the sum of squares. There are several algorithms for adjusting the variables. The most commonly used method was derived by Levenberg and Marquardt (often called simply the Marquardt method), but the details of how this method works cannot be understood without first mastering matrix algebra. However, nonlinear regression can be used to analyze data without knowing anything about these algorithms.
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3. Step 2 is repeated several times. Each time the variables are adjusted by a smaller amount. Stop when the adjustments make virtually no difference in the sum of squares. This typically requires 5 to 20 iterations. 4. Best-fit results are reported. The precise values obtained will depend in part on the initial values and the stopping criteria. This means that repeat analyses of the same data will not always give exactly the same results. Picking a Nonlinear Regression Program When choosing a program to analyze binding data, beware of these three traps: 1. Rather than requesting an equation, some programs automatically fit data to hundreds or thousands of equations and then present the equation(s) that fit the data best. Using such a program is appealing because it frees the user from the need to choose an equation. The problem is that the program has no scientific understanding of the experiment and uses equations that do not correspond to any model of binding. It will not be possible to interpret the best-fit values of the variables in terms of rate constants or affinities. 2. Many so-called curve fitting programs actually fit data to a polynomial equation: y = A + Bx + Cx2 + Dx3…. Since the binding of ligands to receptors cannot be expressed as a polynomial equation, polynomial regression is not useful for analyzing binding data. 3. Some programs fit curves by drawing a cubic spline curve—a smooth curve that goes through every data point. In some cases, a cubic spline curve can look attractive on a graph and it may work well as a standard curve for interpolation. However, the curve does not correspond to any model, so cubic spline is not useful in data analysis. Once the search is narrowed to programs that perform nonlinear regression to fit data to a selected equation, ask the following questions: 1. Can common binding equations be selected from a menu, or must the equations be entered into a file? 2. Does the program provide initial values automatically (see below), or will they have to be entered for each analysis? 3. Can the program automatically compare two models with the F test? 4. Does the program prepare publication-quality graphs of the data and curves? 5. When the data are saved to file, are the analysis choices and results saved as well? This is important to retrace the analysis steps in the future. 6. Is the program designed specifically for analyses of binding data? The program Ligand (Munson and Rodbard, 1980) was designed just for analyses of binding studies, and can handle situations that many other programs cannot (i.e., simultaneous analyses of several experiments; correcting for ligand depletion), although its interface is no longer state-of-the-art. One nonlinear regression program, GraphPad Prism (designed by the authors of this chapter), is particularly well suited for analyses of radioligand binding data (see Analyzing Data with GraphPad Prism, below).
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Decisions That Need to be Made When Fitting Curves with Nonlinear Regression When using a program for nonlinear regression, the following decisions must be made. Which equation? An equation must be chosen that defines y as a function of x and one or more variables. This equation should represent a model, usually the law of mass action. Later sections of this unit show equations that describe the law of mass action in various kinds of binding experiments. Which units? In pure mathematics, it doesn’t matter whether data is entered as 1 picomolar or 10−12 molar, as 100 fmol/mg or 60,000,000,000 receptors/mg. When computers do the calculating, however, it can matter. Calculation problems such as round off errors are far more likely when the values are very high or very low. Scale data to avoid values <10−4 or >104. Estimate initial values Nonlinear regression is an iterative procedure. The program must start with estimated values for each variable that are in the right “ball park”—usually within a factor of five of the actual value. It then adjusts these initial values to improve the fit. It repeats the adjustments until the improvement is no longer significant. Later sections of this unit explain how to choose initial values for various kinds of experiments. The estimates don’t need to be extremely accurate. Nonlinear regression will usually work fine as long as the estimates are within 3 to 5 times their actual values. Some programs (including GraphPad Prism) automatically choose initial values for you. Fix one or more variables to a constant value? In some situations it makes sense to fix some of the variables to constant values. For example, when analyzing specific (rather than total) binding, the bottom plateau of a dissociation experiment should be defined as a constant equal to zero. Weighting In general, the goal of nonlinear regression is to find the values of the variables in the model that make the curve come as close as possible to the data points. Usually this is done by minimizing the sum of the squares of the vertical distances of the data points from the curve. This is appropriate when the scatter of points around the curve is expected to be Gaussian and unrelated to the y values of the points. With many experimental protocols, the experimental scatter is not expected to be the same for all points. Instead, the experimental scatter is expected to be a constant percentage of the y value. If this is the case, points with high y values will have more scatter than points with low y values. When the program minimizes the sum of squares, points with high y values will have a larger influence while points with smaller y values will be relatively ignored. This problem may be avoided by minimizing the sum of the square of the relative distances. This procedure is termed weighting the values by 1/y2. Because it prevents large points from being over-weighted, the term unweighting seems more intuitive. Data may be weighted in other ways. The goal is to obtain a measure of goodness-of-fit that weights all the data points equally. Analyzing Radioligand Binding Data
With binding data, scatter is often proportional to the amount of binding, so relative weighting may be appropriate. Results are usually very similar whether or not you choose to use relative weighting.
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Average replicates? If replicate y values are collected at every value of x, there are two ways to analyze the data: (1) treat each replicate as a separate point, or (2) average the replicate y values and treat the mean as a single point. With radioligand binding data, the first approach is usually best, because all the data are obtained from one tissue preparation and the sources of experimental error are independent for each tube. If one value happens to be a bit high, there is no reason to expect the other replicates to be high as well. Each replicate can be considered an independent data point. Do not treat each replicate as a separate point when the experimental error of the replicates are related. Instead, average the replicates and analyze the averages. This situation doesn’t come up often with radioligand binding data, but here is one example. Assume that you perform an experiment with only a single replicate at each value of y (concentration or time) but count each tube three times. It is not fair to enter the three counts as triplicates, and then analyze each triplicate as a separate value. As the replicates are not independent, any experimental error would appear in all the replicates. Assumptions of Nonlinear Regression The results of nonlinear regression are meaningful only if the following assumptions are true (or nearly true): 1. The model is correct. Nonlinear regression adjusts the variables in the equation you chose to minimize the sum of squares. It does not attempt to find a better equation. 2. The variability of values around the curve follow a Gaussian distribution. Even though no biological variable follows a Gaussian distribution exactly, it is sufficient that the variation be approximately Gaussian. 3. The standard deviation (SD) of the residuals is the same everywhere, regardless of the value of x. In other words, the average scatter of the points around the curve is the same at all parts of the curve. The assumption is termed homoscedasticity. If the SD is not constant but rather is proportional to the value of y, weight the data to minimize the sum of squares of the relative distances. 4. The model assumes that x is known exactly. This is rarely the case, but it is sufficient to assume that any imprecision in measuring x is very small compared to the variability in y. 5. The errors are independent. The deviation of each value from the curve should be random, and should not be correlated with the deviation of the previous or next point. If there is any carryover from one sample to the next, this assumption will be violated. EVALUATING RESULTS OF NONLINEAR REGRESSION Before accepting the results of nonlinear regression, the following questions should be asked: Did the Program Converge on a Solution? A nonlinear regression program will stop its iterations when it can’t improve the fit by adjusting to the values of any of the variables. At that point, the program is said to have converged on the best fit. In some cases, the program gets stuck. It doesn’t know whether the fit would improve by increasing or decreasing the value of a variable. When this happens, the program stops. The exact wording of the error message is unlikely to be
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helpful. In this situation, some programs may still apparently show results, but these “results” do not represent a best-fit curve. Are the Results Scientifically Plausible? The mathematics of curve fitting sometimes yields results that make no scientific sense. For example, noisy or incomplete data can lead to negative rate constants, fractions greater than 1.0, and negative Kd values. If the results make no scientific sense, they are unacceptable, regardless of R2 and of how close the curve comes to the points. Try a simpler equation, or try fixing some variables to constant values. Also check that the best-fit values of the variables are reasonable compared to the range of the data. Don’t trust the results if the top plateau of a sigmoid curve is far higher than the highest data point. Don’t trust the results if an EC50 value is not within the range of the y values. Does the Curve Come Close to the Points? In rare cases, the fit may be far from the data points. This may happen, for example, if the wrong equation is chosen. Look at the graph to make sure this didn’t happen. Goodness of fit can also be evaluated by looking at the value of R2 (known by statisticians as the coefficient of determination). R2 is the fraction of the total variance of y that is explained by the model (equation). Mathematically, it is defined by the equation: R2 = 1.0 − SS/sy2, where sy2 is the variance (standard deviation squared) of y values. The value of R2 is always between 0.0 and 1.0, and it has no units. When R2 equals 0.0, the best-fit curve fits the data no better than a horizontal line going through the mean of all y values. In this case, knowing x does not help you predict y. When R2 = 1.0, all points lie exactly on the curve with no scatter; if x is known, y may be calculated exactly. If R2 is high, the curve comes closer to the points than would a horizontal line through the mean y value, but a high R2 should not be overinterpreted. It does not mean that the chosen equation is the best to describe the data. It also does not mean that the fit is unique—other values of the variables may generate a curve that fits just as well. When comparing one- and two-site models, it is not sufficient to simply compare R2 values. Do the Data Systematically Deviate from the Curve? If the data really follow the model described by the chosen equation, the data points should be randomly scattered above and below the curve. The distance of the points from the curve should also be random, and not be related to the value of x. The best way to look for systematic deviations of the points from the curve is to inspect a graph of the residuals and to look at the runs test.
Analyzing Radioligand Binding Data
Residuals A residual is the distance of a point from the curve. A residual is positive when the point is above the curve, and is negative when the point is below the curve. The residual table has the same x values as the original data, but each y value is replaced by the vertical distance of the point from the curve. An example is shown in Figure A.3H.24. As shown
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150
R 2 = 0.97 Number of runs = 6 P value = 0.0025
Signal
100
50
0 –20
Residuals
–10
0
10
20 0.0
2.5
5.0
7.5 10.0 12.5 15.0 17.5 20.0 Time
Figure A.3H.24 Residuals. The top panel graphs dissociation kinetic data. The bottom panel shows the residuals (i.e., the y axis plots the distance between the point and the curve from the top panel).
in panel A, the data points are not randomly distributed above and below the curve. There are clusters of points all above or all below. This is much easier to see on the graph of the in panel B. The points are not randomly scattered above and below the x axis. The runs test The runs test determines whether the data deviate systematically from the equation you selected. A run is a series of consecutive points that are either all above or all below the regression curve. Another way of saying this is that a run is a series of points whose residuals are either all positive or all negative. If the data points are randomly distributed above and below the regression curve, it is possible to calculate the expected number of runs. If there are fewer runs than expected, it may mean that the regression model is wrong. If the data really follow the equation used to create the curve, the P value from the runs test may be used to determine the chance of obtaining as few (or fewer) runs as observed in the experiment. If the P value is small, it indicates that the data really don’t follow the model. In the example in Fig. A.3H.24, the equation does not adequately match the data. There are only six runs, and the P value for the runs test is very small. This means that the data systematically deviate from the curve, and the data were fit to the wrong equation. Commonly Used Techniques
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Are the Confidence Intervals Wide? In addition to reporting the values of the variables that make the equation fit the data best, nonlinear regression programs also express the uncertainty as a standard error for each variable. Use the standard error to calculate a 95% confidence interval (CI), if the nonlinear regression program doesn’t calculate one. The 95% confidence interval extends from approximately two standard errors below the best-fit value to approximately two standard errors above the best-fit value. (The number 2.0 is approximate. The exact multiplier comes from the t distribution and depends on the number of degrees of freedom which equals the number of data points minus the number of variables fit by the program.) The CI means that if all the assumptions of nonlinear regression are true, there is a 95% chance that the interval contains the true value. More precisely, if a nonlinear regression is performed many times (on different data sets) the expected confidence interval will include the true value 95% of the time, but exclude the true value the other 5% of the time. Three factors can make the confidence interval too narrow: 1. The CI is based only on the scatter of data points around the curve within this one experiment. If the experiment is repeated many times, the scatter between the results is likely to be greater than predicted from the CI determined in one experiment. 2. If any of the assumptions of nonlinear regression are violated, the confidence intervals will probably be too narrow. 3. The confidence intervals from nonlinear regression are calculated using mathematical shortcuts and so are referred to as asymptotic confidence intervals or approximate confidence intervals. In some cases these intervals can be too narrow (too optimistic). Because of these problems, the confidence intervals should not be interpreted too rigorously. Rather than focusing on the CI reported from analysis of a single experiment, repeat the experiment several times. If the confidence interval is extremely wide, do not trust the results. Confidence intervals are wide when the data are very scattered or data have not been collected over a wide enough range of x values. The data in Figure A.3H.25 were fit to a dose-response curve, and the 95% CI for the EC50 extends over six orders of magnitude. The explanation is simple. Since the data do not define plateaus at either the top or the bottom, zero and one hundred are not defined. This makes it impossible to determine the EC50 with precision. In this example, it might make scientific sense to set the bottom plateau to 0% and the top plateau to 100% (if the plateaus were defined by other controls not shown on the graph). If this were done, the equation would fit fine and the confidence interval would be narrow. Note that the problem with the fit is not obvious by inspecting a graph, because the curve goes very close to the points. The value of R2 (0.9999) is also not helpful. That value also indicates that the curve comes close to the points, but does not indicate whether the fit is unique.
Analyzing Radioligand Binding Data
The CI is also wide when data in an important part of the curve has not been collected. The dose-response curve in Figure A.3H.26 has wide confidence intervals. Even when constraining the bottom to be zero and the top to be 100 and the slope to equal 1.0, the 95% CI for the EC50 extends over almost an order of magnitude. The problem is simple.
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The EC50 is the concentration at which the response is half-maximal, and this example has no data near that point. Finally, the CI is wide if one tries to fit data to a two-site model when the data really follow a one-site model. In this case, the program might report very wide confidence intervals, as it will report that the two sites are very similar. Is the Fit a Local Minimum? The nonlinear regression procedure adjusts the variables in small steps in order to improve the goodness-of-fit. If Prism converges on an answer, altering any of the variables a little bit will make the fit worse. But it is theoretically possible that large changes in the
Response
100
50
fit to sigmoidal curve R 2 = 0.9999
0 Log[dose]
Figure A.3H.25 A dose-response curve with data collected over a narrow range of concentrations. When a nonlinear regression program tries to fit the top and bottom plateaus as well as the EC50 and slope, the resulting confidence intervals are very wide. Since there is no data to define zero and one hundred, the program will be very uncertain about the EC50. If the nonlinear regression program is told to set the top and bottom plateaus to constant values (from controls), then it can determine the EC50 with precision.
Percent response
100
50
0
–10
–8
–6
–4
Log[dose]
Figure A.3H.26 A dose response curve with no data in the middle of the curve. Since there are no data points in the middle of the curve, the best-fit value of the EC50 will be uncertain with a wide confidence interval.
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starting here, result will be false minimum
Sum of squares
starting here, result will be best-fit
false minimum best fit
Value of variable
Figure A.3H.27 What is a false minimum? A nonlinear regression program stops when making any small change to a variable will worsen the fit and thus raise the sum of squares. In rare cases, this may happen at a false minimum rather than the true best fit value.
variables might lead to much better goodness-of-fit. Thus, the curve that Prism decides is the “best” may really not be the best. Think of latitude and longitude as representing two variables Prism is trying to fit. Now think of altitude as the sum of squares. Nonlinear regression works iteratively to reduce the sum of squares. This is like walking downhill to find the bottom of the valley. When nonlinear regression has converged, changing any variable increases the sum of squares. When at the bottom of the valley, every direction leads uphill. But there may be a much deeper valley over the ridge that is unknown (see Fig. A.3H.27). In nonlinear regression, large changes in variables might decrease the sum of squares. This problem (called finding a local minimum) is intrinsic to nonlinear regression, no matter what program is used. A local minimum will rarely be encountered if the data have little scatter, data is collected over an appropriate range of x values, and an appropriate equation is chosen. To continue the analogy, the confidence intervals for the variables are very wide when the bottom of the valley is very flat. A great distance can be traveled without changing elevation. The values of the variables can be changed a great deal without changing the goodness-of-fit. To test for the presence of a false minimum: 1. Note the values of the variables and the sum of squares from the first fit. 2. Make a large change to the initial values of one or more variables and run the fit again. Repeat several times. 3. Ideally, Prism will report nearly the same sum of squares and same variables regardless of the initial values. If the values are different, accept the ones with the lowest sum of squares.
Analyzing Radioligand Binding Data
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What to Do When the Fit Is No Good? The previous sections explained how to identify a bad fit. If any of these situations are encountered, Table A.3H.9 describes some things to try. COMPARING TREATMENT GROUPS The results of radioligand binding experiments will often be compared between treatment groups. There are three ways to do this. Compare the Results of Repeated Experiments (Method 1) After repeating the experiment several times, compare the best-fit value of a variable in control and treated preparations using a paired t test (or the analogous Wilcoxon nonparametric test). For example, in Table A.3H.10, the log(Ki) values of results from a competitive binding curve performed in two groups of cells are shown. Compare the results using a paired t test. The t ratio is 16.7, and the P value is 0.0036 (two-tail). If the treatment did not alter the log(Ki), there is only a 0.36% chance that such a large difference (or larger) between log(Ki) is by chance. Since the P value is so low, conclude that the change in Ki was statistically significant. Note that we compare log(Ki) values rather than Ki values. When doing a paired t test, a key assumption is that the distribution of differences (treated vs. control) follow a Gaussian distribution. Since a competitive binding curve (similar to a dose response curve) is conducted with x values (concentration) equally spaced on a log scale, the uncertainty of the EC50 is reasonably symmetrical (and perhaps Gaussian) when expressed on a log scale. It is equally likely that the best-fit value of the log (Ki) is 0.1 log units too high or 0.1 log units too low. In contrast, the uncertainty in Ki is not symmetrical.
Table A.3H.9
Troubleshooting Guide to Evaluating Results of Nonlinear Regression
Potential problem
Solution
The equation simply does not describe the data.
Try a different equation.
The initial values are too far from their correct values.
Enter different initial values. If using a user-defined equation, check the rules for initial values.
The range of x values is too narrow to define the curve completely.
If possible, collect more data. Otherwise, hold one of the variables to a constant value.
There is not enough data collected in a critical range of x values.
Collect more data in the important regions.
The data are very scattered and don’t really define a curve.
Try to collect less scattered data. If combining several experiments, normalize the data for each experiment to an internal control.
The equation includes more than one component, but the data don’t follow a multicomponent model.
Use a simpler equation.
The numbers are too large.
If the y values are very large, change the units. Do not use values greater than ∼104.
The numbers are too small.
If your y values are very small, change the units. Do not use values less than ∼10−4.
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Table A.3H.10 Log(Ki) Values for a Sample Competitive Binding Experiment
Experiment
Control
Treated
1 2 3
−6.13 −6.39 −5.92
−6.53 −6.86 −6.31
Compare the Results within One Experiment: Simple Approach (Method 2) Use a t test to determine whether the difference between best-fit values is greater than would be expected by chance, given the standard errors of the variables. For example, competitive binding curves of control and treated data were compared in an experiment performed once. Nonlinear regression fit three variables, Top, Bottom, and log(EC50). Only the log(EC50) values are of interest. In this example, the control log(EC50) was −6.08 with a standard error of 0.3667. The treated log(EC50) was −6.20 with a standard error of 0.0617. Compare the two groups with an unpaired t test. 1. Calculate the t ratio as the difference between log(EC50) values divided by the standard error of that difference (calculated from the two standard errors). Since the sample size is the same in the two groups, use the equation: t=
log ( EC50 )A − log ( EC50 )B SEMA2 + SEMB2
= 2.292
2. Calculate the number of degrees of freedom (DF), which equals the sum of the number of degrees of freedom in each group. This equals the number of data points minus the number of variables fit by the nonlinear regression procedure. In this example, there were 15 data points, and three variables were fit. So there are 12 DF in each group, and 24 DF altogether. 3. Use a table or program to determine a P value that corresponds to the values of t and DF. For this example, the P value is 0.0309. If the treatment really didn’t alter the EC50, there is only a 3.09% chance that this large of a difference (or more) is by coincidence. Since the P value is so low, it is concluded that the two EC50 values are statistically significantly different. GraphPad Prism, GraphPad InStat and many other programs can compute t and the P value from data entered as mean, SEM, and N. Enter the best-fit value of the log(EC50) (or any other fit variable) instead of the mean, and the SE of that variable instead of the SEM. The trick is figuring out what value to enter as “N” (sample size). Remember that: 1. For nonlinear regression, the number of degrees of freedom equals the number of data points minus the number of variables fit. 2. For an ordinary t test, the number of degrees of freedom for each sample equals one less than the number of data points.
Analyzing Radioligand Binding Data
3. The t test calculations are based on the numbers of degrees of freedom. However, most programs ask for N instead and then compute DF as N − 1. When comparing the results of nonlinear regression, enter N as the number of degrees of freedom plus 1. The program will subtract 1 to determine the DF. All the other calculations are
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based on the value of DF, and N is ignored. In this example, enter N = 12 + 1 = 13 for each group. This method only uses data from one experiment. The SE value is a measure of how precisely the log(EC50) has been determined in this one experiment. It is not a measure of how reproducible the experiment is. Despite the impressive P value, these results should not be trusted until the experiment is repeated. The t test assumes that the uncertainty in the values of the variables follows a Gaussian distribution. This assumption is not necessarily true with the SE values that emerge from nonlinear regression. The only way to assess the validity of this assumption is to simulate many sets of data, fit each with nonlinear regression, and examine the distribution of best-fit values. This has been done with many commonly used equations, and it seems that the assumption is reasonable in many cases. Compare log(EC50), not EC50. You want to express the variables in a form that makes the uncertainty as symmetrical and Gaussian as possible. Since a competitive binding curve (similar to a dose response curve) is conducted with x values (concentration) equally spaced on a log scale, the uncertainty of log(EC50) is reasonably symmetrical (and perhaps Gaussian). It is equally likely that the observed log(Ki) is 0.1 log units too high or 0.1 log units too low. In contrast, the uncertainty in Ki is not symmetrical. Compare the Results Within One Experiment: More Complicated Approach (Method 3) The method of the previous section only compared the value of the log(EC50). This section describes a more general method to compare entire curves to ask whether the data sets differ at all. The idea is to first fit the two curves separately, and then combine the values and fit one curve to all the data. Follow these steps: 1. Fit the two data sets separately as in the previous section. 2. Total the sum of squares and DF from the two fits. For this example the total sum of squares equals 19,560 + 29,320 = 48,880, and the total DF equals 12 + 12 = 24. Since these are the results of fitting the two data sets separately, label these values SSseparate and DFseparate. 3. Combine the two data sets into one. For this example, the combined data set has 30 xy pairs, with each x value appearing twice. 4. Fit the combined data set to the same equation. Note the SS and DF. For this example, SS = 165,200, and DF = 27 (30 data points minus three variables). Call these values SScombined and DFcombined. 5. SSseparate is expected to be smaller than SScombined even if the curves are really identical, simply because the separate fits have more degrees of freedom. The question is whether the SS values are more different than expected by chance. To find out, calculate the F ratio using the equation: SScombined − SSseparate F = SScombined
DFcombined − DFseparate DFseparate
For this example, F = 19.03. Commonly Used Techniques
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6. Determine the P value from F. There are DFcombined − DFseparate degrees of freedom in the numerator, and DFseparate degrees of freedom in the denominator. GraphPad StatMate can calculate the P value (from F and the two DF values), or it may be found in the back of most statistics books. 7. For this example, the P value is <0.0001. If the treatment were really ineffective, there is less than a 0.01% chance that the two curves would differ as much (or more) as they differed in this experiment. Since the P value is low, you’ll conclude that the curves are really different. This method only uses data from one experiment. Despite the impressive P value, these results should not be trusted until the experiment is repeated. This method compares the curves overall. It doesn’t determine which variable(s) are different. Differences might be due to something trivial such as a different baseline, rather than something important such as a different EC50. Advantages and Disadvantages of the Three Approaches If the experiment has been repeated several times, use the first method. There are two advantages. The first is that compared to the other methods discussed below, this method is far easier to understand and communicate to others. Second, the entire test is based on the consistency of the results between repeat experiments. Since there are usually more causes for variability between experiments than within experiments, it makes sense to base the comparison on differences between experiments. The disadvantage of the first method is that information is being discarded. The calculations are based only on the best-fit value from each experiment, and they ignore the SE of those values presented by the curve fitting program. If the experiment has been performed only once, the experiment should be repeated. Regardless of what statistical results are obtained, results from a single experiment should not be trusted. To compare results in a single experiment, use Method 2 or 3. Generally only one variable is of interest (i.e., a rate constant or EC50); the others are less important. Method 2 compares the variable of interest. Method 3 is more general. Since the method compares the entire curve, it does not force a decision regarding which variable(s) to compare. This is both its advantage and disadvantage. CALCULATIONS WITH RADIOACTIVITY Efficiency of Detecting Radioactivity Efficiency is the fraction of radioactive disintegration that is detected by the counter. Efficiency is determined by counting a standard sample under conditions identical to those used in the experiment. With 125I, the efficiency is usually >90%, depending on the geometry of the counter. The efficiency is not 100% because the detector doesn’t entirely surround the tube, which allows a few gamma rays (photons) to miss the detector.
Analyzing Radioligand Binding Data
With 3H, the efficiency of counting is much lower, and usually varies between 40% and 50%. The low efficiency is mostly a consequence of the physics of decay and cannot be improved by better instrumentation or better scintillation fluid. When a tritium atom decays, a neutron converts to a proton and the reaction emits an electron and neutrino. The energy released is always the same, but it is randomly partitioned between the neutrino (not detected) and an electron (detection attempted). When the electron has
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sufficient energy, it will travel far enough to encounter a fluor molecule in the scintillation fluid. This fluid amplifies the signal and gives off a flash of light detected by the scintillation counter. The intensity of the flash (number of photons) is proportional to the energy of the electron. If the electron has insufficient energy, it is not captured by the fluor and is not detected. If it has low energy, it is captured but the light flash has few photons and is not detected by the instrument. Since the decay of many tritium atoms does not lead to a detectable number of photons, the efficiency of counting is much less than 100%. Efficiency of counting 3H is reduced by the presence of any color in the counting tubes, if the mixture of water and scintillation fluid is not homogeneous, or if the radioactivity is trapped in tissue (so emitted electrons don’t travel into the scintillation fluid). Specific Radioactivity Radioligand packaging usually states the specific radioactivity as Curies per millimole (Ci/mmol). Because measurements are expressed in counts per minute (cpm), the specific radioactivity is more useful when stated in cpm. Often the specific radioactivity is expressed as cpm/fmol (1 fmol = 10−15 mole). To convert from Ci/mmol to cpm/fmol, know that 1 Ci equals 2.22 × 1012 disintegrations per minute (dpm). Use this equation to convert Z Ci/mmol to Y cpm/fmol when the counter has an efficiency (expressed as a fraction) equal to E. Y
cpm fmol
=Z
Ci mmol
× 2.22 × 1012
Y = Z × 2.22 × E
dpm Ci
× 10
( in cpm
−12
mmol fmol
×E
cpm dpm
fmol )
For example, the specific activity will be 2190 Ci/mmol if every molecule incorporates exactly one 125I atom. If the counting efficiency is 85%, then the specific activity is 2190 × 2.22 × 0.85 = 4133 cpm/fmol. In many countries, radioligand packaging states the specific radioactivity in GBq/mmol, rather than Ci/mmol. To convert to cpm/fmol, you need to know that 1 Bq (Becquerel) is one radioactive disintegration per second (1 GBq = 109 dps). To convert from GBq/mmol to cpm/fmol, use this equation. Y
cpm fmol
=Z
GBq mmol
× 10 9
dps GBq
sec
× 60 min × 10
−12 mmol fmol
counts
× E disintegrations = Z × 0.06 × E
If every molecule is labeled with 125I, the specific activity is 81,030 GBq/mmol. If the counting efficiency is 85%, then the specific activity can also be expressed as 81,030 × 0.06 × 0.85 = 4133 cpm/fmol. Calculating the Concentration of the Radioligand Rather than trust dilutions, the concentration of radioligand in a stock solution can be accurately calculated. Measure the cpm in a small volume of solution and use the following equation, in which C is cpm counted, V is volume of the solution in ml, and Y is the specific activity of the radioligand in cpm/fmol (calculated in the previous section). concentration in pM =
Y cpm fmol V ml
C cpm 0.001pmol fmol × 0.001liter ml
= CVY Commonly Used Techniques
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Radioactive Decay Radioactive decay is entirely random. The probability of decay at any particular interval is the same as the probability of decay during any other interval. Starting with N0 radioactive atoms, the number remaining at time t is: Nt = N 0 × e
− kdecay t
The rate constant of decay (kdecay) is expressed in units of inverse time. Each radioactive isotope has a different value of kdecay. The value e refers to the base of natural logarithms (2.71828). The half-life (t1⁄2) is the time it takes for half the isotope to decay. Half-life and the decay rate constant are related by this equation: t1 2 =
ln(2) 0.693 = kdecay kdecay
Table A.3H.11 shows the half-lives and rate constants for commonly used radioisotopes. The table also shows the specific activity assuming that each molecule is labeled with one radioactive atom. (This is often the case with 125I and 32P. Tritiated molecules often incorporate two or three tritium atoms, which increases the specific radioactivity.) Radioactive decay can be calculated from a date where you knew the concentration and specific radioactivity using this equation. fraction remaining = e
− kdecay t
For example, after 125I decays for 20 days, the fraction remaining equals 79.5%. Although data appear to be scanty, most scientists assume that the energy released during decay destroys the ligand so it no longer binds to receptors. Therefore the specific radioactivity does not change over time. What changes is the concentration of ligand. After 20 days, the concentration of the iodinated ligand is 79.5% of what it was originally, but the specific radioactivity remains 2190 Ci/mmol. This approach assumes that the unlabeled decay product is not able to bind to receptors and has no effect on the binding. Rather than trust this assumption, use newly synthesized or repurified radioligand for key experiments. Calculations of radioactive decay are straightforward only when each molecule is labeled with a single radioactive isotope, as is usually the case. If a molecule is labeled with several radioactive isotopes, the effective half-life is shorter. If only a fraction of the molecules are labeled with a radioactive isotope, then the decay formula only applies to the labeled portion of the mixture, as the concentration of the unlabeled compound never changes.
Table A.3H.11 Isotopes
Half-Lives and Rate Constants for Commonly Used
Isotope
Half-life
kdecay
Specific radioactivity
3H
12.43 years 59.6 days
0.056 year−1 0.0116 day−1
28.7 Ci/mmol 2190 Ci/mmol
14.3 days 87.4 days
0.0485 day−1 0.0079 day−1
9128 Ci/mmol 1493 Ci/mmol
125I 32P 35S
Analyzing Radioligand Binding Data
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Counting Error and the Poisson Distribution The decay of a population of radioactive atoms is random, and therefore subject to a sampling error. For example, the radioactive atoms in a tube containing 1000 cpm of radioactivity won’t give off exactly 1000 counts in every minute. There will be more counts in some minutes and fewer in others, with the distribution of counts following a Poisson distribution. This variability is intrinsic to radioactive decay and cannot be reduced by more careful experimental controls. There is no way to know the “real” number of counts, but a range of counts can be calculated that is 95% certain to contain the true average value. As long as the number of counts (C) is greater than ∼100, the confidence interval can be calculated using this approximation:
(
) (
95% CI : C − 1.96 C to C + 1.96 C
)
Computer programs can calculate a more exact confidence interval, as becomes necessary when C is less than ∼100. For example, if C = 100, the simple equation above calculates a 95% confidence interval from approximately 80 to 120. A more exact equation calculates an interval from 81.37 to 121.61. When calculating the confidence interval, set C equal to the total number of counts you measured experimentally, not the number of counts per minute. For example, if a radioactive sample is placed into a scintillation counter for 10 min, the counter detects 225 counts per minute. What is the 95% confidence interval? Since the total time was 10 min, the instrument must have detected 2250 radioactive disintegrations. The 95% confidence interval of this number extends from 2157 to 2343. This is the confidence interval for the number of counts in 10 min, so the 95% confidence interval for the average number of counts per minute extends from 216 to 234. That is, there is a 95% certainty that the average cpm value lies within this range. The Poisson distribution explains why it is helpful to count samples longer when the number of counts is small. For example, Table A.3H.12 shows the confidence interval for 100 cpm counted for various times. When longer times are used, the confidence interval is narrower. Table A.3H.12
Determination of Confidence Values
Counts per min (cpm) Total counts 95% CI of counts 95% CI of cpm
1 min
10 min
100 min
100 100 81.4 to 121.6 81.4 to 121.6
100 1000 938 to 1062 93.8 to 106.2
100 10000 9804 to 10196 98.0 to 102.0
Figure A.3H.28 shows percent error as a function of C. Percent error is defined from the width of the confidence interval divided by the number of counts. Of course this graph only shows error due to the randomness of radioactive decay. This is only one source of error in most experiments.
Commonly Used Techniques
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Percent error = 100 x 1.96 √ C C
Percent error
100.0
10.0
1.0
0.1 102
103
104
105
106
Counts
Figure A.3H.28 Counting error. With more counts, the fractional counting error decreases. The x axis shows the number of radioactive decays actually counted (counts per minute times number of minutes).
ANALYZING DATA WITH GRAPHPAD PRISM GraphPad Prism is a general purpose program for scientific graphics, statistics and nonlinear regression, available for both Windows and Macintosh computers. While Prism is not designed especially for analyses of binding data, it is very well suited for analyses. It provides a menu of commonly used equations, including all equations listed in this unit, and can automatically compare one- and two-site models with an F test. When analyzing competitive binding curves, Prism calculates the Ki from the IC50. The program can automatically create a residual plot and calculate the runs test. In addition, Prism’s manual and help screens, like this unit, explain the principles of curve fitting. A trial version of the Windows or Mac versions of Prism can be obtained from the GraphPad web site at http://www.graphpad.com. The trial versions let you analyze data for an unlimited period of time. For the first thirty days, the Windows version is fully functional. After that, data may be analyzed, but the ability to to print, save, and export will be disabled. Contact GraphPad Software (SUPPLIERS APPENDIX) to obtain a brochure and trial disk, or to ask questions.
Analyzing Radioligand Binding Data
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LITERATURE CITED Cheng, Y. and Prusoff, W.H. 1973. Relationship between the inhibition constant (Ki) and the concentration of an inhibitor that causes a 50% inhibition (I50) of an enzymatic reaction. Biochem. Pharmacol. 22:3099-3108. Kenakin, T. 1993. Pharmacologic Analysis of DrugReceptor Interactions. Raven Press, New York. Limbird, L.E. 1996. Cell Surface Receptors: A Short Course in Theory and Methods, 2nd ed. Kluwer Academic Publishers, Boston.
Rosenthal, H.E. 1967. A graphic method for the determination and presentation of binding parameters in complex systems. Anal. Biochem. 20:525-532. Swillens, S. 1995. Interpretation of binding curves obtained with high receptor concentrations: Practical aid for computer analysis. Mol. Pharmacol. 47:1197-1203.
Motulsky, H.J. and Mahan, L.C. 1984. The kinetics of competitive radioligand binding predicted by the law of mass action. Mol. Pharmacol. 25:1-9.
Contributed by Harvey Motulsky GraphPad Software San Diego, California
Munson, P.J. and Rodbard, D. 1980. Ligand: A versatile computerized approach to characterization of ligand binding systems. Anal. Biochem. 107:220-239.
Richard Neubig University of Michigan Ann Arbor, Michigan
Commonly Used Techniques
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APPENDIX 4
Molecular Biology Techniques
Karen L. Elbing1 and roger Brent2 1 2
Clark and Elbing LLP, Boston, Massachusetts The Molecular Science Institute, Berkeley, California
John E. Coligan, Ben M. Dunn, David W. Speicher, and Paul T. Wingfield (eds.) Current Protocols in Protein Science Copyright © 2003 John Wiley & Sons, Inc. All rights reserved. DOI: 10.1002/0471140864.psa04s13 Online Posting Date: May, 2001 Print Publication Date: September, 1998
Some of the units in this manual (e.g., see Chapters 5.0 and 12.0) describe techniques for the cloning, expression, and structural analysis of genes and proteins. We assume that users of these protocols have at least some introductory background in recombinant DNA technology (or are working with a collaborator who does); therefore, we have not provided comprehensive coverage of all of the topics, but rather have concentrated on presenting selected techniques that will be of the most interest and use to the general proteins laboratory. More comprehensive coverage of these techniques can be found in Current Protocols in Molecular Biology or any of a variety of textbooks devoted to recombinant DNA techniques.
About Wiley InterScience | About Wiley | Privacy | Terms & Conditions Copyright© 1999-2005 John Wiley & Sons, Inc. All rights reserved.
2005-01-09
MOLECULAR BIOLOGY TECHNIQUES
APPENDIX 4
Some of the units in this manual (e.g., see Chapters 5 and 12) describe techniques for the cloning, expression, and structural analysis of genes and proteins. We assume that users of these protocols have at least some introductory background in recombinant DNA technology (or are working with a collaborator who does); therefore, we have not provided comprehensive coverage of all of the topics, but rather have concentrated on presenting selected techniques that will be of the most interest and use to the general proteins laboratory. More comprehensive coverage of these techniques can be found in Current Protocols in Molecular Biology or any of a variety of textbooks devoted to recombinant DNA techniques. KEY REFERENCES Ausubel, F.A., Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G., Smith, J.A., and Struhl, K. (eds.). 1998. Current Protocols in Molecular Biology. John Wiley and Sons, New York. Sambrook, J., Fritsch, E.F., and Maniatis, T. 1989. Molecular Cloning: A Laboratory, 2nd ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y.
Media Preparation and Bacteriological Tools
APPENDIX 4A
This appendix presents recipes for basic bacteriological media. The reader is also referred to UNITS 5.2 & 5.3 for media recipes optimized for high-level expression. MINIMAL MEDIA To prepare 5× media, add ingredients to water in a 2-liter flask and heat with stirring until dissolved. Pour into bottles with loosened caps and autoclave 15 min at 15 lb/in2. Cool media to <50°C before adding nutritional supplements and antibiotics (at this temperature the flask still feels hot but can be held continuously in one’s hand). Tighten caps and store concentrated media indefinitely at room temperature. 5× A medium, per liter 5 g (NH4)2SO4 22.5 g KH2PO4 52.5 g K2HPO4 2.5 g sodium citrate⋅2H2O 5× M9 medium, per liter 30 g Na2HPO4 15 g KH2PO4 5 g NH4Cl 2.5 g NaCl 15 mg CaCl2 (optional) 5× M63 medium, per liter 10 g (NH4)2SO4 68 g KH2PO4 2.5 mg FeSO4⋅7H2O Adjust to pH 7 with KOH Molecular Biology Techniques Contributed by Karen L. Elbing and Roger Brent Current Protocols in Protein Science (1998) A.4A.1-A.4A.5 Copyright © 1998 by John Wiley & Sons, Inc.
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Before use, dilute concentrated media to 1× with sterile water and add the following sterile solutions, per liter: 1 ml 1 M MgSO4⋅7H2O 10 ml 20% carbon source (sugar or glycerol) and, if required: 0.1 ml 0.5% vitamin B1 (thiamine) 5 ml 20% Casamino Acids (Difco) or L amino acids to 40 µg/ml or DL amino acids to 80 µg/ml Antibiotic RICH MEDIA Prepare as for minimal media except autoclave 25 min. Do not dilute. Tryptone, yeast extract, agar (Bacto-agar), nutrient broth, and Casamino Acids are from Difco. NZ Amine A (casein hydrolysate) is from ICN Biochemicals. H medium, per liter 10 g tryptone 8 g NaCl Lambda broth, per liter 10 g tryptone 2.5 g NaCl LB medium, per liter 10 g tryptone 5 g yeast extract 5 g NaCl 1 ml 1 N NaOH NZC broth, per liter 10 g NZ Amine A 5 g NaCl 2 g MgCl2⋅6H2O Autoclave 30 min 5 ml 20% Casamino Acids Superbroth, per liter 32 g tryptone 20 g yeast extract 5 g NaCl 5 ml 1 N NaOH TB (terrific broth) 12 g Bacto tryptone 24 g Bacto yeast extract 4 ml glycerol Add H2O to 900 ml and autoclave, then add to above sterile solution 100 ml of a sterile solution of 0.17 KH2PO4 and 0.72 M K2HPO4.
Media Presentation and Bacteriological Tools
Tryptone broth, per liter 10 g tryptone 5 g NaCl
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TY medium, 2×, per liter 16 g tryptone 10 g yeast extract 5 g NaCl TYGPN medium, per liter 20 g tryptone 10 g yeast extract 10 ml 80% glycerol 5 g Na2HPO4 10 g KNO3 SOLID MEDIA Prepare ingredients for minimal and rich plates as described below. Dry plates 2 to 3 days at room temperature, or 30 min with the lids slightly off at 37°C or in a laminar flow hood. Store dried plates wrapped at 4°C. Minimal Plates Autoclave 15 g agar in 800 ml water for 15 min. Add 200 ml sterile 5× minimal medium and carbon source. Cool to 50°C and add supplements and antibiotics. Pour 32 to 40 ml medium/plate to obtain 25 to 30 plates/liter. Rich Plates Add water to ingredients listed below to 1 liter. Autoclave 25 min. Pour 32 to 40 ml medium for LB and H plates, and 45 ml medium for lambda plates (per plate). H plates, per liter 10 g tryptone 8 g NaCl 15 g agar Lambda plates, per liter 10 g tryptone 2.5 g NaCl 10 g agar LB plates, per liter 10 g tryptone 5 g yeast extract 5 g NaCl 1 ml 1 N NaOH 15 g agar or agarose Additives Antibiotics (if required): Ampicillin to 50 µg/ml Tetracycline to 12 µg/ml Galactosides (if required): Xgal to 20 µg/ml IPTG to 0.1 mM Molecular Biology Techniques
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TOP AGAR Top agar is used to distribute phage or cells evenly in a thin layer over the surface of a plate. Prepare top agar in 1-liter batches, autoclave for 15 min to melt, cool to 50°C, swirl to mix, pour into separate 100-ml bottles, reautoclave, cool, and store at room temperature. Before use, melt the agar by heating in a water bath or microwave oven then cool to and hold at 45° to 50°C. Top agar contains less agar than plates and stays molten for days when kept at 45° to 50°C. H top agar, per liter 10 g tryptone 8 g NaCl 7 g agar Lambda top agar, per liter 10 g tryptone 2.5 g NaCl 7 g agar LB top agar, per liter 10 g tryptone 5 g yeast extract 5 g NaCl 7 g agar Top agarose, per liter 10 g tryptone 8 g NaCl 6 g agarose STAB AGAR Stab agar, per liter 10 g nutrient broth 10 mg cysteine⋅Cl 5 g NaCl 10 mg thymine 6 g agar Stab agar is used for storing bacterial strains. Cysteine is thought to increase the amount of time bacteria can survive in stabs. Thymine is included so that thy− bacteria can grow.
TOOLS Inoculating Loops 1. Sterilize the inoculating loop in a bunsen burner flame until it is red hot. 2. Cool by touching to the surface of a sterile agar plate until it stops sizzling. Sterile Toothpicks 1. Autoclave toothpicks in a beaker covered with foil or in the original box. Round wooden toothpicks or the pointed end of flat toothpicks are sometimes used to pick individual colonies or phage plaques. Media Presentation and Bacteriological Tools
2. Reautoclave used toothpicks to reuse.
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making a spreader
Figure A.4A.1
Making a spreader.
Spreaders 1. Make a spreader by heating and bending a piece of 4-mm glass tubing or a Pasteur pipet (see Fig. A.4A.1). 2. Sterilize the spreader by dipping the triangular part into a container of ethanol, removing it from the container, and igniting the ethanol on the spreader by passing it through a gas flame. 3. After the flame goes out, cool the spreader by touching it to the surface of a sterile agar plate.
Contributed by Karen L. Elbing Clark and Elbing LLP Boston, Massachusetts Roger Brent The Molecular Sciences Institute Berkeley, California
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Growth in Liquid or Solid Media
APPENDIX 4B
GROWING AN OVERNIGHT CULTURE 1. Transfer 5 ml liquid medium (APPENDIX 4A) into a sterile 16- or 18-mm culture tube. 2. Inoculate with a single bacterial colony on an inoculating loop (APPENDIX dipping and shaking the loop in the medium.
4A),
BASIC PROTOCOL 1
by
3. Cap the tube and grow at 37°C to saturation (∼6 hr) in a shaker or on a roller drum, 60 rpm. A freshly saturated culture can contain 1–2 × 109 cells/ml.
GROWING LARGER CULTURES 1. Dilute overnight cultures 1:100 in an Erlenmeyer or baffle flask that is ≥5 times the volume of the culture.
BASIC PROTOCOL 2
To grow cells without agitation, use a flask that is ≥20 times the volume of the culture.
2. Grow at 37°C with vigorous agitation, ∼300 rpm. MONITORING GROWTH With a Count Slide 1. Cover a clean count slide or Petraff-Hausser chamber with a clean cover slip. See Figure A.4B.1.
BASIC PROTOCOL 3
2. Touch a pipet containing a small amount of the culture to the edge of the coverslip. 3. Place on a phase-contrast microscope at 400× and calculate concentration based on each cell in a small square being approximately equivalent to 2 × 107 cells/ml. With a Spectrophotometer 1. Dilute culture to achieve an OD600 <1. 2. Calculate concentration based on 0.1 OD being roughly equivalent to 108 cells/ml.
Figure A.4B.1 Culture of E. coli at 2.4 × 108 cells/ml, viewed on a count slide. Molecular Biology Techniques Contributed by Karen L. Elbing and Roger Brent Current Protocols in Protein Science (1998) A.4B.1-A.4B.3 Copyright © 1998 by John Wiley & Sons, Inc.
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BASIC PROTOCOL 4
TITERING AND ISOLATING BACTERIAL COLONIES BY SERIAL DILUTIONS 1. Add 5 ml LB medium (APPENDIX 4A) to three sterile culture tubes and label 1 to 3. 2. Transfer 5 µl of cells to the first tube and vortex, transfer 5 µl from the first tube to the second tube and vortex, and transfer 5 µl from the second tube to the third tube and vortex. Use a new pipet tip with each transfer. Saturated cultures contain ∼109 cells/ml.
3. Spread 100 µl from the culture and from each dilution tube onto separate, labeled, dry LB plates (APPENDIX 4A), and incubate overnight at 37°C. 4. Calculate cells/ml: multiply number of colonies by 10 times the dilution factor. 5. Save for further use by wrapping plates and storing at 4°C. BASIC PROTOCOL 5
ISOLATING SINGLE COLONIES BY STREAKING A PLATE 1. Streak an inoculum across one side of a plate using sterile technique (see Fig. A.4B.2). 2. Resterilize an inoculating loop and streak a sample from the first streak across a fresh part of the plate. Repeat until the plate is covered. 3. Incubate at 37°C until colonies appear. If colonies must be isolated from many bacteria, divide plate into 4, 6, or 8 sectors and streak for single colonies in each sector.
Figure A.4B.2
BASIC PROTOCOL 6
Streaking a plate.
ISOLATING SINGLE COLONIES BY SPREADING A PLATE 1. Pipet 0.05 to 1 ml of the culture onto a dry plate. 2. Spread evenly with a spreader (APPENDIX 4A) using a circular motion. Alternatively, use the edge of the spreader to make a raster pattern (a series of parallel lines) on the plate’s surface; turn plate and repeat at right angle. 3. Incubate at 37°C with plate lid ajar until completely dry.
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REPLICA PLATING Replica plating is a convenient way to test many colonies for their ability to grow under different conditions. Bacterial colonies are transferred from one plate to another in a way that maintains the original pattern of colonies.
SUPPORT PROTOCOL 1
1. Secure a sterile velvet with a metal ring onto a replica block. 2. Press a master plate of well-separated colonies lightly onto the velvet. 3. Press new plates, oriented like the master plate, lightly onto imprinted velvet to transfer colonies. Up to 10 plates/velvet can be replica plated.
STRAIN STORAGE AND REVIVAL Stabs 1. Fill airtight, autoclavable vials with rubber or Teflon caps 2⁄3 full with stab agar.
SUPPORT PROTOCOL 2
2. Inoculate with a single colony, repeatedly poking the inoculating loop deeply into the agar. 3. Incubate at 37°C with the cap loose until cloudy tracks of bacteria become visible (8 to 12 hr). 4. Seal tightly and store in the dark at 15° to 22°C. 5. Revive as follows: using a cool, sterile inoculating loop, remove some bacteria-laden agar, then smear onto an LB plate (APPENDIX 4A) and streak for single colonies. Frozen Stocks 1. Add 1 ml of a freshly saturated culture (or 2 ml of a mid-log culture) to a stab vial or Nunc vial containing 1 ml of 65% (v/v) glycerol/0.1 M MgSO4/0.025 M Tris⋅Cl, pH 8.0, or 1 ml of 7% (v/v) dimethyl sulfoxide (DMSO). The only advantage DMSO seems to have over glycerol for frozen stocks is that it is easier to pipet because it is less viscous.
2. Store at −20° to −70°C. Most strains remain viable much longer when stored at −70°C.
3. Revive by scraping off splinters of solid ice with a sterile toothpick or pipet (being careful not to allow contents to thaw) and streaking onto an LB plate.
Contributed by Karen L. Elbing Clark and Elbing LLP Boston, Massachusetts Roger Brent The Molecular Sciences Institute Berkeley, California
Molecular Biology Techniques
A.4B.3 Current Protocols in Protein Science
Supplement 13
Preparation of Plasmid DNA ALKALINE LYSIS MINIPREP The alkaline lysis procedure is the most commonly used miniprep.
APPENDIX 4C BASIC PROTOCOL 1
Materials LB medium or enriched medium (e.g., superbroth or terrific broth; APPENDIX 4A) containing appropriate antibiotic Glucose/Tris/EDTA (GTE) solution: 50 mM glucose/25 mM Tris⋅Cl (pH 8.0; APPENDIX 2E)/10 mM EDTA (APPENDIX 2E) TE buffer (APPENDIX 2E) NaOH/SDS solution: 0.2 M NaOH/1% (w/v) SDS Potassium acetate solution, pH 4.8: 29.5 ml glacial acetic acid/KOH pellets to pH 4.8 (several)/H2O to 100 ml 95% and 70% ethanol 1. Inoculate 5 ml sterile LB medium with a single bacterial colony and grow to saturation (OD600 ∼4) at 37°C. 2. Microcentrifuge 1.5 ml of cells 20 sec. Resuspend pellet in 100 µl GTE solution and let sit 5 min at room temperature. 3. Add 200 µl NaOH/SDS solution, mix, and place 5 min on ice. 4. Add 150 µl potassium acetate solution, vortex 2 sec, and place 5 min on ice. 5. Microcentrifuge 3 min and transfer 0.4 ml supernatant to a new tube. Add 0.8 ml of 95% ethanol, and let sit 2 min at room temperature. 6. Microcentrifuge 3 min at room temperature, wash pellet with 1 ml of 70% ethanol, and dry under vacuum. 7. Resuspend pellet in 30 µl TE buffer and store as in Support Protocol. Use 2.5 to 5 µl for a restriction digest. Destroy contaminating RNA by adding 1 µl of a 10 mg/ml RNase solution (DNase-free).
ALKALINE LYSIS IN 96-WELL MICROTITER PLATES This procedure makes it possible to perform hundreds of rapid plasmid preps in a day.
ALTERNATE PROTOCOL
Additional Materials (also see Basic Protocol 1) TYGPN medium (APPENDIX 4A) Isopropanol 96-well microtiter plates (Dynatech PS plates or equivalent) Multichannel pipetting device (8-prong Costar; 12-prong Titer Tek) Multitube vortexer Sorvall RT-6000 low-speed centrifuge, or equivalent, with microplate carrier in H-1000B rotor 1. Add 0.3 ml TYGPN medium to each well of a 96-well microtiter plate. Inoculate each with a single plasmid-containing colony and grow to saturation at 37°C (∼48 hr). 2. Centrifuge plates 10 min at 600 × g, 4°C. Flick supernatant from wells.
Contributed by JoAnne Engebrecht and Roger Brent Current Protocols in Protein Science (1998) A.4C.1-A.4C.3 Copyright © 1998 by John Wiley & Sons, Inc.
Molecular Biology Techniques
A.4C.1 Supplement 13
3. Resuspend cells by clamping plate in a multitube vortexer and running it 20 sec at setting 4. 4. Add 50 µl GTE solution and 100 µl NaOH/SDS solution to each well. Wait 2 min and add 50 µl potassium acetate solution to each well. 5. Cover with plate tape, vortex 20 sec at setting 4, and centrifuge 5 min at 600 × g, 4°C. 6. Remove 200 µl from each well and transfer to new plate. Add 150 µl isopropanol to each well, cover, agitate, and chill 30 min at −20°C. 7. Centrifuge 25 min at 600 × g, 4°C. Wash pellets with cold 70% ethanol, gently decant supernatant, wash with 95% ethanol, and again gently decant supernatant. 8. Air dry pellets 30 min, and resuspend in 50 µl TE buffer. Store as in Support Protocol and use 10-µl aliquots for digestion. BASIC PROTOCOL 2
BOILING MINIPREP This procedure is recommended for preparing small amounts of plasmid DNA from 1 to 24 cultures. It is extremely quick, but the quality of DNA produced is lower than that from the alkaline lysis miniprep. Materials LB medium (APPENDIX 4A) containing appropriate antibiotic STET solution: 8% (w/v) sucrose/0.5% (v/v) Triton X-100/50 mM EDTA (APPENDIX 2E)/50 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) Hen egg white lysozyme Isopropanol, ice-cold TE buffer (APPENDIX 2E) 1. Inoculate 5 ml LB medium with a single bacterial colony and grow to saturation at 37°C. 2. Microcentrifuge 1.5 ml of culture 20 sec. Resuspend bacteria by vortexing in 300 µl of STET solution containing 200 µg lysozyme. Place tube 30 sec to 10 min on ice. Be sure cells are completely resuspended to achieve high yields of DNA.
3. Place tube in boiling water bath (100°C) 1 to 2 min, microcentrifuge 15 to 30 min, and pipet supernatant to a new tube. 4. Mix with 200 µl cold isopropanol and place 15 to 30 min at −20°C. 5. Microcentrifuge 5 min. Remove supernatant and dry under a vacuum. 6. Resuspend in 50 µl TE buffer and store as in the support protocol. Use 5 µl for a restriction digest. Destroy contaminating RNA by adding 1 µl of a 10 mg/ml RNase solution (DNase-free) to the digestion mixture.
Preparation of Plasmid DNA
A.4C.2 Supplement 13
Current Protocols in Protein Science
STORAGE OF PLASMID DNA 1. Store plasmids in bacterial strains on selective plates at 4°C for the short-term, and in a glycerol- or DMSO-based solution (APPENDIX 4B) at −70°C for the long-term. Grow cells taken from storage on selective plates (APPENDIX 4A) and check plasmid DNA by restriction analysis (APPENDIX 4I).
SUPPORT PROTOCOL
2. Store plasmid DNA in TE buffer (APPENDIX 2E) at 4°C for short-term storage, and at −20° or −70°C for long-term storage.
Contributed by JoAnne Engebrecht State University of New York Stony Brook, New York Roger Brent The Molecular Sciences Institute Berkeley, California
Molecular Biology Techniques
A.4C.3 Current Protocols in Protein Science
Supplement 13
Introduction of Plasmid DNA into Cells TRANSFORMATION USING CALCIUM CHLORIDE Materials Single colony of E. coli cells LB medium (APPENDIX 4A) CaCl2 solution, ice-cold LB plates containing ampicillin (APPENDIX 4A) Plasmid DNA (APPENDIX 4C) Beckman JS-5.2 rotor or equivalent
APPENDIX 4D BASIC PROTOCOL 1
NOTE: All materials and reagents coming into contact with bacteria must be sterile. 1. Inoculate a single colony of E. coli cells into 50 ml LB medium. Grow overnight at 37°C with shaking (250 rpm; APPENDIX 4B). 2. Inoculate 4 ml of the culture into 400 ml LB medium in a 2-liter flask. Grow at 37°C, with shaking (250 rpm), to an OD590 of 0.375. This procedure requires that cells be growing rapidly (early- or mid-log phase).
3. Aliquot culture into eight 50-ml prechilled, sterile polypropylene tubes and leave the tubes on ice 5 to 10 min. Centrifuge cells 7 min at 1600 × g, 4°C. 4. Resuspend (gently) each pellet in 10 ml ice-cold CaCl2 solution. Centrifuge 5 min at 1100 × g, 4°C. 5. Resuspend each pellet in 10 ml cold CaCl2 solution. Keep resuspended cells on ice for 30 min. Centrifuge 5 min at 1100 × g, 4°C. 6. Resuspend each pellet completely in 2 ml of ice-cold CaCl2 solution. Aliquot 250 µl into prechilled, sterile polypropylene tubes. Freeze immediately at −70°C. 7. Use 10 ng of pBR322 to transform 100 µl of competent cells using the steps below. Plate appropriate aliquots (1, 10, and 25 µl) of transformation culture on LB/ampicillin plates and incubate at 37°C overnight. The number of transformant colonies per aliquot volume (µl) × 105 is equal to the number of transformants per microgram of DNA.
8. Add 10 ng of DNA (10 to 25 µl) into a 15-ml sterile, round-bottom test tube and place on ice. Rapidly thaw competent cells by warming between hands, dispense 100 µl immediately into test tubes containing DNA, swirl, and place on ice 10 min. 9. Heat shock cells by placing tubes 2 min in 42°C water bath. Add 1 ml LB medium to each tube. Incubate 1 hr at 37°C on roller drum (250 rpm). 10. Plate several dilutions on appropriate antibiotic-containing plates, and incubate 12 to 16 hr at 37°C. Store remainder at 4°C for subsequent platings. Transformation efficiencies range from 106 to 108 transformants/µg DNA.
Molecular Biology Techniques Contributed by Christine E. Seidman and Roger Brent Current Protocols in Protein Science (1998) A.4D.1-A.4D.2 Copyright © 1998 by John Wiley & Sons, Inc.
A.4D.1 Supplement 13
ALTERNATE PROTOCOL
ONE-STEP PREPARATION AND TRANSFORMATION OF COMPETENT CELLS Various strains can be made competent by this procedure and the frequency can be as high as that achieved by the basic protocol. However, frequency is considerably lower than can be obtained by electroporation (UNIT 5.10). Additional Materials (also see Basic Protocol 1) 2× transformation and storage solution (TSS), ice-cold: dilute sterile 40% (w/v) polyethylene glycol (PEG) 3350 to 20% in sterile LB medium (APPENDIX 4A) containing 100 mM MgCl2. Add DMSO to 10% and adjust to pH 6.5. LB medium (APPENDIX 4A) containing 20 mM glucose 1. Dilute fresh overnight culture of bacteria 1:100 into LB medium and grow at 37°C to an OD600 of 0.3 to 0.4. 2. Add a volume of ice-cold 2× TSS equal to the cell suspension, and gently mix on ice. For long-term storage, freeze small aliquots of the suspension in a dry ice/ethanol bath and store at −70°C. To use frozen cells for transformation, thaw slowly, then use immediately.
3. Add 100 µl competent cells and 1 to 5 µl DNA (0.1 to 100 ng) to an ice cold polypropylene or glass tube. Incubate 5 to 60 min at 4°C. 4. Add 0.9 ml LB medium/20 mM glucose and incubate 30 to 60 min at 37°C with mild shaking. Select transformants on appropriate plates. The expected transformation frequency is 107 and 108 colonies/µg DNA.
LITERATURE CITED Dower, W.J., Miller, J.F., and Ragdale, C.W. 1988. High efficiency transformation of E. coli by high voltage electroporation. Nucl. Acids Rs. 16:6127-6145. Hanahan, D. 1983. Studies on transformation of Escherichi coli with plasmids. J. Mol. Biol. 166:557-580.
Contributed by Christine E. Seidman and Kevin Struhl Harvard Medical School Boston, Massachusetts
Introduction of Plasmid DNA into Cells
A.4D.2 Supplement 13
Current Protocols in Protein Science
Purification and Concentration of DNA from Aqueous Solutions
APPENDIX 4E
This unit presents basic procedures for manipulating solutions of single- or doublestranded DNA through purification and concentration steps. IMPORTANT NOTE: The smallest amount of contamination of DNA preparations by recombinant phages or plasmids can be disastrous. All materials used for preparation of plasmid or phage DNA should be kept separate from those used for preparation of genomic DNA, and disposable items should be used wherever possible. Particular care should be taken to avoid contamination of commonly used rotors. PHENOL EXTRACTION AND ETHANOL PRECIPITATION OF DNA This protocol describes the most commonly used method of purifying and concentrating DNA preparations using phenol extraction and ethanol precipitation; it is appropriate for the purification of DNA from small volumes (<0.4 ml) at concentrations ≤1 mg/ml.
BASIC PROTOCOL
Materials DNA to be purified (≤1 mg/ml) in 0.1 to 0.4 ml volume 25:24:1 (v/v/v) phenol/chloroform/isoamyl alcohol (made with buffered phenol; APPENDIX 2E) 3 M sodium acetate, pH 5.2 (APPENDIX 2E) 100% ethanol, ice cold 70% ethanol, room temperature TE buffer, pH 8.0 (APPENDIX 2E) Speedvac evaporator (Savant) 1. Add an equal volume of phenol/chloroform/isoamyl alcohol to the DNA solution to be purified in a 1.5-ml microcentrifuge tube. 2. Vortex vigorously 10 sec and microcentrifuge 15 sec at room temperature. 3. Carefully remove the top (aqueous) phase containing the DNA using a 200-µl pipettor and transfer to a new tube. If a white precipitate is present at the aqueous/organic interface, reextract the organic phase and pool aqueous phases. 4. Add 1⁄10 vol of 3 M sodium acetate, pH 5.2, to the solution of DNA. Mix by vortexing briefly or by flicking the tube several times with a finger. 5. Add 2 to 2.5 vol (calculated after salt addition) of ice-cold 100% ethanol. Mix by vortexing and place in crushed dry ice for 5 min or longer. 6. Spin 5 min in a fixed-angle microcentrifuge at high speed and remove the supernatant. 7. Add 1 ml of room-temperature 70% ethanol. Invert the tube several times and microcentrifuge as in step 6. 8. Remove the supernatant. Dry the pellet in a desiccator under vacuum or in a Speedvac evaporator. 9. Dissolve the dry pellet in an appropriate volume of water or TE buffer, pH 8.0.
Molecular Biology Techniques Contributed by David Moore Current Protocols in Protein Science (1998) A.4E.1-A.4E.2 Copyright © 1998 by John Wiley & Sons, Inc.
A.4E.1 Supplement 13
ALTERNATE PROTOCOL
SUPPORT PROTOCOL
PRECIPITATION OF DNA USING ISOPROPANOL Isopropanol may also be used to precipitate DNA. Equal volumes of isopropanol and DNA solution are used, allowing precipitation from a large starting volume (e.g., 0.7 ml) in a single microcentrifuge tube. CONCENTRATION OF DNA USING BUTANOL It is generally inconvenient to handle large volumes or dilute solutions of DNA. Water molecules (but not DNA or solute molecules) can be removed from aqueous solutions by extraction with sec-butanol (2-butanol) to reduce volumes or concentrate dilute solutions. Additional Materials (also see Basic Protocol) sec-butanol 25:24:1 phenol/chloroform/isoamyl alcohol (made with buffered phenol; APPENDIX 2E) Polypropylene tube 1. Add an equal volume of sec-butanol to the sample and mix well by vortexing or by gentle inversion if the DNA is of high molecular weight. Perform extraction in a polypropylene tube, as butanol will damage polystyrene. 2. Centrifuge 5 min at 1200 × g, room temperature, or 10 sec in a microcentrifuge. 3. Remove and discard the upper (sec-butanol) phase. 4. Repeat steps 1 to 3 until the desired volume of aqueous solution is obtained. 5. Extract the lower, aqueous phase with 25:24:1 phenol/chloroform/isoamyl alcohol and ethanol precipitate. LITERATURE CITED Marmur, J. 1961. A procedure for the isolation of desoxyribonucleic acids from microorganisms. J. Mol. Biol. 3:208-218.
Contributed by David Moore Baylor College of Medicine Houston, Texas
Purification and Concentration of DNA from Aqueous Solutions
A.4E.2 Supplement 13
Current Protocols in Protein Science
Agarose Gel Electrophoresis
APPENDIX 4F
RESOLUTION OF LARGE DNA FRAGMENTS ON AGAROSE GELS This protocol is used to separate and purify DNA fragments between 0.5 and 25 kb.
BASIC PROTOCOL
Materials Electrophoresis buffer (TAE or TBE; APPENDIX 2E) Ethidium bromide solution (APPENDIX 2E) Agarose, electrophoresis-grade 10× loading buffer: 20% (w/v) Ficoll 400/0.1 M disodium EDTA, pH 8.0/1% (w/v) SDS/0.25% (w/v) bromphenol blue DNA molecular weight markers (Fig. A.4F.1) Horizontal gel electrophoresis apparatus Gel casting platform Gel combs (slot formers) DC power supply 1. Prepare the gel, using electrophoresis buffer and electrophoresis-grade agarose (see Table A.4F.1) by melting in a microwave oven or autoclave, mixing, cooling to 55°C, pouring into a sealed gel casting platform, and inserting the gel comb. Ethidium bromide can be added to the gel and electrophoresis buffer at 0.5 µg/ml. CAUTION: Ethidium bromide is a potential carcinogen. Wear gloves when handling.
Lambda Bst EII (kb)
Lambda Hin dIII (kb)
8.45 7.24 6.37 5.69 4.82 4.32 3.68 2.32 1.93 1.37 1.26
pBR322 Bst NI (kb)
23.13 9.42 6.56
4.36
2.32 2.03
1.86 1.06 .93
.70 .56
.38
.22 .12
.13
agarose concentration
applied voltage
1%
1V/cm
.12
buffer gel run
TAE 16 hr
Figure A.4F.1 Migration pattern and fragment sizes for commonly used DNA molecular weight markers. Molecular Biology Techniques Contributed by Daniel Voytas Current Protocols in Protein Science (1998) A.4F.1-A.4F.3 Copyright © 1998 by John Wiley & Sons, Inc.
A.4F.1 Supplement 13
Table A.4F.1 Appropriate Agarose Concentrations for Separating DNA Fragments of Various Sizes
Agarose (%) 0.5 0.7 1.0 1.2 1.5
Effective range of resolution of linear DNA fragments (kb) 30 to 1 12 to 0.8 10 to 0.5 7 to 0.4 3 to 0.2
2. After the gel has hardened, remove the seal from the gel casting platform and withdraw the gel comb. Place into an electrophoresis tank containing sufficient electrophoresis buffer to cover the gel ∼1 mm. 3. Prepare DNA samples with an appropriate amount of 10× loading buffer and load samples into wells with a pipettor. Be sure to include appropriate DNA molecular weight markers (Fig. A.4F.1). 4. Attach the leads so that the DNA migrates to the anode or positive lead and electrophorese at 1 to 10 V/cm of gel. 5. Turn off the power supply when the bromphenol blue dye from the loading buffer has migrated a distance judged sufficient for separation of the DNA fragments. Bromphenol blue comigrates with ∼0.5-kb fragments.
6. Photograph a stained gel directly on a UV transilluminator or first stain with 0.5 µg/ml ethidium bromide 10 to 30 min, destaining 30 min in water, if necessary. SUPPORT PROTOCOL
MINIGELS AND MIDIGELS Small gels—minigels and midigels—can generally be run faster than larger gels and are often employed to expedite analytical applications. Because they use narrower wells and thinner gels, minigels and midigels also require smaller amounts of DNA for visualization of the separated fragments. Aside from a scaling down of buffer and gel volumes, the protocol for running minigels or midigels is similar to that for larger gels. Similarly, the parameters affecting the mobility of DNA fragments are the same for both large and small gels. When selecting a mini- or midigel apparatus consider the volume of buffer held by the gel tank. Smaller gels are typically run at high voltages (>10 V/cm) so electrophoresis buffers are quickly exhausted; therefore, it is advantageous to choose a gel apparatus with a relatively large buffer reservoir. Minigel boxes can also be easily constructed from a few simple materials (Sambrook et al., 1989). A small (e.g., 15 cm long × 8 cm wide × 4 cm high) plastic box can be equipped with male connectors and platinum wire electrodes at both ends to serve as a minimal gel tank.
Purification and Concentration of DNA from Aqueous Solutions
Although trays for casting small gels are commercially available, gels can also be poured onto glass lantern slides or other small supports without side walls. Such gels are held on the support simply by surface tension. After pouring the gel (e.g., 10 ml for a 5 cm × 8 cm gel), the comb is placed directly onto the support and held up by metal paper-binding clamps placed to the side. There is no agarose at the bottom of such wells, so extra care must be taken to prevent separation of the gel from the support when removing the comb.
A.4F.2 Supplement 13
Current Protocols in Protein Science
LITERATURE CITED Sambrook, J., Fritsch, E.F., and Maniatis, T. 1989. Molecular Cloning: A Laboratory Manual, 2nd ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. Southern, E. 1979. Gel electrophoresis of restriction fragments. Methods Enzymol. 68:152-176.
Contributed by Daniel Voytas Iowa State University Ames, Iowa
Molecular Biology Techniques
A.4F.3 Current Protocols in Protein Science
Supplement 13
Southern Blotting
APPENDIX 4G
Southern blotting is the transfer of DNA fragments from an electrophoresis gel to a membrane support resulting in immobilization of the DNA fragments, so the membrane carries a semipermanent reproduction of the banding pattern of the gel. SOUTHERN BLOTTING ONTO A NYLON OR NITROCELLULOSE MEMBRANE WITH HIGH-SALT BUFFER The procedure is specifically designed for blotting an agarose gel onto an uncharged or positively charged nylon membrane. With minor modifications, the same protocol can also be used with nitrocellulose membranes.
BASIC PROTOCOL 1
Materials 0.25 M HCl (APPENDIX 2E) Denaturation solution: 1.5 M NaCl/0.5 M NaOH (store at room temperature) Neutralization solution: 1.5 M NaCl/0.5 M Tris⋅Cl, pH 7.0 (store at room temperature) 20× and 2× SSC (APPENDIX 2E) Nylon or nitrocellulose membrane (see Table A.4G.1 for suppliers) UV transilluminator or UV light box (e.g., Stratagene Stratalinker) for nylon membranes CAUTION: Wear gloves from step 2 of the protocol onward to protect your hands from the acid and alkali solutions and to protect the membrane from contamination. Handle the membrane with blunt-end forceps. 1. Digest DNA with appropriate restriction endonuclease(s), run in an agarose gel with appropriate DNA size markers, stain with ethidium bromide, and photograph with a ruler laid alongside the gel so that band positions can later be identified on the membrane. The gel should contain the minimum agarose concentration needed to resolve the bands and should be ≤7 mm thick.
2. Treat the gel as follows, with gentle shaking at room temperature. Be sure the gel is covered completely. Distilled water ∼10 gel volumes 0.25 M HCl—30 min Distilled water ∼10 volumes denaturation solution—20 min, twice Distilled water ∼10 volumes neutralization solution—20 min, twice 3. Using Figure A.4G.1 as a guide, assemble the transfer stack consisting of a sponge or solid support with a Whatman 3MM paper wick in a dish filled with enough 20× SSC to half submerge the sponge, three pieces of Whatman 3MM paper, the gel, pieces of plastic wrap on the edges of the gel, a nylon membrane equilibrated in distilled water (or nitrocellulose membrane wet in distilled water and equilibrated 10 min in 20× SSC), five pieces of Whatman 3MM paper, and a 4-cm stack of paper towels. As each layer is applied, wet it with 20× SCC and remove any trapped air by carefully rolling a 10-ml glass pipet over the surface. Before a sponge is used for the first time, it should be washed thoroughly with distilled water to remove any detergents that may be present. Contributed by Terry Brown Current Protocols in Protein Science (1998) A.4G.1-A.4G.8 Copyright © 1998 by John Wiley & Sons, Inc.
Molecular Biology Techniques
A.4G.1 Supplement 13
Table A.4G.1
Properties of Materials used for Immobilization of Nucleic Acidsa
Nitrocellulose
Supported nitrocellulose
Uncharged nylon
Positively charged nylon
Activated papers
Application
ssDNA, RNA, protein
ssDNA, RNA, protein
ssDNA, dsDNA, RNA, protein
ssDNA, dsDNA, RNA, protein
ssDNA, RNA
Binding capacity (µg nucleic acid/cm2)
80-100
80-100
400-600
400-600
2-40
Tensile strength
Poor
Good
Good
Good
Good
Mode of nucleic acid attachmentb
Noncovalent
Noncovalent
Covalent
Covalent
Covalent
Lower size limit for efficient nucleic acid retention
500 nt
500 nt
50 nt or bp
50 nt or bp
5 nt
Suitability for reprobing
Poor (fragile)
Poor (loss of signal) Good
Good
Good
Commercial examples
Schleicher & Schuell BA83, BA85; Amersham Hybond-C; PALL Biodyne A
Schleicher & Schuell BA-S; Amersham HybondC extra
aThis
Amersham Hybond-N; Stratagene DuralonUV; Du Pont NEN GeneScreen
Schleicher & Schleicher & Schuell Nytran; Schuell APT Amersham papers Hybond-N+; Bio-Rad ZetaProbe; PALL Biodyne B; Du Pont NEN GeneScreen Plus
table is based on Brown (1991), with permission from BIOS Scientific Publishers Ltd.
bAfter
suitable immobilization procedure (see text).
For blots of substantial amounts of plasmid or other very-low-complexity DNA, it is important to lay the membrane down precisely the first time.
4. Lay a glass plate on top of the stack and place a 0.2 to 0.4-kg weight on top to hold everything in place. Leave overnight. 5. Recover the membrane. Mark in pencil the position of the wells on the membrane and cut one corner to mark the orientation. 6. Rinse the membrane in 2× SSC, then place it on a sheet of Whatman 3MM paper and allow to dry completely. 7. Wrap nylon membrane in UV-transparent plastic wrap, place DNA-side-down on a UV transilluminator (254-nm wavelength) and irradiate for the time determined from the Support Protocol. Nitrocellulose membranes should be placed between two sheets of Whatman 3MM paper and baked under vacuum for 2 hr at 80°C.
8. Store membranes dry between sheets of Whatman 3MM paper for several months at room temperature.
Southern Blotting
A.4G.2 Supplement 13
Current Protocols in Protein Science
0.2–0.4-kg weight glass plate
A
paper towels
membrane gel
transfer buffer
Whatman 3MM Whatman 3MM
reservoir
sponge
*
0.2–0.4-kg weight glass plate
paper towels Whatman 3MM
membrane gel plastic wrap
support
Whatman 3MM wick transfer buffer
reservoir
Figure A.4G.1 Transfer stacks for upward capillary transfer. (A) Sponge method. (B) Filter paper wick method.
CALIBRATION OF A UV TRANSILLUMINATOR UV transilluminators must be calibrated to determine their output to ensure the proper intensity of radiation is used for cross-linking.
SUPPORT PROTOCOL
CAUTION: Exposure to UV irradiation is harmful to the eyes and skin. Wear suitable eye protection and avoid exposure of bare skin. 1. Prepare five identical series of DNA dot blots (see Basic Protocol 2) on nylon membrane strips, each with a range of DNA quantity from 1 to 100 pg. 2. Dry the nylon strips, wrap each one in UV-transparent plastic wrap, and place on the UV transilluminator, DNA-side-down. 3. Switch on the transilluminator. Remove individual strips after 30 sec, 45 sec, 1 min, 2 min, and 5 min. 4. Hybridize the strips with a suitable DNA probe labeled to a specific activity of 108 dpm/µg and prepare autoradiograpms (APPENDIX 4H). 5. Determine which nylon strip gives the most intense hybridization signals; the exposure time used for that strip is the optimal exposure time.
Molecular Biology Techniques
A.4G.3 Current Protocols in Protein Science
Supplement 13
ALTERNATE PROTOCOL 1
SOUTHERN BLOTTING ONTO A NYLON MEMBRANE WITH AN ALKALINE BUFFER With a positively charged nylon membrane, the transferred DNA becomes covalently linked to the membrane if an alkaline transfer buffer is used. Additional Materials (also see Basic Protocol 1) 0.4 M NaOH (for charged membrane) or 0.25 M NaOH (for uncharged membrane) 0.25 M NaOH/1.5 M NaCl for uncharged membrane Positively charged or uncharged nylon membrane (see Table A.4G.1 for suppliers) 1. Prepare a gel and treat with 0.25 M HCl as described in steps 1 and 2 of the Basic Protocol. 2. Rinse the gel with distilled water and denature it with 10 gel volumes of 0.4 M NaOH. Shake slowly for 20 min. Use 0.25 M NaOH to denature an uncharged nylon membrane.
3. Carry out the transfer, using 0.4 M NaOH as the transfer solution in place of 20× SSC. Leave to transfer ≥2 hr. Use 0.25 M NaOH/1.5 M NaCl as the transfer solution for an uncharged nylon membrane.
4. Recover the membrane and rinse in 2× SSC. Place membrane on a sheet of Whatman 3MM filter paper, and allow to air dry. Store at room temperature. Baking or UV cross-linking is not necessary with positively charged membranes; however, DNA on uncharged membranes should be cross-linked by UV exposure. ALTERNATE PROTOCOL 2
SOUTHERN BLOTTING BY DOWNWARD CAPILLARY TRANSFER One disadvantage with the upward capillary method (see Basic Protocol 1) is that the gel can become crushed by the weighted filter papers and paper towels that are laid on top of it, reducing capillary action. Simple downward capillary transfer that does not cause excessive pressure to be placed on the gel is described in this protocol. 1. Prepare a gel as described in steps 1 to 4 of the Basic Protocol (high-salt transfer) or steps 1 to 2 of Alternate Protocol 1 (alkaline transfer). 2. Using Figure A.4G.2 as a guide, assemble a transfer pyramid consisting of paper towels (2 to 3 cm high) in a glass dish, four pieces of Whatman 3MM filter paper, a fifth filter paper wet with transfer buffer, the membrane wet with distilled water (nylon) or 20× SSC (nitrocellulose), a plastic wrap skirt, the gel, three pieces of Whatman 3MM paper, and two larger pieces of Whatman 3MM paper prewet with transfer buffer and extending from the stack to the transfer buffer reservoir. 3. Place a light plastic cover (e.g., a gel plate) over the top of the stack to reduce evaporation. Let stand 1 hr. 4. Recover the membrane, rinse in 2× SSC, and air dry. Immobilize the DNA and store the membrane at room temperature.
Southern Blotting
A.4G.4 Supplement 13
Current Protocols in Protein Science
glass plate
Whatman 3MM wick
Whatman 3MM gel membrane Whatman 3MM
paper towels (2-3 cm)
reservoir with transfer buffer
support
Figure A.4G.2 Transfer pyramid for downward capillary transfer. Adapted with permission from Academic Press.
ELECTROBLOTTING FROM A POLYACRYLAMIDE GEL TO A NYLON MEMBRANE
ALTERNATE PROTOCOL 3
Capillary transfer does not work with polyacrylamide gels, whose pore sizes are too small for effective transverse diffusion of DNA. Polyacrylamide gels must therefore be blotted by electrophoretic transfer in a buffer of low ionic strength. Additional Materials (also see Basic Protocol 1) 0.5× TBE electrophoresis buffer (APPENDIX 2E) Scotch-Brite pads (supplied with Trans-Blot apparatus) Trans-Blot electroblotting cell (Bio-Rad) with cooling coil, or other electroblotting apparatus (UNIT 10.7) 1. Run DNA samples in a nondenaturing or denaturing polyacrylamide gel. 2. When electrophoresis is almost complete, float a piece of nylon membrane sufficient in size to cover the relevant parts of the gel on the surface of distilled water, then submerge it and let stand 5 min. 3. Remove one glass plate from the electrophoresis apparatus and stain and photograph the gel (if nondenaturing). Lay a piece of Whatman 3MM filter paper on the surface of the gel, removing any trapped air bubbles. Lift the gel off the glass plate by peeling the filter paper away. 4. Soak two Scotch-Brite pads in 0.5× TBE electrophoresis buffer and remove air pockets by repeated squeezing and agitation. Cut seven pieces of Whatman 3MM paper to the same size as the gel and soak them 15 to 30 min in 0.5× TBE. 5. Place one saturated Scotch-Brite pad on the inner surface of the gray panel of the gel holder. Place three soaked filter papers on the pad, one at a time, removing trapped air bubbles. 6. Flood the filter paper carrying the gel with 0.5× TBE and place on top of the filter-paper stack. Flood the surface of the gel with 0.5× TBE and place the prewetted membrane onto the gel. 7. Flood the surface of the membrane with 0.5× TBE and place the remaining four sheets of saturated Whatman 3MM paper on top, followed by the second saturated ScotchBrite pad. Close the gel holder. 8. Half-fill the Trans-Blot cell with 0.5× TBE and place the gel holder in the cell with the grey panel facing towards the cathode. Fill the cell with 0.5× TBE and electroblot
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at 30 V (∼125 mA), with cooling, for 4 hr under the conditions recommended by the manufacturer. 9. Recover the membrane and mark the orientation of the membrane in pencil or by cutting a corner. 10. Denature the membrane from a nondenaturing gel by placing it, DNA-side-up, 10 min on three pieces of Whatman 3MM paper soaked in 0.4 M NaOH. 11. Rinse the membrane in 2× SSC, place on a sheet of Whatman 3MM paper, and allow to dry. Immobilize the DNA and store membrane at room temperature. BASIC PROTOCOL 2
DOT AND SLOT BLOTTING OF DNA ONTO UNCHARGED NYLON AND NITROCELLULOSE MEMBRANES USING A MANIFOLD Dot and slot blotting (Fig. A.4G.3) are simple techniques for immobilizing bulk unfractionated DNA on a nitrocellulose or nylon membrane for hybridization analysis (APPENDIX 4H) to determine the relative abundance of target sequences in the blotted DNA preparations. Dot and slot blots are usually prepared with the aid of a manifold and suction device. CAUTION: Wear gloves to protect your hands from the alkali solution and to protect the membrane from contamination. Avoid handling nitrocellulose and nylon membranes even with gloved hands—use clean blunt-ended forceps instead. Materials 6× and 20× SSC (APPENDIX 2E) Denaturation solution: 1.5 M NaCl/0.5 M NaOH (store at room temperature) Neutralization solution: 1 M NaCl/0.5 M Tris⋅Cl, pH 7.0 (store at room temperature) Uncharged nylon or nitrocellulose membrane (see Table A.4G.1) Dot/slot blotting manifold (e.g., Bio-Rad Bio-Dot SF or Schleicher and Schuell Minifold II) UV transilluminator for nylon membranes 1. Cut a piece of nylon membrane to the size of the manifold. Place the membrane on the surface of 6× SSC and allow to submerge. Let stand 10 min. Wet nitrocellulose membrane in 20× SSC.
2. Cut a piece of Whatman 3MM filter paper to the size of the manifold. Wet in 6× SSC. Use 20× SSC if transferring onto nitrocellulose.
Figure A.4G.3 Dot (left) and slot (right) blot manifold architectures.
dots
slots
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3. Place the Whatman 3MM paper in the manifold and lay the membrane on top of it. Assemble the manifold according to the manufacturer’s instructions, ensuring that there are no air leaks in the assembly. 4. Add 20× SSC and water to DNA to give a final concentration of 6× SSC in a volume of 200 to 400 µl. Denature 10 min at 100°C, then place in ice. For nitrocellulose membrane, add an equal volume of 20× SSC to each sample after placing in ice. The amount of DNA that should be blotted will depend on the relative abundance of the target sequence that will subsequently be sought by hybridization probing.
5. Switch on the suction to the manifold device and apply 500 µl of 6× SSC to each well. For a nitrocellulose membrane, use 20× SSC. Wells that are not being used can be blocked off by placing masking tape over them,
6. Microcentrifuge DNA samples 5 sec. Apply samples to the wells being careful to avoid touching the membrane with the pipet. Allow the samples to filter through. 7. Dismantle the apparatus and place the membrane on a piece of Whatman 3MM paper soaked in denaturation solution. Leave for 10 min. 8. Transfer the membrane to a piece of Whatman 3MM paper soaked in neutralization solution. Leave for 5 min. 9. Place the membrane on a piece of dry Whatman 3MM paper and allow to dry. 10. Wrap the dry membrane in UV-transparent plastic wrap, place DNA-side-down on a UV transilluminator, and immobilize the DNA by irradiating for the appropriate time. Nitrocellulose membranes should be placed between two sheets of Whatman 3MM paper and baked under vacuum for 2 hr at 80°C.
11. Store the membrane dry between sheets of Whatman 3MM filter paper for several months at room temperature. DOT AND SLOT BLOTTING OF DNA ONTO A POSITIVELY CHARGED NYLON MEMBRANE USING A MANIFOLD
ALTERNATE PROTOCOL 4
Positively charged nylon membranes bind DNA covalently at high pH. Samples for dot or slot blotting can therefore be applied in an alkaline buffer, which promotes both denaturation of the DNA and binding to the membrane. Additional Materials (also see Basic Protocol 2) Positively charged nylon membrane (see Table A.4G.1) 0.4 M and 1 M NaOH (APPENDIX 2E) 200 mM EDTA, pH 8.2 (APPENDIX 2E) 2× SSC (APPENDIX 2E) 1. Cut a piece of positively charged nylon membrane to the appropriate size. Place the membrane on the surface of distilled water and allow to submerge. Leave for 10 min. 2. Prepare the blotting manifold, using distilled water instead of SSC. 3. Add 1 M NaOH and 200 mM EDTA, pH 8.2, to each sample to give a final concentration of 0.4 M NaOH/10 mM EDTA. Denature 10 min at 100°C. Microcentrifuge each tube 5 sec. The alkali/heat treatment denatures the DNA.
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4. Prewash the membrane with 500 µl distilled water. Apply the samples to the membrane. 5. After applying the samples, rinse each well with 500 µl of 0.4 M NaOH and dismantle the manifold. 6. Rinse the membrane briefly in 2× SSC and air dry. 7. Store membrane at room temperature. ALTERNATE PROTOCOL 2
MANUAL PREPARATION OF A DNA DOT BLOT Dot blots can also be set up by hand simply by spotting small aliquots of each sample on to the membrane and waiting for the blot to dry. Repeated applications can be used to apply up to 30 µl of a dilute DNA sample. 1. Cut a strip of uncharged nylon membrane to the desired size and mark out a grid of 0.5-cm × 0.5-cm squares with a blunt pencil. Place membrane on the surface of 6× SSC and allow to submerge. Leave 10 min. Wet nitrocellulose membrane in 20× SSC or a positively charged nylon membrane in distilled water.
2. Add 1⁄2 vol of 20× SSC to DNA to give a final concentration of 6× SSC in the minimum possible volume. Denature the DNA 10 min at 100°C, then place in ice. If using positively charged nylon, add 1 M NaOH and 200 mM EDTA, pH 8.2, to each sample to give a final concentration of 0.4 M NaOH/10 mM EDTA, and denature. If using a nitrocellulose membrane, add an equal volume of 20× SSC to each sample after placing in ice.
3. Place the wetted membrane over the top of an open plastic box so that the bulk of the membrane is freely suspended. 4. Microcentrifuge DNA 5 sec, spot onto the membrane using a pipet, and allow to dry. Do not touch the membrane with the pipet when applying the samples. Up to 2 µl can be spotted in one application.
5. Denature, neutralize, and immobilize DNA on uncharged nylon or nitrocellulose membrane. Rinse and dry a positively charged nylon membrane.
6. Store membrane at room temperature. LITERATURE CITED Dyson, N.J. 1991. Immobilization of nucleic acids and hybridization analysis. In Essential Molecular Biology: A Practical Approach, Vol. 2 (T.A. Brown, ed.) pp. 111-156. IRL Press, Oxford.
Contributed by Terry Brown University of Manchester Institute of Science and Technology Manchester, United Kingdom
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Hybridization Analysis of DNA Blots
APPENDIX 4H
The principle of hybridization analysis is that a single-stranded DNA or RNA molecule of defined sequence (the “probe” which is usually labeled) can base-pair to a second DNA or RNA molecule that is immobilized and contains a complementary sequence (the “target”), with the stability of the hybrid depending on the extent of base pairing that occurs. The technique permits detection of single-copy genes in complex genomes. HYBRIDIZATION ANALYSIS OF A DNA BLOT WITH A RADIOLABELED DNA PROBE This protocol is suitable for hybridization analysis of Southern transfers and dot and slot blots (APPENDIX 4G) with a radioactively labeled DNA probe 100 to 1000 bp in length.
BASIC PROTOCOL
Materials Probe DNA labeled to a specific activity >1 × 108 dpm/µg Aqueous prehybridization/hybridization (APH) solution: 5× SSC/5× Denhardt’s solution (see recipe)/1% (w/v) SDS/100 µg/ml denatured salmon sperm DNA (see Support Protocol 2; added just before use); room temperature and 68°C 2× SSC/0.1% (w/v) SDS 0.2× SSC/0.1% (w/v) SDS, room temperature and 42°C 0.1× SSC/0.1% (w/v) SDS, 68°C 2× and 6× SSC (APPENDIX 2E) Hybridization oven (e.g., Hybridiser HB-1, Techne) or 68°C water bath or incubator Hybridization tube or sealable bag and heat sealer 1. Wet a membrane carrying immobilized DNA in 6× SSC. 2. Place the membrane, DNA-side-up, in a hybridization tube and add ∼1 ml APH solution per 10 cm2 of membrane. Incubate 3 hr in hybridization oven with rotation at 68°C. For a nylon membrane, prehybridize 15 min with 68°C, prehybridization/hybridization solution. See Table A.4H.1 for other hybridization solutions and Table A.4H.2 for alternative blocking reagents.
3. Just before the end of the prehybridization incubation, denature probe DNA 10 min at 100°C. Place in ice. Table A.4H.1
High-Salt Solutions Used in Hybridization Analysis
Stock solution
Composition
20× SSC 20× SSPEa Phosphate solutionb
3.0 M NaCl/0.3 M trisodium citrate 3.6 M NaCl/0.2 M NaH2PO4/0.02 M EDTA, pH 7.7 1 M NaHPO4, pH 7.2c
aSSC
may be replaced with the same concentration of SSPE in all protocols. and hybridize with 0.5 M NaHPO4 (pH 7.2)/1 mM EDTA/7% SDS [or 50% formamide/0.25 M NaHPO4 (pH 7.2)/0.25 M NaCl/1 mM EDTA/7% SDS]; perform moderate-stringency wash in 40 mM NaHPO4 (pH 7.2)/1 mM EDTA/5% SDS; perform high-stringency wash in 40 mM NaHPO4 (pH 7.2)/1 mM EDTA/1% SDS. cDissolve 134 g Na HPO ⋅7H O in 1 liter water, then add 4 ml 85% H PO . The resulting 2 4 2 3 4 solution is 1 M Na+, pH 7.2. bPrehybridize
Contributed by Terry Brown and Kevin Struhl Current Protocols in Protein Science (1998) A.4H.1-A.4H.9 Copyright © 1998 by John Wiley & Sons, Inc.
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4. Pour the APH solution from the hybridization tube and replace with an equal volume of prewarmed (68°C) APH solution. Add denatured probe and incubate with rotation overnight at 68°C. If the specific activity of the probe is 108 dpm/ìg, use 10 ng/ml probe; if it is 1 × 109 dpm/ìg, use 2 ng/ml. See Table A.4H.3 for factors affecting hybrid stability and hybridization rate.
Table A.4H.2 Alternatives to Denhardt/Denatured Salmon Sperm DNA as Blocking Agents in DNA Hybridizationa
Blocking agent
Composition
BLOTTO
5% (w/v) nonfat dried Store at 4°C; use at 4% final milk/0.02% (w/v) NaN3 in H2O concentration 50 mg/ml in 4× SSC Store at 4°C. Use at 500 µg/ml with dextran sulfate or 50 µg/ml without 10 mg/ml in H2O Store at 4°C; use at 100 µg/ml 1 mg/ml poly(A) or poly(C) in Store at 4°C; use at 10 µg/ml in water; H2O appropriate targets: poly(A) for AT-rich DNA, poly(C) for GC-rich DNA
Heparin (porcine grade II) Yeast tRNA Homopolymer DNA aThis
Storage and use
table is based on Brown (1991) with permission from BIOS Scientific Publishers.
Table A.4H.3
Factors Influencing Hybrid Stability and Hybridization Ratea
Factor A. Hybrid stabilityb Ionic strength Base composition Destabilizing agents Mismatched base pairs Duplex length B. Hybridization rateb Temperature Ionic strength Destabilizing agents Mismatched base pairs Duplex length Viscosity Probe complexity Base composition pH
Influence Tm increases 16.6°C for each 10-fold increase in monovalent cations between 0.01 and 0.40 M NaCl AT base pairs are less stable than GC base pairs in aqueous solutions containing NaCl Each 1% of formamide reduces the Tm by about 0.6°C for a DNA-DNA hybrid. 6 M urea reduces the Tm by about 30°C Tm is reduced by 1°C for each 1% of mismatching Negligible effect with probes >500 bp Maximum rate occurs at 20°-25°C below Tm for DNA-DNA hybrids, 10°-15°C below Tm for DNA-RNA hybrids Optimal hybridization rate at 1.5 M Na+ 50% formamide has no effect, but higher or lower concentrations reduce the hybridization rate Each 10% of mismatching reduces the hybridization rate by a factor of two Hybridization rate is directly proportional to duplex length Increased viscosity increases the rate of membrane hybridization; 10% dextran sulfate increases rate by factor of ten Repetitive sequences increase the hybridization rate Little effect Little effect between pH 5.0 and pH 9.0
aThis
Hybridization Analysis of DNA Blots
table is based on Brown (1991) with permission from BIOS Scientific Publishers. have been relatively few studies of the factors influencing membrane hybridization. In several instances extrapolations are made from what is known about solution hybridization. This is probably reliable for hybrid stability, less so for hybridization rate. bThere
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Table A.4H.4
Troubleshooting Guide for DNA Blotting and Hybridization Analysisa
Problem
Possible causeb
Solution
Poor signal
Probe specific activity too low Inadequate depurination Inadequate transfer buffer
Check labeling protocol if specific activity is <108 dpm/µg. Check depurination if transfer of DNA >5 kb is poor. 1. Check that 20× SSC has been used as the transfer solution if small DNA fragments are retained inefficiently when transferring to nitrocellulose. 2. With some brands of nylon membrane, add 2 mM Sarkosyl to the transfer buffer. 3. Try alkaline blotting to a positively charged nylon membrane. Refer to text for recommendations regarding amount of target DNA to load per blot. See recommendations in APPENDIX 4G. See recommendations in APPENDIX 4G. Consider vacuum blotting as an alternative to capillary transfer. 1. Check that the correct amount of DNA has been used in the labeling reaction. 2. Check recovery of the probe after removal of unincorporated nucleotides. 3. Use 10% dextran sulfate in the hybridization solution. 4. Change to a single-stranded probe, as reannealing of a double-stranded probe reduces its effective concentration to zero after hybridization for 8 hr. Denature as described in the protocols. When dot or slot blotting, use the double denaturation methods described in APPENDIX 4G, or blot onto positively charged nylon. If using a nylon membrane, leave the blocking agents out of the hybridization solution. Use a lower temperature or higher salt concentration. If necessary, estimate Tm. Increase hybridization temperature to 65°C in the presence of formamide (see Alternate Protocol). If using formamide with a DNA probe, increase the hybridization time to 24 hr. Check the target molecules are not too short to be retained efficiently by the membrane type (see Table A.4G.1).
Not enough target DNA Poor immobilization of DNA Transfer time too short Inefficient transfer system Probe concentration too low
Incomplete denaturation of probe Incomplete denaturation of target DNA Blocking agents interfering with the target-probe interaction Final wash was too stringent Hybridization temperature too low with an RNA probe Hybridization time too short Inappropriate membrane
continued
5. Pour out the APH solution and add an equal volume of 2× SSC/0.1% SDS. Incubate with rotation 10 min at room temperature. Change the wash solution after 5 min. To reduce background, it may be beneficial to increase the volume of the wash solutions by 100%.
6. Replace the wash solution with an equal volume of 0.2× SSC/0.1% SDS and incubate with rotation 10 min at room temperature. Change the wash solution after 5 min (low-stringency wash). 7. If desired, carry out two 15-min moderate-stringency washes using 42°C 0.2× SSC/0.1% SDS.
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Table A.4H.4
Troubleshooting Guide for DNA Blotting and Hybridization Analysisa, continued
Problem
Possible causeb
Solution
Problems with electroblotting
Make sure no bubbles are trapped in the filter-paper stack. Soak the filter papers thoroughly in TBE before assembling the blot. Used uncharged rather than charged nylon. Remove unincorporated nucleotides by column chromatography. Filter the relevant solution(s). Rinse membrane in 2× SSC after blotting. Rinse membrane in 2× SSC after blotting.
Spotty background Unincorporated nucleotides not removed from labeled probe Particles in the hybridization buffer Agarose dried on the membrane Baking or UV crosslinking when membrane contains high salt Patchy or generally Insufficient blocking agents high background Part of the membrane was allowed to dry out during hybridization or washing Membranes adhered during hybridization or washing
See text for of discussion of extra/alternative blocking agents. Avoid by increasing the volume of solutions if necessary.
Do not hybridize too many membranes at once (ten minigel blots for a hybridization tube, two for a bag is maximum). Bubbles in a hybridization bag If using a bag, fill completely so there are no bubbles. Walls of hybridization bag collapsed Use a stiff plastic bag; increase volume of hybridization on to membrane solution. Not enough wash solution Increase volume of wash solution to 2 ml/10 cm2 of membrane. Hybridization temperature too Increase hybridization temperature to 65°C in the presence low with an RNA probe of formamide (see Alternate Protocol). Formamide needs to be deionized Although commercial formamide is usually satisfactory, background may be reduced by deionizing immediately before use. Labeled probe molecules are too short 1. Use a 32P-labeled probe as soon as possible after labeling, as radiolysis can result in fragmentation. 2. Reduce amount of DNase I used in nick translation. Probe concentration too high Check that the correct amount of DNA has been used in the labeling reaction. Inadequate prehybridization Prehybridize for at least 3 hr with nitrocellulose or 15 min for nylon. Probe not denatured Denature as described in the protocols. Inappropriate membrane type If using a nonradioactive label, check that the membrane is compatible with the detection system. continued
8. If desired, carry out two 15-min high-stringency washes using 68°C 0.1× SSC/0.1% SDS. 9. Pour off the final wash solution, rinse the membrane in 2× SSC at room temperature, and blot excess liquid. Wrap in plastic wrap. Autoradiograph. Do not allow the membrane to dry out if it is to be reprobed. See Table A.4H.4 for troubleshooting. Hybridization Analysis of DNA Blots
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Table A.4H.4
Troubleshooting Guide for DNA Blotting and Hybridization Analysisa, continued
Problem
Possible causeb
Solution
Hybridization with dextran sulfate
Extra bands
Nonspecific background in one or more tracks
Cannot remove probe after hybridization Decrease in signal intensity when reprobed aBased
Dextran sulfate sometimes causes background hybridization. Place the membrane between Schleicher and Schuell no. 589 WH paper during hybridization, and increase volume of hybridization solution (including dextran sulfate) by 2.5%. Not enough SDS in wash solutions Check the solutions are made up correctly. Final wash was not stringent enough Use a higher temperature or lower salt concentration. If necessary, estimate Tm. Probe contains nonspecific sequences Purify shortest fragment that contains the desired (e.g., vector DNA) sequence. Target DNA is not completely Check the restriction digestion (APPENDIX 4I). restriction digested Formamide not used with an RNA RNA-DNA hybrids are relatively strong but are probe destabilized if formamide is used in the hybridization solution. Probe is contaminated with genomic Check purification of probe DNA. The problem is more DNA severe when probes are labeled by random printing. Change to nick translation. Insufficient blocking agents See text for of discussion of extra/alternative blocking agents. Final wash did not approach the Use a higher temperature or lower salt concentration. If desired stringency necessary, estimate Tm. Probe too short Sometimes a problem with probes labeled by random priming. Change to nick translation. Membrane dried out after Make sure the membrane is stored moist between hybridization hybridization and stripping. Poor retention of target DNA during probe stripping
1. Check calibration of UV source if cross-linking on nylon. 2. Use a less harsh stripping method (Support Protocol 1).
on Dyson (1991). each category, possible causes are listed in decreasing order of likelihood.
bWithin
HYBRIDIZATION ANALYSIS OF A DNA BLOT WITH A RADIOLABELED RNA PROBE Purified RNA polymerases from bacteriophages such as SP6, T3, and T7 are very efficient at synthesizing RNA in vitro from DNA sequences cloned downstream of the appropriate phage promoter (Little and Jackson, 1987). If a radiolabeled ribonucleotide is added to the reaction mixture, the polymerase synthesizes several micrograms of uniformly labeled single-stranded RNA with specific activities ≥109 dpm/µg. The hybridization procedure is suitable for both nitrocellulose and nylon membranes, though backgrounds may be higher with nylon. Additional Materials (also see Basic Protocol) TE buffer, pH 8.0 (APPENDIX 2E) Labeling buffer: 200 mM Tris⋅Cl (pH 7.5)/30 mM MgCl2/10 mM spermidine Nucleotide mix: 2.5 mM ATP/2.5 mM CTP/2.5 mM MGTP/20 mM Tris⋅Cl, pH 7.5 (store at −20°C) 200 mM dithiothreitol (DTT), freshly prepared
ALTERNATE PROTOCOL
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20 U/µl human placental ribonuclease inhibitor [α-32P]UTP, 20 mCi/ml (800 Ci/mmol) or 10 mCi/ml (400 Ci/mmol) SP6 or T7 RNA polymerase RNase-free DNase I 0.25 M EDTA, pH 8.0 (APPENDIX 2E) Formamide prehybridization/hybridization (FPH) solution: 5× SSC/5× Denhardt’s solution/50% (w/v) formamide/1% (w/v) SDS/100 µg/ml denatured salmon sperm DNA (see Support Protocol; add just before use) 2× SSC containing 25 µg/ml RNase A + 10 U/ml RNase T1 1. Digest cloned DNA (see Table A.4H.5 for suitable vectors) for the sequence to be transcribed with a restriction endonuclease to linearize and introduce an endpoint for RNA synthesis. Purify the DNA by phenol extraction and ethanol precipitation (APPENDIX 4E) and resuspend in TE buffer, pH 8.0, at a concentration of 1 mg/ml. 2. Mix the following at room temperature: 4 µl labeling buffer 1.5 µl nucleotide mix 1 µl 200 mM DTT 1 µl (20 U) human placental ribonuclease inhibitor 2 µg purified plasmid DNA from step 1 100 to 200 µCi [α-32P]UTP H2O to a final volume of 20 µl. 3. Add 5 U of SP6 or T7 RNA polymerase. Incubate 1 hr at 40°C for SP6 polymerase or at 37°C for T7 polymerase. 4. Add 2 U RNase-free DNase I and incubate 10 min at 37°C. Stop the reaction by adding 2 µl of 0.25 M EDTA, pH 8.0. 5. Measure the specific activity of the RNA by acid precipitation and remove unincorporated nucleotides by the spin-column procedure (Support Protocol 3). Store labeled probe up to 2 days at −20°C. The specific activity should be at least 7 × 108 dpm/ìg, preferably >109 dpm/ìg.
Table A.4H.5 Selection of Cloning Vectors Incorporating Promoters for Bacteriophage RNA Polymerases
Hybridization Analysis of DNA Blots
Vector
Size (bp)
Markersa
Promoters
pBluescript pGEM series pGEMEX-1 pSELECT-1 pSP18, 19, 64, 65 pSP70, 71, 72, 73 pSPORT1 pT3/T7 series pWE15 pWE16
2950 2746-3223 4200 3422 2999-3010 2417-2464 4109 2700, 2950 8800 8800
amp, lacZ′ amp, lacZ′ amp tet, lacZ′ amp amp amp, lacZ′ amp, lacZ′ amp, neo amp, dhfr
T3, T7 SP6, T7 SP6, T3, T7 SP6, T7 SP6 SP6, T7 SP6, T7 T3, T7 T3, T7 T3, T7
aAbbreviations:
amp, ampicillin resistance; dhfr, dihydrofolate reductase; lacZ′, β-galactosidase α-peptide; neo, neomycin phosphotransferase (kanamycin resistance); tet, tetracycline resistance.
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6. Prehybridize in FPH solution and incubate 3 hr at 42°C. 7. Replace the FPH solution with an equal volume of fresh prewarmed solution. Add the labeled probe to a concentration of 1 to 5 ng/ml and incubate overnight with rotation at 42°C. 8. Carry out two 10-min low-stringency washes in 2× SSC/0.1% at room temperature. 9. Replace the wash solution with an equal volume of 2× SSC containing 25 µg/ml RNase A + 10 U/ml RNase T1 and incubate with rotation for 30 min at room temperature. 10. Carry out moderate- and high-stringency washes as desired, rinse the membrane in 2× SSC, and set up autoradiography. REMOVAL OF PROBES FROM HYBRIDIZED MEMBRANES If the DNA has been immobilized on the membrane by UV crosslinking (for uncharged nylon membranes) or by alkaline transfer (for positively charged nylon), the covalent matrix-target DNA interaction is much stronger than the hydrogen-bonded target-probe interaction, so it is possible to remove (or “strip off”) the hybridized probe.
SUPPORT PROTOCOL 1
Additional Materials (also see Basic Protocol) Mild stripping solution: 5mM Tris⋅Cl, pH 8.0/0.2 mM EDTA/0.1× Denhardt’s solution Moderate stripping solution: 200 mM Tris⋅Cl, pH 8.0/0.1× SSC/0.1% (w/v) SDS 0.4 M NaOH 0.1% (w/v) SDS, 100°C 1a. Mild treatment: Wash the membrane in several hundred milliliters of mild stripping solution for 2 hr at 65°C. 1b. Moderate treatment: Wash the membrane in 0.4 M NaOH for 30 min at 45°C. Then rinse twice in several hundred milliliters of moderate stripping solution for 10 min at room temperature. 1c. Harsh treatment: Pour several hundred milliliters of boiling 0.1% SDS onto the membrane. Cool to room temperature. If a membrane is to be reprobed, it must not be allowed to dry out between hybridization and stripping.
2. Place membrane on a sheet of dry Whatman 3MM filter paper and blot excess liquid with a second sheet. Wrap the membrane in plastic wrap and set up autoradiography. If signal is still seen after autoradiography, rewash using harsher conditions.
3. The membrane can now be rehybridized or dried and stored at room temperature for later use. PREPARATION OF DENATURED SALMON SPERM DNA Dissolve 10 mg Sigma type III salmon sperm DNA (sodium salt) in 1 ml water. Pass vigorously through a 17-G needle 20 times to shear the DNA. Place in a boiling water bath for 10 min, then chill. Use immediately or store at −20°C in small aliquots. If stored, reheat to 100°C for 5 min and chill on ice immediately before using.
SUPPORT PROTOCOL 2
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SUPPORT PROTOCOL 3
SPIN-COLUMN PROCEDURE FOR SEPARATING RADIOACTIVELY LABELED DNA FROM UNINCORPORATED dNTP PRECURSORS In this method, the packing and running of the column is accomplished by centrifugation rather than by gravity. It is more rapid than conventional chromatography and is useful when many samples are involved; however, removal of the dNTPs may be less quantitative. Spin columns can be purchased commercially at moderate expense. CAUTION: Use extreme care that centrifuge and work area do not become contaminated with radioactivity. Whenever possible, place samples behind a lucite shield. 1. Plug bottom of a 5-ml disposable syringe with clean, silanized glass wool. Fill syringe with an even suspension of the column resin, and place in a polypropylene tube that is suitable for centrifugation in a tabletop centrifuge (Fig. A.4H.1). Centrifuge 2 to 3 min at a setting of 4 in order to pack the column. The resin should not be packed too tightly or too loosely. Therefore, adjust the time and speed in the tabletop centrifuge according to individual laboratory conditions (∼1200 rpm).
2. Dilute radioactive sample with TE buffer (APPENDIX 2E) to 100 µl and load in the center of the column. Place syringe in a new polypropylene tube, and centrifuge 5 min at a setting of 5 to 6. 3. Save liquid at bottom of tube containing the labeled DNA. Dispose of all radioactive waste properly.
load sample here
5-ml syringe polypropylene centrifuge tube column resin
silanized glass wool
radioactive DNA emerges here after centrifugation
Figure A.4H.1 Preparation of a spin column. A 5-ml syringe containing the column resin is placed upside down in a polypropylene centrifuge tube. Hybridization Analysis of DNA Blots
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REAGENTS AND SOLUTIONS Denhardt solution, 100× 10 g Ficoll 400 10 g polyvinylpyrrolidone 10 g bovine serum albumin (Pentax Fraction V; Miles Laboratories) H2O to 500 ml Filter sterilize and store at −20C in 25-ml aliquots LITERATURE CITED Brown, T.A., ed. 1991. Molecular Biology Labfax. BIOS Scientific Publishers, Oxford. Dyson, N.J. 1991. Immobilization of nucleic acids and hybridization analysis. In Essential Molecular Biology: A Practical Approach, Vol. 2 (T.A. Brown, ed.) pp. 111-156. IRL Press, Oxford. Little, P.F.R. and Jackson, I.J. 1987. Application of plasmids containing promoters specific for phage-encoded RNA polymerases. In DNA Cloning: A Practical Approach, Vol. 3 (D.M. Glover, ed.) pp. 1-18. IRL Press at Oxford University Press, Oxford.
Contributed by Terry Brown University of Manchester Institute of Science and Technology Manchester, United Kingdom Kevin Struhl (spin-column procedure) Harvard Medical School Boston, Massachusetts
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Digestion of DNA with Restriction Endonucleases
APPENDIX 4I
Restriction endonucleases recognize short DNA sequences and cleave double-stranded DNA at specific sites within or adjacent to the recognition sequences. Restriction endonuclease cleavage of DNA into discrete fragments is one of the most basic procedures in molecular biology. Table A.4I.1 describes restriction endonucleases and their properties. A number of factors affect restriction endonuclease reactions. Contaminants found in some DNA preparations (e.g., protein, phenol, chloroform, ethanol, EDTA, SDS, high salt concentration) may inhibit restriction endonuclease activity. The effects of contaminants may be overcome by increasing the number of enzyme units added to the reaction mixture (up to 10 to 20 U per microgram DNA), increasing the reaction volume to dilute potential inhibitors, or increasing the duration of incubation. Digestion of genomic DNA can be facilitated by the addition of the polycation spermidine (final concentration 1 to 2.5 mM), which acts by binding negatively charged contaminants. Larger amounts (up to 20-fold more) of some enzymes are necessary to cleave supercoiled plasmid or viral DNA as compared to the amount needed to cleave linear DNA. In addition, some enzymes cleave their defined sites with different efficiencies, presumably due to differences in flanking nucleotides. DIGESTING A SINGLE DNA SAMPLE WITH A SINGLE RESTRICTION ENDONUCLEASE
BASIC PROTOCOL
Restriction endonuclease cleavage is accomplished simply by incubating the enzyme(s) with the DNA in appropriate reaction conditions. The amounts of enzyme and DNA, the buffer and ionic concentrations, and the temperature and duration of the reaction will vary depending upon the specific application. Materials DNA sample in H2O or TE buffer 10× restriction endonuclease buffers (see Support Protocol) Restriction endonucleases Loading buffer, 10×: 20% (w/v) Ficoll 400/0.1 M Na2 EDTA (pH 8.0)/1.0% (w/v) SDS/0.25% (w/v) bromphenol blue/0.25% (w/v) xylene cyanol (optional) 0.5 M EDTA, pH 8.0 (APPENDIX 2E; optional) 1. Pipet the following into a clean microcentrifuge tube: x µl DNA (0.1 to 4 µg DNA in H2O or TE buffer) 2 µl 10× restriction buffer (Table A.4I.1) 18 − x µl H2O. A 20-ìl reaction is convenient for analysis by electrophoresis in polyacrylamide or agarose gels. The amount of DNA to be cleaved and/or the reaction volume can be increased or decreased provided the proportions of the components remain constant.
2. Add restriction endonuclease (1 to 5 U/µg DNA) and incubate the reaction mixture 1 hr at the recommended temperature (in general, 37°C). In principle, 1 U restriction endonuclease completely digests 1 ìg of purified DNA in 60 min using the recommended assay conditions. The volume of restriction endonuclease added should be less than 1⁄10 the volume of the final reaction mixture, because glycerol in the enzyme storage buffer may interfere with the reaction. Contributed by Kenneth D. Bloch Current Protocols in Protein Science (1998) A.4I.1-A.4I.3 Copyright © 1998 by John Wiley & Sons, Inc.
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A.4I.1 Supplement 13
Table A.4I.1
Cross Index of Recognition Sequences and Restriction Endonucleasesa,b AATT
↓∗∗∗∗
ACGT
AGCT
ATAT
CATG
CCGG CGCG
CTAG
Tsp 509I
∗↓∗∗∗
GATC
GCGC
GGCC
GTAC
TATA
TCGA
TGCA
TTAA
Dpn II Mbo Ic Sau3 AI Msp I Hpa IIc
Mae II
∗∗↓∗∗
Alu I
Bfa I Bst UI
Hin PI Dpn I
∗∗∗↓∗
Csp 61 Hae III
Taq I
Mse I
Rsa I
Hha I
∗∗∗∗↓
Nla III
A↓∗∗∗∗T
Apo I
A∗↓∗∗∗T
Afl III
Hin dIII
Age I Bsr FI Bsa WI
Mlu I Afl III
Spe I
Bgl II Bst YI
Ppu 10I
Cla I Bsp DI
Psp1406I
A∗∗↓∗∗T
Ssp I
Eco 47III
Stu I
Ase I
Sca I
A∗∗∗↓∗T A∗∗∗∗↓T C↓∗∗∗∗G
Nsp HI Mun I
Nco I Sty I Dsa I
C∗↓∗∗∗G
Hae II Xma I Ava I
Dsa I
Sma I
Msp A1I
Avr II Sty I
Nsi I Eag I Eae I
Bsi WI
Sfc I
Xho I Ava I
Afl II
Nde I
C∗∗↓∗∗G
Pml I Bsa AI
Pvu II Msp A1I
C∗∗∗↓∗G
Bsi EI
Pvu I Bsi EI
Sac II
C∗∗∗∗↓G G↓∗∗∗∗C
Sfc I
Pst I
Eco RI Apo I
G∗↓∗∗∗C
Ngo MI Bss HII Bsr Fl
Nhe I
Bam HI Bst YI
Bsp 120I Acc 65I Ban I
Nar I Bsa HI
Bsa HI
G∗∗↓∗∗C
Kas I Ban I
Ecl 136II Eco RV
Acc I
Ehe I
Nae I
Sal I
Apa LI
Acc I Hpa I Hin cII
Bst1107I Hin cII
G∗∗∗↓∗C G∗∗∗∗↓C
Aat II
T↓∗∗∗∗A
Sac I Ban II Bsi HKAI Bsp 1286I
Sph I Nsp H1
Bsp HI
Bbe I Hae II Bsp EI Bsa WI
Xba I
Bcl I
Apa I Ban II Bsp 1286I
Kpn I
Eae I
Bsr GI
T∗↓∗∗∗A T∗∗↓∗∗A
Bsp1286I BsiHKAI
Bst BI Sna BI Bsa AI
Nru I
Fsp I
Msc I
Dra I
T∗∗∗↓∗A T∗∗∗∗↓A
aReprinted
by permission of New England Biolabs.
bSequences at the top of each column are written 5′ to 3′. Asterisks at the left of each row are place holders for nucleotides within a recognition
sequence, and arrows indicate the point of cleavage. Sequences of complementary strands and their cleavage sites are implied. Enzymes written in bold type recognize only one sequence, while those in light type have multiple recognition sequences. cSequence cleaved identically by two or more enzymes that are affected differently by DNA modification at that site.
Digestion of DNA with Restriction Endonucleases
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3. Stop the reaction and prepare it for agarose or acrylamide gel electrophoresis (APPENDIX 4F) by adding 5 µl (20% of reaction vol) gel loading buffer. The reaction can also be stopped by chelating Mg2+ with 0.5 ìl of 0.5 M EDTA (12.5 mM final concentration). Many enzymes can be irreversibly inactivated by incubating 10 min at 65°C; some enzymes that are partially or completely resistant to heat inactivation at 65°C may be inactivated by incubating 15 min at 75°C. When the enzyme(s) is completely resistant to heat inactivation, DNA may be purified from the reaction mixture by extraction with phenol and precipitation in ethanol (APPENDIX 4E) or by using a silica matrix suspension.
RESTRICTION ENDONUCLEASE BUFFERS 10× sodium chloride–based buffers 100 mM Tris⋅Cl, pH 7.5 (APPENDIX 2E) 100 mM MgCl2 10 mM dithiothreitol (DTT) 1 mg/ml bovine serum albumin (BSA) 0, 0.5, 1.0, or 1.5 M NaCl
SUPPORT PROTOCOL
The concentration of NaCl depends upon the restriction endonuclease (Table 3.1.2). The four different NaCl concentrations listed above are sufficient to cover the range for essentially all commercially available enzymes except those requiring a buffer containing KCl instead of NaCl. Autoclaved gelatin (at 1 mg/ml) can be used instead of BSA. Note that most restriction enzymes are provided by the suppliers with the appropriate buffers.
10× KCl-based buffer 60 mM Tris⋅Cl, pH 8.0 (APPENDIX 2E) 60 mM MgCl2 200 mM KCl 60 mM 2-mercaptoethanol (2-ME) 1 mg/ml BSA 10× potassium acetate–based buffer 660 mM potassium acetate 330 mM Tris⋅acetate, pH 7.9 100 mM magnesium acetate 5 mM dithiothreitol (DTT) 1 mg/ml BSA (optional) 2× potassium glutamate–based buffer 200 mM potassium glutamate 50 mM Tris⋅acetate, pH 7.5 100 µg/ml BSA 1 mM 2-mercaptoethanol (2-ME) LITERATURE CITED Danna, A.J. 1980. Determination of fragment order through partial digests and multiple enzyme digests. Methods Enzymol. 65:449-467.
Contributed by Kenneth D. Bloch Harvard Medical School Boston, Massachusetts Molecular Biology Techniques
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The Polymerase Chain Reaction
APPENDIX 4J
The polymerase chain reaction (PCR) is a rapid procedure for in vitro enzymatic amplification of a specific segment of DNA. Like molecular cloning, PCR has spawned a multitude of experiments that were previously impossible. The number of applications of PCR seems infinite—and is still growing. They include direct cloning from genomic DNA or cDNA, in vitro mutagenesis and engineering of DNA, genetic fingerprinting of forensic samples, assays for the presence of infectious agents, prenatal diagnosis of genetic diseases, analysis of allelic sequence variations, analysis of RNA transcript structure, genomic footprinting, and direct nucleotide sequencing of genomic DNA and cDNA. The theoretical basis of PCR is outlined in Figure A.4J.1. There are three nucleic acid segments: the segment of double-stranded DNA to be amplified and two single-stranded oligonucleotide primers flanking it. Additionally, there is a protein component (a DNA polymerase), appropriate deoxyribonucleoside triphosphates (dNTPs), a buffer, and salts. The primers are added in vast excess compared to the DNA to be amplified. They hybridize to opposite strands of the DNA and are oriented with their 3′ ends facing each other so that synthesis by DNA polymerase (which catalyzes growth of new strands 5′→3′) extends across the segment of DNA between them. One round of synthesis results in new strands of indeterminate length which, like the parental strands, can hybridize to
original DNA PCR primer new DNA
DNA + primers + dNTPs + DNA polymerase
denature and synthesize
denature and synthesize
denature and synthesize
etc.
etc.
etc.
Figure A.4J.1 The polymerase chain reaction. DNA to be amplified is denatured by heating the sample. In the presence of DNA polymerase and excess dNTPs, oligonucleotides that hybridize specifically to the target sequence can prime new DNA synthesis. The first cycle is characterized by a product of indeterminate length; however, the second cycle produces the discrete “short product” which accumulates exponentially with each successive round of amplification. This can lead to the many million-fold amplification of the discrete fragment over the course of 20 to 30 cycles.
Contributed by Martha F. Kramer and Donald M. Coen Current Protocols in Protein Science (2002) A.4J.1-A.4J.8 Copyright © 2002 by John Wiley & Sons, Inc.
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the primers upon denaturation and annealing. These products accumulate only arithmetically with each subsequent cycle of denaturation, annealing to primers, and synthesis. However, the second cycle of denaturation, annealing, and synthesis produces two single-stranded products that together compose a discrete double-stranded product which is exactly the length between the primer ends. Each strand of this discrete product is complementary to one of the two primers and can therefore participate as a template in subsequent cycles. The amount of this product doubles with every subsequent cycle of synthesis, denaturation, and annealing, accumulating exponentially so that 30 cycles should result in a 228-fold (270 million–fold) amplification of the discrete product. BASIC PROTOCOL
ENZYMATIC AMPLIFICATION OF DNA BY PCR: STANDARD PROCEDURES AND OPTIMIZATION This protocol describes a method for amplifying DNA enzymatically by the polymerase chain reaction (PCR), including procedures to quickly determine conditions for successful amplification of the sequence and primer sets of interest, and to optimize for specificity, sensitivity, and yield. The first step of PCR simply entails mixing template DNA, two appropriate oligonucleotide primers, Taq or other thermostable DNA polymerases, deoxyribonucleoside triphosphates (dNTPs), and a buffer. Once assembled, the mixture is cycled many times (usually 30) through temperatures that permit denaturation, annealing, and synthesis to exponentially amplify a product of specific size and sequence. The PCR products are then displayed on an appropriate gel and examined for yield and specificity. Many important variables can influence the outcome of PCR. Careful titration of the MgCl2 concentration is critical. Additives that promote polymerase stability and processivity or increase hybridization stringency, and strategies that reduce nonspecific primertemplate interactions, especially prior to the critical first cycle, generally improve amplification efficiency. This protocol, using Taq DNA polymerase, is designed to optimize the reaction components and conditions in one or two stages. The first stage (steps 1 to 7) determines the optimal MgCl2 concentration and screens several enhancing additives. Most suppliers (of which there are many) of Taq and other thermostable DNA polymerases provide a unique optimized MgCl2-free buffer with MgCl2 in a separate vial for user titration. The second stage (steps 8 to 13) compares methods for preventing pre-PCR low-stringency primer extension, which can generate nonspecific products. This has come to be known as “hot start,” whether one omits an essential reaction component prior to the first denaturing-temperature step or adds a reversible inhibitor of polymerase. Hot-start methods can greatly improve specificity, sensitivity, and yield. Use of any one of the hot-start approaches is strongly recommended if primer-dimers or other nonspecific products are generated, or if relatively rare template DNA is contained in a complex mixture, such as viral nucleic acids in cell or tissue preparations. This protocol suggests some relatively inexpensive methods to achieve hot start, and lists several commercial hot-start options which may be more convenient, but of course more expensive. NOTE: Use only molecular biology–grade water (i.e., DNase, RNase, and nucleic acid free) in all steps and solutions.
The Polymerase Chain Reaction
Materials 10× MgCl2-free PCR buffer (see recipe) 50 µM oligonucleotide primer 1: 50 pmol/µl in sterile H2O (store at −20°C) 50 µM oligonucleotide primer 2: 50 pmol/µl in sterile H2O (store at −20°C) Template DNA: 1 µg mammalian genomic DNA or 1.0 to 100.0 pg of plasmid DNA 25 mM 4dNTP mix (see recipe)
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5 U/µl Taq DNA polymerase (native or recombinant) Enhancer agents (optional; see recipe) 15 mM (L), 30 mM (M), and 45 mM (H) MgCl2 Mineral oil TaqStart Antibody (Clontech) Ficoll 400 (optional): prepare as 10× stock; store indefinitely at room temperature Tartrazine dye (optional): prepare as 10× stock; store indefinitely at room temperature 0.5 ml thin-walled PCR tubes Automated thermal cycler NOTE: Do not use DEPC to treat water, reagents, or glassware. NOTE: Reagents should be prepared in sterile, disposable labware, taken directly from its packaging, or in glassware that has been soaked in 10% bleach, thoroughly rinsed in tap water followed by distilled water, and if available, exposed to UV irradiation for ~10 min. Multiple small volumes of each reagent should be stored in screw-cap tubes. This will then serve as the user’s own optimization “kit.” Thin-walled PCR tubes are recommended. Optimize reaction components 1. Prepare four reaction master mixes according to the recipes given in Table A.4J.1. Enhancing agents probably work by different mechanisms, such as protecting enzyme activity and decreasing nonspecific primer binding. However, their effects cannot be readily predicted—what improves amplification efficiency for one primer pair may decrease the amplification efficiency for another. Thus it is best to check a panel of enhancers during development of a new assay.
2. Aliquot 90 µl master mix I into each of three 0.5-ml thin-walled PCR tubes labeled I-L, I-M, and I-H. Similarly, aliquot mixes II through IV into appropriately labeled tubes. Add 10 µl of 15 mM MgCl2 into one tube of each master mix (labeled L; 1.5 mM final). Similarly, aliquot 10 µl of 30 mM and 45 mM MgCl2 to separate tubes of Table A.4J.1
Master Mixes for Optimizing Reaction Components
Components
Final concentration
Master mixa (µl)
Per reaction I
II
III
IV
10× MgCl2-free PCR buffer
1×
10 µl
40.0
40.0
40.0
40.0
Primer 1 Primer 2
0.5 µM 0.5 µM
1 µl 1 µl
4.0 4.0
4.0 4.0
4.0 4.0
4.0 4.0
Template DNAb 25 mM 4dNTP mixc
Undiluted 0.2 mM
1 vol 0.8 µl
4 vol 3.2
4 vol 3.2
4 vol 3.2
4 vol 3.2
Taq polymerase DMSOd (20×)
2.5 U 5%
0.5 µl 5 µl
2.0 —
2.0 20.0
2.0 —
2.0 —
Glycerold (10×) PMPEd (100×)
10% 1%
10 µl 1 µl
— —
— —
40.0 —
— 4.0
H2O
—
To 90 µl
To 360
To 360
aTotal
To 360
To 360
volume = 360 µl (i.e., enough for n + 1 reactions).
bTemplate
DNA volume (“vol”) is generally 1 to 10 µl.
2 mM 4dNTP mix is preferred, use 10 µl per reaction, or 40 µl for each master mix; adjust the volume of water accordingly. cIf
dSubstitute
with other enhancer agents (see recipe in Reagents and Solutions) as available.
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each master mix (labeled M and H, respectively; 3.0 and 4.5 mM final concentrations respectively). It is helpful to set the tubes up in a three-by-four array to simplify aliquotting. Each of the three Mg2+ concentrations is combined with each of the four master mixes.
3. Overlay the reaction mixture with 50 to 100 µl mineral oil (2 to 3 drops). To include hot start in the first step, overlay reaction mixes with oil before adding the MgCl2, heat the samples to 95°C in the thermal cycler or other heating block, and add the MgCl2 once the elevated temperature is reached. Once the MgCl2 has been added, do not allow the samples to cool below the optimum annealing temperature prior to performing PCR. Alternatives to mineral oil include silicone oil and paraffin beads. Additionally, certain cyclers feature heated lids that are designed to obviate the need for an oil overlay.
Choose cycling parameters 4. Using the following guidelines, program the automated thermal cycler according to the manufacturer’s instructions. 30 cycles: 30 sec 30 sec
94°C (denaturation) 55° (GC content ≤50%) or 60°C (GC content >50%) (annealing)
∼60 sec/kb product sequence 72°C
(extension)
Cycling parameters are dependent upon the sequence and length of the template DNA, the sequence and percent complementarity of the primers, and the ramp times of the thermal cycler used. Thoughtful primer design will reduce potential problems. Denaturation, annealing, and extension are each quite rapid at the optimal temperatures. The time it takes to achieve the desired temperature inside the reaction tube (i.e., the ramp time) is usually longer than either denaturation or primer annealing. Thus, ramp time is a crucial cycling parameter. Manufacturers of the various thermal cyclers on the market provide ramp time specifications for their instruments. Ramp times are lower with thin-walled reaction tubes. The optimal extension time also depends on the length of the target sequence. Allow ∼1 min/kb for this step for target sequences >1 kb, and as little as a 2-sec pause for targets <100 bases in length. The number of cycles depends on both the efficiency of the reaction and the amount of template DNA in the reaction. Starting with as little as 100 ng of mammalian genomic DNA (∼104 cell equivalents), after 30 cycles, 10% of the reaction should produce a band that is readily visible on an ethidium bromide–stained gel as a single predominant band. With more template, fewer cycles may suffice. With much less template, further optimization is recommended rather than increasing the cycle number. Greater cycle numbers (e.g., >40) can reduce the polymerase specific activity, increase nonspecific amplification, and deplete substrate (nucleotides). Many investigators lengthen the time for the last extension step—to 7 min, for example—to try to ensure that all the PCR products are full length. These guidelines are appropriate for most commercially available thermal cyclers. For rapid cyclers, consult the manufacturers’ protocols.
Analyze the product 5. Electrophorese 10 µl from each reaction on an agarose, nondenaturing polyacrylamide, or sieving agarose gel appropriate for the PCR product size expected. Stain with ethidium bromide. For resolution of PCR products between 100 and 1000 bp, an alternative to nondenaturing polyacrylamide gels or sieving agarose is a composite 3% (w/v) NuSieve (FMC Bioproducts) agarose/1% (w/v) SeaKem (FMC Bioproducts) agarose gel. SeaKem increases the mechanical strength of the gel without decreasing resolution. The Polymerase Chain Reaction
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Table A.4J.2
Master Mixes for Optimizing First-Cycle Reactions
Components 10× PCR buffer MgCl2 (L, M, or H) Primer 1 Primer 2 Additive Template DNA 25 mM 4dNTP mixb Taq polymerase Taq pol + TaqStart H2O Preparation temperature aV,
Final concentration 1× Optimal 0.5 µM 0.5 µM Optimal —b 0.2 mM 2.5 U 2.5 U To 100 µl
Master mix (µl) A 10 10 1.0 1.0 Va Va 0.8 0.5 — Va Room temperature
B 10 10 1.0 1.0 Va Va 0.8 0.5 — Va Ice slurry
C 10 10 1.0 1.0 Va Va 0.8 — — Va Room temperature
D 10 10 1.0 1.0 Va Va 0.8 — 1.0 Va Room temperature
variable amount (total volume should be 100 µl). undiluted or diluted template DNA based on results obtained in step 6.
bUse
An alternative to ethidium bromide, SYBR Gold Nucleic Acid Gel Stain (Molecular Probes), is 25 to 100 times more sensitive than ethidium bromide, is more convenient to use, and permits optimization of 10- to 100-fold lower starting template copy number.
6. Examine the stained gel to determine which condition resulted in the greatest amount of product. Minor, nonspecific products may be present even under optimal conditions.
7. To ensure that the major product is the correct one, digest an aliquot of the reaction with a restriction endonuclease known to cut within the PCR product. Check buffer compatibility for the restriction endonuclease of choice. If necessary, add Na+ or precipitate in ethanol and resuspend in the appropriate buffer. Electrophorese the digestion product on a gel to verify that the resulting fragments have the expected sizes. Alternatively, transfer the PCR products to a nitrocellulose or nylon filter and hybridize with an oligonucleotide derived from the sequence internal to the primers. With appropriately stringent hybridization and washing conditions, only the correct product (and possibly some minor related products) should hybridize.
Optimize the first cycle These optional steps optimize initial hybridization and may improve efficiency and yield. They are used when primer-dimers and other nonspecific products are detected, when there is only a very small amount of starting template, or when a rare sequence is to be amplified from a complex mixture. For an optimal reaction, polymerization during the initial denaturation and annealing steps should be prevented. Taq DNA polymerase activity can be inhibited by temperature (reaction B), physical separation (reaction C), or reversible antibody binding (reaction D). PCR without hot start is performed for comparison (reaction A). 8. Prepare four reaction mixtures using the optimal MgCl2 concentration and additive requirement determined in step 6. Prepare the mixes according to the recipes in Table A.4J.2. Use the following variations for addition of Taq polymerase. a. Prepare reactions A and C at room temperature.
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b. Chill all components of reaction B in an ice slurry before they are combined. c. For reaction D, combine 1.0 µl TaqStart antibody with 4.0 µl of the dilution buffer provided with the antibody, add 1.0 µl Taq DNA polymerase (for 1:4:1 mixture of these components), mix, and incubate 5 to 10 min at room temperature before adding to reaction mixture D (glycerol and PMPE are compatible with TaqStart antibody but DMSO will interfere with antibody binding). To ensure that the reaction does not plateau and thereby obfuscate the results, use the smallest amount of template DNA necessary for visualization of the PCR product by ethidium bromide staining. Use the results from step 6 to decide how much template to use. If the desired product stains intensely, dilute the starting material as much as 1/100. If only a faint signal is apparent, use undiluted sample. Commercial optimization systems are outlined in Table A.4J.3.
9. Overlay each reaction mixture with 50 to 100 µl mineral oil. 10. Heat all reactions 5 min at 94°C. It is most convenient to use the automated thermal cycler for this step and then initiate the cycling program directly.
11. Cool the reactions to the appropriate annealing temperature as determined in step 4. Add 0.5 µl Taq DNA polymerase to reaction C, making sure the pipet tip is inserted through the layer of mineral oil into the reaction mix. Time is also an important factor in this step. If the temperature drops below the annealing temperature and is allowed to remain low, nonspecific annealing will occur. Taq DNA polymerase retains some activity even at room temperature.
12. Begin amplification of all four reactions at once, using the same cycling parameters as before. 13. Analyze the PCR products on an agarose gel and evaluate the results as in steps 5 and 6. 14. Prepare a batch of the optimized reaction mixture, but omit Taq DNA polymerase, TaqStart antibody, PMPE, and 4dNTP mix—these ingredients should be added fresh just prior to use. If desired, add Ficoll 400 to a final concentration of 0.5% to 1% (v/v) and tartrazine to a final concentration of 1 mM. Adding Ficoll 400 and tartrazine dye to the reaction mix precludes the need for a gel loading buffer and permits direct application of PCR products to agarose or acrylamide gels. At these concentrations, Ficoll 400 and tartrazine do not decrease PCR efficiency and do not interfere with PMPE or TaqStart antibodies. Other dyes, such as bromphenol blue and xylene cyanol, do inhibit PCR. Tartrazine is a yellow dye and is not as easily visualized as other dyes; this may make gel loading more difficult. Ficoll 400 and tartrazine dye may be prepared as 10× stocks and stored indefinitely at room temperature.
The Polymerase Chain Reaction
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Table A.4J.3
PCR Optimization Products
Optimization goal
Supplier
Product
Optimization support Optimization support
Perkin-Elmer Promega
Optimization kits
Boehringer-Mannheim, Invitrogen, Stratagene, Sigma, Epicentre Technologies, Life Technologies Amersham Pharmacia Biotech
Technical information in appendix to catalog PCR troubleshooting program on the Internet: http://www.promega.com/amplification/assistant Several buffers, Mg2+, and enhancers which may include DMSO, glycerol, formamide, (NH4)2SO4, and other unspecified or proprietary agents
Quick startup
Quick startup
Fisher
Quick startup
Life Technologies
Quick startup
Marsh Biomedical
Hot-start/physical barrier Fisher, Life Technologies
Hot-start/separate MgCl2 Invitrogen Hot-start/separate MgCl2 Stratagene Hot Start/separate polymerase
Promega
Hot-start/reversible inactivation of polymerase by antibody binding Hot-start/antibody binding Hot-start/antibody binding Hot-start/reversible chemical modification Hot-start/reversible chemical modification Enhancer
Clontech
Ready-To-Go Beads “optimized for standard PCR” and Ready-To-Go RAPD Analysis Beads (buffer, nucleotides, Taq DNA polymerase) EasyStart PCR Mix-in-a-Tube—tubes prepackaged with wax beads containing buffer, MgCl2, nucleotides, Taq DNA polymerase PCR SuperMix—1.1× conc.—premix containing buffer, MgCl2, nucleotides, Taq DNA polymerase Advanced Biochemicals Red Hot DNA Polymerase—a new rival for Taq polymerase with convenience features Molecular Bio-Products HotStart Storage and Reaction Tubes—preadhered wax bead in each tube; requires manual addition of one component at high temperature HotWax Mg2+ beads—wax beads contain preformulated MgCl2 which is released at first elevated-temperature step StrataSphere Magnesium Wax Beads—wax beads containing preformulated Mg2+ TaqBead Hot Start Polymerase—wax beads encapsulating Taq DNA polymerase which is released at first elevated-temperature step TaqStart Antibody, TthStart Antibody—reversibly inactivate Taq and Tth DNA polymerases until first denaturation at 95°C
Life Technologies
PlatinumTaq—contains PlatinumTaq antibody
Sigma
JumpStart Taq—contains TaqStart antibody
Perkin-Elmer
AmpliTaq Gold—activated at high temperature
Qiagen
HotStarTaq DNA Polymerase—activated at high temperature Tth pyrophosphatase, thermostable
Enhancer Enhancer
Boehringer Mannheim, New England Biolabs Clontech CPG
Enhancer
Fisher
Enhancer Enhancer Enhancer Enhancer Enhancer
Life Technologies Promega Qiagen Stratagene Stratagene
GC-Melt (in Advantage-GC Kits)—proprietary Taq-FORCE Amplification System and MIGHTY Buffer—proprietary Eppendorf MasterTaq Kit with TaqMaster Enhancer—proprietary PCRx Enhancer System—proprietary E.coli Single Stranded Binding Protein (SSB) Q-Solution—proprietary Perfect Match Polymerase Enhancer—proprietary TaqExtender PCR Additive—proprietary
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REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Enhancer agents 5× stocks: 25% acetamide (20 µl/reaction; 5% final) 5 M N,N,N-trimethylglycine (betaine; 20 µl/reaction; 1 M final) 40% polyethylene glycol (PEG) 8000 (20 µl/reaction; 8% final) 10× stocks: Glycerol (concentrated; 10 µl/reaction; 10% final) 20× stocks: Dimethylsulfoxide (DMSO; concentrated; 5 µl/reaction; 5% final) Formamide (concentrated; 5 µl/reaction; 5% final) 100× stocks: 1 U/µl Perfect Match Polymerase Enhancer (Strategene; 1 µl/reaction; 1 U final) 10 mg/ml acetylated bovine serum albumin (BSA) or gelatin (1 µl/reaction; 10 µg/ml final) 1 to 5 U/µl thermostable pyrophosphatase (PPase; Roche Diagnostics; ; 1 µl/reaction; 1 to 5 U final) 5 M tetramethylammonium chloride (TMAC; betaine hydrochloride; 1 µl/reaction; 50 mM final) 0.5 mg/ml E. coli single-stranded DNA-binding protein (SSB; Sigma; 1 µl/reaction; 5 µg/ml final) 0.5 mg/ml Gene 32 protein (Amersham Pharmacia Biotech; 1 µl/reaction; 5 µg/ml final) 10% Tween 20, Triton X-100, or Nonidet P-40 (1 µl/reaction; 0.1% final) 1 M (NH4)2SO4 (1 µl/reaction; 10 mM final; use with thermostable DNA polymerases other than Taq) MgCl2-free PCR buffer, 10× 500 mM KCl 100 mM Tris⋅Cl, pH 9.0 (at 25°C; see APPENDIX 2E) 0.1% Triton X-100 Store indefinitely at −20°C This buffer can be obtained from Promega; it is supplied with Taq DNA polymerase.
4dNTP mix For 2 mM 4dNTP mix: Prepare 2 mM each dNTP in TE buffer, pH 7.5 (APPENDIX 2E). Store up to 1 year at −20°C in 1-ml aliquots. For 25 mM 4dNTP mix: Combine equal volumes of 100 mM dNTPs (Promega). Store indefinitely at −20°C in 1-ml aliquots. Literature Cited Saiki, R.K., Gelfand, D.H., Stoffel, S., Scharf, S.J., Higuchi, R., Horn, G.T., Mullis, K.B., and Erlich, H.A. 1988. Primer-directed enzymatic amplification of DNA with a thermostable DNA polymerase. Science 239:487-491.
The Polymerase Chain Reaction
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Quantitation of Nucleic Acids with Absorption Spectroscopy
APPENDIX 4K
Reliable quantitation of nanogram and microgram amounts of DNA and RNA in solution is essential to researchers in molecular biology. In addition to the traditional absorbance measurements at 260 nm (A260), two more sensitive fluorescence techniques are presented below. These three procedures cover a range from 5 to 10 ng/ml DNA to 50 µg/ml DNA (see Table A.4K.1). Absorbance measurements are straightforward as long as any contribution from contaminants and the buffer components is taken into account. Fluorescence assays are less prone to interference than A260 measurements and are also simple to perform. As with absorbance measurement, a reading from the reagent blank is taken prior to adding the DNA. In instruments where the readout can be set to indicate concentration, a known concentration is used for calibration and subsequent readings are taken in ng/ml or µg/ml DNA. Table A.4K.1 Properties of Absorbance and Fluorescence Spectrophotometric Assays for DNA and RNA
Property Sensitivity (µg/ml) DNA RNA Ratio of signal (DNA/RNA)
Absorbance (A260)
Fluorescence H33258 EtBr
1-50 1-40
0.01-15 n.a.
0.1-10 0.2-10
0.8
400
2.2
DETECTION OF NUCLEIC ACIDS USING ABSORPTION SPECTROSCOPY Absorption of the sample is measured at several different wavelengths to assess purity and concentration of nucleic acids. A260 measurements are quantitative for relatively pure nucleic acid preparations in microgram quantities. Absorbance readings cannot discriminate between DNA and RNA; however, the ratio of A at 260 and 280 nm can be used as an indicator of nucleic acid purity. Proteins, for example, have a peak absorption at 280 nm that will reduce the A260/A280 ratio. Absorbance at 325 nm indicates particulates in the solution or dirty cuvettes; contaminants containing peptide bonds or aromatic moieties such as protein and phenol absorb at 230 nm.
BASIC PROTOCOL
This protocol is designed for a single-beam ultraviolet to visible range (UV-VIS) spectrophotometer. If available, a double-beam spectrophotometer will simplify the measurements, as it will automatically compare the cuvette holding the sample solution to a reference cuvette that contains the blank. In addition, more sophisticated double-beam instruments will scan various wavelengths and report the results automatically. Materials 1× TNE buffer: 0.1 M Tris base/10 mM EDTA/2.0 M NaCl (adjust to pH 7.4 with concentrated HCl) DNA sample to be quantitated Calf thymus DNA standard solutions (Sigma, Hoefer, Pharmacia Biotech) Matched quartz semi-micro spectrophotometer cuvettes (1-cm pathlength) Single- or dual-beam spectrophotometer (ultraviolet to visible) Contributed by Sean Gallagher Current Protocols in Protein Science (1998) A.4K.1-A.4K.3 Copyright © 1998 by John Wiley & Sons, Inc.
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A.4K.1 Supplement 13
1. Pipet 1.0 ml of 1× TNE buffer into a quartz cuvette. Place the cuvette in a single- or dual-beam spectrophotometer, read at 325 nm (note contribution of the blank relative to distilled water if necessary), and zero the instrument. Use this blank solution as the reference in double-beam instruments. For single-beam spectrophotometers, remove blank cuvette and insert cuvette containing DNA sample or standard suspended in the same solution as the blank. Take reading. Repeat this process at 280, 260, and 230 nm. It is important that the DNA be suspended in the same solution as the blank.
2. To determine the concentration (C) of DNA present, use the A260 reading in conjunction with one of the following equations: Single-stranded DNA: C ( pmol /µl) = C (µg / ml) =
A260 10 × S
A260 0.027
Double-stranded DNA: C ( pmol /µl) = C (µg / ml ) =
A260 13.2 × S
A260 0.020
Single-stranded RNA: C (µg / ml) =
A260 0.025
Oligonucleotide: C ( pmol /µl) = A260 ×
100 1.5 NA + 0.71NC + 1.20 NG + 0.84 NT
where S represents the size of the DNA in kilobases and N is the number or residues of base A, G, C, or T. For double- or single-stranded DNA and single-stranded RNA, these equations assume a 1-cm-pathlength spectrophotometer cuvette and neutral pH. The calculations are based on the Lambert-Beer law, A = ECl, where A is the absorbance at a particular wavelength, C is the concentration of DNA, l is the pathlength of the spectrophotometer cuvette (typically 1 cm), and E is the extinction coefficient. For solution concentrations given in mol/liter and a cuvette of 1-cm pathlength, E is the molar extinction coefficient and has units of M−1 cm−1. If concentration units of ìg/ml are used, then E is the specific absorption coefficient and has units of (ìg/ml)−1cm−1. The values of E used here are as follows: ssDNA, 0.027 (ìg/ml)−1cm−1; dsDNA, 0.020 (ìg/ml)−1cm−1; ssRNA, 0.025 (ìg/ml)−1cm−1. Using these calculations, an A260 of 1.0 indicates 50 ìg/ml double-stranded DNA, ∼37 ìg/ml singlestranded DNA, or ∼40 ìg/ml single- stranded RNA (adapted from Applied Biosystems, 1989).
Quantitation of Nucleic Acids with Absorption Spectroscopy
For oligonucleotides: Concentrations are calculated in the more convenient units of pmol/ìl. The base composition of the oligonucleotide has significant effects on absorbance, because the total absorbance is the sum of the individual contributions of each base (Table A.4K.2).
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Table A.4K.2 DNA Basesa
Molar Extinction Coefficients of
Base
ε1M 260 nm
Adenine Cytosine Guanosine Thymine
15,200 7,050 12,010 8,400
aMeasured
at 260 nm; see Wallace and Miyada, 1987.
3. Use the A260/A280 ratio and readings at A230 and A325 to estimate the purity of the nucleic acid sample. Ratios of 1.8 to 1.9 and 1.9 to 2.0 indicate highly purified preparations of DNA and RNA, respectively. Contaminants that absorb at 280 nm (e.g., protein) will lower this ratio. Absorbance at 230 nm reflects contamination of the sample by phenol or urea, whereas absorbance at 325 nm suggests contamination by particulates and dirty cuvettes. Light scatter at 325 nm can be magnified 5-fold at 260 nm (K. Hardy, pers. comm.). Typical values at the four wavelengths for a highly purified preparation are shown in Table A.4K.3.
Table A.4K.3
Spectrophotometric Measurements of Purified DNAa
Wavelength (nm)
Absorbance
A260/A280
Conc. (µg/ml)
325 280 260 230
0.01 0.28 0.56 0.30
— — 2.0 —
— — 28 —
aTypical
absorbancy readings of highly purified calf thymus DNA suspended in 1× TNE buffer. The concentration of DNA was nominally 25 µg/ml.
LITERATURE CITED Applied Biosystems. 1984. Evaluation and Purification of Synthetic Oligonucleotides. User Bulletin, Issue #13. Applied Biosystems, Foster City, Ca. Wallace, R.B. and Miyada, C.G. 1987. Oligonucleotide probes for the screening of recombinant DNA libraries. Methods Enzymol. 152:432-442.
Contributed by Sean Gallagher Motorola Corporation Tempe, Arizona
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Supplement 13
Growth and Manipulation of Yeast
APPENDIX 4L
PREPARATION OF SELECTED YEAST MEDIA Like Escherichia coli, yeast can be grown in either liquid media or on the surface of (or embedded in) solid agar plates. Yeast cells grow well on a minimal medium containing dextrose (glucose) as a carbon source and salts which supply nitrogen, phosphorus, and trace metals. Yeast cells grow much more rapidly, however, in the presence of protein and yeast cell extract hydrolysates, which provide amino acids, nucleotide precursors, vitamins, and other metabolites which the cells would normally synthesize de novo. During exponential or log-phase growth, yeast cells divide every 90 min when grown in such media. Preparation of sterile media of consistently high quality is essential for the genetic manipulation of yeast. Autoclaving is usually carried out for 15 min at 15 lb/in2, but times should be increased when large amounts of media are being prepared (20 min for 4 to 6 liters and 25 min for 6 to 12 liters). Liquid Media Ingredients for liquid media are dissolved in water to 1 liter, mixed until completely dissolved, and autoclaved in 100- or 500-ml media bottles. Alternatively, liquid media can be filter sterilized, resulting in faster preparation, less carmelization (of dextrose), and faster growth of cells. Recipes for “premixes” are provided to minimize the number of materials that must be weighed each time medium is prepared. When preparing premixes, break up any large chunks of dextrose before mixing with other components, then shake the container vigorously until contents are homogenized. It is convenient to make the premix in the empty plastic containers in which 2.5 kg of dextrose is packaged. Throughout this chapter, YNB −AA/AS refers to yeast nitrogen base without amino acids or ammonium sulfate. (See listing of recommended suppliers of ingredients in unit introduction above.) Rich medium YPD medium (YEPD medium) Per liter: 10 g yeast extract 20 g peptone 20 g dextrose
Premix: 250 g yeast extract 500 g peptone 500 g dextrose Use 50 g/liter
Final concentration: 1% yeast extract 2% peptone 2% dextrose
It is preferable to use a 20% (10×) solution of dextrose that has been filter sterilized or autoclaved separately (and added to the other ingredients after autoclaving) to prevent darkening of the media and to promote optimal growth.
Minimal media Minimal medium [synthetic dextrose (SD) medium] Per liter: Premix: 1.7 g YNB −AA/AS 68 g YNB −AA/AS 5 g (NH4)2SO4 200 g (NH4)2SO4 20 g dextrose 800 g dextrose Use 27 g/liter
Final concentration: 0.17% YNB −AA/AS 0.5% (NH4)2SO4 2% dextrose Molecular Biology Techniques
Contributed by Douglas A. Treco, Ann Reynolds, and Victoria Lundblad Current Protocols in Protein Science (1998) A.4L.1-A.4L.6 Copyright © 1998 by John Wiley & Sons, Inc.
A.4L.1 Supplement 14
This minimal medium can support the growth of yeast which have no nutritional requirements. However, it is used most often as a basal medium to which other supplements are added (see CM dropout medium below).
Complete minimal (CM) dropout medium, per liter 1.3 g dropout powder (Table A.4L.1) 1.7 g YNB −AA/AS 5 g (NH4)2SO4 20 g dextrose Alternatively, replace last three ingredients with 27 g minimal (SD) medium premix. CM dropout powder, also known as minus or omission powder, lacks a single nutrient but contains the other nutrients listed in Table A.4L.1. Complete minimal (CM) dropout medium is used to test for genes involved in biosynthetic pathways and to select for gene function in transformation experiments. To test for a gene involved in histidine biosynthesis, determine if the yeast strain in question can grow on CM minus histidine (−His) or “histidine dropout” plates. It is convenient to make several dropout powders, each lacking a single nutrient, to avoid weighing each component separately for all the different dropout plates required in the laboratory. It may be preferable to use a 10× solution of dropout powder (i.e., 13 g of dropout powder in 100 ml water) that has been “sterilized” separately (and added to the other ingredients after autoclaving) to improve the growth rate in this medium.
Table A4.L.1
Nutrient Concentrations for Dropout Powdersa
Nutrientb Adenine (hemisulfate salt) L-arginine (HCl) L-aspartic acide L-glutamic acid (monosodium salt) L-histidine L-leucinef L-lysine (mono-HCl) L-methionine L-phenylalanine L-serine L-threoninee L-tryptophan L-tyrosine L-valine Uracil
Amount in dropout powder (g)c
Final conc. in prepared media (µg/ml)
Liquid stock conc. (mg/100 ml)d
2.5 1.2 6.0 6.0 1.2 3.6 1.8 1.2 3.0 22.5 12.0 2.4 1.8 9.0 1.2
40 20 100 100 20 60 30 20 50 375 200 40 30 150 20
500 240 1200 1200 240 720 360 240 600 4500 2400 480 180f 1800 240
aCM dropout powder lacks a single nutrient but contains the other nutrients listed in this table. Nomenclature
Growth and Manipulation of Yeast
in this manual refers to, e.g., a preparation that omits histidine as histidine dropout powder or CM medium −His. Conditions from Sherman et al. (1979). bAmino acids not listed here can be added to a final concentration of 40 µg/ml (40 mg/liter). cGrind powders into a homogeneous mixture with a clean, dry mortar and pestle. Store in a clean, dry bottle or a covered flask. dUse 8.3 ml/liter of each stock for special nutritional requirements. Store adenine, aspartic acid, glutamic acid, leucine, phenylalanine, tyrosine, and uracil solutions at room temperature. All others should be stored at 4°C. eAlthough these amino acids can be used reliably when included in autoclaved media, they supplement growth better when added after autoclaving. fUse 16.6 ml/liter for L-tyrosine nutritional requirement.
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Solid Media For all plates, agar is added at a concentration of 2% (20 g/liter). A pellet of sodium hydroxide (∼0.1 g) should be added per liter to raise the pH enough to prevent agar breakdown during autoclaving. In addition, add a stir bar to facilitate mixing after autoclaving. After autoclaving, flasks are left for 45 to 60 min at room temperature until cooled to 50° to 60°C. (Drugs and other nutrients are added after 30 min at room temperature.) Just prior to pouring, put the flask on a stir plate at medium to high speed and mix until contents are homogeneous (∼5 min). After pouring, a few bubbles can be removed from the agar surface by passing the flame of a Bunsen burner lightly over the surface of the molten agar (“flaming” the plates). One liter of medium will yield 30 to 35 plates. Most plates can be stored at room temperature for ≤4 months. Plates containing drugs (cycloheximide, 5-fluoroorotic, canavanine, and L-α-aminoadipic acid) or 5-bromo-4chloro-3-indolyl-β-D-galactosidase (Xgal; 20 µg/ml; APPENDIX 4A) are stable for 2 to 3 months when stored at 4°C. Minimal plates and rich plates YPD, minimal, and CM dropout plates Per liter: Follow recipes for liquid medium above, adding 20 g agar and a pellet of NaOH. Premixes: Follow recipes for liquid medium premixes above, adding 500 g agar for YPD premix and 800 g agar for minimal premix. To prepare plates, add one NaOH pellet and the following amounts of premix (per liter): YPD plates—70 g YPD plate premix Minimal plates—47 g minimal plate premix CM dropout plates—47 g minimal plate premix + 1.3 g dropout powder Specialty plates Xgal plates, per liter 1.7 g YNB −AA/AS 5 g (NH4)2SO4 20 g dextrose 20 g agar 0.8 g dropout powder (omitting appropriate amino acids; see Table A4.L.1) NaOH pellet (Alternatively, replace first four ingredients with 47 g minimal plate premix.) Add H2O to 900 ml and autoclave. Add 100 ml of 0.7 M potassium phosphate, pH 7.0, and 2 ml of 20 mg/ml Xgal prepared in 100% N,N-dimethylformamide (stored as frozen stock). STRAIN STORAGE AND REVIVAL Yeast strains can be stored at −70°C in 15% glycerol (viable for >5 years), or at 4°C on slants consisting of rich medium supplemented with potato starch (viable for 1 to 2 years). Preparation and Inoculation of Frozen Stocks Make a solution of 30% (w/v) glycerol. Pipet 1 ml into 15 × 45–mm, 4-ml screw-cap vials. Loosely cap the vials and autoclave 15 min. To inoculate vials for storage, add 1 ml of a late-log or early-stationary phase culture, mix, and set on dry ice. Store at −70°C. Revive by scraping some of the cells off the frozen
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surface and streak onto plates. Do not thaw the entire vial. Cells can also be stored in the same way by adding 80 µl dimethyl sulfoxide (DMSO) to 1 ml cells (8% v/v) and storing at −70°C. BASIC PROTOCOL 1
ASSAY FOR â-GALACTOSIDASE IN LIQUID CULTURES Materials YPD medium (see recipe under Liquid Media) Z buffer (see recipe in Reagents and Solutions) 0.1% sodium dodecyl sulfate (SDS) Chloroform 4 mg/ml O-nitrophenyl-β-D-galactoside (ONPG) in 0.1 M potassium phosphate, pH 7.0 (see APPENDIX 2E for buffer; filter sterilize and store frozen) 1 M Na2CO3 30°C water bath 1. Inoculate 5 ml YPD (or appropriate) medium with single yeast colony. Grow 2 to 3 independent single-colony cultures of a strain containing lacZ fusion protein overnight at 30°C. If fusion is on a plasmid, grow cells in medium that selects for plasmid. 2. Inoculate 5 ml YPD medium (or appropriate selective medium and/or inducing conditions) with 20 to 50 µ1 of each overnight culture. Grow to mid- or late-log phase: 0.5–1 × 108 cells/ml (OD600 = 2.0) for rich medium or 2–5 × 107 cells/ml (OD600 = 0.5 to 1.0) for minimal medium. If goal is to investigate activity of yeast promoter under conditions that induce its expression, grow cells in parallel under inducing and noninducing conditions.
3. Centrifuge cells 5 min at 1100 × g. Resuspend in equal volume of Z buffer and place on ice. Determine OD600 for each sample. If anticipated level of β-galactosidase activity is low, it may be necessary to concentrate cells. For cells demonstrating 100 to 1000 U of activity within 30 min to 4 hr, concentration is not necessary.
4. Set up the following two reaction tubes for each sample (1 ml each), with mixing: (a) 100 µl cells with 900 µl Z buffer and (b) 50 µl cells with 950 µl Z buffer. 5. Add 1 drop of 0.1% SDS and 2 drops chloroform to each sample using a Pasteur pipet. Vortex 10 to 15 sec and equilibrate 15 min in 30°C water bath. 6. Add 0.2 ml of 4 mg/ml ONPG; vortex 5 sec. Place in 30°C water bath and begin timing. When medium-yellow color (OD420 = 0.3 to 0.7) has developed, add 0.5 ml of 1 M Na2CO3 and note time. 7. Centrifuge cells 5 min at 1100 × g. Determine OD420 and OD550 of supernatant. If the cell debris has been well pelleted, the OD550 is usually zero and therefore is not necessary to read.
8. Calculate units with the following equation: U=
Growth and Manipulation of Yeast
1000 × [(OD 420 ) − (1.75 × OD 550 )] (t ) × (v ) × (OD 600 )
where t = time of reaction (min); v = volume of culture used in assay (ml); OD600 = cell density at the start of the assay; OD420 = combination of absorbance by o-ni-
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trophenol and light scattering by cell debris; and OD550 = light scattering by cell debris. TRANSFORMATION USING LITHIUM ACETATE This is the most commonly used method for yeast transformation. It is reasonably fast and provides transformation efficiencies of 105 to 106 transformations/µg.
BASIC PROTOCOL 2
NOTE: All solutions and glassware coming into contact with yeast cells must be sterile. Traces of soap on glassware may decrease the transformation efficiency. In addition, the water used for washes and in solution preparation must be of the highest quality. Materials YPD medium (see recipe under Liquid Media) Yeast strain to be transformed YPAD medium: YPD medium supplemented with 30 mg/liter adenine hemisulfate Highest-quality sterile H2O 10× TE buffer, pH 7.5 (see APPENDIX 2E for 1× recipe), sterile 10× lithium acetate stock solution: 1 M lithium acetate, pH 7.5 (adjust pH with dilute acetic acid), filter sterilized DNA: high-molecular-weight, single-stranded carrier DNA and transforming DNA 50% (w/v) polyethylene glycol (PEG 4000 or 3350 (do not use PEG 8000), filter sterilized CM dropout plates (see recipe under Solid Media) prepared with Difco agar 30°C incubator with shaker Sorvall GSA and SS-34 rotors (or equivalents) 42°C water bath 1. Two days before the experiment, inoculate 5 ml YPD medium with a single yeast colony of the strain to be transformed. Grow overnight to saturation at 30°C. 2. The night before transformation, inoculate a 1-liter sterile flask containing 300 ml YPAD medium with an appropriate amount of the saturated culture and grow overnight at 30°C to 1 × 107 cells/ml (OD600 ≅ 0.3 to 0.5, depending on strain). For 2- to 3-fold higher efficiency, dilute at this point to 2 × 106 cells/ml in fresh YPAD medium and grow for another 1 to 2 generations (2 to 4 hr). Because growth phase and cell density are important in achieving the highest transformation efficiency, the easiest approach is to inoculate three independent flasks with varying amounts of the saturated overnight culture (try 1, 5, and 25 ìl for a 12-hr growth period), then check the OD600 of each flask the next day.
3. Harvest cells by centrifuging 5 min at 4000 × g, room temperature. Resuspend in 10 ml highest-quality sterile water. Transfer to a smaller centrifuge tube and pellet cells by centrifuging 5 min at 5000 to 6000 × g, room temperature. 4. Resuspend in 1.5 ml buffered lithium solution, freshly prepared as follows: 1 vol 10× TE buffer, pH 7.5 1 vol 10× lithium acetate stock solution 8 vol sterile water. 5. For each transformation, mix 200 µg carrier DNA with ≤5 µg transforming DNA in a sterile 1.5-ml microcentrifuge tube. Keep total volume of DNA ≤20 µl. Maximal transformation efficiency is achieved by repeating the denaturation cycle (boiling and chilling) of carrier DNA immediately prior to use.
6. Add 200 µl yeast suspension to each microcentrifuge tube. Add 1.2 ml PEG solution, freshly prepared as follows:
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8 vol 50% PEG 1 vol 10× TE buffer, pH 7.5 1 vol 10× lithium acetate stock solution. Shake 30 min at 30°C. 7. Heat shock exactly 15 min at 42°C. Microcentrifuge 5 sec at room temperature. Resuspend yeast in 200 µl to 1 ml of 1× TE buffer (freshly prepared from 10× stock) and spread up to 200 µl onto CM dropout plates made with Difco agar. Incubate 2 to 5 days at 30°C until transformants appear. REAGENTS AND SOLUTIONS Use Milli-Q water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.
Z buffer 16.1 g Na2HPO4⋅7H2O (60 mM final) 5.5 g NaH2PO4⋅H2O (40 mM final) 0.75 g KC1 (10 mM final) 0.246 g MgSO4⋅7H2O (1 mM final) 2.7 ml 2-mercaptoethanol (50 mM final) Adjust to pH 7.0 and bring to 1 liter with H2O Do not autoclave LITERATURE CITED Sherman, F., Fink, G.R., and Lawrence, C.W. 1979. Methods in Yeast Genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y.
Contributed by Douglas A. Treco Massachusetts General Hospital Boston, Massachusetts Ann Reynolds University of Washington Seattle, Washington Victoria Lundblad Baylor College of Medicine Houston, Texas
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BIOPHYSICAL METHODS: DATA ANALYSIS
APPENDIX 5
Theoretical Aspects of the Quantitative Characterization of Ligand Binding
APPENDIX 5A
Living organisms grow, differentiate, reproduce, and respond to their environment via specific and integrated interactions between biomolecules. The investigation of molecular interactions therefore constitutes a major area of biochemical study, occupying a ubiquitous and central position between molecular physiology on the one hand and structural chemistry on the other. While specificity resides in the details of structural recognition, the dynamic interplay between biomolecules is precisely orchestrated by the thermodynamics of the biomolecular equilibria involved. A consequence is that biochemists of all persuasions are drawn into the quantitative characterization of biomolecular interactions. Fortunately, a common set of physicochemical principles applies to all such phenomena, irrespective of whether the interaction of interest involves an enzyme and its substrate or inhibitor, a hormone or growth factor and its receptor, an antibody and its antigen, or, indeed, the binding of effector molecules that modulate these interactions. The binding affinity, binding specificity, number of binding sites per molecule, as well as the enthalpic and entropic contributions to the binding energy are common parameters that assist an understanding of the biochemical outcome. This unit aims to provide an overview of the design and interpretation of binding experiments. More extensive treatments are provided by two recent monographs (Winzor and Sawyer, 1995; Klotz, 1997). RECTANGULAR HYPERBOLIC BINDING RESPONSES In studies of reversible interactions between two species, the smaller solute is termed the ligand (S) and is usually considered to possess a single site for interaction with the larger species. The latter, designated here as A, is termed either the acceptor or the receptor (if particulate), which may possess several sites for interaction with ligand. For a system entailing the binding of a ligand to p sites on an acceptor, the concentrations of p complexes (AS, AS2, .., ASp) as well as those of free acceptor and free ligand are required, in principle, for complete description of the composition of an acceptor-ligand mixture. However, under favorable circumstances the problem simplifies to quantitative characterization by determining the free concentration of ligand (CS) in a mixture with defined total concentrations of acceptor and ligand (CA and CS, respectively). This simplification arises when the concentration of bound ligand (CS − CS) exhibits a rectangular hyperbolic dependence upon free ligand concentration (CS) for mixtures with a fixed total acceptor concentration (CA). Such a situation is encountered, for example, with enzymes exhibiting Michaelis-Menten behavior, where the observed dependence of initial velocity upon substrate concentration is the manifestation of a rectangular hyperbolic binding response. Although binding theory is usually derived for the general case in which a ligand binds to p sites on an acceptor, this discussion presents an alternative derivation for the simpler case where ligand binding is restricted to two acceptor sites. For this system, the various equilibria are described in terms of molar association constants (Ki) by:
Biophysical Methods: Data Analysis Contributed by William H. Sawyer and Donald J. Winzor Current Protocols in Protein Science (1999) A.5A.1-A.5A.40 Copyright © 1999 by John Wiley & Sons, Inc.
A.5A.1 Supplement 16
− A− + S
K1
¾
−A−S
CAS = K1CA CS
Equation A.5A.1
− A− + S
K2
¾
S−A−
CSA = K 2CA CS
Equation A.5A.2
− A−S + S
K3
¾
CSAS = K3CASCS = K1K3CA CS2
S−A−S
Equation A.5A.3
S−A− + S
K4
¾
CSAS = K4CSA CS = K 2 K 4CA CS2
S−A−S
Equation A.5A.4
Completely general characterization thus requires the determination of four equilibrium constants, the only interrelationship between which is identity of the products K1K3 and K2K4 to maintain independence of the concentration of saturated acceptor-ligand complex (CSAS) upon the pathway of its formation—a thermodynamic necessity because the concentration of saturated complex is either K1K3CACS2 or K2K4CACS2. The total concentrations of acceptor and ligand may therefore be written in the form:
b
g
CA = CA 1 + K1 + K2 CS + K1 K3CS2 Equation A.5A.5
b
g
CS = CS + CA K1 + K 2 CS + 2 K1 K3CS2 Equation A.5A.6
In an experimental context the binding response is usually expressed in terms of a binding function, termed r (Klotz, 1946) or ν (Scatchard, 1949) and defined as the molar ratio of the amount of bound ligand to the total amount of acceptor. On the grounds that these two amounts are contained in the same volume, the binding function for the above system is given by Equation A.5A.7, which is the binding equation in Adair (1925) format.
c
h
r = CS − CS / CA =
bK1 + K2 gCS + 2 K1K3CS2 1 + b K1 + K2 gCS + K1K3CS2
Equation A.5A.7
Quantitative Characterization of Ligand Binding
Simplification of Equation A.5A.7 to a rectangular hyperbolic dependence of the binding function upon free ligand concentration entails the incorporation of two separate assumptions: (1) binding sites on the acceptor are equivalent (K1 = K2), and (2) binding sites on the acceptor are independent (K1 = K4, K2 = K3). The former implies identity of the binding characteristics for the interaction of ligand with either site on (for example) a dimeric enzyme comprising two identical subunits, whereas the latter assumption implies that the same binding constant applies to ligand interaction with a given site, irrespective of the occupancy state of the other. These two assumptions allow the description of binding in
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terms of a rectangular hyperbolic response governed by a single intrinsic (site-binding) constant, KAS = K1 = K2 = K3 = K4. With these substitutions, Equation A.5A.7 becomes: r=
2 K ASCS 1 + KASCS
Equation A.5A.8
which is the particular form (p = 2) of the general expression for a rectangular hyperbolic binding response, namely: r=
pKASCS 1 + KASCS
Equation A.5A.9
where p denotes the number of equivalent and independent ligand-binding sites on the acceptor. Because there is also widespread use of the definition of equilibrium constants in terms of dissociation (Kd = 1/KAS), Equation A.5A.9 can also be written in the form: r=
pCS Kd + CS
Equation A.5A.10
which bears obvious analogies to the Michaelis-Menten equation. The general derivation of Equation A.5A.9 (e.g., Klotz, 1946; Winzor and Sawyer, 1995) is certainly more elegant than that presented here, but its reliance upon statistical considerations dictates the introduction of site equivalence and site independence as a combined assumption. The present treatment emphasizes that there are two separate assumptions, either of which may not be fulfilled. For example, binding sites may be independent but nonidentical—the situation encountered in the binding of protons to the amino and carboxyl groups of glycine. In addition to the possibility of acceptor-site heterogeneity, there may be sites that are equivalent (K1 = K2) but dependent (K1 ≠ K4, K2 ≠ K3)—the basis of a common concept of allostery (Koshland et al., 1966). However, violation of either assumption leads to binding curves that deviate from rectangular hyperbolic form. In Figure A.5A.1, the solid curve depicts the dependence of r upon CS under conditions where both assumptions prevail, whereas the broken lines correspond to examples of acceptor site heterogeneity (- - -) and acceptor site cooperativity with K4 > K1 (– – –). Rectangular hyperbolic responses abound in biology; in many cases the source can be traced to a single equilibrium binding event. There are situations where the parameter being monitored is the actual binding process. On the other hand, the kinetic characterization of an enzyme exhibiting Michaelis-Menten kinetic behavior provides an example where the rectangular hyperbolic dependence of the initial velocity (v) of product formation reflects the interactions of substrate with enzyme and the various enzyme-substrate complexes (ES, ES2,.., ESp−1). In present terms, the maximal velocity, V, is given by the product of the catalytic rate constant (kc) and the total enzyme concentration (CA), whereupon the initial velocity is described by the relationship Biophysical Methods: Data Analysis
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Binding function (r)
2
1
0 0
20
40
60
CS (µM)
Figure A.5A.1 Illustrative dependences of the binding function (r) upon free ligand concentration (CS) for various types of acceptor-ligand interaction. Rectangular hyperbolic dependence (_____) for an acceptor with two equivalent and independent sites (KAS = K1 = K2 = K3 = K4 = 105 M−1). Binding curve (- - -) for an acceptor with two independent but nonequivalent sites (K1 = K4 = 106 M−1; K2 = K3 = 104 M−1). Sigmoidal binding curve (– – –) for an acceptor with two equivalent but dependent sites (K1 = K2 = 104 M−1; K3 = K4 = 106 M−1). See Equations A.5A.1-A.5A.4 for equilibrium constant nomenclature.
v = kc rCA =
pkc CA CS K M + CS
Equation A.5A.11
This emphasizes the fact that the catalytic rate constant and the Michaelis constant (KM) are intrinsic parameters obtained on the basis of equivalent and independent substrate binding to the p active sites on the enzyme. QUANTITATIVE CHARACTERIZATION OF A HYPERBOLIC RESPONSE Quantitative characterization of a rectangular hyperbolic binding response entails the determination of two parameters, p and KAS, in Equation A.5A.9. In the design of a binding experiment it is therefore important to adopt an experimental strategy that will maximize the accuracy with which those two parameters may be estimated from the measured dependence of the binding function (r) upon free ligand concentration (CS). Importance of Studying a Wide Range of Ligand Concentrations
Quantitative Characterization of Ligand Binding
The desired starting point in any binding analysis is a well-defined dependence of the binding function upon CS. Under the equilibrium conditions of a binding assay, the magnitude of r approaches its maximum limiting value (p) very gradually with increasing free ligand concentration (Fig. A.5A.1). A wide range of CS values is therefore required to extract accurate values of both p and KAS from the form of the equilibrium binding curve. Whereas a linear scale in free ligand concentration crowds data at low CS values because of substantial changes in r for small changes in free ligand concentration, a logarithmic scale in CS turns the plot into a sigmoidal curve that is symmetrical about an
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inflection point at r = p/2. For rectangular hyperbolic binding, this inflection point provides an estimate of Kd = 1/KAS. A change by one log unit (base 10) of CS on each side of this inflection point covers a range of 10% to 90% acceptor saturation. Consequently, the dependence of r upon log10CS is an extremely useful means of assessing whether a sufficient range in r has been covered in the binding assay to allow accurate extraction of values of p and KAS (Thompson and Klotz, 1971). The restriction of measurements to a narrow range of CS, and hence of r, also renders difficult the detection of any deviation from rectangular hyperbolic behavior. In situations where either the solubility or insufficiency of ligand precludes measurements of the upper portion of the binding curve, advantage may sometimes be taken of a change to experimental conditions that ensure essentially stoichiometric binding of ligand to acceptor. Such selection of conditions where the binding constant is sufficiently great for the acceptor-ligand mixtures to comprise complex and excess reactant (acceptor or ligand), the binding function (or a parameter related thereto) becomes linear in total ligand concentration to the titration endpoint, and independent of ligand concentration thereafter (Fig. A.5A.2A). Unequivocal determination of the reaction stoichiometry by this means or by use of subsidiary information, such as knowledge of the quaternary state of the acceptor, simplifies analysis of the binding data under equilibrium conditions (Fig. A.5A.2B) to the determination of a single parameter (KAS), and hence enhances the accuracy with which the binding constant may be obtained from a suboptimal set of experimental results. Nonlinear Least-Squares Methods Direct fitting of untransformed data to the appropriate binding expression is the preferred method of evaluating binding parameters. Experimental binding data comprising (r, Cs) coordinate pairs can be fitted directly to Equation A.5A.9 by any of a number of nonlinear least-squares curve-fitting programs that are available in commercial software packages. Good programs allow the fitting equation to be defined by the user, and a choice of the parameters to be floated during the fitting procedure. The program should also provide estimates of the errors in the fitted parameters, as well as assessments of the goodnessof-fit in terms of chi-squared values and plots of the residuals. The first point on which the experimenter must decide is whether the data are indicative of a rectangular hyperbolic function. An objective test of the goodness-of-fit is provided by the value of chi squared. Visual inspection of the residuals plot should reveal a random scatter of experimental data about the fitted curve. Clearly, these tests require that there be sufficient data points or a sufficient sample size for statistical tests to be meaningful. For practical reasons this requirement is not met in many binding assays. However, for many spectroscopic titrations there is no impediment to collecting the required number of data points. The experimenter should also be aware of the limitations of the fitting procedure. Most programs are based on a modified Marquardt algorithm, which may have difficulty converging to a minimum chi squared when initial estimates of the parameters are far removed from those associated with the least squares minimum. Moreover, the experimental points must cover a sufficient range of acceptor saturation. For example, programs experience difficulty returning accurate values of p and KAS when experimental data are only available for a narrow range of acceptor saturation. Knowledge of the error associated with the estimates of both r and CS is also important. Comparison of experimental results with simulated data for various values of KAS (and possibly p) can be very useful for demonstrating the uniqueness of the fit obtained. Biophysical Methods: Data Analysis
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Use of Linear Transforms Before the general availability of computers, linear transforms of the rectangular hyperbolic dependence of r upon CS were used to determine values of p and KAS. By simple algebraic manipulation, Equation A.5A.9 can be transformed into the following expressions: r/CS = pK AS − rK AS Equation A.5A.12
Binding signal
A
80
40
0 0
4
8
12
8
12
CS (µM)
Binding function (r)
B
1.0
0.5
0 0
4 CS (µM)
Quantitative Characterization of Ligand Binding
Figure A.5A.2 Illustrative characterization of an acceptor-ligand interaction for a system with limited ligand solubility. (A) Estimation of the number of binding sites by titration of 3 µM acceptor with ligand under stoichiometric conditions: p = 2 on the grounds that 6 µM ligand is required to titrate 3 µM acceptor. (B) Binding data obtained (for example) by equilibrium dialysis over the limited range of ligand solubility (<10 µM), together with the best-fit rectangular hyperbolic relationship (Equation A.5A.9) for a system with p = 2 (see A).
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b
g
CS /r = 1/ pK AS + CS / p Equation A.5A.13
b
g
1/r = 1/ pK ASCS + 1/ p Equation A.5A.14
which lead to the graphical representations of data shown in Figure A.5A.3 (Scatchard plot, Hames plot, and double-reciprocal plot, respectively). An undesirable characteristic of these plots is the bias introduced by the linear transformation. In the example given in Figure A.5A.3, the data become crowded towards the abscissa in the Scatchard plot, and towards the ordinate in the Hames and double-recip-
r/CS (µM –1)
A
0.3
0.2
pKAS –KAS
0.1
p
0 0
C S /r (µM)
B
0.8 1.6 Binding function (r )
2.4
60
40 1/p 20 1/pKAS 0 0
40
80
120
0.8
1.2
C S (µM)
1/r
C
6
4 1/pKAS 2 1/p 0 0
0.4 1 /C S (µM–1)
Figure A.5A.3 Use of various linear transforms (Equations A.5A.12-A.5A.14) of the rectangular hyperbolic binding equation to characterize an acceptor-ligand interaction with p = 2 and KAS = 105 M−1. (A) Scatchard plot; (B) Hames plot; (C) double-reciprocal plot.
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rocal plots. This distortion of the primary data results in variable weighing of points in a linear regression analysis, a point addressed by Klotz (1983, 1997). The nonlinear least-squares fitting procedure circumvents this problem. Despite the superiority of nonlinear least-squares fitting procedures for determining binding parameters, the plot of binding data in accordance with one of these linear transforms can still serve an important role in the analysis. First, the linearly transformed plot can assist in the decision about the adequacy of the simplest model with binding to equivalent and independent acceptor sites. More complex modes of binding give rise to deviations from rectagular hyperbolic responses (Fig. A.5A.1), and hence from linearity in the transformed manner of presentation. It is easier for the human eye to detect deviations from linearity than subtle deviations from a curvilinear dependence. Second, the linear transform is also useful for providing initial estimates of p and KAS that are required by nonlinear least-squares fitting programs. Experimental design summary 1. Use a semilogarithmic plot of r versus log10CS to establish the adequacy of the range of acceptor saturation covered by the experimental measurements (preferably 10% to 90%). 2. Know the limitations of the experimental method used for determining binding data. Determine the respective (r, CS) coordinate pairs at least in triplicate to allow assessment of the uncertainty associated with each value of r. 3. Collect as many (r, CS) coordinate pairs as practicable to allow the effective use of residuals plots and the statistical evaluation of chi squared. Data analysis summary 1. Analysis of binding data aims, in the first instance, to identify and characterize the most appropriate binding model that is consistent with the experimental data. 2. The simplest model involves the binding of a ligand to equivalent and independent sites on a single acceptor, and results in the rectangular hyperbolic behavior described above. It is therefore the first model to be tested. 3. Deviations from rectangular hyperbolic behavior, detected by unsatisfactory curve fitting of the untransformed or transformed data to the simplest situation, indicate that more complex binding models must be considered (see Binding Responses Deviating from Rectangular Hyperbolic Form). 4. The formulation of a new binding model, together with its mathematical description, frequently requires consideration of complementary information about the acceptorligand system: whether the acceptor undergoes isomerization or self-association, whether there are noninteracting forms of the acceptor, whether the ligand is multivalent, and whether binding is enhanced or inhibited by other effector molecules that may be present. 5. Simulation of the mathematical model can be useful in guiding the design of further experiments. The process of data analysis and interpretation is depicted diagrammatically in Figure A.5A.4. In that regard, it must be appreciated that agreement of the model with the experimental data does not establish the mechanistic legitimacy of the model, but merely the consistency between the two. Quantitative Characterization of Ligand Binding
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binding experiment
experimental design
simulations
curve fitting
interpretation
MODEL
binding equations
complementary experimental data
Figure A.5A.4 Schematic representation of the strategy for the quantitative characterization of a binding response. Taken from Winzor and Sawyer (1995).
ILLUSTRATIVE ANALYSES OF EXPERIMENTAL RESULTS The quantitative characterization of ligand binding is usually envisaged in terms of procedures based on measurement of the free ligand concentration (CS) in mixtures with defined total composition (CA, CS). However, spectral studies afford the opportunity to measure the concentration of complexed ligand (CS − CS), in mixtures with defined total composition. Inasmuch as the concentrations of free and complexed ligand are both required for quantitative analysis of ligand binding, there is no theoretical advantage associated with either type of procedure. Methods Based on Measurement of Free Ligand Concentration In methods such as equilibrium dialysis (Klotz, 1946), steady-state dialysis (Colowick and Womack, 1969; Ford et al., 1984), diafiltration (Blatt et al., 1968), and ultrafiltration (Sophianopoulos et al., 1978), a membrane that is permeable to ligand but not to acceptor is used to create a two-phase system in which one phase comprises the equilibrium mixture with composition (CA, CS) and the other an acceptor-free solution of ligand at a concentration related directly to its equilibrium concentration (CS). Other methods that yield equivalent information include frontal gel chromatography on a matrix that excludes acceptor and hence all acceptor-ligand complexes (Nichol and Winzor, 1964), and liquid-liquid partition studies (Goodman, 1958). Provided that the ligand is sufficiently small that complex formation has no effect on the migration rate of the acceptor, analytical ultracentrifugation also has the potential to generate a sedimentation velocity pattern in which the slower-migrating boundary comprises free ligand at its equilibrium concentration (Steinberg and Schachman, 1966). For the first illustration of the quantitative characterization of ligand binding by measurement of the free ligand concentration in mixtures with known total composition, consider the often-studied interaction between methyl orange and bovine serum albumin. The semilogarithmic plot (Fig. A.5A.5A) summarizes the original equilibrium dialysis data (Klotz et al., 1946) as well as subsequent results obtained by sedimentation velocity (Steinberg and Schachman, 1966). The inset to Figure A.5A.5A shows the double-reciprocal transform of the equilibrium dialysis data that was used to deduce values of 22 and 2,200 M−1 for p and KAS, respectively (Klotz et al., 1946). Nonlinear least-squares analysis of the combined data sets (Fig. A.5A.5B) in terms of Equation A.5A.9 yields
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A
16
12
1.0 1/r
Binding function (r )
1.5
0.5 8 0 0 4
0.08 0.04 1/C S (µM –1)
0 –6
–5
–4
–3
400
600
B
16
Binding function (r )
Log10 C S
12
8
4
0 0
200 CS (µM)
Figure A.5A.5 Results obtained by equilibrium dialysis (filled circles) and sedimentation velocity (open circles) for the interaction of methyl orange (S) with bovine serum albumin (A). (A) Semilogarithmic representation of the results, together with the double-reciprocal transform of the equilibrium dialysis data (inset). (B) Untransformed dependence of the binding function upon free ligand concentration, together with the best-fit description by nonlinear regression analysis according to Equation A.5A.9. Equilibrium dialysis data are taken from Klotz et al. (1946), and sedimentation velocity data from Steinberg and Schachman (1966).
respective estimates (at ±2 SD) of 27 ± 6 and 1900 ± 700 M−1 for the number of sites and the intrinsic binding constant. The relatively large extent of uncertainty associated with this quantitative analysis reflects the limited range of acceptor saturation (0.7% to 52%) covered by the binding measurements. Indeed, this deficiency of the data set is already manifested in the semilogarithmic plot, which shows no sign of the sigmoidal shape that indicates the coverage of a sufficient range of acceptor saturation for accurate definition of the affinity and stoichiometry of the interaction.
Quantitative Characterization of Ligand Binding
A second system is therefore used to better illustrate the potential of nonlinear leastsquares analysis of binding data. The results of a frontal gel chromatographic study of the interaction between NADH and rabbit muscle lactate dehydrogenase (Ward and Winzor, 1983) have been supplemented with additional data to obtain the binding curve shown in Figure A.5A.6, which also establishes the sigmoidal shape of their representation in
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Residual
0.1 0 –0.1
3
4
2 r
Binding function (r )
4
2
1 0 –6
–5 Log10C S
–4
0 0
20
40
60
80
100
C S (µM)
Figure A.5A.6 Characterization of the interaction between NADH and rabbit muscle lactate dehydrogenase (pH 7.4; ionic strength, I, 0.15) by nonlinear regression analysis of the untransformed gel chromatographic binding data in terms of Equation A.5A.9: the upper pattern shows random scatter in the residuals plot. The inset displays the same data in semilogarithmic format. Data taken in part from Ward and Winzor (1983).
semilogarithmic format (inset). The lines indicate the best-fit description obtained by nonlinear least-squares analysis, where p = 4.01 ± 0.04 and KAS = 1.49 (± 0.08) × 105 M−1, about which the experimental points exhibit random scatter (residuals plot in the upper panel). Inasmuch as the range of acceptor saturation ranges between 14.5% and 92.5%, these results exemplify the recommended experimental and analytical protocols that concluded the previous section. Methods Based on Measurement of Complexed Ligand or Acceptor Emphasis is now turned to binding procedures that rely upon the direct measurement of complex formation rather than on its deduction from knowledge of the total and free ligand concentrations. Because the extent of complex formation can be monitored by a response that emanates from either the acceptor or the ligand, there are two situations to consider. A spectral signal arising from the acceptor There are many examples in which a property of an acceptor is perturbed upon binding of a ligand. The property may be a biological activity, as in the case of an enzyme binding a substrate, or a biological response triggered by the occupation of a cell surface receptor. More commonly, a spectral property of the acceptor is changed upon ligand binding, thus providing a convenient method for following the binding reaction when the ligand is titrated into a solution of acceptor. The spectral change in the acceptor is a measure of the concentration of the ligand-acceptor complex (CS − CS). Thus, the concentration of free
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ligand must be found by difference between the total ligand concentration (Cs) and that which is bound, (CS − CS). Besides providing a means of following the binding equilibrium, the spectral change may also provide information about the chemical groups in the acceptor that are located at the acceptor-ligand interface. The use of spectroscopic techniques to monitor binding equilibria requires careful attention to spectroscopic purity. Ideally, the spectroscopic signal should arise only from the acceptor species, so that the signal from the acceptor alone can be used to determine its concentration and ultimately the stoichiometry of the binding reaction. If the spectroscopic signal arises from two or more species, only one of which binds the ligand, the spectral change is overlaid on the background signal from the nonbinding species. The titration is still valid provided that the concentration of the binding form is accurately known. Particular problems arise when an acceptor is labeled with an extrinsic fluorescent probe. Labeling is rarely 100% efficient, and the labeled and unlabeled material may differ in their affinity for the ligand. Much time can be saved by separating the labeled from the unlabeled material before conducting binding experiments. Small amounts of background signal (e.g., solvent fluorescence or scattered excitation light in the case of fluorescence spectroscopy) can be dealt with by using appropriate controls and background subtraction. The ultimate aim of biochemical study is to explain in vivo biochemistry at the molecular level. Consequently, the biochemist frequently proceeds from a controlled reductionist environment to a complex in vivo environment in which experimental rigor may be at least partially compromised. In these complex systems, high-resolution techniques such as nuclear magnetic resonance (NMR) and electron spin resonance (ESR) are particularly valuable, as they can discern a signal from a single acceptor or ligand within extremely heterogeneous solutions. Assuming that each bound ligand causes an equivalent change in the spectroscopic signal, the fractional change in the spectroscopic parameter (fA) is given by:
b
gb
f A = y − yf / y b − yf
g
Equation A.5A.15
where yb and yf are the signals from the acceptor when sites are fully occupied (bound) and unoccupied (free), respectively. Conversion of the fractional saturation (fA) to the binding function (r) requires knowledge of the number of binding sites on the acceptor (p), that is, the stoichiometry of the reaction. The stoichiometry is determined from a separate titration carried out at high acceptor levels such that each amount of ligand added is fully bound to the acceptor. This condition is accomplished when the acceptor concentration is ∼100-fold greater than the dissociation constant (KASCA = 100), whereupon a plot of the fractional saturation against the ratio of total ligand to total acceptor (CS/CA) consists of two linear segments that intersect at the endpoint of the titration, thereby providing the stoichiometry (Fig. A.5A.7, line a). The condition KASCA = 10 provides a lower limit for discerning the two linear segments (Fig. A.5A.7, line b). Once the stoichiometry has been determined, the binding curve can be evaluated from a titration in which the acceptor concentration is close to the dissociation constant for the binding equilibrium (Fig. A.5A.7, line c). The value of r is established at each value of CS, whereupon the concentration of free ligand becomes: Quantitative Characterization of Ligand Binding
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Fractional acceptor saturation ( fA )
a
1.0
b 0.8 c 0.6 0.4
0.2 0 0
1
2
3
4
C S /CA
Figure A.5A.7 Evaluation of the stoichiometry of an acceptor-ligand interaction by stoichiometric titration. Curves have been constructed on the basis of a single acceptor site with a binding constant (KAS) of 106 M−1 and total acceptor concentrations (CA) of 10−4 M (curve a), 10−5 M (curve b), and 10−6 M (curve c).
CS = CS − rCA Equation A.5A.16
The data are now plotted as r versus CS or as one of the linear transforms described previously. An example of this procedure is given by the binding of the Escherichia coli cAMP receptor protein to the primary site on the lac promoter (Heyduk and Lee, 1990). The stoichiometric titration in Figure A.5A.8A establishes the 1:1 stoichiometry, while the equilibrium titration in Figure A.5A.8B provides the data for generating the binding curve in Figure A.5A.8C. The binding analysis therefore requires two titrations—one under stoichiometric conditions, the other under equilibrium conditions. A spectral signal arising from the ligand Spectral properties of the ligand may be perturbed when it becomes bound to an acceptor. For example, changes may occur in the absorption or emission maximum, extinction coefficient, or circular dichroism ellipticity. The first step in binding analysis is determination of the spectral characteristics of the free and bound ligand. The characteristics of the free ligand (yf) can be determined by direct measurement. Those of the bound ligand (yb) can be determined by a separate titration in which acceptor is added to a solution of ligand. In the presence of excess acceptor all the ligand is bound. Plotting the data in double reciprocal form (1/y versus 1/CA) allows the spectral property of the bound ligand to be discerned from the ordinate intercept. This plot is frequently assumed to be linear; however, examination of the theory shows that this need not be the case (Haigh and Sawyer, 1978). In the case of multiple equivalent or nonequivalent binding sites, it is assumed that the binding of each ligand results in an equivalent spectral change. A second titration is now carried out under equilibrium conditions in which a ligand is added to a
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constant concentration of acceptor. The fraction of the ligand bound (fS) may be found from the expression:
b
gb
fS = y − yf / yb − yf
g
Equation A.5A.17
A
0.21
Anisotropy
where y is the measured signal at a particular concentration of ligand. It then follows that
0.19
0.17 0
1
2
3
100
150
100
150
CS/CA
Anisotropy
B
0.200
0.185
0.170 0
50
C
1.0
Binding function (r )
CS (nM)
0.5
0 0
50
CS (nM)
Quantitative Characterization of Ligand Binding
Figure A.5A.8 Fluorescence polarization (anisotropy) study of the interaction between E. coli cAMP receptor protein and a 32-bp DNA fragment of the lac promoter. (A) Stoichiometric titration of the fluorescently labeled DNA fragment with receptor protein in the presence of 0.2 mM cAMP. (B) Equilibrium titration in the presence of 0.5 µM cAMP. (C) Secondary plot of the equilibrium titration data as a binding curve, together with the best fit description (p = 1, KAS = 6.0 × 107 M−1). Data taken from Heyduk and Lee (1990).
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r = fSCS /CA Equation A.5A.18
CS = CS − fSCS Equation A.5A.19
These calculations are repeated at each concentration of ligand added, so that the single titration provides the entire binding curve. Alternative approaches Other procedures have been devised for the analysis of spectroscopic binding data. The dilution titration (Anderson and Weber, 1965) can be applied to situations where the spectroscopic signal originates from either the acceptor or the ligand, and to situations where dilution of the acceptor does not cause any dissociation of an acceptor protein oligomer. An iterative procedure can be applied to situations involving equivalent and independent sites (Holbrook, 1972; Holbrook et al., 1972; Shanley et al., 1985). A model independent procedure, which avoids the restrictive assumption that each ligand causes an equivalent change in the spectroscopic signal, has been devised by Chatelier and Sawyer (1987). Competitive Binding Assays In addition to methods in which an acceptor-ligand interaction is monitored directly, there are also procedures wherein equilibrium parameters for the interaction of interest are inferred from the inhibitory effect of a competing ligand on an acceptor interaction that has already been characterized. Such competitive binding assays abound in the screening of drug and antigen analogs by solid-phase radioimmunoassay and enzyme-linked immunosorbant assay (ELISA) techniques, and also in the characterization of interactions by quantitative affinity chromatography (Winzor, 1997) and biosensor technology (Hall and Winzor, 1998). The analysis of competitive binding assays has been considered most extensively in the context of quantitative affinity chromatography, and is therefore presented in that light. General theoretical aspects In solid-phase competitive binding assays, the matrix-associated reactant (X) is chosen to obtain selective interaction with the solute of interest (acceptor A). For example, the interaction of antithrombin (A) with heparin (S) has been studied (Hogg et al., 1991) using heparin immobilized on Sepharose as the matrix-bound reactant (X). Ligand-facilitated elution of acceptor (A) from the matrix thus occurs because its interactions with ligand (S, soluble heparin) and matrix sites (X, immobilized heparin) are mutually exclusive (Fig. A.5A.9). The first step of a competitive binding assay entails determination of the binding constant for the acceptor-matrix interaction (KAX) and an associated curve-fitting parameter, the = effective total concentration of matrix sites (CX). Provided that A is univalent in its interaction with matrix sites, the extent of acceptor depletion from the liquid phase exhibits a rectangular hyperbolic dependence upon the acceptor concentration in the liquid phase (CA). For a mixture in which the total acceptor concentration in the system = is CA, the rectangular hyperbolic dependence may be written in the form Biophysical Methods: Data Analysis
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X
KAX matrix
XA
KAS A
AS
X Figure A.5A.9 Schematic representation of the competitive interactions of soluble ligand (S) and immobilized affinity sites (X) for acceptor (A) in quantitative affinity chromatography.
c
CA − CA = K AX CX CA / 1 + K AX CA
h
Equation A.5A.20
of which the most convenient version of the Scatchard linear transform is
e CA/ CA j − 1 = KAX CX − KAX CA LMNe CA /CA j − 1OPQ Equation A.5A.21
Expression of the Scatchard transform in that manner allows its ready adaptation to = column chromatographic data, for which the ratio CA/CA may be replaced by VA/VA*, where VA is the measured elution volume of acceptor on an affinity column for which VA* is the corresponding elution volume in the absence of any acceptor-matrix interaction (the elution volume of ligand-saturated A). For frontal chromatographic studies, the counterpart of Equation A.5A.21 thus becomes (Hogg and Winzor, 1984)
eVA /VA* j − 1 = KAX CX − KAXCA eVA /VA* j − 1 Equation A.5A.22
Upon supplementation of the liquid phase with a free concentration (CS) of competing = ligand, the rectangular hyperbolic dependence of (CA− CA) upon CA continues to be described by Equation A.5A.20, except that the acceptor-matrix binding constant so determined is a constitutive parameter (KAX) that is related to KAX by the relationship (Winzor and Jackson, 1993) Q = K AX / K AX = 1 + K ASCS Equation A.5A.23
The binding constant for the competing acceptor-ligand interaction in solution (KAS) may therefore be obtained from the linear dependence of the ratio of acceptor-matrix binding constants (Q) upon free ligand concentration. In instances where the total ligand concentration (CS) is the parameter of known magnitude, Equation A.5A.23 is replaced by the expression Quantitative Characterization of Ligand Binding
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a
f
Q = 1 + K AS qCS − Q − 1 CA /Q Equation A.5A.24
where q is the valence of S in its interaction with A, a consideration necessitated by the enforced design of the experiment whereby A must be univalent in its interaction with matrix sites X. The problem of allowing A to be multivalent is addressed later. Partition equilibrium variants of quantitative affinity chromatography The most direct method of employing Equation A.5A.20 or A.5A.21 is to perform simple partition equilibrium experiments on matrix-acceptor mixtures (with or without S). Measurement of the liquid-phase concentration of A after attainment of equilibrium allows the concentration of matrix-bound solute to be determined as the difference between the total acceptor concentration in the matrix slurry and its corresponding concentration in the liquid phase (Nichol et al., 1974). The problem of ensuring that each mixture contains the same amount of affinity matrix can frequently be overcome by using a recycling partition technique (Ford and Winzor, 1981; Hogg et al., 1991) in which the liquid-phase concentration of acceptor (CA) is monitored spectrophotometrically by means of a flow cell placed in the line returning the liquid phase to the slurry (Fig. A.5A.10A). This procedure has the advantage that a number of partition experiments may be performed with the same sample of matrix by making successive additions of concentrated acceptor solution to the slurry and determining the corresponding value of = CA after each addition. The dilution of the concentration of affinity sites (CX) arising from the stepwise addition of acceptor (and ligand) aliquots with volume δV is accommodated = by multiplying each (CA − CA) result by VA*/(VA*)0: the relevant volume accessible to A after n such additions (VA*) is calculated from the initial value (VA*)0 and the relationship VA* = (VA*)0 + nδV. A Scatchard representation of results obtained by this technique for the interaction of antithrombin with heparin-Sepharose (Hogg et al., 1991) is presented in Figure A.5A.10B. Column chromatographic variants Inspection of Equation A.5A.22 shows that knowledge of CA, the concentration of acceptor in the liquid phase, is a prerequisite for its application to elution profiles obtained by applying acceptor solution (or acceptor-ligand mixture) to an affinity column. However, there have been relatively few chromatographic studies in which the full potential of Equation A.5A.22 has been put to advantage (Winzor, 1997). Far greater use has been made of zonal chromatography, where the lack of a value of CA dictates the truncation of Equation A.5A.22 to:
eVA /VA* j − 1 = KAX CX Equation A.5A.25
which involves the assumption that the total concentration of matrix sites is an acceptable substitute for their free concentration (Bergman and Winzor, 1986). Subject to the validity of that approximation, zonal chromatography of the solute on an affinity column preequilibrated with a range of concentrations of competing ligand (CS) allows the acceptor-ligand binding constant to be evaluated from the expression
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eVA /VA* j − 1 = KAX CX /b1 + KASCS g Equation A.5A.26
=
in which KAS and the product KAXCX are the two parameters to be obtained by curve fitting. Further details of this procedure, introduced by Dunn and Chaiken (1974), are to be found in a recent review of quantitative affinity chromatography (Winzor, 1997). Biosensor studies of ligand binding An alternative means of quantifying the interaction between acceptor and matrix sites is to directly monitor the concentration of matrix-bound A in an equilibrium mixture by
A
pump
monitor
stirrer
matrix slurry
sintered disc
thermostat
[VA*/(VA*)0](CA – CA)/CA
B
15
10 –KAX
5 (C X)0
0 0
0.6
1.2
1.8
2.4
[VA*/(VA*)0 ](CA – CA) (µM)
Quantitative Characterization of Ligand Binding
Figure A.5A.10 Studies of the interaction between acceptor and immobilized affinity sites by the recycling partition variant of quantitative affinity chromatography. (A) Schematic representation of the experimental system. (B) Scatchard plot of results obtained for the interaction of antithrombin with heparin-Sepharose (Hogg et al., 1991).
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biosensor technologies such as those incorporated into the BIAcore and IAsys instruments. Although the major interest in biosensor-based instruments has been their use to determine the kinetics of adsorption and desorption for the acceptor-matrix interaction, attention has also been drawn to the potential of their application in the context of quantitative affinity chromatography (Ward et al., 1995; Hall and Winzor, 1997, 1998). In the BIAcore instrumentation, a solution of acceptor flows through a capillary channel over the biosensor chip, which has been rendered an affinity matrix by the covalent attachment of affinity sites X. Because the solution in contact with the sensor surface is being replaced continually by fresh solution, the time-independent response that is ultimately attained (Re) is directly proportional to the concentration of bound acceptor = (CA − CA) in equilibrium with CA, the concentration of acceptor in the solution being injected into the instrument. Measurement of Re as a function of injected acceptor concentration (CA) allows KAX to be determined from the expression
e
j c
Re = K AX BCX CA / 1 + K AX CA
h
Equation A.5A.27
=
which is essentially a rearranged form of Equation A.5A.20 with (CA − CA) replaced by Re/B, where B is the proportionality constant relating biosensor response to bound acceptor concentration. The other parameter to emerge from curve fitting of the (Re, CA) = data to this rectangular hyperbolic expression is the product BCX, the maximum response corresponding to saturation of all affinity sites. The equilibrium constant for the interaction between acceptor and a competing ligand (S) is then obtained by injecting acceptor-ligand mixtures with= a range of total concentrations of competitive inhibitor (CS). Because the magnitude of BCX has already been established, the substitution of each (Re, CA, CS) combination into Equation A.5A.27 yields the value of KAX pertinent to Equation A.5A.24. KAS is then evaluated from the slope of the dependence of Q = KAX/KAX upon [qCS − (Q − 1)CA/Q]. Analysis of results obtained from the IAsys instrument follows essentially the same protocol, except that CA in Equation A.5A.27 is not the concentration of acceptor placed in the cuvette, because allowance needs to be made for acceptor depletion as the result of AX formation. Thus, the value of CA for use in Equation A.5A.27 (and Equation A.5A.24) must be obtained from the relationship CA = (CA)0 − Re/B, where (CA)0 is the concentration of acceptor placed in the cuvette at the start of the measurement (Hall and Winzor, 1997, 1998; Edwards et al., 1998). Illustrative evaluation of KAS by competitive binding assay The evaluation of the equilibrium constant for an acceptor-ligand interaction by competitive binding assay is illustrated here using a BIAcore study of the interaction between interleukin 6 and the soluble form of its specific receptor (Ward et al., 1995). The recombinant cytokine proved to be dimeric, and has therefore been attached to the biosensor surface so that results could be interpreted in terms of Equation A.5A.27 with the soluble receptor as the univalent acceptor (A). Characterization of the acceptor-matrix interaction is summarized in Figure A.5A.11A, which presents the results in Scatchard format. Values of 2.4 (± 0.2) × 107 M−1 and 2200 (± 100) response units are obtained for = KAX and the product BCX, respectively. Results from the series of experiments on mixtures of acceptor and interleukin 6 are plotted in Figure A.5A.11B according to Equation A.5A.24, with q = 2 because of the dimeric nature of the cytokine. A value of 4.8 (± 0.3) × 107 M−1 is obtained from the slope.
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A
60
Re / CA (nM –1)
40
20
0 0
500
1000
1500
2000
2500
200
250
B
15
Q = KAX/KAX
Re (response units)
10
5
0 0
50
100
150
2CS – (Q – 1)CA/Q (nM)
Figure A.5A.11 Characterization of an acceptor-ligand interaction by competitive binding assay. (A) Scatchard plot of equilibrium responses for the interaction of the soluble domain of interleukin 6 receptor (A) with interleukin 6 immobilized on the sensor surface of a BIAcore instrument. (B) Plot of results obtained with interleukin 6 as the competing ligand (S) in accordance with Equation A.5A.24, with q = 2. Data taken from Ward et al. (1995).
This use of BIAcore data to illustrate the evaluation of equilibrium constants by competitive binding assay is open to criticism on the grounds that partition or chromatographic studies are more traditional forms of quantitative affinity chromatography. However, the selected example serves an important role by emphasizing the assumed univalence of acceptor in the analysis, and by illustrating one means by which the problem of multivalence can be averted. An alternative approach is discussed in the next section. BINDING RESPONSES DEVIATING FROM RECTANGULAR HYPERBOLIC FORM
Quantitative Characterization of Ligand Binding
It has already been emphasized that the rectangular hyperbola represents the simplest form of binding response, and that deviations therefrom signify the need for the incorporation of additional complexity into the model to account for the shape of the binding curves. Possible sources of deviation from the form shown as the broken line (- - -) in Figure A.5A.1 are considered below.
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Effect of Acceptor Site Heterogeneity Equations A.5A.1-A.5A.4 have been used to describe a model involving ligand interaction with two equivalent and noninteracting acceptor sites; this model can now be extended to a situation in which the two binding sites are nonequivalent but still independent. Independence means that the binding constant for one site is the same regardless of whether the other site is occupied or not. Thus, in terms of Equations A.5A.1-A.5A.4, K1 = K4 and K2 = K3. A rearrangement of Equation A.5A.7 shows that the binding to two nonequivalent and independent sites is described by the sum of two rectangular hyperbolae.
b
g
b
K C 1 + K2 CS + K2 CS 1 + K1CS r= 1 S 1 + K1CS 1 + K2 CS
b
gb
g
g
Equation A.5A.28
r=
K1CS K2 CS + 1 + K1CS 1 + K2 CS
b
g b
g
Equation A.5A.29
For two nonequivalent groups of sites (p and q in each group) the general binding equation becomes r=
pK1CS qK2 CS + 1 + K1CS 1 + K2 CS
b
g b
g
Equation A.5A.30
which may be used directly in nonlinear least-squares fitting programs to extract estimates of the two binding constants and the two site numbers. The corresponding Scatchard plot is nonlinear and of the form shown in Figure A.5A.12. Here, the deviation from linearity is useful in recognizing that a simple model involving homogeneous sites is inadequate to describe the experimental data. For a more quantitative analysis, the Scatchard plot is of limited use. A common misconception is that the limiting slopes of this graph provide estimates of the two binding constants while their extrapolation to the abscissa provides the respective site numbers. This difficulty is illustrated in Figure A.5A.12, which shows the Scatchard plots for a binding model involving two independent sites that differ in affinity by factors of 2, 10, and 100. The Scatchard plots for the binding to each independent site are included. The following features are apparent: 1. The Scatchard plot is not the simple sum of the plots describing the individual site contributions. 2. The abscissa intercept represents the total site number (in this case two), but becomes more difficult to determine graphically as the difference between the binding affinities increases. 3. When the binding affinities differ by a factor of two, it becomes very difficult to experimentally identify the deviation from linearity and thus the site heterogeneity. 4. As the difference between the binding affinities increases, the limiting slope of the steepest part of the Scatchard plot provides an estimate of the tighter binding constant.
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1.5
r/CS (×10 – 4 M –1)
1.0
a 0.5
b c
0 0
1
2
Binding function (r )
Figure A.5A.12 Illustrative Scatchard plots for the interaction of ligand with two independent but nonequivalent acceptor sites. The higher-affinity site has been ascribed a binding constant of 105 M−1, whereas the other has been accorded values of 5 × 104 M−1 (curve a), 104 M−1 (curve b), and 103 M−1 (curve c). The broken line describes the interaction of ligand with the higher-affinity binding site, whereas the dotted lines have a common abscissa intercept of 2 and slopes characteristic of the weaker interactions.
However, as has been carefully documented by Klotz (1997), the limiting slopes and intercepts at high and low values of r are complex functions of the two binding constants and site numbers. 5. Besides the difficulty of extracting the binding parameters graphically from the Scatchard plot, it should be recognized that a similar deviation from linearity is observed in cases where multiple sites interact in a negatively cooperative manner. A slightly different situation arises when the two groups of sites (p and q) are on two different noninteracting molecules (A and B) rather than on the same molecule. In addition to being determined by the binding affinities and site numbers (KAS, KBS, p, q), the binding function is determined by the mole fraction (α1) of each acceptor species present: αA = CA/(CA + CB) and αB = (1 − αA) = CB/(CA + CB), where the overbar notation again signifies the total concentration (free and complexed) of a given acceptor. In these circumstances Quantitative Characterization of Ligand Binding
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r=
b
g
α A pK ASCS 1 − α A qK BSCS + 1 + K ASCS 1 + K BSCS
b
g
b
g
Equation A.5A.31
where αA =
b
CA 1 + K ASCS
b
CA 1 + K ASCS
gp
g p + CBb1 + KBSCS gq
Equation A.5A.32
Effect of Ligand Multivalence A limitation of the binding equations presented thus far has been their reliance upon the assumption that the ligand is univalent in its interaction with acceptor sites. Although the use of those binding equations is entirely reasonable for systems in which the ligand is a small, unifunctional metabolite, the validity of their application to (for example) the interactions of antibodies with their specific surface antigens is highly questionable. The problem is that ligand multivalence introduces a deviation from the rectangular hyperbolic response that is of the same form as that for binding of a univalent ligand to heterogeneous acceptor sites (Calvert et al., 1979). Effects of ligand multivalence on the interpretation of binding data were first considered in the context of quantitative affininty chromatography (Nichol et al., 1981), but have been incorporated subsequently into a general counterpart of the Scatchard analysis (Hogg and Winzor, 1985). For a situation in which a ligand possessing f sites binds to an acceptor with p sites for ligand, the binding function needs to be defined (Hogg and Winzor, 1985) as
e
j
r f = CS1/ f − CS1/ f /CA Equation A.5A.33
In terms of this binding function, the multivalent counterpart of the Scatchard equation for equivalent and independent binding governed by the intrinsic association constant KAS is
a f −1f/ f
r f /CS1/ f = pK AS − fK ASr f CS Equation A.5A.34
A linear dependence of rf/CS1/f upon rfCS(f − 1)/f is thus the requirement for equivalence and independence of sites when the ligand is multivalent. Because the application of Equation A.5A.34 to binding data requires the specification of a value for f, there has been a tendency to avoid the necessity of so doing by retaining the conventional Scatchard analysis (Equation A.5A.12). However, such action merely signifies that a value of unity has been chosen as the most appropriate valence of the ligand. Results from a partition equilibrium study of the binding of aldolase to muscle myofibrils (Kuter et al., 1983) serve to emphasize that point. The conventional Scatchard analysis of those results is presented in Figure A.5A.13A, which seemingly signifies heterogeneity of the myofibrillar sites in their interaction with enzyme. However, symptoms of site heterogeneity disappear on reanalysis of the results according to Equation
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A
B
2.0
0.4 r 4/C S1/4 (µM–1)
r /C S (µM–1)
1.6
0.5
1.2
0.8
0.3
0.2
0.4
0.1
0
0 0
0.4
1.2
0.8
0
0.1
0.2
0.3
r 4C S3/4
Binding function (r )
Figure A.5A.13 Analysis of binding data for the interaction of aldolase with rabbit muscle myofibrils. (A) Conventional Scatchard plot. (B) Multivalent Scatchard plot according to Equation A.5A.34 with f = 4. Data taken from Table 1 of Kuter et al. (1983).
A.5A.34 with f = 4 on the grounds that aldolase is a tetramer (Fig. A.5A.13B). The linearity of this plot, particularly in the light of its compliance with the mandatory abscissa intercept of 0.25 (p/f with p = 1), establishes the adequacy of a single intrinsic binding constant of 3.8 (± 0.3) × 105 M−1 to describe the binding of tetravalent enzyme to sites on the myofibrillar matrix. Although this analysis is open to criticism on the grounds of its reliance upon transformed experimental parameters, the finding has been confirmed by analysis of the results in terms of the rectangular hyperbolic expression of which Equation A.5A.34 is the linear transform (Harris et al., 1995a). Use of the expression
a f −1f/ f = fr f C S
a f −1f/ f KASC1/ f S f −1f/ f a K ASC1/ f 1 + fC pfCS
S
S
Equation A.5A.35
allows the magnitudes of p and KAS to be obtained by substituting frfCS(f − 1)/f for r and fCS(f − 1)/fCS1/f for CS in standard computer programs for the evaluation of binding parameters.
Quantitative Characterization of Ligand Binding
In studies of binding to a particulate acceptor there are circumstances under which a rectangular hyperbolic binding response is observed despite multivalence of the ligand (Nichol et al., 1974; Brinkworth et al., 1975)—a finding rationalized intuitively at the time on the grounds that physical restrictions precluded multiple interactions of ligand with acceptor. That the binding constant deduced under such circumstances is the product fKAS has been confirmed by a subsequent mathematical explanation of such behavior (Kalinin et al., 1995), which arises when the total concentration of ligand greatly exceeds that of acceptor sites. Under conditions where the concentration of bound ligand (CS −
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CS) is small relative to the free concentration (CS), the total ligand concentration CS may be written as:
c
h
CS = CS + CS − CS = CS (1 + δ ) Equation A.5A.36
where δ = (CS − CS)/CS is much smaller than unity. From the binomial theorem it follows that CS1/ f = CS1/ f (1 + δ/ f + ...) Equation A.5A.37
whereupon the binding function (Equation A.5A.33) becomes rf = CS1/fδ/(fCA), and Equation A.5A.34 simplifies to
cCS − CS h/CS = fKASCA − fKAS cCS − CS h 1 + ( f − 1)δ/ f Equation A.5A.38
which signifies a slope of f[1 + (f − 1)δ/f]KAS ≈ fKAS for the slope of the conventional Scatchard plot. Equivalent but Cooperative Binding In instances where the acceptor comprises several identical (or very similar) subunits, the binding of a univalent ligand to any one subunit has the potential to induce conformational changes in adjacent subunits and thereby alter their intrinsic affinities for ligand. This concept of equivalent but dependent acceptor sites was introduced by Koshland et al. (1966), who noted the existence of two possible situations. (1) In positive cooperativity, ligand-induced conformational changes in the acceptor enhance subsequent ligand attachments, resulting in a sigmoidal binding response (Fig. A.5A.14A). (2) In negative cooperativity, the alternative situation occurs, with a decreased intrinsic binding constant as the result of conformational change(s) giving rise to a binding response of the same form as that reflecting heterogeneity of acceptor sites (Fig. A.5A.14B). Allowance for nonidentity of successive intrinsic association constants is effected in the Adair (1925) formulation of the binding expression. Thus, Equation A.5A.7 for a system with p = 2 becomes:
b g b g
b gb g b gb g
2 K AS 1 CS + 2 K AS 1 K AS 2 CS2 r= 1 + 2 K AS 1 CS + K AS 1 K AS 2 CS2 Equation A.5A.39
where (KAS)1 is the intrinsic binding constant for the interaction of S with either site on free A, and (KAS)2 is the modified value for occupancy of the remaining site to complete saturation of the acceptor. Multivalence of the ligand introduces the need to consider whether acceptor or ligand sites are the source of the cooperativity (Harris et al., 1995b). For systems in which the cooperativity resides in acceptor sites, the criteria used above for the interpretation of
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binding curves for univalent ligands (Fig. A.5A.14) also apply to data for multivalent ligands provided that results are analyzed in terms of the rectangular hyperbolic relationship for an f-valent ligand (Equation A.5A.35) or its linear transform (Equation A.5A.34). Thus, bivalent Scatchard plots of the form shown in Figure A.5A.15, panels A and B, signify the existence of positive and negative cooperativity, respectively. However, the interpretation must be reversed when ligand sites are the source of cooperativity: the sigmoidal response (Fig. A.5A.15A) now signifies negatively cooperative binding (Harris et al., 1995b).
r /C S
Binding function (r )
B
r /C S
Binding function (r )
A
r
r
Free ligand (C S )
Free ligand (C S )
Figure A.5A.14 Schematic representation of binding curves and their Scatchard transforms reflecting (A) positive cooperativity and (B) negative cooperativity of acceptor sites for a univalent ligand.
frfCS(f –1)/f
rf / C S 1/ f
frfCS(f –1)/f
rfCS(f–1)/f f C S ( f –1) / f C S1/ f
Quantitative Characterization of Ligand Binding
rf / C S 1/ f
B
A
rfCS(f–1)/f
f C S ( f –1) / f C S1/ f
Figure A.5A.15 Conflicting criteria for cooperative binding of multivalent ligands. (A) Schematic binding curve and its multivalent Scatchard counterpart reflecting either positive cooperativity of acceptor sites or negative cooperativity of ligand sites. (B) Corresponding plots reflecting either negative cooperativity of acceptor sites or positive cooperativity of ligand sites.
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Because conflicting criteria apply to the interpretation of cooperativity emanating from acceptor and ligand sites, additional information is clearly required before mechanistic significance can be ascribed to the forms of binding curves or their linear transforms. In that regard, the interpretation of results obtained for the binding of pyruvate kinase to muscle myofibrils (Harris et al., 1995b) serves to illustrate the type of approach that may be used to resolve the dilemma. Interplays of Equilibria Instead of considering the acceptor to undergo ligand-induced conformational changes, Monod et al. (1965) propose that a sigmoidal binding response reflects the preferential binding of a univalent ligand to one of two isomeric acceptor states that coexist in equilibrium. Although the binding sites within a given acceptor state are considered to be equivalent and independent, the intrinsic binding constant and/or the number of binding sites are allowed to differ between the two isomeric states. Even greater generality is accorded this model by allowing the preexisting equilibrium to include acceptor self-association (Nichol et al., 1967). For a system where states A and C possess p and q sites, respectively, with respective intrinsic association constants KAS and KCS, the binding equation becomes: r=
b
g p−1 + nXCAn−1qKCSCS b1 + KCSCS gq−1 b1 + KASCS g p + nXCAn−1b1 + KCSCS gq
pKASCS 1 + K ASCS
Equation A.5A.40
where CA and CC are the concentrations of the respective isomeric states, and X = CC/CAn is the equilibrium constant describing the preexisting coexistence of acceptor states (nA in equilibrium with C). Preferential binding to an isomerizing acceptor A major difference between the models based on acceptor isomerization and self-association is the dependence of the binding function upon acceptor concentration for n > 1, but its independence of acceptor concentration for the Monod et al. (1965) model (n = 1 in Equation A.5A.40). The latter feature is shared by the cooperativity model (Koshland et al., 1966), which is mathematically equivalent to the Monod model of allostery in the absence of a means of distinguishing between an initial state (CS = 0) comprising a single acceptor state (Koshland et al., 1966) and an initial state comprising an equilibrium mixture of isomeric states with one form dominant (Monod et al., 1965). Under favorable circumstances the molecular crowding effect of thermodynamic nonideality has potential for making such distinction (Harris and Winzor, 1988; Bergman and Winzor, 1989; Bergman et al., 1989), but pyruvate kinase is the only allosteric system to which this method has been applied, and thereby established preexistence of an isomerization equilibrium (Harris and Winzor, 1988). However, even that approach is not necessarily definitive, because failure to observe a nonideality-mediated shift towards a smaller isomeric state does not preclude the possibility that the conformation change relating to a preexisting isomerization equilibrium is too subtle for detection as a gross change in size and/or shape. Furthermore, as noted by Saroff (1991), the two models are not mutually exclusive inasmuch as coexisting acceptor states may also exhibit cooperativity of ligand binding. In view of the above considerations, a logical approach is to set aside aspirations of an unequivocal and mechanistically correct characterization of a sigmoidal binding response
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1.00
v/v m ( r/p)
0.75
0.50
0.25
0 0
5
15
10
20
Aspartate (nM)
Figure A.5A.16 Allosteric activation and inhibition of E. coli aspartate transcarbamylase by effectors. Open circles, dependence of initial velocity v (expressed relative to maximal value vm) upon aspartate concentration (CS) in the presence of saturating carbamyl phosphate; filled circles and filled squares, corresponding dependences with 0.5 mM CTP and 2 mM ATP, respectively, included in the reaction mixtures. Data taken from Gerhart (1970).
in favor of an acceptable thermodynamic description in terms of the allosteric model for which parameters are evaluated more readily. The authors’ inclination is towards interpretation in terms of the Monod model, which is also adapted readily to incorporate the consequences of allosteric effectors. The moderating influence of an effector (E) binding to different sites from S on the two isomeric states is accommodated in Equation A.5A.40 with n = 1 by replacing the self-association constant (X) with a constitutive isomerization constant (X) defined as: X=
b
X 1 + KCSCE
gz
b1 + KAECE gw
Equation A.5A.41
Quantitative Characterization of Ligand Binding
where w and z are the respective numbers of sites for effector on A and C isomers, for which intrinsic binding constants of KAE and KCE pertain (Frieden, 1967; Changeux and Rubin, 1968; Nichol et al., 1972b). Preferential binding of E to the isomer with a lower affinity for S gives rise to allosteric inhibition and a more sigmoidal binding curve in the presence of effector. Conversely, the binding curves exhibit activation and a tendency to become rectangular hyperbolic when the effector binds preferentially to the isomer with higher affinity for S. These phenomena are evident in Figure A.5A.16, which illustrates the inhibitory effect of CTP and the activatory effect of ATP on the aspartate transcarbamylase reaction (Gerhart, 1970).
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Fractional saturation (r/p)
1.0
0.5 0.7 1.3 4.5 90 106 0 0
4
8
12
16
20
24
28
CS (µM)
Figure A.5A.17 Concentration dependence of the binding curve for a self-associating acceptor. Binding curves for oxygen uptake by the indicated total concentrations of human hemoglobin (0.7 to 106 µg/ml) at pH 7.4. Data taken from Mills et al. (1976).
Preferential binding to a self-associating acceptor Whereas the binding function for the interaction of a ligand with an isomerizing acceptor is expressed in terms of free ligand concentration (CS) as the single variable, its magnitude also depends upon the free acceptor concentration (CA), and hence the total acceptor concentration when n > 1 in Equation A.5A.40. In that regard, the required concentration of free monomer is given by the physically acceptable solution of the equation expressing mass conservation of acceptor, namely:
b
nX 1 + K CSCS
gq CAn + b1 + KASCS g p CA − CA = 0 Equation A.5A.42
where CA is the base-molar concentration of acceptor (total weight concentration divided by the molecular weight of monomer). Simulations of a binding curve also require the specification of magnitudes for the stoichiometry (n) and equilibrium constant (X) describing the preexisting self-association of acceptor—parameters obtained by physicochemical characterization of the acceptor in the absence of ligand (Nichol et al., 1972a; Tellam et al., 1978, 1979). The importance of maintaining a fixed total acceptor concentration during the experimental delineation of a binding curve for a self-associating acceptor is evident from Figure A.5A.17, which summarizes the interaction of oxygen with hemoglobin over a wide range of protein concentrations (0.7 to 106 µg/ml)—a range where the tetrameric α2β2 species coexists in equilibrium with dimer (αβ), which binds oxygen preferentially (Mills et al., 1976). Clearly, there is a separate binding curve for each fixed total acceptor concentration. Although a change in CA may well enhance the accuracy with which a value of r may be determined at a given CS, a binding curve constructed in this manner would comprise a composite of segments from the binding curves described by Equation A.5A.40 for each total concentration of acceptor used. Biophysical Methods: Data Analysis
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Sigmoidal effects reflecting ligand self-association There is a tendency to regard the acceptor as the species undergoing self-association or isomerization, and therefore to regard allostery as a property emanating from the acceptor. However, there are also situations in which the reactant undergoing self-interaction is the ligand. For the relatively simple situation in which each state of a self-associating ligand (nS ↔ T) interacts competitively with the same p equivalent and independent sites on an acceptor A for which the respective intrinsic binding constants are KAS and KAT, it is relevant to define the binding function as:
c
h
r = CS − CS − nCT /CA Equation A.5A.43
whereupon the binding equation becomes r=
b
p K ASCS + nK AT CT 1 + KASCS + K AT CT
g
Equation A.5A.44
A rectangular hyperbolic response of r upon CS = CS + nCT results from ligand isomerization, whereas a sigmoidal response emanates when n > 1 and KAS = 0, the situation when acceptor binds exclusively to the polymeric form of the ligand. The curves exhibit symptoms of negative cooperativity when monomeric ligand is the exclusive form of ligand bound (Nichol et al., 1969). An example of a situation in which the polymeric form of ligand binds preferentially to acceptor is provided by the interaction of chlorpromazine with tubulin (Hinman and Cann, 1976; Cann et al., 1981)—a system for which the Scatchard plot exhibits two critical points, a maximum and a minimum. Although the unusual form of the Scatchard plot was initially interpreted as signifying relatively strong interaction of chlorpromazine with a single tubulin site and weaker, positively cooperative binding of an additional eight or nine chlorpromazine molecules (Hinman and Cann, 1976), it has subsequently been shown that preferential binding of the chlorpromazine micelle (n = 9 to 11) is a more plausible explanation of the results (Cann et al., 1981). Effect of Ligand Binding on Acceptor Self-Association For an acceptor undergoing preexisting self-association between monomer and a single higher polymeric state (nA in equilibrium with C), the association equilibrium constant can be determined from studies of the acceptor in the absence of ligand. For example, the weight-average_molecular weight (MW) of an equilibrium mixture with total acceptor concentration (ca in mg/ml) is given by:
b
g
M w = cA MA + ca − cA nM A / ca Equation A.5A.45
where MA is the molecular weight of the monomer. This allows the concentration of monomeric acceptor (cA) to be evaluated from the expression
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b a f
c nMA − Mw cA = a n − 1 MA
g
Equation A.5A.46
for an assigned value of n. In fact, this specification of the association stoichiometry can be avoided by direct analysis of the sedimentation equilibrium distributions reflecting acceptor self-association (Milthorpe et al., 1975; Wills et al., 1996). The availability of methods for determining cA as a function of total acceptor concentration allows the association equilibrium constant X (litern−1moln−1) to be determined as:
b
g
n n −1 X = ca − cA /cA MA /n
Equation A.5A.47
This is frequently accomplished from a logarithmic variant of that relationship, namely
b
g
e
j
n −1 + n log cA log ca − cA = log nX / M A
Equation A.5A.48
which allows the magnitudes of n and X _to be determined from the slope and ordinate intercept of the linear dependence of log(ca − cA) upon logcA. Provided that the ligand is sufficiently small in comparison with acceptor for the approximations MASi ≈ MC and MCSi ≈ MC to apply, the perturbation by ligand of the self-association equilibrium affords quantitative information on the relative affinities of the two acceptor states for S (Nichol et al., 1972a; Baghurst and Nichol, 1975; Tellam et al., 1979). Under those circumstances the determination of molecular weight data or direct _ analysis of sedimentation equilibrium distributions yields cA, the weight-concentration of monomer in free and ligand-bound states, whereupon the parameter obtained from Equation A.5A.44 is a constitutive association equilibrium constant (X). Its relationship to parameters within the interplay of equilibria is as follows: X=
b
g
n ca − cA /cA
=
b
X 1 + KCSCS
gq
b1 + KASCS gnp
Equation A.5A.49
The use of this approach is illustrated by a study of the interaction between indole and α-chymotrypsin (Ford and Winzor, 1983), which undergoes reversible dimerization under the conditions of the investigation—a factor evident from the slope of 2 for the plot of sedimentation equilibrium results in the absence of ligand (Fig. A.5A.18, open symbols) according to Equation A.5A.48. Repetition of the experiments with enzyme in dialysis equilibrium with 1 mM (filled circles) and 2 mM (filled squares) indole gives rise to a progressive decrease in X that is commensurate with monomer exhibiting a five-fold greater affinity than dimeric α-chymotrypsin for ligand (p = 1, q = 2, KAS = 5KCS). Interplays of equilibria involving acceptor self-association can be characterized with far greater certainty and precision than those involving acceptor states in isomerization equilibrium. Studies of the acceptor in the absence of ligand can usually be used to guide the distinction between preexisting and ligand-induced self-association as the source of
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–0.4
Log(c a – c A)
–0.8
–1.2
–1.6
–2.0 – 0.6
– 0.4
0
–0.2
+0.2
+0.4
LogcA
Figure A.5A.18 Perturbation of the monomer-dimer equilibrium for α-chymotrypsin (pH 3.9, I 0.11) by the preferential interaction of indole with monomeric enzyme. Plot of results from sedimentation equilibrium experiments according to Equation A.5A.48 for enzyme alone (open symbols) and in the presence of 1 mM (filled circles) and 2 mM (filled squares) ligand.
a sigmoidal binding curve (e.g., in Fig. A.5A.18, where the dimerization of α-chymotrypsin is clearly a preexisting equilibrium). Furthermore, perturbations of the preexisting self-association equilibrium as the result of preferential binding to one acceptor state provides additional quantitative information on the relative affinities of the two acceptor states for ligand. Indeed, measurement of the effect of free ligand concentration on the magnitude of X has formed the sole basis for characterization of the interaction between NADH (an analog of 2,3-bisphosphoglycerate) and methemoglobin—a system for which no meaningful parameters would be extracted from analysis of the binding curve (Jacobsen and Winzor, 1995). On the other hand, the characterization of an interplay of equilibria involving acceptor isomerization relies almost entirely on the analysis of the binding curve—a situation that leads to the isomerization constant, as well as the binding characteristics for the isomer with lower affinity for ligand, being curve-fitting parameters. To that end, the use of thermodynamic nonideality effected by the inclusion of molecular crowding agents to probe preexisting isomerization equilibria can, under favorable circumstances, not only detect the phenomenon but also yield an independent estimate of X (Harris and Winzor, 1988). COMPLICATIONS ARISING FROM NONSPECIFIC BINDING There are many examples in the literature where binding curves deviate from rectangular hyperbolic form as the result of elements of nonspecificity in the overall interaction between ligand and acceptor. There are two major areas where nonspecificity is encountered, and in each the term nonspecific binding has a different connotation. Quantitative Characterization of Ligand Binding
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Membrane-Receptor Interactions The binding of ligands to membrane-bound acceptors (receptors) is a fundamental phenomenon encountered in cell biology, immunology, and endocrinology. Many studies of these ligand-receptor interactions entail the use of radioactively labeled ligand to facilitate measurement of the concentration of unbound ligand in the supernatant after centrifugation to pellet the membranes. Results of such experiments rarely exhibit a rectangular hyperbolic dependence of binding upon free ligand concentration inasmuch as no plateau region is reached. Instead, there is a continuous increase in binding, which is seemingly unsaturable (filled squares, Fig. A.5A.19A). An operative criterion that is used to distinguish specific interaction from other phenomenon, termed nonspecific binding, relies upon an assertion that specifically bound ligand can be displaced by an excess of nonradioactively labeled ligand, whereas the other form of binding is unaffected by this procedure (filled circles, Fig. A.5A.19A).
A
250
Bound [125 I ]insulin (cpm)
The above definition of nonspecific binding only covers ligand-membrane interactions that are irreversible. In that regard, binding curves of the same form as that observed for
125
0 0
Binding function (r )
B
100 150 50 [125 I ] Insulin concentration (ng/ml)
200
10
5
0 0
20
40
60
80
Free metoprolol concentration (µM)
Figure A.5A.19 Nonspecific ligand interactions in studies involving membrane receptors. (A) Binding of [125I]insulin to insulin receptors on Chinese hampster ovary cells (filled squares), and the corrected binding curve (open squares) after allowance for bound radioactivity in the presence of excess nonlabeled insulin (filled circles). Adapted from Winzor and Sawyer (1995). (B) Uptake of metoprolol as the result of physical partition into hepatic microsomes. Data taken from Bogoyevitch et al. (1987).
Biophysical Methods: Data Analysis
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the interaction of insulin with its membrane receptors can also reflect nonspecific uptake of ligand as the result of partition into the membrane phase. Such ligand partitioning has a similar effect on the form of the binding curve because its dependence upon free ligand concentration is also linear (Fig. A.5A.19B). However, inasmuch as the partitioned ligand can be displaced by an excess of nonradioactively labeled ligand, the phenomenon would be classified as specific binding by the above definition. A convenient procedure for the analysis of systems exhibiting both specific binding and partition requires the collection of binding data at concentrations well in excess of those required for saturation of acceptor sites—a region where the expression for binding assumes the form
rCA = CS − CS = K p CS + pCA Equation A.5A.50
where multiplication of the binding function by CA overcomes the problem that the total concentration of receptor sites is one of the parameters to be evaluated from the analysis. From Equation A.5A.50 it is evident that extrapolation of the linear part of the curve at high values of CS to the ordinate provides the value of pCA, the concentration of bound ligand associated with saturation of receptor sites, whereas the partition coefficient (Kp) is obtained from the slope. Once the partition contribution has been defined quantitatively, it can be subtracted from the overall binding curve to allow analysis of the corrected binding curve by standard procedures. Such a procedure can thus be applied to binding curves of the form shown in Figure A.5A.19A, irrespective of the phenomenon responsible for the nonsaturable binding (Winzor and Sawyer, 1995). Protein-Polymer Interactions Interactions between proteins and linear polymer chains such as polynucleotides (Latt and Sober, 1967; McGhee and von Hippel, 1974) and polysaccharides (Olson et al., 1991; Munro et al., 1998) give rise to another form of nonspecificity whenever the site for ligand binding comprises a sequence of two or more consecutive residues on the polymer chain. Because of the similarity of the repeating units, there are no specific acceptor sites, as any residue sequence of the appropriate length is a potential candidate for ligand attachment. Binding curves for such systems are symptomatic of equivalent but negatively cooperative acceptor sites (Fig. A.5A.20A). However, whereas negative cooperativity is usually taken to signify an occupancy-dependent decrease in the affinity of acceptor sites, the phenomenon in protein-polymer interactions merely reflects an occupancy-dependent decrease in the concentration of free acceptor sites that exceeds the concentration of bound ligand. For the binding of ligand to a sequence of m residues of an N-residue acceptor lattice that is governed by intrinsic association constant KAS, the general form of the binding equation (Epstein, 1978) is: i= p
r=
∑ i hCi b KASCS g i =1 i= p
1+
i
∑ i hCi b KASCS g
i
i =1
Equation A.5A.51 Quantitative Characterization of Ligand Binding
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where p is the integer value of N/m, and hCi is the number of possible arrangements of the h items (bound ligands and unoccupied lattice residues) available for distribution on the lattice (Latt and Sober, 1967). The latter is given by: h
a
f a
f
Ci = N − mi + 1 !/ i ! N − mi ! Equation A.5A.52
This combinatorial approach for derivation of the binding equation becomes progressively more cumbersome with increasing length of the acceptor lattice. Furthermore, the need to specify N precludes its use, for example, with DNA as the linear polymer chain because
r/CS (× 10 – 4 M –1)
A
50
25
0 0
0.04
0.12
0.08
Binding function (r )
B
Cooperativity factors in nonspecific binding S
S AS2
ω2KAS
ωKAS
Equilibrium and transient states of acceptor saturation
AS4 AS3
AS2
Figure A.5A.20 Effects of nonspecificity in the binding of ligands to a sequence of residues on a linear polymer chain. (A) Scatchard plot of experimental results (Latt and Sober, 1967) for the interaction of an ε-dinitrophenyl-labeled hexameric lysine oligopeptide with poly(I + C). (B) Schematic representation of thermodynamic and kinetic aspects of nonspecific interaction between a ligand and three-residue sequences on a twelve-residue linear polymer.
Biophysical Methods: Data Analysis
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the number of nucleotides comprising the chain is frequently unknown and almost certainly variable as the result of heterogeneity with respect to polynucleotide chain length. These difficulties may be overcome by switching to a probability approach, which only requires N to be a fairly large number. For the random binding of ligand to m-residue sequences of an infinite lattice, the Scatchard transform of the binding equation (McGhee and von Hippel, 1974) is:
a
r/CS = KAS 1 − mr
FfG 1 − mr IJ m−1 H 1 − am − 1fr K
Equation A.5A.53
which also has the advantage of being a closed solution. Conventional cooperativity is incorporated into the model by designating the modified affinity of a sequence adjacent to a bound ligand as ωKAS, the product of the usual intrinsic binding constant and a dimensionless factor ω, and designating that for a flanked sequence as ω2KAS (Fig. A.5A.20B). The Scatchard formulation of the probability treatment of nonspecific binding (McGhee and von Hippel, 1974) then becomes: r/CS =
a
fa
m −1 2 1 − a m + 1fr + R fa f m −1 2 2aω − 1fa1 − mr f 2a1 − mr f
K AS 1 − mr 2ω + 1 1 − mr + r − R
Equation A.5A.54
where the term R is presented separately for simplification of the typestting of Equation A.5.54.
{ a
f
R = 1− m +1 r
2
a
+ 4ωr 1 − mr
f}1/ 2
Equation A.5A.55
Values of ω greater and less than unity signify classical positive and negative cooperativity, respectively. The application of Equation A.5A.54 is addressed in depth by Kowalczykowski et al. (1986).
Quantitative Characterization of Ligand Binding
Finally, it should be noted that there is a kinetic consideration that may affect the attainment of equilibrium in the binding of ligand to a sequence of acceptor residues—a factor evident from the lower part of Figure A.5A.20B for the binding of ligand to three-residue sequences on a twelve-unit lattice. Clearly, the maximum number of ligands that can attach to the chain is four, but random binding can lead to situations in which the lattice is effectively saturated with only three and even two ligand molecules attached (Fig. A.5A.20B). From the thermodynamic viewpoint, these latter situations are not viable options because they merely represent transient states en route to the equilibrium state of four ligand molecules bound at chain saturation. However, although the time-course of equilibrium attainment is irrelevant to the thermodynamics of nonspecific ligand binding, the experimenter must ensure that sufficient time be allowed for the kinetics of bound ligand rearrangement to take its course before the equilibrium measurements are made for substitution into Equations A.5A.51-A.5A.55 (Epstein, 1979; Munro et al., 1998). In that regard, the kinetic problem is maximal for high levels of acceptor saturation, and therefore impinges on the evaluation of m, the sequence length to which ligand binds. Indeed, because this parameter is often deduced under conditions where the acceptor-li-
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gand interaction is stoichiometric, the endpoint of the titration reflects irreversible rather than equilibrium binding. In such titrations there is effectively no opportunity for the dissociation that is necessary to relocate bound ligate molecules and hence achieve maximal efficiency of site occupancy—the requirement for the equilibrium situation. An algorithm is now available for determining the equilibrium endpoint from the measured, irreversible endpoint of a stoichiometric titration (Munro et al., 1998). CONCLUDING REMARKS In seeking to explain the behavior of plants and animals at a molecular level, biochemists usually adopt a simple reductionist approach and proceed towards an understanding of in vivo complexity. This transition involves a definition of the in vivo state and eventually the matching of in vitro conditions with those existing in biological tissues or fluids. Since specific recognition mechanisms and binding affinities control the biochemical dynamics of living systems, the thermodynamic binding parameters determined under in vitro conditions must be matched to those pertaining to living systems. The acquisition and quantitative analysis of binding data is therefore an essential part of this process. Nature has evolved a variety of complex mechanisms to deal with the interleaving and interdependence of biochemical reactions. While the simple binding function (Equation A.5A.9) remains the starting point for all quantitative analyses of these processes, it is rare that it provides the complete picture. Site heterogeneity, ligand multivalence, ligand and acceptor self-association or isomerization, and the binding of ligands to linear arrays of sites are just a few of the deviations from simple binding behavior that have been discussed in this unit. A common thread in this discussion is the matching of a binding model to experimental results. It has not been the authors’ intention to provide an exhaustive list of the variety of models that have so far been examined in the literature, but rather to illustrate an approach that involves the formulation of a model-dependent binding function and a testing of the experimental data against the derived equations. LITERATURE CITED Adair, G.S. 1925. The hemoglobin system. VI. The oxygen dissociation curve of hemoglobin. J. Biol. Chem. 63:529-545. Anderson, S.R. and Weber, G. 1965. Multiplicity of binding by lactate dehydrogenases. Biochemistry 4:1948-1957. Baghurst, P.A. and Nichol, L.W. 1975. Binding of organic phosphates to human methemoglobin A: Perturbation of the polymerization of proteins by effectors. Biochim. Biophys. Acta 412:168-180. Bergman, D.A. and Winzor, D.J. 1986. Quantitative affinity chromatography: Increased versatility of the technique for studies of ligand binding. Anal. Biochem. 153:380-386. Bergman, D.A. and Winzor, D.J. 1989. Space-filling effects of inert solutes as probes for the detection and study of substrate-mediated conformational changes by enzyme kinetics: Theoretical considerations. J. Theor. Biol. 137:171-191. Bergman, D.A., Shearwin, K.E., and Winzor, D.J. 1989. Effects of thermodynamic nonideality on the kinetics of ester hydrolysis by α-chymotrypsin: A model system with preexistence of the isomerization equilibrium. Arch. Biochem. Biophys. 274:55-63.
Blatt, W.F., Robinson, S.M., and Bixler, H.J. 1968. Membrane ultrafiltration: Diafiltration technique and its application to microsolute exchange and binding phenomena. Anal. Biochem. 26:151-173. Bogoyevitch, M.A., Gillam, E.M.J., Reilly, P.E.B., and Winzor, D.J. 1987. Physical partitioning as the major source of metoprolol uptake by hepatic microsomes. Biochem. Pharmacol. 36:41674168. Brinkworth, R.I., Masters, C.J., and Winzor, D.J. 1975. Evaluation of equilibrium constants for the interaction of lactate dehydrogenase with reduced nicotinamide adenine dinucleotide. Biochem. J. 151:631-636. Calvert, P.D., Nichol, L.W., and Sawyer, W.H. 1979. Binding equations for interacting systems comprising multivalent acceptor and bivalent ligand: Application to antigen-antibody systems. J. Theor. Biol. 80:233-247. Cann, J.R., Nichol, L.W., and Winzor, D.J. 1981. Micellization of chlorpromazine: Implications in the binding of the drug to brain tubulin. Mol. Pharmacol. 20:244-246.
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Changeux, J.P. and Rubin, M.M. 1968. Allosteric interactions in aspartate transcarbamylase. III. Interpretation of experimental data in terms of the model of Monod, Wyman and Changeux. Biochemistry 7:553-561.
Harris, S.J. and Winzor, D.J. 1988. Thermodynamic nonideality as a probe of allosteric mechanisms: Preexistence of the isomerization equilibrium for rabbit muscle pyruvate kinase. Arch. Biochem. Biophys. 265:458-465.
Chatelier, R.C. and Sawyer, W.H. 1987. Isoparametric analysis of binding and partitioning processes. J. Biochem. Biophys. Methods 15:49-61.
Harris, S.J., Jackson, C.M., and Winzor, D.J. 1995a. The rectangular hyperbolic binding equation for multivalent ligands. Arch. Biochem. Biophys. 316:20-23.
Colowick, S.P. and Womack, F.C. 1969. Binding of diffusible molecules: Measurement by rate of dialysis. J. Biol. Chem. 244:774-776. Dunn, B.M. and Chaiken, I.M. 1974. Quantitative affinity chromatography: Determination of binding constants by elution with competitive inhibitors. Proc. Natl. Acad. Sci. U.S.A. 71:2382-2385. Edwards, P.R., Maule, C.H., Leatherbarrow, R.J., and Winzor, D.J. 1998. Second-order kinetic analysis of IAsys biosensor data: Its use and applicability. Anal. Biochem. 263:1-12. Epstein, I.R. 1978. Cooperative and noncooperative binding of large ligands to a finite one-dimensional lattice: A model for ligand-oligonucleotide interactions. Biophys. Chem. 8:327-339. Epstein, I.R. 1979. Kinetics of nucleic acid–large ligand interactions: Exact Monte carlo treatment and limited cases of reversible binding. Biopolymers 18:2037-2050. Ford, C.L. and Winzor, D.J. 1981. A recycling gel partition technique for the study of protein-ligand interactions: The binding of methyl orange to bovine serum albumin. Anal. Biochem. 114:146-152. Ford, C.L. and Winzor, D.J. 1983. Experimental tests of charge conservation in macromolecular interactions. Biochim. Biophys. Acta 756:49-55. Ford, C.L., Winzor, D.J., Nichol, L.W., and Sculley, M.J. 1984. Effects of thermodynamic nonideality in ligand binding studies. Biophys. Chem. 18:1-9. Frieden, C. 1967. Treatment of enzyme kinetic data. II. The multisite case: Comparison of allosteric models and a possible new mechanism. J. Biol. Chem. 242:4045-4052. Gerhart, J.C. 1970. A discussion of the regulatory properties of aspartate transcarbamylase from Escherichia coli. Curr. Top. Cell. Regul. 2:275325. Goodman, D.S. 1958. Interaction of human serum albumin with long-chain fatty acid anions. J. Am. Chem. Soc. 80:3892-3898. Haigh, E.A. and Sawyer, W.H. 1978. Interpretation of double reciprocal plots to determine the spectroscopic parameters of bound ligand for binding assays. Aust. J. Biol. Sci. 31:1-5.
Quantitative Characterization of Ligand Binding
Harris, S.J., Jackson, C.M., and Winzor, D.J. 1995b. Effects of heterogeneity and cooperativity on the forms of binding curves for multivalent ligands. J. Protein Chem. 14:399-407. Heyduk, T. and Lee, J.C. 1990. Application of fluorescence energy transfer and polarization to monitor Escherichia coli cAMP receptor protein and lac promoter interactions. Proc. Natl. Acad. Sci. U.S.A. 87:1744-1748. Hinman, N.D. and Cann, J.R. 1976. Reversible binding of chlorpromazine to brain tubulin. Mol. Pharmacol. 12:769-777. Hogg, P.J. and Winzor, D.J. 1984. Quantitative affinity chromatography: Further developments in the analysis of experimental results from column chromatography and partition equilibrium studies. Arch. Biochem. Biophys. 234:55-60. Hogg, P.J. and Winzor, D.J. 1985. Effects of ligand multivalency in binding studies: A general counterpart of the Scatchard analysis. Biochim. Biophys. Acta 843:159-163. Hogg, P.J., Jackson, C.M., and Winzor, D.J. 1991. Use of quantitative affinity chromatography for characterizing high-affinity interactions: Binding of heparin to antithrombin III. Anal. Biochem. 192:303-311. Holbrook, J.J. 1972. Protein fluorescence of lactate dehydrogenase. Biochem. J. 128:921-931. Holbrook, F.J., Yates, D.W., Reynolds, S.J., Evans, R.W., Greenwood, C., and Gore, M.G. 1972. Protein fluorescence of nicotinamide-dependent dehydrogenases. Biochem. J. 128: 933-940. Jacobsen, M.P. and Winzor, D.J. 1995. Characterization of the interactions of NADH with the dimeric and tetrameric states of methemoglobin. Biochim. Biophys. Acta 1246:17-23. Kalinin, N.L., Ward, L.D., and Winzor, D.J. 1995. Effects of solute valence on the evaluation of binding constants by biosensor technology: Studies with concanavalin A and interleukin-6 as partitioning proteins. Anal. Biochem. 228:238244. Klotz, I.M. 1946. The application of the law of mass action to binding by proteins: Interactions with calcium. Arch. Biochem. 9:109-117.
Hall, D.R. and Winzor, D.J. 1997. Use of a resonant mirror biosensor to characterize the interaction of carboxypeptidase A with an elicited monoclonal antibody. Anal. Biochem. 244:152-160.
Klotz, I.M. 1983. Ligand-receptor interactions: What we can and cannot learn from binding measurements. Trends Pharmacol. Sci. 4:253− 255.
Hall, D.R., and Winzor, D.J. 1998. Potential of biosensor technology for the characterization of interactions by quantitative affinity chromatography. J. Chromatogr., B 715:163−181.
Klotz, I.M. 1997. Ligand-Receptor Energetics: A Guide for the Perplexed. John Wiley & Sons, New York.
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Klotz, I.M., Walker, F.M., and Pivan, R.B. 1946. The binding of organic ions by proteins. J. Am. Chem. Soc. 68:1486-1490. Koshland, D.E., Némethy, G., and Filmer, D. 1966. Comparison of experimental binding data and theoretical models in proteins containing subunits. Biochemistry 5:365-385. Kowalczykowski, S.C., Paul, L.S., Lonberg, N., Newport, J.W., McSwiggen, J.A., and von Hippel, P.H. 1986. Cooperative and noncooperative binding of protein ligands to nucleic acid lattices: Experimental approaches to the determination of thermodynamic parameters. Biochemistry 25:1226-1240. Kuter, M.R., Masters, C.J., and Winzor, D.J. 1983. Equilibrium partition studies of the interaction between aldolase and myofibrils. Arch. Biochem. Biophys. 225:384-389. Latt, S.A. and Sober, H.A. 1967. Protein-oligonucleotide interactions. II. Oligopeptide-polyribonucleotide binding studies. Biochemistry 6:3293-3306. McGhee, J.D. and von Hippel, P.H. 1974. Theoretical aspects of DNA-protein interactions: Cooperative and noncooperative binding of large ligands to a one-dimensional homogeneous lattice. J. Mol. Biol. 86:469-489. Mills, F.C., Johnson, M.L., and Ackers, G.K. 1976. Oxygenation-linked subunit interactions in human hemoglobin: Experimental studies on the concentration dependence of oxygenation curves. Biochemistry 15:5350-5362. Milthorpe, B.K., Jeffrey, P.D., and Nichol, L.W. 1975. Direct analysis of sedimentation equilibrium results obtained with polymerizing systems. Biophys. Chem. 3:169-176. Monod, J., Wyman, J., and Changeux, J.-P. 1965. On the nature of allosteric transitions: A plausible model. J. Mol. Biol. 12:88-118. Munro, P.D., Jackson, C.M., and Winzor, D.J. 1998. On the need to consider kinetic as well as thermodynamic consequences of the parking problem in quantitative studies of nonspecific binding between proteins and linear polymer chains. Biophys. Chem. 71:185-198. Nichol, L.W. and Winzor, D.J. 1964. The determination of equilibrium constants from transport data on rapidly reacting systems of the type A + B ↔ C. J. Phys. Chem. 68:2455-2463. Nichol, L.W., Jackson, W.J.H., and Winzor, D.J. 1967. A theoretical study of the binding of small molecules to a polymerizing protein system: A model for allosteric effects. Biochemistry 6:2449-2456. Nichol, L.W., Smith, G.D., and Ogston, A.G. 1969. Effects of isomerization and polymerization on the binding of ligands to acceptor molecules: Implications in metabolic control. Biochim. Biophys. Acta 184:1-10. Nichol, L.W., Jackson, W.J.H., and Winzor, D.J. 1972a. Preferential binding of competitive inhibitors to the monomeric form of α-chymotrypsin. Biochemistry 11:585-591.
Nichol, L.W., O’Dea, K., and Baghurst, P.A. 1972b. Binding of dissimilar ligand molecules to an interacting acceptor: A model for the action of effectors. J. Theor. Biol. 34:255-263. Nichol, L.W., Ogston, A.G., Winzor, D.J., and Sawyer, W.H. 1974. Evaluation of equilibrium constants by affinity chromatography. Biochem. J. 143:435-443. Nichol, L.W., Ward, L.D., and Winzor, D.J. 1981. Multivalency of the partitioning species in quantitative affinity chromatography: Evaluation of the site-binding constant for the aldolase-phosphate interaction from studies with cellulose phosphate as the affinity matrix. Biochemistry 20:4856-4860. Olson, S.T., Halvorson, H.R., and Björk, I. 1991. Quantitative characterization of the thrombinheparin interaction: Discrimination between specific and nonspecific binding models. J. Biol. Chem. 266:6342-6352. Saroff, H. 1991. Ligand-dependent aggregation and cooperativity : A critique. Biochemistry 30:10085-10090. Scatchard, G. 1949. The attraction of proteins for small molecules and ions. Ann. N.Y. Acad. Sci. 51:660-672. Shanley, B.C., Clarke, K., and Winzor, D.J. 1985. Evaluation of the stoichiometry and strength of chloroquine-porphyrin interactions by difference spectroscopy. Biochem. Pharmacol. 34:141-142. Sophianopoulos, J.A., Durham, S.J., Sophianopoulos, A.J., Ragsdale, H.L., and Cropper, W.P. 1978. Ultrafiltration is theoretically equivalent to equilibrium dialysis but much simpler to carry out. Arch. Biochem. Biophys. 187:132-137. Steinberg, I.Z. and Schachman, H.K. 1966. Ultracentrifugation with absorption optics. V. Analysis of interacting systems involving macromolecules and small molecules. Biochemistry 5:3728-3747. Tellam, R., Winzor, D.J., and Nichol, L.W. 1978. The role of zinc in the stabilization of the dimeric form of α-amylase. Biochem. J. 173:185-190. Tellam, R., de Jersey, J., and Winzor, D.J. 1979. Evaluation of equilibrium constants for the binding of N-acetyl-L-tryptophan to the monomeric and dimeric forms of α-chymotrypsin. Biochemistry 18:5316-5321. Thompson, C.J. and Klotz, I.M. 1971. Macromolecule-small molecule interactions: Analytical and graphical reexamination. Arch. Biochem. Biophys. 147:178-185. Ward, L.D. and Winzor, D.J. 1983. Thermodynamic studies of the activation of rabbit muscle lactate dehydrogenase by phosphate. Biochem. J. 215:685-691. Ward, L.D., Howlett, G.J., Hammacher, A., Weinstock, J., Yasukawa, K., Simpson, R.J., and Winzor, D.J. 1995. Use of a biosensor with surface plasmon resonance detection for the determination of binding constants: Measurement of inter-
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leukin-6 binding to the soluble interleukin-6 receptor. Biochemistry 34:2901-2907.
(T. Kline, ed.) pp. 253-298. Marcel Dekker, New York.
Wills, P.R., Jacobsen, M.P., and Winzor, D.J. 1996. Direct analysis of solute self-association by sedimentation equilibrium. Biopolymers 38:119130.
Winzor, D.J. and Sawyer, W.H. 1995. Quantitative Characterization of Ligand Binding. John Wiley & Sons, New York.
Winzor, D.J. 1997. Quantitative affinity chromatography. In Affinity Separations: A Practical Approach (P. Matejtschuk, ed.) pp. 39-60. IRL Press, Oxford. Winzor, D.J. and Jackson, C.M. 1993. Determination of binding constants by quantitative affinity chromatography: Current and future applications. In Handbook of Affinity Chromatography
Contributed by William H. Sawyer University of Melbourne Melbourne, Australia Donald J. Winzor University of Queensland Brisbane, Australia
Quantitative Characterization of Ligand Binding
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SELECTED SUPPLIERS OF REAGENTS AND EQUIPMENT Listed below are addresses and phone numbers of commercial suppliers who have been recommended for particular items used in our manuals because: (1) the particular brand has actually been found to be of superior quality, or (2) the item is difficult to find in the marketplace. Consequently, this compilation may not include some important vendors of biological supplies. For comprehensive listings, see Linscott’s Directory of Immunological and Biological Reagents (Santa Rosa, CA), The Biotechnology Directory (Stockton Press, New York), the annual Buyers’ Guide supplement to the journal Bio/Technology, as well as various sites on the Internet. A.C. Daniels 72-80 Akeman Street Tring, Hertfordshire, HP23 6AJ, UK (44) 1442 826881 FAX: (44) 1442 826880 A.D. Instruments 5111 Nations Crossing Road #8 Suite 2 Charlotte, NC 28217 (704) 522-8415 FAX: (704) 527-5005 http://www.us.endress.com A.J. Buck 11407 Cronhill Drive Owings Mill, MD 21117 (800) 638-8673 FAX: (410) 581-1809 (410) 581-1800 http://www.ajbuck.com A.M. Systems 131 Business Park Loop P.O. Box 850 Carlsborg, WA 98324 (800) 426-1306 FAX: (360) 683-3525 (360) 683-8300 http://www.a-msystems.com Aaron Medical Industries 7100 30th Avenue North St. Petersburg, FL 33710 (727) 384-2323 FAX: (727) 347-9144 http://www.aaronmed.com Abbott Laboratories 100 Abbott Park Road Abbott Park, IL 60064 (800) 323-9100 FAX: (847) 938-7424 http://www.abbott.com ABCO Dealers 55 Church Street Central Plaza Lowell, MA 01852 (800) 462-3326 (978) 459-6101 http://www.lomedco.com/abco.htm Aber Instruments 5 Science Park Aberystwyth, Wales SY23 3AH, UK (44) 1970 636300 FAX: (44) 1970 615455 http://www.aber-instruments.co.uk ABI Biotechnologies See Perkin-Elmer ABI Biotechnology See Apotex
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Advanced Separation Technologies 37 Leslie Court P.O. Box 297 Whippany, NJ 07981 (973) 428-9080 FAX: (973) 428-0152 http://www.astecusa.com
Advanced Process Supply See Naz-Dar-KC Chicago
Advanced Targeting Systems 11175-A Flintkote Avenue San Diego, CA 92121 (877) 889-2288 FAX: (858) 642-1989 (858) 642-1988 http://www.ATSbio.com Advent Research Materials Eynsham, Oxford OX29 4JA, UK (44) 1865-884440 FAX: (44) 1865-84460 http://www.advent-rm.com Advet Industrivagen 24 S-972 54 Lulea, Sweden (46) 0920-211887 FAX: (46) 0920-13773 Aesculap 1000 Gateway Boulevard South San Francisco, CA 94080 (800) 282-9000 http://www.aesculap.com Affinity Chromatography 307 Huntingdon Road Girton, Cambridge CB3 OJX, UK (44) 1223 277192 FAX: (44) 1223 277502 http://www.affinity-chrom.com Affinity Sensors See Labsystems Affinity Sensors Affymetrix 3380 Central Expressway Santa Clara, CA 95051 (408) 731-5000 FAX: (408) 481-0422 (800) 362-2447 http://www.affymetrix.com Agar Scientific 66a Cambridge Road Stansted CM24 8DA, UK (44) 1279-813-519 FAX: (44) 1279-815-106 http://www.agarscientific.com A/G Technology 101 Hampton Avenue Needham, MA 02494 (800) AGT-2535 FAX: (781) 449-5786 (781) 449-5774 http://www.agtech.com
Suppliers
1 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
Agen Biomedical Limited 11 Durbell Street P.O. Box 391 Acacia Ridge 4110 Brisbane, Australia 61-7-3370-6300 FAX:61-7-3370-6370 http://www.agen.com Agilent Technologies 395 Page Mill Road P.O. Box 10395 Palo Alto, CA 94306 (650) 752-5000 http://www.agilent.com/chem Agouron Pharmaceuticals 10350 N. Torrey Pines Road La Jolla, CA 92037 (858) 622-3000 FAX: (858) 622-3298 http://www.agouron.com Agracetus 8520 University Green Middleton, WI 53562 (608) 836-7300 FAX: (608) 836-9710 http://www.monsanto.com AIDS Research and Reference Reagent Program U.S. Department of Health and Human Services 625 Lofstrand Lane Rockville, MD 20850 (301) 340-0245 FAX: (301) 340-9245 http://www.aidsreagent.org
Alcide 8561 154th Avenue NE Redmond, WA 98052 (800) 543-2133 FAX: (425) 861-0173 (425) 882-2555 http://www.alcide.com Aldevron 3233 15th Street, South Fargo, ND 58104 (877) Pure-DNA FAX: (701) 280-1642 (701) 297-9256 http://www.aldevron.com Aldrich Chemical P.O. Box 2060 Milwaukee, WI 53201 (800) 558-9160 FAX: (800) 962-9591 (414) 273-3850 FAX: (414) 273-4979 http://www.aldrich.sial.com Alexis Biochemicals 6181 Cornerstone Court East, Suite 103 San Diego, CA 92121 (800) 900-0065 FAX: (858) 658-9224 (858) 658-0065 http://www.alexis-corp.com Alfa Aesar 30 Bond Street Ward Hill, MA 10835 (800) 343-0660 FAX: (800) 322-4757 (978) 521-6300 FAX: (978) 521-6350 http://www.alfa.com
AIN Plastics 249 East Sanford Boulevard P.O. Box 151 Mt. Vernon, NY 10550 (914) 668-6800 FAX: (914) 668-8820 http://www.tincna.com
Alfa Laval Avenue de Ble 5 - Bazellaan 5 BE-1140 Brussels, Belgium 32(2) 728 3811 FAX:32(2) 728 3917 or 32(2) 728 3985 http://www.alfalaval.com
Air Products and Chemicals 7201 Hamilton Boulevard Allentown, PA 18195 (800) 345-3148 FAX: (610) 481-4381 (610) 481-6799 http://www.airproducts.com
Alice King Chatham Medical Arts 11915-17 Inglewood Avenue Hawthorne, CA 90250 (310) 970-1834 FAX: (310) 970-0121 (310) 970-1063
ALA Scientific Instruments 1100 Shames Drive Westbury, NY 11590 (516) 997-5780 FAX: (516) 997-0528 http://www.alascience.com Aladin Enterprises 1255 23rd Avenue San Francisco, CA 94122 (415) 468-0433 FAX: (415) 468-5607 Aladdin Systems 165 Westridge Drive Watsonville, CA 95076 (831) 761-6200 FAX: (831) 761-6206 http://www.aladdinsys.com
Allegiance Healthcare 800-964-5227 http://www.allegiance.net
Alltech Associates Applied Science Labs 2051 Waukegan Road P.O. Box 23 Deerfield, IL 60015 (800) 255-8324 FAX: (847) 948-1078 (847) 948-8600 http://www.alltechweb.com
American Association of Blood Banks College of American Pathologists 325 Waukegan Road Northfield, IL 60093 (800) 323-4040 FAX: (847) 8166 (847) 832-7000 http://www.cap.org
Alomone Labs HaMarpeh 5 P.O. Box 4287 Jerusalem 91042, Israel 972-2-587-2202 FAX:972-2-587-1101 US: (800) 791-3904 FAX: (800) 791-3912 http://www.alomone.com
American Bio-Technologies See Intracel Corporation
Alpha Innotech 14743 Catalina Street San Leandro, CA 94577 (800) 795-5556 FAX: (510) 483-3227 (510) 483-9620 http://www.alphainnotech.com
American Cyanamid P.O. Box 400 Princeton, NJ 08543 (609) 799-0400 FAX: (609) 275-3502 http://www.cyanamid.com
Altec Plastics 116 B Street Boston, MA 02127 (800) 477-8196 FAX: (617) 269-8484 (617) 269-1400 Alza 1900 Charleston Road P.O. Box 7210 Mountain View, CA 94043 (800) 692-2990 FAX: (650) 564-7070 (650) 564-5000 http://www.alza.com Alzet c/o Durect Corporation P.O. Box 530 10240 Bubo Road Cupertino, CA 95015 (800) 692-2990 (408) 367-4036 FAX: (408) 865-1406 http://www.alzet.com Amac 160B Larrabee Road Westbrook, ME 04092 (800) 458-5060 FAX: (207) 854-0116 (207) 854-0426
Allelix Biopharmaceuticals 6850 Gorway Drive Mississauga, Ontario L4V 1V7 Canada (905) 677-0831 FAX: (905) 677-9595 http://www.allelix.com
Amaresco 30175 Solon Industrial Parkway Solon, Ohio 44139 (800) 366-1313 FAX: (440) 349-1182 (440) 349-1313
Allentown Caging Equipment Route 526, P.O. Box 698 Allentown, NJ 08501 (800) 762-CAGE FAX: (609) 259-0449 (609) 259-7951 http://www.acecaging.com
Ambion 2130 Woodward Street, Suite 200 Austin, TX 78744 (800) 888-8804 FAX: (512) 651-0190 (512) 651-0200 http://www.ambion.com
American Bioanalytical 15 Erie Drive Natick, MA 01760 (800) 443-0600 FAX: (508) 655-2754 (508) 655-4336 http://www.americanbio.com
American HistoLabs 7605-F Airpark Road Gaithersburg, MD 20879 (301) 330-1200 FAX: (301) 330-6059 American International Chemical 17 Strathmore Road Natick, MA 01760 (800) 238-0001 (508) 655-5805 http://www.aicma.com American Laboratory Supply See American Bioanalytical American Medical Systems 10700 Bren Road West Minnetonka, MN 55343 (800) 328-3881 FAX: (612) 930-6654 (612) 933-4666 http://www.visitams.com American Qualex 920-A Calle Negocio San Clemente, CA 92673 (949) 492-8298 FAX: (949) 492-6790 http://www.americanqualex.com American Radiolabeled Chemicals 11624 Bowling Green St. Louis, MO 63146 (800) 331-6661 FAX: (800) 999-9925 (314) 991-4545 FAX: (314) 991-4692 http://www.arc-inc.com American Scientific Products See VWR Scientific Products American Society for Histocompatibility and Immunogenetics P.O. Box 15804 Lenexa, KS 66285 (913) 541-0009 FAX: (913) 541-0156 http://www.swmed.edu/home pages/ ASHI/ashi.htm
Suppliers
2 CPPS Supplement 45
Current Protocols Selected Suppliers of Reagents and Equipment
American Type Culture Collection (ATCC) 10801 University Boulevard Manassas, VA 20110 (800) 638-6597 FAX: (703) 365-2750 (703) 365-2700 http://www.atcc.org Amersham See Amersham Pharmacia Biotech Amersham International Amersham Place Little Chalfont, Buckinghamshire HP7 9NA, UK (44) 1494-544100 FAX: (44) 1494-544350 http://www.apbiotech.com
Amresco 30175 Solon Industrial Parkway Solon, OH 44139 (800) 829-2805 FAX: (440) 349-1182 (440) 349-1199 Anachem Ltd Anachem House Charles Street Luton, Beds LU2 0EB, UK Anachemia Chemicals 3 Lincoln Boulevard Rouses Point, NY 12979 (800) 323-1414 FAX: (518) 462-1952 (518) 462-1066 http://www.anachemia.com
Amersham Medi-Physics Also see Nycomed Amersham 3350 North Ridge Avenue Arlington Heights, IL 60004 (800) 292-8514 FAX: (800) 807-2382 http://www.nycomed-amersham.com
Ana-Gen Technologies 4015 Fabian Way Palo Alto, CA 94303 (800) 654-4671 FAX: (650) 494-3893 (650) 494-3894 http://www.ana-gen.com
Amersham Pharmacia Biotech 800 Centennial Avenue P.O. Box 1327 Piscataway, NJ 08855 (800) 526-3593 FAX: (877) 295-8102 (732) 457-8000 http://www.apbiotech.com
Analox Instruments USA P.O. Box 208 Lunenburg, MA 01462 (978) 582-9368 FAX: (978) 582-9588 http://www.analox.com
Amgen 1 Amgen Center Drive Thousand Oaks, CA 91320 (800) 926-4369 FAX: (805) 498-9377 (805) 447-5725 http://www.amgen.com Amicon Scientific Systems Division 72 Cherry Hill Drive Beverly, MA 01915 (800) 426-4266 FAX: (978) 777-6204 (978) 777-3622 http://www.amicon.com Amika 8980F Route 108 Oakland Center Columbia, MD 21045 (800) 547-6766 FAX: (410) 997-7104 (410) 997-0100 http://www.amika.com Amoco Performance Products See BPAmoco AMPI See Pacer Scientific Amrad 576 Swan Street Richmond, Victoria 3121, Australia 613-9208-4000 FAX:613-9208-4350 http://www.amrad.com.au
Analytical Biological Services Cornell Business Park 701-4 Wilmington, DE 19801 (800) 391-2391 FAX: (302) 654-8046 (302) 654-4492 http://www.ABSbioreagents.com Analytical Genetics Testing Center 7808 Cherry Creek S. Drive, Suite 201 Denver, CO 80231 (800) 204-4721 FAX: (303) 750-2171 (303) 750-2023 http://www.geneticid.com AnaSpec 2149 O’Toole Avenue, Suite F San Jose, CA 95131 (800) 452-5530 FAX: (408) 452-5059 (408) 452-5055 http://www.anaspec.com Ancare 2647 Grand Avenue P.O. Box 814 Bellmore, NY 11710 (800) 645-6379 FAX: (516) 781-4937 (516) 781-0755 http://www.ancare.com Ancell 243 Third Street North P.O. Box 87 Bayport, MN 55033 (800) 374-9523 FAX: (651) 439-1940 (651) 439-0835 http://www.ancell.com
Anderson Instruments 500 Technology Court Smyrna, GA 30082 (800) 241-6898 FAX: (770) 319-5306 (770) 319-9999 http://www.graseby.com Andreas Hettich Gartenstrasse 100 Postfach 260 D-78732 Tuttlingen, Germany (49) 7461 705 0 FAX: (49) 7461 705-122 http://www.hettich-centrifugen.de Anesthetic Vaporizer Services 10185 Main Street Clarence, NY 14031 (719) 759-8490 http://www.avapor.com Animal Identification and Marking Systems (AIMS) 13 Winchester Avenue Budd Lake, NJ 07828 (908) 684-9105 FAX: (908) 684-9106 http://www.animalid.com Annovis 34 Mount Pleasant Drive Aston, PA 19014 (800) EASY-DNA FAX: (610) 361-8255 (610) 361-9224 http://www.annovis.com Apotex 150 Signet Drive Weston, Ontario M9L 1T9, Canada (416) 749-9300 FAX: (416) 749-2646 http://www.apotex.com Apple Scientific 11711 Chillicothe Road, Unit 2 P.O. Box 778 Chesterland, OH 44026 (440) 729-3056 FAX: (440) 729-0928 http://www.applesci.com Applied Biosystems See PE Biosystems Applied Imaging 2380 Walsh Avenue, Bldg. B Santa Clara, CA 95051 (800) 634-3622 FAX: (408) 562-0264 (408) 562-0250 http://www.aicorp.com Applied Photophysics 203-205 Kingston Road Leatherhead, Surrey, KT22 7PB UK (44) 1372-386537 Applied Precision 1040 12th Avenue Northwest Issaquah, Washington 98027 (425) 557-1000 FAX: (425) 557-1055 http://www.api.com/index.html
Applied Separations 930 Hamilton Street Allentown, PA 18101 (610) 770-0900 FAX: (610) 740-5520 Appligene Oncor Parc d’Innovation Rue Geiler de Kaysersberg, BP 72 67402 Illkirch Cedex, France (33) 88 67 22 67 FAX: (33) 88 67 19 45 http://www.oncor.com/prod-app.htm Applikon 1165 Chess Drive, Suite G Foster City, CA 94404 (650) 578-1396 FAX: (650) 578-8836 http://www.applikon.com Appropriate Technical Resources 9157 Whiskey Bottom Road Laurel, MD 20723 (800) 827-5931 FAX: (410) 792-2837 http://www.atrbiotech.com APV Gaulin 100 S. CP Avenue Lake Mills, WI 53551 (888) 278-4321 FAX: (888) 278-5329 http://www.apv.com Aqualon See Hercules Aqualon Aquarium Systems B141 Tyler Boulevard Mentor, OH 44060 (800) 822-1100 FAX: (440) 255-8994 (440) 255-1997 http://www.aquariumsystems.com Aquebogue Machine and Repair Shop Box 2055 Main Road Aquebogue, NY 11931 (631) 722-3635 FAX: (631) 722-3106 Archer Daniels Midland 4666 Faries Parkway Decatur, IL 62525 (217) 424-5200 http://www.admworld.com Archimica Florida P.O. Box 1466 Gainesville, FL 32602 (800) 331-6313 FAX: (352) 371-6246 (352) 376-8246 http://www.archimica.com Arcor Electronics 1845 Oak Street #15 Northfield, IL 60093 (847) 501-4848 Arcturus Engineering 400 Logue Avenue Mountain View, CA 94043 (888) 446 7911 FAX: (650) 962 3039 (650) 962 3020 http://www.arctur.com
Suppliers
3 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
Ardals Corporation One Ledgemont Center 128 Spring Street Lexington, MA 02421 (781) 274-6420 (781) 274-6421 http://www.ardais.com Argonaut Technologies 887 Industrial Road, Suite G San Carlos, CA 94070 (650) 998-1350 FAX: (650) 598-1359 http://www.argotech.com Ariad Pharmaceuticals 26 Landsdowne Street Cambridge, MA 02139 (617) 494-0400 FAX: (617) 494-8144 http://www.ariad.com Armour Pharmaceuticals See Rhone-Poulenc Rorer Aronex Pharmaceuticals 8707 Technology Forest Place The Woodlands, TX 77381 (281) 367-1666 FAX: (281) 367-1676 http://www.aronex.com Artisan Industries 73 Pond Street Waltham, MA 02254 (617) 893-6800 http://www.artisanind.com ASI Instruments 12900 Ten Mile Road Warren, MI 48089 (800) 531-1105 FAX: (810) 756-9737 (810) 756-1222 http://www.asi-instruments.com Aspen Research Laboratories 1700 Buerkle Road White Bear Lake, MN 55140 (651) 264-6000 FAX: (651) 264-6270 http://www.aspenresearch.com Associates of Cape Cod 704 Main Street Falmouth, MA 02540 (800) LAL-TEST FAX: (508) 540-8680 (508) 540-3444 http://www.acciusa.com Astra Pharmaceuticals See AstraZeneca AstraZeneca 1800 Concord Pike Wilmington, DE 19850 (302) 886-3000 FAX: (302) 886-2972 http://www.astrazeneca.com AT Biochem 30 Spring Mill Drive Malvern, PA 19355 (610) 889-9300 FAX: (610) 889-9304 ATC Diagnostics See Vysis ATCC See American Type Culture Collection
Athens Research and Technology P.O. Box 5494 Athens, GA 30604 (706) 546-0207 FAX: (706) 546-7395 Atlanta Biologicals 1425-400 Oakbrook Drive Norcross, GA 30093 (800) 780-7788 or (770) 446-1404 FAX: (800) 780-7374 or (770) 446-1404 http://www.atlantabio.com Atomergic Chemical 71 Carolyn Boulevard Farmingdale, NY 11735 (631) 694-9000 FAX: (631) 694-9177 http://www.atomergic.com Atomic Energy of Canada 2251 Speakman Drive Mississauga, Ontario L5K 1B2 Canada (905) 823-9040 FAX: (905) 823-1290 http://www.aecl.ca ATR P.O. Box 460 Laurel, MD 20725 (800) 827-5931 FAX: (410) 792-2837 (301) 470-2799 http://www.atrbiotech.com Aurora Biosciences 11010 Torreyana Road San Diego, CA 92121 (858) 404-6600 FAX: (858) 404-6714 http://www.aurorabio.com Automatic Switch Company A Division of Emerson Electric 50 Hanover Road Florham Park, NJ 07932 (800) 937-2726 FAX: (973) 966-2628 (973) 966-2000 http://www.asco.com Avanti Polar Lipids 700 Industrial Park Drive Alabaster, AL 35007 (800) 227-0651 FAX: (800) 229-1004 (205) 663-2494 FAX: (205) 663-0756 http://www.avantilipids.com Aventis BP 67917 67917 Strasbourg Cedex 9, France 33 (0) 388 99 11 00 FAX:33 (0) 388 99 11 01 http://www.aventis.com Aventis Pasteur 1 Discovery Drive Swiftwater, PA 18370 (800) 822-2463 FAX: (570) 839-0955 (570) 839-7187 http://www.aventispasteur.com/usa
Avery Dennison 150 North Orange Grove Boulevard Pasadena, CA 91103 (800) 462-8379 FAX: (626) 792-7312 (626) 304-2000 http://www.averydennison.com Avestin 2450 Don Reid Drive Ottawa, Ontario K1H 1E1, Canada (888) AVESTIN FAX: (613) 736-8086 (613) 736-0019 http://www.avestin.com AVIV Instruments 750 Vassar Avenue Lakewood, NJ 08701 (732) 367-1663 FAX: (732) 370-0032 http://www.avivinst.com Axon Instruments 1101 Chess Drive Foster City, CA 94404 (650) 571-9400 FAX: (650) 571-9500 http://www.axon.com Azon 720 Azon Road Johnson City, NY 13790 (800) 847-9374 FAX: (800) 635-6042 (607) 797-2368 http://www.azon.com BAbCO 1223 South 47th Street Richmond, CA 94804 (800) 92-BABCO FAX: (510) 412-8940 (510) 412-8930 http://www.babco.com Bacharach 625 Alpha Drive Pittsburgh, PA 15238 (800) 736-4666 FAX: (412) 963-2091 (412) 963-2000 http://www.bacharach-inc.com Bachem Bioscience 3700 Horizon Drive King of Prussia, PA 19406 (800) 634-3183 FAX: (610) 239-0800 (610) 239-0300 http://www.bachem.com Bachem California 3132 Kashiwa Street P.O. Box 3426 Torrance, CA 90510 (800) 422-2436 FAX: (310) 530-1571 (310) 539-4171 http://www.bachem.com Baekon 18866 Allendale Avenue Saratoga, CA 95070 (408) 972-8779 FAX: (408) 741-0944 Baker Chemical See J.T. Baker
Bangs Laboratories 9025 Technology Drive Fishers, IN 46038 (317) 570-7020 FAX: (317) 570-7034 www.bangslabs.com Bard Parker See Becton Dickinson Barnstead/Thermolyne P.O. Box 797 2555 Kerper Boulevard Dubuque, IA 52004 (800) 446-6060 FAX: (319) 589-0516 http://www.barnstead.com Barrskogen 4612 Laverock Place N Washington, DC 20007 (800) 237-9192 FAX: (301) 464-7347 BAS See Bioanalytical Systems BASF Specialty Products 3000 Continental Drive North Mt. Olive, NJ 07828 (800) 669-2273 FAX: (973) 426-2610 http://www.basf.com Baum, W.A. 620 Oak Street Copiague, NY 11726 (631) 226-3940 FAX: (631) 226-3969 http://www.wabaum.com Bausch & Lomb One Bausch & Lomb Place Rochester, NY 14604 (800) 344-8815 FAX: (716) 338-6007 (716) 338-6000 http://www.bausch.com Baxter Fenwal Division 1627 Lake Cook Road Deerfield, IL 60015 (800) 766-1077 FAX: (800) 395-3291 (847) 940-6599 FAX: (847) 940-5766 http://www.powerfulmedicine.com Baxter Healthcare One Baxter Parkway Deerfield, IL 60015 (800) 777-2298 FAX: (847) 948-3948 (847) 948-2000 http://www.baxter.com Baxter Scientific Products See VWR Scientific Bayer Agricultural Division Animal Health Products 12707 Shawnee Mission Pkwy. Shawnee Mission, KS 66201 (800) 255-6517 FAX: (913) 268-2803 (913) 268-2000 http://www.bayerus.com
Suppliers
4 CPPS Supplement 45
Current Protocols Selected Suppliers of Reagents and Equipment
Bayer Diagnostics Division (Order Services) P.O. Box 2009 Mishiwaka, IN 46546 (800) 248-2637 FAX: (800) 863-6882 (219) 256-3390 http://www.bayer.com
Beckman Instruments Spinco Division/Bioproducts Operation 1050 Page Mill Road Palo Alto, CA 94304 (800) 742-2345 FAX: (415) 859-1550 (415) 857-1150 http://www.beckman-coulter.com
Bayer Diagnostics 511 Benedict Avenue Tarrytown, NY 10591 (800) 255-3232 FAX: (914) 524-2132 (914) 631-8000 http://www.bayerdiag.com
Becton Dickinson Immunocytometry & Cellular Imaging 2350 Qume Drive San Jose, CA 95131 (800) 223-8226 FAX: (408) 954-2007 (408) 432-9475 http://www.bdfacs.com
Bayer Plc Diagnostics Division Bayer House, Strawberry Hill Newbury, Berkshire RG14 1JA, UK (44) 1635-563000 FAX: (44) 1635-563393 http://www.bayer.co.uk
Becton Dickinson Labware 1 Becton Drive Franklin Lakes, NJ 07417 (888) 237-2762 FAX: (800) 847-2220 (201) 847-4222 http://www.bdfacs.com
BD Immunocytometry Systems 2350 Qume Drive San Jose, CA 95131 (800) 223-8226 FAX: (408) 954-BDIS http://www.bdfacs.com BD Labware Two Oak Park Bedford, MA 01730 (800) 343-2035 FAX: (800) 743-6200 http://www.bd.com/labware BD PharMingen 10975 Torreyana Road San Diego, CA 92121 (800) 848-6227 FAX: (858) 812-8888 (858) 812-8800 http://www.pharmingen.com BD Transduction Laboratories 133 Venture Court Lexington, KY 40511 (800) 227-4063 FAX: (606) 259-1413 (606) 259-1550 http://www.translab.com BDH Chemicals Broom Road Poole, Dorset BH12 4NN, UK (44) 1202-745520 FAX: (44) 1202- 2413720 BDH Chemicals See Hoefer Scientific Instruments BDIS See BD Immunocytometry Systems Beckman Coulter 4300 North Harbor Boulevard Fullerton, CA 92834 (800) 233-4685 FAX: (800) 643-4366 (714) 871-4848 http://www.beckman-coulter.com
Becton Dickinson Labware 2 Bridgewater Lane Lincoln Park, NJ 07035 (800) 235-5953 FAX: (800) 847-2220 (201) 847-4222 http://www.bdfacs.com Becton Dickinson Primary Care Diagnostics 7 Loveton Circle Sparks, MD 21152 (800) 675-0908 FAX: (410) 316-4723 (410) 316-4000 http://www.bdfacs.com Behringwerke Diagnostika Hoechster Strasse 70 P-65835 Liederback, Germany (49) 69-30511 FAX: (49) 69-303-834 Bellco Glass 340 Edrudo Road Vineland, NJ 08360 (800) 257-7043 FAX: (856) 691-3247 (856) 691-1075 http://www.bellcoglass.com Bender Biosystems See Serva Beral Enterprises See Garren Scientific Berkeley Antibody See BAbCO Bernsco Surgical Supply 25 Plant Avenue Hauppague, NY 11788 (800) TIEMANN FAX: (516) 273-6199 (516) 273-0005 http://www.bernsco.com Beta Medical and Scientific (Datesand Ltd.) 2 Ferndale Road Sale, Manchester M33 3GP, UK (44) 1612 317676 FAX: (44) 1612 313656
Bethesda Research Laboratories (BRL) See Life Technologies Biacore 200 Centennial Avenue, Suite 100 Piscataway, NJ 08854 (800) 242-2599 FAX: (732) 885-5669 (732) 885-5618 http://www.biacore.com Bilaney Consultants St. Julian’s Sevenoaks, Kent TN15 0RX, UK (44) 1732 450002 FAX: (44) 1732 450003 http://www.bilaney.com Binding Site 5889 Oberlin Drive, Suite 101 San Diego, CA 92121 (800) 633-4484 FAX: (619) 453-9189 (619) 453-9177 http://www.bindingsite.co.uk BIO 101 See Qbiogene Bio Image See Genomic Solutions Bioanalytical Systems 2701 Kent Avenue West Lafayette, IN 47906 (800) 845-4246 FAX: (765) 497-1102 (765) 463-4527 http://www.bioanalytical.com Biocell 2001 University Drive Rancho Dominguez, CA 90220 (800) 222-8382 FAX: (310) 637-3927 (310) 537-3300 http://www.biocell.com Biocoat See BD Labware BioComp Instruments 650 Churchill Road Fredericton, New Brunswick E3B 1P6 Canada (800) 561-4221 FAX: (506) 453-3583 (506) 453-4812 http://131.202.97.21 BioDesign P.O. Box 1050 Carmel, NY 10512 (914) 454-6610 FAX: (914) 454-6077 http://www.biodesignofny.com
Biofluids Division of Biosource International 1114 Taft Street Rockville, MD 20850 (800) 972-5200 FAX: (301) 424-3619 (301) 424-4140 http://www.biosource.com BioFX Laboratories 9633 Liberty Road, Suite S Randallstown, MD 21133 (800) 445-6447 FAX: (410) 498-6008 (410) 496-6006 http://www.biofx.com BioGenex Laboratories 4600 Norris Canyon Road San Ramon, CA 94583 (800) 421-4149 FAX: (925) 275-0580 (925) 275-0550 http://www.biogenex.com Bioline 2470 Wrondel Way Reno, NV 89502 (888) 257-5155 FAX: (775) 828-7676 (775) 828-0202 http://www.bioline.com Bio-Logic Research & Development 1, rue de l-Europe A.Z. de Font-Ratel 38640 CLAIX, France (33) 76-98-68-31 FAX: (33) 76-98-69-09 Biological Detection Systems See Cellomics or Amersham Biomeda 1166 Triton Drive, Suite E P.O. Box 8045 Foster City, CA 94404 (800) 341-8787 FAX: (650) 341-2299 (650) 341-8787 http://www.biomeda.com BioMedic Data Systems 1 Silas Road Seaford, DE 19973 (800) 526-2637 FAX: (302) 628-4110 (302) 628-4100 http://www.bmds.com Biomedical Engineering P.O. Box 980694 Virginia Commonwealth University Richmond, VA 23298 (804) 828-9829 FAX: (804) 828-1008
BioDiscovery 4640 Admiralty Way, Suite 710 Marina Del Rey, CA 90292 (310) 306-9310 FAX: (310) 306-9109 http://www.biodiscovery.com
Biomedical Research Instruments 12264 Wilkins Avenue Rockville, MD 20852 (800) 327-9498 (301) 881-7911 http://www.biomedinstr.com
Bioengineering AG Sagenrainstrasse 7 CH8636 Wald, Switzerland (41) 55-256-8-111 FAX: (41) 55-256-8-256
Bio/medical Specialties P.O. Box 1687 Santa Monica, CA 90406 (800) 269-1158 FAX: (800) 269-1158 (323) 938-7515
Suppliers
5 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
BioMerieux 100 Rodolphe Street Durham, North Carolina 27712 (919) 620-2000 http://www.biomerieux.com BioMetallics P.O. Box 2251 Princeton, NJ 08543 (800) 999-1961 FAX: (609) 275-9485 (609) 275-0133 http://www.microplate.com Biomol Research Laboratories 5100 Campus Drive Plymouth Meeting, PA 19462 (800) 942-0430 FAX: (610) 941-9252 (610) 941-0430 http://www.biomol.com Bionique Testing Labs Fay Brook Drive RR 1, Box 196 Saranac Lake, NY 12983 (518) 891-2356 FAX: (518) 891-5753 http://www.bionique.com Biopac Systems 42 Aero Camino Santa Barbara, CA 93117 (805) 685-0066 FAX: (805) 685-0067 http://www.biopac.com Bioproducts for Science See Harlan Bioproducts for Science Bioptechs 3560 Beck Road Butler, PA 16002 (877) 548-3235 FAX: (724) 282-0745 (724) 282-7145 http://www.bioptechs.com BIOQUANT-R&M Biometrics 5611 Ohio Avenue Nashville, TN 37209 (800) 221-0549 (615) 350-7866 FAX: (615) 350-7282 http://www.bioquant.com Bio-Rad Laboratories 2000 Alfred Nobel Drive Hercules, CA 94547 (800) 424-6723 FAX: (800) 879-2289 (510) 741-1000 FAX: (510) 741-5800 http://www.bio-rad.com Bio-Rad Laboratories Maylands Avenue Hemel Hempstead, Herts HP2 7TD, UK http://www.bio-rad.com Bioreclamation 492 Richmond Road East Meadow, NY 11554 (516) 483-1196 FAX: (516) 483-4683 http://www.bioreclamation.com
BioRobotics 3-4 Bennell Court Comberton, Cambridge CB3 7DS, UK (44) 1223-264345 FAX: (44) 1223-263933 http://www.biorobotics.co.uk BIOS Laboratories See Genaissance Pharmaceuticals Biosearch Technologies 81 Digital Drive Novato, CA 94949 (800) GENOME1 FAX: (415) 883-8488 (415) 883-8400 http://www.biosearchtech.com BioSepra 111 Locke Drive Marlborough, MA 01752 (800) 752-5277 FAX: (508) 357-7595 (508) 357-7500 http://www.biosepra.com Bio-Serv 18th Street, Suite 1 Frenchtown, NJ 08825 (908) 996-2155 FAX: (908) 996-4123 http://www.bio-serv.com BioSignal 1744 William Street, Suite 600 Montreal, Quebec H3J 1R4 Canada (800) 293-4501 FAX: (514) 937-0777 (514) 937-1010 http://www.biosignal.com
Biotech Instruments Biotech House 75A High Street Kimpton, Hertfordshire SG4 8PU, UK (44) 1438 832555 FAX: (44) 1438 833040 http://www.biotinst.demon.co.uk Biotech International 11 Durbell Street Acacia Ridge, Queensland 4110 Australia 61-7-3370-6396 FAX:61-7-3370-6370 http://www.avianbiotech.com Biotech Source Inland Farm Drive South Windham, ME 04062 (207) 892-3266 FAX: (207) 892-6774 Bio-Tek Instruments Highland Industrial Park P.O. Box 998 Winooski, VT 05404 (800) 451-5172 FAX: (802) 655-7941 (802) 655-4040 http://www.biotek.com Biotecx Laboratories 6023 South Loop East Houston, TX 77033 (800) 535-6286 FAX: (713) 643-3143 (713) 643-0606 http://www.biotecx.com
Biosoft P.O. Box 10938 Ferguson, MO 63135 (314) 524-8029 FAX: (314) 524-8129 http://www.biosoft.com
BioTherm 3260 Wilson Boulevard Arlington, VA 22201 (703) 522-1705 FAX: (703) 522-2606
Biosource International 820 Flynn Road Camarillo, CA 93012 (800) 242-0607 FAX: (805) 987-3385 (805) 987-0086 http://www.biosource.com
Bioventures P.O. Box 2561 848 Scott Street Murfreesboro, TN 37133 (800) 235-8938 FAX: (615) 896-4837 http://www.bioventures.com
BioSpec Products P.O. Box 788 Bartlesville, OK 74005 (800) 617-3363 FAX: (918) 336-3363 (918) 336-3363 http://www.biospec.com
BioWhittaker 8830 Biggs Ford Road P.O. Box 127 Walkersville, MD 21793 (800) 638-8174 FAX: (301) 845-8338 (301) 898-7025 http://www.biowhittaker.com
Biosure See Riese Enterprises Biosym Technologies See Molecular Simulations Biosys 21 quai du Clos des Roses 602000 Compiegne, France (33) 03 4486 2275 FAX: (33) 03 4484 2297 Bio-Tech Research Laboratories NIAID Repository Rockville, MD 20850 http://www.niaid.nih.gov/ncn/repos.htm
B & K Universal 2403 Yale Way Fremont, CA 94538 (800) USA-MICE FAX: (510) 490-3036 BLS Ltd. Zselyi Aladar u. 31 1165 Budapest, Hungary (36) 1-407-2602 FAX: (36) 1-407-2896 http://www.bls-ltd.com Blue Sky Research 3047 Orchard Parkway San Jose, CA 95134 (408) 474-0988 FAX: (408) 474-0989 http://www.blueskyresearch.com Blumenthal Industries 7 West 36th Street, 13th floor New York, NY 10018 (212) 719-1251 FAX: (212) 594-8828 BOC Edwards One Edwards Park 301 Ballardvale Street Wilmington, MA 01887 (800) 848-9800 FAX: (978) 658-7969 (978) 658-5410 http://www.bocedwards.com Boehringer Ingelheim 900 Ridgebury Road P.O. Box 368 Ridgefield, CT 06877 (800) 243-0127 FAX: (203) 798-6234 (203) 798-9988 http://www.boehringer-ingelheim.com Boehringer Mannheim Biochemicals Division See Roche Diagnostics Boekel Scientific 855 Pennsylvania Boulevard Feasterville, PA 19053 (800) 336-6929 FAX: (215) 396-8264 (215) 396-8200 http://www.boekelsci.com Bohdan Automation 1500 McCormack Boulevard Mundelein, IL 60060 (708) 680-3939 FAX: (708) 680-1199
Biozyme Laboratories 9939 Hibert Street, Suite 101 San Diego, CA 92131 (800) 423-8199 FAX: (858) 549-0138 (858) 549-4484 http://www.biozyme.com
BPAmoco 4500 McGinnis Ferry Road Alpharetta, GA 30005 (800) 328-4537 FAX: (770) 772-8213 (770) 772-8200 http://www.bpamoco.com
Bird Products 1100 Bird Center Drive Palm Springs, CA 92262 (800) 328-4139 FAX: (760) 778-7274 (760) 778-7200 http://www.birdprod.com/bird
Brain Research Laboratories Waban P.O. Box 88 Newton, MA 02468 (888) BRL-5544 FAX: (617) 965-6220 (617) 965-5544 http://www.brainresearchlab.com
Suppliers
6 CPPS Supplement 45
Current Protocols Selected Suppliers of Reagents and Equipment
Braintree Scientific P.O. Box 850929 Braintree, MA 02185 (781) 843-1644 FAX: (781) 982-3160 http://www.braintreesci.com
Bristol-Meyers Squibb P.O. Box 4500 Princeton, NJ 08543 (800) 631-5244 FAX: (800) 523-2965 http://www.bms.com
Burleigh Instruments P.O. Box E Fishers, NY 14453 (716) 924-9355 FAX: (716) 924-9072 http://www.burleigh.com
Brandel 8561 Atlas Drive Gaithersburg, MD 20877 (800) 948-6506 FAX: (301) 869-5570 (301) 948-6506 http://www.brandel.com
Broadley James 19 Thomas Irvine, CA 92618 (800) 288-2833 FAX: (949) 829-5560 (949) 829-5555 http://www.broadleyjames.com
Burns Veterinary Supply 1900 Diplomat Drive Farmer’s Branch, TX 75234 (800) 92-BURNS FAX: (972) 243-6841 http://www.burnsvet.com
Branson Ultrasonics 41 Eagle Road Danbury, CT 06813 (203) 796-0400 FAX: (203) 796-9838 http://www.plasticsnet.com/branson B. Braun Biotech 999 Postal Road Allentown, PA 18103 (800) 258-9000 FAX: (610) 266-9319 (610) 266-6262 http://www.bbraunbiotech.com B. Braun Biotech International Schwarzenberg Weg 73-79 P.O. Box 1120 D-34209 Melsungen, Germany (49) 5661-71-3400 FAX: (49) 5661-71-3702 http://www.bbraunbiotech.com B. Braun-McGaw 2525 McGaw Avenue Irvine, CA 92614 (800) BBRAUN-2 (800) 624-2963 http://www.bbraunusa.com B. Braun Medical Thorncliffe Park Sheffield S35 2PW, UK (44) 114-225-9000 FAX: (44) 114-225-9111 http://www.bbmuk.demon.co.uk Brenntag P.O. Box 13788 Reading, PA 19612-3788 (610) 926-4151 FAX: (610) 926-4160 http://www.brenntagnortheast.com Bresatec See GeneWorks Bright/Hacker Instruments 17 Sherwood Lane Fairfield, NJ 07004 (973) 226-8450 FAX: (973) 808-8281 http://www.hackerinstruments.com Brinkmann Instruments Subsidiary of Sybron 1 Cantiague Road P.O. Box 1019 Westbury, NY 11590 (800) 645-3050 FAX: (516) 334-7521 (516) 334-7500 http://www.brinkmann.com
Brookhaven Instruments 750 Blue Point Road Holtsville, NY 11742 (631) 758-3200 FAX: (631) 758-3255 http://www.bic.com Brownlee Labs See Applied Biosystems Distributed by Pacer Scientific Bruel & Kjaer Division of Spectris Technologies 2815 Colonnades Court Norcross, GA 30071 (800) 332-2040 FAX: (770) 847-8440 (770) 209-6907 http://www.bkhome.com Bruker Analytical X-Ray Systems 5465 East Cheryl Parkway Madison, WI 53711 (800) 234-XRAY FAX: (608) 276-3006 (608) 276-3000 http://www.bruker-axs.com Bruker Instruments 19 Fortune Drive Billerica, MA 01821 (978) 667-9580 FAX: (978) 667-0985 http://www.bruker.com BTX Division of Genetronics 11199 Sorrento Valley Road San Diego, CA 92121 (800) 289-2465 FAX: (858) 597-9594 (858) 597-6006 http://www.genetronics.com/btx Buchler Instruments See Baxter Scientific Products Buckshire 2025 Ridge Road Perkasie, PA 18944 (215) 257-0116 Burdick and Jackson Division of Baxter Scientific Products 1953 S. Harvey Street Muskegon, MI 49442 (800) 368-0050 FAX: (231) 728-8226 (231) 726-3171 http://www.bandj.com/mainframe.htm
Burroughs Wellcome See Glaxo Wellcome The Butler Company 5600 Blazer Parkway Dublin, OH 43017 (800) 551-3861 FAX: (614) 761-9096 (614) 761-9095 http://www.wabutler.com Butterworth Laboratories 54-56 Waldegrave Road Teddington, Middlesex TW11 8LG, UK (44)(0)20-8977-0750 FAX: (44)(0)28-8943-2624 http://www.butterworth-labs.co.uk Buxco Electronics 95 West Wood Road #2 Sharon, CT 06069 (860) 364-5558 FAX: (860) 364-5116 http://www.buxco.com C/D/N Isotopes 88 Leacock Street Pointe-Claire, Quebec H9R 1H1 Canada (800) 697-6254 FAX: (514) 697-6148 C.M.A./Microdialysis AB 73 Princeton Street North Chelmsford, MA 01863 (800) 440-4980 FAX: (978) 251-1950 (978) 251-1940 http://www.microdialysis.com Calbiochem-Novabiochem P.O. Box 12087-2087 La Jolla, CA 92039 (800) 854-3417 FAX: (800) 776-0999 (858) 450-9600 http://www.calbiochem.com California Fine Wire 338 South Fourth Street Grover Beach, CA 93433 (805) 489-5144 FAX: (805) 489-5352 http://www.calfinewire.com Calorimetry Sciences 155 West 2050 North Spanish Fork, UT 84660 (801) 794-2600 FAX: (801) 794-2700 http://www.calscorp.com
Caltag Laboratories 1849 Bayshore Highway, Suite 200 Burlingame, CA 94010 (800) 874-4007 FAX: (650) 652-9030 (650) 652-0468 http://www.caltag.com Cambridge Electronic Design Science Park, Milton Road Cambridge CB4 0FE, UK 44 (0) 1223-420-186 FAX:44 (0) 1223-420-488 http://www.ced.co.uk Cambridge Isotope Laboratories 50 Frontage Road Andover, MA 01810 (800) 322-1174 FAX: (978) 749-2768 (978) 749-8000 http://www.isotope.com Cambridge Research Biochemicals See Zeneca/CRB Cambridge Technology 109 Smith Place Cambridge, MA 02138 (6l7) 441-0600 FAX: (617) 497-8800 http://www.camtech.com Camlab Nuffield Road Cambridge CB4 1TH, UK (44) 122-3424222 FAX: (44) 122-3420856 http://www.camlab.co.uk/home.htm Campden Instruments Park Road Sileby Loughborough Leicestershire LE12 7TU, UK (44) 1509-814790 FAX: (44) 1509-816097 http://www.campdeninst.com/home.htm Cappel Laboratories See Organon Teknika Cappel Carl Roth GmgH & Company Schoemperlenstrasse 1-5 76185 Karlsrube Germany (49) 72-156-06164 FAX: (49) 72-156-06264 http://www.carl-roth.de Carl Zeiss One Zeiss Drive Thornwood, NY 10594 (800) 233-2343 FAX: (914) 681-7446 (914) 747-1800 http://www.zeiss.com Carlo Erba Reagenti Via Winckelmann 1 20148 Milano Lombardia, Italy (39) 0-29-5231 FAX: (39) 0-29-5235-904 http://www.carloerbareagenti.com
Suppliers
7 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
Carolina Biological Supply 2700 York Road Burlington, NC 27215 (800) 334-5551 FAX: (336) 584-76869 (336) 584-0381 http://www.carolina.com
Cell Genesys 342 Lakeside Drive Foster City, CA 94404 (650) 425-4400 FAX: (650) 425-4457 http://www.cellgenesys.com
Carolina Fluid Components 9309 Stockport Place Charlotte, NC 28273 (704) 588-6101 FAX: (704) 588-6115 http://www.cfcsite.com
Cell Signaling Technology 166B Cummings Center Boverly, MA 01915 (877) 616-CELL FAX: (978) 867-2388 (978) 867-2488 http://www.cellsignal.com
Cartesian Technologies 17851 Skypark Circle, Suite C Irvine, CA 92614 (800) 935-8007 http://cartesiantech.com
Cell Systems 12815 NE 124th Street, Suite A Kirkland, WA 98034 (800) 697-1211 FAX: (425) 820-6762 (425) 823-1010
Cayman Chemical 1180 East Ellsworth Road Ann Arbor, MI 48108 (800) 364-9897 FAX: (734) 971-3640 (734) 971-3335 http://www.caymanchem.com
Cellmark Diagnostics 20271 Goldenrod Lane Germantown, MD 20876 (800) 872-5227 FAX: (301) 428-4877 (301) 428-4980 http://www.cellmark-labs.com
CB Sciences One Washington Street, Suite 404 Dover, NH 03820 (800) 234-1757 FAX: (603) 742-2455 http://www.cbsci.com
Cellomics 635 William Pitt Way Pittsburgh, PA 15238 (888) 826-3857 FAX: (412) 826-3850 (412) 826-3600 http://www.cellomics.com
CBS Scientific P.O. Box 856 Del Mar, CA 92014 (800) 243-4959 FAX: (858) 755-0733 (858) 755-4959 http://www.cbssci.com CCR (Coriell Cell Repository) See Coriell Institute for Medical Research CE Instruments Grand Avenue Parkway Austin, TX 78728 (800) 876-6711 FAX: (512) 251-1597 http://www.ceinstruments.com Cedarlane Laboratories 5516 8th Line, R.R. #2 Hornby, Ontario L0P 1E0 Canada (905) 878-8891 FAX: (905) 878-7800 http://www.cedarlanelabs.com CEL Associates P.O. Box 721854 Houston, TX 77272 (800) 537-9339 FAX: (281) 933-0922 (281) 933-9339 http://www.cel-1.com Cel-Line Associates See Erie Scientific Celite World Minerals 130 Castilian Drive Santa Barbara, CA 93117 (805) 562-0200 FAX: (805) 562-0299 http://www.worldminerals.com/celite
Celltech 216 Bath Road Slough, Berkshire SL1 4EN, UK (44) 1753 534655 FAX: (44) 1753 536632 http://www.celltech.co.uk Cellular Products 872 Main Street Buffalo, NY 14202 (800) CPI-KITS FAX: (716) 882-0959 (716) 882-0920 http://www.zeptometrix.com CEM P.O. Box 200 Matthews, NC 28106 (800) 726-3331 Centers for Disease Control 1600 Clifton Road NE Atlanta, GA 30333 (800) 311-3435 FAX: (888) 232-3228 (404) 639-3311 http://www.cdc.gov CERJ Centre d’Elevage Roger Janvier 53940 Le Genest Saint Isle France Cetus See Chiron Chance Propper Warly, West Midlands B66 1NZ, UK (44)(0)121-553-5551 FAX: (44)(0)121-525-0139
Charles River Laboratories 251 Ballardvale Street Wilmington, MA 01887 (800) 522-7287 FAX: (978) 658-7132 (978) 658-6000 http://www.criver.com Charm Sciences 36 Franklin Street Malden, MA 02148 (800) 343-2170 FAX: (781) 322-3141 (781) 322-1523 http://www.charm.com Chase-Walton Elastomers 29 Apsley Street Hudson, MA 01749 (800) 448-6289 FAX: (978) 562-5178 (978) 568-0202 http://www.chase-walton.com ChemGenes Ashland Technology Center 200 Homer Avenue Ashland, MA 01721 (800) 762-9323 FAX: (508) 881-3443 (508) 881-5200 http://www.chemgenes.com Chemglass 3861 North Mill Road Vineland, NJ 08360 (800) 843-1794 FAX: (856) 696-9102 (800) 696-0014 http://www.chemglass.com Chemicon International 28835 Single Oak Drive Temecula, CA 92590 (800) 437-7500 FAX: (909) 676-9209 (909) 676-8080 http://www.chemicon.com Chem-Impex International 935 Dillon Drive Wood Dale, IL 60191 (800) 869-9290 FAX: (630) 766-2218 (630) 766-2112 http://www.chemimpex.com Chem Service P.O. Box 599 West Chester, PA 19381-0599 (610) 692-3026 FAX: (610) 692-8729 http://www.chemservice.com Chemsyn Laboratories 13605 West 96th Terrace Lenexa, Kansas 66215 (913) 541-0525 FAX: (913) 888-3582 http://www.tech.epcorp.com/ChemSyn/ chemsyn.htm Chemunex USA 1 Deer Park Drive, Suite H-2 Monmouth Junction, NJ 08852 (800) 411-6734 http://www.chemunex.com
Cherwell Scientific Publishing The Magdalen Centre Oxford Science Park Oxford OX44GA, UK (44)(1) 865-784-800 FAX: (44)(1) 865-784-801 http://www.cherwell.com ChiRex Cauldron 383 Phoenixville Pike Malvern, PA 19355 (610) 727-2215 FAX: (610) 727-5762 http://www.chirex.com Chiron Diagnostics See Bayer Diagnostics Chiron Mimotopes Peptide Systems See Multiple Peptide Systems Chiron 4560 Horton Street Emeryville, CA 94608 (800) 244-7668 FAX: (510) 655-9910 (510) 655-8730 http://www.chiron.com Chrom Tech P.O. Box 24248 Apple Valley, MN 55124 (800) 822-5242 FAX: (952) 431-6345 http://www.chromtech.com Chroma Technology 72 Cotton Mill Hill, Unit A-9 Brattleboro, VT 05301 (800) 824-7662 FAX: (802) 257-9400 (802) 257-1800 http://www.chroma.com Chromatographie ZAC de Moulin No. 2 91160 Saulx les Chartreux France (33) 01-64-54-8969 FAX: (33) 01-69-0968091 http://www.chromatographie.com Chromogenix Taljegardsgatan 3 431-53 Mlndal, Sweden (46) 31-706-20-70 FAX: (46) 31-706-20-80 http://www.chromogenix.com Chrompack USA c/o Varian USA 2700 Mitchell Drive Walnut Creek, CA 94598 (800) 526-3687 FAX: (925) 945-2102 (925) 939-2400 http://www.chrompack.com Chugai Biopharmaceuticals 6275 Nancy Ridge Drive San Diego, CA 92121 (858) 535-5900 FAX: (858) 546-5973 http://www.chugaibio.com Ciba-Corning Diagnostics See Bayer Diagnostics
Suppliers
8 CPPS Supplement 45
Current Protocols Selected Suppliers of Reagents and Equipment
Ciba-Geigy See Ciba Specialty Chemicals or Novartis Biotechnology Ciba Specialty Chemicals 540 White Plains Road Tarrytown, NY 10591 (800) 431-1900 FAX: (914) 785-2183 (914) 785-2000 http://www.cibasc.com CIBA Vision Division of Novartis AG 11460 Johns Creek Parkway Duluth, GA 30097 (770) 476-3937 http://www.cvworld.com Cidex Advanced Sterilization Products 33 Technology Drive Irvine, CA 92618 (800) 595-0200 (949) 581-5799 http://www.cidex.com/ASPnew.htm Cinna Scientific Subsidiary of Molecular Research Center 5645 Montgomery Road Cincinnati, OH 45212 (800) 462-9868 FAX: (513) 841-0080 (513) 841-0900 http://www.mrcgene.com Ciphergen Biosystems, Inc. 6611 Dumbarton Circle Fremont, CA 94555 (888) 864-3770 FAX: (510) 505-2101 (510) 505 2100 http://www.ciphergen.com Cistron Biotechnology 10 Bloomfield Avenue Pine Brook, NJ 07058 (800) 642-0167 FAX: (973) 575-4854 (973) 575-1700 http://www.cistronbio.com Clark Electromedical Instruments See Harvard Apparatus Clay Adam See Becton Dickinson Primary Care Diagnostics CLB (Central Laboratory of the Netherlands) Blood Transfusion Service P.O. Box 9190 1006 AD Amsterdam, The Netherlands (31) 20-512-9222 FAX: (31) 20-512-3332 Cleveland Scientific P.O. Box 300 Bath, OH 44210 (800) 952-7315 FAX: (330) 666-2240 http:://www.clevelandscientific.com
Clonetics Division of BioWhittaker http://www.clonetics.com Also see BioWhittaker Clontech Laboratories 1020 East Meadow Circle Palo Alto, CA 94303 (800) 662-2566 FAX: (800) 424-1350 (650) 424-8222 FAX: (650) 424-1088 http://www.clontech.com Closure Medical Corporation 5250 Greens Dairy Road Raleigh, NC 27616 (919) 876-7800 FAX: (919) 790-1041 http://www.closuremed.com CMA Microdialysis AB 73 Princeton Street North Chelmsford, MA 01863 (800) 440-4980 FAX: (978) 251-1950 (978) 251 1940 http://www.microdialysis.com Cocalico Biologicals 449 Stevens Road P.O. Box 265 Reamstown, PA 17567 (717) 336-1990 FAX: (717) 336-1993 Coherent Laser 5100 Patrick Henry Drive Santa Clara, CA 95056 (800) 227-1955 FAX: (408) 764-4800 (408) 764-4000 http://www.cohr.com Cohu P.O. Box 85623 San Diego, CA 92186 (858) 277-6700 FAX: (858) 277-0221 http://www.COHU.com/cctv Cole-Parmer Instrument 625 East Bunker Court Vernon Hills, IL 60061 (800) 323-4340 FAX: (847) 247-2929 (847) 549-7600 http://www.coleparmer.com Collaborative Biomedical Products and Collaborative Research See Becton Dickinson Labware Collagen Aesthetics 1850 Embarcadero Road Palo Alto, CA 94303 (650) 856-0200 FAX: (650) 856-0533 http://www.collagen.com
Colonial Medical Supply 504 Wells Road Franconia, NH 03580 (603) 823-9911 FAX: (603) 823-8799 http://www.colmedsupply.com Colorado Serum 4950 York Street Denver, CO 80216 (800) 525-2065 FAX: (303) 295-1923 http://www.colorado-serum.com Columbia Diagnostics 8001 Research Way Springfield, VA 22153 (800) 336-3081 FAX: (703) 569-2353 (703) 569-7511 http://www.columbiadiagnostics.com Columbus Instruments 950 North Hague Avenue Columbus, OH 43204 (800) 669-5011 FAX: (614) 276-0529 (614) 276-0861 http://www.columbusinstruments.com Compu Cyte Corp. 12 Emily Street Cambridge, MA 02139 (800) 840-1303 FAX: (617) 577-4501 (617) 492-1300 http://www.compucyte.com Compugen 25 Leek Crescent Richmond Hill, Ontario L4B 4B3 Canada 800-387-5045 FAX: (905) 707-2020 (905) 707-2000 http://www.compugen.com/locations. htm Computer Associates International One Computer Associates Plaza Islandia, NY 11749 (631) 342-6000 FAX: (631) 342-6800 http://www.cai.com Connaught Laboratories See Aventis Pasteur Connectix 2955 Campus Drive, Suite 100 San Mateo, CA 94403 (800) 950-5880 FAX: (650) 571-0850 (650) 571-5100 http://www.connectix.com
Collagen Corporation See Collagen Aesthetics
Contech 99 Hartford Avenue Providence, RI 02909 (401) 351-4890 FAX: (401) 421-5072 http://www.iol.ie/∼burke/contech.html
College of American Pathologists 325 Waukegan Road Northfield, IL 60093 (800) 323-4040 FAX: (847) 832-8000 (847) 446-8800 http://www.cap.org/index.cfm
Continental Laboratory Products 5648 Copley Drive San Diego, CA 92111 (800) 456-7741 FAX: (858) 279-5465 (858) 279-5000 http://www.conlab.com
ConvaTec Professional Services P.O. Box 5254 Princeton, NJ 08543 (800) 422-8811 http://www.convatec.com Cooper Instruments & Systems P.O. Box 3048 Warrenton, VA 20188 (800) 344-3921 FAX: (540) 347-4755 (540) 349-4746 http://www.cooperinstruments.com Cooperative Human Tissue Network (866) 462-2486 http://www.chin.ims.nci.nih.gov Cora Styles Needles ’N Blocks 56 Milton Street Arlington, MA 02474 (781) 648-6289 FAX: (781) 641-7917 Coriell Cell Repository (CCR) See Coriell Institute for Medical Research Coriell Institute for Medical Research Human Genetic Mutant Repository 401 Haddon Avenue Camden, NJ 08103 (856) 966-7377 FAX: (856) 964-0254 http://arginine.umdnj.edu Corion 8 East Forge Parkway Franklin, MA 02038 (508) 528-4411 FAX: (508) 520-7583 (800) 598-6783 http://www.corion.com Corning and Corning Science Products P.O. Box 5000 Corning, NY 14831 (800) 222-7740 FAX: (607) 974-0345 (607) 974-9000 http://www.corning.com Costar See Corning Coulbourn Instruments 7462 Penn Drive Allentown, PA 18106 (800) 424-3771 FAX: (610) 391-1333 (610) 395-3771 http://www.coulbourninst.com Coulter Cytometry See Beckman Coulter Covance Research Products 465 Swampbridge Road Denver, PA 17517 (800) 345-4114 FAX: (717) 336-5344 (717) 336-4921 http://www.covance.com
Suppliers
9 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
Coy Laboratory Products 14500 Coy Drive Grass Lake, MI 49240 (734) 475-2200 FAX: (734) 475-1846 http://www.coylab.com
CWE 124 Sibley Avenue Ardmore, PA 19003 (610) 642-7719 FAX: (610) 642-1532 http://www.cwe-inc.com
CPG 3 Borinski Road Lincoln Park, NJ 07035 (800) 362-2740 FAX: (973) 305-0884 (973) 305-8181 http://www.cpg-biotech.com
Cybex Computer Products 4991 Corporate Drive Huntsville, AL 35805 (800) 932-9239 FAX: (800) 462-9239 http://www.cybex.com
CPL Scientific 43 Kingfisher Court Hambridge Road Newbury RG14 5SJ, UK (44) 1635-574902 FAX: (44) 1635-529322 http://www.cplscientific.co.uk CraMar Technologies 8670 Wolff Court, #160 Westminster, CO 80030 (800) 4-TOMTEC http://www.cramar.com Crescent Chemical 1324 Motor Parkway Hauppauge, NY 11788 (800) 877-3225 FAX: (631) 348-0913 (631) 348-0333 http://www.creschem.com Crist Instrument P.O. Box 128 10200 Moxley Road Damascus, MD 20672 (301) 253-2184 FAX: (301) 253-0069 http://www.cristinstrument.com Cruachem See Annovis http://www.cruachem.com CS Bio 1300 Industrial Road San Carlos, CA 94070 (800) 627-2461 FAX: (415) 802-0944 (415) 802-0880 http://www.csbio.com CS-Chromatographie Service Am Parir 27 D-52379 Langerwehe, Germany (49) 2423-40493-0 FAX: (49) 2423-40493-49 http://www.cs-chromatographie.de Cuno 400 Research Parkway Meriden, CT 06450 (800) 231-2259 FAX: (203) 238-8716 (203) 237-5541 http://www.cuno.com Curtin Matheson Scientific 9999 Veterans Memorial Drive Houston, TX 77038 (800) 392-3353 FAX: (713) 878-3598 (713) 878-3500
Cygnus Technology P.O. Box 219 Delaware Water Gap, PA 18327 (570) 424-5701 FAX: (570) 424-5630 http://www.cygnustech.com Cymbus Biotechnology Eagle Class, Chandler’s Ford Hampshire SO53 4NF, UK (44) 1-703-267-676 FAX: (44) 1-703-267-677 http://[email protected] Cytogen 600 College Road East Princeton, NJ 08540 (609) 987-8200 FAX: (609) 987-6450 http://www.cytogen.com Cytogen Research and Development 89 Bellevue Hill Road Boston, MA 02132 (617) 325-7774 FAX: (617) 327-2405 CytRx 154 Technology Parkway Norcross, GA 30092 (800) 345-2987 FAX: (770) 368-0622 (770) 368-9500 http://www.cytrx.com Dade Behring Corporate Headquarters 1717 Deerfield Road Deerfield, IL 60015 (847) 267-5300 FAX: (847) 267-1066 http://www.dadebehring.com
Dalton Chemical Laboratoris 349 wildcat Road Toronto, Ontario M3J 253 Canada (416) 661-2102 FAX: (416) 661-2108 (800) 567-5060 (in Canada only) http://www.dalton.com Damon, IEC See Thermoquest Dan Kar Scientific 150 West Street Wilmington, MA 01887 (800) 942-5542 FAX: (978) 658-0380 (978) 988-9696 http://www.dan-kar.com DataCell Falcon Business Park 40 Ivanhoe Road Finchampstead, Berkshire RG40 4QQ, UK (44) 1189 324324 FAX: (44) 1189 324325 http://www.datacell.co.uk In the US: (408) 446-3575 FAX: (408) 446-3589 http://www.datacell.com DataWave Technologies 380 Main Street, Suite 209 Longmont, CO 80501 (800) 736-9283 FAX: (303) 776-8531 (303) 776-8214 Datex-Ohmeda 3030 Ohmeda Drive Madison, WI 53718 (800) 345-2700 FAX: (608) 222-9147 (608) 221-1551 http://www.us.datex-ohmeda.com DATU 82 State Street Geneva, NY 14456 (315) 787-2240 FAX: (315) 787-2397 http://www.nysaes.cornell.edu/datu
Dagan 2855 Park Avenue Minneapolis, MN 55407 (612) 827-5959 FAX: (612) 827-6535 http://www.dagan.com
David Kopf Instruments 7324 Elmo Street P.O. Box 636 Tujunga, CA 91043 (818) 352-3274 FAX: (818) 352-3139
Dako 6392 Via Real Carpinteria, CA 93013 (800) 235-5763 FAX: (805) 566-6688 (805) 566-6655 http://www.dakousa.com
Decagon Devices P.O. Box 835 950 NE Nelson Court Pullman, WA 99163 (800) 755-2751 FAX: (509) 332-5158 (509) 332-2756 http://www.decagon.com
Dako A/S 42 Produktionsvej P.O. Box 1359 DK-2600 Glostrup, Denmark (45) 4492-0044 FAX: (45) 4284-1822 Dakopatts See Dako A/S
Decon Labs 890 Country Line Road Bryn Mawr, PA 19010 (800) 332-6647 FAX: (610) 964-0650 (610) 520-0610 http://www.deconlabs.com
Decon Laboratories Conway Street Hove, Sussex BN3 3LY, UK (44) 1273 739241 FAX: (44) 1273 722088 Degussa Precious Metals Division 3900 South Clinton Avenue South Plainfield, NJ 07080 (800) DEGUSSA FAX: (908) 756-7176 (908) 561-1100 http://www.degussa-huls.com Deneba Software 1150 NW 72nd Avenue Miami, FL 33126 (305) 596-5644 FAX: (305) 273-9069 http://www.deneba.com Denley Instruments Ltd Unit 2A, Altbarn Industrial Estate Hawkins Rd Colchester Essex CO2 8LG, UK (44) 1206-798394 FAX: (44) 1206-798466 Deseret Medical 524 West 3615 South Salt Lake City, UT 84115 (801) 270-8440 FAX: (801) 293-9000 Devcon Plexus 30 Endicott Street Danvers, MA 01923 (800) 626-7226 FAX: (978) 774-0516 (978) 777-1100 http://www.devcon.com Developmental Studies Hybridoma Bank University of Iowa 436 Biology Building Iowa City, IA 52242 (319) 335-3826 FAX: (319) 335-2077 http://www.uiowa.edu/∼dshbwww DeVilbiss Division of Sunrise Medical Respiratory 100 DeVilbiss Drive P.O. Box 635 Somerset, PA 15501 (800) 338-1988 FAX: (814) 443-7572 (814) 443-4881 http://www.sunrisemedical.com Dharmacon Research 1376 Miners Drive #101 Lafayette, CO 80026 (303) 604-9499 FAX: (303) 604-9680 http://www.dharmacon.com DiaCheM Triangle Biomedical Gardiners Place West Gillibrands, Lancashire WN8 9SP, UK (44) 1695-555581 FAX: (44) 1695-555518 http://www.diachem.co.uk
Suppliers
10 CPPS Supplement 45
Current Protocols Selected Suppliers of Reagents and Equipment
Diagen Max-Volmer Strasse 4 D-40724 Hilden, Germany (49) 2103-892-230 FAX: (49) 2103-892-222 Diagnostic Concepts 6104 Madison Court Morton Grove, IL 60053 (847) 604-0957 Diagnostic Developments See DiaCheM Diagnostic Instruments 6540 Burroughs Sterling Heights, MI 48314 (810) 731-6000 FAX: (810) 731-6469 http://www.diaginc.com Diamedix 2140 North Miami Avenue Miami, FL 33127 (800) 327-4565 FAX: (305) 324-2395 (305) 324-2300 DiaSorin 1990 Industrial Boulevard Stillwater, MN 55082 (800) 328-1482 FAX: (651) 779-7847 (651) 439-9719 http://www.diasorin.com Diatome US 321 Morris Road Fort Washington, PA 19034 (800) 523-5874 FAX: (215) 646-8931 (215) 646-1478 http://www.emsdiasum.com Difco Laboratories See Becton Dickinson Digene 1201 Clopper Road Gaithersburg, MD 20878 (301) 944-7000 (800) 344-3631 FAX: (301) 944-7121 http://www.digene.com Digi-Key 701 Brooks Avenue South Thief River Falls, MN 56701 (800) 344-4539 FAX: (218) 681-3380 (218) 681-6674 http://www.digi-key.com Digitimer 37 Hydeway Welwyn Garden City, Hertfordshire AL7 3BE, UK (44) 1707-328347 FAX: (44) 1707-373153 http://www.digitimer.com Dimco-Gray 8200 South Suburban Road Dayton, OH 45458 (800) 876-8353 FAX: (937) 433-0520 (937) 433-7600 http://www.dimco-gray.com
Dionex 1228 Titan Way P.O. Box 3603 Sunnyvale, CA 94088 (408) 737-0700 FAX: (408) 730-9403 http://dionex2.promptu.com Display Systems Biotech 1260 Liberty Way, Suite B Vista, CA 92083 (800) 697-1111 FAX: (760) 599-9930 (760) 599-0598 http://www.displaysystems.com Diversified Biotech 1208 VFW Parkway Boston, MA 02132 (617) 965-8557 FAX: (617) 323-5641 (800) 796-9199 http://www.divbio.com DNA ProScan P.O. Box 121585 Nashville, TN 37212 (800) 841-4362 FAX: (615) 292-1436 (615) 298-3524 http://www.dnapro.com DNAStar 1228 South Park Street Madison, WI 53715 (608) 258-7420 FAX: (608) 258-7439 http://www.dnastar.com DNAVIEW Attn: Charles Brenner http://www.wco.com ∼cbrenner/dnaview.htm Doall NYC 36-06 48th Avenue Long Island City, NY 11101 (718) 392-4595 FAX: (718) 392-6115 http://www.doall.com Dojindo Molecular Technologies 211 Perry Street Parkway, Suite 5 Gaitherbusburg, MD 20877 (877) 987-2667 http://www.dojindo.com Dolla Eastern See Doall NYC Dolan Jenner Industries 678 Andover Street Lawrence, MA 08143 (978) 681-8000 (978) 682-2500 http://www.dolan-jenner.com Dow Chemical Customer Service Center 2040 Willard H. Dow Center Midland, MI 48674 (800) 232-2436 FAX: (517) 832-1190 (409) 238-9321 http://www.dow.com
Dow Corning Northern Europe Meriden Business Park Copse Drive Allesley, Coventry CV5 9RG, UK (44) 1676 528 000 FAX: (44) 1676 528 001 Dow Corning P.O. Box 994 Midland, MI 48686 (517) 496-4000 http://www.dowcorning.com Dow Corning (Lubricants) 2200 West Salzburg Road Auburn, MI 48611 (800) 248-2481 FAX: (517) 496-6974 (517) 496-6000 Dremel 4915 21st Street Racine, WI 53406 (414) 554-1390 http://www.dremel.com Drummond Scientific 500 Parkway P.O. Box 700 Broomall, PA 19008 (800) 523-7480 FAX: (610) 353-6204 (610) 353-0200 http://www.drummondsci.com Duchefa Biochemie BV P.O. Box 2281 2002 CG Haarlem, The Netherlands 31-0-23-5319093 FAX:31-0-23-5318027 http://www.duchefa.com Duke Scientific 2463 Faber Place Palo Alto, CA 94303 (800) 334-3883 FAX: (650) 424-1158 (650) 424-1177 http://www.dukescientific.com Duke University Marine Laboratory 135 Duke Marine Lab Road Beaufort, NC 28516-9721 (252) 504-7503 FAX: (252) 504-7648 http://www.env.duke.edu/rnarinelab DuPont Biotechnology Systems See NEN Life Science Products DuPont Medical Products See NEN Life Science Products DuPont Merck Pharmaceuticals 331 Treble Cove Road Billerica, MA 01862 (800) 225-1572 FAX: (508) 436-7501 http://www.dupontmerck.com DuPont NEN Products See NEN Life Science Products
Dynal 5 Delaware Drive Lake Success, NY 11042 (800) 638-9416 FAX: (516) 326-3298 (516) 326-3270 http://www.dynal.net Dynal AS Ullernchausen 52, 0379 Oslo, Norway 47-22-06-10-00 FAX:47-22-50-70-15 http://www.dynal.no Dynalab P.O. Box 112 Rochester, NY 14692 (800) 828-6595 FAX: (716) 334-9496 (716) 334-2060 http://www.dynalab.com Dynarex 1 International Boulevard Brewster, NY 10509 (888) DYNAREX FAX: (914) 279-9601 (914) 279-9600 http://www.dynarex.com Dynatech See Dynex Technologies Dynex Technologies 14340 Sullyfield Circle Chantilly, VA 22021 (800) 336-4543 FAX: (703) 631-7816 (703) 631-7800 http://www.dynextechnologies.com Dyno Mill See Willy A. Bachofen E.S.A. 22 Alpha Road Chelmsford, MA 01824 (508) 250-7000 FAX: (508) 250-7090 E.W. Wright 760 Durham Road Guilford, CT 06437 (203) 453-6410 FAX: (203) 458-6901 http://www.ewwright.com E-Y Laboratories 107 N. Amphlett Boulevard San Mateo, CA 94401 (800) 821-0044 FAX: (650) 342-2648 (650) 342-3296 http://www.eylabs.com Eastman Kodak 1001 Lee Road Rochester, NY 14650 (800) 225-5352 FAX: (800) 879-4979 (716) 722-5780 FAX: (716) 477-8040 http://www.kodak.com ECACC See European Collection of Animal Cell Cultures EC Apparatus See Savant/EC Apparatus
Suppliers
11 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
Ecogen, SRL Gensura Laboratories Ptge. Dos de Maig 9(08041) Barcelona, Spain (34) 3-450-2601 FAX: (34) 3-456-0607 http://www.ecogen.com Ecolab 370 North Wabasha Street St. Paul, MN 55102 (800) 35-CLEAN FAX: (651) 225-3098 (651) 352-5326 http://www.ecolab.com ECO PHYSICS 3915 Research Park Drive, Suite A-3 Ann Arbor, MI 48108 (734) 998-1600 FAX: (734) 998-1180 http://www.ecophysics.com Edge Biosystems 19208 Orbit Drive Gaithersburg, MD 20879-4149 (800) 326-2685 FAX: (301) 990-0881 (301) 990-2685 http://www.edgebio.com Edmund Scientific 101 E. Gloucester Pike Barrington, NJ 08007 (800) 728-6999 FAX: (856) 573-6263 (856) 573-6250 http://www.edsci.com EG&G See Perkin-Elmer Ekagen 969 C Industry Road San Carlos, CA 94070 (650) 592-4500 FAX: (650) 592-4500 Elcatech P.O. Box 10935 Winston-Salem, NC 27108 (336) 544-8613 FAX: (336) 777-3623 (910) 777-3624 http://www.elcatech.com Electron Microscopy Sciences 321 Morris Road Fort Washington, PA 19034 (800) 523-5874 FAX: (215) 646-8931 (215) 646-1566 http://www.emsdiasum.com Electron Tubes 100 Forge Way, Unit F Rockaway, NJ 07866 (800) 521-8382 FAX: (973) 586-9771 (973) 586-9594 http://www.electrontubes.com Elicay Laboratory Products, (UK) Ltd. 4 Manborough Mews Crockford Lane Basingstoke, Hampshire RG 248NA, England (256) 811-118 FAX: (256) 811-116 http://www.elkay-uk.co.uk
Eli Lilly Lilly Corporate Center Indianapolis, IN 46285 (800) 545-5979 FAX: (317) 276-2095 (317) 276-2000 http://www.lilly.com
Erie Scientific 20 Post Road Portsmouth, NH 03801 (888) ERIE-SCI FAX: (603) 431-8996 (603) 431-8410 http://www.eriesci.com
ELISA Technologies See Neogen
ES Industries 701 South Route 73 West Berlin, NJ 08091 (800) 356-6140 FAX: (856) 753-8484 (856) 753-8400 http://www.esind.com
Elkins-Sinn See Wyeth-Ayerst EMBI See European Bioinformatics Institute EM Science 480 Democrat Road Gibbstown, NJ 08027 (800) 222-0342 FAX: (856) 423-4389 (856) 423-6300 http://www.emscience.com EM Separations Technology See R & S Technology Endogen 30 Commerce Way Woburn, MA 01801 (800) 487-4885 FAX: (617) 439-0355 (781) 937-0890 http://www.endogen.com ENGEL-Loter HSGM Heatcutting Equipment & Machines 1865 E. Main Street, No. 5 Duncan, SC 29334 (888) 854-HSGM FAX: (864) 486-8383 (864) 486-8300 http://www.engelgmbh.com Enzo Diagnostics 60 Executive Boulevard Farmingdale, NY 11735 (800) 221-7705 FAX: (516) 694-7501 (516) 694-7070 http://www.enzo.com Enzogenetics 4197 NW Douglas Avenue Corvallis, OR 97330 (541) 757-0288 The Enzyme Center See Charm Sciences
ESA 22 Alpha Road Chelmsford, MA 01824 (800) 959-5095 FAX: (978) 250-7090 (978) 250-7000 http://www.esainc.com Ethicon Route 22, P.O. Box 151 Somerville, NJ 08876 (908) 218-0707 http://www.ethiconinc.com Ethicon Endo-Surgery 4545 Creek Road Cincinnati, OH 45242 (800) 766-9534 FAX: (513) 786-7080
Exciton P.O. Box 31126 Dayton, OH 45437 (937) 252-2989 FAX: (937) 258-3937 http://www.exciton.com Extrasynthese ZI Lyon Nord SA-BP62 69730 Genay, France (33) 78-98-20-34 FAX: (33) 78-98-19-45 Factor II 1972 Forest Avenue P.O. Box 1339 Lakeside, AZ 85929 (800) 332-8688 FAX: (520) 537-8066 (520) 537-8387 http://www.factor2.com Falcon See Becton Dickinson Labware Febit AG Kafertaler Strasse 190 D-68167 Mannheim Germany (49) 621-3804-0 FAX: (49) 621-3804-400 http://www.febit.com Fenwal See Baxter Healthcare
Eurogentec Parc Scientifique du Sart Tilman 4102 Seraing, Belgium 32-4-240-76-76 FAX:32-4-264-07-88 http://www.eurogentec.com
Filemaker 5201 Patrick Henry Drive Santa Clara, CA 95054 (408) 987-7000 (800) 325-2747
European Bioinformatics Institute Wellcome Trust Genomes Campus Hinxton, Cambridge CB10 1SD, UK (44) 1223-49444 FAX: (44) 1223-494468
Fine Science Tools 202-277 Mountain Highway North Vancouver, British Columbia V7J 3P2 Canada (800) 665-5355 FAX: (800) 665 4544 (604) 980-2481 FAX: (604) 987-3299
European Collection of Animal Cell Cultures (ECACC) Centre for Applied Microbiology & Research Salisbury, Wiltshire SP4 0JG, UK (44) 1980-612 512 FAX: (44) 1980-611 315 http://www.camr.org.uk
Enzyme Systems Products 486 Lindbergh Avenue Livermore, CA 94550 (888) 449-2664 FAX: (925) 449-1866 (925) 449-2664 http://www.enzymesys.com
Evergreen Scientific 2254 E. 49th Street P.O. Box 58248 Los Angeles, CA 90058 (800) 421-6261 FAX: (323) 581-2503 (323) 583-1331 http://www.evergreensci.com
Epicentre Technologies 1402 Emil Street Madison, WI 53713 (800) 284-8474 FAX: (608) 258-3088 (608) 258-3080 http://www.epicentre.com
Exalpha Biologicals 20 Hampden Street Boston, MA 02205 (800) 395-1137 FAX: (617) 969-3872 (617) 558-3625 http://www.exalpha.com
Fine Science Tools 373-G Vintage Park Drive Foster City, CA 94404 (800) 521-2109 FAX: (800) 523-2109 (650) 349-1636 FAX: (630) 349-3729 Fine Science Tools Fahrtgasse 7-13 D-69117 Heidelberg, Germany (49) 6221 905050 FAX: (49) 6221 600001 http://www.finescience.com Finn Aqua AMSCO Finn Aqua Oy Teollisuustiez, FIN-04300 Tuusula, Finland 358 025851 FAX:358 0276019 Finnigan 355 River Oaks Parkway San Jose, CA 95134 (408) 433-4800 FAX: (408) 433-4821 http://www.finnigan.com
Suppliers
12 CPPS Supplement 45
Current Protocols Selected Suppliers of Reagents and Equipment
Dr. L. Fischer Lutherstrasse 25A D-69120 Heidelberg Germany (49) 6221-16-0368 http://home.eplus-online.de/ electroporation Fisher Chemical Company Fisher Scientific Limited 112 Colonnade Road Nepean Ontario K2E 7L6, Canada (800) 234-7437 FAX: (800) 463-2996 http://www.fisherscientific.com Fisher Scientific 2000 Park Lane Pittsburgh, PA 15275 (800) 766-7000 FAX: (800) 926-1166 (412) 562-8300 http://www3.fishersci.com W.F. Fisher & Son 220 Evans Way, Suite #1 Somerville, NJ 08876 (908) 707-4050 FAX: (908) 707-4099 Fitzco 5600 Pioneer Creek Drive Maple Plain, MN 55359 (800) 367-8760 FAX: (612) 479-2880 (612) 479-3489 http://www.fitzco.com 5 Prime → 3 Prime See 2000 Eppendorf-5 Prime http://www.5prime.com Flambeau 15981 Valplast Road Middlefield, Ohio 44062 (800) 232-3474 Fax: (440) 632-1581 (400) 632-1631 http://www.flambeau.com Fleisch (Rusch) 2450 Meadowbrook Parkway Duluth, GA 30096 (770) 623-0816 FAX: (770) 623-1829 http://ruschinc.com Flow Cytometry Standards P.O. Box 194344 San Juan, PR 00919 (800) 227-8143 FAX: (787) 758-3267 (787) 753-9341 http://www.fcstd.com Flow Labs See ICN Biomedicals Flow-Tech Supply P.O. Box 1388 Orange, TX 77631 (409) 882-0306 FAX: (409) 882-0254 http://www.flow-tech.com Fluid Marketing See Fluid Metering
Fluid Metering 5 Aerial Way, Suite 500 Sayosett, NY 11791 (516) 922-6050 FAX: (516) 624-8261 http://www.fmipump.com Fluorochrome 1801 Williams, Suite 300 Denver, CO 80264 (303) 394-1000 FAX: (303) 321-1119 Fluka Chemical See Sigma-Aldrich FMC BioPolymer 1735 Market Street Philadelphia, PA 19103 (215) 299-6000 FAX: (215) 299-5809 http://www.fmc.com FMC BioProducts 191 Thomaston Street Rockland, ME 04841 (800) 521-0390 FAX: (800) 362-1133 (207) 594-3400 FAX: (207) 594-3426 http://www.bioproducts.com Forma Scientific Milcreek Road P.O. Box 649 Marietta, OH 45750 (800) 848-3080 FAX: (740) 372-6770 (740) 373-4765 http://www.forma.com Fort Dodge Animal Health 800 5th Street NW Fort Dodge, IA 50501 (800) 685-5656 FAX: (515) 955-9193 (515) 955-4600 http://www.ahp.com Fotodyne 950 Walnut Ridge Drive Hartland, WI 53029 (800) 362-3686 FAX: (800) 362-3642 (262) 369-7000 FAX: (262) 369-7013 http://www.fotodyne.com Fresenius HemoCare 6675 185th Avenue NE, Suite 100 Redwood, WA 98052 (800) 909-3872 (425) 497-1197 http://www.freseniusht.com Fresenius Hemotechnology See Fresenius HemoCare Fuji Medical Systems 419 West Avenue P.O. Box 120035 Stamford, CT 06902 (800) 431-1850 FAX: (203) 353-0926 (203) 324-2000 http://www.fujimed.com Fujisawa USA Parkway Center North Deerfield, IL 60015-2548 (847) 317-1088 FAX: (847) 317-7298
Ernest F. Fullam 900 Albany Shaker Road Latham, NY 12110 (800) 833-4024 FAX: (518) 785-8647 (518) 785-5533 http://www.fullam.com Gallard-Schlesinger Industries 777 Zechendorf Boulevard Garden City, NY 11530 (516) 229-4000 FAX: (516) 229-4015 http://www.gallard-schlessinger.com Gambro Box 7373 SE 103 91 Stockholm, Sweden (46) 8 613 65 00 FAX: (46) 8 611 37 31 In the US: COBE Laboratories 225 Union Blvd. Lakewood, CO 80215 (303) 232-6800 FAX: (303) 231-4915 http://www.gambro.com Garner Glass 177 Indian Hill Boulevard Claremont, CA 91711 (909) 624-5071 FAX: (909) 625-0173 http://www.garnerglass.com
GDB (Genome Data Base, Home) Hospital for Sick Children 555 University Avenue Toronto, Ontario M5G 1X8 Canada (416) 813-8744 FAX: (416) 813-8755 http://www.gdb.org The Gel Company 665 Third Street, Suite 240 San Francisco, CA 94107 (800) 256-8596 (415) 247-8760 Gelman Sciences See Pall-Gelman Gemini BioProducts 5115-M Douglas Fir Road Calabasas, CA 90403 (818) 591-3530 FAX: (818) 591-7084 Gen Trak 5100 Campus Drive Plymouth Meeting, PA 19462 (800) 221-7407 FAX: (215) 941-9498 (215) 825-5115 http://www.informagen.com
Garon Plastics 16 Byre Avenue Somerton Park, South Australia 5044 (08) 8294-5126 FAX: (08) 8376-1487 http://www.apache.airnet.com.au/ ∼garon
Genaissance Pharmaceuticals 5 Science Park New Haven, CT 06511 (800) 678-9487 FAX: (203) 562-9377 (203) 773-1450 http://www.genaissance.com
Garren Scientific 9400 Lurline Avenue, Unit E Chatsworth, CA 91311 (800) 342-3725 FAX: (818) 882-3229 (818) 882-6544 http://www.garren-scientific.com
GENAXIS Biotechnology Parc Technologique 10 Avenue Amp`ere Montigny le Bretoneux 78180 France (33) 01-30-14-00-20 FAX: (33) 01-30-14-00-15 http://www.genaxis.com
GATC Biotech AG Jakob-Stadler-Platz 7 D-78467 Constance, Germany (49) 07531-8160-0 FAX: (49) 07531-8160-81 http://www.gatc-biotech.com Gaussian Carnegie Office Park Building 6, Suite 230 Carnegie, PA 15106 (412) 279-6700 FAX: (412) 279-2118 http://www.gaussian.com
GenBank National Center for Biotechnology Information National Library of Medicine/NIH Building 38A, Room 8N805 8600 Rockville Pike Bethesda, MD 20894 (301) 496-2475 FAX: (301) 480-9241 http://www.ncbi.nlm.nih.gov
G.C. Electronics/A.R.C. Electronics 431 Second Street Henderson, KY 42420 (270) 827-8981 FAX: (270) 827-8256 http://www.arcelectronics.com
Gene Codes 640 Avis Drive Ann Arbor, MI 48108 (800) 497-4939 FAX: (734) 930-0145 (734) 769-7249 http://www.genecodes.com
GDB (Genome Data Base, Curation) 2024 East Monument Street, Suite 1200 Baltimore, MD 21205 (410) 955-9705 FAX: (410) 614-0434 http://www.gdb.org
Genemachines 935 Washington Street San Carlos, CA 94070 (650) 508-1634 FAX: (650) 508-1644 (877) 855-4363 http://www.genemachines.com
Suppliers
13 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
Genentech 1 DNA Way South San Francisco, CA 94080 (800) 551-2231 FAX: (650) 225-1600 (650) 225-1000 http://www.gene.com General Scanning/GSI Luminomics 500 Arsenal Street Watertown, MA 02172 (617) 924-1010 FAX: (617) 924-7327 http://www.genescan.com General Valve Division of Parker Hannifin Pneutronics 19 Gloria Lane Fairfield, NJ 07004 (800) GVC-VALV FAX: (800) GVC-1-FAX http://www.pneutronics.com Genespan 19310 North Creek Parkway, Suite 100 Bothell, WA 98011 (800) 231-2215 FAX: (425) 482-3005 (425) 482-3003 http://www.genespan.com Gene Therapy Systems 10190 Telesis Court San Diego, CA 92122 (858) 457-1919 FAX: (858) 623-9494 http://www.genetherapysystems.com G´en´ethon Human Genome Research Center 1 bis rue de l’Internationale 91000 Evry, France (33) 169-472828 FAX: (33) 607-78698 http://www.genethon.fr Genetic Microsystems 34 Commerce Way Wobum, MA 01801 (781) 932-9333 FAX: (781) 932-9433 http://www.genticmicro.com Genetic Mutant Repository See Coriell Institute for Medical Research Genetic Research Instrumentation Gene House Queenborough Lane Rayne, Braintree, Essex CM7 8TF, UK (44) 1376 332900 FAX: (44) 1376 344724 http://www.gri.co.uk Genetics Computer Group 575 Science Drive Madison, WI 53711 (608) 231-5200 FAX: (608) 231-5202 http://www.gcg.com Genetics Institute/American Home Products 87 Cambridge Park Drive Cambridge, MA 02140 (617) 876-1170 FAX: (617) 876-0388 http://www.genetics.com
Genetix 63-69 Somerford Road Christchurch, Dorset BH23 3QA, UK (44) (0) 1202 483900 FAX: (44)(0) 1202 480289 In the US: (877) 436 3849 US FAX: (888) 522 7499 http://www.genetix.co.uk Gene Tools One Summerton Way Philomath, OR 97370 (541) 9292-7840 FAX: (541) 9292-7841 http://www.gene-tools.com Geneva Bioinformatics (GeneBio) S.A. 25 Avenue de Champel CH–1206 Geneva, Switzerland (41) 22-702-9900 FAX: (41) 22-702-9999 http://www.genebio.com GeneWorks P.O. Box 11, Rundle Mall Adelaide, South Australia 5000, Australia 1800 882 555 FAX: (08) 8234 2699 (08) 8234 2644 http://www.geneworks.com Genome Systems (INCYTE) 4633 World Parkway Circle St. Louis, MO 63134 (800) 430-0030 FAX: (314) 427-3324 (314) 427-3222 http://www.genomesystems.com Genomic Solutions 4355 Varsity Drive, Suite E Ann Arbor, MI 48108 (877) GENOMIC FAX: (734) 975-4808 (734) 975-4800 http://www.genomicsolutions.com Genomyx See Beckman Coulter Genosys Biotechnologies 1442 Lake Front Circle, Suite 185 The Woodlands, TX 77380 (281) 363-3693 FAX: (281) 363-2212 http://www.genosys.com Genotech 92 Weldon Parkway St. Louis, MO 63043 (800) 628-7730 FAX: (314) 991-1504 (314) 991-6034 GENSET 876 Prospect Street, Suite 206 La Jolla, CA 92037 (800) 551-5291 FAX: (619) 551-2041 (619) 515-3061 http://www.genset.fr Gensia Laboratories Ltd. 19 Hughes Irvine, CA 92718 (714) 455-4700 FAX: (714) 855-8210
Genta 99 Hayden Avenue, Suite 200 Lexington, MA 02421 (781) 860-5150 FAX: (781) 860-5137 http://www.genta.com GENTEST 6 Henshaw Street Woburn, MA 01801 (800) 334-5229 FAX: (888) 242-2226 (781) 935-5115 FAX: (781) 932-6855 http://www.gentest.com Gentra Systems 15200 25th Avenue N., Suite 104 Minneapolis, MN 55447 (800) 866-3039 FAX: (612) 476-5850 (612) 476-5858 http://www.gentra.com Genzyme 1 Kendall Square Cambridge, MA 02139 (617) 252-7500 FAX: (617) 252-7600 http://www.genzyme.com See also R&D Systems Genzyme Genetics One Mountain Road Framingham, MA 01701 (800) 255-7357 FAX: (508) 872-9080 (508) 872-8400 http://www.genzyme.com George Tiemann & Co. 25 Plant Avenue Hauppauge, NY 11788 (516) 273-0005 FAX: (516) 273-6199 GeSiM Bautzner Landstraße 45 D-01454 Großerkmannsdorf, Germany (49) 351-2695-322 FAX: (49) 351-2695-320 http://www.gesim.de GIBCO/BRL A Division of Life Technologies 1 Kendall Square Grand Island, NY 14072 (800) 874-4226 FAX: (800) 352-1968 (716) 774-6700 http://www.lifetech.com Gilmont Instruments A Division of Barnant Company 28N092 Commercial Avenue Barrington, IL 60010 (800) 637-3739 FAX: (708) 381-7053 http://barnant.com Gilson 3000 West Beltline Highway P.O. Box 620027 Middletown, WI 53562 (800) 445-7661 (608) 836-1551 http://www.gilson.com
Glas-Col Apparatus P.O. Box 2128 Terre Haute, IN 47802 (800) Glas-Col FAX: (812) 234-6975 (812) 235-6167 http://www.glascol.com Glaxo Wellcome Five Moore Drive Research Triangle Park, NC 27709 (800) SGL-AXO5 FAX: (919) 248-2386 (919) 248-2100 http://www.glaxowellcome.com Glen Mills 395 Allwood Road Clifton, NJ 07012 (973) 777-0777 FAX: (973) 777-0070 http://www.glenmills.com Glen Research 22825 Davis Drive Sterling, VA 20166 (800) 327-4536 FAX: (800) 934-2490 (703) 437-6191 FAX: (703) 435-9774 http://www.glenresearch.com Glo Germ P.O. Box 189 Moab, UT 84532 (800) 842-6622 FAX: (435) 259-5930 http://www.glogerm.com Glyco 11 Pimentel Court Novato, CA 94949 (800) 722-2597 FAX: (415) 382-3511 (415) 884-6799 http://www.glyco.com Gould Instrument Systems 8333 Rockside Road Valley View, OH 44125 (216) 328-7000 FAX: (216) 328-7400 http://www.gould13.com Grace Bio-Labs P.O. Box 228 Bend, OR 97709 (541) 318-1208 FAX: (541) 318-0242 http://www.gracebio.com Gralab Instruments See Dimco-Gray GraphPad Software 5755 Oberlin Drive #110 San Diego, CA 92121 (800) 388-4723 FAX: (558) 457-8141 (558) 457-3909 http://www.graphpad.com Graseby Anderson See Andersen Instruments http://www.graseby.com Grass Instrument A Division of Astro-Med 600 East Greenwich Avenue W. Warwick, RI 02893 (800) 225-5167 FAX: (877) 472-7749 http://www.grassinstruments.com
Suppliers
14 CPPS Supplement 45
Current Protocols Selected Suppliers of Reagents and Equipment
Greenacre and Misac Instruments Misac Systems 27 Port Wood Road Ware, Hertfordshire SF12 9NJ, UK (44) 1920 463017 FAX: (44) 1920 465136 Greer Labs 639 Nuway Circle Lenois, NC 28645 (704) 754-5237 http://greerlabs.com
Hamilton Company 4970 Energy Way P.O. Box 10030 Reno, NV 89520 (800) 648-5950 FAX: (775) 856-7259 (775) 858-3000 http://www.hamiltoncompany.com Hamilton Thorne Biosciences 100 Cummings Center, Suite 102C Beverly, MA 01915 http://www.hamiltonthorne.com
Greiner Maybachestrasse 2 Postfach 1162 D-7443 Frickenhausen, Germany (49) 0 91 31/80 79 0 FAX: (49) 0 91 31/80 79 30 http://www.erlangen.com/greiner
Hampton Research 27631 El Lazo Road Laguna Niguel, CA 92677 (800) 452-3899 FAX: (949) 425-1611 (949) 425-6321 http://www.hamptonresearch.com
GSI Lumonics 130 Lombard Street Oxnard, CA 93030 (805) 485-5559 FAX: (805) 485-3310 http://www.gsilumonics.com
Harlan Bioproducts for Science P.O. Box 29176 Indianapolis, IN 46229 (317) 894-7521 FAX: (317) 894-1840 http://www.hbps.com
GTE Internetworking 150 Cambridge Park Drive Cambridge, MA 02140 (800) 472-4565 FAX: (508) 694-4861 http://www.bbn.com GW Instruments 35 Medford Street Somerville, MA 02143 (617) 625-4096 FAX: (617) 625-1322 http://www.gwinst.com H & H Woodworking 1002 Garfield Street Denver, CO 80206 (303) 394-3764 Hacker Instruments 17 Sherwood Lane P.O. Box 10033 Fairfield, NJ 07004 800-442-2537 FAX: (973) 808-8281 (973) 226-8450 http://www.hackerinstruments.com Haemenetics 400 Wood Road Braintree, MA 02184 (800) 225-5297 FAX: (781) 848-7921 (781) 848-7100 http://www.haemenetics.com Halocarbon Products P.O. Box 661 River Edge, NJ 07661 (201) 242-8899 FAX: (201) 262-0019 http://halocarbon.com Hamamatsu Photonic Systems A Division of Hamamatsu 360 Foothill Road P.O. Box 6910 Bridgewater, NJ 08807 (908) 231-1116 FAX: (908) 231-0852 http://www.photonicsonline.com
Harlan Sera-Lab Hillcrest, Dodgeford Lane Belton, Loughborough Leicester LE12 9TE, UK (44) 1530 222123 FAX: (44) 1530 224970 http://www.harlan.com Harlan Teklad P.O. Box 44220 Madison, WI 53744 (608) 277-2070 FAX: (608) 277-2066 http://www.harlan.com Harrick Scientific Corporation 88 Broadway Ossining, NY 10562 (914) 762-0020 FAX: (914) 762-0914 http://www.harricksci.com Harrison Research 840 Moana Court Palo Alto, CA 94306 (650) 949-1565 FAX: (650) 948-0493 Harvard Apparatus 84 October Hill Road Holliston, MA 01746 (800) 272-2775 FAX: (508) 429-5732 (508) 893-8999 http://harvardapparatus.com Harvard Bioscience See Harvard Apparatus Haselton Biologics See JRH Biosciences Hazelton Research Products See Covance Research Products Health Products See Pierce Chemical
Heat Systems-Ultrasonics 1938 New Highway Farmingdale, NY 11735 (800) 645-9846 FAX: (516) 694-9412 (516) 694-9555
HGS Hinimoto Plastics 1-10-24 Meguro-Honcho Megurouko Tokyo 152, Japan 3-3714-7226 FAX:3-3714-4657
Heidenhain Corp 333 East State Parkway Schaumberg, IL 60173 (847) 490-1191 FAX: (847) 490-3931 http://www.heidenhain.com
Hitachi Scientific Instruments Nissei Sangyo America 8100 N. First Street San Elsa, CA 95314 (800) 548-9001 FAX: (408) 432-0704 (408) 432-0520 http://www.hii.hitachi.com
HEKA Instruments 33 Valley Rd. Southboro, MA 01960 (866) 742-0606 FAX: (508) 481-8945 www.heka.com Hellma Cells 11831 Queens Boulevard Forest Hills, NY 11375 (718) 544-9166 FAX: (718) 263-6910 http://www.helmaUSA.com Hellma Postfach 1163 D-79371 M¨ullheim/Baden, Germany (49) 7631-1820 FAX: (49) 7631-13546 http://www.hellma-worldwide.de Henry Schein 135 Duryea Road, Mail Room 150 Melville, NY 11747 (800) 472-4346 FAX: (516) 843-5652 http://www.henryschein.com Heraeus Kulzer 4315 South Lafayette Boulevard South Bend, IN 46614 (800) 343-5336 (219) 291-0661 http://www.kulzer.com Heraeus Sepatech See Kendro Laboratory Products Hercules Aqualon Aqualon Division Hercules Research Center, Bldg. 8145 500 Hercules Road Wilmington, DE 19899 (800) 345-0447 FAX: (302) 995-4787 http://www.herc.com/aqualon/pharma Heto-Holten A/S Gydevang 17-19 DK-3450 Allerod, Denmark (45) 48-16-62-00 FAX: (45) 48-16-62-97 Distributed by ATR Hettich-Zentrifugen See Andreas Hettich Hewlett-Packard 3000 Hanover Street Mailstop 20B3 Palo Alto, CA 94304 (650) 857-1501 FAX: (650) 857-5518 http://www.hp.com
Hi-Tech Scientific Brunel Road Salisbury, Wiltshire, SP2 7PU UK (44) 1722-432320 (800) 344-0724 (US only) http://www.hi-techsci.co.uk Hoechst AG See Aventis Pharmaceutical Hoefer Scientific Instruments Division of Amersham-Pharmacia Biotech 800 Centennial Avenue Piscataway, NJ 08855 (800) 227-4750 FAX: (877) 295-8102 http://www.apbiotech.com Hoffman-LaRoche 340 Kingsland Street Nutley, NJ 07110 (800) 526-0189 FAX: (973) 235-9605 (973) 235-5000 http://www.rocheUSA.com Holborn Surgical and Medical Instruments Westwood Industrial Estate Ramsgate Road Margate, Kent CT9 4JZ UK (44) 1843 296666 FAX: (44) 1843 295446 Honeywell 101 Columbia Road Morristown, NJ 07962 (973) 455-2000 FAX: (973) 455-4807 http://www.honeywell.com Honeywell Specialty Films P.O. Box 1039 101 Columbia Road Morristown, NJ 07962 (800) 934-5679 FAX: (973) 455-6045 http://www.honeywellspecialtyfilms.com Hood Thermo-Pad Canada Comp. 20, Site 61A, RR2 Summerland, British Columbia V0H 1Z0 Canada (800) 665-9555 FAX: (250) 494-5003 (250) 494-5002 http://www.thermopad.com
Suppliers
15 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
Horiba Instruments 17671 Armstrong Avenue Irvine, CA 92714 (949) 250-4811 FAX: (949) 250-0924 http://www.horiba.com Hoskins Manufacturing 10776 Hall Road P.O. Box 218 Hamburg, MI 48139 (810) 231-1900 FAX: (810) 231-4311 http://www.hoskinsmfgco.com Hosokawa Micron Powder Systems 10 Chatham Road Summit, NJ 07901 (800) 526-4491 FAX: (908) 273-7432 (908) 273-6360 http://www.hosokawamicron.com HT Biotechnology Unit 4 61 Ditton Walk Cambridge CB5 8QD, UK (44) 1223-412583 Hugo Sachs Electronik Postfach 138 7806 March-Hugstetten, Germany D-79229(49) 7665-92000 FAX: (49) 7665-920090 Human Biologics International 7150 East Camelback Road, Suite 245 Scottsdale, AZ 85251 (480) 990-2005 FAX: (480)-990-2155 http://www.humanbiological.com Human Genetic Mutant Cell Repository See Coriell Institute for Medical Research HVS Image P.O. Box 100 Hampton, Middlesex TW12 2YD, UK FAX: (44) 208 783 1223 In the US: (800) 225-9261 FAX: (888) 483-8033 http://www.hvsimage.com Hybaid 111-113 Waldegrave Road Teddington, Middlesex TW11 8LL, UK (44) 0 1784 42500 FAX: (44) 0 1784 248085 http://www.hybaid.co.uk Hybaid Instruments 8 East Forge Parkway Franklin, MA 02028 (888)4-HYBAID FAX: (508) 541-3041 (508) 541-6918 http://www.hybaid.com Hybridon 155 Fortune Boulevard Milford, MA 01757 (508) 482-7500 FAX: (508) 482-7510 http://www.hybridon.com
HyClone Laboratories 1725 South HyClone Road Logan, UT 84321 (800) HYCLONE FAX: (800) 533-9450 (801) 753-4584 FAX: (801) 750-0809 http://www.hyclone.com Hypromatrix 100 Barber Avenue Worcester, MA 01606 (866) 742-6522 FAX: (508) 302-0748 http://www.hypromatrix.com Hyseq 670 Almanor Avenue Sunnyvale, CA 94086 (408) 524-8100 FAX: (408) 524-8141 http://www.hyseq.com IBA GmbH 1508 South Grand Blvd. St. Louis, MO 63104 (877) 422-4624 FAX: (888) 531-6813 http://www.iba-go.com IBF Biotechnics See Sepracor IBI (International Biotechnologies) See Eastman Kodak For technical service (800) 243-2555 (203) 786-5600 ICN Biochemicals See ICN Biomedicals ICN Biomedicals 3300 Hyland Avenue Costa Mesa, CA 92626 (800) 854-0530 FAX: (800) 334-6999 (714) 545-0100 FAX: (714) 641-7275 http://www.icnbiomed.com ICN Flow and Pharmaceuticals See ICN Biomedicals ICN Immunobiochemicals See ICN Biomedicals ICN Radiochemicals See ICN Biomedicals ICONIX 100 King Street West, Suite 3825 Toronto, Ontario M5X 1E3 Canada (416) 410-2411 FAX: (416) 368-3089 http://www.iconix.com ICRT (Imperial Cancer Research Technology) Sardinia House Sardinia Street London WC2A 3NL, UK (44) 1712-421136 FAX: (44) 1718-314991 Idea Scientific Company P.O. Box 13210 Minneapolis, MN 55414 (800) 433-2535 FAX: (612) 331-4217 http://www.ideascientific.com IEC See International Equipment Co.
IITC 23924 Viclory Boulevard Woodland Hills, CA 91387 (888) 414-4482 (818) 710-1556 FAX: (818) 992-5165 http://www.iitcinc.com IKA Works 2635 N. Chase Parkway, SE Wilmington, NC 28405 (910) 452-7059 FAX: (910) 452-7693 http://www.ika.net Ikegami Electronics 37 Brook Avenue Maywood, NJ 07607 (201) 368-9171 FAX: (201) 569-1626 Ikemoto Scientific Technology 25-11 Hongo 3-chome, Bunkyo-ku Tokyo 101-0025, Japan (81) 3-3811-4181 FAX: (81) 3-3811-1960 Imagenetics See ATC Diagnostics Imaging Research c/o Brock University 500 Glenridge Avenue St. Catharines, Ontario L2S 3A1 Canada (905) 688-2040 FAX: (905) 685-5861 http://www.imaging.brocku.ca Imclone Systems 180 Varick Street New York, NY 10014 (212) 645-1405 FAX: (212) 645-2054 http://www.imclone.com IMCO Corporation LTD., AB P.O. Box 21195 SE-100 31 Stockholm, Sweden 46-8-33-53-09 FAX:46-8-728-47-76 http://www.imcocorp.se Imgenex Corporation 11175 Flintkote Avenue Suite E San Diego, CA 92121 (888) 723-4363 FAX: (858) 642-0937 (858) 642.0978 http://www.imgenex.com
Immunochemistry Technologies 9401 James Avenue, South Suite 155 Bloomington, MN 55431 (800) 829-3194 FAX: (952) 888-8988 (952) 888-8788 http://www.immunochemistry.com Immunocorp 1582 W. Deere Avenue Suite C Irvine, CA 92606 (800) 446-3063 http://www.immunocorp.com Immunotech 130, av. Delattre de Tassigny B.P. 177 13276 Marseilles Cedex 9 France (33) 491-17-27-00 FAX: (33) 491-41-43-58 http://www.immunotech.fr Imperial Chemical Industries Imperial Chemical House Millbank, London SW1P 3JF, UK (44) 171-834-4444 FAX: (44)171-834-2042 http://www.ici.com Inceltech See New Brunswick Scientific Incstar See DiaSorin Incyte 6519 Dumbarton Circle Fremont, CA 94555 (510) 739-2100 FAX: (510) 739-2200 http://www.incyte.com Incyte Pharmaceuticals 3160 Porter Drive Palo Alto, CA 94304 (877) 746-2983 FAX: (650) 855-0572 (650) 855-0555 http://www.incyte.com Individual Monitoring Systems 6310 Harford Road Baltimore, MD 21214
IMICO Calle Vivero, No. 5-4a Planta E-28040, Madrid, Spain (34) 1-535-3960 FAX: (34) 1-535-2780
Indo Fine Chemical P.O. Box 473 Somerville, NJ 08876 (888) 463-6346 FAX: (908) 359-1179 (908) 359-6778 http://www.indofinechemical.com
Immunex 51 University Street Seattle, WA 98101 (206) 587-0430 FAX: (206) 587-0606 http://www.immunex.com
Industrial Acoustics 1160 Commerce Avenue Bronx, NY 10462 (718) 931-8000 FAX: (718) 863-1138 http://www.industrialacoustics.com
Suppliers
16 CPPS Supplement 45
Current Protocols Selected Suppliers of Reagents and Equipment
Inex Pharmaceuticals 100-8900 Glenlyon Parkway Glenlyon Business Park Burnaby, British Columbia V5J 5J8 Canada (604) 419-3200 FAX: (604) 419-3201 http://www.inexpharm.com Ingold, Mettler, Toledo 261 Ballardvale Street Wilmington, MA 01887 (800) 352-8763 FAX: (978) 658-0020 (978) 658-7615 http://www.mt.com Innogenetics N.V. Technologie Park 6 B-9052 Zwijnaarde Belgium (32) 9-329-1329 FAX: (32) 9-245-7623 http://www.innogenetics.com Innovative Medical Services 1725 Gillespie Way El Cajon, CA 92020 (619) 596-8600 FAX: (619) 596-8700 http://www.imspure.com
Instrumentarium P.O. Box 300 00031 Instrumentarium Helsinki, Finland (10) 394-5566 http://www.instrumentarium.fi
Intermountain Scientific 420 N. Keys Drive Kaysville, UT 84037 (800) 999-2901 FAX: (800) 574-7892 (801) 547-5047 FAX: (801) 547-5051 http://www.bioexpress.com
Instruments SA Division Jobin Yvon 16-18 Rue du Canal 91165 Longjumeau, Cedex, France (33)1 6454-1300 FAX: (33)1 6909-9319 http://www.isainc.com
International Biotechnologies (IBI) See Eastman Kodak
Instrutech 20 Vanderventer Avenue, Suite 101E Port Washington, NY 11050 (516) 883-1300 FAX: (516) 883-1558 http://www.instrutech.com Integrated DNA Technologies 1710 Commercial Park Coralville, Iowa 52241 (800) 328-2661 FAX: (319) 626-8444 http://www.idtdna.com Integrated Genetics See Genzyme Genetics
Innovative Research 3025 Harbor Lane N, Suite 300 Plymouth, MN 55447 (612) 519-0105 FAX: (612) 519-0239 http://www.inres.com
Integrated Scientific Imaging Systems 3463 State Street, Suite 431 Santa Barbara, CA 93105 (805) 692-2390 FAX: (805) 692-2391 http://www.imagingsystems.com
Innovative Research of America 2 N. Tamiami Trail, Suite 404 Sarasota, FL 34236 (800) 421-8171 FAX: (800) 643-4345 (941) 365-1406 FAX: (941) 365-1703 http://www.innovrsrch.com
Integrated Separation Systems (ISS) See OWL Separation Systems
Inotech Biosystems 15713 Crabbs Branch Way, #110 Rockville, MD 20855 (800) 635-4070 FAX: (301) 670-2859 (301) 670-2850 http://www.inotechintl.com INOVISION 22699 Old Canal Road Yorba Linda, CA 92887 (714) 998-9600 FAX: (714) 998-9666 http://www.inovision.com
IntelliGenetics See Oxford Molecular Group Interactiva BioTechnologie Sedanstrasse 10 D-89077 Ulm, Germany (49) 731-93579-290 FAX: (49) 731-93579-291 http://www.interactiva.de Interchim 213 J.F. Kennedy Avenue B.P. 1140 Montlucon 03103 France (33) 04-70-03-83-55 FAX: (33) 04-70-03-93-60
Instech Laboratories 5209 Militia Hill Road Plymouth Meeting, PA 19462 (800) 443-4227 FAX: (610) 941-0134 (610) 941-0132 http://www.instechlabs.com
Interfocus 14/15 Spring Rise Falcover Road Haverhill, Suffolk CB9 7XU, UK (44) 1440 703460 FAX: (44) 1440 704397 http://www.interfocus.ltd.uk
Instron 100 Royall Street Canton, MA 02021 (800) 564-8378 FAX: (781) 575-5725 (781) 575-5000 http://www.instron.com
Intergen 2 Manhattanville Road Purchase, NY 10577 (800) 431-4505 FAX: (800) 468-7436 (914) 694-1700 FAX: (914) 694-1429 http://www.intergenco.com
International Equipment Co. (IEC) See Thermoquest International Institute for the Advancement of Medicine 1232 Mid-Valley Drive Jessup, PA 18434 (800) 486-IIAM FAX: (570) 343-6993 (570) 496-3400 http://www.iiam.org International Light 17 Graf Road Newburyport, MA 01950 (978) 465-5923 FAX: (978) 462-0759 International Market Supply (I.M.S.) Dane Mill Broadhurst Lane Congleton, Cheshire CW12 1LA, UK (44) 1260 275469 FAX: (44) 1260 276007 International Marketing Services See International Marketing Ventures International Marketing Ventures 6301 Ivy Lane, Suite 408 Greenbelt, MD 20770 (800) 373-0096 FAX: (301) 345-0631 (301) 345-2866 http://www.imvlimited.com International Products 201 Connecticut Drive Burlington, NJ 08016 (609) 386-8770 FAX: (609) 386-8438 http://[email protected] Intracel Corporation Bartels Division 2005 Sammamish Road, Suite 107 Issaquah, WA 98027 (800) 542-2281 FAX: (425) 557-1894 (425) 392-2992 http://www.intracel.com Invitrogen 1600 Faraday Avenue Carlsbad, CA 92008 (800) 955-6288 FAX: (760) 603-7201 (760) 603-7200 http://www.invitrogen.com In Vivo Metric P.O. Box 249 Healdsburg, CA 95448 (707) 433-4819 FAX: (707) 433-2407
IRORI 9640 Towne Center Drive San Diego, CA 92121 (858) 546-1300 FAX: (858) 546-3083 http://www.irori.com Irvine Scientific 2511 Daimler Street Santa Ana, CA 92705 (800) 577-6097 FAX: (949) 261-6522 (949) 261-7800 http://www.irvinesci.com ISC BioExpress 420 North Kays Drive Kaysville, UT 84037 (800) 999-2901 FAX: (800) 574-7892 (801) 547-5047 http://www.bioexpress.com ISCO P.O. Box 5347 4700 Superior Lincoln, NE 68505 (800) 228-4373 FAX: (402) 464-0318 (402) 464-0231 http://www.isco.com Isis Pharmaceuticals Carlsbad Research Center 2292 Faraday Avenue Carlsbad, CA 92008 (760) 931-9200 http://www.isip.com Isolabs See Wallac ISS See Integrated Separation Systems J & W Scientific See Agilent Technologies J.A. Webster 86 Leominster Road Sterling, MA 01564 (800) 225-7911 FAX: (978) 422-8959 http://www.jawebster.com J.T. Baker See Mallinckrodt Baker 222 Red School Lane Phillipsburg, NJ 08865 (800) JTBAKER FAX: (908) 859-6974 http://www.jtbaker.com Jackson ImmunoResearch Laboratories P.O. Box 9 872 W. Baltimore Pike West Grove, PA 19390 (800) 367-5296 FAX: (610) 869-0171 (610) 869-4024 http://www.jacksonimmuno.com The Jackson Laboratory 600 Maine Street Bar Harbor, ME 04059 (800) 422-6423 FAX: (207) 288-5079 (207) 288-6000 http://www.jax.org
Suppliers
17 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
Jaece Industries 908 Niagara Falls Boulevard North Tonawanda, NY 14120 (716) 694-2811 FAX: (716) 694-2811 http://www.jaece.com Jandel Scientific See SPSS Janke & Kunkel See Ika Works Janssen Life Sciences Products See Amersham Janssen Pharmaceutica 1125 Trenton-Harbourton Road Titusville, NJ 09560 (609) 730-2577 FAX: (609) 730-2116 http://us.janssen.com Jasco 8649 Commerce Drive Easton, MD 21601 (800) 333-5272 FAX: (410) 822-7526 (410) 822-1220 http://www.jascoinc.com Jena Bioscience Loebstedter Str. 78 07749 Jena, Germany (49) 3641-464920 FAX: (49) 3641-464991 http://www.jenabioscience.com Jencons Scientific 800 Bursca Drive, Suite 801 Bridgeville, PA 15017 (800) 846-9959 FAX: (412) 257-8809 (412) 257-8861 http://www.jencons.co.uk JEOL Instruments 11 Dearborn Road Peabody, MA 01960 (978) 535-5900 FAX: (978) 536-2205 http://www.jeol.com/index.html Jewett 750 Grant Street Buffalo, NY 14213 (800) 879-7767 FAX: (716) 881-6092 (716) 881-0030 http://www.JewettInc.com
Johnston Matthey Chemicals Orchard Road Royston, Hertfordshire SG8 5HE, UK (44) 1763-253000 FAX: (44) 1763-253466 http://www.chemicals.matthey.com
Karl Storz K¨oningin-Elisabeth Str. 60 D-14059 Berlin, Germany (49) 30-30 69 09-0 FAX: (49) 30-30 19 452 http://www.karlstorz.de
Jolley Consulting and Research 683 E. Center Street, Unit H Grayslake, IL 60030 (847) 548-2330 FAX: (847) 548-2984 http://www.jolley.com
KaVo EWL P.O. Box 1320 D-88293 Leutkirch im Allg¨au, Germany (49) 7561-86-0 FAX: (49) 7561-86-371 http://www.kavo.com/english/ startseite.htm
Jordan Scientific See Shelton Scientific
Keithley Instruments 28775 Aurora Road Cleveland, OH 44139 (800) 552-1115 FAX: (440) 248-6168 (440) 248-0400 http://www.keithley.com
Jorgensen Laboratories 1450 N. Van Buren Avenue Loveland, CO 80538 (800) 525-5614 FAX: (970) 663-5042 (970) 669-2500 http://www.jorvet.com JRH Biosciences and JR Scientific 13804 W. 107th Street Lenexa, KS 66215 (800) 231-3735 FAX: (913) 469-5584 (913) 469-5580 Jule Bio Technologies 25 Science Park, #14, Suite 695 New Haven, CT 06511 (800) 648-1772 FAX: (203) 786-5489 (203) 786-5490 http://hometown.aol.com/precastgel/ index.htm
Kemin 2100 Maury Street, Box 70 Des Moines, IA 50301 (515) 266-2111 FAX: (515) 266-8354 http://www.kemin.com Kemo 3 Brook Court, Blakeney Road Beckenham, Kent BR3 1HG, UK (44) 0181 658 3838 FAX: (44) 0181 658 4084 http://www.kemo.com Kendall 15 Hampshire Street Mansfield, MA 02048 (800) 962-9888 FAX: (800) 724-1324 http://www.kendallhq.com
K.R. Anderson 2800 Bowers Avenue Santa Clara, CA 95051 (800) 538-8712 FAX: (408) 727-2959 (408) 727-2800 http://www.kranderson.com
Kendro Laboratory Products 31 Pecks Lane Newtown, CT 06470 (800) 522-SPIN FAX: (203) 270-2166 (203) 270-2080 http://www.kendro.com
Kabi Pharmacia Diagnostics See Pharmacia Diagnostics
Kendro Laboratory Products P.O. Box 1220 Am Kalkberg D-3360 Osterod, Germany (55) 22-316-213 FAX: (55) 22-316-202 http://www.heraeus-instruments.de
John’s Scientific See VWR Scientific
Kanthal H.P. Reid 1 Commerce Boulevard P.O. Box 352440 Palm Coasl, FL 32135 (904) 445-2000 FAX: (904) 446-2244 http://www.kanthal.com
John Weiss and Sons 95 Alston Drive Bradwell Abbey Milton Keynes, Buckinghamshire MK1 4HF UK (44) 1908-318017 FAX: (44) 1908-318708
Kapak 5305 Parkdale Drive St. Louis Park, MN 55416 (800) KAPAK-57 FAX: (612) 541-0735 (612) 541-0730 http://www.kapak.com
Johnson & Johnson Medical 2500 Arbrook Boulevard East Arlington, TX 76004 (800) 423-4018 http://www.jnjmedical.com
Karl Hecht Stettener Str. 22-24 D-97647 Sondheim Rh¨on, Germany (49) 9779-8080 FAX: (49) 9779-80888
Kent Laboratories 23404 NE 8th Street Redmond, WA 98053 (425) 868-6200 FAX: (425) 868-6335 http://www.kentlabs.com Kent Scientific 457 Bantam Road, #16 Litchfield, CT 06759 (888) 572-8887 FAX: (860) 567-4201 (860) 567-5496 http://www.kentscientific.com Keuffel & Esser See Azon
Keystone Scientific Penn Eagle Industrial Park 320 Rolling Ridge Drive Bellefonte, PA 16823 (800) 437-2999 FAX: (814) 353-2305 (814) 353-2300 Ext 1 http://www.keystonescientific.com Kimble/Kontes Biotechnology 1022 Spruce Street P.O. Box 729 Vineland, NJ 08360 (888) 546-2531 FAX: (856) 794-9762 (856) 692-3600 http://www.kimble-kontes.com Kinematica AG Luzernerstrasse 147a CH-6014 Littau-Luzern, Switzerland (41) 41 2501257 FAX: (41) 41 2501460 http://www.kinematica.ch Kin-Tek 504 Laurel Street LaMarque, TX 77568 (800) 326-3627 FAX: (409) 938-3710 http://www.kin-tek.com Kipp & Zonen 125 Wilbur Place Bohemia, NY 11716 (800) 645-2065 FAX: (516) 589-2068 (516) 589-2885 http://www.kippzonen.thomasregister. com/olc/kippzonen Kirkegaard & Perry Laboratories 2 Cessna Court Gaithersburg, MD 20879 (800) 638-3167 FAX: (301) 948-0169 (301) 948-7755 http://www.kpl.com Kodak See Eastman Kodak Kontes Glass See Kimble/Kontes Biotechnology Kontron Instruments AG Postfach CH-8010 Zurich, Switzerland 41-1-733-5733 FAX:41-1-733-5734 David Kopf Instruments P.O. Box 636 Tujunga, CA 91043 (818) 352-3274 FAX: (818) 352-3139 Kraft Apparatus See Glas-Col Apparatus Kramer Scientific Corporation 711 Executive Boulevard Valley Cottage, NY 10989 (845) 267-5050 FAX: (845) 267-5550 Kulite Semiconductor Products 1 Willow Tree Road Leonia, NJ 07605 (201) 461-0900 FAX: (201) 461-0990 http://www.kulite.com
Suppliers
18 CPPS Supplement 45
Current Protocols Selected Suppliers of Reagents and Equipment
Lab-Line Instruments 15th & Bloomingdale Avenues Melrose Park, IL 60160 (800) LAB-LINE FAX: (708) 450-5830 FAX: (800) 450-4LAB http://www.labline.com Lab Products 742 Sussex Avenue P.O. Box 639 Seaford, DE 19973 (800) 526-0469 FAX: (302) 628-4309 (302) 628-4300 http://www.labproductsinc.com LabRepco 101 Witmer Road, Suite 700 Horsham, PA 19044 (800) 521-0754 FAX: (215) 442-9202 http://www.labrepco.com Lab Safety Supply P.O. Box 1368 Janesville, WI 53547 (800) 356-0783 FAX: (800) 543-9910 (608) 754-7160 FAX: (608) 754-1806 http://www.labsafety.com Lab-Tek Products See Nalge Nunc International Labconco 8811 Prospect Avenue Kansas City, MO 64132 (800) 821-5525 FAX: (816) 363-0130 (816) 333-8811 http://www.labconco.com Labindustries See Barnstead/Thermolyne Labnet International P.O. Box 841 Woodbridge, NJ 07095 (888) LAB-NET1 FAX: (732) 417-1750 (732) 417-0700 http://www.nationallabnet.com LABO-MODERNE 37 rue Dombasle Paris 75015 France (33) 01-45-32-62-54 FAX: (33) 01-45-32-01-09 http://www.labomoderne.com/fr Laboratory of Immunoregulation National Institute of Allergy and Infectious Diseases/NIH 9000 Rockville Pike Building 10, Room 11B13 Bethesda, MD 20892 (301) 496-1124 Laboratory Supplies 29 Jefry Lane Hicksville, NY 11801 (516) 681-7711
Labscan Limited Stillorgan Industrial Park Stillorgan Dublin, Ireland (353) 1-295-2684 FAX: (353) 1-295-2685 http://www.labscan.ie Labsystems See Thermo Labsystems Labsystems Affinity Sensors Saxon Way, Bar Hill Cambridge CB3 8SL, UK 44 (0) 1954 789976 FAX:44 (0) 1954 789417 http://www.affinity-sensors.com Labtronics 546 Governors Road Guelph, Ontario N1K 1E3, Canada (519) 763-4930 FAX: (519) 836-4431 http://www.labtronics.com Labtronix Manufacturing 3200 Investment Boulevard Hayward, CA 94545 (510) 786-3200 FAX: (510) 786-3268 http://www.labtronix.com Lafayette Instrument 3700 Sagamore Parkway North P.O. Box 5729 Lafayette, IN 47903 (800) 428-7545 FAX: (765) 423-4111 (765) 423-1505 http://www.lafayetteinstrument.com Lambert Instruments Turfweg 4 9313 TH Leutingewolde The Netherlands (31) 50-5018461 FAX: (31) 50-5010034 http://www.lambert-instruments.com Lampire Biological Laboratories P.O. Box 270 Pipersville, PA 18947 (215) 795-2538 Fax: (215) 795-0237 http://www.lampire.com Lancaster Synthesis P.O. Box 1000 Windham, NH 03087 (800) 238-2324 FAX: (603) 889-3326 (603) 889-3306 http://www.lancastersynthesis-us.com Lancer 140 State Road 419 Winter Springs, FL 32708 (800) 332-1855 FAX: (407) 327-1229 (407) 327-8488 http://www.lancer.com
LaVision GmbH Gerhard-Gerdes-Str. 3 D-37079 Goettingen, Germany (49) 551-50549-0 FAX: (49) 551-50549-11 http://www.lavision.de
Leica Microsystems Imneuenheimer Feld 518 D-69120 Heidelberg, Germany (49) 6221-41480 FAX: (49) 6221-414833 http://www.leica-microsystems.com
Lawshe See Advanced Process Supply
Leinco Technologies 359 Consort Drive St. Louis, MO 63011 (314) 230-9477 FAX: (314) 527-5545 http://www.leinco.com
Laxotan 20, rue Leon Blum 26000 Valence, France (33) 4-75-41-91-91 FAX: (33) 4-75-41-91-99 http://www.latoxan.com
Leitz U.S.A. See Leica
LC Laboratories 165 New Boston Street Woburn, MA 01801 (781) 937-0777 FAX: (781) 938-5420 http://www.lclaboratories.com
LenderKing Metal Products 8370 Jumpers Hole Road Millersville, MD 21108 (410) 544-8795 FAX: (410) 544-5069 http://www.lenderking.com
LC Packings 80 Carolina Street San Francisco, CA 94103 (415) 552-1855 FAX: (415) 552-1859 http://www.lcpackings.com
Letica Scientific Instruments Panlab s.i., c/Loreto 50 08029 Barcelona, Spain (34) 93-419-0709 FAX: (34) 93-419-7145 www.panlab-sl.com
LC Services See LC Laboratories LECO 3000 Lakeview Avenue St. Joseph, MI 49085 (800) 292-6141 FAX: (616) 982-8977 (616) 985-5496 http://www.leco.com Lederle Laboratories See Wyeth-Ayerst Lee Biomolecular Research Laboratories 11211 Sorrento Valley Road, Suite M San Diego, CA 92121 (858) 452-7700
Leybold-Heraeus Trivac DZA 5700 Mellon Road Export, PA 15632 (412) 327-5700 LI-COR Biotechnology Division 4308 Progressive Avenue Lincoln, NE 68504 (800) 645-4267 FAX: (402) 467-0819 (402) 467-0700 http://www.licor.com Life Science Laboratories See Adaptive Biosystems
The Lee Company 2 Pettipaug Road P.O. Box 424 Westbrook, CT 06498 (800) LEE-PLUG FAX: (860) 399-7058 (860) 399-6281 http://www.theleeco.com
Life Science Resources Two Corporate Center Drive Melville, NY 11747 (800) 747-9530 FAX: (516) 844-5114 (516) 844-5085 http://www.astrocam.com
Lee Laboratories 1475 Athens Highway Grayson, GA 30017 (800) 732-9150 FAX: (770) 979-9570 (770) 972-4450 http://www.leelabs.com
Life Sciences 2900 72nd Street North St. Petersburg, FL 33710 (800) 237-4323 FAX: (727) 347-2957 (727) 345-9371 http://www.lifesci.com
Leica 111 Deer Lake Road Deerfield, IL 60015 (800) 248-0123 FAX: (847) 405-0147 (847) 405-0123 http://www.leica.com
Life Technologies 9800 Medical Center Drive P.O. Box 6482 Rockville, MD 20849 (800) 828-6686 FAX: (800) 331-2286 http://www.lifetech.com
Suppliers
19 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
Lifecodes 550 West Avenue Stamford, CT 06902 (800) 543-3263 FAX: (203) 328-9599 (203) 328-9500 http://www.lifecodes.com Lightnin 135 Mt. Read Boulevard Rochester, NY 14611 (888) MIX-BEST FAX: (716) 527-1742 (716) 436-5550 http://www.lightnin-mixers.com Linear Drives Luckyn Lane, Pipps Hill Basildon, Essex SS14 3BW, UK (44) 1268-287070 FAX: (44) 1268-293344 http://www.lineardrives.com Linscott’s Directory 4877 Grange Road Santa Rosa, CA 95404 (707) 544-9555 FAX: (415) 389-6025 http://www.linscottsdirectory.co.uk Linton Instrumentation Unit 11, Forge Business Center Upper Rose Lane Palgrave, Diss, Norfolk IP22 1AP, UK (44) 1-379-651-344 FAX: (44) 1-379-650-970 http://www.lintoninst.co.uk List Biological Laboratories 501-B Vandell Way Campbell, CA 95008 (800) 726-3213 FAX: (408) 866-6364 (408) 866-6363 http://www.listlabs.com LKB Instruments See Amersham Pharmacia Biotech Lloyd Laboratories 604 West Thomas Avenue Shenandoah, IA 51601 (800) 831-0004 FAX: (712) 246-5245 (712) 246-4000 http://www.lloydinc.com Loctite 1001 Trout Brook Crossing Rocky Hill, CT 06067 (860) 571-5100 FAX: (860)571-5465 http://www.loctite.com Lofstrand Labs 7961 Cessna Avenue Gaithersburg, MD 20879 (800) 541-0362 FAX: (301) 948-9214 (301) 330-0111 http://www.lofstrand.com Lomir Biochemical 99 East Main Street Malone, NY 12953 (877) 425-3604 FAX: (518) 483-8195 (518) 483-7697 http://www.lomir.com
LSL Biolafitte 10 rue de Temara 7810C St.-Germain-en-Laye, France (33) 1-3061-5260 FAX: (33) 1-3061-5234 Ludger Ltd The Oxford BioBusiness Centre Littlemore Park Oxford OX4 4SS, UK (44) 870-085-7011 Ludl Electronic Products 171 Brady Avenue Hawthorne, NY 10532 (888) 769-6111 FAX: (914) 769-4759 (914) 769-6111 http://www.ludl.com Lumigen 24485 W. Ten Mile Road Southfield, MI 48034 (248) 351-5600 FAX: (248) 351-0518 http://www.lumigen.com Luminex 12212 Technology Boulevard Austin, TX 78727 (888) 219-8020 FAX: (512) 258-4173 (512) 219-8020 http://www.luminexcorp.com LYNX Therapeutics 25861 Industrial Boulevard Hayward, CA 94545 (510) 670-9300 FAX: (510) 670-9302 http://www.lynxgen.com Lyphomed 3 Parkway North Deerfield, IL 60015 (847) 317-8100 FAX: (847) 317-8600 M.E.D. Associates See Med Associates Macherey-Nagel 6 South Third Street, #402 Easton, PA 18042 (610) 559-9848 FAX: (610) 559-9878 http://www.macherey-nagel.com Macherey-Nagel Valencienner Strasse 11 P.O. Box 101352 D-52313 Dueren, Germany (49) 2421-969141 FAX: (49) 2421-969199 http://www.macherey-nagel.ch Mac-Mod Analytical 127 Commons Court Chadds Ford, PA 19317 800-441-7508 FAX: (610) 358-5993 (610) 358-9696 http://www.mac-mod.com
Mallinckrodt Baker 222 Red School Lane Phillipsburg, NJ 08865 (800) 582-2537 FAX: (908) 859-6974 (908) 859-2151 http://www.mallbaker.com
Mathsoft 1700 Westlake Avenue N., Suite 500 Seattle, WA 98109 (800) 569-0123 FAX: (206) 283-8691 (206) 283-8802 http://www.mathsoft.com
Mallinckrodt Chemicals 16305 Swingley Ridge Drive Chesterfield, MD 63017 (314) 530-2172 FAX: (314) 530-2563 http://www.mallchem.com
Matreya 500 Tressler Street Pleasant Gap, PA 16823 (814) 359-5060 FAX: (814) 359-5062 http://www.matreya.com
Malven Instruments Enigma Business Park Grovewood Road Malven, Worchestershire WR 141 XZ, United Kingdom Marinus 1500 Pier C Street Long Beach, CA 90813 (562) 435-6522 FAX: (562) 495-3120 Markson Science c/o Whatman Labs Sales P.O. Box 1359 Hillsboro, OR 97123 (800) 942-8626 FAX: (503) 640-9716 (503) 648-0762 Marsh Biomedical Products 565 Blossom Road Rochester, NY 14610 (800) 445-2812 FAX: (716) 654-4810 (716) 654-4800 http://www.biomar.com Marshall Farms USA 5800 Lake Bluff Road North Rose, NY 14516 (315) 587-2295 e-mail: [email protected] Martek 6480 Dobbin Road Columbia, MD 21045 (410) 740-0081 FAX: (410) 740-2985 http://www.martekbio.com Martin Supply Distributor of Gerber Scientific 2740 Loch Raven Road Baltimore, MD 21218 (800) 282-5440 FAX: (410) 366-0134 (410) 366-1696 Mast Immunosystems 630 Clyde Court Mountain View, CA 94043 (800) 233-MAST FAX: (650) 969-2745 (650) 961-5501 http://www.mastallergy.com Matheson Gas Products P.O. Box 624 959 Route 46 East Parsippany, NJ 07054 (800) 416-2505 FAX: (973) 257-9393 (973) 257-1100 http://www.mathesongas.com
Matrigel See Becton Dickinson Labware Matrix Technologies 22 Friars Drive Hudson, NH 03051 (800) 345-0206 FAX: (603) 595-0106 (603) 595-0505 http://www.matrixtechcorp.com MatTek Corp. 200 Homer Ave. Ashland, Massachusetts 01721 (508) 881-6771 FAX: (508) 879-1532 http://www.mattek.com Maxim Medical 89 Oxford Road Oxford OX2 9PD United Kingdom 44 (0)1865-865943 FAX:44 (0)1865-865291 http://www.maximmed.com Mayo Clinic Section on Engineering Project #ALA-1, 1982 200 1st Street SW Rochester, MN 55905 (507) 284-2511 FAX: (507) 284-5988 McGaw See B. Braun-McGaw McMaster-Carr 600 County Line Road Elmhurst, IL 60126 (630) 833-0300 FAX: (630) 834-9427 http://www.mcmaster.com McNeil Pharmaceutical See Ortho McNeil Pharmaceutical MCNC 3021 Cornwallis Road P.O. Box 12889 Research Triangle Park, NC 27709 (919) 248-1800 FAX: (919) 248-1455 http://www.mcnc.org MD Industries 5 Revere Drive, Suite 415 Northbrook, IL 60062 (800) 421-8370 FAX: (847) 498-2627 (708) 339-6000 http://www.mdindustries.com
Suppliers
20 CPPS Supplement 45
Current Protocols Selected Suppliers of Reagents and Equipment
MDS Nordion 447 March Road P.O. Box 13500 Kanata, Ontario K2K 1X8, Canada (800) 465-3666 FAX: (613) 592-6937 (613) 592-2790 http://www.mds.nordion.com
Megazyme Bray Business Park Bray, County Wicklow Ireland (353) 1-286-1220 FAX: (353) 1-286-1264 http://www.megazyme.com
MDS Sciex 71 Four Valley Drive Concord, Ontario Canada L4K 4V8 (905) 660-9005 FAX: (905) 660-2600 http://www.sciex.com
Melles Griot 4601 Nautilus Court South Boulder, CO 80301 (800) 326-4363 FAX: (303) 581-0960 (303) 581-0337 http://www.mellesgriot.com
Mead Johnson See Bristol-Meyers Squibb Med Associates P.O. Box 319 St. Albans, VT 05478 (802) 527-2343 FAX: (802) 527-5095 http://www.med-associates.com Medecell 239 Liverpool Road London N1 1LX, UK (44) 20-7607-2295 FAX: (44) 20-7700-4156 http://www.medicell.co.uk Media Cybernetics 8484 Georgia Avenue, Suite 200 Silver Spring, MD 20910 (301) 495-3305 FAX: (301) 495-5964 http://www.mediacy.com Mediatech 13884 Park Center Road Herndon, VA 20171 (800) cellgro (703) 471-5955 http://www.cellgro.com Medical Systems See Harvard Apparatus Medifor 647 Washington Street Port Townsend, WA 98368 (800) 366-3710 FAX: (360) 385-4402 (360) 385-0722 http://www.medifor.com MedImmune 35 W. Watkins Mill Road Gaithersburg, MD 20878 (301) 417-0770 FAX: (301) 527-4207 http://www.medimmune.com MedProbe AS P.O. Box 2640 St. Hanshaugen N-0131 Oslo, Norway (47) 222 00137 FAX: (47) 222 00189 http://www.medprobe.com
Menzel-Glaser Postfach 3157 D-38021 Braunschweig, Germany (49) 531 590080 FAX: (49) 531 509799 E. Merck Frankfurterstrasse 250 D-64293 Darmstadt 1, Germany (49) 6151-720 Merck See EM Science Merck & Company Merck National Service Center P.O. Box 4 West Point, PA 19486 (800) NSC-MERCK (215) 652-5000 http://www.merck.com Merck Research Laboratories See Merck & Company Merck Sharpe Human Health Division 300 Franklin Square Drive Somerset, NJ 08873 (800) 637-2579 FAX: (732) 805-3960 (732) 805-0300 Merial Limited 115 Transtech Drive Athens, GA 30601 (800) MERIAL-1 FAX: (706) 548-0608 (706) 548-9292 http://www.merial.com Meridian Instruments P.O. Box 1204 Kent, WA 98035 (253) 854-9914 FAX: (253) 854-9902 http://www.minstrument.com Meta Systems Group 32 Hammond Road Belmont, MA 02178 (617) 489-9950 FAX: (617) 489-9952 Metachem Technologies 3547 Voyager Street, Bldg. 102 Torrance, CA 90503 (310) 793-2300 FAX: (310) 793-2304 http://www.metachem.com
Metallhantering Box 47172 100-74 Stockholm, Sweden (46) 8-726-9696 MethylGene 7220 Frederick-Banting, Suite 200 Montreal, Quebec H4S 2A1, Canada http://www.methylgene.com Metro Scientific 475 Main Street, Suite 2A Farmingdale, NY 11735 (800) 788-6247 FAX: (516) 293-8549 (516) 293-9656 Metrowerks 980 Metric Boulevard Austin, TX 78758 (800) 377-5416 (512) 997-4700 http://www.metrowerks.com Mettler Instruments Mettler-Toledo 1900 Polaris Parkway Columbus, OH 43240 (800) METTLER FAX: (614) 438-4900 http://www.mt.com Miami Serpentarium Labs 34879 Washington Loop Road Punta Gorda, FL 33982 (800) 248-5050 FAX: (813) 639-1811 (813) 639-8888 http://www.miamiserpentarium.com Michrom BioResources 1945 Industrial Drive Auburn, CA 95603 (530) 888-6498 FAX: (530) 888-8295 http://www.michrom.com Mickle Laboratory Engineering Gomshall, Surrey, UK (44) 1483-202178 Micra Scientific A division of Eichrom Industries 8205 S. Cass Ave, Suite 111 Darien, IL 60561 (800) 283-4752 FAX: (630) 963-1928 (630) 963-0320 http://www.micrasci.com MicroBrightField 74 Hegman Avenue Colchester, VT 05446 (802) 655-9360 FAX: (802) 655-5245 http://www.microbrightfield.com
Micro-Metrics P.O. Box 13804 Atlanta, GA 30324 (770) 986-6015 FAX: (770) 986-9510 http://www.micro-metrics.com Micro-Tech Scientific 140 South Wolfe Road Sunnyvale, CA 94086 (408) 730-8324 FAX: (408) 730-3566 http://www.microlc.com Microbix Biosystems 341 Bering Avenue Toronto, Ontario M8Z 3A8 Canada 1-800-794-6694 FAX:416-234-1626 1-416-234-1624 http://www.microbix.com MicroCal 22 Industrial Drive East Northampton, MA 01060 (800) 633-3115 FAX: (413) 586-0149 (413) 586-7720 http://www.microcalorimetry.com Microfluidics 30 Ossipee Road P.O. Box 9101 Newton, MA 02164 (800) 370-5452 FAX: (617) 965-1213 (617) 969-5452 http://www.microfluidicscorp.com Microgon See Spectrum Laboratories Microlase Optical Systems West of Scotland Science Park Kelvin Campus, Maryhill Road Glasgow G20 0SP, UK (44) 141-948-1000 FAX: (44) 141-946-6311 http://www.microlase.co.uk Micron Instruments 4509 Runway Street Simi Valley, CA 93063 (800) 638-3770 FAX: (805) 522-4982 (805) 552-4676 http://www.microninstruments.com Micron Separations See MSI
Micro Essential Laboratory 4224 Avenue H Brooklyn, NY 11210 (718) 338-3618 FAX: (718) 692-4491
Micro Photonics 4949 Liberty Lane, Suite 170 P.O. Box 3129 Allentown, PA 18106 (610) 366-7103 FAX: (610) 366-7105 http://www.microphotonics.com
Micro Filtration Systems 7-3-Chome, Honcho Nihonbashi, Tokyo, Japan (81) 3-270-3141
MicroTech 1420 Conchester Highway Boothwyn, PA 19061 (610) 459-3514
Suppliers
21 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
Midland Certified Reagent Company 3112-A West Cuthbert Avenue Midland, TX 79701 (800) 247-8766 FAX: (800) 359-5789 (915) 694-7950 FAX: (915) 694-2387 http://www.mcrc.com
Mini Mitter P.O. Box 3386 Sunriver, OR 97707 (800) 685-2999 FAX: (541) 593-5604 (541) 593-8639 http://www.minimitter.com
Midwest Biotech 12690 Ford Drive Fishers, IN 46038 (800) 772-2798 FAX: (317) 841-7576
Mirus Corporation 505 S. Rosa Road Suite 104 Madison, WI 53719 (608) 441-2852 FAX: (608) 441-2849 http://www.genetransfer.com
Midwest Scientific 280 Vance Road Valley Park, MO 63088 (800) 227-9997 FAX: (636) 225-9998 (636) 225-9997 http://www.midsci.com Miles See Bayer
Misonix 1938 New Highway Farmingdale, NY 11735 (800) 645-9846 FAX: (516) 694-9412 http://www.misonix.com Mitutoyo (MTI) See Dolla Eastern
Miles Laboratories See Serological Miles Scientific See Nunc Millar Instruments P.O. Box 230227 6001-A Gulf Freeway Houston, TX 77023 (713) 923-9171 FAX: (713) 923-7757 http://www.millarinstruments.com MilliGen/Biosearch See Millipore Millipore 80 Ashbury Road P.O. Box 9125 Bedford, MA 01730 (800) 645-5476 FAX: (781) 533-3110 (781) 533-6000 http://www.millipore.com Miltenyi Biotec 251 Auburn Ravine Road, Suite 208 Auburn, CA 95603 (800) 367-6227 FAX: (530) 888-8925 (530) 888-8871 http://www.miltenyibiotec.com Miltex 6 Ohio Drive Lake Success, NY 11042 (800) 645-8000 FAX: (516) 775-7185 (516) 349-0001 Milton Roy See Spectronic Instruments Mini-Instruments 15 Burnham Business Park Springfield Road Burnham-on-Crouch, Essex CM0 8TE, UK (44) 1621-783282 FAX: (44) 1621-783132 http://www.mini-instruments.co.uk
MJ Research Waltham, MA 02451 (800) PELTIER FAX: (617) 923-8080 (617) 923-8000 http://www.mjr.com Modular Instruments 228 West Gay Street Westchester, PA 19380 (610) 738-1420 FAX: (610) 738-1421 http://www.mi2.com Molecular Biology Insights 8685 US Highway 24 Cascade, CO 80809-1333 (800) 747-4362 FAX: (719) 684-7989 (719) 684-7988 http://www.oligo.net Molecular Biosystems 10030 Barnes Canyon Road San Diego, CA 92121 (858) 452-0681 FAX: (858) 452-6187 http://www.mobi.com Molecular Devices 1312 Crossman Avenue Sunnyvale, CA 94089 (800) 635-5577 FAX: (408) 747-3602 (408) 747-1700 http://www.moldev.com Molecular Designs 1400 Catalina Street San Leandro, CA 94577 (510) 895-1313 FAX: (510) 614-3608 Molecular Dynamics 928 East Arques Avenue Sunnyvale, CA 94086 (800) 333-5703 FAX: (408) 773-1493 (408) 773-1222 http://www.apbiotech.com
Molecular Probes 4849 Pitchford Avenue Eugene, OR 97402 (800) 438-2209 FAX: (800) 438-0228 (541) 465-8300 FAX: (541) 344-6504 http://www.probes.com Molecular Research Center 5645 Montgomery Road Cincinnati, OH 45212 (800) 462-9868 FAX: (513) 841-0080 (513) 841-0900 http://www.mrcgene.com Molecular Simulations 9685 Scranton Road San Diego, CA 92121 (800) 756-4674 FAX: (858) 458-0136 (858) 458-9990 http://www.msi.com Monoject Disposable Syringes & Needles/Syrvet 16200 Walnut Street Waukee, IA 50263 (800) 727-5203 FAX: (515) 987-5553 (515) 987-5554 http://www.syrvet.com Monsanto Chemical 800 North Lindbergh Boulevard St. Louis, MO 63167 (314) 694-1000 FAX: (314) 694-7625 http://www.monsanto.com Moravek Biochemicals 577 Mercury Lane Brea, CA 92821 (800) 447-0100 FAX: (714) 990-1824 (714) 990-2018 http://www.moravek.com Moss P.O. Box 189 Pasadena, MD 21122 (800) 932-6677 FAX: (410) 768-3971 (410) 768-3442 http://www.mosssubstrates.com Motion Analysis 3617 Westwind Boulevard Santa Rosa, CA 95403 (707) 579-6500 FAX: (707) 526-0629 http://www.motionanalysis.com Mott Farmington Industrial Park 84 Spring Lane Farmington, CT 06032 (860) 747-6333 FAX: (860) 747-6739 http://www.mottcorp.com
Multiple Peptide Systems See NeoMPS Murex Diagnostics 3075 Northwoods Circle Norcross, GA 30071 (707) 662-0660 FAX: (770) 447-4989 MWG-Biotech Anzinger Str. 7 D-85560 Ebersberg, Germany (49) 8092-82890 FAX: (49) 8092-21084 http://www.mwg biotech.com Myriad Industries 3454 E Street San Diego, CA 92102 (800) 999-6777 FAX: (619) 232-4819 (619) 232-6700 http://www.myriadindustries.com Nacalai Tesque Nijo Karasuma, Nakagyo-ku Kyoto 604, Japan 81-75-251-1723 FAX:81-75-251-1762 http://www.nacalai.co.jp Nalge Nunc International Subsidiary of Sybron International 75 Panorama Creek Drive P.O. Box 20365 Rochester, NY 14602 (800) 625-4327 FAX: (716) 586-8987 (716) 264-9346 http://www.nalgenunc.com Nanogen 10398 Pacific Center Court San Diego, CA 92121 (858) 410-4600 FAX: (858) 410-4848 http://www.nanogen.com Nanoprobes 95 Horse Block Road Yaphank, NY 11980 (877) 447-6266 FAX: (631) 205-9493 (631) 205-9490 http://www.nanoprobes.com Narishige USA 1710 Hempstead Turnpike East Meadow, NY 11554 (800) 445-7914 FAX: (516) 794-0066 (516) 794-8000 http://www.narishige.co.jp
MSI (Micron Separations) See Osmonics
Nasco-Fort Atkinson P.O. Box 901 901 Janesville Ave. Fort Atkinson, WI 53538-0901 (800) 558-9595 FAX: (920) 563-8296 http://www.enasco.com
Multi Channel Systems Markwiesenstrasse 55 72770 Reutlingen, Germany (49) 7121-503010 FAX: (49) 7121-503011 http://www.multichannelsystem.com
National Bag Company 2233 Old Mill Road Hudson, OH 44236 (800) 247-6000 FAX: (330) 425-9800 (330) 425-2600 http://www.nationalbag.com
Suppliers
22 CPPS Supplement 45
Current Protocols Selected Suppliers of Reagents and Equipment
National Band and Tag Department X 35, Box 72430 Newport, KY 41032 (606) 261-2035 FAX: (800) 261-8247 https://www.nationalband.com National Biosciences See Molecular Biology Insights National Diagnostics 305 Patton Drive Atlanta, GA 30336 (800) 526-3867 FAX: (404) 699-2077 (404) 699-2121 http://www.nationaldiagnostics.com National Disease Research Exchange 1880 John F. Kennedy Blvd., 11th Fl. Philadelphia, PA 19103 (800) 222-6374 http://www.ndri.com National Institute of Standards and Technology 100 Bureau Drive Gaithersburg, MD 20899 (301) 975-NIST FAX: (301) 926-1630 http://www.nist.gov National Instruments 11500 North Mopac Expressway Austin, TX 78759 (512) 794-0100 FAX: (512) 683-8411 http://www.ni.com National Labnet See Labnet International National Scientific Instruments 975 Progress Circle Lawrenceville, GA 300243 (800) 332-3331 FAX: (404) 339-7173 http://www.nationalscientific.com National Scientific Supply 1111 Francisco Bouldvard East San Rafael, CA 94901 (800) 525-1779 FAX: (415) 459-2954 (415) 459-6070 http://www.nat-sci.com Naz-Dar-KC Chicago Nazdar 1087 N. North Branch Street Chicago, IL 60622 (800) 736-7636 FAX: (312) 943-8215 (312) 943-8338 http://www.nazdar.com NB Labs 1918 Avenue A Denison, TX 75021 (903) 465-2694 FAX: (903) 463-5905 http://www.nblabslarry.com NEB See New England Biolabs
NEN Life Science Products 549 Albany Street Boston, MA 02118 (800) 551-2121 FAX: (617) 451-8185 (617) 350-9075 http://www.nen.com NEN Research Products, Dupont (UK) Diagnostics and Biotechnology Systems Wedgewood Way Stevenage, Hertfordshire SG1 4QN, UK 44-1438-734831 44-1438-734000 FAX:44-1438-734836 http://www.dupont.com Neogen 628 Winchester Road Lexington, KY 40505 (800) 477-8201 FAX: (606) 255-5532 (606) 254-1221 http://www.neogen.com NeoMPS (formerly Multiple Peptide Systems) 9395 Cabot Drive San Diego, CA 92126 (800) 338-4965 FAX: (800) 654-5592 www.neomps.com
New England Biolabs (NEB) 32 Tozer Road Beverly, MA 01915 (800) 632-5227 FAX: (800) 632-7440 http://www.neb.com New England Nuclear (NEN) See NEN Life Science Products New MBR Gubelstrasse 48 CH8050 Zurich, Switzerland (41) 1-313-0703 New Objective Inc. 2 Constitution Way Woburn, MA 01801 (781) 933-9560 http://www.newobjective.com/ Newark Electronics 4801 N. Ravenswood Avenue Chicago, IL 60640 (800) 4-NEWARK FAX: (773) 907-5339 (773) 784-5100 http://www.newark.com Newell Rubbermaid 29 E. Stephenson Street Freeport, IL 61032 (815) 235-4171 FAX: (815) 233-8060 http://www.newellco.com
Neosystems 380, 11012 Macleod Trail South Calgary, Alberta T2J 6A5 Canada (403) 225-9022 FAX: (403) 225-9025 http://www.neosystems.com
Newport Biosystems 1860 Trainor Street Red Bluff, CA 96080 (530) 529-2448 FAX: (530) 529-2648
Neuralynx 2434 North Pantano Road Tucson, AZ, 85715 (520) 722-6144 FAX: (520) 722-8163 http://www.neuralynx.com
Newport 1791 Deere Avenue Irvine, CA 92606 (800) 222-6440 FAX: (949) 253-1800 (949) 253-1462 http://www.newport.com
Neuro Probe 16008 Industrial Drive Gaithersburg, MD 20877 (301) 417-0014 FAX: (301) 977-5711 http://www.neuroprobe.com Neurocrine Biosciences 10555 Science Center Drive San Diego, CA 92121 (619) 658-7600 FAX: (619) 658-7602 http://www.neurocrine.com
Nexin Research B.V. P.O. Box 16 4740 AA Hoeven, The Netherlands (31) 165-503172 FAX: (31) 165-502291 NIAID See Bio-Tech Research Laboratories
Nevtek HCR03, Box 99 Burnsville, VA 24487 (540) 925-2322 FAX: (540) 925-2323 http://www.nevtek.com
Nichiryo 230 Route 206 Building 2-2C Flanders, NJ 07836 (877) 548-6667 FAX: (973) 927-0099 (973) 927-4001 http://www.nichiryo.com
New Brunswick Scientific 44 Talmadge Road Edison, NJ 08818 (800) 631-5417 FAX: (732) 287-4222 (732) 287-1200 http://www.nbsc.com
Nichols Institute Diagnostics 33051 Calle Aviador San Juan Capistrano, CA 92675 (800) 286-4NID FAX: (949) 240-5273 (949) 728-4610 http://www.nicholsdiag.com
Nichols Scientific Instruments 3334 Brown Station Road Columbia, MO 65202 (573) 474-5522 FAX: (603) 215-7274 http://home.beseen.com technology/nsi technology Nicolet Biomedical Instruments 5225 Verona Road, Building 2 Madison, WI 53711 (800) 356-0007 FAX: (608) 441-2002 (608) 273-5000 http://nicoletbiomedical.com N.I.G.M.S. (National Institute of General Medical Sciences) See Coriell Institute for Medical Research Nikon Science and Technologies Group 1300 Walt Whitman Road Melville, NY 11747 (516) 547-8500 FAX: (516) 547-4045 http://www.nikonusa.com Nippon Gene 1-29, Ton-ya-machi Toyama 930, Japan (81) 764-51-6548 FAX: (81) 764-51-6547 Noldus Information Technology 751 Miller Drive Suite E-5 Leesburg, VA 20175 (800) 355-9541 FAX: (703) 771-0441 (703) 771-0440 http://www.noldus.com Nonlinear Dynamics See NovoDynamics Nordion International See MDS Nordion North American Biologicals (NABI) 16500 NW 15th Avenue Miami, FL 33169 (800) 327-7106 (305) 625-5305 http://www.nabi.com North American Reiss See Reiss Northwestern Bottle 24 Walpole Park South Walpole, MA 02081 (508) 668-8600 FAX: (508) 668-7790 NOVA Biomedical Nova Biomedical 200 Prospect Street Waltham, MA 02454 (800) 822-0911 FAX: (781) 894-5915 http://www.novabiomedical.com Novagen 601 Science Drive Madison, WI 53711 (800) 526-7319 FAX: (608) 238-1388 (608) 238-6110 http://www.novagen.com
Suppliers
23 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
Novartis 59 Route 10 East Hanover, NJ 07936 (800)526-0175 FAX: (973) 781-6356 http://www.novartis.com Novartis Biotechnology 3054 Cornwallis Road Research Triangle Park, NC 27709 (888) 462-7288 FAX: (919) 541-8585 http://www.novartis.com Nova Sina AG Subsidiary of Airflow Lufttechnik GmbH Kleine Heeg 21 52259 Rheinbach, Germany (49) 02226 920-0 FAX: (49) 02226 9205-11 Novex/Invitrogen 1600 Faraday Carlsbad, CA 92008 (800) 955-6288 FAX: (760) 603-7201 http://www.novex.com Novo Nordisk Biochem 77 Perry Chapel Church Road Franklington, NC 27525 (800) 879-6686 FAX: (919) 494-3450 (919) 494-3000 http://www.novo.dk Novo Nordisk BioLabs See Novo Nordisk Biochem Novocastra Labs Balliol Business Park West Benton Lane Newcastle-upon-Tyne Tyne and Wear NE12 8EW, UK (44) 191-215-0567 FAX: (44) 191-215-1152 http://www.novocastra.co.uk NovoDynamics 123 North Ashley Street Suite 210 Ann Arbor, MI 48104 (734) 205-9100 FAX: (734) 205-9101 http://www.novodynamics.com Novus Biologicals P.O. Box 802 Littleton, CO 80160 (888) 506-6887 FAX: (303) 730-1966 http://www.novus-biologicals.com/ main.html NPI Electronic Hauptstrasse 96 D-71732 Tamm, Germany (49) 7141-601534 FAX: (49) 7141-601266 http://www.npielectronic.com NSG Precision Cells 195G Central Avenue Farmingdale, NY 11735 (516) 249-7474 FAX: (516) 249-8575 http://www.nsgpci.com
Nu Chek Prep 109 West Main P.O. Box 295 Elysian, MN 56028 (800) 521-7728 FAX: (507) 267-4790 (507) 267-4689 Nuclepore See Costar Numonics 101 Commerce Drive Montgomeryville, PA 18936 (800) 523-6716 FAX: (215) 361-0167 (215) 362-2766 http://www.interactivewhiteboards.com NYCOMED AS Pharma c/o Accurate Chemical & Scientific 300 Shames Drive Westbury, NY 11590 (800) 645-6524 FAX: (516) 997-4948 (516) 333-2221 http://www.accuratechemical.com Nycomed Amersham Health Care Division 101 Carnegie Center Princeton, NJ 08540 (800) 832-4633 FAX: (800) 807-2382 (609) 514-6000 http://www.nycomed-amersham.com Nyegaard Herserudsvagen 5254 S-122 06 Lidingo, Sweden (46) 8-765-2930 Ohmeda Catheter Products See Datex-Ohmeda
Omega Engineering One Omega Drive P.O. Box 4047 Stamford, CT 06907 (800) 848-4286 FAX: (203) 359-7700 (203) 359-1660 http://www.omega.com Omega Optical 3 Grove Street P.O. Box 573 Brattleboro, VT 05302 (802) 254-2690 FAX: (802) 254-3937 http://www.omegafilters.com Omnetics Connector Corporation 7280 Commerce Circle East Minneapolls, MN 55432 (800) 343-0025 (763) 572-0656 Fax: (763) 572-3925 http://www.omnetics.com/main.htm Omni International 6530 Commerce Court Warrenton, VA 20187 (800) 776-4431 FAX: (540) 347-5352 (540) 347-5331 http://www.omni-inc.com Omnion 2010 Energy Drive P.O. Box 879 East Troy, WI 53120 (262) 642-7200 FAX: (262) 642-7760 http://www.omnion.com Omnitech Electronics See AccuScan Instruments Oncogene Research Products P.O. Box Box 12087 La Jolla, CA 92039-2087 (800) 662-2616 FAX: (800) 766-0999 http://www.apoptosis.com
Optomax 9 Ash Street P.O. Box 840 Hollis, NH 03049 (603) 465-3385 FAX: (603) 465-2291 Opto-Line Associates 265 Ballardvale Street Wilmington, MA 01887 (978) 658-7255 FAX: (978) 658-7299 http://www.optoline.com Orbigen 6827 Nancy Ridge Drive San Diego, CA 92121 (866) 672-4436 (858) 362-2030 (858) 362-2026 http://www.orbigen.com Oread BioSaftey 1501 Wakarusa Drive Lawrence, KS 66047 (800) 447-6501 FAX: (785) 749-1882 (785) 749-0034 http://www.oread.com Organomation Associates 266 River Road West Berlin, MA 01503 (888) 978-7300 FAX: (978)838-2786 (978) 838-7300 http://www.organomation.com Organon 375 Mount Pleasant Avenue West Orange, NJ 07052 (800) 241-8812 FAX: (973) 325-4589 (973) 325-4500 http://www.organon.com
Oncor See Intergen
Organon Teknika (Canada) 30 North Wind Place Scarborough, Ontario M1S 3R5 Canada (416) 754-4344 FAX: (416) 754-4488 http://www.organonteknika.com
Oligos Etc. 9775 S.W. Commerce Circle, C-6 Wilsonville, OR 97070 (800) 888-2358 FAX: (503) 6822D1635 (503) 6822D1814 http://www.oligoetc.com
Online Instruments 130 Conway Drive, Suites A & B Bogart, GA 30622 (800) 852-3504 (706) 353-1972 (706) 353-6547 http://www.olisweb.com
Organon Teknika Cappel 100 Akzo Avenue Durham, NC 27712 (800) 682-2666 FAX: (800) 432-9682 (919) 620-2000 FAX: (919) 620-2107 http://www.organonteknika.com
Olis Instruments 130 Conway Drive Bogart, GA 30622 (706) 353-6547 (800) 852-3504 http://www.olisweb.com
Operon Technologies 1000 Atlantic Avenue Alameda, CA 94501 (800) 688-2248 FAX: (510) 865-5225 (510) 865-8644 http://www.operon.com
Oriel Corporation of America 150 Long Beach Boulevard Stratford, CT 06615 (203) 377-8282 FAX: (203) 378-2457 http://www.oriel.com
Olympus America 2 Corporate Center Drive Melville, NY 11747 (800) 645-8160 FAX: (516) 844-5959 (516) 844-5000 http://www.olympusamerica.com
Optiscan P.O. Box 1066 Mount Waverly MDC, Victoria Australia 3149 61-3-9538 3333 FAX:61-3-9562 7742 http://www.optiscan.com.au
OriGene Technologies 6 Taft Court, Suite 300 Rockville, MD 20850 (888) 267-4436 FAX: (301) 340-9254 (301) 340-3188 http://www.origene.com
Ohwa Tsusbo Hiby Dai Building 1-2-2 Uchi Saiwai-cho Chiyoda-ku Tokyo 100, Japan 03-3591-7348 FAX:03-3501-9001
Oncogene Science See OSI Pharmaceuticals
Suppliers
24 CPPS Supplement 45
Current Protocols Selected Suppliers of Reagents and Equipment
OriginLab One Roundhouse Plaza Northhampton, MA 01060 (800) 969-7720 FAX: (413) 585-0126 http://www.originlab.com
Oxford Biochemical Research P.O. Box 522 Oxford, MI 48371 (800) 692-4633 FAX: (248) 852-4466 http://www.oxfordbiomed.com
Orion Research 500 Cummings Center Beverly, MA 01915 (800) 225-1480 FAX: (978) 232-6015 (978) 232-6000 http://www.orionres.com
Oxford GlycoSystems See Glyco
Ortho Diagnostic Systems Subsidiary of Johnson & Johnson 1001 U.S. Highway 202 P.O. Box 350 Raritan, NJ 08869 (800) 322-6374 FAX: (908) 218-8582 (908) 218-1300 Ortho McNeil Pharmaceutical Welsh & McKean Road Spring House, PA 19477 (800) 682-6532 (215) 628-5000 http://www.orthomcneil.com Oryza 200 Turnpike Road, Unit 5 Chelmsford, MA 01824 (978) 256-8183 FAX: (978) 256-7434 http://www.oryzalabs.com OSI Pharmaceuticals 106 Charles Lindbergh Boulevard Uniondale, NY 11553 (800) 662-2616 FAX: (516) 222-0114 (516) 222-0023 http://www.osip.com Osmonics 135 Flanders Road P.O. Box 1046 Westborough, MA 01581 (800) 444-8212 FAX: (508) 366-5840 (508) 366-8212 http://www.osmolabstore.com Oster Professional Products 150 Cadillac Lane McMinnville, TN 37110 (931) 668-4121 FAX: (931) 668-4125 http://www.sunbeam.com Out Patient Services 1260 Holm Road Petaluma, CA 94954 (800) 648-1666 FAX: (707) 762-7198 (707) 763-1581 OWL Scientific Plastics See OWL Separation Systems OWL Separation Systems 55 Heritage Avenue Portsmouth, NH 03801 (800) 242-5560 FAX: (603) 559-9258 (603) 559-9297 http://www.owlsci.com
Oxford Instruments Old Station Way Eynsham Witney, Oxfordshire OX8 1TL, UK (44) 1865-881437 FAX: (44) 1865-881944 http://www.oxinst.com Oxford Labware See Kendall Oxford Molecular Group Oxford Science Park The Medawar Centre Oxford OX4 4GA, UK (44) 1865-784600 FAX: (44) 1865-784601 http://www.oxmol.co.uk Oxford Molecular Group 2105 South Bascom Avenue, Suite 200 Campbell, CA 95008 (800) 876-9994 FAX: (408) 879-6302 (408) 879-6300 http://www.oxmol.com OXIS International 6040 North Cutter Circle Suite 317 Portland, OR 97217 (800) 547-3686 FAX: (503) 283-4058 (503) 283-3911 http://www.oxis.com Oxoid 800 Proctor Avenue Ogdensburg, NY 13669 (800) 567-8378 FAX: (613) 226-3728 http://www.oxoid.ca Oxoid Wade Road Basingstoke, Hampshire RG24 8PW, UK (44) 1256-841144 FAX: (4) 1256-814626 http://www.oxoid.ca Oxyrase P.O. Box 1345 Mansfield, OH 44901 (419) 589-8800 FAX: (419) 589-9919 http://www.oxyrase.com Ozyme 10 Avenue Amp`ere Montigny de Bretoneux 78180 France (33) 13-46-02-424 FAX: (33) 13-46-09-212 http://www.ozyme.fr
PAA Laboratories 2570 Route 724 P.O. Box 435 Parker Ford, PA 19457 (610) 495-9400 FAX: (610) 495-9410 http://www.paa-labs.com Pacer Scientific 5649 Valley Oak Drive Los Angeles, CA 90068 (323) 462-0636 FAX: (323) 462-1430 http://www.pacersci.com Pacific Bio-Marine Labs P.O. Box 1348 Venice, CA 90294 (310) 677-1056 FAX: (310) 677-1207 Packard Biochip Technologies 39 Manning Road Billerica, MA 01821 (888) 323-9681 FAX: (978) 663-8917 http://www.perkinelmer.com Packard Instrument 800 Research Parkway Meriden, CT 06450 (800) 323-1891 FAX: (203) 639-2172 (203) 238-2351 http://www.packardinst.com Padgett Instrument 1730 Walnut Street Kansas City, MO 64108 (816) 842-1029 Pall Filtron 50 Bearfoot Road Northborough, MA 01532 (800) FILTRON FAX: (508) 393-1874 (508) 393-1800 Pall-Gelman 25 Harbor Park Drive Port Washington, NY 11050 (800) 289-6255 FAX: (516) 484-2651 (516) 484-3600 http://www.pall.com PanVera 545 Science Drive Madison, WI 53711 (800) 791-1400 FAX: (608) 233-3007 (608) 233-9450 http://www.panvera.com Parke-Davis See Warner-Lambert Parr Instrument 211 53rd Street Moline, IL 61265 (800) 872-7720 FAX: (309) 762-9453 (309) 762-7716 http://www.parrinst.com Partec Otto Hahn Strasse 32 D-48161 Munster, Germany (49) 2534-8008-0 FAX: (49) 2535-8008-90
PCR See Archimica Florida PE Biosystems 850 Lincoln Centre Drive Foster City, CA 94404 (800) 345-5224 FAX: (650) 638-5884 (650) 638-5800 http://www.pebio.com Pel-Freez Biologicals 219 N. Arkansas P.O. Box 68 Rogers, AR 72757 (800) 643-3426 FAX: (501) 636-3562 (501) 636-4361 http://www.pelfreez-bio.com Pel-Freez Clinical Systems Subsidiary of Pel-Freez Biologicals 9099 N. Deerbrook Trail Brown Deer, WI 53223 (800) 558-4511 FAX: (414) 357-4518 (414) 357-4500 http://www.pelfreez-bio.com Peninsula Laboratories 601 Taylor Way San Carlos, CA 94070 (800) 650-4442 FAX: (650) 595-4071 (650) 592-5392 http://www.penlabs.com Pentex 24562 Mando Drive Laguna Niguel, CA 92677 (800) 382-4667 FAX: (714) 643-2363 http://www.pentex.com PeproTech 5 Crescent Avenue P.O. Box 275 Rocky Hill, NJ 08553 (800) 436-9910 FAX: (609) 497-0321 (609) 497-0253 http://www.peprotech.com Peptide Institute 4-1-2 Ina, Minoh-shi Osaka 562-8686, Japan 81-727-29-4121 FAX:81-727-29-4124 http://www.peptide.co.jp Peptide Laboratory 4175 Lakeside Drive Richmond, CA 94806 (800) 858-7322 FAX: (510) 262-9127 (510) 262-0800 http://www.peptidelab.com Peptides International 11621 Electron Drive Louisville, KY 40299 (800) 777-4779 FAX: (502) 267-1329 (502) 266-8787 http://www.pepnet.com
Suppliers
25 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
Perbio Science UK Ltd. Unit 9 Atley Way, North Nelson Industrial Estate Cramlington, Northumberland NE23 1WA Tel: 0800.25.21.85 http://www.perbio.com
Pharmacia LKB Nuclear See Wallac
Perceptive Science Instruments 2525 South Shore Blvd., Suite 100 League City, TX 77573 (281) 334-3027 FAX: (281) 538-2222 http://www.persci.com
Pharmed (Norton) Norton Performance Plastics See Saint-Gobain Performance Plastics
Perimed 4873 Princeton Drive North Royalton, OH 44133 (440) 877-0537 FAX: (440) 877-0534 http://www.perimed.se Perkin-Elmer 761 Main Avenue Norwalk, CT 06859 (800) 762-4002 FAX: (203) 762-6000 (203) 762-1000 http://www.perkin-elmer.com See also PE Biosystems PerSeptive Bioresearch Products See PerSeptive BioSystems PerSeptive BioSystems 500 Old Connecticut Path Framingham, MA 01701 (800) 899-5858 FAX: (508) 383-7885 (508) 383-7700 http://www.pbio.com PerSeptive Diagnostic See PE Biosystems (800) 343-1346 Pettersson Elektronik AB Tallbacksvagen 51 S-756 45 Uppsala, Sweden (46) 1830-3880 FAX: (46) 1830-3840 http://www.bahnhof.se/∼pettersson Pfanstiehl Laboratories, Inc. 1219 Glen Rock Avenue Waukegan, IL 60085 (800) 383-0126 FAX: (847) 623-9173 http://www.pfanstiehl.com PGC Scientifics 7311 Governors Way Frederick, MD 21704 (800) 424-3300 FAX: (800) 662-1112 (301) 620-7777 FAX: (301) 620-7497 http://www.pgcscientifics.com Pharmacia Biotech See Amersham Pharmacia Biotech Pharmacia Diagnostics See Wallac Pharmacia LKB Biotech See Amersham Pharmacia Biotech Pharmacia LKB Biotechnology See Amersham Pharmacia Biotech
Pharmaderm Veterinary Products 60 Baylis Road Melville, NY 11747 (800) 432-6673 http://www.pharmaderm.com
PharMingen See BD PharMingen Phenomex 2320 W. 205th Street Torrance, CA 90501 (310) 212-0555 FAX: (310) 328-7768 http://www.phenomex.com PHLS Centre for Applied Microbiology and Research See European Collection of Animal Cell Cultures (ECACC) Phoenix Flow Systems 11575 Sorrento Valley Road, Suite 208 San Diego, CA 92121 (800) 886-3569 FAX: (619) 259-5268 (619) 453-5095 http://www.phnxflow.com Phoenix Pharmaceutical 4261 Easton Road, P.O. Box 6457 St. Joseph, MO 64506 (800) 759-3644 FAX: (816) 364-4969 (816) 364-5777 http://www.phoenixpharmaceutical.com Photometrics See Roper Scientific Photon Technology International 1 Deerpark Drive, Suite F Monmouth Junction, NJ 08852 (732) 329-0910 FAX: (732) 329-9069 http://www.pti-nj.com Physik Instrumente Polytec PI 23 Midstate Drive, Suite 212 Auburn, MA 01501 (508) 832-3456 FAX: (508) 832-0506 http://www.polytecpi.com Physitemp Instruments 154 Huron Avenue Clifton, NJ 07013 (800) 452-8510 FAX: (973) 779-5954 (973) 779-5577 http://www.physitemp.com Pico Technology The Mill House, Cambridge Street St. Neots, Cambridgeshire PE19 1QB, UK (44) 1480-396-395 FAX: (44) 1480-396-296 http://www.picotech.com
Pierce Chemical P.O. Box 117 3747 Meridian Road Rockford, IL 61105 (800) 874-3723 FAX: (800) 842-5007 FAX: (815) 968-7316 http://www.piercenet.com
Polyfiltronics 136 Weymouth St. Rockland, MA 02370 (800) 434-7659 FAX: (781) 878-0822 (781) 878-1133 http://www.polyfiltronics.com
Pierce & Warriner 44, Upper Northgate Street Chester, Cheshire CH1 4EF, UK (44) 1244 382 525 FAX: (44) 1244 373 212 http://www.piercenet.com
Polylabo Paul Block Parc Tertiare de la Meinau 10, rue de la Durance B.P. 36 67023 Strasbourg Cedex 1 Strasbourg, France 33-3-8865-8020 FAX:33-3-8865-8039
Pilling Weck Surgical 420 Delaware Drive Fort Washington, PA 19034 (800) 523-2579 FAX: (800) 332-2308 www.pilling-weck.com
PolyLC 9151 Rumsey Road, Suite 180 Columbia, MD 21045 (410) 992-5400 FAX: (410) 730-8340
PixelVision A division of Cybex Computer Products 14964 NW Greenbrier Parkway Beaverton, OR 97006 (503) 629-3210 FAX: (503) 629-3211 http://www.pixelvision.com P.J. Noyes P.O. Box 381 89 Bridge Street Lancaster, NH 03584 (800) 522-2469 FAX: (603) 788-3873 (603) 788-4952 http://www.pjnoyes.com Plas-Labs 917 E. Chilson Street Lansing, MI 48906 (800) 866-7527 FAX: (517) 372-2857 (517) 372-7177 http://www.plas-labs.com Plastics One 6591 Merriman Road, Southwest P.O. Box 12004 Roanoke, VA 24018 (540) 772-7950 FAX: (540) 989-7519 http://www.plastics1.com Platt Electric Supply 2757 6th Avenue South Seattle, WA 98134 (206) 624-4083 FAX: (206) 343-6342 http://www.platt.com Plexon 6500 Greenville Avenue Suite 730 Dallas, TX 75206 (214) 369-4957 FAX: (214) 369-1775 http://www.plexoninc.com Polaroid 784 Memorial Drive Cambridge, MA 01239 (800) 225-1618 FAX: (800) 832-9003 (781) 386-2000 http://www.polaroid.com
Polymer Laboratories Amherst Research Park 160 Old Farm Road Amherst, MA 01002 (800) 767-3963 FAX: (413) 253-2476 http://www.polymerlabs.com Polymicro Technologies 18019 North 25th Avenue Phoenix, AZ 85023 (602) 375-4100 FAX: (602) 375-4110 http://www.polymicro.com Polyphenols AS Hanabryggene Technology Centre Hanaveien 4-6 4327 Sandnes, Norway (47) 51-62-0990 FAX: (47) 51-62-51-82 http://www.polyphenols.com Polysciences 400 Valley Road Warrington, PA 18976 (800) 523-2575 FAX: (800) 343-3291 http://www.polysciences.com Polyscientific 70 Cleveland Avenue Bayshore, NY 11706 (516) 586-0400 FAX: (516) 254-0618 Polytech Products 285 Washington Street Somerville, MA 02143 (617) 666-5064 FAX: (617) 625-0975 Polytron 8585 Grovemont Circle Gaithersburg, MD 20877 (301) 208-6597 FAX: (301) 208-8691 http://www.polytron.com Popper and Sons 300 Denton Avenue P.O. Box 128 New Hyde Park, NY 11040 (888) 717-7677 FAX: (800) 557-6773 (516) 248-0300 FAX: (516) 747-1188 http://www.popperandsons.com
Suppliers
26 CPPS Supplement 45
Current Protocols Selected Suppliers of Reagents and Equipment
Porphyrin Products P.O. Box 31 Logan, UT 84323 (435) 753-1901 FAX: (435) 753-6731 http://www.porphyrin.com
Prior Scientific 80 Reservoir Park Drive Rockland, MA 02370 (781) 878-8442 FAX: (781) 878-8736 http://www.prior.com
Portex See SIMS Portex Limited
PRO Scientific P.O. Box 448 Monroe, CT 06468 (203) 452-9431 FAX: (203) 452-9753 http://www.proscientific.com
Powderject Vaccines 585 Science Drive Madison, WI 53711 (608) 231-3150 FAX: (608) 231-6990 http://www.powderject.com Praxair 810 Jorie Boulevard Oak Brook, IL 60521 (800) 621-7100 http://www.praxair.com Precision Dynamics 13880 Del Sur Street San Fernando, CA 91340 (800) 847-0670 FAX: (818) 899-4-45 http://www.pdcorp.com Precision Scientific Laboratory Equipment Division of Jouan 170 Marcel Drive Winchester, VA 22602 (800) 621-8820 FAX: (540) 869-0130 (540) 869-9892 http://www.precisionsci.com Primary Care Diagnostics See Becton Dickinson Primary Care Diagnostics Primate Products 1755 East Bayshore Road, Suite 28A Redwood City, CA 94063 (650) 368-0663 FAX: (650) 368-0665 http://www.primateproducts.com 5 Prime → 3 Prime See 2000 Eppendorf-5 Prime http://www.5prime.com Princeton Applied Research PerkinElmer Instr.: Electrochemistry 801 S. Illinois Oak Ridge, TN 37830 (800) 366-2741 FAX: (423) 425-1334 (423) 481-2442 http://www.eggpar.com
Professional Compounding Centers of America 9901 South Wilcrest Drive Houston, TX 77099 (800) 331-2498 FAX: (281) 933-6227 (281) 933-6948 http://www.pccarx.com Progen Biotechnik Maass-Str. 30 69123 Heidelberg, Germany (49) 6221-8278-0 FAX: (49) 6221-8278-23 http://www.progen.de Prolabo A division of Merck Eurolab 54 rue Roger Salengro 94126 Fontenay Sous Bois Cedex France 33-1-4514-8500 FAX:33-1-4514-8616 http://www.prolabo.fr Proligo 2995 Wilderness Place Boulder, CO 80301 (888) 80-OLIGO FAX: (303) 801-1134 http://www.proligo.com Promega 2800 Woods Hollow Road Madison, WI 53711 (800) 356-9526 FAX: (800) 356-1970 (608) 274-4330 FAX: (608) 277-2516 http://www.promega.com Protein Databases (PDI) 405 Oakwood Road Huntington Station, NY 11746 (800) 777-6834 FAX: (516) 673-4502 (516) 673-3939
Princeton Instruments A division of Roper Scientific 3660 Quakerbridge Road Trenton, NJ 08619 (609) 587-9797 FAX: (609) 587-1970 http://www.prinst.com
Protein Polymer Technologies 10655 Sorrento Valley Road San Diego, CA 92121 (619) 558-6064 FAX: (619) 558-6477 http://www.ppti.com
Princeton Separations P.O. Box 300 Aldephia, NJ 07710 (800) 223-0902 FAX: (732) 431-3768 (732) 431-3338
Protein Solutions 391 G Chipeta Way Salt Lake City, UT 84108 (801) 583-9301 FAX: (801) 583-4463 http://www.proteinsolutions.com
Proxeon Staermosegaardsvej 6 DK-5230 Odense M Denmark (45) 6557-2300 FAX: (45) 6557-2301 [email protected]
Quantum Soft Postfach 6613 CH-8023 Z¨urich, Switzerland FAX:41-1-481-69-51 profi[email protected]
Prozyme 1933 Davis Street, Suite 207 San Leandro, CA 94577 (800) 457-9444 FAX: (510) 638-6919 (510) 638-6900 http://www.prozyme.com
Questcor Pharmaceuticals 26118 Research Road Hayward, CA 94545 (510) 732-5551 FAX: (510) 732-7741 http://www.questcor.com
PSI See Perceptive Science Instruments Pulmetrics Group 82 Beacon Street Chestnut Hill, MA 02167 (617) 353-3833 FAX: (617) 353-6766 Purdue Frederick 100 Connecticut Avenue Norwalk, CT 06850 (800) 633-4741 FAX: (203) 838-1576 (203) 853-0123 http://www.pharma.com Purina Mills LabDiet P. O. Box 66812 St. Louis, MO 63166 (800) 227-8941 FAX: (314) 768-4894 http://www.purina-mills.com Qbiogene 2251 Rutherford Road Carlsbad, CA 92008 (800) 424-6101 FAX: (760) 918-9313 http://www.qbiogene.com Qiagen 28159 Avenue Stanford Valencia, CA 91355 (800) 426-8157 FAX: (800) 718-2056 http://www.qiagen.com Quality Biological 7581 Lindbergh Drive Gaithersburg, MD 20879 (800) 443-9331 FAX: (301) 840-5450 (301) 840-9331 http://www.qualitybiological.com Quantitative Technologies P.O. Box 470 Salem Industrial Park, Bldg. 5 Whitehouse, NJ 08888 (908) 534-4445 FAX:534-1054 http://www.qtionline.com Quantum Appligene Parc d’Innovation Rue Geller de Kayserberg 67402 Illkirch, Cedex, France (33) 3-8867-5425 FAX: (33) 3-8867-1945 http://www.quantum-appligene.com Quantum Biotechnologies See Qbiogene
Quidel 10165 McKellar Court San Diego, CA 92121 (800) 874-1517 FAX: (858) 546-8955 (858) 552-1100 http://www.quidel.com R-Biopharm 7950 Old US 27 South Marshall, MI 49068 (616) 789-3033 FAX: (616) 789-3070 http://www.r-biopharm.com R. C. Electronics 6464 Hollister Avenue Santa Barbara, CA 93117 (805) 685-7770 FAX: (805) 685-5853 http://www.rcelectronics.com R & D Systems 614 McKinley Place NE Minneapolis, MN 55413 (800) 343-7475 FAX: (612) 379-6580 (612) 379-2956 http://www.rndsystems.com R & S Technology 350 Columbia Street Peacedale, RI 02880 (401) 789-5660 FAX: (401) 792-3890 http://www.septech.com RACAL Health and Safety See 3M 7305 Executive Way Frederick, MD 21704 (800) 692-9500 FAX: (301) 695-8200 Radiometer America 811 Sharon Drive Westlake, OH 44145 (800) 736-0600 FAX: (440) 871-2633 (440) 871-8900 http://www.rameusa.com Radiometer Analytical 72 rue d’Alsace 69627 Villeurbanne CEDEX Lyon, France (33) 478-03-38-38 FAX: (33) 478-68-88-12 http://www.radiometer-analytical.com Radiometer A/S The Chemical Reference Laboratory kandevej 21 DK-2700 Brnshj, Denmark 45-3827-3827 FAX:45-3827-2727
Suppliers
27 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
Radionics 22 Terry Avenue Burlington, MA 01803 (781) 272-1233 FAX: (781) 272-2428 http://www.radionics.com
Reichert Ophthalmic Instruments P.O. Box 123 Buffalo, NY 14240 (716) 686-4500 FAX: (716) 686-4545 http://www.reichert.com
Radnoti Glass Technology 227 W. Maple Avenue Monrovia, CA 91016 (800) 428-l4l6 FAX: (626) 303-2998 (626) 357-8827 http://www.radnoti.com
Reiss 1 Polymer Place P.O. Box 60 Blackstone, VA 23824 (800) 356-2829 FAX: (804) 292-1757 (804) 292-1600 http://www.reissmfg.com
Rainin Instrument Rainin Road P.O. Box 4026 Woburn, MA 01888 (800)-4-RAININ FAX: (781) 938-1152 (781) 935-3050 http://www.rainin.com
Remel 12076 Santa Fe Trail Drive P.O. Box 14428 Shawnee Mission, KS 66215 (800) 255-6730 FAX: (800) 621-8251 (913) 888-0939 FAX: (913) 888-5884 http://www.remelinc.com
Rank Brothers 56 High Street Bottisham, Cambridge CB5 9DA UK (44) 1223 811369 FAX: (44) 1223 811441 http://www.rankbrothers.com Rapp Polymere Ernst-Simon Strasse 9 D 72072 T¨ubingen, Germany (49) 7071-763157 FAX: (49) 7071-763158 http://www.rapp-polymere.com Raven Biological Laboratories 8607 Park Drive P.O. Box 27261 Omaha, NE 68127 (800) 728-5702 FAX: (402) 593-0995 (402) 593-0781 http://www.ravenlabs.com Razel Scientific Instruments 100 Research Drive Stamford, CT 06906 (203) 324-9914 FAX: (203) 324-5568 RBI See Research Biochemicals Reagents International See Biotech Source
Reming Bioinstruments 6680 County Route 17 Redfield, NY 13437 (315) 387-3414 FAX: (315) 387-3415 RepliGen 117 Fourth Avenue Needham, MA 02494 (800) 622-2259 FAX: (781) 453-0048 (781) 449-9560 http://www.repligen.com Research Biochemicals 1 Strathmore Road Natick, MA 01760 (800) 736-3690 FAX: (800) 736-2480 (508) 651-8151 FAX: (508) 655-1359 http://www.resbio.com Research Corporation Technologies 101 N. Wilmot Road, Suite 600 Tucson, AZ 85711 (520) 748-4400 FAX: (520) 748-0025 http://www.rctech.com Research Diagnostics Pleasant Hill Road Flanders, NJ 07836 (800) 631-9384 FAX: (973) 584-0210 (973) 584-7093 http://www.researchd.com
Research Instruments Kernick Road Pernryn Cornwall TR10 9DQ, UK (44) 1326-372-753 FAX: (44) 1326-378-783 http://www.research-instruments.com Research Organics 4353 E. 49th Street Cleveland, OH 44125 (800) 321-0570 FAX: (216) 883-1576 (216) 883-8025 http://www.resorg.com Research Plus P.O. Box 324 Bayonne, NJ 07002 (800) 341-2296 FAX: (201) 823-9590 (201) 823-3592 http://www.researchplus.com Research Products International 410 N. Business Center Drive Mount Prospect, IL 60056 (800) 323-9814 FAX: (847) 635-1177 (847) 635-7330 http://www.rpicorp.com Research Triangle Institute P.O. Box 12194 Research Triangle Park, NC 27709 (919) 541-6000 FAX: (919) 541-6515 http://www.rti.org Restek 110 Benner Circle Bellefonte, PA 16823 (800) 356-1688 FAX: (814) 353-1309 (814) 353-1300 http://www.restekcorp.com Rheodyne P.O. Box 1909 Rohnert Park, CA 94927 (707) 588-2000 FAX: (707) 588-2020 http://www.rheodyne.com Rhone Merieux See Merial Limited
Receptor Biology 10000 Virginia Manor Road, Suite 360 Beltsville, MD 20705 (888) 707-4200 FAX: (301) 210-6266 (301) 210-4700 http://www.receptorbiology.com
Research Diets 121 Jersey Avenue New Brunswick, NJ 08901 (877) 486-2486 FAX: (732) 247-2340 (732) 247-2390 http://www.researchdiets.com
Rhone-Poulenc 2 T W Alexander Drive P.O. Box 12014 Research Triangle Park, NC 08512 (919) 549-2000 FAX: (919) 549-2839 http://www.Rhone-Poulenc.com Also see Aventis
Regis Technologies 8210 N. Austin Avenue Morton Grove, IL 60053 (800) 323-8144 FAX: (847) 967-1214 (847) 967-6000 http://www.registech.com
Research Genetics 2130 South Memorial Parkway Huntsville, AL 35801 (800) 533-4363 FAX: (256) 536-9016 (256) 533-4363 http://www.resgen.com
Rhone-Poulenc Rorer 500 Arcola Road Collegeville, PA 19426 (800) 727-6737 FAX: (610) 454-8940 (610) 454-8975 http://www.rp-rorer.com
Rhone-Poulenc Rorer Centre de Recherche de Vitry-Alfortville 13 Quai Jules Guesde, BP14 94403 Vitry Sur Seine, Cedex, France (33) 145-73-85-11 FAX: (33) 145-73-81-29 http://www.rp-rorer.com Ribi ImmunoChem Research 563 Old Corvallis Road Hamilton, MT 59840 (800) 548-7424 FAX: (406) 363-6129 (406) 363-3131 http://www.ribi.com RiboGene See Questcor Pharmaceuticals Ricca Chemical 448 West Fork Drive Arlington, TX 76012 (888) GO-RICCA FAX: (800) RICCA-93 (817) 461-5601 http://www.riccachemical.com Richard-Allan Scientific 225 Parsons Street Kalamazoo, MI 49007 (800) 522-7270 FAX: (616) 345-3577 (616) 344-2400 http://www.rallansci.com Richelieu Biotechnologies 11 177 Hamon Montral, Quebec H3M 3E4 Canada (802) 863-2567 FAX: (802) 862-2909 http://www.richelieubio.com Richter Enterprises 20 Lake Shore Drive Wayland, MA 01778 (508) 655-7632 FAX: (508) 652-7264 http://www.richter-enterprises.com Riese Enterprises BioSure Division 12301 G Loma Rica Drive Grass Valley, CA 95945 (800) 345-2267 FAX: (916) 273-5097 (916) 273-5095 http://www.biosure.com Robbins Scientific 1250 Elko Drive Sunnyvale, CA 94086 (800) 752-8585 FAX: (408) 734-0300 (408) 734-8500 http://www.robsci.com Roboz Surgical Instruments 9210 Corporate Boulevard, Suite 220 Rockville, MD 20850 (800) 424-2984 FAX: (301) 590-1290 (301) 590-0055
Suppliers
28 CPPS Supplement 45
Current Protocols Selected Suppliers of Reagents and Equipment
Roche Diagnostics 9115 Hague Road P.O. Box 50457 Indianapolis, IN 46256 (800) 262-1640 FAX: (317) 845-7120 (317) 845-2000 http://www.roche.com
Rubbermaid See Newell Rubbermaid
Roche Molecular Systems See Roche Diagnostics
Safe Cells See Bionique Testing Labs
Rocklabs P.O. Box 18-142 Auckland 6, New Zealand (64) 9-634-7696 FAX: (64) 9-634-7696 http://www.rocklabs.com
Sage Instruments 240 Airport Boulevard Freedom, CA 95076 831-761-1000 FAX:831-761-1008 http://www.sageinst.com
Rockland P.O. Box 316 Gilbertsville, PA 19525 (800) 656-ROCK FAX: (610) 367-7825 (610) 369-1008 http://www.rockland-inc.com Rohm Chemische Fabrik Kirschenallee D-64293 Darmstadt, Germany (49) 6151-1801 FAX: (49) 6151-1802 http://www.roehm.com Roper Scientific 3440 East Brittania Drive, Suite 100 Tucson, AZ 85706 (520) 889-9933 FAX: (520) 573-1944 http://www.roperscientific.com Rosetta Inpharmatics 12040 115th Avenue NE Kirkland, WA 98034 (425) 820-8900 FAX: (425) 820-5757 http://www.rii.com ROTH-SOCHIEL 3 rue de la Chapelle Lauterbourg 67630 France (33) 03-88-94-82-42 FAX: (33) 03-88-54-63-93 Rotronic Instrument 160 E. Main Street Huntington, NY 11743 (631) 427-3898 FAX: (631) 427-3902 http://www.rotronic-usa.com Roundy’s 23000 Roundy Drive Pewaukee, WI 53072 (262) 953-7999 FAX: (262) 953-7989 http://www.roundys.com RS Components Birchington Road Weldon Industrial Estate Corby, Northants NN17 9RS, UK (44) 1536 201234 FAX: (44) 1536 405678 http://www.rs-components.com
SA Instrumentation 1437 Tzena Way Encinitas, CA 92024 (858) 453-1776 FAX: (800) 266-1776 http://www.sainst.com
Sage Laboratories 11 Huron Drive Natick, MA 01760 (508) 653-0844 FAX:508-653-5671 http://www.sagelabs.com Saint-Gobain Performance Plastics P.O. Box 3660 Akron, OH 44309 (330) 798-9240 FAX: (330) 798-6968 http://www.nortonplastics.com San Diego Instruments 7758 Arjons Drive San Diego, CA 92126 (858) 530-2600 FAX: (858) 530-2646 http://www.sd-inst.com Sandown Scientific Beards Lodge 25 Oldfield Road Hampden, Middlesex TW12 2AJ, UK (44) 2089 793300 FAX: (44) 2089 793311 http://www.sandownsci.com Sandoz Pharmaceuticals See Novartis Sanofi Recherche Centre de Montpellier 371 Rue du Professor Blayac 34184 Montpellier, Cedex 04 France (33) 67-10-67-10 FAX: (33) 67-10-67-67 Sanofi Winthrop Pharmaceuticals 90 Park Avenue New York, NY 10016 (800) 223-5511 FAX: (800) 933-3243 (212) 551-4000 http://www.sanofi-synthelabo.com/us Santa Cruz Biotechnology 2161 Delaware Avenue Santa Cruz, CA 95060 (800) 457-3801 FAX: (831) 457-3801 (831) 457-3800 http://www.scbt.com Sarasep (800) 605-0267 FAX: (408) 432-3231 (408) 432-3230 http://www.transgenomic.com
Sarstedt P.O. Box 468 Newton, NC 28658 (800) 257-5101 FAX: (828) 465-4003 (828) 465-4000 http://www.sarstedt.com
Scientific Instruments 200 Saw Mill River Road Hawthorne, NY 10532 (800) 431-1956 FAX: (914) 769-5473 (914) 769-5700 http://www.scientificinstruments.com
Sartorius 131 Heartsland Boulevard Edgewood, NY 11717 (800) 368-7178 FAX: (516) 254-4253 http://www.sartorius.com
Scientific Solutions 9323 Hamilton Mentor, OH 44060 (440) 357-1400 FAX: (440) 357-1416 http://www.labmaster.com
SAS Institute Pacific Telesis Center One Montgomery Street San Francisco, CA 94104 (415) 421-2227 FAX: (415) 421-1213 http://www.sas.com
Scion 82 Worman’s Mill Court, Suite H Frederick, MD 21701 (301) 695-7870 FAX: (301) 695-0035 http://www.scioncorp.com
Savant/EC Apparatus A ThermoQuest company 100 Colin Drive Holbrook, NY 11741 (800) 634-8886 FAX: (516) 244-0606 (516) 244-2929 http://www.savec.com Savillex 6133 Baker Road Minnetonka, MN 55345 (612) 935-5427 Scanalytics Division of CSP 8550 Lee Highway, Suite 400 Fairfax, VA 22031 (800) 325-3110 FAX: (703) 208-1960 (703) 208-2230 http://www.scanalytics.com Schering Laboratories See Schering-Plough Schering-Plough 1 Giralda Farms Madison, NJ 07940 (800) 222-7579 FAX: (973) 822-7048 (973) 822-7000 http://www.schering-plough.com Schleicher & Schuell 10 Optical Avenue Keene, NH 03431 (800) 245-4024 FAX: (603) 357-3627 (603) 352-3810 http://www.s-und-s.de/englishindex.html Science Technology Centre 1250 Herzberg Laboratories Carleton University 1125 Colonel Bay Drive Ottawa, Ontario K1S 5B6, Canada (613) 520-4442 FAX: (613) 520-4445 http://www.carleton.ca/universities/stc
Scott Specialty Gases 6141 Easton Road P.O. Box 310 Plumsteadville, PA 18949 (800) 21-SCOTT FAX: (215) 766-2476 (215) 766-8861 http://www.scottgas.com Scripps Clinic and Research Foundation Instrumentation and Design Lab 10666 N. Torrey Pines Road La Jolla, CA 92037 (800) 992-9962 FAX: (858) 554-8986 (858) 455-9100 http://www.scrippsclinic.com SDI Sensor Devices 407 Pilot Court, 400A Waukesha, WI 53188 (414) 524-1000 FAX: (414) 524-1009 Sefar America 111 Calumet Street Depew, NY 14043 (716) 683-4050 FAX: (716) 683-4053 http://www.sefaramerica.com Seikagaku America Division of Associates of Cape Cod 704 Main Street Falmouth, MA 02540 (800) 237-4512 FAX: (508) 540-8680 (508) 540-3444 http://www.seikagaku.com Sellas Medizinische Gerate Hagener Str. 393 Gevelsberg-Vogelsang, 58285 Germany (49) 23-326-1225 Senn Chemicals USA 11189 Sorrento Valley Road Suite #4 San Diego, CA 92121 (858) 450-6091 FAX. (858) 450-6096 www.sennchem.com
Suppliers
29 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
Sensor Medics 22705 Savi Ranch Parkway Yorba Linda, CA 92887 (800) 231-2466 FAX: (714) 283-8439 (714) 283-2228 http://www.sensormedics.com
Shandon/Lipshaw 171 Industry Drive Pittsburgh, PA 15275 (800) 245-6212 FAX: (412) 788-1138 (412) 788-1133 http://www.shandon.com
Silicon Genetics 2601 Spring Street Redwood City, CA 94063 (866) SIG SOFT FAX: (650) 365 1735 (650) 367 9600 http://www.sigenetics.com
Sonics & Materials 53 Church Hill Road Newtown, CT 06470 (800) 745-1105 FAX: (203) 270-4610 (203) 270-4600 http://www.sonicsandmaterials.com
Sensor Systems LLC 2800 Anvil Street, North Saint Petersburg, FL 33710 (800) 688-2181 FAX: (727) 347-3881 (727) 347-2181 http://www.vsensors.com
Sharpoint P.O. Box 2212 Taichung, Taiwan Republic of China (886) 4-3206320 FAX: (886) 4-3289879 http://www.sharpoint.com.tw
SIMS Deltec 1265 Grey Fox Road St. Paul, Minnesota 55112 (800) 426-2448 FAX: (615) 628-7459 http://www.deltec.com
Sonosep Biotech See Triton Environmental Consultants
SenSym/Foxboro ICT 1804 McCarthy Boulevard Milpitas, CA 95035 (800) 392-9934 FAX: (408) 954-9458 (408) 954-6700 http://www.sensym.com Separations Group See Vydac Sepracor 111 Locke Drive Marlboro, MA 01752 (877)-SEPRACOR (508) 357-7300 http://www.sepracor.com Sera-Lab See Harlan Sera-Lab Sermeter 925 Seton Court, #7 Wheeling, IL 60090 (847) 537-4747 Serological 195 W. Birch Street Kankakee, IL 60901 (800) 227-9412 FAX: (815) 937-8285 (815) 937-8270 Seromed Biochrom Leonorenstrasse 2-6 D-12247 Berlin, Germany (49) 030-779-9060 Serotec 22 Bankside Station Approach Kidlington, Oxford OX5 1JE, UK (44) 1865-852722 FAX: (44) 1865-373899 In the US: (800) 265-7376 http://www.serotec.co.uk Serva Biochemicals Distributed by Crescent Chemical S.F. Medical Pharmlast See Chase-Walton Elastomers SGE 2007 Kramer Lane Austin, TX 78758 (800) 945-6154 FAX: (512) 836-9159 (512) 837-7190 http://www.sge.com
Shelton Scientific 230 Longhill Crossroads Shelton, CT 06484 (800) 222-2092 FAX: (203) 929-2175 (203) 929-8999 http://www.sheltonscientific.com
SIMS Portex 10 Bowman Drive Keene, NH 03431 (800) 258-5361 FAX: (603) 352-3703 (603) 352-3812 http://www.simsmed.com
Sherwood-Davis & Geck See Kendall
SIMS Portex Limited Hythe, Kent CT21 6JL, UK (44)1303-260551 FAX: (44)1303-266761 http://www.portex.com
Sherwood Medical See Kendall
Siris Laboratories See Biosearch Technologies
Shimadzu Scientific Instruments 7102 Riverwood Drive Columbia, MD 21046 (800) 477-1227 FAX: (410) 381-1222 (410) 381-1227 http://www.ssi.shimadzu.com
Skatron Instruments See Molecular Devices
Sialomed See Amika Siemens Analytical X-Ray Systems See Bruker Analytical X-Ray Systems Sievers Instruments Subsidiary of Ionics 6060 Spine Road Boulder, CO 80301 (800) 255-6964 FAX: (303) 444-6272 (303) 444-2009 http://www.sieversinst.com SIFCO 970 East 46th Street Cleveland, OH 44103 (216) 881-8600 FAX: (216) 432-6281 http://www.sifco.com Sigma-Aldrich 3050 Spruce Street St. Louis, MO 63103 (800) 358-5287 FAX: (800) 962-9591 (800) 325-3101 FAX: (800) 325-5052 http://www.sigma-aldrich.com Silenus/Amrad 34 Wadhurst Drive Boronia, Victoria 3155 Australia (613)9887-3909 FAX: (613)9887-3912 http://www.amrad.com.au
SLM Instruments See Spectronic Instruments SLM-AMINCO Instruments See Spectronic Instruments Small Parts 13980 NW 58th Court P.O. Box 4650 Miami Lakes, FL 33014 (800) 220-4242 FAX: (800) 423-9009 (305) 558-1038 FAX: (305) 558-0509 http://www.smallparts.com Smith & Nephew 11775 Starkey Road P.O. Box 1970 Largo, FL 33779 (800) 876-1261 http://www.smith-nephew.com SmithKline Beecham 1 Franklin Plaza, #1800 Philadelphia, PA 19102 (215) 751-4000 FAX: (215) 751-4992 http://www.sb.com Solid Phase Sciences See Biosearch Technologies SOMA Scientific Instruments 5319 University Drive, PMB #366 Irvine, CA 92612 (949) 854-0220 FAX: (949) 854-0223 http://somascientific.com Somatix Therapy See Cell Genesys
Sorvall See Kendro Laboratory Products Southern Biotechnology Associates P.O. Box 26221 Birmingham, AL 35260 (800) 722-2255 FAX: (205) 945-8768 (205) 945-1774 http://SouthernBiotech.com SPAFAS 190 Route 165 Preston, CT 06365 (800) SPAFAS-1 FAX: (860) 889-1991 (860) 889-1389 http://www.spafas.com Specialty Media Division of Cell & Molecular Technologies 580 Marshall Street Phillipsburg, NJ 08865 (800) 543-6029 FAX: (908) 387-1670 (908) 454-7774 http://www.specialtymedia.com Spectra Physics See Thermo Separation Products Spectramed See BOC Edwards SpectraSource Instruments 31324 Via Colinas, Suite 114 Westlake Village, CA 91362 (818) 707-2655 FAX: (818) 707-9035 http://www.spectrasource.com Spectronic Instruments 820 Linden Avenue Rochester, NY 14625 (800) 654-9955 FAX: (716) 248-4014 (716) 248-4000 http://www.spectronic.com Spectrum Medical Industries See Spectrum Laboratories Spectrum Laboratories 18617 Broadwick Street Rancho Dominguez, CA 90220 (800) 634-3300 FAX: (800) 445-7330 (310) 885-4601 FAX: (310) 885-4666 http://www.spectrumlabs.com Spherotech 1840 Industrial Drive, Suite 270 Libertyville, IL 60048 (800) 368-0822 FAX: (847) 680-8927 (847) 680-8922 http://www.spherotech.com
Suppliers
30 CPPS Supplement 45
Current Protocols Selected Suppliers of Reagents and Equipment
SPSS 233 S. Wacker Drive, 11th floor Chicago, IL 60606 (800) 521-1337 FAX: (800) 841-0064 http://www.spss.com
Stephens Scientific 107 Riverdale Road Riverdale, NJ 07457 (800) 831-8099 FAX: (201) 831-8009 (201) 831-9800
SS White Burs 1145 Towbin Avenue Lakewood, NJ 08701 (732) 905-1100 FAX: (732) 905-0987 http://www.sswhiteburs.com
Steraloids P.O. Box 689 Newport, RI 02840 (401) 848-5422 FAX: (401) 848-5638 http://www.steraloids.com
Stag Instruments 16 Monument Industrial Park Chalgrove, Oxon OX44 7RW, UK (44) 1865-891116 FAX: (44) 1865-890562
Sterling Medical 2091 Springdale Road, Ste. 2 Cherry Hill, NJ 08003 (800) 229-0900 FAX: (800) 229-7854 http://www.sterlingmedical.com
Standard Reference Materials Program National Institute of Standards and Technology Building 202, Room 204 Gaithersburg, MD 20899 (301) 975-6776 FAX: (301) 948-3730
Sterling Winthrop 90 Park Avenue New York, NY 10016 (212) 907-2000 FAX: (212) 907-3626
Sumitomo Chemical Sumitomo Building 5-33, Kitahama 4-chome Chuo-ku, Osaka 541-8550, Japan (81) 6-6220-3891 FAX: (81)-6-6220-3345 http://www.sumitomo-chem.co.jp
Sternberger Monoclonals 10 Burwood Court Lutherville, MD 21093 (410) 821-8505 FAX: (410) 821-8506 http://www.sternbergermonoclonals. com
Sun Box 19217 Orbit Drive Gaithersburg, MD 20879 (800) 548-3968 FAX: (301) 977-2281 (301) 869-5980 http://www.sunboxco.com
Stoelting 502 Highway 67 Kiel, WI 53042 (920) 894-2293 FAX: (920) 894-7029 http://www.stoelting.com
Sunbrokers See Sun International
Starna Cells P.O. Box 1919 Atascandero, CA 93423 (805) 466-8855 FAX: (805) 461-1575 (800) 228-4482 http://www.stamacells.com Starplex Scientific 50 Steinway Etobieoke, Ontario M9W 6Y3 Canada (800) 665-0954 FAX: (416) 674-6067 (416) 674-7474 http://www.starplexscientific.com
Stovall Lifescience 206-G South Westgate Drive Greensboro, NC 27407 (800) 852-0102 FAX: (336) 852-3507 http://www.slscience.com
State Laboratory Institute of Massachusetts 305 South Street Jamaica Plain, MA 02130 (617) 522-3700 FAX: (617) 522-8735 http://www.state.ma.us/dph
Stratagene 11011 N. Torrey Pines Road La Jolla, CA 92037 (800) 424-5444 FAX: (888) 267-4010 (858) 535-5400 http://www.stratagene.com
Stedim Labs 1910 Mark Court, Suite 110 Concord, CA 94520 (800) 914-6644 FAX: (925) 689-6988 (925) 689-6650 http://www.stedim.com
Strategic Applications 530A N. Milwaukee Avenue Libertyville, IL 60048 (847) 680-9385 FAX: (847) 680-9837
Steinel America 9051 Lyndate Avenue Bloomingron, MN 55420 (800) 852 4343 FAX: (952) 888-5132 http://www.stelnelamerica.com Stem Cell Technologies 777 West Broadway, Suite 808 Vancouver, British Columbia V5Z 4J7 Canada (800) 667-0322 FAX: (800) 567-2899 (604) 877-0713 FAX: (604) 877-0704 http://www.stemcell.com
Strem Chemicals 7 Mulliken Way Newburyport, MA 01950 (800) 647-8736 FAX: (800) 517-8736 (978) 462-3191 FAX: (978) 465-3104 http://www.strem.com StressGen Biotechnologies Biochemicals Division 120-4243 Glanford Avenue Victoria, British Columbia V8Z 4B9 Canada (800) 661-4978 FAX: (250) 744-2877 (250) 744-2811 http://www.stressgen.com
Structure Probe/SPI Supplies (Epon-Araldite) P.O. Box 656 West Chester, PA 19381 (800) 242-4774 FAX: (610) 436-5755 http://www.2spi.com Sud-Chemie ¨ Performance Packaging 101 Christine Drive Belen, NM 87002 (800) 989-3374 FAX: (505) 864-9296 http://www.uniteddesiccants.com
Sun International 3700 Highway 421 North Wilmington, NC 28401 (800) LAB-VIAL FAX: (800) 231-7861 http://www.autosamplervial.com Sunox 1111 Franklin Boulevard, Unit 6 Cambridge, ON N1R 8B5, Canada (519) 624-4413 FAX: (519) 624-8378 http://www.sunox.ca Supelco See Sigma-Aldrich SuperArray P.O. Box 34494 Bethesda, MD 20827 (888) 503-3187 FAX: (301) 765-9859 (301) 765-9888 http://www.superarray.com Surface Measurement Systems 3 Warple Mews, Warple Way London W3 ORF, UK (44) 20-8749-4900 FAX: (44) 20-8749-6749 http://www.smsuk.co.uk/index.htm SurgiVet N7 W22025 Johnson Road, Suite A Waukesha, WI 53186 (262) 513-8500 (888) 745-6562 FAX: (262) 513-9069 http://www.surgivet.com
Sutter Instruments 51 Digital Drive Novato, CA 94949 (415) 883-0128 FAX: (415) 883-0572 http://www.sutter.com Swiss Precision Instruments 1555 Mittel Boulevard, Suite F Wooddale, IL 60191 (800) 221-0198 FAX: (800) 842-5164 Synaptosoft 3098 Anderson Place Decatur, GA 30033 (770) 939-4366 FAX:770-939-9478 http://www.synaptosoft.com SynChrom See Micra Scientific Synergy Software 2457 Perkiomen Avenue Reading, PA 19606 (800) 876-8376 FAX: (610) 370-0548 (610) 779-0522 http://www.synergy.com Synteni See Incyte Synthetics Industry Lumite Division 2100A Atlantic Highway Gainesville, GA 30501 (404) 532-9756 FAX: (404) 531-1347 Systat See SPSS Systems Planning and Analysis (SPA) 2000 N. Beauregard Street Suite 400 Alexandria, VA 22311 (703) 931-3500 http://www.spa-inc.net 3M Bioapplications 3M Center Building 270-15-01 St. Paul, MN 55144 (800) 257-7459 FAX: (651) 737-5645 (651) 736-4946 T Cell Diagnostics and T Cell Sciences 38 Sidney Street Cambridge, MA 02139 (617) 621-1400 TAAB Laboratory Equipment 3 Minerva House Calleva Park Aldermaston, Berkshire RG7 8NA, UK (44) 118 9817775 FAX: (44) 118 9817881 Taconic 273 Hover Avenue Germantown, NY 12526 (800) TAC-ONIC FAX: (518) 537-7287 (518) 537-6208 http://www.taconic.com
Suppliers
31 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
Tago See Biosource International TaKaRa Biochemical 719 Alliston Way Berkeley, CA 94710 (800) 544-9899 FAX: (510) 649-8933 (510) 649-9895 http://www.takara.co.jp/english Takara Shuzo Biomedical Group Division Seta 3-4-1 Otsu Shiga 520-21, Japan (81) 75-241-5100 FAX: (81) 77-543-9254 http://www.Takara.co.jp/english Takeda Chemical Products 101 Takeda Drive Wilmington, NC 28401 (800) 825-3328 FAX: (800) 825-0333 (910) 762-8666 FAX: (910) 762-6846 http://takeda-usa.com TAO Biomedical 73 Manassas Court Laurel Springs, NJ 08021 (609) 782-8622 FAX: (609) 782-8622 Tecan US P.O. Box 13953 Research Triangle Park, NC 27709 (800) 33-TECAN FAX: (919) 361-5201 (919) 361-5208 http://www.tecan-us.com Techne University Park Plaza 743 Alexander Road Princeton, NJ 08540 (800) 225-9243 FAX: (609) 987-8177 (609) 452-9275 http://www.techneusa.com Technical Manufacturing 15 Centennial Drive Peabody, MA 01960 (978) 532-6330 FAX: (978) 531-8682 http://www.techmfg.com Technical Products International 5918 Evergreen St. Louis, MO 63134 (800) 729-4451 FAX: (314) 522-6360 (314) 522-8671 http://www.vibratome.com
Ted Pella 4595 Mountain Lakes Boulevard P.O. Box 492477 Redding, CA 96049 (800) 237-3526 FAX: (530) 243-3761 (530) 243-2200 http://www.tedpella.com Tekmar-Dohrmann P.O. Box 429576 Cincinnati, OH 45242 (800) 543-4461 FAX: (800) 841-5262 (513) 247-7000 FAX: (513) 247-7050 Tektronix 142000 S.W. Karl Braun Drive Beaverton, OR 97077 (800) 621-1966 FAX: (503) 627-7995 (503) 627-7999 http://www.tek.com Tel-Test P.O. Box 1421 Friendswood, TX 77546 (800) 631-0600 FAX: (281)482-1070 (281)482-2672 http://www.isotex-diag.com TeleChem International 524 East Weddell Drive, Suite 3 Sunnyvale, CA 94089 (408) 744-1331 FAX: (408) 744-1711 http://www.gst.net/∼telechem Terrachem Mallaustrasse 57 D-68219 Mannheim, Germany 0621-876797-0 FAX:0621-876797-19 http://www.terrachem.de Terumo Medical 2101 Cottontail Lane Somerset, NJ 08873 (800) 283-7866 FAX: (732) 302-3083 (732) 302-4900 http://www.terumomedical.com Tetko 333 South Highland Manor Briarcliff, NY 10510 (800) 289-8385 FAX: (914) 941-1017 (914) 941-7767 http://www.tetko.com
Texas Fluorescence Labs 9503 Capitol View Drive Austin, TX 78747 (512) 280-5223 FAX: (512) 280-4997 http://www.teflabs.com The Nest Group 45 Valley Road Southborough, MA 01772 (800) 347-6378 FAX: (508) 485-5736 (508) 481-6223 http://world.std.com/∼nestgrp ThermoCare P.O. Box 6069 Incline Village, NV 89450 (800) 262-4020 (775) 831-1201 Thermo Electron Corporation 81 Wyman Street Waltham, MA 02454 (781) 622-1000 FAX: (781) 622-1207 Thermo Labsystems 8 East Forge Parkway Franklin, MA 02038 (800) 522-7763 FAX: (508) 520-2229 (508) 520-0009 http://www.finnpipette.com Thermometric Spjutvagen 5A S-175 61 Jarfalla, Sweden (46) 8-564-72-200 Thermoquest IEC Division 300 Second Avenue Needham Heights, MA 02194 (800) 843-1113 FAX: (781) 444-6743 (781) 449-0800 http://www.thermoquest.com Thermo Separation Products Thermoquest 355 River Oaks Parkway San Jose, CA 95134 (800) 538-7067 FAX: (408) 526-9810 (408) 526-1100 http://www.thermoquest.com Thermo Shandon 171 Industry Drive Pittsburgh, PA 15275 (800) 547-7429 FAX: (412) 899-4045 http://www.thermoshandon.com
Technicon See Organon Teknika Cappel
TetraLink 4240 Ridge Lea Road Suite 29 Amherst, NY 14226 (800) 747-5170 FAX: (800) 747-5171 http://www.tetra-link.com
Thermo Spectronic 820 Linden Avenue Rochester, NY 14625 (585) 248-4000 FAX: (585) 248-4200 http://www.thermo.com
Techno-Aide P.O. Box 90763 Nashville, TN 37209 (800) 251-2629 FAX: (800) 554-6275 (615) 350-7030 http://www.techno-aid.com
TEVA Pharmaceuticals USA 1090 Horsham Road P.O. Box 1090 North Wales, PA 19454 (215) 591-3000 FAX: (215) 721-9669 http://www.tevapharmusa.com
Thomas Scientific 99 High Hill Road at I-295 Swedesboro, NJ 08085 (800) 345-2100 FAX: (800) 345-5232 (856) 467-2000 FAX: (856) 467-3087 http://www.wheatonsci.com/html/nt/ Thomas.html
Thomson Instrument 354 Tyler Road Clearbrook, VA 22624 (800) 842-4752 FAX: (540) 667-6878 (800) 541-4792 FAX: (760) 757-9367 http://www.hplc.com Thorn EMI See Electron Tubes Thorlabs 435 Route 206 Newton, NJ 07860 (973) 579-7227 FAX: (973) 383-8406 http://www.thorlabs.com Tiemann See Bernsco Surgical Supply Timberline Instruments 1880 South Flatiron Court, H-2 P.O. Box 20356 Boulder, CO 80308 (800) 777-5996 FAX: (303) 440-8786 (303) 440-8779 http://www.timberlineinstruments.com Tissueinformatics 711 Bingham Street, Suite 202 Pittsburgh, PA 15203 (418) 488-1100 FAX: (418) 488-6172 http://www.tissueinformatics.com Tissue-Tek A Division of Sakura Finetek USA 1750 West 214th Street Torrance, CA 90501 (800) 725-8723 FAX: (310) 972-7888 (310) 972-7800 http://www.sakuraus.com Tocris Cookson 114 Holloway Road, Suite 200 Ballwin, MO 63011 (800) 421-3701 FAX: (800) 483-1993 (636) 207-7651 FAX: (636) 207-7683 http://www.tocris.com Tocris Cookson Northpoint, Fourth Way Avonmouth, Bristol BS11 8TA, UK (44) 117-982-6551 FAX: (44) 117-982-6552 http://www.tocris.com Tomtec See CraMar Technologies TopoGen P.O. Box 20607 Columbus, OH 43220 (800) TOPOGEN FAX: (800) ADD-TOPO (614) 451-5810 FAX: (614) 451-5811 http://www.topogen.com
Suppliers
32 CPPS Supplement 45
Current Protocols Selected Suppliers of Reagents and Equipment
Toray Industries, Japan Toray Building 2-1 Nihonbash-Muromach 2-Chome, Chuo-Ku Tokyo, Japan 103-8666 (03) 3245-5115 FAX: (03) 3245-5555 http://www.toray.co.jp Toray Industries, U.S.A. 600 Third Avenue New York, NY 10016 (212) 697-8150 FAX: (212) 972-4279 http://www.toray.com Toronto Research Chemicals 2 Brisbane Road North York, Ontario M3J 2J8, Canada (416) 665-9696 FAX: (416) 665-4439 http://www.trc-canada.com TosoHaas 156 Keystone Drive Montgomeryville, PA 18036 (800) 366-4875 FAX: (215) 283-5035 (215) 283-5000 http://www.tosohaas.com Towhill 647 Summer Street Boston, MA 02210 (617) 542-6636 FAX: (617) 464-0804 Toxin Technology 7165 Curtiss Avenue Sarasota, FL 34231 (941) 925-2032 FAX: (9413) 925-2130 http://www.toxintechnology.com Toyo Soda See TosoHaas Trace Analytical 3517-A Edison Way Menlo Park, CA 94025 (650) 364-6895 FAX: (650) 364-6897 http://www.traceanalytical.com Transduction Laboratories See BD Transduction Laboratories Transgenomic 2032 Concourse Drive San Jose, CA 95131 (408) 432-3230 FAX: (408) 432-3231 http://www.transgenomic.com
Trevigen 8405 Helgerman Court Gaithersburg, MD 20877 (800) TREVIGEN FAX: (301) 216-2801 (301) 216-2800 http://www.trevigen.com
Ultrasound Advice 23 Aberdeen Road London N52UG, UK (44) 020-7359-1718 FAX: (44) 020-7359-3650 http://www.ultrasoundadvice.co.uk
Trilink Biotechnologies 6310 Nancy Ridge Drive San Diego, CA 92121 (800) 863-6801 FAX: (858) 546-0020 http://www.trilink.biotech.com
UNELKO 14641 N. 74th Street Scottsdale, AZ 85260 (480) 991-7272 FAX: (480)483-7674 http://www.unelko.com
Tripos Associates 1699 South Hanley Road, Suite 303 St. Louis, MO 63144 (800) 323-2960 FAX: (314) 647-9241 (314) 647-1099 http://www.tripos.com Triton Environmental Consultants 120-13511 Commerce Parkway Richmond, British Columbia V6V 2L1 Canada (604) 279-2093 FAX: (604) 279-2047 http://www.triton-env.com Tropix 47 Wiggins Avenue Bedford, MA 01730 (800) 542-2369 FAX: (617) 275-8581 (617) 271-0045 http://www.tropix.com TSI Center for Diagnostic Products See Intergen
Unifab Corp. 5260 Lovers Lane Kalamazoo, MI 49002 (800) 648-9569 FAX: (616) 382-2825 (616) 382-2803 Union Carbide 10235 West Little York Road, Suite 300 Houston, TX 77040 (800) 568-4000 FAX: (713) 849-7021 (713) 849-7000 http://www.unioncarbide.com United Desiccants See S¨ud-Chemie Performance Packaging United States Biochemical See USB United States Biological (US Biological) P.O. Box 261 Swampscott, MA 01907 (800) 520-3011 FAX: (781) 639-1768 http://www.usbio.net
2000 Eppendorf-5 Prime 5603 Arapahoe Avenue Boulder, CO 80303 (800) 533-5703 FAX: (303) 440-0835 (303) 440-3705
Universal Imaging 502 Brandywine Parkway West Chester, PA 19380 (610) 344-9410 FAX: (610) 344-6515 http://www.image1.com
Tyler Research 10328 73rd Avenue Edmonton, Alberta T6E 6N5 Canada (403) 448-1249 FAX: (403) 433-0479
Upchurch Scientific 619 West Oak Street P.O. Box 1529 Oak Harbor, WA 98277 (800) 426-0191 FAX: (800) 359-3460 (360) 679-2528 FAX: (360) 679-3830 http://www.upchurch.com
UBI See Upstate Biotechnology
Transonic Systems 34 Dutch Mill Road Ithaca, NY 14850 (800) 353-3569 FAX: (607) 257-7256 http://www.transonic.com
Ugo Basile Biological Research Apparatus Via G. Borghi 43 21025 Comerio, Varese, Italy (39) 332 744 574 FAX: (39) 332 745 488 http://www.ugobasile.com
Travenol Lab See Baxter Healthcare
UltraPIX See Life Science Resources
Tree Star Software 20 Winding Way San Carlos, CA 94070 800-366-6045 http://www.treestar.com
Ultrasonic Power 239 East Stephenson Street Freeport, IL 61032 (815) 235-6020 FAX: (815) 232-2150 http://www.upcorp.com
Upjohn Pharmacia & Upjohn http://www.pnu.com Upstate Biotechnology (UBI) 1100 Winter Street, Suite 2300 Waltham, MA 02451 (800) 233-3991 FAX: (781) 890-7738 (781) 890-8845 http://www.upstatebiotech.com USA/Scientific 346 SW 57th Avenue P.O. Box 3565 Ocala, FL 34478 (800) LAB-TIPS FAX: (352) 351-2057 (3524) 237-6288 http://www.usascientific.com
USB 26111 Miles Road P.O. Box 22400 Cleveland, OH 44122 (800) 321-9322 FAX: (800) 535-0898 FAX: (216) 464-5075 http://www.usbweb.com USCI Bard Bard Interventional Products 129 Concord Road Billerica, MA 01821 (800) 225-1332 FAX: (978) 262-4805 http://www.bardinterventional.com UVP (Ultraviolet Products) 2066 W. 11th Street Upland, CA 91786 (800) 452-6788 FAX: (909) 946-3597 (909) 946-3197 http://www.uvp.com V & P Scientific 9823 Pacific Heights Boulevard, Suite T San Diego, CA 92121 (800) 455-0644 FAX: (858) 455-0703 (858) 455-0643 http://www.vp-scientific.com Valco Instruments P.O. Box 55603 Houston, TX 77255 (800) FOR-VICI FAX: (713) 688-8106 (713) 688-9345 http://www.vici.com Valpey Fisher 75 South Street Hopkin, MA 01748 (508) 435-6831 FAX: (508) 435-5289 http://www.valpeyfisher.com Value Plastics 3325 Timberline Road Fort Collins, CO 80525 (800) 404-LUER FAX: (970) 223-0953 (970) 223-8306 http://www.valueplastics.com Vangard International P.O. Box 308 3535 Rt. 66, Bldg. #4 Neptune, NJ 07754 (800) 922-0784 FAX: (732) 922-0557 (732) 922-4900 http://www.vangard1.com Varian Analytical Instruments 2700 Mitchell Drive Walnut Creek, CA 94598 (800) 926-3000 FAX: (925) 945-2102 (925) 939-2400 http://www.varianinc.com Varian Associates 3050 Hansen Way Palo Alto, CA 94304 (800) 544-4636 FAX: (650) 424-5358 (650) 493-4000 http://www.varian.com
Suppliers
33 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
Vector Core Laboratory/National Gene Vector Labs University of Michigan 3560 E MSRB II 1150 West Medical Center Drive Ann Arbor, MI 48109 (734) 936-5843 FAX: (734) 764-3596 Vector Laboratories 30 Ingold Road Burlingame, CA 94010 (800) 227-6666 FAX: (650) 697-0339 (650) 697-3600 http://www.vectorlabs.com Vedco 2121 S.E. Bush Road St. Joseph, MO 64504 (888) 708-3326 FAX: (816) 238-1837 (816) 238-8840 http://database.vedco.com Ventana Medical Systems 3865 North Business Center Drive Tucson, AZ 85705 (800) 227-2155 FAX: (520) 887-2558 (520) 887-2155 http://www.ventanamed.com Verity Software House P.O. Box 247 45A Augusta Road Topsham, ME 04086 (207) 729-6767 FAX: (207) 729-5443 http://www.vsh.com Vernitron See Sensor Systems LLC Vertex Pharmaceuticals 130 Waverly Street Cambridge, MA 02139 (617) 577-6000 FAX: (617) 577-6680 http://www.vpharm.com Vetamac Route 7, Box 208 Frankfort, IN 46041 (317) 379-3621 Vet Drug Unit 8 Lakeside Industrial Estate Colnbrook, Slough SL3 0ED, UK Vetus Animal Health See Burns Veterinary Supply Viamed 15 Station Road Cross Hills, Keighley W. Yorkshire BD20 7DT, UK (44) 1-535-634-542 FAX: (44) 1-535-635-582 http://www.viamed.co.uk Vical 9373 Town Center Drive, Suite 100 San Diego, CA 92121 (858) 646-1100 FAX: (858) 646-1150 http://www.vical.com
Victor Medical 2349 North Watney Way, Suite D Fairfield, CA 94533 (800) 888-8908 FAX: (707) 425-6459 (707) 425-0294 Virion Systems 9610 Medical Center Drive, Suite 100 Rockville, MD 20850 (301) 309-1844 FAX: (301) 309-0471 http://www.radix.net/∼virion VirTis Company 815 Route 208 Gardiner, NY 12525 (800) 765-6198 FAX: (914) 255-5338 (914) 255-5000 http://www.virtis.com Visible Genetics 700 Bay Street, Suite 1000 Toronto, Ontario M5G 1Z6, Canada (888) 463-6844 (416) 813-3272 http://www.visgen.com Vitrocom 8 Morris Avenue Mountain Lakes, NJ 07046 (973) 402-1443 FAX: (973) 402-1445 Vivascience GmbH Feodor-Lynen-Straße 21 30625 Hannover (49) 511-524-875-0 FAX: (49) 511-524-875-19 [email protected] VTI 7650 W. 26th Avenue Hialeah, FL 33106 (305) 828-4700 FAX: (305) 828-0299 http://www.vticorp.com VWR Scientific Products 200 Center Square Road Bridgeport, NJ 08014 (800) 932-5000 FAX: (609) 467-5499 (609) 467-2600 http://www.vwrsp.com Vydac 17434 Mojave Street P.O. Box 867 Hesperia, CA 92345 (800) 247-0924 FAX: (760) 244-1984 (760) 244-6107 http://www.vydac.com Vysis 3100 Woodcreek Drive Downers Grove, IL 60515 (800) 553-7042 FAX: (630) 271-7138 (630) 271-7000 http://www.vysis.com W&H Dentalwerk Burmoos ¨ P.O. Box 1 A-5111 B¨urmoos, Austria (43) 6274-6236-0 FAX: (43) 6274-6236-55 http://www.wnhdent.com
Wako BioProducts See Wako Chemicals USA Wako Chemicals USA 1600 Bellwood Road Richmond, VA 23237 (800) 992-9256 FAX: (804) 271-7791 (804) 271-7677 http://www.wakousa.com Wako Pure Chemicals 1-2, Doshomachi 3-chome Chuo-ku, Osaka 540-8605, Japan 81-6-6203-3741 FAX:81-6-6222-1203 http://www.wako-chem.co.jp/egaiyo/ index.htm Wallac See Perkin-Elmer Wallac A Division of Perkin-Elmer 3985 Eastern Road Norton, OH 44203 (800) 321-9632 FAX: (330) 825-8520 (330) 825-4525 http://www.wallac.com Waring Products 283 Main Street New Hartford, CT 06057 (800) 348-7195 FAX: (860) 738-9203 (860) 379-0731 http://www.waringproducts.com Warner Instrument 1141 Dixwell Avenue Hamden, CT 06514 (800) 599-4203 FAX: (203) 776-1278 (203) 776-0664 http://www.warnerinstrument.com Warner-Lambert Parke-Davis 201 Tabor Road Morris Plains, NJ 07950 (973) 540-2000 FAX: (973) 540-3761 http://www.warner-lambert.com Washington University Machine Shop 615 South Taylor St. Louis, MO 63310 (314) 362-6186 FAX: (314) 362-6184 Waters Chromatography 34 Maple Street Milford, MA 01757 (800) 252-HPLC FAX: (508) 478-1990 (508) 478-2000 http://www.waters.com Watlow 12001 Lackland Road St. Louis, MO 63146 (314) 426-7431 FAX: (314) 447-8770 http://www.watlow.com Watson-Marlow 220 Ballardvale Street Wilmington, MA 01887 (978) 658-6168 FAX: (978) 988 0828 http://www.watson-marlow.co.uk
Waukesha Fluid Handling 611 Sugar Creek Road Delavan, WI 53115 (800) 252-5200 FAX: (800) 252-5012 (414) 728-1900 FAX: (414) 728-4608 http://www.waukesha-cb.com WaveMetrics P.O. Box 2088 Lake Oswego, OR 97035 (503) 620-3001 FAX: (503) 620-6754 http://www.wavemetrics.com Weather Measure P.O. Box 41257 Sacramento, CA 95641 (916) 481-7565 Weber Scientific 2732 Kuser Road Hamilton, NJ 08691 (800) FAT-TEST FAX: (609) 584-8388 (609) 584-7677 http://www.weberscientific.com Weck, Edward & Company 1 Weck Drive Research Triangle Park, NC 27709 (919) 544-8000 Wellcome Diagnostics See Burroughs Wellcome Wellington Laboratories 398 Laird Road, Guelph Ontario, N1G 3X7, Canada (800) 578-6985 FAX: (519) 822-2849 http://www.well-labs.com Wesbart Engineering Daux Road Billingshurst, West Sussex RH14 9EZ, UK (44) 1-403-782738 FAX: (44) 1-403-784180 http://www.wesbart.co.uk Whatman 9 Bridewell Place Clifton, NJ 07014 (800) 631-7290 FAX: (973) 773-3991 (973) 773-5800 http://www.whatman.com Wheaton Science Products 1501 North 10th Street Millville, NJ 08332 (800) 225-1437 FAX: (800) 368-3108 (856) 825-1100 FAX: (856) 825-1368 http://www.algroupwheaton.com Whittaker Bioproducts See BioWhittaker Wild Heerbrugg Juerg Dedual Gaebrisstrasse 8 CH 9056 Gais, Switzerland (41) 71-793-2723 FAX: (41) 71-726-5957 http://www.homepage.swissonline.net/ dedual/wild heerbrugg
Suppliers
34 CPPS Supplement 45
Current Protocols Selected Suppliers of Reagents and Equipment
Willy A. Bachofen AG Maschinenfabrik Utengasse 15/17 CH4005 Basel, Switzerland (41) 61-681-5151 FAX: (41) 61-681-5058 http://www.wab.ch Winthrop See Sterling Winthrop Wolfram Research 100 Trade Center Drive Champaign, IL 61820 (800) 965-3726 FAX: (217) 398-0747 (217) 398-0700 http://www.wolfram.com World Health Organization Microbiology and Immunology Support 20 Avenue Appia 1211 Geneva 27, Switzerland (41-22) 791-2602 FAX: (41-22) 791-0746 http://www.who.org World Precision Instruments 175 Sarasota Center Boulevard International Trade Center Sarasota, FL 34240 (941) 371-1003 FAX: (941) 377-5428 http://www.wpiinc.com Worthington Biochemical Halls Mill Road Freehold, NJ 07728 (800) 445-9603 FAX: (800) 368-3108 (732) 462-3838 FAX: (732) 308-4453 http://www.worthington-biochem.com
WPI See World Precision Instruments Wyeth-Ayerst 2 Esterbrook Lane Cherry Hill, NJ 08003 (800) 568-9938 FAX: (858) 424-8747 (858) 424-3700 Wyeth-Ayerst Laboratories P.O. Box 1773 Paoli, PA 19301 (800) 666-7248 FAX: (610) 889-9669 (610) 644-8000 http://www.ahp.com Xenotech 3800 Cambridge Street Kansas City, KS 66103 (913) 588-7930 FAX: (913) 588-7572 http://www.xenotechllc.com Xeragon 19300 Germantown Road Germantown, MD 20874 (240) 686-7860 FAX: (240)686-7861 http://www.xeragon.com Xillix Technologies 300-13775 Commerce Parkway Richmond, British Columbia V6V 2V4 Canada (800) 665-2236 FAX: (604) 278-3356 (604) 278-5000 http://www.xillix.com
Xomed Surgical Products 6743 Southpoint Drive N Jacksonville, FL 32216 (800) 874-5797 FAX: (800) 678-3995 (904) 296-9600 FAX: (904) 296-9666 http://www.xomed.com Yakult Honsha 1-19, Higashi-Shinbashi 1-chome Minato-ku Tokyo 105-8660, Japan 81-3-3574-8960 Yamasa Shoyu 23-8 Nihonbashi Kakigaracho 1-chome, Chuoku Tokyo, 103 Japan (81) 3-479 22 0095 FAX: (81) 3-479 22 3435 Yeast Genetic Stock Center See ATCC Yellow Spring Instruments See YSI YMC YMC Karasuma-Gojo Building 284 Daigo-Cho, Karasuma Nisihiirr Gojo-dori Shimogyo-ku Kyoto, 600-8106, Japan (81) 75-342-4567 FAX: (81) 75-342-4568 http://www.ymc.co.jp
Zeneca/CRB See AstraZeneca (800) 327-0125 FAX: (800) 321-4745 Zivic-Miller Laboratories 178 Toll Gate Road Zelienople, PA 16063 (800) 422-LABS FAX: (724) 452-4506 (800) MBM-RATS FAX: (724) 452-5200 http://zivicmiller.com Zymark Zymark Center Hopkinton, MA 01748 (508) 435-9500 FAX: (508) 435-3439 http://www.zymark.com Zymed Laboratories 458 Carlton Court South San Francisco, CA 94080 (800) 874-4494 FAX: (650) 871-4499 (650) 871-4494 http://www.zymed.com Zymo Research 625 W. Katella Avenue, Suite 30 Orange, CA 92867 (888) 882-9682 FAX: (714) 288-9643 (714) 288-9682 http://www.zymor.com Zynaxis Cell Science See ChiRex Cauldron
YSI 1725-1700 Brannum Lane Yellow Springs, OH 45387 (800) 765-9744 FAX: (937) 767-9353 (937) 767-7241 http://www.ysi.com
Suppliers
35 Current Protocols Selected Suppliers of Reagents and Equipment
CPPS Supplement 45
